Current Protocols in Nucleic Acid Chemistry Contents Preface Foreword Chapter 1 Synthesis of Modified Nucleosides 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19.
Introduction Unit 1.1 Palladium-Mediated C5 Substitution of Pyrimidine Nucleosides Unit 1.2 Enzymatic Synthesis of M1G-Deoxyribose Unit 1.3 Synthesis of N2-Substituted Deoxyguanosine Nucleosides from 2-Fluoro-6-O(Trimethylsilylethyl)-2′-Deoxyinosine Unit 1.4 Unnatural Nucleosides with Unusual Base Pairing Properties Unit 1.5 Development of a Universal Nucleobase and Modified Nucleobases for Expanding the Genetic Code Unit 1.6 Syntheses of Specifically 15N-Labeled Adenosine and Guanosine Unit 1.7 Synthesis of Protected 2′-Deoxy-2′-fluoro-β-d-arabinonucleosides Unit 1.8 Synthesis, Characterization, and Application of Substituted Pyrazolopyrimidine Nucleosides Unit 1.9 Synthesis of 1,5-Anhydrohexitol Building Blocks for Oligonucleotide Synthesis Unit 1.10 Synthesis and Properties of 7-Substituted 7-Deazapurine (Pyrrolo[2,3-d]pyrimidine) 2′Deoxyribonucleosides Unit 1.11 Reduction of Ribonucleosides to 2′-Deoxyribonucleosides Unit 1.12 Synthesis of Fluorinated Nucleosides Unit 1.13 Synthesis of Ribonucleosides by Condensation Using Trimethylsilyl Triflate Unit 1.14 Synthesis of 2′-O-β-d-Ribofuranosylnucleosides Unit 1.15 Preparation of 2′-Deoxy-2′-Methylseleno-Modified Phosphoramidites and RNA Unit 1.16 Palladium-Catalyzed Cross-Coupling Reactions in C6 Modifications of Purine Nucleosides Unit 1.17 Nucleobase-Caged Phosphoramidites for Oligonucleotide Synthesis Unit 1.18 Synthesis of Altritol Nucleoside Phosphoramidites for Oligonucleotide Synthesis
Chapter 2 Protection of Nucleosides for Oligonucleotide Synthesis Introduction Unit 2.1 Nucleobase Protection of Deoxyribo- and Ribonucleosides Unit 2.2 Protection of 2′-Hydroxy Functions of Ribonucleosides Unit 2.3 Protection of 5′-Hydroxy Functions of Nucleosides Unit 2.4 A Base-Labile Protecting Group (Fluorenylmethoxycarbonyl) for the 5′-Hydroxy Function of Nucleosides 6. Unit 2.5 2′-Hydroxyl-Protecting Groups that are Either Photochemically Labile or Sensitive to Fluoride Ions 7. Unit 2.6 Deoxyribo- and Ribonucleoside H-Phosphonates 8. Unit 2.7 Deoxyribonucleoside Phosphoramidites 9. Unit 2.8 Regioselective 2′-Silylation of Purine Ribonucleosides for Phosphoramidite RNA Synthesis 10. Unit 2.9 Preparation of 2′-O-[(Triisopropylsilyl)oxy]methyl-protected Ribonucleosides 11. Unit 2.10 Preparation of 5′-Silyl-2′-Orthoester Ribonucleosides for Use in Oligoribonucleotide Synthesis 12. Unit 2.11 Enzymatic Regioselective Levulinylation of 2′-Deoxyribonucleosides and 2′-OMethylribonucleosides 1. 2. 3. 4. 5.
13. Unit 2.12 Nucleobase Protection with Allyloxycarbonyl 14. Unit 2.13 Universal 2-(4-Nitrophenyl)ethyl and 2-(4-Nitrophenyl)ethoxycarbonyl Protecting Groups
for Nucleosides and Nucleotides
Chapter 3 Synthesis of Unmodified Oligonucleotides 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17.
Introduction Unit 3.1 Solid-Phase Supports for Oligonucleotide Synthesis Unit 3.2 Attachment of Nucleosides to Solid-Phase Supports Unit 3.3 Synthetic Strategies and Parameters Involved in the Synthesis of Oligodeoxyribonucleotides According to the Phosphoramidite Method Unit 3.4 Synthesis of Oligodeoxyribo- and Oligoribonucleotides According to the H-Phosphonate Method Unit 3.5 Strategies for Oligoribonucleotide Synthesis According to the Phosphoramidite Method Unit 3.6 Oligoribonucleotides with 2′-O-(tert-Butyldimethylsilyl) Groups Unit 3.7 Synthesis of Oligoribonucleotides Using the 2-Nitrobenzyloxymethyl Group for 2′Hydroxyl Protection Unit 3.8 Chemical Synthesis of RNA Sequences with 2′-O-[(Triisopropylsilyl)oxy]methyl-protected Ribonucleoside Phosphoramidites Unit 3.9 3-(N-tert-Butylcarboxamido)-1-propyl and 4-Oxopentyl Groups for Phosphate/Thiophosphate Protection in Oligodeoxyribonucleotide Synthesis Unit 3.10 DNA Synthesis Without Base Protection Unit 3.11 The 4-Methylthio-1-Butyl Group for Phosphate/Thiophosphate Protection in Oligodeoxyribonucleotide Synthesis Unit 3.12 Nucleoside Phosphoramidites Containing Cleavable Linkers Unit 3.13 Microwave-Assisted Functionalization of Solid Supports for Rapid Loading of Nucleosides Unit 3.14 Solution-Phase Synthesis of Di- and Trinucleotides Using Polymer-Supported Reagents Unit 3.15 DNA Synthesis Without Base Protection Using the Phosphoramidite Approach Unit 3.16 A Universal and Recyclable Solid Support for Oligonucleotide Synthesis
Chapter 4 Synthesis of Modified Oligonucleotides and Conjugates 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15.
Introduction Unit 4.1 A Brief History, Status, and Perspective of Modified Oligonucleotides for Chemotherapeutic Applications Unit 4.2 Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups Unit 4.3 Direct Attachment of Conjugate Groups to the 5′ Terminus of Oligodeoxyribonucleotides Unit 4.4 Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides Unit 4.5 Attachment of Reporter and Conjugate Groups to the 3′ Termini of Oligonucleotides Unit 4.6 3′-Modified Oligonucleotides and their Conjugates Unit 4.7 Synthesis and Purification of Oligonucleotide N3′ P5′ Phosphoramidates and their Phosphodiester and Phosphorothioate Chimeras Unit 4.8 Incorporation of Halogenoalkyl, 2-Pyridyldithioalkyl, or Isothiocyanate Linkers into Ligands Unit 4.9 Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands Unit 4.10 Conjugation of 5′-Functionalized Oligodeoxyribonucleotides with Properly Functionalized Ligands Unit 4.11 Synthesis and Purification of Peptide Nucleic Acids Unit 4.12 Locked Nucleic Acids: Synthesis and Characterization of LNA-T Diol Unit 4.13 Cellular Delivery of Locked Nucleic Acids (LNAs) Unit 4.14 Solid-Phase Synthesis of Branched Oligonucleotides
16. Unit 4.15 Solid-Phase Synthesis of 2′-Deoxy-2′-fluoro- β-d-Oligoarabinonucleotides (2′F-ANA) and
Their Phosphorothioate Derivatives
17. Unit 4.16 Chemistry of CpG DNA 18. Unit 4.17 Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Phosphorothioate
Linkages
19. Unit 4.18 Synthesis of Oligonucleotide Conjugates via Aqueous Diels-Alder Cycloaddition 20. Unit 4.19 5′-Iodination of Solid-Phase-Linked Oligodeoxyribonucleotides 21. Unit 4.20 Reversible Biotinylation of the 5′-Terminus of Oligodeoxyribonucleotides and its
Application in Affinity Purification
22. Unit 4.21 Uridine 2′-Carbamates: Facile Tools for Oligonucleotide 2′-Functionalization 23. Unit 4.22 Stepwise Solid-Phase Synthesis of Nucleopeptides 24. Unit 4.23 Synthesis of Oligoribonucleotides Containing N6-Alkyladenosine and 2-Methylthio-N6-
Alkyladenosine
25. Unit 4.24 Oligodeoxyribonucleotide Analogs Functionalized with Phosphonoacetate and
Thiophosphonoacetate Diesters
26. Unit 4.25 Base-Modified Oligodeoxyribonucleotides: Using Pyrrolo[2,3-d]pyrimidines to Replace
Purines
27. Unit 4.26 An Aminooxy-Functionalized Non-Nucleosidic Phosphoramidite for the Construction of
Multiantennary Oligonucleotide Glycoconjugates on a Solid Support 28. Unit 4.27 Large-Scale Preparation of Conjugated Oligonucleoside Phosphorothioates by the High-
Efficiency Liquid-Phase (HELP) Method
29. Unit 4.28 Disulfide Conjugation of Peptides to Oligonucleotides and Their Analogs 30. Unit 4.29 Methoxyoxalamido Chemistry in the Synthesis of Tethered Phosphoramidites and
Functionalized Oligonucleotides 31. Unit 4.30 Using Morpholinos to Control Gene Expression 32. Unit 4.31 Solid-Phase Oligonucleotide Labeling with DOTA
Chapter 5 Methods for Cross-Linking Nucleic Acids 1. 2. 3. 4. 5. 6. 7. 8.
Introduction Unit 5.1 Engineering Disulfide Cross-Links in RNA Using Thiol-Disulfide Interchange Chemistry Unit 5.2 Chemical and Enzymatic Methods for Preparing Circular Single-Stranded DNAs Unit 5.3 Engineering Specific Cross-Links in Nucleic Acids Using Glycol Linkers Unit 5.4 Engineering Disulfide Cross-Links in RNA Via Air Oxidation Unit 5.5 Use of Electrophilic Substitution to Form Site-Specific Cross-Links in DNA Unit 5.6 Synthesis of Endcap Dimethoxytrityl Phosphoramidites for Endcapped Oligonucleotides Unit 5.7 Engineering Terminal Disulfide Bonds Into DNA
Chapter 6 Chemical and Enzymatic Probes for Nucleic Acid Structure 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.
Introduction Unit 6.1 Probing RNA Structure with Chemical Reagents and Enzymes Unit 6.2 Probing Nucleic Acid Structure with Shape-Selective Rhodium and Ruthenium Complexes Unit 6.3 Probing RNA Structure by Lead Cleavage Unit 6.4 Probing Nucleic Acid Structure with Nickel- and Cobalt-Based Reagents Unit 6.5 Probing RNA Structures with Hydroxyl Radicals Unit 6.6 Chemical Reagents for Investigating the Major Groove of DNA Unit 6.7 Probing DNA Structure with Hydroxyl Radicals Unit 6.8 Probing RNA Structure and Metal-Binding Sites Using Terbium(III) Footprinting Unit 6.9 Probing RNA Structure and Function by Nucleotide Analog Interference Mapping
Chapter 7 Biophysical Analysis of Nucleic Acids
1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.
Introduction Unit 7.1 Biophysical Analysis of Nucleic Acids Unit 7.2 NMR Determination of Oligonucleotide Structure Unit 7.3 Optical Methods Unit 7.4 Calorimetry of Nucleic Acids Unit 7.5 Molecular Modeling of Nucleic Acid Structure Unit 7.6 Methods to Crystallize RNA Unit 7.7 Recent Advances in RNA Structure Determination by NMR Unit 7.8 Molecular Modeling of Nucleic Acid Structure: Energy and Sampling Unit 7.9 Molecular Modeling of Nucleic Acid Structure: Electrostatics and Solvation Unit 7.10 Molecular Modeling of Nucleic Acid Structure: Setup and Analysis Unit 7.11 Characterization of DNA Structures by Circular Dichroism Unit 7.12 Biophysical Analysis of Triple-Helix Formation
Chapter 8 Nucleic Acid Binding Molecules 1. 2. 3. 4. 5. 6.
Introduction Unit 8.1 Determination of Binding Mode: Intercalation Unit 8.2 Determination of Binding Thermodynamics Unit 8.3 A Competition Dialysis Assay for the Study of Structure-Selective Ligand Binding to Nucleic Acids Unit 8.4 Chemistry of Minor Groove Binder–Oligonucleotide Conjugates Unit 8.5 A Fluorescent Intercalator Displacement Assay for Establishing DNA Binding Selectivity and Affinity
Chapter 9 Combinatorial Methods in Nucleic Acid Chemistry 1. 2. 3. 4. 5. 6. 7.
Introduction Unit 9.1 Theoretical Principles of In Vitro Selection Using Combinatorial Nucleic Acid Libraries Unit 9.2 Design, Synthesis, and Amplification of DNA Pools for In Vitro Selection Unit 9.3 In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization Unit 9.4 Selection for Catalytic Function with Nucleic Acids Unit 9.5 In Vitro Selection of RNA Aptamers to a Small Molecule Target Unit 9.6 In Vitro Selection Using Modified or Unnatural Nucleotides
Chapter 10 Purification and Analysis of Synthetic Nucleic Acids and Components Introduction Unit 10.1 Analysis of Oligonucleotides by Matrix-Assisted Laser Desorption/Ionization Time-ofFlight Mass Spectrometry 3. Unit 10.2 Analysis of Oligonucleotides by Electrospray Ionization Mass Spectrometry 4. Unit 10.3 Overview of Purification and Analysis of Synthetic Nucleic Acids 5. Unit 10.4 Polyacrylamide Gel Electrophoresis (PAGE) of Synthetic Nucleic Acids 6. Unit 10.5 Analysis and Purification of Synthetic Nucleic Acids Using HPLC 7. Unit 10.6 Base Composition Analysis of Nucleosides Using HPLC 8. Unit 10.7 Cartridge Methods for Oligonucleotide Purification 9. Unit 10.8 Analysis of Oxidized DNA Fragments by Gel Electrophoresis 10. Unit 10.9 Capillary Electrophoresis of DNA 11. Unit 10.10 Sequencing Oligonucleotides by Enrichment of Coupling Failures Using Matrix-Assisted Laser Desorption/Ionization Time-of-Flight Mass Spectrometry 12. Unit 10.11 Mass Determination of Phosphoramidites 1. 2.
Chapter 11 RNA Folding Pathways
Introduction Unit 11.1 RNA Folding Pathways Unit 11.2 RNA Secondary Structure Prediction Unit 11.3 Thermal Methods for the Analysis of RNA Folding Pathways Unit 11.4 Probing RNA Folding Pathways by RNA Fingerprinting Unit 11.5 Characterization of Tertiary Folding of RNA by Circular Dichroism and Urea Unit 11.6 Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays Unit 11.7 Rapid Magnesium Chelation as a Method to Study Real-Time Tertiary Unfolding of RNA Unit 11.8 Use of Fluorescence Spectroscopy to Elucidate RNA Folding Pathways Unit 11.9 Use of Chemical Modification To Elucidate RNA Folding Pathways Unit 11.10 Probing RNA Structural Dynamics and Function by Fluorescence Resonance Energy Transfer (FRET) 12. Unit 11.11 Site-Specific Fluorescent Labeling of Large RNAs with Pyrene 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.
Chapter 12 Nucleic Acid-Based Microarrays and Nanostructures 1. 2. 3. 4. 5. 6. 7. 8.
Introduction Unit 12.1 Key Experimental Approaches in DNA Nanotechnology Unit 12.2 Preparation of Gold Nanoparticle–DNA Conjugates Unit 12.3 Synthesis of 5′-O-Phosphoramidites with a Photolabile 3′-O-Protecting Group Unit 12.4 Derivatization of Glass and Polypropylene Surfaces Unit 12.5 DNA Microarray Preparation by Light-Controlled In Situ Synthesis Unit 12.6 Preparation of α-Oxo Semicarbazone Oligonucleotide Microarrays Unit 12.7 Synthesis of Covalent Oligonucleotide-Streptavidin Conjugates and Their Application in DNA-Directed Immobilization (DDI) of Proteins
Chapter 13 Nucleoside Phosphorylation and Related Modifications 1. 2. 3. 4. 5. 6. 7.
Introduction Unit 13.1 Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates Unit 13.2 Chemoenzymatic Preparation of Nucleoside Triphosphates Unit 13.3 Synthesis and Polymerase Incorporation of 5′-Amino-2′,5′-Dideoxy-5′-N-Triphosphate Nucleotides Unit 13.4 Nucleoside-5′-Phosphoimidazolides: Reagents for Facile Synthesis of Dinucleoside Pyrophosphates Unit 13.5 Synthesis of Methylenebis(phosphonate) Analogs of Dinucleotide Pyrophosphates Unit 13.6 Chemical Phosphorylation of Deoxyribonucleosides and Thermolytic DNA Oligonucleotides
Chapter 14 Biologically Active Nucleosides 1. 2. 3. 4. 5. 6.
Introduction Unit 14.1 Synthesis of Acyclic Analogs of Adenosine Unit 14.2 Synthesis of Acyclic Nucleoside Phosphonates Unit 14.3 Synthesis of β-l-2′-Deoxythymidine (l-dT) Unit 14.4 Synthesis of Carbovir and Abacavir from a Carbocyclic Precursor Unit 14.5 Synthesis of 2′- and 3′-C-Methylribonucleosides
Chapter 15 Nucleoside Prodrugs and Delivery Strategies 1.
Introduction
2. 3. 4.
Unit 15.1 Synthesis of Amino Acid Phosphoramidate Monoesters via H-Phosphonate Intermediates Unit 15.2 Synthesis of Cidofovir and (S)-HPMPA Ether Lipid Prodrugs Unit 15.3 Chemistry of bisSATE Mononucleotide Prodrugs
Appendix 1 Standard Nomenclature, Data, and Abbreviations 1. 2. 3. 4. 5.
1A Selected Abbreviations Used in This Manual 1B Characteristics of Nucleic Acids 1C IUPAC-IUB Joint Commission on Biochemical Nomenclature Abbreviations and Symbols for the Description of Conformations of Polynucleotide Chains 1D Nucleoside and Nucleotide Nomenclature 1E A Convenient Stereochemical Notation for P-Chiral Nucleotide Analogs
Appendix 2 Laboratory Stock Solutions and Equipment 1.
2A Common Buffers and Stock Solutions
Appendix 3 Commonly Used Techniques 1. 2. 3. 4. 5.
3A References to Commonly Used Techniques 3B Denaturing Polyacrylamide Gel Electrophoresis 3C Introduction to the Synthesis and Purification of Oligonucleotides 3D Thin-Layer Chromatography 3E Column Chromatography
Appendix 4 Resources 1.
4A Useful Nucleic Acid Chemistry Web Sites
Appendix Suppliers 1.
Selected Suppliers of Reagents and Equipment
PREFACE hemistry has played a pivotal role in the development of molecular biology and biotechnology. The value of synthetic oligonucleotides became apparent when they provided the means for deciphering the genetic code. Modern cloning techniques and nucleic acid sequencing depend on the ability to obtain synthetic primers. In recent years intensive research has driven the technological development of high-throughput methods for nucleic acid analysis. Moreover, modified nucleic acids have been extensively studied and identified as highly specific therapeutic agents. At the same time, research efforts have provided new means to probe the structure of nucleic acids for a better understanding of all aspects of nucleic acid function and interactions in biology. Future applications of nucleic acids are likely to extend into material science, as nucleic acid sequences provide a code for information storage and manipulation as well as the means for self-assembly of complex devices and systems on the nanoscale.
C
The pathways by which the nucleoside building blocks of nucleic acids are biosynthesized and incorporated into nucleic acids are important targets for antiviral and chemotherapies. A large variety of modified nucleosides and nucleotides have been designed as tools for studying biochemical processes involving nucleotide binding proteins (as enzyme substrates or as cofactors), and many more have been evaluated as potential drugs. Detailed protocols for much of the early chemistry can be found in a useful series of books starting with Synthetic Procedures in Nucleic Acid Chemistry, Vols. 1 and 2 (Zorbach and Tipson, 1968) and continuing with four volumes, Nucleic Acid Chemistry: Improved and New Synthetic Procedures, Methods and Techniques, Parts I-IV (Townsend and Tipson, 1978, 1986, 1991). The current methods applied to the synthesis of modified nucleosides and nucleic acids for structure-function studies, potential therapeutic agents, and as tools for molecular biology, have spawned a unique set of chemistries that provide the fundamental basis on which this volume of Current Protocols has evolved. In Current Protocols in Nucleic Acid Chemistry, the practical aspects of innovative methods for the preparation of modified nucleosides and nucleic acids are strongly emphasized. Chapters 1 to 4 describe detailed methodology for the synthesis of both natural and modified oligonucleotides. Chapter 1 focuses on the synthesis of a variety of modified nucleosides with the emphasis on compounds that could be used as components of oligonucleotides. Various methods for N-protection of nucleobases and functionalization of 5 -, 3 -, and/or 2 -hydroxyl groups of deoxyribo- and ribonucleosides are presented in Chapter 2 along with the preparation of their H-phosphonate or phosphoramidite derivatives to enable oligonucleotide synthesis. In Chapter 3, the development of solid supports and strategies for solid-phase synthesis of native DNA or RNA oligonucleotides, through a collection of properly functionalized H-phosphonate or phosphoramidite monomers, is emphasized. Such rapid and efficient methods for automated synthesis of oligonucleotides has led to the preparation of modified oligonucleotides for therapeutic applications. Methods for altering native oligonucleotide properties through nucleobase, carbohydrate, and internucleotidic phosphodiester modifications or through conjugation with ligands or reporter groups are the focus of Chapter 4. Chapters 5, 6, and 11 provide methods for probing DNA and RNA structures, including cross-linking strategies to stabilize secondary structure, chemical and enzymatic probes of structure, and techniques for elucidating RNA folding. Two chapters focus on purification Current Protocols in Nucleic Acid Chemistry Contributed by Serge L. Beaucage, Donald E. Bergstrom, Piet Herdewijn, and Akira Matsuda Current Protocols in Nucleic Acid Chemistry (2005) iii-vii C 2005 by John Wiley & Sons, Inc. Copyright
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and analysis. Chapter 7 covers important biophysical and computational methods of nucleic acid analysis, including X-ray crystallography, NMR, and molecular modeling strategies. Mass spectrometric analysis is covered in Chapter 10 along with common purification techniques including HPLC and electrophoresis. Two chapters cover different aspects of the binding of nucleic acids to other molecules. Chapter 8 focuses on protocols for measuring binding mode and affinity of small molecules to DNA, a topic of importance because of the significant numbers of drugs that function by binding to nucleic acids. Chapter 9 includes methods for generating nucleic acid aptamers. Chapter 12 focuses on nucleic acid chemistry as applied to the preparation of oligonucleotide arrays and introduces topics related to the development of nucleic acid related nanotechnology. In Chapter 13, protocols are provided for the synthesis of various phosphorylated derivatives of nucleosides. A newly added Chapter 14 provides protocols for synthesis of modified nucleosides that play an important role in the fight against infectious diseases and cancers.
HOW TO USE THIS MANUAL Format and Organization This publication is available in looseleaf, CD-ROM, Intranet, and online formats. For looseleaf purchasers, binders are provided to accommodate the growth of the manual via the quarterly update service. The looseleaf format of the binder allows easy insertion of new pages, units, and chapters that are added. The index and table of contents are updated with each supplement. Purchasers of the CD-ROM and Intranet versions receive a completely new disc every quarter and should dispose of their outdated discs. The material covered in all formats is identical. Subjects in this manual are organized by chapters, and protocols are contained in units. Units generally describe a method and include one or more protocols with listings of materials, steps and annotations, recipes for unique reagents and solutions, and commentaries on the “hows” and “whys” of the method; there are also “overview” units containing theoretical discussions that lay the foundation for subsequent protocols. Page numbering in the looseleaf version reflects the modular arrangement by unit; for example, page 2.3.5 refers to Chapter 2 (Protection of Nucleosides for Oligonucleotide Synthesis), UNIT 2.3 (Protection of the 5 -Hydroxy Function of Nucleosides), page 5 of that particular unit. Many reagents and procedures are employed repeatedly throughout the manual. Instead of duplicating this information, cross-references among units are used extensively. Crossreferencing helps to ensure that lengthy and complex protocols are not overburdened with steps describing auxiliary procedures needed to prepare raw materials and analyze results. For some widely used techniques (such as gel electrophoresis), readers are referred to APPENDIX 3.
Preface
Introductory and Explanatory Information Because this publication is first and foremost a compilation of laboratory techniques in nucleic acid chemistry, we have not offered extensive instructive material. We have, however, included explanatory information where required to help readers gain an intuitive grasp of the procedures. Some chapters begin with special overview units that describe
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the state of the art of the topic matter and provide a context for the procedures that follow. Chapter and unit introductions describe how the protocols that follow connect to one another, and annotations to the actual protocol steps describe what is happening as a procedure is carried out. Finally, the Commentary that closes each protocol unit describes background information regarding the historical and theoretical development of the method, as well as alternative approaches, critical parameters, troubleshooting guidelines, anticipated results, and time considerations. All units contain cited references and many indicate key references to inform users of particularly useful background reading, original descriptions, or applications of a technique.
Protocols Many units in the manual contain groups of protocols, each presented with a series of steps. The basic protocol is presented first in each unit and is generally the recommended or most universally applicable approach. Alternate protocols are given where different equipment or reagents can be employed to achieve similar ends, where the starting material requires a variation in approach, or where requirements for the end product differ from those in the basic protocol. Support protocols describe additional steps that are required to perform the basic or alternate protocols; these steps are separated from the core protocol because they might be applicable to other uses in the manual, or because they are performed in a time frame separate from the basic protocol steps. Reagents and Solutions Reagents required for a protocol are itemized in the materials list before the procedure begins. Many are common stock solutions, others are commonly used buffers or media, whereas others are solutions unique to a particular protocol. Recipes for the latter solutions are supplied in each unit under the heading Reagents and Solutions. It is important to note that the names of some of these special solutions might be similar from unit to unit (e.g., SDS sample buffer) while the recipes differ; thus, it is essential to ensure that reagents are prepared from the proper recipes. On the other hand, recipes for commonly used stock solutions and buffers are listed once in APPENDIX 2A. These universal recipes are cross-referenced parenthetically in the materials lists rather than repeated with every usage. Commercial Suppliers In some instances throughout the manual, we have recommended commercial suppliers of chemicals, biological materials, or equipment. This has been avoided wherever possible, because preference for a specific brand is subjective and is generally not based on extensive comparison testing. Our guidelines for recommending a supplier are that (1) the particular brand has actually been found to be of superior quality, or (2) the item is difficult to find in the marketplace. The purity of chemical reagents frequently varies with supplier. Generally reagent grade chemicals are preferred. Special care must be paid to procedures that require dry solvents. Different suppliers provide special anhydrous grade solvents which may vary in water content depending on the supplier. Addresses, phone numbers, facsimile numbers, and Web sites of all suppliers mentioned in this manual are provided in the SUPPLIERS APPENDIX.
Safety Considerations Anyone carrying out these protocols will encounter the following hazardous or potentially hazardous materials: (1) radioactive substances, (2) toxic chemicals and carcinogenic or
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teratogenic reagents, (3) pathogenic and infectious biological agents, and (4) recombinant DNA. Most governments regulate the use of these materials; it is essential that they be used in strict accordance with local and national regulations. Cautionary notes are included in many instances throughout the manual, but we emphasize that users must proceed with the prudence and precaution associated with good laboratory practice, and that all materials must be used in strict accordance with local and national regulations.
Reader Response The protocols included in this manual are used routinely by the authors who contribute them, and many are used in our own laboratories. To make them work for you, the authors have annotated critical steps and included critical parameters and troubleshooting guides in the commentaries to most units. However, the successful evolution of this manual depends upon readers’ observations and suggestions. Consequently, a selfmailing reader-response survey is included with each print supplement; we encourage readers to send in their comments. Electronic contact information is available at http://www3.interscience.wiley.com/c p/index.htm. ACKNOWLEDGMENTS This manual is the product of dedicated efforts by many of our scientific colleagues who are acknowledged in each unit, and by the hard work of the Current Protocols editorial staff at John Wiley & Sons, including Joe Ingram, Scott Holmes, Tom Cannon, Susan Lieberman, Allen Ranz, and Joseph White. The publisher’s commitment and continuing support for a nucleic acid chemistry manual have been essential for realizing and continuing this ambitious project. We are extremely grateful for the critical contributions made by Ann Boyle, who played a key role in bringing the initial project to completion, and to Beth Harkins, who continues to keep editors and contributors on track. Finally, we gratefully acknowledge the significant role of past editors Roger Jones and Gary Glick in developing Current Protocols in Nucleic Acid Chemistry during its first half-decade.
LITERATURE CITED Zorbach, W.W. and Tipson, R.S. (eds.) 1968. Synthetic Procedures in Nucleic Acid Chemistry, Vols. 1 and 2. Interscience Publishers, New York. Townsend, L.B. and Tipson, R.S. (eds.) 1978. Nucleic Acid Chemistry: Improved and New Synthetic Procedures, Methods and Techniques, Parts I and II. John Wiley & Sons, New York. Townsend, L.B. and Tipson, R.S. (eds.) 1986. Nucleic Acid Chemistry: Improved and New Synthetic Procedures, Methods and Techniques, Part III. John Wiley & Sons, New York. Townsend, L.B. and Tipson, R.S. (eds.) 1991. Nucleic Acid Chemistry: Improved and New Synthetic Procedures, Methods and Techniques, Part IV. John Wiley & Sons, New York.
RECOMMENDED BACKGROUND READING Blackburn, G.M. and Gait, M.G. 1996. Nucleic Acids in Chemistry Biology, 2nd ed., IRL Press, London General introduction to fundamental aspects of nucleic acid chemistry. Hecht, S.M. 1996. Bioorganic Chemistry: Nucleic Acids. Oxford University Press, New York. Barton, D., Nakanishi, K., and Meth-Cohn, O. 2000. Comprehensive Natural Products Chemistry, Vol. 7: DNA and Aspects of Molecular Biology (E.T. Kool, ed.), Elsevier Science, London. Overviews of many important topics related to nucleic acid chemistry, structure, and biochemistry. De Clercq, E. and Herdewijn, P. 2005. Strategies in the design of antiviral drugs. In Drug Discovery Handbook, pp. 1135-1190. Wiley Intersciences, Hoboken, N.J. Overview of the design and synthesis of nucleoside analog antiviral agents.
Preface
Nelson, D.L. and Cox, M.M. 2004. Lehninger Principles of Biochemistry, 4th ed. W.H. Freeman and Company, New York. Basic biochemistry text that includes fundamentals of nucleic acid biochemistry.
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Neidle, S. 1999. Oxford Handbook of Nucleic Acid Structure. Oxford University Press, Oxford. Neidle, S. 2002. Nucleic Acid Structure and Recognition, Oxford University Press, Oxford. The 1999 Oxford Handbook contains the most extensive and comprehensive coverage on nucleic acid structure. Gesteland, R.F., Cech, T.R., and Atkins, J.F. 1999. The RNA World, 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Reviews on many current topics in RNA biochemistry, chemistry and structure. Hartmann, R.K., Binderelf, A., Sch¨on, A. and Westhof, E. 2005. Handbook of RNA Biochemistry. John Wiley & Sons, Hoboken, N.J. Includes current laboratory techniques and protocols. Zubrick, J.W. 2003. The Organic Chem Lab Survival Manual, A Student’s Guide to Techniques, 6th ed. John Wiley & Sons, Hoboken, N.J. Laboratory manual that describes very basic techniques for synthetic chemistry. Silverstein, R.M., Webster, F.X., and Kiemle, D. 2004. Spectrometric Identification of Organic Compounds, 7th ed. John Wiley & Sons, Hoboken, N.J. Explores the identification of organic compounds by mass spectrometry, infrared spectrometry, and nuclear magnetic resonance spectrometry. Lee, T.A. 1998. A Beginner’s Guide to Mass Spectral Interpretation. John Wiley & Sons, New York. Identification of organic compound by mass spectrometry, emphasizing recognition of typical fragmentation patterns for different types of organic compounds. Macomber, R.S. 1998. A Complete Introduction to Modern NMR Spectroscopy. John Wiley & Sons, New York. Clear, accessible coverage of modern NMR spectroscopy methods.
Serge L. Beaucage, Donald E. Bergstrom, Piet Herdewijn, and Akira Matsuda
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Foreword Nucleic acid chemistry has developed at an accelerating pace in recent years. Today one can rapidly synthesize long DNA and RNA strands of any desired nucleotide sequence on a multigram to microgram scale. “DNA chips” containing thousands of different synthetic oligonucleotides, each at a unique, defined site on a 1 × 1–cm glass plate, can also be made. Elegant methods are available to prepare and incorporate into oligonucleotides a great variety of monomers containing tailored bases, sugars, and bridging units. Powerful analytical techniques, including high performance liquid chromatography, capillary gel electrophoresis, UV and laser spectroscopy, and mass spectrometry, enable rapid separation and characterization of polynucleotides. Advances in NMR spectroscopy and X-ray crystallography have broadened the horizons for structural elucidation, and sophisticated computational programs can now generate useful theoretical models to guide research. These and related developments provide current researchers with unprecedented problem-solving power and opportunities for discovery and invention. Yet the very breadth of the knowledge and the pace of innovation pose significant challenges for serious investigators seeking to capitalize on these advances. This manual, Current Protocols in Nucleic Acid Chemistry, brings together in one volume state-of-the-art protocols for the synthesis, isolation, characterization, modification, and design of a wide variety of compounds in the nucleic acid family. It will prove very useful for experts and practitioners working in the nucleic acid field. An attractive additional feature is a series of quarterly supplements that will describe new and improved methodologies as they emerge. These will be incorporated into the existing manual. Oligonucleotides and related derivatives are unique recognition molecules. They bind reversibly with high selectivity to polymers containing complementary nucleotide sequences, and key rules governing formation of the complexes are well established. Since very large numbers of different oligomers can be generated from different combinations of the four main building blocks, oligonucleotide chemistry offers unique opportunities for rationally designing a wide variety of self-assembling systems exhibiting predesignated properties. Increasingly, scientists in other disciplines are using synthetic oligonucleotides, modified oligonucleotides, or oligonucleotide conjugates as tools in their research. The impact of the chemistry has been greatest in molecular biology, where synthetic oligonucleotides now play an essential role as primers in sequencing and amplifying DNA, as models and probes for elucidating the structure and function of DNA and RNA, and as building units for the synthesis and modification of genes. Applications of oligonucleotide conjugates as nonradioactive probes in diagnostic medicine are expanding rapidly; a variety of modified oligonucleotides show promise as highly selective therapeutic agents; organic and inorganic chemists are finding oligonucleotides useful as tethers in positioning functional groups in complex assemblies; and an active new research area in materials science is directed toward creating novel materials from nanoparticle-oligonucleotide conjugates. Some visionaries even dream of employing oligonucleotides in building molecular-scale machines and computers. For the researchers from other disciplines who wish to utilize nucleic acid chemistry and for those working at the frontiers of several disciplines, this manual will be a special boon. The comprehensive collection of concise, explicit protocols will enable them to capitalize quickly on the opportunities opened by the advances in nucleic acid chemistry.
CHAPTER 1 Synthesis of Modified Nucleosides INTRODUCTION odified nucleosides have a wide variety of uses—often after incorporation into oligonucleotides—such as attachment to reporter groups and studies of basepairing altered by mutagens, by carcinogens, or by design (Beaucage and Iyer, 1993). This chapter will focus on sugar- and base-modified nucleosides. In keeping with Current Protocols format, the goal of this chapter is not to review the subject, but to provide a set of specific and detailed procedures leading to specific compounds. Within this framework, however, the procedures should enable researchers to undertake syntheses of related compounds that have been reported in the chemical literature.
M
The most widely used reaction in nucleoside chemistry is the sugar-base condensation reaction using the Vorbr¨uggen procedure. This reaction is used for synthesis of sugarmodified as well as base-modified nucleosides. It is a very elegant way to introduce heterocyclic bases at the anomeric center of sugars in a stereoselective manner (in case a 2 -O-acyl group is present for neighboring group participation). This reaction is described in UNIT 1.13. In a closely related unit, the synthesis of 2 -O-β-D-ribofuranosyl nucleosides are described in UNIT 1.14. The same principle is used here for the Lewis acid– catalyzed O-glycosylation reaction, in this case leading to disaccharide nucleosides. This procedure demonstrates the reaction conditions needed for the stereospecific formation of O-glycosidic bonds in nucleosides (with base moieties attached at the anomeric center). Disaccharide nucleosides occur widely in nature, especially as one of the modified nucleosides in tRNA. In some cases, modified nucleosides may themselves serve as key intermediates from which other modified nucleosides of interest can be prepared. For example, the palladiummediated C5 substitution of pyrimidines pioneered by Bergstrom has proved over the years to be a uniquely valuable route to these important compounds (Goodchild, 1990). An understanding of the synthetic procedure detailed in UNIT 1.1 will provide access to a large number of C5-modified pyrimidines. Similarly, UNIT 1.8 describes the synthesis and use of aminoalkyl-modified purine analogs. The palladium-catalyzed cross-coupling reaction for C-C bond formation is also used extensively in the purine field, in efforts to generate bioactive compounds. UNIT 1.16 describes the reaction of 6-chloropurine nucleosides with several organometallic reagents using palladium as catalyst, and these reactions are representative for the field. In addition, the 6-hydroxymethyl congener is used as example for further derivatization reactions, mainly for preparation of fluorinated nucleosides. The enzymatic coupling methods in UNITS 1.2 & 1.6 use enzymatic transglycosylation for synthesis of 2 -deoxyribonucleosides and ribonucleosides, respectively. Although nucleoside 2 -deoxyribosyltransferase is not commercially available at this time, the alternative enzymes (thymidine phosphorylase and purine nucleoside phosphorylase) are available. These routes then provide access to a wide variety of both 2 -deoxyribo- and ribonucleosides. UNIT 1.11 describes the widely used Robins procedure for chemically converting ribonucleosides to 2 -deoxyribonucleosides. Modified nucleosides containing reactive functionality, which have been denoted as “convertible” nucleosides when incorporated into oligonucleotides, are an increasingly
Current Protocols in Nucleic Acid Chemistry 1.0.1-1.0.3, September 2007 Published online September 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/0471142700.nc0100s30 C 2007 John Wiley & Sons, Inc. Copyright
Synthesis of Modified Nucleosides
1.0.1 Supplement 30
important class of compounds, of which the 2-fluoro-2 -deoxyinosine derivative described in UNIT 1.3 is a timely example (Huang et al., 1999). UNIT 1.4 provides a discussion on the topic of unnatural nucleotides with unusual base-pairing properties and of universal nucleotides. UNIT 1.5 continues this topic by providing step-by-step procedures for synthesizing N- or C-nucleosides with a number of specific base analogs. Introduction of a fluorine atom in nucleosides has often led to compounds with a potent biological activity. This was demonstrated in the anti-HIV field, and UNIT 1.12 describes the example of 2 -fluoro-2 ,3 dideoxyadenosine. UNIT 1.7 describes the synthesis of 2 -deoxy-2 -fluoroarabinonucleosides. Oligonucleotides containing these modified bases form stable heteroduplexes with RNA that are substrates for RNase H, which is important to antisense applications. UNIT 1.9 addresses the use of nucleosides with a modified sugar to create hexitol nucleic acids (HNAs). This unit decribes the synthesis of 1,5-anhydrohexitol nucleoside monomers as well as their starting sugar compound. These can be used in automated oligonucleotide synthesis with standard phosphoramidite chemistry to synthesize HNAs, which hybridize to RNA and DNA in a sequence-specific manner and have applications in antisense and antiviral approaches. The altritol building blocks, described in UNIT 1.18, differ from the hexitol building blocks by the presence of an additional hydroxyl group on the sugar moiety. Therefore, altritol nucleic acids (ANAs) can be considered the RNA analogs of HNA. The hybridization properties of HNA and ANA are very similar. ANA can also be used for antisense and siRNA applications, and show excellent hybridization properties when used on arrays. UNIT 1.10 describes preparation of pyrrolo[2,3-d]pyrimidine analogs of 2 -deoxyadenosine, 2 -deoxyguanosine, and 2 -deoxyisoguanosine. The incorporation of these compounds into DNA oligomers is described in UNIT 4.25.
A problem encountered in nucleic acid crystallography is the phasing of data, and the use of selenium-labeled DNA and RNA has proven to be very helpful in this process. The preparation of 2 -methylseleno nucleosides and their corresponding phosphoramidites are described in UNIT 1.15. Procedures are also given for synthesis and purification of the modified oligonucleotides as well as their ligation using T4 RNA/DNA ligase. For molecular diagnostics and gene regulation, the development of caged oligonucleotides provides a useful application of nucleoside modification. The principle is that a biologically active oligonucleotide is made inactive by derivatizing some of its nucleobases with a photolabile group, which is the cage. The active oligomer can be released using laser (light) at the desired time and location. These tools can also be used for physicochemical studies (for example, to study oligonucleotide folding) and for biological purposes (for example, for the modulation of protein function using aptamers). UNIT 1.17 describes the synthesis of phosphoramidites derivatized with a 2-nitrophenylethyl or 2-nitrophenylpropyl cage; these provide building blocks for synthesis of caged oligonucleotides on an automated synthesizer. Several other chapters in this book describe specific types of nucleoside modifications or modifications for specific purposes. Chapter 2, for instance, describes nucleoside protection, which is essentially the transient modification of a nucleoside moeity to allow selective modification of other moeities during subsequent reactions. Chapter 13 describes nucleoside phosphorylation and related modifications, and Chapters 14 and 15 describe modifications designed for specific biological functionality and for delivery of nucleosides in a prodrug form.
Introduction
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Current Protocols in Nucleic Acid Chemistry
LITERATURE CITED Beaucage, S.L. and Iyer, R.P. 1993. The synthesis of modified oligonucleotides by the phosphoramidite approach and their applications. Tetrahedron 49:6123-6194. Goodchild, J. 1990. Conjugates of olignucleotides and modified oligonucleotides: A review of their synthesis and properties. Bioconjugate Chem. 1:165-187. Huang, H., Chopra, R., Verdine, G.L., and Harrison, S.C. 1999. Structure of a covalently trapped catalytic complex of HIV-1 reverse transcriptase: Implications for drug resistance. Science 282:1669-1675.
Roger Jones and Piet Herdewijn
Synthesis of Modified Nucleosides
1.0.3 Current Protocols in Nucleic Acid Chemistry
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Palladium-Mediated C5 Substitution of Pyrimidine Nucleosides
UNIT 1.1
One of the most efficient ways to link a reporter group to oligonucleotides is through the incorporation of a modified nucleoside during automated oligonucleotide synthesis. Most techniques, which make use of synthetic oligonucleotides, function by hybridization to a complementary sequence. In order to avoid interference with hybridization, reporter groups should ideally be attached so that they do not interfere with hybridization or destabilize dsDNA. Two different types of tethers are described here—a rigid amidopropynyl linker and a flexible aminoethylthioether linker. The rigid amidopropynyl tether, linked through C5 of deoxyuridine, is sufficiently long and positioned such that a reporter group attached at the distal end lies outside the major groove of a DNA duplex. Basic Protocol 1 describes a detailed procedure for the synthesis of one example of deoxyuridine modified by an amidopropynyl-linked reporter group, 5-(3-nicotinamidopropyn-1-yl)-5′-O-(4,4′-dimethoxytrityl)-2′-deoxyuridine (Fig. 1.1.1). The procedure is general and may be applied to other amidopropynyl-linked functional groups. The nicotinoyl group was used only as an illustration of the strategy for incorporating a functional group on the amimopropynyl tether. For use in oligonucleotide synthesis, the C5-modified deoxyuridine is converted to a 3′-phosphoramite as described in UNIT 3.3. Basic Protocol 2 outlines the synthesis of 5-(3-acetamido-1-thiapropyl)-2′-deoxyuridine (Fig. 1.1.2). In contrast to the amidopropynyl tether, the more conformationally flexible thioether tether was designed to allow positioning of a molecular tool (e.g., chemical cleavage reagent or cross-linking reagent) on a complementary nucleic acid by hybridization of the modified oligonucleotide. The thiapropyl linker is capable of bridging the
O
O l HO
I
NH O
N
4,4'-dimethoxytrityl chloride
O
DMTrO
NH O
N
O
pyridine HO
HO 5-iodo-2'-deoxyuridine
5'-O-(4,4'-dimethoxytrityl)-5-iodo-2'-deoxyuridine
NH2
O
O
propargylamine
Cl N H Cl
N H
nicotinoyl chloride hydrochloride
Pd(PPh3)4 Cul
N
pyridine
N-(3-propyn-1-yl)nicotinamide
O
OCH3
O
N H
NH
N DMTrO
DMTr = H3CO
O
N
O
HO 4,4'-dimethoxytrityl
5-(3-nicotinamidopropyn-1-yl)-5'-O-(4,4'-dimethoxytrityl)-2'-deoxyuridine
Figure 1.1.1 Synthetic scheme for the preparation of 5-(3-nicotinamidopropyn-1-yl)-5′-O-(4,4′-dimethoxytrityl)-2′-deoxyuridine from 5-iodo-2′-deoxyuridine. The structure of 4,4′-dimethoxytrityl (DMTr) is shown in the lower left. Contributed by Mohammad Ahmadian, Douglas A. Klewer, and Donald E. Bergstrom Current Protocols in Nucleic Acid Chemistry (2000) 1.1.1-1.1.18 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Modified Nucleosides
1.1.1
span between two helices. For use in oligodeoxyribonucleotide synthesis, the N-acylated 5-(3-amino-1-thiapropyl)-2′-deoxyuridine is transformed to the 5′-dimethoxytrityl (DMTr) derivative as illustrated in Basic Protocol 1 for 5-iodo-2′-deoxyuridine, and is then converted to the 3′-phosphoramite as described in UNIT 3.3. Support Protocols 1 and 2 describe the preparation of two reagents needed for Basic Protocol 2—N,N′-bis(trifluoroacetyl)cystamine and N-acetoxysuccinimide, respectively. BASIC PROTOCOL 1
SYNTHESIS OF 5-(3-NICOTINAMIDOPROPYN-1-YL)-5′-O-(4,4′DIMETHOXYTRITYL)-2′-DEOXYURIDINE The sequence of reactions outlined here (see Fig. 1.1.1) illustrate conditions that are useful for the synthesis of a wide variety of reporter groups linked through C5 of deoxyuridine. The protocol includes three steps: synthesis of the N-acylated 3-aminopropyne (3-nicotinamidopropyne, in this example), reaction of 5-iodo-2′-deoxyuridine with 4,4′-dimethoxytrityl chloride, and palladium-catalyzed coupling of 3-nicotinamidopropyne with 5′-O-(4,4′-dimethoxytrityl)-5-iodo-2′-deoxyuridine. For the introduction of a different reporter group, 3-aminopropyne can be N-acylated by RCOCl (an acid chloride) or RC(O)OC(O)R (an anhydride) to obtain RC(O)NHCH2C≡CH, in which R is the desired reporter group. CAUTION: All reactions should be run in a suitable fume hood to avoid inhalation of toxic vapors. Materials Nicotinoyl chloride hydrochloride Pyridine, anhydrous Nitrogen (N2) stream Triethylamine, freshly distilled (dried and purified by distillation at atmospheric pressure over calcium hydride; boiling point = 89° to 90°C) Propargylamine, reagent grade (typically 99% pure) Dichloromethane, reagent grade 10% (w/v) hydrochloric acid in water
O
O
O NH
N
dR
(CF3CONHCH2CH2S)2
NH
2. NaCl
O
N
ClHg
1. Hg(OAc)2
F3C
Li2PdCl4
O
O N H
S
dR
2'-deoxyuridine
NH O
N dR
5-[3-(trifluoroacetamido)-1-thiapropyl]-2'-deoxyuridine
5-chloromercurio-2'-deoxyuridine
O DCC
O N-OH
NH4OH
CH3COH
+
acetic acid
O N-hydroxysuccinamide
O
O
N O O HO
O
dR = HO
H3C
O
O N H
O
S
H2N
NH
S
NH
N-succinimidylacetate N
O
dR
N
O
dR
deoxyribosyl
PalladiumMediated C5 Substitution of Pyrimidine Nucleosides
5-(3-acetamido-1-thiapropyl)-2'-deoxyuridine
5-(3-amino-1-thiapropyl)-2'-deoxyuridine
Figure 1.1.2 Synthetic scheme for the preparation of 5-(3-acetamido-1-thiapropyl)-2′-deoxyuridine from 2′-deoxyuridine. The structure of deoxyribosyl (dR) is shown in the lower left. DCC, dicyclohexylcarbodiimide.
1.1.2 Current Protocols in Nucleic Acid Chemistry
Sodium sulfate, anhydrous Silica gel (230 to 400 mesh) Methanol, reagent grade 5-Iodo-2′-deoxyuridine 4,4′-Dimethoxytrityl chloride Diethyl ether, anhydrous N,N-Dimethylformamide, anhydrous Argon gas (optional) Tetrakis(triphenylphosphine)palladium, [(C6H5)3P]4Pd Copper(I) iodide 5% (w/v) Na2EDTA in water Ethyl acetate, reagent grade 25- and 50-mL round-bottom flasks Inert atmosphere/vacuum manifold (see Fig. 1.1.3) 500-µL and 1-mL syringes with stainless steel needles 125- and 250-mL Ehrlenmeyer flask 100-mL separatory funnel Filter funnel and Whatman no. 1 filter paper Chromatotron and radial chromatography plate coated with silica gel (2-mm thickness; Harrison Research) Rotary evaporator with vacuum pump and water aspirator Glass column (2-cm i.d. × ≥20-cm length) with stopcock Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 4D) Synthesize N-(3-propyn-1-yl)nicotinamide 1. In a dry 25-mL round-bottom flask containing a 1⁄2-in. magnetic stir bar, add 1.068 g nicotinoyl chloride hydrochloride (6 mmol) to 10 mL of anhydrous pyridine under a nitrogen stream. It is important that the flask be dry because nicotinoyl chloride reacts with water to give nicotinic acid. Glassware can be effectively dried by heating in a drying oven at 120°C for 2 hr. A small magnetic stir bar is generally dried at the same time as the flask and added to the flask prior to addition of the reagents. A general setup for running small-scale reactions under a dry nitrogen atmosphere is shown in Figure 1.1.3. The apparatus is configured for air-sensitive palladium-catalyzed reactions. For most other reactions, it is not necessary to bubble nitrogen through the reaction mixture at inlet (b).
2. Add 500 µL triethylamine (700 mg, 7 mmol) with a 1-mL syringe and stainless steel needle, and stir the mixture on a magnetic stirrer at room temperature until the triethylamine is completely in solution. CAUTION: Wear reagent-impermeable protective gloves. Triethylamine and propargylamine are corrosive.
3. Add 250 µL propargylamine (365 mg, 6.6 mmol) dropwise to the reaction mixture with a 500-µL syringe and stainless steel needle, and continue stirring under nitrogen at room temperature for 4 hr. 4. Transfer contents of the flask to a 125-mL Ehrlenmeyer flask containing 40 mL water. Stir the mixture briefly, transfer to a 100-mL separatory funnel, and extract three times with 40 mL reagent-grade dichloromethane.
Synthesis of Modified Nucleosides
1.1.3 Current Protocols in Nucleic Acid Chemistry
Figure 1.1.3 Inert atmosphere/vacuum manifold setup for running reactions in a dry, oxygen-free atmosphere. As shown, the inert gas can be introduced into the reaction flask through stopcock (a) and a hypodermic needle inserted at (b; 14/20 standard taper joint). The slow stream of inert gas then passes through stopcock (d) and out through the gas bubbler. Care must be taken to avoid completely closing the system while the insert gas is being introduced under pressure through the manifold. The setup requires a source of dry nitrogen. For very oxygen-sensitive reactions, the solution is purged by bubbling the inert gas (nitrogen or argon) directly through the solution by lowering the stainless steal hypodermic needle into the solution. The needle is then pulled up above the level of the solution and the flask and condenser evacuated through stopcock (a) with stopcock (d) positioned to allow only the inert gas to pass through to the gas bubbler.
5. Wash the combined organic extracts twice with 20 mL of 10% hydrochloric acid and then once with 10 mL water. 6. Transfer the combined dichloromethane solution to a 250-mL Erlenmeyer flask and add 0.5 g anhydrous sodium sulfate. Swirl the solution for a few minutes and then allow to stand for 30 min. 7. Remove the drying agent by gravity filtration through a filter funnel fitted with Whatman no. 1 filter paper. PalladiumMediated C5 Substitution of Pyrimidine Nucleosides
8. Wash the solid with 10 mL dichloromethane and remove solvent under reduced pressure using a rotary evaporator and water aspirator at room temperature to obtain the crude product.
1.1.4 Current Protocols in Nucleic Acid Chemistry
With a water aspirator and a water bath temperature of 25°C, the dichloromethane can generally be completely removed within 30 min. Longer periods of time may be required for complete removal of other higher-boiling-temperature solvents, such as methanol (step 9).
9. Purify the crude product by radial chromatography according to manufacturer’s instructions using a chromatotron plate with 2-mm silica gel thickness. Elute with 96:4 (v/v) dichloromethane/methanol and collect the effluent in ∼10-mL fractions. The silica gel plates may be purchased from the manufacturer or prepared according to the manufacturer’s instructions. Alternatively, the crude product can be purified by column chromatography on silica gel (230 to 400 mesh; 12 × 2 cm) and eluted with the same solvent to give ∼10-mL fractions.
10. Analyze fractions by thin-layer chromatrography (TLC) on silica gel. Develop TLC plates with 96:4 (v/v) dichloromethane/methanol. 11. Combine all fractions that contain the desired product (Rf = 0.26). Evaporate the solvent under reduced pressure (see step 8) to obtain N-(3-propyn-1-yl)nicotinamide (784 mg, 78%) as a white solid. The compound is stable at room temperature and can be stored in a capped amber glass vial that has been purged with nitrogen. The authors do not generally store the product for >1 month, but it may be stable for a longer period of time. Unless otherwise specified, all intermediates synthesized as part of this protocol are stored under these conditions.
12. Analyze the product by mass spectrometry (MS) and by proton and carbon nuclear magnetic resonance (NMR) spectroscopy. N-(3-Propyn-1-yl)nicotinamide has the following spectroscopic characteristics: MS-EI 160 (M+), 106, 78 MS-CI 161 (M + H)+ H NMR 250 MHz (CHCl3-d1) d 9.0 (d, J = 1.6 Hz, H-2 aromatic, 1H), 8.74 (m, H-6, 1H), 8.16 (m, H-5, 1H), 7.41 (m, H-4, 1H), 6.79 (br s, H-N, 1H), 4.28 (dd, J1 = 5.2 Hz, J2 = 2.5 Hz, N-CH2-, 2H), 2.32 (t, J = 2.5 Hz, CCH, 1H)
1
C NMR 62.9 MHz (CHCl3-d1) δ 29.90, 72.32, 79.0, 123.62, 135.26, 147.94, 152.61
13
Analysis calculated for C9H8N2O: C, 67.5; H, 5.0; N, 17.5; observed: C, 67.24; H, 4.82; N, 17.65. All values are given as percentages. The same procedure may be used for the synthesis of other amide derivatives of propargylamine from carboxylic acid chlorides or anhydrides. The Rf and the spectral characteristics will differ depending on the nature of the acyl group.
Synthesize 5′-O-(4,4′-dimethoxytrityl)-5-iodo-2′-deoxyuridine 13. In a 50-mL round-bottom flask containing a 3⁄4-in. egg-shaped magnetic stir bar, dissolve 354 mg of 5-iodo-2′-deoxyuridine (1 mmol) in 10 mL anhydrous pyridine. CAUTION: The reaction must be performed in a well-vented fume hood. This reaction is sensitive to water, and anhydrous solvent(s) must be used under inert atmosphere. Anhydrous pyridine obtained in Sure/Seal bottles (e.g., Aldrich) is suitable for use in this reaction without further drying. Otherwise, the pyridine should be dried over solid KOH and distilled over Linde type 5Å molecular sieves and solid KOH. Pyridine has a fairly high boiling point (115°C).
14. Evaporate approximately half the solvent using a rotary evaporator connected to a vacuum pump.
Synthesis of Modified Nucleosides
1.1.5 Current Protocols in Nucleic Acid Chemistry
It is advisable to use a vacuum pump rather than a water aspirator in order to rapidly evaporate the pyridine. This procedure removes water from the reaction mixture that may have been associated with the nucleoside by way of a pyridine-water azeotrope. With a good vacuum (<1 mmHg) it is possible to concentrate the reaction mixture in 10 to 30 min.
15. Add 406 mg 4,4′-dimethoxytrityl chloride (1.2 mmol) to the resulting solution and stir the mixture overnight at room temperature under a dry nitrogen atmosphere (Figure 1.1.3). In this case it is not necessary to bubble nitrogen through the reaction mixture. The apparatus shown in Figure 1.1.3 provides a means to keep the reaction mixture dry. 4,4′-Dimethoxytrityl chloride may deteriorate if stored for long periods of time. The reagent should be light orange in color; do not use if it appears red. Although the reaction with 5-iodo-2′-deoxyuridine is allowed to run overnight, with pure reagents the reaction is typically compete within 1 to 2 hr.
16. Add 15 mL ice-cold water to the reaction mixture and then extract twice with 20 mL dichloromethane. 17. Combine the two organic extracts, wash with 10 mL water, and dry over anhydrous sodium sulfate (step 6). 18. Remove the drying agent, wash, and remove solvent as described (steps 7 and 8). 19. Dissolve the residue in ∼0.5 mL dichloromethane, introduce the resulting solution to a 10 × 2–cm column of 230- to 400-mesh silica gel, and elute with 98:2 (v/v) dichloromethane/methanol. 20. Collect the effluent in ∼10-mL fractions and analyze by TLC on silica gel. Develop the plates with 98:2 (v/v) dichloromethane/methanol. 21. Combine fractions that contain the desired product (Rf = 0.36) and evaporate the solvent under reduced pressure (step 8). 22. Add 2 mL anhydrous diethyl ether to the residue and evaporate under vacuum (step 8). Repeat this step until a white foam (646 mg, 98.5% yield) is obtained. It is important to continue the evaporation until a foam is obtained. Otherwise, the product will contain unacceptable amounts of solvents that may interfere with the subsequent reaction. This can generally be accomplished by using a rotary evaporator connected to a water aspirator, with the flask partially immersed in a room temperature water bath. The product may be stored under dry nitrogen in an amber bottle in the dark for a few weeks.
23. Analyze the product by mass spectrometry and by proton and carbon NMR spectroscopy. 5′-O-(4,4′-Dimethoxytrityl)-5-iodo-2′-deoxyuridine has the following spectroscopic characteristics: 1 H NMR 250 MHz (CHCl3-d1) d 8.38 (s, N3-H, 1H), 8.13 (s, H-6, 1H), 7.46 to 7.23 (m, DMTr aromatic protons, 9H), 6.85 (d, J = 8.8 Hz, DMTr aromatic protons, 4H), 6.30 (dd, J1 = 7.6 Hz, J2 = 5.5 Hz, H-1′, 1H), 4.54 (m, H-3′, 1H), 4.08 (m, H-4′, 1H), 3.80 (s, OCH3, 6H), 3.40 (m, H-5′, 2H), 2.33 and 2.44 (two sets of multiplets, H-2′, 2H), 1.98 (br s, 3′-OH, 1H)
PalladiumMediated C5 Substitution of Pyrimidine Nucleosides
C NMR 62.9 MHz (CHCl3-d1) d 159.83, 158.66, 149.7, 144.26, 135.39, 135.28, 130.08, 128.11, 128.01, 127.10, 123.80, 113.37, 87.06, 86.45, 85.55, 72.46, 68.51, 63.41, 55.27, 41.46
13
MS-PD m/z calculated for C30H29IN2O7: 656; observed: 656 (M+).
1.1.6 Current Protocols in Nucleic Acid Chemistry
The same procedure may be applied to other C5-substituted deoxyuridine derivatives such as 5-(3-acetamido-1-thiapropyl)-2′-deoxyuridine or 5-(3-trifluoroacetamido-1-thiapropyl)-2′-deoxyuridine (see Basic Protocol 2). The Rf and spectral characteristics will differ depending on the nature of the C5 substituent.
Synthesize 5-(3-nicotinamidopropyn-1-yl)-5′-O-(4,4′-dimethoxytrityl)-2′-deoxyuridine 24. In a two-neck 25-mL round-bottom flask containing a 1⁄2-in. magnetic stir bar (see Fig. 1.1.3), dissolve 328 mg of 5′-O-(4,4′-dimethoxytrityl)-5-iodo-2′-deoxyuridine (0.5 mmol; step 22) in 4 mL anhydrous N,N-dimethylformamide. Anhydrous N,N-dimethylformamide obtained in Sure/Seal bottles (Aldrich) may be used without further drying. Alternatively, N,N-dimethylformamide may be dried by distillation at reduced pressure (boiling point = 76°C at 39 mmHg) over Linde type 4Å molecular sieves.
25. Remove any dissolved oxygen in the solution by alternating three times between a vacuum and inert gas (dry nitrogen or argon) in the reaction apparatus. As shown in Figure 1.1.3, the inert gas can be introduced directly to the solution through a tube attached at stopcock (a) and a hypodermic needle inserted at (b). When pulling a vacuum through stopcock (a), the needle must be lifted above the solution in the flask. The subsequent coupling reaction proceeds by way of a zero-valent palladium complex. Although the dry reagents may be weighed out in the atmosphere, the reaction is very sensitive to oxygen once the reagents are in solution. A complex mixture of products may result if all oxygen is not removed from the system at this point.
26. Add the following reagents in the order indicated and then stir the reaction mixture at room temperature under nitrogen for 8 hr. 300 µL freshly distilled triethylamine (218 mg, 2.15 mmol) 240 mg N-(3-propyn-1-yl)nicotinamide (1.5 mmol; step 10) 58 mg tetrakis(triphenylphosphine)palladium (0.05 mmol) 19 mg copper(I) iodide (0.1 mmol). Experience with many different alkyne-coupling reactions has shown that it is not always possible to predict which palladium reagent will work most effectively. In some cases (e.g., with 3,3-dimethoxypropyne), bis(triphenylphosphine)palladium dichloride is more effective.
27. Add 10 mL of 5% Na2EDTA to the reaction mixture and extract twice with 30 mL dichloromethane. 28. Combine extracts and wash with 10 mL water. Dry the organic solution over anhydrous sodium sulfate (step 6). 29. Remove sodium sulfate by filtration (step 7), wash, and evaporate the solvent (step 8) to obtain the crude product as a foamy solid. 30. Purify the crude product by radial chromatography as in step 9, but elute with 50:50:5 (v/v/v) ethyl acetate/dichloromethane/methanol. Alternatively, the crude product can be purified by column chromatography on silica gel (230 to 400 mesh; 10 × 2 cm) and eluted with 99:1 (v/v) dichloromethane/methanol.
31. Analyze fractions by TLC on silica gel. Develop TLC plates with 99:1 (v/v) dichloromethane/methanol.
Synthesis of Modified Nucleosides
1.1.7 Current Protocols in Nucleic Acid Chemistry
32. Combine all fractions that contain the desired product (Rf = 0.39). Evaporate the solvent under reduced pressure (step 8). 33. Add 2 mL diethyl ether to the residue and evaporate under vacuum (step 8). Repeat this step until a white foam (215 mg, 62.5% yield) is obtained. The product may be stored a short period of time (a few weeks) under nitrogen in an amber bottle. Normally, it is converted immediately to the phosphoramidite for incorporation into an oligonucleotide.
34. Analyze the product by mass spectrometry and by proton and carbon NMR spectroscopy. 5′-O-(4,4′-Dimethoxytrityl)-5-(3-nicotinamidopropyn-1-yl)-2′-deoxyuridine has the following spectroscopic characteristics: 1 H NMR 500 MHz (CHCl3-d1) d 12.84 (s, N3-H, 1H) 8.90 (d, J = 1.5, H-2 pyridine ring, 1H), 8.68 (dd, J1 = 4.5 Hz, J2 = 2 Hz, H-4 pyridine moiety, 1H), 8.22 (s, H-6 pyrimidine moiety, 1H), 7.90 (m, H-6 pyridine moiety, 1H), 7.65 (s, NH-CO-,1H), 7.47 to 7.17 (several sets, m, DMTr aromatic protons overlapped with H-5 pyridine ring, 10H), 6.80 (d, J = 7 Hz, DMTr aromatic protons, 4H), 6.14 (t, J = 10 Hz, H-1′, 1H), 4.59 (m, H-3′, 1H), 4.18 (m, -CH2N- and H-4′, 3H), 3.85 (s, OCH3, 6H), 3.35 (m, H-5′, 1H), 3.45 (m, H-5′′, 1H), 2.56 (m, H-2′, 1H), 2.32 (m, H-2′′, 1H) 13 C NMR 62.9 MHz (CHCl3-d1) d 162.42, 158.30, 151.84, 149.61, 149.18, 148.47, 148.26, 142.75, 135.58, 135.40, 135.19, 134.80, 129.79, 129.50, 127.72, 126.66, 123.33, 123.23, 122.94, 122.54, 113.05, 86.70, 65.61, 70.83, 63.42, 54.95, 41.69, 30.32
MS-CI m/z calculated for C39H36N4O8: 688; observed: 689 (M + H)+ MS-PD m/z observed: 688.8 (M)+, 303 (DMTr)+ Analysis calculated for C39H36N4O8: C, 68.01; H, 5.21; N, 8.13; observed: C, 67.72; H, 5.26; N, 8.31. All values are given as percentage. Other alkynes will give 5-substituted 2′-deoxyuridine products with different Rf values and spectral characteristics. BASIC PROTOCOL 2
SYNTHESIS OF 5-(3-ACETAMIDO-1-THIAPROPYL)-2′-DEOXYURIDINE This protocol describes synthesis of 5-(3-acetamido-1-thiapropyl)-2′-deoxyuridine. The sequence of reactions outlined here (Fig. 1.1.2) illustrates conditions useful for the synthesis of different functional groups linked through an aminothiapropyl tether to C5 of deoxyuridine. The protocol includes four steps: preparation of 5-chloromercurio-2′deoxyuridine, palladium-mediated coupling with N,N′-bis(trifluoroacetyl)cystamine, removal of the trifluoroacetyl protecting group by ammonia, and coupling of an active ester to the pendant amino group. The latter reaction is illustrated with the active ester N-acetoxysuccinimide. Other active esters may be used to link modifying groups to the aminothiapropyl tether. For example, a modified nucleoside that has a bipyridine ligand linked to the amino group of 5-(3-amino-1-thiapropyl)-2′-deoxyuridine was synthesized via an active ester of 4-carboxy-4′-methylbipyridine. This modified nucleotide was used to prepare oligonucleotides that function as metal-mediated sequence-specific nucleases (Bergstrom and Chen, 1996). CAUTION: Mercuric acetate and organomercury compounds are highly toxic. Wear gloves and properly dispose of all waste materials generated by this procedure.
PalladiumMediated C5 Substitution of Pyrimidine Nucleosides
Materials 2′-Deoxyuridine Mercury(II) acetate
1.1.8 Current Protocols in Nucleic Acid Chemistry
30% (w/v) and 0.1 M sodium chloride (reagent grade) in water Ethanol, anhydrous Diethyl ether, anhydrous N,N′-Bis(trifluoroacetyl)cystamine (see Support Protocol 1) 0.1 M Li2PdCl4 solution (see recipe) Hydrogen sulfide (H2S) Methanol, reagent grade Chloroform, reagent grade Silica gel (230 to 400 mesh) Concentrated ammonium hydroxide Dry ice/acetone (for freezing) N-Acetoxysuccinimide (see Support Protocol 2) Triethylamine, freshly distilled (dried and purified by distillation at atmospheric pressure over calcium hydride; boiling point = 89° to 90°C) Tetrahydrofuran, anhydrous Ethyl acetate, reagent grade 100- and 200-mL round-bottom flasks Temperature-controlled magnetic stirrer Mortar and pestle Whatman no. 1 filter paper 100-mm-diameter porcelain Buchner funnel Rotary evaporator with water aspirator Glass chromatography column (2-cm i.d. × ≥20-cm length) with stopcock Lyophilizer Synthesize 5-chloromercurio-2′-deoxyuridine 1. Dissolve 7.094 g of 2′-deoxyuridine (31.09 mmol) in 30 mL water in a 200-mL round-bottom flask and add a 1-in. magnetic stir bar. 2. Dissolve 10.449 g mercury(II) acetate (32.8 mmol) in 45 mL water and add to the deoxyuridine. 3. Add 25 mL water and stir on a magnetic stirrer at 50°C for 2 hr. During the course of the reaction, a dense white suspension of acetoxymercuriodeoxyuridine is formed. This is converted to the chloromercurio derivative in step 4.
4. While stirring, cool the reaction to 40°C and add 15 mL of 30% sodium chloride (4.5 g, 0.077 moles). 5. Cool the reaction mixture to room temperature and filter the solution using a 100-mm-diameter porcelain Buckner funnel and Whatman no. 1 filter paper. 6. Wash the fine white precipitate sequentially with 60 mL of 0.1 M NaCl, 40 mL water, 20 mL ethanol, and 30 mL anhydrous diethyl ether. 7. Dry the white precipitate in a vacuum oven under vacuum at 80°C to obtain 5-chloromercurio-2′-deoxyuridine (13.95 g; 97% yield; melting point = 210.5° to 211°C). 5-Chloromercurio-2′-deoxyuridine is neither water nor air sensitive. It has been successfully stored without decomposition in amber bottles at room temperature for >5 years.
8. Analyze the product by IR, UV, and 1H NMR spectroscopy. 5-Chloromercurio-2′-deoxyuridine has the following spectroscopic characteristics:
Synthesis of Modified Nucleosides
1.1.9 Current Protocols in Nucleic Acid Chemistry
H NMR (1.0 M KCN/D2O) d 7.70 (s, 1H), 6.35 (t, 1H, J = 6.5 Hz), 4.47 (m, 1H), 3.99 (m, 1H), 3.83 (m, 2H), 2.36 (2H, dd, J = 6 Hz) 1
IR (KBr) 3365, 1714, 1642, 1440, 1275, 1089, 1040 cm −1 UV (pH 1.0) lmax 266 (ε = 10,440),lmin 239 (ε = 4,180); (pH 9.0)lmax 266 (e = 10,120), lmin 242 (e = 5,440); (pH 12.3) lmax 267 (e = 8,870),lmin 253 (e = 7,070) Analysis calculated for C9H11N2O5HgCl: C 23.34; H 2.39; N 6.05; observed: C 23.54; H 2.32; N 5.89.
Synthesize 5-[3-(trifluoroacetamido)-1-thiapropyl]-2′-deoxyuridine 9. Grind 5-chloromercurio-2′-deoxyuridine to a fine powder using a mortar and pestle. CAUTION: In the process of grinding, the powder tends to accumulate static electricity and may be difficult to contain in the mortar and pestle. The mortar should be placed on a surface that can be easily cleaned. It is preferable to carry out this step in a hood to avoid breathing the powder. Grind ≥10% more material than required for the subsequent step to make up for loss that occurs during this process.
10. Place 0.926 g finely ground 5-chloromercurio-2′-deoxyuridine (2.0 mmol) and 1.720 g N,N′-bis(trifluoroacetyl)cystamine (5 mmol) in a 100-mL round-bottom flask. 11. Add 40 mL of 0.1 M Li2PdCl4 solution to the flask and stir on a magnetic stir plate for 16 hr at ambient temperature. The mixture turns orange to yellow shortly after the reagents are combined, and usually yields a clear orange-yellow solution within a few hours.
12. Rapidly bubble hydrogen sulfide through the solution for 30 sec. Filter the mixture through a filter funnel by gravity filtration through Whatman no. 1 filter paper, and wash the solid with 20 mL reagent-grade methanol. CAUTION: Hydrogen sulfide gas is highly toxic. All operations should be conducted in a well-ventilated fume hood. Hydrogen sulfide may be obtained in 1⁄2-lb (227-g) lecture bottles.
13. Using a rotary evaporator with a water aspirator, evaporate the solvent from the filtrate under reduced pressure to give an oil. Because the methanol solution still contains hydrogen sulfide gas, this evaporation should be done using a rotary evaporator located inside a fume hood. Water aspirator pressure is normally sufficient to remove the methanol within 30 min at room temperature.
14. Purify the crude product on a 12 × 2–cm, 230- to 400-mesh silica gel column, eluting with a linear chloroform/methanol gradient ranging from 10% to 18% methanol. 15. Combine fractions that contain material with Rf = 0.30 (CH3OH-CHCl3 1:9 v/v) or Rf = 0.71 (CH3OH-CHCl3 1:3 v/v), and evaporate the solvent on a rotary evaporator using a water aspirator to obtain 5-[3-(trifluoroacetamido)-1-thiapropyl]-2′-deoxyuridine (0.41 g; 51% yield). 5-[3-(Trifluoroacetamido)-1-thiapropyl]-2′-deoxyuridine is neither water nor air sensitive. It can be stored without decomposition in amber bottles at room temperature for many years.
16. Analyze the product by MS and by IR, 1H, and 13C NMR spectroscopy. PalladiumMediated C5 Substitution of Pyrimidine Nucleosides
5-[3-(trifluoroacetamido)-1-thiapropyl]-2′-deoxyuridine has the following spectroscopic characteristics: MS-FAB m/z calculated for C13H16F3N3O6S: 399.071; observed: 400.079 (M + H)+
1.1.10 Current Protocols in Nucleic Acid Chemistry
H NMR 300 MHz (CH3OH-d4) d 8.32 (s, 1H, H-6), 6.27 (t, 1H, J = 6.6 Hz, H-1′), 4.43 (m, 1H, H-3′), 3.95 (m, 1H, H-4′), 3.79 (m, 2H, H-5′), 3.47 (t, 2H, J = 6.0 Hz, H-3′′), 2.87 (m, 2H, H-2′), 2.32 (t, 2H, J = 6.0 Hz, H-2′′) 1
13
C NMR 125 MHz (CH3OH-d4) 164.5 (C4), 159.0 (q, J = 286.6 Hz, C5′′), 106.8 (C5), 89.0 (C1′), 86.8 (C4′), 72.0 (C3′), 62.7 (C5′), 41.4 (C2′), 39.7 (C3′′), 33.5 (C2′′) IR (KBr): 3550-2900 (br, O-H), 3423, 3443 (N-H), 1723, 1692, 1651 (C=O), 1660, 1553, 1461 (C=C), 1179, 1271 cm−1 Analysis calculated for C13H16F3N3O6S: C, 39.08; H, 4.01; N, 10.52; S, 8.03; observed: C, 39.35; H, 3.63; N, 10.38; S, 8.22.
Remove trifluoroacetyl protecting group 17. In a 100-mL round-bottom flask containing a 1-in. magnetic stir bar, dissolve 615 mg of 5-[3-(trifluoroacetamido)-1-thiapropyl]-2′-deoxyuridine (1.5 mmol) in 10 mL methanol. 18. Add 30 mL concentrated ammonium hydroxide, cap the reaction container, and stir at room temperature for 16 hr. 19. Remove excess ammonia and methanol under reduced pressure using a rotary evaporator with a water aspirator. Freeze the remainder of the solution with a mixture of dry ice and acetone. The methanol and ammonia can generally be removed on a rotary evaporator in 30 min or less at room temperature.
20. Remove water by lyophilization and dissolve the remaining solid in 3 mL anhydrous ethanol. Synthesize 5-(3-acetamido-1-thiapropyl)-2′-deoxyuridine 21. Prepare a solution of 315 mg N-acetoxysuccinimide (2 mmol) and 350 µL freshly distilled triethylamine (2.5 mmol) in 2 mL anhydrous tetrahydrofuran. Add to ethanolic solution and stir at room temperature for 4 hr. Other active esters may be used in place of N-acetoxysuccinimide to place a different functional group on the tether. The resulting chromatographic characteristics and spectroscopic properties will change accordingly.
22. Remove the solvent under reduced pressure using a rotary evaporator and a water aspirator to obtain the crude product. 23. Purify the crude product by column chromatography using a 12 × 2–cm silica gel column, and eluting with a gradient of 100% ethyl acetate to 80:20 (v/v) ethyl acetate/methanol. 24. Collect effluent in ∼10-mL fractions and analyze by TLC on silica gel, developing the plates with 85:15 (v/v) ethyl acetate/methanol. 25. Combine all fractions that contain the desired product (Rf = 0.32). Evaporate the solvent under reduced pressure using a rotary evaporator and a water aspirator to obtain 5-[3-acetamido-1-thiapropyl]-2′-deoxyuridine (462 mg; 87% yield) as a white solid. The product may be stored indefinitely under nitrogen in an amber bottle.
26. Analyze the product by MS and by UV, 1H, and 13C NMR spectroscopy. 5-(3-Acetamido-1-thiapropyl)-2′-deoxyuridine has the following spectroscopic characteristics:
Synthesis of Modified Nucleosides
1.1.11 Current Protocols in Nucleic Acid Chemistry
MS-FAB m/z calculated for C13H19N3O6S: 345.1073; observed: 346.1072 (M + H)+ H NMR 300 MHz (CH3OH-d4) d 8.33 (s, H-6, 1H), 6.25 (t, J = 7 Hz, H-1′), 4.41 (m, H-3′, 1H), 3.93 (m, H-4′, 1H), 3.78 (m, H-5′, 2H), 3.31 (m, SCH2-, 2H), 2.78 (t, J = 6 Hz, -CH2N-, 2H), 2.29 (m, H-2′, 2H), 1.95 (s, acetyl group’s CH3, 3H) 1
13
C NMR 125 MHz (DMSO-d6) 169.3, 161.65, 150.0, 142.55, 106.8, 87.54, 84.58, 70.23, 61.1, 39.94, 37.88, 32.1, 22.6 UV (methanol) lmax: 282.4, 202.0 nm Analysis calculated for C13H19N3O6S: C, 45.2; H, 5.5; N, 12.2; S, 9.3; observed: C, 44.88; H, 5.37; N, 12.27; S, 9.32.
SUPPORT PROTOCOL 1
SYNTHESIS OF N,N′-BIS(TRIFLUOROACETYL)CYSTAMINE The trifluoroacetyl group has found wide application as a base-sensitive protecting group. It can be removed by ammonia under the same conditions used to deprotect oligodeoxyribonucleotides following synthesis by the phosphoramidite method. Amines are most commonly converted to trifluoracetyl derivatives by treatment with trifluoroacetic anhydride and a tertiary amine. CAUTION: Both triethylamine and trifluoroacetic anhydride are corrosive. Wear gloves and work only in a suitable hood. Materials Chloroform, reagent grade Cystamine dihydrochloride Triethylamine, freshly distilled (dried and purified by distillation at atmospheric pressure over calcium hydride; boiling point = 89° to 90°C) Trifluoroacetic anhydride 10% (w/v) NaHCO3 2 N HCl Sodium sulfate, anhydrous Methanol, reagent grade Ethyl acetate, reagent grade Hexane, reagent grade 1-liter round-bottom flask Drying tube containing Drierite 5-mL syringe 1-liter separatory funnel Rotary evaporator with water aspirator Vacuum oven at 35°C Buchner funnel and Whatman no. 1 filter paper Synthesize N,N′-bis(trifluoroacetyl)cystamine 1. Filter reagent-grade chloroform through a short column of basic alumina and add 500 mL to a 1-liter round-bottom flask containing a magnetic stir bar and capped by a drying tube containing Drierite. Reagent-grade chloroform typically contains ethanol to inhibit decomposition, which produces HCl. The basic alumina removes the ethanol.
PalladiumMediated C5 Substitution of Pyrimidine Nucleosides
2. Add 9.0 g cystamine dihydrochloride (40 mmol) and 20.2 g triethylamine (27.8 mL, 0.2 mol) to the filtered chloroform.
1.1.12 Current Protocols in Nucleic Acid Chemistry
3. Cool the flask in a cold water bath (0° to 5°C) while slowly adding 1.5 mL trifluoroacetic anhydride (18.5 g, 88 mmol) with a 5-mL syringe. Trifluoroacetic anhydride, which hydrolyzes to trifluoroacetic acid when exposed to water, should be stored in a hood and protected from moisture.
4. Stir the reaction mixture at room temperature overnight. 5. Transfer the reaction mixture to a 1-liter separatory funnel and wash sequentially with 250 mL water, 10% NaHCO3, 2 N HCl, and water again. 6. Dry the chloroform solution over anhydrous sodium sulfate for 1 hr (see Basic Protocol 1, step 6). 7. Add 30 mL reagent-grade methanol, filter to remove the solid sodium sulfate (see Basic Protocol 1, step 7), and evaporate the solvent on a rotary evaporator using a water aspirator to yield a light yellow paste. 8. Add 100 mL of 2 N HCl to give a slurry of light yellow crystals, and stir for 20 min at room temperature. 9. Filter the slurry and wash the solid product with 100 mL of 2 N HCl and then with 100 mL water. 10. Dry the solid product in a vacuum oven at 35°C overnight. The product is of suitable purity for use in palladium coupling reactions. To obtain analytically pure product, recrystallize as described below. The product may be stored indefinitely in a closed bottle at room temperature.
Recrystallize product 11. Dissolve product in a minimum of hot methanol. 12. Slowly add ethyl acetate to the warm methanol solution and allow to cool. On cooling to room temperature, a colorless crystalline product separates (melting point = 111°C).
13. Collect crystals by vacuum filtration through a Buchner funnel using Whatman no. 1 filter paper. 14. Collect a second crop by adding hexane to the warmed methanol/ethyl acetate solution, and cooling and collecting crystals again. The total yield of crystalline product is 10.8 g (84%). The product may be stored indefinitely in a closed bottle at room temperature.
Analyze product 15. Analyze the product by 1H and 13C NMR spectroscopy. H NMR (250 MHz, acetone-d6) d 2.70 (t, CH2 4H, J = 10.4 Hz), 2.68 (q, CH2 4H)
1
C NMR (62.9 MHz, acetone-d6) d 37.2 (CH2, J = 10.4 Hz), 39.6 (CH2), 117.0 (CF3, J = 287 Hz), 157.8 (C=O).
13
SYNTHESIS OF N-ACETOXYSUCCINIMIDE N-Hydroxysuccinamide esters are generally prepared from carboxylic acids by reaction with dicyclohexylcarbodiimide and N-hydroxysuccinimide. The procedure outlined below can be applied to the synthesis of other N-hydroxysuccinimide esters. Active esters of some compound classes (e.g., amino acids) are commercially available.
SUPPORT PROTOCOL 2 Synthesis of Modified Nucleosides
1.1.13 Current Protocols in Nucleic Acid Chemistry
Materials N-Hydroxysuccinamide Tetrahydrofuran, anhydrous Nitrogen (N2) gas Glacial acetic acid Dicyclohexylcarbodiimide (DCC) Silica gel (optional; 230 to 400 mesh) Ethyl acetate, reagent grade Methanol, reagent grade Vacuum manifold apparatus (Fig. 1.1.3) modified with a 10-mL conical flask and a 500-µL syringe Filter funnel and Whatman no. 1 filter paper Glass chromatography column (optional; 2-cm i.d. × 10-cm length) with stopcock 1. In a 10-mL conical flask containing a small magnetic stir bar, prepare a solution of 345.3 mg N-hydroxysuccinamide (3 mmol) in 1 mL anhydrous tetrahydrofuran under an inert atmosphere (e.g., nitrogen). The setup shown in Figure 1.1.3 may be used with the inert gas inlet at (b) replaced by a 500-mL syringe containing glacial acetic acid (step 2). The apparatus is maintained under a positive pressure of nitrogen by the inlet line at (c).
2. Add 174 µL glacial acetic acid (3 mmol). 3. Dissolve 620 mg dicyclohexylcarbodiimide (3 mmol) in 1 mL tetrahydrofuran and add to the reaction mixture. Stir the resulting solution at room temperature overnight (12 hr). 4. Remove the white precipitate (dicyclohexylurea) by filtration through Whatman no. 1 filter paper in a filter funnel. 5. Optional: Purify filtrate on a 10 × 2–cm silica gel column, eluting with 9:1 (v/v) ethyl acetate/methanol. Collect appropriate fractions (Rf = 0.45) and evaporate on a rotary evaporator using a water aspirator to give a white powder melting at 120°C. Although N-acetoxysuccinimide can be used without purification, it is generally preferable to purify if the product is to be stored. The product may be stored indefinitely under dry nitrogen in an amber bottle.
6. Analyze by MS and by proton NMR spectroscopy. The purified product has the following spectroscopic characteristics: Rf = 0.45 (CH2Cl2, silica) 1
H NMR (500 MHz, CHCl3-d1) d 2.84 (s, CH2 4H), 2.34 (s, CH3 3H)
MS-CI calculated for C6H7NO4: 157; observed m/z: 158 (M + H)+.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX. PalladiumMediated C5 Substitution of Pyrimidine Nucleosides
0.1 M Li2PdCl4 solution 1.77 g PdCl2 (0.01 mol) 0.84 g LiCl (0.02 mol) 70 mL anhydrous methanol Stir at room temperature for 24 hr
1.1.14 Current Protocols in Nucleic Acid Chemistry
Adjust volume to 100 mL with methanol The solution is generally not stored for more than a few weeks in a stoppered flask at room temperature.
COMMENTARY Background Information The C5 position of pyrimidine nucleosides is nearly ideal as a site for tethering molecular reporter groups and other molecular devices to oligodeoxyribonucleotides, as groups of different sizes may be attached without adversely affecting DNA duplex formation. In recent years, a variety of specialized probe moieties such as biotin (Langer et al., 1981; Shimkus et al., 1985; Cook et al., 1988), fluorophores (Prober et al., 1987; Tesler et al., 1989; Hagmar et al., 1995), paramagnetic probes (Spaltenstein et al., 1988, 1989; Kirchner et al., 1990), pendant catalytic moieties (Dreyer and Dervan, 1985; Bashkin et al., 1994; Kwiatkowski et al., 1994; Bergstrom and Chen, 1996; Shah et al., 1996), and cross-linkers (Gibson and Benkovic, 1987; Tabone et al., 1994; Chaudhuri and Kool, 1995; Meyer and Hanna, 1996) have been coupled to deoxyuridine and then incorporated into nucleic acids. The use of C5 linkers to functionalize nucleic acids has been reviewed (Goodchild, 1990). Although many kinds of linkers have been used to attach reporter groups to C5, alkynyl groups appear to be preferable because they enhance duplex stability (Sagi et al., 1993; Ahmadian et al., 1998). Synthesis of 5-carboxamidopropynyl-2′deoxyuridine derivatives was initiated by 4,4′dimethoxytrityl protection of the readily available 5-iodo-2′-deoxyuridine following a standard procedure (Jones, 1984). Tritylation of 5′-hydroxyl of 5-iodo-2′-deoxyuridine prior to the coupling reaction eliminated the need for toluyl protection and deprotection of the nucleoside hydroxyl groups (Robins and Barr, 1981, 1983). Palladium-mediated coupling reactions were carried out in dimethylformamide following a procedure similar to that reported by Hobbs (1989). The steps outlined in the preparation of 5-(3-nicotinamidopropyn-1-yl)-5′-O-(4,4′-dimethoxytrityl)-2′-deoxyuridine provide a blueprint and strategy for the incorporation of many different kinds of reporter groups. As long as the desired reporter group has a reactive acyl functional group (carboxylic acid anhydride, chloride, or active ester), it is likely that it can be coupled to propargylamine and sub-
sequently linked to deoxyuridine by the organopalladium coupling reaction. The thioether-linked deoxyuridine derivative 5-[3-(trifluoroacetamido)-1-thiapropyl]2′-deoxyuridine was synthesized by the procedure reported by Bergstrom et al. (Bergstrom et al., 1991; Ahmadian et al., 1998) via a palladium-mediated reaction of the disulfide N,N′bis(trifluoroacetyl)cystamine with 5-chloromercurio-2′-deoxyuridine. Trifluoroacetyl functions as a base-sensitive protecting group that can be easily removed with aqueous ammonia. Because the trifluoroacetamido group is a poor ligand for palladium, it does not interfere with the palladium-mediated coupling reaction. Groups that are substantially more electron rich (such as acetamido) interfere with the coupling reaction. For this reason, the coupling reaction must be carried out prior to attaching electron-rich ligands to the cystamine amino group. 5-[3-(Trifluoroacetamido)-1-thiapropyl]-2′deoxyuridine may be incorporated into oligonucleotides as its 5′-O-dimethoxytrityl-3′phosphoramidite derivative (procedure not given). The trifluoroacetyl protecting group is subsequently cleaved during the final ammonia deprotection of the oligonucleotide. This leaves the amino group available for conjugation to reporter groups or molecular tools at the oligonucleotide stage. Alternatively, 5-[3-(trifluoroacetamido)-1-thiapropyl]-2′-deoxyuridine may be deprotected and conjugated to the desired reporter groups or molecular tool prior to transformation to the 5′-O-dimethoxytrityl3′-phosphoramidite derivative and incorporation into the oligonucleotide. Synthesis of 5-[3-(trifluoroacetamido)-1thiapropyl]-2′-deoxyuridine requires 5-chloromercurio-2′-deoxyuridine. This intermediate is obtained in high yield by direct electrophilic mercuration of 2′-deoxyuridine with mercuric acetate in aqueous solution (Bergstrom and Ruth, 1977). An important difference between the synthesis of the alkynyl-linked dU and the thioether-linked dU is that the former requires use of a protected dU and must be done with complete exclusion of oxygen, while the latter is not air sensitive and does not require protecting groups.
Synthesis of Modified Nucleosides
1.1.15 Current Protocols in Nucleic Acid Chemistry
Table 1.1.1
Estimated Times for Completion of Syntheses
Protocol Basic Protocol 1
Time (hr) 18 20 15
Basic Protocol 2
Support Protocol 1 Support Protocol 2
16 21 32 36 14
Synthesis N-(3-Propyn-1-yl)nicotinamide 5′-O-(4,4′-Dimethoxytrityl)-5-iodo-2′-deoxyuridine 5-(3-Nicotinamidopropyn-1-yl)-5′-O-(4,4′-dimethoxytrityl)2′-deoxyuridine 5-Chloromercurio-2′-deoxyuridine 5-[3-(Trifluoroacetamido)-1-thiapropyl]-2′-deoxyuridine 5-(3-Acetamido-1-thiapropyl)-2′-deoxyuridine N,N′-Bis(trifluoroacetyl)cystamine N-Acetoxysuccinimide
There are alternative types of linkers as well as alternative methods for preparing alkynyl and thioalkyl linkers. In addition to the alkynyl and thioether linkages described here, alkyl and alkenyl linkers may also be obtained through organopalladium coupling methodology (Bergstrom and Ogawa, 1978). Examples of reporter groups linked through alkyl and alkenyl linkers are included in Literature Cited. The advantage of the alkynyl linker is the ease of preparation of the C5-substituted deoxyuridine that contains the 5′ protecting group (dimethoxytrityl) needed for subsequent oligonucleotide synthesis via the phosphoramidite methodology. In addition to the nicotinamidopropyne coupling reaction, the preparation of a series of other carboxamidopropynyl derivatives has been described (Ahmadian et al., 1998). Since organopalladium reactions can tolerate a wide variety of functional groups (e.g., hydroxyl, amido, carboxamide, ester, cyano, nitro, keto), there may be relatively few limitations on the nature of the group that can be introduced at C5 as a component of carboxamidopropyne. Deoxyuridine C5 substitution is preferred over deoxycytidine N4 or deoxyadenosine N6 substitution because the latter two modifications destabilize doublestranded DNA.
Critical Parameters
PalladiumMediated C5 Substitution of Pyrimidine Nucleosides
The most critical parameter in each reaction is the purity of the reactants and the reagents. Even fresh commercial reagents should be checked by TLC and 1H NMR for purity and identity. 4,4′-Dimethoxytrityl chloride is sensitive to moisture and will deteriorate over time. The best results are obtained with a freshly opened bottle of the reagent. Alternatively, the authors store and transfer 4,4′-dimethoxytrityl chloride in a Vacuum Atmospheres dry box under a dry nitrogen atmosphere (any dry box
should be suitable). The condensation reaction between 5′-O-(4,4′-dimethoxytrityl)-5-iodo2′-deoxyuridine and the alkyne requires airsensitive tetrakis(triphenylphosphine)palladium. This reagent should also be stored and transferred under nitrogen or argon. Again, it is preferable to use freshly opened reagents, as they are supplied in sealed glass ampules that may be difficult to keep free of oxygen once opened. When planning the construction of a C5modified nucleoside for introduction into oligonucleotides, the intermediates must not contain functional groups that are likely to interfere either with phosphoramidite preparation (e.g., hydroxyl) or oligonucleotide synthesis.
Anticipated Results If the procedures are followed as described in this unit, the yields of isolated product should be comparable to that reported here. The palladium-catalyzed coupling reactions are especially sensitive to reagents and conditions. If an alkyne other than the one described in the protocol is used for the procedure, it may be necessary to try palladium catalysts other than tetrakis(triphenylphosophine)palladium. If the products are to be used to construct phosphoramidites for oligonucleotide synthesis, 50 mg of the C5-substituted 4,4′-dimethoxytrityl (DMTr) derivative is generally sufficient to obtain enough product to accomplish one coupling reaction on a 1-µmol synthesis scale.
Time Considerations The time required to complete each procedure is summarized in Table 1.1.1. The estimated times do not include the time necessary to purify solvents. In some cases, these may be purchased and used without further purification.
1.1.16 Current Protocols in Nucleic Acid Chemistry
Literature Cited Ahmadian, M., Zhang, P., and Bergstrom, D.E. 1998. A comparative study of the thermal stability of oligodeoxyribonucleotides containing 5substituted-2′-deoxyuridines. Nucl. Acids Res. 26:3127-3135. Bashkin, J.K., Sondhi, S.M., Sampath, U., d’Avignon, D.A., and Modak, A.S. 1994. Synthesis and connectivity assignment (by 2D-NMR) of a nucleoside-dipeptide: 5-[3-[[2-[[2-[[[2-Amino]1-oxo-3-[1H-imidazol-4-yl]propyl]amino]-1oxo-3-[1H-imidazol-4-yl]propyl]amino]ethyl] amino]-3-oxopropyl]-2′-deoxyuridine. New J. Chem. 18:305-318. Bergstrom, D.E. and Chen, J. 1996. Sequence-specific oligodeoxyribonucletide cleavage by a major-groove-positioned metal-binding ligand tethered to C-5 of deoxyuridine. Bioorg. Med. Chem. Lett. 6:2211-2214. Bergstrom, D.E. and Ogawa, M.K. 1978. C-5 substituted pyrimidine nucleosides. 2. Synthesis via olefin coupling to organopalladium intermediates derived from uridine and 2′-deoxyuridine. J. Am. Chem. Soc. 100:8106-8112. Bergstrom, D.E. and Ruth, J.L. 1977. Preparation of C-5 mercurated pyrimidine nucleosides. J. Carbohydrates Nucleotides Nucleosides 42:257269. Bergstrom, D.E., Beal, P., Jenson, J., and Lin, X. 1991. Palladium-mediated synthesis of C-5 pyrimidine nucleoside thioethers from disulfides and mercurinucleosides. J. Org. Chem. 56:5598-5602. Chaudhuri, N.C. and Kool, E.T. 1995. Very high affinity DNA recognition by bicyclic and crosslinked oligonucleotides. J. Am. Chem. Soc. 117:10434-10442. Cook, A.F., Vuocolo, E., and Brakel, C.L. 1988. Synthesis and hybridization of a series of biotinylated oligonucleotides. Nucl. Acids Res. 16:4077-4095. Dreyer, G.B. and Dervan, P.B. 1985. Sequence-specific cleavage of single-stranded DNA: Oligodeoxynucleotide-EDTA.Fe(II). Proc. Natl. Acad. Sci. U.S.A. 82:968-972. Gibson, K.J. and Benkovic, S.J. 1987. Synthesis and application of derivatizable oligonucleotides. Nucl. Acids Res. 15:6455-6467. Goodchild, J. 1990. Conjugates of oligonucleotides and modified oligonucleotides: A review of their synthesis and properties. Bioconjugate Chem. 1:165-187. Hagmar, P., Bailey, M., Tong, G., Haralambidis, J., Sawyer, W.H., and Davidson, B.E. 1995. Synthesis and characterization of fluorescent oligonucleotides. Effect of internal labelling on protein recognition. Biochim. Biophys. Acta 1244:259-268. Hobbs, F.W. Jr. 1989. Palladium-catalyzed synthesis of alkynylamino nucleosides. A universal linker for nucleic acids. J. Org. Chem. 54:3420-3422. Jones, R.A. 1984. Preparation of protected deoxyribonucleosides. In Oligonucleotide Synthesis: A Practical Approach (M.J. Gait, ed.) pp. 27-28. IRL Press, Washington, D.C.
Kirchner, J.J., Hustedt, E.J., Robinson, B.H., and Hopkins, P.B. 1990. DNA dynamics from a spin probe: Dependence of probe motion on tether length. Tetrahedron Lett. 31:593-596. Kwiatkowski, M., Samiotaki, M., Lamminmaki, U., Mukkala, V.-M., and Landegren, U. 1994. Solidphase synthesis of chelate-labelled oligonucleoties: Application in triple-color ligase-mediated gene analysis. Nucl. Acids Res. 22:2604-2611. Langer, P.R., Waldrop, A.A., and Ward, D.C. 1981. Enzymatic synthesis of biotin-labeled polynucleotides: Novel nucleic acid affinity probes. Proc. Natl. Acad. Sci. U.S.A. 78:66336637. Meyer, K.L. and Hanna, M.M. 1996. Synthesis and characterization of a new 5-thiol-protected deoxyuridine for site-specific modification of DNA. Bioconjugate Chem. 7:401-412. Prober, J.M., Trainor, G.L., Dam, R.J., Hobbs, F.W., Robertson, C.W., Zagursky, R.J., Cocuzza, A.J., Jensen, M.A., and Baumeister, K. 1987. A system for rapid DNA sequencing with fluorescent chain-terminating dideoxynucleotides. Science 238:336-341. Robins, M.J. and Barr, P.J. 1981. Nucleic acid related compounds. 31. Smooth and efficient palladium-copper catalyzed coupling of terminal alkynes with 5-iodouracil nucleosides. Tetrahedron Lett. 22:421-424. Robins, M.J. and Barr, P.J. 1983. Nucleic acid related compounds. 39. Efficient conversation of 5-iodo to 5-alkynyl and derived 5-substituted uracil bases and nucleosides. J. Org. Chem. 48:1854-1862. Sagi, J., Szemzo, A., Ebinger, K., Szabolcs, A., Sagi, G., Ruff, E., and Otvos, L. 1993. Basemodified oligodeoxynucleotides. I. Effect of 5-alkyl, 5-(1-alkenyl) and 5-(1-alkynyl) substitution of the pyrimidines on duplex stability a n d h y d r o p h o b i c i t y. Tetrahedron Lett. 34:2191-2194. Shah, K., Neenhold, H., Wang, Z., and Rana, T.M. 1996. Incorporation of an artificial protease and nuclease at the HIV-1 Tat binding site of transactivation responsive RNA. Bioconjugate Chem. 7:283-289. Shimkus, M., Levy, J., and Herman, T. 1985. A chemically cleavable biotinylated nucleotide: Usefulness in the recovery of protein-DNA complexes from avidin affinity columns. Proc. Natl. Acad. Sci. U.S.A. 82:2593-2597. Spaltenstein, A., Robinson, B.H., and Hopkins, P.B. 1988. A rigid and nonperturbing probe for duplex DNA motion. J. Am. Chem. Soc. 110:12991301. Spaltenstein, A., Robinson, B.H., and Hopkins, P.B. 1989. Sequence- and structure-dependent DNA base dynamics: Synthesis, structure, and dynamics of site and sequence specifically spin-labeled DNA. Biochemistry 28:9484-9495.
Synthesis of Modified Nucleosides
1.1.17 Current Protocols in Nucleic Acid Chemistry
Tabone, J.C., Stamm, M.R., Gamper, H.B., and Meyer, R.B. Jr. 1994. Factors influencing the extent and regiospecificity of cross-link formation between single-stranded DNA and reactive complementary oligodeoxynucleotides. Biochemistry 33:375-383. Tesler, J., Cruickshank, K.A., Morrison, L.E., and Netzel, T. 1989. Synthesis and characterization of DNA oligomers and duplex containing covalently attached molecular labels: Comparison of biotin, fluorescin, and pyrene labels by thermodynamic and optical spectroscopic measurements. J. Am. Chem. Soc. 111:6966-6976.
Contributed by Mohammad Ahmadian Cerus Corp. Concord, California Douglas A. Klewer Texas A&M University College Station, Texas Donald E. Bergstrom Purdue University West Lafayette, Indiana
PalladiumMediated C5 Substitution of Pyrimidine Nucleosides
1.1.18 Current Protocols in Nucleic Acid Chemistry
Enzymatic Synthesis of M1G-Deoxyribose
UNIT 1.2
M1G-deoxyribose (M1G-dR, 1,N2-pyrimido[1,2-α]purin-10(3H)-one, or pyrimidopurinone; see structure in Fig. 1.2.2, below) is an endogenous exocyclic DNA adduct formed by the reaction of the dicarbonyl compound malondialdehyde (MDA) with a deoxyguanosine residue in DNA. M1G-dR is an intermediate in the synthesis of a class of modified oligodeoxyribonucleotides that are used to study the mutagenicity and repair of M1G. This unit presents two methods for synthesizing M1G-dR using enzymatic coupling. The Basic Protocol describes a procedure for coupling the nucleobase to deoxyribose, in a reaction mediated by the enzyme nucleoside 2′-deoxyribosyltransferase, followed by preparation of the modified base (see Fig. 1.2.1A). Preparation of the enzyme is described in the Support Protocol. The Alternate Protocol uses two commercially available enzymes, purine nucleoside phosphorylase and thymidine phosphorylase (see Fig. 1.2.1B). Although the enzyme preparation step is avoided, additional purification steps are required that increase the time needed to complete the synthesis and decrease the yield (see Commentary). NOTE: Use deionized, distilled water in all recipes and protocol steps.
BASIC PROTOCOL
ENZYMATIC COUPLING USING NUCLEOSIDE 2′-DEOXYRIBOSYLTRANSFERASE Nucleoside 2′-deoxyribosyltransferase (trans-N-deoxyribosylase or nucleoside:purine(pyrimidine) deoxyribosyltransferase; E.C. 2.4.2.6) catalyzes the transfer of the deoxyribosyl moiety from a deoxyribonucleoside to any other nucleoside base (see Fig. 1.2.1A). This enzyme is found exclusively in Lactobacilli and related microorganisms that require deoxynucleosides for growth (Carson and Wasson, 1988). The enzyme’s broad specificity makes it a useful tool for synthesizing modified deoxyribonucleotides. This protocol describes the use of this enzyme (see Support Protocol for preparation) to transfer a deoxyriboside from 2′-deoxycytidine to M1G and produce M1G-dR. The two synthesis steps can be carried out in a single flask, which decreases the time needed to purify M1G-dR and significantly increases the yield of the reaction. A
C
HO
O
HO
transferase
HO
O
M1G
HO
M1G
C
B T
HO
O HO
HO
TPase
O HO
HPO42-
T
HO
PNPase
O
M1G
OPO32M1G
HPO42-
HO
Figure 1.2.1 Enzymatic coupling reactions. (A) Reaction catalyzed by nucleoside 2′-deoxyribosyltransferase (Basic Protocol). (B) Reaction catalysed by thymidine phosphorylase and purine nucleoside phosphorylase (Alternate Protocol). Contributed by Nathalie C. Schnetz-Boutaud, Marie-Christine Chapeau, and Lawrence J. Marnett Current Protocols in Nucleic Acid Chemistry (2000) 1.2.1-1.2.8 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Modified Nucleosides
1.2.1
Materials Guanine hydrochloride (Sigma) 1 N HCl Tetraethoxypropane (Aldrich) Methanol (MeOH; Aldrich) Potassium carbonate (Aldrich) Nanopure water (water purified using Nanopure system from Barnstead/Thermolyne) MES (2-[N-morpholino]ethanesulfonic acid; Sigma) 2′-Deoxycytidine (dC; Sigma) 1 N NaOH Nucleoside 2′-deoxyribosyltransferase (transferase; see Support Protocol) Dichloromethane (CH2Cl2; Fisher) 250-mL round-bottom flask Oil bath, 70°C Magnetic stir plate and stir bar Ice bath pH indicator strips Büchner funnel Whatman No. 1 filter paper Shaking incubator, 37°C Silica-gel thin-layer chromatography (TLC) plates Lyophilizer Silica gel (60 to 100 mesh; Fisher) 8 × 50–cm chromatography column Prepare modified base 1. In a 250-mL round-bottom flask, dissolve 1 g (5.3 mmol) guanine hydrochloride in 100 mL of 1 N HCl that has been heated to 70°C using an oil bath. Stir on a magnetic stir plate until dissolved. The dissolution can take from 30 min to 1 hr.
2. Mix 1.3 mL (5.86 mmol) tetraethoxypropane with 1.25 mL methanol. Add this dropwise to the solution from step 1. Slow addition of tetraethoxypropane favors the formation of the modified base and avoids the polymerization of MDA.
3. Let the reaction mixture stir for 30 min on a magnetic stir plate, then cool to 0°C in an ice bath. Cooling moderates the neutralization reaction to follow.
4. Neutralize by slowly adding potassium carbonate to pH 6, verifying the pH using pH indicator strips. Take special care when approaching the desired pH. 5. Remove unreacted guanine by filtering through Whatman no. 1 filter paper on a Büchner funnel under vacuum. Wash the precipitate twice with 20 mL of Nanopure water. The filtrate contains the modified base. The yield of the reaction is estimated at 30%. Enzymatic Synthesis of M1G-Deoxyribose
6. Add MES to a final concentration of 0.5 M. MES is used to buffer the enzymatic reaction.
1.2.2 Current Protocols in Nucleic Acid Chemistry
7. Add 1.2 g (5.3 mmol) 2′-deoxycytidine. 8. Equilibrate the solution to pH 6.0 with 1 N HCl or 1 N NaOH.
Perform enzymatic coupling 9. Add an appropriate amount of nucleoside 2′-deoxyribosyltransferase solution and incubate overnight at 37°C with shaking. Each enzymatic preparation has its own concentration and activity, which must be tested empirically by assaying the preparation’s ability to convert a small quantity of M1G to M1GdR to estimate the amount to use for the full-scale reaction.
10. Check the progress of the reaction by TLC on silica-gel plates using 9:1 (v/v) CH2Cl2/MeOH as the mobile phase. Add 50 mL methanol to 50 mL of reaction mix and spot on TLC plate using a capillary tube. If the reaction is not complete, add more enzyme and dC, but not more than one-fifth the amount used at the start of the reaction, and incubate an additional 6 to 12 hours.
11. Once the reaction is complete, lyophilize the reaction mixture to dryness. 12. Purify the crude product on a silica-gel column using 9:1 (v/v) CH2Cl2/MeOH for equilibration and elution. Elute by gravity and collect 50-mL fractions. The lyophilized reaction mixture may be added directly to the top of the packed column.
13. Confirm the purity of the product by 1H NMR. Store the product under nitrogen at –20°C. Under these conditions, it is stable for several years. A typical spectrum is presented in Figure 1.2.2. The estimated yield is 10% to 15%. O N N
HO
O
8 7
N N
N
6
HO
Figure 1.2.2
1
H NMR spectrum of M 1-GdR in D2O.
Synthesis of Modified Nucleosides
1.2.3 Current Protocols in Nucleic Acid Chemistry
SUPPORT PROTOCOL
PREPARATION OF NUCLEOSIDE 2′-DEOXYRIBOSYLTRANSFERASE Nucleoside 2′-deoxyribosyltransferase was first isolated by McNutt (1952). Partial purification of the enzyme from Lactobacillus leichmannii has been described by Beck and Levin (1963), and its complete purification and crystallization by Uerkvitz (1971). This protocol is based upon the latter method. The partial purification described here is sufficient for obtaining enzyme to be used in the Basic Protocol. Materials Lactobacillus broth AOAC (see recipe) Lactobacillus helveticus culture 0.15 M NaCl (4°C) 50 mM potassium phosphate buffers, pH 6.0 and 6.9 (see APPENDIX 2A; dilute with Nanopure water to desired molarity) 50 mM potassium phosphate, pH 5.1, containing 10 g/L NaCl 250-mL Erlenmeyer flask Centrifuge and rotors (e.g., Sorvall GS-3 and SS-34) Microtip sonicator (Virsonic 100) BCA Protein Assay (Pierce; optional) or equivalent Purify the enzyme 1. Using aseptic technique, place 25 mL of Lactobacillus broth in an Erlenmeyer flask. Add a loopful of Lactobacillus helveticus commercial stock and incubate overnight at 37°C without shaking. 2. Transfer the 25-mL culture into 1000 mL of fresh Lactobacillus broth. Grow 18 hr at 37°C without shaking. 3. Cool to 4°C and divide into centrifuge tubes. Centrifuge the tubes 10 min at 7000 × g (6500 rpm in a GS-3 rotor), 4°C. 4. Resuspend pellets in 100 mL cold 0.15 M NaCl and centrifuge again as in step 3. Repeat. 5. Suspend cell pellets in a total of 20 mL of 50 mM potassium phosphate, pH 6.0. 6. Sonicate 10 times for 1 min each time with a Virsonic 100 microtip sonicator at a setting of 4 to 5. 7. Centrifuge 30 min at 28,000 × g (15,000 rpm in an SS-34 rotor), 4°C. Save the supernatant. 8. Wash the pellets twice by resuspending in a minimal volume of 0.15 M NaCl and centrifuging as in step 7. Save the supernatants. 9. Combine the supernatants from the previous two steps, and dialyze overnight against 4 L of 50 mM potassium phosphate, pH 5.1, containing 10 g/L NaCl. 10. Heat 10 min at 55°C, then immediately place on ice. The heating denatures all heat-sensitive proteins that are present in the extract, thereby enriching for transferase, which is not heat sensitive.
11. Centrifuge again as in step 7. Enzymatic Synthesis of M1G-Deoxyribose
12. Collect the supernatant and dialyze overnight in 50 mM potassium phosphate, pH 6.9.
1.2.4 Current Protocols in Nucleic Acid Chemistry
To quantify the amount of total protein purified, the BCA Protein Assay (Pierce) can be used. For the purpose of this experiment, this level of purification is sufficient. Store the protein in aliquots at −20°C or proceed to coupling experiments (see Basic Protocol).
Assay the enzyme 13. Prepare five 5-mL aliquots of filtrate containing M1G (see Basic Protocol, step 8). 14. Add to the filtrate 50, 100, 150, 200, and 250 µL of supernatant containing the enzyme (from step 12). 15. Incubate overnight at 37°C with shaking. 16. Spot 10 µL of each reaction mixture three times on a TLC plate using 9:1 (v/v) CH2Cl2/MeOH as the mobile phase. Determine presence of M1G-dR. ENZYMATIC COUPLING USING PURINE NUCLEOSIDE PHOSPHORYLASE AND THYMIDINE PHOSPHORYLASE
ALTERNATE PROTOCOL
Purine nucleoside phosphorylase (PNPase; E.C. 2.4.2.1) catalyzes the displacement of phosphate from deoxyribose-1-phosphate on purines and purine analogs. The stereochemistry of purine attachment produces the naturally occurring β-isomers. Although ribose-1-phosphate is commercially available, it may be more conveniently and cost-effectively generated in situ by thymidine phosphorylase (TPase; E.C. 2.4.2.4)–catalyzed phosphorolysis of thymidine (see Fig. 1.2.1). This Alternate Protocol can be used to avoid the transferase preparation required for the Basic Protocol. First the modified base is synthesized and purified, then the enzymatic coupling is initiated, and finally the product is separated on a medium-performance liquid chromatography (MPLC) column. Due to the instability of M 1G, only 2% to 5% of product is recovered, regardless of the yields achieved in the transribosylation step. Additional Materials (also see Basic Protocol) Thymidine (e.g., Sigma) Purine nucleoside phosphorylase (PNPase; Sigma) Thymidine phosphorylase (TPase; Sigma) 20 mM potassium phosphate, pH 7.3 (APPENDIX 2A) MPLC buffer: 20% methanol in water UV lamp (254 and 365 nm) mPLC column (Baker C18-40 µm, 30 × 500 mm) Prepare modified base 1. Prepare modified base (see Basic Protocol, steps 1 to 5). 2. Concentrate the filtrate under vacuum. 3. Prepare a slurry of the residue with ∼10 g silica gel. 4. Purify by chromatography on a silica-gel column using 9:1 (v/v) CH2Cl2/MeOH for equilibration and elution. 5. Collect and combine the fractions containing M1G as determined by UV fluorescence. M1G base is fluorescent under the long-wavelength (365 nm) of the UV lamp.
Synthesis of Modified Nucleosides
1.2.5 Current Protocols in Nucleic Acid Chemistry
6. Evaporate under vacuum and store under nitrogen at –20°C. 7. Verify the purity of M1G base by 1H NMR. The estimated yield is 2% to 5%.
Perform enzymatic coupling 8. Dissolve 37 mg (0.2 mmol) M1G base and 73 mg (0.6 mmol) thymidine in 200 mL of 20 mM potassium phosphate buffer. Adjust the pH to 7.3 with 1 N HCl or 1 N NaOH. 9. Add 20 U TPase and 30 U PNPase. 10. Incubate the solution 18 hr at 37° to 39°C with shaking. Verify the completion of the enzymatic coupling by thin-layer chromatography (see Basic Protocol, step 10). If the reaction is not complete, continue the purification without further incubation.
11. Concentrate the reaction under vacuum. This and all subsequent vacuum steps may be performed using a side-arm flask with water aspiration.
12. Prepare a slurry of the residue with ∼5 g silica gel. 13. Purify by chromatography on a silica-gel column using 9:1 (v/v) CH2Cl2/MeOH for equilibration and elution. Elute by gravity and collect 50-mL fractions. 14. Collect the fractions containing M1G-dR as determined by yellow fluorescence at 365 nm. M1G and M1G-dR have different elution times.
15. Evaporate to dryness under vacuum. Redissolve two aliquots in 2 mL MPLC buffer each. 16. Purify by MPLC on a Baker C18 40 µm, 30 × 500–mm column, eluting with 20% methanol in water (isocratic) at a flow rate of 3 mL/min. This second column is necessary to separate thymidine from M1G-dR.
17. Collect the fluorescent fractions and evaporate to dryness under vacuum. 18. Verify the purity of the product by 1H NMR. A typical spectrum is presented in Figure 1.2.2 (see Basic Protocol). The estimated yield is 50% to 60%.
REAGENTS AND SOLUTIONS Use Nanopure water (water purified using Nanopure system from Barnstead/Thermolyne) where indicated, and deionized, distilled water in all other recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Enzymatic Synthesis of M1G-Deoxyribose
Lactobacillus broth AOAC Mix 38 g of Lactobacillus broth AOAC (Difco) into 1000 mL Nanopure water. Heat to boiling for 2 min. Autoclave 30 min and allow to cool to room temperature. Prepare fresh for each run.
1.2.6 Current Protocols in Nucleic Acid Chemistry
COMMENTARY Background Information Adducts formed between electrophiles and nucleic acid bases are believed to play a key role in chemically induced mutations and cancer (Singer and Grunnenberger, 1983). Chemical synthesis of deoxynucleoside adducts provides not only authentic standards for comparison to biologically derived materials but also reagents for the synthesis of adducted nucleotides (Basu and Essigmann, 1988). The preparation of certain classes of deoxynucleoside adducts is problematic because of the instability of the intermediates under the conditions used for synthetic manipulations (e.g., the acid lability of purine deoxyribosides). Synthetic approaches to preparing sensitive deoxynucleosides include coupling of adducted bases to activated deoxyribose derivatives (Srivasta et al., 1988) and attachment of deoxyribose moieties to modified bases (Garner and Ramakanth, 1988). However, there are inherent difficulties in controlling both the regiochemistry (e.g., to achieve attack at the N7 versus N9 atom of a purine) and the stereoselectivity (e.g., an SN2 attack on a sugar isomer is needed to generate the desired linkage). Chemical synthesis increases the difficulty of obtaining the correct regioselectivity, whereas enzymatic coupling on nonmodified bases generates only one regioselective isomer. Enzymatic coupling of purine or pyrimidine analogs to deoxyribose has been used to synthesize a number of compounds, including isotopically substituted deoxynucleosides, antitumor agents, and biological active molecules (Holy and Votruba, 1987; Krenitsky et al., 1981, 1986; Muller et al., 1996). When PNPase and TPase are used on M1G (see Alternate Protocol), N9 linkage is shown to be preferred over N7. With the use of transferase (see Basic Protocol), complete regioselectivity (for the N9 isomer only) is obtained.
the Alternate Protocol, which uses a combination of two commercially available enzymes, is more demanding in terms of time and purification. M1G base must be purified; moreover, when the crude mixture is reacted with the combination of PNPase and TPase, transribosylation is inefficient. Also, due to the close polarity of thymidine and M1G-dR (in a variety of solvents system used), a combination reversed-phase and straight-phase column must be employed to obtain pure material. The transferase used in the Basic Protocol and prepared in the Support Protocol has a higher selectivity and produces only the N9 isomer. The two-enzyme method of the Alternate Protocol produces a mixture of N7 and N9, increasing the difficulty of purification. Once a stock solution of transferase is prepared, the Basic Protocol is much easier to perform, less time-consuming, and gives better yields.
Anticipated Results The two protocols described in this unit allow preparation of the desired modified nucleoside in good yields. The Basic Protocol should give yields between 10% and 15% and the Alternate Protocol gives between 2% and 5%. Figure 1.2.2 shows the 1H NMR spectrum of M1G-dR.
Time Considerations Preparation of transferase (see Support Protocol) may take 3 to 4 days. The one-step condensation procedure using transferase (see Basic Protocol), from the synthesis of the modified base to the purification of the nucleoside, should take 24 hr with lyophylization. The alternative method using commercial phosphorylases (see Alternate Protocol) should take 2 or 3 days.
Literature Cited Critical Parameters Since the pyrimidopurinone M1G-dR synthesized in these procedures is base labile, the pH must stay below 7.5 for all protocols. M1G base is even less stable than M1G-dR. The Basic Protocol does not require purification of the modified base; the transferase can be added to the crude mixture. The desired nucleoside can be purified by simple column chromatography. This strategy not only shortens the time required for the synthesis, but also significantly improves the yield. In comparison
Basu, A.K. and Essigmann, J.M. 1988. Site-specifically modified oligonucleotides as probes for the structural and biological effects of DNA-damaging agents. Chem. Res. Toxicol. 1:1-18. Beck, W.S. and Levin, M. 1963. Purification, kinetics, and repression control of bacterial trans-Ndeoxyribosylase. J. Biol. Chem. 238:702. Carson, D.A. and Wasson, D.B. 1988. Synthesis of 2′,3′-dideoxynucleosides by enzymatic transglycosylation. Biochem. Biophys. Res. Comm. 155:829-834. Garner, P. and Ramakanth, S. 1988. A regiocontrolled synthesis of N7- and N9-guanine nucleosides. J. Org. Chem. 53:1294-1298.
Synthesis of Modified Nucleosides
1.2.7 Current Protocols in Nucleic Acid Chemistry
Holy, A. and Votruba, I. 1987. Facile preparation of purine and pyrimidine 2-deoxy-β-D-ribonucleosides by biotransformation on encapsulated cells. Seventh symposium on the Chemistry of Nucleic Acid Components August 30–September 5, 1987. Nucleic Acids Symp. Ser. 18:69-72. Krenitsky, T.A., Kozallka, G.W., and Tuttle, J.V. 1981. Purine nucleoside synthesis, an efficient method employing nucleoside phosphorylases. Biochemistry 20:3615-3621. Krenitsky, T.A., Rideout, J.L., Chao, E.Y., Koszalka, G.W., Gurney, F., Crouch, R.C., Cohn, N.K., Wolberg, G., and Vinegar, R. 1986. Imidazo[4,5c]pyridines (3-deazapurines) and their nucleosides as immunosuppresive and antiinflammatory agents. J. Med. Chem. 29:138-143. McNutt, W.S. 1952. The enzymically catalysed transfer of the deoxyribosyl group from one purine or pyrimidine to another. Biochem. J. 50:384. Muller, M., Hutchinson, L.K., and Guengerich, F.P. 1996. Addition of deoxyribose to guanine and modified DNA bases by Lactobacillus helveticus trans-N-deoxyribosylase. Chem. Res. Toxicol. 9:1140-1144.
Singer, B. and Grunnenberger, D. 1983. Molecular Biology of Mutagens and Carcinogens. Plenum, New York. Srivasta, P.C., Robins, R.K., and Meyer, R.B. 1988. Synthesis and properties of purine nucleosides and nucleotides. In Chemistry of Nucleosides and Nucleotides (L.B. Townsend, ed.) pp. 113281. Plenum, New York. Uerkvitz, W. 1971. Purification of nucleoside 2-deoxyribosyltransferase from Lactobacillus helveticus. Eur. J. Biochem. 23:387-395.
Key Reference Uerkvitz, 1971. See above. Decribes the purification and crystallization of the transferase.
Contributed by Nathalie C. Schnetz-Boutaud, Marie-Christine Chapeau, and Lawrence J. Marnett Vanderbilt University Nashville, Tennessee
Enzymatic Synthesis of M1G-Deoxyribose
1.2.8 Current Protocols in Nucleic Acid Chemistry
Synthesis of N2-Substituted Deoxyguanosine Nucleosides from 2-Fluoro-6-O(Trimethylsilylethyl)-2′-Deoxyinosine
UNIT 1.3
This unit describes the synthesis of 2-fluoro-6-O-(trimethylsilylethyl)-2′-deoxyinosine and gives examples of its use for the preparation of N2-substituted deoxyguanosine nucleosides. Such nucleoside derivatives are used for a variety of purposes including chemotherapy, enzyme mechanism studies, nuclear magnetic resonance (NMR) studies (when isotopically labeled), and as synthetic standards for identification of adducts formed by the reaction of DNA with xenobiotics. In addition, the O6-protected 2-fluoro2′-deoxyinosine compounds can be converted to phosphoramidites and used in the synthesis of oligonucleotides, thus allowing substitution reactions to be carried out after oligonucleotide assembly. 2-Halopurine derivatives have been used for many years for the preparation of N2-substituted guanosine derivatives, with the 2-fluoro substituent being the most easily displaced by nucleophiles (Montgomery and Hewson, 1960; Gerster and Robins, 1965, 1966). The 2-fluoro group is introduced by aqueous diazotization of guanosine in the presence of potassium fluoride or fluoroboric acid. However, these conditions are too harsh for 2′-deoxyguanosine and lead to depurination; hence, different synthetic methodology is needed for the deoxynucleoside. The fluorine atom can be introduced successfully by diazotization under anhydrous conditions with t-butyl nitrite as the diazotizing agent and HF in pyridine as the fluoride source (Robins and Uznanski, 1981; Lee et al., 1990; Harris et al., 1991). Success in the fluoridation step requires protection of the C6 oxygen group, which is done by Mitsunobu alkylation (Mitsunobu, 1981) with trimethylsilylethanol or other alcohols. The O6-protecting group also facilitates displacement of the halogen by nucleophiles. Basic Protocol 1 in this unit describes the synthesis of 2-fluoro-6-O-(trimethylsilylethyl)2′-deoxyinosine and comprises three separate procedures: (1) protection of the 2-NH2, 3′-OH, and 5′-OH groups of 2′-deoxyguanosine to make a triacetyl derivative, (2) protection of the O6 group by Mitsunobu alkylation with trimethylsilylethanol, and (3) introduction of the 2-fluoro group. Alternate Protocol 1 describes the preparation and use of 3′,5′-O-diacetyl-2′-deoxyguanosine and its use in the Mitsunobu reaction described in Basic Protocol 1. Alternate Protocol 1 gives lower yields than Basic Protocol 1, but is quicker and is better for the preparation of 6-O-(p-nitrophenethyl)-2′-deoxyguanosine, another commonly used O6-protected derivative. Basic Protocol 2 describes a general procedure for the synthesis of N2-substituted 2′-deoxyguanosines. Two specific examples are then given in Alternate Protocols 2 and 3, which give detailed directions for synthesis using an unhindered diamine to give a derivative with an alkylamine sidechain, and for using an amino alcohol to yield an N2 hydroxyalkenyl derivative. A Support Protocol outlines the procedure for carrying out reactions in an inert atmosphere. CAUTION: Several of the steps in these protocols involve the use of toxic, corrosive, and flammable chemicals. It is highly recommended that all operations be carried out in a fume hood. Good laboratory safety practices should be observed at all times, including the use of safety goggles, a laboratory coat, and disposable gloves. It is recommended that this synthesis be done only by personnel experienced in the handling of reactive and toxic chemicals. Contributed by Thomas M. Harris and Constance M. Harris Current Protocols in Nucleic Acid Chemistry (2000) 1.3.1-1.3.19 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Modified Nucleosides
1.3.1
NOTE: A variety of methods is used in these procedures to remove volatile solvents and reagents. The choice of method depends upon the boiling points of the volatiles, the stability of the products (e.g., sometimes heat cannot be used to hasten evaporation), and the volume to be removed. For additional details, see Critical Parameters and Troubleshooting. BASIC PROTOCOL 1
SYNTHESIS OF 2-FLUORO-6-O-(TRIMETHYLSILYLETHYL)-2′DEOXYINOSINE USING 2-N-3′,5′-O-TRIACETYL-2′-DEOXYGUANOSINE This protocol describes the synthesis of the 2-fluoro derivative of 6-O-(trimethylsilylethyl)2′-deoxyinosine (Fig. 1.3.1). It is divided into three basic procedures. (1) Protection of the 2-NH2, 3′-OH, and 5′-OH groups of 2′-deoxyguanosine (S.1) is performed by acetylation (S.2). (2) Protection of the O6 group is carried out via Mitsunobu alkylation with trimethylsilylethanol, diethyl azodicarboxylate, and triphenylphosphine. Sodium methoxide and methanol are then added to the reaction to remove the acetyl protecting groups, yielding the 6-O-trimethylsilylethyl (TMSE) derivative (S.3). (3) The 2-NH2 group is converted to the 2-fluoro substituent by performing nonaqueous diazotization and fluoridation at low temperature with t-butyl nitrite and HF/pyridine (Robins and Uznanski, 1981). An Alternate Protocol utilizing 3′,5′-O-diacetyl-2′-deoxyguanosine is also described (see Alternate Protocol 1). NOTE: Anhydrous solvents are required for several steps in these procedures. They can be purchased (e.g., from Aldrich in Sure/Seal bottles) or dried by distillation from appropriate desiccants and stored under nitrogen. Materials 2′-Deoxyguanosine (dG) monohydrate Pyridine, anhydrous (Aldrich; packed under nitrogen in a Sure/Seal bottle) Acetic anhydride, freshly distilled Triethylamine (d 0.726), distilled from calcium hydride 4-Dimethylaminopyridine (DMAP)
N
HO
O
N
NH N
NH2
a
N
AcO
O AcO
HO 1
Me3Si
O
O N
NH N
NHAc
O N
b,c
N
N
HO
N
O
NH2
HO 2
3
d
Me3Si
O N N
HO
O
N N
F
HO 4
Synthesis of N2-Substituted Deoxyguanosine Nucleosides
Figure 1.3.1 Synthesis of 2-fluoro-6-O-(trimethylsilylethyl)-2′-deoxyinosine (S.4) using 2-N-3′,5′O-triacetyl-2′-deoxyguanosine (S.2; see Basic Protocol 1). Reagents: (a) acetic anhydride, pyridine, 4-dimethylaminopyridine (steps 1 to 13): (b) triphenylphosphine, diethyl azodicarboxylate, 2-trimethylsilylethanol (steps 14 to 19); (c) sodium methoxide, methanol (steps 20 to 33); (d) HF/pyridine, t-butyl nitrite (steps 34 to 46).
1.3.2 Current Protocols in Nucleic Acid Chemistry
Dry nitrogen (N2) or argon (Ar) Methanol, anhydrous Methylene chloride (CH2Cl2), anhydrous Anisaldehyde/sulfuric acid spray (see recipe) Acetonitrile Dioxane, anhydrous, distilled from sodium metal before use Triphenylphosphine 2-Trimethylsilylethanol Diethyl azodicarboxylate (DEAD; from a fresh, unopened bottle) 0.35 M sodium methoxide in methanol (see recipe) Aqueous acetic acid: 6.2 mL glacial acetic acid in 30 mL water Sodium sulfate (Na2SO4), anhydrous 63- to 200-mesh silica gel Sand Dry ice/acetonitrile cooling bath (−35° to −40°C) 70% HF/pyridine solution (Aldrich) t-Butyl nitrite Potassium carbonate (K2CO3) Ethyl acetate 2-liter round-bottom flask Rotary evaporator equipped with a condenser cooled with chilled water or a dry ice condenser Reflux condenser with 24/40 joint and gas inlet adapter Temperature-controlled oil bath (up to ∼115°C) 0.25-mm silica gel 60F-254 glass thin-layer chromatography (TLC) plates UV light source Vacuum system (oil pump) capable of creating <1 mmHg pressure, with manifold and cold trap Filter paper (Whatman no. 1, 7-cm diameter) Buchner funnel 500-mL and l-liter Erlenmeyer flasks 1-liter, three-neck flask with 24/40 joints (oven dried) and rubber septa 10-mL glass syringes (oven dried) 1-liter, single-neck flask with 24/40 joint Water aspirator 500-mL separatory funnels Heavy-walled glass column (5-cm i.d. × 40-cm length) Abderhalden apparatus (drying pistol; 78°C) 50-mL polypropylene conical tubes with rubber septa 23-G syringe needles 20-mL plastic syringes with 3-in. (7.6-cm) 20-G needles 1-mL glass syringe (oven dried) Additional reagents and equipment for performing reactions under nitrogen (see Support Protocol) Protection of 2-NH2, 3′-OH, and 5′-OH Groups Remove water from dG 1. Place 8.54 g dG (S.1) in a 2-liter round-bottom flask.
Synthesis of Modified Nucleosides
1.3.3 Current Protocols in Nucleic Acid Chemistry
Table 1.3.1
Synthesis of 2-N-3′,5′-O-Triacetyl-2′-Deoxyguanosine (S.2)
Reagent 2′-Deoxyguanosine⋅H2O Pyridine (for coevaporation) Pyridine (for reaction) Acetic anhydride Triethylamine 4-Dimethylaminopyridine (DMAP) Methanol
Amount 8.54 g 450 mL 1200 mL 28 mL 46 mL 0.36 g
MW (g/mol) 285 79 79 102 101 122
Millimoles 30.0 5600 14900 299 330 2.99
Equivalents 1.0 187 497 10 11 0.1
350 mL
32
8640
288
Table 1.3.1 lists quantities of reagents for the synthesis of N2,O3′,O5′-triacetyl-2′-deoxyguanosine (S.2).
2. Add 150 mL anhydrous pyridine and then remove the pyridine under vacuum using a rotary evaporator equipped with a dry ice condenser and connected to an oil pump. The same oil pump used in step 8, capable of creating <1 mmHg pressure, can be used here; however, it is not necessary. For this step, 1 mmHg pressure would be sufficient.
3. Repeat the addition of pyridine and evaporation twice more. Alternatively, the dG can be dried under vacuum in an Abderhalden drying apparatus at 78°C (refluxing ethanol) for 16 hr.
Acetylate dG 4. Add the following reagents (see Table 1.3.1): 1200 mL anhydrous pyridine 28 mL acetic anhydride 46 mL triethylamine 0.36 g DMAP. 5. Place a large magnetic stir bar in the flask, fit the flask with a reflux condenser, and place the flask in a 55°C temperature-controlled oil bath on top of a magnetic stir plate. Stir the reaction at 55°C for 72 hr under nitrogen (see Support Protocol). 6. Monitor the reaction by occasionally removing a small sample (1 to 2 µL) and analyzing it by TLC. Perform TLC on 0.25-mm silica gel 60F-254 glass plates. Elute with 1:9 (v/v) methanol/methylene chloride. Detect product under UV light and also with anisaldehyde/sulfuric acid spray. Spray (or dip) the plate and heat on a hot plate for several seconds. Anisaldehyde/sulfuric acid spray is used for the detection of sugars. Spots should appear in various shades of blue and purple. The Rf of the product is 0.31 and that of the starting material is <0.1. CAUTION: Anisaldehyde/sulfuric acid spray should be used in a well-ventilated fume hood. It is corrosive and the vapors are highly irritating.
Synthesis of N2-Substituted Deoxyguanosine Nucleosides
Work up and purify 2-N-3′,5′-O-triacetyl-2′-deoxyguanosine 7. When TLC analysis indicates that the reaction is complete, cool in an ice bath and add 350 mL methanol. Stir for 5 min and remove solvents under reduced pressure using a rotary evaporator. A red oily residue should be present at this point.
1.3.4 Current Protocols in Nucleic Acid Chemistry
8. Connect the flask to a vacuum system capable of obtaining a vacuum of <1 mmHg and allow to dry overnight. It is important to remove the solvents as thoroughly as possible before recrystallization.
9. Add 100 mL acetonitrile and incubate at −20°C overnight. 10. While the reaction mixture is still cold, collect the resulting crystals by vacuum filtration using Whatman no. 1 filter paper and a Buchner funnel. 11. Rinse the precipitate with cold acetonitrile (4°C) and allow the crystals to air dry. 12. Recrystallize from a minimum amount of hot methanol. Place the solid in a 500-mL Erlenmeyer flask and add a few milliliters of methanol. Heat with gentle shaking on a hot plate or in a steam bath until boiling. Gradually add methanol until all the solid is dissolved. Remove from heat and allow to cool. Recrystallization can easily be carried out on a larger scale.
13. Cool at −20°C overnight, filter as in step 10, and air dry. The yield of compound S.2 should be 80% to 90% (9.5 to 10.5 g). The product can be stored indefinitely at room temperature.
Protection of the O6 Group by Mitsunobu Alkylation Perform Mitsunobu reaction 14. Equip a 1-liter, 24/40 three-neck flask with rubber septa on the two side necks, a 24/40 reflux condenser in the center neck, and a magnetic stir bar. Place a gas inlet adapter on top of the condenser and connect to a nitrogen source (see Support Protocol). 15. Remove one rubber septum, add the following reagents (see Table 1.3.2), and replace the septum. 430 mL dioxane 5.10 g triacetyl deoxyguanosine (S.2) 6.82 g triphenylphosphine. Table 1.3.2 lists quantities of reagents for the synthesis of 6-O-(trimethylsilylethyl)-2′-deoxyguanosine (S.3).
Table 1.3.2
Synthesis of 6-O-TMSE-2′-Deoxyguanosine (S.3) from S.2a
Reagent Dioxane 2-N-3′,5′-O-Triacetyl-dG (S.2) Triphenylphosphine 2-Trimethylsilylethanol Diethyl azodicarboxylate (DEAD) Methanol Sodium methoxide solution (0.35 M) Glacial acetic acidb
Amount 430 mL 5.10 g 6.82 g 3.75 mL 4.10 mL
MW (g/mol) 88 393 262 118 174
Millimoles 5052 13 26 26 26
Equivalents 389 1.00 2 2 2
120 mL 200 mL
32 54
2940 70
227 5
6.2 mL
60
109
8
aFor synthesis from the diacetyl derivative, use 4.56 g S.7 (MW 351) in place of S.2, and substitute 60 mL concentrated
NH4OH for the 200 mL sodium methoxide solution. bDiluted in 30 mL water.
Synthesis of Modified Nucleosides
1.3.5 Current Protocols in Nucleic Acid Chemistry
16. Transfer the reaction flask to a temperature-controlled oil bath placed on a magnetic stir plate. Start water circulating through the condenser and raise the temperature of the oil bath to ∼115°C. The solvent should begin gently refluxing.
17. Simultaneously add 3.75 mL of 2-trimethylsilylethanol with a 10-mL glass syringe through one rubber septum and 4.10 mL DEAD through the other. Best results are obtained when using DEAD from a fresh unopened bottle. The cloudy yellow suspension should become translucent and darker yellow in color.
18. Continue stirring the reaction mixture for 15 min at 100°C, and then cool to room temperature over ∼2 hr. 19. Transfer the solution to a 1-liter single-neck flask. Remove the solvents under reduced pressure using a rotary evaporator with a dry ice condenser and a water aspirator. Remove acetyl protecting groups 20. Redissolve the viscous red residue in 120 mL anhydrous methanol and slowly add 200 mL of 0.35 M sodium methoxide in methanol. Stir at room temperature for 7 hr. 21. Neutralize the reaction by adding 36.2 mL aqueous acetic acid. Stir for 1 hr at room temperature. 22. Check the pH of the solution using pH paper, and adjust to pH 7.0, if necessary. Work up and purify 6-O-(trimethylsilylethyl)-2′-deoxyguanosine 23. Evaporate the solution under reduced pressure as in step 19. Suspend the residue in 100 mL methylene chloride and transfer to a 500-mL separatory funnel. 24. Extract with 40 mL water. Back extract the aqueous layer with five 100-mL aliquots of methylene chloride. Combine the organic layers. 25. Add sufficient anhydrous Na2SO4 to the extract to cover the bottom of the flask. Swirl gently and allow the salt to settle. If the solution looks cloudy, add more Na2SO4. Allow the solution to stand for 30 to 60 min. 26. Filter by gravity through fluted filter paper into a round-bottom flask to remove the drying agent. Evaporate under reduced pressure (see step 19). 27. While the organic extract is drying, pack a 5 × 20–cm silica gel column using 250 g of 63- to 200-mesh silica gel in methylene chloride containing 0.2% triethylamine in a heavy-walled glass column. Column chromatography is carried out by flash chromatography techniques employing heavy-walled glass columns that can be connected to a gas inlet via a ball joint. Air pressure can be applied to the top of the column via the inlet to increase the flow rate.
28. Dissolve the crude product in 10 mL methylene chloride.
Synthesis of N2-Substituted Deoxyguanosine Nucleosides
29. Load the solution on the column and allow the solution to sink to the level of the column bed. 30. Rinse the flask with two 1-mL aliquots of methylene chloride, add each rinse to the bed, and allow them to sink to the level of the bed.
1.3.6 Current Protocols in Nucleic Acid Chemistry
31. Carefully add a 1-cm layer of sand to the top of the column and begin elution with a gradient of CH2Cl2/methanol/triethylamine ranging from 97.8:2.0:0.2 to 94.8:5.0:0.2 (v/v/v). 32. Collect 10-mL fractions and analyze by TLC (step 6; Rf = 0.41). 33. Combine fractions containing pure product in a round-bottom flask and evaporate to dryness with a rotary evaporator and a dry ice condensor, first under reduced pressure using a water evaporator and then under high vacuum using an oil pump. Impure fractions can be combined, dried, and reanalyzed under the same TLC conditions to yield additional pure product. The product is a pale yellow powder. Approximately 4.2 g (>85% yield) of purified 6-O-TMSE-2′-dG (S.3) should be obtained. It can be stored indefinitely at 4° or –20°C. The Mitsunobu reaction is readily scaled up as much as five fold.
Introduction of the 2-Fluoro Group Fluoridate 6-O-TMSE-2′-dG 34. Dry 0.53 g of 6-O-TMSE-2′-dG (S.3) overnight in an Abderhalden apparatus under vacuum at 78°C. The S.3 product should be quite dry after step 33. The weight should change very little after drying in the Abderhalden apparatus. Table 1.3.3 lists quantities of reagents for the synthesis of 2-fluoro-6-O-TMSE-2′-dI (S.4).
35. Transfer 6-O-TMSE-2′-dG to a 50-mL polypropylene conical tube with a stir bar and a rubber septum. Maintain the reaction under nitrogen using a 23-G syringe needle inserted through the septum. 36. Using a 10-mL glass syringe, place 5.4 mL anhydrous pyridine in a second conical tube, similarly equipped. 37. Place both conical tubes in a dry ice/acetonitrile cooling bath (−35° to −40°C). 38. Using a 20-mL plastic syringe with a 3-in. 20-G needle, add 9.6 mL of 70% HF/pyridine solution to the tube containing the anhydrous pyridine over a period of 3 min (final 45% HF). Stir for 15 min at −35° to −40°C. CAUTION: HF is highly corrosive and should be handled with care. The quality of the 70% HF/pyridine solution is critical to the success of this reaction. The color should be no darker than pale amber. Using a solution that is a dark red or brown leads to increased decomposition products.
Table 1.3.3
Synthesis of 2-Fluoro-6-O-TMSE-2′-Deoxyinosine (S.4) from S.3
Reagent 6-O-TMSE-2′-dG (S.3) Pyridine 70% HF/pyridine t-Butyl nitrite K2CO3
Amount 0.53 g 5.4 mL 9.6 mL 0.43 mL 23 g
a230 neutralizing eq.
MW (g/mol) 368 79 20 103 138
Millimoles 1.44 66 ~336 3.60 166
Equivalents 1.0 46 ~233 2.5 115a
Synthesis of Modified Nucleosides
1.3.7 Current Protocols in Nucleic Acid Chemistry
If the solution appears to be freezing, 2 to 4 mL anhydrous toluene can be added to keep the solution homogeneous.
39. Using another 20-mL plastic syringe and 3-in. 20-G needle, slowly transfer the HF/pyridine solution into the tube containing the dried 6-O-TMSE-2′-dG (S.3). Stir for 5 min at −35° to −40°C. 40. Using a 1-mL glass syringe, add 0.43 mL t-butyl nitrite to the reaction over 5 min while maintaining the bath temperature at −35° to −40°C. Stir the red reaction mixture for 25 min. Work up and purify 2-fluoro-6-O-TMSE-2′-dI 41. Dissolve 23 g K2CO3 in 34 mL water in a 1-liter Erlenmeyer flask. Begin stirring vigorously at 0°C (ice bath). 42. Quench the reaction by slowly pouring the reaction mixture (step 40) into the cold, stirring K2CO3 solution. Rinse the reaction tube with ethyl acetate and add the wash to the neutralized solution. 43. Transfer the solution to a 500-mL separatory funnel and extract five times with 40 mL ethyl acetate. Combine the ethyl acetate extracts. 44. Dry over anhydrous Na2SO4, filter to remove the drying agent, and evaporate to a red oil (steps 25 and 26). Place under high vacuum overnight (see step 8). 45. Purify the crude product by flash chromatography (steps 27 to 31) using an ∼2.5 × 15–cm silica gel column (50 g silica gel). Elute with a gradient of CH2Cl2/methanol/triethylamine ranging from 95:4:1 to 90:9:1 (v/v/v). 46. Analyze by TLC and evaporate appropriate fractions to dryness (steps 32 and 33; Rf = 0.36). Approximately 0.50 g (>90%) of 2-fluoro-6-O-(trimethylsilylethyl)-2′-deoxyinosine (S.4) should be obtained. The product can be stored indefinitely at –20°C under anhydrous conditions. NOTE: This reaction can be carried out starting with 6.7 g of S.3 using a 500-mL round-bottomed flask for the reaction. A mixture of 100 mL anhydrous pyridine and 50 mL toluene is used to dissolve the nucleoside before the addition of the HF/pyridine, which is not diluted before addition at −40°C. The yield of purified S.4 should be ∼70%. ALTERNATE PROTOCOL 1
SYNTHESIS OF 2-FLUORO-6-O-(TRIMETHYLSILYLETHYL)-2′DEOXYINOSINE USING 3′,5′-O-DIACETYL-2′-DEOXYGUANOSINE In this procedure, the Mitsunobu reaction can be carried out starting with 3′,5′-O-diacetyl-2′-deoxyguanosine (Fig. 1.3.2; Zajc et al., 1992), which is synthesized by the procedure of Matsuda et al. (1986). Although the yield of the Mitsunobu product (S.3) is not as high when the diacetyl derivative S.7 is used, the overall synthesis is much quicker because the synthesis of the diacetyl derivative takes only ∼1 hr compared to the 72 hr required for the synthesis of 2-N-3′,5′-O-triacetyl-2′-deoxyguanosine (S.2). This procedure is preferred when 6-O-(p-nitrophenethyl)-2′-deoxyinosine is desired instead of compound S.3, because the p-nitrophenethyl group is not stable in the presence of the sodium methoxide used for deacylation of the triacetyl derivative.
Synthesis of N2-Substituted Deoxyguanosine Nucleosides
Additional Materials (also see Basic Protocol 1) Concentrated ammonium hydroxide (NH4OH) 1-liter, three-neck, round-bottom flask with 24/40 joints
1.3.8 Current Protocols in Nucleic Acid Chemistry
N
HO
N
O
NH
NH N
Me3Si
O
O N
NH2
a
N
AcO
N
O
NH2
b,c
1
N
N
HO
AcO
HO
O N N
O
NH2
HO
7
3
d
Me3Si
O N N
HO
N N
F
O
HO
4
Figure 1.3.2 Synthesis of 2-fluoro-6-O-(trimethylsilylethyl)-2′-deoxyinosine (S.4) using 3′,5′-Odiacetyl-2′-deoxyguanosine (S.7; see Alternate Protocol 1). Reagents: (a) acetic anhydride, 4-dimethylaminopyridine, triethylamine (steps 1 to 5); (b) triphenylphosphine, diethyl azodicarboxylate, 2-trimethylsilylethanol (step 6); (c) NH4OH, methanol (steps 7 and 8); (d) HF/pyridine, t-butyl nitrite (step 9).
Protection of 3′- and 5′-OH Groups Remove water from dG 1. Dry 8.56 g dG in an Abderhalden apparatus (drying pistol) under vacuum at 78°C. Table 1.3.4 lists quantities of reagents used for the synthesis of 3′,5′-O-diacetyl-2′-deoxyguanosine (S.7). Water can also be removed by evaporation with pyridine, and this method is appropriate if the reaction is to be carried out in pyridine. Drying with an Abderhalden apparatus has the advantage of not involving solvent; however, there is a limit to the amount of material that can be effectively dried in a drying pistol.
Acetylate dG 2. Place dried dG in a 1-liter, three-neck, round-bottom flask equipped with a large magnetic stir bar, glass stoppers in two side arms, and a gas inlet tube in the center neck. Attach a nitrogen source to the gas inlet tube as described (see Support Protocol). 3. Add the following reagents (see Table 1.3.4) to produce a yellowish suspension: 0.36 g DMAP 325 mL acetonitrile 85 mL triethylamine 82 mL acetic anhydride. 4. Stir for 30 min to 1 hr at room temperature. During this time the suspension will become whiter. Synthesis of Modified Nucleosides
1.3.9 Current Protocols in Nucleic Acid Chemistry
Table 1.3.4
Synthesis of 3′,5′-O-Diacetyl-2′-Deoxyguanosine (S.7)
Reagent 2′-Deoxyguanosine⋅H2O 4-Dimethylaminopyridine (DMAP) Acetonitrile Triethylamine Acetic anhydride
Amount 8.56 g 0.36 g
MW (g/mol) 285 122
Millimoles 32.0 3.2
Equivalents 1.0 0.1
325 mL 85 mL 82 mL
41 101 102
623 610 869
195 19 27
Work up and purify 3′,5′-O-diacetyl-2′-deoxyguanosine 5. Filter the suspension using Whatman no. 1 filter paper, a Buchner funnel, and a water aspirator. Wash the precipitate with acetonitrile and dry overnight in the Abderhalden apparatus (78°C). This should yield 7.5 to 8.0 g (70%) of the S.7 product. If desired, the compound can be recrystallized from water, but this is not necessary. The compound can be stored indefinitely at room temperature.
Protection of the O6 Group by Mitsunobu Alkylation 6. Perform Mitsunobu reaction as described (see Basic Protocol 1, steps 14 to 19), starting with 4.56 g of the 3′,5′-O-diacetyl compound (S.7; Table 1.3.2). 7. Remove acetyl protecting groups as described (see Basic Protocol 1, steps 20 to 22), but use 60 mL concentrated NH4OH instead of 200 mL sodium methoxide (Table 1.3.2). The yield of the Mitsunobu reaction is not as high (∼50%) when starting with the diacetyl compound, but the overall reaction time is shortened considerably.
8. Work up and purify 6-O-TMSE-2′-dG as described (see Basic Protocol 1, steps 23 to 33). The Mitsunobu reaction is readily scaled up as much as five fold.
Introduction of the 2-Fluoro Group 9. Introduce the 2-fluoro group as described (see Basic Protocol 1, steps 34 to 46). BASIC PROTOCOL 2
GENERAL GUIDELINES FOR SYNTHESIS OF N2-SUBSTITUTED NUCLEOSIDES This protocol outlines a general procedure for the preparation of N2-substituted deoxyguanosines (Fig. 1.3.3A) from the 2-fluoro derivative S.4 synthesized using Basic Protocol 1 or Alternate Protocol 1. Alternate Protocols 2 and 3 give detailed directions for preparing specific aminoalkyl and hydroxyalkanyl derivatives. As can be seen in the Alternate Protocols, considerable latitude can be exercised in the relative amounts of diisopropylethylamine (DIEA) and dimethyl sulfoxide (DMSO) that are used; a greater excess of amine can also be employed without harm.
Synthesis of N2-Substituted Deoxyguanosine Nucleosides
Materials 2-Fluoro-6-O-(trimethylsilylethyl)-2′-deoxyinosine (S.4; see Basic Protocol 1 or Alternate Protocol 1) Amine of choice N,N-Diisopropylethylamine (DIEA), anhydrous Dimethylsulfoxide (DMSO), anhydrous, vacuum distilled from calcium hydride
1.3.10 Current Protocols in Nucleic Acid Chemistry
Me3Si N
A
Basic Protocol 2 Steps 1-3
N
N
HO
Me3Si
O
N
O
F
N
O
HO
Basic Protocol 2 Steps 4-8
N
N
HO
a
O
O N
H NR
N N
HO
b or c
N
O
NH H NR
HO
HO 4 Me3Si
B
4
Alternate Protocol 2 Steps 1-5 a
N
Alternate Protocol 2 Steps 6-13
N
N
HO
O
O
N
O
NH2
N H
b
N N
HO
O
Me3Si Alternate Protocol 3 Steps 1-6 a
O
O N
Alternate Protocol 3 Steps 7-9
N
N
HO
NH2
6
5
4
N H
HO
HO
C
NH N
N
O
N H
c
N
NH
N
HO
N
O
OH
N H
OH
HO
HO 7
8
Figure 1.3.3 (A) General guidelines for synthesis of N2-substituted nucleosides (see Basic Protocol 2). (B) Synthesis of 2-N-(3-aminopropyl)-2′-deoxyguanosine (S.6; see Alternate Protocol 2). (C) Synthesis of 2-N-[2(S)-1-hydroxybut-3-en-2-yl]-2′-deoxyguanosine (S.8; see Alternate Protocol 3). Reagents: (a) amine, diisopropylethylamine, dimethyl sulfoxide; (b) 0.1 M tetrabutylammonium fluoride; (c) 0.1 M acetic acid.
0.1 M aqueous acetic acid or 1 M tetrabutylammonium fluoride (TBAF) in tetrahydrofuran (THF) Methanol, anhydrous 0.1 M sodium bicarbonate (NaHCO3) Vial or test tube with secure cap Temperature-controlled oil bath or heating block at 45° to 60°C Rotary evaporator with water aspirator and oil pump Additional reagents and equipment for thin-layer chromatography (TLC; see Basic Protocol 1), or high-performance liquid chromatography (HPLC) 1. Combine the following in a vial or a test tube that can be securely capped. 1 equivalent 2-fluoro-6-O-(trimethylsilylethyl)-2′-deoxyinosine 1.5 to 2.0 equivalents amine 2 equivalents DIEA 20 µL DMSO per mg of nucleoside. 2. Heat in a temperature-controlled oil bath or heating block at 45° to 60°C. If the reaction is not homogeneous after warming up, add additional anhydrous DMSO. 3. Continue heating for 1 to 2 days, until the fluoronucleoside (S.4) has been completely eliminated. Check the reaction periodically by TLC (see Basic Protocol 1, step 6; S.4 Rf ∼0.36).
Synthesis of Modified Nucleosides
1.3.11 Current Protocols in Nucleic Acid Chemistry
4. Evaporate the solvent under reduced pressure using a rotary evaporator, first with a water aspirator and then under high vacuum with an oil pump. A centrifugal vacuum evaporator (e.g., Speedvac) can also be used and is often more convenient for small-scale syntheses such as those described in Alternate Protocols 2 and 3.
5. Add 20 µL of 0.1 M aqueous acetic acid (or 20 µL of 1 M TBAF in THF) per mg nucleoside and incubate at room temperature for 2 hr to remove the trimethylsilylethyl protecting group. If necessary, add an organic solvent such as methanol to improve solubility. Monitor by TLC until deprotection is complete. Some of the TMSE protecting group will probably have fallen off the product by the end of the reaction. The deprotected product will have a lower Rf than the 6-O-TMSE compound.
6. Neutralize the reaction with 0.1 M NaHCO3 (20 µL if 0.1 M acetic acid was used) and check the pH (6.5 to 7.5) with pH paper. 7. Evaporate solvents under reduced pressure (step 4). 8. Purify the product by flash chromatography on silica gel, by preparative TLC, or by HPLC. The choice of purification technique will depend primarily on the scale of the reaction. The choice of solvent system will depend on the product characteristics. The alkylated nucleosides can be stored indefinitely at 4° or –20°C. ALTERNATE PROTOCOL 2
SYNTHESIS OF 2-N-(3-AMINOPROPYL)-2′-DEOXYGUANOSINE This protocol describes the preparation of a 2-N-aminopropyl derivative of 2′-deoxyguanosine (S.6) using tetrabutylammonium fluoride (TBAF) to remove the 6-Otrimethylsilylethyl group (see Fig. 1.3.3B). Additional Materials (also see Basic Protocol 2) 1,3-Diaminopropane Acetonitrile Concentrated NH4OH Silica gel Sand Tetrahydrofuran (THF), anhydrous 0.1 M ammonium formate in water, pH 6.4 12 × 75–mm glass test tube or conical vial with magnetic stir bar 2.5 × 20–cm column 100-mL round-bottom flask 10 × 250–mm C18 reversed-phase HPLC column (e.g., YMC-ODS-AQ column; YMC) 1. Combine the following in a 12 × 75–mm glass test tube or conical vial with a small magnetic stir bar. 50 mg (0.13 mmol) 2-fluoro-6-O-TMSE-2′-dI (S.4) 16.8 mg (0.13 mmol) DIEA 200 µL DMSO.
Synthesis of N2-Substituted Deoxyguanosine Nucleosides
2. Add 23 mg (0.31 mmol) of 1,3-diaminopropane, cap securely, and heat while stirring in a 45°C oil bath or heating block.
1.3.12 Current Protocols in Nucleic Acid Chemistry
Additional amine (well in excess of the 1 eq that is theoretically required) is used to ensure that only one of the amino groups in the diaminopropane reacts with the fluoronucleoside. It is possible for the diamine to react at both ends to form a bis(nucleoside) cross-linked with a propyl chain. If this is the desired product, the diamine can be used as the limiting reagent (e.g., 1 mmol of diamine to 3 mmol fluoronucleoside).
3. Continue heating until the fluoronucleoside (S.4) has been completely eliminated. Check the reaction periodically by TLC (see Basic Protocol 1, step 6), eluting with 85:8:7 (v/v/v) acetonitrile/H2O/concentrated NH4OH. The reaction should be finished after ∼20 hr. The Rf of 1-(6-O-trimethylsilylethyl)-N2-(3aminopropyl)-2′-deoxyguanosine (S.5) is ∼0.25, and that of S.4 is ~0.8.
4. Evaporate the solvent under reduced pressure (see Basic Protocol 2, step 4). 5. Prepare a column for flash chromatography by packing a 2.5 × 20–cm column with 20 g silica gel using the TLC solvent system (step 3). 6. Place the sample in a 100-mL round-bottom flask and add a few milliliters of methanol to dissolve it. Add ~1 g dry silica gel and evaporate the methanol using a rotary evaporator with a water aspirator. 7. Carefully add the powdery sample/gel mixture as evenly as possible to the top of the column. Add a layer of sand and elute using the TLC solvent mixture. 8. Collect 5- to 7-mL fractions and analyze by TLC (step 3). 9. Combine fractions containing pure product and evaporate to dryness using a rotary evaporator, first with a water aspirator and then under high vacuum with an oil pump. Yields of 70% to 75% (35 to 40 mg) of S.5 should be obtained.
10. Dissolve S.5 in 500 µL anhydrous THF. Add 200 µL of 1 M TBAF in THF and stir the reaction for 12 hr at room temperature. 11. Evaporate the solvent under reduced pressure (see Basic Protocol 2, step 4). 12. Monitor the deprotection reaction by TLC (step 3). The Rf of S.5 is 0.25. The deprotected product will have a much lower Rf .
13. Add 200 µL of 1:1 (v/v) DMSO/methanol and purify by HPLC using the following conditions: Column: 10 × 250–mm YMC-ODS-AQ Solvents: (A) 0.1 M ammonium formate, pH 6.4; (B) acetonitrile Flow rate: 5 mL/min Gradient: 99% to 90% A over 15 min, 90% to 80% A over 5 min, 80% to 0% A over 10 min, return to 99% A over 5 min. 2-N-(3-Aminopropyl)-2′-deoxyguanosine (S.6) elutes at ∼12 min. Yields of ∼50% should be expected for the deprotection and HPLC purification steps. The product can be stored indefinitely at 4° or –20°C.
Synthesis of Modified Nucleosides
1.3.13 Current Protocols in Nucleic Acid Chemistry
ALTERNATE PROTOCOL 3
SYNTHESIS OF 2-N-[2(S)-1-HYDROXYBUT-3-EN-2-YL]-2′DEOXYGUANOSINE This protocol describes the use of 2-amino-3-butenol to prepare an 2-N-hydroxyalkenesubstituted 2′-deoxyguanosine (S.8) using acetic acid to remove the 6-O-trimethylsilylethyl group (Fig. 1.3.3C). Additional Materials (also see Basic Protocol 2) 2(S)-Amino-3-butenol 0.1 M ammonium formate in water, pH 6.4 12 × 75–mm glass test tube or conical vial with a magnetic stir bar Lyophilizer 10 × 250–mm C18 reversed-phase HPLC column (e.g., YMC-ODS-AQ column; YMC) 1. Dry 10 mg of 2-fluoro-6-O-TMSE-2′-dI (S.4; 0.027 mmol) overnight in a 12 × 75–mm glass test tube under vacuum at room temperature using a rotary evaporator and an oil pump. 2. Add a small magnetic stir bar and the following reagents: 35 mg (0.041 mmol) 2(S)-amino-3-butenol 10 µL (0.057 mmol) DIEA 50 µL anhydrous DMSO. 3. Cap the tube securely and heat with stirring in a temperature-controlled oil bath or heating block at 60°C for 20 hr. 4. Monitor the reaction by TLC (see Alternate Protocol 2, step 3). The reaction mixture may contain both product with TMSE (S.7) and desilylated product (S.8). Starting material S.4 has an Rf of 0.8, and S.8 has an Rf of 0.22. S.7 will have an Rf between those of S.4 and S.8.
5. Remove the solvent under reduced pressure (see Basic Protocol 2, step 4). 6. Suspend the residue in 0.5 mL water and lyophilize. 7. Remove remaining TMSE protecting group by treating the product mixture with 200 µL of 0.1 M aqueous acetic acid at room temperature for 2 hr. 8. Neutralize with an equal volume of 0.l M NaHCO3 (final pH 6.5 to 7.5) and lyophilize. 9. Purify by HPLC using the following conditions: Column: 10 × 250-mm YMC-ODS-AQ Solvents: (A) 0.1 M ammonium formate, pH 6.4; (B) methanol Flow rate: 5 mL/min Gradient: 90% to 10% A over 25 min. The deprotected product (S.8) elutes at ∼12 min; protected product (S.7) elutes at ∼26 min. The yield from this reaction is ∼80%. The product can be stored indefinitely at 4° or –20°C. Synthesis of N2-Substituted Deoxyguanosine Nucleosides
1.3.14 Current Protocols in Nucleic Acid Chemistry
SETTING UP A NITROGEN ATMOSPHERE Many of the steps are carried out under a slight positive pressure of inert gas (usually nitrogen, although argon can also be used) to keep moisture and oxygen out of reactions. When a procedure is described as being carried out under nitrogen, this is accomplished by attaching a source of dry nitrogen gas via tubing to a Y- or T-shaped glass connector. One arm of the connector is attached to the inlet of a bubbler that is partially filled with a high-boiling inert liquid such as mineral oil; the outlet of the bubbler is open to the atmosphere. The third arm of the connector is connected via tubing to the reaction flask equipped with either a gas inlet adapter or a syringe needle inserted through a rubber septum in one of the joints of the flask. To ensure a slight positive pressure of inert gas in the system, the flow of nitrogen is adjusted so that a slow stream of bubbles is created in the bubbler. This arrangement is preferable to having nitrogen pass through the flask, which causes evaporation of solvents. A useful guide to handling air-sensitive reagents and working in an inert atmosphere can be found in Technical Bulletin AL-134 (Aldrich Chemical, 1983).
SUPPORT PROTOCOL
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Anisaldehyde/sulfuric acid spray Dissolve 5 mL of p-anisaldehyde in 90 mL of 95% ethanol. Carefully add 5 mL concentrated H2SO4 followed by three drops of glacial acetic acid. Store up to several months at 4° to 6°C or several days at room temperature. Reagent can be applied to TLC plates with a spray bottle or by dipping. CAUTION: Prepare and use this reagent only in a well-ventilated fume hood. It is corrosive and the vapors are highly irritating.
Sodium methoxide in methanol, 0.35 M Prepare a 0.35 M solution by adding 18.9 g solid sodium methoxide (Aldrich) to 1 liter anhydrous methanol. Prepare fresh before use or store for several days at room temperature under an inert atmosphere. Prepared solutions of sodium methoxide in methanol can be purchased from Aldrich. COMMENTARY Background Information Syntheses of N2-substituted guanine nucleosides have been studied for many years, primarily with ribosides. The primary route to these compounds has been via nucleophilic displacement of a halogen at the 2 position of an inosine bearing a substituent (such as O-benzyl or thione) at the 6 position (Montgomery and Hewson, 1960; Gerster and Robins, 1965, 1966). The halogen (bromo, chloro, or fluoro) is introduced by aqueous diazotization of the 2-amino group of a guanosine derivative in the presence of a halide source (Gerster and Robins, 1965), and the O6 substituent is converted by hydrogenation (for the benzyl substituent) or oxidation (for thione) to the 2-fluoro guanine derivative.
This strategy, while successful for ribosides, requires acidic conditions that are too vigorous for the acid-labile deoxyribosides. The current strategy takes advantage of methodology developed by Robins and Uznanski (1981) for low-temperature, nonaqueous diazotization of ribosides in organic solvents using t-butyl nitrite as the diazotizing agent and HF/pyridine as the fluoride source for the preparation of fluoro derivatives. Other fluoridating agents have also been explored (Acedo et al., 1994; Adib et al., 1997). The fluoro derivatives have proved to be the most useful because they are more reactive toward nucleophilic displacement than the other halogen derivatives (Gerster and Robins, 1965). Lee et al. (1990) reported that the nonaqueous diazotization strat-
Synthesis of Modified Nucleosides
1.3.15 Current Protocols in Nucleic Acid Chemistry
Synthesis of N2-Substituted Deoxyguanosine Nucleosides
egy could be used for the preparation of 2fluoro-6-O-benzyl-2′-deoxyinosine. Harris et al. (1991) then reported similar experiments that had been extended to the synthesis of 2fluoro-2′-deoxyinosine itself, and its incorporation into oligonucleotides via phosphoramidite chemistry. However, the 2-fluoro derivative without O6-protection did not react as well with nucleophiles, and its conversion to the corresponding phosphoramidite did not proceed in very high yield. Although the benzyl protecting group is easily introduced, its removal requires catalytic hydrogenation, which can cause problems if a polycyclic aromatic hydrocarbon (PAH) substituent has been introduced at N2 (Lee et al., 1990). Hence, it became apparent that a different O6-protecting group was required. The 4nitrophenethyl (NPE) group, which had been explored by Gaffney and Jones (1982) and Himmelsbach et al. (1984) for the protection of deoxyguanosine during oligonucleotide synthesis, has found wide use for preparation of the 2-fluoroinosine derivative (Zajc et al., 1992; Erlanson et al., 1993; Eritja et al., 1995; Lee et al., 1995; Schmid and Behr, 1995; Diaz et al., 1997). 2-Fluoro-6-O-(4-nitrophenethyl)-2′deoxyinosine can be prepared by the series of reactions described in this unit, starting with either triacetyl (Lee et al., 1995; see Basic Protocol 1) or diacetyl (Zajc et al., 1992; see Alternate Protocol 1) deoxyguanosine. Use of the 3′,5′-O-diacetyl derivative greatly shortens the synthesis, because several days of ammonia/methanol treatment are required for removal of the N2-acetyl group. There is risk of losing the NPE group by β-elimination if sodium methoxide is used for this step. The trimethylsilylethyl group is also very useful for O6-protection (Tsarouhtsis et al., 1995; DeCorte et al., 1996). Both groups can be used for preparation of O6-protected 2-fluoro-2′-deoxyinosine phosphoramidites and for oligonucleotide synthesis. The NPE group is usually removed by treatment with 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU); it may be slowly lost if the substitution reaction requires prolonged heating in the presence of excess amine. The TMSE group, while quite stable in the fluoro derivative under anhydrous neutral or alkaline conditions, is often lost spontaneously once there is an amine substituent at C2, and generally requires only mild acid treatment to achieve complete deprotection. Allerson et al. (1997) have recently reported a synthesis of the 2-fluoro-6-O-(4-nitrophenethyl)-riboside derivative, in which the hydroxyls and N2 were
protected as triethylsilyl (TES) derivatives; the TES groups were removed in the course of the fluoridation reaction. Quite a number of substitution reactions have now been reported using a 2-fluoro-2′-deoxyinosine derivative, either at the nucleoside stage or in oligonucleotides. Amine nucleophiles have been used to attach a wide variety of substituents including amino alcohols (Zajc et al., 1992), polyamines (Schmid and Behr, 1995; Diaz et al., 1997), disulfide-containing diamines (Wolfe and Verdine, 1993), pyrrolizidine amines (Woo et al., 1993; Tsarouhtsis et al., 1995), heterocyclic amines (Wang and Bergstrom, 1993; Ramasamy et al., 1994), benzylic amines bearing large polycyclic substituents (Lee et al., 1990, 1995; Sangaiah et al., 1992; Zajc et al., 1992), and metal-binding ligands (Bergstrom and Gerry, 1994). In addition, 15N is easily introduced by use of 15NH3 at either the nucleoside or oligonucleotide stage (Acedo et al., 1994). An acylated derivative of 2-fluoro-6-ONPE-2′-deoxyinosine has been used in a reaction with the severely hindered triol amines derived from opening of the diol epoxide of benzo[a]pyrene with ammonia (Zajc et al., 1992); however, a high temperature was required for the reaction. The 2-triflate derivative of 2′-deoxyinosine (Steinbrecher et al., 1993; Edwards et al., 1997) offers promise for syntheses requiring very bulky, highly sterically hindered nucleophiles.
Compound Characterization 2-N-3′,5′-O-Triacetyl-2′-deoxyguanosine (S.2): mp 193° to 194°C (soften), 225°C (decompose); TLC Rf 0.31 (9:1 CH2Cl2/MeOH); 1H NMR (DMSO-d ) δ 12.20 to 11.40 (br, 2H, 6 2×NH), 8.23 (s, 1H, H8), 6.22 (dd 1H, J = 7.2 Hz, H1′), 5.33 (d, 1H, J = 5.7 Hz, H3′), 4.22 to 4.20 (m, 3H, H4′, H5′, H5′′), 2.95 (m, 1H, H2′), 2.52 (m, 1H, H2′′), 2.16 (s, 3H, CH3), 2.06 (s, 3H, CH3), 2.00 (s, 3H, CH3). 3′,5′-O-Diacetyl-2′-deoxyguanosine (S.7): mp 300°C (decompose); 1H NMR (DMSO-d6) δ 7.80 (s, 1H, H8), 6.40 (2H, bs, NH2), 6.03 (s, 1H, H1′), 5.19 (1H, m, H3′), 4.15 (1H, m, H5′′), 4.11 (2H, m, H5′′H4′), 2.81 (m, 1H, H2′), 2.33 (m, 1H, H2′′, 1.97 (s, 3H, CH3), 1.94 (s, 3H, CH3). 6-O-(Trimethylsilylethyl)-2′-deoxyguanosine (S.3): mp 66° to 68°C; TLC Rf 0.41 (9:1 CH2Cl2/MeOH); 1H NMR (MeOH-d4) δ 8.00 (s, 1H, H8), 6.30 (dd, 1H, J = 6.1 Hz, J = 2.1 Hz, H1′), 4.61 to 4.53 (m, 3H, OCH2CH2Si, H3′), 4.03 (q, 1H, J = 2.7 Hz, H4′), 3.83 (dd,
1.3.16 Current Protocols in Nucleic Acid Chemistry
1H, J = 3.3, 12.2, H5′), 3.73 (dd, 1H, J = 12.2, 2.9, H5′′), 2.76 (m, 1H, H2′), 2.34 (m, 1H, H2′′), 1.21 (m, 2H, CH2Si), 0.08 (s, 9H, (CH3)3Si); 1H NMR (CDCl3) δ 7.64 (s, 1H, H8), 6.16 (dd, 1H, J = 8.8 Hz, J = 5.7 Hz, H1′), 5.04 (s, 2H, NH2), 4.56 (d, 1H, J = 4.8 H z , H 3′) , 4 . 3 9 ( t , 2 H , J = 8 . 6 H z , OCH2CH2Si), 4.07 (br, 1H, H4′), 3.70 (m, 2H, H5′, H5′′), 2.73 (m, 1H, H2′), 2.21 (m, 1H, H2′′), 1.02 (t, 2H, J = 8.6 Hz, CH2Si), -0.10 (s, 9H, (CH3)3Si). 2-Fluoro-6-O-(trimethylsilylethyl)-2′deoxyinosine (S.4): mp 120° to 122°C; TLC Rf 0.36 (9:1 CH2Cl2/MeOH); 1H NMR (MeOH-d4) δ 8.46 (s, 1H, H8), 6.41 (t, 1H, J = 6.7 Hz, H1′), 4.67 to 4.72 (m, 2H, OCH2CH2Si), 4.56 (m, 1H, H3′), 4.02 (q, 1H, J = 3.6 Hz, H4′), 3.81 (dd, 1H, J = 3.6, H5′′), 3.73 (dd, 1H, J = 2.9, H5′), 2.77 (m, 1H, H2′′), 2.44 (m, 1H, H2′), 1.26 (m, 2H, CH2Si), 0.1 (s, 9H, (CH3)3Si); 1H NMR (CDCl3) δ 8.56 (s, 1H, H8), 6.31 (t, 1H, J = 6.7 Hz, H1′), 5.35 (d, 1H, J = 4.2 Hz, 3′-OH), 4.94 (t, 1H, J = 5.5 Hz, 5′-OH), 4.64 (t, 2H, J = 8.3 Hz, OCH2CH2Si), 4.40 (m, 1H, H3′), 3.87 (q, 1H, J = 4.0 Hz, H4′), 3.55 (m, 2H, H5′ and H5′′), 2.68 (m, 1H, H2′), 2.32 (m, 1H, H2′′), 1.20 (t, 2H, J = 8.3 Hz, CH2Si), 0.08 (s, 9H, (CH3)3Si); 19F NMR (MeOH-d4) δ -51.55; 19F NMR (CDCl ) δ −50.40. 3
Critical Parameters and Troubleshooting Overall, it is important that starting nucleosides for each step in the synthesis be thoroughly dried either by coevaporation with anhydrous pyridine or in an Abderhalden drying apparatus (overnight under vacuum at 78°C). Anhydrous solvents are also important. The solvents should either be freshly distilled and stored under nitrogen or argon, or should be taken from a freshly opened bottle of commercially prepared anhydrous solvent. It is best to purchase anhydrous solvents in the smallest size bottle that will fit the requirements of the synthesis. Acylation. Dry reagents are important for acylation. The main effect of moisture is to decrease the yield of product. Serious problems are seldom encountered in this step. Mitsunobu alkylation. This reaction is most sensitive to the quality of the DEAD reagent. The best yields are obtained when using reagent from a freshly opened bottle. Fluoridation. This reaction requires careful attention to three factors: (1) dryness of the
O6-protected nucleoside, (2) quality of the HF/pyridine reagent, which should be pale amber or lighter in color, and (3) maintenance of low temperature during the course of the reaction. Lack of attention to any of these factors can lead to depurination and failure to obtain any product. Substitution reactions. The nature of the nucleophile is the most critical parameter in this reaction. Sterically hindered amines require longer time and/or higher temperature for reaction. The rate of reaction is also determined by the concentrations of nucleoside and amine. Temperatures as high as 80° to 85°C have been used in certain cases. If the substituent is a bulky hydrocarbon, removal of the TMSE group may require more vigorous deprotection conditions (e.g., 5 mM HCl in methanol at 50°C for several hours). Evaporation of solvents. This is usually accomplished using rotary evaporation under vacuum. It is very helpful if the rotary evaporator is equipped with a dry ice condenser or with a condenser that is cooled with chilled water. The vacuum is normally applied with a water aspirator (25 to 30 mmHg), which is quite sufficient for materials with boiling points <100°C (at 760 mm). However, several of the reagents and solvents used in these procedures have boiling points >100°C. For these steps, it is better to use an oil pump to produce a vacuum of ≤1 mmHg. This allows higher-boiling compounds to be removed with the application of little or no heat. The general procedure is to remove the more volatile materials first using the water aspirator, and then remove the higherboiling compounds using the oil pump. The oil pump should not be used to remove the lowerboiling materials, particularly if there are chlorinated solvents present. If the only volatile material is water, it can also be removed by lyophilizaton or vacuum centrifugal evaporation (e.g., with a Speedvac evaporator). Small volumes of volatile organics can also be removed by this method if the instrument is equipped with a chemically resistant cold trap. This is, for example, an effective way to removed small volumes of DMSO.
Anticipated Results An overall yield of 45% to 55% of 2-fluoro6-O-TMSE-2′-deoxyinosine from 2′-deoxyguanosine can be obtained in this series of reactions. The yields of substitution reactions are generally very high, although the recovered yield of purified product may be considerably
Synthesis of Modified Nucleosides
1.3.17 Current Protocols in Nucleic Acid Chemistry
lower, depending upon the scale and method of purification.
Time Considerations An approximate time scale for the sequence of reactions described in this unit would be 3 to 5 days for acylation, depending upon whether the tri- or diacetyl derivative is chosen as the starting material; 3 days for the Mitsunobu reaction; 2 days for fluoridation; and 2 to 4 days for substitution reactions, depending upon the nucleophile and the mode of purification (e.g., HPLC generally takes longer than preparatory TLC). The reaction can be greatly accelerated by using a large excess of amine. Considerable variation in these times is to be expected and is based upon the time allowed for drying and/or distillation of reagents, recrystallization, chromatographic separation, and analysis, as well as on the experimenter’s level of synthetic chemistry experience.
Literature Cited Acedo, M., Fabrega, C., Avino, A., Goodman, M., Fagan, P., Wemmer, D., and Eritja, R. 1994. A simple method for N-15 labelling of exocyclic amino groups in synthetic oligodeoxynucleotides. Nucl. Acids Res. 22:2982-2989. Adib, A., Potier, P.F., Doronina, S., Huc, I., and Behr, J.-P. 1997. A high-yield synthesis of deoxy-2fluoroinosine and its incorporation into oligonucleotides. Tetrahedron Lett. 38:2989-2992. Aldrich Chemical. 1983. Technical Bulletin AL134: Handling Air-Sensitive Reagents. Aldrich Chemical Co., Milwaukee, Wis. Allerson, C.R., Chen, S.L., and Verdine, G.L. 1997. A chemical method for site-specific modification of RNA: The convertible nucleoside approach. J. Am. Chem. Soc. 119:7423-7433. Bergstrom, D.E. and Gerry, N.P. 1994. Precision sequence-specific cleavage of a nucleic acid by a minor-groove-directed metal-binding ligand linked through N-2 of deoxyguanosine. J. Am. Chem. Soc. 116:12067-12068. DeCorte, B.L., Tsarouhtsis, D., Kuchimanchi, S., Cooper, M.D., Horton, P., Harris, C.M., and Harris, T.M. 1996. Improved strategies for postoligomerization synthesis of oligodeoxynucleotides bearing structurally defined adducts at the N2 position of deoxyguanosine. Chem. Res. Toxicol. 9:630-637. Diaz, A.R., Eritja, R., and Garcia, R.G. 1997. Synthesis of oligodeoxynucleotides containing 2substituted guanine derivatives using 2-fluoro2′-deoxyinosine as common nucleoside precursor. Nucleosides Nucleotides 16:2035-2051. Synthesis of N2-Substituted Deoxyguanosine Nucleosides
Edwards, C., Boche, G., Steinbrecher, T., and Scheer, S. 1997. Synthesis of 2-substituted 2′-deoxyguanosines and 6-O-allylguanines via activation of C-2 by a trifluoromethanesulfonate group. J. Chem. Soc. Perkin Trans 1 1887-1893.
Eritja, R., Acedo, M., Avino, A., and Fabrega, C. 1995. Preparation of oligonucleotides containing non-natural base analogues. Nucleosides Nucleotides 14:821-824. Erlanson, D.A., Chen, L., and Verdine, G.L. 1993. DNA methylation through a locally unpaired intermediate. J. Am. Chem. Soc. 115:12583-12584. Gaffney, B.L. and Jones, R.A. 1982. A new strategy for the protection of deoxyguanosine during oligonucleotide synthesis. Tetrahedron Lett. 23:2257-2260. Gerster, J.F. and Robins, R.K. 1965. Purine nucleosides. X. The synthesis of certain naturally occurring 2-substituted amino-9-β-D-ribofuranosylpurin-6(1H)-ones (N2-substituted guanosines). J. Am. Chem. Soc. 87:3752-3759. Gerster, J.F. and Robins, R.K. 1966. Purine nucleosides. XIII. The synthesis of 2-fluoro- and 2chloroinosine and certain derived purine nucleosides. J. Org. Chem. 31:3258-3262. Harris, C.M., Zhou, L., Strand, E.A., and Harris, T.M. 1991. A new strategy for synthesis of oligonucleotides bearing adducts at exocyclic amino sites of purine nucleosides. J. Am. Chem. Soc. 113:4328-4329. Himmelsbach, F., Schulz, B.S., Trichtinger, T., Charubala, R., and Pfleiderer, W. 1984. The pnitrophenylethyl (NPE) group. A versatile new blocking group for phosphate and aglycone protection in nucleosides and nucleotides. Tetrahedron 40:59-72. Lee, H., Hinz, M., Stezowski, J.J., and Harvey, R.G. 1990. Syntheses of polycyclic aromatic hydrocarbon-nucleoside and oligonucleotide adducts specifically alkylated on the amino functions of deoxyguanosine and deoxyadenosine. Tetrahedron Lett. 31:6773-6776. Lee, H., Luna, E., Hinz, M., Stezowski, J.J., Kiselyov, A.S., and Harvey, R.G. 1995. Synthesis of oligonucleotide adducts of the bay region diol epoxide metabolites of carcinogenic polycyclic aromatic hydrocarbons. J. Org. Chem. 60:56045613. Matsuda, A., Shinozaki, M., Suzuki, M., Watanabe, K., and Miyasaka, T. 1986. A convenient method for the selective acylation of guanine nucleosides. Synthesis 385-386. Mitsunobu, O. 1981. The use of diethyl azodicarboxylate and triphenylphosphine in synthesis and transformation of natural products. Synthesis 1-28. Montgomery, J.A. and Hewson, K. 1960. Synthesis of potential anticancer agents. XX. 2Fluoropurines. J. Am. Chem. Soc. 82:463-468. Ramasamy, K., Zounes, M., Gonzalez, C., Freier, S.M., Lesnik, E.A., Cummins, L.L., Griffey, R.H., Monia, B.P., and Cook, P.D. 1994. Remarkable enhancement of binding affinity of heterocycle-modified DNA to DNA and RNA. Synthesis, characterization and biophysical evaluation of N2-imidazolylpropylguanine and N2-imidazolylpropyl-2-aminoadenine modified oligonucleotides. Tetrahedron Lett. 35:215-218.
1.3.18 Current Protocols in Nucleic Acid Chemistry
Robins, M.J. and Uznanski, B. 1981. Nucleic acid related compounds. 34. Non-aqueous diazotization with tert-butyl nitrite. Introduction of fluorine, chlorine, and bromine at C-2 of purine nucleosides. Can. J. Chem. 59:2608-2611. Sangaiah, R., Gold, A., Ball, L.M., Matthews, D.L., and Toney, G.E. 1992. Synthesis and resolution of putative diastereomeric N2-deoxyguanosine and N6-deoxyadenosine adducts of biologically active cyclopentaPAH. Tetrahedron Lett. 33:5487-5490. Schmid, N. and Behr, J.-P. 1995. Recognition of DNA sequences by strand replacement with polyamino-oligonucleotides. Tetrahedron Lett. 36:1447-1450. Steinbrecher, T., Wameling, C., Oesch, F., and Seidel, A. 1993. Activation of the C2 position of purine by the trifluoromethanesulfonate group: Synthesis of N2-alkylated deoxyguanosines. Angew. Chem. Int. Ed. Engl. 32:404-406. Tsarouhtsis, D., Kuchimanchi, S., DeCorte, B.L., Harris, C.M., and Harris, T.M. 1995. Synthesis of oligonucleotides containing interchain crosslinks of bifunctional pyrroles. J. Am. Chem. Soc. 117:11013-11014. Wang, G. and Bergstrom, D.E. 1993. Synthesis of oligonucleotides containing N2-[2-(imidazol-4ylacetamido)ethyl]-2′-deoxyguanosine. Tetrahedron Lett. 34:6725-6728.
Wolfe, S.A. and Verdine, G.L. 1993. Ratcheting torsional stress in duplex DNA. J. Am. Chem. Soc. 115:12585-12586. Woo, J., Sigurdsson, S.T., and Hopkins, P.B. 1993. DNA interstrand cross-linking reactions of pyrrole-derived, bifunctional electrophiles: Evidence for a common target site in DNA. J. Am. Chem. Soc. 115:3407-3415. Zajc, B., Lakshman, M.K., Sayer, J.M., and Jerina, D.M. 1992. Epoxide and diol epoxide adducts of polycyclic aromatic hydrocarbons at the exocyclic amino group of deoxyguanosine. Tetrahedron Lett. 33:3409-3412.
Key References Gerster and Robins, 1965, 1966. See above. These papers describe the synthesis and some of the reactions of 2-fluoro-6-O-benzyl-inosine. Tsarouhtsis et al., 1995; DeCorte et al., 1996. See above. Synthesis of 2-fluoro-6-O-(trimethylsilylethyl)-2′deoxyinosine and the related phosphoramidite; oligonucleotide synthesis and substitution reactions.
Contributed by Thomas M. Harris and Constance M. Harris Vanderbilt University Nashville, Tennessee
Synthesis of Modified Nucleosides
1.3.19 Current Protocols in Nucleic Acid Chemistry
Unnatural Nucleosides with Unusual Base Pairing Properties Synthetic modified nucleosides designed to pair in unusual ways with the natural nucleic acid bases have many potential applications in nucleic acid biochemistry. These range from biochemical tools for probing nucleic acid structure or protein–nucleic acid interactions to tools for re-engineering DNA and ultimately proteins. Applications as components of nucleic acid–based diagnostic tools for clinical analysis have been envisioned. Furthermore, unnatural bases may be useful as components of antisense or antigene nucleic acid analogs in therapeutic applications. This unit serves to lay the foundation for future protocol units on unnatural base synthesis and application, with particular emphasis on unnatural base analogs that mimic natural bases in size, shape, and biochemical processing. A much more extensive compilation of unnatural nucleobases has recently been published by Luyten and Herdewijn (1998). To design bases that mimic natural bases in function, it is useful to consider the factors that are essential for effective base pairing and stable duplex formation. In addition to structures that are configured to hydrogen bond and base stack within the spatial confines of duplex DNA and RNA, any surrogate base pair must also conform to specific dimensions and geometry if it is to function in roles that require recognition by nucleic acid–processing enzymes. To be isosteric with AT or GC base pairs, the C1′ to C1′ distance must be in the range of 10.8 to 11.0 Å, and λ1 and λ2 should be ~50° (Fig. 1.4.1; Saenger, 1984). Just as it is important to know where to place hydrogen bond donor and acceptor sites, it is important to consider the availability of sur-
Watson-Crick base pair geometry λ
C1' - C1' ~ 10.8 to 11.0 Å
C1'
λ ~ 50 - 54° λ C1'
Figure 1.4.1 Base pair parameters. See APPENDIX 1B and Figure A.1B.4 for other base pairing schemes.
UNIT 1.4
rounding free space when designing new nucleobases. This can generally be determined by inspecting the groove regions of nucleic acid duplex and triplex models. Where the foundation for the design is a natural nucleobase, substitution is allowed at C5 of C, T, or U. N4 of C and N6 of A are also possible attachment sites, but generally compromise base association. N2 of G is acceptable and, unlike all other sites, allows minor groove placement of an appendage. C8 of A and G have been used as sites for attachment of appendages, but substitution here may influence the conformation preference about the glycosidic bond. Replacement of the purine N7 with a carbon provides a site for attachment of appendages in a sterically tolerated position. There are many examples of appendages that, when added to the natural nucleic acid bases, enhance base-pair stability. These include substituents that increase the acidity of proton donor sites (Yu et al., 1993) or increase hydrophobicity and aromatic surface area for enhanced base stacking (Inoue et al., 1985), as well as appended cations for electrostatic interactions with the phosphodiester backbone (Ueno et al., 1998).
BASE PAIRS WITH ALTERNATIVE HYDROGEN BONDING SCHEMES Purine-Pyrimidine-Like Base Pairs Benner and co-workers have described a set of nucleobase analogs that resemble natural bases, but have reconfigured hydrogen bonding patterns (Piccirilli et al., 1990). Six orthogonal base pairs (S.1 to S.6) are shown in Figure 1.4.2. Extensive studies on these nucleosides have revealed that even these subtle changes in structure can cause profound effects on thermodynamic (Voegel and Benner, 1994) and biochemical properties (Switzer et al., 1993; Horlacher et al., 1995). For example, the C-nucleoside pyrimidine mimics appear to base pair more weakly than equivalent base pairs composed of N-nucleosides. As far as biochemical properties, it appears that certain DNA polymerases require the pyrimidine O2 and purine N3 as recognition features for efficient template-mediated oligonucleotide synthesis (Horlacher et al., 1995).
Contributed by Donald E. Bergstrom Current Protocols in Nucleic Acid Chemistry (2001) 1.4.1-1.4.13 Copyright © 2001 by John Wiley & Sons, Inc.
Synthesis of Modified Nucleosides
1.4.1 Supplement 5
C-G N H
N
H
H
N
H N
H
N
H N
N
N
O
H
N
dR
H
N
H
H
H 2
H
N dR N
N
3
N
N
O
O dR
N
H
N
H
N dR N O
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N N
N
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H
1
O N
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O
dR
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H
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4
dR
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H
H
dR
H
N dR
O
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O
N H
dR
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CH3
O H 6
site itself, the duplex was destabilized more than when the triazole was placed opposite each of the natural bases. This result is in line with NMR studies on duplex DNA containing the triazole carboxamide opposite either G or T (Klewer et al., 2001). In both cases, the triazole prefers to adopt a conformation in which the amide group points out into the major groove rather than inward towards the opposing base pair. To achieve specific association through hydrogen bond interactions, one of the two triazole carboxamides in a self-pair would have to face with the amide projecting inward towards the opposing base. This example reflects parameters that one must consider when designing base pairs; the interior of the helix is a less hydrophilic environment and, without sufficient compensation, highly hydrophilic groups will prefer to assume positions that place them in a more hydrophilic environment.
HYDROPHOBIC BASE PAIRS Figure 1.4.2 Structures of six Watson-Cricktype base pairs utilizing mutually exclusive hydrogen bonding schemes.
Self-Complementary Nucleobases Another concept for new base pair development was proposed by Pochet and Marliére (1996). Based on the known ability of the mutagenic base 8-oxoguanine to base pair with A from a syn conformation, these researchers redesigned the base by removing the 6-oxo group. This yields the unnatural base 2-amino-8oxopurine (S.7), which they postulate would pair with itself according to the arrangement shown in Figure 1.4.3. The results of biochemical studies with this unusual base have not yet been reported. One can imagine similar selfpairing potential for azole carboxamide nucleosides, as illustrated in Figure 1.4.3 for 1,2,4triazole-3-carboxamide (S.8). However, Tm studies on a duplex containing 1,2,4-triazole3-carboxamide showed that when placed oppo-
H N H
N
N dR
Unnatural Nucleosides with Unusual Base Pairing Properties
H N
N N
N
O
N H H 7
Figure 1.4.3
N H
N
H O
There are now a significant number of examples of hydrophobic unnatural bases that pair with other hydrophobic bases or with themselves in duplex DNA with higher affinity than with any of the natural bases. 3-Nitropyrrole deoxyribonucleoside (S.26; Fig. 1.4.8), which was originally designed as a universal nucleobase, pairs with almost equal affinity to each of the natural bases, but a 12-mer duplex with nitropyrrole opposite itself is significantly more stable than the same duplex with nitropyrrole opposite each of the natural bases (Bergstrom et al., 1995; Zhang et al., 1998). The significance of the hydrophobic substituent (nitro) in mediating this effect is clear when one compares pyrrole-3-carboxamide, which in duplex DNA yields far more stable duplexes when paired opposite each of the natural bases than when paired opposite itself. Because of these results, hydrophobic base pairs are considered to be attractive candidates for extension of the genetic alphabet. Two themes are possible: (1) the development of a
N
O N
N H H
O
N
dR
N H
N
dR N
N dR 8
Self-pairing bases.
1.4.2 Supplement 5
Current Protocols in Nucleic Acid Chemistry
complementary hydrophobic pair, and (2) the creation of a single self-complementary hydrophobic base. The latter possibility is more attractive, because one need contend with the optimization of DNA replication with only one unnatural nucleoside. A series of recent papers from a research effort at Scripps Research Institute led by Romesberg and Schultz has described considerable progress in the development of hydrophobic self-pairing bases (McMinn et al., 1999; Berger et al., 2000a; Ogawa et al., 2000; Wu et al., 2000). Three of these nucleoside analogs, 1-β-D-deoxyribosyl-7-azaindole (S.10; Ogawa et al., 2000) and two different 7-propynyl isocarbostyril deoxyribonucleosides (S.9; R = H and CH3; McMinn et al., 1999) are shown in Figure 1.4.4. These nucleoside analogs, as assessed by Tm measurements, self-pair significantly more effectively than they pair with any of the natural bases. Furthermore, both contain a lone pair of electrons that may be positioned optimally in the minor groove for interaction with DNA polymerase. Kool and co-workers have designed a set of non-hydrogen-bonding base analogs for A and T (Fig. 1.4.4; S.12 to S15; Schweitzer and Kool, 1995; Guckian et al., 1998). In a 12-bp duplex, virtually all paired combinations of the hydrophobic bases gave higher Tm values than when each hydrophobic base was paired with any of the natural bases. Pyrene deoxyribonucleoside (S.11; Fig. 1.4.4) is noteworthy because of the high specificity with which it pairs opposite abasic sites (Matray and Kool, 1999).
The nonpolar nucleobase difluorotoluene (S.14), a thymine isostere lacking hydrogenbonding functionality, can effectively substitute for thymine in both the template strand and as the incoming nucleoside triphosphate. These results suggest that shape recognition in the absence of hydrogen bonding is an important factor in base pair recognition (Guckian and Kool, 1997; Moran et al., 1997a,b; Guckian et al., 1998; Kool, 1998). Similarly, 4-methylbenzimidazole (S.12) is an effective surrogate for adenine when matched opposite the difluorotoluene nucleobase (Morales and Kool, 1999).
METAL-MEDIATED ASSOCIATION OF LIGAND NUCLEOBASE MIMICS Another way of mediating specific association between nucleobase-like molecules contained on a deoxyribosephosphodiester backbone is to configure them to bind transition metals. The simplest configurations would involve metals that form either linear planar complexes (e.g., Hg) or square planar complexes (e.g., Cu). Two examples of nucleobase mimics that bind a metal in a square planar complex have been published. Meggers et al. (2000) synthesized oligonucleotides containing opposing pyridine and pyridine-2,6-dicarboxylate ligands. The duplex is significantly stabilized in the presence of copper (Fig. 1.4.5; S.16), but not by other metals. Tanaka and Shionoya (1999) have described the synthesis of the diaminophenyl deoxyri-
Hydrophobic self-pairing bases H dR R N dR
N dR
O 9
N P-φ base pair
dR 10 11
R = H, CH3 F N F
N dR
N dR 12
13
Hydrophobic purine analogs
Figure 1.4.4
dR
dR
Hydrophobic nucleobase analogs.
14
15
Hydrophobic pyrimidine analogs
Synthesis of Modified Nucleosides
1.4.3 Current Protocols in Nucleic Acid Chemistry
Supplement 5
O
O P O O
O N
O O P O
N
M
O O O
O
O
O
16
O
P O O H N
O
O
O P O O
H N M
N H
O O
N H 17
Figure 1.4.5 Metal-mediated duplex association through metal-chelating nucleoside analogs. M, metal.
bonucleoside S.17, which binds platinum in a square planar complex to yield a stable structure resembling a natural base pair. Incorporation of this modified base into DNA and metal-mediated assembly of duplexes has not yet been described. However, the concept is noteworthy and could be the prelude to the development of highly novel nucleic acid analogs with unusual applications in material science and nanotechnology.
DEGENERATE BASES Chemical and radiological damage of nucleic acid bases frequently leads to modified bases that exhibit the ability to direct the incorporation of more than one natural base. Harnessing this ability for the development of mo-
P - imino H
O
N dR
O
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H H
N N
H H
N dR
18
K - amino
H
N N
G H
19
MeO
N N dR
N N
H H N
H
O
N
O
C H
N dR
O
N
20
Figure 1.4.6
N
A
N N
N
N
P - amino
O
O
N dR
N dR
H
MeO N H
T
N
N
N N
Unnatural Nucleosides with Unusual Base Pairing Properties
H N
lecular tools could be useful. One example is use of multicoding bases for the introduction of low-level mutations to generate random sequences in protein engineering. Bases that have the ability to pair with one natural base on template-directed primer extension but code for another base on template-directed replication by DNA polymerase have been referred to as convertides. Such bases are potentially useful as components of PCR/ligation-based detection assays (Day et al., 1999a,b). Two types of convertides have been described: (1) analogs that alter hydrogen bonding pattern and base pairing properties through tautomerism, and (2) analogs that alter their hydrogen bonding profile through changes in conformation. Brown and co-workers have described a purine analog that can base pair as either A (S.20) or G (S.21) and a pyrimidine analog that can base pair as C (S.19) or T (S.18; Fig. 1.4.6; Lin and Brown, 1989; Brown and Lin, 1991a; Negishi et al., 1997; Hill et al., 1998). The purine analog can adopt the hydrogen bonding configurations of adenine and guanine (Fig. 1.4.6) and the pyrimidine analog can adopt the hydrogen bonding configurations of thymine and cytosine through tautomerism. Isosteric azole carboxamide nucleobases capable of mimicking different sets of natural bases have been described. These compounds can mimic more than one natural nucleobase by virtue of rotation about bonds α and χ (Fig. 1.4.7). Modeling studies with azole carboxamides paired with the four natural bases in a B-DNA duplex show that two important parameters of base-pair geometry are maintained: the C1′ to C1′ distances are in the range of 10.8 to 11.0 Å, and λ1 and λ2 are ~50°. Models of
N dR
N H
N N
H
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K - imino 21
Tautomerically degenerate base analogs.
1.4.4 Supplement 5
Current Protocols in Nucleic Acid Chemistry
well with all of the natural bases in a nucleic acid duplex. Applications in which a nucleoside analog functions only passively as a partner for a natural base within a duplex represent only one aspect of the behavior necessary for a compound to be truly regarded as a universal nucleoside. Recognition of the triphosphate as a universal nucleotide substrate by DNA polymerase, or suitability of oligonucleotides containing the analog as substrates for enzymes such as restriction endonucleases or ligases, would also be useful functions. Although it is unlikely that a single nucleoside analog can be devised which would function as a universal nucleoside under all circumstances, it is likely that a variety of analogs will be found with universal nucleoside properties optimal for particular techniques.
azole carboxamide base pairs with a natural base pair yield C1′ to C1′ distances that fall within 0.2 Å and λ1 and λ2 within 3° of the natural base pairs (Bergstrom et al., 1996; Johnson et al., 1997; Zhang et al., 1998). 1,2,4Triazole-3-carboxamide deoxyribonucleoside illustrates the difficulty of predicting both structural and biochemical behavior of nucleoside analogs. Based on modeling studies, the author of this unit predicted that the triazole would base pair preferentially with G and T by the pairing motifs S.22 and S.23 (Fig. 1.4.7). Evidence in favor of this hypothesis included Tm data that showed that this analog gives significantly more stable duplexes when paired with G and T than when paired with C and A. However, an NMR study indicated that 1,2,4-triazole-3carboxamide paired opposite T assumes a conformation in which the carboxamide group is not hydrogen bonded to T as shown in Figure 1.4.7 (S.24). On the other hand, 1,2,4-triazole-3-carboxamide directs the incorporation of C and T (but not G) when a template containing this base analog is replicated by Taq DNA polymerase. The geometrically acceptable base pairs in this case would be structures S.25 and S.23.
Universal Base Specifications The specifications for an ideal universal base are the following. 1. It should form equally stable base pairs with each of the natural nucleobases in all sequence contexts as determined by thermal melting experiments (Tm). 2. It should be accepted as template base by DNA polymerase (preferably Taq DNA polymerase) as assessed by PCR replication of universal base–containing template and steadystate kinetic experiments for single-nucleotide primer extension.
UNIVERSAL NUCLEOSIDES Universal Nucleoside Concept The concept of a “wildcard” or “universal” nucleoside is generally understood to mean a nucleoside analog that can base pair equally
O
H
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H N
N
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22
G
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O
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N H
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or
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Thermodynamically favored base pairs
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H
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25 O
C
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O
N N dR
H N
α
N dR
H 23
N dR
O
T
DNA polymerase favored base pairs
Figure 1.4.7
N
Conformationally degenerate base analogs.
Synthesis of Modified Nucleosides
1.4.5 Current Protocols in Nucleic Acid Chemistry
Supplement 5
discrimination (A > G = T > C). By comparison, 3-nitropyrrole (S.29) pairs with little discrimination between each of the four natural bases (Bergstrom et al., 1995, 1997); however, it is significantly destabilizing relative to a natural base. This has been used to advantage in at least two separate applications. 3-Nitropyrrole has been used to increase the selectivity of hybridization-based detection of single-nucleotide polymorphisms (Guo et al., 1997), and as a tool to elevate the fidelity of thermostable Thermus thermophilus (Tth) DNA ligase for the ligation of oligonucleotide primers (Luo et al., 1996). The introduction of a second aromatic ring (e.g., 5-nitroindole; S.30) led to substantial improvement in duplex stability without too great of a loss in pairing nondiscrimination (Loakes and Brown, 1994). Neither 3-nitropyrrole nor 5-nitroindole deoxyribonucleosides are effective substrates or template components for DNA polymerase (Loakes et al., 1995). They preferentially direct the incorporation of the less polar natural bases A and T (Hoops et al., 1997). More recently it has been shown that both nondiscriminatory base pairing and high stability can be achieved with nucleosides based on the quinolone heterocycle (S.31; Berger et al., 2000b). The emphasis of the latter study has been to develop analogs that are effective substrates for DNA polymerases.
3. The corresponding deoxyribonucleoside triphosphate should be suitable as a substrate for DNA polymerase as assessed by steadystate kinetic experiments for single-nucleotide primer extension. 4. The base should be a substrate for DNA ligase as primer component as assessed by ligase chain reaction. This list is by no means complete, but represents those specifications that would be of the greatest use for the manipulation of DNA in conventional molecular biology techniques. The factors that contribute to base pair stability as assessed by thermal denaturation studies are not directly related to template preference by DNA polymerase. This has been illustrated by comparison of a set of azole carboxamide nucleosides for which the order of thermodynamic stabilities differs substantially from their template coding properties (Hoops et al., 1997).
Nonpolar Nucleobase Analogs The development of universal nucleic acid bases has progressed along two parallel lines. The first class of analogs includes molecules that cannot associate through hydrogen bonding, but because of size, shape, and hydrophobicity prefer to occupy the interior of a duplex (Fig. 1.4.8). The first analog of this class, phenyl deoxyribonucleoside (S.28), was reported in 1984 (Millican et al., 1984). It is highly destabilizing and pairs with significant
Evolution of the non-hydrogen bonding, hydrophobic universal base
NO2
NO2
N dR
dR 28
N dR
29
O 30
31
N dR
Hydrogen bonding capable universal base candidates
O N
Unnatural Nucleosides with Unusual Base Pairing Properties
N
H
NH2
N N dR
N
N 26
N dR
27
Deoxyinosine
Figure 1.4.8
Universal base candidates.
1.4.6 Supplement 5
Current Protocols in Nucleic Acid Chemistry
Polar Hydrogen Bonding Nucleobase Analogs The first studies of nucleoside analogs specifically designed to base pair with more than one of the four primary DNA bases appeared over a decade ago (Millican et al., 1984; Eritja et al., 1986; Seela and Kaiser, 1986; Habener et al., 1988; Lin and Brown, 1989; Brown and Lin, 1991b). The most extensively studied example is 2′-deoxyinosine (S.26; Fig. 1.4.8), which has been in use as a putative universal nucleoside in oligonucleotide probes and primers since 1985 (Ohtsuka et al., 1985). Structural studies on deoxyinosine-modified oligonucleotides show that dI can base pair to dC, dA (Corfield et al., 1987; Uesugi et al., 1987), T (Cruse et al., 1989; Carbonnaux et al., 1990), and dG (Oda et al., 1991). However, it is not a true universal nucleoside because the base pairs dI-dX (X = dA, dC, dG, T) differ in stability by as much as 2 to 3 kcal/mol (Martin and Castro, 1985; Kawase et al., 1989). More importantly, primers constructed with multiple sites of deoxyinosine substitution frequently give undecipherable results in sequencing experiments. More recently Seela and Debelak (2000) have developed a nucleoside analog, N8-(2′-deoxyribofuranoside) of 8-aza-7-deazaadenine (S.27; Fig. 1.4.8), which pairs with all four natural bases with significantly less discrimination than inosine, but with relatively high affinity.
TRIPLEX CONSTITUENTS Duplex formation occurs through WatsonCrick pairing of purine and pyrimidine bases, which involves hydrogen bonding of NH3 and O4 of thymine with N1 and NH6 of adenine, and O2, N3, and NH4 of cytosine with NH2, NH1, and O6 of guanine. This leaves two sites on each of the purine bases (N7 and NH6 of adenine and N7 and O6 of guanine) free for hydrogen bonding. A third oligonucleotide strand can associate with a duplex through hydrogen bond formation to these sites within the major groove to form a triplex. Triplex formation with natural nucleotides generally assumes one of two themes: parallel association between a homopyrimidine strand and a homopurine-homopyrimidine duplex following the base association schemes C+•GC (S.32; Fig. 1.4.9) and T•AT (S.33), or antiparallel association involving the triplets G•GC (S.34), A•AT (S.35), and T•AT. In each of these cases, base-base recognition from the third strand involves recognition of N7 and NH6 (adenine) or O6 (guanine). The reason that these base configurations are more stable than other pos-
sible arrangements of bases in a triplet configuration stems from geometry preferences, hydrogen bonding, and base stacking factors. Neither cytosine nor thymine can occupy the center strand, since both have only a single site (O6 of thymine and NH4 of cytosine) available for hydrogen bonding to a base in a third strand located in the major groove. On the other hand, each of the base-triplets illustrated in the figure has two hydrogen bonds between the central purine and the third-strand base. As a result, triplex formation is generally limited to polypurine strings within one stand of the duplex. This significantly limits the number of potential targets in an organism. T•TA, C•CG, C•TA, T•CG, A•TA, A•CG, G•TA, and G•CG, all of which could have at best one hydrogen bond between the third strand base and the central pyrimidine, are not stable. Furthermore, since C must be protonated at N3 in order to hydrogen bond to N7 of G, triplex formation through this motif is favored only at low pH. For these reasons, significant effort has been expended to (1) design third-strand unnatural bases that can bind opposite the pyrimidine bases, and (2) develop structural variations of cytosine that are protonated at neutral pH. Nucleobase design for triplex formation has been extensively reviewed (Ganesh et al., 1996; Doronina and Behr, 1997; Gowers and Fox, 1999). Rather than reiterate the extensive studies that have been done on unnatural nucleosides as third-strand components, this review will only provide a few examples to illustrate the different types of approaches that attempt to solve the problem. The triplets composed of C+•GC and T•AT are isomorphous. On the other hand, the G•GC, A•AT, and T•AT are not, which leads to significant dependence of triplex stability on the duplex sequence as well as on the relative number of GC and AT base pairs. Since the third-strand association of C by G requires that C be protonated, contiguous protonated C’s may lead to some destabilization through charge-charge repulsion. Consequently, nucleic acid chemists have sought nucleobase analogs that are protonated at this site at physiological pH. The approaches include modification of the cytosine to increase the pKa of protonated N3, and replacement of cytosine with unnatural bases that have hydrogen bond donor-donor configuration required to pair with the acceptor G. Figure 1.4.10 (S.36 to S.38) illustrates a few of the reported analogs designed to mimic protonated C.
Synthesis of Modified Nucleosides
1.4.7 Current Protocols in Nucleic Acid Chemistry
Supplement 5
H N H
H N dR N
H
N
O
H
O
H
dR N
H N H
H O
H
N
N
dR N
O
O
N 33
N O
N H N
H N
H
N dR
N
H
H
O
N N A•AT
dR
H N
N dR
N
H N H
N
G•GC
N
H N
N dR
N dR
N
H
N
N H T•AT
N H N
H N
H
32
O
N
N O
H
N
N dR
O
O dR N
O
N
N C+•GC
N dR
N
34
N
N
35
dR
Figure 1.4.9
Base association on triplex formation.
the base is a pyrazopyrimidine (P1) with both hydrogen-bond donating groups contained in the pyrimidine ring (Koh and Dervan, 1992). The third triple, S.38, contains a protonated 2-aminopyridine (designated as P in the figure; Cassidy et al., 1997). This is a more accurate structural mimic of protonated C than M or P1. The greater basicity of the pyridine (pKa ~ 6) means that the equilibrium will be shifted more towards the protonated form at physiological pH than in the case of C (pKa = 4.35).
The three structures shown in Figure 1.4.10 reflect three very different design strategies for mimicking protonated C. The first of these, S.36, contains the protonated-C mimic N6methyl-8-oxo-deoxyadenosine (designated as M in the figure; Krawczyk et al., 1992). The 8-oxo group shifts the equilibrium about the glycosidic bond toward a conformation (shown in the figure) that positions the hydrogen-bond donor sites on M for interaction with N7 and O6 of G. The second triple, S.37, also contains a neutral protonated-C mimic, but in this case
N N
H N H
Me N H
dR N
N
O
H
N
H
N N
N dR
M•GC
H
N
dR
P1•GC
H N H O
H N
Unnatural Nucleosides with Unusual Base Pairing Properties
P•GC
Figure 1.4.10 and P1.
N dR
N
N
N dR
O
H N N
N dR
N
H N H 37
N dR O
H N
O
H N
N H 36
H N
H
N O
H
H N H
H N
N
N N dR
O
N
O
N dR
H N H
38
Protonated cytosine analogs for triplex association. See text for definitions of M, P,
1.4.8 Supplement 5
Current Protocols in Nucleic Acid Chemistry
The expansion of base pair recognition beyond C+•GC, T•AT, G•GC, A•AT, and T•AT is not the only issue. The Tm values for thirdstrand dissociation are typically far lower than for duplex DNA of equivalent length. Consequently, further modifications have been explored to enhance triplex stability. Commonly, this has been achieved by attaching intercalators to one end of the third strand. Significant effort has even been expended to develop triplex-specific intercalators. An alternative is to develop base analogs that provide additional stabilization elements. One example, recently reported by Fox and co-workers, is the use of the nucleoside analog 5-(3-aminopropargyl)2′-deoxyuridine as a component of the third strand in place of thymidine (Bijapur et al., 1999). The increase in triplex stability, presumably due to association of the protonated amino group with the phosphodiester anion, was substantial. A challenge for nucleic acid chemists has been to design and develop modified nucleobases that can recognize CG and TA base pairs through association from the pyrimidine side of the major groove. Two examples of modified nucleosides designed to bind the CG base pair are illustrated in Figure 1.4.11. In S.39, only one hydrogen bond to the cytosine is present (Prévot-Halter and Leumann, 1999). In contrast, S.40 was designed to extend across the base pair and hydrogen bond to G as well as C (Huang et al., 1996). Rothman and Richards have proposed a number of structures from modeling experiments for TA base pair recognition (Rothman and Richards, 1996). The proposed structures, which contain alkyne/alkene spacers to a five-membered ring heterocycle, are predicted to form hydrogen bonds to O4 of T and N6 of A. They should be interesting candidates for future investigation.
MODIFYING NATURAL BASES TO TUNE PAIRING AFFINITY There are applications that would benefit from either decreasing base pair stability or increasing base pair stability while retaining base pair specificity. For natural sequences, CG base pairs contribute more to duplex stability than AT base pairs. This creates a problem in DNA-array-based strategies where it is advantageous for all sequences of the same length to have melting temperatures within a narrow temperature range. One would expect that to accomplish this without complicating sequence-specific effects would be difficult. A number of examples have been published that report progress in this direction. Nguyen et al. (1998) have reported that N4-ethylcytosine destabilizes a CG base pair to the extent that it resembles an AT base pair in stability. Alternatively, it is possible to increase the stability of AT base pairs by appending certain substituents at C5 of T in place of methyl. This is exemplified by the nucleoside analog 5-propynyl-2′deoxyuridine, which has found application in antisense oligonucleotides (Wagner et al., 1993). One would think that it would be possible to increase the affinity of association between A and T by adding an amino group to C2 of A to give 2,6-diaminopurine (2,6-DAP), which should hydrogen bond to T with three hydrogen bonds. However, in practice, this is not the case. The effect of the 2,6-DAP-T base pair on duplex DNA stability is dependent on sequence, and in some instances is not as stabilizing as an AT base pair. However, Matray et al. (2000) have discovered that 2,6-DAP is consistently stabilizing when it is incorporated into oligonucleotides containing the N3′→P5′ phosphoramidate linkage. This effect may be related to the adoption of A-type helices by this backbone modification. This example illustrates the difficulty of designing unnatural bases for manipulating DNA properties. Too many parame-
N dR N N O
H
N
T•CG
Figure 1.4.11
N dR
O H H
4H
39
N
N
N N dR
N H
N N
H
NH O
H
N
H H N
40
N dR
O
H
O
N
N
dR N
N
N H
N N
H
O
dR
Nucleobase analogs for triplex recognition of CG base pairs.
Synthesis of Modified Nucleosides
1.4.9 Current Protocols in Nucleic Acid Chemistry
Supplement 5
ters contribute to base pair stability to enable unambiguous prediction of the effects on duplex properties of even minor structural changes in the heterocycle. Yet another way that modified bases may be used to tune hybridization was reported by Kutyavin et al. (1996). 2-Thiothymine base pairs with adenine, but not with 2,6-DAP. This allowed Kutyavin and co-workers to develop a strategy for invasion of double-stranded DNA with formation of a stable three-arm junction. The complementary invading oligonucleotides contain 2-thiothymine and 2,6-DAP and do not form a stable duplex with each other, but they do hybridize effectively with the complementary natural sequences.
t-Bu
O
t-Bu O
Unnatural Nucleosides with Unusual Base Pairing Properties
O
N
t-Bu
O
N
N
H
H
41 H
N R
N
O
N H N
H N
O
H N
N R
N H
G•G mismatch recognition N
BASE PAIR RECOGNITION The unifying characteristic of the compounds discussed in the previous sections is that they are designed to function as nucleic acid components in place of natural nucleotides. However, the scope and potential uses for new molecules designed for nucleobase recognition extend well beyond those described above for unnatural nucleosides. Work on the recognition of base sequences through azole oligomers that bind to the nucleobases through the minor groove has led to the development of highly specific inhibitors of duplex DNA. Design of analogs that read single base pairs provides the opportunity for development of highly sensitive mismatch detection, which could in turn lead to tools for discovery of single-nucleotide polymorphisms. One example is the bisnaphthyridine intercalator (S.42) that binds specifically to GG base pairs (Fig. 1.4.12; Nakatani et al., 2001). It should be possible to design other analogs that selectively bind other mismatches. A second example illustrated in the figure is the hexylureido isoindolin-1-one derivative (S.41), which can associate with both bases of a CG base pair through hydrogen bonding in the major groove. The molecular details of recognition of multiple base pairs in a sequence through specific association of the bases with DNA-binding molecules such as netropsin are well recognized. Extensive research by Dervan and coworkers has cumulated in the design of polyamides capable of binding long duplex segments with high specificity through hydrogen bonding interactions in the minor groove (Dervan and Burli, 1999; Gottesfeld et al., 2000).
C•G base pair recognition
t-Bu
O N
N
H
H
N
N
N H
H
O N N R
O
N R
O H H N
N
N H
N
H
N
N N N
42
R
Figure 1.4.12 Molecules that recognize and bind combinations of natural bases.
CONCLUSION The objective of this review is to highlight a rapidly developing area of nucleic acid chemistry: unnatural base design. The design of new unnatural base pairs, universal bases, triplex components, and other bases with unusual base pairing specificities will continue to provide an attractive arena for molecular designers. It is hoped that the unit will provide some guidance and inspiration for synthetic chemists seeking problems in nucleic acid chemistry. Many potential uses for unnatural bases have been identified, and in most instances totally successful solutions have not yet been established.
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1.4.10 Supplement 5
Current Protocols in Nucleic Acid Chemistry
Bergstrom, D.E., Zhang, P., Toma, P.H., Andrews, C.A., and Nichols, R. 1995. Synthesis, structure, and deoxyribonucleic acid sequencing with a universal nucleoside: 1-(2′-Deoxy-β-D-ribofuranosyl)-3-nitropyrrole. J. Am. Chem. Soc. 117:1201-1209.
Eritja, R., Horowitz, D.M., Walker, P.A., ZiehlerMartin, J.P., Boosalis, M.S., Goodman, M.F., Itakura, M., and Kaplan, B.E. 1986. Synthesis and properties of oligonucleotides containing 2′-deoxynebularine and 2′-deoxyxanthosine. Nucl. Acids Res. 14:8135-8153.
Bergstrom, D.E., Zhang, P., and Johnson, W.T. 1996. Design and synthesis of heterocyclic carboxamides as natural nucleic acid mimics. Nucleosides Nucleotides 15:59-68.
Ganesh, K.N., Kumar, V.A., and Barawkar, D.A. 1996. Synthetic control of DNA triplex structure through chemical modifications. In Supramolecular Control of Structure and Bonding (A.D. Hamilton, ed.) pp. 263-327. John Wiley & Sons, New York.
Bergstrom, D.E., Zhang, P., and Johnson, W.T. 1997. Comparison of the base pairing properties of a series of nitroazole nucleobase analogs in the oligod eoxyribonucleotide sequence 5′d(CGCXAATTYGCG)-3′. Nucl. Acids Res. 25:1935-1942. Bijapur, J., Keppler, M.D., Bergqvist, S., Brown, T., and Fox, K.R. 1999. 5-(1-Propargylamino)-2′deoxyuridine (Up): A novel thymidine analogue for generating DNA triplexes with increased stability. Nucl. Acids Res. 27:1802-1809. Brown, D.M. and Lin, P.K.T. 1991a. The structure and application of oligodeoxyribonucleotides containing modified, degenerate bases. Nucl. Acids Symp. Ser. 24:209-212. Brown, D.M. and Lin, P.K.T. 1991b. Synthesis and duplex stability of oligonucleotides containing adenine-guanine analogues. Carbohydr. Res. 216:129-139. Carbonnaux, C., Fazakerley, G.V., and Sowers, L.C. 1990. An NMR structural study of deaminated base pairs in DNA. Nucl. Acids Res. 18:40754081. Cassidy, S.A., Slickers, P., Trent, J.O., Capaldi, D.C., Roselt, P.D., Reese, C.B., Neidle, S., and Fox, K.R. 1997. Recognition of GC base pairs by triplex-forming oligonucleotides containing nucleosides derived from 2-aminopyridine. Nucl. Acids Res. 25:4891-4898. Corfield, P.W.R., Hunter, W.N., Brown, T., Robinson, P., and Kennard, O. 1987. Inosine-adenine base pairs in a B-DNA duplex. Nucl. Acids Res. 15:7935-7949. Cruse, W.B.T., Aymani, J., Kennard, O., Brown, T., Jack, A.G.C., and Leonard, G.A. 1989. Refined crystal structures of an octanucleotide duplex with I.T. mismatch base pairs. Nucl. Acids Res. 17:55-72. Day, J.P., Bergstrom, D., Hammer, R.P., and Barany, F. 1999a. Nucleotide analogs facilitate base conversion with 3′-mismatch primers. Nucl. Acids Res. 27:1810-1818. Day, J.P., Hammer, R.P., Bergstrom, D., and Barany, F. 1999b. Nucleotide analogs and new buffers improve a generalized method to enrich for low abundance mutations. Nucl. Acids Res. 27:18191827. Dervan, P.B. and Burli, R.W. 1999. Sequence-specific DNA recognition by polyamides. Curr. Opin. Chem. Biol. 3:688-693. Doronina, S.O. and Behr, J.-P. 1997. Towards a general triple helix mediated DNA recognition scheme. Chem. Soc. Rev. 26:63-71.
Gottesfeld, J.M., Turner, J.M., and Dervan, P.B. 2000. Chemical approaches to control of gene expression. Gene Expr. 9:77-91. Gowers, D.M. and Fox, K.R. 1999. Towards mixed sequence recognition by triple helix formation. Nucl. Acids Res. 27:1569-1577. Guckian, K.M. and Kool, E.T. 1997. Highly precise shape mimicry by a difluorotoluene deoxynucleoside, a replication-competent substitute for thymidine. Angew. Chem. Int. Ed. Engl. 36:2825-2828. Guckian, K.M., Morales, J.C., and Kool, E.T. 1998. Structure and base pairing properties of a replicable nonpolar isostere for deoxyadenosine. J. Org. Chem. 63:9652-9656. Guo, Z., Liu, Q., and Smith, L.M. 1997. Enhanced discrimination of single nucleotide polymorphisms by artificial mismatch hybridization. Nat. Biotechnol. 15:331-335. Habener, J.F., Vo, C.D., Le, D.B., Gryan, G.P., Ercolani, L., and Wang, A.H.J. 1988. 5-Fluorodeoxyuridine as an alternative to the synthesis of mixed hybridization probes for the detection of specific gene sequences. Proc. Natl. Acad. Sci. U.S.A. 85:1735-1739. Hill, F., Loakes, D., and Brown, D.M. 1998. Polymerase recognition of synthetic oligodeoxyribonucleotides incorporating degenerate pyrimidine and purine bases. Proc. Natl. Acad. Sci. U.S.A. 95:4258-4263. Hoops, G.C., Zhang, P., Johnson, W.T., Paul, N., Bergstrom, D.E., and Davisson, V.J. 1997. Template directed incorporation of nucleotide mixtures using azole-nucleobase analogs. Nucl. Acids Res. 25:4866-4871. Horlacher, J., Hottiger, M., Podust, V.N., Hubscher, U., and Benner, S.A. 1995. Recognition by viral and cellular DNA polymerases of nucleosides bearing bases with nonstandard hydrogen bonding patterns. Proc. Natl. Acad. Sci. U.S.A. 92:6329-6333. Huang, C.-Y., Bi, G., and Miller, P.S. 1996. Triplex formation by oligonucleotides containing novel deoxycytidine derivatives. Nucl. Acids Res. 24:2606-2613. Inoue, H., Imura, A., and Ohtsuka, E. 1985. Synthesis and hybridization of dodecadeoxyribonucleotides containing a fluorescent pyridopyrimidine deoxynucleoside. Nucl. Acids Res. 13:7119-7128.
Synthesis of Modified Nucleosides
1.4.11 Current Protocols in Nucleic Acid Chemistry
Supplement 5
Johnson, W.T., Zhang, P., and Bergstrom, D.E. 1997. The synthesis and stability of oligodeoxyribonucleotides containing the deoxyadenosine mimic 1-(2′-deoxy-β- D-ribofuranosyl)imidazole-4carboxamide. Nucl. Acids Res. 25:559-567.
Matray, T., Gamsey, S., Pongracz, K., and Gryaznov, S. 2000. A remarkable stabilization of complexes formed by 2,6-diaminopurine oligonucleotide N3′→P5′ phosphoramidates. Nucleosides Nucleotides Nucleic Acids 19:1553-1567.
Kawase, Y., Iwai, S., and Ohtsuka, E. 1989. Synthesis and thermal stability of dodecadeoxyribonucleotides containing deoxyinosine pairing with four major bases. Chem. Phamacol. Bull. 37:599-601.
McMinn, D.L., Ogawa, A.K., Wu, Y., Liu, J., Schultz, P.G., and Romesberg, F.E. 1999. Efforts towards expansion of the genetic alphabet: DNA polymerase recognition of a highly stable, self pairing hydrophobic base. J. Am. Chem. Soc. 121:11585-11586.
Klewer, D., Zhang, P., Bergstrom, D.E., Davisson, V.J., and Liwang, A.C. 2001. Conformations of nucleoside analog 1-(2′-deoxy-β-D-ribofuranosyl)-1,2,4-triazole-3-carboxamide in DNA duplexes with different sequence contexts. Biochemistry 40:1518-1527. Koh, J.S. and Dervan, P.B. 1992. Design of a nonnatural deoxyribonucleoside for recognition of GC base pairs by oligonucleotide-directed triplehelix formation. J. Am. Chem. Soc. 114:14701478. Kool, E.T. 1998. Replication of non-hydrogen bonded bases by DNA polymerases: A mechanism for steric matching. Biopolymers 48:3-17. Krawczyk, S.H., Milligan, J.F., Wadwani, S., Moulds, C., Froehler, B.C., and Matteucci, M.D. 1992. Oligonucleotide-mediated triple helix formation using an N3-protonated deoxycytidine analog exhibiting pH-independent binding within the physiological range. Proc. Natl. Acad. Sci. U.S.A. 89:3761-3764. Kutyavin, I.V., Lukhtanov, E.A., Gorn, V.V., Meyers, R.B. Jr., and Gamper, H.B. Jr. 1996. Oligonucleotides containing 2-aminoadenine and 2-thiothymine act as selectively binding complementary agents. Biochemistry 35:11170-11176. Lin, T.K.T. and Brown, D.M. 1989. Synthesis and duplex stability of oligonucleotides containing cytosine-thymine analogues. Nucl. Acids Res. 17:10373-10383. Loakes, D. and Brown, D.M. 1994. 5-Nitroindole as an universal base analogue. Nucl. Acids Res. 22:4039-4043. Loakes, D., Brown, D.M., Linde, S., and Hill, F. 1995. 3-Nitropyrrole and 5-nitroindole as universal bases in primers for DNA sequencing and PCR. Nucl. Acids Res. 23:2361-2366. Luo, J., Bergstrom, D.E., and Barany, F. 1996. Improving the fidelity of Thermus thermophilus DNA ligase. Nucl. Acids Res. 24:3071-3078. Luyten, I. and Herdewijn, P. 1998. Hybridization properties of base-modified oligonucleotides within the double and triple helix motif. Eur. J. Med. Chem. 33:515-576. Martin, F.H. and Castro, M.M. 1985. Base pairing involving deoxyinosine: Implications for probe design. Nucl. Acids Res. 13:8927-8938. Matray, T.J. and Kool, E.T. 1999. A specific partner for abasic damage in DNA. Nature 399:704-708.
Meggers, E., Holland, P.L., Tolman, W.B., Romesberg, F.E., and Schultz, P.G. 2000. A novel copper-mediated DNA base pair. J. Am. Chem. Soc. 122:10714-10715. Millican, T.A., Mock, G.A., Chauncey, M.A., Patel, T.P., Eaton, M.A.W., Gunning, J., Cutbush, S.D., Neidle, S., and Mann, J. 1984. Synthesis and biophysical studies of short oligodeoxynucleotides with novel modifications: A possible approach to the problem of mixed base oligodeoxynucleotide synthesis. Nucl. Acids Res. 12:7435-7453. Morales, J.C. and Kool, E.T. 1999. Minor groove interactions between polymerase and DNA: More essential to replication than Watson-Crick hydrogen bonds? J. Am. Chem. Soc. 121:23232324. Moran, S., Ren, R.X.-F., and Kool, E.T. 1997a. A thymidine triphosphate shape analog lacking Watson-Crick pairing ability is replicated with high sequence selectivity. Proc. Natl. Acad. Sci. U.S.A. 94:10506-10511. Moran, S., Ren, R.X.-F., Rumney, S.I., and Kool, E.T. 1997b. Difluorotoluene, a nonpolar isostere for thymine, codes specifically and efficiently for adenine in DNA replication. J. Am. Chem. Soc. 119:2056-2057. Nakatani, K., Sando, S., and Saito, I. 2001. Scanning of guanine-guanine mismatches in DNA by synthetic ligands using surface plasmon resonance. Nat. Biotechnol. 19:51-55. Negishi, K., Williams, D.M., Inoue, Y., Moriyama, K., Brown, D.M., and Hayatsu, H. 1997. The mechanism of mutation induction by a hydrogen bond ambivalent, bicyclic N-4-oxy-2′-deoxycytidine in Escherichia coli. Nucl. Acids Res. 25:1548-1552. Nguyen, N.K., Bonfils, E., Auffray, P., Costaglioli, P., Schmitt, P., Asseline, U., Durand, M., Maurizot, J.C., Dupret, D., and Thuong, N.T. 1998. The stability of duplexes involving AT and/or G(4Et)C base pairs is not dependent on their AT/G(4Et)C ratio content. Implication for DNA sequencing by hybridization. Nucl. Acids Res. 26:4249-4258. Oda, Y., Uesugi, S., Ikehara, M., Kawase, Y., and Ohtsuka, E. 1991. NMR studies for identification of dI:dG mismatch base-pairing structure in DNA. Nucl. Acids Res. 19:5263-5267.
Unnatural Nucleosides with Unusual Base Pairing Properties
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Current Protocols in Nucleic Acid Chemistry
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Tanaka, K. and Shionoya, M. 1999. Synthesis of a novel nucleoside for alternative DNA base pairing through metal complexation. J. Org. Chem. 64:5002-5003. Ueno, Y., Mikawa, M., and Matsuda, A. 1998. Synthesis and properties of oligodeoxynucleotides containing 5-[N-[2[N,N-bis(2-aminoethyl)amino]ethyl]carbamoyl]-2′-deoxyuridine and 5[N-[3-[N,N-Bis(3-aminopropyl)amino]propyl] carbamoyl]-2′- deoxyuridine. Bioconjugate Chem. 9:33-39. Uesugi, S., Oda, Y., Ikehara, M., Kawase, Y., and Ohtsuka, E. 1987. Identification of I-A mismatch base-pairing structure in DNA. J. Biol. Chem. 262:6965-6968. Voegel, J.J. and Benner, S.A. 1994. Nonstandard hydrogen bonding in duplex oligonucleotides. The base pair between an acceptor-donor-donor pyrimidine analog and donor-acceptor-acceptor analog. J. Am. Chem. Soc. 116:6929-6930. Wagner, R.W., Matteucci, M.D., Lewis, J.G., Gutierrez, A.J., Moulds, C., and Froehler, B.C. 1993. Antisense gene inhibition by oligonucleotides containing C-5 propyne pyrimidines. Science 260:1510-1513. Wu, Y., Ogawa, A.K., Berger, M., McMinn, D.L., Schultz, P.G., and Romesberg, F.E. 2000. Efforts toward expansion of the genetic alphabet: Optimization of interbase hydrophobic interactions. J. Am. Chem. Soc. 122:7621-7632. Yu, H., Eritja, R., Bloom, L.B., and Goodman, M.F. 1993. Ionization of bromouracil and fluorouracil stimulates base mispairing frequencies with guanine. J. Biol. Chem. 268:15935-15943. Zhang, P., Johnson, W.T., Klewer, D., Paul, N., Hoops, G., Davisson, V.J., and Bergstrom, D.E. 1998. Exploratory studies on azole carboxamides as nucleobase analogs: Thermal denaturation studies on oligodeoxyribonucleotide duplexes containing pyrrole-3-carboxamide. Nucl. Acids Res. 26:2208-2215.
Contributed by Donald E. Bergstrom Purdue University West Lafayette, Indiana
Synthesis of Modified Nucleosides
1.4.13 Current Protocols in Nucleic Acid Chemistry
Supplement 5
Development of a Universal Nucleobase and Modified Nucleobases for Expanding the Genetic Code
UNIT 1.5
This unit presents protocols for the synthesis and characterization of nucleosides with unnatural bases in order to develop bases for the expansion of the genetic alphabet or for nonselective pairing opposite natural bases. The faithful pairing of nucleobases through complementary hydrogen-bond (H-bond) donors and acceptors forms the foundation of the genetic code. However, there is no reason to assume that the requirements for duplex stability and replication must limit the genetic alphabet to only two base pairs, or, for that matter, hydrogen-bonded base pairs. Expansion of this alphabet to contain a third base pair would allow for the encoding of additional information and would make possible a variety of in vitro experiments using nucleic acids with unnatural building blocks. Previous efforts to generate orthogonal base pairs have relied on H-bonding patterns that are not found with the canonical Watson-Crick pairs. However, in all cases, the unnatural bases were not kinetically orthogonal, and instead competitively paired with natural bases during polymerase-catalyzed DNA synthesis (Horlacher et al., 1995; Lutz et al., 1996, 1998a,b). Tautomeric isomerism, which would alter H-bond donor and acceptor patterns, likely contributes to this kinetic infidelity (Roberts et al., 1997a,b; Robinson et al., 1998; Beaussire and Pochet, 1999). An alternative strategy is centered around developing unnatural bases that form pairs based not on hydrogen bonds, but rather on interbase hydrophobic interactions. Such hydrophobic bases should not pair stably opposite natural bases due to the forced desolvation of the purines or pyrimidines. Additionally, the use of nucleobase analogs, which are not restricted to the shape of H-bonding topologies of natural bases, allows for the use of a wider range of analogs. This unit describes the design, synthesis, and characterization of unnatural base pairs involving 1-β-D-2-deoxyribosyl-N- and -C-nucleosides. To be reasonable candidates for the modification of the genetic code, unnatural nucleosides must meet certain criteria. First, the unnatural bases must pair stably and selectively in duplex DNA. Second, the unnatural bases must be good substrates for DNA polymerases, being replicated with good efficiency and fidelity. Determination of these thermodynamic and kinetic parameters of the unnatural nucleosides is accomplished by incorporation into oligonucleotides and subsequent evaluation as described herein. A general procedure for the synthesis of 1-β-D-2-deoxyribosyl-N-nucleosides containing pyrimidine-like unnatural hydrophobic bases using a modified Silyl-Hilbert-Johnson reaction (Hilbert and Johnson, 1930), namely Vorbrüggen glycosylation (Niedballa and Vorbrüggen, 1970), is first described (see Basic Protocol 1). This methodology is very useful for the synthesis of pyrimidine-like hydrophobic base pairs, because under the necessary reaction conditions the polar (but otherwise often rather insoluble) pyrimidine bases are readily converted by silylation into lipophilic silyl compounds, which are then soluble in organic solvents. Thus, the homogeneous coupling with sugar moieties is permitted. Next, a general procedure is included for the synthesis of 1-β-D-2-deoxyribosyl-N-nucleosides containing purine-like bases using the metal salt procedure (see Basic Protocol 2), which entails the condensation of nucleobase sodium salts with a sugar halide (Kazimierczuk et al., 1984). The sodium salts of acidic heterocyclic systems such as imidazole, purine, triazole, and pyrazole are prepared in situ with NaH or analogous bases. Contributed by Floyd E. Romesberg, Chengzhi Yu, Shigeo Matsuda, and Allison A. Henry Current Protocols in Nucleic Acid Chemistry (2002) 1.5.1-1.5.36 Copyright © 2002 by John Wiley & Sons, Inc.
Synthesis of Modified Nucleosides
1.5.1 Supplement 10
The synthesis of 3,5-dimethylphenyl-C-nucleoside by (1) condensation of in situ–generated Grignard reagent from 1-bromo-3,5-dimethylbenzene with 1-α-chloro-3,5-di-Otoluoyl-2-deoxyribofuranose and (2) methoxide-mediated deprotection of the bistoluoyl groups is also described (see Basic Protocol 3). In addition, the unit includes a procedure for the synthesis of 1,4-dimethylnaphthaleneC-nucleoside (see Basic Protocol 4). The sugar precursor containing the aldehyde group is used; this is synthesized from 2-deoxyribose in seven steps (Eaton and Millican, 1988) and does not require the generation of any stereocenters beyond C1′. The condensation of the aryllithium reagent of 2-bromo-1,4-dimethylnaphthalene with the aldehyde, followed by in situ cyclization and deprotection of hydroxyl groups, affords the desired nucleoside. An Alternate Protocol describes the synthesis of 3-methyl-2-naphthalene-C-nucleoside. A different sugar precursor containing an aldehyde group at C3′ is generated from Felkin-Anh addition of an allylzinc nucleophile to isopropylidene-protected glyceraldehydes with high diastereoselectivity (Solomon and Hopkins, 1993). Mesylation, followed by treatment with excess trifluoroacetic acid, yields a diastereomeric mixture of separable C-nucleosides. The purification of DNA containing these unnatural bases is then presented (see Basic Protocol 5). Purified DNA products are used to characterize the unnatural base pairs as candidate pairs for expanding the genetic code. Finally, the thermodynamic (see Basic Protocol 6) and kinetic characterization (see Basic Protocol 7) of the unnatural bases are described. NOTE: All glassware used for reactions should be evacuated, flame-dried, and flushed with argon before use. All operations should be carried out in a well-ventilated fume hood. All reactions involving moisture-sensitive reagents should be performed under argon atmosphere (see Reagents and Solutions). NOTE: It is recommended that all products in this unit be stored in a desiccator at ≤4°C. BASIC PROTOCOL 1
GENERAL GUIDELINES FOR SYNTHESIS OF PYRIMIDINE-LIKE 1-β-D-2-DEOXYRIBOSYL-N-NUCLEOSIDES This protocol outlines a general procedure for the synthesis of unnatural hydrophobic pyrimidine-like N-nucleosides (Figs. 1.5.1 and 1.5.2; McMinn et al., 1999). Specific protocols are given for the synthesis of 7-propynylisocarbostyril (PICS) triphosphate (S.7), as well as the phosphoramidite of PICS (S.6). Synthesis of the other unnatural nucleosides shown in Figure 1.5.2 can be accomplished using similar synthetic transformations, starting from the appropriate pyrimidine. The introduction of propynyl groups, the deprotection of toluoyl groups, and the formation of the phosphoramidite and triphosphate described here are generally applied to all synthesis protocols in this unit.
Development of a Universal Nucleobase and Unnatural Nucleobases
Materials Argon (see recipe) Isocarbostyril (S.1; Aldrich) or pyrimidine of choice Acetonitrile, anhydrous N,O-Bis(trimethylsilyl)acetamide (Aldrich) Bis-toluoyl-protected chloroglycoside: 1-α-chloro-3,5-di-O-toluoyl-2-deoxyribofuranose (S.2; Berry & Associates; Takeshita et al., 1987)
1.5.2 Supplement 10
Current Protocols in Nucleic Acid Chemistry
TolO O N H 1
O
TolO
a
Cl
+
5'
OTol
HO
d N
O
N
O
O
O
4
TolO
2'
OTol 3
TolO
O 1'
3'
2
b, c
N O
4'
5
OH e, f g
DMTrO N
O
PiPiPiO
O
N
O
O O P
6
N(i -Pr) 2
O
OH
CN
7
Figure 1.5.1 General procedure for synthesis of unnatural hydrophobic pyrimidine-like N-nucleosides (see Basic Protocol 1). Reagents: (A) Bis-acetamide, SnCl4, CH3CN, 0°C to room temperature (steps 1 to 22); (B) ICl, CH2Cl2, reflux to room temperature (steps 23 to 30); (C) propyne, (Ph3P)2PdCl2, CuI, TEA, −78°C to room temperature (steps 31 to 43); (D) 0.5 M sodium methoxide, methanol, room temperature (steps 44 to 49); (E) DMTrCl, pyridine, room temperature (steps 50 to 62); (f) 2-cyanoethyl-diisopropylchlorophosphoramidite, DIPEA, CH2Cl2, 0°C (steps 63 to 71); (g) tetrabutylammonium pyrophosphate, POCl3, tributylamine, Proton-Sponge, trimethyl phosphate, 0°C (steps 72 to 80).
N
O
N
O
N
O
N
O
ICS
MICS
5MICS
Pyridone
(isocarbostyril)
(3-methylisocarbostyril)
(5-methylisocarbostyril)
(3,5-dimethyl-2-pyridone)
N
O
N
O
N
O
PIM
PPyridone
PICS
(7-propynyl-3-methylisocarbostyril)
(7-propynyl-3-methyl-2( 1H )-pyridone)
(7-propynylisocarbostyril)
Figure 1.5.2 Unnatural hydrophobic N-nucleosides (see Basic Protocol 1).
Synthesis of Modified Nucleosides
1.5.3 Current Protocols in Nucleic Acid Chemistry
Supplement 10
SnCl4, anhydrous, freshly distilled in vacuo Ethyl acetate Hexanes Ammonium molybdate solution (see recipe) Saturated sodium bicarbonate (NaHCO3) solution Saturated sodium chloride (NaCl) solution Sodium sulfate (Na2SO4), anhydrous Silica gel, 200 to 400 mesh, 60 Å Dichloromethane (CH2Cl2), freshly distilled from calcium hydride Ethyl ether, anhydrous Iodine monochloride (ICl; Aldrich; packaged under nitrogen in Sure/Seal vials) Saturated sodium thiosulfate (Na2S2O3) solution Magnesium sulfate (MgSO4), anhydrous Triethylamine (TEA), freshly distilled from calcium hydride Dichlorobis(triphenylphosphine) palladium(II) [(Ph3P)2PdCl2] (Aldrich) Copper(I) iodide (CuI) Dry ice/ethyl ether bath (–100°C) and dry ice/acetone bath Propyne Sodium methoxide (Aldrich) Methanol, anhydrous, distilled from magnesium turnings and stored over 3A molecular sieves Ammonium chloride (NH4Cl) Pyridine, anhydrous, distilled and stored on NaOH protected from light 4,4′-Dimethoxytrityl chloride (DMTr-Cl; Aldrich) Diisopropylethylamine (DIPEA), freshly distilled from calcium hydride 2-Cyanoethyl diisopropylchlorophosphoramidite (Aldrich) Trimethyl phosphate Proton-Sponge (Aldrich) Phosphorous oxychloride (POCl3), freshly distilled Tributylamine Tetrabutylammonium pyrophosphate (TBAP), stored over Drierite 1 M triethylammonium bicarbonate (TEAB), pH 7
Development of a Universal Nucleobase and Unnatural Nucleobases
10-, 25-, and 100-mL two-neck round bottom flasks with 14/20 joints, oven-dried Rubber septa Vacuum system (oil pump) with manifold and cold trap Silica gel 60 F254 alumina-backed thin-layer chromatography (TLC) plates (Fisher) Heat gun UV light source 100-, 250-, and 500-mL separatory funnels 300-mL round bottom flask Cotton Glass funnel Rotary evaporator (Büchi) equipped with a dry ice condenser and a vacuum system Heavy-walled glass columns (1.5-cm i.d.; 10-, 15-, and 20-cm length) with glass adapters attached to compressed air or nitrogen source (see flash chromatography steps in APPENDIX 3E) Sea sand Reflux condenser with 14/20 joint 23- and 18-G needles 100-mL three-neck flask with 14/20 joints Propyne inlet adapter (Aldrich, cat. no. Z41.577-4) Teflon caps for 14/20 joints
1.5.4 Supplement 10
Current Protocols in Nucleic Acid Chemistry
Speedvac evaporator (Savant) High-performance liquid chromatography (HPLC) system Glass or disposable plastic syringes 10-mL test tubes Oil bath with temperature controller Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and flash chromatography (APPENDIX 3E) NOTE: Glass or disposable plastic syringes are generally used for addition of liquid reagents. The 10-mL test tubes are used for collection of eluting fractions in column chromatography. Oil baths with temperature controllers are used for heating reactions as necessary. Perform condensation of isocarbostyril (S.1) with bis-toluoyl-protected chloroglycoside (S.2) 1. Place a magnetic stir bar in an oven-dried 100-mL two-neck flask with two rubber septa, and place the flask on top of a magnetic stir plate. 2. Evacuate reaction flask on vacuum line, then flush with argon. Repeat this procedure three times and attach the flask to an argon line on the manifold. 3. Quickly remove the rubber septum on one of the side inlets, add 0.50 g (3.4 mmol) isocarbostyril (S.1) to the flask under argon, then immediately reinsert the septum. 4. Transfer 10 mL acetonitrile to the flask under argon. 5. Add, in dropwise fashion, 0.85 mL (3.4 mmol) N,O-bis(trimethylsilyl)acetamide and stir vigorously for 40 min at room temperature. During this time the suspension is cleared.
6. Add an additional 12 mL of acetonitrile to the clear reaction mixture, followed by 1.10 g (2.80 mmol) bis-toluoyl-protected chloroglycoside (S.2). Cool the reaction mixture in an ice-water bath. 7. Slowly add 0.18 mL (2 mmol) SnCl4 and continue to stir under argon at 0°C. Preparation of S.3 uses 1.0 eq S.1, 0.8 eq S.2, 1.0 eq N,O-bis(trimethylsilyl)acetamide, 0.6 eq SnCl4, and 129.4 eq acetonitrile. Increasing the quantity of SnCl4 from catalytic to substoichiometric amounts may improve yield.
8. Monitor the progress of the reaction by analytical TLC (APPENDIX 3D) as follows: a. Occasionally withdraw a small sample (1 to 5 µL) using a capillary tube and spot on silica gel 60 F254 alumina-backed plates. b. Develop the plate (APPENDIX 3D) with 4:1 (v/v) hexanes/ethyl acetate. c. Visualize by dipping plate into ammonium molybdate solution and heating with a heat gun. Examine with UV light source. For S.3, Rf = 0.4 (see APPENDIX 3D). To analyze the course of reactions by TLC, the anisaldehyde/sulfuric acid staining system (6 g of p-anisaldehyde, 50 mL of absolute ethanol and 2.5 mL of concentrated H2SO4) is useful for nucleosides. For nucleosides containing a DMTr group, 3% TFA/CH2Cl2 is also effective. Ammonium molybdate is used to assay the presence of carbohydrate derivatives and is sufficient for visualization of S.3. Spots generally appear in various shades of purple and blue.
Synthesis of Modified Nucleosides
1.5.5 Current Protocols in Nucleic Acid Chemistry
Supplement 10
Work up and purify S.3 9. When TLC analysis indicates the reaction is complete, transfer the mixture to a 500-mL separatory funnel and add 250 mL ethyl acetate. 10. Extract twice, each time with 50 mL saturated NaHCO3 solution, then extract once with 100 mL saturated NaCl solution. 11. Insert cotton into a glass funnel and add a 2-cm-thick layer of anhydrous Na2SO4 over the cotton. Filter the organic layer by gravity into a 300-mL round-bottom flask. 12. Remove solvents under reduced pressure with a rotatory evaporator equipped with a dry ice condenser. It is recommended that the bath temperature not exceed 45°C.
13. Pack a 1.5 × 20–cm heavy-walled column with 60 g of 200 to 400 mesh silica gel in hexanes for flash column chromatography (APPENDIX 3E). Gas pressure (air or nitrogen) is applied via a glass adapter to the top of the column through the gas inlet to increase the flow rate to about 5 drops/sec.
14. Dissolve the crude product in 1 mL of dichloromethane. 15. Transfer the solution using a 3-mL glass pipet onto the silica gel and let the solution sink into the column bed. 16. Rinse the flask with two 1-mL aliquots of dichloromethane, add each rinse to the surface of silica gel, and allow the rinse to enter the column. 17. Carefully add a 0.5-cm layer of sea sand to the top of the silica gel bed. This step is optional. Keeping the surface of the silica gel flat is important for good separation.
18. Elute with a gradient of 8:1 to 4:1 (v/v) hexanes/ethyl acetate. Use a glass cylinder to freshly mix the solvents.
19. Collect 10-mL fractions and analyze by TLC (see step 8). To achieve the best separation, TLC using 5:1 hexanes/ethyl acetate should give an Rf value of ∼0.35 for the possible product.
20. Transfer fractions containing pure product to a 300-mL round-bottom flask and evaporate to dryness with a rotary evaporator. 21. Add 3 mL of anhydrous ethyl ether to triturate the product. Remove the solvent and evacuate the product until the weight of the flask is not changed. 22. Confirm the desired product by 1H NMR, NOESY, and COSY analysis (see, e.g., Lambert et al., 1998). Flash chromatography gives a pair of diastereoisomers as the α and β anomers. The desired product is the faster-migrating β anomer, which is a white foam. The assignment of β-stereochemistry at C1′ for each nucleoside is based on NOESY data, in which H1′ shows cross-peaks with both α-H4′ and α-H2′ (see Fig. 1.5.1). The optimal yield of 1′-β-3′,5′-O-toluoyl-2′-deoxyribosylisocarbostyril (S.3) is 48%.
Development of a Universal Nucleobase and Unnatural Nucleobases
Iodinate S.3 by Friedel-Crafts iodination 23. Assemble a 25-mL oven-dried two-neck flask with a 14/20 reflux condenser in the center neck, a rubber septum on the side neck, and a magnetic stirring bar. Maintain a smooth flow of argon using a 23-G needle inserted through another septum on top of the reflux condenser.
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Current Protocols in Nucleic Acid Chemistry
24. Mix 0.40 g (0.80 mmol) S.3 (from step 21) with 5 mL freshly distilled dichloromethane (CH2Cl2) and transfer solution to the flask under argon atmosphere. 25. Add, in a dropwise fashion, 0.96 mL of 1 M ICl in CH2Cl2 (0.96 mmol ICl). 26. Reflux the reaction mixture 1 min at 45°C, then cool to room temperature. Perform TLC analysis on the mixture as in step 8. CAUTION: Iodine monochloride is highly toxic and volatile and should be handled with care in a well-ventilated hood.
Work up the iodinated product 27. When TLC analysis indicates that all of the starting material has disappeared during the cooling period, add 10 mL saturated NaHCO3 followed by dropwise addition of saturated Na2S2O3 until the reaction solution is clear. 28. Transfer the mixture to a 100-mL separatory funnel. Extract the aqueous layer three times, each time with 20 mL dichloromethane. 29. Combine the organic extracts and dry over anhydrous MgSO4. 30. Repeat steps 11 and 12 and attach a vacuum line to dry the residue. The resulting crude iodinated product (not shown in the figures) is pure enough for use in the next step. It should be kept in darkness and used immediately.
Introduce propynyl group by Sonogashira coupling 31. Equip a 100-mL, 14/20 three-neck flask with rubber septa on the two side necks, a 14/20 dry-ice condenser-trap in the center neck, and a magnetic stir bar. Place a gas inlet adapter on top of the condenser and attach it to an argon line on the manifold. 32. Evacuate the apparatus and then purge it with argon three times to remove air. 33. Transfer ∼0.80 mmol of the iodinated compound and 15 mL triethylamine (TEA) to the flask and quickly add 0.01 g (0.02 mmol) (Ph3P)2PdCl2 and 0.01 g (0.06 mmol) CuI under argon. (Ph3P)2PdCl2 and CuI are highly sensitive to oxygen and light, respectively. They should be handled very carefully and quickly. High quality of these reagents is essential to the success of Sonogashira coupling reactions.
34. Cool the reaction vessel in dry ice/diethyl ether bath (−100°C). 35. Fill the condenser-trap with pulverized dry ice. 36. Quickly remove the rubber septum on one of the side inlets and insert a propyne inlet adapter. 37. Purge the flask with propyne for 10 min or until final volume is 50 mL. Preparation of S.4 uses 1.0 eq S.3, 1.2 eq ICl, 116.2 eq CH2Cl2, 0.07 eq CuI, 0.02 eq (Ph3P)2PdCl2, 134.5 eq TEA, and >120 eq propyne.
38. Turn off the propyne cylinder. Quickly remove the rubber septa and the propyne inlet, and seal inlets tightly with Teflon caps. 39. Remove dry-ice condenser and seal the inlet tightly with a Teflon cap. Continue to stir at room temperature for 5 hr. Synthesis of Modified Nucleosides
1.5.7 Current Protocols in Nucleic Acid Chemistry
Supplement 10
Work up and purify S.4 40. Cool the reaction mixture in dry ice/acetone bath for 5 min, then remove the cold bath. 41. Remove the caps carefully to vent the remaining propyne at ambient temperature. Evaporate the solvent with a rotary evaporatory. 42. Purify the crude product by flash chromatography (steps 13 to 19) using a 1.5 × 15–cm silica gel column packed with 30 g silica gel. Elute with a gradient of 5:1 to 3:1 (v/v) hexanes/ethyl acetate. Perform TLC on the eluate (see step 8) using 4:1 hexanes/ethyl acetate. For S.4, Rf = ∼0.35.
43. Combine appropriate fractions based on TLC analysis and evaporate to dryness. Flash column chromatography yields 0.11 g (64%) of S.4 as a white foam.
Deprotect 3′- and 5′-OH groups 44. Equip a 10-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G needle threaded through the septum. 45. Add 0.10 g (0.20 mmol) S.4 (from step 43) and 4 mL anhydrous methanol sequentially to the flask. 46. Add dropwise 1.2 mL of 0.5 M sodium methoxide in methanol (0.60 mmol sodium methoxide) to the reaction mixture and continue to stir under argon for 20 min. Preparation of S.5 uses 1.0 eq S.4, 3 eq sodium methoxide, and 617 eq methanol.
Work up and purify S.5 47. Add ∼30 mg NH4Cl to quench the reaction and keep stirring for 10 min. 48. Evaporate solvents and purify the product by flash chromatography (steps 13 to 19) using a 1.5 × 10–cm silica gel column packed with 20 g silica gel. Elute with a gradient of 2:100 to 5:100 (v/v) methanol/CH2Cl2. Perform TLC on the eluate (see step 8) using 4:100 methanol/CH2Cl2. For S.5, Rf = ∼0.35.
49. Combine appropriate fractions based on TLC analysis and evaporate to dryness. The reaction is approximately quantitative and the product, 1-β-2′-deoxyribosyl-7propynylisocarbostyril (S.5), is obtained as a white solid.
Tritylate 5′-OH by Williamson etherification 50. Equip an oven-dried two-neck 10-mL flask with a stir bar and two rubber septa. Keep the vessel under argon using an 18-G needle threaded through the septum. 51. Add 0.046 g (0.154 mmol) S.5 (from step 49) and 4 mL anhydrous pyridine to the flask. 52. Using an 18-G needle, insert a vacuum line through one of the septa to remove most of the pyridine by vacuum evaporation. 53. Purge the reaction flask with argon and remove the vacuum line immediately. Development of a Universal Nucleobase and Unnatural Nucleobases
54. Transfer 0.7 mL anhydrous pyridine to the flask to dissolve S.5.
1.5.8 Supplement 10
Current Protocols in Nucleic Acid Chemistry
55. Prepare a solution of 0.08 g (0.23 mmol) 4,4′-dimethoxytrityl chloride in 0.3 mL pyridine and add in a dropwise fashion to the reaction mixture over a period of 20 min. Tritylation uses 1.0 eq S.5, 1.5 eq DMTr-Cl, and 82.3 eq pyridine.
56. Continue to stir at room temperature for an additional 20 min. Work up and purify tritylated nucleoside 57. Combine the reaction mixture with 20 mL of ethyl acetate, 5 mL saturated NaHCO3, and 5 mL saturated NaCl to partition. 58. Transfer the mixture to a 250-mL separatory funnel. Rinse the reaction vessel twice, each time with 5 mL of ethyl acetate, and transfer to the funnel. 59. Separate the organic layer and extract the aqueous layer twice, each time with 10 mL of ethyl acetate. 60. Filter and dry the organic layer as in steps 11 and 12. 61. Purify the crude product by flash chromatography (steps 13 to 19) using a 1.5 × 10–cm silica gel column packed with 20 g silica gel. Elute with a gradient of 5:1 to 3:1 (v/v) hexanes/ethyl acetate. Perform TLC on the eluate (see step 8) using 3.5:1 hexanes/ethyl acetate. For the DMTr-protected product of the above reaction, Rf = ∼0.35.
62. Combine appropriate fractions based on TLC analysis and evaporate to dryness. Chromatographic purification affords the DMTr-protected product (0.073 g, 79%) as a white foam.
Phosphitylate to give phosphoramidite S.6 63. Equip a 10-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G needle threaded through the septum. 64. Prepare a solution of 0.073 g (0.121 mmol) tritylated product (from step 62) in 1.3 mL of freshly distilled dichloromethane and transfer to the flask. 65. Add 0.084 mL (0.48 mmol) DIPEA to the solution and cool in ice-water bath. 66. Add, in a dropwise fashion, 0.041 mL (0.16 mmol) 2-cyanoethyl diisopropylchlorophosphoramidite and allow the reaction mixture to reach ambient temperature over 15 min. Relative to the starting nucleoside (S.5 in step 51), preparation of S.6 uses 3.2 eq DIPEA, 1.1 eq 2-cyanoethyldiisopropylchlorophosphoramidite, and 135.2 eq dichloromethane.
Work up and purify S.6 67. Dilute the reaction mixture with 50 mL dichloromethane. Transfer the reaction mixture to a 100-mL separatory funnel. 68. Wash with 5 mL saturated NaHCO3 and 5 mL saturated NaCl. 69. Filter and dry as in steps 11 and 12. 70. Perform flash chromatography (steps 13 to 19). Elute with a gradient of 6:1 to 4:1 (v/v) hexanes/ethyl acetate. Perform TLC (see step 8) using 5:1 hexanes/ethyl acetate. For S.6, Rf = ∼0.35.
Synthesis of Modified Nucleosides
1.5.9 Current Protocols in Nucleic Acid Chemistry
Supplement 10
71. Combine appropriate fractions based on TLC analysis and evaporate to dryness using a rotary evaporator. The phosphoramidite product (S.6) is a white foam (63 mg), prepared with 65% yield, and is stored in a desiccator at −20°C. Phosphoramidite compounds are used for automated DNA synthesis; isolation and desalting of oligonucleotides is carried out by PAGE and electrophoretic dialysis (see Basic Protocol 5). Oligonucleotides are used for duplex oligonucleotide denaturation temperature measurements and kinetic studies of DNA extension (see Basic Protocols 6 and 7).
Prepare triphosphate S.7 72. Equip a 10-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G needle threaded through the septum. 73. Quickly remove the septum on one of the side inlets, add 0.008 g (0.027 mmol) S.5 (from step 49) under argon, and insert the rubber septum back to the flask. 74. Coevaporate moisture and methanol twice from 0.1 mL anhydrous pyridine. 75. Add 0.134 mL (0.23 mmol) trimethyl phosphate and 9 mg (0.04 mmol) ProtonSponge, dissolve completely, and bring to 0°C. 76. Add 0.003 mL (0.032 mmol) POCl3 and continue to stir at 0°C for 2 hr. 77. Add 0.042 mL (0.03 mmol) tributylamine and 0.023 g (0.040 mmol) TBAP and stir for 1 min. Work up and purify S.7 78. Add 2.7 mL of 1 M TEAB, pH 7. 79. Stir the reaction mixture for 10 min and concentrate to dryness in a Speedvac evaporator. 80. Dissolve the crude product in 200 µL of 1:1 (v/v) DMSO/isopropanol and purify by HPLC (UNIT 10.5) using the following conditions: Column: Rainin C18-Dynamax 60 Å column (7.8 × 300 mm) Buffer A: 0.1 M TEAB, pH 7 Buffer B: Acetonitrile Gradient: 98% to 90% buffer A over 5 min, 90% to 75% buffer A over 25 min, 75% to 0% buffer A over 5 min, return to 98% A over 5 min. Flow rate: 10 mL/min. Approximately 1 mg of the product triphosphate (S.7) is yielded with a retention time of 27.5 min, where the gradient is ∼65% buffer A. The triphosphate compound could be used directly for kinetic studies of DNA incorporation. BASIC PROTOCOL 2
Development of a Universal Nucleobase and Unnatural Nucleobases
GENERAL GUIDELINES FOR SYNTHESIS OF PURINE-LIKE 1-β-D-2-DEOXYRIBOSYL-N-NUCLEOSIDES This protocol outlines a general procedure for the synthesis of unnatural hydrophobic purine-like N-nucleosides. The coupling strategy between 5H-pyrrolo[2,3-b]pyrazine (PP) and S.2 (Fig. 1.5.1) to generate the nucleoside 1′-β-3′,5′-O-toluoyl-2′-deoxyribosylN-5H-pyrrolo[2,3-b]pyrazine (S.8; Fig. 1.5.3) can be generally applied for the synthesis of other purine-like nucleosides such as those depicted in Figure 1.5.4 by starting with the appropriate purine analog. This protocol describes the synthesis of the 3′,5′-toluoylprotected nucleoside (S.8). Synthesis of the free nucleoside, phosphoramidite, and triphosphate are performed as in Basic Protocol 1.
1.5.10 Supplement 10
Current Protocols in Nucleic Acid Chemistry
N
TolO
N
TolO N
O
+
N H
N
O
Cl OTol
PP
N
OTol
2
8
Figure 1.5.3 General procedure for synthesis of unnatural hydrophobic purine-like N-nucleosides
N N
N
N
N
N
N
7AI
M7AI
ImPy
(7-azaindole)
(6-methyl-7-azaindole)
(imidazole pyridine)
N N
N
N
N
P7AI
PPP
(3-propynyl-7-azaindole)
(3-propynyl-4,7-diazaindole)
Figure 1.5.4 Unnatural hydrophobic purine-like N-nucleosides (see Basic Protocol 2).
Materials Argon (see recipe) 5H-Pyrrolo[2,3-b]pyrazine or purine analog of choice Acetonitrile, anhydrous 60% sodium hydride (NaH; see recipe) Bis-toluoyl-protected chloroglycoside: 1-α-chloro-3,5-di-O-toluoyl-2-deoxyribofuranose (S.2; Berry & Associates; Takeshita et al., 1987) Ethyl acetate Ethyl ether Saturated sodium bicarbonate (NaHCO3) Sodium sulfate (Na2SO4), anhydrous Silica gel, 200-400 mesh, 60 Å Hexanes 10-mL two-neck round-bottom flask with 14/20 joints, oven-dried Rubber septa 18-G needle Cotton Glass funnel 1.5 × 30–cm heavy-walled glass column with glass-adapters attached to compressed air or nitrogen source (see flash chromatography steps in APPENDIX 3E)
Synthesis of Modified Nucleosides
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Sea sand Silica gel 60 F254 alumina-backed thin-layer chromatography (TLC) plates (Fischer) Heat gun UV light source Rotary evaporator (Büchi) equipped with a dry ice condenser and a vacuum system Additional reagents and equipment for purification and analysis (see Basic Protocol 1). 1. Equip a 10-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G syringe needle threaded through the septum. 2. Add 100 mg (0.84 mmol) 5H-pyrrolo[2,3-b]pyrazine and 4.5 mL anhydrous acetonitrile and cool the solution 5 min in an ice-water bath. 3. Add 36 mg of 60% NaH (0.9 mmol) in three aliquots to the reaction mixture. Remove the cold bath and stir at ambient temperature for 10 min. 4. Add 326.8 mg bis-toluoyl-protected chloroglycoside (S.2) in three aliquots to the flask and stir at ambient temperature for 10 min. Preparation of S.8 uses 1.0 eq 5H-pyrrolo[2,3-b]pyrazine, 1.1 eq NaH, 1.2 eq S.2, and 523 eq CH3CN.
5. Add 25 mL of ethyl acetate and 20 mL of saturated aqueous NaHCO3 to partition the reaction mixture. 6. Extract the aqueous layer twice, each time with 20 mL ethyl acetate. 7. Dry and filter the organic layer (see Basic Protocol 1, steps 11 and 12). 8. Perform flash chromatography (see Basic Protocol 1, steps 13 to 19) using a 1.5 × 10–cm column packed with 20 g silica gel. Elute with a gradient of 6:1 to 3:1 (v/v) hexanes/ethyl acetate. Analyze fractions by TLC (see Basic Protocol 1, step 8) using 4:1 hexanes/ethyl acetate. For S.8, Rf = ∼0.35.
9. Combine the appropriate fractions and evaporate to dryness using a rotary evaporator. The bis-protected product (S.8) is prepared as a white foam (161 mg) at 41% yield. BASIC PROTOCOL 3
Development of a Universal Nucleobase and Unnatural Nucleobases
SYNTHESIS OF 3,5-DIMETHYLPHENYL-C-NUCLEOSIDE This protocol describes the synthesis of the 3,5-dimethylphenyl-C-nucleoside, involving the condensation of the Grignard reagent derived from 1-bromo-3,5-dimethylbenzene with the bis-toluoyl-protected chloroglycoside (S.2; Fig. 1.5.1; see Fig. 1.5.5 for the reaction). Deprotection yields the free nucleoside (S.11), followed by generation of the phosphoramidite (S.12) and triphosphate (S.13) derivatives. Two other C-nucleosides are prepared using a similar strategy (TM and 2Nap in Fig. 1.5.6A). Deprotection and formation of the phosphoramidite and triphosphate all follow the general procedures as described in Basic Protocol 1. Materials Argon (see recipe) Magnesium metal turnings Tetrahydrofuran (THF), anhydrous 1-Bromo-3,5-dimethylbenezene (S.9; Aldrich)
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Bis-toluoyl-protected chloroglycoside: 1-α-chloro-3,5-di-O-toluoyl-2-deoxyribofuranose (S.2; Takeshita et al., 1987) Ethyl acetate Saturated ammonium chloride (NH4Cl) solution Sodium sulfate, anhydrous Silica gel, 200-400 mesh, 60 Å Hexanes Ethyl ether, anhydrous Methanol, anhydrous Sodium methoxide (Aldrich or synthesized from sodium metal and methanol) Dichloromethane (CH2Cl2), freshly distilled from calcium hydride Pyridine, anhydrous Triethylamine (TEA), freshly distilled from calcium hydride 4,4′-Dimethoxytrityl chloride (DMTr-Cl; Aldrich) Saturated sodium bicarbonate (NaHCO3) solution 4-Dimethylaminopyridine (DMAP; Aldrich)
Me
Me
TolO O Cl OTol
Br
2
9 a
Me
Me
Me b
TolO
HO O
O OTol
OH
10 c, d
Me
Me
Me
PiPiPiO
O
O
P N(i -Pr)2 O
11 e
Me
DMTr O
O
Me
CN 12
OH 13
Figure 1.5.5 Synthesis of 3,5-dimethylphenyl-C-nucleoside and its phosphoramidite and triphosphate derivatives (see Basic Protocol 3). Reagents: (A) Mg0, then S.9, THF (steps 1 to 9); (B) sodium methoxide, methanol (steps 10 to 15); (C) DMTrCl, TEA, pyridine, CH2Cl2 (steps 16 to 24); (D) 2-cyanoethyl-diisopropylchlorophosphoramidite, triethylamine, CH2Cl2 (steps 25 to 31); (E) ProtonSponge, trimethyl phosphate, POCl3, tributylamine, tributylammonium pyrophosphate, DMF (steps 32 to 38).
Synthesis of Modified Nucleosides
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2-Cyanoethyl diisopropylchlorophosphoramidite (Aldrich) Trimethyl phosphate (Aldrich) Proton-Sponge (Aldrich) Phosphorus oxychloride (POCl3) Tributylamine n-Tetrabutylammonium pyrophosphate (TBAP; Sigma) Dimethylformamide (DMF) Triethylammonium bicarbonate (TEAB; Fluka) Dimethylsulfoxide (DMSO) Isopropanol 25-mL and 100-mL two-neck round-bottom flasks with 14/20 joints, oven-dried Reflux condenser with 14/20 joint Rubber septa 18- and 23-G needles 125-mL separatory funnel Cotton Glass funnel Rotary evaporator (Büchi) equipped with a dry ice condenser and a vacuum system Heavy-walled glass columns (1.5-cm i.d.; 10- and 15-cm length) with glass adapters attached to compressed air or nitrogen source (see flash chromatography steps in APPENDIX 3E) Sea sand 300-mL round-bottom flask Filter paper Lyophilizer (e.g., Labconco freeze-dry system) Additional reagents and equipment for purification and analysis (see Basic Protocol 1) Perform condensation of 1-bromo-3,5-dimethylbenzene (S.9) with bis-toluoyl-protected chloroglycoside (S.2) 1. Assemble a 25-mL oven-dried two-neck round-bottom flask with a 14/20 reflux condenser in the center neck, a rubber septum on the side neck, and a magnetic stirring bar. Maintain a smooth flow of argon using a 23-G needle inserted through another septum on top of the reflux condenser. 2. Quickly add 52 mg (2.139 mmol) magnesium metal and 2 mL THF to the flask under argon atmosphere. 3. Add, in a dropwise fashion, 293 µL (2.156 mmol) 1-bromo-3,5-dimethylbenzene (S.9) and heat the resulting suspension to 50°C for 1 hr using a temperature-controlled oil bath. 4. Prepare a solution of 126 mg (0.324 mmol) bis-toluoyl-protected chloroglycoside (S.2) in 1 mL THF in another round-bottom flask under argon atmosphere. Add all of the prepared Grignard reagent (step 3) to this solution and stir 14 hr at room temperature. 5. During the 14-hr incubation, prepare another batch of Grignard reagent (repeat steps 1 to 3). At the end of the 14-hr incubation, add another 100 µL Grignard reagent to the reaction mixture (step 4) and continue to stir 1 hr at room temperature. Synthesis of S.10 uses 1.0 eq S.9, 0.992 eq magnesium metal, 0.150 eq S.2, and 17.16 eq THF. Development of a Universal Nucleobase and Unnatural Nucleobases
Work up and purify S.10 6. Add 10 mL of ethyl acetate and 10 mL of saturated aqueous NH4Cl and transfer to a 125-mL separatory funnel. Rinse the reaction vessel twice, each time with 5 mL ethyl acetate, and add to the funnel.
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7. Extract separate, and retain the organic layer. Extract the aqueous layer twice, each time with 15 mL of ethyl acetate, and combine these organic layers with the original organic layer. 8. Filter and dry the organic layer (see Basic Protocol, steps 11 to 12). Perform flash chromatography (see Basic Protocol 1, steps 13 to 19). Elute with a gradient of 95:5 to 85:15 (v/v) hexanes/ethyl acetate. Perform TLC (see Basic Protocol 1, step 8) using 9:1 hexanes/ethyl acetate. For S.10, Rf = ∼0.30.
9. Evaporate eluate and analyze product by NMR (see Basic Protocol 1, steps 20 to 22). Approximately 13 mg (9% yield) of purified β-product (S.10) and 48 mg (32%) of α-product should be obtained.
Deprotect 3′- and 5′-OH groups 10. Equip a 100-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G syringe needle threaded through a septum. 11. Add 264 mg (0.576 mmol) S.10 (from step 9) and 10 mL anhydrous methanol to the flask. 12. Add, in a dropwise fashion, 2 mL of 1 M sodium methoxide (2.0 mmol) in methanol to the reaction mixture and keep stirring for 45 min. Synthesis of S.11 uses 1.0 eq S.10, 3.5 eq sodium methoxide, and 514 eq methanol.
Work up and purify S.11 13. Add 100 mg NH4Cl to quench the reaction and continue to stir for 10 min. 14. Evaporate solvents and purify the crude product by flash chromatography (see Basic Protocol 1, steps 13 to 19) using a 1.5 × 10–cm silica gel column packed with 20 g silica gel. Elute with a gradient of 1:99 to 5:95 (v/v) methanol/CH2Cl2. Perform TLC (see Basic Protocol, step 8) using 5:95 methanol/CH2Cl2. For S.11, Rf = ∼0.30.
15. Combine appropriate fractions based on TLC and evaporate to dryness in a rotary evaporator. Flash chromatography gives approximately 105 mg (82% yield) of purified product (S.11) as a white solid.
Tritylate 5′-OH 16. Equip a 100-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G syringe needle threaded through one of the septa. 17. Add 90 mg (0.405 mmol) of S.11 (from step 15), 2 mL anhydrous pyridine, and 2 mL CH2Cl2 to the flask. 18. Add, in a dropwise fashion, 0.30 mL (2.15 mmol) TEA, followed by 175 mg (0.516 mmol) 4,4′-DMTr-Cl in two portions over a 30-min period, and continue to stir for 2 hr at room temperature. Tritylation uses 1.0 eq S.11, 1.27 eq DMTr-Cl, 5.31 eq triethylamine, 61 eq pyridine, and 77 eq dichloromethane.
Work up and purify tritylated product 19. Add 20 mL ethyl acetate and 10 mL saturated NaHCO3 and transfer the mixture to a 125-mL separatory funnel. Rinse the reaction vessel twice, each time with 5 mL ethyl acetate, and add to the funnel.
Synthesis of Modified Nucleosides
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20. Extract separate, and retain the organic layer. Extract the aqueous layer twice, each time with 20 mL ethyl acetate. 21. Combine the organic layers and dry over anhydrous Na2SO4. 22. Filter through filter paper into a 300-mL round-bottom flask and evaporate under reduced pressure in a rotary evaporator. 23. Purify the crude product by flash chromatography (see Basic Protocol 1, steps 13 to 19) using a 1.5 × 10–cm silica gel column packed with 20 g silica gel. Elute with a gradient of 1:1 to 4:1 (v/v) hexanes/ethyl acetate. Perform TLC (see Basic Protocol, step 8) using 1:1 hexanes/ethyl acetate. For the tritylated S.11, Rf = ∼0.3.
24. Combine the appropriate fractions based on TLC analysis and evaporate to dryness in a rotary evaporator. Flash chromatography affords 185 mg (87% yield) of purified DMTr-protected product as a white foam.
Prepare phosphoramidite S.12 by phosphitylation 25. Equip a 100-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G syringe needle threaded through one of the septa. 26. Add 185 mg (0.353 mmol) tritylated product (from step 24) and 3.5 mL anhydrous CH2Cl2 to the flask. 27. Add a catalytic amount (2 mg, 0.016 mmol) of DMAP, followed by 0.350 mL (2.512 mmol) triethylamine and 0.160 mL (0.718 mmol) 2-cyanoethyl diisopropylaminochlorophosphoramidite. 28. Continue to stir for 30 min at room temperature. Synthesis of S.12 uses 1.0 eq tritylated nucleoside, 0.045 eq DMAP, 7.1 eq TEA, 2.0 eq 2-cyanoethyl diisopropylaminochlorophosphoramidite, and 155 eq dichloromethane.
Work up and purify S.12 29. Add 20 mL ethyl acetate and 20 mL saturated NaHCO3 and repeat steps 19 to 22 of this protocol. 30. Purify the crude product by flash chromatography (see Basic Protocol 1, steps 13 to 19). Elute with a gradient of 1:9 to 3:7 (v/v) ethyl acetate/hexane containing 5% triethylamine. Perform TLC using 3:7 ethyl acetate/hexane. For S.12, Rf = ∼0.83.
31. Combine appropriate fractions based on TLC and evaporate to dryness. Flash chromatography affords 238 mg (93% yield) of phosphoramidite product (S.12) as a white foam.
Prepare triphosphate S.13 by phosphorylation of S.11 32. Equip a 25-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G needle threaded through the septum. 33. Add 16 mg (0.072 mmol) free nucleoside (S.11, from step 15) to the flask. 34. Add 0.36 mL (3.08 mmol) trimethyl phosphate and 23 mg (0.107 mmol) ProtonSponge. Cool to 0°C. Development of a Universal Nucleobase and Unnatural Nucleobases
35. Add 8 µL (0.090 mmol) POCl3 in a dropwise fashion and continue to stir for 2 hr at 0°C.
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36. Add 105 µL (0.441 mmol) tributylamine, followed by a solution of 62 mg (0.171 mmol) TBAP in 0.8 mL DMF, and stir for 1 min. Synthesis of S.13 uses 1.0 eq S.11, 43 eq trimethyl phosphate, 1.5 eq Proton-Sponge, 1.3 eq POCl3, 6.1 eq tributylamine, 2.4 eq TBAP, and 143 eq DMF.
Work up and purify S.13 37. Add 7 mL of 1 M TEAB. Dilute the resulting crude solution ∼10 fold with water and lyophilize. 38. Dissolve in 200 mL of 1:1 (v/v) DMSO/isopropanol and purify by HPLC (UNIT 10.5) using the following conditions: Column: Rainin C18-Dynamax 60 Å column Buffer A: 0.1 M TEAB, pH 7.5 Buffer B: Acetonitrile Gradient: 96% to 70% buffer A from 0 to 30 min Flow rate: 10 mL/min. Approximately 1 mg of triphosphate (S.13) is obtained as a white solid after lyophilization.
SYNTHESIS OF 1,4-DIMETHYLNAPHTHALENE-C-NUCLEOSIDE This protocol describes the synthesis of 1,4-dimethylnaphthalene-C-nucleoside (DMN, see Fig. 1.5.6B), as derived from the condensation of the aryllithium species onto an aldehyde followed by a ring-closing reaction, shown in Figure 1.5.7 (Ogawa et al., 2000a). The steps below detail the synthesis of the free nucleoside (S.18). The preparation of the phosphoramidite and triphosphate are performed as described in Basic Protocol 3. A similar strategy may be used to synthesize the 1-methyl-3-naphthalene nucleoside (3MN; Fig. 1.5.6B), starting from 3-bromo-1-methylnaphthalene (Ogawa, et al., 2000b).
BASIC PROTOCOL 4
Materials Argon (see recipe) 2-Bromo-1,4-dimethylnaphthalene (S.14; Aldrich; Sharma, 1993) Tetrahydrofuran (THF), anhydrous n-Butyllithium (Aldrich) Cyclohexane Protected aldehyde of sugar precursor (S.15; Eaton and Millican, 1988) Ethyl acetate Saturated sodium bicarbonate (NaHCO3) solution Sodium sulfate (Na2SO4), anhydrous Silica gel, 200-400 mesh, 60 Å Hexane Ethyl acetate Pyridine, anhydrous Triethylamine (TEA), freshly distilled from calcium Methanesulfonyl chloride Acetic acid, glacial Methanol, anhydrous Trifluoroacetic acid Dichloromethane (CH2Cl2), freshly distilled from calcium hydride 2,3-Dichloro-5,6-dicyano-1,4-benzoquinone (DDQ; Aldrich) 25-mL, 50-mL, and 100-mL two-neck round bottom flasks with 14/20 joints, oven-dried Rubber septa 18-G needles
Synthesis of Modified Nucleosides
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Me
A
Me
4
Me
Me
3
5
2 6
Me
1
TM (trimethylbenzene)
DM (dimethylbenzene)
2Nap (2-naphthalene)
6 7
5
B
Me
4
Me
8
3 2
1
Me
3MN (1-methyl-3-naphthalene)
DMN (dimethylnaphthalene)
C
Me
2MN (3-methyl-2-naphthalene)
Figure 1.5.6 Unnatural hydrophobic C-nucleosides. Synthetic strategies are presented in (A) Basic Protocol 3, (B) Basic Protocol 4, and (C) Alternate Protocol.
OPMB
Me
DMTrO CHO OPMB 15
Me Br 14
a,b
Me
Me Me
DMTrO
c
Me
HO
O OPMB
O 16
OPMB
17
d
Me Me
HO O OH
Development of a Universal Nucleobase and Unnatural Nucleobases
18
Figure 1.5.7 Synthesis of 1,4-dimethylnaphthalene-C-nucleoside (see Basic Protocol 4). Reagents: (A) n-butyllithium, −78°C, then S.15, THF (steps 1 to 8); (B) methanesulfonyl chloride, triethylamine, pyridine, 0°C (steps 9 to 14); (C) acetic acid (steps 15 to 20); (D) DDQ (steps 21 to 26).
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Dry ice/acetone bath (−78°C) Cotton Glass funnel Rotary evaporator (Büchi) equipped with a dry ice condenser and a vacuum system Heavy-walled glass columns (1.5-cm i.d.; 10-, 15-, and 20-cm length) with glass adapters attached to compressed air or nitrogen source (see flash chromatography steps in APPENDIX 3E) Sea sand Additional reagents and equipment for purification and analysis (see Basic Protocol 1) NOTE: n-Butyllithium is flammable and sensitive to moisture. It should be handled carefully and quickly. Perform condensation of 2-bromo-1,4-dimethylnaphthalene (S.14) with protected aldehyde of sugar precursor (S.15) 1. Equip a 100-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G needle threaded through one of the septa. 2. Add 640 mg (2.72 mmol) 2-bromo-1,4-dimethylnaphthalene (S.14) and 15 mL THF to the flask under argon atmosphere and cool to −78°C in a dry ice/acetone bath. 3. Slowly add dropwise 2.0 mL of 2 M n-butyllithium (4.0 mmol) in cyclohexane and continue to stir for 15 min at −78°C. 4. Add, in a dropwise fashion, a solution of 1.214 g (1.801 mmol) of the protected aldehyde of the sugar precursor (S.15) in 5 mL THF and continue to stir for 1 hr at −78°C. 5. Remove the flask from the bath and warm to room temperature over 1.5 hr. Synthesis of the 1,4-dimethylnaphthalene derivative uses 1.0 eq S.14, 1.47 eq n-butyllithium, 0.66 eq S.15, and 90 eq THF.
Work up and purify product 6. Add 10 mL ethyl acetate and 10 mL saturated NaHCO3 to partition. Filter and dry the organic layer (see Basic Protocol 1, steps 11 and 12). 7. Perform flash chromatography (see Basic Protocol 1, steps 13 to 19). Elute with a gradient of 9:1 to 8:2 (v/v) hexane/ethyl acetate. Perform TLC (see Basic Protocol 1, step 8) using 7:3 hexane/ethyl acetate. For this product, Rf ∼0.23.
8. Combine the appropriate fractions based on TLC and evaporate to dryness with a rotary evaporator. Flash chromatography gives the C1 S-isomer (712 mg, 48% yield) and the C1 R-isomer (625 mg, 42% yield). S and R isomers were assigned based on the α or β anomer that was formed after cyclization.
Perform in situ cyclization of mesylate 9. Equip a 25-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G syringe needle threaded through one of the septa. 10. Add 712 mg (0.857 mmol) of the S-diastereomer (step 8) and 9 mL anhydrous pyridine to the flask and cool to 0°C.
Synthesis of Modified Nucleosides
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11. Add 0.57 mL (4.09 mmol) triethylamine followed by 90 µL (1.17 mmol) methanesulfonyl chloride, and warm slowly to room temperature over 1.5 hr with stirring. Synthesis of S.16 uses 1.0 eq S-diastereomer, 4.8 eq triethylamine, 1.37 eq methanesulfonyl chloride, and 130 eq pyridine.
Work up and purify S.16 12. Add 2 mL saturated NaHCO3 to quench the reaction. 13. Perform flash chromatography (see Basic Protocol 1, steps 13 to 19). Elute with a gradient of 95:5 to 85:15 (v/v) hexane/ethyl acetate. Perform TLC using 7:3 hexane/ethyl acetate. For S.16, Rf = ∼0.68.
14. Collect appropriate fractions based on TLC and evaporate to dryness with a rotary evaporator. Flash chromatography yields 176 mg (30% yield) of purified product (S.16).
Detritylate 5′-OH group 15. Equip a 25-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G syringe needle threaded through one of the septa. 16. Add 190 mg (0.274 mmol) S.16, 6 mL acetic acid, and 1 mL methanol to the flask. 17. Add 1 mL trifluoroacetic acid in a dropwise fashion (∼10 drops) and continue stirring for 20 min at room temperature. Detritylation uses 1.0 eq S.16, 383 eq acetic acid, 90 eq methanol, and 47.4 eq trifluoroacetic acid.
Work up and purify S.17 18. Concentrate the reaction solution with a rotary evaporator. 19. Remove solvents (see Basic Protocol, step 12) and perform flash chromatography (see Basic Protocol 1, steps 13 to 19). Elute with a gradient of 4:1 to 1:1 (v/v) hexane/ethyl acetate. Perform TLC using ethyl acetate. For S.17, Rf = ∼0.75.
20. Combine appropriate fractions based on TLC and evaporate to dryness with a rotary evaporator. Flash chromatography affords 93 mg (86% yield) of purified product (S.17).
Remove 3′-PMB group 21. Equip a 50-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G syringe needle threaded through one of the septa. 22. Add 8 mg, 0.020 mmol S.17, 1 mL CH2Cl2, and 1 drop H2O to the flask. 23. Add 7 mg (0.031 mmol) DDQ and continue to stir for 1 hr at room temperature. Deprotection uses 1.0 eq S.17, 780 eq CH2Cl2, 277.5 eq H2O, and 1.55 eq DDQ.
Development of a Universal Nucleobase and Unnatural Nucleobases
Work up and purify S.18 24. Add 15 mL ethyl acetate and 15 mL saturated NaHCO3 to partition. Filter and dry the organic layer (see Basic Protocol 1, steps 11 and 12).
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25. Perform flash chromatography (see Basic Protocol 1, steps 13 to 19). Elute with a gradient of 1:99 to 5:95 (v/v) methanol/CH2Cl2. Perform TLC using 3:97 methanol/CH2Cl2. For S.18, Rf = ∼0.3.
26. Combine appropriate fractions based on TLC and evaporate to dryness using rotary evaporator. Flash chromatography affords 3 mg (55% yield) of pure product (S.18).
SYNTHESIS OF 3-METHYL-2-NAPHTHALENE-C-NUCLEOSIDE This protocol describes the synthesis of the 3-methyl-2-naphthalene-C-nucleoside (Fig. 1.5.6C), which was derived from the condensation of the aryllithium species onto another aldehyde followed by in situ cyclization as shown in Figure 1.5.8 (Ogawa et al., 2000a). The preparation of phosphoramidite and triphosphate follows the general procedures described in Basic Protocol 3.
ALTERNATE PROTOCOL
Additional Materials (also see Basic Protocols 3 and 4) 2-Bromo-3-methylnaphthalene (S.19; prepared according to Lambert et al., 1979) Protected aldehyde of sugar precursor (S.20, prepared according to Solomon and Hopkins, 1993) Perform condensation of 2-bromo-3-methylnaphthalene (S.19) with the protected aldehyde of the sugar precursor (S.20) 1. Place a magnetic stir bar in a dry 100-mL two-neck flask with two rubber septa, and place the flask on top of a magnetic stir plate. 2. Evacuate the reaction flask on vacuum line and then flush it with argon. Repeat this procedure three times and attach the flask to an argon line on the manifold. 3. Add 595 mg (2.70 mmol) 2-bromo-3-methylnaphthalene (S.19) and 8.9 mL THF to the flask under argon atmosphere and cool to −78°C in a dry ice/acetone bath. 4. Add dropwise 1.50 mL of 2 M n-butyllithium (3.0 mmol) in cyclohexane over 10 min and continue to stir 15 min at −78°C. 5. Add dropwise a solution of 502 mg (1.74 mmol) aldehyde (S.20) in 4.4 mL THF over 5 min and continue to stir 30 min at −78°C. 6. Remove the flask from the bath and warm to room temperature. During this time the reaction color changes from deep green to brown. Condensation uses 1.0 eq S.19, 1.1 eq n-butyllithium, 0.64 eq S.20, and 61 eq THF.
O
a,b,c
O Me Br 19
CHO OTBS 20
Me
HO O
21 OH
Figure 1.5.8 Synthesis of 3-methyl-2-naphthalene-C-nucleoside (see Alternate Protocol). Reagents: (A) n-butyllithium, −78°C, then S.2, THF (steps 1 to 9); (B) methanesulfonyl chloride, triethylamine, pyridine, 0°C (steps 10 to 16); (C) TFA (steps 17 to 22).
Synthesis of Modified Nucleosides
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Work up and purify product 7. Add 10 mL ethyl acetate and 10 mL saturated NaHCO3 to partition. Filter and dry the organic layer (see Basic Protocol 1, steps 11 and 12). 8. Perform flash chromatography (see Basic Protocol 1, steps 13 to 19). Elute with a gradient of 9:1 to 5:1 (v/v) hexane/ethyl acetate. Perform TLC (see Basic Protocol 1, step 8) using 9:1 hexane/ethyl acetate. For this product, Rf = ∼0.30.
9. Combine appropriate fractions based on TLC and evaporate to dryness with a rotary evaporator. Flash chromatography gives a crude mixture of both diastereomers (532 mg, 1.24 mmol).
Mesylate hydroxy group 10. Equip a 100-mL two-neck flask with two rubber septa and a stir bar. Keep the vessel under argon using an 18-G syringe needle threaded through one of the septa. 11. Add 510 mg (1.19 mmol) product (step 9, both diastereomers) and 32 mL CH2Cl2 to the flask and cool to 0°C. 12. Add 0.332 mL (2.38 mmol) triethylamine, followed by 0.120 mL (1.55 mmol) methanesulfonyl chloride, and warm slowly to room temperature over 30 min with stirring. Relative to S.19 (step 3), mesylation uses 0.88 eq triethylamine, 0.57 eq methanesulfonyl chloride, and 185 eq CH2Cl2.
Work up and purify product 13. Add 30 mL saturated aqueous NaHCO3 and transfer to a 125-mL separatory funnel. Rinse the reaction vessel twice, each time with 5 mL ethyl acetate and add to the funnel. 14. Separate the organic layer and extract the aqueous layer twice, each time with 15 mL ethyl acetate. 15. Filter and dry the organic layer (see Basic Protocol 1, steps 11 and 12), and perform flash chromatography (see Basic Protocol 1, steps 13 to 19). Elute with a gradient of 95:5 to 85:15 (v/v) hexanes/ethyl acetate. Perform TLC using 9:1 hexanes/ethyl acetate. For this product, Rf = ∼0.30.
16. Evaporate eluate and analyze product by NMR (see Basic Protocol 1, steps 20 to 22). The resulting crude product is pure enough for use in the next step.
Perform in situ cyclization of mesylate 17. Set up the reaction flask as in steps 1 and 2 above. 18. Add 605 mg (1.19 mmol) crude product (step 16) and 56 mL of 4:1 (v/v) TFA/methanol and stir 20 min at room temperature. Relative to S.19 (step 3), cyclization uses 216 eq trifluoroacetic acid and 103 eq methanol.
Development of a Universal Nucleobase and Unnatural Nucleobases
Work up and purify S.21 19. Concentrate the solution with a rotary evaporator and then neutralize the residual TFA with a minimum volume of saturated NaHCO3. 20. Transfer to a separatory funnel and extract four times, each time with 20 mL CH2Cl2.
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21. Filter and dry the organic layer (see Basic Protocol 1, steps 11 and 12), and perform flash chromatography (see Basic Protocol 1, steps 13 to 19). Elute with 3:97 (v/v) methanol/CH2Cl2. Perform TLC using the same solvent For S.21, Rf = ∼0.2.
22. Collect the appropriate fractions and evaporate to dryness with a rotary evaporator. Flash chromatography affords the desired β-anomer (S.21) at 108 mg (0.418 mmol, 24% yield over 3 steps).
PURIFICATION OF DNA OLIGONUCLEOTIDES Oligonucleotides containing an unnatural base are synthesized using an ABI 392 DNA/RNA synthesizer with automatic removal of the 5′-trityl protecting group (so-called trityl-off procedure; APPENDIX 3C). The oligonucleotides are then deprotected and cleaved from the CPG support according to the instrument instruction manual. Figure 1.5.9 shows the sequences of four oligonucleotides that are used in the thermodynamic and kinetic analyses described in Basic Protocols 6 and 7. dC-CPG (500Å, 1.0 µmol) is used for S.22, dG-CPG (500 Å, 1.0 µmol) is used for S.23, dA-CPG (500 Å, 1.0 µmol) is used for S.24, the universal support (500Å, 1.0 µmol) is used for S.25 and dA-CPG (500Å, 1.0 µmol) is used for S.26. To be suitable for use in Basic Protocols 6 and 7, the oligonucleotides are purified by preparative denaturing PAGE (UNIT 10.4 and APPENDIX 3B), excised from the gel, electroeluted using a Schleicher & Schuell Elutrap electrophoresis chamber, and ethanol precipitated to concentrate and desalt the sample. The oligonucleotides are stored at −20 °C.
BASIC PROTOCOL 5
Materials CPG-bound DNA oligonucleotide (APPENDIX 3C) Concentrated ammonium hydroxide 95% (v/v) formamide in 10 mM Tris⋅Cl, pH 8.5 (APPENDIX 2A), with and without 0.05% (w/v) xylene cyanol and bromphenol blue TBE buffer (see APPENDIX 2A) Absolute ethanol, prechilled to –20°C 5 M NaCl, ice cold 80% ethanol, ice cold 1.5-mL screw-cap vial 60° and 80°C Dri-baths (Thermolyne) Speed-Vac evaporator (Savant) 50-mL polypropylene centrifuge tubes UV/Vis spectrophotometer Electroelution unit (Schleicher and Schuell) Silica gel–coated preparative TLC plate UV lamp (hand held) Refrigerated centrifuge and rotor appropriate for sample size Additional reagents and equipment for denaturing PAGE (UNIT 10.4 and APPENDIX 3B) Deprotect oligonucleotide and cleave from support 1. Transfer the CPG-bound DNA oligonucleotide to a 1.5-mL screw-cap vial. 2. Add 1.5 mL concentrated ammonium hydroxide, tighten the cap, and incubate 12 hr at 80°C (for 24-mer) or 60°C (for 13-mer or 45-mer). 3. Cool to 0°C and evaporate to dryness in a Speed-Vac evaporator.
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22 23 24 25 26
5 ′-dCGCATGXGTACGC 5 ′-dGCGTACXCATGCG 5 ′-dTAATACGACTCACTATAGGGAGA 5 ′-dTAATACGACTCACTATAGGGAGAX 5 ′-dCGCTAGGACGGCATTGGATCG XTCTCCCTATAGTGAGTCGTATTA
Figure 1.5.9 Oligonucleotides used for characterization of unnatural nucleobases.
Purify crude oligonucleotide 4. Add 300 µL water and 300 µL of 95% formamide/10 mM Tris⋅Cl, pH 8.5 (without xylene cyanol and bromphenol blue) and mix thoroughly. 5. Prepare a 12-well denaturing PAGE gel (UNIT according to the length of the oligonucleotide:
10.4
and
APPENDIX 3B)
as follows,
13-mer: 15% acrylamide; 42 cm long × 33 cm wide × 1.5 mm thick 24-mer: 20% acrylamide; 42 cm long × 33 cm wide × 1.5 mm thick 45-mer: 10% acrylamide; 42 cm long × 33 cm wide × 1.5 mm thick. 6. Load 60 µL of 95% formamide/10 mM Tris⋅Cl, pH 8.5, with 0.05% (w/v) xylene cyanol and bromphenol blue in each of the two outermost wells; load 60 µL of the sample in each of the ten innermost wells. 7. Purify the DNA to single-nucleotide resolution by denaturing PAGE (UNIT 10.4 and APPENDIX 3B) at 65 W until the full-length oligonucleotide has migrated two-thirds of the way down the gel, as indicated by the dye markers. 8. Excise the full-length DNA product by UV shadowing (UNIT 10.4). Isolate, desalt, and concentrate pure oligonucleotide 9. Place the gel slice in a 50-mL polypropylene centrifuge tube and crush thoroughly with a glass stirring rod. 10. Add 1× TBE electrophoresis buffer mix, and pour the gel slurry into the sample chamber of a properly assembled and oriented electroelution apparatus, according to the instrument instruction manual. 11. Electroelute the sample and monitor its progress by withdrawing small samples and measuring absorbance at 260 nm (A260). 12. Combine the most concentrated fractions. 13. Add 3 vol absolute ethanol (prechilled to –20°C) and 1⁄20 vol 5 M NaCl and freeze at −20°C for at least 30 min. 14. Centrifuge for at least 20 min at >15,000 × g, 4°C. 15. Carefully decant the supernatant, wash the DNA pellet twice with ice cold 80% ethanol, and allow it to air dry. 16. Dissolve DNA in an appropriate volume of sterile deionized water for determination of concentration and downstream applications. Development of a Universal Nucleobase and Unnatural Nucleobases
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DETERMINATION OF THERMODYNAMIC STABILITY OF UNNATURAL BASE PAIRS Thermal stability or thermodynamic stability refers to the melting temperature (Tm) of a DNA duplex that contains the unnatural base pair. Thermodynamic selectivity refers to the difference in Tm between the desired unnatural base pair and the most stable mispair of an unnatural base with a natural base. Oligonucleotides S.22 and S.23 (Fig. 1.5.9) provide the sequence context for determination of the stability and selectivity of the unnatural nucleosides. Duplex oligonucleotide samples are prepared with the unnatural nucleoside in position X of, for example, S.22 and paired against S.23 containing each natural base as well as the same unnatural base (self-pair) and/or any other unnatural base to be tested. The complete set of pairs is also prepared with the unnatural nucleoside in S.23 to determine if there are sequence context effects (e.g., CXT versus TXC). The samples are heated and evaluated for temperature-dependent changes in absorption using a Cary 300 Bio UV-Vis spectrophotometer. Duplex melting curves are collected in triplicate, and melting temperatures are determined by the derivative method contained in the Cary WinUV-Vis software. For a more detailed description of Tm experiments, see UNIT 7.3.
BASIC PROTOCOL 6
Materials Complementary 13-mer oligonucleotides (e.g., S.22 and S.23 in water; see Basic Protocol 5) 2× Tm buffer (see recipe) Variable-temperature double-beam spectrophotometer (Cary 300 BIO UV-Vis spectrophotometer) with optically matched cuvettes 1. Prepare sample cells with 3 µM of each complementary oligonucleotide in 1× Tm buffer. Prepare reference cells with 1× Tm buffer alone. 2. Arrange cells in the spectrophotometer and monitor the sample absorbance at 260 nm while heating at a rate of 0.5°C/min from 20° to 80°C. 3. Collect three curves for each sample and analyze data by the first-derivative method provided with the instrument software. KINETIC ANALYSIS OF UNNATURAL BASE PAIR INCORPORATION, SELECTIVITY AND REPLICABILITY A rapid gel kinetic assay is used for the determination of steady-state parameters for the incorporation and extension of the unnatural base pairs, as well as to evaluate the selectivity, or fidelity of these steps. Initial velocities are determined by radiolabeled primer extension reactions at various concentrations of 5′-nucleoside triphosphate (natural or unnatural). To ensure single completed hit conditions, a final ratio of 30:1 primertemplate to enzyme is used, and less than 20% of the primer is extended. An example of a primer-template set for determining incorporation kinetics is shown in Figure 1.5.9 with S.24 as the primer and S.26 as the template. The reactions can be rapidly initiated and quenched by hand and analyzed by denaturing PAGE; a PhosphorImager (Molecular Dynamics) is used to quantify gel band intensities corresponding to the extended primer. Velocities are plotted against triphosphate concentration and fit to a rectangular hyperbola. Preliminary determination of KM is followed by kinetic analysis using triphosphate concentrations evenly distributed about the KM. Unnatural nucleosides are characterized both in the template strand and as the incoming triphosphate. Kinetic data are collected in triplicate. In general, these kinetic determinations follow the experimental protocols developed by Goodman and co-workers (Creighton et al., 1995).
BASIC PROTOCOL 7
Synthesis of Modified Nucleosides
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Materials 4 µM DNA oligonucleotide primer (e.g., S.24; see Basic Protocol 5) 10 U/µL T4 polynucleotide kinase and 10× buffer (New England Biolabs) 10 mCi/mL [γ-33P]ATP (2500 Ci/mmol) QIAquick Nucleotide Removal Kit (Qiagen) 1 µM DNA oligonucleotide template (e.g., S.26; see Basic Protocol 5) 10 U/µL exonuclease-free Klenow fragment (Amersham Pharmacia Biotech) and 10× random-prime buffer (see recipe) Unnatural triphosphates (see Basic Protocols 1 to 4 and Alternate Protocol) 100 mM stock of each natural dNTP (Amersham Pharmacia Biotech; sequencing grade) Enzyme dilution buffer (see recipe) Quench solution (see recipe) 1× TBE buffer (APPENDIX 2A) 37°, 25°, and 100°C Dri-Baths (Thermolyne) with heating blocks that accommodate 1.5- or 0.5-mL tubes Thin putty knife Geiger counter Chromatography paper (Whatman 3MM CHR, 35 × 45 cm) Gel dryer (Bio-Rad Model 583) Storage Phosphor Screen (Kodak or Molecular Dynamics, 35 × 43 cm) PhosphorImager (e.g., STORM Model 860; Molecular Dynamics) ImageQuant and compatible spreadsheet (MS Excel) and graphing (Kaleidagraph) software Additional reagents and equipment for denaturing PAGE (UNIT 10.4 and APPENDIX 3B) NOTE: Use autoclaved deionized water for all reagents. Radiolabel primer 1. Combine 25 µL of 4 µM oligonucleotide primer with 3.1 µl of 10× T4 polynucleotide kinase buffer, 2 µL of [γ-33P]ATP stock, and 1 µL of T4 polynucleotide kinase. Mix and incubate at 37°C for at least 30 min. 2. Remove excess nucleotide with the QIAquick Nucleotide Removal Kit, according to the manual, and observing proper radioactive waste disposal. 3. Dilute buffer EB (from QIAquick kit) five-fold and use 100 µL to elute the DNA. Store at 4°C. Allow the wetted column to sit for a few minutes before the final spin. Assuming 80% recovery of the labeled primer, this protocol yields 100 ìL of 800 nM 5′-radiolabeled primer.
Prepare analytical gel 5. Pour a 15% denaturing polyacrylamide gel (UNIT 10.4 and APPENDIX 3B) that is 42 cm long, 33 cm wide, and 0.5 mm thick, and has 34 wells. Make sure there are no trapped air bubbles. Allow the gel at least 1 hr to fully polymerize before use.
Development of a Universal Nucleobase and Unnatural Nucleobases
Anneal primer and template 6. Combine 25 µL of water, 10 µL of 10× random-prime buffer, 8 µL of 1 µM template oligonucleotide, and 5 µL of 5′-radiolabeled 800 nM primer oligonucleotide (from step 1) in a 1.5-mL microcentrifuge tube, mix well, then microcentrifuge briefly at maximum speed to bring all the solution to the bottom of the tube. This gives a two-fold excess of template over primer, necessary to ensure that greater than 95% of the primer is annealed to the template. The use of different primer or template sequences may require that this ratio be optimized.
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7. Place the solution in a Dri-bath preheated to 100°C and immediately turn off the heating element. Allow the primer-template to anneal by slow cooling to room temperature in the Dri-bath (∼2 hr). This yields a final volume of 48 ìL, which, when combined with 2 ìL of enzyme, will yield sufficient volume for nine reactions of different triphosphate concentrations. Although it is best to use freshly annealed primer-template, annealed primer-templates can be stored at 4°C overnight. Warm to room temperature before use.
Perform reactions 8. Prepare 5-µL aliquots of 2× concentrations of the desired triphosphate range by combining the appropriate volume of 1 µM, 10 µM, 100 µM, 1 mM, or 10 mM of the previously diluted dNTP and water to a final volume of 5 µL. For the incorporation experiments, the unnatural triphosphate is used (in conjunction with the primer-template complex S.24-S.26) to determine the rate of unnatural base pair synthesis. To measure incorporation selectivity, each of the four natural triphosphates is used in conjunction with this primer-template. For the extension experiments, including both extension efficiency as well as extension fidelity, each of the four natural triphosphates is used in conjunction with primer-template S.25-S.26. It is a good idea to make a reasonably large number of small-volume aliquots of the various concentrations of dNTP to reduce the number of freeze-thaw cycles to which the dNTP is subjected. It is generally desirable to run a control for each experiment with only 5 ìL of water and no triphosphate.
9. Add 2 µL of the appropriate concentration of Klenow fragment freshly diluted in enzyme dilution buffer to cooled annealed primer-template. Mix thoroughly, but do not vortex. This yields 50 ìL of a 2× reaction solution that is 80 nM in primer-template, 2× in all buffer components, and 0.04× of the concentration of diluted enzyme used. This 2× reaction solution will be diluted to 1× when combined with the triphosphate aliquots to initiate the reaction. To ensure single completed hit conditions, make certain that the final enzyme concentration is no more than 1⁄30 that of the final primer-template concentration, final concentration referring to that achieved in step 10.
10. Add 5 µL enzyme-primer-template solution (from step 9) to each 5-µL aliquot of 2× triphosphate (from step 8), mix by pipetting up and down, close the tube, and place in a 25°C Dri-bath. Initiate all reactions in turn as quickly as possible (≤10 sec between starts). 11. Quench the reactions in turn with 20 µL quench solution precisely when the timer reaches the desired length for the experiment. Perform electrophoresis 12. Warm gel (prepared in step 5) by running it at 65 W in 1× TBE buffer until it reaches a temperature of at least 37°C. Wash the wells by pipetting 1× TBE into each a couple of times. 13. Load 8 µL quench solution into the three outermost lanes as well as any unused lanes. Load 8 µL of each quenched reaction into up to 28 of the innermost lanes. Run at 65 W. Up to 28 reactions can be loaded in one row of the gel. Four rows (112 reactions) can be run in one gel by allowing the first row to run at 65W for 30 min, stopping the gel and loading another row. The second row is run for 25 min, the third row for 20 min, and the fourth row until the dark blue dye band of the second row exits the gel. The wells are washed prior to loading each of the four rows.
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14. Remove the gel from the apparatus and cool it under running water. Separate the plates by using a thin putty knife, taking care to notice which plate the gel will remain stuck to and allowing it to do so. 15. Find the region of the gel containing the extended primers by surveying with a Geiger counter. 16. Place a large piece of plastic wrap over the gel, smooth it out, and cut the relevant area. 17. Carefully transfer the gel (covered on one side with plastic wrap) to a 35 × 45–cm sheet of chromatography paper, gel-side down, and smooth it out. Dry the gel completely (time may vary with apparatus) and allow it to cool nearly to room temperature. 18. Cut the chromatography paper down to size, if necessary, and transfer the gel to a storage phosphor screen, plastic-wrap side facing the screen. Allow image to develop overnight. Analyze data 19. Remove the gel from the storage screen and read the plate with a PhosphorImager. 20. Using ImageQuant software, draw rectangles around primer and extended primer and obtain a volume report of the quantitative data from the gel quantitative data. Use the control reaction (no triphosphate) as the background to be subtracted in each series of triphosphate concentrations. 21. Calculate the fraction of primer extended at each triphosphate concentration as: v = Ip/(I0 + Ip) where I0 is the primer band count and Ip is the product band count. 22. Subtract the value obtained for the control reaction from each reaction. This will force the data set to the origin. Make sure that <20% of the primer is extended at the highest triphosphate concentration to ensure single completed hit conditions.
23. Multiply each corrected value by the final concentration of primer template (40 nM) and divide by the final concentration of enzyme and the reaction time. This gives kcat values in min−1. 24. Plot kcat against triphosphate concentration and fit the curve to the nonlinear form of the Michaelis-Menten equation to obtain kcat and KM. 25. After a preliminary determination of KM, adjust the experimental conditions to obtain data points in the Michaelis plot that evenly flank the experimentally determined KM to triphosphate concentrations ∼5-fold under and over this value. 26. Collect data in triplicate and average the values for kcat and KM.
Development of a Universal Nucleobase and Unnatural Nucleobases
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Ammonium molybdate solution 2.0 g ammonium molybdate 7.0 g ceric sulfate 250 mL 10% (v/v) H2SO4 Store indefinitely at 4°C Argon Argon should be purged through two Drierite desiccators and then introduced into the reaction flask via Tygon tubing installed on a manifold. To ensure a slightly positive pressure of argon in the reaction flask, flow rates should be adjusted by regulators on the argon tank so that a slow stream of argon bubbles (one bubble per sec) is obtained in the bubbler connected to the manifold. Enzyme dilution buffer 50 mM potassium phosphate, pH 7.0 (APPENDIX 2A) 1 mM DTT 0.1 mM EDTA 50% (v/v) glycerol Store up to 6 months at −20°C Quench solution 95% formamide 5 mM Tris⋅Cl, pH 8 (APPENDIX 2A) 20 mM EDTA 0.05% (w/v) xylene cyanol 0.05% (w/v) bromphenol blue Random-prime buffer, 10× 500 mM Tris-Cl, pH 7.5 (APPENDIX 2A) 100 mM MgCl2 10 mM DTT 500 µg/mL acetylated BSA Store indefinitely at −20°C Sodium hydride, 60% Purchase sodium hydride (NaH) as a 60% dispersion in mineral oil (Aldrich). Add to a flask and then add hexanes with a pipet. Mix to rinse and remove hexanes with a pipet. Repeat several times until the mineral oil is completely removed, then remove residual hexanes thoroughly using a high-vacuum oil pump. Tm buffer, 2× 20 mM PIPES, pH 7 20 mM MgCl2 200 mM NaCl Store indefinitely at room temperature
Synthesis of Modified Nucleosides
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COMMENTARY Background Information
Development of a Universal Nucleobase and Unnatural Nucleobases
Synthesis of unnatural nucleosides The first synthesis of natural nucleosides was published in the late 1940s. Since then, nucleoside synthesis has enjoyed a long and rich history. To date, there are five typical methods for the synthesis of N-nucleosides (Vorbrüggen and Ruh-Pohlenz, 2001): (1) coupling of heterocyclic salts with halo sugars; (2) SilylHilbert-Johnson reaction followed by Vorbrüggen procedure; (3) addition of heterocycle to glycals; (4) derivation of glycosylamine derivatives; and (5) enzymatic transglycosylation. The first two methods have proven to be the most useful for nucleosidic bond formation. The use of sodium salts (Kazimierczuk et al., 1984) gives considerably higher yield of nucleoside product and is a significant improvement upon the Fischer-Helferich protocol (Fischer and Helferich, 1914), which uses toxic mercuric reagents. The reaction of peracylated sugars with persilylated heterocyclic nucleobases in the presence of a Lewis acids such as SnCl4 (Niedballa and Vorbrüggen, 1974) or trimethylsilyl triflate (TMSOTf; Vorbrüggen et al., 1981) has become a routine method for the synthesis of pyrimidine and purine nucleosides. TMSOTf is often the catalyst of choice, as it gives higher yields of β anomer and avoids the emulsion problems associated with tin residue during workup. Natural and modified C-nucleosides, in addition to being candidates for unnatural bases with interesting thermal or replication properties, have attracted great interest due to their anticancer and antiviral activities (Postema, 1992; Chaudhuri et al., 1997). There are many methods for synthesizing aromatic N- and Cnucleosides. For the purpose of preparing Cnucleoside-containing oligonucleotides, two main approaches have been used: (1) nucleophilic displacement of a halogen group from C1 of the deoxyribose by aryl Grignard reagents (Chaudhuri et al., 1997; Ogawa et al., 2000a) and (2) C-nucleophilic addition to the aldehyde of the deoxyribose precursor (Solomon and Hopkins, 1993; Eaton and Millican, 1988). The merit of the former method is its simplicity and the fact that the desired free nucleoside is directly obtained after deprotection of the hydroxy group. The latter method generally affords the desired product in higher yield than the former. As an alternative method for synthesis of C-nucleoside, ribonolactone
has been used with aryl reagents (Hildbrand et al., 1997). The step of nucleophilic addition in this method results in the intermediate formation of a corresponding hemiacetal, which selectively yields the β anomer upon reduction. Removing the 2′-hydroxy group, however, requires additional steps. Universal bases and unnatural base pairs Unnatural nucleosides that possess hydrophobic base analogs have been shown to form stable pairs in duplex DNA, and have also been demonstrated to be efficiently recognized by DNA polymerases, both when present in a template and as a nucleoside triphosphate. In fact, a wide variety of such unnatural bases, with no H bonding or shape complementarity, are surprisingly able to replace a natural base in duplex DNA or during enzymatic replication. These bases are not constrained to resemble the natural purines or pyrimidines, and a much wider variety of design strategies may be pursued. A variety of predominantly hydrophobic nucleobases have been developed with properties such as stable but nonselective pairing in duplex DNA (universal thermodynamic base), efficient but nonselective polymerase recognition (universal kinetic base), or orthogonal stability and replication (third base pair for expanding the genetic code). This unit describes the synthesis of both Nand C-nucleosides. The N-nucleosides retain the natural glycosidic linkage and maintain the minor groove carbonyl group hydrogen bond acceptor. The minor groove hydrogen bond acceptor may be important for stability in duplex DNA by binding water molecules, and also for polymerase recognition. The C-glycosides are aromatic, in order to retain planarity, and, to date, those examined (i.e., TM, DM, 2Nap, DMN, 3MN, and 2MN) do not possess minor groove hydrogen bond acceptors. The current set of bases has resulted from the derivatization of several scaffoldings that were originally found to be successful: the isocarbostyril framework (Fig. 1.5.2), the 7azaindole framework (Fig. 1.5.4), and the aromatic ring framework (Fig. 1.5.6). The thermodynamic and kinetic characterization of the modified scaffold structures was used to direct their further derivatization in a largely empirical fashion. Although this process has been largely empirical, a few general trends are beginning to emerge (McMinn et al., 1999; Berger
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et al., 2000, 2002; Ogawa et al., 2000a,b; Wu et al., 2000; Tae et al., 2001). The development of universal bases that pair nondiscriminantly opposite any natural base would have useful applications for the design of hybridization probes where sequence ambiguities exist. There has been some success in the design of nucleobase analogs that can hybridize nonselectively to each of the native bases (reviewed in Loakes, 2001), but analogs that can universally hybridize without significant duplex destabilization are rare. Hydrophobic base analogs without H-bonding groups that pack efficiently in duplex DNA show little selectivity in pairing with native bases and are promising universal bases (Berger et al., 2000). For example, MICS and 5MICS (Fig. 1.5.2) form stable base pairs with each of the natural bases, when incorporated into the central position of either oligodeoxyribonucleotide strand of a 13-mer duplex (S.22 and S.23, shown in Fig. 1.5.9). When incorporated into S.22 at position X (Figure 1.5.9) and paired opposite each of the four natural bases in S.23, the average Tm for MICS and 5MICS is 54.6 ± 0.7°C and 55.3 ± 0.6°C, respectively. This compares favorably with the stability of the same duplex with a native dT:dA pair (Tm = 58.7°C). The spread in measured Tm values for both MICS and 5MICS opposite each natural base is the same, 1.4°C. A single MICS or 5MICS paired opposite any natural base is significantly more stable than a mispair in the same sequence context (∆Tm = 3.8° to 17°C for mispairs among the native bases in the same sequence context). When MICS and 5MICS are examined in the opposite strand (position X in S.23), paired opposite natural bases, the measured ∆Tm’s are slightly larger (∆Tm = 1.9° and 2.2°C for MICS and 5MICS, respectively). Less effort has been directed toward the identification of universal bases for DNA replication (Berger et al., 2000). No base analog is known that can act as a universal template for incorporation of each natural nucleoside triphosphate. Such bases would have important applications in the generation of oligonucleotide libraries. A variety of predominantly hydrophobic nucleobases that efficiently direct Klenow fragment to incorporate native triphosphates have been identified. Despite a compromise in absolute rate, when present in template, MICS directs the insertion of each native triphosphate with approximately equal efficiency (Berger et al., 2000). Each natural triphosphate is inserted by Klenow fragment opposite MICS with an efficiency of 5.8 × 103
to 2.4 × 104 M−1 min−1, with dATP and dTTP inserted slightly faster than dCTP and dGTP. Klenow fragment inserts each native triphosphate opposite MICS with only a four-fold variation in efficiency, and can synthesize fulllength DNA with high concentrations of dNTPs. The unnatural nucleoside triphosphate, dPIMTP, behaves as a universal chain terminator. The triphosphate is inserted opposite any natural base in the template with remarkable efficiency (only ∼40-fold reduced relative to insertion of a correct triphosphate) and with only 5-fold discrimination. After incorporation of dPIMTP, Klenow fragment is unable to continue DNA synthesis, even under forcing conditions (0.5-hr incubation time and 1 mM dNTPs). Therefore, PIM inserts efficiently but randomly into the growing oligonucleotide strand and then terminates synthesis. The average length of synthesized oligonucleotide may be tuned by choice of chain terminator concentration. The efficient insertion of dPIMTP opposite any native template results in the efficient generation of random-length oligonucleotides, with low concentrations of a single chain terminator. This contrasts with the conventional dideoxy method of chain termination, in which each dideoxynucleotide must be used in great excess to compete with more efficient incorporation of natural nucleoside triphosphates. For example, Klenow fragment prefers the natural dNTP substrates by several thousand-fold relative to the chain-terminator dideoxy analogs. In this regard, dPIMTP may have practical applications, for example, in the generation of random-length oligonucleotides for sequencing by mass spectrometry. In contrast to nucleobases designed to be universal bases, nucleobases designed as thirdbase-pair candidates should have orthogonal properties relative to the natural bases. They must form stable pairs in duplex DNA, as well as during enzymatic replication, only with their designed base pair partners. Much attention has been focused on unnatural nucleobases with phenyl, naphthyl, isocarbostyril, or 7-azaindole scaffolding. Modifications may be made and then evaluated in terms of base-pair stability (Tm) and replicability (steady-state kinetics). For example, d2MNTP is efficiently inserted by Klenow fragment opposite 2MN (S.21; Fig. 1.5.8) in the template (self-pair formation) with an efficiency of 4.4 × 107 M−1 min−1, but is also efficiently inserted opposite dA (1.0 × 107 M−1 min−1). It was reasoned that this might result from favorable interactions in the developing
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minor groove between the 2MN methyl group and the methine of dA. Therefore, the nucleobase was modified, resulting in 3MN. The mispairing between 3MN and dA was selectively reduced (7.6 × 103 M−1 min−1), resulting in a 3MN:3MN unnatural base pair that is synthesized only 10-fold more slowly than natural base pairs (2.4 × 106 M−1 min−1) and with at least two orders of magnitude selectivity against all possible mispairs. Based on thermodynamic and kinetic characterization, a variety of self-pairs and heteropairs have been identified that are attractive candidates as third base pairs. These include the 7AI:7AI, PICS:PICS, and 3MN:3MN selfpairs, as well as the 7AI:ICS and PP:MICS heteropairs (Figure 1.5.10). It should be noted that the use of a self-pair in no way limits the effort to expand the genetic code. In fact, addition of a self-pair to the genetic alphabet would result in 64 new codons. Each of these unnatural base pairs are stable and replicable with reasonable efficiency and selectivity. The rate-limiting step for the synthesis of DNA with each of the unnatural base pairs is extension, i.e., the continued primer extension after incorporation of an unnatural base. This step may be optimized by design and direct evaluation of steady-state extension rates, or by examining different polymerases. Progress along both lines has been published. For example, it was found that Klenow fragment and polymerase β have complementary recognition of the 7AI:7AI self-pair. Although Klenow fragment efficiently inserts d7AITP opposite 7AI in the template, it is unable to continue primer extension. Polymerase β, while unable to synthesize the base pair, extends it with a
N
N
N
N
N
7AI:7AI
wild-type-like efficiency. The action of both polymerases allows for the synthesis of DNA containing the 7AI:7AI self-pair in addition to dG:dC and dA:dT (Lee et al., 2001). Further derivativization of the unnatural bases is also being evaluated with respect to extension. In particular, the effects of aromatic surface area and a minor groove hydrogen-bond acceptor are being evaluated. It is apparent that hydrophobicity is a suitable force for controlling internucleobase interactions in duplex DNA as well as during enzymatic replication. The approach is expected to be more versatile than one based on purine or pyrimidine analogs because a wider range of nucleobase analogs may be examined. It is perhaps surprising that such a wide range of nucleobase structures are tolerated in the duplex and polymerase environments. Nonetheless, this wide tolerance of structures may be taken advantage of to design nucleobase analogs with desired thermal and kinetic properties for use as a universal base or to expand the genetic code.
Compound Characterization
S.4: 1H NMR (400 MHz, CDCl3) δ 8.39 (1H, dd, J = 8.1, 1.1 Hz), 7.94-7.98 (4H, m), 7.91 (1H, d, J = 7.8 Hz), 7.70 (2H, m), 7.51 (1H, m), 7.20−7.27 (4H, m), 6.86 (1H, dd, J = 8.6, 5.4 Hz), 5.62 (1H, m), 4.79 (1H, dd, J = 12.2, 3.5 Hz), 4.70 (1H, dd, J = 12.1, 3.2 Hz), 4.60 (1H, m), 2.85 (1H, ddd, J = 14.3, 5.5, 1.6 Hz), 2.42 (3H, s), 2.38 (3H, s), 2.32 (1H, ddd, J = 14.6, 8.4, 6.8Hz), 1.94 (3H, s). S.5: 1H NMR (400 MHz, CD3OD) δ 8.28 (1H, m), 7.97 (1H, s), 7.94 (1H, dd, J = 8.1, 0.6 Hz), 7.76 (1H, m), 7.54 (1H, m), 6.63 (1H, dd,
O O PICS:PICS
N 3MN:3MN
N N
N
N O
Development of a Universal Nucleobase and Unnatural Nucleobases
7AI:ICS
N
N
O
N
PP:MICS
Figure 1.5.10 Unnatural base pairs for expansion of the genetic alphabet.
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J = 7.3, 6.2 Hz), 4.4 (1H, m), 3.98 (1H, m), 3.83 (1H, dd, J = 11.9, 3.5 Hz), 3.76 (1H, dd, J = 12.0, 3.8 Hz), 2.41 (1H, ddd, J = 13.7, 6.2, 3.5 Hz), 2.17 (1H, ddd, J = 13.6, 7.4, 6.5 Hz), 2.10 (3H, s). 13C NMR (125 MHz, CD3OD) δ 160.0, 135.2, 131.7, 129.0, 126.0, 125.9, 123.5, 101.5, 87.6, 86.5, 84.1, 71.9, 69.8, 60.3, 39.7. HRMS (MALDI) anal. calcd. for C17H17NO4Na (MNa+) 322.1055; found 322.1045. S.6: 1H NMR (400 M Hz, CDCl3) δ 8.40 (1H, d, J = 8.1 Hz), 7.93 (2H, m), 7.70 (1H, m), 7.50 (3H, m), 7.37-7.41 (4H, m), 7.18-7.30 (4H, m), 6.82 (4H, m), 6.71 (1H, ddd, J = 12.8, 7.6, 5.7 Hz), 4.56 (1H, m), 4.24 (1H, m), 3.743.83 (8H, m), 3.52-3.65 (4H, m), 3.45 (1H, m), 3.33 (1H, m), 2.67 (1H, m), 2.61 (2H, t, J = 6.5 Hz), 2.42 (2H, t, J = 6.5 Hz), 2.26 (1H, m), 1.82 (3H, d, J = 11.9 Hz), 1.17 (9H, m), 1.06 (3H, d, J = 6.8 Hz). 31P NMR (160 MHz, CDCl3) δ 51.4, 50.7. S.7: 31P NMR (160 M Hz, 50 mM Tris (pH 7.5), 2mM EDTA in D2O) δ -6.11 (d, J = 19.2 Hz), -10.38 (d, J = 19.9 Hz), 21.55 (dd, J = 19.9, 19.2 Hz). S.8: 1H NMR (400 M Hz, CDCl3) δ 8.38 (1H, m), 8.08 (2H, d, J = 6.4 Hz), 8.02 (2H, d, J = 6.5 Hz), 7.78 (1H, d, J = 2.1 Hz), 7.36 (2H, d, J = 6.4 Hz), 7.34 (1H, s), 7.33 (2H, d, J = 6.7 Hz), 6.91 (1H, dd, J = 6.9, 4.4 Hz), 6.83 (1H, m), 5.87 (1H, m), 4.80 (1H, dd, J = 9.7, 3.2 Hz), 4.73 (1H, dd, J = 9.4, 3.2 Hz), 4.69 (1H, m), 3.08 (1H, ddd, J = 11.6, 6.6, 5.0 Hz), 2.83 (1H, ddd, J = 11.4, 4.5, 1.8 Hz), 2.52 (3H, s), 2.50 (3H, s). Deprotected S.8: 1H NMR (400 MHz, CD3OD) δ 8.37 (1H, d, J = 2.7 Hz), 8.24 (1H, d, J = 2.7 Hz), 8.05 (1H, d, J = 3.8 Hz), 6.68 (1H, dd, J = 8.1, 6.0 Hz), 6.67 (1H, d, J = 3.8 Hz), 4.53 (1H, m), 3.99 (1H, dd, J = 6.6, 4.1 Hz), 3.77 (1H, dd, J = 12.0, 3.5 Hz), 3.70 (1H, dd, J = 12.0, 4.1 Hz), 2.72 (1H, ddd, J = 13.8, 7.7, 6.0 Hz), 2.35 (1H, ddd, J = 13.5, 6.2, 3.0 Hz); HMRS anal. calcd. for C11H14N3O3 (MH+) 236.1035; found 236.1034. S.11: 1H NMR (400 MHz, CD3OD) δ 7.99 (4H, m), 7.28 (2H, d, J = 8.1 Hz), 7.24 (2H, d, J = 8.1 Hz), 7.02 (2H, s), 6.92 (1H, s), 5.62 (1H, m), 5.20 (1H, dd, J = 10.9, 5.0 Hz), 4.71 (1H, dd, J = 11.8, 4.0 Hz), 4.66 (1H, dd, J = 11.8, 3.6 Hz), 4.53 (1H, m), 2.50 (1H, m), 2.44 (3H, s), 2.41 (3H, s), 2.66 (7H, m). HRMS anal. calcd. for C29H31O5 (MH+) 459.2171; found 459.2179. S.12 31P NMR (140 MHz, CDCl3) δ 148.5, 148.3. S.18: 1H NMR (400 MHz, CDCl3) δ 8.07 (1H, m), 7.98 (1H, m), 7.50 (2H, m), 7.45 (1H,
s), 5.24 (1H, dd, J = 11.5, 1.7 Hz), 4.23 (1H, m), 3.96 (2H, m), 3.81 (1H, m), 2.67 (3H, s), 2.62 (3H, s), 2.10 (1H, m), 1.88 (1H, ddd, J = 14.2, 11.6, 2.3 Hz). 13C NMR (150 MHz, CDCl3) δ 135.5, 132.8, 132.7, 132.1, 127.6, 125.7, 125.3, 124.7, 124.5, 123.8, 76.0, 70.9, 69.2, 68.0, 36.5, 19.5, 13.8. HRMS anal. calcd. for C13H24NO3 (MNH4+) 290.1756; found 290.1764. S.21: 1H NMR (600 MHz, CD3OD) δ 8.03 (1H, s), 7.78 (1H, d, J = 7.1 Hz), 7.71 (1H, d, J = 7.1 Hz), 7.59 (1H, s), 7.37 (2H, m), 5.44 (1H, dd, J = 10.1, 5.3 Hz), 4.35 (1H, m), 3.98 (1H, m), 3.76 (2H, m), 2.47 (3H, s), 2.36 (1H, ddd, J = 13.2, 5.7, 1.8 Hz), 1.90 (1H, m). 13C NMR (150 MHz, CDCl3) δ 139.8, 133.8, 133.6, 133.1, 128.6, 128.2, 127.3, 126.1, 125.6, 124.1,88.2, 78.2, 73.8, 63.5, 42.8, 19.1. HRMS anal. calcd. for C16H18O3 (MNa+) 281.1154; found 281.1158.
Critical Parameters and Troubleshooting Generally, any reaction that involves moisture-sensitive reagents should be handled with care. The reaction equipment, including flasks, rubber septa, adapters, gas or liquid condensers, and syringes, must be predried in an oven or under an infrared lamp and flushed with dry argon before use. Anhydrous solvents are critical to the success of reactions. It is best to freshly distill solvents and use moisture-sensitive reagents in situ. Glycosylation. The Lewis acids used in Silyl-Hilbert-Johnson reactions such as Tin(IV) tetrachloride are generally moisturesensitive. The quality of these reagents dramatically affects the yield of product. Iodination. The reaction time is very critical. Excess time may cause significant side reactions. Propynation. The reaction system should be absolutely air-free, since copper(I) iodide and the palladium catalyst used are generally airand/or light-sensitive. The pressure tube should be sealed very well, otherwise propyne collected at low temperature will evaporate readily when the reaction is stirred at ambient temperature. Deprotection. There are no particular difficulties with this step. Tritylation. The substrate should be thoroughly dried by coevaporation with anhydrous pyridine prior to use. Slow addition of the solution of DMTr-Cl in pyridine or batchwise addition of DMTr-Cl solid is necessary; other-
Synthesis of Modified Nucleosides
1.5.33 Current Protocols in Nucleic Acid Chemistry
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wise, the yield of product will be greatly reduced. Phosphitylation. 2-Cyanoethyl diisopropychlorophosphoramidite should be strictly stored at −20°C and warmed to ambient temperature before use. Quickly transferring this reagent via a short needle to the reaction mixture produces optimal results. Phosphorylation. The moisture contained in the substrate should be removed by azeotropic evaporation from a small amount of anhydrous pyridine before the other reactants are added. SN2 displacement reactions. Sodium hydride should be dried and used in situ and quickly added into the reaction flask in several batches, because dry sodium hydride decomposes very readily during storage. Evaporation of solvents. A rotatory evaporator is routinely used to remove solvents. The efficiency of evaporation is largely dependent on the vacuum (normally between 10 to 20 mm Hg) and the dry-ice condenser. It is better to maintain the bath temperature below 45°C to prevent any chemical change of the crude product. To avoid loss of compound caused by a sudden burst of solvents into the splash-guard adapter, the evaporation flask should be cooled by slow rotary evaporation under low vacuum before it is immersed into the bath. Grignard reaction. This reaction should be carried out with exclusion of moisture. All apparatus must be thoroughly dried in a hot (>120°C) oven before use. Use anhydrous solvents. Deprotection of toluoyl group. There are no particular difficulties with this reaction; it will work effectively. Nucleophilic addition by aryllithium reagents. The reaction is moisture-sensitive. Anhydrous solvent is important for the reaction. Do not use n-butyllithium reagent with a milky precipitate, because such a reagent is decomposed in the presence of water. Removal of protecting group of hydroxy group. Difficulties are seldom encountered in this step.
Anticipated Results
Development of a Universal Nucleobase and Unnatural Nucleobases
An overall good yield of the free nucleoside (48% to 68%) of unnatural hydrophobic nucleobases can be achieved based on the strategies described in this unit. The yields of the nucleosidic glycosylation are generally good eno ug h f or th e pr ep ar atio n of phosphoramidites and triphosphates, although stereoselectivity of the desired β-anomer versus α-anomer may be mediocre in some cases.
The routine deprotection, phosphitylation, and phosphorylation are generally high-yielding. Although the Grignard addition described in Basic Protocol 3 generally favors the αanomer, acid treatment of the corresponding benzylic ethers in refluxing xylene will result in epimerization at C1′ to yield the β-anomer (Ren et al., 1996). Conversely, the aryllithium addition described in Basic Protocol 4 and the Alternate Protocol generally affords the desired product in a good yield. The product is a mixture of diastereomers, but is separable by column chromatography.
Time Considerations The estimated time to complete Basic Protocol 1 in its entirely is 3 weeks. The estimated time required to complete Basic Protocol 2 is 2 days. Basic Protocol 3 should take ∼6 days to complete. For Basic Protocol 4 and Alternate Protocol, allow ∼1 day for the Grignard reaction and several hours for all of the other reactions. For Basic Protocol 5, 2 to 3 days should be sufficient time to prepare the oligonucleotide. On day 1, the oligonucleotide is synthesized and the deprotection should be done overnight. Several hours should be allowed for gel pouring and polymerization; this step may also be done on day 1 or in the morning of day 2. The gel must be kept hydrated, regardless, by wrapping with plastic wrap. The electrophoresis step can take several hours. After separation, the excised gel slice may be stored at −20°C for further purification at a later date. To perform the electroelution, allow <2 hr. Ethanol precipitation can then be done overnight at −20°C and the final steps done the following morning. For Basic Protocol 7, about 1 hr should be allowed for radiolabeling the primer. Allow 2 hr for annealing the primer-template duplex, during which time the gel should be poured and allowed to polymerize. Annealed primer-templates can be stored at 4°C overnight, and the gel can also be stored overnight at room temperature (it must be protected from dehydration by first washing the wells with buffer, then covering with paper towels and wrapping tightly in plastic wrap). Allow the annealed primer-templates to return to room temperature before use. The time it takes to aliquot the triphosphates and run the reactions depends on the number and nature of the reactions. After final loading of the gel, it can be run at 90 W for about 1.5 hr. Excising the portion of the gel containing the radioactive bands and subsequent drying onto filter paper should take no
1.5.34 Supplement 10
Current Protocols in Nucleic Acid Chemistry
more than 1 hr. Finally exposure of the phosphor storage screen is routinely done overnight.
Literature Cited Beaussire, J.J. and Pochet, S. 1999. Recognition of 2′-deoxyisoinosine triphosphate by the Klenow fragment of DNA polymerase I. Nucleosides Nucleotides 18:403-410. Chaudhuri, N.C., Ren, R.X.F., and Kool, E.T. 1997. C-Nucleoside derived from simple aromatic hydrocarbons. Synlett. 341-347. Creighton, S., Bloom, L.B., and Goodman, M.F. 1995. Gel fidelity assay measuring nucleotide misinsertion, exonucleolytic proofreading, and lesion bypass efficiencies. Methods Enzymol. 262:232-256. Eaton, M.A.W. and Millican, T.A. 1988. New methodology for C-nucleoside synthesis: Preparation of 1,2-dideoxy-1-(3-pyridyl)-D-ribofuranose. J. Chem. Soc. Perkin Trans. I. 545-547. Fischer, E. and Helferich, B. 1914. Synthetische Glucoside der Purine. Chem. Ber. 47:210. Hilbert, G.E. and Johnson, T.B. 1930. Researches on pyrimidines. CXV. Alkylation on nitrogen of the pyrimidine cycle by application of a new technique involving molecular rearrangements. J. Am. Chem. Soc. 52:2001. Hildbrand, S., Blaser, A., Parel, S.P., and Leumann, C.J. 1997. 5-Substituted 2-aminopyridine C-nucleosides as protonated cytidine equivalents: Increasing efficiency and selectivity in DNA triplehelix formation. J. Am. Chem. Soc. 119:54995511. Horlacher, J., Hottiger, M., Podust, V.N., Hübscher, U., and Bennier, S.A. 1995. Recognition by viral and cellular DNA polymerases of nucleosides bearing bases with nonstandard hydrogen bonding patterns. Proc. Natl. Acad. Sci. U.S.A. 92:6329-6333. Kazimierczuk, Z., Cottam, H.B., Revankar, G.R., and Robins, R.K. 1984. Synthesis of 2′-deoxytubercidin, 2′-deoxyadenosine, and related 2′-deoxynucleosides via a novel direct stereospecific sodium salt glycosylation procedure. J. Am. Chem. Soc. 106:6379-6382. Kornberg, A. and Baker, T.A. 1992. DNA Replication, 2nd ed. W.H. Freeman and Co., New York. Lambert, J.B., Fabricus, D.M., and Hoard, J.A. 1979. Bond localization approach to the carbon analog of the Claisen rearrangement. Thermolysis of 4-aryl-1-butenes. J. Org. Chem. 44:1480-1485. Lambert, J.B., Shurvell, H.F., Lightner, D.A., and Cooks, R.G. 1998. Organic Structural Spectroscopy. Prentice Hall, Englewood Cliffs, N.J. Lee, E.L., Wu, Y., Xia, G., Schultz, P.G., and Romesberg, F.E. 2001. Efforts toward expansion of the genetic alphabet: Replication of DNA with three base pairs. J. Am. Chem. Soc. 123:7439-7440.
Lutz, M.J., Held, H.A., Hottiger, M., Hübscher, U., and Benner, S.A. 1996. Differential discrimination of DNA polymerases for variants of the non-standard nucleobase pair between xanthosine and 2,4-diaminopyrimidine, two components of an expanded genetic alphabet. Nucl. Acids Res. 24:1308-1313. Lutz, M.J., Horlacher, J., and Benner, S.A. 1998a. Recognition of 2′-deoxyisoguanosine triphosphate by HIV-reverse transcriptase and mammalian cellular DNA polymerases. Bioorg. Med. Chem. Letts. 8:499-504. Lutz, M.J., Horlacher, J., and Benner, S.A. 1998b. Recognition of a non-standard base pair by thermostable DNA polymerases. Bioorg. Med. Chem. Letts. 8:1149-1152. McMinn, D.L., Ogawa, A.K., Wu, Y., Liu, J., Schultz, P.G., and Romesberg, F.E. 1999. Efforts toward expansion of the genetic alphabet: DNA polymerase recognition of a highly stable, selfpairing hydrophobic base. J. Am. Chem. Soc. 121:11585-11586. Niedballa, U. and Vorbrüggen, H. 1970. A general synthesis of pyrimidine nucleosides. Angew. Chem. Int. Ed. Engl. 9:461. Niedballa, U. and Vorbrüggen, H. 1974. A general synthesis of N-glycosides. I. Synthesis of pyrimidine nucleosides. J. Org. Chem. 39:36543660. Ogawa, A.K., Wu, Y., McMinn, D.L., Liu, J., Schultz, P.G., and Romesberg, F.E. 2000a. Efforts toward the expansion of the genetic alphabet: Information storage and replication with unnatural hydrophobic base pairs. J. Am. Chem. Soc. 122:3274-3287. Ogawa, A.K., Wu, Y., Berger, M., Schultz, P.G., and Romesberg, F.E. 2000b. Rational design of an unnatural base pair with increased kinetic selectivity. J. Am. Chem. Soc. 122:8803-8804. Postema, M.H.D. 1992. Recent developments in the synthesis of C-glycosides. Tetrahedron 48:85458599. Ren, R.X.F., Chaudhuri, N.C., Paris, P.L., Rumney S., and Kool, E.T. 1996. Naphthalene, phenanthrene, and pyrene as DNA base analogues: Synthesis, structure, and fluorescence in DNA. J. Am. Chem. Soc. 118:7671-7678. Roberts, C., Chaput, J.C., and Switzer, C. 1997a. Beyond guanine quartets: Cation-induced formation of homogenous and chimeric DNA tetraplexes incorporating iso-guanine and guanine. Chem. & Biol. 4:899-908. Roberts, C., Bandaru, R., and Switzer, C. 1997b. Theoretical and experimental study of isoguanine and isocytosine: Base pairing in an expanded genetic system. J. Am. Chem. Soc. 119:46404649. Robinson, H., Gao, Y.-G., Bauer, C., Roberts, C., Switzer, C., and Wang, A.H. 1998. 2′-Deoxyisoguanosine adopts more than one tautomer to form base pairs with thymidine observed by high-resolution crystal structure analysis. Biochemistry 37:10897-10905.
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1.5.35 Current Protocols in Nucleic Acid Chemistry
Supplement 10
Sharma, P.K. 1993. A new synthesis of trans-3,4-dihydroxy-anti-1,2-epoxy-1,2,3,4-tetrahydro-7, 12-dimethylbenz[a]anthracene. Synth. Commun. 23:389-394. Solomon, M.S. and Hopkins, P.B. 1993. Chemical synthesis and characterization of duplex DNA containing a new base pair: A nondisruptive, benzofused pyrimidine analog. J. Org. Chem. 58: 2232-2243. Takeshita, M., Chang, C.N., Johnson, F., Will, S., and Grollman, A.P. 1987. Oligodeoxynucleotides containing synthetic abasic sites. J. Biol. Chem. 262:10171-10179. Vorbrüggen, H. and Ruh-Pohlenz, C. 2001. Handbook of Nucleoside Synthesis. John Wiley & Sons, New York. Vorbrüggen H., Krolikiewicz, K., and Bennua, B. 1981. Nucleoside synthesis with trimethylsilyl triflate and perchlorate as catalysts. Chem. Ber. 114:1234-1235. Wu, Y., Ogawa, A.K., Berger, M., McMinn, D.L., Schultz, P.G., and Romesberg, F.E. 2000. Efforts toward expansion of the genetic alphabet: Optimization of interbase hydrophobic interactions. J. Am. Chem. Soc. 122:7621-7632.
Key References Berger, M., Wu, Y., Ogawa, A.K., McMinn, D.L., Schultz, P.G., and Romesberg, F.E. 2000. Universal bases for hybridization, replication and chain termination. Nucl. Acids Res. 28:29112914.
This paper describes the evaluation of a variety of unnatural bases as universal bases for hybridization and replication. Berger, M., Luzzi, S.D., Henry, A.A., and Romesberg, F.E. 2002. Stability and selectivity of unnatural DNA with five-membered-ring nucleobase analogues. J. Am. Chem. Soc. 124:12221226. This paper describes the evaluation of 5-membered ring nucleobase analogs. Lee et al., 2001. See above. This paper describes the replication of DNA containing the 7AI self-pair with the Klewnow fragment/pol β binary polymerase system. Ogawa et al., 2000a. See above. This paper describes the synthesis of the C-nucleosides reported herein. Ogawa et al., 2000b. See above. This paper describes the rational design of the 3MN:3MN self-pair.
Contributed by Floyd E. Romesberg, Chengzhi Yu, Shigeo Matsuda, and Allison A. Henry The Scripps Research Institute La Jolla, California
Development of a Universal Nucleobase and Unnatural Nucleobases
1.5.36 Supplement 10
Current Protocols in Nucleic Acid Chemistry
Syntheses of Specifically 15N-Labeled Adenosine and Guanosine
UNIT 1.6
This unit describes the specific incorporation of 15N into the N7 and amino positions of adenosine (Basic Protocol 1), and conversion of the adenosine to guanosine labeled at the N1, N7, and amino positions (Basic Protocol 2). Two variations of the procedures are also presented that include either 12C or 13C at the C8 position of adenosine, and 13C at either the C8 or C2 position of guanosine. These 13C tags permit the incorporation of two 15 N-labeled nucleosides into an RNA strand while ensuring that their nuclear magnetic resonance (NMR) signals can be distinguished from each other by the presence or absence of C-N coupling. While the major application of these specifically 15N-labeled nucleosides is NMR, the additional mass makes them useful in mass spectrometry (MS) as well. The procedures can also be adapted to synthesize the labeled deoxynucleosides. The Support Protocol describes the synthesis of 7-methylguanosine. CAUTION: The procedures in this unit use a number of highly toxic and dangerous reagents. Raney nickel is pyrophoric when dry and may burst into flames if not kept wet. Phosphorous oxychloride (POCl3) is very reactive and hydrolyzes to hydrochloric and phosphoric acids, which are both highly corrosive to skin and tissue. Cyanogen bromide is very dangerous; improper use of this reagent has led to deaths in the laboratory. All reactions must be done with great care in an appropriate chemical fume hood. SYNTHESES OF [7,NH2-15N2]- AND [8-13C-7,NH2-15N2]ADENOSINE As shown in Figure 1.6.1, the procedures described here (Pagano et al., 1995; Zhao, 1997; Shallop and Jones, 2000) start with the inexpensive pyrimidine 4-amino-6-hydroxy-2mercaptopyrimidine and introduce the first 15N label by a direct nitrosation in high yield. Reduction of the nitroso group to an amino group is followed by ring closure using either diethoxymethylacetate in dimethylformamide (DMF) to give 12C at the C8 position, or [13C]sodium ethyl xanthate to give 13C at the C8 position. Removing the thiol group(s) with Raney nickel forms hypoxanthine, which can readily be converted to 6-chloropurine, a good substrate for enzymatic transglycosylation. The second 15N label is then introduced into the nucleoside by displacement of the chloride by 15NH3, which is generated in situ. Materials 4-Amino-6-hydroxy-2-mercaptopyrimidine monohydrate, also called 6-amino-2-thioxo-1,2-dihydro-4(3H)-pyrimidinone (Aldrich) 1 N HCl [15N]Sodium nitrite ([15N] NaNO2; Isotec or Cambridge Isotope Laboratories) 2:98 to 40:60 (v/v) gradient of acetonitrile/0.1 M triethylammonium acetate (TEAA), pH 6.8 95% (v/v) ethanol, 4°C Acetone, 4°C Phosphorous pentoxide (P2O5) Saturated aqueous NaHCO3 Sodium hydrosulfite (Na2S2O4) Glacial acetic acid 96% (v/v) formic acid Nitrogen gas source Dimethylformamide (DMF), anhydrous Diethoxymethyl acetate (DEMA), for 12C synthesis only Acetonitrile, room temperature and 4°C Contributed by Barbara L. Gaffney and Roger A. Jones Current Protocols in Nucleic Acid Chemistry (2002) 1.6.1-1.6.14 Copyright © 2002 by John Wiley & Sons, Inc.
BASIC PROTOCOL 1
Synthesis of Modified Nucleosides
1.6.1 Supplement 10
O
O [ 15N ] N a N O 2
NH H2N
N
O 15N H 2N
SH
1
O
N
H215N
Na 2S 2O 4
NH
NH
H2N
SH
N
2
SH
3 DEMA or [ 13C]NaSCSOEt
Cl
O
15
O
15
N
N
X N H
15
N
POCl 3
6a, X =12C 6b, X =13C
NH
X N H
N
N
RaNi
NH
X N H
N
5a, X =12C 5b, X =13C
N
SH
4a, X =12CH 4b, X =13CSH
7-MeG PNP 15
Cl 15
N
N
X N
HO
O HO
OH
7a, X =12C 7b, X =13C
N
NH2
15
N
[ 15N ] N H 4C l KHCO 3
N
X N
HO
O HO
N
OH
8a, X =12C 8b, X =13C
Figure 1.6.1 Steps for synthesis of [7,NH2-15N2]- and [8-13C-7,NH2-15N2]adenosine using Basic Protocol 1. DEMA, diethoxymethyl acetate; 7-MeG, 7-methylguanosine; [13C]NaSCSOEt, [13C]sodium ethyl xanthate; PNP, purine nucleoside phosphorylase; RaNi, Raney nickel.
[13C]NaSCSOEt (see recipe), for 13C synthesis only NaOH 50% aqueous Raney 2800 nickel (RaNi) slurry (Aldrich) Dipotassium salt of EDTA Boiling water Phosphorous oxychloride (POCl3) N,N-Dimethylaniline 5% (v/v) NH3, (diluted with water from 30% concentrated aqueous ammonia) Ethyl acetate Ethyl ether 1 M HCl 7-Methylguanosine (see Support Protocol) 0.02 M K2HPO4 3 M NaOH Purine nucleoside phosphorylase (Sigma) [15N]Ammonium chloride ([15N]NH4Cl; Isotec or Cambridge Isotope Laboratories) Dimethylsulfoxide (DMSO), anhydrous KHCO3, anhydrous Synthesis of Specifically N-Labeled Adenosine and Guanosine 15
1.6.2 Supplement 10
Current Protocols in Nucleic Acid Chemistry
100- and 250-mL round-bottom flasks Small glass vials 1-, 3-, 10-, and 20-ml syringes Vacuum desiccator Rubber septa fitted with large-bore vent needles Condenser Rotary evaporator, connected to water aspirator and a vacuum pump, the latter with a dry-ice trap Oil bath (silicone oil), 130°C Separatory funnel Continuous extraction apparatus for solvents lighter than water (Aldrich) 30°C oven with shaker 50-mL bomb with Teflon liner (Parr Instrument) 80°C oven Additional reagents and equipment for analytical and preparative reversed-phase high-performance liquid chromatography (HPLC; UNIT 10.5) Synthesize [5-15N]-6-amino-5-nitroso-2-thioxo-1,2-dihydro-4(3H)-pyrimidinone (S.2) 1. Weigh 0.805 g (5.0 mmol) of 4-amino-6-hydroxy-2-mercaptopyrimidine monohydrate into a 100-mL round-bottom flask with a stir bar. 2. Add 25 mL of 1 N HCl and chill the suspension 10 min in an ice bath. 3. Weigh 0.385 g (5.5 mmol, 1.1 eq) [15N]NaNO2 into a small glass vial, dissolve it in ∼1 mL water, and draw the solution into a 3-mL syringe. 4. Slowly add the sodium nitrite to the reaction mixture over ∼5 min. If the addition is too fast, the reaction may bubble over. Before long, the mixture will change from yellow to red.
5. Stir the mixture for ∼7 hr in the ice bath, while monitoring the reaction for completeness by reversed-phase HPLC with a gradient of 2:98 to 40:60 acetonitrile/0.1 M TEAA over 5 min. The peak representing the starting material should diminish to <3%, and a new peak representing the product should appear. Any reversed-phase column can be used. The sample should be injected immediately after mixing for comparison.
6. Collect S.2 by vacuum filtration and wash it with 5 mL cold water, then 5 mL cold 95% ethanol, and finally 5 mL cold acetone. 7. Without removing it from the funnel, dry S.2 over P2O5 in a vacuum desiccator overnight. The yield is usually >95%. See Table 1.6.1 for various data on this and other intermediates and products.
Synthesize [5-15N]-5,6-diamino-2-thioxo-1,2-dihydro-4(3H)-pyrimidinone (S.3) 8. Scrape out most of S.2 from the funnel into a 100-mL round-bottom flask. Rinse the funnel with portions of saturated aqueous NaHCO3 and add them to the flask to give a final volume of 40 mL. 9. Add a stir bar and place the mixture in an ice bath for 10 min. Stir gently.
Synthesis of Modified Nucleosides
1.6.3 Current Protocols in Nucleic Acid Chemistry
Supplement 10
10. Quickly weigh 2.61 g (15 mmol, 3 eq) Na2S2O4 into a small beaker. Using a spatula, gradually add it in portions to the reaction over 20 min. The sodium hydrosulfite smells bad, so it should be kept in the back of the hood. If the addition is too fast, the reaction will bubble over.
11. Insert a rubber septum containing a large-bore vent needle. 12. Stir the mixture for ∼7 hr in the ice bath, while monitoring the reaction for completeness by HPLC as described (step 5). The mixture will change from red to yellow.
13. Slowly add 1.6 mL glacial acetic acid over ∼5 min to neutralize the NaHCO3, and stir another 5 min. 14. Collect the product by vacuum filtration and wash it twice with 5 mL cold water and then twice with 5 mL cold 95% ethanol. 15. Without removing it from the funnel, dry S.3 over P2O5 in a vacuum desiccator overnight. The yield is usually >95%. The synthesis is continued using either diethoxymethylacetate in dimethylformamide (DMF) to give 12C at the 8 position (steps 16a to 26a) or [13C]sodium ethyl xanthate to give 13C at the 8 position (steps 16b to 22b).
Perform ring closure For [7-15N]-2-thioxohypoxanthine (S.4a): 16a. Scrape out most of S.3 from the funnel into a 100-mL round-bottom flask. Rinse the funnel with portions of 96% formic acid and add them to the flask to give a final volume of 25 mL.
Table 1.6.1 Molecular Weights, TLC and HPLC Mobilities, and UV λmax of 15N-Labeled Adenosine and Guanosine Intermediatesa
Compound
mol. wt. (Da)
TLC Rfb
HPLC retention time (min)c
UV λmax (nm)
S.2 S.3 S.4a S.4b S.5a/b S.6a/b S.7a/b S.8a/b S.9a/b S.12a/b S.13a/b
173 159 169 202 137/138 155/156 287/288 269/270 285/286 316 287
0.2 0.0 0.2 0.0 0.1 0.3 0.2 0.1 0.0 0.1 0.0
1.2 0.8 0.7 1.4 0.8 2.4 4.5 2.6 0.9 3.3 1.4
360 301 280 299 250 265 264 259 295 280 253
aAbbreviations: HPLC, high-performance liquid chromatography; mol. wt., molecular weight; TLC, thin-layer chroma-
tography.
Synthesis of Specifically 15 N-Labeled Adenosine and Guanosine
bR values determined with 10:90 (v/v) CH OH/CH Cl . f 3 2 2 cGradient of 2:98 to 40:60 (v/v) acetonitrile/0.1 M triethylammonium acetate, pH 6.8, over 5 min, on a Waters NovaPak
C18 column.
1.6.4 Supplement 10
Current Protocols in Nucleic Acid Chemistry
17a. Add a stir bar, attach a condenser, and reflux the solution for 1 hr to make the formate salt. 18a. Concentrate to dryness using a rotary evaporator and scrape down the sides of the flask with a spatula, if necessary. 19a. Insert a rubber septum and displace the air with nitrogen. 20a. Use syringes to add the following through the septum: 20 mL anhydrous DMF 1.63 mL DEMA (10 mmol, 2 eq) 0.24 mL of 96% formic acid (6 mmol, 1.2 eq). 21a. Heat the mixture for 3 hr in an oil bath set at 130°C. Follow the reaction by HPLC. The flask is lifted from the oil bath and allowed to cool briefly, and then a small syringe with a long, dry needle is used to get a sample for HPLC.
22a. Cool the flask, concentrate the solution to a solid using a rotary evaporator, and loosen it with a spatula if necessary. 23a. Add 15 mL acetonitrile to the flask, attach a condenser, and reflux it for 10 min using the 130°C oil bath. 24a. Cool the flask to room temperature, add 10 mL acetonitrile, and then chill it in an ice bath. 25a. Collect S.4a by vacuum filtration and wash it twice with 5 mL cold acetonitrile. 26a. Without removing it from the funnel, dry S.4a over P2O5 in the vacuum desiccator overnight. Proceed to step 27. The yield is usually >95%.
For [8-13C-7-15N]-2,8-dithioxohypoxanthine (S.4b): 16b. Scrape out most of S.3 from the funnel into a 100-mL round-bottom flask and add 0.80 g (5.5 mmol) [13C]NaSCSOEt. 17b. Insert a condenser into the flask, attach a nitrogen line and vent needle, and displace the air for 5 min. 18b. Add 15 mL DMF and reflux the mixture under nitrogen for ∼3 hr, using HPLC to monitor the reaction. 19b. Cool the mixture in an ice bath and add 50 mL cold acetonitrile to precipitate S.4b. 20b. Collect the solid S.4b by vacuum filtration and wash it twice with 5 mL cold acetonitrile. Save the filtrate and both washes. 21b. Concentrate the filtrate and washes, and purify this portion of S.4b by preparative reversed-phase chromatography. 22b. Dry the combined portions of S.4b over P2O5 in a vacuum desiccator overnight. Continue with step 27. The yield is usually >95%.
Synthesis of Modified Nucleosides
1.6.5 Current Protocols in Nucleic Acid Chemistry
Supplement 10
Synthesize [7-15N]hypoxanthine and [8-13C-7-15N]hypoxanthine (S.5a/b) 27. Scrape out most of S.4a/b from the funnel into a 100-mL round-bottom flask. 28. Rinse the funnel with portions of water and add them to the flask to give a final volume of 30 mL. 29. Add a stir bar and 2 mL of 96% formic acid. 30. Weigh 4.5 g of 50% aqueous RaNi slurry into a small glass vial or beaker. CAUTION: RaNi is pyrophoric (will spontaneously burst into flames) if it is allowed to dry out. To weigh it, shake the bottle and immediately transfer some of the suspended RaNi to the vial. Continue adding the suspension (shaking the bottle each time) until 4.5 g of the 50:50 mixture of RaNi/water has been measured out.
31. Using a dropper, transfer the RaNi suspension over 5 min to the reaction flask, and add 1.5 g dipotassium salt of EDTA. Traces of remaining RaNi in the vial and dropper should be destroyed with 6 M HCl.
32. Connect a condenser to the flask. Using the 130°C oil bath, reflux for ∼2 hr while monitoring the reaction for completeness by HPLC. 33. Remove the flask from the oil bath and allow it to cool only briefly. 34. Remove the condenser from the flask and carefully filter the hot reaction mixture to remove the RaNi. Rinse the flask and then the funnel with three 10-mL portions of boiling water, adding these washes to the filtrate. To destroy the RaNi remaining in the funnel, the funnel should be transferred to a large beaker, and portions of 6 M HCl should be slowly added until no black particles can be observed.
35. Concentrate the filtrate and washes to dryness in a 100-mL round-bottom flask using the rotary evaporator. 36. Dry S.5a/b over P2O5 in the vacuum desiccator overnight. The yield is usually >95%. S.5a/b can be purified by reversed-phase chromatography if desired.
Synthesize [7-15N]- and [8-13C-7-15N]-6-chloropurine (S.6a/b) 37. Weigh 0.69 g (5.0 mmol) S.5a/b into a very dry 100-mL round-bottom flask. 38. Add 20 mL (215 mmol, 43 eq) POCl3 and 2 mL (16 mmol) N,N-dimethylaniline. Use great care with POCl3; it is very reactive.
39. Attach a condenser and reflux 20 min under nitrogen. The resulting solution should be black and homogeneous. It is essential for this reaction to remain anhydrous.
40. Monitor the reaction for completeness by HPLC and continue refluxing the mixture for ≤30 min more. 41. Concentrate the mixture to a very small volume using a rotary evaporator, first with an aspirator and then with a vacuum pump protected with a dry-ice trap. Add 10 mL N,N-dimethylaniline and continue the evaporation. Synthesis of Specifically N-Labeled Adenosine and Guanosine 15
It is necessary to remove all traces of POCl3, or localized heating from later neutralization with NH3 may cause the reaction to reverse. Be extremely careful to dry the evaporator condenser and trap prior to use, to keep dry ice in the traps throughout this process, and then to pour the collected POCl3 into a plastic container of ice to destroy it.
1.6.6 Supplement 10
Current Protocols in Nucleic Acid Chemistry
42. Cool the flask in an ice bath and very slowly add 30 mL of 5% NH3 to dissolve the black gum. As a neutral compound, 6-chloropurine is insoluble in water, but under basic conditions it will ionize and therefore dissolve.
43. Make sure the pH of the solution is >10 (add more NH3 if necessary) and then pour it into a separatory funnel. Wash it first with 30 mL ethyl acetate and then twice with 30 mL ethyl ether. Check the layers by HPLC. 44. Combine all organic layers that contain traces of product in a separatory funnel, backwash with 5% NH3, and add this aqueous layer to the main reaction mixture. 45. Concentrate this aqueous solution to dryness to remove all the NH3. It is necessary to remove all traces of NH3, or localized heating from later neutralization with HCl may cause the reaction to reverse. As the NH3 evaporates, it is very likely to bump, so a large enough flask should be used.
46. Add 20 mL water and chill the flask in an ice bath. Slowly acidify the solution to pH 2 using 1 M HCl. The mixture will turn cloudy.
47. Set up a continuous extraction apparatus for solvents lighter than water. Pour the aqueous layer into the extractor and add ethyl ether until the level is just under the side arm. 48. Fill a 250-mL round-bottom flask with ethyl ether, add a stir bar, connect to the extractor, and place it in an oil bath. Attach a condenser to the top of the extractor, and start heating the oil bath to 45°C. 49. Continue the extraction for 3 to 4 days, using fresh ether each day. Check both the aqueous and ether layers each day by HPLC. Verify that the pH of the aqueous layer is still <2 and, if it is not, adjust it with 1 M HCl. 50. Concentrate the ether layers to a small volume, whereupon a significant amount of S.6a/b should crystallize out. Collect S.6a/b by vacuum filtration and check it for purity by HPLC. Save the filtrate. 51. Concentrate the filtrate to dryness and dissolve it in 10 mL water. Purify by reversed-phase preparative chromatography. 52. Concentrate the fractions containing pure product to dryness. Dry both portions of S.6a/b in a vacuum desiccator over P2O5 overnight. The yield is usually between 80% and 90%.
Synthesize [7-15N]- and [8-13C-7-15N]-6-chloro-9-(β-D-erythropentofuranosyl) purine (S.7a/b) 53. Place 2.12 g (7.5 mmol, 1.5 eq) of 7-methylguanosine and 0.78 g (5 mmol) S.6a/b into a 100-mL round-bottom flask and add 20 mL of 0.02 M K2HPO4. Using pH paper, adjust the pH to 7.4 with 3 M NaOH, if necessary. 54. Add 250 units of purine nucleoside phosphorylase. Insert a septum and heat the mixture ∼3 days in an oven at 30°C with gentle agitation. Monitor the reaction for completeness each day by HPLC. 55. Pour the mixture into 10 mL DMF and stir ∼1 hr at room temperature.
Synthesis of Modified Nucleosides
1.6.7 Current Protocols in Nucleic Acid Chemistry
Supplement 10
Table 1.6.2 Nuclear Magnetic Resonance Chemical Shifts (ppm) for 15N-Labeled Adenosine (S.8) and Guanosine (S.13)a
5′H
N1
N7
NH2 C2
C8
— 8.35 8.14 7.40 5.87 4.60 4.14 3.96 3.6 10.6 7.92 NA 6.44 5.68 4.38 4.07 3.85 3.6
— 148
242 244
83 75
140 135
Compound N1H H8 S.8a/b S.13a/b
H2
NH2 1′H
2′H
3′H
4′H
152 154
aAll NMR samples in DMSO-d . 15N data are relative to 15NH using external 1M [15N]urea in DMSO at 77.0 ppm as a reference. 6 3
Additional data (e.g., for intermediates) are available in Pagano et al. (1995), Zhao et al. (1997), and Shallop and Jones (2000).
Table 1.6.3 Nuclear Magnetic Resonance Coupling Constants (Hz) for 15N-Labeled Adenosine (S.8) and Guanosine (S.13)
Compound
13C8-15N7
13C2-15N1
13C2-15NH
S.8a/b S.13a/b
<1 <1
NA 12
NA 24
2
56. Filter the suspension to remove most of the solid 7-methylguanine. Suspend the solid in 5 mL fresh DMF, stir 15 min, and filter. 57. Concentrate the combined filtrates to a small volume using a rotary evaporator and add 15 mL water (more if necessary to dissolve the solid). 58. Purify the crude S.7a/b by preparative reversed-phase HPLC and dry it over P2O5 in the vacuum desiccator overnight. The yield is usually 85% to 95%.
Synthesize [7,NH2-15N2]- and [8-13C-7,NH2-15N2]adenosine (S.8a/b) 59. Place 1.44 g (5 mmol) S.7a/b into a clean Teflon liner of a bomb and add 0.54 g (10 mmol, 2 eq) of [15N]NH4Cl and 7 mL anhydrous DMSO. 60. Add 1.5 g (15 mmol, 3 eq) KHCO3 and immediately seal the bomb. 61. Heat the bomb 3 days in an 80°C oven, swirling the mixture once or twice each day. 62. Cool the bomb to room temperature and then to −20°C for ≥30 min. Open it carefully and dilute the mixture with 10 mL water. Adjust the pH to 7 with glacial acetic acid. Care should be taken when opening the bomb because CO2 will have been generated.
63. Check the reaction by HPLC. 64. Purify S.8a/b by preparative reversed-phase HPLC and dry it over P2O5 in the vacuum desiccator overnight. The yield is usually 80% to 90%. See Tables 1.6.2 and 1.6.3 for NMR chemical shifts and coupling constants.
Synthesis of Specifically N-Labeled Adenosine and Guanosine 15
1.6.8 Supplement 10
Current Protocols in Nucleic Acid Chemistry
SYNTHESIS OF 7-METHYLGUANOSINE Although 7-methylguanosine can be purchased from Sigma, it is quite easy to make. Commercial 7-methylguanosine is very expensive and the purity may not be as high.
SUPPORT PROTOCOL
Materials Guanosine N,N-Dimethylacetamide Nitrogen gas source Dimethyl sulfate Concentrated aqueous NH3 Acetone, 4°C 95% (v/v) ethanol Ethyl ether 250-mL round-bottom flask Rubber septum 10-mL syringes CAUTION: Dimethyl sulfate is very dangerous because it is a potent alkylating agent; wear gloves and use caution. Synthesize 7-methylguanosine 1. Place 9.91 g (35 mmol) guanosine in a 250-mL round-bottom flask along with a stir bar. 2. Add 80 mL N,N-dimethylacetamide as a solvent, insert a rubber septum, and displace the air with nitrogen gas. 3. Carefully withdraw 7.0 mL (74 mmol, 2.1 eq) dimethyl sulfate with a 10-mL syringe and add it to the suspension. 4. Stir mixture ∼6 hr and monitor the reaction for completeness by HPLC. 5. Slowly add 10 mL concentrated aqueous NH3 and then check the pH using pH paper. Continue to add more NH3 slowly, frequently checking the pH, until the mixture is pH 10. This step is essential and quenches the excess dimethyl sulfate.
6. Slowly add the mixture to 300 mL cold acetone in an ice bath to precipitate the product. 7. Collect the white precipitate by vacuum filtration. 8. Check the acetone filtrate by HPLC for product. If there is a significant amount, concentrate the filtrate to a small volume, chill it, and collect the additional product by vacuum filtration. Combine it with the rest. Purify 7-methylguanosine 9. Suspend the solid crude product in 300 mL of 95% ethanol, stir 5 min, and collect it by filtration. 10. Suspend the solid product in 300 mL ethyl ether, stir 5 min, and collect it by filtration. 11. Dry the pure product over P2O5 in a vacuum desiccator overnight. The yield is usually 80% to 90%. It is very important to keep this compound dry. It should be transferred to a bottle with a tight lid, the air should be displaced with nitrogen, and it should be stored at −20°C. It can be kept for up to 3 months.
Synthesis of Modified Nucleosides
1.6.9 Current Protocols in Nucleic Acid Chemistry
Supplement 10
BASIC PROTOCOL 2
SYNTHESIS OF [2-13C-1,7,NH2-15N3]- AND [8-13C-1,7,NH2-15N3]GUANOSINE As shown in Figure 1.6.2, in the adenosine to guanosine transformation (Zhao, 1997; Shallop and Jones, 2000), the adenosine amino group becomes the guanosine N1, while the guanosine amino and C2 come from potassium cyanide. Thus, to make [2-13C1,7,NH2-15N3]guanosine (S.13a), [7,NH2-15N2]adenosine (S.8a) is used with [13C,15N]KCN, and to synthesize [8-13C-1,7,NH2-15N3]guanosine (S.13b), [8-13C-7,NH215 N2]adenosine (S.8b) is used with [15N]KCN. The first step is the oxidation of adenosine (S.8a/b) to the N1 oxide (S.9a/b), which is followed by a one-flask set of reactions without purification to give S.12a/b. Labeled cyanogen bromide is generated in situ from labeled potassium cyanide and bromine, and its reaction with S.9a/b forms S.10a/b. Treatment with triethylamine opens the oxazolidine ring, allowing the N1 oxide to be methylated by methyl iodide to give S.11a/b. Aqueous sodium hydroxide then opens the pyrimidine ring, which deformylates, rearranges, and closes again upon neutralization and heating to give S.12a/b. Enzymatic deamination then gives the final product, S.13a/b.
15
NH
15NH 15
N N
HO
15
N
N
X
O
N
H N Y 15
NH2
2
N
X HO
15
MCPBA
HO
OH
N
O HO
12
8a, X = C 8b, X =13C
N
O
O
15
N
N
X Br2/[ 13C,15N]KCN or
HO
Br2/[ 15N]KCN
OH
N
O HO
9a, X =12C 9b, X =13C
N
OH
10a, X =12C, Y =13C 10b, X =13C, Y =12C
1) Et 3 N 2) CH 3 I
O 15
N
X HO
O
N
HN 15
15N
NH Y 15 N NH2
X HO
O
N
N
O C H3 15
15
N Y
X
N
15
OH
13a, X =12C, Y =13C 13b, X =13C, Y =12C
Synthesis of Specifically N-Labeled Adenosine and Guanosine 15
HO
NH 2
O
N
Y15N N
O C H3
N
1) NaOH
ADA HO
H 15N
HO
OH
12a, X =12C, Y =13C 12b, X =13C, Y =12C
2) HCl
HO
OH
11a, X =12C, Y =13C 11b, X =13C, Y =12C
Figure 1.6.2 Steps for synthesis of [2-13C-1,7,NH2-15N3]- and [8-13C-1,7,NH2-15N3]guanosine using Basic Protocol 2. ADA, adenosine deaminase; Et3N, triethylamine; MCPBA, 3-chloroperoxybenzoic acid.
1.6.10 Supplement 10
Current Protocols in Nucleic Acid Chemistry
Materials [7,NH2-15N2]Adenosine or [8-13C-7,NH2-15N2]adenosine (S.8a or S.8b; see Basic Protocol 1) 50% (v/v) methanol 3-Chloroperoxybenzoic acid (MCPBA), purified (see recipe) Ethyl ether Phosphorous pentoxide (P2O5) [13C,15N]Cyanogen bromide or [15N]cyanogen bromide, freshly prepared (see recipe) 0.1 M potassium phosphate (KH2PO4), pH 7.5 Dimethyl formamide (DMF), anhydrous Acetonitrile Triethylamine Nitrogen gas source Methyl iodide 0.1 M NaOH 1 M HCl 95% (v/v) ethanol Adenosine deaminase (Sigma) 100-mL round-bottom flasks Rotary evaporator Vacuum desiccator Oil bath (silicone oil), 60°C Additional reagents and equipment for analytical and preparative reversed-phase high-performance liquid chromatography (HPLC; UNIT 10.5) Synthesize [7,NH2-15N2]- and [8-13C-7,NH2-15N2]adenosine-N1-oxide (S.9a/b) 1. Weigh 1.35 g (5.0 mmol) of [7,NH2-15N2]adenosine (S.8a) or [8-13C-7,NH215 N2]adenosine (S.8b) into a 100-mL round-bottom flask. Add a stir bar and 50 mL of 50% methanol. 2. Add 1.72 g (10 mmol, 2 eq) purified MCPBA, cover the flask with aluminum foil, stir 3 to 4 hr, and monitor the reaction for completeness by HPLC. Any reversed-phase column can be used.
3. Dilute the solution with 25 mL water and wash it with three 50-mL portions of ethyl ether. 4. Use a rotary evaporator to concentrate the aqueous solution to a small volume, purify S.9a/b by preparative reversed-phase HPLC, and dry it over P2O5 in a vacuum desiccator overnight in a 100-mL round-bottom flask. The yield is usually 90% to 95%. See Table 1.6.1 for various data on this and other intermediates and products.
Synthesize [2-13C-1,7,NH2-15N3]- and [8-13C-1,7,NH2-15N3]-2-amino-6(methoxyamino)-9-(β-D-ribofuranosyl)purine (S.12a/b) 5. Dissolve 1.43 g (5.0 mmol) S.9a/b in 40 mL water. 6. Add 7.5 mmol freshly prepared [13C,15N]cyanogen bromide or [15N]cyanogen bromide, stir for 2 hr, and monitor the reaction for completeness by HPLC. To make S.13a, [13C,15N]cyanogen bromide is used, and to make S.13b, [15N]cyanogen bromide is used.
Synthesis of Modified Nucleosides
1.6.11 Current Protocols in Nucleic Acid Chemistry
Supplement 10
CAUTION: The waste in the evaporator trap may contain excess cyanogen bromide. Dispose of this waste in an appropriately designated area, following the guidelines provided by the local safety officer.
7. Concentrate the solution to a very small volume using a rotary evaporator, add 10 mL anhydrous DMF and 10 mL acetonitrile, and concentrate again. Repeat this drying process two more times. CAUTION: Be very careful to dispose of the waste from the evaporator trap according to accepted regulations, because it may contain excess cyanogen bromide.
8. Add 25 mL anhydrous DMF and 2.8 mL (20 mmol, 4 eq) triethylamine under nitrogen gas. 9. Stir 45 min and then slowly add 2.5 mL (40 mmol, 8 eq) methyl iodide. Do not wait >1 hr to add the methyl iodide.
10. Cover the flask with aluminum foil and stir 3 to 4 hr while monitoring the reaction for completeness by HPLC. 11. Concentrate the solution to a yellow oil and add 85 mL of 0.1 M NaOH. Do not wait >4 hr to concentrate the solution and add the NaOH. CAUTION: The waste from the evaporator trap must be carefully disposed according to accepted regulations, because it may contain the excess methyl iodide.
12. Stir 20 min and then adjust the pH to 7.4 with 1 M HCl. 13. Add 80 mL of 95% ethanol, attach a condenser, and heat the solution in an oil bath at 60°C for 4 hr while monitoring the reaction for completeness by HPLC. 14. Concentrate the solution to a small volume using a rotary evaporator, purify S.12a/b by reversed-phase HPLC, and dry it over P2O5 in the vacuum desiccator overnight in a 100-mL round-bottom flask. The yield is usually 80% to 90%.
Synthesize [2-13C-1,7,NH2-15N3]- and [8-13C-1,7,NH2-15N3]guanosine (S.13a/b) 15. Dissolve 1.58 g (5 mmol) S.12a/b in 80 mL of 0.1 M KH2PO4, pH 7.5. 16. Add 300 units of adenosine deaminase, stopper the flask, and heat 4 days at 37°C with gentle agitation. Most of the product should crystallize out during this time.
17. Cool the mixture in an ice bath and collect crude S.13a/b by filtration. 18. Recrystallize S.13a/b by recrystallization from water and dry it over P2O5 in a vacuum desiccator overnight. The yield is usually 75% to 85%. See Tables 1.6.2 and 1.6.3 for NMR chemical shifts and coupling constants.
Synthesis of Specifically N-Labeled Adenosine and Guanosine 15
1.6.12 Supplement 10
Current Protocols in Nucleic Acid Chemistry
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
3-Chloroperoxybenzoic acid (MCPBA) Dissolve 10 g MCPBA in 200 mL ether and wash with three 150-mL portions of 0.1 M aqueous potassium phosphate, pH 7.5. Concentrate to dryness and dry over P2O5 in a vacuum desiccator overnight. Store up to 3 months at −20°C in a bottle with a tight cap. CAUTION: MCPBA is potentially explosive and must be handled with care. Commercial MCPBA is contaminated with 40% to 50% 3-chlorobenzoic acid, which can be removed by extraction.
[13C,15N]Cyanogen bromide Place 3 mL water into a small pear-shaped flask and add 1.2 g (0.38 mL, 7.5 mmol) bromine. Chill the flask in an ice bath, slowly add 0.50 g (7.5 mmol, 1 eq) [13C,15N]potassium cyanide dissolved in 20 mL water, and stir 30 min. Draw the solution into a syringe in order to add it to a reaction. CAUTION: Use great care in handling bromine, because it is highly toxic, is fairly volatile, and can cause severe burns. Place a balance in a hood, draw up the required volume of bromine into a preweighed syringe, weigh the syringe, and adjust accordingly. Potassium cyanide and cyanogen bromide are also highly toxic and must be handled with great care.
[15N]Cyanogen bromide Prepare as for [13C,15N]cyanogen bromide (see recipe), but use [15N]potassium cyanide instead of [13C,15N]potassium cyanide. [13C]Sodium ethyl xanthate ([13C]NaSCSOEt) Dissolve 0.40 g (10 mmol) NaOH in 40 mL absolute ethanol. Add 0.77 g (10 mmol, 1 eq) [13C]carbon disulfide ([13C]CS2; Isotec or Cambridge Isotope Laboratories). Cover the flask with aluminum foil and stir the solution overnight at room temperature. Concentrate to dryness and dry over P2O5 in a vacuum desiccator overnight. Store up to 6 months at −20°C. The yield is usually quantitative.
COMMENTARY Background Information 15N NMR studies of specifically 15N-labeled
DNA and RNA fragments have provided significant information about local interactions at nitrogen atoms, such as hydrogen bonding, stacking, and protonation (Wang et al., 1991; Zhang et al., 1997, 1998). The use of one or more multilabeled nucleosides can provide more information than single-labeled nucleosides, but only as long as all signals can be distinguished. The use of 13C tags adjacent to different nitrogens in a pair of nucleosides was designed to allow such differentiation (Zhao, 1997; Abad et al., 1998; Shallop and Jones, 2000). In addition, the 13C chemical shifts can provide valuable information. The route to labeled adenosine described here (Pagano et al., 1995) starts with a direct
nitrosation, which is more convenient than the azo coupling described earlier (Gaffney et al., 1990). This route has been designed so as to remove both thiol groups (in the case of S.4b) in the same step. The enzymatic coupling is done with 6-chloropurine, because the subsequent displacement with [15N]NH3 can be done using much milder conditions. Also, the lipophilicity of the 6-chloro group helps in the purification of the nucleoside. The purine nucleoside phosphorylase uses 7-methylguanosine as a sugar donor and both generates the ribose-α-1-phosphate and couples it to the 6chloropurine. The reaction is driven to completion by precipitation from solution of the very insoluble 7-methylguanine. The final ammination has been optimized so that it only requires two equivalents of [15N]NH4Cl with KHCO3.
Synthesis of Modified Nucleosides
1.6.13 Current Protocols in Nucleic Acid Chemistry
Supplement 10
Although not described here, [7,NH215N ]adenosine can be converted to [1,7,NH 2 2 15N ]adenosine (Pagano et al., 1998). 3 The adenosine to guanosine transformation described here (Zhao, 1997; Shallop and Jones, 2000) is based on a previous method (Goswami and Jones, 1991) that was in turn derived from earlier work (Ueda et al., 1978).
Critical Parameters In the chlorination that converts S.5a/b to S.6a/b, it is essential to remove all excess POCl3, or heat generated during subsequent neutralization with ammonia will cause the reaction to reverse to some extent. During the one-flask set of reactions for the adenosine to guanosine transformation, the intermediates are not particularly stable, and the reactions should not be left for longer than the stated times. After formation of cyanogen bromide, there should be no excess bromine present (as indicated by an orange color). The order of addition during the formation of cyanogen bromide is also important. In the final enzymatic deammination to make S.13a/b, S.12a/b must be purified, or the enzymatic reaction will not work satisfactorily.
Anticipated Results The early steps in these protocols generally give high yields, even for inexperienced workers. The later steps are more challenging, and, with experience, the stated yields can be obtained. Because most of these compounds are fairly polar, the use of TLC to monitor reactions is not very convenient. Reversed-phase HPLC is somewhat more informative, while UV λmax data from an HPLC system with a diode array detector are particularly helpful.
Time Considerations The total time for Basic Protocol 1 is 2 to 3 weeks and that for Basic Protocol 2 is 1 to 2 weeks. The Support Protocol requires 1 to 2 days.
Gaffney, B.L., Kung, P.-P., and Jones, R.A. 1990. Nitrogen-15-labeled deoxynucleosides. 2. Synthesis of [7-15N]-labeled deoxyadenosine, deoxyguanosine, and related deoxynucleosides. J. Am. Chem. Soc. 112:6748-6749. Goswami, B. and Jones, R.A. 1991. Nitrogen-15-labeled deoxynucleosides. 4. Synthesis of [115N]and [2-15N]-labeled 2′-deoxyguanosines. J. Am. Chem. Soc. 113:644-647. Pagano, A.R., Lajewski, W.M., and Jones, R.A. 1995. Synthesis of [6,7-15N]-adenosine, [6,715 N]-2′-deoxyadenosine, and [7-15N]-hypoxanthine. J. Am. Chem. Soc. 117:11669-11672. Pagano, A.R., Zhao, H., Shallop, A., and Jones, R.A. 1998. Synthesis of [1,7-15N2]- and [1,7, NH215 N3]-adenosine and 2′-deoxyadenosine via an N1-alkoxy mediated Dimroth rearrangement. J. Org. Chem. 63:3213-3217. Shallop, A.J. and Jones, R.A. 2000. Use of a 13C “indirect tag” to differentiate two 15N7 specifically labeled nucleosides. Middle Atlantic Regional Meeting, ACS, May 15, 2000, Abstr. 33:215-ORGN. Ueda, T., Miura, K., and Kasai, T. 1978. Synthesis of 6-thioguanine and 2,6-diaminopurine nucleosides and nucleotides from adenine counterparts via a facile rearrangement in the base portion. Chem. Pharm. Bull. 26:2122-2127. Wang, C., Gao, H., Gaffney, B.L., and Jones, R.A. 1991. Nitrogen-15-labeled deoxynucleotides. 3. Protonation of the adenine N1 int he A⋅C and A⋅G m isp a irs of the du plexes an d [d[CG(15N1)AGAATTCCCG]}2 {d[CGGGAATTC(15N1)ACG]}2. J. Am. Chem. Soc. 113:5486-5488. Zhang, X., Gaffney, B.L., and Jones, R.A. 1997. 15N NMR of a specifically labeled RNA fragment containing intrahelical GU wobble pairs. J. Am. Chem. Soc. 119:6432-6433. Zhang, X., Gaffney, B.L., and Jones, R.A. 1998. 15N NMR of RNA fragments containing specifically labeled tandem G⋅A pairs. J. Am. Chem. Soc. 120:6625-6626. Zhao, H., Pagano, A.R., Wang, W., Shallop, A., Gaffney, B.L., and Jones, R.A. 1997. Use of a 13 C atom to differentiate two 15N-labeled nucleosides: syntheses of [15NH2]-adenosine, [1,NH2-15N2] - an d [2-13C-1,NH2-15N2]guanosine, and [1,7,NH2-15N3]- and [2-13C1,7,NH2-15N3]-2′-deoxyguanosine. J. Org. Chem. 62:7832-7835.
Literature Cited Abad, J.-L., Shallop, A.J., Gaffney, B.L., and Jones, R.A. 1998. Use of 13C tags with specifically 15 N-labeled DNA and RNA. Biopolymers 48:5763.
Contributed by Barbara L. Gaffney and Roger A. Jones Rutgers University Piscataway, New Jersey
Synthesis of Specifically N-Labeled Adenosine and Guanosine 15
1.6.14 Supplement 10
Current Protocols in Nucleic Acid Chemistry
Synthesis of Protected 2′-Deoxy-2′-fluoro-β-D-arabinonucleosides
UNIT 1.7
This unit describes in detail the preparation of protected 2′-deoxy-2′-fluoroarabinonucleosides. These building blocks are required for the synthesis of 2′-deoxy-2′-fluoroarabinonucleic acid (2′F-ANA), an oligonucleotide analog exhibiting very promising antisense properties (Damha et al., 1998; Wilds and Damha, 2000; Lok et al., 2002). The preparation of phosphoramidites from these building blocks and the synthesis of 2′F-ANA are described in UNIT 4.15. SYNTHESIS AND CHARACTERIZATION OF N2-ISOBUTYRYL-9[2-DEOXY-2-FLUORO-5-O-(4-METHOXYTRITYL)-β-DARABINOFURANOSYL]GUANINE
BASIC PROTOCOL 1
Synthesis of araF-G (S.6; Figure 1.7.3) was accomplished via the condensation of 2,6-dichloropurine with either 2-deoxy-2-fluoro-1,3,5-tri-O-benzoyl-α-D-arabinofuranose (S.2; Figure 1.7.1) or 2-deoxy-2-fluoro-3,5-di-O-benzoyl-α-D-arabinofuranosyl bromide (S.3) as a key chemical step (Figure 1.7.2). Starting from S.3 gives a higher yield of S.4 and simplifies its purification. The versatile intermediate N9-β-glycoside (S.4) was transformed into araF-G in three steps (Figure 1.7.3). Treatment of S.4 with NaN3 in ethanol gave the 2,6-diazido derivative (Wower et al., 1994). This product was then subjected to reduction with SnCl2 in a mixture of dichloromethane/methanol to give its 2,6-diamino derivative (Tennila et al., 2000). Standard debenzoylation and deamination gave araF-G. Transient protection of the 3′- and 5′-OH of araF-G (Kierzek, 1985), followed by acylation at N2 and 5′-O-tritylation, gave S.7 (Figure 1.7.3) in acceptable yields. Materials Nitrogen gas source 1,3,5-Tri-O-benzoyl-α-D-ribofuranose (S.1; Pfanstiehl) Dichloromethane, dry (see recipe) [Bis(2-methoxyethyl)amino]sulfur trifluoride (MAST; Aldrich) or (diethylamino)sulfur trifluoride (DAST; Aldrich) Saturated aqueous sodium bicarbonate Sodium sulfate (Na2SO4), anhydrous Silica gel (230 to 400 mesh) Chloroform
BzO
O BzO
BzO
OBz CH Cl , 40o-50o C 2 2 HO 1
BzO
O F
MAST or DAST
OBz
BzO
O F
HBr/CH2Cl2 rt, 24 hr
BzO
2 (83%)
Br 3 (98%)
Figure 1.7.1 Synthesis of 2-deoxy-2-fluoro-3,5-di-O-benzoyl-α-D-arabinofuranosyl bromide (S.3). The expected yields are given in parentheses. Bz, benzoyl; DAST, diethylaminosulfur trifluoride; HBr, 30% (w/v) hydrogen bromide in acetic acid; MAST, [bis(2-methoxyethyl)amino]sulfur trifluoride.
Synthesis of Modified Nucleosides
Contributed by Mohamed I. Elzagheid, Ekaterina Viazovkina, and Masad J. Damha
1.7.1
Current Protocols in Nucleic Acid Chemistry (2002) 1.7.1-1.7.19 Copyright © 2002 by John Wiley & Sons, Inc.
Supplement 10
Cl N BzO
N H
O F Br
BzO
N Cl
N
NaH, CH3CN, 50o C, 48 hr
silica gel chromatography (35%-40%)
3 (98%)
Cl N BzO
O F
N
N N
Cl
BzO 4 (N9-β-isomer) Cl N BzO
N N Si(CH3)3
O F OBz
BzO
N
silica gel chromatography (38%)
Cl
TMS-Tfl, CH3CN, reflux
2 (83%)
Figure 1.7.2 Synthesis of 2,6-dichloro-9-(3,5-di-O-benzoyl-2-deoxy-2-fluoro-β-D-arabinofuranosyl)purine (S.4) from S.3 or S.2. The expected yields are given in parentheses. Bz, benzoyl; NaH, 60% (w/v) sodium hydride in oil; TMS-Tfl, trimethylsilyl trifluoromethanesulfonate.
Cl N BzO
O F
N
NH2 N
N N
Cl
BzO
1. NaN3, EtOH (97%) 2. SnCl2, CH2Cl2/CH3OH (83%)
BzO
N
O F
N N
NH2
BzO
4
5 1. NH4OH/CH3OH 2. ADase O N
MMTrO
O F
N
N
NH N
NHi-Bu
1. TMSCl/(CH3)2CHCOCl/Py 2. NH4OH/H2O 3. MMTrCl/Py/4-DMAP
HO 7 (66%)
Synthesis of Protected 2′-Deoxy-2′-fluoroβ-D-arabinonucleosides
O
HO
O F
N
NH N
NH2
HO 6 (araF-G) (79%)
Figure 1.7.3 Synthesis of araF-G (S.6) in four steps from S.4, followed by introduction of isobutyryl (i-Bu) and monomethoxytrityl (MMTr) groups into the N2 and O5′ positions, respectively, to give S.7. The expected yields are given in parentheses. ADase, adenosine deaminase; 4-DMAP, 4-dimethylaminopyridine; EtOH, 95% (v/v) ethanol; NH4OH, 29.7% (w/w) aqueous ammonia; MMTr⋅Cl, p-anisylchlorodiphenylmethane (or monomethoxytrityl chloride); Py, pyridine; TMSCl, chlorotrimethylsilane.
1.7.2 Supplement 10
Current Protocols in Nucleic Acid Chemistry
Sand Merck thin-layer chromatography (TLC) silica plates (Kieselgel 60 F-254; 0.2 mm thick) 30% (w/v) HBr in acetic acid Chlorotrimethylsilane (TMSCl) 2,6-Dichloropurine Hexamethyldisilazane (HMDS) Acetonitrile, dry (see recipe) Trimethylsilyl trifluoromethanesulfonate (TMS-Tfl) 5:1 (v/v) toluene/ethyl acetate Sodium hydride, as a 60% (w/v) solution in mineral oil (Aldrich) Sodium azide (NaN3) 95% (v/v) ethanol Tin dichloride (SnCl2) Methanol 29.7% (w/w) aqueous ammonia (Fisher) 5% and 10% (v/v) methanol in dichloromethane Adenosine deaminase solution in 50% glycerol (from calf intestine mucosa, specific activity 160 to 200 U/mg protein; Sigma) Phosphorus pentoxide (P2O5) Pyridine, dry (see recipe) Isobutyryl chloride 2:1:1 (v/v/v) water/ethyl acetate/ether 4-Dimethylaminopyridine (4-DMAP) p-Anisylchlorodiphenylmethane (monomethoxytrityl chloride or MMTr⋅Cl) 0% to 5% (v/v) gradient of methanol in dichloromethane Oven-dried glassware, including: 25-, 100-, and 250-mL round-bottom flasks 1-L Erlenmeyer flasks Reflux condenser Oil bath, 40° to 50°C 1-L separatory funnels Rotary evaporator equipped with a vacuum pump or water aspirator Chromatography columns: 5 × 50 cm, 3 × 15 cm, and 3 × 20 cm Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Fluorinate 1,3,5-tri-O-benzoyl-α-D-ribofuranose to give S.2 1. In an oven-dried 250-mL round-bottom flask equipped with a reflux condenser, a stir bar, and nitrogen gas source, dissolve 10 g S.1 in 100 mL dry dichloromethane. 2. While stirring, add 8 mL (43.39 mmol) MAST or 9 mL (68.12 mmol) DAST dropwise. It is better to use MAST. It is less expensive, gives higher yields, and makes purification easier. CAUTION: DAST is a flammable and corrosive liquid. MAST is a toxic and corrosive liquid. Both must be handled with gloves in a well-ventilated fume hood. In case of skin contact freely apply calcium gluconate gel (Pharmascience). Reapply and continue application for an additional 10 to 15 min while seeking medical assistance. Synthesis of Modified Nucleosides
1.7.3 Current Protocols in Nucleic Acid Chemistry
Supplement 10
3. Place the flask in an oil bath preheated to 40° to 50°C and stir 24 hr or until the reaction is complete as analyzed by TLC using Merck TLC (APPENDIX 3D) silica plates. The starting material should be run alongside the reaction for comparison. The plates are developed using dichloromethane, and the bands are visualized by UV shadowing and dipping the plate in 10% (v/v) sulfuric acid in methanol followed by heating. Typically, the Rf value of the desired compound (S.2) is 0.34 (dichloromethane).
4. Dilute the reaction mixture with 200 mL dichloromethane and add it carefully to 300 mL saturated aqueous sodium bicarbonate solution. 5. Pour the mixture into a 1-L separatory funnel and allow the phases to separate. Pour the lower, yellow organic phase into a 1-L Erlenmeyer flask. 6. Dry the organic layer over anhydrous Na2SO4. Filter and evaporate the filtrate to dryness in a rotary evaporator equipped with a vacuum pump or water aspirator to give a yellow solid. 7. Prepare a slurry of 350 g silica gel in chloroform. Pour slurry into a 5 × 50–cm chromatography column and carefully layer 2 cm sand on top of slurry (see APPENDIX 3E for column chromatography). 8. Dissolve the crude product (step 6) in a minimal amount of chloroform and layer it carefully on top of the column. 9. Elute with chloroform and collect 200-mL fractions (typically a total of 2 to 4 L). Combine fractions that contain pure product, as determined by TLC. Evaporate to dryness in a rotary evaporator and dry overnight under high vacuum. Typically, the Rf value of the desired product (S.2) is 0.34 (dichloromethane).
10. Check purity of the product. 2-Deoxy-2-fluoro-1,3,5-tri-O-benzoyl-α-D-arabinofuranose (S.2): 8.3 g (83%); TLC (dichloromethane) 0.34; 1H NMR (400 MHz, acetone-d6, tetramethylsilane as internal reference): 8.1 to 7.4 (15 H, m, Bz), 6.7 (1H, d, J1,F = 9.2 Hz, H-1), 5.6 and 5.7 (1H, dd, J3,F = 20 HZ, J3,2 = 3.6 HZ, H-3), 5.5 to 5.7 ( 1H, d, J2,F = 48 Hz, H-2), 4.9 (1H, m, H-4), 4.7 to 4.8 (2H, m, H-5 and H-5′); 19F NMR (300 MHz, dimethyl sulfoxide [DMSO]-d6, 99% trifluoracetic acid as external standard): −113 (ddd). To prepare S.4 from S.2, proceed to step 17a. It is better, however, to brominate as in steps 11 to 16 and use S.3 in the condensation reaction (go to step 17b). This gives a higher yield of S.4, and affords only the N7- and N9-β-isomers, making purification easier.
Brominate to give S.3 11. In an oven-dried 100-mL round-bottom flask, dissolve 2.2 g (4.85 mmol) S.2 in 30 mL dry dichloromethane and add 4 mL of 30% HBr in acetic acid. 12. Stir the reaction mixture 24 hr at room temperature. 13. Analyze the reaction by TLC. The starting material should be run alongside the reaction for comparison. The plates are developed using dichloromethane, and the bands are visualized by UV shadowing and dipping the plate in 10% (v/v) sulfuric acid in methanol followed by heating. Typically, the Rf value of the desired compound (S.3) is ∼0.55 (dichloromethane).
14. Dilute the reaction mixture with 100 mL dichloromethane and add it carefully to 200 mL saturated aqueous sodium bicarbonate. Synthesis of Protected 2′-Deoxy-2′-fluoroβ-D-arabinonucleosides
15. Pour the mixture into a clean 1-L separatory funnel and allow the phases to separate. Pour the lower, brown organic phase into a 1-L Erlenmeyer flask. Dry the organic layer over anhydrous Na2SO4. Filter and evaporate the filtrate to dryness in the rotary evaporator to give a brown oil.
1.7.4 Supplement 10
Current Protocols in Nucleic Acid Chemistry
16. Check the purity of the product. 2-Deoxy-2-fluoro-3,5-di-O-benzoyl-α-D-arabinofuranosyl bromide (S.3): 2.0 g (98%); TLC (dichloromethane) 0.55; 1H NMR (500 MHz, CD2Cl2 ): 8.2 to 7.3 (10 H, m, Bz), 6.8 (1H, d, J1,F = 19 Hz, H-1), 5.6 (1H, d, J2,F = 50 Hz, H-2), 5.4 (1H, dd, H-3), 4.8 (3H, m, H-4, H-5, and H-5′′).
Condense 2,6-dichloropurine with S.2 or S.3 For S.2: 17a. In an oven-dried 250-mL round-bottom flask equipped with a reflux condenser, stir bar, and nitrogen gas source, add 4 mL TMSCl to a suspension of 2.84 g (15 mmol) 2,6-dichloropurine in 40 mL HMDS. Reflux 2 to 3 hr at 120°C, cool down, and evaporate to dryness in the rotary evaporator. 18a. Azeotrope (co-evaporate) the residue with 50 mL dry acetonitrile. To this residue, add 4 g (8.0 mmol) S.2 in 100 mL dry acetonitrile followed by 4 mL TMS-Tfl. Reflux the resulting solution 45 min at 80°C. 19a. Analyze the reaction by TLC. The starting material should be run alongside the reaction for comparison. The plates are developed using 9:1 (v/v) toluene/ethyl acetate, and the bands are visualized by UV shadowing and dipping the plate in 10% (v/v) sulfuric acid in methanol followed by heating. Typically, the Rf value of the desired product, the N9-β-isomer (S.4), is 0.28 (9:1 toluene/ethyl acetate).
20a. Cool the mixture, dilute with 200 mL dichloromethane, and wash it carefully with 300 mL saturated sodium bicarbonate solution. 21a. Dry the dichloromethane layer over anhydrous Na2SO4, filter, and evaporate to dryness. 22a. Prepare a 5 × 50–cm silica chromatography column as described in step 7. Apply the residue (step 21a) to the column and elute with a mixture of 5:1 toluene/ethyl acetate. Collect 50-mL fractions and combine those that contain pure product (S.4) as determined by TLC. 23a. Evaporate combined fractions to dryness in the rotary evaporator and dry overnight under a high vacuum. Proceed to step 24. For S.3: 17b. In a clean oven-dried 250-mL round-bottom flask equipped with reflux condenser, stir bar, and nitrogen gas source, dissolve 0.54 g (2.84 mmol) of 2,6-dichloropurine and 0.07 g (2.95 mmol) sodium hydride in 30 mL dry acetonitrile. Stir 45 min at room temperature. 18b. Dissolve 1.2 g (2.84 mmol) S.3 in 20 mL dry acetonitrile and add it in portions to the flask. Place the flask in an oil bath and stir 48 hr at 50°C or until the reaction is complete as determined by TLC. The starting material should be run alongside the reaction for comparison. The plates are developed using 9:1 (v/v) toluene/ethyl acetate, and the bands are visualized by UV shadowing and dipping the plate in 10% (v/v) sulfuric acid in methanol followed by heating. Typically, the Rf value of the desired N9-β-isomer (S.4) is 0.28 (9:1 toluene/ethyl acetate). Side products have Rf values in the range of 0.20 to 0.23 with the same solvent system.
19b. Vacuum filter the resulting mixture and evaporate to dryness.
Synthesis of Modified Nucleosides
1.7.5 Current Protocols in Nucleic Acid Chemistry
Supplement 12
20b. Prepare a 5 × 50–cm silica chromatography column as described in step 7. Apply the residue (step 19b) to the column and elute with 5:1 toluene/ethyl acetate. Collect 50-mL fractions and combine those that contain the pure product (S.4) as determined by TLC. 21b. Evaporate combined fractions to dryness in the rotary evaporator and dry overnight under high vacuum. Proceed to step 24. 24. Check the purity of the product. 9-(2-Deoxy-2-fluoro-3,5-di-O-benzoyl-β-D-arabinofuranosyl)-2,6-dichloropurine (S.4): average yield of N9-β-isomer is 38% from each condensation (i.e., 2,6-dichloropurine with S.2 or S.3); TLC (9:1 [v/v] toluene/ethyl acetate) 0.28; 1H NMR (400 MHz, CDCl3 ): 8.1 to 7.1 (10 H, m, Bz), 8.4 (1H, d, J8,F = 3.2 Hz, H-8), 6.6 (1H, dd, J1′,2′ = 2.8 Hz, J1′,F = 22 Hz, H-1′), 5.7 (1H, dd, J3′,F = 17 Hz, J3′,4′ = 2.8 Hz, H-3′), 5.4 (1H, dd, J2′, F = 50 Hz, J2′,3′ = 2.0 Hz, H-2′), 4.7 to 4.8 (2H, dd, H-5′ and H-5′′), 4.61 (1H, m, H-4′); 19F NMR (300 MHz, DMSO-d6, 99% [v/v] trifluoroacetic acid as external reference): −113 (ddd). FAB-MS (fast atom bombardment mass spectrometry, NBA-matrix): 531 [M+H+].
Transform N9-β-glycoside (S.4) into araF-G (S.6) 25. In a clean oven-dried 250-mL round-bottom flask equipped with reflux condenser and stir bar, dissolve 0.4 g (0.75 mmol) S.4 and 0.25 g (3.7 mmol) NaN3 in 50 mL of 95% ethanol. Use sodium azide instead of lithium azide to avoid partial debenzoylation of the intermediary 2,6-diazido derivative.
26. Reflux the reaction mixture 2 hr in the 80°C oil bath. Cool mixture and evaporate to dryness. Precipitation of sodium chloride is a good sign of a successful reaction. The Rf values for the 2,6-diazido and 2,6-dichloro derivatives are 0.48 and 0.52, respectively, in 3:1 (v/v) toluene/ethyl acetate.
27. Dissolve the residue in dichloromethane, wash the organic layer with water, dry it over anhydrous Na2SO4, and evaporate to yield a syrupy product. A yield of 0.4 g (97%) is expected. Silica column chromatography is not required after this step.
28. Treat 0.4 g (0.73 mmol) of 2,6-diazidopurine nucleoside (step 27) with 0.42 g (2.22 mmol) SnCl2 in a mixture of 30 mL dichloromethane and 3 mL methanol for 40 min at room temperature. The Rf value for the 2,6-diamino derivative (S.5) is 0.42 in 9:1 (v/v) dichloromethane/methanol.
29. Dilute the mixture with 100 mL dichloromethane, wash the organic layer with water, dry it over anhydrous Na2SO4, and evaporate to yield a white solid. The expected yield is 0.3 g (83%). Silica column chromatography is not required after this step.
30. Treat 0.3 g (0.61 mmol) S.5 with a mixture containing 3 mL of 29.7% aqueous ammonia and 12 mL methanol for 48 hr at room temperature. Synthesis of Protected 2′-Deoxy-2′-fluoroβ-D-arabinonucleosides
The Rf value for the desired product is 0.06 in 9:1 (v/v) dichloromethane/methanol and 0.16 in 3:1 (v/v) chloroform/ethanol.
1.7.6 Supplement 12
Current Protocols in Nucleic Acid Chemistry
31. Evaporate the resulting solution to dryness. Dissolve the crude product in a minimal amount of 5% methanol in dichloromethane and place it carefully on the top of a 3 × 15–cm chromatography column containing 25 g silica gel. 32. Elute with 10% methanol in dichloromethane and collect 50-mL fractions. Combine fractions containing the desired product as determined by TLC and evaporate to dryness in the rotary evaporator to yield a white foam. 33. Check the product. 9-(2-Deoxy-2-fluoro-β-D-arabinofuranosyl)-2,6-diaminopurine (S.5): 0.14 g (83%); TLC (3:1 [v/v] chloroform/ethanol) 0.16, (9:1 [v/v] dichloromethane/methanol) 0.06; UV (H2O) λmax, 280 nm; FAB-MS (NBA-matrix): 285 [M+H+], 307 [M+Na+].
34. Treat 0.14 g (0.49 mmol) of the product with 100 µL adenosine deaminase solution in 5 mL water for 24 hr at room temperature. 35. Collect the precipitated white material by filtration and wash it with 10 mL water and then with 10 mL methanol. Dry the white powder (S.6) over P2O5. This reaction has also been performed on a larger scale using 0.5 g substrate. A yield of 71% was obtained.
36. Check purity of the product. 9-(2-Deoxy-2-fluoro-β-D-arabinofuranosyl)guanine (S.6): 0.11 g (79%); UV (H2O) λmax, 251 nm; 1H NMR (400 MHz, DMSO-d6): 10.6 (1H, s, N-H), 7.8 (1H, d, J8,F = 2.8 Hz, H-8), 6.5 (2H, br s, NH2), 6.1 (1H, dd, J1′,2′ = 4.4 Hz, J1′,F = 16 Hz, H-1′), 5.9 (1H, d, JOH,3′ = 4.8 Hz, HO-C3′), 5.0 and 5.1 (1H, dt or ddd, J2′,F = 33 Hz, J2′,3′ = 3.6 Hz, H-2′), 5.0 (1H, t, HO-C5′), 4.3 (1H, m, J3′,F = 14 Hz, H-3′), 3.8 (1H, m, H-4′), 3.6 (2H, m, H-5′ and H-5′′); 19 F NMR (300 MHz, DMSO-d6, 99% [v/v] trifluoroacetic acid as external reference): −120 (ddd); FAB-MS (NBA-matrix): 286 [M+H+]. See Figure 1.7.4 for 1H NMR.
Isobutyrylate and monomethoxytritylate to give S.7 37. In a clean oven-dried 25-mL round-bottom flask, dissolve 0.3 g (1.05 mmol) araF-G (S.6) in 5 mL dry pyridine, add 3 mL TMSCl, and stir mixture 45 min at room temperature. 38. Add 500 µL isobutyryl chloride and stir 2 hr at room temperature. 39. Immerse the reaction flask in an ice bath and add 1 mL water. After 10 min, add 1 mL of 29.7% aqueous ammonia and stir 20 min at room temperature. The Rf value for N2-isobutyryl-9-(2-deoxy-2-fluoro-β-D-arabinofuranosyl)guanine is 0.45 in 3:1 (v/v) chloroform/ethanol.
40. Carefully evaporate the resulting mixture to near dryness and take up the residue in 120 mL 2:1:1 water/ethyl acetate/ether. Evaporate the aqueous layer to get the desired compound as a colorless powder. 41. Check purity of the product. N2-Isobutyryl-9-(2-deoxy-2-fluoro-β-D-arabinofuranosyl)guanine: 0.35 g (94%); TLC (3:1 [v/v] chloroform/ethanol) 0.45; 1H NMR (300 MHz, DMSO-d6): 12.1 (1H, N-H), 11.7 (1H, N-H), 8.1 (1H, d, J8,F = 2.4 Hz, H-8), 6.17 (1H, dd, J1′,2′ = 5.6 Hz, J1′,F = 14 Hz, H-1′), 6.0 (1H, d, JOH,3′ = 6.0 Hz , HO-C3′), 5.1, 5.2 (1H, dt or ddd, J2’,F = 70 Hz, H-2’), 4.3 (1H, ddd, H-3′), 3.8 (1H, dd, H-4′), 3.6 (2H, m, H-5′ and H-5′′, 2.5(1H, m, H-C[CH3]2), 1.1 (6H, s,[CH3]2- CH).
42. Dissolve 0.3 g (0.85 mmol) of the product in 20 mL dry pyridine and add catalytic amount (∼30 mg) 4-DMAP and 0.4 g (1.3 mmol) MMTr⋅Cl. Stir mixture 48 hr at room temperature. The Rf value for the desired tritylated product is 0.41 in 9:1 (v/v) chloroform/methanol.
Synthesis of Modified Nucleosides
1.7.7 Current Protocols in Nucleic Acid Chemistry
Supplement 12
O N HO
O F
H-1 ′
NH
N
N
NH2
HO 6 (araF-G)
′ 5-OH
6.1
H-8
′ 3-OH
NH2
H-5 ′, H-5 ′′
H-4 ′ H-1 ′
NH
4.4
H-2 ′
7.80
11
Figure 1.7.4
10
7.78
9
4.3
H-3 ′
7.76
3.8
8
7
6
5
4
3
3.7
2
3.6
3.5
1
ppm
1
H NMR spectrum (400 MHz) of 9-(2-deoxy-2-fluoro-β-D-arabinofuranosyl)guanine, araF-G (S.6), in DMSO-d6.
43. Evaporate pyridine, take up the residue in dichloromethane, wash the organic layer with saturated aqueous sodium bicarbonate, dry organic layer over anhydrous Na2SO4, and evaporate to dryness. 44. Dissolve the crude product in a minimal amount of dichloromethane and place it carefully on the top of a 3 × 20–cm chromatography column containing 38 g silica gel. 45. Elute with 0% to 5% methanol in dichloromethane and collect 50-mL fractions. Combine those containing pure product as determined by TLC. 46. Evaporate combined fractions to dryness in the rotary evaporator to get the desired product as a yellow foam. 47. Check purity of the final product.
Synthesis of Protected 2′-Deoxy-2′-fluoroβ-D-arabinonucleosides
N2-Isobutyryl-9-[2-deoxy-2-fluoro-5-O-(4-methoxytrityl)-β-D-arabinofuranosyl]guanine (S.7): 0.35 g (66%); TLC (9:1 [v/v] chloroform/methanol); 1H NMR (400 MHz, acetoned6): 7.8 (1H, d, J8,F = 3.2 Hz, H-8), 7.6 to 6.8 (14H, m, trityl), 6.2 (1H, dd, J1′,2′ = 6 Hz, J1′,F = 16 Hz, H1′), 5.1 (1H, dt or ddd, J2′,F = 54 Hz, H-2′), 4.6 (1H, m, H-3′), 4.2 (1H, m, H-4′), 3.6 (3H, s, CH3O), 3.5 and 3.2 (2H , 2dd, H-5′ and H-5′′), 2.1 (1H, m, H-C[CH3]2), 1.2 and 1.3 (6H, s, [CH3]2-CH).
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BzO
O F
NABs/solvent
BzO
O F
Br
BzO
BzO 3
B 1. deprotection 2. 5’-tritylation 3. N-benzoylation
8a B = N 4-acetylcytosin-1-yl (55%) 8b B = thymin-1-yl (75%) 8c B = N 6-benzoyladenin-9-yl (58%)
MMTrO
O F
B
OH 9a B = N 4-benzoylcytosin-1-yl (58%) 9b B = thymin-1-yl (87%) 9c B = N 6-benzoyladenin-9-yl (52%)
Figure 1.7.5 Synthesis of 2′-deoxy-2′-fluoro-β-D-arabinonucleosides (S.9a-c). The expected yields are given in parentheses. Bz, benzoyl; MMTr, p-anisyldiphenylmethyl; NABs, nucleic acid bases (silylated-N-acetylcytosine or silylated-thymine or N-benzoylated-adenine); solvent, dichloromethane or carbon tetrachloride.
SYNTHESIS AND CHARACTERIZATION OF N4-BENZOYL-1-[2-DEOXY-2FLUORO-5-O-(4-METHOXYTRITYL)-β-D-ARABINOFURANOSYL]CYTOSINE Synthesis of araF-C was accomplished via the condensation of N4-acetylcytosine with 2-deoxy-2-fluoro-3,5-di-O-benzoyl-α-D-arabinofuranosyl bromide (S.3) as a key chemical step (Figure 1.7.5). This was followed by deprotection, benzoylation, and tritylation to give S.9a. Cytosine, N4-benzoylcytosine, or N4-acetylcytosine can be used for the condensation step. Better yields are observed with N4-acetylcytosine.
BASIC PROTOCOL 2
Materials Nitrogen gas source Chlorotrimethylsilane (TMSCl) N4-Acetylcytosine Hexamethyldisilazane (HMDS) 2-Deoxy-2-fluoro-3,5-di-O-benzoyl-α-D-arabinofuranosyl bromide (S.3; see Basic Protocol 1) Carbon tetrachloride (CCl4), anhydrous (Aldrich) Merck thin-layer chromatography (TLC) silica plates (Kieselgel 60 F-254; 0.2 mm thick) Dichloromethane, dry (see recipe) Saturated aqueous sodium bicarbonate Sodium sulfate (Na2SO4), anhydrous 29.7% (w/w) aqueous ammonia (Fisher) Methanol 5% (v/v) ethanol in chloroform Silica gel (230 to 400 mesh) Chloroform 3:1 (v/v) chloroform/ethanol Benzoic anhydride N,N-Dimethylformamide (DMF), anhydrous (Aldrich) Pyridine, dry (see recipe) p-Anisylchlorodiphenylmethane (monomethoxytrityl chloride or MMTr⋅Cl) 4-Dimethylaminopyridine (4-DMAP) 0% to 1% (v/v) gradient of methanol in chloroform 250-mL round-bottom flasks, oven dried Reflux condenser Oil bath, 120°C and 77°C Rotary evaporator equipped with a vacuum pump or water aspirator 3 × 20–cm chromatography column
Synthesis of Modified Nucleosides
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Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Condense N4-acetylcytosine with S.3 1. In an oven-dried 250-mL round-bottom flask equipped with a reflux condenser, stir bar, and nitrogen gas source, add 15 mL TMSCl to a suspension of 11 g (153.14 mmol) N4-acetylcytosine in 80 mL HMDS. 2. Reflux 24 hr in an oil bath at 120°C, cool mixture, and evaporate to dryness under reduced pressure using a rotary evaporator equipped with a vacuum pump or water aspirator. 3. Add 10 g (23.64 mmol) S.3 in 40 mL anhydrous CCl4 and reflux the resulting solution 4 days at 77°C. 4. Analyze the reaction by TLC (APPENDIX 3D) using Merck TLC silica plates. The starting material should be run alongside the reaction for comparison. The plates are developed using 9:1 (v/v) chloroform/methanol, and the bands are visualized by UV shadowing and dipping the plate in 10% (v/v) sulfuric acid in methanol followed by heating. Typically, the Rf value of the desired product N1⋅β-isomer is 0.6 (9:1 chloroform/methanol). The starting material, S.3, has an Rf value of 0.9 under these conditions.
5. Dilute reaction with 400 mL dry dichloromethane and wash it carefully with 500 mL saturated aqueous sodium bicarbonate solution. 6. Dry the dichloromethane layer over anhydrous Na2SO4, filter, and evaporate to dryness. No purification is needed at this step.
Deprotect 3′,5′-O-dibenzoyl araF-CAc 7. Treat 8.5 g crude compound with a mixture of 120 mL of 29.7% aqueous ammonia and 150 mL methanol for 48 hr at room temperature. This step removes O-benzoyl and N-acetyl groups. The Rf value for the desired product is 0.09 in 9:1 (v/v) chloroform/methanol and 0.17 in 3:1 (v/v) chloroform/ethanol.
8. Evaporate the resulting solution to dryness. Dissolve the crude product in a minimal amount of 5% ethanol in chloroform and place it carefully on top of a 3 × 20–cm silica gel chromatography column (see APPENDIX 3E for column chromatography). 9. Elute with 3:1 chloroform/ethanol and collect 50-mL fractions. Combine fractions containing pure product as determined by TLC. 10. Evaporate combined fractions to dryness in the rotary evaporator to yield the desired product as a white foam. A yield of 3.1 g (54% from S.3) is expected.
N4-Benzoylate araF-C 11. Dissolve 3.1 g (12.65 mmol) product and 3.43 g (15.16 mmol) benzoic anhydride in 20 mL anhydrous DMF and stir 35 hr to get araF-CBz. The Rf value of the desired product is 0.63 in 3:1 (v/v) chloroform/ethanol and 0.67 in 3:1 (v/v) chloroform/methanol. Synthesis of Protected 2′-Deoxy-2′-fluoroβ-D-arabinonucleosides
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5′-Tritylate araF-CBz 12. Remove DMF under reduced pressure and co-evaporate the residue with dry pyridine. Add the following and stir 24 hr at room temperature: 20 mL pyridine 4.0 g (12.95 mmol) MMTr⋅Cl 30 mg 4-DMAP (catalytic amount). The Rf value for the desired product (S.9a) is 0.50 in 9:1 (v/v) chloroform/methanol and 0.20 in 20:1 (v/v) chloroform/methanol.
13. Quench with 10 mL methanol, evaporate the resulting mixture, take up the residue in 150 mL chloroform, wash the organic layer with 150 mL saturated aqueous sodium bicarbonate, dry organic layer over anhydrous Na2SO4, and evaporate to dryness. 14. Dissolve the crude product in a minimal amount of chloroform and place it carefully on the top of a 3 × 20–cm silica gel chromatography column. 15. Elute with 0% to 1% methanol in chloroform and collect 50-mL fractions. Combine the fractions containing pure product as determined by TLC. 16. Evaporate combined fractions to dryness in the rotary evaporator to get the desired product as a yellow foam. 17. Check purity of the final product. N4-Benzoyl-1-[2-deoxy-2-fluoro-5-O-(4-methoxytrityl)-β-D-arabinofuranosyl]cytosine (S.9a): 4.6 g (58% from araF-C); TLC (20:1 [v/v] chloroform/methanol); 1H NMR (400 MHz, acetone-d6, tetramethylsilane as internal reference): 8.0 (1H, d, H-6), 7.6 to 7.2 (19H, m, trityl, Bz), 6.9 (1H, d, H-5), 6.3 (1H, dd, J1′,2′ = 3.6 Hz, J1′,F = 14 Hz, H-1′), 5.2 (1H, m, J2′,F = 49 Hz, H-2′), 4.5 (1H, m, H-3′), 4.3 (1H, m, H-4′), 3.8 (3H, s, CH3O), 3.5 (2H, m, H-5′ and H-5′′).
SYNTHESIS AND CHARACTERIZATION OF N6-BENZOYL-9-[2-DEOXY-2FLUORO-5-O-(4-METHOXYTRITYL)-β-D-ARABINOFURANOSYL]ADENINE
BASIC PROTOCOL 3
N6-Benzoyl-9-(2-deoxy-2-fluoro-3,5-di-O-benzoyl-β-D-arabinofuranosyl)adenine was prepared by direct condensation of N6-benzoyladenine with 2-deoxy-2-fluoro-3,5-di-Obenzoyl-α-D-arabinofuranosyl bromide (S.3; Figure 1.7.5; Watanabe et al., 1988). Flash chromatography and the removal of benzoyl groups with ammonia gave araF-A, which was successfully 5′-tritylated and N6-benzoylated in a one-pot procedure to give the title compound, S.9c. Materials 2-Deoxy-2-fluoro-3,5-di-O-benzoyl-α-D-arabinofuranosyl bromide (S.3; see Basic Protocol 1) Dichloromethane, dry (see recipe) N6-Benzoyladenine, anhydrous Activated molecular sieves (type 4A) Pyridine, dry (see recipe) Chloroform Silica gel (230 to 400 mesh) 7:3 (v/v) chloroform/dichloromethane Merck thin-layer chromatography (TLC) silica plates (Kieselgel 60 F-254; 0.2 mm thick) Ethanol 29.7% (w/w) aqueous ammonia (Fisher) p-Anisylchlorodiphenylmethane (monomethoxytrityl chloride or MMTr⋅Cl)
Synthesis of Modified Nucleosides
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Trimethylsilyl chloride (TMSCl) Benzoyl chloride Brine (saturated aqueous NaCl) Magnesium sulfate, anhydrous 33:1 (v/v) dichloromethane/methanol 250- and 500-mL round-bottom flasks, oven dried Reflux condenser Oil bath, 40° to 50°C Rotary evaporator equipped with vacuum pump 7 × 15–cm chromatography column Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Condense N6-benzoyladenine with S.3 1. In a 250-mL round bottom flask equipped with a reflux condenser, dissolve 7.5 g (17.7 mmol) S.3 in 150 mL dry dichloromethane. 2. Add 10.5 g (44 mmol) anhydrous N6-benzoyladenine and 21 g activated molecular sieves (type 4A). 3. Place the flask in an oil bath preheated to 40° to 50°C and reflux for 4 days. 4. Cool reaction mixture to room temperature, add 50 mL dry pyridine, and filter. Rinse molecular sieves with 50 mL pyridine and combine both fractions. 5. Evaporate filtrate to dryness in a rotary evaporator to yield a brown oil. 6. Dissolve the crude product in a minimal amount of chloroform. 7. Apply the resulting solution to a 7 × 15–cm chromatography column packed with 100g silica gel in chloroform. Column chromatography is performed by using a small amount of air pressure at a rate of ∼1 inch of solvent per minute (Still et al., 1978; APPENDIX 3E).
8. Elute the product with chloroform and then continue with 7:3 chloroform/dichloromethane. Collect 100-mL fractions and combine those that contain pure product as determined by TLC (APPENDIX 3D) using Merck TLC silica plates. The Rf value for the desired product is 0.75 in 9:1 (v/v) dichloromethane/methanol.
9. Evaporate combined fractions and dry overnight under a high vacuum to get a white foam. 10. Check the purity of the product. N6-Benzoyl-9-(2-deoxy-2-fluoro-3,5-di-O-benzoyl-β-D-arabinofuranosyl)adenine (S.8c): 6 g as a white foam, (58% from S.3); TLC (9:1 [v/v] dichloromethane/methanol); 1H NMR (270 MHz, DMSO-d6): 8.8 (s, H-2), 8.5 (d, JH8,F = 2 Hz, H-8), 8.1 to 7.3 (m, Bz), 6.8 (dd, J1′, 2′ = 4 Hz, J1′,F = 18 Hz, H-1′), 6.0 (ddd, J3′,-F = 19 Hz, H-3′), 5.8 (ddd, J1′ 2′ = 4 Hz, J2′,3′ = 2 Hz, J2′,F = 52 Hz, H-2′), 4.8 (m, H-4′), 4.7 (m, H-5′ and H-5′′); 19F NMR (270 MHz, DMSO-d6, no external reference was used) –197 (ddd); FAB-MS (NBA-matrix): 582 (M+H+). The purified product contains ∼5% of α-isomer, which is removed in steps 24 to 26. Synthesis of Protected 2′-Deoxy-2′-fluoroβ-D-arabinonucleosides
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Current Protocols in Nucleic Acid Chemistry
Deprotect N6-benzoyl-9-(2-deoxy-2-fluoro-3,5-di-O-benzoyl-β-D-arabinofuranosyl) adenine 11. In a 500-mL round-bottom flask equipped with magnetic stirrer, suspend 6 g (10.33 mmol) S.8c in 150 mL ethanol. 12. Add 150 mL of 29.7% aqueous ammonia and stir 2 days. Use TLC to determine when deprotection is complete. The Rf value for araF-A is 0.20 in 17:3 (v/v) dichloromethane/methanol.
13. Evaporate the resulting solution to dryness. No purification is needed after this step.
Tritylate and benzoylate 9-(2-deoxy-2-fluoro-β-D-arabinofuranosyl)adenine 14. Co-evaporate 9-(2-deoxy-2-fluoro-β-D-arabinofuranosyl)adenine (step 13) twice with 50 mL each dry pyridine. 15. Dissolve the residue in 50 mL dry pyridine. 16. Add 3.7 g (12 mmol) MMTr⋅Cl and stir the reaction mixture overnight at room temperature. Use TLC to ensure that the tritylation is complete. The Rf value for the desired product is 0.40 in 9:1 (v/v) dichloromethane/methanol. An additional portion of MMTr⋅Cl can be added if tritylation is not complete.
17. Add 6.4 mL (50 mmol) TMSCl and stir the resulting solution 30 min at room temperature. 18. Add 5.8 mL (50 mmol) benzoyl chloride and stir 2 hr. 19. Place the reaction flask into an ice bath to cool and add 10 mL water. Stir 5 min. 20. Add 20 mL of 29.7% aqueous ammonia and stir 30 min. 21. Carefully evaporate the reaction mixture to dryness. Avoid formation of foam, which makes evaporation difficult. 22. Redissolve the mixture in 200 mL chloroform, wash with 200 mL brine, dry over anhydrous magnesium sulfate, filter, and evaporate. 23. Dissolve the crude product in a minimal amount of dichloromethane. 24. Place it carefully on the top of a 7 × 15–cm silica gel chromatography column and elute with dichloromethane and then with 33:1 dichloromethane/methanol. Collect 100-mL fractions and combine those that contain pure product as determined by TLC. The Rf value for the desired product is 0.38 in 9:1 (v/v) chloroform/ethanol.
25. Combine fractions that contain pure product and evaporate and dry overnight at high vacuum to get a white foam. 26. Check the purity of the product. N6-Benzoyl-9-[2-deoxy-2-fluoro-5-O-(4-methoxytrityl)-β-D-arabinofuranosyl] adenine (S.9c): 3.8 g (as a white foam, 52% yield from S.8c); TLC (9:1 [v/v] chloroform/ethanol) 0.38; 1H NMR (500 MHz, acetone-d6) 10.0 (br s, N-H), 8.6 (s,H-2), 8.3 (d, JH8-F = 3 Hz, H-8), 8.1 to 6.9 (m, MMTr, Bz), 6.7 (dd, J1′,F = 18 Hz, J1′,2′ = 4 Hz, H-1′), 5.3 (ddd, J1′,H2′ = 4 Hz, J2′,F = 52 Hz, J2′,3′ = 4 Hz, H-2′), 4.7 (ddd, J2′,3′ = 4 Hz, J3′,F = 19 Hz, J3′,4′ = 3 Hz, H-3′), 4.2 (dd, J4′,5′,5′′ = 4 Hz, J3′,4′ = 3 Hz, H-4′), 3.8 (s, CH3O-), 3.5 to 3.4 (m, H-5′ and H-5′′); FAB-MS (NBA-matrix): 646 (M+H+).
Synthesis of Modified Nucleosides
1.7.13 Current Protocols in Nucleic Acid Chemistry
Supplement 10
BASIC PROTOCOL 4
SYNTHESIS AND CHARACTERIZATION OF 1-[2-DEOXY-2-FLUORO-5-O(4-METHOXYTRITYL)-β-D-ARABINOFURANOSYL]THYMINE 1-(Deoxy-2-fluoro-3,5-di-O-benzoyl-2-β-D-arabinofuranosyl)thymine was synthesized by direct coupling of silylated thymine and 2-deoxy-2-fluoro-3,5-di-O-benzoyl-α-Darabinofuranosyl bromide (S.3). Crystallization of the crude product gave pure 1-(2-deoxy-2-fluoro-3,5-di-O-benzoyl-β-D-arabinofuranosyl)thymine, which gave S.9b after deprotection and tritylation using standard procedures. Materials Thymine Ammonium sulfate Phosphorous pentoxide (P2O5) Acetonitrile, dry (see recipe) Hexamethyldisilazane (HMDS) 2-Deoxy-2-fluoro-3,5-di-O-benzoyl-α-D-arabinofuranosyl bromide (S.3; see Basic Protocol 1) Magnesium sulfate, anhydrous Carbon tetrachloride (CCl4), anhydrous (Aldrich) Dichloromethane, dry (see recipe) Ethanol Concentrated aqueous ammonia Merck thin-layer chromatography (TLC) silica plates (Kieselgel 60 F-254; 0.2 mm thick) Pyridine, dry (see recipe) p-Anisylchlorodiphenylmethane (monomethoxytrityl chloride or MMTr⋅Cl) Chloroform Brine (aqueous saturated NaCl) Silica gel (230 to 400 mesh) 19:1 (v/v) dichloromethane/methanol 250- and 500-mL round-bottom flasks, oven dried Oil bath, 100° Rotary evaporator attached to a vacuum pump 7 × 15–cm chromatography column chromatography (APPENDIX 3E) Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Silylate thymine 1. Dry 5.4 g (43 mmol) thymine and 540 mg ammonium sulfate in an oven-dried 250-mL round-bottom flask in a vacuum desiccator over P2O5 overnight. 2. Dissolve dry mixture in 160 mL dry acetonitrile and add 9 mL HMDS. 3. Put flask in a 100°C oil bath and allow reaction mixture to reflux for 4 hr. Once the reaction mixture becomes clear, the reaction is considered to be complete.
4. Let reaction cool to room temperature and evaporate excess solvents under reduced pressure.
Synthesis of Protected 2′-Deoxy-2′-fluoroβ-D-arabinonucleosides
Condense 2,4-bis-O-(trimethylsilyl)thymine with S.3 5. Dissolve 7g (16.5 mmol) S.3 in 70 mL dry CCl4. 6. Carefully transfer solution with a syringe to the flask containing silylated thymine (step 4).
1.7.14 Supplement 10
Current Protocols in Nucleic Acid Chemistry
7. Put flask in an oil bath and leave it to reflux for 4 days. The Rf value for the desired product, S.8b, is 0.49 in 9:1 (v/v) chloroform/methanol.
8. Cool down mixture to room temperature, dilute it with 500 mL dichloromethane, and wash with 1 L water. A large volume of water is necessary to get rid of salts formed during the coupling step.
9. Dry organic layer with anhydrous magnesium sulfate and concentrate under reduced pressure to obtain the crude product as a slightly brown solid. Usually 1-(2-deoxy-2-fluoro-3,5-di-O-benzoyl-2-β-D-arabinofuranosyl)thymine contains ∼2% to 3% of 1-(2-deoxy-2-fluoro-3,5-di-O-benzoyl-2-α-D-arabinofuranosyl)thymine, which cannot be removed by silica gel chromatography at this step. Crystallization (step 10), however, gives the analytically pure compound.
10. Dissolve the crude product in 40 mL dichloromethane and crystallize from 200 mL ethanol. 11. Check purity of the product. 1-(2-Deoxy-2-fluoro-3,5-di-O-benzoyl-2-β-D-arabinofuranosyl)thymine (S.8b) : 5. 8 g (75% yield from S.3); TLC (9:1 [v/v] chloroform/methanol) 0.49; 1H NMR (500 MHz, DMSO-d6): 11.5 (s, N-H), 8.0 to 7.4 (m, H-6 and Bz), 6.3 (dd, J1′,2′ = 4 Hz, J1′-F = 19 Hz, H-1′), 5.7 (ddd, J2′,3′ = 2 Hz, J3′,4′ = 4 Hz, J3′,F = 19 Hz, H-3′), 5.5 (ddd, J1′,2′ = 4 Hz, J2′,3′ = 2 Hz, J2′,F = 53 Hz, H-2′), 4.8 to 4.7 (m, H-5′ and H-5′′), 4.6 (m, H-4′), 1.6 (s, CH3-C5); FAB-MS (NBA-matrix): 468 [M+].
Deprotect 1-(2-deoxy-2-fluoro-3,5-di-O-benzoyl -β-D-arabinofuranosyl)thymine 12. In a 500-mL round bottom flask equipped with magnetic stirrer, suspend 5.8 g (12.38 mmol) S.8b in 150 mL ethanol. 13. Add 150 mL concentrated aqueous ammonia and stir the resulting mixture for 3 days. The Rf value for the desired product is 0.49 in 3:1 (v/v) chloroform/ethanol.
14. When deprotection is complete, as determined by TLC (APPENDIX 3D) using Merck TLC silica plates, evaporate solution to dryness. Tritylate 1-(2-deoxy-2-fluoro-β-D-arabinofuranosyl)thymine 15. Co-evaporate 1-(2-deoxy-2-fluoro-β-D-arabinofuranosyl)thymine (step 14) twice with 50 mL each dry pyridine and dissolve in 50 mL dry pyridine. 16. Add 4.4 g (14.23 mmol) MMTr⋅Cl and stir the reaction overnight. The Rf value for the desired product (S.9b) is 0.3 in 19:1 (v/v) chloroform/methanol.
17. Dilute mixture with 200 mL chloroform, wash with 200 mL brine, dry the organic layer over magnesium sulfate, filter, and evaporate to obtain the crude product. 18. Dissolve the crude product in a minimal amount of dichloromethane. 19. Load it onto a 7 × 15–cm chromatography column packed with 100 g silica gel (APPENDIX 3E). Elute with dichloromethane and then with 19:1 (v/v) dichloromethane/methanol. Collect 100-mL fractions and combine those that contain pure product as determined by TLC. The Rf value for the desired product is 0.3 in 19:1 (v/v) choloroform/methanol.
20. Combine fractions that contain pure product, evaporate, and dry overnight at high vacuum to obtain a white foam.
Synthesis of Modified Nucleosides
1.7.15 Current Protocols in Nucleic Acid Chemistry
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21. Check the purity of the product. 1-[2-Deoxy-2-fluoro-5-O-(4-methoxytrityl)-β-D-arabinofuranosyl]thymine (S.9b): 5.7 g (87%); TLC (9:1 [v/v] chloroform/methanol) 0.3; 1H NMR (500 MHz, DMSO-d6) 11.5 (br s, NH), 7.4 to 6.9 (m, H-6 and MMTr), 6.1 (dd, J1′-2′ = 4.5 Hz, J1′-F = 16 Hz, H-1′), 5.0 (ddd, J1′-2′ = 4 Hz , J2′,3′ = 2.5 Hz, J2′-F = 53 Hz, H-2′), 4.3 (ddd, J2′-3′ = 2.5 Hz, J3′-4′ = 4.5 Hz JH3′-F = 19 Hz, H-3′), 3.9 (m, H-4′), 3.7 (s, CH3O-), 3.3 (m, H-5′ and H-5′), 1.6 (s, CH3-C5); FAB-MS (NBA-matrix): 532 [M+].
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For work up of reaction mixtures and chromatography, use reagent grade solvents. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Acetonitrile Dry acetonitrile by refluxing over calcium hydride (Aldrich) or phosphorus pentoxide (P2O5; Fisher) for 10 hr followed by distillation under inert atmosphere. Dichloromethane Dry dichloromethane by refluxing over calcium hydride (Aldrich) for 24 hr followed by distillation under inert atmosphere. Pyridine Dry pyridine by refluxing over calcium hydride (Aldrich) for 8 hr followed by distillation under inert atmosphere. COMMENTARY Background Information
Synthesis of Protected 2′-Deoxy-2′-fluoroβ-D-arabinonucleosides
The interests of medicinal and organic chemists towards the construction of compounds that contain fluorine is derived from the relative stability of the carbon-fluorine bond, both chemically and metabolically, and from the strong electronegative character of fluorine, which alters the electronic properties of the molecule. In fact, many fluorinated organic compounds exhibit interesting biophysical properties. This has been shown to be the case for anti-inflammatory steroids (Herdewijn et al., 1989), collagen helices (Holmgren et al., 1 99 9) , n ucleo sides (Ma et al., 1997; Pankiewicz, 2000), and nucleic acids (Damha et al., 1998). For example, collagen chains in which fluorine atoms replace the hydroxyl groups form triple helices with extraordinary stability (Eberhardt et al., 1996). A number of highly selective nucleoside antiviral and antileukemic agents based on 2-deoxy-2fluoroarabinose have been prepared over the past 20 years (for reviews, see Burchenal et al., 1983; Pankiewicz, 2000). These compounds also serve as building blocks for the synthesis o f 2′-deoxy-2′-fluoroarabinonucleic acids (2′F-ANAs). It has recently been shown that 2′F-ANA forms a duplex with RNA that is more stable than a DNA:RNA duplex of identical
sequence (Damha et al., 1998; Wilds and Damha, 2000). The authors’ laboratory has also shown that 2′F-ANA:RNA heteroduplexes, like the natural DNA:RNA heteroduplex, are substrates of RNase H, an enzyme implicated in the mechanism of action of antisense oligonucleotides (Damha et al., 1998). This unit describes methods for the synthesis of protected 2′-deoxy-2′-fluoroarabinonucleosides. These methods, based on experience and facilities at hand, allow straightforward synthesis of suitable monomers that can be phosphitylated and used as building blocks for the synthesis of antisense oligonucleotides (Lok et al., 2002). The authors have optimized some of the key steps reported in the literature (Tann et al., 1985; Howell et al., 1988; Pankiewicz et al., 1992; Scharer and Verdine, 1995; Maruyama et al., 1999; Wilds and Damha, 2000), particularly those involved in the synthesis of araF-G (Tennila et al., 2000). A critical step is the fluorination of the ribose sugars, which may lead to loss of water (dehydration) via an elimination reaction. Elimination is virtually avoided by using DAST (or MAST) as the fluorinating reagent instead of HF, SF4, and HF-amine reagents (Middleton, 1975).
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Current Protocols in Nucleic Acid Chemistry
plished with a masked base, namely 2,6-dichloropurine, followed by its transformation into guanine in subsequent steps (Ma et al., 1997; Tennila et al., 2000; Figure 1.7.3). All steps work well, except for the N-glycosylation reaction, which proceeds to give the desired N9β-anomer in 35% to 40% yield (optimized). The authors are currently investigating coupling of S.3 to various purine derivatives in order to simplify and increase the yield of araF-G.
The most convenient method for the synthesis of araF-U, araF-T, and araF-C involves directly coupling 3,5-di-O-benzoyl-2-deoxy-2fluoro-α-D-arabinofuranosyl bromide with silylated pyrimidines (e.g., Howell et al., 1988; Wilds and Damha, 2000). This produces primarily the desired β-anomer with <5% of the α-stereoisomer, which is removed by chromatography and/or crystallization. This is in contrast to the results obtained during the synthesis of 2′-deoxynucleosides (lacking the 2′-fluorine atom) to afford anomeric mixtures that generally favor the α-anomer. This may be accounted for by the electronegative nature of the 2′-F atom, which effectively prevents the ionization of the C1′-leaving group to produce an SN1 oxonium ion intermediate. Therefore, the glycosylation reaction mainly proceeds via the energetically preferred SN2 pathway (Howell et al., 1988; Figure 1.7.6). Originally, the authors synthesized araF-A (Pankiewicz et al., 1992) and araF-G (Chu et al., 1989) starting from the ribonucleosides (rA and rG). Coupling the purines to the fluorinated arabinose sugar S.3, however, largely improves yields and minimizes the number of steps. A presilylation step for the synthesis of araF-A is not required. Synthesis of araF-G is accom-
Critical Parameters and Troubleshooting During the synthesis of araF-G (S.6), two important considerations have to be taken into account. First, the refluxing time for the synthesis of S.4 from S.2 should not exceed 1 hour; otherwise the yield of the desired product (S.4) will be low. Second, purification of the 2,6diaminopurine derivative is very important in order to get good yields of araF-G during the adenosine deaminase step. In the synthesis of araF-C and araF-T, both the coupling time and the equivalents of the heterocyclic bases used in the N-glycosylation reactions are very important factors for obtaining high yields of nucleosides. During synthesis of araF-A and
O
OTMS
NH
N BzO
N
O F Br
BzO
OTMS
SN2
BzO
O F
N
O
BzO
3 β-anomer
α-bromo sugar
SN1 OTMS N
+
BzO
N
O F
OTMS α- + β -anomers
BzO destabilized by 2-F
Figure 1.7.6 Coupling of 2,4-bis(trimethylsilyl)thymine with 2-fluoroarabinose sugar S.3. Under appropriate conditions (e.g., CCl4 solvent, reflux), the reaction proceeds primarily via an SN2 pathway, affording the β and α anomers in a ratio of 98:2. Bz, benzoyl; TMS, trimethylsilane.
Synthesis of Modified Nucleosides
1.7.17 Current Protocols in Nucleic Acid Chemistry
Supplement 10
araF-C, it is better to use N6-benzoyladenine (without silylation) and N4-acetylcytosine (with silylation), respectively, for coupling with S.3.
Anticipated Results The black sheep of the family of nucleosides is guanosine, and araF-G is no exception. The synthesis of araF-G requires more steps than for other araF nucleosides. Nevertheless, these procedures yield araF-G in high purity and moderate yields. Figure 1.7.4 shows the 1H NMR spectrum of araF-G. The purity of the isolated nucleoside (and all others reported here) is usually >98%. The additional splitting of the sugar proton signals is indicative of the presence of fluorine (spin = 1⁄2). The β-configuration of the N-glycosidic bond for all nucleosides was established by two-dimensional nuclear Overhauser effect (2D-NOE) NMR experiments (Wilds and Damha, 2000). A characteristic feature of the araF-A and araF-G spectra (1H NMR) is the doublet corresponding to the H-8 proton. The splitting of the H-8 signal is due to long-range 1H-19F coupling (likely through space via H8...2′-F bonding) that arises only for the βanomers (Wilds and Damha, 2000).
Time Considerations Each of the araF nucleosides presented in this unit can be prepared in 6 to 7 days except for araF-G, for which close to 2 weeks are required.
Literature Cited Burchenal, J.H., Leyland-Jones, B., Watanabe, B., Klein, R., Lopez, C., and Fox, J.J. 1983. Experimental and clinical studies on 2′-fluoroarabinosyl pyrimidines and purine-like C-nucleosides. In Nucleosides, Nucleotides, and Their Biological Applications, Proceedings 5th International Round Table, pp. 47-65. International Society for Nucleosides, Nucleotides, and Nucleic Acids, Montpellier, France. Chu, C.K., Matulic-Adamic, J., Huang, J.-T., Chou, T.-C., Burchenal, J.H., Fox, J.J., and Watanabe, K.A. 1989. Synthesis of some 9-(2-deoxy-2fluoro-β-D-arabinofuranosyl)-9H-purines and their biological activities. Chem. Pharm. Bull. 37:336-339.
Synthesis of Protected 2′-Deoxy-2′-fluoroβ-D-arabinonucleosides
Damha, M.J., Wilds, C.J., Novonha, A., Brunker, I., Borkow, G., Arion, D., and Parniak, M.A. 1998. Hybrids of RNA and arabinonucleic acids (ANA and 2′F-ANA) are substrates of ribonuclease H. J. Am. Chem. Soc. 120:12976-12977.
Eberhardt, E.S., Panasik, N. Jr., and Raines, R.T. 1996. Inductive effects on the energetics of prolyl peptide bond isomerization: Implications for collagen folding and stability. J. Am. Chem. Soc. 118:12261-12266. Herdewijn, P., VanAerschot, A., and Kerremans, L. 1989. Synthesis of nucleosides fluorinated in the sugar moiety. The application of diethylaminosulfur trifluoride to the synthesis of fluorinated nucleosides. Nucleosides Nucleotides 8:65-96. Holmgren, S.K., Bretscher, L.E., Taylor, K.M., and Raines, R.T. 1999. A hyperstable collagen mimic. Chem. Biol. 6:63-70. Howell, H.G., Brodfuehrer, P.R., Brundidge, S.P., Benigni, D.A., and Sapino, C. Jr. 1988. Antiviral nucleosides. A stereospecific, total synthesis of 2′-fluoro-2′-deoxy-β-D-arabinofuranosyl nucleosides. J. Org. Chem. 53:85-88. Kierzek, R. 1985. The synthesis of 5′-O-dimethoxytrityl-N-acetyl-2′-deoxynucleosides. Improved “transient protection” approach. Nucleosides Nucleotides 4:641-649. Ma, T., Lin, J.-S., Newton, M.G., Cheng, Y.-C., and Chu, C.K. 1997. Synthesis and anti hepatitis B virus activity of 9-(2-deoxy-2-fluoro-β-Larabinofuranosyl)purine nucleosides. J. Med. Chem. 40:2750-2754. Maruyama, T., Takamatsu, S., Kozai, S., Satoh, Y., and Izawa, K. 1999. Synthesis of 9-(2-deoxy-2fluoro-β-D-arabinofuranosyl)adenine bearing a selectively removable protecting group. Chem. Pharm. Bull. 47:966-970. Middleton, W.J. 1975. New fluorinating reagents. Dialkylaminosulfur fluorides. J. Org. Chem. 40:574-578. Lok, C.-N., Viazovkina, E., Min, K.-L., Nagy, E., Wilds, C.J., Damha, M.J., and Parniak, M.A. 2002. Potent gene-specific inhibitory properties of mixed-backbone antisense oligonucleotides comprised of 2′-deoxy-2′-fluoro-D-arabinose and 2′-deoxyribose nucleotides. Biochemistry 41:3457-3467. Pankiewicz, K.W. 2000. Fluorinated nucleosides. Carbohydr. Res. 327:87-105. Pankiewicz, K.W., Krzeminski, J., Cizewski, L.A., Ren, W.-Y., and Watanabe, K.A. 1992. Synthesis of 9-(2-deoxy-2-fluoro-β-D-arabinofuranosyl)adenine and hypoxanthine. An effect of C3′endo to C2′-endo conformational shift on the reaction course of 2′-hydroxyl group with DAST. J. Org. Chem. 57:553-559. Scharer, O.D. and Verdine, G.L. 1995. A designed inhibitor of base-excision DNA repair. J. Am. Chem. Soc. 117:10781-10782. Still, W.C., Kahn, M., and Mitra, A. 1978. Rapid chromatographic technique for preparative separation with moderate resolution. J. Org. Chem. 43:2923-2925.
1.7.18 Supplement 10
Current Protocols in Nucleic Acid Chemistry
Tann, C.H., Brodfuehrer, P.R., Brundidge, S.P., Sapino, C. Jr., and Howell, H.G. 1985. Fluorocarbohydrate in synthesis. An efficient synthesis of 1-(2-deoxy-2-fluoro-β-D-arabinofuranosyl)-5iodouracil (β-FIAU) and 1-(2-deoxy-2-fluoroβ-D-arabinofuranosyl)thymine (β-FMAU). J. Org. Chem. 50:3644-3647. Tennila, T., Azhayev, E., Vepsalainen, J., Laatikainen, R., Azhayev, A., and Mikhailopulo, I.A. 2000. Oligonucleotides containing 9-(2-deoxy2-fluoro-β-D-arabinofuranosyl)-adenine and guanine: Synthesis, hybridization and antisense properties. Nucleosides Nucleotides Nucleic Acids 19:1861-1884.
Key References Herdewijn et al., 1989. See above. This article reviews the application of DAST to the synthesis of fluorinated nucleosides. Howell et al., 1988. See above. This article describes key connections and synthetic strategies of araF-nucleosides that form the basis of the procedures outlined in these protocols. Pankiewicz, 2000. See above. This review article focuses on the synthesis of sugar fluorinated nucleosides.
Wilds, C.J. and Damha M.J. 2000. 2′-Deoxy-2′fluoro-β-D-arabinonucleosides and oligonucleotides (2′F-ANA): Synthesis and physicochemical studies. Nucl. Acids Res. 28:36253635.
Tann et al., 1985. See above.
Watanabe, K.A., Chu, C.K., and Fox, J.J. June, 1988. 2-Fluoro-arabinofuranosyl purine nucleosides. U.S. patent 47,551,221.
This article provides procedures for the synthesis of araF-nucleosides and 2′F-ANA.
Wower, J., Hixson, S.S., Sylvers, L.A., Xing, Y., and Zimmermann, R.A. 1994. Synthesis of 2,6-diazido-9-(β-D-ribofuranosyl)purine 3′,5′-bisphosphate: Incorporation into transfer RNA and photochemical labelling of Escherichia coli ribosomes. Bioconjug. Chem. 5:158-161.
This article also describes synthetic strategies of araF-nucleosides. Wilds and Damha, 2000. See above.
Contributed by Mohamed I. Elzagheid, Ekaterina Viazovkina, and Masad J. Damha McGill University Montreal, Canada
Synthesis of Modified Nucleosides
1.7.19 Current Protocols in Nucleic Acid Chemistry
Supplement 10
Synthesis, Characterization, and Application of Substituted Pyrazolopyrimidine Nucleosides
UNIT 1.8
This unit describes, in detail, the preparation of 3-aminopropyl-substituted pyrazolo[3,4d]pyrimidine analogs of the purines deoxyadenosine (dA) and deoxyguanosine (dG). Phosphoramidite reagents of these so-called aminopropyl-PPA and -PPG nucleosides (AP-PPA and AP-PPG, respectively) allow introduction of amino linkers into internal positions of synthetic DNA strands. Synthesis of suitably protected AP-PPA and AP-PPG phosphoramidites (S.1 and S.2) are described in the first two methods (see Basic Protocols 1 and 2). The stepwise alkynylation, hydrogenation, selective protection, and phosphoramidite synthesis is similar for both the PPA and PPG analogs. To demonstrate the application of these reagents, a simple DNA strand is synthesized and conjugated to a lipophilic activated ester (dabcyl-SE) to form a stable amide linkage (see Basic Protocol 3). Utility of this chemistry for preparing internally modified DNA conjugates is discussed. CAUTION: All reactions used for preparation of the phosphoramidites must be performed in a well-ventilated fume hood. These procedures should be performed only by personnel with experience in organic synthesis. In particular, operations involving hydrogenation under pressure are hazardous and require training beyond that described here. Standard precautions to prevent excessive exposure to toxic chemicals and solvents should be followed. All reactions should first be performed on a small scale. PREPARATION OF PROTECTED AMINOPROPYL-PPG PHOSPHORAMIDITE Synthesis of a suitably protected phosphoramidite reagent for incorporating AP-PPG (S.1) into DNA strands is shown in Figure 1.8.1. The starting material is a 3-iodopyrazolopyrimidine nucleoside. Preparation of this iodinated nucleoside is described elsewhere (Seela and Zulauf, 1998; Seela and Becher, 1999). The chemical reactions described here give good yields, and purification of the intermediates is straightforward. Materials 6-Amino-1-(2-deoxy-β-D-erythro-pentofuranosyl)-3-iodopyrazolo[3,4-d]pyrimidin4(5H)-one (Seela and Zulauf, 1998) CuI Tetrakis[triphenylphosphine]palladium[0] (Aldrich) Argon Anhydrous dimethylformamide (DMF), deoxygenated (i.e., argon purged) Anhydrous triethylamine N-Trifluoroacetylpropargylamine (Cruickshank and Stockwell, 1988) Chloroform 230- to 400-mesh (40- to 63-µm) silica gel (EM Sciences) Methanol Ethyl acetate Diethyl ether 20% (w/v) palladium hydroxide on carbon, preactivated (see recipe) 4 M triethylammonium formate buffer, pH 6.5 (see recipe) Hydrogen gas Celite (1.5-cm pad) Contributed by Robert O. Dempcy, Mikhail A. Podyminogin, and Michael W. Reed Current Protocols in Nucleic Acid Chemistry (2002) 1.8.1-1.8.15 Copyright © 2002 by John Wiley & Sons, Inc.
BASIC PROTOCOL 1
Synthesis of Modified Nucleosides
1.8.1 Supplement 11
N,N-Dimethylformamide dimethylacetal (Aldrich) Xylenes Anhydrous pyridine 4,4′-Dimethoxytrityl chloride 5% (w/v) sodium bicarbonate Sodium sulfate Anhydrous methylene chloride N,N-Diisopropylethylamine 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite Hexane 100-mL round-bottom flask Rotary evaporator equipped with a vacuum pump 5 × 35–cm chromatography columns Silica gel 60 F254 aluminum-backed TLC plates (EM Science) 254-nm UV lamp Oil pump (<1 mmHg)
H CF3CO N O
H NH
N N HO
NHCOCF3
O
I
N
NH2
O
N HO
OH
H
H
CF3CO N
CF3CO N
N
MeO N
O
N
H
HO
N 5
NH
N
OMe
N N
O OH
hydrogen (35 psi)
NH
N
NH2
3
20% Pd(OH)2 on carbon 4 M triethylammonium formate, pH 6.5 methanol
O
N
O
OH
HO
NH
N
triethylamine Cul Pd[Ph3P]4 DMF
N
NH2
O OH
DMF
4
DMTr⋅Cl pyridine H
H
CF3CO N
CF3CO N
O NH
N N DMTrO
O OH
N
N N
6
NH
N
i -Pr2NEt, CH2Cl2
N DMTrO
i -Pr2N NC
Pyrazolopyrimidine Nucleosides Substituted
O
i -Pr2NP(Cl)CH2CH2CN N
O
P
O
N N
1
O
Figure 1.8.1 Preparation of AP-PPG phosphoramidite (S.1). Abbreviations: DMF, N,N-dimethylformamide; DMTr⋅Cl, N,N′-dimethoxytrityl chloride.
1.8.2 Supplement 11
Current Protocols in Nucleic Acid Chemistry
500-mL Parr hydrogenation bottle 250-mL separatory funnel Additional reagents and equipment for column chromatography (APPENDIX 3E) and thin-layer chromatography (TLC; APPENDIX 3D) NOTE: 1H NMR spectra were obtained at 300 MHz on a Varian VXR-300 spectrometer. Two-dimensional COSY and NOE experiments assisted in the assignment of proton resonances. Elemental analyses were performed by Quantitative Technologies. NOTE: Like most phosphoramidite reagents, AP-PPA and -PPG are sensitive to hydrolysis and oxidation and must be handled with care after isolation. Desiccated storage at −20°C is recommended. Performance in DNA synthesis depends on good quality reagents; only reagent-grade chemicals should be used to synthesize these phosphoramidites. Characterization of the compounds as described here helps ensure purity. Prepare S.3 1. Weigh a mixture of 6.00 g (15.26 mmol) 6-amino-1-(2-deoxy-β-D-erythro-pentofuranosyl)-3-iodopyrazolo[3,4-d]pyrimidin-4(5H)-one, 297 mg (1.56 mmol) CuI, and 0.91 g (0.79 mmol) tetrakis[triphenylphosphine]palladium[0] into a 100-mL roundbottom flask and purge with argon. 2. Add 30 mL deoxygenated (argon-purged) anhydrous DMF, 3.14 mL anhydrous triethylamine, and 4.29 g (28.41 mmol) N-trifluoroacetylpropargylamine. Stir the reaction solution 40 hr under argon. 3. Evaporate off the DMF on a rotary evaporator equipped with a vacuum pump, and triturate the residual oil in 150 mL chloroform. Vacuum filter the crude solid that forms (S.3) over Whatman no. 1 filter paper, rinse with chloroform, and dry using a rotary evaporator with a vacuum pump. 4. Dissolve the solid in a minimum volume of anhydrous DMF, adsorb onto a minimal amount of 230- to 400-mesh silica gel (so that gel is free flowing), and evaporate. Load the dry mixture onto a 5 × 35–cm chromatography column (5 × 25–cm bed volume) and elute with 10% (v/v) methanol in ethyl acetate (see APPENDIX 3E for column chromatography). 5. Monitor fractions by TLC (APPENDIX 3D) on silica gel 60 F254 aluminum-backed TLC plates using a mobile phase of 20% (v/v) methanol in ethyl acetate and visualize with a 254-nm UV lamp. Pool the product fractions (Rf = 0.59) and evaporate. 6. Precipitate by dissolving in a minimum volume of ethyl acetate and adding to ∼4 volume diethyl ether. Dry overnight at high vacuum on an oil pump (<1 mmHg). The resulting product, 6-amino-1-(2-deoxy-β-D-erythro-pentofuranosyl)-3-(3-trifluoroacetamido-propyn-1-yl)-pyrazolo[3,4-d]pyrimidin-4(5H)-one (S.3), is obtained in 63% yield (4.0 g). 1H NMR (DMSO-d6): δ 10.79 (1H, s, N5-H), 10.16 (1H, t, J = 5.2 Hz, trifluoroacetamido N-H), 6.77 (2H, br s, 6-amino), 6.28 (1H, t, J = 6.3 Hz, 1′-H), 5.23 (1H, d, J = 4.1 Hz, 3′-OH), 4.72 (1H, t, J = 5.1 Hz, 5′-OH), 4.32 (3H, m, -CONH-CH2- and 3′-H), 3.75 (1H, m, 4′-H), 3.50-3.29 (2H, 2 × m, 5′-Hs), 2.65 and 2.15 (2H, 2 × m, 2′-Hs). Anal. calcd. For C15H15F3N6O5.0.74H2O: C, 41.93; H, 3.87; N, 19.56. Found: C, 42.33; H, 3.64; N, 19.13.
Synthesis of Modified Nucleosides
1.8.3 Current Protocols in Nucleic Acid Chemistry
Supplement 11
Prepare S.4 7. In a 500-mL Parr hydrogenation bottle, prepare a solution of 1.0 g (2.40 mmol) S.3 in 100 mL methanol containing 0.12 g of preactivated 20% palladium hydroxide on carbon. Always work with palladium catalysts under an inert atmosphere as oxygen and moisture will reduce yield.
8. Add 2.0 mL of 4 M triethylammonium formate buffer, pH 6.5. Shake the mixture 18 hr under 1810 mmHg (35 psi) hydrogen gas. CAUTION: Hydrogen gas is flammable. Use extreme caution to avoid ignition sources. This step should be supervised by an experienced chemist. During some runs it is necessary to add additional catalyst for complete reduction (as determined by TLC; 20% methanol in ethyl acetate). If necessary, add another 0.06 g catalyst and repeat reaction.
9. Filter the mixture through a 1.5-cm pad of Celite and evaporate the filtrate. 10. Crystallize the residual oil from water and dry overnight at high vacuum (using an oil pump). The resulting product, 6-amino-1-(2-deoxy-β-D-erythro-pentofuranosyl)-3-(3-trifluoroacetamidopropyl) pyrazolo[3,4-d]pyrimidin-4(5H)-one, is obtained in 0.79 g (78%) yield. TLC (20% methanol in ethyl acetate): Rf = 0.52. 1H NMR (DMSO-d6): δ 10.59 (1H, s, N5-H), 9.47 (1H, br t, trifluoroacetimido N-H), 6.64 (2H, br s, 6-amino), 6.27 (1H, t, J = 6.3 Hz, 1′-H), 5.18 (1H, d, J = 4.4 Hz, 3′-OH), 4.75 (1H, t, J = 5.9 Hz, 5′-OH), 4.36 (1H, m, 3′-H), 3.75 (1H, m, 4′-H), 3.48 and 3.61 (2H, 2 × m, 5′-Hs), 3.22, 2.68 and 1.87 (6H, 3 × m, propyl methylene protons), 2.68 and 2.12 (2H, 2 × m, 2′-Hs). Anal. calcd. for C15H19F3N6O5.0.90H2O: C, 41.27; H, 4.80; N, 19.25. Found: C, 41.57; H, 4.50; N, 19.11.
Prepare S.5 11. Dissolve 0.80 g (1.98 mmol) S.4 in 5.0 mL anhydrous DMF. Add 3.1 mL N,N-dimethylformamide dimethylacetal and stir 2.0 hr under argon. 12. Evaporate off solvent, and then evaporate the residue twice from 20 mL xylenes. Place under high vacuum. 13. Triturate the amorphous solid that forms (S.5) in 30 mL diethyl ether, collect by vacuum filtration over no. 1 Whatman paper, and dry overnight at high vacuum. The resulting product, 1-(2-deoxy-β-D-erythro-pentofuranosyl)-6-[(dimethy-lamino) methylidenamino]-3-(3-trifluoroacetamidopropyl)pyrazolo[3,4-d]pyrimidin-4(5H)-one, is obtained in 773-mg (82%) yield. TLC (20% methanol in ethyl acetate): Rf = 0.47. 1H NMR (DMSO-d6): δ 11.22 (1H, br s, N5-H), 9.47 (1H, t, J = 5.5 Hz, trifluoroacetamido N-H), 8.67 (1H, s, N = CH-N), 6.42 (1H, t, J = 6.5 Hz, 1′-H), 5.22 (1H, d, J = 4.3 Hz, 3′-OH), 4.75 (1H, t, J = 6.1 Hz, 5′-OH), 4.40 (1H, m, 3′-H), 3.77 (1H, m, 4′-H), 3.50 and 3.38 (2H, 2 × s, 5′-Hs), 3.18 and 3.05 (6H, 2 × s, N,N-dimethyl Hs), 3.22, 2.72 and 1.89 (6H, 3 × m, propyl methylene Hs), 2.72 and 2.15 (2H, 2 × m, 2′-Hs). Anal. calcd. for C18H24F3N7O5.0.40H2O: C, 44.80; H, 5.18; N, 20.32. Found: C, 45.02; H, 4.96; N, 19.94.
Prepare S.6 14. Prepare a solution of 723 mg (1.52 mmol) S.5 in 9.0 mL anhydrous pyridine. 15. Add 0.61 g (1.80 mmol) 4,4′-dimethoxytrityl chloride and stir the reaction solution 3.0 hr under argon. Pyrazolopyrimidine Nucleosides Substituted
16. Pour the reaction mixture into a 250-mL separatory funnel containing 100 mL of 5% sodium bicarbonate.
1.8.4 Supplement 11
Current Protocols in Nucleic Acid Chemistry
17. Extract the aqueous solution twice with 200 mL ethyl acetate each, and dry the combined extracts over sodium sulfate. 18. Vacuum filter the solution over no. 1 Whatman paper and evaporate the filtrate. 19. Purify the crude product by silica gel chromatography using a 5 × 25–cm bed volume of 230- to 400-mesh silica gel. Elute with a solvent gradient of 0% to 5% (v/v) methanol in ethyl acetate containing 2% triethylamine. 20. Monitor fractions by TLC (see step 5) using a mobile phase of 5% (v/v) methanol/ethyl acetate. Pool the product fractions (Rf = 0.39), evaporate, and dry overnight at high vacuum to give S.6 as an amorphous solid. The resulting product, 1-[5-O-(4,4′-dimethoxytrityl)-2-deoxy-β-D-erythro-pentofuranosyl]-6-[(dimethylamino)methylidenamino]-3-(3-trifluoroacetamidopropyl)pyrazolo[3,4-d] pyrimidin-4(5H)-one, is obtained in 724-mg (61%) yield. 1H NMR (DMSO-d6): δ 11.27 (1H, s, N5-H), 9.43 (1H, t, J = 5.3 Hz, trifluoroacetamido N-H), 8.71 (1H, s, N=CH-N), 7.32, 7.17 and 6.76 (13H, 3 × m, aromatic), 6.45 (1H, t, J = 6.3 Hz, 1′-H), 5.26 (1H, d, J = 5.3 Hz, 3′-OH), 4.45 (1H, m, 3′-H), 3.90 (1H, m, 4′-H), 3.70 (6H, s, OMe Hs), 3.18 and 3.05 (10H, 2 × s, N,N-dimethyl, 5′-Hs and CONH-CH2), 2.62 and 1.65 (4H, 2 × m, methylene Hs), 2.62 and 2.20 (2H, 2 × m, 2′-Hs). Anal. calcd. for C39H42F3N7O7.0.30H2O: C, 59.81; H, 5.48; N, 12.52. Found: C, 59.80; H, 5.39; N, 12.63.
Prepare S.1 21. Prepare a solution of 700 mg (0.900 mmol) S.6 in 22 mL anhydrous methylene chloride containing 0.47 mL N,N-diisopropylethylamine. 22. Add 0.34 mL (1.52 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite and stir 30 min under argon at 25°C. 23. Treat the solution with 3.0 mL methanol and dilute with 200 mL ethyl acetate. 24. Wash the solution with 100 mL of 5% sodium bicarbonate and dry over sodium sulfate. 25. Vacuum filter the solution over no. 1 Whatman paper and evaporate the filtrate. 26. Purify the crude product by silica gel chromatography using a 5 × 25–cm bed volume of 230- to 400-mesh silica gel. Elute with 2% (v/v) triethylamine in ethyl acetate. 27. Monitor fractions by TLC (step 5) using ethyl acetate. Pool the product fractions (Rf = 0.38) and evaporate. 28. Precipitate the residue from 1:2 (v/v) diethyl ether/hexane, evaporate, and dry overnight at high vacuum to give S.1 as an amorphous solid. The resulting product is 1-[5-O-(4,4′-dimethoxytrityl)-2-deoxy-β-D-erythro-pentofuranosyl]-6-[(dimethylamino)methylidenamino]-3-(3-trifluoroacetamidopropyl)pyrazolo[3,4-d] pyrimidin-4(5H)-one 3′-[(2-cyanoethyl) N,N-diisopropylphosphoramidite], is obtained in 583-mg (66%) yield. 31P NMR (DMSO-d6, reference to 85% phosphoric acid): δ 145.50 and 144.72.
Synthesis of Modified Nucleosides
1.8.5 Current Protocols in Nucleic Acid Chemistry
Supplement 11
H CF3CO N NH2
I
NH2
N
N N HO
NHCOCF3 H
O
N HO
OH
7
20% Pd(OH)2 on carbon 4 M triethylammonium formate, pH 6.5 methanol H
hydrogen (35 psi)
H
N
CF3CO N
CF3CO N
N
NH2
N
N N
N
O
OH
HO
N
N
triethylamine Cul Pd[Ph3P]4 DMF
N
MeO
N
N N
O OH
N
N
OMe HO
N
O OH
DMF
DMTr⋅Cl pyridine H
N N
N
i -Pr2NP(Cl)CH2CH2CN
8
N
N
i -Pr2NEt, CH2Cl2
N DMTrO
O OH
N
N
CF3CO N
N N
DMTrO
H
N
CF3CO N
i -Pr2N NC
N
O
P
O
2
O
Figure 1.8.2 Preparation of AP-PPA phosphoramidite (S.2). Abbreviations: DMF, N,N-dimethylformamide; DMTr⋅Cl, N,N′-dimethoxytrityl chloride.
BASIC PROTOCOL 2
Pyrazolopyrimidine Nucleosides Substituted
PREPARATION OF PROTECTED AMINOPROPYL-PPA PHOSPHORAMIDITE To synthesize the AP-PPA phosphoramidite (S.2; Fig. 1.8.2), the method described for the AP-PPG phosphoramidite (see Basic Protocol 1) is utilized with the following exceptions. Some intermediates were not rigorously purified for the protocol described here. To ensure success, a careful step-wise approach on a small scale is advised. The protocol described here allowed successful preparation of the desired AP-PPA phosphoramidite (S.2) and shows the advanced user which intermediates can be taken on in crude form. The reactions described in Figure 1.8.2 were followed by analytical reversedphase HPLC using a 150 × 4.6–mm column, a gradient of acetonitrile (solvent B) in 0.1 M triethylammonium acetate, pH 7.5 (solvent A), and a flow rate of 1 mL/min. Each reaction was monitored by UV detection at 260 nm, and worked up when starting material was consumed. Elution gradients and retention times are noted for each reaction.
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Materials 4-Amino-1-(2-deoxy-β-D-erythro-pentofuranosyl)-3-iodo-1H-pyrazolo[3,4-d]pyrim idine (Seela and Zulauf, 1998) Celite (1.5–cm pad) N,N-Dimethylacetamide dimethylacetal (TCI America) Anhydrous triethylamine Anhydrous dimethylformamide (DMF) Xylenes 4,4′-Dimethoxytrityl chloride Anhydrous pyridine 230- to 400-mesh (40- to 63-µm) silica gel (EM Sciences) Ethyl acetate Methylene chloride Methanol 5 × 35–cm chromatography column Additional reagents and equipment for synthesizing AP-PPG (see Basic Protocol 1) Prepare S.7 1. Alkynylate 1.0 g of 4-amino-1-(2-deoxy-β-D-erythro-pentofuranosyl)-3-iodo-1Hpyrazolo[3,4-d]pyrimidine by the general procedure described for the synthesis of S.3 (see Basic Protocol 1, steps 1 to 6). The resulting product is 4-amino-1-(2-deoxy-β-D-erythro-pentofuranosyl)-3-(3-trifluoroacetamido-propyn-1-yl)-1H-pyrazolo[3,4-d]pyrimidine. Analytical HPLC using 0% to 100% solvent B over 20 min showed complete conversion of 3-iodo-substituted PPA (6 min) to S.7 (7.2 min) after 24 hr.
Prepare S.8 2. Hydrogenate 1.0 g (2.5 mmol) S.7 by the general procedure described for S.4 (see Basic Protocol 1, steps 7 to 10), but reduce the reaction time (step 8) to 1 hr. Analytical HPLC using 0% to 30% solvent B over 20 min showed complete hydrogenation of S.7 (16.5 min) to the desired product (14 min).
3. Filter the reaction mixture through a 1.5–cm Celite pad and evaporate the filtrate to dryness under high vacuum. 4. Dissolve the crude product in 20 mL anhydrous DMF and then add 1.46 mL (10 mmol) N,N-dimethylacetamide dimethylacetal and 2.0 mL triethylamine. Stir for 24 hr. Analytical HPLC using 0% to 60% solvent B over 20 min showed complete conversion of the starting amine (9 min) to the desired dimethylacetal product (11.5 min) after 24 hr. After 5 hr, an intermediate compound (12.3 min) was detected that eventually converted to product.
5. Evaporate the completed reaction solution to dryness, and then evaporate the residue twice from 20 mL xylenes. 6. React the crude product with 0.93 g of 4,4′-dimethoxytrityl chloride in 20 mL anhydrous pyridine for 16 hr. Evaporate the reaction mixture to dryness. Analytical HPLC using 0% to 100% solvent B over 20 min showed complete conversion of the alcohol (9 min) to the desired DMTr product S.8 (16 min).
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7. Prewash a 5 × 25–cm column (bed volume) of 230- to 400-mesh silica gel with 1 column volume of 1:1 (v/v) triethylamine/ethyl acetate. Rinse with 500 mL ethyl acetate. Load the crude product onto the silica gel column in a minimum volume of 9:1 (v/v) ethyl acetate/methylene chloride (see APPENDIX 3E for column chromatography). Rinsing with ethyl acetate removes excess triethylamine and leaves the silica non-acidic. Chromatography can also be performed as in Basic Protocol 1, step 19.
8. Elute the product (S.8) with a gradient of 0% to 10% (v/v) methanol in ethyl acetate and evaporate to give an amorphous solid. The resulting product, 1-[5-O-(4,4′-dimethoxytrityl)-2-deoxy-β-D-erythro-pentofuranosyl]-4-[1-(dimethylamino)ethylidene]amino-3-(3-trifluoroacetamidopropyl)pyrazolo [3,4-d]pyrimidine, is obtained in 90% yield (1.75 g).
Prepare S.2 9. Phosphitylate 1.75 g (2.25 mmol) compound S.8 by the general procedure described for S.1 (see Basic Protocol 1, steps 21 to 28). The resulting product, 1-[5-O-(4,4′-dimethoxytrityl)-2-deoxy-β-D-erythro-pentofuranosyl]-4-[1-(dimethylamino)ethylidene]amino-3-(3-trifluoroacetimidopropyl)pyrazolo[3,4-d] pyrimidine-3′-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite], is obtained in 1.86 g (85%) yield. 31P NMR (DMSO-d6, reference to 85% phosphoric acid): δ 147.43 and 146.76. Analytical HPLC using 30% to 100% solvent B over 10 min followed by 100% solvent B for 10 min showed complete conversion of S.8 (16.5 min) to the desired phosphoramidite S.2 (12 min) after 3 hr. BASIC PROTOCOL 3
Pyrazolopyrimidine Nucleosides Substituted
SYNTHESIS OF DNA CONJUGATES USING AP-PPA AND AP-PPG PHOSPHORAMIDITES This protocol describes the preparation of DNA conjugates from the AP-PPA or AP-PPG modified oligonucleotides as described in Figure 1.8.3. DNA synthesis using these reagents is straightforward for someone trained in DNA synthesis, and is thus not described in detail. The phosphoramidite reagents S.1 and S.2 give good yields of AP-PPG- or AP-PPA-containing DNA strands using standard methods (for reference on standard methods, see UNIT 3.3 and APPENDIX 3C). The introduced aminopropyl linker groups allow reaction of the DNA strands with activated esters of conjugate groups. To demonstrate utility, the N-hydroxysuccinimide ester of a fluorescent quencher molecule (dabcyl-SE) is added to a 9-mer DNA strand containing an internal AP-PPA linker (S.9). As illustrated in Figure 1.8.3, the DNA is converted to the DMSO-soluble triethylammonium salt form (steps 1 to 4) and reacted with dabcyl-SE (step 5 to 8). The crude DNA conjugate is precipitated from DMSO and purified by reversed-phase HPLC (RP-HPLC, steps 9 to 12) to give the desired dabcyl-DNA conjugate (S.10). The protocol described here uses 0.2 µmol of an oligodeoxynucleotide from a 1.0-µmol scale DNA synthesis, but can be modified for synthesis of nucleic acid conjugates at any scale. Materials AP-PPA-modified 9-mer oligonucleotide from a 1-µmol-scale synthesis, trityl-on (e.g., Epoch Biosciences) 3 M sodium acetate Butanol Ethanol Solvent A: 0.1 M triethylammonium acetate, pH 7.5 Solvent B: acetonitrile Anhydrous dimethylsulfoxide (DMSO; e.g., Aldrich Sure-seal; store over calcium hydride) Anhydrous triethylamine (Fluka; store over calcium hydride)
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N-Hydroxysuccinimidyl ester of dabcyl (dabcyl-SE; Molecular Probes) 2% (w/v) sodium perchlorate in acetone Acetone Centrifugal evaporator (i.e., Savant Speedvac) with high vacuum source (<1 mmHg) High-performance liquid chromatograph (HPLC) with: 4.6 × 250–mm C18 analytical column and appropriate guard column (Varian Dynamax) UV-Vis or photodiode array detector 14-mL polypropylene tubes UV-Vis spectrophotometer Additional reagents and equipment for RP-HPLC and detritylation (UNIT 10.5) Prepare starting oligonucleotide 1. Purify the starting AP-PPA-modified 9-mer oligonucleotide (trityl-on) by RP-HPLC and then detritylate with 80% acetic acid (UNIT 10.5). It is important to obtain the starting oligonucleotide (S.9) from a qualified DNA synthesis source. AP-PPA- and AP-PPG-containing oligonucleotides can be purchased from Epoch Biosciences (http://www.epochbio.com).
+
+
H3 N
H3 N
NH2 N
N N dT4-O
N
N
O
dT4-O
Salt Exchange
P
O-dT4
O−
Na+
RP-HPLC with triethylammonium acetate
N
O O
O
9 (Na salt)
P
O-dT4
O−
+NHEt 3
9 (TEA salt)
Acylation
O N Me2N
N
N
O O
NH2
dabcyl-SE anhydrous DMSO
N
N
NH2
H
N
N N dT4-O
O
N
Purification precipitation from DMSO RP-HPLC purification
crude DNA conjugate
O O
P
O-dT4
O−
+NHEt 3
10 (TEA salt)
Figure 1.8.3 Synthesis of dabcyl conjugate at internal AP-PPA linker position in a 9-mer oligonucleotide.
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2. Precipitate with 0.5 mL of 3 M sodium acetate and 4 mL butanol. Wash with ethanol and dry in vacuo using a Speedvac evaporator. The resulting pellet is primarily the sodium salt of the DNA (S.9).
Perform salt exchange 3. Dissolve ∼200 nmol trityl-off S.9 in 500 µL water. Purify by RP-HPLC using a 4.6 × 250–nm C18 analytical column and guard column, with a gradient of 0% to 30% (v/v) solvent B over 20 min at a flow rate of 2 mL/min. Collect the desired fraction (∼15-min retention time) in a 14-mL polypropylene tube. 4. Dry in a centrifugal evaporator (SpeedVac) to dryness to give the pure detritylated oligodeoxynucleotide as a TEA salt. The appropriate amount of DNA can be aliquoted at this stage by dissolving the trityl-off pellet in water, measuring the A260, and drying the appropriate volume in a 1.5-mL microcentrifuge tube.
Perform conjugation 5. Add 30 µL anhydrous DMSO to 200 nmol dried TEA salt of S.9 in a 1.5-mL microcentrifuge tube. Vortex and examine closely to make sure the pellet is dissolved. If the pellet is not completely dissolved, gradually add more DMSO (up to 0.5 mL final). More dilute solutions slow the acylation reaction. Sonication or heating can often be used to help dissolve stubborn pellets.
oligonucleotide
AU
dabcyl conjugate
5
10 min
15
20
AU
0
200
Pyrazolopyrimidine Nucleosides Substituted
250
300
350
400 nm
450
500
550
600
Figure 1.8.4 Reversed-phase HPLC chromatograms and UV-Vis spectra of AP-PPA-modified oligonucleotides (S.9; dashed lines) and dabcyl conjugate (S.10; solid lines). The large difference in retention time simplifies isolation and analysis of the lipophilic conjugate.
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6. Add 7 µL (50 µmol) anhydrous triethylamine. At higher than 5%, the triethylamine is insoluble in DMSO. Triethylamine can be added at up to 10% of the total final volume of DMSO without hurting the conjugation reaction.
7. Dissolve 1.1 mg dabcyl-SE in 100 µL DMSO in a second 1.5-mL microcentrifuge tube. Vortex until homogeneous. CAUTION: DMSO solutions of acylating agents can burn the skin. Wear nitrile (not latex) rubber gloves and handle tubes carefully. Before opening lids of the tubes, spin down liquids in a microcentrifuge. Open cautiously to prevent aerosols. Dispose of excess reagents with waste organic solvents.
8. Add 33 µL (366 µg, 1 µmol) dabcyl-SE solution to the solution of S.9 (step 6). Vortex and incubate 24 hr at room temperature. Monitor the reaction by RP-HPLC as described in step 3 (see Fig. 1.8.4). The reaction rate can be increased by incubating at 37°C. Analytical HPLC using this system can be extremely valuable for identifying the starting oligonucleotide (10.7 min) and desired conjugate peaks (16.7 min). If the acylating agent has a chromophore with an A260 that is unique from that of the oligonucleotide, a photodiode array detector is useful. Hydrolyzed acylating agents can elute in the HPLC collection window and complicate identification of the desired conjugate. In this example with dabcyl-SE, an unreactive peak (20% by A260 integration) remained with a retention time of 10.8 min.
Purify conjugate 9. After the reaction is deemed complete, precipitate the oligonucleotide products with 1.2 mL of 2% sodium perchlorate in acetone. Microcentrifuge the mixture 1 min at maximum speed. If reaction volume is >0.15 mL, precipitate the products by adding to 10 vol of 2% sodium perchlorate in acetone contained in a larger tube.
10. Wash the pellet with 0.5 mL acetone. Microcentrifuge again at maximum speed and dry 30 min in a SpeedVac evaporator. 11. Dissolve the crude DNA conjugate in ∼1 mL solvent A and purify by RP-HPLC as described in step 3. Evaporate the collected fraction to dryness in a Speedvac evaporator. 12. Dissolve the dried conjugate in 0.1 mL water and characterize by RP-HPLC and UV-Vis spectroscopy (Fig. 1.8.4). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. Unless otherwise noted, reagent-grade chemicals should be used. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Palladium hydroxide, 20% (w/v), on carbon Available from Aldrich (also known as Pearlman’s catalyst). Before use, preactivate by gently warming a suspension of the palladium catalyst in 5 mL methanol containing 3 drops of formic acid for 5 min. Use immediately. Always work with palladium catalysts under an inert atmosphere.
Triethylammonium formate buffer, 4 M, pH 6.5 Make 4 M formic acid solution. Add one equivalent of triethylamine. Adjust pH to 6.5. Store tightly capped for up to 1 year at 4°C.
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COMMENTARY Background Information
Pyrazolopyrimidine Nucleosides Substituted
Conjugation of ligands to the nucleobases of synthetic oligonucleotides allows preparation of modified nucleic acids for molecular biology and potential therapeutic applications. Specialty phosphoramidite reagents have been developed that allow preparation of modified DNA on automated DNA synthesizers (Beaucage and Iyer, 1993), but these ligands must survive the harsh ammonia deprotection conditio ns. Preparation of specialty phosphoramidites requires multistep organic synthesis, and these resources are not readily available to most DNA synthesis facilities. Linker phosphoramidites are available which contain functional groups that can be modified with reactive conjugate groups (Goodchild, 1990). Aminoalkyl linkers are usually incorporated into synthetic DNA at terminal positions of synthetic probes or at internal positions via non-nucleotide (abasic) linker groups. These amino linkers can be acylated with various “active esters” to introduce a wide variety of conjugate groups to give stable amide bonds. Internal abasic linkers can disrupt DNA duplex formation or prevent enzyme recognition; therefore, aminoalkyl-modified nucleotide phosphoramidites have been developed. Conjugate groups are well tolerated in the major groove of DNA duplexes and have been shown in some cases to stabilize dsDNA. Suitably protected C5-aminoalkyl-substituted pyrimidines are available as β-cyanoethyl phosphoramidite reagents (Glen Research; http://www.glenres.com). DNA sequences of interest can be synthesized using these reagents to introduce linker sites at various internal dC or dT positions in synthetic DNA probes. Substitution at C5 of the pyrimidine heterocycle places the attached conjugate groups in what will become the major groove in B-form DNA after hybridization with a complementary DNA strand. Unlike the pyrimidine analogs, aminoalkyl-modified purine analogs are not commercially available as phosphoramidite reagents. Deazapurines, modified at the 7 position, also place conjugate groups in the major groove, and are well tolerated in DNA duplexes. Phosphoramidite reagents and NTP analogs of the 7-aminoalkyl deazapurines have been reported (Seela et al., 1996). The 3-modified pyrazolo[3,4-d]pyrimidine analogs of adenosine and guanine (PPA and PPG) are isosteric with the 7-modified deazapurine heterocycles. An early application
of the 3-substituted PPA ring system in DNA probes was introduction of a biotin-labeled PPA nucleoside via the corresponding 5′triphosphate (Petrie et al., 1991). Multiple biotin modifications in DNA probes were tolerated with little decrease in Tm versus natural dA. Advances in the synthesis of 3-substituted PPA and PPG nucleosides allowed their incorporation into DNA probes via phosphoramidite chemistry (Seela and Becher, 1999; Seela and Zulauf, 1999). 3-Alkylamine-modified pyrazolopyrimidines were not prepared, but halogen and alkyne modifications at the 3 position were found to increase stability of DNA duplexes. The 3-aminopropyl-substituted pyrazolo[3,4-d]pyrimidine analogs of adenine (PPA) and guanine (PPG) described here allow postsynthetic modification of DNA sequences with a variety of conjugate groups. The authors briefly described the AP-PPA and AP-PPG nucleoside phosphoramidites (S.1 and S.2) and used these linkers to incorporate the fluorescent quencher molecule TAMRA into probes containing fluorogenic minor groove binders (Kutyavin et al., 2000). Synthesis of these phosphoramidites is described in detail above (see Basic Protocols 1 and 2). The 3-iodo-substituted PPA and PPG nucleosides are particularly valuable intermediates since they are readily alkynylated via Pd-catalyzed cross-coupling reactions. The authors have successfully used the published synthetic procedures for these key intermediates. Trifluoroacetylpropargylamine is a convenient, easily prepared source of the required aminoalkyl linker arm. The authors used the published synthetic procedure for this common intermediate. Alkynylation using trifluoroacetylpropargylamine has been reported for preparation of the C5-substituted dT phosphoramidite (Cruickshank and Stockwell, 1988). The TFA-protected amine survives the organic synthesis steps, yet is readily hydrolyzed during ammonia deprotection after DNA synthesis. Pd-catalyzed coupling to the 3-iodoPPA or -PPG intermediates generally proceeds in good yield, and is easily followed by TLC or analytical HPLC. Use of the aminoalkyne intermediates for synthesis of the corresponding PPA or PPG phosphoramidites is feasible and may be advantageous for some applications. The authors prepared the aminoalkyl linkers since the aliphatic chain is more flexible, and presumably allows attached conjugate groups
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to find low-energy conformations that are less disruptive to DNA duplex formation. Sy nthesis of the aminoalkyl phosphoramidites (S.1 and S.2) from the alkynecoupled products uses well-established chemistry. Hydrogenation of the alkynes to give the aliphatic linkers proceeds in close-to-quantitative yield. Dimethyl formamidine or acetamidine protection of the exocyclic amino groups is readily accomplished by reaction with the corresponding dimethoxyketal. Tritylation of the 5′-hydroxy group uses standard conditions. As noted in the shortened AP-PPA protocol, the crude products from hydrogenation and acetamidine protection are suitably pure for direct use in the tritylation reaction. The tritylated products are easily isolated by silica gel chromatography and converted to the desired phosphoramidites using standard methods. DNA synthesis using S.1 and S.2 uses conventional phosphoramidite chemistry. The TFA and amidine protecting groups are nonbulky, and coupling yields are close to quantitative when introduced at internal positions in synthetic DNA sequences. To demonstrate the use of these linkers, a 9-mer oligonucleotide containing AP-PPA (S.9) was prepared and purified by reversed-phase HPLC. Basic Protocol 3 describes conjugation of a hydrophobic ligand (the NHS ester of dabcyl) to this internally modified 9-mer in anhydrous DMSO solution. Dabcyl-labeled DNA has applications in fluorogenic probes (Tyagi et al., 1998). The authors have used variations of Basic Protocol 3 for synthesis of hundreds of DNA conjugates. The simple method has been described elsewhere (Milesi et al., 1999), and is especially useful for preparation of the internally modified AP-PPA and AP-PPG conjugates described here. The authors have conjugated DNA alkylating agents (Podyminogin et al., 1996), lipophilic aryltin compounds (Karamychev et al., 1997), and sensitive fluorescent tags (Kutyavin et al., 2000) to aminoalkyl-modified nucleobases in DNA probes using the protocols described here.
Critical Parameters Synthesis of the phosphoramidites S.1 and S.2 from the 3-iodo-PPA and -PPG nucleosides is straightforward. Purification and chromatography of the intermediates at each step will ensure success. Likewise, DNA synthesis using S.1 and S.2 uses standard methods (UNIT 3.3 and APPENDIX 3C). The conjugation chemistry can be trickier, and will be the focus of the remainder of this section.
Active ester reagents such as NHS esters are commonly used for aqueous conjugation of ligands to the amino groups of biomolecules such as proteins or synthetic DNA. As a result, many desirable conjugate groups are available as active esters. These acylating agents generally have improved reactivity at nucleophilic aminoalkyl groups that can be introduced into DNA strands. Acylation of aminoalkyl linkers at internal positions in synthetic DNA strands can be difficult. Steric hindrance slows the acylation reaction, and hydrolysis of the acylating agent becomes a major side reaction. Standard aqueous conditions can often be used successfully for attaching hydrophilic conjugate groups to AP-PPA- and AP-PPG-modified oligonucleotides. For example, the authors previously reported addition of a TAMRA-NHS ester to fluorogenic DNA probes using an aqueous buffer system. However, the limited solubility or aqueous stability of some acylating agents presents a problem, especially for internal linker positions. Triethylammonium salts of oligonucleotides are quite soluble in anhydrous DMSO. This is the key to success with the conjugation chemistry presented here. Although oligonucleotides dried in vacuo always contain complexed water, minimizing the water content in the reaction mixture slows hydrolysis of the acylating agents. Nucleophilicity of the alkylamine group is also enhanced in DMSO, and this speeds the desired reaction with the activated ester. Use of the NHS ester of dabcyl (dabcyl-SE) provides an excellent example of the advantages of the DMSO conjugation chemistry. Standard aqueous methods are not feasible since organic solutions of dabcyl-SE precipitate upon addition to aqueous buffers. The TEA salt of the AP-PPA 9-mer (S.9) and dabcyl-SE are both readily soluble in DMSO. Excess triethylamine is added and the reaction proceeds at room temperature. The hydrophobic oligonucleotide conjugate is easily detected by reversed-phase HPLC and the course of the reaction can be followed. HPLC purification before and after the acylation reaction gives great confidence that the desired conjugate has been isolated.
Troubleshooting To ensure that the phosphoramidites perform as expected in DNA synthesis, the authors recommend preparation of a 9-mer probe such as that presented in Basic Protocol 3. Conjugation of a model ligand can give further confi-
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dence that the reagent is in good shape. The conjugation chemistry presented here usually allows isolation of the desired product. However, each DNA probe conjugate has unique properties, and problems associated with aggregation of lipophilic compounds or structural complexes associated with certain sequences can always be a problem. DNA conjugates are complex biomolecules with inherent sequencespecific properties that can be difficult to predict. Long sequences and strands with significant self-complementarity or homopurine runs can lead to unusual aggregation problems, especially with lipophilic conjugate groups. The quality of DNA synthesis and the quality of the activated ester reagent are important factors in successful conjugation chemistry. The key to success with the conjugation chemistry is homogeneity of the DMSO reaction mixture. After reversed-phase HPLC purification of the AP-PPA- or AP-PPG-containing oligonucleotide, it is important that it be dried well in vacuo. Residual triethylammonium acetate or water can adversely affect the desired acylation reaction.
Anticipated Results
Pyrazolopyrimidine Nucleosides Substituted
With good-quality AP-PPA or AP-PPG amidites, DNA synthesis is routine. After tritylon purification by RP-HPLC and sodium-toTEA salt exchange by trityl-off RP-HPLC, a homogeneous product is obtained. For example, the analytical RP-HPLC in Figure 1.8.4 shows a single peak for the starting AP-PPAmodified 9-mer (S.9, TEA salt). Conjugation chemistry using lipophilic activated esters allows the extent of the reaction to be easily tracked by RP-HPLC. The shift in retention time from ∼12 min for the starting AP-PPAcontaining oligo to ∼17 min for the desired dabcyl conjugate is especially dramatic for the example shown in Figure 1.8.4. The yield of DNA conjugates using the anhydrous DMSO method of Basic Protocol 3 is typically 30% to 90%. As shown in Figure 1.8.4, RP-HPLC retention times and UV-Vis spectra are characteristic for oligonucleotide conjugates that are lipophilic or contain unique chromophores. Performance in DNA hybridization–based assays depends on sequence-specific binding with complementary sequences. UV melting studies (UNIT 7.3) can be used to measure the Tm of the AP-PPA- or AP-PPGmodified DNA conjugates. Mass spectral analysis using MALDI-TOF (UNIT 10.1) or elec-
trospray ionization (UNIT 10.2) methods is another approach that can be used to verify the presence of the desired conjugate group. Other assays are ultimately used to detect the presence of conjugate groups in DNA probes. Conjugate groups with specific physical or chemical properties are introduced to enhance performance in hybridization-based diagnostic or therapeutic assays, and these assays are the final screen for oligonucleotide conjugate quality.
Time Considerations Synthesis of the AP-PPA or AP-PPG phosphoramidites takes 2 to 3 days per reaction and purification, depending on the indicated reaction times. Time can be saved by skipping some purification steps as indicated in the preparation of S.2, but this is not advised for the initial synthetic attempts. The conjugation chemistry is not labor intensive, but the HPLC analytical and purification steps can slow the work. Drying in vacuo is generally carried out overnight, and there are several drying steps. Experienced chemists with dedicated instruments can shortcut many of these steps. For example, precipitation of DNA from reversed-phase HPLC buffer has been described (Milesi et al., 1999).
Literature Cited Beaucage, S.L. and Iyer, R.P. 1993. The synthesis of modified oligonucleotides by the phosphoramidite approach and their applications. Tetrahedron 49:6123-6194. Cruickshank, K.A. and Stockwell, D.L. 1988. Oligonucleotide labeling: A concise synthesis of a modified thymidine phosphoramidite. Tetrahedron Lett. 29:5221-5224. Goodchild, J. 1990. Conjugates of oligonucleotides and modified oligonucleotides: A review of their synthesis and properties. Bioconjugate Chem. 1:165-187. Karamychev, V.N., Panyutin, I.G., Reed, M.W., and Neumann, R.D. 1997. Effect of radionuclide linker structure on DNA cleavage by 125I-labeled oligonucleotides. Antisense Nucl. Acid Drug Dev. 7:549-557. Kutyavin, I.V., Afonina, I.A., Mills, A., Gorn, V.V., Lukhtanov, E.A., Belousov, E.S., Singer, M.J., Walburger, D.K., Lokhov, S.G., Gall, A.A., Dempcy, R., Reed, M.W., Meyer, R.B.J., and Hedgpeth, J. 2000. 3′-Minor groove binderDNA probes increase sequence specificity at PCR extension temperatures. Nucl. Acids Res. 28:655-661. Milesi, D., Kutyavin, I.V., Lukhtanov, E.A., Gorn, V.V., and Reed, M.W. 1999. Synthesis of oligonucleotides in anhydrous dimethyl sulfoxide. Methods Enzymol. 313:164-173.
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Petrie, C.R., Adams, A.D., Stamm, M., Van Ness, J., Watanabe, S.M., and Meyer, R.B.J. 1991. A novel biotinylated adenylate analogue derived from pyrazolo[3,4-d]pyrimidine for labeling DNA probes. Bioconjugate Chem. 2:441-446.
Seela, F. and Zulauf, M. 1999. Synthesis of oligonucleotides containing pyrazolo[3,4-d]pyrimidines: The influence of 7-substituted 8-aza-7deazaadenines on the duplex structure and stability. J. Chem. Soc. Perkin Trans. 1 (4):479-488.
Podyminogin, M.A., Meyer, R.B.J., and Gamper, H.B. 1996. RecA-catalyzed, sequence-specific alkylation of DNA by cross-linking oligonucleotides. Effects of length and nonhomologous base substitutions. Biochemistry 35:7267-7274.
Seela, F., Ramzaeva, N., and Becher, G. 1996. 7Deazapurine DNA: Oligonucleotides containing 7-substituted 7-deaza-2′-deoxyguanosine and 8aza-7-deaza-2′-deoxyguanosine. Collect. Czech. Chem. Commun. 61:258-261.
Seela, F. and Becher, G. 1998. Synthesis of 7-halogenated 8-aza-7-deaza-2′-deoxyguanosines and related pyrazolo[3,4-d]pyrimidine 2′-deoxyribonucleotides. Synthesis (9):207-214.
Tyagi, S., Bratu, D.P., and Kramer, F.R. 1998. Multicolor molecular beacons for allele discrimination. Nature Biotech. 16:49-53.
Seela, F. and Becher, G. 1999. Oligonucleotides containing pyrazolo[3,4-d]pyrimidines: The influence of 7-substituted 8-aza-7-deaza-2′-deoxyguanosines on the duplex structure and stability. Helv. Chim. Acta 82:1640-1655. Seela, F. and Zulauf, M. 1998. Synthesis of 7-alkynylated 8-aza-7-deaza-2′-deoxyadenosines via the Pd-catalysed cross-coupling reaction. J. Chem. Soc. Perkin Trans. 1 (19):3233-3240.
Contributed by Robert O. Dempcy, Mikhail A. Podyminogin, and Michael W. Reed Epoch Biosciences Bothell, Washington
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Synthesis of 1,5-Anhydrohexitol Building Blocks for Oligonucleotide Synthesis
UNIT 1.9
Hexitol nucleic acids (HNA) represent a new oligomeric structure able to hybridize as well with DNA and RNA as with itself, and in a sequence-specific manner. In addition, HNA seems to be superior to its DNA and RNA analogs as an antisense construct. This is supported by a study demonstrating that HNA forms highly selective and exceptionally stable duplexes with RNA (Hendrix et al., 1997a) and that these HNA:RNA duplexes are stable towards nuclease degradation (Hendrix et al., 1997b). Furthermore, a high potential for antiviral activities has been reported (Verheggen et al., 1995). Hexitol nucleic acids are made up of phosphorylated 1,5-anhydro-D-arabino-2,3-dideoxyhexitol building blocks with a base moiety positioned in the 2 position. According to the Westheimer model, the base moiety of the hexitol nucleosides is axially oriented, avoiding the sterically unfavorable 1,3-diaxial repulsions (De Winter et al., 1998). This unit describes in detail the preparation of 1,5-anhydrohexitol (see Basic Protocol 1 and Fig. 1.9.1) and the 1,5-anhydrohexitol building blocks for oligonucleotide synthesis (hG, hA, hC, hT; see Basic Protocols 2 to 5 and Figs. 1.9.2 to 1.9.5, respectively). NOTE: Carry out reactions with anhydrous solvents and dried glassware (i.e., 2 hr at 70°C). NOTE: All chemicals are commercially available (e.g., ACROS, Fluka, Aldrich). All starting materials can be synthesized but are also available commercially. The sugar intermediate S.5 is now also commercially available. All reactions can be carried out with standard laboratory equipment and glassware (e.g., round-bottom and Erlenmeyer flasks, consensers, dropping funnels, separatory funnels, desiccators, ultrasonic baths, oil baths, and magnetic stirrers). PREPARATION OF 1,5-ANHYDRO-4,6-O-BENZYLIDINE-3-DEOXY-DGLUCITOL
BASIC PROTOCOL 1
In this protocol, synthesis of the sugar building block starting from tetra-O-acetyl-α-Dbromoglucose is given, as outlined in Figure 1.9.1 (Verheggen et al., 1993). The starting sugar, tetra-O-acetyl-α-D-bromoglucose, can be used to make both tolylsulfonyl and toluoyl intermediates (S.4a and S.4b, respectively). The latter is used to make S.5 (described below), which is then used to make the hG, hT, and hC nucleoside monomers (see Basic Protocols 2, 4, and 5, respectively). The former is used directly to make the hA monomer (see Basic Protocol 3). The chemical reactions described give good yields, and purification of the intermediates is straightforward. Materials 2,3,4,6-Tetra-O-acetyl-α-D-bromoglucose Diethyl ether: reflux overnight on sodium (Na, FeCl2, Et2O) and distill Azoisobutyronitrile [2,2′-azobis(2-methylpropionitrile); AIBN] Tri-n-butyltin hydride Precoated silica gel TLC plates (Alugram Sil G/UV254) Dichloromethane Anisaldehyde/sulfuric acid spray (UNIT 1.3) Potassium fluoride dihydrate Sodium sulfate Silica gel (0.060 to 0.200 nm) Methanol Contributed by Irene M. Lagoja, Arnaud Marchand, Arthur Van Aerschot, and Piet Herdewijn Current Protocols in Nucleic Acid Chemistry (2003) 1.9.1-1.9.22 Copyright © 2003 by John Wiley & Sons, Inc.
Synthesis of Modified Nucleosides
1.9.1 Supplement 14
AcO
HO
AcO O
OAc
Br
OAc
Bu3SnH, Et2O
OAc
KF/H2O
O
NaOMe
1
OH
OAc
O O
Bu2SnO benzene
O
O
OH
H O
TsCl or TolCl resp. dioxane
OR
PhCHO ZnCl2
O
Ph OH
H
O
OH
OAc
OAc
Ph
OH
OH
2
3a R = Tos 3b R = Tol CSCl2, DMAP, 2,4-Cl2C6H3OH, CH2Cl2 Bu3SnH, AIBN, toluene
O
Ph
Ph O
NaOMe
H
O
OR
OH
O H
O O
O
4a R = Tos 4b R = Tol 5 R=H
O
5
OH
Ph
O
5
NaOMe
Figure 1.9.1 Preparation of 1,5-anhydro-4,6-O-benzylidine-3-deoxy-D-glucitol (S.5). Abbreviations: AIBN, azoisobutyronitrile; Bu2SnO, dibutyltinoxide; Bu3SnH, tributyltinhydride; DMAP, 4-(dimethylamino)pyridine; NaOMe, sodium methoxide; PhCHO, benzaldehyde; TolCl, toluoyl chloride; TsCl, tosyl chloride.
0.1 N sodium methoxide, freshly prepared from sodium and dry methanol Acetic acid (glacial not necessary) Toluene Zinc chloride (dry fresh before use) Benzaldehyde Ethyl acetate n-Hexane Dibutyltinoxide Benzene Dioxane (reflux overnight on lithium aluminum hydride and distill) p-Toluenesulfonyl chloride (for S.3a) or p-toluoyl chloride (for S.3b) Delite Celite 4-(Dimethylamino)pyridine (DMAP) Dry ice/isopropanol Thiophosgene 2,4-Dichlorophenol 1 M potassium dihydrogenphosphate solution, pH 5 Nitrogen gas Synthesis of 1,5-Anhydrohexitol Building Blocks
Oil bath and magnetic stirrer Rotary evaporator equipped with a vacuum pump and cooling trap 5 × 35–, 6 × 50–, and 5 × 20–cm chromatography columns
1.9.2 Supplement 14
Current Protocols in Nucleic Acid Chemistry
2 × 18–cm test tubes 1-L round-bottom flask with rubber stopper Dropping funnel Glass funnel Dean-Stark condenser UV lamp, 254 nm Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E) NOTE: The 1H and 13C NMR spectra given as examples were determined with a JEOL FX 90Q spectrometer or 400 MHz Bruker AMX with tetramethylsilane as internal standard. Electron-impact mass spectra (EIMS) and chemical-ionization mass spectra (CIMS) were obtained using a KRATOS Concept 1H mass spectrometer. Abbreviations: s, singlet; d, doublet; dd, double doublet; t, triplet; br s, broad signal; m, multiplet; ddd, double doublet of doublet; and dm, double multiplet. Prepare S.1 1. Dissolve 25.03 g (60.9 mmol) of 2,3,4,6-tetra-O-acetyl-α-D-bromoglucose in 375 mL diethyl ether. 2. Add 0.99 g (6.1 mmol) of AIBN and 25 mL (93.1 mmol) of tri-n-butyltin hydride. 3. Stir the mixture 1 hr at 35°C and then at room temperature until conversion is complete (2 to 12 hr). Monitor by TLC (APPENDIX 3D) by spotting the reaction mixture (100 µL) between two spots of starting material (1 mg diluted to 200 µL) as a reference. Use precoated silica gel plates and develop with dichloromethane. Visualize under a UV lamp (254 nm) and then spray the plate with anisaldehyde/sulfuric acid spray and dry at 150°C (also see Critical Parameters). 4. Dissolve 8.21 g potassium fluoride dihydrate in 40 mL water, add to the suspension, and stir 15 min. 5. Filter off the precipitated tri-n-butyltin hydride and wash the organic retentate (on top of the funnel) with 120 mL water. Set aside the aqueous layer and wash the organic layer two more times with 120 mL water. Combine all of the aqueous layers and wash three times with 60 mL diethyl ether. 6. Dry the combined organic layers from step 5 over sodium sulfate, filter, and evaporate to dryness using a rotary evaporator equipped with a vacuum pump and a cooling trap. 7. Divide the oily residue into two fractions and purify each by column chromatography (APPENDIX 3E) on 350 g of 0.060- to 0.200-nm silica gel in a 5 × 35–cm column. Use a step gradient from 1.5 L of 100% dichloromethane to 1.5 L of 99:1 (v/v) dichloromethane/methanol to elute. Collect fractions in 2 × 18–cm test tubes. See Critical Parameters for additional discussion of column chromatography.
8. Draw a roster of the fraction collector on a TLC plate and apply a drop of each fraction onto the corresponding field of the plate. Spray with anisaldehyde/sulfuric acid spray and dry at 150°C. Combine the product-containing fractions (blue) and remove solvents in vacuo on a rotary evaporator. The resulting product, 1,5-anhydro-2,3,4,6-tetra-O-acetyl-D-glucitol (S.1), should be obtained in a 93% yield (18.82 g, 56.9 mmol). The spectroscopic properties are identical with those previously described (Kocienski and Pant, 1982).
Synthesis of Modified Nucleosides
1.9.3 Current Protocols in Nucleic Acid Chemistry
Supplement 15
Prepare S.2 9. Weigh 30.06 g (90.5 mmol) S.1 into a 1-liter round-bottom flask. Add 400 mL of 0.1 N sodium methoxide (freshly prepared from 0.92 g sodium and 400 mL dry methanol) and stir the reaction 2 hr at room temperature. 10. Neutralize the reaction mixture with ∼2 mL acetic acid (confirm with pH paper) and evaporate the solvent. 11. Coevaporate the residue with 20 mL toluene four times. Add 12.40 g (91.0 mmol) zinc chloride and 46.50 mL (455.0 mmol) benzaldehyde. Close the reaction flask with a rubber stopper and vigorously stir the suspension for two days at room temperature. 12. Pour the reaction mixture into 250 mL ice-water and extract with 100 mL ethyl acetate four times. Dry the combined organic layers over sodium sulfate. 13. Remove excess benzaldehyde on a rotary evaporator in vacuo (bath temperature 70°C) and wash the obtained solid residue with 100 mL n-hexane on a glass funnel. 14. Purify by column chromatography using 700 g silica gel in a 6 × 50–cm column. Elute with a step gradient of 1.5 L each: 1:1 (v/v) n-hexane/dichloromethane 100% dichloromethane 98:2 (v/v) dichloromethane/methanol. Collect fractions in 2 × 18–cm test tubes and monitor by TLC as in step 8. Combine product-containing fractions and remove solvent in vacuo on a rotary evaporator. The resulting product, 1,5-anhydro-4,6-O-benzylidene-D-glucitol (S.2), is obtained in 75% yield (17.1 g, 68.0 mmol). CIMS (iC4H10): m/e 253 (MH+); 1H NMR (DMSO-d6): δ = 3.00-3.90 (m, 7H) and 4.10-4.30 (m, 1H) (H-1′, H-1′′, H-2′, H-3′, H-4′, H-5′, H-6′, H-6′′), 5.03-5.31 (dd, 2H, 2′-OH, 3′-OH), 5.55 (s, 1H, PhCH), 7.20-7.57 (m, 5H, aromatic H); 13 C NMR (DMSO-d6): δ = 68.0 (C-6′), 70.2, 70.4 (C-1′, C-5′), 71.0 (C-2′), 74.4 (C-3′), 81.1 (C-4′), 100.7 (PhCH), 126.3, 127.9, 128.7, 137.8 (arom. C).
Prepare S.3a or S.3b 15. Suspend 8.50 g (33.7 mmol) S.2 and 8.38 g (33.7 mmol) dibutyltinoxide in 250 mL benzene. 16. Reflux the mixture 16 hr with azeotropic removal of water using a Dean-Stark condenser until the volume is reduced to ∼100 mL. Add 150 mL dioxane. 17. For S.3a, add 7.06 g (37.0 mmol) p-toluenesulfonyl chloride and heat the mixture 6 hr at 50°C, until a quantitative conversion to a less polar product (higher Rf value) is demonstrated by TLC. For S.3b, add 4.44 mL (33.7 mmol) p-toluoyl chloride and stir the mixture 5 hr at room temperature. For both, perform TLC using 1:2 (v/v) n-hexane/dichloromethane and visualize the products under a UV lamp. 18. Concentrate the mixture in vacuo using a rotary evaporator, absorb on Delite Celite, and purify by column chromatography using a step gradient from 1:1 (v/v) n-hexane/dichloromethane to 100% dichloromethane. Combine product-containing fractions and evaporate solvent on a rotary evaporator.
Synthesis of 1,5-Anhydrohexitol Building Blocks
1,5-Anhydro-4,6-O-benzylidene-2-O-(p-tolylsulfonyl)-D-glucitol (S.3a) is obtained in 82% yield (11.22 g, 27.6 mmol). EIMS: m/e 406 (MH+); 1H NMR (400 MHz, DMSO-d6): δ = 2.42 (s, 3H, CH3), 3.35-3.42 (m, 2H, H-4′, H-5′), 3.49 (t, 1H, J = 11 Hz, H-1′α), 3.61 (m, 1H, H-6′α), 3.67 (m, 1H, H-3′), 3.87 (dd, J = 5.5 and 11 Hz, 1H, H-1′β), 4.14-4.25 (m, 2H, H-2′, H-6′β), 5.05 (s, 1H, PhCH), 5.12 (d, J = 5.5 Hz, 1H, OH), 7.35-7.50 (m, 7H, H-aromatic), 7.85 (m, 2H, H-aromatic); 13C NMR (90 MHz, DMSO-d6): δ = 21.0 (CH3), 66.9, 67.6 (C-1, C-6), 70.7, 70.8 (C-3, C-5), 79.2, 80.4 (C-2, C-4), 100.7 (PhCH), + aromatic C.
1.9.4 Supplement 15
Current Protocols in Nucleic Acid Chemistry
1,5-Anhydro-4,6-O-benzylidene-2-O-(p-toluoyl)-D-glucitol (S.3b) is obtained in 78% yield (9.73 g, 26.3 mmol). CIMS (iC4H10): m/e 371 (MH+); 1H NMR (DMSO-d6): δ = 2.40, (s, 3H, CH3), 3.19-4.51 (m, 8H, H-1′, H-1′′, H-2′, H-3′, H-4′, H-5′, H-6′, H-6′′), 4.93-5.50 (br, s, 3′-OH), 5.55 (s, 1H, PhCH), 7.05-8.03 (m, 9H, H-aromatic); 13C NMR (DMSO-d6): δ = 21.5 (CH3), 67.2, 68.4 (C-1, C-6),70.9, 71.9, (C-3, C-5), 72.6 (C-2), 80.9( C-4), 101.9 (PhCH), 165.9 (C=O), + aromatic C.
Prepare S.4a or S.4b 19. Dissolve 23.60 g (193.0 mmol) DMAP and either 11.22 g (27.6 mmol) S.3a or 10.21 g (27.6 mmol) S.3b in 400 mL dichloromethane. 20. Cool the reaction mixture to −40°C with a dry ice/isopropanol mixture and add 2.53 mL (33.1 mmol) thiophosgene under vigorous stirring. 21. Allow the mixture to come slowly to room temperature and continue stirring 1 hr at room temperature. 22. Add 6.30 g (38.6 mmol) 2,4-dichlorophenol and continue stirring another 2 hr. Monitor by TLC using 1:5 (v/v) n-hexane/dichloromethane and visualize with a UV lamp. 23. Pour the mixture into 300 mL of 1 M potassium dihydrogenphosphate solution, pH 5, and extract with 300 mL dichloromethane twice. 24. Dry the organic layers over sodium sulfate, filter, and evaporate the solvent. 25. Purify the residue using a 5 × 35–cm flash column and a step gradient from 1.5 L of 2:8 (v/v) n-hexane/dichloromethane to 1.5 L of 100% dichloromethane. Combine product-containing fractions and evaporate solvent. 26. Dissolve the obtained thiocarbonyl compound in 300 mL toluene. 27. Bubble nitrogen gas through the solution for 10 min and then add 7.84 mL (29.1 mmol) tri-n-butyltin hydride and 0.33 g (2.0 mmol) AIBN. 28. Heat the reaction mixture overnight at 80°C. 29. Evaporate the mixture and purify by column chromatography as in step 25. 1,5-Anhydro-4,6-O-benzylidene-3-deoxy-2-O-(p-tolylsulfonyl)-D-ribohexitol (S.4a) is obtained in 64% yield (6.90 g, 17.7 mmol). CIMS (NH3): m/e 391 (MH+); 1H NMR (CDCl3): δ = 1.50-2.10 (m, 2H, H-3′, H-3′′), 2.48 (s, 3H, CH3), 3.06-4.84 (m, 7H, H-1′, H-1′′, H-2′, H-4′, H-5′, H-6′, H-6′′), 5.50 (s, 1H, PhCH), 7.04-7.98 (m, 9H, H aromat.); 13C NMR (CDCl3): δ = 21.4 (CH3), 35.3 (C-3′), 68.7, 69.0 (C-1′, C-6′), 72.9, 73.1 (C-4′, C-5′), 75.7 (C-2′), 101.5 (PhCH) + aromatic C. 1,5-Anhydro-4,6-O-benzylidene-3-deoxy-2-O-(p-toluoyl)-D-ribohexitol (S.4b) is obtained in 75% yield (7.12 g, 20.7 mmol). CIMS (iC4H10): m/e 355 (MH+); 1H NMR (CDCl3): δ = 1.42-2.12 (m, 2H, H-3′, H-3′′), 2.39 (s, 3H, CH3), 3.12-3.92 (m, 4H) and 4.02-4.49 (m, 2H) (H-1′, H-1′′, H-4′, H-5′, H-6′, H-6′′), 4.95-5.43 (m, 1H, H-2′), 5.54 (s, 1H, PhCH), 7.10-8.08 (m, 9H, H aromat.); 13C NMR (CDCl3): δ = 21.5 (CH3), 34.8 (C-3′), 66.9 (C-5′), 69.0, 69.1 (C-1′, C-6′), 73.4 (C-2′), 76.0 (C-4′), 101.5 (PhCH), 165.3 (C=O), + aromatic C.
Synthesis of Modified Nucleosides
1.9.5 Current Protocols in Nucleic Acid Chemistry
Supplement 14
Prepare S.5 30. Weigh 6.79 g (19.7 mmol) S.4b into a 1-liter round-bottom flask. 31. Add 300 mL of 0.1 N sodium methoxide (freshly prepared from 0.70 g sodium and 300 mL dry methanol) and stir the reaction 4 hr at room temperature. 32. Neutralize the reaction mixture with acetic acid (∼6 mL), confirming with pH paper, and evaporate the solvent. 33. Purify by flash chromatography using a 5 × 20–cm column and eluting with 99:1 (v/v) dichloromethane/methanol. Combine product-containing fractions and remove solvent in vacuo on a rotary evaporator. The resulting product of 1,5-anhydro-4,6-O-benzylidene-3-deoxy-D-glucitol (S.5) is obtained in 80% yield (3.72 g, 15.8 mmol). CIMS (iC4H10): m/e 237 (MH+); 1H NMR (DMSO-d6): δ = 1.20-1.66 (m, 1H, H-3′), 2.06-2.42 (m, 1H, H-3′′), 2.99-3.98 (m, 6H) and 4.05-4.30 (m, 1H) (H-1′, H-1′′, H-2′, H-4′, H-5′, H-6′, H-6′′), 5.08 (d, 1H, 2′-OH), 5.57 (s, 1H, PhCH), 7.17-7.67 (m, 5H, H aromatic); 13C NMR (DMSO-d6): δ = 38.3 (C-3′), 65.4 (C-5′), 69.1 (C-6′), 72.3, 73.0 (C-1′, C-2′), 76.3 (C-4′), 101.6 (PhCH), 126.1, 128.2, 129.0, 137.2 (arom. C). BASIC PROTOCOL 2
SYNTHESIS OF 1′,5′-ANHYDRO-2′,3′-DIDEOXY-2′-(N2-ISOBUTYRYLGUANIN-9-YL)-6′-O-MONOMETHOXYTRITYL-D-ARABINOHEXITOL This protocol details the synthesis of the hG 1,5-anhydrohexitol building block S.9 from S.5 (see Fig. 1.9.2 and DeBouvere et al., 1997). Materials 6-Chloro-9H-purine-2-amine 1,5-Anhydro-4,6-O-benzylidene-3-deoxy-D-glucitol (S.5; see Basic Protocol 1) Triphenylphosphine Dioxane (reflux overnight on lithium aluminum hydride and distill) Nitrogen gas Diisopropyl azodicarboxylate (DIAD) n-Hexane Ethyl acetate 10% (v/v) HCl Dichloromethane (store over phosphorous pentoxide and distill before use) Phenolpthalein solution 4 N sodium hydroxide Phosphorous pentoxide Pyridine (reflux overnight over potassium hydroxide distill before use) Bis(trimethylsilyl)acetamide (BSA) Isobutyric anhydride 25% (v/v) ammonia Diethyl ether Dimethylformamide (DMF; remove water by distillation with benzene followed by distillation under vacuum) 4-Monomethoxytrityl chloride (MMTr⋅Cl) Methanol Saturated sodium bicarbonate solution Sodium sulfate Toluene
Synthesis of 1,5-Anhydrohexitol Building Blocks
Dropping funnel Rotary evaporator equipped with a vacuum pump and cooling trap
1.9.6 Supplement 14
Current Protocols in Nucleic Acid Chemistry
Cl
Cl O O Ph
OH +
O
N
N H2 N
DIAD, Ph3P dioxane Ph
N H
N
5
Ph
P
O NH
N
MMTr⋅Cl pyridine DMF
O
N
O
6 O
N
N
HN NH
O MMTrO
N
O N
HN
10% HCl N O
O Ph O
N
N
Ph
N
N
1) BSA, pyridine 2) isobutyric anhydr. 3) NH3, H2O
H2N
HO 9
HO
HO
N
N
O
O
HO
N
HN
8
HO 7
Figure 1.9.2 Preparation of protected hG (S.9). Abbreviations: BSA, bis(trimethylsilyl)acetamide; DIAD, diisopropyl azodicarboxylate; DMF, dimethylformamide; MMTr⋅Cl, 4-monomethoxytrityl chloride; Ph3P, triphenylphosphine.
Oil bath and magnetic stirrer 5 × 35– and 4 × 25–cm chromatography columns Additional reagents and equipment for TLC and column chromatography (see Basic Protocol 1 and Critical Parameters) NOTE: The 1H NMR and 13C NMR spectra were determined with a 200 MHz Varian Gemini spectrometer with tetramethylsilane as internal standard. Abbreviations: s, singlet; d, doublet; dd, double doublet; t, triplet; br s, broad signal; m, multiplet; ddd, double doublet of doublet; dm, double multiplet. Liquid secondary-ion (LSIMS) mass spectra were obtained using a KRATOS Concept 1H mass spectrometer. Prepare S.6 1. Suspend 6.78 g (40.0 mmol) 6-chloro-9H-purine-2-amine, 4.72 g (20.0 mmol) S.5, and 13.12 g (50.0 mmol) triphenylphosphine in 200 mL dioxane. Carry out the reaction under a nitrogen gas atmosphere. 2. Add a solution of 9.84 mL (50.0 mmol) DIAD in 25 mL dioxane via a dropping funnel over a period of 200 min. 3. Stir the mixture overnight at room temperature. Monitor by TLC (see Basic Protocol 1, step 3) using 3:2 (v/v) n-hexane/ethyl acetate. Visualize product using a UV lamp (254 nm). 4. Remove volatiles by evaporation under vacuum using rotary evaporator equipped with a vacuum pump and cooling trap.
Synthesis of Modified Nucleosides
1.9.7 Current Protocols in Nucleic Acid Chemistry
Supplement 14
5. Absorb the crude product on 15.0 g of 0.060- to 0.200-nm silica gel in a 5 × 35–cm chromatography column and elute with a gradient from 1.5 L of 3:2 (v/v) to 1.5 L of 2:3 (v/v) n-hexane/ethyl acetate. Collect fractions in 2 × 18–cm test tubes. 6. Draw a roster of the fraction collector on a TLC plate and apply a drop of each fraction to the corresponding field on the plate. Visualize product-containing fractions using a UV lamp (254 nm). Combine the product-containing fractions and remove solvents in vacuo on a rotary evaporator. The resulting product, 1′,5′-anhydro-4′,6′-O-benzylidene-2′-6-chloro-2[(triphenylphosphoranylidene)amino]-9H-purin-9-yl-2′,3′-dideoxy-D-arabinohexitol (S.6) is obtained in 64% yield (8.31 g, 12.8 mmol). mp: 158°C; LSIMS (thygly): m/z: 648 [M+H]+; 1H NMR (CDCl3): δ = 2.00 (dt ,1H, 3′ax-H, J = 11.6 Hz, J = 4.5 Hz), 2.29 (br, d, 1H, 3′-eq-H, J = 13.0 Hz), 3.57 (m, 2H, 5′-H, 4′-H), 3.76 (t, 1H, 6′ax-H, J = 9.7 Hz), 4.10 (dd, 1H, 1′ax-H, J = 13.2 Hz, J = 2.8 Hz), 4.36 (m, 2H, 1′eq-H, 6′eq-H), 4.73 (br, s, 1H, 2′-H), 5.49 (s, 1H, PhCH), 7.19-7.72 (m, 21H, H aromat.), 8.15 (s, 1H, 8-H); 13C NMR (CDCl3): δ = 32.9 (C-3′), 50.2 (C-2′), 68.9 (C-6′), 69.5 (C-1′), 73.9 (C-4′), 74.5 (C-5′), 101.8 (PhCH), 122.8 (C-5), 125.9 (,2,6-C Ph), 127.5-133.4 (C, CH Ar),137.2 (C-6), 140.2 (C-8), 149.6 (C-4), 158.7 (C-2).
Prepare S.7 7. Suspend 8.20 g (12.6 mmol) S.6 in 160 mL of 10% HCl. Heat the reaction mixture 2 hr at 100°C. 8. Cool to room temperature and wash the yellow-brown solution with 60 mL dichloromethane to remove benzaldehyde and triphenylphosphine. 9. Add a drop of phenolphthalein solution as an indicator and neutralize the aqueous layer with 120 mL of 4 N sodium hydroxide. At pH 7 the product begins to precipitate.
10. Concentrate the suspension in vacuo using a rotary evaporator. 11. Dissolve the obtained white product in 745 mL boiling water and filter hot. 12. Cool to room temperature and incubate the mixture overnight at 4°C. 13. Filter off the obtained crystals and dry over phosphorus pentoxide. The resulting product, 1′,5′-anhydro-2′,3′-dideoxy-2′-(guanin-9-yl)-D-arabinohexitol (S.7) is obtained in 83% yield (2.95 g, 10.5 mmol). mp: >300°C; UV (H2O): λmax ε = 253 nm (9100); LSIMS (thgly): m/z: 282 [M+H]+; 1H NMR (DMSO-d6): δ = 1.80 (m, 1H, 3′ax-H), 2.17 (br, 1H, 3′eq-H), 3.20-3.70 (m, 2H, 5′-H, 4′-H, 6A-H), 3.79 (dd, 1H, 1′ax-H, J = 12.5 Hz, J = 2.2 Hz), 4.05-4.15 (m, 2H, 1′eq-H, 6B-H), 4.52 (s, br, 1H, 2′-H), 4.63 (t, 1H, 6′-OH, J = 6.0 Hz), 4.91 (d, 1H, 4′-OH, J = 5.3 Hz), 6.46 (br, s, 2H, NH2), 7.87 (s, 1H, 8-H); 13C NMR (DMSO-d6): δ = 36.3 (3′-C), 50.2 (2′-C), 61.0 ( 6′-C), 61.2 (4′-C), 68.4 (1′-C), 83.2 (5′-C), 116.3 (C-5), 136.9 (C-8), 151.5 (C-4), 154.1 (C-2), 157.9 (C-6).
Prepare S.8 14. Suspend 3.58 g (12.7 mmol) S.7 in 160 mL pyridine. 15. Add 16.57 mL (63.7 mmol) BSA and reflux 8 hr. 16. Stir the dark-red solution overnight (16 hr) at room temperature. 17. Add 10.00 g (63.7 mmol) isobutyric anhydride and stir 24 hr. 18. Cool the mixture to 0°C and add 20 mL water. Synthesis of 1,5-Anhydrohexitol Building Blocks
19. After 15 min, add 20 mL of 25% ammonia.
1.9.8 Supplement 14
Current Protocols in Nucleic Acid Chemistry
20. Continue stirring 2 hr at room temperature. 21. Evaporate the volatiles under vacuum and add 200 mL water. 22. Stir the suspension 10 min, then filter off the precipitate. Wash once with 50 mL water followed by three washes of 100 mL of 1:1 (v/v) ethyl acetate/diethyl ether. The resulting product, 1′,5′-anhydro-2′,3′-dideoxy-2′-(N2-isobutyrylguanin-9-yl)-D-arabinohexitol (S.8) is obtained in 90% yield (3.74 g, 10.7 mmol). mp: 258°C; LSIMS (thgly): m/z: 352 [M+H]+; 1H NMR (DMSO-d6): δ = 1.13 (d, 6H, 2 × CH3, J = 7 Hz), 1.87 (dt, 1H, 3′ax-H, J = 12.3 Hz, 4.2 Hz), 2.23 (br, d, 1H, 3′eq-H, J = 12.8 Hz), 2.78 (sept, 1H, CH, J = 7 Hz), 3.18 (m, 2H, 5′-H, 4′-H), 3.45-3.75 (m, 2H, 6′-H), 3.85 (dd, 1H, 1′ax-H, J = 12.5 Hz, J = 2.7 Hz), 4.17 (d, 1H, 1′eq-H, J = 12.4 Hz), 4.65 (m, 2H, 2′-H, 6′-OH), 5.01 (br, s, 1H, 4′-OH), 8.15 (s, 1H, 8-H), 10.2 (br, s, 1H, NH); 13C NMR (DMSO-d6): δ = 19.0 (2 × CH3), 34.8 (C-3′), 36.2 (CH), 50.4 (C-2′), 60.6 (C-6′, C-4′), 68.0 (C-1′), 83.1 (C-5′), 119.7 (C-5), 138.6 (C-8), 148.0 (C-2), 148.7 (C-4), 155.1 (C-6), 180.2 (HNC=O).
Prepare S.9 23. Suspend 4.03 g (11.5 mmol) S.8 in a mixture of 60 mL DMF and 60 mL pyridine. 24. Heat the suspension to 120°C until a clear red-brown solution is obtained. 25. Cool to room temperature and add 4.6 g (14.92 mmol, 1.3 eq) MMTr⋅Cl. 26. Stir the reaction overnight at room temperature. 27. Monitor the reaction by TLC using 94:6 (v/v) dichloromethane/methanol and visualize the product using a UV lamp (the product has a higher Rf value than the starting material). 28. Quench the reaction with 200 mL saturated sodium bicarbonate solution. 29. Extract with 100 mL dichloromethane four times. 30. Dry the organic layer over sodium sulfate. 31. Remove the solvent and coevaporate with 10 mL toluene three times. 32. Purify by flash chromatography on a 4 × 25–cm column using a step gradient from 500 mL of 100% dichloromethane to 99:1 (1 L), 98:2 (1 L), and 97:3 (v/v) dichloromethane/methanol. Combine product-containing fractions and evaporate solvents in vacuo on a rotary evaporator. The resulting product, 1′,5′-anhydro-2′,3′-dideoxy-2′-(N2-isobutyrylguanin-9-yl)-6′-Omonomethoxytrityl-D-arabinohexitol (S.9) is obtained in 93% yield (6.65 g, 10.7 mmol). mp: 165°C; LSIMS (thgly): m/z: 646 [M+Na]+; 1H NMR (CDCl3): δ = 1.19 (d, 6H, 2 x CH3, J = 6.8 Hz), 1.80 (t, 1H, 3′ax-H), J = 12.6 Hz), 2.20 (s, 1H, 4′-OH), 2.32 (br, d, 1H, 3′eq-H, J = 12.1 Hz), 2.70 (sept, 1H, CH, J = 6.9 Hz), 3.35-3.50 (m, 4H, 5′-H, 4′-H, 6A-H, 6B-H), 3.75 (m, 4H, 1′ax-H, OCH3), 4.20 (d, 1H, 1′eq-H, J = 13.4 Hz), 4.52 (br, s, 1H, 2′-H), 6.80 (d, 2H, aromatic H), 7.10-7.50 (m, 12H, aromatic H), 8.10 (s, 1H, 8-H), 9.66 (br, s, 1H, NH); 13C NMR (CDCl3): δ = 18.8 (CH3), 18.9 (CH3), 35.8 (3′-C), 36.0 (CH), 50.7 (2′-C), 55.1 (OCH3), 63.3 (4′-C), 63.9 (6′-C), 68.5 (1′-C), 81.1 (5′-C), 86.7 (O-C-Tr), 113.1 (m′-C, 2C), 120.0 (C-5), 126.9 (p-C, 2C), 127.8 (o-C, 4C), 128.3 (m-C, 4C), 130.2 (o′-C, 2C), 135.2 (i′C), 138.6 (C-8), 144.1 (iC, 2C), 147.8 (C-2), 148.8 (C-4), 156.0 (C-6), 158.5 (p’C), 180.0 (HNC=O).
Synthesis of Modified Nucleosides
1.9.9 Current Protocols in Nucleic Acid Chemistry
Supplement 14
NHBz
NH2 N
N adenine, LiH, 12-crown-4 DMF
O OTs O Ph
O
O Ph
4a
N
N
O
N
N BzCl pyridine
N
O Ph
O
N
O O
10
11
80% HOAc
NHBz
NHBz N
N N
N
N
MMTr⋅Cl pyridine
N
N
N
O
O HO
MMTrO
HO
HO 13
12
Figure 1.9.3 Preparation of protected hA (S.13). Abbreviations: BzCl, benzoyl chloride; DMF, dimethylformamide; HOAc, acetic acid; MMTr⋅Cl, monomethoxytrityl chloride.
BASIC PROTOCOL 3
SYNTHESIS OF 1′,5′-ANHYDRO-6′-MONOMETHOXYTRITYL-2′,3′DIDEOXY-2′-(N6-BENZOYLADENIN-9-YL)-D-ARABINOHEXITOL This protocol details the synthesis of the hA 1,5-anhydrohexitol building block S.13 from S.4a (see Fig. 1.9.3 and DeBouvere et al., 1997).
Synthesis of 1,5-Anhydrohexitol Building Blocks
Materials Adenine Lithium hydride 12-Crown-4 Dimethylformamide (DMF; remove water by distillation with benzene followed by distillation under vacuum) Nitrogen gas 1,5-Anhydro-4,6-O-benzylidene-3-deoxy-2-O-(p-tolylsulfonyl)-D-ribohexitol (S.4a; see Basic Protocol 1) n-Hexane Ethyl acetate Dichloromethane Saturated sodium bicarbonate solution Sodium sulfate Methanol Pyridine (reflux over potassium hydroxide overnight and distill) Benzoyl chloride 25% (v/v) ammonia
1.9.10 Supplement 14
Current Protocols in Nucleic Acid Chemistry
Toluene 80% (v/v) acetic acid Diethyl ether: reflux overnight on sodium (Na, FeCl2, Et2O) and distill Pyridine (reflux over potassium hydroxide overnight and distill) 4-Monomethoxytrityl chloride (MMTr⋅Cl) Oil bath and magnetic stirrer Rotary evaporator equipped with a vacuum pump and cooling trap 3 × 30–, 5 × 40–, and 4 × 30–cm chromatography columns Additional reagents and equipment for TLC and column chromatography (see Basic Protocol 1 and Critical Parameters) NOTE: The 1H NMR and 13C NMR spectra were determined with a 200 MHz Varian Gemini spectrometer with tetramethylsilane as internal standard. Abbreviations: s, singlet; d, doublet; dd, double doublet; t, triplet; br s, broad signal; m, multiplet; ddd, double doublet of doublet; dm, double multiplet. Liquid secondary-ion (LSIMS) mass spectra were obtained using a KRATOS Concept 1H mass spectrometer. Prepare S.10 1. Suspend 1.35 g (10.0 mmol) adenine, 0.08 g (9.6 mmol) lithium hydride, and 0.32 mL of 12-crown-4 (2.0 mmol) in 60 mL DMF. Carry out the reaction under a nitrogen gas atmosphere. 2. Heat the reaction mixture 1 hr at 110°C. 3. Add a solution of 1.95 g (5.0 mmol) of S.4a in 13 mL DMF with stirring. 4. Continue stirring at 110°C for 8 hr. 5. Test the reaction by TLC (see Basic Protocol 1, step 3) using 8:2 (v/v) hexane/ethyl acetate as solvent. 6. Cool the reaction mixture to room temperature and quench with 0.09 mL (5.0 mmol) water. 7. Concentrate the mixture under reduced pressure using a rotary evaporator with a vacuum pump. 8. Dissolve the residue in 100 mL dichloromethane and wash with 200 mL saturated sodium bicarbonate followed by two washes with 100 mL water. 9. Dry over sodium sulfate, filter, concentrate, and purify the residue by flash chromatography on a 3 × 30–cm column using a step gradient of 99:1 (1 L), 98:2 (1 L), 97:3 (1 L), and 96:4 (v/v) dichloromethane/methanol. Combine product-containing fractions and evaporate solvent in vacuo on a rotary evaporator. The resulting product, 2′-(adenin-9-yl)-1′,5′-anhydro-4′,6′-O-benzylidene-2′,3′-dideoxyD-arabinohexitol (S.10) is obtained in 82% yield (1.45 g, 4.1 mmol). mp: 227°C; UV (MeOH): λmax (ε) = 262 nm (11300); EIMS: m/z: 353 [M]+; 1H NMR (DMSO-d6): δ = 2.17 (dt, 1H, H- 3′ax, J = 12 Hz, J = 1.3 Hz, J = 4 Hz), 2.46 (m, 1H, H-3′eq), 3.53 (ddd, 1H, H-5′, J = 9 Hz, J = 10 Hz, J = 5 Hz), 3.73 (ddd, 1H, H-4′, J = 12.9 Hz, J = 4 Hz), 3.80 (t, 1H, H-6′ax, J = 10 Hz), 4.10 (dd, 1H, H-1′ax, J = 13 Hz, J = 2.5 Hz), 4.22 (dd, 1H, H-6′eq, J = 5 Hz), 4.44 (d, 1H, H-1′eq, J = 13 Hz), 4.90 (br, s, 1H, H-2′), 5.62 (s, 1H, PhCH), 7.30-7.40 (m, 7H, aromatic H, NH2), 8.18 (s, 1H, H-2), 8.27 (s, 1H, H-8); 13C NMR (DMSO-d6): δ = 32.2 (C-3′), 50.5 (C-2′), 68.1, 68.9 (C-1′, C-6′), 73.4, 73.8 (C-4′, C-5′), 100.9 (PhCH), 118.6 (C-5), 126.2 (2,6-C), 128.1 (3,5-C), 128.9 (4-C), 137.8 (1-C), 139.3 (C-8), 149.5 (C-4), 152.6 (C-2), 156.2 (C-6).
Synthesis of Modified Nucleosides
1.9.11 Current Protocols in Nucleic Acid Chemistry
Supplement 14
Prepare S.11 10. Dissolve 5.90 g (16.7 mmol) S.10 in 100 mL pyridine and coevaporate three times. 11. Add 160 mL pyridine and then cool to 0°C. 12. Add 9.7 mL (83.57 mmol) benzoyl chloride. 13. Allow the mixture to come to room temperature and continue stirring overnight. 14. Cool the orange-brown solution in an ice-bath and add 18 mL water. 15. After 5 min add 35 mL of 25% ammonia and continue stirring 2 hr at room temperature. 16. Remove the volatiles under reduced pressure and coevaporate three times with 20 mL toluene. 17. Dilute the resulting solid with 250 mL dichloromethane and wash with 200 mL saturated sodium bicarbonate solution. 18. Dry the organic layer over sodium sulfate, remove the solvent, and purify by column chromatography using a 5 × 40–cm silica gel column and a step gradient from 1.5 L of 2:8 (v/v) n-hexane/ethyl acetate to 1 L of 1:9 n-hexane/ethyl acetate, to 1 L of 100% ethyl acetate. Combine product-containing fractions and evaporate solvent. The resulting product, 1′,5′-anhydro-2′-(N6-benzoyladenin-9-yl)-4′,6′-O-benzylidene2′,3′-dideoxy-D-arabinohexitol (S.11) is obtained in 94% yield (7.20 g, 15.8 mmol). mp: 150°C; LSIMS (thygly): m/z: 458 [M+H]+; 1H NMR (CDCl3): δ = 2.25 (m, 1H, 3′ax-H, 2.66 (br, d, 1H, 3′eq-H, J = 12 Hz), 3.65 (m, 2H, 5′-H, 4′-H), 3.78 (t, 1H, 6′ax-H, J = 9.8 Hz), 4.16 (dd, 1H, 1′ax-H, J = 13.5 Hz, J = 2.5 Hz), 4.38 (dd, 1H, 6′eq-H, J = 10.2 Hz, J = 3.6 Hz), 4.48 (br, d, 1H, 1′eq-H, J = 13.3 Hz), 5.07 (br, s, 1H, 2′-H, 5.49 (s, 1H, PhCH), 7.50 (m, 8H, aromatic H), 8.04 (d, 2H, aromatic H, J = 6.7 Hz), 8.55 (s, 1H, 8-H), 8.77 (s, 1H, 2-H), 9.30 (s, 1H, NH); 13C NMR (CDCl3): δ = 33.1 (C-3′), 51.0 (C-2′), 68.8 (C-6′), 69.4 (C-1′), 73.8 (C-4′), 74.5 (C-5′), 102.0 (PhCH), 122.8 (C-5), 125.9 (2,6-C Ph), 127.9 (3,5C Ph), 128.2 (3,5-C Bz), 128.8 (2,6-C Bz), 129.1 (4-C Ph), 132.7 (4-C Bz), 133.5 (1-C Bz), 136.9 (1-C Ph), 142.0 (C-8), 149.6 (C-4), 152.0 (C-2), 152.5 (C-6), 164.7 (HNC=O).
Prepare S.12 19. Dissolve 8.06 g (17.6 mmol) S.11 in 450 mL of 80% (v/v) acetic acid. 20. Heat the reaction mixture 6 hr at 60°C. 21. Evaporate the solvent under reduced pressure using a rotary evaporator with a vacuum pump and coevaporate with 25 mL toluene three times. 22. Dissolve the residue in a minimal volume of 1:1 (v/v) dichloromethane/methanol. 23. Slowly add 500 mL diethyl ether while stirring. 24. Filter off the precipitate, wash with diethyl ether, and dry over phosphorous pentoxide.
Synthesis of 1,5-Anhydrohexitol Building Blocks
The resulting product, 1′,5′-anhydro-2′-(N6-benzoyladenin-9-yl)-2′,3′-dideoxy-D-arabinohexitol (S.12) is obtained in 82% yield (5.35 g, 14.5 mmol). mp: 220°C; LSIMS (thygly): m/z: 370 [M+H]+; 1H NMR (DMSO-d6): δ = 1.97 (dt, 1H, 3′ax-H, J = 13 Hz, J = 3.9 Hz), 2.36 (br, d, 1H, 3′eq-H, J = 13.2 Hz), 3.23 (m, 1H, 5′-H), 3.50-3.80 (m, 3H, 6′-H, 4′H), 3.93 (dd, 1H, 1′ax-H, J = 12.7 Hz, J = 2.1 Hz), 4.30 (br, d, 1H, 1′eq-H, J = 12.5 Hz), 4.7 (t, 1H, 6′-OH, J = 6.2 Hz), 4.98 (br, s, 1H, 2′-H), 5.00 (d, 1H, 4′-OH, J = 5.5 Hz), 7.58 (m, 3H, aromatic H), 8.05 (d, 2H, aromatic H, J = 6.9 Hz), 8.62 (s, 1H, 8-H), 8.75 (s, 1H, 2-H), 11.18 (s, 1H, NH); 13C NMR (DMSO-d6): δ = 35.8 (C-3′), 50.7 (C-2′), 60.5 (C-6′, C-4′), 67.9 (C-1′), 83.1 (C-5′)125.1 (C-5), 128.6 (2,3,5,6-C Bz), 132.5 (4-C Bz), 133.5 (1-C Bz), 143.5 (C-8), 150.2 (C-4), 151.5 (C-2), 152.4 (C-6), 165.7 (HNC=O).
1.9.12 Supplement 14
Current Protocols in Nucleic Acid Chemistry
Prepare S.13 25. Dissolve 5.35 g (14.5 mmol) S.12 in 100 mL pyridine and coevaporate three times. 26. Add 340 mL pyridine and dissolve. 27. Add 7.85 g (24.6 mmol) MMTr⋅Cl. 28. Stir the reaction 2 days at room temperature. 29. Monitor by TLC using 94:6 (v/v) dichloromethane/methanol (the product has a higher Rf value than the starting material). 30. Quench the reaction with 300 mL saturated sodium bicarbonate solution. 31. Extract with 400 mL dichloromethane twice. 32. Dry the organic layer over sodium sulfate, evaporate the solvent, and then coevaporate with 25 mL toluene three times. 33. Purify the residue by flash chromatography on a 4 × 30–cm column using a step gradient from 1.5 L of 100% dichloromethane to 99:1 (1 L), 98:2 (1 L), and 97:3 (v/v) dichloromethane/methanol. Combine product-containing fractions and evaporate the solvent. The resulting product, 1′,5′-anhydro-6′-monomethoxytrityl-2′,3′-dideoxy-2′-(N6-benzoyladenin-9-yl)-D-arabinohexitol (S.13) is obtained in 84% yield (7.79 g, 12.2 mmol). mp: 125°C; LSIMS (thygly/NaOAc): m/z: 664 [M+Na]+; 1H NMR (CDCl3): δ = 1.98 (dt, 1H, 3′ax-H, J = 11.6 Hz, J = 4.4 Hz), 2.54 (br, d, 1H, 3′eq-H, J = 12.6 Hz), 2.88 (br, s, 1H, 4′-OH), 3.47 (m, 3H, 5′-H, 4′-H, 6A-H), 3.77 (br, s, 4H, 6B-H, OCH3), 3.96 (dd, 1H, 1′ax-H, J = 12.9 Hz, J = 2.5 Hz), 4.34 (br, d, 1H, 1′eq-H, J = 12.8 Hz), 4.98 (br, s, 1H, 2′-H), 6.84 (d, 2H, aromatic H, J = 8.9 Hz), 7.38 (m, 15H, aromatic H), 8.02 (d, 2H, aromatic H, J = 8.3 Hz), 8.54 (s, 1H, 8-H), 8.77 (s, 1H, 2-H), 9.30 (br, s, 1H, NH); 13C NMR (CDCl3): δ = 35.7 (C-3′), 50.6 (C-2′), 55.2 (OCH3), 64.3 (C-4′, C-6′), 69.0 (C-1′), 80.6 (C-5′), 87.2 (O-C-MMTr), 113.1 (3′5′C MMTr), 120.0 (C-5), 127.1 (4C MMTr 2x), 127.9 (3,5C Bz), 128.0 (2,6C MMTr, 2×), 128.2 (3,5C MMTr, 2x), 128.8 (2,6C Bz), 130.2 (2′6′C MMTr), 132.7 (4C Bz), 133.7 (1C Bz), 134.9 (1′C MMTr), 142.4 (C-8), 143.8 (1C MMTr, 2×), 149.5 (C-4), 152.5 (C-2, C-6), 158.7 (HNC=O).
SYNTHESIS OF 1′,5′-ANHYDRO-6′-O-MONOMETHOXYTRITYL-2′,3′DIDEOXY-2′-(THYMIN-1-YL)-D-ARABINOHEXITOL
BASIC PROTOCOL 4
This protocol details the synthesis of the hT 1,5-anhydrohexitol building block S.16 from S.5 (see Fig. 1.9.4 and DeBouvere et al., 1997). Materials N3-Benzoylthymine 1,5-Anhydro-4,6-O-benzylidene-3-deoxy-D-glucitol (S.5; see Basic Protocol 1) Triphenylphosphine Tetrahydrofuran (THF; reflux overnight on lithium aluminum hydride and distill) Nitrogen gas Diethyl azodicarboxylate (DEAD) n-Hexane Ethyl acetate Saturated ammonia in methanol Dichloromethane Toluene 80% (v/v) acetic acid Methanol
Synthesis of Modified Nucleosides
1.9.13 Current Protocols in Nucleic Acid Chemistry
Supplement 14
O Bz N
Bz
OH O
Ph
O
O
O +
O
DEAD, Ph3P THF
N
O O
5
N H
N
O O
Ph
14
1) MeOH, NH3 2) 80% HOAc
O
O HN
HN O
MMTr⋅Cl pyridine
O
N
O MMTrO
HO 16
N
O HO
HO 15
Figure 1.9.4 Preparation of protected hT (S.16). Abbreviations: DEAD, diethyl azodicarboxylate; THF, tetrahydrofuran; Ph3P, triphenylphosphine; HOAc, acetic acid; MMTr⋅Cl, monomethoxytrityl chloride.
Pyridine (reflux over potassium hydroxide overnight and distill) Monomethoxytrityl chloride (MMTr⋅Cl) Saturated sodium bicarbonate solution Sodium sulfate Oil bath and magnetic stirrer Dropping funnel Rotary evaporator equipped with a vacuum pump and cooling trap 3 × 35– and 4 × 35–cm chromatography columns Additional reagents and equipment for TLC and column chromatography (see Basic Protocol 1 and Critical Parameters) NOTE: The 1H NMR and 13C NMR spectra were determined with a 200-MHz Varian Gemini spectrometer with tetramethylsilane as internal standard. Liquid secondary-ion (LSIMS) mass spectra were obtained using a KRATOS Concept 1H mass spectrometer. Abbreviations: s, singlet; d, doublet; dd, double doublet; t, triplet; br s, broad signal; m, multiplet; ddd, double doublet of doublet; dm, double multiplet.
Synthesis of 1,5-Anhydrohexitol Building Blocks
1.9.14 Supplement 14
Current Protocols in Nucleic Acid Chemistry
Prepare S.14 1. Dissolve 2.40 g (10.5 mmol) N3-benzoylthymine, 1.23 g (5.2 mmol) S.5, and 3.43 g (13.1 mmol) triphenylphosphine in 100 mL THF. Carry out the reaction under a nitrogen atmosphere. 2. Add a solution of 2.06 mL (13.1 mmol) DEAD in 15 mL THF via a dropping funnel over a period of 60 min. 3. Stir the mixture at room temperature overnight. 4. Remove the volatiles under vacuum using a rotary evaporator with a vacuum pump. 5. Absorb the crude product on 20 g of 0.060- to 0.200-nm silica gel and purify by flash chromatography in a 3 × 35–cm column using a step gradient of 1 L each of 8:2, 7:3, and 6:4 (v/v) n-hexane/ethyl acetate. Combine product-containing fractions and evaporate solvent in vacuo on a rotary evaporator. The resulting product, 1′,5′-anhydro-2′-(N3-benzoylthymin-1-yl)-4′,6′-O-benzylidene2′,3′-dideoxy-D-arabinohexitol (S.14) is obtained in 80% yield (1.88 g, 4.2 mmol). mp: 200°C; LSIMS (thygly): m/z: 449 [M+H]+; 1H NMR (CDCl3): δ = 2.02 (s, 3H, CH3), 2.07 (m, 1H, 3′ax-H), 2.47 (d, br, 1H, 3′eq-H, J = 13.9 Hz), 3.53 (dt, 1H, 5′-H, J = 9.6 Hz, J = 4.7 Hz), 3.8 (m, 1H, 4′-H), 3.80 (t, 1H, 6′ax-H, J = 10.2 Hz), 4.02 (dd, 1H, 1′ax-H, J = 13.7 Hz, J = 3.5 Hz), 4.28 (d, br, 1H, 1′eq-H, J = 13.9 Hz), 4.37 (dd, 1H, 6′eq-H, J = 10.5 Hz, J = 4.7 Hz), 4.73 (s, br, 1H, 2′-H), 5.64 (s, 1H, PhCH), 7.30-7.90 (m, 8H, aromatic H), 7.95 (s, + d, 3H, 6-H + aromatic 2H); 13C NMR (CDCl3): δ = 32.9 (C-3′), 51.7 (C-2′), 68.8 (C-6′, C-1′), 73.6 (C-4′), 74.2 (C-5′), 102.0 (PhCH), 110.7 (C-5), 126.0 (2,6C Ph), 128.3 (4C Ph), 129.1 (2,3,56C Bz), 130.4 (3,5 C Ph), 131.5 (1C Bz), 135.0 (4C Bz), 137.0 (1C Ph), 137.7 (C-6), 149.7 (C-2), 162.6 (C-4), 168.9 (HNC = O).
Prepare S.15 6. Dissolve 1.83 g (4.1 mmol) S.14 in 100 mL saturated ammonia in methanol. 7. Once a precipitate has formed, add 50 mL dichloromethane and continue stirring 90 min at room temperature. 8. Check the reaction by TLC (see Basic Protocol 1, step 3) using 1:1 (v/v) n-hexane/ethyl acetate (the starting material has the higher Rf value). 9. Evaporate the volatiles and coevaporate three times with 15 mL toluene. 10. Dissolve the crude product (1.8 g) in 75 mL of 80% (v/v) acetic acid. 11. Heat the reaction mixture 3 hr at 60°C. 12. Evaporate the volatiles on a rotary evaporator and coevaporate three times with 15 mL toluene. 13. Purify the residue by column chromatography using a 3 × 25–cm column and a gradient from 1 L of 95:5 to 1 L of 93:7 (v/v) dichloromethane/methanol. Combine product-containing fractions and evaporate the solvent. The resulting product, 1′,5′-anhydro-2′,3′-dideoxy-2′-(thymin-1-yl)-D-arabinohexitol (S.15) is obtained in 82% yield (0.86 g, 3.4 mmol). mp: 170°C; UV (MeOH): λmax (ε) = 272 nm (9500); LSIMS (thygly): m/z: 357 [M+H]+; 1H NMR (DMSO-d6): δ = 1.75 m, 1H, 3′ax-H), 1.76 (s, 3H, CH3), 2.08 (d, br, 1H, 3′eq-H, J = 13.8 Hz), 3.13 (m, 1H, 5′-H), 3.35 (m, 1H, 4′-H), 3.60 (m, 2H, 6′-H), 3.73 (dd, 1H, 1′ax-H, J = 12.9 Hz, J = 3.4 Hz), 3.99 (d,br, 1H, 1′eq-H, J = 12.8 Hz), 4.50 (s br,, 1H, 2′-H), 4.65 (s, br, 1H, 6′-OH), 4.91 (s, br, 1H, 4′-OH), 7.88 (s, 1H, 6-H), 11.25 (s, 1H, 3-H); 13C NMR (DMSO-d6): δ = 12.5 (CH3), 35.2 (C-3′), 50.1 (C-2′), 60.3 (C′-6′), 60.7 (C-4′), 67.0 (C-1′), 82.4 (C-5′), 108.4 (C-5), 139.0 (C-6), 151.0 (C-2), 163.9 (C-4).
Synthesis of Modified Nucleosides
1.9.15 Current Protocols in Nucleic Acid Chemistry
Supplement 14
Prepare S.16 14. Dissolve 2.77 g (10.8 mmol) S.15 in 40 mL pyridine and coevaporate three times. 15. Dissolve in 100 mL pyridine. 16. Add 5.85 g (18.4 mmol) MMTr⋅Cl. 17. Stir the reaction at room temperature overnight. 18. Check by TLC using 9:1 (v/v) dichloromethane/methanol as the solvent (the reaction mixture has the higher Rf value). 19. Quench the reaction with 200 mL saturated sodium bicarbonate solution. 20. Extract twice with 200 mL dichloromethane. 21. Dry the organic layer over sodium sulfate, evaporate the solvent, and coevaporate with 15 mL toluene three times. 22. Purify the residue by column chromatography on a 4 × 35–cm column using a step gradient from 500 mL of 100% dichloromethane to 1 L each of 99:1, 98:2, and 95:5 (v/v) dichloromethane/methanol. Combine product-containing fractions and evaporate solvents. The resulting product, 1′,5′-anhydro-6′-O-monomethoxytrityl-2′,3′-dideoxy-2′-(thymin-1yl)-D-arabinohexitol (S.16) is obtained in 86% yield (4.90 g, 9.3 mmol). mp: 125°C; LSIMS (thygly / NaOAc): m/z: 551 [M+Na]+; 1H NMR (CDCl3): δ = 1.85 (m, 1H, 3′ax-H), 1.86 (s, 3H, CH3), 2.28 (d, 1H, 4′-OH, J = 3.8 Hz), 2.38 (d, br, 1H, 3′eq-H, J = 14 Hz), 3.31 (m, 1H, 5′-H), 3.45 (m, 2H, 4′H, 6′ax-H), 3.78 (s, 3H, OCH3), 3.81 (dd, 1H, 1′ax-H, J = 13Hz, J = 3.4 Hz), 3.98 (m, 1H, 6′eq-H), 4.18 (d, br, 1H, 1′eq-H, J = 13.2 Hz), 4.67 (s, br, 1H, 2′-H), 6.83 (d, 2H, aromatic H, J = 8.7 Hz), 7.20-7.50 (m, 12H, aromatic H), 8.00 (s, 1H, 6-H), 9.20 (s, br, 1H, NH); 13C NMR (CDCl3): δ = 12.8 (CH3), 35.5 (C-3′), 51.0 (C-2′), 55.2 (OCH3), 63.1 (C-4′), 63.3 (C-6′), 68.5 (C-1′), 80.9 (C-5′), 86.8 (OC MMTr), 110.4 (C-5), 113.2 (3′5′C MMTr), 127.1 (4 MMTr, 2×), 128.0 (2,6C MMTr, 2×), 128.2 (3,5C MMTr, 2×), 130.2 (2′6′C MMtr), 135.0 (1′C MMTr), 138.6 (C-6), 143.9 (1C MMTr, 2×), 151.0 (C-2), 158.7 (4′C MMTr), 163.8 (C-4). BASIC PROTOCOL 5
SYNTHESIS OF 1′,5′-ANHYDRO-2′-(N4-BENZOYLCYTOSIN-1-YL)-2′,3′DIDEOXY-6′-MONOMETHYOXYTRITYL-D-ARABINOHEXITOL This protocol details the synthesis of the hC 1,5-anhydrohexitol building block S.20 from S.5 (see Fig. 1.9.5 and DeBouvere et al., 1997).
Synthesis of 1,5-Anhydrohexitol Building Blocks
Materials N4-Benzoyluracil 1,5-Anhydro-4,6-O-benzylidene-3-deoxy-D-glucitol (S.5; see Basic Protocol 1) Triphenylphosphine Dioxane (reflux overnight on lithium aluminum hydride and distill) Nitrogen gas Diethyl azodicarboxylate (DEAD) Saturated ammonia in methanol Dichloromethane Methanol Toluene Triazole Pyridine (reflux overnight over potassium hydroxide and distill) Phosphoroxy chloride 25% ammonia
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O
O O
Ph
N
+ O
5
O
1. DEAD, Ph3P THF
Bz
OH O
HN
O
2. MeOH/NH3
N H
N
O O
Ph
O
17
1. triazole/POCl3 2. NH4OH NHBz N O O MMTrO
N
MMTr⋅Cl pyridine N
20
O O
HO
HO
NH2
NHBz N
1. BzCl, NH3 2. TFA/CH2Cl2 N
HO
O O
O
Ph
19
N
O 18
Figure 1.9.5 Preparation of protected hC (S.20). Abbreviations: BzCl, benzoylchloride; DEAD, diethyl azodicarboxylate; MMTr⋅Cl, 4-monomethoxy chloride; POCl3, phosphoroxy chloride; TFA, trifluoroacetic acid.
Sodium sulfate Benzoyl chloride Saturated sodium bicarbonate solution Trifluoroacetic acid (TFA) Diethyl ether: reflux overnight on sodium (Na, FeCl2, Et2O) and distill Monomethoxytrityl chloride (MMTr⋅Cl) Dropping funnel Rotary evaporator equipped with a vacuum pump and cooling trap 4 × 30– and 3 × 35–cm chromatography columns Oil bath and magnetic stirrer Drying tube Additional reagents and equipment for TLC and column chromatography (see Basic Protocol 1 and Critical Parameters) NOTE: The 1H NMR and 13C NMR spectra were determined with a 200 MHz Varian Gemini spectrometer with tetramethylsilane as an internal standard. Abbreviations: s, singlet; d, doublet; dd, double doublet; t, triplet; br s, broad signal; m, multiplet; ddd, double doublet of doublet; dm, double multiplet. Liquid secondary-ion (LSIMS) mass spectra were obtained using a KRATOS Concept 1H mass spectrometer.
Synthesis of Modified Nucleosides
1.9.17 Current Protocols in Nucleic Acid Chemistry
Supplement 14
Prepare S.17 1. Suspend 6.48 g (30.0 mmol) N3-benzoyluracil, 4.72 g (20 mmol) S.5, and 7.87 g (30.0 mmol) triphenylphosphine in 200 mL dioxane. Carry out the reaction under a nitrogen atmosphere. 2. Add a solution of 5.90 mL (30.0 mmol) DEAD in 30 mL dioxane via a dropping funnel over a period of 200 min. 3. Stir the mixture overnight at room temperature. 4. Filter off the precipitate formed during the reaction and wash the precipitate with 50 mL dioxane. 5. Remove the volatiles of the filtrate by evaporation under vacuum in a rotary evaporator with a vacuum pump. 6. Dissolve the obtained residue in 100 mL saturated ammonia in methanol. 7. Stir the mixture 90 min at room temperature. 8. Check the reaction by TLC (see Basic Protocol 1, step 3) using 97.5:2.5 (v/v) dichloromethane/methanol. 9. Remove the volatiles and coevaporate with 20 mL toluene three times. 10. Purify the residue by column chromatography on a 4 × 30–cm column using a gradient from 1 L of 100% dichloromethane to 1 L of 97:3 (v/v) dichloromethane/methanol. Combine product-containing fractions and evaporate solvents in vacuo on a rotary evaporator. The resulting product, 1′,5′-anhydro-4′,6′-O-benzylidene-2′,3′-dideoxy-2′-(uracil-1-yl)-Darabinohexitol (S.17) is obtained in 67% yield (4.24 g, 12.8 mmol). mp: 200°C; LSIMS (thygly): m/z: 331 [M+H]+; 1H NMR (CDCl3): δ = 2.12 (dt, 1H, 3′ax-H, J = 11.9 Hz, J = 4.8 Hz), 2.52 (d ,br, 1H, 3′eq-H, J = 12.9 Hz), 3.48-3.74 (m, 2H, 5′-H, 4′-H), 3.79 (t, 1H, 6′ax-H, J = 10.2 Hz), 4.06 (dd, 1H, 1′ax-H, J = 13.7 Hz, J = 3.0 Hz), 4.37 (d, 1H, 1′eq-H, J = 13.8 Hz), 4.36 (dd, 1H, 6′eq-H, J = 10 Hz, J = 4.7 Hz), 4.78 (s, br, 1H, 2′-H), 5.59 (s, 1H, PhCH), 5.80 (d, 1H, J = 8.0 Hz,5-H), 7.34-7.51 (m, 5H, Ph H), 8.09 (d, 1H, 6-H, J = 8.0 Hz), 9.30 (s, br, 1H, NH); 13C NMR (CDCl3): δ = 32.8 (C-3′), 51.5 (C-2′), 68.8 (C-6′), 68.8 (C-1′), 73.6 (C-4′), 74.3 (C-5′), 102.0, 102.2 (PhCH, C-5), 126.0 (2,6-C Ph), 128.4 (3,5-C Ph), 129.3 (4-C Ph), 137.1 (1-C Ph), 142.4 (C-6), 150.9 (C-2), 163.2 (C-4).
Prepare S.18 11. Dissolve 6.23 g (96.0 mmol) triazole in 120 mL pyridine. 12. Add 2.61 mL (28.0 mmol) phosphoroxy chloride and continue stirring at room temperature for 25 min. 13. Add 2.64 g (8 mmol) S.17. The solution should turn yellow.
14. Continue stirring 45 min at room temperature. 15. Remove the volatiles in vacuo on a rotary evaporator and coevaporate with 15 mL toluene three times. 16. Dissolve the brown residue in 80 mL dioxane and add 40 mL of 25% ammonia. 17. Continue stirring another 30 min. Synthesis of 1,5-Anhydrohexitol Building Blocks
18. Evaporate to dryness, dissolve in 100 mL dichloromethane, and wash with 200 mL water.
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19. Dry the organic layer over sodium sulfate and remove the solvent. 20. Purify the crude product by column chromatography using 95:5 (v/v) dichloromethane/methanol as a solvent. Combine product-containing fractions and evaporate solvent. The resulting product, 1′,5′-anhydro-4′,6′-O-benzylidene-2′,3′-dideoxy-2′-(cytosin-1-yl)D-arabinohexitol (S.18) is obtained in 71% yield (1.87 g, 5.7 mmol). mp: 200°C; LSIMS (thygly): m/z: 330 [M+H]+; 1H NMR (CDCl3): δ = 2.05 (dt, 1H, 3′ax-H, J = 11.9 Hz, J = 4.8 Hz), 2.58 (d, br 1H, 3′eq-H, J = 12.9 Hz), 3.43-3.69 (m, 2H, 5′-H, 4′-H), 3.74 (t, 1H, 6′ax-H, J = 10.2 Hz), 4.00 (dd, 1H, 1′ax-H, J = 13.7 Hz, J = 3.0 Hz), 4.20 (d, 1H, 1′eq-H, J = 13.8 Hz), 4.30 (dd, 1H, 6′eq-H, J = 10 Hz, J = 4.7 Hz), 4.82 (s, br, 1H, 2′-H), 5.54 (s, 1H, PhCH), 5.86 (d, 1H, 5-H, J = 7.4 Hz, 7.30-7.47 (m, 5H, aromatic H), 8.05 (d, 1H, 6-H, J = 7.4 Hz); 13C NMR (CDCl3): δ = 32.5 (C-3′), 51.7 (C-2′), 68.8 (C-6′), 69.2 (C-1′), 73.8 (C-4′), 74.1 (C-5′), 94.5 (C-5), 101.9 (PhCH), 126.0 (2,6-C Ph), 128.3 (3,5-C Ph), 129.1 (4-C Ph), 137.2 (1-C Ph), 143.5 (C-6), 156.3 (C-2), 162.0 (C-4), 165.7 (HNC=O).
Prepare S.19 21. Dissolve 2.31 g (7.0 mmol) S.18 in 70 mL pyridine in a 250-mL round-bottom flask equipped with a drying tube. 22. Add 4.1 mL (35.0 mmol, 5 eq) benzoyl chloride and stir the reaction mixture overnight. 23. Analyze by TLC using 95:5 (v/v) dichloromethane/methanol as a solvent. 24. Cool the reaction mixture to 0°C and add 10 mL of 25% ammonia. 25. Continue stirring 1 hr at room temperature. 26. Remove the volatiles, coevaporate with 15 mL toluene three times, and dissolve the residue in 100 mL dichloromethane. 27. Wash with 100 mL saturated sodium bicarbonate solution, dry the organic layer over sodium sulfate, and remove the solvent in vacuo on a rotary evaporator. 28. Dissolve the resulting brownish foam in 70 mL dichloromethane. 29. Slowly add 30 mL TFA and continue stirring 45 min at room temperature. 30. Monitor by TLC using 9:1 (v/v) dichloromethane/methanol. 31. Evaporate the volatiles and coevaporate with 15 mL toluene three times. 32. Purify the resulting foam by column chromatography using a 3 × 35–cm column and a step gradient from 1.5 L of 95:5 to 1.5 L of 93:7 (v/v) dichloromethane/methanol. Collect product-containing fractions. 33. Reduce volatiles in vacuo on a rotary evaporator to a volume of 10 mL. Precipitate by adding 50 mL diethyl ether. The resulting product, 1′,5′-anhydro-2′-(N4-benzoylcytosin-1-yl)-2′,3′-dideoxy-D-arabinohexitol (S.19) is obtained in an overall yield of 67% (1.08 g, 3.1 mmol). mp: 130°C; LSIMS (thygly): m/z: 346 [M+H]+; 1H NMR (DMSO-d6): δ = 1.79 (dt, 1H, 3′ax-H, J = 13.6 Hz, J = 4.4 Hz), 2.20 (d, br, 1H, 3′eq-H, J = 13.7 Hz), 3.14 (m, 1H, 5′-H), 3.50-3.70 (m, 3H, 6′-H, 4′-H), 3.81 (dd, 1H, 1′ax-H, J = 12.9 Hz, J = 2.9 Hz), 4.14 (d, br, 1H, 1′eq-H, J = 13.2 Hz), 4.60 (m, 2H, 6′-OH, + 2′-H), 4.95 (d, 1H, 4′-H, J = 5.1 Hz), 7.31 (d, 1H, 5-H, J = 7.3 Hz), 7.57 (m, 3H, aromatic H), 8.01 (d, 2H, aromatic H, J = 7 Hz), 8.50 (d, 1H, 6-H, J = 7.4 Hz), 11.19 ( s, 1H, HNC=O); 13C NMR (DMSO-d6): δ = 34.8 (C-3′), 52.3 (C-2′), 60.5 (C-6′, C-4′), 67.2 (C-1′), 82.8 (C-5′), 95.9 (C-5), 128.5 (2,3,5,6-C Bz), 132.8 (1,4-C Bz), 148.1 (C-6), 155.0 (C-4), 162.6 (C-4), 167.5 (HNC=O).
Synthesis of Modified Nucleosides
1.9.19 Current Protocols in Nucleic Acid Chemistry
Supplement 15
Prepare S.20 34. Dissolve 2.72 g (7.9 mmol) S.19 in 40 mL pyridine and coevaporate three times (40 mL each). 35. Dissolve in 100 mL pyridine. 36. Add 4.26 g (13.4 mmol) of MMTr⋅Cl. 37. Stir the reaction at room temperature overnight. 38. Monitor by TLC using 95:5 (v/v) dichloromethane/methanol. 39. Quench the reaction with 200 mL saturated sodium bicarbonate solution. 40. Extract twice with 200 mL dichloromethane. 41. Dry the organic layer over sodium sulfate, evaporate the solvent, and coevaporate three times with 20 mL toluene. 42. Purify the residue by column chromatography on a 3 × 35–cm column using a step gradient of 1 L each 99:1, 98:2, 97:3, and 96:4 (v/v) dichloromethane/methanol. Combine product-containing fractions and evaporate solvents. Th e resu lting product, 1′,5′-anhydro-2′-(N4-benzoylcytosin-1-yl)-2′,3′-dideoxy-6′monomethoxytrityl-D-arabinohexitol (S.20) is obtained in 80% yield (3.88 g, 6.3 mmol). mp: 140°C; LSIMS (thygly / NaOAc): m/z: 640 [M+Na]+; 1H NMR (CDCl3): δ = 1.90 (m, 1H, 3′ax-H), 2.55 (br, d, 1H, 3′eq-H, J = 12.2 Hz), 3.08 (s, br, 1H, 5′-H), 3.42 (m, 3H, 4′H, 6′-H), 3.79 (s, 3H, OCH3), 3.89 (dd, 1H, 1′ax-H, J = 13.5 Hz, J = 2.9 Hz), 4.01 (s, br, 1H, 4′-OH), 4.25 (d, br, 1H, 1′eq-H, J = 13.5 Hz), 4.83 (s, br, 1H, 2′-H), 6.85 (d, 2H, aromatic H, J = 8.8 Hz), 7.20-7.70 (m, 18H, aromatic H + 5-H), 8.73 (br, d, 2H, NH + 6-H, J = 7.6 Hz); 13C NMR (CDCl3): δ = 35.0 (C-3′), 52.7 (C-2′), 55.1 (OCH3), 62.1 (C-4′), 62.7 (C-6′), 68.4 (C-1′), 80.9 (C-5′), 86.8 (OC Tr), 96.4 (C-5), 113.2 (3′5′C), 127.1 (4C, 2×), 127.9 (3,5C Bz), 128.0 (2,6C 2×), 128.4 (3,5C 2×), 128.9 (2,6C Bz), 130.0 (2′6′C), 132.7 (4C Bz), 132.9 (1C Bz), 135.0 (1′C), 143.9 (1C 2×), 148.0 (C-6), 155.5 (C-2), 158.6 (4′C), 161.9 (C-4), 166.3 (HNC=O).
COMMENTARY Background Information
Synthesis of 1,5-Anhydrohexitol Building Blocks
Hexitol nucleic acid (HNA) is the first example of an oligonucleotide with a six-membered carbohydrate moiety that hybridizes with natural nucleic acids. The base moiety is connected to the 2′-position of the sugar, which means that the nucleic acids have no anomeric center. These nucleic acids are stable against enzymatic and chemical degradation (Hendrix et al., 1997a,b). HNA can be synthesized by the phosphoramidite method (Hendrix et al., 1997a,b; also see UNIT 3.3) starting from the building blocks described in this protocol. HNAs are RNA-selective oligonucleotides, which means that they hybridize more strongly to RNA as the complement than to DNA (Hendrix et al., 1997a,b). The HNA-RNA duplex is thermally more stable than the DNA-RNA duplex, making HNA suitable as a steric blocking agent for antisense purposes (∆Tm/mod +1° to + 5°C). This higher stability is mainly based on its preorganization (entropy factor).
The NMR structure of an HNA-RNA duplex has been solved (Lescrinier et al., 2000a,b) and it shows typical A-form geometry. The hexitol nucleoside itself is a good mimic of a furanose nucleoside in its 2′-exo,3′-endo conformation (northern type). In a qualitative model, cellular uptake of HNA seems to be similar to that of phoshorothioate oligonucleotides, using cationic lipids as a transfecting agent (Atkins et al., 2000). Hybridization of HNA with RNA is more sequence specific than between DNA and RNA, which is demonstrated by the larger difference in Tm between match and mismatch duplexes of HNA-RNA versus DNA-RNA (Hendrix et al., 1997b). HNA functions as a very efficient template for nonenzymatic oligomerization of activated ribonucleoside monophosphates (Kozlov et al., 1999a,b). HNA templates are chiral-discriminating in that they selectively accept D-nucleotides over L-nucleotides for primer extension
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(Kozlov et al., 1999a,b). Two HNA codons (6 nucleotides) incorporated in an otherwise intact RNA are accepted as messenger in the translation process (Lavrik et al., 2001). Hexitol nucleoside triphosphates may be used by polymerases for enzymatic incorporation into oligodeoxyribonucleotides (Vastmans et al., 2000).
Critical Parameters Overall, it is important that for each step of the syntheses the starting sugar- and basebuilding blocks or nucleosides, respectively, are thoroughly dried either by coevaporation with anhydrous pyridine or in a desiccator over phosphorus pentoxide. The MMtr-protected building blocks (S.9, S.13, S.16, and S.20) are dried over silica blue (not over phosphorous pentoxide) and can be stored at 4°C. For all reactions using anhydrous solvents, the glassware must be predried at 70°C for at least 2 hr. All solvents must be distilled before use. Anhydrous solvents are very important. They should either be freshly distilled and stored under nitrogen or argon, or be taken from a freshly opened bottle of commercially prepared anhydrous solvent. For evaporation of solvents, it is helpful if the rotary evaporator is equipped with a dry ice condenser. For all compounds, a sample of 100 mg should be kept as a reference. In all cases, the reaction progress is followed by TLC. The starting material (1 mg diluted to 200 µL) as well as the reaction mixture (100 µL) are prepared. A baseline is marked and the spots of the solutions are placed at equal distances, with the reaction mixture in the middle surrounded by starting material. After developing in the appropriate solvent, the end line of the TLC is marked, and spots are identified first under a UV lamp (254 nm) and then by spraying the plates with anisaldehyde/sulfuric acid spray and heating to 150°C. The sugar-containing products turn blue. The monomethoxytritylated products (S.9, S.13, S.16, and S.20) turn yellow immediately after spraying (deprotection). After heating, the product-containing spots turn green. Before each column purification, TLC is performed on the reaction mixture to evaluate the completeness of the reaction and the presence of starting materials to be recovered. Silica gel (0.060 to 0.200 nm) is suspended in the first solvent and loaded into the column, which is tightly packed by gentle tapping. The silica
layer is topped by a 1-cm layer of sand. The crude product is dissolved in a minimum volume of solvent and applied to the column. The fractions are collected with a fraction collector and a roster of the collector is drawn on a TLC plate. A drop from each fraction is applied to the corresponding field on the plate, and the plates are evaluated under a UV lamp and by applying anisaldehyde/sulfuric acid spray. The fraction-containing products (or unreacted starting material) are tested for homogeneity by TLC. Elution of the column is continued with the final solvent of the indicated gradient until all product has been isolated. Product-containing fractions are combined and solvents are removed in vacuo on a rotary evaporator.
Troubleshooting Preparation of hC The identical product can be obtained by Mitsunobu reaction of N4-benzoylcytosine with the hexitol building block S.5 as demonstrated in De Bouvere et al. (1997). In this case the yield was rather modest (34%) and turned out to be less reproducible. Furthermore, in some attempts the 1′,5′-anhydro-2′-(N4-benzoylcytosin-2-yl)-4′,6′-O-benzylidene-2′,3′dideoxy-D-arabin ohexitol (O-nucleoside) instead of the desired 1′,5′-anhydro-2′-(N4benzoylcytosin-1-yl)-4′,6′-O-benzylidene-2′, 3′-dideoxy-D-arabinohexitol (N-nucleoside) was obtained. Preparation of hG Due to the high price of 6-iodo-9H-purine2-amine, the synthesis of hG may be performed using the Mitsunobu reaction of 6-chloro-9Hpurine-2-amine with the hexitol building block S.5. If 6-iodo-9H-purine-2-amine is available, another approach for obtaining hG is described in De Bouvere et al. (1997).
Anticipated Results Preparation of appropriately protected anhydrohexitol building blocks is straightforward using the described protocols. Automated oligomer assembly using standard phosphoramidite chemistry cycles—as provided by the manufacturers of automated DNA synthesizers—will yield anhydrohexitol oligomers.
Time Considerations In the planning of the syntheses it has to be considered that most of reactions have to be stirred overnight. Normally each step (includ-
Synthesis of Modified Nucleosides
1.9.21 Current Protocols in Nucleic Acid Chemistry
Supplement 14
ing purification and spectroscopic analysis) can be performed in 1.5 to 2 working days.
Literature Cited Atkins, D., Miller, M., De Bouvere, B., Van Aerschot, A., and Herdewijn, P. 2000. Evaluation of the cellular uptake of hexitol nucleic acids in HeLa cells. Pharmazie 55:615-617. De Bouvere, P., Kerremans, L., Rozenski, J., Janssen, G., Van Aerschot A., Claes, P., Busson, R., and Herdewijn, P. 1997. Improved synthesis of anhydrohexitol building blocks for oligonucleotide synthesis. Liebigs Ann./Recueil 14531461. De Winter, H., Lescrinier, E., Van Aerschot, A., and Herdewijn, P. 1998. Molecular dynamics simulation to investigate differences in minor groove hydration of HNA/RNA hybrids as compared to HNA/DNA complexes. J. Am. Chem. Soc. 120:5381-5394. Hendrix, C., Verheggen, I., Rosemeyer, H., Seela, F., Van Aerschot, A., and Herdewijn, P. 1997a. 1′,5′Anhydrohexitol oligonucleotides: Synthesis, base pairing and recognition by regular oligodeoxyribonucleotides and oligoribonucleotides. Chem. Eur. J. 3:110-119. Hendrix, C., Rosemeyer, H., De Bouvere, B., Van Aerschot A., Seela, F., and Herdewijn, P. 1997b. 1′,5′-Anhydrohexitol oligonucleotides: Hybridization and strand displacement with oligoribonucleotides, interaction with RNase H and HIV reverse transcriptase. Chem. Eur.J. 3:15131520. Kocienski, P. and Pant, C. 1982. A convenient preparation of some 2,3,4,6-tetraacetyl-1,5-anhydroD-hexitols. Carbohydr. Res. 110:330–332. Kozlov, I.A., Politis, P.K., Pitsch, S., Herdewijn, P., and Orgel, L.E. 1999a. A highly enantio-selective hexitol nucleic acid template for nonenzymatic oligoguanylate synthesis. J. Am. Chem. Soc. 121:1108-1109.
Kozlov, I.A., Politis, P.K., Van Aerschot, A., Busson, R., Herdewijn, P., and Orgel, L.E. 1999b. Nonenzymatic synthesis of RNA and DNA oligomers on hexitol nucleic acid templates: The importance of the A structure. J. Am. Chem. Soc. 121:2613-2656. Lavrik, I.N., Avdeeva, O.N., Dontsova, O.A., Froeyen, M., and Herdewijn, P.A. 2001. Translational properties of mHNA, a messenger RNA containing anhydrohexitol nucleotides. Biochemistry 40:11777-11784. Lescrinier, E., Esnouf, R.M., Schraml, J., Busson, R., and Herdewijn, P. 2000a. Solution structure of a hexitol nucleic acid duplex with four consecutive T.T base pairs. Helv. Chim. Acta 83:1291-1310. Lescrinier, E., Esnouf, R.M., Schraml, J., Busson, R., Heus, H.A., Hilbers, C.W., and Herdewijn, P. 2000b. Solution structure of a HNA-RNA hybrid. Chem. & Biol. 7:719-731. Vastmans, K., Pochet, S., Peys, A., Kerremans, L., Van Aerschot, A., Hendrix, C., Marlière, P., and Herdewijn, P. 2000. Enzymatic incorporation in DNA of 1,5-anhydrohexitol nucleotides. Biochemistry 39:12757-12765. Verheggen, I., Van Aerschot, A., Toppet, S. Snoeck, R., Janssen, G., Balzarini, J., De Clercq, E., and Herdewijn, P. 1993. Synthesis and antiherpes virus activity of 1,5-anhydrohexitol nucleosides. J. Med. Chem. 36:2033-2039. Verheggen, I., Van Aerschot, A., Van Meervelt, L., Rozenski, J., Wiebe, L., Snoek, R., Andrei, G., Balzarini, J., Claes, P., De Clercq, E., and Herdewijn, P. 1995. Synthesis, biological evaluation, and structure analysis of a series of new 1,5-anhydrohexitol nucleosides J. Med. Chem. 38:826835.
Contributed by Irene M. Lagoja, Arnaud Marchand, Arthur Van Aerschot, and Piet Herdewijn Rega Institute for Medical Research Leuven, Belgium
Synthesis of 1,5-Anhydrohexitol Building Blocks
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Synthesis and Properties of 7-Substituted 7-Deazapurine (Pyrrolo[2,3-d]pyrimidine) 2-Deoxyribonucleosides
UNIT 1.10
This unit describes the synthesis of 7-deazapurine nucleosides, including their 7-substituted derivatives. (Systematic numbering is used for compound naming in protocol steps, while purine numbering is used elsewhere in the unit.) Initially, the synthesis of 7-deazapurine nucleosides was made difficult by the particular properties of the 7-deazapurine system. While the imidazole nitrogen of a purine base can easily be attacked by an electrophile (e.g., a sugar cation) without affecting the state of aromaticity, eletrophilic reactions at the pyrrole nitrogen of a 7-deazapurine would destroy the aromatic character of the pyrrole ring. Thus, the pyrrole nitrogen is rather inert to glycosylation, with the consequence that the reaction is instead directed to the pyrimidine moiety or to the pyrrole carbons. In 1983, the nucleobase anion method for glycoslyation of 7-deazapurines was developed (Winkeler and Seela, 1983; Seela et al., 1988). From this work, it became apparent that reaction via the pyrrolyl anion occurred regioselectively at N9 and proceeded in a stereocontrolled manner, with the exclusive formation of 2 -deoxy-β-D-ribonucleosides. Figure 1.10.1 shows a selection of 7-deazapurine 2 deoxyribonucleosides and 7-substituted derivatives that have been synthesized by the nucleobase anion glycosylation method. This unit focuses on the synthesis of the 7-substituted 7-deazapurine 2 deoxyribonucleosides S.1d, S.2d, S.3c, S.4c, S.5d, and S.6d. The preparation of the 7-iodo-7-deazapurine 2 -deoxyribonucleosides S.1d, S.2d, S.5d, and S.6d is described first (see Basic Protocol 1 and Basic Protocol 2), followed by the preparation of the brominated nucleosides S.3c and S.4c (see Basic Protocol 3). Selective halogenation reactions are performed either on the nucleobases or on the nucleosides, using N-halosuccinimides. Nucleobase anion glycosylation is carried out in acetonitrile in the presence of powdered potassium hydroxide and a phase-transfer catalyst. The determination of pKa values of 7-deazapurine nucleosides is summarized in Basic Protocol 4. Fluorescence properties are also discussed (see Background Information).
PREPARATION OF 7-SUBSTITUTED 7-DEAZAPURINE NUCLEOSIDES RELATED TO 2 -DEOXYADENOSINE AND 2 -DEOXYINOSINE
BASIC PROTOCOL 1
The syntheses of 7-iodo-7-deaza-2 -deoxyadenosine (S.1d) and the analog 7-iodo-7deaza-2 -deoxyinosine (S.5d) are shown in Figure 1.10.2. The 7-iodinated nucleobase S.7 serves as the starting material. The preparation of this nucleobase is reported elsewhere (Pudlo et al., 1990; Rolland et al., 1997). The chemical conversions described here proceed in high yields, and the purification of intermediates is straightforward.
Materials Potassium hydroxide (KOH) powder (purity, ≥85%; Sigma) Anhydrous acetonitrile (MeCN) Tris[2-(2-methoxyethoxy)ethyl]amine (TDA-1) 4-Chloro-5-iodo-7H-pyrrolo[2,3-d]pyrimidine (S.7; see Pudlo et al., 1990) 2-Deoxy-3,5-di-O-(p-toluoyl)-α-D-erythro-pentofuranosyl chloride (S.8; see Hoffer, 1960; Rolland et al., 1997) Anhydrous dichloromethane (CH2 Cl2 ) Silica gel 60 (particle size, <0.063 mm; Merck)
Contributed by Frank Seela and Xiaohua Peng Current Protocols in Nucleic Acid Chemistry (2005) 1.10.1-1.10.20 C 2005 by John Wiley & Sons, Inc. Copyright
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Figure 1.10.1 The structures of selected 7-deazapurine 2 -deoxyribonucleosides. Purine and systematic numbering systems are shown in the first two structures. Compounds: S.1, 7-deaza-2 -deoxyadenosine; S.2, 7-deaza-2 -deoxyguanosine; S.3, 2-amino-7-deaza-2 -deoxyadenosine; S.4, 7-deaza-2 -deoxyisoguanosine; S.5, 7-deaza-2 -deoxyinosine; S.6, 7deaza-2 -deoxyxanthosine. References: S.1a (Seela and Kehne, 1983); S.1b-e (Seela and Thomas, 1994; Seela and Zulauf, 1996); S.1f (Seela et al., 2005); S.2a (Winkeler and Seela, 1983); S.2b-e (Ramzaeva and Seela, 1995); S.3a (Seela et al., 1987); S.3b-d (Seela and Peng, 2004); 4a (Seela and Wei, 1999); S.4b,c (Seela and Peng, 2004); S.5a (Seela and Menkhoff, 1985); S.5b-d (Ramzaeva et al., 1999); S.6a (Seela et al., 1985); and S.6b-d (Seela and Shaikh, 2004).
7-Substituted 7-Deazapurine (Pyrrolo[2,3d]pyrimidine) 2 -Deoxyribonucleosides
Ethyl acetate (EtOAc) Petroleum ether (b.p. range, 40◦ to 80◦ C) Isopropanol (i-PrOH) Methanol (MeOH) 1,4-Dioxane 25% (v/v) ammonium hydroxide (NH4 OH) Palladium catalyst Pd(PPh3 )4 Copper iodide (CuI) Argon gas Anhydrous dimethylformamide (DMF) Triethylamine (Et3 N) Cyclopentylacetylene Sodium methoxide (NaOMe) 2 M NaOH 1 M HCl 10-, 50-, and 100-mL round-bottom flasks 3-cm-diameter Buchner funnel with filter paper circles Rotary evaporator connected to a vacuum pump
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Figure 1.10.2 Preparation of 7-deaza-7-iodo-2 -deoxyadenosine (S.1d) and 7-deaza-7-iodo-2 -deoxyinosine (S.5d) is carried out by glycosylation of an iodinated nucleobase. S1.d can then be used in a palladium-catalyzed Sonogashira crosscoupling reaction to yield the 7-alkynyl derivative S.10 (Seela et al., 2000). TDA-1, tris[2-(2-methoxyethoxy)ethyl]amine; Tol, p-toluoyl.
3 × 50– and 4 × 50–cm chromatography columns 0.2-mm-thick silica gel 60 F254 aluminum TLC plates (Merck) 254-nm UV lamp High-vacuum pump (final pressure, <1 mmHg) Steel bomb (autoclave) Reflux condenser Heating mantle with controller Additional reagents and equipment for flash chromatography (APPENDIX 3E) and thin-layer chromatography (TLC; APPENDIX 3D) Prepare common precursor S.9 1. In a 100-mL round-bottom flask equipped with a Teflon stir bar, suspend 0.50 g (8.91 mmol) powdered KOH in 60 mL MeCN. 2. Add 75 µL (0.23 mmol) TDA-1 and stir the resulting mixture 10 min at room temperature. 3. Add 1.0 g (3.57 mmol) 4-chloro-5-iodo-7H-pyrrolo[2,3-d]pyrimidine (S.7) and stir 10 min at room temperature.
Synthesis of Modified Nucleosides
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4. Add 1.7 g (4.37 mmol) 2-deoxy-3,5-di-O-(p-toluoyl)-α-D-erythro-pentofuranosyl chloride (S.8) and stir 10 min at room temperature. 5. Remove the insoluble material by vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, washing with three 10-mL aliquots of MeCN. Evaporate the filtrate using a rotary evaporator connected to a vacuum pump. 6. Dissolve the residue in 30 mL CH2 Cl2 , adsorb on ∼3 g silica gel 60, and evaporate the solvent using a rotary evaporator. 7. Load the dry material onto a 4 × 50–cm column packed with silica gel 60 (bed height, 15 cm) and perform flash chromatography (APPENDIX 3E), eluting with 800 mL of 1:4 (v/v) EtOAc/petroleum ether. Collect 15-mL fractions. 8. Monitor fractions by TLC (APPENDIX 3D) on silica gel 60 F254 plates using 1:4 (v/v) EtOAc/petroleum ether as the eluent. Visualize product under 254-nm light using a UV lamp. 9. Pool all product fractions and evaporate the solvent using a rotary evaporator. 10. Crystallize the purified product from i-PrOH, collect the crystals by vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, and dry overnight under high vacuum (<1 mmHg). 4-Chloro-7-[2-deoxy-3,5-di-O-(4-toluoyl)-β-D-erythro-pentofuranosyl]-5-iodo-7Hpyrrolo[2,3-d]pyrimidine (S.9) is obtained as colorless crystals (1.5 g, 66% yield). m.p. 138◦ C. TLC (EtOAc/petroleum ether, 1:4): Rf = 0.5. UV (MeOH): λmax (ε) = 235 nm (25,600). 1 H NMR (DMSO-d6 ): δ 2.39 and 2.41 (2 × s, 6H, 2 × CH3 ), 2.78 (m, 1H, 2 -Hα ), 3.11 (m, 1H, 2 -Hβ ), 4.56 (m, 2H, 5 -H), 4.66 (m, 1H, 4 -H), 5.77 (m, 1H, 3 -H), 6.77 (t, J = 7.0 Hz, 1H, 1 -H), 7.34 and 7.90 (2 × dd, J = 7.8 Hz, J = 7.9 Hz, 8H, 2 × C6 H4 ), 8.18 (s, 1H, 6-H), 8.66 (s, 1H, 2-H). Anal. calcd. for C27 H23 ClIN3 O5 : C, 51.32; H, 3.67; N, 6.65; found: C, 51.55; H, 3.78; N, 6.80.
Prepare 2 -deoxyadenosine analog S.1d 11. In a 100-mL round-bottom flask equipped with a Teflon stir bar, suspend 1.6 g (2.53 mmol) compound S.9 in 40 mL 1,4-dioxane, and then place the flask in an autoclave. 12. Add 80 mL of 25% NH4 OH and stir 15 hr at 110◦ C under pressure. 13. Adsorb reaction mixture onto ∼4 g silica gel 60 and evaporate the solvent using a rotary evaporator. 14. Purify the crude product by flash chromatography (step 7; bed height, 20 cm) using an elution gradient from 95:5 (v/v) CH2 Cl2 /MeOH (300 mL) to 90:10 (v/v) CH2 Cl2 /MeOH (600 mL). Collect 10-mL fractions. 15. Monitor fractions by TLC (step 8) using 9:1 (v/v) CH2 Cl2 /MeOH as the eluent. 16. Pool all product fractions and evaporate the solvent using a rotary evaporator. 17. Crystallize purified product from MeOH, collect the crystals by vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, and dry overnight under high vacuum (<1 mmHg).
7-Substituted 7-Deazapurine (Pyrrolo[2,3d]pyrimidine) 2 -Deoxyribonucleosides
4-Amino-7-(2-deoxy-β-D-erythro-pentofuranosyl)-5-iodo-7H-pyrrolo[2,3-d]pyrimidine (S.1d) is obtained as colorless crystals (612 mg, 64% yield). m.p. 194◦ C. TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.40. UV (MeOH): λmax (ε) = 283 nm (8500). 1 H NMR (DMSO-d6 ): δ 2.16 (m, 1H, 2 -Hα ), 2.46 (m, 1H, 2 -Hβ ), 3.54 (m, 2H, 5 -H), 3.81 (m, 1H, 4 -H), 4.33 (m, 1H, 3 -H), 5.00 (t, J = 5.4 Hz, 1H, 5 -OH), 5.23 (d, J = 3.9 Hz, 1H, 3 -OH), 6.49 (t, J = 7.0 Hz, 1H, 1 -H), 6.65 (br s, 2H, NH2 ), 7.65 (s, 1H, 6-H), 8.10 (s, 1H, 2-H). Anal. calcd. for C11 H13 IN4 O3 : C, 35.12; H, 3.48; N, 14.89; found: C, 35.33; H, 3.60; N, 15.01.
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Prepare S.10 by cross-coupling of S.1d 18. In a 10-mL round-bottom flask equipped with a Teflon stir bar, load 100 mg (0.27 mmol) compound S.1d, 31 mg (0.027 mmol) Pd(PPh3 )4 , and 10.1 mg (0.05 mmol) CuI. After loading, purge flask with argon for 5 min at room temperature. 19. Add 3.0 mL anhydrous DMF, 54 mg (0.53 mmol) Et3 N, and 500 mg (5.31 mmol) cyclopentylacetylene under argon, and stir the resulting mixture 4 hr at room temperature under argon. 20. Evaporate the contents of the flask to dryness using a rotary evaporator. Redissolve the residue in 20 mL MeOH, adsorb on ∼2 g silica gel 60, and evaporate the solvent once more using a rotary evaporator. 21. Purify the crude product by flash chromatography (step 7) using a 3 × 50–cm column (bed height, 12 cm) and an elution gradient from 97:3 (v/v) CH2 Cl2 /MeOH (300 mL) to 90:10 (v/v) CH2 Cl2 /MeOH (300 mL). Collect 5-mL fractions. 22. Monitor fractions by TLC (step 8) using 9:1 (v/v) CH2 Cl2 /MeOH as the eluent. 23. Pool all product fractions, evaporate the solvent using a rotary evaporator, and dry the resulting sample overnight under high vacuum (<1 mmHg). 5-(2-Cyclopentylethynyl)-7-(2-deoxy-β-D-erythro-pentofuranosyl)-7H-pyrrolo[2,3d]pyrimidin-4-amine (S.10) is obtained as a colorless foam (59 mg, 64% yield). TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.46. UV (MeOH): λmax (ε) = 280 nm (10,500), 242 nm (13,900). 1 H NMR (DMSO-d6 ): δ 1.60-2.02 (several m, 8H, 4 × CH2 ), 2.18 (m, 1H, 2 -Hα ), 2.46 (m, 1H, 2 -Hβ ), 2.93 (quintet, J = 7.1 Hz, 1H, CH), 3.54 (m, 2H, 5 -H), 3.82 (m, 1H, 4 -H), 4.34 (m, 1H, 3 -H), 5.03 (t, J = 5.0 Hz, 1H, 5 -OH), 5.24 (d, J = 3.9 Hz, 1H, 3 -OH), 6.48 (t, J = 7.0 Hz, 1H, 1 -H), 6.70 (br s, 2H, NH2 ), 7.64 (s, 1H, 6-H), 8.11 (s, 1H, 2-H). Anal. calcd. for C18 H22 N4 O3 : C, 63.14; H, 6.48; N, 16.36; found: C, 62.74; H, 6.37; N, 16.77.
Prepare 4-methoxy precursor S.11 24. In a 50-mL round-bottom flask equipped with a Teflon stir bar, dissolve 1.26 g (2.0 mmol) compound S.9 in 20 mL of MeOH containing 0.5 M NaOMe. Stir overnight at room temperature. 25. Adsorb reaction mixture onto ∼4 g silica gel 60 and evaporate the solvent using a rotary evaporator. 26. Purify the crude product by flash chromatography (step 7; bed height, 20 cm) using an elution gradient from 95:5 (v/v) CH2 Cl2 /MeOH (600 mL) to 90:10 (v/v) CH2 Cl2 /MeOH (300 mL). Collect 12-mL fractions. 27. Monitor fractions by TLC (step 8) using 9:1 (v/v) CH2 Cl2 /MeOH as the eluent. 28. Pool all product fractions, evaporate the solvent using a rotary evaporator, and dry the resulting sample overnight under high vacuum (<1 mmHg). 7-(2-Deoxy-β-D-erythro-pentofuranosyl)-5-iodo-4-methoxy-7H-pyrrolo[2,3d]pyrimidine (S.11) is obtained as a colorless solid (700 mg, 90% yield). TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.60. UV (MeOH): λmax (ε) = 260 nm (9,400), 282 nm (10,800). 1 H NMR (DMSO-d6 ): δ 2.24 (m, 2H, 2 -H), 3.56 (m, 2H, 5 -H), 3.83 (m, 1H, 4 -H), 4.05 (s, 3H, MeO), 4.35 (m, 1H, 3 -H), 4.96 (t, J = 5.3 Hz, 1H, 5 -OH), 5.26 (d, J = 4.0 Hz, 1H, 3 -OH), 6.58 (t, J = 6.9 Hz, 1H, 1 -H), 7.84 (s, 1H, 6-H), 8.43 (s, 1H, 2-H). Anal. calcd. for C12 H14 IN3 O4 : C, 36.85; H, 3.61; N, 10.74; found: C, 36.70; H, 3.80; N, 10.60. Synthesis of Modified Nucleosides
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Prepare 2 -deoxyinosine analog S.5d 29. In a 100-mL round-bottom flask equipped with a Teflon stir bar, dissolve 1.2 g (3.07 mmol) compound S.11 in 20 mL of 2 M NaOH. 30. Stir reaction mixture 5 hr under reflux. 31. Cool reaction mixture to room temperature. 32. Precipitate product by acidifying with 1 M HCl. Collect precipitate by vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, wash with H2 O, and dry overnight under high vacuum (<1 mmHg). 7-(2-Deoxy-β-D-erythro-pentofuranosyl)-3,7-dihydro-5-iodo-4H-pyrrolo[2,3d]pyrimidin-4-one (S.5d) is obtained as a colorless solid (995 mg, 86% yield). TLC (CH2 Cl2 /MeOH, 4:1): Rf = 0.69. UV (MeOH): λmax (ε) = 265 nm (13,100), 282 nm (12,000). 1 H NMR (DMSO-d6 ): δ 2.20 (m, 1H, 2 -Hα ), 2.38 (m, 1H, 2 -Hβ ), 3.43 (m, 2H, 5 -H), 3.80 (m, 1H, 4 -H), 4.31 (m, 1H, 3 -H), 4.94 (t, J = 6.6 Hz, 1H, 5 -OH), 5.26 (d, J = 4.0 Hz, 1H, 3 -OH), 6.42 (t, J = 6.3 Hz, 1H, 1 -H), 7.53 (s, 1H, 6-H), 7.92 (s, 1H, 2-H), 12.01 (s, 1H, NH). Anal. calcd. for C11 H12 IN3 O4 : C, 35.03; H, 3.21; N, 11.14; found: C, 35.09; H, 3.43; N, 10.93. BASIC PROTOCOL 2
PREPARATION OF 7-IODO-7-DEAZAPURINE NUCLEOSIDES RELATED TO 2 -DEOXYGUANOSINE AND 2 -DEOXYXANTHOSINE In the synthesis of 7-iodo-7-deaza-2 -deoxyguanosine (S.2d) and the analog 7-iodo7-deaza-2 -deoxyxanthosine (S.6d), the 2-amino-protected nucleoside S.12 is used as starting material (Fig. 1.10.3). Regioselective iodination is performed on the protected nucleoside S.13. The protocols described for the preparation of nucleosides S.2d and S.6d give good yields, and the purification of intermediates is straightforward.
Materials
7-Substituted 7-Deazapurine (Pyrrolo[2,3d]pyrimidine) 2 -Deoxyribonucleosides
7-(2-Deoxy-β-D-erythro-pentofuranosyl)-2-(formylamino)-4-methoxy-7Hpyrrolo[2,3-d]pyrimidine (S.12; see Seela and Driller, 1989) Anhydrous acetonitrile (MeCN) Isobutyric anhydride Triethylamine Silica gel 60 (particle size, <0.063 mm; Merck) Anhydrous dichloromethane (CH2 Cl2 ) Acetone Cyclohexane N,N-Dimethylformamide (DMF) N-Iodosuccinimide 5% (w/v) sodium bicarbonate (NaHCO3 ) Anhydrous sodium sulfate (Na2 SO4 ) 2 M NaOH 1 M HCl Methanol (MeOH) Sodium methoxide (NaOMe) 10% (v/v) acetic acid (AcOH) in H2 O Sodium nitrite (NaNO2 ) Ethanol (EtOH) Sodium iodide (NaI) Chlorotrimethylsilane (Me3 SiCl) 0.1 M NaH2 PO4 buffer, pH 7.0 (see recipe) 25-, 50-, and 250-mL round-bottom flasks Rotary evaporator connected to a vacuum pump
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Figure 1.10.3 Preparation of 7-iodo-7-deaza-2 -deoxyguanosine (S.2d) and 7-iodo-7-deaza-2 -deoxyxanthosine (S.6d) is carried out by iodination of a precursor nucleoside. i-Bu2 O, isobutyric anhydride; DMF, N,N-dimethylformamide; NIS, N-iodosuccinimide.
4 × 50–cm chromatography columns 0.2-mm-thick silica gel 60 F254 aluminum TLC plates (Merck) 254-nm UV lamp 3-cm-diameter Buchner funnel with filter paper circles High-vacuum pump (final pressure, <1 mmHg) Separatory funnel 5-cm-diameter funnel with folded 10-cm-diameter Whatman no. 1 filter Reflux condenser Heating mantle with controller Additional reagents and equipment for flash chromatography (APPENDIX 3E) and thin-layer chromatography (TLC; APPENDIX 3D) Prepare common protected nucleoside S.13 1. In a 50-mL round-bottom flask equipped with a Teflon stir bar, suspend 1.0 g (3.24 mmol) 7-(2-deoxy-β-D-erythro-pentofuranosyl)-2-(formylamino)-4-methoxy7H-pyrrolo[2,3-d]pyrimidine (S.12) in 20 mL MeCN. 2. Add 5 mL (30.15 mmol) isobutyric anhydride and 2.3 mL (16.38 mmol) triethylamine and stir the reaction mixture overnight at room temperature to give a clear solution. 3. Evaporate the solution to dryness using a rotary evaporator connected to a vacuum pump. 4. Load the dry material onto a 4 × 50–cm column packed with silica gel 60 (bed height, 15 cm) and perform flash chromatography (APPENDIX 3E), eluting with 800 mL of 95:5 (v/v) CH2 Cl2 /acetone. Collect 12-mL fractions.
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5. Monitor fractions by TLC (APPENDIX 3D) on silica gel 60 F254 plates using 95:5 (v/v) CH2 Cl2 /acetone as the eluent. Visualize products under 254-nm light using a UV lamp. 6. Pool all product fractions and evaporate the solvent using a rotary evaporator. 7. Dissolve the residue in 8 mL CH2 Cl2 and precipitate the purified product by adding 80 mL cyclohexane. Collect the precipitate by vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, and dry overnight under high vacuum (<1 mmHg). 7-[2-Deoxy-3,5-bis-O-(2-methylpropanoyl)-β-D-erythro-pentofuranosyl]-2-(formyl amino)-4-methoxy-7H-pyrrolo[2,3-d]pyrimidine (S.13) is obtained as a colorless solid (1.2 g, 82% yield). m.p. 136◦ -137◦ C. TLC (CH2 Cl2 /acetone, 95:5): Rf = 0.4. 1 H NMR (DMSO-d6 ): δ 1.09 (d, J = 7.0 Hz, 6H, 2 × CH3 ), 1.15 (d, J = 7.0 Hz, 6H, 2 × CH3 ), 2.47, 2.57, 2.62, 2.92 (4 × m, 4H, 2 × CH, 2 × 2 -H), 4.04 (s, 3H, OCH3 ), 4.18 (m, 2H, 5 -H), 4.27 (m, 1H, 4 -H), 5.37 (m, 1H, 3 -H), 6.49 (m, 1H, 1 -H), 6.55 (d, J = 3.7 Hz, 1H, 5 -H), 7.42 (d, J = 3.7 Hz, 1H, 6-H), 9.46 (d, J = 9.9 Hz, 1H, NH), 10.75 (d, J = 9.9 Hz, 1H, HCO). Anal. calcd. for C21 H28 N4 O7 : C, 56.24; H, 6.29; N, 12.49; found: C, 56.44; H, 6.26; N, 12.44.
Perform iodination (prepare S.14) 8. In a 25-mL round-bottom flask equipped with a Teflon stir bar, prepare a solution of 500 mg (1.12 mmol) compound S.13 in 5 mL DMF. 9. Add 264 mg (1.17 mmol) N-iodosuccinimide and stir 23 hr at room temperature. 10. Dilute the reaction mixture with 40 mL CH2 Cl2 and pour into a separatory funnel containing 10 mL of 5% NaHCO3 . Wash the organic layer with 20 mL H2 O. 11. Collect the organic layer and dry over 3 g anhydrous Na2 SO4 . Filter off the Na2 SO4 by gravity filtration through a folded 10-cm-diameter filter. Evaporate the filtrate to dryness using a rotary evaporator. 12. Purify the crude product by flash chromatography (step 4) using 600 mL of 95:5 (v/v) CH2 Cl2 /acetone as the eluent. Collect 12-mL fractions. 13. Monitor fractions by TLC (step 5) using 95:5 (v/v) CH2 Cl2 /acetone as the eluent. 14. Pool all product fractions and evaporate to dryness using a rotary evaporator. 15. Dissolve the residue in 4 mL CH2 Cl2 . Crystallize the purified product by adding 40 mL cyclohexane, collect the crystals by vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, and dry overnight under high vacuum. 7-[2-Deoxy-3,5-bis-O-(2-methylpropanoyl)-β-D-erythro-pentofuranosyl]-2-(formyl amino)-5-iodo-4-methoxy-7H-pyrrolo[2,3-d]pyrimidine (S.14) is obtained as colorless crystals (590 mg, 92% yield). m.p. 129◦ -130◦ C. TLC (CH2 Cl2 /acetone, 95:5): Rf = 0.4. 1 H NMR (DMSO-d6 ): δ 1.09 (d, J = 7.0 Hz, 6H, 2 × CH3 ), 1.15 (d, J = 7.0 Hz, 6H, 2 × CH3 ), 2.46, 2.60, 2.89 (3 × m, 4H, 2 × CH, 2 × 2 -H), 4.06 (s, 3H, OCH3 ), 4.17 (m, 2H, 5 -H), 4.27 (m, 1H, 4 -H), 5.35 (m, 1H, 3 -H), 6.46 (m, 1H, 1 -H), 7.62 (s, 1H, 6-H), 9.55 (d, J = 9.7 Hz, 1H, NH), 10.81 (d, J = 9.9 Hz, 1H, HCO). Anal. calcd. for C21 H27 IN4 O7 : C, 43.91; H, 4.74; N, 9.75; found: C, 43.98; H, 4.75; N, 9.82.
7-Substituted 7-Deazapurine (Pyrrolo[2,3d]pyrimidine) 2 -Deoxyribonucleosides
Prepare 2 -deoxyguanosine analog S.2d 16. Convert 200 mg (0.35 mmol) compound S.14 to nucleoside S.2d using the general procedure described for converting S.11 to nucleoside S.5d (see Basic Protocol 1, steps 29 to 32). 2-Amino-7-(2-deoxy-β-D-erythro-pentofuranosyl)-3,7-dihydro-5-iodo-4H-pyrrolo[2,3d]pyrimidin-4-one (S.2d) is obtained as colorless crystals (126 mg, 92% yield). m.p. 218◦ -220◦ C. TLC (CH2 Cl2 /MeOH, 4:1): Rf = 0.7. UV (MeOH): λmax (ε) = 266 nm
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(12,000), 285 nm (sh 8,400). 1 H NMR (DMSO-d6 ): δ 2.08 (m, 1H, 2 -Hα ), 2.30 (m, 1H, 2 -Hβ ), 3.48 (m, 2H, 5 -H), 3.74 (m, 1H, 4 -H), 4.26 (m, 1H, 3 -H), 4.89 (t, J = 5.7 Hz, 1H, 5 -OH), 5.18 (d, J = 4.3 Hz, 1H, 3 -OH), 6.25 (t, J = 6.5 Hz, 1H, 1 -H), 6.34 (br s, 2H, NH2 ), 7.09 (s, 1H, 6-H), 10.51 (br s, 1H, NH). Anal. calcd. for C11 H13 IN4 O4 : C, 33.69; H, 3.34; N, 14.29; found: C, 33.78; H, 3.42; N, 14.29.
Prepare deprotected iodo precursor S.15 17. In a 250-mL round-bottom flask equipped with a Teflon stir bar, dissolve 2.1 g (3.66 mmol) compound S.14 in 150 mL of MeOH containing 0.5 M NaOMe. Stir overnight at room temperature. 18. Adsorb reaction mixture onto ∼6 g silica gel 60 and evaporate the solvent using a rotary evaporator. 19. Purify the crude product by flash chromatography (step 4; bed height, 10 cm) using an elution gradient from 95:5 (v/v) CH2 Cl2 /MeOH (800 mL) to 90:10 (v/v) CH2 Cl2 /MeOH (300 mL). Collect 15-mL fractions. 20. Monitor fractions by TLC (step 5) using 90:10 (v/v) CH2 Cl2 /MeOH as the eluent. 21. Pool all product fractions, evaporate the solvent using a rotary evaporator, and dry the resulting sample overnight under high vacuum. 2-Amino-7-(2-deoxy-β-D-erythro-pentofuranosyl)-5-iodo-4-methoxy-7H-pyrrolo[2,3d]pyrimidine (S.15) is obtained as a colorless solid (1.17 g, 79% yield). m.p. 159◦ -160◦ C. TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.48. UV (MeOH): λmax (ε) = 233 nm (31,200), 266 nm (2,800), 289 nm (7,300). 1 H NMR (DMSO-d6 ): δ 2.08 (m, 1H, 2 -Hα ), 2.36 (m, 1H, 2 -Hβ ), 3.48 (m, 2H, 5 -H), 3.75 (m, 1H, 4 -H), 3.92 (s, 3H, MeO), 4.27 (m, 1H, 3 -H), 4.94 (t, J = 5.4 Hz, 1H, 5 -OH), 5.22 (d, J = 3.7 Hz, 1H, 3 -OH), 6.37 (t, J = 5.8 Hz, 1H, 1 -H), 6.39 (s, 2H, NH2 ), 7.29 (s, 1H, 6-H). Anal. calcd. for C12 H15 IN4 O4 : C, 35.48; H, 3.72; I, 31.24; N, 13.79; found: C, 35.62; H, 3.84; I, 30.72; N, 13.84.
Perform deamination (prepare S.16) 22. In a 250-mL round-bottom flask equipped with a Teflon stir bar, dissolve 1.0 g (2.46 mmol) compound S.15 in 150 mL of 10% AcOH. 23. Add a solution of 290 mg (4.2 mmol) NaNO2 in 10 mL H2 O dropwise and stir 30 min at room temperature. 24. Adsorb reaction mixture onto ∼4 g silica gel 60 and evaporate the solvent using a rotary evaporator. 25. Purify the crude product by flash chromatography (step 4; bed height, 10 cm), eluting with 900 mL of 95:5 (v/v) CH2 Cl2 /MeOH. Collect 12-mL fractions. 26. Monitor fractions by TLC (step 5) using 90:10 (v/v) CH2 Cl2 /MeOH as the eluent. 27. Pool all product fractions and evaporate the solvent using a rotary evaporator. 28. Crystallize purified product from EtOH, collect the crystals by vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, and dry overnight under high vacuum (<1 mmHg). 7-(2-Deoxy-β-D-erythro-pentofuranosyl)-1,7-dihydro-5-iodo-4-methoxy-2Hpyrrolo[2,3-d]pyrimidin-2-one (S.16) is obtained as colorless crystals (654 mg, 65% yield). m.p. 158◦ -160◦ C. TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.40. UV (MeOH): λmax (ε) = 227 nm (23,200), 286 nm (6,300). 1 H NMR (DMSO-d6 ): δ 2.13 (m, 1H, 2 -Hα ), 2.39 (m, 1H, 2 -Hβ ), 3.52 (m, 2H, 5 -H), 3.79 (m, 1H, 4 -H), 3.96 (s, 3H, MeO), 4.30 (m, 1H, 3 -H), 5.04 (br s, 1H, 3 -OH), 5.25 (t, J = 3.7 Hz, 1H, 5 -OH), 6.37 (dd, J = 6.6 Hz, J = 7.2 Hz, 1H, 1 -H), 7.46 (s, 1H, 6-H), 11.57 (br s, 1H, NH). Anal. calcd. for C12 H14 IN3 O5 : C, 35.40; H, 3.47; I, 31.17; N, 10.32; found: C, 35.48; H, 3.55; I, 31.20; N, 10.34.
Synthesis of Modified Nucleosides
1.10.9 Current Protocols in Nucleic Acid Chemistry
Supplement 21
Prepare 2 -deoxyxanthosine analog S.6d 29. In a 50-mL round-bottom flask equipped with a Teflon stir bar, suspend 620 mg (1.53 mmol) compound S.16 and 450 mg (3.00 mmol) NaI in 20 mL MeCN. 30. Add 442 µL (3.48 mmol) Me3 SiCl and stir 1 hr at room temperature. 31. Subject the reaction mixture to vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, wash the solid with 5 mL MeCN, and collect the filter cake. 32. Crystallize purified product from 10 mL H2 O, collect the crystals by vacuum filtration, and dry overnight under high vacuum (<1 mmHg). 7-(2-Deoxy-β-D-erythro-pentofuranosyl)-1,3,7-trihydro-5-iodo-2H,4H-pyrrolo[2,3d]pyrimidin-2,4-dione (S.6d) is obtained as colorless crystals (543 mg, 90% yield). m.p. >210◦ C. TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.36. UV (0.1 M NaH2 PO4 buffer, pH 7.0): λmax (ε) = 224 nm (21,500), 259 nm (9,700), 285 nm (6,800). 1 H NMR (DMSO-d6 ): δ 2.16 (m, 1H, 2 -Hα ), 2.27 (m, 1H, 2 -Hβ ), 3.57 (m, 2H, 5 -H), 3.82 (m, 1H, 4 -H), 4.28 (m, 1H, 3 -H), 5.28 (d, J = 3.3 Hz, 1H, 3 -OH), 5.53 (br s, 1H, 5 -OH), 6.12 (dd, J = 7.0 Hz, J = 6.5 Hz, 1H, 1 -H), 7.14 (s, 1H, 6-H), 10.70 (s, 1H, NH), 11.70 (s, 1H, NH). Anal. calcd. for C11 H12 IN3 O5 : C, 33.61; H, 3.08; I, 32.28; N, 10.69; found: C, 34.14; H, 3.34; I, 31.56; N, 10.46.
BASIC PROTOCOL 3
PREPARATION OF 7-BROMO-7-DEAZA-2 -DEOXYISOGUANOSINE This protocol describes the preparation of 2-amino-7-bromo-7-deaza-2 -deoxyadenosine (S.3c) and its subsequent oxidation to give 7-bromo-7-deaza-2 -deoxyisoguanosine (S.4c; Fig. 1.10.4). Regioselective bromination is performed on the 2-amino-protected nucleobase S.18. Convergent nucleoside synthesis is accomplished by nucleobase anion glycosylation. The procedure described here allows the successful preparation of the desired nucleoside, S.4c. The chemical reactions give good yields, and the purification of intermediates is straightforward.
Materials
7-Substituted 7-Deazapurine (Pyrrolo[2,3d]pyrimidine) 2 -Deoxyribonucleosides
2-Amino-4-chloro-7H-pyrrolo[2,3-d]pyrimidine (S.17; see Seela et al., 1987) Anhydrous pyridine Pivaloyl chloride Potassium hydroxide (KOH) powder (purity, ≥85%; Sigma) Anhydrous dichloromethane (CH2 Cl2 ) N-Bromosuccinimide Methanol (MeOH) Anhydrous acetonitrile (MeCN) Tris[2-(2-methoxyethoxy)ethyl]amine (TDA-1) 2-Deoxy-3,5-di-O-(p-toluoyl)-α-D-erythro-pentofuranosyl chloride (S.8; see Hoffer, 1960; Rolland et al., 1997) Silica gel 60 (particle size, <0.063 mm; Merck) Petroleum ether (b.p. range, 40◦ to 80◦ C) 1,4-Dioxane 25% (v/v) ammonium hydroxide (NH4 OH) 1:5 (v/v) acetic acid (AcOH)/H2 O Sodium nitrite (NaNO2 ) Isopropanol (i-PrOH) 100- and 250-mL round-bottom flasks Rotary evaporator connected to a vacuum pump
1.10.10 Supplement 21
Current Protocols in Nucleic Acid Chemistry
Figure 1.10.4 Preparation of 7-bromo-7-deaza-2 -deoxyisoguanosine (S.4c) is carried out by glycosylation of a brominated nucleobase followed by amination at C4 and selective deamination at C2 . NBS, N-bromosuccinimide; Piv, pivaloyl; TDA-1, tris[2-(2-methoxyethoxy)ethyl]amine; Tol, p-toluoyl. For structure of S.8, see Figure 1.10.2.
3-cm-diameter Buchner funnel with filter paper circles High-vacuum pump (final pressure, <1 mmHg) 5 × 50–cm chromatography columns 0.2-mm-thick silica gel 60 F254 aluminum TLC plates (Merck) 254-nm UV lamp Steel bomb (autoclave) 5 × 20–cm Serdolit AD-4 column (resin particle size, 0.1 to 0.2 mm; Serva) Additional reagents and equipment for flash chromatography (APPENDIX 3E) and thin-layer chromatography (TLC; APPENDIX 3D) Protect exocyclic amine (prepare S.18) 1. In a 100-mL round-bottom flask equipped with a Teflon stir bar, prepare a solution of 8.70 g (51.61 mmol) 2-amino-4-chloro-7H-pyrrolo[2,3-d]pyrimidine (S.17) in 40 mL anhydrous pyridine. 2. Add 8.0 mL (64.95 mmol) pivaloyl chloride dropwise, with stirring, over the course of 15 min, and then stir the reaction mixture 5 hr at room temperature. 3. Evaporate the reaction mixture to near-dryness using a rotary evaporator equipped with a vacuum pump. 4. Remove traces of pyridine by performing two rounds of coevaporation with 10 mL water. 5. Add 100 mL water to the reaction flask. Collect the resulting solid by vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, wash with three 80-mL aliquots of cold water, and dry in vacuo over KOH. 4-Chloro-2-pivaloylamino-7H-pyrrolo[2,3-d]pyrimidine (S.18) is obtained as a reddish solid (10.9 g, 84% yield). m.p. 226◦ C. TLC (CH2 Cl2 /MeOH, 95:5): Rf = 0.52. 1 H NMR (DMSO-d6 ): δ 1.24 (s, 9H, 3 × CH3 ), 6.53 (m, 1H, 5-H), 7.63 (m, 1H, 6-H), 10.06 (s, 1H, CONH), 12.34 (s, 1H, NH). Synthesis of Modified Nucleosides
1.10.11 Current Protocols in Nucleic Acid Chemistry
Supplement 21
Perform bromination (prepare S.19) 6. In a 250-mL round-bottom flask equipped with a Teflon stir bar, prepare a solution of 5.7 g (22.56 mmol) compound S.18 in 100 mL CH2 Cl2 . 7. Add 4.82 g (27.08 mmol) N-bromosuccinimide in four portions over 10 min and stir the resulting mixture 5 hr at room temperature. 8. Using a rotary evaporator, evaporate the yellow solution to yield an amber residue. 9. Isolate the purified product in the following way. a. Crystallize product from 200 mL MeOH, collect the crystals by vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, and wash crystals twice with 20-mL aliquots of prechilled MeOH. b. Condense filtrate to 80 mL and refrigerate at 5◦ C overnight. Collect the resulting crystals by vacuum filtration, and wash crystals twice with 5-mL aliquots of prechilled MeOH. c. Combine crystals from substeps a and b and dry under high vacuum (<1 mmHg). 5-Bromo-4-chloro-2-pivaloylamino-7H-pyrrolo[2,3-d]pyrimidine (S.19) is obtained as pink crystals (6.0 g, 80% yield). m.p. 204◦ C (dec.). TLC (CH2 Cl2 / MeOH, 95:5): Rf = 0.58. 1 H NMR (DMSO-d6 ): δ 1.23 (s, 9H, 3 × CH3 ), 7.78 (s, 1H, 6-H), 10.16 (s, 1H, CONH), 12.72 (s, 1H, NH). Anal. calcd. for C11 H12 BrClN4 O: C, 39.84; H, 3.65; N, 16.90; found: C, 39.87; H, 3.63; N, 16.90.
Prepare nucleoside precursor S.20 10. In a 100-mL round-bottom flask equipped with a Teflon stir bar, suspend 1.15 g (17.42 mmol) powdered KOH in 60 mL MeCN. 11. Add 0.2 mL (0.63 mmol) TDA-1 and stir the resulting mixture 5 min at room temperature. 12. Under constant stirring, add 1.66 g (5.01 mmol) compound S.19 in three equal portions over the course of 15 min. After addition is complete, stir the reaction mixture for another 15 min at room temperature. 13. Add 2.53 g (6.51 mmol) 2-deoxy-3,5-di-O-(p-toluoyl)-α-D-erythro-pentofuranosyl chloride (S.8) in four portions over 15 min and stir the resulting mixture 30 min at room temperature. 14. Remove the insoluble material by vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, washing with three 20-mL aliquots of MeCN. Evaporate the filtrate to dryness using a rotary evaporator. 15. Load the crude product onto a 5 × 50–cm column packed with silica gel 60 (bed height, 15 cm). Perform flash chromatography (APPENDIX 3E) using an elution gradient from 100% CH2 Cl2 (800 mL) to 99:1 (v/v) CH2 Cl2 /MeOH (400 mL). Collect 15-mL fractions. 16. Monitor fractions by TLC (APPENDIX 3D) on silica gel 60 F254 plates using 99:1 (v/v) CH2 Cl2 /MeOH as the eluent. Visualize products under 254-nm light using a UV lamp. 17. Pool all product fractions and evaporate the solvent using a rotary evaporator.
7-Substituted 7-Deazapurine (Pyrrolo[2,3d]pyrimidine) 2 -Deoxyribonucleosides
18. Dissolve the residue in 20 mL CH2 Cl2 . Crystallize the purified product by adding 100 mL petroleum ether, store 10 hr at 5◦ C, collect the crystals by vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, and dry overnight under high vacuum (<1 mmHg).
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Current Protocols in Nucleic Acid Chemistry
5-Bromo-4-chloro-7-[2-deoxy-3,5-di-O-(p-toluoyl)-β-D-erythro-pentofuranosyl]-2pivaloylamino-7H-pyrrolo[2,3-d]pyrimidine (S.20) is obtained as colorless needles (2.32 g, 68% yield). m.p. 229◦ -230◦ C. TLC (CH2 Cl2 /MeOH, 99:1): Rf = 0.53. UV (MeOH): λmax (ε) = 245 nm (55,300). 1 H NMR (CDCl3 ): δ 1.38-1.47 (m, 9H, 3 × CH3 ), 2.55 (s, 3H, CH3 ), 2.57 (s, 3H, CH3 ), 2.91-2.93 (m, 1H, 2 -H), 3.04-3.08 (m, 1H, 2 -H), 4.69-4.70, 4.76-4.78, and 4.85-4.87 (3 × m, 3H, 4 -H, 5 -H), 5.88-5.91 (m, 1H, 3 -H), 6.84 (t, J = 6.7 Hz, 1H, 1 -H), 7.37, 7.43, 8.03, and 8.10 (4 × d, J = 8.1 Hz, 8H, 2 × C6 H4 ), 8.28 (s, 1H, 6-H), 10.29 (s, 1H, NH). Anal. calcd. for C32 H32 BrClN4 O6 : C, 56.19; H, 4.72; N, 8.19; found: C, 56.12; H, 4.79; N, 8.33.
Prepare 2-amino-2 -deoxyadenosine analog S.3c 19. Convert 1.5 g (2.19 mmol) nucleoside S.20 to compound S.3c using the general procedure described for converting S.9 to nucleoside S.1d (see Basic Protocol 1, steps 11 to 17), but stir the reaction 24 hr at 120◦ C after adding NH4 OH. 5-Bromo-7-(2-deoxy-β-D-erythro-pentofuranosyl)-7H-pyrrolo[2,3-d]pyrimidin-2,4diamine (S.3c) is obtained as colorless crystals (667 mg, 88% yield). m.p. 208◦ C (dec.). TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.33. UV (MeOH): λmax (ε) = 228 nm (27,800), 268 nm (9,300), 288 nm (sh 7,500). 1 H NMR (DMSO-d6 ): δ 2.00-2.06 and 2.29-2.40 (2 × m, 2H, 2 -H), 3.48-3.52 (m, 2H, 5 -H), 3.74-3.78 (m, 1H, 4 -H), 4.25-4.29 (m, 1H, 3 -H), 4.99 (t, J = 5.4 Hz, 1H, 5 -OH), 5.21 (d, J = 3.6 Hz, 1H, 3 -OH), 5.86 (br s, 2H, NH2 ), 6.28 (br s, 2H, NH2 ), 6.34 (dd, J = 6.5 Hz, J = 6.9 Hz, 1H, 1 -H), 7.15 (s, 1H, 6-H). Anal. calcd. for C11 H14 BrN5 O3 : C, 38.39; H, 4.10; N, 20.35; found: C, 38.22; H, 4.08; N, 20.25.
Prepare 2 -deoxyisoguanosine analog S.4c 20. In a 100-mL round-bottom flask equipped with a Teflon stir bar, dissolve 500 mg (1.45 mmol) nucleoside S.3c in 40.0 mL of 1:5 (v/v) AcOH/H2 O. 21. Add a solution of 250 mg (3.62 mmol) NaNO2 in 8.0 mL H2 O dropwise, with stirring, to the reaction flask. Stir the reaction mixture for an additional 30 min at room temperature after all of the solution has been added. 22. Adjust the pH of the dark solution to 7.0 with 25% NH4 OH. 23. Purify the crude product by loading it onto a 5 × 20–cm Serdolit AD-4 column (resin particle size, 0.1 to 0.2 mm) and performing flash chromatography (APPENDIX 3E). Elute with 300 mL H2 O followed by 500 mL of 95:5 (v/v) H2 O/i-PrOH and 400 mL of 90:10 (v/v) H2 O/i-PrOH. Collect 15-mL fractions. 24. Monitor fractions by TLC (step 16) using 5:1 (v/v) CH2 Cl2 /MeOH as the eluent. 25. Pool all product fractions and use a rotary evaporator to reduce the volume to 20% of the original pooled volume. Collect the resulting crystals by vacuum filtration through a 3-cm Buchner funnel equipped with a filter paper circle, and dry under high vacuum (<1 mmHg). 4-Amino-5-bromo-7-(2-deoxy-β-D-erythro-pentofuranosyl)-3,7-dihydro-2H pyrrolo[2,3d]pyrimidin-2-one (S.4c) is obtained as yellowish crystals (325 mg, 65% yield). m.p. 220◦ C (dec.). TLC (25% NH4 OH/i-PrOH/H2 O, 1:7:2): Rf = 0.9. TLC (CH2 Cl2 /MeOH, 5:1): Rf = 0.28 (tailing). UV (MeOH): λmax (ε) = 232 nm (26,800), 262 nm (5,900), 310 nm (5,800). 1 H NMR (DMSO-d6 ): δ = 2.00-2.08 and 2.25-2.33 (2 × m, 2H, 2 -H), 3.503.51 (m, 2H, 5 -H), 3.74-3.77 (m, 1H, 4 -H), 4.25-4.28 (m, 1H, 3 -H), 5.04-5.06 (m, 1H, 5 -OH), 5.23 (d, J = 3.9 Hz, 1H, 3 -OH), 6.25 (dd, J = 5.8 Hz, J = 6.5 Hz, 1H, 1 -H), 6.85 (br s, 2H, NH2 ), 7.21 (s, 1H, 6-H), 10.77 (br s, 1H, NH). Anal. calcd. for C11 H13 BrN4 O4 : C, 38.28; H, 3.80; N, 16.23; found: C, 38.54; H, 3.72; N, 16.45.
Synthesis of Modified Nucleosides
1.10.13 Current Protocols in Nucleic Acid Chemistry
Supplement 21
BASIC PROTOCOL 4
DETERMINATION OF pK a VALUES OF NUCLEOSIDES This protocol describes the determination of nucleoside pKa values by spectrophotometric titration (pH 1.5 to 13.5; Albert and Serjeant, 1971) at 220 to 350 nm. As a representative example, the pKa measurement of compound S.4c is described. The pKa values of several 7-deazapurine nucleosides are summarized in Table 1.10.1. From Table 1.10.1, it is apparent that the 7-halogenated derivatives of 2-amino-7-deaza2 -deoxyadenosine (S.3b-d) have very similar pKa values (4.8 to 4.9), which are lower than that of the parent nucleoside S.3a (pKa = 5.7). This is because electron-withdrawing halogens reduce the basicity of the 7-deazaadenine moiety. Similar results were observed for the 7-deaza-2 -deoxyxanthosines (pKa = 6.7, 6.0, and 6.1 for S.6a, S.6c, and S.6d, respectively). Given their pKa values, deprotonation of S.6a-d occurs under almost neutral conditions, with 7-halogen substituents further enhancing the acidity of these compounds. In the case of the 7-deazaguanine derivatives S.2a and S.2d and the 7-deazaisoguanine analogs S.4a-c, two pKa values are observed. The lower value corresponds to protonation of the nucleobase, whereas the higher one corresponds to the removal of a proton from the 3 position (systematic numbering). The introduction of 7-halogen substituents leads to decreased pKa values (pKa = 3.9 and 9.7 for S.4b and 3.8 and 9.8 for S.4c, compared with 4.6 and 10.5 for the parent nucleoside S.4a). These differences reflect the increased acidity of 7-halogenated 7-deaza-2 -deoxyisoguanosines.
Materials 4-Amino-5-bromo-7-(2-deoxy-β-D-erythro-pentofuranosyl)-3,7-dihydro-2Hpyrrolo[2,3-d]pyrimidin-2-one (S.4c; see Basic Protocol 3) Phosphate buffer solution, pH 4.5 (see recipe) 1 M H3 PO4 3 M NaOH 100-mL volumetric flask 100-mL beaker pH meter UV-Vis spectrophotometer 1. In a 100-mL volumetric flask, dissolve 2.0 mg S.4c in 100 mL phosphate buffer solution, pH 4.5.
Table 1.10.1 pKa Values of Selected 7-Deazapurine 2 -Deoxyribonucleosidesa
Wavelength (nm)b
pKa
Compound
Wavelength (nm)b
pKa
290
1.7
S.4a
255
4.6
295
10.2
275
10.5
S.2d
300
10.0 and <1.5
S.4b
264
3.9 and 9.7
S.3a
282
5.7
S.4c
264
3.8 and 9.8
S.3b
284
4.9
S.6a
250
6.7
S.3c
286
4.9
S.6c
260
6.0
S.3d
288
4.8
S.6d
260
6.1
Compound S.2a
7-Substituted 7-Deazapurine (Pyrrolo[2,3d]pyrimidine) 2 -Deoxyribonucleosides
a Measured in phosphate buffer solution (see recipe) over a pH range of 1.5 to 12.0. b Wavelength used for pK determination (i.e., wavelength at which the absorbance changed most significantly as a function a
of pH).
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2. Transfer a 50-mL aliquot of the S.4c solution to a 100-mL beaker. Vary the pH of the aliquot from 4.5 to 1.5 in steps of 0.5 pH units by adding 1 M H3 PO4 to the beaker in dropwise fashion. At each pH level, obtain a UV absorption spectrum from 220 to 350 nm. 3. Transfer another 50-mL aliquot of the S.4c solution to a clean 100-mL beaker. Vary the pH of the aliquot from 4.5 to 13.5 in steps of 0.5 pH units by adding 3 M NaOH to the beaker in dropwise fashion. At each pH level, obtain a UV absorption spectrum from 220 to 350 nm. 4. Collect all absorbance data obtained at 264 nm (i.e., the wavelength at which the most significant change in UV absorbance as a function of pH is observed) and plot a profile of absorbance versus pH (origin, pH 7.0). 5. Calculate the first derivative of absorbance with respect to pH, dA/d(pH), at each point on the plot. Define the pKa of the nucleoside as the pH that gives a maximal value for dA/d(pH).
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
NaH2 PO4 buffer, 0.1 M, pH 7.0 Dissolve 7.9 g NaH2 PO4 ·H2 O in 500 mL H2 O Neutralize with 1 M NaOH to pH 7.0 Store tightly capped for up to 4 months at room temperature Phosphate buffer solution, pH 4.5 Dissolve 7.8 g NaH2 PO4 ·H2 O in 500 mL H2 O Stir 2 hr at room temperature Store tightly capped for up to 4 months at room temperature COMMENTARY Background Information A series of naturally occurring 7deazapurines (pyrrolo[2,3-d]pyrimidines) have been isolated from natural sources. Among them are monomeric, naturally occurring nucleosides such as tubercidin and its 7-substituted derivatives (Suhadolnik, 1970). Other such nucleosides are found as metabolites or antimetabolites of marine organisms, and some of them carry halogens at the 7 position of the 7-deazapurine moiety. 7-Deazapurine ribonucleosides (e.g., queuosine, archaeosine) are found as constituents of naturally occurring nucleic acids such as tRNA (Kasai et al., 1975; Kilpatrick and Walker, 1982). Several naturally occurring 7-deazapurine nucleosides or chemically designed analogs exhibit antiviral and/or anticancer activity (Martin, 1989; Simons, 2001). As the shape of the 7-deazapurine nucleosides closely resembles that of the
purine nucleosides, these molecules can substitute for the canonical constituents of DNA and RNA. They have also been used in their triphosphate forms for nucleic acid labeling or sequencing (Mizusawa et al., 1986; Prober et al., 1987). The frequent natural occurrence and unusual biological properties of this class of compounds have prompted ample studies directed toward their synthesis, toward analysis of their biological activity, and toward their incorporation into oligonucleotides (Revankar and Robins, 1991). In an effort to investigate the association of these compounds with stabilizing effects on oligonucleotides or with antiviral activity, a number of 7-deazapurine nucleosides have been synthesized. These compounds became synthetically accessible with the development of stereoselective nucleobase anion glycosylation (Winkeler and Seela, 1983;
Synthesis of Modified Nucleosides
1.10.15 Current Protocols in Nucleic Acid Chemistry
Supplement 21
7-Substituted 7-Deazapurine (Pyrrolo[2,3d]pyrimidine) 2 -Deoxyribonucleosides
Seela et al., 1988). A number of 7-deazapurine 2 -deoxyribonucleosides have been synthesized by this method. A similar method was later applied to other base-modified nucleosides (Kazimierczuk et al., 1984), many of which have been incorporated into oligonucleotides. Substituents—in particular, halogen or alkynyl substituents—are well accommodated in the major groove of duplex DNA. Some nucleosides have the potential to be used for further chemical and functional manipulations. Detailed investigations have been performed to study the selective halogenation of 7-deazapurines at C7. In the case of 7-deazapurines containing a 2-amino group, halogenation can also be directed to C8 or to both C7 and C8, yielding 7,8-dihalogenated products. In the absence of a 2-amino group, 7-halogenated 7-deazapurines are formed. It was eventually found that regioselective 7-halogenation with N-halosuccinimides can be accomplished when the 2-amino group is protected (Ramzaeva and Seela, 1995; Balow et al., 1998; Seela and Peng, 2004). From 7iodo derivatives, the 7-alkynyl-7-deazapurine nucleosides are prepared using the palladiumcatalyzed Sonogashira cross-coupling reaction (Seela and Zulauf, 1996). The reaction is performed under argon in anhydrous DMF with tetrakis(triphenylphosphine)palladium(0), copper(I) iodide, and triethylamine. The synthesis of the 7-halogenated derivatives S.1-6 via nucleobase anion glycosylation uses well-established chemistry. Glycosylation of the nucleobases is performed in acetonitrile with 2-deoxy-3,5-di-O-(ptoluoyl)-α-D-erythro-pentofuranosyl chloride (S.8). The glycosylation reaction is stereoselective and gives the β-D anomer exclusively in most cases. Deprotection, amination, deamination, and hydroxylation of 7-deazapurine nucleosides produce close-to-quantitative yields. Two general routes are employed for the synthesis of 7-halogenated 7-deazapurine nucleosides: (1) conversion of a preformed nucleoside precursor to a 7-halogenated nucleoside (see Basic Protocol 2) and (2) halogenation of the nucleobase followed by glycosylation (see Basic Protocol 1 and Basic Protocol 3). Though this unit presents only representative halogenated (i.e., iodo and bromo) derivatives in each protocol, the modifications for preparation of other halogenated analogs (bromo, iodo, chloro, and fluoro) are straightforward, involving primarily the substitution of the appropriate halogenated
reagents. For additional details, refer to the references provided in Figure 1.10.1. The pKa values of the nucleobases reflect the strength of the hydrogen bonds they form in double-stranded DNA, which is an important factor in assessing the biophysical properties of nucleosides. The authors have reported pKa values for a series of 7-modified 7-deazapurine nucleosides. A method for determining pKa values is presented in Basic Protocol 4. A few 7-deazapurine 2 -deoxyribonucleosides show interesting photophysical properties. Among these are the 2-amino and 2-hydroxy derivatives S.21 and S.22 (Fig. 1.10.5), which exhibit strong fluorescence (Seela and Steker, 1984; Seela and Engelke, 1985; Seela et al., 1994). In contrast, 7-deaza-2 -deoxyguanosine (S.2a) and 7-deaza-2 -deoxyisoguanosine (S.4a) serve as efficient fluorescence quenchers (Latimer and Lee, 1991; Li et al., 2004). The fluorescence emission maximum of S.21 is observed at 395 nm (excitation 311 nm), and a quantum yield (φf ) of 0.47 has been determined (Seela and Becher, 2000). Replacement of the amino group with a hydroxyl group (which can take part in keto-enol tautomerization, S.22) results in a bathochromic shift in absorption (331 nm) and emission (444 nm), as well as a decreased fluorescence quantum yield (φf = 0.22; Seela et al., 1998). The strong fluorescence of S.21 is quenched to approximately 1/10 of its original intensity when it is incorporated into a dinucleotide (Seela and Engelke, 1985). This quenching is a result of the strong stacking interactions between nucleobases. When incorporated into oligonucleotides, S.2a and S.4a quench the fluorescence of ethidium bromide (Latimer and Lee, 1991; Li et al., 2004). Normally, the fluorescence of ethidium bromide is enhanced strongly when it intercalates into duplex DNA (LePecq and Paoletti, 1967). However, when S.2a or S.4a is incorporated into duplex DNA in place of deoxyguanosine or deoxyisoguanosine, the fluorescence of the dye is totally quenched (Li et al., 2004). This quenching results from a photoinduced electron transfer from 7-deaza2 -deoxyguanosine to ethidium bromide (Kelley and Barton, 1998).
Compound Characterization 1
H and 13 C NMR spectra were measured for all analogs using AC-250 and AMX500 spectrometers (Bruker); δ values are given in ppm downfield from internal tetramethylsilane (SiMe4 ). The signals are assigned
1.10.16 Supplement 21
Current Protocols in Nucleic Acid Chemistry
Figure 1.10.5 properties.
Structures of 7-deazapurine 2 -deoxyribonucleosides with interesting fluorescence
unambiguously. UV spectra were measured using a U-3200 spectrometer (Hitachi), and extinction coefficients were determined (λmax in nm, ε in M−1 ·cm−1 ). Melting points were measured using a Linstr¨om apparatus. Elemental analyses were performed in all cases by Mikroanalytisches Laboratorium Beller (G¨ottingen, Germany). Mass spectrometric characterization of organic compounds does not provide information on the purity of the material tested. Elemental analyses, UV data, 1 H NMR data, and melting points are given in the protocol steps; 13 C NMR data are shown in Table 1.10.2.
Critical Parameters
The synthesis of 7-deazapurine 2 deoxyribonucleosides via nucleobase anion glycosylation is straightforward. Purification of the intermediates can be performed under regular laboratory conditions. Exposure of the reaction mixture to sunlight should be avoided. The workup must be performed carefully, as protecting groups can be lost due to the alkaline conditions employed in the glycosylation reaction and in the workup. With regard to high glycosylation yields, the purity of the nucleobase is the key to a successful reaction. Impurities that are not removed after halogenation, such as traces of Nhalosuccinimides, will lower the reaction yields significantly. Amination of the 6halogenated intermediates is performed in a
mixture of 1,4-dioxane and aqueous NH3 to reduce the displacement of halogen substituents by hydroxides.
Troubleshooting The solubility of the deprotected nucleosides will sometimes lead to problems during purification (e.g., crystallization on the silica gel column). Crystallization of the deprotected glycosylation product from methanol or other alcohols without column chromatography is often successful. Chromatographic separation on a Serdolit AD-4 column (resin particle size, 0.1 to 0.2 mm; Serva) is useful for purifying guanosine or isoguanosine analogs.
Anticipated Results The nucleobase anion glycosylation reaction allows for large-scale preparation of various 7-deazapurine 2 -deoxyribonucleosides in a regioselective and stereoselective way. Regioselective 7-halogenation can be performed on either the nucleobase or the nucleoside. The position at which the modification occurs and the reactivity observed depend on the reaction conditions and on the structure of the starting material.
Time Considerations The time needed for synthesis and purification of the various reaction intermediates and products depends on the required reaction time, the ease of purification, and the
Synthesis of Modified Nucleosides
1.10.17 Current Protocols in Nucleic Acid Chemistry
Supplement 21
Table 1.10.2
13
C NMR Chemical Shifts (δ) for 7-Deazapurine 2 -Deoxyribonucleosidesa,b
Sys Pur
C(2)c C(2)
C(4)c C(6)
C(4a) C(5)
S.9
150.7
151.2
116.7
S.10 S.1d S.11
151.1
151.4
152.0
157.3
151.2
150.9
116.7 103.2 106.5
C(5) C(7) 53.9 53.3 51.9 162.1
C(6) C(8)
C(7a)c C(4)
C(1 )
C(2 )
C(3 )
C(4 )
C(5 )
133.3
150.5
83.7
—d
74.7
81.5
63.9
85.5
d
70.8
87.8
61.7
d
71.0
87.5
62.0
d
70.7
87.4
61.6
d
133.5 126.9 51.1
150.5 149.8 129.0
83.0 83.6
— — —
S.5d
144.6
147.2
108.0
157.7
55.4
125.6
83.2
—
70.9
87.6
61.8
S.14
152.3
162.8
103.3
52.5
127.3
152.2
82.9
37.8
74.0
81.2
63.5
S.2d
152.7
157.7
99.8
102.2
121.5
150.5
82.2
38.5
70.9
87.0
61.9
82.9
d
70.8
87.9
62.7
d
70.7
87.2
61.6
d
S.15 S.16 S.6d e
S.20 S.3c S.4c
160.2
163.6
159.8
163.8
99.5 100.2
49.4 51.9
124.9 125.5
155.0 151.7
83.0
— —
150.3
158.8
103.1
56.2
122.2
138.7
85.4
—
70.6
87.4
61.3
152.5
151.7
112.0
89.8
125.4
151.3
84.5
40.3
74.8
82.6
63.9
81.9
—
d
70.9
86.9
61.9
—
d
70.8
87.2
61.8
160.1
157.2
153.6
156.0
94.3 91.0
87.4 87.4
117.0
152.1
118.8
f
—
82.5
a Sys, carbon position in systematic numbering scheme; Pur, carbon position in purine numbering scheme. b Measured in DMSO-d unless otherwise indicated. 6 c Peak assignments are tentative. d Superimposed upon DMSO-d peak. 6 e Measured in CDCl . 3
f Not detected.
experience of the person running the experiments. For an experienced chemist, the synthesis of S.1d or S.5d starting from nucleobase S.7 (Basic Protocol 1) and synthesis of S.2d from nucleoside S.12 (Basic Protocol 2) can all be performed in ∼2 weeks. Synthesis of S.6d from S.12 (Basic Protocol 2) and of S.4c from nucleobase S.17 (Basic Protocol 3) require ∼3 weeks. In these syntheses, many intermediates can be purified by crystallization without using column chromatography.
Literature Cited Albert, A. and Serjeant, E.P. 1971. Determination of ionization constants by spectrometry. In The Determination of Ionization Constants, pp. 44-64. Chapman & Hall, London. Balow, G., Mohan, V., Lesnik, E.A., Johnston, J.F., Monia, B.P., and Acevedo, O.L. 1998. Biophysical and antisense properties of oligodeoxynucleotides containing 7-propynyl-, 7-iodo- and 7-cyano-7-deaza-2-amino-2 deoxyadenosines. Nucl. Acids Res. 26:33503357. 7-Substituted 7-Deazapurine (Pyrrolo[2,3d]pyrimidine) 2 -Deoxyribonucleosides
Hoffer, M. 1960. α-Thymidin. Chem. Ber. 93:27772781. Latimer, L.J. and Lee, J.S. 1991. Ethidium bromide does not fluoresce when intercalated adjacent to
7-deazaguanine in duplex DNA. J. Biol. Chem. 266:13849-13851. LePecq, J.B. and Paoletti, C. 1967. A fluorescent complex between ethidium bromide and nucleic acids. Physical-chemical characterization. J. Mol. Biol. 27:87-106. Kasai, H., Ohashi, Z., Harada, F., Nishimura, S., Oppenheimer, N.J., Crain, P.F., Liehr, J.G., von Minden, D.L., and McCloskey, J.A. 1975. Structure of the modified nucleoside Q isolated from Escherichia coli transfer ribonucleic acid. 7-(4,5-cis-Dihydroxy-1-cyclopenten-3ylaminomethyl)-7-deazaguanosine. Biochemistry 14:4198-4208. Kazimierczuk, Z., Cottam, H.B., Revankar, G.R., and Robins, R.K. 1984. Synthesis of 2 deoxytubercidin, 2 -deoxyadenosine, and related 2 -deoxynucleosides via a novel direct stereospecific sodium salt glycosylation procedure. J. Am. Chem. Soc. 106:63796382. Kelley, S.O. and Barton, J.K. 1998. DNA-mediated electron transfer from a modified base to ethidium: Pi-stacking as modulator of reactivity. Chem. Biol. 5:413-425. Kilpatrick, M.W. and Walker, R.T. 1982. The nucleotide-sequence of the transferRNA-met(m) from the archaebacterium thermoplasma-acidophilum. Zentralbl. Bakteriol. Mikrobiol. Hyg. 1 Abt. Orig. C 3: 79-89.
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Current Protocols in Nucleic Acid Chemistry
Li, H., Peng, X., and Seela, F. 2004. Fluorescence quenching of parallel-stranded DNA bound ethidium bromide: The effect of 7deaza-2 -deoxyisoguanosine and 7-halogenated derivatives. Bioorg. Med. Chem. Lett. 14: 6031-6034. Martin, J.C. (ed.) 1989. Nucleotide Analogues as Antiviral Agents. American Chemical Society, Washington, D.C. Mizusawa, S., Nishimura, S., and Seela, F. 1986. Improvement of the dideoxy chain termination method of DNA sequencing by use of deoxy-7deazaguanosine triphosphate in place of dGTP. Nucl. Acids Res. 14:1319-1324. Prober, J.M., Trainor, G.L., Dam, R.J., Hobbs, F.W., Robertson, C.W., Zagursky, R.J., Cocuzza, A.J., Jensen, M.A., and Baumeister, K. 1987. A system for rapid DNA sequencing with fluorescent chain-terminating dideoxynucleotides. Science 238:336-341. Pudlo, J.S., Nassiri, M.R., Kern, E.R., Wotring, L.L., Drach, J.C., and Townsend, L.B. 1990. Synthesis, antiproliferative, and antiviral activity of certain 4-substituted and 4,5-disubstituted 7-[(1,3-dihydroxy-2propoxy)methyl]pyrrolo[2,3-d]pyrimidines. J. Med. Chem. 33:1984-1992. Ramzaeva, N. and Seela, F. 1995. 7-Substituted 7-deaza-2 -deoxyguanosines: Regioselective halogenation of pyrrolo[2,3-d]pyrimidine nucleosides. Helv. Chim. Acta 78:10831090. Ramzaeva, N., Mittelbach, C., and Seela, F. 1999. 7-Halogenated 7-deaza-2 -deoxyinosines. Helv. Chim. Acta 82:12-18. Revankar, G.R. and Robins, R.K. 1991. Pyrrolo[2,3-d]pyrimidine (7-deazapurine) nucleosides. In Chemistry of Nucleosides and Nucleotides (L.B. Townsend, ed.) pp. 200-247. Plenum, New York. Rolland, V., Kotera, M., and Lhomme, J. 1997. Convenient preparation of 2-deoxy-3,5-di-Op-toluoyl-α-D-erythro-pentofuranosyl chloride. Synth. Commun. 27:3505-3511. Seela, F. and Becher, G. 2000. Synthesis, base pairing, and fluorescence properties of oligonucleotides containing 1H-pyrazolo[3,4d]pyrimidin-6-amine (8-aza-7-deazapurin-2amine) as an analogue of purin-2-amine. Helv. Chim. Acta 83:928-942. Seela, F. and Driller, H. 1989. 7-Deaza-2 -deoxyO6 -methylguanosine: Selective N2 -formylation via a formamidine, phosphoramidite synthesis and properties of oligonucleotides. Nucleosides Nucleotides 8:1-21. Seela, F. and Engelke, U. 1985. Ein Purin/7Desazapurin-Dinucleosid-Monophosphat mit 2Amino-7H-pyrrolo[2,3-d]pyrimidin als fluoreszierender Base. Liebigs Ann. Chem. 11751184. Seela, F. and Kehne, A. 1983. 2 -Desoxytubercidin: Synthese eines 2 -Desoxyadenosin-Isosteren durch Phasentransferglycosylierung. Liebigs Ann. Chem. 876-884.
Seela, F. and Menkhoff, S. 1985. Synthese von 7Desaza-2 -desoxyinosin durch Phasentransferglycosylierung. Liebigs Ann. Chem. 1360-1366. Seela, F. and Peng, X. 2004. Regioselective syntheses of 7-halogenated 7-deazapurine nucleosides related to 2-amino-7-deaza-2 -deoxyadenosine and 7-deaza-2 -deoxyisoguanosine. Synthesis 1203-1210. Seela, F. and Shaikh, K. 2004. 7Halogenated 7-deaza-2 -deoxyxanthine 2 -deoxyribonucleosides. Helv. Chim. Acta 87:1325-1332. Seela, F. and Steker, H. 1984. Synthese von 2 -desoxyribofuranosiden des 7H-Pyrrolo[2,3d]pyrimidins: Einfluβ des C-2-Substituenten auf Fluoreszenz. Liebigs Ann. Chem. 1719-1730. Seela, F. and Thomas, H. 1994. Synthesis of certain 5-substituted 2 -deoxytubercidin derivatives. Helv. Chim. Acta 77:897-903. Seela, F. and Wei, C. 1999. The base-pairing properties of 7-deaza-2 -deoxyisoguanosine and 2 deoxyisoguanosine in oligonucleotide duplexes with parallel and antiparallel chain orientation. Helv. Chim. Acta 82:726-745. Seela, F. and Zulauf, M. 1996. Palladium-catalyzed cross coupling of 7-iodo-2 -deoxytubercidin with terminal alkynes. Synthesis 726-730. Seela, F., Driller, H., and Liman, U. 1985. 7Desaza-Isostere von 2 -Desoxyxanthosin und 2 -Desoxyspongosin: Synthese via Glycosylierung von 2,4-Dichlor-7H-pyrrolo[2,3d]pyrimidin. Liebigs Ann. Chem. 312320. Seela, F., Steker, H., Driller, H., and Bindig, U. 1987. 2-Amino-2 -desoxytubercidin und verwandte Pyrrolo[2,3-d]pyrimidinyl-2 desoxyribofuranoside. Liebigs Ann. Chem. 15-19. Seela, F., Westermann, B., and Bindig, U. 1988. Liquid-liquid and solid-liquid phase-transfer glycosylation of pyrrolo[2,3-d]pyrimidines: Stereospecific synthesis of 2-deoxy-βD-ribofuranosides related to 2 -deoxy-7carbaguanosine. J. Chem. Soc., Perkin Trans. I 697-702. Seela, F., Chen, Y., Bindig, U., and Kazimierczuk, Z. 1994. Synthesis of 2 -deoxyisoinosine and related 2 -deoxyribonucleosides. Helv. Chim. Acta 77:194-202. Seela, F., Chen, Y., and Sauer, M. 1998. Synthesis of 2 ,3 -didehydro-2 ,3 -dideoxyisoinosine and oxidation of fluorescent 2-hydroxypurine nucleosides by xanthine oxidase. Nucleosides Nucleotides 17:39-52. Seela, F., Zulauf, M., Sauer, M., and Deimel, M. 2000. 7-Substituted 7-deaza-2 deoxyadenosines and 8-aza-7-deaza-2 deoxyadenosines: fluorescence of DNA-base analogues induced by the 7-alkynyl side chain. Helv. Chim. Acta 83:910-927. Seela, F., Chittepu, P., He, Y., He, J., and Xu, K. 2005. 6-Azapyrimidine and 7-deazapurine Syn2 -deoxy-2 -fluoroarabinonucleosides: thesis, conformation and properties of
Synthesis of Modified Nucleosides
1.10.19 Current Protocols in Nucleic Acid Chemistry
Supplement 21
oligonucleotides. Nucleosides Nucleic Acids In press.
Nucleotides
Simons, C. 2001. Nucleoside Mimetics: Their Chemistry and Biological Properties. Gordon and Breach Science Publishers, Amsterdam. Suhadolnik, R.J. 1970. Pyrrolopyrimidine nucleosides. In Nucleoside Antibiotics, pp. 298-353. Wiley-Interscience, New York. Winkeler, H.D. and Seela, F. 1983. Synthesis of 2-amino-7-(2 -deoxy-β-D-erythropentofuranosyl)-3,7-dihydro-4H-pyrrolo[2,3d]pyrimidin-4-one, a new isostere of 2 deoxyguanosine. J. Org. Chem. 48:31193122.
Contributed by Frank Seela and Xiaohua Peng Universit¨at Osnabr¨uck Osnabr¨uck, Germany and Center for Nanotechnology (CeNTech) M¨unster, Germany
7-Substituted 7-Deazapurine (Pyrrolo[2,3d]pyrimidine) 2 -Deoxyribonucleosides
1.10.20 Supplement 21
Current Protocols in Nucleic Acid Chemistry
Reduction of Ribonucleosides to 2-Deoxyribonucleosides
UNIT 1.11
This unit describes details of a four-step procedure for the conversion of ribonucleosides (RNA components) to 2 -deoxyribonucleosides (DNA components) originally reported in (Robins and Wilson, 1981, 1991; Robins et al., 1983a). These procedures provide an efficient chemical analogy to the biosynthetic reduction of ribonucleoside 5 -di- or 5 -triphosphates to 2 -deoxyribonucleotides, which is catalyzed by ribonucleotide reductases. The general four-step procedure is first illustrated for the conversion of adenosine to 2 -deoxyadenosine (see Basic Protocol). As shown in Figure 1.11.1, the procedure involves (1) simultaneous protection of the 3 - and 5 -hydroxyl groups of adenosine with a bifunctional cyclic disiloxane moiety, (2) thioacylation of the 2 -hydroxyl group with phenoxythiocarbonyl chloride, (3) free radical–induced reductive cleavage of the C2 –O2 bond with tributyltin hydride, and (4) deprotection with tetrabutylammonium fluoride. Using this procedure, adenosine (S.1) is converted to 2 -deoxyadenosine (S.5) with a combined yield of 78%. This approach constitutes a convenient and efficient overall chemical 2 -deoxygenation of a ribonucleoside, with an average yield of 94% for each of the four steps and without the need for chromatographic purification of any intermediates. Analytical samples of intermediates S.2, S.3, and S.4 can be obtained by performing chromatography and/or recrystallization. The use of tributyltin deuteride in the reduction step gives 2 -deuterio-2 -deoxyadenosine (S.7; Fig. 1.11.2) with >85% stereoselectivity for retention of configuration. This variation is presented (see Alternate Protocol 1), as are slightly modified procedures for the synthesis of 2 -deoxyguanosine (see Alternate Protocol 2), 2 -deoxyuridine (see Alternate Protocol 3), and 2 -deoxycytidine (see Alternate Protocol 4). CAUTION: Procedures in this unit use several toxic [e.g., tributyltin hydride, 4-(dimethylamino)pyridine] and sensitive (e.g., phenoxythiocarbonyl chloride) reagents, which require handling with great care. All reactions must be performed in an efficient chemical fume hood. It is highly recommended that all workup (i.e., isolation) and purification operations (including column chromatography on silica gel with organic chemical elution solvents) also be performed in a fume hood. Methods for disposal of all toxic materials must be in compliance with the National Research Council’s Prudent Practices in the Laboratory: Handling and Disposal of Chemicals (NRC, 1995).
PREPARATION OF 2 -DEOXYADENOSINE BY REDUCTION OF ADENOSINE
BASIC PROTOCOL
The first step in the overall 2 -deoxygenation procedure involves synthesis of 3 ,5 -OTPDS-adenosine (3 ,5 -O-TPDS-Ado; S.2; Fig. 1.11.1) by treatment of adenosine (S.1) with 1,3-dichloro-1,1,3,3-tetraisopropyl-1,3-disiloxane (TPDS-Cl2 ) in pyridine. This disiloxane reagent effects simultaneous protection of the 3 - and 5 -hydroxyl groups of ribonucleosides and provides ready access to selective funtionalization of the 2 -hydroxyl group (Markiewicz, 1979; Robins and Wilson, 1981; Robins et al., 1983a,b). The second step involves thioacylation of the 2 -hydroxyl group. To that end, synthesis of 3 ,5 -O-TPDS-2 -O-PTC-Ado (S.3) is accomplished by treatment of 3 ,5 -O-TPDS-Ado with 1.25 molar equivalents of phenoxythiocarbonyl chloride (PTC-Cl) in anhydrous acetonitrile, using 2 molar equivalents of 4-(dimethylamino)pyridine (DMAP) as a catalyst. The development of the thiocarbonyl reagent PTC-Cl was crucial for the efficient overall conversion of ribonucleosides to 2 -deoxyribonucleosides. PTC-Cl is a moderately active
Contributed by Morris J. Robins and Stanislaw F. Wnuk Current Protocols in Nucleic Acid Chemistry (2005) 1.11.1-1.11.14 C 2005 by John Wiley & Sons, Inc. Copyright
Synthesis of Modified Nucleosides
1.11.1 Supplement 21
Figure 1.11.1 Synthesis of 2 -deoxyadenosine from adenosine via a four-step procedure. The expected overall yield of S.5 from S.1 is given in parentheses. The stated yield is achieved if chromatographic purification of the intermediates S.2, S.3, and S.4 is not required. AIBN, α,α -azobisisobutyronitrile; DMAP, 4-(dimethylamino)pyridine; PTC-Cl, phenoxythiocarbonyl chloride; TBAF, tetrabutylammonium fluoride; THF, tetrahydrofuran; TPDS-Cl2 , 1,1,3,3-tetraisopropyl-1,3dichlorodisiloxane; rt, room temperature.
acylating reagent and converts the secondary 2 -hydroxyl group of S.2 to the thionocarbonate ester of S.3 in high yield. This conversion requires catalysis by DMAP (Robins and Wilson, 1981; Robins et al., 1983a). The third step involves free radical–induced reductive cleavage of the C2 –O2 bond in the phenyl thionocarbonate ester using tributyltin hydride. The thionoester undergoes clean homolytic hydrogenolysis (Barton and McCombie, 1975) upon treatment with 1.5 molar equivalents of tributyltin hydride with α,α -azobisisobutyronitrile (AIBN) as the radical initiator (Robins and Wilson, 1981; Robins et al., 1983a), to give 2 -deoxy-3 ,5 -O-TPDSAdo (S.4).
Reduction of Ribonucleosides to 2 -Deoxyribonucleosides
The fourth and final step involves removal of the silyl protection from the 3 - and 5 hydroxyl groups with tetrabutylammonium fluoride (TBAF) in tetrahydrofuran (THF). Treatment of S.4 with TBAF/THF at ambient temperature followed by heating in an oil bath at 75◦ C for an hour produces 2 -deoxyadenosine (S.5). Convenient chromatographic purification with Dowex 1 × 2 (OH− ) ion-exchange resin followed by recrystallization gives 2 -deoxyadenosine in 78% yield overall from adenosine (Robins and Wilson, 1981, 1991; Robins et al., 1983a).
1.11.2 Supplement 21
Current Protocols in Nucleic Acid Chemistry
Materials Adenosine (S.1; Yamasa Shoyu or Aldrich) Nitrogen gas Dried pyridine (see recipe) or anhydrous pyridine in Sure/Seal bottles (Aldrich) 1,3-Dichloro-1,1,3,3-tetraisopropyl-1,3-disiloxane (Aldrich) Chloroform Methanol Ethyl acetate 1 M HCl, ice cold Saturated NaHCO3 solution Saturated NaCl solution (brine) Anhydrous sodium sulfate (Na2 SO4 ) Silica gel (Merck Kieselgel 60, 230 to 400 mesh) Hot acetonitrile (∼70◦ C) Dried acetonitrile (see recipe) 4-(Dimethylamino)pyridine (DMAP; Aldrich) Phenoxythiocarbonyl chloride (phenyl chlorothionoformate; PTC-Cl; Aldrich) Dried toluene (see recipe) α,α -Azobisisobutyronitrile (AIBN; Aldrich) Tributyltin hydride (Aldrich) Oxygen-free nitrogen Ethanol Dried tetrahydrofuran (see recipe) 1 M tetrabutylammonium fluoride (TBAF) in THF (solution purchased from Aldrich) Diethyl ether Dowex 1 × 2 (OH− ) ion-exchange resin (see recipe) Phosphorus pentoxide (P2 O5 ) Heating mantle Reflux condensers Vacuum drying pistol (Ace Glass) Oven-dried 100-mL two- or three-neck round-bottom flasks with septa Syringes and syringe needles TLC plates (Merck Kieselgel 60 F254 aluminum-backed sheets or equivalent) Ultraviolet lamp (254 nm) B¨uchi rotary evaporator with Dewar dry ice condenser, connected to a “house” vacuum system or vacuum pump 50- and 250-mL separatory funnels Glass funnel with Whatman no. 1 fluted filter paper 3 × 40–cm, 2 × 30–cm, and 1 × 20–cm chromatography columns Silicon oil bath Wide-mouth 50-mL Erlenmeyer flask Desiccator or other closed glass vessel Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E) NOTE: All evaporations are effected with a B¨uchi rotary evaporator equipped with a Dewar dry ice condenser. This is connected to a house vacuum system (≤15 Pa) when evaporating more volatile materials and to a mechanical oil pump (<1 Pa) when evaporating less volatile materials in vacuo. The heating bath for the evaporation flask is maintained at ≤40◦ C.
Synthesis of Modified Nucleosides
1.11.3 Current Protocols in Nucleic Acid Chemistry
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Prepare 3 ,5 -O-protected adenosine 1. Dry an adenosine sample (S.1) 24 hr at ≤10 Pa, 100◦ C (refluxing water), in a vacuum drying pistol. 2. Place 267 mg (1.0 mmol) dried adenosine in an oven-dried 100-mL two- or threeneck round-bottom flask containing a stir bar. Seal each neck with a septum, flush the flask and its contents with nitrogen using a slow positive nitrogen flow through a syringe, and maintain a nitrogen atmosphere to protect the contents from moisure. 3. Add 10 mL dried pyridine using a dry syringe inserted through one septum. Alternatively, anhydrous pyridine from a Sure/Seal bottle (Aldrich) can be used.
4. To the resulting suspension, add 320 µL (316 mg, 1.0 mmol) 1,3-dichloro-1,1,3,3tetraisopropyl-1,3-disiloxane over ∼1 min using a dry syringe. 5. Stir the mixture at ambient temperature for ∼6 hr. Monitor reaction progress by TLC (APPENDIX 3D) on silica gel plates. Apply adenosine to the plate beside the reaction mixture for comparison. Develop plates with 9:1 (v/v) chloroform/MeOH, and visualize bands under a UV lamp (254 nm). The Rf value for the product (S.2) is higher than that for adenosine (S.1).
6. When the reaction is complete, evaporate all volatile materials in vacuo using a B¨uchi rotary evaporator equipped with a Dewar dry ice condenser and oil pump. 7. Add 50 ml ethyl acetate to the oily residue in the flask, transfer to a 250-mL separatory funnel, and wash with 30 mL water. 8. Separate the organic layer and wash it successively with ice-cold 1 M HCl (2 × 20 mL), ice-cold water (15 mL), saturated NaHCO3 (15 mL), and saturated NaCl (15 mL). 9. Dry the washed organic layer over ∼70 g anhydrous Na2 SO4 . Filter off the Na2 SO4 by gravity filtration with a glass funnel and Whatman no. 1 fluted filter paper, and evaporate the filtrate to dryness using a rotary evaporator connected to a vacuum system. Dry the resulting amorphous solid overnight in vacuo. The dried amorphous solid S.2 (∼500 mg, ∼98% yield) is of sufficient purity for direct use in the subsequent thioacylation step. If desired, it can be purified further on a silica gel column and recrystallized as described in steps 10 to 15.
10. Prepare a slurry of ∼100 g silica gel in chloroform and pour it into a 3 × 40–cm chromatography column. 11. Dissolve the dried product in a minimal amount of chloroform and layer it carefully on top of the silica gel. 12. Elute with 100 mL chloroform followed by 0.5% to 4.0% MeOH/chloroform, increasing the polarity by 0.5% MeOH for each additional 50 mL chloroform. Collect 25-mL fractions. 13. Evaluate fractions by TLC (as in step 5) and combine fractions that contain only S.2. Evaporate the volatile materials from the combined fractions using a rotary evaporator equipped with a vacuum system, and dry the purified S.2 overnight in vacuo. Reduction of Ribonucleosides to 2 -Deoxyribonucleosides
14. Recrystallize the amorphous glass by adding a small amount of hot (∼70◦ C) acetonitrile (∼30 mL) and chilling the resulting solution at ∼4◦ C. 15. Collect the precipitate by vacuum filtration and dry it in vacuo.
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16. Check the purity of the product if analytically pure material is required. The yield of purified 3 ,5 -O-(1,1,3,3-tetraisopropyldisilox-1,3-diyl)adenosine (S.2) is ∼433 mg (85%). m.p. 98◦ –99.5◦ C. UV (MeOH) max 259 nm (ε 14,900). 1 H NMR (400 MHz, DMSO-d6 ): δ 1.05–1.10 (m, 28H), 3.92 (dd, J = 12.6, 2.6 Hz, 1H), 4.00 (dt, J = 8.0, 3.0 Hz, 1H), 4.05 (dd, J = 12.6, 3.2 Hz, 1H), 4.51 (“t”, J = 5.0 Hz, 1H), 4.78 (dd, J = 8.0, 5.0 Hz, 1H), 5.60 (d, J = 4.7 Hz, 1H), 5.86 (d, J = 0.9 Hz, 1H), 7.30 (br s, 2H), 8.10 (s, 1H), 8.19 (s, 1H). HRMS (EI) m/z 509.2485 (M+ [C22 H39 N5 O5 Si2 ] = 509.2490).
Perform thioacylation of 2 -OH 17. In an oven-dried 100-mL two- or three-neck round-bottom flask containing a stir bar, dissolve the vacuum-dried sample of compound S.2 (from step 9 or step 15) in 15 mL dried acetonitrile. Protect from moisture by flushing with nitrogen and maintaining a nitrogen atmosphere at a positive pressure. 18. Add 250 mg (2.05 mmol) DMAP in one portion. 19. Slowly add 172 µL (215 mg, 1.25 mmol) PTC-Cl using a dry syringe inserted through one of the septa. 20. Stir the mixture at ambient temperature for ∼16 hr. Monitor reaction progress by TLC as in step 5, but develop plates with 19:1 (v/v) chloroform/MeOH. The Rf value for the product (S.3) is higher than that for the starting material (S.2).
21. When the reaction is complete, evaporate all volatile materials in vacuo. 22. Add 50 mL ethyl acetate to the oily residue, transfer to a 250-mL separatory funnel, and wash with 30 mL water. 23. Separate the organic layer and wash it successively with ice-cold 1 M HCl (20 mL), ice-cold water (15 mL), saturated NaHCO3 (15 mL), and saturated NaCl (15 mL). 24. Dry the washed organic layer over ∼60 g anhydrous Na2 SO4 , filter off the Na2 SO4 by gravity filtration, and evaporate the filtrate to dryness using a rotary evaporator connected to a vacuum system. Dry the resulting light-yellow oil overnight in vacuo to give a solid foam. The amorphous product S.3 (∼625 mg, ∼97% yield from S.1) is of sufficient purity for direct use in the subsequent cleavage step. If desired, it can be purified further on a silica gel column as described in steps 25 to 28.
25. Prepare a slurry of 100 g silica gel in chloroform and pour it into a 3 × 40–cm chromatography column. 26. Dissolve the amorphous product in a minimal amount of chloroform and layer it carefully on top of the silica gel. 27. Elute with 100 mL chloroform followed by 0.5% to 2.0% MeOH/chloroform, increasing the polarity by 0.5% MeOH for each additional 75 mL chloroform. Collect 25-mL fractions. 28. Evaluate fractions by TLC (step 5) using 49:1 (v/v) chloroform/MeOH as the eluent, and combine fractions that contain only S.3. Evaporate the combined fractions using a rotary evaporator equipped with a vacuum system, and dry the resulting oily residue overnight in vacuo. Synthesis of Modified Nucleosides
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29. Check the purity of the product if analytically pure material is required. The yield of purified 2 -O-(phenoxythiocarbonyl)-3 ,5 -O-(1,1,3,3-tetraisopropyldisilox1,3-diyl)adenosine (S.3) is ∼586 mg (91% from S.1). UV (MeOH) max 259 and 228 nm, min 245 nm. 1 H NMR (400 MHz, DMSO-d6 ): δ 1.05–1.10 (m, 28H), 3.98 ( br d, J = 13.0 Hz, 1H), 4.00 (m, 1H), 4.05 (dd, J = 13.0, 4.1 Hz, 1H), 5.53 (dd, J = 8.7, 5.5 Hz, 1H), 6.36 (d, J = 1.2 Hz, 1H), 6.53 (dd, J = 5.5, 1.2 Hz, 1H), 7.17 (d, J = 7.8 Hz, 2H), 7.37 (t, J = 7.4 Hz, 1H), 7.43 (br s, 2H), 7.52 (t, J = 7.8 Hz, 2H), 8.08 (s, 1H), 8.32 (s, 1H). HRMS (EI) m/z 602.1924 (M+ − i-Pr [C26 H36 N5 O6 SSi2 ] = 602.1927).
Cleave the C2 –O2 bond 30. In an oven-dried 100-mL two- or three-neck round-bottom flask containing a stir bar, dissolve the vacuum-dried sample of S.3 (from step 24 or step 28) in 20 mL dried toluene. 31. Add 32 mg (0.2 mmol) AIBN in one portion. 32. Slowly add 400 µL (436 mg, 1.5 mmol) tributyltin hydride to the reaction mixture using a dry syringe. 33. To purge (deoxygenate) the reaction mixture, cap each neck of the reaction flask with a septum, insert a syringe needle attached to an oxygen-free nitrogen source deep into the reaction mixture through one septum, insert a second syringe needle that is open to air into another septum (the outlet port), and then flow nitrogen into the flask for at least 20 min. The cleavage reaction is conducted under an atmosphere of oxygen-free nitrogen for best yields.
34. Remove the outlet septum from the flask and replace it with a reflux condenser. Stop the flow of nitrogen, and close the condenser to the external atmosphere by capping it with a septum. 35. Heat the reaction mixture with magnetic stirring for 3 hr in an oil bath at 75◦ C. At this point, if isolation of S.4 is desired, the investigator can add 2 mL (2 mmol) of 1 M TBAF/THF to the reaction mixture, heat an additional 1 hr at 75◦ C, and then skip to step 49.
36. Evaporate all volatile materials using a rotary evaporator equipped with a vacuum system. 37. Add 50 ml ethyl acetate to the oily residue, transfer to a 250-mL separatory funnel, and wash with 20 mL saturated NaHCO3 followed by 20 mL saturated NaCl. 38. Dry the washed organic layer over ∼60 g anhydrous Na2 SO4 . Filter off the Na2 SO4 by gravity filtration, and evaporate the filtrate to dryness using a rotary evaporator connected to a vacuum system. The resulting oil of S.4 (yield >100%, due to traces of tin byproducts) is of sufficient purity for direct use in the subsequent deprotection step. If desired, it can be purified further on a silica gel column and then recrystallized as described in steps 39 to 44.
39. Prepare a slurry of 100 g silica gel in chloroform and pour it into a 2 × 30–cm chromatography column. 40. Dissolve the product in a minimal amount of chloroform and layer it carefully on top of the silica gel. Reduction of Ribonucleosides to 2 -Deoxyribonucleosides
41. Elute with 100 mL chloroform followed by 0.5% to 3.0% MeOH/chloroform, increasing the polarity by 0.5% MeOH with each additional 60 mL chloroform. Collect 25-mL fractions.
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42. Evaluate fractions by TLC (step 5) using 49:1 chloroform/MeOH as the eluent, and combine the fractions that contain only S.4. Evaporate the combined fractions using a rotary evaporator connected to a vacuum system, and dry the residual foam overnight in vacuo. The Rf value for S.4 is lower than that for S.3.
43. Recrystallize the product from ethanol in as small a volume as possible, by heating until dissolved and then cooling slowly to a final temperature of 4◦ C, to give analytically pure S.4. 44. Collect the precipitate by vacuum filtration and dry it in vacuo. 45. Check the purity of the product if analytically pure material is required. The yield of recrystallized 2 -deoxy-3 ,5 -O-(1,1,3,3-tetraisopropyldisilox-1,3-diyl)adenosine (S.4) is ∼385 mg (75% from S.1). m.p. 113◦ –114.5◦ C. UV (MeOH) max 259 nm (ε 14,000). 1 H NMR (400 MHz, DMSO-d6 ): δ 1.0–1.1 (m, 28H), 2.56 (dt, J = 13.3, 8.3 Hz, 1H), 2.82 (ddd, J = 13.3, 7.9, 2.6 Hz, 1H), 3.79 (m, 1H), 3.90 (m, 2H), 5.19 (“q”, J = 7.7 Hz, 1H), 6.28 (dd, J = 8.1, 2.5 Hz, 1H), 7.30 (br s, 2H), 8.07 (s, 1H), 8.21 (s, 1H). HRMS (EI) m/z 493.2538 (M+ [C22 H39 N5 O4 Si2 ] = 493.2544).
Perform deprotection 46. In an oven-dried 100-mL two- or three-neck round-bottom flask containing a stir bar, dissolve the product S.4 (from step 38 or step 44) in 10 mL dried THF under a nitrogen atmosphere. 47. Add 2 mL (2 mmol) of 1 M TBAF in THF using a dry syringe. 48. Stir the reaction mixture 1 hr at ambient temperature, and then 1 hr in an oil bath at 50◦ C. 49. Evaporate all volatile materials to dryness in vacuo. 50. Partition the oily residue between 10 mL H2 O and 10 mL diethyl ether using a 50-mL separatory funnel. 51. Separate the aqueous layer and wash it with an additional 2 mL diethyl ether. 52. Concentrate the aqueous layer to a volume of ∼2 mL. 53. Apply the aqueous concentrate to a 1 × 20–cm chromatography column containing 15 mL of Dowex 1 × 2 (OH– ) ion-exchange resin packed in water. 54. Elute with water and collect 7-mL fractions. Elution volume will vary; monitor fractions by TLC as in step 55 until elution is complete.
55. Monitor fractions by TLC (step 5) using 9:1 (v/v) chloroform/MeOH as the eluent, and combine the fractions that contain only S.5. Evaporate the combined fractions to dryness in vacuo using a rotary evaporator connected to a vacuum system. The Rf value for S.5 is lower than that for S.4.
56. Dissolve the residue in a small amount of ethanol (∼2 mL) in a wide-mouth 50-mL Erlenmeyer flask, applying heat if necessary. Perform diffusion crystallization by placing the flask in a closed chamber (e.g., a desiccator) containing a significant volume of diethyl ether and allowing it to stand for several days. The diffusion crystallization process (Robins et al., 1976) gives better yields of higherpurity crystals than does the usual “heating/cooling” crystallization method with the same solvents.
Synthesis of Modified Nucleosides
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57. Collect the crystals by vacuum filtration and dry in vacuo over P2 O5 at 100◦ C (refluxing water). 58. Check the purity of the product if analytically pure material is required. The yield of purified 2 -deoxyadenosine (S.5) is ∼195 mg (78% overall yield from S.1, without purification of intermediates). m.p. 191◦ –192◦ C. UV (MeOH) max 259 nm (ε 14,100). 1 H NMR (400 MHz, DMSO-d6 ): δ 2.26 (ddd, J = 13.2, 6.1, 2.8 Hz, 1H), 2.73 (ddd, J = 13.3, 7.8, 5.8 Hz, 1H), 3.53 (ddd, J = 11.7, 6.7, 4.3 Hz, 1H), 3.63 (dt, J = 11.8, 4.7 Hz, 1H), 3.88 (m, 1H), 4.41 (m, 1H), 5.25 (dd, J = 4.9, 6.6 Hz, 1H), 5.31 (d, J = 4.0 Hz, 1H), 6.35 (dd, J = 7.7, 6.1 Hz, 1H), 7.31 (s, 2H), 8.13 (s, 1H), 8.32 (s, 1H). HRMS (EI) m/z 251.1073 (M+ [C10 H13 N5 O3 ] = 251.1082). ALTERNATE PROTOCOL 1
PREPARATION OF 2 -DEUTERIO-2 -DEOXYADENOSINE BY REDUCTION OF ADENOSINE The 2 -O-phenoxythiocarbonyl ester S.3 is reduced as described in the Basic Protocol, but with tributyltin deuteride in place of tributyltin hydride to give S.6 (Fig. 1.11.2). Deprotection of S.6 (as described in the Basic Protocol) followed by purification on a Dowex 1 × 2 (OH– ) ion-exchange column and recrystallization gives 2 -deuterio2 -deoxyadenosine (S.7) in 89% overall yield from S.3 (81% from S.1). The ratio of ribo/arabino (2 -R/S) deuterium substitution in S.7 is ∼88:12, based on analysis of 400 MHz 1 H NMR spectra (Robins et al., 1983a). 2 -Deuterio-2 -deoxyadenosine (S.7): 1 H NMR (400 MHz, DMSO-d6 ): δ 2.30 (dd, J2 –1 = 2.8 Hz, J2 –3 = 5.8 Hz, 0.12H, H2 , S epimer), 2.71 (dd, J2 –1 = 7.5 Hz, J2 –3 = 5.8 Hz, 0.88H, H2 , R epimer), 3.57 (dd, J5 –4 = 4.2 Hz, J5 –5 = 12.0 Hz, 1H, H5 ), 3.68 (dd, J5 –4 = 4.2 Hz, J5 –5 = 12.0 Hz, 1H, H5 ), 3.94 (m, 1H, H4 ), 4.46 (dd, J3 –4 = 2.8 Hz, J3 –2 = 5.8 Hz, 1H, H3 ), 5.35 (m, 2H, 3 -OH and 5 -OH), 6.39 (d, J1 –2 = 7.5 Hz, ∼0.9H, H1 , R epimer), 7.29 (s, 2H, NH2 ), 8.18 (s, 1H, H2), 8.36 (s, 1H, H8).
ALTERNATE PROTOCOL 2
PREPARATION OF 2 -DEOXYGUANOSINE BY REDUCTION OF GUANOSINE Guanosine (S.8a; Fig. 1.11.3) is converted to 2 -deoxyguanosine (S.12a) with 57% overall yield, using the steps outlined in the Basic Protocol. Its 3 ,5 -O-TPDS derivative (S.9a) is obtained in 70% yield from S.8a, the lowest of any of the nucleosides examined, but the subsequent three reactions proceed smoothly, giving a combined yield of 81% (Robins et al., 1983a). Purification of 2 -deoxyguanosine (S.12a) is performed on a column of Dowex 1 × 2 (OH– ) ion-exchange resin, but after application of the sample (see Basic Protocol, step 53), the column is washed with water (100 mL) and S.12a is eluted with 0.25 M aqueous triethylammonium bicarbonate (TEAB) buffer, pH ∼9 (see recipe). (Elution with water is precluded by the strong binding of S.12a to the basic resin due to the acidic proton on N1 of the guanine ring.) The fractions containing only S.12a are combined and evaporated (see Basic Protocol, step 55), and then water (∼50 mL) is added and evaporated in vacuo several times (as needed) to remove residual TEAB. The product is recrystallized by adding a minimal volume of hot water and then cooling, instead of using diffusion crystallization with ethanol and diethyl ether (Robins et al., 1983a). The amount of hot water needed varies, but more is required when there is less residual TEAB.
Reduction of Ribonucleosides to 2 -Deoxyribonucleosides
2 -Deoxyguanosine (S.12a): m.p. 251◦ –252◦ C. UV (0.1 M HCl) max 255 nm (ε 12,300), (0.1 M NaOH) max 260 nm (ε 9200). 1 H NMR (400 MHz, DMSO-d6 ): δ 2.20 (ddd, J = 13.0, 6.1, 2.8 Hz, 1H), 2.50 (ddd, J = 13.0, 7.4, 6.0 Hz, 1H), 3.53 (m, 2H), 3.63 (dt, J = 11.8, 4.7 Hz, 1H), 3.80 (m, 1H), 4.34 (m, 1H), 4.96 (t, J = 5.7 Hz, 1H), 5.29 (d, J = 4.0 Hz, 1H), 6.13 (t, J = 6.8 Hz, 1H), 7.45 (s, 2H), 7.93 (s, 1H), 10.96 (br s, 1H). HRMS (EI) m/z 249.0864 (M+ [C10 H13 N5 O4 ] – 18 = 249.0853).
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Figure 1.11.2 Synthesis of 2 -deuterio-2 -deoxyadenosine. The expected overall yield of S.7 from S.3 is given in parentheses. AIBN, α,α -azobisisobutyronitrile; TBAF, tetrabutylammonium fluoride; THF, tetrahydrofuran.
Figure 1.11.3 Synthesis of 2 -deoxyguanosine (S.12a), 2 -deoxyuridine (S.12b), and 2 -deoxycytidine (S.12c). The expected overall yields of these products from S.8a, S.8b, and S.8c , respectively, are given in parentheses. For cytidine, the N-acetyl form S.12c is synthesized from S.8c and then deprotected to give S.12c. AIBN, α,α -azobisisobutyronitrile; DMAP, 4-(dimethylamino)pyridine; PTC-Cl, phenoxythiocarbonyl chloride; TBAF, tetrabutylammonium fluoride; THF, tetrahydrofuran; TPDS-Cl2 , 1,1,3,3-tetraisopropyl-1,3-dichlorodisiloxane; rt, room temperature. Synthesis of Modified Nucleosides
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ALTERNATE PROTOCOL 3
PREPARATION OF 2 -DEOXYURIDINE BY REDUCTION OF URIDINE Uridine (S.8b) is converted to 2 -deoxyuridine (S.12b) with 68% overall yield by following the Basic Protocol. No cyclization of S.10b to give a 2,2 -anhydro byproduct is observed during the free radical–mediated reduction of 3 ,5 -O-TPDS-2 -O-PTC-uridine (S.10b). Chromatographic purification of 2 -deoxyuridine (S.12b) is performed on a column of coarsely ground charcoal with a stepwise elution gradient from 20% to 40% (v/v) ethanol in water. Dowex 1 × 2 (OH– ) ion-exchange resin is not used, as it binds S.12b strongly due to the acidic proton on N3 of the uracil ring (Robins et al., 1983a). 2 -Deoxyuridine (S.12b), chromatographically purified and recrystallized: m.p. 162◦ –163◦ C. UV (MeOH) max 260 nm (ε 9800). 1 H NMR (400 MHz, DMSO-d6 ): δ 2.08 (m, 2H), 3.55 (m, 2H), 3.77 (m, 1H), 4.20 (m, 1H), 5.00 (t, J = 5.5 Hz, 1H), 5.23 (d, J = 4.0 Hz, 1H), 5.61 (d, J = 8.0 Hz, 1H), 6.15 (t, J = 6.4 Hz, 1H), 7.85 (d, J = 8.0 Hz, 1H), 11.30 (br s, 1N). HRMS (EI) m/z 228.0742 (M+ [C9 H12 N2 O5 ] = 228.0746).
ALTERNATE PROTOCOL 4
PREPARATION OF 2 -DEOXYCYTIDINE BY REDUCTION OF CYTIDINE Because of the well-known propensity of cytidine (S.8c) to undergo acylation at the 4-amino group, the overall four-step 2 -deoxygenation procedure begins with 4-Nacetylcytidine (S.8c ). Commercially available S.8c is converted to 2 -deoxycytidine (S.12c) with 65% overall yield. Silylation of 4-N-acetylcytidine (S.8c ) using the procedure described in the Basic Protocol readily gives the 3 ,5 -O-TPDS derivative S.9c . Thioacylation of S.9c with PTC-Cl proceeds slowly and does not go to completion under the usual conditions. However, clean and rapid formation of S.10c is achieved upon increasing the quantity of DMAP to 6 to 9 molar equivalents (see Basic Protocol, step 18). The deprotection protocol requires treatment of S.11c with TBAF/THF (see Basic Protocol, steps 47 and 48) followed by an additional overnight treatment of the dried S.12c (step 49) with methanolic ammonia at room temperature to remove the 4-N-acetyl group and give 2 -deoxycytidine (S.12c). Methanolic ammonia is prepared by cooling reagentgrade methanol to ∼4◦ C in an ice-water bath and then bubbling ammonia gas vigorously through the cold solution for at least 30 min. A tube with a sintered glass tip works well for the introduction of ammonia gas. The product is purified on a Dowex 1 × 2 (OH– ) column (using water for elution) and recrystallized from anhydrous methanol (instead of diffusion crystallization with ethanol and diethyl ether). 2 -Deoxycytidine (S.12c): m.p. 203◦ –206◦ C. UV (0.1 M HCl) max 280 nm (ε 13,200), (0.1 M NaOH) max 271 nm (ε 9000). 1 H NMR (400 MHz, DMSO-d6 ): δ 2.10 (dt, J = 13.4, 6.2 Hz, 1H), 2.21 (ddd, J = 13.4, 5.9, 4.0 Hz, 1H), 3.55 (dd, J = 12.0, 3.7 Hz, 1H), 3.60 (dd, J = 12.0, 3.5 Hz, 1H), 3.82 (m, 1H), 4.25 (m, 1H), 5.15 (br s, 1H), 5.33 (br s, 1H), 6.07 (t, J = 6.4 Hz, 1H), 6.11 (d, J = 7.8 Hz, 1H), 8.18 (d, J = 7.8 Hz, 1H), 8.48 (br s, 1H), 9.40 (br s, 1H). HRMS (EI) m/z 227.0906 (M+ [C9 H13 N3 O4 ] = 227.0907).
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Reduction of Ribonucleosides to 2 -Deoxyribonucleosides
Dowex 1 × 2 (OH– ) ion-exchange resin In a large column, wash a large volume of Dowex 1 × 2 (Cl– ) resin (2% cross-linking, 200 to 400 mesh; Aldrich) with water until the eluate is colorless, and then wash with 4 additional column volumes of water. Wash resin with 4 column volumes of 0.5 M HCl, then wash with water until the pH of the eluate is ∼7, and continued
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continue washing with 4 additional column volumes of water. Next, wash with 1 M NaOH until no chloride anion is detected in the eluate. To test for chloride anion, acidify an aliquot of the eluate to a pH of <4 with HNO3 and then add several drops of aqueous AgNO3 . If no chloride anion is present, no precipitation of AgCl will be observed. Finally, wash resin with water until the pH of the eluate is once again ∼7, and then wash with an additional 2 column volumes of water. After washing, filter under suction and air-dry the OH– form of the resin. Store refrigerated (4◦ C) in a bottle protected from light (i.e., wrapped in aluminum foil) for long periods of time (>2 years).
Dried solvents Acetonitrile Heat reagent-grade acetonitrile at reflux for several hours over a drying agent (P2 O5 or CaH2 ). Distill the dried acetonitrile from the drying agent under an argon atmosphere. Pyridine Allow reagent-grade pyridine to stand with the drying agent CaH2 for 8 hr and then heat at reflux for ∼16 hr over CaH2 . Distill the dried pyridine from the drying agent under an argon atmosphere. Tetrahydrofuran (THF) Allow reagent-grade THF to stand with the drying agent CaH2 for several hours and then heat at reflux for 8 to 16 hr over CaH2 . Distill the dried THF from the drying agent under an argon atmosphere. Toluene Distill ∼25% of an ∼100-mL portion of reagent-grade toluene. Then distill and collect the constant-boiling fraction (∼50 mL) for use in the synthesis protocol. Alternatively, solvent drying systems that filter reagent-grade solvents through an activated alumina column under pressure provide dried solvents of excellent quality. Commercial anhydrous solvents can also be used without disadvantage in most cases.
Triethylammonium bicarbonate (TEAB) buffer Add 34.8 mL (25.3 g, 0.25 mmol) triethylamine to a 1-L volumetric flask, and then fill flask to graduation line with water. Transfer this solution to a 2-L round-bottom flask and cool in an ice-water bath to ∼4◦ C. Bubble carbon dioxide gas through the cold solution for at least 2 hr, until a weakly basic buffer solution (pH ∼9) is achieved. Store in a brown bottle at 4◦ C. COMMENTARY Background Information
Preparation of 2 -deoxyribonucleosides by coupling an activated deoxy sugar derivative with a protected nucleobase or modified precursor produces mixtures of α and β anomers. Regioisomeric mixtures are also formed in some cases. Enzymatic methods that employ transglycosylation of the sugar moiety from a naturally occurring purine or pyrimidine 2 deoxyribonucleoside to a nucleobase analog often give low yields of the desired product. Base analogs with modified structures can fail to function as alternative glycosyl-acceptor substrates. Likewise, a ribonucleotide analog
might fail to function as an alternative substrate for ribonucleotide reductases, and chemical or enzymatic phosphorylation of the precursor nucleoside is required to produce the alternative substrate 5 -di- or 5 -triphosphate. The present four-step procedure overcomes these disadvantages. Conversion of ribonucleosides of known structure to the corresponding 2 -deoxyribonucleosides retains both the regio- and stereochemical integrity of the starting compound. The four steps are generally applicable to a wide range of nucleosides, in contrast to the restrictive specificity of enzymatic approaches. Thus, this approach can be
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Reduction of Ribonucleosides to 2 -Deoxyribonucleosides
applied with a high degree of confidence for syntheses of 2 -deoxy analogs of accessible ribonucleosides. Introduction of hydrogen isotopes at C2 with high retentive stereoselectivity can also be performed. Initial regiospecific approaches for 2 deoxygenation of ribonucleosides were impeded by the lack of convenient methods for differentiating the cis secondary 2 - and 3 hydroxyl groups. This problem was solved by Markiewicz (1979) through the development of a reagent for selective, simultaneous protection of the 3 - and 5 -hydroxyl groups. Treatment of ribonucleosides with the bifunctional compound 1,3-dichloro-1,1,3,3tetraisopropyldisiloxane results in selective initial attack at the primary 5 -hydroxyl group followed by a second, slower reaction at O3 to give the 3 ,5 -O-l,1,3,3-tetraisopropyldisilox1,3-diyl (3 ,5 -O-TPDS) nucleoside derivatives (e.g., S.2, S.9) in high yield. Removal of the 3 ,5 -O-TPDS group is effected with tetrabutylammonium fluoride (TBAF) in tetrahydrofuran (THF). Barton and coworkers employed thionobenzoate, S-methyl xanthate, thiocarbonylimidazolide, and cyclic thionocarbonate derivatives for deoxygenation of secondary alcohols, including carbohydrate derivatives (Barton and McCombie, 1975; Barton and Subramanian, 1977; Barton, 1994). However, the basic reaction conditions used for preparation of the xanthate esters were not applicable to 3 ,5 -O-TPDS-protected nucleosides, and the two-stage preparation of thionobenzoates employed toxic and noxious conditions (phosgene/N,N-dimethylbenzamide followed by hydrogen sulfide). Preparation of the xanthates also required the use of two inconvenient and toxic reagents, carbon disulfide and methyl iodide. Additionally, it had been reported that imidazole (a byproduct of the reduction of thiocarbonylimidazolides) was capable of catalyzing the decomposition of the tributyltin hydride reductant. Phenoxythiocarbonyl chloride (PTC-Cl) is used as a convenient thionoacylation agent. PTC-Cl is moderately reactive with the somewhat hindered secondary 2 -hydroxyl group of 3 ,5 -O-TPDS-nucleosides. Reactions using pyridine as a catalyst are sluggish, but more powerful catalysis by 4(dimethylamino)pyridine (DMAP) results in high yields of the 2 -O-phenoxythiocarbonyl esters in a reasonable time period at ambient temperature. The phenyl thionocarbonate esters (e.g., S.3, S.10) are designed to undergo free radical–mediated hydrogenolysis of the
C2 –O2 bond with complete selectivity. The initial radical intermediate, formed by attack of the tributyltin radical on sulfur, undergoes β-scission exclusively at the secondary carbon (C2 ) in preference to the phenyl carbon, as scission at the latter would produce an aryl radical of much higher energy (Robins and Wilson, 1981; Robins et al., 1983a). The first Barton-McCombie deoxygenations (Barton and McCombie, 1975) employed tributyltin hydride as the hydrogen source in thermal reactions with thionoester derivatives. However, no mention or use of a free radical initiator is found in those first procedures. The introduction of the free radical initiator α,α -azobisisobutyronitrile (AIBN) into the Barton-McCombie deoxygenation yielded clean, radical-mediated homolytic hydrogenolysis of the phenyl thionocarbonate esters S.3 and S.10 with tributyltin hydride in toluene at 75◦ C (Robins and Wilson, 1981). Barton-McCombie deoxygenations now routinely use AIBN or other radical initiators. Reductive 2 -deoxygenation of thionoesters with tributyltin deuteride provides ready access to 2 -deuterio-2 -deoxyribonucleosides (e.g., S.6). The authors obtained >85% retention of configuration under standard conditions at 75◦ C, which compares well with the complete retention of configuration observed in enzymatic 2 -deoxygenations mediated by ribonucleotide reductases. Other hydrogen sources, initiators, substituted phenylthiocarbonyl chloride reagents, and experimental conditions have been developed for the homolytic hydrogenolysis of 2 -thionocarbonate esters derived from nucleosides and carbohydrates. Kawashima et al. (1995) reported triethylborane-initiated reductions of 2 -O-PTC esters using tin deuteride at ambient temperature, and also found that triethylborane-initiated reductions of 2 -bromo-2 -deoxyribonucleosides with tin deuteride at low temperatures (less than −69◦ C) proceed with very high retention of configuration (≥99%). Silyl hydrides have been used as less toxic substitutes for tin hydrides (Chatgilialoglu and Ferreri, 1993). For example, a tris(trimethylsilyl)[2 H]silane/triethylborane system reduces 2 -O-PTC esters of nucleosides in high yields with retention of stereochemistry (2 -R/S ≥99:1) at 0◦ C (Kawashima et al., 1997), and triphenylsilyl radical–mediated hydrogenolysis of 3 -O-PTC esters of nucleosides proceeds in high yields with benzoyl peroxide as the radical initiator (Robins et al., 1996). Hypophosphorous
1.11.12 Supplement 21
Current Protocols in Nucleic Acid Chemistry
acid and dialkyl phosphites have also been employed as radical hydrogen-transfer sources for hydrogenolysis of O-thiocarbonyl groups (or halides) on the sugar moieties of nucleosides (Takamatsu et al., 2001). The directed single-site labeling of nucleosides (i.e., stereoselective introduction of deuterium) using the radical deoxygenation procedure presented here has advantages relative to other methods for the synthesis of selectively deuterated nucleosides, which usually involve reduction of ketonucleosides (Hansske et al., 1984; Robins et al., 1997) or multistep elaboration of sugars followed by coupling with heterocyclic bases (F¨oldesi et al., 2004).
Characterization Data Physical and spectroscopic data for all products and intermediates were reported in Robins et al. (1983a,b). Data for all final products are reported here, and were identical to those for authentic samples of the natural products. Data for intermediates in the reduction of adenosine are also presented here. Data for intermediates of the other nucleosides are similar to those for adenosine (see Robins et al., 1983a).
Critical Parameters It is essential that oven-dried glassware and dried solvents be used in these protocols. Protection of reaction mixtures from atmospheric moisture and oxygen enhances yields. In particular, the reaction mixture for the deoxygenation step must be carefully deoxygenated. In the Alternate Protocols, the ratios of the TLC and column chromatography solvents should be adjusted slightly from those in the Basic Protocol to accommodate differences in the polarities of intermediates with different nucleobases. Guanosine and cytidine analogs are more polar than their adenosine and uridine counterparts, respectively, and greater proportions of methanol relative to chloroform should be used with the more polar compounds. Crystalline intermediates and the final product 2 -deoxyribonucleosides are usually stable at room temperature for extended periods of time. Storage in a refrigerator at ∼4◦ C (in a sealed container that contains a desiccant) markedly extends the storage life. An amorphous intermediate should be subjected to the next step as quickly as convenient. Serious decomposition of an amorphous intermediate is usually not observed if it is stored overnight in a refrigerator at ∼4◦ C (in a sealed container that contains a desiccant).
Anticipated Results The method presented here generally gives high yields even for inexperienced workers. The deoxygenation step and the purification of the final 2 -deoxyribonucleosides on Dowex 1 × 2 (OH– ) resin are more involved. With experience, however, the stated yields can be obtained reliably.
Time Considerations The four-step preparation of each of the 2 -deoxyribonucleosides presented in this unit can be performed in about five days (one week) when the intermediate products (S.2, S.3, and S.4 for adenosine) are used directly in subsequent reactions without further chromatographic purification. Each reaction requires a total of one day (with proper time management), assuming that the first two reactions (protection and thioacylation) are allowed to proceed to completion overnight.
Literature Cited Barton, D.H.R. 1994. The invention of chemical reactions of relevance to the chemistry of natural products. Pure Appl. Chem. 66:1943-1954. Barton, D.H.R. and McCombie, S.W. 1975. A new method for the deoxygenation of secondary alcohols. J. Chem. Soc., Perkin Trans. 1 1574-1585. Barton, D.H.R. and Subramanian, R. 1977. Reactions of relevance to the chemistry of aminoglycoside antibiotics. Part 7. Conversion of thiocarbonates into deoxy-sugars. J. Chem. Soc., Perkin Trans. 1 1718-1723. Chatgilialoglu, C. and Ferreri, C. 1993. Progress of the Barton-McCombie methodology: From tin hydrides to silanes. Res. Chem. Intermed. 19:755-775. F¨oldesi, A., Kundu, M.K., Dinya, Z., and Chattopadhyaya, J. 2004. Synthesis of [2 -2 H1 ]ribonucleosides. Helv. Chim. Acta 87:742-757. Hansske, F., Madej, D., and Robins, M.J. 1984. 2 and 3 -Ketonucleosides and their arabino and xylo reduction products. Convenient access via selective protection and oxidation of ribonucleotides. Tetrahedron 40:125-135. Kawashima, E., Aoyama, Y., Sekine, T., Miyahara, M., Radwan, M.F., Nakamura, E., Kainosho, M., Kyogoku, Y., and Ishido, Y. 1995. Sonochemical and triethylborane-induced tin deuteride reduction for the highly diastereoselective synthesis of (2 R)-2 -deoxy[2 -2 H]ribonucleoside derivatives. J. Org. Chem. 60:6980-6986. Kawashima, E., Uchida, S., Miyahara, M., and Ishido, Y. 1997. Tris(trimethylsilyl)[2 H]silanetriethylborane system producing the highly diastereoselective deuteration (>99:1) of 2 -bromo-2 -deoxy- and 2 -Ophenoxythiocarbonylribonucleosides at 0◦ C. Tetrahedron Lett. 42:7369-7372.
Synthesis of Modified Nucleosides
1.11.13 Current Protocols in Nucleic Acid Chemistry
Supplement 21
Markiewicz, W.T. 1979. Tetraisopropyldisiloxane1,3-diyl, a group for simultaneous protection of 3 - and 5 -hydroxy functions of nucleosides. J. Chem. Res., Synop. 24-25 and J. Chem. Res., Miniprint. 181-197. NRC (National Research Council, Committee on Prudent Practices for Handling, Storage, and Disposal of Chemicals in Laboratories). 1995. Prudent Practices in the Laboratory: Handling and Disposal of Chemicals. National Academy Press, Washington, D.C. Robins, M.J. and Wilson, J.S. 1981. Smooth and efficient deoxygenation of secondary alcohols. A general procedure for the conversion of ribonucleosides to 2 -deoxynucleosides. J. Am. Chem. Soc. 103:932-933. Robins, M.J. and Wilson, J.S. 1991. 2 -Deoxyadenosine and 2 -deoxy-2 -deuterioadenosine. Regiospecific and stereoselective 2 -deoxygenation of a ribonucleoside. In Nucleic Acid Chemistry: Improved and New Synthetic Procedures, Methods and Techniques, Vol. 4 (L.B. Townsend and R.S. Tipson, eds.) pp. 194-200. John Wiley & Sons, New York. Robins, M.J., Mengel, R., Jones, R.A., and Fouron, Y. 1976. Nucleic acid related compounds. 22. Transformation of ribonucleoside 2 ,3 -O-ortho esters into halo, deoxy, and epoxy sugar nucleosides using acyl halides. Mechanism and structure of products. J. Am. Chem. Soc. 98:82048213. Robins, M.J., Wilson, J.S., and Hansske, F. 1983a. Nucleic acid related compounds. 42. A general procedure for the efficient deoxygenation of secondary alcohols. Regiospecific and stereoselective conversion of ribonucleosides to 2 deoxynucleosides. J. Am. Chem. Soc. 105:40594065.
Robins, M.J., Wilson, J., Sawyer, L., and James, M.N.G. 1983b. Nucleic acid related compounds. 41. Restricted furanose conformations of 3 ,5 -O-(1,1,3,3-tetraisopropyldisilox1,3-diyl)nucleosides provide a convenient evaluation of anomeric configuration. Can. J. Chem. 61:1911-1920. Robins, M.J., Wnuk, S.F., Hernandez, A.E., and Samano, M.C. 1996. Nucleic acid related compounds. 91. Biomimetic reactions are in harmony with loss of 2 -substituents as free radicals (rather than anions) during mechanismbased inactivation of ribonucleotide reductases. Differential interactions of C2 azide, halogen, and alkylthio groups with tributylstannane and triphenylsilane. J. Am. Chem. Soc. 118:1134111348. Robins, M.J., Sanker, S., Samano, V., and Wnuk, S.F. 1997. Nucleic acid related compounds. 94. Remarkably high stereoselective reductions of 2 - and 3 -ketonucleoside derivatives to give arabino, ribo, and xylofuranosyl nucleosides with hydrogen isotopes at C2 and C3 . Tetrahedron 53:447-456. Takamatsu, S., Katayama, S., Hirose, N., Naito, M., and Izawa, K. 2001. Radical deoxygenation and dehalogenation of nucleoside derivatives with hypophosphorous acid and dialkyl phosphites. Tetrahedron Lett. 42:7605-7608.
Contributed by Morris J. Robins Brigham Young University Provo, Utah Stanislaw F. Wnuk Florida International University Miami, Florida
Reduction of Ribonucleosides to 2 -Deoxyribonucleosides
1.11.14 Supplement 21
Current Protocols in Nucleic Acid Chemistry
Synthesis of Fluorinated Nucleosides
UNIT 1.12
The fluorination of organic compounds often leads to a dramatic change in their biological activity. As a result, many fluorine-containing pharmaceuticals have been approved by the FDA over the past few decades, including the nucleoside analog gemcitabine. Recently, 9-(2,3-dideoxy-2-fluoro-β-D-threo-pentofuranosyl)adenine (FddA, lodenosine) has attracted much attention due to its high potency against HIV reverse transcriptase and its stability under the acidic conditions present in the human stomach. In principle, there are two routes to produce FddA. The first route, glycosylation of the adenine base with the fluoro-sugar, has two drawbacks: (1) the production of the fluoro-sugar requires several steps and (2) poor selectivity is normally observed in the final glycosylation. The second route, direct fluorination of adenosine, leads to the rearrangement of the base and the formation of an isonucleoside. The fluorination of 3 -deoxyadenosine with DAST results in very low yields. This unit presents two efficient methods for synthesis of FddA in which fluorination proceeds very smoothly, uses inexpensive reagents, and produces high yields. Basic Protocols 1 and 2 describe the synthesis of FddA (as well as FdaraA) via selective protection of 6-chloropurine riboside. The key fluorination is successfully achieved with non-corrosive triethylamine trihydrofluoride via imidazolesulfonate or triflate. The deoxygenation of the 3 -OH group was conveniently carried out using radical reduction. Basic Protocols 3 and 4 describe an alternate synthesis of FddA via 6-chloropurine 3 -deoxyriboside. For the synthesis of 6-chloropurine 3 -deoxyriboside from readily available inosine, the key step is radical debromination using hypophosphorous acid as a reducing agent. The fluorination of 5 -tritylated 6-chloropurine 3 -deoxyriboside is carried out using perfluorobutanesulfonyl fluoride with subsequent conversion to FddA by conventional means. NOTE: All reagents and solvents were purchased from commercial sources and were used without further purification.
PREPARATION OF 6-CHLOROPURINE 3 -O-BENZOYL-5 -OTRITYLRIBOSIDE
BASIC PROTOCOL 1
6-Chloropurine riboside (S.1; Fig. 1.12.1) is converted to the 3 -O-benzoate (S.2) via a stannylene complex. This is further modified to 3 -O-benzoyl-5 -O-tritylriboside (S.3) using trityl chloride in the presence of triethylamine. In both reactions, the 3 -O-benzoates are isolated from their 2 -isomers by crystallization (see Commentary for further discussion).
Materials 6-Chloropurine riboside (S.1), 99% pure (Aldrich) Di-n-butyltin(IV) oxide, 95+% Methanol (MeOH), 99.8+% Triethylamine, 99+% Benzoyl chloride, 99+% Diethyl ether, 99.5+% Magnesium sulfate (MgSO4 ), anhydrous, 99.5+% Dichloromethane (CH2 Cl2 ), 99+% Ethanol (EtOH), 99.5% N,N-Dimethylformamide (DMF), anhydrous, 99.8+%
Synthesis of Modified Nucleosides
Contributed by Satoshi Katayama, Satoshi Takamatsu, Naoko Hirose, Kunisuke Izawa, and Tokumi Maruyama
1.12.1
Current Protocols in Nucleic Acid Chemistry (2006) 1.12.1-1.12.30 C 2006 by John Wiley & Sons, Inc. Copyright
Supplement 25
Figure 1.12.1
Preparation of 6-chloro-(3-O-benzoyl-5-O-trityl-β-D-ribofuranosyl)purine.
Benzene, 99.5+% 4-Dimethylaminopyridine (DMAP), 99+% Trityl chloride, 98+% Chloroform (CHCl3 ), 99+% Pyridine, dehydrated for organic synthesis, 99+% Dimethylsulfoxide (DMSO) 3:2 and 1:4 (v/v) 10 mM phosphoric acid/MeOH Reflux condenser Vacuum filtration unit with glass filter Rotary evaporator equipped with vacuum oil pump 3.5 × 50–cm and 6.5 × 35–cm sintered glass chromatography columns packed with silica gel G TLC plates: silica-coated aluminum plate with fluorescent indicator (Merck silica gel 60 F254 ) 254-nm UV lamp HPLC apparatus with: Cosmosil guard column 5C18-MS (4.6 × 10–mm, Nacalai Tesque) Cosmosil packed column 5C18-MS (4.6 × 150–mm, Nacalai Tesque) CCPD pump (Toso) SPD-M10A photo diode array UV-VIS detector (Shimadzu) Introduce 3 -O-benzoyl group 1. Suspend 14.34 g (50 mmol) 6-chloropurine riboside (S.1) and 12.45 g (50 mmol) di-n-butyltin(IV) oxide in 500 mL MeOH and then reflux for 1 hr at 80◦ C. 2. Cool the solution to room temperature and then add 34.8 mL (250 mmol) triethylamine.
Synthesis of Fluorinated Nucleosides
1.12.2 Supplement 25
3. Add 29.0 mL (250 mmol) benzoyl chloride in a dropwise manner with magnetic stirring. Cool the solution to room temperature and then continue stirring for 30 min. 4. Remove the insoluble materials by vacuum filtration using a glass filter, then evaporate the filtrate on a rotary evaporator equipped with a vacuum oil pump. Current Protocols in Nucleic Acid Chemistry
Table 1.12.1 Selected Data for the 1 H NMR Spectra of S.2 and S.3 in CDCl3
H1
H2
H3
H4
6.20
5.05
5.59
4.37-4.39
d
q
dd
m
J (Hz)
6.6
5.9
5.5, 2.7
ppm
6.48
5.88
4.66
4.15-4.17
d
dd
q
m
J (Hz)
4.9
5.2, 4.9
5.1
ppm
6.20
5.25
5.68
4.48-4.50
d
q
dd
J (Hz)
5.5
5.5
5.5, 4.4
ppm
6.51
6.05-6.10
d
NDa
Compound S.2 (3 -O-Bz)
S.2 (2 -O-Bz)
S.3 (5 -O-Tr-3 -O-Bz)
S.3 (5 -O-Tr-2 -O-Bz)
ppm
J (Hz)
3.6
H5 a
H5 b
2 - or 3 -OH
5 -OH
6.02
5.35
d
t
6.0
5.5
5.81
5.24
d
dd
5.8
5.5, 5.2
3.34-3.43
6.05-6.10
—
m
m
NDa
4.87
4.28
3.34-3.43
5.83
q
dt
m
d
6.1
6.9, 4.4
3.77-3.81 3.71-3.75 m
m
3.66-3.69 3.00-3.83 m
m
6.0
a ND, not determined.
5. Partition the resulting liquor between 400 mL diethyl ether and 250 mL water. Wash the aqueous layer two times with 200 mL diethyl ether. 6. Combine the organic layers and wash with 100 mL water. 7. Dry over MgSO4 until the solution becomes clear, filter the drying agent under vacuum, and concentrate to a small volume on a rotary evaporator. 8. Apply column chromatography (APPENDIX 3E) to the residual solution over a 6.5 × 35–cm column of silica gel G using 1 L CH2 Cl2 and 4 L of 0% to 10% EtOH in CH2 Cl2 as eluents. Monitor and identify correct fractions by TLC (see data in step 10). 9. Concentrate the fraction of the monobenzoyl derivatives to dryness on a rotary evaporator. Crystallize from 50 mL MeOH overnight at room temperature. 10. Characterize the product by TLC (on silica gel), HPLC (see step 21), elemental analysis, UV, MS, and 1 H NMR. The compound is stable for at least 12 months when stored at 4◦ C. 6-Chloro-9-(3-O-benzoyl-β-D-ribofuranosyl)purine (S.2). Yield of pale yellowish crystals: 12.67 g (65%). m.p. 174◦ -177◦ C. TLC (95:5 v/v CH2 Cl2 /EtOH): Rf = 0.40. Anal. calcd. (C17 H15 ClN4 O5 ): C, 51.77; H, 3.94; N, 14.21%; found: C, 51.80; H, 3.99; N, 14.05. MS: m/z 360, 362 (M+ -CH2 O), 385, 387 (M+ -C6 H5 CO). UV λmax (MeOH): 264.5 nm. 1 H NMR (CDCl3 ): δ 8.97, 8.91 (each 1H, s, H2, H8), 7.5-8.1 (5H, m, C6 H5 CO), 6.16 (1H, d, J = 6.5 Hz, H1 ), 5.95 (1H, m, 2 -OH), 5.55 (1H, dd, J = 6.5, 2.4 Hz, H3 ), 5.28 (1H, br s, 5 -OH), 5.00 (1H, m, H2 ), 4.33 (1H, m, H4 ), 3.72 (2H, t, J = 2.8 Hz, H5 ). The 1 H NMR spectrum reveals that the signal of H3 appears in a relatively high field (5.55 ppm), and the two protons exchangeable with deuterium oxide are identified as 2 -OH and 5 -OH, indicating that the benzoyl group is introduced at the 3 -OH position. Analysis of the sample by HPLC indicates the presence of a small amount of the 2 -Obenzoate (3.9%). A comparison of NMR spectra for the 2 - and 3 -isomers can be found in Table 1.12.1.
Synthesis of Modified Nucleosides
1.12.3 Current Protocols in Nucleic Acid Chemistry
Supplement 25
—
Tritylate 5 -hydroxyl 11. Dissolve 10.72 g (25 mmol) S.2 in 220 mL anhydrous DMF and co-evaporate the solution with 100 mL benzene. 12. Add 11.0 mL (78.9 mmol) triethylamine, 1.01 g (8.25 mmol) DMAP, and 23.00 g (82.5 mmol) trityl chloride to the solution with magnetic stirring and keep overnight at 50◦ C. 13. Cool the solution to room temperature. Add 10 mL water and then evaporate to dryness under vacuum. 14. Partition the residue between 300 mL CHCl3 and 150 mL water. Wash the organic layer with 150 mL water. 15. Dry over MgSO4 until the solution becomes clear, filter the drying agent under vacuum, and concentrate to a small volume on a rotary evaporator. 16. Apply column chromatography to the residual solution over a 3.5 × 50–cm column packed with silica gel G using 1 L CH2 Cl2 and 4 L of 0% to 2.5% EtOH in CH2 Cl2 . Monitor and identify correct fractions by TLC (see step 18). 17. Concentrate the solution to dryness on a rotary evaporator, then crystallize the residue from 150 mL MeOH overnight at room temperature. 18. Characterize the product by TLC (on silica gel), HPLC (see step 21), elemental analysis, UV, MS, and 1 H NMR. The compound is stable for at least 12 months when stored at 4◦ C. 6-Chloro-9-(3-O-benzoyl-5-O-trityl-β-D-ribofuranosyl)purine (S.3). Yield of white crystals: 11.21 g (71%). m.p. 102◦ -106◦ . TLC (95:5 v/v CH2 Cl2 /EtOH): Rf = 0.56. Anal. calcd. (C36 H29 ClN4 O5 . 0.8H2 O): C, 66.78; H, 4.76; N, 8.65%; found: C, 66.86; H, 4.81; N, 8.43. MS: m/z 555, 557 (M+ -C6 H5 ), 389, 391 (M+ -trityl). UV λmax (MeOH): 264 nm. 1 H NMR (CDCl3 ): δ 8.69, 8.38 (each 1H, s, H2, H8), 7.2-8.05 (ca 20H, m, C6 H5 CO, trityl), 6.18 (1H, d, J = 6.1 Hz, H1 ), 5.74 (1H, dd, J = 5.5, 2.5 Hz, H3 ), 5.29 (1H, m, H2 ), 4.59 (1H, m, H4 ), 3.95 (1H, d, J = 4.5 Hz, 2 -OH), 3.53 (2H, t, J = 3.6 Hz, H5 ). HPLC confirms the presence of a small amount of the 2 -O-benzoate (2.0%). A comparison of NMR spectra for the 2 - and 3 -isomers can be found in Table 1.12.1.
Check stability of S.3 19. Dissolve 100 mg of S.3 in a solution of 4.1 mL dry CH2 Cl2 and 0.076 mL dehydrated pyridine and reflux for 5 hr at 45◦ C. 20. Cool the solution to room temperature. Dilute 100 µl solution with 2 mL DMSO and then remove the volatile compounds under vacuum. 21. Subject this sample to HPLC analysis using the following conditions:
Columns: Cosmosil guard column 5C18-MS (4.6 × 10–mm) and Cosmosil packed column 5C18-MS (4.6×150–mm) Eluent: 10 mM phosphoric acid/MeOH, 3:2 (v/v) for S.2 or 1:4 (v/v) for S.3 Flow rate: 1 mL/min Column temperature: 50◦ C. The ratio of the 2 -O-benzoate and the 3 -O-benzoate of S.3 is 3.1 to 96.9.
Synthesis of Fluorinated Nucleosides
1.12.4 Supplement 25
Current Protocols in Nucleic Acid Chemistry
SYNTHESIS OF FddA FROM 6-CHLOROPURINE 3 -O-BENZOYL-5 -O-TRITYLRIBOSIDE
BASIC PROTOCOL 2
The common first step in the synthesis of both FddA and FdaraA is fluorination of the C2 -β of S.3. This can be achieved using diethylaminosulfur trifluoride (DAST) in CH2 Cl2 in the presence of pyridine to give the 2 -deoxy-2 -fluoroarabinoside (S.4; Fig. 1.12.2) in good yield, although the non-corrosive triethylamine trihydrofluoride (Et3 N·3HF) is better for large-scale syntheses. The latter method can be performed via an imidazolesulfonate intermediate or a triflate intermediate. All three of these routes are presented here. The 3 -O-benzoyl-5 -O-trityl system is suitable for selective deprotection. Therefore, S.4 can be treated with ammonia in methanol to afford the 5 -O-trityl compound (S.5). Detritylation of S.5 to give FdaraA (S.6) is presented in the Alternate Protocol. To obtain FddA (S.10), the 3 -OH can be removed in two steps: conversion of S.5 to the phenoxythiocarbonyl compound S.7 or the xanthate S.8, followed by radical deoxygenation using tris(trimethylsilyl)silane or hypophosphorous acid as hydrogen donors to produce S.9. Finally, detritylation by acid treatment converts S.9 to FddA (S.10).
Figure 1.12.2
Preparation of FdaraA and FddA from 6-chloropurine 3 -O-benzoyl-5 -O-tritylriboside. Synthesis of Modified Nucleosides
1.12.5 Current Protocols in Nucleic Acid Chemistry
Supplement 25
Materials 6-Chloro-9-(3-O-benzoyl-5-O-trityl-β-D-ribofuranosyl)purine (S.3; see Basic Protocol 1) Dichloromethane (CH2 Cl2 ), anhydrous, 99.8% Pyridine, dehydrated for organic synthesis, 99+% Diethylaminosulfur trifluoride (DAST), 95+% 5% (w/v) and saturated sodium hydrogen carbonate (NaHCO3 ), 99.5+% Chloroform (CHCl3 ), 99+% Magnesium sulfate (MgSO4 ), anhydrous, 99.5+% Ethyl acetate (EtOAc), 99+% n-Hexane, 95+% Sulfuryl chloride, 99+% Imidazole, 99+% Sodium sulfate (Na2 SO4 ), anhydrous, 99+% Toluene, anhydrous, 99.8+% Triethylamine trihydorofluoride (Et3 N·3HF), 98% Acetonitrile (CH3 CN), 99.5+% 70:30 and 50:50 (v/v) 0.02 M aq. KH2 PO4 (pH 3.2) in CH3 CN 4-Dimethylaminopyridine (DMAP), 99+% Trifluoromethanesulfonyl chloride, 99+% Saturated aq. ammonium chloride (NH4 Cl), 99+% Saturated aq. sodium chloride (NaCl), 99+% Triethylamine, 99+% Methanol (MeOH), 99.8+% Ammonia, anhydrous, 99.99+% Phenyl chlorothionoformate, 98+% 2,2 -Azobis(isobutyronitrile) (AIBN), 98+% Tris(trimethylsilyl)silane, 96+% Nitrogen, 99.998+% Dimethylsulfoxide (DMSO), anhydrous, 99.9+% 5 M aq. sodium hydroxide (NaOH) Carbon disulfide, 99.9+% Iodomethane, 99.5+% 1,2-Dimethoxyethane, 99+% 50% (v/v) aq. hypophosphorous acid (H3 PO2 ) 35% to 37% (v/v) hydrochloric acid (conc. HCl) Amberlite IRA-900 (OAc– form; ICN) Reflux condenser Rotary evaporator equipped with a vacuum oil pump 3.5 × 50–cm and 2.5 × 32–cm chromatography columns packed with silica gel G Vacuum filtration unit with glass filter 22 × 3–cm, 230- to 400-mesh silica gel column TLC plates: silica-coated aluminum plate with fluorescent indicator (Merck silica gel 60 F254 ) HPLC with Inertsil ODS 2 4.6 × 150–mm column (GL Science) Glass filter with aspirator Vacuum oil pump 40-mm-diameter Kiriyama filter funnel (glass B¨uchner-type funnel) and no. 5B filter paper ◦ 50 C vacuum oven Synthesis of Fluorinated Nucleosides
CAUTION: DAST should be handled with gloves. This material can cause burns.
1.12.6 Supplement 25
Current Protocols in Nucleic Acid Chemistry
Perform fluorination Using DAST 1a. Dissolve 4.76 g (7.5 mmol) S.3 in a solution of 100 mL anhydrous CH2 Cl2 and 3.6 mL (44.5 mmol) pyridine. Cool the solution on ice. 2a. Add 2.25 mL (17 mmol) DAST in a dropwise manner and warm to room temperature. Reflux for 5 hr at 50◦ C. 3a. Cool the solution to room temperature. Add the solution to 500 mL of 5% NaHCO3 in a dropwise manner with magnetic stirring, and then continue stirring for 20 min. 4a. Separate the organic layer and wash the aqueous layer with 100 mL CHCl3 . Combine the organic layers and wash with 200 mL water. 5a. Dry over MgSO4 until the solution becomes clear, filter the drying agent under vacuum, and concentrate to a small volume on a rotary evaporator equipped with a vacuum oil pump. 6a. Apply column chromatography (APPENDIX 3E) to the residual solution over a 3.5 × 50–cm column packed with silica gel G using 4 L of 0% to 25% EtOAc in n-hexane. Monitor and identify fractions by TLC (see data in step 8a). 7a. Evaporate the desired fraction to dryness on a rotary evaporator. 8a. Characterize the product by TLC (on silica gel), UV, MS, and 1 H NMR. Proceed to step 14. The compound is stable for at least 1 month when stored at 4◦ C. 6-Chloro-9-(3-O-benzoyl-5-O-trityl-2-deoxy-2-fluoro-β - D -arabinofuranosyl)purine (S.4). Yield of a foam: 3.71 g (78%). TLC (50:1 v/v CH2 Cl2 /EtOH): Rf = 0.61. MS: m/z 391, 393 (M+ -trityl), 375, 377 (M+ -trityloxy). UV λmax (MeOH): 263 nm. 1 H NMR (CDCl3 ): δ 8.76 (1H, s, H2), 8.36 (1H, d, J = 3.0 Hz, H8), 7.2-8.1 (ca 20H, m, C6 H5 CO, trityl), 6.66 (1H, dd, J = 21.7, 2.7 Hz, H1 ), 5.70 (1H, dd, J = 17.0, 3.0 Hz, H3 ), 5.28 (1H, m, JHCF = 50.0 Hz, H2 ), 4.42 (1H, m, H4 ), 3.62 (1H, dd, J = 10.4, 5.2 Hz, H5 a), 3.54 (1H, dd, J = 10.4, 4.1 Hz, H5 b). The 1 H NMR spectrum of S.4 shows that the 2 -fluorine causes a downfield shift and a large H2 -C-F geminal coupling (50.0 Hz) of the 2 -proton, suggesting a 2 -fluoro structure.
Using triethylamine trihydrofluoride via imidazolesulfonate 1b. Dissolve 2.0 g (3.1 mmol) S.3 in 15.5 mL anhydrous CH2 Cl2 and then cool to −35◦ C. 2b. Add 0.45 mL (5.57 mmol) pyridine and 0.30 mL (3.72 mmol) sulfuryl chloride and then stir the mixture for 30 min. 3b. Add 0.96 g (13.9 mmol) imidazole and return the solution to room temperature. Stir the solution overnight at room temperature. 4b. Add 10 mL water for phase separation. 5b. Wash the aqueous layer with 10 mL CH2 Cl2 . Combine the organic layers. 6b. Dry over Na2 SO4 until the solution becomes clear, filter the drying agent under vacuum, and evaporate the solution to dryness using a rotary evaporator. 7b. Purify the residue on a 22 × 3–cm, 230- to 400-mesh silica gel column, eluting with a linear EtOAc/n-hexane gradient ranging from 25% to 50% (v/v) EtOAc in a
Synthesis of Modified Nucleosides
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total volume of 2.5 L. Monitor by TLC. Use the activated product (the imidazolesulfonate) immediately for the next reaction without characterization. 6-Chloro-9-(3-O-benzoyl-5-O-trityl-2-O-imidazolesulfonyl-β-D-ribofuranosyl)purine. Yield of oil: 2.28 g (96%). TLC (4:1 v/v toluene/EtOAc): Rf = 0.21.
8b. Dissolve 377 mg (0.494 mmol) of the imidazolesulfonate in 2.4 mL toluene and then add 0.49 mL (3.00 mmol) Et3 N·3HF to the solution. Stir overnight at 70◦ C. Since Et3 N·3HF is non-corrosive, the glassware does not show any indication of corrosion even after several days under these reaction conditions.
9b. Cool to room temperature. Add 10 mL EtOAc and 10 mL saturated NaHCO3 for phase separation. 10b. Wash the aqueous layer with 10 mL EtOAc and combine the organic layers. 11b. Dry over Na2 SO4 until the solution becomes clear, and filter the drying agent under vacuum. Evaporate the solution to dryness using a rotary evaporator. 12b. Dissolve the residue in 5 mL CH3 CN. Remove 100 µL, dilute with 10 mL mobile phase, and analyze by HPLC.
Column: Inertsil ODS 2 4.6 × 150–mm Mobile phase: 70:30 (v/v) 0.02 M aq. KH2 PO4 (pH 3.2) in CH3 CN Flow rate: 1.0 mL/min Column temperature: 40◦ C Injection volume: 10 µL Detection: UV at 262 nm. Under these conditions, the retention time is 12.6 min for S.4 and the yield is calculated as 81% by comparison of the peak area to the external standard sample. The product can also be isolated as in steps 6a and 7a.
13b. Characterize the product by TLC (on silica gel), UV, MS, and 1 H and 13 C NMR. Proceed to step 14. The analytical sample was obtained by purification via preparative TLC using 4:1 (v/v) toluene/EtOAc (Rf = 0.59). The compound is stable for at least 1 month when stored at 4◦ C. 6-Chloro-9-(3-O-benzoyl-5-O-trityl-2-deoxy-2-fluoro-β - D -arabinofuranosyl)purine (S.4). HRMS (FAB+) calcd. for C36 H29 N4 O4 FCl (M + H)+ 635.1861, found 635.1846. Anal. calcd. for C36 H29 N4 O4 FCl: C, 68.08; H, 4.44; N, 8.82; F, 2.99; Cl, 5.58; found: C, 68.11; H, 4.54; N, 8.69; F, 2.65; Cl, 5.82. IR (KBr): ν 3061, 1732, 1567, 1592, 1492, 1450, 1218, 1265, 1092 cm−1 . 1 H NMR (CDCl3 , 300 MHz): δ 8.76 (1H, s, H2), 8.36 (1H, d, J = 3.0 Hz, H8), 8.07 (2H, d, J = 7.2 Hz, C6 H5 CO), 7.66 (1H, t, J = 7.3 Hz, C6 H5 CO), 7.23-7.54 (ca. 17H, m, C6 H5 CO, trityl), 6.66 (1H, dd, J = 21.7, 2.8 Hz, H1 ), 5.67 (1H, dd, J = 16.9, 3.2 Hz, H3 ), 5.28 (1H, dd, J = 50.0, 2.8 Hz, H2 ), 4.41 (1H, m, H4 ), 3.63 (1H, dd, J = 10.3, 5.9 Hz, H5 a), 3.54 (1H, dd, J = 10.3, 4.2 Hz, H5 b). 13 C NMR (DMSO-d6 , 300 MHz): δ 164.9, 152.2, 151.5, 149.8, 146.1 (d, J = 5.1 Hz), 143.6, 134.2, 129.8, 129.0, 128.9, 128.4, 128.1, 127.3, 93.4 (d, J = 193.2 Hz), 86.6, 82.7 (d, J = 16.9 Hz), 79.6, 76.5 (d, J = 27.3 Hz), 63.4.
Using triethylamine trihydrofluoride via triflate 1c. Dissolve 1.3 g (2.0 mmol) S.3 in 20.0 mL toluene and then cool to 0◦ C.
Synthesis of Fluorinated Nucleosides
2c. Add 0.59 g (4.8 mmol) DMAP and 0.51 mL (4.8 mmol) trifluoromethanesulfonyl chloride, and then stir the mixture overnight at room temperature. 3c. Add 10 mL water for phase separation.
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4c. Wash the organic layer with 10 mL saturated aq. NaHCO3 , 10 mL saturated aq. NH4 Cl, and 10 mL saturated aq. NaCl. 5c. Add anhydrous Na2 SO4 until the organic layer becomes clear, and filter the drying agent under vacuum. Evaporate the solution to dryness using a rotary evaporator equipped with a vacuum oil pump. An amount of 1.35 g crude triflate was obtained and half that amount was used in the following reaction. Use the activated product (a triflate) immediately for the next reaction without characterization.
6c. Dissolve the crude triflate in 10 mL EtOAc, then add 0.42 mL (3.0 mmol) triethylamine and 0.98 mL (6.00 mmol) Et3 N·3HF to the solution. 7c. Reflux the mixture overnight at 80◦ C and cool to room temperature. 8c. Evaporate the mixture to a minimum volume using a rotary evaporator, and dissolve the residue in 10 mL MeOH. 9c. Dilute with 5 mL of 1:1 (v/v) CH3 CN/water and quantify 100 µL by HPLC analysis as in step 12b. HPLC analysis indicates the yield is 88% (by comparison to the external standard sample). The product can be isolated as in steps 6a and 7a.
Deprotect 3 -hydroxyl with ammonia 14. Dissolve 3.15 g (4.98 mmol) of S.4 in 100 mL anhydrous MeOH and saturate ammonia into the solution at 0◦ C (∼20% NH3 in MeOH). 15. Keep in a sealed tube (stainless autoclave-type reactor) for 2 days at 100◦ C. 16. Cool on ice and then evaporate carefully to dryness on a rotary evaporator. 17. Dissolve the residue with 100 mL CHCl3 . 18. Remove insoluble materials by diaphragm vacuum filtration using a glass filter, and then concentrate the filtrate to a small volume on a rotary evaporator. 19. Apply column chromatography to the solution over a 3.5 × 50–cm column packed with silica gel G using 4 L of 3% to 10% EtOH in CH2 Cl2 as eluents. Monitor by TLC. 20. Characterize the product by TLC (on silica gel), UV, MS, and 1 H NMR. The compound is stable for at least 12 months when stored at ambient temperature. 9-(5-O-Trityl-2-deoxy-2-fluoro-β-D-arabinofuranosyl)adenine (S.5). Yield of prism-like crystals: 1.87 g (73%). m.p. 210.5◦ -212.5◦ C. TLC (10:1 v/v CH2 Cl2 /EtOH): Rf = 0.45. Anal. calcd. for C29 H26 FN5 O3 : C, 68.09; H, 5.12; N, 13.69%; found: C, 68.06; H, 5.15; N, 13.70. MS: m/z 511 (M+ ), 376 (M+ -adenine), 268 (M+ -trityl). UV λmax (MeOH): 258.5 nm. 1 H NMR ([2 H6 ]DMSO): δ 8.15 (1H, s, H2), 8.05 (1H, d, J = 2.5 Hz, H8), 7.37 (2H, br s, NH2 ), 7.23-7.42 (15H, m, trityl), 6.46 (1H, dd, J = 15.4, 4.7 Hz, H1 ), 6.02 (1H, d, J = 5.2 Hz, 3 -OH), 5.22 (1H, ddd, J = 52.5, 4.7, 3.6 Hz, H2 ), 4.48 (1H, ddt, J = 19.2(d), 5.2(t), 3.6(d) Hz, H3 ), 4.04-4.07 (1H, m, H4 ), 3.24-3.38 (2H, m, H5 ).
Perform radical deoxygenation Via phenoxythiocarbonyl 21a. Dissolve 419 mg (0.82 mmol) S.5 and 273 mg (2.23 mmol) DMAP in 10 mL CH3 CN. 22a. Add 0.22 mL (1.63 mmol) phenyl chlorothionoformate and stir for 3 hr at room temperature.
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23a. Concentrate the solution to a small volume on a rotary evaporator and apply column chromatography over a 2.5 × 32–cm column of silica gel G using 1.2 L of 0% to 10% EtOH in CHCl3 as eluents. Monitor by TLC. TLC (10:1 v/v CH2 Cl2 /EtOH): Rf = 0.58.
24a. Evaporate the desired fraction to dryness on a rotary evaporator. 25a. Characterize the product by UV and 1 H NMR. The compound is stable for at least 12 months when stored at 4◦ C. 9-(3-O-Phenoxythiocarbonyl-5-O-trityl-2-deoxy-2-fluoro-β - D -arabinofuranosyl)adenine (S.7). Yield of a foam: 406 mg (80%). UV λmax (MeOH): 256 nm, 230 nm (sh); λmax (0.05 M HCl): 255 nm, 230 nm (sh). 1 H NMR (CDCl3 ): δ 8.38 (1H, s, H2), 8.08 (1H, d, J = 3.3 Hz, H8), 7.09-7.54 (ca 20H, m, trityl, Ph), 6.60 (1H, dd, J = 23.1, 2.6 Hz, H1 ), 5.95 (1H, dd, J = 17.6, 2.2 Hz, H3 ), 5.80 (1H, br s, NH2 ), 5.35 (1H, dd, J = 52.0, 2.6 Hz, H2 ), 4.45 (1H, m, H4 ), 3.57 (2H, d, J = 4.8 Hz, H5 ).
26a. Dissolve 386 mg (0.63 mmol) S.7 in 4 mL anhydrous toluene. 27a. Add 60 mg (0.37 mmol) AIBN and 0.6 mL (1.95 mmol) tris(trimethylsilyl)silane and heat at 100◦ C under an N2 atmosphere for 30 min. 28a. Cool the solution to room temperature and then collect the product using a glass filter under reduced pressure using an aspirator. 29a. Wash the product with 3 mL toluene and dry the crystals for 15 hr using a vacuum oil pump at 50◦ C. 30a. Characterize the product by TLC (on silica gel), elemental analysis, UV, MS, and 1 H NMR. Proceed to detritylation (step 37). The compound is stable for at least 12 months when stored at 4◦ C. 9-(2,3-Dideoxy-2-fluoro-5-O-trityl-β-D-threo-pentofuranosyl)adenine (S.9). Yield of crystals: 229 mg (73%). m.p. 226◦ -229◦ C. TLC (5:1 v/v toluene/MeOH): Rf = 0.59. Anal. calcd. for C29 H26 FN5 O2 : C, 70.29; H, 5.29; N, 14.13%; found: C, 70.20; H, 5.30; N, 14.24. MS: m/z 252 (M+ -trityl), 135 (adenine). UV λmax (MeOH): 258 nm. 1 H NMR (CDCl3 ): δ 8.36 (1H, s, H2), 8.07 (1H, d, J = 2.6 Hz, H8), 7.1-7.6 (ca 15H, m, trityl), 6.33 (1H, dd, J = 17.0, 3.0 Hz, H1 ), 5.70 (2H, br s, NH2 ), 5.22 (1H, m, JHCF = 43.6 Hz, H2 ), 4.4 (1H, m, H4 ), 3.46 (1H, dd, J = 8.2, 5.9 Hz, H5 a), 3.26 (1H, dd, J = 8.2, 3.2 Hz, H5 b), 2.2-2.65 (2H, m, H3 ).
Via xanthate 21b. Dissolve 6.60 g (12.9 mmol) S.5 in 52.4 mL anhydrous DMSO and cool to 10◦ C. 22b. Add 2.84 mL (14.2 mmol) of 5 M aq. NaOH. 23b. Add 3.19 mL (51.6 mmol) carbon disulfide in a dropwise manner to keep the temperature below 15◦ C. 24b. Add 1.60 mL (25.7 mmol) iodomethane and immediately pour the solution into a stirring mixture of 80 mL EtOAc and 80 mL water. 25b. Separate the layers and wash the organic layer with 80 mL water. 26b. Concentrate the organic layer to a minimum volume using a rotary evaporator and add 50 mL CH3 CN to the residue. 27b. Heat the mixture to 70◦ C and stir mixture 30 min at 70◦ C. The crude xanthate is dissolved and then crystallization occurs. Synthesis of Fluorinated Nucleosides
28b. Cool the mixture to 0◦ C and filter the product under vacuum through a 40-mm Kiriyama funnel (no. 5B filter paper).
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29b. Dry the product in a 50◦ C vacuum oven to obtain the xanthate (7.32 g, 91.8% wt. purity, 86.6% yield). Use the xanthate S.8 immediately for the next reaction.
30b. Dissolve 7.32 g (91.8% wt, 11.2 mmol) S.8 in 30 mL of 1,2-dimethoxyethane. 31b. Add 9.1 mL (65.2 mmol) triethylamine and 6.2 mL (59.4 mmol) 50% aq. H3 PO2 . 32b. Heat the mixture to 90◦ C and add a solution of 0.59 g (3.56 mmol) AIBN in 18 mL of 1,2-dimethoxyethane in three portions at 30-min intervals. Stir an additional 30 min at 90◦ C after the final addition. 33b. Cool the solution to room temperature and then add 300 mL CH2 Cl2 and 200 mL water for phase separation. 34b. Concentrate the organic layer to dryness using a rotary evaporator. Add 80 mL MeOH to the residue, heat the mixture to 60◦ C, and stir for 2 hr at 60◦ C. 35b. Cool the mixture to 0◦ C and filter the resulting slurry under vacuum. 36b. Dry the crystal overnight in a 50◦ C vacuum oven to give S.9 (4.6 g, 92% wt purity, 76% yield). Characterize the product (step 30a) and proceed to detritylation (step 37).
Perform detritylation 37. Suspend 300 mg (0.605 mmol) S.9 in 18 mL MeOH. Add 1.2 mL conc. HCl and stir for 1.5 hr at room temperature. 38. Add 4 mL Amberlite IRA 900 (OAc– form) and stir for 5 min. 39. Remove the resin by filtration using a water aspirator and wash with 10 mL MeOH. Concentrate the filtrate to 2 mL using a rotary evaporator. 40. Dilute the solution with 40 mL water. Wash three times with 10 mL of CHCl3 (total 30 mL). 41. Concentrate the aqueous layer to 5 mL on a rotary evaporator to give crystrals. 42. Characterize the product by TLC (on silica gel), elemental analysis, UV, MS, and 1 H NMR. The compound is stable for at least 12 months when stored at ambient temperature. 9-(2,3-Dideoxy-2-fluoro-β-D-threo-pentofuranosyl)adenine (FddA, S.10). Yield of prism-like crystals: 135 mg (88%). m.p. 226.8◦ -227.7◦ C (lit 227◦ C). TLC (5:1 v/v toluene/MeOH): Rf = 0.17. Anal. calcd. for C10 H12 FN5 O2 : C, 47.43; H, 4.78; N, 27.66%; found: C, 47.60; H, 4.94; N, 27.47. MS: m/z 253 (M+ ), 164 (adenine+CHO). UV λmax (MeOH): 258 nm. 1 H NMR ([2 H6 ]DMSO): δ 8.26 (1H, d, J = 2.5 Hz, H8), 8.16 (1H, s, H2), 7.34 (2H, br s, NH2 ), 6.32 (1H, dd, J = 15.9, 4.1 Hz, H1 ), 5.43 (1H, ddt, J = 54.4(d), 5.8(d), 3.5(t) Hz, H2 ), 5.06 (1H, dd, J = 5.8, 6.0 Hz, 5 -OH), 4.16-4.20 (1H, m, H4 ), 3.58-3.65 (2H, m, H5 ), 2.52-2.62 (1H, m, H3 a), 2.22-2.31 (1H, m, H3 b).
SYNTHESIS OF FdaraA FROM 6-CHLOROPURINE 3 -O-BENZOYL-5 -O-TRITYLRIBOSIDE
ALTERNATE PROTOCOL
Preparation of FdaraA proceeds as described for FddA in the initial fluorination and 3 -deprotection steps. Acid treatment of S.5 yields the detritylated product.
Additional Materials (also see Basic Protocol 2) 9-(5-O-Trityl-2-deoxy-2-fluoro-β-D-arabinofuranosyl)adenine (S.5; see Basic Protocol 2) Trifluoroacetic acid, 98+% Aspirator Current Protocols in Nucleic Acid Chemistry
Synthesis of Modified Nucleosides
1.12.11 Supplement 25
1. Dissolve 100 mg (0.195 mmol) S.5 in 2 mL of 80% trifluoroacetic acid and keep for 10 min at room temperature. 2. Evaporate to dryness under reduced pressure with an aspirator, then co-evaporate with 5 mL water. 3. Partition the residue between 5 mL CHCl3 and 10 mL water. Wash the aqueous layer with 5 mL CHCl3 . 4. Neutralize the aqueous layer with triethylamine and then evaporate to dryness on a rotary evaporator. 5. Crystallize the solid from 150 mL EtOH overnight at room temperature. 6. Characterize the product by elemental analysis, UV, MS, and 1 H NMR. The compound is stable for at least 12 months when stored at ambient temperature. 9-(2-Deoxy-2-fluoro-β-D-arabinofuranosyl)adenine (FdaraA, S.6). Yield of white crystals: 39 mg (74%). m.p. 229◦ -232◦ C (lit 231◦ -234◦ C). Anal. calcd. for C10 H12 FN5 O3 : C, 44.61; H, 4.49; N, 26.01%; found: C, 44.36; H, 4.62; N, 25.57. MS: m/z 269 (M+ ), 164 (adenine+CHO). UV λmax (MeOH): 258 nm. 1 H NMR ([2 H6 ]DMSO): δ 8.24 (1H, d, J = 0.9 Hz, H8), 8.17 (1H, s, H2), 7.35 (2H, br s, NH2 ), 6.41 (1H, dd, J = 14.5, 4.6 Hz, H1 ), 5.96 (1H, d, J = 5.0 Hz, 3 -OH), 5.05-5.36 (2H, m, H2 , 5 -OH), 4.38-4.54 (1H, m, H3 ), 3.82-3.90 (1H, m, H4 ), 3.60-3.74 (2H, m, H5 ). BASIC PROTOCOL 3
Figure 1.12.3
PREPARATION OF 6-CHLOROPURINE 3 -DEOXYRIBOSIDE The sequence of the reactions shown in Figure 1.12.3 illustrates a practical procedure for preparation of 6-chloropurine 3 -deoxyriboside (S.15), an alternate intermediate to FddA. Acetoxybromination of inosine (S.11) and then debromination by radical reduction using hypophosphorous acid and a water-soluble radical initiator gives the purine 3 -deoxyriboside analog S.13. Chlorination of the 6-position of the purine base by Vilsmeier conditions, followed by removal of the acetyl groups, gives 6-chloropurine 3 -deoxyriboside S.15.
Preparation of 6-chloropurine-3 -deoxyriboside from inosine.
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Materials Inosine (S.11), 98+% Acetic acid, 99.5+% Trimethyl orthoacetate, 99+% Acetonitrile (CH3 CN), 99.8+% Acetyl bromide, 97+% 85:15 and 80:20 (v/v) 0.02 M aq. KH2 PO4 (pH 3.2) in CH3 CN 25% (w/v) aq. sodium hydroxide (NaOH) 50% (v/v) aq. hypophosphorous acid (H3 PO2 ) Triethylamine, 99+% 6 M hydrochloric acid (HCl) 2,2 -Azobis[2-(2-imidazolin-2-yl)propane]dihydrochloride, 97+% (VA-044; Wako Pure Chemical Industries) Dichloromethane (CH2 Cl2 ), 99+% Dimethylformamide (DMF), 99.5+% Thionyl chloride, 98+% Saturated aq. sodium hydrogen carbonate (NaHCO3 ), 99.5+% Saturated aq. sodium chloride (NaCl), 99+% Sodium sulfate (Na2 SO4 ), anhydrous, 98+% Methanol (MeOH), 99.8+% 28% (w/w) sodium methoxide solution in methanol 1- and 2-L four-neck round-bottom flasks Mechanical stirrer Thermometers Reflux condensers Vacuum oil pump HPLC with Inertsil ODS 2 4.6 × 150–mm column (GL Science) Dropping funnel Separatory funnel Rotary evaporator with vacuum pump or water aspirator 1-L one-neck round-bottom flask 60-mm-diameter Kiriyama filter funnel (glass B¨uchner-type funnel) and no. 5B filter paper 50◦ C vacuum oven 300-mL three-neck round-bottom flask Perform acetoxybromination 1. To a 1-L four-neck round-bottom flask equipped with a mechanical stirrer, a thermometer, and a reflux condenser, add 150 g (559.2 mmol) inosine (S.11) and 450 mL acetic acid. 2. Add 100.7 g (838.3 mmol) trimethyl orthoacetate and stir for 3 hr at 35◦ C. 3. Concentrate the reaction mixture to half volume under reduced pressure using a vacuum oil pump. Keep the inner temperature at 35◦ C. 4. Add 150 mL acetic acid and concentrate again as in step 3. 5. Dissolve the concentrate in 300 mL CH3 CN and cool to 5◦ C. 6. Slowly add 103.3 mL (1.4 mol) acetyl bromide over 1 hr. This step is exothermic. Keep the inner temperature <10◦ C.
7. Stir the reaction mixture for 3 hr at 10◦ C. 8. Monitor the reaction by HPLC using HPLC condition A as follows:
Synthesis of Modified Nucleosides
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Column: Inertsil ODS 2 4.6 × 150–mm Mobile phase: 85:15 (v/v) 0.02 M aq. KH2 PO4 (pH 3.2) in CH3 CN Flow rate: 1.0 mL/min Column temperature: 40◦ C Injection volume: 10 µL Detection: UV at 262 nm. Under these conditions, the retention times are 14.0 min for S.12 and 12.4 min for the byproduct, triacetylinosine. The ratio of target compound/triacetylinosine is normally ∼7:1 to 9:1 (by HPLC peak area ratio).
9. Use a dropping funnel to add the reaction mixture dropwise into 600 mL of ice-cold 1:1 (v/v) water/CH3 CN while controlling the pH at ∼5 to 6 by dropwise addition of 25% aq. NaOH. 10. Adjust to pH 7 using 25% NaOH. 11. Separate the layers using a separatory funnel, and then extract the aqueous layer with 300 mL CH3 CN. 12. Combine the organic layers and concentrate to a minimum volume using a rotary evaporator with a water aspirator to obtain the target compound (150.3 g, 65% yield, calculated by quantitative HPLC analysis). This results in 30% to 50% wt. of the target compound in a solution of CH3 CN. This can be used without further purification in the next reaction. An analytical sample can be obtained by silica gel column chromatography using 5:1 (v/v) CHCl3 /MeOH, followed by crystallization from CH3 CN.
13. Characterize the product by NMR, IR, UV, and MS. 9-(2,5-Di-O-acetyl-3-bromo-3-deoxy-β - D -xylofuranosyl)-1,9-dihydro-6H-purine6-one (S.12). m.p. 171◦ -172◦ C. 1 H NMR (300 MHz, CDCl3 ): δ 8.34 (s, 1H, H2), 8.24 (s, 1H, H8), 6.20 (bs, 1H, H1 ), 5.74 (bs, 1H, H2 ), 4.4-4.6 (m, 4H, H3 , H4 , H5 ab), 2.20 (s, 3H, 5 O-Ac), 2.14 (s, 3H, 2 O-Ac). 13 C NMR (75 MHz, CDCl3 ): δ 170.4, 158.8, 148.4, 146.0, 138.4, 124.2, 87.9, 83.0, 78.6, 64.7, 20.7. IR (KBr): ν 1750, 1698, 1376, 1226, 1044 cm−1 . UV λmax (MeOH): 206, 245 nm. HRMS (FAB+) calcd. for C14 H16 N4 O6 Br (M + H)+ 415.0253; found 415.0248.
Remove bromine 14. In a 1-L round-bottom flask, dilute 118.6 g (898 mmol) 50% aq. H3 PO2 with 620 mL water, and then add 90.9 g (898 mmol) triethylamine at 18◦ C. 15. To a separate 2-L four-neck round-bottom flask equipped with a mechanical stirrer, a thermometer, and a reflux condenser, add 327 g CH3 CN solution of S.12 (38.0% wt., 299 mmol). 16. Add the H3 PO2 /triethylamine solution to the solution of S.12 and heat to 50◦ C. During heating, adjust pH to 7 to 8 by adding triethylamine and 6 M HCl. 17. Add 9.7 g (30.0 mmol) VA-044 in 50 mL water. 2,2 -Azobis(2-methylpropionamide) dihydrochloride (V-50) is also useful for this reaction. VA-044 and V-50 are known to be water-soluble azo polymerization initiators, and are available from Wako Pure Chemical Industries. A common radical initiator, 2,2 -azobis(2-methylpropionitrile) (AIBN), is also effective for this reaction, but it co-precipitates with the product due to its low solubility in aqueous media. Therefore, a water-soluble radical initiator is preferable. Synthesis of Fluorinated Nucleosides
18. Stir the reaction mixture for 4 hr at 50◦ C, controlling the pH at 3 to 4 by adding 25% NaOH.
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19. Perform HPLC analysis using HPLC condition A (see step 8) to check completion of the reaction. Under these conditions, the retention times are 6.8 min for S.13, 10.9-11.0 min for triacetylinosine, and 11.5-12.3 min for S.12.
20. Cool to 10◦ C on ice and adjust to pH 6 by adding 25% NaOH. After cooling, the target compound precipitates. Because the product obtained in this step contains inorganic salts, it should be purified as described in the following steps.
21. Filter the resulting slurry through a 60-mm Kiriyama funnel (no. 5B filter paper) under vacuum, and wash with 375 mL water. 22. Add the wet crystal to 1 L of 9:1 (v/v) CH3 CN/water. 23. Heat to 60◦ C, stir the slurry for 1 hr, and then cool to 10◦ C. 24. Filter the product through a 60-mm Kiriyama funnel (no. 5B filter paper) under vacuum and rinse with 375 mL water. 25. Dry overnight in a 50◦ C vacuum oven to give the product S.13 (69 g, 94% wt. purity, 65% yield). 26. Characterize the product by MS and NMR. 2 ,5 -Di-O-acetyl-3 -deoxyinosine (S.13). 1 H NMR (300 MHz, CDCl3 ): δ 8.08 (s, 1H, H2), 8.07 (s, 1H, H8), 6.04 (d, 1H, J = 1.1 Hz, H1 ), 5.59 (bd, 1H, J = 5.9 Hz, H2 ), 4.60 (m, 1H, H4 ), 4.39 (dd, 1H, J = 12.3, 2.9 Hz, H5 a), 4.22 (dd, 1H, J = 12.3, 5.2 Hz, H5 b), 2.50 (ddd, 1H, J = 14.0, 10.5, 5.9 Hz, H3 a), 2.16 (ddd, 1H, J = 14.0, 5.8, 1.1 Hz, H3 b), 2.09 (s, 3H, 5 O-Ac), 2.04 (s, 3H, 2 O-Ac). 1 H NMR (300 MHz, DMSO-d6 ): δ 8.26 (s, 1H, H2), 8.10 (s, 1H, H8), 6.11 (d, 1H, J = 1.4 Hz, H1 ), 5.61 (bd, 1H, J = 6.3 Hz, H2 ), 4.52 (m, 1H, H4 ), 4.29 (dd, 1H, J = 12.0, 2.9 Hz, H5 a), 4.16 (dd, 1H, J = 12.0, 5.8 Hz, H5 b), 2.60 (ddd, 1H, J = 14.1, 10.3, 6.3 Hz, H3 a), 2.22 (ddd, 1H, J = 14.1, 5.9, 1.1 Hz, H3 b), 2.10 (s, 3H, 5 O-Ac), 1.99 (s, 3H, 2 O-Ac). 13 C NMR (75 MHz, DMSO-d6 ): δ 170.3, 170.1, 156.7, 147.9, 146.2, 138.9, 124.6, 88.6, 78.2, 77.7, 64.6, 32.6, 20.9, 20.7. HRMS (FAB+) calcd. for C14 H17 N4 O6 (M + H)+ 337.1148; found 337.1153.
Introduce chlorine 27. In a 1 L four-neck round-bottom flask equipped with a thermometer and a reflux condenser, suspend 37.6 g (94% wt., 105 mmol) S.13 in 475 mL CH2 Cl2 . 28. Add 32.8 mL (423 mmol) DMF and 30.9 mL (423 mmol) thionyl chloride and stir with magnetic stirring. The reaction is exothermic. Keep the inner temperature <30◦ C.
29. Reflux the reaction mixture for 6.5 hr at 50◦ C. 30. Check completion of the reaction by HPLC analysis using HPLC condition B as follows:
Column: Inertsil ODS 2 4.6 × 150–mm Mobile phase: 80:20 (v/v) 0.02 M aq. KH2 PO4 (pH 3.2) in CH3 CN Flow rate: 1.0 mL/min Column temperature: 40◦ C Injection volume: 10 µL Detection: UV at 262 nm. Under these conditions, the retention times are 1.7 min for hypoxantine, 2.4 min for 6-chloropurine, 4.0 min for S.13, and 16.8 min for S.14.
31. Cool the reaction mixture to 10◦ C and pour into 400 mL ice-cold water.
Synthesis of Modified Nucleosides
1.12.15 Current Protocols in Nucleic Acid Chemistry
Supplement 25
32. Separate the layers and wash the organic layer with 300 mL water, 400 mL saturated aq. NaHCO3 , and then 400 mL saturated aq. NaCl. 33. Dry over anhydrous Na2 SO4 until the solution becomes clear, and filter off the drying agent through a 60-mm Kiriyama funnel (no. 5A filter paper) under vacuum. 34. Remove solvent under reduced pressure using a rotary evaporator with a water aspirator, and use a vacuum oil pump to obtain the crude product S.14 as a yellow foam (33.5 g, 89.9% yield). 35. Analyze the compound by NMR, IR, and MS. 6-Chloro-9-(2,5-di-O-acetyl-3-deoxy-β-D-erythro-pentofuranosyl)-9H-purine (S.14). 1 H NMR (300 MHz, CDCl3 ): δ 8.77 (s, 1H, H2), 8.32 (s, 1H, H8), 6.15 (d, 1H, J = 1.3 Hz, H1 ), 5.74 (d, 1H, J = 5.8 Hz, H2 ), 4.63-4.74 (m, 1H, H4 ), 4.46 (dd, 1H, J = 12.3, 2.8 Hz, H5 a), 4.30 (dd, 1H, J = 12.3, 5.1 Hz, H5 b), 2.65 (ddd, 1H, J = 14.1, 10.6, 6.0 Hz, H3 a), 2.27 (ddd, 1H, J = 14.1, 5.6, 1.3 Hz, H3 b), 2.17 (s, 3H, 5 O-Ac), 2.08 (s, 3H, 2 O-Ac). 1 H NMR (300 MHz, DMSO-d6 ): δ 8.85 (s, 2H, H2, H8), 6.31 (s, 1H, H1 ), 5.76 (s, 1H, H2 ), 4.55-4.64 (m, 1H, H4 ), 4.32 (dd, 1H, J = 12.1, 2.8 Hz, H5 a), 4.22 (dd, 1H, J = 12.1, 5.8 Hz, H5 b), 2.67 (ddd, 1H, J = 14.2, 10.3, 6.1 Hz, H3 a), 2.01-2.32 (m, 1H, H3 b), 2.13 (s, 3H, 5 O-Ac), 1.97 (s, 3H, 2 O-Ac). 13 C NMR (75 MHz, DMSO-d6 ): δ 170.3, 170.1, 152.0, 151.3, 149.6, 140.6, 131.5, 89.2, 78.6, 77.4, 64.5, 32.4, 20.9, 20.6. IR (neat): ν 1745, 1674, 1592, 1562, 1387, 1339, 1232, 1094, 1053 cm−1 . HRMS (FAB+) calcd. for C14 H16 N4 O5 Cl (M + H)+ 355.0809; found 355.0816.
Remove acetyl protection 36. In a 300-mL three-neck round-bottom flask equipped with a thermometer, dissolve 26.3 g (74.1 mmol) S.14 in 39.5 mL MeOH. 37. Cool the mixture at 5◦ C and add 2.8 g of 28% sodium methoxide in methanol (14.5 mmol). 38. Stir the mixture for 3 hr at ambient temperature. The clear solution turns to a thick, white slurry.
39. Employ HPLC analysis using HPLC condition A (see step 8) to check completion of the reaction. Under these conditions, the retention times are 3.4-3.5 min for the fully deprotected S.15 and 9.7 min for a monoacetyl intermediate.
40. Cool the reaction mixture to 5◦ C and stir for 2 hr. 41. Filter the product through a 40-mm Kiriyama funnel (no. 5B filter paper) under vacuum and rinse with cold MeOH. 42. Dry overnight in a 50◦ C vacuum oven to give the target compound S.15 (14.7 g, 98% wt. purity, 71% yield). It is possible to recover a second crop from the mother liquor. After concentrating the mother liquor to about half the volume, filter the solid precipitate and purify by recrystallization from 25% (v/v) aqueous MeOH to produce pure S.15 (2.4 g, ∼45% recovery). An analytical sample is obtainable by recrystallization from 1:3 (v/v) MeOH/water.
43. Analyze the compound by melting point, UV, IR, MS, and NMR.
Synthesis of Fluorinated Nucleosides
1.12.16 Supplement 25
6-Chloro-9-(3-deoxy-β-D-erythro-pentofuranosyl)-9H-purine (S.15). m.p. 180◦ -181◦ C. 1 H NMR (300 MHz, CDCl3 ): δ 8.68 (s, 1H, H2), 8.33 (s, 1H, H8), 5.83 (d, 1H, J = 4.6 Hz, H1 ), 4.92 (ddd, 1H, J = 7.2, 6.5, 4.6 Hz, H2 ), 4.53-4.59 (m, 1H, H4 ), 3.98 (dd, 1H, J = 12.5, 2.1 Hz, H5 a), 3.60 (dd, 1H, J = 12.5, 2.6 Hz, H5 b), 2.53 (ddd, 1H, J = 12.9, 7.2, 5.7 Hz, H3 a), 2.18 (ddd, 1H, J = 12.9, 8.0, 6.5 Hz, H3 b). 1 H NMR (300 MHz, DMSO-d6 ): δ 8.97 (s, 1H, H2), 8.82 (s, 1H, H8), 6.06 (d, 1H, J = 1.4 Hz, H1 ), 5.80 (d, Current Protocols in Nucleic Acid Chemistry
1H, J = 3.9 Hz, 2 -OH), 5.12 (dd, 1H, J = 5.3, 5.2 Hz, 5 -OH), 4.62-4.68 (m, 1H, H2 ), 4.42-4.50 (m, 1H, H4 ), 3.78 (ddd, 1H, J = 12.1, 5.3, 3.2 Hz, H5 a), 3.59 (ddd, 1H, J = 12.1, 5.2, 3.8 Hz, H5 b), 2.28 (ddd, 1H, J = 13.3, 9.6, 5.3 Hz, H3 a), 1.93 (ddd, 1H, J = 13.3, 6.0, 2.2 Hz, H3 b). 13 C NMR (75 MHz, DMSO-d6 ): δ 151.8, 151.3, 149.3, 145.4, 131.5, 91.6, 81.9, 75.2, 62.1, 33.6. IR (KBr): ν 3331, 1596, 1562, 1442, 1405, 1391, 1337, 1207, 1129, 1079, 1068 cm−1 . UV λmax (MeOH): 204, 265 nm. HRMS (FAB+) calcd. for C10 H12 N4 O3 Cl (M + H)+ 271.0598; found 271.0584.
PREPARATION OF FddA FROM 6-CHLOROPURINE 3 -DEOXYRIBOSIDE The sequence of reactions shown in Figure 1.12.4 illustrates a useful procedure for a practical synthesis of FddA (S.20) from 6-chloropurine 3 -deoxyriboside (S.15). Trityl protection of the C5 -OH of S.15 followed by fluorination at the C2 -β position by perfluorobutanesulfonyl fluoride affords the fluorinated compound S.17 along with the byproduct S.18. Selective decomposition of S.18 in an acidic medium allows for isolation of S.17. Finally, amination of the C6 position of the purine base and subsequent detritylation gives FddA in good yields.
Figure 1.12.4
BASIC PROTOCOL 4
Preparation of FddA from 6-chloro-9-(3-deoxy-β-D-erythro-pentofuranosyl)-9H-purine.
Materials 6-Chloro-9-(3-deoxy-β-D-erythro-pentofuranosyl)-9H-purine (S.15) Acetonitrile (CH3 CN), 99.8+% Nitrogen 2,4,6-Collidine, 99+% Trityl chloride, 98+% Mobile phase: HPLC-grade MeOH, far-UV-grade CH3 CN, and 0.1% H3 PO4 (see Tables 1.12.2 and 1.12.3 for gradients) Methanol (MeOH), 99.8+% Dichloromethane (CH2 Cl2 ), 99+% 6 M hydrochloric acid (HCl) Saturated aq. sodium hydrogencarbonate (NaHCO3 ), 99.5+% Sodium sulfate (Na2 SO4 ), anhydrous, 98+% Benzotrifluoride, 98+% n-Hexane, 95+% Toluene, 99.5+%
Synthesis of Modified Nucleosides
1.12.17 Current Protocols in Nucleic Acid Chemistry
Supplement 25
Perfluorobutanesulfonyl fluoride, 90+% N,N-Dimethylcyclohexylamine, 98+% 5.0% aq. ammonium chloride (NH4 Cl), 99.5+% 80% aq. acetic acid, 99.7+% Cyclohexane, 99.5+% Tetrahydrofuran, 99.5+% Ammonia (NH3 ), anhydrous, 99.99+% 35% to 37% hydrochloric acid (conc. HCl) 25% aq. ammonia (NH3 ) Ethyl acetate (EtOAc), 99.5+% 500-mL and 1-L round-bottom flasks Reflux condensers HPLC with Zorbax SB-phenyl 4.6 × 250–mm column (Chrompack) Rotary evaporator with vacuum pump and water aspirator Separatory funnel 60- and 40-mm Kiriyama filter funnels (glass B¨uchner-type funnel) with no. 5A and 5B filter papers 50◦ C vacuum oven 500-mL four-neck round-bottom flasks 1-L stainless steel pressure vessel equipped with a pressure gauge Water trap unit Glass filter holder with PTFE membrane filter Perform 5 -tritylation 1. In a 500-mL round-bottom flask equipped with a reflux condenser, suspend 20 g (99% purity, 73.1 mmol) S.15 in 140 mL anhydrous CH3 CN and stir with magnetic stirring under a nitrogen stream. As trityl chloride is moisture sensitive, the reaction should be carried out under dry conditions.
2. Add 11.6 mL (87.7 mmol) 2,4,6-collidine and 22.4 g (80.4 mmol) trityl chloride. 3. Heat the reaction mixture to 45◦ C and stir for 3 hr under a nitrogen atmosphere. 4. Monitor the reaction by HPLC using HPLC condition C as follows:
Column: Zorbax SB-phenyl 4.6 × 250–mm (Chrompack) Mobile phase: see Table 1.12.2 for gradient Flow rate: 1.0 mL/min Column temperature: ambient Injection volume: 10 µL Detection: UV at 262 nm (220 nm for analysis of non-nucleoside impurities). Under these conditions, the retention times are 5.0-5.5 min for S.15, 13.5-14.5 min for toluene, 18.4 min for triphenylmethyl alcohol (TrOH), 20.0 min for S.16, 23.0 min for methyltriphenylmethyl ether (TrOMe), and 29.0 min for a 2 ,5 -di-O-trityl impurity.
5. Add 1.5 mL (37.1 mmol) MeOH and stir the mixture for 1 hr at 45◦ C. Quenching with methanol leads to the formation of TrOMe, which is easily removed during crystallization.
6. Cool to room temperature. Concentrate the reaction mixture to a minimum volume using a rotary evaporator with a water aspirator. Synthesis of Fluorinated Nucleosides
1.12.18 Supplement 25
Current Protocols in Nucleic Acid Chemistry
Table 1.12.2 Gradient Program for HPLC Condition Ca
Time (min)
% aq. H3 PO4
% CH3 CN
% MeOH
0
60
10
30
5
60
10
30
8
30
60
10
16
30
60
10
19
10
80
10
32
10
80
10
35
60
10
30
45
60
10
30
a Using HPLC-grade MeOH, far-UV-grade CH CN, and 0.1% H PO 3 3 4
(2.94 mL 85% H3 PO4 adjusted to 2.5 L with deionized water).
7. Add 100 mL water and 200 mL CH2 Cl2 . Adjust to pH 3 with 6 M HCl, and separate the layers using a separatory funnel. The aqueous layer contains 2,4,6-collidine, HCl, and CH3 CN.
8. Add 100 mL water to the organic layer and adjust to pH 3 with 6 M HCl, then wash and separate the layers. 9. Add 100 mL water to the organic layer and adjust to pH 7 with saturated aq. NaHCO3 , then wash and separate the layers. 10. Dry the organic layer over anhydrous Na2 SO4 until the solution becomes clear, filter off the drying agent through a 60-mm Kiriyama funnel (no. 5A filter paper) under vacuum, and remove the solvent under reduced pressure using a rotary evaporator with a water aspirator. 11. Add 220 mL benzotrifluoride and concentrate to a minimum volume under reduced pressure. CH2 Cl2 is removed by co-evaporation with benzotrifluoride.
12. Add 180 mL benzotrifluoride, heat to 80◦ C, and then cool to room temperature. The product S.16 is crystallized from benzotrifluoride.
13. Filter the crystals over a 60-mm Kiriyama funnel (no. 5B filter paper) under vacuum and wash with 50 mL benzotrifluoride and 50 mL n-hexane. 14. Dry overnight in a 50◦ C vacuum oven to obtain the product S.16 (31.9 g, 85.0% yield). 15. Analyze the product by melting point, UV, IR, MS, and NMR. 6-Chloro-9-[3-deoxy-5-O-(triphenylmethyl)-β - D-erythro-pentofuranosyl]-9H-purine (S.16). m.p. 205◦ -206◦ C. 1 H NMR (300 MHz, CDCl3 ): δ 8.64 (s, 1H, H2), 8.40 (s, 1H, H8), 7.21-7.41 (m, 15H, 5 -OTr), 6.04 (d, 1H, J = 2.2 Hz, H1 ), 4.85-4.91 (m, 1H, H2 ), 4.68-4.78 (m, 1H, H4 ), 3.44 (dd, 1H, J = 10.6, 3.1 Hz, H5 a), 3.33 (dd, 1H, J = 10.6, 4.6 Hz, H5 b), 2.30 (ddd, 1H, J = 13.3, 7.7, 5.6 Hz, H3 a), 2.17 (ddd, 1H, J = 13.3, 6.5, 3.9 Hz, H3 b). 13 C NMR (75 MHz, CDCl3 ): δ 151.4, 151.2, 150.4, 146.7, 143.3, 132.4, 128.5, 127.8, 127.2, 93.1, 87.0, 80.7, 76.0, 64.3, 33.9. IR (KBr): ν 3354, 3059, 1592, 1562, 1491, 1449, 1440, 1338, 1206, 1130, 1078, 1018 cm−1 . UV λmax (MeOH): 207, 265 nm. HRMS (FAB+) calcd. for C29 H26 N4 O3 Cl (M + H)+ 513.1693; found 513.1717.
Synthesis of Modified Nucleosides
1.12.19 Current Protocols in Nucleic Acid Chemistry
Supplement 25
Perform fluorination 16. In a 500-mL four-neck round-bottom flask equipped with a mechanical stirrer, a thermometer, and a reflux condenser, suspend 30.0 g (58.5 mmol) S.16 in 300 mL toluene. 17. Add 23.3 mL (117 mmol) perfluorobutanesulfonyl fluoride and 17.7 mL (117 mmol) N,N-dimethylcyclohexylamine at 20◦ C. Although conventional fluorinating reagents such as DAST and MOST can be effective for this reaction, for safety purposes, perfluorobutanesulfonyl fluoride is highly preferable (see Commentary).
18. Heat to an internal temperature of 50◦ C and stir the reaction mixture for 23 hr. 19. Monitor the reaction by HPLC using HPLC condition D as follows:
Column: Zorbax SB-phenyl 4.6 × 250–mm (Chrompack) Mobile phase: see Table 1.12.3 for gradient Flow rate: 1.0 mL/min Column temperature: ambient Injection volume: 10 µL Detection: UV at 262 nm (220 nm for the analysis of non-nucleoside impurities). Under these conditions, the retention times are 3.4 min for 6-chloropurine (a deribose byproduct), 4.6-4.7 min for toluene, 5.0 min for N,N-dimethylcyclohexylamine, 7.7-7.8 min for S.16, 10.9-11.0 min for S.18, 11.5-12.3 min for S.17, 13.5 min for 2 -(trityloxymethyl)furane, 16.5 min for a 5 -O-trityl-2 -perfluorobuntanesulfonate intermediate, and 19.5 min for a 2 ,5 -di-O-trityl impurity. The elimination product (S.18) is obtained as a byproduct. The ratio of S.17 to S.18 is 7:3 (HPLC peak area ratio).
20. Add 75 mL of 5.0% aq. NH4 Cl with stirring at 50◦ C and then separate the layers. During extraction at 50◦ C, the mixture forms three layers. The lowest layer contains mostly perfluoro compounds, the middle layer contains water, and the upper layer contains the organic compounds.
21. Extract the lower layer with 30 mL toluene at 50◦ C. 22. Wash the combined toluene layer with 75 mL of 5.0% aq. NH4 Cl at 50◦ C and separate the layers. 23. Wash the organic layer with 75 mL water at 50◦ C and separate the layers. 24. Concentrate the organic layer to half the volume under reduced pressure using a rotary evaporator with a water aspirator. Table 1.12.3 Gradient Program for HPLC Condition Da
Synthesis of Fluorinated Nucleosides
1.12.20 Supplement 25
Time (min)
% aq. H3 PO4
% CH3 CN
% MeOH
0
30
60
10
8
30
60
10
10
10
80
10
20
10
80
10
22
30
60
10
30
30
60
10
a Using HPLC-grade MeOH, far-UV-grade CH CN, and 0.1% H PO 3 3 4
(2.94 mL 85% H3 PO4 adjusted to 2.5 L with deionized water). Current Protocols in Nucleic Acid Chemistry
25. Cool to 10◦ C. Add 150 mL of 80% aq. acetic acid and stir the reaction mixture for 3 hr at 10◦ C. The fluorinated compound (S.17) is stable in an acidic medium due to the effect of the C2 -β fluoride. In contrast, the elimination product (S.18) is acid-labile due to acidolysis of the glycosyl bond. Therefore, S.17 can be purified by selective decomposition of S.18 by means of treatment with acetic acid.
26. Separate the layers and wash the organic layer with 150 mL water. 27. Add 150 mL cyclohexane to crystallize the product and stir the mixture for 1 hr at 20◦ C. 28. Cool to 0◦ C and stir for 1 hr. 29. Filter the crystals over a 60-mm Kiriyama funnel (no. 5B filter paper) under vacuum and wash with 150 mL cyclohexane. 30. Dry overnight in a 50◦ C vacuum oven to obtain the product S.17 (15.0 g, 49.6% yield). 31. Characterize the product by UV, IR, MS, and NMR. 6-Chloro-9-[2,3-dideoxy-2-fluoro-5-O-(triphenylmethyl)-β-D-threo-pentofuranosyl]9H-purine (S.17). 1 H NMR (300 MHz, CDCl3 ): δ 8.73 (s, 1H, H2), 8.34 (d, 1H, J = 2.8 Hz, H8), 7.22-7.52 (m, 15H, 5 -OTr), 6.41 (dd, 1H, J = 19.1, 3.1 Hz, H1 ), 5.25 (dddd, 1H, J = 53.7, 5.2, 3.1, 2.0 Hz, H2 ), 4.42-4.50 (m, 1H, H4 ), 3.48 (dd, 1H, J = 9.9, 6.6 Hz, H5 a), 3.30 (dd, 1H, J = 9.9, 3.8 Hz, H5 b), 2.57 (dddd, 1H, J = 35.0, 14.8, 9.0, 5.6 Hz, H3 a), 2.36 (dddd, 1H, J = 27.5, 15.1, 5.1, 1.7 Hz, H3 b). 13 C NMR (75 MHz, CDCl3 ): δ 152.0, 151.2 (d, J = 16.8 Hz), 144.7 (d, J = 5.8 Hz), 143.6, 128.6, 127.9, 127.2, 90.6 (d, J = 188.9 Hz), 86.9, 85.0 (d, J = 16.4 Hz), 76.8, 65.7, 33.8 (d, J = 20.5 Hz). IR (KBr): ν 1593, 1567, 1492, 1220, 1206, 1079 cm−1 . UV λmax (MeOH): 206, 264 nm. HRMS (FAB+) calcd. for C29 H25 N4 O2 FCl (M + H)+ 515.1650; found 515.1658. Anal. calcd. for C29 H24 N4 O2 FCl: C, 67.64; H, 4.70; N, 10.88; Cl, 6.88%; found: C, 67.34; H, 4.81; N, 10.61; Cl, 6.98.
Perform amination 32. Fill a 1-L stainless steel pressure vessel with 15.0 g (29 mmol) S.17 and 250 mL tetrahydrofuran. The starting material dissolves in the tetrahydrofuran.
33. Blow NH3 gas into the solution at a pressure of 2.5 bar. The NH3 gas slowly dissolves into the solution. It may take ∼1 hr to add NH3 at a pressure of 2.5 bar. Stir for an additional 1 hr to make sure the pressure stabilizes.
34. Heat to 70◦ C and stir the reaction mixture for 72 hr. The pressure will rise from 2.5 bar to a maximum of 8 bar. During the reaction some crystallization of NH4 Cl occurs.
35. Cool to 20◦ C and release NH3 gas to a water trap unit filled with water. 36. Employ HPLC analysis using HPLC condition D (see step 19) to check that the reaction has proceeded to completion. Under these conditions, the retention times are 6.5 min for S.19, 10.9-11.0 min for S.18, 11.5-12.3 min for S.17, and 13.5 min for 2-(trityloxymethyl)furane.
37. Concentrate the solution to a minimum volume. This concentration results in a very thick slurry that can be used directly in the next reaction.
38. Characterize the product S.19 by UV, IR, MS, and NMR.
Synthesis of Modified Nucleosides
An analytical sample can be obtained by recrystallization from methanol.
1.12.21 Current Protocols in Nucleic Acid Chemistry
Supplement 25
9-[2,3-Dideoxy-2-fluoro-5-O-(triphenylmethyl)-β - D -threo-pentofuranosyl]adenine (S.19). m.p. 235◦ -236◦ C. 1 H NMR (300 MHz, CDCl3 ): δ 8.33 (s, 1H, H2), 8.06 (d, 1H, J = 3.0 Hz, H8), 7.20-7.52 (m, 15H, 5 -OTr), 6.33 (dd, 1H, J = 19.9, 2.9 Hz, H1 ), 6.18 (bs, 2H, 6-NH2 ), 5.20 (dm, 1H, J = 53.8 Hz, H2 ), 4.35-4.45 (m, 1H, H4 ), 3.46 (dd, 1H, J = 10.0, 6.5 Hz, H5 a), 3.27 (dd, 1H, J = 10.0, 4.1 Hz, H5 b), 2.50 (dddd, J = 35.5, 14.9, 9.0, 5.4 Hz, H3 a), 2.31 (dddd, 1H, J = 27.5, 14.9, 4.8, 1.4 Hz, H3 b). 13 C NMR (75 MHz, CDCl3 ): δ 155.4, 152.9, 149.6, 143.6, 140.1 (d, J = 6.9 Hz), 128.6, 127.9, 127.1, 119.0, 90.6 (d, J = 188.6 Hz), 86.7, 84.7 (d, J = 16.7 Hz), 76.2, 65.8, 33.9 (d, J = 20.6 Hz). IR (KBr): ν 3151, 1649, 1599, 1578, 1403, 1063 cm−1 . UV λmax (MeOH): 208, 259 nm. HRMS (FAB+) calcd. for C29 H27 N5 O2 F (M+H)+ 496.2149; found 496.2141.
Perform detritylation 39. In a 1-L round-bottom flask, dilute the thick slurry of S.19 with 144 mL MeOH. 40. Add 4.8 mL of conc. (37%) HCl. Stir the solution for 4 hr at 20◦ C. This reaction is slightly exothermic. The reaction mixture turns to a clear solution.
41. Employ HPLC analysis using HPLC condition C (see step 4) to check that the reaction has proceeded to completion. Under these conditions, the retention times are 3.5 min for S.20 (FddA) and 18.1 min for S.19.
42. Neutralize the reaction mixture with a slight excess of 25% aq. NH3 to adjust the pH to ∼6. 43. Evaporate the solvent to a minimum volume using a rotary evaporator with a water aspirator. Keep the bath temperature below 35◦ C.
44. Add 43 mL EtOAc to the residue and evaporate again to a minimum volume. MeOH may cause product loss because S.20 dissolves to a certain extent in EtOAc/MeOH.
45. Add 43 mL EtOAc and 43 mL water to the residue. 46. Add 2.0 mL conc. HCl to the solution to adjust the pH to ∼2. Stir and then separate the layers. 47. Wash the aqueous layer with 14.4 mL EtOAc two times. Filter the aqueous layer on a membrane filter. 48. Add 25% aq. NH3 to the filtrate to adjust the pH value to 7 to 8. 49. Stir the suspension overnight and then cool to 0◦ C. 50. Filter the product through a 40-mm Kiriyama funnel (no.5B filter paper) under vacuum at 0◦ C, and wash three times with 14.4 mL water. 51. Dry the product for 24 hr in a 50◦ C vacuum oven to obtain FddA (S.20; 5.3 g, 99.1% purity, 73% yield in two steps). 52. Characterize of the product by UV, IR, MS, and NMR.
Synthesis of Fluorinated Nucleosides
1.12.22 Supplement 25
9-(2,3-Dideoxy-2-fluoro-β-D-threo-pentofuranosyl)adenine (FddA, lodenosine, S.20). m.p. 226◦ -227◦ C. 1 H NMR (300 MHz, DMSO-d6 ): δ 8.27 (d, 1H, J = 2.3 Hz, H8), 8.16 (s, 1H, H2), 7.33 (bs, 2H, 6-NH2 ), 6.32 (dd, 1H, J = 16.1, 3.9 Hz, H1 ), 5.43 (dm, 1H, J = 54.4 Hz, H2 ), 5.07 (t, 1H, J = 5.9 Hz, 5 -OH), 4.13-4.23 (m, 1H, H4 ), 3.54-3.69 (m, 2H, H5 ab), 2.47-2.68 (m, 1H, H3 a), 2.17-2.36 (m, 1H, H3 b). 13 C NMR (75 MHz, DMSO-d6 ): δ 156.2, 152.9, 149.3, 139.7 (d, J = 4.7 Hz), 118.4, 91.5 (d, J = 185.9 Hz), 83.8 (d, J = 16.1 Hz), 77.9, 63.1, 32.6 (d, J = 19.3 Hz). IR (KBr): ν 3328, 3208, 3136, 1661, 1076, 1061 cm−1 . UV λmax (MeOH): 204, 259 nm. HRMS (FAB+) calcd. for C10 H13 N5 O2 F (M + H)+ 254.1053; found 254.1048. Anal. calcd. for C10 H12 N5 O2 F: C, 47.43; H, 4.78; N, 27.66; F, 7.50%; found: C, 47.45; H, 4.84; N, 27.46; F, 7.13. Current Protocols in Nucleic Acid Chemistry
COMMENTARY Background Information Lodenosine activity Lodenosine (FddA) is an anti-HIV agent that is a C2 -β-fluorinated dideoxynucleoside (Herdewijn et al., 1987; Marquez et al., 1987, 1990; Ruxrungtham et al., 1996; Graul et al., 1998). Like other nucleoside derivatives such as zidovudine (AZT), 2 ,3 -dideoxyadenosine (ddA), and 2 ,3 -dideoxyinosine (ddI), FddA acts as an inhibitor of HIV reverse transcriptase (RT). FddA was chosen for evaluation as one of the most selective inhibitors in a series of 2 ,3 -dideoxyadenosines with either an azido, fluorine, or hydroxyl group substituted in the “up” or “down” position of the C2 and C3 of the ribose moiety. Although it is less active than the parent compound ddA (Herdewijn et al., 1987), FddA is stable in the acidic conditions found in the human stomach, which are known to decompose ddA and ddI (Marquez et al., 1987, 1990). Furthermore, FddA does not exhibit cross-resistance to AZT, zalcitabine (ddC), and ddI (Driscoll et al., 1997), and even has synergetic activity with AZT. FddA also acts synergistically with other anti-HIV agents including HIV protease inhibitors such as Ritonavir, thus providing new possible combination therapy protocols for HIV. Consequently, FddA has progressed into development as a novel antiHIV agent exhibiting significant advantages in terms of high stability, high oral bioavailability (Ruxrungtham et al., 1996), and effectiveness against strains resistant to existing dideoxynucleosides (Ueno and Mitsuya, 1997). Synthetic approaches The considerable attention that FddA received from antiviral researchers because of its biological activity and chemical and metabolic stability (Chu et al., 1989; Marquez et al., 1990) has led many groups to strive to develop efficient methods for synthesizing C2 -β-fluorinated purine deoxynucleosides. Although condensation of fluorinated sugar derivatives with a nucleobase is a wellknown means of obtaining C2 -β-fluorinated nucleosides (Wysocki et al., 1991; Siddiqui et al., 1994; Chou et al., 1996), the synthesis of the fluorinated sugar derivatives themselves requires many steps, and the condensation remains subject to α-anomer formation (Barchi et al., 1991). The introduction of fluorine at the 2 -(S)ara site of a nucleoside is difficult. The first problem is the poor yields associated with
3 ,5 -di-O-protection by a trityl group (Pankiewicz et al., 1992a,b). Pankiewicz et al. (1992a) reported an excellent method for the fluorination of ribosides at the C2 -β position employing advantageous conformational control to synthesize 2 -fluorinated araA (FaraA). They found that the C2 -β-fluorinated com pound could be obtained from O3 ,O5 ,N6 tritrityladenosine in 30% yield, but only in the presence of an unexpected and undesired isonucleoside in 51% yield, due to migration caused by an intramolecular nucleophilic attack of N3 at the electrophilic C2 . Recently, a new synthetic approach to FddA was reported via dehydration of a 2 deoxy-2 -fluoroadenosine and subsequent hydrogenation of the vinyl fluoride intermediate (Siddiqui et al., 1998). However, the preparation of the 2 -deoxy-2 -fluoroadenosine starts with a costly compound, 9-(β-Darabinofuranosyl)adenine (Kawasaki et al., 1993), and requires a multi-step synthesis. This unit presents methods for fluorination of 6-chloropurine derivatives, which are subsequently aminated to give the desired FddA. Fluorination of a 6-chloropurine riboside The first approach (see Basic Protocols 1 and 2) is an efficient method for preparing FddA and FdaraA (S.6; Maruyama et al., 1996) from a 3 -O-benzoyl-5 -O-tritylriboside (S.3; Fig. 1.12.2; Maruyama et al., 1999). FdaraA is an important fluorinated nucleoside that has potent anti-tumor activities (Sato et al., 1984; Parker et al., 2003). One of the major advantages of this method is that these biologically important nucleosides, FddA and FdaraA, can be synthesized via a common intermediate, 9-(5-O-trityl-2-deoxy-2-fluoro-βD-arabinofuranosyl)adenine (S.5). The protocol begins with 6-chloropurine riboside (S.1; Fig. 1.12.1), which is commercially available and can also be prepared from inosine (Zemlicka and Sorm, 1965). 3 -O-Benzoylation. The treatment of ribonucleosides with acyl chloride usually produces a mixture of monoacyl, diacyl, or triacyl derivatives. However, the 2 ,3 -O-din-butylstannylene complex is a useful intermediate for the introduction of a single benzoyl group onto the 2 - and/or 3 -OH (Wagner et al., 1974; Maruyama et al., 1998). Thus, 6-chloropurine riboside (S.1; Zemlicka and Owens, 1978) can be successively treated with di-n-butyltin oxide and excess benzoyl chloride in the presence of triethylamine in
Synthesis of Modified Nucleosides
1.12.23 Current Protocols in Nucleic Acid Chemistry
Supplement 25
methanol. Workup and purification by silica gel chromatography give an inseparable mixture of the 2 -O- and 3 -O-benzoates. Crystallization from methanol produces the 3 -Obenzoate (S.2) as pale yellowish crystals in 65% yield. The purity at this point can be checked by HPLC, which reveals that the product is the 3 -O-benzoate (96.1%) mixed with the 2 -O-benzoate (3.9%). 5 -O-Tritylation. Trityl (triphenylmethyl) chloride has been employed to achieve selective protection of primary alcohols. A base such as pyridine promotes this reaction. However, the reaction of S.2 in pyridine causes nucleophilic displacement at the 6-chloro moiety and the solution becomes colored. To overcome this undesired reaction, triethylamine is used in an amount less than that of trityl chloride. Under such reaction conditions, however, the 3 -O-benzoyl group migrates to the 2 -OH position, rapidly reaching equilibrium. TLC of the mixture shows spots of both 5 -Otrityl-2 -O-benzoylriboside and 5 -O-trityl-3 O-benzoylriboside (S.3). Therefore, the mixture is crystallized from methanol to afford S.3 in 71% yield. Analysis by HPLC reveals S.3 to be of 98% purity (Fig. 1.12.5). Acyl migration. The 2 -OH of a ribonucleoside is preferentially acetylated over the 3 -OH group. However, because 2 -acyl derivatives are unstable with respect to their 3 -isomers, subsequent equilibrium results in acetyl migration, with the 3 -O-acylate being obtainable by crystallization from the equilibrium mixture (Brown et al., 1956; Reese and Trentham, 1965; Johnston, 1968; Reese et al., 1975). The acidity of the 2 -OH group explains this tendency (Brown and Robins, 1965; Gin and Dekker, 1968). In this manner, the 3 O-benzoates (S.2, S.3) can be isolated from
Synthesis of Fluorinated Nucleosides
mixtures of the 3 - and 2 -isomers. Acyl migration can be controlled during fluorination. Migration was explored using the 3 -O-benzoate S.2 in methanol under acidic and basic conditions by means of HPLC analysis. S.2 reached equilibrium rapidly in a 1 mM triethylamine solution. However, the rate of migration was slow in a 1 mM pyridine solution and was similar to that in methanol alone. In a 1 mM imidazole solution, an intermediate value was obtained. Under acidic conditions, no migration was observed even after 1 day (Fig. 1.12.6). The acyl migration of S.3 was also examined since the fluorination reaction of S.3 with diethylaminosulfur trifluoride (DAST) should be carried out in the presence of an amine to protect the acid-labile 5 -O-trityl group. S.3 was heated under reflux in the presence of 6 eq of pyridine in CH2 Cl2 for 5 hr. Only 3.1% of the 2 -O-benzoate was observed in the mixture. This result indicates that the 3 -O-benzoyl group does not migrate to the 2 -hydroxl group in CH2 Cl2 containing a small amount of pyridine at reflux. Fluorination with DAST. Fluorination using DAST liberates acidic HF and cleaves the 5 -O-trityl group. To prevent deprotection, the addition of a base is effective. However, a strong base such as triethylamine accelerates the migration of the 3 -O-benzoate. Pyridine in CH2 Cl2 prevents deprotection without acyl migration as mentioned above. This enables the fluorination of S.3 using DAST in the presence of pyridine. S.4 is obtained in 78% yield. Fluorination with Et3 N·3HF via imidazolesulfonate. DAST is corrosive and should be avoided in large-scale industrial production if possible. Chou and colleagues reported a large-scale synthesis of C2-(arabino)-fluorinated sugar derivatives
Figure 1.12.5 Chromatogram of 5 -O-trityl-3 -O-benzoyl-6-chloropurine riboside (S.3) by HPLC. ◦ Operating conditions: detector λ, 265 nm; column, 5C18-MS (4.6 A, 150 mm); eluent, 1:4 (v/v) 10 mM H3 PO4 /MeOH.
1.12.24 Supplement 25
Current Protocols in Nucleic Acid Chemistry
Figure 1.12.6 Ratio of the 2 -O-benzoate mixed with 3 -O-benzoate S.2 as a function of elapsed time in MeOH alone (×), and 1 mM solution of Et3 N (circle), imidazole (up triangle), pyridine (down triangle), and HCl (square). Table 1.12.4 Fluorination of the Imidazolesulfonate Derived from S.3
Temperature (◦ C) Time (hr) Yield (%)a
Run
Solvent
Et3 N·3HF (eq.)
Et3 N (eq.)
1
EtOAc
6.0
—
70
24
67
2
EtOAc
6.0
3.0
70
24
54
3
Toluene
6.0
—
70
24
78
4
CH3 CN
6.0
—
70
24
62
5
CH2 Cl2
6.0
—
70
24
60
a Yields include imidazolesulfonation of S.4a.
using non-corrosive triethylamine trihydrofluoride (Et3 N·3HF; McClinton, 1995; Chou et al., 1996; Haufe, 1996). The only previously reported use of Et3 N·3HF in the synthesis of nucleoside derivatives was for 3 -β-fluorinated thymidine (von Janta-Lipinski et al., 1983). For fluorination of the sugar moiety of S.3, the 2 -OH should first be converted to an imidazolesulfonate, and then reacted with Et3 N·3HF in several solvents at 70◦ C (Table 1.12.4). HPLC analysis shows that the yield of S.4 varies from 54% to 78%, including imidazolesulfonation. The best result is obtained using toluene as the solvent. Fluorination with Et3 N·3HF via triflate. Fluorination with Et3 N·3HF can also be achieved via a trifluoromethanesulfonate, which is obtained quantitatively from S.3. S.3 is similarly reacted with 6 eq Et3 N·3HF and 3 eq triethylamine in ethyl acetate at reflux temperature overnight. HPLC analysis indicates that the yield of S.4 is 88%. However,
the activating agents imidazole and sulfuryl chloride are much less expensive than trifluoromethansulfonyl chloride. Therefore, from an industrial point of view, the imidazolesulfonate route may be more suitable. Treatment of S.4 with ammonia. Nucleophilic displacement of the 6-chloro function and removal of the 3 -O-benzoyl group is achieved by treatment of S.4 with ammonia in methanol to produce S.5 as prism-like crystals. Since S.5 exhibits a nuclear Overhauser effect (NOE) between H1 and H2 in the two-dimensional NOE (NOESY) spectrum, the configuration of S.5 can be identified as a 2 -(S)-arabinoside structure. Part of the product is converted to the known product FdaraA (S.6). The measured data for S.6 are identical with the published values (Pankiewicz et al., 1992a). Radical deoxygenation. The 3 -OH of S.5 is esterified using phenyl chlorothionoformate to give S.7 in 80% yield. S.7 is then hydrogenated
Synthesis of Modified Nucleosides
1.12.25 Current Protocols in Nucleic Acid Chemistry
Supplement 25
with tris(trimethylsilyl)silane in the presence of 2,2 -azobis(isobutyronitrile) (AIBN) in toluene under a nitrogen atmosphere to give S.9 as white crystals in 73% yield. The 3 deoxygenation can also be accomplished by preparation of the xanthate S.8 (87%) and radical reduction by hypophosphorous acid (93%). Preparation of FddA. Treatment of S.9 with concentrated HCl gives FddA (S.10) as prismlike crystals in good yield (Table 1.12.4). This sample was identical in all respects to the published data (Marquez et al., 1987), and the NOESY spectrum supported the structure. Thus, the first synthesis of FddA is accomplished from 6-chloropurine riboside, which is obtainable from inosine.
Synthesis of Fluorinated Nucleosides
Fluorination of a 6-chloropurine 3 -deoxyriboside The disadvantage to the above approach is that it requires deoxygenation of the 3 -OH group after the crucial fluorination step, which results in a loss of the precious fluorinated compound. To develop a more direct approach to the synthesis of FddA that can be performed on a large scale, direct fluorination at the C2 -β position of a 3 -deoxynucleoside was chosen as a potentially more straightforward approach. Marquez et al. (1987) studied the fluorination of protected 3 -deoxyadenosine with tetrabutylammonium fluoride, but a fluorinated compound was not obtained. Herdewijn et al. (1987) and Shiragami et al. (1992) reported the fluorination of 5 -Oprotected 3 -deoxyadenosine with DAST, but the yields of the fluorinated compounds were only 10% in both cases. In contrast, fluorination of 5 -O-protected 6-chloropurine 3 deoxyriboside proceeds in good yield, and the substrate is easily prepared from the readily available inosine (Takamatsu et al., 2001a,b; Izawa et al., 2003). This approach is presented in Basic Protocols 3 and 4. Acetoxybromination and radical debromination. Synthesis of the 6-chloropurine 3 deoxyriboside (S.15; Fig. 1.12.3) from inosine is achieved by acetoxybromination chemistry already developed for large-scale synthesis of ddA (Shiragami et al., 1996), followed by radical debromination using hypophosphorous acid as a reducing agent in the presence of a water-soluble radical initiator such as VA-044 or V-50 (Wako Pure Chemical Industries). Although the acetoxybromination yield of inosine is less than that of adenosine due to its acid lability, S.12 can nevertheless be obtained in moderate yield. Subsequent debromination is carried out by radical re-
duction (Takamatsu et al., 2001c). Though conventional n-Bu3 SnH/AIBN conditions are applicable to this reaction, the removal of the residual Sn compound is generally troublesome, and it is difficult to remove the AIBN that co-precipitates with the product. Barton et al. (1992, 1993) reported that the commercially available and inexpensive hypophosphorous acid could be used for radical reduction instead of n-Bu3 SnH. Although Barton et al. used these reagents for rather simple substrates, the application of this reaction in the deoxygenation or dehalogenation of sugar moieties of nucleosides proved successful. Furthermore, water-soluble radical initiators can be used in the aqueous reaction medium and, since the product S.13 is crystallized from aqueous medium, it proved easy to remove both the reducing agent and the radical initiator. Chlorination. Chlorination of the 6position of the purine base is accomplished under Vilsmeier conditions. Though acidic cleavage of the glycoside bond is a major side reaction, it can be decreased to a negligible level by monitoring the reaction carefully and by cooling the reaction mixture immediately at the end of the reaction. Finally, 2 ,5 -Oacetyl groups can be removed by treatment with sodium methoxide in methanol to give S.15. 5 -O-Tritylation. Because tritylation of primary alcohols is much faster than that of secondary alcohols, the reaction selectively proceeds at the 5 -OH group. Pyridine-type bases such as 2,4-6-collidine or 2-picoline are preferable for this reaction, as the reaction is a type of SN 1 conversion (trityl chloride reacts as a trityl cation, which is stabilized by an aromatic base). The crystallization of the 5 -Otrityl product S.16 is rather tricky, as it can be crystallized only from benzotrifluoride. Fluorination. The fluorination is carried out using a combination of perfluorobutanesulfonyl fluoride (nonaflate) and N, N-dimethylcyclohexylamine in toluene to give S.17 (Fig. 1.12.4; Takamatsu et al., 2002). Although treatment of S.16 with DAST gives S.17 in a moderate yield (Takamatsu et al., 2001a,b), DAST is not suitable for large-scale synthesis because of its high toxicity and corrosiveness. Moreover, it is not commercially available in large quantities due to its explosive nature. Fluorination of the triflate of S.16 can be performed using triethylamine trihydrogenfluoride (Et3 N·3HF; Izawa et al., 2003); however, this requires activation of the hydroxyl group as a triflate with the expensive
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Current Protocols in Nucleic Acid Chemistry
reagent triflic anhydride. Among the reagents that can achieve dehydroxy-fluorination in a single step, perfluoroalkanesulfonyl fluoride (Beyl et al., 1970) appears to be the reagent of choice, since it is only mildly corrosive and is readily available from commercial sources. This reagent is stable at room temperature and is less moisture-sensitive, thus rendering it suitable for use in large-scale syntheses. Fluorination using perfluoroalkanesulfonyl fluoride had previously been applied only to simple compounds (BennuaSkalmowski, 1994; Bennua-Skalmowski and Vorbruggen, 1995). For fluorination of the C2 -β position of a 3 -deoxy compound such as S.16, an eliminated byproduct S.18 is obtained together with the fluorinated compound S.17. The choice of base and solvent has a big impact on the reactivity and the S.17/S.18 selectivity. The combination of N,N-dimethylcyclohexylamine as the base and toluene as the solvent was selected after many screening tests, which indicated that an apolar condition is required to reduce the eliminated byproduct. Using optimized conditions to isolate the fluorinated compound, S.17 can be isolated by selective decomposition of S.18 in an acetic acid solution, because S.18 is very unstable under acidic conditions while S.17 is stable. Amination. The subsequent amination of the 6-position of the purine base is a very clean reaction, although somewhat slow. An autoclave reactor is required due to the pressure of the NH3 gas (8 bar at the maximum). The choice of solvent may be important; THF has, thus far, given the best result. Detritylation. The final deprotection proceeds under normal acidic conditions, such as HCl in methanol. Methanol causes losses of FddA in the organic layer, so it must be distilled completely before extraction. Hydrophobic impurities are removed in the organic layer, while hydrophilic material such as FddA, adenine, and residual S.17 (where the reaction has not run to completion) stays in the aqueous layer. Neutralization of the aqueous layer with NH4 OH causes precipitation of FddA. The purity of FddA obtained is normally >99%.
Compound Characterization Via 6-chloropurine riboside. Melting points were determined on a Yanagimoto micromelting point apparatus (hot-stage type). UV spectra were obtained on a Shimadzu UV190 digital spectrometer. Low-resolution mass spectra were obtained on a Shimadzu-LKB
9000B mass spectrometer in the direct-inlet mode, while high-resolution spectra were obtained on a JMS AX-500 spectrometer in the direct-inlet mode. 1 H NMR spectra were obtained on a Varian Unity 200 (200 MHz) or Varian Unity 600 (600 MHz) in CDCl3 (or DMSO-d6 ) with tetramethylsilane as an internal standard. Via 6-chloropurine 3 -deoxyriboside. All intermediates and products were characterized by standard NMR, IR, and MS techniques as listed in the protocols. In addition, the use of HPLC is recommended to monitor the reactions and to check the purity of the compounds. Conditions (two isocratic conditions and two gradient conditions) and retention times are also listed in the protocols. Because this approach was developed as an industrial process with the HPLC control, TLC was not used to monitor reactions. Melting points were determined on a Yanagimoto micro-melting point apparatus (hot-stage type). UV spectra were obtained on a Shimadzu UV-1600 spectrometer. NMR spectra were obtained on a Varian XL-300 (300 MHz) in CDCl3 (or DMSO-d6 ) with tetramethylsilane as an internal standard. Infrared (IR) spectra were obtained on a Perkin Elmer PARAGON-1000 FT-IR spectrometer. High-resolution mass spectra were obtained on a JOEL JMS-700V spectrometer using FAB (fast atom bombardment) ionization.
Critical Parameters and Troubleshooting General laboratory safety is of primary concern because hazardous materials are involved in the course of the synthesis. Knowledge of the basic procedures of organic synthesis and basic analytical methods is also required. One of the most critical parameters of the synthesis is the purity of the reactants and reagents. Although all the reagents and solvents can be readily purchased and used without further purification, checking the purity of the reactants or intermediates by suitable methods such as TLC, HPLC, and NMR is highly recommended. In particular, it is better to analyze the isolatable intermediates by NMR, IR, MS, UV, and HPLC. If the purity is not high enough, further purification is recommended. S.2 and S.3 can be purified by crystallization from MeOH. S.5, S.13, S.14, and the products FddA (S.6) and FddA (S.10 = S.20) can be purified by recrystallization from aqueous MeOH. S.16 and S.17 can be recrystallized by repeating their respective isolation procedures.
Synthesis of Modified Nucleosides
1.12.27 Current Protocols in Nucleic Acid Chemistry
Supplement 25
Table 1.12.5 Estimated Operation Time for Synthesis of FddA from Inosine
Reaction 1 2 3 4 5
2 ,5 -OAc-3 -Br-3 -deoxyinosine (S.12)
2
2
2
2 ,5 -OAc-3 -deoxyinosine (S.13) 2 ,5 -OAc-3 -deoxy-6-chloropurine riboside (S.14)
3 -deoxy-6-chloropurine riboside (S.15)
5 -OTr-3 -deoxy-6-chloropurine riboside (S.16)
2 3
6
5 -OTr-2 -fluoro-2 ,3 -dideoxy-6-chloropurine riboside (S.17)
4
7
5 -OTr-2 -fluoro-ddA (S.19)
4
8
FddA (S.20)
2
From inosine to FddA
21
Total
Anticipated Results Via 6-chloropurine riboside. Adherence to the methods outlined in Basic Protocols 1 and 2 will yield the total synthesis of FdaraA and FddA on a large scale (e.g., 10 to 20 g scale with 20% to 30% overall yield of FddA from S.1). Many 6-substituted derivatives of FddA could also be obtainable from 6-chloro-9-(3-O-benzoyl-5-O-trityl-2deoxy-2-fluoro-β-D-arabinofuranosyl)purine (S.4). However, the second approach may be preferable for FddA preparation, as there is less loss of product after the fluorination reaction. Via 6-chloropurine 3 -deoxyriboside. Adherence to the methods outlined in Basic Protocols 3 and 4 will yield the total synthesis of FddA on a large scale. As this method is robust and reproducible, highly pure FddA may be obtained in a short period of time. The overall yield from 6-chloro-9-(3-deoxy-β-D-erythropentofuranosyl)-9H-purine (S.15) is ∼31%.
Time Considerations Following the procedures in Basic Protocol 1 and the Alternate Protocol, the synthesis of FdaraA can be accomplished in ∼2 weeks. The synthesis of lodenosine may be accomplished in ∼1 month using either approach. The time required to complete each step from inosine to FddA (see Basic Protocols 3 and 4) is summarized in Table 1.12.5. The estimated time does not include time for analysis.
Literature Cited Synthesis of Fluorinated Nucleosides
Estimated time (days)
Product
Barchi, J.J. Jr., Marquez, V.E., Driscoll, J.S., Ford, H. Jr., Mitsuya, H., Shirasaka, T., Aoki, S., and Kelley, J.A. 1991. Potential anti-AIDS drugs. Lipophilic, adenosine deaminase-activated prodrugs. J. Med. Chem. 34:1647-1655.
Barton, D.H.R., Jang, D.O., and Jaszberenyi, J.Cs. 1992. Hypophosphorous acid and its salts: New reagents for radical chain deoxygenation, dehalogenation and deamination. Tetrahedron Lett. 33:5709-5712. Barton, D.H.R., Jang, D.O., and Jaszberenyi, J.Cs. 1993. The invention of radical reactions. 32. Radical deoxygenations, dehalogenations, and deaminations with dialkyl phosphates and hypophosphorous acid as hydrogen sources. J. Org. Chem. 58:6838-6842. Bennua-Skalmowski, B. and Vorbruggen, H. 1995. A facile conversion of primary or secondary alcohols with n-perfluorobutanesulfonyl fluoride/1,8-diazabicyclo[5.4.0]undec-7-ene into their corresponding fluorides. Tetrahedron Lett. 36:2611-2614. Bennua-Skalmowski, B., Krolikiewicz, K., and Vorbruggen, H. 1994. The reaction of perfluorobutanesulfonyl fluoride with alcohols in the presence of 4-dialkylaminopyridines. Bull. Soc. Chim. Belg. 103:453-461. Beyl, V., Niederpruem, H., and Voss, P. 1970. New reaction of perfluoroalkylsulfonyl fluorides. Liebigs Ann. Chem. 731:58-66. Brown, A.D. and Robins, R.K. 1965. The direct preparation of 2 -O-methyladenosine from adenosine. J. Amer. Chem. Soc. 87:1145-1146. Brown, D.M., Todd, A., and Varadarajan, S. 1956. Nucleotides. XXXVII The structure of uridylic acids a and b, and a synthesis of spongouridine (3-β-D-arabofuranosyluracil). J. Chem. Soc. 2388-2393. Chou, T.S., Becke, L.M., O’Toole, J.C., Carr, M.A., and Parker, B.E. 1996. Triethylamine poly(hydrogen fluorides) in the synthesis of a fluorinated nucleoside glycon. Tetrahedron Lett. 37:17-20. Chu, C.K., Matulic-Adamic, J., Huang, J.T., Chou, T.C., Burchenal, J.H., Fox, J.J., and Watanabe, K.A. 1989. Nucleosides. CXXXV. Synthesis of some 9-(2-deoxy-2-fluoro-βD-arabinofuranosyl)-9H-purines and their
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biological activities. Chem. Pharm. Bull. 37:336-339. Driscoll, J.S., Mayers, D.L., Barder, J.P., Weislow, O.S., Johns, D.G., and Buckheit, R.W. Jr. 1997. 2-Fluoro-2 ,3-dideoxyarabinosyladenine (F-ddA): Activity against drug-resistant human immunodeficiency virus strains and clades A-E. Antivir. Chem. Chemother. 8:107-111. Gin, J.B. and Dekker, C.A. 1968. The preparation and properties of O-methylated adenosine derivatives. Biochemistry 7:1413-1420. Graul, A., Silvestre, J., and Castaner, J. 1998. Lodenosine, anti-HIV reverse transcriptase inhibitor. Drugs Future 23:1176-1189. Haufe, G. 1996. Triethylamine trishydrofluoride in synthesis. J. Prakt. Chem. 338:99-113. Herdewijn, P., Pauwels, R., Baba, M., Balzarini, J., and De Clercq, E. 1987. Synthesis and anti-HIV activity of various 2 - and 3 -substituted 2 ,3 dideoxyadenosines: A structure-activity analysis. J. Med. Chem. 30:2131-2137. Izawa, K., Takamatsu, S., Katayama, S., Hirose, N., Kozai, S., and Maruyama, T. 2003. An industrial process for synthesizing Lodenosine (FddA). Nucleosides Nucleotides Nucleic Acids 22:507517. Johnston, G.A.R. 1968. Acetylation of nucleosides and acetyl migration. Tetrahedron 24:69876993. Kawasaki, A.M., Casper, M.D., Freier, S.M., Lesnik, E.A., Zounes, M.C., Cummins, L.L., Gonzalez, C., and Cook, P.D. 1993. Uniformly modified 2 -deoxy-2 -fluoro-phosphorothioate oligonucleotides as nuclease-resistant antisense compounds with high affinity and specificity for RNA targets. J. Med. Chem. 36:831-841. Marquez, V.E., Tseng, C.K.-H., Kelley, J.A., Mitsuya, H., Broder, S., Roth, J.S., and Driscoll, J.S. 1987. 2 ,3 -Dideoxy-2 -fluoro-ara-A, an acid stable purine nucleoside active against human immunodeficiency virus (HIV). Biochem. Pharmacol. 36:2719-2722. Marquez, V.E., Tseng, C.K.-H., Mitsuya, H., Aoki, S., Kelley, J.A., Ford, H. Jr., Roth, J.S., Border, S., Johns, D.G., and Driscoll, J.S. 1990. Acid stable 2 -fluoro purine dideoxynucleoside as active agents against HIV. J. Med. Chem. 33:978985. Maruyama, T., Sato, Y., Oto, Y., Takahashi, Y., Snoeck, R., Andrei, G., Witvrouw, M., and De Clercq, E. 1996. Synthesis and antiviral activity of 6-chloropurine arabinoside and its 2 deoxy-2 -fluoro derivative. Chem. Pharm. Bull. 44:2331-2334. Maruyama, T., Sato, Y., and Sakamoto, T. 1998. Synthesis of 8-substituted analogs of 3 -deoxy3 -fluoroadenosine. Nucleosides Nucleotides 17:115-122. Maruyama, T., Takamatsu, S., Kozai, S., Satoh, Y., and Izawa, K. 1999. Synthesis of 9-(2-deoxy-2fluoro-β-D-arabinofuranosyl)adenine bearing a selectively removable protecting group. Chem. Pharm. Bull. 47:966-970.
McClinton, M.A. 1995. Triethylamine tris(hydrogen fluoride): Applications in synthesis. Aldrichimica Acta 28:31-35. Pankiewicz, K.W., Krzeminski, J., Ciszewski, L.A., Ren, W.-Y., and Watanabe, K.A. 1992a. A synthesis of 9-(2-deoxy-2-fluoro-βD-arabinofuranosyl)adenine and -hypoxanthine. An effect of C3 -endo to C2 -endo conformational shift on the reaction course of 2 -hydroxyl group with DAST. J. Org. Chem. 57:553-559. Pankiewicz, K.W., Krzeminski, J., and Watanabe, K.A. 1992b. Synthesis of 2 -β-Fluoro- and 3 -α-Fluoro-substituted guanine nucleosides. Effects of sugar conformational shifts on nucleophilic displacement of 2 -hydroxy and 3 hydroxy group with DAST. J. Org. Chem. 57:7315-7321. Parker, W.B., Allan, P.W., Hassan, A.E.A., Secrist, J.A. 3rd, Sorscher, E.J., and Waud, W.R. 2003. Antitumor activity of 2-fluoro-2 deoxyadenosine against tumors that express Escherichia coli purine nucleoside phosphorylase. Cancer Gene Ther. 10:23-29. Reese, C.B. and Trentham, D.R. 1965. 2 -O-Acyl ribonucleoside derivatives. Tetrahedron Lett. 2459-2465. Reese, C.B., Stewart, J.C.M., van Boom, J.H., de Leeuw, H.P.M., Nagel, J., and de Rooy, J.F.M. 1975. Synthesis of oligoribonucleotides. XI. Preparation of ribonucleoside 2 -acetal 3 -esters by selective deacylation. J. Chem. Soc., Perkin Trans. 1 934-942. Ruxrungtham, K., Bonne, E., Ford, H. Jr., Driscoll, J.S., Davey, R.T. Jr., and Lane, H.C. 1996. Potent activity of 2 -β-fluoro-2 ,3 -dideoxyadenosine against human immunodeficiency virus type 1 infection in hu-PBL-SCID mice. Antimicrob. Agents Chemother. 40:2369-2374. Sato, A., Montgomery, J.A., and Cory, J.G. 1984. Synergistic inhibition of leukemia L1210 cell growth in vitro by combinations of 2fluoroadenine nucleosides and hydroxyurea or 2,3-dihdro-1H-pyrazole[2,3-a]imidazole. Cancer Res. 44:3286-3290. Shiragami, H., Tanaka, Y., Uchida, Y., Iwagami, H., Izawa, K., and Yukawa, T. 1992. A novel method for the synthesis of ddA and F-ddA via regioselective 2 -O-deacetylation of 9-(2,5-di-O-acetyl-3-bromo-3-deoxy-β-Dxylofuranosyl)adenine. Nucleosides Nucleotides 11:391-400. Shiragami, H., Amino, Y., Honda, Y., Arai, M., Tanaka, Y., Iwagami, H., Yukawa, T., and Izawa, K. 1996. Synthesis of 2 ,3 dideoxypurinenucleosides via the palladium catalyzed reduction of 9-(2,5-di-O-acetyl-3bromo-3-deoxy-β- D -xylofuranosyl)purine derivatives. Nucleosides Nucleotides 15:31-45. Siddiqui, M.A., Marquez, V.E., Driscoll, J.S., and Barchi, J.J. Jr. 1994. A diastereoselective synthesis of (S,S)-α-fluoro-2,2-dimethyl1,3-dioxolane-4-propanoic acid methyl ester, a key intermediate for the preparation of antiHIV effective fluorodideoxynucleosides. Tetrahedron Lett. 35:3263-3266.
Synthesis of Modified Nucleosides
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Siddiqui, M.A., Driscoll, J.S., and Marquez, V.E. 1998. A new synthetic approach to the clinically useful, anti-HIV-active nucleoside, 9-(2,3-dideoxy-2-fluoro-β-D-threopentofuranosyl)adenine (β-FddA). Introduction of a 2 -β-fluoro substituent via inversion of a readily obtainable 2 -α-fluoro isomer. Tetrahedron Lett. 39:1657-1660.
Wagner, D., Verheyden, J.P.H., and Moffat, J.G. 1974. Preparation and synthetic utility of some organotin derivatives of nucleosides. J. Org. Chem. 39:24-30. Wysocki, R.J. Jr., Siddiqui, M.A., Barchi, J.J. Jr., Driscoll, J.S., and Marquez, V.E. 1991. A more expedient approach to the synthesis of anti-HIV-active 2,3-dideoxy-2-fluoro-βD-threo-pentofuranosyl nucleosides. Synthesis 11:1005-1008.
Takamatsu, S., Maruyama, T., Katayama, S., Hirose, N., Naito, M., and Izawa, K. 2001a. Synthesis of 9-(2,3-dideoxy-2-fluoro-β-Dthreo-pentofuranosyl)adenine (FddA) via a purine 3 -deoxynucleoside. J. Org. Chem. 66:7469-7477.
Zemlicka, J. and Owens, J. 1978. 6-Chloro-9-β-Dribofuranosylpurine. A versatile intermediate in the synthesis of purine ribonucleosides. Nucleic Acid Chem. 2:611-614.
Takamatsu, S., Maruyama, T., Katayama, S., Hirose, N., Naito, M., and Izawa, K. 2001b. Practical synthesis of 9-(2,3-dideoxy-2-fluoroβ-D-threo-pentofuranosyl)adenine (FddA) via a purine 3 -deoxynucleoside. Tetrahedron Lett. 42:2325-2328.
Zemlicka, J. and Sorm, F. 1965. Nucleic acids components and their analogs. LX. The reaction of chloromethylenedimethylammonium chloride with 2 ,3 ,5 -tri-O-acetylinosine. A new synthesis of 6-chloro-9-(β-D-ribofuranosyl)purine. Collect. Czech. Chem. Commun. 30:1880-1889.
Takamatsu, S., Katayama, S., Hirose, N., Naito, M., and Izawa, K. 2001c. Radical deoxygenation and dehalogenation of nucleoside derivatives with hypophosphorous acid and dialkyl phosphates. Tetrahedron Lett. 42:7605-7608. Takamatsu, S., Katayama, S., Hirose, N., De Cook, E., Schelkens, G., Demillequand, M., Brepoels, J., and Izawa, K. 2002. Convenient synthesis of fluorinated nucleosides with perfluoroalkanesulfonyl fluoride. Nucleosides Nucleotides Nucleic Acids 21:849-861. Ueno, T. and Mitsuya, H. 1997. Comparative enzymatic study of HIV-1 reverse transcriptase resistant to 2 ,3 -dideoxynucleotide analogs using the single-nucleotide incorporation assay. Biochemistry 36:1092-1099.
Contributed by Satoshi Katayama, Satoshi Takamatsu, Naoko Hirose, and Kunisuke Izawa AminoScience Laboratories Ajinomoto Co. Kanagawa, Japan Tokumi Maruyama Tokushima Bunri University Kagawa, Japan
von Janta-Lipinski, M., Langen, P., and Czech, D. 1983. Cleavage of 2,3 -anhydronucleosides with hydrohalic acids. Z. Chem. 23:335.
Synthesis of Fluorinated Nucleosides
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Synthesis of Ribonucleosides by Condensation Using Trimethylsilyl Triflate
UNIT 1.13
This unit provides protocols for the synthesis of each of the four common ribonucleosides adenosine (S.16, Fig. 1.13.2), cytidine (S.14), guanosine (S.18), and uridine (S.12) from the heterocycles uracil (S.1, Fig. 1.13.1), cytosine (S.2), N6 -benzoyladenine (S.3), and N2 -acetylguanosine (S.4). A procedure for silylation of each of the heterocycles is provided (see Support Protocols 1 and 2) and glycosylation mediated by trimethylsilyl triflate (S.10) of the silylated heterocycles by 1-O-acetyl-2,3,5-tri-O-benzoylribofuranose (S.9) is described (see Basic Protocols 1 through 4). The protocols may be applied to many modified pyrimidines and purines as well as other related heterocycles to construct modified nucleosides (Vorbr¨uggen and Ruh-Pohlenz, 2000, 2001). The chemical reactions described here produce good yields, and purification of the intermediates is straightforward. Mechanistic details of the glycosylation reaction are outlined in the Commentary.
Figure 1.13.1 Heterocyclic bases (S.1-4) and silylated intermediates (S.5-8) obtained by treatment of bases with hexamethyldisilazane (HMDS; see Support Protocol 1) or N,Obis(trimethylsilyl)acetamide (BSA; see Support Protocol 2).
¨ Contributed by Helmut Vorbruggen, Irene M. Lagoja, and Piet Herdewijn Current Protocols in Nucleic Acid Chemistry (2006) 1.13.1-1.13.16 C 2006 by John Wiley & Sons, Inc. Copyright
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Figure 1.13.2 Glycosylation of silylated bases using TMSOTf (S.10) to yield (after ammonia deprotection) the natural ribonucleosides uridine (S.12), cytidine (S.14), adenosine (S.16), and guanosine (S.18).
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NOTE: All reagents and solvents must be anhydrous. HMDS and other silyl derivatives can be obtained from Fa. Bucher. All starting materials are commercially available (e.g., Acros, Aldrich, Fluka, Lancaster).
PREPARATION OF URIDINE Uracil (S.1) is first converted to 2,4-bis(trimethylsilyloxy)pyrimidine (S.5; Fig. 1.13.1) following the procedure outlined in Support Protocol 1. S.5 is then allowed to react with 1-O-acetyl-2,3,5-tri-O-benzoylribofuranose (S.9) and trimethylsilyl triflate (S.10) in dry 1,2-dichloroethane (Fig. 1.13.2). The resulting uridine 2 ,3 ,5 -tri-O-benzoate (S.11) is purified by chromatography on silica gel, and the benzoyl protecting groups removed by treatment with methanolic ammonia to yield uridine (S.12).
BASIC PROTOCOL 1
Materials 2,4-Bis(trimethylsilyloxy)pyrimidine (S.5; see Support Protocol 1) Anhydrous 1,2-dichloroethane (reflux overnight on P2 O5 and distill before use) 1-O-Acetyl-2,3,5-tri-O-benzoylribofuranose (S.9) Trimethylsilyltrifluoromethanesulfonate (TMSOTf; S.10) Dichloromethane (CH2 Cl2 ) Methanol (MeOH) Saturated sodium bicarbonate (sat. NaHCO3 ) solution, ice cold Sodium sulfate (Na2 SO4 ) Silica gel (0.060 to 0.200 nm) Benzene (optional) Saturated ammonia in methanol (sat. NH3 /MeOH) Ethanol (EtOH) 100- and 250-mL round-bottom flask Balloon filled with nitrogen or argon Reflux condenser Precoated TLC plates (e.g., Alugram Sil G/UV254 , Macherey-Nagel) 254-nm UV lamp (for TLC) 500-mL separatory funnel 500-mL Erlenmeyer flask Buchner funnel Filter paper 250-mL filter flask Rotary evaporator equipped with a vacuum pump 5 × 35–cm chromatography column Desiccator Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare uridine 2 ,3 ,5 -tri-O-benzoate (S.11) 1. Dissolve 1.32 g (5.15 mmol) of 2,4-bis(trimethylsilyloxy)pyrimidine (S.5) in 18 mL of 1,2-dichloroethane in a 100-mL round-bottom flask. 2. Add 2.57 g (5.0 mmol) 1-O-acetyl-2,3,5-tri-O-benzoylribofuranose (S.9). 3. Add 0.55 g (2.5 mmol) TMSOTf (S.10) and heat the mixture under nitrogen protection under reflux (90◦ -95◦ C) for 4 hr. CAUTION: TMSOTf is toxic and reacts violently with water, and should be protected from moisture.
Synthesis of Modified Nucleosides
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4. Monitor by TLC (APPENDIX 3D) using 95:5 (v/v) CH2 Cl2 /MeOH (Rf = 0.64). When the starting material is completely depleted, cool the mixture to room temperature and dilute the clear yellow solution with 50 mL CH2 Cl2 . 5. Extract with 50 mL ice-cold sat. NaHCO3 solution in a 500-mL separatory funnel. 6. Wash the organic layer three times with 20 mL water. 7. Collect the organic layers in a 500-mL Erlenmeyer flask and dry over Na2 SO4 . Filter off the drying agent using filter paper and 250-mL round-bottom flask, and evaporate to dryness (2.80 g colorless foam) on a rotary evaporator. 8. To remove traces of impurities, purify by column chromatography (APPENDIX 3E) using a 5 × 35–cm column with 250 g silica gel, eluting with 1.5 L of 9:1 (v/v) CH2 Cl2 /MeOH, or crystallize 2.80 g crude product from 40 mL benzene for 2 hr at 24◦ C. Yield of uridine 2 ,3 ,5 -tri-O-benzoate (S.11) following chromatography: 2.63 g (95%). Yield following crystallization: 2.25 g (81%). m.p. 138◦ -140◦ C. TLC (95:5 CH2 Cl2 /MeOH): Rf 0.64. FAB+ : C30 H25 N2 O9 [M+H] 557.1. 1 H NMR (300 MHz, CDCl3 ): δ 4.66-4.73 (2H, 2 × H5 ), 4.84 (1H, H4 ), 5.91 (1H, H3 ), 6.17 (1H, H2 ), 6.34 (1H, d, J = 5.6 Hz, 5-H), 6.60 (1H, d, J = 3.3 Hz, H1 ), 7.27-8.13 (16H, 3 × Bz, 6-H), 8.78 (1H, s, br, ex, NH). 13 C NMR (75 MHz, CDCl3 ): δ = 63.7 (C5 ), 71.1 (C2 ), 73.7 (C3 ), 80.5 (C4 ), 88.1 (C1 ), 103.4 (5-C), 128.3-133.7 (3 × C Bz), 139.6 (6-C), 149.9 (2-C), 162.5 (4-C), 165.3, 165.4, 166.0 (3 × C=O Bz).
Prepare uridine (S.12) 9. Dissolve 2.00 g (3.59 mmol) S.11 in 50 mL sat. NH3 /MeOH. Keep the reaction mixture overnight at room temperature. 10. Evaporate the volatiles on a rotary evaporator to obtain a colorless crystalline material. 11. Stir material in 10 mL of 1:1 (v/v) CH2 Cl2 /MeOH for 2 hr at room temperature. 12. Filter off the crystals using a Buchner funnel and 250-mL filter flask, and wash with 20 mL CH2 Cl2 . 13. Crystallize from a minimum amount of boiling ethanol and then dry the product in a desiccator. Yield of uridine (S.12): 0.82 g (89%). m.p. 165◦ -166◦ C. The spectroscopic properties (NMR, MS, UV, m.p.) of the obtained nucleosides are identical with those previously described. TLC (2:1 CH2 Cl2 /MeOH): Rf 0.67. FAB+ : C9 H13 N2 O6 [M+H]: 245.1. 1 H NMR (300 MHz, DMSO-d6 ): δ = 3.54-3.61 (2H, 2 × H5 ), 3.84 (1H, H4 ), 3.96 (1H, H3 ), 4.01 (1H, m, H2 ), 5.06, 5.08, 5.37 (3H, (d), (t), (d), J ∼ 5.6 Hz, OH), 5.77 (1H, d, J = 8.1 Hz, 5-H), 5.78 (1H, d, J = 5.3 Hz, H1 ), 7.89 (1H, d, J = 8.1 Hz, 6-H), 11.29 (1H, s, br, ex, NH). 13 C NMR (75 MHz, DMSO-d6 ): δ = 61.3 (C5 ), 70.3 (C2 ), 74.0 (C3 ), 85.3 (C4 ), 88.1 (C1 ), 102.2 (5-C), 141.2 (6-C), 151.2 (2-C), 163.6 (4-C). UV (H2 O) λmax 261 nm. BASIC PROTOCOL 2
Synthesis of Ribonucleosides by Condensation Using Trimethylsilyl Triflate
PREPARATION OF CYTIDINE Cytosine is first converted to 4-(trimethylsilylamino)-2-(trimethylsilyloxy)pyrimidine (S.6) following the procedure outlined in Support Protocol 1. S.6 is then allowed to react with 1-O-acetyl-2,3,5-tri-O-benzoylribofuranose (S.9) and trimethylsilyl triflate (S.10) in dry 1,2-dichloroethane. The resulting cytidine 2 ,3 ,5 -tri-O-benzoate (S.13) is treated with methanolic ammonia to remove the protecting groups, and the resulting cytidine (S.14) is purified by recrystallization from ethanol.
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Materials 4-(Trimethylsilylamino)-2-(trimethylsilyloxy)pyrimidine (S.6; Support Protocol 1) Anhydrous 1,2-dichloroethane (reflux overnight over P2 O5 and distill before use) 1-O-Acetyl-2,3,5-tri-O-benzoylribofuranose (S.9) Trimethylsilyltrifluoromethanesulfonate (TMSOTf, S.10) Dichloromethane (CH2 Cl2 ) Methanol (MeOH) Saturated sodium bicarbonate (sat. NaHCO3 ) solution, ice cold Sodium sulfate (Na2 SO4 ) Silica gel (0.060 to 0.200 nm) Ethanol (EtOH), hot Charcoal Celite Saturated ammonia in methanol (sat. NH3 /MeOH) 250-mL round-bottom flask Balloon filled with nitrogen or argon Reflux condenser Precoated TLC plates (e.g., Alugram Sil G/UV254 , Macherey-Nagel) 254-nm UV lamp (for TLC) 500-mL separatory funnel 500-mL Erlenmeyer flask Filter paper Buchner funnel 250-mL filter flask Rotary evaporator equipped with a vacuum pump Desiccator Additional reagents and equipment for TLC (APPENDIX 3D) Prepare cytidine 2 ,3 ,5 -tri-O-benzoate (S.13) 1. Dissolve 2.56 g (5.15 mmol) 4-(trimethylsilylamino)-2-(trimethylsilyloxy)pyrimidine (S.6) in 35 mL of 1,2-dichloroethane in a 250-mL round-bottom flask. 2. Add 5.04 g (10.0 mmol) 1-O-acetyl-2,3,5-tri-O-benzoylribofuranose (S.9). 3. Add 2.67 g (12 mmol) TMSOTf (S.10) and heat the mixture under nitrogen protection under reflux (90◦ -95◦ C) for 1 hr. CAUTION: TMSOTf is toxic and reacts violently with water, and should be protected from moisture.
4. Monitor by TLC (APPENDIX 3D) using 9:1 (v/v) CH2 Cl2 /MeOH (Rf = 0.57). When all starting material is depleted, cool the mixture to room temperature and dilute the clear yellow solution with 100 mL CH2 Cl2 . 5. Extract with 50 mL ice-cold sat. NaHCO3 solution using a 500-mL separatory funnel. 6. Wash the organic layer three times with 20 mL water. 7. Collect the organic layers in a 500-mL Erlenmeyer flask and dry over Na2 SO4 . Filter off the drying agent using filter paper and 250-mL round-bottom flask, and evaporate to dryness (brownish foam) on a rotary evaporator. 8. Dissolve the obtained foam in 150 mL boiling ethanol. Treat with 0.2 g charcoal and continue refluxing for 15 min. Synthesis of Modified Nucleosides
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9. Filter hot over a Buchner filter equipped with a bed (4-mm layer) of celite and evaporate the volatiles on a rotary evaporator. Yield of cytidine 2 ,3 ,5 -tri-O-benzoate (S.13): 4.50 g (98%). TLC (9:1 CH2 Cl2 /MeOH): Rf 0.57. FAB+ : C30 H26 N3 O8 [M+H]: 556.1. 1 H NMR (300 MHz, CDCl3 ): δ 4.51-4.56 (2H, 2 × H5 ), 4.83 (1H, H4 ), 5.84 (2H, H3 , H2 ), 5.82 (2H, s, br ex, NH2 ), 5.99 (1H, d, J = 4.2 Hz, 5-H), 6.44 (1H, d, J = 1.3 Hz, H1 ), 7.33-8.10 (16H, 3 × Bz, 6-H). 13 C NMR (75 MHz, CDCl3 ): δ 63.8 (C5 ), 71.1 (C2 ), 74.5 (C3 ), 79.7 (C4 ), 90.5 (C1 ), 95.9 (5-C), 128.4-133.6 (3 × C Bz), 141.5 (6-C), 155.5 (2-C), 165.3 (4-C), 165.4, 166.0, 166.2 (3 × C = O Bz).
Prepare cytidine (S.14) 10. Dissolve 4.00 g (7.2 mmol) S.13 in 100 mL sat. NH3 /MeOH. Keep the reaction mixture overnight at room temperature. 11. Evaporate the volatiles on a rotary evaporator to obtain a colorless crystalline material. 12. Stir material in 15 mL of 1:1 (v/v) CH2 Cl2 /MeOH for 2 hr at room temperature. 13. Filter off the crystals using a Buchner funnel and 250-mL filter flask, and wash with 20 mL CH2 Cl2 . 14. Crystallize from a minimum amount of boiling EtOH and then dry the product in a desiccator. Yield of cytidine (S.14): 1.62 g (91%). m.p. 210◦ -212◦ C. The spectroscopic properties (NMR, MS, UV, m.p.) of the obtained nucleosides are identical with those previously described. TLC (2:1 CH2 Cl2 /MeOH): Rf 0.54. FAB+ : C9 H13 N3 O5 [M+H]: 244.1. 1 H NMR (300 MHz, DMSO-d6 ): δ 3.51-3.57 (2H, 2 × H5 ), 3.68 (1H, H4 ), 3.82-3.95 (2H, H3 , H2 ), 4.99, 5.04, 5.30 (3H, (d), (t) (d), J ∼ 4.5 Hz, OH), 5.70 (1H, d, J = 7.4 Hz, 5-H), 5.77 (1H, d, J = 3.7 Hz, H1 ), 7.19 (2H, s, br, ex, NH2 ), 7.83 (1H, d, J = 7.4 Hz, 6-H). 13 C NMR (75 MHz, DMSO-d6 ): δ 61.1 (C5 ), 69.9 (C2 ), 74.4 (C3 ), 84.5 (C4 ), 89.6 (C1 ), 94.4 (5-C), 142.0 (6-C), 155.9 (2-C), 166.0 (4-C). UV (H2 O) λmax 272 nm. BASIC PROTOCOL 3
PREPARATION OF ADENOSINE N6 -Benzoyladenine is first converted to N6,9 -bis(trimethylsilyl)-N6 -benzoyladenine (S.7) following the procedure outlined in Support Protocol 1. Silylated adenine (S.7) is then allowed to react with 1-O-acetyl-2,3,5-tri-O-benzoylribofuranose (S.9) and trimethylsilyl triflate (S.10) in dry 1,2-dichloroethane. The resulting N6 -benzoyladenine 2 ,3 ,5 -tri-Obenzoate (S.15) is treated with methanolic ammonia to remove the protecting groups, and the resulting adenosine is purified by recrystallization from methanol/water.
Materials
Synthesis of Ribonucleosides by Condensation Using Trimethylsilyl Triflate
N6,9 -Bis(trimethylsilyl)-N6 -benzoyladenine (S.7) obtained from 10 mmol N6 -benzoyladenine (see Support Protocol 1) 1-O-Acetyl-2,3,5-tri-O-benzoylribofuranose (S.9) Anhydrous 1,2-dichloroethane (reflux overnight on P2 O5 and distill before use) Trimethylsilyltrifluoromethanesulfonate (TMSOTf, S.10) Dichloromethane (CH2 Cl2 ) Methanol (MeOH) Saturated sodium bicarbonate (sat. NaHCO3 ) solution, ice cold Sodium sulfate (Na2 SO4 ) Silica gel (0.060 to 0.200 nm) Petroleum ether (optional) Saturated ammonia in methanol (sat. NH3 /MeOH)
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250-mL round-bottom flasks Balloon filled with nitrogen or argon Reflux condenser Precoated TLC plates (e.g., Alugram Sil G/UV254 , Macherey-Nagel) 254-nm UV lamp (for TLC) 500-mL separatory funnel 500-mL Erlenmeyer flask Filter paper Buchner funnel 250-mL filter flask Rotary evaporator equipped with a vacuum pump 5 × 50–cm chromatography column Desiccator Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare N6 -benzoyladenosine 2 ,3 ,5 -tri-O-benzoate (S.15) 1. Dissolve the solid yellowish N 6,9 -bis(trimethylsilyl)-N 6 -benzoyladenine (S.7) obtained from 10 mmol N6 -benzoyladenine in Support Protocol 1, plus 5.04 g (10 mmol) 1-O-acetyl-2,3,5-tri-O-benzoylribofuranose (S.9), in 25 mL dry 1,2-dichloroethane in a 250-mL round-bottom flask. 2. Add 0.22 g (1.0 mmol) TMSOTf (S.10) and heat the mixture under nitrogen protection under reflux (90◦ -95◦ C) for 4 hr. CAUTION: TMSOTf is toxic and reacts violently with water, and should be protected from moisture.
3. Monitor by TLC (APPENDIX 3D) using 95:5 (v/v) CH2 Cl2 /MeOH (Rf = 0.82). When all starting material is depleted, cool the mixture to room temperature and then dilute the clear yellow solution with 100 mL CH2 Cl2 . 4. Extract with 50 mL ice-cold sat. NaHCO3 solution using a 500-mL separatory funnel. 5. Wash the organic layer three times with 20 mL water. 6. Collect the organic layers in a 500-mL Erlenmeyer flask and dry over Na2 SO4 . Filter off the drying agent using filter paper and 250-mL round-bottom flask, and evaporate to dryness (brownish foam) on a rotary evaporator. Yield of crude N6 -benzoyladenosine 2 ,3 ,5 -tri-O-benzoate (S.15): 7.10 g (quant.).
7. To remove traces of impurities, purify by column chromatography (APPENDIX 3E) using a 5 × 50–cm column with 500 g silica gel, eluting with 2.5 L of 95:5 (v/v) CH2 Cl2 /MeOH, or crystallize the crude product from a minimum amount of petroleum ether. Yield of N6 -benzoyladenosine 2 ,3 ,5 -tri-O-benzoate (S.15) after chromatography: 6.73 g (95%.). m.p. 88◦ -92◦ C. TLC (95:5 CH2 Cl2 /MeOH): Rf 0.82. FAB+ : C38 H30 N5 O8 [M+H]: 684.2. 1 H NMR (300 MHz, CDCl3 ): δ 4.73-4.97 (3H, 2 × H5 , H4 ), 6.31 (1H, H3 ), 6.46 (1H, H2 ), 6.53 (1H, s, H1 ), 7.38-8.13 (20H, 4 × Bz), 8.23 (1H, s, 2-H), 8.71 (1H, s, 8-H), 9.31 (1H, s, br, ex, NH). 13 C NMR (75 MHz, CDCl3 ): δ 63.5 (C5 ), 71.5 (C2 ), 73.9 (C3 ), 80.9 (C4 ), 87.0 (C1 ), 123.6 (5-C), 127.9-133.8 (4 × C Bz), 141.7 (8-C), 149.8 (6-C), 151.7 (4-C), 152.9 (2-C), 165.1, 165.3, 165.4, 166.1 (4 × C=O Bz).
Prepare adenosine (S.16) 8. Dissolve 7.00 g (10 mmol) S.15 in 250 mL sat. NH3 /MeOH. Keep the reaction mixture overnight at room temperature.
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9. Evaporate the volatiles on a rotary evaporator to obtain a crystalline material. 10. Stir material in 10 mL of 1:1 (v/v) CH2 Cl2 /MeOH for 2 hr at room temperature. 11. Filter off the crystals using a Buchner funnel and 250-mL filter flask, and wash with 20 mL CH2 Cl2 . 12. Crystallize from 200 mL of 2:1 (v/v) MeOH/water. After cooling, filter off crystals with a Buchner funnel, and then dry the product in a desiccator. Yield of pure adenosine (S.16): 2.16 g (80.9%). m.p. 143◦ -145◦ C. The spectroscopic properties (NMR, MS, UV, m.p.) of the obtained nucleosides are identical with those previously described. TLC (2:1 CH2 Cl2 /MeOH): Rf 0.51. FAB+ : C10 H14 N5 O4 [M+H]: 268.1. 1 H NMR (300 MHz, DMSO-d6 ): δ 3.52-3.59 (2H, 2 × H5 ), 3.97 (1H, H3 ), 4.15 (1H, H4 ), 4.60 (1H, H2 ), 5.18, 5.44 (3H, OH), 5.88 (1H, d, J = 6.2 Hz, H1 ), 7.35 (2H, s, br, ex, NH2 ), 8.14 (1H, s, 2-H), 8.35 (1H, s, 8-H). 13 C NMR (75 MHz, DMSO-d6 ): δ 62.1 (C5 ), 71.1 (C2 ), 73.9 (C3 ), 86.4 (C4 ), 88.4 (C1 ), 119.8 (5-C), 140.4 (8-C), 149.5 (4-C), 152.8 (2-C), 156.6 (6-C). UV (H2 O) λmax 259 nm. BASIC PROTOCOL 4
PREPARATION OF GUANOSINE N2 -Acetylguanine (S.4) is first converted to 6-(trimethylsilyloxy)-2-(N-trimethylsilylacetamido)-(N9 -trimethylsilyl)purine (S.8) following the procedure outlined in Support Protocol 1. Silylated guanine S.8 is then allowed to react with 1-O-acetyl-2,3,5-tri-Obenzoylribofuranose (S.9) and trimethylsilyl triflate (S.10) in dry 1,2-dichloroethane. The resulting N2 -acetylguanosine 2 ,3 ,5 -tri-O-benzoate (S.17) is treated with methanolic ammonia to remove the protecting groups, and the resulting guanosine (S.18) purified by recrystallization from water.
Materials
Synthesis of Ribonucleosides by Condensation Using Trimethylsilyl Triflate
6-(Trimethylsilyloxy)-2-(N-trimethylsilylacetamido)-(N9 -trimethylsilyl)purine (S.8) obtained from 4.09 mmol N2 -acetylguanine (S.4) (see Support Protocol 1) 1-O-Acetyl-2,3,5-tri-O-benzoylribofuranose (S.9) Anhydrous 1,2-dichloroethane (reflux overnight on P2 O5 and distill before use) Trimethylsilyltrifluoromethanesulfonate (TMSOTf, S.10) Dichloromethane (CH2 Cl2 ) Methanol (MeOH) Saturated sodium bicarbonate solution (sat. NaHCO3 solution), ice cold Sodium sulfate (Na2 SO4 ) Silica gel (0.060 to 0.200 nm) Saturated ammonia in methanol (sat. NH3 /MeOH) 250-mL round-bottom flask Balloon filled with nitrogen or argon Reflux condenser Precoated TLC plates (e.g., Alugram Sil G/UV254 , Macherey-Nagel) 254-nm UV lamp (for TLC) 500-mL separatory funnel 500-mL Erlenmeyer flask Filter paper Buchner funnel 250-mL filter flask Rotary evaporator equipped with a vacuum pump 5 × 35–cm chromatography column Desiccator Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E)
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Prepare N2 -acetylguanosine 2 ,3 ,5 -tri-O-benzoate (S.17) 1. Dissolve the solid yellowish 6-(trimethylsilyloxy)-2-(N-trimethylsilylacetamido)(N9 -trimethylsilyl)purine (S.8) obtained from 4.09 mmol N2 -acetylguanine (S.4) in Support Protocol 1, and 1.86 (3.7 mmol) 1-O-acetyl-2,3,5-tri-O-benzoylribofuranose (S.9), in 35 mL dry 1,2-dichloroethane in a 250-mL round-bottom flask. 2. Add 0.99 g (4.46 mmol) TMSOTf (S.10) and heat the mixture under nitrogen protection under reflux (90◦ -95◦ C) for 1.5 hr. CAUTION: TMSOTf is toxic and reacts violently with water, and should be protected from moisture.
3. Monitor by TLC (APPENDIX 3D) using 9:1 CH2 Cl2 /MeOH (Rf = 0.72). When all starting material is depleted, cool the mixture to room temperature, and dilute the clear yellow solution with 20 mL CH2 Cl2 . 4. Extract with 20 mL ice-cold sat. NaHCO3 solution using a 500-mL separatory funnel. 5. Wash the organic layer three times with 20 mL water. 6. Collect the organic layers in a 500-mL Erlenmeyer flask and dry over Na2 SO4 . Filter off the drying agent using filter paper and 250-mL round-bottom flask, and evaporate to dryness on a rotary evaporator. Yield: 2.32 g (98%) crude product.
7. To remove traces of impurities (i.e., the N7-isomer), purify by column chromatography (APPENDIX 3E) using a 5 × 35–cm column with 250 g of silica gel, eluting with 1.5 L of 9:1 (v/v) CH2 Cl2 /MeOH. Yield of N2 -acetylguanosine 2 ,3 ,5 -tri-O-benzoate (S.17): 2.32 g as a yellowish foam. TLC (9:1 CH2 Cl2 /MeOH): Rf 0.72 (0.60 for N7 isomer). FAB+ : C33 H28 N5 O9 [M+H]: 638.2. 1 H NMR (300 MHz, CDCl3 ): δ 2.32 (3H, s, CH3 ), 4.70-5.00 (3H, 2 × H5 , H4 ), 6.24 (1H, H3 ), 6.35 (2H, H2 , H1 ), 7.27-8.10 (15H, 3 × Bz), 7.91 (1H, s, 8-H), 9.78 (1H, s, br, ex, 1-NH), 12.0 (1H, s, br, ex, NHAc). 13 C NMR (75 MHz, CDCl3 )): δ 24.3 (CH3 ), 63.0 (C5 ), 71.5 (C2 ), 74.0 (C3 ), 80.0 (C4 ), 88.3 (C1 ), 122.8 (5-C), 128.4-133.9 (3 × C Bz), 133.9 (8-C), 138.9 (2-C), 147.5 (4-C), 155.5 (6-C), 165.2, 165.4, 166.8 (3 × C=O Bz), 172.0 (C=O Ac).
Prepare guanosine (S.18) 8. Dissolve 2.30 g (3.60 mmol) S.17 in 125 mL sat. NH3 /MeOH. Keep the reaction mixture for 42 hr at room temperature. 9. Evaporate the volatiles on a rotary evaporator to obtain a colorless crystalline material. 10. Stir material in 10 mL of 1:1 (v/v) CH2 Cl2 /MeOH for 2 hr at room temperature. 11. Filter off the crystals using a Buchner funnel and 250-mL filter flask, and wash with 20 mL CH2 Cl2 . 12. Crystallize from a minimum amount of boiling water and then dry the product in a desiccator. Yield of guanosine (S.18): 0.69 g (66%). m.p. 238◦ -239◦ C. The spectroscopic properties (NMR, MS, UV, m.p.) of the obtained nucleosides are identical with those previously described. TLC (2:1 CH2 Cl2 /MeOH): 0.46. FAB+ : C10 H14 N5 O5 [M+H]: 284.1. 1 H NMR (300 MHz, DMSO-d6 ): δ 3.52-3.60 (2H, 2 × H5 ), 3.86 (1H, H3 ), 4.08 (1H, H4 ),4.39 (1H, H2 ), 5.02, 5.11, 5.39 (3H, (t), (d) (d), J ∼ 6.0 Hz, OH), 5.69 (1H, d, J = 6.0 Hz, H1 ), 6.46 (2H, s, br, ex, NH2 ), 7.94 (1H, s, 8-H), 10.65 (1H, s, br, ex, NH). 13 C NMR (75 MHz, DMSO-d6 ): δ = 61.8 (C5 ), 70.8 (C2 ), 74.1 (C3 ), 85.6 (C4 ), 86.8 (C1 ), 117.1 (5-C), 136.0 (8-C), 151.8 (4-C), 154.1 (2-C), 157.2 (6-C). UV (H2 O) λmax 251 nm.
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SUPPORT PROTOCOL 1
PREPARATION OF SILYLATED HETEROCYCLES USING HMDS This procedure is used for conversion of uracil (S.1) to 2,4-bis(trimethylsilyloxy)pyrimidine (S.5), cytosine (S.2) to 4-(trimethylsilylamino)-2-(trimethylsilyloxy)pyrimidine (S.6), N6 -benzoyladenine (S.3) to N6,9 -bis(trimethylsilyl)-N6 -benzoyladenine (S.7), and N2 -acetylguanine (S.4) to 6-(trimethylsilyloxy)-2-(N-trimethylsilylacetamido)-(N9 trimethylsilyl)purine (S.8). It is applicable to most other heterocyclic bases.
Materials Heterocyclic base, e.g., uracil (S.1), cytosine (S.2), N6 -benzoyladenine (S.3), or N2 -acetylguanine (S.4) Hexamethyldisilazane (HMDS) Ammonium chloride (NH4 Cl) Trimethylchlorosilane (TCS) Pyridine Xylenes (mixture of isomers, extra pure; Acros) 50-mL round-bottom flask Reflux condenser 1. In a 50-mL round-bottom flask with a reflux condenser, heat 10 mmol heterocyclic base in 20 to 30 mL HMDS and a few crystals of NH4 Cl to 145◦ C. 2. If the base does not dissolve after 0.5 to 2 hr, add either 0.1 mL trimethylchlorosilane (TCS; e.g., for 5-nitrouracil, lumazine) or 10 mL pyridine (e.g., for 4-aminopyridine, N6 -benzoyladenine, N2 -acetylguanine, xanthine) to accelerate the silylation. 3. After a clear solution is obtained, remove excess HMDS by co-distillation with two 25- to 50-mL portions of xylenes. 4. Use the obtained silylated base immediately (no purification) for the condensation reaction. SUPPORT PROTOCOL 2
PREPARATION OF SILYLATED HETEROCYCLES USING BSA This silylation procedure is applicable to nucleic acid bases and other closely related substituted pyrimidines and purines. The advantage of this method is that the silylated product is prepared in situ, making the evaporation of the solvent and redissolving of the silylated base unnecessary.
Materials Heterocyclic base, e.g., uracil (S.1), cytosine (S.2), N6 -benzoyladenine (S.3), or N2 -acetylguanine (S.4) Anhydrous 1,2-dichloroethane Nitrogen source N,O-Bis(trimethylsilyl)acetamide (BSA) 1. Suspend 10 mmol heterocyclic base in 25 mL of 1,2-dichloroethane. 2. Under a nitrogen atmosphere, add BSA at 1.1 eq. per NH of the base (i.e., 22 mmol for uracil, 44 mmol for 6-aminouracil). 3. Stir at room temperature until a clear solution is obtained (30 to 60 min). Synthesis of Ribonucleosides by Condensation Using Trimethylsilyl Triflate
4. Add the protected sugar and TMSOTf and continue as described in Basic Protocols 1 through 4.
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COMMENTARY Background Information The protocols in this unit provide the most efficient procedure for synthesis of ribonucleosides from heterocyclic bases. There are a wide variety of methods for synthesis of nucleosides from purine and pyrimidine as well as many other heterocycles; these have been extensively reviewed elsewhere (Vorbr¨uggen and Ruh-Pohlenz, 2000, 2001). The Lewis acid– mediated glycosylation of silylated pyrimidines and purines is one of the most efficient high-yield reactions in nucleoside chemistry. The introduction of Friedel-Crafts catalysts such as SnCl4 (S.20; Niedballa and Vorbr¨uggen, 1970, 1974a-e) for the sugar-base condensation reaction as a more effective version of the silyl-Hilbert-Johnson synthesis of nucleosides, was followed by the use of the Lewis acid trimethylsilyl triflate (TMSOTf; S.10) in the condensation reaction, which is the subject of this unit (Vorbr¨uggen and Kroliliewicz, 1975; Vorbr¨uggen et al., 1981). Understanding the mechanism of this reaction is essential for its effective application. The details of the mechanism have been worked out for the two C5-substituted pyrimidine nucleosides 5-methoxy- and 5-morpholinouridine, which are reviewed here to illustrate the pathway likely followed for most silylated pyrimidine bases upon TMSOTf-mediated glycosylation, including the two syntheses outlined in Basic Protocols 1 and 2. The TMSOTfmediated glycosylation of silylated purines is more complex. The proposed mechanistic pathways are outlined below for the reactions in Basic Protocols 3 and 4. Synthesis of pyrimidine ribonucleosides Due to their basic character, silylated nucleobases form σ-complexes with Lewis acids (Fig. 1.13.3). In the case of stronger basic heterocycles, an equivalent amount of the Lewis acid is “neutralized or deactivated” as σcomplex with the silylated base. Therefore, any further reaction of these σ-complexes is much slower. In the 13 C NMR spectra of the (rather basic) silylated 5-methoxyuracil (S.22, R = OMe) in the presence of TMSOTf (S.10) or SnCl4 (S.20), the signal of C6 at 139 ppm is broadened for the SnCl4 -σ-complex. This proves a tight binding of SnCl4 to N1, the center of highest electron density. The corresponding C6 signal of the σ-complex S.25 is sharp and indicates a rather loose binding and a rapid exchange of the weaker Lewis acid with
N1 (Vorbr¨uggen et al., 1978; Vorbr¨uggen and Hoefle, 1981; Fig. 1.13.3). As a result, TMSOTf, which is a much weaker Lewis acid than SnCl4 , is a superior if not optimal catalyst for nucleoside synthesis. TMSOTf converts stable O-acylated sugars such as S.9 via O-silylation of the 1-β-Oacetyl group into trimethylsilyl acetate and reactive cyclic sugar salts (S.21). However, the decreased acidity of TMSOTf leads to lower amounts of the σ-complex S.25 (formed between S.22 and TMSOTf) than of the σcomplex S.27 (formed between S.22 and the stronger Lewis acid SnCl4 ) (Vorbr¨uggen et al., 1978; Vorbr¨uggen and Hoefle, 1981). Whereas only free persilylated base S.22 reacts with the activated sugar salt S.21 to give the desired silylated N1 -nucleoside S.23, the σ-complex S.25 (in which N1 is blocked) can only condense in a slower reaction with S.21 to deliver the undesired N3 -nucleoside S.26. Particularly, in the rather apolar solvent 1,2-dichlorethane, less of the N1 -nucleoside S.23 and more of the N3 -nucleoside S.26 is obtained, since much more of the σ-complex S.27 is formed during reaction with SnCl4 . Both the silylated N1 -nucleoside S.23 and, in particular, the undesired silylated N3 -nucleoside S.26 can further react with S.21 to give the N1 ,N3 bisriboside S.24. The more polar solvent acetonitrile (in comparison to 1,2-dichlorethane) also forms σ-complexes with S.10, or with S.20 (Vorbr¨uggen and Ruh-Pohlenz, 2001), as depicted in Figure 1.13.3. Therefore, the polar acetonitrile as solvent competes with the silylated base S.22 for the Lewis acids S.10 and S.20, resulting in formation of reduced amounts of the σ-complexes S.25 or S.27, and consequently a diminished formation of the N3 -nucleoside S.26 and increased formation of the desired natural protected N1 -nucleoside S.23 (Vorbr¨uggen et al., 1978; Vorbr¨uggen and Hoefle, 1981; Fig. 1.13.3). In particular, in the case of basic silylated pyrimidines such as silylated 5methoxyuracil S.22 (R = OMe) or silylated 5-morpholinouracil S.22 (R = C4 H8 NO), the use of TMSOTf instead of SnCl4 results (after workup with aqueous bicarbonate) in much higher yields of the desired protected natural N1 -nucleosides S.23 (R = OMe and R = C4 H8 NO) and a dramatically reduced formation of the undesired protected N3 -nucleosides S8 (R = OMe and R = C4 H8 NO) and N1 ,N3 -bisribosides S.24 (Vorbr¨uggen et al.,
Synthesis of Modified Nucleosides
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Figure 1.13.3 Pathway for Lewis acid–mediated condensation of 1-O-acetyl-2,3,5-tri-O-benzoylribofuranose (S.9) with 5-methoxy- and 5-morpholino-2,4-bis(trimethylsilyloxy)pyrimidine (S.22).
Synthesis of Ribonucleosides by Condensation Using Trimethylsilyl Triflate
1978; Vorbr¨uggen and Hoefle, 1981). Reactions in acetonitrile (after aqueous workup) lead to higher yields of the desired protected N1 -nucleosides S.23 and thus decreased amounts of the undesired N3 -nucleosides S26 (R = OMe and R = C4 H8 NO) for previously mentioned reasons. In the less polar solvent 1,2-dichloroethane, much more σcomplex between the silylated bases and the Lewis acids is formed, resulting in more N3 nucleosides and N1 ,N3 -bisribosides. The variation in yields of the N1 -riboside, N3 -riboside, and N1 ,N3 -bisriboside as a function of Lewis acid and solvent is summarized in Figure
1.13.4 (Vorbr¨uggen et al., 1978; Vorbr¨uggen and Hoefle, 1981). Silylated cytosine forms strong σcomplexes with Lewis acids S.10 and S.20, in analogy to the silylated uracils S.23. Therefore, the less basic silylated N4 -acylated cytosine should always be used in acetonitrile for the TMSOTf- or SnCl4 -catalyzed silyl-Hilbert-Johnson synthesis of cytidines (Vorbr¨uggen and Ruh-Pohlenz, 2000, 2001). Synthesis of purine ribonucleosides The advantage of employing TMSOTf as catalystis even more evident for the
1.13.12 Supplement 27
Current Protocols in Nucleic Acid Chemistry
N3
OSiMe3 R
R
1
Me3SiO
1) Lewis acid 10 or 20 solvent
N
22
BzO
HN
BzO
BzO
BzO
N
O BzO
OBz
BzO
R
N
N 1, N 3-bisribosides O
O
OAc O
1 3
O
NH O
2) NaHCO3
O
24
OBz 29
28 OBz
9
a) R = -OMe
b) R = -N
Figure 1.13.4
O
28
29
24
TMSOTf (10), ClCH2CH2Cl SnCl4 (20), ClCH2CH2Cl SnCl4 (20), CH3CN
89% 53% 90%
27% 3%
13% -
TMSOTf (10), ClCH2CH2Cl SnCl4 ( 20), ClCH2CH2Cl SnCl4 ( 20), CH3CN
89% 53% 90%
27% 3%
13% -
Variation of N 1 and N 3 glycosylation yields with Lewis acid and solvent.
synthesis of purine nucleosides. On the basis of 13 C NMR studies (Vorbr¨uggen and Hoefle, 1981) following the downfield shift of C8, it is assumed that the most basic N1 in persilylated N6 -benzoyladenine (S.7) is trimethylsilylated by TMSOTf to form the σ-complex S.30 (Fig. 1.13.5). This complex eliminates TMSOTf to form the activated putative isomeric silylated N6 -benzoyladenine, S.31. It was suggested (Vorbr¨uggen and Ruh-Pohlenz, 2000) that compounds S.7, S.30, and S.31 react reversibly with the sugar salt S.21 in an initially kinetically controlled reaction to give the protected N1 -nucleoside S.32 and the N3 nucleoside S.33, the latter of which can be isolated at the start of the reaction. However, S.32 and S.33 occur in equilibrium with S.31 and S.21, and subsequent reaction of S.31 with S.21 furnishes primarily a mixture of the undesired protected N7 -nucleoside S.34 and the desired N9 -nucleoside S.35. On boiling the reaction mixture in the rather unpolar solvent 1,2-dichloroethane (compared to acetonitrile), σ-complex formation of the undesired products S.32, S.33, and S.34 with TMSOTf is favored, and thus rearrangement and formation of equilibria between S.32, S.33, and S.34 with S.31 and S.21 occurs (cf. the depicted TMSOTf-catalyzed cleavage of S.34 to S.31
and S.21 shown in the box at the bottom of Fig. 1.13.5). This eventually results in the nearly exclusive formation of the desired and thermodynamically favored protected natural N9 nucleoside adenosine S.35. On ammonolysis of S.35 with methanolic ammonia, crystalline adenosine (S.16) is obtained in 81% overall yield starting from S.7 and S.9 (Vorbr¨uggen et al., 1981; Fig. 1.13.5). Framski et al. (2006) have demonstrated that on reacting N6 -benzoyladenine S.3 in the presence of TMSOTf with 1,2,3,5-tetraO-acetyl-β-D-ribofuranose (whose bridged cation corresponding to S.9 is less stable than that of the benzoylated intermediate S.9 and whose nucleosides consequently rearrange more slowly), the initially formed N1 -nucleoside can be isolated as the major product, together with the corresponding N9 -nucleoside as well as the N1 ,N9 bisriboside. Persilylated N2 -acetylguanine S.4 affords, on reaction with the protected sugar S.9 in the presence of TMSOTf in boiling 1,2dichloroethane and subsequent ammonolysis with methanolic ammonia, crystalline natural guanosine S.18 in 66% overall yield (Niedballa and Vorbr¨uggen, 1976). These results were confirmed by others, and the ratio between the desired natural N9 -guanosine S.18
Synthesis of Modified Nucleosides
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Bz
N
SiMe3 N
N1
7 9
3
Bz
TMSOTf (10) ClCH2CH2Cl ∆
Me3Si
SiMe3
N 30
Bz
TMSOTf (10) ClCH2CH2Cl ∆
N
N 3
N SiMe3
N
N
7 9
Me3Si
N SiMe3
31
± 21
± 21
BzO
N O BzO
Bz
N SiMe3 N
N
N
N
3
1
N
BzO
N Bz SiMe3
O BzO
32
BzO
N
O
Bz
N SiMe3
OBz
BzO
O
N
OBz
35
O 31
O O
BzO 21
BzO
N
BzO
N
N
BzO
Me3Si ........OTf
N O
SiMe3
34
33
N
N
SiMe3
9
BzO
O
N
N N
N
N
OBz
N
7
BzO
Bz
OBz
N
N
Bz N
N
7 9
3
± 21
N
SiMe3
N1
7
± 21
N
OTf Ph
O Ph
Figure 1.13.5
Pathway for TMSOTf (S.10)-mediated glycosylation of N 6,9 -bis(trimethylsilyl)-N 6 -benzoyladenine (S.7).
Synthesis of Ribonucleosides by Condensation Using Trimethylsilyl Triflate
and the undesired N7 -guanosine S.19 was estimated to be 6:1 (Fig. 1.13.2). The use of SnCl4 in acetonitrile or 1,2-dichloroethane leads to an excess of the undesired N7 -guanosine S.20 (Garrner and Ramakanth, 1988). 1,2-Dichloroethane (which favors σcomplex formation of the purine moieties with TMSOTf and also favors the rearrangement of the initially formed kinetically controlled intermediates to give the thermodynamically most stable products) is thus the preferred solvent for the synthesis of purine nucleosides (Vorbr¨uggen et al. 1981; Vorbr¨uggen and RuhPohlenz, 2000, 2001). Finally, in comparing the TMSOTf with SnCl4, the workup using TMSOTf is much simpler. By addition of an ice-cold solution of aqueous sodium bicarbonate to the reaction mixtures, no emulsions are obtained using TMSOTf as they nearly always are upon aqueous workup of reactions employing
SnCl4 (Vorbr¨uggen et al., 1981; Vorbr¨uggen and Ruh-Pohlenz, 2000, 2001). Alternative procedures Working under careful exclusion of moisture during the silylation of polar heterocyclic bases with hexamethyldisilazane (HMDS) and during the subsequent nucleoside synthesis is often rather difficult for scientists with only limited training in preparative organic chemistry. One can, therefore, simplify the nucleoside synthesis by combining the reactions for (1) acid-catalyzed silylation of the heterocyclic bases, (2) silylation of free triflic acid to give TMSOTf, and (3) nucleoside synthesis in acetonitrile (Vorbr¨uggen and Bennua, 1981). In this one-pot reaction, purines such as adenine and N6 -benzoyladenine, or pyrimidines such as N4 -acylcytidines, can also be reacted with protected sugars such as S.9 in the presence of excess SnCl4 . The catalyst
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Current Protocols in Nucleic Acid Chemistry
apparently forms partially soluble σcomplexes with these bases in ethylene chloride, producing protected nucleosides in yields of 70% to 90%. The Lewis acid–catalyzed condensation of nucleobases and acyl-protected sugars has become a well-established method over the years, allowing, e.g., the preparation of the naturally occurring β-nucleosides in good yields. The method has not only been applied to the preparation of ribosides, but also for sugar and/or base-modified nucleosides (Vorbr¨uggen and Ruh-Pohlenz, 2000).
Critical Parameters and Troubleshooting For all syntheses, freshly prepared dry solvents and water-free reagents are used. The reactions are carried out in oven-dried glassware under exclusion of moisture, preferably under nitrogen or argon protection. Neutralization of the reaction mixture with a saturated solution of sodium bicarbonate must be done very carefully (foaming). Reactions should be monitored by TLC, especially if the kinetic reaction products are desired. Silylation with HMDS can be replaced by silylation with N,O-bis(trimethylsilyl)acetamide (BSA), which appears to be a very gentle working silylating agent. However, when using BSA, it has to be taken into consideration that the N-monosilylacetamide formed can interact as a “base” in the nucleoside synthesis (Vorbr¨uggen and Ruh-Pohlenz, 2000). In the case of very poorly soluble nuclebases, an excess of nucleobase is used for the glycosylation reaction to obtain a better yield of the desired nucleoside. In these cases, after quenching the reaction mixture with sat. NaHCO3 solution, the mixture should first be filtered (paper filter) to remove the unreacted base. The resulting residue is then washed with CH2 Cl2 . The organic and aqueous layers of the filtrate are separated and further treated as described.
Anticipated Results The synthesis of ribonucleosides by condensation using trimethylsilyl triflate is a generally applicable method for the synthesis of the naturally occurring β-nucleosides. The quality of the reagents and dryness of the solvents will have a significant impact on product yield and purity. This method is also applicable for the synthesis of base-modified ribonucleosides or for sugar-modified nucleosides. In the latter case, a 1,2-diacyl (acyl = acetyl, benzoyl. . .) proCurrent Protocols in Nucleic Acid Chemistry
tection is necessary to obtain β-nucleosides following the herein discussed reaction mechanism. For the naturally occurring nucleosides and closely related bases, coupling yields in the range of 80% to 95% (pyrimidines) and 60% to 85% (purines) can be expected. All reactions described in this unit can be scaled up to yield ∼500 mmol of the nucleosides by linearly increasing the amounts of all materials used in the procedures. Reaction times have to be adjusted by TLC monitoring.
Time Considerations Deprotected β-nucleosides (A, G, C, U) and closely related nucleosides can be obtained in 2 days. On the first day, the glycosylation reaction is carried out (silylation ∼1 hr, condensation 2 to 4 hr) followed by flash chromatography to obtain the pure, protected nucleoside derivatives. The deprotection reaction is preferably carried out overnight. The second day is used for drying the product in a desiccator and for spectroscopic characterization. When glycosylating various unnatural heterocyclic bases, the time needed will depend on the reaction time of the glycosylation reaction (in most cases, between 2 and 10 hr).
Literature Cited Framski, G., Gdaniec, Z., Gdaniec, M., and Boryski, J. 2006. A reinvestigated mechanism of ribosylation of adenine under silylating conditions. Tetrahedron 62:10123-10129. Garner, P. and Ramakanth, S. 1988. A regioncontrolled synthesis of N7 - and N9 -guanine nucleosides. J. Org. Chem. 53:1294-1298. Niedballa, U. and Vorbr¨uggen, H. 1970. A general synthesis of pyrimidine nucleosides. Angew. Chem. Int. Ed. 9:461-462. Niedballa, U. and Vorbr¨uggen, H. 1974a. Synthesis of nucleosides. 9. General synthesis of Nglycosides. I. Synthesis of pyrimidine nucleosides. J. Org. Chem. 39:3654-3659. Niedballa, U. and Vorbr¨uggen, H. 1974b. Synthesis of nucleosides. 10. General synthesis of N-glycosides. II. Synthesis of 6-methyluridines. J. Org. Chem. 39:3660-3663. Niedballa, U. and Vorbr¨uggen, H. 1974c. Synthesis of nucleosides. 11. General synthesis of N-glycosides. III. Simple synthesis of pyrimidine disaccharide nucleosides. J. Org. Chem. 39:3664-3667. Niedballa, U. and Vorbr¨uggen, H. 1974d. Synthesis of nucleosides. 12. General synthesis of N-glycosides. IV. Synthesis of nucleosides of hydroxy and mercapto nitrogen heterocycles. J. Org. Chem. 39:3668-3671. Niedballa, U. and Vorbr¨uggen, H. 1974e. Synthesis of nucleosides. 13. General synthesis of N-glycosides. V. Synthesis of 5-azacytidines. J. Org. Chem. 39:3672-3673.
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Niedballa, U. and Vorbr¨uggen, H. 1976. Synthesis of nucleosides. 17. A general synthesis of Nglycosides. 6. On the mechanism of the stannic chloride catalyzed silyl Hilbert-Johnson reaction. J. Org. Chem. 41:2084-2086. Vorbr¨uggen, H. and Bennua, B. 1981. Nucleoside syntheses. XXV. A new simplified nucleoside synthesis. Chem. Ber. 114:1279-1286. Vorbr¨uggen, H. and Hoefle, G. 1981. Nucleoside syntheses. XXIII. On the mechanism of nucleoside synthesis. Chem. Ber. 114:1256-1268. Vorbr¨uggen, H. and Krolikiewicz, K. 1975. New catalysts for nucleoside synthesis. Angew. Chem. 87:417. Vorbr¨uggen, H. and Ruh-Pohlenz, C. 2000. Synthesis of nucleosides. Organic Reactions 55:1630. Vorbr¨uggen, H. and Ruh-Pohlenz, C. 2001. Handbook of Nucleoside Synthesis. Wiley Interscience, New York.
Vorbr¨uggen, H., Niedballa, U., Krolikiewicz, K., Bennua, B., and H¨ofle, G. 1978. On the mechanism of nucleoside synthesis. In Chemistry and Biology of Nucleosides and Nucleotides (R.E. Harman,R.K. Robins, andL.B. Townsend, eds.) pp. 251-266. Academic Press, New York. Vorbr¨uggen, H., Krolikiewicz, K., and Bennua, B. 1981. Nucleoside syntheses. XXII. Nucleoside synthesis with trimethylsilyl triflate and perchlorate as catalysts. Chem. Ber. 114:1234-1255.
Contributed by Helmut Vorbr¨uggen Free University Berlin Berlin, Germany Irene M. Lagoja and Piet Herdewijn Rega Institute for Medical Research Leuven, Belgium
Synthesis of Ribonucleosides by Condensation Using Trimethylsilyl Triflate
1.13.16 Supplement 27
Current Protocols in Nucleic Acid Chemistry
Synthesis of 2 -O-β-dRibofuranosylnucleosides
UNIT 1.14
This unit describes a three-step procedure for the preparation of 2 -O-β-D-ribofuranosylnucleosides (Mikhailov et al., 1997a; Markiewicz et al., 1998). First, the complete synthesis of 2 -O-β-D-ribofuranosyladenosine is described (see Basic Protocol). As shown in Figure 1.14.1, the procedure involves (1) condensation of a small excess of 1-O-acetyl2,3,5-tri-O-benzoyl-β-D-ribofuranose activated with tin tetrachloride with an N-protected 3 ,5 -O-tetraisopropyldisiloxane-1,3-diyl-ribonucleoside in 1,2-dichloroethane, (2) removal of silyl protecting group with tetrabutylammonium fluoride, and (3) deacylation with ammonia in methanol. Using these procedures, 2 -O-β-D-ribofuranosyladenosine is prepared in a 61% overall yield. The 2 -O-ribosylation proceeds stereospecifically with the formation of a β-glycosidic bond. The presence of a participating 2-O-benzoyl group leads exclusively to 1,2-trans-ribofuranoside. This reaction is carried out under mild conditions (0◦ C, 1,2-dichloroethane, 2 hr for pyrimidine nucleosides, 7 to 16 hr for purine derivatives) and yields of the target compounds are 72% to 80% (Mikhailov et al., 1997a). At room temperature, the reaction occurs faster (30 min), but the yields in the coupling steps are lower (40% to 77%; Markiewicz et al., 1998). These variations are presented as slightly modified procedures for the preparation of four other 2 -O-β-D-ribofuranosylnucleosides (see Alternate Protocols 1 through 4; Fig. 1.14.3). CAUTION: Carry out all operations involving organic solvents and reagents in a wellventilated fume hood, and wear gloves and protective glasses.
PREPARATION OF 2 -O-β -D-RIBOFURANOSYLADENOSINE The first step of preparation of 2 -O-β-D-ribofuranosyladenosine involves the condensation of N6 -benzoyl-3 ,5 -(1,1,3,3-O-tetraisopropyldisiloxane-1,3-diyl)adenosine (S.1; Markiewicz and Wiewiorowski, 1986; UNIT 2.4) with 1-O-acetyl-2,3,5-tri-O-benzoyl-β-Dribofuranose (S.2) in the presence of tin tetrachloride in 1,2-dichloroethane at 0◦ C (Fig. 1.14.1). To obtain the product S.3 in high yield and to simplify its isolation, it is recommended to run the reaction under nitrogen and to perform preactivation of the sugar with tin tetrachloride; usually the reaction is complete in 7 to 8 hr. During workup of the reaction mixture with saturated sodium bicarbonate solution, a suspension is formed, which should be filtered through a 2- to 3-cm layer of Hyflo Super Cel before the separation of organic and aqueous layers. The second step involves removal of the silyl protection from the 3 - and 5 -hydroxyl groups with tetrabutylammonium fluoride (TBAF) in tetrahydrofuran (THF) for 15 to 20 min at room temperature followed by silica gel column chromatography. In the last step, all benzoyl groups from S.4 are removed with 5 M ammonia (half-saturated at 0◦ C) in methanol for 2 to 3 days at room temperature followed by recrystallization from methanol to give 2 -O-β-D-ribofuranosyladenosine (S.5) in an overall yield of 61%.
BASIC PROTOCOL
Materials 1-O-Acetyl-2,3,5-tri-O-benzoyl-β-D-ribofuranose (S.2) N6 -Benzoyl-3 ,5 -(1,1,3,3-O-tetraisopropyldisiloxane-1,3-diyl)adenosine (S.1; UNIT 2.4; Fig. 2.4.5) Phosphorus pentoxide (P2 O5 ) Balloon of nitrogen or argon 1,2-Dichloroethane, anhydrous Tin tetrachloride (SnCl4 ) Methanol (MeOH), analytical grade
Synthesis of Modified Nucleosides
Contributed by Sergey N. Mikhailov, Ekaterina V. Efimtseva, Andrei A. Rodionov, Georgii V. Bobkov, Irina V. Kulikova, and Piet Herdewijn
1.14.1
Current Protocols in Nucleic Acid Chemistry (2006) 1.14.1-1.14.19 C 2006 by John Wiley & Sons, Inc. Copyright
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Methylene chloride (CH2 Cl2 ), reagent grade Saturated sodium bicarbonate solution (sat. NaHCO3 ) Hyflo Super Cel (Fluka) Sodium sulfate, anhydrous (Na2 SO4 ) Silica gel (e.g., Kieselgel 60, 0.06 to 0.20 mm; Merck) Tetrabutylammonium fluoride trihydrate (TBAF) Tetrahydrofuran (THF), reagent grade Chloroform (CHCl3 ), reagent grade 5 M ammonia in methanol (half-saturated at 0◦ C) Diethyl ether, reagent grade 50- and 250-mL round-bottom flasks Vacuum desiccator Vacuum oil pump TLC plate: silica-coated aluminum plate with fluorescent indicator (Merck silica gel 60 F254 ) 254-nm UV lamp Long disposable capillaries 100-mL funnels with sintered glass disc filters (porosity 3) 100- and 250-mL separatory funnel Rotary evaporator equipped with a water aspirator 3 × 20–cm and 3 × 15–cm sintered glass chromatography columns, porosity 3 Stainless steel spatula Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare S.3 1. Weigh 1.26 g (2.5 mmol) of 1-O-acetyl-2,3,5-tri-O-benzoyl-β-D-ribofuranose (S.2) into a 250-mL round-bottom flask containing a stir bar and weigh 1.23 g (2 mmol) of nucleoside S.1 into a separate flask. 2. Put both flasks in a vacuum desiccator with phosphorus pentoxide and evacuate using a vacuum oil pump for 10 to 15 min. Close the stopcock of the desiccator and leave overnight at room temperature. 3. Connect the desiccator with a balloon of nitrogen or argon and open the stopcock of the desiccator. Open the desiccator and quickly close the flasks. 4. Dissolve 1-O-acetyl-2,3,5-tri-O-benzoyl-β-D-ribofuranose (S.2) in 25 mL of 1,2dichloroethane, close with stopcock and balloon of nitrogen or argon, and put the flask in the ice-water bath (Fig. 1.14.2A). 5. Add 0.35 mL (3 mmol) of tin tetrachloride in one portion under stirring and keep the reaction mixture for 10 min at 0◦ C. 6. Add 1.23 g (2 mmol) of nucleoside S.1 in one portion and stir the reaction mixture for 7 hr at 0◦ C. 7. Monitor reaction by TLC (APPENDIX 3D) using 2% (v/v) MeOH in CH2 Cl2 . To protect the reaction mixture from atmospheric moisture, remove the sample using a long capillary as shown in Figure 1.14.2B. The starting compound S.1 (Rf = 0.21) usually disappears after 6 to 7 hr at 0◦ C. The product S.3 (Rf = 0.31) moves faster in the same solvent. Synthesis of 2 -O-β β-dRibofuranosylnucleosides
1.14.2 Supplement 27
8. Add 10 mL saturated sodium bicarbonate solution and stir the suspension for 20 min at 0◦ C. 9. Lay a 2- to 3-cm layer of Hyflo Super Cel on a 100-mL funnel with sintered disc (porosity 3) and wash it with 20 mL CH2 Cl2 . Filter the suspension using a vacuum Current Protocols in Nucleic Acid Chemistry
Figure 1.14.1 Synthesis of 2 -O-β-D-ribofuranosyladenosine (S.5). The expected overall yield of S.5 from S.1 is given in parentheses. DCE, 1,2-dichloroethane; TBAF, tetrabutylammonium fluoride; THF, tetrahydrofuran.
pump, and wash the layer with 20 mL CH2 Cl2 and 20 mL of 5% (v/v) MeOH in CH2 Cl2 . 10. Separate the organic layer using a 250-mL separatory funnel and wash the organic layer with 20 mL water. 11. Dry the organic layer over ∼10 g Na2 SO4 , filter off Na2 SO4 by gravity filtration, wash the precipitate with 20 mL CH2 Cl2 , and evaporate the combined filtrates using a rotary evaporator connected to vacuum system. 12. Prepare a slurry of 50 g silica gel in CH2 Cl2 and pour into a 3 × 20–cm chromatography column (APPENDIX 3E). 13. Dissolve the residue in a minimal amount of CH2 Cl2 and layer it carefully on top of the silica gel. 14. Wash the column with 300 mL CH2 Cl2 and 300 mL of 0.5% (v/v) MeOH in CH2 Cl2 . Elute with 1% (v/v) MeOH in CH2 Cl2 . Collect 25-mL fractions.
Synthesis of Modified Nucleosides
1.14.3 Current Protocols in Nucleic Acid Chemistry
Supplement 27
Figure 1.14.2 Protection from atmospheric moisture for synthesis of 2 -O-β-D-ribofuranosylnucleosides. (A) A 250-mL round-bottom flask with adaptor and stopcock connected with a balloon of nitrogen or argon in an ice-water bath. (B) Checking the reaction mixture with a long capillary (diameter ∼1 mm). The diameter of the hole in the stopcock is ∼3 mm.
15. Evaluate fractions by TLC using 2% (v/v) MeOH in CH2 Cl2 and combine the fractions that contain only S.3. Evaporate the volatile materials from the combined fractions using a rotary evaporator connected to vacuum system, and dry the residual foam for 2 to 3 hr using a vacuum oil pump. 16. Grind the foam with a stainless steel spatula and dry the resulting powder 12 to 16 hr using a vacuum oil pump. 17. Characterize the product by TLC, 1 H NMR, and 13 C NMR. The compound is stable for at least 12 months at ambient temperature.
Synthesis of 2 -O-β β-dRibofuranosylnucleosides
N6-Benzoyl-9-[3,5-(1,1,3,3-tetraisopropyldisiloxane-1,3-diyl)-2-O-(2,3,5-tri-O-benzoylβ-D-ribofuranosyl)-β-D-ribofuranosyl]adenine (S.3). Yield of white amorphous solid 1.57 g (74%). TLC: Rf 0.31 (98:2 v/v CH2 Cl2 /MeOH).1 H NMR (400 MHz, CDCl3 ): 9.02 brs (1H, NH), 8.72 s (1H, H8), 8.15 s (1H, H2), 8.03-7.87 m (8H, Bz), 7.61-7.28 m (12H, Bz), 6.09 d (1H, J1 ,2 = 1.0 Hz, H1 , Ado), 5.96 dd (1H, J3 ,2 = 4.8 Hz, J3 ,4 = 6.2 Hz, H3 , Rib), 5.88 d (1H, H2 , Rib), 5.82 s (1H, H1 , Rib), 4.96 dd (1H, J3 ,2 = 4.7 Hz, J3 ,4 = 9.0 Hz, H3 , Ado), 4.90 dd (1H, H2 , Ado), 4.81-4.65 m (3H, H4 , Ado; H4 ,5 a, Rib), 4.18 d (1H, J5 a,5 b = −13.4 Hz, H5 a, Ado), 4.13 dd (1H, J5 b,4 = 1.0 Hz, J5 b,5 a = −9.5 Hz, H5 b, Rib), 4.03 dd (1H, J5 b,4 = 1.5 Hz, H5 b, Ado), 1.08-1.03 m (28H, iPr). 13 C NMR (CDCl3 ): 166.01, 165.37, 164.97 and 164.43 (C=O), 152.70 (C2), 150.76 (C6), 149.35 (C4), 141.80 (C8), 133.41, 133.16, 132.69, 129.71, 129.64, 129.10, 128.83, 128.34 and 127.78 (Bz), 123.40 (C5), 105.65 (C1 , Rib), 88.81 (C1 , Ado), 81.34 (C4 , Ado), 79.55 (C4 , Rib), 78.47 (C2 , Ado), 75.51 (C2 , Rib), 72.53 (C3 , Rib), 69.84 (C3 , Ado), 65.27 (C5 , Rib), 59.70 (C5 , Ado), 17.26, 17.04, 16.87, 16.77, 13.31, 12.89, 12.74 and 12.59 (iPr).
1.14.4 Supplement 27
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Prepare S.4 18. Weigh 1.06 g (1 mmol) S.3 into a 50-mL round-bottom flask, add 5 mL of 0.5 M TBAF in THF, stopper the flask, and keep the solution for 20 to 30 min at room temperature. Monitor deprotection by TLC using 5% (v/v) MeOH in CH2 Cl2 . The starting compound S.3 (Rf = 0.93) usually disappears after 20 to 30 min at 20◦ C. The product S.4 (Rf = 0.22) moves slower in the same solvent.
19. When the reaction is complete, evaporate all volatile material to dryness in vacuo using a rotary evaporator. Add 10 mL chloroform to the residue and evaporate to dryness. 20. Dissolve the residue in a minimal amount (3 to 4 mL) of chloroform and apply on a 3 × 15–cm column containing 30 g silica gel. Wash column with 300 mL CH2 Cl2 and 200 mL of 1% (v/v) MeOH in CH2 Cl2 . Elute with 2% (v/v) MeOH in CH2 Cl2 and collect 25-mL fractions. 21. Evaluate fractions by TLC using 5% (v/v) MeOH in CH2 Cl2 and combine the fractions that contain only S.4. Evaporate the combined fractions using a rotary evaporator connected to a vacuum system, and dry the residual foam for 2 to 3 hr using a vacuum oil pump. 22. Grind the foam with a stainless steel spatula and dry the resulting powder 12 to 16 hr using a vacuum oil pump. 23. Characterize the compound by TLC, 1 H NMR, and 13 C NMR. The compound is stable for at least 12 months at 0◦ C. N6 -Benzoyl-9-[2-O-(2,3,5-tri-O-benzoyl-β - D-ribofuranosyl)-β-D-ribofuranosyl]adenine (S.4). Yield of white amorphous solid 0.74 g (91%). TLC: Rf 0.22 (95:5 v/v CH2 Cl2 /MeOH). 1 H NMR (400 MHz, CDCl3 ): 9.52 brs (1H, NH), 8.78 s (1H, H8), 8.18 s (1H, H2), 8.06-7.88 m (8H, Bz), 7.53-7.34 m (12H, Bz), 6.05 d (1H, J1 ,2 = 7.2 Hz, H1 , Ado), 5.72 dd (1H, J3 ,2 = 5.3 Hz, J3 ,4 = 5.0 Hz, H3 , Rib), 5.64 dd (1H, J2 ,1 = 2.2 Hz, H2 , Rib), 5.21 d (1H, H1 , Rib), 5.20 dd (1H, J2 ,3 = 4.7 Hz, H2 , Ado), 4.59 brd (1H, H3 , Ado), 4.54 dd (1H, J5 a,4 = 4.1 Hz, J5 a,5 b = −11.8 Hz, H5 a, Rib), 4.48 ddd (1H, J4 ,5 b = 4.2 Hz, H4 , Rib), 4.30 brs (1H, H4 , Ado), 4.11 dd (1H, H5 b, Rib), 3.96 brd (1H, J5 a,5 b = −12.7 Hz, H5 a, Ado), 3.75 brd (1H, H5 b, Ado). 13 C NMR (CDCl3 ): 165.71, 165.21 and 164.66 (C=O), 151.96 (C2), 150.26 (C6), 150.11 (C4), 143.85 (C8), 133.47, 133.36, 133.16, 132.54, 129.51, 129.41, 128.94, 128.54 and 127.82 (Bz), 123.80 (C5), 106.19 (C1 , Rib), 88.86 (C1 , Ado), 86.99 (C4 , Ado), 80.48 (C4 , Rib), 79.53 (C2 , Ado), 75.61 (C2 , Rib), 72.18 (C3 , Rib), 71.06 (C3 , Ado), 64.12 (C5 , Rib), 62.73 (C5 , Ado).
Prepare S.5 24. Dissolve 408 mg (0.5 mmol) S.4 in 15 mL of 5 M ammonia in methanol (halfsaturated at 0◦ C) in a 50-mL round-bottom flask, stopper the flask, and keep the solution for 2 to 3 days at room temperature. 25. Evaporate all volatile material under reduced pressure using a rotary evaporator. 26. Partition the residue between 10 mL CH2 Cl2 and 20 mL water using a 100-mL separatory funnel. Separate the aqueous layer and wash it two additional times with 10 mL CH2 Cl2 . 27. Concentrate the aqueous layer to a volume of ∼1 mL, add 7 mL MeOH, and keep the mixture 16 hr at 0◦ C. 28. Collect the precipitate by vacuum filtration on a glass filter (porosity 3), wash with 2 to 3 mL MeOH followed by 5 mL diethyl ether, and dry in a vacuum desiccator with phosphorus pentoxide for 24 hr at room temperature.
Synthesis of Modified Nucleosides
1.14.5 Current Protocols in Nucleic Acid Chemistry
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29. Characterize the compound by TLC, UV spectroscopy, 1 H NMR, and 13 C NMR. The compound is stable for at least 12 months at 0◦ C. 9-(2-O-β-D-Ribofuranosyl-β-D-ribofuranosyl)adenine (S.5): Yield of white crystals 190 mg (91%). According to elemental analysis, S.5 is obtained as a monohydrate. m.p. 212◦ -214◦ C (softening at 162◦ -164◦ C). TLC: Rf 0.12 (8:2 v/v CH2 Cl2 /MeOH). [α]20 D −97◦ (c 0.76, DMSO). UV (pH 7-13): λmax 261 nm (ε 14200); (pH 1): λmax 261 nm (ε 13700). LSIMS (C15 H21 N5 O8 + H): calcd. 400.1468, found 400.1465. 1 H NMR (400 MHz, D2 O): 8.32 s (1H, H8, Ade), 8.19 s (1H, H2, Ade), 6.12 d (1H, J1 ,2 = 6.4 Hz, H1 , Ado), 5.07 s (1H, H1 , Rib), 4.80 dd (1H, J2 ,3 = 5.0 Hz, H2 , Ado), 4.55 dd (1H, J3 ,4 = 3.3 Hz, H3 , Ado), 4.29 ddd (1H, J4 ,5 a = 2.6 Hz, J4 ,5 b = 3.6 Hz, H4 Ado), 4.13 d (1H, J2 ,3 = 4.5 Hz, H2 , Rib), 3.99 dd (1H, J3 ,4 = 7.4 Hz, H3 , Rib), 3.92 dd (1H, J5 a,5 b = −13.0 Hz, H5 a, Ado), 3.83 dd (1H, H5 b, Ado), 3.82 ddd (1H, J4 ,5 a = 3.7 Hz, J4 ,5 b = 6.8 Hz, H4 Rib), 3.32 dd (1H, J5 a,5 b = −12.0 Hz, H5 a, Rib), 2.75 dd (1H, H5 b, Rib). 13 C NMR (D2 O): 156.14 (C6), 153.06 (C2), 149.74 (C4), 141.08 (C8), 119.42 (C5), 106.42 (C1 , Rib), 87.44 (C1 , Ado), 86.84 (C4 , Ado), 83.11 (C4 , Rib), 78.64 (C2 , Ado), 74.75 (C2 , Rib), 71.31 (C3 , Rib), 69.43 (C3 , Ado), 63.16 (C5 , Rib), 61.88 (C5 , Ado). ALTERNATE PROTOCOL 1
PREPARATION OF 2 -O-β -D-RIBOFURANOSYLURIDINE 3 ,5 -(1,1,3,3-O-Tetraisopropyldisiloxane-1,3-diyl)uridine (S.6a; Markiewicz and Wiewiorowski, 1986; UNIT 2.10; Fig. 2.10.2) is converted to 2 -O-β-D-ribofuranosyluridine (S.9a) with an overall yield of 56% (Mikhailov et al., 1997a) using the steps outlined in the Basic Protocol (Fig. 1.14.3). The reaction uses the same molar equivalent of starting nucleoside (i.e., 2 mmol S.6a) and the same amounts of other reagents as in the Basic Protocol. The condensation reaction of pyrimidine nucleoside S.6a with 1O-acetyl-2,3,5-tri-O-benzoyl-β-D-ribofuranose (S.2) is carried out in 1,2-dichloroethane for 2 hr at 0◦ C. 2 -O-β-D-Ribofuranosyluridine (S.9a) is isolated by crystallization from a minimal amount of water. 1-[3,5-(1,1,3,3-Tetraisopropyldisiloxane-1,3-diyl)-2-O-(2,3,5-tri-O-benzoyl-β-Dribofuranosyl)-β-D-ribofuranosyl]uracil (S.7a). Yield of white amorphous solid 76%. TLC: Rf 0.30 (98:2 v/v CH2 Cl2 /MeOH). 1 H NMR (400 MHz, CDCl3 ): 8.35 brs (1H, NH), 8.05-7.85 m (6H, Bz), 7.80 d (1H, J6,5 = 8.0 Hz, H6), 7.56-7.26 m (9H, Bz), 5.88 dd (1H, J3 ,2 = 5.0 Hz, J3 ,4 = 6.5 Hz, H3 , Rib), 5.82 d (1H, H2 , Rib), 5.81 s (1H, H1 , Urd), 5.80 s (1H, H1 , Rib), 5.65 d (1H, H5), 4.80-4.75 m (3H, H3 ,4 , Urd; H4 , Rib), 4.41 d (1H, J2 ,3 = 4.8 Hz, H2 , Urd), 4.30 dd (1H, J5 a,4 = 4.0 Hz, J5 a,5 b = −9.6 Hz, H5 a, Rib), 4.23 d (1H, J5 a,5 b = −13.4 Hz, H5 a, Urd), 4.08 dd (1H, J5 b,4 = 1.0 Hz, H5 b, Rib), 3.95 dd (1H, J5 ,4 = 1.5 Hz, H5 b, Urd), 1.09-0.96 m (28H, iPr). 13 C NMR (400 MHz, CDCl3 ): 166.01, 165.22 and 164.95 (C=O), 163.51 (C4), 149.68 (C2), 139.41 (C6), 133.31, 133.21, 133.00, 132.83, 129.64, 129.10, 128.87, 128.32 and 128.21 (Bz), 105.44 (C1 , Rib), 101.45 (C5), 89.25 (C1 , Urd), 81.52 (C4 , Urd), 79.33 (C4 , Rib), 78.47 (C2 , Urd), 75.47 (C2 , Rib), 72.96 (C3 , Rib), 68.71 (C3 , Urd), 65.60 (C5 , Rib), 59.22 (C5 , Urd), 17.22, 17.12, 17.01, 16.77, 16.67, 13.26, 12.91, 12.75 and 12.42 (iPr).
Synthesis of 2 -O-β β-dRibofuranosylnucleosides
1-[2-O-(2,3,5-Tri-O-benzoyl-β-D-ribofuranosyl)-β-D-ribofuranosyl]uracil (S.8a). Yield of white amorphous solid 92%. TLC: Rf 0.20 (95:5 v/v CH2 Cl2 /MeOH). 1 H NMR (400 MHz, CDCl3 ): 8.47 brs (1H, NH), 8.08-7.91 m (6H, Bz), 7.54-7.36 m (10H, H6, Bz), 5.82 dd (1H, J3 ,2 = 5.3 Hz, J3 ,4 = 5.9 Hz, H3 , Rib), 5.74 dd (1H, J2 ,1 = 1.9 Hz, H2 , Rib), 5.70 d (1H, J1 ,2 = 4.7 Hz, H1 , Urd), 5.65 d (1H, J5,6 = 8.0 Hz, H5), 5.48 d (1H, H1 , Rib), 4.78-4.69 m (3H, H2 , Urd; H4 ,5 a, Rib), 4.49-4.42 m (2H, H3 , Urd; H5 b, Rib), 3.93-3.87 m (2H, H4 ,5 a, Urd), 3.75 dd (1H, J5 b,4 = 2.3 Hz, J5 b,5 a = −12.4 Hz, H5 b, Urd). 13 C NMR (400 MHz, CDCl3 ): 166.12 and 165.46 (C=O), 163.78 (C4), 150.41 (C2), 142.54 (C6), 133.51, 133.47, 133.28, 129.69, 129.34, 128.68 and 128.42 (Bz), 106.82 (C1 , Rib), 102.08 (C5), 91.10 (C1 , Urd), 84.51 (C4 , Urd), 80.65 (C4 , Rib), 79.75 (C2 , Urd), 75.77 (C2 , Rib), 72.40 (C3 , Rib), 69.08 (C3 , Urd), 64.65 (C5 , Rib), 61.14 (C5 , Urd).
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Figure 1.14.3 Synthesis of 2 -O-β-D-ribofuranosyluridine (S.9a), 2 -O-β-D-ribofuranosylthymidine (S.9b), 2 -O-β-D-ribofuranosylcytidine (S.9c), and 2 -O-β-D-ribofuranosylguanosine (S.9d). The expected overall yields are given in parentheses. DCE, 1,2-dichloroethane; TBAF, tetrabutylammonium fluoride; THF, tetrahydrofuran.
Synthesis of Modified Nucleosides
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Supplement 27
1-(2-O-β-D-Ribofuranosyl-β-D-ribofuranosyl)uracil (S.9a): Yield of white crystals 80%. m.p. 224◦ -225◦ C (water). TLC: Rf 0.15 (8:2 v/v CH2 Cl2 /MeOH). [α]20 D −36◦ (c 0.68, DMSO). UV (pH 1-7): λmax 262 nm (ε 9400); (pH 13): λmax 262 nm (ε 6900). LSIMS (C14 H20 N2 O10 + H): calcd. 377.1196, found 377.2801. 1 H NMR (400 MHz, D2 O): 7.87 d (1H, J5,6 = 8.1 Hz, H6, Ura), 6.04 d (1H, J1 ,2 = 5.0 Hz, H1 , Urd), 5,92 d (1H, H5, Ura), 5.15 s (1H, H1 , Rib), 4.44 dd (1H, J2 ,3 = 5.4 Hz, H2 , Urd), 4.36 dd (1H, J3 ,4 = 5.1 Hz, H3 , Urd), 4.19 dd (1H, J3 ,2 = 4.8 Hz, J3 ,4 = 6.9 Hz, H3 , Rib), 4.16 d (1H, H2 , Rib), 4.11 ddd (1H, J4 ,5 a = 2.9 Hz, J4 ,5 b = 4.5 Hz, H4 , Urd), 3.99 ddd (1H, J4 ,5 a = 3.4 Hz, J4 ,5 b = 6.6 Hz, H4 , Rib), 3.88 dd (1H, J5 a,5 b = −12.7 Hz, H5 a, Urd), 3.79 dd (1H, H5 b, Urd), 3.75 dd (1H, J5 a,5 b = −12.1 Hz, H5 a, Rib), 3.48 dd (1H, H5 b, Rib). 13 C NMR (D2 O): 166.66 (C4), 152.17 (C2), 142.55 (C6), 107.28 (C1 , Rib), 103.06 (C5), 88.47 (C1 , Urd), 85.11 (C4 , Urd), 83.35 (C4 , Rib), 78.93 (C2 , Urd), 74.85 (C2 , Rib), 71.14 (C3 , Rib), 68.87 (C3 , Urd), 63.34 (C5 , Rib), 61.17 (C5 , Urd). ALTERNATE PROTOCOL 2
PREPARATION OF 2 -O-β -D-RIBOFURANOSYLTHYMIDINE 3 ,5 -(1,1,3,3-O-Tetraisopropyldisiloxane-1,3-diyl)thymidine (S.6b; Markiewicz and Wiewiorowski, 1986; see Support Protocol) is converted to 2 -O-β-D-ribofuranosylthymidine (S.9b) with an overall yield of 48% (Mikhailov et al., 1997a) using the steps outlined in the Basic Protocol (Fig. 1.14.3). The reaction uses the same molar equivalent of starting nucleoside (i.e., 2 mmol S.6b) and the same amounts of other reagents as in the Basic Protocol. The condensation reaction of pyrimidine nucleoside S.6b with 1-O-acetyl-2,3,5-tri-O-benzoyl-β-D-ribofuranose (S.2) is carried out in 1,2dichloroethane for 2 hr at 0◦ C. 2 -O-β-D-Ribofuranosylthymidine (S.9b) is isolated by crystallization from a minimal amount of 9:1 (v/v) ethanol/water. 1-[3,5-(1,1,3,3-Tetraisopropyldisiloxane-1,3-diyl)-2-O-(2,3,5-tri-O-benzoyl-β-Dribofuranosyl)-β-D-ribofuranosyl]thymine (S.7b). Yield of white amorphous solid 74%. TLC: Rf 0.30 (98:2 v/v CH2 Cl2 /MeOH). 1 H NMR (400 MHz, CDCl3 ): 8.21 brs (1H, NH), 8.00-7.83 m (6H, Bz), 7.55-7.25 m (10H, Bz, H6), 5.85 dd (1H, J3 ,2 = 5.0 Hz, J3 ,4 = 6.5 Hz, H3 , Rib), 5.78 d (1H, H2 , Rib), 5.76 s (1H, H1 , Thd), 5.75 s (1H, H1 , Rib), 4.77-4.69 m (3H, H3 ,4 , Thd; H4 , Rib), 4.39 d (1H, J2 ,3 = 4.3 Hz, H2 , Thd), 4.30 dd (1H, J5 a,4 = 4.4 Hz, J5 a,5 b = −9.5 Hz, H5 a, Rib), 4.18 d (1H, J5 a,5 b = −13.4 Hz, H5 a, Thd), 4.04 dd (1H, J5 b,4 = 2.0 Hz, H5 b, Rib), 3.92 dd (1H, J5 b,4 = 2.6 Hz, H5 b, Thd), 1.86 d (3H, J5,6 = 1.2 Hz, Me5), 1.07-0.92 m (28H, iPr). 13 C NMR (CDCl3 ): 166.11 and 165.32 (C=O), 165.01 (C4), 149.55 (C2), 135.27 (C6), 133.39, 133.30, 132.96, 129.72, 129.18, 128.94, 128.41, 128.29 and 128.25 (Bz), 110.09 (C5), 105.51 (C1 , Rib), 89.64 (C1 , Thd), 81.48 (C4 , Thd), 79.38 (C4 , Rib), 78.52 (C2 , Thd), 75.53 (C2 , Rib), 73.05 (C3 , Rib), 69.02 (C3 , Thd), 65.77 (C5 , Rib), 59.25 (C5 , Thd), 17.41, 17.33, 17.22, 17.13, 17.01, 16.88, 16.77, 13.42, 12.86 and 12.68 (iPr), 12.57 (Me5). 1-[2-O-(2,3,5-Tri-O-benzoyl-β-D-ribofuranosyl)-β-D-ribofuranosyl]thymine (S.8b). Yield of white amorphous solid 87%. TLC: Rf 0.20 (95:5 v/v CH2 Cl2 /MeOH). 1 H NMR (400 MHz, CDCl3 -CD3 OD): 7.98-7.81 m (6H, Bz), 7.42 q (1H, J6,5 = 1.2 Hz, H6), 7.497.25 m (9H, Bz), 5.76 d (1H, J1 ,2 = 4.2 Hz, H1 , Thd), 5.74 dd (1H, J3 ,2 = 5.1 Hz, J3 ,4 = 6.2 Hz, H3 , Rib), 5.69 dd (1H, J2 ,1 = 1.2 Hz, H2 , Rib), 5.44 d (1H, H1 , Rib), 4.53 ddd (1H, J4 ,5 a = 4.8 Hz, J4 ,5 b = 5.9 Hz, H4 , Rib), 4.41 dd (1H, J5 a,5 b = −11.8 Hz, H5 a, Rib), 4.30 dd (1H, H5 b, Rib), 4.28 dd (1H, J2 ,3 = 5.3 Hz, H2 , Thd), 4.13 dd (1H, J3 ,4 = 5.6 Hz, H3 , Thd), 3.77 ddd (1H, J4 ,5 a = 2.3 Hz, J4 ,5 b = 2.5 Hz, H4 , Thd), 3.66 dd (1H, J5 a,5 b = −12.4 Hz, H5 a, Thd), 3.52 dd (1H, H5 b, Thd), 1.76 d (3H, Me5). 13 C NMR (CDCl3 -CD3 OD): 166.15 and 165.48 (C=O), 164.33 (C4), 150.55 (C2), 137.74 (C6), 133.62, 133.47, 133.21, 129.62, 129.54, 129.18, 128.55, 128.48, 128.38, 128.31 and 128.28 (Bz), 110.48 (C5), 106.31 (C1 , Rib), 89.92 (C1 , Thd), 84.36 (C4 , Thd), 79.31 (C4 , Rib), 77.32 (C2 , Thd), 75.57 (C2 , Rib), 72.36 (C3 , Rib), 68.89 (C3 , Thd), 64.69 (C5 , Rib), 60.79 (C5 , Thd), 11.96 (Me5).
Synthesis of 2 -O-β β-dRibofuranosylnucleosides
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Current Protocols in Nucleic Acid Chemistry
1-(2-O-β-D-Ribofuranosyl-β-D-ribofuranosyl)thymine (S.9b). Yield of white crystals 75%. m.p. 236◦ -237◦ C (aq. EtOH). TLC: Rf 0.16 (8:2 v/v CH2 Cl2 /MeOH). [α]20 D −52◦ (c 0.82, water). UV (pH 1-7): λmax 269 nm (ε9400); (pH 13): λmax 262 nm (ε7100). LSIMS (C15 H22 N2 O10 + H): calcd. 391.1352, found 391.1362. 1 H NMR (400 MHz, D2 O): 7.66 q (1H, J6,CH3 = 1.2 Hz, H6, Thy), 6.03 d (1H, J1 ,2 = 5.5 Hz, H1 , Thd), 5.12 s (1H, H1 , Rib), 4.44 dd (1H, J2 ,3 = 5.5 Hz, H2 , Thd), 4.38 dd (1H, J3 ,4 = 4.7 Hz, H3 , Thd), 4.14 dd (1H, J2 ,3 = 4.7 Hz, J3 ,4 = 6.7 Hz, H3 , Rib), 4.13 dd (1H, H2 , Rib), 4.11 ddd (1H, J4 ,5 a = 3.1 Hz, J4 ,5 b = 4.5 Hz, H4 , Urd), 3.99 ddd (1H, J4 ,5 a = 3.6 Hz, J4 ,5 b = 6.7 Hz, H4 , Rib), 3.88 dd (1H, J5 a,5 b = −12.7 Hz, H5 a, Thd), 3.81 dd (1H, H5 b, Urd), 3.71 dd (1H, J5 a,5 b = −12.1 Hz, H5 a, Rib), 3.44 dd (1H, H5 b, Rib), 1.91 d (1H, Me5). 13 C NMR (D2 O): 167.28 (C4), 152.71 (C2), 138.49 (C6), 112.82 (C5), 107.71 (C1 , Rib), 88.50 (C1 , Thd), 85.60 (C4 , Thd), 83.88 (C4 , Rib), 79.17 (C2 , Thd), 75.32 (C2 , Rib), 71.74 (C3 , Rib), 69.32 (C3 , Thd), 64.03 (C5 , Rib), 61.71 (C5 , Thd), 12.47 (Me5).
PREPARATION OF 2 -O-β -D-RIBOFURANOSYLCYTIDINE N4-Benzoyl-3,5 -(1,1,3,3-O-tetraisopropyldisiloxane-1,3-diyl)cytidine (S.6c; Markiewicz and Wiewiorowski, 1986; UNIT 2.4; Fig. 2.4.4) is converted to 2 -O-β-D-ribofuranosylcytidine (S.9c) with an overall yield of 72% (Mikhailov et al., 1997a) using the steps outlined in the Basic Protocol (Fig. 1.14.3). The reaction uses the same molar equivalent of starting nucleoside (i.e., 2 mmol S.6c) and the same amounts of other reagents as in the Basic Protocol. The condensation reaction of pyrimidine nucleoside S.6c with 1-O-acetyl-2,3,5-tri-O-benzoyl-β-D-ribofuranose (S.2) is carried out in 1,2dichloroethane for 2 hr at 0◦ C. After step 26, the aqueous layer containing product S.9c is evaporated to dryness in vacuo, the residue is dissolved in 5 mL methanol, and is evaporated to dryness again. The residual foam is dried for 2 to 3 hr using a vacuum oil pump, and is finally dried in a vacuum desiccator with phosphorus pentoxide.
ALTERNATE PROTOCOL 3
N4-Benzoyl-1-[3,5-(1,1,3,3-tetraisopropyldisiloxane-1,3-diyl)-2-O-(2,3,5-tri-O-benzoylβ-D-ribofuranosyl)-β-D-ribofuranosyl]cytosine (S.7c). Yield of white amorphous solid 80%. TLC: Rf 0.32 (98:2 v/v CH2 Cl2 /MeOH). 1 H NMR (400 MHz, CDCl3 ): 8.77 brs (1H, NH), 8.30 d (1H, J6,5 = 7.6 Hz, H6), 8.02-7.82 m (8H, Bz), 7.55-7.24 m (13H, H5, Bz), 5.96 s (1H, H1 , Cyd), 5.92 dd (1H, J3 ,2 = 5.0 Hz, J3 ,4 = 6.3 Hz, H3 , Rib), 5.89 s (1H, H1 , Rib), 5.81 d (1H, H2 , Rib), 4.83-4.78 m (3H, H3 ,4 , Cyd; H4 , Rib), 4.49 d (1H, J2 ,3 = 4.1 Hz, H2 , Cyd), 4.28 dd (1H, J5 a,4 = 3.8 Hz, J5 a,5 b = −9.4 Hz, H5 a, Rib), 4.24 d (1H, J5 a,5 b = −13.4 Hz, H5 a, Cyd), 4.17 dd (1H, J5 b,4 = 1.0 Hz, H5 b, Rib), 3.98 dd (1H, J5 ,4 = 1.5 Hz, H5 b, Cyd), 1.10-0.96 m (28H, iPr). 13 C NMR (CDCl3 ): 166.09, 165.28 and 165.05 (C=O) 162.40 (C4), 155.02 (C2), 144.36 (C6), 133.31, 133.18, 132.80, 129.74, 129.23, 128.98, 128.37, 128.21 and 127.53 (Bz), 105.48 (C1 , Rib), 96.02 (C5), 90.14 (C1 , Cyd), 81.81 (C4 , Cyd), 79.07 (C4 , Rib), 78.51 (C2 , Cyd), 75.63 (C2 , Rib), 73.10 (C3 , Rib), 68.71 (C3 , Cyd), 65.51 and 59.32 (C5 ), 17.41, 17.28, 17.06, 16.91, 16.77, 13.31, 13.02, 12.85 and 12.50 (iPr). N4 -Benzoyl-1-[2-O-(2,3,5-tri-O-benzoyl-β-D-ribofuranosyl)-β-D-ribofuranosyl]cytosine (S.8c). Yield of white amorphous solid 95%. TLC: Rf 0.25 (95:5 v/v CH2 Cl2 /MeOH). 1 H NMR (400 MHz, CDCl3 ): 9.18 brs (1H, NH), 8.05-7.89 m (9H, H6, Bz), 7.58-7.33 m (13H, H5, Bz), 5.84 dd (1H, J3 ,2 = 5.2 Hz, J3 ,4 = 5.3 Hz, H3 , Rib), 5.77 d (1H, J1 ,2 = 3.8 Hz, H1 , Cyd), 5.74 dd (1H, J2 ,1 = 1.6 Hz, H2 , Rib), 5.60 d (1H, H1 , Rib), 4.92 dd (1H, J2 ,3 = 5.0 Hz, H2 , Cyd), 4.76 dd (1H, J5 a,4 = 4.0 Hz, J5 a,5 b = −11.9 Hz, H5 a, Rib), 4.69 ddd (1H, J4 ,3 = 5.3 Hz, J4 ,5 a = 4.3 Hz, H4 , Rib), 4.49-4.45 m (2H, H3 , Cyd; H5 b, Rib), 4.03 brd (1H, J4 ,3 = 5.3 Hz, H4 , Cyd), 3.94 brd (1H, J5 a,5 b = −12.8 Hz, H5 a, Cyd), 3.77 brd (1H, H5 b, Cyd). 13 C NMR (CDCl3 ): 166.66, 166.05 and 165.37 (C=O), 162.77 (C4), 155.31 (C2), 147.17 (C6), 133.46, 133.35, 133.14, 132.95, 129.67, 129.42, 128.76, 128.35 and 127.71 (Bz), 106.66 (C1 , Rib), 96.87 (C5), 92.38 (C1 , Cyd), 84.87 (C4 , Cyd), 80.67 (C4 , Rib), 79.51 (C2 , Cyd), 75.87 (C2 , Rib), 72.57 (C3 , Rib), 68.34 (C3 , Cyd), 64.82 (C5 , Rib), 60.56 (C5 , Cyd). Synthesis of Modified Nucleosides
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1-(2-O-β-D-Ribofuranosyl-β-D-ribofuranosyl)cytosine (S.9c): Yield of white amorphous solid 95%. TLC: Rf 0.04 (8:2 v/v CH2 Cl2 /MeOH). [α]20 D −29◦ (c 0.53, water). UV (pH 7-13): λmax 271 nm (ε8200); (pH 1): λmax 279 nm (ε 12200). LSIMS (C14 H21 N3 O9 + H): calcd. 376.1355, found 376.1338. 1 H NMR (400 MHz, D2 O): 7.81 d (1H, J5,6 = 7.6 Hz, H6, Cyt), 6.06 d (1H, H5, Cyt), 6.04 d (1H, J1 ,2 = 5.1 Hz, H1 , Cyd), 5.12 s (1H, H1 , Rib), 4.37 dd (1H, J2 ,3 = 5.5 Hz, H2 , Cyd), 4.33 dd (1H, J3 ,4 = 4.7 Hz, H3 , Cyd), 4.17 dd (1H, J3 ,2 = 4.6, J3 ,4 = 6.8, H3 , Rib), 4.16 d (1H, H2 , Rib), 4.11 ddd (1H, J4 ,5 a = 3.0 Hz, J4 ,5 b = 4.4 Hz, H4 , Cyd), 3.97 ddd (1H, J4 ,5 a = 3.4 Hz, J4 ,5 b = 6.7 Hz, H4 , Rib), 3.88 (1H, J5 a,5 b = −12.7 Hz, H5 a, Cyd), 3.79 dd (1H, H5 b, Cyd), 3.71 dd (1H, J5 a,5 b = −12.2 Hz, H5 a, Rib), 3.37 dd (1H, H5 b, Rib). 13 C NMR (D2 O): 166.57 (C4), 158.03 (C2), 142.28 (C6), 106.96 (C1 , Rib), 97.17 (C5), 88.70 (C1 , Cyd), 85.08 (C4 , Cyd), 83.30 (C4 , Rib), 78.92 (C2 , Cyd), 74.84 (C2 , Rib), 71.23 (C3 , Rib), 68.90 (C3 , Cyd), 63.51 (C5 , Rib), 61.35 (C5 , Cyd). ALTERNATE PROTOCOL 4
PREPARATION OF 2 -O-β -D-RIBOFURANOSYLGUANOSINE N2 -Isobutyryl-3 ,5 -(1,1,3,3-O-tetraisopropyldisiloxane-1,3-diyl)guanosine (S.6d; Markiewicz and Wiewiorowski, 1986; UNIT 2.4; Fig. 2.4.6) is converted to 2 -O-β-Dribofuranosylguanosine (S.9d) with an overall yield of 46% (Mikhailov et al., 1997a) using the steps outlined in the Basic Protocol (Fig. 1.14.3) with some modifications described here. The reaction uses the same molar equivalent of starting nucleoside (i.e., 2 mmol S.6d) and the same amounts of other reagents as in the Basic Protocol. The condensation reaction of guanosine nucleoside S.6d with 1-O-acetyl-2,3,5-tri-O-benzoyl-βD-ribofuranose (S.2) is carried out in 1,2-dichloroethane for 16 hr at 0◦ C. The guanosine derivatives form stable complexes with tin tetrachloride. After the condensation reaction (step 6), the reaction mixture is taken up with a long capillary (Fig. 1.14.2B) and is added to a microcentrifuge tube containing 0.2 mL saturated sodium bicarbonate solution and 0.2 mL ethyl acetate. The tube is shaken and kept for 5 to 10 min at 20◦ C to destroy the tin tetrachloride complexes. A sample from the upper organic layer is then checked by TLC using 2% (v/v) MeOH in CH2 Cl2 (step 7). The starting compound S.6d (Rf = 0.14) usually disappears after 16 hr at 0◦ C, and the product S.7d (Rf = 0.25) moves faster in the same solvent. After TLC, the purification of S.7d resumes with step 8. After step 26, the aqueous layer containing 2 -O-β-D-ribofuranosylguanosine (S.9d) is evaporated to dryness in vacuo. The residue is dissolved in 10 mL ethanol and the mixture is kept for 16 hr at 0◦ C. The resulting precipitate is collected by vacuum filtration on a glass filter (porosity 3), washed with 5 ml diethyl ether, and dried for 24 hr in a vacuum desiccator with phosphorus pentoxide. N2 - Isobutyryl-9-[3,5-(1,1,3,3-tetraisopropyldisiloxane-1,3-diyl)-2-O-(2,3,5-triO-benzoyl-β-D-ribofuranosyl)-β-D-ribofuranosyl]guanine (S.7d). Yield of white amorphous solid 72%. TLC: Rf 0.25 (98:2 v/v CH2 Cl2 /MeOH). 1 H NMR (400 MHz, CDCl3 ): 12.19 brs (1H, NH), 9.81 brs (1H, NH), 8.08 s (1H, H8), 8.00-7.82 m (6H, Bz), 7.55-7.26 m (9H, Bz), 6.10 dd (1H, J3 ,2 = 5.1 Hz, J3 ,4 = 5.8 Hz, H3 , Rib), 5.86 dd (1H, J2 ,1 = 0.9 Hz, H2 , Rib), 5.79 d (1H, H1 , Rib), 5.72 s (1H, H1 , Guo), 4.87-4.53 m (5H, H2 ,3 ,4 , Guo; H4 ,5 a, Rib), 4.23 d (1H, J5 a,5 b = −13.5 Hz, H5 a, Guo), 4.15 dd (1H, J5 b,4 = 2.2 Hz, J5 b,5 a = −9.4 Hz, H5 b, Rib), 3.96 dd (1H, J5 ,4 = 2.5 Hz, H5 b, Guo), 2.86 sep (1H, J = 6.8 Hz, CH, iBu), 1.32 d (3H, Me, iBu), 1.22 d (3H, Me, iBu), 1.07-0.92 m (28H, iPr). 13 C NMR (CDCl3 ): 179.42, 167.87, 165.34 and 165.00 (C=O), 155.42 (C6), 148.27 (C2), 147.09 (C4), 135.91 (C8), 134.01, 133.57, 129.71, 128.95, 128.83, 128.66 and 128.45 (Bz), 121.60 (C5), 105.46 (C1 , Rib), 87.32 (C1 , Guo), 81.17 (C4 , Guo), 79.38 (C4 , Rib), 78.75 (C2 , Guo), 75.81 (C2 , Rib), 73.29 (C3 , Rib), 69.31 (C3 , Guo), 65.65 (C5 , Rib), 59.41 (C5 , Guo), 36.10 (CH, iBu), 19.21 (Me, iBu), 18.92 (Me, iBu), 17.48, 17.31, 17.13, 16.96, 16.80, 16.71, 13.32, 13.06, 12.82 and 12.53 (iPr).
Synthesis of 2 -O-β β-dRibofuranosylnucleosides
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N2 -Isobutyryl-9-[2-O-(2,3,5-tri-O-benzoyl-β-D-ribofuranosyl)-β-D-ribofuranosyl]guanine (S.8d). Yield of white amorphous solid 84%. TLC: Rf 0.15 (95:5 v/v CH2 Cl2 /MeOH). 1 H NMR (400 MHz, CDCl3 ): 12.27 brs (1H, NH), 9.67 brs (1H, NH), 8.62 s (1H, H8), 7.99-7.88 m (6H, Bz), 7.55-7.32 m (9H, Bz), 6.07 dd (1H, J3 ,2 = 5.0 Hz, J3 ,4 = 5.9 Hz, H3 , Rib), 5.88 d (1H, J1 ,2 = 1.9 Hz, H1 , Guo), 5.85 d (1H, H2 , Rib), 5.80 s (1H, H1 , Rib), 4.76 dd (1H, J5 a,4 = 5.9 Hz, J5 a,5 b = −11.3 Hz, H5 a, Rib), 4.73-4.66 m (2H, H3 , Guo; H4 , Rib), 4.53 dd (1H, J5 b,4 = 3.7 Hz, H5 b, Rib), 4.40 dd (1H, J2 ,3 = 4.4 Hz, H2 , Guo), 4.17 brd (1H, J4 ,3 = 8.2 Hz, H4 , Guo), 4.06brs (2H, H5 a,5 b, Guo), 2.82 sep (1H, J = 6.8 Hz, CH, iBu), 1.31 d (3H, Me, iBu), 1.24 d (3H, Me, iBu). 13 C NMR (CDCl3 ): 179.73, 167.77 and 165.27 (C=O), 155.95 (C6), 148.03 (C2), 147.48 (C4), 138.98 (C8), 133.93, 133.48, 129.87, 129.73, 129.69, 128.99, 128.83, 128.64, 128.45 and 128.39 (Bz), 120.50 (C5), 105.48 (C1 , Rib), 87.97 (C1 , Guo), 83.67 (C4 , Guo), 79.25 (C4 , Rib), 77.32 (C2 , Guo), 75.82 (C2 , Rib), 73.07 (C3 , Rib), 68.79 (C3 , Guo), 65.57 (C5 , Rib), 59.83 (C5 , Guo), 36.05 (CH, iBu), 19.13 (Me, iBu), 18.85 (Me, iBu). 9-(2-O-β-D-Ribofuranosyl-β-D-ribofuranosyl)guanine (S.9d): Yield of white crystals 76%. m.p. 215◦ -216◦ C (EtOH). TLC: Rf 0.09 (8:2 v/v CH2 Cl2 /MeOH). [α]20 D −75◦ (c 0.92, water). UV (pH 1): λmax 257 nm (ε 10800); (pH 7): λmax 254 nm (ε 12000); (pH 13): λmax 263 nm (ε 10000). LSIMS (C15 H21 N5 O9 + H): calcd. 416.1417, found 416.1413. 1 H NMR (400 MHz, D2 O): 8.03 s (1H, H6, Guo), 6.01 d (1H, J1 ,2 = 6.3 Hz, H1 , Guo), 5,12 d (1H, J1 ,2 = 0.8 Hz, H1 , Rib), 4.81 dd (1H, J2 ,3 = 5.3 Hz, H2 , Guo), 4.56 dd (1H, J3 ,4 = 3.3 Hz, H3 , Guo), 4.26 ddd (1H, J4 ,5 a = 3.1 Hz, J4 ,5 b = 4.1 Hz, H4 , Guo), 4.17 dd (1H, J2 ,3 = 4.6 Hz, H2 , Rib), 4.08 dd (1H, J3 ,4 = 7.3 Hz, H3 , Rib), 3.92 ddd (1H, J4 ,5 a = 3.8 Hz, J4 ,5 b = 6.8 Hz, H4 , Rib), 3.91 dd (1H, J5 a,5 b = −12.8 Hz, H5 a, Guo), 3.85 dd (1H, H5 b, Guo), 3.47 dd (1H, J5 a,5 b = −11.9 Hz, H5 a, Rib), 3.01 dd (1H, H5 b, Rib). 13 C NMR (D2 O): 159.89 (C6), 154.86 (C2), 150.85 (C4), 139.28 (C8), 117.67 (C5), 107.18 (C1 , Rib), 87.52 (C1 , Guo), 87.03 (C4 , Guo), 83.80 (C4 , Rib), 78.98 (C2 , Guo), 75.37 (C2 , Rib), 72.06 (C3 , Rib), 69.96 (C3 , Guo), 64.02 (C5 , Rib), 62.41 (C5 , Guo).
PREPARATION OF THE 3 ,5 -PROTECTED RIBOTHYMIDINE 1-[3,5-(1,1,3,3-Tetraisopropyldisiloxane-1,3-diyl)-β-D-ribofuranosyl]thymine (S.6b) is prepared starting from thymine and 1-O-acetyl-2,3,5-tri-O-benzoyl-β-D-ribofuranose (Fig. 1.14.4). A procedure proposed by Vorbr¨uggen and Ruh-Pohlenz (2000) has completely replaced the previously employed methods for the synthesis of nucleosides and their analogs. The method involves glycosylation of trimethylsilyl derivatives of heterocyclic bases with peracylated sugars in aprotic solvents in the presence of Lewis acids such as SnCl4 or trimethylsilyl trifluoromethanesulfonate (TMSOTf; UNIT 1.13). The use of this approach has significantly simplified the reaction procedure and increased the yields of the target products. The presence of a 2-O-acyl group in a carbohydrate residue is believed to be crucial for the stereochemistry of this reaction, since it stabilizes C1 carbonium ion generated via an intermediate 1,2-acyloxonim ion and yields 1,2-trans derivatives. In the first step of this procedure, crystalline 1-(2,3,5-tri-O-benzoyl-β-D-ribofuranosyl)thymine (S.11) is prepared from thymine derivative S.10 and 1-O-acetyl-2,3,5-tri-Obenzoyl-β-D-ribofuranose (S.2) in a yield of 82%. In the second step, the cleavage of the benzoyl protecting groups is achieved using sodium methoxide in methanol under mild conditions, and 1-(β-D-ribofuranosyl)thymine (S.12) is purified by crystallization from ethanol. In the third step, nucleoside S.12 is simultaneously protected on its 3 - and 5 -hydroxy groups by the bifunctional protecting reagent 1,3-dichloro1,1,3,3-tetraisopropyldisiloxane developed by Markiewicz and Wiewiorowski (1986). The derivative S.6b is formed in high yield (86% from S.12) due to the higher reactivity of the primary 5 -hydroxy group and the subsequently favorable cyclization to the 3 hydroxy group. Chromatography is required only in the last step of the sequence. Using this three-step procedure, S.6b is prepared in an overall yield of 55%.
SUPPORT PROTOCOL
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Figure 1.14.4 Synthesis of 1-[3,5-(1,1,3,3-tetraisopropyldisiloxane-1,3-diyl)-β-D-ribofuranosyl]thymine (S.6b) The expected overall yield is given in parentheses. DCE, 1,2dichloroethane; TMSOTf, trimethylsilyl trifluoromethanesulfonate; TPDSCl2 , 1,3-dichloro-1,1,3,3tetraisopropyldisiloxane.
Additional Materials (also see Basic Protocol) Thymine Ammonium sulfate [(NH4 )2 SO4 ] 1,1,1,3,3,3-Hexamethyldisilazane, reagent grade Calcium chloride (CaCl2 ), anhydrous Trimethylsilyl trifluoromethanesulfonate (TMSOTf) 0.2 N sodium methoxide (NaOMe), freshly prepared from sodium and dry methanol Dowex 50 × 4 (100 to 200 mesh) in H+ form Pyridine, anhydrous Markiewicz reagent: 1,3-dichloro-1,1,3,3-tetraisopropyldisiloxane, 96% pure (Wacker) Toluene, reagent grade 100-, 250-, and 500-mL round-bottom flasks Reflux condensers CaCl2 protection tubes Oil bath with temperature control Adapters with stopcocks and vacuum pump (Fig. 1.14.5) 250- and 500-mL separatory funnels Glass filters (porosity 3) 3 × 20–cm chromatography columns Synthesis of 2 -O-β β-dRibofuranosylnucleosides
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Prepare S.11 1. Weigh 3.78 g (30 mmol) of thymine and 20 mg of (NH4 )2 SO4 into a 250-mL round-bottom flask, add 50 mL of 1,1,1,3,3,3-hexamethyldisilazane (b.p. 125◦ C), and attach a condenser equipped with CaCl2 protection tube. 2. Reflux the mixture in an oil bath set at 130◦ C until complete dissolution of thymine (10 to 12 hr). 3. Cool the flask, remove the condenser, and attach an adaptor with a stopcock (Fig. 1.14.5A). Concentrate the solution to a viscous oil using a rotary evaporator connected to a vacuum system (bath temperature ∼30◦ to 35◦ C). The resulting bis-O-trimethylsilylthymine S.10 is very hygroscopic.
4. Close the stopcock of adaptor and attach a second adaptor with stopcock to the flask. Connect the system to a vacuum pump (Fig. 1.14.5B), evacuate the system, and close the upper stopcock. Connect the system with a balloon of nitrogen or argon and open the upper stopcock to fill the adaptor with nitrogen. Repeat this procedure (evacuation and flash with nitrogen) and then open both stopcocks to fill the entire system with nitrogen (Fig. 1.14.5C). 5. Dissolve the residue in 30 mL of anhydrous 1,2-dichloroethane and concentrate the solution to a viscous oil using a rotary evaporator with a bath temperature ∼30◦ to 35◦ C. 6. Repeat step 4. 7. Dissolve the residue in 120 mL of anhydrous 1,2-dichloroethane. Add 12.6 g (25 mmol) of 1-O-acetyl-2,3,5-tri-O-benzoyl-β-D-ribofuranose (S.2) in one portion and 5.79 mL (32 mmol) of trimethylsilyl trifluoromethanesulfonate (TMSOTf) with gentle hand stirring. 8. Attach a condenser equipped with CaCl2 protection tube and heat the clear solution for 1 hr in a 100◦ C oil bath. Monitor reaction by TLC in 98:2 (v/v) CH2 Cl2 /MeOH. The starting S.2 (Rf = 0.91) usually disappears after 1 hr at 100◦ C. The product S.11 (Rf = 0.45) moves slower in the same solvent.
9. Cool the flask to room temperature. Remove the condenser, add 50 mL of saturated sodium bicarbonate solution, and stir the suspension for 20 min at 20◦ C. 10. Put a 2- to 3-cm layer of Hyflo Super Cel on a 100-mL funnel with a sintered disc (porosity 3) and wash with 20 mL CH2 Cl2 . Filter the suspension using vacuum pump and wash the layer with 50 mL CH2 Cl2 . 11. Separate the organic layer using a 250-mL separatory funnel and wash the organic layer with 50 mL of water. 12. Dry the organic layer over ∼20 g Na2 SO4 , filter off Na2 SO4 by gravity filtration, and wash the precipitate with 40 mL CH2 Cl2 . 13. Concentrate the combined filtrates to a solid using a rotary evaporator connected to vacuum system. 14. Add 100 mL of ethanol, attach a condenser, and reflux (79◦ C) until the solid is completely dissolved. Cool the flask to room temperature and keep the mixture for 16 hr at 0◦ C. 15. Collect the precipitate by vacuum filtration on a glass filter (porosity 3), wash the precipitate with 10 mL ethanol, and dry S.11 over phosphorus pentoxide in a vacuum desiccator overnight.
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Figure 1.14.5 Protection from atmospheric moisture for synthesis of the 3 ,5 -protected starting thymidine nucleoside. (A) Evaporation of volatile solvents from a 250-mL round-bottom flask with adaptor and stopcock using a rotary evaporator connected to a vacuum system. (B) A 250-mL round-bottom flask with two adaptors and stopcocks connected to a vacuum system. The upper stopcock is opened. (C) Flashing the whole reaction system with nitrogen or argon using a 250-mL round-bottom flask with two adaptors connected to a balloon of nitrogen or argon with both stopcocks opened.
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16. Characterize the compound by TLC and 1 H NMR. The compound is stable for at least 12 months at 20◦ C. 1-(2,3,5-Tri-O-benzoyl-β-D-ribofuranosyl)thymine (S.11): Yield of white crystals 11.7 g (82%). m.p. 158◦ -159◦ C (EtOH). TLC: Rf 0.45 (98:2 v/v CH2 Cl2 /MeOH). 1 H NMR (400 MHz, CDCl3 ): 8.32 brs (1H, NH), 8.15-7.32 m (15H, Ph), 7.13 q (1H, J6,Me = 1.2, H6), 6.39 d (1H, J1 ,2 = 6.4, H1 ), 5.87 dd (1H, J3 ,2 = 6.0, J3 ,4 = 3.7, H3 ), 5.74 dd (1H, H2 ), 4.83 dd (1H, J5 a,4 = 2.6, J5 a,5 b = −12.2, H5 a), 4.63 ddd (1H, J4 ,5 b = 3.4, H4 ), 4.61 dd (1H, H5 b), 1.55 d (3H, Me).
Prepare S.12 17. Weigh 11.4 g (20 mmol) S.11 into a 500-mL round-bottom flask. Add 200 mL of 0.2 N NaOMe, stopper the flask, and stir the reaction for 30 min at room temperature. 18. Monitor reaction by TLC in 98:2 (v/v) CH2 Cl2 /MeOH. The starting S.11 (Rf = 0.45) usually disappears after 30 min at 20◦ C. The product S.12 (Rf = 0.01) moves slower in the same solvent.
19. Remove MeOH under reduced pressure using a rotary evaporator in vacuo and dissolve the residue in 100 mL water and 100 mL chloroform. 20. Separate the aqueous layer using a 500-mL separatory funnel and wash with 50 mL chloroform. 21. Apply aqueous solution to a 3 × 20–cm chromatography column containing 50 mL of Dowex 50 × 4 (100 to 200 mesh) in H+ form packed in water. 22. Elute with water and collect 50-mL UV-containing fractions until elution is complete (elution volume will vary from 200 to 400 mL). Monitor fractions by TLC using 9:1 (v/v) CH2 Cl2 /MeOH (Rf = 0.11.). 23. Combine the fractions that contain S.12 in a 500-mL round-bottom flask and evaporate to dryness using a rotary evaporator connected to a vacuum system. 24. Add 30 mL EtOH, stopper the flask, and keep the mixture for 16 hr at 0◦ C. 25. Collect the precipitate by vacuum filtration on a glass filter (porosity 3), wash the precipitate with 10 mL EtOH followed by 10 mL diethyl ether, and dry S.12 over phosphorus pentoxide in a vacuum desiccator overnight. 26. Characterize the compound by TLC, UV, and 1 H NMR. The compound is stable for at least 12 months at 20◦ C. 1-(β-D-Ribofuranosyl)thymine (S.12): Yield of white crystals 4.0 g (78%). m.p. 180◦ 181◦ C (EtOH). TLC: Rf 0.11 (9:1 v/v CH2 Cl2 /MeOH). UV (pH 1-7): λmax 269 nm (ε9200); (pH 13): λmax 264 nm (ε7300). 1 H NMR (400 MHz, D2 O): 7.62 s (1H, H6), 5.85 d (1H, J1 ,2 = 4.9 Hz, H1 ), 4.29 dd (1H, J2 ,3 = 5.2 Hz, H2 ), 4.19 dd (1H, J3 ,4 = 5.4 Hz, H3 ), 4.08 ddd (1H, J4 ,5 a = 3.0 Hz, J4 ,5 b = 4.3 Hz, H4 ), 3.87 dd (1H, J5 a,5 b = −12.8 Hz, H5 a), 3.78 dd (1H, H5 b), 1.85 s (3H, Me).
Prepare S.6b 27. Weigh 2.6 g (10 mmol) of dry S.12 in a 100-mL round-bottom flask and add 20 mL anhydrous pyridine. Evaporate using a rotary evaporator equipped with a water aspirator and then apply a dry nitrogen atmosphere (steps 3 to 4, Fig. 1.14.5). 28. Dissolve the residue in 40 mL pyridine with magnetic stirring. Rapidly add 3.14 mL (10 mmol) 1,3-dichloro-1,1,3,3-tetraisopropyldisiloxane (Markiewicz reagent) in one portion, stopper the flask, and keep 4 hr at 20◦ C.
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29. Monitor reaction by TLC using 98:2 (v/v) CH2 Cl2 /MeOH. The starting S.12 (Rf = 0.01) usually disappears after 4 to 6 hr at 20◦ C. The product S.6b (Rf = 0.23) moves faster in the same solvent.
30. Remove nearly all the pyridine using a rotary evaporator connected to a vacuum system and dissolve the residue in 50 mL water and 100 mL CH2 Cl2 . 31. Separate the organic layer using a 250-mL separatory funnel and wash with 20 mL saturated sodium bicarbonate solution and then with 20 mL water. 32. Dry the organic layer over ∼10 g Na2 SO4 , filter off Na2 SO4 by gravity filtration, and wash the precipitate with 20 mL CH2 Cl2 . 33. Evaporate the combined filtrates using a rotary evaporator connected to a vacuum system. Remove traces of pyridine by co-evaporating two times with 20 mL toluene. 34. Prepare a slurry of 50 g of silica gel in CH2 Cl2 and pour it into a 3 × 20–cm chromatography column. 35. Dissolve the obtained residue in a minimal amount CH2 Cl2 and layer it carefully on top of the silica gel. 36. Wash column with 300 mL CH2 Cl2 and 300 mL of 0.5% (v/v) MeOH in CH2 Cl2 . Elute with 1% (v/v) MeOH in CH2 Cl2 . Collect 25-mL fractions. 37. Evaluate fractions by TLC and combine the fractions that contain only S.6b. Evaporate the volatile materials from the combined fractions using a rotary evaporator connected to a vacuum system, and dry the residual foam for 2 to 3 hr using a vacuum oil pump. 38. Grind the foam with a stainless steel spatula and dry the resulting powder 12 to 16 hr using a vacuum oil pump. 39. Characterize the product by TLC and 1 H NMR. The compound is stable for at least 12 months at ambient temperature. 1-[3,5-(1,1,3,3-Tetraisopropyldisiloxane-1,3-diyl)-β-D-ribofuranosyl]thymine (S.6b): Yield of white amorphous solid 4.30 g (86%). TLC: Rf 0.23 (98:2 v/v CH2 Cl2 /MeOH). 1 H NMR (400 MHz, CDCl3 ): 9.18 brs (1H, NH), 7.44 s (1H, H6), 5.70 s (1H, H1 ), 4.38 dd (1H, J3 ,2 = 5.1, Hz J3 ,4 = 8.6 Hz, H3 ), 4.18 dd (1H, J5 a,4 = 2.5 Hz, J5 a,5 b = −13.2 Hz, H5 a), 4.17 d (1H, H2 ), 3.95 ddd (1H, J4 ,5 b = 2.9 Hz, H4 ), 4.01 dd (1H, H5 b), 1.89 s (3H, Me), 1.09-1.02 m (28H, iPr). 13 C NMR (CDCl3 ): 163.93 (C4), 150.21 (C2), 135.99 (C6), 110.74 (C5), 91.42 (C1 ), 82.14 (C4 ), 75.15 (C2 ), 69.57 (C3 ), 60.81 (C5 ), 17.57, 17.50, 17.41, 17.38, 17.22, 17.14, 17.10, 17.02, 13.59, 13.15, 12.91 and 12.76 (iPr), 12.63 (Me).
COMMENTARY Background Information
Synthesis of 2 -O-β β-dRibofuranosylnucleosides
Disaccharide nucleosides belong to an important group of natural compounds that form poly(ADP-ribose) and are found in tRNA, antibiotics, and other physiologically active compounds. To date, ∼100 disaccharide nucleosides and related derivatives have been isolated from a variety of natural sources. These compounds contain an extra carbohydrate residue linked to one of the nucleoside hydroxyl groups via an O-glycosidic bond. The disaccharide residue and heterocyclic base make their properties similar to those of carbo-
hydrates and nucleosides. These compounds manifest a broad spectrum of biological activities, including antibacterial, fungicidal, herbicidal, antitumoral, and antiviral (Lerner, 1991; Efimtseva and Mikhailov, 2004; Nauwelaerts et al., 2004). An effective and simple synthesis of 2 -Oβ-D-ribofuranosylnucleosides, minor tRNA components, has been recently elaborated (Mikhailov et al., 1997a; Markiewicz et al., 1998). The method consists of the condensation of a small excess of 1-O-acetyl-2,3,5-triO-benzoyl-β-D-ribofuranose activated with
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tin tetrachloride with N-protected 3 ,5 -Otetraisopropyldisiloxane-1,3-diyl-ribonucleosides in 1,2-dichloroethane. O-Glycosylation proceeds stereospecifically with the formation of a β-glycosidic bond. Yields of disaccharide nucleosides reach 70% to 80% if the reaction is performed at 0◦ C (Mikhailov et al., 1997a). O-Glycosylation proceeds stereospecifically with formation of a β-glycosidic bond. After complete deprotection, 2 -O-βD-ribofuranosylnucleoside disaccharide nucleosides are obtained in good overall yields (Mikhailov et al., 1997a; Markiewicz et al., 1998). This method has been used for the preparation of pyrimidine 3 -O-β-Dribofuranosyl-2 -deoxyribonucleosides and 3 -O-β-D-ribopyranosyl-2 -deoxyribonucleosides (Mikhailov et al., 1996), and for 5 -O-β-D-ribofuranosyl-2 -deoxyribonucleosides and 5 -O-β-D-ribofuranosylnucleosides (Mikhailov et al., 1997b, 1998). To study the broad applicability of the method, some other sugars such as fully acylated D- and L-arabinofuranose, D-ribopyranose, and D-erythrofuranose have been used in the Oglycosylation reaction, and the corresponding disaccharide nucleosides have been incorporated into oligodeoxyribonucleotides and oligoribonucleotides (Efimtseva et al., 2001). Melting point determinations demonstrated that modified oligonucleotides containing disaccharide nucleosides form stable duplexes with complementary RNA (Tm = 0◦ C; Efimtseva et al., 2001). The duplex RNA maintains an A-type helical geometry with the extra 2 -O-ribose moiety located in the minor groove. This implies that the voluminous extra sugar moiety has almost no effect on the stability of the RNA duplex (Luyten et al., 2000). Oligonucleotides containing 2 -O-ribofuranosylnucleosides have been used as modified primers in RNA-templated DNA synthesis catalyzed by HIV reverse transcriptase. It was shown that an additional 2 -ribofuranose residue in a specific position (-3-4) of the primer prevents elongation (Andreeva et al., 2002). An important feature of these oligonucleotides is the presence of an additional cis-diol group, which may be readily oxidized with sodium periodate to give oligonucleotide derivatives with aldehyde groups placed anywhere in the sequence (Efimtseva and Mikhailov, 2002). Such oligonucleotides were used for affinity labelling of different enzymes (Tunitskaya et al., 1999; Gritsenko et al., 2002). The incorporation of disaccharide nucleosides into oligonucleotides opens
new possibilities for the functionalization of nucleic acids. The extra sugar ring attached to a nucleoside can serve several purposes. For the synthetic work, it is important that acyl blocking groups used in disaccharide synthesis are compatible with the chemistry of automated oligonucleotide synthesis.
Compound Characterization The structure of the compound is supported by NMR spectroscopy and mass spectrometry. The 1 H NMR spectra of the obtained compounds are rather complicated due to the presence of two ribofuranose residues. In spite of this, most of the chemical shifts and coupling constants may be calculated directly from the NMR spectra. In some cases, comparison with the published spectra of disaccharide nucleosides and 1 H-13 C correlation and COSY spectra were used for assignment. The chemical shifts were assigned using double resonance techniques and COSY experiments. Several conclusions were drawn from the 1 H NMR spectral analysis. In nucleosides S.1 and S.6 and disaccharide nucleosides S.3 and S.7 with a 1,1,3,3-tetraisopropyldisiloxane1,3-diyl group, the coupling constants J1 ,2 of nucleoside moieties are <0.5 Hz (Robins et al., 1983). The O-glycosylation reaction proceeded stereospecifically with the formation of 1 ,2 -trans-substituted disaccharide nucleosides S.3 and S.7. The coupling constants (J1 ,2 ) in the additional ribose moiety are <0.5 Hz. As in the case of disaccharide nucleosides (Mikhailov et al., 1997a) and 2 O-methylnucleosides (Robins et al., 1981), incorporation of the 2 -O-substitutent results in a low field shift (+2+3 ppm) of C2 of the nucleoside moiety in 13 C NMR spectra.
Critical Parameters and Troubleshooting
The synthesis of 2 -O-β-D-ribofuranosylnucleosides is short, straightforward, and fairly efficient. However, careful attention to details of basic organic synthesis procedures is required. Preparation of the various compounds requires prior experience with routine chemical laboratory techniques such as solvent evaporation, extraction, TLC, and column chromatography. Characterization of the products demands knowledge of 1 H and 13 C NMR, UV, and mass spectroscopy. General laboratory safety is also of primary concern when hazardous materials are involved. Strict adherence to the outlined methods is therefore highly recommended.
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All solvents should be distilled before use. Anhydrous solvents are very important. They should be either freshly distilled and stored over molecular sieves, or be taken from a freshly opened bottle of commercially prepared anhydrous solvent. It is important for each step of the syntheses that the starting compounds be thoroughly dried either by coevaporation with anhydrous pyridine or in a desiccator over P2 O5 . Protection of reaction mixtures from atmospheric moisture (as shown in Figs. 1.14.2 and 1.14.5) will enhance yields, especially in the case of ribosylation and silylation reactions. For all compounds, a sample of 50 to 100 mg should be kept as a reference. In all cases, the reaction progress is followed by TLC. A baseline is marked and the spots (∼1 optical unit) of the reaction mixture and starting material are placed at equal distances. For the preparation of S.3 and S.7, where the Rf values of the starting compounds and products are very similar, a mixed sample containing both starting compound and reaction mixture may be placed on the TLC plate. After developing in the appropriate solvent, the end-line of the TLC is marked, and spots are identified under a UV lamp (254 nm). Before each column purification, TLC is performed on the reaction mixture to evaluate the completeness of the reaction and to choose the optimal elution system for column chromatography. The silica gel for column chromatography (0.06 to 0.20 mm) is suspended in methylene chloride and loaded into the column, which is tightly packed by gentle tapping until there is a constant volume. The crude product is dissolved in a minimal volume of solvent and is carefully applied using a glass Pasteur pipet. After sample application, the flask and column walls are washed with a minimal volume of solvent. For protection, the silica layer is topped by a 1-cm layer of sand.
Anticipated Results
Synthesis of 2 -O-β β-dRibofuranosylnucleosides
The method presented here generally produces high yields even for inexperienced workers. Good overall yields (46% to 72%) of the final 2 -O-ribofuranosylnucleosides (S.5 and S.9) from the Markiewicz-protected nucleosides (S.1 and S.6) are expected following these procedures. 2 -O-Ribosylation proceeds stereospecifically with formation of a β-glycosidic bond. The first ribosylation reaction is carried out under mild conditions (0◦ C, 1,2-dichloroethane, 2 hr for pyrimidine nucleosides, 7 to 16 hr for purine derivatives) and
the yields of the target compounds are 72% to 80% (Mikhailov et al., 1997a). At room temperature, the reaction occurs faster (30 min), but the yields in the coupling steps are lower (40% to 77%; Markiewicz et al., 1998).
Time Considerations
The three-step preparation of 2 -Oβ-D-ribofuranosylnucleosides starting from Markiewicz-protected nucleosides can be accomplished in 1 week. Each column chromatography step requires 3 to 5 hr. Normally the first two steps (ribosylation and silylation) may be performed in 3 to 4 working days, including purification and spectroscopic analysis. The final deacylation reaction normally proceeds in 2 to 3 days at room temperature and may be run over a weekend. The three-step preparation of the protected thymine starting nucleoside (S.6b) may be accomplished in 1 week.
Literature Cited Andreeva, O.I., Golubeva, A.S., Kochetkov, S.N., Van Aerschot, A., Herdewijn, P., Efimtseva, E.V., Ermolinsky, B.S., and Mikhailov, S.N. 2002. An additional 2 -ribofuranose residue at a specific position of DNA primer prevents its elongation by HIV-1 reverse trancriptase. Bioorg. Med. Chem. Lett. 12:681-684. Efimtseva, E.V. and Mikhailov, S.N. 2002. Disaccharide nucleosides and oligonucleotides on their bases. New tools for the study of enzymes of nucleic acid metabolism. Biochemistry 67:1136-1144. Efimtseva, E.V. and Mikhailov, S.N. 2004. Disaccharide nucleosides. Russ. Chem. Rev. 73:401414. Efimtseva, E.V., Bobkov, G.V., Mikhailov, S.N., Van Aerschot, A., Schepers, G., Busson, R., Rozenski, J., and Herdewijn, P. 2001. Oligonucleotides containing disaccharide nucleosides. Helv. Chim. Acta 84:2387-2397. Gritsenko, O.M., Koudan, E.V., Mikhailov, S.N., Ermolinsky, B.S., Van Aerschot, A., Herdewijn, P., and Gromova, E.S. 2002. Affinity modification of EcoRII DNA methyltransferase by the dialdehyde-substituted DNA duplexes: Mapping the enzyme region that interacts with DNA. Nucleosides Nucleotides Nucleic Acids 21:753764. Lerner, L.M. 1991. Synthesis and Properties of Various Disaccharide Nucleosides. In Chemistry of Nucleosides and Nucleotides, Vol. 2 (L.B. Townsend, ed.) pp. 27-79. Plenum Press, New York. Luyten, I., Esnouf, R.M., Mikhailov, S.N., Efimtseva, E.V., Michiels, P., Heus, H.A., Hilbers, C.W., and Herdewijn, P. 2000. Solution structure of a RNA decamer duplex, containing 9-[(2-O-(β-D-ribofuranosyl)-(β-Dribofuranosyl]adenine, a special residue in lower
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eukaryotic initiator tRNAs. Helv. Chim. Acta 83:1278-1289. Markiewicz, W.T. and Wiewiorowski, M. 1986. 1-[3,5-(1,1,3,3-Tetraisopropyldisiloxane-1,3diyl)ribonucleosides. In Nucleic Acid Chemistry. Improved and New Synthetic Procedures and Techniques, Part 3: (L.B. Townsend and R.S. Tipson, eds.) pp. 229-231. John Wiley & Sons, Hoboken. Markiewicz, W.T., Niewczyk, A., Gdaniec, Z., Adamiak, D.A., Dauter, Z., Rypniewski, W., and Chmielewski, M. 1998. Studies on synthesis and structure of O-β-D-ribofuranosyl (1 →2 )ribonucleosides and oligonucleotides. Nucleosides Nucleotides Nucleic Acids 17:411424. Mikhailov, S.N., De Clercq, E., and Herdewijn, P. 1996. Ribosylation of pyrimidine 2 deoxynucleosides. Nucleosides Nucleotides Nucleic Acids 15:1323-1334. Mikhailov, S.N., Efimtseva, E.V., Gurskaya, G.V., Zavodnik, V.E., De Bruyn, A., Janssen, G., Rozenski, J., and Herdewijn, P. 1997a. An efficient synthesis and physico-chemical properties of 2 -O-β-D-ribofuranosyl-nucleosides, minor tRNA components. J. Carbohydr. Chem. 16:75-92. Mikhailov, S.N., Rodionov, A.A., Efimtseva, E.V., Fomitcheva, M.V., Padyukova, N.Sh., Herdewijn, P., and Oivanen, M. 1997b. Preparation of pyrimidine 5 -O-β-D-ribofuranosylnucleosides, and hydrolytic stability of O-Dribofuranosyl-nucleosides. Carbohydrate Lett. 2:321-328. Mikhailov, S.N., Rodionov, A.A., Efimtseva, E.V., Ermolinsky, B.S., Fomitcheva, M.V., Padyukova, N.Sh., Rothenbacher, K., Lescrinier, E., and Herdewijn, P. 1998. Formation of trisaccharide nucleoside during disaccharide nucleoside synthesis. Eur. J. Org. Chem. 2193-2199.
Nauwelaerts, K., Efimtseva, E.V., Mikhailov, S.N., and Herdewijn, P. 2004. Disaccharide nucleosides, an important group of natural compounds In Frontiers in Nucleosides and Nucleic Acids (R.F. Schinazi and D.C. Liotta, eds.) pp 187-220. IHL Press, Arlington, MA. Robins, M.J., Hansske, F., and Bermier, S.E. 1981. Nucleic acid related compounds. 36. Synthesis of the 2 -O-methyl and 3 -O-methyl ethers guanosine and 2-amino adenosine and correlation of O -methylnucleoside C13 NMR spectral shifts. Can. J. Chem. 59:3360-3364. Robins, M.J., Wilson, J.S., Sawyer, L., and James, M.N.G. 1983. Nucleic acid related compounds. 41. Restricted furanose conformations of 3 ,5 -(1,1,3,3-tetraisopropyldisilox-1,3diyl)nucleosides provide a convenient evaluation of anomeric configuration. Can. J. Chem. 59:1911-1920. Tunitskaya, V.L., Rusakova, E.E., Memelova, L.V., Kochetkov, S.N., Van Aerschot, A., Herdewijn, P., Efimtseva, E.V., Ermolinsky, B.S., and Mikhailov, S.N. 1999. The mapping of T7 RNA polymerase active site with novel reagents—Oligonucleotides with reactive dialdehyde groups. FEBS Lett. 442:20-24. Vorbr¨uggen, H. and Ruh-Pohlenz, C. 2000. Synthesis of nucleosides. Org. React. 55:1-630.
Contributed by Sergey N. Mikhailov, Ekaterina V. Efimtseva, Andrei A. Rodionov, Georgii V. Bobkov, and Irina V. Kulikova Russian Academy of Sciences Engelhardt Institute of Molecular Biology Moscow, Russia Piet Herdewijn Rega Institute for Medical Research Leuven, Belgium
Synthesis of Modified Nucleosides
1.14.19 Current Protocols in Nucleic Acid Chemistry
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Preparation of 2 -Deoxy-2 -MethylselenoModified Phosphoramidites and RNA
UNIT 1.15
This unit contains procedures for the preparation of RNA possessing site-specifically incorporated ribose 2 -deoxy-2 -methylseleno (2 -Se-methyl) groups. The unit begins with the preparation of 5 -O-DMTr-3 -phosphoramidites of 2 -deoxy-2 -methylselenouridine (see Basic Protocol 1), N4 -acetyl-2 -deoxy-2 -methylselenocytidine (see Alternate Protocol 1), N6 -acetyl-2 -deoxy-2 -methylselenoadenosine (see Alternate Protocol 2), and N2 acetyl-2 -deoxy-2 -methylselenoguanosine (see Alternate Protocol 3). Methods are also presented for solid-phase synthesis of oligoribonucleotides labeled with 2 -methylseleno groups using 2 -O-[(triisopropylsilyl)oxy]methyl (TOM) phosphoramidites (see Basic Protocol 2; also see UNITS 2.9 & 3.8), along with their deprotection and purification (see Basic Protocol 3). The final protocols describe enzymatic ligation of the oligonucleotides to produce biologically interesting RNA targets (see Basic Protocol 4 and Alternate Protocol 4). CAUTION: Carry out all operations involving organic solvents and reagents in a wellventilated fume hood, and wear gloves and protective glasses. Procedures involving trifluoromethanesulfonyl chloride and dimethyl diselenide should be performed only by personnel trained and experienced in organic synthesis. Standard precautions to prevent excessive exposure to toxic chemicals should be followed. All reactions should first be performed on a small scale.
PREPARATION OF 2 -DEOXY-2 -METHYLSELENOURIDINE PHOSPHORAMIDITE
BASIC PROTOCOL 1
This protocol describes the synthesis of 5 -O-(4,4 -dimethoxytrityl)-2 -deoxy-2 methylselenouridine 3 -O-(2-cyanoethyl)-N,N-diisopropylphosphoramidite. The synthesis involves four steps: (1) transformation of uridine into 2,2 -anhydrouridine (S.1), (2) regioselective introduction of the 5 -O-DMTr-group to give S.2, (3) SN 2-ring opening by sodium methylselenide to produce S.3, and (4) 3 -OH phosphitylation using 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite to give S.4. These steps are illustrated in Figure 1.15.1.
Materials Diphenyl carbonate (Fluka) N,N-Dimethylformamide (DMF) Uridine (Fluka) Sodium bicarbonate (NaHCO3 ) Methanol (MeOH) Balloon of argon Anhydrous pyridine 4,4 -Dimethoxytrityl chloride (DMTr-Cl; Fluka) Dichloromethane (CH2 Cl2 ) 5% (w/v) citric acid solution in water Half-saturated sodium bicarbonate solution in water Sodium sulfate (Na2 SO4 ) Silica gel 60 (230 to 400 mesh; Fluka) Triethylamine (TEA) Anisaldehyde reagent (see recipe) Sodium borohydride (NaBH4 ; Fluka)
Synthesis of Modified Nucleosides
Contributed by Ronald Micura, Claudia H¨obartner, Renate Rieder, Christoph Kreutz, Barbara Puffer, Kathrin Lang, and Holger Moroder
1.15.1
Current Protocols in Nucleic Acid Chemistry (2006) 1.15.1-1.15.34 C 2006 by John Wiley & Sons, Inc. Copyright
Supplement 27
Figure 1.15.1 Preparation of a 2 -deoxy-2 -methylselenouridine phosphoramidite building block for RNA solid-phase synthesis (see Basic Protocol 1). DMF, N,N-dimethylformamide; DMTr, 4,4 -dimethoxytrityl.
Anhydrous tetrahydrofuran (THF) Dimethyl diselenide (Aldrich) 30% (v/v) H2 O2 solution Anhydrous ethanol 0.2 M triethylammonium acetate (TEAA) buffer, pH 7.0 Ethyl acetate Saturated aqueous sodium chloride (NaCl) solution N-Ethyldimethylamine (Fluka) 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite (Aldrich) Hexanes Chloroform (CHCl3 ) 25-mL two-neck round-bottom flask Reflux condenser Glass-fritted B¨uchner funnel (pore size 3) Filter paper (Macherey-Nagel MN 615) Vacuum oil pump (rotary vane vaccuum pump; for all steps requiring drying in vacuo) 50-mL one-neck round-bottom flasks equipped with rubber septa Rotary evaporator equipped with a membrane pump (for all evaporation steps) 250-, 500-, and 1000-mL separatory funnels 100-mL two-neck round-bottom flasks with rubber septa Glass filter tube 1- and 20-mL syringes and 21-G needles 500-µL Hamilton syringe Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) 2 -MethylselenoModified Phosphoramidites and RNA
Prepare 2,2 -anhydrouridine 1. Prepare a stirred suspension of 1.92 g (9.0 mmol) diphenyl carbonate in 2.2 mL dry DMF in a 25-mL two-neck flask equipped with a condenser and stir bar.
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2. Add 2.0 g (8.2 mmol) uridine and heat the slurry to 80◦ C with stirring. 3. Add 25 mg (0.3 mmol) NaHCO3 , heat to 120◦ C, and continue stirring for 3 to 4 hr. Gas evolution is observed and a clear solution is obtained from which the solid product soon precipitates.
4. Cool the solution to room temperature and isolate the precipitate by filtration using a glass-fritted B¨uchner funnel. Wash the precipitate with 15 mL MeOH. 5. Dry the resulting white powder 2 hr in vacuo using a vacuum oil pump. 2,2 -O-Anhydro-(β-D-arabinofuranosyl)uracil (S.1). Yield: 1.4 g (75%). TLC (85:15 CH2 Cl2 /MeOH): Rf 0.2. 1 H NMR (300 MHz, DMSO- d6 ): δ 3.17-3.28 (m, 2H, H5 ); 4.06 (m, 1H, H4 ); 4.37 (m, 1H, H3 ); 4.95 (triplettoid, 1H, 5 -OH); 5.18 (d, J = 6.2 Hz, 1H, H2 ); 5.84 (m, 2H, H5 + 3 -OH); 6.29 (d, J = 6.2 Hz, 1H, H1 ); 7.82 (d, J = 7.4 Hz, 1H, H6).
Protect 5 -hydroxy group 6. In a 50-mL one-neck flask equipped with a rubber septum, an argon-filled balloon, and a stir bar, suspend 1.0 g (4.4 mmol) S.1 in 10 mL dry pyridine. 7. Add 1.65 g (4.8 mmol) DMTr-Cl in three 0.55-g portions at 20-min intervals. Continue stirring at room temperature until a clear solution is obtained (4 to 6 hr). 8. Add 1.0 mL MeOH, stir for 30 min, and evaporate the solvents. 9. Dissolve the residue in 300 mL CH2 Cl2 . 10. Wash two times each with 200 mL of 5% citric acid, 200 mL water, and 200 mL of half-saturated sodium bicarbonate solution in a 1000-mL separatory funnel. 11. Dry the organic phase with 70 g Na2 SO4 , remove the solid by means of a glassfritted B¨uchner funnel and filter paper, and evaporate the filtrate to dryness. 12. Purify the residue on a chromatography column (APPENDIX 3E) containing 50 g of 230 to 400-mesh silica gel, using a solvent system of 2% to 10% (v/v) MeOH/1% (v/v) TEA in CH2 Cl2 . 13. Combine fractions containing product S.2 as determined by TLC (APPENDIX 3D) using 9:1 (v/v) CH2 Cl2 /MeOH as eluent and visualizing the product with anisaldehyde reagent. Evaporate and then coevaporate with 20 mL CH2 Cl2 . 14. Dry the residue in vacuo to obtain the product as a colorless foam. 2,2 -O-Anhydro-5 -O-(4,4 -dimethoxytrityl)-(β-D-arabinofuranosyl)uracil (S.2). Yield: 1.55 g (67%). TLC (9:1 CH2 Cl2 /MeOH): Rf 0.6. 1 H NMR (500 MHz, DMSO-d6 ): δ 2.80 (dd, J = 10.0, 7.5 Hz, 1H, H5 a); 2.93 (dd, J = 10.0, 4.0 Hz, 1H, H5 b); 3.72 (s, 6H, 2 × OCH3 ); 4.20 (m, 1H, H4 ); 4.29 (m, 1H, H3 ); 5.19 (d, J = 6.0 Hz, 1H, H2 ); 5.86 (d, J = 7.5 Hz, 1H, H5); 5.94 (s, br, 1H, 3 -OH); 6.31 (d, J = 6.0 Hz, 1H, H1 ); 6.82 (m, 4H, H-C(ar)); 7.12-7.28 (m, 9H, H-C(ar)); 7.93 (d, J = 7.5 Hz, 1H, H6).
Introduce 2 -methylseleno group 15. In a 100-mL two-neck round-bottom flask, place 430 mg (11.3 mmol) NaBH4 and a stir bar. 16. Attach a glass filter tube on the flask and connect a second 100-mL two-neck round-bottom flask on top (needed for filtration under inert conditions in step 20). Evacuate using a vacuum oil pump to deplete all oxygen, and expand argon into the apparatus from a balloon by injection through one of the rubber septa. 17. Using a 20-mL syringe and 21-G needle, inject 14 mL dry THF through the septum to suspend the NaBH4 .
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18. Inject 0.36 mL (3.8 mmol) dimethyl diselenide through the septum using a 500-µL Hamilton syringe. Rinse the injection needle, syringe, and all glassware contaminated by the selenide reagent with 30% H2 O2 solution and collect the waste in a separate bottle.
19. Add anhydrous ethanol dropwise by injection through the septum until gas bubbles start to appear in the yellow mixture (requires ∼0.3 to 0.4 mL). Stir the solution for 1 hr at room temperature. The solution turns almost colorless when the reaction is complete.
20. Filter the solution under argon from remaining NaBH4 by cautiously turning the apparatus upside down and pressurizing via the argon balloon. Optionally, apply a soft vacuum on the second flask to accelerate the filtration process.
21. Via the septum, transfer the filtered solution into a 20-mL syringe and slowly inject it into a stirred solution of 1.0 g (1.89 mmol) S.2 in 20 mL dry THF under argon. 22. Monitor the reaction by TLC using 92:8 (v/v) CH2 Cl2 /MeOH. The reaction is usually complete after ∼1 to 2 hr.
23. Add 30 mL of aqueous 0.2 M TEAA buffer, pH 7.0. 24. Reduce the solution to half-volume by evaporation. 25. Dilute with 100 mL water and extract the product three times with 20 mL ethyl acetate. 26. Wash the combined ethyl acetate phases with 200 mL saturated NaCl solution. 27. Dry the organic phase with 20 g Na2 SO4 , remove the solid by means of a glassfritted B¨uchner funnel and filter paper, and evaporate the filtrate to dryness. 28. Purify the residue on a 40-g silica gel column using a solvent system of 1% to 2% (v/v) MeOH in CH2 Cl2 . 29. Combine the fractions containing the product S.3, as determined by TLC using 92:8 (v/v) CH2 Cl2 /MeOH. Evaporate and then coevaporate with 15 mL CH2 Cl2 . 30. Dry the residue in vacuo to obtain the product as a colorless foam. 5 -O-(4,4 -Dimethoxytrityl)-2 -deoxy-2 -methylselenouridine (S.3). Yield: 1.08 g (92%). TLC (92:8 CH2 Cl2 /MeOH): Rf 0.75. 1 H NMR (300 MHz, CDCl3 ): δ 2.12 (s, 3H, SeCH3 ); 2.62 (d, J = 3.0 Hz, 1H, 3 -OH); 3.51-3.61 (m, 3H, 2 × H5 , H2 ); 3.81 (s, 6H, 2 × OCH3 ); 4.19 (m, 1H, H4 ); 4.39 (m, 1H, H3 ); 5.38 (d, J = 8.3 Hz, 1H, H5); 6.18 (d, J = 7.5 Hz, 1H, H1 ); 6.85 (m, 4H, H-C(ar)); 7.25-7.38 (m, 9H, H-C(ar)); 7.78 (d, J = 8.3 Hz, 1H, H6); 8.11 (s, br, 1H, NH). FTICR ESI-MS (m/z): calcd. for C31 H32 N2 O7 Se [M+Na]+ 647.12756, found 647.12756.
Prepare phosphoramidite 31. In a 50-mL one-neck flask equipped with a stir bar, a rubber septum, and an argon balloon, prepare a stirred mixture of 1.04 mL (9.6 mmol) N-ethyldimethylamine and 10 mL dry CH2 Cl2 . 32. Add 0.6 g (0.96 mmol) S.3 and dissolve.
2 -MethylselenoModified Phosphoramidites and RNA
33. Slowly add 340 mg (1.44 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite using a 1-mL syringe and continue stirring for 2 hr at room temperature.
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34. Monitor the reaction by TLC using 7:3 (v/v) ethyl acetate/hexanes. The reaction is usually complete after 2 to 4 hr.
35. Add 0.4 mL MeOH to quench the excess phosphine. Continue stirring for 15 min. 36. Dilute the reaction mixture with 50 mL of CH2 Cl2 and wash with 40 mL of halfsaturated sodium bicarbonate solution. 37. Dry the organic phase with 10 g Na2 SO4 , remove the solid by means of a glassfritted B¨uchner funnel and filter paper, and evaporate the filtrate to dryness. 38. Purify the residue on a 14-g silica gel column using a solvent system of 50% to 70% (v/v) ethyl acetate/1% (v/v) TEA in hexanes. 39. Combine the fractions containing product S.4, as determined by TLC using 7:3 (v/v) ethyl acetate/hexanes. Evaporate and then coevaporate with 10 mL CHCl3 . 40. Dry the residue in vacuo to obtain the product as a colorless foam. 5 -O-(4,4 -Dimethoxytrityl)-2 -deoxy-2 -methylselenouridine 3 -O-(2-cyanoethyl)-N,Ndiisopropylphosphoramidite (S.4). Yield: 0.74 g (93%; mixture of diastereoisomers). TLC (7:3 ethyl acetate/hexanes): Rf 0.6. 1 H NMR (500 MHz, CDCl3 ): δ 1.05-1.28 (m, 24H, ((CH3 )2 CH)2 N); 2.04, 2.08 (2s, 6H, SeCH3 ); 2.37, 2.62 (2m, 4H, CH2 CN); 3.413.69 (m, 10H, ((CH3 )2 CH)2 N, POCH2 , 2 × H5 ); 3.79 (2s, 12H, 2 × OCH3 ); 3.90, 3.93 (2m, 2H, POCH2 ); 4.24, 4.30 (2m, 2H, H4 ); 4.63, 4.68 (2m, 2H, H3 ); 5.25, 5.31 (2d, 2H, J = 8.0 Hz, H5); 6.33 (d, J = 6.8 Hz, 2H, H1 ); 6.84 (m, 8H, H-C(ar)); 7.24-7.37 (m, 18H, H-C(ar)); 7.80 (d, 2H, J = 8.0 Hz, H6); 8.36 (s, br, 2H, NH). 31 P NMR (121 MHz, CDCl3 ): δ 151.1, 151.9. FTICR ESI-MS (m/z): calcd. for C40 H49 N4 O8 PSe [M+Na]+ 847.23560, found 847.23285; calcd [M+K]+ 863.20954, found 863.20837.
PREPARATION OF 2 -DEOXY-2 -METHYLSELENOCYTIDINE PHOSPHORAMIDITE
ALTERNATE PROTOCOL 1
This protocol describes the synthesis of N4 -acetyl-5 -O-(4,4 -dimethoxytrityl)-2 deoxy-2 -methylselenocytidine 3 -O-(2-cyanoethyl)-N,N-diisopropylphosphoramidite. The synthesis starts from the uridine precursor S.3 (see Basic Protocol 1) and proceeds in six steps: (1) 3 -OH silylation of S.3 to give S.5, (2) O4 -sulfonation to S.6, (3) NH3 -mediated conversion of S.6 to S.7, (4) N4 -acetylation to give S.8, (5) 3 O-desilylation to give S.9, and (6) 3 -OH phosphitylation using 2-cyanoethyl-N,Ndiisopropylchlorophosphoramidite to give S.10. These steps are illustrated in Figure 1.15.2.
Additional Materials (also see Basic Protocol 1) S.3 (see Basic Protocol 1) Imidazole (Fluka) tert-Butyldimethylsilyl chloride (TBDMS-Cl; Fluka) 4-(N,N-Dimethylamino)pyridine (DMAP; Fluka) 2,4,6-Triisopropylbenzenesulfonyl chloride (TPS-Cl; Fluka) Saturated sodium bicarbonate solution in water 32% (v/v) aqueous ammonia Acetic anhydride (Fluka) Tetrabutylammonium fluoride trihydrate (TBAF; Fluka) Acetic acid (AcOH) 25-mL one-neck round-bottom flasks equipped with rubber septa Protect 3 -hydroxyl group of S.3 1. In a 50-mL one-neck flask equipped with a stir bar, a rubber septum, and an argon balloon, dissolve 1.0 g (1.60 mmol) S.3 in 8 mL DMF.
Synthesis of Modified Nucleosides
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Supplement 27
Figure 1.15.2 Preparation of a 2 -deoxy-2 -methylselenocytidine phosphoramidite building block for RNA solid-phase synthesis (see Alternate Protocol 1). DMAP, 4-(N,N-dimethylamino)pyridine; DMF, N,N-dimethylformamide; DMTr, 4,4 dimethoxytrityl; TBAF, tetra-n-butylammonium fluoride; TBDMS, tert-butyldimethylsilyl; THF, tetrahydrofuran; TPS, 2,4,6triisopropylbenzenesulfonyl.
2. While stirring, add 0.44 g (6.40 mmol) imidazole and then 0.48 g (3.20 mmol) TBDMS-Cl. Keep stirring for 18 hr at room temperature. 3. Evaporate the solvents and precipitate the product from the resulting oil by adding 35 mL water. 4. Isolate the precipitate by filtration on a glass-fritted B¨uchner funnel with filter paper. Wash the precipitate with 20 mL water and 20 mL hexanes, and dry in vacuo using a vacuum oil pump.
2 -MethylselenoModified Phosphoramidites and RNA
3-O-tert-Butyldimethylsilyl-5-O-(4,4-dimethoxytrityl)-2-deoxy-2 -methylselenouridine (S.5). Yield: 1.12 g (95%). TLC (98:2 CH2 Cl2 /MeOH): Rf 0.65. UV (MeOH): λ(ε) 260 (11600) nm (L mol−1 cm−1 ). 1 H NMR (300 MHz, CDCl3 ): δ −0.02 (s, 3H, SiCH3 ); 0.12 (s, 3H, SiCH3 ); 0.86 (s, 9H, SiC(CH3 )3 ); 2.04 (s, 3H, SeCH3 ); 3.38 (dd, J = 11.0, 2.3 Hz, 1H, H5 a); 3.47 (m, 1H, H2 ); 3.56 (dd, J = 11.0, 3.0 Hz, 1H, H5 b); 3.81 (s, 6H, 2 × OCH3 ); 4.09 (m, 1H, H4 ); 4.48 (m, 1H, H3 ); 5.33 (d, J = 8.3 Hz, 1H, H5); 6.34 (d, J = 7.1 Hz, 1H, H1 ); 6.85 (d, J = 8.7 Hz, 4H, H-C(ar)); 7.25-7.38 (m, 9H, H-C(ar)); 7.88 (d, J = 8.3 Hz, 1H, H6); 7.95 (s, br, 1H, NH). 13 C NMR (75 MHz, CDCl3 ): δ −4.8 (SiCH3 ); −4.7 (SiCH3 ); 3.3 (SeCH3 ); 18.1 (SiC(CH3 )3 ); 25.7 (SiC(CH3 )3 ); 48.6 (C2 ); 55.2 (OCH3 ); 62.4 (C5 ); 73.5 (C3 ); 85.6 (C4 ); 87.3; 90.0 (C1 ); 102.5 (C5); 113.3 (C(ar)); 127.2, 128.0, 128.2, 130.1 (C(ar)); 134.9, 135.1, 140.2 (C6); 144.0, 150.3, 158.8, 163.1. FTICR ESI-MS (m/z): calcd. for C37 H46 N2 O7 SeSi [M+Na]+ 761.21421, found 761.21496.
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Sulfonate O4 and displace adduct with ammonia 5. In a 50-mL one-neck flask equipped with a stir bar, a rubber septum, and an argon balloon, dissolve 1.0 g (1.36 mmol) S.5, 20 mg (0.16 mmol) DMAP, and 1.9 mL (13.6 mmol) TEA in 10 mL dry CH2 Cl2 . 6. Slowly add 0.62 g (2.04 mmol) 2,4,6-triisopropylbenzenesulfonyl chloride and stir for 1 hr at room temperature. 7. Dilute the reaction mixture with 100 mL CH2 Cl2 and wash with 80 mL saturated sodium bicarbonate solution. 8. Dry over 20 g Na2 SO4 , remove the solid by means of a glass-fritted B¨uchner funnel and filter paper, and evaporate the solvents. 9. Dry the resulting intermediate S.6 (yellow foam) for 1 hr in vacuo using a vacuum oil pump. 10. Dissolve S.6 in 20 mL THF, add 30 mL of 32% aqueous ammonia, and stir overnight at room temperature. 11. Evaporate the solvents and purify the residue on a chromatography column (APPENDIX 3E) containing 40 g silica gel, using a solvent system of 2% to 8% (v/v) MeOH in CH2 Cl2 . 12. Combine the fractions containing product S.7, as determined by TLC (APPENDIX 3D) using 95:5 (v/v) CH2 Cl2 /MeOH. Evaporate on a rotary evaporator and then coevaporate with 10 mL CH2 Cl2 . 13. Dry the residue in vacuo using a vacuum oil pump to obtain the product as a colorless foam. 3-O-tert-Butyldimethylsilyl-5-O-(4,4-dimethoxytrityl)-2-deoxy-2 -methylselenocytidine (S.7). Yield: 0.79 g (79%). TLC (95:5 CH2 Cl2 /MeOH): Rf 0.45. UV (MeOH): λ(ε) 260 (10300) nm (L mol−1 cm−1 ). 1 H NMR (300 MHz, CDCl3 ): δ −0.08 (s, 3H, SiCH3 ); 0.08 (s, 3H, SiCH3 ); 0.81 (s, 9H, SiC(CH3 )3 ); 2.13 (s, 3H, SeCH3 ); 3.30 (dd, J = 2.3, 10.5 Hz, 1H, H5 a); 3.58 (m, 1H, H2 ); 3.63 (dd, J = 3.0, 10.5 Hz, 1H, H5 b); 3.80 (s, 6H, 2 × OCH3 ); 4.14 (m, 1H, H4 ); 4.46 (m, 1H, H3 ); 5.30 (d, J = 7.5 Hz, 1H, H5); 6.38 (d, J = 3.9 Hz, 1H, H1 ); 6.84 (d, J = 7.6 Hz, 4H, H-C(ar)); 7.26-7.40 (m, 9H, H-C(ar)); 8.12 (d, J = 7.5 Hz, 1H, H6). 13 C NMR (75 MHz, CDCl3 ): δ −4.9 (SiCH3 ); −4.7 (SiCH3 ); 3.1 (SeCH3 ); 18.1 (SiC(CH3 )3 ); 25.7 (SiC(CH3 )3 ); 49.1 (C2 ); 55.3 (OCH3 ); 61.9 (C5 ); 72.1 (C3 ); 84.6 (C4 ); 87.0; 90.7 (C1 ); 94.9 (C5); 113.2, 113.3 (C(ar)); 127.1, 127.9, 128.3, 130.2 (C(ar)); 135.3, 135.3; 141.2 (C(6)); 144.3, 158.8, 165.4. FTICR ESI-MS (m/z): calcd. for C37 H47 N3 O6 SeSi [M+Na]+ 760.23018, found 760.22880.
Acetylate N4 14. In a 25-mL one-neck flask equipped with a stir bar, a rubber septum, and an argon balloon, dissolve 0.4 g (0.54 mmol) S.7 in 1.0 mL dry pyridine and cool to 0◦ C. 15. Add 130 µL (136 mmol) acetic anhydride, allow solution to warm up to room temperature, and stir for 45 min. 16. Quench the reaction by adding 0.6 mL MeOH and evaporate the solvents. 17. Dissolve the oily residue in 100 mL CH2 Cl2 . 18. Wash two times each with 50 mL of 5% citric acid, 50 mL water, and 50 mL saturated sodium bicarbonate solution. 19. Dry over 15 g Na2 SO4 , remove the solid by means of a glass-fritted B¨uchner funnel and filter paper, and evaporate the solvents.
Synthesis of Modified Nucleosides
1.15.7 Current Protocols in Nucleic Acid Chemistry
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20. Purify the residue on a 16-g silica gel column using a solvent system of 1% to 5% (v/v) MeOH in CH2 Cl2 . 21. Combine the fractions containing product S.8, as determined by TLC using 97:3 (v/v) CH2 Cl2 /MeOH. Evaporate and then coevaporate with 5 mL CH2 Cl2 . 22. Dry the residue in vacuo to obtain the product as a colorless foam. N4 -Acetyl-3 -O-tert-butyldimethylsilyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxy-2 -methylselenocytidine (S.8). Yield: 0.39 g (94%). TLC (97:3 CH2 Cl2 /MeOH): Rf 0.65. UV (MeOH): λ(ε) 272 (6000), 300 (5800) nm (L mol−1 cm−1 ). 1 H NMR (300 MHz, CDCl3 ): δ −0.06 (s, 3H, SiCH3 ); 0.08 (s, 3H, SiCH3 ); 0.82 (s, 9H, SiC(CH3 )3 ); 2.15 (s, 3H, SeCH3 ); 2.27 (s, 3H, COCH3 ); 3.38 (dd, J = 2.0, 10.5 Hz, 1H, H5 a); 3.58 (m, 1H, H2 ); 3.67 (dd, J = 2.0, 10.5 Hz, 1H, H5 b); 3.82 (s, 6H, 2 × OCH3 ); 4.20 (m, 1H, H4 ); 4.44 (m, 1H, H3 ); 6.39 (d, J = 3.9 Hz, 1H, H1 ); 6.86 (d, J = 8.0 Hz, 4H, H-C(ar)); 7.08 (d, J = 7.5 Hz, 1H, H5); 7.28-7.40 (m, 9H, H-C(ar)); 8.47 (d, J = 7.5 Hz, 1H, H6); 10.37 (s, br, 1H, NH). 13 C NMR (75 MHz, CDCl3 ): δ −5.0 (SiCH3 ); −4.7 (SiCH3 ); 3.2 (SeCH3 ); 18.0 (SiC(CH3 )3 ); 24.8 (COCH3 ); 25.6 (SiC(CH3 )3 ); 49.3 (C2 ); 55.2 (OCH3 ); 61.5 (C5 ); 71.7 (C3 ); 85.0 (C4 ); 87.2; 91.1 (C1 ); 97.0 (C5); 113.3 (C(ar)); 127.3, 128.0, 128.3, 130.2 (C(ar)); 135.0, 135.1, 143.9; 144.7 (C6); 155.1, 158.8, 163.1, 171.0. FTICR ESI-MS (m/z): calcd. for C39 H49 N3 O7 SeSi [M+Na]+ 802.24080, found 802.24121.
Deprotect 3 -hydroxyl group 23. In a 25-mL one-neck flask equipped with a stir bar, a rubber septum, and an argon balloon, dissolve 0.41 g (0.52 mmol) S.8 in 1.0 mL of 1 M TBAF/0.5 M AcOH in THF. Stir for 2.5 hr at room temperature. 24. Evaporate the solvents and purify the residue on a 14-g silica gel column using a solvent system of 2% to 6% (v/v) MeOH in CH2 Cl2 . 25. Combine the fractions containing product S.9, as determined by TLC using 97:3 (v/v) CH2 Cl2 /MeOH. Evaporate and then coevaporate with 5 mL CH2 Cl2 . 26. Dry the residue in vacuo to obtain the product as colorless foam. N4 -Acetyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxy-2 -methylselenocytidine (S.9). Yield: 0.30 g (87 %). TLC (97:3 CH2 Cl2 /MeOH): Rf 0.40. UV (MeOH): λ(ε) 272 (7200), 300 (7400) nm (L mol−1 cm−1 ). 1 H NMR (300 MHz, CDCl3 ): δ 2.17 (s, 3H, SeCH3 ); 2.19 (s, 3H, COCH3 ); 3.00 (s, br, 1H, 3 -OH); 3.54 (2dd, J = 3.0, 11.3 Hz, 2H, H5 ); 3.66 (m, 1H, H2 ); 3.81 (s, 6H, 2 × OCH3 ); 4.15 (m, 1H, H4 ); 4.39 (m, 1H, H3 ); 6.30 (d, J = 4.5 Hz, 1H, H1 ); 6.86 (d, J = 9.2 Hz, 4H, H-C(ar)); 7.14 (d, J = 7.5 Hz, 1H, H5); 7.26-7.42 (m, 9H, H-C(ar)); 8.30 (d, J = 7.5 Hz, 1H, H6); 9.48 (s, br, 1H, NH). 13 C NMR (75 MHz, CDCl3 ): δ 4.6 (SeCH3 ); 24.7 (COCH3 ); 51.5 (C2 ); 55.2 (OCH3 ); 62.2 (C5 ); 70.0 (C3 ); 84.4 (C4 ); 87.2; 90.2 (C1 ); 97.0 (C5); 113.4 (C(ar)); 127.2, 128.0, 128.1, 130.1 (C(ar)); 135.2, 135.4, 144.2; 144.5 (C6); 155.3, 158.8, 162.9, 170.8. FTICR ESI-MS (m/z): calcd. for C33 H35 N3 O7 Se [M+H]+ 666.17217, found 666.17208; calcd. [M+Na]+ 688.15414, found 688.15344.
Prepare phosphoramidite 27. Apply Basic Protocol 1, steps 31 to 40, to 0.64 g (0.96 mmol) S.9. For column purification of S.10, use 14 g silica gel and a solvent system of 70% (v/v) ethyl acetate/1% (v/v) TEA in hexanes.
2 -MethylselenoModified Phosphoramidites and RNA
N4 -Acetyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxy-2 -methylselenocytidine 3 -O-(2-cyanoethyl)-N,N-diisopropylphosphoramidite (S.10). Yield: 0.68 g (82%; mixture of diastereoisomers). TLC (96:4 CH2 Cl2 /MeOH): Rf 0.55. UV (MeOH): λ(ε) 274 (7830), 300 (7950) nm (L mol−1 cm−1 ). 1 H NMR (500 MHz, CDCl3 ): δ 0.98-1.25 (m, 24H, ((CH3 )2 CH)2 N); 2.11 (s, 6H, COCH3 ); 2.15, 2.17 (2s, 6H, SeCH3 ); 2.33, 2.59 (2m, 4H, CH2 CN); 3.41-3.70 (m, 12H, ((CH3 )2 CH)2 N, POCH2 , 2 × H5 , H2 ); 3.78 (2s, 12H, 2 × OCH3 ); 3.80-3.93 (m, 2H, POCH2 ); 4.29 (m, 2H, H4 ); 4.55, 4.61 (2m, 2H, H3 ); 6.36 (m, 2H, H1 ); 6.81 (m, 8H, H-C(ar)); 6.89, 6.95 (2d, br, 2H, H5); 7.22-7.37 (m,
1.15.8 Supplement 27
Current Protocols in Nucleic Acid Chemistry
18H, H-C(ar)); 8.30 (m, 2H, H6); 8.78 (s, br, 2H, NH). 31 P NMR (121 MHz, CDCl3 ): δ 151.23, 151.49. FTICR ESI-MS (m/z): calcd. for C42 H52 N5 O8 PSe [M+H]+ 866.28022, found 866.27866.
PREPARATION OF 2 -DEOXY-2 -METHYLSELENOADENOSINE PHOSPHORAMIDITE
ALTERNATE PROTOCOL 2
This protocol describes the synthesis of N6 -acetyl-5 -O-(4,4 -dimethoxytrityl)-2 deoxy-2 -methylselenoadenosine 3 -O-(2-cyanoethyl)-N,N-diisopropylphosphoramidite (S.17). The synthesis starts from the precursor 5 -O-(4,4 -dimethoxytrityl)-3 -O{[(triisopropylsilyl)oxy]methyl}adenosine S.11 (Pitsch et al., 2001; also see UNITS 2.9 & 3.8) and proceeds in six steps: (1) triflation of the 2 -hydroxyl group to give S.12, (2) SN 2-displacement of the 2 -O-triflate ester by potassium trifluoroacetate to produce S.13, (3) 2 -OH activation to S.14, (4) SN 2-displacement of the 2 -O-triflate ester by sodium methylselenide to give S.15, (5) fluoride-assisted removal of the 3 OTOM group to give S.16, and (6) 3 -OH phosphitylation using 2-cyanoethyl-N,Ndiisopropylchlorophosphoramidite to give S.17. These steps are illustrated in Figure 1.15.3.
Figure 1.15.3 Preparation of a 2 -deoxy-2 -methylselenoadenosine phosphoramidite building block for RNA solid-phase synthesis (see Alternate Protocol 2). DMAP, 4-(N,N-dimethylamino)pyridine; DMTr, 4,4 -dimethoxytrityl; TBAF, tetra-nbutylammonium fluoride; Tf, trifluoromethanesulfonyl; THF, tetrahydrofuran; TOM, [(triisopropylsilyl)oxy]methyl. Synthesis of Modified Nucleosides
1.15.9 Current Protocols in Nucleic Acid Chemistry
Supplement 27
Additional Materials (also see Basic Protocol 1) 5 -O-(4,4 -Dimethoxytrityl)-3 -O-{[(triisopropylsilyl)oxy]methyl}adenosine (S.11; Pitsch et al., 2001; UNIT 2.9) 4-(N,N-Dimethylamino)pyridine (DMAP; Fluka) Trifluoromethanesulfonyl chloride (Tf-Cl; Fluka) Saturated sodium bicarbonate solution in water Anhydrous toluene N-Ethyldiisopropylamine (Fluka) Potassium trifluoroacetate (Fluka) 18-Crown-6-ether (Fluka) Tetrabutylammonium fluoride trihydrate (TBAF; Fluka) Acetic acid (AcOH) 25- and 100-mL one-neck round-bottom flasks equipped with rubber septa Activate 2 -hydroxyl group 1. In a 100-mL one-neck flask equipped with a stir bar, a rubber septum, and an argon balloon, dissolve 1.46 g (1.82 mmol) S.11, 334 mg (2.74 mmol) DMAP, and 635 µL (4.56 mmol) TEA in 35 mL dry CH2 Cl2 . 2. Cool the stirred solution to 0◦ C, then slowly add 289 µL (2.74 mmol) Tf-Cl. Continue stirring for 20 min at room temperature. 3. Dilute the reaction mixture with 250 mL CH2 Cl2 and wash with 250 mL saturated sodium bicarbonate solution. 4. Dry over 40 g Na2 SO4 , remove the solid using a glass-fritted B¨uchner funnel and filter paper, and evaporate the solvents. 5. Purify the residue on a chromatography column (APPENDIX 3E) containing 35 g silica gel, using a solvent system of 0.5% (v/v) MeOH in CH2 Cl2 . 6. Combine the fractions containing product S.12, as determined by TLC (APPENDIX 3D) using 94:6 (v/v) CH2 Cl2 /MeOH. Evaporate and then coevaporate with 15 mL CH2 Cl2 . 7. Dry the residue in vacuo to obtain the product as a colorless foam. N6 -Acetyl-5 -O-(4,4 -dimethoxytrityl)-2 -O-trifluoromethanesulfonyl-3 -O-{[(triisopropylsilyl)oxy]methyl}adenosine (S.12). Yield: 1.02 g (60%). TLC (94:6 CH2 Cl2 /MeOH): Rf 0.51. 1 H NMR (300 MHz, CDCl3 ): δ 1.02 (m, 21H, (iPr)3 Si); 2.65 (s, 3H, COCH3 ); 3.40 (dd, J = 3.9, 10.8 Hz, 1H, H5 a); 3.62 (dd, J = 2.7, 10.8 Hz, 1H, H5 b); 3.79 (s, 6H, 2 × OCH3 ); 4.51 (m, 1H, H4 ); 4.89 (m, 1H, H3 ); 4.96 (d, J = 5.0 Hz, 1H, OCH2 O); 5.11 (d, J = 5.0 Hz, 1H, OCH2 O); 6.21 (m, 1H, H2 ); 6.36 (d, J = 5.1 Hz, 1H, H1 ); 6.80 (d, J = 8.7 Hz, 4H, H-C(ar)); 7.28 (m, 7H, H-C(ar)); 7.40 (m, 2H, H-C(ar)); 8.10 (s, 1H, H8); 8.57 (s, 1H, H2); 8.68 (s, 1H, HN6 ). 13 C NMR (75 MHz, CDCl3 ): δ 11.8 (SiCH(CH3 )2 ); 17.7 (SiCH(CH3 )2 ); 25.6 (COCH3 ); 55.1 (OCH3 ); 62.6 (C5 ); 74.4 (C3 ); 83.3 (C4 ); 84.2 (C2 ); 85.6 (C1 ); 86.9; 89.9 (OCH2 O); 113.2, 122.0, 127.0, 127.8, 128.1, 130.0, 130.1, 135.2, 135.3 (C(ar)); 141.4 (C8); 144.2, 149.4, 150.8 (C(ar)); 152.6 (C2); 158.7 (C(ar)); 170.3 (COCH3 ). ESI-MS (m/z): [M+H]+ calcd. for C44 H54 F3 N5 O10 SSi, 930.1, found 929.5.
Perform inversion of configuration at C2 8. In a 100-mL two-neck flask equipped with a stir bar, a rubber septum, a reflux condenser, and an argon balloon, dissolve 1.02 g (1.10 mmol) S.12 in 50 mL dry toluene. 2 -MethylselenoModified Phosphoramidites and RNA
9. Add 470 µL (2.74 mmol) N-ethyldiisopropylamine, 1.39 g (9.12 mmol) potassium trifluoroacetate, and 965 mg (3.65 mmol) 18-crown-6-ether, and stir the reaction mixture 11 hr at 80◦ C.
1.15.10 Supplement 27
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10. Evaporate the solvents, dissolve the residue in 200 mL CH2 Cl2 , and wash with 150 mL of half-saturated sodium bicarbonate. 11. Dry over 30 g Na2 SO4 , remove the solid using a glass-fritted B¨uchner funnel and filter paper, and evaporate again. 12. Purify the residue on a 40-g silica gel column using a solvent system of 0.2% to 1.0% (v/v) MeOH in CH2 Cl2 . 13. Combine the fractions containing product S.13, as determined by TLC using 92:8 (v/v) CH2 Cl2 /MeOH. Evaporate and then coevaporate with 10 mL CH2 Cl2 . 14. Dry the residue in vacuo to obtain the product as sligthly yellow foam. N6 -Acetyl-5 -O-(4,4 -dimethoxytrityl)-3 -O-[(triisopropylsilyl)oxy]methyl-(-D-arabinofuranosyl)adenine (S.13). Yield: 0.79 g (89%). TLC (92:8 CH2 Cl2 /MeOH): Rf 0.50. 1 H NMR (300 MHz, CDCl3 ): δ 1.09 (m, 21H, (iPr)3 Si); 2.63 (s, 3H, COCH3 ); 3.42 (dd, J = 3.5, 10.7 Hz, 1H, H5 a); 3.66 (dd, J = 2.6, 10.7 Hz, 1H, H5 b); 3.80 (s, 6H, 2 × OCH3 ); 4.25 (m, 1H, H4 ); 4.41 (m, 2H, H3 , 2 -OH); 4.48 (m, 1H, H2 ); 4.96 (d, J = 5.3 Hz, 1H, OCH2 O); 5.05 (d, J = 5.3 Hz, 1H, OCH2 O); 6.42 (d, J = 3.9 Hz, 1H, H1 ); 6.84 (d, J = 8.4 Hz, 4H, H-C(ar)); 7.27 (m, 7H, H-C(ar)); 7.42 (d, J = 6.9 Hz, 2H, H-C(ar)); 8.43 (s, 1H, H8); 8.66 (s, 1H, H2); 8.71 (s, 1H, HN6 ). 13 C NMR (75 MHz, CDCl3 ): δ 11.9 (SiCH(CH3 )2 ); 17.7 (SiCH(CH3 )2 ); 25.5 (COCH3 ); 55.2 (OCH3 ); 63.2 (C5 ); 74.8 (C2 ); 81.9 (C4 ); 82.8 (C3 ); 84.9 (C1 ); 87.6; 89.6 (OCH2 O); 113.3, 121.4, 127.1, 128.0, 128.2, 130.07, 130.12, 135.0, 135.2 (C(ar)); 142.7 (C8); 143.8, 149.0, 151.2 (C(ar)); 152.2 (C2); 158.7 (C(ar)); 170.3 (COCH3 ). UV (MeOH): λmax 271 nm. ESI-MS (m/z): [M+H]+ calcd. for C43 H55 N5 O8 Si, 798.0, found 797.5.
Activate 2 -hydroxyl group 15. In a 50-mL one-neck flask equipped with a stir bar, a rubber septum, and an argon balloon, dissolve 0.45 g (0.56 mmol) S.13, 104 mg (0.86 mmol) DMAP, and 196 µL (1.42 mmol) TEA in 12 mL of dry CH2 Cl2 . 16. Cool the stirred solution to 0◦ C, slowly add 90 µL (0.86 mmol) Tf-Cl, and continue stirring for 20 min at room temperature. 17. Dilute the reaction mixture with 120 mL CH2 Cl2 and wash with 80 mL saturated sodium bicarbonate solution. 18. Dry over 15 g Na2 SO4 , remove the solid using a glass-fritted B¨uchner funnel and filter paper, and evaporate the solvent. The product can be used without further purification for the next step. For analysis, the product was purified by silica gel column chromatography using a solvent system of 0.1% to 0.6% (v/v) MeOH in CH2 Cl2 . N6 -Acetyl-5-O-(4,4-dimethoxytrityl)-2 -O-trifluoromethanesulfonyl-3 -O-[(triisopropylsilyl)oxy]methyl-(β-D-arabinofuranosyl)adenine (S.14). Yield: 0.51 g (98%). TLC (94:6 CH2 Cl2 /MeOH): Rf 0.51. 1 H NMR (300 MHz, CDCl3 ): δ 1.07 (m, 21H, (iPr)3 Si); 2.65 (s, 3H, COCH3 ); 3.52 (m, 2H, H5 ); 3.81 (s, 6H, 2 × OCH3 ); 4.34 (m, 1H, H4 ); 4.72 (m, 1H, H3 ); 5.04 (s, 2H, OCH2 O); 5.53 (m, 1H, H2 ); 6.60 (d, J = 3.3 Hz, 1H, H1 ); 6.85 (d, J = 8.7 Hz, 4H, H-C(ar)); 7.29 (m, 7H, H-C(ar)); 7.48 (d, J = 7.2 Hz, 2H, H-C(ar)); 8.10 (s, 1H, H8); 8.62 (s, 1H, HN6 ); 8.67 (s, 1H, H2). 13 C NMR (75 MHz, CDCl3 ): δ 11.8 (SiCH(CH3 )2 ); 17.7 (SiCH(CH3 )2 ); 25.6 (COCH3 ); 55.2 (OCH3 ); 62.7 (C5 ); 80.5 (C4 ); 82.9 (C3 ); 83.2 (C1 ); 86.3 (C2 ); 86.6; 89.5 (OCH2 O); 113.2, 121.2, 126.9, 127.9, 128.0, 130.0, 135.49, 135.53, (C(ar)); 141.1 (C8); 144.4, 149.2, 150.5 (C(ar)); 152.6 (C2); 158.7 (C(ar)); 170.2 (COCH3 ). UV (MeOH): λmax 270 nm. ESI-MS (m/z): [M+H]+ calcd. for C44 H54 F3 N5 O10 SSi 930.1, found 929.5. Synthesis of Modified Nucleosides
1.15.11 Current Protocols in Nucleic Acid Chemistry
Supplement 27
Introduce 2 -methylseleno group 19. Apply Basic Protocol 1, steps 15 to 24, for 1.79 g (1.89 mmol) S.14 in place of S.2. 20. Dilute with 200 mL water and extract the product three times with 100 mL CH2 Cl2 . 21. Wash the combined CH2 Cl2 phases with 200 mL of 0.2 M TEAA solution and then with 200 mL saturated NaCl solution. 22. Dry the organic phase with 30 g Na2 SO4 , filter using a glass-fritted B¨uchner funnel and filter paper, and evaporate the filtrate to dryness. 23. Purify the residue on a 50-g silica gel column using a solvent system of 0.2% to 1.0% (v/v) MeOH in CH2 Cl2 . 24. Combine the fractions containing product S.15, as determined by TLC using 94:6 (v/v) CH2 Cl2 /MeOH. Evaporate and then coevaporate with 10 mL CH2 Cl2 . 25. Dry the residue in vacuo to obtain the product as a colorless foam. N6 -Acetyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxy-2 -methylseleno-3 -O-{[(triisopropylsilyl)oxy]methyl}adenosine (S.15). Yield: 0.81 g (49%). TLC (94:6 CH2 Cl2 /MeOH): Rf 0.45. 1 H NMR (300 MHz, CDCl3 ): δ 1.08 (m, 21H, (iPr)3 Si); 1.69 (s, 3H, SeCH3 ); 2.64 (s, 3H, COCH3 ); 3.44 (dd, J = 4.4, 10.4 Hz, 1H, H5 a); 3.50 (dd, J = 4.2, 10.4 Hz, 1H, H5 b); 3.80 (s, 6H, 2 × OCH3 ); 4.38 (m, 1H, H2 ); 4.51 (m, 1H, H4 ); 4.56 (m, 1H, H3 ); 5.01 (d, J = 5.1 Hz, 1H, OCH2 O); 5.09 (d, J = 5.1 Hz, 1H, OCH2 O); 6.36 (d, J = 8.4 Hz, 1H, H1 ); 6.81 (d, J = 8.4 Hz, 4H, H-C(ar)); 7.28 (m, 7H, H-C(ar)); 7.46 (d, J = 6.6 Hz, 2H, H-C(ar)); 8.11 (s, 1H, H8); 8.56 (s, 1H, H2); 8.62 (s, 1H, HN6 ). 13 C NMR (75 MHz, CDCl3 ): δ 3.5 (SeCH3 ); 11.9 (SiCH(CH3 )2 ); 17.8 (SiCH(CH3 )2 ); 25.6 (COCH3 ); 45.0 (C2 ); 55.1 (OCH3 ); 63.6 (C5 ); 79.8 (C3 ); 84.5 (C4 ); 86.6; 89.8 (OCH2 O); 91.1 (C1 ); 113.1, 122.2, 126.9, 127.8, 128.2, 130.07, 130.10, 135.66, 135.70 (C(ar)); 142.0 (C8); 144.6, 149.2, 151.3 (C(ar)); 152.3 (C2); 158.6 (C(ar)); 170.3 (COCH3 ). UV (MeOH): λmax 276 nm. ESI-MS (m/z): [M+H]+ calcd. for C44 H57 N5 O7 SeSi 875.0, found 875.5.
Deprotect 3 -hydroxyl group 26. In a 25-mL one-neck flask equipped with a stir bar, a rubber septum, and an argon balloon, dissolve 0.68 g (0.78 mmol) S.15 in 3.4 mL of 1 M TBAF/0.5 M AcOH in THF and stir for 2.5 hr at room temperature. 27. Evaporate the solvents and purify the residue on a 40-g silica gel column using a solvent system of 0.5% to 2.0% (v/v) MeOH in CH2 Cl2 . 28. Combine the fractions containing product S.16, as determined by TLC using 92:8 (v/v) CH2 Cl2 /MeOH. Evaporate and then coevaporate with 5 mL CH2 Cl2 . 29. Dry the residue in vacuo to obtain the product as colorless foam.
2 -MethylselenoModified Phosphoramidites and RNA
N6 -Acetyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxy-2 -methylselenoadenosine (S.16). Yield: 0.44 g (82%). TLC (92:8 CH2 Cl2 /MeOH): Rf 0.49. 1 H NMR (300 MHz, CDCl3 ): δ 1.95 (s, 3H, SeCH3 ); 2.63 (s, 3H, COCH3 ); 2.90 (s, br, 1H, 3 -OH); 3.45 (dd, J = 4.2, 10.5 Hz, 1H, H5 a); 3.53 (dd, J = 4.4, 10.5 Hz, 1H, H5 b); 3.80 (s, 6H, 2 OCH3 ); 4.34 (m, 1H, H4 ); 4.41 (m, 1H, H2 ); 4.50 (m, 1H, H3 ); 6.22 (d, J = 8.7 Hz, 1H, H1 ); 6.82 (d, J = 8.7 Hz, 4H, H-C(ar)); 7.28 (m, 7H, H-C(ar)); 7.45 (d, J = 1.5 Hz, 2H, H-C(ar)); 8.12 (s, 1H, H8); 8.58 (s, 1H, H2); 8.79 (s, 1H, HN6 ). 13 C NMR (75 MHz, CDCl3 ): δ 4.5 (SeCH3 ); 25.6 (COCH3 ); 48.8 (C2 ); 55.2 (OCH3 ); 63.6 (C5 ); 72.6 (C3 ); 85.1 (C4 ); 86.7; 88.8 (C1 ); 113.2, 122.2, 127.0, 127.8, 128.1, 130.0, 135.7 (C(ar)); 141.9 (C8); 144.5, 149.2, 151.3 (C(ar)); 152.3 (C2); 158.6 (C(ar)); 170.4 (COCH3 ). UV (MeOH): λmax 271 nm. FTICR ESI-MS (m/z): [M+H]+ calcd. for C34 H35 N5 O6 Se 690.18341, found 690.18238.
1.15.12 Supplement 27
Current Protocols in Nucleic Acid Chemistry
Prepare phosphoramidite 30. Apply Basic Protocol 1, steps 31 to 40, to 0.66 g (0.96 mmol) S.16. For column purification of S.17, use 14 g silica gel and a solvent system of 50% to 70% (v/v) ethyl acetate/1% (v/v) TEA in hexanes. N6 -Acetyl-5-O-(4,4-dimethoxytrityl)-2 -deoxy-2-methylselenoadenosine 3 -O-(2-cyanoethyl)-N,N-diisopropylphosphoramidite (S.17). Yield: 0.78 g (91%; mixture of diastereoisomers). TLC (94:6 CH2 Cl2 /MeOH): Rf 0.44. 1 H NMR (500 MHz, CDCl3 ): δ 1.09-1.29 (m, 24H, ((CH3 )2 CH)2 N); 1.61 (s, 3H, SeCH3 ); 1.69 (s, 3H, SeCH3 ); 2.34 (m, 2H, CH2 CN); 2.66 (m, 8H, COCH3 , CH2 CN); 3.38 (m, 2H, H5 a); 3.47-3.75 (m, 8H, ((CH3 )2 CH)2 N, H5 b, POCH2 ); 3.78, 3.79 (2s, 12H, OCH3 ); 3.90-3.99, 4.09-4.22 (2m, 2H, POCH2 ); 4.41 (m, 4H, H2 , H4 ); 4.67, 4.73 (2m, 2H, H3 ); 6.31 (m, 2H, H1 ); 6.80, 7.22-7.45 (m, 26H, H-C(ar)); 8.11 (s, 1H, H8); 8.15 (s, 1H, H8); 8.49 (s, br, 2H, HN6 ); 8.54, 8.55 (2s, 2H, H2). 31 P NMR (121 MHz, CDCl3 ): δ 150.8, 152.4. UV (MeOH): λmax 271 nm. FTICR ESI-MS (m/z): [M+H]+ calcd. for C43 H52 N7 O7 PSe 890.29147, found 890.29005.
PREPARATION OF 2 -DEOXY-2 -METHYLSELENOGUANOSINE PHOSPHORAMIDITE
ALTERNATE PROTOCOL 3
This protocol describes the synthesis of N2 -acetyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxy2 -methylselenoguanosine 3 -O-(2-cyanoethyl)-N,N-diisopropylphosphoramidite (S.27). The synthesis starts from 9-[β-D-arabinofuranosyl]guanine (S.18) and proceeds in nine steps: (1) protection of the 3 - and 5 -hydroxyl groups to give S.19, (2) N2 - and 2 O-acetylation to give S.20, (3) O6 -protection with 2-(4-nitrophenyl)ethyl group under Mitsunobu conditions to give S.21, (4) removal of the 2 -O-acetyl group by selective ester cleavage to give S.22, (5) triflation of the 2 -OH to give S.23, (6) SN 2displacement of the 2 -O-triflate ester by sodium methylselenide to produce S.24, (7) fluoride-assisted removal of the O6 -, 3 -OH, and 5 -OH protecting groups to give S.25, (8) 5 -O-tritylation to give S.26, and (9) 3 -O-phosphitylation using 2-cyanoethyl-N,Ndiisopropylchlorophosphoramidite to give S.27. These steps are illustrated in Figure 1.15.4.
Additional Materials (also see Basic Protocol 1) 9-[β-D-Arabinofuranosyl]guanine (S.18; Metkinen Oy) 1,3-Dichloro-1,1,3,3-tetraisopropyldisiloxane (TIPSCl2 ; Fluka) Acetic anhydride (Fluka) Triphenylphosphine (PPh3 ; Fluka) 2-(4-Nitrophenyl)ethanol (NPE-OH; Fluka) Anhydrous dioxane Diisopropyl azodicarboxylate (DIAD; Aldrich) Saturated sodium bicarbonate solution in water 0.5 M sodium hydroxide (NaOH) solution in water Acetic acid 4-(N,N-Dimethylamino)pyridine (DMAP; Fluka) Trifluoromethanesulfonyl chloride (Tf-Cl; Fluka) Tetrabutylammonium fluoride trihydrate (TBAF; Fluka) 250- and 500-mL one-neck round-bottom flasks 500-, 1000-, and 2000-mL separatory funnels Protect 3 - and 5 -hydroxyl groups 1. In a 500-mL one-neck flask, coevaporate 4.16 g (14.7 mmol) S.18 three times with 30 mL dry pyridine. Then, suspend in 160 mL DMF and 14 mL pyridine. 2. Add 4.86 g (15.4 mmol) TIPSCl2 dropwise and keep the reaction mixture stirring 16 hr at room temperature.
Synthesis of Modified Nucleosides
1.15.13 Current Protocols in Nucleic Acid Chemistry
Supplement 27
Figure 1.15.4 Preparation of a 2 -deoxy-2 -methylselenoguanosine phosphoramidite building block for RNA solidphase synthesis (see Alternate Protocol 3). DIAD, diisopropyl azodicarboxylate; DMAP, 4-(N,N-dimethylamino)pyridine; DMF, N,N-dimethylformamide; DMTr, 4,4 -dimethoxytrityl; NPE, 2-(4-nitrophenyl)ethyl; PPh3 , triphenylphosphine; TBAF, tetra-n-butylammonium fluoride; Tf, trifluoromethanesulfonyl; THF, tetrahydrofuran; TIPSCl2 , 1,3-dichloro-1,1,3,3tetraisopropyldisiloxane. 2 -MethylselenoModified Phosphoramidites and RNA
1.15.14 Supplement 27
Current Protocols in Nucleic Acid Chemistry
3. Precipitate the reaction mixture by pouring into 2.6 L ice-water under vigorous stirring. 4. Collect the crude product on a glass-fritted B¨uchner funnel with filter paper, and dry the residue in vacuo. The dried residue is used in the next step without further purification. For analysis, a small portion of the crude product was crystallized from ethanol. 9-[3 ,5 -O-(1,1,3,3-Tetraisopropyldisiloxane-1,3-diyl)-β-D-arabinofuranosyl]guanine (S.19). Yield: 7.60 g (98%). TLC (94:6 CH2 Cl2 /MeOH): Rf 0.65. 1 H NMR (300 MHz, DMSO-d6 ): δ 1.06 (m, 28H, (iPr)4 Si2 O); 3.74 (m, 1H, H4 ); 3.95 (m, 2H, H5 a and H5 b); 4.30 (m, 1H, H3 ); 4.42 (m, 1H, H2 ); 5.78 (d, J = 5.9 Hz, 1H, 2 -OH); 5.95 (d, J = 6.4 Hz, 1H, H1 ); 6.43 (s, 2H, NH2 ); 7.60 (s, 1H, H8); 10.57 (s, 1H, HN1 ). 13 C NMR (75 MHz, DMSO-d6 ): δ 11.9, 12.3, 12.4, 12.8, 16.7, 16.8, 16.9, 17.1, 17.11, 17.13, 17.3 (iPr)4 Si2 O); 61.2 (C5 ); 74.5 (C2 ); 75.3 (C3 ); 79.4 (C4 ); 80.5 (C1 ); 115.8 (C(ar)); 135.9 (C8); 151.2, 153.6, 156.7 (C(ar)). UV (MeOH): λmax 252 nm. ESI-MS (m/z): calcd. for C22 H39 N5 O6 Si2 [M+H]+ 526.76, found 526.10; calcd. [M+Na]+ 548.74, found 548.17.
Acetylate 2 -hydroxyl and exocyclic amino group 5. In a 500-mL one-neck flask, coevaporate S.19 three times with 30 mL pyridine. 6. Suspend the residue in 80 mL DMF, 80 mL pyridine, and 80 mL acetic anhydride under an argon atmosphere. Stir the reaction mixture for 16 hr at 80◦ C. 7. Remove the solvents under vacuum and partition the brown viscous oil between 500 mL CH2 Cl2 and 500 mL aqueous 5% citric acid. 8. Separate the organic layer, and extract the aqueous layer four times with 700 mL CH2 Cl2 . 9. Wash the combined organic layers with 600 mL saturated aqueous NaCl. 10. Dry the organic phase with 50 g Na2 SO4 , filter using a glass-fritted B¨uchner funnel and filter paper, and evaporate the filtrate to dryness. 11. Purify the residue on a chromatography column containing 200 g silica gel, using a solvent system of 1.0% to 8.0% (v/v) MeOH in CH2 Cl2 . 12. Combine the fractions containing product S.20, as determined by TLC using 9:1 (v/v) CH2 Cl2 /MeOH. Evaporate and then coevaporate with 40 mL CH2 Cl2 . 13. Dry the residue in vacuo to obtain the product as a colorless foam. A mixture of N2 -acetyl-9-[2 -O-acetyl-3 ,5 -O-(1,1,3,3-tetraisopropyldisiloxane-1,3diyl)-β-D-arabinofuranosyl]guanine (S.20a) and the N2 ,N2 -diacetyl byproduct (S.20b) is obtained and can be used in the next reaction step. Yield: 2.91 g S.20a; 5.20 g S.20a + S.20b (64%). TLC (9:1 CH2 Cl2 /MeOH): Rf = 0.57 (S.20b), 0.63 (S.20a). The higher Rf value of S.20a, though unexpected, is correct. For S.20a: 1 H NMR (300 MHz, CDCl3 ): δ 0.95-1.05 (m, 28H, (iPr)4 Si2 O); 1.75 (s, 3H, COCH3 ); 2.30 (s, 3H, NHCOCH3 ); 3.87 (m, 1H, H4 ); 4.08 (m, 2H, H5 a and H5 b); 4.64 (m, 1H, H3 ); 5.49 (m, 1H, H2 ); 6.26 (d, J = 6.4 Hz, 1H, H1 ); 7.97 (s, 1H, H8); 9.58 (s, 1H, HN1 ); 12.03 (s, 1H, H2). 13 C NMR (75 MHz, CDCl3 ): δ 12.4, 12.8, 13.0, 13.3, 16.70, 16.72, 16.8, 16.9, 17.26, 17.34, 17.4 (iPr)4 Si2 O); 20.1 (COCH3 ); 24.3 (NHCOCH3 ); 60.6 (C5 ); 71.1 (C3 ); 76.4 (C2 ); 80.4 (C4 ); 80.4 (C1 ); 120.8 (C(ar)); 137.6 (C8); 147.5, 148.1, 155.7 (C(ar)); 169.8, 172.1 (2 × COCH3 ). UV (MeOH): λmax 256 nm. ESI-MS (m/z): calcd. for C26 H43 N5 O8 Si2 [M+H]+ 610.82, found 610.10; calcd. [M+Na]+ 632.81, found 632.26. Synthesis of Modified Nucleosides
1.15.15 Current Protocols in Nucleic Acid Chemistry
Supplement 27
Protect O6 group 14. In a 250-mL one-neck flask, coevaporate 4.65 g (7.48 mmol) S.20, 2.94 g (11.2 mmol) triphenylphosphine, and 1.88 g (11.2 mmol) 2-(4-nitrophenyl)ethanol with 30 mL dry dioxane. 15. Suspend in 150 mL dioxane and stir for 30 min at room temperature. 16. Add 2.36 mL (12.0 mmol) DIAD and continue stirring for 45 min. 17. Remove the solvents under vacuum and partition the residue between 500 mL CH2 Cl2 and 500 mL saturated sodium bicarbonate solution. 18. Separate the organic layer, and extract the aqueous layer two times with 700 mL CH2 Cl2 . 19. Dry the combined organic layers with 40 g Na2 SO4 , filter using a glass-fritted B¨uchner funnel and filter paper, and evaporate the filtrate to dryness. 20. Purify the residue on a 120-g silica gel column using a solvent system of 10% to 50% (v/v) EtOAc in hexanes. 21. Combine the fractions containing product S.21, as determined by TLC using 2:8 hexanes/EtOAc. Evaporate and then coevaporate with 30 mL CH2 Cl2 . 22. Dry the residue in vacuo to obtain the product as a colorless foam. A mixture of N2 -acetyl-9-[2 -O-acetyl-3 ,5 -O-(1,1,3,3-tetraisopropyldisiloxane-1,3diyl)-β-D-arabinofuranosyl]-O6 -[2-(4-nitrophenyl)ethyl]guanine (S.21a) and the N2 ,N2 -diacetyl byproduct (S.21b) is used in the next step. Yield: 3.63 g S.21 as yellowish foam (64%). TLC (2:8 hexanes/EtOAc): Rf = 0.55 (S.21a), 0.72 (S.21b). For S.21a: 1 H NMR (300 MHz, CDCl3 ): δ 0.98-1.25 (m, 28H, (iPr)4 Si2 O); 1.71 (s, 3H, COCH3 ); 2.56 (s, 3H, NHCOCH3 ); 3.31 (t, J = 6.8 Hz, 2H, CH2 -C6 H4 -NO2 ); 3.90 (m, 1H, H4 ); 4.10 (m, 2H, H5 a and H5 b); 4.69 (m, 1H, H3 ); 4.74 (m, 2H, O6 -CH2 ); 5.56 (m, 1H, H2 ); 6.41 (d, J = 6.2 Hz, 1H, H1 ); 7.50 (d, J = 8.5 Hz, 2H, 4-nitrophenyl H2/H6); 7.84 (s, 1H, HN2 ); 8.09 (s, 1H, H8); 8.18 (d, J = 8.5 Hz, 2H, 4-nitrophenyl H3/H5). 13 C NMR (75 MHz, CDCl3 ): δ 12.4, 12.97, 13.03, 13.4, 16.7, 16.8, 16.9, 17.26, 17.32, 17.39, 17.42 ((iPr)4 Si2 O); 20.0 (COCH3 ); 25.1 (NHCOCH3 ); 35.0 (CH2 -C6 H4 NO2 ); 60.5 (C5 ); 66.9 (O6 -CH2 ); 71.3 (C3 ); 76.4 (C2 ); 80.5 (C4 ); 80. 6 (C1 ); 117.3 (C(ar)); 123.8 (4-nitrophenyl C2/C6); 129.9 (4-nitrophenyl C3/C5); 140.3 (C8); 145.5, 147.0, 152.2, 152. 8, 160.5 (C(ar)); 169.3, 172.0 (COCH3 ). UV (MeOH): λmax 269 nm. ESI-MS (m/z): calcd. for C34 H5 0N6 O10 Si2 [M+H]+ 759.97, found 759.26; calcd. for [M+Na]+ 781.96, found 781.42. For S.21b: 1 H NMR (300 MHz, CDCl3 ): δ 0.99-1.17 (m, 28H, (iPr)4 Si2 O); 1.69 (s, 3H, COCH3 ); 2.27 (s, 6H, 2 × COCH3 ); 3.30 (t, J = 6.9 Hz, 2H, CH2 -C6 H4 -NO2 ); 3.91 (m, 1H, H4 ); 4.10 (m, 2H, H5 a and H5 b); 4.78 (m, 1H, H3 ); 4.79 (m, 2H, O6 -CH2 ); 5.55 (dd, J = 6.4, 8.0 Hz, 1H, H2 ); 6.44 (d, J = 6.4 Hz, 1H, H1 ); 7.46 (d, J = 8.7 Hz, 2H, 4-nitrophenyl H2/H6); 7.84 (s, 1H, HN2 ); 8.15 (d, J = 8.7 Hz, 2H, 2H, 4-nitrophenyl H3/H5); 8.26 (s, 1H, H8). 13 C NMR (75 MHz, CDCl3 ): δ 12.4, 12.97, 12.99, 13.3, 16.7, 16.8, 16.9, 17.26, 17.33, 17.38, 17.42 ((iPr)4 Si2 O); 19.9 (COCH3 ); 26.0 (2 × COCH3 ); 35.0 (CH2 -C6 H4 -NO2 ); 61.1 (C5 ); 67.3 (O6 -CH2 ); 72.0 (C3 ); 76.7 (C2 ); 80.8 (C4 ); 81.3 (C1 ); 120.4 (C(ar)); 123.8 (4-nitrophenyl C2/C6); 129.9 (4-nitrophenyl C3/C5); 142.7 (C8); 145.3, 147.0, 152.5, 152.9, 161.3 (C(ar)); 169.4, 171.9 (COCH3 ). UV (MeOH): λmax 260 nm. ESI-MS (m/z): calcd. for C36 H52 N6 O11 Si2 [M+H]+ 802.00, found 801.07; calcd. for [M+Na]+ 823.99, found 823.31.
2 -MethylselenoModified Phosphoramidites and RNA
1.15.16 Supplement 27
Deprotect 2 -hydroxyl group 23. In a 250-mL one-neck flask, dissolve 3.41 g (4.49 mmol) S.21 in 130 mL of 5:4 (v/v) THF/MeOH and cool the stirred solution to 0◦ C. 24. Add 150 mL of 0.5 M NaOH under vigorous stirring for 7 min. 25. Add 160 mL of 0.5 M acetic acid under vigorous stirring. Current Protocols in Nucleic Acid Chemistry
26. Evaporate the solvents. Dissolve the residue in 400 mL CH2 Cl2 and wash with 400 mL water. 27. Dry the combined organic layers with 20 g Na2 SO4 , filter using a glass-fritted B¨uchner funnel and filter paper, and evaporate the filtrate to dryness. 28. Purify the residue on a 240-g silica gel column using a solvent system of 1.0% to 2.0% (v/v) MeOH in CH2 Cl2 . 29. Combine the fractions containing product S.22, as determined by TLC using 2:8 (v/v) hexanes/EtOAc. Evaporate and then coevaporate with 20 mL CH2 Cl2 . 30. Dry the residue in vacuo to obtain the product as a colorless foam. N2-Acetyl-9-[3 ,5 -O-(1,1,3,3-tetraisopropyldisiloxane-1,3-diyl)-β-D-arabinofuranosyl]O6 -[2-(4-nitrophenyl)ethyl]guanine (S.22). Yield: 1.84 g (57%). TLC (2:8 hexanes/EtOAc): Rf = 0.46. 1 H NMR (300 MHz, CDCl3 ): δ 1.01-1.14 (m, 28H, (iPr)4 Si2 O); 2.37 (s, 3H, COCH3 ); 2.29 (t, J = 6.8 Hz, 2H, CH2 -C6 H4 -NO2 ); 3.86 (m, 1H, H4 ); 4.05 (m, 2H, H5 a and H5 b); 4.51 (t, J = 7.3 Hz, 1H, H3 ); 4.30 (s, 1H, 2 -OH); 4.64(m, 1H, H2 ); 4.74 (m, 2H, O6 -CH2 ); 6.14 (d, J = 5.7 Hz, 1H, H1 ); 7.49 (d, J = 8.7 Hz, 2H, 4-nitrophenyl H2/H6); 8.07 (s, 1H, HN2 ); 8.14 (s, 1H, H8); 8.16 (d, J = 8.7 Hz, 2H, 4-nitrophenyl H3/H5). 13 C NMR (75 MHz, CDCl3 ): δ 12.5, 13.0, 13.1, 13.5, 16.89, 16.95, 17.0, 17.3, 17.4, 17.5 (iPr)4 Si2 O); 24.9 (COCH3 ); 35.0 (CH2 -C6 H4 -NO2 ); 61.5 (C5 ); 67.0 (O6 -CH2 ); 74.6 (C3 ); 76.7 (C2 ); 81.7 (C4 ); 83.8 (C1 ); 117.7 (C(ar)); 123.8 (4-nitrophenyl C2/C6); 129.9 (4-nitrophenyl C3/C5); 141.4 (C8); 145.4, 147.0, 151.6, 152.5, 160.2 (C(ar)), 169.9 (COCH3 ). UV (MeOH): λmax 260 nm. ESI-MS (m/z): calcd. for C32 H48 N6 O9 Si2 [M+H]+ 717.93, found 717.11; calcd. for [M+Na]+ 739.92, found 739.29.
Activate 2 -hydroxyl group 31. In a 250-mL one-neck flask, coevaporate 0.60 g (0.837 mmol) S.22 with 10 mL dry pyridine and dissolve in 100 mL CH2 Cl2 . 32. Add 153 mg (1.26 mmol) DMAP and 169 mg (233 µL, 1.67 mmol) TEA. 33. Cool the stirred solution to 0◦ C, add 211 mg (158 µL, 1.26 mmol) Tf-Cl, and continue stirring for 2 hr at room temperature. 34. Dilute the reaction mixture with 300 mL CH2 Cl2 and wash with 300 mL saturated sodium bicarbonate solution. 35. Dry over 10 g Na2 SO4 , filter using a glass-fritted B¨uchner funnel and filter paper, and evaporate the solvent. 36. Purify the residue on a 34-g silica gel column using a solvent system of 10% to 50% (v/v) EtOAc in hexanes. 37. Combine the fractions containing product S.23, as determined by TLC using 1:1 (v/v) hexanes/EtOAc. Evaporate and then coevaporate with 5 mL CH2 Cl2 . 38. Dry the residue in vacuo to obtain the product as a colorless foam. N2-Acetyl-9-{2-O-[(trifluoromethyl)sulfonyl]-3 ,5 -O-(1,1,3,3-tetraisopropyldisiloxane1,3-diyl)-β-D-arabinofuranosyl}-O6 -[2-(4-nitrophenyl)ethyl]guanine (S.23). Yield: 490 mg (69%). TLC (1:1 hexanes/EtOAc): Rf = 0.44. 1 H NMR (300 MHz, CDCl3 ): 1.00-1.16 (m, 28H, (iPr)4 Si2 O); 2.55 (s, 3H, COCH3 ); 3.35 (t, J = 6.8 Hz, 2H, CH2 -C6 H4 -NO2 ); 3.94 (m, 1H, H4 ); 4.12 (m, 2H, H5 a and H5 b); 4.83 (t, J = 6.8 Hz, 2H, O6 -CH2 ); 4.95 (m, 1H, H3 ); 4.30 (s, 1H, 2 -OH); 5.46 (m, 1H, H2 ); 6.42 (d, J = 5.9 Hz, 1H, H1 ); 7.49 (d, J = 8.7 Hz, 2H, 4-nitrophenyl H2/H6); 7.88 (s, 1H, HN2 ); 8.05 (s, 1H, H8); 8.17 (d, J = 8.7 Hz, 2H, 4-nitrophenyl H3/H5). 13 C NMR (75 MHz, CDCl3 ): δ 12.6, 13.0, 13.1, 13.2, 16.6, 16.7, 16.8, 17.2, 17.3, 17.4 ((iPr)4 Si2 O); 25.1 (COCH3 ); 35.0 (CH2 -C6 H4 -NO2 ); 61.0 (C5 ); 67.0 (O6 -CH2 ); 72.7 (C3 ); 80.0
Synthesis of Modified Nucleosides
1.15.17 Current Protocols in Nucleic Acid Chemistry
Supplement 27
(C1 ); 80.9 (C4 ); 87.7 (C2 ); 117.4 (C(ar)); 123.8 (4-nitrophenyl C2/C6); 129.9 (4-nitrophenyl C3/C5); 139.7 (C8); 145.4, 147.0, 152.3, 152.8, 160.7 (C(ar)); 170.6 (COCH3 ). UV (MeOH): λmax 268 nm. ESI-MS (m/z): calcd. for C33 H47 F3 N6 O11 SSi2 [M+H]+ 849.99, found 849.37; calcd. for [M+Na]+ 871.98, found 871.27.
Introduce 2 -methylseleno group 39. Apply Basic Protocol 1, steps 15 to 24, for 230 mg (0.27 mmol) S.23 instead of S.2, scaling down all quantities used by a factor of 7.0. 40. Extract the product three times with 150 mL CH2 Cl2 . 41. Wash the combined CH2 Cl2 phases with 200 mL of 0.2 M TEAA solution and then with 200 mL saturated NaCl solution. 42. Dry the organic phase with 3 g Na2 SO4 , filter using a glass-fritted B¨uchner funnel and filter paper, and evaporate the filtrate to dryness. 43. Purify the residue on a 17-g silica gel column using a solvent system of 15% to 60% (v/v) EtOAc in hexanes. 44. Combine the fractions containing product S.24, as determined by TLC using 4:6 (v/v) hexanes/EtOAc. Evaporate and then coevaporate with 5 mL CH2 Cl2 . 45. Dry the residue in vacuo to obtain the product as a colorless foam. N2 -Acetyl-3 ,5 -O-(1,1,3,3-tetraisopropyldisiloxane-1,3-diyl)-O6 -[2-(4-nitrophenyl)ethyl]-2 -deoxy-2 -methylselenoguanosine (S.24). Yield: 186 mg (87%). TLC (4:6 hexanes/EtOAc): Rf = 0.49. 1 H NMR (300 MHz, CDCl3 ): δ 0.97-1.10 (m, 28H, (iPr)4 Si2 O); 1.93 (s, 3H SeCH3 ); 2.55 (s, 3H, COCH3 ); 3.31 (t, J = 6.8 Hz, 2H, CH2 -C6 H4 -NO2 ); 3.94 (dd, J = 4.4, 4.5 Hz, 1H, H2 ); 4.06 (m, 2H, H5 a and H5 b); 4.16 (m, 1H, H4 ); 4.75 (s, 1H, H3 ); 4.78 (m, 2H, O6 -CH2 ); 6.21 (d, J = 4.5 Hz, 1H, H1 ); 7.49 (d, J = 8.7 Hz, 2H, 4-nitrophenyl H2/H6); 7.87 (s, 1H, HN2 ); 8.07 (s, 1H, H8); 8.16 (d, J = 8.7 Hz, 2H, 4-nitrophenyl H3/H5). 13 C NMR (75 MHz, CDCl3 ): δ 12.7, 13.0, 13.1, 13.5, 16.8, 16.96, 16.97, 17.1, 17.27, 17.28, 17.3, 17.4 ((iPr)4 Si2 O); 25.1 (COCH3 ); 35.0 (CH2 -C6 H4 -NO2 ); 47.0 (C2 ); 61.8 (C5 ); 66.9 (O6 -CH2 ); 71.7 (C3 ); 84.6 (C4 ); 89.7 (C1 ); 118.3 (C(ar)); 123.8 (4-nitrophenyl C2/C6); 129.9 (4-nitrophenyl C3/C5); 139.9 (C8); 145.5, 147.0, 152.0, 152.3, 160.6 (C(ar)); 170.6 (COCH3 ). UV (MeOH): λmax 269 nm. ESI-MS (m/z): calcd. for C33 H50 N6 O8 SeSi2 [M+H]+ 794.92, found 794.97; calcd. for [M+Na]+ 816.93, found 817.15.
Remove O6 - and 3 ,5 -OH protecting groups 46. In a 250-mL one-neck flask, dissolve 735 mg (0.73 mmol) S.24 in 25 mL THF and add 6 mL of 1 M TBAF in THF. Stir the solution 2.5 hr at room temperature. 47. Evaporate the solvent and purify the residue on a 40-g silica gel column using a solvent system of 3% to 7% (v/v) MeOH in CH2 Cl2 . 48. Combine the fractions containing product S.25, as determined by TLC using 85:15 (v/v) CH2 Cl2 /MeOH. Evaporate and then coevaporate with 5 mL CH2 Cl2 . 49. Dry the residue in vacuo to obtain the product as colorless foam.
2 -MethylselenoModified Phosphoramidites and RNA
N2 -Acetyl-2 -deoxy-2 -methylselenoguanosine (S.25). Yield: 230 mg (79%). TLC (85:15 CH2 Cl2 /MeOH): Rf = 0.45. 1 H NMR (300 MHz, DMSO-d6 ): δ 1.63 (s, 3H SeCH3 ); 2.18 (s, 3H, COCH3 ); 3.57 (m, 2H, H5 a and H5 b); 3.97 (m, 1H, H4 ); 4.01 (m, 1H, H2 ); 4.31 (s, 1H, H3 ); 5.02 (t, J = 5.5 Hz, 1H, 5 -OH); 5.82 (d, J = 4.6 Hz, 1H, 3 -OH); 6.14 (d, J = 9.1 Hz, 1H, H1 ); 8.30 (s, 1H, H8); 11.68 (s, 1H, HN2 ); 12.05(s, 1H, HN1 ). 13 C NMR (75 MHz, DMSO-d6 ): δ 2.9 (SeCH3 ); 24.2 (COCH3 ); 46.8 (C2 ); 62.0 (C5 ); 73.2 (C3 ); 87.7 (C4 ); 88.9 (C1 ); 120.5 (C(ar)); 138.2 (C8); 148.5, 149.3, 155.2, (C(ar)); 173.9 (COCH3 ). UV (MeOH): λmax 277 nm. ESI-MS (m/z): calcd. for C13 H17 N5 O5 Se [M+H]+ 403.26, found 403.76; calcd. for [M+Na]+ 425.25, found 426.04.
1.15.18 Supplement 27
Current Protocols in Nucleic Acid Chemistry
Tritylate 5 -hydroxyl group 50. In a 25-mL one-neck flask, coevaporate 178 mg (0.44 mmol) S.25 with dry pyridine, then dissolve in 2 mL pyridine. 51. Add 165 mg (0.48 mmol) DMTr-Cl in two portions within 1 hr to the stirred solution. Continue stirring overnight at room temperature. 52. Evaporate the solvents and dissolve the residue in 200 mL CH2 Cl2 . Wash with 100 mL of 5% citric acid, 100 mL water, and then 100 mL of half-saturated sodium bicarbonate solution in a 500-mL separatory funnel. 53. Dry the organic phase with 10 g Na2 SO4 , remove the solid using a glass-fritted B¨uchner funnel and filter paper, and evaporate the filtrate to dryness. 54. Purify the residue on a 15-g silica gel column using a solvent system of 2% to 5% (v/v) MeOH in CH2 Cl2 . 55. Combine the fractions containing product S.26, as determined by TLC using 9:1 (v/v) CH2 Cl2 /MeOH. Evaporate and then coevaporate with 3 mL CH2 Cl2 . 56. Dry the residue in vacuo to obtain the product as colorless foam. N2 -Acetyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxy-2 -methylselenoguanosine (S.26). Yield: 184 mg (59%). TLC (9:1 CH2 Cl2 /MeOH): Rf = 0.56. 1 H NMR (300 MHz, CDCl3 ): δ 1.76 (s, 3H SeCH3 ); 1.87 (s, 3H, COCH3 ); 3.27 (m, 1H, H5 a); 3.45 (m, 1H, H5 b); 3.73, 3.74 (2s, 6H, OCH3 ); 3.90 (m, 1H, 3 -OH); 4.11 (m, 1H, H2 ); 4.29 (m, 1H, H4 ); 4.57 (s, 1H, H3 ); 5.98 (d, J = 9.0 Hz, 1H, H1 ); 7.18-7.47 (m, 13H, trityl-H); 7.87 (s, 1H, H8); 9.63 (s, 1H, HN2 ); 12.11(s, 1H, HN1 ). 13 C NMR (75 MHz, CDCl3 ): δ 4.05 (SeCH3 ); 23.8 (COCH3 ); 49.0 (C2 ); 55.2 (2 × OCH3 ); 64.1 (C5 ); 72.9 (C3 ); 85.4 (C4 ); 96.5 (C(ar)); 88.9 (C1 ); 113.2, 121.5, 127.1, 127.9, 128.0, 130.0, 135.5, 135.0 (C(ar)); 138.4 (C8); 144.8, 147.5, 148.8, 155.8, 158.7 (C(ar)); 172.5 (COCH3 ). UV (MeOH): λmax 260 nm. FTICR ESI-MS (m/z): calcd. for C34 H35 N5 O7 Se [M+H]+ 706.17833, found 706.17730.
Prepare phosphoramidite 57. Apply Basic Protocol 1, steps 31 to 40, to 220 mg (0.32 mmol) S.26, scaling down all quantities used by a factor of 3. For column purification of S.27, use 15 g silica gel and a solvent system of 0.5% to 1.5% (v/v) MeOH in CH2 Cl2 . N2 -Acetyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxy-2 -methylselenoguanosine 3 -O-(2cyanoethyl)-N,N- diisopropylphosphoramidite (S.27). Yield: 248 mg (88%). TLC (97:3 CH2 Cl2 /MeOH): Rf = 0.63. 1 H NMR (500 MHz, CDCl3 ): δ 1.00-1.27 (m, 24H, 2 × ((CH3 )2 CH)2 N); 1.37, 1.48 (s, 6H, 2 × SeCH3 ); 1.69, 1.73 (s, 6H, COCH3 ); 2.24, 2.28 (2 × m, 2H, CH2 CN); 2.62, 2.66 (2 × t, 2H, CH2 CN); 3.15 (m, 2H, H5 a); 3.54-3.63 (m, 4H, ((CH3 )2 CH)2 N, 2H, H5 b, 2H, POCH2 ); 3.77, 3.78 (2s, 12H, OCH3 ); 3.91-3.99, 4.10-4.22 (2m, 2H, POCH2 ); 4.36, 4.41 (m, 2H, H4 ); 4.45 (m, 2H, H2 ); 4.71, 4.76 (m, 1H, H3 ); 6.01, 6.06 (2 × d, J = 9.0 Hz, 9.34 Hz, 2H, H1 ); 6.80, 7.26, 7.40, 7.55 (4 × m, 26H, trityl-H); 7.80 (s, 2H, H8); 7.67 (s, 2H, HN2 ); 11.93 (s, 2H, HN1 ). 31 P NMR (121 MHz, CDCl3 ): δ 151.3, 150.4. UV (MeOH): λmax 235 nm. FTICR ESI-MS (m/z): calcd. for C34 H35 N5 O7 Se [M+H]+ 906.28639, found 906.28685.
SYNTHESIS OF RNA OLIGONUCLEOTIDES CONTAINING 2 -METHYLSELENO NUCLEOSIDES
BASIC PROTOCOL 2
This protocol describes the automated solid-phase assembly of RNA sequences using 2 -O-TOM-phosphoramidites (UNIT 3.8) and 2 -methylseleno phosphoramidites for sitespecific incorporation of 2 -methylseleno labels. The protocol was developed for a 1µmol synthesis on a Gene Assembler (Amersham Pharmacia Biotech), but can easily be adapted to other automated DNA/RNA synthesizers. Coupling times of 3.0 to 4.0 min are recommended to achieve stepwise coupling yields >98%.
Synthesis of Modified Nucleosides
1.15.19 Current Protocols in Nucleic Acid Chemistry
Supplement 27
Although 2 -methylseleno nucleoside phosphoramidites are stable in the solid state over prolonged storage at −20◦ C, solutions of 2 -methylseleno nucleoside phosphoramidites in CH3 CN should not be stored >20 hr at room temperature (23◦ C). For reliable preparation of selenium-containing oligoribonucleotides, in particular for sequences with >10 to 15 nt, treatment with threo-1,4-dimercapto-2,3-butanediol (DTT) is required during automated assembly (after each step of oxidation; Table 1.15.1) and during deprotection and ligation. This treatment guarantees a uniformly labeled product without oxidation byproducts.
Materials 2 -Methylseleno phosphoramidites (S.4, S.10, S.17, S.27; see Basic Protocol 1 and Alternate Protocols 1 through 3) 2 -O-TOM-phosphoramidites (Glen Research; also see UNITS 2.9 & 3.8) 5-Benzylthio-1H-tetrazole (BTT; Glen Research) Anhydrous acetonitrile (CH3 CN; <30 ppm water; Biosolve) ◦ 4-A molecular sieves (optional) Detritylation solution (see recipe) Oxidation solution (see recipe) Capping solutions A and B (see recipes) 100 mM DTT solution (see recipe) 1,2-Dichloroethane (Fluka) Solid support: e.g., long-chain alkylamine controlled-pore glass (LCAA-CPG; 500 ◦ ◦ A for <40 nt; 1000 A for >40 nt) derivatized with 5 -O-(4,4 -dimethoxytritylated) nucleosides (Glen Research) or polystyrene base supports (PS200; GE Biosciences) Automated DNA synthesizer (e.g., Gene Assembler; Amersham Pharmacia Biotech) Syringes Synthesis column for 1-µmol scale Additional reagents and equipment for automated oligonucleotide synthesis (APPENDIX 3C) 1. Calculate the amounts of phosphoramidites and activator (BTT) required to assemble the desired sequence (∼12 mg phosphoramidite and ∼21 mg BTT per coupling) and add an extra 30 mg phosphoramidite/50 mg BTT to give enough material to purge the lines of the synthesizer. 2. Place the calculated amounts of phosphoramidites and BTT into the appropriate vials of the DNA synthesizer. 3. Use a syringe to add the required amount of dry acetonitrile to obtain ∼0.1 M phosphoramidite solutions (i.e., 1 mL/100 mg solid phosphoramidite) and ∼0.30 M BTT solution (i.e., 1 mL/58 mg solid BTT). ◦
When working with Gene Assembler, 4-A molecular sieves may also be added to these reagents.
4. Create methods for the assembly of the desired sequence. In principle, methods employed for synthesis of DNA sequences can be used here. The steps listed in Tables 1.15.1 and 1.15.2 are recommended.
2 -MethylselenoModified Phosphoramidites and RNA
1.15.20 Supplement 27
Current Protocols in Nucleic Acid Chemistry
Table 1.15.1 Oligoribonucleotide Assembly on a 1.3-µmol Scale with 2 -Methylseleno-Modified Phosphoramidites
Meaninga
Comments
Go to DETRIT method
Deprotection of the last added nucleotide
2.1
Set valve 2 to pos 1
Dichloroethane wash
VALVE POS
2.3
Set valve 2 to pos 3
Acetonitrile wash
K
INTEGRATE
0
4.00
A
ML/MIN
6
4.00
F
LOOP TIMES
7
4.00
B
VALVE POS
8
4.05
A
9
4.15
10
Line
Time
Command
Function
1
0.00
D
CALL METHOD
2
1.50
B
VALVE POS
3
2.50
B
4
2.70
5
Value DETRIT
Stop integrator, peak Peak data can be data available displayed on screen
0.00
Stop the flow
2
Start of loop
Do following steps two times
1.8
Set valve 1 to pos 8
Addition of 90 µL BTT solution
ML/MIN
0.90
Set flow rate 0.9 mL/min
A
ML/MIN
0.00
Stop the flow
4.15
B
VALVE POS
1.#
Set valve 1 to pos #
11
4.20
A
ML/MIN
0.60
Set flow rate 0.6 mL/min
12
4.30
A
ML/MIN
0.00
Stop the flow
13
4.30
B
VALVE POS
1.8
Set valve 1 to pos 8
14
4.35
A
ML/MIN
0.90
Set flow rate 0.9 mL/min
15
4.45
A
ML/MIN
0.00
Stop the flow
16
4.45
G
END OF LOOP
17
4.45
B
VALVE POS
1.1
Set valve 1 to pos 1
18
4.50
A
ML/MIN
1.00
Set flow rate 1 mL/min
19
4.60
A
ML/MIN
0.00
Stop the flow
20
4.60
F
LOOP TIMES
7
Start the loop
21
4.60
C
STEP VALVE
3
Step valve 3 to next pos
Set valve 3 in recycle position
22
4.60
G
END OF LOOP
23
4.65
A
ML/MIN
2.50
Set flow rate 2.5 mL/min
Coupling for 3 min
24
7.65
A
ML/MIN
0.00
Stop the flow
25
7.65
C
STEP VALVE
3
Step valve 3 to next pos
Addition of 60 µL amidite solution
Addition of 90 µL BTT solution
Push BTT and amidite into the recycle loop with acetonitrile
Set valve 3 in flow through pos continued
Synthesis of Modified Nucleosides
1.15.21 Current Protocols in Nucleic Acid Chemistry
Supplement 27
Table 1.15.1 Oligoribonucleotide Assembly on a 1.3-µmol Scale with 2 -Methylseleno-Modified Phosphoramidites, continued
Line
Time
Command
Function
Value
Meaninga
Comments
26
7.70
A
ML/MIN
2.50
Set flow rate 2.5 mL/min
Acetonitrile wash for 0.3 min
27
8.00
A
ML/MIN
0.00
Stop the flow
28
8.05
B
VALVE POS
2.5
Set valve 2 to pos 5
29
8.10
A
ML/MIN
1.00
Set flow rate 1 mL/min
30
8.10
F
LOOP TIMES
4
Start the loop
31
8.10
B
VALVE POS
2.5
Set valve 2 to pos 5
Capping A for 0.1 min
32
8.20
B
VALVE POS
2.6
Set valve 2 to pos 6
Capping B for 0.1 min
33
8.30
G
END OF LOOP
34
8.35
B
VALVE POS
2.3
Set valve 2 to pos 3
Acetonitrile wash for 0.5 min
35
8.35
A
ML/MIN
2.00
Set flow rate 2 mL/min
36
8.85
B
VALVE POS
2.4
Set valve 2 to pos 4
Oxidation for 1 min
37
9.85
B
VALVE POS
2.3
Set valve 2 to pos 3
Acetonitrile wash for 1.5 min
38
11.35
A
ML/MIN
0.00
Stop the flow
39
11.40
B
VALVE POS
2.5
Set valve 2 to pos 5
40
11.45
A
ML/MIN
1.00
Set flow rate 1 mL/min
41
11.45
F
LOOP TIMES
2
Start the loop
42
11.45
B
VALVE POS
2.5
Set valve 2 to pos 5
Capping A for 0.1 min
43
11.55
B
VALVE POS
2.6
Set valve 2 to pos 6
Capping B for 0.1 min
44
11.65
G
END OF LOOP
45
11.65
A
ML/MIN
0.00
Stop the flow
46
11.70
B
VALVE POS
2.3
Set valve 2 to pos 3
47
11.75
A
ML/MIN
2.50
Set flow rate 2.5 mL/min
48
12.25
A
ML/MIN
0.00
Stop the flow
49
12.30
B
VALVE POS
2.7
Set valve 2 to pos 7
50
12.35
A
ML/MIN
0.50
Set flow rate 0.5 mL/min
51
14.35
A
ML/MIN
0.00
Stop the flow
Acetonitrile wash for 0.5 min
Treatment with 1 mL DTT solution in 2 min (100 mM DTT in 2:3 v/v EtOH/H2 O)
continued 2 -MethylselenoModified Phosphoramidites and RNA
1.15.22 Supplement 27
Current Protocols in Nucleic Acid Chemistry
Table 1.15.1 Oligoribonucleotide Assembly on a 1.3-µmol Scale with 2 -Methylseleno-Modified Phosphoramidites, continued
Meaninga
Comments
2.3
Set valve 2 to pos 3
Push residual DTT over column with acetonitrile for 0.55 min
ML/MIN
0.50
Set flow rate 0.5 mL/min
A
ML/MIN
2.00
Set flow rate 2 mL/min
16.50
A
ML/MIN
0.00
Stop the flow
56
16.55
B
VALVE POS
2.5
Set valve 2 to pos 5
57
16.60
A
ML/MIN
1.00
Set flow rate 1 mL/min
58
16.60
F
LOOP TIMES
2
Start the loop
59
16.60
B
VALVE POS
2.5
Set valve 2 to pos 5
Capping A for 0.1 min
60
16.70
B
VALVE POS
2.6
Set valve 2 to pos 6
Capping B for 0.1 min
61
16.80
G
END OF LOOP
62
16.80
A
ML/MIN
0.00
Stop the flow
63
16.85
B
VALVE POS
2.3
Set valve 2 to pos 3
64
16.90
A
ML/MIN
2.5
Set flow rate 2.5 mL/min
65
17.40
A
ML/MIN
0.00
Stop the flow
66
17.40
F
LOOP TIMES
7
Start of loop
67
17.40
C
STEP VALVE
3
Set valve 3 to next pos
68
17.40
G
END OF LOOP
Line
Time
Command
Function
52
14.40
B
VALVE POS
53
14.45
A
54
15.00
55
Value
Acetonitrile wash for 1.5 min
Acetonitrile wash for 0.5 min
Set valve 3 in recycle position
a # = 2 for A-amidite, 3 for C-amidite, 4 for G-amidite, 5 for U-amidite, 6 for custom X-amidite, 7 for custom Y-amidite.
5. Connect the following reagents to the synthesizer according to the manufacturer’s instructions:
Detritylation solution Oxidation solution Capping solutions A and B DTT solution 1,2-Dichloroethane Dry acetonitrile Phosphoramidite solutions BTT solution 6. Calculate the required amount of solid support based on its nucleoside load (as reported by the manufacturer) and the synthesis scale. Fill an empty synthesis column with the RNA support and connect it to the synthesizer. For example, a 1-µmol RNA synthesis requires a column filled with 25 mg of support carrying a nucleoside load of 40 µmol/g.
Synthesis of Modified Nucleosides
1.15.23 Current Protocols in Nucleic Acid Chemistry
Supplement 27
Table 1.15.2 Detritylation Subroutine of the Method File for the Pharmacia Gene Assembler
Line
Time
Command
Function
Value
Meaning
Comments
1
0.00
B
VALVE POS
1.1
Set valve 1 to pos 1
2
0.00
B
VALVE POS
2.1
Set valve 2 to pos 1
3
0.00
C
STEP VALVE
3
4
0.00
A
ML/MIN
2.50
5
1.00
M
PORT SET
9.1
6
1.00
B
VALVE POS
2.2
7
1.10
M
PORT SET
9.0
8
1.10
H
CLEAR DATA
9
1.10
I
MONITOR
1
Selects monitor
Should be 1, only one monitor connected
10
1.10
J
LEVEL%
4
Specifies the value that the monitor signal has to pass to start/stop the integrator
Should be 4
11
1.10
K
INTEGRATE
1
Start integration
12
1.10
E
PS CALL
13
4.00
B
VALVE POS
2.1
Set valve 2 to pos 1
14
6.00
A
ML/MIN
0.00
Stop the flow
15
6.00
B
VALVE POS
2.7
Set valve 2 to pos 7
16
6.00
E
PS CALL
17
6.00
L
HOLD
18
6.00
A
ML/MIN
2.50
Set flow rate 2.5 mL/min
19
6.00
B
VALVE POS
2.2
Set valve 2 to pos 2
Dichloroethane wash
Step valve 3 to next pos Set flow rate 2.5 mL/min Auto zero the UV monitor Set valve 2 to pos 2
Start detritylation Initialization for new peak data
DETR-RET Peak Start Call
Conditional jump to method DETR-RETa if the start of a peak is detected
The software will indicate a HOLD instruction has been reached and will ask whether to continue
0
a This is a method for returning from the detritylation subroutine to the chain elongation method after a peak start was detected.
7. Purge the lines of the synthesizer with all solutions and solvents. 8. Carry out the assembly according to the manufacturer’s instructions. Choose the DMTr-OFF option in the setup menu. After the tenth coupling cycle, coupling yields of >99% can be expected when assayed by the detritylation colorimetric test. 2 -MethylselenoModified Phosphoramidites and RNA
9. Remove the synthesis column from the synthesizer and dry it in vacuo or under a stream of argon. 10. Deprotect the assembled, still fully protected oligoribonucleotide (see Basic Protocol 3).
1.15.24 Supplement 27
Current Protocols in Nucleic Acid Chemistry
DEPROTECTION, PURIFICATION, AND ANALYSIS OF RNA OLIGONUCLEOTIDES CONTAINING 2 -METHYLSELENO LABELS
BASIC PROTOCOL 3
This protocol describes the three-step deprotection of Se-modified RNA sequences assembled according to Basic Protocol 2. In the first step, the fully protected oligoribonucleotide attached to the solid-support is treated with a freshly prepared DTT solution in ethanol/H2 O to reduce potentially formed selenoxides. In the second step, methylamine in 50% ethanol is used to release the sequence from the solid support, eliminate the cyanoethyl groups, and remove the acetyl-protecting groups from the nucleobases. In the third step, TBAF in THF is used to remove the 2 -O-TOM protecting groups. This reaction is quenched by addition of aqueous TEAA solution (pH 7.4). A final desalting procedure with commercially available size-exclusion columns yields the crude RNA oligonucleotide. Additionally, this protocol describes purification by anion-exchange HPLC and determination of molecular weights of Se-modified RNA by LC-ESI mass spectrometry.
Materials Se-modified, protected oligoribonucleotide attached to the solid support (see Basic Protocol 2) 150 mM and 2 M DTT solutions (see recipe) 8 M methylamine in ethanol (Fluka) 40% (v/v) aqueous methylamine (Fluka) 50% (v/v) ethanol in water 1 M tetrabutylammonium fluoride trihydrate (TBAF·3H2 O; Fluka) in dry tetrahydrofuran (THF) N-Methylpyrrolidone (NMP) or N,N-dimethylformamide (DMF; optional) 1 M triethylammonium acetate (TEAA) buffer, pH 7.4, or 1 M Tris·Cl buffer, pH 7.4 (RNase-free, sterile; Fluka, for molecular biology) Eluant A: 25 mM Tris·Cl (pH 8.0; APPENDIX 2A)/6 M urea Eluant B: 25 mM Tris·Cl (pH 8.0)/0.5 M NaClO4 /6 M urea 0.1 M triethylammonium bicarbonate (TEAB) buffer Acetonitrile (CH3 CN) Ethylenediaminetetraacetic acid (EDTA) Eluant C: 8.6 mM triethylamine (TEA)/100 mM 1,1,1,3,3,3-hexafluoroisopropanol (HFIP) in H2 O, pH 8.3 Eluant D: MeOH 1.5- or 2.0-mL microcentrifuge tubes (twist-top and/or screw-cap) 10-mL one-neck round-bottom flasks Rotary evaporator Speedvac evaporator ¨ ¨ HPLC system (e.g., Amersham AKTAprime or AKTApurifier system) with: 2.6 × 10–cm Amersham HiPrep 26/10 desalting column (Sephadex G25) 4 × 250– and 9 × 250–mm Dionex NucleoPac PA-100 or 200 columns C18 SepPak cartridges (Waters/Millipore) Lyophilizer LC-ESI mass spectrometer (e.g., Finnigan LCQ Advantage MAX ion trap) 2.1 × 100–mm Amersham µRPC C2/C18 column Perform DTT treatment 1. Remove solid support containing the Se-modified, protected oligoribonucleotide from the synthesis cartridge and transfer it into a 2.0-mL sealable screw-cap vial. 2. Add 200 µL of 150 mM DTT solution. Keep the closed vial 1 to 3 hr at room temperature with periodic shaking.
Synthesis of Modified Nucleosides
1.15.25 Current Protocols in Nucleic Acid Chemistry
Supplement 27
Remove acetyl and cyanoethyl groups and release oligoribonucleotide from support 3. Add 0.6 mL of 8 M methylamine in ethanol, 0.6 mL of 40% methylamine in H2 O, and 95 µL of 2 M DTT solution (final DTT concentration 150 mM). 4. Incubate 6 hr at room temperature with periodic shaking. 5. Centrifuge 3 min at 12,500 × g, room temperature, and transfer supernatant from the 2.0-mL vial into a 10-mL one-neck flask. Alternatively, filter the solution through a small glass frit.
6. Wash solid support two times with 0.6 mL of 50% ethanol and add the washes to the flask. 7. Evaporate the solution to dryness on a rotary evaporator and dry briefly in vacuo.
Remove TOM groups 8. Add 0.95 mL of 1 M TBAF in dry THF to the 10-mL flask, and dissolve the crude 2 -O-TOM-protected oligoribonucleotide by vortexing. If the oligoribonucleotide is not dissolved after 10 min, add a few drops of NMP or DMF. Increase solution volume up to 1.6 mL for better solubility of long RNA sequences.
9. Incubate the solution overnight at room temperature with periodic shaking. 10. Quench the reaction by adding an equal volume of 1 M TEAA buffer, pH 7.4 (or 1 M Tris·Cl buffer, pH 7.4). 11. Concentrate to a volume of 0.8 mL. IMPORTANT NOTE: Do not overevaporate to dryness. Check the remaining volume frequently.
Desalt crude oligoribonucleotide 12. Apply the remaining solution to size-exclusion chromatography using a 2.6 × ¨ 10–cm Amersham HiPrep desalting column on an Amersham AKTAprime or ¨ AKTApurifier system, with simultaneous detection by conductivity and UV absorption. Elute with sterile water, pool the oligonucleotide-containing fractions, and evaporate to dryness. This step is required to remove excess TBAF, TEAA, and DTT. Alternatively, desalting can be performed on NAP-10 cartridges.
13. Dissolve the residue in 1.0 mL sterile water in a 2.0-mL screw-cap vial. This stock solution of the crude oligonucleotide can be stored for up to 1 week at −20◦ C.
Purify oligoribonucleotide 14. Prior to purification, analyze the crude product by anion-exchange chromatography. Use 1 to 2 µL of stock solution under the following conditions: Column: 4 × 250–mm Dionex NucleoPac PA-100 or 200 Temperature: 80◦ C Flow rate: 1 mL/min Eluant A: 25 mM Tris·Cl (pH 8.0)/6 M urea Eluant B: 25 mM Tris·Cl (pH 8.0)/0.5 M NaClO4 /6 M urea Gradient: 0% to 60% B in A over 45 min UV detection at 260 nm. 2 -MethylselenoModified Phosphoramidites and RNA
15. Purify the oligonucleotide by anion-exchange chromatography. Use 50- to 150-µL fractions of stock solution to avoid overloading the column. Combine productcontaining fractions.
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Column: 9 × 250–mm Dionex NucleoPac PA-100 Temperature: 80◦ C Flow rate: 2 mL/min Eluant A: 25 mM Tris·Cl (pH 8.0)/6 M urea Eluant B: 25 mM Tris·Cl (pH 8.0)/0.5 M NaClO4 /6 M urea Gradient: 8% to 25% B in A over 20 min UV detection at 275 nm. Desalt purified oligonucleotide 16. Dilute product-containing fractions with an approximately equal volume of 0.1 M TEAB buffer. 17. Load on a conditioned C18 SepPak cartridge. 18. Wash with 10 mL of 0.1 M TEAB buffer and then with 20 mL H2 O. 19. Elute product with 30 mL of 6:4 (v/v) H2 O/CH3 CN. 20. Lyophylize RNA-containing fractions.
Determine molecular weight 21. Apply 250 pmol Se-modified RNA oligonucleotide dissolved in 20 µL of 20 mM EDTA solution to an LC-ESI mass spectrometer (injection volume: 10 to 20 µL). Determine the molecular weight of the oligoribonucleotide in the negative-ion mode. Column: 2.1 × 100–mm Amersham µRPC C2/C18 Temperature: 21◦ C Flow rate: 100 µL/min Eluant C: 8.6 mM TEA/100 mM HFIP in H2 O, pH 8.3 Eluant D: MeOH Gradient: 0% to 100% D in C over 30 min UV detection at 254 nm. The above conditions were used with a Finnigan LCQ Advantage MAX ion trap instrument connected to an Amersham Ettan micro HPLC system with –4 kV applied to the spray needle.
ENZYMATIC LIGATION OF SELENIUM-MODIFIED RNA OLIGONUCLEOTIDES USING T4 RNA LIGASE
BASIC PROTOCOL 4
This protocol describes the enzymatic ligation of 2 -methylseleno-modified oligoribonucleotides at a nanomolar scale using T4 RNA ligase, as exemplified by the preparation of an adenine deaminase (add) adenine riboswitch aptamer domain (Fig. 1.15.5).
Materials 2 -Methylseleno-modified RNA oligonucleotide (see Basic Protocol 3) T4 RNA ligase 1 (ssRNA ligase, 20,000 U/mL) and 10× ligation buffer (New England Biolabs) Phenol (water-saturated) Chloroform Isoamyl alcohol 1.5-mL screw-cap vials 21◦ and 90◦ C heating block HPLC and LC-ESI-MS systems (see Basic Protocol 3)
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Figure 1.15.5 Enzymatic ligation of the adenine deaminase (add) adenine riboswitch aptamer domain (Vibrio vulnificus) with 2 -methylseleno labels. Ligation of the 71-nt RNA was achieved using T4 RNA ligase (0.20 U/µL; cRNA = 40 µM each strand; donor/acceptor = 1/1) at 21◦ C. DTT in ligation buffer is dithiothreitol. Analysis of the experiment by anion-exchange HPLC, denaturating PAGE, and LC-ESI-MS.
1. In a 1.5-mL screw-cap vial, mix 16 nmol of each of the 2 -methylseleno-modified RNA donor/acceptor strands drawn from their respective aqueous stock solutions. 2. Adjust the volume to 360 µL with water. 3. Heat to 90◦ C for 3 min, then allow to cool to 25◦ C within 15 min. 4. Add 40 µL of 10× ligation buffer and 4.0 µL (80 U) of T4 RNA ligase (in storage solution). This results in a final RNA concentration of 40 µM for each RNA fragment and a final ligase concentration of 0.2 U/µL. The 1× ligation buffer is 50 mM Tris·Cl, 10 mM MgCl2 , 10 mM dithiothreitol, 1 mM ATP, pH 7.8 at 25◦ C. For alternative ligation sites, optimize ligase concentration on a small scale.
5. Incubate the reaction solution 2 hr at 21◦ C. For alternative ligation sites, optimize ligation temperature and time on a small scale (using 400 pmol of each RNA fragment).
6. Inactivate the reaction solution by heating to 90◦ C for 3 min. 2 -MethylselenoModified Phosphoramidites and RNA
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7. Extract the reaction mixture two times with an equal volume of 25:24:1 (v/v/v) phenol/chloroform/isoamyl alcohol and once with an equal volume of 24:1 (v/v) chloroform/isoamyl alcohol. This step is necessary to remove ligase protein, which may impair analysis and purification by HPLC in the following step.
8. Analyze and purify the ligation product by anion-exchange HPLC (see Basic Protocol 3, step 15). Alternatively, analyze and purify by denaturating polyacrylamide gel electrophoresis (APPENDIX 3B). HPLC purification is preferred as it leads to higher isolated yields.
9. Determine the molecular weight of the ligation product by LC-ESI mass spectrometry (see Basic Protocol 3, step 21).
ENZYMATIC LIGATION OF SELENIUM-MODIFIED RNA OLIGONUCLEOTIDES USING T4 DNA LIGASE
ALTERNATE PROTOCOL 4
This protocol describes the enzymatic ligation of 2 -methylseleno-modified oligoribonucleotides at a nanomolar scale using T4 DNA ligase, as exemplified by the preparation of an adenine deaminase (add) adenine riboswitch aptamer domain (Fig. 1.15.6). Because T4 DNA ligase catalyzes ligation of nicks in a duplex, a complementary oligonucleotide splint is used to bring the donor/acceptor sites together and form a duplex substrate.
Additional Materials (also see Basic Protocol 4) 2 -O-Methyl RNA oligonucleotide splint T4 DNA ligase (5 U/µL) and 10× ligation buffer (Fermentas) 50% (w/v) polyethylene glycol (PEG) 4000 (Fermentas) 1. In a 1.5-mL screw-cap vial, mix equimolar amounts of the splint oligonucleotide and each of the 2 -methylseleno-modified RNA donor/acceptor strands drawn from their respective aqueous stock solutions. 2. Dilute the resulting mixture with water to achieve a concentration of 20 to 40 µM for each oligonucleotide. 3. Heat to 90◦ C for 3 min, then allow to cool to 25◦ C within 15 min. 4. Calculate the final volume needed to bring each oligonucleotide to a concentration of 10 µM. Add 1/10 of this volume of 10× DNA ligase buffer (final 1× buffer = 40 mM Tris·Cl) and 1/10 vol of 50% (w/v) PEG 4000 (final 5%). 5. Adjust the reaction volume with water as needed to give 19/20 of the final volume. 6. Add 1/20 vol of 5 U/µL T4 DNA ligase (final 0.25 U/µL). 7. Incubate the reaction solution 4 hr at 37◦ C. For alternative ligation sites, optimize ligation temperature and time on a small scale.
8. Perform Basic Protocol 4, steps 6 to 9.
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Figure 1.15.6 Enzymatic ligation of the adenine deaminase (add) adenine riboswitch aptamer domain (Vibrio vulnificus) with 2 -methylseleno labels. Ligation of the 71-nt RNA was achieved using T4 DNA ligase and a 2 -Omethyl RNA splint oligonucleotide (0.25 U/µL; cRNA = 10 µM each strand; donor/acceptor/splint = 1/1/1) at 37◦ C. DTT in ligation buffer is dithiothreitol. (*Slight degradation of the product at 37◦ C was accepted for higher reaction rates).
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Anisaldehyde reagent In a clean container, mix 10 mL anisaldehyde with 180 mL ethanol. While stirring, add 10 mL concentrated sulfuric acid, followed by 2 mL acetic acid. Store up to 3 months at 4◦ C in the dark; avoid contamination with acetone. For staining, TLC plates are dipped into the solution and heated with a heat-gun until reaction compounds appear as dark spots.
Capping solution A 100 mL acetonitrile 6 g DMAP (0.5 M) Store in a dark well-sealed bottle up to 2 months at 25◦ C 2 -MethylselenoModified Phosphoramidites and RNA
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Capping solution B 50 mL acetonitrile 30 mL sym-collidine 20 mL acetic anhydride Store in a dark well-sealed bottle up to 2 months at 25◦ C Detritylation solution 40 mL dichloroacetic acid 960 mL 1,2-dichloroethane (or toluene) Store in a dark well-sealed bottle up to 2 months at 25◦ C DTT solution for oligonucleotide deprotection, 150 mM and 2 M 0.07 g or 0.92 g threo-1,4-dimercapto-2,3-butanediol (DTT) 1.5 mL ethanol 1.5 mL water Prepare fresh (preferred) or store in a vial up to 2 to 3 days in the dark at 4◦ C DTT solution for oligonucleotide synthesis, 100 mM 0.77 g threo-1,4-dimercapto-2,3-butanediol (DTT) 20 mL ethanol 30 mL water Prepare fresh (preferred) or store in a dark well-sealed bottle up to 2 to 3 days in the dark at 4◦ C Oxidation solution 0.40 g iodine (10 mM) 100 mL acetonitrile 9.2 mL sym-collidine 46 mL water Dissolve iodine in acetonitrile, then add sym-collidine followed by water. Store in a dark well-sealed bottle up to 1 week at 25◦ C.
COMMENTARY Background Information Application of 2 -methylseleno RNA In nucleic acid crystallography, seleniumlabeled DNA and RNA oligonucleotides were recently recognized to represent useful derivatives for convenient phasing of X-ray crystallographic data. Pioneering work by Egli, Huang, and co-workers led to the successful MAD-phasing of a short Z-form DNA duplex via phosphoroselenoate backbone modifications and of an A-form DNA duplex via 2 -methylselenouridine modifications (Du et al., 2002; Wilds et al., 2002). Advanced procedures for the preparation of 2 -methylseleno-modified RNA labeled at specific bases have since been introduced (H¨obartner and Micura, 2004; H¨obartner et al., 2005; Moroder et al., 2006) and are described in this unit. The use of threo-1,4-dimercapto-
2,3-butanediol (DTT) during all crucial steps of RNA preparation, including the solid-phase synthesis cycle, represents a major breakthrough. This innovation enables high performance in the preparation of high-purity RNAs with up to 100 nucleotides containing multiple Se-modified residues. Successful applications of the Se-derivatized RNAs have been reported recently for the X-ray structure determination of the group I intron with both exons (Adams et al., 2004) and for the structure determination of the Diels-Alder ribozyme by the SAD technique via site-specific incorporation of 2 -methylseleno pyrimidine derivatives (Serganov et al., 2005). Positioning of labels The replacement of the native 2 -hydroxyls with 2 -methylseleno groups does not significantly interfere with RNA secondary or
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tertiary structure if the labels are properly positioned. As long as the 2 -methylseleno moieties reside in the covariant, double-helical regions of the target fold, and as long as the labeled and native RNAs comprise the same functional groups at their 5 and 3 ends (phosphates, triphosphates, cyclophosphates, or hydroxyl groups), crystallization behavior and structure of the Se-modified RNAs compare well with their nonlabeled counterparts (Du et al., 2002; Teplova et al., 2002; H¨obartner et al., 2005; Moroder et al., 2006). Comparison with other methods of derivatization Although heavy metal ion derivatization, achieved by soaking crystals in heavy atom salt solutions, is the established way to obtain crystals with anomalous scattering centers, labeling with covalent 2 -methylseleno groups is most competitive for structure analysis of RNAs comprising 30 to ∼80 nucleotides. Heavy atom search is a time-consuming process that requires soaking of the RNA crystals with dozens of compounds at various concentrations to obtain reasonably good crystals. Derivatization of RNA with selenium has more impact on nucleic acid crystallography than derivatization with 5-halogenated pyrimidines. Because of the availability of all four nucleoside phosphor2 -methylseleno amidites, there is greater flexibility for adequate positioning of the labels within an RNA target. In addition, nucleobase stacking interactions can be severely perturbed by 5-halogenated pyrimidines (Teplova et al., 2002). Moreover, in contrast to 2 -methylseleno nucleosides, 5-halogenated pyrimidine derivatives are highly photoreactive species (Gott et al., 1991; Xu and Sugiyama, 2004; Zeng and Wang, 2004); inherent radiation damage of 5-halogenmodified nucleic acids during MAD data collection has been documented as a limitation (Ennifar et al., 2002).
Critical Parameters
2 -MethylselenoModified Phosphoramidites and RNA
Nucleoside synthesis There are two critical features in the synthesis of 2 -methylseleno phosphoramidite building blocks. The first is the preparation of the sodium methyl selenoate in THF using dimethyl diselenide and an excess of sodium borohydride. Starting the reaction with anhydrous ethanol (see Basic Protocol 1, step 19) and filtering under inert conditions at the end of the reduction (see Basic Protocol 1, steps
20 and 21) is required to avoid interference by any nucleophiles other than the sodium methyl selenoate that would result in the generation of unwanted products. The second critical feature is the activation of 2 -hydroxyl groups required in the synthesis of the adenosine derivative S.12. Commercial trifluoromethanesulfonyl chloride of the highest quality should be used (see Alternate Protocol 2, step 2). Once the reagent bottle has been opened, it should be used for no longer than 1 month when stored at 4◦ C under argon. In addition, one should not use a larger excess of triethylamine than is described in Alternate Protocol 2 (steps 1 and 15; i.e., 2.5 equivalents) to prevent formation of elimination side products. For small-scale preparation, the equivalent-amounts of triethylamine can be reduced or can be replaced with N-ethyldiisopropylamine (see Alternate Protocol 2, step 1). Automated RNA solid-phase synthesis The DTT treatment step in the solid-phase synthesis cycle, as implemented in the Pharmacia Gene Assembler instrumentation (see Tables 1.15.1 and 1.15.2), is highly recommended. Furthermore, 2 -methylseleno phosphoramidite solutions should not be stored for more than 20 hr at room temperature. Contingent with RNA target design, it is advantageous to place the 2 -methylseleno labels closer to the 5 end than to the 3 end to minimize their exposure to the oxidation solution during synthesis. RNA deprotection, purification, and mass analysis The use of DTT during deprotection is required for high reproducibility (reflected in HPLC chromatograms of crude deprotected materials) and for high yields of the isolated 2 -methylseleno-modified RNA. The authors have evidence that DTT treatment is responsible for the conversion of 2 -methylselenoxides back to 2 -methylseleno groups (Moroder et al., 2006; unpub. observ.). During RNA oligonucleotide purification by anion-exchange chromatography, oligoribonucleotides with 2 -methylselenoxides elute faster than the desired 2 -methylseleno products under the conditions provided in Basic Protocol 3. If other RNA by-products are detected in minor amounts by mass analysis of the isolated product (e.g., 2-cyanoethyl adducts or partially deprotected RNAs), the authors recommend additional purification by ion-exchange chromatography (Dionex
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DNAPac column) using a LiClO4 buffer system at 40◦ to 80◦ C without urea (eluant A: 25 mM Tris·Cl, pH 8.0; eluant B: 25 mM Tris·Cl, pH 8.0, 0.5 M LiClO4 ). Under these conditions, the elution of 2-cyanoethyl adducts and other partially deprotected RNAs is significantly slower and thus permits the contaminants to be well separated. RNA ligation The choice of ligation site is critical to obtain high overall yields when using T4 RNA ligase. Specifically, the site of ligation is best positioned within a single-stranded region comprising 1 to ∼10 nucleotides. The two strands to be ligated should be involved in strong inter-strand pairing interactions that are entropically favorable. Moreover, donor strands (providing the 5 -phosphate) with 5 -terminal pyrimidines are preferred slightly over purines by T4 RNA ligase, whereas 3 -terminal adenosines are the best acceptors. Cytidines and guanosines show intermediate reactivity, and terminal uridines are poor substrates (Arn and Abelson, 1998). It is also recommended to use donor strands that are 5 ,3 -bisphosphates to avoid cyclization of the donor strands and oligomerization by repeated ligation of the donor with previously formed ligation products. When using T4 DNA ligase, annealing of the two ligation fragments is supported by a splint oligonucleotide. Usually a 40- to 60-nt DNA splint is used; however, 16 to 26 nt are recommended for 2 -O-methyl RNA splints. Base-pairing between 2 -O-methyl RNA and RNA is thermodynamically favored compared to base-pairing of RNA with DNA. Therefore, competitive intramolecular secondary structure interactions are more easily broken up with a 2 -O-methyl RNA oligonucleotide compared to a DNA oligonucleotide of the same length. After ligation, during final HPLC purification under denaturing conditions, the ligation product can easily be separated from the splint oligonucleotide by the significant size difference.
Anticipated Results The isolated yields reported in the protocols for each step during phosphoramidite synthesis are representative of the yields typically obtained. The quality of reagents and the dryness of the solvents used will have a significant impact on product yield and purity, as will the amount of care taken during chromatographic purifications.
For a 1-µmol solid-phase synthesis, typical yields of HPLC-purified 2 -methylselenomodified RNA oligonucleotides are 50 to 200 nmol for sequences up to 50 nt with up to 6 methylseleno groups. When using T4 RNA ligase for ligation reactions, isolated yields of HPLC-purified 2 methylseleno-modified RNA are typically in the range of 40% to 60%. A total amount up to 40 nmol is easily obtained for sequences up to 90 nt. Yields are slightly lower for ligations using T4 DNA ligase (30% to 50%).
Time Considerations Each of the intermediates described in this unit can be prepared within 1 or 2 days. Synthesis of the 2 -methylselenouridine phosphoramidite is by far the shortest (5 days) compared to that of the 2 -methylselenocytidine phosphoramidite and 2 methylselenoadenosine phosphoramidite (2 to 3 weeks each), and that of the 2 methylselenoguanosine phosphoramidite (3 to 4 weeks). Solid-phase synthesis (1-µmol scale), deprotection, and purification of three to four 2 methylseleno-modified oligoribonucleotides of 25 to 30 nt each can typically be achieved within 5 days. Ligations of 2 -methylseleno-modified RNAs (product <100 nt), including optimization of ligation conditions on a small scale, are typically completed within 3 to 5 days, although several testing rounds may be needed to find the optimal ligation site.
Literature Cited Adams, P.L., Stahley, M.R., Kosek, A.B., Wang, J., and Strobel, S.A. 2004. Crystal structure of a self-splicing group I intron with both exons. Nature 430:45-50. Arn, E.A. and Abelson, J. 1998. RNA ligases: Function, mechanism, and sequence conservation. In RNA Structure and Function (R.W. Simons and M. Grunberg-Manago, eds.) pp. 695-726. CSHL Press, Cold Spring Harbor, N.Y. Du, Q., Carrasco, N., Teplova, M., Wilds, C.J., Egli, M., and Huang, Z. 2002. Internal derivatization of oligoribonucleotides with selenium for X-ray crystallography using MAD. J. Am. Chem. Soc. 124:24-25. Ennifar, E., Carpentier, P., Ferrer, J.L., Walter, P., and Dumas, P. 2002. X-ray-induced debromination of nucleic acids at the Br K absorption edge and implications for MAD phasing. Acta Crystallogr. D Biol. Crystallogr. 58:1262-1268. Gott, J.M., Wu, H., Koch, T.H., and Uhlenbeck, O.C. 1991. A specific, UV-induced RNAprotein cross-link using 5-bromouridinesubstituted RNA. Biochemistry 30:6290-6295.
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H¨obartner, C. and Micura, R. 2004. The chemical synthesis of selenium-modified oligoribonucleotides and their enzymatic ligation leading to an U6 snRNA stem-loop segment. J. Am. Chem. Soc. 126:1141-1149. H¨obartner, C., Rieder, R., Kreutz, C., Puffer, B., Lang, K., Polonskaia, A., Serganov, A., and Micura, R. 2005. Combined chemical and enzymatic syntheses of RNAs with up to 100 nucleotides containing site-specific 2 -Se-methyl labels for use in X-ray crystallography. J. Am. Chem. Soc. 127:12035-12045. Moroder, H., Kreutz, C., Lang, K., Serganov, A., and Micura, R. 2006. Synthesis, oxidation behavior, crystallization and structure of 2 -methylseleno guanosine containing RNAs. J. Am. Chem. Soc. 128:9909-9918. Pitsch, S., Weiss, P.A., Jenny, L., Stutz, A., and Wu, X. 2001. Reliable chemical synthesis of oligoribonucleotides (RNA) with 2 O-[(triisopropylsilyl)oxy]methyl (2’-O-TOM) protected phosphoramidites. Helv. Chim. Acta 84:3773-3795. Serganov, A., Keiper, S., Malinina, L., Tereshko, V., Skripkin, E., H¨obartner, C., Polonskaia, A., Phan, A.T., Wombacher, R., Micura, R., Dauter, Z., Jaschke, A., and Patel, D.J. 2005. Structural basis for Diels-Alder ribozyme catalyzed carbon-carbon bond formation. Nat. Struct. Mol. Biol. 12:218-224.
Teplova, M., Wilds, C.J., Wawrzak, Z., Tereshko, V., Du, Q., Carrasco, N., Huang, Z., and Egli, M. 2002. Covalent incorporation of selenium into oligonucleotides for X-ray crystal structure determination via MAD: Proof of principle. Biochimie 84:849-858. Wilds, C.J., Pattanayek, R., Pan, C., Wawrzak, Z., and Egli, M. 2002. Selenium-assisted nucleic acid crystallography: Use of phosphoroselenoates for MAD phasing of a DNA structure. J. Am. Chem. Soc. 124:14910-14916. Xu, Y. and Sugiyama, H. 2004. Highly efficient photochemical 2 -deoxyribonolactone formation at the diagonal loop of a 5-iodouracilcontaining antiparallel G-quartet. J. Am. Chem. Soc. 126:6274-6279. Zeng, Y. and Wang, Y. 2004. Facile formation of an intrastrand cross-link lesion between cytosine and guanine upon pyrex-filtered UV light irradiation of d(Br CG) and duplex DNA containing 5-bromocytosine. J. Am. Chem. Soc. 126:65526553.
Contributed by Ronald Micura, Claudia H¨obartner, Renate Rieder, Christoph Kreutz, Barbara Puffer, Kathrin Lang, and Holger Moroder Leopold-Franzens University Innsbruck, Austria
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Palladium-Catalyzed Cross-Coupling Reactions in C6 Modifications of Purine Nucleosides
UNIT 1.16
The most efficient methodology for introducing carbon substituents to carbon atoms of purine derivatives (positions 2, 6, and 8) consists of palladium-catalyzed cross-coupling reactions of halopurines with organometallics. The protocols in this unit describe typical examples of reactions for modification of protected 6-chloropurine nucleosides. Basic Protocol 1 describes cross-coupling reactions of nucleoside S.1 with arylboronic acids, hetarylstannanes, trialkylaluminium, and benzylzinc chloride (Fig. 1.16.1), while Basic Protocol 2 describes the reaction of S.10 with (acetyloxy)methylzinc iodide (Fig. 1.16.2). The latter method affords tris-O-toluoyl-protected 6-(acetyloxymethyl)purine nucleoside S.12, which can be selectively deacetylated by ammonia in the presence of zinc chloride. The resulting sugar-protected 6-(hydroxymethyl)purine nucleoside S.13 can be converted to 6-(fluoromethyl)-, 6-(formyl)-, and 6-(difluoromethyl)purine nucleosides (see Basic Protocols 3 and 4). All the sugar-acylated nucleosides are easily deprotected by treatment with catalytic sodium methoxide in methanol.
CROSS-COUPLING OF AN ACETYL-PROTECTED 6-CHLOROPURINE NUCLEOSIDE
BASIC PROTOCOL 1
C6 modification of the acetyl-protected 6-chloropurine S.1 is described using four different cross-coupling reagents and one of two palladium catalysts, as shown in Figure 1.16.1. The four acetyl-protected products S.2-S.5 are purified and deprotected using a common set of steps.
Materials 6-Chloro-9-(2,3,5-tri-O-acetyl-β-D-ribofuranosyl)purine (S.1; Buck and Reese, 1990) Cross-coupling reagent (Sigma-Aldrich): Phenylboronic acid 2-(Tributylstannyl)thiophene Trimethylaluminium (AlMe3 , 2 M solution in toluene) Benzylzinc chloride (1 M solution in THF) Potassium carbonate (K2 CO3 , anhydrous, Sigma-Aldrich) Palladium catalyst (Sigma-Aldrich): Tetrakis(triphenylphosphine)palladium (Pd(PPh3 )4 ) Bis(triphenylphosphine)palladium dichloride (PdCl2 (PPh3 )2 ) Argon Toluene (dried, degassed, and flushed with argon) Ethyl acetate N,N-Dimethylformamide (DMF, anhydrous under argon, Sigma-Aldrich) Tetrahydrofuran (THF, anhydrous, freshly distilled from Na/benzophenone under argon) Ammonium chloride (NH4 Cl) Anhydrous magnesium sulfate (MgSO4 ) Silica gel (200 to 400 mesh) Hexanes Chloroform Anhydrous methanol
ˇ ar Contributed by Michal Hocek and Peter Silh´ Current Protocols in Nucleic Acid Chemistry (2007) 1.16.1-1.16.17 C 2007 by John Wiley & Sons, Inc. Copyright
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Figure 1.16.1
Synthesis of 6-substituted purine nucleosides.
1 M sodium methoxide (NaOMe) in methanol Heptane 50-, 100-, and 250-mL round-bottom flasks with 14/20 joints, flame dried Rubber septa 21-G needles Vacuum system (oil pump with manifold and cold trap) Oil bath Glass frit (10- to 20-µm) Rotary evaporator (B¨uchi) equipped with a vacuum system (membrane pump Vacuubrand) Glass columns (3.5-cm i.d., 40-cm length) Silica gel 60 F254 aluminium TLC sheets (Merck) UV lamp and heat gun Vacuum pump Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) Perform cross-coupling With phenylboronic acid 1a. Add 413 mg (1 mmol) S.1, 244 mg (2 mmol) phenylboronic acid, 200 mg (1.5 mmol) K2 CO3 , and 59 mg (0.05 mmol) Pd(PPh3 )4 to a flame-dried 100-mL round-bottom flask. 2a. Place a magnetic stir bar in the flask and seal it with a rubber septum. Place the flask on a magnetic stir plate, and connect to an argon/vacuum line via a 21-G needle inserted through the septum. 3a. Evaporate using the vacuum line, then flush with argon very slowly to prevent scattering of the powders. Repeat this procedure three times and leave the flask attached to the argon line on the manifold. 4a. Add 10 mL toluene through the septum. 5a. Place in an oil bath on the magnetic stir plate and heat the reaction flask to 90◦ C. Stir the reaction mixture 8 hr at 90◦ C. PalladiumCatalyzed C6 Modifications of Purine
6a. Cool the reaction mixture to ambient temperature.
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7a. Dilute with 50 mL ethyl acetate and filter through a 10- to 20-µm frit. Wash the frit with 20 mL ethyl acetate. 8a. Evaporate the solution in vacuo. Proceed to step 10.
With 2-(tributylstannyl)thiophene 1b. Add 413 mg (1 mmol) S.1, 560 mg (1.5 mmol) 2-(tributylstannyl)thiophene, and 35 mg (0.05 mmol) PdCl2 (PPh3 )2 to a flame-dried 50-mL round-bottom flask. 2b. Place a magnetic stir bar in the flask and seal it with a rubber septum. Place the flask on a magnetic stir plate, and connect to an argon/vacuum line via a 21-G needle inserted through the septum. 3b. Evaporate using the vacuum line, then flush with argon very slowly to prevent scattering of the powders. Repeat this procedure three times and leave the flask attached to the argon line on the manifold. 4b. Add 5 mL DMF through the septum. 5b. Place in an oil bath on the magnetic stir plate and heat the reaction flask to 90◦ C. Stir the reaction mixture 8 hr at 90◦ C. 6b. Cool the reaction mixture to ambient temperature. 7b. Evaporate the solution in vacuo. Proceed to step 10.
With trimethylaluminium 1c. Add 413 mg (1 mmol) S.1 and 59 mg (0.05 mmol) Pd(PPh3 )4 to a flame-dried 100-mL round-bottom flask. 2c. Place a magnetic stir bar in the flask and seal it with a rubber septum. Place the flask on a magnetic stir plate, and connect to an argon/vacuum line via a 21-G needle inserted through the septum. 3c. Evaporate using the vacuum line, then flush with argon very slowly to prevent scattering of the powders. Repeat this procedure three times and leave the flask attached to the argon line on the manifold. 4c. Add 10 mL THF through the septum. 5c. While stirring, add dropwise 1 mL of 2 M trimethylaluminium (2 mmol). 6c. Place in an oil bath on the magnetic stir plate and heat the reaction flask to 70◦ C. Stir the reaction mixture 8 hr at 70◦ C. 7c. Cool the reaction mixture to ambient temperature. 8c. Pour the mixture onto a mixture of 200 mL crushed ice and 5 g NH4 Cl, and extract three times with 150 mL ethyl acetate. 9c. Dry the collected organic phases with anhydrous MgSO4 , filter, and evaporate in vacuo. Proceed to step 10.
With benzylzinc chloride 1d. Add 413 mg (1 mmol) S.1 and 59 mg (0.05 mmol) Pd(PPh3 )4 to a flame-dried 100-mL round-bottom flask. 2d. Place a magnetic stir bar in the flask and seal it with a rubber septum. Place the flask on a magnetic stir plate, and connect to an argon/vacuum line via a 21-G needle inserted through the septum.
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3d. Evaporate using the vacuum line, then flush with argon very slowly to prevent scattering of the powders. Repeat this procedure three times and leave the flask attached to the argon line on the manifold. 4d. Add 10 mL THF through the septum. 5d. While stirring, add dropwise 2 mL of 1 M benzylzinc chloride (2 mmol). 6d. Place in an oil bath on the magnetic stir plate and heat the reaction flask to 70◦ C. Stir the reaction mixture 8 hr at 70◦ C. 7d. Cool the reaction mixture to ambient temperature. 8d. Pour the mixture onto a mixture of 200 mL crushed ice and 5 g NH4 Cl, and extract three times with 150 mL ethyl acetate. 9d. Dry the collected organic phases with anhydrous MgSO4 , filter, and evaporate in vacuo. Proceed to step 10.
Purify by chromatography 10. Pack a 3.5 × 40–cm glass column with 150 g of 200 to 400 mesh silica gel in 3:1 (v/v) hexanes/ethyl acetate. 11. Dissolve the evaporated residue of the crude acylated nucleoside (S.2-S.5) in 2 to 3 mL chloroform and apply it to the top of the silica gel column using a glass Pasteur pipet. Let the solution settle into the silica gel. 12. Elute with 500 mL each of 3:1, 2:1, and 1:1 (v/v) hexanes/ethyl acetate, followed by 500 mL pure ethyl acetate. Collect 40- to 50-mL fractions. 13. Analyze fractions by TLC using 2:1 (v/v) hexanes/ethyl acetate. 14. Transfer fractions containing pure product to a 250-mL round-bottom flask, evaporate, and dry under vacuum. 15. Confirm the structure of the product by NMR. 9-(2,3,5-Tri-O-acetyl-β-D-ribofuranosyl)-6-phenylpurine (S.2). Colorless amorphous solid, yield 79%. FAB MS, m/z (rel. %): 455 (22) [M+H], 197 (100) [M+H-AcRf]. 1 H NMR (400 MHz, CDCl3 ): 2.09, 2.13, and 2.16 (3 × s, 3 × 3H, CH3 ); 4.37-4.50 (m, 3H, H4 and 2 × H5 ); 5.71 (dd, 1H, J = 3.6 and 5.4 Hz, H3 ); 6.02 (t, 1H, J = 5.4 Hz, H2 ); 6.30 (d, 1H, J = 5.3 Hz, H1 ); 7.52-7.58 (m, 3H, HPh); 8.28 (s, 1H, H8); 8.76 (dd, 2H, J = 1.6 and 8.0 Hz, HPh); 9.03 (s, 1H, H2). 6-(2-Thienyl)-9-(2,3,5-tri-O-acetyl-β-D-ribofuranosyl)purine (S.3). Colorless amorphous solid, yield 89%. FAB MS, m/z (rel. %): 461 (100) [M+H]. 1 H NMR (400 MHz, CDCl3 ): 2.09, 2.14, and 2.16 (3 × s, 3 × 3H, CH3 CO); 4.37-4.50 (m, 3H, H4 and 2 × H5 ); 5.70 (dd, 1H, J = 5.4 and 4.4 Hz, H3 ); 6.00 (t, 1H, J = 5.4 Hz, H2 ); 6.28 (d, 1H, J = 5.4 Hz, H1 ); 7.27 (m, 1H, H-arom. overlapped with CHCl3 ); 7.63 (dd, 1H, J = 4.9 and 0.9 Hz, H-arom.); 8.26 (s, 1H, H8); 8.66 (dd, 1H, J = 3.7 and 0.9 Hz, H-arom.); 8.90 (s, 1H, H2). 13 C NMR (100 MHz, CDCl3 ): 20.37, 20.49, and 20.74 (3 × CH3 ); 63.04 (C5 ); 70.67 (C3 ); 73.11 (C2 ); 80.42 (C4 ); 86.33 (C1 ); 128.82 (CH-arom.); 129.44 (C5); 131.11 and 132.95 (CH-arom.); 139.70 (C-i-arom.); 142.47 (C8); 150.64 and 151.58 (C4 and C6); 152.75 (C2); 169.32, 169.54, and 170.27 (3 × CO). HRMS (FAB): calcd. for C20 H21 N4 O7 S [M+H], 461.1131; found, 461.1137.
PalladiumCatalyzed C6 Modifications of Purine
6-Methyl-9-(2,3,5-tri-O-acetyl-β-D-ribofuranosyl)purine (S.4). Colorless oil, yield 66%. 1 H NMR (400 MHz, CDCl3 ): 2.09, 2.13, and 2.16 (3 × s, 3 × 3H, CH3 CO); 2.87 (s, 3H, CH3 -6); 4.38 (dd, 1H, Jgem = 12.8, J5 b,4 = 5.2 Hz, H5 b); 4.44-4.49 (m, 2H, H4 and H5 a); 5.69 (dd, 1H, J3 ,2 = 5.6, J3 ,4 = 4.5 Hz, H3 ); 5.98 (t, 1H, J2 ,3 = 5.6, J2 ,1 = 5.2 Hz, H2 ); 6.24 (d, 1H, J1 ,2 = 5.2 Hz, H1 ); 8.18 (s, 1H, H8); 8.87 (s, 1H, H2). 13 C NMR (100.6 MHz, CDCl3 ): 19.51 (CH3 -6); 20.37, 20.52, and 20.74 (CH3 CO);
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63.01 (C5 ); 70.61 (C3 ); 73.06 (C2 ); 80.35 (C4 ); 86.43 (C1 ); 133.65 (C5); 142.01 (C8); 150.12 (C4); 152.56 (C2); 159.94 (C6); 169.35, 169.56, and 170.29 (CO). 6-Benzyl-9-(2,3,5-tri-O-acetyl-β-D-ribofuranosyl)purine (S.5). Colorless amorphous solid, yield 62%. FAB MS, m/z (rel. %): 469 (14) [M+H], 211 (100). 1 H NMR (400 MHz, CDCl3 ): 2.07, 2.10, and 2.15 (3 × s, 3 × 3H, CH3 CO); 4.35-4.47 (m, 3H, H4 and 2 × H5 ); 4.52 (s, 2H, CH2 -Ph); 5.68 (dd, 1H, J = 4.3, 5.5 Hz, H3 ); 5.98 (dd, 1H, J = 5.3, 5.5 Hz, H2 ); 6.23 (d, 1H, J = 5.3 Hz, H1 ); 7.17-7.49 (m, 5H, H-arom.); 8.20 (s, 1H, H8); 8.90 (s, 1H, H2). 13 C NMR (100 MHz, CDCl3 ): 20.40, 20.54, and 20.74 (3 × CH3 ); 39.57 (CH2 Ph); 63.08 (C5 ); 70.69 (C3 ); 73.07 (C2 ); 80.45 (C4 ); 86.43 (C1 ); 126.75, 128.62, and 129.39 (CH-arom.); 133.17 and 137.62 (C5 and C-i-Ph); 142.52 (C8); 150.80 (C4); 152.92 (C2); 161.34 (C6); 169.36, 169.57, and 170.31 (3 × CO). HRMS (FAB): calcd. for C23 H25 N4 O7 [M+H], 469.1723; found, 469.1694.
Deprotect hydroxyl groups 16. Dissolve 0.5 to 0.9 mmol of purified acylated nucleoside (S.2-S.5) in 100 mL anhydrous methanol in a 250-mL round-bottom flask. 17. Add 100 µL of 1 M NaOMe (0.1 mmol) and allow to stand overnight at ambient temperature. Monitor the progress of the reaction by TLC using ethyl acetate. 18. Evaporate the solvent. Recrystallize the product from a mixture of 5 mL methanol and 5 mL toluene and subsequent addition of 20 mL hot heptane. 19. Confirm the structure by NMR. 6-Phenyl-9-(β-D-ribofuranosyl)purine (S.6). Colorless crystals, yield 66%. m.p. 228◦ 230◦ C. [α]D −56.1 (c 0.5 DMF). FAB MS, m/z (rel. %): 329 (35) [M+H], 197 (75) [M+H-Rf]. 1 H NMR (400 MHz, DMSO-d6 ): 3.31 (s, 3H, CH3 ); 3.59-3.65 and 3.70-3.76 (2 × m, 2H, CH2 -5 ); 4.02 (m, 1H, H4 ); 4.23 (m, 1H, H3 ); 4.67 (ddd, 1H, J = 4.8, 5.5 and 5.7 Hz, H2 ); 5.13 (t, 1H, J = 5.4 Hz, 5 -OH); 5.25 (d, 1H, J = 5.0 Hz, 3 -OH); 5.56 (d, 1H, J = 5.7 Hz, 2 -OH); 6.11 (d, 1H, J = 5.5 Hz, H1 ); 7.55-7.65 (m, 3H, H-Ph); 8.84 (d, 2H, J = 8.0 Hz, H-Ph); 8.93 (s, 1H, H8); 9.02 (s, 1H, H2). UV λmax (ε): methanol, 290 (18300); water, pH 7, 289 (16800); pH 2, 298 (15700); pH 11, 289 (17100). Anal. calcd. for C16 H16 N4 O (280.3): C, 68.55; H, 5.75; N, 19.99; found: C, 68.82; H, 4.79; N, 20.04. 6-(2-Thienyl)-9-(β-D-ribofuranosyl)purine (S.7). Colorless crystals, yield 66%. m.p. 126◦ -129◦ C (96% aq. EtOH/toluene). [α]D −55.26 (c 0.2, DMF). FAB MS, m/z (rel. %): 335 (19) [M+H], 93 (100). 1 H NMR (400 MHz, DMSO-d6 ): 3.58-3.64 and 3.703.75 (2 × m, 2H, CH2 -5 ); 4.01 (m, 1H, H4 ); 4.22 (m, 1H, H3 ); 4.65 (ddd, 1H, J = 4.7, 5.4 and 5.9 Hz, H2 ); 5.13 (t, 1H, J = 5.5 Hz, 5 -OH); 5.24 (d, 1H, J = 5.0 Hz, 3 -OH); 5.55 (d, 1H, J = 5.9 Hz, 2 -OH); 6.07 (d, 1H, J = 5.4 Hz, H1 ); 7.35 (dd, 1H, J = 3.2 and 4.9 Hz, H-arom.); 7.94 (d, 1H, J = 4.9 Hz, H-arom.); 8.65 (d, 1H, J = 3.2 Hz, H-arom.); 8.87 and 8.90 (2 × s, 2 × 1H, H8 and H2). 13 C NMR (100.6 MHz, DMSO-d6 ): 61.23 (C5 ); 70.27 (C3 ); 73.80 (C2 ); 85.71 (C4 ); 87.76 (C1 ); 128.63 (C5); 129.13, 131.87 and 132.54 (CH-arom.); 139.59 (C-i-arom.); 145.06 (C8); 148.76 (C6); 151.67 (C4); 152.02 (C2). UV (MeOH) λmax (εmax ): 323 (23300), 270 (6400), 228 (8900). Anal. calcd. for C14 H16 N4 O5 S·H2 O (352.4): C, 47.72%; H, 4.58%; N, 15.90%; found: C, 47.81%; C, 4.60%; N, 15.67%. 6-Methyl-9-(β-D-ribofuranosyl)purine (S.8). Colorless crystals, yield 80%. m.p. 207◦ 209◦ C. [α]D −51.2 (c 0.2, DMF) [Lit.: m.p. 209◦ C, [α]D −52.1 (MeOH)]. 1 H NMR (400 MHz, DMSO-d6 ): 2.73 (s, 3H, CH3 ); 3.57 (ddd, 1H, Jgem =12.0, J5 b,OH =6.1, J5 b,4 = 4.0 Hz, H5 b); 3.69 (ddd, 1H, Jgem = 12.0, J5 a,OH = 5.2, J5 a,4 = 4.0 Hz, H5 a); 3.98 (q, 1H, J4 ,5 = 4.0, J4 ,3 = 3.6 Hz, H4 ); 4.19 (td, 1H, J3 ,2 = J3,OH = 4.9, J3 ,4 = 3.6 Hz, H3 ); 4.63 (q, 1H, J2 ,OH = 6.0, J2 ,1 = 5.7, J2 ,3 = 4.9 Hz, H2 ); 5.13 (t, 1H, JOH,5 = 6.1, 5.2 Hz, 5 -OH); 5.25 (d, 1H, JOH,3 = 4.9 Hz, 3 -OH); 5.53 (d, 1H, JOH,2 = 6.0 Hz, 2 -OH); 6.02 (d, 1H, J1 ,2 = 5.7 Hz, H1 ); 8.76 and 8.79 (2 × s, 2 × 1H, H2, H8). 13 C NMR (100.6 MHz, DMSO-d6 ): 19.28 (CH3 ); 61.48 (C5 ); 70.51 (C3 ); 73.84 (C2 ); 85.87 (C4 ); 87.81 (C1 ); 133.09 (C5); 144.19 (C8); 150.26 (C4); 151.84 (C2); 158.53 (C6).
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6-Benzyl-9-(β-D-ribofuranosyl)purine (S.9). Colorless crystals, yield 66%. m.p. 97◦ 100◦ C (EtOH/toluene/heptane). [α]D −47.73 (c 0.2, DMF). FAB MS, m/z (rel. %): 343 (16) [M+H], 52 (32), 185 (45), 93 (100). 1 H NMR (500 MHz, DMSO-d6 ): 3.55-3.60 and 3.66-3.71 (2 × m, 2H, CH2 -5 ); 3.98 (m, 1H, H4 ); 4.19 (m, 1H, H3 ); 4.42 (s, 2H, CH2 Ph); 4.65 (m, 1H, H2 ); 5.09 (t, 1H, J = 5.5 Hz, 5 -OH); 5.22 (d, 1H, J = 4.8 Hz, 3 -OH); 5.51 (d, 1H, J = 5.9 Hz, 2 -OH); 6.02 (d, 1H, J = 5.7 Hz, H1 ); 7.15-7.39 (m, 5H, H-arom.); 8.81 and 8.83 (2 × s, 2 × 1H, H8 and H2). 13 C NMR (100 MHz, DMSO-d6 ): 38.57 (CH2 Ph); 61.23 (C5 ); 70.29 (C3 ); 73.52 (C2 ); 85.67 (C4 ); 87.58 (C1 ); 126.32, 128.34 and 129.02 (CH-arom.); 132.36 and 137.92 (C5 and C-i-arom.); 144.52 (C8); 151.88 (C2); 150.67 and 159.75 (C4 and C6). UV (MeOH), λmax (εmax ): 263 (5600). Anal. calcd. for C17 H18 N4 O4 (342.3): C, 59.64%; H, 5.30%; N, 16.37%; found: C, 59.25%; H, 5.31%; N, 15.99%. BASIC PROTOCOL 2
CROSS-COUPLING OF A TOLUOYL-PROTECTED 6-CHLOROPURINE NUCLEOSIDE C6-modification of the toluoyl-protected 6-chloropurine S.10 with (acetyloxymethyl)zinc iodide is shown in Figure 1.16.2. The acetylated product S.12 can be deacetylated and deprotected to yield S.14, or can be deacetylated to yield S.13, which is the starting compound for further substitution reactions described in Basic Protocols 3 and 4.
Materials ˇ ar et al., 6-Chloro-9-[2,3,5-tri-O-(p-toluoyl)-β-D-ribofuranosyl]purine S.10 (Silh´ 2005b) Tetrakis(triphenylphosphine)palladium (Pd(PPh3 )4 ) Argon Tetrahydrofuran (THF, anhydrous, freshly distilled from Na/benzophenone under argon) 1 M (acetyloxymethyl)zinc iodide in THF (S.11, see Support Protocol) 1 M NaH2 PO4 Chloroform Anhydrous magnesium sulfate (MgSO4 ) Silica gel (200 to 400 mesh) Hexanes Ethyl acetate Dichloromethane Ethanol Zinc chloride Anhydrous methanol 25%wt. aq. ammonia 1 M sodium methoxide (NaOMe) in methanol 25- and 250-mL round-bottom flasks with 14/20 joints, flame dried Rubber septum Vacuum system (oil pump with manifold and cold trap) 21-G needle Rotary evaporator (B¨uchi) equipped with a vacuum system (membrane pump Vacuubrand) Glass columns (3-cm i.d., 20-cm length) Silica gel 60 F254 aluminium TLC sheets (Merck) UV lamp and heat gun Vacuum pump PalladiumCatalyzed C6 Modifications of Purine
Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D)
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Figure 1.16.2
Synthesis and transformations of 6-(hydroxymethyl)purines.
Perform cross-coupling with (acetyloxymethyl)zinc iodide 1. Add 641 mg (1 mmol) S.10 and 59 mg (0.05 mmol) Pd(PPh3 )4 to a flame-dried 25-mL round-bottom flask. 2. Place a magnetic stir bar in the flask and seal it with a rubber septum. Place the flask on a magnetic stir plate, and connect to an argon/vacuum line via a 21-G needle inserted through the septum. 3. Evaporate using the vacuum line, then flush with argon very slowly to prevent scattering of the powders. Repeat this procedure three times and leave the flask attached to the argon line on the manifold. 4. Add 10 mL THF through the septum. 5. While stirring, add 3 mL of 1 M (acetyloxymethyl)zinc iodide (S.11, 3 mmol) in THF. Continue stirring 8 hr at ambient temperature. 6. Pour the mixture into 50 mL of 1 M NaH2 PO4 and extract four times with 50 mL chloroform. 7. Dry the collected organic phases with anhydrous MgSO4 , filter, and evaporate.
Synthesis of Modified Nucleosides
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8. Pack a 3 × 20–cm glass column with 200 to 400 mesh silica gel in 3:1 (v/v) hexanes/ethyl acetate. 9. Dissolve the evaporated residue in 2 to 3 mL chloroform and apply it to the top of the silica gel column using a glass Pasteur pipet. Let the solution settle into the silica gel. 10. Elute with 500 mL each of 3:1, 2:1, 1:1, and 1:2 (v/v) hexanes/ethyl acetate. Collect 40- to 60-mL fractions. 11. Analyze fractions by TLC using 1:1 (v/v) hexanes/ethyl acetate. Beside the desired product S.12, a small amount of more polar deacetylated (hydroxymethyl)purine S.13 forms.
12. Transfer fractions containing pure products to a 250-mL round-bottom flask and evaporate. 13. Co-evaporate the residues with 50 mL dichloromethane. Dry the resulting foams under vacuum. 14. Confirm the structure of the products by NMR. 6-(Acetyloxymethyl)-9-[2,3,5-tri-O-(p-toluoyl)-β-D-ribofuranosyl]purine (S.12). Yield: 550 mg (81%) of yellowish foam. Exact mass (FAB HRMS) found: 679.2415; calcd. for C37 H35 N4 O9 : 679.2404. FAB MS m/z (%): 679 (MH+ , 1); 487 (10); 193 (4); 151 (5); 119 (100). 1 H NMR (400 MHz, CDCl3 ): 2.21 (s, 3H, CH3 -Ac); 2.38 and 2.42 (2 × s, 9H, CH3 -Tol); 4.66 (dd, 1H, Jgem = 12.2, J5 b,4 = 4.1 Hz, H5 b); 4.82 (td, 1H, J4 ,3 = 4.6, J4 ,5 = 4.1, 3.1 Hz, H4); 4.89 (dd, 1H, Jgem = 12.2, J5 a,4 = 3.1 Hz, H5 a); 5.60 (s, 2H, O-CH2 ); 6.20 (dd, 1H, J3 ,2 = 5.8, J3 ,4 = 4.6 Hz, H3 ); 6.38 (t, 1H, J2 ,3 = 5.8, J2 ,1 = 5.5 Hz, H2 ); 6.48 (d, 1H, J1 ,2 = 5.5 Hz, H1 ); 7.16, 7.22, and 7.26 (3 × m, 3 × 2H, H-m-Tol); 7.81, 7.91, and 7.99 (3 × m, 3 × 2H, H-o-Tol); 8.23 (s, 1H, H8); 8.85 (s, 1H, H2). 13 C NMR (100.6 MHz, CDCl3 ): 20.80 (CH3 -Ac); 21.70 and 21.73 (CH3 -Tol); 62.42 (O-CH2 ); 63.41 (C5 ); 71.39 (C3 ); 73.66 (C2 ); 81.06 (C4 ); 86.84 (C1 ); 125.60, 125.99, and 126.56 (C-i-Tol); 129.22, 129.26, and 129.34 (CH-m-Tol); 129.77, 129.86, and 129.88 (CH-o-Tol); 132.35 (C5); 143.41 (C8); 144.24, 144.59, and 144.71 (C-p-Tol); 151.27 (C4); 152.65 (C2); 155.69 (C6); 165.15, 165.37, and 166.18 (CO-Tol); 170.74 (CO-Ac). 6-(Hydroxymethyl)-9-[2,3,5-tri-O-(p-toluoyl)-β-D-ribofuranosyl]purine (S.13). Yield: 63 mg (10%) of yellowish foam. Exact mass (FAB HRMS) found: 673.2313; calcd. for C35 H33 N4 O8 : 673.2298. FAB MS m/z (%): 637 (MH+ , 4); 487 (5); 151 (4); 119 (100). 1 H NMR (500 MHz, DMSO-d6 ): 2.34, 2.38, and 2.39 (3 × s, 3 × 3H, CH3 -Tol); 4.63 (dd, 1H, Jgem = 12.3, J5 b,4 = 4.8 Hz, H5 b); 4.78 (dd, 1H, Jgem = 12.3, J5 a,4 = 3.6 Hz, H5 a); 4.85 (ddd, 1H, J4 ,3 = 5.7, J4 ,5 = 4.8, 3.6 Hz, H4); 4.89 (d, 2H, JCH2,OH = 6.3 Hz, CH2 -OH); 5.46 (t, 1H, JOH,CH2 = 6.3 Hz, OH); 6.23 (dd, 1H, J3 ,2 = 6.0, J3 ,4 = 5.7 Hz, H3 ); 6.51 (dd, 1H, J2 ,3 = 6.0, J2 ,1 = 4.7 Hz, H2 ); 6.66 (d, 1H, J1 ,2 = 4.7 Hz, H1 ); 7.25, 7.30, and 7.31 (3 × m, 3 × 2H, H-m-Tol); 7.76, 7.85 and 7.87 (3 × m, 3 × 2H, H-o-Tol); 8.76 (s, 1H, H2); 8.79 (s, 1H, H8). 13 C NMR (125.8 MHz, DMSO-d6 ): 21.36, 21.37, and 21.39 (CH3 -Tol); 60.14 (CH2 -OH); 63.19 (C5 ); 70.76 (C3 ); 72.91 (C2 ); 79.59 (C4 ); 86.75 (C1 ); 125.75, 126.07, and 126.72 (C-i-Tol); 129.45, 129.52, 129.60, and 129.62 (CH-Tol); 131.81 (C5); 144.06, 144.57, and 144.69 (C-p-Tol); 145.59 (C8); 150.61 (C4); 152.07 (C2); 160.36 (C6); 164.66, 164.86, and 165.59 (CO-Tol).
Selectively deacetylate S.12 15. Dissolve 500 mg (0.736 mmol) of 6-(acetyloxymethyl)purine derivative S.12 in 25 mL ethanol. PalladiumCatalyzed C6 Modifications of Purine
16. Add 120 mg (0.883 mmol) zinc chloride and 0.4 mL of 25 %wt. aqueous ammonia (6 mmol).
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17. Stir the reaction mixture for 36 hr at ambient temperature and monitor the reaction by TLC using 1:1 (v/v) hexanes/ethyl acetate. Even after 36 hr a small amount of starting compound S.12 is present.
18. Add 3 g silica gel to the reaction mixture and evaporate solvents. 19. Pack a 3 × 20–cm glass column with 200 to 400 mesh silica gel in 2:1 (v/v) hexanes/ethyl acetate. 20. Add the adsorbed product on the top of the silica gel column. 21. Elute with 500 mL each of 1:1, 1:2, and 1:3 (v/v) hexanes/ethyl acetate. Collect 40to 60-mL fractions. 22. Analyze fractions by TLC using 1:2 (v/v) hexanes/ethyl acetate. Beside the desired deacetylated product S.13, a small amount (∼10%) of less polar starting compound S.12 can be eluted.
23. Transfer fractions containing pure product to a 250-mL round-bottom flask and evaporate. 24. Co-evaporate the residue with 50 mL dichloromethane. Dry the resulting foam under vacuum. 25. Characterize the product by NMR. Yield of S.13: 277 mg (59%) as white foam.
Deprotect S.12 26. Dissolve 0.5 to 0.9 mmol S.12 in 100 mL anhydrous methanol in a 250-mL roundbottom flask. 27. Add 100 µL of 1 M NaOMe (0.1 mmol) and allow the solution to stand overnight at ambient temperature. Monitor the progress by TLC using 10:1 (v/v) ethyl acetate/ methanol. 28. Add 3 g silica gel to the reaction mixture and evaporate solvents. 29. Pack a 3 × 20–cm glass column with silica gel in 1:3 (v/v) hexanes/ethyl acetate. 30. Add the adsorbed product on the top of the silica gel column and elute with 300 mL of 10:1 and 600 mL of 8:2 (v/v) ethyl acetate/methanol. Collect 40- to 60-mL fractions. 31. Analyze by TLC using 10:1 (v/v) ethyl acetate/methanol. 32. Transfer fractions containing pure product to a 250-mL round-bottom flask and evaporate solvents. 33. Confirm the structure by NMR. 6-(Hydroxymethyl)-9-(β-D-ribofuranosyl)purine (S.14). Yield: 80% of white hygroscopic solid (lyophilized from water). 1 H NMR (400 MHz, MeOD): 3.77 (dd, 1H, Jgem = 12.3, J5 b,4 = 3.3 Hz, H5 b); 3.89 (dd, 1H, Jgem = 12.3, J5 a,4 = 3.0 Hz, H5 a); 4.16 (q, 1H, J4 ,3 = 3.6, J4 ,5 b = 3.3, J4 ,5 a = 3.0 Hz, H4 ); 4.36 (dd, 1H, J3 ,2 = 5.1, J3 ,4 = 3.6 Hz, H3 ); 4.74 (t, 1H, J2 ,1 = 5.6, J2 ,3 = 5.1 Hz, H2 ); 5.09 (s, 2H, O-CH2 ); 6.14 (d, 1H, J1 ,2 = 5.6 Hz, H1 ); 8.74 (s, 1H, H8); 8.89 (s, 1H, H2). 13 C NMR (100.6 MHz, MeOD): 61.51 (O-CH2 ); 62.99 (C5 ); 72.18 (C3 ); 75.76 (C2 ); 87.62 (C4 ); 90.71 (C1 ); 132.82 (C5); 146.32 (C8); 152.10 (C4); 153.05 (C2); 161.07 (C6). Anal. calcd. for C11 H14 N4 O5 ·H2 O: C, 44.00; H, 5.37; N, 18.66; found: C, 44.40; H, 5.29; N, 18.37.
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SUPPORT PROTOCOL
PREPARATION OF (ACETYLOXYMETHYL)ZINC IODIDE Preparation of (acetyloxymethyl)zinc iodide is performed in three reactions and is illustrated in the top of Figure 1.16.2.
Materials Paraformaldehyde (Sigma-Aldrich) Zinc chloride (ZnCl2 , Sigma-Aldrich) Dichloromethane Acetyl chloride (AcCl; Sigma-Aldrich) Sodium iodide (NaI; Sigma-Aldrich) Argon Dry acetone Hexanes Zinc dust (Sigma-Aldrich) Dry tetrahydrofuran (THF) 1,2-Dibromoethane (Sigma-Aldrich) Chlorotrimethylsilane (Sigma-Aldrich) Iodomethyl acetate 25-mL, 250-mL, and 1-L round-bottom flasks, flame dried Reflux condenser Dropping funnel with a by-pass Vacuum system (oil pump with manifold and cold trap) Distillation head Vacuum pump Glass frit Rotary evaporator (B¨uchi) equipped with a vacuum system (membrane pump Vacuubrand) Rubber septum 21-G needles Prepare chloromethyl acetate 1. Add 15 g (0.5 mol) paraformaldehyde, 0.68 g (5 mmol) dry ZnCl2 , and a magnetic stir bar to a 250-mL round-bottom flask. 2. Add 30 mL dichloromethane and fit the flask with a reflux condenser. Place on a magnetic stir plate and mix the suspension. Place a dropping funnel with a by-pass on top of the condenser. 3. Add 36 mL (0.51 mol) acetyl chloride dropwise from the dropping funnel into the stirred suspension. CAUTION: The reaction is considerably exothermic. Add AcCl slowly. The exothermic reaction starts after addition of ∼5 to 10 mL of AcCl.
4. After complete addition of AcCl, stir the reaction mixture for 8 hr at ambient temperature. 5. Fit the flask with a distillation head and distill the chloromethyl acetate in vacuo. Collect the fraction with a boiling point of 48◦ to 53◦ C/100 mbar. Yield: 24 g (45%). This crude chloromethyl acetate is used directly for preparation of iodomethyl acetate. PalladiumCatalyzed C6 Modifications of Purine
Prepare iodomethyl acetate 6. Add 43 g (0.288 mol) NaI to a 1-L round-bottom flask and heat it to ∼200◦ C for 5 min. Dry under vacuum. Fill the flask with argon.
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7. Add a magnetic stir bar and 150 mL dry acetone to the flask. 8. Add the crude chloromethyl acetate (24 g, 0.221 mol) to the NaI solution (solid NaCl begins to separate immediately) and stir the reaction mixture 5 hr at ambient temperature. 9. To crystalize unreacted NaI, add 600 mL hexanes to the reaction mixture and filter off the solids through a glass frit. 10. Evaporate the solvents in vacuo. 11. Purify iodomethyl acetate by vacuum distillation. Collect the fraction with a boiling point of 33◦ to 35◦ C/5 mbar. Yield: 26.5 g (60%). 1 H NMR (200 MHz, CDCl3 ): 2.10 (s, 3H, Ac); 5.90 (s, 2H, CH2 ).
12. Store iodomethyl acetate at −20◦ C. Iodomethyl acetate is a slightly unstable compound; after 1 to 2 months it turns dark.
Prepare (acetyloxymethyl)zinc iodide 13. Add 1.31 g (20 mmol) zinc dust to a flame-dried 25-mL round-bottom flask. 14. Place a magnetic stir bar in the flask and seal it with a rubber septum. Place the flask on a magnetic stir plate, and connect to an argon/vacuum line via a 21-G needle inserted through the septum. 15. Evaporate using the vacuum line, then flush with argon. Repeat this procedure three times and leave the flask attached to the argon line on the manifold. 16. Add 4 mL dry THF, 20 µL of 1,2-dibromoethane, and 20 µL chlorotrimethylsilane through the septum and stir 15 min at ambient temperature. 17. While stirring, add dropwise a solution of 2.00 g (10 mmol) iodomethyl acetate in 5 mL dry THF at 0◦ to 5◦ C. 18. Stir the reaction mixture 1 hr at 0◦ to 5◦ C. 19. Allow nonreacted zinc to settle, leaving a clear solution of (acetyloxymethyl)zinc iodide (S.11) in THF. This solution can be stored at −20◦ C in a septum-sealed flask under an argon atmosphere for 1 to 2 weeks. If desired, the concentration of organozinc can be determined by complexometric titration of Zn(II) ions in a hydrolyzed aliquot (average value ∼1.0 M).
PREPARATION OF 6-(FLUOROMETHYL)PURINE Deoxo-fluorination of the 6-(hydroxymethyl)purine derivative S.13 is performed in a single reaction, followed by purification and deprotection to yield the 6-(fluoromethyl)purine S.16 (Fig. 1.16.2).
BASIC PROTOCOL 3
Materials 6-(Hydroxymethyl)purine (S.13; see Basic Protocol 2) Argon Dry dichloromethane Deoxo-Fluor ([bis-(2-methoxyethyl)amino]sulfur trifluoride, Sigma-Aldrich) 5% aq. NaHCO3 Chloroform Anhydrous magnesium sulfate (MgSO4 ) Ethyl acetate Methanol
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25-mL round-bottom flask Rubber septum 21-G needles Vacuum system (oil pump with manifold and cold trap) Rotary evaporator (B¨uchi) equipped with a vacuum system (membrane pump Vacuubrand) Additional reagents and equipment for deprotection and purification (see Basic Protocol 2) Perform deoxo-fluorination 1. Add 320 mg (0.5 mmol) of 6-(hydroxymethyl)purine S.13 to a 25-mL round-bottom flask. 2. Place a magnetic stir bar in the flask and seal it with rubber septum. Place the flask on a magnetic stir plate, and connect to an argon/vacuum line via a 21-G needle inserted through the septum. 3. Evaporate using the vacuum line, then flush with argon very slowly to prevent scattering of the powders. Repeat this procedure three times and leave the flask attached to the argon line on the manifold. 4. Add 20 mL dry dichloromethane through the septum and cool the solution to −20◦ C. 5. While stirring, add 185 µL (1 mmol) Deoxo-Fluor and allow the reaction mixture to warm to ambient temperature. Stir the reaction mixture 8 hr at ambient temperature. 6. Pour the mixture into 40 mL of cold 5% aq. NaHCO3 and extract four times with 40 mL chloroform. 7. Combine the organic phases, dry with anhydrous MgSO4 , filter, and evaporate. 8. Purify the fluorinated nucleoside S.15 by chromatography as in Basic Protocol 2, steps 8 through 14. For TLC analysis, the desired product S.15 is less polar than the starting compound. 6-(Fluoromethyl)-9-[2,3,5-tri-O-(p-toluoyl)-β-D-ribofuranosyl]purine (S.15). Yield: 130 mg (41%) of white foam. Exact mass (FAB HRMS) found: 639.2228; calcd. for C35 H32 FN4 O7 : 639.2255. FAB MS m/z (%): 639 (M+ , 2); 621 (2); 487 (10); 119 (100). 1 H NMR (400 MHz, CDCl3 ): 2.38, 2.42, and 2.43 (3 × s, 3 × 3H, CH3 -Tol); 4.66 (dd, 1H, Jgem = 12.3, J5 b,4 = 4.1 Hz, H5 b); 4.83 (td, 1H, J4 ,3 = 4.6, J4 ,5 = 4.1, 3.1 Hz, H4); 4.91 (dd, 1H, Jgem = 12.3, J5 a,4 = 3.1 Hz, H5 a); 5.86 (d, 2H, JH,F = 46.6 Hz, CH2 -F); 6.22 (dd, 1H, J3 ,2 = 5.8, J3 ,4 = 4.6 Hz, H3 ); 6.40 (t, 1H, J2 ,3 = 5.8, J2 ,1 = 5.4 Hz, H2 ); 6.49 (d, 1H, J1 ,2 = 5.4 Hz, H1 ); 7.16, 7.23, and 7.26 (3 × m, 3 × 2H, H-m-Tol); 7.81, 7.92, and 7.98 (3 × m, 3 × 2H, H-o-Tol); 8.27 (s, 1H, H8); 8.88 (s, 1H, H2). 13 C NMR (100.6 MHz, CDCl3 ): 21.71 and 21.75 (CH3 -Tol); 63.30 (C5 ); 71.37 (C3 ); 73.68 (C2 ); 80.79 (d, JC,F = 173 Hz, CH2 -F); 81.11 (C4 ); 87.00 (C1 ); 125.57, 125.98, and 126.52 (C-i-Tol); 129.24, 129.29, and 129.35 (CH-m-Tol); 129.76 and 129.88 (CH-o-Tol); 132.11 (d, JC,F = 1 Hz, C5); 143.86 (C8); 144.29, 144.63, and 144.76 (C-p-Tol); 151.55 (C4); 152.69 (C2); 154.98 (d, JC,F = 18 Hz, C6); 165.18, 165.39, and 166.17 (CO-Tol).
PalladiumCatalyzed C6 Modifications of Purine
Deprotect S.15 9. Perform deprotection and purification as in Basic Protocol 2, steps 26 to 33, but use 400 mL of 30:1 and 500 mL of 10:1 (v/v) ethyl acetate/methanol to elute the column in step 30. 6-(Fluoromethyl)-9-(β-D-ribofuranosyl)purine (S.16). Yield: 82% of white solid (crystalization from MeOH/EtOAc/heptane, m.p. 184◦ -185◦ C). 1 H NMR (400 MHz,
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DMSO-d6 ): 3.58 (ddd, 1H, Jgem = 12.0, J5 b,OH = 5.9, J5 b,4 = 4.1 Hz, H5 b); 3.70 (dt, 1H, Jgem = 12.0, J5 a,OH = 5.2, J5 a,4 = 4.1 Hz, H5 a); 3.99 (q, 1H, J4 ,5 = 4.1, J4 ,3 = 3.8 Hz, H4 ); 4.20 (bq, 1H, J3 ,2 = 5.1, J3 ,OH = 5.0, J3 ,4 = 3.8 Hz, H3 ); 4.63 (q, 1H, J2 ,OH = 5.7, J2 ,1 = 5.6, J2 ,3 = 5.1 Hz, H2 ); 5.10 (t, 1H, JOH,5 = 5.9, 5.2, 5 -OH); 5.26 (d, 1H, JOH,3 = 5.0 Hz, 3 -OH); 5.56 (d, 1H, JOH,2 = 5.7 Hz, 2 -OH); 5.84 (d, 2H, JH,F = 46.6 Hz, CH2 -F); 6.07 (d, 1H, J1 2 = 5.6 Hz, H1 ); 8.90 (s, 1H, H8); 8.99 (s, 1H, H2). 13 C NMR (100.6 MHz, DMSO-d6 ): 61.36 (C5 ); 70.42 (C3 ); 73.94 (C2 ); 80.71 (d, JC,F = 167 Hz, CH2 -F); 85.90 (C4 ); 87.87 (C1 ); 131.97 (d, JC,F = 2 Hz, C5); 145.74 (C8); 151.76 (C4); 152.08 (C2); 153.62 (d, JC,F = 17 Hz, C6). 19 F NMR (100.6 MHz, DMSO-d6 ): −216.76 (t, JF,H = 46.6 Hz).
PREPARATION OF 6-(DIFLUOROMETHYL)PURINE Oxidation of the 6-(hydroxymethyl)purine derivative S.13 is performed using the DessMartin reagent to give 6-(formyl)purine S.17 (Fig. 1.16.2). This product can undergo deoxo-fluorination and deprotection to yield the 6-(difluoromethyl)purine S.19.
BASIC PROTOCOL 4
Materials 6-(Hydroxymethyl)purine (S.13; see Basic Protocol 2) Dry dichloromethane Dess-Martin reagent [1,1,1-Tris(acetyloxy)-1,1-dihydro-1,2-benziodoxol-3-(1H)-one, Sigma-Aldrich] Saturated aq. NaHCO3 Chloroform Anhydrous magnesium sulfate (MgSO4 ) Silica gel (200 to 400 mesh) Hexanes Acetone Argon Deoxo-Fluor ([bis-(2-methoxyethyl)amino]sulfur trifluoride, Sigma-Aldrich) Ethyl acetate Methanol 25-, 50-, and 250-mL round-bottom flasks Vacuum system (oil pump with manifold and cold trap) Rotary evaporator (B¨uchi) equipped with a vacuum system (membrane pump Vacuubrand) 3 × 20–cm glass columns Vacuum pump TLC plates Vacuum system (oil pump with manifold and cold trap) 21-G needles Rubber septum Additional reagents and equipment for column chromatography (APPENDIX 3E), TLC (APPENDIX 3D), and deprotection and purification (see Basic Protocol 2) Perform oxidation 1. Add 640 mg (1.0 mmol) 6-(hydroxymethyl)purine S.13 to a 50-mL round-bottom flask. 2. Add 15 mL dry dichloromethane and 510 mg (1.2 mmol) Dess-Martin periodinane. 3. Stir the reaction mixture 2 hr at ambient temperature. 4. Pour the mixture into 50 mL saturated aqueous NaHCO3 and extract four times with 50 mL chloroform.
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5. Dry the collected organic phases with anhydrous MgSO4 , filter, and evaporate. 6. Pack a 3 × 20–cm glass column with 200 to 400 mesh silica gel in 4:1 (v/v) hexanes/acetone. 7. Dissolve the evaporated residue in 2 to 3 mL chloroform and add it to the top of the silica gel column using glass Pasteur pipet. Let the solution settle into the silica gel. 8. Elute with 500 mL of 3:1 (v/v) hexanes/acetone followed by 2:1 hexanes/acetone. Collect 40- to 60-mL fractions. 9. Analyze by TLC using 2:1 (v/v) hexanes/acetone. The desired product S.17 is slightly less polar than the starting compound.
10. Transfer fractions containing pure product to a 250-mL round-bottom flask, evaporate solvents, and dry amorphous solid on vacuum. 11. Confirm the structure of the product by NMR. 6-Formyl-9-[2,3,5-tri-O-(p-toluoyl)-β-D-ribofuranosyl]purine (S.17). Yield: 621 mg (98%) of white amorphous solid. Exact mass (FAB HRMS) found: 635.2175; calcd. for C35 H31 N4 O8 : 635.2142. FAB MS m/z (%): 635 (MH+ , 1); 487 (42); 369 (5); 149 (7); 119 (100). 1 H NMR (500 MHz, CDCl3 ): 2.38, 2.42, and 2.43 (3 × s, 3 × 3H, CH3 -Tol); 4.67 (dd, 1H, Jgem = 12.3, J5 b,4 = 4.0 Hz, H5 b); 4.85 (td, 1H, J4 ,3 = 4.6, J4 ,5 = 4.0, 3.1 Hz, H4 ); 4.93 (dd, 1H, Jgem = 12.3, J5 a,4 = 3.1 Hz, H5 a); 6.21 (dd, 1H, J3 ,2 = 5.8, J3 ,4 = 4.6 Hz, H3 ); 6.41 (t, 1H, J2 ,3 = 5.8, J2 ,1 = 5.4 Hz, H2 ); 6.52 (d, 1H, J1 ,2 = 5.4 Hz, H1 ); 7.16, 7.24, and 7.25 (3 × m, 3 × 2H, H-m-Tol); 7.80, 7.93, and 7.97 (3 × m, 3 × 2H, H-o-Tol); 8.46 (s, 1H, H8); 9.05 (s, 1H, H2); 10.45 (s, 1H, HCO). 13 C NMR (125.8 MHz, CDCl3 ): 21.71 and 21.75 (CH3 -Tol); 63.17 (C5 ); 71.35 (C3 ); 73.66 (C2 ); 81.28 (C4 ); 87.23 (C1 ); 125.46, 125.91, and 126.43 (C-i-Tol); 129.26, 129.31, and 129.38 (CH-m-Tol); 129.73 and 129.87 (CH-oTol); 131.97 (C5); 144.38, 144.71, and 144.84 (C-p-Tol); 147.06 (C8); 147.16 (C6); 152.94 (C2); 154.16 (C4); 165.17, 165.39, and 166.13 (CO); 191.31 (CHO).
Perform deoxo-fluorination 12. Add 320 mg (0.5 mmol) 6-formylpurine S.17 to a 25-mL round-bottom flask. 13. Place a magnetic stir bar in the flask and seal it with a rubber septum. Place the flask on a magnetic stir plate, and connect to an argon/vacuum line via a 21-G needle inserted through the septum. 14. Evaporate using a vacuum line, then flush with argon. Repeat this procedure three times and leave the flask attached to the argon line on the manifold. 15. Add 10 mL dry dichloromethane through the septum and cool the solution to −20◦ C. 16. While stirring, add 320 µL (1.75 mmol) Deoxo-Fluor and allow the reaction mixture to warm to ambient temperature. Stir the reaction mixture 14 hr at ambient temperature. 17. Pour the mixture into 40 mL cold 5% aq. NaHCO3 and extract four times with 40 mL chloroform. 18. Dry the collected organic phases with anhydrous MgSO4 , filter, and evaporate. 19. Pack a 3 × 20–cm glass column with silica gel in 3:1 (v/v) hexanes/ethyl acetate. PalladiumCatalyzed C6 Modifications of Purine
20. Dissolve the evaporated residue in 2 to 3 mL chloroform and add it to the top of the silica gel column using a glass Pasteur pipet. Let the solution settle into the silica gel.
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21. Elute with 500 mL each 3:1, 2:1, and 1:1 (v/v) hexanes/ethyl acetate. Collect 40- to 60-mL fractions. 22. Analyze by TLC using 1:1 (v/v) hexanes/ethyl acetate. The desired product S.18 is less polar than the starting compound.
23. Transfer fractions containing pure product to a 250-mL round-bottom flask and evaporate. 24. Co-evaporate the residues with 50 mL dichloromethane. Dry the resulting foam under vacuum. 25. Confirm the structure of the product by NMR. 6-(Difluoromethyl)-9-[2,3,5-tri-O-(p-toluoyl)-β-D-ribofuranosyl]purine (S.18). Yield: 157 mg (48%) of white foam. Exact mass (FAB HRMS) found: 657.2145; calcd. for C35 H31 F2 N4 O7 : 657.2161. FAB MS m/z (%): 657 (MH+ , 0.5); 487 (16), 119 (100). 1 H NMR (500 MHz, CDCl3 ): 2.38, 2.42, and 2.43 (3 × s, 3 × 3H, CH3 -Tol); 4.67 (dd, 1H, Jgem = 12.4, J5 b,4 = 4.1 Hz, H5 b); 4.86 (td, 1H, J4 ,3 = 4.6, J4 ,5 = 4.1, 3.1 Hz, H4 ); 4.92 (dd, 1H, Jgem = 12.4, J5 a,4 = 3.1 Hz, H5 a); 6.20 (dd, 1H, J3 ,2 = 5.8, J3 ,4 = 4.6 Hz, H3 ); 6.40 (t, 1H, J2 ,3 = 5.8, J2 ,1 = 5.4 Hz, H2 ); 6.50 (d, 1H, J1 ,2 = 5.4 Hz, H1 ); 7.01 (t, 1H, JH,F = 53.8 Hz, CH-F2 ); 7.16, 7.23, and 7.25 (3 × m, 3 × 2H, H-m-Tol); 7.80, 7.92, and 7.98 (3 × m, 3 × 2H, H-o-Tol); 8.36 (s, 1H, H8); 8.92 (s, 1H, H2). 13 C NMR (100.6 MHz, CDCl3 ): 21.71 and 21.75 (CH3 -Tol); 63.22 (C5 ); 71.33 (C3 ); 73.63 (C2 ); 81.22 (C4 ); 87.10 (C1 ); 111.79 (t, JC,F = 242 Hz, CH-F2 ); 125.49, 125.93, and 126.46 (C-i-Tol); 129.25, 129.30, and 129.37 (CH-m-Tol); 129.73 and 129.87 (CH-o-Tol); 131.46 (C5); 144.36, 144.68, and 144.81 (C-p-Tol); 145.27 (C8); 150.11 (t, JC,F = 26 Hz, C6); 152.54 (C2); 152.70 (C4); 165.17, 165.38, and 166.15 (CO-Tol).
Deprotect S.18 26. Perform deprotection and purification as in Basic Protocol 2, steps 26 to 33, but use 400 mL of 30:1 and 500 mL of 10:1 (v/v) ethyl acetate/methanol to elute the column in step 30. 6-(Difluoromethyl)-9-(β-D-ribofuranosyl)purine (S.19). Yield: 83% of white solid (crystalization from MeOH/EtOAc/heptane; m.p. 181◦ -182◦ C). 1 H NMR (400 MHz, DMSOd6 ): 3.59 (ddd, 1H, Jgem = 12.1, J5 b,OH = 5.7, J5 b,4 = 4.0 Hz, H5 b); 3.71 (dt, 1H, Jgem = 12.1, J5 a,OH = 5.3, J5 a,4 = 4.0 Hz, H5 a); 4.00 (q, 1H, J4 ,5 = 4.0, J4 ,3 = 3.8 Hz, H4 ); 4.21 (bq, 1H, J3 ,OH = 5.2, J3 ,2 = 4.9, J3 ,4 = 3.8 Hz, H3 ); 4.62 (q, 1H, J2 ,OH = 5.7, J2 ,1 = 5.4, J2 ,3 = 4.9 Hz, H3 ); 5.11 (t, 1H, JOH,5 = 5.7, 5.3 Hz, 5 -OH); 5.28 (d, 1H, JOH,3 = 5.2 Hz, 3 -OH); 5.60 (d, 1H, JOH,2 = 5.7 Hz, 2 -OH); 6.10 (d, 1H, J1 ,2 = 5.4 Hz, H1 ); 7.36 (t, 1H, JH,F = 53.3 Hz, CH-F2 ); 9.01 (s, 1H, H8); 9.09 (s, 1H, H2). 13 C NMR (100.6 MHz, DMSO-d6 ): 61.27 (C5 ); 70.35 (C3 ); 74.10 (C2 ); 85.94 (C4 ); 88.05 (C1 ); 112.28 (t, JC,F = 240 Hz, CH-F2 ); 131.00 (C5); 147.11 (C8); 148.87 (t, JC,F = 25, C6); 152.05 (C2); 153.02 (C4). 19 F NMR (188.2 MHz, DMSO-d6 ): −118.80 (t, JF,H = 53.3 Hz).
COMMENTARY Background Information Several types of purines bearing Csubstituents in position 6 are biologically active. 6-Aryl-, 6-hetaryl-, and 6-benzylpurine (Hocek et al., 2000, 2001) as well as 6trifluoromethylpurine (Hockova et al., 1999) ribonucleosides display significant cytostatic activity, and some 6-hetarylpurine ribonucleosides (Hocek et al., 2005) exert potent antiviral activity against hepatitis C virus (HCV). 6Methylpurine and its ribonucleoside are highly
cytotoxic (Montgomery and Hewson, 1968), and the liberation of the nucleobase from its non-toxic deoxyribonucleoside by purine nucleoside phosphorylases was proposed as a novel principle in gene therapy for cancer (Parker et al., 1997). Recently, it has been found that 6-(hydroxymethyl)purine ribonucleoside exerts a very high cytostatic effect against leukemia cell lines (IC50 = 10 to 100 nM) and moderate inhibition of ˇ ar et al., 2004, adenosine deaminase (Silh´
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2005a), while 6-(fluoromethyl)purine and 6(difluoromethyl)purine ribonucleosides possess lower cytostatic effect and do not inhibit ˇ ar et al., 2005b, 2006). this enzyme (Silh´ C-substituted purines can be prepared by several approaches, including heterocyclization, radical substitution, and nucleophilic substitution. However, the use of crosscoupling reactions of halopurines with diverse organometallics is the most versatile, efficient, and practical approach. The limitations of the previous methodologies and the advantages of the cross-coupling reactions are reviewed in detail in Hocek (2003). The synthesis of purine nucleosides bearing simple aryl or alkyl substituents in position 6 is very easily performed by the Pd-catalyzed cross-coupling reactions of acetyl-protected 6chloropurine riboside S.1 (Fig. 1.16.1). Its reaction with phenyl boronic acid in toluene in the presence of Pd(PPh3 )4 gives the corresponding protected 6-phenylpurine nucleoside in good yield (Hocek et al., 2000). Analogous Stille coupling of S.1 with 2thienyl(tributyl)tin in DMF gives the 6-(2thienyl)purine nucleoside (Hocek et al., 2001). Reaction of S.1 with trimethylaluminium or benzylzinc chloride in THF gives the corresponding 6-methyl- and 6-benzylpurine nucleosides (Hocek et al., 2001, 2006). In all cases, the treatment of the acyl-protected nucleosides with catalytic NaOMe in methanol cleanly produces the desired free nucleosides. Introduction of a hydroxymethyl group to position 6 of the purine ring represents an entry to the field of functionalized substituents since it is easily transformed to many other types of functional groups. A new methodology of hydroxymethylation of purines consists of the cross-coupling reactions of halopurines with (acyloxymethyl)zinc iodides, easily available from chloromethyl esters (Fig. 1.16.2; ˇ ar et al., 2004). The reaction of this Silh´ organozinc reagent with 6-chloropurine nucleoside S.10 in the presence of Pd(PPh3 )4 gives a mixture of 6-(acetyloxymethyl)- and 6-(hydroxymethyl)purine nucleosides. The acetyl group at the hydroxymethyl group can be selectively cleaved, in the presence of toluoyl protection of the sugar moiety, by using aqueous ammonia in the presence of ZnCl2 ˇ ar et al., 2005b). The resulting protected (Silh´ 6-(hydroxymethyl)purine nucleoside is converted to 6-(fluoromethyl)purine nucleoside ˇ ar et al., using the Deoxo-Fluor reagent (Silh´ 2005b). To prepare 6-(difluoromethyl)purines, the 6-(hydroxymethyl)purine is first oxidized to 6-formylpurine by the Dess-Martin reagent,
followed by deoxofluorination by Deoxoˇ ar et al., 2006). Again, all toluoylFluor (Silh´ protected nucleosides are deprotected by NaOMe in methanol.
Critical Parameters The most critical parameter in each crosscoupling reaction is ensuring the absence of oxygen, which destroys the catalyst. Therefore, all reactions must be carried out in argonpurged flasks using septum techniques and a vacuum line. In the case of organozinc and trialkyl aluminium reagents, the absence of any trace of water is necessary for successful reactions; therefore, all reagents and solvents must be perfectly pure and anhydrous. The catalyst Pd(PPh3 )4 should be stored under an argon atmosphere at 4◦ C; however, it can be manipulated in air if it is done reasonably quickly and cleanly.
Anticipated Results The yields of all the cross-coupling reactions should be from 60% to 90% and the separation of the products by column chromatography is very easy. The cleavage of the protecting groups usually proceeds with very high yields. The deoxofluorination of hydroxymethyl- and formylpurines with the Deoxo-Fluor reagent produces only moderate yields of ∼40% to 50%.
Time Considerations The cross-coupling reactions with simple commercial organometallics are usually completed within 8 to 10 hr, and subsequent separation and characterization take an additional 3 to 5 hr. The deprotections are performed overnight, and again the purification and characterization of the products requires an additional 3 to 5 hr. Altogether, it should be feasible to accomplish the two steps within 3 days. In the case of hydroxymethylation, synthesis of 6-(acetyloxymethyl)purine starting from protected 6-chloropurine ribonucleoside can be accomplished in 3 to 4 days. Subsequent selective deacetylation and transformation of the hydroxymethyl group can be accomplished in another 3 to 4 days.
Literature Cited Buck, I.M. and Reese, C.B. 1990. An unambiguous synthesis of adenylosuccinic acid and its constituent nucleoside. J. Chem. Soc., Perkin Trans. 1, 2937-2942. Hocek, M. 2003. Syntheses of purines bearing carbon substituents in positions 2, 6 or 8 by metalor organometallics-mediated C-C bond forming reactions. Eur. J. Org. Chem. 245-254.
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Hocek, M., Hol´y, A., Votruba, I., and Dvoˇra´ kov´a, H. 2000. Synthesis and cytostatic activity of substituted 6-phenylpurine bases and nucleosides: Application of the Suzuki-Miyaura crosscoupling reactions of 6-chloropurine derivatives with phenylboronic acids. J. Med. Chem. 43:1817-1825. Hocek, M., Hol´y, A., Votruba, I., and Dvoˇra´ kov´a, H. 2001. Cytostatic 6-arylpurine nucleosides III. Synthesis and structure-activity relationship study in cytostatic activity of 6-aryl-, 6-hetaryland 6-benzylpurine ribonucleosides. Collect. Czech. Chem. Commun. 66:483-499. Hocek, M., Nauˇs, P., Pohl, R., Votruba, I., Furman, P.A., Tharnish, P.M., and Otto, M.J. 2005. Cytostatic 6-arylpurine nucleosides. 6. SAR in antiHCV and cytostatic activity of extended series of 6-hetarylpurine ribonucleosides. J. Med. Chem. 48:5869-5873. ˇ ar, P., Shih, I., Mabery, E., and Hocek, M., Silh´ Mackman, R. 2006. Cytostatic and antiviral 6-arylpurine ribonucleosides. Part 7: Synthesis and evaluation of 6-substituted purine L-ribonucleosides. Bioorg. Med. Chem. Lett. 16:5290-5293. Hockov´a, D., Hocek, M., Dvoˇra´ kov´a, H., and Votruba, I. 1999. Synthesis and cytostatic activity of nucleosides and acyclic nucleoside analogues derived from 6-(trifluoromethyl)purines. Tetrahedron 55:11109-11118. Montgomery, J.A. and Hewson, K. 1968. Analogs of 6-methyl-9-β-D-ribofuranosylpurine. J. Med. Chem. 11:48-52.
Parker, W.B., King, S.A., Allan, P.W., Bennett, L.L. Jr., Secrist, J.A. III, Montgomery, J.A., Gilbert, K.S., Waud, W.R., Wells, A.H., Gillespie, G.Y., and Sorscher, E.J. 1997. In vivo gene therapy of cancer with E. coli purine nucleoside phosphorylase. Hum. Gene Ther. 8:1637-1644. ˇ ar, P., Pohl, R., Votruba, I., and Hocek, Silh´ M. 2004. Facile and efficient synthesis of 6-(hydroxymethyl)purines. Org. Lett. 6:32253228. ˇSilh´ar, P., Pohl, R., Votruba, I., and Hocek, M. 2005a. Synthesis of 2-substituted 6(hydroxymethyl)purine bases and nucleosides. Collect. Czech. Chem. Commun. 70:1669-1695. ˇSilh´ar, P., Pohl, R., Votruba, I., and Hocek, M. 2005b. The first synthesis and cytostatic activity of novel 6-(fluoromethyl)purine bases and nucleosides. Org. Biomol. Chem. 3:3001-3007. ˇSilh´ar, P., Pohl, R., Votruba, I., and Hocek, M. 2006. Synthesis and cytostatic activity of novel 6-(difluoromethyl)purine bases and nucleosides. Synthesis 1848-1852.
Contributed by Michal Hocek and Peter ˇ ar Silh´ Institute of Organic Chemistry and Biochemistry Academy of Sciences of the Czech Republic Prague, Czech Republic
Synthesis of Modified Nucleosides
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Nucleobase-Caged Phosphoramidites for Oligonucleotide Synthesis
UNIT 1.17
Caged compounds are biologically active compounds that have been made temporarily inactive via modification with a photolabile group (Pelliccioli and Wirz, 2002; Goeldner and Givens, 2005; Mayer and Heckel, 2006). The caged derivative can then be applied, for example, to a model organism and, with the help of (laser) light and a confocal microscope, the active compound can be released with exact spatiotemporal and dosage control. More simple setups are also possible. Due to the localized and ideally infinitesimal jump in concentration of the active form, elaborate kinetic experiments become possible. This unit focuses on the preparation of phosphoramidites that bear such a caging group, and their incorporation into oligonucleotides by standard solid-phase synthesis. The resulting nucleobase-caged oligonucleotides add an element of spatiotemporal control to oligonucleotide-based applications, such as gene regulation, modulation of protein function (e.g., via aptamers), molecular diagnostics, or DNA nanoarchitectures. The protocols in this unit describe the synthesis of the protected precursors S.1a, S.1b, S.2, and S.3 for the introduction of TNPE , TNPP , dGNPP , and dCNPE residues into oligonucleotides, respectively (Fig. 1.17.1). Each of these residues bears its respective caging group on the nucleobase, which makes the formation of the corresponding Watson-Crick base pair difficult or prevents it entirely. Nucleobase-caged nucleotides can be seen as temporary mismatches. The residues shown have been applied in the light-regulation of transcription (Kr¨ock and Heckel, 2005), the light-control of protein function via caged aptamers (Heckel and Mayer, 2005; Mayer et al., 2005), and the light-induction of oligonucleotide folding (Mayer et al., 2005). By applying an antisense strategy (Heckel et al., 2006), it becomes possible not only to liberate an active oligonucleotide (switch
Figure 1.17.1 Protected phosphoramidites used for solid-phase synthesis of oligonucleotides containing TNPE , TNPP , dGNPP , or dCNPE residues. DMTr, 4,4 -dimethoxytrityl; NPE, 1-(2-nitrophenyl)ethyl, NPP, 2-(2-nitrophenyl)propyl.
Contributed by Alexander Heckel Current Protocols in Nucleic Acid Chemistry (2007) 1.17.1-1.17.26 C 2007 by John Wiley & Sons, Inc. Copyright
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it on), but also to sequester it (switch it off). The NPE and NPP cages yield different cleavage products with different toxicological profiles. NOTE: Since most of the reaction steps are sensitive to moisture, great care must be taken to use anhydrous solvents and dry argon or nitrogen as a protecting gas. However, it is sufficient to close the reaction apparatus with a rubber septum and provide a protecting gas balloon for pressure equalization. CAUTION: All reactions must be carried out in a well-ventilated chemical fume hood, and protective gloves, a laboratory coat, and safety glasses must be worn. BASIC PROTOCOL 1
PREPARATION OF PROTECTED NPE- AND NPP-CAGED THYMIDINE PHOSPHORAMIDITES The synthesis scheme for two thymidine phosphoramidites, S.1a and S.1b, is shown in Figure 1.17.2. These phosphoramidites are used for synthesis of oligonucleotides containing TNPE and TNPP caged residues, respectively. The NPE and the NPP groups have different photocleavage products and hence different toxicological profiles. The cleavage of the NPP group (Walbert et al., 2001) from oligonucleotides has been shown to occur more quickly and cleanly than that of the NPE group (Kr¨ock and Heckel, 2005).
Materials Thymidine (S.4) Acetic anhydride (Ac2 O) Pyridine Hexane Acetone Chloroform 1 M aqueous sulfuric acid, 4◦ C Saturated aqueous NaHCO3 Anhydrous Na2 SO4 Diethyl ether Ethanol Acetonitrile Argon source 4-(N,N-Dimethylamino)pyridine (DMAP) Triethylamine (Et3 N) 2,4,6-Triisopropylbenzenesulfonyl chloride (iPr3 C6 H2 SO2 Cl) Cyclohexane 1-(2-Nitrophenyl)ethan-1-ol (NPE-OH; S.22; see Support Protocol 1) or 2-(2-nitrophenyl)propan-1-ol (NPP-OH; S.24; see Support Protocol 2) 1 M aqueous HCl Anhydrous MgSO4 Methanol (MeOH) 33% (v/v) aqueous ammonium hydroxide solution (NH4 OH) 4,4 -Dimethoxytrityl chloride (DMTr-Cl) Dichloromethane (CH2 Cl2 ) N,N-Diisopropylethylamine (DIPEA, H¨unig’s base) 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite
Nucleobase-Caged Phosphoramidites
50-, 100-, and 250-mL round-bottom flasks Reflux condenser 90◦ C oil bath Rotary evaporator connected to vacuum pump High vacuum system
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Figure 1.17.2 Overview of the synthesis of the protected phosphoramidites S.1a and S.1b, for the introduction of TNPE and TNPP into oligonucleotides, starting from thymidine (S.4). DMAP, 4-(N,N-dimethylamino)pyridine; DMTr-Cl, 4,4 -dimethoxytrityl chloride; NPE, 1-(2-nitrophenyl)ethyl; NPP, 2-(2-nitrophenyl)propyl.
500-µL syringes Separatory funnels Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Acetylate thymidine 1. In a 250-mL round-bottom flask equipped with a reflux condenser, suspend 10.0 g (41.3 mmol) thymidine (S.4) in 70 mL acetic anhydride and add 5 mL pyridine. 2. Add a stir bar and heat the reaction mixture slightly in a 90◦ C oil bath until it becomes clear (∼30 min). 3. Allow the reaction to cool to room temperature and continue stirring for an additional 1 hr. 4. Check for complete conversion by TLC using 1:1 (v/v) hexane/acetone as eluent. Traces of pyridine can be removed before developing the TLC by placing the TLC plate in a round-bottom flask of appropriate size and applying vacuum for several minutes. Inspection of the developed TLC plate under UV light (254 nm) should show almost complete conversion (Rf of S.5 ∼0.5, Rf of thymidine <0.1).
5. Pour the reaction mixture into 400 mL cold (4◦ C) water and stir the mixture for 30 min. Two layers will form.
6. Extract the mixture three times with 250 mL chloroform. 7. Combine the organic extracts and wash with 100 mL of 1 M cold (4◦ C) sulfuric acid. During this step and the next, great care must be exercised to prevent spillage of the contents of the separation funnel due to overpressure. Current Protocols in Nucleic Acid Chemistry
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8. Wash the organic phase with 100 mL saturated aqueous NaHCO3 . 9. Dry the organic extracts with Na2 SO4 . Filter off the drying agent. 10. Remove the organic solvents from the filtrate, first on a rotary evaporator connected to a vacuum pump then in a high vacuum. This affords a colorless syrup.
11. Add 100 mL diethyl ether to initiate precipitation of the product, then remove the organic solvent again on a rotary evaporator connected to a vacuum pump. 12. Recrystallize the crude product from 20 to 30 mL ethanol. 13. Characterize the product by melting point determination, TLC, and 1 H NMR. 3 ,5 -Di-O-acetylthymidine (S.5): Yield 11.7 g (86%). m.p. 125◦ C. TLC (silica, 1:1 v/v hexane/acetone): Rf = 0.5. 1 H NMR (CDCl3 /TMS): δ 9.35 (1H, br s), 7.26 (1H, s), 6.25 (1H, m), 5.15 (1H, m), 4.27 (2H, m), 4.18 (1H, m), 2.40 (1H, m), 2.1 (1H, m), 2.07 (6H, s), 1.85 (3H, s).
Add caging group by O4 -modification 14. In a 50-mL round-bottom flask dissolve 1.5 g (4.6 mmol) S.5 in 20 mL acetonitrile under an argon atmosphere. 15. Add 1.12 g (9.2 mmol) DMAP and 1.28 mL Et3 N, and stir the reaction mixture for 10 min at room temperature. 16. Add 2.8 g (9.2 mmol) 2,4,6-triisopropylbenzylsulfonyl chloride and stir at room temperature. The reaction mixture turns yellow and after some minutes a precipitate forms. If the reaction mixture becomes too thick for the stir bar, dilute with a small amount of acetonitrile (20 mL maximum).
17. After 3 hr, check by TLC using 1:1 (v/v) cyclohexane/acetone to verify that all starting material has reacted. Upon inspection under UV light (254 nm) there should be no spot visible from the starting material (Rf = 0.5).
18. Dilute 2.3 g (13.8 mmol) NPE-OH (S.22) or 2.5 g (13.8 mmol) NPP-OH (S.24) with a small amount of acetonitrile and 1.92 mL (13.8 mmol) Et3 N, and add to the reaction mixture. During this procedure, the precipitate can disappear and reappear later.
19. Stir the reaction mixture overnight at room temperature. 20. Add 200 mL chloroform and 200 mL water to the reaction mixture and separate the layers. Extract the aqueous layer two additional times with 200 mL chloroform each time. 21. Combine the organic phases and wash with 200 mL of 1 M HCl. 22. Wash the organic phases with saturated aqueous NaHCO3 and dry with MgSO4 . Filter off the drying agent. During this step, great care must be exercised to prevent spillage of the contents of the separation funnel. TLC control of all the phases at this point should show product only in the organic phase.
Nucleobase-Caged Phosphoramidites
23. Remove the solvent from the filtrate on a rotary evaporator connected to a vacuum pump to afford a brown oil.
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24. Perform column chromatography using a 5 × 24–cm silica gel bed and 2 L of 2:1 (v/v) cyclohexane/acetone as eluent. Analyze fractions by TLC. Isolate the spot that runs just below the spot of the respective alcohol used. The excess alcohol can be recovered.
25. Combine the pure fractions and remove the solvents on a rotary evaporator. 26. Characterize the product by TLC, 1 H NMR, and MS. 3 ,5 -Di-O-acetyl-TNPE (S.6a): Yield 1.3 g (60%). TLC (silica, 1:1 v/v hexane/acetone): Rf = 0.5. 1 H NMR (CDCl3 /TMS): δ 7.85 (1H), 7.58 (2H), 7.50 (1H), 7.37 (1H), 6.73 (1H), 6.20 (1H), 5.14 (1H), 4.30 (2H), 4.22 (1H), 2.60 (1H), 2.00 (10H), 1.70 (3H). HRMS (EI): calcd. for C22 H25 N3 O9 (M+ ) 475.1591; found 475.1591. 3 ,5 -Di-O-acetyl-TNPP (S.6b): Yield 1.6 g (58%). TLC (silica, 1:1 v/v cyclohexane/ acetone): Rf = 0.38. 1 H NMR (CDCl3 /TMS): δ 7.71 (1H), 7.52 (1H), 7.47 (2H), 7.33 (1H), 6.27 (1H), 5.16 (1H), 4.50 (2H), 4.31 (2H), 4.24 (1H), 3.80 (1H), 2.64 (1H), 2.01 (7H), 1.79 (3H), 1.37 (3H). HRMS (EI): calcd. for C23 H27 N3 O9 (M+ ) 489.175; found 489.174.
Deacetylate and tritylate 27. Prepare a mixture of 20 mL MeOH and 6 mL of 33% aq. NH4 OH. 28. In a 100-mL round-bottom flask, dissolve 2.65 g (5.58 mmol) S.6a or 2.73 g (5.58 mmol) S.6b in this mixture. Add a stir bar and stir for 2.5 hr at room temperature. 29. Check for complete deprotection by TLC using 1:1 (v/v) cyclohexane/acetone. Upon inspection under UV light (254 nm), there should be a single spot at Rf = 0.2.
30. Remove the solvent using a rotary evaporator connected to a vacuum pump. The bath temperature should be <40◦ C. Ensure that the exhaust fumes of the vacuum pump are led into the fume hood.
31. Dissolve the residue in 30 mL MeOH and evaporate the solvents again completely on a rotary evaporator connected to a vacuum pump. Repeat this step at least two additional times. After this step, clean the rotary evaporator well to ensure that no traces of NH4 OH are left.
32. Dissolve the residue in 30 mL pyridine and add a stir bar and a few crystals of DMAP. 33. Cool the solution by putting the flask in an ice-water bath. 34. Suspend 3.8 g (11.2 mmol) DMTr-Cl in 20 mL pyridine and add the suspension to the reaction mixture. Continue stirring overnight in the ice-water bath. The ice will melt overnight and the reaction temperature will gradually rise to room temperature.
35. Check for an almost complete conversion by TLC using 1:2 (v/v) cyclohexane/acetone with 1% Et3 N as eluent. Traces of pyridine can be removed before developing the TLC by putting the TLC plate in a round-bottom flask of appropriate size and applying a vacuum for several minutes. Under UV light (254 nm), the TLC will show three spots: the top spot is excess DMTr-Cl, the middle spot is the desired product, and the bottom spot is traces of remaining starting material.
36. Add 20 mL ethanol and evaporate the solvents using a rotary evaporator connected to a vacuum pump.
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37. Perform column chromatography using a 5 × 27–cm silica gel bed and 2.5 L of 1:1 (v/v) cyclohexane/acetone with 1% Et3 N as eluent. Analyze fractions by TLC. 38. Combine fractions that contain the main product and remove the solvent with a rotary evaporator connected to a vacuum pump. 39. Characterize the product by TLC, 1 H NMR, and MS. 5 -O-DMTr-TNPE (S.8a): Yield 3.3 g (84%). TLC (silica, 1:1 v/v hexane/acetone + 1% Et3 N): Rf = 0.34. 1 H NMR (CDCl3 /TMS): δ 7.80 (2H), 7.53 (2H), 7.30 (3H), 7.17 (7H), 6.73 (4H), 6.67 (1H), 6.2 (1H), 4.45 (1H), 4.05 (1H), 3.70 (6H), 3.32 (2H), 2.88 (1H, br s), 2.52 (1H), 2.12 (1H), 1.66 (3H), 1.53 (3H). MS (EI): 693.5 (M+ ). 5 -O-DMTr-TNPP (S.8b): Yield 3.7 g (94%). TLC (silica, 1:2 v/v cyclohexane/acetone + 1% Et3 N): Rf = 0.5. 1 H NMR (CDCl3 /TMS): δ 7.75 (1H), 7.65 (1H), 7.45 (2H), 7.27 (3H), 7.16 (7H), 6.72 (4H), 6.28 (1H), 4.45 (3H), 4.05 (1H), 3.70 (7H),3.35 (2H), 2.95 (1H, br s), 2.55 (1H), 2.15 (1H), 1.35 (6H). MS (FAB): 708.2 (MH+ ).
Prepare phosphoramidite 40. In a 50-mL round-bottom flask, dissolve 208 mg (0.3 mmol) S.8a or 212 mg (0.3 mmol) S.8b in 5 mL dichloromethane and add a stir bar. In steps 40 to 43, special care must be taken that pure and dry reagents are used and that the reaction mixture is kept under argon atmosphere at all times. Failure to do so will prevent any reaction.
41. Add 282 µL (1.65 mmol) DIPEA via a 500-µL syringe and allow to stir for 10 min at room temperature. 42. Add 134 µL (0.6 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite and continue stirring for 1.5 hr at room temperature. Ideally, this reagent should be completely clear. Proper handling is critical. Upon contact with air, the reagent becomes cloudy in a matter of minutes.
43. Check for complete conversion by TLC using 1:1 (v/v) cyclohexane/acetone with 1% Et3 N as eluent. Upon inspection under UV light (254 nm), no starting material (Rf = 0.3) should be detectable; rather, there should be a new spot at Rf = 0.6. At this step, the reaction mixture can again be handled in air, but steps 44 to 52 should be performed without unnecessary interruption.
44. Dilute the reaction mixture with 80 mL dichloromethane and transfer it into a separatory funnel. 45. Add 50 mL saturated aqueous NaHCO3 , shake gently, and separate the layers. Extract the aqueous phase two additional times with 50 mL dichloromethane each time. 46. Combine the organic phases and dry them with MgSO4 . Filter off the drying agent. 47. Remove the solvent from the filtrate using a rotary evaporator connected to a vacuum pump. 48. Perform column chromatography using a 3 × 22–cm silica gel bed and 0.5 L of 3:2 (v/v) cyclohexane/acetone with 1% Et3 N as eluent. Analyze fractions by TLC. 49. Combine fractions that contain the main product and remove the solvent with a rotary evaporator connected to a vacuum pump.
Nucleobase-Caged Phosphoramidites
50. To remove traces of volatile components, dissolve the product in 30 mL dichloromethane and remove the solvents again, first on a rotary evaporator
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connected to a vacuum pump and then under high vacuum. Repeat this procedure at least two additional times. This will result in a colorless brittle foam.
51. Characterize the product by TLC, 1 H NMR, 31 P NMR, and MS. 5 -O-DMTr-TNPE -3 -O-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (S.1a): Yield 255 mg (95%). TLC (silica, 1:1 v/v hexane/acetone + 1% Et3 N): Rf = 0.6. 1 H NMR (CDCl3 /TMS): δ 7.57 (14H), 6.80 (5H), 6.30 (1H), 4.60 (1H), 4.16 (1H), 3.77 (7H), 3.55 (4H), 3.33 (1H), 2.65 (2H), 2.41 (1H), 2.22 (1H), 1.75 (3H), 1.58 (3H), 1.11 (12H). 31 P NMR (CDCl3 /H3 PO4 ): δ 151.45, 151.47, 152.07, 152.16. 5 -O-DMTr-TNPP -3 -O-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (S.1b): Yield 256 mg (94%). TLC (silica, 1:1 v/v cyclohexane/acetone + 1% Et3 N): Rf = 0.6. 1 H NMR (CDCl3 /TMS): δ 7.50 (14H), 6.83 (4H), 6.38 (1H), 4.58 (3H), 4.16 (1H), 3.80 (8H), 3.54 (4H), 3.33 (1H), 2.67 (2H), 2.40 (1H), 2.25 (1H), 1.41 (6H), 1.12 (12H). 31 P NMR (CDCl3 /H3 PO4 ): δ 151.46, 151.94, 152.05, 152.05. MS (FAB): 908.5 (MH+ ).
52. Dissolve the foam under argon in the appropriate amount of dry acetonitrile to form a 0.1 M solution (in this case 2.8 mL). To ensure that there are no insoluble residues, the solution can be centrifuged or filtered through a syringe filter. The clear solution is ready for use on an automated DNA synthesizer (APPENDIX 3C). Due to the base lability of the caging groups on O4 of thymidine, a different set of protecting groups must be used for the uncaged nucleobases so that conditions not harsher than NH4 OH (33%) at room temperature will be sufficient for deprotection. The Ultramild protecting groups, for example, satisfy this requirement (phosphoramidites are available from Glen Research).
PREPARATION OF A PROTECTED NPP-CAGED DEOXYGUANOSINE PHOSPHORAMIDITE
BASIC PROTOCOL 2
The synthesis scheme for the deoxyguanosine phosphoramidite S.2 is shown in Figure 1.17.3. This phosphoramidite is used for synthesis of oligonucleotides containing dGNPP caged residues.
Materials 2 -Deoxyguanosine (S.9) Pyridine N,N-Dimethylformamide (DMF) Imidazole tert-Butyldimethylsilyl chloride (TBDMS-Cl) Dichloromethane (CH2 Cl2 ) Methanol (MeOH) Ethanol (EtOH), cold (4-Isopropylphenoxy)acetyl chloride (iPrPacCl; S.30; see Support Protocol 4) Cyclohexane Acetone Tetrahydrofuran (THF) 2-(2-Nitrophenyl)propan-1-ol (NPP-OH; S.24; see Support Protocol 2) Triphenylphosphine 40% diethylazodicarboxylate (DEAD) in toluene Ethyl acetate Brine (saturated NaCl) Anhydrous MgSO4 Glacial acetic acid
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Figure 1.17.3 Overview of the synthesis of the protected phosphoramidite S.2, for the introduction of dGNPP into oligonucleotides, starting from deoxyguanosine (S.9). DEAD, diethylazodicarboxylate; DMTr-Cl, 4,4 -dimethoxytrityl chloride; NPP, 2-(2-nitrophenyl)propyl; Pac, phenoxyacetyl; TBAF, tetrabutylammonium fluoride; TBDMS-Cl, tert-butyldimethylsilyl chloride.
1 M tetrabutylammonium fluoride (TBAF) in THF 4,4 -Dimethoxytrityl chloride (DMTr-Cl) Triethylamine (Et3 N) N,N-Diisopropylethylamine (DIPEA, H¨unig’s base) 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite Saturated aqueous NaHCO3 Dry acetonitrile 50-, 100-, and 250-mL round-bottom flasks Rotary evaporator connected to a vacuum pump High vacuum B¨uchner funnel connected to a vacuum pump Water aspirator Syringes Separatory funnels Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Silylate deoxyguanosine 1. In a 100-mL round-bottom flask, suspend 5 g (18.6 mmol) 2 -deoxyguanosine (S.9) in 50 mL pyridine and remove the solvent with a rotary evaporator connected to a vacuum pump. Repeat this procedure two additional times and dry the residue well using a high vacuum. Nucleobase-Caged Phosphoramidites
This step is for the azeotropic removal of any water.
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2. Suspend the residue in 40 mL DMF and add 8.3 g (122.6 mmol) imidazole and 10.6 g (70.6 mmol) TBDMS-Cl. Add a stir bar and stir 20 hr at room temperature. The deoxyguanosine will not dissolve completely at the beginning, but the reaction mixture will slowly become clear upon stirring.
3. Check for complete conversion by TLC using 9:1 (v/v) CH2 Cl2 /MeOH as eluent. Traces of pyridine can be removed before developing the TLC by placing the TLC plate in a round-bottom flask of appropriate size and applying vacuum for several minutes. Upon inspection under UV light (254 nm), the starting material should appear as a baseline spot and the product at Rf = 0.6.
4. Add 50 mL EtOH and place the flask in an ice-water mixture. The formation of some precipitate can already be observed in the reaction mixture, and this significantly intensifies upon addition of EtOH.
5. Filter the precipitate off using a B¨uchner funnel connected to a vacuum pump and wash it with a little cold (4◦ C) EtOH (10 mL). 6. Dry the colorless powder first with a water aspirator and later under high vacuum. 7. Characterize the product by TLC and 1 H NMR. 3 ,5 -Di-O-TBDMS-2 -dG (S.10): Yield 7.1 g (77%). TLC (silica, 9:1 v/v CH2 Cl2 /MeOH): Rf = 0.6. 1 H NMR (DMSO/DMSO): δ 10.60 (1H, s), 7.88 (1H, s), 6.47 (1H, s), 6.10 (1H, dd, J = 7.6, 6.4 Hz), 4.48 (1H, m), 3.80 (1H, m), 3.67 (2H, m), 2.64 (1H, m), 2.22 (1H, m), 0.88 (9H, s), 0.86 (9H, m), 0.10 (6H, s), 0.04 (3H, s), 0.03 (3H, s).
Perform N2 -protection 8. In a 250-mL round-bottom flask dissolve 3 g (6.05 mmol) S.10 in 40 mL pyridine. Add a stir bar and place the flask in an ice-water bath. 9. Add 1.93 g (9.1 mmol) of (4-isopropylphenoxy)acetyl chloride (S.30) dropwise using a syringe. 10. Stir for 2 hr, keeping the reaction flask in the ice-water bath. Then, take the flask out of the bath and continue stirring for an additional 1 hr. 11. Check for complete conversion by TLC using 19:1 (v/v) CH2 Cl2 /MeOH as eluent. Traces of pyridine can be removed before developing the TLC by placing the TLC plate in a round-bottom flask of appropriate size and applying vacuum for several minutes. Upon inspection under UV light (254 nm), the product will show up at Rf = 0.3. The spot above results from excess acid chloride (S.30).
12. Cool the reaction mixture by placing the flask in an ice-water bath. Then, add 120 mL MeOH dropwise. 13. Remove the solvents using a rotary evaporator connected to a vacuum pump. 14. Dissolve the residue in 50 mL MeOH and remove the solvent using the rotary evaporator. This step helps to remove traces of pyridine.
15. Perform column chromatography using a 5 × 30–cm silica gel bed and a gradient from 3:1 to 2:1 (v/v) cyclohexane/acetone as eluent. Analyze fractions by TLC. 16. Combine fractions that contain the main product and remove the solvent with a rotary evaporator connected to a vacuum pump. Synthesis of Modified Nucleosides
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17. Characterize the product by TLC, 1 H NMR, and MS. 3 ,5 -Di-O-TBDMS-N-(iPrPac)-2 -dG (S.11): Yield 2.2 g (54%). TLC (silica, 95:5 v/v CH2 Cl2 /MeOH): Rf = 0.33. 1 H NMR (CDCl3 /TMS): δ 11.75 (1H, br s), 9.06 (1H, br s), 7.99 (1H, s), 7.22 (2H, d, J = 8.6 Hz), 6.93 (2H, J = 8.6 Hz), 6.28 (1H, t, J = 6.5 Hz), 4.66 (2H, s), 4.49 (1H, m), 3.87 (1H, m), 3.76 (2H, m), 2.88 (1H, sept, J = 6.9 Hz), 2.42 (2H, m), 1.22 (6H, d, J = 6.9 Hz), 0.89 (18H, s), 0.07 (12H, s). HRMS (EI): calcd. for C33 H53 N5 O6 Si2 (M+ ) 671.3534; found 671.3515.
Add caging group by O6 -modification 18. In a 100-mL round-bottom flask dissolve 2 g (2.98 mmol) S.11 in 15 mL THF and add a stir bar to the flask. 19. Add 0.81 g (4.47 mmol) NPP-OH (S.24) and 1.17 g (4.47 mmol) triphenylphosphine. 20. While stirring, slowly add 2.05 mL (4.47 mmol) of 40% DEAD in toluene. 21. Continue stirring for 20 min at room temperature and then check for complete conversion by TLC using 3:1 (v/v) cyclohexane/ethyl acetate as eluent. Upon inspection under UV light (254 nm), the product spot is visible at Rf = 0.37.
22. Transfer the reaction mixture to a separatory funnel with a total of 150 mL CH2 Cl2 . Wash the organic phase with 100 mL water and then with 100 mL brine. 23. Dry the organic phase using MgSO4 and filter off the drying agent. 24. Remove the organic solvent from the filtrate using a rotary evaporator connected to a vacuum pump. 25. Purify the residue by column chromatography using a 5 × 30–cm silica gel bed and a gradient from 3:1 to 1:1 (v/v) cyclohexane/ethyl acetate as eluent. Analyze fractions by TLC. 26. Combine fractions that contain the main product and remove the solvent with a rotary evaporator connected to a vacuum pump. 27. Characterize the product by TLC, 1 H NMR, and MS. 3 ,5 -Di-O-TBDMS-N-(iPrPac)-2 -dGNPP (S.12): Yield 1.7 g (69%). TLC (silica, 3:1 v/v cyclohexane/ethyl acetate): Rf = 0.37. 1 H NMR (CDCl3 /TMS): δ 8.85 (1H, br s), 8.13 (1H), 7.72 (1H), 7.62 (1H), 7.53 (1H), 7.33 (1H), 7.19 (2H), 6.96 (2H), 6.31 (1H), 4.72 (5H), 4.00 (2H), 3.84 (2H), 2.88 (1H), 2.55 (2H), 1.53 (3H), 1.24 (6H), 0.91 (18H), 0.09 (12H). MS (EI): 834.4 (M+ ).
Desilylate 28. In a 50-mL round-bottom flask, dissolve 0.5 g (0.6 mmol) S.12 in 15 mL THF. Add a stir bar and chill the reaction mixture with stirring in an ice-water bath. 29. Add 0.2 mL (3.6 mmol) glacial acetic acid and then slowly add 1.8 mL (1.8 mmol) of 1 M TBAF in THF. 30. Remove the ice-water bath and continue stirring for 24 hr at room temperature. 31. Check for complete conversion by TLC using 9:1 (v/v) CH2 Cl2 /MeOH as eluent. Upon inspection under UV light (254 nm), the product spot is visible at Rf = 0.32.
32. Remove the organic solvents using a rotary evaporator connected to a vacuum pump.
Nucleobase-Caged Phosphoramidites
33. Purify the residue by column chromatography using a 3 × 26–cm silica gel bed and a gradient from 99:1 to 92.5:7.5 (v/v) CH2 Cl2 /MeOH as eluent. Analyze fractions by TLC.
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34. Combine fractions that contain the main product and remove the solvent with a rotary evaporator connected to a vacuum pump. 35. Characterize the product by TLC, 1 H NMR, and MS. N-(iPrPac)-2 -dGNPP (S.13): Yield 0.36 g (99%). TLC (silica, 9:1 v/v CH2 Cl2 /MeOH): Rf = 0.32. 1 H NMR (CDCl3 /TMS): δ 9.04 (1H), 8.03 (1H), 7.71 (1H), 7.62 (1H), 7.53 (1H), 7.33 (1H), 7.18 (2H), 6.96 (2H), 6.35 (1H), 4.96 (1H), 4.72 (4H), 4.13 (1H), 3.91 (3H), 2.99 (1H), 2.88 (1H), 2.41 (1H), 1.49 (3H), 1.22 (6H). HRMS (EI): calcd. for C30 H34 N6 O8 (M+ ) 606.2438; found 606.2439.
Tritylate 36. In a 50-mL round-bottom flask, dissolve 250 mg (0.41 mmol) S.13 in 5 mL pyridine and remove the solvent using a rotary evaporator connected to a vacuum pump. 37. Repeat this procedure two additional times, then dry the residue well under a high vacuum. 38. Dissolve the residue in 8 mL pyridine and add 168 mg (0.49 mmol) DMTr-Cl. Add a stir bar and stir 15 hr at room temperature. 39. Check for complete conversion by TLC using 95:5 (v/v) CH2 Cl2 /MeOH with 1% Et3 N as eluent. Upon inspection under UV light (254 nm), the product spot should be visible at Rf = 0.29.
40. Add 10 mL MeOH to the reaction mixture and remove the solvents in a rotary evaporator connected to a vacuum pump. 41. Add another 10 mL MeOH to the residue and remove the solvent on the rotary evaporator. This will help to remove traces of pyridine for the subsequent chromatography step.
42. Purify the residue by column chromatography using a 2 × 26–cm silica gel bed and 99:1 (v/v) CH2 Cl2 /MeOH with 1% Et3 N as eluent. Analyze fractions by TLC. 43. Combine fractions that contain the main product and remove the solvent with a rotary evaporator. 44. Characterize the product by TLC, 1 H NMR, and MS. 5 -O-DMTr-N-(iPrPac)-2 -dGNPP (S.14): Yield 319 mg (85%). TLC (silica, 95:5 v/v CH2 Cl2 /MeOH + 1% Et3 N): Rf = 0.29. 1 H NMR (CDCl3 /TMS): δ 8.93 (1H), 8.03 (1H), 7.71 (1H), 7.61 (1H), 7.53 (1H), 7.30 (12H), 6.95 (2H), 6.77 (4H), 6.65 (1H), 4.70 (5H), 4.23 (1H), 4.00 (1H), 3.74 (6H), 3.50 (1H), 3.37 (2H), 2.87 (1H), 2.67 (2H), 1.49 (3H), 1.22 (6H). MS (FAB): 909.4 (MH+ ).
Prepare phosphoramidite 45. In a 50-mL round-bottom flask dissolve 100 mg (0.11 mmol) S.14 in 2 mL THF and add a stir bar. In steps 45 to 48, special care must be taken that pure and dry reagents are used and the reaction mixture must be kept under argon atmosphere at all times. Failure to do so will prevent any reaction.
46. Add 66 µL (0.39 mmol) DIPEA via syringe and stir 10 min at room temperature. 47. Add 49 µl (0.22 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite and continue stirring for 1.5 hr at room temperature. Ideally, this reagent should be completely clear. Proper handling is critical. Upon contact with air, the reagent becomes cloudy in a matter of minutes.
Synthesis of Modified Nucleosides
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48. Check for complete conversion by TLC using 1:4 (v/v) cyclohexane/ethyl acetate with 1% Et3 N as eluent. Upon inspection under UV light (254 nm), the product spot will be visible at Rf = 0.5. At this step, the reaction mixture can again be handled in air, but steps 49 to 57 should be performed without unnecessary interruption.
49. Transfer the reaction mixture to a separatory funnel using 100 mL ethyl acetate. 50. Add 100 mL saturated aqueous NaHCO3 solution, shake gently, and separate the layers. Extract the aqueous phase two additional times with 50 mL ethyl acetate each time. 51. Combine the organic phases, wash with brine, and dry with MgSO4 . Filter off the drying agent. 52. Remove the solvent from the filtrate using a rotary evaporator connected to a vacuum pump. 53. Perform column chromatography using a 2 × 20–cm silica gel bed and a gradient from 4:1 to 1:4 (v/v) cyclohexane/ethyl acetate with 1% Et3 N as eluent. Analyze fractions by TLC. 54. Combine fractions that contain the main product and remove the solvent with a rotary evaporator connected to a vacuum pump. 55. To remove traces of volatile components, dissolve the product in 30 mL dichloromethane and remove the solvents again, first on a rotary evaporator and then under high vacuum. Repeat this procedure at least two times. This will result in a colorless brittle foam.
56. Characterize the product by TLC, 1 H NMR, 31 P NMR, and MS. 5 -O-DMTr-2 -dGNPP -3 -O-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (S.2): Yield 99 mg (81%). TLC (silica, 1:4 v/v cyclohexane/ethyl acetate + 1% Et3 N): Rf = 0.5. 1 H NMR (CDCl3 /TMS): δ 8.78 (1H), 8.01 (1H), 7.73 (1H), 7.63 (1H), 7.53 (1H), 7.30 (12H), 6.96 (2H), 6.78 (4H), 6.43 (1H), 4.76 (5H), 4.27 (1H), 4.00 (1H), 3.76 (6H), 3.76 (2H), 3.58 (2H), 3.36 (2H), 2.89 (1H), 2.62 (1H), 2.72 (2H), 2.45 (1H), 1.52 (3H), 1.24 (6H), 1.14 (12H). 31 P NMR (CDCl3 /H3 PO4 ): δ 151.84. MS (FAB): 1109.5 (MH+ ).
57. Dissolve the foam under argon in the appropriate amount of dry acetonitrile to form a 0.1 M solution (in this case 0.9 mL). To ensure that there are no insoluble residues, the solution can be centrifuged or filtered through a syringe filter. The clear solution is ready for use on an automated DNA synthesizer (APPENDIX 3C). BASIC PROTOCOL 3
PREPARATION OF A PROTECTED NPE-CAGED DEOXYCYTIDINE PHOSPHORAMIDITE The scheme for synthesizing the deoxycytidine phosphoramidite S.3 from 2 deoxyuridine is shown in Figure 1.17.4. This phosphoramidite is used for synthesis of oligonucleotides containing dCNPE caged residues.
Materials
Nucleobase-Caged Phosphoramidites
2 -Deoxyuridine (S.15) Pyridine 4,4 -Dimethoxytrityl chloride (DMTr-Cl) Dichloromethane (CH2 Cl2 ) Methanol (MeOH) Ethanol (EtOH)
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O
O NH
HO
N
O
O
O NH
DMTr-Cl pyridine
N
DMTrO
OH 15
O
O
NH Ac2O pyridine
N
DMTrO
OH 16
OAc 17
NO2
O S O O
HN N
N Me3C6H2SO2Cl DMAP, Et 3N
N
DMTrO
O
NPE-NH2
O
N
DMTrO
OAc 18
O
O OAc 19
NO2
HN-NPE
HN
N N
MeNH2 EtOH
DMTrO
N
O
O
O
N
DMTrO
CEOP(Cl)NiPr2
O
O
O OH 20
NC
O
O P NiPr2
3
Figure 1.17.4 Overview of the synthesis of the protected phosphoramidite S.3, for the introduction of dCNPE into oligonucleotides, starting from deoxyuracil (S.15). DMAP, 4-(N,N-dimethylamino)pyridine; DMTr-Cl, 4,4 dimethoxytrityl chloride; NPE, 1-(2-nitrophenyl)ethyl.
Ethyl acetate Saturated aqueous NaHCO3 Brine (saturated NaCl) Anhydrous MgSO4 n-Hexane 4-(N,N-Dimethylamino)pyridine (DMAP) Acetic anhydride (Ac2 O) Triethylamine (Et3 N) 5% (w/v) aqueous citric acid 2-Mesitylenesulfonyl chloride (Me3 C6 H2 SO2 Cl) N,N-Dimethylformamide (DMF) 1-(2-Nitrophenyl)ethylamine (NPE-NH2 ; S.26; see Support Protocol 3) 33% methylamine in ethanol N,N-Diisopropylethylamine (DIPEA, H¨unig’s base) 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite Cyclohexane Acetone Acetonitrile 50-mL, 250-mL, and 1-L round-bottom flasks Rotary evaporator connected to a vacuum pump Glass sinter filter High vacuum Syringes
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Reflux condenser 90◦ C oil bath Silicone septa, optional Separatory funnel Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Tritylate deoxyuridine 1. In a 250-mL round-bottom flask, dissolve 5 g (22 mmol) 2 -deoxyuridine (S.15) in 160 mL pyridine, and remove ∼60 mL with a rotary evaporator connected to a vacuum pump. This step is for the azeotropic removal of any water.
2. Add 8.91 g (27 mmol) DMTr-Cl and a stir bar. Stir 4 hr at room temperature. 3. Check for complete conversion by TLC using 95:5 (v/v) CH2 Cl2 /MeOH as eluent. Traces of pyridine can be removed before developing the TLC by placing the TLC plate in a round-bottom flask of appropriate size and applying vacuum for several minutes. Upon inspection under UV light (254 nm), the starting material would appear as a baseline spot and the product will show up at Rf = 0.25.
4. Add 25 mL ethanol and continue stirring 15 min at room temperature. 5. Remove all solvents using a rotary evaporator connected to a vacuum pump. 6. Dissolve the residue with 300 mL ethyl acetate. Wash the organic phase two times with 300 mL saturated aqueous NaHCO3 and then one time with 300 mL brine. 7. Dry the organic phase with anhydrous MgSO4 and filter off the drying agent. 8. Remove the organic solvent from the filtrate using a rotary evaporator connected to a vacuum pump. 9. Dissolve the residue in 100 mL ethyl acetate. 10. Prepare a 1-L round-bottom flask with 500 mL n-hexane, add a stir bar, and begin stirring vigorously. Add the product dropwise to this flask. A colorless precipitate will form.
11. Collect the precipitate by filtration using a glass sinter filter and dry well under high vacuum. 12. Characterize the product by TLC and 1 H NMR. 5 -O-DMTr-2 -dU (S.16): Yield 11.3 g (97%). TLC (silica, 95:5 v/v CH2 Cl2 /MeOH): Rf = 0.25. 1 H NMR (DMSO/DMSO): δ 11.31 (1H), 7.63 (1H), 7.31 (10H), 6.89 (3H), 6.14 (1H), 5.37 (1H), 5.32 (1H), 4.28 (1H), 3.87 (1H), 3.73 (6H), 3.20 (2H), 2.18 (2H).
Acetylate 13. In a 250-mL round-bottom flask, dissolve 10.25 g (19.3 mmol) S.16 in 60 mL pyridine. 14. Add 244 mg (2.0 mmol) DMAP followed by 2.7 mL (29 mmol) acetic anhydride. Place a stir bar into the flask and stir the reaction mixture 3 hr at room temperature. 15. Check for complete conversion by TLC using 98:2 (v/v) CH2 Cl2 /MeOH with 1% Et3 N as eluent. Nucleobase-Caged Phosphoramidites
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Traces of pyridine can be removed before developing the TLC by placing the TLC plate in a round-bottom flask of appropriate size and applying vacuum for several minutes. Upon inspection under UV light (254 nm), the starting material would appear at Rf = 0.07 and the product will show up at Rf = 0.24. Current Protocols in Nucleic Acid Chemistry
16. Add 2 mL methanol to the reaction mixture and remove the solvent in a rotary evaporator connected to a vacuum pump. 17. Dissolve the residue in 200 mL dichloromethane and wash this organic phase with 200 mL of 5% aqueous citric acid, followed by 200 mL water, and then 200 mL saturated aqueous NaHCO3 . In the last extraction, care must be taken to prevent spillage due to overpressure in the separatory funnel.
18. Dry the organic phase with MgSO4 and filter off the drying agent. 19. Remove the solvent from the filtrate in a rotary evaporator connected to a vacuum pump. When most of the solvent has been removed, continue drying under high vacuum. This affords compound S.17 as a yellowish foam that does not need to be purified further.
20. Characterize the product by TLC, 1 H NMR, and MS. 5 -O-DMTr-3 -O-acetyl-2 -dU (S.17): Yield 11.6 g (quant.). TLC (silica, 98:2 v/v CH2 Cl2 /MeOH + 1% Et3 N): Rf = 0.24. 1 H NMR (DMSO/DMSO): δ 11.38 (1H), 7.63 (1H), 7.14 (13H), 6.14 (1H), 5.45 (1H), 5.24 (1H), 4.07 (1H), 3.73 (6H), 3.29 (2H), 2.38 (2H), 2.04 (3H). HRMS (EI): calcd. for C32 H32 N2 O8 (M+ ) 572.2158; found 572.2156.
Add caging group by O4 -modification 21. In a 50-mL round-bottom flask, dissolve 1.0 g (1.75 mmol) S.17 in 10 mL dichloromethane and add 21.3 mg (0.17 mmol) DMAP. Add a stir bar and start stirring at room temperature. 22. Add 1.22 mL (8.75 mmol) Et3 N, followed by 498 mg (2.28 mmol) 2mesitylenesulfonyl chloride. Continue stirring for 2 hr at room temperature. The reaction solution will become dark red.
23. Check for complete conversion by TLC using 98:2 (v/v) CH2 Cl2 /MeOH with 1% Et3 N as eluent. Upon inspection under UV light (254 nm), the starting material would appear at Rf = 0.24 and the product will show up at Rf = 0.60.
24. Dilute the reaction mixture with 150 mL dichloromethane and wash this organic phase with saturated aqueous NaHCO3 . 25. Dry the organic phase with MgSO4 and filter of the drying agent. 26. Remove the solvent from the filtrate using a rotary evaporator connected to a vacuum pump. 27. Perform column chromatography using a 3 × 22–cm silica gel bed and a gradient from CH2 Cl2 to 99:1 (v/v) CH2 Cl2 /MeOH to 98:2 (v/v) CH2 Cl2 /MeOH, each with 1% Et3 N, as eluent. Analyze fractions by TLC. It is sufficient if this purification step separates the remaining 2-mesitylenesulfonyl chloride with Rf = 0.90 (silica, 98:2 v/v CH2 Cl2 /MeOH + 1% Et3 N). Under these conditions, the intermediate S.18 runs at Rf = 0.60.
28. Combine all product-containing fractions and remove the solvent using a rotary evaporator connected to a vacuum pump. This crude product (750 mg) can be used without any further purification and characterization.
Synthesis of Modified Nucleosides
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29. Dissolve the crude product in 4.5 mL DMF, add a stir bar, and start stirring at room temperature. 30. Dissolve 415 mg (2.5 mmol) NPE-NH2 (S.26) in 7.5 mL DMF and add this solution dropwise with a syringe to the stirring solution. 31. Place a reflux condenser on the reaction flask and heat the reaction mixture with stirring for 45 min in a 90◦ C oil bath. 32. Check for complete conversion by TLC using ethyl acetate with 1% Et3 N as eluent. Traces of DMF can be removed before developing the TLC by placing the TLC plate in a round-bottom flask of appropriate size and applying vacuum for several minutes. Upon inspection under UV light (254 nm), the intermediate S.18 would appear at Rf = 0.66 and the product S.19 will show up at Rf = 0.17.
33. Remove the organic solvent with a rotary evaporator connected to a high vacuum pump. Dissolve the residue in 100 mL dichloromethane. 34. Wash the organic solution with 100 mL of 5% aqueous citric acid, then 100 mL water, and finally 100 mL saturated aqueous NaHCO3 . 35. Dry with MgSO4 and filter off the drying agent. 36. Remove the solvent from the filtrate using a rotary evaporator connected to a vacuum pump. 37. Perform column chromatography using a 3 × 25–cm silica gel bed and ethyl acetate with 1% Et3 N as eluent. Analyze fractions by TLC. 38. Combine fractions that contain the main product and remove the solvent with a rotary evaporator connected to a vacuum pump. 39. Characterize the product by TLC, 1 H NMR, and MS. 5 -O-DMTr-3 -O-acetyl-2 -dCNPE (S.19): Yield 453 mg (62%). TLC (ethyl acetate + 1% Et3 N): Rf = 0.17. 1 H NMR (DMSO/DMSO): δ 8.40 (1H), 7.93 (1H), 7.62 (4H), 7.13 (13H), 6.09 (1H), 5.69 (1H), 5.51 (1H), 5.20 (1H), 4.05 (1H), 3.75 (6H), 3.25 (2H), 2.24 (2H), 2.02 (3H), 1.49 (3H). MS (FAB): 721.2 (MH+ ).
Perform deprotection 40. In a 50-mL round-bottom flask dissolve 427 mg (0.592 mmol) S.19 in 5 mL of 33% methylamine in ethanol. Add a stir bar and close the reaction flask, preferably with a silicone septum. Stir the reaction mixture for 1 hr at room temperature. 41. Check for complete conversion by TLC using 96:4 (v/v) CH2 Cl2 /MeOH with 1% Et3 N as eluent. Upon inspection under UV light (254 nm), the intermediate S.19 would appear at Rf = 0.27 and the product S.20 will show up at Rf = 0.16.
42. Remove the solvent using a rotary evaporator connected to a vacuum pump. Make sure that the outlet of the vacuum pump leads into a fume hood, and rinse the rotary evaporator well after this procedure to remove remaining methylamine.
43. Perform column chromatography using a 3 × 25–cm silica gel bed and 98:2 (v/v) CH2 Cl2 /MeOH with 1% Et3 N as eluent. Analyze fractions by TLC. 44. Combine fractions that contain the main product and remove the solvent with a rotary evaporator connected to a vacuum pump. Nucleobase-Caged Phosphoramidites
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45. Characterize the product by TLC, 1 H NMR, and MS. 5 -O-DMTr-2 -dCNPE (S.20): Yield 402 mg (quant.). TLC (96:4 v/v CH2 Cl2 /MeOH + 1% Et3 N): Rf = 0.16. 1 H NMR (DMSO/DMSO):δ 8.35 (1H), 7.93 (1H), 7.61 (4H), 7.13 (13H), 6.08 (1H), 5.65 (1H), 5.50 (1H), 5.26 (1H), 4.22 (1H), 3.85 (1H), 3.75 (6H), 3.18 (2H), 1.96 (2H), 1.49 (3H). HRMS (ESI): calcd. for C38 H39 N4 O8 (MH+ ) 679.2768; found 679.2756.
Prepare phosphoramidite 46. In a 50-mL round-bottom flask, dissolve 129 mg (0.19 mmol) S.20 in 4 mL dichloromethane and add a stir bar. In steps 46 to 49, special care must be taken that pure and dry reagents are used and that the reaction mixture is kept under argon atmosphere at all times. Failure to do so will prevent any reaction.
47. Add 146 µL (0.89 mmol) DIPEA via a syringe and allow to stir for 10 min at room temperature. 48. Add 85 µl (0.38 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite and continue stirring for 1.5 hr at room temperature. Ideally, this reagent should be completely clear. Proper handling is critical. Upon contact with air the reagent becomes cloudy in a matter of minutes.
49. Check for complete conversion by TLC using 1:1 (v/v) cyclohexane/acetone with 1% Et3 N as eluent. No starting material (Rf = 0.1) should be detectable; there should be a new spot at Rf = 0.36. At this step, the reaction mixture can again be handled in air, but steps 50 to 58 should be performed without unnecessary interruption.
50. Dilute the reaction mixture with 100 mL dichloromethane and transfer to a separatory funnel. 51. Add 100 mL saturated aqueous NaHCO3 solution, shake gently, and separate the layers. Extract the aqueous phase two additional times with 50 mL dichloromethane each time. 52. Combine the organic phases, wash with brine, and then dry with MgSO4 . Filter off the drying agent. 53. Remove the solvent from the filtrate using a rotary evaporator connected to a vacuum pump. 54. Perform column chromatography using a 2 × 22–cm silica gel bed and 2:1 (v/v) cyclohexane/acetone with 1% Et3 N as eluent. Analyze fractions by TLC. 55. Combine fractions that contain the main product and remove the solvent with a rotary evaporator connected to a vacuum pump. 56. To remove traces of volatile components, dissolve the product in 30 mL dichloromethane and remove the solvents again, first on a rotary evaporator and then under high vacuum. Repeat this procedure at least two times. This will result in a colorless brittle foam.
57. Characterize the product by TLC, 1 H NMR, 31 P NMR, and MS. 5 -O-DMTr-2 -dCNPE -3 -O-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (S.3): Yield 115 mg (68%). TLC (silica, 1:1 v/v cyclohexane/acetone + 1% Et3 N): Rf = 0.36. 1 H NMR (DMSO/DMSO):δ 8.40 (1H), 7.93 (1H), 7.69 (3H), 7.5 (1H), 7.3 (9H), 6.9 (4H),
Synthesis of Modified Nucleosides
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6.10 (1H), 5.70 (1H), 5.52 (1H), 4.48 (1H), 4.00 (1H), 3.75 (6H), 3.70 (1H), 3.60 (1H), 3.50 (2H), 3.25 (2H), 2.75 (1H), 2.65 (1H), 2.30 (1H), 2.10 (1H), 1.50 (3H), 1.10 (12H). 31 P NMR: (DMSO/H3 PO4 ):δ 145.53, 145.18. MS (FAB): 879.4 (MH+ ).
58. Dissolve the foam under argon in the appropriate amount of dry acetonitrile to form a 0.1 M solution (in this case 1.3 mL). To ensure that there are no insoluble residues, the solution can be centrifuged or filtered through a syringe filter. The clear solution is ready for use on an automated DNA synthesizer (APPENDIX 3C). SUPPORT PROTOCOL 1
PREPARATION OF THE CAGING PRECURSOR 1-(2-NITROPHENYL)ETHAN-1-OL This protocol is taken from the “progenitor” caging paper by Kaplan et al. (1978). The single reaction for preparation of NPE-OH (S.22) from 2-nitroacetophenone is shown in Figure 1.17.5.
Materials 2-Nitroacetophenone (S.21) 1,4-Dioxane Methanol (MeOH) Sodium borohydride (NaBH4 ) Dichloromethane (CH2 Cl2 ) Anhydrous MgSO4 250-mL round-bottom flask Rotary evaporator connected to a vacuum pump Separatory funnel Additional reagents and equipment for TLC (APPENDIX 3D)
Figure 1.17.5 Overview of the synthesis of caging group precursors S.22, S.24, and S.26, as well as the acid chloride S.30 used for N-protection of guanosine. NBS, N-bromosuccinimide. Nucleobase-Caged Phosphoramidites
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1. In a 250-mL round-bottom flask, dilute 5 g (30.2 mmol) 2-nitroacetophone (S.21) in 75 mL of 2:3 (v/v) 1,4-dioxane/MeOH. Add a stir bar and begin stirring. 2. Cool the reaction mixture in an ice/water bath. 3. Add 3.5 g (92.5 mmol) NaBH4 in small portions. After addition is complete, continue stirring in the ice/water bath for an additional 30 min. Strong bubbles are observed after each addition. A precipitate will form after addition is complete.
4. Remove the cooling bath and continue stirring for 2 hr at room temperature. 5. Remove the solvent using a rotary evaporator connected to a vacuum pump. 6. Add 150 mL water and stir the mixture for 1.5 hr at room temperature. 7. Add 100 mL dichloromethane and transfer the mixture to a separatory funnel. Separate the layers and extract the aqueous phase two additional times with 100 mL dichloromethane. 8. Combine the organic layers and dry them with MgSO4 . Filter off the drying agent. 9. Remove the organic solvents from the filtrate using a rotary evaporator connected to a vacuum pump. 10. Characterize the product by TLC and 1 H NMR. 1-(2-Nitrophenyl)ethan-1-ol (S.22): Yield 4.64 g (92%). TLC (silica, chloroform): Rf = 0.20. 1 H NMR (CDCl3 /TMS):δ 7.82 (2H), 7.61 (1H), 7.39 (1H), 5.36 (1H), 2.76 (1H, br s), 1.51 (3H).
PREPARATION OF THE CAGING PRECURSOR 2-(2-NITROPHENYL)PROPAN-1-OL
SUPPORT PROTOCOL 2
The single reaction for preparation of NPP-OH (S.24) from 1-ethyl-2-nitrobenzene is shown in Figure 1.17.5. This protocol is adapted from Tsuji et al. (1990).
Materials 1-Ethyl-2-nitrobenzene (S.23) Dimethylsulfoxide (DMSO) Paraformaldehyde 40% Triton B (BuMe3 NOH) in methanol Cyclohexane Acetone 100-mL round-bottom flask Reflux condenser 90◦ C oil bath Distillation apparatus Additional reagents and equipment for TLC (APPENDIX 3D) 1. In a 100-mL round-bottom flask, dissolve 22.7 g (150 mmol) 1-ethyl-2-nitrobenzene (S.23) in 20 mL DMSO. Add a stir bar and begin stirring at room temperature. 2. Add 1.85 g (62 mmol) paraformaldehyde and 1.8 ml of 40% Triton B in MeOH. The reaction mixture will change color several times and become clear.
3. Add a reflux condenser and heat the reaction mixture 3 hr in a 90◦ C oil bath.
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4. Perform TLC with 3:2 (v/v) cyclohexane/acetone as eluent. Upon inspection under UV light (254 nm), apart from the spot resulting from excess starting material (Rf = 0.6), the product will result in a spot at Rf = 0.4.
5. Perform distillation under reduced pressure (1 mbar). Collect the main fraction, which distills at ∼140◦ C vapor temperature. 6. Characterize the product by TLC and 1 H NMR. 2-(2-Nitrophenyl)propan-1-ol (S.24): Yield 7.61 g (68%). TLC (3:2 v/v cyclohexane/acetone): Rf = 0.40. 1 H NMR (CDCl3 /TMS): δ 7.67 (1H), 7.48 (1H), 7.41 (1H), 7.28 (1H), 3.70 (2H), 3.45 (1H), 1.26 (3H). SUPPORT PROTOCOL 3
PREPARATION OF THE CAGING PRECURSOR 1-(2-NITROPHENYL)ETHYLAMINE The preparation of NPE-NH2 (S.26) is performed in two steps starting from 1-ethyl-2nitrobenzene and is shown in Figure 1.17.5.
Materials 1-Ethyl-2-nitrobenzene (S.23) Tetrachlorocarbon (CCl4 ) N-Bromosuccinimide (NBS) Dibenzoyl peroxide [(BzO)2 ] Hexane Acetone Cyclohexane N,N-Dimethylformamide (DMF) Potassium phthalimide Ethyl acetate Dichloromethane (CH2 Cl2 ) Saturated aqueous NaHCO3 Anhydrous MgSO4 Ethanol 80% hydrazinium hydroxide solution in water Methanol Triethylamine (Et3 N) 100- and 500-mL round-bottom flasks Reflux condensers 60◦ , 77◦ , and 155◦ C oil baths B¨uchner funnel connected to a vacuum pump Rotary evaporator connected to a vacuum pump Separatory funnels High vacuum Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Perform bromination 1. In a 500-mL round-bottom flask, dissolve 30 g (198 mmol) 1-ethyl-2-nitrobenzene (S.23) in 300 mL CCl4 . Add a stir bar and begin stirring at room temperature. CAUTION: Make sure to avoid all contact with the liquid or vapor, as CCl4 is toxic. Dispose of all waste properly. Nucleobase-Caged Phosphoramidites
2. Add 53 g (298 mmol) N-bromosuccinimide and 720 mg (3 mmol) dibenzoyl peroxide. Equip the flask with a reflux condenser and heat the reaction mixture to reflux (77◦ C) for 4 hr.
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3. Check for complete conversion by TLC using 9:1 (v/v) hexane/acetone as eluent. Upon inspection under UV light (254 nm), there should be no visible spot of the starting material (Rf = 0.88), and the product should be visible at Rf = 0.40.
4. Remove the oil bath and let the reaction mixture cool to room temperature. 5. Filter off any solid material using a B¨uchner funnel connected to a vacuum pump. This will result in a clear orange solution.
6. Remove the organic solvent using a rotary evaporator connected to a vacuum pump. This will afford the crude product as an orange oil.
7. Perform column chromatography using an 8 × 20–cm silica gel bed and 1.2 L of 9:1 (v/v) cyclohexane/acetone as eluent. Analyze fractions by TLC. 8. Combine fractions that contain the main product and remove the solvent with a rotary evaporator connected to a vacuum pump. 9. Characterize the product by TLC and 1 H NMR. 1-(2-Nitrophenyl)ethylbromide (S.25): Yield 46.5 g (quant.). TLC (9:1 v/v cyclohexane/acetone): Rf = 0.40. 1 H NMR (CDCl3 /TMS): δ 7.88 (1H), 7.80 (1H), 7.6 (1H), 7.40 (1H), 5.80 (1H), 2.08 (3H).
Perform substitution 10. In a 100-mL round-bottom flask, dissolve 5 g (21.7 mmol) S.25 in 20 mL DMF. Add a stir bar and begin stirring at room temperature. 11. Add 4.03 g (21.7 mmol) potassium phthalimide. 12. Add a reflux condenser and heat the reaction mixture 1 hr in a 155◦ C oil bath. 13. Remove the oil bath and allow it to cool to room temperature. 14. Check for complete conversion by TLC using 1:1 (v/v) cyclohexane/ethyl acetate as eluent. Upon inspection under UV light (254 nm), there should be no spots resulting from either the starting material (Rf = 0.9) or the phthalimide (Rf = 0.7), and a new spot should be visible at Rf = 0.8.
15. Add 400 mL dichloromethane and transfer the solution to a separatory funnel. Wash this organic phase first with saturated aqueous NaHCO3 and then with water. 16. Dry the organic phase with anhydrous MgSO4 and filter off the drying agent. 17. Remove the solvent from the filtrate using a rotary evaporator connected to a vacuum pump, and dry the residue under high vacuum. 18. Dissolve the residue in 80 mL ethanol, add a stir bar, and begin stirring at room temperature. 19. Add 1.36 mL of 80% hydrazinium hydroxide solution. Equip the flask with a reflux condenser and heat the reaction mixture 1.5 hr at 60◦ C. CAUTION: Make sure to avoid any contact with the liquid or vapor, as hydrazine is toxic.
20. Add an additional 1.36 mL of 80% hydrazinium hydroxide solution and continue heating for an additional 1 hr. 21. Allow solution to cool to room temperature.
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22. Check for complete conversion by TLC using 95:5 (v/v) CH2 Cl2 /MeOH with 0.5 % Et3 N as eluent. Upon inspection under UV light (254 nm), there should be a prominent product spot at Rf = 0.10 and only a few spots above it.
23. Remove any solid by filtering the suspension using a B¨uchner funnel connected to a vacuum pump. Wash the solid well with ethanol. 24. Remove the solvent from the combined filtrates using a rotary evaporator connected to a vacuum pump. 25. Dissolve the residue in 150 mL dichloromethane and transfer the solution to a separatory funnel. Wash this organic phase with 150 mL water. 26. Dry the organic phase using MgSO4 and filter off the drying agent. 27. Remove the solvent from the filtrate using a rotary evaporator connected to a vacuum pump. 28. Perform column chromatography using a 5 × 22–cm silica gel bed and 95:5 (v/v) CH2 Cl2 /MeOH with 0.5 % Et3 N as eluent. Analyze fractions by TLC. 29. Combine fractions that contain the main product and remove the solvent with a rotary evaporator connected to a vacuum pump. 30. Characterize the product by TLC and 1 H NMR. 1-(2-Nitrophenyl)ethylamine (S.26): Yield 1.3 g (36%). TLC (95:5 v/v CH2 Cl2 /MeOH + 0.5% Et3 N): Rf = 0.10. 1 H NMR (DMSO/DMSO): δ 7.90 (1H), 7.77 (1H), 7.68 (1H), 7.44 (1H), 4.32 (1H), 2.02 (2H), 1.03 (3H). SUPPORT PROTOCOL 4
PREPARATION OF (4-ISOPROPYLPHENOXY)ACETYL CHLORIDE FOR PROTECTION OF DEOXYGUANOSINE The acid chloride S.30 is prepared from 4-isopropylphenol in three steps as illustrated in Figure 1.17.5.
Materials 60% NaH in paraffin Tetrahydrofuran (THF) 4-Isopropylphenol (S.27) Bromoacetic acid methyl ester Diethyl ether Anhydrous MgSO4 1,4-Dioxane Lithium hydroxide Half-concentrated HCl Ethyl acetate Thionyl chloride (SOCl2 ) 100-, 250-, and 500-mL round-bottom flasks Dropping funnel Rotary evaporator connected to a vacuum pump Separatory funnels Reflux condenser 90◦ C oil bath Distillation bridge Nucleobase-Caged Phosphoramidites
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Perform acetylation 1. In a 250-mL round-bottom flask, suspend 5.88 g of 60% NaH in 50 mL THF. Add a stir bar and begin stirring at room temperature. Be extremely careful when handling NaH, and be certain to exclude any water from the reaction.
2. Equip the flask with a dropping funnel. Dissolve 20 g (147 mmol) 4-isopropylphenol (S.27) in 160 mL THF and add this solution carefully, dropwise, to the reaction mixture using the dropping funnel. 3. Continue stirring for 1.5 hr at room temperature. 4. Cool the reaction mixture using an ice/water bath. 5. Transfer 14.4 mL (147 mmol) bromoacetic acid methyl ester to the dropping funnel and carefully add dropwise to the reaction mixture. 6. Continue stirring overnight at room temperature. 7. Remove the solvent using a rotary evaporator connected to a vacuum pump. 8. To the residue, add 150 mL water and 250 mL diethyl ether. Transfer the mixture to a separatory funnel and separate the layers. Extract the aqueous phase two additional times with 250 mL diethyl ether each time. 9. Combine the organic extracts and dry with anhydrous MgSO4 . Filter off the drying agent. 10. Remove the solvent from the filtrate using a rotary evaporator connected to a vacuum pump. 11. Purify the crude product by distillation under reduced pressure (6 mbar). Collect the fraction that distills at ∼130◦ C vapor temperature. 12. Characterize the product by 1 H NMR. (4-Isopropylphenoxy)acetic acid methyl ester (S.28): Yield 29.5 g (97%). 1 H NMR (CDCl3 /TMS): δ 7.05 (2H), 6.77 (2H), 4.53 (2H), 3.73 (3H), 2.80 (1H), 1.12 (6H).
Perform saponification 13. In a 500-mL round-bottom flask, dissolve 29.5 g (142 mmol) S.28 in 300 mL of 1,4-dioxane, add a stir bar, and begin stirring at room temperature. 14. Dissolve 4.06 g lithium hydroxide in 200 mL water and add this solution to the reaction mixture. Continue stirring for 13 hr at room temperature. 15. Remove the solvents using a rotary evaporator connected to a vacuum pump. 16. Adjust the reaction mixture to pH 1 by careful dropwise addition of half-concentrated HCl. 17. Transfer the reaction mixture to a separatory funnel and extract it three times with 300 mL ethyl acetate each time. 18. Combine the organic phases and dry with anhydrous MgSO4 . Filter off the drying agent. 19. Remove the solvent from the filtrate using a rotary evaporator connected to a vacuum pump. This results in a colorless solid.
Synthesis of Modified Nucleosides
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20. Characterize the product by 1 H NMR. (4-Isopropylphenoxy)acetic acid (S.29): Yield 27.1 g (98%). 1 H NMR (CDCl3 /TMS): δ 9.0 (br s, 1H), 7.08 (2H), 6.78 (2H), 4.6 (2H), 2.8 (1H), 1.16 (6H).
Convert to acid chloride 21. In a 100-mL round-bottom flask, dissolve 5 g (25.7 mmol) S.29 in 50 mL thionyl chloride. Add a stir bar and begin stirring at room temperature. CAUTION: Make sure to handle thionyl chloride very carefully and avoid any contact with liquid or vapor.
22. Equip the reaction flask with a reflux condenser and heat the reaction mixture 16 hr in a 90◦ C oil bath. 23. Allow reaction to cool to room temperature and exchange the reflux condenser with a distillation bridge. First distill off the excess thionyl chloride under ambient pressure. 24. Cooling to room temperature again, then carefully apply vacuum (1 mbar) to the reaction setup and continue distilling under reduced pressure. Collect the main fraction, which will distill at ∼112◦ C vapor temperature. 25. Characterize the product by 1 H NMR. (4-Isopropylphenoxy)acetyl chloride (S.30): Yield 5 g (91%). 1 H NMR (CDCl3 /TMS): δ 7.08 (2H), 6.73 (2H), 4.85 (2H), 2.77 (1H), 1.12 (6H).
COMMENTARY Background Information
Nucleobase-Caged Phosphoramidites
The term “caged compounds” for biologically active substances that have been temporarily inactivated and can be triggered by light-irradiation is a rather unfortunate choice. It is misleading because most chemists will think of cage compounds in a topological sense, and the lack of semantic precision in this term makes literature searches in the field difficult. The origin of the field dates back to the late 1970s, when Engels and co-workers prepared a cAMP bearing a photolabile group on the phosphate (Engels and Schlaeger, 1977), and J.F. Hoffman and co-workers synthesized ATP with a photolabile group on the γ -phosphate (Kaplan et al., 1978). Only the latter group called their compounds “caged.” Since that time, the concept of caging has been applied to a whole variety of compounds, including many small molecules, peptides, proteins, and, only recently, nucleic acids. For a deeper insight, especially in the field of caged oligo(deoxy)nucleotides, the reader is referred to available reviews (Pelliccioli and Wirz, 2002; Goeldner and Givens, 2005; Mayer and Heckel, 2006). Using light-reactive compounds (either irreversibly triggerable, caged, or reversibly switchable ones based, e.g., on azo benzene), intriguing experiments have become possible, such as the spatiotemporally controlled release of calcium or the light-driven transport of cal-
cium across membranes (Bennett et al., 2002). Trauner, Isacoff, and colleagues succeeded in making ion channels that can be activated by light (Volgraf et al., 2006), and Feringa, Meijberg, and colleagues were successful in a similar approach (Koc¸er et al., 2005), to name only a few recent highlights. The applicability of caging technology depends on the broad spectrum of tools used to generate and manipulate light, also with high spatiotemporal precision. In addition to expensive but powerful lasers, LEDs are now available that have excellent technical specifications at a price that is lower by two orders of magnitude. Experiments have already shown that they can, in fact, compete with the more established light sources (Bernardinelli et al., 2005).
Critical Parameters and Troubleshooting Phosphoramidite synthesis The flask-scale synthesis procedures reported in this unit are all very straightforward. The only step that is sometimes unsuccessful, especially for novice researchers, is the preparation of the phosphoramidites. It is crucial that the flask and the solvents be completely dry and that fresh reagents be used. As noted in the protocols, fresh 2-cyanoethylN,N-diisopropylchlorophosphoramidite is a
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colorless and clear liquid that becomes cloudy upon contact with air in a matter of minutes. It is best to use as little of this reagent as possible, and it is very important to remove any excess in the purification of the phosphoramidite. It is a good idea to always check the purity of the phosphoramidite via 31 P NMR. For NMR, some batches of CDCl3 contain traces of acid, which will result in spectra of decomposed phosphoramidite despite the fact that the amidite is ready to be used. As a solution, DMSO-d6 can be used as a substitute, or the CDCl3 can be filtered through neutral aluminium oxide. One should not be surprised by splitting of signals in the NMR spectra or by spots in the TLC resulting from the formation of diastereomeric mixtures upon introduction of the respective caging groups. However, the sole disadvantage of having slightly more complicated NMR spectra does not justify a stereoselective synthesis. For optimal results, the phosphoramidites described here should not be stored for more than several days. It is therefore advisable to keep a stock of the precursors (S.8a, S.8b, S.14, and S.20), and always prepare the respective phosphoramidites fresh just before the DNA synthesis. With storage at −20◦ C, no decomposition of precursors has been observed. Oligonucleotide synthesis The phosphoramidites can be used in regular solid-phase synthesis protocols. Prolonged coupling times are possible but not necessary. Due to the lability of the caging group under basic conditions, the presence of a TNPE or TNPP residue requires that a different set of nucleobase-protecting groups be used in the solid-phase synthesis. Ultramild protection has been shown to work very well. Ultramildprotected phosphoramidites for the uncaged nucleobases are commercially available from Glen Research. The deprotection conditions should not be harsher than 33% NH4 OH at room temperature. The dGNPP and dCNPE residues are much more tolerant towards base treatment and do not require any additional measures. For higher purity, a DMTr-on synthesis is advisable, followed by solution-phase DMTr-deprotection and repurification. Purification is best performed by RP-HPLC (UNIT 10.5) using a gradient between 0.1 M triethylammonium acetate (TEAA) and acetonitrile. One need not worry about unwanted uncaging due to the light from the detector. The same is true for melting point studies, which are performed by following the temperature-
dependent absorbance. Characterization of the caged oligonucleotides is conveniently performed by ESI-MS, since the samples can be injected right away after the aforementioned HPLC purification with TEAA buffer. Much better results are obtained, however, using an LC-ESI-MS coupling (in which case the aqueous buffer should have a concentration of only 5 mM TEAA). Caged oligonucleotides do not need to be handled in a dark room. In fact, the authors have never observed any unwanted uncaging. Nonetheless, caged oligonucleotides are stored in brown microcentrifuge tubes at –20◦ C without deleterious effects.
Anticipated Results With few exceptions, the reported procedures can be performed with good yields so that it is possible to make a sufficiently large stock of phosphoramidite precursors. The introduction of the caged phosphoramidites in the solid-phase synthesis is also usually unproblematic. The many different reaction steps in the solid-phase synthesis do lead to some loss of caging groups, but since each caging group leads to a very significant time shift in the RP-HPLC, these sequences can be separated very easily, and the peak with the highest retention time is usually by far the biggest peak.
Time Considerations The preparation of each caged phosphoramidite can be completed in ∼2 weeks. An additional 2 weeks are required for synthesis of the precursors described in the Support Protocols (S.22, S.24, S.26, and S.30). As mentioned previously, the phosphoramidites should not be stored but rather used right away. Thus, it is better to keep a stock of the respective precursors (S.8a, S.8b, S.14, and S.20) and prepare the phosphoramidites fresh. The solid-phase synthesis with subsequent deprotection, purification, DMTr-cleavage and repurification, characterization, and quantification will take ∼3 days.
Literature Cited Bennett, I.M., Farfano, H.M.V., Bogani, F., Primak, A., Liddell, P.A., Otero, L., Sereno, L., Silber, J.J., Moore, A.L., Moore, T.A., and Gust, D. 2002. Active transport of Ca2+ by an artificial photosynthetic membrane. Nature 420:398401. Bernardinelli, Y., Haeberli, C., and Chatton, J.Y. 2005. Flash photolysis using a light emitting diode: An efficient, compact, and affordable solution. Cell Calcium 37:565-572.
Synthesis of Modified Nucleosides
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Engels, J. and Schlaeger, E.J. 1977. Synthesis, structure, and reactivity of adenosine cyclic 3 ,5 -phosphate benzyl triesters. J. Med. Chem. 20:907-911.
Mayer, G., Kr¨ock, L., Mikat, V., Engeser, M., and Heckel, A. 2005. Light-induced formation of G-quadruplex DNA secondary structures. ChemBioChem 6:1966-1970.
Goeldner, M. and Givens, R. 2005. Dynamic studies in biology. Wiley-VCH, Weinheim, Germany.
Pelliccioli, A.P. and Wirz, J. 2002. Photoremovable protecting groups: Reaction mechanisms and applications. Photochem. Photobiol. Sci. 1:441458.
Heckel, A. and Mayer, G. 2005. Light-regulation of aptamer activity: An anti-thrombin aptamer with caged thymidine nucleobases. J. Am. Chem. Soc. 127:822-823. Heckel, A., Buff, M.C.R., Raddatz, M.-S.L., M¨uller, J., P¨otzsch, B., and Mayer, G. 2006. An anticoagulant with light-triggered antidote activity. Angew. Chem. Int. Ed. 45:6748-6750. Kaplan, J.H., Forbush, B. III, and Hoffman, J.F. 1978. Rapid photolytic release of adenosine 5 triphosphate from a protected analogue: Utilization by the Na:K pump of human red blood cell ghosts. Biochemistry 17:1929-1935. Koc¸er, A., Walko, M., Meijberg, W., and Feringa, B.L. 2005. A light-actuated nanovalve derived from a channel protein. Science 309:755-758. Kr¨ock, L. and Heckel, A. 2005. Photoinduced transcription by using temporarily mismatched caged oligonucleotides. Angew. Chem. Int. Ed. 44:471-473. Mayer, G. and Heckel, A. 2006. Biologically active molecules with a “light switch”. Angew. Chem. Int. Ed. 45:4900-4921.
Tsuji, Y., Kotachi, S., Huh, K.-T., and Watanabe, Y. 1990. 2-Aminophenethyl alcohols and 2-nitrophenyl alcohols. J. Org. Chem. 55:580584. Volgraf, M., Gorostiza, P., Numano, R., Kramer, R.H., Isacoff, E.Y., and Trauner, D. 2006. Allosteric control of an ionotropic glutamate receptor with an optical switch. Nat. Chem. Biol. 2:47-52. Walbert, S., Pfleiderer, W., and Steiner, U.E. 2001. Photolabile protecting groups for nucleosides: Mechanistic studies on the 2-(2nitrophenyl)ethyl group. Helv. Chim. Acta 84:1601-1611.
Contributed by Alexander Heckel University of Bonn Bonn, Germany
Nucleobase-Caged Phosphoramidites
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Synthesis of Altritol Nucleoside Phosphoramidites for Oligonucleotide Synthesis
UNIT 1.18
Mikhail Abramov1 and Piet Herdewijn1 1
Rega Institute for Medical Research, Leuven, Belgium
ABSTRACT This unit describes in detail the optimized preparations of altritol nucleoside phosphoramidite building blocks for oligonucleotide synthesis (aA, aG, aC, aU). D-Altritol nucleosides with adenine and uracil bases are obtained by nucleophilic opening of the epoxide ring of 1,5:2,3-dianhydro-4,6-O-benzylidene-D-allitol using the 1,8diazabicyclo[5.4.0]undec-7-ene salts of the above-mentioned salts, while phase transfer catalysis (18-crown-6, K2 CO3 ) is optimal for alkylation of 2-amino-6-chloropurine. The cytosine nucleoside is synthesized starting from the uracil congener. The 3 -hydroxyl function of hexitol sugar is protected with the benzoyl group. Curr. Protoc. Nucleic Acid C 2007 by John Wiley & Sons, Inc. Chem. 30:1.18.1-1.18.21. Keywords: altritol nucleic acid (ANA) r altritol nucleoside phosphoramidites r aA r aG r aC r aU
INTRODUCTION Altritol nucleic acids (ANA) represent a new oligomeric structure able to hybridize in a sequence-selective manner with RNA and with itself. In contrast to most chemically modified nucleic acids, ANA possess a free secondary hydroxyl group, which contributes to the stability of RNA-ANA duplexes. The hybridization strength with RNA is higher than with DNA (Allart et al., 1999b; Abramov et al., 2007b). ANA has superior siRNA activity (Fisher et al., 2007) and is an efficient template for nonenzymatic catalyzed polymerization reactions (Kozlov et al., 2000). ANAs are composed of phosphorylated 1,5-anhydro-2-deoxy-D-altro-hexitol building blocks with a base moiety in the 2 -position. During recent years the synthesis of hexitol nucleic acids (HNA) has been developed and described in detail (De Bouvere et al., 1997; UNIT 1.9). ANA nucleosides are very similar to HNA; they differ only by the presence of a supplementary hydroxyl group in the 3 −α-position. Due to the presence of this 3 -hydroxyl function, which requires selective protection, the synthesis of the D-altritol phosphoramidite building blocks takes more time than the synthesis of the 1,5-anhydrohexitol analogs. This unit describes in detail the preparation of the phosphoramidite building blocks (aU, aC, aA, and aG) used for ANA oligonucleotide synthesis (Allart et al., 1999a). Synthesis of the ANA nucleosides is presented in Basic Protocols 1 through 4, followed by a general procedure in Basic Protocol 5 for their phosphitylation, yielding the phosphoramidite derivatives. The chemical reactions described here have optimized yields, and purification of the intermediates is straightforward.
SYNTHESIS OF 1 ,5 -ANHYDRO-3 -O-BENZOYL-6 -OMONOMETHOXYTRITYL-2 -DEOXY-2 -(URACIL-1-YL)-D-ALTROHEXITOL Synthesis of the protected nucleoside from uracil is performed in four steps that are illustrated in Figure 1.18.1. Reaction of the DBU salt of uracil with 1,5:2,3-dianhydro-4,6O-benzylidene-D-allitol yields the altro-derivative (S.1). Treatment with excess benzoyl Current Protocols in Nucleic Acid Chemistry 1.18.1-1.18.21, September 2007 Published online September 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/0471142700.nc0118s30 C 2007 John Wiley & Sons, Inc. Copyright
BASIC PROTOCOL 1
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Figure 1.18.1 Preparation of protected aU. Bz, benzoyl; DBU, 1,8-diazabicyclo[5.4.0]undec-7ene; DCE, 1,2-dichloroethane; DMF, N,N-dimethylformamide; MMTr, monomethoxytrityl; Py, pyridine; TFA, trifluoroacetic acid.
chloride leads to protection of the 3 -hydroxyl group. Benzylidene cleavage with TFA followed by monomethoxytritylation provides aU (S.4).
Materials Uracil, 99+% 1,5:2,3-Dianhydro-4,6-O-benzylidene-D-allitol (CMS Chemical, http://www.cms-chemicals.com, or Brockway et al., 1984) N,N-Dimethylformamide (DMF), anhydrous Nitrogen gas 1,8-Diazabicyclo[5.4.0]undec-7-ene (DBU) Dry ice (solid CO2 for cooling bath) Acetic acid (AcOH) Brine (saturated aqueous sodium chloride solution) Ethyl acetate (EtOAc) Sodium sulfate (Na2 SO4 )Silica gel (0.060- to 0.200-nm) Dichloromethane PA (DCM) Methanol (MeOH) Anhydrous pyridine Benzoyl chloride (BzCl) Toluene Saturated aqueous sodium bicarbonate solution (sat. aq. NaHCO3 ) Chloroform Hexane PA 1,2-Dichloroethane, 99.8+% (DCE) Trifluoroacetic acid (TFA) Monomethoxytrityl chloride (MMTr-Cl) Synthesis of Altritol Nucleoside Phosphoramidites
100-, 250-, and 500-mL round-bottom flasks Gas balloon Dropping funnel
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Rotary evaporator equipped with a vacuum pump 5 × 35–cm and 5 × 70–cm chromatography columns 500- and 1000-mL separatory funnels Glass filters (porosity 3) Vacuum desiccator Additional reagents and equipment for column chromatography (APPENDIX 3E) NOTE: All reactions are carried out with anhydrous solvents. All starting materials are commercially available. All evaporations are accomplished in vacuo using a rotary evaporator at 40◦ C. All filtrations are performed using a glass filter under vacuum.
Perform condensation 1. Dissolve 8.0 g uracil (71.4 mmol) and 10 g 1,5:2,3-dianhydro-4,6-O-benzylideneD-allitol (42.7 mmol) in 80 mL anhydrous DMF in a 500-mL flask. Carry out the reaction under N2 atmosphere using a gas balloon. 2. Add 12 mL DBU (79 mmol) in 20 mL anhydrous DMF via a dropping funnel over a period of 30 min. 3. Stir the reaction mixture at 95◦ C for 4 hr. 4. Cool the light-brown solution to room temperature and remove the volatiles in vacuo. 5. Add 300 mL water and suspend the crude product. 6. Cool suspension with an ice-water bath and neutralize with 5 to 6 mL AcOH. 7. Add 200 mL brine and extract three times with 200 mL EtOAc in a 1000-mL separatory funnel. 8. Dry the organic layer over Na2 SO4 , filter off the drying agent under vacuum using a glass filter, and evaporate the solvent from the filtrate using a rotary evaporator equipped with a vacuum pump. 9. Purify the residue by column chromatography (APPENDIX 3E) using 200 g of silica gel with gradient elution (98:2, 96:4 DCM/MeOH). 1 ,5 -Anhydro-4 ,6 -O-benzylidene-2 -deoxy-2 -(uracil-1-yl)-D-altro-hexitol (S.1): 85% yield (12.5 g). TLC (96:4 DCM/MeOH): Rf = 0.2. UV (MeOH): λmax = 266 nm. 1 H NMR (CDCl3 ) δ 3.65 (dd, 1H, J = 9.6 and 2.5 Hz, 4 -H); 3.72 (t, 1H, J= 10.6 Hz, 6 -Ha); 3.85 (br s, 1H, 3 -OH); 3.89 (app d, J = 13.6 Hz, 1 -Ha); 4.10 (dt, 1H, J= 4.7 and 9.9 Hz; 5 -H); 4.26-4.44 (m, 3H, 6 -He, 1 -He, 3 -H); 4.43 (t, 1H, 2 -H); 5.60 (s, 1H, Ph-CH); 5.78 (dd, 1H, J = 8.1 and 2.0 Hz, 5-H); 7.33-7.47 (m, 5H, ar-H), 7.99 (d, 1H, J = 8.1 Hz, 6-H). 13 C NMR (CDCl3 ) δ 56.60 (C-2 ); 64.01 (C-1 ); 65.83 (C-3 ); 66.20 (C-5 ); 68.80 (C-6 ); 76.51 (C-4 ); 102.13 (Ph-CH); 102.86 (C-5); 126.17 (ar-Co ); 128.26 (ar-Cm ); 129.23 (ar-Cp ); 137.00 (ar-Ci ); 142.19 (C-6); 151.36 (C-2); 163.41 (C-4). HRMS (thgly) calcd. for C17 H18 N2 O6 (MH)+ 347.1243, found 347.1291.
Benzoylate 3-hydroxy 10. Dissolve 8.5 g of S.1 (25 mmol) in 100 mL dry pyridine in a 500-mL flask and cool to 0◦ C. 11. Add 5.9 mL BzCl (50 mmol) and allow the mixture to come to room temperature. Continue stirring overnight. 12. Cool the brown solution with an ice bath and add 10 mL water. 13. Remove the volatiles under reduced pressure using a rotary evaporator and coevaporate three times with 50 mL toluene.
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14. Dissolve the resulting solid in 700 mL DCM and wash two times with 200 mL saturated NaHCO3 in a 1000-mL separatory funnel. 15. Dry the organic layer over Na2 SO4 , filter off the drying agent using a glass filter, and evaporate the solvent from the filtrate using a rotary evaporator at 40◦ C. 16. Absorb the crude product on silica gel by suspending the product in 250 mL chloroform in a 500-mL flask with stirring, then adding 50 g silica gel and evaporating the suspension carefully to dryness. Purify by column chromatography with gradient elution (40:60, 10:90 hexane/EtOAc and 100% EtOAc). 1 ,5-Anhydro-3-O-benzoyl-4 ,6 -O-benzylidene-2 -deoxy-2 -(uracil-1-yl)-D-altro-hexitol (S.2): 80% yield (8.7 g). TLC (50:50 hexane/EtOAc): Rf = 0.25. 1 H NMR (DMSO-d6 ) δ 3.72-3.90 (m, 1H, 4 -H); 4.10 (m, 2H, 5 -H, and 6 -Ha); 4.22-4.45 (m, 3H, 6 -He, 1 -Ha, and 1 -He); 4.48 (t, 1H, 2 -H); 5.69 (s, 1H, Ph-CH); 5.71 (d, 1H, J = 8.1 Hz, 5-H); 5.73 (br s, 1H, 3 -H); 7.20-7.35 (m, 5H, Bn); 7.54-7.85 (m, 3H, Bz m+p ); 7.96 (d, 1H, J = 8.1 Hz, 6-H); 8.13 (d, 2H, J = 7.0 Hz, Bzo ); 11.42 (br s, 1H, N3-H). HRMS (thgly) calcd. for C24 H23 N2 O7 (MH)+ 451.1505, found 451.1510.
Cleave benzylidene protecting group 17. Suspend 3.6.g of S.2 (8 mmol) in dry 50 mL DCE in a 100-mL flask and cool to 0◦ C. 18. Add 4 mL TFA (50 mmol) and stir yellow-clear solution for 30 min at 0◦ C. 19. Add 160 µL water (9 mmol) and continue stirring for 20 min at 0◦ C. 20. Neutralize the reaction mixture with 4.8 mL pyridine (60 mmol) and continue stirring the colorless solution at 0◦ C for 2-6 hr. 21. Filter off white crystalline solid using a glass filter, wash twice with 5 mL cold DCM, and dry overnight in a vacuum desiccator. 1 ,5 -Anhydro-3 -O-benzoyl-2 -deoxy-2 -(uracil-1-yl)-D-altro-hexitol (S.3): 82% yield (2.4 g). TLC (95:5 DCM/MeOH): Rf = 0.05. 1 H NMR (DMSO-d6 ) δ 3.60-3.80 (m, 3H, 4 -H, 6 -Ha, and 6 -He); 3.95 (m, 1H, 5 -H); 4.0-4.13 (m, 2H, 1 -Ha+e); 4.66 (dd, 1H, 2 -H); 4.77 (t, 1H, 6 -OH); 5.33 (d, 1H, J = 5.5 Hz, 4 -OH); 5.50 (dd, 1H, J = 6.0 and 2.9Hz, 3 -H); 5.59 (dd, 1H, J = 8.1 and 2.1 Hz, 5-H); 7.50-7.75 (m, 3H, Bzm+p ); 7.98-8.10 (m, 3H, Bzo , 6-H); 11.35 (br s, 1H, N3-H). HRMS (thgly) calcd. for C17 H18 N2 O7 (MH+ ) 363.1192, found 363.1185.
Tritylate 6-hydroxy 22. Co-evaporate 2.4 g of S.3 (6.6 mmol) with 15 mL dry pyridine in a 250-mL flask. 23. Add 5 mL dry pyridine and 50 mL DCM and cool to 0◦ C. 24. Add 2.25 g MMTr-Cl (7.2 mmol), allow the mixture to come to room temperature, and then stir for 3 hr. 25. Add 1 mL MeOH with stirring for 15 min to decompose excess MMTr-Cl. 26. Add 150 mL DCM and wash three times with 100 mL water in a 500-mL separatory funnel. 27. Dry the organic layer over Na2 SO4 , filter off the drying agent using a glass filter, and evaporate the solvent using a rotary evaporator at 40◦ C until 30 to 40 mL remains. 28. Add the solution dropwise over 30 min to 200 mL hexane at 0◦ C with stirring. Continue stirring 1 hr at 0◦ C. Synthesis of Altritol Nucleoside Phosphoramidites
29. Filter off the white fine powder using a glass filter, wash twice with 10 mL cold hexane, and dry overnight in a vacuum desiccator. The dry product can be stored in the dark at –20◦ C for 6 months.
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1 ,5 -Anhydro-3 -O-benzoyl-6 -O-monomethoxytrityl-2 -deoxy-2 -(uracil-1-yl)-D-altrohexitol (S.4): 90% yield (3.8 g). TLC (80:20 EtOAc/hexane): Rf = 0.7. 1 H NMR (CDCl3 ) δ 3.42 (dd, J = 10.4 and 3.1 Hz, 6 -Ha); 3.54 (dd, 1H, J = 10.4 and 2.7 Hz, 6 -He); 3.77 (s, 3H, O-CH3 ); 3.75-3.85 (m, 1H, 4 -H); 4.12 ( dd, 1H, J = 14.0 and 7.2 Hz, 1 -Ha); 4.31 (dd, 1H, J = 14.0 and 3.6 Hz, 1 -He); 4.17-4.38 (m, 1H, 5 -H); 4.73 (app t, 1H, 2 -H); 5.69-5.80 (m, 2H, 3 -H and 5-H); 6.82 (d, 2H, J = 8.9 Hz, ar-H); 7.18-7.60 (m, 15H, ar-H); 7.99 (d, 2H, J = 7.0 Hz, ar-H); 8.29 (d, 1H, J = 8.2 Hz, 6-H); 8.80 (br s, 1H, N3-H). 13 C NMR (CDCl3 ) δ 53.70 (C-2 ); 55.20 (O-CH3 ); 62.12 (C-6 ); 64.50 (C-4 ); 64.69 (C-1 ); 69.92 (C-3 ); 76.37 (C-5 ); 86.65 (CTr -O); 102.89 (C-5); 113.21, 127.07, 127.91, 128.14, 128.32, 128.52, 129.17, 129.85, 130.39, 133.55, 135.08, 144.12, 158.55 (ar-C); 142.43 (C-6); 150.77 (C-2); 162.72 (C-4); 165.63 (Bz-CO). HRMS (thgly:NaOAc) calcd. for C37 H34 N2 O8 Na (M+Na)+ 657.2213 found 657.2219.
SYNTHESIS OF 1 ,5 -ANHYDRO-3 -O-BENZOYL-6 -OMONOMETHOXYTRITYL-2 -DEOXY-2 -(N4 -BENZOYLCYTOSIN-1-YL)-DALTRO-HEXITOL
BASIC PROTOCOL 2
Synthesis of the protected nucleoside starts with the uracil intermediate S.2 and is illustrated in Figure 1.18.2. The cytosine altro-nucleoside (S.5) is prepared from S.2 via ammonium treatment of the 1,2,4-triazolyl intermediate. Treatment of S.5 with excess benzoyl chloride leads to protection of 3 -hydroxyl group and the exocyclic amino group of cytosine. Benzylidene cleavage with TFA followed by monomethoxytritylation provides aC (S.8).
Figure 1.18.2 Preparation of protected aC. Bz, benzoyl; DCE, 1,2-dichloroethane; MMTr, monomethoxytrityl; Py, pyridine; TFA, trifluoroacetic acid.
Synthesis of Modified Nucleosides
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Materials 1,2,4-1H-Triazole, 99.5% Phosphorus oxychloride (POCl3 ) Anhydrous pyridine Dry ice (solid CO2 for cooling bath) Triethylamine 1 ,5 -Anhydro-3 -O-benzoyl-4 ,6 -O-benzylidene-2 -deoxy-2 -(uracil-1-yl)-D-altrohexitol (S.2; see Basic Protocol 1) Toluene PA Dichloromethane PA (DCM) Brine (saturated aqueous sodium chloride solution) Sodium sulfate (Na2 SO4 )Dioxane Concentrated aqueous ammonia (25% NH3 ) Methanol (MeOH) Celite Silica gel (0.060- to 0.200-nm) Benzoyl chloride (BzCl)1,2-Dichloroethane, 99.8+% (DCE) Trifluoroacetic acid (TFA) Monomethoxytrityl Chloride (MMTr-Cl) Hexane PA 100-, 250-, and 500-mL round-bottom flasks Dropping funnel 5 × 35–cm and 5 × 70–cm chromatography columns Gas balloon Rotary evaporator equipped with a vacuum pump Glass filters (porosity 3) Vacuum desiccator Additional reagents and equipment for column chromatography (APPENDIX 3E) NOTE: All reactions are carried out with anhydrous solvents. All starting reagents are commercially available. All evaporations are accomplished in vacuo using a rotary evaporator at 40◦ C. All filtrations are performed using a glass filter under vacuum.
Convert uracil to cytosine 1. Prepare a mixture of 5.2 g 1,2,4-1H-triazole (75 mmol) and 1.5 mL phosphorus oxychloride (15.8 mmol) in 40 mL dry pyridine at 0◦ C in a 500-mL flask. 2. Add 10 mL triethylamine (72 mmol) dropwise and continue stirring at 0◦ C for 30 min. 3. Add a solution of 3.2 g S.2 (8.25 mmol) in 40 mL dry pyridine and stir the reaction mixture at room temperature for 2 hr. The solution turns yellow.
4. Remove the volatiles under vacuum using a rotary evaporator at 40◦ C and coevaporate three times, each time with 25 mL toluene. 5. Dissolve the residue in 250 mL DCM and wash two times with 100 mL brine using a 500-mL separatory funnel. 6. Wash the combined aqueous layer with 50 mL DCM. Synthesis of Altritol Nucleoside Phosphoramidites
7. Combine organic layers, dry over Na2 SO4 , filter off the drying agent under vacuum using a glass filter, and evaporate to dryness using a rotary evaporator equipped with a vacuum pump.
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8. Dissolve the brown residue in 180 mL dioxane in a 500-mL flask and cool to 0◦ C. 9. Add 60 mL concentrated aqueous ammonia and leave the solution overnight at room temperature. 10. Evaporate to dryness using a rotary evaporator under vacuum and co-evaporate three times, each time with 50 mL toluene. 11. Dissolve in 100 mL of 50:50 DCM/MeOH and filter through a small Celite pad. 12. Absorb the crude product on silica gel and purify by column chromatography (APPENDIX 3E), eluting with 98:2 to 95:5 DCM/MeOH. Unreacted starting material S.2 (0.1 to 0.3 g) can be recovered. 1 ,5 -Anhydro-4 ,6 -O-benzylidene-2 -deoxy-2 -(cytosin-1-yl)-D-altro-hexitol (S.5): 70% yield by NMR (3.0 g) as a yellow-orange foam (contains benzamide), used without further purification for the next step. TLC (90:10 DCM/MeOH): Rf = 0.25. UV (MeOH): λmax = 276 nm. 1 H NMR (DMSO-d6 ) δ 3.60 (dd, 1H, J = 2.3 and 9.6 Hz, 4 -H); 3.64 (t, 1H, J = 10.2 Hz, 6 -Ha); 3.91 (dd, 1H, J = 4.9 and 9.6 Hz, 5 -H); 4.00 (m, 1H, 3 -H); 4.00-4.26 (m, 3H, 1 -Ha, 1 -He, and 6 -He); 4.29 (m, 1H, 2 -H); 5.65 (s, 1H, Ph-CH); 5.72 (d, 1H, J = 4.2 Hz, 3 -OH); 5.77 (d, 1H, J = 7.5 Hz, 5-H); 7.05 and 7.19 (2 br s, 2H, 4-NH2 ); 7.30-7.45 (m, 5H, ar-H); 7.94 (d, 1H, J = 7.5 Hz, 6-H). 13 C NMR (DMSO-d6 ) δ 57.46 (C-2 ); 64.00 (C-1 ); 64.87 (C-3 ); 65.79 (C-5 ); 68.28 (C-6 ); 76.50 (C-4 ); 94.09 (C-5); 101.20 (Ph-CH); 126.50 (2C, ar-Co ); 128.10 (2C, ar-Cm ); 128.95 (ar-Cp ); 137.93 (ar-Ci ); 143.75 (C-6); 154.98 (C-2); 165.19 (C-4). HRMS (thgly) calcd. for C17 H20 N3 O5 (MH)+ 346.1403, found 346.1380.
Benzoylate 3-hydroxy and nucleobase 13. Dissolve 3.0 g of S.5 in 50 mL dry pyridine in a 250-mL round-bottom flask equipped with a gas balloon. 14. Add 2.5 mL benzoyl chloride (20.0 mmol) and stir the reaction mixture overnight at room temperature. 15. Cool the reaction mixture to 0◦ C, add 1 mL water, and stir for 20 min. 16. Add 10 mL concentrated aqueous ammonia and continue stirring at room temperature for 1 hr. 17. Remove the volatiles under reduced pressure using a rotary evaporator, and coevaporate three times, each time with 15 mL toluene. 18. Absorb the crude product on silica gel and purify by column chromatography (APPENDIX 3E), eluting with 98:2 DCM/MeOH. The title product crystallized in tubes during collection of the fractions. 1 ,5-Anhydro-3 -O-benzoyl-4 ,6 -O-benzylidene-2 -deoxy-2 -(N4 -benzoylcytosin-1-yl)-Daltro-hexitol (S.6): overall yield 61% (2.8 g, 5.0 mmol) from S.2. TLC (95:5 DCM/MeOH): Rf = 0.25. 1 H NMR (DMSO-d6 ) δ 3.85 (m, 1H, 4 -H); 4.00-4.60 (m, 5H, 5 -H, 1 -Ha, 1 -He, 6 -Ha, and 6 -He); 4.63 (br s, 1H, 2 -H); 5.70 (s, 1H, Ph-CH); 5.81 (br s, 1H, 3 -H); 7.20-7.35 (m, 5H, ar-H); 7.45-7.80 (m, 7H, 2 × Bz, 5-H); 8.04 (d, 2H, J = 7 Hz, Bzo ); 8.15 (d, 2H, J = 7 Hz, Bzo ); 8.48 (d, 1H, J = 7.7 Hz, 6-H); 11.35 (br s, 1H, N4-H). 13 C NMR (DMSO-d6 ) δ 55.83 (C-2 ); 64.84 (C-1 ); 66.20 and 67.23 (C-3 , C-5 ); 68.08 (C-6 ); 74.21 (C-4 ); 97.14 (C-5); 100.81 (Ph-CH); 126.03, 128.18, 128.64, 128.97, 129.08, 129.44, 129.76, 132.95, 133.29, 133.88, 137.60 (ar-C); 147.71 (C-6); 155.20 (C-2); 163.24 (C-4); 164.11 (PhCONH and PhCOO). HRMS (thgly) calcd. for C31 H28 N3 O7 (MH)+ 554.1927, found 554.1892.
Cleave benzylidene protecting group 19. Suspend 2.8 g of S.6 (5.0 mmol) in 25 mL dry DCE in a 100-mL flask and cool to 0◦ C.
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20. Add 2.5 mL TFA (30 mmol) and stir yellow-clear solution for 30 min at 0◦ C. 21. Add 110 µl water (6 mmol) and continue stirring for 20 min at 0◦ C. 22. Neutralize the reaction mixture with 3.2 mL pyridine (40 mmol) and continue stirring the colorless solution at 0◦ C for 2 to 4 hr. 23. Filter off white crystalline solid using a glass filter, wash twice with 5 mL cold DCM, and dry overnight on a vacuum desiccator. 1 ,5 -Anhydro-3 -O-benzoyl-2 -deoxy-2 -(N4 -benzoylcytosin-1-yl)-D-altro-hexitol (S.7): 80% yield (1.8 g). TLC (95:5 DCM/MeOH): Rf = 0.25. 1 H NMR (DMSO-d6 ) δ 3.60-4.00 (m, 3H, 4 -H, 6 -Ha, and 6 -He); 4.10-4.40 (m, 3H, 5 -H, 1 -Ha, and 1 -He); 4.65-4.85 (m, 2H, 2 -H and 6 -OH); 5.25 (br s, 1H, 4 -OH); 5.65 (m, 1H, 3 -H); 7.35 (d, 1H, J = 7.8 Hz, 5-H); 7.45-7.80 (m, 6H, ar-H); 7.90-8.15 (m, 4H, ar-H); 8.48 (d, 1H, J = 7.8 Hz, 6-H); 11.35 (br s, 1H, N4-H). LSIMS (thgly, negative mode) 466.2 (MH)+ . HRMS (thgly) calcd. for C24 H23 N3 O7 (MH+ ) 466.1614, found 466.1603.
Tritylate 6-hydroxy 24. Co-evaporate 1.8 g S.7 (4.0 mmol) with 10 mL dry pyridine in a 250-mL flask. 25. Add 3 mL dry pyridine and 20 mL DCM and cool to 0◦ C. 26. Add 1.4 g MMTr-Cl (4.4 mmol) in two portions over 30 min. Allow the mixture to come to room temperature and stir for 3 hr. 27. Add 1 mL MeOH to decompose excess MMTr-Cl and stir for 15 min. 28. Add 150 mL DCM and extract three times, each time with 100 mL water. 29. Dry the organic layer over Na2 SO4 , filter off the drying agent using a glass filter, and evaporate the solvent until 30 to 40 mL of solution remains. 30. Add solution dropwise over 30 min to 150 mL hexane at 0◦ C with stirring. Continue stirring for 1 hr at 0◦ C. 31. Filter off the white fine powder using a glass filter, wash two times with 10 mL cold hexane, and dry overnight in a vacuum desiccator. The dry product can be stored in the dark at –20◦ C for 6 months. 1’,5’-Anhydro-3’-O-benzoyl-6’-O-monomethoxytrityl-2’-deoxy-2’-(N4-benzoylcytosin-1-yl)D-altro-hexitol (S.8): 75% yield (2.2 g). TLC (80:20 DCM/acetone): Rf = 0.58. 1 H NMR (DMSO-d6) δ 3.43 (dd, 1H, J = 10.1 and 3.1 Hz, 6’-Ha); 3.53 (dd, 1H, 6’-He); 3.65 (m, 1H, 4’-H); 3.79 (s, 3H, OCH3 ); 3.75-3.90 (m, 1H, 5’-H); 4.05-4.42 (m, 3H, 1’-Ha, 1’-He, 4’-OH); 4.93 (br s, 1H, 2’-H); 5.84 (app t, 1H, J= 3.1 Hz, 3’-H); 6.86 (d, 2H, J = 8.8 Hz, ar-H); 7.15-7.70 (m, 19H, ar-H, 5-H); 7.90-8.05 (m, 4H, ar-H); 8.78 (d, 1H, J = 7.7 Hz, 6-H). 13 C NMR (CDCl3 ) δ 55.07 and 55.20 (C-2’, OCH3 ); 62.08 (C-6’); 64.24 (C-4’); 64.82 (C-1’); 69.66 (C-3’); 75.96 (C-5’); 86.59 (CTr -O); 97.28 (C-5); 113.21; 127.07, 127.65, 127.91, 128.46, 129.02, 129.32, 129.86, 130.16, 133.02, 133.17, 133.47, 135.57, 143.99, 144.13 and 158.62 (ar-C); 147.56 (C-6); 155.0 (C-2); 162.20 (C-4); 165.83 (CO-O and CO-N). HRMS (thgly) calcd. for C44 H39 N3 O8 Na (M+Na)+ 760.2635 found 760.2642. BASIC PROTOCOL 3
Synthesis of Altritol Nucleoside Phosphoramidites
SYNTHESIS OF 1 ,5 -ANHYDRO-3 -O-BENZOYL-6 -OMONOMETHOXYTRITYL-2 -DEOXY-2 -(N6 -BENZOYLADENIN-9-YL)-DALTRO-HEXITOL Synthesis of the protected nucleoside from adenine is illustrated in Figure 1.18.3. The key product is prepared in four steps. Reaction of the DBU salt of adenine with 1,5:2, 3-dianhydro-4,6-O-benzylidene-D-allitol yields the altro-derivative S.9. Treatment with excess benzoyl chloride leads to protection of the 3 -hydroxyl group. Benzylidene cleavage with TFA followed by monomethoxytritylation provides aA (S.12).
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Figure 1.18.3 Preparation of protected aA. Bz, benzoyl; DBU, 1,8-diazabicyclo[5.4.0]undec-7ene; DCE, 1,2-dichloroethane; DMF, N,N-dimethylformamide; MMTr, monomethoxytrityl; Py, pyridine; TFA, trifluoroacetic acid.
Materials Adenine, 99+% 1,5:2,3-Dianhydro-4,6-O-benzylidene-D-allitol (CMS Chemical, http://www.cms-chemicals.com, or Brockway et al., 1984) N,N-Dimethylformamide (DMF), dry Nitrogen gas 1,8-Diazabicyclo[5.4.0]undec-7-ene (DBU)Dry ice (solid CO2 for cooling bath) Acetic acid (AcOH) Anhydrous pyridine Benzoyl chloride (BzCl) Concentrated aqueous ammonia (25% NH3 ) Toluene PA Dichloromethane PA (DCM) Silica gel (0.060- to 0.200-nm) Hexane PA Ethyl acetate (EtOAc) 1,2-Dichloroethane 99.8+% (DCE), dry Trifluoroacetic acid (TFA) Monomethoxytrityl chloride (MMTr-Cl) Methanol (MeOH) Sodium sulfate (Na2 SO4 ) 100-, 250-, and 500-mL round-bottom flasks Gas balloon Dropping funnel Ice bath 500-mL and 1-L separatory funnels 5 × 35–cm and 5 × 70–cm chromatography columns Glass filters (porosity 3)
Synthesis of Modified Nucleosides
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Oven at 70◦ to 80◦ C Rotary evaporator equipped with a vacuum pump Vacuum desiccator Additional reagents and equipment for column chromatography (APPENDIX 3E) NOTE: All reactions are carried out with anhydrous solvents. All starting reagents are commercially available. All evaporations are accomplished in vacuo using a rotary evaporator at 40◦ C. All filtrations are performed using a glass filter under vacuum.
Perform condensation 1. Dissolve 9.0 g adenine (67 mmol) and 10.0 g 1,5:2,3-dianhydro-4,6-O-benzylideneD-allitol (mmol) in 80 mL anhydrous DMF in a 500-mL flask. Carry out the reaction under N2 atmosphere using a gas balloon. 2. Add 12 mL DBU (79 mmol) in anhydrous 20 mL DMF via a dropping funnel over a period of 30 min. Continue stirring the reaction mixture for 4 hr at 95◦ C. 3. Cool the light-brown solution to room temperature and remove the volatiles in vacuo. 4. Add 300 mL water and suspend the crude product. 5. Cool the suspension with an ice-water bath and neutralize with 5 to 6 ml AcOH. 6. Filter off white crystalline solid using a glass filter and wash three times, each time with 50 mL cold water. 7. Dry the white solid in an oven overnight at 70◦ to 80◦ C. 1,5-Anhydro-4,6-O-benzylidene-2-deoxy-2-(adenin-9-yl)-D-altro-hexitol (S.9): 85% yield (13.4 g). TLC (90:10 DCM/MeOH): Rf = 0.35. UV (MeOH) : λmax = 262 nm. 1 H NMR (DMSO-d6 ) δ 3.60 (dd, 1H, 4 -H); 3.80 (t, 1H, 6 -Ha); 4.0 (m, 1H, 5 -H); 4.20-4.40 (m, 4H, 1 -Ha, 1 -He, 3 -H, and 6 -He); 4.60 (br s, 1H, 2 -H); 5.62 (s, 1H, Ph-CH); 5.99 (d, 1H, J = 4.4 Hz, 3 -OH); 7.25-7.40 (m, 5H, ar-H); 8.18 (s, 1H, 2-H); 8.26 (s, 1H, 8-H); 8.55 (br s, 2H, 6-NH2 ). 13 C NMR (DMSO-d6 ) δ 56.17 (C-2 ); 64.55 (C-1 ); 65.28 (C-3 ); 66.61 (C-5 ); 68.34 (C-6 ); 76.35 (C-4 ); 101.30 (Ph-CH); 118.54 (C-5); 126.65 (2C, ar-Co ); 128.32 (2C, ar-Cm ); 129.17 (ar-Cp ); 138.03 (ar-Ci ); 139.49 (C-8); 149.96 (C4); 152.99 (C-2); 156.42 (C-6). HRMS (thgly) calcd. for C18 H20 N5 O4 (MH)+ 370.1515, found 370.1559.
Benzoylate 3-hydroxy and nucleobase 8. Dissolve 13.4 g of S.9 (36 mmol) in 100 mL dry pyridine in a 500-mL flask and cool to 0◦ C. 9. Add 24 mL BzCl (200 mmol) and allow the mixture to come to room temperature. Continue stirring overnight. 10. Cool the brown solution with an ice bath and add 10 mL water. 11. After 5 min add 35 mL aqueous 25% NH3 and continue stirring 1 hr at 0◦ C. 12. Remove the volatiles under reduced pressure and co-evaporate three times, each time with 50 mL toluene. 13. Suspend the resulting solid in 250 mL DCM in a 500-mL flask and filter off precipitate using a glass filter.
Synthesis of Altritol Nucleoside Phosphoramidites
14. Co-evaporate mother liquid with 50 g silica gel and purify the crude product by column chromatography (APPENDIX 3E) with gradient elution (40:60, 10:90 hexane/EtOAc, 100% EtOAc).
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1 ,5 -Anhydro-3 -O-benzoyl-4 ,6 -O-benzylidene-2 -deoxy-2 -(N6 -benzoyladenin-9-yl)-Daltro-hexitol (S.10): 75% yield (15.6 g). TLC (95:5 DCM/MeOH): Rf = 0.5. 1 H NMR (DMSO-d6 , 500 MHz) δ 3.82 (t, 1H, J = 10.25 Hz, 6 -Ha); 3.91 (dd, 1H, J = 9.7 and 2.6 Hz, 4 -H); 4.28 (dt, 1H, J= 5.1 and 9.7 Hz, 5 -H); 4.46 (m, 2H, 1 -Ha, and 6 -He); 4.57 (dd, 1H, J = 2.6 and 13.3 Hz, 1 -He); 5.08 (t, 1H, J = 2.6 Hz, 2 -H); 5.52 (s, 1H, PhCH); 5.99 (t, 1H, J = 2.6 Hz, 3 -H); 7.22-7.27 and 7.30-7.34 (m, 5H, Bn); 7.48-7.54 (m, 4H, two Bzo ); 7.58-7.64 (m, 2H, two Bzp ); 8.05 (d, 2H, J = 7.3 Hz); 8.16 (d, 2H, J = 8.4 Hz, Bzm ); 8.63 (s, 1H, 2-H); 8.85 (s, 1H, 8-H); 9.19 (br s, 2H, 6-NH2 ). 13 C NMR (DMSO-d6 ) δ 53.42 (C-2 ); 65.74 (C-1 ); 67.02 (C-5 ); 68.41 (C-3 ); 68.96 (C-6 ); 74.79 (C-4 ); 102.13 (Ph-CH); 122.83 (C-5); 125.99, 128.02, 128.20, 128.66, 128.87, 129.12, 129.35, 129.96, 132.88, 133.69 and 136.70 (ar-C); 141.77 (C-8); 149.78 (C-4); 152.51 (C-2); 153.21 (C-6); 164.89 (two Bz-Co ). HRMS (thgly) calcd. for C32 H28 N5O6 (MH)+ 578.2040, found 578.2073.
Cleave benzylidene protecting group 15. Suspend 5.0 g of S.10 (8.7 mmol) in 50 mL dry DCE in a 100-mL flask and cool to 0◦ C. 16. Add 4 mL TFA (50 mmol) and stir yellow clear solution for 30 min at 0◦ C. 17. Add 180 µl water (10 mmol) and continue stirring for 20 min at 0◦ C. 18. Neutralize reaction mixture with 4.8 mL pyridine (60 mmol) and continue stirring at 0◦ C for 2 to 6 hr until the precipitation is complete. 19. Filter off white crystalline solid using a glass filter, wash two times with 5 mL cold DCM, and dry overnight in a vacuum desiccator. 1 ,5 -Anhydro-3 -O-benzoyl-2 -deoxy-2 -(N6 -benzoyladenin-9-yl)-D-altro-hexitol (S.11): 90% yield (3.8 g). TLC (95:5 DCM/MeOH): Rf = 0.05. 1 H NMR (DMSO-d6 ) δ 3.704.00 (m, 4H, 4 -H, 5 -H, 6 -Ha, and 6 -He); 4.28 (dd, 1H, J = 2.9 and 12.9 Hz, 1 -Ha); 4.39 (dd, 1H, J = 2.2 and 12.9 Hz, 1 -He); 4.87 (br s, 1H, 6 -OH); 5.06 (br d, 1H, 2 -H); 5.34 (br d, 1H, 4 -OH); 5.74 (dd, 1H, J = 2.5 Hz and 4.5 Hz, 3 -H); 7.50-7.75 (m, 6H, two Bzm+p ); 8.05 (d, 4H, two Bzo ); 8.76 (s, 1H, 2-H); 8.80 (s, 1H, 8-H); 11.21 (br s, 2H, 6-NH2 ). 13 C NMR (DMSO-d6 ) δ 52.62 (C-2 ); 59.85 (C-6 ); 62.61 (C-4 ); 63.82 (C-1 ); 70.38 (C-3 ); 78.34 (C-5 ); 125.43 (C-5); 128.59, 128.85, 128.20, 129.65, 132.57, 133.47 and 133.68 (12C, ar-C); 143.56 (C-8); 150.42 (C-4); 151.85 (C-2); 152.66 (C-6); 164.78, 165.71 (two Bz-Co). HRMS (thgly) calcd. for C25 H24 N5 O6 (MH)+ 490.1727, found 490.1740.
Tritylate 6-hydroxy 20. Co-evaporate 4.0 g of S.11 (8.2 mmol) with 15 mL dry pyridine in a 250-mL flask. 21. Add 5 mL dry pyridine and 50 mL DCM and cool to 0◦ C. 22. Add 2.7 g MMTr-Cl (8.7 mmol) in three portions over 30 min. Allow the mixture to come to room temperature and stir for 3 hr. 23. Add 1 mL MeOH to decompose excess MMTr-Cl and stir for 15 min. 24. Add 150 mL DCM and wash three times, each time with 100 mL water in a 500-mL separatory funnel. 25. Dry the organic layer over Na2 SO4 , filter off the drying agent under vacuum using a glass filter, and evaporate the solvent using a rotary evaporator equipped with a vacuum pump until 30 to 40 mL remains. 26. Add solution dropwise over 30 min to 200 mL hexane at 0◦ C with stirring. Continue stirring suspension for 1 hr at 0◦ C. 27. Filter off white fine powder using a glass filter, wash two times with 10 mL cold hexane, and dry overnight in a vacuum desiccator.
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The dry product can be stored in the dark at –20◦ C for 6 months. 1,5-Anhydro-3-O-benzoyl-6-O-monomethoxytrityl-2 -deoxy-2 -(N6 -benzoyladenin-9-yl)D-altro-hexitol (S.12): 70% yield (4.4 g). TLC (80:20 EtOAc/hexane) : Rf = 0.45. 1 H NMR (DMSO-d6 ) δ 3.20-3.50 (m, 2H, 6 -Ha, and 6 -He); 3.71 (s, 3H, OCH3 ); 4.01 (br s, 2H, 4 -H, and 5 -H); 4.34 (br d, J = 12.5 Hz, 1 -Ha); 4.52 (br d, 1H, J = 12.5 Hz, 1 -He); 5.06 (m, 1H, 2 -H); 5.27 (t, 1H, 4 -OH); 5.85 (d, 1H, J = 4.7 Hz, 3 -H); 6.91 (d, 2H, J = 9.2 Hz, ar-H); 7.17-7.75 (m, 18H, ar-H); 8.07 (d, 4H, J = 7.7 Hz, ar-H); 8.79 (s, 2H, 2-H and 8-H); 11.27 (br s, 1H, 6-NH). 13 C NMR (DMSO-d6 ) δ 52.94 (C-2 ); 55.08 (OCH3 ); 62.34 (C-6 ); 62.87 (C-4 ); 63.88 (C-1 ); 70.03 (C-3 ); 76.42 (C-5 ); 85.65 (CTr -O); 125.24 (C-5); 113.29, 126.90, 127.97, 128.22, 128.58, 128.80, 129.58, 129.70, 130.08, 132.56, 133.48, 133.65, 135.46, 144.36 and 158.20 (ar-C); 143.39 (C-8); 150.57 (C-4); 151.85 (C-2); 152.62 (C-6); 164.77 (CO-O); 165.75 (CO-N). HRMS (thgly) calcd. for C45 H39 N5 O7 Na (M+Na)+ 784.2747, found 784.2743. BASIC PROTOCOL 4
SYNTHESIS OF 1 ,5 -ANHYDRO-3 -O-BENZOYL-6 -OMONOMETHOXYTRITYL-2 -DEOXY-2 -(N2 (DIMETHYLAMINO)METHYLENE-GUANIN-9-YL)-D-ALTRO-HEXITOL Synthesis of the protected nucleoside starts from 2-amino-6-chloropurine and is completed in six steps, as illustrated in Figure 1.18.4. Reaction of the salt of 2-amino6-chloropurine with 1,5:2,3-dianhydro-4,6-O-benzylidene-D-allitol yields the altroderivative S.13. The displacement of chlorine in S.13 by sodium hydroxide, assisted by DABCO, affords the guanine analog S.14. The N2 -protecting group is introduced using N,N-dimethylformamide diethylacetal. Treatment of S.15 with excess benzoyl cyanide leads to protection of the 3 -hydroxyl group. Benzylidene cleavage with TFA followed by monomethoxytritylation provides aG (S.18).
Materials
Synthesis of Altritol Nucleoside Phosphoramidites
2-Amino-6-chloropurine, 99+% 1,5:2,3-Dianhydro-4,6-O-benzylidene-D-allitol (CMS Chemical, http://www.cms-chemicals.com, or Brockway et al., 1984) Hexamethylenephosphorotriamide (HMPA), anhydrous Nitrogen gas Aqueous potassium carbonate (K2 CO3 ), anhydrous 18-Crown-6 Dichloromethane PA (DCM) Sodium sulfate (Na2 SO4 ) Silica gel (0.060- to 0.200-nm) Methanol (MeOH) 1,8-Diazabicyclo[2.2.2]octane (DABCO) 1 N NaOH Dry ice (solid CO2 for cooling bath) 1 N HCl N,N-Dimethylformamide diethyl acetal N,N-Dimethylformamide (DMF), anhydrous p-Xylene Silica gel (0.060 to 0.200 nm) Chloroform (CHCl3 ) Tri-n-butylamine Benzoyl cyanide 1,2-Dichloroethane, 99.8+% (DCE) Trifluoroacetic acid (TFA) Pyridine (Py), anhydrous Monomethoxytrityl chloride (MMTr-Cl) Hexane PA
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Figure 1.18.4 Preparation of protected aG. Bz, benzoyl; DCE, 1,2-dichloroethane; dmf, dimethylaminomethylene; DMF, N,N-dimethylformamide; HMPA, hexamethylenephosphorotriamide; MMTr, monomethoxytrityl; Py, pyridine; TFA, trifluoroacetic acid.
100-, 250-, and 500-mL round-bottom flasks Gas balloon 5 × 35–cm and 5 × 70–cm chromatography columns Glass filters (porosity 3) Vacuum desiccator Rotary evaporator equipped with a vacuum pump 500-mL and 1-L separatory funnels Additional reagents and equipment for column chromatography (APPENDIX 3E) NOTE: All reactions are carried out with anhydrous solvents. All starting reagents are commercially available. All evaporations are accomplished in vacuo using a rotary evaporator at 40◦ C. All filtrations are performed using a glass filter under vacuum.
Perform condensation 1. Suspend 5.4 g 2-amino-6-chloropurine (60 mmol) and 10.0 g 1,5:2,3-dianhydro4,6-O-benzylidene-D-allitol (42.7 mmol) in 150 mL anhydrous HMPA in a 500-mL flask. Carry out the reaction under N2 atmosphere using a gas balloon.
Synthesis of Modified Nucleosides
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2. Add 6 g K2 CO3 (43 mmol) and 5.0 g 18-crown-6 (519 mmol), and stir the reaction mixture at 90◦ C for 3 hr. 3. Cool the light-brown solution to room temperature. 4. Pour into 500 mL ice-water and stir suspension for 1 hr. 5. Filter off white-crystalline solid using a glass filter and wash three times, each time with 50 mL cold water. 6. Dissolve wet paste in 500 mL DCM, separate organic layer using a 1000-mL separatory funnel, and dry over Na2 SO4 . 7. Filter off the drying agent under vacuum using a glass filter, evaporate to dryness, and purify crude material by column chromatography (APPENDIX 3E) on silica gel using 2% MeOH in DCM. 1,5-Anhydro-4 ,6-O-benzilidene-2 -deoxy-(2-amino-6-chloropurin-9-yl)-D-altro-hexitol (S.13): 65% yield (11.3 g). TLC (97.5:2.5 DCM/MeOH): Rf = 0.2. 1 H NMR (DMSO-d6 , 500 MHz) δ 3.69 (1H, dd, J = 2.4 and 9.5 Hz, 4 -H); 3.80 (t, 1H, J = 10.2 Hz, 6 ax-H), 4.00 (td, 1H, J = 5.1 and 10.2 Hz, 5 -H); 4.23-4.30 (m, 4H, 1 eq-H, 1 ax-H, 3 -H, and 6 eq-H); 4.46 (br s, 1H, 2 -H); 5.64 (s, 1H, PhCH); 5.88 (d, 1H, J = 4.5 Hz, 3 -OH), 6.98 (s, 2H, NH2 ); 7.32-7.34 (m, 3H, ar-H); 7.38-7.40 (m, 2H, ar-H); 8.23 (s, 1H, 8-H). 13 C NMR (DMSO-d6 , 125 MHz): δ 55.87 (2 -C); 64.29 (1 -C); 64.77 (3 -C); 66.46 (5 -C); 68.13 (6 -C); 76.27 (4 -C); 101.06 (PhCH); 123.02 (5-C); 126.41-137.75 (4 × C arom); 141.37 (8-C); 149.73 (2-C); 154.26 (4-C); 159.93 (6-C). HRMS: calcd for C18 H19 ClN5 O4 (M+H)+ 404.1125, found 404.1119.
Convert nucleobase to guanine 8. Prepare a suspension of 11.3 g of S.13 (28 mmol) and 1 g DABCO in 100 mL 1 N NaOH using a 500-mL flask. Stir the reaction mixture 2 hr at 90◦ C. 9. Cool the resulting clear colorless solution with an ice bath. 10. Adjust the pH to neutral with 1 N HCl at 0◦ C. 11. Filter off formed solid using a glass filter and wash three times, each time with 50 mL cold water. 12. Dry the fine heavy solid in a desiccator under reduced pressure at room temperature. 1 ,5 -Anhydro-4 ,6 -O-benzilidene-2 -deoxy-(guanin-9-yl)-D-altro-hexitol (S.14): 82% yield (10.0 g). 1 H NMR (DMSO-d6 ) δ 3.67 (dd, 1H, J = 1.8 and 9.5 Hz, 4 -H); 3.79 (t, 1H, J = 9.9 Hz; 6 -Ha); 3.98 (dt, 1H, J = 4.8 and 9.9 Hz, 5 -H); 4.07-4.31 (m, 4H, 1 -Ha, 1 -He, 6 -He, and 3 -H); 4.34 (br s, 1H, 2 -H); 5.66 (s, 1H, Ph-CH); 5.81 (d, 1H, J = 4.3 Hz, 3 -OH); 6.55 (br s, 2H, 2-NH2 ); 7.28-7.49 (m, 5H, ar-H); 7.88 (s, 1H, 8-H). 13 C NMR (DMSO-d6 ) δ 55.50 (C-2 ); 64.68 (C-1 ); 65.36 (C-3 ); 66.45 (C-5 ); 68.25 (C-6 ); 76.37 (C-4 ); 101.17 (Ph-CH); 116.29 (C-5); 126.55 (ar-Co ); 128.11 (ar-Cm ); 128.98 (ar-Cp ); 135.88 (C-8); 137.88 (ar-Ci ); 151.49 (C-4); 153.88 (C-2); 157.04 (C-6). HRMS (thgly) calcd for C18 H19 N5 O5 (MH)+ 386.1464, found 386.1496.
Perform nucleobase protection 13. Prepare a mixture of 10.0 g of S.14 (26 mmol) and 20 mL N,N-dimethylformamide diethyl acetal in 100 mL DMF in a 500-mL flask. Stir the reaction mixture at room temperature overnight. 14. Cool the resulting clear light-yellow solution with an ice bath. 15. Add 20 mL water and stir for 30 min. Synthesis of Altritol Nucleoside Phosphoramidites
16. Evaporate to dryness in vacuo and co-evaporate three times, each time with 30 mL xylene.
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17. Absorb the crude product on 50 g silica gel and purify by column chromatography (APPENDIX 3E) with gradient elution (98:2; 90:10 chloroform/MeOH). 1,5-Anhydro-4 ,6-O-benzylidene-2 -deoxy-2 -(N2 -(dimethylamino)methylene-guanin-9yl)-D-altro-hexitol (S.15): 88% yield (10.0 g). TLC (93:7 DCM/MeOH): Rf = 0.5. 1 H NMR (CDCl3 ) δ 3.04 and 3.13 (two s, 6H), (N(CH3 )2 ); 3.60-4.60 (m, 7H, 1 -Ha, 1 -He, 3 -H, 4 -H, 5 -H, 6 -Ha, and 6 -He); 4.68 (br s, 1H, 2 -H); 5.30 (s, 1H, 3 -OH); 5.52 (s, 1H, Ph-CH); 7.20-7.48 (m, 5H, ar-H); 8.10 (s, 1H, 8-H); 8.61 (s, 1H, N=CH-N); 10.06 (N1-H). 13 C NMR (CDCl3 ) δ 35.09 and 41.34 (N(CH3 )2 );54.72 (C-2 ); 65.10 (C-1 ); 66.38 and 66.80 (C-3 ,C-5 ); 68.90 (C-6 ); 76.36 (C-4 ); 101.98 (Ph-CH); 119.61 (C-5); 126.05 (ar-Co ); 128.14 (ar-Cm ); 129.08 (ar-Cp ); 137.00 (ar-Ci ), 137.25; 150.63 (C-4); 156.97 (C-2,C-6); 158.37 (N=CH-N). LSIMS (thgly) 441 (MH)+ . HRMS (thgly) calcd. for C21 H24 N6 O5 (MH+ ) 441.1886, found 441.1879.
Benzoylate 3-hydroxy 18. Prepare a solution of 10.0 g of S.15 (23 mmol) and 4.6 mL tri-n-butylamine (19 mmol) in 80 mL dry DMF in a 500-mL flask. 19. Add 6.0 g benzoyl cyanide (45 mmol) and stir reaction mixture under nitrogen at room temperature for 3 hr. 20. Add 10 mL MeOH and stir 30 min to destroy the excess reagent. 21. Evaporate the solvent in vacuo and co-evaporate two times, each time with 50 mL p-xylene. 22. Dissolve the residue in 250 mL CHCl3 and adsorb on 25 g silica. 23. Purify by column chromatography (APPENDIX 3E) with gradient elution (100:0; 98:2 chloroform/MeOH). 1,5-Anhydro-3-O-benzoyl-4,6-O-benzylidene-2-deoxy-2-(N2-(dimethylamino)methylene -guanin-9-yl)-D-altro-hexitol (S.16): 80% yield (10.0 g). TLC (97:3 DCM/MeOH): Rf = 0.1. Product can be crystallized from EtOH. 1 H NMR (CDCl3 ) δ 3.16 and 3.32 (two s, 6H, N(CH3 )2 ); 3.80 (t, 1H, J = 10 Hz, 6 -Ha); 3.83 (dd, 1H, J = 10.6 and 2.6 Hz, 4 -H); 4.25 (dt, 1H, J = 4.8 and 9.9 Hz, 5 -H); 4.39-4.53 (m, 3H, J = 5 Hz, 1 -Ha, 1 -He, and 6 -He); 4.65 (br d, 1H, 2 -H); 5.52 (s, 1H, Ph-CH); 6.20 (t, 1H, J = 2.9 Hz, 3 -H); 7.20-7.40 (m, 5H, Bn); 7.42-7.67 (m, 3H, BZm+p ); 8.05 (s, 1H, 8-H); 8.13 (d, 2H, Bzo ); 9.15 (s, 1H, N=CH-N); 9.38 (N1-H). 13 C NMR (CDCl3 ) δ 35.06 and 41.40 (N(CH3 )2 );53.72 (C-2 ); 65.01 (C-1 ); 67.14 (C-5 ); 68.30 (C-3 ); 68.99 (C-6 ); 74.42 (C-4 ); 102.19 (Ph-CH); 119.89 (C-5); 126.08, 128.20, 128.72, 129.11 (ar-C); 129.75, 133.69, 136.85, 136.24 (C-8), 150.66 (C-4); 157.67 (C-2); 158.00 (C-6); 160.25 (N=CH-N); 165.14 (Bz-CO). LSIMS (thgly) 545 (MH)+ . HRMS (thgly) calcd. for C28 H28 N6 O6 (MH+ ) 545.2148, found 545.2145.
Cleave benzylidene protecting group 24. Suspend 5.3 g of S.16 (10 mmol) in 50 mL dry DCE in a 100-mL flask and cool to 0◦ C. 25. Add 5 mL TFA (62.5 mmol) and stir yellow clear solution for 30 min at 0◦ C. 26. Add 270 µl water (15 mmol) and continue stirring for 20 min at 0◦ C. 27. Neutralize reaction mixture with 5.2 mL pyridine (65 mmol) and continue stirring at 0◦ C for 1 to 2 hr until the precipitation is complete. 28. Filter off white heavy crystalline solid using a glass filter, wash two times with 10 mL cold DCE, and dry overnight in a vacuum desiccator. 1,5 -Anhydro-3-O-benzoyl-2-deoxy-2-(N2-(dimethylamino)methyleneguanin-9-yl)-D-altrohexitol (S.17): 90% yield (4.1 g). TLC (95:5 DCM/MeOH): Rf = 0.05. 1 H NMR (DMSO-d6 ) δ 3.09 and 3.27 (two s, 6H, N(CH3 )2 ); 3.57-3.84 (m, 4H, 4 -H, 5 -H, 6 -Ha, and 6 -He); 4.21 (dd, 1H, 1 -Ha); 4.38 (br d, 1H, 1 -He); 4.74 (br d, 1H, 2 -H); 5.65 (br
Synthesis of Modified Nucleosides
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s, 1H, 6 -OH); 5.86 (br s, 1H, 3 -H); 6.20 (br s, 1H, 4 -OH); 7.50-7.78 (m, 3H,Bzm+p ); 8.07 (s, 1H, 8-H); 8.08 (d, 2H, J = 7.0 Hz, Bzo ); 8.43 (br s, 1H, N1-H); 8.93 (s, 1H, N=CH-N). 13 C NMR (DMSO-d6 ) δ 34.83 and 40.85 (N(CH3 )2 );53.29 (C-2 ); 60.18 (C-6 ); 62.18 (C-4 ); 63.46 (C-1 ); 70.04 (C-3 ); 78.21 (C-5 ); 126.13 (C-5); 128.95 (ar-Co ); 129.75 (ar-Cm ); 133.78 (ar-Cp ); (ar-Ci not detected); 144.49 (C-8); (C-4 not detected); 157.82 (C-2); 158.12 (C-6); 159.34 (N=CH-N); 165.13 (Bz-CO). HRMS (thgly) calcd. for C21 H24 N6 O6 (MH+ ) 457.1835, found 457,1827.
Tritylate 6-hydroxy 29. Co-evaporate 4.1 g of S.17 (9.0 mmol) with 15 mL dry pyridine in a 250-mL flask. 30. Add 5 mL dry pyridine and 50 mL DCM and cool to 0◦ C. 31. Add 3.0 g MMTr-Cl (9.7 mmol) in three portions over 1 hr. Allow the mixture to come to room temperature and then stir for 3 hr. 32. Add 1 mL MeOH and stir for 15 min to decompose excess MMTr-Cl. 33. Add 150 mL DCM and wash three times, each time with 100 mL water using a 500-mL separatory funnel. 34. Dry the organic layer over Na2 SO4 , filter off the drying agent under vacuum using a glass filter, and evaporate the solvent using a rotary evaporator equipped with a vacuum pump until 30 to 40 mL remains. 35. Add solution dropwise over 30 min to 200 mL hexane at 0◦ C with stirring. Continue stirring suspension for 1 hr at 0◦ C. 36. Filter off white fine powder using a glass filter, wash two times, each time with 10 mL cold hexane, and dry overnight in a vacuum desiccator. The dry product can be stored in the dark at –20◦ C for 6 months. 1 ,5 -Anhydro-3 -O-benzoyl-6 -O-monomethoxytrityl-2 -deoxy-2 -(N2 -(dimethylamino)methylene-guanin-9-yl)-D-altro-hexitol (S.18): 75% yield (4.9 g). TLC (50:50 CH2 Cl2 /acetone): Rf = 0.33. 1 H NMR (CDCl3 , 200 MHz) δ 3.08 and 3.24 (two s, 3H, N(CH3 )2 ); 3.36-3.65 (m, 2H, 6 -Ha, and 6 -He); 3.77 (s, 3H, OCH3 ); 3.85-4.10 (m, 2H, 4 -H, 5 -H); 4.23-4.48 (m, 2H, 1 -Ha and 1 -He); 4.60 (br d, 1H, J = 2.9 Hz, 2 -H); 6.02 (t, 1H, J = 3.0 Hz, 3 -H); 6.84 (d, 2H, ar-H); 7.10-7.70 (m, 15H, ar-H); 8.06 (d, 2H, J = 5.8 Hz, ar-H); 8.08 (s, 1H, 8-H); 8.97 (s, 1H, N=CH-N); 9.26 (1-NH). 13 C NMR (CDCl3 , 50 MHz) δ 34.97 and 41.34 (N(CH3 )2 );53.18 (C-2 ); 55.15 (OCH3 ); 63.77 (C-6 ); 64.47 (C-4 ); 64.80 (C-1 ); 69.60 (C-3 ); 76.03 (C-5 ); 86.93 (CTr -O); 119.67 (C-5); 113.27, 127.08, 128.02, 128.35, 128.63, 129.75, 130.33, 133.57 and 135.18; 136.85 (C-8); 144.11 (MMTr); 150.69 (C-4); 157.31 (C-2); 158.10 (C-6); 158.73 (MMTr); 159.83 (N=CH-N); 165.44 (PhCO/OBz). HRMS (thgly:NaOAc) calcd. for C41 H40 N6 O7 Na (MNa)+ 751.2856 found 751.2878. BASIC PROTOCOL 5
GENERAL PROCEDURE FOR PHOSPHORAMIDITE SYNTHESIS Preparation of altritol nucleoside phosphoramidites from the protected nucleosides is illustrated in Figure 1.18.5.
Materials
Synthesis of Altritol Nucleoside Phosphoramidites
Altritol nucleoside S.4 (Basic Protocol 1), S.8 (Basic Protocol 2), S.12 (Basic Protocol 3), or S.18 (Basic Protocol 3) Dioxane, dry 2,4,6-Collidine N-Methylimidazole N,N-Diisopropyl(cyanoethyl)phosphonamidic chloride (CEPA) Dichloromethane PA (DCM)
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Figure 1.18.5 Preparation of protected altritol amidites. Bz, benzoyl; CEPA, N,N-diisopropyl(cyanoethyl)phosphonamidic chloride; DCM, dichloromethane; dmf, dimethylaminomethylene; MMTr, monomethoxytrityl; Py, pyridine.
Acetone PA Triethylamine PA (TEA) 5% Aqueous sodium bicarbonate solution (NaHCO3 ) Sodium sulfate (Na2 SO4 ) Silica gel (0.035- to 0.060-mm) Hexane PA Glass syringe Gas balloon Glass filter (porosity 3) Rotary evaporator equipped with a vacuum pump Vacuum desiccator 2 × 35–cm chromatography column 100-mL round bottom flasks 250-mL separatory funnel Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) NOTE: The reactions are carried out with dry dioxane under nitrogen in 1 to 2 mmol scale. All starting reagents are commercially available. All evaporations are accomplished in vacuo using a rotary evaporator at 25◦ C. All filtrations are performed using a glass filter under vacuum. 1. Dissolve dry protected nucleoside S.4, S.8, S.12, or S.18 (1 mmol) in 5 mL dry dioxane in a 100-mL flask. 2. Add 990 µL 2,4,6-collidine (7.5 mmol) followed by N-methylimidazole (0.5 mmol). 3. Add 560 µL N,N-diisopropylamino(cyanoethyl)phosphonamidic chloride (2.5 mmol) dropwise over 5 min via a glass syringe under nitrogen (using a gas balloon). 4. Stir the reaction for 1 to 2 hr and monitor the progress of the reaction by TLC (APPENDIX 3D) using the solvent systems in step 10. 5. Add 100 µl water and stir for 10 min. 6. Dilute the reaction mixture with 50 mL DCM. Wash DCM solution once with 50 mL 5% NaHCO3 and twice with 100 mL water using a 250-mL separatory funnel.
Synthesis of Modified Nucleosides
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7. Dry over Na2 SO4 , filter off the drying agent using a glass filter, and remove solvent in vacuo using a rotary evaporator. 8. Co-evaporate viscous oil twice with 10 mL toluene. 9. Purify the crude phosphoramidite by flash column chromatography (APPENDIX 3E) on 40 g silica gel using the solvent systems indicated for TLC in step 10. 10. Dissolve oil in 5 mL DCM and add dropwise over 5 min to 100 mL hexane at –60◦ C with stirring. Continue stirring suspension for 30 min at –60◦ C. 11. Filter off white fine powder using a glass filter, wash two times with 10 mL cold (0◦ C) hexane, and dry product overnight in a vacuum desiccator at room temperature. The dry product can be stored in the dark at –20◦ C for 6 months. 1 ,5 -Anhydro-3 -O-benzoyl-6 -O-monomethoxytrityl-2 -deoxy-2 -(uracil-1-yl)-D-altrohexitol 3 -(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (S.19): yield 85%. TLC (59:40:1 hexane/acetone/Et3 N): Rf = 0.39. 31 P NMR: 151.16/150.21. LSIMS (thgly:NaOAc) 856 [M + Na]+ . 1 ,5-Anhydro-3-O-benzoyl-6-O-monomethoxytrityl-2 -deoxy-2 -(N4 -benzoylcytosin-1-yl)3 -(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (S.20): yield 75%. TLC (64:35:1 hexane/acetone/Et3 N): Rf = 0.27. 31 P NMR: 151.45/150.07. LSIMS (thgly:NaOAc) 960 [M + Na]+ .
D-altro-hexitol
1 ,5-Anhydro-3-O-benzoyl-6-O-monomethoxytrityl-2-deoxy-2 -(N6 -benzoyladenin-9-yl)3 -(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (S.21): yield 95%. TLC (59:40:1 hexane/acetone/Et3 N): Rf = 0.28. 31 P NMR: 151.64/149.54. LSIMS (thgly:NaOAc) 984 [M + Na]+ .
D-altro-hexitol
1 ,5 -Anhydro-3 -O-benzoyl-6 -O-monomethoxytrityl-2 -deoxy-2 -(N2 -(dimethylamino)methylene-guanin-9-yl)-D-altro-hexitol 3 -(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (S.22): yield 85%. TLC (19:80:1 hexane/acetone/Et3 N): Rf = 0.33. 31 P NMR: 151.49/149.97. LSIMS (thgly:NaOAc) 929 [M + Na]+ .
COMMENTARY Background Information Altritol nucleic acid (ANA) is an example of an oligonucleotide with a six-membered carbohydrate moiety that hybridizes with natural nucleic acids. These nucleic acids are stable against enzymatic and chemical degradation because they do not have a glycosyl bond. ANA can be synthesized by the phosphoramidite method starting from the building blocks described in this protocol.
Synthesis of Altritol Nucleoside Phosphoramidites
Synthesis The problems in ANA oligonucleotide synthesis are defined by the need to protect the additional 3 -hydroxyl group of altritol nucleosides for oligonucleotide synthesis. These problems have been largely overcome by the use of the benzoyl protecting group for the 3 hydroxyl group (Allart et al., 1999a). However, the problem of 3 →4 benzoyl migration during synthesis of the protected building blocks results in difficulties for large-scale preparation of isomerically pure phosphoramidites.
Therefore, the use of the 3 -O-TBDMS protecting group in ANA oligonucleotide synthesis has also been investigated (Abramov et al., 2004). Although RNA can be produced using this process, the deprotection steps are much more difficult for the preparation of ANA sequences than for RNA sequences. Steric hindrance in the ANA amidites, as of the 3 axial TBDMS group, requires longer coupling times, which increases the formation of side products. Base deprotection with ammonia requires a longer reaction time, which may cause internucleotide cleavage. Desilylation with TBAF is very sensitive to water, and produces salts that must be removed prior to analysis. Triethylamine trihydrogen fluoride (TEA-3HF) has been used as an alternative to TBAF, but is likewise unsuccessful for long stretches of ANA. Because of the variable and not always reproducible deprotection results of the 3 -OTBDMS group in ANA oligonucleotides synthesis, an Fmoc protecting strategy for ANA
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synthesis has been studied as well. The synthesis of four phosphoramidite building blocks of altritol nucleosides with Fmoc protection on the nucleobase and sugar moieties has been accomplished successfully (Abramov et al., 2007a). Coupling yields in oligonucleotide synthesis are generally lower than with 3 -Obenzoyl protected building blocks, but the excellent compatibility with Pac-RNA chemistry for synthesis of chimeric oligonucleotides has been shown, although complete deprotection of the synthesized oligonucleotides proves to be cumbersome at this moment. This could be due to steric hindrance and/or to the low solubility of the growing oligonucleotides in the solvents used for synthesis and deprotection. In conclusion, the benzoyl group is a better protecting group than TBDMS or Fmoc (for the 3 -hydroxy function) in the synthesis of ANA. Further research has focused on the synthesis of 3 -O-benzoyl-protected altritol nucleoside phosphoramidite building blocks, and reaction protocols for large-scale synthesis have been optimized to avoid migration reactions. The results are presented in this unit. Properties of altritol oligonucleotides Oligonucleotides composed of a phosphorylated D-altritol backbone, with nucleobases introduced in the 2 position, hybridize strongly and in a sequence-selective manner with RNA in an antiparallel way. The order of hybridization strength is: dsANA> ANA:RNA>ANA:DNA. Complexes between ANA and RNA or DNA are more stable than those between natural oligonucleotides or hexitol nucleic acids (HNA; UNIT 1.9). The dsANA hybrid is extremely stable. When compared
with an identical dsDNA sequence, the Tm per modification is more than +10◦ C for a hexamer (Table 1.18.1). The Tm /base decreases as a function of oligonucleotide length. CD spectral analysis indicates that ANA complexes are very similar to the A-form dsRNA duplex. ANA are stable in alkaline medium up to pH 12, and are not degraded in human serum (Allart et al., 1999b). ANA oligomers are also superior to the corresponding DNA, RNA, and HNA for nonenzymatic template-directed synthesis of complimentary RNAs (Kozlov et al., 2000). They have also been shown to be useful for developing siRNA with enhanced pharmacological effectiveness (Fisher et al., 2007). Novel analytical platforms based on ANA units have been evaluated for their potential to be used as chimeric oligonucleotide microarrays on glass solid supports for single nucleotide polymorphism detection in DNA and RNA targets. It has been shown that the intensity and selectivity of hybridization signals for RNA targets is higher than for DNA and increases when applying to ANA arrays (ANA > HNA > DNA). Certainly in the new field of RNA detection based on miRNA levels, ANA arrays could be very beneficial (Abramov et al., 2007b).
Compound Characterization TLC was performed on precoated Alugram Sil G/UV254 plates, and compounds were detected using sulfuric acid/anisaldehyde spray and a 254-nm UV lamp. The 1 H and 13 C NMR spectra for nucleosides were determined with a Varian Gemini 200 MHz spectrometer with tetramethylsilane as the internal standard. Abbreviations are: s = singlet, d = doublet,
Table 1.18.1 Melting Temperatures of Fully Modified Hexamers and Dodecamers
Sequencea 6 -h(AGG AGA)
Tm DNA complimentb,c
Tm RNA complimentb,c
31.2
44.8
39.2
59.7
10.0
—
64.8
84.0
73.5
88.1
49.0
47.6
6 -a(AGG AGA) 5 -AGG AGA 6 -h(AGG GAG AGG AGA) 6 -a(AGG GAG AGG AGA) 5 -AGG GAG AGG AGA
a 1,5-Anhydrohexitol oligomers (h); altrohexitol oligomers (a). ◦ m values given in C at 260 nm, and determined in 0.1 M NaCl, 20 mM KH2 PO4 , pH 7.5, 0.1 mM EDTA with 4 µM each oligonucleotide. c Duplexes formed with complimentary sequences 5 -TCT CCT, 5 -TCT CCT CTC CCT as the respective DNA complements and 5 -TT-r(UCU CCU)-TT and 5 -TT-r(UCU CCU CUC CCU) as the respective RNA compliments.
bT
Synthesis of Modified Nucleosides
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dd = double doublet, t = triplet, br s = broad signal, m = multiplet, ddd = double doublet of doublet, dm = double multiplet, app= apparent, a = axial, e = equatorial, ar = aryl. The 31 P NMR spectra for phosphoramidites were determined with a Varian Unity-500 spectrometer (500 MHz for 1 H). 31 P NMR chemical shifts quoted are downfield from 85% H3 PO4 (external). All liquid secondary-ion mass spectra (LSIMS) were obtained using a KRATOS Concept 1H mass spectrometer. Thgly = thioglycerol.
Critical Parameters Overall, it is important for each step of the syntheses that the starting sugar and base building blocks or starting nucleosides are dried thoroughly, either by co-evaporation with anhydrous pyridine or in a desiccator. Anhydrous solvents are very important. They should be either freshly distilled or stored under nitrogen or argon, or be taken from a freshly opened bottle of commercially prepared anhydrous solvent. For all reactions using anhydrous solvents, the glassware should be predried at 70◦ C for at least 2 hr. For evaporation of solvents, it is helpful if the rotary evaporator is equipped with a dryice trap.
Troubleshooting
Synthesis of Altritol Nucleoside Phosphoramidites
A major drawback of the 3 -O-benzoyl group is its migration from the 3 -axial to the 4 -equatorial position during deprotection of the benzylidene group, mandating a change of conditions for benzylidene cleavage and subsequent monomethoxytritylation. Whereas treatment of the benzylidene-protected compounds 2 and 6 with 80% aqueous acetic acid at 80◦ C, or 90% aqueous trifluoroacetic acid at ambient temperature, provides a mixture of 3 - and 4 -O-benzoyl regioisomers, a short treatment of 2 and 6 with anhydrous trifluoroacetic acid at low temperature, followed by neutralization of the acid and precipitation of the product, afforded the expected 3 -Obenzoylated derivatives 3 and 7. Benzoyl migration from the 3 -position to the 4 -position occurred rapidly upon chromatography on silica gel applying protic as well as nonprotic solvents. Using precise amounts of reagent and careful monitoring of reactions allows one to exclude the chromatography purification step and minimize benzoyl migration. Benzoylation of S.15. Treatment of S.15 with BzCl yields S.16 (40%) together with an N2 -benzoyl derivative. This transamidation
can be avoided when the reaction is carried out with benzoyl cyanide in the presence of tributylamine, as described in this unit. Deprotection with TFA. In large-scale experiments, starting nucleoside derivatives are contaminated with 5% to 10% wt of benzamide, which forms on the previous step and is difficult to remove by column chromatography. The presence of this by-product does not influence the yield of the reaction. The deprotected products could contain up to 10% wt of pyridinium trifluoroacetate. Monomethoxytritylation. In large-scale experiments, the starting nucleoside derivatives contained pyridinium trifluoroacetate, which forms in the previous step. The presence of this impurity does not influence the yield of the reaction. During purification, most of the monomethoxytrityl alcohol and its methyl ether could be removed by careful repeated precipitations, but reaction products might contain up to 5 % wt of these contaminants, which are more easily removed on the next step.
Anticipated Results The synthesized altritol nucleoside phosphoramidite building blocks have been successfully used in synthesis of fully modified ANA oligonucleotides as well as for incorporation of ANA units into DNA and RNA sequences (Allart et al., 1999b; Abramov et al., 2007a). The synthesis of oligonucleotides was accomplished by the standard phosphoramidite method on an Exedite synthesizer (Applied Biosystem) with an extended 12-min coupling step. A 15-fold excess of nucleobase acetonitrile solution (200 µl of 0.07 M) and an excess of activator (200 µl) relative to CPGbound 5 -hydroxyl group was used in each coupling cycle, and the synthesis scale was 1.0 µmol. Average coupling yields were monitored by an online colorimeter by colorimetric quantitation of the trityl fractions. Detritylation solution was 3% TCA in DCM, capping was performed with Ac2 O-THF and 10% 1methylimidazole-THF-PyH. Oxidation solution was 0.02 M I2 in THF-H2 O-PyH. Tetrazole solution was 0.45 M in acetonitrile. SEthylthioterazole solution (0.25 M in MeCN) or pyridinium trifluoroacetate solution (0.22 M in MeCN) was used as an activator.
Time Considerations Each step (including purification and spectroscopic analysis) can usually be performed in 1.5 to 2 working days. All synthesized and carefully purified and dried nucleosides
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including phosphoramidites can be stored up to 6 months in the dark at –20◦ C or below under argon or nitrogen. In the planning of the syntheses, it has to be considered that some of reactions must be stirred overnight.
Allart, B., Khan, K., Rosemeyer, H., Schepers, G., Hendrix, C., Rothenbacher, K., Seela, F., Van Aerschot, A., and Herdewijn, P. 1999b. D-Altritol nucleic acids (ANA): Hybridization properties, stability, and initial structural analysis. Chem. Eur. J. 5:2424-2431.
Literature Cited
Brockway, C., Kocienski, P., and Pant, C. 1984. Unusual stereochemistry in the copper-catalysed ring opening of a carbohydrate oxirane with vinylmagnesium bromide. J. Chem. Soc. Perkin Trans. I 875-878.
Abramov, M., Marchand, A., and Herdewijn, P. 2004. Synthesis of D-altritol nucleosides with a 3 -O-tert-butyldimethylsilyl protecting group. Nucleosides, Nucleotides and Nucleic Acids 23:439-455. Abramov, M., Schepers, G., Van Aerschot, A., and Herdewijn, P. 2007a. Fmoc-protected altritol phosphoramidite building blocks and their application in the synthesis of altritol nucleic acids (ANAs). Eur. J. Org. Chem. 14461456. Abramov, M., Schepers, G., Van Aerschot, A., and Herdewijn, P. 2007b. HNA and ANA highaffinity arrays for detections of DNA and RNA single-base mismatches. Biosensors and Bioelectronics, in press. Allart, B., Busson, R., Rozenski, J., Van Aerschot, A., and Herdewijn, P. 1999a. Synthesis of protected D-altritol nucleosides as building blocks for oligonucleotide synthesis. Tetrahedron 55:6527-6546.
De Bouvere, B., Kerremans, L., Rozenski, J., Janssen, G., Van Aerschot, A., Claes, P., Busson, R., and Herdewijn, P. 1997. Improved synthesis of anhydrohexitol building blocks for oligonucleotide synthesis. Liebigs Ann. 1453-1461. Fisher, M., Abramov, M., Van Aerschot, A., Xu, D., Juliano, R.L., and Herdewijn, P. 2007. Inhibition of MDR1 expression with altritol-modified siRNAs. Nucl. Acids Res. 35:1064-1074. Kozlov, I.A., Zielinski, M., Allart, B., Kerremans, L., Van Aerschot, A., Busson, R., Herdewijn, P., and Orgel, L.E. 2000. Nonenzymatic templatedirected reactions on altritol oligomers, preorganized analogues of oligonucleotides. Chem. Eur. J. 6:151-155.
Synthesis of Modified Nucleosides
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CHAPTER 2 Protection of Nucleosides for Oligonucleotide Synthesis INTRODUCTION ince the discovery of the helical structure of DNA by Watson and Crick in 1953, tremendous strides in the chemical synthesis of oligonucleotides have been made in an attempt to produce substantial quantities of synthetic DNA or RNA of defined sequences. The availability of synthetic oligonucleotides has facilitated investigations on gene structure and function, and has intensified research efforts in deciphering the nature of nucleic acid interactions in complex cellular functions.
S
The essence of DNA and RNA synthesis is the correct formation of internucleotide phosphodiester linkages. This is complicated by the presence of reactive functional groups within nucleotide monomers that require protection prior to incorporation into DNA or RNA oligonucleotides. The presence of two and three hydroxyls in deoxyribonucleosides and ribonucleosides, respectively, demands proper protection of these groups to ensure formation of internucleotide linkages with precise directional polarity during chain assembly. In addition to hydroxy groups, the functional groups of the nucleobases require protection to prevent formation of side products during internucleotide coupling reactions. One objective of this chapter is to provide literature overviews of the functional groups that have been used for nucleobase protection along with those employed as protecting groups for the 5 - and 2 -hydroxyls of nucleosides. For example, UNIT 2.1 outlines protective groups for the imide/lactam function of thymine/uracil and guanine, respectively. This class of protecting groups prevents irreversible nucleobase modifications that may occur in the presence of alkylating or condensing reagents; these are commonly used in nucleoside protection strategies and during oligonucleotide synthesis. UNIT 2.1 also identifies protecting groups for the exocyclic amino function of cytosine, adenine, and guanine. In this context, the overview addresses oligonucleotide depurination during synthesis, and examines purine N-protecting groups that have been shown to minimize this problem. Finally, UNIT 2.1 explores recent trends in nucleobase protection that would permit reliable oligonucleotide synthesis and facile removal of N-protecting groups under very mild conditions. relates to 2 -hydroxyl protection of oligoribonucleotides, which, in fact, dictates the selection of both nucleobase and 5 -hydroxyl-protecting groups. The unit surveys in exquisite detail the various types of protecting groups that have been used in the past and those that are currently being used in the synthesis of oligoribonucleotides. The requirements that a protective group must satisfy to become the 2 -hydroxyl-protecting group of choice, in regard to effective oligoribonucleotide synthesis, are delineated in the unit. Furthermore, the issue of 2 -O-acyl and 2 -O-silyl group migration to the 3 hydroxy function of ribonucleosides during protection along with the consequences of the conditions used for their removal on the stability of internucleotide linkages are authoritatively presented. The wealth of information emerging from UNIT 2.2 shall allow one to undertake oligoribonucleotide synthesis with confidence. UNIT 2.2
Current Protocols in Nucleic Acid Chemistry 2.0.1-2.0.3, September 2007 Published online September 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/0471142700.nc0200s30 C 2007 John Wiley & Sons, Inc. Copyright
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2.0.1 Supplement 30
Methods for protecting the 5 -hydroxy function of nucleosides are equally well reviewed in UNIT 2.3. Acid-labile protecting groups that can be used as hydrophobic ligands in the purification of synthetic oligonucleotides are particularly useful and compatible with the most popular oligonucleotide synthesis protocols. Alternatively, 5 -hydroxyl protection of nucleosides with base-labile protecting groups is particularly attractive because it eliminates the risk of oligonucleotide depurination encountered in the stepwise deprotection of acid-labile 5 -O-protecting groups during synthesis. Base-labile 5 -O-protecting groups enable the development of orthogonal protection systems for oligoribonucleotides by expanding the choice of acid-labile 2 -O-protecting groups that can be used. The utilization of the 9-fluorenylmethoxycarbonyl (Fmoc) group for 5 O-protection and 4-methoxytetrahydropyran-4-yl for 2 -O-protection of ribonucleosides is an excellent example of orthogonal protection in oligoribonucleotide synthesis. This method is detailed in UNIT 2.4.
Introduction
In addition to acid- and base-labile protecting groups, 5 -O-silyl-blocking groups that are removable by fluoride ions, photolabile 5 -O-protecting groups, and other functional groups that can be removed under near neutral conditions have been carefully examined in UNIT 2.3. The overviews presented in this chapter will provide anyone interested in oligonucleotide synthesis with the basic knowledge to proceed with the preparation of properly protected nucleoside phosphoramidites or H-phosphonate monomers for incorporation into oligonucleotides. In this regard, a number of synthetic protocols delineating the stepby-step preparation of nucleosides functionalized with N-, 5 -O-, and 2 -O-protecting groups will eventually be included in this chapter to illustrate the versatility of protecting group combinations that allow, in a number of ways, the synthesis of oligonucleotides with well-defined physicochemical properties. Syntheses of N-protected ribonucleosides having a 4,4 -dimethoxytrityl (DMTr) group for 5 -O-protection and a photolabile 2nitrobenzyloxymethyl (NBOM) group or a fluoride-sensitive tert-butyldimethylsilyl (TBDMS) group for 2 -O-protection are outlined in UNIT 2.5, and are representative examples of such synthetic protocols. The preparation of deoxyribonucleosides and ribonucleosides functionalized with a DMTr, TBDMS, and allyloxycarbonyl (AOC) group for 5 -O-, 2 -O- and nucleobase N-protection, respectively, is featured in UNIT 2.12 along with the synthesis of their 3 -phosphoramidite derivatives. Given that the N-AOC protecting group can eventually be removed under near neutral conditions by treatment with an organopalladium(0) catalyst in the presence of an appropriate nucleophile, the preparation of oligonucleotides modified with base-labile nucleobases and/or internucleosidic linkages can now be achieved. The 2-(4-nitrophenyl)ethyl (NPE) group for protection of thymine/uracil at O4 and guanine at O6, along with the 2-(4-nitrophenyl)ethoxycarbonyl (NPEOC) group for protection of cytosine, adenine, and guanine at N4, N6, and N2, respectively, also offer an attractive nucleobase protection strategy for deoxyribonucleosides and ribonucleosides. As described in UNIT 2.13, 5 -O-dimethoxytritylation and 3 -O-phosphinylation of NPE/NPEOC-protected deoxyribonucleosides provide phosphoramidite monomers suitable for solid-phase oligonucleotide synthesis. The cleavage of the NPE/NPEOC groups requires only treatment with a strong, non-nucleophilic base. This step can be performed while the oligonucleotide is still attached to the solid support, thereby allowing removal of all deprotection side products and reagents. An elegant method for the highly regioselective synthesis of 2 -O-TBDMS purine ribonucleosides is presented in UNIT 2.8. A creative modification of the NBOM group has led to the development of the fluoride-sensitive [(triisopropylsilyl)oxy]methyl (TOM) group for 2 -O-protection of ribonucleosides. The detailed preparation of N-protected 5 -O-DMTr-2 -O-TOM ribo-nucleosides is provided in UNIT 2.9. Alternatively, UNIT 2.10 addresses the preparation of N-protected 5 -O-benzhydroxy-bis(trimethylsilyloxy)silyl2 -O-bis(acetoxyethyloxy)-methyl ribonucleosides and their phosphoramidite derivatives as innovative precursors to the solid-phase synthesis of oligoribonucleotides.
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The production of oligonucleotides on a large scale for therapeutic applications presents a challenge that may be dealt with advantageously through a solution-phase oligonucleotide synthesis approach rather than a traditional solid-phase approach. For this purpose, the levulinyl group for 5 -/3 -hydroxyl protection of nucleoside precursors in the synthesis of oligonucleotides is attractive given its compatibility with nucleobase-protecting groups, stability during oligonucleotide assembly, and rapid removal at neutral pH. The details of a regioselective “levulinylation” of 2 -deoxyribonucleoside or 2 -O-methylribonucleoside derivatives using commercial lipases are included in UNIT 2.11. The conversion of suitably protected deoxyribonucleosides and ribonucleosides to Hphosphonate derivatives is carefully described in UNIT 2.6. Alternatively, the conversion of protected deoxyribonucleosides to phosphoramidite derivatives functionalized with groups different than the conventional 2-cyanoethyl group for P-protection is delineated in UNIT 2.7. The generality of these methods ensures the reliable preparation of monomeric nucleoside building blocks for oligonucleotide syntheses. It should be noted that with the advent of oligonucleotide microarrays as powerful diagnostic tools, deoxyribonucleoside phosphoramidites functionalized with photosensitive groups for 5 -/3 -hydroxyl protection have been developed to enable the synthesis of oligonucleotides on planar glass surfaces. To this end, the preparation of 3 -O-[2-(2nitrophenyl)propoxycarbonyl] deoxyribonucleoside 5 -phosphoramidites is reported in UNIT 12.3. Serge L. Beaucage
Protection of Nucleosides for Oligonucleotide Synthesis
2.0.3 Current Protocols in Nucleic Acid Chemistry
Supplement 30
Nucleobase Protection of Deoxyribo- and Ribonucleosides SALIENT FEATURES OF OLIGONUCLEOTIDE SYNTHESIS DNA and RNA are biopolymers comprised of nucleotide units. The synthesis of DNA and RNA as segments consisting of several nucleotides each, referred to as oligonucleotides, is quite a challenging endeavor, requiring the rapid and quantitative coupling of nucleosidic units to form internucleotidic linkages. In order to achieve high efficiency in the coupling reactions, it is necessary to direct the reaction to the desired site in the nucleoside. This is achieved by using appropriate protecting group strategies. To date, five methods of oligonucleotide synthesis have been reported, all of which use protected nucleosides: (1) The H-phosphonate approach (Michelson and Todd, 1955; Hall et al., 1957; Froehler et al., 1986, and references therein; Garegg et al., 1986, and references therein; UNIT 3.4); (2) the phosphodiester approach, pioneered by Khorana and co-workers (Gilham and Khorana, 1958; Khorana, 1979); (3) the phosphotriester approach, which heralded a new era in nucleic acid synthesis by enabling more rapid synthesis and purification (Letsinger and Ogilvie, 1969, and references therein; Reese, 1978); (4) the phosphite strategy (Letsinger and Lunsford, 1976, and references therein; Matteucci and Caruthers, 1980); and (5) the phosphoramidite approach, presently the most popular method for oligonucleotide synthesis (Beaucage and Caruthers, 1981; UNIT 3.3). Over the years, improvements in coupling chemistries, along with advances in methods of solid-phase assembly, have revolutionized the art of oligonucleotide synthesis. In the synthesis of oligonucleotides, persistent as well as transient protection of amino, imido, and hydroxy groups of nucleosides are employed. Transient protecting groups are used to temporarily block one of the hydroxy groups. For example, the 4,4′-dimethoxy trityl group (DMTr) is used for transient protection of the 5′ hydroxyl of a nucleoside (UNIT 2.3). The transient protecting group is removed at the beginning of each coupling cycle. This enables the coupling reaction to be directed to the desired hydroxy group during the synthesis cycle. On the other hand, persistent protecting groups are those that remain on the nucleoside throughout chain assembly and are only removed at the end
UNIT 2.1
of synthesis. For example, the exocyclic amino groups of nucleobases, the 2′-hydroxy groups of sugars (UNIT 2.2), and the internucleotidic phosphodiester functions, are protected using persistent protecting groups. The present commentary addresses issues related to the persistent protection of nucleobases. The commentary is not a comprehensive review of published literature but is intended to highlight the salient features of nucleobase protection. Only selected references are cited, and readers should refer to recent reviews (Beaucage and Iyer, 1992; Iyer and Beaucage, 1999) and primary literature for more comprehensive citations.
GENERAL ASPECTS OF NUCLEOBASE PROTECTION The choice of nucleobase protecting groups and deprotection protocols is of paramount importance in oligonucleotide synthesis, and is dictated by many considerations. The protecting groups for nucleosides should be designed on the basis of several criteria, which have been summarized by Reese (1978). (1) It should be possible to introduce the protecting group using a stable reagent that is readily obtainable. (2) The group should be achiral. (3) The nuclear magnetic resonance (NMR) spectrum of the protected nucleoside should be simple to interpret. (4) The protecting group should be readily installed on the nucleoside. (5) The group should enhance the solubility of the nucleoside in organic solvents so that it can be adapted for coupling reactions. (6) The group should be stable to reagents employed in oligonucleotide synthesis. (7) The group should not cause other structural changes in the nucleoside during its installation or removal, or during oligonucleotide assembly. A number of nucleobase protecting groups have been reported. Improved procedures for their installation and removal have been published. Nevertheless, unforeseen problems have been reported when certain protecting groups are employed in oligonucleotide synthesis. For example, N-acylated purine nucleosides, particularly N-acylated deoxyadenosine, are more predisposed to depurination than their unprotected counterparts, making the growing oligonucleotide chain susceptible to cleavage during the synthesis cycle. In fact, each pro-
Contributed by Radhakrishnan P. Iyer Current Protocols in Nucleic Acid Chemistry (2000) 2.1.1-2.1.17 Copyright © 2000 by John Wiley & Sons, Inc.
Protection of Nucleosides for Oligonucleotide Synthesis
2.1.1
Table 2.1.1 pKa Values of Nucleoside Nucleobases
tected purine and pyrimidine nucleobase exhibits different patterns of reactivity compared with the unprotected nucleoside. Certain protecting groups are also sensitive to reagents used in oligonucleotide synthesis, resulting either in their premature removal or in production of base-modified products (reviewed by Beaucage and Iyer, 1992). The sections that follow attempt to highlight the structural basis for the nucleophilicity and reactivity of nucleosides. An overview of the strategies employed for nucleobase protection is also provided.
Nucleobase (site of protonation/ pKa deprotonation)a 2′-Deoxythymidine (N3) Guanosine (N1) Uridine (N3) Cytidine (N3) Adenosine (N1)
aFor numbering of nucleoside positions, see Figure 2.1.1. For additional pKa values, see Clauwaert and Stockx (1986); Dunn and Hall (1975).
tions, the tautomeric equilibrium could shift to where both imino and enol forms could exist (Saenger, 1984). In turn, these factors influence the reactivity of the nucleosides.
NUCLEOSIDE TAUTOMERISM AND pKa VALUES The nucleophilicity of nucleobases (Fig. 2.1.1) is dictated by the pKa of the amino and amido functions and their tautomeric forms. Table 2.1.1 lists the pKa values of nucleobases. The amide-like nitrogens (N3 of uridine and N1 of guanosine) are acidic in character, whereas the ring nitrogens are basic. Therefore, at strongly alkaline pH, the proton at N3 of uridine and thymidine and that at N1 of guanosine are removed. Under acidic conditions (at pH ∼3), the sites of protonation are N1 of adenosine and N3 of cytidine. At more acidic pH, the N7 of guanosine and adenosine and O4 of uridine are protonated. Thus, all the bases remain mostly uncharged in the physiological range of pH 5 to 9 (Saenger, 1984). It is noteworthy that each of the nucleosides A, C, and G becomes protonated at one of the ring nitrogens rather than on the exocyclic amino group. Thus, the electron pair of the amino group can delocalize into the heteroaromatic ring. Indeed the C–NH2 bonds of A, C, and G are ∼1.34 Å long and have 40% to 50% double-bond character (Saenger, 1984). The charge on a nucleobase, its tautomeric structures, and its ability to form and accept hydrogen bonds are also determined by its pKa values as well as by the pH of the medium. At physiological pH, the major nucleobases exist almost exclusively in the amino and keto tautomeric forms. However, under appropriate pH condi-
NH2
7
6
N 5 8
HO
Nucleobase Protection of Deoxyribo- and Ribonucleosides
O
9N
4
N
2
8
HO
O
3
HO
REACTIVITY OF NUCLEOSIDES Nucleosides participate in electrophilic and nucleophilic substitution reactions as well as addition reactions (Shabarova and Bogdanov, 1994). Quite clearly, protecting groups and the protocols for their installation and removal should be designed to avoid various side reactions. Nucleobases undergo substitution reactions with electrophilic reagents. For example, both N- and O-alkylation of the imide and lactam groups occur with alkylating agents. The N7 position of purines is also a potential site for electrophilic attack (Fig. 2.1.5). Because of these competing reactions, simple alkylation of exocyclic amino function is not a viable protection strategy for nucleobases. On the other hand, it is possible to chemoselectively acylate the exocyclic amino group. Thus, acyl-type protecting groups are widely used for the protection of the exocyclic amino groups of nucleosides (Fig. 2.1.7). The imide/lactam NH of thymidine, uridine (pKa, 9.38), and guanosine (pKa, 9.42) is weakly acidic and can deprotonate under basic conditions. The resulting nucleophilic anion can react with a variety of reagents such as activated phosphates, dicyclohexylcarbodi-
O
O 7 1 N 5 6 NH 9N
H3C 5
2 4
N 3
HO
deoxyadenosine
Figure 2.1.1
N1
deoxyguanosine
NH2
9.93 9.42 9.38 4.17 3.52
6
HO
O
4
NH2 4
3
NH
1 2
N
O
5'1 5'2
H H 6 HO 5' O 4' 3'
HO deoxythymidine
5
HO
O
N3
5'1 5'2
1 2
N 2'
1'
H2'1
deoxycytosine
O
3 4 NH 1 2
5
H H 6 HO 5' O 4'
3'
HO
N 2'
O
1'
OH
uridine
Structures and numbering of nucleosides.
2.1.2 Current Protocols in Nucleic Acid Chemistry
Tri-O-acetyluridine (S.1) reacts with MSNT to produce the triazolo derivative S.2 (Fig. 2.1.2; Reese and Ubasawa, 1980). During deprotection with ammonium hydroxide, S.2 gives cytidine (S.3). Interestingly, during deprotection with either the tetramethylguanidinium salt of syn-4-nitrobenzaldoxime or tetrabutylammonium fluoride (TBAF), S.2 reverts to uridine (den Hartog et al., 1982). The triazolo derivative S.2 is also known to react with other nucleophiles. For example, in the presence of methanol and 1,8-diazabicyclo[3.4.0]undecene-7 (DBU), S.2 gives the corresponding 4-OMe derivative (Li et al., 1987). S.1 reacts with phosphorylating reagents at O4 and undergoes 4-triazolation in the presence of the phosphorylating reagent 2- or 4-chlorophenyl phosphorodi-1,2,4-triazolide. The triazolo derivative is converted to the fluorescent pyridinium salt S.4 in the presence of pyridine (Divakar and Reese, 1982; Sung, 1982; Huynh-Dinh et al., 1985). Further reaction with triazole gives S.5 (Fig. 2.1.3). The purine nucleosides also react with electrophilic reagents. For example, guanine nucleosides react with MSNT to form the corresponding triazolo derivatives. Reaction of the N2-protected guanosine nucleoside S.6 with mesitylene sulfonyl chloride gives the crystal-
imide (DCC), mesitylene sulfonyl chloride, 1(mesitylene-2-sulfonyl)-3-nitro-1,2,4-triazole (MSNT), acid chlorides, phosphitylating reagents, and electrophilic reagents that are employed during coupling reactions. These side reactions result in nucleobase-derived N- and O-products. Nucleosides also react with a variety of nucleophilic reagents. For example, cytosine reacts with hydroxylamine at neutral and acidic pH to give the corresponding hydroxylamine derivative. It reacts with hydrazine at neutral pH to give the corresponding hydrazide. Indeed, nucleophilic substitution reaction of the exocyclic amino groups of cytosine proceeds rapidly in the presence of alkali and amines. Under oxidative conditions, certain nucleobases (for example, adenine and cytosine) can form N-oxides. Also, the C8 position of guanosine is vulnerable to hydrolytic attack under either strongly acidic or strongly alkaline conditions. The 5,6 double bond of pyrimidine nucleosides also reacts with halogens and halohydrins to give the corresponding addition products (Shabarova and Bogdanov, 1994). Selected examples of the side reactions that occur during oligonucleotide synthesis are given below.
NO2 N N O
AcO
N
O AcO
NH2
N NH O
N
N NH3
MSNT AcO
O
(PhO)2P(O)OH/C5H5N OAc
AcO
1
N
O
HO
O HO
OAc
N
O
OH 3
2
MSNT, 1-(mesitylene-2-sulfonyl)-3-nitro-1,2,4-triazole
Figure 2.1.2 Conversion of tri-O-acetyluridine to cytidine via a nitrotriazolide intermediate. Reprinted from Iyer and Beaucage (1999) with permission from Elsevier Science Publishing.
N N O NH AcO
O AcO
N
N
N
O
N
N 1,2,4-triazole
MSNT AcO
O
4-ClPhOP(O)Cl2/C5H5N OAc
1
AcO
N
OAc 4
O
AcO
O AcO
N
O
OAc 5
Figure 2.1.3 Formation of the triazolo derivative S.5 via the pyridinium intermediate S.4. MSNT, 1-(mesitylene-2-sulfonyl)-3-nitro-1,2,4-triazole. Reprinted from Iyer and Beaucage (1999) with permission from Elsevier Science Publishing.
Protection of Nucleosides for Oligonucleotide Synthesis
2.1.3 Current Protocols in Nucleic Acid Chemistry
R O S O O
O N AcO
AcO
NH
N
O
N
N
R-SO2-Cl
AcO
NHTr
O
DMAP AcO
OAc
N
N
N
NHTr
OAc 7
6
DMAP, 4-dimethylaminopyridine; Tr, triphenylmethyl
Figure 2.1.4 Formation of an O6-sulfonated derivative of guanosine. R⋅SO2⋅Cl, mesitylene sulfonyl chloride. Reprinted from Iyer and Beaucage (1999) with permission from Elsevier Science Publishing. R, 2,4,6-trimethylphenyl.
line O6-sulfonated derivative S.7 (Fig. 2.1.4; Bridson et al., 1977; Francois et al., 1985). In the presence of pyridine, S.7 is converted to the corresponding C6 pyridinium compound. The side reactions that are predominant in phosphodiester and phosphotriester chemistry have been summarized (Reese and Ubasawa, 1980). These side reactions are of concern when the H-phosphonate method is employed in oligonucleotide synthesis in conjunction with phosphorochloridates as coupling reagents. The imide and lactam functions of nucleosides also react with phosphitylating reagents. For example, the 2′-deoxyguanosine derivative S.20 (Fig. 2.1.5) reacts with methyl phosphoramidites to give the O6-phosphitylated product S.8 (Pon et al., 1985a; Nielsen et al.,
1987). During oxidation of S.8 with iodine, Oto N-phosphoryl migration occurs (S.9 → S.10; Pon et al., 1985b). Once formed, S.10 induces depurination resulting in chain-cleaved products upon treatment with ammonium hydroxide during the final step of deprotection. It is pertinent that base modifications can also occur during oligonucleotide deprotection. For example, thymine residues are significantly modified as N3-methylthymine during deprotection of oligonucleoside methyl phosphotriesters with aqueous ammonium hydroxide (Urdea et al., 1986). However, when deprotection of the methyl phosphate backbone of oligonucleotides is accomplished by treatment with thiophenol or 2-mercaptobenzothiazole, followed by aqueous ammonium hydroxide to
OR′ O N DMTrO
N
O
RO P O
N NH NHi-Bu
N
N
RO P
N
OR′ DMTrO
AcO
N
O
NHi-Bu
N
AcO 8
20
[31P NMR, d 133.95, 133.79 ppm]
I2
OR′ O
RO P
N DMTrO
O
N
AcO
I OR′ RO P O
O
N
NH N
NHi-Bu
DMTrO
O
N N
NHi-Bu
AcO 10
Nucleobase Protection of Deoxyribo- and Ribonucleosides
N
9 DMTr, 4,4'-dimethoxytriphenylmethyl; i-Bu, isobutyryl
Figure 2.1.5 Reaction of an N2-isobutyryl-2′-deoxyguanosine derivative with phosphitylating reagents. R, Me; R′, nucleoside.
2.1.4 Current Protocols in Nucleic Acid Chemistry
remove nucleobase protecting groups, the formation of N3-methylthymine is considerably reduced (McBride et al., 1987; Andrus and Beaucage, 1988). The reactivity of nucleosides and the attendant risk of base modification suggest an apparent need to protect imide and lactam groups of nucleosides in oligonucleotide synthesis. It is expected that the use of protecting groups at
Protecting group
the imide/lactam function of nucleosides can potentially avert the side reactions. This has been borne out by success in the synthesis of a number of polynucleotides (Reese, 1978).
PROTECTION OF IMIDE AND LACTAM FUNCTIONS A number of groups have been proposed for the protection of thymine and uracil and N3 and
Nucleobase (protection site)
Deprotection conditions
References
U (O4)
syn-4-Nitrobenzaldoxime/ tetramethylguanidine
Nylias et al., 1987 MacMillan and Verdine, 1991
G (O6) U (O4)
syn-4-Nitrobenzaldoxime/ tetramethylguanidine
Jones et al., 1981 Zhou and Chattopadhyaya, 1986 Zhou et al., 1986 Reese and Skone, 1984
T (O4) U (O4)
syn-4-Nitrobenzaldoxime/ tetramethylguanidine
Jones et al., 1981 Reese and Skone, 1984 Scalfi-Happ et al., 1987 Rao et al., 1987
G (O6)
syn-4-Nitrobenzaldoxime/ tetramethylguanidine
Reese and Skone, 1984 Rao et al., 1987 Hagen and Chladek, 1989
G (N1, O6) T/U (N3)
NH4OH, heat
Mag and Engels, 1988
T/U (N3)
10% Pd/H2
Krecmerová et al., 1990
G (O6) T (O4)
Pyridine-2-aldoxime/ tetramethylguanidine 0.5 M DBU/pyr
Pfister et al., 1988 Van Aerschot et al., 1988
G (O6)
R=Me3Si: n-Bu4NF/THF R=PhS: 4-NO2PhS: NaIO4/ NH4OH, heat; R=CN; DBU, or NH4OH
Hagen and Chladek, 1989 Gaffney and Jones, 1982 Engels and Mag, 1982
G (O6) T (O4)
Pd(PPh3)4/PPh3, n-BuNH3+HCO2-
2,4,6-trimethylphenyl NO2
2-nitrophenyl R
R R = H or Me Cl
R R = H or Cl
O
O S
(4-NO2)Ph
2-(4-nitrophenylsulfonyl)ethyl
Ph
O
benzyloxymethyl
R
R = NO2 or CN
R
R = Me3Si; PhS; CN; 4-NO2PhS
Hayakawa et al., 1993
allyl
Figure 2.1.6 Protecting groups for the imide/lactam functions of guanine, uracil, and thymine. DBU, 1,8-diazabicyclo[3.4.0]undecene-7; NalO4, sodium periodate; pyr, pyridine; THF, tetrahydrofuran.
Protection of Nucleosides for Oligonucleotide Synthesis
2.1.5 Current Protocols in Nucleic Acid Chemistry
Protecting group
Ph3C S
Nucleobase (protection site)
Deprotection conditions
References
U (N3)
0.1 M I2/THF/collidine/H2O
Takaku et al., 1988
G (O6)
NH4OH
Kamimura et al., 1983b
U (N3)
DBU/morpholine/pyr
Nyilas et al., 1988
U (N3, O4)
NH4OH
Sekine, 1989, Kamimura et al., 1983a Kamaike et al., 1988
U (N3)
Pyr/H2O or 0.2 M n-Bu4NF/ THF
Sekine, 1989 Welch et al., 1985
G (N2, O6); U (N3)
NH4OH/MeOH
Fujii et al., 1987
Pd[PPh3]4/PPh3, n-BuNH2/THF
Hayakawa et al., 1986
triphenylmethyl sulfenyl O Ph2N diphenylcarbamoyl O O O S O R R = H; Me
O
R R = H; Me; OMe; Cl
NO2 S
R
R = H; Me; NO2
O
n-BuS butylthiocarbonyl O
U (N3, O4)
O allyloxycarbonyl MeO
U (N3)
O
NH4OH, heat, or Ph3C+BF4-/MeCN/H2O
Ito et al., 1986
(methoxyethoxy)methyl
Figure 2.1.6
Nucleobase Protection of Deoxyribo- and Ribonucleosides
(Continued)
O4, and of guanine at O6. A representative list of these groups is shown in Figure 2.1.6. For the benefit of the readers, references that include experimental details are cited. The imide- and lactam-protecting groups can be installed in two steps: sulfonylation or triazolation, followed by displacement of the resulting O-sulfonate or triazolide by the nucleophilic protecting group. The protecting groups can also be installed directly—for example, via Mitsunobu alkylation (UNIT 1.2) or via the transient protection approach. The preparation of these protected nucleosides appears to be straightforward (Jones et al., 1981; Gaffney and Jones, 1982; Trichtinger et al., 1983; Nyilas et al., 1987; Kamaike et al., 1988). However, it is still a matter of debate whether imide and lactam protection is necessary in oligonucleotide synthesis. There are a number of factors to be considered. (1) As noted before,
when using the phosphotriester method in oligonucleotide synthesis, O6-modified guanine and O4-modified uracil revert to guanine and uracil residues upon “oximate” treatment that follows chain assembly. (2) When the phosphoramidite method is used, O6-phosphitylated deoxyguanosine reverts to deoxyguanosine on contact with water or acetate ions (Mag and Engels, 1988). Thus, during the synthesis cycle, if capping is performed (using acetic anhydride) after coupling, any O6-modified deoxyguanosine can potentially revert back to deoxyguanosine. (3) Base modifications generated during oligonucleotide chain assembly depend on the reagents employed and the contact time. Solid-phase oligonucleotide synthesis uses automated pulsed delivery of reagents, resulting in shorter reaction times compared with solution-phase synthesis. Consequently, side
2.1.6 Current Protocols in Nucleic Acid Chemistry
reactions during oligonucleotide chain assembly appear to be minimal. Thus, in principle, appropriate adjustment of synthesis protocols may obviate the necessity for imide and lactam protection. Nevertheless, β-cyanoethyl protection is generally recommended for protection of guanosine at O6 or uridine at O4, and anisoyl for uridine at N3.
PROTECTION OF EXOCYCLIC AMINO GROUPS On the basis of a number of criteria for protecting groups, as outlined previously, Nacyl-type protection for the exocyclic amino groups has emerged as a logical choice. Figure 2.1.7 shows selected examples of N-acyl protecting groups and the reagents used to effect their deprotection. The N-acyl groups are introduced into nucleosides by a number of procedures. (1) They
Protecting group
can be installed by peracylation of the nucleoside followed by chemoselective O-deacylation (Schaller et al., 1963). (2) The recent trend has been to use the “transient” protection approach (Ti et al., 1982). In this procedure, the nucleoside is persilylated using a silylating agent, and the acyl function is installed on the amino group using the corresponding acid chloride or acid anhydride. (3) Some acyl functions such as benzoyl, α-phenylcinnamoyl, and naphthaloyl are directly incorporated on the nucleobase using the corresponding anhydride (Watanabe and Fox, 1966; Bhat et al., 1989). (4) The exocyclic amino group of cytidine and deoxycytidine can be directly acylated using activated esters (Igolen and Morin, 1980), acid chlorides (Mishra and Misra, 1986, and references therein), or alkyloxycarbonylbenzotriazoles (Himmelsbach et al., 1984). (5) Recent reports suggest that site-selective incorporation
Nucleobase (protection site)
Deprotection conditions
References
7-Deaza-6-methyl-G (N2)
NH4OH
Seela and Driller, 1989
C (N4)
NH3/EtOH (1:1)
Köster et al., 1981 Chaix et al., 1989
G (N2); C (N4)
NH4OH, heat
Köster et al., 1981 Büchi and Khorana, 1972 Schulhof et al. 1987
G (N2)
NH4OH
Schulhof et al., 1987
G (N2); A (N6); C (N4)
Pd[PPh3]4/HCO2H/Et2NH
Hayakawa et al., 1986 Hayakawa et al., 1990
G (N2); A (N6); C (N4)
NH4OH
Uznanski et al., 1989
G (N2); A (N6); C (N4)
0.5 M NH2NH2·H2O/ pyr/AcOH (4:1)
Ogilvie et al., 1982
G (N2); A (N6); C (N4)
Multiple protocols
Iyer et al., 1997
O H formyl O
acetyl O
isobutyryl O MeO methoxyacetyl O O allyloxycarbonyl O
i-PrO isopropoxyacetyl O
O
levulinyl O
4-pentenoyl
Figure 2.1.7 Protecting groups for the exocyclic amino function of nucleobases. RT, room temperature; pyr, pyridine.
Protection of Nucleosides for Oligonucleotide Synthesis
2.1.7 Current Protocols in Nucleic Acid Chemistry
Protecting group O
Nucleobase (protection site)
Deprotection conditions
References
G (N2); A (N6); C (N4)
DBU or DBU/pyr
Himmelsbach et al., 1984 Pfleiderer et al., 1985 Pfister and Pfleiderer, 1989
G (N2)
5 M NH3/ MeOH
Jones et al., 1981 Rao et al., 1987 Köster et al., 1981 Balgobin et al., 1981
G (N2); A (N6); C (N4)
NH4OH/pyr Et3N/pyr
Scalfi-Happ et al., 1987 Hagen and Chládek, 1989 Heikkilä and Chattopadhyaya, 1983
G (N2); A (N6); C (N4)
NH4OH, heat
Nagaich and Misra, 1989
G (N2); A (N6)
NH4OH
Chaix et al., 1989 Schulhof et al., 1987 Singh and Misra, 1988
G (N2); A (N6); C (N4)
0.2 M NaOH/MeOH or NH4OH or NH3 gas
(4-NO2)Ph
O 4-nitrophenylethyloxycarbonyl R
O
R = H, C(CH3)3
O O
9-fluorenylmethoxycarbonyl O Ph Ph H α-phenylcinnamoyl R
O O
R = H, Cl
R
O O
CH3C
Köster et al., 1981 Sinha et al., 1993 Boal et al., 1996
R = H, Cl
4-(tert-butyl)phenoxyacetyl O Brown et al., 1989 Rao et al., 1987 Tanimura et al., 1988 Schaller et al., 1963 Balgobin et al., 1981
G (N2); A (N6); C (N4)
NH4OH
G (N2); A (N6); C (N4)
TetramethylguaniNyilas et al., 1988 dine/morpholine/pyr; DBU/morpholine/pyr; NH4OH/morpholine/pyr.
G (N2)
NH4OH, heat
Marugg et al., 1984
G (N2)
0.2 M NaOH/MeOH
Koster et al., 1981
G (N2); A (N6); C (N4)
NH4OH, heat
Mishra and Misra, 1986
R R = H, OMe, Cl, NO2, NMe2, CMe3
O
O
O S
O
R R =H; Cl; NO2
O Ph Ph diphenylacetyl O Cl Cl 3,4-dichlorobenzoyl O MeO PhO 3-methoxy-4-phenoxybenzoyl
Nucleobase Protection of Deoxyribo- and Ribonucleosides
Figure 2.1.7
(Continued)
2.1.8 Current Protocols in Nucleic Acid Chemistry
Protecting group
Nucleobase (protection site)
Deprotection conditions
References
C (N4)
0.2 M NaOH/MeOH (1:1)
Köster et al., 1981
G (N2); A (N6); C (N4)
0.05 M K2CO3/MeOH; NH4OH, RT
Kuijpers et al., 1990
G (N2); A (N6); C (N4)
NH4OH, heat
Dikshit et al., 1988
G (N2); A (N6); C (N4)
n-Bu4NF/pyr/H2O
Dreef-Tromp et al., 1990
O
PhN N 4-(phenylazo)benzoyl O
O
R O
R = Ac, Ph
O
O
1,8-naphthaloyl O
O
Ph Si C(CH3)3 Ph
2-(tert-butyldiphenylsilyloxymethyl)benzoyl
Figure 2.1.7
(Continued)
and removal of N-acyl protecting groups can also be achieved by enzymatic methods (Prasad and Wengel, 1996). Overall, most N-acyl protecting groups are stable in neutral or acidic medium and moderately stable at high pH (pH >13). However, many N-acyl protecting groups are readily removed by ammonolysis. This observation forms the basis for the widespread use of 28% NH4OH as a deprotection reagent when N-acylprotected nucleosides are employed in oligonucleotide synthesis. A salient feature of N-acyl protecting groups is that their stability in alkaline pH can be modulated by the steric and electronic characteristics of specific acyl groups. A study comparing the stability of various N-acyl nucleosides in alkaline medium (0.2 N NaOH/MeOH) has been reported (Köster et al., 1981). Importantly, the stability of the acyl function towards alkaline hydrolysis is determined by the nature of the heterocyclic base. For example, the rate of deacylation of N-acyl derivatives of deoxycytidine is faster than that of
deoxyadenosine or deoxyguanosine. The hydrolytic lability is also determined by inductive, resonance, and steric effects. For example, in a series of N-acyl nucleosides, N-benzoyl nucleosides are hydrolyzed sixteen times faster than N-(2,4-dimethyl)benzoyl nucleosides, presumably because of steric effects. Similarly, N-(2,4dimethoxy)benzoyl nucleoside is hydrolyzed eight times faster than N-(4-dimethylamino)benzoyl nucleoside, perhaps due to a combination of inductive and resonance effects. The choice of a particular N-acyl protecting group also depends on the type of coupling chemistry that is employed. For example, when phosphodiester and phosphotriester chemistries are used in oligonucleotide synthesis, it is necessary to select sturdy N-acyl protecting groups that can withstand the harsh reagents and conditions employed during synthesis. This requirement is met by the benzoyl group for adenine and cytosine, and the isobutyryl group for guanine. However, removal of these protecting groups requires prolonged heating
Protection of Nucleosides for Oligonucleotide Synthesis
2.1.9 Current Protocols in Nucleic Acid Chemistry
(12 to 14 hr) with 28% NH4OH at 55°C. In spite of this limitation, these protecting groups have remained popular even with the advent of automated solid-phase oligonucleotide synthesis using phosphoramidite and H-phosphonate chemistries.
mechanism for depurination of deoxyadenosine (Fig. 2.1.8) involves initial protonation of the nucleoside to produce the N1-protonated form S.11, and then equilibration (prototropic shift) to the N7-protonated species S.12 or S.13, followed by cleavage of the glycosidic bond to give S.15 via the oxonium S.14 (Zoltewicz et al., 1970; Zoltewicz and Clark, 1972). It is also conceivable that, at lower pH, depurination can occur via protonation of the purine nucleobase at both N1 and N7. Presence of the 2′-OH has a significant effect on the nucleoside’s susceptibility to depurination. For example, guanosine and adenosine are more resistant to depurination compared with deoxyadenosine and deoxyguanosine. Deoxyadenosine itself depurinates 1200 times faster than adenosine (York, 1981). Interestingly, N-acyl-protected purine nucleosides (particularly deoxyadenosine) are more prone to depurination than unprotected nucleosides. Among N-acyl-protected deoxyadenosines, protection at N6 with αphenylcinnamoyl, naphthaloyl, 3-methoxy-4phenoxybenzoyl, 9-fluorenylmethoxycarbonyl (FMOC), and tert-butylphenoxyacetyl (t-PAC) groups (Fig. 2.1.7) provides greater resistance to depurination than with N6-benzoyl (reviewed in Beaucage and Iyer, 1992). It is believed that in the case of acyl-protected
PROTECTION OF PURINE NUCLEOBASES: THE PROBLEM OF DEPURINATION The development of suitable protecting groups for purine nucleobases has been an area of considerable interest because purine nucleosides rapidly depurinate under acidic conditions. The problem is compounded by the acidlabile DMTr group used for protection of the 5′-OH in solid-phase oligonucleotide synthesis. Prior to each coupling step in the synthesis cycle, the DMTr group is removed by exposure to a strong acid such as 2% dichloroacetic acid in dichloromethane. Consequently, the growing oligonucleotide chain is repeatedly exposed to strongly acidic conditions, potentially resulting in depurination and reduced yield of the desired “full-length” product. The kinetics and mechanisms of nucleoside depurination have been investigated by several research groups (Romero et al., 1978; Oivanen et al., 1987; Suzuki et al., 1994). The presumed
7
N
NH2 6
NH2 N
N1
9
HO
O
N
H
N
HO
3
HO
N
O
NH N
HO 11
H N HO
O HO 15
HO OH
HO
O
HO oxonium ion 14
O
N
NH2 N N
HO 12
H N HO
O
N
O NH N
NH2
HO 13
Nucleobase Protection of Deoxyribo- and Ribonucleosides
Figure 2.1.8 Scheme showing the proposed depurination mechanism for 2′-deoxyguanosine and 2′-deoxyadenosine catalyzed by protic acids. Modified from Iyer and Beaucage (1999) with permission from Elsevier Science Publishing.
2.1.10 Current Protocols in Nucleic Acid Chemistry
Cl
Cl
Cl
OCOPh OCOPh
Cl Ph
O
N
RO
O
O
N
N
O
N
N
RO
RO
O
O
O
N
N
N
N
O
HN N
N N
RO
RO 16
N
RO
O
N
N
N
Ph
N
RO
O
N
OCOPh
N
RO 18
17
C
19
Figure 2.1.9 Examples of N6-protecting groups for 2′-deoxyadenosine derivatives that reduce depurination. Reprinted from Iyer and Beaucage (1999) with permission from Elsevier Science Publishing. R, DMTr.
purine nucleosides under acidic conditions, the initial site of protonation is N7, rendering the protonated species more prone to glycosidic cleavage. Naturally, caution should be exercised in the synthesis of oligonucleotides whose sequence contains deoxyadenosines at the 3′ terminus. Bis-acylation has also been studied as a strategy to reduce depurination. Imide protecting groups S.16 and S.17, as well as the diamide protecting group S.18, have been investigated (Fig. 2.1.9; Kume et al., 1982, 1984). However, these groups were labile to aqueous pyridine, a
Protecting group
solvent used during the oxidation step in solidphase oligonucleotide synthesis. Thus, alternate oxidants have to be employed for the oxidation step. Since N-acyl-protected purine nucleosides are sensitive to deacylation, alternate protecting groups have been investigated. Prominent among these are the amidine protecting groups, which are introduced using an exchange reaction with appropriate amidine acetals. Interestingly, N-amidine-protected nucleosides (Fig. 2.1.10) resist depurination 20-fold better than the corresponding N-benzoyl nucleosides
Nucleobase (protection site)
Deprotection conditions
References
G (N2), A (N6)
NH4OH, heat
Smrt and Sorm, 1967 Vu et al., 1990 Sproat et al., 1991 Caruthers et al., 1985 McBride et al., 1986
G (N2), A (N6)
NH4OH, heat; 0.5 M NH2NH2·H2O/pyr/AcOH
McBride et al., 1986 Froehler and Matteucci, 1983
A (N6)
NH4OH/10% NH4OAc/heat; 0.5 M NH2NH2·H2O/pyr/AcOH
Froehler and Matteucci, 1983
G (N2), A (N6)
NH4OH, heat
McBride et al., 1986
A (N6)
NH4OH/10% NH4OAc/heat; 0.5 M NH2NH2·H2O/pyr/AcOH
Froehler and Matteucci, 1983
C (N4)
NH4OH, heat
McBride et al., 1986
H Me2N (dimethylamino)methylene
H
n-Bu2N (di-n-butylamino)methylene
H
i-Pr2N (diisopropylamino)methylene
Me2N 1-(dimethylamino)ethylidene
H
i-Bu2N (diisobutylamino)methylene
N Me N-methylpyrrolidin-2-ylidene
Figure 2.1.10
Amidine protecting groups for exocyclic amino functions. Pyr, pyridine.
Protection of Nucleosides for Oligonucleotide Synthesis
2.1.11 Current Protocols in Nucleic Acid Chemistry
(Smrt and Sorm, 1967; Holy and Zemlicka, 1969, and references therein; Froehler and Matteucci, 1983; Caruthers et al., 1985; Vu et al., 1990; Sproat et al., 1991). It is presumed that the protonation sites of amidine-protected nucleosides are N1 and N6 instead of N1 and N7, respectively, resulting in a slower rate of depurination. Indeed, amidineprotected nucleoside phosphoramidites are frequently employed in solid-phase oligonucleotide synthesis. However, changes are required in the oxidation step to avoid nucleobase modifications (Mullah et al., 1995). Nucleosides protected with O-nitrophenylsulfonyl and tris(benzoyloxy)trityl (S.19) groups also appear to be more resistant to depurination (Shimidzu and Letsinger, 1968; Honda et al., 1984, and references therein). However, the derived nucleoside phosphoramidites couple less efficiently than the N-acyl-protected nucleoside phosphoramidites (Sekine et al., 1985). Depurination is also influenced by other factors such as the nature of the solid support (controlled-pore glass versus polystyrene; see UNIT 3.1), the composition of the deblocking solution and deblocking time, and the washing solvent and washing time that are employed in solid-phase synthesis of oligonucleotides (Paul and Royappa, 1996). Depurination is faster at terminal sites than at internal sites in an oligonucleotide chain (Suzuki et al., 1994). Interestingly, a solution of 15% dichloroacetic acid (DCA) in methylene chloride was ideal as a detritylating reagent that induced minimal depurination compared to the traditionally used 2% DCA/methylene chloride. It is pertinent that with modern DNA synthesizers the pulsed delivery of reagents to the synthesis columns, in conjunction with optimized synthesis programs, results in short contact times and has greatly minimized the depurination problem. Thus, N-acyl-protected nucleoside (benzoyl for dA and dC, and isobutyryl for dG) phosphoramidites and H-phosphonates can be used for the efficient synthesis of oligonucleotides.
RECENT TRENDS IN NUCLEOBASE PROTECTION
Nucleobase Protection of Deoxyribo- and Ribonucleosides
Over the past few years, new applications of oligonucleotides in diagnostics and therapeutics have emerged, necessitating the expeditious synthesis of large numbers of oligonucleotides. In order to speed the synthesis process, more “labile” N-acyl protecting groups for nucleosides have been sought. As the simplest
member of the family of N-acyl protecting groups, the N4-acetyl of cytosine has been used for “ultrafast” DNA synthesis using the phosphoramidite approach. Rapid deprotection rates were achieved using methylamine/ammonia (Reddy et al., 1994). This group is unsuitable for use in phosphodiester and phosphotriester chemistries, however. The synthesis of oligonucleotide analogs carrying sensitive backbones requires protecting groups that (1) withstand the synthetic rigors of chain assembly, and (2) can be removed chemoselectively under mild conditions. Nucleobases protected by phenoxyacetyl (Singh and Misra, 1988; Chaix et al., 1989; Sproat et al., 1991), and their derivatives such as t-PAC, show accelerated deacylation (Köster et al., 1981; Sinha et al., 1993) under mildly basic conditions. It is presumed that the inductive effect of the phenoxy group renders the amide carbonyl group more susceptible to nucleophilic attack, facilitating rapid base-catalyzed hydrolysis. Thus, PAC- and t-PAC-phosphoramidites have been employed for rapid synthesis of oligonucleotides and certain analogs. As a rule, following oligonucleotide assembly on solid support, the PAC and t-PAC groups are removed under milder conditions (28% NH4OH, room temperature; Sinha et al., 1993) or, according to a recent report, using gaseous amines under pressure (Boal et al., 1996). However, the use of t-PAC-protected nucleoside phosphoramidites results in trans acylation of the t-PAC protecting groups during the capping step when acetic anhydride is employed as a capping reagent. Thus, tert-butylphenoxyacetic anhydride should be used for the capping step in solid-phase oligonucleotide synthesis (Sinha et al., 1993). The concept of neighboring-group participation has also been used to design acyl protecting groups and to accelerate the deprotection step (Dreef-Tromp et al., 1990; Kuijpers et al., 1990). For example, removal of the 2-(acetoxymethyl)benzoyl group from nucleosides under basic conditions is accelerated by intramolecular participation of the deacylated hydroxymethyl group. New protecting groups have also been introduced that can be chemoselectively removed under neutral or mildly basic conditions. Indeed, the allyloxycarbonyl protecting group (Hayakawa et al., 1986, 1990) is chemoselectively removed using Pd(0), whereas the (p-nitrophenyl)ethoxycarbonyl group (Trichtinger et al., 1983; Pfleiderer et al., 1985; Pfister et al., 1988) and the 2-dansylethoxy cabonyl group
2.1.12 Current Protocols in Nucleic Acid Chemistry
(Wagner and Pfleiderer, 1997) are selectively removed using DBU. Consequently, supportbound oligonucleotides can be prepared using building blocks that carry these protecting groups. Recently, the N-pent-4-enoyl (PNT) group has been introduced as a new acyl protecting group for situations where multiple deprotection protocols are used (Iyer et al., 1997; see also references therein). PNT-protected nucleoside phosphoramidites can potentially be used for the rapid synthesis of oligonucleotides and oligonucleotide analogs, as well as for the preparation of support-bound oligonucleotides. Several groups have been reported for nucleobase protection and may be particularly valuable in the synthesis of oligonucleotides intended for specific applications. Figure 2.1.7 shows a partial list of protecting groups. A complete list of such groups is covered in other reviews (Sonveaux, 1986; Beaucage and Iyer, 1992; Iyer and Beaucage, 1999). Nonetheless, it should be noted that many of these still need to be evaluated in routine synthesis. Because of various issues associated with nucleobase protection, oligonucleotide synthesis has been evaluated using building blocks bearing unprotected nucleobases. However, these efforts have met with only limited success (Narang et al., 1972; Fourrey and Varenne, 1985; Gryaznov and Letsinger, 1991; Uchiyama et al., 1993, and references therein). Clearly, more work is necessary in this area.
CONCLUSION Evidently, a nucleobase protecting group should meet several criteria before it can be adapted for routine oligonucleotide synthesis. It is imperative, therefore, that the potential for side reactions be closely examined when designing new reagents, evaluating new protecting groups, and implementing modifications of established protocols during oligonucleotide synthesis, as well as during the manufacture of oligonucleotides and their analogs. The development of nucleobase protecting groups and deprotection protocols has been crucial for the successful synthesis of oligonucleotides, functionalized oligonucleotides, oligonucleotide analogs, ribonucleotides, and phosphorylated biomolecules (reviewed in Beaucage and Iyer, 1992, 1993a,b,c). As in the past, oligonucleotides are expected to play a dominant role in fostering advances in functional genomics, proteomics, diagnostics, and therapeutics. Consequently, continued demand exists for simultaneous synthesis of large numbers of oligonucleotides in miniature formats,
and for the synthesis and manufacture of novel analogs. It is hoped that the present commentary will serve as a framework for developing new protecting group strategies for oligonucleotide synthesis that could meet these challenges.
ACKNOWLEDGMENT The author wishes to thank Dr. WenQiang Zhou for his help with the artwork in this manuscript.
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Protection of Nucleosides for Oligonucleotide Synthesis
2.1.13 Current Protocols in Nucleic Acid Chemistry
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2.1.14 Current Protocols in Nucleic Acid Chemistry
Holy, A. and Zemlicka, J. 1969. Oligonucleotidic compounds. XXXIII. A study on hydrolysis of N-dimethylaminomethylenecytidine, -adenosine, -guanosine, and related 2′-deoxy compounds. Collect. Czech. Chem. Commun. 34:2449-2458. Honda, S., Urakami, K., Koura, K., Terada, K., Sato, Y., Kohno, K., Sekine, M., and Hata, T. 1984. Synthesis of oligoribonucleotides by use of S,Sdiphenyl N-monomethoxytrityl ribonucleoside 3′-phosphorodithioates. Tetrahedron 40:153163. Huynh-Dinh, T., Langlois d’Estaintot, B., Allard, P., and Igolen, J. 1985. Synthèse simplifiée de sondes mixtes avec des triazolo-nucléosides. Tetrahedron Lett. 26:431-434. Igolen, J. and Morin, C. 1980. Rapid synthesis of protected 2′-deoxycytidine derivatives. J. Org. Chem. 45:4802-4804. Ito, T., Ued a, S., and Takaku, H. 19 86. (Methoxyethoxy)methyl group: New amide and hydroxyl protecting groups of uridine in oligonucleotide synthesis. J. Org. Chem. 51:931933. Iyer, R.P. and Beaucage, S.L. 1999. Oligonucleotide synthesis. In Comprehensive Natural Products Chemistry, Vol. 7: DNA and Aspects of Molecular Biology (E.T. Kool, ed.) pp. 105-152. Elsevier Science Publishing, New York. Iyer, R.P., Yu, D., Habus, I., Ho, N.H., Johnson, S., Devlin, T., Jiang, Z., Zhou, W., Xie, J., and Agrawal, S. 1997. N-Pent-4-enoyl (PNT) group as a universal nucleobase protector: Applications in the rapid and facile synthesis of oligonucleotides, analogs and conjugates. Tetrahedron 53:2731-2750. Jones, S.S., Reese, C.B., Sibanda, S., and Ubasawa, A. 1981. The protection of uracil and guanine residues in oligonucleotide synthesis. Tetrahedron Lett. 22:4755-4758.
Krecmerová, M., Hrebabecky, H., and Holy, A. 1990. Synthesis of 5′-O-phosphonomethyl derivatives of pyridine 2′-deoxynucleosides. Collect. Czech. Chem. Commun. 55:2521-2536. Kuijpers, W.H.A., Huskens, J., and van Boeckel, C.A.A. 1990. The 2-(acetoxymethyl)benzoyl (AMB) group as a new base-protecting group, designed for the protection of phosphate modified oligonucleotides. Tetrahedron Lett. 31:6729-6732. Kume, A., Sekine, M., and Hata, T. 1982. Phthaloyl group: A new amino protecting group of deoxyadenosine in oligonucleotide synthesis. Tetrahedron Lett. 23:4365-4368. Kume, A., Iwase, R., Sekine, M., and Hata, T. 1984. Cyclic diacyl groups for protection of the N6amino group of deoxyadenosine in oligodeoxynucleotide synthesis. Nucl. Acids Res. 12:8525-8538. Letsinger, R.L. and Ogilvie, K.K. 1969. Synthesis of oligothymidylates via phosphotriester intermediates. J. Am. Chem. Soc. 91:3350-3355. Letsinger, R.L. and Lunsford, W.B. 1976. Synthesis of thymidine oligonucleotides by phosphite triester intermediates. J. Am. Chem. Soc. 98:36553661. Li, B.F.L., Reese, C.B., and Swann, P.F. 1987. Synthesis and characterization of oligodeoxynucleotides containing 4-O-methylthymine. Biochemistry 26:1086-1093. MacMillan, A.M. and Verdine, G.L. 1991. Engineering tethered DNA molecules by the convertible nucleoside approach. Tetrahedron 47:26032616. Mag, M. and Engels, J.W. 1988. Synthesis and structure assignments of amide protected nucleosides and their use as phosphoramidites in deoxyoligonucleotide synthesis. Nucl. Acids Res. 16:3525-3543. Marugg, J.E., Tromp, M., Jhurani, P., Hoyng, C.F., van der Marel, G.A., and van Boom, J.H. 1984. Synthesis of DNA fragments by the hydroxybenzotriazole phosphodiester approach. Tetrahedron 40:73-78.
Kamaike, K., Hasegawa, Y., and Ishido, Y. 1988. A simple, preparative procedure for N3-anisoyluridine and O6-diphenylcarbamoylguanosine 2′O-(tetrahydropyran-2-yl) derivatives via the corresponding 3′,5′-dibenzoates. Nucleosides Nucleotides 7:37-43.
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Protection of Nucleosides for Oligonucleotide Synthesis
2.1.15 Current Protocols in Nucleic Acid Chemistry
Mullah, B., Andrus, A., Zhao, H., and Jones, R.A. 1995. Oxidative conversion of N-dimethylformamidine nucleosides to N-cyano nucleosides. Tetrahedron Lett. 36:4373-4376.
Prasad, A.K. and Wengel, J. 1996. Enzyme-mediated protecting group chemistry on the hydroxyl groups of nucleosides. Nucleosides Nucleotides 15:1347-1359.
Nagaich, A.K. and Misra, K. 1989. Highly efficient synthesis of oligodeoxyribonucleotides using αphenyl cinnamoyl group for selective amino protection. Nucl. Acids Res. 17:5125-5134.
Rao, T.S., Reese, C.B., Serafinowska, H.T., Takaku, H., and Zappia, G. 1987. Solid-phase synthesis of the 3′-terminal nonadecaribonucleoside octadecaphosphate sequence of yeast alanine transfer ribonucleic acid. Tetrahedron Lett. 28:48974900.
Narang, S.A., Itakura, K., and Wightman, R.H. 1972. A simplification in the synthesis of deoxyribooligonucleotides. Can. J. Chem. 50:769770. Nielsen, J., Dahl, O., Remaud, G., and Chattopadhyaya, J. 1987. Phosphitylation of guanine or inosine bases during the preparation of nucleoside phosphoramidites. Isolation of model products as thiophosphoric amide derivatives and structure elucidation by 15N NMR spectroscopy. Acta. Chem. Scand. B41:633-639. Nyilas, A., Zhou, X.-X., Welch, C.J., and Chattopadhyaya, J. 1987. A versatile strategy for the O4protection and modification of the lactam function of uridine and uridylic acid. Nucl. Acids Res. Symp. Ser. 18:157-160. Nyilas, A., Földesi, A., and Chattopadhyaya, J. 1988. Arenesulfonylethoxycarbonyl—A set of amino protecting groups for DNA and RNA synthesis. Nucleosides Nucleotides 7:787-793. Ogilvie, K.K., Nemer, M.J., Hakimelahi, G.H., Proba, Z.A., and Lucas, M. 1982. N-Levulination of nucleosides. Tetrahedron Lett. 23:26152618. Oivanen, M., Lönnberg, H., Zhou, X.X., and Chattopadhyaya, J. 1987. Acidic hydrolysis of 6-substituted 9-(2-deoxy-β-D-erythro-pentofuranosyl)purines and their 9-(1-alkoxyethyl) counterparts: Kinetics and mechanism. Tetrahedron 43:1133-1140. Paul, C.H. and Royappa, A.T. 1996. Acid binding and detritylation during oligonucleotide synthesis. Nucl. Acids Res. 24:3048-3052. Pfister, M. and Pfleiderer, W. 1989. New results in oligoribonucleotide synthesis. Nucleosides Nucleotides 8:1001-1006. Pfister, M., Farkas, S., Charubala, R., and Pfleiderer, W. 1988. Recent progress in oligoribonucleotide synthesis. Nucleosides Nucleotides 7:595-600. Pfleiderer, W., Himmelsbach, F., Charubala, R., Schirmeister, H., Beiter, A., Schulz, B., and Trichtinger, T. 1985. The p-nitrophenylethyl group—An universal blocking group in nucleoside and nucleotide chemistry. Nucleosides Nucleotides 4:81-94. Pon, R.T., Damha, M.J., and Ogilvie, K.K. 1985a. Necessary protection of the O6-position of guanosine during the solid phase synthesis of oligonucleotides by the phosphoramidite approach. Tetrahedron Lett. 26:2525-2528. Nucleobase Protection of Deoxyribo- and Ribonucleosides
Pon, R.T., Damha, M.J., and Ogilvie, K.K. 1985b. Modification of guanine bases by nucleoside phosphoramidite reagents during the solid phase synthesis of oligonucleotides. Nucl. Acids Res. 13:6447-6465.
Reddy, M.P., Hanna, N.B., and Farooqui, F. 1994. Fast cleavage and deprotection of oligonucleotides. Tetrahedron Lett. 35:4311-4314. Reese, C.B. 1978. The chemical synthesis of oligoand poly-nucleotides by the phosphotriester approach. Tetrahedron 34:3143-3179. Reese, C.B. and Ubasawa, A. 1980. Reaction between 1-arenesulphonyl-3-nitro-1,2,4-triazoles and nucleoside residues. Elucidation of the nature of side-reactions during oligonucleotide synthesis. Tetrahedron Lett. 21:2265-2268. Reese, C.B. and Skone, P.A. 1984. The protection of thymine and guanine residues in oligodeoxyribonucleotide synthesis. J. Chem. Soc. Perkin Trans. 1:1263. Romero, R., Stein, R., Bull, H.G., and Cordes, E.H. 1978. Secondary deuterium isotope effects for acid-catalyzed hydrolysis of inosine and adenosine. J. Am. Chem. Soc. 100:7620-7624. Saenger, W. 1984. Principles of Nucleic Acids Structure. Springer-Verlag, New York. Scalfi-Happ, C., Happ, E., and Chládek, S. 1987. New approach to the synthesis of 2′(3′)-O-aminoacyl-oligoribonucleotides related to the 3′-terminus of aminoacyl transfer ribonucleic acid. Nucleosides Nucleotides 6:345-348. Schaller, H., Weimann, G., Lerch, W.B., and Khorana, H.G. 1963. Studies on polynucleotides. XXIV. The stepwise synthesis of specific deoxyribopolynucleotides (4). Protected derivatives of deoxyribonucleosides and new syntheses of deoxyribonucleoside-3′ phosphates. J. Am. Chem. Soc. 85:3821-3827. Schulhof, J.C., Molko, D., and Teoule, R. 1987. Facile removal of new base protecting groups useful in oligonucleotide synthesis. Tetrahedron Lett. 28:51-54. Seela, F. and Driller, H. 1989. 7-Deaza-2′-deoxyO6-methylguanosine: Selective N2-formylation via a formamidine, phosphoramidite. Synthesis and properties of oligonucleotides. Nucleosides Nucleotides 8:1-21. Sekine, M. 1989. General method for the preparation of N3- and O4-substituted uridine derivatives by phase-transfer reactions. J. Org. Chem. 54:2321-2326. Sekine, M., Masuda, N., and Hata, T. 1985. Introduction of the 4,4′,4′′-tris(benzoyloxy)trityl group into the exo amino groups of deoxyribonucleosides and its properties. Tetrahedron 41:5445-5453.
2.1.16 Current Protocols in Nucleic Acid Chemistry
Shabarova, Z. and Bogdanov, A. 1994. Advanced organic chemistry of nucleic acids. VCH Publishers, New York. Shimidzu, T. and Letsinger, R.L. 1968. Synthesis of deoxyguanylyl-deoxyguanosine on an insoluble polymer support. J. Org. Chem. 33:708-711. Singh, R.K. and Misra, K. 1988. Improvements in oligodeoxyribonucleotide synthesis using phenoxyacetyl as amino protecting group. Indian J. Chem. 27B:409-417. Sinha, N.D., Davis, P., Usman, N., Pérez, J., Hodge, R., Kremsky, J., and Casale, R. 1993. Labile exocyclic amine protection of nucleosides in DNA, RNA and oligonucleotide analog synthesis facilitating N-deacylation, minimizing depurination and chain degradation. Biochimie 75:1323. Smrt, J. and Sorm, F. 1967. Oligonucleotidic compounds. XVIII. Synthesis of guanylyl-(3′-5′)uridylyl-(3′-5′)-arabinofuranosyluracil and guanylyl-(3′-5′)-uridylyl-(3′-5′)-arabinofurano -sylcytosine. Collect. Czech. Chem. Commun. 32:3169-3176. Sonveaux, E. 1986. The organic chemistry underlying DNA synthesis. Bioorganic Chem. 14:274325. Sproat, B.S., Iribarren, A.M., Guimil Garcia, R., and Beijer, B. 1991. New synthetic routes to synthons suitable for 2′-O-allyloligoribonucleotide assembly. Nucl. Acids Res. 19:733-738. Sung, W.L. 1982. Synthesis of 4-(1,2,4-triazol-1yl)pyrimidin-2(1H)-one-ribonucleotide and its application in synthesis of oligoribonucleotides. J. Org. Chem. 47:3623-3628. Suzuki, T., Ohsumi, S., and Makino, K. 1994. Mechanistic studies on depurination and apurinic site chain breakage in oligodeoxyribonucletides. Nucl. Acids Res. 22:4997-5003. Takaku, H., Imai, K., and Nagai, M. 1988. Triphenylmethanesulfenyl group. A new protecting group for the uracil residue in oligoribonucleotide synthesis. Chem. Lett. 857-860. Tanimura, H., Fukazawa, T., Sekine, M., Hata, T., Efcavitch, J.W., and Zon, G. 1988. The practical synthesis of RNA fragments in the solid phase approach. Tetrahedron Lett. 29:577-578. Ti, G.S., Gaffney, B.L., and Jones, R.A. 1982. Transient protection: Efficient one-flask synthesis of protected deoxynucleosides. J. Am. Chem. Soc. 104:1316-1319. Trichtinger, T., Charubala, R., and Pfleiderer, W. 19 83. Synth esis of O6-p-nitrophenylethyl guanosine and 2′-deoxyguanosine derivatives. Tetrahedron Lett. 24:711-714. Uchiyama, M., Aso, Y., and Noyori, R. 1993. O-Selective phosphorylation of nucleosides without N-protection. J. Org. Chem. 58:373-379.
Urdea, M.S., Ku, L., Horn, T., Gee, Y.G., and Warner, B.D. 1986. Base modification and cloning efficiency of oligodeoxyribonucleotides synthesized by the phosphoramidite method: Methyl versus cyanoethyl phosphorous protection. Nucl. Acids Res. Symp. Ser. 16:257-260. Uznanski, B., Grajkowski, A., and Wilk, A. 1989. The isopropoxyacetic group for convenient base protection during solid-support synthesis of oligodeoxyribonucleotides and their triester analogs. Nucl. Acids Res. 17:4863-4871. Van Aerschot, A., Herdewijn, P., Janssen, G., and Vanderhaeghe, H. 1988. Protection of the lactam function of 2′-deoxyinosine with a 2-(4-nitrophenyl)-ethyl moiety. Nucleosides Nucleotides 7:519-536. Vu, H., McCollum, C., Jacobson, K., Theisen, P., Vinayak, R., Spiess, E., and Andrus, A. 1990. Fast oligonucleotide deprotection phosphoramidite chemistry for DNA synthesis. Tetrahedron Lett. 31:7269-7272. Wagner, T. and Pfleiderer, W. 1997. Aglycone protection by the (2-dansylethoxy)carbonyl (= {2{[5-(dimethylamino)naphthalen-1-yl] sulfonyl}ethoxy}carbonyl; dnseoc) group—A new variation in oligodeoxyribonucleotide synthesis. Helv. Chim. Acta 80:200-212. Watanabe, K.A. and Fox, J.J. 1966. A simple method for selective acylation of cytidine on the 4-amino group. Angew. Chem. Intl. Ed. Engl. 5:579-580. Welch, C.J., Bazin, H., Heikkilä, J., and Chattopadhyaya, J. 1985. Synthesis of C-5 and N-3 arenesulfenyl uridines. Preparation and properties of a new class of uracil protecting group. Acta Chem. Scand. B39:203-212. York, J.L. 1981. Effect of structure of the aglycon on the acid-catalyzed hydrolysis of adenine nucleosides. J. Org. Chem. 46:2171-2173. Zhou, X.-X. and Chattopadhyaya, J. 1986. Site-specific modification of the pyrimidine residue during the deprotection of the fully-protected diuridylic acid. Tetrahedron 42:5149-5156. Zhou, X.X., Sandström, A., and Chattopadhyaya, J. 1986. A convenient preparation of 2-N-(4-tbutylbenzoyl)-6-O-(2-nitrophenyl)guanosine and its application in the synthesis of 5′(GpGpGpU)3′ constituting the 3′-anticodon stem of E.coli tRNAIle. Chem. Scr. 26:241-249. Zoltewicz, J.A. and Clark, D.F. 1972. Kinetics and mechanism of the hydrolysis of guanosine and 7-methylguanosine nucleosides in perchloric acid. J. Org. Chem. 37:1193-1197. Zoltewicz, J.A., Clark, D.F., Sharpless, T.W., and Grahe, G. 1970. Kinetics and mechanism of the hydrolysis of some purine nucleosides. J. Am. Chem. Soc. 92:1741-1750.
Contributed by Radhakrishnan P. Iyer OriGenix Technologies Laval, Quebec, Canada
Protection of Nucleosides for Oligonucleotide Synthesis
2.1.17 Current Protocols in Nucleic Acid Chemistry
Protection of 2′-Hydroxy Functions of Ribonucleosides The methods used to protect 2′-hydroxy functions of ribonucleosides have recently been reviewed (Beaucage and Iyer, 1992; Sonveaux, 1994; Beaucage and Caruthers, 1996). In addition, there have been earlier brief reviews (Ohtsuka and Iwai, 1987; Reese, 1989). The main purpose of this article is to discuss 2′-protection in the context of effective oligoribonucleotide synthesis. For this reason, emphasis will be placed on what are now, or are likely to become, the 2′-protecting groups of choice in the synthesis of oligo- and poly-ribonucleotides (RNA sequences). As a result, only some of the protecting groups that have been suggested for this purpose are considered in detail here, and some interesting chemistry has necessarily been omitted.
CONSIDERATIONS FOR 2′-PROTECTING GROUPS IN OLIGORIBONUCLEOTIDE SYNTHESIS There are three main general criteria that all protecting groups should fulfill (Reese, 1978). (1) They should be easy to introduce and, as part of this criterion, the reagents involved in their introduction should be readily accessible. (2) They should be stable and remain intact until it is appropriate to remove them. (3) They should be removable at the appropriate time using conditions under which the desired product is completely stable. In the case of chiral substrates such as ribonucleosides, achiral protecting groups are desirable for analytical (e.g., NMR, TLC, and HPLC) purposes. In the case of all substrates, it is desirable that the introduction of protecting groups should not result in unduly complex NMR spectra.
O
The successful chemical synthesis of polynucleotides (including RNA sequences) depends on the choice of suitable protecting groups and effective phosphorylation procedures. Arguably the most crucial single decision that has to be made in oligoribonucleotide synthesis is the choice of the protecting group (R; see S.1) for the 2′-hydroxy functions (Reese, 1978). This protecting group must remain intact until the very last step of the synthesis (Fig. 2.2.1), and must then be removable under conditions that are mild enough to avoid subsequent attack of the released 2′-hydroxy functions (see S.2) on the vicinal phosphodiester internucleotide linkages, thereby leading to their cleavage or migration. Protecting groups are often removed hydrolytically under either basic or acidic conditions. Cleavage of interribonucleotide linkages can occur under relatively mild basic conditions (Järvinen et al., 1991; Kuusela and Lönnberg, 1994). This process, illustrated in Figure 2.2.2A, essentially involves an ester exchange reaction between the 2′-hydroxy function of the 3′-linked nucleoside residue and the 5′-hydroxy function of the 5′-linked nucleoside residue, leading to a (2′,3′)-cyclic phosphate (S.3). This intermediate then undergoes further basecatalyzed hydrolysis to give a mixture of isomeric 2′- and 3′-phosphates (S.5 and S.6, respectively). Under acidic conditions (Fig. 2.2.2B), internucleotide cleavage and migration can both occur (Griffin et al., 1968). These processes are both believed to proceed via a phosphorane intermediate (S.7; Järvinen et al., 1991). If the P-O(2′) bond is then cleaved, starting material S.2 is regenerated. If the PO(3′) bond is cleaved, the isomeric product S.8
B
O
O
O P O O
O O
B'
Figure 2.2.1
OH
O P O O
OR
1
B
O O
OR
O
UNIT 2.2
O O
B'
OH
2
Scheme showing protected 2′-hydroxy functions. B and B′ are bases.
Contributed by Colin B. Reese Current Protocols in Nucleic Acid Chemistry (2000) 2.2.1-2.2.24 Copyright © 2000 by John Wiley & Sons, Inc.
Protection of Nucleosides for Oligonucleotide Synthesis
2.2.1
O O
B
O
O O
O
OH
B
O
A
P
O
O
HO
O
O
O P O
3
HO
B
O
OH
5
HO
O P O O
O O
B' HO
B'
O
O
B
O
OH O
2
OH
O
OH
O P O
4
OH 6
B O
B
O 3' 2' O HO
O
B
O O
OH
P
O
O OH
5'
O
O
H3O
B' O OH
H3O
7
O P O O
O O
B
O HO
O O P O
B'
O H3O
OH
O O
B'
OH
8
2
O
B
O O O
P
HO O O
3
O O
B'
OH 4
Figure 2.2.2 Scheme showing cleavage of interribonucleotide linkages under (A) basic and (B) acidic conditions. Although only shown in panel A, the hydrolysis of S.3 can yield either the 2′-phosphate (S.5) or the 3′-phosphate (S.6) under either basic or acidic conditions.
Protection of 2′Hydroxy Functions of Ribonucleosides
with the migrated internucleotide linkage is obtained. Finally, if the P-O(5′) bond is cleaved, the (2′,3′)-cyclic phosphate S.3 is obtained. The cyclic phosphate S.3 undergoes further hydrolysis to give an isomeric mixture of the corresponding 2′- and 3′-phosphates (S.5 and S.6, respectively) under acidic as well as under basic conditions. The significance of these reactions in the context of oligo- and poly-ribonucleotide synthesis will be considered later. However, it is clearly of crucial importance that the 2′-protecting group should strictly satisfy the above criteria (1) and (2). As will become apparent, this is a very demanding requirement, as the 2′-pro-
tecting group must also be fully compatible with the groups that are used to protect the 5′-terminal hydroxy function, the base residues, and the internucleotide linkages. It is therefore appropriate to consider these other protecting group requirements at the outset.
Protection of the 5′-Terminal Hydroxy Function Figure 2.2.3 illustrates a number of groups (R′ in S.9) used to protect the 5′-terminal hydroxy function. Although a good deal of work has been carried out on the synthesis of oligoribonucleotides in solution, most of the recent studies in this area have been concerned with
2.2.2 Current Protocols in Nucleic Acid Chemistry
R'O
B
O O
Ph
Ph C
R
OMe O
OR
9
10a, R = H (MMTr) 10b, R = OMe (DMTr)
CHBr2
O O
11 (Px)
O O
O
O 12 (Fmoc)
Figure 2.2.3
13 (Lev)
14 (Dbmb)
S
O
15 (Ptmt)
Several protecting groups for 5′-terminal hydroxy functions.
solid-phase synthesis. The 5′-terminal protecting group (R′) that has been used most widely for this purpose is the (di-p-anisyl)phenylmethyl group (also known as 4,4′-dimethoxytrityl or DMTr; S.10b; Schaller et al., 1963; UNIT 2.3). The 9-phenylxanthen-9-yl (pixyl or Px, S.11) group has very similar properties to the DMTr group and is equally suitable (Chattopadhyaya and Reese, 1978). The somewhat less labile p-anisyl(diphenyl)methyl group (4-monomethoxytrityl or MMTr; S.10a; Schaller et al., 1963) has also been used, but its greater stability to acid makes it generally less suitable. The great advantage of the DMTr (S.10b) and Px (S.11) protecting groups in solid-phase synthesis, and perhaps also in solution-phase synthesis, is that they can be rapidly and quantitatively removed by treatment with acids, such as diand tri-chloroacetic acids, in anhydrous dichloromethane solution (Sproat and Gait, 1984). A further advantage shared by all three of these protecting groups is that with acid treatment they give rise to colored carbocations that can easily be assayed spectrophotometrically. This permits coupling efficiencies to be monitored. Clearly, if one of these three protecting groups is used in oligoribonucleotide synthesis, the 2′-protecting group (R in formula S.9) must be completely stable under the acidic conditions required for 5′-deprotection. Numerous other 5′-protecting groups have been suggested (Sonveaux, 1994; UNIT 2.3), some of which are removable under mildly basic or virtually neutral conditions. Protecting groups in this latter category include 9-fluorenylmethoxycarbonyl (Fmoc; S.12; Pathak and
Chattopadhyaya, 1985), levulinyl (Lev; S.13; van Boom and Burgers, 1976), 2-(dibromomethyl)benzoyl (Dbmb; S.14; Chattopadhyaya et al., 1979), and 2-(isopropylthiomethoxymethyl)benzoyl (Ptmt; S.15; Brown et al., 1989a). None of these protecting groups has found widespread use in the solidphase synthesis of RNA sequences, but some have proved to be useful in solution-phase synthesis.
Protection of Base Residues The protection of base residues is illustrated in Figure 2.2.4. In the solid-phase synthesis of RNA sequences (Rao et al., 1993), adenine, cytosine, and guanine residues are generally protected by N-acylation (as in S.16, S.18, and S.19, respectively), while uracil residues are left unprotected (as in S.23; UNIT 2.1). The N-acyl protecting groups are usually removed by ammonolysis in the step before the removal of the 2′-protecting groups. As RNA can undergo internucleotide cleavage (Fig. 2.2.2A) under ammonolytic conditions, the base-protecting groups must be removable using conditions under which the 2′-protecting groups are completely stable. Thus, the choice of an N-acyl protecting group for a particular base residue is, to some extent, dependent on the 2′-protecting group used. The dimethylaminomethylene protecting group, which is also removable under ammonolytic conditions, has been recommended for the protection of adenine and guanine residues (as in S.17 and S.20, respectively; Vinayak et al., 1992). Particularly in the solution-phase synthesis of RNA sequences, it may be desir-
Protection of Nucleosides for Oligonucleotide Synthesis
2.2.3 Current Protocols in Nucleic Acid Chemistry
O
O HN N N
N
R N
N
N
N
HN
NMe2
N
N
N O
N
NH
N
N
N
18
17
16
O
R
O
N H
R
19 O
O N N
NH N
N
N
N
NMe2
OAr
HN
N N
N H
N
R
21
O
O
N
O
N
20
O
O R
N
N H
N
R
O NO2
N O
24
O
N
22
N
N N
23
NPh2
O
OAr N
N
OH
26
25
OH
27
Figure 2.2.4 Several protecting groups for base residues (A: S.16 and S.17; C: S.18; G: S.19 to S.22; U: S.23 to S.25). S.26 and S.27 are used in oximate treatment for the removal of aryl (Ar) groups.
able to protect guanine residues on O6 as well as on N2 (as in S.21 and S.22). Aryl protecting groups are particularly suitable for this purpose (as in S.21; Ar = 2-nitrophenyl, 3-chlorophenyl, and 3,5-dichlorophenyl; Jones et al., 1981; Reese and Skone, 1984; Brown et al., 1989a); they may readily be removed by treatment with the N1,N1,N3,N3-tetramethylguanidinium salt of (E)-2-nitrobenzaldoxime S.26 or of (E)pyridine-2-carboxaldoxime S.27 (oximate treatment; Reese and Zard, 1981) before the ammonolytic removal of the N-acyl protecting groups. The N,N-diphenylcarbamoyl group (as in S.22; Kamimura et al., 1984) is removable by ammonolysis (UNIT 2.1). In the solution-phase synthesis of RNA sequences, it may also be desirable to protect
O
NC
B
O
O O
Protection of 2′Hydroxy Functions of Ribonucleosides
28
Figure 2.2.5 group.
Protection of Internucleotide Linkages Virtually all of the groups commonly used to protect the internucleotide linkages in both solid- and solution-phase oligo- and poly-ribonucleotide synthesis are removed under basic conditions (Fig. 2.2.5). The 2-cyanoethyl group (as in S.28; Sinha et al., 1983) is by far
O
O OR O P O O
uracil residues on O4 with an aryl group (as in S.24; Ar = 2,4-dimethylphenyl; Jones et al., 1981) or on N3 with an acyl group (as in S.25; R = 4-MeO.C6H4; Kamimura et al., 1984). O4-Aryl and N3-acyl protecting groups may be removed from uracil residues by oximate treatment and by ammonolysis, respectively (UNIT 2.1). It should be noted that the ammonolytic and oxime treatment conditions are both basic.
O
OR
O P O O
B'
O
Cl OR
O 29
O
B
O
B'
B
O
O OR ArS P O O O O
OR
B'
OR
30
Several protecting groups for internucleotide linkages. Ar is phenyl or another aryl
2.2.4 Current Protocols in Nucleic Acid Chemistry
the most commonly used protecting group for the internucleotide linkages in the solid-phase synthesis of RNA sequences, and the 2-chlorophenyl group (as in S.29; Reese, 1970) has been widely used for this purpose in solutionphase synthesis. Another approach to the synthesis of oligoribonucleotides was pioneered by Hata and co-workers (Honda et al., 1984) and involves intermediate S-aryl phosphorothioates (S.30, Ar = Ph). 2-Cyanoethyl protecting groups are usually removed at the same time as N-acyl base-protecting groups by treatment with ammonia (Sinha et al., 1983). 2-Chlorophenyl-protected oligo- and poly-ribonucleotides are best unblocked by treatment with the conjugate base of (E)-2-nitrobenzaldoxime S.26 or (E)-pyridine-2-carboxaldoxime S.27 (Reese et al., 1978; Reese and Zard, 1981). S-Aryl phosphorothioates (S.30), which are masked phosphodiesters, may also be unblocked by oximate treatment (Kamimura et al., 1984). In order to avoid internucleotide cleavage (Fig. 2.2.2A), it is necessary that the 2′-protecting group (R in S.28, S.29, and S.30) should be completely stable under the basic conditions used in the unblocking of the internucleotide linkages.
tions required to remove the 5′-terminal DMTr protecting group S.10b. The 2′-protecting groups must also be stable under the basic conditions (i.e., concentrated aqueous ammonia and oximate ions) required to unblock the base residues and the internucleotide linkages. It is further desirable that 2′-protecting groups not be excessively bulky and thereby impede the coupling process. For the successful removal of the 2′-protecting groups, it must always be borne in mind that RNA is a very sensitive material that is unstable under both acidic and basic conditions and in the presence of various hydrolytic enzymes. It is therefore desirable that manipulation should be kept to a minimum in the isolation of fully unblocked RNA.
Ether Protecting Groups The protection of the 2′-hydroxy functions as readily cleavable ether groups would appear at first sight to be an attractive proposition. Indeed, the possibility of using the benzyl protecting group was first examined over 30 years ago (Griffin et al., 1966). Uridine was converted into its 2′-O-benzyl derivative S.31, which was then successfully converted via the protected dinucleoside phosphate S.32 into uridylyl(3′→5′)-uridine (S.33; Fig. 2.2.6). The benzyl protecting group, which is stable both to acidand base-catalyzed hydrolysis, was removed by catalytic hydrogenolysis in the presence of palladized charcoal. However, it was subsequently reported that concomitant hydrogenation of the uracil 5,6-double bond can occur (Reitz and Pfleiderer, 1975). There is also a danger that the total removal of all of the 2′-protecting groups of per-2′-O-benzylated RNA sequences may not always be possible. The 2-nitrobenzyl group (as in S.34; Fig. 2.2.7), which was introduced by Ikehara and
PROTECTION OF THE 2′-HYDROXY FUNCTION IN OLIGORIBONUCLEOTIDE SYNTHESIS It is clear from the above discussion that the requirements for a 2′-protecting group in oligoand polyribonucleotide synthesis are very demanding indeed. With regard to solid-phase synthesis, in addition to meeting the above general criteria for protecting groups, it is crucially important that 2′-protecting groups be stable to repeated exposure to the acidic condi-
Me3C O
N
O HO
Ura
O
HO
O
O
OBn
O P O O
O
Ura
O
O
H
OMe
32
31
OH
O
O
Ura
O
O NH
HO
O
O P O O
O HO
Ura
OH
33
Ura = uracil-1-yl Bn = PhCH2
Figure 2.2.6 uridine.
Scheme showing the preparation of uridylyl-(3′→5′)-uridine from 2′-O -benzyl (Bn)-
Protection of Nucleosides for Oligonucleotide Synthesis
2.2.5 Current Protocols in Nucleic Acid Chemistry
O
B
O O O O P O O
O
O
B OMe
O O O P O
NO2
R
O 35a, R = H 35b, R = OMe
34
Figure 2.2.7 The 2-nitrobenzyl (S.34), 4-methoxybenzyl (S.35a), and 3,4-dimethoxybenzyl (S.35b) protecting groups.
co-workers (Ohtsuka et al., 1978), is potentially a more useful 2′-protecting group. Like the benzyl group, it is stable both to acid- and base-catalyzed hydrolysis. However, it may be cleaved photochemically by irradiation with ultraviolet light (λ > 280 nm). It was later reported that photolytic cleavage of the 2-nitrobenzyl protecting group proceeds more efficiently in slightly acidic (pH 3.5) 0.1 mol dm−3 ammonium formate solution (Hayes et al., 1985). A serious drawback to the use of the 2-nitrobenzyl protecting group is that the photolytic cleavage reaction does not always proceed quantitatively (Ohtsuka and Iwai, 1987), especially in the unblocking of relatively highmolecular-weight RNA sequences. Takaku and co-workers have used the 4methoxybenzyl (as in S.35a; Takaku and Kamaike, 1982; Takaku et al., 1984) and 3,4dimethoxybenzyl (S.35b; Takaku et al., 1986) groups to protect 2′-hydroxy functions in solution-phase oligoribonucleotide synthesis. The 4-methoxybenzyl protecting groups were removed from a hexaribonucleoside pentaphosphate (Takaku et al., 1984) by treatment for 3 hr at room temperature with a reagent prepared by adding triphenylmethyl tetrafluoroborate (∼0.10 mmol/mL) to acetonitrile/water (4:1 v/v). However, Takaku et al. (1986) reported that incomplete unblocking and some cleavage of the glycosidic linkages can occur under these presumably rather acidic conditions. Some cleavage and migration of the internucleotide linkages might also be expected to occur. The 3,4-dimethoxybenzyl protecting group may be
removed (Takaku et al., 1986) under somewhat milder conditions by treatment with 2,3-dichloro-5,6-dicyano-1,4-benzoquinone (DDQ) in wet dichloromethane; this group would appear to be potentially more promising than the 4-methoxybenzyl group for the protection of the 2′-hydroxy functions in oligoribonucleotide synthesis.
The tert-Butyldimethylsilyl Protecting Group The tert-butyldimethylsilyl group (TBDMS; as in S.37 and S.38; Fig. 2.2.8) was originally suggested by Stork and Hudrlik (1968) for the protection of enols, and was first used by Corey and Venkateswarlu (1972) for the protection of alcoholic hydroxy functions. Ogilvie and coworkers (Ogilvie et al., 1974) then introduced it as a protecting group for the 2′-hydroxy functions of ribonucleoside building blocks. The TBDMS group is at present the most widely used 2′-protecting group in solid-phase oligoribonucleotide synthesis (Damha and Ogilvie, 1993). It meets some but by no means all of the above general requirements for protecting groups. It may be readily introduced (Usman et al., 1987), for example, by treating a 5′-O-DMTr-ribonucleoside derivative (S.36) with tert-butylchlorodimethylsilane and imidazole in N,N-dimethylformamide (DMF) solution (Fig.2.2.8). Although the regiochemistry of the silylation reaction can be controlled to some extent (Hakimelahi et al., 1982), a mixture of 2′- and 3′-isomers (S.37 and S.38, respectively) is in-
DMTrO DMTrO
O
B
HO
Protection of 2′Hydroxy Functions of Ribonucleosides
36
Figure 2.2.8 group.
OH
DMTrO
O 3'
2'
Me2Si(t-Bu)Cl imidazole
B
O
HO O Me Si Me t-Bu 37
B
OH O Me Si Me t-Bu 38
Scheme showing introduction of the tert-butyldimethylsilyl (TBDMS) protecting
2.2.6 Current Protocols in Nucleic Acid Chemistry
AcO
O
Ade
AcO
O
Ade
OH O Me Si Me t-Bu
HO
O Me Si Me t-Bu 39
40 Ade = adenin-9-yl
Figure 2.2.9 Scheme showing the interconversion of the 2′-O - (S.39) and 3′-O - (S.40) TBDMS adenosine derivatives.
variably obtained. Fortunately, such isomeric mixtures can usually be separated by chromatography on silica gel. However, great care has to be taken in the purification and isolation of the 2′-protected ribonucleoside building blocks (S.37), as the TBDMS group readily migrates from the 2′- to the 3′-hydroxy function and vice versa. Interconversion between the adenosine derivatives S.39 and S.40 (Fig. 2.2.9) was found to be a base-catalyzed first-order equilibration reaction (Jones and Reese, 1979). Equilibration rates were observed to be the same in both directions, and the equilibrium constant was estimated to be 1.0. The half time (t1/2) for equilibration in anhydrous pyridine solution at 36°C was 19 hr. The equilibration rate was increased by a factor of 3.0 when 0.1 mol equiv. (with respect to substrate) of benzylamine (pKa 9.34) was added. Equilibration was faster still (t1/2 = ∼1 hr at 36°C) in methanol-d4 solution without added base. When 0.1 mol equiv. of triethylamine (pKa 10.87) was added to the methanol-d4 solution at 20°C, equilibration was complete within ∼5 min. Precautions must be taken to avoid migration of the TBDMS protecting group during the purification and isolation of 2′-O-TBDMS-5′O-DMTr-ribonucleoside derivatives (S.37) and during the course of their conversion into the required monomeric building blocks. Otherwise, the resulting synthetic RNA sequences will be contaminated with material containing
DMTrO
O HO
(2′→5′)-internucleotide linkages. Thus, in the preparation of 3′-phosphoramidite building blocks (S.41; Fig. 2.2.10), it is advisable that the presence of a strong base such as diisopropylethylamine be avoided. Usman and coworkers (Scaringe et al., 1990) have recommended that a mixture of 2,4,6-collidine and 1-methylimidazole be used. Although the presence of contaminating isomeric 2′-phosphoramidites (S.42) can be detected above a certain level by 31P NMR spectroscopy, these impurities cannot readily be removed. It was recently reported that the oligoribonucleotide r[(Up)20U], prepared by treating its per-2′-O-TBDMS derivative with tetra-n-butylammonium fluoride, contained an average of 1.3% (2′→5′)-internucleotide linkages (Morgan et. al., 1995). As acid was not used either during or after the unblocking process, a reasonable explanation for this observation is that the phosphoramidite S.41 (B = uracil-1-yl) used in its synthesis was contaminated with 1.3% of its 2′-isomer (S.42; B = uracil-1-yl). Using an analytical procedure similar to that described by Morgan et al., it was later concluded that some commercially supplied 2′-OTBDMS-protected ribonucleoside phosphoramidites (S.41) were contaminated with comparable amounts (i.e., >1%) of isomeric 2′-phosphoramidites (S.42; Reese et al., unpub. observ.). However, some other batches of commercially supplied material were estimated to contain smaller quantities of the corresponding
DMTrO
B
OTBDMS
NC
B
O
i O
OTBDMS
B
O
TBDMSO
O
O P N(i-Pr)2
37
DMTrO
41
P O
CN
(i-Pr)2N 42
Figure 2.2.10 Scheme showing conversion of 2′-O -TBDMS-5′-O -DMTr-ribonucleoside into its corresponding 3′-phosphoramidite (S.41) and the structure of the possibly contaminating isomeric 2′-phosphoramidite (S.42). Reagents (i): NCCH2CH2OPN(i-Pr)2Cl, base.
Protection of Nucleosides for Oligonucleotide Synthesis
2.2.7 Current Protocols in Nucleic Acid Chemistry
2′-isomer (S.42; Reese et al., unpub. observ.). It would therefore appear that migration of the TBDMS group can to a large extent be controlled if careful manufacturing protocols are observed. The use of the TBDMS protecting group in the solid-phase synthesis of RNA sequences can lead to long coupling times and unsatisfactory coupling efficiencies, possibly due to the bulk of this protecting group. However, the use of 5-ethylthio-1H-tetrazole (S.43b; Fig. 2.2.11) instead of 1H-tetrazole (S.43a) as the phosphoramidite activator can result in shorter coupling times and higher quality products (Sproat et al., 1995; UNIT 3.5). One important advantage of the TBDMS protecting group is that it appears to be stable under the acidic conditions used to remove the 5′-terminal DMTr protecting group in solid-phase synthesis. Problems have arisen in the unblocking of 2′-O-TBDMS-protected RNA sequences. The standard unblocking procedure used in the solid-phase synthesis of DNA sequences involves heating the fully loaded solid support with concentrated aqueous ammonia at 55°C overnight (Brown and Brown, 1991). These ammonolytic conditions lead to the release of the oligodeoxyribonucleotides from the solid support, the removal of the 2-cyanoethyl protecting groups from the internucleotide linkages, and the removal of N-acyl protecting groups from the base residues (in oligodeoxyribonucleotide synthesis, the adenine and cytosine residues are usually protected with benzoyl groups as in S.16 and S.18, R = Ph, and the guanine residues are usually protected with isobutyryl groups as in S.19, R = Me2CH). If these ammonolysis conditions are also used in the unblocking of 2′-O-TBDMS-protected oligoribonucleotides, appreciable loss of the 2′O-TBDMS protecting groups and concomitant cleavage of the internucleotide linkages are likely to occur (Stawinski et al., 1988). This problem of premature removal of 2′-OTBDMS protecting groups has been largely
N N N N H 43a, R = H 43b, R = SEt R
Protection of 2′Hydroxy Functions of Ribonucleosides
Et3N • 3HF
44
Figure 2.2.11 Phosphoramidite activators 5ethylthio-1H-tetrazole (S.43b) and 1H-tetrazole (S.43a), and the unblocking reagent triethylamine trihydrofluoride (S.44).
overcome by protecting the base residues with more labile acyl groups (Chaix et al., 1989), and by replacing concentrated aqueous ammonia with a more selective reagent such as 35% aqueous ammonia/ethanol (3:1 v/v) ammonia/ethanol (Mullah and Andrus, 1996), anhydrous ethanolic ammonia (Goodwin et al., 1994), or aqueous methylamine (Wincott et al., 1995). Téoule and co-workers (Chaix et al., 1989) have recommended that adenine and guanine residues be protected with phenoxyacetyl groups (as in S.16 and S.19, R = CH2OPh) and that cytosine residues be acetylated (as in S.18, R = Me). These workers found that the half times for removal of the latter protecting groups in aqueous ammonia/ethanol (1:1 v/v) at room temperature ranged from 10 to 15 min. It should therefore be possible completely to unblock base residues that are protected in this way without any significant loss of the 2′-O-TBDMS protecting groups and without internucleotide cleavage. In the final unblocking step, the TBDMS protecting groups are removed from the 2′-hydroxy functions of the synthetic RNA sequences. Until recently, a solution of tetra-nbutylammonium fluoride in tetrahydrofuran (Damha and Ogilvie, 1993) was almost always used for this unblocking process. However, it has been reported that the use of the latter reagent results in an inconvenient work-up procedure and can lead to incomplete unblocking (Sproat et al., 1995). More recently, it has been suggested that triethylamine trihydrofluoride (S.44; Fig. 2.2.11; Gasparutto et al., 1992; Westman and Strömberg, 1994) is a more suitable reagent for this purpose. Both the neat reagent S.44, which is slightly acidic as evidenced by the concomitant loss of 5′-O-DMTr protecting groups (Mullah and Andrus, 1996), and a solution of S.44 and triethylamine in 1-methylpyrrolidone (Wincott et al., 1995) have been used. There now seems to be little doubt that, if the above precautions are taken and the most suitable base-protecting groups and reagents are used, the TBDMS group may be used effectively for the protection of the 2′-hydroxy functions in the solid-phase synthesis of RNA sequences. 2′-O-TBDMS-ribonucleoside 3′H-phosphonate building blocks (S.45; Fig. 2.2.12) have also been used successfully in the solid-phase synthesis of oligoribonucleotides (Rozners et al., 1994). It is reasonable to assume that the same precautions and considerations that apply to solid-phase synthesis based on phosphoramidite building blocks (S.41) should
2.2.8 Current Protocols in Nucleic Acid Chemistry
DMTrO
sequences containing one or more (2′→5′)-internucleotide linkages.
B
O
O OTBDMS O P O H 45
Figure 2.2.12 2′-O -TBDMS-ribonucleoside 3′-H-phosphonate.
be taken into account if good quality RNA sequences are to be obtained from the corresponding H-phosphonates (S.45).
Acetal Protecting Groups In general, acetal groups have several distinct advantages over the TBDMS group as far as the protection of the 2′-hydroxy functions in oligoribonucleotide synthesis is concerned. First, acetal protecting groups can usually be placed regiospecifically on the 2′-hydroxy functions (see below) and, once in position, they cannot migrate. Secondly, they are completely stable under the basic conditions that normally obtain during the unblocking of internucleotide linkages and base residues. Thirdly, the 2′-protected RNA sequences obtained after the removal of the other protecting groups (Rao et al., 1993) can be purified under neutral or basic conditions without any danger of endonuclease-promoted digestion. However, there is one important drawback to the use of acetal protecting groups in that they are generally removed by acid-catalyzed hydrolysis. Unless the acidic conditions used are particularly mild, both cleavage and migration of the internucleotide linkages can occur (Griffin et al., 1968; Capaldi and Reese, 1994; Fig. 2.2.2B). While cleavage of internucleotide linkages is clearly highly undesirable, migration is a very much more serious matter as it is virtually impossible to free even a relatively low-molecular-weight RNA sequence from contaminating isomeric
AcO
O
B
HO
B
O
i, ii OH
AcO
The tetrahydropyran-2-yl (Thp) group In the 1960s, the use of the 2′-O-tetrahydropyran-2-yl protecting group (Thp) in oligoribonucleotide synthesis was examined (Smith et al., 1962; Smrt and Šorm, 1962; Griffin and Reese, 1964). Pure 2′-O-Thp derivatives of uridine and adenosine (S.47a and S.47b, respectively) were prepared according to the procedure indicated in Figure 2.2.13, and were converted into dinucleoside phosphates by the methods then available (Griffin and Reese, 1964; Griffin et al., 1968). Careful unblocking studies were carried out in 0.01 mol dm−3 hydrochloric acid (pH 2.0) at 24°C (Griffin et al., 1968), and the half time (t1/2) for the conversion of 2′-O-Thp-UpU (S.49; Fig. 2.2.14) into completely unprotected uridylyl(3′→5′)-uridine (UpU; S.50) was found to be 29 min. It can therefore be estimated that >99.9% removal of the Thp group would occur in <5 hr under these conditions. It was also found that after UpU (S.50) had been allowed to stand in 0.01 mol dm−3 hydrochloric acid (pH 2.0) at 25°C for 216 hr, it underwent ∼99% ribonuclease A–catalyzed digestion to uridine-3′-phosphate and uridine, thereby indicating that not more than 1% isomerization to uridylyl-(2′→5′)-uridine (S.51) had occurred (Griffin et al., 1968). Under these conditions (i.e., pH 2.0, 25°C, 216 hr), UpU also underwent ∼0.5% hydrolytic cleavage (Fig. 2.2.2B). It can therefore be estimated that in the time required for >99.9% removal of the Thp protecting group from 2′-O-ThpUpU, not more than 0.02% phosphoryl migration and 0.01% internucleotide cleavage would be expected to occur. Thus, it seemed reasonable to conclude from the data then available that the Thp group was suitable for the protection of the 2′-hydroxy functions in oligoribonucleotide synthesis.
HO
O
O
O (Thp)
46
47
48
a, B = uracil-1-yl b, B = adenin-9-yl
Figure 2.2.13 Scheme showing preparation of 2′-O -Thp derivatives of uridine and adenosine. Reagents: (i) 3,4-dihydro-2H-pyran (S.48), toluene-4-sulfonic acid (TsOH), dioxane; (ii) NaOMe, MeOH.
Protection of Nucleosides for Oligonucleotide Synthesis
2.2.9 Current Protocols in Nucleic Acid Chemistry
HO
O
Ura
HO
O OThp O P O O
Ura
HO
O
O HO
OH
49
Ura
O HO
O OH O P O
H3O Ura
O HO
O
Ura
O O P O O
OH
HO
O
Ura
OH
51
50
Ura = uracil-1-yl
Figure 2.2.14 Scheme showing conversion of 2′-O -Thp-UpU into unprotected uridylyl-(3′→5′)uridine (S.50) and the structure of its (2′→5′)-isomer (S.51).
A particular disadvantage of the Thp group is that it is chiral, and therefore its use in the protection of ribonucleoside derivatives and other chiral compounds leads to mixtures of diastereoisomers. Thus, two diastereoisomers each of 2′-O-Thp-uridine (S.47a) and 2′-OThp-adenosine (S.47b) were obtained (Fig. 2.2.13; Griffin et al., 1968). Although both pairs of diastereoisomers were easily separable and all four compounds were obtained as pure crystalline solids, this is clearly an undesirable complication. The 4-methoxytetrahydropyran-4-yl (Mthp) group A search for an achiral alternative to the Thp protecting group led to the introduction of the 4-methoxytetrahydropyran-4-yl group (Mthp; Reese et al., 1967; 1970). 2′-O-Mthp derivatives of ribonucleosides (S.54) were first prepared from 3′,5′-di-O-acyl-ribonucleosides (S.53; Fig. 2.2.15). However, they are more
R
conveniently prepared from the corresponding 3′,5′-O-(1,1,3,3-tetraisopropyl- disiloxan-1,3diyl) derivatives (S.56; Brown et al., 1989a). 2′-O-Mthp derivatives (S.54) are usually obtained as pure crystalline solids in satisfactory to good yields (Reese et al., 1970). The half times for the hydrolysis of 2′-OMthp-uridine and 2′-O-Mthp-adenosine (S.54, B = uracil-1-yl and adenosine-9-yl, respectively) in 0.01 mol dm−3 hydrochloric acid at 22°C were found to be 18.7 and 34 min, respectively (Norman et al., 1984). It is interesting to note that the removal of the Mthp protecting group from 2′-O-Mthp-uridylyl-(3′→5′)-uridine (S.57a) and 2′-O-Mthp-adenylyl-(3′→5′)adenosine (S.57b; Fig. 2.2.16) under the same conditions was found to proceed at significantly faster rates (t1/2 = 6.1 and 19.9 min, respectively; Norman et al., 1984). The rate of removal of the Thp protecting group from 2′O-Thp-UpU (S.49) is also faster than from 2′-O-Thp-uridine (S.47a; Griffin et al., 1968).
B
O
O
O R'
O
O HO
OH
i, ii
O 53
B
HO
OH
B
HO
HO
iii
OMe
O O
OMe O
i, iv
52
O (i-Pr)2Si O (i-Pr)2Si O
B
O
55
O 54
(Mthp)
OH
56
Protection of 2′Hydroxy Functions of Ribonucleosides
Figure 2.2.15 Scheme showing preparation of 2′-O -Mthp ribonucleoside derivatives (S.54) via 3′,5′-di-O-acyl-ribonucleosides (S.53) or 3′,5′-O-(1,1,3,3-tetraisopropyldisiloxan-1,3-diyl) derivatives (S.56). Reagents: (i) 4-methoxy-5,6-dihydro-2 H-pyran (S.55), toluene-4-sulfonic acid (TsOH), dioxane; (ii) NH3, MeOH; (iii) (i-Pr)2Si(Cl)OSi(Cl)(i-Pr)2, imidazole, MeCN; (iv) Et4NF, MeCN.
2.2.10 Current Protocols in Nucleic Acid Chemistry
HO
O
B
HO
O
O OMthp O P O O
O HO
B
O
OH
O P O
H3O B
O
OH
O HO
57
B
OH
58
a, B = uracil-1-yl b, B = adenin-9-yl
Figure 2.2.16
Scheme illustrating the removal of the Mthp protecting group.
The fact that the presence of a vicinal phosphodiester internucleotide linkage appears to facilitate the acid-catalyzed unblocking of a 2′-O-Mthp- or 2′-O-Thp-protected hydroxy function is clearly advantageous if migration and cleavage of the internucleotide linkages (Fig. 2.2.2B) in the final unblocking step of oligoribonucleotide synthesis are to be kept to a minimum. As well as being achiral, Mthp has an additional advantage over Thp in that it is more labile to acidic hydrolysis. The Mthp and Thp protecting groups have been used in both solution- and solid-phase synthesis of RNA sequences. The Mthp group was introduced particularly for solution-phase synthesis, and it has been used successfully in the preparation of the 3′-terminal decamer, nonadecamer, and heptatriacontamer (37-mer) sequences (r[UpCpGpUpCpCpApCpCpA], r[ApUpUpCpCpGpGpApCpUpCpGpUpCpCpApCpCpA], and r[GpGpApGpApGpGpUpCpUpCpCp GpGpTpψpCpGpApUpUpCpCpGpGpApCpUpCpGpUpCpCpApCpCpA], respectively) of yeast alanine transfer RNA (tRNAAla; Jones et al., 1980, 1983; Brown et al., 1989a,b). This work has already been reviewed (Reese, 1989). The tetrahydrofuran-2-yl (Thf) and 1,5dimethoxycarbonyl-3-methoxypentan-3-yl (Mdmp) groups The above approach to the solution-phase synthesis of RNA sequences was successful largely because treatment with acid was completely avoided until the final unblocking step. However, other workers (Ohtsuka et al., 1984) reported a solution-phase block synthesis of a tritriacontamer (33-mer) sequence of E. coli tRNA2Gly using tetrahydrofuran-2-yl (Thf; S.59; Fig. 2.2.17) and DMTr groups for the protection of the 2′- and 5′-hydroxy functions, respectively. The 5′-terminal DMTr protecting groups were removed by treatment with zinc
bromide in dry dichloromethane/isopropanol solution rather than with a protic acid. Although Thf is more labile than Thp (and probably also Mthp) to acid-catalyzed hydrolysis (Kruse et al., 1979), the latter combination of protecting groups was apparently effective. There are a number of reports in the literature relating to solid-phase RNA synthesis in which the 2′-hydroxy functions are protected by Thp, Mthp, or Thf groups and the 5′-hydroxy functions are also protected with acid-labile groups (Tanaka et al., 1986; Kierzek et al., 1986; Iwai et al., 1987; Tanimura et al., 1989; Tanimura and Imada, 1990). Such acid-labile groups include DMTr (S.10b), 9-phenylxanthen-9-yl (S.11), and 9-(4-methoxyphenyl)xanthen-9-yl (S.60; Fig. 2.2.17; UNIT 2.3). Although some sequences appear to have been prepared successfully in this way, other reports suggest that this is an unsound strategy, particularly for the synthesis of comparatively high-molecular-weight RNA sequences (Reese and Skone, 1985; Christodoulou et al., 1986; Kierzek, 1994). Even when precautions are taken to maintain stringently anhydrous conditions, the repeated exposure of the growing protected oligoribonucleotide to di- or tri-chloroacetic acid in order to remove the 5′-protecting group in each synthetic cycle is likely to lead to some loss of such relatively labile 2′-
MeO
O
O
59
60
Figure 2.2.17 Tetrahydrofuran-2-yl (S.59) and 9-(4-methoxyphenyl)xanthen-9-yl (S.60) protecting groups.
Protection of Nucleosides for Oligonucleotide Synthesis
2.2.11 Current Protocols in Nucleic Acid Chemistry
O O (Lev)
B
O
HO
B
O
O NC
O
OThf
i
NC
O P O O
B
O O
O O
(Fmoc) NC
O
OThf
O
61
O
OThf
O P O B
OThf
62
O
B
O
HO
O
B
O
ii O
OMthp
NC
O P O O
O O
B
OMthp
63
O
OMthp
O P O O
O O
B
OMthp
64
Figure 2.2.18 Scheme showing removal of levulinyl (top) and Fmoc (bottom) protecting groups. Reagents: (i) N2H4⋅H2O, C5H5N, AcOH; (ii) 0.1 M 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU), MeCN.
Protection of 2′Hydroxy Functions of Ribonucleosides
protecting groups, resulting both in cleavage and migration of internucleotide linkages (Pathak and Chattopadhyaya, 1985; Reese and Skone, 1985; Kierzek, 1994). If Thp, Mthp, or Thf groups are to be used to protect the 2′-hydroxy functions, it would appear to be a better strategy in solid-phase synthesis to protect the 5′-terminal hydroxy function with a group that is readily removable under virtually neutral or mildly basic conditions. Thus, following van Boom’s use of the levulinyl group (as in S.61; Fig. 2.2.18) for the protection of the 5′-hydroxy functions in solution-phase synthesis (den Hartog et al., 1981), other workers (Iwai and Ohtsuka, 1988) successfully used Lev in conjunction with Thf in solid-phase oligoribonucleotide synthesis. The Lev group was removed (Fig. 2.2.18) in the usual way (den Hartog et al., 1981) by treatment with hydrazine hydrate in pyridine/acetic acid solution. A number of RNA sequences, including a heneicosamer (21mer), were prepared in this way. In another study (Lehmann et al., 1989), the Fmoc group (S.12) was used to protect the 5′-hydroxy functions in solid-phase oligoribonucleotide synthesis. These workers protected the 2′-hydroxy functions with Mthp groups (as in S.63) and removed the Fmoc group with 1,8- diazabicyclo[5.4.0]undec-7-ene (DBU) in acetonitrile solution (Fig. 2.2.18). As the authors (Lehmann
et al., 1989) pointed out, it is very likely that some concomitant loss of the 2-cyanoethyl protecting groups from the partially protected oligoribonucleotides (S.63) occurs during DBU treatment. A nonadecamer and an icosamer (20-mer) RNA sequence were prepared successfully in this way. Although the above approach using either 2′-O-Thf and 5′-O-Lev protection or 2′-OMthp and 5′-O-Fmoc protection (Fig. 2.2.18) was successful, it is much more convenient to use the acid-labile DMTr (S.10b) or Px (S.11) groups to protect the 5′-hydroxy functions in solid-phase RNA synthesis. The latter modified trityl groups can be rapidly and quantitatively removed under anhydrous conditions and the released carbocations can easily be assayed spectrophotometrically (Brown and Brown, 1991). For this reason, attempts have been made to develop somewhat more sophisticated acetal protecting groups that are stable under normal “detritylation” conditions and are also sufficiently labile to acidic hydrolysis in the final unblocking step for cleavage and migration of the internucleotide linkages (Fig. 2.2.2B) to be avoided. Chattopadhyaya and co-workers (Sandström et al., 1985) showed that the 1,5-dimethoxycarbonyl-3-methoxypentan-3-yl (Mdmp) group (as in S.65; Fig. 2.2.19), derived from dimethyl 4-ketopimelate, was
2.2.12 Current Protocols in Nucleic Acid Chemistry
HO
B
O HO
HO
OMe
O
CO2Me
concentrated aqueous NH3
B
O HO
O
OMe
CO2Me
CONH 2 CONH 2
(Mdmp) 66
65
X
O
OMe O
O
B
O
Thy O
O
OMe
HO 67a, X = O 67b, X = S 67c, X = SO2
Figure 2.2.19
O
B
O
H O
O
N
NAr 68
OMe H Ar
69
Thy = thymin-1-yl Ar = aryl
Acetal protecting groups labile to acidic hydrolysis.
converted under the standard ammonolytic unblocking conditions used in solid-phase synthesis into the corresponding bis-amide (S.66), which was seventeen times more labile to acidic hydrolysis than the bis-ester (S.65). However, the Mdmp protecting group itself is unlikely to find application in solid-phase synthesis of oligoribonucleotides as it undergoes hydrolysis in 4:1 (v/v) acetic acid/water solution even more rapidly than the Mthp group.
The 1-Aryl-4-methoxypiperidin-4-yl (Ctmp and Fpmp) groups Acetal hydrolysis is a second-order reaction; its rate, which is proportional to the concentrations both of substrate and hydrogen ions, is very sensitive to inductive effects (Kreevoy and Taft, 1955). Thus 5′-O-(4methoxytetrahydrothiopyran-4-yl)-thymidine (S.67b; Fig. 2.2.19) was found to be ∼5 times more labile to acidic hydrolysis than the corresponding Mthp derivative (S.67a) and was estimated to be >2000 times more labile than the corresponding sulfone (S.67c; van Boom et al., 1972). It seemed possible that, if the aryl substituent Ar were selected carefully, a 1-aryl-4methoxypiperidin-4-yl protecting group (as in S.68) could be identified that would be almost fully protonated (as in S.69) under detritylation conditions (i.e., in dichloromethane containing, for instance, 2% to 3% trichloroacetic
acid), but would be virtually unprotonated (as in S.68) under the milder conditions of acidic hydrolysis obtaining in the final unblocking step of oligoribonucleotide synthesis. Although it would, of course, depend on the aryl substituent, it seemed possible that the rate of hydrolysis of the unprotonated and protonated piperidinyl species might correspond approximately to those of the Mthp (S.67a) and sulfone (S.67c) derivatives, respectively. In the overall rate expression for the hydrolysis of a 1-aryl4-methoxypiperidin-4-yl derivative, it is reasonable to assume that the component relating to the hydrolysis of the conjugate acid S.69 is likely to be negligible in comparison with that relating to the unprotonated S.68 and that, as a first approximation, it can be ignored. If this is the case, the observed rate of hydrolysis of the 1-aryl-4-methoxypiperidin-4-yl acetal system should be pH independent. For example, if the pH of the hydrolytic medium is lowered by one unit, the concentration of the unprotonated acetal S.68 will decrease by an order of magnitude, and at the same time the rate of hydrolysis of the remaining unprotonated acetal will increase by an order of magnitude. Despite the very limited synthetic methodology available at the outset, it was possible to prepare 2′-O-[1-(2-chloro-4-methylphenyl)-4methoxypiperidin-4-yl] (Ctmp) ribonucleoside derivatives (S.71a; Fig. 2.2.20A) and
Protection of Nucleosides for Oligonucleotide Synthesis
2.2.13 Current Protocols in Nucleic Acid Chemistry
A B
O
HO
O
(i-Pr)2Si O (i-Pr)2Si O
OH
B
O
i, ii HO
O
OMe R
1
N
56
2
R 71
B O Cl
O
iii Cl
Cl
Cl
72
MeO
OMe
OMe
73 iv
N
v R
N
1
1
R
NH2 R
1 2
R
2
2
R
R
75
70
74 1
2
a, R = Cl, R = Me 1 2 b, R = F, R = H
Figure 2.2.20 (A) Scheme showing preparation of Ctmp (S.71a) and Fpmp (S.71a) ribonucleoside derivatives. (B) Preparation of the 1-aryl-4-methoxy-1,2,5,6-tetrahydropyridines (S.70) required in (A). Reagents (i) S.70, CF3CO2H, CH2Cl2; (ii) Et4NF, MeCN; (iii) ethylene, AlCl3, CH2Cl2; (iv) toluene-4-sulfonic acid monohydrate (TsOH⋅H2O), MeOH, and reflux followed by (MeO)3CH; (v) (i-Pr)2NEt, Et2O→BF3, CH2Cl2, 0°C.
Protection of 2′Hydroxy Functions of Ribonucleosides
show that the Ctmp protecting group had the desired properties (Reese et al., 1986). Thus it can be seen from Figure 2.2.21 that the rate of hydrolysis of 2′-O-Ctmp-uridine (S.71a, B = uracil-1-yl) at 30°C is only 1.75 times faster at pH 0.5 than it is at pH 2.5. At 25°C, 2′-O-Ctmpuridine is ∼40 times more stable than 2′-OMthp-uridine (S.54, B = uracil-1-yl) at pH 1.0 (Reese et al., 1986), but it is nearly 1.6 times more labile than 2′-O-Mthp-uridine at pH 3.0. The first general criterion that all protecting groups should meet (see above) is that they should be easy to introduce, and an important part of this criterion is that the reagent required should be readily accessible. The Ctmp and related piperidine-derived protecting groups are easy to introduce, but until recently the preparation of the enol ether reagents (such as S.70) involved a number of steps. However, these 1-aryl-4-methoxy-1,2,5,6-tetrahydropyridine derivatives can now be readily prepared (Fig. 2.2.20B) in two steps and in good overall yields (Faja et al., 1997) from 1,5-dichloropentan-3-one (S.73; Owen and Reese, 1970) and the appropriate primary aromatic amine (S.74). The procedure for the preparation of 2′-O-(1-aryl-4-methoxypiperidin-4-yl) ribonucleoside derivatives (such as S.71; Fig.
Figure 2.2.21 Dependence of half times (t1/2) on pH for hydrolysis of 2′-O -Ctmp-uridine (S.71a) and 2′-O -Fpmp-uridine (S.71b) at 30°C.
2.2.14 Current Protocols in Nucleic Acid Chemistry
PxO
O
NC
B
O
DMTrO
OCtmp
N(i-Pr)2
77
Ctmp-protected phosphoramidite (S.76) and H-phosphonate (S.77).
2.2.20A; Rao et al., 1987, 1993) is closely similar to that used in the preparation of the corresponding 2′-O-Mthp derivatives (S.54; Fig. 2.2.15), except that a much smaller excess of the enol ether reagent S.70 is needed. The 2′-O-Ctmp protecting group was used in conjunction with the 5′-O-Px protecting group (Rao et al., 1987) or the 5′-O-DMTr protecting group (Sakatsume et al., 1989) in the solid-phase synthesis of oligoribonucleotides. Phosphoramidite building blocks (S.76; Fig. 2.2.22) were successfully used in the preparation of the 3′-terminal nonadecamer sequence r[ApUpUpCpCpGpGpApCpUp CpGpUpCpCpApCpCpA] of yeast tRNA Ala (Rao et al., 1987), and H-phosphonate building blocks (S.77) were used successfully in the preparation of the octadecamer sequence, r[ApGpUpApUpApApGpApGpGpApCpApUp ApUpG] (Sakatsume et al., 1989). However, the required enol ether reagent (S.70) was difficult to prepare by the original procedure (Reese et al., 1986), and its preparation by the improved protocol (Fig. 2.2.20B; Faja et al., 1997) involves either the use of an expensive aromatic amine (S.74a) or an additional chlorination step. It was later found that several other 1-aryl4-methoxypiperidin-4-yl groups were also suitable for the protection of the 2′-hydroxy functions in solid-phase oligoribonucleotide synthesis. Among these is the 1-(2fluorophenyl)-4-methoxypiperidin-4-yl (Fpmp) protecting group (as in S.71b; Reese and Thompson, 1988). The Fpmp protecting group has two distinct advantages over the Ctmp group. First, the enol ether reagent S.70b,
PxO
NC
B
OCtmp O O P O Et3NH H
O P 76
Figure 2.2.22
O
B
O O
OFpmp
O P
which is a low-melting solid, is readily prepared (Faja et al., 1997) from 2-fluoroaniline (S.74b), which is an inexpensive starting material. Secondly, the Fpmp group is somewhat more stable than the Ctmp group to acidic hydrolysis in the pH range of 0.5 to 1.0 (∼1.4 times at 30°C; Fig. 2.2.21), and therefore the risk of concomitant 2′-unblocking in the detritylation steps is even smaller. However, removal of the 2′-O-Fpmp protecting group occurs more slowly than removal of the 2′-OCtmp group in the final unblocking step of oligoribonucleotide synthesis, and this can be disadvantageous (see below). It can be seen from Figure 2.2.21 that at 30°C the rate of hydrolysis of 2′-O-Fpmp-uridine (S.71b; B = uracil-1-yl) is only about twice as fast at pH 0.5 as at pH 2.5. The 2′-O-Fpmp protecting group has been widely used in solid-phase oligoribonucleotide synthesis (Beijer et al., 1990; Rao et al., 1993; Capaldi and Reese, 1994; Pieles et al., 1994; Sproat et al., 1994; Rao and Macfarlane, 1995; McGregor et al., 1996). The 5′-O-Px2′-O-Fpmp phosphoramidite building blocks S.78 (Fig. 2.2.23) were used successfully in the synthesis of r[UpCpGpUpCpCpApCpCpA], r[ApUpUpCpCpGpGpApUpCpGpUpCpCp ApCpCpA], and r[GpGpApGpApGpGpUp CpUpCp CpGpGpUpUpCpGpApUpUpCpCpGp GpApCpUpCpGpUpCpCpApCpCpA], the 3′-decamer, nonadecamer, and heptatriacontamer (37-mer) sequences, respectively, of unmodified yeast tRNAAla (Rao et al., 1993). Sproat and co-workers (Pieles et al., 1994) carried out the solid-phase synthesis of some modified oligoribonucleotides containing
DMTrO
NC
O
OFpmp
O P
DMTrO
NC
79
B
O O
OMe
O P
Ni-Pr2
Ni-Pr2 78
B
O
Ni-Pr2 80
Figure 2.2.23 5′-O -Px-2′-O -Fpmp phosphoramidite (S.78), 5′-O -DMTr-2′-O -Fpmp phosphoramidite (S.79), and 5′-O -DMTr-2′-O -methyl-ribonucleoside phosphoramidite (S.80).
Protection of Nucleosides for Oligonucleotide Synthesis
2.2.15 Current Protocols in Nucleic Acid Chemistry
O
B
O
O O
O
B
O
O
CO2R
O P O
R
O
1
O
O
O
O P O O R 1
RO2C
2
2
81a, R = NO2, R = H 1 2 81b, R = H, R = NO2
82a, R = Me 82b, R = H
Figure 2.2.24 Additional acetal protecting groups: (2-nitrobenzyloxy)methyl (S.81a), (4-nitrobenzyloxy)methyl (S.81b), (2,6-dimethoxycarbonyl)phenoxymethyl (S.82a), and (2,6-dicarboxy)phenoxymethyl (S.82b).
Protection of 2′Hydroxy Functions of Ribonucleosides
pseudouridine, 2′-O-methylpseudouridine, and some other 2′-O-methyl-ribonucleoside residues starting from the appropriate 5′-ODMTr-2′-O-Fpmp and 5′-O-DMTr-2′-Omethyl-ribonucleoside phosphoramidites (S.79 and S.80, respectively). In all of the early work (Rao et al., 1993), the 2′-O-Fpmp and 5′-terminal Px (or DMTr) protecting groups were removed by treatment with 0.01 mol dm−3 hydrochloric acid (pH ∼2) at room temperature. However, it soon became clear that the susceptibility of the internucleotide linkages of oligoribonucleotides to acid-catalyzed cleavage and migration was sequence dependent, and that certain sequences were unstable at pH 2 and room temperature (Capaldi and Reese, 1994). Thus, despite the relative stability of uridylyl(3′→5′)-uridine (S.50; Griffin et al., 1968) at pH 2 and room temperature, r[(Up)9U] and r[(Up)19U] both underwent virtually complete degradation in the course of the removal of the 2′-O-Fpmp protecting groups under the same conditions (Capaldi and Reese, 1994). However, when unblocking was carried out at room temperature above pH 3.0, virtually no internucleotide cleavage or migration could be detected (see Conclusions). Other workers subsequently reported that no cleavage or migration of the internucleotide linkages could be detected after r[(Up)20U] had been allowed to stand at pH 3.25 in 0.5 M sodium acetate buffer solution at room temperature for 96 hr (Rao and Macfarlane, 1995), which is very much more than the time required to remove the 2′-O-Fpmp protecting groups. These workers went on to recommend that 2′-O-Fpmp protecting groups be removed at pH 3.25 and 30°C in 0.5 M sodium acetate buffer solution. They successfully unblocked RNA sequences containing up to 50 nucleoside residues under these conditions, and obtained oligoribonucleotides that
were active as ribozymes and ribozyme substrates. Other acetal groups Three other interesting and potentially useful acetal groups have recently been suggested for the protection of the 2′-hydroxy functions in solid-phase oligoribonucleotide synthesis. Like the 2-nitrobenzyl group (as in S.34, see above), the (2-nitrobenzyloxy)methyl group (as in S.81a; Schwartz et al., 1992; Fig. 2.2.24) is removable photochemically; however, possibly for steric reasons, its use leads to faster and more efficient coupling reactions. The (2-nitrobenzyloxy)methyl protecting group has been used successfully in the solid-phase synthesis of a number of RNA sequences including a dodecamer, a hexadecamer, and a tritriacontamer (33-mer) sequence that are all components of ribozyme structures. The related (4-nitrobenzyloxy)methyl protecting group (as in S.81b; Gough et al., 1996), which has also been used successfully in solid-phase oligoribonucleotide synthesis, is removable by treatment with tetra-n-butylammonium fluoride in THF solution. Finally the (2,6-dimethoxycarbonyl)phenoxymethyl protecting group (as in S.82a; Rastogi and Usher, 1995), which has been used in the solid-phase synthesis of two dinucleoside phosphates, is extremely (over 100 times more than the Fpmp group) stable under standard detritylation conditions. After the assembly of the desired RNA sequences, the two methoxycarbonyl groups are saponified by treatment with aqueous sodium hydroxide, which also releases the product from the solid support and removes base-labile protecting groups. The resulting (2,6-dicarboxy)phenoxymethyl acetal system (as in S.82b) is estimated to be >1300 times more labile to acidic hydrolysis at pH 3.0 than the original (2,6-di-
2.2.16 Current Protocols in Nucleic Acid Chemistry
R2O
HO
R2O
B
O O
O
O
R1
O
OH
R1
83
Figure 2.2.25 derivatives.
B
O
84
Scheme showing interconversion of isomeric 2′- and 3′-O -acyl-ribonucleoside
methoxycarbonyl)phenoxymethyl acetal system; however, it is still ∼2.3 times more stable at pH 3.0 than the Fpmp protecting group.
with 2′,3′-di-O-acetyluridine 5′-phosphate (S.86) by the now obsolete phosphodiester approach in solution to give, after deprotection, guanylyl-(3′→5′)-uridine (S.87; B = guanin-9yl) and cytidylyl-(3′→5′)-uridine (S.87; B = cytosin-1-yl), respectively. Both of the latter dinucleoside phosphates were apparently free from their (2′→5′)-isomers. Two later studies relating to the use of 2′-Oacyl protecting groups in solid-phase oligoribonucleotide synthesis are also of interest. In one study (Kempe et al., 1982), oligoribonucleotides and chimeric RNA:DNA sequences were prepared from 2′-O-benzoyl-protected phosphoramidites (S.88; Fig. 2.2.27). However, as these phosphoramidites (S.88) were contaminated with 1% to 3% of the isomeric 2′-phosphoramidites, the integrity of the internucleotide linkages in the target RNA sequences was to some extent compromised. The other study (Rozners et al., 1992) described the solid-phase synthesis of oligoribonucleotides from 2′-O-(2-chlorobenzoyl)3′-H-phosphonate building blocks (S.89). This is a more promising approach for two reasons. First, it was possible to separate the isomeric
Ester Protecting Groups It has been known for many years that isomeric 2′- and 3′-O-acyl-ribonucleoside derivatives (S.83 and S.84, respectively; Fig. 2.2.25) interconvert under mildly basic conditions, and that the equilibrium mixture eventually obtained is generally somewhat richer in the 3′isomer (Reese and Trentham, 1965). Unlike corresponding mixtures of 2′- and 3′-OTBDMS derivatives (e.g., S.39 and S.40; Fig. 2.2.9), it is usually very difficult or even impossible to separate isomeric mixtures of 2′- and 3′-esters (S.83 and S.84) by standard chromatographic methods. Furthermore, acyl migration can occur during chromatography. For these reasons, 2′-O-acyl protecting groups have only very rarely been used in oligoribonucleotide synthesis. However, in an early study (Fromageot et al., 1968), N2,O2′,O5′-tribenzoylguanosine (S.85a; Fig. 2.2.26) and N4,O2′,O5′triacetylcytidine (S.85b), two pure crystalline compounds, were both successfully coupled
R
O
B
O
O HO
O
HO
O
O
B
R 85 i, ii O HO P O
O
O AcO
O OH O P O O O
Ura
OAc
HO
Ura
OH
87
86 Ura = uracil-1-yl a, B = 2-N-benzoylguanin-9-yl, R = Ph b, B = 4-N-acetylcytosin-1-yl, R = Me
Figure 2.2.26 Scheme showing preparation of guanylyl-(3′→5′)-uridine and cytidylyl-(3′→5′)uridine using a 2′-O -acyl protecting group. Reagents: (i) mesitylene-2-sulfonyl chloride, C5H5N; (ii) MeNH2, EtOH, or NH3, MeOH.
Protection of Nucleosides for Oligonucleotide Synthesis
2.2.17 Current Protocols in Nucleic Acid Chemistry
DMTrO DMTrO
O
B
O
B
O
O O O P O H
OBz
MeO P NMe2
DMTrO
O
O Cl
O
Cl
89
88
B
O O P H
90
Figure 2.2.27 2′-O -Benzoyl-protected 3′-phosphoramidite (S.88), 2′-O -(2-chlorobenzoyl)-protected 3′-H-phosphonate (S.89), and the isomeric 2′-H-phosphonate (S.90).
2′- and 3′-H-phosphonates (S.90 and S.89, respectively) by chromatography on silica gel, and thereby obtain isomerically pure building blocks (S.89). Secondly, after the desired RNA sequences had been assembled, the 2′-protecting groups could be removed by ammonolysis under conditions that were mild enough to avoid cleavage of the internucleotide linkages. A number of RNA sequences of moderate length were prepared successfully by this approach.
CONCLUSIONS At present, the TBDMS group (as in S.37; Fig. 2.2.28) is the most widely used protecting group for 2′-hydroxy functions in solid-phase oligoribonucleotide synthesis. The Fpmp (as in S.71b; Fig. 2.2.28) group is also widely used. Both 2′-O-TBDMS- and 2′-O-Fpmp-protected phosphoramidites (S.41 and S.79, respectively) are commercially available. So far, there is no report in the literature that constitutes a thorough and definitive comparison of these two protecting groups. However, it is worthwhile discussing how they meet the necessary criteria
DMTrO
O
HO
B
B
O HO
for protecting groups in oligoribonucleotide synthesis, and also whether improvements could be made by modifying them. With regard to the introduction of these two protecting groups, the reagents required, namely tert-butylchlorodimethylsilane and 1(2-fluorophenyl)-4-methoxy-1,2,5,6-tetrahydropyridine (S.70b; Fig. 2.2.28) are both readily available. However, as far as the introduction of these protecting groups is concerned, the Fpmp group has the edge over the TBDMS group inasmuch as it can be introduced regiospecifically (Fig. 2.2.20A) and cannot then migrate. As indicated above, great care has to be exercised in the preparation of TBDMS-protected building blocks (S.41) in order to avoid contamination with the isomeric 2′-phosphoramidites (S.42; Fig. 2.2.10), the presence of which will inevitably lead to (2′→5′)-internucleotide linkages in the final product. In solid-phase synthesis involving phosphoramidite building blocks, it seems clear that coupling rates and efficiencies are generally lower when ribonucleoside rather than 2′-deoxyribonucleoside building blocks are used
DMTrO O
O
OMe F
HO O Me Si Me t-Bu
NC
N
O
B
OTBDMS
O P N(i-Pr)2 41
71b
37
OMe DMTrO
NC
O O
B
OFpmp
N F
O P N(i-Pr)2
Protection of 2′Hydroxy Functions of Ribonucleosides
79
70b
Figure 2.2.28 Structures relating to a discussion of the relative merits of the TBDMS and Fpmp protecting groups.
2.2.18 Current Protocols in Nucleic Acid Chemistry
DMTrO
B
O O
HO
O
OFpmp
B
O
OH
O P O
O P O O
B
O O
O
OFpmp
n-2
O HO
O
n-2
OH
O P O
O P O O
B
O
H3O
B
O
HO
OH
91
O
B
OH
92
Figure 2.2.29 Unblocking of 2′-O -Fpmp-protected oligoribonucleotides under mild conditions of acidic hydrolysis.
(Hayakawa et al., 1996). It is not yet clear whether TBDMS-protected or Fpmp-protected phosphoramidites (S.41 or S.79; Fig. 2.2.28) are the more hindered. Although TBDMS ethers (Kawahara et al., 1996) and Fpmp acetals are both susceptible to acid-catalyzed hydrolysis, the available evidence suggests that both groups remain intact under the anhydrous acidic conditions used during the detritylation steps. The Fpmp group has advantages over the TBDMS protecting group in the ammonolytic unblocking step at the end of the synthesis. First, in the Fpmp approach, adenine, cytosine, and guanine base residues are protected with relatively stable acyl groups (as in S.16, R = Me3C, S.18, R = Ph, and S.19, R = PhCH2, respectively; Fig. 2.2.4; Rao et al., 1993). However, it is advisable to use much more labile acyl protecting groups in the TBDMS approach (Chaix et al., 1989). More importantly, as the Fpmp protecting group is completely stable under the ammonolytic conditions, “Fpmp-on” RNA sequences (S.91; Fig. 2.2.29) are obtained (Rao et al., 1993). Such “Fpmp-on” oligoribonucleotides are stable to endonucleases and base, and may be conveniently purified and stored. On treatment with aqueous acid under very mild conditions (see below), they are readily converted into unprotected RNA sequences (S.92). “TBDMS-on” RNA sequences do not appear to have been purified and isolated in this way. The one clear advantage that the TBDMS approach has over the Fpmp approach is that removal of the TBDMS protecting group in the final unblocking step does not normally involve acidic hydrolysis, and therefore cannot lead to migration of the internucleotide linkages. However, such migration in the Fpmp approach can be virtually eliminated by carefully controlling
the unblocking conditions. Hecht and co-workers (Morgan et al., 1995) reported that when S.91 (B = uracil-1-yl, n = 21; Fig.2.2.29) was unblocked in 0.5 mol dm−3 sodium acetate buffer, pH 3.25, at 25°C for 20 hr, analysis of the resulting r[(Up)20U] (S.92; B = uracil-1-yl, n = 21) revealed that an average of 0.40% migration per internucleotide linkage had occurred. However, Reese et al. (unpub. observ.) have found that under somewhat milder unblocking conditions (0.5 mol dm−3 sodium acetate buffer, pH 4.0, at 35°C), unblocking of S.91 (B = uracil-1-yl, n = 20) was complete after 9 hr and no migration of internucleotide linkages could be detected in the resulting r[(Up)19U] (S.92; B = uracil-1-yl, n = 20). As has been suggested before (Capaldi and Reese, 1994), it cannot be concluded from the results obtained by Strömberg and co-workers (Rozners et al., 1994) in connection with the use of 2′-O-Ctmp5′-O-DMTr-uridine 3′-H-phosphonate (S.77; B = uracil-1-yl; Fig. 2.2.22) and the corresponding Fpmp-protected H-phosphonate building block in the synthesis of r[(Up)11U] and r[(Up)11A] that Ctmp and Fpmp are unsuitable protecting groups for the 2′-hydroxy functions in the H-phosphonate approach to the solid-phase RNA synthesis. A much more likely explanation for Strömberg’s observations is that r[(Up)11U] and r[(Up)11A], like r[(Up)9U] and r[(Up)19U] (Capaldi and Reese 1994), are particularly labile at pH 2.0 and room temperature. Although it is clear that the solid-phase synthesis of relatively high-molecular-weight RNA sequences using TBDMS, Fpmp, or other groups to protect the 2′-hydroxy functions is now a feasible proposition, it is likely that even better protecting groups will be identified in the
Protection of Nucleosides for Oligonucleotide Synthesis
2.2.19 Current Protocols in Nucleic Acid Chemistry
future. The next generation of 2′-protecting groups could include modifications of TBDMS, Fpmp, and some of the other groups described above, and it could also include completely different groups. Any alternative silyl protecting group (S.93; Fig. 2.2.30) would need to be bulky to be sufficiently stable, and so far there is no evidence that any such group is likely to have superior properties to those of the TBDMS group itself. However, modification of the Fpmp group could well lead to improvements. A choice is already available between the Fpmp group, which has the advantage of being more stable at low pH (Fig. 2.2.21) during the detritylation steps, and the Ctmp group, which has the advantage of being more labile at high pH during the final unblocking step. It might well be possible, by a careful choice of R1 and R2, to identify a 1-aryl-4-alkoxypiperidin-4-yl protecting group (S.94; Fig. 2.2.30) that is as stable as (or perhaps even more stable than) the Fpmp group at low pH and as labile as (or perhaps even more labile than) the Ctmp group at high pH. Most of the above discussion has been concerned with the small-scale synthesis of RNA sequences on a solid support. In the light of recent developments in the possible use of oligonucleotide analogs in chemotherapy, a demand has arisen for the development of methods for large-scale synthesis. This may well involve a shift from solid-phase to solutionphase methodology. While this need not necessarily affect the strategy of 2′-protection, the cost of the requisite monomeric building blocks is likely to become a matter of crucial importance. Therefore, particular emphasis will need to be laid on the first general criterion for protecting groups–that they should be easy to introduce and that the reagents involved should be readily accessible. It is not envisaged that this will present a problem for the Fpmp and most other related 1-aryl-4-alkoxypiperidin-4yl protecting groups (S.94). Apart from the practical problems associated with the prepara-
N
R3
Protection of 2′Hydroxy Functions of Ribonucleosides
93
94
ACKNOWLEDGMENT The author would like to acknowledge the huge contributions that his co-workers have made over a period of more than 30 years to studies on the chemical synthesis of oligo- and poly-ribonucleotides, and especially to those studies relating to the problem of 2′-protection. Some of their names appear in the references below; they are all owed an enormous debt of gratitude.
LITERATURE CITED Beaucage, S.L. and Caruthers, M.H. 1996. The chemical synthesis of DNA/RNA. In Bioorganic Chemistry: Nucleic Acids (S.M. Hecht, ed.) pp. 36-74. Oxford University Press, New York and Oxford. Beaucage, S.L. and Iyer, R.P. 1992. Advances in the synthesis of oligonucleotides by the phosphoramidite approach. Tetrahedron 48:22232311. Beijer, B., Sulston, I., Sproat, B.S., Rider, P., Lamond, A.I., and Neuner, P. 1990. Synthesis and applicatons of oligoribonucleotides with selected 2′-O-methylation using the 2′-O-[1-(2fluorophenyl)-4-methoxypiperidin-4-yl] protecting group. Nucl. Acids Res. 18:5143-5151. Brown, T. and Brown, D.J.S. 1991. Modern machine-aided methods of oligoribonucleotide synthesis. In Oligonucleotides and Analogues. A Practical Approach (F. Eckstein, ed.) pp. 1-24. IRL Press, Oxford. Brown, J.M., Christodoulou, C., Jones, S.S., Modak, A.S., Reese, C.B., Sibanda, S., and Ubasawa, A. 1989a. Synthesis of the 3′-terminal half of yeast alanine transfer ribonucleic acid (tRNAAla) by the phosphotriester approach in solution. Part 1. Preparation of nucleoside building blocks. J. Chem. Soc. Perkin Trans. 1 1735-1750. Brown, J.M., Christodoulou, C., Modak, A.S., Reese, C.B., and Serafinowska, H.T. 1989b. Synthesis of the 3′-terminal half of yeast alanine transfer ribonucleic acid (tRNAAla) by the phosphotriester approach in solution. Part 2. J. Chem. Soc. Perkin Trans. 1 1751-1767.
R1O R2 R1 Si
tion of very large quantities of 2′-O-TBDMS5′-O-DMTr-protected 3′-phosphoramidites (S.41) that are free from their 2′-isomers (S.42), there is no obvious reason why the TBDMS group should not also be used to protect 2′-hydroxy functions in the large-scale synthesis of RNA sequences in solution.
R2
Figure 2.2.30 Substituted silyl and 1-aryl-4alkoxypiperidin-4-yl protecting groups.
Capaldi, D.C. and Reese, C.B. 1994. Use of the 1-(2-fluorophenyl)-4-methoxypiperidin-4-yl (Fpmp) and related protecting groups in oligoribonucleotide synthesis: Stability of internucleotide linkages to aqueous acid. Nucl. Acids Res. 22:2209-2216.
2.2.20 Current Protocols in Nucleic Acid Chemistry
Chaix, C., Molko, D., and Téoule, R. 1989. The use of labile base protecting groups in oligoribonucleotide synthesis. Tetrahedron Lett. 30:71-74. Chattopadhyaya, J.B. and Reese, C.B. 1978. The 9-phenylxanthen-9-yl protecting group. J.Chem. Soc., Chem. Commun. 639-640. Chattopadhyaya, J.B., Reese, C.B., and Todd, A.H. 1979. 2-Dibromobenzoyl: An acyl protecting group removable under exceptionally mild conditions. J. Chem. Soc., Chem. Commun. 987988. Christodoulou, C., Agrawal, S., and Gait, M.J. 1986. Incompatibility of acid-labile 2′ and 5′ protecting groups for solid-phase synthesis of oligoribonucleotides. Tetrahedron Lett. 27:1521-1522. Corey, E.J. and Venkateswarlu, A. 1972. Protection of hydroxyl groups as tert-butyldimethylsilyl derivatives. J. Am. Chem. Soc. 94:6190-6191. Damha, M.J. and Ogilvie, K.K. 1993. Oligoribonucleotide synthesis. The silyl-phosphoramidite method. In Protocols for Oligonucleotides and Analogs (S. Agrawal, ed.) pp. 81-114. Humana Press, Totowa, N.J. den Hartog, J.A.J., Wille, G., and van Boom, J.H. 1981. Synthesis of oligoribonucleotides with sequences identical to the nucleation region of tobacco mosaic virus RNA: Preparation of AAG, AAGAAG and AAGAAGUUG via phosphotriester methods. Rec. Trav. Chim. 100:320-330. Faja, M., Reese, C.B., Song, Q., and Zhang, P.-Z. 1997. Facile preparation of acetals and enol ethers derived from 1-arylpiperidin-4-ones. J. Chem. Soc. Perkin Trans. 1 191-194. Fromageot, H.P.M., Reese, C.B., and Sulston, J.E. 1968. The synthesis of oligoribonucleotides. VI. 2′-O-Acyl ribonucleoside derivatives as intermediates in the synthesis of dinucleoside phosphates. Tetrahedron 24:3533-3540. Gasparutto, D., Livache, T., Bazin, H., Duplaa, A.M., Guy, A., Khorlin, A., Molko, D., Roget, A., and Téoule, R. 1992. Chemical synthesis of a biologically active natural RNA with its minor bases. Nucl. Acids Res. 20:5159-5166. Goodwin, J.T., Stanick, W.A., and Glick, G.D. 1994. Improved solid-phase synthesis of long oligoribonucleotides. Application to tRNAPhe and tRNAGly. J. Org. Chem. 59:7941-7943. Gough, G.R., Miller, T.J., and Mantick, N.A. 1996. p-Nitrobenzyloxymethyl: A new fluoride-removable protecting group for ribonucleoside 2′hydroxyls. Tetrahedron Lett. 37:981-982. Griffin, B.E. and Reese, C.B. 1964. Oligoribonucleotide synthesis via 2,5-protected ribonucleoside derivatives. Tetrahedron Lett. 29252931. Griffin, B.E., Reese, C.B., Stephenson, G.F., and Trentham, D.R. 1966. Oligoribonucleotide synthesis from nucleoside 2′-O-benzyl ethers. Tetrahedron Lett. 4349-4354.
Griffin, B.E., Jarman, M., and Reese, C.B. 1968. The synthesis of oligoribonucleotides. IV. Preparation of dinucleoside phosphates from 2′,5′-protected ribonucleoside derivatives. Tetrahedron 24:639-662. Hakimelahi, G.H., Proba, Z.A., and Ogilvie, K.K. 1982. New catalysts and procedures for the dimethoxytritylation and selective silylation of ribonucleosides. Can. J. Chem. 60:1106-1113. Hayakawa, Y., Kataoka, M., and Noyori, R. 1996. Benzimidazolium triflate as an efficient promoter for nucleotide synthesis via the phosphoramidite method. J. Org. Chem. 61:79967997. Hayes, J.A., Brunden, M.J., Gilham, P.T., and Gough, G.R. 1985. High-yield synthesis of oligoribonucleotides using o-nitrobenzyl protection of 2′-hydroxyls. Tetrahedron Lett. 26:24072410. Honda, S., Urakami, K., Koura, K., Terada, K., Sato, Y., Kohno, K., Sekine, M., and Hata, T. 1984. Synthesis of oligoribonucleotides by the use of S,S-diphenyl N-monomethoxytrityl ribonucleoside 3′-phosphorodithioates. Tetrahedron 40:153-163. Iwai, S. and Ohtsuka, E. 1988. 5′-Levulinyl and 2′-tetrahydrofuranyl protection for the synthesis of oligoribonucleotides by the phosphoramidite approach. Nucl. Acids Res. 16:9443-9456. Iwai, S., Yamada, E., Asaka, M., Hayasa, Y., Inone, H., and Ohtsuka, E. 1987. A new solid phase synthesis of oligoribonucleotides by the phosphoro-p-anisidate method using tetrahydrofuranyl protection of 2′-hydroxyl groups. Nucl. Acids Res. 15:3761-3772. Järvinen, P., Oivanen, M., and Lönnberg, H. 1991. Interconversion and phosphoester hydrolysis of 2′,5′ and 3′,5′-dinucleoside monophosphates: Kinetics and mechanisms. J. Org. Chem. 56:5396-5401. Jones, S.S. and Reese, C.B. 1979. Migration of t-butyldimethylsilyl protecting groups. J. Chem. Soc. Perkin Trans. 1 2762-2764. Jones, S.S., Rayner, B., Reese, C.B., Ubasawa, A., and Ubasawa, M. 1980. Synthesis of the 3′-terminal decaribonucleoside nonaphosphate of yeast alanine transfer ribonucleic acid. Tetrahedron 36:3075-3085. Jones, S.S., Reese, C.B., Sibanda, S., and Ubasawa, A. 1981. The protection of uracil and guanine residues in oligonucleotide synthesis. Tetrahedron Lett. 22:4755-4758. Jones, S.S., Reese, C.B., and Sibanda, S. 1983. Studies directed towards the synthesis of yeast alanine tRNA. In Current Trends in Organic Synthesis (H. Nozaki, ed.) pp. 71-81. Pergamon Press, Oxford. Kamimura, T., Tsuchiya, M., Urakami, K., Koura, K., Sekine, M., Shinozaki, K., Miura, K., and Hata, T. 1984. Synthesis of a dodecaribonucleotide GUAUCAAUAAUG by use of fully protected ribonucleotide building blocks. J. Am. Chem. Soc. 106:4552-4557.
Protection of Nucleosides for Oligonucleotide Synthesis
2.2.21 Current Protocols in Nucleic Acid Chemistry
Kawahara, S., Wada, T., and Sekine, M. 1996. Unprecedented mild acid-catalyzed desilylation of the 2′-O-tert-butyldimethylsilyl group from chemically synthesized oligoribonucleotide intermediates via neighbouring group participation of the internucleotide phosphate residue. J. Am. Chem. Soc. 118:9461-9468. Kempe, T., Chow, F., Sundquist, W.I., Nardi, T.J., Paulson, B., and Peterson, S.M. 1982. Selective 2′-benzoylation at the cis 2′,3′-diols of protected ribonucleosides. New solid phase synthesis of RNA and DNA-RNA mixtures. Nucl. Acids Res. 10:6695-6714. Kierzek, R. 1994. The stability of trisubstituted internucleotide bond in the presence of vicinal 2′hydroxyl. Chemical synthesis of uridylyl(2′phosphate)-(3′→5′)-uridine. Nucleosides Nucleotides 13:1757-1768. Kierzek, R., Caruthers, M.H., Longfellow, C.E., Swinton, D., Turner, D.H., and Freier, S.M. 1986. Polymer-supported RNA synthesis and its application to test the nearest-neighbour model for duplex stability. Biochemistry 25:7840-7846. Kreevoy, M.M. and Taft, R.W. Jr. 1955. The evaluation of inductive and resonance effects on reactivity. I. Hydrolysis rates of acetals of non-conjugated aldehydes and ketones. J. Am.Chem. Soc. 77:5590-5595. Kruse, C.G., Jonkers, F.L., Dert, V., and van der Gen, A. 1979. Synthetic applications of 2-chlorotetrahydrofuran: Protection of alcohols as tetrahydro2-furanyl (THF) ethers. Rec.Trav. Chim. 98:371380. Kuusela, S. and Lönnberg, H. 1994. Hydrolysis and isomerisation of the internucleosidic phosphodiester bonds of polyuridylic acids: Kinetics and mechanism. J. Chem. Soc. Perkin Trans. 2 21092113. Lehmann, C., Xu, Y.-Z., Christodoulou, C., Tan, Z.-K., and Gait, M.J. 1989. Solid-phase synthesis of oligoribonucleotides using 9-fluorenylmethoxycarbonyl (Fmoc) for 5′-hydroxyl protection. Nucl. Acids Res. 17:2379-2390. McGregor, A., Rao, M.V., Duckworth, G., Stockley, P.G., and Connolly, B.A. 1996. Preparation of oligoribonucleotides containing 4-thiouridine using Fpmp chemistry. Photo crosslinking to RNA bridging proteins using 350 nm irradiation. Nucl. Acids Res. 24 : 3173-3180. Morgan, M.A., Kazakov, S.A., and Hecht, S.M. 1995. Phosphoryl migration during the chemical synthesis of RNA. Nucl. Acids Res. 23:3949-3953. Mullah, B. and Andrus, A. 1996. Purification of 5′-O-trityl-on oligoribonucleotides. Investigation of phosphate migration during purification and detritylation. Nucleosides Nucleotides 15:419-430. Protection of 2′Hydroxy Functions of Ribonucleosides
Norman, D.G., Reese, C.B., and Serafinowska, H.T. 1984. The protection of 2′-hydroxy functions in oligoribonucleotide synthesis. Tetrahedron Lett. 25:3015-3018.
Ogilvie, K.K., Sadana, K.L., Thompson, A.E., Quillian, M.A., and Westmore, J.B. 1974. The use of silyl groups in protecting the hydroxyl functions of ribonucleosides. Tetrahedron Lett. 28612863. Ohtsuka, E. and Iwai, S. 1987. Chemical synthesis of RNA. In Synthesis and Applications of DNA and RNA (S.A. Narang, ed.) pp. 115-136. Academic Press, San Diego. Ohtsuka, E., Tanaka, S., and Ikehara, M. 1978. Synthesis of the heptanucleotide corresponding to a eukaryotic initiator tRNA loop sequence. J. Am. Chem. Soc. 100:8210-8213. Ohtsuka, E., Yamane, A., Doi, T., and Ikehara, M. 1984. Chemical synthesis of the 5′-half molecule of E.coli tRNA2Gly. Tetrahedron 40:47-57. Owen, G.R. and Reese, C.B. 1970. A convenient preparation of tetrahydro-4H-pyran-4-one. J. Chem. Soc. C 2401-2403. Pathak, T. and Chattopadhyaya, J. 1985. The 2′-hydroxy function assisted cleavage of the internucleotide phosphotriester bond of a ribonucleotide under acidic conditions. Acta Chem. Scand. B 39:799-806. Pieles, U., Beijer, B., Bohmann, K., Weston, S., O’Loughlin, S., Adam, V., and Sproat, B.S. 1994. New and convenient protection system for pseudouridine, highly suitable for solid phase oligoribonucleotide synthesis. J. Chem. Soc. Perkin Trans. 1 3423-3429. Rao, M.V. and Macfarlane, K. 1995. Improvements to the chemical synthesis of biologically-active RNA using 2′-O-Fpmp chemistry. Nucleosides Nucleotides 14:911-915. Rao, T.S., Reese, C.B., Serafinowska, H.T., Takaku, H., and Zappia, G. 1987. Solid phase synthesis of the 3′-terminal nonadecaribonucleoside octadecaphosphate sequence of yeast alanine transfer ribonucleic acid. Tetrahedron Lett. 28:48974900. Rao, M.V., Reese, C.B., Schehlmann, V., and Yu, P.S. 1993. Use of the 1-(2-fluorophenyl)-4methoxypiperidin-4-yl (Fpmp) protecting group in the solid phase synthesis of oligo- and polyribonucleotides. J. Chem. Soc. Perkin Trans. 1 43-55. Rastogi, H. and Usher, D.A. 1995. A new 2′-hydroxyl protecting group for the automated synthesis of oligoribonucleotides. Nucl. Acids Res. 23:4872-4877. Reese, C.B. 1970. A systematic approach to oligoribonucleotide synthesis. Colloq. Int. Cent. Natl. Rech. Sci. 182:319-328. Reese, C.B. 1978. The chemical synthesis of oligoand poly-nucleotides by the phosphotriester approach. Tetrahedron 34:3143-3179. Reese, C.B. 1989. The chemical synthesis of oligoand poly-ribonucleotides. In Nucleic Acids and Molecular Biology, Vol. 3 (F. Eckstein and D.M.J. Lilley, ed.) pp. 164-181. Springer-Verlag, Berlin.
2.2.22 Current Protocols in Nucleic Acid Chemistry
Reese, C.B. and Skone, P.A. 1984. The protection of thymine and guanine residues in oligodeoxyribonucleotide synthesis. J. Chem. Soc. Perkin Trans. 1 1263-1271. Reese, C.B. and Skone, P.A. 1985. Action of acid on oligoribonucleotide phosphotriester intermediates. Effects of released vicinal hydroxy functions. Nucl. Acids Res. 13:5215-5231. Reese, C.B. and Thompson, E.A. 1988. A new synthesis of 1-arylpiperidin-4-ols. J. Chem. Soc. Perkin Trans. 1 2881-2885. Reese, C.B. and Trentham, D.R. 1965. Acyl migration in ribonucleoside derivatives. TetrahedronLett. 2467-2472. Reese, C.B. and Zard, L. 1981. Some observations relating to oximate ion promoted unblocking of oligonucleotide aryl esters. Nucl. Acids Res. 9:4611-4626. Reese, C.B., Saffhill, R., and Sulston, J.E. 1967. A symmetrical alternative to the tetrahydropyranyl protecting group. J. Am. Chem. Soc. 89:33663368. Reese, C.B., Saffhill, R., and Sulston, J.E. 1970. 4-Methoxytetrahydropyran-4-yl. A symmetrical alternative to the tetrahydropyranyl protecting group. Tetrahedron 26:1023-1030. Reese, C.B., Titmas, R.C., and Yau, L. 1978. Oximate ion promoted unblocking of oligonucleotide phosphotriester intermediates. Tetrahedron Lett. 30:2727-2730. Reese, C.B., Serafinowska, H.T., and Zappia, G. 1986. An acetal group suitable for the protection of 2′-hydroxy functions in rapid oligoribonucleotide synthesis. Tetrahedron Lett. 27:22912294. Reitz, G. and Pfleiderer, W. 1975. Synthese und Eigenschaften von O-benzyl substituierten Diuridylphosphaten. Chem. Ber. 108:2878-2894. Rozners, E., Renhofa, R., Petrova, M., Popelis, J. Kumpins, V., and Bizdena, E., 1992. Synthesis of oligoribonucleotides by the H-phosphonate approach using base labile 2′-O-protecting groups. V. Recent progress in development of the method. Nucleosides Nucleotides 11:579-1593. Rozners, E., Westman, W., and Strömberg, R. 1994. Evaluation of 2’-hydroxyl protection in RNA synthesis using the H-phosphonate approach. Nucl. Acids Res. 22:94-99. Sakatsume, O., Ohtsuki, M., Takaku, H., and Reese, C.B. 1989. Solid phase synthesis of oligoribonucleotides using the 1-[(2-chloro-4-methyl)]-4methoxypiperidin-4-yl (Ctmp) group for the protection of the 2′-hydroxy functions and the H-phosphonate approach. Nucl. Acids Res. 17:3689-3697. Sandström, A., Kwiatkowski, M., and Chattopadhyaya, J. 1985. Chemical synthesis of a pentaribonucleoside tetraphosphate constituting the 3′acceptor stem sequence of E. coli tRNAIle using 2′-O-(3-methoxy-1,5-dicarbomethoxypentan3-yl)-ribonucleoside building blocks. Acta Chem. Scand. B 39:273-290.
Scaringe, S.A., Francklyn, C., and Usman, N. 1990. Chemical synthesis of biologically active oligoribonucleotides using β-cyanoethyl protected ribonucleoside phosphoramidites. Nucl.Acids Res. 18:5433-5441. Schaller, H., Weimann, G., Lerch, B., and Khorana, H.G. 1963. Protected derivatives of deoxyribonucleosides and new syntheses of deoxyribonucleoside-3′ phosphates. J. Am.Chem. Soc. 85:3821-3827. Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., deBear, J.S., and Gough, G.R. 1992. Rapid synthesis of oligoribonucleotides using 2′-O-(o-nitrobenzyloxymethyl)-protected monomers. Bioorg. Med. Chem. Lett. 2:1019-1024. Sinha, N.O., Biernat, J., and Köster, H. 1983. β-Cyanoethyl N,N-dialkylamino/N-morpholinomonochlorophosphoramidites, new phosphitylating agents facilitating ease of deprotection and work-up of synthesized oligonucleotides. Tetrahedron Lett. 24:5843-5846. Smith, M., Rammler, D.H., Goldberg, I.H., and Khorana, H.G. 1962. Studies on polynucleotides XIV. Specific synthesis of the C-3′-C-5′ inter ribonucleotide linkage. Synthesis of uridylyl(3′→5′)-uridine and uridylyl-(3′→5′)-adenosine. J. Am. Chem. Soc. 84:430-440. Smrt, J. and Šorm, F. 1962. Oligonucleotidic compounds I. The direct blocking of 2′-hydroxyl in ribonucleoside-3′ phosphates. The synthesis of 6-azauridylyl-(5′→3′)-uridine. Coll. Czech. Chem. Commun. 27:73-86. Sonveaux, E. 1994. Protecting groups in oligonucleotide synthesis. In Protocols for Oligonucleotide Conjugates: Synthesis and Analytical Techniques (S. Agrawal, ed.) pp. 1-71. Humana Press, Totowa, N.J. Sproat, B.S. and Gait, M.J. 1984. Solid-phase synthesis of oligodeoxyribonucleotides by the phosphotriester method. In Oligonucleotide Synthesis. A Practical Approach (M.J. Gait, ed.) pp. 83-115. IRL Press, Oxford. Sproat, B.S., Beijer, B., Groetli, M., Ryder, U., Morand, K.L., and Lamond, A.I. 1994. Novel solid-phase synthesis of branched oligoribonucleotides including a substrate for RNA debranching enzyme. J. Chem. Soc. Perkin Trans. 1 419-431. Sproat, B.S., Calonna, F., Mullah, B., Tsou, D., Andrus, A., Hampel, A., and Vinayak, R. 1995. An efficient method for the isolation and purification of oligoribonucleotides. Nucleosides Nucleotides 14:255-273. Stawinski, J., Strömberg, R., Thelin, M., and Westman, E. 1988. Studies on the t-butyldimethylsilyl group as 2′-O-protection in oligoribonucleotide synthesis via the H-phosphonate approach. Nucl. Acids Res. 16:9285-9298. Stork, G. and Hudrlik, P.F. 1968. Isolation of ketone enolates as trialkylsilyl ethers. J. Am.Chem. Soc. 90:4462-4464.
Protection of Nucleosides for Oligonucleotide Synthesis
2.2.23 Current Protocols in Nucleic Acid Chemistry
Takaku, H. and Kamaike, K. 1982. Synthesis of oligoribonucleotides using 4-methoxybenzyl group as a new protecting group of the 2′-hydroxyl group of adenosine. Chem. Lett. 189-192.
van Boom, J.H. and Burgers, P.M.J. 1976. Use of levulinic acid in the protection of oligonucleotides via the modified phosphotriester method: Synthesis of the decaribonucleotide UAUAUAUAUA. Tetrahedron Lett. 4875-4878.
Takaku, H., Kamaike, K., and Tsuchiya, H. 1984. Synthesis of ribooligonucleotides using the 4methoxybenzyl group as a new protecting group for the 2′-hydroxyl group. J. Org. Chem. 49:5156.
van Boom, J.H., van Deursen, P., Meeuse, J., and Reese, C.B. 1972. Two sulphur-containing protecting groups for alcoholic hydroxyl functions. J. Chem. Soc., Chem. Commun. 766-767.
Takaku, H., Ito, T. and Iwai, K. 1986. Use of the 3,4-dimethoxybenzyl group as a protecting group for the 2′-hydroxyl group in the synthesis of oligoribonucleotides. Chem. Lett. 1005-1008.
Vinayak, R., Anderson, P., McCollum, C., and Hampel, A. 1992. Chemical synthesis of RNA using fast oligonucleotide deprotection chemistry. Nucl. Acids Res. 20:1265-1269.
Tanaka, T., Fujino, K., Tamatsukuri, S., and Ikehara, M. 1986. Synthesis of oligoribonucleotides via the phosphite triester approach on a solid support. Chem. Pharm. Bull. Jpn. 34:4126-4132.
Westman, E. and Strömberg, R. 1994. Removal of t-butyldimethylsilyl protection in RNA synthesis. Triethylamine trihydrofluoride (TEA,3HF) is a more reliable alternative to tetrabutylammonium fluoride (TBAF). Nucl. Acids Res. 22:2430-2431.
Tanimura, H. and Imada, T. 1990. The utility of 2′-Thp group in the synthesis of the relatively long RNA fragments on the solid support. Chem. Lett. 2081-2084. Tanimura, H., Mieda, M., Fukazawa, T., Sekine, M., and Hata, T. 1989. Chemical synthesis of the 24 RNA fragments corresponding to hop stunt viroid. Nucl. Acids Res. 17:8135-8147. Usman, N., Ogilvie, K.K., Jiang, M.Y., and Cedergren, R.J. 1987. Automated chemical synthesis of long oligoribonucleotides using 2′-O-silylated ribonucleoside 3′-phosphoramidites on a controlled pore glass support: Synthesis of a 43-nucleotide sequence similar to the 3′-half molecule of Escherichia coli formylmethionine tRNA. J. Am. Chem. Soc.109:7845-7854.
Wincott, F., DiRenzo, A., Shaffer, C., Grimm, S., Tracz, D., Workman, C., Sweedler, D., Gonzalez, C., Scaringe, S., and Usman, N. 1995. Synthesis, deprotection, analysis and purification of RNA and ribozymes. Nucl. Acids Res. 23:2677-2684.
Contributed by Colin B. Reese King’s College London London, United Kingdom
Protection of 2′Hydroxy Functions of Ribonucleosides
2.2.24 Current Protocols in Nucleic Acid Chemistry
Protection of 5′-Hydroxy Functions of Nucleosides The 5′-OH group is the primary hydroxy group of nucleosides. It is the least influenced by the electron-withdrawing effects of the other substituents on the sugar moiety. Moreover, it is the least sterically hindered hydroxy function, and shows the highest reactivity of all nucleoside hydroxy groups in nucleophilic substitutions. Although nucleobases can eventually be left unprotected, and nucleosides with free 3′-hydroxy groups have been used in some triester syntheses, it is mandatory to protect 5′ hydroxyls in all methods of oligonucleotide synthesis that require nucleoside synthons. For chemical oligonucleotide synthesis, the blocking groups for the 5′-hydroxy function must be integrated into an orthogonal protection system. Several such systems have been proposed and are in use for deoxyribo- and ribooligonucleotide synthesis as well as for the preparation of structurally modified oligonucleotide analogs. These protection schemes, details of which are described elsewhere (e.g., UNITS 2.1-2.5 and 3.1-3.4), can be distinguished by requiring, in principle, three alternative methods for 5′-deprotection: (1) acid conditions, (2) alkaline or ammoniacal conditions, or (3) selective deblocking reagents applied essentially in the absence of acid or base. Protecting groups for 5′-hydroxy functions can be broadly classified into these three categories, and the subsequent sections of this unit will follow this division. Additional criteria governing the choice of 5′-hydroxyl-protecting groups include (1) the direction of chain lengthening, (2) the use of polymer supports and/or other purification handles, and (3) additional features related to molecular instability or chemical reactivity in the case of oligonucleotides deviating from biological structure. In particular, the direction of chain extension determines whether a 5′-hydroxyl-protecting group will be permanent (i.e., remain attached to the growing oligonucleotide chain) or intermediary (i.e., will have to be removed prior to each chain extension). If lengthening occurs from the 5′ to the 3′ end, a permanent protecting group is used. If chain extension is done from the 3′ to the 5′ end, which is currently most common, the 5′-hydroxy function must be substituted by an intermediary protecting group. In polymer support synthesis, the poly-
UNIT 2.3
meric carrier assumes the role of a permanent protecting group and is now usually attached to the 3′-hydroxy end. However, there are a number of mostly earlier publications that describe carrier fixation through the 5′ end (see Miscellaneous Acid-Labile 5′-Substituents and see 5′-Hydroxyl-Protecting Groups Cleaved Under Nonacidic and Nonalkaline Conditions). In polymer support synthesis and in some solution methods, it is desirable to simplify the workup of the crude product obtained after oligonucleotide chain extensions. A variety of 5′-hydroxyl-protecting groups have been designed that serve as “purification handles” for this purpose (see Triaryl-methyl Groups as Affinity Ligands). The underlying idea is to single out the product chain from a complex admixture of truncated and failure sequences, although this may still be an unattainable goal. Nevertheless, such “handle” methods may significantly reduce the time and effort for oligonucleotide purification, especially on a preparative scale. A final point in these introductory remarks is that there is no protecting group exclusively in use for the 5′-hydroxy function. Only the conditions of the reaction determine whether the same group will serve for the protection of the 5′, the 3′, or the 2′ hydroxyl, or even of functional groups at nucleobases, because there are only subtle differences in the reactivity of all of these functions. To master these subtle differences is an art of regioselective substitution and comprises much of the challenge of oligonucleotide synthesis. Additionally, there are differences in the approach used for deoxyribo- versus ribooligonucleotide synthesis, and these will be discussed in the subsequent sections.
SCOPE OF THIS OVERVIEW This unit will deal with substituents for the 5′-hydroxy function that fulfill the following criteria of protecting groups. (1) The substituents can be affixed to/removed from the 5′hydroxy function of a growing oligonucleotide chain before/after chain elongation or in the context of other reactions of the oligonucleotide chain. (2) The substituents remain bound during chain elongation and do not interfere with other oligonucleotide reactions. Substituents that serve to permanently modify
Contributed by H. Seliger Current Protocols in Nucleic Acid Chemistry (2000) 2.3.1-2.3.34 Copyright © 2000 by John Wiley & Sons, Inc.
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.1
the 5′ position are not in the scope of this unit. This includes chemical modifications of the 5′-hydroxy function as well as the substitution of the 5′-hydroxy function by linkers, labels, spacers, and other groups that are meant to remain an integral part of the completed oligonucleotide chain (e.g., UNITS 4.2 & 4.3). Also, protecting groups used specifically for the preparation of certain oligonucleotide analogs will not be treated here, as such syntheses are discussed in further units (e.g., see Chapter 4). Protected nucleoside-5′-phosphates have played a role especially in the early days of oligonucleotide synthesis. Essentially, a 5′phosphate residue can be used for protection of the terminus, because enzymatic hydrolysis can easily reconvert to a 5′ hydroxyl. However, only those cases when this was the declared reason for 5′-phosphorylation will be mentioned in the following text (see examples given in Fig. 2.3.10). The topic of 5′-hydroxyl protection is essential to oligonucleotide synthesis and, therefore, is generally included in all textbooks on preparative nucleotide chemistry, as well as in books and articles dealing specifically with the field of protective groups. A few recent reviews have dealt more extensively with groups for protection of 5′ hydroxyls, including articles by Sonveaux (1986) and Beaucage and Iyer (1992, 1993). More extensive reviews in the earlier literature are given by Kössel and Seliger (1975) and Reese (1978).
ACID-LABILE PROTECTING GROUPS Triarylmethyl and Related Substituents
Protection of 5′-Hydroxy Functions of Nucleosides
Introduction of trityl and substituted trityl groups The triphenylmethyl (trityl or Tr) group (S.1; Fig. 2.3.1), a well-known sugar protecting group, was first used for nucleoside 5′-protection in Lord Todd’s laboratory (Andersen et al., 1954) during hydrogenolytic removal. However, this work had limited success. The breakthrough came from H.G. Khorana’s group, who discovered that detritylation in acidic medium is greatly facilitated if the trityl group is modified with one to three p-methoxy substituents (Gilham and Khorana, 1958; Smith et al., 1962). In 80% acetic acid, the rate of hydrolysis increased roughly 10-fold with each introduction of an additional methoxy group.
R
R'
C
R'' 1 2 3 4
R = R' = R'' = H R = R' = H; R'' = OCH3 R = H; R' = R'' = OCH3 R = R' = R'' = OCH3
Figure 2.3.1 Trityl and p-methoxy-substituted trityl protecting groups.
Since their introduction as acid-labile protecting groups for ribo- (Smith et al., 1962) and deoxyribonucleotide (Schaller et al., 1963) chemistry, the mono- and dimethoxytrityl groups (MMTr, S.2, and DMTr, S.3, respectively; Fig. 2.3.1) have become standard for the protection of the 5′-hydroxy function. The DMTr group has especially proven its value in automated solid-phase deoxyribooligonucleotide synthesis, for five main reasons (Sonveaux, 1986). (1) It can be introduced regiospecifically and in high yield at the 5′-hydroxy function of (base-protected) nucleosides. (2) It can be readily and quantitatively removed from the growing oligonucleotide chain by nonaqueous acid. (3) It is sufficiently stable to tetrazole, which is used as an activator in the chain extension step. (4) Its deprotection in nonaqueous acid gives an intense color reaction (ascribed to a cationic species), which can be monitored by spectroscopy to estimate yields of chain elongation. (5) A terminal 5′-DMTr group conveys a certain hydrophobicity to the longest oligonucleotide chain in the crude product released from the polymer support. This hydrophobicity is often used to isolate the target oligonucleotide from the mixture of truncated and failure chains. These considerations (1 to 5) will be elaborated on further throughout this unit. Other trityl protecting groups are less useful for automated synthesis. Unsubstituted trityl and MMTr require conditions that are too harsh for multistep removal with the complication of depurination (see below). The trimethoxytrityl group (TMTr; S.4; Fig. 2.3.1) is extremely sensitive. It can be introduced readily and in high yield at the 5′-position of deoxyribonucleosides and was found to be completely stable when stored at −20°C for ∼6 months; however, it is partially removed in a mixture of tetrazole
2.3.2 Current Protocols in Nucleic Acid Chemistry
for tritylation. This method is no longer in use. The more elegant route B relies on transient silyl protection to apply acyl groups regioselectively to the nucleobases in a procedure that can be carried out in a single reaction tube (Ti et al., 1982; also see “silylation first” procedure in Fritz et al., 1982). The alternative route C uses initial 5′-tritylation followed by silylation and then acylation (“tritylation first” procedure, Fritz et al., 1982; for a recent report see Wada et al., 1998a,b). In most cases, pyridine serves both to dissolve the reactants and to neutralize the ensuing hydrochloric acid. If necessary, trityl groups can be substituted at the unprotected 5′-hydroxy function of oligonucleotide chains, either postsynthesis or in exchange for other protecting groups. This can be done by treating a support-bound oligonucleotide with 4,4′-DMTr chloride in pyridine/4-dimethylaminopyridine for 1 hr, which restores ∼95% of the previously removed DMTr (Kotschi, 1987; Reddy et al., 1987). This reaction could even be performed in the presence of unprotected internucleotidic bonds due to the lability of phosphoric acid trityl esters (Reddy et al., 1987). With support-bound nucleosides, however, the reaction was more sluggish and had to be
and acetonitrile (Kotschi, 1987). This detritylation occurred more readily with purine than with pyrimidine nucleosides. Also, an oligoadenylate prepared with TMTr-deoxyadenosine contained a significant admixture of longer chains, obviously arising from multiple monomer addition. The trityl protecting groups are generally introduced by treatment of nucleosides with the respective trityl chloride. The reactivity of these trityl chlorides increases with increasing number of p-methoxy substituents. If the reaction is run at room temperature and with not more than a slight excess of trityl chloride, the substitution will be highly regiospecific at the 5′ hydroxyl. If the reagent is in higher excess and the temperature is raised, the substitution also occurs on the more sterically hindered 3′ hydroxyl. Exocyclic amino groups are usually protected prior to tritylation, because they would otherwise react with trityl chlorides. Three routes to base-protected, 5′-tritylated nucleosides are described in Figure 2.3.2. The classical approach (route A) from Khorana’s laboratory uses per-acylation of all hydroxy and amino functions, followed by treatment with strong alkali, which selectively cleaves acyl esters and thus liberates 5′ and 3′ hydroxyls
route A AcylO
O
route B
B
Acyl
HO
AcylO
B
O
acylation
route C DMTrO
HO
B
OH 3',5'-silylation
Me3SiO
O
tritylation
B
O
3'-silylation DMTrO
O
B
alkaline deesterification Me3SiO
Me3SiO acylation
HO
Acyl
O
B
Acyl
Me3SiO
O
desilylation
OH
B
DMTrO
DMTrO
O
B
Acyl
Me3SiO
Me3SiO
tritylation
acylation
Acyl
O
B
desilylation
OH
Figure 2.3.2 Three routes to N-protected, 5′-O-tritylated 2′-deoxyribonucleosides (modified from Fritz et al., 1982, with permission from Verlag Chemie).
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.3 Current Protocols in Nucleic Acid Chemistry
activated by addition of tetra-n-butylammonium nitrate and 2,4,6-collidine in dimethylformamide (DMF). The retritylation procedure has recently been applied to the preparation of oligonucleotide-polyamide conjugates (Tong et al., 1993) and to the postsynthetic introduction of the 4-(17-tetrabenzo(a,c,g,i)fluorenylmethyl)-4′,4′′-dimeth oxytrityl protecting group (TBF-DMTr; S.25; Fig. 2.3.8; Ramage and Wahl, 1993; see Triarylmethyl Groups as Affinity Ligands). Other methods of tritylation can be applied if this is required by the sensitivity of modified nucleoside or oligonucleotide reactants. Reports have described the application of powdered molecular sieves as acid scavengers (Kohli et al., 1980), and the use of 4-N,N-dimethylaminopyridine in a mixture of triethylamine and DMF as an alternative solvent (Chaudhary and Hernandez, 1979). As alternatives to trityl chlorides, N-tritylpyridinium fluoroborate (Fersht and Jencks, 1970) and DMTr tetrafluoroborate (Lakshman and Zajc, 1996) have been described.
Removal of trityl and substituted trityl groups The removal of trityl groups was initially performed with 80% aqueous acetic acid (e.g., Smith et al., 1962; Schaller et al., 1963), and such aqueous media are still in use if the deprotection is carried out in the last step of oligonucleotide workup. Nuclear magnetic resonance (NMR) studies have shown that the chemical shifts of all nucleoside protons change upon removal of 5′-trityl groups. On this basis, rate constants were determined for the hydrolysis of different methoxy-substituted trityl groups from the 5′ and 3′ positions of deoxythymidine in aqueous tetrahydrofuran solution brought to different pHs with HCl; a linear relationship between pH and hydrolysis rate was established (Regel et al., 1974).
If detritylations are done as intermediate steps during machine-aided polymer-support synthesis, nonaqueous acidic media are generally applied. The most common are solutions of strong protic acids, such as trichloro- or dichloroacetic acid (Adams et al., 1983; for a recent survey of detritylation conditions see Habus and Agrawal, 1994). Methylene chloride is routinely used as a solvent, although there is a tendency to avoid chlorinated hydrocarbons. For this reason, toluene has been recommended as an alternative for detritylation with dichloroacetic acid (Krotz et al., 1999; Table 2.3.1). This was the result of extensive kinetic studies that showed that the rate of detritylation decreases in the series DMTrdGi-Bu > DMTrdABz > DMTrdCBz > DMTrdT (where Bz is benzoyl), and is highest not only in haloaliphatic but also in aromatic solvents, but is slow in DMF, hexane, ethylacetate, tetrahydrofuran, or tertbutylmethyl ether. Unwanted retritylation can occur through reversal of the equilibrium generated by nonaqueous detritylation, between the colored cationic species and the DMTr-oligonucleotide (Fig. 2.3.3; Dellinger et al., 1998). The same problem was reported for the 9-phenylxanthen9-yl group (see discussion of pixyl and related protecting groups, below; Reese et al., 1986). This is usually not a problem in solid-support oligonucleotide synthesis, since this equilibrium is shifted by washing and filtration; however, it can lead to incomplete deblocking in solution-phase synthesis. Detritylation steps in solid-phase oligonucleotide synthesis are generally believed to be complete when appropriate treatment with nonaqueous acid and extensive washing leave resins and wash solutions colorless. Minute deviations from this scheme of quantitative deblocking, which can occur, for example, through trace impurities in the deblocking solution, are listed among the reasons for the occurrence of truncated sequences in the crude
OCH3
H3CO
Protection of 5′-Hydroxy Functions of Nucleosides
C O
P
Figure 2.3.3
OCH3
O
B
CI2CHCOOH
H3CO
C
CI2CHCOO
HO
O
O
O
P
P
B
= polymer support
Detritylation equilibrium.
2.3.4 Current Protocols in Nucleic Acid Chemistry
products released from the polymer support (e.g., Fearon et al., 1995). Whether this is a potential reason for failure cannot be decided without distinguishing between the efficiencies of the individual steps in the elongation cycle. The literature appears undecided about whether such failures occur statistically throughout all cycles (Fearon et al., 1995) or with higher probability during the initial chain elongation (Temsamani et al., 1995). Depurination as a side reaction during detritylation The most stringent problem, however, is the avoidance of depurination on removal of trityl groups. N-Acylated nucleosides and N-acylated units in oligonucleotides are especially susceptible to deglycosidation. This leads to the formation of apurinic sites, with subsequent chain cleavage during ammoniacal deprotection. This side reaction becomes more and more problematic with longer oligonucleotide chains or larger-scale preparations. The incentive to overcome the depurination problem has led to the development of a wide variety of finely tuned acid-deprotection conditions; examples are given in Table 2.3.1. Although most publications list only the optimum detritylation conditions without giving background data, recent studies, stimulated by large-scale oligonu-
cleotide support synthesis, have been accompanied by extensive analyses of detritylation versus depurination kinetics (Paul and Royappa, 1996; Septak, 1996). The essence of their findings is that haloacetic acids bind strongly to immobilized growing oligonucleotide chains. If, as usual, very dilute acid solutions are applied, detritylation is slowed by depletion of acid from the medium, whereas depurination is allowed to proceed through acid saturation. The authors, therefore, recommend using a short pulse of more concentrated acid (e.g., 15% dichloroacetic acid in methylene chloride) and avoiding any acetonitrile contamination. The length of the acid treatment must be adjusted to the length of the growing oligonucleotide chain. In some cases, ion exchange resins in the H+ form may be a good choice for detritylation (Patil et al., 1994), especially to substitute for acetic acid in large-scale preparations (Iyer et al., 1995). A long treatment with silica gel was found advantageous in the preparation of sensitive nucleosides (Rosowsky et al., 1989). Lewis acid deprotection of trityl and substituted trityl groups and miscellaneous detritylation methods Great expectations to solve the depurination problem had accompanied the introduction of
Table 2.3.1 Examples of Acidic Deprotection Conditions of 5′-Trityl Groupsa
Protecting group DMTr DMTr DMTr DMTr DMTr DMTr DMTr DMTr DMTr DMTr DMTr
Deprotection conditions
Application
Reference
Benzene sulfonic acid in 9:1 (v/v) DMF/DCM 3% (w/v) TCA in DCM
ODN solid phase
Patel et al. (1982)
3% (w/v) TCA in 95:5 (v/v) DCM/CH3OH 0.1 M p-toluene sulfonic acid in THF 0.1 M p-toluene sulfonic acid in acetonitrile 2% (v/v) DCA/0.1% (v/v) CH3OH in DCM 3% (v/v) DCA in 1,2-dichloroethane 3% (v/v) DCA in toluene 3% (w/v) TCA in 1% CH3OH/nitromethane 2% (w/v) benzene sulfonic acid in 7:3 (v/v) DCM/CH3OH 15% (v/v) DCA in DCM
A-rich ODN solid phase Tanaka and Oishi (1985) Protected dA Takaku et al. (1983) ODN solid phase
ODN solid phase
Matteucci and Caruthers (1981) Seliger et al. (1987); Septak (1996) Habus and Agrawal (1994) Sproat and Gait (1984)
ODN solid phase ODN solid phase
Krotz et al. (1999) Sinha et al. (1984)
ODN solution
Gaffney et al. (1984)
ODN solid phase
Habus and Agrawal (1994)
ODN solid phase ODN solid phase
aAbbreviations: DCA, dichloroacetic acid; DCM, dichloromethane; DMF, dimethylformamide; DMTr, dimethoxytrityl; ODN, conditions applied in oligodeoxynucleotide synthesis; TCA, trichloroacetic acid; THF, tetrahydrofuran.
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.5 Current Protocols in Nucleic Acid Chemistry
OCH3
CH3O
MeOH
Ph
HN N
C O
ZnBr2
O
B
OH
Protection of 5′-Hydroxy Functions of Nucleosides
N
N
HO
O HO
O ZnBr2
N
H
Figure 2.3.4 Proposed mechanism for the selective removal of 5′-O-DMTr groups by ZnBr2 in organic solvent (according to Matteucci and Caruthers, 1980).
Figure 2.3.5 Deacylation of nucleobases as a side reaction in the removal of 5′-O-trityl protecting groups with zinc bromide (Kierzek et al., 1981).
zinc bromide as a detritylating agent (Kohli et al., 1980; Matteucci and Caruthers, 1980). The mechanism, described for βmethoxyethoxymethyl ethers by Corey et al. (1976), involves the formation of a bidentate complex with O-C5′ and intracyclic oxygen (Fig. 2.3.4). This explains the selectivity for 5′-hydroxy over 3′-hydroxy detritylation, which was found to disappear upon addition of an alcohol (Waldmeier et al., 1982). This does not play a role in oligonucleotide support synthesis, since the 3′-hydroxy function, conventionally, is anchored to a support. Here the objective is to ensure the most efficient and quantitative deblocking of immobilized growing chains. Initially, a saturated (∼0.1 M) solution of zinc bromide in nitromethane was applied (Matteucci and Caruthers, 1980, 1981). Under these conditions, depurination of N6benzoyl-deoxyadenosine was found to be insignificant over a 24-hr period. However, the formation of an unreactive chain end (presumably a zinc compound) was observed under anhydrous conditions, requiring a hydrolytic wash after detritylation. Additionally, the time required for complete deblocking was relatively long (∼15 min for purine, ∼30 min for pyrimidine; Matteucci and Caruthers, 1981). Therefore, it was found advantageous to add 1% water (Seliger et al., 1982; Winnacker and Dörper, 1982) or 5% methanol (Caruthers, 1982) to the nitromethane solution, which serves to increase the zinc bromide concentration. Applying zinc bromide in a number of protic solvents, Itakura and colleagues (Kierzek et al., 1981) found that a 0.7 M solution of zinc bromide in 9:1 (v/v) chloroform/methanol would lead to complete detritylation within 1 min. Depurination was not detected; however, on prolonged treatment, N-acyl groups were found to be removed, especially from the adenine moiety (Fig. 2.3.5). Since this was also attributed to a nucleophilic attack on a chelate
of zinc bromide with the protected base (Kierzek et al., 1981; Beaucage and Iyer, 1992), sterically hindered alcohol components were found to suppress this side reaction. A 1 M solution of zinc bromide in 85:15 (v/v) dichloromethane/isopropanol was found to be an optimal deblocking reagent (Kierzek et al., 1981; Ito et al., 1982; Itakura et al., 1984). Addition of zinc bromide to amide groups linking the oligonucleotide chains to the support was also postulated (Ito et al., 1982; Adams et al., 1983). In spite of these extensive investigations, neither zinc bromide nor other Lewis acids— such as TiCl4, AlCl3 (Matteucci and Caruthers, 1981), diethyl and diisopropyl aluminum chloride (Köster and Sinha, 1982), or boron trifluoride, applied as etherate (Engels, 1979) or methanol complex (Mitchell et al., 1990)—are of importance in current automated solid-phase deoxyribooligonucleotide synthesis. One of the reasons may be the deposition or adsorption of reagents and by-products (e.g., zinc salts) within the oligonucleotide/solid support system, resulting in the necessity for extensive and time-consuming washes. Outside the mainstream, a number of studies have reported the application of unusual detritylation reagents. Examples are formic acid (Bessodes et al., 1986), 1,1,1,3,3,3-hexafluoro2-propanol (Leonard and Neelima, 1995), chlorine/chloroform solution (Fuentes et al., 1994), or diethyl oxomalonate/methanol solution (Sekine, 1994); some earlier reports are summarized in other reviews (e.g., Beaucage and Iyer, 1992). Other noteworthy detritylation alternatives, such as reductive cleavage with radical anion (Letsinger and Finnan, 1975) or electrochemical deblocking (Mairanovsky, 1976), may not be easily adaptable to automated oligonucleotide synthesis. In routine solid-phase oligonucleotide synthesis, the detritylation solution usually goes to waste after yield monitoring. However, this is
2.3.6 Current Protocols in Nucleic Acid Chemistry
not tolerable for syntheses scaled up to kilogram dimensions and beyond, because the DMTr group comprises ∼35% of the total weight of constituent-protected nucleoside phosphoramidites. Recently, a process has been reported for the workup and neutralization of the detritylation solution, coupled with the reconversion of the ensuing dimethoxytrityl alcohol (DMTrOH) to DMTr chloride. This process resulted in the recycling of 89% of the weight of DMTr residues. At the same time, >90% of the hazardous solvent dichloromethane was recovered with >95% purity (Guo et al., 1998). Pixyl and related protecting groups As a structural analog to the trityl group, the 9-phenylxanthen-9-yl (pixyl or Px) protecting group (S.5, Fig. 2.3.6) has been described (Chattopadhyaya and Reese, 1978; Chattopadhyaya, 1980). This protecting group is removed by acid at approximately the same rate as the DMTr group; however, the pixyl derivatives of nucleosides can be more readily purified by crystallization. The preparation of monomeric (Christodoulou and Reese, 1983) and all sixteen dimeric (Balgobin et al., 1981b) building blocks for deoxyriboolignucleotide synthesis in solution has been done using the pixyl group, and the combination of such blocks to deoxyribooligonucleotides of biological interest has been described (Josephson and Chattopadhyaya, 1981; Balgobin and Chattopadhyaya, 1982b). A more acid-labile variant is the 9-(panisyl)xanthen-9-yl (MOX) group (S.6; Kwiatkowski et al., 1983; Kwiatkowski and Chattopadhyaya, 1984; Tanimura et al., 1988, 1989; Tanimura and Imada, 1990). Alternatively, the 9-phenylthioxanthen-9-yl (S-pixyl; S.7) and 9-phenyl-7-chlorothioxanthen-9-yl (S.8) groups were introduced to modulate deprotection (Balgobin and Chattopadhyaya, 1982a). The recent finding that the pixyl substituent is susceptible to photochemical cleav-
R
R' X 5 6 7 8
X = O; R = H; R' = H X = O; R = OCH3; R' = H X = S; R = H; R' = H X = S; R = H; R' = Cl
Figure 2.3.6 9-Phenylxanthen-9-yl and related 5′-hydroxyl-protecting groups.
age (Misetic and Boyd, 1998) may revive interest in this 5′-protecting group. Trityl and related groups for 5′-hydroxyl protection in oligoribonucleotide synthesis Somewhat different considerations apply to the use of trityl and related groups for 5′-hydroxyl protection in oligoribonucleotide synthesis. Depurination is not very problematic in this case, which allowed the application of MMTr as the preferred protecting group in earlier studies focusing on the preparation of relatively short sequences by solution methods (for reviews, see Reese, 1978, 1989; Ohtsuka and Iwai, 1987). Detailed procedures for the chemical preparation of small oligoribonucleotides were described by van Boom and Wreesmann (1984). In such oligoribonucleotide syntheses where the conditions of acid deprotection are not of great concern, essentially all of the previously described trityl-derived protecting groups should be applicable in RNA synthesis. This has, in fact, been shown in a number of publications cited earlier; however, for simplicity, most protecting groups have been tested first (and often only) in DNA chemistry. Nonetheless, modern strategies of oligoribonucleotide synthesis, in particular the preparation of long sequences on solid phase, require orthogonality of the complete set of protecting groups used for all functionalities of the RNA synthons. In particular, the intermediate protecting group for the 5′ hydroxyl must blend with the groups used for 2′-hydroxyl protection. The latter must be stable throughout all chain elongations, and their removal after completion of synthesis, in the presence of unprotected phosphodiester internucleotide bonds, should not be done in an alkaline medium for risk of isomerization (e.g., Beaucage and Iyer, 1992). Therefore, 2′-hydroxyl-protecting groups should be removed in acidic solution or by a specific reagent in near-neutral medium. Hence, acid-labile protecting groups of the trityl type can be used for 5′-protection of ribonucleotide synthons only under the following conditions. (1) If an acid-labile protecting group is used for the 2′ hydroxyl, there must be a significant difference in the deprotection rates for the 5′ and 2′ functionalities. (2) If the 2′-hydroxyl-protecting groups are stable to acid and released by a specific reagent in a close-to-neutral medium, there is free choice of acid-labile 5′-hydroxyl protection. (3) An acid-labile protecting group can be used for 2′-hydroxyl protection in combination with a substituent for
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.7 Current Protocols in Nucleic Acid Chemistry
Protection of 5′-Hydroxy Functions of Nucleosides
5′-hydroxyl protection that can be removed under nonacidic conditions. This alternative will be discussed below (see Blocking Groups Labile to Nonacidic Conditions). It is beyond the scope of this unit to give credit to all publications that have described solutions to the above protection alternatives (1) and (2); these are treated in detail in recent reviews (Beaucage and Iyer, 1992, 1993). As a general guideline, it can be said that the combination of the most common acid-labile protecting groups (i.e., DMTr or pixyl for 5′ hydroxyls, and tetrahydropyranyl or 4-methoxytetrahydropyranyl for 2′ hydroxyls) is not fully compatible with approach (1). This can be remedied by changing the conditions or properties of the 5′-hydroxyl-protecting groups or by changing the nature of the 2′-hydroxyl-protecting group. For instance, it was found that the 5′-DMTr group could be deprotected selectively at 0°C without cleavage of 2′-tetrahydropyranyl (Seliger et al., 1983; Caruthers et al., 1986). Since this condition does not lend itself to automated support synthesis, the use of the 4,4′,4′′-trimethoxytrityl protecting group was investigated for the 5′ hydroxyl (Seliger et al., 1986); however, the lability of this substituent caused substantial loss on chromatographic purification of the synthons. A number of 2′-hydroxyl-protecting groups with modulated or increased acid stability have been described that are compatible with 5′DMTr protection. Examples are the 1-(2chloro-4-methylphenyl)-4-methoxypiperidin -4-yl (CTMP) and 1-(2-fluorophenyl)-4methoxypiperidin-4-yl (FPMP) groups (Reese et al., 1986; Reese and Thompson, 1988), and the 1-(2-chloro-ethoxy)ethyl (Yamakage et al., 1989) and 3-methoxy-1,5dicarbomethoxypentan-3-yl (Sandström et al., 1985) groups. However, none of these approaches is of widespread application in current solid-support oligoribonucleotide synthesis. The approach that has currently won the competition for a DMTr-compatible system of 2′- and 5′-hydroxyl protection comes from alternative (2) above, namely the introduction of appropriate silyl groups at the 2′ hydroxyl. This approach, which was mainly developed in the laboratory of Ogilvie (Usman et al., 1987, and references therein; Ogilvie et al., 1988), has been extensively investigated during recent years (for discussion, see Beaucage and Iyer, 1992) and is at present the most commonly used method for automated solid-phase oligoribonucleotide preparations.
Triarylmethyl Groups as Affinity Ligands Up to now, the modification of trityl protecting groups has been discussed only with respect to steering the acid lability by introduction of one, two, or three p-methoxy substituents. In an attempt to simplify the workup of the crude product of polymer-support synthesis, it was first shown that MMTr or DMTr groups can serve to single out the target chains by hydrophobic chromatography if they remain bound to the last monomer unit after completion of chain elongations (“trityl-on purification”; Seliger et al., 1977a, 1978). Subsequent to purification, these groups are cleaved with 80% acetic acid to generate the unprotected oligonucleotide. Such groups are referred to as “purification handles.” More recently, a number of trityl groups substituted with longer alkyl chains, ranging from C8H17 to C16H33 (S.9 to S.13; Table 2.3.2), have been described as lipophilic protecting groups (Görtz and Seliger, 1981) and used for separation on reversed-phase columns (Seliger and Görtz, 1981). The structures of substituted trityl groups tailored for purification assistance and other special applications are summarized in Table 2.3.2. The 4-decyloxytrityl (C10Tr or DTr) group (S.10, Fig. 2.3.7; Seliger and Schmidt, 1987) was found to be especially useful for the purification of genes and gene fragments with lengths up to 147 bases (Seliger et al., 1987; Schmidt et al., 1988). The C16Tr group (S.13) has the highest lipophilic affinity; however, the solubility of C16Tr-nucleosides in acetonitrile and other solvents common for oligonucleotide synthesis is somewhat decreased. In order to minimize the risk of depurination during terminal acid deprotection, a series of more labile 4-methoxy-4′-alkoxytrityl groups (S.14 to S.20; Table 2.3.2) was proposed for purification assistance (Gupta et al., 1991). Of these, the 4-methoxy-4′-octyloxytrityl group (S.19; MOTr; Fig. 2.3.7) was found most suitable. The 4,4′-dianisyl-2′′-hexadecyloxyphenyl group (S.21; Table 2.3.2) has mainly been used for oligoribonucleotide purification (van Boom and Wreesman, 1984). The pixyl (S.5), MOX (S.6; Kwiatkowski et al., 1983; Kwiatkowski and Chattopadhyaya, 1984; Tanimura et al., 1988, 1989; Tanimura and Imada, 1990), and especially the 9-(4-octadecyloxyphenyl)xanthen-9-yl (C18Px; S.22; Fig. 2.3.7; Welch et al., 1986) groups have been used as purification handles in solution- and solid-phase oligonucleotide synthesis.
2.3.8 Current Protocols in Nucleic Acid Chemistry
Table 2.3.2 5′-Hydroxyl-Protecting Groups (R1, R2, and R3) of the Substituted Triarylmethyl Type as Purification Handles or for Other Special Applicationsa,b
StrucR1 ture S.2 Phenyl S.3
Phenyl
p-Anisyl
p-Anisyl
S.5
Phenyl
Xanthen-9-yl
Xanthen-9-yl
AbbreviaDeprotection tion MMTr Acid, ZnBr2, etc. DMTr Acid, ZnBr2, etc. Px Acid
S.6
p-Anisyl
Xanthen-9-yl
Xanthen-9-yl
C1Px
Acid
S.9 Phenyl S.10 Phenyl
Phenyl Phenyl
4-Octyloxyphenyl 4-Decyloxyphenyl
Acid Acid
S.11 Phenyl
Phenyl
Acid
Seliger and Schmidt (1987)
S.12 Phenyl
Phenyl
C14Tr
Acid
Seliger and Schmidt (1987)
S.13 Phenyl
Phenyl
C16Tr
Acid
Seliger and Schmidt (1987)
S.14 S.15 S.16 S.17 S.18 S.19 S.20
p-Anisyl p-Anisyl p-Anisyl p-Anisyl p-Anisyl p-Anisyl p-Anisyl
4-Dodecyloxyphenyl 4-Tetradecyloxyphenyl 4-Hexadecyloxyphenyl 4-Propyloxyphenyl 4-Butyloxyphenyl 4-Pentyloxyphenyl 4-Hexyloxyphenyl 4-Heptyloxyphenyl 4-Octyloxyphenyl 4-Dodecyloxyphenyl 2-Hexadecyloxyphenyl Xanthen-9-yl
C8Tr C10Tr or DTr C12Tr
Chattopadhyaya and Reese (1978) Kwiatkowski and Chattopadhyaya (1984) Seliger and Schmidt (1987) Seliger and Schmidt (1987)
Acid Acid Acid Acid Acid Acid Acid
Gupta et al. (1991) Gupta et al. (1991) Gupta et al. (1991) Gupta et al. (1991) Gupta et al. (1991) Gupta et al. (1991) Gupta et al. (1991)
Acid
Phenyl Phenyl Phenyl Phenyl Phenyl Phenyl Phenyl
S.21 p-Anisyl
R2
R3
Phenyl
p-Anisyl
p-Anisyl
S.22 4-Octadecyl- Xanthen-9-yl oxyphenyl S.23 p-Anisyl p-Anisyl S.24 Phenyl Phenyl S.25 p-Anisyl
p-Anisyl
S.26 p-Anisyl
p-Anisyl
S.30 Phenyl S.31 o-Anisyl S.32 Phenyl S.33 Phenyl S.34 Phenyl S.35 p-Anisyl
p-Anisyl o-Anisyl p-Fluorophenyl Phenyl o-Anisyl p-Anisyl
S.36 p-Anisyl
p-Anisyl
Pyrenyl 4-(17-Tetrabenzo(a,c,g,i)fluorenylmethyl)phenyl 4-(17-Tetrabenzo(a,c,g,i)fluorenylmethyl)phenyl 4-[(Succinimidyl-Noxy)carbonyl]phenyl 1-Naphthyl 1-Naphthyl 1-Naphthyl p-Tolyl o-Anisyl 3-(Imidazolyl-1methyl)phenyl 3-(Imidazolyl-1ethylcarbamoyl) phenyl
MOTr
Reference Seliger et al. (1978) Seliger et al. (1978)
C18Px
Acid
BMPM TBF-Tr
Acid Acid
van Boom and Wreesman (1984) Kwiatkowski and Chattopadhyaya (1984) Fourrey et al. (1987) Ramage and Wahl (1993)
TBFDMTr
Acid
Ramage and Wahl (1993)
Acid
Gildea et al. (1990)
Acid, ZnBr2 Acid, ZnBr2 Acid, ZnBr2
Fisher and Caruthers (1983) Fisher and Caruthers (1983) Fisher and Caruthers (1983)
IDTr
Acid, ZnBr2 Acid, ZnBr2 Acid
Fisher and Caruthers (1983) Fisher and Caruthers (1983) Sekine and Hata (1987)
IETr
Acid
Sekine et al. (1993)
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.9 Current Protocols in Nucleic Acid Chemistry
Table 2.3.2 Continued
StrucR1 ture S.37 p-Anisyl
R2
R3
p-Anisyl
S.38 p-Anisyl
p-Anisyl
S.39 p-Anisyl
p-Anisyl
S.40 p-Anisyl
p-Anisyl
3-(Imidazolyl-1propylcarbamoyl) phenyl 3-(Imidazolyl-1IBTr butylcarbamoyl)phenyl 3-(Imidazolyl-1IHTr hexylcarbamoyl)phenyl 3-(N-MethylimidIMTr azolyl-2-ethylcarbamoyl)phenyl p-Benzoyloxyphenyl
S.41 p-Benzoyl- p-Benzoyloxyl oxyphenyl S.42 p-(4,5p-(4,5-DichloroDichlorophthalimido) phthalimido) phenyl phenyl S.43 p-(Levulin p-(Levulin yloxy)phenyl yloxy)phenyl S.44 p-Anisyl
p-Anisyl
S.45 p-Anisyl
p-Anisyl
p-(4,5-Dichlorophthalimido) phenyl
AbbreviaDeprotection tion IPTr Acid
CPTr
p-(Levulinyloxy)phenyl p-(Fluorenyl-9-methoxycarbonyl)phenyl p-(Fluorenyl-9-methoxy carbonyl)aminophenyl
Reference Sekine et al. (1993)
Acid
Sekine et al. (1993)
Acid
Sekine et al. (1993)
Acid
Sekine et al. (1993)
Alkali
Sekine and Hata (1983)
Hydrazine in Sekine and Hata (1984); pyridine/acetic Happ and Scalfi-Happ acid (1988) Hydrazine in Sekine and Hata (1985) pyridine/acetic acid β- elimination Happ and Scalfi-Happ (1988) β-elimination Happ and Scalfi-Happ (1988)
aAbbreviations: BMPM, 1,1-bis(4-methoxyphenyl)-1-pyrenylmethyl; CPTr, tris-(4,5-dichlorophthalimido)trityl; DMTr, dimethoxytrityl; DTr, 4-decyloxytrityl; IBTr, (imidazolyl-1-butylcarbamoyl)dimethoxytrityl; IDTr, (imidazolyl-1-methylcarbamoyl)dimethoxytrityl; IETr, (imidazolyl-1-ethylcarbamoyl)dimethoxytrityl; IHTr, (imidazolyl-1-hexylcarbamoyl)dimethoxytrityl; IMTr, N-methylimidazolyl-(Z-ethylcarbamoyl)dimethoxytrityl; IPTr, (imidazolyl-1-propylcarbamoyl)dimethoxytrityl; MOTr, 4-methoxy-4′-octyloxytrityl; MMTr, monomethoxytrityl; Px, 9-phenylxanthen-9-yl (pixyl); Tr, triphenylmethyl (trityl). bOther applications might include purification, fluorescent labeling, visible absorption, biotin substitution, color coding, 3′-phosphate activation, and nonacidic cleavage.
Protection of 5′-Hydroxy Functions of Nucleosides
The 1,1-bis(4-methoxyphenyl)-1-pyrenylmethyl (BMPM) group (S.23; Fig. 2.3.8; Table 2.3.2; Fourrey et al., 1987) not only allows the easy purification of target sequences from the solid-phase preparation of deoxyribooligonucleotides and their methyl phosphonate analogs, but also allows their detection by thinlayer chromatography (TLC) or gel electrophoresis down to the picomole level by virtue of the fluorescence of the 5′ substituent in the visible range. Ramage and Wahl (1993) have described the 4-(17-tetrabenzo(a,c,g,i)fluorenylmethyl)trityl (TBF-Tr; S.24) and 4-(17tetrabenzo(a,c,g,i)fluorenylmethyl)-4′,4′′dimethoxytrityl (TBF-DMTr; S.25; Fig. 2.3.8; Table 2.3.2) protecting groups. The latter group was especially found interesting for the purification of long oligonucleotides. Its acid hydrolysis is about twice as fast as that of DMTr, and the product is easily identified through the visible absorption of the substituent. The yields
of TBF-DMTr-protected synthons in phosphoramidite synthesis were only between 70% and 88%, but this problem could be circumvented by postsynthetic treatment of the 5′-deprotected, support-bound oligonucleotide with TBF-DMTrCl. Gildea et al. (1990) have described a DMTr group substituted with a hydroxysuccinimide active ester residue (S.26; Fig. 2.3.9). This linker allowed the addition of, for example, biotin, allowing the possibility of purifying a target oligonucleotide from a crude solid-phase product on a streptavidin-agarose column (Fig. 2.3.9). The 5′-hydroxyl-protecting group could subsequently be released by acid treatment to elute the unprotected oligonucleotide. This trityl-on purification scheme can be coupled with chemical 5′ phosphorylation, as demonstrated by Lönnberg and collaborators (see protecting group S.27 in Figure 2.3.10; Guzaev et al., 1995). Bannwarth and Wippler have de-
2.3.10 Current Protocols in Nucleic Acid Chemistry
OC18H37
C10H21O
CH3O
C
C
O OC8H17
10 (DTr)
22 (C18Px)
19 (MOTr)
Figure 2.3.7 Examples of trityl or pixyl groups modified to serve as purification handles (for complete list see Table 2.3.2).
scribed an interesting approach to combined purification and phosphorylation by adding a protected uridine-5′-phosphate at the 5′ terminus of an oligonucleotide chain. The uridine base was modified with a DMTr-protected thioether residue (S.28 in Fig. 2.3.10). After detritylation, the target sequence was selectively retained and purified by disulfide formation with the activated thiol function of a resin. The oligonucleotide was released by periodate treatment with formation of a terminal 5′ phosphate (Bannwarth and Wippler, 1990). Of course, the “dual applications” described above for many modified or substituted trityl groups can be similarly found in oligoribonucleotide chemistry. The trityl-on purification, for instance, is adaptable in the ribonucleotide series, and a recent publication has established that the deprotection of DMTr does not cause migration of the internucleotidic linkages (Mullah and Andrus, 1996). The 4,4′-dimethoxy-2′′-hexadecyloxytrityl group (S.21; van Boom and Wreesmann, 1984) and the 9(4-octadecyloxyphenyl)xanthen-9-yl group (S.22; Welch et al., 1986) were alternatively introduced to promote the separation of RNA fragments. A problem inherent to polymer-support synthesis is that the product is always a more or less complex mixture of the desired chain with
CH3O
truncated and failure sequences (Földes-Papp et al., 1998, and references therein). In view of the efficiency and selectivity of affinity techniques, a solution-phase synthesis using “affinity protecting groups” would be an interesting alternative to solid-phase preparations (Seliger, 1993). An example of such an affinity separation–based solution synthesis was described (Seliger et al., 1977b) using a combination of the 5′-MMTr and 3′-lipoyl groups. Very recently, an approach to large-scale oligonucleotide synthesis was described in which the chain elongation was done in solution and the extended chain was intermediately anchored to a polymer via a Diels-Alder reaction, so as to allow filtration of educts and by-products (product-anchored sequential synthesis or PASS; Pieken, 1997). A purification-oriented oligonucleotide synthesis in solution has also been achieved through the use of a bridged bis-DMTr 5′-protecting group as soluble carrier (S.29 in Fig. 2.3.11; Biernat et al., 1983).
Color-Coded Triarylmethyl Groups and Triarylmethyl Groups with a Catalytic Function In order to modulate the visible absorption of the species obtained after nonaqueous acid deprotection, a variety of groups of the triarylmethyl type has been constructed (see S.30 to
OCH3
OCH3
C
C
OCH3
23 (Bmpm)
25 (Tbf-DMTr)
Figure 2.3.8 Examples of trityl groups tailored to serve as combined purification handles and visible-absorbing fluorescent markers.
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.11 Current Protocols in Nucleic Acid Chemistry
OCH3
CH3O
C
O
O
N
O
O
26
OCH3
HN
C O oligonucleotide−OH
CH3O
O NH H N
S
N H
O
O
Figure 2.3.9 A hydroxysuccinimide-substituted DMTr group and its biotinyl derivative for affinity purification of solid-phase oligonucleotide products.
S.34 in Table 2.3.2). This should allow the “color coding” of different synthons (Fisher and Caruthers, 1983), a principle that has not found application in routine automated oli-
EtO2C CO2Et DMTrO
DMTrO
B
O
EtO2C CO2Et
O
O P O O
NC O
gonucleotide synthesis, but may be useful to monitor the mixed simultaneous addition of two or more nucleotides.
O
O O
DMTrO
B
O
O P O − O
NH3
O
B
OH
O 27
P
P AcOH / H2O 1. Cl2CHCO2H / CH2Cl2 2. NH3
−
EtO2C CO2Et
O
O P O − O
O
HO
B base
O
O P O − O
OH
O
B
OH
BzO O OBz O MMTrS
O P OR
N 11
Protection of 5′-Hydroxy Functions of Nucleosides
O
N
O O
O
B
OH 28
Figure 2.3.10
5′-Phosphorylation via DMTr- or MMTr-protected protecting groups.
2.3.12 Current Protocols in Nucleic Acid Chemistry
Cl3CCH2O Cl3CCH2O P O O O
B
O C O
O C O
O
B
O O P OCH2CCl3 OCH2CCl3
29
Figure 2.3.11 Example of modified trityl protection for a purification-oriented solution synthesis (Biernat et al., 1983).
tuted derivatives, other acid-labile residues do not play a role as protecting groups in current oligonucleotide chemistry. Acetal groups have, for example, been used in early syntheses (e.g., Grams and Letsinger, 1970), and 5′-O-tetrahydropyranyl or 5′-O-methoxytetrahydropyranyl derivatives have been mentioned (Reese, 1978). Although a thorough discussion is not in the scope of this unit, there are also approaches to solid-phase oligonucleotide synthesis that use a support anchored to the 5′-hydroxy function. A number of studies appeared, especially in the 1980s, in which support-bound trityl groups were used as anchors for oligonucleotide preparations in the 5′-to-3′ direction (Shabarova, 1980; Belagaje and Brush, 1982; BirchHirschfeld et al., 1983; Rosenthal et al., 1983). The availability of 3′-protected nucleoside-5′phosphoramidites or -H-phosphonates allows
Through another structural modification, namely the introduction of an imidazol-1-ylmethyl residue, the DMTr substituent could be transformed into a protecting group that serves to activate a protected 3′-phosphate moiety (S.35; Fig. 2.3.12; Sekine and Hata, 1987). This concept was recently exended to the introduction of a number of 3-imidazolylalkylcarbamoylphenyl-4,4′-dianisylmethyl substituents (S.36 to S.39, Fig. 2.3.12), as well as a corresponding N-methylimidazolyl derivative (S.40; Sekine et al., 1993; Wada et al., 1998b). In addition to accelerating the rate of internucleotide bond formation, a shift in the ratio of diastereomeric triester was observed in some cases.
Miscellaneous Acid-Labile 5′-Substituents In view of the multiple advantages of trityl groups, as well as their modified and substi-
OCH3
OCH3
CH3O
C
O
O
B
CH3O
O N
+
N P O
−
Cl
C
OCH3
O
O O
O
O P SPh
HN
SPh
n
SPh
CH3O
C
O
O
B
O
O
O P SPh
HN
SPh
N MeN
N 35
B
36 37 38 39
n=2 n=3 n=4 n=6
N
40
Figure 2.3.12 DMTr groups substituted with imidazolyl residues for 3′-phosphate activation (S.36 to S.40), and a proposed activated intermediate (S.35).
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.13 Current Protocols in Nucleic Acid Chemistry
the use of all timely methods of solid-phase synthesis in both directions of chain growth. Finally, it may be noteworthy that Tr or DMTr groups present at the 5′ terminus of deoxyribooligonucleotides were found to enhance their anti-HIV activity (Furukawa et al., 1994; Hotoda et al., 1994), a fact that was attributed to the enhancement of membrane permeability through the remaining protecting group.
BLOCKING GROUPS LABILE TO NONACIDIC CONDITIONS Acyl Substituents and Base-Labile Triarylmethyl Groups
Protection of 5′-Hydroxy Functions of Nucleosides
In spite of their obvious advantages, acidlabile protecting groups for the 5′-hydroxy function have some limitations to their application in oligonucleotide synthesis. In the deoxyribonucleotide series, even a small number of statistically distributed depurination sites may seriously impair the quality of a solid-phase or large-scale synthesis product. In the ribonucleotide series, multiple acid deprotection steps may interfere with the stability of 2′-hydroxylprotecting groups. This has stimulated the search for new orthogonal protection schemes involving 5′-hydroxyl-protecting groups that can be deblocked under nonacidic conditions. Of course, there are limitations to the use of simple acyl groups such as acetyl or benzoyl as long as the exocyclic amino groups of nucleobases are blocked by acyl residues as well. Thus, acetyl protecting groups are found in the recent literature only for syntheses at the monomer level—e.g., in the preparation of 15N-labeled (Kamaike et al., 1995, 1996) or 2H-labeled nucleosides (Kawashima et al., 1995, 1997), or of components for oligodeoxyribonucleotides with base-modified or mutagenic units (Matsuda et al., 1993; Ozaki et al., 1994; for earlier literature see, e.g., Kössel and Seliger, 1975; Reese, 1978; Sonveaux, 1986). From the preparative standpoint, methods for rapid O-acylation of nucleoside hydroxy groups through phase-transfer catalysis may be noteworthy (Sekine, 1993). Isobutyryl (Gaffney and Jones, 1982a,b) or methoxyacetyl groups (Reese and Skone, 1984) were introduced at sugar hydroxy functions in order to allow the additional protection of the 6-oxo group of deoxyguanosine as well as the 4-oxo group of thymidine. Acyl groups that can be removed under virtually neutral conditions are more attractive. Earlier examples, such as methoxy- and phenoxyacetyl (Reese and Stewart, 1968), did not
completely fulfill the expectations. A more interesting alternative was the o-bromomethylbenzoyl group (S.46; Fig. 2.3.13; Chattopadhyaya et al., 1979), which was used for the preparation of an SV40-specific deoxyribooligonucleotide (Chattopadhyaya and Reese, 1980). Derivatives of the 4-hydroxybutyryl and 2-hydroxymethylbenzoyl groups (Brown et al., 1984; van Boom and Wreesman, 1984; Reese, 1985; Brown et al., 1989a,b) are similarly of interest, since the neighboring participation of the hydroxy groups allows their hydrolysis under extremely mild conditions. Of course, the hydroxy groups have to be protected during chain elongation. In case of the 4-(methylthiomethoxy)butyryl (MTMB; S.51), 2-(methylthiomethoxymethyl)benzoyl (MTMT; S.47), and 2-(isopropylthiomethoxymethyl)benzoyl (DTMT; S.48) groups (Fig. 2.3.13), the deblocking of the methylthiomethyl residue was done with mercury(II) perchlorate and 2,4,6-collidine within 3 hr at room temperature, and the subsequent treatment with K2CO3 in tetrahydrofuran/water released the 2-hydroxymethylbenzoyl substituent within 30 sec (Fig. 2.3.14). As an alternative, the 2-(2,4-dinitrophenylsulfenyloxymethyl) benzoyl (DNBSB) group (S.49; Fig. 2.3.13) was initiated by removal of the sulfenyl residue with p-toluenethiol (Christodoulou et al., 1987a,b). Although these “protected protecting groups” are appealing from their deprotection conditions, difficulties in in-
O R'
R 46 R = Br; R' = H 47 R = OCH2SCH3; R' = H 48 R = OCH2SCH(CH3)2; R' = H 49 R = O S
NO2
R' = H
O2N 50 R = OCOCH2CH2COCH3; R' = NO2 O
R'' 51 R'' = OCH2SCH3
Figure 2.3.13 Acyl substituents for removal at close-to-neutral pH.
2.3.14 Current Protocols in Nucleic Acid Chemistry
S O O
O
OH
Hg(CIO4)2
T
O
collidine
O
T
K2CO3 THF/H2O
HO
O
T
O
O
HO
HO
HO
Figure 2.3.14 Example of two-step removal of an ortho-substituted benzoyl protecting group (Brown et al., 1984; Reese, 1985).
troduction and, in particular, the two-step procedure of removal make them less practical for current automated oligonucleotide synthesis. The β-benzoylpropionyl (S.52; Letsinger et al., 1967) and the levulinyl (S.53; van Boom and Burgers, 1976; Iwai and Ohtsuka, 1988; Iwai et al., 1990) groups can be deprotected by hydrazine in a pyridine/acetic acid mixture (Fig. 2.3.15). However, a partial deprotection of acyl groups from the nucleobases was also observed under the conditions of hydrazinolysis of the β-benzoylpropionyl group (Letsinger and Miller, 1969). Especially mild conditions of hydrazinolytic deprotection apply to the 2-levulinyloxymethyl-5-nitrobenzoyl group (S.50; Fig. 2.3.13; Kamaike et al., 1997). Appropriate substitution with electrondonating substituents that stabilize a trityl cation may strongly modify the conditions of cleavage. Based upon the earlier finding that the 4-hydroxytrityl group is hydrolyzed much more easily than the corresponding 4-acetoxytrityl residue (Taunton-Rigby et al., 1972), the 4,4′,4′′-tris(benzoyloxy)trityl substituent (S.41; Table 2.3.2) was prepared as an acid-stable but base-labile “protected 5′-protecting group” (Sekine and Hata, 1983), the removal of which is shown in Figure 2.3.16. This principle was further extended to 4,4′,4′′-tris(4,5dichlorophthalimido)trityl (S.42; Sekine and Hata, 1984) and 4,4′,4′′-tris(levulinyloxy)trityl (S.43; Table 2.3.2; Sekine and Hata, 1985) as protecting groups labile to hydrazine treatment in pyridine/acetic acid. Based on the same general concept, the 4-(9-fluorenylmethoxycarbonyl)oxy-4′,4′′-dimethoxytrityl (S.44) and 4-(9-fluorenyl-
methoxycarbonyl)amino-4′,4′′-dimethoxytrityl) (S.45; Table 2.3.2) groups were introduced. Their release could be triggered through β-elimination (Scalfi-Happ et al., 1987; Happ and Scalfi-Happ, 1988). The usefulness of hydrazine-labile modified trityl groups such as 4,4′,4′′-tris(4,5-dichlorophthalimido)trityl (S.42; Sekine and Hata, 1984, 1986; ScalfiHapp et al., 1987) and the S.44 and S.45 groups for β-elimination-triggered deprotection (Happ and Scalfi-Happ, 1988) was also demonstrated for large-scale oligoribonucleotide synthesis and for the preparation of 2′(3′)-Oaminoacyl-oligoribonucleotides (Scalfi-Happ et al., 1987).
Carbonate-Type Protecting Groups Protecting groups of the carbonate type, most popular in peptide chemistry, have also received much attention in oligonucleotide synthesis. In earlier work, the isobutyloxycarbonyl group (S.54; Fig. 2.3.17; Ogilvie and Letsinger, 1967) and the p-nitrophenyloxycarbonyl (NPOC) group (S.55; Letsinger and Ogilvie, 1967) were shown to be introduced rather selectively at 5′-hydroxy functions en route to 3′-tritylated thymidine and uridine derivatives. However, their removal in dilute sodium hydroxide/dioxane would not be compatible with exocyclic acyl protection of nucleobases. p-Nitrophenyloxycarbonyl and a number of other carbonate protecting groups could also be introduced into partially protected thymidine and uridine via the corresponding 5′-chloroformates (Seliger, 1972). These studies later led to the introduction of the 5′-p-phenylazophenyl-
O O
R
O
B
O HO
NH2NH2
HO
N NH R
O
O
B
pyridine / acetic acid HO
52 R = phenyl 53 R = methyl
Figure 2.3.15 groups.
Hydazinolytic deprotection of β-benzoylpropionyl (S.52) and levulinyl (S.53)
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.15 Current Protocols in Nucleic Acid Chemistry
O Ph
O
O
O
O
Ph O O
C X
OH
O
C X
C
HX HX
O Ph
O
O
X = nucleoside-5'-oxy
O 41
Figure 2.3.16 group.
Protection of 5′-Hydroxy Functions of Nucleosides
4,4′,4′′-Tris(benzoyloxy)trityl as an acid-stable, base-labile 5′-hydroxyl-protecting
oxycarbonyl (PAPOC) group (S.56; Kössel and Seliger, 1975), which was introduced via pphenylazophenylchloroformate (Seliger and Kotschi, 1985; Seliger et al., 1986). This group could be released by a two-step treatment with β-cyanoethanol/triethylamine and diazabicycloundecene (DBU). This group retains some of the features of trityl- or pixyl-type substituents (i.e., its deprotection gives a visible-absorbing solution), and the PAPOC nucleosides are readily crystalline; however, the two-step deprotection procedure is less advantageous. More attention has been given to the 9-fluorenylmethoxycarbonyl (FMOC) group (S.57, Fig. 2.3.17), a standard in peptide protection. FMOC-protected synthons allowed an alternative protocol for solid-support deoxyribooligonucleotide synthesis, retaining the standard acyl protection for the bases and avoiding acid deblocking steps (Gioeli and Chattopadhyaya, 1982; Balgobin and Chattopadhyaya, 1987; Ma and Sonveaux, 1987, 1989). The potential of 5′-FMOC protection for oligoribonucleotide synthesis was demonstrated by Fukuda et al. (1988) and by the group of Gait (Lehmann et al., 1989). The FMOC group was also used in acetal-linked solid-support oligonucleotide synthesis (Palom et al., 1993). Recently, the p-chlorophenyloxycarbonyl group (S.57a, Fig. 2.3.17) was found to be removed in <2 min in a solution of peroxyanions buffered to pH 9.6 (10 ml LiOH⋅H2O, 12 ml 30% H2O2, 1.78 g m-chloroperbenzoic acid, 15 ml 1.5 M AMP buffer, pH 10.3, in 50 ml dioxane; Caruthers et al., 1994; Dellinger et al., 1998). This allowed the design of a solid-phase two-step chain-elongation cycle consisting of only a coupling step and the combined deprotection/oxidation step (excluding capping), reducing the phosphoramidite cycle time to 6 min.
Other carbonate protecting groups that could be selectively released in the presence of baseprotecting acyl groups include 2-(trimethylsilyl)ethoxycarbonyl (S.58; Fig. 2.3.17; Gioeli et al., 1981) and 2-(phenylsulfonyl)ethoxycarbonyl (S.59; Balgobin et al., 1981a). More recently, Pfleiderer and colleagues have developed an orthogonal protection scheme for solution- and solid-phase deoxyribo- and ribooligonucleotide synthesis, with avoidance of acid-deprotection steps. The main feature is the exclusive use of protecting groups that can be cleaved by β-elimination. For instance, in combination with the p-nitrophenylethyl (NPE) phosphate protecting group, the 2-(4-nitrophenyl)ethoxycarbonyl (NPEOC; S.60) or 2-(2,4-dinitrophenyl)ethoxycarbonyl (DNPEOC; S.61) substituents were applied for 5′-protection (Schirmeister et al., 1993; Fig. 2.3.18). The NPE/NPEOC strategy was further extended to large-scale oligonucleotide synthesis (Weiler and Pfleiderer, 1995). Particularly attractive is the dansylethoxycarbonyl group (S.62; Bergmann and Pfleiderer, 1994a,b,c), which
O R
O
O
O
B
HO
54 55 56 57 57a 58 59 88
R = isobutyl R = p-nitrophenyl R = p-phenylazophenyl R = fluoren-9-methyl R = p-chlorophenyl R = 2-trimethylsilylethyl R = 2-phenylsulfonylethyl R = o-nitrophenyl
Figure 2.3.17 Carbonate protecting groups for the 5′-hydroxy function.
2.3.16 Current Protocols in Nucleic Acid Chemistry
RO2C O
O
B NO2
O (i-Pr)2N
P
O
60 R = NO2
61 R = O2N
NO2
O S 62 R =
O NMe2
Figure 2.3.18 Carbonate substituents for 5′OH as part of an orthogonal scheme of protecting groups that can be cleaved by β-elimination.
can easily be cleaved with dilute DBU in aprotic solvent and allows yield monitoring by far UV or fluorescence detection at 530 nm, even in small-scale synthesis of long RNA molecules (Pfleiderer et al., 1995). A further advantage of this scheme of deprotection via β-elimination in solid-phase synthesis is the possibility of retaining a deprotected oligonucleotide product on the support, which facilitates purification of the target sequence.
5′-O-Silyl Protecting Groups The use of silyl groups for 5′-hydroxyl protection does not have any major advantages over trityl in the deoxyribonucleotide series; examples in the literature are concerned with modeling the behavior of different silyl groups (e.g., S.63 to S.67; Fig. 2.3.19; Ogilvie et al., 1974) or the preparation of structurally modified nucleosides (e.g., keto-nucleosides; Robins et al., 1990; McEldoon and Wiemer, 1995). In oligoribonucleotide synthesis, however, silyl protecting groups are nearly essential in developing orthogonal protection systems. The alternative of 2′-silyl protection combined with 5′-MMTr or 5′-DMTr has been discussed above (see Triarylmethyl and Related Substituents). A new route to orthogonal ribonucleoside protection uses acid-labile groups for the 2′-hydroxy function in combination with silyl sub-
stituents at the 5′-hydroxy function. In this case, the tetra(isopropyl)disiloxane-1,3-diyl group (S.68; Fig. 2.3.19; Markiewicz, 1979, 1980) can be elegantly used as a transient bifunctional protecting group for 3′ and 5′ hydroxyls, which ensures the regioselective introduction of an acid-labile group at the 2′ position. Subsequent to deblocking of the disiloxane-diyl group, another silyl substituent is selectively attached to protect the 5′-hydroxy function. Two approaches of this kind have recently been described. The 1,1,3,3-tetraisopropyl-3[2-(triphenylmethoxy)ethoxy]disiloxane-1-yl (TES) group (S.69; Fig. 2.3.19; Hirao et al., 1998), a “trityl-protected silyl protecting group,” can be deblocked with 0.1 M tetrabutylammonium fluoride in tetrahydrofuran within 1 min at room temperature. The resultant deblocking solution will be colored on addition of acid. In solid-phase synthesis, using terephthalate as an anchor to the support, the silyl deprotection conditions do not lead to removal of other blocking groups except for the β-cyanoethyl residue at the internucleotidic linkages, which is lost during this treatment. A better solution to this orthogonality problem has recently been reported (Caruthers, 1998, and pers. comm.). A number of silyl substituents (S.70 to S.86; Fig. 2.3.20) can be regioselectively introduced and deprotected in <30 sec with tetrabutylammonium fluoride in tetrahydrofuran. On testing for stability in acid and base (Table 2.3.3), this author found that the bis(trimethylsiloxyl)cyclooctyloxysilyl group (S.83) was best for 5′-hydroxyl protection, as the resulting fully protected phosphoramidite synthons (Fig. 2.3.21) could be prevented from oiling out on workup. The internucleotidic bonds, in this case, were protected by the methoxy group, which was found to be resistant to fluoride cleavage, but is released with a thiol reagent. Alternatively, the bis(trimethylsiloxyl)-1,3-(trityloxy)propyl-2-oxysilyl group (S.87; Fig. 2.3.20) can be designated for monitoring the deprotection step.
Photosensitive 5′-Hydroxyl-Protecting Groups Groups that can be cleaved by photochemical methods have not been of relevance to 5′-hydroxyl protection in earlier times, although photolabile phosphate-protecting groups such as o-nitrobenzyl were in use (Hasan et al., 1997, and references therein). Only recently has the photochemical cleavage of 5′-hydroxyl-protecting groups become an
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.17 Current Protocols in Nucleic Acid Chemistry
R2
O
R1 Si O
O
R3
O
B
Si
B
O
TrO
O Si
HO
Si O
Si O
OH
63 R1 = CH3; R2, R3 = isopropyl
69
68
64 R1 = isopropyl; R2, R3 = cyclopentyl 65 R1, R2 = CH3; R3 = t-butyl 66 R1, R2, R3 = isopropyl 67 R1 = t-butyl; R2, R3 = cyclopentyl
Figure 2.3.19
Silyl protecting groups for the 5′-hydroxy function.
of relevant oligonucleotide spots in a technique similar to photolithography. The efficiency of this technique relies very much on the ease of photochemical deprotection and the extent to which it can be quantified. Several new protecting groups been developed for this purpose. Based on the experience of investigations on carbonate protecting groups, the o-nitrophenyloxycarbonyl (NPOC; S.88;
interesting option, since Fodor and collaborators pointed out the possibility of producing DNA arrays on glass substrates (DNA “chips”) by light-directed solid-phase synthesis (Fodor et al., 1991; Pease et al., 1994). Arrays containing up to 106 unique oligonucleotide sequences per square centimeter can be constructed by the elongation of each sequence triggered through photodeprotection of the 5′-hydroxy function R1 R2 Si O
O
R3
B
HO
O a
Si O PAP
b
Me3SiO
Me d
c
e
O O
TrO
O
O O
g
f
h
i OAc
Ph3SiO j
Protection of 5′-Hydroxy Functions of Nucleosides
O
l
k
Structure
R1
R2
R3
70 71 72 73 74 75 76 77
a a a a a a b b
b h j
78 79
d j
80 81 82 83 84 85 86 87 104 105
a j
a a a a a b b b j k a j
e e e k i e methyl isopropyl
e e k k e e methyl isopropyl
f g b b j j k c k e k k k e i i i
Figure 2.3.20 Silyl protecting groups that are rapidly deblocked by tetrabutylammonium fluoride (Caruthers, 1998). PAP, 4-phenylazophenyl.
2.3.18 Current Protocols in Nucleic Acid Chemistry
Table 2.3.3 Fluoride Deprotection and Acid/Base Degradation of Silyl Protecting Groups S.70 to S.86a
Structure
R1, R2, R3
S.70 S.71 S.72 S.73 S.74 S.75 S.76 S.77 S.78 S.79 S.80 S.81 S.82 S.83 S.84 S.85 S.86
a,a,b a,a,h a,a,j a,a,f a,a,g a,b,b b,b,b b,b,j d,j,j j,k,k a,a,c j,j,k e,e,e e,e,k e,k,k k,k,k i,e,e
Fluoride deprotectionb <1 min <15 sec <15 sec <30 sec <30 sec <30 sec <15 sec <15 sec <15 sec <15 sec <15 sec <15 sec <15 sec <15 sec <15 sec <15 sec <15 sec
Acid degradation (%)c 1% 24 hr 100% 24 hr 1% 24 hr 1% 24 hr 5% 13 hr NA 5% 5 hr 1% 24 hr NA NA NA NA NA NA NA NA NA
Base degradation (%)d 0% 24 hr 75% 24 hr 0% 24 hr 0% 24 hr 5% 13 hr NA 5% 5 hr 0% 24 hr NA NA NA NA NA NA NA NA NA
aFrom Caruthers (1998 and pers. comm.). See Figure 2.3.20 for structures and substituents (R1, R2, R3). bTime for complete removal of protecting group with 5 eq tetrabutylammonium fluoride in tetrahydrofuran. cPercent degradation in 0.01 M HCl within specified time. NA, data not available. dPercent degradation in 10% aqueous triethylamine within specified time. NA, data not available.
Fig. 2.3.17) and o-nitrophenylethoxycarbonyl (NPEOC; S.89; Fig. 2.3.22) groups were reported from Pfleiderer’s laboratory (Hasan et al., 1997). Their photochemical cleavage leads to the formation of o-nitrobenzaldehyde and CO2 in the case of NPOC, or o-nitrostyrene and CO2 in the case of NPEOC (Fig. 2.3.23). The parent nucleosides were recovered in a clean reaction after irradiation at 365 nm, with a half-life of 7.5 min for NPEOC (Bühler et al., 1999; Table 2.3.4). Such long deprotection times are, of course, less suitable if a great
Me3SiO
O Si O
B
O
Me3SiO MeO
P i-Pr2N
O
O
O O
OAc OAc
B = uracil-yl
N4-acetyl-cytosin-yl N6-benzoyl-adenin-yl N2-isobutyryl-guanin-yl
Figure 2.3.21 5′-O-Silyl-protected synthons for oligoribonucleotide synthesis (Caruthers, 1998).
number of parallel oligonucleotide chain extensions are to be performed in a fast and efficient way. Therefore, a number of new protecting groups have been described that are either of the o-NPEOC type (S.90 to S.94, Fig. 2.3.22) or of the o-nitrophenylethylsulfonyl type (S.95 to S.101, Fig. 2.3.22) (Bühler et al., 1999), for which half-lives of photochemical deprotection are reported to be between 10 min and 50 sec (Table 2.3.4). Up to 85% of unprotected nucleosides were recovered after 10 min irradiation time. Other protecting groups such as methylnitropiperonyloxycarbonyl (MeNPOC; S.102; Fig. 2.3.22; Pease et al., 1994) and 3′,5′-dimethoxybenzoinoxycarbonyl (DMBOC; S.103; Fig. 2.3.22; Pirrung and Bradley, 1995a,b) have been tested in solid-support oligonucleotide synthesis. A comparison of results obtained with both protecting groups and with conventional DMTr protection showed that cycle yields are diminished with photochemical deprotection with respect to cycles including conventional detritylation, and some by-product was obtained due to damage of N-benzoyl cytosine bases. In a parallel study, the removal of the MeNPOC group (Fig. 2.3.24) was found to be independent of the length and terminal base of the growing oligonucleotide chain, and was most rapid in nonpolar solvents or in absence of solvent (t1/2 =
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.19 Current Protocols in Nucleic Acid Chemistry
2
1
R
R
O O
3
NO2 R 1
2
3
Structure
R
R
R
89 90 91 92 93 94
H Cl H H H H
H H H Cl Br I
H H CH3 CH3 CH3 CH3
1
R 2
R
3
O S O
R NO2 1
3
R
R
R
95 96 97 98 99 100 101
H Cl H H H H
H H Cl H Cl Br I
H H H CH3 CH3 CH3 CH3
H
O O O
2
Structure
CH3O
OCH3 O
O O
NO2 O (MeNPOC)
(DMBOC)
102
103
Figure 2.3.22 Photochemically cleaved 5′-hydroxyl-protecting groups of the carbonate or sulfonate type. Taken in part from Bühler et al. (1999) with permission from Marcel Dekker.
Protection of 5′-Hydroxy Functions of Nucleosides
10 to 13 sec at A = 365 nm, irradiation intensity 27.5 mW/cm2). The average yield of chain elongation was 92% to 94% in solid-support reactions and ∼96% on photolysis in solution, as compared to 98% in conventional cycles (McGall et al., 1997). A thorough investigation on the DMBOC group (Pirrung et al., 1998) showed half times of deprotection on a glass surface in the presence of dioxane (326.5 nm, irradiation with 44.5 mW/cm2) to be between 5 and 13 sec. The use of shorter wavelengths decreases the deprotection time, but is discouraged due to the risk of damage to the deoxyribooligonucleotide chain. The yields per chain extension cycle were found to be base dependent, ranging from 91% to 98% for T, 82% to 95% for C, 79% to 92% for G, and 74% to 84% for A.
Recently, McGall and collaborators have proposed a technique for the preparation of oligonucleotide arrays that combines photolithography with acid deprotection (McGall et al., 1996; Wallraff et al., 1997). In this case, spots of growing oligonucleotide chains are grafted to the surface of a functionalized carrier (chip), which is then covered by a photosensitive polymer coating (a “photoresist”). Oligonucleotide spots that are meant for chain lengthening are first exposed to further chain elongation by photoetching of the polymer coating. They can then be subjected to conventional 5′-deprotection and chain lengthening. It is likely that, in further pursuit of light-directed combinatorial oligonucleotide synthesis, new protecting groups will have to be developed. Some of these may deviate from the
2.3.20 Current Protocols in Nucleic Acid Chemistry
Table 2.3.4 Photolabile 5′-Protecting Groups of the o-Nitrophenylethoxycarbonyl and o-Nitrophenylethylsulfonyl Typesa
Structure Carbonate groups S.89 S.90 S.91 S.92 S.93 S.94 Sulfonate groups S.95 S.96 S.97 S.98 S.99 S.100 S.101
R1
R2
R3
t1/2b
H Cl H H H H
H H H Cl Br I
H H CH3 CH3 CH3 CH3
7.5 min 82 min 56 sec 62 sec 60 sec 53 sec
H Cl H H H H H
H H C1 H C1 Br I
H H H CH3 CH3 CH3 CH3
10.9 min 16.3 min 10.9 min 82 sec 66 sec 70 sec 60 sec
aData from Bühler et al. (1999) with permission from Marcel Dekker. Deprotection conditions: irradiation with a 200 W mercury lamp (365 nm) in 1:1 (v/v) methanol/water. See Figure 2.3.22 for structures and substituents (R 1, R2, R3). bValues given for 5′-protected thymidine derivatives.
structural concept of carbonate esters, which are now the first choice. Examples include photochemically cleavable silyl groups such as hydroxystyryldimethylsilyl (HSDMS; S.104; Fig. 2.3.20) and hydroxystyryldiisopropylsilyl (HSDIS; S.105; Fig. 2.3.20; Pirrung and Lee, 1993). However, these do not seem to have been
NO2
O O
O R
hv
O N OH
O O
O R
NO2 H C
CH2
CO2
ROH
Figure 2.3.23 Proposed mechanism for photodeprotection of the o-nitrophenylethoxycarbonyl group (Hasan et al., 1997).
extensively tested for oligonucleotide synthesis.
5′-Hydroxyl-Protecting Groups Cleaved Under Nonacidic and Nonalkaline Conditions The general interest in 5′-hydroxyl-protecting groups that are cleaved under nonacidic and nonalkaline conditions has yielded a number of interesting solutions. Structural modifications of trityl that allow deprotection under nonacidic conditions (e.g., in a two-step procedure involving hydrazine in a near-neutral solution) have been discussed in the previous section (see Acyl Substituents and Base-Labile Triarylmethyl Groups). 2,4-Dinitrobenzenesulfenyl residues (S.106, Fig. 2.3.25) have been introduced by reaction with the corresponding sulfenyl chloride (Grams and Letsinger, 1968). This protecting group can be removed by treatment with a nucleophilic reagent such as thiophenol in pyridine (Letsinger et al., 1964). Stereochemical and mechanistic aspects of introduction and release of sulfenyl groups have been investigated (Bazin et al., 1985). The protection of hydroxy functions of sugar derivates by p-methoxybenzyl (S.107) and 3,4-dimethoxybenzyl groups (S.108; Fig. 2.3.25), which are susceptible to oxidative cleavage with 2,3-dichloro-5,6-dicyanobenzoquinone (DDQ), has been described (Oikawa et al., 1982), but these groups are not in use in oligonucleotide chemistry. The p-methoxybenzyl group could also be released from the 5′ hydroxyl in 68% yield by treatment with ceric
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.21 Current Protocols in Nucleic Acid Chemistry
NO2
O O
O
O
hv
O O
B
O O
O P OMe
Figure 2.3.24
O
CO2
O
O O
HO
NO
B
O
O P OMe O
CN
CN
Proposed photocleavage of the MeNPOC protecting group.
ammonium nitrate in aqueous acetonitrile (Luzzio and Menes, 1994); attempts at hydrogenolytic removal of the benzyl group did not give satisfactory results. The 1,1-dianisyl-2,2,2-trichloroethyl residue (S.109, Fig. 2.3.25) was introduced in the laboratory of Ugi as a protecting group that is stable to acid and base, but is readily removed by reductive fragmentation with the supernucleophile lithium cobalt(I)-phthalocyanine (Karl et al., 1995; Klösel et al., 1996). The application of this group in the preparation of a dinucleoside-trifluoromethyl-phosphonate was demonstrated (Karl et al., 1996). Again, it is beyond the scope of this unit to review all publications that describe the linkage of the 5′ hydroxyl to polymer supports with a non-acid-labile anchor. Examples of anchoring through ester linkage (e.g., Shimidzu and Letsinger, 1968) or, more often, through succinate groups (Ohtsuka et al., 1984; Weiss et al., 1984) are in the literature; in each case, the chain extension was carried out from the 5′ to the 3′ end.
done with acetic anhydride/4-dimethylaminopyridine (e.g., Köster et al., 1983, 1984). In order to blend with a set of protecting groups removable under β-elimination conditions, Reiner and Pfleiderer have proposed a number of β-substituted ethylsulfonylchlorides as capping reagents. The resulting sulfonyl groups are removed with DBU together with the protecting groups for bases and internucleotide linkages, as well as the terminal 5′-NPEOC group (Reiner and Pfleiderer, 1987). In oligonucleotide synthesis with 5′-DMTrprotected phosphoramidite synthons, capping with acetic anhydride activated by 4-dimethylaminopyridine or N-methylimidazole has been the standard procedure from the beginning (Atkinson and Smith, 1984). Because the yields of chain extension in phosphoramidite support synthesis average 98% to 99%, only ∼1% to 2% of unreacted ends are left over for capping NO2 S O2 N
Capping of the 5′-Hydroxy Terminus in Oligonucleotide Synthesis
Protection of 5′-Hydroxy Functions of Nucleosides
Capping refers to the introduction of a protecting group at the 5′-hydroxy function as a postcondensation step during solid-phase oligonucleotide synthesis. The capping step is meant to mask growing chain ends that have escaped elongation and, therefore, will remain as truncated or failure sequences. For this reason, the groups substituted during capping should not be removed before the end of chain growth, if at all. Hence, capping groups should (1) be easily attached even to longer oligonucleotide chains through a highly reactive and sterically unhindered reagent, and (2) be compatible with the chemistry chosen for chain elongation. For solid-phase preparations of oligonucleotides according to the phosphotriester method, phenylisocyanate has been used as a capping reagent in earlier procedures (e.g., Gait et al., 1982). Subsequently, capping was mostly
106
R CH3O 107 R = H 108 R = OCH3
CCl3 CH3O
OCH3 109
Figure 2.3.25 Miscellaneous 5′-hydroxylprotecting groups labile to nonacidic conditions.
2.3.22 Current Protocols in Nucleic Acid Chemistry
during each cycle. The acetylation is generally assumed to be nearly quantitative, but there are no experimental methods to detect average capping yields. Recently, an estimation of capping efficiency has been available through the correlation of calculations using fractal mathematics with the results of capillary electrophoretic separation of the crude product of oligonucleotide preparations of up to 60 bases in length. For support syntheses using the common controlled-pore glass (CPG) or polystyrene supports, the best correlation between calculated and experimentally determined chain length distributions was observed if capping was assumed to proceed with not more than 70% to 80% of the non-elongated chain ends (Földes-Papp et al., 1996, 1998). When the phenoxyacetyl group was used for the protection of exocyclic amino groups, partial replacement of this group by acetyl on the guanine base was observed during conventional capping. This could be avoided by using phenoxyacetic anhydride in place of acetic anhydride (Chaix et al., 1989). The choice of capping procedure necessarily influences the workup of the solid support product. The capping procedures described so far have the common feature that the protecting groups attached to the truncated chains are hydrolyzed on removal of the polymer support and deprotection of bases and internucleotidic bonds. This, again, is prerequisite to the tritylon purifications described in a previous section (see Triarylmethyl Groups as Affinity Ligands). Horn and Urdea have described an alternative route where bases, internucleotidic bonds, and truncated chain ends are deprotected while the crude product—with DMTr on the 5′ end of the full-length oligonucleotide— remains bound to the support. This allows the use of phosphodiesterase to selectively digest the truncated chains and, thus, further simplify the workup (Horn and Urdea, 1985). Other capping alternatives result in an irreversible modification of the truncated chains. This applies to the application of highly reactive phosphite derivatives, for instance, to a mixture of diethoxy-N,N-diisopropylphosphoramidite with tetrazole (Yu et al., 1994), or to diethoxytetrazolophosphane, a stable tetrazole derivative of diethoxyphosphorous acid (Berner et al., 1989). Also, bis(1,1,1,3,3,3hexafluoro-2-propyl)-2-propylphosphite, activated by N-methylimidazole, has been described as a capping reagent to blend with oligonucleotide syntheses using deoxyribonucleoside-3′-bis(1,1,1,3,3,3-hexafluoro-2-pro
pyl)phosphite synthons (Hosaka et al., 1991). In these cases, an adjustment of workup and purification procedures may be required. Recently, a capping procedure that leads to complete reversal of the removal of truncated chains has been described (Natt and Häner, 1997). In this case, capping is done using β-cyanoethoxy-(n-octyloxy)-phosphoramidite/tetrazole as a lipophilic phosphitylating reagent (“lipocap”). If the synthesis is conducted to generate a crude "trityl-off" product, reversedphase high-performance liquid chromatography (RP-HPLC) will lead to a stronger retention of the more lipophilic truncated chains. The target sequence was eluted first and found to be recovered in higher quantity than through “trityl-on” purification. In oligonucleotide syntheses with H-phosphonate intermediates, the average yields of chain elongation are slightly lower due to “selfcapping” with the acyl chloride activating agent. Therefore, an additional capping step is generally unnecessary. Yet, in order to avoid further elongation of reactive truncated chain ends, especially in large-scale preparations, some authors have recommended a capping procedure with, for example, triethylammonium isopropyl phosphite (Andrus et al., 1988) or β-cyanoethyl-H-phosphonate (Gaffney and Jones, 1988).
ENZYMATIC METHODS FOR 5′-HYDROXYL PROTECTION AND DEPROTECTION Compared to the efficiency of automated chemical solid-phase preparation techniques, enzymatic methods play only a minor role in current oligonucleotide synthesis. This is all the more true for 5′-hydroxyl protection or deprotection methods, which are often considered more or less trivial steps in the elongation cycle. Nevertheless, there are some noteworthy arguments in favor of performing protection/deprotection steps via enzyme catalysis. Enzymatic reactions are regio- and stereospecific and often proceed with good yields and high selectivity. Such points are particularly important, for instance, for large-scale oligonucleotide synthesis, an area where enzymatic routes may be worth consideration. The state of enzymatic protecting-group techniques for biomolecules has been reviewed by Waldmann and Sebastian (1994). A specialized survey of enzymatic oligonucleotide acylation/deacylation reactions has recently been published by Prasad and Wengel (1996). A summary of enzymatic reactions leading to the
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.23 Current Protocols in Nucleic Acid Chemistry
introduction of 5′-protecting groups is given in Table 2.3.5. Essentially, two classes of enzymes are known to introduce acyl or alkoxycarbonyl substituents at the 5′-hydroxy function of nucleosides, namely subtilisin and lipases. The best results, so far, were obtained with subtilisin 8350, an enzyme prepared from subtilisin via site-specific mutagenesis. Catalysis of acetyl-group transfer from isopropenyl acetate in anhydrous DMF occurred selectively to the 5′-hydroxy group of various deoxyribo- or ribonucleosides in yields ranging from 65% to nearly 100% (Wong et al., 1990). Somewhat lower yields were found for acyl transfer from trichloro- or trifluoroethylbutyrate to ribonucleosides catalyzed by subtilisin in DMF or pyridine (Riva et al., 1988; Singh et al., 1994). Lipases from various sources have alternatively been applied, but the yields are generally lower and the conditions must be chosen very carefully to ensure the introduction of acyl or alkoxycarbonyl groups selectively or predominantly at the 5′-hydroxy position (Nozaki et al., 1990; Moris and Gotor, 1992a,b, 1993; GarciaAlles et al., 1993; Garcia-Alles and Gotor, 1995; Ozaki et al., 1995). Enzyme-catalyzed deacylations were of interest in early times of nucleotide chemistry, when an enzymatic approach to oligonucleotide chain lengthening seemed to be a feasible alternative to the still cumbersome methods of phosphodiester chemistry. The dihydrocinnamoyl protecting group received some attention at that time, because 3′-dihydrocinnamoyl-nucleoside-5′-diphosphates were dis-
covered as monoaddition substrates for polynucleotide phosphorylase (Kaufmann et al., 1971), and the same protecting group could be removed by chymotrypsin (Sachdev and Starkovsky, 1969; Taunton-Rigby, 1973). If, however, a 3′,5′-disubstituted nucleoside derivative was the substrate, a distinct preference for the hydrolysis of the 5′-O-acyl group was demonstrated (Taunton-Rigby, 1973). The regioselective 5′-O-deacetylation of 2′,3′,5′-triacetyl nucleosides was described by Singh et al. (1993). In recent times, the tendency has been to reverse the action of the before-mentioned enzymes, subtilisin and lipase. Whereas lipase from Pseudomonas fluorescens has a preference for the removal of secondary O-acyl groups, subtilisin attacks first the acyl substituent on the 5′-hydroxy moiety (Uemura et al., 1989a,b). In both cases, however, the reaction can proceed to complete deprotection. Selective deprotection of 5′-O-acyl was found with lipase from porcine pancreas, which yielded 3′-O-acetylthymidine from the 3′,5′-disubstituted nucleoside in 98% yield (Wong et al., 1990). Similarly, 2′,3′,5′-tri-O-acetylated pyrimidine and purine ribonucleosides (A, C, G, U, and modified bases) were selectively 5′-deprotected by subtilisin to give 2′,3′-di-Oacyl nucleosides in 40% to 92% yield (Singh et al., 1993). In general, it appears that the potential of enzymatic protection/deprotection reactions for the 5′-hydroxy function remains to be further exploited. The isolation of enzymes from different sources, as well the adaptation of enzyme specificity through mutagenesis, protein
Table 2.3.5 Enzymatic Introduction of Protecting Groups at the 5′-Hydroxy Function
Acyl group transferred Butyryl Butyryl Acetyl Acyl Alkoxycarbonyl Protection of 5′-Hydroxy Functions of Nucleosides
Acyl
Enzyme Subtilisin
Substrate acyl donor
Trichloroethylbutyrate in DMF Subtilisin Trifluoroethylbutyrate in pyridine Subtilisin 8350 Isopropenyl acetate
Product
5′-O-Acyl predominant 67%-82% 5′-O-acyl 80%-100% 5′-O-acyl Lipase SP 435 Alkane carbonyl oxime 5′-O-Acyl esters r(d)A, G, T, U (Candida antarctica) Lipase SP 435 Alkoxy carbonyl oxime 5′-O-Alkoxy(Candida esters r(d)A, G, T, U carbonyl antarctica) 31%-75% Pentanoic acid Lipase 5′-O-Acyl (Pseudomonas anhydride, preferably dC fluorescens)
Reference Riva et al. (1988) Singh et al. (1994) Wong et al. (1990) Moris and Gotor (1992a,b, 1993) Moris and Gotor (1992a,b, 1993) Nozaki et al. (1990)
2.3.24 Current Protocols in Nucleic Acid Chemistry
engineering, and in vitro evolution and selection techniques, may provide tools tailored to perform highly specific reactions and, thus, to match chemical methods for introduction and removal of protecting groups.
PROTECTION OF 5′-HYDROXY FUNCTIONS: REMAINING PROBLEMS, CONSIDERATIONS, AND OPTIONS Protection and deprotection of the 5′-hydroxy group remain essential operations for oligonucleotide synthesis, independent of the chemistry used for the formation of the internucleotidic bond. To many researchers involved in routine small-scale oligonucleotide synthesis, questions connected with 5′-hydroxyl-protecting groups appear to be solved, although the protection chemistry used in >95% of all syntheses is now 35 years old. Failures that arise through incomplete deprotection or capping during the elongation cycles have been discussed above. Many attempts have been made to minimize these failures through the design of new protecting groups for the 5′ hydroxyl, nucleobases, or other functional groups of the oligonucleotide, or through the development of an altogether new orthogonal protection scheme. However, no method exists, so far, that allows detection and quantitative analysis of, for example, small numbers of depurination sites statistically distributed within a population of long-chain oligonucleotide molecules. Techniques still have to be developed that might help assess such homogeneity problems, which are of importance especially when oligonucleotides are produced in large numbers, large scale, or extended length. Generally, more consideration should be given to the avoidance of acid deprotection steps. A particularly interesting new deprotection chemistry comes from the application of photolabile protecting groups. Considerable efforts are still necessary to establish quantitative and selective removal of such groups, and new developments are under way at several laboratories. For solid-phase bulk synthesis, other deprotection alternatives that use neither acid nor alkaline/ammoniacal media may have to be investigated. The availability of silyl protecting groups that can be released in <1 min by fluoride treatment at room temperature may find application beyond oligoribonucleotide synthesis. Recent years have seen rapid progress in the preparation of structurally modified oligonucleotides, among them structures highly sensitive to acid or base. These syntheses often
require a new kind of protection chemistry; such developments may feed back into routine preparations of unmodified oligonucleotides. One of the reasons that trityl groups have been the first choice for several decades is that, beyond their application for the protection of the growing chain, they can be endowed by appropriate substitution with additional properties useful for oligonucleotide synthesis and purification. Examples—such as the monitoring of chain extension yields, the simplification of the separation of support products, or the activation of a 3′-phosphate moiety—have been discussed above. There will be a general interest to also endow non-acid-labile protecting groups with similar properties; examples are in the literature, but there are still options for new developments. Such projects may also yield new solutions to simple practical questions, e.g., the choice of nontoxic, environmentally unproblematic coupling and deprotection solvents. The development of synthetic oligonucleotides that are ready to move into the drug market has led to the budding of a new field: the technology of industrial oligonucleotide production. In this frame, protection and deprotection steps are part of a series of unit operations that have to be carefully analyzed and optimized with respect to technical performance. In the previous sections, reference has occasionally been given to investigations that have been performed on this technical background, with the idea in mind that these investigations may well add new experience to small-scale routine synthesis as well. Future reviews may have to give broader attention to technical aspects as this field develops into a science of its own. In this context, it is interesting to see that oligonucleotide synthesis in the past years has developed in the direction of ever new chemistry, while chemical production in general looks more and more to biotechnology as a tool to solve problems connected with the utilization of fossil carbon sources, environmental pollution, or waste disposal. The impact of such problems is already recognized and will be increasingly felt as oligonucleotide synthesis moves more towards bulk production. Enzymatic methods, which may be in the foreground of future developments, necessarily require a new approach to questions of protection and deprotection.
Protection of Nucleosides for Oligonucleotide Synthesis
2.3.25 Current Protocols in Nucleic Acid Chemistry
LITERATURE CITED Adams, S.P., Kavka, K.S., Wykes, E.J., Holder, S.B., and Galluppi, G.R. 1983. Hindered dialkylamino nucleoside phosphite reagents in the synthesis of two DNA 51-mers. J. Am. Chem. Soc. 105:661663. Andersen, W., Hayes, D.H., Michelson, A.M., and Todd, A.R. 1954. Deoxyribonucleosides and related compounds. Part IV. The configuration at the glycosidic centre in deoxyadenosine and deoxycytidine. J. Chem. Soc. 1882-1887. Andrus, A., Efcavitch, J.W., McBride, L.J., and Giusti, B. 1988. Novel activating and capping reagents for improved hydrogen-phosphonate DNA synthesis. Tetrahedron Lett. 29:861-864. Atkinson, T. and Smith, M. 1984. Solid-phase synthesis of oligodeoxyribonucleotides by the phosphate triester method. In A Practical Approach to Oligonucleotide Synthesis (M.J. Gait, ed.) pp. 35-81. IRL Press, Oxford. Balgobin, N. and Chattopadhyaya, J.B. 1982a. Two sulfur containing protecting groups for alcoholic hydroxyl function. Chem. Scr. 19:143-144. Balgobin, N. and Chattopadhyaya, J.B. 1982b. An efficient chemical synthesis of a biologically functional DNA molecule, 5′d(A-T-G-G-G-T-TT-C-T-T-C-G-C-)3′, through the phospho-triester approach. Chem. Scr. 20:133-138. Balgobin, N. and Chattopadhyaya, J. 1987. Solid phase synthesis of DNA under a non-depurinating condition with a base labile 5′-protecting group (Fmoc) using phosphiteamidite approach. Nucleosides Nucleotides 6:461-463. Balgobin, N., Josephson, S., and Chattopadhyaya, J.B. 1981a. The 2-phenylsulfonylethylcarbonyl (PSEC) group for the protection of the hydroxyl function. Tetrahedron Lett. 22:3667-3670. Balgobin, N., Josephson, S., and Chattopadhyaya, J.B. 1981b. A general approach to the chemical synthesis of oligodeoxyribonucleotides. Acta Chem. Scand. B 35:201-212. Bannwarth, W. and Wippler, J. 1990. A new combined purification/phosphorylation procedure for oligodeoxynucleotides. Helv. Chim. Acta 73:1139-1147. Bazin, H., Heikkilä, J., and Chattopadhyaya, J. 1985. Some aspects of the reaction of arenesulfenyl chlorides with hydroxyl functions of ribonucleosides. Acta Chem. Scand. B 39:391400. Beaucage, S.L. and Iyer, R.P. 1992. Advances in the synthesis of oligonucleotides by the phosphoramidite approach. Tetrahedron 48:22232311. Protection of 5′-Hydroxy Functions of Nucleosides
Beaucage, S.L. and Iyer, R.P. 1993. The synthesis of modified oligonucleotides by the phosphoramidite approach and their applications. Tetrahedron 49:6123-6194.
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Protection of Nucleosides for Oligonucleotide Synthesis
2.3.27 Current Protocols in Nucleic Acid Chemistry
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2.3.28 Current Protocols in Nucleic Acid Chemistry
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Kawashima, E., Aoyama, Y., Sekine, T., Miyahara, M., Radwan, M.F., Nakamura, E., Kainosho, M., Kyogoku, Y., and Ishido, Y. 1995. Sonochemical and triethylborane-induced tin deuteride reduction for the highly diastereoselective synthesis of (2′R)-2′-deoxy[2′-2H]ribonucleoside derivatives. J. Org. Chem. 60:6980-6986. Kawashima, E., Toyama, K., Ohshima, K., Kainosho, M., Kyogoku, Y., and Ishido, Y. 1997. Novel approach to diastereoselective synthesis of 2′-deoxy[ 5′-2H1]ribonucleoside derivatives by reduction of the corresponding 5′-Oacetyl-2′-deoxy-5′-phenylselenoribonucleoside derivatives with a Bu3Sn2H-Et3B system. Chirality 9:435-442. Kierzek, R., Ito, H., Bhatt, R., and Itakura, K. 1981. Selective N-deacylation of N,O-protected nucleosides by zinc bromide. Tetrahedron Lett. 22:3761-3764. Klösel, R. König, S., and Lehnhoff, S. 1996. The 1,1-dianisyl-2,2,2-trichloroethyl group as 2′hydroxyl protection of ribonucleotides. Tetrahedron 52:1493-1502. Kohli, V., Blöcker, H., and Köster, H. 1980. The triphenylmethyl (trityl) group and its uses in nucleotide chemistry. Tetrahedron Lett. 21:2683-2686. Kössel, H. and Seliger, H. 1975. Recent advances in polynucleotide synthesis. Fortschr. Chem. Org. Naturst. 32:297-508. Köster, H. and Sinha, N.D. 1982. Dialkyl aluminium chloride: A reagent for removal of trityl group from trityl ethers of deoxynucleosides, deoxynucleotides, and oligodeoxynucleotides. Tetrahedron Lett. 23:26412644. Köster, H., Stumpe, A., and Wolter, A. 1983. Polymer support oligonucleotide synthesis 13: Rapid and efficient synthesis of oligodeoxynucleotides on porous glass support using triester approach. Tetrahedron Lett. 24:747750. Köster, H., Biernat, J., McManus, J., Wolter, A., Stumpe, A., Narang, C.K., and Sinha, N.D. 1984. Polymer support oligonucleotide synthesis. XV. Synthesis of oligodeoxynucleotides on controlled pore glass (CPG) using phosphate and a new phosphite triester approach. Tetrahedron 40:103-112. Kotschi, U. 1987. Präparative und apparative Beiträge zur Festphasensynthese von Nucleinsäure-Fragmenten. Dissertation, University of Ulm, Germany. Krotz, A.H., Cole, D.L., and Ravikumar, V.T. 1999. Dimethoxytrityl removal in organic medium: Efficient oligonucleotide synthesis without chlorinated solvents. Nucleosides Nucleotides 18:1207-1209. Protection of Nucleosides for Oligonucleotide Synthesis
2.3.29 Current Protocols in Nucleic Acid Chemistry
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Protection of 5′-Hydroxy Functions of Nucleosides
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Protection of Nucleosides for Oligonucleotide Synthesis
2.3.31 Current Protocols in Nucleic Acid Chemistry
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Protection of Nucleosides for Oligonucleotide Synthesis
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Contributed by H. Seliger University of Ulm Ulm, Germany
Protection of 5′-Hydroxy Functions of Nucleosides
2.3.34 Current Protocols in Nucleic Acid Chemistry
A Base-Labile Protecting Group (Fluorenylmethoxycarbonyl) for the 5′-Hydroxy Function of Nucleosides
UNIT 2.4
Once the exocyclic amino functions of a nucleoside have been protected (UNIT 2.1), further functionalization aimed towards assembly into oligonucleotides requires consideration of the specific regioselective protection of the hydroxy groups present in the monomeric building block. Some years ago it was found that, in the case of 2′-deoxynucleosides, the two hydroxy groups (3′- and 5′-) could be satisfactorily differentiated by the application of the triarylmethyl (trityl) family of acid-labile protecting groups for temporary protection of the primary 5′-hydroxy function (UNIT 2.3). In the case of the ribonucleoside series, however, the differentiation of the three hydroxy groups (2′-, 3′-, and 5′-) has to be achieved at the extremely high level of efficiency required for the iterative steps involved in oligonucleotide assemblies carried out on synthesizer machines. This increased complexity has led to a number of alternative synthetic pathways to oligoribonucleotides, some of which are now commercialized; other pathways remain to be explored more profoundly. Most of the present oligoribonucleotide synthesis strategies adhere to the popular 5′-Otrityl chemistry and aim at finding solutions for 2′-O-protection (UNIT 2.2), such as the t-butyldimethylsilyl (TBDMS) group, which can be removed by treatment with fluoride ion. In contrast, the authors chose to take advantage of the well-established ketal-type group for 2′-O-protection, which allows final deprotection under particularly mild and isomerization-free acidolytic aqueous conditions at the end of the synthesis (UNIT 2.2), and investigated for 5′-protection the base-labile 9-fluorenylmethoxycarbonyl (FMOC) group as a valuable orthogonal complement. The following Basic Protocol presents a reliable procedure for the regioselective introduction of the FMOC group at the 5′-hydroxy position of nucleosides, starting with deoxythymidine or with 4-methoxytetrahydropyran-4-yl (MTHP)–protected ribonucleosides. The Basic Protocol includes steps for the isolation and characterization of all four 5′-O-FMOC-2′-O-MTHP-protected ribonucleosides and of 5′-O-FMOC-2′-deoxythymidine, which serves as a reference for the deoxyribonucleoside series. No modification is required for 5′-O-FMOC-2′-dABz, -dCBz, or -dGi-Bu (yields are ~65% for dGi-Bu and 70% to 80% for dABz and dCBz; Balgobin and Chattopadhyaya, 1987; Ma and Sonveaux, 1987). Additionally, four Support Protocols describe the preparation of the starting N-protected-2′-O-MTHP-ribonucleosides, which are not commercially available and are considered key intermediates for the preparation of the 5′-O-FMOC-protected ribonucleoside building blocks. Characterization of the various products requires experience in thin-layer chromatography, short column (flash) chromatography, and proton NMR. ACYLATION OF THE 5′-HYDROXY GROUP OF 2′-O-(4-METHOXYTETRAHYDROPYRAN-4-YL)-RIBONUCLEOSIDES WITH 9-FLUORENYLMETHOXYCARBONYL CHLORIDE
BASIC PROTOCOL
The procedure describes the acylation of the 5′-hydroxy function of N-base- and 2′-Oprotected ribonucleosides with 9-fluorenylmethoxycarbonyl (FMOC) chloride in pyridine at moderately low temperature. The FMOC workup procedure and the chromatographic isolation and 1H-NMR spectroscopic characterization of the resulting 5′-O-(9-fluorenylmethyl)carbonates S.2a to S.2e (Fig. 2.4.1) are presented in this protocol. The starting intermediates are the four 2′-O-MTHP-protected ribonucleotides rUMTHP, rCBzMTHP, rABzMTHP, and rGi-BuMTHP (S.1a to S.1d; Fig. 2.4.1 and Fig. 2.4.2) and
Protection of Nucleosides for Oligonucleotide Synthesis
Contributed by Michael J. Gait and Christian Lehmann
2.4.1
Current Protocols in Nucleic Acid Chemistry (2000) 2.4.1-2.4.22 Copyright © 2000 by John Wiley & Sons, Inc.
HO
O
4' 3'
HO
[Fmoc]
B
5'
1' 2'
O
Fmoc chloride OCH3
O
pyridine, 0ºC
B
O
O
O
O HO
[Mthp]
O
OCH3
O 1a: 1b: 1c: 1d: 1e:
B = U = uracil-1-yl Bz 4 B = C = N -benzoylcytosin-1-yl Bz 6 B = A = N -benzoyladenin-9-yl -Bu 2 B = Gi = N -isobutyrylguanin-9-yl B = T = thymin-1-yl; 2'-deoxy
2a: 2b: 2c: 2d: 2e:
B=U Bz B=C Bz B=A -Bu B = Gi B = T; 2'-deoxy
Figure 2.4.1 General procedure for the preparation of 5′-O-fluorenylmethoxycarbonyl-2′-O-(4methoxytetrahydropyran-4-yl)-ribonucleosides (S.2a-d) and of 5′-O-fluorenylmethoxycarbonyl-2′deoxythymidine (S.2e).
deoxythymidine (dT; S.1e; Fig. 2.4.1). Support Protocols 1 to 4 provide a summary for the preparation of the starting key intermediates S.1a to S.1d. Materials 2′-Deoxythymidine or 2′-O-MTHP-ribonucleoside: 2′-O-(4-methoxytetrahydropyran-4-yl)-rU, -rCBz, -rABz, or -rGi-Bu (see Support Protocols 1 to 4) Dry pyridine, freshly distilled from calcium hydride (5 g/L) after refluxing for 2 hr under an inert atmosphere Nitrogen gas 9-Fluorenylmethoxycarbonyl (FMOC) chloride (Cambridge Research Biochemicals), recrystallized from ether/pentane in a large desiccator (vapor diffusion method) Diethyl ether (analytical grade) Chloroform stabilized with 1% ethanol (commercially available analytical grade) Methanol (analytical grade) Anisaldehyde reagent (see recipe) Ethane-1,2-diol (analytical grade) Saturated aqueous sodium bicarbonate solution Sodium sulfate (anhydrous) Toluene (analytical grade) Ethanol (analytical grade) Pentane (analytical grade) Silica-coated thin-layer chromatography (TLC) plate with fluorescent indicator Kieselgel 60F254 (Merck 5554 glass plates or 5744 aluminium foils) UV light source D4 glass-filter crucible Short column (diameter 5 cm; length ~10 cm) containing 50 g Kieselgel 60H without calcium sulfate (Merck 7736 or Fluka 60770, particle size 5-40 µm; or Merck 11677, particle size 15 µm), preconditioned with chloroform Glass-fiber tissue A Base-Labile Protecting Group
CAUTION: Wear gloves and perform all operations involving TLC solvents and reagents in a well-ventilated fume hood.
2.4.2 Current Protocols in Nucleic Acid Chemistry
[Bz]
O HN
O 4 5
2
6
N1
O
4'
6
O
O
4'
2'
3'
5'
HO
1'
HO
3
5
NH
5'
HO
4
3
2
O
N1 1' 2'
3'
HO
OMthp
N
OMthp
1a
1b
from uridine
from cytidine
O HN
7
6
N 5 5'
O
4' 3'
HO
N 4 N 9 1'
1
2
3
2'
5'
O
4' 3'
OMthp
6 1
N 5
NH
8
HO
HO
[i-Bu]
O
7
N
8
HO
[Bz]
N 4 N 9 3
1'
O
2
N H
2'
OMthp
1c
1d
from adenosine
from guanosine
Figure 2.4.2 2′-O-(4-Methoxytetrahydropyran-4-yl)-ribonucleosides used as starting intermediates for FMOC protection. Preparation of these intermediates is shown in Figures 2.4.3 to 2.4.6.
Introduce FMOC group 1. Coevaporate 3.0 mmol of 2′-O-MTHP-ribonucleoside or deoxythymidine derivative (S.1a to S.1e) twice from 25 mL dry pyridine under reduced pressure using a rotary evaporator, and then apply a dry nitrogen atmosphere. 2. Dissolve the substrate in a third 25-mL portion of dry pyridine and cool to 0°C. 3. Stir the solution and add 3.6 mmol crystalline FMOC chloride. Stir under nitrogen for 30 min at 0°C. Normally, all the FMOC chloride will have dissolved by this time.
Check reaction by TLC 4. Resolve products on a silica-coated TLC plate with fluorescent indicator. Develop with diethyl ether followed by 9:1 (v/v) chloroform/methanol. Predevelopment in diethyl ether is used to disperse the pyridine.
5. Visualize under UV light (254 nm). 6. Stain by spraying with anisaldehyde reagent and heating on a hot plate at 80°C. Coloring plates containing nucleoside materials (summarized in Jork et al., 1990) is necessary for compounds that do not absorb strongly in the 254-nm region (i.e., unprotected rUMTHP and dT). TLC shows complete conversion into products with higher Rf values than the starting nucleosides. The major component is the 5′-O-FMOC derivative (see respective Rf values for each nucleoside following step 18; when starting with S.1b, the 5′-derivative Rf = 0.54). A very minor component at slightly higher Rf is identified as the 3′-O-FMOC derivative
Protection of Nucleosides for Oligonucleotide Synthesis
2.4.3 Current Protocols in Nucleic Acid Chemistry
(Rf = 0.68 when starting with S.1b). Some 3′,5′-di-O-FMOC derivative can be found at a very much higher Rf (0.80 when starting with S.1b). If complete conversion is not observed, the reaction in step 3 can be continued. It should not exceed 2 hr, however, as this leads to increased production of the 3′,5′-di-O-FMOC derivative.
Isolate product by chromatography 7. Terminate the reaction by adding 200 µL of ethane-1,2-diol and concentrate the mixture to an oil using a rotary evaporator with a vacuum pump. 8. Dissolve the oil in 150 mL chloroform and wash with 50 mL saturated aqueous sodium bicarbonate solution. Remove the aqueous phase and wash twice with 50 mL chloroform. 9. Dry the combined chloroform extracts over ~20 g anhydrous sodium sulfate. 10. Filter the dried extract under vacuum with a D4 glass filter crucible, and concentrate under reduced pressure (step 7). 11. Coevaporate the residual oil from ∼50 mL each toluene (twice), ethanol (once), and chloroform (once). This removes traces of pyridine that are still contained in the dried, concentrated extract (step 10).
12. Apply as concentrated as possible (i.e., in ∼5 mL) to a short 50-g Kieselgel 60H column preconditioned with chloroform. Apply a circular piece of glass-fiber tissue (not sea sand) to the top of the column. Short-column chromatography is performed according to Hunt and Rigby (1967) and is similar to the flash chromatography described by Still et al. (1978).
13. Elute minor side products first with chloroform (first 1 to 2 column volumes; 20-mL fraction size), then elute the major product-containing fractions with a gradient of 1% to 5% (v/v) ethanol/chloroform. Do not use >1 atm pressure for elution (Hunt and Rigby, 1967; Still et al., 1978). 14. Combine the product-containing fractions and concentrate to a foam. 15. Dissolve foam in 5 mL chloroform, precipitate by dropwise addition to 500 mL rapidly stirring pentane, and decant off most of the supernatant (∼450 mL). 16. Collect solids by centrifuging 10 min at 3000 rpm g, 15°C, and decant the pentane. 17. Wash twice by resuspending in 50 mL fresh pentane and repeating the centrifugation. Dry the resulting white powder (S.2a to S.2e) in vacuo using a high vacuum from an oil pump. Characterize the product 18. Characterize the final product by TLC and 1H-NMR. Chemical characterization data is provided below for S.2a to S.2e. 1H-NMR spectra (250 MHz) were measured on a Bruker WM 250 instrument in DMSO-d6. Chemical shifts (δ) are given in ppm downfield from tetramethylsilane (TMS, 0 ppm). Coupling constants (J) are given in Hz. D2O exchange was performed on all samples.
A Base-Labile Protecting Group
5′-O-FMOC-2′-O-MTHP-uridine (S.2a): yield 64%; Rf (silica, chloroform/methanol 9:1): 0.43; 1H-NMR (250 MHz, DMSO-d6): 1.6-1.9 (m×m, 4H; -CH2-(3,5) of MTHP), 2.98 (s, 3H; H3CO- of MTHP), 3.3-3.7 (m×m, 4H; -CH2-(2,6) of MTHP), 3.9-4.6 (m×m, 8H; -CH-(4′, 3′, 2′), -CH2-(5′), fluorenyl-H9 and -CH2-), 5.4 (dbr, 1H, exchangeable with D2O;
2.4.4 Current Protocols in Nucleic Acid Chemistry
3′-OH), 5.63 (d, J = 8.0 , 1H; uridyl-H5), 5.99 (d, J = 6.3, 1H; -CH-(1′)), 7.29-7.95 (m×m, 9H; uridyl-H6, fluorenyl-aromatic H), 11.4 (sbr, 1H, exchangeable; uridyl-N3-H). 5′-O-FMOC-2′-O-MTHP-4-N-benzoyl-cytidine (S.2b): yield 61%; Rf (silica, chloroform/methanol 9:1): 0.54; 1H-NMR (250 MHz, DMSO-d6): 1.5-1.9 (m×m, 4H; -CH2-(3,5) of MTHP), 2.95 (s, 3H; H3CO- of MTHP), 3.2-3.7 (m×m, 4H; -CH2-(2,6) of MTHP), 4.00-4.35 (m×m, 5H; -CH-(4′, 3′, 2′), -CH2-(5′)), 4.47 (t, J = 6.0, 1H; fluorenyl-H9), 4.54 (d, J = 6.0, 2H; fluorenyl-CH2-), 5.4 (d, J = 5, 1H, exchangeable with D2O; 3′-OH), 6.09 (d, J = 6.4, 1H; -CH-(1′)), 7.3-8.3 (m×m, 15H; cytidyl-H5 and -H6, fluorenyl- and benzoyl-aromatic H), 11.3 (sbr, 1H, exchangeable; cytidyl-N4-H). 5′-O-FMOC-2′-O-MTHP-6-N-benzoyl-adenosine (S.2c): yield 65%; Rf (silica, chloroform/methanol 9:1): 0.56; 1H-NMR (250 MHz ,DMSO-d6): 1.4-1.9 (m×m, 4H; -CH2-(3,5) of MTHP), 2.62 (s, 3H; H3CO- of MTHP), 3.2-3.7 (m×m, 4H; -CH2-(2,6) of MTHP), 4.18-4.48 (m×m, 4H; -CH-(4′, 3′), -CH2-(5′)), 4.30 (t, J = 6.0, 1H; fluorenyl-H9), 4.51 (d, J = 6.0, 2H; fluorenyl-CH2-), 5.08 (d×d, J2′-1′ = 6.8, J2′-3′ = 5.0; 1H; -CH-(2′)), 5.6 (d, J = 4.5, 1H, exchangeable with D2O; 3′-OH), 6.21 (d, J = 6.8, 1H; -CH-(1′)), 7.2-8.4 (m×m, 13H; fluorenyl- and benzoyl-aromatic H), 8.73 and 8.74 (2s, 2H; adenyl-H2 and H8), 11.3 (sbr, 1H, exchangeable; adenyl-N6-H). 5′-O-FMOC-2′-O-MTHP-2-N-isobutyryl-guanosine (S.2d): yield 52%; Rf (silica, chloroform/methanol 9:1): 0.37; 1H-NMR (250 MHz, DMSO-d6): 1.12 (2d, J = 7.0, 6H; (H3C)2CH-)), 1.4-1.9 (m×m, 4H; -CH2-(3,5) of MTHP), 2.67 (s, 3H; H3CO- of MTHP), 2.75 (septet, J = 7.0, 1H; (H3C)2CH-)), 3.2-3.7 (m×m, 4H; -CH2-(2,6) of MTHP), 4.12-4.37 (m×m, 4H; -CH-(4′, 3′), -CH2-(5′)), 4.34 (m(t), 1H; fluorenyl-H9), 4.53 (d, J = 6.0, 2H; fluorenyl-CH2-), 4.81 (d×d, J2′-1′ = 7.5, J2′-3′ = 4.7, 1H; -CH-(2′)), 5.4 (d, J = 4, 1H, exchangeable with D2O; 3′-OH), 5.99 (d, J = 7.5, 1H; -CH-(1′)), 7.2-8.0 (m×m, 8H; fluorenyl-aromatic H), 8.25 (s, 1H; guanyl-H8), 11.6 and 12.1 (2sbr, 2H, exchangeable; guanyl-N1-H and N2-H). 5′-O-FMOC-2′-deoxythymidine (S.2e): obtained in analytically pure form by subsequent crystallization from ethyl acetate (76%; mp. 183°-185°C (decomp.); calc.: C 64.65, H 5.21, N 6.03%; found: C 64.54, H 5.09, N 5.82%); Rf (silica, chloroform/methanol 9:1): 0.44; 1 H-NMR (250 MHz, DMSO-d6): 1.69 (s, 3H; thymidyl-5-CH3), 2.10 (m, 2H; -CH2-(2′)), 3.90 (m, 1H; fluorenyl-H9), 4.10-4.40 (m×m, 4H; -CH-(4′, 3′), -CH2-(5′)), 4.58 (m(d), 2H; fluorenyl-CH2-), 5.44 (d, J = 4.4, 1H, exchangeable with D2O; 3′-OH), 6.17 (t, J = 6.3, 1H; -CH-(1′)), 7.29-7.91 (m×m, 9H; thymidyl-H6, fluorenyl-aromatic H), 11.3 (sbr, 1H, exchangeable; thymidyl-N3-H).
PREPARATION OF 2′-O-(4-METHOXYTETRAHYDROPYRAN-4-YL)URIDINE (S.1a) FROM URIDINE
SUPPORT PROTOCOL 1
Starting from uridine (S.3; Fig. 2.4.3), the desired key intermediate S.1a is obtained by a series of well-described reactions that handle the three ribonucleoside hydroxy groups differentially without protection of the base (Fromageot et al., 1967; Reese et al., 1970; van Boom and Wreesmann, 1984). Chromatography is necessary only in the last step of the sequence, which makes the procedure particularly useful for the preparation of large amounts of material. In the first step, reaction of uridine (S.3) with trimethyl orthoacetate gives the 2′,3′-methoxyethylidene derivative S.4, which is then acylated at the 5′-OH group to yield S.5. This derivative is cleaved by partial acidic hydrolysis to give a mixture of 2′,5′-di-O- and 3′,5′-di-O-acetates, of which the latter (S.6) can be fractionally crystallized in good yield. Acid-catalyzed ketalization with 5,6-dihydro-4-methoxy-2Hpyran followed by ammonolysis subsequently gives 2′-O-MTHP-uridine (S.1a). Additional Materials (also see Basic Protocol) Uridine (Sigma or Fluka), dried before use for 2 hr at 50°C over phosphorus pentoxide in vacuo Toluene-p-sulfonic acid monohydrate (analytical grade)
Protection of Nucleosides for Oligonucleotide Synthesis
2.4.5 Current Protocols in Nucleic Acid Chemistry
O
O 4
3
5 6 5'
HO
O
4'
O NH
NH
NH
2
N1
O
steps 1-2
N
HO
O
steps 3-9
1'
N
AcO
O
O
O
2'
3'
HO
O
OH
O
H3C
3
O H3C
OCH3
5
4
O
O
NH
NH steps 10-19
N
AcO
O
steps 20-31
O AcO
O OCH3
N
HO
HO
OH
O
O O
OCH3
O
6 1a
Figure 2.4.3 Scheme showing the preparation of 2′-O-MTHP-uridine (S.1a) from uridine (see Support Protocol 1).
Dioxane (analytical grade), dried by keeping over activated basic aluminum oxide, then distilled before use from sodium/benzophenone after refluxing until the blue color indicates complete dryness Trimethyl orthoacetate (purum, 97%), dried by refluxing over calcium hydride (5 g/L) and distilled under inert atmosphere Pyridine (analytical grade), dried by refluxing over calcium hydride (5 g/L) and then distilled under inert atmosphere Acetic anhydride (analytical grade) Methylene chloride (analytical grade) Saturated (~1 M) aqueous sodium hydrogen carbonate Magnesium sulfate Formic acid (analytical grade) 5,6-Dihydro-4-methoxy-2H-pyran (analytical grade; Sigma or Fluka pract., ~90%; b.p.20 59° to 61°C) Half-saturated methanolic ammonia solution (see recipe) 500-mL separatory funnel Prepare 2′,3′-methoxyethylidene derivative (S.4) 1. Prepare a mixture of 24.2 g dry uridine (S.3; 100 mmol) 100 mg toluene-p-sulfonic acid monohydrate 150 mL dry dioxane 17 mL dry trimethyl orthoacetate and stir for 6 hr at room temperature. 2. Check the reaction by TLC (see Basic Protocol, steps 4 to 6). A Base-Labile Protecting Group
TLC should show complete conversion of the starting material (Rf = 0.04) to the less polar cyclic orthoester S.4 (Rf = 0.38).
2.4.6 Current Protocols in Nucleic Acid Chemistry
Prepare 5′-acetylated derivative (S.5) 3. Combine 150 mL dry pyridine and 50 mL acetic anhydride. Add this acylating solution dropwise over 15 min at room temperature to the stirred reaction mixture (step 1), and leave overnight. 4. Check the reaction by TLC. TLC should show that the acetylation has gone to completion to yield a product (S.5) with higher mobility (Rf = 0.63).
5. Add 50 mL methanol dropwise to the reaction mixture at 0°C to destroy excess acetic anhydride. 6. Concentrate the mixture to a colorless oil under reduced pressure using a rotary evaporator. 7. Dissolve in 250 mL methylene chloride and wash with 100 mL of 1 M aqueous sodium hydrogen carbonate. 8. Extract the aqueous layer five times with 100 mL methylene chloride for each extraction. 9. Dry the combined organic layers over 25 g magnesium sulfate and concentrate to a thick oil. Prepare 3′,5′-di-O-acetyluridine (S.6) 10. Coevaporate the resulting oil twice from 100 mL ethanol to remove traces of pyridine. 11. Dissolve S.5 in 150 mL of 9:1 (v/v) formic acid/water and incubate 15 min at room temperature. 12. Check the reaction by TLC. TLC should show that the cleavage of the methoxyethylidene function is complete, yielding 2′,5′-di-O- and 3′,5′-di-O-acetyluridines (Rf ≈ 0.4).
13. Evaporate the aqueous formic acid under reduced pressure with a rotary evaporator, and coevaporate the residue twice with 100 mL toluene and twice with 100 mL ethanol. 14. Dissolve in 250 mL chloroform and transfer to a 500-mL separatory funnel containing 150 mL saturated aqueous sodium hydrogen carbonate solution. Shake carefully and separate out the chloroform layer. 15. Extract the aqueous layer twelve times with 50-mL portions of chloroform. 16. Dry the combined chloroform extracts over ~80 g anhydrous sodium sulfate, filter (see Basic Protocol, step 10), and evaporate to a glass. 17. Dissolve the crude product in 500 to 1000 mL boiling ethanol and store the solution for 2 days at room temperature. 18. Filter off the crystals and dry at high vacuum. 19. Determine the isomeric purity by 1H-NMR spectroscopy. Recrystallize the product if the 2′,5′-isomer is present. 3′,5′-Di-O-acetyluridine (S.6): yield 23.9 g (73% from S.3; m.p. = 152°-155°C); Rf (silica, chloroform/methanol 9:1): 0.42; 1H-NMR (250 MHz, DMSO-d6 /D2O, 1:1, containing three drops of acetic acid per mL): 7.57 (d, J = 7.5, 1H; uridyl-H6), 5.89 (d, J = 7.5, 1H; uridyl-H5), 5.88 (d, J = 5.5, 1H; -CH-(1′)); the 2′,5′-isomer shows a slightly different
Protection of Nucleosides for Oligonucleotide Synthesis
2.4.7 Current Protocols in Nucleic Acid Chemistry
1
H-NMR spectrum (same conditions): 7.57 (d, J = 7.5, 1H; uridyl-H6), 5.89 (d, J = 7.5, 1H; uridyl-H5), 5.86 (d, J = 6.0, 1H; -CH-(1′)).
Prepare 2′-O-MTHP-uridine (S.1a) 20. Combine 138 mg toluene-p-sulfonic acid monohydrate and 250 mL dry dioxane and stir until clear. 21. Add 32.8 g of 3′,5′-di-O-acetyluridine (S.6; 100 mmol) and 57 g of 5,6-dihydro4-methoxy-2H-pyran (57 g, 5.0 eq) to the mixture and stir for 20 hr at room temperature. 22. Check the reaction by TLC. TLC should indicate complete reaction of the starting material.
23. Cool the clear solution in an ice water bath. The solution should be below 0°C to absorb exothermicity during neutralization.
24. Neutralize the reaction mixture by adding 800 µl of half-saturated methanolic ammonia solution. 25. Concentrate under reduced pressure and redissolve in 500 mL methanolic ammonia. Let stand overnight and check the reaction by TLC. TLC should show complete deacetylation.
26. Evaporate off methanolic ammonia and apply the residue to a column of silica gel (300 g; 12 × 7 cm) packed in methylene chloride. Short-column chromatography is performed according to Hunt and Rigby (1967) and is similar to the flash chromatography described by Still et al. (1978). Do not use air pressure >1 atm.
27. Elute the column with the following methylene chloride/methanol solutions: 500 mL of 98:2 (v/v); 500 mL of 96:4 (v/v); 1000 mL of 9:1 (v/v); and 1000 mL of 8:2 (v/v). Combine the product-containing fractions and evaporate to a colorless glass. 28. Dissolve the residue in 25 mL chloroform, precipitate by dropwise addition to 2500 mL rapidly stirring pentane, and decant off most of the supernatant (>2000 mL). 29. Collect solids by centrifuging 10 min at 3000 rpm, 15°C, and decant the pentane. 30. Wash twice by resuspending in 250 mL fresh pentane and repeating the centrifugation. Dry the resulting white powder (S.1a) in vacuo. 31. Characterize the final product by TLC and 1H-NMR. 2′-O-MTHP-uridine (S.1a): yield 26.1 g (73% from S.6); Rf (silica, chloroform/methanol 9:1): 0.24; 1H-NMR (250 MHz, DMSO-d6): 1.50-1.90 (m×m, 4H; -CH2-(3,5) of MTHP), 2.95 (s, 3H; H3CO- of MTHP), 3.35-3-50 (m, 2H, -CH2-(5′)), 3.5-3.8 (m×m, 4H; -CH2-(2,6) of MTHP), 3.90 (m, 1H; -CH-(4′)), 3.96 (m, 1H, -CH-(3′)), 4.25-4.36 (m, 1H; -CH-(2′)), 5.10-5.25 (d×tbr, 2H, exchangeable with D2O; 3′/5′-OH), 5.73 (d, J = 8.1, 1H; uridyl-H5), 6.00 (d, J = 7.8, 1H; -CH-(1′)), 7.93 (d, 1H; uridyl-H6), 11.4 (sbr, 1H, exchangeable; uridyl-N3-H).
A Base-Labile Protecting Group
2.4.8 Current Protocols in Nucleic Acid Chemistry
PREPARATION OF 2′-O-(4-METHOXYTETRAHYDROPYRAN-4-YL)4-N-BENZOYLCYTIDINE (S.1b) FROM CYTIDINE
SUPPORT PROTOCOL 2
In the first step of this procedure, cytidine (S.7; Fig. 2.4.4) is simultaneously protected on its 3′- and 5′-hydroxy groups by the bifunctional protecting reagent 1,3-dichloro1,1,3,3-disiloxane (Markiewicz and Wiewerowski, 1985). The derivative S.8 is formed exclusively, due to the higher reactivity of the sterically more accessible 5′-hydroxy group and the subsequently favorable cyclization to the 3′-hydroxy group. Selective benzoylation on N4 of the base is then achieved by treatment with the active ester 1-hydroxybenzotriazolyl benzoate to give S.9. The latter compound is ketalized at the 2′-hydroxy group upon reaction with 5,6-dihydro-4-methoxy-2H-pyran. Then a solution of n-tetrabutylammonium fluoride in acetonitrile removes the disiloxane protecting group to give 2′-OMTHP-4-N-benzoylcytidine (S.1b) in good yield (cf. Reese et al., 1970; van Boom and Wreesmann, 1984). Chromatography is required only for the last step of the sequence. Additional Materials (also see Basic Protocol and Support Protocol 1) Cytidine (S.7; Sigma or Fluka), dried before use for 2 hr at 50°C over phosphorus pentoxide in vacuo N,N-Dimethylformamide, dried by stirring overnight at room temperature with calcium hydride (5 g/L) and subsequent distillation under reduced pressure (b.p. 70° to 80°C, 20 to 30 mmHg) Dry pyridine (see Support Protocol 1 for drying procedure) 1,3-Dichloro-1,1,3,3-tetraisopropyldisiloxane (see Support Protocol 5) 2 M triethylammonium bicarbonate buffer (see recipe) Acetone (analytical grade) 1-Hydroxybenzotriazole (Fluka), dried before use for 72 hr at 50°C over phosphorus pentoxide in vacuo Triethylamine, dried by refluxing 2 hr over calcium hydride (5 g/L) followed by distillation Benzoyl chloride (Fluka, puriss.) Acetonitrile (analytical grade) 1 M n-tetrabutylammonium fluoride (Aldrich/Fluka) in acetonitrile CAUTION: 1-Hydroxybenzotriazole may explode at higher temperatures.
NH2 4
N N1
5'
O
4'
N 2
6
HO
NH2
3
5
steps 1-7
O
Si
1' 2'
3'
HO
N
O
steps 8-15
O
O
O Si O
OH
OH
7 O
O
8
HN
HN N
Si
N
O
O
N O
steps 16-30
N
HO
O
O
O Si
O
OH
HO
O
OCH3
O 9
1b
Figure 2.4.4 Scheme showing the preparation of 2′-O-MTHP-4-N-benzoylcytidine (S.1b) from cytidine (see Support Protocol 2).
Protection of Nucleosides for Oligonucleotide Synthesis
2.4.9 Current Protocols in Nucleic Acid Chemistry
Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)cytidine (S.8) 1. Prepare and stir a suspension of the following: 12.12 g dry cytidine (S.7; 50 mmol) 200 mL dry N,N-dimethylformamide 40 mL dry pyridine (40 mL). 2. Add 18 mL of 1,3-dichloro-1,1,3,3-tetraisopropyldisiloxane (62.5 mmol) to 50 mL dry N,N-dimethylformamide and add dropwise to the stirred cytidine solution over a period of 15 min. Stir the reaction mixture for 1 hr at room temperature. 3. Check the reaction by TLC (see Basic Protocol, steps 4 to 6). TLC should indicate complete reaction of the starting material (Rf < 0.05) to S.8 (Rf = 0.28).
4. Neutralize with 75 mL of 2 M triethylammonium bicarbonate buffer and concentrate under reduced pressure to a small volume (100 mL) using a rotary evaporator. 5. Dissolve in 1000 mL methylene chloride and wash with 500 mL of 1 M aqueous sodium hydrogen carbonate followed by 500 mL water. 6. Dry the organic layer over 50 g magnesium sulfate and concentrate to a colorless oil. 7. Crystallize from 500 mL acetone to produce pure 3′,5′-O-(tetraisopropyldisiloxane1,3-diyl)cytidine. 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)cytidine (S.8): yield 19.5 g (85%), m.p. 226-228 (decomp.), Rf = 0.28. 1H-NMR (CDCl3/CD3OD): 8.00 (d, J = 7.5, 1H; cytidyl-H6), 7.58 (d, J = 7.5, 1H; cytidyl-H5), 5.64 (d, J = 5.5, 1H; -CH-(1′)); 13C-NMR (CDCl3/CD3OD): 166.2 (C4), 156.3 (C2), 140.6 (C6), 94.7 (C5), 91.5 (C1′), 81.6 (C4′), 75.1 (C3′), 68.2 (C2′), 60.0 (C5′), 17.4, 17.0 13.5, 13.0, 12.5 (TIPS).
Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-4-N-benzoyl-cytidine (S.9) 8. Prepare and stir a solution of the following: 10.0 g dry 1-hydroxybenzotriazole (75 mmol) 300 mL dry dioxane 21 mL dry triethylamine (150 mmol). 9. Combine 8.70 mL benzoyl chloride (75 mmol) and 50 mL dry dioxane, and add dropwise to the above mixture (step 8) over 15 min. Stir for 1 hr at room temperature. 10. Filter off the precipitated triethylammonium chloride salts under inert atmosphere using a glass filter crucible, and add the filtrate to a solution containing 24.25 g of 2′-O-(tetraisopropyldisiloxane-1,3-diyl)cytidine (S.8; 50 mmol) in 150 mL dry N,Ndimethylformamide. 11. Evaporate off a volume of ∼100 mL under reduced pressure and stir the residue at room temperature for 3 days. Check the reaction by TLC. TLC should indicate complete reaction of the starting material.
12. Add 5 mL water and concentrate the reaction mixture under reduced pressure to a small volume (100 mL). 13. Dissolve in 1000 mL methylene chloride and wash as in step 5. A Base-Labile Protecting Group
14. Dry the organic layer over 50 g magnesium sulfate and concentrate to a light brown oil.
2.4.10 Current Protocols in Nucleic Acid Chemistry
15. Crystallize from a minimal amount of refluxing acetonitrile to yield analytically pure 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-4-N-benzoylcytidine (S.9). 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)-4-N-benzoylcytidine (S.9): yield 22.6 g (77% in first crop). Rf = 0.40 (silica, chloroform/methanol 9:1). Elemental analysis: calc.; C 57.04, H 7.30, N 7.26%; found: C 56.36, H 7.08, N 7.26%.
Prepare 2′-O-MTHP-4-N-benzoylcytidine (S.1b) 16. Dissolve 4.75 g toluene-p-sulfonic acid monohydrate (25 mmol) in 125 mL dry dioxane and evaporate under reduced pressure to a colorless oil to remove traces of water. 17. Under an inert atmosphere, add 29.45 g of 3′,5′-O-(tetraisopropyldisiloxane-1,3diyl)-4-N-benzoylcytidine (S.9; 50 mmol) followed by 250 mL dry dioxane. 18. To the resulting clear solution, add 29 mL of 5,6-dihydro-4-methoxy-2H-pyran (28.5 g; 250 mmol). Stir 20 hr at room temperature and check the reaction by TLC. TLC should show complete reaction of the starting material.
19. Neutralize the reaction with 3.5 mL half-saturated methanolic ammonia solution (check with moist pH indicator paper) and concentrate immediately under reduced pressure to a small volume (100 mL). 20. Dissolve in 1000 mL methylene chloride and wash as in step 5. 21. Dry the organic layer over 50 g magnesium sulfate and concentrate to a glass. 22. Add 250 mL of 1 M n-tetrabutylammonium fluoride in acetonitrile, stir for 1 hr at room temperature, and check the reaction by TLC. TLC should show complete removal of the silyl protecting group. n-Tetrabutylammonium fluoride in acetonitrile must be prepared by the experimenter. Commercially available solutions in tetrahydrofuran can also be used.
23. Concentrate under reduced pressure to a small volume (100 mL). Dissolve in methylene chloride, wash, and dry as in steps 20 and 21. 24. Dissolve with a minimal amount of dichloromethane and apply to a column of silica gel (300 g; 12 × 7 cm) packed in methylene chloride. Short-column chromatography is performed according to Hunt and Rigby (1967) and is similar to the flash chromatography described by Still et al. (1978). Do not use air pressure >1 atm.
25. Elute the column with the following methylene chloride/methanol solutions: 500 mL of 98:2 (v/v); 500 mL of 96:4 (v/v); and 1000 mL of 9:1 (v/v). 26. Collect the main product-containing fractions and evaporate to a glass. 27. Dissolve the residue in 25 mL chloroform, precipitate by dropwise addition to 2500 mL rapidly stirring pentane, and decant off most of the supernatant (>2000 mL). 28. Collect the solids by centrifuging 10 min at 3000 rpm g, 15°C, and decant the pentane. 29. Wash twice by resuspending in 250 mL fresh pentane and repeating the centrifugation. Dry the resulting white powder (S.1b) in vacuo. 30. Characterize the final product by TLC and 1H-NMR. 2′-O-MTHP-4-N-benzoylcytidine (S.1b): yield 16.1 g (70% from S.9); Rf (silica, chloroform/methanol 9:1): 0.43; 1H-NMR (250 MHz, DMSO-d6): 1.50-1.90 (m×m, 4H; -CH2-
Protection of Nucleosides for Oligonucleotide Synthesis
2.4.11 Current Protocols in Nucleic Acid Chemistry
(3,5) of MTHP), 2.93 (s, 3H; H3CO- of MTHP), 3.40-3.60 (m, 2H, -CH2-(5′)), 3.5-3.8 (m×m, 4H; -CH2-(2,6) of MTHP), 3.96 (m, 1H; -CH-(4′)), 4.02 (m, 1H, -CH-(3′)), 4.39 (m(d×d), 1H; -CH-(2′)), 5.19 (d, J = 6.5, 1H, exchangeable with D2O; 3′-OH), 5.25 (t, J = 2.7, 1H, exchangeable with D2O; 5′-OH), 6.14 (d, J = 6.8, 1H; -CH-(1′)), 7.36 (d, J = 7.7, 1H; cytidyl-H5), 7.4-8.1 (m×m, 5H; benzoyl-aromatic H), 8.43 (d, 1H; cytidyl-H6), 11.3 (sbr, 1H, exchangeable with D2O; cytidyl-N4-H). SUPPORT PROTOCOL 3
PREPARATION OF 2′-O-(4-METHOXYTETRAHYDROPYRAN-4-YL)6-N-BENZOYLADENOSINE (S.1c) FROM ADENOSINE In this protocol, adenosine (S.10, Fig. 2.4.5) is base protected prior to 3′,5′-disilylation by benzoylation of its crystalline 2′,3′,5′-tri-O-acetyl-derivative (S.11), yielding 6-N-benzoyladenosine (S.12) after deacetylation in situ. The chosen route (cf. Schaller et al., 1963; Reese et al., 1970; Büchi and Khorana, 1972; Jones, 1984; van Boom and Wreesmann, 1984; Markiewicz and Wiewerowski, 1985) is then comparable to the cytidine case (see Support Protocol 2): 3′,5′-disilylation to yield S.13, followed by acid-catalyzed ketalization of the 2′-OH by 5,6-dihydro-4-methoxy-2H-pyran, and cleavage of the silyl protecting group by fluoride ion to yield 2′-O-MTHP-6-N-benzoyladenosine (S.1c) in good chromatographically isolated yield. Additional Materials (also see Basic Protocol and Support Protocols 1 and 2) Adenosine (S.10; Sigma or Fluka), dried before use for 2 hr at 50°C over phosphorus pentoxide in vacuo Dry pyridine (see Support Protocol 1 for drying procedure) Acetic anhydride, fractionally distilled with 10% toluene to remove acetic acid 25% (w/v) sodium methoxide in methanol (pract., Aldrich, Fluka) Acetic acid (analytical grade) Diethyl ether (analytical grade) Imidazole (analytical grade) 0.1 M hydrochloric acid
O NH2
7
6
N 5 HO
N 4 N 9
5'
O
4' 3'
HO
1
N
N
8
steps 1-5
2
3
1'
HN
NH2
N
AcO
O
N
N steps 6-18
N
N
HO
O
N N
steps 19-27
2'
OH
AcO
10
HO
OAc
O
O HN
HN N O Si
N O
N
N N
OH
12
11
step 28
N
HO
O
N N
O Si O
OH 13
HO
O
OCH3
O 1c
A Base-Labile Protecting Group
Figure 2.4.5 Scheme showing the preparation of 2′-O-MTHP-6-N-benzoyladenosine (S.1c) from adenosine (see Support Protocol 3). The final step from S.13 to S.1c is performed as in steps 16 to 30 of Support Protocol 2 (see Fig. 2.4.4).
2.4.12 Current Protocols in Nucleic Acid Chemistry
Prepare 2′,3′,5′-tri-O-acetyladenosine (S.11) 1. Prepare and stir a suspension of 10.0 g adenosine (S.10; 37.4 mmol) in 50 mL dry pyridine. 2. Add 23 mL acetic anhydride (0.24 mol) and stir overnight at room temperature. 3. Cool in an ice bath and quench with 40 mL methanol. 4. Remove from the ice bath and stir 1 hr at room temperature. 5. Concentrate to dryness, and recrystallize the white solid residue twice from 100 mL hot ethanol to yield 12.0 g (82%) tri-O-acetyladenosine (S.11). Check the product by TLC (see Basic Protocol, steps 4 to 6). The product is sufficiently pure (Rf = 0.31) for use in the subsequent steps.
Prepare 6-N-benzoyladenosine (S.12) 6. Dissolve crystalline S.11 in 50 mL pyridine, cool in an ice bath, and add 5.3 mL benzoyl chloride (45.8 mmol). 7. Let the reaction mixture warm to room temperature and stir for 2 hr. 8. Quench with 6 mL water and stir for another 30 min. 9. Concentrate to a syrup using a rotary evaporator. 10. Partition the residue between 100 mL chloroform and 100 mL saturated aqueous sodium hydrogen carbonate solution. Reextract the aqueous phase with two 50-mL portions of chloroform. 11. Dry the combined organic extracts over ~20 g anhydrous sodium sulfate and evaporate to a glass (Rf = 0.71). 12. Redissolve in 50 mL of 1:1 (v/v) methanol/pyridine, cool on ice, and then add 15 mL (∼3 eq) of 25% sodium methoxide in methanol. 13. Let the reaction mixture warm to room temperature and monitor by TLC. Deacetylation to the more polar product (Rf = 0.24) occurs in <15 min.
14. Neutralize with 4.0 mL acetic acid and evaporate to a small volume (~13 ) under reduced pressure. 15. Pour the concentrated product mixture into 100 mL saturated aqueous sodium hydrogen carbonate and extract with 200 mL chloroform. 16. Collect the precipitate that settles between the chloroform and aqueous layers by centrifuging 10 min at 3000 rpm, 15°C. 17. Resuspend in ∼100 mL water and centrifuge again. Repeat with ∼100 mL each ethanol and then diethyl ether. 18. Dry the white solid first in air and then under high vacuum to yield 7.15 g (63%) S.12 as a white solid. Check the product by TLC. The product is sufficiently pure (Rf = 0.24) for use in the subsequent steps.
Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-6-N-benzoyladenosine (S.13) 19. Dissolve 2.40 g of 6-N-benzoyladenosine (S.12; 6.46 mmol) in 32 mL dry pyridine. 20. Add 2.16 g imidazole followed by 2.50 mL of 1,3-dichloro-1,1,3,3-tetraisopropyldisiloxane (7.9 mmol; 1.2 eq).
Protection of Nucleosides for Oligonucleotide Synthesis
2.4.13 Current Protocols in Nucleic Acid Chemistry
21. Stir the reaction overnight. Check the reaction by TLC. TLC should show complete reaction of the starting material.
22. Add 10 mL methanol, stir for another 15 min, and concentrate under reduced pressure. 23. Take up the residue in 50 mL chloroform and wash sequentially with 25 mL saturated sodium hydrogen carbonate solution, 50 mL of 0.1 M hydrochloric acid, and 30 mL water. 24. Evaporate the organic layer. 25. Remove traces of pyridine by coevaporating with ∼50 mL (once each) toluene, ethanol, and chloroform. 26. Purify the crude product by short-column chromatography (see Basic Protocol, step 12). Elute with chloroform until the product appears, then with chloroform containing 2% (v/v) ethanol (∼700 mL total in 20-mL fractions). Do not use >1 atm pressure for elution (Hunt and Rigby, 1967; Still et al., 1978). 27. Combine the product-containing fractions, evaporate the solvent, and dry under high vacuum to yield 3.21 g (81%) chromatographically pure S.13 as a colorless foam. Check by TLC (Rf = 0.58). Prepare 2′-O-MTHP-6-N-benzoyladenosine (S.1c) 28. Introduce the MTHP protecting group and perform desilylation (see Support Protocol 2, steps 16 to 30), replacing S.9 (29.45 g; 50 mmol) with the corresponding adenosine derivative S.13 (30.84 g; 50 mmol) to yield S.1c as a dry white powder. 2′-O-MTHP-6-N-benzoyladenosine (S.1c): yield 18.9 g (78% from S.13); Rf (silica, chloroform/methanol 9:1): 0.30; 1H-NMR (250 MHz, DMSO-d6): 1.40-1.90 (m×m, 4H; -CH2(3,5) of MTHP), 2.55 (s, 3H; H3CO- of MTHP), 3.20-3.80 (m×m, 2H, -CH2-(5′)), 3.3-3.8 (m×m, 4H; -CH2-(2,6) of MTHP), 4.05 (m, 1H; -CH-(4′)), 4.18 (m, 1H, -CH-(3′)), 5.00 (m(d×d), 1H; -CH-(2′)), 5.27 (t, J = 5.5, 1H, exchangeable with D2O; 5′-OH), 5.33 (d, J = 4.5, 1H, exchangeable with D2O; 3′-OH), 6.19 (d, J = 7.4, 1H; -CH-(1′)), 7.5-8.1 (m×m, 5H; benzoyl-aromatic H), 8.77 (2s, 2H; adenyl-H2/8), 11.3 (sbr, 1H, exchangeable with D2O; adenyl-N6-H). SUPPORT PROTOCOL 4
PREPARATION OF 2′-O-(4-METHOXYTETRAHYDROPYRAN-4-YL)-2-NISOBUTYRYLGUANOSINE (S.1d) FROM GUANOSINE Guanosine (S.14; Fig. 2.4.6) is base protected on N2 (S.16) by reaction with isobutyryl chloride after transiently blocking the 2′-, 3′-, and 5′-hydroxy groups by persilylation (S.15). The chosen route is then comparable to the cytidine and adenosine cases: 3′,5′-disilylation to S.17, followed by acid-catalyzed ketalization of the 2′-OH by 5,6-dihydro-4-methoxy-2H-pyran and cleavage of the silyl protecting group by fluoride ion to yield 2′-O-MTHP-2-N-isobutyrylguanosine (S.1d) in satisfactory isolated yield. Generally, the higher basicity and polarity of guanosine derivatives renders them more difficult to handle, and the yields are frequently compromised by loss of material in aqueous extraction media or on polar silica gel phases. Nevertheless, the proposed sequence is a viable route that is based on well-described literature procedures (Reese et al., 1970; Ti et al., 1982; Jones, 1984; van Boom and Wreesmann, 1984; Markiewicz and Wiewerowski, 1985; McLaughlin et al., 1985).
A Base-Labile Protecting Group
Additional Materials (also see Basic Protocol and Support Protocols 1 and 2) Guanosine (S.14; Sigma or Fluka), dried before use for 2 hr at 50°C over phosphorus pentoxide in vacuo Dry pyridine (see Support Protocol 1 for drying procedure)
2.4.14 Current Protocols in Nucleic Acid Chemistry
O
O 7 N 5
6 1
N 4 N 9
5'
HO
O
4' 3'
HO
1'
N
NH
8 3
2
NH2
steps 1-2
N
(H3C)3SiO
O
O N
NH N
NH2
steps 3-9
NH
N
HO
N
O
O
N H
steps 10-16
2'
(H3C)3SiO
OH
OSi(CH3)3
HO
15
14
O
O N O Si
N O
NH N
OH
16
N H
N
O steps 17-33
N
HO
O
NH N
O
N H
O Si O
OH
HO
O
OCH3
17 O 1d
Figure 2.4.6 Scheme showing the preparation of 2′-O-MTHP-2-N-isobutyrylguanosine (S.1d) from guanosine (see Support Protocol 4).
Trimethylsilyl chloride (analytical grade) Isobutyryl chloride (analytical grade) Concentrated aqueous ammonia (~25%) Methylene chloride (analytical grade) Phosphorus pentoxide Prepare 2-N-isobutyrylguanosine (S.16) 1. Suspend 14.15 g dry guanosine (S.14; 50 mmol) in 200 mL dry pyridine. 2. Add 47.5 mL trimethylsilyl chloride (375 mmol) and stir for 3 hr at ambient temperature to produce S.15. 3. Cool to 0°C and add 15.6 mL isobutyryl chloride (150 mmol), dropwise, over a period of 30 min. 4. Allow to warm to room temperature and stir overnight. 5. Cool again to 0°C and quench by adding 50 mL water. Stir for another 10 min at room temperature. 6. Add 100 mL concentrated aqueous ammonia to the clear solution and stir for an additional 30 min. 7. Pour the mixture into 800 mL water and extract with 150 mL methylene chloride. 8. Evaporate the aqueous phase to dryness and crystallize three times from 50 to 100 mL boiling water. 9. Dry the resulting solids for 2 days under high vacuum over phosphorus pentoxide to obtain 7.25 g (41%) 2-N-isobutyrylguanosine (S.16) as a grayish-white, chromatographically homogenous powder. Check by TLC (see Basic Protocol, steps 4 to 6; Rf = 0.074). Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-2-N-isobutyrylguanosine (S.17) 10. Dissolve 7.05 g of 2-N-isobutyrylguanosine (S.16; 20.0 mmol) in 125 mL dry pyridine. 11. Add 6.6 mL of 1,3-dichloro-1,1,3,3-tetraisopropyldisiloxane (24 mmol) and stir overnight. Check the reaction by TLC. TLC shows complete reaction of the starting material.
Protection of Nucleosides for Oligonucleotide Synthesis
2.4.15 Current Protocols in Nucleic Acid Chemistry
12. Evaporate the suspension and partition between 250 mL chloroform and 250 mL saturated aqueous sodium hydrogen carbonate solution. 13. Reextract the aqueous layer twice with 125 mL chloroform, dry the organic layer over ~30 g sodium sulfate, and evaporate to a glass using a rotary evaporator. 14. Remove traces of pyridine by azeotropic rotoevaporation from 100 mL toluene (twice), 100 mL ethanol (once), and 100 mL chloroform (once). 15. Purify the crude product by short-column chromatography using 200 g silica gel (see Basic Protocol, step 12). Elute with chloroform until the product appears, then with chloroform containing 2% (v/v) ethanol (∼1500 mL total in 30-mL fractions). Do not use >1 atm pressure for elution (Hunt and Rigby, 1967; Still et al., 1978). 16. Combine the product-containing fractions, evaporate the solvent, and dry under high vacuum to give 7.25 g (61%) chromatographically pure S.17 as a colorless foam. Check by TLC (Rf = 0.55). Prepare 2′-O-MTHP-2-N-isobutyrylguanosine (S.1d) 17. Dissolve 0.95 g of toluene-p-sulfonic acid monohydrate (5 mmol) in 25 mL dry dioxane and evaporate under reduced pressure to a colorless oil to remove traces of water. 18. Under an inert atmosphere, add 5.95 g of 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)2-N-isobutyrylguanosine (S.17; 10 mmol) followed by 50 mL dry dioxane. 19. To the resulting clear solution, add 5.8 mL of 5,6-dihydro-4-methoxy-2H-pyran (5.7 g, 50 mmol). Stir 20 hr at room temperature and check the reaction by TLC. TLC should show complete reaction of the starting material.
20. Neutralize the reaction with 0.8 mL half-saturated methanolic ammonia solution and concentrate immediately under reduced pressure to a small volume (20 mL). 21. Dissolve the residue in 200 mL chloroform and wash with 100 mL of 1 M aqueous sodium hydrogen carbonate solution. 22. Dry the organic layer over 50 g magnesium sulfate and concentrate to a glass. 23. Add 50 mL of 1 M n-tetrabutylammonium fluoride in acetonitrile, stir for 1 hr at room temperature, and check the reaction by TLC. TLC should show complete removal of the silyl group.
24. Concentrate under reduced pressure to a small volume (20 mL). Dissolve in chloroform and wash as in step 21. 25. Reextract the aqueous phase twice with 100 mL chloroform. 26. Dry the organic layer over 10 g anhydrous sodium sulfate and concentrate to a glass. 27. Dissolve the residue with minimal amount of chloroform and apply to a column of silica gel (100 g; 8 × 5 cm) packed in chloroform.
A Base-Labile Protecting Group
28. Elute the column with chloroform until the product appears, and then collect 20-mL fractions with the following chloroform/ethanol solutions: 300 mL of 98:2 (v/v); 300 mL of 96:4 (v/v), 600 mL of 9:1 (v/v). 29. Collect the main product-containing fractions and evaporate to a glass.
2.4.16 Current Protocols in Nucleic Acid Chemistry
30. Dissolve the residue in 25 mL chloroform, precipitate by dropwise addition to 500 mL rapidly stirring pentane, and decant off most of the supernatant (>400 mL). 31. Collect the solids by centrifuging 10 min at 3000 rpm, 15°C, and decant the pentane. 32. Wash the solids twice by resuspending in 100 mL fresh pentane and repeating the centrifugation. Dry the resulting white powder (S.1d) in vacuo. 33. Characterize the final product by TLC and 1H-NMR. 2′-O-MTHP-2-N-isobutyrylguanosine (S.1d): yield 2.42 g (52% from S.17); Rf (silica, chloroform/methanol 9:1): 0.29; 1H-NMR (250 MHz, DMSO-d6): 1.12 (2d, J = 6.8, 6H; (H3C)2CH-)), 1.40-1.90 (m×m, 4H; -CH2-(3,5) of MTHP), 2.55 (s, 3H; H3CO- of MTHP), 2.75 (septet, J = 6.8, 1H; (H3C)2CH-)), 3.19-3.33 (m×m, 2H, -CH2-(5′)), 3.4-3.7 (m×m, 4H; -CH2-(2,6) of MTHP), 3.98 (m(t), 1H; -CH-(4′)), 4.10 (m(t), 1H, -CH-(3′)), 4.74 (m(d×d), 1H; -CH-(2′)), 5.77 (m(t), 1H, exchangeable with D2O; 5′-OH), 5.19 (m(d), 1H, exchangeable with D2O; 3′-OH), 6.98 (d, J = 7.9, 1H; -CH-(1′)), 7.5-8.1 (m×m, 5H; benzoyl-aromatic H), 8.31 (s, 1H; guanyl-H8), 11.7/12.1 (2sbr, 2H, exchangeable with D2O; guanyl-N1-H and N2-H).
PREPARATION OF 1,3-DICHLORO-1,1,3,3TETRAISOPROPYLDISILOXANE
SUPPORT PROTOCOL 5
Although this reagent is commercially available (e.g., Fluka), this protocol provides a convenient procedure for its preparation. Materials Magnesium curls Dry diethyl ether, distilled from phosphorus pentoxide (30 g/L) Isopropyl bromide, distilled from calcium hydride (5 g/L) Trichlorosilane, freshly distilled 0.1 N hydrochloric acid Magnesium sulfate Methylene chloride (analytical grade), dried by passage through activated basic alumina Chlorine gas, dried over concentrated sulfuric acid NaCl plates for infrared (IR) spectroscopy Additional reagents and equipment for IR spectroscopy Prepare 1,1,3,3-tetraisopropyldisiloxane 1. Combine 64 g magnesium curls and 200 mL dry diethyl ether. 2. Add a solution of 270 mL isopropyl bromide in 400 mL dry diethyl ether dropwise to the solution. 3. Stir the reaction mixture mechanically and heat under reflux for 3.5 hr. 4. Add a solution of 100 mL trichlorosilane and 400 mL dry diethyl ether dropwise to the stirred solution (step 3) and heat under reflux overnight. 5. Quench by adding 800 mL of 0.1 N hydrochloric acid in a dropwise fashion. 6. Stir the mixture and heat under reflux for another 3.5 hr. 7. Separate out the organic layer and extract the aqueous layer three times with 300 mL diethyl ether.
Protection of Nucleosides for Oligonucleotide Synthesis
2.4.17 Current Protocols in Nucleic Acid Chemistry
8. Dry the combined organic layers over ~50 g magnesium sulfate and concentrate to a colorless oil under reduced pressure. 9. Distill the residue to produce pure 1,1,3,3-tetraisopropyldisiloxane (100 g; b.p. 80° to 90°C, 10 mmHg) as a colorless oil. Prepare 1,3-dichloro-1,1,3,3-tetraisopropyldisiloxane 10. Dissolve the above product (100 g of 1,1,3,3-tetraisopropyldisiloxane) in 500 mL methylene chloride and pass a stream of dry chlorine gas through the solution. Protect the apparatus from atmospheric moisture using a paraffin oil bubbler. Calcium chloride drying tubes are entirely inefficient and will lead to reaction failure.
11. When the temperature rises to ∼27° to 30°C, cool the reaction mixture to 17° to 20°C by immersing it in an ice water bath while continuing the stream of chlorine gas. 12. After 2 hr, and then after every hour, withdraw a small sample of the reaction mixture and analyze by IR spectroscopy using NaCl plates. 13. Stop the chlorination when the IR spectrum indicates the disappearance of the absorption band at 2100 cm−1 (Si-H). 14. Evaporate off the volatile compounds and distill the residue under diminished pressure (b.p. 85° to 90°C, 2 mmHg) to give 1,3-dichloro-1,1,3,3-tetraisopropyldisiloxane (70 g) as a colorless oil. Store dry at 0°C for up to several months or even years. REAGENTS AND SOLUTIONS Use distilled, deionized water for all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Anisaldehyde reagent 1 mL 4-methoxybenzaldehyde 2 mL concentrated sulfuric acid 10 mL glacial acetic acid 87 mL methanol (analytical grade) Store up to 6 months at room temperature All analytical-grade reagents are preferred if available.
Methanolic ammonia solution, half-saturated (∼8 M) Pass dry ammonia gas (passed over solid potassium hydroxide pellets) through 500 mL analytical-grade methanol at −20°C until saturation. Dilute with 500 mL analytical-grade methanol. Store up to 4 weeks at –18°C. Triethylammonium bicarbonate buffer, 2 M Combine 825 mL analytical-grade triethylamine and 2175 mL water. Saturate with carbon dioxide gas at 0°C until the pH of the clear solution reaches 7.5. Store up to 4 weeks at 4°C. COMMENTARY Background Information
A Base-Labile Protecting Group
Within the field of solid-phase peptide synthesis, the introduction of a base-labile protecting group for the nucleophilic (amino-) center has been very successful, exceeding by far the status of alternative and becoming a standard
technique in many laboratories (Atherton and Sheppard, 1989, and references cited therein). The base-sensitive amino-protecting 9-fluorenylmethoxycarbonyl (FMOC) group, first introduced by Carpino and Han in 1970 (for a review see Carpino, 1987), is strictly orthogo-
2.4.18 Current Protocols in Nucleic Acid Chemistry
nal to the acid-labile protecting groups used for the side chains. Orthogonality is fulfilled when permanent side-chain protecting groups remain completely stable during peptide assembly while being repeatedly exposed to the conditions applied for removal of the temporary terminal protecting group. For solid-phase oligonucleotide assemblies, the use of 5′-OFMOC chemistry may provide an opportunity for such orthogonality, and also avoids the problem of depurination occasionally observed during repeated acidolytic steps required for 5′-O-detritylation. Compared to solid-phase peptide synthesis, functional protection is less demanding in oligonucleotide synthesis due to the smaller number of building blocks. In the deoxyribonucleotide series, very satisfactory protecting groups have resulted from the pioneering efforts in the chemical synthesis of genes by the group of Khorana (e.g., Schaller et al., 1963; Büchi and Khorana, 1972). For example, permanent protection of the exocyclic amino functions as N-acyl and N-aroyl derivatives, respectively, is compatible with nonnucleophilic conditions, but the protecting groups may be cleaved at the very end of the synthesis by hydrolysis in aqueous ammonium hydroxide at elevated temperature. On the other hand, an adequate treatment of the protecting group strategy in RNA synthesis requires the differential handling of the 2′-, 3′-, and 5′-hydroxy groups. If phosphoramidite chemistry is chosen for chain extension, the central problem is the selection of a suitable combination of the permanent 2′-hydroxylprotecting group together with a temporary 5′-hydroxyl-protecting group. The former must remain intact until the very end of the synthesis, including base deprotection, whereafter it must be removed very cleanly and under conditions that leave the phosphate-ribose backbone in complete constitutional and regiochemical integrity. This was convincingly shown both for the acetal (tetrahydropyranyl or THP) as well as the presently applied 4-methoxytetrahydropyranyl (MTHP) ketal-type of 2′-hydroxylprotecting groups some years ago by the research group of Reese (Norman et al., 1984). It was demonstrated by HPLC analysis that diuridine and diadenosine phosphates are stable in 0.01 N hydrochloric acid for time periods over ten times longer than that required for complete 2′-deprotection. Moreover, the authors (Christodoulou et al., 1986) and others (Reese and Skone, 1985) showed that if a conventional acid-labile group such as di-O-(p-an-
isyl)phenylmethyl (dimethoxytrityl or DMTr) or 9-phenylxanthen-9-yl (pixyl or Px) is used to protect the 5′ position, concomitant cleavage of the 2′-O-THP or 2′-O-MTHP takes place to an unacceptable extent. For orthogonal protection of the 5′-hydroxy group, the authors therefore chose the base-labile FMOC protecting group (Fig. 2.4.1 and Fig. 2.4.7). FMOC was first proposed for 5′-O-protection in the deoxyribonucleotide series for solidphase synthesis of an octathymidylic acid fragment (Gioeli and Chattopadhyaya, 1982), and was later used in the same laboratory for the regioselective construction of a 2′-O-pixylprotected dinucleotide (Pathak and Chattopadhyaya, 1985). Later, the results reported for oligothymidylic acids were extended to deoxyribonucleotide sequences containing 4-Nbenzoyl-2′-deoxycytidine and 6-N-benzoyl-2′deoxyadenosine, whereby the above procedure for the regioselective introduction of the FMOC group was again confirmed (Ma and Sonveaux, 1987; Balgobin and Chattopadhyaya, 1987). An approach for a solution-phase RNA synthesis combining an acid-labile acetal group (1ethoxyethyl) for 2′-protection with 5′-OFMOC was proposed by Fukuda et al. (1988). However, the drawback of using chiral protecting groups on chiral nucleoside building blocks has been commented on with crystallographic data (Lehmann et al., 1991). Further efforts to incorporate the FMOC strategy within the framework of solid-phase
B = U; CBz; ABz; Gi-Bu
O [Fmoc] temporary base-labile 5'-O-protection
O O RO
B
5'
O 3'
2'
O O P N(i-Pr)2
OCH3
O [Mthp]
permanent acid-labile 2'-O-protection
phosphate protection R = CH2CH2CN R = CH3 R = CH2CHCH2
Figure 2.4.7 Orthogonal protection scheme for solid-phase RNA synthesis applying a temporary base-labile (FMOC) 5′-O-protecting group and a permanent acid-labile (MTHP) 2′O-protecting group within the phosphoramidite strategy. Compatibility with current variants of phoshate internucleotide protection as well as anchorage to the solid support are discussed in the text.
Protection of Nucleosides for Oligonucleotide Synthesis
2.4.19 Current Protocols in Nucleic Acid Chemistry
A Base-Labile Protecting Group
oligonucleotide chemistry were aimed at avoiding unwanted side reactions during deprotection. Primarily, it was reported (Gao et al., 1985) that, under strongly basic deprotection conditions (5% 1,8-diazabicyclo-[5.4.0]undec7-ene [DBU] in acetonitrile), methyl group transfer to the N3 position of thymine (and presumably uracil) occurs via an intramolecular SN2-reaction from the phosphotriester linkage if the common methyl phosphoramidite chemistry is applied. It was not possible to use H-phosphonates, since the latter are decomposed immediately under the action of base (Lehmann et al., 1989). The authors therefore first applied cyanoethyl phosphoramidites, presently in use for DNA synthesis, conscious of the fact that probably some or all of the phosphate protection is lost during FMOC cleavage. Nevertheless, the results were remarkably encouraging, and it was possible to assemble oligomers of up to 20 residues containing all four ribonucleosides in good yield and in regiochemically homogenous form. The fact that phosphoramidite building blocks are of a significantly basic nature and are prone to hydrolysis to the corresponding H-phosphonates on normal silica gel surfaces demands deactivation of such surfaces by the addition of base to the eluent. If the kind of base (which should be nonnucleophilic and of a low pKa) is chosen inadequately or the 5′-OFMOC-nucleoside-3′-O-phosphoramidite is exposed to higher concentrations of base at elevated temperatures, some loss of the protecting group may occur during isolation, which may lead to a danger of double couplings. The authors therefore recommend (Lehmann et al., 1989) repeated coevaporation of the derived phosphoramidite fractions from toluene under high vacuum followed by immediate precipitation of the product. An evaluation of the less basic N-morpholino phosphoramidite derivatives (McBride and Caruthers, 1983) may be advantageous in the present context. From the more recent literature, two improvements to the FMOC strategy emerge as particularly noteworthy. First, it was possible to apply a safer protection of the phosphate moiety by use of the allyl group as described for a variety of solid-phase protocols by Hayakawa et al. (1990). Moreover it was found (Bergmann et al., 1995) that the final deprotection conditions normally used for removal of the base amino protecting groups (concentrated ammonia, several hours at 55°C) simultaneously take off the allyl groups on the internucleotide phosphate linkage. A second improve-
ment consists of the development of a different linker unit to the solid support, which is of the acetal type and hence is acid labile (2′,3′-Omethylidene-4-phenoxyacetyl; Palom et al., 1993). This allows cleavage of the FMOC group under nucleophilic conditions (10% piperidine in N,N-dimethylformamide) that are significantly less basic and are therefore compatible with an oligonucleotide assembly via methyl phosphoramidites. In addition, two recently introduced alternatives to the FMOC group (BSMOC and MSPOC; Carpino and Mansour, 1999), which are cleaved with lower concentrations of weak nucleophilic bases such as piperidine or morpholine, may be particularly well suited for this oligonucleotide assembly scheme. The linker arm to the support previously used by the authors (succinyl-sarcosyl; Brown et al., 1989) is stable under strongly basic conditions (up to 5% DBU in CH3CN), but is cleaved by exposure to nucleophiles. Finally, the authors would like to encourage the exploitation of new chemistries potentially compatible with the FMOC strategy. For instance, it might be possible to combine the photolabile o-nitrobenzyloxymethyl group (Schwartz et al., 1992; Pitsch, 1997) with the base-labile FMOC group to further increase the scope of this remarkably versatile tool in the chemical synthesis of biooligomers.
Critical Parameters and Troubleshooting The synthesis outlined in the Basic Protocol is fairly short and straightforward. Other than careful attention to basic organic synthesis techniques, little troubleshooting advice needs to be offered. Critical to achieving the expected yields is strict adherence to the given reaction and isolation conditions. The syntheses of the starting materials are considerably more involved, although the expected yields are quite good. Success is dependent upon careful operational planning inherent to multistep syntheses.
Anticipated Results Following the synthesis strategy outlined in the Basic Protocol, yields of between 52% and 65% can be achieved for ribonucleosides protected at the 5′ hydroxyl by FMOC, and 76% can be achieved for 5′-O-FMOC-2′-deoxythymidine. Similarly, good to excellent yields are reported in the literature for deoxyguanosine, deoxyadenosine, and deoxycytidine derivatives (Balgobin and Chattopadhyaya, 1987; Ma and Sonveaux, 1987). The four Support
2.4.20 Current Protocols in Nucleic Acid Chemistry
Protocols for the preparation of the starting materials (2′-O-MTHP-protected ribonucleosides) provide yields of ∼70%, except for the synthesis of 2′-O-MTHP-2-N-isobutyrylguanosine, which provide a yield of ∼52%.
Time Considerations The procedure described in the Basic Protocol may be carried out within 1 to 2 days starting from 2′-O-MTHP-protected ribonucleosides. The preparation of the MTHP-protected starting intermediates can take 2 to 3 weeks per nucleoside derivative. The authors suggest starting with uridine (the simplest case) and allowing time for crystallization of the 3′,5′diacetyl intermediate.
Literature Cited Atherton, E. and Sheppard, R.C. 1989. Solid-Phase Peptide Synthesis: A Practical Approach. IRL Press, Oxford. Balgobin, N. and Chattopadhyaya, J.B. 1987. Solid phase synthesis of DNA under a non-depurinating condition with a base labile 5′-protecting group (Fmoc) using phosphite-amidite approach. Nucleosides Nucleotides 6:461-463. Bergmann, F., Kueng, E., Iaiza, P., and Bannwarth, W. 1995. Allyl as internucleotide protecting group in DNA synthesis to be cleaved off by ammonia. Tetrahedron 51:6971-6976. Brown, T., Pritchard, C.E., Turner, G., and Salisbury, S.A. 1989. A new base-stable linker for solidphase oligonucleotide synthesis. J. Chem. Soc., Chem. Commun. 891-893. Büchi, H. and Khorana, H.G. 1972. Total synthesis of the structural gene for an alanine transfer ribonucleic acid from yeast. Chemical synthesis of an icosadeoxyribonucleotide corresponding to the nucleotide sequence 31 to 50. J. Mol. Biol. 72:251-288. Carpino, L.A. 1987. The 9-fluorenylmethoxycarbonyl family of base-sensitive amino-protecting groups. Acc. Chem. Res. 20:401-407.
Fukuda, T., Hamana, T., and Marumoto, R. 1988. Synthesis of RNA oligomers using 9-fluorenylmethoxycarbonyl (Fmoc) group for 5′-hydroxyl protection. Nucl. Acids Res. 16:13-16. Gao, X., Gaffney, B.L., Senior, M., Riddle, R.R., and Jones, R.A. 1985. Methylation of thymine residues during oligonucleotide synthesis. Nucl. Acids Res. 13:573-584. Gioeli, C. and Chattopadhyaya, J.B. 1982. The fluoren-9-ylmethoxycarbonyl group for the protection of hydroxy-groups; its application in the synthesis of an octathymidylic acid fragment. J. Chem. Soc. Chem. Commun. 1982:672-674. Hayakawa, Y., Uchiyama, M., Kato, H., and Noyori, R. 1990. Allylic protection of internucleotide linkage. Tetrahedron Lett. 26:6505-6508. Hunt, B.J. and Ribgy, W. 1967. Short column chromatography. Chem. Ind. 1967:1868-1869. Jones, R.A. 1984. Preparation of protected deoxyribonucleotides. In Oligonucleotide Synthesis: A Practical Approach (M.J. Gait, ed.) pp. 23-34. IRL Press, Oxford. Jork, H., Funk, W., Fischer, W., and Wimmer, H. 1990. Thin-Layer Chromatography, Vol. Ia: Reagents and Detection Methods, pp. 195-198. VCH-Verlagsgesellschaft, Weinheim. Lehmann, C., Xu, Y.-Z., Christodoulou, C., Tan, Z.-K., and Gait, M.J. 1989. Solid-phase synthesis of oligoribonucleotides using 9-fluorenylmethoxy-carbonyl (Fmoc) for 5′-hydroxyl protection. Nucl. Acids Res. 17:2379-2390. Lehmann, C., Xu, Y.-Z., Christodoulou, C., Gait, M.J., Van Meervelt, L., Moore, M., and Kennard, O. 1991. 3′/5′-Regioselectivity of introduction of the 9-fluorenylmethoxy-carbonyl group to 2′O-tetrahydropyran-2-yl and 2′-O-(4methoxytetrahydropyran-4-yl)-nucleosides: Useful intermediates for solid-phase RNA synthesis. Nucleosides Nucleotides 10:1599-1614. Ma, Y. and Sonveaux, E. 1987. The 9-fluorenylmethyloxycarbonyl (Fmoc) group as a 5′-O base labile protecting group in solid supported oligonucleotide synthesis. Nucleosides Nucleotides 6:491-493.
Carpino, L.A. and Han, G.Y. 1970. the 9-fluorenylmethoxycarbonyl function, a new base-sensitive amino-protecting group. J. Am. Chem. Soc. 92:5748-5749.
Markiewicz, W.T. and Wiewerowski, M. 1985. Simultaneous protection of 3′- and 5′-hydroxyl groups of nucleosides. In Nucleic Acid Chemistry, Section III: Nucleosides (L.B. Townsend and R.S. Tipson, eds.) pp. 229-231. John Wiley & Sons, New York.
Carpino, L.A. and Mansour, E.M.E. 1999. The 2methylsulfonyl-3-phenyl-1-prop-2-enyloxycarbonyl (MSPOC) amino-protecting group. J. Org. Chem. 64:8399-8401.
McBride, L.J. and Caruthers, M.H. 1983. An investigation of several deoxynucleoside phosphoramidites useful for synthesizing deoxyoligonucleotides. Tetrahedron Lett. 24:245-248.
Christodoulou, C., Agrawal, S., and Gait, M.J. 1986. Incompatibility of acid-labile 2′ and 5′ protecting groups for solid-phase synthesis of oligoribonucleotides. Tetrahedron Lett. 27:1521-1522.
McLaughlin, L.W., Piel, N., and Hellmann, T. 1985. Preparation of protected ribonucleotides suitable for chemical oligoribonucleotide synthesis. Synthesis 1985:322-323.
Fromageot, W.P.M., Griffin, B.E., Reese, C.B., and Sulston, J.E. 1967. Monoacylation of ribonucleosides and derivatives via orthoester exchange. Tetrahedron 23:2315-2331.
Norman, D.G., Reese, C.B., and Serafinowska, H.T. 1984. The protection of 2′-hydroxy functions in oligoribonucleotide synthesis. Tetrahedron Lett. 25:3015-3018.
Protection of Nucleosides for Oligonucleotide Synthesis
2.4.21 Current Protocols in Nucleic Acid Chemistry
Palom, Y., Alazzouzi, E-M., Gordillo, F., Grandas, A., and Pedroso, E. 1993. An acid-labile linker for solid-phase oligoribonucleotide synthesis using Fmoc group for 5′-hydroxyl protection. Tetrahedron Lett. 34:2195-2198. Pathak, T. and Chattopadhyaya, J. 1985. The 2′-hydroxyl function assisted cleavage of the internucleotide phosphotriester bond of a ribonucleotide under acidic conditions. Acta Chem. Scand. B39:799-806.
van Boom, J.H. and Wreesmann, C.T.J. 1984. Chemical synthesis of small oligoribonucleotides in solution. In Oligonucleotide Synthesis: A Practical Approach (M.J. Gait, ed.) pp. 153-183. IRL Press, Oxford.
Key-References Blackburn, M. and Gait, M.J. (eds.) 1996. Nucleic Acids in Chemistry and Biology. Oxford University Press, New York.
Pitsch, S. 1997. An efficient synthesis of enantiomeric ribonucleic acid from D-glucose. Helv. Chim. Acta 80:2286-2314.
In particular, chapter 3 on chemical synthesis is very recommendable as an illustrative overview to the present topics.
Reese, C.B. and Skone, P.A. 1985. Action of acid on oligoribonucleotide phosphotriester intermediates. Effect of released vicinal hydroxy functions. Nucl. Acids Res. 13:5215-5231.
Gait, M.J. (ed.) 1984. Oligonucleotide Synthesis: A Practical Approach, IRL Press, Oxford.
Reese, C.B., Saffhill, R., and Sulston, J. 1970. 4Methoxytetrahydropyran-4-yl: A symmetrical alternative to the tetrahydropyranyl group. Tetrahedron 26:1023-1030. Schaller, H., Weiman, G., Lerch, B., and Khorana, H.G. 1963. Protected derivatives of deoxyribonucleotides and new syntheses of deoxyribonucleotide-3′ phosphates. J. Am. Chem. Soc. 85:3821-3827. Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., deBear, J.S., and Gough, G.R. 1992. Rapid synthesis of oligoribonucleotides using 2′-O-(o-nitrobenzyloxymethyl)-protected monomers. Bioorg. Med. Chem. Lett. 2:1019-1024. Still, W.C., Kahn, M., and Mitra, A. 1978. Rapid chromatographic technique for preparative separations with moderate resolution. J. Org. Chem. 43:2923-2925. Ti, G.S., Gaffney, B.L., and Jones, R.A. 1982. Transient protection: Efficient one-flask synthesis of protected deoxynucleosides. J. Am. Chem. Soc. 104:1316-1319.
Basic principles of oligonucleotide synthesis are illustrated by practical advice through further stepby-step protocols; some of the chapters may be regarded as primers for units in the present volume. Lehmann et al., 1989. See above. The procedure in the Basic Protocol is first described for the four 2′-O-MTHP-protected ribonucleosides. For more background information on particular complications arising when the chiral 2′-O-tetrahydropyranyl (THP) group is used, see Lehmann et al., 1991.
Contributed by Michael J. Gait MRC Laboratory of Molecular Biology Cambridge, United Kingdom Christian Lehmann Institute of Organic Chemistry University of Lausanne Lausanne, Switzerland
A Base-Labile Protecting Group
2.4.22 Current Protocols in Nucleic Acid Chemistry
2′-Hydroxyl-Protecting Groups That Are Either Photochemically Labile or Sensitive to Fluoride Ions
UNIT 2.5
Automated chemical synthesis of RNA, like that of DNA, uses protected nucleotide monomers for construction of oligonucleotide chains. RNA monomers, however, are more difficult to obtain than their deoxy counterparts, because of the necessity of protecting their 2′-hydroxyl functions. This unit describes the stepwise preparation, starting from uridine, cytidine, adenosine, and guanosine, of some suitably 2′-protected ribonucleosides. In addition, details are given for protecting the 5′-hydroxyl and the nucleobase, affording ribonucleosides that can be easily converted into either phosphoramidite or phosphonate derivatives, ready to be used in a synthesizer for making RNA. Two alternative sets of protected ribonucleosides are represented here. They are distinguished by the differing conditions required for removal of their 2′-protecting groups. The first consists of uridine, cytidine, adenosine, and guanosine derivatives carrying 2-nitrobenzyloxymethyl (NBOM) groups on their 2′-hydroxyls. The synthesis of these four ribonucleosides is described in Basic Protocols 1 to 4. Oligoribonucleotides synthesized from these components are deprotected by exposure to long-wave UV light (UNIT 3.7). The second set of nucleosides has its 2′-hydroxyls protected with tert-butyldimethylsilyl (TBDMS) groups; these can be removed from product oligoribonucleotides by treatment with tetra-n-butylammonium fluoride. Their syntheses are described in Alternate Protocols 1 to 4. CAUTION: The syntheses in these protocols involve the use of chemicals that have various degrees and kinds of toxicity. Avoid skin contact and inhalation of dusts or vapors. Most operations should be carried out in a well-vented fume hood. NOTE: These reactions should be carried out under strictly anhydrous conditions, using anhydrous solvents and reagents. All glassware should be dried in an oven prior to use. All connections to atmospheric pressure should be through a drying tower containing a desiccant. NOTE: For general information regarding thin-layer chromatography (TLC) or column chromatography, see APPENDIX 3D and APPENDIX 3E, respectively. PREPARATION OF N-PROTECTED 5′-O-(4,4′-DIMETHOXYTRITYL)2′-O-(2-NITROBENZYLOXYMETHYL) NUCLEOSIDES These protocols describe the preparation of ribonucleoside derivatives that incorporate 2-nitrobenzyloxymethyl groups for protection of their 2′-hydroxyls (Figure 2.5.1). In all cases, the 2-nitrobenzyloxymethyl group is introduced into the nucleosides by means of an alkylation reaction utilizing the reagent 2-nitrobenzyl chloromethyl ether, which is prepared as required from its precursor, 2-nitrobenzyl methylthiomethyl ether (see Support Protocol 1). Before any of the nucleoside syntheses described below are carried out, an adequate stock of the methylthiomethyl ether should be accumulated. In addition to the synthesis details, data pertaining to the Rf values from TLC (Table 2.5.1), proton nuclear magnetic resonance (1H NMR; Tables 2.5.2 and 2.5.3), and 13C NMR (Table 2.5.4) of the various 2-nitrobenzyloxymethyl-protected nucleosides are shown.
Contributed by Tod J. Miller, Miriam E. Schwartz, and Geoffrey R. Gough Current Protocols in Nucleic Acid Chemistry (2000) 2.5.1-2.5.36 Copyright © 2000 by John Wiley & Sons, Inc.
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.1 Supplement 3
CH3O DMTrO
B
O 3'
2'
DMTr = OH
O
C
O NO2 CH3O O
O NH
B= N
O
HN
O
Uracil-1-yl
HN N
N N
N
O
N4-benzoylcytosin-1-yl
O N
N
N
N
N6-benzoyladenin-9-yl
NH N
O
N H
N2-isobutyrylguanin-9-yl
Figure 2.5.1 The four 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl) ribonucleosides. The general structure of these ribonucleosides is in the upper-left corner and the four bases (B) are shown below. DMTr is the 4,4′-dimethoxytrityl group.
These data should prove useful in confirming the identities of the products and their synthetic intermediates. NOTE: 2-Nitrobenzyloxymethyl groups are designed to be removable by irradiation with long-wave UV light. Sunlight, or even the light emitted by standard overhead fluorescent bulbs, contains enough UV light to cause slow loss of these protecting groups. Minimize exposure of sensitive compounds by carrying out operations in an area, such as a fume hood, fitted with yellow fluorescent tubes (GE or Sylvania Golds). As an added precaution, wrap flasks and chromatography columns in aluminum foil.
Table 2.5.1 Rf Values of 2-Nitrobenzyloxymethyl-Protected Ribonucleosides on Merck Silica Gel 60 F254 TLC Plates
2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
Ribonucleosidea
Rf valueb
Solvent system
U2′NBOM U3′NBOM A2′NBOM A3′NBOM ABz2′NBOM Gi-Bu2′NBOM 5′DMTrU2′NBOM 5′DMTrU3′NBOM 5′DMTrC2′NBOM 5′DMTrCBz2′NBOM 5′DMTrABz2′NBOM 5′DMTrGi-Bu2′NBOM
0.31 0.31 0.42 0.33 0.56 0.24 0.51 0.29 0.05 0.42 0.56 0.38
9:1 (v/v) chloroform/methanol 9:1 (v/v) chloroform/methanol 9:1 (v/v) chloroform/methanol 9:1 (v/v) chloroform/methanol 9:1 (v/v) chloroform/methanol 9:1 (v/v) chloroform/methanol 2:1 (v/v) ethyl acetate/hexane 2:1 (v/v) ethyl acetate/hexane 2:1 (v/v) ethyl acetate/hexane 2:1 (v/v) ethyl acetate/hexane 2:1 (v/v) ethyl acetate/hexane 2:1 (v/v) ethyl acetate/hexane
a Abbreviations: A, adenosine; C, cytidine; G, guanosine; U, uridine; Bz, benzoyl; DMTr, 4,4′-dimethoxytrityl; i-Bu, isobutyryl; NBOM, 2-nitrobenzyloxymethyl. b TLC Rf values are notoriously irreproducible unless measured under strictly controlled conditions. However, the relative mobilities of substances run on the same plate in the same solvent are valuable guides to compound identification.
2.5.2 Supplement 3
Current Protocols in Nucleic Acid Chemistry
Table 2.5.2 Derivativesb
1H-NMR
Chemical Shiftsa of Nontritylated 2-Nitrobenzyloxymethyl Ribonucleoside
Hydrogen(s)
A2′NBOM
A3′NBOM
ABz2′NBOM
Gi-Bu2′NBOM
H1′ H2′ H3′ H4′ H5′ H2 or H5 H6 or H8 -CH2-O -CH2-Ar Ar(NBOM) 2′- or 3′-OH 5′-OH N6-NH2 Ar (Bz)
6.07 d 4.87 m 4.34 bs 4.03 bs 3.67 m 8.02 s 8.35 s 4.75 s 4.56 s 7.3-8.0 m 5.39 d 5.53 bs 7.29 bs NA
5.96 d 4.85 m 4.34 bs 4.12 bs 3.65 m 8.20 s 8.42 s 5.04 s 5.01 s 7.6-8.1 m 5.71 d 5.63 bs 7.46 bs NA
6.12 d 5.26 m 4.59 d 4.40 bs 3.98 d, 3.78 d 8.60 s 8.69 s 4.73 d 4.90 m 7.4-8.1 m ND ND 7.26 bs 7.3-8.1 m
5.97 d 5.16 m 4.30 bs 3.97 bs 3.61 d 8.27 s ND 4.87 m 4.71 bs 7.2-8.1 m ND ND NA NA
The internal reference for 1H-NMR spectra was tetramethylsilane at 0 ppm. The solvents were dimethy sulfoxide-d6 for the A derivatives and acetonitrile-d3 for the G derivative. All shifts are measured in ppm. b Abbreviations: A, adenosine; G, guanosine; Ar, aromatic; Bz, benzoyl; i-Bu, isobutyryl; NBOM, 2-nitrobenzyloxymethyl; bs, broad singlet; d, doublet; m, multiplet; s, singlet; NA, not applicable; ND, not determined. a
Table 2.5.3 1H-NMR Chemical Shiftsa of 5′-Dimethoxytritylated 2-Nitrobenzyloxymethyl Ribonucleoside Derivativesb
Hydrogen(s)
5′DMTrU2′ NBOM
5′DMTrC2′ NBOM
5′DMTrCBz2′ 5′DMTrABz2′ 5′DMTrGi-Bu NBOM NBOM 2′NBOM
H1′ H2′ H3′ H4′ H5′ H2 or H5 H6 or H8 -OCH3 Ar (DMTr) -CH2-O-CH2-Ar Ar (NBOM) 2′- or 3′-OH N1- or N3-H N4- or N6-H Ar (Bz) i-Bu-CH(CH3)2
6.06 d 4.40 dd 4.51 m 4.11 bs 3.56 bs 5.21 d 7.96 d 3.82 s 6.80-7.40 m 5.00-5.15 bs 5.00-5.15 bs 7.40-8.41 m 5.19 bs 10.00 s NA NA NA
5.87 d 4.15 d 4.26 m 4.00 d 3.28 bs 5.44 d 7.79 d 3.73 s 6.87-7.40 m 4.85-5.05 m 4.85-5.05 m 7.57-8.08 m 5.30 d NA ND NA NA
6.10 bs 4.58 m 4.43 d 4.19 d 3.64 bs 5.46 d 7.93 d 3.86 s 6.80-7.50 m 5.12 m 5.12 m 7.50-8.20 m ND NA ND 7.10-8.70 m NA
5.99 d 5.26 m 4.78 d 4.40 bs 3.91 m 8.72 s 8.68 bs 3.80 s 6.75-7.40 m 4.60 m 4.83 m 7.40-8.10 m ND NA 10.00 s 7.10-8.10 m NA
5.99 d 5.20 m 4.60 dd 4.26 d 3.48 d, 3.21 dd 8.65 bs ND 3.79 s 6.70-7.40 m 4.81 m 4.90 m 7.40-8.70 m ND 9.28 s NA NA 0.92 dd
a The internal reference for 1H-NMR spectra was tetramethylsilane at 0 ppm. The solvent was chloroform-d. All shifts are measured in ppm. b Abbreviations: A, adenosine; C, cytidine; G, guanosine; U, uridine; Ar, aromatic; Bz, benzoyl; DMTr, 4,4′-dimethoxytrityl; i-Bu, isobutyryl; NBOM, 2-nitrobenzyloxymethyl; bs, broad singlet; d, doublet; dd, doublet of doublets; m, multiplet; s, singlet; NA, not applicable; ND, not determined.
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.3 Current Protocols in Nucleic Acid Chemistry
Supplement 3
Table 2.5.4
13C-NMR
Chemical Shiftsa of 2-Nitrobenzyloxymethyl Ribonucleoside Derivativesb
Carbon
A2′NBOM
U2′NBOM
Gi-Bu2′NBOM
C1′ C2′ C3′ C4′ C5′ C2 C4 C5 C6 C8 -OCH2-OCH2-Ar NBOM (C1) NBOM (C2) NBOM (C3) NBOM (C4) NBOM (C5) NBOM (C6) i-Bu (CH3) i-Bu (CH3) i-Bu (CH) i-Bu C=O
86.5 78.3 69.3 86.2 61.5 152.2 148.7 119.2 156.0 139.7 65.6 94.0 133.6 146.6 133.3 128.1 124.3 128.3 NA NA NA NA
86.4 78.5 68.7 85.3 60.6 150.2 163.1 102.0 140.4 NA 65.9 94.1 133.9 147.1 133.7 128.6 124.6 128.8 NA NA NA NA
86.7 81.0 70.3 85.0 61.8 149.0 148.4 120.3 154.9 137.7 66.2 95.2 134.3 146.5 134.0 127.9 124.7 128.7 19.1 19.3 35.1 180.4
a The internal reference for 13C-NMR spectra was dimethyl sulfoxide-d6 at 39.7 ppm. The solvent was dimethyl sulfoxide-d6. While not exhaustive, these useful and commonly observed resonances are listed with tentative assignments. All shifts are measured in ppm. b Abbreviations: A, adenosine; G, guanosine; U, uridine; Ar, aromatic; i-Bu, isobutyryl; NBOM, 2-nitrobenzyloxymethyl; NA, not applicable.
BASIC PROTOCOL 1
2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
Synthesis of 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)uridine In this protocol, uridine is converted into its dibutylstannylene derivative, then alkylated using 2-nitrobenzyl chloromethyl ether. The resulting mixture of 2′- and 3′-O-(2-nitrobenzyloxymethyl)uridine is treated with 4,4′-dimethoxytrityl chloride. The dimethoxytrityl derivatives are separated by silica-gel chromatography, affording pure 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)uridine. The overall reaction sequence is illustrated in Figure 2.5.2. Materials Methanol Uridine Dibutyltin oxide Phosphorus pentoxide Anhydrous tetra-n-butylammonium bromide Anhydrous dimethylformamide, store over 4A molecular sieves 2-Nitrobenzyl chloromethyl ether, make fresh (see Support Protocol 1) Anhydrous pyridine, store over coarse granules of calcium hydride 9:1 and 95:5 (v/v) chloroform/methanol 66% (v/v) aqueous pyridine Silica gel 60, 70 to 230 mesh ASTM (e.g., EM Science)
2.5.4 Supplement 3
Current Protocols in Nucleic Acid Chemistry
O
O
NH
NH HO
N
O
O
HO MeOH
OH
OH
N
O
Dibutyltin oxide O
O 2-Nitrobenzyl chloromethyl ether Bu4NBr/DMF
O Sn
Bu
Bu
O
O NH
HO
N
O OH
NH
O
DMTrO 1. DMTrCl/Pyridine
O
2. Separation of isomers
O NO2
+
N
O OH
O
O
O NO2
3'-isomer
Figure 2.5.2 Preparation of 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)uridine from uridine. Bu is n-butyl.
2.5 × 30–cm and 3 × 60–cm glass chromatography columns packed with silica gel 60, 70 to 230 mesh ASTM, in chloroform to a bed height of 25 and 50 cm, respectively (see Support Protocol 3) Chloroform Anhydrous triethylamine, store over coarse granules of calcium hydride 4,4′-Dimethoxytrityl chloride Ethyl acetate 1 M NaHCO3 2 M NaCl Anhydrous sodium sulfate 2:1 (v/v) ethyl acetate/hexane 0% to 1% (v/v) methanol in chloroform, containing 0.25% (v/v) pyridine 250-mL and 2-L round-bottom flasks Boiling chips Water-cooled reflux condenser Heating mantle Rotary evaporator connected interchangeably to water aspirator and vacuum pump 50-mL pressure-equalizing dropping funnel Large petri plate Filter paper, coarse porosity and fast flow rate (e.g., Quantitative Q8, Fisher) Additional reagents and equipment for TLC and column chromatography (see Support Protocol 3) Prepare 2′,3′-O-(dibutylstannylene)uridine 1. Add a stir-bar to 1 L methanol in a 2-L round-bottom flask and begin stirring. Add 4.88 g (20 mmol) uridine and 5.00 g (20 mmol) dibutyltin oxide.
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.5 Current Protocols in Nucleic Acid Chemistry
Supplement 3
2. Remove stir-bar, add boiling chips, and fit flask with a water-cooled reflux condenser. Heat mixture to a boil using a heating mantle, reflux 45 min to 1 hr, and let cool to room temperature. 3. Decant solution from boiling chips into a second 2-L flask and remove methanol using a rotary evaporator connected to a water aspirator. 4. Dry crystalline residue to constant weight in a vacuum desiccator over phosphorus pentoxide. A yield of 9 g 2′,3′-O-(dibutylstannylene)uridine (with a melting point of 232° to 234°C) is expected. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
Alkylate to form 2′- and 3′-O-(2-nitrobenzyloxymethyl)uridine 5. Add 5.10 g (10.7 mmol) dry 2′,3′-O-(dibutylstannylene)uridine and 1.72 g (5.35 mmol) anhydrous tetra-n-butylammonium bromide to 40 mL anhydrous dimethylformamide in a 250-mL round-bottom flask, while stirring. Fit flask with a 50-mL pressure-equalizing dropping funnel. 6. Use dropping funnel to add a freshly prepared solution of 2-nitrobenzyl chloromethyl ether (from 16 mmol 2-nitrobenzyl methylthiomethyl ether; see Support Protocol 1) dropwise over a 5-min period while stirring, then seal flask and stir 2 hr at room temperature. 7. Add 5 mL anhydrous pyridine followed by 2 mL water and continue stirring 20 min at room temperature. 8. Check reaction by TLC (see Support Protocol 3) using 9:1 chloroform/methanol. The unresolved 1:1 mixture of 2′- and 3′-isomers will be visible under UV light as a dark band about a third of the way up the plate (Table 2.5.1).
9. Remove volatile solvents using the rotary evaporator connected to water aspirator, then continue evaporation under high vacuum (i.e., with a vacuum pump) to remove as much of the dimethylformamide as possible. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
Purify 2′- and 3′-O-(2-nitrobenzyloxymethyl)uridine 10. Dissolve residue in the minimum volume of 66% aqueous pyridine and, using a Pasteur pipet, transfer it dropwise onto 10 g silica gel 60 in a large petri plate. Allow wet silica to dry in the moving airstream inside a fume hood. An overnight drying time should be adequate if most of the dimethylformamide was removed and the minimum amount of aqueous pyridine was used.
11. Pulverize the dry, coated silica. Divide the powdery material into halves and layer them onto two 2.5 × 30–cm glass chromatography columns. 12. Wash each column with 100 mL chloroform, elute with 95:5 chloroform/methanol, and collect fractions as described (see Support Protocol 3). 2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
The volume of 95:5 chloroform/methanol should be determined from TLC results.
13. Monitor fractions by TLC using 9:1 chloroform/methanol, comparing selected fractions against samples of the original reaction mixture to locate the products.
2.5.6 Supplement 3
Current Protocols in Nucleic Acid Chemistry
14. Pool and evaporate the appropriate fractions using the rotary evaporator connected to a water aspirator. A yield of 4.2 g of a 1:1 mixture of the 2′- and 3′-isomers of (2-nitrobenzyloxymethyl)uridine is expected. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
Prepare 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)uridine 15. Dry 4.2 g (10.3 mmol) mixed 2′- and 3′-O-(2-nitrobenzyloxymethyl)uridine isomers by three co-evaporations with 30 mL anhydrous pyridine using the rotary evaporator connected to a vacuum pump. 16. Dissolve residue in 114 mL anhydrous pyridine, then add 1.60 mL (11.4 mmol) anhydrous triethylamine and 3.87 g (11.4 mmol) 4,4′-dimethoxytrityl chloride. Stir 3 hr at room temperature. 17. Add 5 mL methanol, allow to stand 10 min, then remove solvents on the rotary evaporator connected to a vacuum pump. 18. Dissolve residue in 250 mL ethyl acetate. Extract three times with 100 mL of 1 M NaHCO3 and once with 100 mL of 2 M NaCl. CAUTION: Carbon dioxide is released during the sodium bicarbonate extractions.
19. Dry ethyl acetate layer by addition of 5 g anhydrous sodium sulfate, filter through filter paper to remove salt, and rotary evaporate filtrate to a yellow foam using a water aspirator. 20. Analyze this material by TLC in 2:1 ethyl acetate/hexane. Two major bands should be visible. The upper (faster-running) one is the 2′-isomer (Table 2.5.1). If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
Purify 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)uridine 21. Dissolve foam (∼8 g) in 5 to 10 mL chloroform and add it to the top of a 3 × 60–cm silica gel column. Elute products from column using a 1.2-L stepwise gradient of 0% to 1% methanol in chloroform containing 0.25% pyridine as described. 22. Monitor fractions by TLC using 2:1 ethyl acetate/hexane. Any mixed fractions containing both isomers can be rechromatographed to obtain more pure 2′-isomer. Gradients of ethyl acetate in hexane may also be used for purification of these dimethoxytrityl nucleosides, as preliminary results with this alternative solvent mixture have been encouraging.
23 Combine those fractions containing the pure 2′-isomer, and rotary evaporate them to a foam using a water aspirator. A yield of 40% 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)uridine, based on the starting quantity of uridine, is expected.
24. Store protected nucleoside in the dark at −20°C in a sealed flask containing a trace of pyridine (stable for many months).
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.7 Current Protocols in Nucleic Acid Chemistry
Supplement 3
BASIC PROTOCOL 2
Synthesis of 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N4benzoylcytidine In this protocol, 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)uridine (see Basic Protocol 1) is converted into its corresponding cytidine derivative, then selectively benzoylated with pentafluorophenyl benzoate at the N4 position. The reaction sequence is illustrated in Figure 2.5.3. Materials 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)uridine (see Basic Protocol 1) Anhydrous pyridine, stored over coarse granules of calcium hydride Acetic anhydride Ethyl acetate 1 M NaHCO3 2 M NaCl Anhydrous sodium sulfate 1,2,4-Triazole 4-Chlorophenyl phosphorodichloridate 3:1 (v/v) pyridine/concentrated ammonium hydroxide 2.5 × 30–cm glass chromatography column packed with silica gel 60, 70 to 230 mesh ASTM, in chloroform to a bed height of 25 cm (see Support Protocol 3) 0% to 1% and 0% to 4% (v/v) methanol in chloroform, containing 0.25% (v/v) pyridine 2:1 (v/v) ethyl acetate/hexane Pentafluorophenyl benzoate (see Support Protocol 2)
O
O NH
DMTrO
N
O OH
NH
O
DMTrO
O
O
N
O
Acetic anhydride/Pyridine AcO
O
O
O
NO2
NO2 O O P Cl Cl
1. Cl
2. NH4OH
Triazole/Pyridine O NH2
HN
N
N DMTrO
2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
N
O OH
O
DMTrO Pentafluorophenyl benzoate
O
Pyridine
O NO2
N
O OH
O
O
O NO2
Figure 2.5.3 Preparation of 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N4-benzoylcytidine from 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)uridine.
2.5.8 Supplement 3
Current Protocols in Nucleic Acid Chemistry
250- and 500-mL separatory funnels Filter paper, coarse porosity and fast flow rate (e.g., Quantitative Q8, Fisher) Rotary evaporator connected interchangeably to water aspirator and vacuum pump 10- and 25-mL round-bottom flasks Additional reagents and equipment for TLC and column chromatography (see Support Protocol 3) Protect 3′-hydroxyl of 5′-O-(4,4′-dimethoxytrityl)-2′-O(2-nitrobenzyloxymethyl)uridine 1. Add 580 mg (0.82 mmol) 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)uridine to a flask containing 3 mL anhydrous pyridine and stir to dissolve. 2. Add 1.64 mL (17 mmol) acetic anhydride and allow reaction mixture to stand 2 hr at room temperature. 3. Cool flask in an ice bath and add 2.5 mL anhydrous pyridine, followed by dropwise addition over a 5-min period of 4 mL water, while stirring and cooling continuously. 4. Add an additional 1 mL anhydrous pyridine to redissolve any precipitate, and allow solution to stand 18 hr at room temperature in the dark. 5. Transfer mixture to a 250-mL separatory funnel using 50 mL ethyl acetate and extract the organic layer three times with 50 mL of 1 M NaHCO3 and three times with 50 mL of 2 M NaCl. CAUTION: Carbon dioxide is released during the sodium bicarbonate extractions. The 3′-hydroxyl is temporarily protected as an acetate ester.
Prepare 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)cytidine 6. Dry ethyl acetate layer over 3 g anhydrous sodium sulfate, filter through filter paper to remove salt, and evaporate filtrate to a foam using a rotary evaporator connected to a water aspirator. 7. Co-evaporate the product three times with 10 mL anhydrous pyridine in a 25-mL round-bottom flask using the rotary evaporator connected to a vacuum pump. Dissolve final residue in 2.7 mL anhydrous pyridine. 8. Add 314 mg (4.54 mmol) 1,2,4-triazole, stir to dissolve (∼1 hr), then add 0.36 mL (2.2 mmol) 4-chlorophenyl phosphorodichloridate. Stir 1 hr and then let stand 72 hr at room temperature in the dark. 9. Cool flask in an ice bath and add 0.2 mL water while stirring. Stir 30 min at 0°C, then add 3 mL anhydrous pyridine. 10. Use a disposable Pasteur pipet to add the resulting solution dropwise to 80 mL of 3:1 pyridine/concentrated ammonium hydroxide while stirring. Loosely stopper the container and allow reaction mixture to stand 48 hr at room temperature in the dark. 11. Concentrate solution to a slurry by rotary evaporation using a water aspirator and transfer to a 500-mL separatory funnel using 150 mL ethyl acetate. 12. Extract three times with 100 mL of 1 M NaHCO3, then three times with 100 mL of 2 M NaCl. 13. Repeat step 6.
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.9 Current Protocols in Nucleic Acid Chemistry
Supplement 3
14. Layer residue on a 2.5 × 30–cm silica gel column and elute products using a 1-L stepwise gradient of 0% to 4% methanol in chloroform containing 0.25% pyridine as described (see Support Protocol 3). 15. Monitor fractions by TLC (see Support Protocol 3) using 2:1 ethyl acetate/hexane. 16. Evaporate appropriate fractions using the rotary evaporator connected to a water aspirator to produce the pure product, 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)cytidine. A 71% yield, based on the starting quantity of 5′-O-(4,4’-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)uridine, is expected. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
Prepare 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N4benzoylcytidine 17. Dissolve 412 mg (0.58 mmol) 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)cytidine in 1 mL anhydrous pyridine in a 10-mL round-bottom flask. 18. Add 251 mg (0.87 mmol) pentafluorophenyl benzoate and swirl flask to dissolve. Seal flask and allow solution to stand 6 days at room temperature in the dark. 19. Examine mixture by TLC using 2:1 ethyl acetate/hexane, comparing it to the starting material to confirm complete conversion to the N-benzoyl derivative. 20. Transfer reaction mixture to a 250-mL separatory funnel using 100 mL ethyl acetate, then extract as in step 5. 21. Repeat step 6. 22. Repeat steps 14 to 16, using a 1.2-L stepwise gradient of 0% to 1% methanol in chloroform containing 0.25% pyridine in step 14. A 91% yield based on the starting quantity of 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)cytidine is expected.
23. Store protected nucleoside in the dark at −20°C in a sealed flask containing a trace of pyridine (stable for many months). BASIC PROTOCOL 3
Synthesis of 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N6-benzoyladenosine In this protocol, adenosine is converted into its dibutylstannylene derivative, then alkylated with 2-nitrobenzyl chloromethyl ether, and the resulting 2′- and 3′-O-(2-nitrobenzyloxymethyl)adenosines are separated by column chromatography. Benzoylation at the N6 position (to protect the nucleobase), followed by dimethoxytritylation (to protect the 5′-hydroxyl), completes the synthesis. The reaction sequence is illustrated in Figure 2.5.4.
2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
Materials Methanol Adenosine Dibutyltin oxide Phosphorous pentoxide Anhydrous tetra-n-butylammonium bromide Anhydrous dimethylformamide, stored over 4A molecular sieves 2-Nitrobenzyl chloromethyl ether, make fresh (see Support Protocol 1)
2.5.10 Supplement 3
Current Protocols in Nucleic Acid Chemistry
NH2 N HO
NH2 N
N
N
N
N
HO
O
O
Dibutyltin oxide MeOH
OH OH
N N 2-Nitrobenzyl chloromethyl ether
O
Bu4NBr/DMF
O Sn Bu
Bu
O NH2 N HO
O
HN N
N
N
HO
N 1. Separation of isomers
OH O
2. Me3SiCl/Pyridine 3. Benzoyl chloride 4. NH4OH
O NO2
+
O
N
N
OH O
N
O NO2
3'-isomer
O DMTrCl/Pyridine HN N DMTrO
O OH
N
N
O
N
O NO2
Figure 2.5.4 Preparation of 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N6-benzoyladenosine from adenosine. Bu is n-butyl.
Anhydrous pyridine, store over coarse granules of calcium hydride 9:1 and 95:5 (v/v) chloroform/methanol 66% (v/v) aqueous pyridine Silica gel 60, 70 to 230 mesh ASTM (e.g., EM Science) 2.5 × 25–cm, 2.5 × 30–cm, and 3 × 60–cm glass chromatography columns packed with silica gel 60, 70 to 230 mesh ASTM, in chloroform (see Support Protocol 3) 1% to 5% (v/v) methanol in chloroform 80% (v/v) aqueous acetonitrile Acetonitrile Trimethylchlorosilane Benzoyl chloride Concentrated ammonium hydroxide Ethyl acetate 1 M NaHCO3 2 M NaCl Anhydrous sodium sulfate Chloroform Triethylamine, stored over coarse granules of calcium hydride 4,4′-Dimethoxytrityl chloride 0% to 1% (v/v) methanol in chloroform containing 0.25% (v/v) pyridine 2:1 (v/v) ethyl acetate/hexane 100-mL, 250-mL, and 2-L round-bottom flasks Boiling chips Water-cooled reflux condenser Heating mantle
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.11 Current Protocols in Nucleic Acid Chemistry
Supplement 3
Rotary evaporator connected interchangeably to water aspirator and vacuum pump Oil bath, 60°C 50-mL pressure-equalizing dropping funnel 500-mL separatory funnel Filter paper, coarse porosity and fast flow rate (e.g., Quantitative Q8, Fisher) Additional reagents and equipment for TLC and column chromatography (see Support Protocol 3) Prepare 2′,3′-O-(dibutylstannylene)adenosine 1. Add a stir-bar to 700 mL methanol in a 2-L round-bottom flask and begin stirring. Add 6.7 g (25 mmol) adenosine and 6.25 g (25 mmol) dibutyltin oxide. 2. Remove stir-bar, add boiling chips, and fit flask with a water-cooled reflux condenser. Heat mixture to boiling using a heating mantle, reflux 45 min, and let cool to room temperature. 3. Decant solution from boiling chips into a second 2-L flask and remove methanol using a rotary evaporator connected to a water aspirator. 4. Dry residue to constant weight in a vacuum desiccator over phosphorus pentoxide. A yield of 10.5 g is expected. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
Alkylate to form 2′- and 3′-O-(2-nitrobenzyloxymethyl)adenosine 5. Add 8 g (16 mmol) dry 2′,3′-O-(dibutylstannylene)adenosine and 7.7 g (16 mmol) anhydrous tetra-n-butylammonium bromide to 65 mL anhydrous dimethylformamide in a 250-mL round-bottom flask while stirring. 6. Heat to 60°C in an oil bath and fit flask with a 50-mL pressure-equalizing dropping funnel. 7. Use the dropping funnel to add freshly prepared 2-nitrobenzyl chloromethyl ether (from 16 mmol 2-nitrobenzyl methylthiomethyl ether; see Support Protocol 1) dropwise over a 5-min period while stirring at 60°C. Seal flask and stir 1 hr at 60°C. 8. Add 20 mL anhydrous pyridine and stir 25 min at room temperature. 9. Examine reaction mixture by TLC (see Support Protocol 3) in 9:1 chloroform/methanol. The 2′- and 3′-isomers are visible as two closely spaced bands in the lower half of the plate. The upper band of this pair is the 2′-isomer (Table 2.5.1).
10. Remove solvents by rotary evaporation, using high vacuum (i.e., with a vacuum pump) to remove as much dimethylformamide as possible. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
Purify 2′-O-(2-nitrobenzyloxymethyl)adenosine 11. Dissolve residue in the minimum volume of 66% aqueous pyridine and, using a Pasteur pipet, transfer it dropwise onto 15 g silica gel 60 in a large petri plate. Allow wet silica to dry in the moving stream of air inside a fume hood.
2.5.12 Supplement 3
Current Protocols in Nucleic Acid Chemistry
An overnight drying time should be adequate if most of the dimethylformamide was removed and the minimum amount of aqueous pyridine was used.
12. Pulverize the dry, coated silica. Layer powder onto a 3 × 60–cm glass chromatography column and elute with 95:5 chloroform/methanol, collecting fractions as described (see Support Protocol 3). The volume of 95:5 chloroform/methanol should be determined from TLC results.
13. Monitor fractions by TLC in 9:1 chloroform/methanol. 14. Combine all fractions containing the 2′-isomer and rotary evaporate to dryness using a water aspirator. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
15. Dry material on 6 g silica gel 60 as in step 11 and repeat step 12 using a 1-L stepwise gradient of 1% to 5% methanol in chloroform (beginning with 200 mL of 1% methanol, followed by 200 mL of 2% methanol, and so on). 16. Repeat steps 13 and 14. 2′-O-(2-Nitrobenzyloxymethyl)adenosine contaminated with a trace of the 3′-isomer is left after solvent removal. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
17. Dissolve residue in a minimal amount of 80% aqueous acetonitrile with heating, then use the rotary evaporator connected to water aspirator to repeatedly co-evaporate with acetonitrile to a final volume of ∼5 mL until crystallization of the 2′-isomer occurs. A 24% yield of white crystals is obtained, based on the starting quantity of 2′,3′-O-(dibutylstannylene)adenosine. This material may be further recrystallized from water. The melting point of the product is 175° to 177°C. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
Prepare 2′-O-(2-nitrobenzyloxymethyl)-N6-benzoyladenosine 18. Add 1.08 g (2.5 mmol) 2′-O-(2-nitrobenzyloxymethyl)adenosine to 10 mL anhydrous pyridine in a 100-mL round-bottom flask while stirring to dissolve. Add 1.59 mL (12.5 mmol) trimethylchlorosilane, seal flask, and stir 2 hr at room temperature. 19. Add 1.45 mL (12.5 mmol) benzoyl chloride and reseal flask. Stir 2 hr, then cool to 0°C and add 2.5 mL water. Stir 10 min, then add 5 mL concentrated ammonium hydroxide. Cover (do not seal) flask and stir 30 min at room temperature. Ammonium hydroxide from a newly opened bottle should be used, and the reaction should be checked with pH paper to make certain it remains alkaline throughout the 30 min. More ammonium hydroxide should be added in 1-mL aliquots, if necessary, to maintain alkalinity.
20. Remove ammonia and solvents by rotary evaporation using a water aspirator. 21. Transfer reaction mixture to a 500-mL separatory funnel using 150 mL ethyl acetate and extract three times with 100 mL of 1 M NaHCO3, then three times with 100 mL of 2 M NaCl. 22. Dry ethyl acetate layer over 5 g anhydrous sodium sulfate, filter through filter paper to remove salt, and rotary evaporate filtrate using a water aspirator.
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.13 Current Protocols in Nucleic Acid Chemistry
Supplement 3
If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
23. Layer residue on a 2.5 × 25–cm silica gel column and elute with 100 mL chloroform, followed by 300 mL of 1% methanol in chloroform and 800 mL of 2% methanol in chloroform as described (see Support Protocol 3). 24. Monitor fractions by TLC using 9:1 chloroform/methanol. 25. Use the rotary evaporator connected to a water aspirator to evaporate appropriate fractions and produce 2′-O-(2-nitrobenzyloxymethyl)-N 6-benzoyladenosine. A yield of 88%, based on the starting quantity of 2′-O-(2-nitrobenzyloxymethyl)adenosine, is expected. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
Prepare 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N6benzoyladenosine 26. Dry 1.05 g (1.96 mmol) 2′-O-(2-nitrobenzyloxymethyl)-N6-benzoyladenosine by three co-evaporations with 30 mL anhydrous pyridine using the rotary evaporator connected to a vacuum pump. 27. Dissolve residue in 30 mL anhydrous pyridine, then add 0.34 mL (2.4 mmol) triethylamine and 0.80 g (2.4 mmol) 4,4′-dimethoxytrityl chloride. Stir 3 hr at room temperature. 28. Add 1 mL methanol and, after 10 min, remove solvents by rotary evaporation using a vacuum pump. 29. Repeat steps 21 to 25 with the following exceptions: a. Extract three times with NaHCO3 but only once with NaCl in step 21. b. Use a 2.5 × 30–cm silica gel column and a 1.2-L stepwise gradient of 0% to 1% methanol in chloroform containing 0.25% pyridine in step 23. c. Use 2:1 ethyl acetate/hexane for TLC in step 24. The yield of 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N6-benzoyladenosine is expected to be 91% based on the starting quantity of 2′-O-(2-nitrobenzyloxymethyl)-N6-benzoyladenosine.
30. Store 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N6-benzoyladenosine in the dark at −20°C in a sealed container containing a trace of pyridine (stable for many months). BASIC PROTOCOL 4
2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
Synthesis of 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N2isobutyrylguanosine In this protocol, guanosine is protected at its N2 position with the isobutyryl group, then converted into its dibutylstannylene derivative and alkylated with 2-nitrobenzyl chloromethyl ether. The resulting 2′-O-(2-nitrobenzyloxymethyl) nucleoside is purified by silica-gel chromatography, then crystallized from an acetonitrile/water mixture. Dimethoxytritylation for protection of the 5′-hydroxyl is the final step in the synthesis. The reaction sequence is illustrated in Figure 2.5.5.
2.5.14 Supplement 3
Current Protocols in Nucleic Acid Chemistry
O N HO
O N
NH
N
N
O
HO
NH2
O
1. Isobutyryl chloride/Pyridine 2. Aq. NaOH in ethanol
OH OH
N
NH N
O
N H
OH OH
Dibutyltin oxide MeOH
O
O N HO
O OH
O
O
NH
N
N
N
N H
HO
2-Nitrobenzyl chloromethyl ether
O
O O
N
NH N
O
N H
O Sn
NO2
Bu
Bu
O DMTrCl/Pyridine
N DMTrO
N
O OH
NH
O
N
O
N H
O NO2
2 Figure 2.5.5 Preparation of 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N -isobutyrylguanosine from guanosine. Bu is n-butyl.
Materials Guanosine hydrate Anhydrous pyridine, stored over coarse granules of calcium hydride Isobutyryl chloride Ethyl acetate 1 M NaHCO3 2 M NaCl Anhydrous sodium sulfate Ethanol 2 N NaOH, ice cold Dowex AG50W-X8 ion exchange resin (pyridinium form) 5 × 60–cm glass chromatography column filled with 50 mL Dowex AG50W-X8 ion-exchange resin (pyridinium form) in water 4:1 (v/v) chloroform/methanol Phosphorus pentoxide Dibutyltin oxide Methanol Anhydrous dimethylformamide, stored over 4A molecular sieves 2-Nitrobenzyl chloromethyl ether, make fresh (see Support Protocol 1) Silica gel 60, 70 to 230 mesh ASTM (e.g., EM Science) 2.5 × 25–cm and 2.5 × 30–cm glass chromatography columns packed with silica gel 60, 70 to 230 mesh ASTM, in chloroform to a bed height of 20 and 25 cm, respectively (see Support Protocol 3) Chloroform 2%, 4%, 6%, and 8% (v/v) methanol in chloroform
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.15 Current Protocols in Nucleic Acid Chemistry
Supplement 4
9:1 (v/v) chloroform/methanol 80% (v/v) aqueous acetonitrile Acetonitrile Triethylamine, stored over coarse granules of calcium hydride 4,4′-Dimethoxytrityl chloride 0% to 1% (v/v) methanol in chloroform, containing 0.25% (v/v) pyridine 2:1 (v/v) ethyl acetate/hexane Oven, 130°C 500-mL and 2-L round-bottom flasks 100-mL pressure-equalizing dropping funnel Rotary evaporator connected interchangeably to water aspirator and vacuum pump 1-L separatory funnel Filter paper, coarse porosity and fast flow rate (e.g., Quantitative QB, Fisher) Water-cooled reflux condenser Heating mantle Additional reagents and equipment for TLC and column chromatography (see Support Protocol 3) NOTE: The pyridinium form of Dowex AG50W-X8 ion-exchange resin is made by washing the hydrogen form with several changes of 20% aqueous pyridine. Prepare tetraisobutyrylguanosine 1. Dry guanosine hydrate to constant weight in an oven at 130°C. A couple of hours drying is sufficient to drive off the water of crystallization.
2. Add 28.3 g (100 mmol) dry guanosine to 1 L anhydrous pyridine in a 2-L round-bottom flask while stirring. Fit flask with a 100-mL pressure-equalizing dropping funnel. The guanosine will form a suspension in the pyridine.
3. Cool mixture to 0°C in an ice bath and use the dropping funnel to add 100 mL isobutyryl chloride dropwise over a 10-min period with vigorous stirring at 0°C. Stir 2 hr at 0°C. 4. Use the dropping funnel to add 100 mL water dropwise to the stirring solution, slowly enough that the temperature is maintained <5°C. Allow mixture to stand overnight at 0°C. 5. Evaporate reaction mixture to ∼200 mL using a rotary evaporator connected to a vacuum pump. 6. Transfer slurry to a 4-L Erlenmeyer flask using 500 mL ethyl acetate. Stir vigorously while slowly adding 2 L of 1 M NaHCO3. Stir overnight at room temperature. CAUTION: The mixture froths violently as carbon dioxide is evolved.
7. Transfer mixture in portions to a 1-L separatory funnel, removing the water layer at each transfer until only the ethyl acetate layer remains in the funnel. Extract three times with 300 mL of 1 M NaHCO3 and three times with 300 mL of 2 M NaCl. 2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
8. Dry ethyl acetate solution over 10 g anhydrous sodium sulfate, filter through filter paper to remove salt, and rotary evaporate filtrate using a water aspirator to a foam consisting of tetraisobutyrylguanosine.
2.5.16 Supplement 4
Current Protocols in Nucleic Acid Chemistry
If desired, this intermediate can be stored at –20°C in a sealed dry container (stable for many months).
Prepare N2-isobutyrylguanosine 9. Add 11.3 g (20 mmol) tetraisobutyrylguanosine to 165 mL ethanol and stir to dissolve completely. Cool solution to 0°C in an ice bath. 10. Add 165 mL ice-cold 2 N NaOH to mixture while stirring at 0°C. Stir 5 min, then warm mixture quickly to room temperature and let stand 10 min. The timing of this step is critical.
11. Immediately pour solution into a 1.5-L beaker that contains 450 mL Dowex AG50WX8 ion-exchange resin (pyridinium form) while stirring. The pH should now be near neutral.
12. Pour contents of the beaker into a 5 × 60–cm glass chromatography column filled with 50 mL Dowex AG50W-X8 ion-exchange resin (pyridinium form) and wash column with 3 L water, collecting eluate. 13. Using the rotary evaporator connected to water aspirator, evaporate eluate to dryness, co-evaporate twice with 100 mL water, and then crystallize the residue from water. 14. Examine reaction mixture by TLC (see Support Protocol 3) in 4:1 chloroform/methanol. An 80% to 90% yield (based on the starting quantity of guanosine) of N2-isobutyrylguanosine is expected. A band with an Rf of 0.4 should be visible following TLC.
15. Dry the N2-isobutyrylguanosine in a vacuum desiccator over phosphorus pentoxide before use. If desired, this intermediate can be stored at –20°C in a sealed dry container (stable for many months).
Prepare 2′,3′-O-dibutylstannylene-N 2-isobutyrylguanosine 16. Add 5.3 g (15 mmol) dry N2-isobutyrylguanosine and 3.75 g (15 mmol) dibutyltin oxide to 750 mL methanol in a 2-L round-bottom flask while stirring. 17. Attach a water-cooled reflux condenser, heat to boiling with a heating mantle, reflux mixture 45 min, and allow to cool. 18. Rotary evaporate resulting clear solution using a water aspirator to remove methanol, then dry the white solid, 2′,3′-O-dibutylstannylene-N2-isobutyrylguano-sine, in a vacuum desiccator over phosphorus pentoxide. A yield of 8 g is expected. If desired, this intermediate can be stored at –20°C in a sealed dry container (stable for many months).
Prepare 2′-O-(2-nitrobenzyloxymethyl)-N2-isobutyrylguanosine 19. Add 6.51 g (11.2 mmol) 2′,3′-O-dibutylstannylene-N2-isobutyrylguanosine to 100 mL anhydrous dimethylformamide in a 500-mL round-bottom flask, while stirring to dissolve. Heat mixture to 60°C with a heating mantle. 20. Add freshly prepared 2-nitrobenzyl chloromethyl ether (from 16 mmol 2-nitrobenzyl methylthiomethyl ether as described; see Support Protocol 1) dropwise over a 5-min period while stirring at 60°C. Seal flask and stir 2 hr at room temperature.
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.17 Current Protocols in Nucleic Acid Chemistry
Supplement 3
21. Add 5 mL anhydrous pyridine, followed by 2 mL water while stirring. Stir 20 min at room temperature. 22. Concentrate mixture using the rotary evaporator connected to a vacuum pump to remove as much dimethylformamide as possible. If desired, this intermediate can be stored at –20°C in a sealed dry container (stable for many months).
Purify 2′-O-(2-nitrobenzyloxymethyl)-N2-isobutyrylguanosine 23. Dissolve residue in a minimum volume of anhydrous pyridine and, using a Pasteur pipet, transfer it dropwise onto 10 g silica gel 60 in a large petri plate. Allow wet silica to dry in the moving airstream inside a fume hood. An overnight drying time should be adequate if most of the dimethylformamide was removed and the minimum amount of pyridine was used.
24. Pulverize the dry, coated, crusty silica. Layer powder onto a 2.5 × 25–cm glass chromatography column and elute with 100 mL chloroform, followed by 200 mL of 2% methanol in chloroform, 200 mL of 4% methanol in chloroform, and 6% methanol in chloroform until all product is off the column (see Support Protocol 3). 25. Monitor fractions by TLC in 9:1 chloroform/methanol. 26. Combine appropriate fractions and rotary evaporate to dryness using a water aspirator. 27. Repeat steps 23 to 26, except use 2.5 g silica gel 60 in a 2.5 × 30–cm glass chromatography column and elute with 100 mL of 2% methanol in chloroform, 100 mL of 4% methanol in chloroform, 200 mL of 6% methanol in chloroform, then 8% methanol in chloroform. A yield of 1.7 g (30% of the starting quantity of 2′,3′-O-dibutylstannylene-N2-isobutyrylguanosine) is expected. If desired, this intermediate can be stored at –20°C in a sealed dry container (stable for many months).
28. Dissolve dry 2′-O-(2-nitrobenzyloxymethyl)-N 2-isobutyrylguanosine in a minimal amount of 80% aqueous acetonitrile with heating. 29. Using the rotary evaporator connected to a water aspirator, co-evaporate several times with acetonitrile to a volume of ~5 mL until crystallization of the desired 2′-isomer occurs. Additional crops may be obtained that require recrystallization. The 2′-isomer has a melting point of 176° to 177°C. If desired, this intermediate can be stored at –20°C in a sealed dry container (stable for many months).
Prepare 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N2isobutyrylguanosine 30. Using the rotary evaporator connected to vacuum pump, dry 1.2 g (2.32 mmol) 2′-O-(2-nitrobenzyloxymethyl)-N2-isobutyrylguanosine by two co-evaporations with 30 mL anhydrous pyridine. 2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
31. Dissolve residual oil in 30 mL anhydrous pyridine, then add 0.39 mL (2.8 mmol) triethylamine and 0.94 g (2.8 mmol) 4,4′-dimethoxytrityl chloride. Stir 3 hr at room temperature.
2.5.18 Supplement 3
Current Protocols in Nucleic Acid Chemistry
32. Add 0.5 mL methanol and, after 10 min, rotary evaporate using a vacuum pump to remove solvents. 33. Dissolve residue in 150 mL ethyl acetate and extract three times with 100 mL of 1 M NaHCO3 and once with 100 mL of 2 M NaCl. 34. Repeat step 8, using 5 g of anhydrous sodium sulfate. 35. Layer ~1-g portions of this material dissolved in a minimal amount of chloroform on 2.5 × 30–cm silica gel columns and elute with a 1-L stepwise gradient of 0% to 1% methanol in chloroform containing 0.25% pyridine. 36. Monitor fractions by TLC in 2:1 ethyl acetate/hexane. 37. Using the rotary evaporator connected to a water aspirator, evaporate appropriate fractions to produce 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N2isobutyrylguanosine. A yield of 1.8 g is expected.
38. Store the pure protected nucleoside in the dark at −20°C in a sealed container containing a trace of pyridine (stable for many months). PREPARATION OF N-PROTECTED 5′-O-(4,4′-DIMETHOXYTRITYL)2′-O-(tert-BUTYLDIMETHYLSILYL) NUCLEOSIDES The following protocols describe the preparation of an alternative set of ribonucleoside derivatives, utilizing the tert-butyldimethylsilyl group for protection of their 2′-hydroxyls (Figure 2.5.6). In the cases of adenosine and guanosine, the exocyclic amine functions of the nucleobases are protected by phenoxyacetyl groups; these form part of a quick deprotection strategy (see Commentary). As an adjunct to the synthesis procedures detailed below, information regarding the properties of the tert-butyldimethylsilyl-protected nucleosides can be found in Tables 2.5.5 and 2.5.6; these data should prove useful in characterizing the title compounds and their synthetic intermediates. CAUTION: Chromatographic solvents used in these protocols are flammable. Avoid spills and work in a well-vented fume hood away from potential ignition sources.
DMTrO
B
O
H3C TBDMS =
HO
OTBDMS
H3C
O HN
O B= N
N
O
uracil-1-yl
CH3
O Ph N O
4
N -benzoylcytosin-1-yl
OPh
HN
N
NH
CH3
H3C C Si
N 6
O N
N
N
N
N -phenoxyacetyladenin-9-yl
NH N
N H
O OPh
2
N -phenoxyacetylguanin-9-yl
Figure 2.5.6 The four 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl) ribonucleosides. The general structure of these ribonucleosides is in the upper-left corner and the four bases (B) are shown below. DMTr is the 4,4′-dimethoxytrityl group (Fig. 2.5.1), and TBDMS is the tert-butyldimethylsilyl group.
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.19 Current Protocols in Nucleic Acid Chemistry
Supplement 3
Table 2.5.5 Rf Values of tert-Butyldimethylsilyl-Protected Ribonucleosides and Their Precursors on Merck Silica Gel 60 F254 TLC Platesa
Ribonucleosideb
Rf value
Solvent system
APa
0.60 0.32 0.28 0.30 0.82 0.55 0.22 0.77 0.82 0.45
4:1 methylene chloride/methanol 4:1 methylene chloride/methanol 9:1 chloroform/methanol 9:1 chloroform/methanol 9:1 methylene chloride/methanol 9:1 methylene chloride/methanol 3:1 diethyl ether/hexane 19:1 chloroform/methanol 1:1 methylene chloride/ethyl acetate 1:1 methylene chloride/ethyl acetate
GPa 5′DMTrU 5′DMTrCBz 5′DMTrAPa 5′DMTrGPa 5′DMTrU2′TBDMS 5′DMTrCBz2′TBDMS 5′DMTrAPa2′TBDMS 5′DMTrGPa2′TBDMS a
Data are from Hakimelahi et al. (1982) and Chaix et al. (1989). Abbreviations: A, adenosine; C, cytidine; G, guanosine; U, uridine; Bz, benzoyl; DMTr, 4,4′-dimethoxytrityl; Pa, phenoxyacetyl; TBDMS, tert-butyldimethylsilyl. b
CAUTION: Use fresh diethyl ether when removal of this solvent by evaporation is required, otherwise explosive peroxides may develop. NOTE: Excellent chromatographic separations are obtained using the fine grade of silica gel recommended in Alternate Protocols 1 through 4. However, with small column diameters, solvents must be pushed through the silica by the application of external pressure, so flash chromatography columns are required. In other respects, the separation methods are based on information provided in Hakimelahi et al. (1982), Chaix et al. (1989), and Sinha et al. (1993). ALTERNATE PROTOCOL 1
Synthesis of 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)uridine In this protocol, uridine is converted into its 5′-O-(4,4′-dimethoxytrityl) derivative, which, upon reaction with tert-butyldimethylsilyl chloride in the presence of silver nitrate as catalyst, forms predominantly the desired 2′-O-(tert-butyldimethylsilyl) isomer. The reaction scheme is shown in Figure 2.5.7.
Table 2.5.6 1H-NMR Chemical Shiftsa of tert-Butyldimethylsilyl-Protected Purine Ribonucleosides and Their Synthetic Intermediatesb
2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
Hydrogen
APa
GPa
5′DMTrAPa 5′DMTrGPa
5′DMTrAPa2′ TBDMS
5′DMTrGPa2′ TBDMS
H1′ H2′ H3′ H4′ H5′′ H5′ H2 H8
6.21 4.98 4.55 4.30 3.92 3.92 8.70 8.76
6.01 4.78 4.54 4.22 3.91 3.91 NA 8.27
6.27 5.12 4.76 4.39 3.54 3.54 8.70 8.61
6.27 5.24 4.64 4.40 3.58 3.57 8.71 8.62
6.06 4.93 4.52 4.34 3.57 3.52 NA 8.18
6.05 4.87 4.59 4.33 3.51 3.51 NA 8.12
a The internal reference for 1H-NMR spectra was tetramethylsilane at 0 ppm. The solvent was acetone-d6. All shifts are measured in ppm. These data, from Chaix et al. (1989), are useful and commonly observed resonances. b Abbreviations: A, adenosine; G, guanosine; DMTr, 4,4′-dimethoxytrityl; Pa, phenoxyacetyl; TBDMS, tert-butyldimethylsilyl; NA, not applicable.
2.5.20 Supplement 3
Current Protocols in Nucleic Acid Chemistry
O
O NH
HO
O
N
NH
O
DMTrO
DMTrCl
O
N
O
Pyridine HO
OH
HO
OH
O NH DMTrO
HO
Figure 2.5.7 uridine.
N
O
1. TBDMS-Cl/AgNO3/THF/Pyridine 2. Separation of isomers
O
OTBDMS
Preparation of 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)uridine from
Materials Uridine Anhydrous pyridine, stored over coarse granules of calcium hydride 4,4′-Dimethoxytrityl chloride Methanol Methylene chloride 1 M NaHCO3 Anhydrous sodium sulfate 1:20 (v/v) chloroform/hexane 3 × 30–cm and 3 × 60–cm flash chromatography columns (with reservoirs and flow controller), packed with silica gel 60, 230 to 400 Mesh ASTM (see Support Protocol 3) Diethyl ether containing 0.25% (v/v) pyridine Ethyl acetate containing 0.25% (v/v) pyridine 9:1 (v/v) chloroform/methanol Anhydrous tetrahydrofuran Silver nitrate tert-Butyldimethylsilyl chloride 3:1 (v/v) diethyl ether/hexane 2 M NaCl Diethyl ether 30% (v/v) ethyl acetate in hexane 100- and 250-mL round-bottom flasks Rotary evaporator connected interchangeably to water aspirator and vacuum pump 250-mL separatory funnels Filter paper, coarse porosity and fast flow rate (e.g., Quantitative Q8, Fisher) Additional reagents and equipment for TLC and column chromatography (see Support Protocol 3) Prepare 5′-O-(4,4′-dimethoxytrityl)uridine 1. Add 2.44 g (10 mmol) uridine to 20 mL anhydrous pyridine in a 250-mL round-bottom flask, while stirring to dissolve. Seal flask and cool solution in an ice bath. Protection of Nucleosides for Oligonucleotide Synthesis
2.5.21 Current Protocols in Nucleic Acid Chemistry
Supplement 3
2. Add 0.5 g (1.5 mmol) 4,4′-dimethoxytrityl chloride and reseal flask. Stir at 0°C. Each hour thereafter for 7 hr, add 0.5 g more 4,4′-dimethoxytrityl chloride, for a total of 4 g (∼12 mmol). Stir an additional 3 hr at 0°C. 3. Add 3 mL methanol and, after 10 min, concentrate mixture to a small volume using a rotary evaporator connected to a vacuum pump. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
4. Add 50 mL methylene chloride and stir. Slowly add 50 mL of 1 M NaHCO3 and stir until evolution of carbon dioxide ceases. CAUTION: Carbon dioxide is evolved.
5. Transfer layers to a 250-mL separatory funnel with 50 mL methylene chloride and run off the organic layer into a second funnel. Wash organic layer with 50 mL water, dry it over 5 g anhydrous sodium sulfate, filter through filter paper to remove salt, and rotary evaporate filtrate to a thick oil using a water aspirator. 6. Triturate oil with 10 mL of 1:20 chloroform/hexane to remove most of the residual pyridine. Decant the wash. Purify 5′-O-(4,4′-dimethoxytrityl)uridine 7. Apply residue to a 3 × 60–cm silica gel flash chromatography column packed in diethyl ether. Wash with 300 mL diethyl ether containing 0.25% pyridine, elute with ethyl acetate containing 0.25% pyridine at a rate of 3 mL/min, and collect 15-mL fractions as described (see Support Protocol 3). CAUTION: Diethyl ether is a highly flammable solvent that should be handled with extreme care.
8. Assay fractions by TLC (see Support Protocol 3) using 9:1 chloroform/methanol. 5′-O-(4,4′-Dimethoxytrityl)uridine has an Rf of 0.28 (Table 2.5.5).
9. Pool appropriate fractions, rotary evaporate to dryness using a water aspirator, and crystallize the residue (∼4.5 g) from diethyl ether. 5′-O-(4,4′-Dimethoxytrityl)uridine has a melting point of 111° to 112°C. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
Prepare 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)uridine 10. Using the rotary evaporator connected to vacuum pump, dry 2.73 g (5 mmol) 5′-O-(4,4′-dimethoxytrityl)uridine by two co-evaporations with 25 mL anhydrous pyridine in a 100-mL round-bottom flask. 11. Dissolve residue in 50 mL anhydrous tetrahydrofuran, then add 1.5 mL (18.5 mmol) anhydrous pyridine and 1 g (6 mmol) silver nitrate. Stir until silver nitrate dissolves (∼5 min). 12. Add 1 g (6.6 mmol) tert-butyldimethylsilyl chloride and stir 5 hr at room temperature. 13. Examine reaction mixture by TLC in 3:1 diethyl ether/hexane. 2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
The 2′-isomer has an Rf of 0.22 and should be visible as the major product, running ahead of the 3′-isomer .
2.5.22 Supplement 3
Current Protocols in Nucleic Acid Chemistry
14. Filter reaction mixture through filter paper into 50 mL of 1 M NaHCO3, then extract with 300 mL methylene chloride. Extract organic layer with 100 mL of 2 M NaCl. 15. Dry organic layer over 10 g anhydrous sodium sulfate, filter through filter paper to remove salt, and rotary evaporate filtrate to an oil using a water aspirator. 16. Dissolve residue in 50 mL diethyl ether, filter to remove undissolved material, then remove solvent by rotary evaporation using a water aspirator without heating. 17. Layer mixture on a 3 × 30–cm silica gel flash chromatography column packed in hexane, elute using 30% ethyl acetate in hexane, and collect fractions as described. Some experimentation may be required to determine the precise mixture or gradient of ethyl acetate in hexane that gives optimum separation of isomers.
18. Monitor fractions by TLC using 3:1 diethyl ether/hexane and rotary evaporate fractions containing the 2′-isomer using a water aspirator to produce the pure product, 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)uridine. A yield of ∼2.3 g, or 70% based on the starting quantity of 5′-O-(4,4′-dimethoxytrityl)uridine, is expected. The product can be stored at –20°C in a sealed, dry container (stable for many months).
Synthesis of 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N4-benzoylcytidine
ALTERNATE PROTOCOL 2
In this protocol, cytidine is selectively benzoylated at its N4 position, then dimethoxytritylated to protect its 5′-hydroxyl. Subsequent reaction with tert-butyldimethylsilyl chloride in the presence of silver nitrate gives predominantly the 2′-O-(tert-butyldimethylsilyl) derivative. The reaction sequence is illustrated in Figure 2.5.8. Additional Materials (also see Alternate Protocol 1) Cytidine Benzoic anhydride Chloroform
O HN
NH2
N
N HO
HO
O
N
O
Benzoic anhydride/MeOH
HO
OH
O HO
N
O
OH
DMTrCl Pyridine
O
O
HN
HN N
DMTrO
O HO
N
N O
DMTrO
O
1. TBDMS-Cl/AgNO3/THF/Pyridine OTBDMS
2. Separation of isomers
HO
N
O
OH
Figure 2.5.8 Preparation of 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N4-benzoylcytidine from cytidine.
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.23 Current Protocols in Nucleic Acid Chemistry
Supplement 3
19:1 (v/v) chloroform/methanol 1-L round-bottom flasks Water-cooled reflux condenser Heating mantle Coarse sintered funnel Prepare N4-benzoylcytidine 1. Add 5 g (20.6 mmol) cytidine and 5 g (22.3 mmol) benzoic anhydride to 500 mL methanol in a 1-L round-bottom flask, while stirring. Fit flask with a water-cooled reflux condenser and heat mixture to a boil using a heating mantle. 2. At the end of each hour for the first 3 hr, briefly cool mixture and add an additional 5 g benzoic anhydride. Reflux for a total of 5 hr. Let mixture cool to 0°C. 3. Collect crystals of N4-benzoylcytidine by filtration through a coarse sintered funnel. Wash with 25 mL ice-cold methanol, then with diethyl ether to remove residual benzoic acid. A yield of 4.5 g product (with a melting point of 218° to 219°C) is expected. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
Prepare 5′-O-(4,4′-dimethoxytrityl)-N4-benzoylcytidine 4. Using a rotary evaporator connected to a vacuum pump, dry 3.5 g (10 mmol) N4-benzoylcytidine by two co-evaporations with 20 mL anhydrous pyridine. 5. Dissolve residue in 20 mL anhydrous pyridine in a 100-mL round-bottom flask while stirring. Seal flask and cool solution in an ice bath to 0°C. 6. Continue with dimethoxytritylation (see Alternate Protocol 1, steps 2 to 6), with the following exceptions: a. Increase methanol to 5 mL in step 3. b. Decrease methylene chloride and 1 M NaHCO3 to 25 mL in step 4. c. Wash organic layer with 50 mL of 2 M NaCl instead of water in step 5. d. Increase 1:20 chloroform/hexane to 50 mL in step 6. 7. Dissolve residue in 15 mL chloroform and apply to a 3 × 60–cm silica gel flash chromatography column packed in chloroform. Wash column with 100 mL chloroform and elute with 9:1 chloroform/methanol as described (see Support Protocol 3). 8. Assay fractions by TLC (see Support Protocol 3) in 9:1 chloroform/methanol. Combine fractions containing pure 5′-O-(4,4′-dimethoxytrityl)-N4 -benzoylcytidine and rotary evaporate to dryness using a water aspirator. This product can be crystallized (see Alternative Protocol 1, step 9), but this is probably not required. A yield of ∼5.5 g product (with a melting point of 109° to 111°C) is expected. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months). 2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
2.5.24 Supplement 3
Current Protocols in Nucleic Acid Chemistry
Prepare 5′-O-(4,4′-dimethoxytrityl)-2 ′-O-(tert-butyldimethylsilyl)-N4-benzoylcytidine 9. Prepare the 2′-O-tert-butyldimethylsilyl isomer as described (see Alternate Protocol 1, steps 10 to 18), but dry 3.25 g (5 mmol) 5′-O-(4,4′-dimethoxytrityl)-N4 -benzoylcytidine in step 10, and use 19:1 chloroform/methanol for TLC in steps 13 and 18. A yield of 2.5 g is expected.
Synthesis of 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N6-phenoxyacetyladenosine
ALTERNATE PROTOCOL 3
In this protocol, the exocyclic amine at the N6 position of adenosine is protected as a phenoxyacetyl amide, then a dimethoxytrityl group is added to the 5′-hydroxyl. Treatment with tert-butyldimethylsilyl chloride and imidazole produces a mixture of the 2′- and 3′-O-tert-butyldimethylsilyl derivatives, which are separated by column chromatography. The reaction sequence is shown in Figure 2.5.9. Materials Adenosine Anhydrous pyridine, stored over coarse granules of calcium hydride Trimethylchlorosilane 1-Hydroxybenzotriazole Anhydrous acetonitrile Phenoxyacetyl chloride Concentrated ammonium hydroxide Chloroform 90% (v/v) ethanol 4,4′-Dimethoxytrityl chloride 9:1 (v/v) methylene chloride/methanol Methanol Ethyl acetate 1 M NaHCO3, 5°C 2 M NaCl Anhydrous sodium sulfate
O
N HO
O HO
N
N
N
N
1. Me3SiCl/Pyridine
O
N
O HO
N N
OH
O OPh N 1. TBDMS-Cl/Imidazole/Pyridine
OPh
HN
N N
N
DMTrCl Pyridine
O HN
DMTrO
HO
2. Phenoxyacetyl chloride, 1-Hydroxybenzotriazole, MeCN-Pyridine (1:1 v/v) 3. NH4OH
OH
N
OPh
HN
NH2
DMTrO
O
N
N N
2. Separation of isomers HO
OTBDMS
HO
OH
Figure 2.5.9 Preparation of 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N6-phenoxyacetyladenosine from adenosine.
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.25 Current Protocols in Nucleic Acid Chemistry
Supplement 3
3 × 30–cm and 3 × 60–cm flash chromatography columns (with reservoirs and flow controller), packed with silica gel 60, 230 to 400 Mesh ASTM (see Support Protocol 3) 2% to 10% (v/v) ethanol in ethyl acetate Imidazole tert-Butyldimethylsilyl chloride 1:1 (v/v) methylene chloride/ethyl acetate 1% and 35% (v/v) ethyl acetate in hexane Rotary evaporator connected interchangeably to water aspirator and vacuum pump 250-mL round-bottom flasks with rubber septa 5- and 50-mL syringes and vent needles 100-mL pressure-equalizing dropping funnel 500-mL separatory funnel Filter paper, coarse porosity and fast flow rate (e.g., Quantitative Q8, Fisher) Additional reagents and materials for TLC and column chromatography (see Support Protocol 3) Prepare N6-phenoxyacetyladenosine 1. Using a rotary evaporator connected to a vacuum pump, dry 5.34 g (20 mmol) adenosine by two co-evaporations with 40 mL anhydrous pyridine in a 250-mL round-bottom flask. 2. Suspend residue in 100 mL anhydrous pyridine and add 19 mL (150 mmol) trimethylchlorosilane dropwise while stirring. Stir 25 min at room temperature. 3. Using the rotary evaporator connected to a water aspirator, dry 4.16 g (31.6 mmol) 1-hydroxybenzotriazole in a second 250-mL round-bottom flask by three co-evaporations with 30 mL anhydrous acetonitrile. Add 10 mL anhydrous acetonitrile and 10 mL anhydrous pyridine and stir to suspend residue. The formation of colored side products is prevented by the use of 1-hydroxybenzotriazole.
4. Fit flask with a rubber septum. Using a 5-mL syringe and vent needle, add 4.2 mL (30 mmol) phenoxyacetyl chloride dropwise through the septum, and stir 5 min at room temperature. 5. Cool both flasks in an ice bath to 0°C, then, using a 50-mL syringe and a vent needle, add the contents of the first flask (containing the adenosine) dropwise through the septum to the acylating agent in the second flask. Stir overnight at room temperature. 6. Cool flask to 5°C and add 20 mL cold water, followed by 10 mL concentrated ammonium hydroxide. Stir 15 min at 5°C. 7. Immediately remove ammonia and solvents by rotary evaporation using a water aspirator and dissolve residual gum in 600 mL water. Extract aqueous solution twice with 300 mL chloroform, then rotary evaporate to dryness using a water aspirator. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
8. Crystallize product from 90% ethanol. 2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
A yield of 5.2 g N6-phenoxyacetyladenosine (or 65% based on the starting quantity of adenosine) is expected. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
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Prepare 5′-O-(4,4′-dimethoxytrityl)-N6-phenoxyacetyladenosine 9. Using the rotary evaporator connected to a vacuum pump, dry 4 g (10 mmol) N6-phenoxyacetyladenosine by two co-evaporations with 10 mL anhydrous pyridine in a 250-mL round-bottom flask. 10. Dissolve residue in 60 mL anhydrous pyridine. Put a stir-bar in the flask, fit it with a 100-mL pressure-equalizing dropping funnel, and cool mixture to 5°C. 11. Dissolve 3.73 g (11 mmol) 4,4′-dimethoxytrityl chloride in 40 mL anhydrous pyridine, then add it dropwise over a 2-hr period to the N6-phenoxyacetyladenosine while stirring. Let stand overnight at 5°C. 12. Examine solution by TLC (see Support Protocol 3) in 9:1 methylene chloride/methanol. The 5′-O-(4,4′-dimethoxytrityl) derivative is the major product.
13. Add 5 mL methanol, stir 5 min, then rotary evaporate reaction mixture to a slurry using a vacuum pump. 14. Using 200 mL ethyl acetate, transfer residue to a 500-mL separatory funnel. Extract twice with 100 mL cold (5°C) 1 M NaHCO3 and once with 100 mL of 2 M NaCl. 15. Dry organic layer over 5 g anhydrous sodium sulfate, filter through filter paper to remove salt, and rotary evaporate filtrate to a yellow foam using a water aspirator. If desired, this intermediate can be stored at –20°C in a sealed, dry container (stable for many months).
16. Layer material on a 3 × 60–cm silica gel flash chromatography column packed in ethyl acetate and elute initially with ethyl acetate, then with a stepwise gradient of 2% to 10% ethanol in ethyl acetate as described (see Support Protocol 3). Use 200-mL portions of ethyl acetate, then 2% ethanol, 4% ethanol, and so on. 17. Monitor fractions by TLC using 9:1 methylene chloride/methanol and rotary evaporate appropriate fractions using a water aspirator to produce the pure product, 5′-O-(4,4′-dimethoxytrityl)-N6-phenoxyacetyladenosine. A 55% yield is expected based on the starting quantity of N6-phenoxyacetyladenosine.
Prepare 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N6-phenoxyacetyladenosine 18. Using the rotary evaporator connected to a vacuum pump, dry 2.8 g (4 mmol) 5′-O-(4,4′-dimethoxytrityl)-N6-phenoxyacetyladenosine by two co-evaporations with 10 mL anhydrous pyridine. Dissolve residue in 40 mL anhydrous pyridine. 19. Add 0.8 g (11.8 mmol) imidazole, followed by 0.35 g (2.4 mmol) tert-butyldimethylsilyl chloride. Stir 24 hr at room temperature. 20. Add another 0.35 g tert-butyldimethylsilyl chloride and stir 24 hr at room temperature. 21. Repeat step 20. 22. Check reaction by TLC in 1:1 methylene chloride/ethyl acetate. The starting material should be completely converted into its 2′-silyl, 3′-silyl, and 2′,3′-disilyl derivatives. The 2′,3′-disilyl derivative is the fastest running product, followed by the 2′-silyl derivative, and then the 3′-silyl derivative.
23. Cool mixture to 0°C and add 5 mL water.
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24. Rotary evaporate mixture to dryness using a vacuum pump, dissolve residue in 300 mL chloroform, and extract chloroform solution twice with 150 mL cold 1 M NaHCO3 and once with 150 mL water. Remove chloroform by rotary evaporation using a water aspirator. 25. Layer residual mixture on a 3 × 30–cm silica gel flash chromatography column packed in 1% ethyl acetate in hexane and elute with 35% ethyl acetate in hexane. Some experimentation may be required to determine the precise mixture or gradient of ethyl acetate in hexane that gives optimum separation of isomers.
26. Monitor fractions by TLC using 1:1 methylene chloride/ethyl acetate, combine fractions containing pure 2′-isomer, and rotary evaporate them using a water aspirator to produce the pure product, 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N6-phenoxyacetyladenosine. A yield of 35%, based on the starting quantity of 5′-O-(4,4′-dimethoxytrityl)-N6-phenoxyacetyladenosine, is expected. The product can be stored at –20°C in a sealed, dry container (stable for many months). ALTERNATE PROTOCOL 4
Synthesis of 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N2-phenoxyacetylguanosine In this protocol, the exocyclic amine at the N2 position of guanosine is protected as a phenoxyacetyl amide, then a 4,4′-dimethoxytrityl group is added to the 5′-hydroxyl. Treatment with tert-butyldimethylsilyl chloride and imidazole gives a mixture of the 2′- and 3′-O-tert-butyldimethylsilyl derivatives, which are separated by column chromatography. The reaction sequence is shown in Figure 2.5.10. This protocol is performed by following steps 1 to 26 of Alternate Protocol 3, with the exceptions outlined below. Additional Materials (also see Alternate Protocol 3) Guanosine hydrate 50% (v/v) ethyl acetate in hexane Oven, 130°C
O N HO
O HO
O N
NH
N
N
NH2
1. Me3SiCl/Pyridine
HO
2. Phenoxyacetyl chloride, 1-Hydroxybenzotriazole, MeCN-Pyridine (1:1 v/v) 3. NH4OH
OH
O OH
N
NH N
O OPh
N H
OH
DMTrCl Pyridine O N N
DMTrO
2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
O
O NH
N
N H
N
O OPh 1. TBDMS-Cl, Imidazole/Pyridine
DMTrO
O
N
NH N
N H
O OPh
2. Separation of isomers HO
OTBDMS
HO
OH
Figure 2.5.10 Preparation of 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N2-phenoxyacetylguanosine from guanosine.
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1a. Dry guanosine hydrate to constant weight in an oven at 130°C. Dry guanosine can be stored for many months in a sealed container at room temperature.
2a. Suspend 5.66 g (20 mmol) dry guanosine in 100 mL anhydrous pyridine and add 19 mL (150 mmol) trimethylchlorosilane dropwise while stirring. Stir 25 min at room temperature. 9a. Using a rotary evaporator connected to a vacuum pump, dry 4.12 g (10 mmol) of N2-phenoxyacetylguanosine by two co-evaporations with 10 mL anhydrous pyridine in a 250-mL round-bottom flask. 18a. Using the rotary evaporator connected to a vacuum pump, dry 2.9 g (4 mmol) 5′-O-(4,4′-dimethoxytrityl)-N2-phenoxyacetylguanosine by two co-evaporations with 10 mL anhydrous pyridine. Dissolve residue in 40 mL anhydrous pyridine. 25a. Layer residual mixture on a 3 × 30–cm silica gel flash chromatography column packed in hexane and elute with 50% ethyl acetate in hexane as described. Expected yields are: 60% N2-phenoxyacetylguanosine, 50% 5′-O-(4,4′-dimethoxytrityl)N2-phenoxyacetylguanosine, and 32% 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N2-phenoxyacetylguanosine, all based on the amount of starting material.
PREPARATION OF 2-NITROBENZYL CHLOROMETHYL ETHER 2-Nitrobenzyl chloromethyl ether is the alkylating agent used to convert nucleosides to their 2′-O-(2-nitrobenzyloxymethyl) derivatives. It is moderately unstable and must be prepared fresh each time it is used. The reaction is shown in Figure 2.5.11. The precursor, 2-nitrobenzyl methylthiomethyl ether, can be synthesized in quantity and stored for many months in the dark at −20°C.
SUPPORT PROTOCOL 1
CAUTION: A foul-smelling aqueous layer is produced in step 7. Work in a fume hood with yellow lights. Materials 2-Nitrobenzyl alcohol Dimethyl sulfoxide, dried over 4A molecular sieves Acetic anhydride Acetic acid 1:1 (v/v) diethyl ether/hexane Sodium bicarbonate 1:1 (v/v) ethyl acetate/hexane Saturated sodium bicarbonate 2 M NaCl Anhydrous sodium sulfate 5 × 60–cm chromatography column packed with silica gel 60, 70 to 230 mesh ASTM, in hexane to a bed height of 56 cm (see Support Protocol 3) 1%, 5%, 10%, and 15% (v/v) diethyl ether in hexane
NO2 OH
DMSO Ac2O/AcOH
NO2 O
SCH3
2-Nitrobenzyl methylthiomethyl ether
Figure 2.5.11
SO2Cl2
NO2
CH2Cl2
O
Cl
2-Nitrobenzyl chloromethyl ether
Preparation of 2-nitrobenzyl chloromethyl ether from 2-nitrobenzyl alcohol.
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Anhydrous methylene chloride Ethanol (for dry ice/ethanol bath) 1 M SO2Cl2 in methylene chloride Anhydrous dimethylformamide, store over 4A molecular sieves 500-mL separatory funnel Filter paper, coarse porosity and fast flow rate (e.g., Quantitative Q8, Fisher) Rotary evaporator connected to water aspirator 250-mL round-bottom flask 50-mL pressure-equalizing dropping funnel Additional reagents and equipment for TLC and column chromatography (see Support Protocol 3) CAUTION: The sulfuryl chloride (SO2Cl2) solution is corrosive and extremely toxic. Prepare 2-nitrobenzyl methylthiomethyl ether 1. Dissolve 23 g (150 mmol) 2-nitrobenzyl alcohol in 150 mL dry dimethyl sulfoxide in a 1-L flask. 2. Add 108 mL acetic anhydride and 77 mL acetic acid and stir to dissolve. Allow the homogeneous, yellow-tinted solution to stand 24 hr at room temperature in the dark. 3. Examine the mixture by TLC (see Support Protocol 3) using 1:1 diethyl ether/hexane. The fastest-running of the three UV-absorbing bands is the desired product.
4. Pour reaction mixture into a 500-mL separatory funnel, then add it dropwise over a 2-hr period to a vigorously stirred slurry of 330 g sodium bicarbonate in 1.5 L water. Stir 24 hr in the dark. The evolution of carbon dioxide is well controlled by the dropwise addition of the mixture.
5. Allow the oily product to settle to the bottom of the container, then decant the aqueous supernatant. 6. Dissolve oil in 400 mL of 1:1 ethyl acetate/hexane and extract this solution three times with 100 mL saturated sodium bicarbonate and once with 100 mL of 2 M NaCl. 7. Dry organic layer over 10 g anhydrous sodium sulfate, filter through filter paper to remove salt, and remove volatile organic solvents on a rotary evaporator connected to a water aspirator. The large volume of foul-smelling aqueous supernatant can be deodorized by treatment with household bleach before disposal.
Purify 2-nitrobenzyl methylthiomethyl ether 8. Layer yellow residue on a 5 × 60–cm silica gel column packed in 1% diethyl ether in hexane and elute at a rate of 5 to 10 mL/min using a stepwise gradient consisting of 500 mL of 1% diethyl ether in hexane followed by 1 L each of 5%, 10%, and 15% diethyl ether in hexane (see Support Protocol 3). Allow the first liter of eluate to run into a flask, then collect 250-mL fractions until the product begins to emerge, as judged by TLC using 1:1 diethyl ether/hexane. Thereafter, collect 100-mL fractions. 2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
CAUTION: These solvents are flammable.
9. Assay fractions by TLC using 1:1 diethyl ether/hexane. Pool fractions containing the pure product, 2-nitrobenzyl methylthiomethyl ether.
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10. Remove volatile solvents on the rotary evaporator to afford a viscous, yellow oil. A yield of 15 to 20 g is expected.
11. Store this stock reagent at −20°C in a sealed container in the dark (stable for many months). 1
H- and 13C-NMR data for this material can be found in Schwartz et al. (1992).
Prepare 2-nitrobenzyl chloromethyl ether 12. Dissolve 3.4 g (16 mmol) 2-nitrobenzyl methylthiomethyl ether in 30 mL anhydrous methylene chloride in a 250-mL round-bottom flask and fit flask with a 50-mL pressure-equalizing dropping funnel. 13. Cool mixture to −78°C using a dry ice/ethanol bath. 14. Using the pressure-equalizing dropping funnel, add 16 mL of 1 M SO2Cl2 in methylene chloride dropwise over a 5-min period while stirring at −78°C. Continue to cool the mixture and stir 30 min. 15. Quickly transfer flask to rotary evaporator and remove solvent and volatile products. 16. Repeat rotary evaporation three times with 20 mL anhydrous methylene chloride, then add 15 mL anhydrous dimethylformamide and use the clear solution immediately. PREPARATION OF PENTAFLUOROPHENYL BENZOATE Pentafluorophenyl benzoate is used to selectively protect cytidine at N4 in Basic Protocol 2.
SUPPORT PROTOCOL 2
Materials 2,3,4,5,6-Pentafluorophenol Anhydrous dimethylformamide, store over 4A molecular sieves Benzoic acid 1,3-Dicyclohexylcarbodiimide Ethyl acetate Ethanol Phosphorus pentoxide Filter paper, coarse porosity and fast flow rate (e.g., Quantitative Q8, Fisher) Rotary evaporator connected interchangeably to water aspirator and vacuum pump Coarse sintered funnel CAUTION: Pentafluorophenol, pentafluorophenyl benzoate, and 1,3-dicyclohexylcarbodiimide are toxic irritants. Avoid skin contact. 1. Add 2.9 g (15.7 mmol) 2,3,4,5,6-pentafluorophenol to a flask containing 7 mL anhydrous dimethylformamide and stir to dissolve. Stopper flask to minimize exposure to atmospheric moisture and cool in an ice bath. 2. Add 1.92 g (15.7 mmol) benzoic acid, followed by 3.5 g (17.9 mmol) 1,3-dicyclohexylcarbodiimide. Stir 1 hr at 0°C. A white suspension will be visible.
3. Warm mixture to room temperature and remove precipitate by suction filtration through filter paper.
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Supplement 3
4. Evaporate filtrate to dryness using a rotary evaporator under high vacuum (i.e., with a vacuum pump) and a water-bath temperature <40°C. 5. Dissolve residue in 20 mL ethyl acetate, filter through filter paper to remove undissolved solid, and evaporate filtrate to dryness using the rotary evaporator connected to a water aspirator. 6. Dissolve residue in ∼20 mL ethanol with gentle warming (<50°C). Evaporate to ∼10 mL using the rotary evaporator connected to water aspirator and cool to 0 °C. 7. Collect crystals of pentafluorophenyl benzoate by filtration through a coarse sintered funnel. Dry material over phosphorus pentoxide in a vacuum desiccator. 8. Store in an amber bottle at −20°C and make sure moisture is excluded (stable for many months). A yield of ∼2.5 g product (with a melting point of 76° to 78°C) is expected. SUPPORT PROTOCOL 3
2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
CHROMATOGRAPHIC TECHNIQUES Some skill in various techniques of chromatography, both analytical and preparative, are required in these protocols. Reaction mixtures are invariably analyzed by thin-layer chromatography (TLC) on 2 × 8–cm or 6 × 8–cm aluminum-backed sheets of Merck silica gel 60 F254 (e.g., EM Science). A 2-µL sample of the mixture to be analyzed is applied with a micropipettor in a thin 1-cm line to the origin on the TLC plate. After blowing off excess solvent in a rapidly moving stream of cool air, chromatography is carried out using an appropriate mixture of solvents in a developing jar (e.g., Eastman Kodak). The plate is then dried and placed under short-wave UV light in a viewing box. The products should be visible as well-defined bands against the fluorescent background of the TLC plate. If they are not, the original sample may need to be diluted with acetonitrile or ethyl acetate and reanalyzed. It should also be noted that when the original mixture contains pyridine, as is often the case, the residue of this solvent is visible on the chromatogram as a dark, diffuse band, which can be removed by heating the developed plate in an oven at 140°C for a few minutes. This treatment also causes dimethoxytrityl compounds, of which there are many in these protocols, to develop a characteristic yellow color. As a general rule, all reactions and all column fractions should be monitored by TLC. Moreover, it is very important that a reference sample (100 to 200 µL) of each reaction mixture be removed and stored (at −20°C) for checking against column fractions during the purification process. For additional details on TLC, see APPENDIX 3D. Preparative chromatographic separations to purify products are carried out on silica gel (e.g., EM Science) packed in glass chromatography columns using the starting solvent. The type and mesh size of the silica gel is important and is specified in each protocol. Columns are generally packed to a bed height of 5 to 10 cm below their full length to allow loading of the compound mixtures. For additional details on column chromatography, including general packing instructions, see APPENDIX 3E. The sample is applied as a narrow band at the top of the column, and products are eluted with increasingly polar solvent mixtures. Many of the separations in this unit are effected using stepwise gradients. A gradient described as, for example, 1.2 L of 0% to 10% methanol in chloroform refers to elution with 200 mL chloroform, then with 200 mL of 2% methanol in chloroform, then with 200 mL of 4% methanol in chloroform, and so on up to 10% methanol in chloroform. At that point, elution is continued with the 10% solvent mixture, if necessary, until all product is removed from the column, as determined by TLC. The columns are run at such a rate that elution is complete in 6 to 8 hr. The eluate emerging from the column is collected as 15- to 20-mL fractions in 18 × 150–mm disposable glass
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culture tubes using a fraction collector. The fractions are then assayed by TLC to locate the desired product. To reduce the number of analyses, every fifth fraction, for example, can be analyzed by TLC, and then these results can be used to narrow down the remaining fractions to assay. COMMENTARY Background Information Chemical synthesis of RNA is now almost as simple as that of DNA. However, it requires a specialized set of monomers derived from the four common ribonucleosides U, C, A, and G; each of these monomers must carry an extra protecting group to prevent reaction at its 2′hydroxyl. The protocols in this unit describe the syntheses of two useful sets of protected ribonucleosides. These products are ready to be converted into their corresponding phosphoramidites for use in RNA synthesis (UNIT 3.7; Sinha et al., 1993). 2-Nitrobenzyloxymethyl protection The first set consists of the 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl) derivatives of U and of N-protected C, A, and G. In Basic Protocols 1 and 3, respectively, the readily available nucleosides U and A are first converted into their 2′,3′-O-dibutylstannylene derivatives (Wagner et al., 1974). Their 2′,3′glycol systems are susceptible to monoalkylation after this step. The alkylation is carried out using 2-nitrobenzyl chloromethyl ether and a catalyst, tetra-n-butylammonium bromide. This reaction is nonselective, however, and results in formation of the 2′-O-(2-nitrobenzyloxymethyl) derivatives of U and A along with equal amounts of their respective 3′-isomers. In the case of adenosine, the 2′- and 3′-isomers can be separated chromatographically. The pure 2′-isomer is isolated and protected at its N6 position by benzoylation, using the transient protection method of Ti et al. (1982). Its 5′-hydroxyl is then protected by reaction with 4,4′-dimethoxytrityl chloride, to give 5′-O-(4,4′dimethoxytrityl)-2′ -O-(2-nitrobenzyloxymethyl)N6-benzoyladenosine. In contrast, the mixture of 2′- and 3′-O-(2nitrobenzyloxymethyl)uridines is first protected at the 5′-hydroxyls, and then chromatographically separated, affording pure 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)uridine. In Basic Protocol 4, guanosine is alkylated after protection of its exocyclic amine with an isobutyryl group (Ohtsuka et al., 1978).
Subsequent conversion to its dibutylstannylene derivative and alkylation in the absence of a catalyst leads to exclusive production of the 2′-O-(2-nitrobenzyloxymethyl) isomer. This material is purified and 5′-protected to give 5′-O(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-N2-isobutyrylguanosine. Cytidine, because of the susceptibility of its aminopyrimidone ring to side-alkylation (Wagner et al., 1974), cannot be derivatized in the same way as the other ribonucleosides. Accordingly, some of the 5′-O-(4,4′-dimethoxytrityl)2′-O-(2-nitrobenzyloxymethyl)uridine obtained in Basic Protocol 1 is converted directly into its corresponding cytidine derivative in Basic Protocol 2, using the substitution procedure of Sung (1982). Its N4 amino group is then selectively benzoylated using pentafluorophenyl benzoate (Igolen and Morin, 1980) to give 5′-O-(4,4′-dimethoxytrityl)-2′-O(2-nitrobenzyloxymethyl)-N4-benzoylcytidine. After conversion to their respective phosphoramidites, the protected nucleosides can be used in the construction of RNA (UNIT 3.7). The particular advantage of these amidites lies in their short coupling times; they allow RNA to be synthesized almost as rapidly as DNA (Schwartz et al., 1992). Although alternative groups for protecting the 2′-hydroxyls of ribonucleosides are now available that allow coupling times to be reduced nearly to those for protected deoxyribonucleoside phosphoramidites, this was not the case as recently as 3 to 5 years ago. Upon completion of synthesis, the 2-nitrobenzyloxymethyl groups are removed from the resulting oligoribonucleotide products by irradiation with long-wave UV light. tert-Butyldimethylsilyl protection The second set of protected ribonucleosides has the 2′-hydroxyl protected with the tertbutyldimethylsilyl group (Alternate Protocols 1 to 4). This particular group has been used extensively in RNA synthesis (Wincott et al., 1995), and phosphoramidite monomers incorporating it are widely available commercially. The strategy for preparing 2′-O-(tert-butyldimethylsilyl) ribonucleosides in the laboratory
Protection of Nucleosides for Oligonucleotide Synthesis
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Supplement 3
differs in a number of respects from that used for the corresponding nitrobenzyloxymethyl derivatives. Again, the starting materials are the four common ribonucleosides, but in this case the exocyclic amines on C, A, and G are all protected first. Cytidine is selectively benzoylated at its N4 position with benzoic anhydride in boiling methanol (Watanabe and Fox, 1966); adenosine and guanosine are phenoxyacetylated at their N6 and N2 positions, respectively, using the transient protection method (Ti et al., 1982). These N-acyl derivatives and uridine are then dimethoxytritylated to protect their 5′ positions (Hakimelahi et al., 1982; Chaix et al., 1989). The silyl groups are introduced last, by treating the 5′-O-(4,4′-dimethoxytrityl)-N-acyl nucleosides with tert-butyldimethylsilyl chloride (Hakimelahi et al., 1982; Chaix et al., 1989). Mixtures of the 2′- and 3′-isomers are formed in each case, from which the pure 2′O-(tert-butyldimethylsilyl) nucleosides are isolated by column chromatography. They can then be converted into their phosphoramidites using standard procedures (Sinha et al., 1984, 1993). The tert-butyldimethylsilyl groups can be quantitatively removed from oligoribonucleotides by treatment with tetra-n-butylammonium fluoride (Usman et al., 1987). They are also somewhat sensitive to ammonia, however, and their loss during the ammonia treatment that is part of the standard oligoribonucleotide deprotection procedure has caused problems in the past by leading to strand cleavage of the product RNA. Much effort has been expended, therefore, in finding ways to make the ammonolysis conditions less severe. One such method is replacement of the traditional but relatively stable benzoyl and isobutyryl groups on A and G, respectively, with more base-labile ones such as phenoxyacetyl (Chaix et al., 1989) and tert-butylphenoxyacetyl (Sinha et al., 1993). It is the phenoxyacetyl derivatives whose preparation is described in this unit.
Compound Characterization
2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
Relevant Rf values and available NMR data are shown in Tables 2.5.1 to 2.5.6. Other properties, such as the melting points of crystalline compounds, can be found at the appropriate places in the protocols, and further data are provided in the cited literature. Methods for distinguishing and identifying 2′- and 3′-isomers of derivatized ribonucleosides by NMR are described in Fromageot et al. (1966) and Chaix et al. (1989).
Critical Parameters The success of the operations described here can be affected by a number of critical factors. For example, many of the reagents and reaction mixtures are sensitive to water, and precautions should be taken to exclude moisture. All reactions, unless otherwise specified, should be conducted with dry starting material and anhydrous solvents. The air inlet to the rotary evaporator should be connected to a drying tower, so that no moist air is drawn into the apparatus. If necessary, materials should be co-evaporated with dry solvent a few times before being redissolved in the required reaction volume. Flasks should be opened only long enough for addition or removal of materials, and cold flasks or reagent bottles should always be allowed to reach room temperature before being opened. Some of the protecting groups used in these protocols are labile, as indeed they are usually designed to be. The 4,4′-dimethoxytrityl group, for example, is removed rapidly under acidic conditions such as those found on some types of silica TLC plates, in silica-gel columns, and as contaminants in solvents such as chloroform. Detritylation and loss of product can be prevented under these circumstances by adding trace amounts of pyridine. The phenoxyacetyl group, on the other hand, is quite base-labile, and alkaline treatments should be strictly limited. The 2′-O-(tert-butyldimethylsilyl) ribonucleosides should also be treated with care. The tert-butyldimethylsilyl group is known to isomerize between the 2′- and 3′-hydroxyls of the ribose ring in protic solvents such as methanol and in aqueous solutions such as pyridine in water; these environments should be avoided. The 2-nitrobenzyloxymethyl group is sensitive to UV light and is slowly removed even under normal conditions of laboratory illumination. Exposure of nitrobenzyloxmethyl compounds to UV should be minimized by replacing the overhead white fluorescent bulbs in the work area with yellow ones. Flasks and chromatography columns should be wrapped in foil, and storage containers should be kept in the dark. The most important consideration of all is the requirement for absolute isomeric purity of the 2′-protected ribonucleosides described here. These products should be examined scrupulously by loading them heavily on TLC plates and looking for the presence of traces of contaminating 3′-isomer after chromatogra-
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phy. If any of the 3′-isomer is found, the suspect nucleoside should be repurified; it is better to sacrifice a small amount of yield for absolute purity. The consequences of isomeric contamination can be put in perspective by realizing that the presence of the 3′-isomer at a level of 1% in the monomers means that, for an oligonucleotide 100 units long, not a single molecule is likely to possess all natural 3′→5′ linkages.
Anticipated Results The experimentalist can expect to obtain one to several grams of each protected nucleoside, ready for phosphitylation and use in an automated synthesizer. By repetition and/or judicious scaling up of the reactions, one can accumulate multigram quantities. Each gram of material corresponds to upwards of 100 nucleotide incorporations on a 0.2-µmol synthesis scale.
Time Considerations The many partially protected intermediates in these protocols can be safely stored either as crude mixtures ready for chromatographic purification or as purified materials awaiting the next synthesis step. They should be stored in sealed dry containers at −20°C and are stable for many months under these conditions. Within the protocols, each set of preparation and purification steps can be completed in 1 to 2 days, with the following exceptions: preparing the cytidine and benzoylcytidine derivatives in Basic Protocol 2 requires 6 and 7 days, respectively; purifying the isobutyryl guanosine derivative in Basic Protocol 4 requires 3 days; and preparing the tert-butyldimethylsilyl derivatives in Alternate Protocols 3 and 4 requires 4 days. Support Protocols 1 and 2 can be completed in 3 to 4 days and 1 day, respectively. After some familiarity with the methods has been achieved, ∼2 weeks should be required to complete each protocol.
Literature Cited Benneche, T., Strande, P., and Undheim, K. 1983. A new synthesis of chloromethyl benzyl ethers. Synthesis 1983:762-763. Chaix, C., Duplaa, A.M., Molko, D., and Teoule, R. 1989. Solid phase synthesis of the 5′-half of the initiator t-RNA from B. subtilis. Nucl. Acids Res. 17:7381-7393. Fromageot, H.P.M., Griffin, B.E., Reese, C.B., Sulston, J.E., and Trentham, D.R. 1966. Orientation of ribonucleoside derivatives by proton magnetic resonance spectroscopy. Tetrahedron 22:705710.
Hakimelahi, G.H., Proba, Z.A., and Ogilvie, K.K. 1982. New catalysts and procedures for the dimethoxytritylation and selective silylation of ribonucleosides. Can. J. Chem. 60:1106-1113. Igolen, J. and Morin, C. 1980. Rapid synthesis of protected 2′-deoxycytidine derivatives. J. Org. Chem. 45:4802-4804. Ohtsuka, E., Nakagawa, E., Tanaka, T., Markham, A.F., and Ikehara, M. 1978. Studies on transfer ribonucleic acids and related compounds. XXI. Synthesis and properties of guanine rich fragments from E. coli tRNAfMet 5′ end. Chem. Pharm. Bull. 26:2998-3006. Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., deBear, J.S., and Gough, G.R. 1992. Rapid synthesis of oligoribonucleotides using 2′-O-(2-nitrobenzyloxymethyl)-protected monomers. BioMed. Chem. Lett. 2:1019-1024. Sinha, N.D., Biernat, J., McManus, J., and Köster, H. 1984. Polymer support oligonucleotide synthesis XVIII: Use of β-cyanoethyl-N,N-dialkylamino-/N-morpholino phosphoramidite of deoxynucleosides for the synthesis of DNA fragments simplifying deprotection and isolation of the final product. Nucl. Acids Res. 12:45394557. Sinha, N.D., Davis, P., Usman, N., Perez, J., Hodge, R., Kremsky, J., and Casale, R. 1993. Labile exocyclic amine protection of nucleosides in DNA, RNA and oligonucleotide analog synthesis facilitating N-deacylation, minimizing depurination and chain degradation. Biochimie 75:1323. Sung, W.L. 1982. Synthesis of 4-(1,2,4-triazol-1yl)pyrimidin-2(1H)-one ribonucleotide and its application in synthesis of oligoribonucleotides. J. Org. Chem. 47:3623-3628. Ti, G.S., Gaffney, B.L., and Jones, R.A. 1982. Transient protection: Efficient one-flask synthesis of protected deoxynucleosides. J. Am. Chem. Soc. 104:1316-1319. Usman, N., Ogilvie, K.K., Jiang, M.-Y., and Cedergren, R.J. 1987. Automated chemical synthesis of long oligoribonucleotides using 2′-O-silylated ribonucleoside 3′-O-phosphoramidites on a controlled pore glass support: Synthesis of a 43-nucleotide sequence similar to the 3′-half molecule of an Escherichia coli formylmethionine tRNA. J. Am. Chem. Soc. 109:7845-7854. Wagner, D., Verheyden, J.P.H., and Moffatt, J.G. 1974. Preparation and synthetic utility of some organotin derivatives of nucleosides. J. Org. Chem. 39:24-30. Watanabe, K.A. and Fox, J.J. 1966. A simple method for selective acylation of cytidine on the 4-amino group. Angew. Chem., Int. Ed. Engl. 5:579-580. Wincott, F., DiRenzo, A., Shaffer, C., Grimm, S., Tracz, D., Workman, C., Sweedler, D., Gonzalez, C., Scaringe, S., and Usman, N. 1995. Synthesis, deprotection, analysis and purification of RNA and ribozymes. Nucl. Acids Res. 23:2677-2684.
Protection of Nucleosides for Oligonucleotide Synthesis
2.5.35 Current Protocols in Nucleic Acid Chemistry
Supplement 3
Key References
Wincott et al., 1995. See above.
Schwartz et al., 1992. See above.
This publication describes some recent advances in RNA synthesis using the tert-butyldimethylsilyl group.
The general strategy involved in using the 2′-O-(2nitrobenzyloxymethyl protecting group is presented by this method’s developers. Usman et al., 1987. See above. The general strategy involved in using the 2′-O(tert-butyldimethylsilyl) protecting group is presented by this method’s developers.
Contributed by Tod J. Miller, Miriam E. Schwartz, and Geoffrey R. Gough Purdue University West Lafayette, Indiana
2′-OH-Protecting Groups That Are Photochemically Labile or Sensitive to Fluoride Ions
2.5.36 Supplement 3
Current Protocols in Nucleic Acid Chemistry
Deoxyribo- and Ribonucleoside H-Phosphonates
UNIT 2.6
This unit includes a collection of four synthetic methods for nucleoside H-phosphonate building blocks and a variant on one of these syntheses. All the methods are quite efficient and experimentally simple, and they use readily available reagents. Depending on reactivity and reaction conditions, different methods may be preferred for preparation of different building blocks. The phosphonylation of protected nucleosides with the in situ–generated tris-(1-imidazolyl)phosphine (from phosphorus trichloride and imidazole), followed by hydrolysis of the formed nucleoside diimidazolyl phosphite intermediate, is described in Basic Protocol 1. This method (or its variants with other azoles) is probably the most widely used and can be recommended as a general basic method applicable for the preparation of both deoxyribo- and ribonucleoside 3′-H-phosphonates. Imidazole from the first published procedure (Garegg et al., 1986c) is preferred over the other azoles (e.g., triazole; Froehler et al., 1986) because it is less expensive and forms the least reactive species. Since tris-(1-imidazolyl)phosphine is a trifunctional reagent, it should be used in excess in the phosphonylation reaction to avoid the formation of symmetrical H-phosphonate diesters. There are several variants of this protocol that differ in the kind of azoles, external bases, or solvents used, including procedures for ribonucleoside building blocks that combine 2′-O-protection and 3′-O-phosphonylation. The Alternate Protocol describes one of these procedures (also see Commentary). In Basic Protocol 2, pyridinium H-pyrophosphonate, generated in situ from phosphorous acid and a condensing agent, is used. This one-step reaction is most likely carried out by a nucleophilic attack of nucleoside on a phosphorus center of H-pyrophosphonate. Usually 5 mol eq in excess of the nucleoside is used to speed up the reaction. The chemical nature of a condensing agent is of minor importance for the final yield of the H-phosphonate monoesters. Phosphonylation of ribonucleosides protected with 2′-O-tertbutyldimethylsilyl (2′-O-TBDMS) occurs very slowly with this reagent, which makes it less suitable for use in preparation of ribonucleoside building blocks. Basic Protocol 3 describes a convenient approach for the preparation of nucleoside H-phosphonate monoesters based on transesterification of diphenyl H-phosphonate with protected nucleosides. Diphenyl H-phosphonate is relatively inexpensive, commercially available, stable, and easy to handle. It also gives high yields of nucleoside H-phosphonates. This is the newest of the presented methods and thus the least used, but some clear advantages of the method could make it a generally preferred choice. It should, however, be noted that synthesis of ribonucleoside building blocks with this method requires more basic work-up conditions, which may prevent its use with N-protections that are quite base labile. Basic Protocol 4 makes use of 2-chloro-4H-1,3,2-benzo-dioxaphosphinan-4-one (salicylchlorophosphite) as the phosphonylating reagent. The reaction with nucleosides is usually fast and quantitatively produces a phosphite intermediate that is converted into the H-phosphonate monoester upon hydrolysis. Since salicylchlorophosphite is practically a monofunctional phosphonylating agent, it can be used in nearly stoichiometric amounts. The formation of symmetrical dinucleoside H-phosphonate diesters is usually negligible, but removal of byproducts from the reagent can sometimes be a problem. Protection of Nucleosides for Oligonucleotide Synthesis Contributed by Jacek Stawinski and Roger Strömberg Current Protocols in Nucleic Acid Chemistry (2001) 2.6.1-2.6.15 Copyright © 2001 by John Wiley & Sons, Inc.
2.6.1 Supplement 4
NOTE: These reactions should be carried out under strictly anhydrous conditions, using anhydrous solvents and reagents. All glassware should be dried in an oven prior to use. The nucleosidic component should be rendered anhydrous by consecutive evaporation of added pyridine or acetonitrile depending on solvent used for the reaction. All connections to atmospheric pressure should be through a drying tower containing a desiccant. BASIC PROTOCOL 1
SYNTHESIS OF PROTECTED DEOXYRIBONUCLEOSIDE OR RIBONUCLEOSIDE 3′-H-PHOSPHONATES USING THE PHOSPHORUS TRICHLORIDE/IMIDAZOLE/TRIETHYLAMINE REAGENT This is a general-purpose protocol, applicable to both ribo- and deoxyribonucleosides bearing standard protecting groups. This reaction is shown in Figure 2.6.1. Materials Imidazole, made anhydrous by repeated evaporation of added dry acetonitrile Dichloromethane (99.9+%; HPLC grade), stored over 3A molecular sieves Acetone (for dry ice/acetone bath) Phosphorus trichloride, freshly distilled Triethylamine, distilled, stored over calcium hydride Protected deoxyribo- or ribonucleoside (see Chapter 2), dried by repeated evaporation of added pyridine 9:1 (v/v) chloroform/methanol 2 M TEAB, pH 7.5 (see recipe) Sodium sulfate 5 × 25–cm glass chromatography column, packed with Merck silica gel 60, 230 to 400 mesh ASTM (e.g., EM Science) or matrix silica, 35 to 70 µm (Millipore) in chloroform (bed height, 15 to 20 cm) Methanol Chloroform 0.5:1:8.5 or 1:2:7 (v/v/v) concentrated ammonia/water/isopropanol 1:1 (v/v) hexane/diethyl ether TLC silica-gel plates (e.g., Merck silica gel 60 F254, EM Science) Rotary evaporator connected to water aspirator Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E)
Deoxyribo- and Ribonucleoside H-Phosphonates
Figure 2.6.1
Synthesis of nucleoside 3′-H-phosphonates using PCl3/Im/TEA reagent system.
2.6.2 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Prepare phosphorus trichloride/imidazole/triethylamine reagent 1. Dissolve 5.8 g imidazole (86 mmol) in 300 mL dichloromethane and cool solution to approximately −10°C in a dry ice/acetone bath. Dry acetonitrile and tetrahydrofuran can be used as alternative solvents, but with dichloromethane the work-up procedure is more convenient.
2. While stirring, add dropwise 2.45 mL phosphorus trichloride (28 mmol) and then 12.5 mL triethylamine (90 mmol) mixed with 10 mL dichloromethane. Stir 15 to 30 min at 0° to −10°C and then cool to −78°C in dry ice/acetone bath. Prepare 3′-H-phosphonates 3. Add a protected deoxyribo- or ribonucleoside (8 mmol) dissolved in 200 mL dichloromethane dropwise over ∼30 min while stirring at −78°C. Stir 1 hr at −78°C. Steps 1 to 3 can be somewhat simplified by cooling the mixture with an ice/water bath instead; however, this is not recommended for guanosine and deoxyguanosine derivatives because substantial amounts of side products, and thus a lower yield, can be produced following subsequent reactions (presumably with the lactam system).
4. Monitor reactions for disappearance of the starting nucleoside and the formation of a baseline material by TLC (APPENDIX 3D) on silica-gel plates using 9:1 chloroform/methanol as the solvent. 5. When the reaction is complete (as determined by TLC analysis), pour the cold mixture directly onto 300 mL of 2 M TEAB and wash the organic layer with an additional 300 mL of 2 M TEAB. With acetonitrile as solvent, 20 mL of 0.5 M TEAB should be added to the cold mixture. The mixture is then concentrated under reduced pressure (at 30° to 40°C) and the residue is dissolved in 300 mL dichloromethane and washed twice with 150 mL of 2 M TEAB.
6. Separate the organic layer, dry it over ~20 g sodium sulfate, and evaporate solvent under reduced pressure using a rotary evaporator connected to a water aspirator. Purify 3′-H-phosphonates 7. Purify crude product by short-column silica-gel chromatography (APPENDIX 3E) on a 5 × 25–cm glass chromatography column using a stepwise gradient of methanol in chloroform to elute products. For example, a stepwise gradient of 0% to 12% methanol in chloroform containing 0.1% triethylamine can be used. Unwanted isomers of the ribonucleoside H-phosphonates, that is 3′-TBDMS (or 2-ClBz) 2′-H-phosphonates, if present, are readily removed by silica-gel chromatography.
8. Check eluate fractions for H-phosphonate purity by TLC using 0.5:1:8.5 or 1:2:7 (v/v/v) concentrated ammonia/water/isopropanol as the solvent. The polarity, and thus the mobility on TLC, of an H-phosphonate will be affected by its protecting group. The latter solvent is probably better for more polar compounds. For compounds bearing more base-labile N-protection, the use of 3:7 or 2:8 (v/v) isopropanol/1 M TEAB can be advantageous.
9. Combine fractions containing the desired product and concentrate them to a white foam using the rotary evaporator. Store at <4°C. A yield of 70% to 95% 3′-H-phosphonate monoesters (triethylammonium salts), based on the staring quantity of nucleoside, is expected.
Protection of Nucleosides for Oligonucleotide Synthesis
2.6.3 Current Protocols in Nucleic Acid Chemistry
Supplement 4
Occasionally, DBUH+ salts of H-phosphonate monoesters can be preferred to TEAH+ salts. The transformation can be effected by washing a solution of nucleoside H-phosphonate monoester (triethylammonium salt) in dichloromethane with 0.2 M 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) bicarbonate buffer (pH 8.5).
10. Optional: Dissolve foam in a small amount of dichloromethane and add it dropwise while stirring vigorously to 100 to 150 mL of 1:1 hexane/diethyl ether per mmol product. Store at <4°C. The nucleoside H-phosphonate is stored as a microcrystalline solid. Although stable for several months at room temperature, it should be stored in the freezer to prevent decomposition due to residual solvents. Under such conditions, no decomposition is seen even after several months. ALTERNATE PROTOCOL
SYNTHESIS OF RIBONUCLEOSIDE 3′-H-PHOSPHONATES VIA IN SITU 2′-O-2-CHLOROBENZOYLATION FOLLOWED BY PHOSPHONYLATION This procedure is specific for ribonucleosides bearing a 2′-O-chlorobenzoyl group and permits simultaneous 2′-O-benzoylation and 3′-O-H-phosphorylation as a “one pot” reaction. Additional Materials (also see Basic Protocol 1) Protected ribonucleoside with free 2′- and 3′-hydroxyls (see Chapter 2), dried by repeated evaporation of added pyridine Pyridine (HPLC grade; e.g., LabScan), dried by and stored over 4A molecular sieves 2-Chlorobenzoyl chloride 1. Dissolve a protected ribonucleoside with free 2′- and 3′-hydroxyls (8 mmol) in 95 mL dichloromethane and add 5 mL pyridine. Cool to −78°C in a dry ice/acetone bath while stirring. 2. Mix 1.18 mL 2-chlorobenzoyl chloride (8.8 mmol) and 10 mL dichloromethane, and add dropwise over a 15-min period. Incubate 30 min at −78°C with stirring. 3. Prepare phosphorus trichloride/imidazole/triethylamine reagent as described (see Basic Protocol 1, steps 1 and 2). 4. Add phosphorus trichloride/imidazole/triethylamine reagent dropwise over 15 to 30 min to nucleoside mixture while stirring at −78°C. Stir 1 hr at −78°C. 5. Continue with synthesis and purification as described (see Basic Protocol 1, steps 4 to 10). A yield of 40% to 70%, based on the starting quantity of nucleoside, is expected.
BASIC PROTOCOL 2
SYNTHESIS OF PROTECTED DEOXYRIBONUCLEOSIDE 3′-H-PHOSPHONATES USING H-PYROPHOSPHONATE This procedure is suitable for phosphorylation of hydroxyl groups in deoxyribonucleosides or 5′-hydroxyl groups in ribonucleosides. This reaction is shown in Figure 2.6.2.
Deoxyribo- and Ribonucleoside H-Phosphonates
Materials Protected deoxyribonucleoside (see Chapter 2), dried by repeated evaporation of added pyridine 2 M phosphorous acid (H3PO3) solution (see recipe)
2.6.4 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Figure 2.6.2
Synthesis of nucleoside 3′-H-phosphonates using H-pyrophosphonate.
Condensing agent (select one): 5,5-Dimethyl-2-oxo-2-chloro-1,3,2-dioxaphosphinane (NEPCl; see recipe) Pivaloyl chloride (PV⋅Cl), commercial grade (e.g., Aldrich), distilled before use (store <2 months at −20°C) 9:1 (v/v) chloroform/methanol 1 M and 2 M TEAB, pH 7.5 (see recipe) Dichloromethane (HPLC grade), stored over 3A molecular sieves Sodium sulfate 4 × 25–cm glass chromatography column, packed with Merck silica gel 60, 230 to 400 mesh ASTM (e.g., EM Science) or Matrix silica, 35 to 70 µm (Millipore) in chloroform (bed height, 10 to 15 cm) Methanol Chloroform 0.5:1:8.5 or 1:2:7 (v/v/v) concentrated ammonia/water/isopropanol 1:1 (v/v) hexane/diethyl ether TLC silica-gel plates (e.g., Merck silica gel 60 F254, EM Science) Rotary evaporator connected to water aspirator Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) 1. Dissolve protected deoxyribonucleoside (2 mmol) in 10 mL of 2 M H3PO3 solution (20 mmol). 2. Stir reaction mixture and slowly add 2.0 g NEPCl (11 mmol) or 1.35 mL PV⋅Cl (11 mmol). Stir ∼3 hr. The reaction can be left overnight at 20°C (room temperature). The specified amount of a condensing agent should not be exceeded, otherwise more reactive species can be generated from phosphorous acid, which may result in a significant formation of symmetrical H-phosphonate diesters. Instead of generating H-pyrophosphonate in situ from phosphorous acid and a condensing agent, a 1 M stock solution of this phosphonylating reagent can be prepared by adding 0.5 mol eq PV⋅Cl or NEPCl to 2 M H3PO3 in pyridine. The stock solution of pyridinium H-pyrophosphonate in pyridine is stable for several months at room temperature.
3. Monitor reactions for disappearance of the starting nucleoside and the formation of a baseline material by TLC (APPENDIX 3D) on silica-gel plates using 9:1 chloroform/methanol as the solvent.
Protection of Nucleosides for Oligonucleotide Synthesis
2.6.5 Current Protocols in Nucleic Acid Chemistry
Supplement 4
4. When the reaction is complete (as determined by TLC analysis), add 2 mL of 1 M TEAB and partition reaction mixture between 75 mL dichloromethane and 50 mL of 2 M TEAB. 5. Separate the organic layer, dry it over ~10 g sodium sulfate, and evaporate solvent under reduced pressure using a rotary evaporator connected to a water aspirator. 6. Purify crude product by short-column silica-gel chromatography (APPENDIX 3E) on a 4 × 25–cm glass chromatography column using a stepwise gradient of methanol in chloroform to elute products. For example, a stepwise gradient of 0% to 12% methanol in chloroform containing 0.1% triethylamine can be used. Unwanted isomers of the ribonucleoside H-phosphonates, that is 3′-TBDMS (or 2-ClBz) 2′-H-phosphonates, if present, are readily removed by silica-gel chromatography.
7. Check eluate fractions for H-phosphonate purity by TLC using 0.5:1:8.5 or 1:2:7 (v/v/v) concentrated ammonia/water/isopropanol as the solvents. The polarity, and thus the mobility on TLC, of an H-phosphonate will be affected by its protecting group. The latter solvent is probably better for more polar compounds. For compounds bearing more base-labile N-protection, the use of 3:7 or 2:8 (v/v) isopropanol/1M TEAB can be advantageous.
8. Combine fractions containing the desired product and concentrate them to a white foam using the rotary evaporator. Store at <4°C. A yield of 85% to 95% 3′-H-phosphonate monoesters (triethylammonium salts), based on the starting quantity of nucleoside, is expected. Occasionally, DBUH+ salts of H-phosphonate monoesters can be preferred to TEAH+ salts. The transformation can be effected by washing a solution of nucleoside H-phosphonate monoester (triethylammonium salt) in dichloromethane with 0.2 M 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) bicarbonate buffer (pH 8.5). When NEPCl is used to generate H-pyrophosphonate from phosphorous acid, the isolated nucleoside H-phosphonate monoesters can be occasionally contaminated by hydrolysis products of the condensing agent. If this is the case (as shown by 31P NMR), the product should be dissolved in 20 mL dichloromethane and washed twice with 5 mL of 0.05 M TEAB buffer.
9. Optional: Dissolve the foam in a small amount of dichloromethane and add it dropwise while stirring vigorously to 100 to 150 mL of 1:1 hexane/diethyl ether per mmol product. Store at <4°C. The nucleoside H-phosphonate is stored as a microcrystalline solid. Although stable for several months at room temperature, it should be stored in the freezer to prevent decomposition due to residual solvents. Under such conditions, no decomposition is seen even after several months. BASIC PROTOCOL 3
SYNTHESIS OF PROTECTED NUCLEOSIDE 3′-H-PHOSPHONATES USING DIPHENYL H-PHOSPHONATE This is a general-purpose procedure, applicable to both ribo- and deoxyribonucleosides. One should exercise some caution when base-sensitive protecting groups are present in the substrates (see Commentary). This reaction is shown in Figure 2.6.3.
Deoxyribo- and Ribonucleoside H-Phosphonates
2.6.6 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Figure 2.6.3
Synthesis of nucleoside 3′-H-phosphonates using diphenyl H-phosphonate.
Materials Protected deoxyribo- or ribonucleoside (see Chapter 2), dried by repeated evaporation of added pyridine Pyridine (HPLC grade; e.g., Labscan), dried by and stored over 4A molecular sieves Diphenyl H-phosphonate, commercial grade (e.g., Aldrich) 9:1 (v/v) chloroform/methanol Triethylamine, distilled, stored over calcium hydride 5% (w/v) sodium bicarbonate Dichloromethane, distilled Sodium sulfate 4 × 25–cm glass chromatography column, packed with Merck silica gel 60, 230 to 400 mesh ASTM (e.g., EM Science) or Matrix chloroform (bed height, 10 to 15 cm) Methanol Chloroform 0.5:1:8.5 or 1:2:7 (v/v/v) concentrated ammonia/water/isopropanol 1:1 (v/v) hexane/diethyl ether TLC silica-gel plates (e.g., Merck silica gel 60 F254, EM Science) Rotary evaporator connected to water aspirator Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) 1. Add a protected deoxyribo- or ribonucleoside (1 mmol) to 5 mL pyridine and stir. Add 1.34 mL diphenyl H-phosphonate (7 mmol) in one portion. Stir ∼15 min. An excess of phosphonylating agents is necessary to avoid the formation of symmetrical dinucleoside H-phosphonate diesters. Diphenyl H-phosphonate may be listed by some suppliers as diphenyl phosphite. In the instance of 2′-O-protected ribonucleosides, the amount of diphenyl H-phosphonate can be reduced to 3 mol eq.
2. Monitor reactions for disappearance of the starting nucleoside and the formation of a baseline material by TLC (APPENDIX 3D) on silica-gel plates using 9:1 chloroform/methanol as the solvent. 3. When the reaction is complete (as determined by TLC analysis), add 1 mL water and 1 mL triethylamine to quench the reaction, and let mixture stand 15 min.
Protection of Nucleosides for Oligonucleotide Synthesis
2.6.7 Current Protocols in Nucleic Acid Chemistry
Supplement 4
4. Use a rotary evaporator connected to a water aspirator to evaporate solvents, and partition the residue between 20 mL dichloromethane and 20 mL of 5% sodium bicarbonate. 5. Extract organic phase twice with 20 mL of 5% sodium bicarbonate, dry it over ~10 g sodium sulfate, and evaporate the solvent to produce an oil. Most phenol and phenyl H-phosphonates formed during the reaction and from hydrolysis of diphenyl H-phosphonate are removed by the aqueous sodium bicarbonate extraction. Crude reaction products (after extraction with aqueous sodium bicarbonate) can be precipitated from dichloromethane into 1:1 (v/v) hexane/diethyl ether to produce nucleoside H-phosphonates of >95% purity, without chromatography.
6. Purify crude product by short-column silica-gel chromatography (APPENDIX 3E) on a 4 × 25–cm glass chromatography column using a stepwise gradient of methanol in chloroform to elute products. For example, a stepwise gradient of 0% to 12% methanol in chloroform containing 0.1% triethylamine can be used. Unwanted isomers of the ribonucleoside H-phosphonates, that is 3′-TBDMS (or 2-ClBz) 2′-H-phosphonates, if present, are readily removed by silica-gel chromatography.
7. Check eluate fractions for H-phosphonate purity by TLC using 0.5:1:8.5 or 1:2:7 concentrated ammonia/water/isopropanol. The polarity, and thus the mobility on TLC, of an H-phosphonate will be affected by its protecting group. The latter solvent is probably better for more polar compounds. For compounds bearing more base-labile N-protection, the use of 3:7 or 2:8 (v/v) isopropanol/1M TEAB can be advantageous.
8. Combine fractions containing the desired product and concentrate them in the rotary evaporator to a white foam. Store at <4°C. A yield of 75% to 90% 3′-H-phosphonate monoesters (triethylammonium salts), based on the starting quantity of nucleoside, is expected.
9. Optional: Dissolve the foam in a small amount of dichloromethane and add it dropwise while stirring vigorously to 100 to 150 mL of 1:1 hexane/diethyl ether per mmol product. Store at <4°C. The nucleoside H-phosphonate is stored as a microcrystalline solid. Although stable for several months at room temperature, it should be stored in the freezer to prevent decomposition due to residual solvents. Under such conditions, no decomposition is seen even after several months. BASIC PROTOCOL 4
SYNTHESIS OF PROTECTED NUCLEOSIDE 3′-H-PHOSPHONATES USING 2-CHLORO-4H-1,3,2-BENZO-DIOXAPHOSPHINAN-4-ONE This is a general-purpose protocol applicable to both ribo- and deoxyribonucleotides. This reaction is shown in Figure 2.6.4.
Deoxyribo- and Ribonucleoside H-Phosphonates
Materials Protected deoxyribo- or ribonucleoside (see Chapter 2), dried by repeated evaporation of added pyridine 2:1 (v/v) dichloromethane/pyridine 1.25 M 2-chloro-4H-1,3,2-benzo-dioxaphosphinan-4-one solution (see recipe) 9:1 (v/v) chloroform/methanol 1 M TEAB, pH 7.5 (see recipe)
2.6.8 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Figure 2.6.4
Synthesis of nucleoside 3′-H-phosphonates using salicylchlorophosphite.
Dichloromethane (99%; HPLC grade), stored over 3A molecular sieves Sodium sulfate 5 × 25–cm glass chromatography column, packed with Merck silica gel 60, 230 to 400 mesh ASTM (e.g., EM Science) or Matrix silica, 35 to 70 µm (Millipore) in chloroform (bed height, 10 to 15 cm) Methanol Chloroform 0.5:1:8.5 or 1:2:7 (v/v/v) concentrated ammonia/water/isopropanol 1:1 (v/v) hexane/diethyl ether TLC silica-gel plates (e.g., Merck silica gel 60 F254, EM Science) Rotary evaporator connected to water aspirator Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) 1. Dissolve a protected deoxyribo- or ribonucleoside (1 mmol) in 20 mL of 2:1 dichloromethane/pyridine while stirring and cool solution with an ice/water bath. 2. Add 1 mL of 1.25 M 2-chloro-4H-1,3,2-benzo-dioxaphosphinan-4-one solution in one portion, while stirring. Remove from bath and stir 15 min. 3. Monitor reactions for disappearance of the starting nucleoside and the formation of a baseline material by TLC (APPENDIX 3D) on silica-gel plates using 9:1 chloroform/methanol as the solvent. 4. When the reaction is complete (as determined by TLC analysis), add 100 mL of 1 M TEAB to quench reaction and extract the aqueous phase three times with 50 mL dichloromethane. Collect and pool organic phases. 5. Dry organic phase over ~10 g sodium sulfate, and evaporate solvent under reduced pressure using a rotary evaporator connected to a water aspirator. 6. Purify crude product by short-column silica-gel chromatography (APPENDIX 3E) on a 5 × 25–cm glass chromatography column using a stepwise gradient of methanol in chloroform to elute products. For example, a stepwise gradient of 0% to 12% methanol in chloroform containing 0.1% triethylamine can be used.
Protection of Nucleosides for Oligonucleotide Synthesis
2.6.9 Current Protocols in Nucleic Acid Chemistry
Supplement 4
Unwanted isomers of the ribonucleoside H-phosphonates, that is 3′-TBDMS (or 2-ClBz) 2′-H-phosphonates, if present, are readily removed by silica-gel chromatography.
7. Check eluate fractions for H-phosphonate purity by TLC using 0.5:1:8.5 or 1:2:7 concentrated ammonia/water/isopropanol as the solvents. The polarity, and thus the mobility on TLC, of an H-phosphonate will be affected by its protecting group. The latter solvent is probably better for more polar compounds. For compounds bearing more base-labile N-protection, the use of 3:7 or 2:8 (v/v) isopropanol/1M TEAB can be advantageous. With nucleoside H-phosphonate monoesters produced in this way, it can sometimes be troublesome to remove contaminants resulting from the decomposition products of the phosphonylating reagent (these are visible as fluorescent spots on TLC plates under a UV lamp). In such cases, the samples should be rechromatographed.
8. Combine fractions containing the desired product and concentrate them in the rotary evaporator to a white foam. Store at <4°C. A yield of 80% to 90% 3′-H-phosphonate monoesters (triethylammonium salts), based on the starting quantity of nucleoside, is expected. Occasionally, DBUH+ salts of H-phosphonate monoesters can be preferred to TEAH+ salts. The transformation can be effected by washing a solution of nucleoside H-phosphonate monoester (triethylammonium salt) in dichloromethane with 0.2 M 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) bicarbonate buffer (pH 8.5).
9. Optional: Dissolve the foam in a small amount of dichloromethane and add it dropwise while stirring vigorously to 100 to 150 mL of 1:1 hexane/diethyl ether per mmol product. Store at <4°C. The nucleoside H-phosphonate is stored as a microcrystalline solid. Although stable for several months at room temperature, it should be stored in the freezer to prevent decomposition due to residual solvents. Under such conditions, no decomposition is seen even after several months.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
2-Chloro-4H-1,3,2-benzo-dioxaphosphinan-4-one solution, 1.25 M Prepare crystalline 2-chloro-4H-1,3,2-benzo-dioxaphosphinan-4-one (salicylchlorophosphite) as described (Young, 1952) or purchase commercially (e.g., Aldrich). Dissolve 2.52 g in 6 mL dioxane and adjust final volume to 10 mL. Prepare fresh. Store crystalline product several months at 4°C. Salicylchlorophosphite was previously called 2-chloro-4H-1,3,2-benzodioxaphosphorin-4one and may still be listed as such by some suppliers.
5,5-Dimethyl-2-oxo-2-chloro-1,3,2-dioxaphosphinane (NEPCl) Prepare this crystalline reagent as described (McConnell and Coover, 1959). Store several months at room temperature. Phosphorous acid (H3PO3) solution, 2 M Evaporate 1.62 g H3PO3 (commercial grade, e.g., Aldrich) from dry pyridine (HPLC grade, dried and stored over 4A molecular sieves) and dissolve residue in 8 mL dry pyridine. Adjust final volume to 10 mL. Stored several months at room temperature. Deoxyribo- and Ribonucleoside H-Phosphonates
Phosphorous acid in pyridine is prepared according to Stawinski and Thelin (1990a).
2.6.10 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Triethylammonium bicarbonate (TEAB), 2 M, pH 7.5 Bubble CO2 through a suspension of 280 mL triethylamine (2 mol) in 700 mL water until the pH of the solution reaches 7.5. Adjust final volume to 1 L. Store buffer up to a few weeks at 4°C and check pH before use. Dilute as needed to appropriate molarity. COMMENTARY Background Information The methods for the preparation of H-phosphonate monoesters of natural products can be divided into four major types. They are based on: (1) esterification of phosphorous acid in the presence of various condensing reagents (Hata and Sekine, 1974; Holy et al., 1965; Schofield and Todd, 1961; Sekine and Hata, 1975); (2) reactions of phosphorus trichloride (or reactive species derived thereof) with alcohols (Holy and Sorm, 1966; Honjo et al., 1966; Sekine et al., 1979); (3) transesterification of trialkyl or triaryl phosphites with appropriate hydroxylic components (Holy and Smrt, 1966); and (4) transesterification of β-substituted dialkyl Hphosphonate diesters with alcohols, followed by hydrolysis of the produced mixed diesters (Gibbs and Larsen, 1984; Takaku et al., 1988). Most of these methods, unfortunately, suffer from variable yields and are often incompatible with common protecting groups used in oligonucleotide synthesis. This unit describes four variants of the above general types for the preparation of nucleoside H-phosphonate monoesters: Basic Protocol 2 is a type 1 reaction; Basic Protocol 1, the Alternate Protocol, and Basic Protocol 4 are type 2 reactions; and Basic Protocol 3 is related to the type 4 reaction. These methods consistently give high yields of the desired products (75% to 90%), and taken together they provide an arsenal of phosphonylation (phosphitylation) procedures that are compatible with most common protecting groups used in oligodeoxyribo- and oligoribonucleotide synthesis. Phosphonylation using phosphorus trichloride Phosphonylation of protected nucleosides using the phosphorus trichloride/imidazole/triethylamine reagent system (Basic Protocol 1) is a mild and efficient method (with yields of 75% to 90%) for the preparation of protected deoxyribo- (Garegg et al., 1986a,c) and ribonucleoside 3′-H-phosphonates (Garegg et al., 1986b; Stawinski et al., 1988; Rozners et al., 1995b, 1998) as well as for arabinonucleoside H-phosphonates (Rozners et al., 1995a). A popular variant of this proce-
dure involves the replacement of imidazole by 1,2,4-triazole (Froehler, 1993; Froehler et al., 1986). Gaffney and Jones (1988) also reported that using N-methylmorpholine instead of triethylamine as an acid scavenger offers some advantages. Acetonitrile and dichloromethane are commonly used as solvents in this procedure. The phosphonylating reagent is generated in situ immediately prior to synthesis and the reaction is frequently carried out in a dry ice/acetone bath (or a similar bath well below 0°C), which is particularly beneficial with guanine-containing nucleosides as these are prone to side reactions. (The side products that can be formed are readily removed, but they will lower the yields.) The reagent is probably made less suitable for a large-scale production of building blocks, because of the use of a sub-zero cold bath. A convenient procedure for one-flask 2′-hydroxyl protection (2-chlorobenzoylation) and subsequent phosphonylation using this reagent (Alternate Protocol) has also been developed (Rozners et al., 1990, 1992, 1995b, 1998) and used for the preparation of formacetal-linked (Rozners and Strömberg, 1997) as well as amide-linked (D. Katkevica, E. Rozners, E. Bizdena, and R. Strömberg, unpub. observ.) dimeric H-phosphonate building blocks. Another recent procedure for preparation of purine-containing ribonucleoside building blocks is based on the reaction of tris(triazolyl)phosphine with 2′,3′-unprotected purine ribonucleosides to produce, after hydrolysis, an isomeric mixture of 2′- and 3′-H-phosphonates. Upon silylation with TBDMS⋅Cl, this mixture gives a remarkable selectivity of 85% to 90% for 2′-O-silylation (Zhang et al., 1997). Phosphonylation using phosphorous acid esterification Direct condensation of phosphorous acid with appropriate alcohols in the presence of an arene sulfonyl derivative (Sekine and Hata, 1975) to produce H-phosphonate monoesters is of little synthetic value due to the concomitant formation of H-phosphonate diesters (Sekine et al., 1988) and oxidation of H-phosphonate monoesters (Garegg et al.,
Protection of Nucleosides for Oligonucleotide Synthesis
2.6.11 Current Protocols in Nucleic Acid Chemistry
Supplement 4
1987). However, by using a limited amount of a condensing agent such as pivaloyl chloride or 5,5-dimethyl-2-oxo-2-chloro-1,3,2dioxaphosphinane (McConnell and Coover, 1959; Basic Protocol 2), it is possible to convert phosphorous acid into its H-pyrophosphonate (Stawinski and Thelin, 1990b), which in turn may act as a mild phosphonylating agent (Stawinski and Thelin, 1990a). The procedure is experimentally simple and reliable, and produces deoxyribonucleoside 3′-H-phosphonate in yields of 86% to 92% (Stawinski and Thelin, 1990a). Pyridinium H-pyrophosphonate can also be prepared and stored for several months at room temperature as a stock solution in pyridine. Reaction mixtures containing a nucleoside and 5 mol eq of pyrophosphonate can be left overnight without the danger of side reactions with heterocyclic base residues because of this reagent’s moderate reactivity (Stawinski and Thelin, 1990a). However, this reagent is rather slow to react with sterically hindered hydroxyl functions such as the 3′-hydroxyl group in 2′O-TBDMS ribonucleosides (Stawinski and Thelin, 1990a), and thus cannot be recommended for preparation of ribonucleoside Hphosphonate building blocks. The mild reaction conditions, experimental simplicity, and inexpensive starting materials make this method a strong candidate for large-scale preparations of deoxyribonucleotide H-phosphonates.
Deoxyribo- and Ribonucleoside H-Phosphonates
Phosphonylation by transesterification Transesterification of diphenyl H-phosphonate with protected nucleosides (Basic Protocol 3) is a most convenient approach for the preparation of H-phosphonate building blocks. Diphenyl H-phosphonate is an inexpensive and commercially available reagent that is stable and easy to handle. H-Phosphonate monoesters are synthesized with a purity usually >95% even without column chromatography (Jankowska et al., 1994). No side reactions involving the heterocyclic bases were detected. The same observation has also been reported for nucleosides with unprotected nucleobases (Wada et al., 1997). Although the reagent in pyridine undergoes disproportionation (Kers et al., 1996), this process is much slower than the transesterification reaction, and the side products formed (phenyl H-phosphonate and triphenyl phosphite) are easily removed during subsequent purification steps. Diphenyl H-phosphonate usually has to be used in a 3 to 7 mol eq excess of the nucleoside
to minimize the formation of symmetrical Hphosphonate diesters. However, by introducing some changes into the protocol (Jankowska et al., 1994), it is possible to reduce significantly the excess of diphenyl H-phosphonate. The method has been used for the preparation of both deoxyribo- and ribonucleoside H-phosphonates. It may, however, pose some problems when applied to ribonucleosides carrying more base-labile protection (e.g., N-phenoxyacetyl protection), due to the relatively basic conditions required for hydrolysis of 2′-O-TBDMSprotected nucleoside 3′-(phenyl H-phosphonate). Phosphonylation using salicylchlorophosphite Nucleoside H-phosphonate monoesters are also easily accessible via the salicylchlorophosphite method (Basic Protocol 4), which utilizes 2-chloro-4H-1,3,2-benzo-dioxaphosphinan-4-one as the phosphonylating reagent (Marugg et al., 1986). Salicylchlorophosphite is crystalline, stable, readily prepared (Young, 1952), and also commercially available. Although the phosphonylation is virtually quantitative, some amounts of the hydroxylic component (∼5%) are occasionally regenerated during hydrolysis of the phosphite intermediate (Froehler, 1993). The most serious inconvenience of the method is that it often can be difficult to separate the nucleoside Hphosphonate from hydrolysis products of the phosphonylating reagent. Some other synthetic methods that have been used for preparation of nucleoside Hphosphonate monoesters are: (1) phosphonylation of nucleosides with 2-cyanoethyl phosphordiamidite, hydrolysis to the H-phosphonate diester, and β-elimination of the 2-cyanoethyl group (Garegg et al., 1985); (2) phosphonylation of nucleosides with 2-cyanoethyl H-phosphonate, followed by β-elimination (Szabó et al., 1995); (3) condensation of aryl H-phosphonates with nucleosides, followed by hydrolysis (Ozola et al., 1996); (4) anionic debenzylation of nucleoside benzyl Hphosphonates (Hall et al., 1957); (5) oxidative phosphonylation with phosphinic acid in the presence of mesitylenedisulfonyl chloride (Sekine et al., 1988); (6) reaction of nucleoside bis-(N,N-di-isopropylamino)chlorophosphine, followed by acidolysis (Marugg et al., 1986); and (7) condensation of phosphorous acid with nucleosides promoted by triphosgene (Bhongle and Tang, 1995).
2.6.12 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Critical Parameters For these protocols, anhydrous reaction conditions must be used, especially if the phosphonylating reagent is present in almost stoichiometric amounts. Solvents should be stored over molecular sieves (or be freshly distilled). All glassware should be dried in an oven prior to use. The air inlets of apparatuses used in syntheses should be connected to atmospheric pressure through a drying tower. All nucleosides should be anhydrous, which is usually most conveniently accomplished by repeated evaporation of added pyridine. The purity of the produced compounds should be checked by NMR spectroscopy (31P and 1H), but TLC is also most convenient, especially for the detection of minor amounts of impurities. For example, the contamination of ribonucleoside H-phosphonate building blocks with the 3′-O-protected 2′-H-phosphonate isomers can be conveniently detected by TLC analysis, as these separate well in ammonia/water/isopropanol systems (and usually also in isopropanol/TEAB). In the phosphorus trichloride/imidazole method, the slow addition of nucleoside (see Basic Protocol 1, step 3) is important to reduce the formation of symmetrical H-phosphonate diesters. It is also important to note that a low temperature is required to avoid side reactions on guanine-containing nucleosides. Although the procedure is rather insensitive to the kind of azole used, there is a higher risk of formation of the symmetrical H-phosphonate esters using triazole (or tetrazole) rather than imidazole due to the higher reactivity of the former reagents. Imidazole is also less costly. If overly concentrated solutions or too small an excess of reagent is used, or if the nucleoside solution is added too quickly, smaller or larger amounts of symmetrical H-phosphonate diesters can be formed. These can be easily hydrolyzed during subsequent purification (giving monoester product and nucleoside), but their formation will lower the yield of the H-phosphonate monoesters. For the pyrophosphonate method, pyridine seems to be an optimal solvent. Strong bases (e.g., triethylamine) or basic nucleophilic catalysts (e.g., N-methylimidazole) should be avoided, because they usually slow down the phosphonylation. When NEPCl is used to generate H-pyrophosphonate, the resulting Hphosphonate monoesters should be checked for impurities arising from the condensing agent (e.g., by 31P NMR). During the phosphonylation of nucleosides, two species are formed
from the condensing agent: 5,5-dimethyl-2oxo-2-hydroxy-1,3,2-dioxaphosphinane and its pyrophosphate. The first product is usually completely removed during the extraction step. The second product is easy to remove during chromatography, but may slowly hydrolyze on silica gel, releasing 5,5-dimethyl-2-oxo-2-hydroxy-1,3,2-dioxaphosphinane. Transesterification of diphenyl H-phosphonate also works best when pyridine is used as solvent and base. The reaction in neutral solvents in the presence of a strong base (e.g., triethylamine) may fail due to rapid disproportionation of the phosphonylation reagent under such conditions. Formation of symmetrical H-phosphonates is usually not detected when salicylchlorophosphite is used as a phosphonylating reagent. The purity of nucleoside H-phosphonates prepared by this method should be carefully checked, however, for the presence of fluorescent spots visible on TLC plates under UV light. These are due to contamination by hydrolysis products of the phosphonylating reagent, which may be nontrivial to remove by silica-gel chromatography.
Anticipated Results All methods described in this unit afford, after silica-gel column chromatography, nucleoside H-phosphonate monoesters of comparable quality and in comparable yields, but they vary in their suitability for the preparation of different nucleoside derivatives. All methods can be scaled up to ~25-mmol reactions. A 1-mmol scale generates 0.5 to 1 g product and a 25-mmol scale generates 15 to 25 g. Compounds prepared by any of these methods are suitable for solid-phase as well as solution synthesis of oligonucleotides and their analogs. For deoxyribonucleoside H-phosphonates bearing quite stable N-acyl protection, all methods can produce good results. For ribonucleoside H-phosphonates, the pyrophosphonate method seems to be too slow, although it may be preferred in large-scale preparation of deoxyribonucleoside building blocks (another possible choice for this being the diphenyl H-phosphonate method). For use with ribonucleosides carrying more base-labile N-protection, the phosphorus trichloride/imidazole method is still preferred, with the salicylchlorophosphite approach being the second alternative. Protection of Nucleosides for Oligonucleotide Synthesis
2.6.13 Current Protocols in Nucleic Acid Chemistry
Supplement 4
Time Considerations None of the procedures described in this unit are very time consuming. The whole synthesis procedure including workup usually does not take more than a few hours. The pyrophosphonate method is the slowest one, but in this instance the reaction mixtures can conveniently be left overnight. Together with chromatographic purification, it will take from half a day to a day, depending on method, to obtain the purified nucleoside H-phosphonates. The method that probably involves the least manual effort and technical skill is the diphenyl Hphosphonate procedure.
Literature Cited Bhongle, N.N. and Tang, J.Y. 1995. A convenient synthesis of nucleoside 3′-H-phosphonate monoesters using triphosgene. Tetrahedron Lett. 36:6803-6806. Froehler, B.C. 1993. Oligodeoxynucleotide synthesis. H-Phosphonate approach. In Protocols for Oligonucleotides and Analogs (S. Agrawal, ed.) pp. 63-80. Humana Press, Totowa, N.J. Froehler, B.C., Ng, P.G., and Matteucci, M.D. 1986. Synthesis of DNA via deoxynucleoside H-phosphonate intermediates. Nu cl. Acids Res. 14:5399-5407. Gaffney, B.L. and Jones, R.A. 1988. Large-scale oligonucleotide synthesis by the H-phosphonate method. Tetrahedron Lett. 29:2619-2622. Garegg, P.J., Regberg, T., Stawinski, J., and Strömberg, R. 1985. Formation of internucleotidic bond via phosphonate intermediates. Chem. Scr. 25:280-282. Garegg, P.J., Lindh, I., Regberg, T., Stawinski, J., Strömberg, R., and Henrichson, C. 1986a. Nucleoside H-Phosphonates. III. Chemical synthesis of oligodeoxyribonucleotides by the hydrogenphosphonate approach. Tetrahedron Lett. 27:4051-4054. Garegg, P.J., Lindh, I., Regberg, T., Stawinski, J., Strömberg, R., and Henrichson, C. 1986b. Nucleoside H-phosphonates. IV. Automated solid phase synthesis of oligoribonucleotides by the hydrogenphosphonate approach. Tetrahedron Lett. 27:4055-4058.
Deoxyribo- and Ribonucleoside H-Phosphonates
Hall, R.H., Todd, A., and Webb, R.F. 1957. Nucleotides. Part XLI. Mixed anhydrides as intermediates in the synthesis of dinucleoside phosphates. J. Chem. Soc. (1957):3291-3296. Hata, T. and Sekine, M. 1974. Oxidation of nucleoside phosphites by means of 2,2′-dipyridilyl disulfide via nucleoside silylphosphite intermediates. Tetrahedron Lett. (1974):3943-3946. Holy, A. and Smrt, J. 1966. Oligonucleotidic compounds. XI. Synthesis of ribonucleoside 2′,3′cyclophosphates from nucleosides via nucleoside 2′,3′-phosphites. Collect. Czech. Chem. Commun. 31:1528-1534. Holy, A. and Sorm, F. 1966. Nucleic acids components and their analogues. LXXX. Preparation of nucleoside phosphites by the reaction of nucleosides with triphenyl phosphites. Collect. Czech. Chem. Commun. 31:1544-1561. Holy, A., Smrt, J., and Sorm, F. 1965. Nucleic acids components and their analogues. LIX. The preparation and properties of nucleoside phosphites. Collect. Czech. Chem. Commun. 30:1635-1641. Honjo, M., Marumoto, R., Kobayashi, K., and Yoshioka, Y. 1966. Phosphorylation of ribonucleosides with phosphorus trichloride. Tetrahedron Lett. (1966):3851-3856. Jankowska, J., Sobkowski, M., Stawinski, J., and Kraszewski, A. 1994. Studies on aryl H-phosphonates. I. Efficient method for the preparation of deoxyribo- and ribonucleoside 3′-H-phosphonate monoesters by transesterification of diphenyl H-phosphonate. Tetrahedron Lett. 35:3355-3358. Kers, A., Kers, I., Stawinski, J., Sobkowski, M., and Kraszewski, A. 1996. Studies on aryl H-phosphonates. 3. Mechanistic investigations related to the disproportionation of diphenyl H-phosphonate under anhydrous basic conditions. Tetrahedron 52:9931-9944. Marugg, J.E., Tromp, M., Kuyl-Yeheskiely, E., van der Marel, G.A., and van Boom, J.H. 1986. A convenient and general approach to the synthesis of properly protected d-nucleoside-3′ hydrogenphosphonates via phosphite intermediates. Tetrahedron Lett. 27:2661-2664. McConnell, R.L. and Coover, H.W. 1959. Phosphorus-containing derivatives of 2,2-dimethyl-1,3propanediol. J. Org. Chem. 24:630-635.
Garegg, P.J., Regberg, T., Stawinski, J., and Strömberg, R. 1986c. Nucleoside hydrogenphosphonates in oligonucleotide synthesis. Chem. Scr. 26:59-62.
Ozola, V., Reese, C.B., and Song, Q.L. 1996. Use of ammonium aryl H-phosphonates in the preparation of nucleoside H-phosphonate building blocks. Tetrahedron Lett. 37:8621-8624.
Garegg, P.J., Stawinski, J., and Strömberg, R. 1987. Activation of nucleoside hydrogenphosphonates by use of aryl sulfonyl chlorides. Nucleosides Nucleotides 6:425-427.
Rozners, E. and Strömberg, R. 1997. Synthesis and properties of oligoribonucleotide analogs having formacetal internucleoside linkages. J. Org. Chem. 62:1846-1850.
Gibbs, D.E. and Larsen, C. 1984. Bis[2,2,2-trifluoroethyl] phosphite, a new reagent for synthesizing mono- and diesters of phosphorus acid. Synthesis (1984):410-413.
Rozners, E., Kumpins, V., Rekis, A., and Bizdena, E. 1988. Solid phase synthesis of oligoribonucleotides by the H-phosphonate method using 2′-O-benzoyl protective group. Bioorg. Khim. 14:1580-1582.
2.6.14 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Rozners, E., Rekis, A., Kumpins, V., and Bizdena, E. 1990. Synthesis of oligoribonucleotides by the H-phosphonate method using base-labile 2′O-protecting groups. II. Some aspects of use of 2′-O-benzoyl and anisoyl protecting groups. Bioorg. Khim. 16:1531-1536.
Takaku, H., Yamakage, S., Sakatsume, O., and Ohtsuki, M. 1988. A convenient approach to the synthesis of deoxyribonucleoside 3′-hydrogenphosphonates via bis(1,1,1,3,3,3-hexafluoro-2propyl) phosphonate intermediate. Chem. Lett. (1988):1675-1678.
Rozners, E., Renhofa, R., Petrova, M., Popelis, J., Kumpins, V., and Bizdena, E. 1992. Synthesis of oligoribonucleotides by the H-phosphonate approach using base labile 2′-O-protecting groups. 5. Recent progress in development of the method. Nucleosides Nucleotides 11:15791593.
Wada, T., Sato, Y., Honda, F., Kawahara, S., and Sekine, M. 1997. Chemical synthesis of oligodeoxyribonucleotides using N-unprotected Hphosphonate monomers and carbonium and phosphonium condensing reagents: O-Selective phosphonylation and condensation. J. Am. Chem. Soc. 119:12710-12721.
Rozners, E., Strömberg, R., and Bizdena, E. 1995a. Synthesis of oligoarabinonucleotides using Hphosphonates. Nucleosides Nucleotides 14:851853.
Young, R.W. 1952. A re-examination of the reaction between phosphorus trichloride and salicylic acid. J. Am. Chem. Soc. 74:1672-1673.
Rozners, E., Strömberg, R., and Bizdena, E. 1995b. Synthesis of RNA fragments using the H-phosphonate method and 2′-(2-chlorobenzoyl) protection. Nucleosides Nucleotides 14:855-857. Schofield, J.A. and Todd, A. 1961. Nucleotides. Part XLVI. A new method for the preparation of nucleoside phosphites. J. Ch em. Soc. (1961):2316-2320. Sekine, M. and Hata, T. 1975. Phenylthio group as a protecting group of phosphates in oligonucleotide synthesis via phosphotriester approach. Tetrahedron Lett. (1975):1711-1714. Sekine, M., Mori, H., and Hata, T. 1979. New type of chemical oxidative phosphorylation: Activation of phosphonate function by use of triisopropylbenzenesulfonyl chloride. Tetrahedron Lett. (1979):1145-1148.
Zhang, X., Abad, J.-L., Huang, Q., Zeng, F., Gaffney, B., and Jones, R. 1997. High yield protection of purine ribonucleosides for H-phosphonate RNA synthesis. Tetrahedron. Lett. 38:7135-7138.
Key References Froehler et al., 1986. See above. Garegg et al., 1986c. See above. Rozners et al., 1995b. See above. Stawinski et al., 1988. See above. Provide primary sources with complete experimental details for Basic Protocol 1. Stawinski and Thelin, 1990a. See above. Provides primary sources with complete experimental details for Basic Protocol 2.
Sekine, M., Narui, S., and Hata, T. 1988. A convenient method for the synthesis of deoxyribonucleoside 3′-hydrogenphosphonates. Tetrahedron Lett. 29:1037-1040.
Jankowska et al., 1994. See above.
Stawinski, J. and Thelin, M. 1990a. Nucleoside H-Phosphonates. XI. A convenient method for the preparation of nucleoside H-phosphonates. Nucleosides Nucleotides 9:129-135.
Marugg et al., 1986. See above.
Stawinski, J. and Thelin, M. 1990b. Studies on the activation pathway of phosphonic acid using acyl chlorides as activators. J. Chem. Soc., Perkin Trans. 2. (1990):849-853.
Stawinski, J. 1992. Some Aspects of H-Phosphonate Chemistry. In Handbook of Organophosphorus Chemistry (R. Engel, ed.) pp. 377-434. Marcel Dekker, New York.
Stawinski, J., Strömberg, R., Thelin, M., and Westman, E. 1988. Studies on the t-butyldimethylsilyl group as 2′-O-protection in oligoribonucleotide synthesis via the H-phosphonate approach. Nucl. Acids Res. 16:9285-9298.
Stawinski, J. and Strömberg, R. 1993. H-Phosphonates in Oligonucleotide Synthesis. Trends Org. Chem. 4:31-67.
Szabó, T., Almer, H., Strömberg, R., and Stawinski, J. 1995. 2-Cyanoethyl H-phosphonate. A reagent for the mild preparation of nucleoside H-phosphonate monoesters. Nucleosides Nucleotides 14:715-716.
Provides primary sources with complete experimental details for Basic Protocol 3.
Provides primary sources with complete experimental details for Basic Protocol 4.
Provide general aspects of the underlying chemistry in the context of nucleotide and oligonucleotide synthesis.
Contributed by Jacek Stawinski Stockholm University Stockholm, Sweden Roger Strömberg Karolinska Institutet Stockholm, Sweden
Protection of Nucleosides for Oligonucleotide Synthesis
2.6.15 Current Protocols in Nucleic Acid Chemistry
Supplement 4
Deoxyribonucleoside Phosphoramidites
UNIT 2.7
This unit provides methods for converting N- and 5′-O-protected deoxyribonucleosides to deoxyribonucleoside phosphoramidites, which are the building blocks in solid-phase DNA oligonucleotide synthesis. Deoxyribonucleoside phosphoramidites carrying the 2-cyanoethyl group for P(III)-protection are extensively used in oligonucleotide synthesis and are commercially available. The preparation of deoxyribonucleoside phosphoramidites bearing a different P(III)-protecting group will be described in this unit; phosphoramidites functionalized with the 4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl groups will be synthesized. Considering the importance of phosphordiamidite precursors in the preparation of deoxyribonucleoside phosphoramidites, two syntheses of these phosphinylating agents will be reported. These detailed methods provide a convenient and reliable approach to the synthesis of various deoxyribonucleoside phosphoramidites, assuming compatibility of the nucleosidic N- and 5′-O-protecting groups with the reagents used for such syntheses. PREPARATION OF 5′-O-(4,4′-DIMETHOXYTRITYL)-3′-O(N,N-DIISOPROPYLAMINO)-{4-[N-METHYL-N-(2,2,2-TRIFLUOROACETYL) AMINO]BUTOXY}PHOSPHINYL-2′-DEOXYRIBONUCLEOSIDES
BASIC PROTOCOL
This protocol describes a general method for the synthesis of deoxyribonucleoside phosphoramidites S.3a-d from commercially available N- and 5′-O-protected deoxyribonucleosides S.1a-d and N,N,N′,N′-tetraisopropyl-O-{4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl}phosphordiamidite S.2 (see Support Protocol 1) in the presence of 1H-tetrazole. This approach to the preparation of deoxyribonucleoside phosphoramidites is illustrated in Figure 2.7.1. Materials 5′-O-(4,4′-Dimethoxytrityl)-2′-deoxythymidine (S.1a; Chem-Impex International) N4-Benzoyl-5′-O-(4,4′-dimethoxytrityl)-2′-deoxycytidine (S.1b; Chem-Impex International) 6 N -Benzoyl-5′-O-(4,4′-dimethoxytrityl)-2′-deoxyadenosine (S.1c; Chem-Impex International) N2-Isobutyryl-5′-O-(4,4′-dimethoxytrityl)-2′-deoxyguanosine (S.1d; Chem-Impex International) Anhydrous methylene chloride (Aldrich)
i
DMTrO
B
Pr2N P i O Pr2N
O OH
1a b c d
CH3 N CF3 2
O
1H-Tetrazole/CH2Cl2
B = Thymin-1-yl 4 B = N -benzoylcytosin-1yl 6 B = N -benzoyladenin-9-yl 2 B = N -isobutyrylguanin-9-yl
B
DMTrO
i
Pr2N
P
O O CF3
O
N O CH3
3a-d
DMTr: 4,4'dimethoxytrityl; iPr: isopropyl
Figure 2.7.1 Preparation of deoxyribonucleoside phosphoramidites from N- and 5′-O-protected deoxyribonucleosides.
Protection of Nucleosides for Oligonucleotide Synthesis
Contributed by Andrzej Wilk, Andrzej Grajkowski, Marcin K. Chmielewski, Lawrence R. Phillips, and Serge L. Beaucage
2.7.1
Current Protocols in Nucleic Acid Chemistry (2001) 2.7.1-2.7.12 Copyright © 2001 by John Wiley & Sons, Inc.
Supplement 4
N,N,N′,N′-Tetraisopropyl-O-{4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl}phosphordiamidite (S.2; see Support Protocol 1) Sublimed 1H-tetrazole (Aldrich) 9:1 (v/v) benzene/triethylamine (both available from Aldrich) 230- to 400-mesh silica gel 60Å (Merck) 50-mL three-necked round-bottom flask and rubber septa 15-mL powder addition funnel (Labglass) Vacuum desiccator Rotary evaporator High vacuum pump Dry argon gas cylinder 10-mL syringe 2.5 × 20–cm disposable Flex chromatography columns (Kontes) Fraction collector Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E) 1. Dry 2 mmol of a suitably protected deoxyribonucleoside (S.1a-d) in a 50-mL three-necked round-bottom flask for 2 hr at 25°C under high vacuum in a desiccator. Seal the flask with a 15-mL powder addition funnel containing 140 mg (2 mmol) dry 1-H tetrazole, and rubber septa. 2. Under an argon atmosphere, using a 10-mL syringe, add 10 mL anhydrous methylene chloride followed by 900 mg (2.1 mmol) N,N,N′,N′-tetraisopropyl-O-{4[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl}phosphordiamidite S.2 and stir with magnetic stir bar. 3. Add the 140 mg (2 mmol) 1H-tetrazole in the 15-mL powder addition funnel over a 0.5-hr period. 4. Monitor the progress of the reaction by TLC (APPENDIX 3D) using 9:1 benzene/triethylamine as an eluent. Phosphinylation of suitably protected 2′-deoxynucleosides S.1a-c is usually complete within 1 hr at ambient temperature. For best results, phosphinylation of properly protected 2′-deoxyguanosine S.1d should be allowed to proceed for 12 hr at 25°C.
5. Concentrate the reaction mixture to a foam using a rotary evaporator/vacuum pump system. 6. Suspend the crude product in a minimum amount (∼3 mL) of 9:1 benzene/triethylamine and apply the suspension to a 2.5 × 20–cm disposable Flex column containing 40 g silica gel (APPENDIX 3E) that has been equilibrated in 9:1 benzene/triethylamine. 7. Elute the column with 9:1 benzene/triethylamine. Collect in a fraction collector, pool appropriate fractions, and concentrate using a rotary evaporator/vacuum pump system. Each deoxyribonucleoside phosphoramidite S.3a-d is isolated as a white amorphous foam in yields ranging from 92% to 98%. Pure S.3a-d can be stored indefinitely at –20°C in sealed amber glass vials.
Deoxyribonucleoside Phosphoramidites
5′-O-(4,4′-Dimethoxytrityl)-3′-O-(N,N-diisopropylamino)-{4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butoxy}phosphinyl-2′-deoxythymidine S.3a: 31P-NMR (121 MHz, C6D6): δ 143.3, 143.4, 143.66, 143.75. FAB-HRMS: calcd. for C44H56F3N4O9P (M+Na)+ 895.3635, found 895.3599.
2.7.2 Supplement 4
Current Protocols in Nucleic Acid Chemistry
N4-Benzoyl-5′-O-(4,4′-dimethoxytrityl)-3′-O-(N,N-diisopropylamino)-{4-[(N-methyl-N(2,2,2-trifluoroacetyl)amino]butoxy}phosphinyl-2′-deoxycytidine S.3b: 31P-NMR (121 MHz, C6D6): δ 143.82, 143.87, 143.92. FAB-HRMS: calcd. for C50H59F3N5O9P (M+Na)+ 984.3900, found 984.3908. N6-Benzoyl-5′-O-(4,4′-dimethoxytrityl)-3′-O-(N,N-diisopropylamino)-{4-[N-methyl-N(2,2,2-trifluoroacetyl)amino]butoxy}phosphinyl-2′-deoxyadenosine S.3c: 31P-NMR (121 MHz, C6D6): δ 143.5, 143.6, 143.8, 144.0. FAB-HRMS: calcd. for C51H59F3N7O8P (M+Na)+ 1008.4010, found 1008.3970. N2-Isobutyryl-5′-O-(4,4′-dimethoxytrityl)-3′-O-(N,N-diisopropylamino)-{4-[N-methyl-N(2,2,2-trifluoroacetyl)amino]butoxy}phosphinyl-2′-deoxyguanosine S.3d: 31P-NMR (121 MHz, C6D6): δ 142.9, 143.1, 143.5, 143.7. FAB-HRMS: calcd. for C48H61F3N7O9P (M+Na)+ 990.4118, found 990.4099.
PREPARATION OF N,N,N′,N′-TETRAISOPROPYL-O-{4-[N-METHYL-N(2,2,2-TRIFLUOROACETYL)AMINO]BUTYL}PHOSPHORDIAMIDITE
SUPPORT PROTOCOL 1
4-[N-Methyl-N-(2,2,2-trifluoroacetyl)amino]butan-1-ol S.4 (see Support Protocol 2; Fig. 2.7.2) is treated with phosphorus trichloride to generate the corresponding O-{4-[Nmethyl-N-(2,2,2-trifluoroacetyl)amino]butyl}phosphordichloridite S.5, which is then purified by vacuum distillation. Condensation of the phosphordichloridite S.5 with N,N-diisopropylamine affords N,N,N′,N′-tetraisopropyl-O-{4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl}phosphordiamidite S.2, which can be purified further by silica gel chromatography. As shown in Figure 2.7.1, the phosphordiamidite S.2 is required for the phosphinylation of N- and 5′-O-protected deoxyribonucleosides S.1a-d (see Basic Protocol). Materials Phosphorus trichloride (Aldrich), freshly distilled Anhydrous acetonitrile (Aldrich) 4-[N-Methyl-N-(2,2,2-trifluoroacetyl)amino]butan-1-ol (S.4; see Support Protocol 2) Anhydrous petroleum ether (Aldrich), freshly distilled from phosphorus pentoxide Drierite, 8 mesh (Aldrich) Anhydrous N,N-diisopropylamine (Aldrich) 250- and 1000-mL round-bottom flasks 25-mL pressure-equalizing dropping funnel Vacuum distillation head 24/40 and appropriate thermometer Three-way stopcock Reflux condenser
CH3 N
F3C
PCl3 OH
O
MeCN
Pr2NH
P
Cl
5
i
Pr2N P Pr2N O
CH3 N CF3
i
Petroleum ether
O
O
4
i
Cl
CH3 N
F3C
O
2
Figure 2.7.2 Preparation of N,N,N′,N′-tetraisopropyl-O-{4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl}phosphordiamidite.
Protection of Nucleosides for Oligonucleotide Synthesis
2.7.3 Current Protocols in Nucleic Acid Chemistry
Supplement 4
Drying tube Rubber septum 250-mL sintered glass funnel (coarse porosity) Rotary evaporator/vacuum pump system NOTE: Phosphordichloridites are very sensitive to moisture. Reaction yields depend on the dryness of the reaction conditions. It is recommended that all glassware be oven-dried overnight at 120°C. The dried glassware should then be cooled to ambient temperature under an inert gas atmosphere in a desiccator. Acetonitrile is refluxed over calcium hydride for ≥2 hr prior to distillation. Prepare O-{4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl}phosphordichloridite 1. Dissolve 6.8 g (50 mmol) freshly distilled phosphorus trichloride in 50 mL anhydrous acetonitrile in an oven-dried 250-mL round-bottom flask. 2. Place 8 g (40 mmol) of 4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butan-1-ol S.4 and 10 mL anhydrous acetonitrile in a 25-mL pressure-equalizing dropping funnel connected to a vacuum line via a three-way stopcock. 3. Add the S.4 solution dropwise over a 2-hr period to the magnetically stirred phosphorus trichloride solution from step 1. CAUTION: During the course of the addition, a slight vacuum should be applied through the dropping funnel to maintain an internal pressure of ∼750 mmHg. This lower-than-atmospheric pressure ensures the removal of gaseous hydrogen chloride that is being generated.
4. Stir the reaction mixture for an additional 3 hr at 40°C and ∼40 mmHg. 5. Fractionally distill the remaining material under reduced pressure. O-{4-[N-Methyl-N-(2,2,2-trifluoroacetyl)amino]butyl}phosphordichloridite S.5 is a colorless liquid boiling at 110°C/0.1 mmHg. Yield 11 g (37 mmol, 92%). 1H-NMR (300 MHz, C6D6): δ [1.01(bm) (33%) and 1.11 (bm) (67%) (4H)], [2.38 (bs) (67%) and 2.45 (bs) (33%), (3H)], [2.75 (m) (33%) and 2.88 (m) (67%) (2H)], 3.77 (m, 2H). 13C-NMR (75 MHz, C6D6): δ 22.3, 23.9, 26.0, 26.3, 33.2, 33.5, 47.9, 48.0, 67.5 (d, 2JPC = 10.6 Hz), 67.7 (d, 2JPC = 10.6 Hz), 116.9 (q, 1JCF = 288 Hz), 117.0 (q, 1JCF = 288 Hz), 156.6 (q, 2JCF = 36.0 Hz), 156.7 (q, 2JCF = 36.0 Hz). 31P-NMR (121 MHz, C6D6): δ 174.2, 174.3. Although S.5 can be stored for a week at –20°C without significant decomposition, it is recommended to use S.5 when freshly distilled.
Prepare N,N,N′,N′-tetraisopropyl-O-{4 -[N-methyl-N-(2,2,2trifluoroacetyl)amino]butyl}phosphordiamidite 6. Add 300 mL anhydrous petroleum ether and 11 g (37 mmol) S.5 to an oven-dried 1000-mL round-bottom flask equipped with a reflux condenser fitted with a drying tube filled with Drierite. 7. Add in portions over a 15-min period a solution of 28 mL (200 mmol) anhydrous N,N-diisopropylamine in 100 mL anhydrous petroleum ether through the reflux condenser to the vigorously stirred phosphordichloridite solution. Large quantities of white precipitate (N,N-diisopropylammonium hydrochloride) are produced.
Deoxyribonucleoside Phosphoramidites
8. As the resulting suspension cools to ambient temperature, remove the reflux condenser and seal the flask with a rubber septum. Allow the reaction mixture to stir for an additional 48 hr at 25°C.
2.7.4 Supplement 4
Current Protocols in Nucleic Acid Chemistry
9. Filter off the precipitated N,N-diisopropylammonium hydrochloride salt using a 250-mL sintered glass funnel of coarse porosity. 10. Concentrate the filtrate on a rotary evaporator/vacuum pump system and keep under high vacuum for 12 hr. As prepared, compound S.2 is sufficiently pure for phosphinylating properly protected deoxyribonucleosides. However, S.2 can be purified by silica gel chromatography (APPENDIX 3E) using ∼20 g of silica gel per gram of crude product, and 9:1 (v/v) benzene/triethylamine as the eluent. Fractions containing pure S.2 are detected on analytical TLC plates (APPENDIX 3D) by staining with a saturated solution of silver nitrate in ethanol and then briefly heating the TLC plate at 50°C. N,N,N′,N′-Tetraisopropyl-O-{4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl}phosphordiamidite S.2. Yield 15 g (35 mmol, 95%). 1H-NMR (300 MHz, C6D6): δ 1.21 (m, 24H), 1.40 (m, 4H), [2.41 (q, 5JHF = 1.6 Hz) (67%) and 2.50 (q, 5JHF = 0.7 Hz, (33%), (3H)], [2.89 (tq, J = 7.4 Hz, 5JHF = 1.0 Hz) (33%) and 3.04 (t, J = 6.9 Hz) (67%), (2H)], 3.43 (m, 2H), 3.50 (m, 4H). 13C-NMR (75 MHz, C6D6): δ 23.2, 23.7, 23.8, 24.3, 24.5, 25.0, 28.3, 28.4, 28.5, 28.6 33.3, 33.5 (q, 4JCF = 4.2 Hz), 44.3, 44.5, 48.5, 48.7 (q, 4JCF = 4.2 Hz), 63.4 (d, 2JCP = 6.4 Hz), 63.7 (d, 2JCP = 6.4 Hz), 117.0 (q, 1JCF = 288 Hz), 117.1 (q, 1JCF = 288 Hz), 156.1 (q, 2JCF = 36 Hz). 31P-NMR (121 MHz, C6D6): δ 117.9, 118.2. FAB-MS: calcd. for C19H39F3N3O2P (M+H)+ 430, found 430. Pure S.2 can be stored indefinitely at –20°C in sealed amber glass vials.
PREPARATION OF 4-[N-METHYL-N-(2,2,2-TRIFLUOROACETYL)AMINO]BUTAN-1-OL
SUPPORT PROTOCOL 2
As shown in Figure 2.7.3, 4-(N-methyl)aminobutan-1-ol S.8 is prepared from the reaction γ-butyrolactone S.6 with gaseous methylamine followed by reduction of the resulting amido alcohol S.7 with lithium aluminum hydride. Chemoselective N-trifluoroacetylation is performed by condensing 4-(N-methyl)aminobutan-1-ol S.8 with methyl trifluoroacetate. 4-[N-Methyl-N-(2,2,2-trifluoroacetyl)amino]butan-1-ol S.4 is required in the preparation of N,N,N′,N′-tetraisopropyl-O-{4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl}phosphordiamidite S.2 (see Support Protocol 1). Materials γ-Butyrolactone (S.6; Aldrich) Anhydrous gaseous methylamine (Aldrich) Diethyl ether (Malinckrodt), freshly distilled from sodium Drierite (Aldrich) Lithium aluminum hydride powder (LAH, Aldrich)
O
O
CH3NH2
O HO
2.5 bar 6
7
HO
CF3CO2CH3 NHCH3
LAH NHCH3
Et2O
O HO
N
CF3
CH3
8
4
LAH: lithium aluminum hydride
Figure 2.7.3
Preparation of 4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butan-1-ol.
Protection of Nucleosides for Oligonucleotide Synthesis
2.7.5 Current Protocols in Nucleic Acid Chemistry
Supplement 4
Triethanolamine (Aldrich) Methyl trifluoroacetate (Aldrich) 250-mL pressure vessel (Barrskogen) 2-L three-necked round-bottom flask Mechanical stirrer (Arrow) Pressure-equalizing dropping funnel Reflux condenser connected to a drying tube Dry argon gas cylinder 250-mL sintered glass funnel (coarse porosity) Rotary evaporator/water aspirator system 50-mL round-bottom flask Prepare N-methyl-4-hydroxybutyramide 1. Place 17.2 g (199 mmol) γ-butyrolactone S.6 in a 250-mL pressure vessel. 2. Evacuate the vessel to ∼2 mmHg and fill it with anhydrous gaseous methylamine. 3. Stir the reaction mixture overnight under a positive pressure of methylamine (∼2.5 bar). CAUTION: This reaction involves highly irritating gaseous methylamine under pressure. Ensure that the pressure vessel is tight and placed in a well-ventilated chemical fume hood. Use moist pH-indicator paper to check for leaks. Even trace amounts of strongly alkaline methylamine can be detected using this test.
4. Carefully release excess methylamine to the atmosphere of a well-ventilated chemical fume hood. 5. Evacuate the vessel and maintain the reaction product under vacuum (2 mmHg) for 1 hr. N-Methyl-4-hydroxybutyramide S.7 is isolated as a pure (>98%) crystalline mass (mp 32° to 34°C) in near quantitative yield (23.1 g, 197 mmol). 1H-NMR (300 MHz, DMSO-d6): δ 1.63 (dt, J = 7.6, 6.4 Hz, 2H), 2.10 (t, J = 7.6 Hz, 2H), 2.56 (d, J = 4.6 Hz, 3H), 3.37 (t, J = 6.4 Hz, 2H), 3.58 (b, ∼1H), 7.75 (b, 1H). 13C-NMR (75 MHz, DMSO-d6): δ 25.6, 28.7, 32.2, 60.5, 173.1. S.7 can be stored indefinitely at –20°C in sealed amber glass vials.
Reduce N-methyl-4-hydroxybutyramide 6. Add 1 L diethyl ether to an oven-dried 2-L three-necked round-bottom flask equipped with a mechanical stirrer, a pressure-equalizing dropping funnel, and a reflux condenser connected to a drying tube filled with Drierite. 7. Add, under an argon atmosphere, 36 g (0.94 mol) lithium aluminum hydride powder. 8. Melt 58.5 g (0.50 mol) N-methyl-4-hydroxybutyramide S.7 and place in the dropping funnel. 9. Add the neat supercooled liquid (S.7) to the mechanically stirred LAH suspension, dropwise, over a 2-hr period at 25°C. 10. Reflux the reaction mixture for 6 hr and stir for an additional 6 hr at 25°C.
Deoxyribonucleoside Phosphoramidites
CAUTION: Diethyl ether is highly flammable. Lithium aluminum hydride is extremely sensitive to water; traces of water from the solvent and the environment will result in the release of highly flammable hydrogen gas. Consequently, steps 7 to 10 must be performed with the utmost care to exclude moisture from the reaction mixture. A flow of anhydrous argon gas through the reaction vessel greatly reduces the danger of fire.
2.7.6 Supplement 4
Current Protocols in Nucleic Acid Chemistry
11. Quench the reduction reaction while stirring by first adding 125 mL (0.94 mol) triethanolamine dropwise over a 1-hr period (Powell et al., 1986; Gardrat et al., 1990), followed by 50 mL (2.8 mol) water dropwise over a 1-hr period. 12. Stir the resulting slurry for 12 hr at 25°C. 13. Filter the slurry through a 250-mL sintered glass funnel (coarse porosity) and wash the solid cake three times with 200 mL diethyl ether. 14. Pool the etheral filtrates and evaporate on a rotary evaporator/water aspirator system. Distill the residue under reduced pressure. The amino alcohol S.8 (34.3 g ,0.33 mol, 67%) is obtained as a colorless liquid boiling at 47°C/0.25 mmHg. 1H-NMR (300 MHz, CDCl3): δ 1.64 (m, 4H), 2.42 (s, 3H), 2.62 (dd, J = 6.0, 5.3 Hz, 2H), 3.57 (dd, J = 5.3, 4.6 Hz, 2H). 13C-NMR (75 MHz, CDCl3): δ 28.3, 32.3, 35.7, 51.7, 62.5. S.8 can be stored indefinitely at –20°C in sealed amber glass vials.
Perform chemoselective trifluoroacetylation of 4-(N-methyl)aminobutan-1-ol 15. To 5 g (48 mmol) of 4-(N-methyl)aminobutan-1-ol S.8 in a 50-mL round-bottom flask, add 7.8 g (60 mmol) methyl trifluoroacetate. 16. Stir the reaction mixture for 12 hr at 25°C. 17. Remove methanol and excess methyl trifluoroacetate by distillation at atmospheric pressure. 18. Distill the residue under reduced pressure. 4-[N-Methyl-N-(2,2,2-trifluoroacetyl)amino]butan-1-ol S.4 is obtained as a slightly viscous colorless liquid (8.9 g, 45 mmol, 94%) boiling at 71°C/0.36 mmHg. 1H-NMR (300 MHz, CDCl3): δ 1.56 (m, 2H), 1.70 (m, 2H), [3.00 (q, 5JHF = 0.7 Hz) (33%) and 3.14 (q, 5 JHF = 1.6 Hz) (67%), (3H)], 3.46 (m, J = 7.4 Hz, 2H), 3.65 (t, J = 6.1 Hz, 2H). 13C-NMR (75 MHz, CDCl3): δ 22.8, 24.7, 29.2, 29.3, 34.1, 34.6, 49.1, 49.4, 61.7, 61.8, 116.5 (q, 1JCF = 288 Hz), 116.6 (q, 1JCF = 288 Hz), 156.7 (q, 2JCF = 36.0 Hz), 156.8 (q, 2JCF = 36.0 Hz). Pure S.4 can be stored indefinitely at –20°C in sealed amber glass vials.
PREPARATION OF N,N,N′,N′-TETRAISOPROPYL-O{2-[(N-FORMYL-N-METHYL)AMINO]ETHYL}PHOSPHORDIAMIDITE
ALTERNATE PROTOCOL
Depending on the protecting group used for P(III), the preparation of phosphordichloridites may be problematic and could complicate the synthesis of phosphordiamidites, which are required for the preparation of deoxyribonucleoside phosphoramidites. A number of phosphordichloridites have been reported to decompose violently when heated (Beaucage, 1993). This protocol describes the generation of bis(N,N-diisopropylamino)chlorophosphine S.9 in situ from phosphorus trichloride and N,N-diisopropylamine, and its reaction with 2-(N-formyl-N-methyl)aminoethan-1-ol S.10 to give the desired phosphordiamidite S.11 (Fig. 2.7.4). The method thus alleviates the need of distilling potentially hazardous phosphordichloridites. Additional Materials (also see Basic Protocol) 2-(Methylamino)ethanol (Aldrich) Ethyl formate (Aldrich) Anhydrous benzene Freshly distilled phosphorous trichloride Anhydrous N,N-diisopropylamine 95:5 (v/v) anhydrous benzene/triethylamine Toluene
Protection of Nucleosides for Oligonucleotide Synthesis
2.7.7 Current Protocols in Nucleic Acid Chemistry
Supplement 4
O HO
PCl3
i-Pr2NH dry C6H6
i-Pr2N
P i-Pr2N
Cl
N H CH3
10
O
i-Pr2N
P i-Pr2N
O
N H CH3
11
9
Figure 2.7.4 Preparation of N,N,N′,N′-tetraisopropyl-O-{2-[(N-methyl-N-formyl)amino]ethyl}phosphordiamidite.
100- and 250-mL round-bottom flasks equipped with a reflux condenser Heating mantle 60-mL sintered glass funnel (coarse porosity) 3 × 20–cm chromatography column (APPENDIX 3E) Prepare 2-(N-formyl-N-methyl)aminoethan-1-ol 1. Place 51.0 g (0.68 mol) of 2-(methylamino)ethanol into a 250-mL round-bottom flask equipped with a reflux condenser, and cool to 5°C by immersion in an ice bath. 2. Add, in portions through the condenser, 75.0 g (1.01 mol) ethyl formate to the stirred amino alcohol over a 5-min period at 5°C. CAUTION: This reaction is exothermic.
3. Remove the ice bath and bring the solution to reflux for 1 hr using a heating mantle. 4. Distill the solution at atmospheric pressure to remove excess ethyl formate, and then carefully distill the remaining residue under high vacuum. 2-(N-Formyl-N-methyl)aminoethan-1-ol S.10 was obtained as a clear colorless liquid (63.1 g, 0.61 mol, 90%) boiling at 120° to 122°C at 0.15 mmHg. 1H-NMR (300 MHz, DMSO-d6): δ [2.75 (s) and 2.94 (s, 30%) (3H)], 3.27 (m, 2H), 3.47 (m, 3H), [7.94 (s) and 7.99 (s, 30%) (1H)]. 13C-NMR (75 MHz, DMSO-d6): δ 29.2, 34.9, 46.2, 51.2, 57.8, 57.9, 58.1, 58.2, 162.7, 163.0. EI-MS: calcd. for C4H9NO2 (M•+) 103, found 103. Pure S.10 can be stored indefinitely at –20°C in sealed amber glass vials.
Prepare N,N,N′,N′-tetraisopropyl-O-{2 -[(N-formyl-N-methyl)amino]ethyl}phosphordiamidite 5. To an oven-dried 100-mL round-bottom flask containing 50 mL anhydrous benzene under a dry argon atmosphere, add by syringe through a rubber septum 876 µL (10 mmol) freshly distilled phosphorus trichloride. 6. Cool the stirred solution to 5°C by immersing in an ice bath and then, over a 30-min period, add 7.7 mL (55 mmol) of anhydrous N,N-diisopropylamine under argon using a 10-mL syringe. 7. Remove the ice bath and allow the stirred reaction to warm to 25°C under a positive pressure of argon until the formation of bis(N,N-diisopropylamino)chlorophosphine S.9 is complete.
Deoxyribonucleoside Phosphoramidites
The rate of the reaction is monitored by 31P-NMR spectroscopy; after ∼48 hr, the presence of S.9 (132.0 ppm downfield relative to a phosphoric acid external standard) is observed as the major (>96%) reaction product.
2.7.8 Supplement 4
Current Protocols in Nucleic Acid Chemistry
8. Add 1.03 mL (10 mmol) of 2-(N-formyl-N-methyl)aminoethan-1-ol S.10 to the suspension. Stir resulting mixture for 2 hr at 25°C under a positive pressure of argon. 31
P-NMR indicates that the reaction is essentially quantitative (∼96%). Spectrum of the product in benzene-d6 shows two singlets at 118.0 and 118.7 ppm.
9. Filter the suspension through a 60-mL sintered glass funnel (coarse porosity) and wash the collected salt with 20 mL anhydrous benzene. 10. Evaporate the filtrates under reduced pressure to an oil and dissolve with a minimum amount (∼3 mL) of 95:5 anhydrous benzene/triethylamine. 11. Apply the viscous solution uniformly to the top of a 3 × 20–cm chromatography column (APPENDIX 3E) packed with 30 g of a 60 Å silica gel slurry in a solution of 95:5 benzene/triethylamine. 12. Elute the column isocratically with 95:5 benzene/triethylamine collecting 8-mL fractions. 13. Analyze the fractions by TLC (APPENDIX 3D) and pool the fractions that contain the product. Evaporate fractions containing the phosphordiamidite S.11 to an oil on a rotary evaporator/vacuum pump system. Using 95:5 benzene/triethylamine as the eluent, S.11 has an Rf ~ 0.49. S.11 is visible only by staining the TLC with silver nitrate as recommended in Support Protocol 1, step 10.
14. Co-evaporate the oil four times with 10 mL toluene to remove traces of triethylamine. Leave under high vacuum for ≥3 hr. N,N,N′,N′-tetraisopropyl-O-{2 -[(N-formyl-N-methyl)amino]ethyl}phosphordiamidite S. 11. Yield: 2.43 g (7.3 mmol, 73%). 1H-NMR (300 MHz, C6D6): δ [1.14 (d, J = 6.9 Hz), 1.16 (d, J = 6.7 Hz) 1.18 (d, J = 6.7 Hz) (24H)], [2.40 (s, 34%) and 2.64 (s, 66%) (3H)], 2.80 (t, J = 5.4 Hz, 2H), 3.43 (m, 4H), [3.29 (dt, J =5.4 Hz, JHP = 6.6 Hz) and 3.60 (dt, J =5.4 Hz, JHP = 6.6 Hz)(2H)], [7.82 (s, 34%) and 7.98 (s, 66%) (1H)]. 13C-NMR (75 MHz, C6D6): δ 24.1, 24.2, 24.6, 24.7, 44.7, 44.9, 45.8 (d, 2JCP = 8.5 Hz), 50.4(d, 2JCP = 8.5 Hz), 61.3, 61.5, 61.9, 62.2, 161.9, 162.3. 31P-NMR (121 MHz, C6D6): δ 118.0, 118.7. EI-MS: calcd. for C16H36N3O2P (M•+) 333.2545, found 333.2528. Pure S.11 can be stored indefinitely at –20°C in sealed amber glass vials.
COMMENTARY Background Information Since its inception in the early 1980s, the phosphoramidite method for oligonucleotide synthesis (Beaucage and Caruthers, 1981; Beaucage and Iyer, 1992; UNIT 3.3) has been extensively used to produce polynucleotides (Pon et al., 1994) on solid supports. The method is most convenient when using the 2-cyanoethyl group for phosphate protection (Sinha et al., 1984) on small-scale syntheses of oligonucleotides. This group is eventually removed along with nucleobase-protecting groups during oligonucleotide deprotection by treatment with either concentrated ammonium hydroxide or pressurized ammonia gas (Boal et al., 1996). Under these conditions, 2-cyanoethyl groups undergo β-elimination, generating acrylonitrile as a side-product (Tener,
1961). Acrylonitrile is a potent carcinogen (Solomon et al., 1984), and alkylation of the nucleobase of nucleosides and oligonucleotides by this reagent is well documented (Chambers, 1965; Solomon et al., 1984; Prokopczyk et al., 1988; Crippa et al., 1993). In an effort to obviate nucleobase modification during oligonucleotide deprotection at any scale level, an alternative to the 2-cyanoethyl group for phosphate protection is recommended. The use of the 4-[N-methyl-N-(2,2,2trifluoroacetyl)]aminobutyl group for phosphodiester protection in the synthesis of oligodeoxyribonucleotides was recently reported (Wilk et al., 1999). Polydeoxyribonucleotides carrying this phosphodiester protecting group are easily deprotected by treatment with concentrated ammonium hydroxide. Cleavage of
Protection of Nucleosides for Oligonucleotide Synthesis
2.7.9 Current Protocols in Nucleic Acid Chemistry
Supplement 4
that, unlike the 2-cyanoethyl group, it can be removed in the absence of base under neutral conditions (authors’ unpub. observ.). The use of (N,N-diisopropylamino)-O-alkylchlorophosphine (Sinha et al., 1984) in the preparation of deoxyribonucleoside phosphoramidites will not be discussed in this unit as these phosphinylating reagents are, because of their inherent reactivity, less stable to moisture than the corresponding phosphordiamidites, and are consequently much less convenient to handle.
the 2,2,2-trifluoroacetyl groups is rate limiting, and is followed by rapid cyclo-deesterification of the resulting 4-(N-methyl)aminobutyl phosphotriesters to give the corresponding phosphodiesters with the concomitant formation of N-methylpyrrolidine (Fig. 2.7.5). When compared to the potential DNA-alkylating properties of acrylonitrile, 2,2,2-trifluoroacetamide and N-methylpyrrolidine, which are produced during oligonucleotide deprotection, are quite innocuous. The high coupling efficiency of the deoxyribonucleoside phosphoramidites S.3a-d (Fig. 2.7.1), their stability in acetonitrile solutions, and the relative ease of 4-(N-methyl)aminobutyl phosphotriester deprotection prompted the authors to describe the preparation of these phosphoramidites in this unit as alternatives to the commercially available 2-cyanoethyl deoxyribonucleoside phosphoramidites. Given that the procedure for preparing phosphoramidites S.3a-d is similar to that of the 2-cyanoethyl phosphoramidites, it shall serve as a relevant model for the synthesis of deoxyribonucleoside phosphoramidites, which carry different P(III) protecting groups. In this context, depending on the group selected for P(III) protection, the method used for the preparation of either the phosphordiamidite S.2 or the analogous N,N,N′,N′-tetraisopropyl-O-(2-cyanoethyl)phosphordiamidite may not be adequate. For example, the phosphordiamidite S.11 (Fig. 2.7.4) could not be conveniently prepared from a phosphordichloridite precursor. Thus, the phosphordiamidite S.11 was prepared from bis(N,N-diisopropylamino)chlorophosphine, which provides added versatility to the synthesis of phosphordiamidites as phosphinylating agents. The advantage of the 2-[(N-formyl-N-methyl)amino]ethyl group for oligonucleotide phosphodiester protection is
O
X
O
O CH3
O O P
N
O
O
O
X
CF3 O
NH4OH − CF3CONH2
O
Critical Parameters and Troubleshooting Since 4-(N-methyl)aminobutan-1-ol S.8 (see Support Protocol 2) is not commercially available, its synthesis has been studied for careful optimization. The lithium aluminum hydride reduction of the parent amido alcohol S.7 is demanding in that it requires strict anhydrous conditions and a mechanical stirrer for optimal performance. In addition, quenching the reduction reaction by adding triethanolamine followed by water to destroy aluminum complexes (as recommended by Powell et al., 1986, and Gardrat et al., 1990), and carefully washing the solid aluminate cake with diethyl ether (see Support Protocol 2, steps 11 to 13) must be performed to ensure a maximum recovery of S.8. The selection of acetonitrile as the solvent for converting 4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butan-1-ol S.4 to its phosphordichloridite S.5 under lower than atmospheric pressure conditions (see Support Protocol 1) is required for optimal production of S.5. The use of other solvents will result in a considerably lower yield of S.5. Performing the reaction at atmospheric instead of reduced pres-
B
O
CH3
O O P
O
HN
O O O
O NH4OH
O B
B
O
O P
O
−
O
+
NH4 O
+
CH3N
B
O
X = Thy or N-protected nucleobase; B = Thy, Ade, Cyt, or Gua
Deoxyribonucleoside Phosphoramidites
Figure 2.7.5 Deprotection of oligodeoxyribonucleotides carrying the 4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl phosphate protecting group under standard basic conditions.
2.7.10 Supplement 4
Current Protocols in Nucleic Acid Chemistry
sure will also result in unwanted formation of side-products. It is important to mention that the preparation of phosphordiamidites from the bis(N,Ndiisopropylamino)chlorophosphine intermediate S.9 (see Alternate Protocol) should be performed under strictly anhydrous conditions to minimize the formation of side-products resulting from hydrolysis. These side-products unfortunately co-elute with deoxyribonucleoside phosphoramidites during silica gel chromatography. Depending on the amount of contaminating side-products, the stoichiometry of deoxyribonucleoside phosphoramidites required for solid-phase oligonucleotide synthesis may be difficult to determine accurately. Furthermore, the chemical reactivity of the phosphorus-containing hydrolysis side-products is not known; these side-products may react with activated phosphoramidites and further decrease the effective concentration of the amidites. Occasionally, deoxyribonucleoside phosphoramidites may not produce the expected coupling efficiency during conventional solidphase oligonucleotide synthesis. A number of parameters may be responsible for such an apparent lack of reactivity. (1) Phosphoramidites may be contaminated with the triethylamine used with benzene as an eluent during purification on silica gel chromatography. Triethylamine stoichiometrically neutralizes 1H-tetrazole, the weak acid required for the activation of deoxyribonucleoside phosphoramidites in solid-phase oligonucleotide synthesis. Thus, it will prevent optimal coupling efficiency of the phosphoramidites to an extent corresponding to its concentration. It is therefore critical that the phosphoramidites be subjected to high-vacuum foaming after chromatographic purification to ensure complete removal of triethylamine contaminants. Alternatively, lyophilization of a frozen benzene solution of deoxyribonucleoside phosphoramidites is an efficient method for the removal of residual triethylamine. (2) Phosphoramidites may be contaminated with adventitious moisture upon storage, which will result in lower coupling efficiency. It is recommended that deoxyribonucleoside phosphoramidites be dried overnight under high vacuum in a desiccator containing phosphorus pentoxide as a drying agent. (3) The overall purity of deoxyribonucleoside phosphoramidites, as determined by 31P-NMR spectroscopy, should be >95%; phosphoramidites contaminated with >15% of phosphorus-containing hydrolysis products invariably exhibit lower coupling efficiency
when compared to that of pure deoxyribonucleoside phosphoramidites.
Anticipated Results It should be emphasized that deviation from Support Protocol 1 in regard to preparing phosphordichloridite S.5 with a solvent other than acetonitrile at atmospheric pressure will generally cause the yield of the product to drop to 50% to 60%. Caution should be exercised when attempting the distillation of unknown phosphordichloridites; a number of these compounds may decompose or even explode on heating (Beaucage, 1993). In this context, the preparation of phosphordiamidites from bis(N,N-diisopropylamino)chlorophosphine S.9 generated in situ (see Alternate Protocol) is a much safer process and is easier to perform experimentally. Furthermore, the purification of phosphordiamidites on silica gel chromatography is milder than distillation to the phosphonylating reagents and safer to perform. Isolated yields of phosphordiamidites after silica gel chromatography range between 70% and 80% based on the starting amido alcohol S.10; it should be noted that formation of some hydrolysis side-products cannot be avoided, only minimized by careful attention to experimental detail. The synthesis of deoxyribonucleoside phosphoramidites S.3a-d from phosphordiamidite S.2 (see Basic Protocol) is performed essentially as recommended by Barone et al. (1984). More reaction time is required for the preparation of S.3d to optimize yields. It is possible that phosphinylation at O6 of guanine by S.2 might occur and serve as a secondary, slower, phosphinylating path (Barone et al., 1984) for the nucleoside S.1d. The phosphoramidites S.3a-d are very soluble and stable in acetonitrile. For example, an acetonitrile solution of S.3d, unlike the corresponding 2-cyanoethyl phosphoramidite, does not precipitate upon storage exceeding 3 days. In fact, the coupling efficiency of phosphoramidites S.3a-d is essentially unchanged after being in acetonitrile solution for 1 week (Wilk et al., 1999).
Time Considerations The preparation and purification of 4-(Nmethyl)aminobutan-1-ol S.8 can be accomplished in ∼3 days. The conversion of S.8 to the amido alcohol S.4 can be done in one day, and the purified phosphordiamidite S.2 can be obtained from S.4 within 3 days. The synthesis and purification of deoxyribonucleoside phosphoramidites S.3a-d require ∼2 days to com-
Protection of Nucleosides for Oligonucleotide Synthesis
2.7.11 Current Protocols in Nucleic Acid Chemistry
Supplement 4
plete. Alternatively, the synthesis and purification of the amido alcohol S.10 takes a half day, and its conversion to the purified phosphordiamidite S.11 takes ∼3 days.
Literature Cited Barone, A.D., Tang, J.-T., and Caruthers, M.H. 1984. In situ activation of bis-dialkylaminophosphines—A new method for synthesizing deoxyoligonucleotides on polymer supports. Nucl. Acids Res. 12:4051-4061. Beaucage, S.L. 1993. Oligodeoxyribonucleotides synthesis—Phosphoramidite approach. In Methods in Molecular Biology, Vol. 20: Protocols for Oligonucleotides and Analogs (S. Agrawal, ed.) pp. 33-61. Humana Press, Totowa, N.J. Beaucage, S.L. and Caruthers, M.H. 1981. Deoxynucleoside phosphoramidite—A new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Lett. 22:1859-1862. Beaucage, S.L. and Iyer, R.P. 1992. Advances in the synthesis of oligonucleotides by the phosphoramidite approach. Tetrahedron 48:22232311. Boal, J.H., Wilk, A., Harindranath, N., Max, E.E., Kempe, T., and Beaucage, S.L. 1996. Cleavage of oligodeoxyribonucleotides from controlledpore glass supports and their rapid deprotection by gaseous amines. Nucl. Acids Res. 24:31153117. Chambers, R.W. 196 5. The chemistry of pseudouridine. IV. Cyanoethylation. Biochemistry 4:219-226. Crippa, S., Di Gennaro, P., Lucini, R., Orlandi, M., and Rindone, B. 1993. Characterization of adducts of nucleic bases and acrylic-monomers. Gazz. Chim. Ital. 123:197-203. Gardrat, C., Latxague, L., and Picard, J.P. 1990. A new sy nthesis of dl-5-vinyloxazolidine-2thione, a natural antithyroid factor. J. Het. Chem. 27:811-812. Grajkowski, A., Wilk, A., Chmielewski, M.K., and Beaucage, S.L. 2000, unpublished results.
Powell, J., James, N., and Smith, S.J. 1986. Lithium aluminum hydride reductions: A new hydrolysis method for intractable products. Synthesis 338340. Prokopczyk, B., Bertinato, P., and Hoffman, D. 1988. Synthesis and kinetics of decomposition of 7-(2-cyanoehtyl)guanine and O-6-(2-cyanoethyl)guanine, markers for reaction of acrylonitrile and 3-(methylnitrosamino)propionitrile with DNA. Carcinogenesis 9:2125-2128. Sinha, N.D., Biernat, J., McManus, J., and Köster, H. 1984. Polymer support oligonucleotide synthesis. 18. Use of β-cyanoethyl-N,N,-dialkylamino/-N-morpholino phosphoramidite of deoxynucleosides for the synthesis of DNA fragments simplifying deprotection and isolation of the final product. Nucl. Acids Res. 12:45394557. Solomon, J.J., Cote, I.L., Wortman, M., Decker, K., and Segal, A. 1984. In vitro alkylation of calf thymus DNA by acrylonitrile—Isolation of cyanoethyl adducts of guanine and thymine and carboxyethyl adducts of adenine and cytosine. Chem.-Biol. Interactions 51:167-190. Tener, G.M. 1961. 2-Cyanoethyl phosphate and its use in the synthesis of phosphate esters J. Am. Chem. Soc. 83:159-168. Wilk, A., Grajkowski, A., Phillips, L.R., and Beaucage, S.L. 1999. The 4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl group as an alternative to the 2-cyanoethyl group for phosphate protection in the synthesis of oligodeoxyribonucleotides. J. Org. Chem. 64:7515-7522.
Contributed by Andrzej Wilk, Andrzej Grajkowski, Marcin K. Chmielewski, and Serge L. Beaucage Food and Drug Administration Bethesda, Maryland Lawrence R. Phillips National Cancer Institute Frederick, Maryland
Pon, R.T., Buck, G.A., Niece, R.L., Robertson, M., Smith, A.J., and Spicer, E. 1994. A survey of nucleic-acid services in core laboratories. BioTechniques 17:526-534.
Deoxyribonucleoside Phosphoramidites
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Current Protocols in Nucleic Acid Chemistry
Regioselective 2′-Silylation of Purine Ribonucleosides for Phosphoramidite RNA Synthesis
UNIT 2.8
This unit describes high-yield procedures for protection of purine ribonucleosides based on a reaction that allows highly regioselective 2′-silylation. The H-phosphonate monoester group produced in the silylation reaction is then cleaved, without silyl migration (Song et al., 1999; Zhang et al., 1997), to give intermediates ready for phosphitylation to yield the phosphoramidites. This method gives overall yields that are three times the best yields available by conventional procedures for adenosine (see Basic Protocol 1) and guanosine (see Basic Protocol 2), but offers no advantage for cytidine or uridine. SYNTHESIS OF 5′-O-(4,4′-DIMETHOXYTRITYL)-2′-O-tertBUTYLDIMETHYLSILYL-6-N-ACYLADENOSINE
BASIC PROTOCOL 1
This protocol makes use of transient protection of the 2′,3′-diol moiety of a ribonucleoside (S.2 in Fig. 2.8.1) by reaction with N,N-dimethylformamide dimethylacetal (Zemlicka, 1963) to prevent the small, but potentially troublesome, tritylation of the 2′-hydroxyl that otherwise accompanies tritylation of the 5′-hydroxyl (Zhang et al., 1997). The 2′,3′-Odimethylaminomethylene group is cleaved by any protic solvent. The N-dimethylaminomethylene group is cleaved by treatment with either aqueous ammonia or methylamine. The phenoxyacetylation reaction is carried out using the hydroxybenzotriazole active ester of phenoxyacetic acid after transient hydroxyl protection with trimethylchlorosilane. The regioselective silylation of the N- and 5′-O-protected adenosine and guanosine derivatives (S.4 and S.13, respectively) presumably occurs by a reaction sequence in which the phenyl-H-phosphonate reacts first with tert-butyldimethylchlorosilane to generate the corresponding diester, which then undergoes a transesterification with S.4/S.13 to generate a mixture of isomers (S.5/S.14). Subsequent transfer of the tert-butyldimethylsilyl (TBDMS) group predominantly to the more acidic 2′-hydroxyl gives S.6a/S15a along with 10% to 15% of the 3′-O-TBDMS isomers. The H-phosphonate moiety is removed by reaction of the isomers of S.6a,b/S15a,b with ethylene glycol or glycerol. The extraordinarily facile transesterification of H-phosphonate diesters in the presence of a vicinal hydroxyl group effects the conversion to S.8a,b/S.17a,b quantitatively within minutes, presumably via the intermediate S.7a,b/S.16a,b. After careful purification, S8a/S.17a are converted to the phosphoramidites S.9/S.18 by reaction with 2-cyanoethyl tetraisopropylphosphorodiamidite using diisopropylammonium tetrazolide as a catalyst. A short silica gel column removes the excess reagent. For H-phosphonate synthesis, monomers like S.6a/S.15a but without amino protection can be prepared by a similar route (Zhang et al., 1997). The labile phenoxyacetyl group used here does not survive the polar conditions required for purification of the charged H-phosphonates. Materials Adenosine Pyridine (reagent grade or better) Dimethylformamide dimethyl acetal Nitrogen source Contributed by Barbara L. Gaffney and Roger A. Jones Current Protocols in Nucleic Acid Chemistry (2001) 2.8.1-2.8.13 Copyright © 2001 by John Wiley & Sons, Inc.
Protection of Nucleosides for Oligonucleotide Synthesis
2.8.1 Supplement 6
Figure 2.8.1 Dimethoxytritylation N-phenoxyacetylation, silylation, dephosphonylation, and phosphitylation of adenosine. Abbreviations: Ad, adamantanecarbonyl; DBU, 1,8-diazabyclyclo[5.4.0]undex-7-ene; DMTr, 4,4′-dimethoxytrityl; HOBT, 1-hydroxybenzotriazole; Ph, phenyl; i-Pr, isopropyl; TMS, trimethylsilyl; TBDMS, tert-butyldimethylsilyl.
Regioselective 2′-Silylation of Purine Ribonucleosides for Phosphoramidite RNA Synthesis
Acetonitrile (anhydrous, dried over 3Å molecular sieves) 0.1 M triethylammonium acetate (TEAA) 4,4′-Dimethoxytrityl chloride (DMTr-Cl) 5% (v/v) methanol in dichloromethane Dichloromethane Concentrated aqueous ammonium hydroxide N-Methylmorpholine Trimethylchlorosilane Adenosine phenoxyacetylating reagent (see recipe) Sodium bicarbonate Ethyl acetate
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Current Protocols in Nucleic Acid Chemistry
Petroleum ether Ammonium phenyl-H-phosphonate (see recipe) 1,8-Diazabicyclo[5.4.0]undec-7-ene (DBU) tert-Butyldimethylsilyl chloride (TBDMS⋅Cl) 0.5 M potassium phosphate buffer, pH 7.0 (APPENDIX 2A) Glycerol 1-Adamantanecarbonyl chloride Diisopropylammonium tetrazolide Argon source 2-Cyanoethyl tetraisopropylphosphorodiamidite Methylene chloride (anhydrous) Triethylamine (anhydrous) 250-mL and 25-mL round-bottom flask Rotary evaporator Silica gel 60F TLC plates (Merck) Waters XTerra 2.5-µm C18 chromatography column Vacuum pump Septum Vent needle Desiccator with P2O5 Additional reagents and equipment for TLC (APPENDIX 3D), column chromatography (APPENDIX 3E), and HPLC (UNIT 10.5) Tritylate adenosine 1. Suspend 1.34 g (5 mmol) adenosine in 10 mL pyridine in a 250-mL round-bottom flask with a magnetic stir bar and concentrate to dryness using a rotary evaporator and vacuum pump. Repeat this azeotropic drying process two times with 10-mL portions of pyridine. 2. Suspend the dry adenosine in 60 mL pyridine, reduce the volume to ∼50 mL, and add 2.6 mL (20 mmol) dimethylformamide dimethyl acetal. 3. Seal the flask with a septum and displace the air with nitrogen through a vent needle in the septum. After a few minutes, remove the nitrogen line and vent needle. Allow to sit for 1 hr. 4. Concentrate to an oil, dissolve the oil in 60 mL pyridine, and concentrate to ∼50 mL. 5. Add 2.03 g (6 mmol) 4,4′-dimethoxytrityl chloride and stir 2 to 3 hr. 6. Check the reaction by HPLC using a gradient of 2:98 to 80:20 acetonitrile: 0.1 M TEAA, pH 6.8 on a C18 column or by TLC (APPENDIX 3D) on silica gel plates using 60F methanol in dichloromethane (Table 2.8.1). Examine the plate under UV light, and then hold it over an open container of fresh aqueous HCl to observe trityl-containing spots. When the TLC plate is held over an open container of HCl, the fumes will cause any trityl-containing spots to turn a bright orange. If the reaction is not complete after 1 hr, add 0.5 mmol more of 4,4′-dimethoxytrityl chloride and wait an additional 1 hr. HPLC and TLC mobility of compounds in the reaction are listed in Table 2.8.1.
Protection of Nucleosides for Oligonucleotide Synthesis
2.8.3 Current Protocols in Nucleic Acid Chemistry
Supplement 6
Table 2.8.1
HPLC and TLC Mobility of Compounds Described in Figures 2.8.1 and 2.8.2
Adenosine TLC Rf (methanol:dichloromethane) (5:95)
(10:90)
HPLCa Retention time, min
0.09 0.08 0.01 minor 0.03 major 0.55 0.48 0.80, 0.84
0.27 0.37 0.08 minor 0.13 major 0.83 0.75 0.73, 0.90
11.3 12.5 12.7 major 13.4 minor 15.7 16.2 —b
11 13 15a,b
0.00 0.05 —
0.10 0.44 —
17a 17b 18
0.41 0.32 0.38, 0.43
0.58 0.53 0.58, 0.83
— 11.0 11.1 major 12.2 minor 13.9 14.5 —b
Compound (structure number in figures) 3 4 6a,6b 8a 8b 9 Guanosine
a
Gradient of 2:98 to 80:20 acetonitrile:0.1 M TEAA (pH 6.8) over 10 min, remaining at 80:20 for 5 min, then back down to 2:98 over 2 min, at 1 mL/min on a Waters XTerra 2.5 µm C18 column. b Too hydrophobic to analyze conveniently on HPLC.
7. Add 10 mL methanol to quench the excess reagent, wait 5 min, and then pour the solution into 100 mL water containing 1 g (12 mmol) sodium bicarbonate. 8. Extract the solution two times with 80-mL portions of dichloromethane, concentrate the combined organic layers, dissolve the residue in 25 mL pyridine, and add 25 mL concentrated aqueous ammonium hydroxide. Seal the flask tightly and heat 5 hr at 60°C. Use caution in heating this closed system. Use a shield and carefully inspect the flask for cracks or defects that might weaken it.
9. Cool to room temperature, open the flask carefully, and check the mixture by HPLC or TLC to have a record of the retention time or Rf value of S.3 (Table 2.8.1). 10. Concentrate the mixture with frequent additions of pyridine so that the water is removed azeotropically to give S.3 as an oil. Dry by evaporation of pyridine as in step 1, leaving dry S.3 in ∼50 mL pyridine. Phenoxyacetylate 11. To the solution of S.3 in 50 mL dry pyridine, add 5.5 mL (50 mmol) N-methylmorpholine. Seal the flask with a septum and displace the air with nitrogen through a vent needle in the septum. Keep the nitrogen flowing slowly.
Regioselective 2′-Silylation of Purine Ribonucleosides for Phosphoramidite RNA Synthesis
12. Cool this mixture in an ice-bath and add 3.2 mL (25 mmol) trimethylchlorosilane over 3 to 5 min. Trimethylchlorosilane is highly moisture sensitive. Use of special bottles such as the Aldrich SURE-SEAL system will help to prevent deterioration of the reagent.
13. Remove the flask from the ice-bath and maintain for 1 hr at room temperature.
2.8.4 Supplement 6
Current Protocols in Nucleic Acid Chemistry
14. Add all of the freshly prepared adenosine phenoxyacetylating reagent using a syringe. Remove the nitrogen line and vent needle and stir 12 to 18 hr. 15. Check the reaction by HPLC or TLC. If the reaction is not complete, add 0.5 mmol more of the phenoxyacetylating reagent and wait an additional 2 hr.
16. Pour the mixture into 100 mL water containing 2.5 g (30 mmol) sodium bicarbonate, extract two times with 100-mL portions of dichloromethane, and concentrate the combined organic layers to dryness. 17. Dissolve the residue in 50 mL pyridine, add 25 mL water, and stir 12 to 18 hr. 18. Concentrate the solution and check the mixture by HPLC or TLC. 19. Purify the residue by column chromatography (APPENDIX 3E) on silica gel using 0:100 to 15:85 (v/v) methanol/dichloromethane to give pure S.4 in yields of up to ∼90%. Silylate 20. To 2.63 g (15 mmol) of ammonium phenyl-H-phosphonate, add 2.3 mL (15 mmol) DBU and co-evaporate with 50 mL pyridine. 21. Dissolve the residue in 120 mL pyridine, concentrate to ∼100 mL, add 2.26 g (15 mmol) TBDMS-Cl and mix. Place 3.52 g (5 mmol) S.4 in a dry 250-mL round-bottom flask and dry by evaporation of pyridine as in step 1. Using a syringe, add the mixture prepared in step 21 to S.4, followed by 3.8 mL (25 mmol) DBU. The silylation reaction can be performed using other N- and 5′-O-protected adenosine derivatives.
22. Stir 5 to 8 hr and check the reaction by HPLC or TLC. If the reaction is not complete, add 1 mmol more of DBU and wait another 2 hr.
23. Pour the mixture into 100 mL of 0.5 M aqueous potassium phosphate buffer, pH 7.0, and extract two times with 100-mL portions of dichloromethane. Concentrate the combined organic layers to dryness. Dephosphonylate 24. Add 1.38 g (15 mmol) glycerol to the residue and dry the mixture by co-evaporation with 50 mL pyridine. 25. Dissolve the residue in 60 mL pyridine, concentrate to ∼50 mL, and add 2.98 g (15 mmol) of 1-adamantanecarbonyl chloride. Stir 10 min. 26. Pour the solution into 100 mL of 0.5 M aqueous potassium phosphate buffer and extract two times with 100-mL portions of dichloromethane. Concentrate the combined organic layers to dryness. 27. Check the mixture by HPLC or TLC and purify the residue by column chromatography in silica gel using 60:40 to 100:0 (v/v) ethyl acetate petroleum ether to give S.8a in yields of ∼65% from S.4. The isomers can be distinguished because the 2′-O-silyl isomer runs faster than the 3′-O-silyl isomer both on silica gel and C18 reversed-phase chromatography. See Table 2.8.2 for NMR data. If desired, the identity can be verified by two-dimensional COSY NMR: for the 2′-O-silyl isomer, the hydroxyl resonance only shows a cross-peak to the 3′-H resonance, while for the 3′-O-silyl isomer, the hydroxy resonance only shows a cross-peak to the 2′-H resonance.
Protection of Nucleosides for Oligonucleotide Synthesis
2.8.5 Current Protocols in Nucleic Acid Chemistry
Supplement 6
Phosphitylate 28. In an oven-dried 25-mL round-bottom flask, place 3.05 g (3.0 mmol) of pure S.8a and 0.28 g (1.5 mmol) of diisopropylammonium tetrazolide. Dry the flask and a rubber septum (not inserted) in an evacuated desiccator over P2O5 overnight. It is critical that the reaction be absolutely anhydrous. Further, the starting material must be pure, since the product can only tolerate a very fast chromatographic purification to remove the excess reagent.
29. Open the desiccator under argon and immediately insert the septum. Displace any air with argon through a vent needle in the septum. Add 15 mL of dry dichloromethane through the septum and swirl 5 to 10 min to dissolve the solids completely. 30. Cool the flask in an ice bath at 0° and add 1.00 mL (3.0 mmol) of 2-cyanoethyl tetraisopropylphosphorodiamidite. Keep the mixture at 0° for one hour and swirl it every 15 min or so. 31. Remove a small sample carefully with a syringe with an oven-dried needle to check by HPLC or TLC. Normally, the reaction will be about 75% to 85% done. 32. Remove the flask from the ice bath and keep it at room temperature. Add another 0.5 mL (1.5 mmol) of the phosphitylating reagent and allow the reaction to proceed for several more hours. Check by HPLC or TLC no more than once per hr, each time using a dry syringe needle. Note that the product exists as a pair of diastereomers, since the phosphorus atom is chiral. Any inadvertent hydrolysis of the product results in a pair of hydrogen phosphonate diesters that are much more polar than the product. Some hydrolysis will occur during HPLC and TLC analysis.
33. Prepare a small glass column containing about 10 cm of silica gel packed in 98:2 dry methylene chloride:triethylamine. 34. Place the reaction mixture directly onto this column and load it using nitrogen pressure. Wash the column using nitrogen pressure with about 30 mL of 98:2 dry methylene chloride:triethylamine, followed by 49:1:50 dry methylene chloride:triethylamine:dry acetonitrile. The product normally elutes after 20 to 50 mL, sometimes just after a yellow impurity. It is very important to work as quickly as possible so as not to leave the product in solution any longer than necessary, since it will start to degrade immediately. A quick way to check the fractions for product is to spot them on a grid marked on a TLC plate. The plate does not have to be developed, just checked for UV-active material. Then the first and last fractions can be checked by HPLC or on a developed TLC plate.
36. Combine the fractions containing pure product and evaporate to a foam. Dry the product in a desiccator over P2O5. 37. Check the product for purity by 31P NMR (Table 2.8.2). BASIC PROTOCOL 2
Regioselective 2′-Silylation of Purine Ribonucleosides for Phosphoramidite RNA Synthesis
SYNTHESIS OF 5′-O-(4,4′-DIMETHOXYTRITYL)-2′-O-tertBUTYLDIMETHYSILYL-2-N-ACYLGUANOSINE In this protocol, N- and 5′-O-protected guanosine (S.13 in Fig. 2.8.2) is generated in a similar fashion as S.4 in Basic Protocol 1, except that guanosine is first amino protected (whereas adenosine is first tritylated) because the N-phenoxyacetyl derivative is crystalline and therefore easier to isolate. Silylation, dephosphorylation, and phosphitylation are performed following the procedures for adenosine.
2.8.6 Supplement 6
Current Protocols in Nucleic Acid Chemistry
Figure 2.8.2 N-Phenoxylacetylation, dimethoxytritylation, silylation, dephosphonylation, and phosphitylation of guanosine. Abbreviations: Ad, adamantanecarbonyl; DBU, 1,8-diazabyclyclo[5.4.0]undec-7-ene; DMTr, 4,4′-dimethoxytrityl; HOBT, 1-hydroxybenzotriazole; Ph, phenyl; i-Pr, isopropyl; TMS, trimethylsilyl; TBDMS, tert-butyldimethylsilyl.
Materials Guanosine Pyridine Nitrogen source Trimethylchlorosilane Guanosine phenoxyacetylating reagent (see recipe) 2-Propanol Methanol Dichloromethane Acetonitrile (anhydrous, dried over 3Å molecular sieves) 0.1 M triethylammonium acetate (TEAA) Dimethylformamide dimethyl acetal 4,4′-Dimethoxytrityl chloride (DMTr-Cl) Methanol
Protection of Nucleosides for Oligonucleotide Synthesis
2.8.7 Current Protocols in Nucleic Acid Chemistry
Supplement 6
Sodium bicarbonate Dichloromethane Ammonium phenyl-H-phosphonate (see recipe) 1,8-Diazabicyclo[5.4.0]undec-7-ene (DBU) tert-Butyldimethylsilyl chloride (TBDMS-Cl) 0.5 M potassium phosphate buffer, pH 7.0 (APPENDIX 2A) Glycerol 1-Adamantanecarbonyl chloride 5:95 to 15:85 (v/v) acetone/dichloromethane (optional; for phosphoramidite synthesis) 250-mL round-bottom flasks Magnetic stir bar Rotary evaporator Silica gel 60F TLC plates (Merck) Waters XTerra 2.5-µm C18 chromatography column Vacuum pump Septum Vent needle Desiccator with P2O5 Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Phenoxyacetylate guanosine 1. Suspend 1.42 g (5 mmol) guanosine in 10 mL pyridine in a 250-mL round-bottom flask with a magnetic stir bar and concentrate to dryness using a rotary evaporator and vacuum pump. Repeat this azeotropic drying process two times with 10-mL portions of pyridine. 2. Suspend the dry guanosine in 60 mL pyridine and concentrate to ∼50 mL. 3. Seal the flask with a septum and displace the air with nitrogen through a vent needle in the septum. Keep the nitrogen flowing slowly. 4. Cool this mixture in an ice-bath and add 3.8 mL (30 mmol) trimethylchlorosilane over 3 to 5 min. Trimethylchlorosilane is highly moisture sensitive. Use of special bottles such as the Aldrich SURE-SEAL system will help to prevent deterioration of the reagent.
5. Remove the flask from the ice bath and maintain 1 hr at room temperature. 6. Add all the freshly prepared guanosine phenoxyacetylating reagent using a syringe. Remove the nitrogen line and vent needle and stir 36 hr. 7. Add 30 mL water, concentrate the solution, and co-evaporate two times with 30-mL portions of water to a final volume of ~15 mL. 8. Filter the slurry. The solid contains both the product (S.11) and some 1-hydroxybenzotriazole from the guanosine phenoxyacetylating reagent. Regioselective 2′-Silylation of Purine Ribonucleosides for Phosphoramidite RNA Synthesis
9. To remove the 1-hydroxybenzotriazole, shake the solid thoroughly with a 50-mL portion of water and filter. Repeat with 20 mL water followed by three 20-mL portions of 2-propanol to give S.11 as a colorless solid in yields of up to 95%.
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10. Check the product by HPLC using a gradient of 2:98 to 80:20 acetonitrile: 0.1 M TEAA pH 6.8 on a C18 column or TLC (APPENDIX 3D) on silica gel 60F plates using the appropriate concentration of methanol in dichloromethane (Table 2.8.1). 11. Dry the product in a desiccator over P2O5 at least overnight and check that the yield is <100%. A yield >100% means that the product is contaminated with hydroxybenzotriazole and one should repeat the thorough shaking with 2-propanol.
Tritylate 12. Dissolve the dry S.11 in 60 mL pyridine, concentrate to ∼50 mL, and add 0.8 mL (6 mmol) dimethylformamide dimethyl acetal. 13. Seal the flask with a septum and displace the air with nitrogen through a vent needle in the septum. After a few minutes, remove the nitrogen line and vent needle. Allow to sit for 1 hr. The dimethylformamide dimethyl acetal will slowly replace the phenoxyacetyl group so the time and the amount of this reagent used is kept to a minimum.
14. Concentrate to an oil, dissolve the oil in 60 mL pyridine, and concentrate to ∼50 mL. 15. Add 2.03 g (6 mmol) 4,4′-dimethoxytrityl chloride and stir 2 to 3 hr. 16. Check the reaction by HPLC or TLC. Examine the plate under UV light, and then hold it over an open container of fresh aqueous HCl to observe trityl-containing spots. When the TLC plate is held over an open container of HCl, the fumes will cause any trityl-containing spots to turn a bright orange. If the reaction is not complete after 1 hr, add an additional 0.5 mmol of 4,4′-dimethoxytrityl chloride and wait another 1 hr.
17. Add 10 mL methanol to quench the excess reagent, wait 5 min, and then pour the solution into 100 mL water containing 1 g (12 mmol) sodium bicarbonate. 18. Extract the solution two times with 80-mL portions of dichloromethane and concentrate the combined organic layers to dryness. 19. Check the mixture by HPLC or TLC. Purify the residue by column chromatography (APPENDIX 3E) on silica gel using 0:100 to 10:90 (v/v) methanol/dichloromethane to give pure S.13 in yields of up to ∼90%. Silylate 20. To 2.63 g (15 mmol) ammonium phenyl-H-phosphonate, add 2.3 mL (15 mmol) DBU and co-evaporate with 50 mL pyridine. 21. Dissolve the residue in 120 mL pyridine, concentrate to ∼100 mL, add 2.26 g (15 mmol) TBDMS-Cl and mix. Place 3.52 g (5 mmol) S.4 in a dry 250-mL round-bottom flask and dry by evaporation of pyridine as in step 1. Using a syringe, add the mixture prepared in step 21 to S.4, followed by 3.8 mL (25 mmol) DBU. The silylation reaction can be performed using other N- and 5′-O-protected guanosine derivatives.
22. Stir 5 to 8 hr and check the reaction by HPLC or TLC. If the reaction is not complete, add 1 mmol more of DBU and wait another 2 hr.
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23. Pour the mixture into 100 mL of 0.5 M aqueous potassium phosphate buffer, pH 7.0, and extract two times with 100-mL portions of dichloromethane. Concentrate the combined organic layers to dryness. Dephosphonylate 24. Add 1.38 g (15 mmol) glycerol to the residue and dry the mixture by co-evaporation with 50 mL pyridine. 25. Dissolve the residue in 60 mL pyridine, concentrate to about ∼50 mL, and add 2.98 g (15 mmol) of 1-adamantanecarbonyl chloride. Stir 10 min. 26. Pour the solution into 100 mL of 0.5 M potassium phosphate buffer, pH 7.0, and extract two times with 100-mL portions of dichloromethane. Concentrate the combined organic layers to dryness. 27. Check the mixture by HPLC or TLC and purify the residue by column chromatography on silica gel using 5:95 to 15:85 acetone/dichloromethane to give S.17a in yields of ∼65% from S.13. The isomers can be distinguished because the 2′-O-silyl isomer runs faster than the 3′-O-silyl isomer both on silica gel and C18 reversed-phase chromatography. See Table 2.8.2 for NMR data. If desired, the identity can be verified by two-dimensional COSY NMR: for the 2′-O-silyl isomer, the hydroxyl resonance only shows a cross-peak to the 3′-H resonance, while for the 3′-O-silyl isomer, the hydroxyl resonance only shows a cross-peak to the 2′-H resonance.
Phosphitylate 28. In an oven-dried 25-mL round-bottom flask, place 3.05 g (3.0 mmol) of pure S.17a and 0.28 g (1.5 mmol) of diisopropylammonium tetrazolide. Dry the flask and a rubber septum (not inserted) in an evacuated desiccator over P2O5 overnight. It is critical that the reaction be absolutely anhydrous. Further, the starting material must be pure, since the product can only tolerate a very fast chromatographic purification to remove the excess reagent.
29. Open the desiccator under argon and immediately insert the septum. Displace any air with argon through a vent needle in the septum. Add 15 mL of dry dichloromethane through the septum and swirl 5 to 10 min to dissolve the solids completely. 30. Cool the flask in an ice bath at 0° and add 1.00 mL (3.0 mmol) of 2-cyanoethyl tetraisopropylphosphorodiamidite. Keep the mixture at 0° for 1 hr and swirl it every 15 min or so. 31. Remove a small sample carefully with a syringe with an oven-dried needle to check by HPLC or TLC. Normally, the reaction will be about 75% to 85% done. 32. Remove the flask from the ice bath and keep it at room temperature. Add another 0.5 mL (1.5 mmol) of the phosphitylating reagent and allow the reaction to proceed for several more hours. Check by HPLC or TLC no more than once per hr, each time using a dry syringe needle.
Regioselective 2′-Silylation of Purine Ribonucleosides for Phosphoramidite RNA Synthesis
Note that the product exists as a pair of diastereomers, since the phosphorus atom is chiral. Any inadvertent hydrolysis of the product results in a pair of hydrogen phosphonate diesters that are much more polar than the product. Some hydrolysis will occur during HPLC and TLC analysis.
33. Prepare a small glass column containing about 10 cm of silica gel packed in 98:2 dry methylene chloride:triethylamine.
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Protection of Nucleosides for Oligonucleotide Synthesis
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34. Place the reaction mixture directly onto this column and load it using nitrogen pressure. Wash the column using nitrogen pressure with about 30 mL of 98:2 dry methylene chloride:triethylamine, followed by 49:1:50 dry methylene chloride:triethylamine:dry acetonitrile. The product normally elutes after 20 to 50 mL, sometimes just after a yellow impurity. It is very important to work as quickly as possible so as not to leave the product in solution any longer than necessary, since it will start to degrade immediately. A quick way to check the fractions for product is to spot them on a grid marked on a TLC plate. The plate does not have to be developed, just checked for UV-active material. Then the first and last fractions can be checked by HPLC or on a developed TLC plate.
36. Combine the fractions containing pure product and evaporate to a foam. Dry the product in a desiccator over P2O5. 37. Check the product for purity by 31P NMR (Table 2.8.2). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Adenosine phenoxyacetylating reagent Co-evaporate 2.03 g (15 mmol) of 1-hydroxybenzotriazole and 2.2 mL (20 mmol) N-methylmorpholine two times with 20-mL portions of acetonitrile. Dissolve the residue in 50 mL dichloromethane. Add 2.1 mL (15 mmol) phenoxyacetyl chloride and shake for 10 min. Prepare fresh. Ammonium phenyl-H-phosphonate Add 38.3 mL (0.20 mol) diphenyl phosphite over 10 min to 400 mL of 7.4 M aqueous ammonia. Stir for 1 hr. Concentrate to dryness and co-evaporate the residue two times with 100-mL portions of absolute ethanol. Stir the residue with 400 mL ethyl ether for 30 min to give up to 85% of the colorless crystalline product. Store up to 1 month at –20°C. Diisopropylammonium tetrazolide Place 2.00 g (28.5 mmol) of solid tetrazole in a dry 250-mL round-bottom flask containing a dry stir bar and add 130 mL of dry acetonitrile. Stir until dissolved, then add 9.0 mL (63.9 mmol) of freshly distilled diisopropylamine. After 2 min of stirring, collect the white precipitate by filtration in a dry glass funnel and wash it four times with 10-mL portions of dry acetonitrile. Dry the solid in a desiccator over P2O5. Store up to 3 months at –20°C. Guanosine phenoxyacetylating reagent Co-evaporate 1.35 g (10 mmol) of 1-hydroxybenzotriazole and 1.1 mL (10 mmol) N-methylmorpholine two times with 20-mL portions of acetonitrile. Dissolve the residue in 25 mL dichloromethane. Add 1.39 mL (10 mmol) phenoxyacetyl chloride and shake for 10 min. Prepare fresh. COMMENTARY Background Information Regioselective 2′-Silylation of Purine Ribonucleosides for Phosphoramidite RNA Synthesis
The procedures described here use adenosine and guanosine that are 5′-protected with the 4,4′-dimethoxytriyl (DMTr) group and amino-protected with the labile phenoxyacetyl group (Wu et al., 1988; Chaix et al., 1989; Singh
and Nahar, 1995; Sinha et al., 1995). They should also be applicable to nucleosides containing most other amino-protecting groups. Amino protection of adenosine with the benzoyl group and guanosine with the isobutyryl group can be carried out by standard literature
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procedures, and these derivatives are more easily handled than are the more-labile phenoxyacetylated compounds. A general discussion of amino-protecting groups and literature references is given in UNIT 2.1. The procedures developed by Ogilvie for 2′-silylation proceed with only modest regioselectivity (Ogilvie, 1978, 1979). Although use of silver nitrate improves the selectivity, the results are variable and do not approach the selectivity of the above protocols (Hakimelahi, 1982). A general discussion of 2′-protecting groups, including the 2′-TBDMS group, is given in UNIT 2.2.
Critical Parameters The most difficult of the above protocols to carry out are the tritylation and phenoxyacetylation steps used for preparation of S.4 and S.13. Skill and practice are required to achieve high yields on these protection reactions, largely because of the phenoxyacetyl group. In contrast, the regioselective silylation and dephosphonylation reactions work well even in the hands of unskilled researchers.
Anticipated Results For the reasons discussed above, some experience is required to achieve high yields for the phenoxyacetylation reactions, and initial efforts are likely be give more modest yields (e.g., 50%). With experience, yields of ~90% can be expected. The regioselectivity of the silylation reaction is invariably 85% to 90% regardless of experience.
Time Considerations The total time for conversion of adenosine or guanosine to the fully protected derivatives S.8a and S.17a, respectively, is ∼1 week.
Literature Cited Chaix, C., Duplaa, A.M., Molko, D., and Téoule, R. 1989. Solid phase synthesis of the 5′-half of the initiator t-RNA from B. subtilis. Nucl. Acids Res. 17:7381-7393.
Hakimelahi, G.H., Proba, Z.A., and Ogilvie, K.K. 1982. New catalysts and procedures for the dimethoxytritylation and selective silylation of ribonucleosides. Can. J. Chem. 60:1106-1113. Ogilvie, K.K., Beaucage, S.L., Schifman, A.L., Theriault, N.Y., and Sadana, K.L. 1978. The synthesis of oligoribonucleotides. II. The use of silyl protecting groups in nucleoside and nucleotide chemistry. VII. Can. J. Chem. 56:27682780. Ogilvie, K.K., Schifman, A.L., and Penney, C.L. 1979. The synthesis of oligoribonucleotides. III. The use of silyl protecting groups in nucleoside and nucleotide chemistry. VIII. Can. J. Chem. 57:2230-2238. Singh, K.K. and Nahar, P. 1995. An improved method for the synthesis of N-phenoxyacetylribonucleosides. Synth. Commun. 25:1997-2003. Sinha, N.D., Davis, P., Schultze, L.M., and Upadhya, K. 1995. A simple method for N-acylation of adenosine and cytidine nucleosides using carboxylic acids activated in situ with carbonyldiimidazole. Tetrahedron Lett. 36:9277-9280. Song, Q., Wang, W., Fischer, A., Zhang, X., Gaffney, B.L., and Jones, R.A. 1999. High yield protection of purine ribonucleosides for phosphoramidite RNA synthesis. Tetrahedron Lett. 40:4153-4156. Wu, T., Ogilvie, K.K., and Pon, R.T. 1988. N-Phenoxyacetylated guanosine and adenosine phosphoramidites in the solid phase synthesis of oligoribonucleotides: Synthesis of a ribozyme sequence. Tetrahedron Lett. 29:4249-4252. Zemlicka, J. 1963. Reactions of dimethylformamide acetals with some heterocyclic systems. Collect. Czech. Chem. Commun. 28:1060-1062. Zhang, X., Abad, J.-L., Huang, Q., Zeng, F., Gaffney, B.L., and Jones, R.A. 1997. High yield protection of purine ribonucleosides for H-phosphonate RNA synthesis. Tetrahedron Lett. 38:7135-7138.
Contributed by Barbara L. Gaffney and Roger A. Jones Rutgers, The State University of New Jersey Piscataway, New Jersey
Protection of Nucleosides for Oligonucleotide Synthesis
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Preparation of 2′-O-[(Triisopropylsilyl)oxy]methylprotected Ribonucleosides
UNIT 2.9
This unit describes, in detail, the preparation of 2′-O-[(triisopropylsilyl)oxy]methyl (TOM)–protected phosphoramidite building blocks derived from the ribonucleosides adenosine, cytidine, guanosine, and uridine (see Basic Protocol). These building blocks are suitable for the chemical synthesis of RNA oligonucleotides under DNA-coupling conditions. Additionally, protocols are presented for the preparation of suitably protected purines, N6-acetyl-5′-O-(4,4′-dimethoxytrityl)adenosine and N2-acetyl-5′-O-(4,4′-dimethoxytrityl)guanosine (see Support Protocols 1 and 2, respectively), and for the protecting group reagent [(triisopropylsilyl)oxy]methyl chloride (TOM⋅Cl; see Support Protocol 3). The two corresponding pyrimidine derivatives are commercially available (e.g., ChemGenes). The preparation of phosphoramidites from these building blocks and the synthesis of oligoribonucleotides is described in UNIT 3.8. CAUTION: All reactions must be performed in a well-ventilated fume hood to avoid exposure to dibutyltin dichloride and TOM⋅Cl. These procedures should be performed only by personnel trained and experienced in organic synthesis. Standard precautions to prevent excessive exposure to toxic chemicals and solvents should be followed. All reactions should first be performed on a small scale. INTRODUCTION OF THE [(TRIISOPROPYLSILYL)OXY]METHYL GROUP INTO N-ACETYLATED, 5′-O-DIMETHOXYTRITYLATED RIBONUCLEOSIDES
BASIC PROTOCOL
The introduction of the TOM group into the four N-acetylated, 5′-O-dimethoxytritylated ribonucleosides is presented in Figure 2.9.1. Alkylation of the 2′,3′-diols of ribonucleosides is achieved by first forming their cyclic 2′,3′-di-O-dibutylstannyl derivatives under basic conditions with n-dibutyltin dichloride and N-ethyl-N,N-diisopropylamine in 1,2-dichloroethane for 1 hr at 25°C. These activated intermediates are subsequently treated with 1.1 to 1.3 equivalents of TOM⋅Cl for 20 min at 80°C. After the aqueous workup, the 2′-O-alkylated and 3′-O-alkylated nucleosides (S.3a-d and S.2a-d, respectively) are isolated in pure form by chromatography on silica gel (unreacted starting materials can also be recovered). Under these optimized reaction conditions, side reactions (alkylation of the base moieties) are observed only to a very small extent (<5%). Fair to good yields of the desired 2′-O-alkylated products S.3a-d are obtained (40% to 60%), along with various amounts of the 3′-O-alkylated isomers S.2a-d (5% to 30%). In all cases, the predominant first-eluting isomer is the desired 2′-O-alkylated compound. In the following protocol, 5′-O-dimethoxytritylated, N-acetylated ribonucleoside derivatives are used as starting materials. However, employing the identical reaction conditions, the TOM group can also be introduced with similar results to 5′-O-dimethoxytritylated ribonucleosides that are protected with other N-acyl protecting groups (e.g., benzoyl, isobutyryl).
Protection of Nucleosides for Oligonucleotide Synthesis Contributed by Stefan Pitsch and Patrick A. Weiss Current Protocols in Nucleic Acid Chemistry (2001) 2.9.1-2.9.14 Copyright © 2001 by John Wiley & Sons, Inc.
2.9.1 Supplement 7
Figure 2.9.1 Preparation of the N-acetylated-5′-O-DMTr-2′-O-TOM-protected ribonucleosides S.3a-d. n-Bu2SnCl2, n-dibutyltin chloride; DMTr, 4,4′-dimethoxytrityl; i-Pr2NEt, N-ethyl-N,N-diisopropylamine.
Materials Nitrogen source 5′-O-Dimethoxytritylated, N-acetylated ribonucleosides: N6-Ac-5′-O-DMTr-adenosine (S.1a; see Support Protocol 1) N2-Ac-5′-O-DMTr-guanosine (S.1b; see Support Protocol 2) N4-Ac-5′-O-DMTr-cytidine (S.1c; ChemGenes) 5′-O-DMTr-uridine (S.1d; ChemGenes) 1,2-Dichloroethane (reagent grade) N-Ethyl-N,N-diisopropylamine n-Dibutyltin dichloride [(Triisopropylsilyl)oxy]methyl chloride (TOM⋅Cl; S.6; see Support Protocol 3) Ethyl acetate (for chromatography, technical grade) Saturated aqueous sodium bicarbonate (NaHCO3) 9:1 (v/v) ethyl acetate/hexane (for A, C, and U) 19:1 (v/v) dichloromethane/methanol (for G) Anisaldehyde reagent (see recipe) Dichloromethane (CH2Cl2) with and without 5% triethylamine (with TEA for G only) Magnesium sulfate Celite Silica gel (230 to 400 mesh) 6:4, 5:5, 4:6, and so on (v/v) hexane/ethyl acetate containing 2% triethylamine (for A, C, and U) Sand Methanol (for G) 95:5:0.1 (v/v/v) dichloromethane/methanol/triethylamine
Preparation of 2′-O[(Triisopropylsilyl)oxy]methylprotected Ribonucleosides
500-mL two-neck flask equipped with a reflux condenser and stir bar Balloon Rubber septum 80°C water bath TLC plates (Merck silica gel 60, 4 × 10–cm) 254-nm UV lamp 1-L separatory funnel
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1-L Erlenmeyer flask 6-cm glass filter Rotary evaporator equipped with a vacuum pump or water aspirator 5-cm-diameter chromatography column Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare TOM-protected ribonucleosides 1. In a 500-mL two-neck flask equipped with a reflux condenser, a stir bar, a balloon filled with N2, and a rubber septum, dissolve 50 mmol of 5′-O-dimethoxytritylated, N-acetylated ribonucleoside in 200 mL of 1,2-dichloroethane. 2. While stirring, add 29.2 mL (180 mmol) N-ethyl-N,N-diisopropylamine and then slowly add 18.8 g (50 mmol) solid n-dibutyltin dichloride. Stir 1 hr at room temperature. It is extremely important that the dibutyltin dichloride be added after the N-ethyl-N,N-diisopropylamine. CAUTION: Dibutyltin dichloride is very toxic and quite volatile. It must be handled in a well-ventilated fume hood.
3. Place the flask in an 80°C water bath and stir 5 to 10 min. 4. Add 14 mL (65 mmol) TOM⋅Cl (S.6) with a syringe and continue stirring and heating for an additional 15 min at 80°C. Remove the flask from the water bath. 5. Using a syringe, transfer ∼50 µL of the clear reaction mixture to a small tube containing 50 µL ethyl acetate/50 µl saturated aqueous NaHCO3 and mix thoroughly. 6. Analyze the clear supernatant by TLC (APPENDIX 3D). Co-spot the starting material for comparison. Elute with 9:1 (v/v) ethyl acetate/hexane for adenosine, cytidine, and uridine derivatives, and with 19:1 (v/v) dichloromethane/methanol for the guanosine derivative. 7. Visualize by exposing the plate to a 254-nm UV lamp and staining it with anisaldehyde reagent. Typically, the Rf value of the desired 2′-O-alkylated compound S.3a-d is ∼0.55 and the Rf value of the 3′-O-alkylated byproduct S.2a-d is ∼0.45. The Rf value of the starting material S.1a-d is ∼0.1, and the Rf value of the nucleobase-alkylated byproduct is ∼0.9.
8. Optional: If the turnover is less than ∼70% to 80% (∼20% remaining starting material), return the reaction mixture to the 80°C water bath and add an additional 0.2 eq TOM⋅Cl. Stir an additional 15 min at 80°C and analyze again by TLC. No more TOM⋅Cl should be added if the amount of nucleobase-alkylated byproducts (Rf ∼0.9) exceeds 5% to 10%.
9. Dilute the reaction mixture (step 4) with 250 mL dichloromethane and add it under stirring to 400 mL saturated aqueous NaHCO3. Stir the biphasic mixture for 10 to 15 min. During this time a white solid precipitates.
10. Pour the mixture into a 1-L separatory funnel and allow the phases to separate. Pour the lower, yellow-colored organic phase (which contains some colorless, solid material) into a 1-L Erlenmeyer flask.
Protection of Nucleosides for Oligonucleotide Synthesis
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11. Extract the aqueous phase once more with 250 mL dichloromethane. Combine this organic phase with first organic phase in the 1-L Erlenmeyer flask. 12. Add 5 to 10 g solid magnesium sulfate and vacuum filter the organic phase by means of a 6-cm glass filter covered with 1.5 cm Celite. 13. Wash the Celite with dichloromethane and evaporate the filtrate to dryness in a rotary evaporator. Isolate 2′-O-TOM-protected ribonucleosides 14. Prepare a slurry of 400 g silica gel in 6:4 (v/v) hexane/ethyl acetate containing 2% triethylamine for A, C, and U (S.3a,c,d) or in dichloromethane containing 5% triethylamine for G (S.3b). For detailed procedures for column chromatography, see APPENDIX 3E.
15. Pour slurry into a 5-cm-diameter column and carefully layer ∼3 cm sand on top. 16. Dissolve the crude product in a minimal amount of dichloromethane and place it carefully on top of the column. 17. Elute first with 1 L of the solvent mixture used to pack the column, and then with gradually more polar solvent, collecting 200-mL fractions. a. For A, C, and U, use 1 L each of hexane/ethyl acetate at serial dilutions of 5:5, 4:6, and so on. b. For G, use 1-L portions of dichloromethane with increased methanol at 0.5% increments (1000:5, 1000:10, and so on). Always include the amount of triethylamine indicated in step 14.
18. Check fractions periodically by TLC. As soon as the two alkylation products are completely eluted, wash column with 3 L of 95:5:0.1 (v/v/v) dichloromethane/methanol/triethylamine in order to isolate unreacted starting material. 19. Pool fractions that contain pure 2′-O-TOM-substituted ribonucleosides (the firsteluting main product, Rf typically 0.55). 20. Pool fractions that contain impure product separately and repeat chromatography with these fractions on an appropriate smaller scale. 21. Combine fractions that contain pure product, evaporate to dryness in a rotary evaporator, and dry overnight at high vacuum (<0.05 mbar). Typically, ∼15 to 20 g (40% to 60%) pure 2′-O-TOM-protected ribonucleoside derivatives are obtained in the form of a solid foam.
22. Check purity of the material by 1H-NMR. Only pure products (purity >95%) should be used for phosphoramidite and oligoribonucleotide synthesis (UNIT 3.8).
Preparation of 2′-O[(Triisopropylsilyl)oxy]methylprotected Ribonucleosides
N6-Ac-5′-O-DMTr-2′-O-TOM-adenosine (S.3a): 15.9 g (40%). TLC (hexane/ethyl acetate 1:9) 0.60; [a]D25 +11.5 (c = 1.36, CDCl3); λmax (methanol) 272 (20,500), 234 (24,700); δH (CDCl3) 0.99-1.08 (21 H, m, i-Pr3Si); 2.61 (s, CH3CO), 3.07 (br. d, J ≈ 3 Hz, HO-3′), 3.40 (dd, J = 3.9, 10.6 Hz, H-5′), 3.53 (dd, J = 3.5, 10.6 Hz, H′-5′), 3.78 (s, 2 OCH3), 4.30 (q, J = 3.7 Hz, H-4′), 4.57 (br. q, J ≈ 3 Hz, H-3′), 4.96 (t, J = 5.3 Hz, H-2′), 4.98 and 5.14 (2d, J = 5.0 Hz, OCH2O), 6.21 (d, J = 5.3 Hz, H-1′), 6.78-6.81 (4 H, m, DMTr), 7.23-7.44 (9 H, m, DMTr), 8.16 (s, H-2), 8.60 (s, H-8), 8.65 (br. s, HN); δC (CDCl3) 11.8 (d), 17.8 (q), 25.9 (q), 55.3 (q), 63.4 (t), 71.0 (d), 82.2 (d), 84.6 (d), 86.8 (s), 87.4 (d), 91.0 (t), 113.4 (d), 122.5 (s), 127.2 (d), 128.1 (d), 128.4 (d), 130.3 (d), 135.9, (s), 142.1 (d), 144.8 (s), 149.4 (s), 151.4 (s), 152.7 (d), 158.9 (s), 170.7 (s); m/z 798 (29, MH+), 303 (100). Anal. calcd. for C43H55N5O8Si: C, 64.72; H, 6.95; N, 8.78. Found: C, 64.64; H, 7.06; N, 8.74.
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N2-Ac-5′-O-DMTr-2′-O-TOM-guanosine (S.3b): 24.4 g (60%). TLC (CH2Cl2/methanol 19:1) 0.55; [a]D25 +7.7 (c = 0.85, CDCl3); λmax (methanol) 282sh (13,500), 275 (14,300), 260sh (17,500), 251 (19,200), 235 (26,500); δH (CDCl3) 0.94-1.12 (21 H, m, i-Pr3Si), 1.46 (s, CH3CO), 3.03 (br. d, J ≈ 3 Hz, HO-3′), 3.13 (dd, J = 2.8, 10.6 Hz, H-5′), 3.53 (dd, J = 2.0, 10.6 Hz, H′-5′), 3.76 and 3.78 (2s, 2 OCH3), 4.24 (q, J = 2.0 Hz, H-4′), 4.57 (ddd, J = 1.5, 5.0 Hz, 3.0, H-3′), 4.95 and 5.14 (2d, J = 4.7 Hz, OCH2O), 5.09 (dd, J = 5.3, 7.5 Hz, H-2′), 5.90 (d, J = 7.5 Hz, H-1′), 6.78-6.83 (4 H, m, DMTr), 7.18-7.57 (9 H, m, DMTr), 7.75 (br. s, NH), 7.81 (s, H-8), 11.82 (br. s, NH); δC (CDCl3) 11.8 (d), 17.8 (q), 23.7 (q), 55.4 (q), 63.9 (t), 70.8 (d), 81.5 (d), 84.5 (d), 86.5 (d), 86.9 (s), 91.1 (t), 113.5 (d), 122.4 (s), 127.3 (d), 128.2 (d), 128.3 (d), 130.3 (d), 135.9 (s), 136.3 (s), 139.1 (d), 145.3 (s), 147.4 (s), 148.7 (s), 156.3 (s), 159.0 (s), 171.8 (s). m/z 814 (17, MH+), 303 (100). Anal. calcd. for C43H55N5O9Si: C, 63.45; H, 6.81; N, 8.60. Found: C, 63.34; H, 6.84; N, 8.35. N4-Ac-5′-O-DMTr-2′-O-TOM-cytidine (S.3c): 17.4 g (45%). TLC (hexane/ethyl acetate 1:9) 0.55; [a]D25 +46.8 (c = 0.94, CDCl3); λmax (methanol) 300 (6,800), 283 (6,800), 236 (27,200); δH (CDCl3) 1.04-1.15 (21 H, m, i-Pr3Si), 2.21 (s, CH3CO), 3.34 (d, J = 8.4 Hz, HO-3′), 3.53 (dd, J = 2.5, 10.9 Hz, H-5′), 3.62 (dd, J = 1.8, 10.9 Hz, H’-5′), 3.81 and 3.82 (2s, 2 OCH3), 4.09 (br. dt, J ≈ 9, 2 Hz, H-4′), 4.23 (d, J = 5.0 Hz, H-2′), 4.37 (m, H-3′), 5.15 and 5.28 (2d, J = 4.6 Hz, OCH2O), 5.97 (s, H-1′), 6.84-6.88 (4 H, m, DMTr), 7.04 (d, J = 7.1 Hz, H-5), 7.24-7.44 (9 H, m, DMTr), 8.48 (d, J = 7.4 Hz, H-6), 8.62 (br. s, HN); δC (CDCl3) 11.9 (d), 17.8 (q), 24.9 (q), 55.3 (q), 61.4 (t), 67.9 (d), 83.50 (d), 83.55 (d), 87.2 (s), 90.2 (d), 90.9 (t), 96.8 (d), 113.5 (d), 127.4 (d), 128.3 (d), 128.4 (d), 128.5 (d), 130.4 (d), 135.6 (s), 135.8 (s), 144.6 (s), 145.1 (d), 155.3 (s), 159.0 (s), 163.1 (s), 170.6 (s); m/z 774 (23, MH+), 303 (100). Anal. calcd. for C42H55N3O9Si: C, 65.18; H, 7.16; N, 5.43. Found: C, 64.89; H, 7.18; N, 5.42. 5′-O-DMTr-2′-O-TOM-uridine (S.3d): 14.6 g (40%). TLC (hexane/ethyl acetate 1:4) 0.65; [a]D25 +42.9 (c = 1.00, CDCl3); λmax (methanol) 267 (13,100), 226 (28,600); δH (CDCl3) 1.05-1.15 (21 H, m, i-Pr3Si), 3.17 (br. d, J ≈ 3 Hz, HO-3′), 3.52 (br. d, J ≈ 11 Hz, H-5′), 3.55 (br. d, J ≈ 11 Hz, H′-5′), 3.80 (s, 2 OCH3); 4.11 (m, H-4′), 4.26 (dd, J = 3.5, 5.0 Hz, H-2′), 4.55 (br. t, J ≈ 3 Hz, H-3′), 5.03 and 5.23 (2d, J = 5.0 Hz, OCH2O), 5.29 (d, J = 8.1 Hz, H-5), 6.03 (d, J = 3.1 Hz, H-1′), 6.83-6.87 (4 H, m, DMTr); 7.24-7.40 (9 H, m, DMTr), 7.94 (d, J = 8.1 Hz, H-6), 8.38 (br. s, HN); δC (CDCl3) 11.9 (d), 17.8 (q), 55.3 (q), 62.3 (t), 69.5 (d), 83.0 (d), 83.8 (d), 87.3 (s), 88.0 (d), 90.8 (t), 102.4 (d), 113.5 (d), 127.4 (d), 128.2 (d), 128.4 (d), 130.4 (d), 135.4 (s), 135.6 (s), 140.4 (d), 144.6 (s), 150.4 (s), 159.0 (s), 159.1 (s), 163.3 (s); m/z 732 (21, M+), 303 (100). Anal. calcd. for C40H52N2O9Si: C, 65.55; H, 7.15; N, 3.82. Found: C, 65.29; H, 7.08; N, 4.02.
PREPARATION OF N6-ACETYL-5′-O-(4,4′-DIMETHOXYTRITYL)ADENOSINE
SUPPORT PROTOCOL 1
In this protocol, the preparation of N6-acetyl-5′-O-(4,4′-dimethoxytrityl)adenosine (S.1a) is presented. In this two-step process (Figure 2.9.2), N6-acetyladenosine (S.4a) is synthesized by first silylating the hydroxyl groups with trimethylsilyl chloride and then acetylating the amino group with acetyl chloride. Extractive work-up and cleavage of the trimethylsilyl ethers is carried out with acetic acid in methanol. The introduction of the 5′-O-(4,4′-dimethoxytrityl) group into the intermediates is carried out under standard conditions with 4,4′-dimethoxytrityl chloride/pyridine, giving the title compound (S.1a) in good yields. The intermediate (S.4a) is isolated by crystallization and the dimethoxytritylated compound (S.1a) is isolated by chromatography on silica gel. Materials Adenosine Pyridine (reagent grade) Nitrogen source Trimethylsilyl chloride (TMS⋅Cl) Acetonitrile (reagent grade), dry Acetyl chloride
Protection of Nucleosides for Oligonucleotide Synthesis
2.9.5 Current Protocols in Nucleic Acid Chemistry
Supplement 7
Figure 2.9.2 Preparation of the N-acetylated-5′-O-dimethoxytritylated purine ribonucleosides S.1a and S.1b. Ac, acetyl; DMTr, 4,4′-dimethoxytrityl.
Dichloromethane, precooled Magnesium sulfate Toluene Acetic acid Dimethoxytrityl chloride (DMTr⋅Cl) Triethylamine Silica gel (230 to 400 mesh) Sand Saturated sodium bicarbonate solution in water 250- and 500-mL two-neck flasks Pressure-equilizing dropping funnel Balloon Rubber septum Stir bar 1-L separatory funnels Rotary evaporator with vacuum pump 1-L Erlenmeyer flasks 3-cm glass filter 3-cm-diameter chromatography column Additional reagents and equipment for thin-layer chromatography (TLC, APPENDIX 3D) and column chromatography (APPENDIX 3E)
Preparation of 2′-O[(Triisopropylsilyl)oxy]methylprotected Ribonucleosides
Acetylate adenosine 1. In an ice bath, prepare a suspension of 10 g adenosine (0.037 mol) in 75 mL pyridine in a 500-mL two-neck flask equipped with a pressure-equilizing dropping funnel, nitrogen-filled balloon, rubber septum, and stir bar. 2. Add 47.3 mL trimethylsilyl chloride (0.37 mol) dropwise to the precooled (4°C) suspension.
2.9.6 Supplement 7
Current Protocols in Nucleic Acid Chemistry
3. Remove the cooling bath and stir suspension 14 hr at room temperature. 4. Dilute with 110 mL dry acetonitrile and cool to 4°C using an ice bath. 5. Slowly add 4 mL (0.056 mol) acetyl chloride by syringe and stir 1 hr at 4°C. 6. Pour reaction mixture into a 1-L Erlenmeyer flask. While stirring, add 220 mL precooled (4°C) dichloromethane and 150 mL precooled (4°C) water. Pour into a 1-L separatory funnel. 7. Pour the lower organic layer into another 1-L separatory funnel. Add 200 mL ice-water to the funnel, shake, and pour the lower organic layer into a 1-L Erlenmeyer flask. 8. Add magnesium sulfate until a clear solution is obtained and vacuum filter by means of a 3-cm glass filter. 9. Evaporate the filtrate to dryness using a rotary evaporator. 10. Add 200 mL toluene and evaporate to dryness using a rotary evaporator. Repeat. Evaporation with toluene removes the pyridine.
11. Dissolve residue in 75 mL methanol and add 22.5 mL acetic acid. Keep the mixture at −20°C for 2 days to crystallize product. 12. Collect crystals by vacuum filtration on a 3-cm glass filter, wash them with a small amount of methanol, and dry them at high vacuum. N6-Acetyladenosine (S.4a): 9.1 g (80%) as off-white powder. TLC (CH2Cl2/methanol 7:3) 0.50; m.p. 175°C; [a]D25 -53.7 (c = 0.98, H2O/methanol 1:1); λmax (H2O) 273 (16,700), 209 (21,700); δH (CD3OD/D2O 1:1) 2.36 (s, CH3CO), 3.80 (dd, J = 3.3, 12.6 Hz, H-5′), 3.92 (dd, J = 2.8, 12.6 Hz, H′-5′), 4.23 (br. q, J ≈ 3 Hz, H-4′), 4.39 (dd, J = 3.4, 5.5 Hz, H-3′), 4.77 (t, J = 5.6 Hz, H-2′), 6.10 (d, J = 5.9 Hz, H-1′), 8.58 and 8.65 (2s, H-2 and H-8); δC (CD3OD/D2O 1:1) 24.9 (q), 63.2 (t), 72.4 (d), 75.7 (d), 87.8 (d), 90.8 (d), 125.1 (s), 145.3 (d), 150.9 (s), 153.0 (s), 153.5 (d), 173.4 (s); m/z 310 (100, MH+), 178 (55), 107 (40). Anal. calcd. for C12H15N5O5.(0.25H2O): C, 45.93; H, 4.98; N, 22.31. Found: C, 46.00; H, 4.87; N, 22.23.
Tritylate N6-acetyladenosine 13. In a 250-mL two-neck flask equipped with a rubber septum, nitrogen-filled balloon, and stir bar, dissolve 7.7 g (0.025 mol) N6-acetyladenosine (S.4a) in 80 mL pyridine. 14. Add 10.2 g (0.03 mol) dimethoxytrityl chloride in four 2.55-g portions at 20-min intervals. Keep the solution 1 hr at room temperature. 15. Pour reaction mixture into a 1-L Erlenmeyer flask. While stirring, add 250 mL dichloromethane and 150 mL water. Pour the mixture into a 1-L separatory funnel. Pour the lower organic layer into a 1-L Erlenmeyer flask. 16. Wash the aqueous phase with 100 mL dichloromethane. Allow phases to separate and then pour the organic phase into the same 1-L Erlenmeyer flask. 17. Add magnesium sulfate until a clear solution is obtained, filter by means of a 3-cm glass filter, and evaporate the filtrate to dryness in a rotary evaporator. 18. Add 100 mL toluene and evaporate again to dryness in a rotary evaporator. Repeat. Evaporation with toluene removes the pyridine. Protection of Nucleosides for Oligonucleotide Synthesis
2.9.7 Current Protocols in Nucleic Acid Chemistry
Supplement 7
19. Prepare a slurry of 40 g silica gel in 98:2 (v/v) dichloromethane/triethylamine and pour into a 3-cm-diameter chromatography column. Add ∼2-cm layer of sand on top of the column. 20. Dissolve the crude product in a minimal amount of dichloromethane and place it carefully on top of the column. 21. Start eluting with 250 mL of 98:2 (v/v) dichloromethane/triethylamine and continue eluting with gradually more polar solvent by adding stepwise 1% methanol (250 mL for each step). Collect 100-mL fractions. Always add 2% triethylamine to solvents.
22. Check fractions periodically by TLC (APPENDIX 3D). Pool the fractions that contain the main product [Rf = 0.55 in 9:1 (v/v) dichloromethane/methanol] and evaporate to dryness using a rotary evaporator. 23. Dissolve residue in 150 mL dichloromethane, add 100 mL saturated sodium bicarbonate solution, and stir 5 min. 24. Transfer mixture to a 1-L separatory funnel. Pour the lower organic layer into a 1-L Erlenmeyer flask. 25. Wash the aqueous phase with 100 mL dichloromethane. Allow phases to separate and pour the organic phase into the same 1-L Erlenmeyer flask. Add magnesium sulfate until a clear solution is obtained and vacuum filter by means of a 3-cm glass filter. 26. Evaporate to dryness using a rotary evaporator and dry overnight at high vacuum (<0.05 mbar). N6-Ac-5′-O-DMTr-adenosine (S.1a): 11.4 g (75%) as yellow, solid foam. TLC (CH2Cl2/methanol 9:1) 0.50; [a]D25 −7.4 (c = 0.87, CDCl3); λmax (methanol) 272 (18,700), 234 (22,700); δH (CDCl3) 2.58 (s), 3.31 (dd, J = 3.4, 10.6 Hz, H-5′), 3.44 (dd, J = 3.1, 10.6 Hz, H′-5′), 3.53 (br. s, OH), 3.75 (s, 2OCH3), 4.43 (br. q, J ≈ 2.5 Hz, H-4’), 4.49 (dd, J = 2.5, 5.3 Hz, H-3′), 4.88 (t, J = 5.3 Hz, H-2′), 5.74 (br. s, OH), 6.05 (d, J = 5.6 Hz, H-1′), 6.72-6.75 (4 H, m, DMTr), 7.16-7.29 (9 H, m, DMTr), 8.23 (s, H-2), 8.60 (s, H-8), 8.94 (br. s, NH); δC (CDCl3) 25.7 (q), 55.3 (q), 63.7 (t), 72.7 (d), 76.0 (d), 86.1 (d), 86.8 (s), 90.7 (d), 113.4 (d), 122.2 (s), 127.2 (d), 128.1 (d), 128.2 (d), 130.1 (d), 130.2 (d), 135.7 (s), 141.7 (d), 144.6 (s), 149.6 (s), 150.8 (s), 152.2 (d), 158.9 (s), 170.7 (s); m/z 612 (17, MH+), 303 (100). Anal. calcd. for C33H33N5O7: C, 64.80; H, 5.44; N, 11.45. Found: C, 64.56; H, 5.55; N, 11.18. SUPPORT PROTOCOL 2
PREPARATION OF N2-ACETYL-5′-O-(4,4′-DIMETHOXYTRITYL)GUANOSINE As with the protected adenosine (see Support Protocol 1), the preparation N2-acetyl-5′O-(4,4′-dimethoxytrityl)guanosine (S.1b) is a two-step process. N2-Acetylguanosine (S.4b) is prepared by peracetylation of guanosine with acetic anhydride in dimethylformamide/pyridine, followed by cleavage of the O-bound acetyl groups with NaOH in tetrahydrofuran/methanol/water. Tritylation is performed as in Support Protocol 1, although dimethylformamide is used as a co-solvent in the reaction with the guanosine derivative. Isolation of the intermediate (S.4b) is performed by chromatography and crystallization, and isolation of the product (S.1b) is performed by chromatography.
Preparation of 2′-O[(Triisopropylsilyl)oxy]methylprotected Ribonucleosides
Additional Materials (also see Support Protocol 1) Guanosine Dimethylformamide Acetic anhydride 1 M hydrochloric acid Methanol
2.9.8 Supplement 7
Current Protocols in Nucleic Acid Chemistry
Tetrahydrofuran 10 M NaOH 250-mL two-neck flasks Reflux condenser 135°C bath Distillation apparatus (15 × 2.9–cm Vigreux column equipped with a condenser and water aspirator) 500-mL flask 4A molecular sieves Acetylate guanosine 1. In a 250-mL two-neck flask equipped with a reflux condenser, stir bar, and nitrogenfilled balloon, prepare a mixture of: 5.7 g (0.02 mol) guanosine 25 mL dimethylformamide 25 mL pyridine 25 mL acetic anhydride. 2. Heat 3 hr at reflux in a 135°C bath. 3. Remove all liquids by distillation on a 15 × 2.9–cm Vigreux column equipped with a condenser and water aspirator, at 100°C and 30 mbar, and dissolve the residue in 100 mL dichloromethane. 4. Extract with 100 mL of 1 M hydrochloric acid. 5. Extract the organic phase with 100 mL saturated sodium bicarbonate solution. 6. Dry organic phase with magnesium sulfate until a clear solution is obtained and vacuum filter by means of a 3-cm glass filter. 7. Add 15 g silica gel to the filtrate and evaporate to dryness using a rotary evaporator. 8. Place the powder on top of a chromatography column containing 20 g silica gel in dichloromethane. Elute with 500 mL dichloromethane, and then with 250 mL each (in order) 98:2, 96:4, 94:6, 92:8, and 90:10 (v/v) dichloromethane/methanol. Collect 100-mL fractions. 9. Check fractions periodically by TLC (APPENDIX 3D). Pool the fractions containing the intermediate N2-acetyl-2′,3′, 5′-tri-O-acetylguanosine (Rf = 0.5 in 95:5 [v/v] dichloromethane/methanol) in a 500-mL flask and evaporate them to dryness using a rotary evaporator. 10. Dissolve the residue in 200 mL of 10:8:7 (v/v/v) tetrahydrofuran/methanol/water and add 2 mL of 10 M NaOH. Stir solution 20 min at room temperature. 11. Add 2.5 mL acetic acid and concentrate the solution to 40 mL using a rotary evaporator. Keep the solution 1 day at 4°C to crystallize the product. 12. Collect crystals by vacuum filtration by means of a 3-cm glass filter, wash with a small amount of water, and dry at high vacuum (<0.05 mbar). N2-Acetylguanosine (S.4b): 5.05 g (77%) as an off-white powder. TLC (CH2Cl2/methanol 7:3) 0.30; m.p. 226°C; [a]D25 -19.0 (c = 0.94, H2O); λmax (H2O) 280sh (11,200), 259 (16,300); δH (D2O) 2.28 (s, CH3CO), 3.83 (dd, J = 4.0, 12.6 Hz, H-5′), 3.92 (dd, J = 1.0, 12.6 Hz, H′-5′), 4.21 (m, H-4′), 4.45 (t, J = 4.7 Hz, H-3′), 4.72 (t, J = 4.6 Hz, H-2′), 5.97 (d, J = 4.4 Hz, H-1′), 8.18 (s, H-8). δC (D2O) 26.2 (q), 64.0 (t), 72.9 (d), 76.8 (d), 87.8 (d), 90.9 (d), 122.9 (s), 142.5 (d), 150.6 (s), 152.1 (s), 160.2 (s), 178.3 (s); m/z 326 (100, MH+). Anal. calcd. for C12H15N5O6.(0.33H2O): C, 43.51; H, 4.77; N, 21.14. Found: C, 43.65; H, 5.14; N, 21.16.
Protection of Nucleosides for Oligonucleotide Synthesis
2.9.9 Current Protocols in Nucleic Acid Chemistry
Supplement 7
Tritylate N2-acetylguanosine 13. In a 250-mL two-neck flask equipped with a rubber septum, nitrogen-filled balloon, and stir bar, dissolve 8.3 g (0.026 mol) N2-acetylguanosine (S.4b) in 75 mL pyridine and 25 mL dimethylformamide. 14. Add 5 g of 4A molecular sieves and stir mixture 1 hr at room temperature. 15. Perform tritylation reaction and purification of product as described for adenosine (see Support Protocol 1, steps 14 to 26). N2-Ac-5′-O-DMTr-guanosine (S.1b): 13.4 g (81%) as a yellow powder. TLC (CH2Cl2/methanol 9:1) 0.35; m.p. 174°C; [a]D25 + 15.5 (c = 0.93, methanol); λmax (methanol) 282 sh (13,700) 274 (14,600), 260 sh (18,300), 251 sh (19,600), 235 (25,800); δH (CDCl3/CD3OD 3:1) 2.02 (s, CH3CO), 3.20 (dd, J = 4.3, 10.5 Hz, H-5′), 3.26 (dd, J = 2.8, 10.5 Hz, H-5′), 3.62 (s, 2 OCH3), 3.69 (br. s, H-4′), 4.22 (t, J = 4.9 Hz, H-3′), 4.45 (t, J = 4.9 Hz, H-2′), 5.76 (d, J = 4.8 Hz, H-1′), 6.64 (6 H, m, DMTr), 7.03-7.26 (9 H, m, DMTr), 7.74 (s, H-8); δC (CDCl3/CD3OD 3:1) 23.4 (q), 55.1 (q), 63.6 (t), 71.0 (d), 74.8 (d), 84.2 (d), 86.6 (d), 88.6 (s), 113.2 (d), 120.8 (s), 127.0 (d), 127.9 (d), 128.2 (d), 130.1 (d), 135.8 (s), 137.8 (d), 144.6 (s), 147.9 (s), 148.8 (s), 156.2 (s), 158.7 (s), 173.3 (s); m/z 628 (74, MH+), 303 (100). Anal. calcd. for C32H35N4O8: C, 63.15; H, 5.30; N, 11.16; Found: C, 63.08; H, 5.35; N, 10.91. SUPPORT PROTOCOL 3
PREPARATION OF [(TRIISOPROPYLSILYL)OXY]METHYL CHLORIDE This protocol describes the two-step synthesis of the reagent TOM⋅Cl, required for introduction of the 2′-O-TOM group into 5′-O-dimethoxytritylated, nucleobase-protected ribonucleosides. The two reactions are illustrated in Figure 2.9.3. In the first reaction, the adduct formed from formaldehyde and ethanethiol in situ according to a procedure by Gundersen et al. (1989) is converted with triisopropylsilyl chloride (TIPS⋅Cl)/imidazole into the corresponding silylated S,O-acetal (S.5). Upon treatment of this intermediate with sulfuryl chloride, TOM⋅Cl (S.6) is formed. The reaction can easily be performed on a larger scale than that presented; however, the authors recommend that it be carried out first on the scale described here. Both the intermediate (S,O-acetal) and TOM⋅Cl are isolated by distillation. After distillation, TOM⋅Cl can be stored for at least 1 year in pure form at −20°C. Materials Paraformaldehyde Ethanethiol Nitrogen or argon gas 10 M aqueous NaOH
Preparation of 2′-O[(Triisopropylsilyl)oxy]methylprotected Ribonucleosides
Figure 2.9.3 Preparation of [(triisopropylsilyl)oxy]methyl chloride (TOM⋅Cl; S.6). Et, ethyl; i-Pr, isopropyl.
2.9.10 Supplement 7
Current Protocols in Nucleic Acid Chemistry
Dichloromethane (CH2Cl2; reagent grade) Imidazole Triisopropylsilyl chloride (TIPS⋅Cl) Hexane (technical grade) 10% (w/v) NaH2PO4 Anhydrous magnesium sulfate (MgSO4) Sulfuryl chloride (SO2Cl2) 250- and 1000-mL one-neck round-bottom flasks Balloon Oil bath, 40°C 1-L separatory funnel 3-cm glass filter Rotary evaporator with a vacuum pump 10-cm 14.5-mm Vigreux distillation column Vacuum distillation equipment (head, thermometer, and so on) with high-vacuum pump 250-mL two-neck flask Rubber septum Syringes Prepare [(triisopropylsilyl)oxy]methylthioethyl ether 1. In a well-ventilated fume hood, mix 3.06 g (0.102 mol) paraformaldehyde with 7.5 mL (0.102 mol) ethanethiol in a 250-mL one-neck round-bottom flask equipped with a magnetic stir bar and a balloon filled with nitrogen or argon gas. 2. Place the suspension in an ice bath and stir until the temperature reaches 4°C (∼5 min). 3. While the solution stirs, remove the gas-filled balloon, add one drop 10 M aqueous NaOH and then quickly replace the balloon. Let the reaction mixture warm up slowly by removing it from the ice bath. At a temperature between 15° and 23°C, a sudden exothermic reaction occurs, forming a colorless solution. The violent reaction results from depolymerization of the paraformaldehyde. If it does not occur spontaneously, heat the reaction mixture gently until the depolymerization reaction starts. Do not heat above 40°C.
4. Place the flask in a preheated oil bath and stir 1 hr at 40°C. 5. Cool the reaction mixture to room temperature and dilute with 100 mL dichloromethane. 6. Add 13.9 g (0.204 mol) solid imidazole, quickly followed by 20.5 mL (0.097 mol) triisopropylsilyl chloride. Replace the balloon. The imidazole does not need to be completely dissolved before addition of the triisopropylsilyl chloride. It is very important that the triisopropylsilyl chloride be added quickly.
7. Stir the resulting colorless suspension overnight at room temperature. 8. Pour the reaction mixture into a 1-L flask containing a stirred mixture of 250 mL hexane and 150 mL of 10% NaH2PO4. 9. Separate the two phases in a 1-L separatory funnel. 10. Re-extract the aqueous phase with 200 mL hexane and combine the organic extracts.
Protection of Nucleosides for Oligonucleotide Synthesis
2.9.11 Current Protocols in Nucleic Acid Chemistry
Supplement 7
11. Dry the extract over anhydrous magnesium sulfate, vacuum filter by means of a 3-cm glass filter, and evaporate the solvents in a rotary evaporator with a vacuum pump. 12. Distill the residue in vacuo by means of a 10-cm, 14.5-mm Vigreux column using a vacuum that is <0.1 mbar and a heating bath temperature that does not exceed 100°C. Collect the main fraction, which typically distills at 70°C (0.05 mbar). 13. Check the purity of the product by 1H-NMR. If the purity of the product is <95%, repeat the distillation carefully. Typically, the product is obtained in yields between 80% and 90%. {[(Triisopropylsilyl)oxy]methyl}thioethyl ether (S.5): 20.6 g (85%) as a colorless liquid, b.p. 72°C (0.05 torr); δH (CDCl3) 1.08-1.18 (21 H, m, i-Pr3Si), 1.31 (t, J = 7.4 Hz, CH3), 2.71 (q, J = 7.4 Hz, CH2), 4.88 (s, CH2); δC (CDCl3) 12.0 (CH), 15.0 (CH3), 17.9 (CH3), 24.7 (CH2), 66.1 (CH2).
Prepare [(triisopropylsilyl)oxy]methyl chloride 14. In a 250-mL two-neck round-bottom flask equipped with a rubber septum and a magnetic stir bar, and connected to a continuous nitrogen flow, dissolve 17.6 g (0.07 mol) {[(triisopropylsilyl)oxy]methyl}thioethyl ether (S.5) in 50 mL dry dichloromethane. Place the colorless solution in an ice bath and stir for 10 min. CAUTION: This reaction must be carried out in a well-ventilated hood, since one byproduct of this reaction (ethanesulfenyl chloride) is toxic and an irritant.
15. Over a 15-min period, add 5.75 mL (0.07 mol) sulfuryl chloride by syringe, remove the ice bath, and continue stirring. The solution turns gradually more yellow. During the reaction, sulfur dioxide and ethanesulfenyl chloride are produced.
16. Stir 1 hr at room temperature. 17. Connect the flask to a dry rotary evaporator and start to evaporate slowly and carefully. The yellow ethanesulfenyl chloride distills together with the dichloromethane and the residue turns colorless again.
18. Remove the solvent completely. The reaction mixture contains the partially dissolved gaseous compounds sulfur dioxide and ethanesulfenyl chloride; apply initially only a gentle vacuum.
19. Distill the residue in vacuo by means of a 10-cm, 14.5-mm Vigreux column. Use a vacuum that is <0.05 mbar and a heating bath temperature that does not exceed 80°C. Collect the main fraction, which typically distills at 40°C (0.01 mbar). 20. Check the purity of the product by 1H-NMR. If the purity of the product is <85%, repeat the distillation carefully. Typically, the product is obtained in yields between 90% and 95%. [(Triisopropylsilyl)oxy]methyl chloride (S.6): 14.8 g (95 %) as a colorless liquid, b.p. 40°C (0.01 torr); δH (CDCl3) 1.08-1.10 (21 H, m, i-Pr3Si), 5.66 (s, CH2); δC (CDCl3) 11.8 (CH), 17.7 (CH3), 76.6 (CH2).
Preparation of 2′-O[(Triisopropylsilyl)oxy]methylprotected Ribonucleosides
2.9.12 Supplement 7
Current Protocols in Nucleic Acid Chemistry
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. Use dry solvents (reagent grade) for all reactions. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Anisaldehyde reagent In a clean container, mix 10 mL anisaldehyde with 180 mL ethanol. Add slowly, while stirring, 10 mL concentrated sulfuric acid, followed by 2 mL acetic acid. Store in the dark for up to 3 months at 25°C. Contamination with acetone must be avoided. For staining, TLC plates are dipped into this mixture and heated with a heat gun until dark spots, corresponding to the reaction products, appear on a pink to red background.
COMMENTARY Background Information The chemical synthesis of DNA sequences b ased on 5′-O-dimethoxytritylated phosphoramidite building blocks is straightforward and very reliable. This extremely powerful methodology can, in principle, also be applied to the synthesis of RNA oligonucleotides, which are structurally very similar. However, compared to DNA, each RNA nucleoside contains an additional 2′-OH group that must be protected during chemical assembly of the nucleotide chain. Since RNA products are base labile, the removal of these supplementary protecting groups is carried out separately, after removing all other protecting groups under basic, nucleophilic conditions (Beaucage and Caruthers, 1996). Among the large number of 2′-O-protecting groups investigated thus far, the fluoride-labile tert-butyldimethylsilyl (TBDMS) group (Ogilvie et al., 1974; UNIT 3.6) has the widest application. However, several factors have limited the length of routinely synthesized RNA sequences using this protecting group. In particular, relatively low coupling yields of ≤98% are obtained with these RNA phosphoramidites despite rather long coupling times of ~15 min, com pared to >99% with DNA phosphoramidites using coupling times of ~2 min. A very attractive alternative was the photolabile 2-nitrobenzyloxymethyl (NBOM) group (Schwartz et al., 1992; UNIT 3.7), which leads to coupling yields of ~99% under DNA coupling conditions. The superior coupling behavior of these building blocks is presumably the result of the minimal steric demand of the NBOM group, achieved by connecting the photocleavable 2-nitrobenzyl protecting group via a sterically nondemanding formaldehyde acetal linker to the 2′-O position of the ribonucleosides (Schwartz et al., 1992; Pitsch, 1997). The 2′-O-TOM protecting group represents an
advantageous combination of the TBDMS and NBOM protecting groups. First, it is completely stable towards all reaction conditions applied during the assembly and the first (basic) deprotection step. Second, it does not interfere with the coupling reaction and leads to very good coupling yields under DNA coupling conditions (99% with a coupling time of 2.5 min). Third, its final cleavage occurs quantitatively without concomitant destruction of the RNA product sequences (UNIT 3.8). The introduction of the TOM group into partially protected ribonucleosides is carried out according to a method originally developed for the introduction of the related NBOM group (Pitsch, 1997, Wu and Pitsch, 1998; Pitsch et al., 1999). The separation of the two regioisomeric TOM-substituted ribonucleosides, formed during the dibutyltin-mediated alkylation reaction, is straightforward and leads to uniform products. In contrast to the TBDMS protecting group, the TOM group does not migrate from the 2′-O to the 3′-O position, even under strongly basic conditions. These factors allow the routine preparation of very pure phosphoramidites, which result in the exclusive formation of 3′→5′ phosphodiester moieties. Furthermore, due to the stability of the 2′-O-TOM group, a variety of nucleobase and sugar manipulations can be carried out after its introduction (Wu and Pitsch, 1998; Stutz et al., 2000). Although the Basic Protocol in this unit uses N-acetylated-5′-O-dimethoxytritylated starting nucleosides, the authors have also introduced the TOM group into a large number of different nucleosides, including nucleosides with different acyl-type nucleobase-protecting groups, unnatural backbones, or noncanonical nucleobases. To date, the reaction has failed only with 5′-O-dimethoxytritylated inosine, which consequently was prepared from 2′-OTOM-substituted adenosine by deamination.
Protection of Nucleosides for Oligonucleotide Synthesis
2.9.13 Current Protocols in Nucleic Acid Chemistry
Supplement 7
Alkylation of dialkyl-stannylated nucleosides with TOM⋅Cl occurred under a variety of reaction conditions. Performing the reaction at 80°C and using 1,2-dichloroethane as solvent favors the formation of the 2′-O-alkylated products, whereas formation of the 3′-O-alkylated products can predominate at lower temperatures and with other solvents (tetrahydrofuran, acetonitrile, benzene). Slightly higher yields (5% to 10%) of 2′-O-TOM-protected nucleosides can sometimes be obtained by forming the stannylated nucleosides with tert-Bu2SnCl2 instead of n-Bu2SnCl2. However, since tertBu2SnCl2 is very expensive, the authors use it only for the alkylation of precious nucleosides.
Critical Parameters
Preparation of 2′-O[(Triisopropylsilyl)oxy]methylprotected Ribonucleosides
In general, there are two important considerations for a successful preparation of 2′-OTOM-protected ribonucleosides. The reaction always results in the formation of a mixture of 2′-O- and 3′-O-alkylated products. Fortunately, the chromatographic behavior (on silica gel) of these two regioisomers differs substantially (∆Rf ≥ 0.1). In all experiments to date, the more quickly migrating isomer has always been the desired 2′-O-alkylated product, which by careful chromatography could be obtained in pure form. The unambiguous identification of the products is conveniently carried out by 1HNMR spectroscopy. Irradiation experiments provide the identification of the signals from the sugar-bound protons and the remaining hydroxyl protons. By irradiation (or D2O-exchange) of the latter, their connection to either C3′ or C2′ can be established, and therefore the alkylation sites can be identified (Pitsch, 1997). Under the authors’ preferred reaction conditions, employing only 1.1 to 1.3 eq of TOM⋅Cl, a significant portion of starting material can be recovered. It is recommended that a larger excess of TOM⋅Cl not be used, since this generally results not in better product yields, but only in the formation of nucleobase-alkylated byproducts, which are quite difficult to remove from the desired product. As a rule, the addition of extra equivalents of TOM⋅Cl should be stopped when more than ~5% of nucleobase-alkylated byproducts (eluting substantially faster then the 2′-O-alkylated product) are detected by TLC analysis. The unreacted starting materials can be isolated by elution with 95:5 (v/v) dichloromethane/methanol. If required, the 3′-O-alkylated regioisomers can be transformed into the starting materials by short treatment (e.g., 30 min) with 0.25 M Bu4NF/THF or 0.25 M, tetraethylammonium fluoride/acetonitrile.
Anticipated Results Introduction of the TOM group into nucleosides depends on the structure of the starting material and on the reaction conditions. In the authors’ experience, yields may range from 25% and 60%. The successful separation of the two diastereoisomers formed can require careful optimization of the chromatography conditions. However, the unique properties of TOMprotected phosphoramidites for RNA synthesis justify the efforts required for their introduction.
Time Considerations Each of the intermediates and products described in this unit can be prepared and isolated within 1 or 2 days.
Literature Cited Beaucage, S.L. and Caruthers, M.H. 1996. The chemical synthesis of DNA/RNA. In Bioorganic Chemistry: Nucleic Acids (S.M. Hecht, ed.) pp. 36-74. Oxford University Press, Oxford. Gundersen, L.L., Benneche, T., and Undheim, K.A. 1989. Chloromethoxysilanes as protecting reagents for sterically hindered alcohols. Acta Chem. Scand. 43:706-709. Ogilvie, K.K., Sadana, K.L., Thompson, E.A., Quilliam, M.A., and Westmore, J.B. 1974. The use of silyl protecting groups in protecting the hydroxyl functions of ribonucleosides. Tetrahedron Lett. 15:2861-2863. Pitsch, S. 1997. An efficient synthesis of enantiomeric oligoribonucleotides from D-glucose. Helv. Chim. Acta 80:2286-2314. Pitsch, S., Weiss, P.A., and Jenny, L. Nov. 1999. Ribonucleoside-derivative and method for preparing the same. US Patent 5,986,084. Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., deBear, J.S., and Gough, G.R. 1992. Rapid synthesis of oligoribonucleotides using 2′-O-(ortho-nitrobenzyloxymethyl)-protected monomers. Bioorg. Med. Chem. Lett. 2:1019-1024. Stutz, A., Höbartner, C., and Pitsch, S. 2000. Synthesis of 3′-O-aminoacylated RNA-fragments with novel, fluoride-labile base-protecting groups. Helv. Chim. Acta 83:2477-2503. Wu, X. and Pitsch, S. 1998. Synthesis and pairing properties of oligoribonucleotide analogues containing a metal-binding site attached to β-D-allofuranosyl cytosine. Nucl. Acids Res. 26:43154323.
Contributed by Stefan Pitsch Institut de Chimie Organique, EPFL Lausanne, Switzerland Patrick A. Weiss Xeragon AG Zürich, Switzerland
2.9.14 Supplement 7
Current Protocols in Nucleic Acid Chemistry
Preparation of 5′-Silyl-2′-Orthoester Ribonucleosides for Use in Oligoribonucleotide Synthesis
UNIT 2.10
This unit describes the synthesis of chemically protected ribonucleoside monomers that can be used to prepare oligoribonucleotides using either manual or automated synthesis approaches. The protection strategy employs two orthogonal protecting groups—a fluoride ion–labile silyl ether on the 5′-hydroxyl and an acid-labile orthoester on the 2′-hydroxyl—as described by Scaringe et al. (1998). The structures of the protected phosphoramidite derivatives of the four standard ribonucleosides (rA, rC, rG, and U) are shown in Figure 2.10.1. These reagents are used with appropriate modifications of standard solid-phase synthesis methods (Matteucci and Caruthers, 1981) to efficiently and rapidly assemble oligoribonucleotides.
O O O Me3Si O Si
N
O
Me3Si O MeO
O
NH O
P
O
Me3Si O Si
O
N O
O O
N
MeO
OAc
N
O
Me3Si O O
CH3
HN
P
O
O
O
OAc
O
N
OAc
O
OAc
cytidine
uridine
O HN N
O Me3Si O Si
O
MeO
O
Me3Si O Si
N
Me3Si P N
O
O
O O
N
O
N
N
O
Me3Si O
Ph
OAc OAc
adenosine
O O
O MeO
P N
O
NH
N
O
N
O O
O
N H
OAc OAc
guanosine
Figure 2.10.1 Protected ribonucleoside phosphoramidites for 5′-O-silyl-2′-O-orthoester RNA synthesis.
Protection of Nucleosides for Oligonucleotide Synthesis
Contributed by Stephen A. Scaringe, David Kitchen, Robert J. Kaiser, and William S. Marshall
2.10.1
Current Protocols in Nucleic Acid Chemistry (2004) 2.10.1-2.10.16 Copyright © 2004 by John Wiley & Sons, Inc.
Supplement 16
The choice of protecting groups described above is predicated on the following requirements. (1) The 2′-hydroxyl must be protected with a species that is stable to the conditions of chain assembly. (2) The synthesis procedure must protect against product cleavage from the solid support and against removal of the protecting groups on the nucleobases. (3) The protecting group must be easily removed at the end of the procedure under mildly acidic aqueous conditions. (4) Given items 1 to 3, the acid-labile 4,4′-dimethoxytrityl (DMTr) group that is traditionally used to protect the 5′-hydroxyl in deoxyribonucleotide synthesis must be replaced with a moiety that does not require acidic or basic conditions for its rapid removal. The 5′-O-silyl ether and 2′-O-orthoester groups fit these criteria and have been structurally optimized to provide crude oligoribonucleotides of excellent purity and biological activity. Using commercial synthesizers, the 5′-O-silyl-2′-O-orthoester chemistry enables routine synthesis of oligoribonucleotides up to 80 bases in length, regardless of sequence or secondary structure. Synthesis of the required monomers begins from the standard base-protected ribonucleosides rABz (benzoyl), rCAc (acetyl), rGi-Bu (isobutyryl), and U (unprotected). The procedure for synthesizing the protected uridine phosphoramidite is presented in detail (see Basic Protocol). The syntheses of the other three phosphoramidites—adenosine, guanosine, and cytidine—are essentially identical. Modifications to the purification procedures, as well as compound characterization data, are detailed below (see Alternate Protocols 1 to 3). BASIC PROTOCOL
SYNTHESIS OF 2′-ACE-PROTECTED URIDINE RIBONUCLEOSIDE PHOSPHORAMIDITE This protocol describes the synthesis of 5′-O-benzhydroxy-bis(trimethylsilyloxy)silyl-2′O-bis(acetoxyethyloxy)methyl-3′-O-(N,N-diisopropylamino)methoxyphosphinyl uridine. The general strategy for preparation of the 5′-O-silyl-2′-O-orthoester-ribonucleoside-3′-O-phosphoramidites consists of five steps: (1) simultaneous temporary protection of the 5′- and 3′-hydroxyls, (2) conversion of the 2′-hydroxyl to the 2′-O-orthoester using tris(acetoxyethyl)orthoformate (ACE orthoformate), (3) removal of the temporary 5′- and 3′-hydroxyl-protecting groups, (4) regioselective introduction of the 5′-O-silyl-ether protecting group using benzhydroxy-bis(trimethylsilyloxy)chlorosilane (BzH-Cl), and (5) preparation of the 3′-O-phosphoramidite. These steps are illustrated for the uridine derivatives in Figures 2.10.2 to 2.10.6. NOTE: All reactions must be carried out using proper ventilation (fume hood) and personal protective wear (e.g., gloves, eye protection). All reagents without specified vendors should be of the highest purity and dryness possible. Glassware for performing the synthetic transformations should be thoroughly cleaned and dried in an oven at 110°C for ≥30 min before use.
Preparation of 5′-Silyl-2′Orthoester Ribonucleosides
Materials Uridine (S.1; 244 g/mol; Monomer Sciences) Pyridine 1,3-Dichloro-1,1,3,3-tetraisopropyldisiloxane (TIPS-Cl2; 315 g/mol; d = 0.986 g/mL; Monomer Sciences) Toluene Dichloromethane 5% and 8% (w/v) aqueous sodium bicarbonate (NaHCO3) Saturated aqueous sodium chloride (NaCl) Anhydrous sodium sulfate (Na2SO4)
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Current Protocols in Nucleic Acid Chemistry
Silica gel 60 (for column chromatography) Hexane Ethyl acetate (EtOAc) Silica gel thin-layer chromatography (TLC) plates Methanol (MeOH) Acetonitrile (MeCN) Tris(2-acetoxyethyl) orthoformate (322 g/mol; Dharmacon Inc.) Pyridinium p-toluenesulfonate (251 g/mol) Triethylamine (TEA) 4-tert-Butyldimethylsiloxy-3-penten-2-one (238 mL/mol; Silar) N,N,N′,N′-Tetramethylethylene diamine (TEMED; 150 mL/mol) 48% (w/v) aqueous hydrofluoric acid (HF; 32 mL/mol) Diisopropylamine (140 mL/mol) Benzhydroxy-bis(trimethylsilyloxy)chlorosilane (BzH-Cl; 425 g/mol; Dharmacon Inc.) Acetone Bis(N,N-diisopropylamino)methoxyphosphine (262 g/mol; Monomer Sciences) 0.45 M 1H-tetrazole in MeCN (AIC) Ethanol (EtOH) Rotary evaporator 1-L separatory funnel Glass-fritted Buchner funnel (coarse porosity) and side-arm Erlenmeyer flask High-vacuum pump Water aspirator 50 × 600–mm chromatography column Additional reagents and equipment for flash chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) NOTE: For solvent evaporation on a rotary evaporator, the required vacuum source will depend on the boiling point of the solvent involved. For dichloromethane and hexane, a water aspirator or diaphragm pump will suffice. For all other solvents, a high-vacuum oil pump is needed. NOTE: The efficiency of reactions as well as the results of column chromatography at each step are assessed by TLC on silica gel plates. It is suggested that the TLC plates be spotted in three horizontal locations at the origin of the plate as follows: (1) starting material alone, (2) co-spot of starting material plus crude reaction mixture or product fraction, and (3) crude reaction mixture or product fraction alone. In this way, the conversion of starting material to product, or the purity of the product fraction, can be conveniently monitored. Protect 3′- and 5′-OH (S.2) 1. Co-evaporate 40 mmol (9.76 g) uridine (S.1; Fig. 2.10.1) with 50 mL pyridine on a rotary evaporator equipped with a vacuum pump. Dissolve residue in 50 mL fresh pyridine and chill reaction mixture in an ice bath. 2. Add 13.86 mL (44 mmol; 1.1 equiv per equiv uridine) TIPS-Cl2 dropwise over 60 min to the stirred, cold uridine solution. Remove ice bath and stir solution at room temperature for an additional 30 min. Add 2 mL water to quench the reaction. 3. Remove pyridine on the rotary evaporator, and then co-evaporate the residue once with 100 mL toluene to remove as much pyridine as possible.
Protection of Nucleosides for Oligonucleotide Synthesis
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4. Add 300 mL dichloromethane to the residue and swirl to dissolve. Transfer the solution to a 1-L separatory funnel and wash once with 300 mL of 5% aqueous NaHCO3. 5. Separate the layers and wash the aqueous phase with an additional 150 mL dichloromethane. 6. Combine the dichloromethane extracts and wash once with 200 mL saturated aqueous NaCl. 7. Wash the NaCl solution with 50 mL dichloromethane. 8. Combine the dichloromethane extracts and dry over 50 g anhydrous Na2SO4. Remove the solid by vacuum filtration using a glass-fritted Buchner funnel (coarse porosity) and a side-arm Erlenmeyer flask. 9. Remove the dichloromethane on the rotary evaporator, and then co-evaporate the residue once with 100 mL toluene to remove any remaining pyridine. Dry the crude product under high vacuum (with a high-vacuum pump) for 1 hr. 10. Prepare a flash chromatography (APPENDIX 3E) column using 250 g silica gel 60 in a 50 × 600–mm chromatography column and a 50:50 (v/v) hexane/EtOAc solvent system. 11. Dissolve the dry product in 50 mL of 50:50 hexane/EtOAc and load onto the column. Elute with the same solvent system (typically 2 L). Collect 20-mL fractions and monitor the results of the separation by TLC (APPENDIX 3D) on silica gel plates in 6% (v/v) MeOH/CH2Cl2. Visualize TLC separation using UV light. The product has an Rf of 0.5 under these conditions.
12. Pool the fractions containing pure product and evaporate to dryness on the rotary evaporator to obtain a white crystalline powder. The molecular weight of 3′,5′-O-TIPS-uridine (S.2; Fig. 2.10.2) is 486 g/mol. The yield is 15.57 g (80%). 1H NMR (CDCl3): δ 10.41 (b, 1H), 7.81 (d, J = 8.4 Hz, 1H), 5.72–5.69 (m, 1H), 5.66 (d, J = 8.4 Hz, 1H), 4.38 (b, 1H), 4.23–4.18 (m, 4H), 3.96 (d, J = 12.8 Hz, 1H), 1.05–0.90 (m, 28H). 13C NMR (CDCl3): δ 164.0, 150.7, 139.9, 102.1, 91.1, 81.8, 75.1, 68.4, 60.0, 17.6, 17.5, 17.4, 17.3, 17.1, 17.02, 17.01, 16.9, 13.4, 13.1, 13.0, 12.5. Exact mass calculated for C21H38N2O7Si2: 486.2; found by ESMS (M + H): 487.
O
O NH
HO
O HO
O
O TIPS-Cl2
OH
1 Preparation of 5′-Silyl-2′Orthoester Ribonucleosides
N
NH
pyridine
O
Si O Si
O
N
O
OH
2
Figure 2.10.2 3′,5′-Protection of the nucleoside. Synthesis of 3′,5′-O-(tetraisopropyldisiloxane1,3-diyl)uridine from uridine. TIPS-Cl2, 1,3-dichloro-1,1,3,3-tetraisopropyldisiloxane.
2.10.4 Supplement 16
Current Protocols in Nucleic Acid Chemistry
O
O NH
O
O
Si O Si
O
N
NH
O CH(OCH 2CH2OAc) 3, CH2Cl2
OH
pyridinium p -toluenesulfonate
O O Si
N
O
Si O
O
O O
2
O
OAc OAc
3
Figure 2.10.3 2′-ACE protection. Synthesis of 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-2′-O-bis(acetoxyethyloxy)methyl uridine from 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)uridine.
Perform 2′-O-ACE protection (S.3) 13. Dry the product S.2 by co-evaporating twice with 100-mL portions of MeCN. 14. Add, in order, tris(2-acetoxyethyl) orthoformate (2.3 equiv per equiv S.2), dichloromethane (2 mL per mmol S.2), and pyridinium p-toluenesulfonate (0.2 equiv per equiv S.2). Stir reaction at room temperature. 15. To check the progress of the reaction by TLC, dilute a 40-µL aliquot of the reaction solution with 40 µL TEA and 200 µL CH2Cl2. Analyze 1 to 5 µL of this solution using a solvent system of 5% (v/v) MeOH in CH2Cl2. The product, 3′,5′-O-TIPS-2′-O-ACE-uridine (S.3; Fig. 2.10.3), typically migrates with a slightly faster Rf than the starting material.
16. After TLC analysis shows that the reaction is progressing well (∼60 min), add 4-tert-butyldimethylsiloxy-3-penten-2-one (1.8 equiv per equiv S.2) and stir the reaction for an additional 36 to 48 hr at room temperature. If necessary, the reaction can be accelerated by gently refluxing at 38° to 40°C. The solution should appear clear, with a slightly yellow or orange color. If the reaction still appears to be sluggish or stagnant, an additional aliquot of pyridinium p-toluenesulfonate (up to 1.0 equiv total) can be added. The 4-tert-butyldimethylsiloxy-3-penten-2-one is added to drive the reaction to completion. The 2′-ACE reaction is an equilibrium exchange of the 2′-hydroxyl with one of the three alcohols in the orthoester reagent. 4-tert-Butyldimethylsiloxy-3-penten-2-one reacts selectively with the primary hydroxyl of the liberated 2-acetoxy-ethanol byproduct, rendering it unable to react further, and thereby forcing the equilibrium to the 2′-ACE product.
17. When TLC indicates that the reaction is ≥95% complete, add TEMED (0.5 equiv per equiv S.2) to neutralize the reaction mixture. 4-tert-Butyldimethylsiloxy-3-penten-2-one is UV absorbing and volatile. It appears on the TLC plate as a dark spot or as a pair of spots (two isomers) migrating at the solvent front. Upon standing, the spots will slowly disappear. If additional pyridinium p-toluenesulfonate was added to accelerate the reaction, additional TEMED (up to 2.5 equiv) must be added to compensate. Protection of Nucleosides for Oligonucleotide Synthesis
2.10.5 Current Protocols in Nucleic Acid Chemistry
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18. Prepare a flash chromatography column with 250 g silica gel 60 using a solvent mix of 75:25 (v/v) hexane/EtOAc containing 0.5% (v/v) TEMED. Dilute the reaction solution with 2 vol (90 mL) hexane and load this mixture directly onto the column. Only TEMED should be used as the base. TEA or another tertiary amine should not be used. TEMED-HF is used to deprotect the 5′- and 3′-hydroxyls in the next step. If TEA is present in that step, the HF salt of TEA will be generated. The TEA-HF salt migrates on silica gel and is difficult or impossible to separate from the 2′-O-ACE nucleoside product. This problem does not occur with the more polar TEMED-HF salt.
19. Elute the product using 49:50:0.5:0.5 (v/v/v/v) hexane/EtOAc/MeOH/TEMED (typically 2 L). Collect 20-mL fractions and monitor by TLC. It is very important to remove the several small impurities that elute just ahead of the desired product. If present, they severely complicate subsequent purifications. Therefore, when the product begins to elute, collect smaller fractions and analyze carefully by TLC. Because the excess orthoester reagent and any remaining starting material often co-elute with the product, the product cannot be fully characterized at this point. Carry it through to the next reaction, in which the TIPS protecting group will be removed. After the next reaction, all impurities can be easily removed by column chromatography, and the pure product can be readily characterized.
Remove 3′,5′-TIPS protection (S.4) 20. To the semi-pure S.3, add 2 mL MeCN per mmol of starting protected uridine in step 13).
(S.2
21. Prepare TEMED-HF reagent in a separate flask as follows. Place 2 mL MeCN per mmol S.2 in a separate vessel on ice. Add 5 equiv TEMED (per equiv S.2). Then slowly add (over 5 min) 48% (w/v) aqueous HF (3.5 equiv per equiv S.2). Stir on ice for 5 min. 22. Slowly add TEMED-HF solution to the nucleoside solution over 2 min, with stirring. Stir reaction at room temperature for ∼4 hr. Monitor the progress of the reaction by TLC on silica gel using 8% (v/v) MeOH in CH2Cl2 until it is complete. The desired product moves considerably more slowly on the TLC plate than does the starting material in this solvent systems.
23. Remove MeCN from the reaction on the rotary evaporator. When most of the solvent is removed, add 100 mL dichloromethane, 20 mL hexane, and 2 mL TEMED. Mix well. 24. Slowly load this solution directly onto a 50 × 600–mm flash chromatography column containing 250 g silica gel poured in 15:85 (v/v) hexane/EtOAc containing 0.5% (v/v) TEMED. The solution should be loaded slowly onto the column using gravity flow, as heat tends to be generated and may disrupt the silica gel bed.
25. Elute the product using 94:6 (v/v) EtOAc/MeOH containing 0.5% (v/v) TEMED (typically 2 L). Collect 20-mL fractions.
Preparation of 5′-Silyl-2′Orthoester Ribonucleosides
The molecular weight of 2′-O-ACE-uridine (S.4; Fig. 2.10.4) is 462 g/mol. The yield is 11.83 g (65% to 80%) overall (steps 13 to 25 combined). 1H NMR (CDCl3): δ 7.79 (d, J = 8.4 Hz, 1H), 5.80 (d, J = 4.0 Hz, 1H), 5.66 (d, J = 8.0 Hz, 1H), 5.42 (s, 1H), 4.45–4.43 (m, 1H), 4.26–4.23 (m, 1H), 4.15–4.10 (m, 4H), 4.02–4.00 (m, 1H), 3.87–3.82 (m, 1H), 3.74–3.66 (m, 5H), 1.98 (s, 3H), 1.97 (s, 3H). 13C NMR (CDCl3): δ 171.25, 171.22, 164.2, 150.7, 141.8, 112.4, 102.2, 89.2, 84.9, 76.2, 69.4, 63.1, 63.0, 62.97, 62.8, 61.2, 30.9, 20.84, 20.82. Exact mass calculated for C18H26N2O12: 462.2; found by ESMS (M + H): 463.
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Current Protocols in Nucleic Acid Chemistry
O
O
NH
NH O Si O Si
N
O O
HO
O TEMED-HF
O
O O
3
MeCN OAc
N
O HO
O
O
O O
OAc
OAc OAc
4
Figure 2.10.4 Removal of 3′,5′-protection. Synthesis of 2′-O-bis(acetoxyethyloxy)methyl uridine from 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-2′-O-bis(acetoxyethyloxy)methyl uridine. TEMED, N,N,N′,N′-tetramethylethylenediamine.
Perform 5′-O-silylation (S.5) 26. Dissolve S.4 in 5 mL CH2Cl2 (per mmol S.4) and add diisopropylamine (1 equiv per equiv S.4). Cool solution to 0°C with an ice bath. 27. In a separate flask, dissolve 1.5 equiv BzH-Cl (per equiv S.4) in CH2Cl2 (2 mL/g BzH-Cl). 28. Add 1 equiv diisopropylamine (per equiv BzH-Cl) to the silyl chloride solution (step 27) slowly over 1 min. Shake resulting solution for 5 min and ensure that it remains clear. If the solution does not clear after several minutes, the mixture is not sufficiently anhydrous, and the solution should be made again using fresh, dry dichloromethane and N,N-diisopropylamine. These materials can be conveniently dried by storing them over 4A molecular sieves for at least 24 hr.
29. Slowly add the diisopropylamine-activated silylating reagent (step 28) to the nucleoside solution (step 26) while stirring at 0°C. Begin by adding 0.5 equiv aliquots of the silylating reagent, and then taper to smaller aliquots as the reaction proceeds to completion. Add each aliquot over a period of 10 to 15 min. Monitor the reaction by TLC on silica gel after each addition using 5% (v/v) MeOH/CH2Cl2. Continue to add the silylating solution until the start material disappears, and then add slightly more to ensure that any 3′-O-silylated nucleoside is converted to the 3′,5′-O-disilylated nucleoside. Keep the total reaction time <3 hr. The reaction will start out clear, but will turn cloudy as the silylating solution is added. The silylating reagent appears to react slightly with the nucleobases on uridine and guanosine, requiring >1.0 equiv to 5′-protect all of the starting material. The silylating reagent reacts preferentially with the primary 5′-hydroxyl but will also react with the 3′-hydroxyl. The 3′-O-silylated nucleoside contaminant chromatographs immediately ahead of the desired 5′-O-silylated product, and is difficult to separate. However, the 3′,5′-bis-silylated product runs very quickly in the TLC solvent given, near the solvent front. It is well separated from the desired 5′-O-silylated product and can be removed easily during the chromatography step. Therefore, the reaction is continued until all of the starting material is consumed, in order to also 5′-O-silylate any 3′-O-silylated nucleoside that has formed. It is best to do this reaction steadily, without pausing for an extended time between additions of silyl reagent. As soon as TLC shows the reaction is complete, immediately work up the reaction.
Protection of Nucleosides for Oligonucleotide Synthesis
2.10.7 Current Protocols in Nucleic Acid Chemistry
Supplement 16
O
O NH
HO
N
O HO
O
Me3Si
O
BzH-Cl O
O O 4
NH
O
CH2Cl2, DIPA
Me3Si
Si O
N
O
O
O HO
OAc
O
O O
OAc
OAc OAc
5
Figure 2.10.5 5′-Silylation. Synthesis of 5′-O-benzhydroxy-bis(trimethylsilyloxy)silyl-2′-O-bis(acetoxyethyloxy)methyl uridine from 2′-O-bis(acetoxyethyloxy)methyl uridine. BzH-Cl, benzhydroxy-bis(trimethylsilyloxy)silyl chlorosilane; DIPA, N,N-diisopropylamine.
30. Immediately after the reaction is complete according to TLC, extract the reaction mixture with 200 mL of 8% (w/v) aqueous NaHCO3, followed by 200 mL saturated aqueous NaCl. Dry the organic phase over 50 g Na2SO4 and concentrate on the rotary evaporator. 31. Dissolve the residue in a solution of 50 mL CH2Cl2/150 mL hexane/1 mL TEA. Load onto a 50 × 600–mm flash chromatography column filled with 250 g silica gel 60 in 80:20 (v/v) hexane/acetone containing 0.5% (v/v) TEA. 32. Elute the product using 50:30:20 (v/v/v) hexane/EtOAc/acetone containing 0.5% (v/v) TEA (typically 2 L). Collect 20-mL fractions. The molecular weight of 5′-O-BzH-2′-O-ACE-uridine (S.5; Fig. 2.10.5) is 850 g/mol. The yield is 17.43 g (75% to 90%). 1H NMR (CDCl3): δ 7.79 (d, J = 8.0 Hz, 1H), 7.32–7.18 (m, 10H), 5.99 (d, J = 3.2 Hz, 1H), 5.92–5.89 (m, 1H), 5.48–5.47 (m, 1H), 5.46 (s, 1H), 4.25–4.13 (m, 6H), 4.02–3.84 (m, 2H), 3.83–3.74 (m, 5H), 2.03 (s, 3H), 2.01 (s, 3H), 1.89 (s, 1H), 0.04 (s, 9H), 0.03 (s, 9H). 13C NMR (CDCl3): δ 171.0, 163.9, 150.8, 143.8, 139.9, 128.4, 127.5, 126.4, 126.3, 112.7, 102.4, 87.6, 84.2, 77.1, 68.8, 63.2, 63.1, 63.06, 63.02, 61.6, 20.94, 20.90, 1.6. Exact mass calculated for C37H54N2O15Si3: 850.3; found by ESMS (M + H): 851.
Prepare 3′-phosphoramidite (S.6) 33. Co-evaporate S.5 once with 2.5 mL MeCN per mmol S.5. Add 2 mL CH2Cl2 per mmol S.5, cap the reaction flask with a rubber septum, and stir the solution at 0°C in an ice bath. 34. In a separate flask under anhydrous conditions, dilute the phosphinylating reagent bis(N,N-diisopropylamino)methoxyphosphine (1.25 equiv per equiv S.5) with CH2Cl2 (2 mL per mmol phosphine). Add a solution of 0.45 M 1H-tetrazole in MeCN (0.5 equiv tetrazole per equiv S.5) and shake for 5 min. 35. Add the solution of activated phosphinylating reagent to the well-stirred solution of nucleoside at room temperature.
Preparation of 5′-Silyl-2′Orthoester Ribonucleosides
36. After 15 min, remove a 40-µL aliquot of the crude reaction mixture and add to 200 µL of CH2Cl2 containing 10% (v/v) TEA. Spot on a TLC plate pre-run in 60:30:10 (v/v/v) hexane/CH2Cl2/TEA. Develop the plate in this solvent mixture to check that the reaction is progressing. Continue to stir at room temperature until the reaction is complete by TLC analysis (12 to 18 hr). The desired product will move more quickly on the plate than the starting material.
2.10.8 Supplement 16
Current Protocols in Nucleic Acid Chemistry
O
O
Me3Si
O
NH
O Me3Si
O
Si O
N
O
Me3Si O
O i
O HO
O
O O
5
OAc OAc
Me3Si
NH
Si O
MeO
N
O
O P
O
O
O
O O
N
OAc OAc
6
Figure 2.10.6 Preparation of the phosphoramidite monomer for oligoribonucleotide synthesis. Synthesis of 5′-O-benzhydroxy-bis(trimethylsilyloxy)silyl-2′-O-bis(acetoxyethyloxy)methyl-3′-O-(N,N-diisopropylamino)methoxyphosphinyl uridine from 5′-O-benzhydroxy-bis(trimethylsilyloxy)silyl-2′-O-bis(acetoxyethyloxy)methyl uridine. i: bis(N,N-diisopropylamino)methoxyphosphine; 1H-tetrazole, CH2Cl2, 12 hr.
37. Add 10 equiv EtOH per equiv S.5 to quench the excess phosphine. Continue stirring for 30 min. Dry reaction on the rotary evaporator. 38. Purify the product on silica gel. Take up the residue from a 20-mmol reaction in a solution of 25 mL CH2Cl2/125 mL hexane/1 mL TEA and load onto a 50 × 600–mm flash chromatography column filled with 25 mL silica/g product in a solvent mix of 94:5:1 (v/v/v) hexane/CH2Cl2/TEA. 39. Elute the product with 75:25 (v/v) hexane/acetone containing 0.5% (v/v) TEA (typically 2 L). Collect 20-mL fractions, monitor by TLC (step 36), and pool fractions containing purified product. 40. Co-evaporate the pooled fractions with 100 mL toluene to ensure the amidite is not concentrated down into TEA. Finish with two co-evaporations using 100 mL portions of MeCN under high vacuum. Th e product, 5′-O-benzhydroxy-bis(trimethylsilyloxy)silyl-2′-O-bis(acetoxyethyloxy)methyl-3′-O-(N,N-diisopropylamino)methoxyphosphinyl uridine (S.6; Fig. 2.10.6), is a clear syrup, often pale yellow in color. The molecular weight is 1011 g/mol; the yield is 18.65 g (75% to 95%). 1H NMR (CDCl3, mixture of diasteromers): δ 8.08 (b, 1H), 7.81 and 7.80 (each as d, J = 8.0 Hz, 1H), 7.34–7.21 (m, 10H), 6.05 and 6.01 (each as d, J = 4.4 Hz, 1H), 5.93 and 5.92 (each as s, 1H), 5.58 and 5.45 (each as s, 1H), 5.41 and 5.35 (each as d, J = 8.0 Hz, 1H), 4.34–4.13 (m, 8H), 3.99–3.96 and 3.92–3.89 (each as m, 1H), 3.82–3.73 (m, 5H), 3.60–3.57 (m, 2H), 3.38 and 3.31 (each as d, J = 12.0 Hz, 3H), 2.08 and 2.06 (each as s, 6H), 1.20–1.15 (m, 12H), and 0.06, 0.06, 0.04, and 0.02 (each as s, 18H). 31P NMR (CDCl3, mixture of diasteromers): δ 152.2, 152.0. Exact mass calculated for C44H70N3O16PSi3: 1011.4; found by ESMS (M + H): 1012.
Protection of Nucleosides for Oligonucleotide Synthesis
2.10.9 Current Protocols in Nucleic Acid Chemistry
Supplement 16
ALTERNATE PROTOCOL 1
SYNTHESIS OF 2′-ACE-PROTECTED ADENOSINE RIBONUCLEOSIDE PHOSPHORAMIDITE The procedure for the synthesis of the N6-benzoyl-5′-O-silyl-2′-O-ACE-adenosine 3′-Ophosphoramidite is generally the same as for the uridine derivative (see Basic Protocol). The only significant changes occur in the solvent mixtures used for flash chromatographic purification of the various intermediates. Isolated yields per step are generally within ±10% of the yields obtained for the uridine derivatives. The TLC conditions for the analysis of the crude reaction mixtures described in the Basic Protocol may be used for analysis in this protocol as well. Although the Rf values of the adenosine derivatives will differ slightly from those of the uridine derivatives, the differences between starting material and desired product will be similar at each step. Additional Materials (also see Basic Protocol) N6-Benzoyladenosine (337 g/mol; Monomer Sciences) 1. Synthesize N6-benzoyl-3′,5′-O-TIPS-adenosine from N6-benzoyladenosine as described (see Basic Protocol, steps 1 to 12). For flash chromatography (steps 10 and 11), use a solvent system of 40:60 (v/v) hexane/EtOAc. The product has an Rf of 0.5 under these conditions. Its molecular weight is 579 g/mol. 1H NMR (CDCl3): δ 9.78 (b, 1H), 8.58 (s, 1H), 8.27 (s, 1H), 6.02 (s, 1H), 4.94–4.91 (m, 1H), 4.56 (d, J = 5.2 Hz, 1H), 4.39 (b, 1H), 4.09–4.06 (m, 2H), 3.96–3.92 (m, 1H), 3.28–3.24 (m, 1H), 1.20–1.17 (m, 6H), 1.03–0.84 (m, 28H). 13C NMR (CDCl3): δ 176.9, 152.3, 150.4, 142.0, 122.7, 81.9, 80.8, 74.9, 70.2, 61.1, 35.6, 19.1, 19.07, 17.3, 17.15, 17.14, 17.13, 16.9, 16.8, 16.79, 16.7, 13.1, 12.8, 12.6, 12.4. Exact mass calculated for C26H45N5O6Si2: 579.3; found by ESMS (M + H): 580.
2. Synthesize N6-benzoyl-3′,5′-O-TIPS-2′-O-ACE-adenosine as described (see Basic Protocol, steps 13 to 19). Elute the product (step 19) using 55:42:3:0.5 (v/v/v/v) hexane/EtOAc/MeOH/TEMED. Characterization data are not provided, because the crude product is directly carried to the next synthesis step.
3. Synthesize N6-benzoyl-2′-O-ACE-adenosine as described (see Basic Protocol, steps 20 to 25). Elute the product (step 25) using 95:5 (v/v) EtOAc/MeOH containing 0.5% (v/v) TEMED. The molecular weight of the product is 555 g/mol. 1H NMR (CDCl3): δ 9.23 (b, 1H), 8.63 (s, 1H), 8.14 (s, 1H), 5.99 (d, J = 7.6 Hz, 1H), 5.97 (b, 1H), 5.18 (s, 1H), 5.02 (dd, J = 4.8 Hz, 7.2 Hz, 1H), 4.48 (d, J = 4.8 Hz, 1H), 4.31–4.28 (m, 1H), 4.00–3.90 (m, 5H), 3.72–3.68 (m, 1H), 3.60–3.55 (m, 3H), 3.43–3.38 (m, 2H), 3.24–3.20 (m, 1H), 1.98 (s, 3H), 1.97 (s, 3H), 1.23 (d, J = 7.2 Hz, 6H). 13C NMR (CDCl3): δ 176.7, 170.9, 152.1, 150.2, 143.3, 123.5, 112.5, 89.2, 87.7, 76.3, 71.9, 63.3, 63.1, 62.7, 62.6, 62.57, 36.1, 20.8, 19.2. Exact mass calculated for C23H33N5O11: 555.2; found by ESMS (M + H): 556.
4. Synthesize N6-benzoyl-5′-O-BzH-2′-O-ACE-adenosine as described (see Basic Protocol, steps 26 to 32). Elute the product (step 32) using 45:35:20 (v/v/v) hexane/EtOAc/acetone containing 0.5% (v/v) TEA.
Preparation of 5′-Silyl-2′Orthoester Ribonucleosides
The molecular weight of the product is 943 g/mol. 1H NMR (CDCl3): δ 8.68 (b, 1H), 8.67 (s, 1H), 8.29 (s, 1H), 7.33–7.15 (m, 10H), 6.21 (d, J = 5.2 Hz, 1H), 5.91 (s, 1H), 5.31 (s, 1H), 4.83–4.79 (m, 1H), 4.36–4.33 (m, 1H), 4.13–4.09 (m, 3H), 4.05–4.03 (m, 2H), 3.91–3.87 (m, 1H), 3.78–3.75 (m, 1H), 3.69–3.62 (m, 3H), 3.52–3.46 (m, 1H), 3.28–3.24 (m, 1H), 3.05 (m, 1H), 1.99 (s, 6H), 1.27 (d, J = 6.8 Hz, 6H), 0.04 (s, 9H), 0.03 (s, 9H). 13 C NMR (CDCl3): δ 176.4, 170.8, 152.6, 151.2, 149.4, 144.0, 141.7, 128.3, 127.3, 126.4, 122.3, 112.5, 88.6, 84.9, 77.0, 76.97, 70.5, 63.1, 63.0, 62.8, 62.7, 62.6, 36.1, 20.8, 19.3, 1.5. Exact mass calculated for C42H61N5O14Si3: 943.4; found by ESMS (M + H): 944.
2.10.10 Supplement 16
Current Protocols in Nucleic Acid Chemistry
5. Synthesize N6-benzoyl-5′-O-BzH-2′-O-ACE-3′-O-(N,N-diisopropylamino)methoxyphosphinyl adenosine as described (see Basic Protocol, steps 33 to 40). Elute the product (step 39) with 70:30 (v/v) hexane/acetone containing 0.5% (v/v) TEA. The product (Fig. 2.10.1) is a clear syrup, often pale yellow in color. The molecular weight is 1105 g/mol. 1H NMR (CDCl3, mixture of diasteromers): δ 8.68 and 8.67 (each as s, 1H); 8.40 (b, 1H); 8.29 and 8.25 (each as s, 1H); 7.35–7.17 (m, 10H); 6.25 and 6.23 (each as d, J = 4.4 Hz, 1H); 5.93 and 5.91 (each as s, 1H); 5.41 and 5.31 (each as s, 1H); 4.85–4.82 (m, 1H); 4.59–4.55 (m, 1H); 4.29–4.21 (m, 1H); 4.10–3.91 (m, 5H); 3.78–3.54 (m, 7H); 3.40 and 3.33 (each as d, J = 12.0 Hz, 3H); 3.32–3.26 (m, 1H); 2.02 and 2.00 (each as s, 6H); 1.31 (d, J = 6.8 Hz, 6H); 1.28–1.14 (m, 12H); 0.06, 0.04, 0.03, and 0.02 (each as s, 18H). 31P NMR (CDCl3, mixture of diasteromers): δ 152.2, 151.4. Exact mass calculated for C49H77N6O15PSi3: 1104.5; found by ESMS (M + H): 1106.
SYNTHESIS OF 2′-ACE-PROTECTED GUANOSINE RIBONUCLEOSIDE PHOSPHORAMIDITE
ALTERNATE PROTOCOL 2
The procedure for the synthesis of the N2-isobutyryl-5′-O-silyl-2′-O-ACE-guanosine 3′-O-phosphoramidite is generally the same as for the uridine derivative (see Basic Protocol). The only significant changes occur in the solvent mixtures used for flash chromatographic purification of the various intermediates. Isolated yields per step are generally within ±10% of the yields obtained for the uridine derivatives. The TLC conditions for the analysis of the crude reaction mixtures described in the Basic Protocol may be used for analysis in this protocol as well. Although the Rf values of the guanosine derivatives will differ slightly from those of the uridine derivatives, the differences between starting material and desired product will be similar at each step. Additional Materials (also see Basic Protocol) N2-Isobutyrylguanosine (353 g/mol; Monomer Sciences) 1. Synthesize N2-isobutyryl-3′,5′-O-TIPS-guanosine from N2-isobutyrylguanosine as described (see Basic Protocol, steps 1 to 12). For flash chromatography (steps 10 and 11), use a solvent system of 30:70 (v/v) hexane/EtOAc. The product has an Rf of 0.5 under these conditions. Its molecular weight is 595 g/mol. 1H NMR (CDCl3): δ 12.30 (b, 1H), 10.66 (b, 1H), 7.94 (s, 1H), 5.72 (m, 1H), 4.45–4.42 (m, 1H), 4.28–4.26 (m, 1H), 4.14–4.09 (m, 2H), 4.01–3.93 (m, 1H), 2.94–2.88 (m, 1H), 1.26–1.21 (m, 6H), 1.08–0.86 (m, 28H). 13C NMR (CDCl3): δ 180.1, 156.1, 148.4, 148.1, 136.8, 121.3, 89.1, 81.7, 75.3, 69.6, 60.6, 36.2, 31.7, 36.2, 19.2, 19.1, 17.6, 17.43, 17.41, 17.3, 17.1, 16.9, 13.5, 13.1, 13.0, 12.6. Exact mass calculated for C26H45N5O7Si2: 595.3; found by ESMS (M + H): 596.
2. Synthesize N2-isobutyryl-3′,5′-O-TIPS-2′-O-ACE-guanosine as described (see Basic Protocol, steps 13 to 19). Elute the product (step 19) using 47:50:3:0.5 (v/v/v/v) hexane/EtOAc/MeOH/TEMED. Characterization data are not provided, because the crude product is directly carried to the next synthesis step.
3. Synthesize N2-isobutyryl-2′-O-ACE-guanosine as described (see Basic Protocol, steps 20 to 25). Elute the product (step 25) using 64:30:6 (v/v/v) EtOAc/acetone/MeOH containing 0.5% (v/v) TEMED. The molecular weight of the product is 571 g/mol. 1H NMR (CDCl3): δ 12.30 (b, 1H), 10.38 (b, 1H), 8.15 (s, 1H), 5.93 (d, J = 4.4 Hz, 1H), 5.37 (s, 1H), 4.92 (b, 1H), 4.73–4.70 (m, 1H), 4.57–4.45 (m, 1H), 4.37 (b, 1H), 4.12–4.02 (m, 5H), 3.89–3.85 (m, 1H), 3.77–3.74 (m, 1H), 3.67–3.56 (m, 4H), 2.82–2.74 (m, 1H), 1.97 (s, 3H), 1.95 (s, 3H), 1.17–1.14 (m, 6H). 13C NMR (CDCl3): δ 180.3, 171.3, 171.2, 155.7, 148.3, 139.1, 120.9, 112.4, 87.5, 85.6, 76.7, 70.0, 63.2, 63.1, 62.9, 62.3, 61.5, 35.9, 31.0, 20.9, 19.0. Exact mass calculated for C23H33N5O12: 571.2; found by ESMS (M + H): 572.
Protection of Nucleosides for Oligonucleotide Synthesis
2.10.11 Current Protocols in Nucleic Acid Chemistry
Supplement 16
4. Synthesize N2-isobutyryl-5′-O-BzH-2′-O-ACE-guanosine as described (see Basic Protocol, steps 26 to 32). Elute the product (step 32) using 45:35:30 (v/v/v) hexane/EtOAc/acetone containing 0.5% (v/v) TEA. The molecular weight of the product is 959 g/mol. 1H NMR (CDCl3): δ 12.29 (b, 1H), 10.04 (b, 1H), 7.96 (s, 1H), 7.31–7.12 (m, 10H), 5.96 (d, J = 5.2 Hz, 1H), 5.90 (s, 1H), 5.22 (s, 1H), 4.53–4.50 (m, 1H), 4.36–4.30 (m, 1H), 4.20–4.14 (m, 1H), 4.10–4.06 (m, 4H), 3.82–3.80 (m, 1H), 3.77–3.75 (m, 1H), 3.68–3.61 (m, 3H), 3.52–3.50 (m, 1H), 4.41–3.38 (m, 1H), 2.83–2.78 (m, 1H), 2.00 (s, 3H), 1.99 (s, 3H), 1.19 (t, J = 7.2 Hz, 6H), 0.01 (s, 9H), 0.00 (s, 9H). 13C NMR (CDCl3): δ 179.8, 171.3, 171.0, 155.9, 148.6, 148.1, 143.9, 143.8, 137.3, 128.24, 128.23, 127.2, 126.32, 126.29, 121.0, 112.4, 86.2, 84.8, 77.8, 76.9, 70.5, 63.15, 63.07, 62.9, 62.80 62.6, 36.0, 20.9, 20.8, 19.00, 18.98, 14.8, 14.7. Exact mass calculated for C42H61N5O15Si3: 959.4; found by ESMS (M + H): 960.
5. Synthesize N2-isobutyryl-5′-O-BzH-2′-O-ACE-3′-O-(N,N-diisopropylamino)methoxyphosphinyl guanosine as described (see Basic Protocol, steps 33 to 40). Elute the product (step 39) with 70:30 (v/v) hexane/acetone containing 0.5% (v/v) TEA. The product (Fig. 2.10.1) is a clear syrup, often pale yellow in color. The molecular weight is 1120 g/mol. 1H NMR (CDCl3, mixture of diasteromers): δ 12.06 (b, 1H); 9.05 and 8.97 (each as b, 1H); 8.00 and 7.96 (each as s, 1H); 7.35–7.17 (m, 10H); 6.09 and 6.03 (each as d, J = 4.0 Hz, 1H); 5.93 and 5.91 (each as s, 1H); 5.39 and 5.26 (each as s, 1H); 4.50–4.45 (m, 2H); 4.24–4.11 (m, 8H); 3.86–3.83 (m, 2H); 3.76–3.56 (m, 9H); 3.38 and 3.34 (each as d, J = 12.0 Hz, 3H); 2.69–2.64 (m, 1H); 2.07, 2.06, and 2.05 (each as s, 6H); 1.29–1.17 (m, 12H); 0.06, 0.04, 0.04, and 0.02 (each as s, 18H). 31P NMR (CDCl3, mixture of diasteromers): δ 152.0, 151.8. Exact mass calculated for C49H77N6O16PSi3: 1120.4; found by ESMS (M + H): 1121. ALTERNATE PROTOCOL 3
SYNTHESIS OF 2′-ACE-PROTECTED CYTIDINE RIBONUCLEOSIDE PHOSPHORAMIDITE The procedure for the synthesis of the N4-acetyl-5′-O-silyl-2′-O-ACE-cytidine 3′-Ophosphoramidite is generally the same as for the uridine derivative (see Basic Protocol). The only significant changes occur in the solvent mixtures used for flash chromatographic purification of the various intermediates. Isolated yields per step are generally within ±10% of the yields obtained for the uridine derivatives. The TLC conditions for the analysis of the crude reaction mixtures described in the Basic Protocol may be used for analysis in this protocol as well. Although the Rf values of the cytidine derivatives will differ slightly from those of the uridine derivatives, the differences between starting material and desired product will be similar at each step. Additional Materials (also see Basic Protocol) N4-Acetylcytidine (285 g/mol; Monomer Sciences) 1. Synthesize N4-acetyl-3′,5′-O-TIPS-cytidine from N4-acetylcytidine as described (see Basic Protocol, steps 1 to 12). For flash chromatography (steps 10 and 11), use a solvent system of 45:55 (v/v) hexane/EtOAc. The product has an Rf of 0.5 under these conditions. Its molecular weight is 527 g/mol. 1H NMR (CDCl3,): δ 10.63 (b, 1H), 8.14 (d, J = 7.6 Hz, 1H), 7.35 (d, J = 7.6 Hz, 1H), 5.71 (s, 1H), 4.19–4.13 (m, 4H), 3.92–3.80 (m, 2H), 2.21 (s, 3H), 0.99–0.79 (m, 28H). 13C NMR (CDCl3): δ 171.6, 163.4, 155.0, 144.2, 96.7, 91.5, 81.8, 75.0, 63.3, 59.8, 24.8, 17.4, 17.3, 17.23, 17.19, 16.9, 16.8, 16.7, 16.6, 13.3, 12.9, 12.8, 12.3. Exact mass calculated for C23H41N3O7Si2: 527.3; found by ESMS (M + H): 528.
Preparation of 5′-Silyl-2′Orthoester Ribonucleosides
2.10.12 Supplement 16
Current Protocols in Nucleic Acid Chemistry
2. Synthesize N4-acetyl-3′,5′-O-TIPS-2′-O-ACE-cytidine as described (see Basic Protocol, steps 13 to 19). Elute the product (step 19) using 46:50:4:0.5 (v/v/v/v) hexane/EtOAc/MeOH/TEMED. Characterization data are not provided, because the crude product is directly carried to the next synthesis step.
3. Synthesize N4-acetyl-2′-O-ACE-cytidine as described (see Basic Protocol, steps 20 to 25). Elute the product (step 25) using 67:30:3 (v/v/v) EtOAc/acetone/MeOH containing 0.5% (v/v) TEMED. The molecular weight of the product is 503 g/mol. 1H NMR (CDCl3): δ 9.65 (b, 1H), 8.27 (d, J = 7.6 Hz, 1H), 7.41 (d, J = 7.2 Hz, 1H), 5.77 (d, J = 2.8 Hz, 1H), 5.60 (s, 1H), 4.62–4.59 (m, 1H), 4.38–4.36 (m, 1H), 4.23–4.13 (m, 5H), 4.03–3.98 (m, 2H), 3.85–3.79 (m, 5H), 3.52 (b, 1H), 2.23 (s, 3H), 2.05 (s, 3H), 2.04 (s, 3H). 13C NMR (CDCl3): δ 171.2, 171.16, 163.1, 155.6, 146.8, 112.9, 97.1, 92.4, 85.3, 68.6, 63.25, 63.17, 63.10, 62.9, 60.8, 25.1, 21.1, 21.0. Exact mass calculated for C20H29N3O12: 503.2; found by ESMS (M + H): 504.
4. Synthesize N4-acetyl-5′-O-BzH-2′-O-ACE-cytidine as described (see Basic Protocol, steps 26 to 32). Elute the product (step 32) using 30:50:20 (v/v/v) hexane/EtOAc/acetone containing 0.5% (v/v) TEA. The molecular weight of the product is 891 g/mol. 1H NMR (CDCl3): δ: 10.23 (b, 1H), 8.30 (δ, J = 7.6 Hz, 1H), 7.36–7.14 (m, 10H), 5.93–5.91 (m, 2H), 5.67 (s, 1H), 5.23 (s, 1H), 4.24–4.16 (m, 5H), 4.11–3.98 (m, 3H), 3.88–3.76 (m, 5H), 3.01 (δ, J = 8.0 Hz, 1H), 2.26 (s, 3H), 2.03 (s, 3H), 2.02 (s, 3H), 0.07 (s, 9H), 0.05 (s, 9H). 13C NMR (CDCl3): δ 171.0, 163.2, 155.0, 144.6, 143.91, 143.87, 128.3, 127.3, 126.35, 126.29, 112.8, 96.7, 89.4, 83.8, 78.5, 77.0, 67.2, 63.2, 63.1, 63.0, 62.9, 62.7, 60.6, 53.5, 24.9, 20.9, 20.8, 1.6. Exact mass calculated for C39H57N3O15Si3: 891.3; found by ESMS (M + H): 892.
5. Synthesize N4-acetyl-5′-O-BzH-2′-O-ACE-3′-O-(N,N-diisopropylamino)methoxyphosphinyl cytidine as described (see Basic Protocol, steps 33 to 40). Elute the product (step 39) with 65:35 (v/v) hexane/acetone containing 0.5% (v/v) TEA. The product (Fig. 2.10.1) is a clear syrup, often pale yellow in color. The molecular weight is 1052 g/mol. 1H NMR (CDCl3, mixture of diasteromers): δ 8.77 (b, 1H); 8.36 and 8.34 (each as d, J = 4.8 Hz, 1H); 7.36–7.15 (m, 10H); 6.00 (d, J = 6.8 Hz, 1H); 5.94 (d, J = 4.8 Hz, 1H); 5.81 and 5.76 (each as s, 1H); 4.30–4.15 (m, 8H); 4.10–4.02 (m, 1H); 3.96–3.77 (m, 5H); 3.60–3.52 (m, 3H); 3.34 and 3.31 (each as d, J = 12.0 Hz, 3H); 2.23 (s, 3H); 2.05, 2.05, and 2.04 (each as s, 6H); 1.20–1.13 (m, 12H); 0.08, 0.07, 0.06, and 0.04 (each as s, 18H). 31P NMR (CDCl3, mixture of diasteromers): δ 152.2, 151.4. Exact mass calculated for C46H73N4O16PSi3: 1052.4; found by ESMS (M + H): 1053.
COMMENTARY Background Information Until recently, most advancements in oligonucleotide synthesis technologies focused on DNA. One reason for this “DNA-centricity” is that DNA is easier to work with than RNA, making advancements in DNA technologies easier to develop. Another, more important, reason is that biologists have traditionally attributed DNA with far more therapeutic significance than RNA. As a result, the market for synthetic RNA has traditionally been small and highly specialized. The recent discovery that small interfering RNAs (siRNAs) induce gene suppression in mammalian cells (Elbashir et al., 2001) has,
however, sparked tremendous interest in developing siRNA-based assays and high-throughput screens to identify and study disease targets. Reliable, efficient, and high-quality RNA synthesis methods are now critically important for enabling cutting-edge research in drug discovery and therapeutic intervention. As a result, a substantial and rapidly growing market for synthetic RNA has been created. Three methods for chemically synthesizing siRNAs are now well established: 2′-TBDMS, 2′- TOM, and 2′-ACE. 2′-O-tert-Butyldimethylsilyl (TBDMS) chemistry was originally developed in the 1980s, when Ogilvie and Usman applied a 2′-TBDMS strategy for 2′-hy-
Protection of Nucleosides for Oligonucleotide Synthesis
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Preparation of 5′-Silyl-2′Orthoester Ribonucleosides
droxyl protection (Usman et al., 1987) to classical phosphoramidite-based DNA synthesis (Caruthers, 1987; also see UNIT 2.5). The more recently developed 2′-O-triisopropylsilyloxymethyl (TOM) 2′-hydroxyl chemistry implements a 2′-silyl protecting group to achieve improved synthesis efficiencies relative to 2′TBDMS (Wu and Pitsch, 1998; also see UNIT 2.9). 2′-TOM chemistry is still limited, however, by such factors as oligonucleotide length and yield. Significant improvements in RNA synthesis were ach ieved with the 5′-silyl-2′-acetoxyethoxy (ACE) chemistry (Scaringe et al., 1998) described in this unit. For synthetic RNA to be useful to biological practitioners studying gene function, it has to meet certain physical, quality, and convenience criteria. Some of the characteristics of synthetic RNA that are important to the user are: 1. Sequence integrity: assurance that the sequence desired is actually the one produced. 2. Scalability: the ability to obtain large quantities of RNA for use in high-throughput screens or animal models. 3. Strand length: the ability to obtain long strands of RNA (e.g., to make hairpin structures). 4. Chemical modifications: the ability to obtain RNA featuring chemical labels or preservatives (e.g., to improve pharmacokinetics in animal tissues). 5. Robustness: the ability of the RNA protecting groups to repel nucleases, facilitating handling and long-term storage. 6. Labor intensiveness: the ease of deprotecting, annealing, purifying, and/or preparing the RNA for use in a biological assay. 7. Short- and long-term cost: as measured in both time and money. 2′-ACE chemistry is the only method capable of meeting all of the criteria listed above. First, 2′-ACE chemistry provides superior sequence integrity; 99% efficiencies for 21-mers are routinely achieved at coupling speeds as fast as 2 nucleotides/min. By comparison, 2′-TOM chemistry achieves 99% efficiency when one nucleotide is coupled every 2 to 6 min. Second, 2′-ACE chemistry delivers a crude product of high purity. 2′-ACE chemistry produces milligram to kilogram quantities of >90% pure material for high-throughput or in vivo efforts in less time, and at a lower cost, than any other method. Third, 2′-ACE is the only chemical method that can be used to economically synthesize long double-stranded RNA (dsRNA) or siRNA hairpins >30 bases in length; oligori-
bonucleotides of up to 80 bases can be synthesized at efficiencies of ∼60%, far greater than for any other method. By comparison, 2′-TOM chemistry achieves 36% efficiencies for synthesis of 50-mers. Fourth, the 2′-ACE protecting scheme enables virtually any chemical modification to be easily incorporated into the oligoribonucleotide. For in vivo applications, such chemical modifications are required to increase RNA stability in tissues and fluids. The 2′-TBDMS and 2′-TOM methods are more restricted in the type and number of modifications that can be added. Fifth, 2′-ACE-protected RNA provides superior handling properties relative to other methods. As with other protecting strategies, the 2′-ACE group renders the oligoribonucleotide resistant to nuclease and other forms of degradation during handling and purification. 2′-ACE chemistry is the only method, however, in which the last deprotection step can be carried out in a mild, aqueous solution. This step is typically done using 100 mM acetic acid, pH 3.8 (conditions under which the RNA is quite stable) and takes only 30 min to complete. Finally, when considering cost, 2′-ACE chemistry provides better economies of scale than the other methods. For smallscale production runs, 2′-ACE is slightly more expensive than 2′-TBDMS and 2′-TOM; however, when larger-scale production runs are required, fast coupling times and high coupling efficiencies ensure that 2′-ACE chemistry is cheaper than competing technologies.
Critical Parameters and Troubleshooting For all of these protocols, anhydrous reaction conditions must be maintained. Traces of water will severely affect the yield and purity of the product at each step. Solvents should be stored over molecular sieves or should be freshly distilled from an appropriate drying agent. All glassware must be thoroughly cleaned and dried in an oven for 30 min at 110° to 120°C prior to use. Thin-layer chromatography (TLC) on silica gel is conveniently used throughout these protocols to monitor the progress of the various reactions. It is also used to provide a rapid initial assessment of the purity of the chromatographically isolated products. The absolute purity of the product compounds, however, should be determined by NMR spectroscopy (1H, 13C, or 31P, where appropriate) and/or mass spectrometry. During the synthesis of 3′,5′-O-TIPS-2′-OACE-nucleosides from 3′,5′-O-TIPS-nucleo-
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sides (e.g., S.3; see Basic Protocol, steps 13 to 19), an additional aliquot of pyridinium p-toluenesulfonate (up to 1.0 equiv total) can be added to complete the reaction. If additional pyridinium p-toluenesulfonate is added, however, additional TEMED (up to 2.5 equiv) must be added to compensate. This reaction typically takes 36 to 48 hr to complete at room temperature, but it can be accelerated by gently refluxing at 38° to 40°C. The solution should eventually appear clear, with a slight yellow or orange color. When analyzing the progress of this reaction with TLC, beware that 4-tertbutyldimethylsiloxy-3-penten-2-one is UV absorbing and volatile. It appears on the TLC plate as a dark spot or pair of spots (two isomers) migrating at the solvent front. Upon standing, the spots will slowly disappear. When preparing a silica gel flash chromatography column to purify the desired product (see Basic Protocol, step 18), only use TEMED as the base. Do not use TEA or another tertiary amine. TEMED-HF is used to deprotect the 5′and 3′-hydroxyls in the next step. If TEA is present in that step, the HF salt of TEA will be generated. The TEA-HF salt migrates on silica gel and is difficult or impossible to separate from the 2′-O-ACE nucleoside product. This problem does not occur with the more polar TEMED-HF salt. When eluting, it is very important to remove the several small impurities that elute just ahead of the desired product. If present, they severely complicate subsequent purifications. Therefore, when the product begins to elute, collect smaller fractions and analyze carefully by TLC. During the synthesis of 2′-ACE-nucleosides f rom 3′,5′-O-TIPS-2′-O-ACE-nucleosides (see Basic Protocol, steps 20 to 25), it is important to take care when loading the silica gel flash chromatography column. If the product is not loaded slowly, heat can be generated, which may disrupt the silica gel bed. During the synthesis of 5′-O-BzH-2′-OACE-nucleosides from 2′-O-ACE-nucleosides (see Basic Protocol, steps 26 to 32), it is important to add the diisopropylamine-activated silylating reagent slowly to the nucleoside solution. Begin by adding 0.5 equiv of the silylating reagent, and then taper to smaller aliquots as the reaction proceeds to completion. It is best to do this reaction steadily, over a period of 10 to 15 min, without pausing for an extended time between additions of silyl reagent. The silylating reagent appears to react somewhat with the nucleobases on uridine and guanosine, requiring more than 1.0 equiv to 5′-protect all of the
starting material. As soon as TLC shows that the reaction is complete, immediate work-up of the reaction is required. At the end of the final synthesis step, yielding 5′-O-BzH-2′-O-ACE-3′-O-(N,N-diisopropylamino)methoxyphosphinyl nucleosides from 5′-O-BzH-2′-O-ACE-nucleosides, elute the final product from the chromatography column, and then co-evaporate the pooled fractions with toluene to ensure the amidite is not concentrated down into TEA. Finish with one or two co-evaporations using acetonitrile on high vacuum.
Anticipated Results The isolated yields presented in the protocols represent typical values obtained. The quality of the reagents and the dryness of the solvents will have a significant impact on product yield and purity, as will the amount of care taken during the chromatographic purifications. The product yields are explicitly given at each step for uridine in the Basic Protocol. These values are also representative of the yields obtained for the corresponding adenosine, guanosine, and cytidine derivatives, within about ±10%. The TIPS-protected nucleosides obtained in the first step of the synthetic scheme (S.2) are generally white solids; all other products are generally viscous syrups, often pale yellow in color. It is possible to analyze all products by reversed-phase high-performance liquid chromatography (RP-HPLC). Typical conditions are as follows: analytical C18 column (available from a variety of vendors); linear gradient of HPLC-grade acetonitrile in aqueous 0.05 M ammonium acetate, pH 8.5; and UV detector monitoring at 260 nm. Typical HPLC analytical data for the final phosphoramidites (Waters NovaPak C18 column, 3.9 × 150–mm plus guard cartridge; flow rate, 1 mL/min; linear gradient from 25% to 100% [v/v] acetonitrile over 30 min): retention times of 33 to 35 min; purities ≥97% (by integration of all peaks in the 260-nm chromatogram); ≤0.1% desilylated material (retention times of 16 to 18 min); ≤1% hydrolyzed material (retention times of 25 to 27 min). All methods described in this unit can be scaled up to yield ∼300 mmol (∼300 g) of phosphoramidites simply by linearly increasing the amounts of all materials used in the procedures. The protected phosphoramidite monomers prepared using these methods are ideal for solid-phase RNA oligonucleotide synthesis.
Protection of Nucleosides for Oligonucleotide Synthesis
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Time Considerations Chemical synthesis of 2′-ACE-protected phosphoramidites is an involved process that, from start to finish, takes experienced chemists days to weeks to complete. The following time estimates include reagent preparation, labor, reaction, evaporation, purification, and drying, and assume that all reactions proceed smoothly. Synthesis of 3′,5′-O-(tetraisopropyldisiloxane1,3-diyl)uridine from uridine (steps 1 to 12 of the Basic Protocol) takes ∼2 days. Synthesis of 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-2′O-ACE-uridine (steps 13 to 19) takes ∼3 days. Synthesis of 2′-O-ACE-uridine (steps 20 to 25) and synthesis of 5′-O-BzH-2′-O-ACE-uridine (steps 26 to 32) each take 1 day. Synthesis of 5′-O-BzH-2′-O-ACE-3′-O-(N,N-diisopropylamino)methoxyphosphinyl uridine (steps 33 to 40) takes ∼2 days. As such, synthesis of 2′-OACE-protected uridine ribonucleoside phosphoramidites takes ∼9 days in total. Times for Alternate Protocols 1 to 3 are similar.
Elbashir, S.M., Harborth, J., Lendeckel, W., Yalcin, A., Weber, K., and Tuschl, T. 2001. Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells. Nature 411:494498. Matteucci, M.D. and Caruthers, M.H. 1981. Synthesis of deoxyoligonucleotides on a polymer support. J. Am. Chem. Soc. 103:3185-3191. Scaringe, S.A., Wincott, F.E., and Caruthers, M.H. 1998. Novel RNA synthesis method using 5′silyl-2′-orthoester protecting groups. J. Am. Chem. Soc. 120:11820-11821. Usman, N.O., Ogilvie, K.K., Jiang, M.Y., and Cedergren, R.J. 1987. The automated chemical synthesis of long oligoribonucleotides using 2′-Osilylated ribonucleoside 3′-O-phosphoramidites on a controlled-pore glass support: Synthesis of a 43-nucleotide sequence similar to the 3′-half molecule of an Escherichia coli formylmethionine tRNA. J. Am. Chem. Soc. 109:7845-7854. Wu, X. and Pitsch, S. 1998. Synthesis and pairing properties of oligoribonucleotide analogues containing a metal-binding site attached to β-D-allofuranosyl cytosine. Nucl. Acids Res. 26:43154323.
Literature Cited Caruthers, M.H. 1985. Gene synthesis machines: DNA chemistry and its uses. Science 230:281285.
Contributed by Stephen A. Scaringe, David Kitchen, Robert J. Kaiser, and William S. Marshall Dharmacon Inc. Lafayette, Colorado
Preparation of 5′-Silyl-2′Orthoester Ribonucleosides
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Enzymatic Regioselective Levulinylation of 2-Deoxyribonucleosides and 2-O-Methylribonucleosides
UNIT 2.11
With the recent success of various aptamers, ribozymes, and other oligonucleotides (e.g., antisense, immunostimulatory CpG-containing, and RNAi-related) undergoing human clinical trials, the possibility of their commercial launch in the near future may require production of very large quantities of these products. Thus, development of methods for their large-scale synthesis has become a high priority (Pon et al., 1999). Particularly when ton quantities of oligonucleotides are required, solution-phase synthesis appears to be a preferred alternative to traditional solid-phase synthesis. The key building blocks for solution-phase oligonucleotide synthesis are appropriately 3 - and/or 5 -protected nucleosidic monomers. Of the limited number of protecting groups available for 3 and/or 5 -hydroxyl protection, the levulinyl group is frequently chosen, as it is stable under oligonucleotide coupling reaction conditions and can be selectively cleaved at neutral pH without affecting other protecting groups in the same molecule (Greene and Wuts, 1999; Kocie´nski, 2004; also see other units in this chapter). Until recently, the preparation of these building blocks had been carried out through several tedious chemical protection/deprotection steps (Kumar and Poonian, 1984; Iwai and Ohtsuka, 1988; Iwai et al., 1990; Krotz et al., 1997; Eleuteri et al., 1999; Reese and Song, 1999; March´an et al., 2004). Clearly, such methods for the synthesis of 3 - and/or 5 -levulinyl-protected nucleosides are not suitable for the large-scale production of these key building blocks. Selective protection and deprotection of compounds such as nucleosides that contain multiple hydroxyl groups is a challenging problem in organic synthesis. For the manipulation of protecting groups, application of biocatalysts in organic synthesis has become an attractive alternative to conventional chemical methods (Bornscheuer and Kazlauskas, 1999; Carrea and Riva, 2000; Patel, 2000; Klibanov, 2001; Koeller and Wong, 2001). Enzymes have been reported to catalyze reactions with high chemo-, regio-, and stereoselectivity (Ferrero and Gotor, 2000a,b). Furthermore, enzyme-catalyzed reactions are less hazardous, polluting, and energy consuming than their conventional counterparts. Among various commercial enzymes, the authors have had success with the use of lipases as versatile biocatalysts in organic synthesis (Garc´ıa et al., 2002, 2003, 2004a,b). Lipases are attractive because they are readily available, do not require cofactors, are inexpensive and highly stable, exhibit broad substrate specificity, and retain a high degree of activity in organic solvents. The following Basic Protocols present reliable procedures using commercial lipases for the regioselective introduction of the levulinyl group at the 3 - or 5 -hydroxy position of a 2 -deoxyribonucleoside or a 2 -O-methylribonucleoside. Additionally, Support Protocols describe the preparation of O-levulinyl acetonoxime from levulinic acid and acetonoxime (see Support Protocol 1) and the synthesis of 3 ,5 -di-O-levulinyl esters (see Support Protocols 2 and 3), which serve as starting material in several methods.
SYNTHESIS OF 3 -O-LEVULINYL-2 -DEOXYRIBONUCLEOSIDES The preparation of 3 -O-levulinyl-2 -deoxyribonucleosides involves regioselective acylation of the 3 -hydroxyl function of deoxyribonucleosides with O-levulinyl acetonoxime using immobilized Pseudomonas cepacia lipase (PSL-C), as illustrated in Figure 2.11.1. Protection of Nucleosides for Oligonucleotide Synthesis Contributed by Iv´an Lavandera, Javier Garc´ıa, Susana Fern´andez, Miguel Ferrero, Vicente Gotor, and Yogesh S. Sanghvi
2.11.1
Current Protocols in Nucleic Acid Chemistry (2005) 2.11.1-2.11.36 C 2005 by John Wiley & Sons, Inc. Copyright
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Figure 2.11.1 Regioselective levulinylation of deoxyribonucleosides using Pseudomonas cepacia lipase (PSL-C) or Candida antarctica lipase B (CAL-B). Abbreviations: ABz , N 6 -benzoyladenin9-yl; CBz , N 4 -benzoylcytosin-1-yl; Gi-Bu , N 2 -isobutyrylguanin-9-yl; T, thymin-1-yl.
Figure 2.11.2 Regioselective hydrolysis of N 2 -isobutyryl-3 ,5 -di-O-levulinyl-2 -deoxyguanosine (S.4d) using CAL-B. Abbreviations: DCC, 1,3-dicyclohexylcarbodiimide; DMAP, 4-(dimethylamino)pyridine; Et3 N, triethylamine; Lev, levulinyl; LevOH, levulinic acid.
This is a general procedure for T (see Basic Protocol 1), dC (see Basic Protocol 2), and dA (see Basic Protocol 3). Preparation of O-levulinyl acetonoxime from levulinic acid and acetonoxime is presented in Support Protocol 1. Enzymatic Regioselective Levulinylation of Nucleosides
Because direct regioselective enzymatic acylation of N2 -isobutyryl-2 -deoxyguanosine using PSL-C does not work efficiently, another lipase-catalyzed approach is used to prepare the dG monomer (see Basic Protocol 4). The dG monomer is prepared from N2 isobutyryl-3 ,5 -di-O-levulinyl-2 -deoxyguanosine via regioselective hydrolysis of the
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5 -O-levulinyl ester, catalyzed by Candida antarctica lipase B (CAL-B) as depicted in Figure 2.11.2. Preparation of 3 ,5 -di-O-levulinylated deoxyguanosine using levulinic acid is presented in Support Protocol 2. CAUTION: Wear safety glasses and gloves, and perform all operations involving solvents and reagents in a well-ventilated fume hood.
Preparation of 3 -O-Levulinylthymidine via Regioselective Enzymatic Acylation with PSL-C
BASIC PROTOCOL 1
Materials Thymidine (Rasayan) O-Levulinyl acetonoxime (see Support Protocol 1) Immobilized Pseudomonas cepacia lipase (PSL-C; 904 propyl laurate units per gram [PLU/g]; Amano Enzyme) Anhydrous nitrogen Anhydrous tetrahydrofuran (THF), freshly distilled from sodium and benzophenone Methanol (MeOH) Dichloromethane (CH2 Cl2 ) p-Anisaldehyde solution (see recipe) Saturated sodium bicarbonate (NaHCO3 ) solution Sodium sulfate (Na2 SO4 ) Diethyl ether (Et2 O), cold (4◦ C) Silica gel Ethyl acetate (EtOAc) 1-L Erlenmeyer flask equipped with a three-way glass stopcock Rubber septum Orbital shaker, 30◦ C Microcentrifuge tubes Capillary tubes Silica gel 60 F254 aluminum-backed TLC plates UV light source Heat gun Buchner funnels with filter paper circles 250- and 500-mL round-bottom flasks Rotary evaporator equipped with a cooling condenser 500-mL separatory funnel 250-mL Erlenmeyer flasks 1-L filter flask Glass funnel with Whatman no. 1 filter paper Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) Perform levulinylation 1. Add 10 g (41.28 mmol) thymidine, 21.2 g (123.86 mmol) O-levulinyl acetonoxime, and 20 g PSL-C to a 1-L Erlenmeyer flask equipped with a three-way glass stopcock. 2. Evacuate the Erlenmeyer flask on a vacuum line, and then flush the flask with nitrogen. Repeat this procedure three times. 3. Add 206 mL dry THF. 4. Seal the flask with a rubber septum and place in an orbital shaker kept at 30◦ C. Shake the reaction mixture at 250 rpm.
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5. Monitor the progress of the reaction by analytical TLC (APPENDIX 3D) as follows: a. Take a sample (usually 20 to 50 µL) under nitrogen using a capillary tube. b. Transfer the sample to a microcentrifuge tube, dilute with 50 to 100 µL MeOH, and spot on a silica gel 60 F254 aluminum-backed plate. c. Develop plate using 10% (v/v) MeOH/CH2 Cl2 as the eluent. d. Visualize spots first under UV light and then by dipping the plate into panisaldehyde solution and warming with a heat gun. At this point in the synthesis, the solution becomes clear, with immobilized lipase that will be filtered off. The levulinylation reaction takes between 14 and 16 hr to complete. To analyze the course of reactions by TLC, the p-anisaldehyde solution is useful for visualizing nucleosides.
Work up and purify product 6. When the reaction is complete, filter off the lipase under vacuum using a Buchner funnel and filter paper circle, and collect the filtrate in a 500-mL round-bottom flask. 7. Wash PSL-C twice with 50 mL CH2 Cl2 , collecting washes in the same round-bottom flask. 8. Concentrate the filtrate under reduced pressure using a rotary evaporator. The resulting oil will develop an orange color due to the presence of excess oxime ester.
9. Dissolve the residue in 100 mL CH2 Cl2 and transfer to a 500-mL separatory funnel. 10. Remove unreacted thymidine by extracting the organic layer four times with 20 mL saturated NaHCO3 solution, collecting the phases in 250-mL Erlenmeyer flasks. Verify that thymidine has been completely removed from the organic phase by analytical TLC of both phases. 11. Extract the aqueous phases six to eight times with 20 mL CH2 Cl2 to recover any extracted product, again collecting in a 250-mL Erlenmeyer flask. 12. Pool the organic phases from step 11 and verify by TLC that they contain no residual thymidine. If thymidine is present, re-extract with 20 mL saturated NaHCO3 solution. 13. Pool the organic phases from steps 10 and 12 and dry over 4 g Na2 SO4 for 10 to 15 min. Remove the drying agent by gravity filtration through Whatman no. 1 filter paper in a glass funnel, and collect the filtrate in a 500-mL round-bottom flask. 14. Wash the Na2 SO4 four times with 25 mL CH2 Cl2 , and collect all washes in the same round-bottom flask. 15. Concentrate the dried organic extract under reduced pressure using a rotary evaporator. The orange color may persist.
16. Dissolve the residue in 10 mL CH2 Cl2 . Gradually add the solution to a beaker containing 50 mL cold (4◦ C) Et2 O to precipitate the product as a white solid. Cool the precipitate at 4◦ C overnight.
Enzymatic Regioselective Levulinylation of Nucleosides
17. Filter off the precipitate under vacuum using a Buchner funnel and filter paper circle, collecting the filtrate in a 250-mL round-bottom flask. Wash the white solid twice with 50 mL cold Et2 O into the same round-bottom flask. Dry the white solid under vacuum. The average yield of levulinylated product obtained in this step is 75% to 80%.
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Recover product from filtrate 18. Evaporate Et2 O washes under reduced pressure using a rotary evaporator. 19. To recover product by dry column chromatography, weigh out 50 g silica gel and place in a Buchner funnel with filter paper circle, attached to a 1-L filter flask connected to a vacuum line. Wash the silica with 200 mL of 25% (v/v) EtOAc/Et2 O. 20. Using a Pasteur pipet, spread the residue carefully on top of the silica bed. 21. Elute the residual oxime ester with ∼500 to 700 mL of 25% (v/v) EtOAc/Et2 O. 22. Using clean filter flasks to collect 100-mL fractions, elute the desired product with ∼800 to 1200 mL of 10% (v/v) MeOH/CH2 Cl2 . Monitor fractions by analytical TLC. 23. Transfer fractions containing pure levulinylated product to a 500-mL round-bottom flask, and remove solvents under reduced pressure using a rotary evaporator. 24. Add 5 mL Et2 O and triturate the product. Evaporate the solvent on a rotary evaporator, collect the product, and combine it with the product from step 17. An additional 5% to 15% of the product can be recovered by the process outlined in steps 18 to 24. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
25. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. 3 -O-Levulinylthymidine (S.2a) is obtained in an overall combined yield of ∼85% to 95%. ◦ TLC (10% MeOH/CH2 Cl2 ): Rf = 0.40. m.p. 50◦ -52◦ C. [α]20 D = −4.7 (c 0.8, CHCl3 ). IR (KBr): υ 3449, 3065, 2927, 1706 cm−1 . 1 H NMR (MeOH-d4 , 200 MHz): δ 2.09 (d, 4 JHH = 1.3 Hz, 3H, Me), 2.39 (s, 3H, Me-Lev), 2.57 (m, 2H, H2 ), 2.80 (t, 3 JHH = 6.0 Hz, 2H, CH2 -Lev), 3.05 (t, 3 JHH = 6.2 Hz, 2H, CH2 -Lev), 4.01 (m, 2H, H5 ), 4.29 (m, 1H, H4 ), 5.02 (m, 1H, H3 ), 6.50 (dd, 3 JHH = 8.1, 6.5 Hz, 1H, H1 ), 8.04 (d, 4 JHH = 1.3 Hz, 1H, H6). 13 C NMR (CDCl3 , 75 MHz): δ 12.4 (Me), 27.8 (CH2 -Lev), 29.6 (Me-Lev), 37.1 (C2 ), 37.7 (CH2 -Lev), 62.2 (C5 ), 74.9 and 85.0 (C3 + C4 ), 85.7 (C1 ), 111.1 (C5), 136.5 (C6), 150.6 (C2), 164.3 (C4), 172.4 (C=O Lev), 206.8 (C=O Lev). ESI-MS (m/z): 363 [(M + Na)+ , 100%], 379 [(M + K)+ , 30%]. Anal. calcd. (%) for C15 H20 N2 O7 : C, 52.92; H, 5.93; N, 8.23; found: C, 53.0; H, 5.8; N, 8.2.
Preparation of O-Levulinyl Acetonoxime from Levulinic Acid and Acetonoxime O-Levulinyl acetonoxime is now commercially available form Sai Life Sciences (http://www.sailifesciences.com). The main advantage to preparing it is cost.
SUPPORT PROTOCOL 1
Additional Materials (also see Basic Protocol 1) Acetone oxime (Aldrich) Levulinic acid (Aldrich) 1,3-Dicyclohexylcarbodiimide (DCC; Aldrich) 25-L three-neck round-bottom flask equipped Overhead stirrer Thermometer Addition funnel Cooling bath 1. In a 25-L three-neck round-bottom flask equipped with an overhead stirrer, a thermometer, an addition funnel, and a stir bar, dissolve 500 g (6.84 mol) acetone oxime in 12 L anhydrous CH2 Cl2 . Cool the flask to −5◦ C using an external cooling bath.
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2. With the solution still stirring, add 794 g (6.84 mol) levulinic acid over a period of 1 hr under a nitrogen atmosphere, maintaining the internal reaction temperature at 0◦ C. 3. Dissolve 1.41 kg (6.86 mol) DCC in 3 L CH2 Cl2 and add to the reaction mixture slowly over a period of 2 hr, still at 0◦ C. After 2 hr, the reaction mixture will become heterogeneous due to the precipitation of 1,3-dicyclohexylurea, indicating the completion of the reaction.
4. Stir for an additional 2 hr at 5◦ to 10◦ C. 5. Filter off the 1,3-dicyclohexylurea under vacuum, collecting the filtrate in a filter flask. Wash the precipitate twice with 1 L CH2 Cl2 , collecting the washes in the same vessel. 6. Concentrate the combined filtrate under vacuum in a rotary evaporator to give the product as a pale yellow oil. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
7. Confirm product purity by TLC; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. O-Levulinyl acetonoxime is obtained in a yield of 1.14 kg (97%). TLC (EtOAc): Rf = 0.37. IR (NaCl): υ 1756, 1719 cm−1 . 1 H NMR (CDCl3 , 300 MHz): δ 1.99 (s, 3H, Me), 2.01 (s, 3H, Me), 2.19 (s, 3H, MeCO), 2.66 (t, 3 JHH = 6.6 Hz, 2H, CH2 -Lev), 2.82 (t, 3 JHH = 6.8 Hz, 2H, CH2 -Lev). 13 C NMR (CDCl3 , 75 MHz): δ 16.9 (Me), 21.8 (Me), 26.5 (CH2 -Lev), 29.8 (Me), 37.7 (CH2 -Lev), 163.9 (C=N), 170.4 (C=O), 206.3 (C=O). ESI-MS (m/z): 172 [(M + H)+ , 30%], 194 [(M + Na)+ , 100%]. Anal. calcd. (%) for C8 H13 NO3 : C, 56.13; H, 7.65; N, 8.18; found: C, 56.3; H, 7.4; N, 8.3. The product is visualized on the TLC plate in an iodine chamber (APPENDIX 3D). BASIC PROTOCOL 2
Preparation of N4 -Benzoyl-3 -O-Levulinyl-2 -Deoxycytidine via Regioselective Enzymatic Acylation with PSL-C Materials N4 -Benzoyl-2 -deoxycytidine (Rasayan) O-Levulinyl acetonoxime (see Support Protocol 1) Immobilized Pseudomonas cepacia lipase (PSL-C; 904 propyl laurate units per gram; Amano Enzyme) Anhydrous nitrogen Anhydrous tetrahydrofuran (THF), freshly distilled from sodium and benzophenone Methanol (MeOH), hot (65◦ C) Diethyl ether (Et2 O), cold (4◦ C) 1-L Erlenmeyer flask equipped with a three-way glass stopcock Rubber septum Orbital shaker, 40◦ C Buchner funnels with filter paper circles 500-mL round-bottom flasks Rotary evaporator Additional reagents and equipment for TLC (see Basic Protocol 1)
Enzymatic Regioselective Levulinylation of Nucleosides
Perform levulinylation 1. Add 10 g (30.19 mmol) N4 -benzoyl-2 -deoxycytidine, 15.5 g (90.57 mmol) Olevulinyl acetonoxime, and 20 g PSL-C to a 1-L Erlenmeyer flask equipped with a three-way glass stopcock.
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2. Evacuate the Erlenmeyer flask on a vacuum line, and then flush the flask with nitrogen. Repeat this procedure three times. 3. Add 302 mL dry THF. 4. Seal the flask with a rubber septum and place on an orbital shaker kept at 40◦ C. Shake the reaction mixture at 250 rpm. 5. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). At this point in the synthesis, the reaction mixture will appear as a white suspension. The levulinylation reaction will take between 14 and 16 hr to complete.
Work up and purify product 6. When the reaction is complete, filter off the lipase under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 500-mL round-bottom flask. 7. Wash the PSL-C ten times with 100 mL hot (65◦ C) MeOH. When the 500-mL flask is full, evaporate the solvent under vacuum on a rotary evaporator and continue washes. Because of the poor solubility of the product in MeOH, additional washes may be required for complete recovery.
8. Remove solvents under reduced pressure using a rotary evaporator. The excess oxime ester will impart an orange color to the residual product.
9. Filter under vacuum using a Buchner funnel with a filter paper circle. Wash the white solid (i.e., the purified product) three times with 50 mL cold Et2 O, and then dry the solid under vacuum. Since the product is relatively insoluble in Et2 O, additional extraction is not required. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
10. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. N4 -Benzoyl-3 -O-levulinyl-2 -deoxycytidine (S.2b) is obtained in 80% to 85% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.30. m.p. 189◦ -191◦ C. [α]20 D = +20.5 (c 1.0, MeOH). IR (KBr): υ 3289, 2929, 1727, 1709 cm−1 . 1 H NMR (CDCl3 , 300 MHz): δ 2.20 (s, 3H, MeLev), 2.44 (m, 1H, H2 ), 2.59 (m, 2H, CH2 -Lev), 2.73 (m, 3H, H2 + CH2 -Lev), 3.97 (m, 2H, H5 ), 4.20 (m, 1H, H4 ), 5.37 (m, 1H, H3 ), 6.26 (apparent t, 3 JHH = 6.8 Hz, 1H, H1 ), 7.58 (m, 4H, H5 + Hm + Hp ), 7.90 (apparent d, 3 JHH = 7.3 Hz, 2H, Ho ), 8.30 (d, 3 JHH = 7.4 Hz, 1H, H6). 13 C NMR (DMSO-d6 , 75 MHz): δ 27.8 (CH2 -Lev), 29.6 (Me-Lev), 37.5 (CH2 -Lev), 38.2 (C2 ), 61.2 (C5 ), 74.9 and 85.6 (C3 + C4 ), 86.4 (C1 ), 96.3 (C5), 128.5 (Co + Cm ), 132.8 (Ci ), 133.1 (Cp ), 145.0 (C6), 154.5 (PhC=O), 163.3 (C4), 167.4 (C2), 172.1 (C=O), 207.0 (C=O). ESI-MS (m/z): 430 [(M + H)+ , 30%], 452 [(M + Na)+ , 52%], 468 [(M + K)+ , 95%]. Anal. calcd. (%) for C21 H23 N3 O7 : C, 58.74; H, 5.40; N, 9.79; found: C, 58.7; H, 5.5; N, 9.8.
Preparation of N6 -Benzoyl-3 -O-Levulinyl-2 -Deoxyadenosine via Regioselective Enzymatic Acylation with PSL-C
BASIC PROTOCOL 3
Materials N6 -Benzoyl-2 -deoxyadenosine (Rasayan) O-Levulinyl acetonoxime (see Support Protocol 1) Immobilized Pseudomonas cepacia lipase (PSL-C; 904 propyl laurate units per gram; Amano Enzyme) Anhydrous nitrogen
Protection of Nucleosides for Oligonucleotide Synthesis
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Anhydrous tetrahydrofuran (THF), freshly distilled from sodium and benzophenone Dichloromethane (CH2 Cl2 ) Diethyl ether (Et2 O), cold (4◦ C) 1-L Erlenmeyer flask equipped with a three-way glass stopcock Rubber septum Orbital shaker, 40◦ C Buchner funnels with filter paper circles 500-mL round-bottom flasks Rotary evaporator Additional reagents and equipment for TLC (see Basic Protocol 1) Perform levulinylation 1. Add 10 g (28.15 mmol) N6 -benzoyl-2 -deoxyadenosine, 14.4 g (84.45 mmol) Olevulinyl acetonoxime, and 30 g PSL-C to a 1-L Erlenmeyer flask equipped with a three-way glass stopcock. 2. Evacuate the Erlenmeyer flask on a vacuum line, and then flush the flask with nitrogen. Repeat this procedure three times. 3. Add 282 mL dry THF. 4. Seal the flask with a rubber septum and place in an orbital shaker kept at 40◦ C. Shake the reaction mixture at 250 rpm. 5. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). At this point in the synthesis, the reaction mixture will appear as a white suspension. The levulinylation reaction will take between 16 and 18 hr to complete.
Work up and purify product 6. When the reaction is complete, filter off the lipase under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 500-mL round-bottom flask. 7. Wash the PSL-C three times with 50 mL CH2 Cl2 . 8. Remove solvents under reduced pressure using a rotary evaporator. The excess oxime ester will impart an orange color to the residual product.
9. Filter under vacuum using a Buchner funnel with a filter paper circle. Wash the white solid (i.e., the purified product) three times with 50 mL cold Et2 O, and then dry the solids under vacuum. Since the product is relatively insoluble in Et2 O, additional extraction is not required. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
10. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis.
Enzymatic Regioselective Levulinylation of Nucleosides
N6 -Benzoyl-3 -O-levulinyl-2 -deoxyadenosine (S.2c) is obtained in 80% to 85% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.44. m.p. 125◦ -127◦ C. [α]20 D = −26.7 (c 0.9, CHCl3 ). IR (KBr): υ 2360, 2341, 1733, 1715, 1687 cm−1 . 1 H NMR (CDCl3 , 200 MHz): δ 2.22 (s, 3H, Me-Lev), 2.61 (m, 3H, CH2 -Lev + H2 ), 2.82 (m, 2H, CH2 -Lev), 3.18 (m, 1H, H2 ), 3.97 (m, 2H, H5 ), 4.31 (s, 1H, H4 ), 5.57 (apparent d, 3 JHH = 5.4 Hz, 1H, H3 ), 6.38 (dd, 3 JHH = 9.8, 5.2 Hz, 1H, H1 ), 7.57 (m, 3H, Hm + Hp ), 8.05 (m, 2H, Ho ), 8.17 (s, 1H, H2 or H8), 8.79 (s, 1H, H8 or H2). 13 C NMR (CDCl3 , 75 MHz): δ 27.7 (CH2 Lev), 29.6 (Me-Lev), 37.6 (CH2 -Lev + C2 ), 62.8 (C5 ), 76.1 and 86.8 (C3 + C4 ), 87.0
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(C1 ), 124.3 (C5), 127.8 and 128.6 (Co + Cm ), 132.7 (Cp ), 133.2 (Ci ), 142.3 (C2 or C8), 150.1 (C4), 150.6 (C6), 151.9 (C8 or C2), 164.8 (PhC=O), 172.1 (C=O), 206.4 (C=O). ESI-MS (m/z): 454 [(M + H)+ , 20%], 476 [(M + Na)+ , 33%], 492 [(M + K)+ , 5%]. Anal. calcd. (%) for C22 H23 N5 O6 : C, 58.27; H, 5.11; N, 15.44; found: C, 58.2; H, 5.2; N, 15.5.
Preparation of N2 -Isobutyryl-3 -O-Levulinyl-2 -Deoxyguanosine via Regioselective Enzymatic Hydrolysis with CAL-B
BASIC PROTOCOL 4
Materials N2 -Isobutyryl-3 ,5 -di-O-levulinyl-2 -deoxyguanosine (S.4d; see Support Protocol 2) 1,4-Dioxane 0.15 M KH2 PO4 buffer (see recipe) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram [PLU/g]; Novo Nordisk) Dichloromethane (CH2 Cl2 ) Saturated sodium bicarbonate (NaHCO3 ) solution Chloroform (CHCl3 ) Sodium sulfate (Na2 SO4 ) Diethyl ether (Et2 O) 500-mL Erlenmeyer flask with rubber septum Orbital shaker, 40◦ C Buchner funnel with filter paper circles 500-mL round-bottom flasks Rotary evaporator 500-mL separatory funnel 250-mL Erlenmeyer flasks Glass funnel with Whatman no. 1 filter paper Additional reagents and equipment for TLC (see Basic Protocol 1) Perform regioselective hydrolysis 1. Add 5 g (9.37 mmol) N2 -isobutyryl-3 ,5 -di-O-levulinyl-2 -deoxyguanosine, 16.4 mL 1,4-dioxane, 77.4 mL of 0.15 M KH2 PO4 buffer, and 5 g CAL-B to a 500-mL Erlenmeyer flask, and then seal the flask with a rubber septum. 2. Place the flask on an orbital shaker kept at 40◦ C. Shake the reaction mixture at 250 rpm. 3. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5), but using 20% (v/v) MeOH/CH2 Cl2 as the eluent. After spotting, it will be necessary to dry the TLC plate with a heat gun to evaporate the water in the sample. At this point in the synthesis, the reaction mixture will be clear. The hydrolysis reaction typically takes between 13 and 15 hr to complete.
Work up and purify product 4. When the reaction is complete, filter off the lipase under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 500-mL round-bottom flask. 5. Wash the CAL-B twice with 50 mL CH2 Cl2 , collecting washes in the same flask. 6. Remove solvents under reduced pressure using a rotary evaporator.
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7. Dissolve the residue in 100 mL CH2 Cl2 and transfer to a 500-mL separatory funnel. 8. Extract the organic layer four times with 25 mL saturated NaHCO3 solution, using two 250-mL Erlenmeyer flasks to separate the phases. Monitor the progress of the extraction by performing analytical TLC on both phases. N2 -Isobutyryl-2 -deoxyguanosine and levulinic acid are extracted into the aqueous phase. Additional extractions may be needed for complete removal from the organic phase. Alternatively, brine can be used to separate the phases effectively.
9. Pool the aqueous phases and recover product from the aqueous phase by extracting four times with 25 mL CH2 Cl2 and four times with 25 mL CHCl3 , collecting the combined organic phases in a 250-mL Erlenmeyer flask. 10. Pool the organic phases from step 9 and verify by TLC that they contain no residual N2 -isobutyryl-2 -deoxyguanosine. If any is present, re-extract with 20 mL saturated NaHCO3 solution. Re-extraction is unnecessary if N2 -isobutyryl-2 -deoxyguanosine is already absent from the organic phase.
11. Pool the organic phases from steps 8 and 10 and dry over 4 g Na2 SO4 for 10 to 15 min. Remove the drying agent by gravity filtration through Whatman no. 1 filter paper in a glass funnel, and collect the filtrate in a 500-mL round-bottom flask. 12. Wash the Na2 SO4 four times with 25 mL CH2 Cl2 , and collect all washes in the same round-bottom flask. 13. Evaporate solvents under reduced pressure using a rotary evaporator. 14. Add 10 mL Et2 O and triturate the product. Evaporate the solvent on a rotary evaporator and collect the product. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
15. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. N2 -Isobutyryl-3 -O-levulinyl-2 -deoxyguanosine (S.2d) is obtained in ∼70% to 75% yield. TLC (20% MeOH/CH2 Cl2 ): Rf = 0.75. m.p. 170◦ -172◦ C. [α]20 D = +1.0 (c 1.1, DMSO). IR (KBr): υ 3415, 2961, 2929, 2859, 1725, 1686, 1614 cm−1 . 1 H NMR (MeOH-d4 , 200 MHz): δ 1.41 (d, 3 JHH = 6.8 Hz, 6H, Me-iBu), 2.38 (s, 3H, Me-Lev), 2.70-3.09 (m, 7H, 2CH2 -Lev + 2H2 + CH-iBu), 3.98 (d, 3 JHH = 3.6 Hz, 2H, 2H5 ), 4.32 (m, 1H, H4 ), 5.57 (m, 1H, H3 ), 6.51 (dd, 3 JHH = 6.1, 8.0 Hz, 1H, H1 ), 8.43 (s, 1H, H8). 13 C NMR (MeOHd4 , 50 MHz): δ 19.3 (Me-iBu), 28.9 (CH2 -Lev), 29.7 (Me-Lev), 36.9 (CH-iBu), 38.6 and 39.0 (C2 + CH2 -Lev), 63.0 (C5 ), 76.6 and 85.6 (C3 + C4 ), 87.0 (C1 ), 121.2 (C5), 139.6 (C8), 149.7 and 150.2 (C2 + C4), 157.3 (C6), 173.9 (C=O), 181.7 (iBu-C=O), 209.5 (C=O). ESI-MS (m/z): 436 [(M + H)+ , 15%], 458 [(M + Na)+ , 50%]. Anal. calcd. (%) for C19 H25 N5 O7 : C, 52.41; H, 5.79; N, 16.08; found: C, 52.6; H, 5.8; N, 16.2. SUPPORT PROTOCOL 2
Enzymatic Regioselective Levulinylation of Nucleosides
Preparation of N2 -Isobutyryl-3 ,5 -Di-O-Levulinyl-2 -Deoxyguanosine Additional Materials (also see Basic Protocol 1) N2 -Isobutyryl-2 -deoxyguanosine (Rasayan) Triethylamine (Et3 N) Anhydrous 1,4-dioxane, freshly distilled from sodium and benzophenone Levulinic acid 1,3-Dicyclohexylcarbodiimide (DCC) 4-(Dimethylamino)pyridine (DMAP; Aldrich)
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Perform levulinylation 1. Add a magnetic stir bar and 5.0 g (14.82 mmol) N2 -isobutyryl-2 -deoxyguanosine to a 500-mL round-bottom flask equipped with a three-way glass stopcock. Seal the flask with a rubber septum. 2. Evacuate the flask on a vacuum line, and then flush the flask with nitrogen. Repeat this procedure three times. 3. Add 12.60 mL (88.92 mmol) Et3 N, 150 mL dry 1,4-dioxane, 7.9 mL (77.06 mmol) levulinic acid, 15.86 g (77.06 mmol) DCC, and 148 mg (1.19 mmol) DMAP. Close the three-way glass stopcock and place the flask on a magnetic stirrer. 4. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5), but using 20% (v/v) MeOH/CH2 Cl2 as the eluent. The levulinylation reaction is complete in 2 to 3 hr. After 2 hr the reaction mixture becomes heterogeneous due to the precipitation of 1, 3-dicyclohexylurea (DCU).
Work up and purify product 5. When the reaction is complete, filter the suspension under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 500-mL round-bottom flask. This filtration step is performed to remove the DCU formed during the reaction.
6. Wash DCU twice with 50 mL CH2 Cl2 , collecting washes in the same round-bottom flask. 7. Remove solvents under reduced pressure using a rotary evaporator. The product that results will be an orange oil.
8. Dissolve the residue in 50 mL CH2 Cl2 and transfer to a 500-mL separatory funnel. 9. Remove levulinic acid by extracting the organic layer two to three times with 20 mL saturated NaHCO3 solution. Separate the layers into two 250-mL Erlenmeyer flasks, and verify that all levulinic acid has been removed from the organic phase by performing analytical TLC on both phases. Additional extractions may be needed for complete removal. Alternatively, brine can be used to clarify both phases.
10. Dry the organic layer over 4 g Na2 SO4 for 10 to 15 min, and then remove the drying agent by gravity filtration using a glass funnel and Whatman no. 1 filter paper. Collect the filtrate in a 500-mL round-bottom flask. 11. Wash the Na2 SO4 four times with 25 mL CH2 Cl2 , and collect all washes in the same round-bottom flask. 12. Evaporate solvents under reduced pressure using a rotary evaporator. The product that results will again be an orange oil.
13. To recover product by dry column chromatography, weigh out 100 g silica gel and place in a Buchner funnel with a filter paper circle, attached to a 1-L filter flask connected to a vacuum line. Wash the silica with 200 mL of 25% (v/v) Et2 O/EtOAc. 14. Using a Pasteur pipet, spread the residue carefully on top of the silica bed. 15. Elute any residual 1,3-dicyclohexyl-1-levulinylurea with ∼500 to 800 mL of 25% (v/v) Et2 O/EtOAc.
Protection of Nucleosides for Oligonucleotide Synthesis
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16. Using clean filter flasks to collect 100-mL fractions, elute the desired product with ∼800 to 1200 mL of 10% (v/v) MeOH/CH2 Cl2 . Monitor fractions by analytical TLC. 17. Transfer fractions containing pure product to a 500-mL round-bottom flask, and remove solvents under reduced pressure using a rotary evaporator. If desired, solvents can be recovered by distillation.
18. Add 30 mL Et2 O and triturate the product. Evaporate the solvent on a rotary evaporator and collect the pale yellow product. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
19. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. N2 -Isobutyryl-3 ,5 -di-O-levulinyl-2 -deoxyguanosine (S.4d) is obtained in ∼85% to 90% yield. TLC (20% MeOH/CH2 Cl2 ): Rf = 0.85. m.p. 45◦ -47◦ C. [α]20 D = −8.8 (c 1.0, DMSO). IR (KBr): υ 3413, 2935, 1740, 1714, 1680, 1613 cm−1 . 1 H NMR (DMSO-d6 , 200 MHz): δ 1.23 (d, 3 JHH = 6.5 Hz, 6H, Me-iBu), 2.15 (s, 3H, Me-Lev), 2.20 (s, 3H, Me-Lev), 2.55-3.19 (several m, 11H, 4CH2 -Lev + 2H2 + CH-iBu), 4.32 (m, 3H, H4 + 2H5 ), 5.35 (m, 1H, H3 ), 6.35 (apparent t, 3 JHH = 7.2 Hz, 1H, H1 ), 8.35 (s, 1H, H8), 11.80 (br s, 1H, NH), 12.20 (br s, 1H, NH). 13 C NMR (DMSO-d6 , 50 MHz): δ 18.86 (Me-iBu), 18.91 (Me-iBu), 27.5 (CH2 -Lev), 27.6 (CH2 -Lev), 29.5 (Me-Lev), 29.6 (Me-Lev), 34.8 (CH-iBu), 35.5 (C2 ), 37.38 (CH2 -Lev), 37.45 (CH2 -Lev), 63.7 (C5 ), 74.6 and 81.7 (C3 + C4 ), 82.9 (C1 ), 120.3 (C5), 137.3 (C8), 148.3 and 148.7 (C2 + C4), 154.8 (C6), 172.1 (C=O), 172.2 (C=O), 180.2 (iBu-C=O), 206.9 (C=O), 207.1 (C=O). ESI-MS (m/z): 534 [(M + H)+ , 100%], 556 [(M + Na)+ , 60%], 572 [(M + K)+ , 27%]. Anal. calcd. (%) for C24 H31 N5 O9 : C, 54.01; H, 5.86; N, 13.13; found: C, 54.2; H, 5.8; N, 13.2.
SYNTHESIS OF 5 -O-LEVULINYL-2 -DEOXYRIBONUCLEOSIDES The preparation of 5 -O-levulinyl-2 -deoxyribonucleosides consists of regioselective acylation of the 5 -hydroxyl function of deoxyribonucleosides with O-levulinyl acetonoxime using Candida antarctica lipase B (CAL-B) as illustrated in Figure 2.11.1. This is a general procedure for the T, C, A, and G monomers. BASIC PROTOCOL 5
Preparation of 5 -O-Levulinylthymidine via Regioselective Enzymatic Acylation with CAL-B Materials Thymidine (Rasayan) O-Levulinyl acetonoxime (see Support Protocol 1) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram; Novo Nordisk) Anhydrous nitrogen Anhydrous tetrahydrofuran (THF), freshly distilled from sodium and benzophenone 1-L Erlenmeyer flask equipped with a three-way glass stopcock Rubber septum Orbital shaker, 10◦ C Additional reagents and equipment for analysis and work up of regioselectively levulinated thymidine (see Basic Protocol 1)
Enzymatic Regioselective Levulinylation of Nucleosides
Perform levulinylation 1. Add 10 g (41.28 mmol) thymidine, 21.2 g (123.86 mmol) O-levulinyl acetonoxime, and 10 g CAL-B to a 1-L Erlenmeyer flask equipped with a three-way glass stopcock.
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2. Evacuate the Erlenmeyer flask on a vacuum line and then flush with nitrogen. Repeat this procedure three times. 3. Add 206 mL dry THF. 4. Seal the flask with a rubber septum and place in an orbital shaker kept at 10◦ C. Shake the reaction mixture at 250 rpm. 5. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). The reaction mixture containing immobilized lipase will remain clear throughout the course of the reaction, which takes 5 to 7 hr to complete.
Work up and purify product 6. Work up, purify, and recover the product as described for compound S.2a (see Basic Protocol 1, steps 6 to 24). The initial isolated yield (steps 6 to17) is ∼50% to 60%, and the recovery process (steps 18 to 24) yields another ∼15% to 25%. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
7. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. 5 -O-Levulinylthymidine (S.3a) is obtained in an overall combined yield of ∼75% to 80%. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.22. m.p. 141◦ -143◦ C. [α]20 D = +21.5 (c 0.5, MeOH). IR (KBr): υ 3393, 3215, 2934, 1737, 1724, 1643, 1629 cm−1 . 1 H NMR (DMSOd6 , 200 MHz): δ 1.91 (s, 3H, Me), 2.27 (s, 3H, Me-Lev), 2.30 (m, 2H, H2 ), 2.66 (m, 2H, CH2 -Lev), 2.89 (t, 3 JHH = 6.2 Hz, 2H, CH2 -Lev), 4.07 (m, 1H, H4 ), 4.35 (m, 3H, H3 + 2H5 ), 5.55 (d, J = 4.8 Hz, 1H, OH), 6.32 (t, 3 JHH = 7.0 Hz, 1H, H1 ), 7.6 (s, 1H, H6), 11.45 (s, 1H, NH). 13 C NMR (DMSO-d6 , 50 MHz): δ 12.02 (Me), 27.4 (CH2 -Lev), 29.4 (Me-Lev), 37.2 (C2 ), 38.4 (CH2 -Lev), 63.8 (C5 ), 70.1 and 83.5 (C3 + C4 ), 83.6 (C1 ), 109.7 (C5), 135.7 (C6), 150.3 (C4), 163.6 (C2), 172.1 (C=O Lev), 206.7 (C=O Lev). ESI-MS (m/z): 341 [(M + H)+ , 40%], 363 [(M + Na)+ , 100%], 379 [(M + K)+ , 80%]. Anal. calcd. (%) for C15 H20 N2 O7 : C, 52.92; H, 5.93; N, 8.23; found: C, 52.7; H, 5.8; N, 8.2.
Preparation of N4 -Benzoyl-5 -O-Levulinyl-2 -Deoxycytidine via Regioselective Enzymatic Acylation with CAL-B
BASIC PROTOCOL 6
Materials N4 -Benzoyl-2 -deoxycytidine (Rasayan) O-Levulinyl acetonoxime (see Support Protocol 1) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram; Novo Nordisk) Anhydrous nitrogen Anhydrous tetrahydrofuran (THF), freshly distilled over sodium and benzophenone Dichloromethane (CH2 Cl2 ) Methanol (MeOH) Diethyl ether (Et2 O), cold (4◦ C) 1-L Erlenmeyer flask equipped with a three-way glass stopcock Rubber septum Orbital shaker, 30◦ C Buchner funnels with filter paper circles 500-mL round-bottom flask
Protection of Nucleosides for Oligonucleotide Synthesis
2.11.13 Current Protocols in Nucleic Acid Chemistry
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Rotary evaporator 1-L filter flask 250-mL Erlenmeyer flask Horst heating mantle Additional reagents and equipment for TLC (see Basic Protocol 1) Perform levulinylation 1. Add 10 g (30.19 mmol) N4 -benzoyl-2 -deoxycytidine, 15.5 g (90.57 mmol) Olevulinyl acetonoxime, and 10 g CAL-B to a 1-L Erlenmeyer flask equipped with a three-way glass stopcock. 2. Evacuate the Erlenmeyer flask on a vacuum line and then flush with nitrogen. Repeat this procedure three times. 3. Add 302 mL dry THF. 4. Seal the flask with a rubber septum and place in an orbital shaker kept at 30◦ C. Shake the reaction mixture at 250 rpm. 5. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). The reaction mixture is a white suspension, and the reaction is complete in 16 to 18 hr.
Work up and purify product 6. When the reaction is complete, filter off the lipase under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 500-mL round-bottom flask. 7. Wash the CAL-B twice with 50 mL CH2 Cl2 and then twice with 50 mL MeOH. Because of the poor solubility of the product in MeOH, additional washings may be required for complete recovery.
8. Remove solvents under reduced pressure using a rotary evaporator. The excess oxime ester will impart an orange color to the product.
9. Dissolve the residue in 20 mL CH2 Cl2 and precipitate the product from 40 to 60 mL Et2 O as a white solid. Cool the precipitate at 4◦ C overnight. 10. Transfer the precipitate to a Buchner funnel with a filter paper circle, attached to a 1-L filter flask connected to a vacuum line. Wash twice with 50 mL cold (4◦ C) Et2 O, and then dry the white solid under vacuum. Since the levulinylation reaction goes to completion, no further extractions are required.
11. Crystallize the product from 50 to 60 mL MeOH using a Horst heating mantle, and collect the filtrate in a 250-mL Erlenmeyer flask. This separates the 5 -acylated product from its 3 -regioisomer (5% to 10% of the crude product).
12. Collect the solid by vacuum filtration through a Buchner funnel with a filter paper circle, and then wash twice with 40 mL cold Et2 O. Dry the white solid under vacuum. Et2 O removes traces of MeOH and dissolves the undesired 3 -acylated regioisomer. Enzymatic Regioselective Levulinylation of Nucleosides
If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
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13. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. N4 -Benzoyl-5 -O-levulinyl-2 -deoxycytidine (S.3b) is obtained in ∼70% to 75% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.37. m.p. 50◦ -52◦ C. [α]20 D = +71.1 (c 0.5, MeOH). IR (KBr): υ 3410, 2919, 1738, 1701, 1650 cm−1 . 1 H NMR (CDCl3 , 300 MHz): δ 2.20 (s, 3H, Me-Lev), 2.25 (m, 1H, H2 ), 2.58 (m, 2H, CH2 -Lev), 2.75 (m, 1H, H2 ), 2.82 (m, 2H, CH2 -Lev), 3.35 (s, 1H, OH), 4.25 (m, 1H, H3 ), 4.40 (m, 3H, 2H5 + H4 ), 6.30 (apparent t, 3 JHH = 6.2 Hz, 1H, H1 ), 7.55 (m, 4H, H5 + 2Hm + Hp ), 7.90 (apparent d, 3 JHH = 7.1 Hz, 2H, Ho ), 8.20 (d, 3 JHH = 7.4 Hz, 1H, H6), 8.78 (s, 1H, NH). 13 C NMR (CDCl3 , 75 MHz): δ 27.7 (CH2 -Lev), 29.6 (Me-Lev), 37.7 (CH2 -Lev), 41.3 (C2 ), 63.7 (C5 ), 70.6 and 84.8 (C3 + C4 ), 87.4 (C1 ), 96.8 (C5), 127.6 and 128.7 (Co + Cm ), 132.8 (Ci ), 133.0 (Cp ), 144.2 (C6), 155.1 (C2), 162.4 (C4), 166.7 (PhC=O), 172.6 (C=O), 206.8 (C=O). ESI-MS (m/z): 430 [(M + H)+ , 20%], 452 [(M + Na)+ , 65%], 468 [(M + K)+ , 40%]. Anal. calcd. (%) for C21 H23 N3 O7 : C, 58.74; H, 5.40; N, 9.79; found: C, 58.8; H, 5.6; N, 9.7.
Preparation of N6 -Benzoyl-5 -O-Levulinyl-2 -Deoxyadenosine via Regioselective Enzymatic Acylation with CAL-B
BASIC PROTOCOL 7
Materials N6 -Benzoyl-2 -deoxyadenosine (Rasayan) O-Levulinyl acetonoxime (see Support Protocol 1) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram; Novo Nordisk) Anhydrous nitrogen Anhydrous tetrahydrofuran (THF), freshly distilled over sodium and benzophenone Methanol (MeOH), hot (65◦ C) Diethyl ether (Et2 O), cold (4◦ C) 1-L Erlenmeyer flask equipped with a three-way glass stopcock Rubber septum Orbital shaker, 30◦ C Buchner funnels with filter paper circles 500-mL round-bottom flask Rotary evaporator 1-L filter flask Additional reagents and equipment for TLC (see Basic Protocol 1) Perform levulinylation 1. Add 10 g (28.15 mmol) N6 -benzoyl-2 -deoxyadenosine, 14.4 g (84.45 mmol) Olevulinyl acetonoxime, and 10 g CAL-B to a 1-L Erlenmeyer flask equipped with a three-way glass stopcock. 2. Evacuate the Erlenmeyer flask on a vacuum line and flush with nitrogen. Repeat this procedure three times. 3. Add 140 mL dry THF. 4. Seal the flask with a rubber septum and place in an orbital shaker kept at 30◦ C. Shake the reaction mixture at 250 rpm. 5. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). The reaction mixture will remain a white suspension and the reaction is complete in 24 to 26 hr.
Protection of Nucleosides for Oligonucleotide Synthesis
2.11.15 Current Protocols in Nucleic Acid Chemistry
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Work up and purify product 6. When the reaction is complete, filter off the lipase under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 500-mL round-bottom flask. 7. Wash the CAL-B five times with 100 mL hot (65◦ C) MeOH. When the flask becomes full, evaporate the solvent under vacuum on a rotary evaporator and continue washes. Because of the poor solubility of the product in MeOH, additional washes may be required for complete recovery.
8. Remove solvents under reduced pressure using a rotary evaporator. The excess oxime ester will impart an orange color to the product.
9. Transfer the solid to a Buchner funnel with a filter paper circle, attached to a 1-L filter flask connected to a vacuum line, and then triturate and wash the solid three times with 50 mL cold (4◦ C) Et2 O. Dry the white solid under vacuum. Since the levulinylation reaction goes to completion, no further extractions are required. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
10. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. N6 -Benzoyl-5 -O-levulinyl-2 -deoxyadenosine (S.3c) is obtained in 70% to 75% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.50. m.p. 137◦ -139◦ C. [α]20 D = −5.9 (c 1.1, MeOH). IR (KBr): υ 3413, 2959, 2928, 1726, 1637, 1616 cm−1 . 1 H NMR (DMSO-d6 , 300 MHz): δ 2.06 (s, 3H, Me-Lev), 2.45 (m, 3H, CH2 -Lev + H2 ), 2.66 (m, 2H, CH2 -Lev), 2.90 (m, 1H, H2 ), 4.03 (m, 1H, H4 ), 4.20 (m, 2H, 2H5 ), 4.51 (m, 1H, H3 ), 5.55 (s, 1H, OH), 6.50 (apparent t, 3 JHH = 6.5 Hz, 1H, H1 ), 7.58 (m, 3H, 2Hm + Hp ), 8.03 (m, 2H, 2Ho ), 8.69 (s, 1H, H2 or H8), 8.74 (s, 1H, H8 or H2), 11.20 (s, 1H, NH). 13 C NMR (DMSO-d6 , 75 MHz): δ 27.7 (CH2 -Lev), 29.8 (Me-Lev), 37.6 and 38.7 (CH2 -Lev + C2 ), 64.2 (C5 ), 70.8 and 83.8 (C3 + C4 ), 84.5 (C1 ), 126.1 (C5), 128.7 and 128.8 (Co + Cm ), 132.7 (Cp ), 133.6 (Ci ), 143.4 (C2 or C8), 150.7 (C4), 151.9 (C8 or C2), 152.1 (C6), 165.9 (PhC=O), 172.5 (C=O), 207.1 (C=O). ESI-MS (m/z): 476 [(M + Na)+ , 100%], 492 [(M + K)+ , 53%]. Anal. calcd. (%) for C22 H23 N5 O6 : C, 58.27; H, 5.11; N, 15.44; found: C, 58.4; H, 4.9; N, 15.3. BASIC PROTOCOL 8
Preparation of N2 -Isobutyryl-5 -O-Levulinyl-2 -Deoxyguanosine via Regioselective Enzymatic Acylation with CAL-B Materials
Enzymatic Regioselective Levulinylation of Nucleosides
N2 -Isobutyryl-2 -deoxyguanosine (Rasayan) O-Levulinyl acetonoxime (see Support Protocol 1) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram; Novo Nordisk) Anhydrous nitrogen Anhydrous tetrahydrofuran (THF), freshly distilled over sodium and benzophenone Methanol (MeOH) Dichloromethane (CH2 Cl2 ) Saturated sodium bicarbonate (NaHCO3 ) solution Chloroform (CHCl3 ) Sodium sulfate (Na2 SO4 ) Silica gel Ethyl acetate (EtOAc) Diethyl ether (Et2 O)
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1-L Erlenmeyer flask equipped with a three-way glass stopcock Rubber septum Orbital shaker, 40◦ C Buchner funnels with filter paper circles 500-mL round-bottom flasks Rotary evaporator 500-mL separatory funnel Glass funnel with Whatman no. 1 filter paper 1-L filter flask Additional reagents and equipment for TLC (see Basic Protocol 1) Perform levulinylation 1. Add 10 g (29.64 mmol) N2 -isobutyryl-2 -deoxyguanosine, 15.2 g (88.94 mmol) Olevulinyl acetonoxime, and 10 g CAL-B to a 1-L Erlenmeyer flask equipped with a three-way glass stopcock. 2. Evacuate the Erlenmeyer flask on a vacuum line and flush with nitrogen. Repeat this procedure three times. 3. Add 148 mL dry THF. 4. Seal the flask with a rubber septum and place in an orbital shaker kept at 40◦ C. Shake the reaction mixture at 250 rpm. 5. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5), but using 20% (v/v) MeOH/CH2 Cl2 as the eluent. At this point, the reaction mixture will be clear. The levulinylation reaction usually takes 16 to 18 hr to complete.
Work up and purify product 6. When the reaction is complete, filter off the lipase under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 500-mL round-bottom flask. 7. Wash the CAL-B six times with 50 mL MeOH. 8. Remove solvents under reduced pressure using a rotary evaporator. The product is an orange-colored oil due to the presence of excess oxime ester.
9. Dissolve the residue in 100 mL CH2 Cl2 and transfer to a 500-mL separatory funnel. 10. Extract the organic layer four times with 25 mL saturated NaHCO3 solution, using two 250-mL Erlenmeyer flasks to separate the phases. Monitor the progress of the extraction by performing analytical TLC on both phases. N2 -Isobutyryl-2 -deoxyguanosine is extracted into the aqueous phase. Additional extractions may be needed for complete removal from the organic phase. Alternatively, brine can be used to separate the phases effectively.
11. Pool the aqueous phases and recover product from the aqueous phase by extracting four times with 25 mL CH2 Cl2 and four times with 25 mL CHCl3 . 12. Pool the organic phases from step 11 and verify by TLC that they contain no residual N2 -isobutyryl-2 -deoxyguanosine. If any is present, re-extract with 20 mL saturated NaHCO3 solution. 2
Re-extraction is unnecessary if N -isobutyryl-2 -deoxyguanosine is already absent from the organic phase.
Protection of Nucleosides for Oligonucleotide Synthesis
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13. Pool the organic phases from steps 10 and 12 and dry over 4 g Na2 SO4 for 10 to 15 min. Remove the drying agent by gravity filtration through Whatman no. 1 filter paper in a glass funnel, and collect the filtrate in a 500-mL round-bottom flask. 14. Wash the Na2 SO4 four times with 25 mL CH2 Cl2 , and collect all washes in the same round-bottom flask. 15. Evaporate solvents under reduced pressure using a rotary evaporator. The product is an orange oil, which fails to precipitate from Et2 O, perhaps due to oxime ester contamination. Separation by chromatography is therefore necessary.
16. To recover product by dry column chromatography, weigh out 100 g silica gel and place in a Buchner funnel with a filter paper circle, attached to a 1-L filter flask connected to a vacuum line. Wash the silica with 200 mL of 25% (v/v) EtOAc/Et2 O. 17. Using a Pasteur pipet, spread the residue carefully on top of the silica bed. 18. Elute the residual oxime ester with ∼1500 to 1800 mL of 25% (v/v) EtOAc/Et2 O. 19. Using clean filter flasks to collect 100-mL fractions, elute the desired product with ∼1700 to 2200 mL of 10% (v/v) MeOH/CH2 Cl2 . Monitor fractions by analytical TLC. 20. Transfer fractions containing pure levulinylated product to a 500-mL round-bottom flask, and remove solvents under reduced pressure using a rotary evaporator. If desired, solvents can be recovered by distillation.
21. Add 10 mL Et2 O and triturate the product. Evaporate the solvent on a rotary evaporator and collect the product. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
22. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. N2 -Isobutyryl-5 -O-levulinyl-2 -deoxyguanosine (S.3d) is obtained in 75% to 80% yield. TLC (20% MeOH/CH2 Cl2 ): Rf = 0.60. m.p. 45◦ -47◦ C. [α]20 D = +3.0 (c 1.0, MeOH). IR (KBr): υ 3415, 2930, 1720, 1685 cm−1 . 1 H NMR (MeOH-d4 , 300 MHz): δ 1.42 (d, 3 JHH = 6.8 Hz, 6H, Me-iBu), 2.32 (s, 3H, Me-Lev), 2.59-3.07 (m, 7H, 2CH2 -Lev + 2H2 + CHiBu), 4.32 (m, 1H, H4 ), 4.50 (m, 2H, H5 ), 4.75 (m, 1H, H3 ), 6.53 (apparent t, 3 JHH = 6.5 Hz, 1H, H1 ), 8.33 (s, 1H, H8). 13 C NMR (MeOH-d4 , 75 MHz): δ 19.7 (Me-iBu), 29.0 (CH2 -Lev), 29.9 (Me-Lev), 37.2 (CH-iBu), 38.9 and 41.1 (C2 + CH2 -Lev), 65.3 (C5 ), 72.6 and 86.1 (C3 + C4 ), 86.5 (C1 ), 121.8 (C5), 139.8 (C8), 150.0 and 150.5 (C2 + C4), 157.8 (C6), 174.5 (C=O), 182.0 (iBu-C=O), 209.7 (C=O). ESI-MS (m/z): 436 [(M + H)+ , 20%], 458 [(M + Na)+ , 100%], 474 [(M + K)+ , 50%]. Anal. calcd. (%) for C19 H25 N5 O7 : C, 52.41; H, 5.79; N, 16.08; found: C, 52.3; H, 5.8; N, 16.2.
SYNTHESIS OF 3 -O-LEVULINYL-2 -O-METHYLRIBONUCLEOSIDES Because the enzymatic acylation of 2 -O-methyl derivatives with PSL-C is not regioselective, 3 -O-levulinyl-2 -O-methylribonucleosides are prepared via regioselective enzymatic hydrolysis of 3 ,5 -di-O-levulinyl-2 -O-methylribonucleosides with CAL-B as illustrated in Figure 2.11.3. This is a general procedure for the U, C, A, and G monomers. Synthesis of the 3 ,5 -di-O-levulinyl-2 -O-methylribonucleosides using levulinic acid is presented in Support Protocol 3. Enzymatic Regioselective Levulinylation of Nucleosides
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Figure 2.11.3 Regioselective hydrolysis of 3 ,5 -di-O-levulinyl-2 -O-methylribonucleosides (S.6a-d) using CAL-B. Abbreviations: ABz , N 6 -benzoyladenin-9-yl; CBz , N 4 -benzoylcytosin-1-yl; DCC, 1,3-dicyclohexylcarbodiimide; DMAP, 4-(dimethylamino)pyridine; Et3 N, triethylamine; Gi-Bu , N 2 -isobutyrylguanin-9-yl; Lev, levulinyl; LevOH, levulinic acid; U, uracil-1-yl.
Preparation of 3 -O-Levulinyl-2 -O-Methyluridine via Regioselective Enzymatic Hydrolysis with CAL-B
BASIC PROTOCOL 9
Materials 3 ,5 -Di-O-levulinyl-2 -O-methyluridine (S.6a; see Support Protocol 3) 1,4-Dioxane 0.15 M KH2 PO4 buffer (see recipe) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram; Novo Nordisk) 500-mL Erlenmeyer flask Rubber septum Orbital shaker, 30◦ C Additional reagents and equipment for TLC (see Basic Protocol 1) and for workup and purification of the product (see Basic Protocol 4) 1. Add 1 g (2.20 mmol) 3 ,5 -di-O-levulinyl-2 -O-methyluridine, 3.9 mL 1,4-dioxane, 18.2 mL of 0.15 M KH2 PO4 buffer, and 1 g CAL-B to a 500-mL Erlenmeyer flask. 2. Seal the flask with a rubber septum and place on an orbital shaker kept at 30◦ C. Shake the reaction mixture at 250 rpm. 3. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). After spotting, it will be necessary to dry the TLC plate with a heat gun to evaporate the water in the sample. The reaction mixture remains a white suspension and the reaction is usually complete in 9 to 11 hr.
4. Work up and purify the product as described for compound S.2d (see Basic Protocol 4, steps 4 to 14). If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months. Protection of Nucleosides for Oligonucleotide Synthesis
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5. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR analysis; mass spectrometry; and elemental analysis. 3 -O-Levulinyl-2 -O-methyluridine (S.7a) is obtained in 75% to 80% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.32. m.p. 106◦ -108◦ C. [α]20 D = −2.6 (c 1.0, DMSO). IR (NaCl): υ 3452, 2934, 1693 cm−1 . 1 H NMR (DMSO-d6 , 300 MHz): δ 2.10 (s, 3H, Me-Lev), 2.54 (m, 2H, CH2 -Lev), 2.73 (m, 2H, CH2 -Lev), 3.24 (s, 3H, OMe), 3.59 (m, 2H, H5 ), 4.03 (m, 2H, H2 + H4 ), 5.23 (m, 1H, H3 ), 5.34 (br s, 1H, OH), 5.70 (d, 3 JHH = 8.0 Hz, 1H, H5), 5.87 (d, 3 JHH = 6.5 Hz, 1H, H1 ), 7.89 (d, 3 JHH = 8.2 Hz, 1H, H6). 13 C NMR (DMSO-d6 , 75 MHz): δ 31.7 (CH2 -Lev), 33.6 (Me-Lev), 41.4 (CH2 -Lev), 62.1 (OMe), 64.8 (C5 ), 74.9, 84.6, 87.2, 89.6 (C1 + C2 + C3 + C4 ), 106.6 (C5), 144.3 (C6), 154.7 (C4), 167.0 (C2), 175.8 (C=O), 210.7 (C=O). ESI-MS (m/z): 357 [(M + H)+ , 25%], 379 [(M + Na)+ , 100%], 395 [(M + K)+ , 21%]. Anal. calcd. (%) for C15 H20 N2 O8 : C, 50.56; H, 5.66; N, 7.86; found: C, 50.8; H, 5.9; N, 7.7. BASIC PROTOCOL 10
Preparation of N4 -Benzoyl-3 -O-Levulinyl-2 -O-Methylcytidine via Regioselective Enzymatic Hydrolysis with CAL-B Materials N4 -Benzoyl-3 ,5 -di-O-levulinyl-2 -O-methylcytidine (S.6b; see Support Protocol 3) 1,4-Dioxane 0.15 M KH2 PO4 buffer (see recipe) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram; Novo Nordisk) 500-mL Erlenmeyer flask Rubber septum Orbital shaker, 30◦ C Additional reagents and equipment for TLC (see Basic Protocol 1) and for workup and purification of the product (see Basic Protocol 4) 1. Add 1 g (1.79 mmol) N4 -benzoyl-3 ,5 -di-O-levulinyl-2 -O-methylcytidine, 3.6 mL 1,4-dioxane, 14.3 mL of 0.15 M KH2 PO4 buffer, and 1 g CAL-B to a 500-mL Erlenmeyer flask. 2. Seal the flask with a rubber septum and place on an orbital shaker kept at 30◦ C. Shake the reaction mixture at 250 rpm. 3. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). After spotting, it will be necessary to dry the TLC plate with a heat gun to evaporate the water in the sample. The reaction mixture remains a white suspension and the reaction is usually complete in 9 to 11 hr.
4. Work up and purify the product as described for compound S.2d (see Basic Protocol 4, steps 4 to 14). Since the product is hygroscopic, storage under a nitrogen atmosphere at −20◦ C is recommended. Under such conditions, no decomposition is seen even after several months.
5. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. Enzymatic Regioselective Levulinylation of Nucleosides
N4 -Benzoyl-3 -O-levulinyl-2 -O-methylcytidine (S.7b) is obtained in 70% to 75% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.48. m.p. not available (hygroscopic foam). [α]20 D = +10.1 (c 0.6, DMSO). IR (NaCl): υ 3318, 2931, 2359, 2339, 1739, 1715, 1698 cm−1 . 1 H NMR (CDCl3 , 300 MHz): δ 2.16 (s, 3H, Me-Lev), 2.63 (m, 2H, CH2 -Lev), 2.76 (m, 2H,
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CH2 -Lev), 3.48 (s, 3H, OMe), 3.78 (m, 1H, H5 ), 4.05 (m, 1H, H5 ), 4.28 (m, 2H, H2 + H4 ), 5.26 (apparent t, 3 JHH = 5.5 Hz, 1H, H3 ), 5.87 (d, 3 JHH = 3.4 Hz, 1H, H1 ), 7.437.54 (m, 4H, H5 + Hm + Hp ), 7.86 (m, 2H, Ho ), 8.37 (d, 3 JHH = 7.7 Hz, 1H, H6), 9.08 (br s, 1H, NH). 13 C NMR (CDCl3 , 75 MHz): δ 29.6 (CH2 -Lev), 29.5 (Me-Lev), 37.6 (CH2 -Lev), 58.6 (OMe), 60.2 (C5 ), 69.9, 81.6, 82.7, 89.9 (C1 + C2 + C3 + C4 ), 97.0 (C5), 127.6 and 128.6 (Co + Cm ), 132.7 (Ci ), 132.8 (Cp ), 145.8 (C6), 154.9 (C2), 162.6 (PhC=O), 166.8 (C4), 172.1 (C=O), 206.5 (C=O). ESI-MS (m/z): 460 [(M + H)+ , 100%], 482 [(M + Na)+ , 20%], 498 [(M + K)+ , 7]. ESI-HRMS (m/z): calcd. for C22 H25 N3 O8 , 459.1642; found, 459.1646. Anal. calcd. (%) for C22 H25 N3 O8 : C, 57.51; H, 5.48; N, 9.15; found: C, 57.3; H, 5.4; N, 8.9.
Preparation of N6 -Benzoyl-3 -O-Levulinyl-2 -O-Methyladenosine via Regioselective Enzymatic Hydrolysis with CAL-B
BASIC PROTOCOL 11
Materials N6 -Benzoyl-3 ,5 -di-O-levulinyl-2 -O-methyladenosine (S.6c; see Support Protocol 3) 1,4-Dioxane 0.15 M KH2 PO4 buffer (see recipe) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram; Novo Nordisk) Dichloromethane (CH2 Cl2 ) Silica gel Acetone Diethyl ether (Et2 O) 500-mL Erlenmeyer flask Rubber septum Orbital shaker, 30◦ C Buchner funnels with filter paper circles 500-mL round-bottom flasks Rotary evaporator 1-L filter flask Additional reagents and equipment for TLC (see Basic Protocol 1) Perform hydrolysis 1. Add 1 g (1.72 mmol) N6 -benzoyl-3 ,5 -di-O-levulinyl-2 -O-methyladenosine, 3.5 mL 1,4-dioxane, 13.7 mL of 0.15 M KH2 PO4 buffer, and 1 g CAL-B to a 500-mL Erlenmeyer flask. 2. Seal the flask with a rubber septum and place on an orbital shaker kept at 30◦ C. Shake the reaction mixture at 250 rpm. 3. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). After spotting, it will be necessary to dry the TLC plate with a heat gun to evaporate the water in the sample. The reaction mixture remains a white suspension and the reaction is usually complete in 13 to 15 hr.
Work up and purify product 4. When the reaction is complete, filter off the lipase under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 500-mL round-bottom flask. 5. Wash the CAL-B twice with 50 mL CH2 Cl2 , and collect all washes in the same round-bottom flask. Current Protocols in Nucleic Acid Chemistry
Protection of Nucleosides for Oligonucleotide Synthesis
2.11.21 Supplement 21
6. Remove solvents under reduced pressure using a rotary evaporator. 7. To recover product by dry column chromatography, weigh out 50 g silica gel and place in a Buchner funnel with a filter paper circle, attached to a 1-L filter flask connected to a vacuum line. Wash the silica with 200 mL of 25% (v/v) acetone/CH2 Cl2 . Acetone must be used in this case, as the product is unstable when exposed to MeOH or when subjected to extraction with saturated NaHCO3 solution.
8. Using a Pasteur pipet, spread the residue carefully on top of the silica bed. 9. Elute residual levulinic acid and N6 -benzoyl-3 ,5 -di-O-levulinyl-2 -O-methyladenosine with ∼200 to 400 mL of 25% (v/v) acetone/CH2 Cl2 . 10. Using clean filter flasks to collect 50-mL fractions, elute the desired product with ∼600 to 800 mL of 40% (v/v) acetone/CH2 Cl2 . Monitor fractions by analytical TLC. 11. Transfer fractions containing pure product to a 500-mL round-bottom flask, and remove solvents under reduced pressure using a rotary evaporator. 12. Add 10 mL Et2 O and triturate the product. Evaporate the solvent on a rotary evaporator and collect the product. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
13. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. N6 -Benzoyl-3 -O-levulinyl-2 -O-methyladenosine (S.7c) is obtained in 80% to 85% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.47. m.p. 56◦ -58◦ C. [α]20 D = −11.4 (c 1.0, DMSO). IR (NaCl): υ 3331, 1720, 1708, 1611cm−1 . 1 H NMR (CDCl3 , 200 MHz): δ 2.21 (s, 3H, Me-Lev), 2.71 (m, 2H, CH2 -Lev), 2.80 (m, 2H, CH2 -Lev), 3.27 (s, 3H, OMe), 3.84 (m, 2H, H5 ), 4.35 (m, 1H, H4 ), 4.74 (m, 1H, H2 ), 5.64 (apparent d, 3 JHH = 4.9 Hz, 1H, H3 ), 5.90 (d, 3 JHH = 8.0 Hz, 1H, H1 ), 7.46-7.61 (m, 3H, 2Hm + Hp ), 8.02 (m, 2H, 2Ho ), 8.12 (s, 1H, H2 or H8), 8.79 (s, 1H, H8 or H2). 13 C NMR (CDCl3 , 50 MHz): δ 27.7 (CH2 -Lev), 29.7 (Me-Lev), 37.7 (CH2 -Lev), 59.0 (OMe), 62.7 (C5 ), 72.1, 80.9, 86.3, 89.4 (C1 + C2 + C3 + C4 ), 124.6 (C5), 127.9 and 128.7 (Co + Cm ), 132.8 (Cp ), 133.2 (Ci ), 143.1 (C2 or C8), 150.4 (C4), 151.9 (C6 + C8 or C2), 164.6 (PhC=O), 171.8 (C=O), 206.3 (C=O). ESI-MS (m/z): 484 [(M + H)+ , 100%], 506 [(M + Na)+ , 7%]. ESI-HRMS (m/z): calcd. for C23 H25 N5 O7 , 483.1754; found, 483.1750. Anal. calcd. (%) for C23 H25 N5 O7 : C, 57.14; H, 5.21; N, 14.49; found: C, 57.2; H, 5.3; N, 14.7.
BASIC PROTOCOL 12
Preparation of N2 -Isobutyryl-3 -O-Levulinyl-2 -O-Methylguanosine via Regioselective Enzymatic Hydrolysis with CAL-B Materials
Enzymatic Regioselective Levulinylation of Nucleosides
N2 -Isobutyryl-3 ,5 -di-O-levulinyl-2 -O-methylguanosine (S.6d; see Support Protocol 3) 1,4-Dioxane 0.15 M KH2 PO4 buffer (see recipe) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram; Novo Nordisk) 500-mL Erlenmeyer flask Rubber septum Orbital shaker, 30◦ C Additional reagents and equipment for TLC (see Basic Protocol 1) and for workup and purification of the product (see Basic Protocol 4)
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Current Protocols in Nucleic Acid Chemistry
1. Add 1 g (1.76 mmol) N2 -isobutyryl-3 ,5 -di-O-levulinyl-2 -O-methylguanosine, 3.2 mL 1,4-dioxane, 14.4 mL of 0.15 M KH2 PO4 buffer, and 1 g CAL-B to a 500-mL Erlenmeyer flask. 2. Seal the flask with a rubber septum and place on an orbital shaker kept at 30◦ C. Shake the reaction mixture at 250 rpm. 3. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). After spotting, it will be necessary to dry the TLC plate with a heat gun to evaporate the water in the sample. The reaction mixture remains a white suspension and the reaction is usually complete in 9 to 11 hr.
4. Work up and purify the product as described for compound S.2d (see Basic Protocol 4, steps 4 to 14). If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
5. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. N2 -Isobutyryl-3 -O-levulinyl-2 -O-methylguanosine (S.7d) is obtained in 85% to 90% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.49. m.p. 82◦ -84◦ C. [α]20 D = −6.2 (c 1.0, DMSO). IR (NaCl): υ 3352, 2972, 2885, 1747, 1682, 1615, 1563 cm−1 . 1 H NMR (CDCl3 , 300 MHz): δ 1.20 (d, 3 JHH = 6.7 Hz, 6H, Me-iBu), 2.17 (s, 3H, Me-Lev), 2.63-2.82 (several m, 5H, 2CH2 -Lev + CH-iBu), 3.21 (s, 3H, OMe), 3.73-3.94 (m, 2H, H5 ), 4.21 (br s, 1H, H4 ), 4.55 (m, 1H, H2 ), 5.18 (br s, 1H, OH), 5.48 (m, 1H, H3 ), 5.77 (d, 3 JHH = 6.8 Hz, 1H, H1 ), 8.01 (s, 1H, H8), 9.85 (br s, 1H, NH), 12.30 (br s, 1H, NH). 13 C NMR (CDCl3 , 75 MHz): δ 18.7 (Me-iBu), 18.8 (Me-iBu), 27.7 (CH2 -Lev), 29.6 (Me-Lev), 36.0 (CH-iBu), 37.7 (CH2 -Lev), 58.8 (OMe), 61.8 (C5 ), 71.7, 81.3, 84.4, and 87.3 (C1 + C2 + C3 + C4 ), 121.5 (C5), 138.8 (C8), 147.8 and 147.9 (C2 + C4), 155.4 (C6), 171.9 (C=O), 179.4 (iBu-C=O), 206.5 (C=O). ESI-MS (m/z): 466 [(M + H)+ , 100%], 488 [(M + Na)+ , 24%]. Anal. calcd. (%) for C20 H27 N5 O8 : C, 51.61; H, 5.85; N, 15.05; found: C, 51.7; H, 6.0; N, 14.8.
Preparation of 3 ,5 -Di-O-Levulinyl-2 -O-Methylribonucleosides Additional Materials (also see Basic Protocol 1)
SUPPORT PROTOCOL 3
2 -O-Methylribonucleoside (Rasayan): 2 -O-methyluridine, N4 -benzoyl-2 -O-methylcytidine, N6 -benzoyl-2 -O-methyladenosine, or N2 -isobutyryl-2 -O-methylguanosine Triethylamine (Et3 N) Anhydrous 1,4-dioxane, freshly distilled from sodium and benzophenone Levulinic acid Dicyclohexylcarbodiimide (DCC) 4-(Dimethylamino)pyridine (DMAP) 200-mL round-bottom flask equipped with a three-way glass stopcock Perform levulinylation 1. Add a magnetic stir bar and 1 g of 2 -O-methylribonucleoside to a 200-mL roundbottom flask equipped with a three-way glass stopcock. One gram corresponds to 3.87 mmol 2 -O-methyluridine, 2.76 mmol N4 -benzoyl2 -O-methylcytidine, 2.59 mmol N6 -benzoyl-2 -O-methyladenosine, or 2.72 mmol N2 isobutyryl-2 -O-methylguanosine.
2. Evacuate the flask on a vacuum line and flush with nitrogen. Repeat this procedure three times.
Protection of Nucleosides for Oligonucleotide Synthesis
2.11.23 Current Protocols in Nucleic Acid Chemistry
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3. Add the following reagents: a. For 2 -O-mU: 1.30 mL (9.67 mmol) Et3 N, 25 mL dry 1,4-dioxane, 2.05 mL (20.13 mmol) levulinic acid, 4.15 g (20.13 mmol) DCC, and 38 mg (0.31 mmol) DMAP. b. For 2 -O-mCBz : 0.96 mL (6.90 mmol) Et3 N, 20 mL dry 1,4-dioxane, 1.47 mL (14.39 mmol) levulinic acid, 2.96 g (14.39 mmol) DCC, and 27 mg (0.22 mmol) DMAP. c. For 2 -O-mABz : 0.90 mL (6.47 mmol) Et3 N, 20 mL dry 1,4-dioxane, 1.38 mL (13.50 mmol) levulinic acid, 2.78 g (13.50 mmol) DCC, and 26 mg (0.21 mmol) DMAP. d. For 2 -O-mGi-Bu : 1.00 mL (6.80 mmol) Et3 N, 20 mL dry 1,4-dioxane, 1.45 mL (14.15 mmol) levulinic acid, 2.90 g (14.15 mmol) DCC, and 27 mg (0.22 mmol) DMAP. 4. Close the stopcock and begin stirring on a magnetic stirrer. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). The reaction is complete in 2 to 3 hr. After 2 hr, the reaction mixture will become heterogeneous due to the precipitation of 1, 3-dicyclohexylurea (DCU).
Work up and purify product 5. When the reaction is complete, filter the reaction mixture under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 500-mL roundbottom flask. This filtration step removes the DCU formed during the reaction.
6. Wash the DCU twice with 50 mL CH2 Cl2 and collect all washes in the same roundbottom flask. 7. Remove solvents under reduced pressure using a rotary evaporator. The product is an orange oil.
8. Dissolve the residue in 50 mL CH2 Cl2 and transfer to a 500-mL separatory funnel. 9. Remove levulinic acid by extracting the organic layer two to three times with 20 mL saturated NaHCO3 solution, using two 250-mL Erlenmeyer flasks to separate the layers. Verify that all levulinic acid has been removed from the organic phase by performing analytical TLC on both phases. Additional extractions may be needed for complete removal. Alternatively, brine can be used to clarify both phases.
10. Dry the organic layer over 4 g Na2 SO4 for 10 to 15 min. Remove the drying agent by gravity filtration using a glass funnel and Whatman no. 1 filter paper, collecting the filtrate in a 500-mL round-bottom flask. 11. Wash the Na2 SO4 four times with 25 mL CH2 Cl2 , collecting all washes in the same round-bottom flask. 12. Evaporate solvents under reduced pressure using a rotary evaporator. The product is obtained as orange oil due to 1,3-dicyclohexyl-1-levulinylurea contamination. Enzymatic Regioselective Levulinylation of Nucleosides
13. To recover product by dry column chromatography, weigh out 50 g silica gel and place in a Buchner funnel with a filter paper circle, attached to a 1-L filter flask connected to a vacuum line. Wash the silica with 100 mL of 25% (v/v) Et2 O/EtOAc.
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14. Using a Pasteur pipet, spread the residue carefully on top of the silica bed. 15. Elute 1,3-dicyclohexyl-1-levulinylurea with ∼200 to 300 mL of 25% (v/v) Et2 O/EtOAc. 16. Using clean filter flasks to collect 50-mL fractions, elute the desired product with ∼300 to 500 mL of 10% (v/v) MeOH/CH2 Cl2 . Monitor fractions by analytical TLC. 17. Transfer fractions containing pure product to a 500-mL round-bottom flask, and remove solvents under reduced pressure using a rotary evaporator. If desired, solvents can be recovered by distillation.
18. Add 10 mL Et2 O and triturate the product. Evaporate the solvent on a rotary evaporator and collect the orange product. If desired, products S.6b-d can be stored at −20◦ C. Since product S.6a is hygroscopic, storage of this compound under a nitrogen atmosphere at −20◦ C is recommended. Under such conditions, no decomposition is seen even after several months.
19. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. 3 ,5 -Di-O-levulinyl-2 -O-methyluridine (S.6a) is obtained in 90% to 95% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.51. m.p. not available (hygroscopic foam). [α]20 D = +2.5 (c 1.1, DMSO). IR (NaCl): υ 3209, 2933, 1738, 1714 cm−1 . 1 H NMR (CDCl3 , 200 MHz): δ 2.18 (s, 3H, Me-Lev), 2.19 (s, 3H, Me-Lev), 2.55-2.90 (m, 8H, 4CH2 -Lev), 3.44 (s, 3H, OMe), 4.03 (m, 1H, H4 ), 4.27-4.45 (m, 3H, H2 + 2H5 ), 5.09 (apparent t, 3 JHH = 5.3 Hz, 1H, H3 ), 5.81 (d, 3 JHH = 8.2 Hz, 1H, H5), 5.97 (d, 3 JHH = 4.6 Hz, 1H, H1 ), 7.64 (d, 3 JHH = 8.2 Hz, 1H, H6), 9.55 (br s, 1H, NH). 13 C NMR (CDCl3 , 50 MHz): δ 27.7 (CH2 -Lev), 27.7 (CH2 -Lev), 29.6 (Me-Lev), 29.7 (Me-Lev), 37.7 (CH2 -Lev), 37.8 (CH2 -Lev), 58.9 (OMe), 62.7 (C5 ), 70.1, 79.6, 81.3, and 87.6 (C1 + C2 + C3 + C4 ), 102.9 (C5), 139.4 (C6), 150.2 (C4), 163.1 (C2), 171.9 (C=O), 172.1 (C=O), 206.1 (C=O), 206.4 (C=O). ESI-MS (m/z): 455 [(M + H)+ , 66%], 477 [(M + Na)+ , 100%], 493 [(M + K)+ , 12%]. ESI-HRMS (m/z): calcd. for C20 H26 N2 O10 , 454.1587; found, 454.1565. Anal. calcd. (%) for C20 H26 N2 O10 : C, 52.86; H, 5.77; N, 6.16; found: C, 52.6; H, 5.8; N, 6.2. N4 -Benzoyl-3 ,5 -di-O-levulinyl-2 -O-methylcytidine (S.6b) is obtained in 90% to 95% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.61. m.p. 220◦ -222◦ C (decomposed). [α]20 D = +10.5 (c 1.1, DMSO). IR (NaCl): υ 2359, 2339, 1742, 1715, 1667 cm−1 . 1 H NMR (CDCl3 , 300 MHz): δ 2.20 (s, 3H, Me-Lev), 2.22 (s, 3H, Me-Lev), 2.59 (m, 4H, 2CH2 -Lev), 2.87 (m, 4H, 2CH2 -Lev), 3.60 (s, 3H, OMe), 4.13 (m, 1H, H4 ), 4.36-4.51 (m, 3H, H2 + 2H5 ), 4.93 (dd, 3 JHH = 7.8, 5.6 Hz, 1H, H3 ), 6.04 (m, 1H, H1 ), 7.50-7.65 (m, 4H, H5 + Hm + Hp ), 7.90 (m, 2H, Ho ), 8.23 (d, 3 JHH = 7.4 Hz, 1H, H6), 8.65 (br s, 1H, NH). 13 C NMR (DMSO-d6 , 75 MHz): δ 27.6 (CH2 -Lev), 29.6 (Me-Lev), 37.5 (CH2 -Lev), 58.3 (OMe), 63.1 (C5 ), 70.4, 79.4, 80.7, and 89.2 (C1 + C2 + C3 + C4 ), 97.0 (C5), 128.6, 128.7 (Co + Cm ), 132.5 (Cp ), 132.9 (Ci ), 145.3 (C6), 154.4 (C2), 163.5 (PhC=O), 167.5 (C4), 171.8 (C=O), 172.3 (C=O), 206.8 (C=O), 207.0 (C=O). ESI-MS (m/z): 558 [(M + H)+ , 100%], 580 [(M + Na)+ , 19%], 596 [(M + K)+ , 3%]. Anal. calcd. (%) for C27 H31 N3 O10 : C, 58.16; H, 5.60; N, 7.54; found: C, 58.2; H, 5.4; N, 7.7. N6 -Benzoyl-3 ,5 -di-O-levulinyl-2 -O-methyladenosine (S.6c) is obtained in 90% to 95% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.55. m.p. 52◦ -54◦ C. [α]20 D = −23.1 (c 1.1, DMSO). IR (NaCl): υ 2932, 2341, 1741, 1715, 1610 cm−1 . 1 H NMR (CDCl3 , 300 MHz): δ 2.19 (s, 3H, Me-Lev), 2.21 (s, 3H, Me-Lev), 2.26 (m, 2H, CH2 -Lev), 2.71 (m, 2H, CH2 -Lev), 2.80 (m, 4H, 2CH2 -Lev), 3.42 (s, 3H, OMe), 4.44 (m, 3H, H4 + 2H5 ), 4.74 (apparent t, 3 JHH = 5.3 Hz, 1H, H2 ), 5.43 (apparent t, 3 JHH = 4.4 Hz, 1H, H3 ), 6.16 (d, 3 JHH = 5.4 Hz, 1H, H1 ), 7.53-7.61 (m, 3H, 2Hm + Hp ), 8.02 (m, 2H, 2Ho ), 8.30 (s, 1H, H2 or H8), 8.81 (s, 1H, H8 or H2), 9.05 (br s, 1H, NH). 13 C NMR (CDCl3 , 75 MHz): δ 27.7 (CH2 -Lev), 29.7 (Me-Lev), 37.7 (CH2 -Lev), 37.8 (CH2 -Lev), 59.1 (OMe), 63.0 (C5 ), 70.8, 80.5, 81.1, and 87.0 (C1 + C2 + C3 + C4 ), 123.6 (C5), 127.8 and 128.7 (Co + Cm ), 132.7 (Cp ), 133.5 (Ci ), 141.6 (C2 or C8), 149.6 (C4), 151.6 (C6), 152.7 (C8 or
Protection of Nucleosides for Oligonucleotide Synthesis
2.11.25 Current Protocols in Nucleic Acid Chemistry
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C2), 164.7 (PhC=O), 171.8 (C=O), 172.2 (C=O), 206.1 (C=O), 206.4 (C=O). ESI-MS (m/z): 582 [(M + H)+ , 100%], 604 [(M + Na)+ , 3%]. Anal. calcd. (%) for C28 H31 N5 O9 : C, 57.83; H, 5.37; N, 12.04; found: C, 58.1; H, 5.2; N, 12.2. N2 -Isobutyryl-3 ,5 -di-O-levulinyl-2 -O-methylguanosine (S.6d) is obtained in 90% to 95% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.59. m.p. 55◦ -57◦ C. [α]20 D = −10.1 (c 1.0, DMSO). IR (NaCl): υ 3374, 2972, 2931, 1719, 1689, 1611, 1467, 1380, 1266 cm−1 . 1 H NMR (CDCl3 , 300 MHz): δ 1.22 (d, 3 JHH = 6.8 Hz, 6H, Me-iBu), 2.13 (s, 3H, Me-Lev), 2.18 (s, 3H, Me-Lev), 2.55-2.80 (several m, 9H, 4CH2 -Lev + CH-iBu), 3.30 (s, 3H, OMe), 4.39-4.56 (m, 4H, H2 + H4 + 2H5 ), 5.39 (m, 1H, H3 ), 5.81 (d, 3 JHH = 5.7 Hz, 1H, H1 ), 7.83 (s, 1H, H8), 9.75 (br s, 1H, NH), 12.18 (br s, 1H, NH). 13 C NMR (CDCl3 , 75 MHz): δ 18.7 (Me-iBu), 18.8 (Me-iBu), 27.6 (CH2 -Lev), 27.8 (CH2 -Lev), 29.5 (Me-Lev), 29.6 (Me-Lev), 36.0 (CH-iBu), 37.6 (CH2 -Lev), 37.7 (CH2 -Lev), 58.9 (OMe), 63.1 (C5 ), 71.0, 80.2, 80.9, and 87.4 (C1 + C2 + C3 + C4 ), 121.9 (C5), 138.0 (C8), 147.8 and 147.9 (C2 + C4), 155.5 (C6), 171.8 (C=O), 172.8 (C=O), 179.2 (iBu-C=O), 206.2 (C=O), 206.4 (C=O). ESI-MS (m/z): 564 [(M + H)+ , 100%], 586 [(M + Na)+ , 7%]. Anal. calcd. (%) for C25 H33 N5 O10 : C, 53.28; H, 5.90; N, 12.43; found: C, 53.3; H, 5.9; N, 12.2.
SYNTHESIS OF 5 -O-LEVULINYL-2 -METHYLRIBONUCLEOSIDES The preparation of 5 -O-levulinyl-2 -O-methylribonucleosides involves regioselective acylation of the 5 -hydroxyl function of 2 -O-methylribonucleosides with O-levulinyl acetonoxime using CAL-B as shown in Figure 2.11.4. This is a general procedure for the U, C, A, and G monomers. BASIC PROTOCOL 13
Preparation of 5 -O-Levulinyl-2 -O-Methyluridine via Regioselective Enzymatic Acylation with CAL-B Materials 2 -O-Methyluridine (Rasayan) O-Levulinyl acetonoxime (see Support Protocol 1) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram; Novo Nordisk) Anhydrous nitrogen Anhydrous tetrahydrofuran (THF), freshly distilled over sodium and benzophenone Dichloromethane (CH2 Cl2 ) Diethyl ether (Et2 O), cold (4◦ C) 1-L Erlenmeyer flask equipped with a three-way glass stopcock Rubber septum Orbital shaker, 40◦ C
Enzymatic Regioselective Levulinylation of Nucleosides
Figure 2.11.4 Regioselective levulinylation of 2 -O-methylribonucleosides (S.5a-d) using CAL-B. Abbreviations: ABz, N 6 -benzoyladenin-9-yl; CBz, N 4 -benzoylcytosin-1-yl; Gi-Bu, N 2 -isobutyrylguanin9-yl; Lev, levulinyl; U, uracil-1-yl.
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Buchner funnels with filter paper circles 500-mL round-bottom flask Rotary evaporator Additional reagents and equipment for TLC (see Basic Protocol 1) Perform levulinylation 1. Add 5 g (19.36 mmol) 2 -O-methyluridine, 9.9 g (58.08 mmol) O-levulinyl acetonoxime, and 5 g CAL-B to a 1-L Erlenmeyer flask equipped with a three-way glass stopcock. 2. Evacuate the Erlenmeyer flask on a vacuum line and flush with nitrogen. Repeat this procedure three times. 3. Add 97 mL dry THF. 4. Seal the flask with a rubber septum and place on an orbital shaker kept at 40◦ C. Shake the reaction mixture at 250 rpm. 5. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). The reaction mixture remains a white suspension and the reaction is complete in 5 to 7 hr.
Work up and purify product 6. When the reaction is complete, filter off the lipase under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 500-mL round-bottom flask. 7. Wash the CAL-B twice with 50 mL CH2 Cl2 , collecting all washes in the same roundbottom flask. 8. Remove solvents under reduced pressure using a rotary evaporator. 9. Dissolve the residue in 5 mL CH2 Cl2 and slowly add to 30 to 40 mL Et2 O to precipitate the product as a white solid. Cool the solution 2 to 3 days at 4◦ C. 10. Filter off the precipitated product under vacuum using a Buchner funnel with a filter paper circle. Wash the precipitate two to three times with 50 mL cold (4◦ C) Et2 O, and then dry the solid under vacuum. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
11. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. 5 -O-Levulinyl-2 -O-methyluridine (S.8a) is obtained in 90% to 95% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.38. m.p. 114◦ -116◦ C. [α]20 D = +9.1 (c 1.0, DMSO). IR (NaCl): υ 2359, 2340, 1698 cm−1 . 1 H NMR (CDCl3 , 300 MHz): δ 2.19 (s, 3H, Me-Lev), 2.56 (m, 2H, CH2 -Lev), 2.81 (m, 2H, CH2 -Lev), 3.63 (s, 3H, OMe), 3.84 (m, 1H, H4 ), 4.14 (m, 2H, H2 + H3 ), 4.43 (m, 2H, H5 ), 5.79 (d, 3 JHH = 8.0 Hz, 1H, H5), 5.91 (m, 1H, H1 ), 7.72 (d, 3 JHH = 8.0 Hz, 1H, H6), 9.54 (br s, 1H, NH). 13 C NMR (CDCl3 , 75 MHz): δ 27.7 (CH2 -Lev), 29.6 (Me-Lev), 37.8 (CH2 -Lev), 58.7 (OMe), 62.4 (C5 ), 68.6, 81.6, 83.1, and 87.8 (C1 + C2 + C3 + C4 ), 102.4 (C5), 139.5 (C6), 150.1 (C4), 163.3 (C2), 172.3 (C=O), 206.5 (C=O). ESI-MS (m/z): 357 [(M + H)+ , 12%], 379 [(M + Na)+ , 100%], 395 [(M + K)+ , 7%]. Anal. calcd. (%) for C15 H20 N2 O8 : C, 50.56; H, 5.66; N, 7.86; found: C, 50.4; H, 5.7; N, 7.8.
Protection of Nucleosides for Oligonucleotide Synthesis
2.11.27 Current Protocols in Nucleic Acid Chemistry
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BASIC PROTOCOL 14
Preparation of N4 -Benzoyl-5 -O-Levulinyl-2 -O-Methylcytidine via Regioselective Enzymatic Acylation with CAL-B Materials N4 -Benzoyl-2 -O-methylcytidine (Rasayan) O-Levulinyl acetonoxime (see Support Protocol 1) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram; Novo Nordisk) Anhydrous nitrogen Anhydrous tetrahydrofuran (THF), freshly distilled over sodium and benzophenone Dichloromethane (CH2 Cl2 ) Saturated sodium bicarbonate (NaHCO3 ) solution Sodium sulfate (Na2 SO4 ) Diethyl ether (Et2 O), cold (4◦ C) 1-L Erlenmeyer flask equipped with a three-way glass stopcock Rubber septum Orbital shaker, 40◦ C Buchner funnels with filter paper circles 500-mL round-bottom flasks Rotary evaporator 250-mL separatory funnel Glass funnel with Whatman no. 1 filter paper Filter flask Additional reagents and equipment for TLC (see Basic Protocol 1) Perform levulinylation 1. Add 5 g (13.84 mmol) N4 -benzoyl-2 -O-methylcytidine, 7.1 g (41.52 mmol) Olevulinyl acetonoxime, and 5 g CAL-B to a 1-L Erlenmeyer flask equipped with a three-way glass stopcock. 2. Evacuate the Erlenmeyer flask on a vacuum line and flush with nitrogen. Repeat this procedure three times. 3. Add 69 mL dry THF. 4. Seal the flask with a rubber septum and place on an orbital shaker kept at 40◦ C. Shake the reaction mixture at 250 rpm. 5. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). The reaction mixture remains a white suspension and the reaction is complete in 7 to 9 hr.
Work up and purify product 6. When the reaction is complete, filter off the lipase under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 500-mL round-bottom flask. 7. Wash the CAL-B twice with 50 mL CH2 Cl2 , and collect all washes in the same round-bottom flask. 8. Remove solvents under reduced pressure using a rotary evaporator. 9. Dissolve the residue in 50 mL CH2 Cl2 and transfer to a 250-mL separatory funnel. Enzymatic Regioselective Levulinylation of Nucleosides
10. Remove unreacted starting material by extracting the organic layer three times with 10 mL saturated NaHCO3 solution, using two 250-mL Erlenmeyer flasks to
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separate the phases. Verify that N4 -benzoyl-2 -O-methylcytidine has been removed completely from the organic phase by performing analytical TLC on both phases. N4 -Benzoyl-2 -O-methylcytidine is partially soluble in the aqueous phase. Alternatively, brine can be used to separate the phases effectively.
11. Pool the aqueous phases and extract twice with 10 mL CH2 Cl2 to recover any product. 12. Pool the organic phases from step 11 and verify by TLC that they contain no residual N4 -benzoyl-2 -O-methylcytidine. If any is present, re-extract with 10 mL saturated NaHCO3 solution. This extraction is unnecessary if N4 -benzoyl-2 -O-methylcytidine is absent from the organic phase.
13. Pool the organic phases from steps 10 and 12 and dry over 3 g Na2 SO4 for 10 to 15 min. Remove the drying agent by gravity filtration with a glass funnel and Whatman no. 1 filter paper, collecting the filtrate in a 500-mL round-bottom flask. 14. Wash the Na2 SO4 five times with 15 mL CH2 Cl2 , and collect all washes in the same round-bottom flask. 15. Evaporate solvents under reduced pressure using a rotary evaporator. The product is an orange oil due to oxime ester contamination.
16. Cool the crude product at 4◦ C overnight to precipitate the product. 17. Transfer the solid to a Buchner funnel with a filter paper circle, attached to a filter flask connected to a vacuum line. Wash the orange solid (i.e., the final product) five to six times with 20 mL cold Et2 O, and then dry the solid under vacuum. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
18. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. N4 -Benzoyl-5 -O-levulinyl-2 -O-methylcytidine (S.8b) is obtained in 80% to 85% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.52. m.p. 134◦ -136◦ C. [α]20 D = +68.5 (c 1.0, DMSO). IR (NaCl): υ 3318, 2947, 2359, 2339, 1738, 1698, 1651 cm−1 . 1 H NMR (CDCl3 , 300 MHz): δ 2.16 (s, 3H, Me-Lev), 2.58 (m, 2H, CH2 -Lev), 2.79 (m, 2H, CH2 -Lev), 3.66 (s, 3H, OMe), 3.83 (m, 1H, H3 ), 4.13 (m, 2H, H2 + H4 ), 4.47 (m, 2H, H5 ), 5.92 (m, 1H, H1 ), 7.46-7.57 (m, 4H, H5 + Hm + Hp ), 7.89 (m, 2H, Ho ), 8.26 (d, 3 JHH = 7.4 Hz, 1H, H6), 9.26 (br s, 1H, NH). 13 C NMR (CDCl3 , 75 MHz): δ 27.8 (CH2 -Lev), 29.7 (Me-Lev), 37.8 (CH2 -Lev), 58.8 (OMe), 62.0 (C5 ), 68.0, 81.4, 83.2, and 88.9 (C1 + C2 + C3 + C4 ), 96.8 (C5), 127.5 and 128.9 (Co + Cm ), 132.8 (Ci ), 133.1 (Cp ), 144.2 (C6), 154.5 (C2), 162.5 (PhC=O), 166.8 (C4), 172.4 (C=O), 206.4 (C=O). ESI-MS (m/z): 460 [(M + H)+ , 100%], 482 [(M + Na)+ , 22%], 498 [(M + K)+ , 6%]. ESI-HRMS (m/z): calcd. for C22 H25 N3 O8 , 459.1642; found, 459.1639. Anal. calcd. (%) for C22 H25 N3 O8 : C, 57.51; H, 5.48; N, 9.15; found: C, 57.3; H, 5.7; N, 9.4.
Preparation of N6 -Benzoyl-5 -O-Levulinyl-2 -O-Methyladenosine via Regioselective Enzymatic Acylation with CAL-B
BASIC PROTOCOL 15
Materials N6 -Benzoyl-2 -O-methyladenosine (Rasayan) O-Levulinyl acetonoxime (see Support Protocol 1) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram; Novo Nordisk) Anhydrous nitrogen
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Anhydrous tetrahydrofuran (THF), freshly distilled over sodium and benzophenone Dichloromethane (CH2 Cl2 ) Silica gel Acetone Diethyl ether (Et2 O), cold (4◦ C) 1-L Erlenmeyer flask equipped with a three-way glass stopcock Rubber septum Orbital shaker, 40◦ C Buchner funnels with filter paper circles 500-mL round-bottom flasks Rotary evaporator 1-L filter flask Additional reagents and equipment for TLC (see Basic Protocol 1) Perform levulinylation 1. Add 5 g (12.97 mmol) N6 -benzoyl-2 -O-methyladenosine, 6.7 g (38.93 mmol) Olevulinyl acetonoxime, and 5 g CAL-B to a 1-L Erlenmeyer flask equipped with a three-way glass stopcock. 2. Evacuate the Erlenmeyer flask on a vacuum line and flush with nitrogen. Repeat this procedure three times. 3. Add 65 mL dry THF. 4. Seal the flask with a rubber septum and place on an orbital shaker kept at 40◦ C. Shake the reaction mixture at 250 rpm. 5. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). The reaction mixture remains a white suspension and the reaction is complete in 18 to 20 hr.
Work up and purify product 6. When the reaction is complete, filter off the lipase under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 500-mL round-bottom flask. 7. Wash the CAL-B twice with 50 mL CH2 Cl2 , and collect all washes in the same round-bottom flask. 8. Remove solvents under reduced pressure using a rotary evaporator. 9. To recover product by dry column chromatography, weigh out 100 g silica gel and place in a Buchner funnel with a filter paper circle, attached to a 1-L filter flask connected to a vacuum line. Wash the silica with 200 mL of 10% (v/v) acetone/CH2 Cl2 . Acetone must be used in this protocol, as the product is unstable when exposed to MeOH or when subjected to extraction with saturated NaHCO3 solution.
10. Using a Pasteur pipet, spread the residue carefully on top of the silica bed. 11. Elute the excess oxime ester with ∼500 to 700 mL of 10% (v/v) acetone/CH2 Cl2 .
Enzymatic Regioselective Levulinylation of Nucleosides
12. Using clean filter flasks to collect 100-mL fractions, elute the desired product with ∼1200 to 1500 mL of 25% (v/v) acetone/CH2 Cl2 . Monitor fractions by analytical TLC.
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13. Transfer fractions containing pure product to a 500-mL round-bottom flask, and remove solvents under reduced pressure using a rotary evaporator. 14. Transfer the solid to a Buchner funnel with a filter paper circle, attached to a filter flask connected to a vacuum line. Wash the white solid (i.e., the final product) three to four times with 20 mL cold (4◦ C) Et2 O, and then dry the solid under vacuum. If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
15. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis. N6 -Benzoyl-5 -O-levulinyl-2 -O-methyladenosine (S.8c) is obtained in 75% to 80% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.44. m.p. 51◦ -53◦ C. [α]20 D = −6.7 (c 1.0, DMSO). IR (NaCl): υ 2931, 2359, 1716, 1612 cm−1 . 1 H NMR (DMSO-d6 , 200 MHz): δ 2.09 (s, 3H, Me-Lev), 2.50 (m, 2H, CH2 -Lev), 2.71 (m, 2H, CH2 -Lev), 3.41 (s, 3H, OMe), 4.15-4.59 (m, 5H, H2 + H3 + H4 + 2H5 ), 5.56 (br s, 1H, OH), 6.20 (d, 3 JHH = 3.2 Hz, 1H, H1 ), 7.53-7.65 (m, 3H, 2Hm + Hp ), 8.06 (m, 2H, 2Ho ), 8.70 (s, 1H, H2 or H8), 8.78 (s, 1H, H8 or H2). 13 C NMR (DMSO-d6 , 50 MHz): δ 27.5 (CH2 -Lev), 29.5 (Me-Lev), 37.4 (CH2 -Lev), 57.8 (OMe), 63.8 (C5 ), 68.9, 81.8, 82.3, and 85.8 (C1 + C2 + C3 + C4 ), 125.8 (C5), 128.5, 128.6 (Co + Cm ), 132.5 (Cp ), 133.3 (Ci ), 143.1 (C2 or C8), 150.6 (C4), 151.8 (C8 or C2), 152.0 (C6), 165.8 (PhC=O), 172.2 (C=O), 206.8 (C=O). ESI-MS (m/z): 484 [(M + H)+ , 100%], 506 [(M + Na)+ , 5%], 522 [(M + K)+ , 1%]. Anal. calcd. (%) for C23 H25 N5 O7 : C, 57.14; H, 5.21; N, 14.49; found: C, 57.3; H, 5.7; N, 14.5.
Preparation of N2 -Isobutyryl-5 -O-Levulinyl-2 -O-Methylguanosine via Regioselective Enzymatic Acylation with CAL-B
BASIC PROTOCOL 16
Materials N2 -Isobutyryl-2 -O-methylguanosine (Rasayan) O-Levulinyl acetonoxime (see Support Protocol 1) Immobilized Candida antarctica lipase B (CAL-B, under the trade name Novozyme 435; 10,000 propyl laurate units per gram; Novo Nordisk) Anhydrous nitrogen Anhydrous tetrahydrofuran (THF), freshly distilled over sodium and benzophenone Dichloromethane (CH2 Cl2 ) Silica gel Ethyl acetate (EtOAc) Diethyl ether (Et2 O) Methanol (MeOH) 500-mL Erlenmeyer flask equipped with a three-way glass stopcock Rubber septum Orbital shaker, 40◦ C Buchner funnels with filter paper circles 250- and 500-mL round-bottom flasks Rotary evaporator 1-L filter flask Additional reagents and equipment for TLC (see Basic Protocol 1) Perform levulinylation 1. Add 1 g (2.72 mmol) N2 -isobutyryl-2 -O-methylguanosine, 1.40 g (8.16 mmol) Olevulinyl acetonoxime, and 1 g CAL-B to a 500-mL Erlenmeyer flask equipped with a three-way glass stopcock. 2. Evacuate the Erlenmeyer flask on a vacuum line and flush with nitrogen. Repeat this procedure three times.
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3. Add 14 mL dry THF. 4. Seal the flask with a rubber septum and place on an orbital shaker kept at 40◦ C. Shake the reaction mixture at 250 rpm. 5. Monitor the progress of the reaction by analytical TLC as described for the synthesis of compound S.2a (see Basic Protocol 1, step 5). The reaction mixture remains a white suspension and the reaction is complete in 6 to 8 hr.
Work up and purify product 6. When the reaction is complete, filter off the lipase under vacuum using a Buchner funnel with a filter paper circle, and collect the filtrate in a 250-mL round-bottom flask. 7. Wash the CAL-B twice with 20 mL CH2 Cl2 , and collect all washes in the same round-bottom flask. 8. Remove solvents under reduced pressure using a rotary evaporator. The excess oxime ester imparts an orange color to the crude product. Because the product does not precipitate, a chromatography step is necessary.
9. To recover product by dry column chromatography, weigh out 50 g silica gel and place in a Buchner funnel with a filter paper circle, attached to a 1-L filter flask connected to a vacuum line. Wash the silica with 200 mL of 25% (v/v) EtOAc/Et2 O. 10. Using a Pasteur pipet, spread the residue carefully on top of the silica bed. 11. Elute the excess oxime ester with ∼300 to 500 mL of 25% (v/v) EtOAc/Et2 O. 12. Using clean filter flasks to collect 50-mL fractions, elute the desired product with ∼500 to 800 mL of 10% (v/v) MeOH/CH2 Cl2 . Monitor fractions by analytical TLC. 13. Transfer fractions containing pure product to a 500-mL round-bottom flask, and remove solvents under reduced pressure using a rotary evaporator. 14. Add 10 mL Et2 O and triturate the product. Evaporate the solvent on a rotary evaporator and collect the white compound (i.e., the final product). If desired, the product can be stored at −20◦ C. Under such conditions, no decomposition is seen even after several months.
15. Confirm product purity by TLC; m.p. analysis; optical rotation analysis; infrared (IR), 1 H NMR, and 13 C NMR spectroscopy; mass spectrometry; and elemental analysis.
Enzymatic Regioselective Levulinylation of Nucleosides
N2 -Isobutyryl-5 -O-levulinyl-2 -O-methylguanosine (S.8d) is obtained in 90% to 95% yield. TLC (10% MeOH/CH2 Cl2 ): Rf = 0.40. m.p. 57◦ -59◦ C. [α]20 D = −1.5 (c 1.0, DMSO). IR (NaCl): υ 3374, 2972, 1712, 1688, 1609, 1467, 1380, 1161 cm−1 . 1 H NMR (CDCl3 , 300 MHz): δ 1.24 (d, 3 JHH = 6.5 Hz, 6H, Me-iBu), 2.15 (s, 3H, Me-Lev), 2.56-2.92 (several m, 5H, 2CH2 -Lev + CH-iBu), 3.44 (s, 3H, OMe), 4.26-4.63 (several m, 5H, H2 + H3 + H4 + 2H5 ), 5.90 (d, 3 JHH = 3.4 Hz, 1H, H1 ), 7.96 (s, 1H, H8), 10.38 (br s, 1H, NH), 12.39 (br s, 1H, NH). 13 C NMR (CDCl3 , 50 MHz): δ 18.8 (Me-iBu), 19.0 (Me-iBu), 27.8 (CH2 -Lev), 29.6 (Me-Lev), 36.0 (CH-iBu), 37.8 (CH2 -Lev), 58.7 (OMe), 63.7 (C5 ), 69.7, 82.2, 83.2, and 87.5 (C1 + C2 + C3 + C4 ), 121.4 (C5), 138.2 (C8), 148.1 (C2 + C4), 155.7 (C6), 172.8 (C=O), 179.9 (iBu-C=O), 206.9 (C=O). ESI-MS (m/z): 466 [(M + H)+ , 100%], 488 [(M + Na)+ , 23%], 504 [(M + K)+ , 7%]. Anal. calcd. (%) for C20 H27 N5 O8 : C, 51.61; H, 5.85; N, 15.05; found: C, 51.4; H, 6.1; N, 15.2.
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
p-Anisaldehyde solution Prepare 2.5% (v/v) anisaldehyde in ethanol. Dissolve and then add 3.5% (v/v) H2 SO4 and 1% (v/v) acetic acid. Store up to 2 months at room temperature. Apply to TLC plates with a spray bottle or by dipping the plate in the solution. CAUTION: Prepare and use this reagent only in a well-ventilated fume hood. It is corrosive, and its vapors are highly irritating.
KH2 PO4 buffer, 0.15 M To prepare 100 mL of buffer, dissolve 2.04 g KH2 PO4 in 70 mL distilled water and gradually add 0.1 N KOH until pH 7 is attained. Add water to 100 mL. Store up to 4 months at room temperature. COMMENTARY Background Information The increasing demand for oligonucleotides in therapeutic, diagnostic, and target validation has impelled research aimed at the development of improved synthetic protocols. Among these protocols, solid-phase synthesis is the current method of choice due to its speed and high coupling efficiency. In recent years, however, interest in solution-phase synthesis of oligonucleotides has been steadily growing due to much anticipated success with therapeutic oligonucleotides, which may trigger a very large demand for such products (Reese and Yan, 2001). In order to assemble oligonucleotides on a large scale, it is essential that investigators have access to appropriately protected nucleosides as building blocks. Since nucleosides have multiple reactive sites, such as primary and secondary hydroxyl groups and exocyclic amino groups, these compounds must be protected in an orthogonal way, such that one group is selectively excised over others. This is a serious problem in multifunctional molecules such as oligonucleotides, which must be fully protected from unwanted reactions during their assembly. A number of 3 - and 5 -hydroxyl-protecting groups have been reported to date for synthesis of oligonucleotides (see Chapter 2). Among the arsenal of available protecting groups, the levulinyl group has been well recognized and well established as a very useful orthogonal protecting group for hydroxyl groups. The reason for its popularity is easy to understand: it is stable to phosphorus coupling and is cleaved under mild conditions without affecting other protecting groups in the molecule,
such as acyl groups on exocyclic amines, internucleotidic phosphate-protecting groups, and the dimethoxytrityl group. Importantly, the levulinyl group is unmasked under essentially neutral conditions: 0.5 M hydrazine hydrate in 4:1 (v/v) pyridine/acetic acid at room temperature within 2 to 4 min in solution-phase synthesis protocols (van Boom and Burgers, 1976), or 10 to 15 min in solid-phase syntheses (Iwai and Ohtsuka, 1988; Iwai et al., 1990). Until recently, syntheses of 3 - and/or 5 -Olevulinyl-protected nucleosides were carried out using multiple tedious protection and deprotection steps and resulted in relatively low yields. In 2002, the authors reported an improved synthesis of such nucleosides via regioselective enzymatic hydrolysis of 3 ,5 -diO-levulinyl nucleosides (Garc´ıa et al., 2002). Although that protocol works very well, it requires a two-step procedure: chemical bisacylation of the nucleoside followed by selective hydrolysis of one of the levulinyl groups. In 2003, the authors further improved the protocol by regioselectively acylating nucleosides (Garc´ıa et al., 2003). Arguably, one-step direct acylation using lipases constitutes the shortest method that is both cost-efficient and environmentally friendly. Today, lipases have been established as valuable reagents for industrial-scale organic synthesis. Lipases promote the hydrolysis of carboxylic esters as well as acyl transfer onto hydroxyl groups with unparalleled stereo-, regio-, and chemoselectivity. Furthermore, the authors have utilized lipases that are immobilized, providing additional advantages such as enhanced stability with ease of handling, ability to recycle and reuse, convenient
Protection of Nucleosides for Oligonucleotide Synthesis
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separation from the reaction mixture by simple filtration, and commercial availability on a large scale. The protocols in this unit describe the lipase-mediated syntheses of two sets of 3 -levulinyl-protected and two sets of 5 -levulinyl-protected nucleosides that are essential raw materials for oligonucleotide assembly. 3 -O-Levulinyl nucleosides The direct protection of the secondary 3 hydroxyl group in an unprotected nucleoside is a challenging problem due to the competing primary hydroxyl group in the same molecule. Although several chemical methods are available for regioselective acylation of a primary hydroxyl group in a nucleoside, a method for direct chemical acylation of the 3 -hydroxyl group in a nucleoside is nonexistent. In contrast, selective introduction of an acyl group onto the 3 -hydroxyl group of a nucleoside has been successfully accomplished with lipases. This unit presents syntheses of 3 -O-levulinyl derivatives of T, dCBz , dABz , dGi-Bu as well as 2 -O-Me analogs of U, CBz , ABz and Gi-Bu . In Basic Protocols 1 to 3, the readily available nucleosides T, dCBz , and dABz are treated with O-levulinyl acetonoxime (prepared in Support Protocol 1) in the presence of PSL-C at 30◦ C in THF for 14 to 18 hr. The resulting 3 -Olevulinylated derivatives are isolated in >80% yields without column chromatography. Because all of the authors’ attempts to acylate dGi-Bu in a single step resulted in formation of a mixture of products with low selectivity, 3 -O-levulinyl dGi-Bu is prepared using a twostep protocol that involves bis-acylation of the nucleoside followed by CAL-B-mediated hydrolysis (see Basic Protocol 4). Similarly, the preparation of 3 -O-levulinyl-2 -O-Me nucleosides resulted in a mixture of products under direct acylation conditions. As a result, this set of nucleosides is prepared via CAL-B-driven hydrolysis of di-O-levulinylated nucleosides (S.6a-d) to furnish 3 -O-levulinylated products (S.7a-d) in 75% to 90% yields. To the best of the authors’ knowledge, this is the first synthesis of S.7a-d to be reported in the literature. Furthermore, these protocols demonstrate that lipases are able to accept a broad range of substrates with structural diversity.
Enzymatic Regioselective Levulinylation of Nucleosides
5 -O-Levulinyl nucleosides Enzymatic acylation methods have shown an increasing tendency to proceed preferentially at the 5 -hydroxyl group of a nucleoside. Until recently, however, this technique had not been exploited for the installation of levulinyl
groups. Highly regioselective enzymatic levulinyl transfer to the 5 -hydroxyl group has been studied in the authors’ laboratory. In Basic Protocols 5 to 8, CAL-B-catalyzed syntheses of 5 -O-levulinyl derivatives of T, dCBz , dABz , and dGi-Bu that result in 70% to 90% yields are described. The scalability of this acylation method has been clearly demonstrated by performing the syntheses of S.3a-d on a 10-g scale. Similarly, Basic Protocols 13 to 16 describe the CAL-B-catalyzed syntheses of 5 -O-levulinyl-2 -O-Me ribonucleosides in excellent yields. Importantly, the syntheses of the 5 -O-levulinyl derivatives S.8a-d are the first that have been reported in the literature.
Critical Parameters The success of the various reactions described here can be affected by several factors. For example, all starting materials, including nucleosides, must be thoroughly dried under vacuum prior to use in acylation reactions, which are carried out in freshly distilled anhydrous THF. It is important to note that some levulinyl-protected nucleosides are partially soluble in water. Thus, after the initial extraction, back-extraction of the aqueous layer may be required for complete recovery of the desired product. Yields are invariably improved after performing back-extraction. Although these protocols use excess oxime ester and lipase, both of these reagents are easy to recapture and recycle. A large-scale (25-g) study on the 3 -O-acylation of thymidine demonstrated that PSL-C can be recycled more than four times without loss of enzyme activity. Since the cost of lipases is relatively high, recycling of both CAL-B and PSL-C is recommended. Furthermore, excess oxime ester could potentially be captured and recycled should there be a need. It is noteworthy that the use of excess oxime ester during acylation reactions results in formation of orange-colored products. This color is easily eliminated via simple precipitation of the product from ether or hexanes. The color is also eliminated by chromatography. Generally, all products described in this unit are obtained as white solids; should the orange color persist, there is a possibility that a trace amount of oxime ester is present as an impurity. Although the protocols described in this unit may appear to be very similar, they differ in subtle but important ways. For example, the amount of THF required for each reaction is dependent on the solubility of the starting material. Also, the speed and temperature of the orbital shaker are critical to the successful
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outcome of these reactions. The use of dry column chromatography (Pedersen and Rosenbohm, 2001) results in significantly reduced solvent consumption and minimizes the waste stream. It should be emphasized that a magnetic stirrer is not recommended for enzymatic reactions involving immobilized lipases due to the possibility of grinding the beads. As indicated, best results are obtained when reactions are carried out on an orbital shaker at constant temperature.
Anticipated Results Lipase activity and selectivity are strongly influenced by the quality of solvents used, the substrate concentration, the reaction time, the temperature, and the moisture content. As indicated, hydrolytic reactions can be carried out in buffered aqueous solutions, whereas acylation is achieved in anhydrous solvents. More importantly, the type of acyl donor has a great influence on reactivity and selectivity. Use of acetonoxime levulinate as an acyl donor has been found to be best for all acylation protocols described in this unit. Some of these protocols have been scaled up without serious issues and show great potential for adaptation into industrial processes. Since the shelf lives of both CAL-B and PSL-C are excellent, it is advisable to purchase a large lot of lipase with good activity that can be used for an extended period of time. This will eliminate the lot-to-lot variation in enzyme activity and help produce consistent results. Also, it is more cost-effective to buy reagents in bulk. In the authors’ experience, all protocols described in this unit are safe, scalable, robust, and easy to perform, and all provide consistent results when good-quality raw materials and reagents are used. Furthermore, the use of enzymes for organic synthesis is ecofriendly and creates less waste compared with chemical methods.
Time Considerations All protocols described in this unit can be completed in 1 to 3 days with common laboratory equipment. It is noteworthy that these reactions are highly specific and that extending the reaction times for a few extra hours does not compromise the yield or quality of the products. In fact, one of the key advantages of working with lipases is that their specificity is fine-tuned, minimizing the potential for product damage. The partially protected nucleoside intermediates and reagents used in these protocols are now commercially available, which
may save significant time in the synthesis of the target products.
Literature Cited Bornscheuer, U.T. and Kazlauskas, R.J. 1999. Hydrolases in Organic Synthesis. Regioand Stereoselective Biotransformations. WileyVCH, Weinheim, Germany. Carrea, G. and Riva, S. 2000. Properties and synthetic applications of enzymes in organic solvents. Angew. Chem. Int. Ed. Engl. 39:22262254. Eleuteri, A., Cheruvallath, Z.S., Capaldi, D.C., Cole, D.L., and Ravikumar, V.T. 1999. Synthesis of dimer phosphoramidite synthons for oligodeoxyribonucleotide phosphorothioates using diethyldithiocarbonate disulfide as an efficient sulfurizing reagent. Nucleosides Nucleotides Nucleic Acids 18:1803-1807. Ferrero, M. and Gotor, V. 2000a. Biocatalytic selective modifications of conventional nucleosides, carbocyclic nucleosides, and C-nucleosides. Chem. Rev. 100:4319-4347. Ferrero, M. and Gotor, V. 2000b. Chemoenzymatic transformations in nucleoside chemistry. Monatsh. Chem. 131:585-616. Garc´ıa, J., Fern´andez, S., Ferrero, M., Sanghvi, Y.S., and Gotor, V. 2002. Building blocks for the solution phase synthesis of oligonucleotides: Regioselective hydrolysis of 3 ,5 -diO-levulinylnucleosides using an enzymatic approach. J. Org. Chem. 67:4513-4519. Garc´ıa, J., Fern´andez, S., Ferrero, M., Sanghvi, Y.S., and Gotor, V. 2003. Novel enzymatic synthesis of levulinyl protected nucleosides useful for solution phase synthesis of oligonucleotides. Tetrahedron Asymmetr. 14:3533-3540. Garc´ıa, J., Fern´andez, S., Ferrero, M., Sanghvi, Y.S., and Gotor, V. 2004a. Mild, efficient and regioselective enzymatic procedure for 5 -Obenzoylation of 2 -deoxynucleosides. Tetrahedron Lett. 45:1709-1712. Garc´ıa, J., Fern´andez, S., Ferrero, M., Sanghvi, Y.S., and Gotor, V. 2004b. Regioselective enzymatic acylation of β-L-2 -deoxynucleosides: Application in resolution of β-D/L-2 -deoxynucleosides. Org. Lett. 6:3759-3762. Greene, T.W. and Wuts, P.G.M. 1999. Protective Groups in Organic Synthesis, 3rd ed., p. 168. John Wiley & Sons, New York. Iwai, S. and Ohtsuka, E. 1988. 5 -Levulinyl and 2 tetrahydrofuranyl protection for the synthesis of oligoribonucleotides by the phosphoramidite approach. Nucl. Acids Res. 16:9443-9456. Iwai, S., Toshiro, S., and Ohtsuka, E. 1990. Large scale synthesis of oligoribonucleotides on a solid support: Synthesis of a catalytic RNA duplex. Tetrahedron 46:6673-6688. Klibanov, A.M. 2001. Improving enzymes by using them in organic solvents. Nature 409:241-246. Kocie´nski, P.J. 2004. Protecting Groups, 3rd ed., pp. 464 and 475. Georg Thieme Verlag, Stuttgart, Germany.
Protection of Nucleosides for Oligonucleotide Synthesis
2.11.35 Current Protocols in Nucleic Acid Chemistry
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Koeller, K.M. and Wong, C.H. 2001. Enzymes for chemical synthesis. Nature 409:232-240. Krotz, A.H., Klopchin, P., Cole, D.L., and Ravikumar, V.T. 1997. Improved impurity profile of phosphorothioate oligonucleotides through the use of dimeric phosphoramidite synthons. Nucleosides Nucleotides Nucleic Acids 16:16371640. Kumar, G. and Poonian, M.S. 1984. Improvements in oligodeoxyribonucleotide synthesis: Methyl N,N-dialkylphosphoramidite dimer units for solid support phosphite methodology. J. Org. Chem. 49:4905-4912. March´an, V., Cie´slak, J., Livengood, V., and Beaucage, S.L. 2004. 2,2,5,5Tetramethylpyrrolidin-3-one-1-sulfinyl group for 5 -hydroxyl protection of deoxyribonucleoside phosphoramidites in the solid-phase preparation of DNA oligonucleotides. J. Am. Chem. Soc. 126:9601-9610. Patel, R.N. 2000. Stereoselective Biocatalysis. Marcel Dekker, New York. Pedersen, D.S. and Rosenbohm, C. 2001. Dry column vacuum chromatography. Synthesis 24312434.
Reese, C.B. and Song, Q. 1999. The H-phosphonate approach to the solution phase synthesis of linear and cyclic oligoribonucleotides. Nucl. Acids Res. 27:963-971. Reese, C.B. and Yan, H. 2001. Solution phase synthesis of ISIS 2922 (Vitravene) by the modified H-phosphonate approach. J. Chem. Soc., Perkin Trans. 1 2619-2633. van Boom, J.H. and Burgers, P.M.J. 1976. Use of levulinic acid in the protection of oligonucleotides via the modified phosphotriester method: Synthesis of decaribonucleotide U-A-U-A-U-A-UA-U-A. Tetrahedron Lett. 17:4875-4878.
Contributed by Iv´an Lavandera, Javier Garc´ıa, Susana Fern´andez, Miguel Ferrero, and Vicente Gotor Universidad de Oviedo Oviedo, Spain Yogesh S. Sanghvi Rasayan, Inc. Encinitas, California
Pon, R.T., Yu, S., Guo, Z., and Sanghvi, Y.S. 1999. Multiple oligodeoxyribonucleotide syntheses on a reusable solid-phase CPG support via the hydroquinone-O,O -diacetic acid (Q-Linker) linker arm. Nucl. Acids Res. 27:1531-1538.
Enzymatic Regioselective Levulinylation of Nucleosides
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Current Protocols in Nucleic Acid Chemistry
Nucleobase Protection with Allyloxycarbonyl
UNIT 2.12
This unit includes protocols for preparation of four N-allyloxycarbonyl-protected 2 deoxyribonucleosides and three N-allyloxycarbonyl-protected ribonucleosides from the corresponding parent nucleosides, including steps for OH-protection with 4,4 -dimethoxytrityl (DMTr) and tert-butyldimethylsilyl (TBDMS). The protected 2 -deoxyribonucleosides include N6 -AOC-5 -O-DMTr-2 -deoxyadenosine, N4 -AOC-5 O-DMTr-2 -deoxycytidine, N2 -AOC-5 -O-DMTr-2 -deoxyguanosine, and N2 -AOC-O6 allyl-5 -O-DMTr-2 -deoxyguanosine. The protected ribonucleosides include N6 -AOC2 -O-TBDMS-5 -O-DMTr-adenosine, N4 -AOC-2 -O-TBDMS-5 -O-DMTr-cytidine, and N2 -AOC-O6 -allyl-2 -O-TBDMS-5 -O-DMTr-guanosine. CAUTION: All operations involving organic solvents and reagents should be carried out in a well-ventilated chemical fume hood, and gloves and protective glasses should be worn.
SYNTHESIS OF N6 -ALLYLOXYCARBONYL-5 -O(4,4 -DIMETHOXYTRITYL)-2 -DEOXYADENOSINE
BASIC PROTOCOL 1
This compound is prepared from 2 -deoxyadenosine through a reaction sequence consisting of (1) 5 -O-dimethoxytritylation using 4,4 -dimethoxytrityl chloride in the presence of triethylamine and 4-(dimethylamino)pyridine in pyridine to give 5 -O-DMTr-2 deoxyadenosine, (2) 3 -O-protection by treatment with tert-butyldimethylsilyl chloride in the presence of imidazole to afford 3 -O-TBDMS-5 -O-DMTr-2 -deoxyadenosine, (3) N-allyloxycarbonylation using 1-(allyloxycarbonyl)tetrazole to provide N6 -AOC-3 O-TBDMS-5 -O-DMTr-2 -deoxyadenosine, and (4) removal of the TBDMS protecting group by exposure to tetrabutylammonium fluoride. The overall reaction sequence is illustrated in Figure 2.12.1. All operations are carried out at room temperature unless otherwise noted.
Materials 2 -Deoxyadenosine (S.1), 99% (Mitsui Chemicals) Triethylamine, distilled from CaH2 4-(Dimethylamino)pyridine (DMAP), 99% (Aldrich, TCI) Pyridine, distilled from CaH2 4,4 -Dimethoxytrityl chloride (DMTr-Cl), 97% (Aldrich, TCI) Ethyl acetate, 99% (Aldrich, Kishida) Sodium sulfate, 99% (Aldrich, Nacarai Tesque) N,N -Dimethylformamide (DMF), distilled from 4A molecular sieves Imidazole, 99% (Aldrich, Nacarai Tesque) tert-Butyldimethylsilyl chloride (TBDMS-Cl), 99% (Aldrich, Shinetsu) Hexane, 99% (Aldrich, Kishida) Brine: saturated solution of sodium chloride (99%; Aldrich, Wako) Silica gel 60 (spherical, neutrality, particle size 75 µm; Merck, Nacarai Tesque) Methanol, 99% (Aldrich, Kishida) Dichloromethane, 99% (Aldrich, Nacarai Tesque) 1-(Allyloxycarbonyl)tetrazole (AOC-Tet; see recipe) Tetrahydrofuran (THF), distilled from sodium benzophenone ketyl Aqueous saturated sodium hydrogencarbonate (NaHCO3 ; 99%; Aldrich, Nacarai Tesque) 1.0 M tetrabutylammonium fluoride in THF (Aldrich) Chloroform, 99% (Aldrich, Wako)
Contributed by Mamoru Hyodo and Yoshihiro Hayakawa Current Protocols in Nucleic Acid Chemistry (2005) 2.12.1-2.12.26 C 2005 by John Wiley & Sons, Inc. Copyright
Protection of Nucleosides for Oligonucleotide Synthesis
2.12.1 Supplement 23
Figure 2.12.1
Preparation of N 6 -AOC-5 -O-DMTr-2 -deoxyadenosine.
Glass column Diaphragm pump Rotary evaporator equipped with a diaphragm pump and cooling unit Vacuum oil pump TLC plate: silica-coated glass plate with fluorescent indicator (Merck silica gel 60 F254 ) 254-nm UV lamp Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) Prepare 3 -O-TBDMS-5 -O-DMTr-2 -deoxyadenosine 1. Prepare a solution of 3.05 g (12.1 mmol) 2 -deoxyadenosine (S.1), 2.50 mL (17.9 mmol) triethylamine, and 80.5 mg (0.659 mmol) DMAP in 100 mL pyridine. 2. Add 5.72 g (16.9 mmol) DMTr-Cl and stir the mixture at 13◦ C for 12 hr in the dark. 3. Pour the reaction mixture into 650 mL water. 4. Extract the whole mixture once with 150 mL and then twice with 100 mL ethyl acetate. Combine the organic layers and dry the organic solution over ∼15 g sodium sulfate. Remove the drying agent by filtration through a glass column with a cotton stopper under vacuum from a diaphragm pump. 5. Evaporate the resulting organic solution to dryness using a rotary evaporator with a diaphragm pump and a cooling unit. 6. Dry residue for 5 hr using a vacuum oil pump. 7. Dissolve the dried material in 50 mL DMF. Add 1.02 g (15.0 mmol) imidazole and 2.65 g (17.6 mmol) TBDMS-Cl, and stir the resulting mixture at 13◦ C for 4.5 hr. 8. Quench the reaction by adding 250 mL water. 9. Extract twice with 100 mL and then once with 50 mL of 1:1 (v/v) hexane/ethyl acetate, and combine the organic extracts. 10. Wash the organic extracts with 50 mL brine and dry the organic solution over sodium sulfate. 11. Concentrate the dried organic solution using a rotary evaporator.
Nucleobase Protection with Allyloxycarbonyl
12. Purify residue by column chromatography (APPENDIX 3E) on a column of 150 g silica gel with 1:30 (v/v) methanol/dichloromethane as eluent. Monitor fractions by TLC (APPENDIX 3D) using 1:20 (v/v) methanol/dichloromethane (Rf = 0.25). Visualize under UV light (254 nm).
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13. Collect all fractions containing the desired product. Evaporate to dryness using a rotary evaporator. 3 -O-(tert-Butyldimethylsilyl)-5 -O-(4,4 -dimethoxytrityl)-2 -deoxyadenosine (S.2): 5.1 g (63%) as white powder. m.p. 151◦ –152◦ C. TLC (1:20 methanol/dichloromethane): Rf = 0.25. IR (KBr) 3330, 3160, 2960, 2940, 2860, 1670, 1650, 1510, 1260, 1180 cm–1 . UV λmax : 236 nm (ε 23,400), 260 (16,100). 1 H NMR: 0.03, 0.33 (2 s, 6H, Si(CH3 )2 ), 0.89 (s, 9H, Si-tert-C4 H9 ), 2.01 (ddd, 1H, J = 3.6, 6.0, and 13.2 Hz, H2 ), 2.81 (dt, 1H, J = 6.0 and 13.2 Hz, H2 ), 3.36 (t, 2H, J = 4.5 Hz, H5 ), 3.82 (s, 6H, 2 × OCH3 ), 4.12 (dd, 1H, J = 4.5 and 7.5 Hz, H4 ), 4.63 (m, 1H, H3 ), 5.83 (br, 2H, NH2 ), 6.46 (t, 1H, J = 6.0 Hz, H1 ), 6.73–6.95 (m, 4H, protons ortho to OCH3 of DMTr), 7.20–7.53 (m, 9H, arom. protons of DMTr), 8.06 (s, 1H, H2), 8.35 (s, 1H, H8).
Prepare N6 -AOC-5 -O-DMTr-2 -deoxyadenosine 14. Add 18.0 g (0.027 mol) S.2 to a solution of ∼0.1 mol AOC-Tet in 150 mL THF, and stir the resulting mixture at 70◦ C for 3 hr. 15. Cool the reaction mixture to 25◦ C and rotary evaporate. 16. Dissolve residue in 300 mL ethyl acetate and pour the resulting solution into 100 mL of an aqueous solution saturated with NaHCO3 . 17. Extract once with 300 mL ethyl acetate, wash the organic layer with 50 mL brine, and dry over sodium sulfate. 18. Evaporate the solvent using a rotary evaporator. 19. Dissolve residue in 100 mL THF and add 50 mL (0.05 mol) of 1.0 M TBAF in THF. Stir the mixture for 12 hr. 20. Evaporate the solvent with a rotary evaporator and dissolve residue in 150 mL ethyl acetate. 21. Wash the organic solution with 50 mL brine and dry over sodium sulfate. 22. Concentrate the dried organic solution using a rotary evaporator. 23. Purify residue by column chromatography over 300 g silica gel using an eluent of trace:1:50 (v/v/v) to trace:1:30 (v/v/v) triethylamine/methanol/chloroform. Monitor fractions by TLC using 1:20 (v/v) methanol/chloroform (Rf = 0.33). 24. Collect fractions including the desired product and concentrate using a rotary evaporator. N6 -Allyloxycarbonyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxyadenosine (S.3): 14.4 g (84%) as white powder. m.p. 84◦ –86◦ C. TLC (1:20 methanol/chloroform): Rf = 0.33. IR (KBr) 3410, 2960, 2940, 1760, 1620, 1515, 1470, 1255, 1220, 1180 cm−1 . UV λmax : 236 nm (ε 23,300), 268 (19,100). 1 H NMR: 2.37–3.04 (m, 2H, H2 ), 3.43 (br d, 2H, J = 5.1 Hz, H5 ), 3.76 (s, 6H, 2 × OCH3 ), 4.24 (br d, 1H, J = 3.0 Hz, H4 ), 4.74 (d, 3H, J = 5.7 Hz, CH2 CH= =CH2 and H3 ), 5.25 (dd, 1H, J = 1.5 and 10.2 Hz, cis-CH2 CH= =CH2 ), 5.37 (dd, 1H, J = 1.5 and 17.4 Hz, trans-CH2 CH= =CH2 ), 6.00 (ddt, 1H, J = 10.2, 17.4, and 5.7 Hz, CH2 CH= =CH2 ), 6.52 (t, 1H, J = 6.0 Hz, H1 ), 6.80 (d, 4H, J = 9.0 Hz, protons ortho to OCH3 of DMTr), 7.13–7.50 (m, 9H, arom. protons of DMTr), 8.18 (s, 1H, H2), 8.73 (s, 1H, H8).
SYNTHESIS OF N4 -ALLYLOXYCARBONYL-5 -O(4,4 -DIMETHOXYTRITYL)-2 -DEOXYCYTIDINE This compound is synthesized from 2 -deoxycytidine hydrochloride via (1) conversion of the starting nucleoside to N4 -AOC-2 -deoxycytidine by successive treatment with trimethylsilyl chloride in pyridine, allyl 1-benzotriazolyl carbonate, and an aqueous NaHCO3 solution without isolation of the intermediary that is formed, and (2) dimethoxytritylation using 4,4 -dimethoxytrityl chloride in pyridine to afford
BASIC PROTOCOL 2
Protection of Nucleosides for Oligonucleotide Synthesis
2.12.3 Current Protocols in Nucleic Acid Chemistry
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Figure 2.12.2
Preparation of N 4 -AOC-5 -O-DMTr-2 -deoxycytidine.
N4 -AOC-5 -O-DMTr-2 -deoxycytidine. The overall reaction sequence is illustrated in Figure 2.12.2. All operations are carried out at room temperature unless otherwise noted.
Materials 2 -Deoxycytidine hydrochloride (S.4), 99% (Aldrich, Mitsui Chemicals) Pyridine, distilled from CaH2 Trimethylsilyl chloride (TMS-Cl), 99% (Aldrich, Shinetsu) Allyl 1-benzotriazolyl carbonate (AOCOBT; see recipe) Saturated aqueous sodium hydrogencarbonate (NaHCO3 ), 99% (Aldrich, Nacarai Tesque) Dichloromethane, 99% (Aldrich, Nacarai Tesque) Sodium sulfate, 99% (Aldrich, Nacarai Tesque) 4,4 -Dimethoxytrityl chloride (DMTr-Cl), 97% (Aldrich, TCI) Hexane, 99% (Aldrich, Kishida) Ethyl acetate, 99% (Aldrich, Kishida) Brine: saturated aqueous sodium chloride (99%; Aldrich, Wako) Silica gel 60 (spherical, neutrality, particle size 75 µm; Merck, Nacarai Tesque) Chloroform, 99% (Aldrich, Wako) Methanol, 99% (Aldrich, Kishida) Glass column Diaphragm pump Rotary evaporator equipped with a diaphragm pump and cooling unit Vacuum oil pump TLC plate: silica-coated glass plate with fluorescent indicator (Merck silica gel 60 F254 ) 254-nm UV lamp Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) Prepare crude N4 -AOC-2 -deoxycytidine 1. Add 7.5 mL (59.1 mmol) trimethylsilyl chloride to a suspension of 3.09 g (11.7 mmol) 2 -deoxycytidine hydrochloride (S.4) in 30 mL pyridine. Stir the mixture at 10◦ C for 1.5 hr. 2. Add 5.14 g (23.5 mmol) AOCOBT and stir the mixture for another 24 hr at 0◦ C. 3. Add 50 mL saturated NaHCO3 solution and continue stirring for an additional 30 min at room temperature. Nucleobase Protection with Allyloxycarbonyl
4. Pour the reaction mixture into 100 mL water and extract the resulting aqueous mixture with 30 mL dichloromethane.
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5. Separate the aqueous layer and extract four times with 50 mL of 1:1 (v/v) dichloromethane/pyridine. 6. Dry the organic layer over ∼15 g sodium sulfate. Remove the drying agent by filtration through a glass column with a cotton stopper under vacuum from a diaphragm pump. Concentrate the organic solution using a rotary evaporator with a diaphragm pump and a cooling unit.
Prepare N4 -AOC-5 -O-DMTr-2 -deoxycytidine 7. Dissolve the residue of crude S.5 in 30 mL pyridine. 8. Add 3.96 g (11.7 mmol) DMTr-Cl and stir the resulting solution at 10◦ C for 24 hr. 9. Pour the reaction mixture into 300 mL water and extract three times with 1:1 (v/v) hexane/ethyl acetate. 10. Combine the organic layers and wash with 100 mL brine. 11. Dry the organic solution over sodium sulfate and concentrate the dried solution with a rotary evaporator. 12. Purify the crude residue on a 100 g silica gel column (APPENDIX 3E) with 1:50 (v/v) methanol/chloroform as eluent. Monitor fractions by TLC (APPENDIX 3D) using 1:20 (v/v) methanol/chloroform (Rf = 0.24). Visualize the product under UV light (254 nm). 13. Collect fractions containing the desired product and concentrate using a rotary evaporator. N4 -Allyloxycarbonyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxycytidine (S.6): 4.97 g (69%) as white solid. m.p. 98◦ –102◦ C (4.97 g, 69%). TLC (1:20 methanol/chloroform): Rf = 0.24. IR (KBr) 3400, 2940, 1750, 1650, 1630, 1610, 1560, 1510, 1255, 1210, 1100, 1040 cm−1 . UV λmax : 237 nm (ε 30,600), 285 (8,400). 1 H NMR: 2.28 (m, 1H, H2 ), 2.81 (m, 1H, H2 ), 3.46 (br, 2H, H5 ), 3.80 (s, 6H, 2 × OCH3 ), 4.21 (m, 1H, H4 ), 4.40–4.80 (m, 3H, CH2 CH= =CH2 and H3 ), 5.15–5.51 (m, 2H, CH2 CH= =CH2 ), 5.96 (m, 1H, CH2 CH= =CH2 ), 6.32 (t, 1H, J = 5.7 Hz, H1 ), 6.70–7.55 (m, 14H, H5 and arom. protons of DMTr), 8.27 (d, 1H, J = 7.5 Hz, H6).
SYNTHESIS OF N2 -ALLYLOXYCARBONYL-5 -O(4,4 -DIMETHOXYTRITYL)-2 -DEOXYGUANOSINE
BASIC PROTOCOL 3
This compound is prepared starting from 2 -deoxyguanosine via a reaction sequence consisting of (1) 5 -O-tert-butyldimethylsilylation using tert-butyldimethylsilyl chloride in the presence of imidazole to afford 3 ,5 -di-O-TBDMS-2 -deoxyguanosine, (2) N-allyloxycarbonylation by treatment with tert-butylmagnesium chloride, HMPA, and allyloxycarbonyl chloride to produce N2 -AOC-3 ,5 -di-O-TBDMS-2 -deoxyguanosine, (3) removal of the silyl protecting groups using tetrabutylammonium fluoride to give N2 -AOC-2 -deoxyguanosine, and (4) 5 -O-dimethoxytritylation by treatment with 4,4 dimethoxytrityl chloride in pyridine to provide N2 -AOC-5 -O-DMTr-2 -deoxyguanosine. The overall reaction sequence is illustrated in Figure 2.12.3. All operations are carried out at room temperature unless otherwise noted.
Materials 2 -Deoxyguanosine (S.7), 99% (Aldrich, Mitsui Chemicals) Imidazole, 99% (Aldrich, Nacarai Tesque) N,N -Dimethylformamide (DMF), distilled from 4A molecular sieves tert-Butyldimethylsilyl chloride (TBDMS-Cl), 99% (Aldrich, Shinetsu) Ethanol, 99.5% (Aldrich, Nacarai Tesque) tert-Butylmagnesium chloride, 99% (Aldrich)
Protection of Nucleosides for Oligonucleotide Synthesis
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Figure 2.12.3
Preparation of N 2 -AOC-5 -O-DMTr-2 -deoxyguanosine.
Tetrahydrofuran (THF), distilled from sodium benzophenone ketyl Hexamethylphosphoric triamide (HMPA), 99% (Aldrich) Allyloxycarbonyl chloride (AOC-Cl), 99% (Aldrich, TCI) Methanol, 99% (Aldrich, Kishida) Diethyl ether, distilled from sodium benzophenone ketyl 0.15 M ethylenediamine tetraacetic acid (EDTA; 99%; Aldrich, Nacarai Tesque) Saturated aqueous sodium hydrogencarbonate (NaHCO3 ; 99%; Aldrich, Nacarai Tesque) Brine: saturated aqueous sodium chloride (99%; (Aldrich, Wako) Sodium sulfate, 99% (Aldrich, Nacarai Tesque) Silica gel 60 (spherical, neutrality, particle size 75 µm; Merck, Nacarai Tesque) Ethyl acetate, 99% (Aldrich, Kishida) Hexane, 99% (Aldrich, Kishida) 1.0 M tetrabutylammonium fluoride (TBAF) in THF (Aldrich) Pyridine, distilled from CaH2 Dichloromethane, 99% (Aldrich, Nacarai Tesque) 4,4 -Dimethoxytrityl chloride (DMTr-Cl), 97% (Aldrich, TCI) Chloroform, 99% (Aldrich, Wako) Rotary evaporator equipped with a diaphragm pump and cooling unit Glass column Diaphragm pump Vacuum oil pump TLC plate: silica-coated glass plate with fluorescent indicator (Merck silica gel 60 F254 ) 254-nm UV lamp Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) Nucleobase Protection with Allyloxycarbonyl
2.12.6 Supplement 23
Prepare 3 ,5 -di-O-TBDMS-2 -deoxyguanosine 1. Prepare a solution of 10.0 g (35 mmol) 2 -deoxyguanosine (S.7) and 17 g (247 mmol) imidazole in 20 mL DMF. 2. Add 21 g (139 mmol) TBDMS-Cl and stir at room temperature for 24 hr. Current Protocols in Nucleic Acid Chemistry
3. Quench the reaction by adding 50 mL ethanol and evaporate the mixture on a rotary evaporator with a diaphragm pump and a cooling unit. 4. Wash the residual solid with 200 mL cold (4◦ C) ethanol and then 200 mL water. Dry the solid under reduced pressure with a vacuum pump. 3 ,5 -Di-O-(tert-butyldimethylsilyl)-2 -deoxyguanosine (S.8): 14.9 g (86%). 1 H NMR (DMSO-d6 ): 0.02, 0.08 (2 s, 12H), 0.85, 0.87 (2 s, 18H), 2.17–2.22 (m, 1H), 2.26 (s, 1H), 2.59–2.66 (m, 1H), 3.60–3.77 (m, 3H), 3.77–3.83 (m, 1H), 6.05–6.08 (m, 1H), 6.43 (br, 2H), 7.84 (s, 1H), 10.57 (br, 1H).
Prepare N2 -AOC-3 ,5 -di-O-TBDMS-2 -deoxyguanosine 5. Add dropwise 16.0 mL (20.3 mmol) of 1.27 M tert-butylmagnesium chloride in THF to a solution of 3.40 g (6.86 mmol) S.8 and 10 mL HMPA in 70 mL THF. Stir the mixture at 25◦ C for 20 min. CAUTION: HMPA is a suspected carcinogen. Handle with gloves in a well-ventilated chemical fume hood.
6. Add a solution of 1.80 mL (17.0 mmol) AOC-Cl in 10 mL THF and continue stirring for an additional 15 min. 7. Quench the reaction by adding 5 mL methanol and evaporate the reaction mixture on a rotary evaporator. 8. Dissolve the residue in 200 mL diethyl ether. 9. Wash the ether solution once with 150 mL of 0.15 M EDTA, once with 50 mL saturated aqueous NaHCO3 , and once with 50 mL brine. 10. Dry the ether solution over ∼15 g sodium sulfate. Remove the drying agent by filtration through a glass column with a cotton stopper under vacuum from a diaphragm pump. Concentrate the dried solution. 11. Subject the residue to column chromatography (APPENDIX 3E) using a 100 g silica gel column and 3:2 (v/v) hexane/ethyl acetate as eluent. Monitor fractions by TLC (APPENDIX 3D) with 1:1 (v/v) hexane/ethyl acetate (Rf = 0.36). Visualize the product under UV light (254 nm). 12. Collect fractions containing the desired product and concentrate the fractions using a rotary evaporator. N2 -Allyloxycarbonyl-3 ,5 -di-O-(tert-butyldimethylsilyl)-2 -deoxyguanosine (S.9): 3.09 g (78%) as a foam. TLC (1:1 ethyl acetate/hexane): Rf = 0.36. IR (KBr) 3250, 2950, 2930, 2850, 1715, 1690, 1610, 1240, 1100 cm−1 . UV λmax : 257 nm (ε 14,700), 280 (sh). 1 H NMR: 0.06, 0,08 (2 s, 12H, 2 × Si(CH3 )2 ), 0.90 (s, 18H, 2 × Si-tert-C4 H9 ), 1.90 (m, 1H, H2 ), 2.40 (m, 1H, H2 ), 3.76 (d, 2H, J = 3.6 Hz, H5 ), 3.98 (dd, 1H, J = 3.6 and 6.6 Hz, H4 ), 4.57 (dd, 1H, J = 3.9 and 7.5 Hz, H3 ), 4.73 (d, 2H, J = 6.0 Hz, CH2 CH= =CH2 ), =CH2 ), 5.37 (dd, 1H, J = 1.8 and 17.4 Hz, 5.32 (dd, 1H, J = 1.8 and 10.5 Hz, cis-CH2 CH= trans-CH2 CH= =CH2 ), 5.94 (ddt, 1H, J = 10.5, 17.4, and 6.0 Hz, CH2 CH= =CH2 ), 6.23 (t, 1H, J = 6.3 Hz, H1 ), 7.95 (s, 1H, H8).
Prepare N2 -AOC-5 -O-DMTr-2 -deoxyguanosine 13. Add 20 mL (20 mmol) of 1.0 M TBAF in THF to a solution of 2.20 g (3.79 mmol) S.9 in 20 mL THF. Stir the resulting mixture at 10◦ C for 30 min. 14. Concentrate the mixture and add 20 mL water to the resulting residue. 15. Extract the aqueous mixture three times with 50 mL of 1:1 (v/v) dichloromethane/pyridine and combine the organic layers. 16. Dry the organic extracts over sodium sulfate and then concentrate to dryness with a rotary evaporator.
Protection of Nucleosides for Oligonucleotide Synthesis
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17. Dissolve the residual oil in 30 mL pyridine and add 1.32 g (3.90 mmol) DMTr-Cl. Stir the resulting mixture at 10◦ C for 15 hr. 18. Pour the mixture into 300 mL water and extract three times with 100 mL of 1:1 (v/v) hexane/ethyl acetate. 19. Dry the combined organic layers over sodium sulfate and concentrate them using a rotary evaporator. 20. Subject the residue to column chromatography on a 70 g silica gel column and elute with 1:30 (v/v) methanol/chloroform. Monitor fractions by TLC using 1:10 (v/v) methanol/chloroform (Rf = 0.72). 21. Collect fractions containing the desired product and concentrate them using a rotary evaporator. N2 -Allyloxycarbonyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxyguanosine (S.10): 1.62 g (65%) as a white solid. m.p. 124◦ –127◦ C. TLC (1:10 methanol/chloroform): Rf = 0.72. IR (KBr) 3400, 3250, 2970, 2940, 1700, 1615, 1570, 1515, 1250, 1180, 1100, 1040 cm−1 . UV λmax : 234 nm (ε 33,300), 276 (sh). 1 H NMR: 2.47–2.90 (m, 2H, H2 ), 3.40 (br, 2H, H5 ), 3.72 (s, 6H, 2 × OCH3 ), 4.28 (m, 1H, H4 ), 4.80 (br d, 2H, J = 4.8 Hz, CH2 CH= =CH2 ), 5.05 =CH2 ), 5.74–6.40 (m, 2H, CH2 CH= =CH2 and (m, 1H, H3 ), 5.18–5.53 (m, 2H, CH2 CH= H1 ), 6.74 (d, 4H, J = 9.9 Hz, protons ortho to OCH3 of DMTr), 7.02–7.53 (m, 9H, arom. protons of DMTr), 7.77 (s, 1H, H8). BASIC PROTOCOL 4
SYNTHESIS OF N2 -ALLYLOXYCARBONYL-O6 -ALLYL-5 -O(4,4 -DIMETHOXYTRITYL)-2 -DEOXYGUANOSINE This O6 -protected compound is synthesized from N2 -AOC-3 ,5 -di-O-TBDMS-2 deoxyguanosine (S.9) prepared as in the previous protocol. The reaction sequence includes (1) O6 -allylation of S.9 by treatment with 2-mesitylenesulfonyl chloride in the presence of triethylamine and 4-(dimethylamino)pyridine followed by allyl alcohol in the presence of trimethylamine and 1,8-diazabicyclo[5,4,0]undec-7-ene to afford N2 AOC-O6 -allyl-3 ,5 -di-O-TBDMS-2 -deoxyguanosine, (2) removal of the silyl protecting groups using tetrabutylammonium fluoride to give N2 -AOC-O6 -allyl-2 -deoxyguanosine, and (3) O-dimethoxytritylation by treatment with 4,4 -dimethoxytrityl chloride in pyridine to provide N2 -AOC-O6 -allyl-5 -O-DMTr-2 -deoxyguanosine. The overall reaction sequence is illustrated in Figure 2.12.4. All operations are carried out at room temperature unless otherwise noted.
Figure 2.12.4
Preparation of N 2 -AOC-O 6 -allyl-5 -O-DMTr-2 -deoxyguanosine.
Nucleobase Protection with Allyloxycarbonyl
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Materials N2 -Allyloxycarbonyl-3 ,5 -di-O-(tert-butyldimethylsilyl)-2 -deoxyguanosine (S.9; see Basic Protocol 3) Triethylamine, distilled from CaH2 4-(Dimethylamino)pyridine (DMAP), 99% (Aldrich, TCI) Dichloromethane, 99% (Aldrich, Nacarai Tesque) 2-Mesitylenesulfonyl chloride, 99% (Aldrich, TCI) Saturated aqueous sodium hydrogencarbonate (NaHCO3 ; 99%, Aldrich, Nacarai Tesque) Sodium sulfate, 99% (Aldrich, Nacarai Tesque) Trimethylamine, anhydrous gas (Aldrich) Allyl alcohol, distilled from Mg 1,8-Diazabicyclo[5,4,0]undec-7-ene (DBU), 99% (Aldrich, Nacarai Tesque) Brine: saturated aqueous sodium chloride (99%; Aldrich, Wako) Silica gel 60 (spherical, neutrality, particle size 75 µm; Merck, Nacarai Tesque) Ethyl acetate, 99% (Aldrich, Kishida) Hexane, 99% (Aldrich, Kishida) 1.0 M tetrabutylammonium fluoride (TBAF) in THF (Aldrich, Aldrich) Tetrahydrofuran (THF), distilled from sodium benzophenone ketyl Pyridine, distilled from CaH2 4,4 -Dimethoxytrityl chloride (DMTr-Cl), 97% (Aldrich, TCI) Methanol, 99% (Aldrich, Kishida) Chloroform, 99% (Aldrich, Wako) Glass column Diaphragm pump Rotary evaporator equipped with a diaphragm pump and cooling unit Vacuum oil pump TLC plate: silica-coated glass plate with fluorescent indicator (Merck silica gel 60 F254 ) 254-nm UV lamp Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) Prepare N2 -AOC-O6 -allyl-3 ,5 -di-O-TBDMS-2 -deoxyguanosine 1. Prepare a mixture of 11.2 g (19.4 mmol) N2 -AOC-3 ,5 -di-O-TBDMS-2 -deoxyguanosine (S.9), 11 mL (7.99 g, 78.9 mmol) triethylamine, and 278 mg (2.27 mmol) DMAP in 100 mL dichloromethane. 2. Add 5.54 g (25.3 mmol) 2-mesitylenesulfonyl chloride and stir the mixture at room temperature for 3 hr. 3. Pour the mixture into 200 mL dichloromethane. Wash twice with 100 mL water and twice with 100 mL of saturated aqueous NaHCO3 solution. 4. Dry the organic layer over ∼15 g sodium sulfate. Remove the drying agent by filtration through a glass column with a cotton stopper under vacuum from a diaphragm pump. Remove the organic solvent using a rotary evaporator with a diaphragm pump and a cooling unit. Dry the residue with a vacuum oil pump. 5. Dissolve the dried material in 100 mL dichloromethane. 6. Add 27 mL (17.7 g, 300 mmol) trimethylamine followed by 20 mL (17.1 g, 290 mmol) allyl alcohol at 0◦ C with stirring. Continue stirring at the same temperature for 10 min. 7. Add 3 mL (3.05 g, 20.1 mmol) DBU and stir the resulting mixture overnight at 0◦ C. Current Protocols in Nucleic Acid Chemistry
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8. Warm the mixture to room temperature and remove the resulting precipitates by filtering through a glass column with a cotton stopper under vacuum with a diaphragm pump. 9. Dilute the filtrate with 200 mL dichloromethane. 10. Wash the dichloromethane solution once with 200 mL water and twice with 200 mL brine. 11. Concentrate the dichloromethane solution with a rotary evaporator. 12. Purify the residue by column chromatography (APPENDIX 3E) on a 100 g silica gel column using 1:2 (v/v) ethyl acetate/hexane as eluent. Monitor fractions by TLC (APPENDIX 3D) using 1:1 (v/v) ethyl acetate/hexane (Rf = 0.40). Visualize under UV light (254 nm). 13. Combine fractions containing the desired product and evaporate them with a rotary evaporator. N 2 -Allyloxycarbonyl-O6 -allyl-3 ,5 -di-O-(tert-butyldimethylsilyl)-2 -deoxyguanosine (S.11): 9.64 g (80%) as a foam. TLC (1:1 ethyl acetate/hexane): Rf = 0.40. IR 3418, 1661, 1537 cm−1 . UV λmax : 284 nm (ε 7,300), 207 (sh, 20,000). 1 H NMR: 0.06 (s, 6H, (CH3 )2 Si), 0.87 (s, 9H, t-C4 H9 Si), 1.95 (s, 3H, CH3 C= =), 2.27 (ddd, 1H, J = 13.4, 6.4, and 3.9 Hz, H2 ), 2.47 (dt, 1H, J = 13.4 and 6.4 Hz, H2 ), 3.23 (dd, 1H, J = 6.2 and 3.1 Hz, 5 -OH), 3.71–3.78 (m, 1H, H5 ), 3.91–3.95 (m, 2H, H4 and H5 ), 4.50 (dt, 1H, J = 6.2 and 3.9 Hz, H3 ), 4.88 (d, 2H, J = 5.9 Hz, CH2=CHCH2 ), 5.26 (dd, 1H, J = 10.4 and 1.5 Hz, cis-CH2=CHCH2 ), 5.37 (dd, 1H, J = 17.3 and 1.5 Hz, trans-CH2=CHCH2 ), 5.95–6.10 (m, 2H, CH2=CHCH2 and H1 ), 7.62 (s, 1H, H6). Anal. calcd. for C19 H32 N2 O5 Si: C, 57.53; H, 8.15; N, 7.06; found: C, 57.35; H, 8.29; N, 7.23.
Prepare N2 -AOC-O6 -allyl-5 -O-DMTr-2 -deoxyguanosine 14. Add 20 mL (20 mmol) of 1.0 M TBAF in THF to a solution of 2.35 g (3.79 mmol) S.11 in THF (20 mL). Stir the resulting mixture at 10◦ C for 30 min. 15. Concentrate the mixture with a rotary evaporator and add 20 mL water to the residue. 16. Extract the aqueous mixture three times with 50 mL of 1:1 (v/v) dichloromethane/pyridine and dry the combined organic extracts over sodium sulfate. 17. Concentrate to dryness using a vacuum oil pump. 18. Dissolve the residue in 30 mL pyridine and add 1.32 g (3.90 mmol) DMTr-Cl. Stir the mixture at 10◦ C for 15 hr. 19. Pour the reaction mixture into 300 mL water and extract three times with 100 mL of 1:1 (v/v) hexane/ethyl acetate. 20. Dry the combined organic extracts with sodium sulfate and concentrate with a rotary evaporator. 21. Subject the residue to column chromatography with a 70 g silica gel column and 1:30 (v/v) methanol/chloroform as eluent. Monitor fractions by TLC with 1:10 (v/v) methanol/chloroform (Rf = 0.35). 22. Collect fractions containing the desired product and concentrate using a rotary evaporator.
Nucleobase Protection with Allyloxycarbonyl
N 2 -Allyloxycarbonyl-O6 -allyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxyguanosine (S.12): 1.58 g (60%) as an amorphous solid. m.p. = 76◦ C. TLC (1:10 methanol/chloroform): Rf = 0.35. UV λmax : 254 nm (shoulder, ε 12,900), 268 (14,000). 1 H NMR (400 MHz, DMSO-d6 ): 2.23–2.29 (m, 1H), 2.69–2.75 (m, 1H), 3.47–3.61 (m, 2H), 3.82–3.85 (m, 1H), 4.41 (m, 1H), 4.61–4.62 (m, 2H), 4.85 (t, 1H, J = 5.6 Hz), 5.05 (d, 2H, J = 5.9 Hz),
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5.21–5.48 (m, 5H), 5.92–6.02 (m, 1H), 6.10–6.20 (m, 1H), 6.20–6.32 (m, 1H), 8.41 (s, 1H), 10.32 (s, 1H). Anal. calcd. for C17 H21 N5 O6 : C, 52.17; H, 5.41; N, 17.89; found: C, 52.18; H, 5.65; N, 17.80.
SYNTHESIS OF N6 -ALLYLOXYCARBONYL-2 -O-(tertBUTYLDIMETHYLSILYL)-5 -O-(4,4 -DIMETHOXYTRITYL)ADENOSINE
BASIC PROTOCOL 5
This compound is prepared from adenosine via the following sequence of reactions: (1) N6 -allyloxycarbonylation by successive treatment with hexamethyldisilazane in the presence of ammonium sulfate as a catalyst, with allyloxycarbonyl chloride in the presence of N-methylimidazole, and with triethylamine in methanol to give N6 -AOC-adenosine, (2) 5 -O-dimethoxytritylation by reaction with 4,4 -dimethoxytrityl chloride in pyridine to provide N6 -AOC-5 -O-DMTr-adenosine, (3) tert-butyldimethylsilylation using tert-butyldimethylsilyl chloride in the presence of imidazole to afford a mixture of N6 AOC-2 -O-TBDMS-5 -O-DMTr-adenosine and its 3 -O-TBDMS isomer, and (4) chromatographic separation of this mixture on a silica gel column to furnish N6 -AOC-2 -OTBDMS-5 -O-DMTr-adenosine. Following separation, the 3 -O-silylated byproduct is isomerized to increase the yield of the desired 2 -isomer. The overall reaction sequence is illustrated in Figure 2.12.5. All operations are carried out at room temperature unless otherwise noted.
Materials Hexamethyldisilazane (HMDS), 99% (Aldrich, Shinetsu) Adenosine (S.13), 99% (Sigma) Ammonium sulfate, 99% (Aldrich, Wako) 1,4-Dioxane, distilled from sodium benzophenone ketyl Toluene, distilled from sodium benzophenone ketyl
Figure 2.12.5
Preparation of N 6 -AOC-2 -O-TBDMS-5 -O-DMTr-adenosine. Protection of Nucleosides for Oligonucleotide Synthesis
2.12.11 Current Protocols in Nucleic Acid Chemistry
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Dichloromethane, 99% (Aldrich, Nacarai Tesque) N-Methylimidazole (Aldrich, Nacarai Tesque) Allyloxycarbonyl chloride (AOC-Cl), 99% (Aldrich, TCI) 1.0 M phosphate buffer, pH 7 Sodium sulfate, 99% (Aldrich, Nacarai Tesque) Methanol, 99% (Aldrich, Kishida) Triethylamine, distilled from CaH2 Diethyl ether, 99% (Aldrich, Nacarai Tesque) Diphosphorus pentoxide (P2 O5 ), 98% (Aldrich, Nacarai Tesque) Pyridine, distilled from CaH2 N,N -Dimethylformamide (DMF), distilled from 4A molecular sieves 4,4 -Dimethoxytrityl chloride (DMTr-Cl), 97% (Aldrich, TCI) Silica gel 60 (spherical, neutrality, particle size 75 µm; Merck, Nacarai Tesque) Ethyl acetate, 99% (Aldrich, Kishida) Hexane, 99% (Aldrich, Kishida) Chloroform, 99% (Aldrich, Wako) Imidazole, 99% (Aldrich, Nacarai Tesque) tert-Butyldimethylsilyl chloride (TBDMS-Cl), 99% (Aldrich, Shinetsu) Glass column Diaphragm pump Reflux condenser Rotary evaporator equipped with a diaphragm pump and cooling unit Vacuum oil pump TLC plate: silica-coated glass plate with fluorescent indicator (Merck silica gel 60 F254 ) 254-nm UV lamp Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) Prepare N6 -AOC-adenosine 1. Add 130 mL HMDS to a suspension of 15.0 g (56 mmol) adenosine (S.13) and a catalytic amount of ammonium sulfate in 140 mL dry dioxane. Heat under reflux for 3 hr (b.p. 125◦ C; oil bath 145◦ C). 2. Evaporate the solvent on a rotary evaporator with a diaphragm pump and a cooling unit, and then dry the residue by co-evaporating twice with 50 mL dry toluene. 3. Dissolve residue in 500 mL dichloromethane and then add 14.5 mL (14.9 g, 180 mmol) N-methylimidazole followed by 18.5 mL (21.0 g, 170 mmol) AOC-Cl. Stir the mixture overnight. 4. Add more 4.50 mL (4.60 g, 56.0 mmol) N-methylimidazole and 6.00 mL (6.80 g, 56.0 mmol) AOC-Cl and stir the mixture for an additional 12 hr. 5. Pour into 300 mL of 1.0 M phosphate buffer, pH 7, and extract three times with 50 mL dichloromethane. 6. Combine the organic extracts and dry over ∼15 g sodium sulfate. Remove the drying agent by filtration through a glass column with a cotton stopper under vacuum from a diaphragm pump. Evaporate with a rotary evaporator. 7. Dissolve the residue in 500 mL methanol and add 130 mL triethylamine. Stir the mixture at room temperature for 12 hr. Nucleobase Protection with Allyloxycarbonyl
8. Collect precipitates by vacuum filtration through a glass column with a cotton stopper, and wash with 300 mL diethyl ether.
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9. Concentrate the filtrate to a volume of 300 mL. 10. Collect the resulting precipitates by vacuum filtration as above, and wash with 300 mL diethyl ether. 11. Combine all precipitates and dry them over P2 O5 under reduced pressure with a vacuum oil pump. N6 -(Allyloxycarbonyl)adenosine (S.14): 17.5 g (89%) as a white solid. m.p. 164◦ –166◦ C. TLC (1:9 methanol/dichloromethane): Rf = 0.3. UV λmax : 267 nm (ε 18,000). 1 H NMR (270 MHz, DMSO-d6 ): 3.55–3.71 (m, 2H), 3.95–3.97 (m, 1H), 4.17 (m, 1H), 4.61–4.66 (m, 3H), 5.12 (t, 1H, J = 5.6 Hz), 5.21–5.22 (m, 1H), 5.23 (d, 1H, J = 10.6 Hz), 5.38 (dd, 1H, J = 1.4 and 15.8 Hz), 5.52 (d, 1H, J = 6.3 Hz), 5.90–6.04 (m, 2H), 8.63 (s, 1H), 8.68 (s, 1H), 10.66 (s, 1H). Anal. calcd. for C14 H17 N5 O6 : C, 47.86; H, 4.88; N, 19.93; found: C, 47.86; H, 4.86; N, 20.14.
Prepare N6 -AOC-5 -O-DMTr-adenosine 12. Prepare a solution of 15.0 g (42.7 mmol) S.14 in 150 mL of 1:1 (v/v) pyridine/DMF and cool to 0◦ C. 13. Add 15.6 g (46.0 mmol) DMTr-Cl in small portions over 20 min. Stir at the same temperature for an additional 20 min and then at room temperature for 12 hr. 14. Pour into 1.2 L ice-cold water. Collect the precipitates by vacuum filtration as above, and wash with 200 mL water. 15. Dry at 50◦ C under vacuum with a vacuum oil pump. 16. Purify the solid by column chromatography (APPENDIX 3E) using a 300 g silica gel column. Monitor fractions by TLC (APPENDIX 3D) using 1:9 (v/v) methanol/dichloromethane. Visualize under UV light (254 nm). Elute the column first with 1.2 L of 4:6 (v/v) ethyl acetate/hexane to remove the byproducts, and then with 1:9 (v/v) methanol/ethyl acetate to collect the desired product (Rf = 0.76). 17. Evaporate the fractions containing the desired product using a rotary evaporator. N6 -Allyloxycarbonyl-5 -O-(4,4 -dimethoxytrityl)adenosine (S.15): 21.5 g (77%) as an amorphous solid. TLC (1:9 methanol/dichloromethane): Rf = 0.76. 1 H NMR (270 MHz, DMSO-d6 ): 3.22–3.24 (m, 2H), 3.71 (s, 6H), 4.06–4.11 (m, 1H), 4.30–4.34 (m, 1H), 4.64–4.66 (m, 2H), 4.73–4.78 (m, 1H), 5.25–5.27 (m, 2H), 5.42 (dd, 1H, J = 1.5 and 17.3 Hz), 5.60 (d, 1H, J = 5.3 Hz), 5.90–6.04 (m, 2H), 6.79–6.84 (m, 4H), 7.18–7.36 (m, 9H), 8.56 (s, 2H), 10.65 (s, 1H). Anal. calcd. for C35 H35 N5 O8 ·1/2 H2 O: C, 63.44; H, 5.48; N, 10.57; found: C, 63.31; H, 5.30; N, 10.30.
Prepare N6 -AOC-2 -O-TBDMS-5 -O-DMTr-adenosine 18. Add 4.4 g (64.6 mmol) imidazole to a solution of 19.8 g (30.3 mmol) S.15 in DMF (35 mL). 19. Add 4.85 g (32.2 mmol) TBDMS-Cl and stir overnight. 20. Dilute the reaction mixture with 400 mL ethyl acetate and wash three times with 150 mL water. 21. Dry the organic layer over sodium sulfate and remove the organic solvent using a rotary evaporator. 22. Subject the crude material to column chromatography using a 500 g silica gel column. Monitor fractions by TLC using 3:2 (v/v) ethyl acetate/hexane. 23. Elute first with 2:8 (v/v) ethyl acetate/hexane to remove all undesired 2 ,3 -di-Osilylated product. In this preparation, 3.2 g (12% yield) of the 2 ,3 -di-O-silylated product are obtained (Rf = 0.45; 1:3 ethyl acetate/hexane). Current Protocols in Nucleic Acid Chemistry
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24. Elute next with 35:65 (v/v) ethyl acetate/hexane and collect fractions containing first the desired N6 -AOC-2 -O-TBDMS-5 -O-DMTr-adenosine (S.16; Rf = 0.64) and then its 3 -O-TBDMS isomer (Rf = 0.40). 25. Finally, elute with ethyl acetate to collect the starting material (S.15). 26. Dissolve the 3 -isomer in 1.1 L of 1:5:5 (v/v/v) triethylamine/methanol/ethyl acetate and leave at room temperature for 12 hr to isomerize to the 2 -isomer. 27. Concentrate the resulting solution. Subject the residue to column chromatography using 50 g silica gel and elute the 2 - and 3 -isomers as described above. 28. Concentrate fractions including the desired product S.16 with a rotary evaporator. N6 -Allyloxycarbonyl-2 -O-(tert-butyldimethylsilyl)-5 -O-(4,4 -dimethoxytrityl)adenosine (S.16): 7.68 g (33%) as an amorphous solid. m.p. = 95◦ C. TLC (3:2 ethyl acetate/hexane): Rf = 0.64. UV λmax : 236 nm (ε 24,800), 268 (20,800). 1 H NMR (270 MHz, DMSO-d6 ): –0.15 (s, 3H), –0.05 (s, 3H), 0.73 (s, 9H), 3.27–3.68 (m, 2H), 3.71 (s, 6H), 4.11–4.28 (m, 2H), 4.63–4.65 (m, 2H), 4.86 (t, 1H, J = 4.8 Hz), 5.20 (s, 1H), 5.21 (dd, 1H, J = 1.7 and 10.55 Hz), 5.36 (dd, 1H, J = 1.7 and 17.3 Hz), 5.89–6.04 (m, 2H), 7.19–7.47 (m, 4H), 7.80–7.84 (m, 9H), 8.54 (s, 1H), 8.62 (s, 1H), 10.65 (s, 1H). Anal. calcd. for C41 H49 N5 O8 Si: C, 64.13; H, 6.43; N, 9.12; found: C, 64.12; H, 6.40; N, 8.72. After carrying out the isomerization procedure one time, 10.5 g (45% yield) of the 3 -Osilylated byproduct (Rf = 0.40; 3:2 ethyl acetate/hexane) remain. BASIC PROTOCOL 6
SYNTHESIS OF N4 -ALLYLOXYCARBONYL-2 -O(tert-BUTYLDIMETHYLSILYL)-5 -O-(4,4 -DIMETHOXYTRITYL)CYTIDINE This compound is prepared via (1) N4 -allyloxycarbonylation of cytidine by successive treatment with hexamethyldisilazane in the presence of ammonium sulfate as a catalyst, with allyloxycarbonyl chloride in the presence of N-methylimidazole, and with triethylamine in methanol to produce N4 -AOC-cytidine, (2) 5 -O-dimethoxytritylation by exposure to 4,4 -dimethoxytrityl chloride in pyridine to give N4 -AOC-5 -O-DMTr-cytidine, (3) tert-butyldimethylsilylation using tert-butyldimethylsilyl chloride in the presence of imidazole to afford a mixture of N4 -AOC-2 -O-TBDMS-5 -O-DMTr-cytidine and its 3 O-TBDMS isomer, and (4) chromatographic separation of this mixture on a silica gel column to furnish N4 -AOC-2 -O-TBDMS-5 -O-DMTr-cytidine. Following separation, the 3 -O-silylated byproduct is isomerized to increase the yield of the desired 2 -isomer. The overall reaction sequence is illustrated in Figure 2.12.6. All operations are carried out at room temperature unless otherwise noted.
Materials
Nucleobase Protection with Allyloxycarbonyl
Hexamethyldisilazane (HMDS), 99% (Aldrich, Shinetsu) Cytidine (S.17), 99% (Sigma) Ammonium sulfate, 99% (Aldrich, Wako) 1,4-Dioxane, distilled from sodium benzophenone ketyl Toluene, distilled from sodium benzophenone ketyl Dichloromethane, 99% (Aldrich, Nacarai Tesque) N-Methylimidazole, (Aldrich, Nacarai Tesque) Allyloxycarbonyl chloride (AOC-Cl), 99% (Aldrich, TCI) Methanol, 99% (Aldrich, Kishida) Triethylamine, distilled from CaH2 Ethyl acetate, 99% (Aldrich, Kishida) Diphosphorus pentoxide (P2 O5 ), 98% (Aldrich, Nacarai Tesque) Pyridine, distilled from CaH2 N,N -Dimethylformamide (DMF), distilled from 4A molecular sieves
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Figure 2.12.6
Preparation of N 4 -AOC-2 -O-TBDMS-5 -O-DMTr-cytidine.
4,4 -Dimethoxytrityl chloride (DMTr-Cl), 97% (Aldrich, TCI) Silica gel 60 (spherical, neutrality, particle size 75 µm; Merck, Nacarai Tesque) Imidazole, 99% (Aldrich, Nacarai Tesque) tert-Butyldimethylsilyl chloride (TBDMS-Cl), 99% (Aldrich, Shinetsu) Sodium sulfate, 99% (Aldrich, Nacarai Tesque) Hexane, 99% (Aldrich, Kishida) Rotary evaporator equipped with a diaphragm pump and cooling unit. Vacuum oil pump TLC plate: silica-coated glass plate with fluorescent indicator (Merck silica gel 60 F254 ) 254-nm UV lamp Glass column Diaphragm pump Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) Prepare N4 -AOC-cytidine 1. Add 150 mL HMDS to a mixture of 15.0 g (61.7 mmol) cytidine (S.17) and a catalytic amount of ammonium sulfate in 150 mL dioxane. Reflux the mixture for 3 hr (b.p. 125◦ C, oil bath 145◦ C). 2. Evaporate the solvent on a rotary evaporator with a diaphragm pump and a cooling unit, and then dry the residue by co-evaporating twice with 100 mL dry toluene. 3. Dissolve the residue in 500 mL dichloromethane and add 6.70 mL (6.90 g, 84.0 mmol) N-methylimidazole followed by 8.90 mL (10.1 g, 83.9 mmol) AOCCl. Stir the mixture for 12 hr. 4. Evaporate the reaction mixture and dissolve the residue in a mixture of 500 mL methanol and 130 mL triethylamine. Stir the solution at room temperature for 12 hr.
Protection of Nucleosides for Oligonucleotide Synthesis
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5. Evaporate the reaction mixture with a rotary evaporator and collect the resulting solid residue. 6. Recrystallize the solid residue from a mixture of 30 mL methanol and 250 mL ethyl acetate. 7. Collect the crystals by filtering through a glass column with a cotton stopper under vacuum from a diaphragm pump. Wash with ethyl acetate. 8. Dry the crystals over P2 O5 at 50◦ C under vacuum with a vacuum oil pump. N4 -(Allyloxycarbonyl)cytidine (S.18): 20.0 g (99%) as colorless powdery crystals. m.p. 103◦ –107◦ C. TLC (1:9 methanol/dichloromethane): Rf = 0.3. 1 H NMR (400 MHz, DMSOd6 ): 3.57–3.97 (m, 5H), 4.62–4.64 (m, 2H), 5.03 (d, 1H, J = 5.4 Hz), 5.15 (t, 1H, J = 4.9 Hz), 5.22–5.38 (m, 2H), 5.47 (d, 1H, J = 4.4 Hz), 5.77 (d, 1H, J = 2.4 Hz), 5.91–5.98 (m, 1H), 7.00 (d, 1H, J = 7.3 Hz), 8.40 (d, 1H, J = 7.3 Hz), 10.80 (s, 1H). Anal. calcd. for C13 H17 N3 O7 ·1/2 H2 O: C, 46.43; H, 5.39; N, 12.49; found: C, 46.03; H, 5.16; N, 12.49.
Prepare N4 -AOC-5 -O-DMTr-cytidine 9. Prepare a solution of 9.52 g (33.4 mmol) S.18 in 150 mL of 1:1 (v/v) pyridine/DMF and cool to 0◦ C. 10. Add 15.6 g (46.0 mmol) DMTr-Cl in small portions over 20 min. Stir the mixture at the same temperature for 20 min and then at room temperature for 12 hr. 11. Pour the reaction mixture into 1.2 L of ice-cold water. 12. Collect the resulting precipitates by vacuum filtration as above, and wash with 300 mL water. 13. Dry the precipitates at 50◦ C under vacuum with a vacuum oil pump. 14. Dissolve the dried solid material in 50 mL dichloromethane and apply to a 500 g silica gel column (APPENDIX 3E). Monitor fractions by TLC (APPENDIX 3D) with 1:9 methanol/dichloromethane. Visualize under UV light (254 nm). Elute the column first with 1.5 L of 1:20 (v/v) methanol/dichloromethane to remove byproducts, and then with 1:9 (v/v) methanol/dichloromethane to collect the product (Rf = 0.64). 15. Combine fractions containing the desired product and concentrate using a rotary evaporator. 16. Crystallize the product from 200 mL methanol and collect the resulting crystals by vacuum filtration as above. N4 -Allyloxycarbonyl-5 -O-(4,4 -dimethoxytrityl)cytidine (S.19): 15.7 g (26.7 mmol, 80%) as colorless powdery crystals. m.p. 118◦ –120◦ C. TLC (1:9 methanol/dichloromethane): Rf = 0.64. 1 H NMR (400 MHz, CDCl3 ): 2.01 (br 1H), 3.38–3.48 (m, 2H), 3.79 (s, 6H), 4.38 (s, 3H), 4.67 (d, 2H, J = 5.4 Hz), 5.27–5.38 (m, 2H), 5.85–5.98 (m, 3H), 6.81–6.84 (m, 4H), 7.04 (d, 1H, J = 7.3 Hz), 7.20–7.36 (m, 9H), 8.23 (d, 1H, J = 7.3 Hz).
Prepare N4 -AOC-2 -O-TBDMS-5 -O-DMTr-cytidine 17. Prepare a solution of 15.4 g (26.1 mmol) S.19 in 25 mL DMF. Add 3.9 g (57.5 mmol) imidazole and begin stirring the mixture. 18. Add 4.33 g (28.8 mmol) TBDMS-Cl and continue stirring overnight. 19. Dilute the reaction mixture with 400 mL ethyl acetate and wash three times with 150 mL water. Nucleobase Protection with Allyloxycarbonyl
20. Dry the organic layer over ∼15 g sodium sulfate. Remove the drying agent by filtration through a glass column with a cotton stopper under vacuum from a diaphragm pump. Evaporate with a rotary evaporator.
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21. Subject the residue to column chromatography on a 500 g silica gel column. Monitor fractions by TLC using 3:7 (v/v) ethyl acetate/hexane. 22. Elute first with 1:30:30 (v/v/v) methanol/ethyl acetate/hexane to collect the undesired 2 ,3 -di-O-silylated product (Rf = 0.70). In this preparation, 6.13 g (29% yield) of the 2 ,3 -di-O-silylated product are obtained.
23. Elute next with 1:20:20 (v/v/v) methanol/ethyl acetate/hexane and collect fractions containing first the desired N6 -AOC-2 -O-TBDMS-5 -O-DMTr-cytidine (S.20; Rf = 0.52) and then its 3 -O-TBDMS isomer (Rf = 0.37). 24. Finally, elute with ethyl acetate to collect the starting material (S.19). 25. Dissolve the 3 -isomer in 1.1 L of 1:5:5 (v/v/v) triethylamine/methanol/ethyl acetate and leave at room temperature for 12 hr to isomerize to the 2 -isomer. 26. Concentrate the resulting solution. Subject the residue to column chromatography using 50 g silica gel and elute the 2 - and 3 -isomers as described above. 27. Evaporate fractions containing S.20 with a rotary evaporator. N4 -Allyloxycarbonyl-2 -O-(tert-butyldimethylsilyl)-5 -O-(4,4 -dimethoxytrityl)cytidine (S.20): 4.05 g (22%) as an amorphous solid. m.p. = 110◦ C. TLC (3:7 ethyl acetate/ hexane): Rf = 0.52. 1 H NMR (400 MHz, CDCl3 ): 0.19 (s, 3H), 0.32 (s, 3H), 0.93 (s, 9H), 1.62 (br, 1H), 2.35 (d, J = 9.8 Hz, 1H), 3.51–3.60 (m, 2H), 3.81 (s, 6H), 4.08 (d, J = 7.8 Hz, 1H), 4.28–4.35 (m, 2H), 4.66 (d, J = 5.8 Hz, 2H), 5.28–5.38 (m, 2H), 5.89–5.97 (m, 2H), 6.85 (d, J = 8.8 Hz, 4H), 7.25–7.42 (m, 10H), 8.45 (d, J = 7.8 Hz, 1H). After carrying out the isomerization procedure one time, 2.58 g (14% yield) of the 3 -Osilylated byproduct (Rf = 0.37; 3:7 ethyl acetate/hexane) remain.
SYNTHESIS OF N2 -ALLYLOXYCARBONYL-O6 -ALLYL-2 -O-(tertBUTYLDIMETHYLSILYL)-5 -O-(4,4 -DIMETHOXYTRITYL)GUANOSINE
BASIC PROTOCOL 7
This compound is synthesized by (1) tri-O-acetylation of guanosine using acetic anhydride in pyridine to give 2 ,3 ,5 -tri-O-(acetyl)guanosine, (2) O6 -allylation by treatment with allyl alcohol in the presence of triphenylphosphine, dioxane, and diethyl azodicarboxylate to afford O6 -allyl-2 ,3 ,5 -tri-O-(acetyl)guanosine, (3) N-allyloxycarbonylation by treatment with allyloxycarbonyl chloride in the presence of tert-butylmagnesium chloride and tetrahydrofuran to produce N2 -AOC-O6 -allyl-2 ,3 ,5 -tri-O-(acetyl)guanosine, (4) removal of the acetyl protecting groups using sodium hydroxide in ethanol to give N2 -AOC-O6 -(allyl)guanosine, (5) 5 -O-dimethoxytritylation by treatment with 4,4 dimethoxytrityl chloride in pyridine and N,N-dimethylformamide to provide N2 -AOCO6 -allyl-5 -O-DMTr-guanosine, and (6) O-silylation with tert-butyldimethylsilyl in the presence of imidazole and N,N-dimethylformamide. Following silylation, the desired 2 -O-silylated product, N2 -AOC-O6 -allyl-2 -O-TBDMS-5 -O-DMTr-guanosine, is separated from the 3 -O-silylated and 2 ,3 -di-O-silylated byproducts by chromatography, and the 3 -O-silylated product is isomerized to increase the yield of the desired 2 -O-silylated product. The overall reaction sequence is illustrated in Figure 2.12.7. All operations are carried out at room temperature unless otherwise noted.
Materials Guanosine (S.21), 99% (Aldrich) Pyridine, distilled from CaH2 N,N -Dimethylformamide (DMF), distilled from 4A molecular sieves Acetic anhydride, 99% (Aldrich, Kishida) 2-Propanol, 99% (Aldrich, Kishida) Triphenylphosphine, recrystallized from hexane
Protection of Nucleosides for Oligonucleotide Synthesis
2.12.17 Current Protocols in Nucleic Acid Chemistry
Supplement 23
Figure 2.12.7
Preparation of N 2 -AOC-O 6 -allyl-2 -O-TBDMS-5 -O-DMTr-guanosine.
Allyl alcohol, distilled from Mg 1,4-Dioxane, distilled from sodium benzophenone Diethyl azodicarboxylate (DEAD), 40% solution in toluene (Aldrich, TCI) Dichloromethane, 99% (Aldrich, Nacarai Tesque) Silica gel 60 (spherical, neutrality, particle size 75 µm; Merck, Nacarai Tesque) Ethyl acetate, 99% (Aldrich, Kishida) Hexane, 99% (Aldrich, Kishida) Tetrahydrofuran (THF), distilled from sodium benzophenone ketyl Allyloxycarbonyl chloride (AOC-Cl), 99% (Aldrich, TCI) tert-Butylmagnesium chloride, 99% (Aldrich) Methanol, 99% (Aldrich, Kishida) Saturated aqueous ammonium chloride (99%; Aldrich, Kishida) Aqueous saturated sodium hydrogencarbonate (NaHCO3 ; 99%; Aldrich, Nacarai Tesque) Brine: aqueous saturated sodium chloride Ethanol, 99.5% (Aldrich, Nacarai Tesque) Sodium hydroxide 80% (v/v) aqueous acetic acid 4,4 -Dimethoxytrityl chloride (DMTr-Cl), 97% (Aldrich, TCI) Sodium sulfate, 99% (Aldrich, Nacarai Tesque) Imidazole, 99% (Aldrich, Nacarai Tesque) tert-Butyldimethylsilyl chloride (TBDMS-Cl), 99% (Aldrich, Shinetsu) Triethylamine, distilled from CaH2
Nucleobase Protection with Allyloxycarbonyl
Vacuum oil pump Rotary evaporator equipped with a diaphragm pump and cooling unit TLC plate: silica-coated glass plate with fluorescent indicator (Merck silica gel 60 F254 ) 254-nm UV lamp Glass column Diaphragm pump Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D)
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Prepare 2 ,3 ,5 -tri-O-(acetyl)guanosine 1. Prepare a suspension of 15 g (52.9 mmol) guanosine (S.21) in a mixture of 15 mL pyridine and 37.5 mL DMF. 2. Add 30 mL (317 mmol) acetic anhydride, heat to 70◦ C, and stir for 3 hr at this temperature. 3. Cool the mixture to room temperature. 4. Add 100 mL 2-propanol and stir for 30 min. 5. Collect the resulting precipitates by filtering through a glass column with a cotton stopper under vacuum from a diaphragm pump. Wash with 300 mL 2-propanol. 6. Dry the precipitates under reduced pressure with a vacuum oil pump. 2 ,3 ,5 -Tri-O-(acetyl)guanosine (S.22): 17.4 g (79%) as a white solid. m.p. 230◦ –234◦ C. 1 H NMR (400 MHz, DMSO-d6 ): 2.04 (2 s, 6H), 2.11 (s, 3H), 4.23–4.39 (m, 3H), 5.48–5.51 (m, 1H), 5.79 (t, J = 5.9 Hz, 1H), 5.98 (d, J = 6.1 Hz, 1H), 7.92 (s, 1H), 10.72 (br, 1H).
Prepare N2 -AOC-O6 -(allyl)guanosine 7. Add 5.40 g (20.6 mmol) triphenylphosphine and 7.50 mL (6.41 g, 110 mmol) allyl alcohol to a suspension of 4.90 g (12.0 mmol) S.22 in 90 mL dry dioxane. Heat the mixture at 80◦ C for 45 min. 8. Cool the reaction mixture to room temperature. Add 3.50 mL (3.71 g, 21.3 mmol) diethyl azodicarboxylate and then heat at 60◦ C for 3 hr. 9. Evaporate the reaction mixture and dissolve the residue in 15 mL dichloromethane. Keep the solution in a refrigerator (at 4◦ C) for 2 to 3 hr. If no precipitate forms during this time, cool for another 2 to 3 hr.
10. Remove the resulting precipitates by vacuum filtration as above, and wash with 50 mL cold dichloromethane. 11. Combine the filtrates and concentrate on a rotary evaporator with a diaphragm pump and a cooling unit. 12. Subject the residue to column chromatography (APPENDIX 3E) using 250 g silica gel. Monitor fractions by TLC (APPENDIX 3D) using 3:1 (v/v) ethyl acetate/hexane. Visualize under UV light (254 nm). Elute the column first with 1.5 L of 3:7 (v/v) ethyl acetate/hexane to remove undesired byproducts, and then with 3:2 (v/v) ethyl acetate/hexane obtain the target compound (Rf = 0.48). 13. Evaporate the fractions containing the desired product with a rotary evaporator. O6 -Allyl-2 ,3 ,5 -tri-O-(acetyl)guanosine: 4.30 g (80%) as a colorless amorphous solid. TLC (3:1 ethyl acetate/hexane): Rf = 0.48. 1 H NMR (400 MHz, DMSO-d6 ): 2.03 (s, 6H), 2.12 (s, 3H), 4.25–4.42 (m, 3H), 4.95 (d, 2H, J = 5.9 Hz), 5.26 (d, 1H, J = 10.3 Hz), 5.40 (dd, 1H, J = 1.5 and 19.0 Hz), 5.54–5.56 (m, 1H), 5.86–5.89 (m, 1H), 6.07–6.12 (m, 2H), 6.55 (s, 2H), 8.11 (s, 1H).
14. Dissolve the residue in 60 mL THF and add 3.00 mL (28.3 mmol) AOC-Cl. 15. Cool the solution to 5◦ C and add dropwise (over 10 min) 18.0 mL (25.2 mmol) of 1.4 M tert-butylmagnesium chloride in THF. Stir at the same temperature for an additional 60 min. 16. Quench the reaction by adding 3 mL methanol. 17. Dilute the mixture with 100 mL ethyl acetate and wash with 100 mL aqueous saturated ammonium chloride.
Protection of Nucleosides for Oligonucleotide Synthesis
2.12.19 Current Protocols in Nucleic Acid Chemistry
Supplement 23
18. Wash the organic layer with 100 mL aqueous saturated NaHCO3 followed by 100 mL brine. 19. Evaporate the organic solution with a rotary evaporator. 20. Purify residue by column chromatography using 250 g silica gel with 2:3 (v/v) ethyl acetate/hexane as eluent. Monitor by TLC using 1:1 ethyl acetate/hexane (Rf = 0.53). 21. Concentrate fractions containing the desired compound with a rotary evaporator. N2 -Allyloxycarbonyl-O6 -allyl-2 ,3 ,5 -tri-O-(acetyl)guanosine: 3.90 g (66%) as an oily product. TLC (1:1 ethyl acetate/hexane): Rf = 0.53. 1 H NMR (400 MHz, DMSO-d6 ): 1.96 (s, 3H), 2.02 (s, 3H), 2.10 (s, 3H), 4.40–4.18 (m, 3H), 4.68–4.67 (m, 4H), 5.04 (d, 2H, J = 5.4 Hz), 5.60–5.15 (m, 6H), 5.74–5.73 (m, 1H), 5.91–5.81 (m, 3H), 6.12–6.06 (m, 1H), 6.27 (d, 1H, J = 3.4 Hz), 8.69 (s, 1H). FAB-MS: m/z 618 (M+ + H).
22. Dissolve residue in 35 mL ethanol and add 200 mL of 0.1 M sodium hydroxide in 95% ethanol. Stir the resulting mixture for 30 min. 23. Add the reaction mixture to 30 mL of 80% aqueous acetic acid and evaporate with a rotary evaporator. 24. Purify residue by chromatography on a short 50 g silica gel column with 1:5:5 (v/v/v) methanol/ethyl acetate/hexane as eluent. Monitor fractions by TLC using 1:10 (v/v) methanol/dichloromethane (Rf = 0.35). 25. Concentrate fractions containing the target product with a rotary evaporator. N2 -Allyloxycarbonyl-O6 -(allyl)guanosine (S.23): 2.55 g (52% overall yield) as an amorphous solid. TLC (1:10 methanol/dichloromethane): Rf = 0.35. 1 H NMR (400 MHz, DMSO-d6 ): 3.52–3.54 (m, 1H), 3.62–3.65 (m, 1H), 3.90–3.91 (m, 1H), 4.17 (m, 1H), 4.61–4.62 (m, 3H), 4.93 (br s, 1H), 5.04–5.05 (m, 2H), 5.16–5.48 (m, 6H), 5.88 (d, 1H, J = 5.9 Hz), 5.91–6.00 (m, 1H), 6.12–6.19 (m, 1H), 8.43 (s, 1H), 10.38 (s, 1H). FAB-MS (C17 H21 N5 O7 ): m/z 408 (M+ + H).
Prepare N2 -AOC-O6 -allyl-5 -O-DMTr-guanosine 26. Add 1.54 g (4.55 mmol) DMTr-Cl to a solution of 1.73 g (4.25 mmol) S.23 in 20 mL of 1:1 (v/v) pyridine/DMF. Heat at 55◦ C for 1.5 hr. 27. Pour into 100 mL of saturated aqueous NaHCO3 and extract three times with 30 mL ethyl acetate. 28. Dry the combined organic extracts over ∼15 g sodium sulfate. Remove the drying agent by filtration through a glass column with a cotton stopper under vacuum from a diaphragm pump. Evaporate with a rotary evaporator. 29. Subject residue to column chromatography using 100 g silica gel. Monitor elution by TLC using 3:1 (v/v) ethyl acetate/hexane. Elute first with 1.5 L of 3:7 (v/v) ethyl acetate/hexane to remove byproducts, and then with 3:2 (v/v) ethyl acetate/hexane to collect fractions containing the target product (Rf = 0.61). 30. Concentrate the fractions with a rotary evaporator.
Nucleobase Protection with Allyloxycarbonyl
N2 -Allyloxycarbonyl-O6 -allyl-5 -O-(4,4 -dimethoxytrityl)guanosine (S.24): 2.31 g (77%) as colorless foam. TLC (3:1 ethyl acetate/hexane): Rf = 0.61. UV λmax : 237 nm (ε 26,800), 269 (17,400). 1 H NMR (400 MHz, DMSO-d6 ): 3.16–3.56 (m, 2H), 3.69 (s, 3H), 3.70 (s, 3H), 4.02–4.03 (m, 1H), 4.33–4.37 (m, 1H), 4.59–4.60 (m, 2H), 4.69–4.72 (m, 1H), 5.04 (d, 2H, J = 4.9 Hz), 5.10 (d, 1H, J = 5.9 Hz), 5.22 (d, 1H, J = 10.7 Hz), 5.29 (d, 1H, J = 10.7 Hz), 5.38 (d, 1H, J = 17.6 Hz), 5.47 (d, 1H, J = 17.1 Hz), 5.57 (d, 1H, J = 5.9 Hz), 5.92–5.99 (m, 2H), 6.13–6.20 (m, 1H), 6.72–6.78 (m, 4H), 7.16 (m, 9H), 8.33 (s, 1H), 10.34 (s, 1H). Anal. calcd. for C38 H39 N5 O9 : C, 64.31; H, 5.54; N, 9.87; found: C, 64.32; H, 5.46; N, 9.69.
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Prepare N2 -AOC-O6 -allyl-2 -O-TBDMS-5 -O-DMTr-guanosine 31. Add 0.29 g (4.3 mmol) imidazole to a solution of 1.38 g (1.94 mmol) S.24 in 2 mL DMF. 32. Add 0.32 g (2.1 mmol) TBDMS-Cl and stir at 60◦ C for 1 hr. 33. Pour the reaction mixture into 50 mL ice-cold water and extract three times with 20 mL ethyl acetate. 34. Combine the organic extracts, dry over sodium sulfate, and evaporate with a rotary evaporator. 35. Purify residue by column chromatography over 75 g silica gel. Monitor fractions by TLC using 3:7 (v/v) ethyl acetate/hexane. 36. Elute first with 2:8 (v/v) ethyl acetate/hexane to separate the 2 ,3 -di-O-silylated product (Rf = 0.67). In this preparation, 613 mg (38% yield) of the 2 ,3 -di-O-silylated product are obtained.
37. Elute next with 1:3 (v/v) ethyl acetate/hexane and collect fractions containing first the desired N2 -AOC-O6 -allyl-2 -O-TBDMS-5 -O-DMTr-guanosine (S.25; Rf = 0.34) and then its 3 -O-TBDMS isomer (Rf = 0.22). 38. Finally, elute with ethyl acetate to collect the starting material (S.24). 39. Dissolve the 3 -isomer in 1.1 L of 1:5:5 (v/v/v) triethylamine/methanol/ethyl acetate and leave at room temperature for 12 hr to isomerize to the 2 -isomer. 40. Concentrate the resulting solution. Subject the residue to column chromatography using 50 g silica gel and elute the 2 - and 3 -isomers as described above. 41. Combine all fractions containing the desired product (S.25) and concentrate with a rotary evaporator. N2 -Allyloxycarbonyl-O6 -allyl-2 -O-(tert-butyldimethylsilyl)-5 -O-(4,4 -dimethoxytrityl) guanosine (S.25): 457 mg (29% yield) as an amorphous solid. m.p. = 81◦ C. TLC (3:7 ethyl acetate/hexane): Rf = 0.34. UV λmax : 237 nm (ε 24,300), 269 (15,900). 1 H NMR (400 MHz, DMSO-d6 ): –0.16 (s, 3H), –0.05 (s, 3H), 0.74 (s, 9H), 3.21–3.24 (m, 1H), 3.35–3.56 (m, 1H), 3.70 (s, 3H), 3.71 (s, 3H), 3.74–4.05 (m, 1H), 4.24–4.28 (m, 1H), 4.57–4.58 (m, 2H), 4.88–4.89 (m, 1H), 4.96–4.98 (m, 1H), 5.04 (d, 2H, J = 5.9 Hz), 5.21 (dd, 1H, J = 1.71 and 10.5 Hz), 5.30 (d, 1H, J = 10.2 Hz), 5.37 (dd, 1H, J = 1.5 and 17.1 Hz), 5.45 (dd, 1H, J = 1.5 and 17.1 Hz), 5.92–5.99 (m, 2H), 6.12–6.19 (m, 1H), 6.75–6.80 (m, 4H), 7.15–7.34 (m, 9H), 8.33 (s, 1H), 10.25 (s, 1H). Anal. calcd. for C44 H53 N5 O9 Si: C, 64.14; H, 6.48; N, 8.50; found: C, 64.13; H, 6.43; N, 8.35. After carrying out the isomerization procedure one time, 213 mg (12% yield) of the 3 -O-silylated byproduct (Rf = 0.22; 3:7 ethyl acetate/hexane) remain.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
1-(Allyloxycarbonyl)tetrazole (AOC-Tet) Add 12.0 mL (0.113 mol) allyloxycarbonyl chloride (AOC-C1; 99%; Aldrich, TCI) dropwise to a solution of 7.0 g (0.100 mol) 1H-tetrazole (99%; Dozindo) and 15.0 mL (0.108 mol) triethylamine in 150 mL tetrahydrofuran (THF) at 0◦ C over 15 min. Stir the resulting mixture for 30 min. Remove the resulting precipitates by vacuum filtration through a pad of Celite 545 (Aldrich, Nacarai Tesque) and wash with 150 mL THF. Concentrate the resulting THF solution containing AOC-Tet to ∼30 mL. Prepare immediately before use. AOC-Tet is used for N-allyloxycarbonylation of 2 -deoxyadenosine. Current Protocols in Nucleic Acid Chemistry
Protection of Nucleosides for Oligonucleotide Synthesis
2.12.21 Supplement 23
Allyl 1-benzotriazolyl carbonate (AOCOBT) To a solution of 13.5 g (0.10 mol) 1-hydroxybenzotriazole and 16.7 mL (0.12 mol) triethylamine in 350 mL tetrahydrofuran, add 11.8 mL (0.11 mol) allyloxycarbonyl chloride (AOC-C1; 99%; Aldrich, TCI) dropwise with vigorous stirring at 0◦ C. Continue stirring for an additional 10 min. Remove the resulting precipitates by vacuum filtration through a pad of Celite 545 (Aldrich, Nacarai Tesque). Wash the pad three times with 30 mL ethyl acetate. Combine the filtrate and washes, and evaporate the solvent using a rotary evaporator with a diaphragm pump and a cooling unit. Recrystallize the resulting crystalline residue from 100 mL ethyl acetate. Prepare immediately before use. AOCOBT is used for N-allyloxycarbonylation of 2 -deoxycytidine. Yield: (19.9 g, 91%) as a white solid. m.p. 107◦ –111◦ C. IR (KBr) 1760, 1430, 1390, 1260, 770 cm−1 . UV λmax : 236 nm (ε 13,900), 321 (11,800), 334 (10,000). 1 H NMR: 5.07 (dt, 2H, J = 5.4 and 1.8 Hz, =CH2 ), 5.4–5.7 (m, 2H, CH2 CH= =CH2 ), 6.14 (ddt, 1H, J = 10.2, 17.1, and 5.4 Hz, CH2 CH= CH2 CH= =CH2 ), 7.5–8.3 (m, 4H, arom. protons).
COMMENTARY Background Information
Nucleobase Protection with Allyloxycarbonyl
Currently, the synthesis of oligodeoxyribonucleotides and oligoribonucleotides is most generally achieved via the phosphoramidite method, in which acyl and aroyl protecting groups (including benzoyl, panisoyl, and isobutyryl groups) are normally used for protection of nucleobases (Letsinger et al., 1975; Beaucage and Caruthers, 1981; Matteucci and Caruthers, 1981; McBride and Caruthers, 1983; Sinha et al., 1983; Dorman et al., 1984). However, the acyl and aroyl N-protecting groups are not useful for the synthesis of compounds with base-labile functions, because these protecting groups require heating, usually at 55◦ C, for several hours with concentrated aqueous ammonia. In order to improve on the acyl/aroyl protection of nucleobases, alternative protecting groups have been developed; the representative ones are phenoxyacetyl (Schulhof et al., 1987), isopropoxyacetyl (Uznanski et al., 1989; Vinogradov et al., 1993), tertbutylphenoxyacetyl (K¨oster et al., 1981), 2-(acetoxymethyl)benzoyl (Kuijpers et al., 1990), 2-(tert-butyldiphenylsilyloxymethyl) benzoyl (Dreef-Tromp et al., 1990), formamidine (Vu et al., 1990), 9-fluorenylmethoxycarbonyl (Koole et al., 1989), and 2-(pnitrophenyl)ethoxycarbonyl (Himmelsbach et al., 1984). Removal of these protecting groups can be achieved under rather mild conditions, but still requires basic reagents, thus involving a risk of undesirable decomposition in the synthesis of base-sensitive derivatives. In addition, some of these protecting groups are themselves labile to bases, and are
thus at risk from basic treatment during synthesis. There is thus a strong demand for a nucleobase-protecting group that can be removed under non-basic and mild conditions but is stable under conditions employed for the oxidation of the phosphite intermediate and the removal of the 5 -O-DMTr group using dichloroacetic acid. The allyloxycarbonyl (AOC) group is one example of an N-protecting group that meets these requirements. The AOC group is quite stable to acids, including dichloroacetic acid and trichloroacetic acid; to bases, including concentrated ammonia; and to oxidizing agents such as an I2 /H2 O/pyridine mixture, tert-butyl hydroperoxide (Hayakawa et al., 1986a), bis(trimethylsilyl) peroxide in the presence of trimethylsilyl triflate (Hayakawa et al., 1986a), and 2-butanone peroxide (Kataoka et al., 2001). These are reagents that are normally used for the synthesis of nucleotides via the phosphoramidite method. By contrast, the AOC group is very sensitive to organic Pd(0) complexes such as =CH)2 CO]3 · Pd[P(C6 H5 )3 ]4 or Pd2 [(C6 H5 CH= CHCl3 in the presence of a nucleophile, leading to clean and smooth removal of the protecting group. As the nucleophile, butylammonium formate, diethylammonium hydrogencarbonate, diethylammonium carbonate, or dimedone can be used. The deprotection is performed under nearly neutral conditions (Hayakawa et al., 1986b). The AOC group is stable under conditions normally employed for conversion of the protected nucleoside to the 3 -phosphoramidite using an (alkoxy)bis(diisopropylamino)phosphine
2.12.22 Supplement 23
Current Protocols in Nucleic Acid Chemistry
in the presence of isopropylammonium tetrazolide (Hayakawa et al., 1990; Heidenhain and Hayakawa, 1999).
Compound Characterization Chemical characterization data are provided for all compounds. NMR spectra were recorded on a JEOL JNM-α400 or ECA-500 instrument. The 1 H, 13 C, and 31 P NMR chemical shifts are described as δ values in ppm relative to (CH3 )4 Si (for 1 H and 13 C) and 85% H3 PO4 (for 31 P). Infrared (IR) spectra were measured with a JASCO FT/IR-460 spectrometer. Ultraviolet (UV) spectra were obtained in methanol, unless otherwise noted, on a JASCO V-550 spectrometer.
Critical Parameters and Troubleshooting
For 2 -deoxycytidine, adenosine, and cytidine, allyloxycarbonylation of the parent nucleoside takes place at the amino group of the nucleobase in a chemoselective manner to give the desired product in a satisfactory yield. Therefore, preparation of N4 -AOC-5 -O-DMTr-2 -deoxycytidine, N6 AOC-5 -O-DMTr-adenosine, and N4 -AOC5 -O-DMTr-cytidine can be carried out by N-allyloxycarbonylation followed by dimethoxytritylation, which is the shortest and most convenient route. In contrast, allyloxycarbonylation of 2 -deoxyadenosine takes place competitively at the amino group of the nucleobase and at the 5 - and 3 hydroxy groups, so that the desired Nallyloxycarbonylated product is not obtained in a high yield. Dimethoxytritylation of deoxyadenosine, however, is achieved in a highly 5 -O-selective manner. For this reason, although it is more tedious, N6 -AOC-5 -ODMTr-2 -deoxyadenosine should be prepared by 5 -O-dimethoxytritylation, 3 -O-silylation, N-allyloxycarbonylation, and finally removal of the 3 -O-TBDMS protecting group. In Basic Protocol 1, dimethoxytritylation of 2 -deoxyadenosine takes place regioselectively at the 5 -hydroxyl under the conditions described in the text, but the reaction does not go to completion, leaving a small amount of unmodified 2 -deoxyadenosine. Nevertheless, the amount of 4,4 -dimethoxytrityl chloride should never be increased. When extra dimethoxytrityl chloride is added, 3 O-dimethoxytritylation leads to formation of 3 ,5 -di-O-DMTr-2 -deoxyadenosine, and separation of this compound from the target 5 O-DMTr derivative is rather tedious. The allyloxycarbonyl tetrazolide (AOC-Tet) reagent
used for the N-allyloxycarbonylation of S.2 is rather sensitive to moisture; thus, the reaction should be carried out with all possible precautions for maintaining anhydrous conditions. If the reaction is not satisfactory, dry the AOCTet reagent by co-evaporation with toluene before use. In trimethylsilylation of 2 -deoxycytidine, adenosine, and cytidine (in Basic Protocol 2, 5, and 6), the parent nucleoside is not completely dissolved in the reaction solvent, and thus the reaction mixture is heterogeneous at the initial stage. However, since the di-O-silylated 2 deoxycytidine and the tri-O-silylated adenosine and cytidine are all soluble in the reaction solvent, the mixtures become homogeneous as the reactions go to completion. For the N-allyloxycarbonylation of di-Osilylated 2 -deoxycytidine, the use of the AOCOBT reagent is crucial for obtaining the desired product in a satisfactory yield. When AOC-Tet is used, a complex mixture is formed. In most cases, the trimethylsilyl protecting groups of the product resulting from N-allyloxycarbonylation are completely eliminated during aqueous workup (see Basic Protocol 2, step 4). However, in some cases the desilylation is not completely achieved, so it is important to check by TLC or 1 H NMR whether the silyl groups have been removed completely. If desilylation was incomplete, an excess of triethylamine in methanol (25◦ C, 30 min) can be used to complete the deprotection. In the preparation of N6 -AOC-2 -OTBDMS-5 -O-DMTr-adenosine, the compound is separated by silica column chromatography from its 3 -O-TBDMS isomer, which is produced in an almost equal amount. Efficiency of the separation strongly depends on the activity of the silica gel. Because it is very difficult to use silica gel of the same activity in every experiment, reproducibility of this separation is not very high. If chromatography under conditions described in this unit do not allow fine separation, repeat the separation using a larger amount of silica gel. The same attention is required in the purification of N4 -AOC-2 -O-TBDMS-5 -O-DMTrcytidine and N2 -AOC-O6 -allyl-2 -O-TBDMS5 -O-DMTr-guanosine. The undesired 3 -Osilylated isomers of these three ribonucleoside derivatives can be partially (∼50%) converted to the desired 2 -O-silylated compounds by treatment (25◦ C, overnight) with a 1:5:5 mixture of triethylamine, methanol, and ethyl acetate (Heidenhain and Hayakawa, 1999).
Protection of Nucleosides for Oligonucleotide Synthesis
2.12.23 Current Protocols in Nucleic Acid Chemistry
Supplement 23
Nucleobase Protection with Allyloxycarbonyl
Several methods have been developed for preparation of N-allyloxycarbonylated 5 O-DMTr-guanosine and -2 -deoxyguanosine derivatives. Among them, the approaches described in this text provide the best yields of the target compounds. Because the C(2)NH2 group of guanine has low reactivity to acylation, the amino group does not undergo allyloxycarbonylation using AOC-Cl, AOC-Tet, and AOC-OBT. As a result, Nallyloxycarbonylation of guanosine and 2 deoxyguanosine derivatives requires a tedious and tricky procedure. The NH2 group is first activated by treatment with tertbutylmagnesium chloride to give the magnesium amide, which then is reacted with AOCCl to produce the desired compound. In this procedure, three equivalents of the Grignard reagent should be used, because two equivalents are consumed for neutralizing two protons of higher acidity than C(2)-NH2 , i.e., N(1)H and C(2)-NHCOOCH2 CH= =CH2 of the allyloxycarbonylated product. The Grignard reagent is extremely sensitive to moisture and oxygen (air). Therefore, this reaction should be carried out with great care to exclude moisture. Dryness of the THF used as the reaction solvent is critical. The solvent should be freshly distilled from sodium benzophenone immediately before use. In some cases (not always), it is necessary to dry the starting materials S.8 and S.22 by co-evaporation with toluene. For the derivatives presented in this unit, it is recommended that an alkyl group be introduced to the O6 position (although this is not always necessary), because it greatly increases the solubility of guanosine derivatives in organic solvents, reducing the required volume of solvents in later reaction steps. Use of the allyl group is most strongly recommended, as it can be removed at the same time as the N-allyloxycarbonyl group by the organopalladium-catalyzed reaction. In the O6 -allylation of the guanine base described in Basic Protocol 4, the O6 -sulfonylguanosine derivative produced as an intermediate should be used as soon as possible (within a few days) after preparation, because this compound is not absolutely stable, particularly to moisture. Trimethylamine, which is used for the next step (the conversion of the O6 sulfonylguanosine derivative to the O6 -allyl compound S.11), is a toxic gas (b.p. 3◦ –4◦ C) with an unpleasant odor, and should be used in a well-ventilated hood. Recently, it was reported that the use of a mixture of triethylamine (b.p. 88◦ –89◦ C) and DABCO in place of the trimethylamine and DBU described in
this text serves as a useful promoter for the reaction of the O6 -sulfonylguanosine and an alcohol to give the O6 -alkyl compound (Grotli et al., 1997). Use the former mixture might be preferred. The allylation of S.22 by means of Mitsunobu reaction takes place specifically at the O6 position. No N(1)-allylated product is formed. In this reaction, perfect separation of the desired product and triphenylphosphine oxide is considerably difficult, and the allylated product often contains a small amount of the phosphine oxide following a single purification by chromatography. In such cases, it is necessary to repeat the chromatography to obtain pure material. Alternatively, an effective method for removing the phosphine oxide is to dissolve the chromatographed product in ethyl acetate, leave it in a refrigerator at 4◦ C, and remove the resulting precipitates (triphenylphosphine oxide) by vacuum filtration. For protection of hydroxyl groups on the carbohydrate moiety, use of the TBDMS and acetyl groups are recommeded for 2 deoxyguanosine and guanosine derivatives, respectively. In the authors’ experience, use of acetyl for protection of deoxyguanosine hydroxyls or TBDMS for protection of guanosine hydroxyls leads to markedly decreased yields of the desired products. In triacetylation of guanosine in Basic Protocol 7, guanosine is not sufficiently dissolved in a mixture of DMF and pyridine, and thus the reaction mixture is heterogeneous at the starting point. As the reaction proceeds, the mixture becomes nearly homogeneous, perhaps because the monoacetylated product is soluble, although this has not been determined. Finally, the mixture becomes heterogeneous again because the target is not completely soluble. In the authors’ experience, no N2 acetylated compound was produced in this reaction. However, since N-acetylation can occur, the reaction time and temperature should not be modified from the conditions given in the protocol, in order to avoid this side reaction.
Anticipated Results The allyloxycarbonyl group serves as a protecting group for not only nucleobases but also the sugar hydroxy groups of nucleosides. Further, in the synthesis of nucleotides, the allyl group has been developed as a protecting group for internucleotide linkages. This protecting group is also removed by means of the abovementioned organopalladium method under mild, nearly neutral
2.12.24 Supplement 23
Current Protocols in Nucleic Acid Chemistry
conditions (Hayakawa et al., 1985). Accordingly, the combined use of allyloxycarbonyl and allyl groups for the protection of nucleobases and/or hydroxyls and internucleotide linkages, respectively, has allowed for the efficient synthesis of a variety of natural and artificial nucleic acid– related compounds with base-sensitive structures and functions, producing them in higher yields and at greater levels of purity than methods that use acyl or aroyl and 2cyanoethyl groups for protection of nucleobases and internucleotide linkages, respectively. Compounds prepared by means of the allyl-protection method include O-(cytidine5 -monophosphono)-N-acetylneuraminic acid (CMP-Neu5Ac; Makino et al., 1993), 5 cytosinearabinosyl 5-nitrofurufuryl N-methylN-(4-chlorobutyl) phosphoramidate (Tobias and Borch, 2004), adenylyl(2 –5 )adenylyl(2 – 5 )adenosine (2–5A core; Hayakawa et al., 2001), oligodeoxyribonucleotides with the enzyme-labile 2-(acetylthio)ethyl group on the internucleotide linkage (Spinelli et al., 2002), nucleotide-peptide conjugates with base-labile phosphodiester linkers (Sakakura and Hayakawa, 2000; Jeyaraj and Waldmann, 2001), a DNA oligomer with sitespecific addition of alkaline-labile and mutagenic and/or cytotoxic (5R)-5,6-dihydro5-hydroxythymidine (Matray and Greenberg, 1994), N-acetylcytidine-incorporated RNA oligomers (Bogdan and Chow, 1998), and 2 -deoxy-3-isoadenosine-incorporated DNA oligomers (Bhat et al., 1996). The allylprotection method also allows for high yield and high purity in the synthesis of normal DNA (Hayakawa et al., 1990) and RNA (Hayakawa et al., 2001) oligomers.
Time Considerations Approximate times required for preparation, workup, and purification of key compounds are as follows: S.2: 3 days; S.3: 2 days; S.6: 3 days; S.8: 2 days; S.9: 1 days, S.10: 3 days, S.11: 2 days, S.12: 3 days, S.14: 2 days, S.15: 2 days, S.16: 3 days, S.18: 2 days, S.19: 2 days, S.20: 3 days, S.22: 1 days, S.23: 3 days, S.24: 2 days, S.25: 3 days.
Literature Cited Beaucage, S.L. and Caruthers, M.H. 1981. Deoxynucleoside phosphoramidites—a new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Lett. 22:1859-1862. Bhat, B., Neelima, H., Leonard, N.J., Robinson, H., and Wang, A.H.-J. 1996. 2 -Deoxy-3isoadenosine forms Hoogsteen-type base pairs
with thymidine in the d(CG[iA]TCG)2 duplex. J. Am. Chem. Soc. 118:3065-3066. Bogdan, F.M. and Chow, C.S. 1998. The synthesis of allyl- and allyloxycarbonyl-protected RNA phosphoramidites. Useful reagents for solidphase synthesis of RNAs with base-labile modifications. Tetrahedron Lett. 39:1897-1900. Dorman, M.A., Noble, S.A., McBride, L.J., and Caruthers, M.H. 1984. Synthesis of oligodeoxynucleotides and oligodeoxynucleotide analogs using phosphoramidite intermediates. Tetrahedron 40:95-102. Dreef-Tromp, C.M., Hoogerhout, P., van der Marel, G.A., and van Boom, J.H. 1990. A new protected acyl protecting group for exocyclic amino functions of nucleobases. Tetrahedron Lett. 31:427430. Grotli, M., Douglas, M., Beijer, B., Garcia, G., Eritja, R., and Sproat, B. 1997. Protection of the guanine residue during synthesis of 2 O-alkylguanosine derivatives. J. Chem. Soc., Perkin Trans. 1 18:2779-2788. Hayakawa, Y., Uchiyama, M., Kato, H., and Noyori, R. 1985. Allyl protection of internucleotide linkage. Tetrahedron Lett. 26:6505-6508. Hayakawa, Y., Uchiyama, M., and Noyori, R. 1986a. Nonaqueous oxidation of nucleoside phosphites to the phosphates. Tetrahedron Lett. 27:4191-4194. Hayakawa, Y., Kato, H., Uchiyama, M., Kajino, H., and Noyori, R. 1986b. Allyloxycarbonyl group: A versatile blocking group for nucleotide synthesis. J. Org. Chem. 51:2400-2402. Hayakawa, Y., Wakabayashi, S., Kato, H., and Noyori, R. 1990. The allylic protection method in solid-phase oligonucleotide synthesis. An efficient preparation of solid-anchored DNA oligomers. J. Am. Chem. Soc. 112:1691-1696. Hayakawa, Y., Kawai, R., Hirata, A., Sugimoto, J., Kataoka, M., Sakakura, A., Hirose, M., and Noyori, R. 2001. Acid/azole complexes as highly effective promoters in the synthesis of DNA and RNA oligomers via the phosphoramidite method. J. Am. Chem. Soc. 123:81658176. Heidenhain, S.B. and Hayakawa, Y. 1999. Improved methods for the preparation of 2 deoxyribonucleoside and ribonucleoside 3 phosphoramidites with allylic protectors. Nucleosides Nucleotides Nucleic Acids 18:1771-1787. Himmelsbach, F., Schulz, B.S., Trichtinger, T., Charubala, R., and Pfleiderer, W. 1984. The pnitrophenylethyl (NPE) group: A versatile new blocking group for phosphate and aglycone protection in nucleosides and nucleotides. Tetrahedron 40:59-72. Jeyaraj, D.A. and Waldmann, H. 2001. Synthesis of nucleopeptides by an enzyme labile urethane protecting group. Tetrahedron Lett. 42:835-837. Kataoka, M., Hattori, A., Okino, S., Hyodo, M., Asano, M., Kawai, R., and Hayakawa, Y. 2001. 2-Butanone peroxide as an efficient reagent for the oxidation of nucleoside phosphites into
Protection of Nucleosides for Oligonucleotide Synthesis
2.12.25 Current Protocols in Nucleic Acid Chemistry
Supplement 23
phosphates under anhydrous conditions. Org. Lett. 3:815-818. Koole, L.H., Moody, H.M., Broeders, N.L.H.L., Quaedflieg, P.J.L.M., Kuijpers, W.H.A., van Genderen, M.H.P., Coenen, A.J.J.M., van der Wal, S., and Buck, H.M. 1989. Synthesis of phosphate-methylated DNA fragments using 9-fluorenylmethoxycarbonyl as transient base protecting group. J. Org. Chem. 54:1657-1664. K¨oster, H., Kulikowski, K., Liese, T., Heikens, W., and Kohli, V. 1981. N-Acyl protecting groups for deoxynucleosides: A quantitative and comparative study. Tetrahedron 37:363-369. Kuijpers, W.H.A., Huskens, J., and van Boeckel, C.A.A. 1990. The 2-(acetoxymethyl)benzoyl (AMB) group as a new base–protecting group, designed for the protection of (phosphate) modified oligonucleotides. Tetrahedron Lett. 46:6729-6732. Letsinger, R.L., Finann, J.L., Heavner, G.A., and Lunsford, W.B. 1975. Phosphite coupling procedure for generating internucleotide links. J. Am. Chem. Soc. 97:3278-3279. Makino, S., Ueno, Y., Ishikawa, M., Hayakawa, Y., and Hata, T. 1993. Chemical synthesis of cytidine-5 -monophosphono-Nacetylneuraminic acid (CMP-Neu5Ac). Tetrahedron Lett. 34:2775-2778. Matray, T.J. and Greenberg, M.M. 1994. Sitespecific incorporation of the alkaline labile, oxidative stress product (5R)-5,6-dihydro-5hydroxythymidine in an oligonucleotide. J. Am. Chem. Soc. 116:6931-6932. Matteucci, M.D. and Caruthers, M.H. 1981. Synthesis of deoxyribonucleotides on a polymer support. J. Am. Chem. Soc. 103:3185-3191. McBride, L.J. and Caruthers, M.H. 1983. An investigation of several deoxynucleoside phosphoramidites useful for synthesizing deoxyoligonucleotides. Tetrahedron Lett. 24:245-248.
Schulhof, J.C., Molko, D., and Teoule, R. 1987. The final deprotection step in oligonucleotide synthesis is reduced to a mild and rapid ammonia treatment by using labile base–protecting groups. Nucl. Acids Res. 15:397-416. Sinha, N.D., Biernat, J., and K¨oster, H. 1983. β-Cyanoethyl N,N-dialkylamino/Nmorpholinomonochloro phosphoramidites, new phosphitylating agents facilitating ease of deprotection and work-up of synthesized oligonucleotides. Tetrahedron Lett. 24:58435846. Spinelli, N., Meyer, A., Hayakawa, Y., Imbach, J.-L., and Vasseur, J.-J. 2002. Use of allylic protecting groups for the synthesis of base-sensitive prooligonucleotides. Eur. J. Org. Chem. 49-56. Tobias, S.C. and Borch, R.F. 2004. Synthesis and biological evaluation of a cytarabine phosphoramidite prodrug. Mol. Pharm. 1:112-116. Uznanski, B., Grajkowski, A., and Wilk, A. 1989. The isopropoxyacetic group for convenient base protection during solid–support synthesis of oligodeoxyribonucleotides and their triester analogs. Nucl. Acids Res. 17:4863-4871. Vinogradov, S., Asseline, U., and Thuong, N.T. 1993. Synthesis and physicochemical studies of partially phosphate-methylated oligodeoxyribonucleotides. Tetrahedron Lett. 34:5899-5902. Vu, H., McCollum, C., Jacobson, K., Theisen, P., Vinayak, R., Spiess, E., and Andrus, A. 1990. Fast oligonucleotide deprotection phosphoramidite chemistry for DNA synthesis. Tetrahedron Lett. 31:7269-7272.
Contributed by Mamoru Hyodo and Yoshihiro Hayakawa Nagoya University Nagoya, Japan
Sakakura, A. and Hayakawa, Y. 2000. A novel synthesis of oligonucleotide–peptide conjugates with a base-labile phosphate linker between the two components according to the allyl-protected phosphoramidite strategy. Tetrahedron 56:44274435.
Nucleobase Protection with Allyloxycarbonyl
2.12.26 Supplement 23
Current Protocols in Nucleic Acid Chemistry
Universal 2-(4-Nitrophenyl)ethyl and 2-(4-Nitrophenyl)ethoxycarbonyl Protecting Groups for Nucleosides and Nucleotides
UNIT 2.13
Wolfgang Pfleiderer1 1
Konstanz University, Konstanz, Germany
ABSTRACT A universal blocking group strategy for nucleobases is described, using the 2-(4-nitrophenyl)ethyl (NPE) group for O4 -T-, O4 -U-, O6 -dG-, and O6 -G-protection as well as the 2-(4-nitrophenyl)ethoxycarbonyl (NPEOC) group for amino protection in dC, C, dA, A, dG, and G. Conversion into the corresponding 5 -Odimethoxytrityl derivatives and subsequent phosphitylation to form the fully protected 3 O-(2-cyanoethyl-N,N-diisopropylphosphoramidites) and 3 -O-(2-(4-nitrophenyl)ethylN,N-diisopropylphosphoramidites) produces a new class of interesting building blocks for oligonucleotide synthesis. Curr. Protoc. Nucleic Acid Chem. 30:2.13.1C 2007 by John Wiley & Sons, Inc. 2.13.25. Keywords: NPE and NPEOC protection r Mitsunobu reaction r dimethoxytritylation r phosphitylation r fully protected building blocks
INTRODUCTION The well-established solid-support approach to oligo-2 -deoxyribonucleotides by the phosphoramidite methodology (Caruthers et al., 1982) generally uses monomeric building blocks that have been standardized regarding the functional groups at the sugar moiety (5 -O-dimethoxytrityl-3 -O-(2-cyanoethyl-N,N-diisopropylphosphoramidite), but vary broadly where the protection of the nucleobases is concerned. Usually, the amino groups are blocked by various acyl groups, whereas the amide functions in thymidine, uridine, and guanosine are considered to be chemically inert in the applied strategies. Often, however, solubility problems arise, especially with 2 -deoxyguanosine and guanosine derivatives, which form strong intermolecular hydrogen bonds due to the unprotected amide groups. This unit describes a strategy to overcome these disadvantages using a new universal protecting group that fulfills all anticipated expectations. The outstanding properties of the 2-(4-nitrophenyl)ethyl (NPE) group first applied in the phosphotriester approach (Uhlmann et al., 1981) suggested a more general application of this interesting group for protection of various other functions present in the aglycones and sugar moieties of nucleosides and nucleotides. The NPE group can be used for O-protection of uracil and thymine (see Basic Protocol 1) and of guanine (see Basic Protocols 4 and 6). The related 2-(4-nitrophenyl)ethoxycarbonyl (NPEOC) group is used to protect the amino groups of cytosine (see Basic Protocol 2), adenine (see Basic Protocol 3 and Alternate Protocols 1 and 2), and guanine (see Basic Protocols 5 and 6). The advantage of a universal protection strategy over the multiplicity of previously used protecting group combinations appears obvious at least during the final deprotection steps, which are considerably simplified by this approach. Deprotection generally requires only a DBU treatment to cleave the NPE and NPEOC groups by a ß-elimination process and an acid cleavage of the dimethoxytrityl group to form the free oligonucleotide from its
Current Protocols in Nucleic Acid Chemistry 2.13.1-2.13.25, September 2007 Published online September 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/0471142700.nc0213s30 C 2007 John Wiley & Sons, Inc. Copyright
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2.13.1 Supplement 30
fully protected synthetic precursor. These deblocking steps can even be performed while the oligonucleotide is still attached to the solid support, allowing the clean removal of all protecting groups and reagents. Final ammonia treatment then yields the clean oligomer cleaved from the solid support. BASIC PROTOCOL 1
O4 -p-NPE PROTECTION OF THYMIDINE AND URIDINE Extended studies of the phosphotriester approach revealed some side reactions at the aglycone residues of thymidine and uridine (Reese and Ubasawa, 1980). Condensing agents of the type of arenesulfonyl 1,2,4-triazolide, nitrotriazolide, and tetrazolide, as well as phosphorylating agents, activate the amide function and lead to 4-azol-1-ylpyrimidin2-one nucleosides. This base modification introduces the possibility of nucleophilic displacement reactions leading to cytidine derivatives with ammonia and amines. A successful amide protection has been achieved by phenol and 2,4-dimethylphenol (Jones et al., 1981), two blocking groups that can be cleaved simultaneously with the substituted aryl phosphotriester functions by the oximate procedure. This protocol describes the introduction of the 2-(4-nitrophenyl)ethyl group onto O4, and is shown in Figure 2.13.1. Since the Mitsunobu reaction does not afford O4 but rather N3 substitution, the silver-catalyzed SN 1-type alkylation with 2-(4-nitrophenyl)ethyl iodide in toluene is applied. 3 ,5 -Di-O-acetylthymidine (S.1; Beltz and Visser, 1955) and 2 ,3 ,5 -tri-O-acetyluridine (S.2; Brown et al., 1956) were chosen as starting materials to give the corresponding O4 -2-(4-nitrophenyl)ethyl derivatives S.3 and S.4, which can be deacetylated with ammonia in methanol or dioxane to give O4 -2-(4nitrophenyl)ethylthymidine (S.5) and -uridine (S.6), respectively. The synthesis of the starting reagent, 2-(4-nitrophenyl)ethyl iodide, is as described in Himmelsbach et al. (1984).
Materials Sodium iodide (NaI) Ethyl methyl ketone 2-(4-Nitrophenyl)ethyl chloride Acetone Chloroform (CHCl3 ) 1% aqueous sodium hydrosulfite (Na2 S2 O3 ) Anhydrous sodium sulfate (Na2 SO4 ) Diethyl ether 3 ,5 -Di-O-acetylthymidine (S.1; Beltz and Visser, 1955) or 2 ,3 ,5 -tri-O-acetyluridine (S.2; Brown et al., 1956)
Figure 2.13.1
2-(4-Nitrophenyl)ethylation of 3 ,5 -di-O-acetyl-thymidine (S.1) and 2 ,3 ,5 -tri-O-acetyl-uridine (S.2).
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Current Protocols in Nucleic Acid Chemistry
Absolute toluene Silver carbonate (Ag2 CO3 ) Silica gel 60 (220 to 440 mesh; Fluka) n-Hexane 1,4-Dioxane Saturated ammonia in MeOH Ethyl acetate (EtOAc) Methanol (MeOH) 100- and 250-mL round-bottom flasks Reflux condensers Oil bath G4 glass filters Rotary evaporator equipped with water aspirator Filter paper 30 × 4–cm and 11 × 3–cm chromatography columns 20 × 20 × 0.2–cm TLC silica-gel plates (silica gel PF60254 , Fluka) Silica-coated TLC-plate with fluorescent indicator UV light (254 nm) Vacuum desiccator with P2 O5 Prepare 2-(4-nitrophenyl)ethyl iodide 1. Dissolve 8.8 g (59 mmol) NaI in 100 mL ethyl methyl ketone in a 250-mL roundbottom flask with magnetic stir bar and condenser. 2. Add 10 g (54 mmol) of 2-(4-nitrophenyl)ethyl chloride and reflux 24 hr at 80◦ C. 3. Filter off the solid by suction through a G4 glass filter, wash with 20 mL acetone, and evaporate the combined filtrates on a rotatory evaporator. 4. Dissolve the residue in 150 mL CHCl3 , wash two times with 50 mL of 1% aqueous Na2 S2 O3 solution, and separate the two layers. 5. Dry the organic phase over Na2 SO4 , filter through filter paper, and evaporate again under reduced pressure to dryness. 6. Recrystallize the residue from 30 mL diethyl ether and 3 mL CHCl3 to give 2-(4nitrophenyl)ethyl iodide as yellowish crystals after standing overnight at 7◦ C. 2-(4-Nitrophenyl)ethyl iodide: yield 13.9 g (93%). m.p. 100o -101◦ C. C8 H8 INO2 (277.0): C 34.68, H 2.91, N 5.06; found: C 34.73, H 2.88, N 4.97.
Perform 2-(4-nitrophenyl)ethylation 7. Prepare a suspension of 10 mmol (3.26 g) 3 ,5 -di-O-Ac-T (S.1) or 10 mmol (3.6 g) 2 ,3 ,5 -tri-O-Ac-U (S.2) in 50 mL absolute toluene and 15 mmol (2.5 g) Ag2 CO3 in a 100-mL round-bottom flask with magnetic stir bar and condenser. Heat 2 hr under reflux to 110◦ C. 8. Cool to 50◦ C. Add 15 mmol (4.16 g) 2-(4-nitrophenyl)ethyl iodide and stir 48 hr at 50◦ C. 9. Filter off the precipitate through a G4 glass filter by suction, wash with CHCl3 , and evaporate the filtrate on a rotary evaporator. 10. Dissolve the residue in CHCl3 and load the solution onto a silica gel column (30 × 4–cm, 100 g silica gel 60). Elute first with 100 mL CHCl3 followed by 300 mL of 4:1 CHCl3 /acetone. 11. Collect the product fractions (check by TLC) and evaporate under reduced pressure to give a solid foam. Current Protocols in Nucleic Acid Chemistry
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2.13.3 Supplement 30
12. Further purify a small sample by preparative TLC on a silica gel plate (20 × 20 × 0.2–cm) with CHCl3 . Elute the main zone, evaporate, and dissolve the residue in 5 mL CHCl3 . 13. Precipitate dissolved residue by adding it dropwise to 100 mL of rapidly stirring n-hexane, yielding an amorphous solid. 14. Dry 1 day in a vacuum desiccator over P2 O5 . 3 ,5 -Di-O-acetyl-O4 -NPE-T (S.3): yield 3.47 g (73%). Rf (CHCl3 /acetone 10:1): 0.41. UV (MeOH): 276 (4.19). 1 H NMR (CDCl3 ): 8.14 (d, 2H, H o to NO2 ); 7.51 (s, 1H, H-C6); 7.39 (d, 2H, H m to NO2 ); 6.29 (m, 1H, H-C1 ); 5.17 (m, 1H, H-C3 ); 4.63 (t, 2H, OCH2 CH2 ); 4.33 (m, 2H, H-C5 ); 4.26 (m, 1H, H-C4 ); 3.16 (t, 2H, OCH2 CH2 ); 2.67 (m, 1H, H-C2 ); 2.06 + 2.07 (2s, 6H, 2CH3 ); 2.00 (m, 1H, H-C2 ); 1.87 (s, 3H, H3 C-C5). C22 H25 N3 O9 (475.5): C 55.58, H 5.30, N 8.84; found: C 55.39, H 5.39, N 8.62. 2 ,3 ,5 -Tri-O-acetyl-O4 -NPE-U (S.4): yield 3.84 g (74%). Rf (toluene/EtOAc 1:1): 0.37. UV (MeOH): 204 (4.47), 273 (4.20). 1 H NMR (CDCl3 ): 8.18 (d, 2H, H o to NO2 ); 7.71 (s, 1H, H-C6); 7.42 (d, 2H, H m to NO2 ); 6.13 (d, 1H, H-C1 ); 5.91 (d, 1H, H-C5); 5.34 (m, 2H, H-C2 ,3 ); 4.66 (t, 2H, OCH2 CH2 ); 4.40 (m, 3H, H-C4 ,5 ); 3.16 (t, 2H, OCH2 CH2 ); 2.13 (s, 3H, CH3 ); 2.11 (s, 6H, 2CH3 ). C23 H25 N3 O11 (519.5): C 53.18, H 4.85, N 8.09; found: C 53.70, H 4.71, N 7.67.
Deacetylate 15. Prepare a solution of 1.5 mmol (1.15 g) S.3 or 1.5 mmol (1.9 g) S.4 in 5 mL of 1,4-dioxane. Add 10 mL saturated ammonia in MeOH and stir 24 hr at room temperature. 16. Evaporate on a rotary evaporator. 17. Either recrystallize the residue from EtOAc or purify by column chromatography. For chromatography, dissolve the residue in CHCl3 , put onto a silica gel column for chromatography (11 × 3–cm, 30 g), and elute first with 100 mL CHCl3 and then with 500 mL of 20:1 CHCl3 /MeOH. 18. Collect the product fractions (check by TLC) and evaporate in a vacuum to give a solid foam. 19. Recrystallize from EtOAc to give colorless crystals after standing overnight at 7◦ C. O4 -NPE-T (S.5): yield 0.55 g (94%). m.p. 143o -148◦ C. Rf (toluene/EtOAc/MeOH 5:4:1): 0.23. UV (MeOH): 276 (4.19). 1 H NMR (CDCl3 ): 8.14 (d, 2H, H o to NO2 ); 7.68 (s, 1H, H-C6); 7.40 (d, 2H, H m to NO2 ); 6.10 (pt, 1H, H-C1 ); 4.63-4.55 (m, 3H, H-C3 , OCH2 CH2 ); 4.03 (m, 1H, H-C4 ); 3.85 (m, 2H, H-C5 ); 3.27-3.13 (m, 4H, OCH2 CH2 , HO-C3 , HO-C5 ); 2.40 (m, 2H, H-C2 ,2 ); 1.86 (s, 3H, H3 C-C5). C18 H21 N3 O7 (391.4): C 55.24, H 5.41, N 10.74; found: C 55.26, H 5.21, N 10.71. O4 -NPE-U (S.6): yield 0.50 g (85%). m.p. 147o -148◦ C. Rf (CHCl3 /MeOH 95:5): 0.78. UV (MeOH): 204 (4.43), 275 (4.22). 1 H NMR (DMSO-d6 ): 8.29 (s, 1H, H-C6); 8.17 (d, 2H, H o to NO2 ); 7.58 (d, 2H, H m to NO2 ); 5.98 (d, 1H, H-C5); 5.76 (pd, 1H, H-C1 ); 5.44 (d, 1H, HO-C2 ); 5.14 (d, 1H, HO-C5 ); 5.02 (d, 1H, HO-C3 ); 4.52 (t, 2H, OCH2 CH2 ); 3.90 (m, 3H, H-C2 ,3 ,4 ); 3.70-3.56 (m, 2H, H-C5 ); 3.16 (t, 3H, OCH2 CH2 ). C17 H19 N3 O8 (393.4): C 51.91, H 4.87, N 10.68; found: C 51.42, H 4.90, N 10.69. BASIC PROTOCOL 2
Universal NPE and NPEOC Protecting Groups
N4 -p-NPEOC PROTECTION OF 2 -DEOXYCYTIDINE AND CYTIDINE The amino group in 2 -deoxycytidine (S.7) and cytidine (S.8) is highly nucleophilic, which allows direct 2-(4-nitrophenyl)ethoxycarbonylation with the appropriate acylating agent, e.g., 1-[2-(4-nitrophenyl)ethoxycarbonyl]benztriazole, without protection of the sugar hydroxy groups. The method is shown in Figure 2.13.2.
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Figure 2.13.2
2-(4-Nitrophenyl)ethoxycarbonylation of 2 -deoxycytidine (S.7) and cytidine (S.8).
Materials 2-(4-Nitrophenyl)ethanol (Sigma-Aldrich) Methylenechloride (CH2 Cl2 , analytical grade), cold 1-(Chlorocarbonyl)benztriazole Triethylamine Anhydrous sodium sulfate (Na2 SO4 ) Benzene (analytical grade) Cytidine or 2 -deoxycytidine (Sigma-Aldrich, Pharma Waldhof) 1-[2-(4-Nitrophenyl)ethoxycarbonyl]benztriazole N,N-Dimethylformamide (DMF, anhydrous) 100- and 250-mL round-bottom flasks Reflux condenser 60◦ C oil bath G4 glass filters 50◦ C oven Prepare 1-[2-(4-nitrophenyl)ethoxycarbonyl]benztriazole 1. Dissolve 8.35 g (50 mmol) 2-(4-nitrophenyl)ethanol in 70 mL cold absolute CH2 Cl2 (0◦ to 5◦ C) in a 250-mL round-bottom flask with magnetic stir bar. Add 9.05 g (50 mmol) 1-(chlorocarbonyl)benztriazole. 2. Add slowly (dropwise) a solution of 10 mL triethylamine in 10 mL CH2 Cl2 under stirring. 3. Warm to room temperature with stirring for 1 hr, and continue stirring for 30 min at room temperature. 4. Add 250 mL CH2 Cl2 and shake the solution three times with 200 mL ice water. 5. Separate the organic layer, dry over Na2 SO4 , filter through filter paper, and evaporate to dryness. 6. Recrystallize the residue from 120 mL dry benzene to give yellowish crystals after standing overnight at room temperature. 1-[2-(4-Nitrophenyl)ethoxycarbonyl]benztriazole: yield 14.4 g (92%). m.p. 135o -136◦ C. 1 H NMR (CDCl3 ): 8.22 (d, 2H, arom. H); 8.05 (d, 2H, arom. H); 7.65 (m, 1H, arom. H); 7.54 (d, 2H, arom. H); 7.52 (m, 1H, arom. H); 4.88 (t, 2H, CH2 ); 3.37 (t,
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2H, CH2 ). Rf (toluene/EtOAc 7:3): 0.65. C15 H12 N4 O4 (312.3): C 57.69, H 3.87, N 17.94; found: C 57.59, H 3.79, N 17.95.
Perform nitrophenylethoxycarbonylation 7. Dissolve 15 mmol (3.43 g) 2 -deoxycytidine (S.7) or 15 mmol (3.63 g) cytidine (S.8) and 4.7 g (15 mmol) 1-[2-(4-nitrophenyl)ethoxycarbonyl]benztriazole in a 100-mL round-bottom flask with condenser in 50 mL absolute DMF and heat 20 hr in a 60◦ C oil bath with magnetic stirring. 8. Distill off the DMF under high vacuum and treat the oily residue with a mixture of 30 mL H2 O and 30 mL CH2 Cl2 with vigorous shaking. 9. Collect the resulting precipitate by suction through a G4 glass filter, wash with H2 O, and dry in a 50◦ C oven to give colorless crystals. N4 -NPEOC-2 -dC (S.9): yield 5.86 g (91%). m.p. 114o -120◦ C. Rf (CHCl3 /MeOH 19:1): 0.28. UV: 242 (4.24), 281 (4.16). 1 H NMR (DMSO-d6 ): 10.72 (s, 1H, H-N); 8.28 (d, 1H, H-C6); 8.15 (d, 2H, H o to NO2 ); 7.59 (d, 2H, H m to NO2 ); 6.97 (d, 1H, H-C5); 6.08 (pt, 1H, H-C1 ); 5.26 (d, 1H, HO-C3 ); 5.04 (t, 1H, HO-C5 ); 4.35 (t, 2H, OCH2 CH2 ); 4.20 (m, 1H, H-C3 ); 3.84 (m, 1H, H-C4 ); 3.57 (m, 2H, H-C5 ); 3.08 (t, 2H, OCH2 CH2 ); 2.26 (m, 1H, H-C2 ); 2.00 (m, 1H, H-C2 ). C18 H20 N4 O8 ·H2 O (438.4): C 50.35, H 4.93, N 13.05; found: C 50.63, H 4.67, N 13.03. N4 -NPEOC-C (S.10): yield 4.77 g (70%). m.p. 85o -89◦ C. Rf (CHCl3 /MeOH 9:1): 0.30. UV: 242 (4.23), 280 (4.14). 1 H NMR (DMSO-d6 ): 10.76 (bs, 1H, H-N); 8.39 (d, 1H, H-C6); 8.17 (d, 2H, H o to NO2 ); 7.60 (d, 2H, H m to NO2 ); 6.96 (d, 1H, H-C5); 5.76 (d, 1H, H-C1 ); 5.49 (d, 1H, HO-C2 ); 5.16 (t, 1H, HO-C5 ); 5.05 (d, 1H, HO-C3 ); 4.35 (t, 2H, OCH2 CH2 ); 4.02-3.82 (m, 3H, H-C4 ,3 ,2 ); 3.80-3.52 (m, 2H, H-C5 ); 3.08 (t, 2H, OCH2 CH2 ). C18 H20 N4 O9 ·H2 O (454.4): C 47.58, H 4.88, N 12.33; found: C 47.36, H 4.51, N 12.29. BASIC PROTOCOL 3
N6 -p-NPEOC PROTECTION OF ACETYLATED 2 -DEOXYADENOSINE AND ADENOSINE Direct 2-(4-nitrophenyl)ethoxycarbonylation of 2 -deoxyadenosine and adenosine can not be achieved with 1-[2-(4-nitrophenyl)ethoxycarbonyl]benztriazole in the same manner as in the cytidine series due to the lower nucleophilic potential of the amino function of adenosine. The more reactive 2-(4-nitrophenyl)ethyl chloroformate in pyridine, however, smoothly acylates 3 ,5 -di-O-acetyl-2 -deoxyadenosine (S.13; Hayes et al., 1955) and 2 ,3 ,5 -tri-O-acetyladenosine (S.14; Bredereck, 1947), respectively, to form mixtures of N6 -mono-(S.15, S.16) and N6 -di-2-(4nitrophenyl)ethoxycarbonyl derivatives (S.17, S.18; Fig. 2.13.3). These can be converted with ammonia into N6 -2-(4-nitrophenyl)ethoxycarbonyl-2 -deoxyadenosine (S.19) and N6 -2-(4-nitrophenyl)ethoxycarbonyladenosine (S.20). Two strategies for monoalkylation of adenosine are presented in Alternate Protocols 1 and 2.
Materials
Universal NPE and NPEOC Protecting Groups
Toluene (anhydrous) Phosgene 2-(4-Nitrophenyl)ethanol (Sigma-Aldrich) Methylenechloride (CH2 Cl2 ) 3 ,5 -Di-O-acetyl-2 -deoxyadenosine (S.13; Hayes et al., 1955) or 2 ,3 ,5 -tri-O-acetyladenosine (S.14; Bredereck, 1947) Pyridine (anhydrous) Chloroform (CHCl3 ) Phosphate buffer, pH 7 Na2 SO4 Toluene
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Figure 2.13.3 2-(4-Nitrophenyl)ethoxycarbonylation of 3 ,5 -di-O-acetyl-2 -deoxy- (S.13) and 2 ,3 ,5 -tri-O-acetyladenosine (S.14).
Silica gel Methanol (MeOH) Concentrated aqueous ammonia Ethanol (EtOH) 500-mL three-neck round-bottom flasks 50◦ C oil bath Condenser with drying tube Gas inlet tubes 20 × 4–cm chromatography column 250- and 500-mL round-bottom flasks Rotatory evaporator Prepare 2-(p-nitrophenyl)ethyl chloroformate 1. Using a 500-mL three-neck round-bottom flask with condenser and gas inlet tube, cool 200 mL of dry toluene in an ice bath to 0◦ to 5◦ C and bubble 70 g (0.7 mol) phosgene from a storage tank into the solution under stirring. 2. Prepare a solution of 33.4 g (0.2 mol) 2-(4-nitrophenyl)ethanol in a mixture of 100 mL CH2 Cl2 and 50 mL toluene, and add dropwise with stirring to the phosgene solution. 3. Allow to warm to room temperature, then stir 1 hr at room temperature, and finally raise slowly to 50◦ C (within 5 hr) in an oil bath. 4. After 5 hr at 50◦ C, cool the mixture to room temperature (∼1 hr) and evaporate under vacuum. 5. Let stand overnight to allow the residue to crystallize. Dry under high vacuum at room temperature to give chromatographically pure material that can be used for further reactions. 6. Obtain an analytically pure sample by recrystallization from toluene to give yellowish crystals. 2-(4-Nitrophenyl)ethyl chloroformate: yield 45.0 g (98%). m.p. 42◦ C. 1 H NMR (CDCl3 ): 8.15 (d, 2H, arom. H); 7.38 (d, 2H, arom. H); 4.53 (t, 2H, CH2 ); 3.14 (t, 2H, CH2 ).
Perform nitrophenylethoxycarbonylation 7. Prepare an ice-cold solution of 5 mmol (1.68 g) 3 ,5 -di-O-Ac-2 -dA (S.13) or 5 mmol (2.02 g) 2 ,3 ,5 -tri-O-Ac-A (S.14) in 10 mL absolute pyridine.
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8. With magnetic stirring, add 12.5 mmol (2.88 g) 2-(4-nitrophenyl)ethyl chloroformate. Continue stirring 1 hr at 0◦ to 5◦ C and then 2 hr at room temperature. 9. Dilute with 200 mL CHCl3 and treat with 100 mL phosphate buffer, pH 7, three times with intensive shaking, separating the organic layer each time. 10. Wash the CHCl3 phase with H2 O, dry over Na2 SO4 , evaporate under reduced pressure to dryness, and coevaporate three times with 20 mL toluene. 11. Dissolve the residue in 5 mL CHCl3 , place onto to a silica gel column (20 × 4–cm), elute the reagents first with CHCl3 , and then co-elute with 49:1 (v/v) CHCl3 /MeOH the mixed N6 -mono- and N6 -di-NPEOC derivatives S.15 plus S.17 and S.16 plus S.18, respectively. See Alternate Protocol 1 for characterization data for S.15 and S.16.
Remove acetyl groups and additional NPE group 12. Evaporate the mixture of S.15 plus S.17 or S.16 plus S.18, respectively, in a 500-mL round-bottom flask, dissolve the residue in 200 mL MeOH, and then treat with 60 mL concentrated aqueous ammonia under magnetic stirring for 1 hr at room temperature. 13. Evaporate and recrystallize the residue from 20 mL EtOH and 2 mL H2 O to give 1.78 g (80%) of S.19 and 1.9 g (83%) of S.20, respectively, as colorless crystals. See Alternate Protocol 1 for characterization data for S.19 and S.20. ALTERNATE PROTOCOL 1
MONO-N6 -p-NPEOC PROTECTION OF ACETYLATED 2 -DEOXYADENOSINE AND ADENOSINE By analogy to various reports (Watkins et al., 1982), monoalkoxycarbonylation of S.13 and S.14 can be effected directly with 1-methyl-3-[2-(4-nitrophenyl)ethoxycarbonyl]imidazolium chloride (prepared from 2-(4-nitrophenyl)ethyl chloroformate and N-methylimidazole in CH2 Cl2 ) to give S.15 and S.16, respectively, in 85% yield. Deacetylation with triethylamine in MeOH produces S.19 and S.20 almost quantitatively.
Materials 2-(4-Nitrophenyl)ethyl chloroformate (see Basic Protocol 3) Methylenechloride (CH2 Cl2 anhydrous) N-Methylimidazole N2 atmosphere 3 ,5 -Di-O-acetyl-2 -deoxyadenosine (S.13; Hayes et al., 1955) or 2 ,3 ,5 -tri-O-acetyladenosine (S.14; Bredereck, 1947) 1,2-Dichloroethane Silica gel Chloroform (CHCl3 ) Methanol (MeOH) Ethyl acetate (EtOAc) P2 O5 Triethylamine Ethanol
Universal NPE and NPEOC Protecting Groups
100- and 500-mL round-bottom flasks Reflux condensers Drying tube G4 glass filters High-vacuum pump Drying pistol Rotatory evaporator
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Chromatography column (8 × 30–cm) Silica-coated TLC plate with fluorescent indicator 50◦ C oven Prepare 1-methyl-3-[2-(4-nitrophenyl)ethoxycarbonyl]imidazolium chloride 1. In a 100-mL round-bottom flask with condenser and drying tube, dissolve 20 mmol (4.6 g) 2-(4-nitrophenyl)ethyl chloroformate in 40 mL dry CH2 Cl2 . Cool in an ice bath and then add 20 mmol (1.7 mL) N-methylimidazole under magnetic stirring. 2. Stir the mixture 15 min at 0◦ to 5◦ C and then another 30 min at room temperature. 3. Collect the precipitate by suction under a dry N2 atmosphere, wash with absolute CH2 Cl2 , and dry under high vacuum in a drying pistol. 1-Methyl-3-[2-(4-nitrophenyl)ethoxycarbonyl]imidazolium chloride: yield 5.0 g (80%). m.p. 92o -95◦ C. 1 H NMR (DMSO-d6 ): 10.15 (s, 1H, H-C2); 8.17 (d, 2H, arom. H); 8.09 (d, 1H, H-C4); 7.98 (d, 1H, H-C5); 7.70 (d, 2H, arom. H); 4.74 (t, 2H, CH2 ); 4.00 (s, 3H, CH3 ); 3.27 (t, 2H, CH2 ). C13 H14 ClN3 O4 (311.7): C 50.09, H 4.53, N 13.48; found: C 49.98, H 4.44, N 13.38.
Perform nitrophenylethoxycarbonylation 4. Dissolve 10 mmol (3.35 g) 3 ,5 -di-O-Ac-2 -dA (S.13) or 10 mmol (4.0 g) 2 ,3 ,5 tri-O-Ac-A (S.14) in 60 mL dry CH2 Cl2 . Add 14 mmol (4.36 g) 1-methyl-3-[2-(4nitrophenyl)ethoxycarbonyl]imidazolium chloride and stir 18 hr at room temperature. 5. Filter off the precipitate from the suspension by suction through a G4 glass filter, wash the residue with CH2 Cl2 , combine the filtrates, and evaporate under vacuum. 6. Dissolve the resulting residue in 1,2-dichloroethane for short-column silica-gel (50 g) chromatography using an 8 × 30–cm chromatography column (Hunt and Rigby, 1967) and subsequently elute with 1,2-dichloroethane, CHCl3 , and 50:1 (v/v) CHCl3 /MeOH. Do not use >1 atm pressure for elution. 7. Collect the product fractions (check by TLC), evaporate, and treat the resulting solid with EtOAc to give colorless crystals. 8. Dry under vacuum in a drying pistol over P2 O5 . 3 ,5 -Di-O-acetyl-N6 -NPEOC-2 -dA (S.15): yield 4.15 g (78%). m.p. 134o -136◦ C. Rf (CHCl3 /MeOH 19:1): 0.63. UV (MeOH): 267 (4.45). 1 H NMR (CDCl3 ): 8.70 (bs, 1H, H-N); 8.66 (s, 1H, H-C8); 8.11 (d, 2H, H o to NO2 ); 8.11 (s, 1H, H-C2); 7.39 (d, 2H, H m to NO2 ); 6.44 (dd, 1H, H-C1 ); 5.42 (m, 1H, H-C3 ); 4.53 (t, 2H, OCH2 CH2 ); 4.35 (m, 3H, H-C4 ,5 ); 3.14 (t, 2H, OCH2 CH2 ); 2.95 (m, 1H, H-C2 ); 2.70-2.58 (m, 1H, H-C2 ); 2.13 (s, 3H, ac); 2.06 (s, 3H, ac). C23 H24 N6 O9 (528.5): C 52.27, H 4.58, N 15.90; found: C 52.51, H 4.59, N 15.93. 2 ,3 ,5 -Tri-O-acetyl-N6 -NPEOC-A (S.16): yield 5.1 g (87%). m.p. 149o -151◦ C. Rf (CHCl3 /MeOH 19:1): 0.69. UV (MeOH): 267 (4.46). 1 H NMR (CDCl3 ): 8.70 (s, 1H, H-C8); 8.47 (bs, 1H, H-N); 8.14 (d, 2H, H o to NO2 ); 8.09 (s, 1H, H-C2); 7.40 (d, 2H, H m to NO2 ); 6.18 (dd, 1H, H-C1 ); 5.90 (pt, 1H, H-C2 ); 5.64 (m, 1H, H-C3 ); 4.52 (t, 2H, OCH2 CH2 ); 4.48-4.30 (m, 3H, H-C4 ,5 ); 3.14 (t, 2H, OCH2 CH2 ); 2.14 (s, 3H, ac); 2.10(s, 3H, ac); 2.06 (s, 3H, ac). C25 H26 N6 O11 (586.5): C 51.20, H 4.47, N 14.33; found: C 51.19, H 4.36, N 14.58.
Deacetylate 9. Dissolve 5 mmol (2.64 g) S.15 or 5 mmol (2.93 g) S.16 in 300 mL MeOH and add 60 mL triethylamine at room temperature. 10. Stir for 18 hr, filter off the resulting precipitate by suction through a G4 glass filter, wash the solid with MeOH, and dry overnight in a 50◦ C oven.
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11. Evaporate the reaction filtrate to dryness in vacuum, shake the residue with a mixture of 40 mL H2 O and 40 mL of 1,2-dichloroethane to get a second crop of chromatographically pure crystals. 12. Recrystallize from EtOH to get an analytical sample if necessary. N6 -NPEOC-2 -dA (S.19): yield: 2.17 g (94%). m.p. 100o -110◦ C. Rf (CHCl3 /MeOH 9:1) 0.37. UV(MeOH): 267 (4.44). 1 H NMR (DMSO-d6 ): 10.56 (s, 1H, H-N); 8.62 (s, 1H, H-C8); 8.60 (s, 1H, H-C2); 8.16 (d, 2H, H o to NO2 ); 7.60 (d, 2H, H m to NO2 ); 6.42 (pt, 1H, H-C1 ); 5.37 (d, 1H, HO-C3 ); 5.03 (t, 1H, HO-C5 ); 4.50-4.30 (m, 3H, H-C3 , OCH2 CH2 ); 3.89 (m, 1H, H-C4 ); 3.70-3.45 (m, 2H, H-C5 ); 3.11 (t, 2H, OCH2 CH2 ); 2.76 (m, 1H, H-C2 ); 2.32 (m, 1H, H-C2 ). C19 H20 N6 O7 × H2 O (462.4): C 49.35, H 4.80, N 18.17; found: C 49.29, H 4.74, N 18.04. N6 -NPEOC-A (S.20): yield: 1.96 g (85%). m.p. 152o -160◦ C. Rf (CHCl3 /MeOH 9:1) 0.37. UV (MeOH): 267 (4.45). 1 H NMR (DMSO-d6 ): 10.60 (s, 1H, H-N); 8.86 (s, 1H, H-C8); 8.62 (s, 1H, H-C2); 8.17 (d, 2H, H o to NO2 ); 7.61 (d, 2H, H m to NO2 ); 5.99 (pt, 1H, H-C1 ); 5.53 (d, 1H, HO-C2 ); 5.24 (d, 1H, HO-C3 ); 5.14 (t, 1H, HO-C5 ); 4.62 (m, 1H, H-C2 ); 4.40 (t, 2H, OCH2 CH2 ); 4.18 (m, 1H, H-C3 ); 3.98 (m, 1H, H-C4 ); 3.75-3.50 (m, 2H, H-C5 ); 3.11 (t, 2H, OCH2 CH2 ). C19 H20 N6 O8 (460.4): C 49.57, H 4.38, N 18.25; found: C 49.42, H 4.26, N 18.30. ALTERNATE PROTOCOL 2
N6 -p-NPEOC PROTECTION OF TRIMETHYLSILYLATED 2 -DEOXYADENOSINE An even simpler one-pot reaction leads directly to S.19 and S.20, respectively, starting from 2 -deoxyadenosine (S.11) and adenosine (S.12). The starting nucleosides are first silylated by hexamethyldisilazane to form 3 ,5 -bis-O-trimethylsilyl-2 -deoxyadenosine (S.21) and 2 ,3 ,5 -tri-O-trimethylsilyladenosine (S.22) as sugar-protected intermediates. Addition of 1-methyl-3-[2-(4-nitrophenyl)ethoxycarbonyl]imidazolium chloride causes monoacylation, and work-up with triethylamine in methanol leads to S.19 and S.20, respectively, in 86% to 90% yield.
Materials 2 -Deoxyadenosine or adenosine (Sigma-Aldrich, Pharma Waldhof) Dioxane (anhydrous) Hexamethyldisilazane (Sigma-Aldrich) Ammonium sulfate ((NH4 )2 SO4 ) Toluene (anhydrous) Methylenechloride (CH2 Cl2 , anhydrous) 1-Methyl-3-(2-(4-nitrophenyl)ethoxycarbonyl)imidazolium chloride Triethylamine Methanol (MeOH) 100-mL round-bottom flasks Reflux condenser with drying tube 120◦ C oil bath G4 glass filters Rotatory evaporator 50◦ C drying oven
Universal NPE and NPEOC Protecting Groups
Perform trimethylsilylation 1. In a 100-mL round-bottom flask with stir bar and a condenser with a drying tube, heat a mixture of 10 mmol (2.54 g) dry 2 -deoxyadenosine (S.11) or 10 mmol (2.7 g) dry adenosine (S.12) in 25 mL dry dioxane with 25 mL hexamethyldisilazane and 50 mg (NH4 )2 SO4 under reflux 3 hr in a 120◦ C oil bath to achieve trimethylsilylation of the hydroxyl functions.
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2. Evaporate under vacuum, dissolve the residue in 100 mL dry toluene, and filter off the insoluble material through a G4 glass filter. 3. Evaporate the filtrate to give S.21 and S.22, respectively, as as a colorless syrup.
Perform nitrophenylethyloxycarbonylation 4. Dissolve S.21 or S.22 in 200 mL dry CH2 Cl2 , then add 20 mmol (6.24 g) 1-methyl3-[2-(4-nitrophenyl)ethoxycarbonyl]imidazolium chloride and stir 18 hr at room temperature. 5. Filter off the insoluble material from the suspension through a G4 glass filter and evaporate the filtrate to dryness under vacuum.
Desilylate 6. Dissolve the residue in 100 mL MeOH, add 25 ml triethylamine, and stir overnight at room temperature. 7. Collect the precipitate solid, wash with MeOH, and dry overnight in a 50◦ C drying oven to give S.19 in 86% yield or S.20 in 90% yield. See Alternate Protocol 1 for characterization data for S.19 and S.20.
O6 -p-NPE PROTECTION OF ACYLATED 2 -DEOXYGUANOSINE AND GUANOSINE DERIVATIVES
BASIC PROTOCOL 4
A long-standing problem in oligonucleotide synthesis is dealing with the various side reactions of the guanine residue as well as its low solubility in organic solvents. Protection of the amide function in the guanine moiety seems to be an obvious necessity to overcome these difficulties. Various efforts have been undertaken to protect the O6 position using the 2-nitrophenyl (Jones et al., 1981), silyl, sulfonyl, phosphoryl, and phosphinothioyl groups (Daskalov et al., 1981), as well as substituted ethyl groups that are prone to ß-elimination cleavage reactions (Gaffney and Jones, 1982). The initial introduction of the 2-(4-nitrophenyl)ethyl group in the O6 position was a multistep procedure and the expected nucleophilic displacement of a 6-chloro or 6-methanesulfonyl substituent by 2-(4nitrophenyl)ethanol was unsuccessful. However, as reported in Trichtinger et al. (1983), the Mitsunobu reaction (Mitsunobu, 1981) offers a good approach to direct O-alkylation if the 2-amino group of the guanine moiety is acylated. In this method, outlined in Figure 2.13.4, treatment of N2 ,3 ,5 -triisobutyryl (S.23), N2 ,2 ,3 ,5 -tetraisobutyryl (S.24), and N2 ,2 ,3 ,5-tetrabenzoyl (S.25) guanine residues with 1.5 molar equivalent each of diethyl azodicarboxylate, triphenylphosphine, and 2-(4-nitrophenyl)ethanol, respectively, in 1,2-dioxane leads directly to the corresponding O6 -2-(4-nitrophenyl)ethyl derivatives S.26 to S.28 in high yields. Cleavage of the sugar acyl groups in conc. ammonia/MeOH proceeds in 3 hr to give S.31 to S.33. Finally, cleavage of the N2 -acyl group, without harming the O6 -NPE substituent, is best performed by prolonged treatment with aqueous ammonia in methanol (up to 5 to 6 days), yielding S.34 and S.35, respectively.
Materials N2 ,3 ,5 -Triisobutyryl-2 -deoxyguanosine (S.23; Flockerzi et al., 1981), N2 ,2 ,3 ,5 -tetraisobutyrylguanosine (S.24; Pharma Waldhof), or N2 ,2 ,3 ,5 -tetrabenzoylguanosine (S.25; Himmelsbach et al., 1984) Triphenylphosphine 2-(4-Nitrophenyl)ethanol (Sigma-Aldrich) 1,2-Dioxane (anhydrous) Diethyl azodicarboxylate (DEAD; Sigma-Aldrich) Chloroform (CHCl3 ) Methanol (MeOH)
Protection of Nucleosides for Oligonucleotide Synthesis
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Figure 2.13.4
Mitsunobu reaction with acylated 2 -deoxyguanosine and guanosine derivatives.
n-Hexane Tetrachloromethane (CCl4 ) Conc. aqueous ammonia Saturated methanolic ammonia Ethanol (EtOH) Silica-gel PF 60254 (Sigma-Aldrich) Toluene Ethyl acetate (EtOAc) Silica gel 60, 220 to 440 mesh (Sigma-Aldrich) 100- and 250-mL round-bottom flasks G4 glass filters 60 × 2.5–cm and 20 × 5–cm chromatography columns Preparative silica-gel plates (20 × 20 × 0.2–cm) Silica-coated TLC plate with fluorescent indicator Rotatory evaporator Drying pistol 50◦ C oven
Universal NPE and NPEOC Protecting Groups
Perform nitrophenylethylation 1. Dissolve a mixture of 2 mmol (0.95 g) N2 ,3 ,5 -triisobutyryl-2 -dG (S.23), 2 mmol (1.13 g) N2 ,2 ,3 ,5 -tetraisobutyryl-G (S.24), or 2 mmol (1.4 g) N2 ,2 ,3 ,5 tetrabenzoyl-G (S.25) plus 3 mmol triphenylphosphine (0.77 g) and 3 mmol 2(4-nitrophenyl)ethanol (0.5 g) in 40 mL dry dioxane.
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2. Add 3 mmol diethyl azodicarboxylate (0.52 g) and stir magnetically 24 hr at room temperature (Himmelsbach et al., 1984). 3. Evaporate to a smaller volume and filter off the precipitated diethyl hydrazinedicarboxylate. 4. Evaporate the filtrate to dryness, dissolve the residue in CHCl3 , and put onto a silica gel column (20 × 5–cm) for chromatography, eluting with 300 mL CHCl3 followed by 99:1 (v/v) CHCl3 /MeOH. 5. Collect the product fractions (check by TLC), evaporate to dryness, and recrystallize the resulting solid either from CHCl3 /n-hexane or from CCl4 . N2 ,3 ,5 -Triisobutyryl-O6 -NPE-2 -dG (S.26): yield: 1.06 g (85%). Rf (CHCl3 /MeOH 19:1) 0.49. UV: 218 (4.42), 269 (4.40). 1 H NMR (CDCl3 ): 8.13 (s, 2H, H o to NO2 ); 7.94 (s, 1H, H-N); 7.92 (s, 1H, H-C8); 7.49 (d, 2H, H m to NO2 ); 6.34 (m, 1H, H-C1 ); 5.39 (m, 1H, H-C3 ); 4.81 (t, 2H, OCH2 CH2 ); 4.48-4.35 (m, 2H, H-C5 ); 4.30 (m, 1H, (H-C4 ); 3.18 (t, 2H, OCH2 CH2 ); 2.99-2.88 (m, 2H, H-C2 ,2 ); 2.61-2.51 (m, 3H, H-CMe2 ); 1.27-1.11 (m, 18H, 3 (H3 C)2 CH). C30 H37 N6 O9 (625.7): C 57.59, H 5.96, N 13.43; found: C 57.10, H 5.91, N 13.11. N2 ,2 ,3 ,5 -Tetraisobutyryl-O6 -NPE-G (S.27): yield: 1.01 g (70%). Rf (CHCl3 /MeOH 19:1) 0.87. UV: 217 (4.46), 268 (4.46). 1 H NMR (CDCl3 ): 8.14 (s, 2H, H o to NO2 ); 7.96 (s, 1H, H-N); 7.91 (s, 1H, H-C8); 7.50 (d, 2H, H m to NO2 ); 6.06 (m, 1H, HC1 ); 5.86 (m, 1H, H-C2 ); 5.76 (m, 1H, H-C3 ); 4.82 (t, 2H, OCH2 CH2 ); 4.41 (m, 3H, H-C4 ,5 ); 3.18 (t, 2H, OCH2 CH2 ); 2.63-2.50 (m, 3H, H-CMe2 ); 1.26-1.08 (m, 18H, 3 (H3 C)2 CH). C34 H44 N6 O11 (712.7): C 57.29, H 6.22, N 11.79; found: C 57.18, H 6.12, N 12.01. N2 ,2 ,3 ,5 -Tetrabenzoyl-O6 -NPE-G (S.28): yield: 1.23 g (71%). m.p. 168◦ C. Rf (CHCl3 /MeOH 19:1) 0.91. UV: 230 (4.77), 270 (4.55). 1 H NMR (DMSO-d6 ): 10.89 (s, 1H, H-N); 8.48 (s, 1H, H-C8); 8.19 (d, 2H, H o to NO2 ); 7.96-7.86 (m, 8H, arom. H); 7.68-7.55 (m, 6H, arom. H); 7.51-7.40 (m, 8H, arom. H, H m to NO2 ); 6.57 (d, 1H, HC1 ); 6.50 (m, 1H, H-C2 ); 6.39 (m, 1H, H-C3 ); 4.91-4.71 (m, 5H, H-C4 ,5 , OCH2 CH2 ); 3.34 (t, 2H, OCH2 CH2 ). 1 H NMR (CDCl3 ): 8.93 (s, 1H, H-N); 8.18 (d, 2H, H o to NO2 ); 8.05-7.88 (m, 9H, H-C8, 8 arom. H); 7.59-7.26 (m, 14H, arom. H, H m to NO2 ); 6.76 (d, 1H, H-C1 ); 6.43 (m. 1H, H-C2 ); 6.28 (m, 1H, H-C3 ); 4.97-4.75 (m, 5H, H-C4 ,5 , OCH2 CH2 ); 3.36 (t, 2H, OCH2 CH2 ). C46 H36 N6 O11 × H2 O (866.8): C 63.74, H 4.42, N 9.70; found: C 63.52, H 4.12, N 9.83.
Remove 3 ,5 -hydroxyl protection 6. Dissolve 1 mmol S.26 (0.626 g), S.27 (0.713 g), or S.28 (0.867 g) in 20 mL of 1,2-dioxane. Add 20 mL conc. aqueous ammonia or saturated methanolic ammonia at room temperature and stir for 1 to 2 days. Check progress of reaction by TLC. 7. Evaporate to dryness and recrystallize the residue from MeOH or EtOH. 8. Prepare an analytical sample by chromatography on a preparative silica gel plate (20 × 20 × 0.2–cm) and develop with 4:1 (v/v) CHCl3 /MeOH. 9. Cut out the main band, elute with 1:1 (v/v) CHCl3 /MeOH, evaporate to dryness, and dry in a 50◦ C oven. N2 -Isobutyryl-O6 -NPE-2 -dG (S.31): yield: 0.414 g (85%). Rf (CHCl3 /MeOH 9:1) 0.13. UV: 218 (4.44), 269 (4.42). 1 H NMR (DMSO-d6 ): 9.45 (s, 1H, H-N); 8.16 (d, 2H, H o to NO2 ); 7.64 (d, 2H, H m to NO2 ); 7.57 (s, 1H, H-C8); 6.31 (m, 1H, H-C1 ); 4.66 (t, 2H, OCH2 CH2 ); 4.47 (d, 1H, HO-C3 ); 4.06 (t, 1H, HO-C5 ); 3.56 (m, 1H, H-C3 ); 3.20 (t, 2H, OCH2 CH2 ); 3.00 (m, 1H, H-C4 ); 2.74-2.66 (m, 2H, H-C5 ); 2.01 (m, 1H, H-C2 ); 1.85 (m, 1H, H-C2 ); 1.49-1.39 (m, 1H, H-CMe2 ); 0.24 (d, 6H, (H3 C)2 CH). C22 H26 N6 O7 (486.5): C 54.23, H 5.39, N 17.28; found: C 54.09, H 5.31, N 17.18.
Protection of Nucleosides for Oligonucleotide Synthesis
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N2 -Isobutyryl-O6 -NPE-G (S.32): yield: 0.382 g (76%). Rf (CHCl3 /MeOH 9:1) 0.39. UV: 228 (4.43), 268 (4.40). 1 H NMR (DMSO-d6 ): 10.45 (s, 1H, H-N); 8.45 (s, 1H, H-C8); 8.18 (d, 2H, H o to NO2 ); 7.66 (d, 2H, H m to NO2 ); 5.89 (m, 1H, H-C1 ); 5.51 (d, 1H, HO-C2 ); 5.21 (d, 1H, HO-C3 ); 4.99 (t, 1H, HO-C5 ); 4.79 (t, 2H, OCH2 CH2 ); 4.55 (m, 1H, H-C2 ); 4.16 (m, 1H, H-C3 ); 3.92 (m, 1H, H-C4 ); 3.60 (m, 2H, H-C5 ); 3.32 (t, 2H, OCH2 CH2 ); 2.84 (m, 1H, H-CMe2 ); 1.09 (d, 6H, (H3 C)2 CH). C22 H26 N6 O8 (502.5): C 52.59, H 5.22, N 16.73; found: C 52.64, H 5.14, N 16.60. N2 -Benzoyl-O6 -NPE-G (S.33): yield: 0.311 g (57%). m.p. 194◦ C. Rf (CHCl3 /MeOH 9:1) 0.41. UV: 230 (4.31), 271 (4.49). 1 H NMR (DMSO-d6 ): 10.93 (s, 1H, H-N); 8.51 (s, 1H, H-C8); 8.18 (d, 2H, H o to NO2 ); 7.95 (d, 2H, arom. H); 7.57 (d, 2H, H m to NO2 ); 7.51 (m, 3H, arom. H); 5.93 (m, 1H, H-C1 ); 5.50 (d, 1H, HO-C2 ); 5.22 (d, 1H, HO-C3 ); 5.00 (t, 1H, HO-C5 ); 4.77 (t, 2H, OCH2 CH2 ); 4.60 (m, 1H, H-C2 ); 4.17 (m, 1H, H-C3 ); 3.92 (m, 1H, H-C4 ); 3.70-3.52 (m, 2H, H-C5 ); 3.33 (t, 2H, OCH2 CH2 ). C25 H24 N6 O8 ·0.5 H2 O (554.5): C 55.04, H 4.62, N 15.41; found: C 54.99, H 4.49, N 15.24.
Remove N2 protection 10. Suspend 2 mmol S.31 (0.97 g), S.32 (1.0 g), or S.33 (1.09 g) in 75 mL MeOH. Add 75 mL conc. aqueous ammonia and stir 5 to 6 days at room temperature. Check progress of deacylation by TLC. 11. Evaporate to dryness on a rotatory evaporator. 12. Recrystallize the residue from H2 O or purify by silica gel column chromatography (60 × 2.5–cm) using 10:8:3 (v/v/v) toluene/EtOAc/MeOH. 13. Collect the product fractions and evaporate to dryness. 14. Dry under vacuum at 40◦ C in a drying pistol. O6 -NPE-2 -dG (S.34): yield: 0.66 g (76%). m.p. amorphous solid. Rf (CHCl3 /MeOH 9:1) 0.28. UV: 250 (4.16), 279 (4.25). 1 H NMR (DMSO-d6 ): 8.20 (d, 2H, H o to NO2 ); 8.18 (s, 1H, H-C8); 7.63 (d, 2H, H m to NO2 ); 6.48 (s, 2H, NH2 ); 6.21 (d, 1H, H-C1 ); 5.28 (d, 1H, HO-C3 ); 5.00 (t, 1H, HO-C5 ); 4.65 (t, 2H, OCH2 CH2 ); 4.34 (dd, 1H, H-C3 ); 3.81 (m, 1H, H-C4 ); 3.53 (m, 2H, H-C5 ); 3.25 (t, 2H, OCH2 CH2 ); 2.57 (m, 1H, H-C2 ); 2.19 (m, 1H, H-C2 ). C18 H20 N6 O6 ·H2 O (434.4): C 49.76, H 5.10, N 19.34; found: C 50.12, H 4.65, N 19.15. O6 -NPE-G (S.35): yield: 0.51 g (59%). m.p. > 320◦ C. Rf (CHCl3 /MeOH 9:1) 0.48. UV: 251 (4.17), 278 (4.26). 1 H NMR (DMSO-d6 ): 8.19 (d, 2H, H o to NO2 ); 8.10 (s, 1H, H-C8); 7.64 (d, 2H, H m to NO2 ); 6.49 (s, 2H, NH2 ); 5.77 (d, 1H, H-C1 ); 5.41 (d, 1H, HO-C2 ); 5.15 (d, 1H, HO-C3 ); 5.10 (t, 1H, HO-C5 ); 4.67 (t, 2H, OCH2 CH2 ); 4.46 (dd, 1H, H-C2 ); 4.09 (dd, 1H, H-C3 ); 3.89 (m, 1H, H-C4 ); 3.56 (m, 2H, H-C5 ); 3.25 (t, 2H, OCH2 CH2 ). C18 H20 N6 O7 (432.4): C 50.00, H 4.66, N 19.44; found: C 50.18, H 4.55, N 19.72. BASIC PROTOCOL 5
N2 -p-NPEOC PROTECTION OF 2 -DEOXYGUANOSINE The N2 -NPEOC-protected nucleoside S.40 is synthesized from 2 -deoxyguanosine (S.36) by the transient protection method (Ti et al., 1982) using trimethylsilyl chloride for intermediary blocking of the sugar hydroxyl groups, followed by acylation with 2-(4nitrophenyl)ethyl chloroformate and desilylation by aqueous sodium bicarbonate.
Materials
Universal NPE and NPEOC Protecting Groups
2 -Deoxyguanosine (S.36; Sigma-Aldrich, Pharma Waldhof) Pyridine (anhydrous) Trimethylsilyl chloride (Sigma-Aldrich) 2-(4-Nitrophenyl)ethyl chloroformate (see Basic Protocol 3) Chloroform (CHCl3 ) Methanol (MeOH)
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10% aqueous sodium bicabonate solution (NaHCO3 ) Diethyl ether Silica-coated TLC plate with fluorescent indicator Rotatory evaporator Drying pistol Vacuum pump 1. Coevaporate 1 mmol (0.267 g) 2 -dG (S.36) three times with 3 mL dry pyridine. Suspend the residue in 20 mL dry pyridine. 2. Add 0.64 mL (5 mmol) trimethylsilyl chloride (0.543 g) with stirring at room temperature. Continue stirring 30 min. Check progess of reaction by TLC. 3. Cool the mixture consisting of S.37 in an ice-bath and then slowly add 2 mmol 2-(4nitrophenyl)ethyl chloroformate (0.46 g) dissolved in 4 mL CHCl3 dropwise with stirring. Continue stirring 1 hr at 0◦ to 5◦ C, then stir for 2 days at room temperature. 4. Add 10 mL MeOH to the reaction mixture and stir for 5 hr at room temperature. 5. Evaporate to dryness and then treat the residue overnight with 20 mL 10% aqueous NaHCO3 under stirring. 6. Collect the resulting colorless precipitate, S.40. Wash with 20 mL diethyl ether and dry in a drying pistol at 50◦ C under vacuum. N2 -NPEOC-2 -dG (S.40): yield: 0.23 g (50%). m.p. 185o -190◦ C. Rf (CHCl3 /MeOH 4:1) 0.36. UV: 251 (sh 4.37), 258 (4.41), 272 (sh 4.26). 1 H NMR (DMSO-d6 ): 11.46, 11.32 (2s, 2H, NH); 8.21 (s, 1H, H-C8); 8.17 (d, 2H, o to NO2 ); 7.63 (d, 2H, m to NO2 ); 6.21 (t, 1H, H-C1 ); 5.32 (d, 1H, HO-C3 ); 4.95 (t, 1H, HO-C5 ); 4.48 (t, 2H, OCH2 CH2 ); 4.36 (m, 1H, H-C3 ); 3.82 (m, 1H, H-C4 ); 3.59-3.45 (m, 2H, H-C5 ); 3.14 (t, 2H, OCH2 CH2 ); 2.99 (m, 1H; H-C2 ); 2.55 (m, 1H, H-C2 ). C18 H20 N6 O8 (460.4): C 49.57, H 4.38, N 18.25; found: C 49.21, H 4.19, N 18.20.
O6 -p-NPE AND N2 -p-NPEOC PROTECTION OF 2 -DEOXYGUANOSINE AND GUANOSINE
BASIC PROTOCOL 6
An interesting combination in the universal blocking group strategy is seen in the application of N2 -NPEOC protection of guanosine derivatives in combination with O6 NPE protection (Lang et al., 1999). The preparation of compounds S.41 and S.42 is achieved in a one-pot reaction starting from 3 ,5 -di-O-acetyl-2 -deoxyguanosine (S.38) or 2 ,3 ,5 -tri-O-acetylguanosine (S.39), respectively. These are first subjected to a Mitsunobu reaction with 2-(4-nitrophenyl)ethanol to give the O6 -NPE-protected S.29 and S.30, respectively. Subsequent 2-(4-nitrophenyl)ethoxycarbonylation and final treatment with ammonia afford S.41 and S.42 in good overall yields.
Materials 3 ,5 -Di-O-acetyl-2 -deoxyguanosine (S.38; Schaller et al., 1963) or 2 ,3 ,5 -tri-O-acetylguanosine (S.39; Pharma Waldhof) Triphenylphosphine 2-(4-Nitrophenyl)ethanol (Sigma-Aldrich) Dioxane (anhydrous) Diethyl azodicarboxylate (Sigma-Aldrich) Pyridine (anhydrous) 2-(4-Nitrophenyl)ethyl chloroformate (see Basic Protocol 3) Chloroform (CHCl3 ; anhydrous) Sodium sulfate (Na2 SO4 ) Toluene
Protection of Nucleosides for Oligonucleotide Synthesis
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Silica gel (Merck 60) Methylenechloride (CH2 Cl2 ) Methanol (MeOH) Conc. aqueous ammonia 100-mL round-bottom flasks Rotary evaporator Chromatography column (20 × 2.5–cm) Silica-coated TLC plate with fluorescent indicator Drying pistol, 50◦ C Perform three-step synthesis 1. In a 100-mL round-bottom flask, prepare a solution of 2 mmol (0.69 g) 3 ,5 -di-Oacetyl-2 -dG (S.38) or 2 mmol (0.82 g) 2 ,3 ,5 -tri-O-acetyl-G (S.39) with 3.2 mmol triphenylphosphine (0.84 g) and 3 mmol 2-(4-nitrophenyl)ethanol (0.50 g) in 40 mL dry dioxane. Stir for 10 min at room temperature. 2. Add 3.2 mmol diethyl azodicarboxylate (0.558 g) and stir for 1 hr to obtain a clear solution. 3. Evaporate under vacuum in a rotatory evaporator and coevaporate with 40 mL dry pyridine. 4. Dissolve the residue in 10 mL dry pyridine and cool 30 min in an ice bath. 5. While stirring, add a solution of 6 mmol 2-(4-nitrophenyl)ethyl chloroformate (1.38 g) in 10 mL dry CHCl3 slowly dropwise. Stir 1 hr at 0◦ to 5◦ C and then 3 hr at room temperature. 6. Dilute with 100 mL H2 O and extract four times with 50 mL CHCl3 . 7. Combine the organic layers and dry over Na2 SO4 . Filter off the drying agent through filter paper, evaporate the filtrate, and coevaporate two times with 10 mL toluene. 8. Purify the resulting residue by silica gel column chromatography (20 × 2.5–cm), eluting first with CH2 Cl2 , followed by CHCl3 . 9. Collect the product fractions (check by TLC), evaporate again, and treat the residue in 25 mL dioxane and 25 mL MeOH with 25 mL conc. aqueous ammonia, keeping the reaction mixture for 24 hr at 7◦ C. 10. Evaporate under vacuum and recrystallize the resulting solid from MeOH. Dry under vacuum in the drying pistol at 50◦ C. N2 -NPEOC-O6 -NPE-2 -dG (S.41): yield: 0.81 g (66%). m.p. 179o -182◦ C. Rf (CHCl3 /MeOH 9:1) 0.53. UV: 216 (4.63), 269 (4.54). 1 H NMR (DMSO-d6 ): 10.33 (s, 1H, NH); 8.40 (s, 1H, H-C8); 8.17 (d, 4H, o to NO2 ); 7.64 (d, 2H, m to NO2 ); 7.61 (d, 2H, H m to NO2 ); 6.30 (t, 1H, H-C1 ); 5.32 (d, 1H, HO-C3 ); 4.89 (t, 1H, HO-C5 ); 4.73 (t, 2H, OCH2 CH2 ); 4.41 (m, 1H, H-C3 ); 4.37 (t, 2H, OCH2 CH2 ); 3.83 (m, 1H, H-C4 ); 3.67-3.42 (m, 2H, H-C5 ); 3.30 (t, 2H, OCH2 CH2 ); 3.11 (t, 2H, OCH2 CH2 ); 2.71 (m, 1H; H-C2 ); 2.25 (m, 1H, H-C2 ). C27 H27 N7 O10 × 0.5 H2 O (618.5): C 52.43, H 4.56, N 15.85; found: C 52.32, H 4.67, N 15.63.
Universal NPE and NPEOC Protecting Groups
N2 -NPEOC-O6 -NPE-G (S.42): yield: 0.834 g (65%). m.p. 172o -174◦ C (decomp.) Rf (CHCl3 /MeOH 9:1) 0.50. UV (MeOH): 216 (4.62), 269 (4.53). 1 H NMR (DMSO-d6 ): 10.37 (s, 1H, NH); 8.42 (s, 1H, H-C8); 8.17 (d, 4H, o to NO2 ); 7.64 (d, 2H, m to NO2 ); 7.61 (d, 2H, m to NO2 ); 5.89 (t, 1H, H-C1 ); 5.49 (d, 1H, HO-C2 ); 5.20 (d, 1H, HOC3 ); 4.98 (t, 1H, HO-C5 ); 4.77 (t, 2H, OCH2 CH2 ); 4.60 (m, 1H, H-C2 ); 4.38 (t, 2H, OCH2 CH2 ); 4.18 (m, 1H, H-C3 ); 3.92 (m, 1H, H-C4 ); 3.72-3.47 (m, 2H, H-C5 ); 3.30 (t, 2H, OCH2 CH2 ); 3.10 (t, 2H, OCH2 CH2 ). C27 H27 N7 O11 ·H2 O (643.5): C 50.38, H 4.54, N 15.23; found: C 50.40, H 4.38, N 15.25.
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PREPARATION OF NPE- AND NPEOC-PROTECTED 2 -DEOXYRIBONUCLEOSIDE 3 -O-PHOSPHORAMIDITES
BASIC PROTOCOL 7
The most common and highly successful oligo-2 -deoxyribonucleotide synthesis is based on the phosphoramidite approach initiated and developed by Caruthers (Caruthers et al., 1982; UNIT 3.3). The general synthetic strategy involves adding mononucleotides sequentially to a nucleoside that is covalently attached to an insoluble solid support. The monomers of choice are 5 -O-dimethoxytrityl-2 -deoxyribonucleoside-3 -O(2-cyanoethyl-N,N-diisopropylphosphoramidites) in which the nucleobases are protected by various acyl groups. Using the universal NPE/NPEOC approach, the sugar moiety can be substituted as in S.48 to S.52 (Fig. 2.13.5), or can be modified by the 3 -O-NPE-N,Ndiisopropylphosphoramidite functionality as in S.53 to S.56 (Lang et al., 1999). In this protocol, dimethoxytritylation of the unprotected sugar moiety is performed using a standard procedure, leading selectively to the corresponding 5 -O-dimethoxytrityl derivatives S.43 to S.47. Phosphitylation to give phosphoramidites S.48 to S.52 is performed by standard methods using 2-cyanoethyl-N,N,N ,N -tetraisopropylphosphordiamidite (Barone et al., 1984; Sinha et al., 1984; Zon et al., 1985) and 1H-tetrazole in an aprotic solvent. Phosphitylation to give S.53 to S.56 can be performed with bis-[N,N-diisopropylamino2-(4-nitrophenyl)ethoxyphosphane] (Sobol et al., 1995) and 1H-tetrazole or with chloro-N,N-diisopropyl-2-(4-nitrophenyl)ethoxyphosphane (Sobol et al., 1995) and triethylamine in acid-free CH2 Cl2 under argon atmosphere.
Figure 2.13.5
NPE- and NPEOC-protected 2 -deoxyribonucleoside-3 -O-phosphoramidites.
Protection of Nucleosides for Oligonucleotide Synthesis
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Materials Protected nucleoside: O4 -NPE-thymidine (S.5) N4 -NPEOC-2 -deoxycytidine (S.10) N6 -NPEOC-2 -deoxyadenosine (S.19) N2 -NPEOC-2 -deoxyguanosine (S.40) N2 -NPEOC-O6 -NPE-2 -deoxyguanosine (S.41) Pyridine (anhydrous) 4,4 -Dimethoxytrityl chloride (Sigma-Aldrich) Methanol (MeOH) Methylenechloride (CH2 Cl2 , anhydrous and acid-free) Sodium sulfate (Na2 SO4 ) Toluene Silica-gel (Merck 60, 63 to 200 mesh) Chloroform (CHCl3 ) Triethylamine (Et3 N) 2-Cyanoethyl-N,N,N ,N -tetraisopropylphosphordiamidite (Sigma-Aldrich) 1H-Tetrazole (Sigma-Aldrich) Argon pressure bottle Saturated NaHCO3 solution Ethyl acetate (EtOAc) Trichlorophosphane (PCl3 , freshly distilled) Diethyl ether (anhydrous) Nitrogen pipeline 2-(4-Nitrophenyl)ethanol (Sigma-Aldrich) N,N-Diisopropylamine Isopropanol N,N-Diisopropyltrimethylsilylamine (Sigma-Aldrich) 50- and 100-mL round-bottom flasks Rotatory evaporator Chromatography columns (40 × 2.5–cm, 50 × 2.5–cm, and 60 × 2.5–cm) 100- and 250-mL three-neck round-bottom flasks Condenser with drying tube Gas inlet tube G4 glass filter High-vacuum pump 100-mL two-neck round-bottom flasks Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Perform 5 -O-dimethoxytritylation 1. Dissolve in a 100-mL round-bottom flask 5 mmol of dry nucleoside O4 -NPE-T (S.5) (1.96 g), N4 -NPEOC-dC (S.10) (2.15 g), N6 -NPEOC-dA (S.19) (2.31 g), N2 NPEOC-dG (S.40) (2.30 g), or N2 -NPEOC-O6 -NPE-dG (S.41) (3.09 g) in 25 mL pyridine. Add 6 mmol of 4,4 -dimethoxytrityl chloride (2.0 g) and stir 2 to 20 hr at room temperature. Monitor by TLC (APPENDIX 3D). 2. Add 10 mL MeOH to the mixture after completion of the reaction and evaporate to 1/4 of its volume on a rotary evaporator. Universal NPE and NPEOC Protecting Groups
2.13.18 Supplement 30
3. Dilute with 25 mL CH2 Cl2 , wash two times with 10 mL H2 O, and separate the organic layer. 4. Dry over Na2 SO4 , filter through filter paper, evaporate the filtrate, and then coevaporate with toluene. Current Protocols in Nucleic Acid Chemistry
5. Purify the residue by silica gel column or flash chromatography (APPENDIX 3E) using a 50 × 2.5–cm column and the appropriate solvent mixture: S.43: flash chromatography (50:1 v/v CHCl3 /MeOH) S.44: flash chromatography (CH2 Cl2 to 95:5 CH2 Cl2 /MeOH) S.45: flash chromatography (25:1 v/v CHCl3 /MeOH) S.46: column chromatography (100:0.2 v/v CHCl3 /Et3 N to 90:10:02 CHCl3 / MeOH/Et3 N) e. S.47: column chromatography (100:0.2 v/v CHCl3 /Et3 N to 99:1:02 v/v/v CHCl3 / MeOH/Et3 N)
a. b. c. d.
6. Collect the product fractions and evaporate under vacuum to give a colorless foam. 7. Dry under high vacuum at room temperature. 5 -O-DMTr-O4 -NPE-T (S.43): yield: 3.46 g (95%). Rf (CHCl3 /MeOH 95:5) 0.82. UV: 232 (sh 4.40), 275 (4.26), 280 (sh 4.25). 1 H NMR (CDCl3 ): 8.14 (d, 2H, o to NO2 ); 7.92 (s, 1H, H-C6); 7.30 (m, 9H, arom. H); 6.81 (d, 4H, o to MeO); 6.40 (pt, 1H, H-C1 ); 4.61 (m, 3H, H-C3 , OCH2 CH2 ); 4.16 (m, 1H, H-C4 ); 3.77 (s, 6H, MeO); 3.73 (d, 1H, HO-C3 ); 3.41 (m, 2H, H-C5 ); 3.15 (t, 2H, OCH2 CH2 ); 2.66 (m, 1H, H-C2 ); 2.23 (m, 1H, H-C2 ); 1.46 (s, 3H, H3 C-C5). C39 H38 N3 O9 ·2H2 O (728.7): C 64.28, H 5.81, N 5.76; found: C 64.30, H 5.46, N 5.81. 5 -O-DMTr-N4 -NPEOC-2 -dC (S.44): yield: 3.11 g (86%). Rf (CHCl3 /MeOH 95:5) 0.36. UV: 235 (4.56), 275 (4.25), 280 (sh 4.25). 1 H NMR (CDCl3 ): 8.25-8.15 (m, 4H, 2H o to NO2 , H-N, H-C6); 7.42-7.19 (m, 11H, arom. H); 6.95 (m, 1H, H-C5): 6.86-6.81 (m, 4H, o to MeO); 6.29 (m, 1H, H-C1 ); 4.51 (m, 3H, H-C3 ); 4.41 (t, 2H, OCH2 CH2 ); 4.15 (m, 1H, H-C4 ); 3.78 (s, 6H, MeO); 3.60-3.36 (m, 3H, HO-C3 , H-C5 ); 3.09 (t, 2H, OCH2 CH2 ); 2.75 (m, 1H, H-C2 ); 2.27-2.16 (m, 1H, H-C2 ). C39 H38 N4 O10 (722.8): C 64.81, H 5.30, N 7.75; found: C 64.89, H 5.56, N 7.67. 5 -O-DMTr-N6 -NPEOC-2 -dA (S.45): yield: 3.17 g (84%). Rf (CHCl3 /MeOH 95:5) 0.34. UV: 236 (4.43), 268 (4.47), 276 (sh 4.41). 1 H NMR (CDCl3 ): 8.66 (s, 1H, H-C8); 8.36 (bs, 1H, H-N); 8.16 (d, 2H, o to NO2 ); 8.11 (s, 1H, H-C2); 7.44-7.15 (m, 9H, arom. H); 6.80 (d, 4H, o to MeO); 6.44 (pt, 1H, H-C1 ); 4.71 (m, 3H, H-C3 ); 4.53 (t, 2H, OCH2 CH2 ); 4.24 (m, 1H, H-C4 ); 3.77 (s, 6H, MeO); 3.73 (d, 1H, HO-C3 ); 3.52-3.40 (m, 2H, H-C5 ); 3.14 (t, 2H, OCH2 CH2 ); 2.91-2.51 (m, 2H, H-C2 ,2 ). C40 H38 N6 O9 ·0.5H2 O (755.8): C 63.57, H 5.20, N 11.12; found: C 63.62, H 5.21, N 10.95. 5 -O-DMTr-N2 -NPEOC-2 -dG (S.46): yield: 3.01 g (79%). Rf (CHCl3 /MeOH 95:5) 0.40. UV: 237 (4.46), 249 (4.42), 258 (4.42), 272 (sh 4.38). 1 H NMR (CDCl3 ): 11.27 (s, 1H, NH); 9.76 (s, 1H, NH); 8.00 (d, 2H, o to NO2 ); 7.75 (s, 1H, H-C8); 7.32-7.03 (m, 9H, arom. H); 6.67 (d, 4H, o to MeO); 6.15 (pt, 1H, H-C1 ); 4.80 (m, 3H, H-C3 ); 4.44 (m, 3H, HO-C3 , OCH2 CH2 ); 4.15 (m, 1H, H-C4 ); 3.65 (s, 6H, MeO); 3.30 (m, 2H, H-C5 ); 3.01 (t, 2H, OCH2 CH2 ); 2.60-2.46 (m, 2H, H-C2 ,2 ). C40 H38 N6 O10 (762.8): C 62.99, H 5.02, N 11.02; found: C 62.43, H 5.14, N 10.95. 5 -O-DMTr-N2 -NPEOC-O6 -NPE-2 -dG (S.47): yield: 3.88 g (85%). Rf (CHCl3 /MeOH 95:5) 0.43. UV: 235 (4.47), 268 (4.55). 1 H NMR (CDCl3 ): 8.15-800 (m, 3H, NH, o to NO2 ); 7.96 (s, 1H, H-C8); 7.50-7.13 (m, 9H, arom. H); 6.77 (d, 4H, o to MeO); 6.48 (m, 1H, H-C1 ); 4.81-4.74 (m, 3H, H-C3 , OCH2 CH2 ); 4.45 (t, 2H, OCH2 CH2 ); 4.16 (m, 1H, H-C4 ); 3.75 (s, 6H, MeO); 3.56-3.25 (m, 4H, H-C5 , OCH2 CH2 ); 3.09-3.03 (t, 3H, HO-C3 , OCH2 CH2 ); 2.75 (m, 1H, H-C2 ); 2.55 (m, 1H, H-C2 ). C48 H45 N7 O12 (911.9): C 63.22, H 4.97, N 10.75; found: C 63.08, H 5.14, N 10.67.
Prepare 3 -O-(2-cyanoethyl-N,N-diisopropyl)phosphoramidites 8. Dissolve in a 50-mL round-bottom flask 1 mmol S.43 (0.729 g), S.44 (0.719 g), S.45 (0.756 g), S.46 (0.763 g), or S.47 (0.912 g) and 1.5 mmol of 2-cyanoethylN,N,N ,N -tetraisopropylphosphordiamidite in 15 mL acid-free CH2 Cl2 . Add 0.5 mmol of 1H-tetrazole (35 mg) and stir 2.5 hr at room temperature under argon atmosphere.
Protection of Nucleosides for Oligonucleotide Synthesis
2.13.19 Current Protocols in Nucleic Acid Chemistry
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9. Dilute the solution with 100 mL CH2 Cl2 , extract two times with 50 mL saturated NaHCO3 solution, and reextract the aqueous phase with CH2 Cl2 . 10. Combine the organic phases, dry over Na2 SO4 , filter through filter paper, and evaporate under vacuum. 11. Purify the resulting residue by column chromatography or flash chromatography using a 40 × 2.5–cm column and the appropriate solvent system: a. b. c. d.
S.48 flash chromatography (1:1 v/v toluene/EtOAc) S.49 flash chromatography (2:1 and 1:1 v/v toluene/EtOAc) S.50 flash chromatography (3:7 and 1:1 v/v toluene/EtOAc) S.51 column chromatography (97.5:2:0.5 to 91.5:8:0.5 v/v/v CH2 Cl2 /MeOH/ Et3 N) e. S.52 column chromatography (99:1 v/v EtOAc/Et3 N) 12. Evaporate under vacuum to give a diastereoisomeric mixture as a solid foam. Dry under high vacuum at room temperature. 5 -O-DMTr-O4 -NPE-T-3 -O-(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.48): yield: 0.662 g (74%). Rf (toluene/EtOAc 1:1): 0.42. UV (MeOH): 232 (sh 4.39), 275 (4.25), 280 (sh 4.24). 1 H NMR (CDCl3 ): 8.16 (d, 2H o to NO2 ); 7.90 (2s, 1H, H-C6); 7.30 (m, 11H, arom. H); 6.82 (dd, 4H o to MeO); 6.38 (m, 1H, H-C1 ); 4.63 (m, 3H, H-C3 , OCH2 CH2 ); 4.15 (m, 1H, H-C4 ); 3.78 (m, 7H, 2 MeO, OCH2 CH2 CN); 3.54 (m, 4H, 2 Me2 CHN), OCH2 CH2 CN, H-C5 ); 3.32 (m, 1H, H-C5 ); 3.16 (t, OCH2 CH2 ); 2.64 (m, 2H, OCH2 CH2 CN); 2.32 (m, 2H, H-C2 ,2 ); 1.41 (2s, 3H, H3 C-C5); 1.11 (m. 12H, 2 MeCH). 31 P NMR (CDCl3 ): 149.71, 149.16. C48 H56 N5 O12 P (894.0): C 64.49, H 6.31, N 7.83; found: C 64.45, H 6.54, N 7.31. 5 -O-DMTr-N4 -NPEOC-2 -dC-3 -O-(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.49): yield: 0.785 g (85%). Rf (n-hexane/EtOAc/Et3 N 3:7:1): 0.24. UV (MeOH): 236 (4.65), 274 (4.42), 280 (sh 4.40). 1 H NMR (CDCl3 ): 8.20 (m, 2H o to NO2 ); 8.20 (2s, 1H, H-C6); 7.51-7.12 (m, 11H, arom. H); 6.74 (dd, 4H o to MeO); 6.35 (m, 2H, H-C1 , H-C5); 4.80 (m, 1H, H-C3 ); 4.80 (t, 2H, OCH2 CH2 ); 4.43 (t, 2H, OCH2 CH2 ); 4.21 (m, 1H, H-C4 ); 3.90 (m, 2H, OCH2 CH2 ); 3.73 (2s, 6H, 2 MeO); 3.70-3.47 (m, 4H, 2 Me2 CHN, OCH2 CH2 CN); 3.32 (m, 2H, H-C5 ); 3.08 (m, OCH2 CH2 ); 2.92 (m, 1H, H-C2 ); 2.75 (m, 1H, H-C2 ); 2.64 (m, 2H, OCH2 CH2 CN); 1.10-0.91 (m, 12H, 2 MeCH). 31 P NMR (CDCl3 ): 150.0, 149.9. C48 H55 N6 O11 P (923.0): C 62.46, H 6.00, N 9.10; found: C 62.46, H 6.08, N 8.83. 5 -O-DMTr-N6 -NPEOC-2 -dA-3 -O-(2-cyanoethyl-N,N-diisopropylphosphor-amidite) (S.50): yield: 0.871 g (92%). Rf (n-hexane/EtOAc/Et3 N 3:7:1): 0.25. UV (MeOH): 236 (4.45), 266 (4.47), 275 (sh 4.41). 1 H NMR (CDCl3 ): 8.68 (s, 1H, H-C8); 8.25-8.15 (m, 2H o to NO2 ); 8.01 (m, 1H, NH); 7.35 (d, 2H m to NO2 ); 7.35-7.18 (m, 11H, arom. H); 6.74 (dd, 4H o to MeO); 6.45 (m, 1H, H-C1 ); 4.74 (m, 1H, H-C3 ); 4.51 (t, 2H, OCH2 CH2 ); 4.43 (t, 2H, OCH2 CH2 ); 4.29 (m, 1H, H-C4 ); 3.75 (m, 8H, OCH2 CH2, 2 MeO); 3.52-3.43 (m, 4H, 2 Me2 CHN)); 3.31 (m, 2H, H-C5 ); 3.13 (m, OCH2 CH2 ); 2.82 (m, 1H, H-C2 ); 2.60 (m, 1H, H-C2 ); 2.59, 2.44 (2t, 2H, OCH2 CH2 CN); 1.27-1.08 (m. 12H, 2 MeCH). 31 P NMR (CDCl3 ): 149.6, 149.5. C49 H55 N8 O10 P (947.0): C 62.14, H 5.85, N 11.83; found: C 61.93, H 6.06, N 11.32.
Universal NPE and NPEOC Protecting Groups
5 -O-DMTr-N2 -NPEOC-2 -dG-3 -O-(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.51): yield: 0.568 g (60%). Rf (n-hexane/EtOAc/Et3 N 95:5:2): 0.52. UV (MeOH): 237 (4.46), 249 (4.42), 258 (4.42), 272 (4.37), 281 (sh 4.31). 1 H NMR (CDCl3 ): 8.15-8.11 (m, 2H o to NO2 ); 7.76, 7.72 (2s, 1H, H-C8); 8.01 (m, 1H, NH); 7.35 (d, 2H m to NO2 ); 7.38-7.18 (m, 10H, NH, arom. H); 6.74 (d, 4H o to MeO); 6.16 (m, 1H, H-C1 ); 4.74-4.61 (m, 1H, H-C3 ); 4.44 (t, 2H, OCH2 CH2 ); 4.24 (m, 1H, H-C4 ); 3.74, 3.73 (2s, 6H, 2 MeO); 4.14-3.45 (m, 4H, OCH2 CH2 CN, 2 Me2 CHN)); 3.34-3.20 (m, 2H, H-C5 ); 3.09 (t, 2H, OCH2 CH2 ); 2.81-2.32 (m, 4H, H-C2 ,2 , OCH2 CH2 CN); 1.30-1.06 (m, 12H, 2 MeCH). 31 P NMR (CDCl3 ): 149.48, 148.89. C49 H55 N8 O11 P (963.0): C 61.12, H 5.76, N 11.64; found: C 60.80, H 5.90, N 11.40.
2.13.20 Supplement 30
Current Protocols in Nucleic Acid Chemistry
5 -O-DMTr-N2 -NPEOC-O6 -NPE-2 -dG-3 -O-(2-cyanoethyl-N,N-diisopropyl-phosphoramidite) (S.52): yield: 0.956 g (86%). Rf (EtOAc/Et3 N 99:1): 0.81, 0.72. UV (MeOH): 236 (4.48), 269 (4.55). 1 H NMR (CDCl3 ): 8.17-8.11 (m, 4H o to NO2 ); 7.96, 7.95 (2s, 1H, H-C8); 7.51-7.13 (m, 12H, H m to NO2 , NH, arom. H); 6.74 (d, 4H o to MeO); 6.36 (m, 1H, H-C(1 )); 4.82-4.70 (m, 3H, H-C3 , OCH2 CH2 ); 4.42 (t, 2H, OCH2 CH2 ); 4.24 (m, 1H, H-C4 ); 3.74 (2s, 6H, 2 MeO); 3.86-3.48 (m, 4H, OCH2 CH2 CN, 2 Me2 CHN); 3.37-3.26 (m, 4H, H-C5 , OCH2 CH2 ); 3.09 (t, 2H, OCH2 CH2 ); 2.87-2.55 (m, 2H, H-C2 ,2 ); 2.62, 2.41 (2t, 2H, OCH2 CH2 CN); 1.26-1.07 (m, 12H, 2 MeCH). 31 P NMR (CDCl3 ): 149.59, 140.38. C57 H62 N9 O13 P (1112.2): C 61.56, H 5.62, N 11.33; found: C 61.76, H 5.65, N 11.05.
Prepare phosphitylating reagent for NPE phosphoramidites For bis-(N,N-diisopropylamino)-2-(4-nitrophenyl)ethoxyphosphane 13a. Dissolve in a 100-mL round-bottom flask 28 mL (280 mmol) of freshly distilled PCl3 in 30 mL anhydrous diethyl ether under nitrogen atmosphere and cool down to −5◦ C. 14a. Add 8.35 g (50 mmol) 2-(4-nitrophenyl)ethanol in small portions over a period of 30 min, and then continue stirring 15 min at −5◦ C and finally 1.5 hr at room temperature. 15a. Evaporate under high vacuum and dissolve the resulting syrup in 200 mL anhydrous diethyl ether. 16a. Cool to −10◦ C, add slowly dropwise 64 mL (450 mmol) N,N-diisopropylamine under N2 atmosphere, and stir 15 min at −10◦ C and then 16 hr at room temperature. 17a. Filter off the precipitate of N,N-diisopropylamine hydrochloride through a G4 glass funnel and evaporate the filtrate under high vacuum to give 17.6 g (89%) of a yellowish syrup, which crystallizes on storage at −20◦ C. The product is pure enough for phosphitylation reactions. Bis-(N,N-diisopropylamino)-2-(4-nitrophenyl)ethoxyphosphane: 1 H NMR (CHCl3 ): 8.11 (d, 2H, H o to NO2 ); 7.38 (d, 2H, H m to NO2 ); 3.79 (q, 2H, P-OCH2 ); 3.513.36 (m, 2H, N-CHMe2 ); 2.97 (t, 2H, P-O-CH2 CH2 ); 1.09 (2d, 12H, N-CH(CH3 )2 ). 31 P NMR (CHCl3 ): 123.53.
For chloro-N,N-diisopropylamino-2-(4-nitrophenyl)ethoxyphosphane 13b. Dissolve in a 100-mL round-bottom flask 14 mL freshly distilled PCl3 in 40 mL anhydrous diethyl ether and cool under N2 atmosphere to −30◦ C in an isopropanol/dry ice bath. 14b. Under magnetic stirring, add 4.16 g (25 mmol) 2-(4-nitrophenyl)ethanol in small portions over a period of 45 min. 15b. Stir 1.5 hr at room temperature and evaporate under high vacuum. 16b. Cool the residue to 0◦ C and add dropwise under N2 atmosphere 4.33 g (25 mmol) N,N-diisopropyltrimethylsilylamine. Stir 30 min at 0◦ C and then 20 hr at room temperature. 17b. Evaporate under high vacuum to give 7.1 g (85%) of a yellowish syrup, which crystallizes on storage at −20◦ C. The product is pure enough for further use. Chloro-N,N-diisopropylamino-2-(4-nitrophenyl)ethoxyphosphane: 1 H NMR (CDCl3 ): 8.15 (m, 2H, H o to NO2 ); 7.41 (m, 2H, H m to NO2 ); 4.11 (m, 2H, P-OCH2 ); 3.71 (m, 2H, N-CHMe2 ); 3.10 (t, 2H, P-OCH2 CH2 ); 1.27-1.14 (2d, 12H, N-CH(CH3 )2 ). 31 P NMR (CH2 Cl2 ): 181.60.
Protection of Nucleosides for Oligonucleotide Synthesis
2.13.21 Current Protocols in Nucleic Acid Chemistry
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Prepare 3 -O-(2-(4-nitrophenyl)ethylphosphoramidites 18a. Under argon atmosphere, dissolve in a 50-mL round-bottom flask a mixture of 1 mmol tritylated nucleoside S.43 (0.729 g), S.44 (0.719 g), S.45 (0.756 g), or S.47 (0.912 g) and 1.5 mmol bis-(N,N-diisopropylamino)-2-(4-nitrophenyl) ethoxyphosphane (0.6 g) in 15 mL acid-free CH2 Cl2 . Add 0.5 mmol 1H-tetrazole (35 mg) and stir 2.5 hr at room temperature. Monitor by TLC. 18b. Alternatively, treat with 1.2 mmol chloro-N,N-diisopropylamino-2-(4nitrophenyl)ethoxyphosphane (0.416 g) and 3.8 mmol triethylamine (0.7 mL) in 15 mL acid-free CH2 Cl2 . Stir for 30 min at room temperature. Monitor by TLC. 19. Dilute the reaction mixture with 100 mL CH2 Cl2 , extract two times with 50 mL saturated NaHCO3 solution, and reextract the aqueous phase with CH2 Cl2 . 20. Combine the organic phases, dry over Na2 SO4 , filter, and evaporate. 21. Purify either by flash chromatography using a 60 × 2.5–cm column and the appropriate solvent system: a. b. c. d.
S.53 flash chromatography (99:1 v/v EtOAc/Et3 N) S.54 flash chromatography (96:4 EtOAc/Et3 N) S.55 flash chromatography (70:30:1 to 50:50:1 v/v/v toluene/EtOAc/Et3 N) S.56 flash chromatography (96:4 v/v EtOAc/Et3 N)
22. Evaporate under high vacuum to give a diastereoisomeric mixture as a solid foam. 5 -O-DMTr-O4 -NPE-T-3 -O-(2-(4-nitrophenyl)ethyl-N,N-diisopropylphosphoramidite) (S.53): yield: 0.871 g (88%). Rf (EtOAc): 0.77. UV (MeOH): 234 (sh 4.33), 275 (4.32), 281 (sh 4.30). 1 H NMR (CDCl3 ): 8.63 (m, 4H o to NO2 ); 7.61 (m, 1H, H-C6); 7.42-7.21 (m, 13H, arom. H); 6.76 (m, 4H o to MeO); 6.43 (m, 1H, H-C1 ); 4.64 (m, 3H, H-C3 , OCH2 CH2 ); 4.16 (m, 1H, H-C4 ); 3.79 (2s, 6H, 2 MeO); 3.93-3.62 (m, 4H, OCH2 CH2 ); 3.60-3.26 (m, 6H, H-C5 , 2 MeCHN, OCH2 CH2 ); 3.05-2.82 (2t, 2H, OCH2 CH2 ); 2.51 (m, 1H, H-C2 ); 2.17 (m, 1H, H-C2 ); 1.42 (s, 3H, H3 C-C5); 1.17-0.99 (m, 12H, 2 MeCH). 31 P NMR (CDCl3 ): 148.50, 147.23. C53 H60 N5 O12 P (990.1): C 64.29, H 6.18, N 7.07; found: C 63.92, H 5.96, N 6.89. 5 -O-DMTr-N4 -NPEOC-dC-3 -O-(2-(4-nitrophenyl)ethyl-N,N-diisopropylphosphoramidite) (S.54): yield: 0.927 g (91%). Rf (EtOAc/Et3 N 96:4): 0.68. UV (MeOH): 236 (4.56), 276 (4.41), 282 (sh 4.41). 1 H NMR (CDCl3 ): 8.46 8s, 1H, NH); 8.19 (2s, 1H, H-C6); 8.14-8.04 (m, 4H o to NO2 ); 7.42-7.02 (m, 13H, arom. H); 6.84 (m, 1H, H-C5, 4H o to MeO); 6.23 (m, 2H, H-C1 ); 4.58 (m, 1H, H-C3 ); 4.42 (t, 2H, OCH2 CH2 ); 4.15 (m, 1H, H-C4 ); 3.88-3.61 (m, 8H, 2 MeO, OCH2 CH2 ); 3.47 (m, 4H, H-C5 , 2 Me2 CHN); 3.10 (t, 2H, OCH2 CH2 ); 2.87 (2t, 2H, OCH2 CH2 ); 2.69 (m, 1H, H-C2 ); 2.18 (m, 1H, H-C2 ); 1.20-1.01 (m, 12H, 2 MeCHN). 31 P NMR (CDCl3 ): 148.81, 148.47. C53 H60 N6 O13 P (1019.0): C 62.46, H 5.83, N 8.24; found: C 62.24, H 5.81, N 8.00.
Universal NPE and NPEOC Protecting Groups
5-O-DMTr-N6 -NPEOC-dA-3 -O-(2-(4-nitrophenyl)ethyl-N,N-diisopropylphosphoramidite) (S.55): yield: 0.907 g (87%). Rf (EtOAc): 0.66. UV (MeOH): 236 (4.45), 267 (4.58), 274 (sh 4.55). 1 H NMR (CDCl3 ): 8.68 (s, 1H, H-C8); 8.31 (bs, 1H, NH); 8.25 (s, 1H, H-C2); 8.15 (m, 4H o to NO2 ); 7.42-7.12 (m, 13H, arom. H); 6.75 (d, 4H o to MeO); 6.35 (m, 1H, H-C1 ); 4.65 (m, 1H, H-C3 ); 4.42 (t, 2H, OCH2 CH2 ); 4.21 (m, 1H, H-C4 ); 3.90-3.70 (m, 2H, OCH2 CH2 ); 3.73 (2s, 6H, 2 MeO); 3.52-3.43 (m, 4H, 2 Me2 CHN); 3.31 (m, 2H, H-C5 ); 3.08 (t, OCH2 CH2 ); 3.00-2.82 (m, 3H, H-C2 , OCH2 CH2 ); 2.62 (m, 1H, H-C2 ); 1.25-1.02 (m. 12H, 2 MeCH). 31 P NMR (CDCl3 ): 148.67, 148.37. C54 H59 N8 O12 P (1043,1): C 62.18, H 5.70, N 10.74; found: C 61.78, H 6.07, N 9.96.
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Current Protocols in Nucleic Acid Chemistry
5-O-DMTr-N2 -NPEOC-O6 -NPE-dG-3 -O-(2-(4-nitrophenyl)ethyl-N,N-diisopropylphosphoramidite) (S.56): yield: 1.1 g (91%). Rf (EtOAc/Et3 N 96:4): 0.83. UV (MeOH): 237 (4.46), 269 (4.60). 1 H NMR (CDCl3 ): 8.17-8.05 (m, 6H o to NO2 ); 7.95 (s, 1H, H-C8); 7.51-7.14 (m, 15H, arom. H); 6.74 (d, 4H o to MeO); 6.35 (m, 1H, H-C1 ); 4.80 (t, 2H, OCH2 CH2 ); 4.65 (m, 1H, H-C3 ); 4.43 (t, 2H, OCH2 CH2 ); 4.20 (m, 1H, H-C4 ); 3.90 (m, 2H, OCH2 CH2 ); 3.73 (2s, 6H, 2 MeO); 3.52-3.43 (m, 4H, 2 Me2 CHN); 3.31 (2m, 4H, H-C5 , OCH2 CH2 ); 3.09 (t, 2H, OCH2 CH2 ); 3.00-2.84 (m, 2H, OCH2 CH2 ); 2.82 (m, 1H, H-C2 ); 2.45 (m, 1H, H-C2 ); 1.25-1.02 (m, 12H, 2 MeCH). 31 P NMR (CDCl3 ): 148.37, 148.28. C62 H66 N9 O15 P (1208.2): C 61.63, H 5.51, N 10.43; found: C 60.77, H 5.75, N 10.20.
COMMENTARY Background Information The chemical synthesis of oligonucleotides during the last 50 years was marked by a tremendous effort to develop new protecting groups for the various funtionalities of nucleosides and nucleotides. It is common practice to use different protecting groups for each functional group of the nucleobase, the sugar moiety, and the phosphite and phosphate functions. This approach works very well in many instances, but has the disadvantage of a multistep reaction sequence to remove, in the final procedure, all blocking groups and yield the unprotected oligonucleotide. The deprotection process can be simplified by using universal protecting groups (Himmelsbach et al., 1984), especially for the functional groups of the nucleobases and, if necessary, for the hydroxyl groups of the sugar moiety as well as the phosphite and phosphate functions. These universal protecting groups are removed in a one-step reaction at the end of oligonucleotide synthesis. This unit describes the use of an improved protection methodology in the phosphotriester and phosphoramidite approach of oligonucleotide synthesis. The 2-(4-nitrophenyl)ethyl group (Uhlmann and Pfleiderer, 1978, 1980, 1981) provides an advanced phosphateprotecting group due to its chemical inertness in the condensation steps, its stability towards mild acid and base hydrolyses, and its easy cleavage in aprotic solvents by DBU and DBN, respectively, in a β-elimination reaction (Pfleiderer, 1978; Pfleiderer et al., 1980; Pfleiderer and Ichiba, 1981; Uhlmann et al., 1981; Charubala and Pfleiderer 1982; Himmelsbach and Pfleiderer, 1982). In addition, the phenylethyl group can be tuned regarding its stability in the β-elimination process by virtue of the nature of the electron-attracting and electron-donating substituents at the phenyl moiety. In its initial development, the 2-(4-nitrophenyl)ethyl group turned out to fulfill all criteria best in
oligonucleotide synthesis and was considered the most appropriate universal protecting group (Uhlmann and Pfleiderer, 1981).
Compound Characterization 1
H NMR spectra (250 MHz) were measured on a Bruker WM 250 instrument and 31 P NMR spectra on a Jeol 400 (400 MHz) instrument in DMSO-d6 or CDCl3 . Chemical shifts are given in ppm. UV spectra were performed on a Perkin-Elmer Lambda 15 instrument in MeOH (λ max in nm; log ε in brakets; sh = shoulder).
Critical Parameters The protection procedures are straightforward and can be performed by a skilled researcher without problems when following the protocols as outlined. The phosphitylations are more problematic due to the sensitivity of the reaction products, and require some practice and experience with this type of chemistry. The described reaction conditions should not be modified, and the applied starting materials and synthesized intermediates should always be of high purity. Phosphoramidites should be stored in sealed vials in a deep freezer at −20◦ C. Crystalline substances are always more stable than amorphous solids or syrupy materials. Preparative scales for the synthesis of the NPE- and NPEOC-protected nucleosides and their 5 -O-dimethoxytrityl derivatives can be increased without loss of the anticipated yields.
Anticipated Results Following the detailed protocols coupled with experience in nucleic acid chemistry will lead to the reported high yields. The advantage of the NPE- and NPEOC-protected nucleosides is seen in their relatively high stability towards weak acidic and basic reaction conditions allowing chemical modifications at the sugar moiety. The synthesized nucleoside derivatives are very stable and can be stored
Protection of Nucleosides for Oligonucleotide Synthesis
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at room temperature for a long period of time without decomposition.
Time Considerations The total time from a starting nucleoside to its fully protected nucleoside-3 -Ophosphoramidite will be at least 1 week.
Literature Cited Barone, A.D., Tang, J.Y., and Caruthers, M.H. 1984. In situ activation of bis-dialkylaminophosphines—A new method for synthesizing deoxyoligonucleotides on polymer supports. Nucl. Acids Res. 12:4051-4061. Beltz, R.E. and Visser, D.W. 1955. Growth inhibition of Escherichia coli by new thymidine analogs. J. Am. Chem. Soc. 77:736-738. Bredereck, H. 1947. Methylierung von nucleosiden durch diazomethan. Chem. Ber. 80:401-405. Brown, D.M., Todd, A., and Varadarajan, S. 1956. The structure of uridylic acids a and b, and a synthesis of spongouridine (3-β-D-arabofuranosyluracil). J. Chem. Soc. 1956:2388-2393. Caruthers, M.H., Beaucage, S.L., Becker, C., Efcavitch, W., Fisher, E.F., Galluppi, G., DdeHaseth, P., Martin, F., Mateucci, M., and Stabinsky, Y. 1982. Genetic Engineering Principles and Methods (J.K. Setlow and A. Hollaender, eds.) Vol. 4, p. 1. Plenum Press, New York.
Jones, S.S., Reese, C.B., Sibanda, S., and Ubasawa, A. 1981. The protection of uracil and guanine residues in oligonucleotide synthesis. Tetrahedron Lett. 22:4755-4758. Lang, H., Gottlieb, M., Schwarz, H., Farkas, S., Schulz, B.S., Himmelsbach, F., Charubala, R., and Pfleiderer, W. 1999. New 2-(4-nitrophenyl)ethyl (NPE)- and 2-(4-nitrophenyl)ethoxycarbonyl (NPEOC)protected 2 -deoxyribonucleosides and their 3 -phosphoramidites—Versatile building blocks for oligonucleotide synthesis. Helv. Chim. Acta 82:2172-2185. Mitsunobu, O. 1981. The use of diethyl azodicarboxylate and triphenylphosphine in synthesis and transformation of natural products. Synthesis 1-28. Pfleiderer, W. 1978. Modern strategy in oligonucleotide chemistry. Les Colloques de l’INSERM. Nucleosides Nucleotides 81:177193. Pfleiderer, W. and Ichiba, M. 1981. Chemical synthesis of the ribo-hexamer CpApApCpCpA. Nucl. Acids Res. 9:169-172. Pfleiderer, W., Uhlmann, E., Charubala, R., Flockerzi, D., Silber, G., and Varma, R.S. 1980. Recent progress in oligonucleotide synthesis. Nucl. Acids Res. 7:61-71.
Charubala, R. and Pfleiderer, W. 1982. Synthesis of inosinate trimer I2 p5 I2 p5 I and tetramer I2 p5 I2 p5 I2 p5 I. Tetrahedron Lett. 23:47894792.
Reese, C.B. and Ubasawa, A. 1980. Reaction between 1-arenesulphonyl-3-nitro-1,2,4-triazoles and nucleoside base residues. Tetrahedron Lett. 22:2265-2268.
Daskalov, H.P., Sekine, M., and Hata, T. 1981. Synthesis and properties of O6-substituted guanosine derivatives. Bull. Chem. Soc. Jpn. 54: 3076-3083.
Schaller, H., Weimann, G., Lerch, B., and Khorana, H.G. 1963. The stepwise synthesis of specific deoxyribopolynucleotides. Protected derivatives of deoxyribonucleosides and new syntheses of deoxyribonucleoside-3 -phosphates. J. Am. Chem. Soc. 85:3821-3827.
Flockerzi, D., Silber, G., Charubala, R., Schlosser, W., Varma, R.S., Creegan, F., and Pfleiderer, W. 1981. Synthese und eigenschaften von 2 -O-und 3 -O-(tert.Butyldimethyl-silyl)-5 -O(4-methoxytrityl)-sowie 2 ,3 -Bis-O-(tert.butyldimethylsilyl(ribonucleosiden—Ausgangssubstanzen f¨ur oligoribonucleotid-synthesen. Liebigs Ann. Chem. 1981:1568-1585. Gaffney, B.L. and Jones, R.A. 1982. A new strategy for the protection of deoxyguanosine during oligonucleotide synthesis. Tetrahedron Lett. 23:2257-2260. Hayes, D.H., Michelson, A., and Todd, A.R. 1955. Mononucleotides derived from deoxyadenosine and deoxyguanosine. J. Chem. Soc. 1955:803815. Himmelsbach, F. and Pfleiderer, W. 1982. Bis-(pnitrophenylethyl)phosphoromono-chloridate, a versatile phosphorylating agent. Tetrahedron Lett. 23:4793-4796.
Universal NPE and NPEOC Protecting Groups
Hunt, B.J. and Rigby, W. 1967. Short column chromatography. Chem. Ind. 65:1868-1869.
Himmelsbach, F., Schulz, B.S., Trichtinger, T., Charubala, R., and Pfleiderer, W. 1984. The pnitrophenylethyl (NPE) group, a versatile new blocking group for phosphate and aglycone protection in nucleosides and nucleotides. Tetrahedron 40:59-72.
Sinha, N.D., Biernat, J., and K¨oster, H. 1984. Polymer support oligonucleotide synthesis XVIII: Use of β-cyanoethyl-N,Ndialkylamino/N-morpholino phosphoramidite of deoxynucleosides for the synthesis of DNA fragments simplifying deprotection and isolation of the final product. Nucl. Acids Res. 12:4539-4557. Sobol, R.W., Henderson, E.E., Kon, N., Shao, J., Hitzges, P., Mordechai, E., Reichenbach, N.L., Charubala, R., Pfleiderer, W., and Suhadolnik, R.J. 1995. Inhibition of HIV1 replication and activation of RNase L by phosphorothioate/phosphodiester 2 ,5 oligoadenylate derivatives. J. Biol. Chem. 270:5963-5678. Ti, G.S., Gaffney, B.L., and Jones, R. 1982. Transient protection: Efficient one-flask syntheses of protected deoxynucleosides. J. Am. Chem. Soc. 104:1316-1319. Trichtinger, T., Charubala, R., and Pfleiderer, W. 1983. Synthesis of O6 -p-nitrophenyl-ethyl guanosine and 2 -deoxyguanosine derivatives. Tetrahedron Lett. 25:711-714.
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Uhlmann, E. and Pfleiderer, W. 1978. New phosphate protecting groups in nucleotide synthesis. Nucl. Acids Res. 4:s25-s28. Uhlmann, E. and Pfleiderer, W. 1980. New improvements in oligonucleotide synthesis by use of the p-nitrophenylethyl phosphate blocking group and its deprotection by DBU or DBN. Tetrahedron Lett. 22:1181-1184. Uhlmann, E. and Pfleiderer, W. 1981. Substituierte β-phenylethyl-gruppen. Neue schutzgruppen f¨ur oligonucleotide-synthesen nach dem phosphors¨auretriester-verfahren. Helv. Chim. Acta 64:1688-1703. Uhlmann, E., Charubala, R., and Pfleiderer, W. 1981. Synthesis of 3 -terminal nucleoside phosphates using the p-nitrophenylethyl blocking group. Nucl. Acids Res. 9:131-134. Watkins, B.E., Kiely, J.S., and Rapoport, H. 1982. Synthesis of oligodeoxyribo-nucleotides using N-benzyloxycarbonyl-blocked nucleosides. J. Am. Chem. Soc. 104:5702-5708. Zon, G., Gallo, K.A., Samson, C.J., Shao, K., Byrd, R.A., and Summers, M.F. 1985. Analytical studies of ‘mixed sequence’ oligodeoxyribonucleotides synthesized by competitive coupling of either methyl- or β-cyanoethyl-N,Ndiisopropylamino phosphoramidite reagents, including 2 -deoxyinosine. Nucl. Acids Res. 13:8181-8196.
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CHAPTER 3 Synthesis of Unmodified Oligonucleotides INTRODUCTION ecause of the significant progress achieved in the chemical synthesis of oligonucleotides during the past four decades, synthetic oligonucleotides have become readily available and have fueled the biotechnology revolution that has irreversibly changed biomedical research and the pharmaceutical industry. For example, without the ability to rapidly and efficiently synthesize DNA oligonucleotides, the development of the polymerase chain reaction (PCR) and its multiple applications would have been difficult, if not impossible, because this technology is completely dependent upon the use of DNA primers. Similarly, the availability of synthetic oligonucleotides has been instrumental in the development of automated DNA sequencing, which is also dependent on the use of DNA primers. An important biological application of synthetic oligonucleotides relates to site-specific mutagenesis of genes. Mutagenesis of this type has been utilized to study protein structure-function relationships, and to alter the therapeutic spectrum of pharmaceutically active proteins. Synthetic oligonucleotides have also found application as diagnostic hybridization probes and, more recently, in the use of DNA chips for highthroughput detection of genetic diseases and identification of genomic single-nucleotide polymorphisms. Furthermore, the availability of synthetic RNA oligonucleotides has led to the preparation of ribozymes, which are RNAs with catalytic activity. In this regard, it is being speculated that RNA may have been the primary self-replicating molecule from which life originated. Collectively, the multiple biomedical applications of synthetic DNA and RNA oligonucleotides are a direct measure of the colossal impact the methods for rapid and efficient preparation of these biomolecules have had on the life sciences.
B
Like Chapter 2, this chapter is composed of a number of overviews that provide investigators with a fundamental knowledge of solid-phase oligonucleotide synthesis, as well as a number of step-by-step protocols delineating the preparation of various oligonucleotides from suitably protected nucleoside phosphoramidite or H-phosphonate derivatives. browses through the physical and chemical properties of liquid-phase and solidphase supports that have been developed over the years, and compares their suitability for oligonucleotide synthesis. The unit also examines the properties of linkers for transient or permanent attachment of properly protected nucleosides to the derivatized support of interest. Conditions for the release of synthetic oligonucleotides from specific supports have been discussed in detail. A step-by-step protocol for the attachment of nucleosides to commonly functionalized supports via either a succinyl or a hydroquinone-O,O -diacetyl linker is provided in UNIT 3.2. In this context, an ultra fast and efficient functionalization of solid supports has been achieved using microwave-assisted protocols. The details of these innovative procedures are addressed in UNIT 3.13. To facilitate synthesis in multiwell plates, deoxyribonucleoside phosphoramidites that have a cleavable linker between the 3 -hydroxyl of the nucleoside and the phosphoramidite moiety have been developed to functionalize underivatized amino supports in any well. This approach eliminates the error-prone placement of nucleoside-specific prederivatized supports into the correct wells for the preparation of specific DNA sequences. Moreover, this approach enables one to synthesize and isolate oligonucleotides with a free 3 -hydroxyl without the requirement for additional cleavage or deprotection steps. UNIT 3.12 outlines the preparation of these UNIT 3.1
Current Protocols in Nucleic Acid Chemistry 3.0.1-3.0.3, September 2007 Published online September 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/0471142700.nc0300s30 C 2007 John Wiley & Sons, Inc. Copyright
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novel phosphoramidite synthons and their use in the synthesis of oligonucleotides in multi-well plates or in the tandem synthesis of multiple oligonucleotides in a single strand. UNIT 3.16 features the use of 5 -O-DMTr-deoxyribonucleoside tert-butyl phosphoramidites in the phosphitylation of a hydroxylated solid support. Cleavage of the 5 -O-DMTr group from the nucleoside phosphite triester under acidic conditions allows standard solidphase oligonucleotide synthesis to proceed while generating a strategic H-phosphonate diester function linking the oligonucleotide to the solid support. This H-phosphonate diester linkage enables post-synthesis release of the oligonucleotide with free 5 - and 3 hydroxyls from the universal and recyclable solid support via a simple transesterification reaction. Solid-phase synthesis of oligonucleotides is governed by a number of parameters. For example, UNIT 3.3 reviews the mechanism of activation of deoxyribonucleoside phosphoramidites, the type of activators required for such activation, and the consequences of an activator’s acidity on the production of synthetic oligonucleotides. Steric factors affecting the condensation rates of deoxyribonucleoside phosphoramidites and the significance of “capping” and oxidation reactions in the chemical synthesis of oligonucleotides, along with oligonucleotide deprotection strategies, are discussed at length in the unit. Finally, UNIT 3.3 deals with the application of dinucleotide and trinucleotide phosphoramidites as alternatives to monomeric deoxyribonucleoside phosphoramidites in the synthesis of DNA oligonucleotides. In this regard, UNIT 3.14 provides a detailed procedure for the solution-phase synthesis of short oligonucleotides using deoxyribonucleoside phosphoramidites with solid-supported reagents such as polyvinyl pyridinium tosylate as an activator for the coupling reaction and polymer-bound periodate or tetrathionate as an oxidant. The method is particularly attractive for the clean preparation of di- and trinucleotides, which can be converted to 3 -phosphoramidite or 3 -H-phosphonate monoester building blocks and used in solid-phase oligonucleotide synthesis. Additional synthetic strategies and parameters related to the synthesis of oligodeoxyriboand oligoribonucleotides according to the H-phosphonate method are discussed in UNIT 3.4. An optimized synthetic protocol for oligoribonucleotide synthesis is described to further validate the potential of the method. The chemical synthesis of DNA oligonucleotides without nucleobase protection has been and is still an elusive goal. An overview of the strategies that have been developed so far to achieve this goal is presented in UNIT 3.10. A successful synthetic approach to the preparation of deoxyribonucleoside phosphoramidites without nucleobase protecting groups is summarized in UNIT 3.15. These monomeric phosphoramidites are amenable to automated solid-phase oligonucleotide synthesis and may thus become invaluable tools. Strategies in the selection of N-, 5 -O-, and 2 -O-protecting groups for efficient synthesis of oligoribonucleotides according to the phosphoramidite approach are discussed in depth in UNIT 3.5. Critical synthesis parameters such as the type of support being used and selection of the optimal activator for ribonucleoside phosphoramidites, as well as oligonucleotide deprotection protocols, are also reviewed in the unit to provide a comprehensive account of RNA phosphoramidite chemistry. As a logical extension of UNIT 3.5, the solid-phase synthesis of oligoribonucleotides with 2 -O-tert-butyldimethylsilyl, 2 -O-(2nitrobenzyloxymethyl), or 2 -O-[(triisopropylsilyl)oxy]methyl groups according to the phosphoramidite approach is delineated in UNITS 3.6–3.8, respectively, along with detailed methods for the deprotection of these oligonucleotides.
Introduction
In future supplements, this chapter will also incorporate step-by-step protocols describing new methods for the synthesis of unmodified DNA oligonucleotides that employ either new phosphate-protecting groups or novel coupling procedures to accommodate specific nucleic acid applications. In this regard, the use of 3-(N-tert-butylcarboxamido)-1-propyl
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Current Protocols in Nucleic Acid Chemistry
and 4-oxopentyl groups for phosphate/thiophosphate protection in solid-phase DNA synthesis is presented in UNIT 3.9, along with that of the 4-methylthio-l-butyl group in UNIT 3.11. These protecting groups may find application in large-scale preparations of therapeutic oligonucleotides. Likewise, oligoribonucleotide synthesis protocols comparing the properties of selected 2 -O-protecting groups, coupling methods, and deprotection conditions will be included in an effort to contribute the best nucleic acid tools to investigators pushing further the frontiers of knowledge. Serge L. Beaucage
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Solid-Phase Supports for Oligonucleotide Synthesis INTRODUCTION TO SOLID-PHASE SYNTHESIS The quest to understand and create biological molecules has long challenged synthetic chemists. In particular, the chemical synthesis of peptides and nucleic acids has always been a major pursuit. The primary structure of these molecules is a linear assembly of repeating units linked together in a defined orientation. Although solution-phase synthetic methods for coupling small units together were developed many years ago, the large number of couplings needed to assemble useful sequences was daunting. This was because each step required some type of workup, extraction, or purification, and the labor and cumulative loss of material from all the manipulations rapidly became significant problems. Indeed, the pioneering work by Khorana (1979) on total gene synthesis was not considered of practical importance by some researchers because of the enormous effort involved. The problems involved in performing so many repetitive steps were addressed by Merrifield (1965) with the introduction of solidphase synthesis (Fig. 3.1.1). In this strategy, a large insoluble support is covalently linked to the end of the sequence being assembled. The product on the surface of the support is available to react with reagents in the surrounding solution phase. The extended products remain covalently linked to the insoluble support while unreacted reagents remain free in solution. Therefore, at the completion of each step, the products can be rapidly and conveniently isolated by simply washing the unbound reagents away from the support. This can be performed as easily as filtering off the support and washing it with solvent. The support with its attached product is then ready for immediate use in the next step, as long as moisture contamination has not been introduced (in which case the support must be dried before use). In practice, it is convenient to handle the supports inside sealed reactors or columns so exposure to the atmosphere is minimized. This is also ideal for automation and the necessary reagent additions and solvent washes are readily mechanized. The process of adding each unit is repeated over and over until the desired sequence has been assembled on the surface of the support. The product can then be released from the support
UNIT 3.1
by cleavage of the covalent attachment (linker arm), and after removing the protecting groups, the synthesis is complete. This strategy was originally applied to peptide synthesis, but it is also applicable to other linear macromolecules, such as DNA and RNA (Beaucage and Iyer, 1992) and oligosaccharides (Adinolfi et al., 1996). Recently, there has been a great deal of interest in applying this strategy to the combinatorial synthesis of small molecules and a new field of solid-phase organic chemistry (SPOC) is rapidly developing (Fruchtel and Jung, 1996; Porco et al., 1997). In this review, the main focus is on supports for oligodeoxyribonucleotide and oligoribonucleotide synthesis. The synthetic strategies are often similar, particularly when synthetic libraries are prepared.
ADVANTAGES OF SOLID-PHASE SYNTHESIS The principal advantage of solid-phase synthesis is the ease with which immobilized products can be separated from other reactants and by-products. The simple filtration and washing steps are readily automated, and the method is ideal for the synthesis of linear molecules, which require the repetition of the same steps for every chain extension cycle. The use of insoluble solid-phase supports also permits relatively small quantities of material to be synthesized, because the additional physical bulk of the support, which is ~10 to 100 times the mass of the attached nucleoside, can be handled more easily than the nucleoside alone. Also confinement of the support inside a synthesis column eliminates handling losses. A small synthesis scale is important because of the high cost of reagents. Very little material is required for many biochemical applications and most syntheses actually prepare much more material than required. Therefore, as instrumentation has improved, the synthesis scale has decreased. Presently, synthesis on a 40-nmol scale, instead of a 0.2- to 1-µmol scale, is preferred for may applications. Oligonucleotide synthesis on a picomole scale or less may eventually become more common (Weiler and Hoheisel, 1997). It is already possible to synthesize molecules on single beads and to characterize the picomole quantities of synthetic peptides (Rapp, 1997) or oligonu-
Contributed by Richard T. Pon Current Protocols in Nucleic Acid Chemistry (2000) 3.1.1-3.1.28 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Unmodified Oligonucleotides
3.1.1
Solid-Phase Supports for Oligonucleotide Synthesis
Figure 3.1.1 The general strategy for solid-phase oligonucleotide synthesis. The first step is attachment of a mononucleoside/tide (N1) to the surface of an insoluble support (P) through a covalent bond. Excess monomers, which are not chemically attached to the support, are washed away. Before chain elongation can proceed, the terminal-protecting group (=) on the nucleoside must be removed. This exposes a free 5′-OH or 3′-OH group, depending on the orientation of the synthesis. Usually synthesis proceeds from the 3′- to 5′-direction and the terminal protecting group is an acid-labile DMTr group. The next nucleotide unit (N2) can then be added using the appropriate synthesis chemistry (usually phosphoramidite). An excess of reagent is used to force the coupling reaction to occur on as many of the immobilized nucleotides as possible. After the coupling reaction, excess reagents are washed away. Depending on the coupling chemistry, the reaction is followed by a capping step, to block off nonextended sites, and an oxidation step (these steps are not shown; see UNIT 3.3 for details) to complete the chain-extension cycle. The process of terminal-protecting group removal and chain extension is then repeated, using different bases, until the desired sequence has been assembled. Some or all of the protecting groups may optionally be removed, and then the covalent attachment to the support is hydrolyzed to release the product. After removal of any remaining protecting groups, the oligonucleotide is ready for purification and use.
3.1.2 Current Protocols in Nucleic Acid Chemistry
cleotides (Seliger et al., 1997) present on single beads. The simplicity and similarity of the steps required for each chain extension reaction also greatly facilitate synthesis of modified oligonucleotides. As long as the modified substituents do not require any incompatible chemical treatments (i.e., to remove protecting groups), the inclusion of different bases and nucleosides, linkage inversions, branch points, non-nucleotide units, and end modifications can be readily accomplished. This is particularly so when the modified substituents are available as phosphoramidite derivatives, which use the same coupling chemistry as do regular bases (Beaucage and Iyer, 1993). Chimeric oligonucleotides containing peptide or peptide nucleic acid (PNA) sequences can, however, also be prepared (Bergmann and Bannwarth, 1995; Hyrup and Nielsen, 1996; van der Laan et al., 1997). Although, in these cases, the different coupling conditions and protecting groups require much more attention to ensure overall compatibility. Finally, combinatorial methods can be used to create large numbers of different sequences. In the simplest application, multiple bases (“mixed bases”) can be incorporated at defined positions by using a mixture of different monomers, instead of a single monomer, in the chain extension reaction. This procedure was originally developed to prepare oligonucleotide probes from peptide sequences when the exact codon usage was unknown. Later, this method became important when large libraries of degenerate or random sequences were required for in vitro selection experiments, such as the systematic evolution of ligands by exponential enrichment (SELEX) technique (Gold et al., 1995; see Chapter 9). Although DNA synthesizers can prepare mixed-base sites by on-line mixing, large numbers of degenerate sites are best made up by manually preparing solutions containing the desired ratio of nucleotides and incorporating the premixed reagents on the synthesizer. This is also the procedure used in base doping, when only one base, at random, within a particular section needs to be mutated (Hermes et al., 1989). Another combinatorial approach was developed to simplify the synthesis of large numbers of oligonucleotides. This procedure used cellulose disks of filter paper as the insoluble support and became known as filter disk or segmented solid-phase synthesis (Frank et al., 1983; Matthes et al., 1984; Ott and Eckstein, 1984; Frank, 1993). In this procedure, multiple filter disks
(each producing one unique oligonucleotide) are stacked together and handled at once. Reagents can be easily passed through the stack from top to bottom, and the number of oligonucleotides synthesized is limited only by the maximum stack height that can be manipulated. A different oligonucleotide sequence is prepared on each disk by interrupting the synthesis after each chain extension step. The individual filter disks are then sorted into separate piles according to the next base to be added. The insoluble support in this case provides the means to sort the products and to separate them from the excess reagents. For normal oligonucleotides, the sorting results in a maximum of four piles, because only dA, dC, dG, and T base additions are required. Thus an operator manipulating four concurrent syntheses can produce a large number of oligonucleotides per day. This method is not limited to paper filter disks; stackable “synthesis wafers” containing packets of support in bead form have also been used. The sorting step, however, is quite difficult to automate; and although semimechanized instruments have been reported (Seliger et al., 1987; Beattie et al., 1988), the segmented approach has not been widely adopted. The ease with which immobilized oligonucleotides can be manipulated has also lead to the development of combinatorial strategies for the synthesis of oligonucleotide libraries. Unlike the above strategies, which release the oligonucleotide product from the support at the end of the synthesis, the oligonucleotides are left attached to the insoluble support (Markiewicz et al., 1994). This method can be used to create dispersed libraries, when the sequences are prepared on separate beads, or integrated libraries, when one- or two-dimensional arrays of sequences are prepared on a single surface. The sequence identity of each element in an integrated array is known from its spatial coordinates, whereas the sequence of elements in a dispersed library must be deduced from either direct sequencing (Seliger et al., 1997) or other sequence tags. The most elegant and powerful demonstration of this technique is the synthesis of high-density arrays on small (1.28 cm2) glass chips using photolithography and light-sensitive protecting groups (Fodor et al., 1991). With the appropriate masking, any set of oligonucleotides of length N can be performed using only 4N coupling steps, and this technique can produce arrays of >106 different sequences (Lipshutz et al., 1995; McGall et al., 1996). Other combinatorial strategies using either glass plates (Milner et al., 1997) or
Synthesis of Unmodified Oligonucleotides
3.1.3 Current Protocols in Nucleic Acid Chemistry
polypropylene sheets (Matson et al., 1995; Weiler and Hoheisel, 1996) as the insoluble support have been described for the synthesis of oligonucleotide arrays, although the array densities were much lower.
DISADVANTAGES OF SOLID-PHASE SUPPORTS Although a powerful technique, solid-phase synthesis has some drawbacks. The main limitation is the need for very high coupling yields in every chain extension step. This is because the overall yield of product decreases rapidly as the number of consecutive chain extension steps increases (Fig. 3.1.2). For example, if each base addition step had a yield of 90%, then the amount of dinucleotide produced (one base addition) is 90%. The yield of trinucleotide (two base additions) is 0.90 × 0.90 × 100% = 81%; the yield of tetranucleotide (three base additions) is 0.90 × 0.90 × 0.90 × 100% = 73%; and so on. Note that the first nucleoside is attached to the insoluble support before the start of oligonucleotide synthesis and the efficiency of that step is not included in the calculation. The mathematical relationship between the overall yield (OY) and the average coupling efficiency (AY) is either n
AY OY = × 100% 100
or N−1
AY OY = 100
Solid-Phase Supports for Oligonucleotide Synthesis
× 100%
where n is the number of coupling steps and N is the length of the oligonucleotide. The second equation assumes that the synthesis was performed by extending the product by one base at a time, as is usual. The consequence of the exponential relationship between overall yield and average coupling efficiency is that long oligonucleotides cannot be prepared without very high yields in every step. The most difficult step is usually the coupling reaction; but in some strategies (e.g., light-directed synthesis of arrays or the use of liquid-phase supports), quantitative removal of the terminal-protecting group is also problematic. Coupling yields that would be acceptable for most solution-phase reactions (e.g., the 90% yield assumed in the above example) are not adequate; only yields >98% are acceptable. The lack of a coupling reaction that could reliably produce such high efficiencies was the major reason why solid-phase oligonucleotide synthesis was not successful until the early 1980s. After the discovery of trivalent phosphite–coupling chemistry and phosphoramidite derivatives (Caruthers, 1991), however, average coupling efficiencies of 99% or more were possible. Such high coupling efficiencies now allow oligonucleotides as long as 200 bases to be prepared (Bader et al., 1997b). Another consequence of producing less than 100% coupling efficiencies is the accumulation of failure sequences containing deletions. The number of these failure products can be greatly reduced by the addition of a capping step after each chain extension reaction. This step, which typically uses acetic anhydride to acetylate nonextended molecules, prevents the failure
Figure 3.1.2 Overall yield vs. number of couplings. The overall yield of full-length product decreases with the number of coupling reactions for average coupling efficiencies of 90%, 95%, 98%, and 99%.
3.1.4 Current Protocols in Nucleic Acid Chemistry
sequences from participating in any further reactions; however, a series of failure sequences, each one base shorter than the desired full-length product, will be present at the end of the synthesis. Separating the full-length product (of length N) from the shorter failure sequences and especially the N – 1 failure sequence is another significant problem. This purification step becomes more difficult as oligonucleotide length increases, and for oligonucleotides greater than ~30 bases long, only polyacrylamide gel electrophoresis (PAGE) has sufficient resolving power to separate the full-length product from the N – 1 component. Fortunately, however, many biochemical applications do not have stringent purity requirements; and if the coupling efficiency was high enough, the mixture of products produced can often be used with either minimal (desalting) or no purification (Pon et al., 1996). Analysis of the synthetic products still attached to the surface of the insoluble support also presents a major difficulty for researchers developing new techniques or new solid-phase supports. This is an especially significant problem for applications using immobilized arrays, because removal of the products for characterization is often difficult, if not impossible. Nuclear magnetic resonance (NMR) studies of immobilized products on solvent-swollen (gelphase) polymers (Bardella et al., 1993) can be performed; but because such supports are not preferred for oligonucleotide synthesis, there have been few studies relating to oligonucleotide synthesis. Rigid supports can be studied using NMR and magic angle spinning, but there has been only one report of 31P NMR performed on controlled-pore glass (CPG) particles with oligonucleotides (Macdonald et al., 1996). Recently, ellipsometry, interferometry, and optical wave guides have been used to study oligonucleotide arrays (Stimpson et al., 1995; Gray et al., 1997), but these techniques do not provide specific information about the fidelity of the oligonucleotide synthesis. Finally, the cost of the support is a major factor when performing large-scale syntheses because, even with high loading supports, ∼3 g of support is required for each gram of oligonucleotide product. Because most supports are expensive and can be used only once, there is a strong economic incentive to develop methods for regenerating and reusing the supports, especially when tonne quantities of products are required. Recently, examples of up to 12 syntheses of oligonucleotide phosphodiester
(Pon et al., 1999) and phosphorothioate (Pon et al., 1998) sequences on the same reusable supports have appeared, and further improvements in this area are expected.
CHEMICAL REQUIREMENTS FOR SOLID-PHASE SUPPORTS A wide variety of different insoluble support materials have been developed for different applications. The ideal support should contain an appropriate chemical group on its surface that can be selectively coupled, usually through a linker arm, to the first monomer unit. Normally, supports for oligonucleotide synthesis are purchased with primary amino group functionality, although hydroxyl and carboxyl derivatized supports may also be obtained. The amount of surface derivatization (loading) on the support determines the maximum amount of product that can be prepared. Supports with loadings of 100 to 1000 µmol/g or more are available for the synthesis of either peptides or small molecules (Winter, 1996). Oligonucleotide synthesis, however, is almost always performed using nucleoside loadings of less than ~100 µmol/g; optimum results are obtained on supports with less than ~40 µmol/g of nucleoside, because the efficiency of coupling decreases as the number of molecules on the surface increases. Because the amount of support can usually be increased to accommodate the scale required, supports with loadings >50 µmol/g are not commonly used. The lower coupling efficiencies obtained with higher loaded supports can actually make it counterproductive to use these materials in most automated DNA synthesizers. The structure of the compound(s) used to join the surface of the support to the first nucleoside is also of critical importance (Fig. 3.1.3). This attachment is generally composed of two distinct portions. The first portion is the spacer that connects the active functional group (usually NH2 or OH groups) to the matrix of the insoluble support. This spacer can be as simple as a single methylene group (e.g., aminomethyl polystyrene) or it can be a lengthy alkyl or alkyloxy chain (e.g., long-chain alkylamine CPG). Generally, a long chain is preferred to distance the terminal functional group from the support’s surface. Usually, the supports are sold with a satisfactory spacer, but sometimes additional spacers are added to change the terminal functional group or to increase the overall length (Katzhendler et al., 1987; van Aerschot et al., 1988; Arnold et al., 1989).
Synthesis of Unmodified Oligonucleotides
3.1.5 Current Protocols in Nucleic Acid Chemistry
A second difunctional molecule is then required to connect the amino group on the support to the first nucleoside unit. In oligonucleotide synthesis, this structure is commonly referred to as a linker or linker arm. In peptide synthesis and combinatorial synthesis, this structure is referred to as either a handle or anchor, terms not usually associated with oligonucleotide synthesis. In oligonucleotide synthesis, the linker arms are usually dicarboxylic acids, such as succinic acid (Pon et al., 1988; Damha et al., 1990; Bhongle and Tang, 1995) or hydroquinone-O,O-diacetic acid (Pon and Yu, 1997a), that connect the nucleoside to the support via ester and amide bonds. The length, rigidity, and hydrophobicity of the linker arm can affect coupling efficiency (Katzhendler et al., 1989), and the chemical stability restricts the conditions that can be used during synthesis. This affects the choice of protecting groups. Most linkers for oligonucleotide synthesis are resistant to acidic conditions and cleavable by basic conditions. This allows the most popular combination of protecting groups—acid-labile 5′-dimethoxytrityl (DMTr) groups and baselabile N-acyl and cyanoethyl phosphate protecting groups—to be used. As will be discussed later, the speed with which the linker arm can be cleaved is also an important consideration. Strategies that require removal of oligonucleotide-protecting groups without cleavage from the support also require linker arms that are either very stable or removable using
DMTO
O
conditions orthogonal to the deprotection conditions. Finally, different linker arms can be used to prepare oligonucleotides with terminal end modifications, such as 3′-phosphate, amino, carboxyl, thiol, or other substituents. The chemical properties of the rest of the surface should be either inert or capable of being made inert by silylation, benzoylation, or other similar passivating treatment (Pon, 1993; Tang and Tang, 1997). This is because residual groups, such as amino, hydroxyl, or silanol groups, can also react with phosphoramidite derivatives. This creates failure sequences coupled to the support through either phosphoramidate or phosphodiester linkages and lacking the correct 3′-nucleoside. When coupling reactions are monitored by quantitation of the orange dimethoxytrityl cation released during detritylation, the formation of such failure sequences can be deduced from apparent coupling yields of >100%. Fortunately, the phosphodiester linkages to the surface of the support are difficult to hydrolyze, and these failure sequences are not released from the support by the usual cleavage conditions (Pon et al., 1988). The hydrophobicity of the support’s surface is another consideration. Supports that are quite hydrophobic, such as polystyrene and benzoylated polymethacrylate, are sometimes preferred over supports such as CPG and polyethylene glycol (PEG), which have hydrophilic surfaces (McCollum and Andrus, 1991; Tang
B O
O
N H
A O
OR
H N
O
O
O
succinyl linker long-chain alkyl amine spacer
DMTO
O
Si
CPG support
B O
O B O succinyl linker
Solid-Phase Supports for Oligonucleotide Synthesis
N H aminomethylpolystyrene support
Figure 3.1.3 The structure of the two most commonly used solid-phase supports for oligonucleotide synthesis. (A) LCAA-CPG. (B) Aminomethyl polystyrene (nonswelling and highly cross-linked). In both cases, a nucleoside is attached to the amino group of the support through a succinic acid linker, which can be cleaved by ammonium hydroxide after the synthesis. DMT, 5′-dimethoxytrityl (DMTr).
3.1.6 Current Protocols in Nucleic Acid Chemistry
and Tang, 1997). All phosphoramidite-coupling reactions are sensitive to moisture contamination, and such contamination is presumed to be more easily washed off the hydrophobic supports. This may allow greater synthesis efficiency with smaller excesses of reagent; however, quantitative comparisons are difficult to make, and both rigid polystyrene and CPG supports are widely used.
PHYSICAL AND CHEMICAL PROPERTIES OF SOLID-PHASE SUPPORTS The accessibility of the support’s surface to incoming reagents is probably the most important consideration when choosing the physical properties of the insoluble support. Although a greater surface area provides higher capacity, increased porosity must be balanced against steric restrictions and rate-limiting diffusion. The following sections discuss the major classes of solid-phase supports, categorized by the type of surface accessibility.
is about 100 mg per gram of starting support (Bonora, 1995). The advantages of having a homogeneous solution include lower costs, because less reagent excess is required, and the ability to use spectroscopic methods (UV/VIS, NMR, Fourier transform-IR) to monitor the reactions and quality of the immobilized products. Furthermore, the method does not require any elaborate instrumentation because the reactions and precipitation/filtration steps are performed in ordinary glassware. Consequently, liquid-phase supports were some of the earliest supports to be used (Cramer et al., 1966; Hayatsu and Khorana, 1966). The method, however, is not easily automated, and each chain extension cycle requires several hours to perform. Moisture contamination must also be scrupulously avoided, because the PEG supports are very hydrophilic. Nevertheless, this method is suitable for larger scale oligonucleotide synthesis, when cost is more important than speed.
Gelatinous Polymer Supports Liquid-Phase Supports Liquid-phase supports are high molecular weight polymers that can be completely dissolved in the solvents required for synthesis but can be precipitated or crystallized in other solvents or solvent conditions. When they are dissolved in solution, coupling reactions on liquid-phase supports are performed in the same manner as conventional solution-phase synthesis. After completion of the coupling reaction, however, the liquid-phase support is precipitated by adding a solvent in which it is insoluble. The resulting precipitated support can then be filtered off and washed free of excess reagents in the same manner as other insoluble supports. After the washing step, the support is redissolved in the appropriate solvent, and the synthesis continued. Alternately, dialysis or ultrafiltration can also be used to remove low molecular weight impurities. The most widely used liquid-phase supports are PEG polymers (Bonora, 1995) with average molecular weights varying between 5,000 and 20,000, although cellulose acetate (Kamaike et al., 1988) and poly(N-acryloylmorpholine) polymers (Bonora et al., 1996) have also been used. These supports are soluble in solvents such as dichloromethane, pyridine, and acetonitrile but insoluble in solvents such as ethers and alcohols. Nucleoside loadings of ∼100 to 200 µmol/g are generally obtained, and the purified yield of oligonucleotide 8- to 20-mers
The first insoluble supports developed were polystyrene-divinylbenzene polymers with only a small amount (1% to 5%) of cross-linking (Pon, 1993). These supports could swell up to five times their dry volume in nonpolar solvents, such as dichloromethane, to provide a large surface area and loading capacity (0.1 to 1.5 mmol/g). Other swellable polyacrylamide-containing supports, with loadings up to 5 mmol/g, have also been developed (Winter, 1996). In these supports, up to 99% of the reactive sites are located inside the bead. They are classified as gelatinous polymer supports because of the gel-like environment in which the reactions take place (Rapp, 1996). These supports perform very poorly when used for solid-phase oligonucleotide synthesis because they don’t swell satisfactorily in the polar solvents required and because reagent diffusion into and out of the supports is slow. Therefore, the swollen beads can be used only in batch reactors and not in continuous flow synthesizers (Belagaje and Brush, 1982; Ito et al., 1982; Ohsima et al., 1984). Only the very earliest oligonucleotide syntheses were attempted on these types of supports; and with one exception (Montserrat et al., 1994), their present use is restricted to peptide synthesis.
Macroporous Supports The difficulties mentioned above were overcome by the development of rigid macroporous
Synthesis of Unmodified Oligonucleotides
3.1.7 Current Protocols in Nucleic Acid Chemistry
Solid-Phase Supports for Oligonucleotide Synthesis
supports. These supports are based on inorganic materials, such as silica gel and CPG, or highly cross-linked polymers, such as polystyrene or polymethacrylate. Well-defined pores are created in these supports to increase the surface area and loading capacity. They do not become swollen with solvent and have permanent porosity. Their rigidity allows them to be used in packed continuous-flow columns, and their properties are very similar to the packing materials used in HPLC separations. The maximum loading possible on the rigid supports, however, is much less than on the swellable supports. Silica gel and porous glass supports are ideal nonswelling materials that are readily derivatized using techniques developed by the glass fiber and chromatography industries. CPG beads, which are stronger and easier to handle than is silica gel, are preferred and are available in three particle sizes—125 to 177 µm, 74 to 125 µm, and 37 to 74 µm—with large (75 to 4000 Å in diameter) and very uniformly sized pores. The maximum pore diameter distribution is only ±10% for 80% of the pore volume. A variety of chemically derivatized CPG supports with different functional groups is available, including magnetic CPG beads. The surface area and loading depend on the pore size; the beads with larger pores have lower loadings. Long-chain alkylamine (LCAA) derivatized CPG supports with 500-Å pores and amino loadings of ∼100 µmol/g are the most commonly used. These supports are usually derivatized with 30 to 40 µmol/g of nucleoside (Pon, 1993) and are suitable for the synthesis of oligonucleotides of up to 50 to 60 bases. The 500-Å pore size begins to restrict the coupling efficiency of longer oligonucleotides because of steric factors; however, much longer oligonucleotides (100 to 150 bases) can be prepared on 1000-Å CPG supports (Efcavitch et al., 1986). Synthesis of very long oligonucleotides also benefits from a support with a low surface loading (~5 µmol/g), because it contributes to greater coupling efficiency. Highly cross-linked rigid polymer beads with large pores have also been developed as an alternative to CPG. These were developed primarily to overcome cost and supply problems associated with CPG supports. The greater inertness of the polymers relative to CPG, especially during alkaline deprotection conditions, was also an advantage. The first rigid synthetic polymer, introduced by Perkin-Elmer/Applied Biosystems Division (PE/ABD), was a highly cross-linked polysty-
rene support with 1000-Å pores (McCollum and Andrus, 1991). These supports produce excellent quality oligonucleotides, and prepacked ABI LV40 (40 nmol) and ABI LV200 (200 nmol) columns are widely used. The nucleoside capacity of the supports, however, is lower than that of CPG, and prepacked columns >200 nmol are not available. A second rigid polymer based on a polymethacrylate vinyl alcohol copolymer with 1000-Å pores has also been used (Reddy et al., 1994b). This copolymer is sold as a chromatographic medium by both Merck and TosoHaas, respectively, under the trade names Fractogel and Toyopearl. It can be purchased with either hydroxyl functional groups (for size exclusion chromatography) or amino functional groups (for affinity chromatography). These supports are easy to handle, durable, and inexpensive, and their loading capacity is much higher (up to 135 µmol of nucleoside/g) than either rigid polystyrene or CPG supports. Composite macroporous supports Composite supports have been prepared that combine the advantages of gelatinous and rigid supports. These supports are prepared by polymerizing a low cross-linked polyacrylamide inside the pores of a rigid macroporous substrate, such as silica gel, or highly cross-linked polystyrene beads. Typical capacities are between 100 and 500 µmol/g. The soft gelatinous phase is protected by the rigid carrier, and these supports can be used in a continuous flow column system. The supports, however, are fragile, and swelling differences can create unwanted fines. Although this type of support was once used for oligonucleotide synthesis by the phosphotriester method (Gait et al., 1982), present use of commercially available composite supports is limited to other fields.
Grafted Polymeric Carriers Another method of creating hybrid supports, which combine the advantages of gelatinous supports and rigid supports, is to covalently couple or graft long polymeric chains onto the surface of a rigid support. The surface polymers are not cross-linked and are readily solvated, whereas the rigid core remains insoluble. The low thickness of the surface layer and the absence of cross-linking increase the rate of mass exchange and allow a large number of functional sites to be introduced for nucleoside attachment (up to 160 µmol/g). These supports can, therefore, be used in continuous flow column synthesis because the supports are me-
3.1.8 Current Protocols in Nucleic Acid Chemistry
chanically stable, do not show significant swelling, and allow reagents to be removed using short wash steps. The first example of this type of support in oligonucleotide synthesis (by the phosphodiester method) was a polystyrenepolytetrafluoroethylene (PS-PTFE) graft copolymer. This was prepared by 60Co irradiation of PTFE beads and vapor-phase styrene deposition (Potapov et al., 1979). Another coated Teflon support, in the form of fibers not beads, was also commercially available in the 1980s (Bower et al., 1987; Duncan and Cavalier, 1988). More recently, PS-PTFE beads have been found very satisfactory for oligonucleotide synthesis using the phosphoramidite method (Birch-Hirschfield et al., 1996). Another group of widely used graft copolymers are the polyethylene glycol-polystyrene (PEG-PS) tentacle polymers produced by Rapp Polymere under the TentaGel trade name (Rapp, 1996). These supports are prepared by anionic polymerization of ethylene glycol on hydroxyl derivatized cross-linked polystyrene. Copolymers with PEG chains of about 3000 Da are considered optimal, but the polymerization process can produce PEG chains as large as 20,000 Da. Unlike the previous hydrophobic PS-PTFE copolymers, the PEG-PS copolymers have an insoluble polystyrene core and a hydrophilic PEG coating. The relative amounts of material in the coating and the core are also quite different. The PS-PTFE supports have between 2% and 10% polystyrene as the surface coating, whereas the PEG-PS supports have 70% PEG as the coating and only 30% polystyrene as the core. Therefore, the properties of the TentaGel resins are mostly dictated by the PEG coating. The supports swell considerably (3 to 5 times dry volume) in solvents that dissolve PEG, but owing to the insoluble core, they are suitable for both batch and continuous flow processing. The gel-like environment surrounding these supports allows coupling reactions to proceed in a manner similar to solution-phase reactions. This environment presumably allows cyclization reactions to proceed much more efficiently than on CPG supports, and circular oligonucleotides of up to 32 bases have been prepared (Alazzouzi et al., 1997). A number of different TentaGel resins are commercially available with different functional end groups, particle sizes (ranging from uniformly sized 10 µm beads to 750 µm macrobeads), and loading capacities (0.25 to 1.3 mmol/g) for all types of solid-phase and combinatorial synthesis (Winter, 1996). The high
capacity of these supports has found particular use in large-scale (200 to 1000 µmol) oligonucleotide synthesis (Wright et al., 1993).
Nonporous Supports Rigid nonporous beads without surface copolymerization have also been used as supports of oligonucleotide synthesis, although the capacity is two to three orders of magnitude less than similar porous supports. Nonporous silica beads allow long oligonucleotides to be prepared with high coupling yields because of the absence of restrictive pores (Seliger et al., 1989, 1995). The very small diameter (1.5 µm) particles required to provide an acceptable surface loading (2 µmol/g), however, made this support very difficult to work with. Similar handling problems also occurred when nonporous 4.5µm magnetic Dynabeads were used (Albretsen et al., 1990). A more practical application for nonporous supports is the synthesis of immobilized oligonucleotides. Although a large number of methods have been developed to immobilize previously synthesized oligonucleotides on insoluble supports, it is simpler to synthesize the oligonucleotide directly onto the support required for the final hybridization assay. This type of synthesis requires a stable linker that can withstand the conditions used to remove all of the protecting groups (typically 55°C NH4OH, 16 hr) after completion of the synthesis. The deprotected oligonucleotides left attached to the support can then be used as hybridization probes. Both nonporous glass (Maskos and Southern, 1992) and polystyrene beads, with respective loadings of 50 to 70 and 150 nmol/g, have been used. In the latter case, time-resolved fluorescence detection on single Dynosphere beads was performed (Hakala et al., 1997). Hybridization assays using two-dimensional formats, however, are much more common than assays using beads. Consequently, a great deal of effort has gone into the synthesis of oligonucleotide arrays on flat glass and polypropylene supports. The surfaces of glass slides can be derivatized using the same techniques developed for silica gel and CPG supports. Typical surface loadings of 10 to 40 pmol/cm2 are obtained (Maskos and Southern, 1992; McGall et al., 1997), although one account of ∼166 pmol/cm2 was reported when phosphoramidite reagents were reacted directly with surface silanol groups (Cohen et al., 1997). Oligonucleotide synthesized with permanent linkages to quartz fibers has also been used as a DNA sensor (Uddin et al., 1997).
Synthesis of Unmodified Oligonucleotides
3.1.9 Current Protocols in Nucleic Acid Chemistry
Recently, polypropylene sheets have been used as solid-phase supports for oligonucleotide synthesis (Matson et al., 1994, 1995 Wehnert et al., 1994). Polypropylene has the advantages of greater flexibility, physical strength, and chemical stability at high pH. It also has low nonspecific adsorption of biomolecules. One report, however, has mentioned an incompatibility between polypropylene supports and the tetrahydrofuran (THF) solvent commonly used in capping and oxidation reagents (Weiler and Hoheisel, 1996). The inertness of this polymer makes chemical derivatization difficult. Amino-modified supports are prepared by exposure of the polypropylene surfaces to ammonia vapor inside a radio frequency plasma (RFP) generator (Chu et al., 1992). This results in an amino group loading of about 15 to 25 nmol/cm2. Oligonucleotides have been attached to these amino groups, either directly via phosphoramidate linkages (Matson et al., 1994) or through intermediate spacers with terminal hydroxyl (Shchepinov et al., 1997) or amino groups (Weiler and Hoheisel, 1996) to give supports with respective loadings of 10, 0.3, and 0.03 to 0.09 nmol/cm2. Efficient hybridization requires optimization of both loading density and spacer length, because duplex formation can be inhibited by too close spacing and by spacers that are either too short or too long. Polypropylene can also be chemically derivatized without requiring an RFP generator. Bromination using N-bromosuccinimide and 2,2′-azobisisobutyronitrile followed by amination with long-chain diamines or amino alcohols has recently been described, but no surface loadings were reported (Seliger et al., 1995). Oxidation of polypropylene with chromium(VI) oxide followed by borane-tetrahydrofuran complex and H2O2/NaOH treatment has also been used to produce hydroxyl-derivatized polypropylene tapes. Direct reaction of phosphoramidites to this tape yielded a nucleotide loading of 7 nmol/cm2. These polypropylene tapes have been used to prepare a 200-base-long polythymidylic acid sequence and overlapping one-dimensional arrays (Bader et al., 1997a,b).
Filter Disks, Membranes, and Sintered Blocks Solid-Phase Supports for Oligonucleotide Synthesis
This section deals with supports whose physical properties are not easily categorized. Paper filter disks probably represent the cheapest and most readily accessible insoluble support for oligonucleotide synthesis. These sup-
ports contain cellulose fibers that have many hydroxyl groups available for oligonucleotide attachment and that are resistant to all of the chemical conditions required for oligonucleotide synthesis. Paper filter disks were ideal supports for synthesizing multiple oligonucleotides simultaneously using the “segmental solid-phase” procedure, because they could be easily labeled and sorted (Frank et al., 1983; Matthes et al., 1984; Ott and Eckstein, 1984; Frank, 1993). Another important innovation was the development of commercially available MemSyn DNA synthesis supports (Perseptive Biosystems). These contain a membrane-based support made of porous PTFE and coated with an aminopropyl linker. These membranes are sealed inside specially designed low dead volume disposable housings, which resemble common syringe filters, and have loadings of either 50 or 200 nmol. The membranes are easier to handle than are particulate supports, and mass-produced synthesis cartridges are presumably more reliable and easier to manufacture. The large pore diameter of these filter membranes (0.2 µm or 2000 Å) allow both large and small oligonucleotides to be synthesized. Finally a new process, developed at NASA, has been used to derivatize polyethylene (Devivar et al., 1999). In this process, gaseous amine radicals are used to aminate porous polyethylene sintered blocks (FlowGenix, Webster, TX). The amino functionalized sites can then be derivatized with nucleosides for oligonucleotide synthesis. This technology allows supports in the form of plugs and disks to be produced, which should be more convenient to handle than are particulate supports. This discussion clearly indicates that a wide range of insoluble supports have been developed for an increasing number of different applications. Selection of an appropriate support requires consideration of both the chemical properties and the physical characteristics of the support. The size and shape of support beads can vary from small uniformly sized or irregularly sized particles to large macrobeads. Supports in sheet and plate form are amenable to extremely sensitive isotopic or fluorescent detection schemes. High-density arrays or “gene chips” are also emerging as important tools for gene expression studies. Supports in membrane or foam formats provide the synthesis capacity of porous beads but greatly simplify the handling and manufacturing steps required to mass produce ready-to-use synthesis cartridges.
3.1.10 Current Protocols in Nucleic Acid Chemistry
SUPPORT DERIVATIZATION: NUCLEOSIDE AND LINKER ARM COUPLING STRATEGIES Nucleosides are attached to the surface of the support through an intermediate linker arm, whose design must be carefully considered. The linker arm should allow easy nucleoside attachment to the support and be compatible with all of the conditions required for synthesis. Furthermore, the linker must be designed to accommodate different cleavage and deprotection strategies. The many different types of possible linkages can be classified into four groups for strategies that require either: 1.
cleavage from the support with concomitant or postcleavage deprotection;
2.
deprotection of the immobilized products with optional postdeprotection cleavage;
3.
deprotection of the immobilized products with no cleavage from the support; or
4.
linkers that impart terminal end modifications to the oligonucleotide products.
The following section discusses how different linkers have been use in the first three strategies. The use of different linkers and insoluble supports in the synthesis of end-modified oligonucleotides will be discussed in future units.
Linker Arms Cleaved after Synthesis Succinyl linker arm The most commonly used linker arm in oligonucleotide synthesis is succinic acid (Fig. 3.1.3). This linker was used in the early 1970s and has remained very popular because of low cost and ease of incorporation (Yip and Tsou, 1971). Both succinyl dichloride (Sharma et al., 1992) and succinic anhydride have been used as starting materials, but the anhydride is greatly preferred because of its easier handling. A suitably protected 2′-deoxyribonucleoside can be succinylated at either the 5′- or 3′hydroxyl position, and the resulting 5′- or 3′O-hemisuccinate is coupled to an amino- or hydroxyl-derivatized support. Alternatively, the support can be succinylated first and then coupled to a nucleoside (Damha et al., 1990). This method has the advantage of not requiring the synthesis of an inventory of succinylated
nucleosides. Coupling of a nucleoside to a succinylated support, however, is more difficult and usually gives lower nucleoside loadings than does attachment of a presynthesized nucleoside 3′-O-hemiester. The coupling reactions between the succinate and the nucleoside or support have usually been performed using carbodiimide coupling reagents, such as dicyclohexylcarbodiimide (Montserrat et al., 1993), 1-(3-dimethylaminopropyl)ethylcarbodiimide (Pon et al., 1988), and diisopropylcarbodiimide (Bhongle and Tang, 1995), and required coupling times between 1 and 24 hr. A faster coupling reaction—involving reaction of a nucleoside-3′-O-hemisuccinate with 2,2′dithiobis(5-nitropyridine) (DTNP) and dimethylaminopyridine (DMAP) followed by addition of triphenylphosphine (TPP) and LCAA-CPG—can reduce the coupling time to 2 to 30 min (Kumar et al., 1996). Extremely fast coupling of a nucleoside-3′-O-hemisuccinate to LCAA-CPG can be obtained using a variety of phosphonium or uronium coupling agents and DMAP. These reactions are complete in the time required to add the reagent to the support ~4 sec) and allow for the possibility of automated on-line support derivatization (Pon and Yu, 1997b). After completion of the oligonucleotide assembly, the protected products can be cleaved from the support by hydrolysis of the succinyl linker arm with either concentrated aqueous ammonium hydroxide (1 to 2 hr) or gaseous ammonia at 10 bar pressure (15 min; Boal et al., 1996). Faster hydrolysis can be performed by including stronger reagents, such as methylamine (Reddy et al., 1994a) or sodium hydroxide (Chow and Kempe, 1997), with the ammonium hydroxide. These reagents can reduce the cleavage time to 5 min and speed up the removal of base-protecting groups. There are, however, potential problems with the modification of cytosine bases through either aminoalkylation (Macmillan and Verdine, 1991) or deamination (Debear et al., 1987) with these reagents. Although the succinic acid linker has been widely used for a long time, the succinyl linker is unnecessarily stable for oligonucleotide synthesis. The relatively harsh conditions required to hydrolyze the succinyl linker are incompatible with a number of base-sensitive minor bases, backbone modifications, and dye labels; and the time required to cleave the succinyl linker with NH4OH is unnecessarily long (Alul et al., 1991; Avino et al., 1996; Pon and Yu, 1997a). Therefore, a number of more easily
Synthesis of Unmodified Oligonucleotides
3.1.11 Current Protocols in Nucleic Acid Chemistry
DMTO
O
T O2N P
HN
A
O CH2
O
O O
DMTO B
O O
T O2N
O O
O CH2 O
C
O2N
DMTO
D
OMe O H O P N OCE
DMTO
O
N H
P
P
T
O
O
Si O
H N
O
P
O DMTO
E
O
B
O
Si O Si O P O OCE DMTO F
O O
O
H N
O
N H
O
P
B
O
O P O CH2CH2 S CH2 OClPh O
P
Figure 3.1.4 Structure of labile linker arms that can be cleaved under milder conditions than a succinic acid linker. (A) o-Nitrobenzyl carbonate photolabile linker arm (Greenberg and Gilmore, 1994). (B) 5-Methoxy-2-nitrobenzyl carbonate photolabile linker arms (Venkatesan and Greenberg, 1996). (C) o-Nitrophenyl-1,3-propanediol base photolabile linker for 3′-phosphorylated oligonucleotides (Dell’Aquila et al., 1997). (D) Fluoride ion labile diisopropylsilyl linker arm (Routledge et al., 1995). (E) Fluoride ion labile disiloxyl phosphoramidite linker arm (Kwiatkowski et al., 1996). (F) Benzenesulfonylethyl linker arm cleavable with triethylamine/dioxane (Efimov et al., 1983). (G) NPE carbonate linker arm cleavable with DBU/pyridine (Eritja et al., 1991). (H) 9-Fluorenylmethyl linker cleavable with DBU (Avino et al., 1996). (I) Phthaloyl linker arm cleavable with DBU (Avino et al., 1996; Brown et al., 1989). (J) Oxalyl linker, cleavable under very mild conditions (Alul et al., 1991). (K) Malonic acid linker for the synthesis of 3′-phosphorylated oligonucleotides (Guzaev and Lonnberg, 1997). (L) Diglycolic acid linker used to make 3′-TAMRA dye-labeled oligonucleotides (Mullah et al., 1998). (M) Hydroquinone-O,O′-diacetic acid (Q-linker), which can be used for routine oligonucleotides to improve synthesis productivity or to synthesize base-labile products (Pon and Yu, 1997a). DMT, 5′-dimethoxytrityl (DMTr).
cleavable linker arms have been investigated (Fig. 3.1.4).
Solid-Phase Supports for Oligonucleotide Synthesis
Labile linker arms Photolysis offers a very mild method for cleavage. Photolabile linker arms based on onitrobenzyl groups (Greenberg and Gilmore,
1994) have been used to synthesize oligonucleotides with 3′-hydroxyl (Fig. 3.1.4A and B), 3′-phosphate (Fig. 3.1.4C), and other 3′-end modifications. The photolysis can, however, cause small amounts (<3%) of thyminethymine photodimers, and alkaline or other conditions still need to be employed to remove
3.1.12 Current Protocols in Nucleic Acid Chemistry
DMTO
B
O O
G
DMTO
NO2
H
DMTO
N H
O
O
P
B
J O
HN
NH
O
O
H N O
P
L
O
DMTO M
O
B
O
O
RO
O O
P
O
O
DMTO
K
H N
O
O
P
O
HN
O
O O
I
B
O
RO
P
H N
O O
P
O
B
O O
H N
O
P
O
Figure 3.1.4 Continued
base-protecting groups. In addition, N-benzoyl-protected dA and dC nucleosides must also be avoided. Very mild cleavage, under non-nucleophilic conditions and neutral pH, can be obtained through silyl- or disiloxyl-based linker arms (Fig. 3.1.4D and E), which are cleavable with fluoride ion. Triethylamine has been used to cleave a benzylsulfonylethanol linker arm to yield 3′-phosphorylated oligonucleotide blocks suitable for solution-phase coupling (Fig. 3.1.4F). An even more labile 2-(4-carboxyphenylsulfonyl)ethanol linker arm was considered unsuitable for phosphotriester synthesis but was not evaluated using phosphoramidite synthesis (Schwyzer et al., 1984). The non-nucleophilic base, 1,8-diazabicyclo[5.4.0]-undec-7-ene (DBU), can also be used to cleave the 2-(o-nitrophenyl)ethyloxycarbonyl (NPE) linker, N-[9-(hydroxymethyl)-2fluorenyl]-succinamic acid (Fmoc) linker, and phthaloyl linker arms (Fig. 3.1.4G to I). When using DBU cleavage, however, thymine and
guanine modification can occur if methoxy- or cyanoethyl phosphate–protecting groups are not removed before the DBU treatment. Furthermore, oligonucleotides with terminal TT sequences are not efficiently cleaved. All of the above linker arms were either difficult to prepare or did not offer any speed advantage. In addition, the requirement for deprotection conditions or reagents different from the simple NH4OH cleavage used in standard oligonucleotide synthesis procedures was an obstacle to the widespread adoption of any of these linker arms. More satisfactory alternatives would be other dicarboxylic acid linkers, especially if they were readily available and compatible with the derivatization and cleavage methods used for succinic acid. The most labile dicarboxylic acid linker reported has been the oxalyl linker (Fig. 3.1.4J). This was completely cleaved by concentrated NH4OH in only a few seconds, and cleavage with a number of other milder reagents was also possible. The oxalyl linker, however, was too
Synthesis of Unmodified Oligonucleotides
3.1.13 Current Protocols in Nucleic Acid Chemistry
Solid-Phase Supports for Oligonucleotide Synthesis
labile for routine use, and oxalyl derivatized supports had to be used within a few weeks because significant spontaneous nucleoside loss occurred (Pon and Yu, 1997a). More stable linkages have been created using either malonic acid or diglycolic acid as the linker arm (Fig. 3.1.4K and L). Treatment of diglycolic acid (Pon and Yu, 1997a) and malonic acid (Guzaev and Lonnberg, 1997) linkers with room temperature concentrated NH4OH for 10 min was sufficient to respectively hydrolyze 68% and 90% of these linker arms, conditions that caused only 15% cleavage of the succinyl linker. The malonic acid linker arm was used in combination with a diethyl 2,2-bis(hydroxymethyl)malonate spacer to afford 3′phosphorylated methyl phosphotriester and methylphosphonate analogues. The diglycolic acid linker has principally been used in combination with a branching spacer, such as 2amino-1,3-propanediol, to prepare 3′-tetramethylrhodamine (TAMRA) labeled oligonucleotides, which are damaged by conventional ammonium hydroxide hydrolysis (Mullah et al., 1998). In this case t-butylamine/methanol/water (1:1:2) was used for cleavage from the support (20 to 60 min at room temperature) and subsequent base deprotection (1 to 3 hr at 65° to 85°C). A more satisfactory replacement for succinic acid is hydroquinone-O,O′-diacetic acid, which is used to create a Q-linker arm (Fig. 3.1.4M). This linker is sufficiently stable so decomposition during room temperature storage is not a problem. The Q-linker, however, can be cleaved much faster than either the succinyl or diglycolic acid linkers (Pon and Yu, 1997a). For example, cleavage using NH4OH required only 2 to 3 min and cleavage using t-butylamine/methanol/water was performed in only 5 min, instead of the 45 min described above. Moreover, for routine use, supports derivatized with the Q-linker can be used without any modifications to either protecting groups, reagents, or synthesis procedures (other than a reduction in cleavage time). Thus the Q-linker can serve as a general replacement for the succinyl linker in the synthesis of either unmodified or base-sensitive oligonucleotides. The main advantage of the Q-linker, however, is the improved productivity that results from the decreased cleavage time. Unlike postsynthesis deprotection, which is performed off the automated synthesizer, the cleavage step is usually performed by the instrument; and subsequent runs cannot be started until the cleavage is complete. Because typical oligonu-
cleotide syntheses are usually complete within 2 hr, waiting an additional 1 or 2 hr for cleavage of a succinyl linker represents a significant bottleneck. Recently, the Q-linker arm has also been included in a strategy for oligonucleotide synthesis on reusable solid-phase supports (Pon et al., 1998, 1999). In this approach, an hydroxyl derivatized support is used to form ester linkages, which can be easily cleaved and regenerated for subsequent use. The mild cleavage conditions required to release the oligonucleotide reduce damage to the support’s surface and reduce the time required to prepare the support for another use. This approach will be very useful in the large-scale (>1000 kg/y) manufacturing of oligonucleotide pharmaceuticals in which the support is the most expensive single consumable. Universal linkers In the supports discussed above, attachment of the first nucleoside is always done separately from the actual oligonucleotide synthesis, because of the different chemistry and long coupling times required. In the past, maintaining an inventory of prederivatized supports was not problematic because of the limited number of common nucleosides. The recent development of high throughput DNA synthesizers. however, has created a need for universal supports that have the terminal nucleoside added as part of the automated synthesis. This is required not so much for inventory purposes but because manual setup of prederivatized supports is time-consuming and error prone. Universal linkers are also an advantage for the synthesis of oligonucleotides containing rare or modified bases that one wishes to incorporate at internal sites and at the 3′-end. In this way it is necessary to synthesize or purchase only the phosphoramidite derivative of the rare or modified base. Automation of the nucleoside-coupling reaction using very fast uronium coupling reagents and DMAP is one possible approach (Pon and Yu, 1997b); however, implementation of this strategy requires construction of new DNA synthesizers with additional reagent reservoirs. A simpler approach would be to design a linker arm that could use a conventional nucleoside-3′-phosphoramidite as the first monomeric unit. It is fairly simple to design universal supports that can produce oligonucleotides with 3′-phosphate ends using either amino or hydroxyl end functions (Fig. 3.1.4A to F). Furthermore, a sulfonyldiethanol phos-
3.1.14 Current Protocols in Nucleic Acid Chemistry
a neighboring hydroxyl group so a cyclic phosphate can form via an intramolecular attack (Fig. 3.1.6). A linker containing a single ribonucleoside in an inverted orientation, so chain extension occurs from the 2′- (or 3′-) hydroxyl position and not the 5′-position, will allow cleavage of the phosphate group via a mechanism similar to the alkaline cleavage of RNA. This will produce the desired oligonucleotide with a 3′-hydroxyl terminus and a uridine-2′, 3′-cyclic phosphate. This strategy was first used on cellulose supports (Crea and Horn, 1980; van der Marel et al., 1982); however, because dinucleotide units with inverted 3′-2′ (3′) linkages were
phoramidite, usually used for 5′-phosphorylation, can be used to synthesize 3′-phosphorylated sequences on any amino- or hydroxylderivatized support (Fig. 3.1.5G). Obtaining an oligonucleotide with a free 3′-hydroxyl terminus instead of a 3′-phosphate is essential, however, if the oligonucleotides are to be enzymatically extended (e.g., used as DNA sequencing or polymerase chain reaction primers). Removal of the terminal 3′-phosphate group introduced by the 3′-phosphoramidite reagents is usually difficult, because the negative charge(s) on phosphodiester and phosphomonoester bonds make them very resistant to hydrolysis. This difficulty can be overcome by introducing
O A
B
C
N H
NH2
HO
H N
O S O
Si
P
O
O Si
P
CH3
DMTO
S S
DMTO
S S
Si
P
O D
O
N H
O O
H N
P
P
E HO NO2
DMTO
O
F
G
N H
DMTO
O
O
S O
O P O OCE
P
P
Figure 3.1.5 Structure of universal linkers for 3′-phosphorylated oligonucleotides (see Fig. 3.1.4C and K). (A) Benzidine linker arm (Markiewicz and Wyrzykiewicz, 1989). (B) Hydroxyethylsulfonyl linker arm (Markiewicz and Wyrzykiewicz, 1989). (C) Hydroxyethyl disulfide linker (Kumar et al., 1991). (D) Hydroxyethyl disulfide linker (Asseline and Thuong, 1989; Gupta et al., 1991). (E) NPE linker (Eritja et al., 1991). (F) Universal allyl linker, 9-O-(4,4′-dimethoxytrityl)-10-undecenoic (Zhang and Jones, 1996). (G) Dimethoxytrityl sulfonyldiethanol phosphoramidite linker (Bader et al., 1997b; Shchepinov et al., 1997). DMT, 5′-dimethoxytrityl (DMTr).
Synthesis of Unmodified Oligonucleotides
3.1.15 Current Protocols in Nucleic Acid Chemistry
oligonucleotide
oligonucleotide
O
B
O O O
O O
O
O
O
B
O
O
1
O
NCCH2CH2O P O
oligonucleotide
2
O
O P O R
U
OH
O
HO
O
O P
O O
O O
HO
U
B
O
U
O
O P
NH
3
O
oligonucleotide
O P
O
O HO
O
O
B
HO
O
U
Figure 3.1.6 Mechanism of terminal phosphate cleavage in universal supports. (1) The first step in the cleavage/deprotection process is hydrolysis of the ester from the hydroxyl group adjacent to the first phosphate linkage. This step occurs concurrently with the removal of the cyanoethyl groups on the phosphate linkages and hydrolysis of the ester attachment to the support. (2) The deprotected hydroxyl group can then cyclize by attacking the phosphorus atom. (3) Formation of a 2′,3′-cyclic phosphodiester releases the oligonucleotide sequence with a free 3′-OH group.
Solid-Phase Supports for Oligonucleotide Synthesis
prepared in solution before attachment to the support the method was not truly universal (Fig. 3.1.7A). The universal support concept was first fully examined when uridine mononucleosides were attached to CPG through 5′-succinate linkages (Fig. 3.1.7Ba to c). Oligonucleotide synthesis, using nucleoside-3′-phosphoramidites, can then be performed from the 2′ (or 3′) hydroxyl position of the uridine linker in the normal manner. Cleavage from the linker, however, involves two steps: hydrolysis of the succinyl linker to release the material from the support and elimination of the terminal uridine as the 2′,3′-cyclic phosphate. Although both steps, along with removal of base-protecting groups, can be performed simultaneously, elimination of the terminal cyclic phosphate is the rate-limiting step. After normal NH4OH deprotection, treatment at neutral pH with lead acetate (18 hr at 37°C) can complete the terminal deblocking (Gough et al., 1983). Complete deblocking can be performed with extended NH4OH hydrolysis, although the rate depends on the nature of the adjacent nucleoside with all ribonucleosides > dA, dG > T > dC (Debear et al., 1987). Therefore, NH4OH deprotection conditions ranging from 16 hr at 50°C to 24 hr at 65°C were first proposed, and a sub-
sequent paper has used 48 hr at 65°C (Schwartz et al., 1995). Attachment of a ribonucleotide through the N4-position of a cytosine base (Fig. 3.1.7C), with subsequent chain extension from the 3′-phosphate group, has also been used (Pochet et al., 1987). In this case, cleavage was performed using 2N NaOH (10 min at 60°C); however, use of alkali hydroxides is not recommended because of possible damage to cytosine bases. A reversed uridine phosphoramidite reagent can also be used to convert previously derivatized supports into universal supports (Fig. 3.1.7D). Universal supports with N-benzoylcytidine linkers (Fig. 3.1.7Bd) are commercially available (Biogenex, San Ramon, CA), and the addition of 0.5 M lithium chloride to the NH4OH reagent has been recommended for their cleavage and deprotection (15 hr at 55°C). A ribonucleoside is not essential for a universal linker, and other diol linkers have been used. An acyclic 3-amino-1,2-propanediol linker has been reported (Lyttle et al., 1996) that uses neighboring group participation by the amino group to cleave the oligonucleotide under mild conditions (0.1 M triethylamine acetate [1 mL] plus 3% NH4OH [40 µL], 2 hr at room temperature; Fig. 3.1.7E). This linker,
3.1.16 Current Protocols in Nucleic Acid Chemistry
DMTO
B
O
OR2
R1O O O P OR1 OR2 O
A
O P
P
O
O P O
O
B
U OR3 Aa, R1 = R3 = 4-chlorophenyl, R2 = acetyl Ab, R1 = 2-chlorophenyl, R2 = methoxytetrahydropyranyl, R3 = 4-t-butyl-2-chlorophenyl
Ba, B = uracil, R1 = H, R2 = benzoyl Bb, B = uracil, R1 = H, R2 = acetyl 3 Bc, B = N -anisoyluracil, R1 = H, R2 = benzoyl 4 Bd, B = N -benzoylcytosine, R1 = acetyl, R2 = DMT Be, B = H, R1 = DMT, R2 = chloroacetyl
O O C
DMTO
O O
O P O
C
O C O
AcO
O
O D
N
U
O P OCE O
O
O
B O
N O
P
HN
N H
O O
O DMTO
N H
E
F
H N
P
O
O P O OCE
P
O
O
O
O O
B
O
Cl
O
O
O
N H
O O
O O
O
O P O OCE
O O G
DMTO
H N
O
P
O
Figure 3.1.7 Universal linker arms for the synthesis of oligonucleotides with free 3′-OH ends. (A) First 3′-2′(3′) inverted linkages (Crea and Horn, 1980; van der Marel et al., 1982). (B) Universal supports based on: (a) 2′(3′)-O-benzoyluridine (Gough et al., 1983); (b) 2′(3′)-O-acetyluridine (Cosstick and Eckstein, 1985); (c) N3-anisoyluridine (Debear et al., 1987); (d) commercially available N4-benzoylcytidine universal support (Biogenex, San Ramon, CA); (e) 1,4-anhydroribitol (Scheuerlarsen et al., 1997). (C) Inverted linkage attached through N4-position of cytidine (Pochet et al., 1987). (D) Inverted linkage obtained through use of a “universal” 5′-phosphoramidite (Schwartz et al., 1995). (E) Acyclic universal linker with a 3-N-allyloxycarbonyl-protecting group (Lyttle et al., 1996). (F) Special phosphoramidite linkage to allow the synthesis of a second oligonucleotide on the 5′-end of another (Hardy et al., 1998). (G) 1,4-Anhydroerythreitol coupled via a succinyl linker to CPG (Nelson et al., 1997). DMT, 5′-dimethoxytrityl (DMTr).
however, required a preliminary step with tetrakis(triphenylphosphine) palladium to remove the N-allyloxycarbonyl-protecting group, and not all of the oligonucleotide was released from the support. The rate of cleavage has been found to depend strongly on the stereochemistry of the attacking hydroxyl group, as reported in studies
that used the cyclic cis diol 1,4-anhydroerythritol (Scott et al., 1994; Hardy et al., 1998). These studies developed a novel linker, phosphoramidite, that allowed consecutive oligonucleotide sequences to be synthesized on the same support (Fig. 3.1.7F). Cleavage with NH4OH (16 hr at 80°C) or NH4OH/MeNH2 (16 hr at 60°C) was recommended. The 1,4-anhy-
Synthesis of Unmodified Oligonucleotides
3.1.17 Current Protocols in Nucleic Acid Chemistry
droerythritol linker has also been coupled to CPG supports through a succinic acid linker (Fig. 3.1.7G); these Rainbow Universal CPG supports are commercially available (Clontech, Palo Alto, CA). Deprotection and cleavage from the Rainbow Universal CPG supports was performed using either 0.5 M LiCl/NH4OH (16 hr at 55°C) or 0.23 M triethylamine/0.5 M LiCl/NH4OH (1 hr at 80°C) as the cleavage reagent (Nelson et al., 1997). Deprotection of the hydroxyl group adjacent to the phosphate linkage is a requirement before this group can participate in the cyclization and phosphate group elimination reactions. Therefore, faster deprotection can be obtained when the very labile chloroacetyl-protecting group is used instead of acetyl, benzoyl, or succinyl groups (Scheuerlarsen et al., 1997). This strategy was first applied to a 1,4-anhydroribitol linker arm using a specific 2′-5′ linkage (Fig. 3.1.7Be), and cleavage of the terminal cyclic phosphate was achieved under “normal” deprotection conditions (NH4OH for either 12 hr at 55°C or 72 hr at 22°C).
Linker Arms for the Deprotection of Immobilized Products with Optional Postdeprotection Cleavage
Solid-Phase Supports for Oligonucleotide Synthesis
A number of applications require the removal of all or some of the protecting groups before the oligonucleotide is cleaved from the support (i.e., deprotection conditions orthogonal to cleavage conditions). Removal of the terminal 4′-dimethoxytrityl-protecting group is easily performed because the acidic conditions do not affect the acid-resistant linker arms most commonly used. In contrast, removal of the phosphate-protecting groups from the internucleotide linkages or the amino-protecting groups from the adenine, cytosine, and guanine bases requires special consideration if these groups are to be removed without cleaving the product from the solid-phase support. If, however, these deprotection steps are performed while the product remains immobilized (i.e., attached to the support), then the solid-phase support can provide the same handling and workup advantages as realized during the solidphase synthesis. After these steps, the linker arm can be cleaved to release the products. In certain cases, the final product will be used while immobilized on the support, but it is still helpful to be able to cleave samples off for characterization and quality analysis. Generally, succinyl linkers are not sufficiently stable to allow removal of any protecting groups other than O-methyl- or cyanoethyl
phosphate–protecting groups. Succinyl linkers, however, can be left intact if different protecting group and deprotection schemes are employed. For example, hydrazine hydrate/pyridine/acetic acid can be used as the deprotection reagent if the N6-isobutyryl-protecting group on deoxyguanosine is replaced with an N6(N′,N′-dibutylformamidine)-protecting group (Urdea and Horn, 1986). Base deprotection using ethanolamine can also be accomplished, but the N4-protection on deoxycytidine must be modified to prevent N4-hydroxyethylation (Berner et al., 1989). Allylic protection on all of the bases and on the phosphate linkages can also be removed using palladium reagents without affecting the succinyl linker (Hayakawa et al., 1990). Attachment of a succinyl linker to a secondary N-methyl amino group on an intermediate linker such as N-methyl glycine (sarcosine), bis-1,6-dimethylaminohexane, or N-propyl polyethylene glycol (Fig. 3.1.8A to C) instead of a primary amino group (such as LCAACPG) creates linkages that are resistant to cleavage by the non-nucleophilic base DBU. These linkages have been used with base labile 5′-Fmoc-protecting groups to make acid-sensitive oligodeoxyribonucleotides and oligoribonucleotides (Brown et al., 1989) or with 2-(4-nitrophenyl)ethoxycarbonyl/NPE baseprotecting groups to allow on-column deprotection (Stengele and Pfleiderer, 1990; Weiler and Pfleiderer, 1995). A triethylamine-resistant sarcosine-succinic acid linker arm (Fig. 3.1.8D) has also been used to prepare branched RNA and DNA/RNA chimeras (Grotli et al., 1997). In each case, after synthesis or deprotection, the products are easily released from the support by conventional hydrolysis with NH4OH. A stronger linkage can also be created by attaching the succinyl linker to the amino group of a base instead of a hydroxyl group (Figs. 3.1.7C and 3.1.8E and F). Alkaline hydrolysis or oxidative cleavage with NaIO4 can then release the product. This strategy, however, requires two orthogonal-protecting groups for the 5′- and 3′(2′)-positions and does not offer any advantage, except for the synthesis of cyclic oligonucleotides. A number of different linker arms have also been developed that take advantage of the resistance of phosphate ester and amide bonds to alkaline hydrolysis and allow protecting group removal before cleavage from the support. Thus thioether and thiophosphate linkers cleavable by oxidative cleavage (Fig. 3.1.8G and H), allyl
3.1.18 Current Protocols in Nucleic Acid Chemistry
DMTO
O
B O
A
O
N CH3 O
O DMTO
O
H N
P
B O
B
O
N CH3
O DMTO
O
CH3 N O
P
O
B O
O
C
O
N Pr
O
P
O CH2
n n = 40-70 O O O
D
O
O O
H N
P
O O
N CH3
O
O
O
H N
HN
DMTO
O
N
P H N
O
N E
B
HN O
N
F DMTO
O O P O OCE
O
N
N
O
NH O
N
OAc OAc Cl P
Figure 3.1.8 Linker arms that allow on-column deprotection and then optional cleavage. (A) Succinic acid linked to an N-methylglycine (sarcosine) derivatized support (Brown et al., 1989). (B) Succinic acid linked to 1,6-bis methylaminohexane spacer (Stengele and Pfleiderer, 1990). (C) Succinic acid linked to N-propyl polyethylene glycol Tentagel support (Weiler and Pfleiderer, 1995). (D) Succinyl-sarcosine linkage for the solid-phase synthesis of branched oligonucleotides (Grotli et al., 1997). (E) Linkage through the amino group of cytosine for branched and cyclic oligonucleotide synthesis (De Napoli et al., 1995). (F) Oxidizable solid support (Bower et al., 1987; Markiewicz et al., 1994). (G) Phenyl thioether linker, which is stable until oxidized into a phenylsulfone (Felder et al., 1984). (H) Thiophosphate linker, cleavable by iodine/water oxidation or acetic acid hydrolysis (Tanaka et al., 1989). (I) 3-Chloro-4-hydroxyphenyl linker for the solid-phase synthesis of cyclic oligonucleotides (Alazzouzi et al., 1997). (J) Linker arm produced from tolylene 2,6-diisocyanate with more stable carbamate and urethane linkages (Kumar, 1994; Sproat and Brown, 1985). DMT, 5′-dimethoxytrityl (DMTr).
linkers cleavable with tetrakis(triphenylphosphine) palladium (Fig. 3.1.5F), and phosphoroamidate linkages cleavable by acidic hydrolysis have been developed (Gryaznov and
Letsinger, 1992). These supports can also be considered as universal supports because the first nucleotide is added as part of the automated synthesis. All of these methods, how-
Synthesis of Unmodified Oligonucleotides
3.1.19 Current Protocols in Nucleic Acid Chemistry
DMTO
T
O
G O O P OCE O
H
O S
O O P S O
C HN
O
H N
N H
T
O
O
O
I
P
O
Cl
DMTO
P
N H
O P O OCE
P
Cl
DMTO
B
O
J
H N
O O
H N
H N
P
O
Figure 3.1.8 Continued
Solid-Phase Supports for Oligonucleotide Synthesis
ever, produced oligonucleotides with a terminal 3′-phosphate instead of a 3′-hydroxyl group. An exception was the solid-phase synthesis of cyclic oligonucleotides. In this approach, 3chloro-4-hydroxylphenylacetic acid served as both the linker arm and a phosphate-protecting group (Fig. 3.1.8I). The phosphotriester linkage was converted into a phosphodiester by selective removal of the cyanoethyl-protecting group, and then cyclization of the 5′-end to the 3′-terminal phosphodiester group was achieved using 1-mesitylenesulfonyl-3-nitro-1,2,4-triazole as the coupling agent. The linker was then cleaved with tetramethylguanidinium syn-2-aldoximate (8 hr), and a circular oligonucleotide was released (Alazzouzi et al., 1997). The requirement for a 3′-phosphate has also been avoided by insertion of a disiloxyl linkage between the nucleoside and the phosphate linkage (Fig. 3.1.4E). This linkage was stable to mild base, and preliminary purification by selective cleavage of apurinic sites using triethylamine/ethanol (1:1, 3 hr at 20°C), was possible
(Kwiatkowski et al., 1996). After this treatment, the products were cleaved from the support with tetrabutylammonium fluoride and then deprotected with NH4OH. Isocyanate reagents can react with hydroxyl groups and amino groups to produce carbamate and urethane bonds, respectively, which are more resistant to hydrolysis than are esters. Although acyclic diisocyanates have not been successful, the more rigid tolylene 2,6-diisocyanate reagent (Sproat and Brown, 1985) has been used to create a stable linker arm (Fig. 3.1.8J). The carbamate linkage can be hydrolyzed by long exposure to NH4OH (24 to 48 hr at 55°C), whereas deprotection under milder conditions allowed the product to remain attached to the support. Two methods have been developed for removing failure or depurinated sequences from the full-length product while both are still attached to the support. In the first method (Urdea and Horn, 1986), spleen phosphodiesterase was used to selectively degrade non-full-length oli-
3.1.20 Current Protocols in Nucleic Acid Chemistry
A
O
H N
HO
DMTO
B
P
N H
P
O C
DMTO O
DMTO
D
O
O
O
B
a, R = b, R =
O O O P O R O P
H N
P
O n CE
O CE
n = 0-30 (8-10 optimum)
P
N CH2 H
c, R =
(CH2)3 CH2CH2 O CH2CH2 CH2CH2 O CH2CH2 O CH2CH2
d, R =
CH2 CH CH2
e, R =
CH2 CH CH2 NH O
NH3
NH3 DMTO
E
O
B
O
O
O P O O
S O
CE
DMTO F
O
O O H O P O R O P N O O n CE CE
P
n = 0-30 (8-10 optimum)
B
O O P O Si OCE
P
Figure 3.1.9 Linker arms for permanent attachment to solid-phase supports. (A) Hydroxy propylamine linker (Seliger et al., 1995). (B) Dimethoxytrityl glycolic acid linker (Hakala et al., 1997). (C) Dimethoxytrityl-4,7,10,13-tetraoxatridecanoate linker (Markiewicz et al., 1994). (D) Long spacer linkages prepared using repetitive coupling of various phosphoramidites (Shchepinov et al., 1997). (E) Cleavable spacer linkage used in conjunction with the preceding phosphoramidites to control the surface oligonucleotide density (Shchepinov et al., 1997), (F) Direct phosphate linkage to surface silanol groups (Cohen et al., 1997). (G) Diol linker formed from 3-glycidoxypropyl trimethoxysilane (Maskos and Southern, 1992). (H) Polyethylene glycol linkers (Maskos and Southern, 1992). (I) Bis-(2-hydroxethyl)-aminopropylsilane linker with hexaethylene glycol spacer phosphoramidites (Pease et al., 1994). (J) N-(3-(triethoxysilyl)-propyl)-4-hydroxybutyramide linker (McGall et al., 1997). (K) Linkage through the N4-position of cytosine (Markiewicz et al., 1994). (L) Triethylene glycol ethylacrylamide linker (Markiewicz et al., 1994). DMT, 5′-dimethoxytrityl (DMTr).
gonucleotides. This method required a careful selection of protecting groups and a special capping reagent (levulinic anhydride) that could be selectively removed without exposing the full-length product (blocked by a DMTr or 5′-benzoyl group) to enzymatic digestion. In a second approach (Horn and Urdea, 1988; Kwiatkowski et al., 1996), the purification of 5′-dimethoxytritylated oligonucleotides was
improved by selective fragmentation of sequences containing apurinic defects, while still attached to the support. This treatment greatly reduced the proportion of DMTr-labeled oligonucleotides that were not full-length and allowed oligonucleotides as long as 118 bases to be purified by a simple reverse-phase cartridge procedure.
Synthesis of Unmodified Oligonucleotides
3.1.21 Current Protocols in Nucleic Acid Chemistry
OH G
HO
HO H
O
P
Si
O
O
O
Si
n
n = 0, 4, 5
O O P O 6 OCE
DMTO I
P
N
Si
P
O O P O OCE 6
DMTO
O J
HO
N H
P
Si
Si
HN
P
NH K DMTO
O
N
O
AcO
O L
DMTO
O
O
O
N H
Si
P
Figure 3.1.9 Continued
Linker Arms for Permanent Attachment to the Support
Solid-Phase Supports for Oligonucleotide Synthesis
Linker arms for permanent attachment of the oligonucleotide to the support must be very resistant to the hydrolysis conditions used to remove protecting groups. The most common method for permanent attachment is through phosphodiester or phosphoroamidate linkages by the direct coupling of phosphoramidite reagents, respectively, to hydroxyl or amino derivatized supports. The phosphoramidate linkages are particularly common when aminated polypropylene films are used as supports (Matson et al., 1994, 1995; Wehnert et al., 1994; Shchepinov et al., 1997). Other derivatized supports have been converted into hydroxyl functionalized materials using either 3-amino-1propanol, dimethoxytrityl glycolic acid, sodium 13-O-dimethoxytrityl-4,7,10,13-tetra-oxatrid ecanoate, or a variety of consecutive spacer
phosphoramidites (Fig. 3.1.9A to D). These spacer phosphoramidites may also contain cleavable sulfonyldiethanol groups (Fig. 3.1.9E) or positively charged 2-amino-1,2propandiol groups so that the surface density and charge can be modulated as well as the spacer chain length. Although a recent publication has described the direct coupling of phosphoramidite reagents to the silanol groups on acid-washed glass slides (Fig. 3.1.9F), most glass surfaces are derivatized with alkoxysilanes to produced linker arms extending away from the surface. Several hydroxy-derivatized linkers on glass based on 3-glycidoxypropyl trimethoxysilane and ethylene glycol ethers have been described (Fig. 3.1.9G and H). Unfortunately, however, these linkers were not completely resistant to NH4OH (5 to 10 hr at 55°C), and much of the product was lost. Other hydroxyl linker arms on glass plates have been based on bis-(2-hy-
3.1.22 Current Protocols in Nucleic Acid Chemistry
droxyethyl)-aminopropylsilane or N-(3(triethoxysilyl)-aminopropyl)-4-hydroxybutyramide and used in the synthesis of high-density oligonucleotide arrays (Fig. 3.1.9I and J). In these reports, base deprotection was accomplished using 1,2-diaminoethane/ethanol (1:1, 2 to 6 hr at room temperature) instead of hot NH4OH, and no linker cleavage was reported. Linkages to CPG beads using attachments through the N4-position of 2′-deoxycytidine or through a triethylene glycol ethylacrylamide linker have also been reported (Fig. 3.1.9K and L); however, the researchers noted that the lability of the disiloxane bond (Si-O-Si) between any silica-based support and silane linker arm is a limiting factor and, although certain resistance to aqueous NH4OH is possible, use of pyridine/NH4OH always results in substantial cleavage (Markiewicz et al., 1994).
CONCLUSIONS Solid-phase oligonucleotide synthesis was once considered to be a “mature” technology limited to incremental improvements; however, new and ingenious applications for oligonucleotide based materials continue to be developed, and the role of solid-phase synthesis is clearly going to be very important in making these new materials available. Quite remarkably, the scale of these applications spans the range from the extremely minute (i.e., single molecule detection and nanoengineering) to large-scale oligonucleotide pharmaceuticals. Success in any of these areas requires a firm understanding of the chemical requirements of every step involved. It is hoped that this short review provides enough introduction to convey the power and diversity of solid-phase oligonucleotide synthesis techniques and convince the reader that new technology can be developed in even mature fields.
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Synthesis of Unmodified Oligonucleotides
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Sproat, B.S. and Brown, D.M. 1985. A new linkage for solid phase synthesis of oligodeoxyribonucleotides. Nucl. Acids Res. 13:2979-2987.
Routledge, A., Wallis, M.P., Ross, K.C., and Fraser, W. 1995. A new deprotection strategy for automated oligonucleotide synthesis using a novel silyl-linked solid support. Bioorgan. Med. Chem. Lett. 5:2059-2064. Scheuerlarsen, C., Rosenbohm, C., Jorgensen, T.J.D., and Wengel, J. 1997. Introduction of a universal solid support for oligonucleotide synthesis. Nucleos. Nucleot. 16:67-80. Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., and Gough, G.R. 1995. A universal adapter for chemical synthesis of DNA or RNA on any single type of solid support. Tetrahedron Lett. 36:27-30. Schwyzer, R., Felder, E., and Failli, P. 1984. 148. The CAMET and CASET links for the synthesis of protected oligopeptides and oligodeoxynucleotides on solid and soluble supports. Helv. Chim. Acta 67:1316-1327. Scott, S., Hardy, P., Sheppard, R.C., and McLean, M.J. 1994. A universal support for oligonucleotide synthesis. In Innovation and Perspectives in Solid-Phase Synthesis. Peptides, Proteins, and Nucleic Acids, Biological and Biomedical Applications (R. Epton, ed.) pp. 115124. Mayflower Worldwide, Ltd., Birmingham, UK. Seliger, H., Herold, A., Kotschi, U., Lyons, J., and Schmidt, G. 1987. Semi-mechanized simultaneous synthesis of multiple oligonucleotide fragments. In Biophosphates and Their Analogues— Synthesis, Structure, Metabolism and Activity (K.S. Bruzik and W.J. Stec, eds.) pp. 43-58. Elsevier Science Publishers, New York.
Stengele, K.P. and Pfleiderer, W. 1990. Improved synthesis of oligodeoxyribonucleotides. Tetrahedron Lett. 31:2549-2552. Stimpson, D.I., Hoijer, J.V., Hsieh, W.T., Jou, C., Gordon, J., Theriault, T., Gamble, R., and Baldeschwieler, J.D. 1995. Real-time detection of DNA hybridization and melting on oligonucleotide arrays by using optical wave guides. Proc. Natl. Acad. Sci. U.S.A. 92:6379-6383. Tanaka, T., Yamada, Y., Uesugi, S., and Ikehara, M. 1989. Preparation of a new phosphorylating agent: S-(N-monomethoxytritylaminoethyl)-O(o-chlorophenyl)phosphorothioate and its application in oligonucleotide synthesis. Tetrahedron 45:651-660. Tang, J.Y. and Tang, J.X. 1997. Passivated polymer supports for nucleic acid synthesis. United States Patent #5,668,268. Uddin, A.H., Piunno, P.A., Hudson, R.H., Damha, M.J., and Krull, U.J. 1997. A fiber optic biosensor for fluorimetric detection of triple-helical DNA. Nucl. Acids Res. 25:4139-4146. Urdea, M.S. and Horn, T. 1986. Solid-supported synthesis, deprotection and enzymatic purification of oligodeoxyribonucleotides. Tetrahedron Lett. 27:2933-2936. van Aerschot A., Herdewijn, P., and Vanderhaeghe, H. 1988. Silica gel functionalised with different spacers as solid support for oligonucleotide synthesis. Nucleos. Nucleot. 7:75-90. van der Laan, A.C., Brill, R., Kuimelis, R.G., Kuylyeheskiely, E., Vanboom, J.H., Andrus, A., and Vinayak, R. 1997. A convenient automated solid-phase synthesis of PNA-(5′)-DNA-(3′)PNA chimera. Tetrahedron Lett. 38:2249-2252.
Synthesis of Unmodified Oligonucleotides
3.1.27 Current Protocols in Nucleic Acid Chemistry
van der Marel, G.A., Marugg, J.E., de Vroom, E., Wille, G., Tromp, M., van Boeckel, C.A.A., and van Boom, J.H. 1982. Phosphotriester synthesis of DNA fragments on cellulose and polystyrene solid supports. Recl. Trav. Chim. Pays-Bas 101:234-241. Venkatesan, H. and Greenberg, M.M. 1996. Improved utility of photolabile solid phase synthesis supports for the synthesis of oligonucleotides containing 3′-hydroxyl termini. J. Org. Chem. 61:525-529. Wehnert, M.S., Matson, R.S., Rampal, J.B., Coassin, P., and Caskey, C.T. 1994. A rapid scanning strip for tri- and dinucleotide short tandem repeats. Nucl. Acids Res. 22:1701-1704. Weiler, J. and Hoheisel, J.D. 1996. Combining the preparation of oligonucleotide arrays and synthesis of high-quality primers. Anal. Biochem. 243:218-227. Weiler, J. and Hoheisel, J.D. 1997. Picomole syntheses of high quality oligonucleotide primers in combination with the preparation of oligonucleotide arrays. Nucleos. Nucleot. 16:17931796.
Weiler, J. and Pfleiderer, W. 1995. An improved method for the large scale synthesis of oligonucleotides applying the NPE/NPEOC strategy. Nucleos. Nucleot. 14:917-920. Winter, M. 1996. Supports for solid-phase organic synthesis. In Combinatorial Peptide and NonPeptide Libraries: A Handbook (G. Jung, ed.) pp. 465-510. VCH, Weinheim. Wright, P., Lloyd, D., Rapp, W., and Andrus, A. 1993. Large scale synthesis of oligonucleotides via phosphoramidite nucleosides and a highloaded polystyrene support. Tetrahedron Lett. 34:3373-3376. Yip, K.F. and Tsou, K.C. 1971. A new polymer support method for the synthesis of ribooligonucleotide. J. Am. Chem. Soc. 93:3272-3276. Zhang, X.H. and Jones, R.A. 1996. A universal allyl linker for solid-phase synthesis. Tetrahedron Lett. 37:3789-3790.
Contributed by Richard T. Pon University of Calgary Calgary, Alberta, Canada
Solid-Phase Supports for Oligonucleotide Synthesis
3.1.28 Current Protocols in Nucleic Acid Chemistry
Attachment of Nucleosides to Solid-Phase Supports
UNIT 3.2
The first step in any solid-phase synthesis is the covalent attachment of a monomeric unit (i.e., a nucleoside or mononucleotide) to the surface of an insoluble support. For oligonucleotide synthesis, this process is performed separately from the automated coupling steps used for chain elongation because of the different coupling chemistry involved. This unit provides protocols for using amide and ester linkages to attach nucleosides to long-chain alkylamine–controlled-pore glass (LCAA-CPG) bead supports. Protocols and descriptions of the coupling chemistry used by automated synthesizers to create internucleotide phosphodiester linkages are contained in subsequent units, and the properties and applications of a large variety of solid supports are reviewed in UNIT 3.1. Although prederivatized solid supports are commercially available from a number of sources, preparation of these materials in the lab is a very reasonable alternative to their purchase: the procedures are very simple, and a single support synthesis can produce sufficient material for a great number of oligonucleotide syntheses. Significant savings can also be obtained by filling one’s own synthesis columns instead of purchasing ready-to-use columns. In addition, many minor-nucleoside- and special non-nucleosidederivatized supports are either not commercially available or quite costly. This unit provides protocols for attaching nucleosides to supports via two different dicarboxylic acid linkers: succinic anhydride, S.4, and hydroquinone-O,O′-diacetic acid (HQDA or Q-linker), S.8. Basic Protocol 1 describes how commercially available nucleoside-3′-O-succinate hemiesters can be attached to LCAA-CPG through an amide linkage. Alternate Protocol 1 describes how nucleosides with free 3′-OH groups can be attached to succinylated LCAA-CPG through an ester linkage. Basic Protocol 2 describes how to synthesize nucleoside-3′-O-hydroquinone diacetyl hemiesters from nucleosides and HQDA and how they are coupled to LCAA-CPG. Alternate Protocol 2 describes the preparation of an HQDA-derivatized LCAA-CPG support and how nucleosides with free 3′-OH groups can be coupled to it through an ester linkage. Protocols for quantitating dimethoxytrityl (DMTr) groups attached to a support (Support Protocol 1), for converting surface amino or hydroxyl groups into DMTr derivatives for quantitation (Support Protocol 2), and for quantitating surface carboxylic acid groups as either p-nitrophenol (Support Protocol 3) or N-monomethoxytrityl-6-amino-1-hexanol (Support Protocol 4) derivatives are also included. The succinic acid linker has the advantage of being the most widely used linker arm, and both succinic anhydride and protected nucleoside-3′-O-succinyl hemiester starting materials are readily available. On the other hand, the hydroquinone-O,O′-diacetic acid linker offers greater compatibility with base-sensitive sequence modifications and greater synthetic throughput, because it can be cleaved under milder and faster conditions than can the succinyl linker. No changes in oligonucleotide synthesis protocols, other than reducing the cleavage time (i.e., from 60 to 90 min to only 2 to 3 min), are required when Q-linker is substituted for a succinyl linker. Almost any application requiring synthetic oligonucleotides can be satisfied using one of these linker arms. The major exceptions are applications that require leaving the final oligonucleotide immobilized on the support after synthesis and deprotection, i.e., the direct synthesis of immobilized oligonucleotide libraries. Researchers interested in this application should consult UNIT 3.1 for the appropriate noncleavable linker arms.
Synthesis of Unmodified Oligonucleotides
Contributed by Richard T. Pon
3.2.1
Current Protocols in Nucleic Acid Chemistry (2000) 3.2.1-3.2.23 Copyright © 2000 by John Wiley & Sons, Inc.
STRATEGIC PLANNING: SELECTING A LINKER ARM AND COUPLING PROTOCOL The protocols in this unit involving the two linker arms (succinic acid and HQDA) are each divided into two separate procedures, depending on the site where the linker is first attached. When the linker arm is attached to the 3′-hydroxyl position of a suitably protected nucleoside to produce a nucleoside-3′-O-hemiester, such as S.2 or S.9, a separate synthetic preparation is necessary for each different nucleoside. The alternate approach, which first couples the linker arm to the support to give a carboxyl-derivatized support, such as S.5 or S.12, eliminates the need for a separate hemiester preparation for each nucleoside. The coupling reaction between a carboxyl-derivatized support and the secondary 3′-OH of a nucleoside (i.e., S.6), however, is more difficult; this method is also slower and generates less nucleoside attached to the surface of the support. The selection of the best protocol depends on the cost and availability of the required nucleosides as well as each researcher’s experience with organic synthesis. Researchers who want to perform only minimal synthetic manipulations should choose Basic Protocol 1 (Fig. 3.2.1). This method consists of only one coupling step, if commercially available nucleoside succinate, S.2, is purchased. Researchers requiring supports derivatized with minor or unusual nucleosides, which are often very costly, and who will be preparing oligonucleotides with no unusual sensitivity to basic cleavage conditions should use Alternate Protocol 1, involving succinylated LCAA-CPG (Fig. 3.2.2). The succinylated LCAA-CPG support, S.5, is very easy to prepare, and this approach minimizes consumption of rare and expensive nucleosides. The two protocols using the Q-linker require a slightly greater familiarity with synthesis methodology, although these protocols are still within the ability of most “beginners.” Basic Protocol 2 describes the preparation of nucleoside hemiesters of HQDA (which are not yet commercially available) and their subsequent attachment to LCAA-CPG (Fig. 3.2.3). This method has the advantage of providing higher nucleoside loadings and is particularly important for synthesis on reusable supports (Pon et al., 1998, 1999). Researchers with more limited chemical resources should use Alternate Protocol 2 to prepare HQDA-derivatized LCAA-CPG (Fig. 3.2.4), at least until nucleoside-3′-O-hemiesters of HQDA become commercially available. As with Alternate Protocol 1 for the succinyl linker, this method is simple and eliminates the need for nucleoside-3′-O-hemiesters of HQDA, S.9. This method is also attractive because it avoids any use of pyridine. Use of the Q-linker rather than the succinyl linker is highly recommended for facilities wishing to increase their daily synthesis throughput or for projects requiring base-sensitive sequences, such as methyl phosphonate or methyl phosphotriester backbones, basesensitive nucleosides, and 3′-linked fluorescent dye labels. BASIC PROTOCOL 1
COUPLING OF NUCLEOSIDE-3′-O-SUCCINYL HEMIESTERS TO LCAA-CPG SUPPORT This protocol (Fig. 3.2.1) attaches 5′-dimethoxytrityl-N-protected nucleoside-3′-O-succinate hemiesters, S.2, directly to long-chain alkylamine–controlled-pore glass (LCAACPG; S.1). This protocol has the advantage of using commercially available succinate DMTO P
NH2 1
Attachment of Nucleosides to Solid-Phase Supports
O
O +
O
HO O 2
B
DMTO DEC/DMAP pyridine
O
B
O P
O
N H
O 3
Figure 3.2.1 Coupling of LCAA-CPG, S.1, directly to a nucleoside-3′-O-succinyl hemiester, S.2, as described in Basic Protocol 1. DMT, dimethoxytrityl group.
3.2.2 Current Protocols in Nucleic Acid Chemistry
P
O
O
NH2
O
+
1
4
DMAP/pyridine
O P DMTO
OH
N H
O
B
O
5 1 pentachlorophenol DEC/DMAP/pyridine 2 piperidine
HO DEC/DMAP pyridine
6
DMTO
O
B O
O P
N
N H
P
O
N H
O
O 7
3
Figure 3.2.2 Synthesis of succinylated support, S.5, by reaction of succinic anhydride, S.4, with LCAA-CPG, S.1. Nucleosides with a free 3′-OH group, S.6, can then be esterified to S.5 to yield support S.3. Unreacted succinic acid groups are blocked by conversion into an active pentachlorophenyl ester followed by treatment with piperidine to yield the unreactive site shown as support S.7. DMT, dimethoxytrityl group.
O
DMTO
O
HO C CH2 O
O CH2 C OH
O
B
+ HO
8
6
DEC/DMAP/pyridine DMTO O
B
O
O
HO C CH2 O
O CH2 C O 9
DMTO
O
+
B
DMTO
O O
O
B
O
C CH2 O
O CH2 C O 10
P
NH2
DEC/DMAP/pyridine
1 DMTO
P
H O N C CH2 O
O
B
O O CH2 C O 11
Figure 3.2.3 Synthesis of nucleoside-3′-O-hydroquinone diacetyl hemiester, S.9, from HQDA, S.8, and a protected nucleoside, S.6. A nucleoside diester, S.10, is also produced as a minor byproduct. The mixture of S.9 and S.10, however, is used directly in the coupling reaction with LCAA-CPG, S.1, to produce nucleoside-derivatized support S.11. DMT, dimethoxytrityl group.
Synthesis of Unmodified Oligonucleotides
3.2.3 Current Protocols in Nucleic Acid Chemistry
O P
NH2
O
HO C CH2 O
+
1
O CH2 C OH 8
HBTU/DMAP acetonitrile
H O N C CH2 O
P
O O CH2 C OH 12
DMTO
O
B HBTU/DMAP acetonitrile
HO 6
DMTO
P
H O N C CH2 O
O
B
O O CH2 C O 11
Figure 3.2.4 Coupling of LCAA-CPG, S.1, to HQDA, S.8, using HBTU and DMAP to produce HQDA-derivatized support S.12. A suitably protected nucleoside, S.6, with a 3′-OH group is then esterified to S.12 to generate nucleoside-derivatized support S.11. DMT, dimethoxytrityl group.
hemiester starting materials, S.2, and so the synthesis of these reagents is not included in this protocol. Researchers requiring derivatives of nucleosides other than the four common deoxyribonucleosides (dA, dC, dG, and T) may need to prepare them by coupling succinic anhydride with the appropriately protected rare nucleoside, according to established procedures (Gait et al., 1980; Miyoshi et al., 1980; Chow et al., 1981; Kumar et al., 1993; Markiewicz et al., 1997).
Materials LCAA-CPG beads (other amino-derivatized nonswelling support materials may also be used; see recipe) 5′-Dimethoxytrityl-N-protected nucleoside 3′-O-succinyl hemiester (see recipe) 4-Dimethylaminopyridine (DMAP) 1-(3-Dimethylaminopropyl)-3-ethylcarbodiimide hydrochloride (DEC; see recipe) Triethylamine, reagent grade Anhydrous pyridine, for coupling reactions (see recipe) Methanol, reagent grade Dichloromethane, reagent grade Pyridine, reagent-grade, for washing supports Cap A capping solution (see recipe) Cap B capping solution (see recipe) Screw-capped glass vials or round-bottom flasks Pasteur pipet Buchner funnel Whatman no. 1 filter paper Shaker, wrist action or similar Attachment of Nucleosides to Solid-Phase Supports
Additional reagents and solutions for trityl analysis (see Support Protocol 1)
3.2.4 Current Protocols in Nucleic Acid Chemistry
CAUTION: Carbodiimide reagents, such as DEC, can cause severe skin or eye irritation and allergic reactions. Use appropriate skin and eye protection, and wipe up all work areas and utensils with a wet cloth immediately after use (DEC is water soluble). NOTE: If the amino group loading of the starting support is not known, it is recommended that dimethoxyl derivatization to assay for amino and hydroxyl groups (see Support Protocol 2) be performed to verify the support’s capacity before starting the nucleoside derivatization.
Couple nucleoside 3′-O-succinyl hemiesters to LCAA-CPG 1. Combine in a small glass screw-capped vial or round-bottom flask: 1.0 g LCAA-CPG 0.2 mmol 5′-dimethoxytrityl-N-protected nucleoside 3′-O-succinyl hemiester 12 mg (0.1 mmol) DMAP 382 mg (2.0 mmol) DEC. 2. Add 80 µL triethylamine and ∼5 to 10 mL anhydrous pyridine. Quickly seal the vial, and shake at room temperature. The reaction time used depends on the nucleoside loading desired. Reactions as short as 1 hr can yield loadings of ∼20 to 30 mmol/g, whereas overnight reactions can yield loadings of up to 60 mmol/g. A coupling time of 2 to 3 hr, which usually gives a loading of between 30 and 40 mmol/g, is generally most satisfactory.
3. If desired, periodically monitor the course of the coupling reaction by removing a small sample of support (10 to 20 mg) with a Pasteur pipet and applying it to a Buchner funnel. Wash with 20 mL methanol and then 20 mL dichloromethane; dry uncovered at room temperature, ~15 min. Quickly determine the nucleoside loading by trityl analysis (see Support Protocol 1). 4. After sufficient nucleoside loading has been obtained, filter off the support, working in the fume hood. 5. Wash successively with ∼5 to 10 mL reagent-grade pyridine, ∼10 to 20 mL methanol, and finally ∼20 to 30 mL dichloromethane. Allow the support to dry at room temperature.
Cap unreacted amino groups 6. Return the support to a clean vial or flask, and add equal volumes (∼5 mL) of Cap A and Cap B capping reagents. Seal the vial and shake 1 to 2 hr at room temperature. 7. Filter off the support, and wash with ∼10 to 20 mL methanol and then ∼20 to 30 mL dichloromethane. Allow to air dry. 8. Determine the nucleoside loading by trityl analysis of a small portion of the final product (see Support Protocol 1). Store at room temperature in a sealed screw-capped vial. Supports can be stored for several years at room temperature.
Synthesis of Unmodified Oligonucleotides
3.2.5 Current Protocols in Nucleic Acid Chemistry
ALTERNATE PROTOCOL 1
NUCLEOSIDE COUPLING TO SUCCINYLATED LCAA-CPG This protocol describes the coupling of succinic anhydride, S.4, to LCAA-CPG, S.1, to produce a succinylated support, S.5 (Fig. 3.2.2). Suitably protected nucleosides, S.6, with a free 3′-OH group are then esterified to the succinylated support. Once the succinylated support has been prepared, only 5′-dimethoxytrityl-N-protected nucleosides (with free 3′-OH groups) are required. This provides an advantage over Basic Protocol 1, because a greater variety of nucleosides are commercially available as 5′- and N-protected derivatives than as 5′- and N-protected 3′-O-succinates. Additional Materials (also see Basic Protocol 1) Succinic anhydride, reagent grade Ninhydrin solution: 2% (w/v) ninhydrin in ethanol Pentachlorophenol, reagent grade Piperidine, reagent grade Water aspirator 13 × 75–mm glass test tube Additional reagents and solutions for quantitative nitrophenol assay (see Support Protocol 3), N-monomethoxytrityl-6-amino-1-hexanol assay (see Support Protocol 4), and trityl analysis (see Support Protocol 1) CAUTION: Carbodiimide reagents, such as DEC, can cause severe skin or eye irritation and allergic reactions. Use appropriate skin and eye protection, and wipe up all work areas and utensils with a wet cloth immediately after use (DEC is water soluble). CAUTION: Pentachlorophenol is highly toxic. CAUTION: Piperidine has a strong unpleasant odor and is highly toxic. Wear gloves and always handle in a fume hood. CAUTION: Avoid skin contact with the ninhydrin solution; blue stains will form. Couple succinic anhydride to LCAA-CPG 1. Combine in a screw-capped vial (such as a glass scintillation vial) or small roundbottom flask: 1.0 g LCAA-CPG 200 mg (2.0 mmol) succinic anhydride 24 mg (0.2 mmol) DMAP. Immediately add ∼5 to 10 mL anhydrous pyridine and seal the vial. Perform the pyridine addition quickly, and immediately seal the vial tightly, to minimize exposure to moisture in the air. Laboratories with very high humidity can avoid moisture contamination by sealing the vial with a septum and using a syringe for the pyridine transfer, but this is not necessary in most (air-conditioned) laboratories.
2. Shake the vial at room temperature overnight, using a speed sufficient to keep the support in a gentle suspension. Attachment of Nucleosides to Solid-Phase Supports
Do not use a magnetic stirrer for this or any subsequent steps, because the stir bar can break up the LCAA-CPG particles and produce unwanted fines.
3. Inside a fume hood, filter off the LCAA-CPG using a Buchner funnel, Whatman no. 1 filter paper, and vacuum provided by a water aspirator.
3.2.6 Current Protocols in Nucleic Acid Chemistry
4. Wash the support successively with ∼5 to 10 mL pyridine, then ∼10 to 20 mL methanol, and finally ∼20 to 30 mL dichloromethane. 5. Transfer the support to a piece of smooth, white paper and allow to sit uncovered at room temperature until dry, ∼15 to 30 min. The succinylated LCAA-CPG support can be stored indefinitely at room temperature.
6. Place 2 to 3 mg succinylated LCAA-CPG in a clean 13 × 75–mm glass test tube, and add 1 or 2 drops of ninhydrin solution. Warm the mixture until the solvent evaporates and the beads are dry. Any unreacted amino groups will form a very dark blue color on the beads, whereas the succinylated support should only have a faint blue color. This is a qualitative test, and it is helpful to treat a sample of LCAA-CPG alongside the test material to observe the color difference.
7. Verify the extent of the succinylation reaction by performing the quantitative nitrophenol (see Support Protocol 3) or N-monomethoxytrityl-6-amino-1-hexanol (see Support Protocol 4) assay for carboxylic acid loading. If the test results are unsatisfactory, repeat the succinylation reaction on the support. The carboxyl loading obtained on succinyl-derivatized LCAA-CPG should be close to the amino loading determined by the LCAA-CPG manufacturer. Owing to differences in the nature of the amino and carboxyl determinations, it is normal to observe some difference.
Couple nucleoside to the succinylated LCAA-CPG 8. Combine in a small screw-capped glass vial or round-bottom flask: 1.0 g succinylated LCAA-CPG 0.1 mmol 5′-dimethoxytrityl-N-protected 2′-deoxyribonucleoside 12 mg (0.1 mmol) DMAP 192 mg (1.0 mmol) DEC. Add 80 µL triethylamine and ∼5 to 10 mL anhydrous pyridine. Quickly seal the vial, and shake overnight at room temperature. 9. Verify the progress of the coupling reaction by removing ∼10 to 20 mg of support using a Pasteur pipet. Apply the sample to a Buchner funnel, and wash with 10 to 20 mL methanol and then 20 to 30 mL dichloromethane. Allow the support to dry, and then determine the nucleoside loading by trityl analysis (see Support Protocol 1). If the loading is satisfactory (i.e., a minimum of 20 and preferably 30 to 40 µmol/g), then proceed to step 10. Otherwise, extend the shaking for another day or repeat the reaction by adding more of the reagents (nucleoside, DEC, DMAP, and triethylamine) to the flask. Cap unreacted carboxyl and amino sites on the support 10. Add 134 mg (0.5 mmol) pentachlorophenol to the reaction mixture, and shake 16 to 24 hr at room temperature. Pentachlorophenol is added to convert any unreacted carboxyl groups into activated pentachlorophenyl esters. When treated with the cyclic amine piperidine (step 11), the pentachlorophenyl esters are converted into an unreactive amide, S.7.
11. Add 5 mL piperidine and continue shake 5 min at room temperature. Prolonged exposure to piperidine will also hydrolyze the succinyl linker and decrease the nucleoside loading on the support.
Synthesis of Unmodified Oligonucleotides
3.2.7 Current Protocols in Nucleic Acid Chemistry
12. Inside a fume hood, filter off the LCAA-CPG beads and wash successively with ∼5 to 10 mL reagent-grade pyridine, ∼10 to 20 mL methanol, and finally ∼20 to 30 mL dichloromethane. Allow the support to sit at room temperature until dry. 13. Return the support to a clean glass screw-capped vial or small round-bottom flask and add equal amounts (∼5 mL) of Cap A and Cap B capping reagents. Seal the vial and shake 1 to 2 hr at room temperature. This reaction is performed to acetylate any amino groups on the support that may not have been succinylated in the first reaction.
14. Filter off the LCAA-CPG on a Buchner funnel and wash with ∼10 to 20 mL methanol and then ∼20 to 30 mL dichloromethane. Allow to dry at room temperature. 15. Determine the nucleoside loading by trityl analysis of a small portion of the final product (see Support Protocol 1). Store at room temperature in a sealed screw-capped vial. Supports can be stored for several years. BASIC PROTOCOL 2
COUPLING OF NUCLEOSIDE-3′-O-HYDROQUINONE-O,O′-DIACETYL HEMIESTERS TO LCAA-CPG In this protocol, the hydroquinone-O,O′-diacetic acid (HQDA), S.8, is first attached to the 3′-OH of a suitably protected nucleoside, S.6, by using 1-(3-dimethylaminopropyl)3-ethylcarbodiimide hydrochloride (DEC) as the coupling reagent. Unfortunately, selective coupling to only one end of the diacid linker is not possible, and a mixture of nucleoside hemiesters, S.9, and diesters, S.10, always occurs (Fig. 3.2.3). Fortunately, chromatographic separation of the hemiesters and diesters is not necessary, because only the hemiester can couple to the long-chain alkylamine–controlled-pore glass (LCAACPG) support. Thus the preparation of the HQDA hemiesters is not much more difficult than the synthesis of nucleoside-3′-O-succinyl hemiesters. Coupling of the HQDA hemiesters to LCAA-CPG is performed in the same manner as the succinyl compounds.
Attachment of Nucleosides to Solid-Phase Supports
Materials 5′-Dimethoxytrityl-N-protected deoxyribonucleoside (see recipe) Hydroquinone-O,O′-diacetic acid (HQDA; Lancaster Synthesis) 4-Dimethylaminopyridine (DMAP), reagent grade 1-(3-Dimethylaminopropyl)-3-ethylcarbodiimide hydrochloride (DEC; see recipe) Triethylamine, reagent grade Anhydrous pyridine, for coupling reactions (see recipe) Toluene, reagent grade Chloroform, reagent grade 5% (v/v) methanol/chloroform, for TLC analysis Saturated aqueous sodium bicarbonate solution Anhydrous magnesium sulfate Pyridine, reagent-grade, for washing supports AG 50W-X4 H+ form cation-exchange resin (Bio-Rad) LCAA-CPG beads (other amino-derivatized nonswelling support materials may also be used; see recipe) Methanol, reagent grade Dichloromethane, reagent grade Cap A capping solution (see recipe) Cap B capping solution (see recipe)
3.2.8 Current Protocols in Nucleic Acid Chemistry
50- and 100-mL round-bottom flasks Magnetic stir bar Pasteur pipet Fluorescent silica gel thin-layer chromatography (TLC) plates, ∼2.5 cm × 10 cm (Merck 60 or similar) Covered jar for TLC Shortwave UV (254-nm) lamp Separatory funnel Shaker, wrist action or similar Rotary evaporator Additional reagents and solutions for dimethoxyl derivatization to assay for amino and hydroxyl groups present (see Support Protocol 2; optional) and trityl analysis (see Support Protocol 1) CAUTION: Carbodiimide reagents, such as DEC, can cause severe skin or eye irritation and allergic reactions. Use appropriate skin and eye protection, and wipe up all work areas and utensils with a wet cloth immediately after use (DEC is water soluble). Synthesize nucleoside-3′-O-HQDA hemiesters 1. Combine in a 100-mL round-bottom flask with a magnetic stir bar: 10 mmol 5′-dimethoxytrityl-N-protected deoxyribonucleoside 3.39 g (15 mmol) HQDA 122 mg (1 mmol) DMAP 2.88 g (15 mmol) DEC. Add 0.8 mL triethylamine and 50 mL anhydrous pyridine. Seal the flask and stir overnight at room temperature. 2. Remove 50 to 100 µL of the solution using a Pasteur pipet and place in a 50-mL round-bottom flask. Add ∼1 to 2 mL toluene and evaporate to dryness using a rotary evaporator. 3. Redissolve in ∼100 to 200 µL chloroform. 4. Spot the sample near the bottom edge of an ∼2.5 × 10–cm TLC plate, alongside samples of the starting nucleoside and HQDA. Place the TLC plate in a jar containing a small amount of 5% (v/v) methanol/chloroform (the liquid should be ∼2 to 4 mm deep), and allow the solvent to rise most of the way up the TLC plate. 5. Remove plate and examine under a shortwave UV (254-nm) lamp. The desired hemiester will appear as a UV-absorbing spot near the baseline, and the diester product will be a UV-absorbing spot near the top of the plate. The starting nucleoside will migrate to near the middle of the plate. The coupling reaction is complete when all the starting nucleoside is converted into the faster- (diester) and slower-migrating (hemiester) products. If more than a trace of starting nucleoside is visible, add more DEC (2 to 5 mmol) to the reaction, and continue stirring for another day. Repeat steps 2 to 5.
6. When TLC shows complete disappearance of the starting nucleoside, concentrate the solution on the rotary evaporator until a thick oil forms. 7. Redissolve the oil in ∼200 mL chloroform and transfer to a separatory funnel. Wash the solution twice with ∼100 mL saturated aqueous sodium bicarbonate. CAUTION: Carbon dioxide gas is released during this extraction step. Shake the separatory funnel slowly with frequent venting until gas evolution stops.
Synthesis of Unmodified Oligonucleotides
3.2.9 Current Protocols in Nucleic Acid Chemistry
8. Wash the chloroform solution three times with ∼100 mL water each time. Collect the chloroform phase and discard the aqueous phase. Do not shake the separatory funnel vigorously at this step, because the mixture will form an emulsion that is slow to separate. Invert the funnel slowly to mix the phases. If an inseparable emulsion does form, then either (for small volumes) centrifugation or (for large volumes) precipitation by addition of hexanes followed by filtration and redissolution of the sticky precipitate back into chloroform can be performed.
9. Add the chloroform solution to ∼5 g anhydrous magnesium sulfate, and mix to remove residual moisture from the solution. Filter off the magnesium sulfate, wash with a small amount of chloroform, and then evaporate the chloroform solution to dryness using the rotary evaporator. A light brown foam should form and solidify. The crude product, which contains a mixture of diester and nucleoside hemiester sodium salt, is suitable for coupling directly to LCAA-CPG using the DEC/pyridine coupling procedure that follows. The sodium salt, however, may not be sufficiently soluble in other solvents for other types of coupling reactions. In this case, the sodium salt can be easily converted into the more soluble pyridinium salt by the procedure outlined in step 10.
10. Conversion into pyridinium salt (optional): Dissolve the crude sodium salt in ∼50 to 100 mL of reagent-grade pyridine, and add 2 eq AG 50W-X4 H+ cation-exchange resin. Stir ∼5 to 10 min; then filter off the ion-exchange resin. Evaporate the pyridine solution to dryness; a light brown foam will form and solidify. Dry under vacuum overnight to remove excess pyridine. Store in sealed, moisture-proof containers at −20°C. Samples are stable for at least 1 to 2 years. Older samples should be checked by TLC to determine the extent of decomposition, if any.
Couple nucleoside 3′-O-HQDA hemiesters to LCAA-CPG 11. If the amino-group loading of the starting support is not known: Verify the support’s capacity by dimethoxyl derivatization to determine the number of amino and hydroxyl groups present (see Support Protocol 2) before starting the nucleoside derivatization. 12. Combine in a round-bottom flask: 5.0 g LCAA-CPG 2 g (2.0 mmol) crude nucleoside 3′-O-HQDA hemiester (step 9 or 10) 122 mg (0.1 mmol) DMAP 1.44 g (7.5 mmol) DEC. Add 1 mL triethylamine and 30 mL anhydrous pyridine and quickly seal the flask. Shake overnight at room temperature. 13. In the fume hood, filter off the support and wash successively with ∼25 to 50 mL reagent-grade pyridine, ∼50 to 100 mL methanol, and finally ∼100 to 150 mL dichloromethane. Allow to dry at room temperature. 14. Remove a small aliquot for trityl analysis to determine the nucleoside loading (see Support Protocol 1).
Attachment of Nucleosides to Solid-Phase Supports
15. If the nucleoside loading is acceptable, return the support to a clean flask, and add equal amounts (∼25 mL) of Cap A and Cap B capping solutions. Seal the flask and shake 1 hr at room temperature. 16. Filter off the support and wash with ∼50 to 100 mL methanol and then ∼100 to 150 mL dichloromethane. Allow to dry at room temperature.
3.2.10 Current Protocols in Nucleic Acid Chemistry
17. Verify the nucleoside loading by performing trityl analysis (see Support Protocol 1) on a sample of the final product. Supports derivatized with the Q-linker have been stored for 3 years at room temperature without any significant loss of nucleoside.
NUCLEOSIDE COUPLING TO HYDROQUINONE-O,O′-DIACETIC ACID DERIVATIZED LCAA-CPG
ALTERNATE PROTOCOL 2
In this protocol HQDA, S.8, is coupled to LCAA-CPG, S.1, to give a carboxyl-derivatized support, S.12 (Fig. 3.2.4). This coupling reaction must be performed in the presence of a coupling reagent because HQDA is not available as an acid anhydride (as succinic anhydride is for use in succinylation approaches). In addition, the coupling reactions involving immobilized HQDA—especially the formation of esters to the secondary hydroxyl group of protected nucleosides, S.6—are best performed using a more reactive coupling reagent than the carbodiimide used in the above protocols. Therefore, 2-(1Hbenzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HBTU), which is a coupling reagent developed for peptide synthesis (Dourtoglou et al., 1984), is used instead of DEC along with an increased amount of DMAP. Another advantage to using HBTU is that anhydrous acetonitrile can be substituted for pyridine as the solvent. Additional Materials (also see Basic Protocol 2) 2-(1H-Benzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HBTU) Anhydrous diisopropylethylamine (DIEA; see recipe) Anhydrous acetonitrile (see recipe) Acetonitrile, reagent grade Screw-capped glass vials or round-bottom flasks Syringes Buchner funnel Whatman no. 1 filter paper Additional reagents and solutions for N-monomethoxytrityl-6-amino-1-hexanol assay of carboxylic acid loading (see Support Protocol 4) and trityl analysis (see Support Protocol 1) CAUTION: All coupling reagents, such as HBTU, can potentially cause adverse reactions and lead to severe allergic reactions. Although specific data on HBTU are not available, safe laboratory practices (i.e., no skin or eye contact or inhalation of powders) should always be followed when handling this type of reagent. Couple HQDA to LCAA-CPG 1. Combine in a glass screw-capped vial or small glass-round bottom flask: 1.0 g LCAA-CPG 90 mg (0.4 mmol) HQDA 46 mg (0.38 mmol) DMAP 144 mg (0.36 mmol) HBTU. Via syringe, add 140 µL (0.8 mmol) DIEA and 10 mL anhydrous acetonitrile. Seal the vial and shake 2 hr at room temperature. 2. Filter off the support using a Buchner funnel and wash the LCAA-CPG successively with ∼10 to 20 mL acetonitrile, ∼10 to 20 mL methanol, and finally ∼20 to 30 mL dichloromethane. Allow the support to dry at room temperature.
Synthesis of Unmodified Oligonucleotides
3.2.11 Current Protocols in Nucleic Acid Chemistry
3. Verify the extent of the HQDA derivatization by performing the N-monomethoxytrityl-6-amino-1-hexanol assay of carboxylic acid loading (see Support Protocol 4). A carboxyl loading of 30 to 40 mmol/g is typical. Supports can be stored indefinitely at room temperature. Store the support at room temperature in a sealed glass vial.
Couple nucleoside to the HQDA-derivatized LCAA-CPG 4. Combine in a screw-capped glass vial or small glass round-bottom flask: 1.0 g HQDA-derivatized LCAA-CPG 0.2 g (0.2 mmol) 5′-dimethoxytrityl-N-protected deoxyribonucleoside 24 mg (0.2 mmol) DMAP 76 mg (0.2 mmol) HBTU. Via syringe, add 140 µL (0.8 mmol) DIEA and 5 mL anhydrous acetonitrile. Quickly seal the vial, and shake 2 to 3 hr at room temperature. 5. Filter off the support using a Buchner funnel and wash successively with ∼10 to 20 mL acetonitrile, ∼10 to 20 mL methanol, and finally ∼10 to 20 mL dichloromethane. Allow the support to dry at room temperature. 6. Determine the nucleoside loading by trityl analysis of a small sample of support (see Support Protocol 1). Cap unreacted amino groups 7. Return the support into a clean vial or flask and add equal amounts (∼5 mL) of Cap A and Cap B capping reagents. Seal the vial and shake 1 hr at room temperature. 8. Filter off the support and wash with ∼10 to 20 mL methanol and then with ∼10 to 20 mL dichloromethane. Allow to dry at room temperature. 9. Determine the nucleoside loading by trityl analysis of a small portion of the final product (see Support Protocol 1). Label the support with this loading value and the date of synthesis, and store at room temperature in a sealed screw-capped vial. Supports can be kept for several years. SUPPORT PROTOCOL 1
QUANTITATIVE TRITYL ANALYSIS OF SURFACE LOADING OF DERIVATIZED SUPPORTS This procedure describes how the surface loading of supports containing trityl groups can be determined by colorimetric analysis. This method detects the 5′-trityl protecting group on nucleoside-derivatized supports or trityl groups added to derivatize aminoor hydroxyl-functionalized supports (see Support Protocol 2). This assay is based on the fact that acid-labile dimethoxytrityl (from nucleoside-derivatized supports) or monomethoxytrityl protecting groups (from N-monomethoxytrityl-6-amino-1-hexanol–derivatized supports) can be rapidly and quantitatively hydrolyzed to form intensely orangeand yellow-colored solutions.
Attachment of Nucleosides to Solid-Phase Supports
Materials Derivatized LCAA-CPG support (see Basic Protocols 1 and 2 and Alternate Protocols 1 and 2) Detritylation reagent (see recipe)
3.2.12 Current Protocols in Nucleic Acid Chemistry
10-mL volumetric flask Analytical balance accurate to 0.1 mg Quartz cuvettes UV/visible spectrophotometer or colorimeter 1. Tare an empty 10-mL volumetric flask on an analytical balance. 2. Place ∼4 mg dry, derivatized LCAA-CPG support directly into the flask and weigh. Record the weight to at least 2 significant figures. 3. Add detritylation reagent to the volumetric flask to the mark. Seal the flask and mix. An orange-colored dimethoxytrityl cation (yellow if monomethoxytrityl groups are being measured) will immediately form. The orange color is stable if moisture contamination and evaporation are avoided and the solution can be kept at room temperature indefinitely.
4. Fill a reference cuvette with detritylating reagent or, if using a single-beam spectrophotometer, run a baseline. The dichloromethane solvent requires quartz rather than disposable polystyrene cuvettes.
5. Measure the spectrum of the sample from 400 to 600 nm. Record the absorbance of the peak maximum at ∼503 to 505 nm (dimethoxytrityl group) or ∼470 nm (monomethoxytrityl group). If the absorbance exceeds the maximum reliable value for the spectrophotometer (usually 2 to 3 absorbance units), prepare another sample with less support or perform a serial dilution using detritylation reagent as the diluent.
6. Calculate the loading of the support using the following equation (for a cuvette with a 1-cm path length): Loading =
volume × absorbance 1000 × weight ε
where loading is in micromoles per gram, the solution volume in milliliters, and the support weight is in milligrams; and ε is the extinction coefficient of either the dimethoxytrityl group (76 mL cm−1 µmol−1) or the monomethoxytrityl group (56 mL cm−1 µmol−1). DIMETHOXYTRITYL DERIVATIZATION OF AMINO- AND/OR HYDROXYL-FUNCTIONALIZED SUPPORTS
SUPPORT PROTOCOL 2
A variety of methods have been developed to determine the primary amino group content of insoluble supports (Horn and Novak, 1987). The most convenient method, however, uses the dimethoxytrityl chloride/tetrabutylammonium perchlorate reagent developed for the solid-phase tritylation of nucleosides and nucleotides (Reddy et al., 1987). This reagent will also tritylate amino groups, and it has been used to monitor coupling efficiency in solid-phase peptide synthesis (Reddy and Voelker, 1988). This is a very fast, sensitive, and general method for determining the number of amino and hydroxyl groups on insoluble supports. Materials LCAA-CPG support 0.25 M dimethoxytrityl (see recipe) chloride stock solution (see recipe) 0.25 M tetrabutylammonium perchlorate stock solution (see recipe) Methanol, reagent grade
Synthesis of Unmodified Oligonucleotides
3.2.13 Current Protocols in Nucleic Acid Chemistry
Dichloromethane, reagent grade Screw-capped glass vials 1. Place ∼10 to 20 mg of LCAA-CPG support in a small screw-capped glass vial and add equal amounts (∼500 µL) of 0.25 M dimethoxytrityl chloride and 0.25 M tetrabutylammonium perchlorate. Seal the vial and shake 10 min at room temperature. 2. Filter off the support and wash first with 10 to 20 mL dichloromethane, then with 10 to 20 mL methanol, and finally with 20 to 30 mL dichloromethane again. The methanol wash is important to remove any adsorbed trityl reagent from the support.
3. Allow the support to dry and then perform the colorimetric trityl analysis (see Support Protocol 1). SUPPORT PROTOCOL 3
DETERMINATION OF CARBOXYLIC ACID GROUPS ON INSOLUBLE SUPPORTS BY p-NITROPHENOL DERIVATIZATION The amount of carboxylic acid attached to the surface on an insoluble support can be determined by titration, but this consumes large amounts (gram quantities) of support. A better and more sensitive method is to derivatize the support with an appropriate reporter molecule. p-Nitrophenol is a convenient and readily available label that has been widely used. When this label is removed from the support, by hydrolysis with aqueous sodium hydroxide, a yellow-green solution is produced that can be quantitated by colorimetry (Matteucci and Caruthers, 1981; Damha et al., 1990). Although the extinction coefficient for the p-nitrophenoxide group is only one-fifth as large as for the dimethoxytrityl cation (see Support Protocol 1), this test is useful for measuring the succinic acid surface loading produced in Alternate Protocol 1. This method does not, however, work with HQDA-derivatized LCAA-CPG (see Support Protocol 4 instead). Additional Materials (see Basic Protocol 1 and Alternate Protocol 1) p-Nitrophenol, reagent grade 0.1 M aqueous NaOH solution N-Monomethoxytrityl-6-amino-1-hexanol 10-mL volumetric flask Analytical balance UV/visible spectrophotometer or colorimeter Additional reagents and solutions for DEC/DMAP coupling (see Alternate Protocol 1) and trityl analysis (see Support Protocol 1) 1. Combine in a small screw-capped glass vial or round-bottom flask: 1.0 g succinylated LCAA-CPG 14 mg (0.1 mmol) p-nitrophenol 190 mg (0.1 mmol) DEC. Via syringe, add 40 µL triethylamine and 1 mL anhydrous pyridine. Seal the vial, and shake overnight at room temperature.
Attachment of Nucleosides to Solid-Phase Supports
2. Filter off the support and wash sequentially with 5 mL pyridine, 10 mL methanol, and 10 to 20 mL dichloromethane. Leave the support uncovered at room temperature to dry. The capping steps (see Alternate Protocol 1, steps 10 to 15) are unnecessary or this protocol.
3.2.14 Current Protocols in Nucleic Acid Chemistry
3. Accurately weigh ∼10 mg of support directly into a 10-mL volumetric flask. Add 0.1 M NaOH solution to the mark and mix. 4. Determine the absorbance of the solution at 400 nm. Calculate the support loading using an extinction coefficient (ε) of 15.7 mL cm−1 µmol−1 for the nitrophenoxide group; see Support Protocol 1, step 6, for formula. DETERMINATION OF CARBOXYLIC ACID GROUPS ON INSOLUBLE SUPPORTS BY MONOMETHOXYTRITYL DERIVATIZATION
SUPPORT PROTOCOL 4
This protocol is an alternate to Support Protocol 3. Unlike Support Protocol 3, this method can be used to determine the carboxyl loading of both succinyl- and HQDA-derivatized LCAA-CPG. This method requires synthesis of a custom reporter molecule, Nmonomethoxytrityl-6-amino-1-hexanol, developed by Pon and Yu (1997a). The carboxyl loading determination is more sensitive, because the yellow color produced from the monomethoxytrityl group (in acid solution; i.e., see Support Protocol 1) is more than three times as intense as that produced by the p-nitrophenoxide group. Additional Materials (see Basic Protocol 2 and Alternate Protocol 2) 6-Amino-1-hexanol Monomethoxytrityl chloride N-Monomethoxytrityl-6-amino-1-hexanol (see recipe) Silica gel, Merck 60, for flash chromatography Glass chromatography column for flash chromatography, ~4 to 5 cm diameter × ~50 to 60 cm long Silica-gel TLC plates (Merck, ~2.5 × 8 cm) CAUTION: All coupling reagents, such as HBTU, can potentially cause adverse reactions and lead to severe allergic reactions. Although specific data on HBTU are not available, safe laboratory practices (i.e., no skin or eye contact or inhalation of powders) should always be followed when handling this type of reagent. 1. Dissolve 2.42 g (20 mmol) 6-amino-1-hexanol and 6.37 g (20 mmol) monomethoxytrityl chloride in 100 mL anhydrous pyridine, and stir 1 to 2 days at room temperature. 2. Concentrate the solution on a rotary evaporator to 10 to 15 mL. 3. Redissolve in 75 mL chloroform, wash with 25 mL aqueous sodium bicarbonate solution, and wash twice with 25 mL water. Dry over anhydrous magnesium sulfate. 4. Concentrate the chloroform solution to an oil. 5. Apply to a flash chromatography column packed ~20 cm high with Merck 60 silica gel. Begin the purification by eluting the column with 5% methanol/chloroform and collect the eluate in ~50- to 100-mL fractions. 6. Monitor the separation by spotting a small amount (~2 to 5 µL) of each fraction on a silica-gel TLC plate (~2.5 × 8 cm). Develop the plate in 20% methanol/chloroform and detect the products either by UV or by spraying with detritylation reagent and then gently warming the plate (e.g., with a hair dryer) for a few seconds: monomethoxytrityl-containing products will appear as yellow spots. Once all the unreacted monomethoxytrityl chloride has eluted from the column or when the desired product begins to appear, change the solvent to 15% methanol/chloroform.
Synthesis of Unmodified Oligonucleotides
3.2.15 Current Protocols in Nucleic Acid Chemistry
Initially, hydrolyzed, unreacted monomethoxytrityl chloride will appear as an impurity moving near the solvent front. The desired product will have a much slower mobility (Rf ~ 0.15). Unreacted aminohexanol will remain bound to the silica gel. 7. Combine fractions containing the desired product. Evaporate on a rotary evaporator to obtain pure N-monomethoxytrityl-6-amino-1-hexanol as a thick brown liquid (5.5 g, 71% yield). 8. Combine in a screw-capped glass vial or small glass round-bottom flask: 0.1 g succinyl- or HQDA-derivatized LCAA-CPG 38 mg (0.1 mmol) N-monomethoxytrityl-6-amino-1-hexanol 24 mg (0.02 mmol) DMAP 38 mg (0.1 mmol) HBTU. Via syringe, add 14 µL DIEA (0.08 mmol) and ∼0.5 mL anhydrous acetonitrile. Quickly seal the vial and shake 2 to 3 hr at room temperature. 9. Filter off the support using a Buchner funnel, and wash successively with ∼10 to 20 mL acetonitrile, ∼10 to 20 mL methanol, and finally ~20 to 30 mL dichloromethane. Allow the support to dry uncovered at room temperature. 10. Determine the monomethoxytrityl loading by trityl analysis (see Support Protocol 1). NOTE: The monomethoxytrityl group generates a yellow rather than an orange color, and the measurement must be made with the wavelength and extinction coefficient specified in Support Protocol 1.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Anhydrous acetonitrile Distill from calcium hydride or obtain from commercially available “low-water” reagents suitable for automated oligonucleotide synthesis. Store over molecular sieves in tightly sealed bottles. Discard or redistill after each day’s use. Anhydrous diisopropylethylamine (DIEA) Purchase as the 99.5% redistilled grade (Aldrich), and keep anhydrous by storage over type 4A molecular sieves. Store in septum-sealed bottles at room temperature, protected from light. Anhydrous pyridine Prepare by distillation from calcium hydride, and keep anhydrous by storage over type 4A molecular sieves in tightly sealed bottles. Store up to 1 year at room temperature. Alternatively, this may be purchased ready to use in Sure/Seal bottles (Aldrich).
Attachment of Nucleosides to Solid-Phase Supports
Cap A and Cap B capping reagents Cap A: 94 mL 1 M acetic anhydride/116 mL 1 M 2,6-lutidine or pyridine in 790 mL tetrahydrofuran (THF). Store indefinitely at room temperature. Cap B: 160 mL 2 M N-methylimidazole in 840 mL THF. Store indefinitely at room temperature. These are the same reagents used as capping reagents on automated DNA synthesizers, and may be purchased commercially.
3.2.16 Current Protocols in Nucleic Acid Chemistry
Detritylation reagent 5% Dichloroacetic acid (v/v) or 3% trichloroacetic acid (w/v) in 1,2-dichloroethane or dichloromethane. Store indefinitely at room temperature. There is no significant difference between these reagents in the trityl analysis, although dichloroacetic acid reagents are more suitable for the synthesis of long oligonucleotides. To reduce solvent vapors, it is recommended that the less volatile 1,2-dichloroethane be substituted for the more commonly used dichloromethane. Detritylation reagent may be purchased commercially (sometimes referred to as “deblock” reagent). CAUTION: Many older references suggest the use of perchloric acid solutions to dilute the trityl colors. Such a strong acid is not required, and for safety reasons, perchloric acid solutions should not be used. The same detritylation reagent used on the DNA synthesizers can be used to dilute the colors for measurement.
Dimethoxytrityl chloride stock solution, 0.25 M Dissolve 847 mg (2.5 mmol) dimethoxytrityl chloride in 10 mL dichloromethane. Store in a sealed glass vial up to 3 to 4 weeks at room temperature. 5′-Dimethoxytrityl-N-protected nucleosides Purchase 5′-dimethoxytrityl nucleosides with a free 3′-OH or, for ribonucleosides, free 2′- and 3′-OH groups (Chem-Impex, Sigma, and ChemGenes). N6-benzoyl-5′dimethoxytrityl-2′-deoxyadenosine, N4-benzoyl-5′-dimethoxytrityl-2′-deoxycytidi ne, N2-isobutyryl-5′-dimethoxytrityl-2′-deoxyguanosine, and 5′-dimethoxytritylthymidine are the most commonly used standard nucleosides and are compatible with the protocols described in this unit. Store indefinitely at room temperature. 5′-Dimethoxytrityl-N-protected nucleoside succinyl hemiesters Purchase nucleosides with the standard protecting groups—N6-benzoyl-5′-dimethoxytrityl-2′-deoxyadenosine-3′-O-succinate, N4-benzoyl-5′-dimethoxytrityl2′-deoxycytidine-3′-O-succinate, N2-isobutyryl-5′-dimethoxytrityl-2′-deoxy-guanosine3′-O-succinate, and 5′-dimethoxytritylthymidine-3′-O-succinate (Chem-Impex and Sigma). Store indefinitely at room temperature. Only the standard succinates are available for purchase; more unusual or complicated succinates must be made up.
1-(3-Dimethylaminopropyl)-3-ethylcarbodiimide hydrochloride (DEC) Also referred to as 3-ethyl-1-(3-dimethylaminopropyl)-carbodiimide (EDC) and water soluble carbodiimide (WSC). Readily available from commercial suppliers. Store indefinitely at –20°C. Long-chain alkylamine–controlled-pore glass (LCAA-CPG) Available in a variety of pore and particle sizes and sieved into three sets of particle sizes: 80 to 120 mesh (125 to 177 µm), 120 to 200 mesh (74 to 125 µm), and 200 to 400 mesh (37 to 74 µm); the two largest size ranges are the most suitable for oligonucleotide synthesis. Generally, 500-Å-diameter pores are used for oligonucleotides of up to ∼50 bases in length, and 1000-Å-diameter pores are required for longer oligonucleotides. Store indefinitely at room temperature. N-Monomethoxytrityl-6-amino-1-hexanol Prepare from monomethoxytrityl chloride and 6-amino-1-hexanol (see Support Protocol 4 or Pon and Yu, 1997a). Store indefinitely at room temperature. Tetrabutylammonium perchlorate stock solution, 0.25 M Dissolve 854 mg (2.5 mmol) tetrabutylammonium perchlorate (Sigma) in 0.3 mL 2,6-lutidine and 9.7 mL dichloromethane. Store in a sealed glass vial up to 3 to 4 weeks at room temperature.
Synthesis of Unmodified Oligonucleotides
3.2.17 Current Protocols in Nucleic Acid Chemistry
COMMENTARY Background Information
Attachment of Nucleosides to Solid-Phase Supports
The succinic acid linker was one of the earliest linkages used to immobilize nucleosides to insoluble supports (Ogilvie and Kroeker, 1971; Yip and Tsou, 1971), and it has remained widely used since. Although a variety of different linkages have been developed for specialized applications (UNIT 3.1; Pon, 1993), routine oligonucleotide synthesis has been well served by the succinyl linker, primarily because succinic anhydride, S.4, does not need any activation to react with hydroxyl (or amino) groups and selectively produces only hemiesters (or amides). The low cost and easy handling of this reagent are also advantageous. Once the anhydride has been opened, however, coupling to the terminal free carboxylic acid group is more difficult. This group requires activation by conversion to an acid chloride (Ogilvie and Kroeker, 1971; Sharma et al., 1992), an activated ester, or a symmetrical anhydride (Gait et al., 1980) or by reaction with a coupling reagent such as dicyclohexylcarbodiimide (DCC; Chow et al., 1981; Matteucci and Caruthers, 1981; Montserrat et al., 1993). Because acid chlorides are difficult to work with and the DCC coupling reagent does not perform well with LCAA-CPG supports, most early work was performed using activated nucleoside-3′-O-succinyl pentachlorophenyl (Miyoshi et al., 1980), p-nitrophenyl (Koster et al., 1983), or pentafluorophenyl esters (Efimov et al., 1993). These intermediates, however, require an additional coupling step to prepare, and the support derivatization reactions are slow. Later it was discovered that the carbodiimide reagent DEC could rapidly couple unactivated nucleoside succinates directly to LCAA-CPG (Pon et al., 1988). Given that coupling reactions with DEC are completed in as little as 1 hr, this technique has become the preferred approach. Recently, however, improved coupling methods have been described using diisopropylcarbodiimide and hydroxybenzotriazole (Bhongle and Tang, 1995) or DCC and 3,4-dihydro-3-hydroxy-4-oxo-1,2,3-benzotriazine (Walsh et al., 1997), which allow satisfactory derivatization of LCAA-CPG supports. Other new coupling reagents have also been used to speed up the derivatization reactions. Use of an oxidation-reduction coupling reagent combination, triphenylphosphine and 2,2′dithiobis(5-nitropyridine), reduces the coupling time to between 2 and 30 min (Kumar et
al., 1996). Extremely fast derivatization (4 sec) is obtained by using either uronium- or phosphonium-coupling reagents and DMAP (Pon and Yu, 1997b); however, none of these methods has been widely adopted. Another benefit of the DEC coupling reagent is that it can also be used to create a secondary ester linkage between the 3′ (or 2′) hydroxyl group of a nucleoside and a succinylated support (Damha et al., 1990). Because forming a secondary ester is more difficult than creating a primary amide, the coupling reaction is slower and support loadings are often not as high as obtained from coupling a pre-formed nucleoside-3′-succinate to LCAA-CPG. Not having to prepare nucleoside succinates is a significant advantage, however, especially when limited starting materials are available. Recently, the huge success of solid-phase oligonucleotide synthesis has created a great demand for large numbers of synthetic oligonucleotides, and a number of productivity enhancements have been implemented. New high-throughput DNA synthesizers have been developed, and more labile protecting groups and faster deprotection conditions have been introduced to speed up processing times. A significant increase in productivity can also be obtained by replacing the succinyl linker with a more rapidly cleavable linker. This is particularly so with instrumentation that performs the cleavage step automatically after completion of oligonucleotide synthesis. In this case, the automated synthesizer essentially remains idle for the 60 to 90 min required for the cleavage of the succinyl linker arm by ammonium hydroxide. A very labile oxalic acid linker arm, which can be cleaved in only a few seconds, was introduced for the synthesis of base-sensitive oligonucleotide modifications (Alul et al., 1991). Unfortunately, the oxalyl linker proved to be too labile for routine use, because of spontaneous cleavage during storage at room temperature. Therefore, a linker arm that was more stable than oxalic acid but more easily cleaved than succinic acid was sought. This resulted in the introduction of hydroquinoneO,O′-diacetic acid (HQDA or Q-linker), S.8, as a new all-purpose linker arm (Pon and Yu, 1997a). Using Q-linker, ammonium hydroxide cleavage requires only 2 min for oligodeoxyribonucleotides, 5 min for oligoribonucleotides, and 10 min for phosphorothioate-modified oligonucleotides. Cleavage can also be performed using a number of other reagents (Table 3.2.1)
3.2.18 Current Protocols in Nucleic Acid Chemistry
Table 3.2.1 Conditions for Cleavage of HQDA
Amount of cleavage (%) at room temperature Reagent 40% aqueous MeNH2/NH4OH, 1:1 (v/v) 0.05 M K2CO3 in MeOH 9.3 bar (140 psi) ammonia gas Triethylamine trihydrofluoride (neat) 1 M tetrabutylammonium fluoride in THF 1:2:1 (v/v/v) t-butylamine/MeOH/H2O NH3 in MeOH (saturated) 5% NH4OH in MeOH 20% piperidine/DMF 0.5 M (7.5%) DBU in pyridine 1:1 (v/v) triethylamine/EtOH
1 min 100 100 94 46 36 97 27 9
to accommodate base-sensitive oligonucleotides. The stability of HQDA is sufficient for long-term storage at room temperature (>3 years) and for virtually complete resistance to oligonucleotide synthesis conditions. Prepacked synthesis columns with Q-linker are now commercially available (Glen Research), and if the current emphasis on faster production and milder deprotection continues, HQDA may eventually replace the succinyl linker.
Compound Characterization Characterization of products attached to the surface of insoluble supports is very difficult, if not impossible. This is especially so with a rigid, nonswelling, inorganic support such as controlled-pore glass beads. Therefore, the success of nucleoside derivatization reactions is usually determined indirectly by examining cleavage products. This can range from the comprehensive characterization of nucleoside or oligonucleotide products hydrolyzed from the support to the simple trityl analysis described in this unit. The dimethoxytrityl protecting group, commonly used to block the 5′-OH of nucleosides, provides a convenient and highly sensitive marker (Support Protocol 1). There is a 1:1 relationship between the amount of orange-colored dimethoxytrityl cation released and the amount of nucleoside present, so nucleoside loading of any support can be easily determined by a colorimetric measurement. Because the trityl group is easily hydrolyzed (by mild acid) from the support and generates such a distinctive and easily quantitated species, it is much simpler and faster to perform a trityl analysis than a nucleoside analysis.
5 min
15 min
100 100 83
99
75 27
98 63
2
4
60 min
98 15 7 5
t1/2 <10 sec <10 sec ≈10 sec ≈1 min ≈2 min ≈1 min ≈3 min ≈11 min ≈3 hr ≈16 hr
The trityl results can be quite accurate and reproducible, as long as care is taken to avoid the following three possibilities. First, the trityl colors are most stable in the presence of excess acid. Therefore, dilutions should be performed by adding acidic deprotection reagent and not plain solvent. Second, the orange color is quenched by traces of protic solvents, such as water or alcohols. Thus glassware and cuvettes should be clean and dry before use. Finally, the trityl assay does not distinguish between nucleosides (or trityl groups) that are covalently and noncovalently bound. Thus it is possible for supports that have not been adequately washed to give false trityl results. This is not usually a problem, because most unbound reagents are easily removed by a methanol wash. Positively charged amino supports can, however, sometimes bind negatively charged nucleoside hemiesters strongly enough to resist washing. This is one reason why the derivatization protocols recommend checking the trityl loading both before and after acetylation of the surface amino groups (i.e., the capping step with Cap A and Cap B reagents). The number of functional groups on the surface of the support can also be determined by trityl analysis, if they can be derivatized with trityl groups. Amino and/or hydroxyl surface loadings are easily determined through the dimethoxytrityl derivatization procedure described in Support Protocol 2 (Reddy and Voelker, 1988). Surface carboxylic acids groups cannot be directly tritylated, so an ester derivative containing a chromophore is made instead. p-Nitrophenyl ester derivatives (Matteucci and Caruthers, 1981; Damha et al., 1990) can be used to measure succinic acid groups
Synthesis of Unmodified Oligonucleotides
3.2.19 Current Protocols in Nucleic Acid Chemistry
(Support Protocol 3), but this method does not work for HQDA groups. Instead, a ligand bearing a monomethoxytrityl group on the amino end of 6-amino-1-hexanol is used as the chromophore (Support Protocol 4). It is generally useful to know the extent of surface modification before proceeding with any nucleoside derivatization. This ensures that the support functionalization was successful and that the desired degree of nucleoside loading can be achieved. Characterization of nucleoside hemiesters, either succinyl or HQDA, can be most easily performed by thin-layer chromatography (TLC), especially if samples of authentic material are available for comparison. Even if authentic samples are not available, observation of the complete conversion of the starting nucleoside into either slower-moving products (hemiesters) or faster-moving products (diesters) is sufficient for most casual users. TLC analysis of the nucleoside hemiesters along with satisfactory trityl results from the derivatized support are usually sufficient if the identity of the starting nucleosides is well established (i.e., commercially available material). Of course, researchers working with novel compounds will also require the usual NMR, UV, and mass or elemental composition analyses to sufficiently confirm structures for publication.
Critical Parameters and Troubleshooting Handling of the support Insoluble supports must be handled carefully to prevent particle fracturing and the formation of fines because these small particles can plug filters or create higher back pressures in synthesis columns. Although CPG particles are much more durable than the silica gel supports previously used, it is still important to avoid the use of stirrers (magnetic or otherwise) when performing coupling reactions. A shaker, such as a wrist-action shaker, is the preferred method for keeping reaction mixtures agitated.
Attachment of Nucleosides to Solid-Phase Supports
Quality reagents and solvents Lower-than-expected support loadings are obtained if the coupling reagents (DEC or HBTU) are more than several years old or have been exposed to moisture contamination. These reagents should be kept at −20°C for long-term storage and not opened until warmed to room temperature to prevent contamination from moisture condensation. A more common prob-
lem is the presence of moisture in the anhydrous reactions. Both pyridine and acetonitrile are quite hygroscopic, and storage over molecular sieves is strongly recommended. Although the coupling reactions described in this unit are not extremely sensitive to moisture contamination, because a large excess of coupling reagent is used in each protocol, excessive moisture will cause poor results. Reagent concentration One of the most important factors in reactions involving solid-phase supports is the solution concentration, and the easiest way to improve results, without consuming more of the reagents, is to increase the concentration of either nucleoside or coupling reagent by reducing the reaction volume. This method is limited, however, by the volume required to suspend the support. Typically, 4 to 5 mL of solution per gram of LCAA-CPG is the minimum required. Generally, the protocols described in this unit do not depend strongly on reagent concentration, and reasonable results can be obtained without careful measurement of solvent volume. Researchers should be aware, however, that excessive solvent will lower results through both dilution effects and increased moisture content (if solvents are not completely anhydrous). Ribonucleosides Although, for simplicity, the figures and text in this unit refer only to 2′-deoxyribonucleosides, the coupling procedures are also applicable to ribonucleosides. When a ribonucleoside is attached to an insoluble support, it is of no consequence whether the linkage is through the 2′ or 3′ hydroxyl group. Therefore, ribonucleosides with a single unprotected 2′ or 3′ hydroxyl group can be used, as well as mixtures of the 2′ and 3′ isomers. It is also possible to use ribonucleosides with both 2′ and 3′ hydroxyl groups unprotected (i.e., only 5′and N-protecting groups). In this case, after attachment of the nucleoside to the support through one hydroxyl group, the adjacent unreacted hydroxyl group is acetylated by the same capping step used to acetylate unreacted amino groups. HBTU side reaction with amines Researchers should also be aware that the HBTU coupling reagent can form a Schiff base with amino groups (Gausepohl et al., 1992; Story and Aldrich, 1994). Therefore, this reagent should never be added to LCAA-CPG in
3.2.20 Current Protocols in Nucleic Acid Chemistry
the absence of the carboxylic acid component of the coupling reaction (which normally, and very rapidly, reacts preferentially with HBTU). Thus only 0.9 eq of HBTU are used in the protocol for coupling HQDA to LCAA-CPG. Accidental incorporation of >1 eq of HBTU into this reaction results in conversion of surface amino groups to unreactive Schiff bases. Capping of unreacted amino groups on the support An essential part of each nucleoside coupling protocol is the blocking or capping of unreacted functional groups on the surface of the support. These unreacted groups are present because the amount of nucleoside added is almost always less than the total capacity of the support. In particular, it is very important to cap off any unreacted amino groups by acetylating them with acetic anhydride. Otherwise they will react with the phosphoramidite reagents delivered during subsequent oligonucleotide synthesis cycles to produce oligonucleotide fragments with a 3′ deletion (i.e., the terminal nucleoside will be missing). These deletion fragments will be attached to the solid support via a 3′-phosphoramidate linkage and can be released from the support by either acidic hydrolysis (cleavage of the phosphoramidate bond) or prolonged ammonium hydroxide hydrolysis (cleavage of the silyl bonds joining the linker arm to the LCAA-CPG surface). Although normal cleavage conditions (NH4OH, ≤1 hr, room temperature) do not release significant amounts of these 3′-deletion products, they still reduce coupling yields and oligonucleotide quality by steric hindrance and competition for reagents. The effectiveness of the support capping can be determined by comparing the amount of trityl color released from the first nucleoside with the amount of color released after the first phosphoramidite-coupling cycle (Pon et al., 1988). The second amount should be the same as, or slightly less than, the first amount, if coupling occurred only on the immobilized nucleoside (i.e., the yield of the coupling is ≤100%); however, if the second color is more intense than the first (i.e., apparent coupling yields of >100%), then unwanted coupling to the surface of the support is occurring. In this case, the supports should be capped again before further use. The pore size of the support For rigid, macroporous supports, such as the LCAA-CPG particles described in this unit, the pore size is the most important parameter. This
is because the pore size affects the amount of nucleoside loading that can be obtained and the coupling efficiency of subsequent chain extension steps during oligonucleotide synthesis. The smaller the pore size, the greater the surface area and the higher the nucleoside loading. Too small a pore size, however, results in poor coupling efficiency and limits the oligonucleotide length that can be synthesized. LCAACPG supports with 500-Å pores are good choices for most oligonucleotide syntheses (shorter than ∼50 bases), and LCAA-CPG supports with 1000-Å pores are recommended for the synthesis of oligonucleotides longer than ∼50 bases (see UNIT 3.1).
Anticipated Results The amount of nucleoside attached to the surface of the support is generally not very critical, because most oligonucleotide synthesis columns can produce a great deal more product than required. Generally, however, LCAA-CPG supports are prepared with nucleoside loadings in the 30- to 40-µmol/g range; and the protocols in this unit should produce supports like these. This amount of loading gives excellent results for all but the very longest oligonucleotides. The greater the nucleoside loading, the more crowded the surface of the support; and this crowding appears to have an effect on coupling efficiency, especially with nucleoside loadings >50 µmol/g. Therefore, it is not necessary, and actually detrimental, to try to obtain nucleoside loadings greater than ∼40 to 50 µmol/g. For very long oligonucleotides (75 to 150 bases), use 1000-Å-pore-sized LCAA-CPG supports with only 5 to 10 µmol/g of nucleoside. The lower nucleoside loading presumably improves coupling efficiency by reducing steric hindrance and increasing the amount of excess coupling reagents. With this low loading, however, the maximum capacity of a synthesis column is reduced. It is also important to ensure that the synthesis reagents are of the highest quality and that the acidic detritylation conditions are minimized (to avoid unwanted depurination and chain cleavage) when preparing long oligonucleotides.
Time Considerations The total time required to perform each support derivatization varies between 1 and 4 days; however, the time required for the actual setup and workup is much less. Setup time for most reactions is very short (<15 min), because the reactants just need to be weighed and added
Synthesis of Unmodified Oligonucleotides
3.2.21 Current Protocols in Nucleic Acid Chemistry
to a single flask. Workup of the support derivatization reactions is also very fast (<10 min), because this involves only filtering off the support and washing it with solvent. Use of a vacuum aspirator to pull the solvent through the support during the wash steps makes very quick work. After the final dichloromethane wash recommended for each procedure, the LCAACPG beads are left to dry uncovered at room temperature. Evaporation of the dichloromethane solvent is quite fast, and the beads are usually sufficiently dry for trityl analysis or other use within 10 to 20 min. Also, preparations of the succinyl- or HQDA-derivatized LCAA-CPG supports should be performed in bulk, and the intermediate carboxyl derivatized supports saved for future nucleoside coupling reactions. Thus conversion of the LCAA-CPG support to a carboxylic acid–derivatized support does not need to be repeated each time. The time required for the synthesis of nucleoside hemiesters (of either succinic acid or HQDA) is more substantial, because these products must be worked up by conventional solution phase methods. An experienced chemist will require at least 2 to 3 days to synthesize and purify each reaction, and this process must be repeated for each nucleoside required. Therefore, these products are best purchased, if synthetic resources are not readily available. Once the support derivatization methods are mastered, it is very easy to prepare sufficient material for a large number of oligonucleotide syntheses. For example, a 5-g support preparation is sufficient for about 500 0.2-µmol-scale syntheses or about 2000 40-nmol-scale syntheses.
Literature Cited
E fi mov, V. A . , K a l in k in a , A.L., an d Chakhmakhcheva, O.G. 1993 . Dipentafluorophenyl carbonate—A reagent for the synthesis of oligonucleotides and their conjugates. Nucl. Acids Res. 21:5337-5344. Gait, M.J., Singh, M., and Sheppard, R.C. 1980. Rapid synthesis of oligodeoxyribonucleotides IV. Improved solid phase synthesis of oligodeoxyribonucleotides through phosphotriester intermediates. Nucl. Acids Res. 8:1081-1097. Gausepohl, H., Pieles, U., and Frank, R.W. 1992. Schiff’s base analog formation during in situ activation by HBTU and TBTU. In Peptides: Chemistry and Biology (J.A. Smith and J.E. Rivier, eds.) pp. 523-524. ESCOM, Leiden. Horn, M. and Novak, C. 1987. A monitoring and control chemistry for solid-phase peptide synthesis. Am. Biotechnol. Lab. 5:12-21. Koster, H., Stumpe, A., and Wolter, A. 1983. Polymer support oligonucleotide synthesis 13: Rapid and efficient synthesis of oligodeoxynucleotides on porous-glass support using triester approach. Tetrahedron Lett. 24:747-750. Kumar, P., Ghosh, N.N., Sadana, K.L., Garg, B.S., and Gupta, K.C. 1993. Improved methods for 3′-O-succinylation of 2′-deoxyribo- and ribonucleosides and their covalent anchoring on polymer supports for oligonucleotide synthesis. Nucleos. Nucleot. 12:565-584. Kumar, P., Sharma, A.K., Sharma, P., Garg, B.S., and Gupta, K.C. 1996. Express protocol for functionalization of polymer supports for oligonucleotide synthesis. Nucleos. Nucleot. 15:879888. Markiewicz, W.T., Groger, G., Rosch, R., Zebrowska, A., Markiewicz, M., Klotz, M., Hinz, M., Godzina, P., and Seliger, H. 1997. A new method of synthesis of fluorescently labelled oligonucleotides and their application in DNA sequencing. Nucl. Acids Res. 25:3672-3680.
Alul, R.H., Singman, C.N., Zhang, G.R., and Letsinger, R.L. 1991. Oxalyl-CPG—A labile support for synthesis of sensitive oligonucleotide derivatives. Nucl. Acids Res. 19:1527-1532.
Matteucci, M.D. and Caruthers, M.H. 1981. Synthesis of deoxyoligonucleotides on a polymer support. J. Am. Chem. Soc. 103:3185-3191.
Bhongle, N.N. and Tang, J.Y. 1995. A convenient and practical method for derivatization of solid supports for nucleic acid synthesis. Synth. Commun. 25:3671-3679.
Miyoshi, K.-I., Miyake, T., Hozumi, T., and Itakura, K. 1980. Solid-phase synthesis of polynucleotides. II. Synthesis of polythymidylic acids by the block coupling phosphotriester method. Nucl. Acids Res. 8:5473-5489.
Chow, F., Kempe, T., and Palm, G. 1981. Synthesis of oligodeoxyribonucleotides on silica gel support. Nucl. Acids Res. 12:2807-2817. Attachment of Nucleosides to Solid-Phase Supports
Dourtoglou, V., Gross, B., Lambropoulou, V., and Zio u d r ou , C. 1 984. O-BenzotriazolylN,N,N′,N′-tetramethyluronium hexafluorophosphate as a coupling reagent for the synthesis of peptides of biological interest. Synthesis 572-574.
Damha, M.J., Giannaris, P.A., and Zabarylo, S.V. 1990. An improved procedure for derivatization of controlled pore glass beads for solid-phase oligonucleotide synthesis. Nucl. Acids Res. 18:3813-3821.
Montserrat, F.X., Grandas, A., and Pedroso, E. 1993. Predictable and reproducible yields in the anchoring of DMT-nucleoside-succinates to highly loaded aminoalkyl-polystyrene. Nucleos. Nucleot. 12:967-971. Ogilvie, K.K. and Kroeker, K. 1971. Synthesis of oligothymidylates on an insoluble polymer support. Can. J. Chem. 50:1211-1215.
3.2.22 Current Protocols in Nucleic Acid Chemistry
Pon, R.T. 1993. Solid-phase supports for oligonucleotide synthesis. In Protocols for Oligonucleotides and Analogs (S. Agrawal, ed.) pp. 465496. Humana Press, Totowa, N.J.
Reddy, M.P. and Voelker, P.J. 1988. Novel method for monitoring the coupling efficiency in solid phase peptide synthesis. Int. J. Pept. Protein Res. 31:345-348.
Pon, R.T. and Yu, S. 1997a. Hydroquinone-O,O′diacetic acid (‘Q-linker’) as a replacement for succinyl and oxalyl linker arms in solid phase oligonucleotide synthesis. Nucl. Acids Res. 25:3629-3635.
Reddy, M.P., Rampal, J.B., and Beaucage, S.L. 1987. An efficient procedure for the solid phase tritylation of nucleosides and nucleotides. Tetrahedron Lett. 28:23-26.
Pon, R.T. and Yu, S. 1997b. Rapid automated derivatization of solid-phase supports for oligonucleotide synthesis using uronium or phosphonium coupling reagents. Tetrahedron Lett. 38:3331-3334. Pon, R.T., Usman, N., and Ogilvie, K.K. 1988. Derivatization of controlled pore glass beads for solid phase oligonucleotide synthesis. Biotechniques 6:768-775. Pon, R.T., Yu, S., Guo, Z., Yang, X., and Sanghvi, Y.S. 1999. Reusable solid-phase supports for oligonucleotide synthesis using hydroquinoneO,O′-diacetic acid (Q-Linker). Nucleos. Nucleot. In press. Pon, R.T., Yu, S., Guo, Z., and Sanghvi, Y.S. 1999. Multiple oligodeoxyribonucleotide syntheses on a reusable solid-phase CPG support via the hydroquinone-O,O′-diacetic acid (Q-Linker) linker arm. Nucleic Acids Res. 27:1531-1538.
Sharma, P., Sharma, A.K., Malhotra, V.P., and Gupta, K.C. 1992. One pot general method for the derivatization of polymer support for oligonucleotide synthesis. Nucl. Acids Res. 20:4100-4100. Story, S.C. and Aldrich, J.V. 1994. Side-product formation during cyclization with HBTU on a solid support. Int. J. Pept. Protein Res. 43:292296. Walsh, A.J., Clark, G.C., and Fraser, W. 1997. A direct and efficient method for derivatisation of solid supports for oligonucleotide synthesis. Tetrahedron Lett. 38:1651-1654. Yip, K.F. and Tsou, K.C. 1971. A new polymer support method for the synthesis of ribooligonucleotide. J. Am. Chem. Soc. 93:3272-3276.
Contributed by Richard T. Pon University of Calgary Calgary, Alberta, Canada
Synthesis of Unmodified Oligonucleotides
3.2.23 Current Protocols in Nucleic Acid Chemistry
Synthetic Strategies and Parameters Involved in the Synthesis of Oligodeoxyribonucleotides According to the Phosphoramidite Method The study of nucleic acids has, over the years, driven the development of fundamental methodologies necessary for the examination of their structure and chemistry. The ability to produce substantial quantities of sequence-defined synthetic DNA has been invaluable to nucleic acid research. Synthetic oligonucleotides have facilitated structural investigations and provided a better understanding of the interactions between nucleic acids and, for example, binding and/or modifying proteins. In addition, synthetic oligodeoxyribonucleotides have been extensively applied to the preparation of primers for enzymatic synthesis, amplification, and DNA sequencing. They have also been commonly used in site-directed mutagenesis experiments and as hybridization probes for diagnostic purposes. More recently, modified synthetic oligonucleotides have been targeted to cellular messenger RNAs in an attempt to control gene expression and develop therapeutic agents against various types of cancer and human infectious diseases (Beaucage and Iyer, 1993; Crooke and Bennett, 1996). On this basis, the availability of synthetic oligonucleotides has undoubtedly paved the way to the biotechnology revolution. The basic strategy in oligonucleotide synthesis resembles that of the stepwise synthesis of polypeptides. Typically, a functionally protected nucleotide monomer is linked to a growing oligonucleotide chain that is then chemoselectively deprotected and allowed to couple with the next nucleotide monomer of the desired sequence. Although the coupling of nucleotide monomers was traditionally carried out in solution according to the phosphodiester (Khorana, 1968) and phosphotriester (Letsinger and Ogilvie, 1969) methods, these strategies ultimately culminated in the development of modern automated techniques using the phosphoramidite method (Beaucage and Caruthers, 1981; Caruthers et al., 1987a). Given the major impact the phosphoramidite approach has had on the synthesis of oligonucleotides since its inception in the early 1980s, this commentary provides an overview of the parameters affecting the performance of the method and a number of compatible
UNIT 3.3
strategies for the preparation of oligodeoxyribonucleotides. Milestones that led to the discovery of the phosphoramidite method for oligodeoxyribonucleotide synthesis are chronologically reported. Alternate strategies to the preparation of deoxyribonucleoside phosphoramidites are then described to underscore the versatility with which these synthons can be obtained. The mechanisms of deoxyribonucleoside phosphoramidite activation and the factors affecting condensation rates are discussed along with the importance of “capping” and oxidation reactions toward defining optimal performance conditions for oligonucleotide synthesis. Finally, alternate phosphoramidite methods to oligodeoxyribonucleotide synthesis and additional deprotection strategies are presented to demonstrate facile access to synthetic oligonucleotides for a variety of applications. This unit is intended to complement APPENDIX 3C on the synthesis and purification of oligonucleotides.
ACCOUNTS OF CHEMICAL RESEARCH IN DNA OLIGONUCLEOTIDE SYNTHESIS A major advance in the chemical synthesis of oligodeoxyribonucleotides was accomplished in the mid-1970s through the development of the “phosphite triester” method (Letsinger and Lunsford, 1976). This approach relied on the use of P(III) chemistry and entailed the reaction of 5′-O-phenoxyacetyl thymidine S.1 with 2,2,2-trichloroethyl phosphorodichloridite S.2 to generate the corresponding deoxyribonucleoside-3′-O-phosphorochloridite S.3 and variable amounts of (3′→3′)-dinucleoside phosphite triester S.4 (Fig. 3.3.1). Addition of 3′-O-monomethoxytrityl thymidine to the reaction mixture gave, after treatment with aqueous iodine, the dinucleoside phosphate triester S.5 along with the symmetrical (3′→3′)- and (5′→5′)dinucleoside phosphate triesters S.6 and S.7, respectively. The formation of S.7 presumably resulted from the reaction of 3′-Omonomethoxytrityl thymidine with unreacted S.2. It should be noted that the phosphite triester
Contributed by Serge L. Beaucage and Marvin H. Caruthers Current Protocols in Nucleic Acid Chemistry (2000) 3.3.1-3.3.20 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Unmodified Oligonucleotides
3.3.1
Thy
PhOCH2COO Thy
PhOCH2COO
O
Cl3CCH2OPCl2 2
Thy
PhOCH2COO
2,6-lutidine/THF
OH 1
Cl
O
O O P
Cl3CCH2O
O P
O
OCH2CCl3
O
PhOCH2COO
Thy
3 4
Thy
HO
O
Thy
PhOCH2COO
PhOCH2COO
O
MMTrO
Thy O
1. O
MMTrO 2.
O
I2/H2O
Thy
O
PhOCH2COO
MMTrO MMTr, 4-methoxytrityl Ph, phenyl
O
O O P OCH2CCl3 O
O P OCH2CCl3
5
O Thy
O P OCH2CCl3 O
O
Thy
O MMTrO
Thy 6
7
Figure 3.3.1 The phosphite triester method to oligodeoxyribonucleotide synthesis.
and inert atmosphere. Although nucleoside chlorophosphites/tetrazolides led to the rapid and efficient preparation of DNA oligonucleotides on silica supports, the technical hardship associated with handling these synthons impeded automation of oligodeoxyribonucleotide syntheses. These problems vanished when deoxyribonucleoside phosphoramidites were developed in the early 1980s (Beaucage and Caruthers, 1981). These synthons were stable to air oxidation and hydrolysis under normal laboratory conditions but readily activated under mild acidic conditions to form internucleotide linkages in near-quantitative yields. Because of these properties, deoxyribonucleoside phosphoramidites have been conducive to automated oligonucleotide synthesis on polymeric supports and are still the preferred synthons for such syntheses. The preparation of these reagents was straightforward and involved the reaction of protected deoxyribonucleosides, such as S.11, with chloro-(N,N-dimethy-
approach to oligonucleotide synthesis was remarkably rapid; the preparation of S.5 was complete within 1 hr. In the mid-1970s, the formation of internucleotide linkages with such kinetics was unmatched. Later, the deoxyribonucleoside chlorophosphite S.8 (Fig. 3.3.2) was synthesized and reacted with deoxyribonucleosides covalently attached to a silica support (S.9) through a 3′-O-succinate linkage (Matteucci and Caruthers, 1981; see also UNITS 3.1 and 3.2). The dinucleoside phosphate triester S.10 was produced in yields > 90%. Addition of 1H-tetrazole to chlorophosphite S.8 significantly improved condensation rates and coupling yields. The application of nucleoside chlorophosphites or the corresponding tetrazolides to solid-phase synthesis of oligonucleotides was, however, tedious. Because of inherent sensitivity to moisture and air oxidation, the preparation of these reagents from reactive bifunctional phosphitylating agents had to be performed at −78°C under rigorously anhydrous conditions
B
DMTrO B
DMTrO
O
B
HO
O
I2/H2O
O P
O OMe
O
8
9
O O P OMe
O
Cl
O
N H
P
B
O
O O O O
Oligodeoxyribonucleotide Synthesis Using the Phosphoramidite Method
DMTr, 4,4'-dimethoxytrityl B, Thy or any N-protected nucleobase Me, methyl P, solid support
N H
P
10
Figure 3.3.2 Application of the phosphite triester approach to solid-phase DNA synthesis.
3.3.2 Current Protocols in Nucleic Acid Chemistry
Cl DMTrO
B O OH
Me2N
P
OMe
B
DMTrO
O
12
i-Pr2NEt
Me2N
11
O P
OMe 13
Et, ethyl i-Pr, isopropyl
Figure 3.3.3 Preparation of deoxyribonucleoside phosphoramidite monomers.
lamino)methoxyphosphine S.12 and N,N-diisopropylethylamine (Fig. 3.3.3). The rapid reaction afforded deoxyribonucleoside phosphoramidites S.13 without the wasteful formation of (3′→3′)-dinucleoside phosphite triesters. The phosphoramidites S.13 were isolated by conventional laboratory techniques and stored as dry powders (Beaucage and Caruthers, 1981). The phosphoramidite method is distinctive in that it enables the conversion of relatively stable deoxyribonucleoside phosphoramidite derivatives into highly reactive intermediates suitable for DNA oligonucleotide synthesis. For example, addition of 1H-tetrazole to the phosphoramidite S.13 and 3′-O-levulinyl thymidine S.14 in dry acetonitrile generated the dinucleoside phosphite triester S.15 (Fig. 3.3.4) in almost quantitative yields within a few minutes, according to 31P-NMR spectroscopy (Beaucage and Caruthers, 1981). This methodology has been successfully applied to the solid-phase synthesis of oligodeoxyribonucleotides of varying chain lengths (Caruthers et al., 1982; Josephson et al., 1984). The sensitivity of phosphoramidite S.13 to a slightly acidic environment, however, precluded silica gel purification. Consequently, the chemical stability of crude S.13 in acetonitrile was variable, and the use of this type of phosphoramidite in automated systems not always reliable. Thus deoxyribonucleoside phosphoramidites with N,N-dialkylamino groups different from N,Ndimethylamino were investigated as potential alternatives to S.13 in automated solid-phase
DNA synthesis (Adams et al., 1983; McBride and Caruthers, 1983). For instance, the deoxyribonucleoside phosphoramidite S.16 (Fig. 3.3.5), unlike S.13, survived silica gel purification and, as a result, consistently showed stability in acetonitrile solutions for several days without significant decomposition (Adams et al., 1983). This class of phosphoramidites enabled reliable oligodeoxyribonucleotide syntheses (51-mers) on controlled-pore glass (CPG). In the early 1980s, these 51-mers were the largest DNA segments ever chemically synthesized. One drawback to the use of S.16 in automated oligonucleotide synthesis is the noxious thiophenolate treatment required for postsynthesis removal of the methyl phosphate protecting groups (Daub and van Tamelen, 1977). Although demethylation of methyl phosphotriesters can be effected by 2-mercaptobenzothiazole (S.17; Andrus and Beaucage, 1988) or disodium 2-carbamoyl-2-cyanoethylene-1,1dithiolate (S.18; Dahl et al., 1990) under odorless conditions, this deprotection step was inconvenient because it added to the time and reagents needed for oligonucleotide processing (Fig. 3.3.5). In an effort to simplify and shorten postsynthesis deprotection protocols, the deoxyribonucleoside phosphoramidite S.19 (Sinha et al., 1984) was prepared under conditions similar to those originally reported by Beaucage and Caruthers (Fig. 3.3.5). This phosphoramidite derivative was more stable than the methyl phosphoramidite S.16 in wet acetonitrile (Zon et al., 1985). Furthermore, phosphoramidite S.19 generated oligonucleotides from which the 2-cyanoethyl phosphate protecting groups are cleaved under the basic conditions required for deprotection of the nucleobase protecting groups (Tener, 1961; Letsinger and Mahadevan, 1966; Letsinger and Olgilvie, 1969). Such a convenient “single step–single reagent” postsynthesis deprotection protocol led to the widespread acceptance of phosphoramidite S.19 for automated solidphase oligonucleotide synthesis.
B
DMTrO HO
O
Thy O
1H-tetrazole
13
O P OCH3
MeCN
LevO
O
O
Thy
14 LevO Lev, levulinyl
15
Figure 3.3.4 Activation of deoxyribonucleoside phosphoramidites toward oligonucleotide synthesis.
Synthesis of Unmodified Oligonucleotides
3.3.3 Current Protocols in Nucleic Acid Chemistry
B
DMTrO
O
SH
i-Pr2N
O P
B
DMTrO
O
Na
− +Na
O
S OMe
− +
S
NC
N
S NH2
17
O P
i-Pr2N
18
OCH2CH2CN 19
16
Figure 3.3.5 Deoxyribonucleoside phosphoramidite monomers with improved stability properties and reagents for the deprotection of methyl phosphate triesters.
ALTERNATE STRATEGIES TO THE PREPARATION OF DEOXYRIBONUCLEOSIDE PHOSPHORAMIDITES The basicity and/or nucleophilicity of aminophosphine derivatives such as S.20 depends on the nature of the functional groups bound to phosphorus and on the interactions each of these groups might have with the vacant d orbital of the phosphorus atom through pπ-dπ overlap (Boudjebel et al., 1975). The bis-(dialkylamino) phosphine S.20 (Fig. 3.3.6) would, therefore, be more prone to protonation by a weak acid than would the deoxyribonucleoside phosphoramidite S.21, because it has been demonstrated that an alkoxy group contributes less to pπ-dπ interactions than does a dialkylamino group (Mathis et al., 1974). This concept was convincingly tested when the deoxyribonucleoside S.11 was reacted with bis-(pyrrolidino)methoxyphosphine S.20 and the weak acid 4,5-dichloroimidazole (Beaucage, 1984; Moore and Beaucage, 1985) to generate the deoxyribonucleoside phosphoramidite S.21 within 10 min, in yields exceeding 86% (Fig. 3.3.6). Presumably because of weaker pπ-dπ interactions, further activation of S.21 by 4,5-
dichloroimidazole produced only a small amount of (3′→3′)-dinucleoside phosphite triester (< 10%). Thus, as soon as it was generated, phosphoramidite S.21 was immediately activated by addition of 1H-tetrazole and used in the manual synthesis of an oligodeoxyribonucleotide (22-mer). This strategy eliminated the tedious isolation and purification of deoxyribonucleoside phosphoramidites and problems associated with the stability of phosphoramidite solutions. Despite these appealing features, this novel approach to the synthesis of deoxyribonucleoside phosphoramidites and oligonucleotides has never been automated, perhaps because of engineering limitations. In this context, it has been shown that reaction of the deoxyribonucleoside S.11 with bis-(N,Ndiisopropylamino)methoxyphosphine S.22 (Fig. 3.3.7) and limiting amounts of 1H-tetrazole or its N,N-diisopropylammonium salt afforded the corresponding deoxyribonucleoside phosphoramidite S.16 in isolated yields varying between 82% and 92% (Barone et al., 1984; Lee and Moon, 1984). This method is definitely recommended for the preparation of a variety of 2-cyanoethyl deoxyribonucleoside phosphoramidite derivatives (like S.19) from phos-
B
DMTrO
11 N
N P
O
4,5-dichloroimidazole O OMe
N-methylpyrrolidone N
P
OMe 21
20
Figure 3.3.6 Chemoselective preparation of deoxyribonucleoside phosphoramidites in situ using bis-(pyrrolidino)methoxyphosphine and 4,5-dichloroimidazole.
B
DMTrO 11
i-Pr2N P i-Pr2N OR
22 R = CH3 23 R = CH2CH2CN
Oligodeoxyribonucleotide Synthesis Using the Phosphoramidite Method
O
N,N-diisopropylammonium tetrazolide or 1H-tetrazole
i-Pr2N
O P
OR
16 R = CH3 19 R = CH2CH2CN
Figure 3.3.7 Chemoselective preparation of deoxyribonucleoside phosphoramidites using bis(N,N-diisopropyl)alkoxyphosphine and limiting amounts of 1H-tetrazole or its N,N-diisopropylammonium salt.
3.3.4 Current Protocols in Nucleic Acid Chemistry
NEt2 Et2N
P
NEt2
B
DMTrO
B
DMTrO
O
24
O
CF3CONH(CH2)4OH
11 O
N,N-diethylammonium tetrazolide
P
Et2N
NEt2
25
N,N-diethylammonium tetrazolide
O Et2N
P
O(CH2)4NHCOCF3 26
Figure 3.3.8 Preparation of deoxyribonucleoside phosphoramidites using hexaethylphosphorus triamide and limiting amounts of N,N-diethylammonium tetrazolide.
phorodiamidite S.23 (Fig. 3.3.7) because of the mildness and high chemoselectivity of the phosphinylation reaction. Along similar lines, reaction of S.11 with hexaethylphosphorus triamide S.24 (Fig. 3.3.8) and an equimolar amount of the N,N-diethylammonium salt of 1H-tetrazole cleanly gave the nucleoside phosphorodiamidite S.25 (Yamana et al., 1989). Addition of, for example, 4-(N-trifluoroacetylamino)butan-1-ol to the reaction mixture produced the deoxyribonucleoside phosphoramidite S.26 (Fig. 3.3.8) in isolated yields exceeding 90% (Wilk et al., 1997). The versatility of this procedure allows facile access to phosphoramidites different from the conventional methyl or 2-cyanoethyl deoxyribonucleoside phosphoramidites.
ACTIVATION OF DEOXYRIBONUCLEOSIDE PHOSPHORAMIDITES The elegance of the phosphoramidite approach to oligodeoxyribonucleotide synthesis emanates from the transformation of relatively stable deoxyribonucleoside phosphoramidite derivatives to highly reactive intermediates for rapid and efficient chain extension reactions. Such a conversion is catalyzed by weak acids. For example, activation of phosphoramidite S.13 with N,N-dimethylaniline hydrochloride (pKa ≈ 5.15) generated the deoxyribonucleoside chlorophosphite S.27 (Fig. 3.3.9) in quantitative yield according to 31P-NMR spectroscopy (Beaucage and Caruthers, 1981). Because most tertiary amine hydrochlorides are hygroscopic, these could not be used in routine
B
DMTrO
O
N,N-dimethylaniline hydrochloride 13 O
CHCl3 Cl
P
OMe 27
Figure 3.3.9 Activation of deoxyribonucleoside phosphoramidites with N,N-dimethylaniline hydrochloride.
automated oligodeoxyribonucleotide synthesis because anhydrous conditions are absolutely necessary for optimum coupling reactions. Instead, the nonhygroscopic weak acid 1H-tetrazole has been and is still extensively used as an activator for deoxyribonucleoside phosphoramidites. In an incisive experiment, Seliger and Gupta (1985) demonstrated that activation of the solid-phase-linked deoxyribonucleoside phosphoramidite S.28 with 1Htetrazole generated the putative phosphorotetrazolide derivative S.29 (Fig. 3.3.10). Immediate condensation of S.29 with a deoxyribonucleoside covalently attached to a solid support (such as S.9) gave the dinucleoside phosphite triester S.30 in yields greater than 95%. The generation of S.29 during activation of deoxyribonucleoside phosphoramidites by 1Htetrazole was further investigated using diethoxy(N,N-diisopropylamino)phosphine S.31 and diethoxy-(N-tetrazolyl)phosphine S.32 (Fig. 3.3.11) as models (Berner et al., 1989). 31P-NMR spectrum of S.32 shows a resonance at 126 ppm. This resonance is also apparent when S.31 or phosphoramidite S.16 is mixed with 1H-tetrazole in acetonitrile (McBride and Caruthers, 1983). It should be noted that P-diastereomerically pure S.16 (Sp or Rp) rapidly epimerized at phosphorus upon activation with 1H-tetrazole and produced a diastereomeric deoxyribonucleoside phosphite triester when reacted with ethanol (Stec and Zon, 1984). These data indicate that activation of deoxyribonucleoside phosphoramidites with 1H-tetrazole occurs through a rapid and reversible protonation followed by a slower and reversible formation of the phosphorotetrazolide intermediate S.29. Paradoxically, activation of the nucleoside bicyclic phosphoramidites S.33-S.35 (Fig. 3.3.12) with 1H-tetrazole proceeded with low epimerization at phosphorus and led to highly stereoselective syntheses of oligonucleoside phosphorothioates (Guo et al., 1998; Iyer et al., 1998). A higher energy barrier to pseudorotation is apparently responsible for the reduced P-epimerization of these oxazaphospholidi-
Synthesis of Unmodified Oligonucleotides
3.3.5 Current Protocols in Nucleic Acid Chemistry
B
DMTrO
MeO
O P
MeCN MeO
O
B
DMTrO
O
1H-tetrazole Et N CH2
B
DMTrO
O
O P
O
9 MeO
P
O
O
N N N N
B O
O
29 P
O
28
P
N H
30
Figure 3.3.10 Mechanism of the activation of deoxyribonucleoside phosphoramidites by 1Htetrazole during solid-phase oligonucleotide synthesis.
nes during activation (Guo et al., 1998). It is somehow surprising that, although the conformationally restricted bicyclic oxazaphospholidines S.33 a n d S.34 allowed highly stereoselective syntheses of dinucleoside phosphorothioates (SP:RP > 9:1), the bicyclic phosphoramidite S.36 produced the same dinucleotides with only moderate stereoselectivity (RP:SP ≈ 3:1) (Guo et al., 1998). The bicyclic phosphoramidites S.33-S.35 a r e nonetheless promising candidates for the preparation of P-diastereomerically enriched oligonucleoside phosphorothioates. Reagents other than 1H-tetrazole have also been used to activate deoxyribonucleoside phosphoramidites, including N-methylimidazolium trifluoromethanesulfonate (Arnold et al., 1989); N-methylimidazole hydrochloride (Hering et al., 1985); pyridinium tetrafluoroborate (Brill et al., 1991); pyridinium chloride, pyridinium bromide, and pyridinium 4-methylbenzinesulfonate (Beier and Pfleiderer, 1999); N-methylanilinium trifluoroacetate (Fourrey and Varenne, 1984); N-methylanilinium trichloroacetate (Fourrey et al., 1987); benzimidazolium triflate (Hayakawa et al., 1996); imidazolium triflate (Hayakawa and Kataoka, 1998); pyridine hydrochloride/imidazole (Gryaznov and Letsinger, 1992); 5-trifluoromethyl-1H-tetrazole (Hering et al., 1985); 5-(4-nitrophenyl)-1H-tetrazole (Froehler and Matteucci, 1983); 5-(2-nitrophenyl)1H-tetrazole (Pon, 1987; Montserrat et al., 1994); 1-hydroxybenzotriazole (Claesen et al., 1984); 2,4,5-tribromo- and 2-nitro-imidazoles
(Xin and Just, 1996); benzotriazole and 5-chlorobenzotriazoles (Xin and Just, 1996); and 4,5dichloro-, 2-bromo-4,5-dicyano-, and 4,5-dicyano-imidazoles (Xin and Just, 1996). 5Ethylthio- and 5-benzylthio-1H-tetrazoles are also potent in the activation of phosphoramidites; these activators have been particularly useful in RNA synthesis (Wincott et al., 1995; Wu and Pitsch, 1998). It is important to note, however, that 1H-tetrazole and those more acidic activators (pKa < 4.8) can cleave the 5′-dimethoxytrityl (DMTr) group of deoxyribonucleoside phosphoramidites to a small extent and trigger the formation of activated dimers. The coupling of these dimers during chain extension produced oligonucleotides longer (n + 1) than the expected size (n) (Krotz et al., 1997a). The rates of DMTr deprotection by an activator depend on its acidity, exposure time, and nature of the nucleobase carrying the DMTr group (purines deprotect faster than pyrimidines). Typically, when the activation of deoxyribonucleoside phosphoramidites is performed by 1H-tetrazole for a period of 100 sec, ∼0.3%–0.9% of (n + 1)-mers is observed; however, when 1H-tetrazole is replaced by 5ethylthio-1H-tetrazole under similar conditions, ∼1.0%–4.3% of (n + 1)-mers are generated. It should, therefore, be understood that extended coupling times and the use of more acidic activators during solid-phase oligode-
B
DMTrO
O
O
O P
OEt OEt
Oligodeoxyribonucleotide Synthesis Using the Phosphoramidite Method
31
N N N
OEt N P OEt
O
X N
S
i-Pr2N P
B
DMTrO
33 X = H 34 X = OMe
O
O P
X N
R
35 X = H 36 X = OMe
32
Figure 3.3.11 Model compounds used in the study of phosphoramidite activation by 1Htetrazole.
Figure 3.3.12 Nucleoside bicyclic phosphoramidites for the preparation of P-diastereomerically enriched oligonucleoside phosphorothioates.
3.3.6 Current Protocols in Nucleic Acid Chemistry
oxyribonucleotide synthesis will result in lower recovery of full-length oligomers, because longer than full-length oligonucleotides will be produced. These observations prompted an extensive search for less-acidic, more-nucleophilic activators. It has been reported that 4,5-dicyanoimidazole (Xin and Just, 1996) is less acidic (pKa 5.2), more soluble, and more nucleophilic than 1H-tetrazole (Vargeese et al., 1998). Moreover, the usefulness of 4,5-dicyanoimidazole relates not only to efficient synthesis of oligonucleotides but also to the preparation of nucleoside phosphoramidites from phosphorodiamidite S.23. Even though 1H-tetrazole is still very popular as an activator for deoxyribonucleoside phosphoramidites, 4,5dicyanoimidazole is an attractive option for the activation of deoxyribonucleoside and ribonucleoside phosphoramites in solid-phase oligonucleotide synthesis.
pling yields averaging 45% on a CPG support (Sekine et al., 1986). Under similar conditions, activated S.38 required a coupling time of 60 min to produce yields of 65%–70% (Casale and McLaughlin, 1990). Such coupling yields are far below those obtained with conventional deoxyribonucleoside phosphoramidites (∼99%) and thus underscore the importance of steric hindrance when designing nucleobase protecting groups and/or modified nucleobases toward the synthesis of oligodeoxyribonucleotides and their analogues. One effective approach to lessening steric interferences is to increase the distance between the bulky entity and the phosphoramidite function by the use of flexible linkers. For example, unlike S.38, the deoxyribonucleoside phosphoramidite S.39 (Fig. 3.3.13) has been efficiently incorporated into oligonucleotides under the conditions used for standard 2-cyanoethyl deoxyribonucleoside phosphoramidites (Bergstrom and Gerry, 1994). Other factors influencing the coupling rates of activated deoxyribonucleoside phosphoramidites have been investigated by Dahl and his colleagues (1987). In a systematic study, it was observed that condensation rates varied with the nature of phosphoramidite Oalkyl and N,N-dialkylamino groups. Typically, coupling rates decreased according to the following order: O-methyl > O-(2-cyanoethyl) > O-(1-methyl-2-cyanoethyl) > O-(1,1-dimethyl-2-cyanoethyl) and N,N-diethylamino >
FACTORS AFFECTING THE CONDENSATION RATES OF DEOXYRIBONUCLEOSIDE PHOSPHORAMIDITES The steric bulk of specific guanine N-2 functional groups has been shown to affect significantly condensation rates and coupling efficiency of these deoxyribonucleoside phosphoramidite derivatives. Two classic examples illustrating this fact are the activation of phosphoramidites S.37 and S.38 with 1H-tetrazole (Fig. 3.3.13). In the case of activated S.37, a condensation time of 10 min generated cou-
O
O N N
O O
O
NH
N
DMTrO
N
P
N OBz
N
PixO
NH
O
NH N
NH
O
BzO OBz
OMe
i-Pr2N
P
OCH2CH2CN 38
37 4-NO2PhCH2CH2O N N
DMTrO
O
N N
N H
O
N N
O
i-Pr2N
P
Bz, benzoyl 4-NO2Ph, 4-nitrophenyl Pix, 9-phenylxanthen-9-yl
OCH2CH2CN 39
Figure 3.3.13 Deoxyribonucleoside phosphoramidites functionalized with nucleobase bulky groups.
Synthesis of Unmodified Oligonucleotides
3.3.7 Current Protocols in Nucleic Acid Chemistry
N,N-diisopropylamino > N-morpholino > Nmethylanilino. The effect of steric hindrance on coupling rates is further illustrated by activation of the 2′-substituted nucleoside phosphoramidites S.40-S.43 (Fig. 3.3.14) and their competitive condensations with thymidine covalently attached to a solid support (Kierzek et al., 1987). Dimer formation was quantitated and correlated with the condensation rates of S.40-S.43. The amount of dimers formed decreased when the groups at C-2′ increased in sizes; thus 2′-H > 2′-O-methyl > 2′-O-tetrahydropyranyl > 2′O-tert-butyldimethylsilyl (Kierzek et al., 1987). In agreement with these findings, the deoxyribonucleoside phosphoramidites S.44 (Polushin, 1996) and S.45 (Jørgensen et al., 1994) have also exhibited significantly lower condensation rates and coupling efficiency because of steric factors (Fig. 3.3.14). Like sterically hindered groups attached to nucleobases (vide supra), it is also possible to decrease the steric demand of 2′-O-bulky protecting groups by increasing the distance between these groups and the phosphoramidite function. Specifically, the ribonucleoside 2′-O-triisopropylsilyloxymethyl phosphoramidite S.46 (Weiss, 1998; Wu and Pitsch, 1998; Fig. 3.3.15) allows much faster coupling reactions (2 min) in solid-phase oligoribonucleotide synthesis than do ribonucleoside 2′-O-tertbutyldimethysilyl phosphoramidites (5–8 min) under essentially identical conditions (see UNIT 3.4). Similarly, coupling reactions of t h e r i b o n u c l e o s i d e 2′-O-(o-nitrobenzyloxymethyl) phosphor- amidite S.47 (Fig. 3.3.15) are faster (2 min) than those effected b y r i b o n u c l e o s i d e 2′-O-(o-nitrobenzyl) phosphoramidites (10 min) under the same conditions (deBear et al., 1987; Schwartz et al., 1992). Thus functional groups generating
B
DMTrO
i-Pr2N
O
P
O
O
O
Si
NCCH2CH2O 46
B
DMTrO
O O
i-Pr2N
P
O2N O
O
OCH2CH2CN 47
Figure 3.3.15 Efficient ribonucleoside phosphoramidites for solid-phase RNA synthesis.
steric bulk near the nucleosidic phosphoramidite moiety are likely to interfere with coupling rates and should be given consideration when developing novel phosphoramidite monomers.
SIGNIFICANCE OF THE “CAPPING” REACTION IN THE CHEMICAL SYNTHESIS OF OLIGODEOXYRIBONUCLEOTIDES The phosphoramidite approach to oligodeoxyribonucleotide synthesis is renowned for its high coupling efficiency. Nonetheless, oligonucleotide chain extension does not occur quantitatively even under optimum conditions. As a result, the desired n-mer oligodeoxyribonucleotide is contaminated in the final product with a population of shorter (n − 1)-oligomers. Separation of the n-mer oligonucleotide from (n − 1)-mers can be challenging; however, this problem is almost completely eliminated by acetylation of the remaining unphosphitylated oligonucleotides after each condensation step. This “capping” reaction terminates the elongation of the unphosphitylated oligomers
NHBz N N
DMTrO
O
O
DMTrO
Ura
Thy
DMTrO TBDMSO
O
O
O
i-Pr2N
O P
40 41 42 43
Oligodeoxyribonucleotide Synthesis Using the Phosphoramidite Method
R OCH2CH2CN R=H R = OMe R = OThp R = OTBDMS
O i-Pr2N P NCCH2CH2O
HN
OMe O
44
i-Pr2N
O P
OCH2CH2CN 45
Thp, tetrahydropyran-2-yl TBDMS, tert-butyldimethylsilyl
Figure 3.3.14 Nucleoside phosphoramidites functionalized with 2′- or 3′-sterically demanding groups.
3.3.8 Current Protocols in Nucleic Acid Chemistry
that would otherwise occur during the next coupling step. A very effective capping reagent is a solution of acetic anhydride, 2,6-lutidine, and N-methylimidazole in tetrahydrofuran (Farrance et al., 1989). Such a capping formulation not only prevents extension of unphosphitylated oligomers but also efficiently reduces the concentration of O6-phosphitylated guanine residues that are generated during the condensation step (Pon et al., 1986; Eadie and Davidson, 1987). The capping formulation is rich in acetate ions; these nucleophiles efficiently cleave O6-phosphitylated guanine adducts by attacking tricoordinated “enol phosphites” (like S.48; Fig. 3.3.16) and releasing unmodified guanine residues. Adducts such as S.48 or its oxidized form S.49, if not destroyed, can serve as secondary sites for oligonucleotide synthesis and lead to the formation of a complex mixture of branched oligodeoxyribonucleotides. The generation of these adducts is most efficiently minimized when the capping reaction is performed before the oxidation step. O,O-Diethyl-N,N-diisopropyl phosphoramidite has also been reported as an improved capping reagent in oligonucleotide synthesis (Yu et al., 1994). On the basis of the data presented, this phosphoramidite exhibited a capping efficiency that was only modestly superior to that of the standard acetic anhydride/N-methylimidazole/2,6-lutidine capping formulation. The phosphoramidite capping reagent was also claimed to not produce nucleobase modification; supporting data were, however, not shown. Interestingly, recent use of the lipophilic O-(2-cyanoethyl),O-octyl-N,N-diisopropyl phosphoramidite as a capping re-
B O O O
O
B
DMTrO
O
O O P
N
O
B
DMTrO
O
O P OCH2CH2CN O
OCH2CH2CN
N
O P OR N
Oxidation of a newly generated phosphite triester linkage to the corresponding phosphate triester function is an essential step in automated synthesis of oligodeoxyribonucleotides by the phosphoramidite approach. Without oxidation, an internucleoside phosphite triester function decomposes under the acidic conditions required for cleavage of the 5′-dimethoxytrityl group (Matteucci and Caruthers, 1981). Thus oxidation of phosphite triesters is absolutely necessary to ensure consistent highyielding oligodeoxyribonucleotide syntheses. An aqueous solution of iodine (0.05–0.1 M)
O
N
O
THE OXIDATION REACTION IN THE SYNTHESIS OF OLIGODEOXYRIBONUCLEOTIDES ACCORDING TO THE PHOSPHORAMIDITE METHOD
B
DMTrO
DMTrO
agent in solid-phase oligonucleotide synthesis has allowed facile separation of capped failure sequences from trityl-off full-length oligonucleotides by reverse-phase HPLC (RP-HPLC; Natt and Häner, 1997). This capping method simplified RP-HPLC purification of synthetic oligonucleotides and resulted in higher isolated yields. Phosphoramidite capping reagents may, however, like nucleoside phosphoramidites, phosphitylate nucleobases (especially at O-6 of guanosines) and eventually lead to the formation of, for example, 2,6-diaminopurine residues (Eadie and Davidson, 1987). This potential problem has not, as yet, been thoroughly investigated. Until this issue is resolved, use of the standard and well-studied acetic anhydride/N-methylimidazole/2,6-lutidine capping formulation during solid-phase oligonucleotide synthesis is recommended.
N
N
O P OR NHi-Bu
O
O
O
O
P
P 48 R = CH2CH2CN
N
N
NHi-Bu
49 R = CH2CH2CN
i-Bu, isobutyryl
6
Figure 3.3.16 Postulated O -guanine adducts generated during the chain extension step of the synthesis cycle according to the phosphoramidite method.
Synthesis of Unmodified Oligonucleotides
3.3.9 Current Protocols in Nucleic Acid Chemistry
and 2,6-lutidine (Letsinger and Lunsford, 1976) or pyridine (Usman et al., 1985) in tetrahydrofuran is generally used for this task; this formulation is stable and provides rapid-reaction kinetics, usually without formation of side products. When the N-2 amino function of guanine is protected with a N,N-dimethylformamidine group (Zemlicka and Holy, 1967; McBride et al., 1986; Vu et al., 1990) during automated oligonucleotide synthesis, however, use of the traditional iodine formulation as oxidant led to cyanation of guanine at N-2 (Mullah et al., 1995). It has also been shown that this side reaction is completely eliminated by the use of a lower-concentration (0.02 M) iodine oxidation reagent without losing speed and efficiency in the conversion of internucleoside phosphite triesters to phosphate triesters (Mullah et al., 1995). Thus the latter aqueous iodine formulation is recommended for standard oligonucleotide synthesis. For specific applications, however, nonaqueous oxidizing reagents may advantageously offer an alternative to aqueous iodine for the oxidation of oligodeoxyribonucleoside phosphite triesters. For example, m-chloroperbenzoic acid (Tanaka and Letsinger, 1982); iodobenzene diacetate and tetra-n-butylammonium periodate (Fourrey and Varenne, 1985); tert-butyl hydroperoxide (Hayakawa et al., 1986; Hayakawa and Kataoka, 1998); ditert-butyl hydroperoxide; cumene hydroperoxide; hydrogen peroxide; bis-trimethylsilyl peroxide, and catalytic amounts of trimethylsilyl triflate (Hayakawa et al., 1986); dinitrogen tetroxide and molecular oxygen in the presence of 2,2′-azobis(2-methylpropionitrile) under thermal or photochemical conditions (Bentrude et al., 1989); and (1S)-(+)-(10-camphorsulfonyl) oxaziridine (S.50; Ugi et al., 1988) have been effective. The oxaziridine S.50 (Fig. 3.3.17) is particularly useful for the synthesis of oligonucleotide containing multiple 7deaza-2′-deoxyguanosine residues. Incorpora-
Thy
DMTrO
N S O O O 50
Oligodeoxyribonucleotide Synthesis Using the Phosphoramidite Method
i-Pr2N
O O P
CH3 O 51
Figure 3.3.17 An oxaziridine derivative as a useful oxidant in the synthesis of oligonucleotides containing iodine sensitive residues, and a benzylic deoxyribonucleoside phosphoramidite suitable for the preparation of oligonucleotide analogues.
tion of this modified 2′-deoxyguanosine into oligonucleotides via the phosphoramidite approach is sensitive to iodine-containing solutions regardless of iodine concentration (Anonymous, 1996). In this context, it should be noted that when applied to oligonucleotide synthesis, the benzylic deoxyribonucleoside phosphoramidite S.51 (Fig. 3.3.17) generated internucleoside o-methylbenzyl phosphite esters that were sensitive to aqueous iodine oxidation. This sensitivity to iodine resulted in the loss of benzylic phosphate protection (Caruthers et al., 1987b). The absence of phosphate protecting groups did not, however, impair subsequent additions of S.51 to the DNA chain. In fact, an oligothymidylic acid (20-mer) was prepared by the iterative incorporation of S.51 with an average coupling efficiency of 96%. It was speculated that phosphate-phosphite mixed anhydrides could have been generated from the interaction of phosphate diesters with activated S.51 and then cleaved by excess 1H-tetrazole to regenerate the deoxyribonucleoside phosphorotetrazolide intermediates needed for chain extension. Because of the inherent hazards involved with handling peroxides, the use of oxaziridine S.50 is, therefore, recommended for the oxidation of phosphite triesters of those modified oligonucleotides that are reactive to iodine and/or necessitate rigorously anhydrous conditions. Moreover, the use of S.50 in oligonucleotide synthesis does not lead to detectable nucleobase modifications (Anonymous, 1996). It has also been shown that oxidation of the dinucleoside 2-cyano-1,1-dimethylethyl phosphite triester S.52 (Fig. 3.3.18) with iodine in the presence of water, alcohols, and amines produced the corresponding dinucleoside phosphate S.54, phosphate triester S.55, and phosphoramidate S.56, respectively (Nielsen and Caruthers, 1988). It is postulated that under these oxidative Arbuzov-type conditions, elimination of the 2-cyano-1,1-dimethylethyl group led to the dinucleoside phosphoryl iodide intermediate S.53. The formation of S.53 is supported by 31P-NMR data and thus provides a versatile pathway to the synthesis of oligodeoxyribonucleotide analogues from deoxynucleoside 3′-O-(2-cyano-1,1-dimethylethyl) or o-methylbenzyl phosphoramidites. Another example of the importance of P(III) oxidation in oligodeoxyribonucleotide synthesis according to the phosphoramidite approach is the incorporation of internucleotide phosphorothioates linkages into these bio-
3.3.10 Current Protocols in Nucleic Acid Chemistry
P
O
I2
O P O
O
Thy
THF
H2O I
O
AcO
O
Thy
or ROH
O O
O P OR O
O
Thy
AcO
AcO
52
Thy
DMTrO
O
O O
NCCH2C(CH3)2O
Thy
DMTrO
Thy
DMTrO
53
54 R = H 55 R = CH3
R'NH2
Thy
DMTrO
O O
O P NHR' O
O
Thy
AcO Ac, acetyl
56 R' = n-butyl
Figure 3.3.18 Access to oligodeoxyribonucleotide analogues from deoxynucleoside (2-cyano1,1-dimethylethyl) phosphoramidites.
molecules. Oligonucleotides carrying internucleotide phosphorothioate diesters display enhanced resistance to hydrolysis catalyzed by nucleases (Eckstein, 1985). Because of this property, oligodeoxyribonucleoside phosphorothioates have been extensively used as antisense molecules in the inhibition of gene expression. Automated synthesis of these modified oligonucleotides via the phosphoramidite method consists of replacing the aqueous iodine oxidation step by a sulfurization reaction that had originally been effected by elemental sulfur. Given the poor solubility of elemental sulfur in organic solvents, its use in automated systems has been difficult. This problem was eliminated when phenylacetyl disulfide (S.57; Kamer et al. 1989; Roelen et al., 1991) and 3H-1,2-benzodithiol-3-one 1,1-dioxide (S.58; Iyer et al., 1990; Regan et al., 1992) were employed as sulfurization reagents (Fig. 3.3.19). These compounds are soluble in organic solvents and produce efficient and rapid sulfurization kinetics. For example, S.58 converted the dinucleoside phosphite triester S.59 to the corresponding phosphorothioate dimer S.60 in yields exceeding 99% within 30 sec at 25°C (Iyer et al., 1990; Regan et al., 1992). Deprotection of S.60 afforded the dinucleoside phosphorothioate S.61 (Fig. 3.3.19). Thus the sulfur-transfer reagent S.58 has enabled reliable automated synthesis of phosphorothioated oligomers carrying either exclusively or a predetermined number of phos-
phorothioate groups (Iyer et al., 1990). Given the biological significance of oligonucleoside phosphorothioates, application of S.58 to the synthesis of these modified oligonucleotides has spurred interest in the development of additional sulfurizing reagents. The most notable sulfur-transfer agents that have been reported during this decade include N,N,N′,N′tetraethylthiuram disulfide (Vu and Hirschbein, 1991), dibenzoyl tetrasulfide (Rao et al., 1992), bis-(O,O-diisopropoxyphosphinothioyl) disulfide (Stec et al., 1993), benzyltriethylammonium tetrathiomolybdate (Rao and Macfarlane, 1994), bis(p-toluenesulfonyl)disulfide (Efimov et al., 1995), 3-ethoxy1,2,4-dithiazoline-5-one (Xu et al., 1996), thiiranes (Arterburn and Perry, 1997), bis(ethoxythiocarbonyl)tetrasulfide (Zhang et al., 1998), and 3-methyl-1,2,4-dithiazoline-5one (Zhang et al., 1999). Out of these sulfurtransfer reagents, 3H-1,2-benzodithiol-3-one 1,1-dioxide and 3-ethoxy-1,2,4-dithiazoline-5one are currently the most extensively used in solid-phase synthesis of oligonucleoside phosphorothioates.
STRATEGIES IN THE DEPROTECTION OF SYNTHETIC OLIGODEOXYRIBONUCLEOTIDES The efficiency of the phosphoramidite method for solid-phase synthesis of oligodeoxyribonucleotides is such that oligonucleotides up to 50 bases long can be synthesized
Synthesis of Unmodified Oligonucleotides
3.3.11 Current Protocols in Nucleic Acid Chemistry
O
O S
S
S S
O
O
57
B
DMTrO O
S O
O
O
58
O
O O
P − + S
O S
P OCH2CH2CN
O
B
DMTrO
O O
S
O
58
B
O
OCH2CH2CN
O
O
O
O
P
P
B
59
B
HO
O
−
deprotection
O P S O
B
DMTrO
O
O O
S P OCH2CH2CN O
B
OH
O
O
B
O
61 P 60
Figure 3.3.19 Preparation of oligodeoxyribonucleoside phosphorothioates according to the solidphase phosphoramidite method.
Oligodeoxyribonucleotide Synthesis Using the Phosphoramidite Method
within a few hours. While the cleavage of these oligonucleotides from solid supports is normally accomplished by treatment with concentrated ammonium hydroxide for ∼1 hr at ambient temperature, it will take ∼10 hr at elevated temperature (55°C) to deprotect the N-isobutyryl group of guanines, and N-benzoyl group of cytosines and adenines (see UNIT 2.1). This time-consuming deprotection step clashed with the urgent demand for synthetic oligodeoxyribonucleotides and thus provided an incentive to improve the chemistry involved with postsynthesis oligonucleotide processing. Specifically, methods for rapid removal of oligonucleotide protecting groups have attracted considerable attention and motivated the development of novel base-labile blocking groups for nucleobases (Schulhof et al., 1987; Uznanski et al., 1989; Kuijpers et al., 1990; Vu et al., 1990; Beaucage and Iyer, 1992; Sinha et al., 1993; Iyer et al., 1997; see also UNIT 2.1). Concentrated solutions of ammonia in water, ethanol, or methanol have been used for the cleavage of these groups. Alternatively, an aqueous
solution of methylamine and ammonium hydroxide has been employed for the deprotection of oligonucleotides carrying N-acetyl cytosines, N-benzoyl adenines, and N-isobutyryl guanines (Reddy et al., 1994). With this reagent, oligonucleotides were cleaved from solid supports in 5 min at ambient temperature, and complete deprotection was accomplished in 5 min at 65°C. It should, however, be emphasized that an aqueous solution of methylamine and ammonium hydroxide cannot be used for the deprotection of oligonucleotides bearing conventional N-benzoyl cytosines because primary amines have been reported to attack N4-anisoyl- or N4-benzoyl-2′-deoxycytidine at C-4 to produce N4-alkylated 2′-deoxycytidine derivatives (Weber and Khorana, 1972; Reddy et al., 1997). Gaseous amines such as ammonia or methylamine have also been employed under pressure to achieve mild and rapid deprotection conditions (Boal et al., 1996). For example, oligodeoxyribonucleotides having cytosines, adenines, and guanines N-protected with a tert-
3.3.12 Current Protocols in Nucleic Acid Chemistry
butylphenoxyacetyl group were released from CPG supports and fully deprotected at 25°C by pressurized ammonia or methylamine within 35 or 2 min, respectively. It has also been shown that when the N-benzoyl group is used for protection of cytosines and adenosines, and N-isobutyryl for guanines, complete deprotection of oligodeoxyribonucleotides by ammonia gas will take ∼7 hr at 25°C. At that temperature, it would take ∼36 hr for concentrated aqueous ammonium hydroxide to accomplish the same task (Boal et al., 1996). The use of ammonia or methylamine gas allows the simultaneous deprotection of a large number of oligodeoxyribonucleotides. In fact, the number of oligonucleotides or CPG columns that can be deprotected is limited only by the size of the pressure vessel employed. Because no water is present during deprotection, fully deblocked oligonucleotides remain adsorbed to CPG and thus prevent cross-contamination between columns. Oligonucleotides can then be eluted from individual columns with a minimum amount of water for further purification, if desired. This deprotection procedure eliminates hazards inherent to the handling and heating of aqueous amine solutions in glass vials and, more important, the time-consuming evaporation of these solutions. The gas-phase deprotection methodology is recommended when oligonucleotides carrying base-sensitive nucleobases demand mild deprotection conditions or when rapid deprotection is needed to accelerate the production of synthetic oligonucleotides.
ALTERNATE STRATEGIES TO THE SYNTHESIS OF OLIGODEOXYRIBONUCLEOTIDES ACCORDING TO THE PHOSPHORAMIDITE METHOD The versatility of the phosphoramidite approach to oligodeoxyribonucleotide synthesis has been further demonstrated by the use of deoxyribonucleoside phosphoramidites with unprotected nucleobases. The success of this strategy depends on a modified synthesis cycle protocol that involves treatment of the solid support with an equimolar solution (0.1 M) of pyridine hydrochloride and aniline in acetonitrile (Gryaznov and Letsinger, 1991) or benzimidazolium triflate in methanol (Hayakawa and Kataoka, 1998) immediately after each condensation reaction. This treatment destroys nucleobase adducts that are forming on the oligonucleotidic chain during each coupling step. This procedure should facilitate the syn-
thesis of oligonucleotides bearing base-sensitive functional groups because treatment with concentrated ammonium hydroxide at elevated temperature will no longer be required for oligonucleotide deprotection. Furthermore, depurination of adenine and guanine residues under the acidic conditions required for the removal of the 5′-O-DMTr group will become even less likely. More data are still needed to assess whether the synthesis of oligodeoxyribonucleotides according to the phosphoramidite method without nucleobase protection is trouble-free. The method is promising in that it may significantly expedite the production of synthetic oligonucleotides by shortening postsynthesis oligonucleotide processing time. Another strategy toward the preparation of oligodeoxyribonucleotides entails the stepwise condensation of dinucleotide phosphoramidite blocks such as S.62-S.65 (Fig. 3.3.20) instead of conventional monomeric deoxyribonucleoside phosphoramidites for chain extension. Activation of S.62 with 1H-tetrazole produced coupling yields (∼99%) similar to those generated by monomeric phosphoramidites (Kumar and Poonian, 1984). The incorporation of S.63 into oligonucleotides allowed syntheses of randomized DNA sequences containing the 20 codons corresponding to all natural amino acids (Neuner et al., 1998). The efficiency of dinucleotide phosphoramidites to solid-phase oligonucleotide synthesis has been further demonstrated by the preparation of a large oligomer (101-mer) through repetitive condensations of the dimeric phosphoramidite S.64 (Wolter et al., 1986). Furthermore, the impurity profile of oligonucleoside phosphorothioates synthesized by iterative coupling of the thioated dinucleotide phosphoramidite S.65 (Krotz et al., 1997b) showed at least 70% reduction of the (n − 1)-mers and a ∼50% reduction of phosphodiester formation when compared to profiles obtained by standard monomer phosphoramidite couplings. The use of dimeric phosphoramidites in the synthesis of unmodified oligodeoxyribonucleotides has not been widely adopted, probably because a library of up to 16 combinatorial dimers had to be prepared to accomplish the synthesis of one oligonucleotide. Conversely, the application of dimeric phosphoramidites to oligonucleotide analogue synthesis has been popular especially for the incorporation of modified internucleotide bridges. For example, the dimeric 5′-phosphonate–linked thymidine phosphoramidite S.66 (Zhao and Caruthers, 1996) and S.67 (Kofoed and Caruthers, 1996)
Synthesis of Unmodified Oligonucleotides
3.3.13 Current Protocols in Nucleic Acid Chemistry
B
DMTrO
Thy
DMTrO
O
O O
O
X P OCH2CH2CN
O P OMe O
O
R2N
O P
B
O
i-Pr2N
OR'
Thy
O O P
OCH2CH2CN
64 X = O 65 X = S
62 R = R' = CH3 63 R = CH(CH3)2, R' = CH2CH2CN
Figure 3.3.20 Solid-phase oligonucleotide synthesis using dinucleotide phosphoramidite derivatives.
have been prepared and incorporated into oligodeoxyribonucleotides to assess the physicochemical and biochemical properties imparted by such modifications (Fig. 3.3.21). For similar purposes, and given the growing interest in the development of therapeutic oligonucleotides, a plethoric number of dimeric phosphoramidites structurally related to S.66-S.68 have been prepared in recent years. Because of the intense activity in this area of research, only a fraction of the work has, so far, been reviewed (see Beaucage and Iyer, 1993; Sanghvi and Cook, 1994; Agrawal and Iyer, 1995; Iyer et al., 1999). Oligodeoxyribonucleotides have also been prepared by the condensation of trinucleotide phosphoramidite blocks to enable oligonucleotide-directed mutagenesis. More and more, oligonucleotides of mixed composition are being used to generate combinatorial libraries of variants in the search for peptides and proteins with improved properties. The most direct route to controlled mutagenesis is indeed the use of trinucleotide synthons that correspond to the amino acid codons needed. The synthesis of trinucleotide phosphoramidites S.69 (Sondek
and Shortle, 1992; Virnekäs et al., 1994), S.70 (Lyttle et al., 1995), S.71 (Ono et al., 1995; Kayushin et al., 1996; Zehl et al., 1996), and S.72 (Gaytán et al., 1998; Fig. 3.3.22) representing the codons for all 20 amino acids has been achieved. The incorporation of S.69 into oligonucleotides was accomplished by allowing a coupling time of 1 min and performing the trinucleotide condensation step twice. Under these conditions, coupling yields averaged 96%–98.5% (Virnekäs et al., 1994). Considering that each trinucleotide condensation adds three nucleobases to the growing oligonucleotide chain, these coupling yields are equivalent to three individual monomeric phosphoramidite condensations, each with a coupling efficiency of 98%–99.5%. Incorporation of the trinucleotide phosphoramidites S.71 and S.72 into oligonucleotides via automated solidphase synthesis occurred in yields that varied with the sequence of the trinucleotide block used. Nonetheless, the incorporation of these trinucleotide phosphoramidite blocks into synthetic DNA in the controlled, codon-by-codon construction of combinatorial libraries of struc-
B
DMTrO DMTrO
Thy
O
O
1 O
O
2 O
O P OPhCl-2
O O P
O
Thy O
BnO O
i-Pr2N
O
Thy
DMTrO
P
OCH2CH2CN
i-Pr2N
O P
Thy
3 O 4 O
OCH2CH2CN
i-Pr2N 66
Oligodeoxyribonucleotide Synthesis Using the Phosphoramidite Method
67
O P
O
B
OCH2CH2CN
68
Bn, benzyl PhCl-2, o-chlorophenyl
n , atom or group of atoms O
Figure 3.3.21 Solid-phase synthesis of oligonucleotide analogues from dimeric phosphoramidites carrying modified internucleotidic linkages.
3.3.14 Current Protocols in Nucleic Acid Chemistry
tural genes will be invaluable in creating molecular diversity by mutagenesis.
CONCLUDING REMARKS Owing to the high performance of the phosphoramidite method, synthetic oligodeoxyribonucleotides became readily available and fueled the biotechnology revolution that has irreversibly changed biomedical research and the pharmaceutical industry. For example, without the ability to rapidly and efficiently synthesize DNA oligonucleotides, the development of the polymerase chain reaction (PCR) and its multiple applications would have been difficult, if not impossible, because this technology completely depends on the use of DNA primers. Similarly, the phosphoramidite method has been instrumental in the development of automated DNA sequencing, which also requires rapid and efficient synthesis of fluorescent DNA primers. Another important biological application for oligodeoxyribonucleotides generated by the phosphoramidite method relates to site-specific mutagenesis of protein genes. Mutagenesis of this type has been used to study protein structure-function relationships and to alter the therapeutic spectrum of pharmaceutically active proteins. In addition, the phosphoramidite method has been particularly useful in the synthesis of modified oligonucleotides for diagnostic applications and as potential therapeutic drugs. Although the latter research area is relatively new, several oligonucleotide-based drugs have already reached the clinic, and others are under preclinical investigation to benefit public health and push further the frontiers of knowledge.
B
DMTrO
O
Adams, S.P., Kavka, K.S., Wykes, E.J., Holder, S.B., and Galluppi, G.R. 1983. Hindered dialkylamino nucleoside phosphite reagents in the synthesis of two DNA 51-mers. J. Am. Chem. Soc. 105:661663. Agrawal, S. and Iyer, R.P. 1995. Modified oligonucleotides as therapeutic and diagnostic agents. Curr. Opin. Biotechnol. 6:12-19. Andrus, A. and Beaucage, S.L. 1988. 2-Mercaptobenzothiazole—An improved reagent for the removal of methyl phosphate protecting groups from oligodeoxynucleotide phosphotriesters. Tetrahedron Lett. 29:5479-5482. Anonymous. 1996. Non-aqueous oxidation with 10camphorsulfonyl-oxaziridine. The Glen Report 9:8-9. Arnold, L., Tocik, Z., Bradkova, E., Hostomsky, Z., Paces, V., and Smrt, J. 1989. Automated chloridite and amidite synthesis of oligodeoxyribonucleotides on a long chain support using amidine protected purine nucleosides. Collect. Czech. Chem. Commun. 54:523-532. Arterburn, J.B. and Perry, M.C. 1997. Rhenium catalyzed sulfurization of phosphorus(III) compounds with thiiranes: New reagents for phosphorothioate ester synthesis. Tetrahedron Lett. 38:7701-7704. Barone, A.D., Tang, J.-Y., and Caruthers, M.H. 1984. In situ activation of bis-dialkylaminophosphines—A new method for synthesizing deoxyoligonucleotides on polymer supports. Nucl. Acids Res. 12:4051-4061. Beaucage, S.L. 1984. A simple and efficient preparation of deoxynucleoside phosphoramidites in situ. Tetrahedron Lett. 25:375-378. Beaucage, S.L. and Caruthers, M.H. 1981. Deoxynucleoside phosphoramidites—A new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Lett. 22:1859-1862. Beaucage, S.L. and Iyer, R.P. 1992. Advances in the synthesis of oligonucleotides by the phosphoramidite approach. Tetrahedron 48:22232311.
B
R'O
O
LITERATURE CITED
O O
O P OR B
O
Beaucage, S.L. and Iyer, R.P. 1993. The synthesis of modified oligonucleotides by the phosphoramidite approach and their applications. Tetrahedron 49:6123-6194.
O P OPhCl-2
O
O O
O P OR O
i-Pr2N
B
O
O
O P OPhCl-2 O
O P
B
OR
69 R = CH3 70 R = CH2CH2CN
O
i-Pr2N
O O P
B
Beier, M. and Pfleiderer, W. 1999. Pyridinium salts–An effective class of catalysts for oligonucleotide synthesis. Helv. Chim. Acta 82:879887.
OCH2CH2CN
71 R' = DMTr 72 R' = Fmoc
Fmoc, 9-fluorenylmethoxycarbonyl PhCl-2, o-chlorophenyl
Figure 3.3.22 Trinucleotide phosphoramidite blocks for the controlled, codon-by-codon, construction of combinatorial gene libraries.
Bentrude, W.G., Sopchik, A.E., and Gajda, T. 1989. Stereo- and regiochemistries of the oxidations of 2-methoxy-5-tert-butyl-1,3,2-dioxaphosphori nanes and the cyclic methyl 3′,5′-phosphite of thymidine by H2O/I2 and O2/AIBN to P-chiral phosphates. 17O NMR assignment of phosphorus configuration to the diastereomeric thymidine cyclic methyl 3′,5′-monophosphates. J. Am. Chem. Soc. 111:3981-3987.
Synthesis of Unmodified Oligonucleotides
3.3.15 Current Protocols in Nucleic Acid Chemistry
Bergstrom, D.E. and Gerry, N. 1994. Precision sequence-specific cleavage of a nucleic acid by a minor-groove-directed metal-binding ligand linked through N-2 of deoxyguanosine. J. Am. Chem. Soc. 116:12067-12068. Berner, S., Mühlegger, K., and Seliger, H. 1989. Studies on the role of tetrazole in the activation of phosphoramidites. Nucl. Acids Res. 17:853864. Boal, J.H., Wilk, A., Harindranath, N., Max, E.E., Kempe, T., and Beaucage, S.L. 1996. Cleavage of oligodeoxyribonucleotides from controlledpore glass supports and their rapid deprotection by gaseous amines. Nucl. Acids Res. 24:31153117. Boudjebel, H., Gonçalves, H., and Mathis, F. 1975. Étude de la liaison P—N dans le motif S2P— NMe3 en résonance magnétique nucléaire et par la réaction d’échange avec le trifluoroacétate de méthyle. Bull. Chem. Soc. Chim. France 628634. Brill, W.K.-D., Nielsen, J., and Caruthers, M.H. 1991. Synthesis of deoxydinucleoside phosphorodithioates. J. Am. Chem. Soc. 113:39723980. Caruthers, M.H., Beaucage, S.L., Becker, C., Efcavitch, W., Fisher, E.F., Galluppi, G., Goldman, R., deHaseth, P., Martin, F., Matteucci, M., and Stabinsky, Y. 1982. New methods for synthesizing deoxyoligonucleotides. In Genetic Engineering: Principles and Methods, Vol. 4 (J.K. Setlow and A. Hollaender, eds.) pp. 1-17. Plenum, New York. Caruthers, M.H., Barone, A.D., Beaucage, S.L., Dodds, D.R., Fisher, E.F., McBride, L.J., Matteucci, M., Stabinsky, Z., and Tang, Y.-Y. 1987a. Chemical synthesis of deoxyoligonucleotides by the phosphoramidite method. In Methods and Enzymology; Vol. 154 (R. Wu and L. Grossman, eds.) pp. 287-313. Academic Press, San Diego; and references therein. Caruthers, M.H., Kierzek, R., and Tang, J.Y. 1987b. Synthesis of oligonucleotides using the phosphoramidite method. In Biophosphates and Their Analogues—Synthesis, Structure, Metabolism and Activity (K.S. Bruzik and W.J. Stec, eds.) pp. 3-21. Elsevier/North Holland, Amsterdam. Casale, R. and McLaughlin, L.W. 1990. Synthesis and properties of an oligodeoxynucleotide containing a polycyclic aromatic hydrocarbon site specifically bound to the N2 amino group of a 2′-deoxyguanosine residue. J. Am. Chem. Soc. 112:5264-5271.
Oligodeoxyribonucleotide Synthesis Using the Phosphoramidite Method
Dahl, B.H., Nielsen, J., and Dahl, O. 1987. Mechanistic studies on the phosphoramidite coupling reaction in oligonucleotide synthesis. I. Evidence for nucleophilic catalysis by tetrazole and rate variations with the phosphorus substituents. Nucl. Acids Res. 15:1729-1743. Dahl, B.H., Bjergårde, K., Henriksen, L., and Dahl, O. 1990. A highly reactive, odourless substitute for thiophenol/triethylamine as a deprotection reagent in the synthesis of oligonucleotides and their analogues. Acta Chem. Scand. 44:639-641. Daub, G.W. and van Tamelen, E.E. 1977. Synthesis of oligoribonucleotides based on the facile cleavage of methyl phosphotriester intermediates. J. Am. Chem. Soc. 99:3526-3528. deBear, J.S., Hayes, J.A., Koleck, M.P., and Gough, G.R. 1987. A universal glass support for oligonucleotide synthesis. Nucleosides Nucleotides 6:821-830. Eadie, J.S. and Davidson, D.S. 1987. Guanine modification during chemical DNA synthesis. Nucl. Acids Res. 15:8333-8349. Eckstein, F. 1985. Nucleoside phosphorothioates. Annu. Rev. Biochem. 54:367-402. Efimov, V.A., Kalinkina, A.L., Chakhmakhcheva, O.G., Schmaltz Hill, T. and Jayaraman, K. 1995. New efficient sulfurizing reagents for the preparation of oligodeoxyribonucleotide phosphorothioate analogues. Nucl. Acids Res. 23:4029-4033. Farrance, I.K., Eadie, J.S., and Ivarie, R. 1989. Improved chemistry for oligodeoxyribonucleotide synthesis substantially improves restriction enzyme cleavage of a synthetic 35 mer. Nucl. Acids Res. 17:1231-1245. Fourrey, J.-L. and Varenne, J. 1984. Improved procedure for the preparation of deoxynucleoside phosphoramidites: Arylphosphoramidites as new convenient intermediates for oligodeoxynucleotide synthesis. Tetrahedron Lett. 25:45114514. Fourrey, J.-L. and Varenne, J. 1985. Introduction of a nonaqueous oxidation procedure in the phosphite triester route for oligonucleotide synthesis. Tetrahedron Lett. 26:1217-1220. Fourrey, J.-L., Varenne, J., Fontaine, C., Guittet, E., and Yang, Z.W. 1987. A new method for the synthesis of branched ribonucleotides. Tetrahedron Lett. 28:1769-1772. Froehler, B. and Matteucci, M.D. 1983. Substituted 5-phenyltetrazoles: Improved activators of deoxynucleoside phosphoramidites in deoxyoligonucleotide synthesis. Tetrahedron Lett. 24:3171-3174.
Claesen, C., Tesser, G.I., Dreef, C.E., Marugg, J.E., van der Marel, G.A., and van Boom, J.H. 1984. Use of 2-methylsulfonylethyl as a phosphorus protecting group in oligonucleotide synthesis via a phosphite triester approach. Tetrahedron Lett. 25:1307-1310.
Gaytán, P., Yañez, J., Sánchez, F., Mackie, H., and Soberón, X. 1998. Combination of DMTmononucleotide and Fmoc-trinucleotide phosphoramidites in oligonucleotide synthesis affords an automatable codon-level mutagenesis method. Chem. Biol. 5:519-527.
Crooke, S.T. and Bennett, C.F. 1996. Progress in antisense oligonucleotide therapeutics. Annu. Rev. Pharmacol. Toxicol. 36:107-129.
Gryaznov, S.M. and Letsinger, R.L. 1991. Synthesis of oligonucleotides via monomers with unprotected bases. J. Am. Chem. Soc. 113:5876-5877.
3.3.16 Current Protocols in Nucleic Acid Chemistry
Gryaznov, S.M. and Letsinger, R.L. 1992. Selective O-phosphitilation with nucleoside phosphoramidite reagents. Nucl. Acids Res. 20:18791882. Guo, M., Yu, D., Iyer, R.P., and Agrawal, S. 1998. Solid-phase stereoselective synthesis of 2′-Omethyl oligoribonucleoside phosphorothioates using nucleoside oxazaphospholidines. Bioorg. Med. Chem. Lett. 8:2539-2544. Hayakawa, Y. and Kataoka, M. 1998. Facile synthesis of oligodeoxyribonucleotides via the phosphoramidite method without nucleoside base protection. J. Am. Chem. Soc. 120:12395-12401. Hayakawa, Y., Uchiyama, M., and Noyori, R. 1986. Nonaqueous oxidation of nucleoside phosphites to the phosphates. Tetrahedron Lett. 27:41914194. Hayakawa, Y., Kataoka, M., and Noyori, R. 1996. Benzimidazolium triflate as an efficient promoter for nucleotide synthesis via the phosphoramidite method. J. Org. Chem. 61:79967997. Hering, G., Stöcklein-Schneiderwind, R., Ugi, I., Pathak, T., Balgobin, N., and Chattopadhyaya, J. 1985. Preparation and properties of chloroN,N-dialkylamino-2,2,2-trichloroethoxy- and chloro-N,N-dialkylamino-2,2,2-trichloro-1,1dimethylethoxyphosphines and their deoxynucleoside phosphiteamidates. Nucleosides Nucleotides 4:169-171. Iyer, R.P., Phillips, L.R., Egan, W., Regan, J.B., and Beaucage, S.L. 1990. The automated synthesis of sulfur-containing oligodeoxyribonucleotides using 3H-1,2-benzodithiol-3-one 1,1-dioxide as a sulfur-transfer reagent. J. Org. Chem. 55:46934698. Iyer, R.P., Yu, D., Habus, I., Ho, N.-H., Johnson, S., Devlin, T., Jiang, Z., Zhou, W., Xie, J., and Agrawal, S. 1997. N-Pent-4-enoyl (PNT) group as a universal nucleobase protector: Applications in the rapid and facile synthesis of oligonucleotides, analogs, and conjugates. Tetrahedron 53:2731-2750. Iyer, R.P., Guo, M.-J., Yu, D., and Agrawal, S. 1998. Solid-phase stereoselective synthesis of oligonucleoside phosphorothioates: The nucleoside bicyclic oxazaphospholidines as novel synthons. Tetrahedron Lett. 39:2491-2494. Iyer, R.P., Roland, A., Zhou, W., and Ghosh, K. 1999. Modified oligonucleotides—Synthesis, properties and applications. Curr. Opin. Mol. Ther. 1:344-358. Jørgensen, P.N., Stein, P.C., and Wengel, J. 1994. Synthesis of 3′-C-(hydroxymethyl) thymidine: Introduction of a novel class of dexoynucleosides and oligodeoxynucleotides. J. Am. Chem. Soc. 116:2231-2232. Josephson, S., Lagerholm, E., and Palm, G. 1984. Automatic synthesis of oligodeoxynucleotides and mixed oligodeoxynucleotides using the phosphoramidite method. Acta Chem. Scand. B38:539-545.
Kamer, P.C.J., Roelen, H.C.P.F., van den Elst, H., van der Marel, G.A., and van Boom, J.H. 1989. An efficient approach toward the synthesis of phosphorothioate diesters via the Schönberg reaction. Tetrahedron Lett. 30:6757-6760. Kayushin, A.L., Korosteleva, M.D., Miroshnikov, A.I., Kosch, W., Zubov, D., and Piel, N. 1996. A convenient approach to the synthesis of trinucleotide phosphoramidites—Synthons for the generation of oligonucleotide/peptide libraries. Nucl. Acids Res. 24:3748-3755. Khorana, H.G. 1968. Nucleic acid synthesis. Pure Appl. Chem. 17:349-381. Kierzek, R., Rozek, M., and Markiewicz, W.T. 1987. Some steric aspects of synthesis of oligoribonucleotides by phosphoramidite approach on solid support. Nucl. Acids Res. Symp. Ser. No. 18:201204. Kofoed, T. and Caruthers, M.H. 1996. Synthesis of 5′-phosphonate linked thymidine deoxyoligonucleotides. Tetrahedron Lett. 37:6457-6460. Krotz, A.H., Klopchin, P.G., Walker, K.L., Srivatsa, G.S., Cole, D.L., and Ravikumar, V.T. 1997a. On the formation of longmers in phosphorothioate oligodeoxyribonucleotide synthesis. Tetrahedron Lett. 38:3875-3878. Krotz, A.H., Klopchin, P., Cole, D.L., and Ravikumar, V.T. 1997b. Improved purity profile of phosphorothioate oligonucleotides through the use of dimeric phosphoramidite synthons. Nucleosides Nucleotides 16:1637-1640. Kuijpers, W.H.A., Huskens, J., and van Boeckel, C.A.A. 1990. The 2-(acetoxymethyl)benzoyl (AMB) group as a new base-protecting group, designed for the protection of (phosphate) modified oligonucleotides. Tetrahedron Lett. 31:6729-6732. Kumar, G. and Poonian, M.S. 1984. Improvements in oligodeoxyribonucleotide synthesis: Methyl N,N-dialkylphosphoramidite dimer units for solid support phosphite methodology. J. Org. Chem. 49:4905-4912. Lee, H.-J. and Moon, S.-H. 1984. Bis-(N,N-dialkylamino)-alkoxyphosphines as a new class of phosphite coupling agent for the synthesis of oligonucleotides. Chem. Lett. 1229-1232. Letsinger, R.L. and Lunsford, W.B. 1976. Synthesis of thymidine oligonucleotides by phosphite triester intermediates. J. Am. Chem. Soc. 98:36553661. Letsinger, R.L. and Mahadevan, V. 1966. Stepwise synthesis of oligodeoxyribonucleotides on an insoluble polymer support. J. Am. Chem. Soc. 88:5319-5324. Letsinger, R.L. and Ogilvie, K.K. 1969. Synthesis of oligothymidylates via phosphotriester intermediates. J. Am. Chem. Soc. 91:3350-3355. Lyttle, M.H., Napolitano, E.W., Calio, B.L., and Kauvar, L.M. 1995. Mutagenesis using trinucleotide β-cyanoethyl phosphoramidites. BioTechniques 19:274-280.
Synthesis of Unmodified Oligonucleotides
3.3.17 Current Protocols in Nucleic Acid Chemistry
Mathis, R., Lafaille, L., and Burgada, R. 1974. Fréquence d’absorption de la liaison P-N dans des composés du phosphore tricoordonné. Spectrochim. Acta, Part A 30:357-370. Matteucci, M.D. and Caruthers, M.H. 1981. Synthesis of deoxyoligonucleotides on a polymer support. J. Am. Chem. Soc. 103:3185-3191. McBride, L.J. and Caruthers, M.H. 1983. An investigation of several deoxynucleoside phosphoramidites useful for synthesizing deoxyoligonucleotides. Tetrahedron Lett. 24:245-248. McBride, L.J., Kierzek, R., Beaucage, S.L., and Caruthers, M.H. 1986. Amidine protecting groups for oligonucleotide synthesis. J. Am. Chem. Soc. 108:2040-2048. Montserrat, F.X., Cerandas, A., Eritja, R., and Pedroso, E. 1994. Criteria for the economic large scale solid-phase synthesis of oligonucleotides. Tetrahedron 50:2617-2622. Moore, M.F. and Beaucage, S.L. 1985. Conceptual basis of the selective activation of bis(dialkylamino)methoxyphosphines by weak acids and its application toward the preparation of deoxynucleoside phosphoramidites in situ. J. Org. Chem. 50:2019-2025. Mullah, B., Andrus, A., Zhao, H., and Jones, R.A. 1995. Oxidative conversion of N-dimethylformamidine nucleosides to N-cyano nucleosides. Tetrahedron Lett. 36:4373-4376. Natt, F. and Häner, R. 1997. Lipocap: A lipophilic phosphoramidite-based capping reagent. Tetrahedron 53:9629-9636. Neuner, P., Cortese, R., and Monaci, P. 1998. Codon-based mutagenesis using dimer-phosphoramidites. Nucl. Acids Res. 26:1223-1227. Nielsen, J. and Caruthers, M.H. 1988. Directed Arbuzov-type reactions of 2-cyano-1,1-dimethylethyl deoxynucleoside phosphites. J. Am. Chem. Soc. 110:6275-6276. Ono, A., Matsuda, A., Zhao, J., and Santi, D.V. 1995. The synthesis of blocked triplet-phosphoramidites and their use in mutagenesis. Nucl. Acids Res. 23:4677-4682. Polushin, N.N. 1996. Synthesis of functionally modified oligonucleotides from methoxyoxalamido precursors. Tetrahedron Lett. 37:32313234. Pon, R.T., Usman, N., Damha, M.J., and Ogilvie, K.K. 1986. Prevention of guanine modification and chain cleavage during the solid phase synthesis of oligonucleotides using phosphoramidite derivatives. Nucl. Acids Res. 14:6453-6470.
Oligodeoxyribonucleotide Synthesis Using the Phosphoramidite Method
Pon, R.T. 1987. Enhanced coupling efficiency using 4-dimethylamino pyridine (DMAP) and either tetrazole, 5-(o-nitrophenyl) tetrazole or 5-(pnitrophenyl) tetrazole in the solid phase synthesis of oligoribonucleotides by the phosphoramidite procedure. Tetrahedron Lett. 28: 3643-3646.
Rao, M.V. and Macfarlane, K. 1994. Solid phase synthesis of phosphorothioate oligonucleotides using benzyltriethylammonium tetrathiomolybdate as a rapid sulfur transfer reagent. Tetrahedron Lett. 35:6741-6744. Rao, M.V., Reese, C.B., and Zhengyun, Z. 1992. Dibenzoyl tetrasulphide—A rapid sulphur transfer agent in the synthesis of phosphorothioate analogues of oligonucleotides. Tetrahedron Lett. 33:4839-4842. Reddy, M.P., Hanna, N.B., and Farooqui, F. 1994. Fast cleavage and deprotection of oligonucleotides. Tetrahedron Lett. 35:4311-4314. Reddy, M.P., Hanna, N.B., and Farooqui, F. 1997. Ultrafast cleavage and deprotection of oligonucleotides—Synthesis and use of CAc derivatives. Nucleosides Nucleotides 16:1589-1598. Regan, J.B., Phillips, L.R., and Beaucage, S.L. 1992. Large-scale preparation of the sulfurtransfer reagent 3H-1,2-benzodithiol-3-one 1,1dioxide. Org. Prep. Proc. Int. 24:488-492. Roelen, H.C.P.F., Kamer, P.C.J., van den Elst, H., van der Marel, G.A., and van Boom, J.H. 1991. A study on the use of phenylacetyl disulfide in the solid-phase synthesis of oligodeoxynucleoside phosphorothioates. Recl. Trav. Chim. Pays-Bas 110:325-331. Sanghvi, Y.S. and Cook, P.D. 1994. Carbohydrates: Synthetic methods and applications in antisense therapeutics. In ACS Symposium Series 580— Carbohydrate Modifications in Antisense Research (Y.S. Sanghvi and P.D. Cook, eds.) pp. 1-22. American Chemical Society, Washington, D.C. Schulhof, J.C., Molko, D., and Teoule, R. 1987. The final deprotection step in oligonucleotide synthesis is reduced to a mild and rapid ammonia treatment by using labile base-protecting groups. Nucl. Acids Res. 15:397-416. Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., deBear, J.S., and Gough, G.R. 1992. Rapid synthesis of oligoribonucleotides using 2′-O-(o-nitrobenzyloxymethyl)-protected monomers. Bioorg. Med. Chem. Lett. 2:1019-1024. Sekine, M., Masuda, N., and Hata, T. 1986. Synthesis of oligodeoxyribonucleotides involving a rapid procedure for the removal of base-protecting groups by use of the 4,4′,4′′-tris(benzoyloxy)trityl (TBTr) group. Bull. Chem. Soc. Jpn. 59:1781-1789. Seliger, H. and Gupta, K.C. 1985. Three-phase synthesis of oligonucleotides. Angew. Chem. Int. Ed. Engl. 24:685-687. Sinha, N.D., Biernat, J., McManus, J., and Köster, H. 1984. Polymer support oligonucleotide synthesis XVIII: Use of β-cyanoethyl-N,N-dialkylamino-/N-morpholino phosphoramidite of deoxynucleosides for the synthesis of DNA fragments simplifying deprotection and isolation of the final product. Nucl. Acids Res. 12:45394557.
3.3.18 Current Protocols in Nucleic Acid Chemistry
Sinha, N.D., Davis, P., Usman, N., Pérez, J., Hodge, R., Kremsky, J., and Casale, R. 1993. Labile exocyclic amine protection of nucleosides in DNA, RNA and oligonucleotide analog synthesis facilitating N-deacylation, minimizing depurination and chain degradation. Biochimie 75:1323. Sondek, J. and Shortle, D. 1992. A general strategy for random insertion and substitution mutagenesis: Substoichiometric coupling of trinucleotide phosphoramidites. Proc. Natl. Acad. Sci. U.S.A. 89:3581-3585. Stec, W.J. and Zon, G. 1984. Stereochemical studies of the formation of chiral internucleotide linkages by phosphoramidite coupling in the synthesis of oligodeoxyribonucleotides. Tetrahedron Lett. 25:5279-5282. Stec, W.J., Uznanski, B., Wilk, A., Hirschbein, B.L., Fearon, K.L., and Bergot, B.J. 1993. Bis(O,O-diisopropoxy phosphinothioyl) disulfide—A highly efficient sulfurizing reagent for cost-effective synthesis of oligo(nucleoside phosphorothioate)s. Tetrahedron Lett. 34:5317-5320. Tanaka, T. and Letsinger, R.L. 1982. Syringe method for the stepwise chemical synthesis of oligonucleotides. Nucl. Acids Res. 10:32493260. Tener, G.M. 1961. 2-Cyanoethyl phosphate and its use in the synthesis of phosphate esters. J. Am. Chem. Soc. 83:159-168. Ugi, I., Jacob, P., Landgraf, B., Rupp, C., Lemmen, P., and Verfürth, U. 1988. Phosphite oxidation and the preparation of five-membered cyclic phosphorylating reagents via the phosphites. Nucleosides Nucleotides 7:605-608. Usman, N., Pon, R.T., and Ogilvie, K.K. 1985. Preparation of ribonucleoside 3′-O-phosphoramidites and their application to the automated solid phase synthesis of oligonucleotides. Tetrahedron Lett. 26:4567-4570. Uznanski, B., Grajkowski, A., and Wilk, A. 1989. The isopropoxyacetic group for convenient base protection during solid-support synthesis of oligodeoxyribonucleotides and their triester analogs. Nucl. Acids Res. 17:4863-4871. Vargeese, C., Carter, J., Yegge, J., Krivjansky, S., Settle, A., Kropp, E., Peterson, K., and Pieken, W. 1998. Efficient activation of nucleoside phosphoramidites with 4,5-dicyanoimidazole during oligonucleotide synthesis. Nucl. Acids Res. 26:1046-1050. Virnekäs, B., Ge, L., Plückthun, A., Schneider, K.C., Wellnhofer, G., and Moroney, S.E. 1994. Trinucleotide phosphoramidites: ideal reagents for the synthesis of mixed oligonucleotides for random mutagenesis. Nucl. Acids Res. 22:5600-5607. Vu, H. and Hirschbein, B.L. 1991. Internucleotide phosphite sulfurization with tetraethylthiuram disulfide. Phosphorothioate oligonucleotide synthesis via phosphoramidite chemistry. Tetrahedron Lett. 32:3005-3008.
Vu, H., McCollum, C., Jacobson, K., Theisen, P., Vinayak, R., Spiess, E., and Andrus, A. 1990. Fast oligonucleotide deprotection phosphoramidite chemistry for DNA synthesis. Tetrahedron Lett. 31:7269-7272. Weber, H. and Khorana, H.G. 1972. CIV. Total synthesis of the structural gene for an alanine transfer ribonucleic acid from yeast. Chemical synthesis of an icosadeoxynucleotide corresponding to the nucleotide sequence 21 to 40. J. Mol. Biol. 72:219-249. Weiss, P. 1998. TOM-protecting group—A major improvement in RNA synthesis. The Glen Report 11:2-4. Wilk, A., Srinivasachar, K., and Beaucage, S.L. 1997. N-Trifluoroacetylamino alcohols as phosphodiester protecting groups in the synthesis of oligodeoxyribonucleotides. J. Org. Chem. 62:6712-6713. Wincott, F., DiRenzo, A., Shaffer, C., Grimm, S., Tracz, D., Workman, C., Sweedler, D., Gonzalez, C., Scaringe, S., and Usman, N. 1995. Synthesis, deprotection, analysis and purification of RNA and ribosymes. Nucl. Acids Res. 23:2677-2684. Wolter, A., Biernat, J., and Köster, H. 1986. Polymer support oligonucleotide synthesis XX: Synthesis of a henhectacosa deoxynucleotide by use of a dimeric phosphoramidite synthon. Nucleosides Nucleotides 5:65-77. Wu, X. and Pitsch, S. 1998. Synthesis and pairing properties of oligoribonucleotide analogues containing a metal-binding site attached to β-D-allofuranosyl cytosine. Nucl. Acids Res. 26:43154323. Xin, Z. and Just, G. 1996. Diastereoselective synthesis of phosphite triesters. Tetrahedron Lett. 37:969-972. Xu, Q., Barany, G., Hammer, R.P., Musier-Forsyth, K. 1996. Efficient introduction of phosphorothioates into RNA oligonucleotides by 3ethoxy-1,2,4-dithiazoline-5-one (EDITH). Nucl. Acids Res. 24:3643-3644. Yamana, K., Nishijima, Y., Oka, A., Nakano, H., Sangen, O., Ozaki, H., and Shimidzu, T. 1989. A simple preparation of 5′-O-dimethoxytrityl deoxyribonucleoside 3′-O-phosphor-bisdiethylamidites as useful intermediates in the synthesis of oligodeoxyribonucleotides and their phosphorodiethylamidate analogs on a solid support. Tetrahedron 45:4135-4140. Yu, D., Tang, J.-Y., Iyer, R.P., and Agrawal, S. 1994. Diethoxy N,N-diisopropyl phosphoramidite as an improved capping reagent in the synthesis of oligonucleotides using phosphoramidite chemistry. Tetrahedron Lett. 35:8565-8568. Zehl, A., Starke, A., Cech, D., Hartsch, T., Merkl, R., and Fritz, H.-J. 1996. Efficient and flexible access to fully protected trinucleotides suitable for DNA synthesis by automated phosphoramidite chemistry. Chem. Commun. 26772678.
Synthesis of Unmodified Oligonucleotides
3.3.19 Current Protocols in Nucleic Acid Chemistry
Zemlicka, J. and Holy, A. 1967. Preparation of N-dimethylaminomethylene derivatives—A new method of a selective substitution of nucleoside amino groups. Coll. Czech. Chem. Commun. 32:3159-3168. Zhang, Z., Nichols, A., Alsbeti, M., Tang, J.X., and Tang, J.Y. 1998. Solid phase synthesis of oligonucleotide phosphorothioate analogues using bis(ethoxythiocarbonyl)tetrasulfide as a new sulfur-transfer reagent. Tetrahedron Lett. 39:24672470. Zhang, Z., Nichols, A., Tang, J.X., Han, Y., and Tang, J.Y. 1999. Solid phase synthesis of oligonucleotide phosphorothioate analogues using 3methyl-1,2,4-dithiazolin-5-one (MEDITH) as a new sulfur-transfer reagent. Tetrahedron Lett. 40:2095-2098.
Zon, G., Gallo, K.A., Samson, C.J., Shao, K., Summers, M.F., and Byrd, R.A. 1985. Analytical studies of ‘mixed sequence’ oligodeoxyribonucleotides synthesized by competitive coupling of either methyl- or β-cyanoethyl-N,N-diisopropylamino phosphoramidite reagents, including 2′deoxyinosine. Nucl. Acids Res. 13:8181-8196.
Contributed by Serge L. Beaucage Center for Biologics Evaluation and Research Food and Drug Administration Bethesda, Maryland Marvin H. Caruthers University of Colorado Boulder, Colorado
Zhao, Z. and Caruthers, M.H. 1996. Synthesis and preliminary biochemical studies with 5′-deoxy5′-methylidyne phosphonate linked thymidine oligonucleotides. Tetrahedron Lett. 37:62396242.
Oligodeoxyribonucleotide Synthesis Using the Phosphoramidite Method
3.3.20 Current Protocols in Nucleic Acid Chemistry
Synthesis of Oligodeoxyribo- and Oligoribonucleotides According to the H-Phosphonate Method This protocol outlines a general procedure for the preparation of oligodeoxyribo- and oligoribonucleotides using H-phosphonate monomers. It is followed by an in-depth discussion of the advantages of the H-phosphonate approach as well as its underlying chemistry (see Commentary). The preparation of the H-phosphonate monomers can be achieved by a variety of methods; these are presented in UNIT 2.6.
UNIT 3.4
BASIC PROTOCOL
Oligonucleotide synthesis employing H-phosphonates is considerably simpler than synthesis using the phosphotriester or phosphoramidite procedures. The elongation cycle includes only two chemical steps: deprotection of the terminal 5 -OH function of the support-bound oligonucleotide, and coupling of the 5 -OH with a nucleoside 3 -Hphosphonate in the presence of a condensing agent (Fig. 3.4.1). After completion of the desired number of elongation cycles (i.e., assembly of the oligomeric chain), a single oxidation cycle is performed to convert the internucleoside H-phosphonate functions to phosphodiesters (or some analog, such as phosphorothioates). Finally, the linkage between the oligomer and the support is cleaved under ammonolytic conditions, which is also the final deprotection step for oligodeoxyribonucleotide and oligoribonucleotide synthesis with 2 -O-2-chlorobenzoyl groups (also see UNIT 2.6). Purification by standard methods is then carried out to isolate the oligonucleotides. The synthetic protocol described below was optimized for oligoribonucleotide synthesis with 5 -O-MMTr and 2 -O-TBDMS protection on a modified Gene Assembler (Pharmacia) with a polystyrene support. It also gives good results with 2 -O-alkyl RNA and fairly good results for oligodeoxyribonucleotide synthesis. Controlled-pore glass (CPG) can be used, but has proven to be less reliable in the authors’ experience. The efficiency of each elongation step in solid-phase oligonucleotide synthesis is usually high, but technical aspects of the procedure may have to be adjusted for each particular machine. The most important of these are probably (1) the time of H-phosphonate preactivation before it reaches the solid support, (2) the concentration of the condensing agent, and (3) the proportion of pyridine in the solvent mixture. Some recently introduced condensing agents—e.g., bis(pentafluorophenyl) carbonate (Efimov et al., 1993) and carbonium- and phosphonium-based condensing agents (Wada et al., 1997)—seem to make the condensation less sensitive to these factors. The reaction conditions for the removal of the acid-labile 5 -O-MMTr or 5 -O-DMTr groups (usually 1% to 2% of various haloacetic acids in an anhydrous chlorinated solvent) have been shown not to affect the integrity of the H-phosphonate linkages within a relevant time (Stawinski et al., 1988). The most important factors affecting the condensation and oxidation steps are discussed later (see Commentary).
Materials 1,2-Dichloroethane (DCE; BDH) over 4-Å molecular sieves Trifluoroacetic acid (TFA; Fluka) Dichloroacetic acid (DCA; Lancaster, 99%), distilled Acetonitrile (MeCN; Lab-Scan) over 3-Å molecular sieves Synthesis of Unmodified Oligonucleotides Contributed by Roger Str¨omberg and Jacek Stawinski Current Protocols in Nucleic Acid Chemistry (2004) 3.4.1-3.4.15 C 2004 by John Wiley & Sons, Inc. Copyright
3.4.1 Supplement 19
Figure 3.4.1 Condensation of protected nucleoside H-phosphonate monoester with a nucleoside and conversion to the dinucleoside phosphate or backbone-modified analog.
Pyridine (Py; Lab-Scan, Anhydroscan) over 4-Å molecular sieves Protected nucleoside 3 -H-phosphonate building blocks (triethylammonium salts; see Commentary and UNIT 2.6) Pivaloyl chloride (Pv-Cl; Acros Organics, 99%), freshly distilled Polystyrene support (PE Applied Biosystems) loaded at 20 to 30 µmol/g with a 5 -O-(4,4 -dimethoxytrityl) (DMTr)– or 5 -O-(4-monomethoxytrityl) (MMTr)–protected nucleoside succinate (or equivalent nucleoside-loaded solid support) I2 Diethyl ether Concentrated (28% to 32%) aqueous ammonium hydroxide (NH4 OH) or 3:1 (v/v) concentrated NH4 OH/ethanol Triethylamine trihydrofluoride (for RNA synthesis with 2 -O-TBDMS protection) n-Butanol 20 mM sodium acetate buffer, pH 6.5, containing 30% and 10% MeCN LiClO4 0.1 M triethylammonium acetate (TEAA) buffer, pH 6.5 Automated oligonucleotide synthesizer (Gene Assembler, Pharmacia) 5-mL syringes with Luer lock 1.5-mL cryovials with screw caps Glass sintered funnel of coarse porosity Speedvac evaporator and a vacuum pump C18 cartridge (Waters Sep-Pac) Syringe filters (Millex-GV13 filter, 0.22-µm, 13 mm) and disposable syringes 4 × 250–mm Dionex NucleoPac PA-100 column Lyophilizer 4.6 × 150–mm Supelcosil LC-18 (3 µm) column Additional reagents and equipment for automated oligonucleotide synthesis (APPENDIX 3C) and purification by ion-exchange and reversed-phase HPLC (UNIT 10.5) Synthesis of Oligonucleotides According to the H-Phosphonate Method
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Table 3.4.1 Stepwise Oligonucleotide Synthesis Program
Step
Reagenta
Wash
DCE
Detritylation
Time (min)
Flow rate (mL/min)
2.0
2
1.0
2
2.0
2
3.5% DCA/DCE
2.5
2
DCE
2.0
2
MeCN
1.0
2
3:1 (v/v) MeCN/Py
1.0
2
50 mM phosphonate
0.1
1
225 mM Pv-Cl
0.1
1
50 mM phosphonate
0.1
1
225 mM Pv-Cl
0.1
1
50 mM phosphonate
0.1
1
Pump forward
0.4
1
Pump reverse
1.0
0.5
3:1 (v/v) MeCN/Py
1.0
2
MeCN
1.0
2
For 5 -O-MMTr-RNA or 2 -O-alkyl-RNA: 1% TFA/DCE
For 5 -O-DMTr-DNA: 3.5% DCA/DCE
For mixed 5 -O-MMTr-RNA/2 -O-alkyl-RNA and DNA: Wash
Coupling
Wash Total time per cycle
10.9–12.4
a Abbrevations: DCA, dichloroacetic acid; DCE, 1,2-dichloroethane; DMTr, 4,4 -dimethoxytrityl; MeCN, acetonitrile;
MMTr, 4-methoxytrityl; Pv-Cl, pivaloyl chloride; Py, pyridine; TFA, trifluoroacetic acid.
Prepare reagents 1. Charge the synthesizer with solvents and detritylation solution needed for the steps shown in Table 3.4.1. These can usually be kept on the synthesizer until they are consumed.
2. Dissolve the protected nucleoside 3 -H-phosphonates (triethylammonium salts) in pyridine and evaporate the solvent (two times) under reduced pressure in a rotary evaporator. Use 15 µmol/coupling plus 15 µmol extra for margins and priming of solutions. 3. Dissolve the residue in 3:1 (v/v) MeCN/pyridine to a concentration of 50 mM and transfer to appropriate vessels for attachment to the synthesizer. 4. Prepare a solution of 225 mM pivaloyl chloride in 3:1 (v/v) MeCN/pyridine directly in the vessel that will be used for synthesis. Prepare 0.2 mL/coupling plus 1 to 2 mL extra for margins. Attach the vessel to the synthesizer. 5. Prime the solutions and connect a column/cartridge filled with 0.2 to 1 µmol nucleoside-loaded support. 6. Immediately before the final oxidation step, prepare an iodine solution by dissolving 0.4 g I2 in 20 mL of 98:2 (v/v) pyridine/water.
Synthesis of Unmodified Oligonucleotides
The solution should be prepared no more than 30 min before use.
3.4.3 Current Protocols in Nucleic Acid Chemistry
Supplement 19
Table 3.4.2 Final Oxidation Program
Step
Reagenta
Wash
Time (min)
Flow rate (mL/min)
DCE
2
2
Detritylation
1% TFA/DCE
1
2
Wash
DCE
2
2
MeCN
1
2
Oxidation
2% I2 in 98:2 (v/v) Py/H2 O
30
0.5
Wash
MeCN
10
2
Total time
46
a Abbrevations: DCE, 1,2-dichloroethane; MeCN, acetonitrile; Py, pyridine; TFA, trifluoroacetic acid.
Synthesize oligonucleotide 7. Perform automated oligonucleotide synthesis (APPENDIX 3C) using the synthesis cycle shown in Table 3.4.1, with a final oxidation step as shown in Table 3.4.2. In the coupling step, program the synthesis so that the H-phosphonate and pivaloyl chloride are taken up in alternating 100-µL portions. The volume of the tubing between the valve and the column (including the pump hose of the peristaltic pump, through which the reagents flow) is 0.4 mL, which means that the front of the condensation mixture reaches the column when the last segment is taken up from the reagent bottles. The segments are then passed through the column (0.4 min at 1 mL/min), and then the flow is lowered and the direction reversed (1 min at 0.5 mL/min), giving a total condensation cycle time of 1.9 min, with an effective condensation time of ∼1.5 min. The effective time of preactivation for the nucleoside H-phosphonate when it first reaches the column is ∼0.3 min, for a total of 1.8 min at the end of condensation.
Purify oligonucleotide 8. Remove the cartridge/column from the synthesizer and wash the support with ∼10 mL diethyl ether using a 5-mL syringe with a Luer fitting. 9. Air dry the support using the above 5-mL syringe with a Luer fitting by pushing air through the column a few times. 10. Remove the support from the cartridge and transfer it to a 1.5-mL cryovial (with screw cap). 11. Carry out ammonolysis by adding 3:1 (v/v) concentrated NH4 OH/ethanol (∼1 mL for up to a 1-µmol scale synthesis) and incubating 8 to 16 hr at 20◦ to 25◦ C (8-hr incubations for sequences up to 25-mers; 16-hr incubations for longer sequences, e.g., 50- to 60-mers). The above conditions are for RNA oligomers with N2 -phenoxyacetyl guanosine protection, N6 -butyryl adenosine protection, and N4 -propionyl cytidine protection, which are recommended to avoid cleavage of TBDMS groups (and subsequent cleavage of RNA) upon ammonolysis at elevated temperatures (Stawinski et al., 1988), which is required for more stable base protection. Other reagents and conditions may be needed for other protecting groups. For DNA or for 2 -O-alkyl-RNA, concentrated NH4 OH should be used alone for ∼20 hr at 55◦ to 60◦ C.
12. Filter off the support using a glass sintered funnel of medium porosity. Synthesis of Oligonucleotides According to the H-Phosphonate Method
13. Wash support with 1 mL of 3:1 (v/v) concentrated NH4 OH/ethanol (or concentrated NH4 OH alone) and combine the filtrates. 14. Concentrate the oligonucleotide using a Speedvac evaporator and a vacuum pump.
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15. For RNA synthesis with 2 -O-TBDMS protection, perform deprotection step with triethylamine trihydrofluoride (Westman and Str¨omberg, 1994) using the following steps: a. Dissolve the residue in 0.3 mL neat triethylamine trihydrofluoride and incubate 14 to 16 hr at room temperature. b. Add 30 µL water and 1 mL n-butanol, and incubate 1 hr at –20◦ C. c. Centrifuge and remove the liquid supernatant. Dissolve the pellet in HPLC buffer (see step 16). 16. Purify by ion-exchange and reversed-phase HPLC (UNIT 10.5). a. Dissolve the deprotected oligoribonucleotides in 0.5 mL of 20 mM sodium acetate buffer, pH 6.5, containing 30% MeCN and filter through a disposable C18 cartridge. b. Wash the cartridge with 1 mL of buffer, then combine fractions and filter through a disposable syringe attached to a 0.22-µm filter before HPLC purification. c. Purify by anion-exchange HPLC on a 4 × 250–mm Dionex NucleoPac PA-100 column using a linear gradient of LiClO4 in 20 mM sodium acetate buffer, pH 6.5, containing 10% MeCN at a flow rate of 1 mL/min. For analysis, inject 0.2 OD260 units; for purification, inject 15 to 30 OD260 units of crude oligoribonucleotides. d. Lyophilize the collected fractions, dissolve in 1 mL of 0.1 M triethylammonium acetate (TEAA) buffer, pH 6.5, and filter through a disposable syringe attached to a 0.22-µm filter. e. Further purify by reversed-phase HPLC on a 4.6 × 150–mm Supelcosil LC-18 column using a linear gradient of MeCN in 0.1 M TEAA buffer, pH 6.5, at a flow rate of 1 mL/min. f. Collect the fractions containing the product, lyophilize, dissolve in 1 mL water, and lyophilize again.
COMMENTARY Background Information General information and synthetic strategies Although the most common method today for synthesis of oligonucleotides and their analogs is the phosphoramidite approach (Beaucage and Iyer, 1993, see also UNIT 3.3), the newer H-phosphonate methodology can often be a preferred alternative (Garegg et al., 1985, 1986a,b,c; Froehler and Matteucci, 1986; Froehler et al., 1986). The use of H-phosphonates in nucleotide synthesis was pioneered by Sir Todd’s group in Cambridge, UK, who in 1952 demonstrated the formation of H-phosphonate diesters in a condensation reaction of H-phosphonate monoesters with a protected nucleoside, promoted by diphenyl phosphorochloridate (Corby et al., 1952; Hall et al., 1957). This chemical principle was, however, not explored further; it was rediscovered three decades later (Garegg et al., 1985, 1986c) and explored for oligonu-
cleotide synthesis (Froehler and Matteucci, 1986; Froehler et al., 1986; Garegg et al., 1986a,b,c, 1987a). The method consists of condensing a protected nucleoside H-phosphonate monoester (UNIT 2.6) with a nucleoside in the presence of a coupling agent to produce the corresponding dinucleoside H-phosphonate diester. This, under various experimental conditions, can be converted to the dinucleoside phosphate or to a variety of backbone-modified analogs, e.g., phosphorothioates, phosphoramidates, etc. (Fig. 3.4.1). The condensation step of the elongation cycles (i.e., the formation of an internucleoside H-phosphonate linkage between a nucleoside 3 -H-phosphonate monoester and the supportbound 5 -hydroxylic component) is usually carried out in pyridine-acetonitrile mixtures. Out of the various condensing agents initially tested, pivaloyl chloride (Pv-Cl) gave the best results in automated solid-support synthesis
Synthesis of Unmodified Oligonucleotides
3.4.5 Current Protocols in Nucleic Acid Chemistry
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Synthesis of Oligonucleotides According to the H-Phosphonate Method
of oligonucleotides, and it is still the most frequently used reagent. The reaction in pyridine or acetonitrile-pyridine mixtures using 2 to 5 equiv of Pv-Cl is usually fast and goes to completion in <1 min. Nowadays, an array of other condensing agents (see below) is available; these reagents can, in some instances, be superior to Pv-Cl. The H-phosphonate methodology became commercially available (for use in automated synthesizers from Applied Biosystems and Biosearch) soon after the initial reports on oligonucleotide synthesis using this approach were published. The research group at Applied Biosystems also introduced 1adamantanecarbonyl chloride as an alternative to Pv-Cl (Andrus et al., 1988). Although a capping step during oligonucleotide synthesis via H-phosphonate intermediates initially seemed to be superfluous, its incorporation into the synthetic protocol was shown to potentially improve the overall performance of the method. The procedure simply consists of an additional condensation step using isopropyl H-phosphonate after each nucleoside H-phosphonate coupling (Andrus et al., 1988). The use of 2-cyanoethyl Hphosphonate for capping was also reported, although this reagent has the disadvantage of being less accessible. The capping procedure routinely used in the phosphoramidite approach (acylation with acetic anhydride in the presence of nucleophilic catalysts more powerful than pyridine) is not compatible with H-phosphonate chemistry, owing to the occurrence of P-acylation (see below). Since the introduction of the H-phosphonate methodology for synthesis of oligonucleotides, a number of improvements have been introduced. These involve new condensing agents and new reaction conditions. Oligodeoxynucleotide synthesis via H-phosphonates has been scaled up to 10 to 14 µmol using only a few equivalents of monomeric building blocks relative to a support-bound oligomer per condensation (Gaffney and Jones, 1988). The H-phosphonate approach was used in a cartridgebased procedure for simultaneous synthesis of multiple oligodeoxynucleotides (Seliger and R¨osch, 1990). The H-phosphonate methodology has been used in the synthesis of a number of different oligonucleotide analogs, for example, phosphoramidates (Froehler, 1986) and phosphorothioates (Agrawal and Tang, 1990; Stein et al., 1990), including those with all-Rp -linkages (Almer et al., 1996). It has also been used in the synthesis of oligonu-
cleotides bearing modified heterocyclic bases (Ramzaeva et al., 1997; Seela and Wei, 1997). One particularly interesting feature of oligonucleotide synthesis when using the Hphosphonate methodology is that it can be performed without protection of the nucleobases (Kung and Jones, 1992; Wada et al., 1997; UNIT 3.10). RNA synthesis is perhaps where the advantages of the H-phosphonate approach are most apparent. In the first report, the t-butyldimethylsilyl (TBDMS) group was used for protection of the 2 -OH and the protocol was similar to that described for DNA synthesis (Garegg et al., 1986a,b). A number of subsequent reports have differed in the choice of 2 -OH protection (see UNITS 2.2 and 3.5). These include the use of photolabile groups (e.g., o-nitrobenzyl; Tanaka et al., 1987), acid-sensitive groups (such as 1-(2chloro-4-methylphenyl)-4-methoxypiperidin4-yl; Ctmp; Sakatsume et al., 1989), and base-labile benzoyl derivatives (Rozners et al., 1988, 1990, 1992). The H-phosphonate approach seems to be even more suited for RNA than for DNA synthesis. Potential problems such as double activation (i.e., the bis-acyl phosphite formation) and P-acylation are significantly suppressed when using ribonucleoside Hphosphonates (Stawinski et al., 1991b). The condensation reaction is virtually as fast as for DNA synthesis and at least as efficient, even with bulky 2 -O-protection. Synthesis of oligoribonucleotides of up to 50 to 60 nucleotide residues in length is readily achieved (Agrawal and Tang, 1990; Rozners et al., 1998), and the deprotection steps have been adjusted to more base-labile N-protection that gives better performance (Rozners et al., 1998). The most common choice for 2 -OH protection in oligoribonucleotide synthesis is the TBDMS group (see UNIT 3.5). Studies on the stability of this group under the reaction conditions used for removal of N-acyl groups have been carried out (Stawinski et al., 1988; Wu and Ogilvie, 1990). A conclusion (relevant not only to the H-phosphonate approach) was that synthesis of longer RNA oligonucleotides with 2 -O-TBDMS protection would benefit from the use of more labile N-acylprotecting groups for the heterocyclic base moieties. These were later duly introduced to the synthesis of oligoribonucleotides, both by the phosphoramidite approach (Wu and Ogilvie, 1988; Chaix et al., 1989) and the H-phosphonate method (Westman et al. 1993, 1994; Rozners et al., 1998).
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Although Pv-Cl (Froehler et al., 1986; Garegg et al., 1986a) and perhaps adamantane carbonyl chloride (Andrus et al., 1988) are the most popular condensing agents used in solidphase synthesis of oligonucleotides via the H-phosphonate approach, several others that may be considered for different applications have also been evaluated. These include various pivalic (Str¨omberg and Stawinski, 1987) or other carboxylic acid derivatives (J¨ager et al., 1987), diphenyl phosphorochloridate (Hall et al., 1957; Garegg et al., 1985), 2-chloro5,5-dimethyl-2-oxo-1,3,2-dioxaphosphinane (Str¨omberg and Stawinski, 1987), bis(2oxo-3-oxazolidinyl)phosphinic chloride (Str¨omberg and Stawinski, 1987), various dialkyl phosphorochloridates (Str¨omberg and Stawinski, 1987; Stawinski et al., 1988), arene sulfonic acid derivatives (Garegg et al., 1985; J¨ager et al., 1987; Dan’kov et al., 1988), 1,3-dimethyl-2-chloro-imidazolium
chloride (Sakatsume et al., 1989, 1990), bis(pentafluorophenyl)carbonate (Efimov et al., 1993), and a number of carbonium and phosphonium derivatives (Wada et al., 1997). Parameters of the underlying chemistry Activation of H-phosphonate monoesters The formation of an internucleoside linkage by the H-phosphonate approach, using acyl chlorides, proceeds via a mixed phosphonocarboxylic anhydride that reacts with the nucleosidic component to produce an Hphosphonate diester (Garegg et al., 1987c; Fig. 3.4.2). The order in which the reactants are added in solution synthesis with near equimolar amounts of phosphonate and nucleoside was found to be important for the efficiency of H-phosphonate diester formation (Garegg et al., 1985, 1986c). The coupling reaction is virtually quantitative when the condensing
Figure 3.4.2 Activation of H-phosphonate monoesters. R1 = protected nucleoside-3 -yl; R2 = protected nucleoside-5 -yl.
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Synthesis of Oligonucleotides According to the H-Phosphonate Method
agent is added to a solution containing both the alcohol and the H-phosphonate monoester. If, on the other hand, the H-phosphonate is preactivated with a condensing agent before the addition of an alcohol, the reaction is considerably less efficient. This phenomenon was investigated and traced back to the further activation of the initially formed intermediates in the absence of an alcohol (Garegg et al., 1987c,d,e). Irrespective of the condensing agent used, tricoordinated phosphite derivatives were formed; upon reaction with an alcohol, these species (the chemical nature of which depends on the coupling agent used) always gave various unwanted products in addition to the H-phosphonate diesters. Preactivation of H-phosphonate monoesters with acyl chlorides produces bis-acyl phosphites (Garegg et al., 1987c). The consequence of such preactivation is that, in solution synthesis with near equimolar amounts of H-phosphonate and alcohol, formation of phosphite triesters is unavoidable (at least to some extent). It is quite clear that such species must also be formed to some extent when the condensing agent and H-phosphonate are mixed before entering the column in a synthesizer. No side products that could arise from this species have so far been detected in solid-phase synthesis products, but it has been shown that too long of a preactivation substantially slows down the coupling reaction, and the competing reaction of alcoholic functions with the condensing agent becomes more pronounced (Gaffney and Jones, 1988). It has also been shown that even if condensation with nucleoside bis-acyl phosphites gave the correct product on a solid support, the reaction of a bis-acyl phosphite with a nucleoside is substantially slower than that of the corresponding H-phosphonate which is postulated to proceed via the mixed phosphono-carboxylic anhydride (shown in Fig. 3.4.2; Garegg et al., 1987c; Efimov and Dubey, 1990). Because an excess of H-phosphonate to alcohol is normally used in solid-phase synthesis, some formation of bis-acyl phosphite is tolerable. It is clear that the preactivation time should be minimized so that the mixture reaching the column has the highest ratio of mixed phosphono-carboxylic anhydride to bis-acyl phosphite as possible. Otherwise, the oligonucleotide synthesis will lose in efficiency. A preactivation time that is too long can probably not be compensated to the full extent by longer condensation time, because competing reactions become more prominent. Apart from minimizing preactivation time, it is
also possible to reduce or change the amount of condensing agent (which is usually used in excess) and control the reaction by varying solvent composition (see below). It is convenient to use enough condensing agent to achieve conditions that are not too moisture sensitive; but over a certain level, additional excess of acyl chloride increases the level of side reactions that may lower the coupling efficiency and overall yield. Particularly noteworthy was the observation that H-phosphonate monoesters when activated in the absence of a base (like pyridine) produce only monoactivated species (Garegg et al., 1987c). Efimov and co-workers found that replacing pyridine by the slightly weaker base quinoline is sufficient to suppress the formation of bis-acyl phosphite and keep its amount <5% to 10% within the period of time necessary for a condensation (Efimov and Dubey, 1990; Efimov et al., 1990). This novel solvent mixture (acetonitrile-quinoline, 4:1 v/v) was used in the synthesis of oligodeoxynucleotides up to 39 units in length (Efimov and Dubey, 1990). Although the replacement of pyridine by quinoline in oligonucleotide synthesis via the H-phosphonate approach offers some advantages, the rate of condensation is usually lower and quinoline gives some side reactions with activated H-phosphonate monoesters (Stawinski et al., 1991a). The influence of base strength on preactivation of ribonucleoside H-phosphonates was examined, and substituted pyridines that were less basic than pyridine itself were found to suppress formation of tricoordinated species (Stawinski et al., 1991b). With these less basic pyridine derivatives, the side reactions observed with quinoline could not be detected. Steric factors are also likely to be important for double activation, because, for example, the formation of tricoordinated species is slower for 2 -O-TBDMS-protected ribonucleoside H-phosphonates than for the corresponding 2 deoxyribo derivatives. Recently, bis(pentafluorophenyl) carbonate (PFPC) was advocated as a condensing agent for H-phosphonate diesters formation (Efimov et al., 1993). The reagent may be as reactive as Pv-Cl in promoting condensations and compares favorably with Pv-Cl in regard to its reactivity toward heterocyclic bases and the 5 hydroxyl function of a nucleosidic component (Efimov et al., 1993). Although the reagent produces double-activated species from Hphosphonate monoesters, these are as reactive as the initially formed monoactivated intermediate. Truncated sequences are claimed to be
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significantly reduced compared to the protocol that uses Pv-Cl as a condensing agent. The utility of the reagent was demonstrated in the synthesis of several oligodeoxyribonucleotides (chain length 22 to 40) and oligonucleotidephospholipid conjugates (Efimov et al., 1993). Competing O-acylation Direct reaction of a condensing agent with hydroxylic functions in competition with the desired coupling reaction is another potential source of inefficiency in oligonucleotide synthesis. The suggestion that capping is important when using the H-phosphonate approach for oligonucleotide synthesis points to a rather low extent of acylation during the condensation step (Andrus et al., 1988; Gaffney and Jones, 1988). Self-capping by the acyl chlorides used for condensation was estimated to account for only ∼10% to 50% of nonphosphonylated hydroxyls (Gaffney and Jones, 1988), which represents 0.1% to 0.5% of an average coupling yield of 99%. When a fourfold excess of H-phosphonate to nucleoside and 5 equiv Pv-Cl were reacted for 1.5 min, however, the total amount of 5 -O-acylation was ∼0.8% (Gaffney and Jones, 1988). The extent of O-acylation depends heavily on reaction conditions, such as ratios and concentrations of reagents, and is likely to be substantially lower when a larger excess of H-phosphonate is used in conjunction with fewer equivalents of Pv-Cl. The authors assert that the most important reason for the occurrence of competing 5 -O-acylation in machine-assisted synthesis is that the condensation is slowed down owing to formation of bis-acyl phosphites (see
Figure 3.4.3
P-acylation side reaction.
above). Acylation of 5 -OH functions has never been observed when the condensation reaction is carried out in solution without preactivation. Minimizing formation of the less reactive tricoordinated species (from preactivation) should result in higher coupling efficiency. Using fewer equivalents of condensing agent relative to H-phosphonate monoester is likely to be the most efficient way to decrease 5 -Oacylation, because it will affect the rate of both O-acylation and bis-acyl phosphite formation. Changes in solvent composition may also be beneficial (e.g., less pyridine). P-acylation Another side reaction that potentially can occur when using acyl chlorides as condensing agents in the H-phosphonate approach to oligonucleotide synthesis is the formation of acylphosphonates (Fig. 3.4.3). When the synthesis of an H-phosphonate diester is carried out with all reactants in solution under standard conditions (approximately equimolar amounts of H-phosphonate and nucleosidic components together with 2 to 3 equiv of coupling agent), the reaction can be quenched long before any P-acylation can be detected. In solid-support oligonucleotide synthesis, however, because of iterative condensation steps, reaction of H-phosphonate functions with the condensing agent could become a problem. Acylphosphonate formation was not detected when H-phosphonate diesters were exposed to 3 to 5 equiv of Pv-Cl in pyridine for 30 min (Regberg et al., 1988). In a study of P-acylation, however, a slow conversion of a di(deoxyribonucleoside) H-phosphonate to the corresponding pivaloylphosphonate went
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to completion in the presence of 5 equiv of PvCl in pyridine-acetonitrile after 15 hr at 20◦ C (Kuyl-Yeheskiely et al., 1986). The P-acylation during a condensation reaction should in principle be less pronounced, owing to the inhibitory effect of pyridinium ions that are formed (Str¨omberg, 1987). In a condensation mixture containing 5 equiv of Pv-Cl (in pyridine-acetonitrile 1:1 v/v), only ∼15% of the produced H-phosphonate diester is pivaloylated in 14 hr (Str¨omberg, 1987). In the presence of bases stronger than pyridine (e.g., N-methylimidazole, triethylamine), the P-acylation is considerably faster (Str¨omberg, 1987). Note, however, that even if P-acylation occurred to some extent during oligonucleotide synthesis, this modification could not be easily carried through to the fully deprotected oligonucleotide. Oligonucleotidic chains bearing acylphosphonate modifications will be cleaved during final deprotection upon treatment with aqueous ammonia, and thus only a decrease in the yield of oligomers with the desired chain length is expected. Reactions of protected ribonucleoside Hphosphonate diesters with Pv-Cl are considerably slower than those of deoxyribonucleoside derivatives (Stawinski et al., 1991b). The presence of a bulky 2 -O-TBDMS-protecting group vicinal to the H-phosphonate linkage in the ribonucleoside derivatives is likely to be one reason for the observed difference. In a typical condensation reaction (5 equiv of Pv-Cl in pyridine-acetonitrile; 1:1 v/v), only a few percent of the produced H-phosphonate diester is pivaloylated within 22 hr (Stawinski et al., 1991b), which corresponds to the time required for 880 standard condensations reactions. As a rough estimate, a 50-residue-long 2 -O-TBDMSoligoribonucleoside H-phosphonate synthesized with 5 equiv of Pv-Cl and 50% pyridineacetonitrile (1:1 v/v) gives truncated sequences (owing to P-acylation and subsequent cleavage) to a level of 0.1%; it is further estimated that decreasing the excess Pv-Cl to 3 equiv in pyridine-acetonitrile (1:3 v/v), would only produce ∼0.025% P-acylation. For oligodeoxyribonucleotide synthesis, the corresponding levels should be roughly 0.9% and 0.2% to 0.3% for a coupling time of 1 min. This suggests that P-acylation could well be responsible for a substantial part of the observation that DNA synthesis with H-phosphonates, in contradistinction to oligoribonucleotide synthesis, is not quite as efficient as with phosphoramidites (when using 5 equiv of PvCl in pyridine-acetonitrile, 1:1 v/v). It seems
clear that oligodeoxyribonucleotide synthesis would benefit even more than oligoribonucleotide synthesis from reducing the excess of coupling agent and pyridine (and possibly the basicity by using a substituted pyridine; see below). In the earlier discussion on preactivation, some advantages connected with using a slightly weaker base than pyridine as a co-solvent during the condensation step were mentioned. Such conditions are also advantageous when considering P-acylation. The extent of this reaction is substantially decreased when quinoline or some substituted pyridines are used instead of pyridine (Stawinski et al., 1991b). In the case of deoxyribonucleoside H-phosphonate diesters, only traces of P-acylation can be detected (31 P NMR) after 22 hr when 5 equiv of Pv-Cl in quinoline-acetonitrile (1:3 v/v) is used. For ribonucleoside H-phosphonate diesters, this side reaction cannot be detected even after 22 hr under the same conditions (Stawinski et al., 1991b). Other potential side reactions Not many studies on the extent of base modification when using the H-phosphonate approach for oligonucleotide synthesis have been reported. Error rates in oligodeoxynucleotides that were produced using the H-phosphonate methodology have been estimated by sequencing (Vasser et al., 1990). The average number of substitutions was found to be 1/1783 bases. These are not necessarily caused by base modification during synthesis, but they at least indicate the maximum level of modification one may anticipate to be present in purified oligomers. Guanosine-containing dinucleoside Hphosphonates have been reported to react with some condensing agents (Regberg et al., 1988). The guanine residue reacts with Pv-Cl to produce a modification (most likely pivaloylation of the lactam function) that is reversed by treatment with ammonia for a much shorter time than that used for the deprotection of oligonucleotides. Because this reaction is also considerably slower than the condensation, protection of the heteroaromatic lactam system in guanine seems to be unnecessary for most applications when using H-phosphonate chemistry. It may, however, be advisable to analyze for base modifications if novel condensing reagents are being used. Some concern has been expressed about the possibility of a partial loss of the 4,4 dimethoxytrityl group from 5 -protected
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H-phosphonates upon standing in vials on automated synthesizers (Froehler et al., 1986; Froehler and Matteucci, 1987). It was estimated that 5 -O-(4,4 -dimethoxytrityl)thymidine H-phosphonate (triethylammonium salt (TEAH+ )) dissolved in pyridine-acetonitrile (1:1 v/v) loses 2% of its 5 -protection within 4 weeks (Froehler and Matteucci, 1987). Assuming a first-order reaction, this would translate to ∼0.05% detritylation per 1000 min. If fresh solutions are prepared before synthesis, this can be considered a measure of the extent to which the loss of 5 -protection could be for the last added nucleotide unit in the synthesis of a 100-mer (assuming a 10-min cycle time). The replacement of TEAH+ by 2,3,4,6,7,8,9,10-octahydropyrimido[1,2a]azeponium (DBUH+ ) salts of nucleoside H-phosphonate monoesters was also claimed to decrease the amount of oligomers with a chain length of n + 1 (Froehler and Matteucci, 1987) owing to suppression of partial detritylation during the condensation step. It is, therefore, conceivable that the loss of the 5 -O-(4-monomethoxytrityl group) frequently used in RNA synthesis should probably be negligible under these conditions. Although the detritylation step has been shown to be completely compatible with H-phosphonate linkages (Stawinski et al., 1988), be aware that using acid-labile 2 -O-protecting groups in H-phosphonate-based oligoribonucleotide synthesis may cause problems (Rozners et al., 1994). These can be alleviated by using the fluoride-labile 2 -O-TBDMS or base-labile 2 -O-(2-chlorobenzoyl) groups. The former group can be recommended for the synthesis of longer oligoribonucleotides, whereas the latter can be selected for shorter fragments, owing to the simplicity and cost of preparation of starting materials. The oxidation step In the phosphoramidite approach, the oxidation step is carried out after each coupling reaction (Beaucage and Caruthers, 1981; Matteucci and Caruthers, 1981; see also UNIT 3.3). It may be feasible to do so via the H-phosphonate method, because the condensation reaction can be accomplished in the presence of phosphodiester functions (F¨oldesi et al., 1989; Gryaznov and Potapov, 1990). Given that H-phosphonate functions remain intact throughout the oligomeric chain assembly, the oxidation reaction can be carried out in a single step upon completion of the synthesis. This can be advantageous for the preparation of
uniformly modified oligonucleotides (e.g., elemental sulfur can produce phosphorothioate oligomers in one step from H-phosphonate oligonucleotides). The final oxidation step in the Hphosphonate method for oligonucleotide synthesis is usually performed using a solution of iodine in aqueous pyridine (Garegg et al., 1986a,b, 1987b) or with iodine-water in the presence of another base (triethylamine, N-methylimidazole; Froehler and Matteucci, 1986; Froehler et al., 1986). Completeness of the oxidation reaction is, of course, important for efficient synthesis of oligonucleotides. Any oligomeric chain containing an H-phosphonate function will be immediately cleaved in the subsequent ammonia treatment step (Hammond, 1962). Incomplete oxidation will produce shorter than expected oligomers, thus reducing the yield of the desired product. An oxidation time of 10 min with 2% iodine in pyridine-water (98:2) seems to be sufficient for solid-phase synthesis of 20-mers (Garegg et al., 1986a,b). Because the oxidation reaction needs to be carried out only once after chain assembly, one can extend the oxidation time (rather than using stronger bases) to ensure complete reaction within a marginal time loss. A longer oxidation time may still be advisable for longer oligonucleotides. Competitive hydrolysis of H-phosphonate diesters during aqueous oxidation may be considered a potential problem. This has not yet been reported, but it may be justified to discuss this issue because H-phosphonate diesters are very unstable under aqueous basic conditions (Nyl´en, 1937; Westheimer et al., 1988). The rate constant (kOH ) for hydroxide-catalyzed oxidation with iodine was reported to be ∼1400 to 4300 times larger than that for hydrolysis of a series of dialkyl H-phosphonates (Nyl´en, 1937, 1938). These values have been used to estimate that an aqueous pyridine solution containing 2% iodine should give no more than ∼5 to 25 ppm H-phosphonate diester cleavage (Stawinski and Str¨omberg, 1993). This suggests that hydrolysis during oxidation is not a serious problem in oligonucleotide synthesis according to the Hphosphonate approach. The rate of oxidation of nucleoside Hphosphonate diesters with iodine-pyridinewater can be increased by including a presilylation step (e.g., treatment with trimethylsilyl chloride) in the synthetic protocol (Hata and Sekine, 1974; Garegg et al., 1987b). This has, however, been used infrequently in oligonucleotide synthesis.
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Critical Parameters
Synthesis of Oligonucleotides According to the H-Phosphonate Method
The detritylation step has been shown to be completely compatible with H-phosphonate linkages (Stawinski et al., 1988). However, using acid-labile 2 -O-protecting groups in Hphosphonate-based oligoribonucleotide synthesis may cause problems (Rozners et al., 1994). These problems can be alleviated by using the fluoride-labile 2 -O-TBDMS or baselabile 2 -O-2-chlorobenzoyl groups. The former group can be recommended for the synthesis of longer oligoribonucleotides, while the latter can be a choice for shorter fragments, due to the simplicity and cost of preparing the starting materials. The preactivation time, i.e., contact of a condensing agent with nucleoside 3 -Hphosphonates before they reach the reaction chamber, should be minimized as much as possible, since an extended preactivation time adversely affects rate and efficiency of internucleotide bond formation. The amount of condensing agent and proportion of pyridine in the solvent is also important. It is convenient to use enough condensing agent to obtain conditions that are not too moisture sensitive. Over a certain level, however, excess acyl chloride increases the extent of side reactions that may lower the coupling efficiency and overall yield. The quality of the pivaloyl chloride is important for the outcome of the synthesis. Since the reagent decomposes with time, it should be distilled approximately once a month. The distilled material, divided into a number of smaller vials to avoid too frequent opening and closing of the same bottle, can be safely stored at −18◦ to –20◦ C. The best results are obtained if the reagent solutions are prepared fresh just before the start of oligonucleotide synthesis. This is most beneficial for the pivaloyl chloride solution and the oxidation solution. The latter degrades with time (particularly when a base stronger than pyridine is used in the oxidation solution) and it is recommended that a fresh oxidation solution be used for each synthesis. A shorter oxidation time than that stated in the protocol can be used for shorter oligonucleotides, but since this is only carried out once, the time savings are not significant and a standard time of 30 min that appears to result in complete oxidation (with some margin for 50- to 60-mers) may be best.
Anticipated Results
Since 2 -protected ribonucleoside 3 -Hphosphonates are less susceptible to double
activation with a condensing agent, and diribonucleoside H-phosphonate diesters are more resistant to P-acylation by acyl chlorides used as condensing agent, the H-phosphonate approach is particularly suited for the preparation of oligoribonucleotides. Because oxidation is carried out as a final synthetic step, the Hphosphonate approach can also be a method of choice for the preparation of uniformly modified oligonucleotides. With the reported protocol, one should be able to routinely synthesize oligoribonucleotides of up to ∼50 to 60 residues with high yield when using 5 -MMTr and 2 -O-TBDMS protection (and N4 -propionyl for cytidine, N6 butyryl for adenosine, and N2 -phenoxyacetyl for guanosine). For oligodeoxyribonucleotides, one could expect similar results. When using 2 -O-2-chlorobenzoyl protection, oligoribonucleotides of up to ∼20- to 30-mers can be produced with fairly good yields. Average coupling yields per step are usually 98% to 99+% depending on the building blocks used.
Time Considerations Using the protocol in this unit, the total time for the machine-assisted part of synthesis of a 20-mer and 50-mer RNA is 4 hr 13 min and 9 hr 40 min, respectively. The total time will depend upon protecting groups, linker to support, deprotection conditions, and purification procedures used. The procedure should not be stopped before the oxidation step, because the H-phosphonate linkages are rather labile. One can take a break after oxidation. At that point, the oligonucleotide is more stable than at the corresponding stage in the phosphoramidite method, since phosphodiesters are obtained after this step. The support-bound oligonucleotide can be stored at –20◦ C until the ammonolysis step is carried out to release the oligonucleotide from the support and remove the N-acyl protection.
Literature Cited Agrawal, S. and Tang, J.-Y. 1990. Efficient synthesis of oligoribonucleotide and its phosphorothioate analog using H-phosphonate approach. Tetrahedron Lett. 31:7541-7544. Almer, H., Stawinski, J., and Str¨omberg, R. 1996. Solid support synthesis of all Rpoligoribonucleoside phosphorothioates. Nucl. Acids Res. 24:3811-3820. Andrus, A., Efcavitch, J.W., McBride, L.J., and Giusti, B. 1988. Novel activating and capping reagent for improved hydrogen-phosphonate DNA synthesis. Tetrahedron Lett. 29:861-864.
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Beaucage, S.L. and Caruthers, M.H. 1981. Deoxynucleoside phosphoramidites—a new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Lett. 22:1859-1862. Beaucage, S.L. and Iyer, R.P. 1993. The synthesis of modified oligonucleotides by the phosphoramidite approach and their applications. Tetrahedron 49:6123-6194. Chaix, C., Molko, D., and Teoule, R. 1989. The use of labile base protecting groups in oligoribonucleotide synthesis. Tetrahedron Lett. 30:71-74. Corby, N.S., Kenner, G.W., and Todd, A.R. 1952. Nucleotides. Part XVI. Ribonucleoside-5 phosphites. A new method for the preparation of mixed secondary phosphites. J. Chem. Soc. 3669-3675. Dan’kov, Y.V., Batchikova, N.V., Scaptsova, N.V., Besidsky, E.S., and Azhayev, A.V. 1988. H-Phosphonate solid-phase synthesis of oligodeoxyribonucleotides in syringe. Bioorg. Khim. 14:615-620. Efimov, V.A. and Dubey, I.Y. 1990. Modification of the H-phosphonate oligonucleotide synthesis on polymer support. Bioorg. Khim. 16:211-218. Efimov, V.A., Dubey, I.Y., and Chakhmakhcheva, O.G. 1990. NMR study and improvement of Hphosphonate oligonucleotide synthesis. Nucleosides Nucleotides 9:473-477. Efimov, V.A., Kalinkina, A.L., and Chakhmakhcheva, O.G. 1993. Dipentafluorophenyl carbonate–a reagent for the synthesis of oligonucleotides and their conjugates. Nucl. Acids Res. 21:5337-5344. F¨oldesi, A., Balgobin, N., and Chattopadhyaya, J. 1989. Synthesis of a “branched” trinucleotide using the H-phosphonate chemistry. Tetrahedron Lett. 30:881-884. Froehler, B.C. 1986. Deoxynucleoside Hphosphonate diester intermediates in the synthesis of internucleotide phosphate analogs. Tetrahedron Lett. 27:5575-5578. Froehler, B.C. and Matteucci, M.D. 1986. Nucleoside H-phosphonates: Valuable intermediates in the synthesis of deoxyoligonucleotides. Tetrahedron Lett. 27:469-472. Froehler, B.C. and Matteucci, M.D. 1987. The use of nucleoside H-phosphonates in the synthesis of deoxyoligonucleotides. Nucleosides Nucleotides 6:287-291. Froehler, B.C., Ng, P.G., and Matteucci, M.D. 1986. Synthesis of DNA via deoxynucleoside H-phosphonate intermediates. Nucl. Acids Res. 14:5399-5407. Gaffney, B.L. and Jones, R.A. 1988. Large-scale oligonucleotide synthesis by the H-phosphonate method. Tetrahedron Lett. 29:2619-2622. Garegg, P.J., Regberg, T., Stawinski, J., and Str¨omberg, R. 1985. Formation of internucleotidic bond via phosphonate intermediates. Chem. Scr. 25:280-282. Garegg, P.J., Lindh, I., Regberg, T., Stawinski, J., Str¨omberg, R., and Henrichson, C. 1986a. Nucleoside H-phosphonates. III. Chemical
synthesis of oligodeoxyribonucleotides by the hydrogenphosphonate approach. Tetrahedron Lett. 27:4051-4054. Garegg, P.J., Lindh, I., Regberg, T., Stawinski, J., Str¨omberg, R., and Henrichson, C. 1986b. Nucleoside H-phosphonates. IV. Automated solid phase synthesis of oligoribonucleotides by the hydrogenphosphonate approach. Tetrahedron Lett. 27:4055-4058. Garegg, P.J., Regberg, T., Stawinski, J., and Str¨omberg, R. 1986c. Nucleoside hydrogenphosphonates in oligonucleotide synthesis. Chem. Scr. 26:59-62. Garegg, P.J., Henrichson, C., Lindh, I., Regberg, T., Stawinski, J., and Str¨omberg, R. 1987a. Automated solid phase synthesis of DNA and RNA fragments by the hydrogenphosphonate approach. In Biophosphates and Their Analogs— Synthesis, Structure, Metabolism and Activity (K.S. Bruzik and W.J. Stec, eds.) pp. 93-98. Elsevier/North-Holland, Amsterdam. Garegg, P.J., Regberg, T., Stawinski, J., and Str¨omberg, R. 1987b. Nucleoside Hphosphonates. VII. Studies on the oxidation of nucleoside hydrogenphosphonate esters. J. Chem. Soc. Perkin Trans. I:1269-1273. Garegg, P.J., Regberg, T., Stawinski, J., and Str¨omberg, R. 1987c. Nucleosides H-phosphonates. V. The mechanism of hydrogenphosphonate diester formation using acyl chlorides as coupling agents in oligonucleotide synthesis by the hydrogenphosphonate approach. Nucleosides Nucleotides 6:655662. Garegg, P.J., Stawinski, J., et al. 1987d. Nucleoside H-phosphonates. VI. Reaction of nucleoside hydrogenphosphonates with arenesulfonyl chlorides. J. Chem. Soc. Perkin Trans. II:12091214. Garegg, P.J., Stawinski, J., and Str¨omberg, R. 1987e. Nucleoside H-phosphonates. VIII. activation of hydrogenphosphonate monoesters by chlorophosphates and aryl sulfonyl derivatives. J. Org. Chem. 52:284-287. Gryaznov, S.M. and Potapov, V.K. 1990. New approach in synthesis of natural and modified oligodeoxyribonucleotides by H-phosphonate method. Bioorg. Khim. 16:1419-1422. Hall, R.H., Todd, A., and Webb, R.F. 1957. Nucleotides. Part XLI. Mixed anhydrides as intermediates in the synthesis of dinucleoside phosphates. J. Chem. Soc. 3291-3296. Hammond, P.R. 1962. A simple preparation of alkyl ammonium phosphonates and some comments on the reaction. J. Chem. Soc. 2521-2522. Hata, T. and Sekine, M. 1974. Silyl phosphites. I. The reaction of silyl phosphites with diphenyl disulphides. Synthesis of S-phenyl nucleoside phosphorothioates. J. Am. Chem. Soc. 96:73637364. J¨ager, A., Charubala, R., and Pfleiderer, W. 1987. Synthesis and characterization of deoxy and ribo H-phosphonate dimers. Nucl. Acids Symp. Ser. 18:197-200.
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3.4.13 Current Protocols in Nucleic Acid Chemistry
Supplement 19
Kung, P.-P. and Jones, R.A. 1992. H- phosphonate DNA synthesis without amino protection. Tetrahedron Lett. 33:5869-5872. Kuyl-Yeheskiely, E., Spierenberg, M., van den Elst, H., Tromp, M., van der Marel, G.A., and van Boom, J.H. 1986. Reaction of pivaloyl chloride with internucleosidic H-phosphonate diesters. Recl. Trav. Chim. Pays-Bas 105:505-506. Matteucci, M.D. and Caruthers, M.H. 1981. Synthesis of deoxyoligonucleotides on a polymer support. J. Am. Chem. Soc. 103:3185-3191. Nyl´en, P. 1937. Die Kinetik der Verseifung von Dialkylphosphiten. II. Allgemeine S¨aureund Basekatalyse bei der Hydrolyse von Di¨athylphosphit. Svensk Kem. Tidskr. 49:7996. Nyl´en, P. 1938. Die S¨aure- und Basekatalyse bei der Reaktion zwischen Dialkylphosphiten und Jod. Z. Anorg. Allg. Chem. 235:161-182. Ramzaeva, N., Mittelbach, C., and Seela, F. 1997. 7deazaguanine DNA: Oligonucleotides with hydrophobic or cationic side chains. Helv. Chim. Acta 80:1809-1822. Regberg, T., Stawinski, J., and Str¨omberg, R. 1988. Nucleoside H-phosphonates. IX. Possible sidereactions during hydrogenphosphonate diester formation. Nucleosides Nucleotides 7:23-35. Rozners, E., Kumpins, V., Rekis, A., and Bizdena, E. 1988. Solid phase synthesis of oligoribonucleotides by the H-phosphonate method using 2 -O-benzoyl protective group. Bioorg. Khim. 14:1580-1582. Rozners, E., Rekis, A., Kumpins, V., and Bizdena, E. 1990. Synthesis of oligoribonucleotides by the H-phosphonate method using base-labile 2 O-protecting groups. II. Some aspects of use of 2 -O-benzoyl and anisoyl protecting groups. Bioorg. Khim. 16:1531-1536. Rozners, E., Renhofa, R., Petrova, M., Popelis, J., Kumpins, V., and Bizdena, E. 1992. Synthesis of oligoribonucleotides by the H-phosphonate approach using base labile 2 -O-protecting groups. 5. Recent progress in development of the method. Nucleosides Nucleotides 11:15791593. Rozners, E., Westman, E., and Str¨omberg, R. 1994. Evaluation of 2 -hydroxyl protection in RNAsynthesis using the H-phosphonate approach. Nucl. Acids Res. 22:94-99. Rozners, E., Westman, E., Stawinski, J., Garegg, P.J., Bizdena, E., and Str¨omberg, R. 1998. Oligoribonucleotide synthesis with H-phosphonates. An improved procedure with base labile N-acyl protection and two alternative 2 -O-protecting groups. Manuscript in preparation.
Synthesis of Oligonucleotides According to the H-Phosphonate Method
Sakatsume, O., Ohtsuka, M., Takaku, H., and Reese, C.B. 1989. Solid phase synthesis of oligoribonucleotides using the 1-[2-chloro4-methyl)phenyl]-4-methoxypiperidin-4-yl (Ctmp) group for the protection of the 2 hydroxy functions and the H-phosphonate approach. Nucl. Acids Res. 17:3689-3696. Sakatsume, O., Yamane, H., Takaku, H., and Yamamoto, N. 1990. Use of new phospho-
nylating and coupling agents in the synthesis of oligodeoxyribonucleotides via the H-phosphonate approach. Nucl. Acids Res. 18:3327-3331. Seela, F. and Wei, C.F. 1997. 7-Deazaisoguanine quartets: Self-assembled oligonucleotides lacking the Hoogsteen motif. Chem. Commun. 18691870. Seliger, H. and R¨osch, R. 1990. Simultaneous synthesis of multiple oligonucleotides using nucleoside H-phosphonate intermediates. DNA Cell Biol. 9:691-696. Stawinski, J. and Str¨omberg, R. 1993. Hphosphonates in oligonucleotide synthesis. Trends Org. Chem. 4:31-67. Stawinski, J., Str¨omberg, R., Thelin, M., and Westman, E. 1988. Studies on the tbutyldimethylsilyl group as 2 -O-protection in oligoribonucleotide synthesis via the H-phosphonate approach. Nucl. Acids Res. 16:9285-9298. Stawinski, J., Str¨omberg, R., and Westman, E. 1991a. Ribonucleoside H-phosphonates. Pyridine vs quinoline–influence on condensation rate. Nucleosides Nucleotides 10:519520. Stawinski, J., Str¨omberg, R., and Westman, E. 1991b. Studies on reaction conditions for ribonucleotide synthesis via the H-phosphonate approach. Nucl. Acids Symp. Ser. 24:228. Stein, A., Iversen, P.L., Subasinghe, C., Cohen, J.S., Stec, W.J., and Zon, G. 1990. Preparation of 35 S-labelled polyphosphorothioate oligodeoxyribonucleotides by use of hydrogenphosphonate chemistry. Anal. Biochem. 188:11-16. Str¨omberg, R. 1987. Nucleoside H-phosphonates. Chemical studies directed towards oligonucleotide synthesis. Chem. Commun. Stockholm Univ. 1:1-54. Str¨omberg, R. and Stawinski, J. 1987. Evaluation of some new condensing reagents for hydrogenphosphonate diester formation. Nucl. Acids Symp. Ser. 18:185-188. Tanaka, T., Tamatsukuri, S., and Ikehara, M. 1987. Solid phase synthesis of oligoribonucleotides using the o-nitrobenzyl group for 2 -hydroxyl protection and H-phosphonate chemistry. Nucl. Acids Res. 15:7235-7248. Vasser, M., Ng, P.G., Jhurani, P., and Bischofberger, N. 1990. Error rates in oligodeoxynucleotides synthesized by the H-phosphonate method. Nucl. Acids Res. 18:3089. Wada, T., Sato, Y., Honda, F., Kawahara, S., and Sekine, M. 1997. Chemical synthesis of oligodeoxyribonucleotides using N-unprotected H-phosphonate monomers and carbonium and phosphonium condensing reagents: O-Selective phosphonylation and condensation. J. Am. Chem. Soc. 119:12710-12721. Westheimer, F.H., Huang, S., and Covitz, F. 1988. Rates and mechanisms of hydrolysis of esters of phosphorous acid. J. Am. Chem. Soc. 110:181185.
3.4.14 Supplement 19
Current Protocols in Nucleic Acid Chemistry
Westman, E. and Str¨omberg, R. 1994. Removal of tbutyldimethylsilyl protection in RNA-synthesis. Triethylamine trihydrofluoride (TEA, 3HF) is a more reliable alternative to tetrabutylammonium fluoride (TBAF). Nucl. Acids Res. 22:24302431. Westman, E., Stawinski, J., and Str¨omberg, R. 1993. RNA-synthesis using H-phosphonates. Synchronizing 2 -OH and N-protection. Collect. Czech. Chem. Commun. 58:236-237. Westman, E., Sigurdsson, S., Stawinski, J., and Str¨omberg, R. 1994. Improving the Hphosphonate approach to oligoribonucleotide synthesis. Nucl. Acids Symp. Ser. 31:25.
Wu, T. and Ogilvie, K.K. 1990. A study on the alkylsilyl groups in oligoribonucleotide synthesis. J. Org. Chem. 55:4717-4724.
Contributed by Roger Str¨omberg Karolinska Institute Stockholm, Sweden Jacek Stawinski Stockholm University Stockholm, Sweden
Wu, T. and Ogilvie, K.K. 1988. NPhenoxyacetylated guanosine and adenosine phosphoramidites in the solid phase synthesis of oligoribonucleotides: synthesis of a ribozyme sequence. Tetrahedron Lett. 29:4249-4251.
Synthesis of Unmodified Oligonucleotides
3.4.15 Current Protocols in Nucleic Acid Chemistry
Supplement 19
Strategies for Oligoribonucleotide Synthesis According to the Phosphoramidite Method Research in the many roles of ribonucleic acids was hindered by limited means of producing such biologically relevant molecules (Gold, 1988; Francklyn and Schimmel, 1989; Cook et al., 1991; Cech, 1992). Although enzymatic methods existed, protocols that allowed one to probe structure-function relationships were limited. Only uniform postsynthetic chemical modification (Karaoglu and Thurlow, 1991) or site-directed mutagenesis (Johnson and Benkovic, 1990) was available. In the latter case, researchers were limited to use of natural bases. Fortunately, adaptation of the phosphoramidite protocol for DNA synthesis to RNA synthesis has greatly accelerated our understanding of RNA. Site-specific introduction of modified nucleotides to any position in a given RNA has now become routine. Furthermore, one is not confined to a single modification but can include many variations in each molecule. It is seemingly out of proportion that one small structural modification could cause such a dilemma; however, the presence of a single hydroxyl at the 2′-position of the ribofuranose ring has been the major reason that research in the RNA field has lagged so far behind comparable DNA studies. Progress has been made in improving methods for DNA synthesis that have enabled the production of large amounts of antisense deoxyoligonucleotides for structural and therapeutic applications. Only recently have similar gains been achieved for ribonucleotides (Sproat et al., 1995; Wincott et al., 1995; Vargeese et al., 1998). The chasm between DNA and RNA synthesis is the result of the difficulty of identifying orthogonal protecting groups for the 5′- and 2′-hydroxyls. Historically, two standard approaches were taken by scientists attempting to solve the RNA synthesis problem: (1) developing a method that is compatible with state-ofthe-art DNA synthesis and (2) designing an approach specifically suited for RNA. Although adaptation of the DNA process provides a more universal procedure in which non-RNA amidites can easily be incorporated into RNA oligomers, the advantage to the latter approach is that one can develop a process that is best for RNA synthesis, allowing better yields to be realized. In both cases, however, similar issues are faced; for example identifying protecting
UNIT 3.5
groups that are compatible with synthesis conditions and yet can be removed at the appropriate juncture. This problem refers not only to the 2′- and 5′-OH groups but also to the base- and phosphate-protecting groups. Consequently, the accompanying deprotection steps, in addition to the choice of ancillary agents, are affected. Another shared issue is the need for efficient synthesis of the monomer building blocks.
BASIC CHEMISTRY OF OLIGORIBONUCLEOTIDE SYNTHESIS Solid-phase synthesis of oligoribonucleotides follows the same pathway as DNA synthesis (UNIT 3.3). A solid support with an attached nucleoside is subjected to removal of the protecting group on the 5′-hydroxyl. The incoming amidite is coupled to the growing chain in the presence of an activator. Any unreacted 5′-hydroxyl is capped, and the phosphite triester is then oxidized to provide the desired phosphotriester linkage. The process is then repeated until an oligomer of the desired length results. The actual reagents used may vary according to the 5′- and 2′-protecting groups. Other ancillary reagents may also differ. Once the oligoribonucleotide has been synthesized, it must then be deprotected. This is typically a two-step process that entails cleavage of the oligomer from the support and deprotection of the base- and phosphate-blocking groups, followed by removal of the 2′-protecting groups. Occasionally, a different order of reactions or separate deprotection of the phosphate groups is required. In all cases, it is imperative that indiscriminate removal of protecting groups not occur; this is particularly an issue in the classic situation wherein the first step is base mediated. In this case, if the 2′-hydroxyl is revealed under these conditions, strand scission will result because of attack of the vicinal hydroxyl group on the neighboring phosphate backbone (UNIT 2.2). Two other concerns that are prevalent in RNA synthesis but play no part in DNA synthesis are the propensity for (3′→2′)-phosphodiester migration to provide undesired (2′→5′)-linkages and the susceptibility of oligoribonucleotides to degradation by ribonucleases. The latter fact has led many researchers to develop 2′-protecting
Contributed by Francine E. Wincott Current Protocols in Nucleic Acid Chemistry (2000) 3.5.1-3.5.12 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Unmodified Oligonucleotides
3.5.1
groups that can remain in place until the oligomer is required for the desired experiment. Obviously, the parameters of 2′-deprotection are dictated by the protecting groups used; they will be discussed in the appropriate sections.
SOLID SUPPORTS As in DNA synthesis, the solid-phase synthesis of RNA by the phosphoramidite approach requires that one start with a solid support that is functionalized with the appropriate nucleoside corresponding to the 3′-end of the desired oligoribonucleotide. Typically, the nucleoside is attached to the support through a succinate linkage that is cleaved under alkaline conditions at the end of the synthesis (UNITS 3.1 & 3.2). Controlled-pore glass (CPG; Pon et al., 1988; Damha et al., 1990) and polystyrene (McCollum and Andrus, 1991) are the most commonly used solid supports for the synthesis of RNA. There have been reports that polystyrene resins are optimal for the synthesis of RNA on small scales, <50 µmol (Sproat et al., 1995; Wincott et al., 1995); however, recent literature describes excellent syntheses of RNA on CPG on scales of ≥100 µmol (Vargeese et al., 1998). In both cases, the best results are obtained with loadings of ∼30 µmol/g.
ACTIVATION OF RIBONUCLEOSIDE PHOSPHORAMIDITES
Strategies for Oligoribonucleotide Synthesis According to the Phosphoramidite Method
A drawback to solid-phase RNA synthesis has been the coupling step. (See Fig. 3.5.1 for the activators discussed below.) An activator reacts with the incoming amidite to produce a reactive electrophilic intermediate that is attacked by the 5′-hydroxyl of the growing polymer chain. Because of the usually bulky 2′-protecting group, coupling reactions between ribonucleoside phosphoramidites are typically sluggish. Reaction times of as much as 1 hr have been reported, although incubation times of anywhere from 10 to 30 min have usually been achieved with 1H-tetrazole (S.1) as the activator (Usman et al., 1987; Scaringe et al., 1990). This is in contrast to the extremely short coupling times required for DNA, typically on the order of 30 sec. Furthermore, coupling is usually not as efficient as observed with DNA, average stepwise yields (ASY) of 97% are common, whereas ASY of 99% are regularly achieved with DNA. Changes in concentration and/or coupling time result only in additional side products, not increased coupling yields. Recently the use of substituted 1H-tetrazoles (Leiber and Enkoju, 1961) as activators
for DNA and RNA synthesis was reported (Andrus et al., 1986; Vinayak et al., 1994). 5Ethylthio-1H-tetrazole (S.2) was found to be a more effective activator than 1H-tetrazole because of its higher solubility in acetonitrile and greater acidity (Sproat et al., 1995; Wincott et al., 1995). There are also a number of reports of the use of 5-(3-nitrophenyl)-1H-tetrazole (S.3) as an effective activator for RNA synthesis. Coupling times as short as 6 min have been reported (Rao et al., 1993). Two additional activators were recently described in the literature: 4,5-dicyanoimidazole (S.4; Vargeese et al., 1998) and benzimidazolium triflate (S.5; Hayakawa et al., 1996). These reagents rely on increased nucleophilicity to enhance the rate of the coupling reaction without increasing acidity. It should also be emphasized that choosing the appropriate protecting groups is the key to successful oligoribonucleotide synthesis. Although the 2′- and 5′-protecting groups will be discussed in depth, one cannot ignore the importance of the base- and phosphate-blocking groups. The interplay between the protecting groups is crucial. Some base- and phosphateblocking groups are not stable to the conditions required for the repetitive removal of the 5′blocking group during each nucleotide addition cycle. In other cases, the protecting group may not be stable to the conditions required to produce the monomers. Specific cases will be discussed in this unit; however, generic protection strategies are delineated below.
NUCLEOBASE-PROTECTING GROUPS Standard DNA nucleobase-protecting groups (benzoyl for A and C and isobutyryl for G) can be easily removed by treatment with
H N N N N
X
1, X = H 2, X = S-CH2CH3
NC N NC
N H 4
NO2
H N N N N 3
H N N+ H
CF3SO3
5
Figure 3.5.1 Activators commonly used for the synthesis of oligoribonucleotides.
3.5.2 Current Protocols in Nucleic Acid Chemistry
concentrated NH4OH. Many of the 2′-protecting groups used in RNA synthesis, however, are unstable to this harsh reagent. Although milder deprotection conditions have been developed—NH4OH/EtOH (3/1, v/v; Usman et al., 1987) or EtOH/NH3 (Scaringe et al., 1990)— long incubation times are still required (∼16 hr at 55°C to 65°C). Consequently, protecting groups with increased base sensitivity were developed: phenoxyacetyl (S.6; Wu et al., 1988), (4-isopropylphenoxy)acetyl (S.7), (4-tbutylphenoxy)acetyl (S.8; Chaix et al., 1989; Sinha et al., 1993), and N,N-dialkylformamidines (S.9; Theisen et al., 1993). See Figure 3.5.2 for the structures. These nucleobase-protecting groups can be removed by treatment with NH4OH/EtOH (3/1, v/v) within 4 hr at 55°C. Because of this short deprotection protocol, better yields of higher-quality RNA product were obtained. An even faster deprotection method for synthetic RNA oligonucleotides was recently reported (Reddy et al., 1995; Wincott et al., 1995). The method entails the use of aqueous methylamine at 65°C, which reduces oligonucleotide deprotection time to 10 min and produces full-length product in yields higher than those obtained with the standard NH4OH/EtOH (3/1, v/v) deprotection protocol (Wincott et al., 1995). With this reagent, however, the cytosines of any given oligonucleotide must by N-protected with an acetyl group to prevent transamination during deprotection (Reddy et al., 1994). Another family of nucleobase-protecting groups for RNA synthesis relates to the 2-(4nitrophenyl)ethyl (Npe) S.10 and 2-(4-nitrophenyl)ethoxycarbonyl (Npeoc) S.11 groups (Himmelsbach et al., 1984). They are stable to both weak acids and weak bases and yet can be readily removed with a non-nucleo-
O N
O N
Another approach to optimizing the coupling step in RNA synthesis has been to modify either the dialkylamine or the protecting group component of the ribonucleoside phosphoramidite function. In most cases, the 2-cyanoethyl/N,N-diisopropylamino combination is used for the synthesis of oligoribonucleotides. The use of lower-alkyl-substituted phosphoramidites, such as diethylamino, instead of diisopropylamino has been reported to improve coupling yields in RNA synthesis when measured by dimethoxytrityl cation quantitation (Lyttle et al., 1991). These compounds, however, have not been used extensively because of their instability. In regard to the phosphate-protecting groups, the 2-cyanoethyl is most favored; however, in some instances other protecting groups must be employed. For example, different phosphate-protecting groups may be required because of incompatibility of the 2-cyanoethyl group with synthesis or deprotection conditions or for increasing coupling rates by offsetting the bulky 2′-protecting group with a smaller phosphorous moiety. In these cases, the Npe (Himmelsbach et al., 1984) or methyl (Usman et al., 1985) group may advantageously replace the 2-cyanoethyl group for phosphate protection. The selection of phosphate protecting groups along with appropriate deprotection conditions will be discussed on a case-by-case basis in conjunction with the groups being used for 5′ and 2′ protection. Now that the general parameters regarding the synthesis of oligoribonucleotides have been reviewed, specific synthetic strategies that affect the choice of 5′- and 2′-protecting groups will be discussed.
2′-HYDROXYL PROTECTION 9
N
O
NO2
O O 10
PHOSPHATE PROTECTION
NMe2
R 6, R = H 7, R = i-Pr 8, R = t-Bu
NO2
philic base such as 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU). The use of these β-eliminating blocking groups is usually indicated when an acid-labile 5′-protecting group is present.
11
Figure 3.5.2 Base-protecting groups typically used for the synthesis of oligoribonucleotides.
The most common paradigm has been to adapt DNA synthesis to the preparation of RNA oligonucleotides. As a result, a 2′-hydroxylprotecting group must be identified that is compatible with DNA-protecting groups and easily be removed once the oligomer is synthesized (UNIT 2.2). See Figure 3.5.3 for the 2′-hydroxylprotecting groups discussed below. Owing to constraints placed by the existing amide-protecting groups on the bases and the 5′-O-dimethoxytrityl (DMTr) group—or in some
Synthesis of Unmodified Oligonucleotides
3.5.3 Current Protocols in Nucleic Acid Chemistry
DMTrO
B
O O P
R'2N
R = 12-24 R' = i-Pr/Me R" = CE/Me/Npe
OR OR"
OMe
N
OMe
O
Me Me Si Me
Cl
O
Me Me
Me 15
14
13
12
F
OMe CO2Me O
N F
16
O
Cl
Me
O
O O
Me O
MeO2C
17
18
19 NO2
OMe X1
O2 N
X2 O
CCl3
O
NO2
S O OMe
20
21, X1 = NO2, X2 = H 22, X1 = H, X2 = NO2
23
24
Figure 3.5.3 2′-Protecting groups that are compatible with solid-phase oligonucleotide synthesis protocols.
cases the 5-O-(9-(phenyl)xanthen-9-yl (Px) group—the 2′-blocking group must be stable to both acid and base. In addition, the group must also be inert to the oxidizing and capping reagents. Although the most widely used 2′-hydroxyl-protecting group is the tert-butyldimethylsilyl (TBDMS) group, many others have been explored because of the longer coupling times required when the bulky 2′-O-TBDMS substituent is used. Other types of 2′ protection will be covered in the following sections (a number of these can also be found in Gait et al., 1991, and Beaucage and Iyer, 1992).
The TBDMS Group Strategies for Oligoribonucleotide Synthesis According to the Phosphoramidite Method
By far the most popular 2′-protecting group is the TBDMS (S.12), developed principally by Ogilvie and co-workers (Usman et al., 1987). Synthesis of 2′-O-TBDMS nucleoside derivatives can be quite readily accomplished in good yields (Scaringe et al., 1990). The
chemistry used in the construction of oligoribonucleotides is completely compatible with the DNA synthesis cycle, thereby allowing for the simple preparation of DNA/RNA chimeras. In addition, many of the earlier disadvantages of the TBDMS group no longer exist. Although amidite coupling times are not as short as with DNA, through the use of 5-ethylthio-1H-tetrazole (S.2), coupling times have been reduced from 30 min to as low as 5 min (Wincott et al., 1995). Furthermore, new deprotection protocols have not only reduced incubation times but also greatly increased the quality of the product to the extent that contaminating oligonucleotides with (2′→5′) phosphodiester linkages can be eliminated completely. Deprotection of oligoribonucleotides containing 2′-O-TBDMS groups was once a twostep process that first entailed a basic step similar to that used for the deprotection of DNA oligonucleotides, in which an oligomer was
3.5.4 Current Protocols in Nucleic Acid Chemistry
cleaved from the support and the base and phosphate groups were removed. As stated earlier, through the use of methylamine this step has been reduced to 10 min. The second step was the removal of the 2′-O-TBDMS groups from the oligonucleotide. In the past, this was accomplished by treatment with 1 M ntetrabutyl ammonium fluoride (TBAF) in tetrahydrofuran (THF) at room temperature over 24 hr (Usman et al., 1987; Scaringe et al., 1990). Unfortunately, the use of this deprotecting agent produces salts that must be removed before analysis and purification. In addition, the long exposure time required for complete removal of the protecting group, coupled with the reagent’s sensitivity to adventitious water (Hogrefe et al., 1994), made it less than ideal. Although a few reports were published regarding the use of neat triethylamine trihydrofluoride (TEA•3HF) as a desilylating reagent (Gasparutto et al., 1992; Westman and Strömberg, 1994), results were mixed. A solution of TEA•3HF in N-methyl pyrrolidinone (NMP; Wincott et al., 1995) or N,N-dimethylformamide (DMF; Sproat et al., 1995) has also been described from which full deprotection can be achieved in 30 to 90 min at 65°C or 4 to 8 hr at room temperature. As an added advantage, because no salts are produced, the product can be directly precipitated from the desilylating reagent. More recently, a further improved procedure was reported in which both the basic deprotection and desilylation reaction can be accomplished in one pot using a mixture of methylamine in ethanol followed by the addition of TEA•3HF (Bellon, 1999). This protocol allows for the complete deprotection of an oligoribonucleotide in <2 hr without any evidence of (3′→2′)-phosphodiester migration.
Acetal-Protecting Groups Because of concerns about conversion of the desired (3′→5′)-internucleotidic linkages to (2′→5′) linkages, acid-labile acetals were thought to be the ideal 2′-protecting groups. They are stable to alkaline conditions and can be hydrolyzed with dilute acids; therefore, there are no residual reagents to complicate purification. Furthermore, the oligonucleotide can be isolated with the 2′-protecting group intact, thereby allowing one to store the oligonucleotide in a nuclease-resistant form. A number of different acetals have been investigated. The 2′-O-tetrahydropyranyl (Thp) S.13 and 2′-O-methoxytetrahydropyranyl (Mthp) S.14 groups proved to be
unstable to the conditions required for iterative removal of the 5′-O-DMTr group (Reese and Skone, 1985; Christodoulou et al., 1986). Although some successful syntheses have resulted from the use of these acetals, they have been limited to very short oligomers. As a result, aryl-substituted piperidines were developed. The 2′-O-[1-(2-chloro-4-methylphenyl)4-methoxypiperidin-4-yl] (Ctmp) group (S.15) and the 2′-O-[1-(2-fluorophenyl)-4-methoxypiperidin-4-yl] (Ftmp) group (S.16) were first investigated by Reese and co-workers (1986; UNIT 2.2). The Fpmp group is relatively easy to prepare and readily incorporated into ribonucleosides to provide the required phosphoramidite monomers. The 2′-O-Fpmp group is more stable to acidic hydrolysis than 2′-OThp and 2′-O-Mthp acetals (Beijer et al., 1990) and can, therefore, be used in conjunction with either a 5′-O-DMTr or a 5′-O-Px group in solid-phase RNA synthesis. Coupling times of 3 to 12 min have been reported for ribonucleoside 2′-O-Fpmp phosphoramidite derivatives upon activation with nitrophenyl-substituted 1H-tetrazoles (Beijer et al., 1990; Rao et al., 1993; Capaldi and Reese, 1994). Early reports indicated that deprotections of 2′-OFpmp oligonucleoties was optimal at pH 2.0 for 20 hr at room temperature (Rao et al., 1993). It has since, however, been determined that the rate of acid-catalyzed hydrolysis of internucleotidic linkages is sequence dependent. To avoid hydrolytic cleavage and phosphodiester migration, the removal of the 2′-O-Fpmp groups should be performed at a pH above 3.0 for 24 hr at room temperature (Capaldi and Reese, 1994; UNIT 2.2). Another approach to acetal protection of the 2′-hydroxy function led to the development of the 1-(2-chloroethoxy)ethyl group (S.17; Sakatsume et al., 1991a,b). Oligoribonucleotides of up to 20 residues in length have been prepared using S.17 for 2′-OH protection. This protecting group is stable under the acidic conditions required for removing the 5′-ODMTr group and yet can be removed postsynthetically within 30 hr upon hydrolysis with 0.01 N HCl (pH 2.0) at room temperature. No base modification or phosphodiester migration was detected. In light of the results of Capaldi and Reese (1994), deprotection at pH 3.0 might be worth investigating. Furthermore, synthesis of the 2′-O-protected ribonucleosides proceeds quite smoothly from the corresponding Markiewicz-protected nucleosides. More recently, a new 2′-protecting group was reported: the 2-hydroxyisophthalate for-
Synthesis of Unmodified Oligonucleotides
3.5.5 Current Protocols in Nucleic Acid Chemistry
maldehyde acetal (S.18; Rastogi and Usher, 1995). This is a convertible protecting group that, as the diester, is stable to acidic treatment during synthesis but is converted upon treatment with ammonia to a diacid that is more labile under acidic conditions than the parent diester. The half-life for the deprotection of the resulting diacid is ∼390 min at pH 3 compared to 166 min for the cleavage of S.16 under the same conditions. At this time only UpU and UpG dimers and the corresponding uridine phosphoramidite have been synthesized. Finally, Pfleiderer et al. (1996) designed a new acetal for the solid-phase synthesis of oligoribonucleotides that is used successfully in conjunction with a 5′-O-DMTr group. Like the 2′-protecting group S.18, the 2′-O-acetal, 1-{4[2-(4-nitrophenyl)ethoxycarbonyloxy]-3fluorobenzyloxy}ethyl (S.19), is a convertible, or “protected protecting,” group. Cleavage of the (4-nitrophenyl)ethoxycarbonyl group from S.19 results in a 2′-O-acetal that can be hydrolyzed within 4 hr under acidic conditions; this is in sharp contrast to the 24 hr required for hydrolysis of the parent 2′-O-acetal S.19 under the same conditions. Ribonucleoside phosphoramidites functionalized with the 2′-Oacetal S.19 and Npe/Npeoc (S.10/S.11) baseprotecting groups require a 20 min coupling time for optimal solid-phase RNA synthesis. Oligoribonucleotides were treated with DBU to remove base- and phosphate-protecting groups as well as the (4-nitrophenyl)ethoxycarbonyl group of the 2′-O-acetal S.19. After cleavage from the support, the deprotected 2′O-acetal was cleaved from RNA oligomers upon acidification with 0.5% AcOH for 18 hr at room temperature. The use of 80% AcOH (pH <3) led to (3′→2′)-phosphodiester migration along with strand scission (Capaldi and Reese, 1994).
Strategies for Oligoribonucleotide Synthesis According to the Phosphoramidite Method
amidite was accomplished in 15 min with tetrazole as the activator or 2.5 min using 5-(p-nitrophenyl)1H-tetrazole in conjunction with the methyl phosphate–protecting group (Tanaka et al., 1986). After oligonucleotide deprotection under basic conditions, the remaining 2′-O-(o-nitrobenzyl) groups were cleaved upon irradiation of the oligomer with longwave UV light at pH 3.5 for 1 hr in solutions that have been purged with N2. At higher pH, the formation of side products occurred. Because the 2′-O-(o-nitrobenzyl) group (in conjunction with the 2-cyanoethyl phosphate group) requires extended amidite coupling times (deBear et al., 1987), the o-nitrobenzyloxymethyl (S.21) group was proposed as an alternative 2′-photolabile-protecting group (Schwartz et al., 1992). It was postulated that the extended arm present in this group might ease steric crowding, thereby reducing amidite coupling times. Synthesis of the corresponding ribonucleoside phosphoramidite monomers proceeds similarly to that of the 2′-O-TBDMS amidites. Unlike silyl-protected ribonucleotides, however, these amidites required only a 2-min coupling time. After a standard basic deprotection protocol—pyridine:NH4OH (1:4, v/v), 50°C, 24 hr—the 2′protected oligomers were exposed to longwave UV light at pH 3.7 for 4.5 hr at room temperature to remove the o-nitrobenzyloxymethyl groups. More recently, the p-nitrobenzyloxymethyl (S.22) group was recommended for 2′-OH protection (Gough et al., 1996). This protecting group behaves almost identically to S.21 in regard to amidite monomers synthesis and coupling times; however, it can be removed from 2′-protected oligoribonucleotides within 24 hr upon reaction with TBAF at room temperature.
Photolabile Groups
The 1,1-Dianisyl-2,2,2-Trichloroethyl Group
Another approach to the protection of the ribonucleoside 2′-hydroxyl is the use of photolabile-protecting groups. This strategy has many advantages. The protecting groups are completely orthogonal, because they are resistant to both acid and base and, as a result, remain intact throughout synthesis and final deprotection. Furthermore, incorporation of such protecting groups into ribonucleosides is accomplished quite readily without any migration. Originally, the 2′-O-(o-nitrobenzyl) group (S.20) was the photolabile-protecting group of choice (Ohtsuka et al., 1981; Hayes et al., 1985). Coupling of the corresponding
Klosel et al. (1996) described a completely new protecting group for 2′ protection of ribonucleosides: the 2′-O-1,1-dianisyl-2,2,2trichloroethy group (S.23). This β-haloalkyl group is stable to acid and base and yet is cleavable under mild, neutral conditions via reductive fragmentation. Furthermore, there is no migration after the protecting group between the 2′- and the 3′-hydroxyls. Only synthesis of the uridine phosphoramidite and the corresponding UpT dimer has been described. Synthesis of the amidite monomer is fairly straightforward; the product is prepared in five steps from uridine. A 15-min
3.5.6 Current Protocols in Nucleic Acid Chemistry
coupling time is required for the activated amidite to form the dimer. After treatment with ammonia, the dimer was exposed to lithium cobalt(I)phthalocyanine and phenol in MeOH (O2 free) for 14 hr at room temperature to effect cleavage of the 2′-protecting group. The reaction mixture was then quenched by the addition of buffer; upon analysis it was shown that the backbone was intact and that no (2′→5′)-phosphodiester linkage was present.
when the Npeoc and 2-cyanoethyl groups are used for base and phosphate protection, respectively. Treatment of 2′-protected oligonucleotides with DBU results in the removal of all protecting groups. Unfortunately, this 2′protection strategy is not compatible with uridine that is unprotected at O4 because of concomitant anhydro nucleoside formation. As a result, protection of O4 with a 2-cyanoethyl group was explored. This group can also be removed upon exposure to DBU, however only at elevated temperatures (50°C).
The p-Nitrophenylethyl Sulfonyl Group
ALTERNATIVE TO 5′-TRITYL DERIVATIVES FOR THE 5′-OH PROTECTION OF RIBONUCLEOSIDES
The p-nitrophenylethyl sulfonyl group (S.24) has also been proposed as a 2′-protecting group for ribonucleosides (Pfister et al., 1988). The advantages of this sulfonate-derived group are acid stability and the absence of (2′→3′) migration. This protecting group works best
B
PO
R'2N
In many cases, researchers have chosen a de novo approach to the synthesis of oligoribonucleotides in which the focus is not on develop-
O O P
R' = i-Pr/Me R" = CE/Me/Npe OR
OR"
P Me
R Me
N
OMe
O
S
O
O
O
O 14
25 O
O Me O 27
26
O Me
Me Me
O 28
O
14 or 29 O
TMSO OTMS Si O
O O
O O
Me O
Me 30
31
Figure 3.5.4 5′(P)- and 2′(R)-protecting group combinations for the synthesis of oligoribonucleotides.
Synthesis of Unmodified Oligonucleotides
3.5.7 Current Protocols in Nucleic Acid Chemistry
ing a method that is compatible with DNA synthesis but rather evolving a process that is best for RNA synthesis. A number of different approaches have been explored. In most cases, the decision was made to proceed with an acetal-protecting group for 2′-protection. This choice was made for all the reasons stated earlier: there is no (2′→3′) migration during amidite monomer synthesis; no residual reagents are present after deprotection; and oligomers can be stored with the 2′-protecting groups in place until the RNA product is needed, thereby protecting it against nuclease degradation. Because these groups are acid labile, the 5′-protecting groups that have been developed are typically base labile or sensitive to fluoride ions. Obviously, these constraints further affect the choice of suitable nucleobases and phosphate-protecting groups. Some of the options that have been explored are presented in this unit (see Fig. 3.5.4).
The 2-Dansylethoxycarbonyl Group
Strategies for Oligoribonucleotide Synthesis According to the Phosphoramidite Method
One approach to resolving the incompatibility of 2′-O-acetal-protecting groups with the standard acid-labile 5′-O-DMTr and 5′-O-Px groups in oligoribonucleotide synthesis was the development of the 2-dansylethoxycarbonyl group for 5′ protection (S.25; Bergmann and Pfleiderer, 1994a). This base-labile group can be readily removed (in 140 sec) using dilute DBU (Bergmann and Pfleiderer, 1994b). As a result, the more stable 2-(4-nitrophenyl)ethyl phosphate-protecting group is required in place of the traditional 2-cyanoethyl group; Npe (S.10) and Npeoc (S.11) are used for base protection. When this protecting group scheme was used in conjunction with a 2′-O-Mthp group, the coupling reaction time of the corresponding phosphoramidite was 12 min with 1H-tetrazole. Oligomers of 20 nucleotides were synthesized; and in special cases, polymers of 40 nucleotides were prepared. It was determined that the best results were obtained with the N,N-diethylphosphoramidite rather than with the N,N-diisopropyl analog. Deprotection of the oligoribonucleotide first required a 10-hr treatment with 0.5 to 1 M DBU to remove the nucleobase- and phosphate-protecting groups, followed by cleavage from the support by ammonolysis (200 min). The oligomer could be stored at this point or exposed to acid to remove the 2′-O-Mthp group (S.14).
The Levulinyl Group Oligoribonucleotides (21-mers) were synthesized using a 5′-O-levulinyl (S.26)/2′-O-
tetrahydrofuranyl (S.27) protecting group combination (Iwai et al., 1987; Iwai and Ohtsuka, 1988). The 5′-O-levulinyl group is removed during solid-phase synthesis by hydrozinolysis. After ammonia treatment, the base- and phosphate-deprotected oligomer is then treated with 0.01 N HCl (pH 2) for 24 hr to effect removal of the 2′-O-acetal. Although, no base modification is observed, there are some drawbacks to this scheme. As in the case of a number of 5′-protecting groups, removal of the levulinyl group cannot be monitored. Furthermore, because of the prolonged amount of time required for full removal of the 5′-O-levulinyl group, cycle times are very long. Finally, introduction of the levulinyl group to the 5′-position of ribonucleosides is not selective, thereby reducing yields.
The 9-Fluorenylmethoxycarbonyl Group The lability of acetal groups to iterative acidic treatment led to the development of the 9-fluorenylmethoxycarbonyl (Fmoc) group (S.28) as an alternative to the DMTr group for 5′-hydroxyl protection (Lehmann et al., 1989). The Fmoc group is readily introduced to the 5′-position of the 2′-O-Mthp-protected nucleosides. During solid-phase oligonucleotide synthesis, the 5′-O-Fmoc group is removed before each chain elongation step by a brief treatment (2 min) with 0.1 M DBU in acetonitrile. The release of the Fmoc group can be monitored by UV spectroscopy, thereby allowing quantitation of the ribonucleoside phosphoramiditecoupling efficiency. The coupling reaction time of incoming amidites activated by 5-(pnitrophenyl)-1H-tetrazole ranges between 2 and 10 min. Upon completion of the synthesis, the oligoribonucleotide is treated with ammonia to remove nucleobase- and phoshate-protecting groups and release it from the support. At this point, the 2′-protected oligoribonucleotide can be purified, if so desired. Deprotection of the 2′-O-Mthp groups takes place at pH 2.0 within 4 hr at room temperature. After careful analysis, it was shown that all internucleotidic linkages were (3′→5′) and that no base modification occurred. Oligomers of up to 20 residues have been successfully synthesized using this combination of protecting groups. Ogawa et al. (1991) substituted the acidlabile 1-(isopropoxy) ethyl (IPE) group (S.29) for the 2′-O-Mthp group. The desired nucleosides were prepared from the corresponding Markiewicz-protected intermediates in a fourstep procedure in good yields. Removal of the
3.5.8 Current Protocols in Nucleic Acid Chemistry
5′-O-Fmoc group during solid-phase synthesis was accomplished with 0.1 M piperidine in acetonitrile (2 min), whereas amidite coupling was effected with 1H-tetrazole over a period of 20 to 25 min. Oligonucleotide deprotection consisted of a treatment with ammonia for 6 to 12 hr at 55°C; the 2′-O-IPE group was removed last at pH 2.0 within 3 hr at room temperature. Again, no (2′→5′)-phosphodiester isomerization or base modification was observed under these conditions. Oligomers up to 21 residues were reported using this combination of protecting groups.
oligomers of up to 36 residues in length have been reported. Careful analysis of the deprotected oligomers showed there was no base modification and no sign of (2′→5′)-phosphodiester migration. Furthermore, appropriate molecular weights and enzymatic activity were observed for the oligomers that were synthesized. It should be noted that when oligoribonucleotides are identically produced by either 5′-O-DMTr/2′-O-TBDMS or the 5′-OSIL/2′-O-ACE phosphoramidite method, better yields of RNA oligonucleotides were obtained with the 5′-O-SIL/2′-O-ACE phosphoramidite protocol.
The Bis(trimethylsiloxy) cyclooctyloxysilyl Group
SUMMARY
A completely different approach to 5′-protection was taken by Scaringe et al. (1998), wherein a 5′-O-silyl ether was used in tandem with a 2′-O-orthoester. The 5′-O-bis(trimethylsiloxy)cyclooctyloxysilyl ether (SIL; S.30) can be removed by fluoride ion treatment under conditions that will not affect an acid-labile 2′-protecting group. The 2′-O-bis(2-acetoxyethoxy) methyl orthoester (ACE; S.31) is a convertible protecting group that is stable to all synthesis conditions but is modified during nucleobase deprotection under basic conditions. The resulting 2′-O-bis(2-hydroxyethoxy)methyl orthoester is 10 times more acid labile than the original orthoesterprotecting group. The 2′-O-protected ribonucleosides can be produced in four steps from the Markiewicz-protected nucleosides in overall yields of 45% to 55%. Because 2-cyanoethyl groups are not compatible with repeated exposure to fluoride ion, methyl N,N-diisopropylphosphoramidite derivatives are used. The 5′O-silyl group is removed in 35 sec upon reaction with 1.1 M HF in TEA/DMF. Amidite coupling is complete after 90 sec when 5ethylthio-1H-tetrazole (S.2) is used as the activator; coupling yields were reported as >99%. Once the oligomer has been synthesized, deprotection of the methyl phosphate group is effected by disodium-2-carbamoyl-2-cyanoethylene-1,1-dithiolate (10 min;- Dahl et al., 1990); followed by treatment with aqueous 40% methylamine at 55°C for 10 min. The 2′-protected oligomer can then be analyzed; purified, if necessary; and then stored. To remove the modified 2′-O-orthoester, the oligoribonucleotide is heated to 55°C for 10 min in a pH 3 buffer, followed by incubation at pH 7.7 to 8.0 for 10 min at 55°C. This final step cleaves any remaining 2′-O-formyl groups that result from the orthoester deprotection. Syntheses of
Significant advances in RNA biology and biochemistry can be achieved only through concomitant advances in RNA chemistry. The current state of the art in ribozyme research would not have been possible without the recent improvements in RNA synthesis. The current technology, however, is still limiting. There is no report of routine syntheses of tRNAs or even hairpin ribozymes. Until RNA synthesis chemistry can provide oligoribonucleotides as readily as DNA, the search for new and better methods for the synthesis of RNA will continue. Currently, the 5′-O-DMTr/2′-O-TBDMS is the benchmark for the synthesis of oligoribonucleotides (Usman et al., 1987; Scaringe et al., 1990; Sproat et al., 1995; Wincott et al., 1995). The use of the TBDMS-protecting group (S.12) was first described in the 1970s. In the ensuing years, many other methods for the synthesis of RNA were developed, but none has gained the popularity of the TBDMS chemistry. Recent advances in the use of this silyl chemistry in terms of synthesis (Sproat et al., 1995; Wincott et al., 1995; Vargeese et al., 1998) and deprotection (Sproat et al., 1995; Wincott et al., 1995; Bellon, 1999) have made it an even more viable approach to the production of oligoribonucleotides. In the early 1990s the 5′-O-DMTr/2′O-Fpmp strategy to RNA synthesis showed great promise (Rao et al., 1993; Capaldi and Reese, 1994). Since that time, however, there have been very few reports of successful RNA syntheses using this protocol, although these monomers are commercially available. The results obtained with the 2′-O-(o-nitrobenzyloxymethyl) (S.21; Schwartz et al., 1992) and 2′-O-(p-nitrobenzyloxymethyl) (S.22; Gough et al., 1996) groups also appeared quite encouraging. Again, since the initial reports describing this chemistry, there have been few follow-ups,
Synthesis of Unmodified Oligonucleotides
3.5.9 Current Protocols in Nucleic Acid Chemistry
and the use of these 2′-protecting groups does not appear to have gained an appreciable audience beyond its initial developers. Other 2′-protecting groups that, like TBDMS and Fpmp, are compatible with current DNA synthesis protocols are the convertible protecting groups S.18 (Rastogi and Usher, 1995) and S.19 (Pfleiderer et al., 1996). These 2′-O-acetal-derived groups look interesting, but there have been few reports since the initial publications. Of the synthetic methods that have been designed specifically for RNA synthesis, none is currently commercially available. Around 1990, there were reports citing the combination of 5′-O-Fmoc and either 2′-O-Mthp (Lehmann et al., 1989) or 2′-O-IPE (Ogawa et al., 1991) that provided good-quality oligoribonucleotides; however, the longest oligomer synthesized was a 21-mer. No further communication regarding oligoribonucleotide synthesis with these protecting groups have surfaced. The 5′-O-SIL/2′-O-ACE protocol, however, looks very attractive (Scaringe et al., 1998). The quality of the product is excellent, and oligomers of up to 36 residues have been synthesized. Currently, none of these amidites is commercially available, although efforts are under way to commercialize the 5′-O-SIL/2′-O-ACE method. It seems clear that TBDMS chemistry is the current choice for the synthesis of oligoribonucleotides. The amidites are commercially available, and quality products can be produced on a reasonable scale. RNA synthesis chemistry using the 2′-O-TBDMS group, however, has not yet reached the level achieved by DNA synthesis. As a result, the search for improved protocols or new approaches altogether persists.
LITERATURE CITED Andrus, A., Beaucage, S., Ohms, J., and Wert, K. 1986. American Chemical Society Meeting, New York, Organic Division, Abstract 333. Beaucage, S.L. and Iyer, R.P. 1992. Advances in the synthesis of oligonucleotides by the phosphoramidite approach. Tetrahedron 48:22232311. Beijer, B., Sulston, I., Sproat, B.S., Rider, P., Lamond, A.I., and Neuner, P. 1990. Synthesis and applications of oligoribonucleotides with selected 2′-O-methylation using the 2′-O-[1-(2fluorophenyl)-4-methoxypiperidin-4-yl] protecting group. Nucl. Acids Res. 18:5143-5151. Strategies for Oligoribonucleotide Synthesis According to the Phosphoramidite Method
Bellon, L. 2000. Oligoribonucleotides with 2′-O-(tbutyldimethylsilyl) groups. In Current Protocols in Nucleic Acid Chemistry (S.L. Beaucage, D.E. Bergstrom, G.D. Glick and R.A. Jones, eds.) in press. John Wiley & Sons, New York.
Bergmann, F. and Pfleiderer, W. 1994a. Solid-phase synthesis of oligoribonucleotides using the 2dansylethoxycarbonyl group for 5′-hydroxy protection. Helvetica Chim. Acta 77:481-500. Bergmann, F. and Pfleiderer, W. 1994b. The 2-dansylethoxycarbonyl group for the protection of the 5′-hydroxy function in oligoribonucleotide synthesis. Helvetica Chim. Acta 77:989-998. Capaldi, D.C. and Reese, C.B. 1994. Use of the 1-(2-fluorophenyl)-4-methoxypiperidin-4-yl (Fpmp) and related protecting groups in oligoribonucleotide synthesis: Stability of internucleotide linkages to aqueous acid. Nucl. Acids Res. 22:2209-2216. Cech, T. 1992. Ribozyme engineering. Curr. Opin. Struct. Biol. 2:605-609. Chaix, C., Duplaa, A.M., Molko, D., and Téoule, R. 1989. Solid phase synthesis of the 5′-half of the initiator t-RNA from B. subtilis. Nucl. Acids Res. 17:7381-7393. Christodoulou, C., Agrawal, S., and Gait, M.J. 1986. Incompatibility of acid-labile 2′ and 5′ protecting groups for solid-phase synthesis of oligoribonucleotides. Tetrahedron Lett. 27:1521-1522. Cook, K.S., Fisk, G.J., Hauber, J., Usman, N., Daly, T.J., and Rusche, J.R. 1991. Characterization of HIV-1 REV protein: Binding stoichiometry and minimal RNA substrate. Nucl. Acids Res. 19:1577-1583. Dahl, B.J., Bjergarde, K., Henriksen, L., and Dahl, O. 1990. Deoxyribonucleoside phosphorodithioates. Preparation of dinucleoside phosphorodithioates from nucleoside thiophosphoramidites. Acta Chem. Scand. 44:639-641. Damha, M.J., Giannaris, P.A., and Zabarylo, S.V. 1990. An improved procedure for derivitization of controlled-pore glass beads for solid-phase oligonucleotide synthesis. Nucl. Acids Res. 18:3813-3820. deBear, J.S., Hayes, J.A., Koleck, M.P., and Gough, G.R. 1987. A universal glass support for oligonucleotide synthesis. Nucleosides Nucleotides 6:821-830. Francklyn, C. and Schimmel, P.R. 1989. Aminoacylation of RNA minihelixes with alanine. Nature 337:478-481. Gait, M.J., Pritchard, C., and Slim, G. 1991. Oligoribonucleotide synthesis. In Oligonucleotides and Analogues, A Practical Approach (F. Eckstein, ed.) pp 25-48. Oxford University Press, Oxford. Gasparutto, D., Livache, T., Bazin, H., Duplaa, A.M., Guy, A., Khorlin, A., Molko, D., Roget, A., and Téoule, R. 1992. Chemical synthesis of a biologically active natural tRNA with its minor bases. Nucl. Acids Res. 20:5159-5166. Gold, L. 1988. Posttranscriptional regulatory mechanisms in Escherichia coli. Annu. Rev. Biochem. 57:199-233. Gough, G.R., Miller, T.J., and Mantick, N.A. 1996. p-Nitrobenzyloxymethyl: A new fluoride-removable protecting group for ribonucleoside 2′hydroxyls. Tetrahedron Lett. 37:981-982.
3.5.10 Current Protocols in Nucleic Acid Chemistry
Hayakawa, Y., Kataoka, M., and Noyori, R. 1996. Benzimidazolium triflate as an efficient promoter for nucleotide synthesis via the phosphoramidite approach. J. Org. Chem. 61:79967997.
Ogawa, T., Hosaka, H., Makita, T., and Takaku, H. 1991. Solid-phase synthesis of oligoribonucleotides using 5′-9-fluorenylmethoxycarbonyl and 2′-1-(isopropoxyl)ethyl protection. Chem. Lett. 1169-1172.
Hayes, J.A., Brunden, M.J., Gilham, P.T. and Gough, G.R. 1985. High-yield synthesis of oligoribonucleotides using o-nitrobenzyl protection of 2′-hydroxyls. Tetrahedron Lett. 26:24072410.
Ohtsuka, E., Fujiyama, K., and Ikehara, M. 1981. Studies on transfer ribonucleic acids and related compounds. XL. Synthesis of an eicosaribonucleotide corresponding to residues 35-54 of tRNAfMet from E. coli. Nucl. Acids Res. 9:35033522.
Himmelsbach, F., Schulz, B.S., Trichtinger, T., Charubala, R., and Pfleiderer, W. 1984. The pnitrophenylethyl (Npe) group. Tetrahedron 40:59-72. Hogrefe, R.I., McCaffrey, A.P., Borozdina, L.U., McCampbell, E.S., and Vaghefi, M.M. 1994. Effect of excess water on the desilylation of oligoribonucleotides using tetrabutylammonium fluoride. Nucl. Acids Res. 21:4739-4741. Iwai, S. and Ohtsuka, E. 1988. 5′-Levulinyl and 2′-tetrahydrofuranyl protection for the synthesis of oligoribonucleotides by the phosphoramidite approach. Nucl. Acids Res. 16:9443-9456. Iwai, S., Yamada, E., Asaka, M., Hayase,Y., Inoue, H., and Ohtsuka, E. 1987. A new solid-phase synthesis of oligoribonucleotides by the phosphoro-p-anisidate method using tetrahydrofuranyl protection of 2′-hydroxyl groups. Nucl. Acids Res. 15:3761-3772. Johnson, K.A. and Benkovic, S.J. 1990. Analysis of protein function by mutagenesis. In The Enzymes, Vol. 19 (Sigman, D.S. and Boyer, P.D., eds) pp. 159-211. Academic Press, San Diego. Karaoglu, D. and Thurlow, D.L. 1991 A chemical interference study on the interaction of ribosomal protein L11 from in Escherichia coli with RNA molecules containing its binding site from 23S rRNA. Nucl. Acids Res. 19:5293-5300. Klosel, R., Konig, S., Lehnhoff, S., and Karl, R.M. 1996. The 1,1-dianisyl-2,2,2-trichloroethyl group as a 2′-hydroxyl protection of ribonucleotides. Tetrahedron 52:1493-1502. Lehmann, C., Xu, Y.Z., Christodoulou, C., Tan, Z.K., and Gait, M.J. 1989. Solid-phase synthesis of oligoribonucleotides using 9-fluorenylmethoxycarbonyl (Fmoc) for 5′-hydroxyl protection. Nucl. Acids Res. 17:2379-2390. Leiber, E. and Enkoju, T. 1961. Synthesis and properties of 5-(substituted) mercaptotetrazoles. J. Org. Chem. 26:4472-4479. Lyttle, M.H., Wright, P.B., Sinha, N.D., Bain, J.D., and Chamberlin, A.R. 1991. New nucleoside phosphoramidites and coupling protocols for solid-phase RNA synthesis. J. Org. Chem. 56:4608-4615. McCollum, C. and Andrus, A. 1991. An optimized polystyrene support for rapid, efficient oligonucleotide synthesis. Tetrahedron Lett. 32:40694072.
Pfister, M., Farkas, S., Charubala, R., and Pfleiderer, W. 1988. Recent progress in oligoribonucleotide synthesis. Nucleosides Nucleotides 7:595-600. Pfleiderer, W., Matysiak, S., Bergmann, F., and Schnell, R. 1996. Recent progress in oligonucleotide synthesis. Acta Biochim. Polonica 43:37-44. Pon, R.T., Usman, N., and Ogilvie, K.K. 1988. Derivitization of controlled pore glass beads for solid phase oligonucleotide synthesis. BioTechniques 6:768-774. Rao, M.V., Reese, C.B., Schehlmann, V., and Yu, P.S. 1993. Use of the 1-(2-fluorophenyl)-4methoxypiperidin-4-yl (Fpmp) protecting group in the solid-phase synthesis of oligo- and polyribonucleotides. J. Chem. Soc. Perkin Trans. I:43-55. Rastogi, J. and Usher, D. 1995. A new 2′-hydroxyl protecting group for the automated synthesis of oligoribonucleotides. Nucleic Acids Res. 23:4872-4877. Reddy, M.P., Hanna, N.B., and Farooqui, F. 1994. Fast cleavage and deprotection of oligonucleotides. Tetrahedron Lett. 35:4311-4314. Reddy, M.P., Hanna, N.B., and Farooqui, F. 1995. Methylamine deprotection provides increased yield of oligoribonucleotides. Tetrahedron Lett. 36:8929-8932. Reese, C.B. and Skone, P.A. 1985. Action of acid on oligoribonucleotide phosphotriester intermediates. Effect of released vicinal hydroxy functions. Nucl. Acids Res. 13:3501-. Reese, C.B., Serafinowska, H.T., and Zappia, G. 1986. An acetal group suitable for the protection of 2′-hydroxy functions in rapid oligoribonucleotide synthesis. Tetrahedron Lett. 27:22912294. Sakatsume, O., Yamaguchi, T., Ishikawa, M., Hirao, I., Miura, K., and Takaku, H. 1991a. Solid phase synthesis of oligoribonucleotides by the phosphoramidite approach using 2′-O-1-(2-chloroethox y)ethyl p ro tection. Tetrahedron 47:8717-8728. Sakatsume, O., Ogawa, T., Hosaka, H., Kawashima, M., Takaki, M., and Takaku, H. 1991b. Synthesis and properties of non-hammerhead RNA using 1-(2-chloroethoxy)ethyl group for the protection of 2′-hydroxyl function. Nucleosides Nucleotides 10:141-153. Synthesis of Unmodified Oligonucleotides
3.5.11 Current Protocols in Nucleic Acid Chemistry
Scaringe, S.A., Francklyn, C., and Usman, N. 1990. Chemical synthesis of biologically active oligoribonucleotides using β-cyanoethyl protected ribonucleoside phosphoramidites. Nucl. Acids Res. 18:5433-5341. Scaringe, S.A., Wincott, F.E., and Caruthers, M.H. 1998. Novel RNA synthesis method using 5′-Osilyl-2′-O-orthoester protecting groups. J. Am. Chem Soc. 120:11820-11821. Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., deBear, J.S., and Gough, G.R. 1992. Rapid synthesis of oligoribonucleotides using 2′-O-(o-nitrobenzyloxymethyl)-protected monomers. Bioorg. Med. Chem. Lett. 2:1019-1024. Sinha, N.D., Davis, P., Usman, N., Pérez, J., Hodge, R., Kremsky, J., and Casale, R. 1993. Labile exocyclic amine protection in DNA, RNA and oligonucleotide analog synthesis facilitating Ndeacylation, minimizing depurination and chain degradation. Biochimie 75:13-23. Sproat, B., Colonna, F., Mullah, B., Tsou, D., Andrus, A., Hampel, A., and Vinayak, R. 1995. An efficient method for the isolation and purification of oligoribonucleotides. Nucleosides Nucleotides 14:255-273. Tanaka, T., Tamatsukuri, S., and Ikehara, M. 1986. Solid phase synthesis of oligoribonucleotides using o-nitrobenzyl protection of 2′-hydroxyl via a phosphite triester approach. Nucl. Acids Res. 14:6265-6279. Theisen, P., McCollum, C., and Andrus, A. 1993. N-6-Dialkylformamidine-2′-deoxyadenosine phosphoramidites in oligodeoxynucleotide synthesis. Rapid deprotection of oligodeoxynucleotides. Nucleosides Nucleotides 12:10331046. Usman, N., Pon, R.T., and Ogilvie, K.K. 1985. Preparation of ribonucleoside 3′-O-phosphoramidites and their application to the automated solid phase synthesis of oligonucleotides. Tetrahedron Lett. 26:4567-4570.
Usman, N., Ogilvie, K.K., Jiang, M.-Y., and Cedergren, R.J. 1987. Automated chemical synthesis of long oligoribonucleotides using 2′-O-silylated ribonucleoside 3′-O-phosphoramidites on a controlled-pore glass support: Synthesis of a 43-nucleotide sequence similar to the 3′-half molecule of an in Escherichia coli formylmethionine tRNA. J. Am. Chem. Soc. 109:78457854. Vargeese, C., Carter, J., Yegge, J., Krivjansky, S., Settle, A., Kropp, E., Peterson, K., and Pieken, W. 1998. Efficient activation of nucleoside phosphoramidites with 4,5-dicyanoimidazole during oligonucleotide synthesis. Nucl. Acids Res. 26:1046-1050. Vinayak, R., Ratmeyer, L., Wright, P., Andrus, A., and Wilson, D. 1994. Chemical synthesis of biologically active RNA using labile protecting groups. In Innovations and Perspectives in Solid Phase Synthesis (R. Epton, ed.) pp 45-50. Mayflower Worldwide, Birmingham. Westman, E. and Strömberg, R. 1994. Removal of t-butyldimethylsilyl protection in RNA-synthesis. Triethylamine trihydrofluoride (TEA, 3HF) is a more reliable alternative to tetrabutylammonium fluoride (TBAF). Nucl. Acids Res. 22:2430-2431. Wincott, F., DiRenzo, A., Shaffer, C., Grimm, S., Tracz, D., Workman, C., Sweedler, D., Gonzalez, C., Scaringe, S., and Usman, N. 1995. Synthesis, deprotection, analysis and purification of RNA and ribozymes. Nucl. Acids Res. 23:2677-2684. Wu, T., Ogilvie, K.K., and Pon, R.T. 1988. N-Phenoxyacetylated guanosine and adenosine phosphoramidites in the solid phase synthesis of oligoribonucleotides: Synthesis of a ribozyme sequence. Tetrahedron Lett. 34:4249-4252.
Contributed by Francine E. Wincott Ribozyme Pharmaceuticals, Inc. Boulder, Colorado
Strategies for Oligoribonucleotide Synthesis According to the Phosphoramidite Method
3.5.12 Current Protocols in Nucleic Acid Chemistry
Oligoribonucleotides with 2′-O-(tert-Butyldimethylsilyl) Groups
UNIT 3.6
This unit describes current methodologies to chemically synthesize oligoribonucleotides on solid support by means of automated DNA synthesizers. It also includes an updated collection of protocols describing the deprotection of base-labile phosphate, and nucleobase protecting groups, and fluoride-labile 2′-O-(tert-butyldimethylsilyl) protecting groups of crude synthetic oligoribonucleotides. Small-scale synthesis (i.e., 0.2 µmol scale) of oligoribonucleotide provides ∼400 to 600 µg of purified material, which is usually enough for a wide range of biochemical applications. Such synthesis can be readily achieved using the exocyclic amine-protected 5′-O-dimethoxytrityl-2′-O-tert-butyldimethylsilyl-3′-O-(2-cyanoethyl-N,N-diisopropyl) -ribonucleoside phosphoramidites in combination with the acidic 5-ethylthio-1H-tetrazole (SET, pKa = 4.28) as the activator (Wincott et al., 1995; Vinayak et al., 1995; see Basic Protocol 1). Alternatively, the use of less acidic activators such as 4,5-dicyanoimidazole (DCI, pKa = 5.2; Vargeese et al., 1998) or 1H-tetrazole (TET, pKa = 4.8; Usman et al., 1987; Scaringe et al., 1990; Usman and Cedergren, 1992) also allows for efficient oligoribonucleotide synthesis at comparable scales. DCI is better suited for larger-scale (i.e., >500 µmol) syntheses due to lesser acidity (Vargeese et al., 1998) that may limit the activator-induced detritylation of incoming phosphoramidite during extended coupling time (Krotz et al., 1997). Historically, crude oligoribonucleotides have been fully deprotected with a 3:1 mixture of concentrated ammonium hydroxide/ethanol followed by an n-tetrabutylammonium fluoride (TBAF) treatment (Usman et al., 1987; Stawinsky et al., 1988; see Basic Protocol 2). The use of aqueous methylamine followed by treatment with the triethylamine trihydrofluoride complex (see Alternate Protocol 1) constitutes a significant improvement in the deprotection process (Wincott et al., 1995; Vinayak et al., 1995), alleviating premature deprotection of the 2′-hydroxyl during the basic treatment, eliminating the well-known sensitivity of the TBAF to water (Hogrefe et al., 1993), and shortening the overall deprotection time considerably. Crude oligoribonucleotides can also be efficiently deprotected in a “one-pot” reaction using anhydrous methylamine and neat triethylamine trihydrofluoride (see Alternate Protocol 2). This alternate deprotection protocol eliminates the time-consuming evaporation step, thereby reducing the overall deprotection time to 45 min, which allows for a high-throughput production mode (Bellon, 2000). AUTOMATED OLIGORIBONUCLEOTIDE SYNTHESIS This protocol describes automated chemical synthesis of oligoribonucleotides by means of the phosphoramidite method (UNIT 3.5) according to the synthetic scheme pictured in Figure 3.6.1. The procedure described below was developed for the ABI 394 DNA/RNA synthesizer at the 0.2 µmol scale although it can be modified to utilize any standard synthesizer.
BASIC PROTOCOL 1
Materials Aminomethyl polystyrene (RNA primer solid support) derivatized with 5′-O-DMTr-2′-O-TBDMS-3′-O-succinyl ribonucleosides (Amersham Pharmacia Biotech) Synthesis of Unmodified Oligonucleotides Contributed by Laurent Bellon Current Protocols in Nucleic Acid Chemistry (2000) 3.6.1-3.6.13 Copyright © 2000 by John Wiley & Sons, Inc.
3.6.1 Supplement 1
Phosphoramidite Coupling Reaction DMTrO
B DMTrO
Cycle Entry
5'-Hydroxyl Deprotection
DMTrO
B
O
HO
O
i-Pr2N
B
O
OR
O
N H
O
O P
B
O
OR O
OCE
OR
P OCE
O O
N H
O
OR
EtS
O
O
H N
O
HO
+
B
O
N N N
O
N H
O
B
O O
N H
OR
OR
O
O
To Next Cycle Capping 5'-Unreacted Hydroxyl DMTrO
B
O
DMTrO O
AcO
OR
+
O P OCE O
O
B
O N H
O O
O
PhosphiteTriester Oxidation
O N H
O O
B
O
B
OR
OR
P OCE O
O
OR
O
O N H
O
AcO
+ B
OR
O
B
O N H
O
OR
O
O
Cycle End
DMTr, 4,4′-dimethoxytrityl; R, tert-butyldimethylsilyl; CE, 2-cyanoethyl; Ac, acetyl; B, uracil-1-yl (U); N 4-acetylcytosin-1-yl (CAc); N 6-phenoxyacetyladenin-9-yl (APAC); N2-[(4-isopropyl)phenoxy]acetylguanin-9-yl (GiPrPAC)
Figure 3.6.1 Synthesis of oligoribonucleotide with 2′-O-TBDMS groups on solid-support via the phosphoramidite approach.
RNA phosphoramidites (Amersham Pharmacia Biotech; diluted on the synthesizer to 0.1 M in acetonitrile, using automated protocols) 5′-O-DMTr-N6-(phenoxyacetyl)-2′-O-TBDMS-adenosine-3′-O-(β-cyanoethylN,N-diisopropyl) phosphoramidite 5′-O-DMTr-N2-(isopropylphenoxyacetyl)-2′-O-TBDMS-guanosine-3′-O-(βcyanoethyl-N,N-diisopropyl) phosphoramidite 5′-O-DMTr-N4-(acetyl)-2′-O-TBDMS-cytidine-3′-O-(β-cyanoethyl-N,Ndiisopropyl) phosphoramidite 5′-O-DMTr-2′-O-TBDMS-uridine-3′-O-(β-cyanoethyl-N,N-diisopropyl) phosphoramidite. 3% (v/v) TCA in methylene chloride (PE Biosystems) Cap A: 10% (v/v) acetic anhydride/10% (v/v) 2,6-lutidine in THF (PE Biosystems) Cap B: 16% (v/v) 1-methyl imidazole in THF (PE Biosystems) Iodine solution: 16.9 mM I2/49 mM pyridine/9% (v/v) water in THF (PE Biosystems) Synthesis grade acetonitrile (Burdick & Jackson) Activator (prepare in acetonitrile): 0.25 M 5-ethylthio-1H-tetrazole (SET), made from solid (American International Chemical) or 0.5 M 4,5-dicyanoimidazole solution (DCI), made from solid (Proligo) or 0.45 M 1H-tetrazole (TET; Glen Research). Synthesis columns for 0.2-µmol-scale syntheses (PE Biosystems) ABI 394 DNA/RNA synthesizer (PE Biosystems)
Oligoribonucleotides with 2′-O(tert-Butyldimethylsilyl) Groups
1. Load an empty synthesis column with ∼8 mg of the RNA primer solid-support (i.e., ∼25 µmol/g) corresponding to the first nucleotide at the 3′- end of the oligoribonucleotide.
3.6.2 Supplement 1
Current Protocols in Nucleic Acid Chemistry
Table 3.6.1 Equivalents to the Synthesis Scale and Wait Time Required for Optimal Solid-Phase Synthesis
Reagentsa
Equivalents Wait time (sec)
Phosphoramidites 15 Activator: SET 39 DCI 80 TET 70 Acetic anhydride (Cap A) 655 1-Methylimidazole (Cap B) 1245 TCA 700 Iodine solution 21
465 465 465 465 5 5 10 15
aDescriptions of these reagents have been shortened; see Basic Protocol 1 materials list for complete information.
2. Perform synthesis on an ABI 394 synthesizer according to the cycle outlined in Tables 3.6.1 and 3.6.2. See materials list above as well as Tables 3.6.1 and 3.6.2 for synthesis materials.
3. At the end of the synthesis, perform a manual detritylation cycle on the synthesizer (optional) to remove the dimethoxytrityl group at the 5′-end of the oligonucleotide. 4. Remove the synthesis column from the synthesizer and dry it either under a stream of argon or in a vacuum dessicator for 10 to 15 min. 5. Deprotect the oligoribonucleotide (see Basic Protocol 2, Alternate Protocol 1, or Alternate Protocol 2). OLIGORIBONUCLEOTIDE DEPROTECTION WITH NH4OH/ETHANOL AND TBAF
BASIC PROTOCOL 2
This protocol describes a deprotection scheme using a 3:1 cocktail of concentrated ammonium hydroxide and ethanol to cleave the oligoribonucleotide from the solid support, perform the β-elimination of the cyanoethyl phosphodiester protecting group, and cleave the exocyclic N-acyl protecting groups. A subsequent treatment with ntetrabutylammonium fluoride effects cleavage of the tert-butyldimethylsilyl group protecting the 2′-hydroxyl functionality. (Fig. 3.6.2). Materials Oligoribonucleotide attached to solid support (see Basic Protocol 1) 3:1 (v/v) 29% ammonium hydroxide (Mallinckrodt Baker)/100% ethanol (prepare immediately before use) 3:1:1 (v/v/v) ethanol/acetonitrile/H2O 1.0 M n-tetrabutylammonium fluoride (TBAF) in THF (Aldrich) 50 mM and 2 M triethylammonium bicarbonate (TEAB), pH 7.8 (see recipe) Heating blocks 4-mL glass screw-top vial with Teflon lined lid (Wheaton) 14-mL centrifuge tubes (Falcon) Qiagen-tip 500 column (Qiagen) 1. Transfer the dried oligoribonucleotide on solid support from the synthesis column (see Basic Protocol 1) to a 4-mL glass screw top vial with Teflon-lined lid.
Synthesis of Unmodified Oligonucleotides
3.6.3 Current Protocols in Nucleic Acid Chemistry
Supplement 1
Table 3.6.2 Summary of a 0.2 µmol–Scale Cycle for Oligoribonucleotide Synthesis on ABI 394 DNA/RNA Synthesizera
Step
Function
Wash steps 1 Acetonitrile to waste 2 Acetonitrile to column 3 Argon reverse flush 4 Argon block flush Chain extension steps 5 Activator to waste 6 Amidite + activator to column 7 Push to column 8 Wait 9 Push to column 10 Wait 11 Repeat steps 9-10 (6 times) 12 Argon flush to waste Wash steps 13 Acetonitrile to waste 14 Repeat steps 3 and 4 Capping steps 15 Cap A and B to column 16 Wait Wash steps 17 Repeat steps 13 and 14 Oxidation steps 18 Iodine to column 19 Wait Wash steps 20 Repeat steps 13 and 14 21 Acetonitrile to column 22 Argon flush to waste 23 Acetonitrile to column 24 Repeat steps 3 and 4 25 Repeat steps 21, 23 and 24 Detritylation steps 26 TCA/DCM to column 27 Wait 28 Argon trityl flush 29 Repeat steps 26-28 Wash steps 30 Acetonitrile to column 31 Argon trityl flush 32 Repeat steps 2, 3, and 4 33 End
Time (sec) 3 10 8 4 1.7 1.2 NA 150 0.1 45 0.1 4
4 5
4 15
10 4 10
6 5 5
10 5
aDelivery flow rate are ∼3.1 mL/min for phosphoramidites and activators and ∼3.6 mL/min for all other reagents.
Oligoribonucleotides with 2′-O(tert-Butyldimethylsilyl) Groups
3.6.4 Supplement 1
Current Protocols in Nucleic Acid Chemistry
2. Add 2 mL of a 3:1 ammonium hydroxide/ethanol solution to the vial, screw cap on tightly, and place in a heat block for 4 hr at 65°C. 3. Remove the vial from the heat block, place it in a block kept at room temperature, and place the block in a –20°C freezer until cooled (i.e., ∼30 min). IMPORTANT NOTE: To avoid loss of contents, it is important to cool the sample vial in step 4 before opening the screw cap.
4. Decant the solution (containing the deprotected oligonucleotide) into a 14-mL Falcon tube. Add 1 mL of ethanol/acetonitrile/H2O solution, vortex well, and allow the support to settle. Decant wash and add to deprotected oligonucleotide solution. Repeat wash twice. Due to the presence of the hydrophobic 2′-O-TBDMS groups on the RNA, this organic wash helps increase the recovery yield.
5. Evaporate the combined supernatant from step 4 in the 14-mL tube on a Speedvac evaporator (i.e., ∼2.5 hr on medium heat). 6. Add 1 mL of 1.0 M TBAF to the 14-mL tube containing the dried RNA and allow to react at room temperature for 24 hr. 7. Quench the desilylation reaction by adding 9 mL of 50 mM TEAB, then refrigerate at 4°C until ready for desalting. Deprotected oligoribonucleotides are highly sensitive to nuclease degradation. Therefore, gloves should always be worn when manipulating deprotected synthetic RNA; sterile disposable containers, nuclease-free laboratory reagents, and Milli-Q water should always be used to limit potential exposure to nucleases.
8. Prewash the Qiagen-tip 500 cartridge with 10 mL of 50 mM TEAB. 9. Load the quenched reaction in TEAB onto the Qiagen-tip 500 anion-exchange cartridge. 10. Wash the loaded cartridge with 10 mL of 50 mM TEAB and discard the eluent. Elute the RNA with 10 mL of 2 M TEAB into a sterile tube, and dry to a white powder on a Speedvac evaporator. OLIGORIBONUCLEOTIDE DEPROTECTION WITH AQUEOUS METHYLAMINE AND TRIETHYLAMINE TRIHYDROFLUORIDE
ALTERNATE PROTOCOL 1
This protocol describes a deprotection scheme using aqueous methylamine and triethylamine trihydrofluoride as alternate reagents to effect nucleobase, 2′-hydroxyl and phosphodiester deprotection (Fig. 3.6.2.). Also see APPENDIX 3C, Basic Protocol 3, for general discussion of RNA oligonucleotide of deprotection. Additional Materials (also see Basic Protocol 2) 40% (w/v) aqueous methylamine (Aldrich) Triethylamine trihydrofluoride/NMP/TEA solution (see recipe) 3 M aqueous sodium acetate (e.g., Fluka) n-butanol 70% aqueous ethanol 1. Transfer the oligoribonucleotide attached to the solid-support from the synthesis column (see Basic Protocol 1) to a 4-mL glass screw top vial. 2. Add 1 mL of 40% aqueous methylamine to the vial, screw the cap on tightly, and place in a heat block at 65°C for 10 min.
Synthesis of Unmodified Oligonucleotides
3.6.5 Current Protocols in Nucleic Acid Chemistry
Supplement 1
3. Remove the vial from the heat block, place it in block kept at room temperature, and place the block in −20°C freezer until cooled (e.g., ~20 min). 4. Decant the solution into a 14-mL tube. Add 1 mL of 3:1:1 ethanol/acetonitrile/H2O solution, vortex well, and allow the support to settle. Decant wash solution and add to deprotection solution. Repeat wash twice, for a total of three washes. 5. Evaporate the combined supernatants in the tube on a Speedvac evaporator (i.e., ∼2.5 hr on medium heat). 6. Add 0.3 mL of triethylamine trihydrofluoride/NMP/TEA solution to the tube containing the dried RNA, cap the tube, and place on a heat block for 90 min at 65°C. After incubation, bring to room temperature. 7. Precipitate the oligoribonucleotide directly from the desilylation reaction by adding 25 µL of 3 M aqueous sodium acetate, followed by 1 mL of n-butanol. If the oligoribonucleotide has been synthesized trityl-off, proceed to quenching and desalting steps (see Basic Protocol 2, steps 7 to 10).
8. Cool the mixture to −20°C for 2 hr to overnight and centrifuge 30 min at 4000 × g (3750 rpm in a Beckman GS-GR rotor), 4°C. 9. Decant the solution and wash the pellet with 70% ethanol. Centrifuge 10 min at 4000 × g (3750 rpm in a Beckman GS-GR rotor), 4°C. Decant the supernatant and dry the oligoribonucleotide pellet by using a Speedvac evaporator. This precipitation procedure cannot be applied to the TBAF procedure (see Basic Protocol 2) because of the high organic content of the desilylation reaction. Alternately, if a trityl-on deprotected oligoribonucleotide is sought, quench the desilylation reaction by adding 5 mL of 1.5 M ammonium bicarbonate, pH 7.5 (see recipe in Reagents and Solutions). ALTERNATE PROTOCOL 2
“ONE-POT” OLIGORIBONUCLEOTIDE DEPROTECTION WITH ANHYDROUS METHYLAMINE AND NEAT TRIETHYLAMINE TRIHYDROFLUORIDE This protocol describes an expedited deprotection scheme for oligoribonucleotides using anhydrous ethanolic methylamine and triethylamine trihydrofluoride to effect nucleobase, 2′-hydroxyl, and phosphodiester deprotection (Fig. 3.6.2).
HO
B
O O
R
HO
OTBDMS
Nucleobase Deprotection
O
B
NH4OH:EtOH (3:1) or 40% aq. CH3NH2 or 33% CH3NH2 in EtOH
R
O
n
2'-O-Deprotection
−
O
H
B
O O
OH
O P O O
O
OTBDMS
O
HO
OTBDMS
O O P O
O P OCE O
B
O
1.0 M TBAF in THF or TEA•3HF/NMP/TEA or TEA•3HF
B
−
O
O O
OTBDMS n
H
B
OH n
O NH
Oligoribonucleotides with 2′-O(tert-Butyldimethylsilyl) Groups
BR = U, CAc, APAC, GiPrPAC B = U, C, A, G TBDMS, tert-butyldimethylsilyl; TBAF, tetra-n-butylammonium fluoride; NMP, 1-methyl-2-pyrrolidinone; TEA, triethylamine
Figure 3.6.2 Deprotection of chemically synthesized oligoribonucleotides with 2′-O-TBDMS groups.
3.6.6 Supplement 1
Current Protocols in Nucleic Acid Chemistry
Additional Materials (also see Basic Protocol 2) 1:1 (v/v) mixture of 33% ethanolic methylamine and anhydrous DMSO 1.5 M ammonium bicarbonate, pH 7.5 (see recipe) Acetonitrile 1:1:1 (v/v/v) acetonitrile/methanol/H2O RNase-free H2O/DEPC-treated C18 SepPak cartridges (Waters) 1. Transfer the dried solid support-bound oligoribonucleotide from the synthesis column to a 4-mL glass screw top vial. 2. Add 0.8 mL of a 1:1 (v/v) mixture of 33% methylamine/DMSO to the vial, screw the cap on tightly, and place on a heat block at 65°C for 15 min. DMSO is useful for solubilizing the partially deprotected oligoribonucleotide and helps prevent alkaline hydrolysis of the fully deprotected RNA.
3. Remove the vial from the heat block and place it in a block kept at room temperature. 4. Add 0.1 mL of neat triethylamine trihydrofluoride, vortex well, and place on a heating block for 15 min at 65°C. Cool the sample vial at room temperature and then at –20°C for 10 min. The solution usually gels after the addition of triethylamine trihydrofluoride.
5. Quench the reaction by adding 1 mL of 1.5 M ammonium bicarbonate, pH 7.5. Allow the support to settle and decant the supernatant. If sample is not cool enough, addition of ammonium bicarbonate solution may lead to significant effervescence.
6. Prewash the C18 Sep-pak cartridge successively with 10 mL of acetonitrile, 10 mL of 1:1:1 acetonitrile/methanol/H2O solution, and 20 mL of RNase free H2O. 7. Apply the quenched solution to the prewashed C18 Sep-pak cartridge and wash the loaded cartridge with 10 mL of RNase-free H 2O to remove salts. 8. Elute the product from the column by using 10 mL of 1:1:1 acetonitrile/methanol/H2O and evaporate the eluate to dryness on a Speedvac. This desalting step will detritylate a trityl-on deprotected oligoribonucleotide. The precipitation procedure of Alternate Protocol 1 (steps 7 to 9) cannot be applied to the “one-pot” deprotection protocol because of the high organic content.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Ammonium bicarbonate,1.5 M (pH 7.5) Weigh 118.59 g of solid NH4HCO3 and bring up to 1 L with Milli-Q water. Store the buffer solution up to 2 months at 4°C. Triethylamine trihydrofluoride/NMP/TEA solution, 1.5:0.75:1 (v/v/v) Combine, in the following order, 0.75 mL 1-methyl-2-pyrrolidinone (NMP), 1.0 mL triethylamine (TEA), and 1.5 mL triethylamine trihydrofluoride. If the reagent is not used immediately after preparation, store it in a capped container on a warm (55° to 65°C) heat block. The reagent will form an intractable gel if allowed to stand at room temperature.
Synthesis of Unmodified Oligonucleotides
3.6.7 Current Protocols in Nucleic Acid Chemistry
Supplement 1
Triethylammonium bicarbonate (TEAB), 50 mM and 2 M (pH 7.8) Place a 4-L bottle containing 3 L of Super-Q water in an ethanol/ice bath. Bubble carbon dioxide through the water. After 15 min add 279 mL of neat triethylamine every 2 hr for 8 hr (total 4 additions). After 8 hr of CO 2 saturation, check the pH on an aliquot. Continue bubbling CO2 until the pH reaches 7.8 then bring to 4 L with water. Store the buffer solution up to 6 months at 4°C. To prepare 50 mM TEAB, dilute 2M TEAB solution 40-fold with water. COMMENTARY Background Information
Oligoribonucleotides with 2′-O(tert-Butyldimethylsilyl) Groups
Synthesis of oligoribonucleotides Similar to that of oligodeoxyribonucleotides, the chemical synthesis of oligoribonucleotides on solid support is routinely performed via the phosphoramidite method (Fig. 3.6.1; also see UNITS 3.3 & 3.5). However, the additional 2′-hydroxyl function of the ribofuranosyl sugar requires suitable protection during oligoribonucleotide synthesis. Among the various protecting groups available, trialkylsilyl ethers (Ogilvie et al., 1976, 1977; Wincott and Usman, 1994), and particularly the tertbutyldimethylsilyl ether (TBDMS; Usman et al., 1985), have been the most extensively studied, since fluoride-mediated silyl ether deprotection is quite orthogonal to the other acid- and base-labile protecting groups commonly used in oligonucleotide chemistry. Because of the steric bulk introduced by the TBDMS group, coupling 2′-O-TBDMS protected ribophosphoramidites to a growing oligonucleotide chain is notoriously more sluggish as compared to their 2′-deoxy analogs. The elongation cycle for automated oligoribonucleotide synthesis is similar to that of oligodeoxynucleotides and consists of the detritylation, coupling, capping, and oxidation steps. Each of these steps is critical for successful RNA synthesis. One important difference regarding the detritylation step is the possibility of using higher concentrations of haloacids due to the reduced sensitivity of ribofuranosylnucleotides to acid-mediated depurination. The oxidation and capping steps do not differ significantly from traditional oligodeoxyribonucleotide synthesis. Conversely, the coupling step has been the focus of much attention through the development of a number of phosphate protecting groups (Usman et al., 1987; Sinha et al., 1983; Schwarz et al., 1984; Hamamoto et al., 1986; Kayakawa et al., 1990; Wada and Sekine, 1994; Ravikumar and Cole, 1994) and different phosphoramidite dialkylamino functions (Lyttle et al., 1991; Sinha et al., 1984; Beaucage and Caruthers,
1981) aimed at providing faster coupling rates. Although some of these modifications have been commercialized, the phosphoramidites carrying the 2-cyanoethyl-N,N-diisopropyl combination (Sinha et al., 1983, 1984) are predominantly used. According to this chemistry, the coupling time for 2′-O-TBDMS ribophosphoramidites is considerably longer than that of the 2′-deoxy series for synthesis scales ranging from 0.2 to 2.5 µmol. Particularly important to commercially available ribophosphoramidites is the acidic activation step that allows rapid conversion of stable N,N-diisopropyl phophoramidite to azolide intermediates. These highly reactive transient species are then coupled to the free 5′-hydroxyl of any oligonucleotide bound to a support to form phosphite triester internucleotidic bonds which are further oxidized to the acid-stable pentavalent phosphotriester linkages. Dramatic improvements in oligoribonucleotide synthesis are achieved when activators more acidic than the standard 1-H-tetrazole (pKa = 4.8) are used. Such activators may include 5-(4-nitrophenyl)1H-tetrazole (pKa = 3.7) (Sproat et al., 1989) or 5-ethylthio-1H-tetrazole (pKa = 4.28) (Wincott et al., 1995). These tetrazole derivatives are supposedly more efficient at protonating the trivalent phosphorus. Once protonated, this electrophilic phosphorus center reacts with a tetrazole molecule to displace an N,N-diisopropylamino group. 5-Ethylthio-1H-tetrazole (SET) became the preferred activator for oligoribonucleotides, and has been used in a number of reports describing successful RNA synthesis. However, the acidic properties of 5ethylthio-1H-tetrazole can cripple larger-scale oligoribonucleotide syntheses because of concomitant acidic activator–mediated detritylation of nucleoside phosphoramidites, occurring as a result of an extended coupling reaction time required for satisfactory coupling efficiency (Krotz et al., 1997). This problem spearheaded the development of less acidic but more nucleophilic activators (Vargeese et al., 1998) like 4,5-dicyanoimidazole (pKa = 5.2) or benzimi-
3.6.8 Supplement 1
Current Protocols in Nucleic Acid Chemistry
dazolium triflate (pKa = 4.5) (Hayakawa et al., 1996). Although no mechanistic studies have been presented, these activators are reported to speed up coupling reaction rates due to the increased nucleophilicity of their conjugated base while exhibiting sufficient acidity to protonate the tricoordinated phosphorus center. Automated oligoribonucleotide synthesis can be easily performed on long chain alkylamine controlled pore glass (CPG; Pon et al., 1988; UNITS 3.1 & 3.2). Non-swellable highly cross-linked polystyrene solid-supports have shown to be generally superior to CPG due to their increased mechanical and chemical resistance (McCollum and Andrus, 1991). Deprotection of oligoribonucleotides The deprotection of crude oligoribonucleotides traditionally requires a basic treatment, which allows for the transamination of the N-acyl exocyclic protecting groups, βelimination of the 2-cyanoethyl phosphate protecting group, and cleavage of the succinic ester bond linking the oligoribonucleotide to the solid support (Fig. 3.6.2). Of these three concomitant reactions, base deprotection is by far the most rate-limiting step. Once these reactions are accomplished, a fluoride treatment removes the substituted silyl ether function protecting the 2′-hydroxyl of the ribofuranose ring. Nucleobase deprotection In the early days of oligoribonucleotide chemistry, the heterocyclic amino function of each nucleobase was almost exclusively protected with the benzoyl (rA and rC) and isobutyryl (rG) groups. Although these protecting groups on synthetic DNA are efficiently removed by treatment with concentrated ammonium hydroxide, the presence of the 2′-OTBDMS group in synthesized RNA necessitated the use of ethanolic ammonia (Lyttle, 1993) or a solution of ammonium hydroxide in ethanol (3:1, v/v) for 12 to 16 hours at 55°C to minimize the cleavage of the silyl group (Stawinski, 1988; Wu et al., 1989). Premature deprotection of the 2′-hydroxyl under basic conditions results in extensive oligoribonucleotide chain cleavage from intramolecular nucleophilic attack of the 2′-hydroxyl group on the phophodiester function. To circumvent this unwanted side reaction and shorten the deprotection time, two strategies were investigated over the last decade. One of these strategies involves the use of various combinations of hydrazine, ethano-
lamine, and alcohol (Polushin et al., 1991) or more nucleophilic alkylamines (Wincott et al., 1995; Reddy et al., 1995) for oligoribonucleotide deprotection. For example, 40% aqueous methylamine, in place of or in addition to 30% ammonium hydroxide, cleaves the nucleobase N-acyl protecting groups in a few minutes at 65°C or ~1 hr at room temperature. However, this procedure requires the use of N4-acetyl cytidine phosphoramidite derivatives to avoid a well documented transamination side reaction (Reddy et al., 1994). The second strategy relates to the development of base-labile amino-protecting groups that would further shorten exposure of the 2′O-TBDMS group to basic conditions. These groups belong to the phenoxyacetyl (PAC) (Wu et al., 1988; Chaix et al., 1989) and amidinetype (McBride et al., 1986) protecting groups, mainly used for ribopurine phosphoramidites given that the N4-acetyl protection of cytidine appears quite optimal. Amidine protecting groups include acetamidines (McBride et al., 1986) and dialkylformamidines (Vinayak et al., 1992), which can be cleaved in 2 to 3 hours at 55°C using concentrated ammonium hydroxide/ethanol (3:1, v/v). The main advantage of the dimethylformamidine protecting group when used in conjunction with 2′-deoxyribonucleoside phosphoramidites (Vu et al., 1990) is that it confers an increased resistance towards depurination during the detritylation step. However, because oligoribonucleotides are inherently less sensitive to depurination, N-phenoxyacetyl protection has been preferred for the commercial manufacturing of these biomolecules. The phenoxyacetyl and 4-tert-butyl- or (4-isopropylphenoxy)acetyl protecting groups (Sinha et al., 1993) are considerably more labile than amidines under basic conditions. Typically, these can be quantitatively cleaved from the exocyclic amino function of nucleobases after a 15 min to 1 hr incubation in concentrated ammonium hydroxide/ethanol (3:1) at 65°C, or 2 to 4 hr at room temperature. The use of “fast deprotecting groups” of the PAC family does not preclude the concomitant use of more nucleophilic alkylamines. Indeed, combining these two strategies allows for expedited base-labile removal of protecting groups as exemplified in Alternate Protocol 1. 2′-O-tert-butyldimethylsilyl deprotection The fluoride-sensitive tert-butyldimethylsilyl group allows for an efficient orthogonal deprotection of the 2′-hydroxyl of synthetic RNA. The use of fluoride-based reagents is the
Synthesis of Unmodified Oligonucleotides
3.6.9 Current Protocols in Nucleic Acid Chemistry
Supplement 1
Oligoribonucleotides with 2′-O(tert-Butyldimethylsilyl) Groups
preferred methodology for the removal of the 2′-O-TBDMS group, although Sekine and colleagues (Kawahara et al., 1996) recently reported the application of an acid-catalyzed desilylation scheme to oligoribonucleotides. After completion of nucleobase and phosphotriester deprotection, and subsequent evaporation of the basic solution, addition of n-tetrabutylammonium fluoride (TBAF) in THF to the partially deprotected RNA cleaves the 2′-O-TBDMS group at room temperature within 24 hr (Basic Protocol 2). However, limited solubility of the RNA in this apolar solvent hampers the efficient deprotection of longer oligoribonucleotides. To circumvent this problem, RNA dissolution in either dimethylsulfoxide (Gasparutto et al., 1992) or 50% ethanol (Scaringe, 1995) has been investigated. The notorious water sensitivity of TBAF (Hogrefe et al., 1993) and the desalting step required after quenching the desilylation reaction have led to the development of other fluoride-based reagents. Triethylamine trihydrofluoride (Alternate Protocol 1) has been shown to be a superior reagent either neat (Pirrung et al., 1994; Westman and Stromberg, 1994) or in combination with polar aprotic solvents such as dimethylformamide (Vinayak et al., 1995) or 1-methyl2-pyrrolidinone (Wincott et al., 1995). This reagent promotes 2′-O-TBDMS deprotection within 30 to 90 min at 65°C or 4 to 8 hr at room temperature. Using triethylamine trihydrofluoride, the time-consuming desalting step may be replaced by a sodium acetate/1-butanol precipitation procedure (Wincott et al., 1995) that is not compatible with the high organic content present in TBAF deprotection mixtures. Further application of neat triethylamine trihydrofluoride in a “one-pot deprotection” procedure (Alternate Protocol 2; Bellon, 2000) that employs a mixture containing anhydrous ethanolic methylamine allows for expedititious RNA deprotection. This procedure requires a quenching step with ammonium bicarbonate if one desires to retain the trityl group on the oligonucleotide, and a subsequent desalting/purification on a reverse-phase cartridge because the presence of ethanol and dimethylsulfoxide prevent RNA precipitation from butanol. HPLC analysis and purification of synthetic RNA is now well-documented (Wincott et al., 1995; Sproat et al., 1995; and Vinayak et al., 1995). In particular, the use of perchloratebased buffer (Na or Li form) in conjunction with anion-exchange Nucleo Pac columns, allows for easy purification of full-length RNA product from truncated sequences.
Critical Parameters Automated oligoribonucleotide synthesis according to the phosphoramidite method has not significantly evolved over the last decade. Therefore, all precautions mentioned in the major textbooks (Gait, 1984; Eckstein, 1991; Agrawal, 1993) still remain valid. It is particularly important to emphasize that phosphoramidite chemistry is highly water sensitive. Great care should therefore be taken to ensure that the phosphoramidites and activator are dissolved in strictly anhydrous acetonitrile. All ancillary reagents (i.e., acetonitrile, detrilylation, capping, and oxidation solutions) are commercially available, guaranteeing high performance reproducibility in the syntheses and relieving the chemist from time-consuming anhydrous distillations. Sterile, disposable pipet tips and plastic tubes should be used for storing and handling RNA. Troubleshooting an oligoribonucleotide synthesis is a relatively easy task when the trityl assay is used to spectrophotometrically monitor the synthesis (Gait, 1984; see also APPENDIX 3C, Basic Protocol 1, Support Protocol 1). Deprotection of oligoribonucleotides according to the three protocols presented in this unit is quite straightforward. However, Alternate Protocol 2 or the “one-pot” deprotection protocol should not be used in conjunction with controlled-pore-glass (CPG) synthesized RNA because of the inherent incompatibility between triethylamine trihydrofluoride and the silyl components of CPG. Because all the basic solutions used are composed of gaseous amines dissolved in water or ethanol, freshly opened bottles will ensure that the effective concentration of the amine (i.e., 29% NH4OH or 40% methylamine in water) is close to its stated nominal value. Typically, reagent bottles should be replaced every two weeks if opened on a regular basis. This may be especially true for the TBAF solution in THF (used in Basic Protocol 2) because a low water content is critical for efficient desilylation. The deprotection times at 65°C are suggested for 2 mL of basic solution, and need to be extended if larger amounts of reagents are used. Finally, deprotected oligoribonucleotides are highly sensitive to nuclease degradation. Therefore, gloves should always be worn when manipulating deprotected synthetic RNA; sterile disposable containers and Milli-Q water should also always be used to limit potential exposure to nucleases. DEPC treated water should be used when desalting RNA on C18 cartridges.
3.6.10 Supplement 1
Current Protocols in Nucleic Acid Chemistry
Anticipated Results Because of the iterative nature of solidphase oligoribonucleotide synthesis, only modest chemical yields of oligoribonucleotide can be produced, especially for longer synthetic RNAs (i.e., >30 residues). When applying Basic Protocol 1 using 5-ethylthio-1H-tetrazole, a realistic averaged stepwise chemical yield (ASWY) of 97.5% can be routinely obtained for RNA synthesis. This ASWY is determined by the ratio (µmol FLR/µmol scale)1/n × 100 where µmol FLR is the amount of full length RNA in the crude mixture, µmol scale is the synthesis scale, and n is the number of synthesis cycles. This corresponds to an isolated yield of 41% for the all-RNA 36-mer pictured in Figure 3.6.3.A. Figure 3.6.3.B and C shows HPLC profiles of the same sequence synthesized according to Basic Protocol 1 using the alternate activators DCI and TET, respectively. Figure 3.6.3 indicates that 5-ethylthio-1H-tetrazole is the activator of choice for small scale RNA synthesis. At small scale (i.e., <10 µmol), the quality of oligoribonucleotides generated from the presented deprotection protocols does not vary much. Therefore, one should consider the time requirement as an important parameter in se-
lecting a particular protocol. However, given that oligoribonucleotide synthesis often incorporates one or more modified synthons that may be more or less sensitive to the basic or fluoride treatments used, a careful examination of chemical compatibility should be performed prior to selecting the optimal deprotection method.
Time Considerations
Typically, the duration of the 0.2 µmol elongation cycle of an oligoribonucleotide is ∼12 min. It will thus take ∼8 hr to synthesize an all-RNA 37-mer. It requires about 32 hr to perform oligoribonucleotide deprotection according to Basic Protocol 2, whereas processing time can be cut to <8 hrs when adopting Alternate Protocol 1. When using Alternate (“one-pot”) Protocol 2, deprotection time can be shortened to a mere 45 min from start to finish.
Acknowlegments The author gratefully acknowledges Chris Workman, Lara Maloney, Chris Shaffer, and Victor Mokler for their efforts in the development of the second alternative deprotection
Figure 3.6.3 HPLC chromatograms of a 36-mer oligoribonucleotide (5′- GUU UUC CCU GAU GAG GCC GAA AGG CCG AAA UUC UCC -3′) synthesized at the 0.2 µmol scale according to the basic synthesis protocol and deprotected using the “one-pot” deprotection protocol (Alternate Protocol 2). (A) 0.25 M SET, (B) 0.5 M DCI, (C) 0.45M TET. Results obtained using Dionex NucleoPac PA-100 22 × 250–mm column at 50°C. Buffer A (1 mM Tris, 20 mM NaClO4), buffer B (1 mM Tris, 300 mM NaClO4). Gradient 40% B to 70% B in 12 min, flow rate = 1.5 mL/min (see UNIT 10.5).
Synthesis of Unmodified Oligonucleotides
3.6.11 Current Protocols in Nucleic Acid Chemistry
Supplement 1
protocol, and Drs. Fran Wincott and Nassim Usman for continuous support.
Literature Cited Agrawal, S. 1993. Protocols for oligonucleotides and analogs: Synthesis and properties. Humana Press, Totowa, N.J. Beaucage, S.L. and Caruthers, M.H. 1981. Deoxynucleoside phosphoramidites: A new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Lett. 37:1859-1862. Bellon, L. and Workman, C. 2000. Deprotection of RNA. US patent 6,054,576. Chaix, C., Molko, D., and Teoule, R. 1989. The use of labile base protecting groups in oligoribonucleotide synthesis. Tetrahedron Lett. 30:71-74. Ecktein, F. 1991. Oligonucleotides and analogues: A practical approach. IRL Press, New York. Gait, M.J. 1984. Oligonucleotide synthesis: A practical approach. IRL Press, Washington D.C. Gasparutto, D., Livache, T., Bazin, H., Duplaa, A.M., Guy, A., Khorlin, A., Molko, D., Roget, D., and Teoule, R. 1992. Chemical synthesis of a biologically active natural tRNA with its minor bases. Nucl. Acids Res. 20:5159-5166. Hamamoto, S. and Takaku, H. 1986. New approach to the synthesis of deoxyribonucleoside phosphoramidite derivatives. Chem. Lett. 1401-1404. Hayakawa, Y., Kataoka, M., and Noyori, R. 1996. Benzimidazolium triflate as an efficient promoter for nucleotide synthesis via the phosphoramidite method. J. Org. Chem. 61:79967997. Hogrefe, R., McCaffrey, A., Borodzine, L., McCampbell, E., and Vaghefi, M. 1993. Effect of excess water on the desilylation of oligoribonucleotides using tetrabutylammonium fluoride. Nucl. Acids Res. 21:4739-4741. Kawahara, S., Wada, T., and Sekine, M. 1996. Unprecedented mild acid-catalyzed desilylation of 2′-O-tert-butyldimethylsilyl group from chemically synthesized oligoribonucleotides intermediates via neighboring group participation of the internucleotidic phosphate residue. J. Am. Chem. Soc. 118:9461-9468. Kayakawa, Y., Wakabayashi, S., Kato, H., and Noyori, R. 1990. The allylic protection in solid-phase oligonucleotide synthesis: An efficient preparation of solid-anchored DNA oligomers. J. Am. Chem. Soc. 112:1691-1696. Krotz, A., Klopchin, P., Walker, K., Srivatsa, S., Cole, D., and Ravikumar, V. 1997. On the formation of longmers in phosphorothioate oligodeoxyribonucleotide synthesis. Tetrahedron Lett. 38:3875-3878.
Oligoribonucleotides with 2′-O(tert-Butyldimethylsilyl) Groups
Lyttle, M., Wright, P., Sinha, N., Bain, J., and Chamberlin, A. 1991. New nucleoside phosphoramidites and coupling protocols for solidphase RNA synthesis. J. Org. Chem. 56:46084615.
Lyttle, M. 1993. Chain cleavage during deprotection of RNA synthesized by the 2′-O-trialkylsilyl protection strategy. Nucleosides Nucleotides. 12:95-106. McBride, L.J., Kierzek, R., Beaucage, S.L., and Caruthers M.H. 1986. Amidine protecting groups in oligonucleotide synthesis. J. Am. Chem. Soc. 108:2040-2048. McCollum, C. and Andrus, A. 1991. An optimized polystyrene support for rapid efficient oligonucleotide synthesis. Tetrahedron Lett. 32:40694072. Ogilvie, K.K., Beaucage, S.L., Entwistle, D.W., Thompson, E.A., Quilliam, M.A., and Westmore, J.B. 1976. Alkylsilyl groups in nucleoside and nucleotide chemistry. J. Carbohydr. Nucleosides Nucleotides. 3:197-227. Ogilvie, K.K., Theriault, N., and Sadana, K. 1977. Synthesis of oligoribonucleotides. J. Am. Chem. Soc. 99:7741-7743. Pirrung, M., Shuey, S., Lever, D., and Fallon, L. 1994. A convenient procedure for the deprotection of silylated nucleosides and nucleotides using triethylamine trihydrofluoride. Tetrahedron Lett. 4:1345-1346. Polushin, N., Pashkova, I., and Efimov, V. 1991. Rapid deprotection procedures for synthetic oligonucleotides. Nucl. Acids Symp. Series. 24:4950. Pon, R., Usman, N., and Ogilvie, K. 1988. Derivatization of controlled pore glass beads for solidphase oligonucleotide synthesis. BioTechniques 6:768-775 Ravikumar, V., and Cole, D. 1994. 2-diphenylmethylsilylethyl (DPSE): A versatile protecting group for oligonucleotide synthesis. Gene 149:157-161. Reddy, M.P., Hanna, N.B., and Farooqui, F. 1994. Fast cleavage and deprotection of oligonucleotides. Tetrahedron Lett. 35:4311-4314. Reddy, M.P., and Hanna, N.B., Farooqui, F. 1995. Methylamine deprotection provides increased yield of oligoribonucleotides. Tetrahedron Lett. 35:4311-4314. Scaringe, S.A., Francklyn, C., and Usman, N. 1990. Chemical synthesis of biologically active oligoribonucleotides using β-cyanoethyl protected ribonucleoside phosphoramidites. Nucl. Acids Res. 18:5433-5441. Schwarz, M., and Pfleiderer, W. 1984. Solution synthesis of fully protected thymidine dimers using various phosphoramidites. Tetrahedron Lett. 24:5513-5516. Sinha, N., Biernat, J., and Koster, H. 1983. Cyanoethyl N,N-dialkylamino-N-morpholinomonochloro phosphoramidites: New phosphitylating agents facilitating ease of deprotection and work-up of synthesized oligonucleotides. Tetrahedron Lett. 24:5843-5846.
3.6.12 Supplement 1
Current Protocols in Nucleic Acid Chemistry
Sinha, N., Biernat, J., McManus, J., and Koster, H. 1984. Use of cyanoethyl-N,N-dialkylamine-/Nmorpholino phosphoramidite of deoxynucleosides for the synthesis of DNA fragments simplifying deprotection and isolation of the final product. Nucl. Acids Res. 12:4539-4557. Sinha, N., Davis, P., Usman, N., Perez, J., Hodge, R., Kremsky, J., and Casale, R. 1993. Labile exocyclic amine protection of nucleosides in DNA, RNA and oligonucleotide analog synthesis facilitating deacylation, minimizing depurination and chain degradation. Biochimie. 75:1323. Sproat, B., Lamond, A., Beijer, B., Neuner, P., and Ryder, U. 1989. Highly efficient chemical synthesis of 2′-O-methyloligoribonucleotides and tetrabiotinylated derivatives: Novel probes that are resistant to degradation by RNA or DNA specific nucleases. Nucl. Acids Res. 17:33733386. Sproat, B., Colonna, F., Mullah, B., Tsou, D., Andrus, A., Hampel, A., and Vinayak, R. 1995. An efficient method for the isolation and purification of oligoribonucleotides. Nucleosides Nucleotides. 14:255-273. Stawinski, J., Stromberg, R., Thelin, M., and Westman, E. 1988. Studies on the t-butyldimethylsilyl group as 2′-O-protection in oligoribonucleotide synthesis via the H-phosphonate approach. Nucl. Acids Res. 16:9285-9298. Usman, N., Pon, R., and Ogilvie, K. 1985. Preparation of ribonucleoside 3′-O-phosphoramidites and their application to the automated solidphase synthesis of oligonucleotides. Tetrahedron Lett. 26:4567-4570. Usman, N., Ogilvie, K., Jiang, M.Y., and Cedergren, R. 1987. Automated chemical synthesis of long oligoribonucleotides using 2′-O-silylated ribonucleoside 3′-O-phosphoramidtes on CPG. J. Am. Chem. Soc. 109:7845-7854. Usman, N. and Cedergren, R. 1992. Exploiting the chemical synthesis of RNA. Trends Biochem. Sci. 17:334-339. Vargeese, C., Carter, J., Yegge, J., Krivjansky, S., Settle, A., Kropp, E., Peterson, K., and Pieken, W. 1998. Efficient activation of nucleoside phosphoramidites with 4,5-dicyanoimidazole during oligonucleotide synthesis. Nucl. Acids Res. 26:1046-1050.
Vinayak, R., Anderson P., McColum C., and Hampel, A. 1992. Chemical synthesis of RNA using fast oligonucleotide deprotection chemistry. Nucl. Acids Res. 20:1265-1269. Vinayak, R., Andrus, A., Mullah, B., and Tsou, D. 1995. Advances in the chemical synthesis and purification pf RNA. Nucl. Acids Symp. Series 33:123-125. Vu, H., McColum C., Jacobson K., Theisen P., Vinayak R., Spiess E., and Andrus, A. 1990. Fast oligonucleotide deprotection phosphoramidite chemistry for DNA synthesis. Tetrahedron Lett. 31:7269-7272. Wada, T. and Sekine, M. 1994. 2-(trimethylsilyl)ethyl as a phosphate protecting group in oligonucleotide synthesis. Tetrahedron Lett. 35:757-760. Westman, E. and Stromberg, R. 1994. Removal of t-butyldimethylsilyl protection in RNA synthesis: Triethylamine trihydrofluoride is a more reliable alternative to tetrabutylammonium fluoride. Nucl. Acids Res. 22:2430-2431. Wincott, F. and Usman, N. 1994. 2′-(trimethylsilyl)ethoxymethyl protection of the 2′-hydroxyl group in oligoribonucleotide synthesis. Tetrahedron Lett. 35:6827-6830. Wincott, F., DiRenzo, A., Shaffer, C., Grimm, S., Tracz, D., Workman, C., Sweedler, D., Gonzalez, C., Scaringe, S., and Usman, N. 1995. Synthesis, deprotection, analysis and purification of RNA and ribozymes. Nucl. Acids Res. 23:2677-2684. Wu, T., Ogilvie, K.K., and Pon, R.T. 1988. N-phenoxyacetylated guanosine and adenosine phosphoramidites in the solid phase synthesis of oligoribonucleotides: synthesis of a ribozyme sequence. Tetrahedron Lett. 29:4249-4252. Wu, T., Ogilvie, K.K., and Pon, R.T. 1989. Prevention of chain cleavage in the chemical synthesis of 2′-O-silylated oligoribonucleotides. Nucl. Acids Res. 17:3501-3517.
Contributed by Laurent Bellon Ribozyme Pharmaceuticals Boulder, Colorado
Synthesis of Unmodified Oligonucleotides
3.6.13 Current Protocols in Nucleic Acid Chemistry
Supplement 1
Synthesis of Oligoribonucleotides Using the 2-Nitrobenzyloxymethyl Group for 2′-Hydroxyl Protection
UNIT 3.7
The methodology for oligoribonucleotide construction described in the Basic Protocol uses a DNA synthesizer to add protected ribonucleoside phosphoramidite monomers one after another in defined sequence to a 3′-terminal nucleoside that is attached to a solid support. The monomers used are the 3′-O-(2-cyanoethyl-N,N-diisopropyl) phosphoramidites of Nprotected 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl) ribonucleosides. Their structures are shown in Figure 3.7.1. After a target oligoribonucleotide has been synthesized, it is treated with ammonia to release it from the support and to remove protecting groups from its nucleobases and phosphates. Finally, it is irradiated with long-wave UV light, which removes the 2-nitrobenzyloxymethyl groups from the 2′-hydroxyls, to give the unmodified RNA. The preparation of the 3′-O-(2-cyanoethyl-N,N-diisopropyl) phosphoramidites is described in the Support Protocol. SYNTHESIS AND DEPROTECTION OF N-PROTECTED 2′-O-(2-NITROBENZYLOXYMETHYL) OLIGORIBONUCLEOTIDES
BASIC PROTOCOL
This protocol describes the steps involved in constructing oligoribonucleotides using a DNA synthesizer and starting with phosphoramidites of the 2′-O-(2-nitrobenzyloxymethyl) derivatives of the four common ribonucleosides U, C, A, and G. Details of those parts of the operation that are carried out on the synthesizer will vary depending on the make and model used. However, the above monomers are compatible with all the usual reagents used in these instruments and, with the few exceptions noted below, the manufacturer’s directions for setup and synthesis should be followed in each specific case. A generalized discussion of the use of DNA synthesizers can be found in APPENDIX 3C. In contrast, the deprotection methodology that constitutes the second section of the protocol is unique to oligoribonucleotides constructed from these particular monomers, and is set out in detail below. Materials 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-3′-O-(2-cyanoethylN,N-diisopropyl) phosphoramidites (U, C, A, and G; see Support Protocol) Phosphorus pentoxide Anhydrous acetonitrile (Aldrich) 0.45 M tetrazole (Amersham Pharmacia Biotech) in anhyrdous acetonitrile Pyridine, HPLC grade (Aldrich) Concentrated ammonium hydroxide, 4°C 50% (v/v) aqueous pyridine 50% (v/v) aqueous 2-methyl-2-propanol, HPLC grade 0.2 M formic acid 0.2 M ammonium hydroxide DNA synthesizer with ancillary reagents Amidite vials fitted with septa 5-mL syringes and 21-G needles Synthesis column containing support derivatized with O-acyl ribonucleosides (Glen Research; see Critical Parameters) 10.2-cm pressure tube (e.g., Ace Glass) fitted with Teflon screw-in cap and Teflon-encapsulated O ring Contributed by Tod J. Miller, Miriam E. Schwartz, and Geoffrey R. Gough Current Protocols in Nucleic Acid Chemistry (2000) 3.7.1-3.7.8 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Unmodified Oligonucleotides
3.7.1 Supplement 3
O O
HN NH
DMTrO
5'
N
O
DMTrO
O 3'
(i-Pr)2N
N N
O
O
2'
O
O
O
P
O
(i-Pr)2N NO2
OCH2CH2CN
O
P
O NO2
OCH2CH2CN
O O
HN N N
DMTrO
(i-Pr)2N
P
O O
O
OCH2CH2CN
N
N
N
DMTrO
N
O
(i-Pr)2N NO2
P
NH N
O O
O
O
N H
O
OCH2CH2CN
NO2
CH3O
DMTr =
C
CH3O
i-Pr, isopropyl
Figure 3.7.1 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-3′-O-phosphoramidites of the four common ribonucleosides. Chemical structures are shown for (clockwise from upper-left corner): uridine phosphoramidite, cytidine phosphoramidite, guanosine phosphoramidite, and adenosine phosphoramidite. DMTr, dimethoxytrityl group (bottom).
Synthesis of Oligoribonucleotides Using the 2-Nitrobenzyloxymethyl Group
Yellow lights Covered water bath, 55°C 30-mL Buchner funnel (medium porosity) 500-mL filter flask Vacuum aspirator 500-mL round-bottom flask Rotary evaporator connected to water aspirator, with bath temperature set <40°C 250-mL Pyrex Erlenmeyer flasks Long-wave (365-nm) UV lamp, containing two 15-W black-light tubes behind a UV-transmitting glass filter measuring ∼10 × 30 cm NOTE: The amidites are sensitive to UV light (see Critical Parameters).
3.7.2 Supplement 3
Current Protocols in Nucleic Acid Chemistry
Synthesize oligoribonucleotides 1. Determine volumes of amidite solutions needed to carry out synthesis of the target oligoribonucleotide, following the manufacturer’s instructions for the DNA synthesizer. 2. Use the following concentrations to calculate the required weight of each amidite. U C A G
137 mg/mL 152 mg/mL 156 mg/mL 153 mg/mL.
These values correspond to a concentration of 0.15 M for each amidite.
3. Weigh out the calculated quantity of each 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl)-3′-O-(2-cyanoethyl-N,N-diisopropyl) phosphoramidite into a separate amidite vial. 4. Dry amidites 12 to 16 hr in a vacuum desiccator over phosphorus pentoxide, then cap vials with rubber septa. The phosphorus pentoxide should be allowed to absorb a little atmospheric moisture before being placed in the desiccator, so that the amidites are not contacted by loose dust from the compound’s surface.
5. Dissolve each amidite in the required volume of anhydrous acetonitrile (as determined in step 1), using a fresh 5-mL syringe and 21-G needle to deliver the solvent through the septum of the vial. A syringe and needle should be used to avoid moisture contamination.
6. Fit amidite vials onto the DNA synthesizer as part of the general setup procedure, following the manufacturer’s specific directions. 7. Use 0.45 M tetrazole in anhydrous acetonitrile as the phosphoramidite activator solution. 8. Set the coupling time at 3 min. 9. Install a synthesis column containing support derivatized with O-acyl ribonucleosides and carry out the synthesis. The amidites and growing oligoribonucleotide must be protected from light.
10. Perform the final detritylation on the machine, then remove the synthesis column and either begin the deprotection process immediately, or store the column up to 1 week at –20°C. Deprotect oligoribonucleotides 11. Empty derivatized support from the synthesis column into a 10.2-cm pressure tube containing 2 mL pyridine. Rinse column with pyridine if necessary, but do not allow total amount of pyridine in the pressure tube to exceed 4 mL. NOTE: Steps 11 through 16 should be carried out under yellow lights (see Critical Parameters).
12. Cool pressure tube in ice and fill with cold concentrated ammonium hydroxide to just below the level of the screw cap. A volume (∼16 mL) of ammonium hydroxide should be used that minimizes air space in the closed tube. Only full-strength ammonium hydroxide from a recently opened bottle should be used. The bottle should be sealed and stored at 4°C when not in use.
Synthesis of Unmodified Oligonucleotides
3.7.3 Current Protocols in Nucleic Acid Chemistry
Supplement 3
13. Screw in a Teflon cap fitted with a Teflon-encapsulated O ring, making sure seal is tight. Wrap tube in aluminum foil and heat 24 hr in a covered water bath at 55°C. Invert tube occasionally to mix contents. 14. Chill tube on ice, open carefully, and quickly filter contents through a 30-mL medium-porosity Buchner funnel into a 500-mL filter flask using a vacuum aspirator. CAUTION: This should be performed in a fume hood because the mixture bubbles violently as ammonia is released.
15. Transfer filtrate into a 500-mL round-bottom flask, using two 10-mL washes of 50% aqueous pyridine. 16. Evaporate solution to ∼10 mL using a rotary evaporator connected to a water aspirator. Add 100 mL water and again evaporate to 10 mL. The evaporation should be carried out at the lowest bath temperature (<40°C) consistent with efficient removal of solvent. The solution should not be evaporated to dryness.
17. Add 100 mL of 50% aqueous 2-methyl-2-propanol and evaporate to ∼5 mL. Repeat this process once more to remove traces of pyridine. 18. Transfer solution into a graduated beaker and dilute with 50% aqueous 2-methyl-2propanol so that the final volume corresponds to a concentration of 1 A260 unit of oligonucleotide/mL. Stir solution. The volume can be roughly calculated using the formula V = 10L × S, where V is the final volume in milliliters, L is the number of nucleotides in the oligomer, and S is the scale of the reaction in micromoles.
19. Adjust pH of the solution to 3.7 by careful dropwise addition of 0.2 M formic acid. Do not overshoot. Removal of the 2′-hydroxyl-protecting groups is pH dependent, and is most efficient at pH 3.7.
20. Place 50-mL aliquots of the solution in 250-mL Pyrex Erlenmeyer flasks. Pyrex is transparent to long-wave UV light.
21. In a 4°C cold room, set up flasks on the flat glass filter of a long-wave UV lamp. Irradiate solutions 4.5 hr with intermittent swirling. A brilliant sky-blue fluorescence develops as photolysis products derived from the 2-nitrobenzyl group accumulate in the solution. CAUTION: UV light can cause eye damage. Wear protective glasses and shield the irradiation setup with a sheet of aluminum foil to prevent inadvertent exposure of others.
22. Recombine solutions and adjust pH back up to 7 to 8 by careful addition of 0.2 M ammonium hydroxide with stirring. RNA is degraded by base at a pH above 8. Step 23 should be carried out immediately if too much ammonium hydroxide is added by mistake.
23. Evaporate solution to a volume of 1 to 2 mL under vacuum using the rotary evaporator. The deprotected oligoribonucleotide can now be purified by anion-exchange HPLC or by gel electrophoresis (UNITS 10.4 & 10.5). Synthesis of Oligoribonucleotides Using the 2-Nitrobenzyloxymethyl Group
3.7.4 Supplement 3
Current Protocols in Nucleic Acid Chemistry
SYNTHESIS AND PURIFICATION OF 3′-O-(2-CYANOETHYL-N,N-DIISOPROPYL) PHOSPHORAMIDITES OF 5′-O-(4,4′-DIMETHOXYTRITYL)-2′-O-(2-NITROBENZYLOXYMETHYL) RIBONUCLEOSIDES
SUPPORT PROTOCOL
This section describes a method for obtaining the phosphoramidite monomers of U, C, A, and G used in the Basic Protocol. Materials N-Protected 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl) ribonucleosides (UNIT 2.5) Pyridine, HPLC grade (Aldrich) Anhydrous toluene (Aldrich) Anhydrous tetrahydrofuran (Aldrich) N,N-Diisopropylethylamine (Aldrich) 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite (Aldrich) 99:1 (v/v) chloroform/methanol containing 3% (v/v) triethylamine Ethyl acetate 1 M sodium carbonate (for U, A, G) or 1 M sodium bicarbonate (for C), ice cold Merck 60 silica gel (230 to 400 mesh ASTM) 4:1 (v/v) ethyl acetate/hexane containing 0.1% (v/v) triethylamine (for A, C, and U) 0.5%, 1.0%, 1.5%, 2.0%, 2.5%, and 3.0% (v/v) methanol in dichloromethane containing 5% (v/v) triethylamine (for G) Anhydrous diethyl ether Dry argon 100-mL flask fitted with a septum Rotary evaporator, with its air inlet connected to a Drierite gas-drying unit, connected interchangeably to water aspirator and vacuum pump 1-mL syringe and 21-C needle 3 × 60–cm flash chromatography column, with reservoir and flow controller 100-mL round-bottom flask Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) NOTE: Triethylamine, N,N-diisopropylethylamine, and pyridine should be kept dry over calcium hydride granules. Synthesize phosphoramidites 1. Using a rotary evaporator connected to a water aspirator, dry 1 mmol of any N-protected 5′-O-(4,4′-dimethoxytrityl)-2′-O-(2-nitrobenzyloxymethyl) ribonucleoside in a 100-mL flask by two successive co-evaporations with each of the following solvents: 20 mL pyridine, 20 mL anhydrous toluene, and 20 mL anhydrous tetrahydrofuran. 2. Dissolve residue in 3 mL anhydrous tetrahydrofuran and add 0.7 mL (4 mmol) N,N-diisopropylethylamine. Add a stir-bar, seal flask with a septum, and begin stirring. 3. Using a 1-mL syringe and 21-G needle, add 0.44 mL (2 mmol) 2-cyanoethyl-N,Ndiisopropylchlorophosphoramidite dropwise over a 2-min period through the septum. Continue stirring 2 hr at room temperature.
Synthesis of Unmodified Oligonucleotides
3.7.5 Current Protocols in Nucleic Acid Chemistry
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4. Examine reaction mixture by TLC (APPENDIX 3D) in 99:1 chloroform/methanol containing 3% triethylamine. The product should appear under short-wave UV light as two closely spaced bands of approximately equal intensity, corresponding to its two diastereomeric forms.
5. Dissolve reaction mixture in 150 mL ethyl acetate and extract three times with 75 mL ice-cold 1 M sodium carbonate (1 M sodium bicarbonate for phosphoramidite of C). 6. Evaporate ethyl acetate solution to an oil using the rotary evaporator with the water aspirator. Purify phosphoramidites 7. Prepare a 3 × 60–cm flash chromatography column with a 100-mL bed volume of Merck 60 silica gel packed in starting solvent. 8a. For phosphoramidites of U, C, and A: Dissolve phosphoramidite oil in 2 to 3 mL starting solvent and apply to chromatography column. Use 600 mL of 4:1 ethyl acetate/hexane containing 0.1% triethylamine to elute. Collect 15-mL fractions and assay by TLC in 99:1 chloroform/methanol containing 3% triethylamine. 8b. For phosphoramidites of G: Dissolve phosphoramidite oil in 2 to 3 mL starting solvent and apply to chromatography column. Use a stepwise gradient of 50 mL each of 0.5%, 1.0%, 1.5%, 2.0%, 2.5%, and 3.0% methanol in dichloromethane containing 5% triethylamine to elute. Collect 15-mL fractions and assay by TLC in 99:1 chloroform/methanol containing 3% triethylamine. 9. Combine fractions containing pure product in a 100-mL round-bottom flask and evaporate solution to an oil using the rotary evaporator with the water aspirator. 10. Dissolve residue in 1 to 2 mL anhydrous diethyl ether. Concentrate solution on the rotary evaporator first using the water aspirator, and finally under high vacuum (i.e., with a vacuum pump) until a stable foam is formed. 11. Fill flask with dry argon, seal it, and wrap in aluminum foil. Store amidite up to 1 year at −80°C in the dark. COMMENTARY Background Information
Synthesis of Oligoribonucleotides Using the 2-Nitrobenzyloxymethyl Group
Ribonucleoside phosphoramidite monomers with their 2′-hydroxyls protected by 2-nitrobenzyloxymethyl groups (Schwartz et al., 1992) were introduced at a time when phosphoramidites with other modes of 2′-protection required very long coupling times on synthesizers, often >10 min per condensation. The nitrobenzyloxymethyl compounds were unique in needing coupling times not much longer than those of DNA amidites (i.e., ∼2 min). This enhanced reactivity may be due to the presence of the methylenedioxy “arm” that forms part of the 2-nitrobenzyloxymethyl group and allows it the flexibility to remain out of the way of the reactive phosphoramidite center. However, problems of low reactivity with other types of amidites have been largely
overcome by the introduction of newer, more powerful activating agents such as 5-ethylthioH1-tretrazole (Vinayak et al., 1994), and machine-assisted synthesis of RNA is now generally as rapid and simple as that of DNA. The protocol described in this unit, for example, has been used to prepare biologically functional RNA of up to 33 nucleotides in length.
Compound Characterization The 31P nuclear magnetic resonance (31PNMR) chemical shifts for the 2′-O-(2-nitrobenzyloxymethyl)-protected phosphoramidites of U, C, A, and G are given in Table 3.7.1. If these monomers have been properly prepared and purified, no extraneous peaks should be present in their NMR spectra. Oligoribonucleotides made from these monomers can be charac-
3.7.6 Supplement 3
Current Protocols in Nucleic Acid Chemistry
Table 3.7.1 31P-NMR Chemical Shifts of 2′-O-(2-Nitrobenzyloxymethyl)-Protected Ribonucleoside Phosphoramiditesa
Ribonucleoside
Downfield (ppm)
Upfield (ppm)
A C G U
151.359 152.552 151.408 151.501
151.095 151.127 151.141 150.471
a
In acetonitrile-d3, relative to 85% H3PO4 as external standard at 0 ppm.
terized, after deprotection and purification, using an RNA sequencing kit (nuclease method, e.g., USB) or by digestion with snake venom phosphodiesterase and bacterial alkaline phosphatase followed by reversed-phase HPLC analysis of the digest.
Critical Parameters The techniques for RNA synthesis described in this protocol have been thoroughly tested using polystyrene supports in conjunction with an easily prepared universal adapter that allows any sequence, DNA or RNA, to be assembled on any kind of support (Schwartz et al., 1995). Consequently, this system is strongly recommended. Use of controlled-pore glass has been less well studied. Supports of this type should be chosen with caution as they are attacked by hot ammonia solutions, and the effects of significant amounts of dissolved silica on the deprotection of longer oligoribonucleotides by the methods described here are unknown. In any case, synthesis columns that contain a support derivatized with base-labile O-acyl ribonucleosides (e.g., the 3′-RNA supports supplied by Glen Research) should be used in this protocol. Only these nucleosides will be completely deprotected under the conditions specified. The ribonucleoside phosphoramidites are prepared using the method of Sinha et al. (1984). As in all such syntheses, it is essential to maintain anhydrous conditions. Make sure all glassware is dry and avoid exposing amidite reagents or their solutions to atmospheric moisture. Take further precautions when using these particular amidites because of their sensitivity to UV light. Enough of this radiation exists as a component of standard laboratory lighting to cause slow loss of 2-nitrobenzyloxymethyl groups. Loss of 2′-protecting groups in RNA leads to chain cleavage during the ammonia treatment that is part of the deprotection process. Take steps to minimize exposure not only
of the amidites but also of the synthesized oligoribonucleotides to UV light. This means replacing overhead fluorescent bulbs in a designated work area with yellow bulbs, such as GE or Sylvania Golds. Wrap amidite vials, synthesis columns, and the vessels used in the deprotection process (up to the point where ammonia is removed from the mixtures) in aluminum foil. Once the oligomers are deprotected, they become vulnerable to the action of ribonucleases, which are everywhere in the laboratory, including on the experimenter’s skin. Always wear latex gloves when handling RNA, heat all glassware at 250°C for a few hours before use, and maintain a supply of freshly distilled water in covered containers at 4°C. Store pH electrodes in solutions containing thymol crystals.
Anticipated Results Yields of ribonucleoside phosphoramidites should be 80% to 90%. In oligoribonucleotide syntheses, the amounts of isolated, deprotected oligoribonucleotides that are obtained decrease with increasing length. However, for products up to 20 nucleotides long, yields in the range of 10% to 30%, based on support-bound starting nucleoside, can be expected.
Time Considerations The ribonucleoside phosphoramidites each require one day for preparation and purification. Oligoribonucleotide synthesis takes a day, and deprotection requires a further 2 to 3 days.
Literature Cited Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., deBear, J.S., and Gough, G.R. 1992. Rapid synthesis of oligoribonucleotides using 2′-O-(2-nitrobenzyloxymethyl)-protected monomers. BioMed. Chem. Lett. 2:1019-1024. Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., and Gough, G.R. 1995. A universal adapter for chemical synthesis of DNA or RNA on any single type of solid support. Tetrahedron Lett. 36:27-30.
Synthesis of Unmodified Oligonucleotides
3.7.7 Current Protocols in Nucleic Acid Chemistry
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Sinha, N.D., Biernat, J., McManus, J., and Köster, H. 1984. Polymer support oligonucleotide synthesis XVIII: Use of β-cyanoethyl-N,N-dialkylamino-/N-morpholino phosphoramidite of deoxynucleosides for the synthesis of DNA fragments simplifying deprotection and isolation of the final product. Nucl. Acids Res. 12:45394557. Vinayak, R., Colonna, F., Tsou, D., Mullah, B., Andrus, A., and Sproat, B. 1994. Large scale chemical synthesis and purification of RNA. Nucl. Acids Symp. Ser. 31:165–166.
Key References Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., deBear, J.S., and Gough, G.R. 1992. Rapid synthesis of oligoribonucleotides using 2′-O-(2- nitrobenzyloxymethyl)-protected monomers. Bio. Med. Chem. Lett. 2:1019-1024. Provides an overview of the use of 2-nitrobenzyloxymethyl as a 2′-hydroxyl-protecting group in oligoribonucleotide synthesis.
Contributed by Tod J. Miller, Miriam E. Schwartz, and Geoffrey R. Gough Purdue University West Lafayette, Indiana
Synthesis of Oligoribonucleotides Using the 2-Nitrobenzyloxymethyl Group
3.7.8 Supplement 3
Current Protocols in Nucleic Acid Chemistry
Chemical Synthesis of RNA Sequences with 2′-O-[(Triisopropylsilyl)oxy]methylprotected Ribonucleoside Phosphoramidites
UNIT 3.8
This unit describes the chemical synthesis of oligoribonucleotides from 2′-O-[(triisopropylsilyl)oxy]methyl-protected phosphoramidites (TOM-phosphoramidites), including TOM-phosphoramidite preparation (see Basic Protocol 1), assembly on DNA synthesizers (see Basic Protocol 2), and subsequent deprotection (see Basic Protocol 3). The preparation of the TOM-protected phosphoramidite building blocks is carried out by introducing a 2-cyanoethyl-N,N-diisopropylaminophosphinyl group to the 3′-O position of N-acetyl-5′-O-(4,4′-dimethoxytrityl)-2′-O-[(triisopropylsilyl)oxy]methyl ribonucleosides. The preparation of these TOM-protected ribonucleosides is described in UNIT 2.9. The assembly of RNA sequences with TOM-phosphoramidites can be carried out under essentially the same conditions as the assembly of DNA sequences. With very minor exceptions, the same chemistry is employed and no additional equipment is required. Due to the minimal steric hindrance of the TOM protecting group, RNA syntheses with TOM-phosphoramidites are more efficient than syntheses with the traditional 2′- O-(tertbutyldimethylsilyl)phosphoramidites (TBDMS-phosphoramidites, UNIT 3.6). Specifically, better coupling yields are obtained with much shorter coupling times (99% at 2.5 min). This property, together with a reliable deprotection behavior, allows the efficient preparation of relatively long RNA sequences, even on a large scale and within a short time. The assembly of oligoribonucleotides is carried out by stepwise addition of phosphoramidite building blocks to an immobilized nucleoside until the desired sequence has been obtained. Each addition of a new building block requires four reactions (detritylation, coupling, capping, oxidation) that make up one cycle. After the assembly, the detachment of the sequence from the solid support and the removal of the nucleobaseand phosphodiester-protecting groups are carried out under basic nucleophilic conditions with methylamine/water/ethanol, followed by removal of the remaining 2′-O-TOM protecting groups with tetra-n-butylammonium fluoride. After workup by size-exclusion chromatography, crude sequences are obtained, which ultimatively can be purified by high-performance liquid chromatography (HPLC; UNIT 10.5) or polyacrylamide gel electrophoresis (PAGE; UNIT 10.4). CAUTION: All reactions must be performed in a well-ventilated fume hood to avoid exposure to 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite. NOTE: Always use the highest quality of solvents and reagents. All reagents and solvents used for deprotection must be sterile and free of RNases (e.g., Fluka reagents for molecular biology). PREPARATION OF 2′-O-TOM-PROTECTED PHOSPHORAMIDITES The conversion of N-acetyl-5′-O-DMTr-2′-O-TOM-protected ribonucleosides into the corresponding phosphoramidite building blocks is shown in Figure 3.8.1. The structure of the TOM protecting group prevents its migration from the 2′-O to the 3′-O position, even under basic reaction conditions. Without extraction, the products are directly purified by chromatography on silica gel. In order to obtain pure phosphoramidite building blocks, the starting materials and the reaction solvent should be as dry as possible. For a
BASIC PROTOCOL 1
Synthesis of Unmodified Oligonucleotides Contributed by Stefan Pitsch and Patrick A. Weiss Current Protocols in Nucleic Acid Chemistry (2001) 3.8.1-3.8.15 Copyright © 2001 by John Wiley & Sons, Inc.
3.8.1 Supplement 7
Figure 3.8.1 Preparation of the four 2′-O-TOM-protected ribonucleoside phosphoramidites S.2a-S.2d. D M Tr, 4,4′-dimethoxytrityl; i-Pr2NE t, N-ethyl-N,N-diisopropylamine; iPr2NP(Cl)OCH2CH2CN, 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite.
straightforward isolation, the reaction should be complete and the minimal amount of phosphitylation agent should be employed. NOTE: The products are very acid sensitive; any contact with acids must be completely avoided. Materials Nitrogen or argon gas source N-Acetyl-5′-O-(4,4′-dimethoxytrityl)-2′-O-[(triisopropylsilyl)oxy]methyl ribonucleosides (UNIT 2.9): N6-Ac-5′-O-DMTr-2′-O-TOM-adenosine (S.1a) N2-Ac-5′-O-DMTr-2′-O-TOM-guanosine (S.1b) N4-Ac-5′-O-DMTr-2′-O-TOM-cytidine (S.1c) 5′-O-DMTr-2′-O-TOM-uridine (S.1d) Dichloromethane N-Ethyl-N,N-diisopropylamine 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite (Aldrich) Ethyl acetate Hexane Methanol Anisaldehyde reagent (see recipe) Silica gel (230 to 400 mesh, Fluka or Merck) Sand 250-mL one-neck flask equipped with a stir bar Rubber septum Balloon 50-µL syringe TLC plates (Merck silica gel 60, 4 × 10–cm) 254-nm UV lamp 5-cm-diameter chromatography column Rotary evaporator with a vacuum pump or water aspirator Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E)
Synthesis of RNA Sequences with 2′-O-TOM-protecte d Ribonucleoside Phosphoramidites
3.8.2 Supplement 7
Current Protocols in Nucleic Acid Chemistry
Prepare phosphoramidites 1. In a 250-mL one-neck flask equipped with a stir bar, a balloon filled with nitrogen or argon gas, and a rubber septum, dissolve 20 mmol of each dry N-Ac-5′-O-DMTr2′-O-TOM-ribonucleoside (S.1a-d) in 70 mL dry dichloromethane. 2. While stirring, add 8.1 mL (50 mmol) N-ethyl-N,N-diisopropylamine and then slowly add 5.4 mL (25 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite. 3. Stir the clear reaction mixture overnight at room temperature. 4. Take ∼50 µL of the clear reaction mixture with a syringe and dilute it with 0.5 mL dichloromethane in a small tube. Analyze by TLC (APPENDIX 3D) using 1:4 (v/v) hexane/ethyl acetate for A, C, and U, and 19:1 (v/v) dichloromethane/methanol for G. Co-spot the starting material for comparison. Visualize by exposure to a 254-nm UV lamp and stain with anisaldehyde reagent. The product typically migrates slightly faster than the starting material. In this reaction, two epimeric products are formed; consequently, the two pyrimidines show two distinct product spots. Addition of triethylamine to solvents is not necessary at this step.
5. Optional: If the reaction is not nearly complete (>5% starting material remaining), add an additional portion (∼0.2 eq) of 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite, stir 2 hr at room temperature, and repeat TLC analysis. Try to limit the amount of phosphitylating agent as much as possible. If too much phosphitylating agent is used, it might be difficult to remove hydrolysis products (H-phosphonates) from the ribonucleoside phosphoramidites.
Isolate phosphoramidites 6. Prepare a slurry of 400 g silica gel (230 to 400 mesh) in 4:1 (v/v) hexane/ethyl acetate containing 3% triethylamine for A, C, and U, and in 1:1 (v/v) hexane/ethyl acetate containing 3% triethylamine for G. For detailed steps for column chromatography, see APPENDIX 3E.
7. Pour into a 5-cm-diameter chromatography column and add a layer of ∼3 cm sand on top of the column. 8. Concentrate the reaction mixture (step 3) to a volume of ∼40 mL in a rotary evaporator with a vacuum pump. 9. Load the reaction mixture carefully on top of the column and start the elution slowly using the same solvent mixture used to prepare the slurry. Continue until the dichloromethane (clearly visible as a transparent plug) has been eluted. 10. Continue eluting with gradually more polar solvent and collect 100-mL fractions. For A, C, and U, use 1 L each of hexane/ethyl acetate at 7:3, 6:4, 5:5, and so on. For G, use 1 L each of hexane/ethyl acetate at 1:1 and 1:3, then 100% ethyl acetate, and then add stepwise 1% methanol to the ethyl acetate. Always add 3% triethylamine to solvents.
11. Monitor fractions by TLC and pool fractions that contain pure 2′-O-TOM-substituted phosphoramidites. 12. Repeat column chromatography with the impure fractions on an appropriate smaller scale and combine fractions that contain pure products.
Synthesis of Unmodified Oligonucleotides
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13. Evaporate to dryness in a rotary evaporator and dry overnight at high vacuum (<0.05 mbar). Typically, ∼90% pure 2′-O-TOM-protected phosphoramidites are obtained as a colorless, solid foam.
14. Check the purity of the material by 1H- and 31P-NMR. Only pure products should be employed for RNA synthesis (see Basic Protocol 2). Small amounts of an H-phosphonate species (hydrolysis product of the phosphitylation agent) with a characteristic signal around δ = 10 ppm (31P-NMR) can be tolerated. In pure and dry form, the products can be stored at −20°C for years without decomposition. N6-Ac-5′-O-DMTr-2′-O-TOM-adenosine 3′-(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.2a): 17.9 g (90%). TLC (hexane/ethyl acetate 3:7) 0.75; λmax (acetonitrile): 270 (21,800), 235 (25,400); δH (CDCl3) 0.88-0.90 (21 H, m, i-Pr3Si), 1.09 and 1.18 and 1.19 (12 H, 3d, J = 6.8 Hz, Me), 2.38 (1 H, t, J = 6.4 Hz, CH2), 2.62 (3 H, s, CH3CO), 2.65 (1 H, dt, J = 3.3, 6.6 Hz, CH2), 3.35 (1 H, ddd, J = 4.6, 7.5 Hz, 11.7, CH2), 3.51-3.75 (5 H, m, CH2, CH, H-5′), 3.77 and 3.78 (6 H, 2s, 2 OCH3), 3.85-3.97 (1 H, m, CH2), 4.36 and 4.41 (2q, J = 4.0 Hz, H-4′), 4.68 (m, H-3′), 4.92 and 4.95 and 4.99 (2 H, 3d, J = 5.0 Hz, OCH2O), 5.17 and 5.20 (2t, J = 5.5 Hz, H-2′), 6.17 and 6.20 (2d, J = 5.8 Hz, H-1′), 6.76-6.80 (4 H, m, DMTr), 7.18-7.42 (9 H, m, DMTr), 8.12 and 8.15 (2s, H-2), 8.53 (br. s, HN), 8.54 and 8.56 (2s, H-8); δC (CDCl3) 11.8 (d), 17.5 (q), 17.6 (q), 17.7 (q), 20.1 and 20.3 (2t, JCP 7 Hz), 24.52 (q), 24.56 (q), 24.58 (q), 24.62 (q), 25.6 (q), 43.2 and 43.4 (2d, JCP 13 Hz), 55.1 (q), 55.2 (q), 58.0 and 58.9 (2t, JCP 18 Hz), 62.8 (t), 63.2 (t), 71.2 and 71.8, (2d, JCP 16 Hz), 77.0 and 77.6 (2d, JCP 4 Hz), 84.1 (d), 84.2 (d), 86.5 (s), 86.6 (s), 87.4 (d), 87.5 (d), 89.2 (t), 89.5 (t), 113.0 (d), 113.1 (d), 117.3 (s), 117.6 (s), 122.2 (s), 126.8 (d), 126.9 (d), 127.8 (d), 128.1 (d), 128.2 (d), 128.3 (d), 130.0 (d), 130.1 (d), 130.2 (d), 135.6 (s), 135.7 (s), 135.8 (s), 142.1 (d), 142.2 (d), 144.4 (s), 144.5 (s), 149.0 (s), 149.1 (s), 151.0 (s), 151.1 (s), 152.2 (d), 158.5 (s), 158.6 (s), 170.3 (s); δP (CDCl3) 150.8, 151.5; m/z 998 (30, MH+), 821 (63), 303 (100). Anal. calcd. for C52H72N7O9SiP: C, 62.57; H, 7.27; N, 9.82. Found: C, 62.62; H, 7.25; N, 9.72. N2-Ac-5′-O-DMTr-2′-O-TOM-guanosine 3′-(2-cyanoethyl-N,N-diisopropylphosphoramidite] (S.2b): 19.0 g (94%). TLC (CH2Cl2/methanol 19:1) 0.60; λmax (acetonitrile): 276 (13,500), 250sh (19,700), 237 (24,600); δH (CDCl3) 0.91-0.94 (21 H, m, Pri3Si), 1.02-1.19 (12 H, 5d, J = 7 Hz, Me), 1.73 and 1.82 (2s, COCH3), 2.25 and 2.75 (2m, 2 CH2), 3.22 (0.5 H, dd, J = 3.7, 10.7 Hz, H-5′), 3.28 (0.5 H, dd, J = 5.3, 10.6 Hz, H-5′), 3.47-3.62 (3.5 H, m, CH, H-5′, CH2), 3.68 (0.5H, m, CH2), 3.75 and 3.76 and 3.77 and 3.78 (4s, 4 OCH3), 3.92 and 4.04 (1.5 H, 2m, CH2), 4.23 (0.5 H, q, J = 2.5 Hz, H-4′), 4.32 (0.5 H, br. dt, J ≈ 5, 2 Hz, H-4′), 4.52 (0.5 H, ddd, J = 2.0, 4.8, 12.1 Hz, H-3′), 4.62 (0.5 H, ddd, J = 4.8, 5.8, 10.6 Hz, H-3′), 4.91 (1 H, s, OCH2O), 4.90 and 5.00 (1 H, 2d, J = 5.2 Hz, OCH2O), 5.02 (0.5H, dd, J = 4.8, 7.4 Hz, H-2′), 5.05 (0.5 H, t, J = 5.8 Hz, H-2′), 5.87 (0.5 H, d, J = 5.7 Hz, H-1′), 5.97 (0.5 H, d, J = 7.4 Hz, H-1′), 6.76-6.82 (4 H, m, DMTr), 7.14-7.52 (9 H, m, DMTr), 7.74 and 7.80 (1 H, 2s, H-8), 8.29 and 8.57 (1 H, 2br. s, HN), 11.89 (1 H, br. s, HN); δC (CDCl3) 11.8 (d), 17.6 (q), 17.7 (q), 17.8 (q), 20.1 (t, JCP 3 Hz), 20.2 (t), 23.5 (q), 23.6 (q), 24.5 (q), 24.6 (q), 24.7 (q), 43.1 and 43.3 (2d, JCP 13 Hz), 55.2 (q), 55.3 (q), 56.9 (t, JCP 19 Hz), 58.8 (t, JCP 13 Hz), 63.5 (t), 63.9 (t), 70.7 (d, JCP 17 Hz), 71.7 (d, JCP 14 Hz), 76.9 (d), 78.3 (d), 84.2 (d), 84.3 (d, JCP 4 Hz), 86.2 (d), 86.3 (s), 86.7 (s), 88.9 (d), 89.4 (t), 89.5 (t), 113.1 (d), 113.2 (d), 117.5 (s), 118.1 (s), 122.0 (s), 122.7 (s), 127.0 (d), 127.1 (d), 127.9 (d), 128.0 (d), 128.1 (d), 130.0 (d), 130.1 (d), 130.2 (d), 135.6 (s), 135.8 (s), 136.0 (s), 136.3 (s), 137.7 (d), 139.1 (d), 144.6 (s), 145.0 (s), 146.8 (s), 147.1 (s), 148.0 (s), 148.5 (s), 155.6 (s), 158.6 (s), 158.7 (s), 171.5 (s), 171.6 (s); δP (CDCl3) 149.9, 150.5; m/z 1014 (62, MH+), 303 (100). Anal. calcd. for C52H72N7O10SiP: C, 61.58; H, 7.15; N, 9.67. Found C, 61.22; H, 7.19; N, 9.55.
Synthesis of RNA Sequences with 2′-O-TOM-protecte d Ribonucleoside Phosphoramidites
N4-Ac-5′-O-DMTr-2′-O-TOM-cytidine 3′-(2-cyanoethyl-N,N-diisopropylphosphoramidite] (S.2c): 18.1 g (93%). TLC (hexane/ethyl acetate 3:7) 0.75; λmax (acetonitrile) 305 (6,700), 279 (6,200), 237 (29,000); δH (CDCl3) 0.87-1.11 (21 H, m, i-Pr3Si), 1.13 and 1.16 (2d, J = 6.8, 4 Hz, Me), 2.22 and 2.33 (2s, CH3CO), 2.38 and 2.59 (2 H, 2t, J = 6.5 Hz, CH2), 3.42-3.68 (5 H, m, CH2, CH, H-5′), 3.80 and 3.81 and 3.814 and 3.82 (6 H, 4s, OCH3), 3.91 (1 H, m, CH2), 4.28-4.41 (2 H, m, H-2′,4′), 4.50 (ddd, J = 4.8, 8.0, 9.9 Hz, H-3′),
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5.15-5.22 (2 H, m, OCH2O), 6.15 (0.5 H, d, J = 1.8 Hz, H-1′), 6.16 (0.5 H, d, J = 2.4 Hz, H-1′), 6.82-6.88 (4 H, m, DMTr), 6.96 and 7.03 (2d, J = 7.4 Hz, H-5), 7.24-7.47 (9 H, m, DMTr), 8.36 and 8.45 (2d, J = 7.4 Hz, H-6), 9.52 and 9.60 (2br. s, HN); δC (CDCl3) 12.0 (d), 17.8 (q), 17.9 (q), 20.2 and 20.3 (2t, JCP 7 Hz), 24.48 (q), 24.54 (q), 24.58 (q), 24.61 (q), 24.64 (q), 24.7 (q), 24.9 (q), 43.2 and 43.3 (2d, JCP 7 Hz), 55.2 (q), 55.3 (q), 58.2 and 58.7 (2t, JCP 20 Hz), 60.9 (t), 61.6 (t), 69.4 and 69.7, (2d, JCP 14 Hz), 78.7 (d, JCP 3 Hz), 78.9 (d), 82.3 (d, JCP 3 Hz), 82.5 (d, JCP 5 Hz), 87.0 (s), 87.2 (s), 89.6 (d), 89.7 (d), 89.8 (t), 96.5 (d), 113.2 (d), 113.3 (d), 117.4 (s), 117.7 (s), 127.2 (d), 128.0 (d), 128.2 (d), 128.4 (d), 130.1 (d), 130.2 (d), 130.3 (d), 130.4 (d), 135.2 (s), 135.3 (s), 135.4 (s), 135.5 (s), 144.1 (s), 144.3 (s), 145.0 (d), 145.1 (d), 154.9 (s), 158.7 (s), 162.6 (s), 162.7 (s), 170.5 (s); δP (CDCl3) 150.6, 150.9; m/z 974 (21, MH+), 303 (100). Anal. calcd. for C51H72N5O10SiP: C, 62.88; H, 7.45; N, 7.19. Found: C, 62.74; H, 7.56; N, 7.00. 5′-O-DMTr-2′- O-TOM-u ridine 3′-(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.2d): 17.1 g (92%). TLC (hexane/ethyl acetate 1:1) 0.65; λmax (acetonitrile) 264 (12,100), 236 (23,700); δH (CDCl3) 1.02-1.05 (21 H, m, i-Pr3Si), 1.06 and 1.17 (12 H, m, Me), 2.39 and 2.63 (2t, J = 6.7 Hz, CH2), 3.39 (m, H-5′), 3.53-3.70 (4 H, m, CH, H′-5′, CH2), 3.79 and 3.80 (6 H, 2 OCH3), 3.81-3.96 (m, CH2), 4.19 and 4.27 (2br. dt, J ≈ 5, 2 Hz, H-4′), 4.40-4.55 (m, H-2′,3′), 4.96-5.06 (2 H, m, OCH2O), 5.32 and 5.36 (2d, J = 8.1 Hz, H-5), 6.12 (0.5 H, d, J = 5.1 Hz, H-1′), 6.14 (0.5 H, d, J = 4.9 Hz, H-1′), 6.82-6.85 (4 H, m, DMTr), 7.05-7.47 (9 H, m, DMTr), 7.80 and 7.86 (2d, J = 8.1 Hz, H-6), 8.33 (br. s, HN); δC (CDCl3) 11.9 (d) 12.0 (d), 17.7 (q), 17.8 (q), 17.9 (q), 20.2 and 20.4 (2t, JCP 7 Hz), 24.52 (q), 24.55 (q), 24.58 (q), 24.62 (q), 43.2 and 43.4 (2d, JCP 12 Hz), 55.2 (q), 55.3 (q), 57.8 and 58.9 (2t, JCP 18 Hz), 62.1 and 62.5 (2t), 70.4 and 70.9, (2d, JCP 15 Hz), 77.4 and 78.1 (2d, JCP 4 Hz), 83.3 (d, JCP 4 Hz), 83.5 (d), 87.1 (s), 87.2 (s), 87.4 (d), 89.0 (d), 89.3 (t), 89.4 (t), 102.3 (d), 102.4 (d), 113.1 (d), 113.2 (d), 113.3 (d), 117.3 (s), 117.7 (s), 127.2 (d), 127.9 (d), 128.0 (d), 128.2 (d), 128.3 (d), 130.2 (d), 130.3 (d), 130.4 (d), 135.1 (s), 135.3 (s), 135.4 (s), 135.5 (s), 140.3 (d), 140.4 (d), 144.2 (s), 144.3 (s), 150.1 (s), 158.7 (s), 162.8 (s), 162.9 (s); δP (CDCl3) 150.9, 151.3; m/z 933 (57, MH+), 303 (100). Anal. calcd. for C49H69N4O10SiP: C, 63.07; H, 7.45; N, 6.00. Found: C, 62.84; H, 7.49; N, 5.98.
ASSEMBLY OF 2′-O-TOM-PROTECTED PHOSPHORAMIDITES ON DNA SYNTHESIZERS
BASIC PROTOCOL 2
This protocol describes the setup and steps required for automated assembly of RNA sequences from TOM-phosphoramidites on 1-µmol and 10-µmol scales. The protocol was developed for GeneAssemblers (Amersham Pharmacia Biotech), but can easily be adapted to other automated DNA synthesizers. The authors recommend coupling times of 2.5 to 3.5 min and 6 to 9 min for 1.0-µmol and 10-µmol syntheses, respectively. The TOM-phosphoramidites allow preparation of RNA sequences consisting of up to 100 nt; however, the authors recommend first preparing shorter sequences (e.g., 20 nt) to gain experience in the handling of the intermediate and product sequences. The reactions carried out in each cycle are shown in Figure 3.8.2. Any commercially available solid support containing the first immobilized ribonucleoside (or some modified ribonucleoside) can be employed. In the authors’ hands, controlledpore glass (CPG) supports give the best results, but polystyrene-based materials may be used with similar results. CPG supports with pore sizes of 500 and 1000 Å should be used for the synthesis of ≤47-mers and >47-mers, respectively. Materials 2′-O-TOM-phosphoramidites (0.1 M in acetonitrile; Glen Research; see Basic Protocol 1) Activator: 0.25 M 5-ethylthio-1H-tetrazole (SET, Glen Research) in acetonitrile Argon source Dry acetonitrile (<30 ppm water) 4A molecular sieves (optional)
Synthesis of Unmodified Oligonucleotides
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Figure 3.8.2 One coupling cycle—including detritylation, coupling, capping, and oxidation— required for the attachment of one nucleotide to the growing oligonucleotide chain. Additionally, after the last coupling cycle, the terminal dimethoxytrityl group is removed from the immobilized oligonucleotide. B, any N-protected nucleobase; Bz, benzoyl; CPG, controlled-pore glass; DMTr, 4,4′-dimethoxytrityl; TOM, [(triisopropylsilyl)oxy]methyl.
Detritylation solution (see recipe) Oxidation solution (see recipe) Capping solutions A and B (see recipes) 1,2-Dichloroethane (reagent grade) Solid support, e.g., long-chain alkylamine controlled-pore glass (CPG) supports (500 or 1000 Å) for RNA synthesis derivatized with 5′-O-4,4′-dimethoxytritylated ribonucleosides (Glen Research) Vials and bottles for attachment of the phosphoramidites and reagents to the synthesizer Automated DNA synthesizer (e.g., GeneAssembler; Amersham Pharmacia Biotech) Synthesis column for 1-µmol or 10-µmol synthesis Additional reagents and equipment for automated DNA synthesis (APPENDIX 3C) 1. Calculate the amount of phosphoramidites and activator (SET) required to assemble the desired sequence by multiplying the following by the number of nucleotides in the sequence, and adding an extra 50 mg phosphoramidites/50 mg SET to give enough material to purge the lines of the synthesizer. Synthesis of RNA Sequences with 2′-O-TOM-protecte d Ribonucleoside Phosphoramidites
1-µmol scale: 12 mg phosphoramidite, 12 mg SET/coupling 10-µmol scale: 40 mg phosphoramidite, 20 mg SET/coupling. 2. Place the calculated amounts of phosphoramidites and SET into the appropriate vials in an automated DNA synthesizer and flush with argon.
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3. Use a syringe to add the amount of dry acetonitrile required to obtain ∼0.1 M phosphoramidite (i.e., 1 mL/100 mg solid phosphoramidite) and ∼0.25 M SET (1 mL/32 mg solid SET). Allow solids to dissolve. When working with the GeneAssembler, beads of 4A molecular sieves may also be added.
4. Create methods for the assembly of the desired sequence. In principle, methods employed for synthesis of DNA sequences can be used here. The authors recommend the steps listed in Table 3.8.1.
5. Connect the following reagents to the synthesizer according to the manufacturer’s instructions: detritylation solution oxidation solution capping solutions A and B 1,2-dichloroethane dry acetonitrile phosphoramidite solution SET solution. 6. Calculate the required amount of solid support (e.g., CPG support) according to its individual loading (as reported by the manufacturer) and the synthesis scale. Use a CPG support with a pore size of 500 Å for the synthesis of ≤47-mers, and a support with a pore size of 1000 Å for >47-mers. Load an empty synthesis column with the RNA support and connect it to the synthesizer. For example, when the loading is 40 ìmol/g, use 25 mg and 250 mg for 1-ìmol and 10-ìmol syntheses, respectively.
7. Purge the lines of the synthesizer with all solutions and solvents. 8. Carry out the assembly according to manufacturer’s instructions. Choose the option “DMTr off” in the setup menu. After the tenth coupling cycle, coupling yields of >99% can be expected by detritylation assay. If lower yields are obtained, see Troubleshooting.
9. Remove the synthesis column from the synthesizer and dry it by vacuum or under a stream of argon. 10. Deprotect the assembled, still fully protected oligoribonucleotide (see Basic Protocol 3). DEPROTECTION OF RNA SEQUENCES ASSEMBLED FROM 2′-O-TOM-PROTECTED PHOSPHORAMIDITES
BASIC PROTOCOL 3
This protocol describes the two-step deprotection of RNA sequences assembled according to Basic Protocol 2. The two steps are presented schematically in Figure 3.8.3. In the first step, 10 M methylamine in 50% ethanol is used to detach the sequence from the solid support, eliminate the cyanoethyl groups, and remove the acetyl protecting groups from the nucleobases. In the second step, 1 M tetra-n-butylammonium fluoride in tetrahydrofuran is used to remove the 2′-O-TOM protecting groups. This reaction is quenched by addition of aqueous Tris⋅Cl buffer, pH 7.4. A final desalting procedure with commercially available size-exclusion cartridges yields the crude RNA sequence.
Synthesis of Unmodified Oligonucleotides
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Table 3.8.1 Methods for Oligoribonucleotide Assembly on 1.0-µmol and 10-µmol Scales with TOM-Phosphoramiditesa
1.0-µmol scale Time (min)
Step
Detritylation 0.00 DETRIT 1.50 VALVE POS 2.50 VALVE POS 2.70 INTEGRATE 4.00 ML/MIN Coupling 4.00 LOOP TIMES 4.00 VALVE POS 4.05 ML/MIN 4.15 ML/MIN 4.15 VALVE POS 4.20 ML/MIN 4.30 ML/MIN 4.30 VALVE POS 4.35 ML/MIN 4.45 ML/MIN 4.45 END OF LOOP 4.45 VALVE POS 4.50 ML/MIN 4.60 ML/MIN 4.60 LOOP TIMES 4.60 STEP VALVE 4.60 END OF LOOP 4.65 ML/MIN 7.15 ML/MIN 7.15 STEP VALVE 7.20 ML/MIN 7.50 ML/MIN Capping 7.55 VALVE POS 7.60 ML/MIN 7.60 LOOP TIMES 7.60 VALVE POS 7.70 VALVE POS 7.80 END OF LOOP 7.80 ML/MIN 7.85 VALVE POS 7.85 ML/MIN Oxidation 8.15 VALVE POS 8.75 VALVE POS 10.25 ML/MIN 10.25 LOOP TIMES 10.25 STEP VALVE 10.25 END OF LOOP
10-µmol Valueb
2.1 2.3 0 0.00 2 1.8 0.90 0.00 1.X 0.60 0.00 1.8 0.90 0.00 1.1 1.00 0.00 7 3 2.50 0.00 3 2.50 0.00 2.5 1.00 4 2.5 2.6 0.00 2.3 2.50 2.4 2.3 0.00 7 3
Valueb Description
Time (min)
Step
0.00 3.00 4.00 5.00 8.00
10DETRIT VALVE POS VALVE POS INTEGRATE ML/MIN
8.00 8.00 8.05 8.15 8.15 8.20 8.30 8.30 8.35 8.45 8.45 8.45 8.50 9.00 9.00 9.00 9.00 9.05 16.05 16.05 16.10 17.10
LOOP TIMES VALVE POS ML/MIN ML/MIN VALVE POS ML/MIN ML/MIN VALVE POS ML/MIN ML/MIN END OF LOOP VALVE POS ML/MIN ML/MIN LOOP TIMES STEP VALVE END OF LOOP ML/MIN ML/MIN STEP VALVE ML/MIN ML/MIN
4 1.8 0.75 0.00 1.X 1.00 0.00 1.8 0.75 0.00
17.15 17.20 17.20 17.20 17.30 17.40 17.40 17.45 17.45
VALVE POS ML/MIN LOOP TIMES VALVE POS VALVE POS END OF LOOP ML/MIN VALVE POS ML/MIN
2.5 1.00 12 2.5 2.6
18.45 20.45 23.45 23.45 23.45 23.45
VALVE POS VALVE POS ML/MIN LOOP TIMES STEP VALVE END OF LOOP
2.4 2.3 0.00 7 3
2.1 2.3 0 0.00
Detritylation Dichloroethane wash Acetonitrile wash Trityl assay
Activator to column
Amidite to column
Activator to column
1.1 1.00 0.00 7 3 2.50 0.00 3 2.50 0.00
0.00 2.3 2.50
Coupling reaction
Acetonitrile wash
Capping A
Capping A Capping B
Acetonitrile wash
Oxidation Acetonitrile wash
Synthesis performed on a GeneAssembler (Amersham Pharmacia Biotech). For 1.0-µmol scale: synthesis: 120 µl phosphoramidite solution (12 eq) and 360 µl activator solution (90 eq); detritylation: 1.5 min; coupling: 2.5 min; capping: 0.8 min; oxidation: 0.3 min. For 10-µmol scale: synthesis: 400 µl phosphoramidite solution (4 eq) and 600 µl activator solution (15 eq); detritylation: 3.0 min; coupling: 7.0 min; capping: 2.4 min; oxidation: 1.0 min. b Description: VALVE POS 1.X: phosphoramidite (A, C, G, or U); 1.8: activator; 2.1: dichloroethane; 2.2: detritylation; 2.3: acetonitrile; 2.4: oxidation; 2.5: capping A; 2.6: capping B. a
Synthesis of RNA Sequences with 2′-O-TOM-protecte d Ribonucleoside Phosphoramidites
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Figure 3.8.3 Two-step deprotection of oligoribonucleotide sequences assembled from TOMphosphoramidites. B, any nucleobase; BAc, any N-acetylated nucleobase; CPG, controlled-pore glass; TBAF, tetra-n-butylammonium fluoride; TOM, [(triisopropylsilyl)oxy]methyl.
Materials TOM-protected oligoribonucleotide attached to solid support (see Basic Protocol 2), in synthesis cartridge 12 M aqueous methylamine 8 M methylamine in ethanol (Fluka) 50% and 100% (v/v) ethanol 1 M tetra-n-butylammonium fluoride trihydrate (TBAF.3H2O; Fluka) in tetrahydrofuran (THF) N-Methylpyrrolidone (NMP) or dimethylformamide (DMF; optional) 1 M Tris⋅Cl, pH 7.4, RNase free, sterile (Fluka, for molecular biology) 3 M sodium acetate (APPENDIX 2A) 1.5-, 2-, and 5-mL twist-top vials 35°C incubator or heating block Speedvac evaporator NAP-10 columns (Amersham Pharmacia Biotech) Additional reagents and equipment for oligoribonucleotide purification by HPLC (UNIT 10.5) or polyacrylamide gel electrophoresis (PAGE; UNIT 10.4)
Synthesis of Unmodified Oligonucleotides
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Remove acetyl and cyanoethyl groups and solid support 1. Remove solid support from the synthesis cartridge and transfer it into 1.5-mL twist-top vials. For syntheses performed on a 1-ìmol scale, a single 1.5-mL vial should suffice. For a 10-ìmol scale, divide the solid support into four portions and add each to a separate 1.5-mL vial.
2. Add 0.5 mL of 12 M aqueous methylamine and 0.5 mL of 8 M methylamine in ethanol to each 1.5-mL vial. 3. Incubate 6 hr at 35°C with periodic shaking. Sequences with <60 nt can be incubated 6 hr at room temperature.
4. Centrifuge 3 min at 10,000 × g, 25°C, and transfer supernatant from each 1.5-mL vial into a 2-mL twist-top vial. 5. Wash solid support two times with 0.3 mL of 50% ethanol and place the washing solutions in the same 2-mL vial. 6. Evaporate to dryness in a Speedvac evaporator. Remove TOM groups 7. To each 2-mL vial, add 1 mL of 1 M TBAF/THF and dissolve the crude intermediate sequence by vortexing or ultrasonicating. If intermediates are not dissolved after 10 min, add a few drops of NMP or DMF. The intermediates usually dissolve almost immediately. Short adenosine-rich RNA sequences or RNA hybrid sequences containing a majority of non-RNA nucleosides (e.g., DNA/2′-OMe-RNA-nucleosides) must be deprotected with 1 M TBAF in 1:1 THF/NMP.
8. Incubate the solution overnight at 35°C with periodic shaking. Sequences with <60 nt can be incubated overnight at room temperature.
9. Quench the reaction by adding 1 mL of 1 M Tris⋅Cl solution, pH 7.4. IMPORTANT NOTE: Deprotected oligoribonucleotides are highly sensitive to nuclease degradation. Therefore, gloves should always be worn when manipulating deprotected RNA. Sterile vials and pipets, nuclease-free reagents, and UV-treated sterile water should always be used to limit potential exposure to nucleases. Work quickly and store the oligoribonucleotides at –20°C.
10. Evaporate to a volume of 1 mL in a Speedvac evaporator. IMPORTANT NOTE: This evaporation takes 20 to 40 min (depending on the setup). Do not overevaporate to dryness. Check the remaining volume frequently.
Desalt oligoribonucleotide 11. Wash a NAP-10 column with 10 mL sterile water. 12. Place the oligoribonucleotide solution on top of the column and allow the solution to sink in slowly by gravity. 13. Place 1.5 mL sterile water on top of the column and collect the eluate (1.5 mL) in a 5-mL vial. Synthesis of RNA Sequences with 2′-O-TOM-protecte d Ribonucleoside Phosphoramidites
This step is required to remove excess TBAF. Alternatively, desalting can be carried out by ion-exchange chromatography according to the protocol in UNIT 3.6. If the oligoribonucleotide will be used in its crude form, it should first be precipitated (steps 14 to 16). If it will be purified before use (step 17), precipitation is not required.
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Precipitate (optional) 14. Evaporate eluate to dryness in a Speedvac evaporator and dissolve the residue in 0.5 mL sterile water. 15. Add 50 µl of 3 M sodium acetate solution followed by 3.5 mL of 100% ethanol. Store overnight at −20°C. 16. Centrifuge 3 min at 10,000 × g, 25°C, and carefully remove the supernatant. The precipitate consists of crude oligoribonucleotide in its sodium form.
Purify (optional) 17. Purify the oligoribonucleotide by HPLC (UNIT 10.5) or polyacrylamide gel electrophoresis (UNIT 10.4). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Anisaldehyde reagent In a clean container, mix 10 mL anisaldehyde with 180 mL 100% ethanol. Add slowly, while stirring, 10 mL concentrated sulfuric acid, followed by 2 mL acetic acid. Store reagent up to 3 months in the dark at 25°C; avoid contamination with acetone. For staining, TLC plates are dipped into this mixture and heated with a heat-gun until dark spots, corresponding to the reaction products, appear on a pink to red background.
Capping solution A 100 mL acetic anhydride 800 mL tetrahydrofuran (THF) 100 mL 2,6-lutidine Store in a well-sealed bottle in a dark and cool (25°C) environment for up to 2 months Capping solution B 160 mL N-methylimidazole 840 mL tetrahydrofurant (THF) Store in a well-sealed bottle in a dark and cool (25°C) environment for up to 2 months Detritylation solution 40 mL dichloroacetic acid 960 mL dry 1,2-dichloroethane Store in a well-sealed bottle in a dark and cool (25°C) environment for up to 2 months Oxidation solution 12.7 g iodine 700 mL tetrahydrofuran (THF) 100 mL pyridine 200 mL water Dissolve iodine in THF, then add pyridine followed by water. Store in a well-sealed bottle in a dark and cool (25°C) environment for up to 2 months. Synthesis of Unmodified Oligonucleotides
3.8.11 Current Protocols in Nucleic Acid Chemistry
Supplement 7
COMMENTARY Background Information
Synthesis of RNA Sequences with 2′-O-TOM-protecte d Ribonucleoside Phosphoramidites
During the last decade, many research groups have contributed to the development of a reliable chemical synthesis of nucleic acids. The synthesis of oligodeoxyribonucleotides by the phosphoramidite method (UNITS 3.3, 3.5 & 3.6) is probably the most evolved chemical process known thus far, and has almost reached perfection in terms of efficiency and automation. This extremely powerful methodology can, in principle, also be applied to the synthesis of the structurally very similar RNA oligonucleotides. Compared to DNA, however, each nucleotide within an RNA strand contains an additional 2′-OH group, which is responsible for the instability of RNA under basic conditions (pH >12 at 25°C). This hydroxyl group must be protected during oligoribonucleotide assembly. Since the RNA products are base labile, the removal of these supplementary protecting groups is carried out separately, after removal of the 2-cyanoethyl groups and nucleobase-protecting groups and cleavage of the oligoribonucleotide from the support under basic nucleophilic conditions. As a consequence, the removal of these additional 2′-O-protecting groups must be completely orthogonal to all other acid-promoted and base-promoted deprotection conditions. The large number of reported 2′-O ribonucleoside-protecting groups can essentially be divided into acid-labile (Griffin and Reese, 1964; Reese et al., 1991; Scaringe et al., 1998), photo-labile (Ohtsuka et al., 1974; Schwarz et al., 1992; Pitsch, 1997; Pitsch et al., 1999a), and fluoride-labile groups (Ogilvie et al., 1974; Usman et al., 1985; Pitsch et al., 1999a, b). Among them, the fluoride-labile tert-butyldimethylsilyl (TBDMS) group (Ogilvie et al., 1974) has the widest application. This is well reviewed in Beaucage and Caruthers (1996) and in UNIT 3.6. However, several factors—including the relatively sluggish coupling observed with RNA phosphoramidites as compared to the corresponding DNA phosphoramidites—has limited the length of routinely synthesized RNA sequences (UNIT 3.6). A very attractive alternative that leads to better coupling yields under shorter coupling times was the photolabile 2-nitrobenzyloxymethyl (NBOM) group (Schwartz et al., 1992; UNIT 3.7). In a formal sense, the 2′-O-TOM protecting group represents a combination of the TBDMS and the NBOM protecting groups. It also displays some unique properties that render it a
very valuable 2′-O-protecting group for the chemical synthesis of oligoribonucleotides. It is completely stable under all reaction conditions required for assembly and deprotection of RNA sequences. Its excellent stability towards both strongly acidic conditions (employed during detritylation) and strongly basic conditions (employed for deprotection of the nucleobase) is a consequence of the sterically very hindered triisopropylsilyl group. In contrast, the TOM group is very labile towards TBAF and is completely removed even in the presence of up to 20% water (Pitsch et al., 2001). Complete removal of the TBDMS group, in contrast, occurs only in the absence of water and requires drying of the TBAF solution with molecular sieves (Hogrefe et al., 1993). Many interesting RNA sequences, such as tRNAs, ribozymes, and aptamers, consist of 60 to 80 nt. In order to obtain a reasonable overall product yield of ∼50%, an individual coupling yield of ∼99.3% must be achieved for each cycle in the assembly. Lower coupling yields result not only in small product yields, but also in complex crude products that are often very difficult to purify to homogeneity. The average coupling yields obtained with 2′-O-TOM phosphoramidites are generally >99% and, therefore, even relatively long RNA sequences (>70 nt) can be prepared routinely. Additionally, the short coupling times (<5 min) minimize side reactions during the coupling reaction. In addition to high coupling yields, a reliable deprotection scheme is crucial for the efficient synthesis of longer sequences. The combination of 2′-O-TOM and N-acetyl protecting groups allows, for both deprotection processes, reaction times that correspond to >100 individual half-lives (with respect to the removal of one protecting group) without destruction of the oligonucleotide product. Therefore, it is possible to carry out the deprotection reactions even for extremely prolonged periods of time, assuring complete deprotection without risking concomitant destruction of the product (Pitsch et al., 1999a, 2001). The chemical synthesis of oligonucleotides allows the more or less unrestricted incorporation of nucleobase, sugar, and backbone modifications, the preparation of hybrid sequences, and labeling with specific reporter groups. In this context, a great number of useful modified building blocks have been developed and are commercially available. By adapting the presented RNA chemistry to the established DNA
3.8.12 Supplement 7
Current Protocols in Nucleic Acid Chemistry
and RNA chemistry, the 2′-O-TOM-protected building blocks can be combined with all of these modifications and even with TBDMSprotected RNA phosphoramidites.
Critical Parameters Phosphoramidite chemistry is extremely water-sensitive. The phosphoramidites and the activator should be completely dry. Only acetonitrile that contains <30 ppm water should be used as solvent for the coupling reaction or to dissolve the phosphoramidites and the activator. If working with the GeneAssembler from Amersham Pharmacia Biotech, molecular sieves may be added to the corresponding vials and bottles. The synthesizer should be in good working condition. All lines should be washed after completion of each synthesis round. To prevent particles from reaching the valves, dust-free bottles, high-quality solvents and reagents, and appropriate filters should be used. Reagents should be replaced periodically with fresh supplies and should be stored in well-sealed bottles in a dark, cool environment. Only sterile, RNase-free solvents, reagents, containers, vials, pipets, and other equipment should be used. Any contact between the oligoribonucleotides and body fluids (e.g., sweat, saliva) should be avoided. In all aqueous solutions and buffers, RNase-producing microorganisms may live and grow. Keep such solutions as sterile as possible and replace them often. Good coupling yields should be obtained using the presented coupling conditions. Coupling times should not be significantly increased. Specifically, do not use the coupling conditions used for the assembly of 2′-OTBDMS phosphoramidites in combination with SET as activator. The deprotection under the presented conditions is straightforward. A longer deprotection time can be used, but the temperature should not exceed 35°C. Methylamine solutions should be stored at 4°C to prevent loss of concentration. The bottles should be opened only for a very brief time and should be replaced periodically. The TBAF solution should not be dried with molecular sieves, since this may lead to its decomposition. Ensure that the sequence is dissolved during TBAF treatment. If it is not, add a few drops of N-methylpyrrolidone (NMP), dimethylformamide (DMF), or dimethylsulfoxide (DMSO). Always quench the second deprotection reaction with Tris⋅Cl buffer according to the presented protocol.
The crude product should not be isolated by direct precipitation from the TBAF/THF solution. It should always be desalted by chromatography on Sephadex columns, NAP columns (see Basic Protocol 3), or ion-exchange cartridges (UNIT 3.6). When the analysis of the crude products by HPLC or PAGE shows not one main peak (band), but a variety of peaks (or bands) or broad features, the product may exist in a variety of secondary structures. For complete denaturation, HPLC analysis can be performed at a higher temperature (with NucleoPac columns at neutral pH values, up to 90°C). Alternatively, 6 M urea can be added to HPLC and PAGE buffers.
Troubleshooting Low coupling yield. Several conditions can lead to coupling yields that are <99% after the tenth coupling. (1) The acetonitrile may contain water. Substitute it with acetonitrile of better quality; add molecular sieves. (2) The synthesizer may be malfunctioning. Determine whether it is working properly by performing a DNA synthesis; also check the flow rate. (3) The phosphoramidites may contain water. Dry them under high vacuum (<0.05 mbar) overnight in a desiccator containing KOH pellets. Multiple product peaks. Several conditions can also lead to the observation of several main product peaks or spots when the crude product is analyzed. (1) The sequence may exist in several secondary structures. Perform analysis at higher temperature and/or in the presence of 6 M urea. Isolate materials corresponding to the different features, heat them briefly at 95°C (at neutral pH), and reanalyze them separately. If the same pattern is observed, the sequence is not folding uniformly. (2) Deprotection may be incomplete. Use new deprotection solutions and reagents. Check (e.g., by carefully smelling) whether vials are sealed tightly so that methylamine can not evaporate. Check whether the intermediate product obtained after the first deprotection step completely dissolves in TBAF/THF. If it does not, add some polar solvent (NMP, DMF, DMSO). (3) The sequence was degraded by RNases. All materials and liquids that come into contact with the sequence must be free of contamination by RNases or microorganisms. Replace the Tris⋅Cl buffer. Use a different source of sterile water. Autoclave all vials, pipet, and other materials. Synthesis of Unmodified Oligonucleotides
3.8.13 Current Protocols in Nucleic Acid Chemistry
Supplement 7
Figure 3.8.4 Capillary gel electrophoresis analysis of the 34-mer RNA sequence r(GGCGACCCUGAUGAGGCCGAAAGGCCGAAACCGU) prepared at the 10-µmol scale according to Basic Protocols 2 and 3; (A) crude product; (B) product after HPLC purification. Chromatography performed with BioFocus 3000 (Bio-Rad), coated BioCap XL-column 75 µm × 40 cm, “run-buffer” containing 6 M urea, elution with 15 kV at 40°C, detection at 260 nm.
Anticipated Results With TOM-phosphoramidites, employing the methods presented here and observing the Critical Parameters, it should be possible to routinely obtain individual coupling yields >99% and complete deprotection. These properties allow routine preparation of oligoribonucleotides containing up to ∼80 nt. The purification efficiency with such long oligomers depends strongly on the individual structure, and
Synthesis of RNA Sequences with 2′-O-TOM-protecte d Ribonucleoside Phosphoramidites
therefore on the sequence. With up to ∼50 nt, HPLC purification with NucleoPac columns leads to very uniform products. Figure 3.8.4 shows the results from capillary gel electrophoresis analysis of a crude 34-mer and the HPLC-purified 34-mer (prepared on a 10-µmol scale). Oligoribonucleotides >50mers can be purified by PAGE; the purity of such oligoribonucleotide products usually exceeds 90%. In Figure 3.8.5, the gel analysis of
Figure 3.8.5 Polyacrylamide gel electrophoresis (PAGE) of five crude RNA sequences (sequence length indicated) prepared at the 1-µmol scale according to Basic Protocols 2 and 3. PAGE-conditions: 15% (w/v) acrylamide (29:1 acrylamide/bisacrylamide), 0.75 × 100 mm, 90 mM Tris-borate, pH 8.3, 7 M urea, 2 mM EDTA, elution at 200 V, stained with ethidium bromide.
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five crude products with sequence lengths ranging from 40 to 81 nt is presented (all products were synthesized on a 1.5-µmol scale). Depending on the purification method, an overall yield of 10% to 40% purified RNA (based on solid support) can be expected.
Time Considerations Preparation of TOM-phosphoramidites takes ∼1 day each. Depending on the scale and the sequence length, the typical time for the assembly of an RNA sequence is 5 to 12 hr. It takes ∼1.5 days to perform the deprotections and the final desalting according to Basic Protocol 3, although at least five deprotections can be carried out in parallel during this time. Purification of an RNA sequence typically requires 1 day (1-µmol synthesis scale).
Pitsch, S. 1997. An efficient synthesis of enantiomeric oligoribonucleotides from D-glucose. Helv. Chim. Acta 80:2286-2314. Pitsch, S., Weiss, P.A., Wu, X., Ackermann, D., and Honegger, T. 1999a. Fast and reliable automated synthesis of RNA and partially 2′-O-protected precursors (‘caged RNA’) based on two novel, orthogonal 2′-O-protecting groups. Helv. Chim. Acta 82:1753-1761. Pitsch, S., Weiss, P.A., and Jenny, L. Nov. 1999b. Ribonucleoside-derivative and method for preparing the same. US Patent 5,986,084. Pitsch, S., Weiss, P.A., Jenny, L., Stutz, A., and Wu, X. 2001. Reliable chemical synthesis of oligoribonucleotides (RNA) with 2′-O-[(triisopropylsilyl)oxy]methyl (2′-O-TOM) protected phosphoramidites. Helv. Chim. Acta In press.
Literature Cited
Reese, C.B., Rao, M.V., Serafinowska, H.T., Thompson, E.A., and Yu, P.S. 1991. Studies in the solid phase synthesis of oligo- and poly-ribonucleotides. Nucleosides Nucleotides 10:8197.
Beaucage, S.L. and Caruthers, M.H. 1996. The chemical synthesis of DNA/RNA. In Bioorganic Chemistry: Nucleic Acids (S.M. Hecht, ed.) pp. 36-74. Oxford University Press, Oxford.
Scaringe, S.A., Wincott, F.E., and Caruthers, M.H. 1998. Novel RNA synthesis method using 5′-Osilyl-2′-O-orthoester protecting groups. J. Am. Chem. Soc. 120:11820-11821.
Griffin, B.E. and Reese, C.B. 1964. Oligoribonucleotide synthesis via 2′,5′-protected ribonucleoside derivatives. Tetrahedron Lett. 5:29252931.
Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., deBear, J.S., and Gough, G.R. 1992. Rapid synthesis of oligoribonucleotides using 2′-O-(ortho-nitrobenzyloxymethyl)-protec ted monomers. Bioorg. Med. Chem. Lett. 2:1019-1024.
Hogrefe, R.I., McCaffrey, A.P., Borozdine, L.U., McCampbell, E.S., and Vaghefi, M.M. 1993. Effect of excess water on the desilylation of oligoribonucleotides using tetrabutylammonium fluoride. Nucl. Acids Res. 21:4739-4741. Ogilvie, K.K., Sadana, K.L, Thompson, E.A., Quilliam, M.A., and Westmore, J.B. 1974. The use of silyl protecting groups in protecting the hydroxyl functions of ribonucleosides. Tetrahedron Lett. 15:2861-2863. Ohtsuka, E., Tanaka, S., and Ikehara, M. 1974. Ribooligonucleotide synthesis using a photosensitive o-nitrobenzyl protection at the 2′-hydroxyl group. Nucl. Acids Res. 1:1351-1357.
Usman, N., Pon, R.T., and Ogilvie, K.K. 1985. Preparation of ribonucleoside 3′-O-phosphoramidites and their application to the automated solid-phase synthesis of oligonucleotides. Tetrahedron Lett. 26:4567-4570.
Contributed by Stefan Pitsch Institut de Chimie Organique, EPFL Lausanne, Switzerland Patrick A. Weiss Xeragon AG Zürich, Switzerland
Synthesis of Unmodified Oligonucleotides
3.8.15 Current Protocols in Nucleic Acid Chemistry
Supplement 7
3-(N-tert-Butylcarboxamido)-1-propyl and 4-Oxopentyl Groups for Phosphate/Thiophosphate Protection in Oligodeoxyribonucleotide Synthesis
UNIT 3.9
This unit provides procedures for the preparation of deoxyribonucleoside phosphoramidites and appropriate phosphordiamidite precursors with P(III) protecting groups different than the standard 2-cyanoethyl group. Specifically, these phosphoramidites are functionalized with the 3-(N-tert-butylcarboxamido)-1-propyl or 4-oxopentyl groups. The usefulness of these novel deoxyribonucleoside phosphoramidites in the solid-phase synthesis of a 20-mer DNA oligonucleotide and its phosphorothioated analog will be demonstrated. It will also be shown that removal of the 3-(N-tert-butylcarboxamido)-1propyl phosphate/thiophosphate-protecting group from these oligonucleotides is rapidly effected under thermolytic conditions at neutral pH, whereas the 4-oxopentyl group is preferably removed by treatment with pressurized ammonia gas or concentrated ammonium hydroxide at ambient temperature. These detailed methods constitute an economical and alkylation-free approach to large-scale preparations of therapeutic oligonucleotides. PREPARATION OF OLIGODEOXYRIBONUCLEOTIDES USING 5′-O-(4,4′-DIMETHOXYTRITYL)-3′-O-(N,N-DIISOPROPYLAMINO)-[3-(N-tertBUTYLCARBOXAMIDO)-1-PROPYLOXY]PHOSPHINYL-2′-DEOXYRIBONUCLEOSIDES
BASIC PROTOCOL
The 3-(N-tert-butylcarboxamido)-1-propyl group for phosphate/thiophosphate protection in oligonucleotide synthesis is convenient in that it is removable under thermolytic conditions at a rate considerably faster than the 2-(N-formyl,N-methyl)aminoethyl group (Grajkowski et al., 2001; Wilk et al., 2002). Since these deprotection conditions produce alkylation-free oligodeoxyribonucleotides, they are recommended for large-scale production of therapeutic oligonucleotides. This protocol delineates a general method for the synthesis of deoxyribonucleoside phosphoramidites (S.3a-d) from N- and 5′-O-protected deoxyribonucleosides (S.1a-d) using N,N,N′,N′-tetraisopropyl-O-[3-(N-tert-butylcarboxamido)-1-propyl]phosphordiamidite (S.2; see Support Protocol) and 1H-tetrazole.
i -Pr2N DMTrO
O
B
P
i -Pr2N
H N
O
DMTrO 2
OH 1a b c d
O
O
1H -tetrazole/CH2Cl2
B = thymin-1-yl B = N 4-benzoylcytosin-1-yl B = N 6-benzoyladenin-9-yl B = N 2-isobutyrylguanin-9-yl
i -Pr2N
B
O P
O
O 3a-d
N H
Figure 3.9.1 Preparation of deoxyribonucleoside phosphoramidites S.3a-d. DMTr, 4,4′-dimethoxytrityl; i-Pr, isopropyl. Adapted from Wilk et al. (2002) with permission from the American Chemical Society.
Synthesis of Unmodified Oligonucleotides
Contributed by Andrzej Wilk, Marcin K. Chmielewski, Andrzej Grajkowski, Lawrence R. Phillips, and Serge L. Beaucage
3.9.1
Current Protocols in Nucleic Acid Chemistry (2002) 3.9.1-3.9.16 Copyright © 2002 by John Wiley & Sons, Inc.
Supplement 11
This approach to the preparation of S.3a-d is illustrated in Figure 3.9.1. In addition, procedures are described for using these precursors for DNA synthesis, and for analyzing the resulting oligonucleotides. Materials Protected deoxyribonucleoside (S.1a-d; Chem-Impex International): 5′-O-(4′,4′-Dimethoxytrityl)-2′-deoxythymidine N4-Benzoyl-5′-O-(4,4′-dimethoxytrityl)-2′-deoxycytidine N6-Benzoyl-5′-O-(4,4′-dimethoxytrityl)-2′-deoxyadenosine N2-Isobutyryl-5′-O-(4,4′-dimethoxytrityl)-2′-deoxyguanosine Sublimed 1H-tetrazole (Aldrich) Argon gas Anhydrous methylene chloride (Aldrich) N,N,N′,N′-Tetraisopropyl-O-[3-(N-tert-butylcarboxamido)-1-propyl]phosphordiamidite (S.2; see Support Protocol) Benzene (Aldrich; optional) Triethylamine (Aldrich) Methylene chloride Hexane D2O (Aldrich) Silica gel (60 Å, 230 to 400 mesh; Merck) Anhydrous acetonitrile Reagents recommended for automated solid-phase oligonucleotide synthesis (PE Biosystems): Standard 2-cyanoethyl deoxyribonucleoside phosphoramidites (T, CBz, ABz, Gi-Bu) Activator solution (1H-tetrazole in acetonitrile) Oxidation solution (0.02 M iodine in tetrahydrofuran/pyridine/water) Cap A solution (acetic anhydride in tetrahydrofuran/pyridine) Cap B solution (1-methylimidazole in tetrahydrofuran) Deblocking solution (trichloroacetic acid in dichloromethane) 3H-1,2-Benzodithiol-3-one-1,1-dioxide (Glen Research) Ammonia gas (LB, Aldrich) 1× PBS, pH 7.2 (APPENDIX 2A) Loading buffer: 1:4 (v/v) 10 × TBE electrophoresis buffer (APPENDIX 2A) in formamide containing 2 mg/mL bromphenol blue Staining buffer: 1:5:20:0.1 (v/v/v/v) formamide/isopropyl alcohol/ddH2O/3.0 M Tris⋅Cl, pH 8.8 1 mg/mL Stains-All in formamide (both from Aldrich) 1.0 M Tris⋅Cl buffer, pH 9.0 (APPENDIX 2A) 1.0 M MgCl2 (Sigma) Snake venom phosphodiesterase (SVP, Crotallus durissus, Boehringer) Bacterial alkaline phosphatase (BAP, E. coli, Sigma) 0.1 M triethylammonium acetate buffer, pH 7.0 (2 M stock from Applied Biosystems)
3-(N-tert-Butylcarboxamido)-propyl and 4-Oxopentyl Protecting Groups
50-mL three-necked round-bottom flask with rubber septa High vacuum pump 15-mL powder-addition funnel 10-mL syringe 2.5 × 7.5–cm Whatman TLC plates precoated with a 250-µm layer of Diamond MK6F silica gel (60 Å) 2.5 × 20–cm disposable Flex chromatography columns (Kontes) DNA synthesizer (e.g., Applied Biosystems 380B) 200-mL pressure vessel container (Barrskogen)
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1-mL syringes with Luer tip 4-mL screw-capped glass vials Heating block (VWR), 90° ± 2°C UV spectrophotometer 1.5-mL microcentrifuge tubes 37°C water bath 5-µm Supelcosil LC-18S HPLC column (25 cm × 4.6 mm, Supelco) Additional reagents and equipment for polyacrylamide gel electrophoresis (PAGE; UNIT 10.4 and APPENDIX 3B), thin-layer chromatography (TLC; APPENDIX 3D), column chromatography (APPENDIX 3E), solid-phase oligonucleotide synthesis (APPENDIX 3C), SVP and BAP digestion (UNIT 10.6), and reversed-phase HPLC (RP-HPLC; UNIT 10.5) NOTE: The composition of DNA synthesis reagents (i.e., activator, oxidation, capping, and deblocking solutions) varies by manufacturer and by instrument. It is important to use the DNA synthesis reagents recommended for the specific synthesizer used, according to manufacturer’s instructions. Prepare deoxyribonucleoside phosphoramidites S.3a-d 1. Dry 2 mmol of suitably protected deoxyribonucleoside (S.1a-d) in a 50-mL threenecked round-bottom flask for 2 hr at 25°C under high vacuum. Attach a 15-mL powder-addition funnel containing 140 mg (2 mmol) sublimed 1H-tetrazole, and seal remaining necks with rubber septa. 2. Under an argon atmosphere, using a 10-mL syringe, add 10 mL anhydrous methylene chloride followed by 818 mg (2.1 mmol) N,N,N′,N′-tetraisopropyl-O-[3-(N-tertbutylcarboxamido)-1-propyl]phosphordiamidite (S.2) and stir with a magnetic stirrer and stir bar. 3. While still stirring, add the 1H-tetrazole in the addition funnel over a period of 1 hr. 4. Monitor the progress of the reaction by TLC (APPENDIX 3D) using 9:1 (v/v) benzene/triethylamine or 3:6:1 (v/v/v) methylene chloride/hexane/triethylamine as the eluent. Phosphinylation of suitably protected 2′-deoxynucleosides S.1a-c is usually complete within 3 hr at ambient temperature. For best results, phosphinylation of S.1d should be allowed to proceed for 12 hr at 25°C. Rf [9:1(v/v) benzene/triethylamine]: S.3a: 0.35, 0.43; S.3b: 0.47, 0.55; S.3c: 0.18, 0.29; S.3d: 0.10, 0.13. Rf [3:6:1(v/v/v) methylene chloride/hexane/triethylamine]: S.3a: 0.25; S.3b: 0.21; S.3c: 0.14; S.3d: 0.08.
5. Concentrate the reaction mixture to a foam using a rotary evaporator connected to a vacuum pump. 6. Suspend the crude product in 1 mL of 1:8:1 methylene chloride/hexane/triethylamine and apply the suspension to a 2.5 × 20–cm disposable Flex chromatography column containing ∼20 g silica gel that has been equilibrated in 1:8:1 methylene chloride/hexane/triethylamine (APPENDIX 3E). 7. Elute the column first with 100 mL of 10:80:10 (v/v/v) methylene chloride/hexane/triethylamine and collect 10-mL fractions. Using 100 mL of each concentration and collecting 10-mL fractions throughout, increase the concentration of methylene chloride in 5% increments until final ratios of 30:60:10 (S.3a), 50:40:10 (S.3b), 70:20:10 (S.3c), and 80:10:10 (S.3d) elute pure phosphoramidites. Analyze fractions by TLC (step 4), pool appropriate fractions, and concentrate under reduced pressure (∼20 min).
Synthesis of Unmodified Oligonucleotides
3.9.3 Current Protocols in Nucleic Acid Chemistry
Supplement 11
Each deoxyribonucleoside phosphoramidite S.3a-d is isolated as a white amorphous foam in yields ranging from 70% to 90%. Phosphoramidites can be stored up to several months at −20°C. 5′-O-(4,4′-Dimethoxytrityl)-3′-O-(N,N-diisopropylamino)-[3-(N-tert-butylcarboxamido)1-propyloxy]phosphinyl-2′-deoxythymidine (S.3a): 31P-NMR (121 MHz, C6D6): δ 148.3, 148.5. Fast atom bombardment/high-resolution mass spectrometry (FAB-HRMS): anal. calcd. for C45H61N4O9P (M+Na)+ 855.4074, found 855.4083. N4-Benzoyl-5′-O-(4,4′-dimethoxytrityl)-3′-O-(N,N-diisopropylamino)-[3-(N-tert-butylcarboxamido)-1-propyloxy]phosphinyl-2′-deoxycytidine (S.3b): 31P-NMR (121 MHz, C6D6): δ 146.3, 148.3. FAB-HRMS: anal. calcd. for C51H64N5O9P (M+Cs)+ 1054.3496, found 1054.3480. N6-Benzoyl-5′-O-(4,4′-dimethoxytrityl)-3′-O-(N,N-diisopropylamino)-[3-(N-tert-butylcarboxamido)-1-propyloxy]phosphinyl-2′-deoxyadenosine (S.3c): 31P-NMR (121 MHz, C6D6): δ 148.6, 148.7. FAB-HRMS: anal. calcd. for C52H64N7O8P (M+Na)+ 968.4453, found 968.4430. N2-Isobutyryl-5′-O-(4,4′-dimethoxytrityl)-3′-O-(N,N-diisopropylamino)-[3-(N-tert-butylcarboxamido)-1-propyloxy]phosphinyl-2′-deoxyguanosine (S.3d): 31P-NMR (121 MHz, C6D6): δ 148.2, 149.1. FAB-HRMS: anal. calcd. for C49H66N7O9P (M+Na)+ 950.4559, found 950.4560.
Synthesize oligonucleotides 8. Perform a 0.2-µmol-scale solid-phase synthesis of the desired oligonucleotide, or a 1-µmol-scale synthesis for its phosphorothioate analog, on an Applied Biosystems 380B DNA synthesizer (in trityl-off mode) according to the manufacturer’s recommendations (also see APPENDIX 3C). To compare coupling efficiency and the quality of the synthetic DNA produced using this protecting group, perform a parallel synthesis using standard 2-cyanoethyl deoxyribonucleoside phosphoramidites. Synthesis of d(ATCCGTAGCTAAGGTCATGC) is described in Wilk et al. (2002). 2-Cyanoethyl deoxyribonucleoside phosphoramidites and all the reagents pertaining to the automated preparation of oligonucleotides were purchased from Perkin-Elmer and used as recommended by the manufacturer. Like 2-cyanoethyl deoxyribonucleoside phosphoramidites, phosphoramidites S.3a-d are used as 0.1 M solutions in dry acetonitrile. The synthetic cycle used in the preparation of unmodified DNA oligonucleotides is slightly modified for the preparation of phosphorothioated DNA oligonucleotides in that the capping step is performed after the sulfurization reaction, which replaces the iodine oxidation step (see Iyer et al., 1990). The sulfurization reaction is effected by treatment with 0.05 M 3H-1,2-benzodithiol-3-one-1,1-dioxide (Glen Research) in acetonitrile using a wait step of 30 sec.
Deprotect oligonucleotide and release from support 9. Place the synthesis column in a 200-mL pressure vessel and fill the container with ammonia gas (∼10 bar, 25°C). Perform N-deprotection for 10 hr under these conditions. 10. Release gas from the pressure vessel and apply vacuum briefly (1 min) to the pressure vessel to remove traces of ammonia. 11. Using a 1-mL syringe with a Luer tip, carefully push back and forth 1.0 mL of 1× PBS, pH 7.2, through the synthesis column. Collect the solution in a 4-mL screwcapped glass vial. 3-(N-tert-Butylcarboxamido)-propyl and 4-Oxopentyl Protecting Groups
DNA oligonucleotides synthesized from commercial 2-cyanoethyl deoxyribonucleoside phosphoramidites are subjected to deprotection steps 9 though 11 only. Proceed to step 13.
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12. Place the capped vial containing the oligonucleotide solution in a heating block preheated to 90° ± 2°C and continue heating at this temperature for 30 min to remove the phosphate-protecting group. Cool to room temperature. 13. Determine the concentration of the oligonucleotide by UV spectrophotometry at 260 nm. Analyze oligonucleotide by polyacrylamide gel electrophoresis 14. Pipet 0.25 OD260 units of the oligonucleotide solution into a 1.5-mL microcentrifuge tube and evaporate to dryness using a stream of air.
A
B
Figure 3.9.2 Polyacrylamide gel electrophoresis analysis of d(ATCCGTAGCTAAGGTCATGC) under denaturing conditions (7 M urea, 1× TBE buffer, pH 8.3). (A) A crude 20-mer synthesized from commercial 2-cyanoethyl deoxyribonucleoside phosphoramidites and deprotected as described in the Basic Protocol, steps 9 through 11. (B) A crude 20-mer synthesized from S.3a-d and deprotected as described in steps 9 through 12. Bromphenol blue is used as a marker and shows as a large band in each lane at the bottom of the gel. Reprinted from Wilk et al. (2002) with permission from the American Chemical Society.
A
B
0
10
20 Retention time (min)
30
40
Figure 3.9.3 R P - HP LC analysis of the enzymatic hydrolysis of cr ude d(ATCCGTTGCTAAGGTCATGC) by snake venom phosphodiesterase and bacterial alkaline phosphatase. (A) Chromatogram of the digest obtained from the crude 20-mer that was synthesized from commercial 2-cyanoethyl deoxyribonucleoside phosphoramidites and deprotected as described in the Basic Protocol, steps 9 through 11. (B) Chromatogram of the digest obtained from the crude 20-mer that was synthesized from S.3a-d and deprotected as described in steps 9 through 12. Identities of the RP-HPLC peaks from left to right are dC, dG, dT, and dA when compared to authentic commercial samples. Reprinted from Wilk et al. (2002) with permission from the American Chemical Society.
Synthesis of Unmodified Oligonucleotides
3.9.5 Current Protocols in Nucleic Acid Chemistry
Supplement 11
15. Add 10 µL of loading buffer. Vortex well and then centrifuge 5 sec at 14,000 × g. 16. Load the sample in a 2-cm-wide well of a 20 × 40–cm, 7 M urea/20% polyacrylamide gel (UNIT 10.4 and APPENDIX 3B). Electrophorese at 350 V until the bromphenol blue dye travels 75% of the length of the gel. 17. Dismantle the gel apparatus and immerse the gel in 250 mL staining buffer containing 10 mL of 1 mg/mL Stains-All in formamide. Agitate the gel for ∼1 hr in the dark. 18. Discard the staining solution and rinse the gel three times with 250 mL distilled water. 19. Expose the gel to natural light until the purple background disappears and photograph the blue bands against a white background (see Fig. 3.9.2). Characterize oligonucleotide by enzymatic hydrolysis 20. Pipet 1 OD260 unit of crude oligonucleotide in a 1.5-mL microcentrifuge tube and evaporate to dryness using a stream of air. 21. Add 6 µL of 1.0 M Tris⋅Cl buffer, pH 9.0, 8 µL of 1.0 M MgCl2, and 75 µL water. Mix well. 22. Add 5 µL (0.015 U) SVP and 6 µL (0.7 U) BAP and heat 16 hr in a 37°C water bath (also see UNIT 10.6). 23. Heat deactivate the enzyme 3 min at 90°C. Centrifuge 5 min at 14,000 × g, 25°C. 24. Analyze a 50-µL aliquot of the digest by RP-HPLC (UNIT 10.5) using a 5-µm Supelcosil LC-18S column and a linear gradient of 1% acetonitrile/min, starting from 0.1 M triethylammonium acetate, pH 7.0, at a flow rate of 1 mL/min. An RP-HPLC profile of a digest is shown in Figure 3.9.3.
Analyze phosphorothioated oligonucleotides by 31P-NMR spectroscopy 25. Analyze phosphorothioated oligonucleotide (synthesized from S.3a-d and deprotected as in steps 9 through 12) in D2O by 31P-NMR spectroscopy. A 31P-NMR spectrum of the phosphorothioated oligonucleotide d(ATCCGTAGCTAAGGTCATGC) is shown in Figure 3.9.4. The crude oligomer is shown as a broad singlet at ∼55 ppm. Insignificant amounts of partially desulfurized 20-mer are detected near 0 ppm.
200 190 180 170 160 150 140 130 120 110 100 90
80 70 60 50 40 30 20 10
0 −10
ppm
3-(N-tert-Butylcarboxamido)-propyl and 4-Oxopentyl Protecting Groups
Figure 3.9.4 A 121-MHz 31P-NMR spectrum of d(APSTPSCPSCPSGPSTPSAPSGPSCPSTPSAPSAPS GPSGPSTPSCPSAPSTPSGPSC) in D2O. The crude 20-mer is prepared from S.3a-d and deprotected as described in the Basic Protocol, steps 9 through 12. Reprinted from Wilk et al. (2002) with permission from the American Chemical Society.
3.9.6 Supplement 11
Current Protocols in Nucleic Acid Chemistry
PREPARATION OF N,N,N′,N′-TETRAISOPROPYL-O-[3-(N-tert-BUTYLCARBOXAMIDO)-1-PROPYL]PHOSPHORDIAMIDITE This protocol describes the generation of bis(N,N-diisopropylamino)chlorophosphine (S.4) in situ upon mixing phosphorus trichloride and N,N-diisopropylamine, and its subsequent reaction with (N-tert-butyl)-4-hydroxybutyramide (S.5) to give the desired phosphordiamidite S.2 (see Fig. 3.9.5). The amidoalcohol S.5 is prepared from γ-butyrolactone (S.6) and tert-butylamine (Fig. 3.9.6).
SUPPORT PROTOCOL
Additional Materials (also see Basic Protocol) γ-Butyrolactone (S.6; Aldrich) tert-Butylamine (Aldrich) Benzene Phosphorus trichloride (Aldrich), freshly distilled N,N-Diisopropylamine (Aldrich), anhydrous 90°C water bath 100-mL round-bottom flask, oven dried 60-mL sintered glass funnel (coarse porosity) Prepare N-tert-butyl-4-hydroxybutyramide (S.5) 1. Place 8.60 g (100 mmol) γ-butyrolactone (S.6) in a 200-mL glass-lined pressure vessel. 2. Add 11.0 g (150 mmol) tert-butylamine and stir 3 days at 90°C. CAUTION: The above reaction involves heating highly irritating tert-butylamine above its boiling point. Ensure that the pressure vessel is sealed tightly and is placed in a well-ventilated chemical fume hood. Use moist pH indicator paper to check for leaks.
O HO
PCl3
i -Pr2N
i -Pr2NH
P
C6H6
Cl
N H 5
2
+ i -PrNH2 ⋅ HCl
i -Pr2N 4
Figure 3.9.5 Preparation of N,N,N′,N′-tetraisopropyl-O-[3-(N-tert-butylcarboxamido)-1-propyl]phosphordiamidite (S.2).
O
O O
t -BuNH2
HO
N H
90°C, 3 days 6
5
Figure 3.9.6 Preparation of N-tert-butyl-4-hydroxybutyramide (S.5). Adapted from Wilk et al. (2002) with permission from the American Chemical Society.
Synthesis of Unmodified Oligonucleotides
3.9.7 Current Protocols in Nucleic Acid Chemistry
Supplement 11
3. Cool the vessel to ambient temperature and release residual pressure in a well-ventilated chemical fume hood. 4. Apply vacuum to the pressure vessel to remove excess tert-butylamine, and keep the remaining material under reduced pressure (2 mmHg) for 1 hr, yielding a semi-crystalline mass. N-tert-Butyl-4-hydroxybutyramide (S.5) is recrystallized from chloroform/hexane (mp 73°C) and is isolated in near quantitative yield (15.5 g, 97.0 mmol). 1H-NMR (300 MHz, DMSO-d6): δ 1.23 (s, 9H), 1.59 (dt, J = 7.5, 6.6 Hz, 2H), 2.04 (t, J = 7.5 Hz, 2H), 3.34 (t, J = 6.6 Hz, 2H), 3.38 (b, 1H), 7.35 (b, 1H). 13C-NMR (75 MHz, DMSO-d6): δ 28.5, 28.7, 32.9, 49.8, 60.3, 171.8. Electrospray ionization mass spectrometry (ESI-MS; 70 eV): m/z (% relative abundance), 159 (M+, 4), 144 (7), 115 (31), 87 (11), 86 (14), 59 (19), 58 (100), 57 (29).
Prepare N,N,N′,N′-tetraisopropyl-O-[3-(N-tert-butylcarboxamido)-1-propyl]phosphordiamidite (S.2) 5. To an oven-dried 100-mL round-bottom flask containing 50 mL dry benzene under a dry argon atmosphere, add 876 µL (10 mmol) freshly distilled phosphorus trichloride by syringe and needle through a rubber septum. 6. Cool to 5°C by immersion in an ice bath. While stirring, add by syringe 8.8 mL (70 mmol) anhydrous N,N-diisopropylamine over a 30-min period under an argon atmosphere. 7. Remove the ice bath and allow the stirred reaction to warm to 25°C under a positive pressure of argon until the formation of bis(N,N-diisopropylamino) chlorophosphine (S.4) is complete. The rate of the reaction is monitored by 31P-NMR spectroscopy; after ∼3 to 4 days, complete conversion of (N,N-diisopropylamino) dichlorophosphine (singlet, 168.2 ppm in benzened6 downfield relative to a phosphoric acid external standard) to S.4 (singlet, 134.1 ppm) is achieved.
8. Add 1.9 g (12 mmol) S.5 and stir 2 hr at 25°C under a positive pressure of argon. The 31P-NMR spectrum of the reaction mixture in benzene-d6 shows a singlet at 123.2 ppm, indicating the formation of S.2 in ∼95% yield.
9. Filter the suspension through a 60-mL sintered glass funnel (coarse porosity) and wash the collected salt with 20 mL dry benzene. 10. Evaporate the filtrates under reduced pressure to afford 3.5 g (9 mmol, 90%) S.2 as an oil. Crude S.2 can be used for the phosphinylation of S.1a-d without further purification. It is, however, recommended to purify S.2 by silica gel chromatography using 9:1 (v/v) benzene/triethylamine as an eluent. This purification step allows S.2 to be isolated as a white crystalline solid (mp 71°-72°C). 1H-NMR (300 MHz, C6D6): δ 1.19 (d, J = 6.8 Hz, 12H), 1.22 (d, J = 6.8 Hz, 12H), 1.25 (s, 9H), 2.00 (m, 2H), 2.05 (m, 2H), 3.48 (sept, J = 6.8 Hz, 2H), 3.52 (sept, J = 6.8 Hz, 2H), 3.63 (dt, J = 6.0 Hz, 3JPH = 7.3 Hz, 2H). 13C-NMR (75 MHz, C6D6): δ 24.0, 24.1, 24.7, 24.8, 28.1 (d, 3JPC = 8.5 Hz), 28.8, 34.0, 44.6, 44.7, 50.7, 63.71 (d, 2JPC = 21.2 Hz), 171.1. 31P-NMR (121 MHz, C6D6): δ 123.2
3-(N-tert-Butylcarboxamido)-propyl and 4-Oxopentyl Protecting Groups
3.9.8 Supplement 11
Current Protocols in Nucleic Acid Chemistry
PREPARATION OF OLIGODEOXYRIBONUCLEOTIDES USING 5′-O-(4,4′-DIMETHOXYTRITYL)-3′-O-(N,N-DIISOPROPYLAMINO)-(4-OXO-1PENTYLOXY)PHOSPHINYL-2′-DEOXYRIBONUCLEOSIDES
ALTERNATE PROTOCOL
Like the 3-(N-tert-butylcarboxamido)-1-propyl group (see Basic Protocol), the 4-oxopentyl group for phosphate protection in DNA oligonucleotide synthesis is convenient because it can be removed under thermolytic conditions with comparable deprotection rates. This group, however, has the additional convenience of being labile under pressurized ammonia, methylamine gas, or concentrated ammonium hydroxide (Wilk et al., 2001). The latter conditions are especially recommended for the removal of the 4-oxopentyl group from oligonucleoside phosphorothioates. This protocol thus delineates a general method for the synthesis of deoxyribonucleoside phosphoramidites (S.8a-d) from the reaction of N- and 5′-O-protected deoxyribonucleosides (S.1a-d) with N,N,N′,N′-tetraisopropyl-O-(4-oxopentyl)phosphordiamidite (S.7) and 1H-tetrazole. This approach to the preparation of S.8a-d is illustrated in Figure 3.9.7. Synthesis and characterization of oligonucleotides follows the general procedures described in the Basic Protocol. Additional Materials (also see Basic Protocol and Support Protocol) 3-Acetyl-1-propanol (Aldrich) Prepare N,N,N′,N′-tetraisopropyl-O-(4-oxopentyl)phosphordiamidite S.7 1. Prepare S.7 (Fig. 3.9.7) from S.4 and 3-acetyl-1-propanol using the procedure described for synthesis of S.2 (see Support Protocol, steps 5 through 10). Perform on a scale that is two times that described for S.2. The 31P-NMR spectrum of S.7 shows a singlet at 123.5 ppm in C6D6. Like S.2, S.7 is used without further purification in the synthesis of the deoxyribonucleoside phosphoramidites S.8a-d.
Prepare deoxyribonucleoside phosphoramidites S.8a-d 2. Prepare phosphoramidites S.8a-d as described for S.3a-d (see Basic Protocol, steps 1 to 5), but reduce sublimed 1H-tetrazole to 120 mg (1.6 mmol) and use 980 µL (∼3 mmol) S.7 in place of S.2. As for S.3a-d, the coupling time for S.8d is 12 hr, compared to 3 hr for S.8a-c.
i -Pr2N DMTrO
O
B
P
i -Pr2N
CH3
O 7
OH 1a b c d
B = thymin-1-yl B = N 4-benzoylcytosin-1-yl B = N 6-benzoyladenin-9-yl B = N 2-isobutyrylguanin-9-yl
DMTrO
O
O
1H -tetrazole/CH2Cl2
i -Pr2N
B
O P
O
O
CH3 8a-d
Figure 3.9.7 Preparation of deoxyribonucleoside phosphoramidites S.8a-d using N,N,N′,N′tetraisopropyl-O-(4-oxopentyl)phosphordiamidite (S.7). DMTr, 4,4′-dimethoxytrityl; i-Pr, isopropyl. Adapted from Wilk et al. (2001) with permission from Elsevier Science.
Synthesis of Unmodified Oligonucleotides
3.9.9 Current Protocols in Nucleic Acid Chemistry
Supplement 11
Rf [9:1 (v/v) benzene/triethylamine]: S.8a: 0.37, 0.41; S.8b: 0.45, 0.54; S.8c: 0.16, 0.27; S.8d: 0.09, 0.12. Rf [3:6:1(v/v/v) methylene chloride/hexane/triethylamine]: S.8a: 0.22; S.8b: 0.18; S.8c: 0.12; S.8d: 0.05.
3. Perform column chromatography (see Basic Protocol, steps 6 and 7), but use the following final solvent ratios: 50:40:10 (S.8a), 60:30:10 (S.8b), 70:20:10 (S.8c), and 90:00:10 (S.8d) (v/v/v) methylene chloride/hexane/triethylamine. Each deoxyribonucleoside phosphoramidite S.8a-d is isolated as a white amorphous foam in yields ranging from 68% to 80%. 5′-O-(4,4′-Dimethoxytrityl)-3′-O-(N,N-diisopropylamino)-(4-oxo-1-pentyloxy)phosphinyl-2′-deoxythymidine (S.8a): 31P-NMR (121 MHz, C6D6 ): δ 148.4, 149.0. FAB-HRMS: anal. calcd. for C42H54N3O9P (M+Na)+ 798.3496, found 798.3541. N4-Benzoyl-5′-O-(4,4′-dimethoxytrityl)-3′-O-(N,N-diisopropylamino)-(4-oxo-1-pentyloxy)phosphinyl-2′-deoxycytidine (S.8b): 31P-NMR (121 MHz, C6D6 ): δ 148.9, 149.2. FAB-HRMS: anal. calcd. for C48H57N4O9P (M+Na)+ 887.3761, found 887.3682. N6-Benzoyl-5′-O-(4,4′-dimethoxytrityl)-3′-O-(N,N-diisopropylamino)-(4-oxo-1-pentyloxy)phosphinyl-2′-deoxyadenosine (S.8c): 31P-NMR (121 MHz, C6D6 ): δ 148.6, 148.8. FAB-HRMS: anal. calcd. for C49H57N6O8P (M+Na)+ 911.3873, found 911.3881. N2-Isobutyryl-5′-O-(4,4′-dimethoxytrityl)-3′-O-(N,N-diisopropylamino)-(4-oxo-1-pentyloxy)phosphinyl-2′-deoxyguanosine (S.8d): 31P-NMR (121 MHz, C6D6 ): δ 148.0, 148.5. FAB-HRMS: anal. calcd. for C46H59N6O9P (M+Na)+ 893.3980, found 893.3965.
Synthesize oligonucleotides 4. Perform solid-phase synthesis as described in Basic Protocol (step 8), but use S.8a-d as 0.2 M solutions in dry acetonitrile, and increase the wait time for the coupling reaction to 150 sec.
A
B
C
0
3-(N-tert-Butylcarboxamido)-propyl and 4-Oxopentyl Protecting Groups
10
20 Retention time (min)
30
40
Figure 3.9.8 RP-HPLC analysis of crude d(ATCCGTAGCTAAGGTCATGC). (A) Crude 20-mer synthesized from standard 2-cyanoethyl deoxyribonucleoside phosphoramidites and deprotected by treatment with concentrated ammonium hydroxide as described in the Alternate Protocol, step 5 annotation. (B) Crude 20-mer synthesized from S.8a-d and deprotected as described in (A). (C) Crude 20-mer synthesized from S.8a-d and deprotected under pressurized ammonia gas as delineated in the Alternate Protocol, step 5. Adapted from Wilk et al. (2001) with permission from Elsevier Science.
3.9.10 Supplement 11
Current Protocols in Nucleic Acid Chemistry
0
10
20 Retention time (min)
30
40
Figure 3.9.9 R P - HP LC analysis of the enzymatic hydrolysis of cr ude d(ATCCGTTGCTAAGGTCATGC) by snake venom phosphodiesterase and bacterial alkaline phosphatase. The crude 20-mer is synthesized from S.8a-d and deprotected under pressurized ammonia gas as described in the Alternate Protocol, step 5. Identities of the RP-HPLC peaks from left to right are dC, dG, dT, dA, and benzamide (∼17 min) when compared to authentic commercial samples. Reprinted from Wilk et al. (2001) with permission from Elsevier Science.
A
200
150
100
50
ppm
200
150
100
50
ppm
B
Figure 3.9.10 Two 121-MHz 31P-NMR spectra of crude 20-mer d(APSTPSCPSCPSGPS TPSAPSGPSCPSTPSAPSAPSGPSGPSTPSCPSAPSTPSGPSC) in aqueous solvents. (A) A 20-mer synthesized from commercial 2-cyanoethyl deoxyribonucleoside phosphoramidites and deprotected by treatment with concentrated NH4OH (55°C, 10 hr). (B) A 20-mer prepared from S.8a-d and deprotected under pressurized NH3 as described in the Alternate Protocol, step 5. Reprinted from Wilk et al. (2001) with permission from Elsevier Science.
Synthesis of Unmodified Oligonucleotides
3.9.11 Current Protocols in Nucleic Acid Chemistry
Supplement 11
Perform oligonucleotide deprotection 5. Deprotect as described (see Basic Protocol, steps 9 and 10). Under these conditions, both nucleobase- and phosphate-protecting groups are removed, and the oligonucleotide is released from the support. Alternatively, push 1 mL of concentrated ammonium hydroxide back and forth through the synthesis column over a 1-hr period using two syringes with Luer tips. Then transfer the solution to a 4-mL screw-capped glass vial and heat at 55°C in the closed container for 10 hr. Cool to room temperature and proceed to step 7.
6. Using a 1-mL syringe with Luer tip, carefully push back and forth 0.5 mL of 0.1 M triethylammonium acetate buffer through the synthesis column. Collect the solution in a 4-mL screw-capped glass vial. 7. Determine the concentration of the oligonucleotide by UV spectrophotometry at 260 nm. Analyze oligonucleotide by RP-HPLC 8. Analyze ∼1 OD260 unit of the oligonucleotide on a 5-µm Supelcosil LC-18S HPLC column (UNIT 10.5) using a linear gradient of 1% acetonitrile/min starting from 0.1 M triethylammonium acetate, pH 7.0, at a flow rate of 1 mL/min. RP-HPLC profiles of oligonucleotides synthesized from S.8a-d or standard 2-cyanoethyl deoxyribonucleoside phosphoramidites, which were deprotected by treatment with pressurized ammonia gas or concentrated ammonium hydroxide, are presented in Figure 3.9.8.
Characterize oligonucleotide by enzymatic hydrolysis 9. Pipet 1 OD260 unit of crude oligonucleotide synthesized from S.8a-d in a 1.5-mL microcentrifuge tube and evaporate to dryness using a stream of air. Analyze as described (see Basic Protocol, steps 21 to 24). A RP-HPLC profile of the enzymatic hydrolysates is shown in Figure 3.9.9.
Analyze phosphorothioated oligonucleotide by 31P-NMR spectroscopy 10. Elute phosphorothioated oligonucleotide (synthesized from S.8a-d and deprotected as in step 5) from the synthesis column with 0.5 mL D2O and analyze by 31P-NMR spectroscopy. A 31P-NMR spectrum of the phosphorothioated oligonucleotide d(ATCCGTAGCTAAGGTCATGC) is shown in Figure 3.9.10.
COMMENTARY Background Information
3-(N-tert-Butylcarboxamido)-propyl and 4-Oxopentyl Protecting Groups
The demand for oligonucleotide syntheses has increased dramatically over the years, and the scales at which these syntheses are performed typically ranges from nanomole or micromole scale to millimole scale for therapeutic applications. The large-scale preparation of therapeutic oligonucleotides has challenged the suitability of 2-cyanoethyl deoxyribonucleoside phosphoramidites (Sinha et al., 1984) for such syntheses because acrylonitrile is produced as a side-product during oligonucleotide deprotection (Tener, 1961). Acrylonitrile is a potent DNA alkylating agent that can compromise the potency of oligonucleotide drug prod-
ucts (Wilk et al., 1999a). The authors have earlier reported a phosphoramidite approach to the solid-phase synthesis of DNA oligonucleotides that employs the 4-[N-methyl-N(2,2,2-trifluoroacetyl)amino]butyl group for phosphate/thiophosphate protection (Wilk et al, 1999a,b; UNIT 2.7). This group is easily removed from oligonucleotides by standard treatment with concentrated ammonium hydroxide or pressurized ammonia gas (Boal et al., 1996), and generates 2,2,2-trifluoroacetamide and Nmethylpyrrolidine as nonmutagenic side-products. Although these phosphoramidites lead to alkylation-free DNA oligonucleotide products, 4-(N-methylamino)butan-1-ol, which is a pre-
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Current Protocols in Nucleic Acid Chemistry
properties (Grajkowski et al., 2001; Wilk et al., 2002). This protecting group is prepared economically from γ -butyrolactone and tert-butylamine, and then converted to the phosphinylating reagent S.2 upon reaction with the chlorophosphine S.4 (see Support Protocol; Fig. 3.9.5). Condensation of suitably protected nucleosides S.1a-d with S.2 in the presence of 1H-tetrazole affords the 3-(N-tert-butylcarboxamido)-1-propyl deoxyribonucleoside phosphoramidites S.3a-d (see Basic Protocol; Fig. 3.9.1). These have been successfully used in the so lid-phase synthesis of the 20-mer d(ATCCGTAGCTAAGGTCATGC) and its phosphorothioated analog (Wilk et al., 2002). Following nucleobase deprotection and release of the oligonucleotide from the support, removal of the 3-(N-tert-butylcarboxamido)-1propyl phosphate/thiophosphate-protecting group is accomplished within 1 hr under thermolytic conditions at neutral pH. The deprotection reaction is considerably faster than that of the 2-(N-formyl-N-methyl)aminoethyl protecting group (3 hr) under identical conditions, and proceeds with the participation of the amidic carbonyl group in an intramolecular cyclodeesterification reaction, which, as shown in Figure 3.9.11, releases the phosphate/thiophosphate diester as its 2-tert-butyliminotetra-
cursor in the synthesis of the phosphoramidites, is not commercially available and must therefore be synthesized. Because of this added expenditure, the 4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl deoxyribonucleoside phosphoramidites might not effectively compete with other phosphoramidites to produce oligonucleotide drugs at the most affordable cost per dose. This shortcoming prompted the investigation of P(III)-protecting groups for cost-effective syntheses of deoxyribonucleoside phosphoramidites. The authors found the 2-(N-formyl-N-methyl)aminoethyl group to be adequate for this purpose, and syntheses of oligodeoxyribonucleotides, such as dT18 and d(AG)10, have been reported (Grajkowski et al., 2001). Although the 2-(Nformyl-N-methyl)aminoethyl phosphate/thiophosphate-protecting group cannot be removed reliably by standard ammonolysis, it is easily cleaved from oligonucleotides via a unique thermolytic cyclodeesterification process (Wilk et al., 2000; Grajkowski et al., 2001). While the group is cleanly removed from phosphate or phosphorothioate triesters under neutral conditions, its deprotection is excessively time consuming (3 hr). The 3-(N-tert-butylcarboxamido)-1-propyl group also exhibits thermolytic deprotection
O
O
B
B
O
X
O
pH 7.0
P O
90°C, 1 hr
O
B O
HN
HN O
O
O
B
O X
P O
O
O
B
O
O
O
O
P X−
O
B
O
O
HN +
O O
O
Figure 3.9.11 Proposed mechanism for the removal of the 3-(N-tert-butylcarboxamido)-1-propyl phosphate/thiophosphate-protecting group under thermolytic conditions at neutral pH. X = O or S; B = any of thymin-1-yl, cytosin-1-yl, adenin-9-yl, or guanin-9-yl.
Synthesis of Unmodified Oligonucleotides
3.9.13 Current Protocols in Nucleic Acid Chemistry
Supplement 11
B1
O
O
B
O
X
O
NH3 (g)
P O
or NH4OH
O
B O CH3
CH3
CH3
O
O
O
B1
O X
H2 N
P O
O
O
B
O
O
O
OH
P
X−
O
N B
O
O
+ O
O
Figure 3.9.12 Proposed mechanism for the removal of the 4-oxopentyl phosphate/thiophosphateprotecting group under mild basic conditions. X = O or S; B1 = any of thymin-1-yl or N-protected nucleobases; B = any of thymin-1-yl, cytosin-1-yl, adenin-9-yl, or guanin-9-yl.
3-(N-tert-Butylcarboxamido)-propyl and 4-Oxopentyl Protecting Groups
hydrofuran salt. Neither nucleobase alkylation nor significant phosphorothioate desulfurization were detected under conditions that would mimic millimole-scale oligonucleotide deprotection (Wilk et al., 2002). The removal of phosphate/thiophosphate-protecting groups from DNA oligonucleotides in the absence of concentrated ammonium hydroxide is advantageous in that the handling of hazardous ammonium hydroxide can be completely eliminated from oligonucleotide processing. This advantage should certainly be appreciated when performing large-scale therapeutic oligonucleotide syntheses, as large quantities of concentrated ammonium hydroxide are needed for releasing oligonucleotides from the solid support and for nucleobase deprotection. Another attractive phosphate/thiophosphate-protecting group for synthetic oligodeoxyribonucleotides is the 4-oxopentyl group (Wilk et al., 2001). This protecting group is incorporated into deoxyribonucleoside phosphoramidites S.8a-d under conditions identical to those used for the preparation of S.3a-d (see Alternate Protocol; Fig. 3.9.7). The phosphoramidites S.8a-d are as efficient as S.3a-d or 2-cyanoethyl deoxyribonucleoside phosphoramidites in the solid-phase synthesis of the 20-mer d(ATCCGTAGCTAAGGTCATGC) and its phosphorothioated analog (Wilk et al., 2001). Unlike the 3-(N-tert-butylcarboxamido)-1-propyl and 2-(N-formyl-Nmethyl)aminoethyl groups, the 4-oxopentyl
group was rapidly and completely cleaved from these oligonucleotides under the conditions required for nucleobase deprotection using either pressurized ammonia gas or concentrated ammonium hydroxide (Wilk et al., 2001). The reaction of ammonia with the ketone function of the phosphate/thiophosphate-protecting group most likely leads to the formation of a hemiaminal intermediate as shown in Figure 3.9.12. This intermediate underwent rapid cyclodeesterification to release the phosphate/thiophosphate diester with the concomitant formation of 2-methyl-1-pyrroline as the major deprotection side-product (Wilk et al., 2001). This deprotection pathway is consistent with the removal of 4-N-methylaminobutyl phosphate/thiophosphate-protecting groups reported earlier under similar conditions (Wilk et al., 1999a,b). The generation of 2-methyl-1pyrroline during the course of oligonucleotide deprotection did not produce detectable nucleobase alkylation or significant phosphorothioate desulfurization even under conditions that would simulate large-scale oligonucleotide deprotection (Wilk et al., 2001). Thus, use of the 3-(N-tert-butylcarboxamido)-1-propyl or 4-oxopentyl group for phosphate/thiophosphate protection in the large-scale production of therapeutic oligonucleotides is an obvious choice as these groups offer economy in the manufacture of such biopolymers, provide versatility in the selection of oligonucleotide deprotection condi-
3.9.14 Supplement 11
Current Protocols in Nucleic Acid Chemistry
tions, and ensure alkylation-free drug product for optimal potency.
Critical Parameters and Troubleshooting Since the phosphordiamidites S.2 and S.7 are prepared using the bis(N,N-diisopropylamino) chlorophosphine intermediate S.4 (see Support Protocol, Fig. 3.9.5) in a manner similar to that reported for the N,N,N′,N′-tetraisopropyl-O-{2-[(N-formyl-N-methyl)amino]-ethylphosphordiamidite (UNIT 2.7, Fig. 2.7.4), the readers are referred to the section on Critical Parameters and Troubleshooting of UNIT 2.7, which addresses issues pertaining to the synthesis of phosphordiamidites. It is critically important that the formation of S.4 be carefully monitored by 31P-NMR spectroscopy to ensure complete exhaustion of phosphorus trichloride prior to adding the amidoalcohol S.5 or 3-acetyl-1-propanol. It is recommended that the purification of the deoxyribonucleoside phosphoramidites S.3a-d and S.8a-d be performed on a silica gel column using first a 1:8:1 (v/v/v) solution of methylene chloride/hexane/triethylamine as the eluent, and to carefully monitor by TLC the elution of fast-moving impurities. The concentration of methylene chloride should then be increased to elute the remaining fast-moving impurities until the final composition of the eluent is reached to elute the desired deoxyribonucleoside phosphoramidites. In order to ensure optimal coupling efficiency of individual deoxyribonucleoside phosphoramidites during conventional solidphase oligonucleotide synthesis, a number of parameters must be taken into consideration. These are addressed in the section on Critical Parameters and Troubleshooting of UNIT 2.7.
Anticipated Results The synthesis of deoxyribonucleoside phosphoramides S.3a-d and S.8a-d from their respective phosphordiamidites S.2 and S.7 is performed essentially as recommended by Barone et al. (1984). The condensation time is extended for the preparation of S.3d and S.8d to ensure optimal yields. The coupling efficiency of phosphoramidites S.3a-d and S.8a-d is comparable to that of standard 2-cyanoethyl deoxyribonucleoside phosphoramidites.
phosphordiamidites S.2 and S.7 also takes 4 days. The synthesis and purification of each of the deoxyribonucleoside phosphoramidites S.3a-d and S.8a-d requires 1 to 2 days to complete.
Literature Cited Barone, A.D., Tang, J.-T., and Caruthers, M.H. 1984. In situ activation of bis-dialkylaminophosphines—a new method for synthesizing deoxyoligonucleotides on polymer supports. Nucl. Acids Res. 12:4051-4061. Boal, J.H., Wilk, A., Harindranath, N., Max, E.E., Kempe, T., and Beaucage, S.L. 1996. Cleavage of oligodeoxyribonucleotides from controlledpore glass supports and their rapid deprotection by gaseous amines. Nucl. Acids Res. 24:31153117. Grajkowski, A., Wilk, A., Chmielewski, M.K., Phillips, L.R., and Beaucage, S.L. 2001. The 2-(Nformyl,N-methyl)aminoethyl group as a potential phosphate/thiophosphate protecting group in solid-phase oligodeoxyribonucleotide synthesis Organic Lett. 3:1287-1290. Iyer, R.P., Phillips, L.R., Egan, W., Regan, J.B., and Beaucage, S.L. 1990. The automated synthesis of sulfur-containing oligodeoxyribonucleotides using 3H-1,2-benzodithiol-3-one-1,1-dioxide as a sulfur-transfer reagent. J. Org. Chem. 55:46934699. Sinha, N.D., Biernat, J., McManus, J., and Köster, H. 1984. Polymer support oligonucleotide synthesis. 18. Use of β-cyanoethyl-N,N-dialkylamino/N-morpholino phosphoramidite of deoxynucleosides for the synthesis of DNA fragments simplifying deprotection and isolation of the final product. Nucl. Acids Res. 12:45394557. Tener, G.M. 1961. 2-Cyanoethyl phosphate and its use in the synthesis of phosphate esters. J. Am. Chem. Soc. 83:159-168. Wilk, A., Grajkowski, A., Phillips, L.R., and Beaucage, S.L. 1999a. The 4-[N-methyl-N-(2,2,2-trifluoroacetyl)amino]butyl group as an alternative to the 2-cyanoethyl group for phosphate protection in the synthesis of oligodeoxyribonucleotides. J. Org. Chem. 64:7515-7522. Wilk, A., Grajkowski, A., Srinivasachar, K., and Beaucage, S.L. 1999b. Improved chemistry for the production of synthetic oligodeoxyribonucleotides. Antisense Nucleic Acid Drug Dev. 9:361-366. Wilk, A., Grajkowski, A., Phillips, L.R., and Beaucage, S.L. 2000. Deoxyribonucleoside cyclic Nacylphosphoramidites as a new class of monomers for the stereocontrolled synthesis of oligothymidylyl- and oligocytidylyl-phosphorothioates. J. Am. Chem. Soc. 122:21492156.
Time Considerations The preparation and purification of (N-tertbutyl)-4-hydroxybutyramide S.5 is accomplished within 4 days. The synthesis of the
Synthesis of Unmodified Oligonucleotides
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Wilk, A., Chmielewski, M.K., Grajkowski, A., Phillips, L.R., and Beaucage, S.L. 2001. The 4oxopentyl group as a labile phosphate/thiophosphate protecting group for synthetic oligodeoxyr ibonucleotides. Te trahedron Lett. 42:5635-5639. Wilk, A., Chmielewski, M.K., Grajkowski, A., Phillips, L.R., and Beaucage, S.L. 2002. The 3-(Ntert-butylcarboxamido)-1-propyl group as an attractive phosphate/thiophosphate protecting group for solid-phase oligodeoxyribonucleotide synthesis. J. Org. Chem. 67:6430-6438.
Contributed by Andrzej Wilk, Marcin K. Chmielewski, Andrzej Grajkowski, and Serge L. Beaucage Food and Drug Administration Bethesda, Maryland Lawrence R. Phillips National Cancer Institute Frederick, Maryland
3-(N-tert-Butylcarboxamido)-propyl and 4-Oxopentyl Protecting Groups
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DNA Synthesis Without Base Protection An ultimate goal in the chemical synthesis of DNA is the ability to form O-selective internucleotide bonds without the use of baseprotecting groups, as can be done by enzymatic synthesis using DNA polymerases. The current chemical synthesis of DNA fragments requires multistep procedures for the introduction and removal of protecting groups to guarantee the fidelity of the synthesis (Caruthers, 1991). Satisfactory results have in fact been reported for the synthesis of unmodified DNA oligomers (UNIT 3.1). In these post-genome days, however, there are increasing demands to accelerate manipulation for gene detection and diagnosis, so that it is increasingly desirable to find more straightforward methods for the synthesis of DNA fragments (Shena, 1999). For this purpose, a number of improvements in the widespread phosphoramidite approach, which is used as the most reliable method, have been reported (Beaucage and Iyer, 1992; UNIT 3.3). They involve the use of more base-labile N-phenoxyacetyl protecting groups (Schulhof et al., 1987) and methylamine (Boal et al., 1996) as a powerful deacylating reagent for rapid deprotection, as well as the use of protecting groups that can be removed under neutral conditions (Hayakawa et al., 2001). As an alternative to these types of improvements, O-selective internucleotide bond formation provides an entirely different approach; the methods described here will offer new insight into modern DNA synthesis. The feasibility of O-selective internucleotide bond formation using non-N-acylated nucleoside derivatives has been suggested based on the unique properties of the amino and hydroxy groups of nucleosides. Baddiley and Todd (1947) reported that 2 ,3 O-isopropylideneadenosine underwent 5 -Oselective phosphorylation with dibenzyl phosphorochloridate in pyridine (Fig. 3.10.1). In phosphorylation of nucleosides, it is well known that phosphorus oxychloride selectively reacts with nucleosides in triethyl phosphate to give 5 -O-phosphorylated nucleosides (Yoshikawa and Kato, 1967; Ikemoto et al., 1995). Partial acetylation of adenosine with acetic anhydride gives 3 ,5 -di-Oacetyladenosine (Kool et al., 1987). Silylation of deoxyribonucleotides gives exclusively Osilylated products (Ogilvie, 1973; Sekine et al.,
UNIT 3.10
1985). In contrast, it was also reported that deoxycytidine underwent selective N-acylation with various acylating reagents (Watanabe and Fox, 1966; Nikolenko et al., 1967; Hata and Kurihara, 1973). These apparently contradictory results could be explained through a comprehensive understanding that the difference in reactivity between the hydroxy and amino groups of nucleosides varies depending on the inherent Soft-Hard (Pearson, 1987, 1990) nature of nucleobases and electrophilic reagents, as well as reaction conditions such as solvents and additives. Therefore, to realize oligonucleotide synthesis without base protection, it is important to create the most reasonable intermediates that are capable of O-selective phosphorylation in the reaction system.
LIQUID-PHASE SYNTHESIS Several precedents concerning oligonucleotide synthesis without N-protecting groups have been reported for liquid-phase oligonucleotide synthesis (Fig. 3.10.1). Using his original phosphite triester approach (Yuodka et al., 1976), Letsinger found that the amino group of the adenine base moiety exhibited rather poor reactivity toward tervalent phosphitylating intermediates (Letsinger et al., 1976; Finnan et al., 1980). Using this method, these investigators synthesized an oligonucleotide with the sequence TAAAT. Adamiak reported the synthesis of tRNA anticodon loop fragments containing ureidonucleosides without protection of the adenine moiety by using 2,4,6triisopropylbenzensulfonyl 1H-tetrazolide as the condensing reagent in the phosphotriester approach (Adamiak and Stawinski, 1977; Adamiak et al., 1978). Fourrey and Varenne (1985) reported the synthesis of DNA dimers using N-unprotected dA and dC phosphoramidite building blocks with 2-chlorophenyl and morpholino groups. Hayakawa reported several methods for activation of alcoholic components by use of metalating reagents such as t-BuLi (Hayakawa et al., 1983a), tBuMgCl (Hayakawa et al., 1984, 1985), and LiAl(NiPr2 )4 (Hayakawa et al., 1983b). These agents accelerate displacement with active esters of nucleoside phosphotriester components without introduction of any protecting groups on the adenine base. Synthesis of Unmodified Oligonucleotides
Contributed by Mitsuo Sekine Current Protocols in Nucleic Acid Chemistry (2004) 3.10.1-3.10.15 C 2004 by John Wiley & Sons, Inc. Copyright
3.10.1 Supplement 18
Figure 3.10.1 Structure of oligonucleotide derivatives obtained by liquid-phase synthesis without base protection. Abbreviations: Ade, adenin-9-yl; Cyt, cytosin-1-yl; DMTr, 4,4 -dimethoxytrityl; Gua, guanin-9-yl; MMTr, 4-monomethoxytrityl; TBDMS, tert-butyldimethylsilyl; Thy, thymin-1-yl; Ura, uracil-1-yl. References: (1) Letsinger et al. (1976); (2) Adamiak and Stawinski (1977); (3) Fourrey and Varenne (1985); (4) Hayakawa et al. (1983a,b, 1984); (5) Hayakawa et al. (1987).
SOLID-PHASE SYNTHESIS USING THE PHOSPHORAMIDITE APPROACH Synthesis of Common Starting Nucleosides 5 -O-Dimethoxytrityldeoxyribonucleosides
DNA Synthesis Without Base Protection
There are two methods for the synthesis of deoxyribonucleosides with 5 -O-(4,4 dimethoxytrityl) (DMTr) protection. One is the direct introduction of the DMTr group into deoxyribonucleosides. The 5 -O-selective tritylation of dC in pyridine gives rise to 5 -ODMTr-dC in 89% yield, as first described by Michelson and Todd (1954). Jones reported that direct dimethoxytritylation of dA with 1.5 equiv of DMTr-Cl in pyridine gave 5 -ODMTr-dA in 77% yield (Fig. 3.10.2, method
A), and that the use of 2 equiv of DMTr-Cl for dC gave 5 -O-DMTr-dC and 5 -O,N-bisDMTrdC in 32% and 40% yields, respectively (Ti et al., 1982). Levin and colleagues also reported that dimethoxytritylation of dA, dC, and dG gave the corresponding 5 -O-DMTr derivatives in 61%, 48%, and 55% yields, respectively (Levin et al., 1991), although no details were described in the paper. In connection with 5 -O-selective dimethoxytritylation, Adamiak’s early paper concerning 5 -O-selective monomethoxytritylation of A, C, and G is quite positive (Okupniak et al., 1981). Reaction of dA, dC, or dG with 1.2 equiv of 4-monomethoxytrityl chloride (MMTr-Cl) in dimethylformamide (DMF) or dimethyl sulfoxide (DMSO) in the presence of 6-nitroquinoline (pKa = 2.7) or 3-bromopyridine (pKa = 2.9) gave
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Figure 3.10.2 Synthesis of 5 -O-DMTr-N-unprotected deoxyribonucleosides using direct approaches (methods A to C) or transient dimethylaminomethylene (Dmf) base protection (method D). DMF, dimethylformamide.
5 -O-MMTr-dC, 5 -O-MMTr-dA, and 5 -OMMTr-dG in 85%, 92%, and 66% yields, respectively. The use of bases that are weaker than the dA and dC nucleobases is essential for high 5 -O-selectivity, because the nucleobases must be protonated to be converted to inert species under the reaction conditions used. Further application of this method to a series of deoxyribonucleosides should be studied in the future. Ishido reported that 5 -O-selective dimethoxytritylation of dC in the presence of dichloroacetic acid gave 5 -O-DMTr-dC in 71% yield (Ishido, 1989; Nishino et al., 1991; Fig. 3.10.2, method B). This procedure is applicable to the synthesis of 5 -O-DMTrdA but not 5 -O-DMTr-dG (A. Ohkubo, K. Seio, and M. Sekine, unpub. observ.). Hayakawa reported an improved method for the selective 5 -O-dimethoxytritylation of deoxyribonucleosides using DMTr-Cl/ triethylamine/imidazole in DMF, which gave 5 -O-DMTr-dC, 5 -O-DMTr-dA, and 5 -O-DMTr-dG in 76% to 85%, 82% to 89%,
and 74% to 78% yields, respectively (Kataoka and Hayakawa, 1999; Fig. 3.10.2, method C). A different method employed at an earlier stage by Jones involves transient Nprotection with the dimethylaminomethylene (Dmf) group (Froehler and Matteucci, 1983a) by reaction of deoxyribonucleosides with dimethylformamide dimethylacetal (Kung and Jones, 1992; Fig. 3.10.2, method D). Although this approach requires a three-step procedure to obtain the desired 5 -O-DMTrdeoxyribonucleosides, the transient protection and deprotection using the Dmf group proceed cleanly and quantitatively, so that purification is necessary for only the final product. This procedure has proved to be reliable and reproducible. In the case of dG, 5 -O,2-Nbis(DMTr)deoxyguanosine was used as the precursor of the 3 -phosphoromorpholidite derivative (Fourrey and Varenne, 1985). This ditritylated species can be obtained by reaction of dG with excess DMTr-Cl in pyridine (Schaller et al., 1963).
Synthesis of Unmodified Oligonucleotides
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Figure 3.10.3 (left).
Synthesis of 5 -O,N-bis(DMTr)deoxyribonucleosides by successive tritylation (right) or direct tritylation
5 -O,N-Bis(dimethoxytrityl)deoxyribonucleosides for the 3 terminus
DNA Synthesis Without Base Protection
The general synthesis of oligodeoxyribonucleotides with appropriate sequences requires the synthesis of 3 -terminal deoxyribonucleoside building blocks that can be attached to resins via linkers. To introduce dA, dC, and dG into the 3 -terminal site, their 3 -succinate ester derivatives or other equivalents are required. Because succinic anhydride reacts with the nucleobases (Kume et al., 1984), the amino functions of these deoxyribonucleosides should be protected to avoid such side reactions. For this purpose, the best approach is to use 5 -O,N-bis(DMTr)deoxyribonucleoside derivatives (S.7g-S.7i; Fig. 3.10.3). Several approaches have been tested for the synthesis of these compounds. NDimethoxytritylation with DMTr-Cl in the presence of Me3 SiCl followed by successive 5 -O-dimethoxytritylation (right side of figure) gave N-DMTr-deoxyribonucleosides in high yields (Wada et al., 1998b). The direct preparation of these substances from deoxyribonucleosides (left) resulted in complex mixtures including the desired materials. To introduce these 5 -O,N-bis(DMTr) derivatives onto polymer supports, they were converted to 3 -succinate half-esters, which were linked to resins in situ by condensation using a phosphonium-type condensing reagent, 2-(benzotriazol-1-yloxy)1,1-dimethyl-2-(pyrrolidin-1-yl)-1,3,2diazaphospholidinium hexafluorophosphate
(BOMP; Wada et al., 1997). To mask unreacted amino groups on the resin, careful treatment with suitable acylating reagents is very important. When an N-DMTr-dG derivative was treated with acetic anhydride in pyridine in the presence of 4-(dimethylamino)pyridine, complete replacement of the DMTr group with an acetyl group took place (Fig. 3.10.4; Wada et al., 1998b). This inherent side reaction was overcome by using 1-acetylimidazole as the acylating reagent. This reagent did not react with the guanosine moiety but completely reacted with the unreacted amino groups on the resin. The N-DMTr group is generally more stable than the O-DMTr group. Complete deprotection for dA and dG required only 5 min when using 1% trifluoroacetic acid (TFA) in CH2 Cl2 . dC derivatives required a more prolonged time of 30 min for complete removal of the DMTr group using 3% TFA in CH2 Cl2 , but the glycosyl bond of these derivatives is sufficiently stable that this is not a problem. Several short oligodeoxyribonucleotides containing dA, dC, or dG at the 3 -terminal site were successfully synthesized using the Hphosphonate approach without base protection (Fig. 3.10.5; Wada et al., 1998b). Although the syntheses of relatively longer oligodeoxyribonucleotides (20- to 21-mers) having dG (Gryaznov and Letsinger, 1992; Hayakawa and Kataoka, 1998) and dC (Hayakawa and Kataoka, 1998) at the 3 -terminal position were reported, no details about how to solve the aforementioned problem were described.
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Figure 3.10.4 The capping reaction prescribed for masking the unreacted amino groups on the resin also results in an undesired N-acetylation side reaction. This can be prevented by using 1-acetylimidazole (AcIm) instead of acetic anhydride (Ac2 O) and 4-(dimethylamino)pyridine (DMAP).
Figure 3.10.5 Synthesis of polymer-supported deoxyribonucleoside monomer building blocks. B and B2 are unprotected and N-DMTr-protected nucleobases, respectively.
Synthesis of Deoxyribonucleoside 3 -Phosphoramidite Building Blocks
The 3 -(O-methyl-N,N -tetraisopropylphosphoramidite) building blocks of Nunprotected deoxyribonucleosides (dC, dA, and dG) were synthesized in 85% to 90% yield by 3 -O-selective phosphitylation of the corresponding N-unprotected 5 -ODMTr-deoxyribonucleosides with methyl (N,N -tetraisopropyl)phosphorodiamidite in the presence of diisopropylammonium 1Htetrazolide (Fig. 3.10.6, method B; Gryaznov and Letsinger, 1991, 1992). The N-unprotected deoxyribonucleoside 3 -(O-cyanoethyl-N,N tetraisopropylphosphoramidite) building blocks were also synthesized by reaction of 5 -O-DMTr-deoxyribonucleosides with (i-Pr)2 NP(Cl)OCH2 CH2 CN in tetrahydrofuran in the presence of diisopropylethylamine (method A; Hayakawa and Kataoka, 1998). As far as the solubility of these synthetic units is
concerned, the guanosine 3 -(O-cyanoethyl)phosphoramidite is not completely soluble in acetonitrile, which is the most suitable solvent for DNA synthesizers because of its low viscosity. It is necessary to take this poor solubility into account when using an automated DNA synthesis approach, as it may cause clogging of the tubing.
Oligodeoxyribonucleotide Synthesis Using the Phosphoramidite Approach The first report of a solid-phase DNA synthesis without base protection was made by Ogilvie and Nemer (1981), who synthesized a short oligoribonucleotide, UGCA, in 80% yield using a polymer-bound non-N-acylated adenosine derivative and the phosphite triester approach. Later, Gryaznov and Letsinger (1991, 1992) reported a general method for the synthesis
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Figure 3.10.6 Synthesis of N-unprotected deoxyribonucleoside 3 -O-phosphoramidite building blocks. DIPAT, diisopropylammonium 1H-tetrazolide; r.t., room temperature; THF, tetrahydrofuran.
Figure 3.10.7 Synthesis of oligodeoxyribonucleotides without base protection using the phosphoramidite approach, and postcondensation treatment for N-P bond cleavage of branched chains resulting from concomitant side reactions on dA and dC bases. BIT, benzimidazolium triflate; IMT, imidazolium triflate; Pyr-HCl, pyridinium hydrochloride.
DNA Synthesis Without Base Protection
of oligodeoxyribonucleotides using pyridinium hydrochloride as the activator and their original syringe method (Tanaka and Letsinger, 1982; Fig. 3.10.7). With this approach, they succeeded in synthesizing 20mers. Because they observed serious modifications of the dA and dC bases, the modified sites were reconverted to free amino groups after each condensation by treatment with aniline in the presence of pyridinium hydrochloride. When internucleotide bonds are formed using the phosphoramidite approach, it is noteworthy that the guanine moiety is not modified with activated phosphoramide species (Gryaznov and Letsinger, 1991, 1992). This is in sharp contrast to the fact that significant 6-O-phosphorylation of dG residues has been observed when the phosphotriester approach was employed (Sekine et al., 1982).
A newer phosphoramidite approach was reported using imidazolium triflate (IMT) as a new activator and using a somewhat swelled Tentagel as the resin (Hayakawa and Kataoka, 1998; Hayakawa et al., 2001). In this approach, a more powerful activator, benzimidazolium triflate (BIT; Hayakawa et al., 1996; Hayakawa, 2001), was used in methanol as a nucleophile to eliminate 5 O-DMTr-deoxyribonucleoside 3 -phosphityl residues from unavoidably modified base sites. By this method, 32-mer, 60-mer, and two 21-mer oligonucleotides were synthesized as the major products using a DNA synthesizer. The reproducibility of this approach has not been confirmed, however, as no applications of this method have appeared to date. As the solvent used for this synthesis was acetonitrile, it seems to be somewhat difficult to
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Figure 3.10.8
Outline of the proton-block method using the activator 5-nitrobenzimidazolium triflate (NBT).
completely dissolve the guanosine building block. On the other hand, it has recently been reported that the BIT/MeOH treatment causes serious internucleotide bond fission (∼50%; Sekine et al., 2003) when highly cross-linked polystyrene (HCP; McCollum and Andrus, 1991) or controlled-pore glass (CPG; K¨oster et al., 1983) were employed as the resin in place of Tentagel. Another new strategy for oligodeoxyribonucleotide synthesis without base protection takes a completely different approach. It is called the “proton-block” method, because the simple proton is used for protection of base moieties (Fig. 3.10.8; Sekine et al., 2003). This strategy was developed based on the principle of acid-base reactions. It is well known that once amino functions are protonated they lose nucleophilicity because of the formation of inert ammonium salts. Adamiak’s successful 5 -O-monomethoxytritylation of ribonucleosides (Adamiak, 1981; see Synthesis of Common Starting Nucleosides) is the only precedent that utilizes this principle in nucleic acid chemistry. Based on this principle, a new activator, 5-nitrobenzimidazolium triflate (NBT), was synthesized. This new promoter has a pKa value of 2.76 and thus enables sufficient protonation of cytosine and adenine bases, which have pKBH+ values of 4.3 and 3.8, respectively (Hall, 1971). To date, such uses of a more acidic species as the promoter for the phosphoramidite approach have not been reported. It has been reported that 5-(4-nitrophenyl)-1H-tetrazole (Froehler and Matteucci, 1983b) is so acidic (pKa = 3.7) that it causes serious elimina-
tion of the DMTr group. When NBT was used in acetonitrile for the solid-phase synthesis of d[ApT], 3% of the product was obtained as d[ApApT]. This side reaction could be significantly suppressed, however, by using THF as the solvent. This solvent effect can be explained by the difference in basicity between acetonitrile and THF, which have pKBH+ values of −10 and −2, respectively (see Internet Resource). The 5 -DMTr ether linkage (DMTrO-CH2 -) was estimated to have a pKBH+ value that is between those of acetonitrile and THF, and probably nearer to THF, so that NBT cannot protonate the DMTr ether linkage in the presence of excess amounts of the more basic THF. It is important to choose the correct solvent in this strategy. The oligos d[CCCCCT] and d[AAAAAT] were synthesized without base protection and without postcondensation treatment with P-N bond-cleaving reagents. In the synthesis of d[CAGTCAGTCAGT], a mixed solvent of 9:1 (v/v) acetonitrile/Nmethylpyrrolidone (NMP) was used to avoid the competitive elimination of the DMTr group during the condensation. NMP has a pKBH+ value of −0.71, so that the excess promoter protonates this solvent predominantly over the 5 -etheral oxygen. Consequently, the 12-mer was obtained in 19% overall yield. The superiority of this strategy is evidenced by the fact that, when IMT was used as the promoter without the postcondensation treatment with BIT-MeOH, an extremely complicated HPLC profile was obtained (Sekine et al., 2003). Finally, a new strategy called the HOBT method has been developed (Fig. 3.10.9;
Synthesis of Unmodified Oligonucleotides
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Figure 3.10.9 The HOBT method for oligodeoxyribonucleotide synthesis without base protection and without postcondensation treatment. HOBT, 1-hydroxybenzotriazole; NMP, N-methylpyrrolidone; r.t., room temperature.
DNA Synthesis Without Base Protection
Ohkubo et al., 2002). 1-Hydroxy-benzotriazole (HOBT) was previously used as the promoter in the phosphoramidite approach by van Boom (Claesen et al., 1984). However, because of the poor solubility of this reagent in acetonitrile, which is widely used for automated DNA synthesizers, syntheses using HOBT had not appeared until this HOBT strategy was reported. Based on the mechanism of the H-phosphonate approach without base protection, HOBT was the reagent of choice for the purpose. Because the intermediate of a previous H-phosphonate approach without base protection was thought to be a tervalent phosphorus species (Wada et al., 1997; UNIT 3.4), it was believed that this kind of tervalent phosphorus species would be essential for O-selective internucleotide bond formation, and suitable reagents that can convert phosphoramidite derivatives to tervalent phosphite triester intermediates were studied. Among the reagents tested, HOBT was found to be the best reagent for O-selective internucleotide bond formation. Because HOBT (pKa = 5.3) is more basic than 1H-tetrazole (pKa = 4.8), there is no risk of elimination of the DMTr group during condensation, unlike the proton-block method. The only drawback of this reagent is its inherent insolubility in acetonitrile, but this was overcome by using a mixed solvent of acetonitrile and NMP. Moreover, the insolubility of the dG phosphoramidite, another common problem in the Nunprotected strategy, was also overcome by the use of the mixed acetonitrile/NMP solvent, in which the dG building block is easily soluble. With the help of these improvements,
d[CAGTCAGTCAGT] was synthesized as the major product in high yield. The O-selectivity in the synthesis of dimers and trimers exceeded 99.8%, a most excellent result. This approach enabled the synthesis of up to 20-mer oligodeoxyribonucleotides. The coupling efficiency decreased to ∼98%, however, when the dG monomer was used. If this problem could be solved, the HOBT method would be more realistic and practical.
SOLID-PHASE SYNTHESIS USING THE H-PHOSPHONATE APPROACH Synthesis of Deoxyribonucleoside 3 -Phosphonate Building Blocks
Synthesis of 5 -O-DMTr-deoxyribo nucleoside 3 -H-phosphonates was reported by several research groups. Levin et al. (1991) reported synthesis using salicyl chlorophosphite (Marugg et al., 1986; Ven’yaminova et al., 1988) as the phosphitylating reagent (Fig. 3.10.10, method B). Phosphonylation of N-dimethylaminomethylene5 -O-DMTr-deoxyribonucleosides with tris(triazolyl)phosphine (method A) gave the 3 -phosphonylated products as 1,8diazabicyclo-[5,4,0]undec-7-ene (DBU) salts (Kung and Jones, 1992). In the case of dC, the N-unprotected product was obtained directly, as the amidine group was completely lost during silica gel column chromatography. In the case of dA and dG, however, concomitant elimination of the amidine group occurred, giving a mixture of N-protected and N-unprotected products.
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Figure 3.10.10 Synthesis of N-unprotected deoxyribonucleoside 3 -O-phosphonate building blocks for oligonucleotide synthesis using the H-phosphonate approach.
Therefore, pretreatment of the crude products with ammonia before chromatography is necessary. Wada and Sekine reported the selective 3 -O-phosphonylation of 5 -ODMTr-deoxyribonucleosides with excess diphenyl phosphonate or with phosphonic acid in the presence of bis(2-oxo-3-oxazolidinyl)phosphinic chloride (methods C and D, respectively; Wada et al., 1997). The use of method C gave somewhat better yields of the 3 -O-phosphonylated products as DBU salts compared to method D (88% to 94% versus 78% to 87%, respectively).
Oligodeoxyribonucleotide Synthesis Using the H-Phosphonate Approach Levin et al. (1991) first reported the synthesis of oligodeoxyribonucleotides by the H-phosphonate method using N-unprotected 5 -O-DMTr-deoxyribonucleoside 3 -phosphonates. They synthesized d[TATTTGGCT], but no details concerning the reaction conditions were described. Kung and Jones (1992) reported the synthesis of DMTr-d[TCAGT] using the H-phosphonate approach with
pivaloyl chloride or adamantyl chloride as the coupling reagent (Fig. 3.10.11). In this synthesis, it was found that pivaloyl chloride reacted with base residues. Although the pivaloyl groups were removed by standard ammonia treatment, the necessity for such ammonia treatment eliminates one of the advantages to not using amino-protecting groups. In 1997, Wada and Sekine also reported the use of the H-phosphonate approach without base-protecting groups (Wada et al., 1997; Fig. 3.10.12). In their approach, a phosphoniumtype condensing reagent, BOMP, was developed and shown to be very effective for selective internucleotide bond formation. The synthesis of d[CAGTCAGTCAGT] was accomplished with an overall isolated yield of 19%. Of particular interest, a plausible explanation as to why the selective O-phosphorylation occurred was also reported. The detailed molecular orbital (MO) calculations of possible intermediates suggest that two sets of HOMO-LUMO (highest and lowest molecular orbital) interactions between an activated Hphosphonate species and the 5 -terminal HO function are involved. In the reaction of the
Synthesis of Unmodified Oligonucleotides
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Figure 3.10.11 Use of pivaloyl chloride (trimethylacetyl chloride, R = t-Bu) for synthesis of oligodeoxyribonucleotides by the H-phosphonate approach without base protection.
Figure 3.10.12 Use of phosphonium-type condensing reagents for synthesis of oligodeoxyribonucleotides by the Hphosphonate approach without base protection. B is the unprotected nucleobase. BOMP, (2-benzotriazol-1-yloxy)-1,3dimethyl-2-pyrrolidin-1-yl-1,3,2-diazaphospholidinium hexafluorophosphate; BSA, N,O-bis(trimethylsilyl)acetamide; DMTrCl, 4,4 -dimethoxytrityl chloride.
DNA Synthesis Without Base Protection
activated species with amino functions of the nucleobases, the HOMO orbital on the hydrogen atom of the HO group is lacking, so that only a weaker interaction between the nitrogen atom and the phosphorus atom of the activated species is available. Later, it was reported that this strategy was accompanied by a serious side reaction that caused elimination of one nucleotide block during the condensation, resulting in the formation of significant amounts of (n-1)-mers (Wada et al., 1999).
ADVANTAGES AND PROSPECTS OF THE N-UNPROTECTED APPROACH Stability of N-Unprotected Deoxyribonucleoside Derivatives The stability of the glycosidic bond of Nunprotected deoxyribonucleoside derivatives is important. Several research groups have reported the kinetic data of N-protected and N-unprotected deoxyadenosine derivatives
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Current Protocols in Nucleic Acid Chemistry
(Butkus et al., 1983; Septak, 1996; Krotz et al., 2003). Froehler and Matteucci (1983a) reported that N-unprotected deoxyadenosine is more resistant to acids than is N-benzoyldeoxyadenosine. Specifically, Nbenzoyldeoxyadenosine has a T1/2 of 4 hr in 80% acetic acid at 25◦ C, whereas Nunprotected deoxyadenosine has a T1/2 of >5 days. The stability of the glycosidic bond of deoxyadenosine is similar to that of 6N-diisopropylformamidinedeoxyadenosine. It was also reported that unprotected deoxyguanosine is somewhat more unstable than deoxyadenosine under acidic conditions (Venner, 1964, 1966). This is in reverse order of the same series of N-acylated purine deoxyribonucleosides. In the current phosphoramidite approach using N-protected deoxyribonucleosides, it is important to balance the contradictory goals of obtaining maximum detritylation with minimum depurination at the final stage (Septak, 1996; Krotz et al., 2003).
Prospects of N-Unprotected Approach One promising advantage of the Nunprotected phosphoramidite approach is to make it possible to synthesize oligodeoxyribonucleotides directly on a resin or even on microchips or microarrays. The straightforward synthesis of DNA fragments on DNA microchips or microarrays will be realized in the near future. For this purpose, O-selectivity at the present level of internucleotide bond formation can be sufficiently utilized without difficulty. In gene diagnosis, success relies on the ability of a complementary sequence to bind to the patients’ target sequence on the DNA chip and not other sequences. In this respect, achievement of 99.8% to 99.9% O-selectivity (Ohkubo et al., 2002) in the chemical synthesis of DNA fragments means the high presence of the correct sequence relative to incorrect sequences. Because O-selectivity is nearly perfect in the HOBT method, the ratio of the correct sequence to each failure sequence is highly dependent on the efficiency of condensation at each cycle. As no capping treatment is carried out, the correct sequence is unavoidably contaminated with failure sequences lacking one nucleotide unit. If the coupling efficiency is 99%, then 1% failure sequences are obtained at each coupling. The ratio of the correct sequence to each shorter sequence is calculated to range from 89.5:1 to 99:1. Therefore, rather high base recognition can be expected. It is apparent that, with increasing
chain length of the DNA fragments, formation of N-branched DNA molecules increases exponentially. Therefore, the N-unprotected strategy has a certain limitation in this respect, although it seems that a 20-mer synthesis is at least achievable without significant difficulty. In addition to these advantages, there is a possibility that because the HOBT method can provide oligodeoxyribonucleotides with phosphotriester linkages directly on DNA microchips or microarrays, O-protected oligodeoxyribonucleotides can be used as neutral DNA probes that can bind to target DNA or RNA fragments of patients for gene diagnosis. This possibility should be tested in the near future. Several kinds of O-alkylated oligodeoxyribonucleotide derivatives maintain hybridization properties when bound to complementary oligodeoxyribonucleotides (Yamana et al., 1991; Iyer et al., 1995, 1996). For gene diagnosis using DNA microchips or microarrays, unlike gene therapy, there is no need to consider the reactivity of phosphotriester linkages as potential alkylating reagents toward target nucleic acids or other molecules such as proteins and lipids. In previous approaches to the synthesis of DNA oligomers containing base-labile functional groups, various N-protecting groups have been developed that are removable under neutral conditions (Dreef-Tromp et al., 1991; Iyer et al., 1995; Alvarez et al., 1999; Spinelli et al., 2002). The N-unblocked strategy described here would be useful as a more straightforward approach to base-labile modified DNA derivatives. For example, the Hphosphonate approach without base protection was successfully applied to the synthesis of oligodeoxyribonucleotides having base-labile N-acetyldeoxycytidine residues, which proved to form a Watson-Crick base pair with guanine more effectively than the unmodified base (Wada et al., 1998a). Several papers related to this application have also appeared (Wada et al., 2001a; Kobori et al., 2002b). Because the phosphoramidite approach uses tervalent phosphorous compounds as the monomer units, it is impossible to synthesize oligodeoxyribonucleotides having functions that react with them or the activated intermediates. For example, the azido group is a typical functional group that reacts with phosphoramidites or phosphites to undergo the Staudinger reaction. In such a case, the Hphosphonate approach without base protection is useful. An oligodeoxyribonucleotide derivative having 2-azidodeoxyadenosine was successfully synthesized by the BOMP strategy
Synthesis of Unmodified Oligonucleotides
3.10.11 Current Protocols in Nucleic Acid Chemistry
Supplement 18
(Wada et al., 2001b). On the other hand, the H-phosphonate method was used not only for isolation of base-labile H-phosphonate DNA oligomers (Wada et al., 1999) but also for the synthesis of hydroxymethylphosphonate DNA oligomers (Wada and Sekine, 1995). In the near future, the aforementioned N-unprotected strategies could be used for the synthesis of other targets, such as oligodeoxyribonucleotides having ester or aldehyde groups, that cannot be obtained directly by the use of modified deoxyribonucleoside 3 -phosphoramidites or 3 -phosphonates. For this purpose, suitable phosphate-protecting groups as well as linkers that are removable under neutral conditions should be developed. Some strategies described in recent papers (Kobori et al., 2002a; Ushioda et al., 2002) would be useful for further studies in this direction.
Literature Cited Adamiak, R.W. and Stawinski, J. 1977. A highly effective route to N,N -disubstituted ureas under mild conditions. An application to the synthesis of tRNA anticodon loop fragments containing ureidonucleosides. Tetrahedron Lett. 18:19351936. Adamiak, R.W., Biala, E., Grzeskowiak, K., Kierzek, R., Kraszewski, A., Markiewicz, W.T., Okupniak, J., Stawinski, J., and Wiewiorowski, M. 1978. The chemical synthesis of the anticodon loop of an eukaryotic initiator tRNA containing the hypermodified nucleoside N6 -/Nthreonylcarbonyl/-adenosine/t6 A/. Nucl. Acids Res. 5:1889-1905. Alvarez, K., Vasseur, J.-J., and Imbach, J.-L. 1999. Use of photolabile amino-protecting groups in the synthesis of base-sensitive DNA SATE-phosphotriesters. Nucleosides Nucleotides 18:1435-1436. Baddiley, J. and Todd, A.R. 1947. Nucleotides. I. Muscle adenylic acid and adenosine diphosphate. J. Chem. Soc. 648-651. Beaucage, S.L. and Iyer, R.P. 1992. Advances in the synthesis of oligonucleotides by the phosphoramidite approach. Tetrahedron 48:2223-2311. Boal, J.H., Wilk, A., Harindranath, N., Max, E.E., Kempe, T., and Beaucage, S.L. 1996. Cleavage of oligodeoxyribonucleotides from controlledpore glass supports and their rapid deprotection by gaseous amines. Nucl. Acids Res. 24:31153117. Butkus, V., Kayushin, A.L., Berlin, Y.A., Kolosov, M.N., and Smirnov, I.V. 1983. Cleavage of 5 -Oprotecting trityl groups in oligodeoxynucleotide synthesis. Effect of substrate structure and reaction conditions on detritylation and depurination rates. Bioorg. Khim. 9:1518-1530. DNA Synthesis Without Base Protection
Caruthers, M.H. 1991. Chemical synthesis of DNA and DNA analogues. Acc. Chem. Res. 24:278284.
Claesen, C., Tesser, G.I., Dreef, C.E., Marugg, J.E., van der Marel, G.A., and van Boom, J.H. 1984. Use of 2-methylsulfonylethyl as a phosphorus protecting group in oligonucleotide synthesis via a phosphite triester approach. Tetrahedron Lett. 25:1307-1310. Dreef-Tromp, C.M., Van Dam, E.M.A., van den Elst, H., van den Boogaart, J.E., van der Marel, G.A., and van Boom, J.H. 1991. Solid-phase synthesis of RNA via a silyl-protecting-group strategy. Recl. Trav. Chim. Pays Bas 110:378383. Finnan, J.L., Varshney, A., and Letsinger, R.L. 1980. Developments in the phosphite-triester method of synthesis of oligonucleotides. Nucl. Acids Res. Symp. Ser. 7:133-145. Fourrey, J.-L. and Varenne, J. 1985. Preparation and phosphorylation reactivity of N-nonacylated nucleoside phosphoramidites. Tetrahedron Lett. 26:2663-2666. Froehler, B.C. and Matteucci, M.D. 1983a. Dialkylformamidines: Depurination resistant N6 protecting group for deoxyadenosine. Nucl. Acids Res. 11:8031-8036. Froehler, B.C. and Matteucci, M.D. 1983b. Substituted 5-phenyltetrazoles: Improved activators of deoxynucleoside phosphoramidites in deoxyoligonucleotide synthesis. Tetrahedron Lett. 24:3171-3174. Gryaznov, S.M. and Letsinger, R.L. 1991. Synthesis of oligonucleotides via monomers with unprotected bases. J. Am. Chem. Soc. 113:5876-5877. Gryaznov, S.M. and Letsinger, R.L. 1992. Selective O-phosphitylation with nucleoside phosphoramidite reagents. Nucl. Acids Res. 20:18791882. Hall, R.H. 1971. The Modified Nucleosides in Nucleic Acids. Columbia University Press, New York. Hata, T. and Kurihara, T. 1973. Preparation of intermediates in the synthesis of polynucleotides. II. N4 -Benzoylation of deoxycytidylic and cytidylic acids by 2-chloromethyl-4-nitrophenyl benzoate. Chem. Lett. 8:859-862. Hayakawa, Y. 2001. Toward an ideal synthesis of oligonucleotides: Development of a novel phosphoramidite method with high capability. Bull. Chem. Soc. Jpn. 74:1547-1565. Hayakawa, Y. and Kataoka, M. 1998. Facile synthesis of oligodeoxyribonucleotides via the phosphoramidite method without nucleoside base protection. J. Am. Chem. Soc. 120:1239512401. Hayakawa, Y., Aso, Y., Uchiyama, M., and Noyori, R. 1983a. Facile nucleoside phosphorylation via hydroxyl activation. Tetrahedron Lett. 24:11651168. Hayakawa, Y., Aso, Y., Uchiyama, M., and Noyori, R. 1983b. Chemoselective phosphorylation of N-unprotected nucleosides via aluminum alkoxides. Tetrahedron Lett. 24:5641-5644. Hayakawa, Y., Uchiyama, M., and Noyori, R. 1984. A convenient method for the formation of
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internucleotide 25:4003-4006.
linkage.
Tetrahedron
Lett.
Hayakawa, Y., Uchiyama, M., Nobori, T., and Noyori, R. 1985. A convenient synthesis of 2 -5 linked oligoribonucleotides. Tetrahedron Lett. 26:761-764. Hayakawa, Y., Nobori, T., Noyori, R., and Imai, J. 1987. Synthesis of 2 -5 ,3 -5 linked triadenylates. Tetrahedron Lett. 28:2623-2626. Hayakawa, Y., Kataoka, M., and Noyori, R. 1996. Benzimidazolium triflate as an efficient promoter for nucleotide synthesis via the phosphoramidite method. J. Org. Chem. 61:7996-7997. Hayakawa, Y., Kawai, R., and Kataoka, M. 2001. Nucleotide synthesis via methods without nucleoside-base protection. Eur. J. Pharm. Sci. 13:5-16. Ikemoto, T., Haze, A., Hatano, H., Kitamoto, Y., Ishida, M., and Nara, K. 1995. Phosphorylation of nucleosides with phosphorus oxychloride in trialkyl phosphate. Chem. Pharm. Bull. 43:210215.
Ishido, R. 1989. Protection of 5 -hydroxy groups of cytidine derivatives. Jpn. Kokai Tokkyo Koho Jp Pat. 01308294; Chem. Abstr. 1990. 112:700. Iyer, R.P., Yu, D., Ho, N.-H., Devlin, T., and Agrawal, S. 1995. O- and S-Methyl phosphotriester oligonucleotides: Facile synthesis using N-pent-4-enoyl nucleoside phosphoramidites. J. Org. Chem. 60:8132-8133. Iyer, R.P., Yu, D., Devlin, T., Ho, N.-H., Johnson, S., and Agrawal, S. 1996. Synthesis, biophysical properties, and stability studies of mixed backbone oligonucleotides containing novel non-ionic linkages. Nucleosides Nucleotides 16:1491-1495. Kataoka, M. and Hayakawa, Y. 1999. A convenient method for the synthesis of N-free 5 -O(p,p -dimethoxytrityl)-2 -deoxyribonucleosides via 5 -O-selective tritylation of the parent substances. J. Org. Chem. 64:6087-6089. Kobori, A., Miyata, K., Ushioda, M., Seio, K., and Sekine, M. 2002a. A new silyl ether-type linker useful for the automated synthesis of oligonucleotides having base-labile protective groups. Chem. Lett. 31:16-17. Kobori, A., Miyata, K., Ushioda, M., Seio, K., and Sekine, M. 2002b. A new method for the synthesis of oligodeoxyribonucleotides containing 4N-alkoxycarbonyldeoxycytidine derivatives and their hybridization properties. J. Org. Chem. 67:476-485. Koole, L.H., Buck, H.M., Kanters, J.A., and Schouten, A. 1987. Molecular conformation of 2 -deoxy-3 ,5 -di-O-acetyladenosine. Crystal structure and high resolution proton nuclear magnetic resonance investigations. Can. J. Chem. 65:326-331. K¨oster, H., Stumpe, A., and Wolter, A. 1983. Polymer support oligonucleotide synthesis. 13. Rapid and efficient synthesis of oligodeoxynucleotides on porous glass support using triester approach. Tetrahedron Lett. 24:747-750.
Krotz, A.H., McElroy, B., Scozzari, A.N. Cole, D.L., and Ravikumar, V.T. 2003. Controlled detritylation of antisense oligonucleotides. Org. Process Res. Dev. 7:47-52. Kume, A., Iwase, R., Sekine, M., and Hata, T. 1984. Cyclic diacyl groups for protection of the N6 -amino group of deoxyadenosine in oligodeoxynucleotide synthesis. Nucl. Acids Res. 12:8525-8538. Kung, P.-P. and Jones, R.A. 1992. H-Phosphonate DNA synthesis without amino protection. Tetrahedron Lett. 33:5869-5872. Letsinger, R.L., Finnan, J.L., Jacobs, S.A., Juodka, B.A., and Varshney, A.K. 1976. Exploration of new procedures for the synthesis of polynucleotides. In Proceedings of the International Conference on Synthesis, Structure and Chemistry of Transfer Ribonucleic Acids and Their Components, Dymaczewo, Poland, September 13-17, 1976. Institute of Organic Chemistry, Polish Academy of Sciences. Levin, A.S., Tabatadze, E.R., and Komarova, N.I. 1991. Synthesis of oligodeoxyribonucleotides by H-phosphonate method using N-unblocked synthons. Sibirskii Khim. Z. 6:142-144 Marugg, J.E., Tromp, M., Kuyl-Yeheskiely, E., van der Marel, G.A., and van Boom, J.H. 1986. A convenient and general approach to the synthesis of properly protected d-nucleoside 3 hydrogen phosphonates via phosphite intermediates. Tetrahedron Lett. 27:2661-2664. McCollum, C. and Andrus, A. 1991. An optimized polystyrene support for rapid, efficient oligonucleotide synthesis. Tetrahedron Lett. 32:40694072. Michelson, M. and Todd, A.D. 1954. Nucleotides. Part XXIII. Mononucleotides derived from deoxycytidine. Note on the structure of cytidylic acids a and b. J. Chem. Soc. 34-40. Nikolenko, L.N., Nezavibat’ko, V.N., and Tolmacheva, N.S. 1967. Selective N-benzoylation of deoxycytidine and cytidine. Khim. Prir. Soedin. 3:359. Nishino, S., Nagato, Y., Hasegawa, Y., Yamamoto, H., Kamaike, K., and Ishido, Y. 1991. Partial protection of carbohydrate derivatives. Part 27. Efficient deanilidation of phosphoranilidates by the use of nitrites and acetic anhydride. Heteroatom. Chem. 2:187-196. Ogilvie, K.K. 1973. The tert-butyldimethylsilyl group as a protecting group in deoxynucleosides. Can. J. Chem. 51:3799-3807. Ogilvie, K.K. and Nemer, M. 1981. Nonaqueous oxidation of phosphites to phophates in nucleotide synthesis. Tetrahedron Lett. 22:2531-2532. Ohkubo, A., Ezawa, Y., Seio, K., and Sekine, M. 2002. A new strategy for the synthesis of oligodeoxynucleotides in the phosphoramidite method without base protection via phosphite intermediates. Nucl. Acids Res. Suppl. 2:2930. Okupniak, J., Adamiak R.W., and Wiewiorowski, M. 1981. New conditions for a selective
Synthesis of Unmodified Oligonucleotides
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introduction of trityl type protective group into adenosine and cytidine 3 -phosphates. Pol. J. Chem. 55:679-682.
Venner, H. 1964. Studies on nucleic acids. IX. Stability of the N-glycosidic linkage in nucleosides. Z. Physiol. Chem. 339:14-27.
Pearson, R.G. 1987. Recent advances in the concept of hard and soft acids and bases. J. Chem. Educ. 64:561-567.
Venner, H. 1966. Research on nucleic acids. XII. Stability of the N-glycoside bond of nucleotides. Z. Physiol. Chem. 344:189-196.
Pearson, R.G. 1990. Hard and soft acids and bases— the evolution of a chemical concept. Coord. Chem. Rev. 100:403-425.
Ven’yaminova, A.G., Komarova, N.I., Levin, A.S., and Repkova, M.N. 1988. Synthesis of ribonucleoside-3 - and -5 -H-phosphonates via salicylchlorophosphine. Bioorg. Khim. 14:484489.
Schaller, H., Weimann, G., Lerch, B., and Khorana, H.G. 1963. Studies on polynucleotides. XXIV. The stepwise synthesis of specific deoxyribopolynucleotides. Protected derivatives of deoxyribonucleosides and new syntheses of deoxyribonucleoside 3 -phosphates. J. Am. Chem. Soc. 85:3821-3827. Schulhof, J.C., Molko, D., and Teoule, R. 1987. Facile removal of new base protecting groups useful in oligonucleotide synthesis. Tetrahedron Lett. 28:51-54. Sekine, M., Matsuzaki, J., Satoh, M., and Hata, T. 1982. Improved 3 -O-phosphorylation of guanosine derivatives by O6 -oxygen protection. J. Org. Chem. 47:571-573. Sekine, M., Masuda, N., and Hata, T. 1985. Introduction of the 4,4 ,4 -tris(benzoyloxy)trityl group into the exo amino groups of deoxyribonucleosides and its properties. Tetrahedron 41:5445-5453. Sekine, M., Ohkubo, A., and Seio, K. 2003. Protonblock strategy for the synthesis of oligodeoxynucleotides without base protection, capping reaction, and P-N bond cleavage reaction. J. Org. Chem. 68:5478-5492. Septak, M. 1996. Kinetic studies on depurination and detritylation of CPG-bound intermediates during oligonucleotide synthesis. Nucl. Acids Res. 24:3053-3058. Shena, M. 1999. DNA Microarrays: A practical approach. Oxford University Press, London. Spinelli, N., Meyer, A., Hayakawa, Y., Imbach, J.L., and Vasseur, J.-J. 2002. Use of allylic protecting groups for the synthesis of base-sensitive prooligonucleotides. Eur. J. Org. Chem. 4956. Tanaka, T. and Letsinger, R.L. 1982. Syringe method for stepwise chemical synthesis of oligonucleotides. Nucl. Acids Res. 10:32493260. Ti, G.S., Gaffney, B.L., and Jones, R.A. 1982. Transient protection: Efficient one-flask syntheses of protected deoxynucleosides. J. Am. Chem. Soc. 104:1316-1319.
DNA Synthesis Without Base Protection
Ushioda, M., Kadokura, M., Moriguchi, T., Kobori, A., Aoyagi, M., Seio, K., and Sekine, M. 2002. Unique participation of unprotected internucleotidic phosphodiester residues on unexpected cleavage reaction of the Si-O bond of the diisopropylsilandiyl group used as a linker for the solid-phase synthesis of 5 -terminal guanylated oligodeoxynucleotides. Helv. Chim. Acta 85:2930-2945.
Wada, T. and Sekine, M. 1995. Chemical synthesis of oligothymidylate having hydroxymethylphosphonate internucleotidic linkages. Tetrahedron Lett. 36:845-848. Wada, T., Sato, Y., Honda, F., Kawahara, S., and Sekine, M. 1997. Chemical synthesis of oligodeoxyribonucleotides using N-unprotected H-phosphonate monomers and carbonium and phosphonium condensing reagents: O-Selective phosphonylation and condensation. J. Am. Chem. Soc. 119:12710-12721. Wada, T., Kobori, A., Kawahara, S., and Sekine, M. 1998a. Synthesis and properties of oligodeoxyribonucleotides containing 4-N-acetylcytosine bases. Tetrahedron Lett. 39:6907-6910. Wada, T., Mochizuki, A., Sato, Y., and Sekine, M. 1998b. Functionalization of solid supports with N-unprotected deoxyribonucleosides. Tetrahedron Lett. 39:5593-5596. Wada, T., Honda, F., Sato, Y., and Sekine, M. 1999. First synthesis of H-phosphonate oligonucleotides bearing N-unmodified bases. Tetrahedron Lett. 40:915-918. Wada, T., Kobori, A., Kawahara, S., and Sekine, M. 2001a. Synthesis and hybridization ability of oligodeoxyribonucleotides incorporating N-acyldeoxycytidine derivatives. Eur. J. Org. Chem. 4583-4593. Wada, T., Mochizuki, A., Higashiya, S., Tsuruoka, H., Kawahara, S., Ishikawa, M., and Sekine, M. 2001b. Synthesis and properties of 2-azidodeoxyadenosine and its incorporation into oligodeoxynucleotides. Tetrahedron Lett. 42:9215-9219. Watanabe, K.A. and Fox, J.J. 1966. Simple method for selective acylation of cytidine on the 4-amino group. Angew. Chem. Int. Ed. Engl. 5:579-580. Yamana, K., Nishijima, Y., Negishi, K., Yoshiki, T., Nishio, K., and Nakano, H. 1991. Deoxyribonucleoside 3 -phosphorobisamidites in the synthesis of isopropyl phosphotriester oligodeoxyribonucleotide analogs. Tetrahedron Lett. 32:4721-4724. Yoshikawa, M. and Kato, T. 1967. Studies of phosphorylation. I. Phosphorylation of 2 ,3 -Oisopropylidene nucleoside by phosphoryl chloride. Bull. Chem. Soc. Jpn. 40:2849-2853. Yuodka, Y., Lunsford, W.B., and Letsinger, R.L. 1976. Oligonucleotides and nucleotidepeptides. XXV. New method for the synthesis of oligonucleotides. Bioorg. Chem. 2:12181324.
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Current Protocols in Nucleic Acid Chemistry
Internet Resource http://www.chfi.unipd.it/-chor/wisor/ lecture notes/Bagno acidbase.pdf A list of the basicity of common solvents, maintained by A. Bagno, Centro CNR, Padova, Italy.
Contributed by Mitsuo Sekine Tokyo Institute of Technology Yokohama, Japan
Synthesis of Unmodified Oligonucleotides
3.10.15 Current Protocols in Nucleic Acid Chemistry
Supplement 18
The 4-Methylthio-1-Butyl Group for Phosphate/Thiophosphate Protection in Oligodeoxyribonucleotide Synthesis
UNIT 3.11
This unit provides a detailed preparation of deoxyribonucleoside phosphoramidites functionalized with a 4-methylthio-1-butyl group for P(III) protection, and describes their use in the solid-phase synthesis of DNA oligonucleotides. Specifically, the efficiency of these phosphoramidites in the synthesis of a 20-mer DNA oligonucleotide and its phosphorothioated analog is demonstrated. The thermolytic cleavage of the 4-methylthio-1-butyl protecting group from these oligonucleotides, at neutral pH or under the standard basic conditions used for nucleobase deprotection, proceeds through a cyclodeesterification mechanism with the concomitant formation of a cyclic sulfonium salt (Cie´slak et al., 2004). The 4-methylthio-1-butyl protecting group is distinct from previously studied thermolytic groups (UNITS 2.7 & 3.9; Grajkowski et al., 2001) in terms of structural simplicity and ease of deprotection. Thus, this unit describes an attractive method for preparation of alkylation-free oligonucleotides for potential therapeutic applications, while bringing closer to reality the implementation of a “heat-driven” process for oligonucleotide synthesis on microarrays.
PREPARATION OF 4-METHYLTHIO-1-BUTYL DEOXYRIBONUCLEOSIDE PHOSPHORAMIDITES AND THEIR USE IN SOLID-PHASE DNA SYNTHESIS
BASIC PROTOCOL
The 4-methylthio-1-butyl group for phosphate/thiophosphate protection can be thermolytically cleaved from oligonucleotides at a lower temperature and at a faster rate than the 4-oxopentyl (UNIT 3.9; Wilk et al., 2001), 3-(N-tert-butylcarboxamido)-1-propyl (UNIT 3.9; Wilk et al., 2002), and 2-(N-formyl,N-methyl)aminoethyl (Grajkowski et al., 2001) groups under neutral conditions. Moreover, the 4-methylthio-1-butyl group can also be removed from oligonucleotides under conditions (NH4 OH, 10 hr at 55◦ C) that are not suitable for removal of 3-(N-tert-butylcarboxamido)-1-propyl and 2-(N-formyl,Nmethyl)aminoethyl groups. As illustrated in Figure 3.11.1, this protocol outlines a general method for the preparation of deoxyribonucleoside phosphoramidites (S.3a-d) from commercial 5 -O- and N-protected deoxyribonucleosides (S.1a-d), O-(4-methylthio-1butyl)-N,N,N ,N -tetraisopropylphosphorodiamidite (S.2; see Support Protocol), and 1Htetrazole. Procedures for the use of S.3a-d in solid-phase DNA synthesis and for the analysis of synthetic oligonucleotides are also described.
Materials Protected deoxyribonucleosides (S.1a-d; Chem-Impex International): 5 -O-(4,4 -dimethoxytrityl)-2 -deoxythymidine N4 -benzoyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxycytidine N6 -benzoyl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxyadenosine N2 -isobutyryl-5 -O-(4,4 -dimethoxytrityl)-2 -deoxyguanosine Argon source Anhydrous dichloromethane (CH2 Cl2 ; Aldrich) O-(4-Methylthio-1-butyl)-N,N,N ,N -tetraisopropylphosphorodiamidite (S.2; see Support Protocol) 1H-Tetrazole, sublimed Triethylamine (TEA; Aldrich) Benzene (Aldrich)
Contributed by Jacek Cie´slak, Andrzej Grajkowski, Victor Livengood, and Serge L. Beaucage Current Protocols in Nucleic Acid Chemistry (2004) 3.11.1-3.11.14 C 2004 by John Wiley & Sons, Inc. Copyright
Synthesis of Unmodified Oligonucleotides
3.11.1 Supplement 19
Figure 3.11.1 Preparation of deoxyribonucleoside phosphoramidites S.3a-d. DMTr, 4,4 dimethoxytrityl; i-Pr, isopropyl. Adapted and reprinted from Cie´slak et al. (2004) with permission from the American Chemical Society.
4-Methylthio-1Butyl Group for Phosphate/ Thiophosphate Protection
Silica gel (60 Å, 230 to 400 mesh; Merck) Hexane (−20◦ C) Dry ice/acetone bath Reagents recommended for automated solid-phase oligonucleotide synthesis (Applied Biosystems and/or Glen Research): Standard 2-cyanoethyl deoxyribonucleoside phosphoramidites (T, CBz , ABz and Gi-Bu ) Activator solution: 1H-tetrazole in acetonitrile Oxidation solution: 0.02 M iodine in THF/pyridine/water Cap A solution: acetic anhydride in THF/pyridine Cap B solution: 1-methylimidazole in THF Deblocking solution: trichloroacetic acid in dichloromethane Acetonitrile 3H-1,2-Benzodithiol-3-one-1,1-dioxide (Glen Research) Concentrated ammonium hydroxide (NH4 OH; Fisher Scientific) 2 M triethylammonium acetate (TEAA) buffer, pH 7.0 (Applied Biosystems) 80% acetic acid (AcOH) Loading buffer: 1:4 (v/v) 10× TBE electrophoresis buffer (APPENDIX 2A) in formamide containing 2 mg/mL bromphenol blue 20 × 40–cm, 7 M urea/20% polyacrylamide gel (UNIT 10.4 and APPENDIX 3B) Staining buffer: 1:5:20:0.1 (v/v/v/v) formamide/isopropyl alcohol/ddH2 O/3.0 M Tris·Cl, pH 8.8 1 mg/mL Stains-All (Aldrich) in formamide 1.0 M Tris·Cl, pH 9.0 (APPENDIX 2A) 1.0 M MgCl2 (Sigma) Snake venom phosphodiesterase (SVPD, Crotalus adamanteus; Sigma) Bacterial alkaline phosphatase (BAP, E. coli; Sigma) 50-, 100-, and 250-mL round-bottom flasks Rubber septa for 14/20- and 24/40-glass joints Vacuum desiccator High-vacuum oil pump 1- and 10-mL glass syringes 5-mm NMR tube Rotary evaporator connected to a vacuum pump 2.5 × 20–cm disposable Flex chromatography columns (Kontes) 2.5 × 7.5–cm EMD TLC plates precoated with a 250-µm layer of silica gel 60 F254 Lyophilizer 392 DNA/RNA synthesizer (Applied Biosystems) 1-mL Luer-tipped syringes
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4-mL screw-cap glass vials Heating block (VWR) 25 cm × 4.6–mm Supelcosil LC-18S HPLC column (5-µm; Supelco) 1.5-mL microcentrifuge tubes 37◦ C water bath Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D), column chromatography (APPENDIX 3E), automated DNA synthesis (APPENDIX 3C), DNA quantification by UV spectrophotometry (UNIT 10.3), reversed-phase HPLC (UNIT 10.5), and polyacrylamide gel electrophoresis (PAGE; UNIT 10.4 & APPENDIX 3B) Prepare deoxyribonucleoside phosphoramidites 1. Dry 2.2 mmol of a suitably protected deoxyribonucleoside (S.1a-d) by exposure to high vacuum for 2 hr at 25◦ C in a 50-mL round-bottom flask. Add a stir bar and seal flask with a rubber septum. 2. Under an argon atmosphere, add 10 mL anhydrous dichloromethane using a 10-mL glass syringe and ∼700 mg (∼2 mmol) O-(4-methylthio-1-butyl)-N,N,N ,N tetraisopropylphosphorodiamidite (S.2) via a 1-mL glass syringe. Stir the solution with a magnetic stirrer. 3. While stirring the solution under a positive pressure of argon, remove the rubber septum from the flask and quickly add 112 mg (1.6 mmol) sublimed 1H-tetrazole using a spatula. Seal the flask with the rubber septum. Other activators such as ethylthio-1H-tetrazole and dicyanoimidazole could theoretically be used, but have not been tested.
4. Carefully transfer ∼0.5 mL of the reaction mixture by syringe to a dry 5-mm NMR tube. Monitor the progress of the reaction in the NMR tube by 31 P NMR spectroscopy. Upon completion of the reaction, as seen by the disappearance of S.2, return the NMR sample to the main reaction mixture before proceeding with the next step. The 31 P NMR signal corresponding to S.2 (δ P 121 ppm) disappears within 2 hr at ambient temperature and thus indicates complete phosphinylation of S.1a-d.
5. Add 1 mL triethylamine and immediately concentrate the reaction mixture to a syrup using a rotary evaporator connected to a vacuum pump. 6. Suspend the crude product in ∼3 mL of 9:1 (v/v) benzene/triethylamine and apply the suspension to a 2.5 × 20–cm disposable Flex chromatography column containing ∼20 g silica gel that has been equilibrated in 9:1 (v/v) benzene/triethylamine (APPENDIX 3E). 7. For A, C, and T derivatives, elute the column with 9:1 (v/v) benzene/triethylamine and collect 6-mL fractions. For the G derivative, elute with 45:45:10 (v/v/v) benzene/dichloromethane/triethylamine. 8. Analyze fractions by TLC (APPENDIX 3D) on a 2.5 × 7.5–cm EMD silica gel 60 F254 TLC plate using 9:1 (v/v) benzene/triethylamine as the eluent. Pool appropriate fractions and rotoevaporate under reduced pressure (∼20 mmHg) until a white amorphous solid is obtained. TLC analysis of diastereomeric S.3a-d shows two tight spots. The Rf of each pair of spots ranges between 0.1 and 0.6 when 9:1 (v/v) benzene/triethylamine is used as the eluent. For the G derivative using 45:45:10 benzene/dichloromethane/triethylamine, the Rf is ∼0.3.
9. Dissolve the solid in ∼3 mL benzene and add the solution to ∼100 mL vigorously stirred cold (−20◦ C) hexane in a 250-mL round-bottom flask.
Synthesis of Unmodified Oligonucleotides
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10. Allow the suspension to settle and carefully decant off most of the supernatant. 11. Rotoevaporate the wet material to dryness under reduced pressure and then dissolve in ∼10 mL benzene. 12. Transfer the solution to a 100-mL round-bottom flask, freeze in a dry ice/acetone bath, and lyophilize under high vacuum to afford triethylamine-free S.3a-d as white powders in yields ranging from 72% to 85%. 5 -O-(4,4 -Dimethoxytrityl)-3 -O-[(N,N-diisopropylamino)(4-methylthio-1-butyloxy)] phosphinyl-2 -deoxythymidine S.3a: 31 P NMR (121 MHz, CDCl3 ): δ 147.6, 147.9 Fast-atom bombardment high-resolution mass spectrometry (FAB-HRMS): anal. calcd. for C42 H56 N3 O8 PS (M+Cs)+ 926.2580, found 926.2537. N4 -Benzoyl-5 -O-(4,4 -dimethoxytrityl)-3 -O-[(N,N-diisopropylamino)(4-methylthio-1butyloxy)]phosphinyl-2 -deoxycytidine S.3b: 31 P NMR (121 MHz, CDCl3 ): δ 147.5, 147.9. FAB-HRMS: anal. calcd. for C48 H59 N4 O8 PS (M+Cs)+ 1015.2846, found 1015.2870. N6 -Benzoyl-5 -O-(4,4 -dimethoxytrityl)-3 -O-[(N,N-diisopropylamino)(4-methylthio1-butyloxy)]phosphinyl-2 -deoxyadenosine S.3c: 31 P NMR (121 MHz, CDCl3 ): δ 147.3, 147.7. FAB-HRMS: anal. calcd. for C49 H59 N6 O7 PS (M+Cs)+ 1039.2958, found 1039.2996. N2 -Isobutyryl-5 -O-(4,4 -dimethoxytrityl)-3 -O-[(N,N-diisopropylamino)(4-methylthio1-butyloxy)]phosphinyl-2 -deoxyguanosine S.3d: 31 P-NMR (121 MHz, CDCl3 ): δ 147.4, 147.8. FAB-HRMS: anal. calcd. for C46 H61 N6 O8 PS (M+Cs)+ 1021.3064, found 1021.3011. It is common practice to mix cesium iodide, potassium iodide, and/or sodium iodide with a sample to be analyzed by FAB-HRMS. The accuracy in mass determination can be better ascertained through the association of cesium (M+Cs)+ , potassium (M+K)+ , or sodium (M+Na)+ with the parent ion. The accurate mass is not always easy to determine as a (M+H)+ ion when the sample is dissolved in the traditional glycerol matrix in the absence of metallic salts.
Synthesize oligonucleotides 13. Perform a 0.2-µmol scale solid-phase synthesis of both d(ATCCGTAGCTAAGGTCATGC) and its phosphorothioate analog using an Applied Biosystems 392 DNA/RNA synthesizer in “trityl-on” mode according to the manufacturer’s recommendations (also see APPENDIX 3C). Perform parallel syntheses employing exclusively commercial 2-cyanoethyl deoxyribonucleoside phosphoramidites to serve as reference standards for comparing the overall quality of the synthetic oligonucleotides. As in the case of 2-cyanoethyl deoxyribonucleoside phosphoramidites, S.3a-d are used as 0.1 M solutions in dry acetonitrile. All ancillary reagents required for the automated preparation of oligonucleotides were purchased from Applied Biosystems and/or Glen Research and used as recommended by the manufacturer(s). The synthetic cycle recommended for the preparation of phosphorothioated DNA oligonucleotides differs from the conventional cycle used for the synthesis of unmodified oligonucleotides in that the “capping step” is performed after the oxidative sulfurization reaction (Iyer et al., 1990). The sulfur transfer reaction is effected by treatment with 0.05 M 3H1,2-benzodithiol-3-one-1,1-dioxide in acetonitrile using a wait step of 30 sec.
4-Methylthio-1Butyl Group for Phosphate/ Thiophosphate Protection
Cleave oligonucleotide from support 14. Using two 1-mL Luer-tipped syringes, one attached to each end of the synthesis column, push back and forth 0.5 mL of concentrated NH4 OH three times through the column every 10 min over a 30-min period. Transfer the solution to a 4-mL screwcap glass vial. Push back and forth 0.5 mL of fresh NH4 OH for 1 min through the synthesis column and pour the wash into the glass vial.
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Complete oligonucleotide deprotection 15. Place the tightly capped vial containing each oligonucleotide solution in a heating block pre-heated to 55◦ C ± 2◦ C and continue heating at this temperature for 10 hr. Cool each sample to room temperature. 16. Determine the concentration of the oligonucleotide by UV spectrophotometry at 260 nm (UNIT 10.3).
Analyze oligonucleotide by RP-HPLC 17. Analyze ∼0.25 OD260 of each oligonucleotide on a 5-µm Supelcosil LC-18S HPLC column (UNIT 10.5) using a linear gradient of 1% acetonitrile/min starting from 0.1 M triethylammonium acetate, pH 7.0, at a flow rate of 1 mL/min for 40 min. Hold the gradient isocratically for 20 min. RP-HPLC profiles of oligonucleotides synthesized from S.3a-d and conventional 2-cyanoethyl deoxyribonucleoside phosphoramidites are presented in Figure 3.11.2. The two peaks for the PS-oligonucleotides are caused by the presence of a terminal 5 -DMTr group, which facilitates the hydrophobic resolution of two populations of diastereomers (RP -rich and SP -rich). The resolution of these diastereomeric populations is dependent
Figure 3.11.2 RP-HPLC chromatograms of crude 5 -O-DMTr-d(ATCCGTAGCTAAGGTCATGC) and 5 -O-DMTr-d(APS TPS CPS CPS GPS TPS APS GPS CPS TPS APS APS GPS GPS TPS CPS APS TPS GPS C). (A) 5 -O-DMTr-20-mer and (C) PS-5 -O-DMTr-20-mer synthesized via phosphoramidites S.3a-d and deprotected as described in the Basic Protocol. (B) 5 -O-DMTr-20-mer and (D) PS-5 -O-DMTr20-mer synthesized from commercial 2-cyanoethyl deoxyribonucleoside phosphoramidites and deprotected as described in Basic Protocol, steps 14 and 15. RP-HPLC analyses were performed as described in Basic Protocol, step 17. DMTr, 4,4 -dimethoxytrityl; PS, phosphorothioate diester. Adapted and reprinted from Cie´slak et al. (2004) with permission from the American Chemical Society.
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on the sequence and size of the oligomer and will completely vanish upon removal of the 5 -DMTr group under acidic conditions. This phenomenon is not well understood, although it has consistently been observed in several laboratories.
Analyze oligonucleotide by PAGE 18. Pipet 0.25 OD260 of each oligonucleotide solution into a 1.5-mL microcentrifuge tube. Add 100 µL of 80% AcOH. After 30 min, evaporate to dryness using a Speedvac. 19. Add 10 µL of loading buffer. Vortex well and then centrifuge 5 sec at 14,000 × g. 20. Load the sample in a 2-cm-wide well of a 20 × 40–cm, 7 M urea/20% polyacrylamide gel (UNIT 10.4 and APPENDIX 3B). Electrophorese at 350 V until the bromphenol blue dye travels 75% of the length of the gel. 21. Dismantle the gel apparatus and immerse the gel in 250 mL staining buffer containing 10 mL of 1 mg/mL Stains-All in formamide. Agitate the gel for ∼1 hr in the dark. 22. Discard the staining solution and rinse the gel three times with 250 mL distilled water. 23. Expose the gel to natural light until the purple background disappears and photograph the blue bands against a white background. A photograph of the gel is presented in Figure 3.11.3.
4-Methylthio-1Butyl Group for Phosphate/ Thiophosphate Protection
Figure 3.11.3 PAGE analysis of d(ATCCGTAGCTAAGGTCATGC) and its phosphorothioated analog under denaturing conditions (7 M urea, 1× TBE buffer, pH 8.3). (A) Crude oligomer synthesized from commercial 2-cyanoethyl deoxyribonucleoside phosphoramidites. (B) Crude oligomer synthesized from phosphoramidites S.3a-d. (C) Crude phosphorothioated oligomer synthesized from S.3a-d. All were deprotected as described in Basic Protocol, steps 14, 15, and 18. Unmodified oligonucleotides are visualized as blue bands and fully thioated oligonucleotides as purple bands upon staining the gel with Stains-All. Bromphenol blue is used as a marker and shows as a large band at the bottom of each lane. Adapted and reprinted from Cie´slak et al. (2004) with permission from the American Chemical Society.
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Characterize oligonucleotide by enzyme hydrolysis 24. Pipet 1 OD260 of each oligonucleotide solution into a 1.5-mL microcentrifuge tube. Add 100 µL of 80% AcOH. After 30 min, evaporate to dryness using a Speedvac. 25. Add 6 µL of 1.0 M Tris·Cl, pH 9.0, 8 µL of 1.0 M MgCl2 , and 75 µL water. Mix well. 26. Add 5 µL (0.015 U) SVPD and 6 µL (0.7 U) BAP and heat 16 hr in a 37◦ C water bath (also see UNIT 10.6). 27. Heat deactivate the enzyme 3 min at 90◦ C. Centrifuge 5 min at 14,000 × g, 25◦ C. 28. Analyze a 50-µL aliquot of the digest by RP-HPLC as in step 17. The RP-HPLC profile of the enzyme hydrolysates is shown in Figure 3.11.4.
Figure 3.11.4 RP-HPLC analysis of crude d(ATCCGTAGCTAAGGTCATGC) after digestion with SVPD and BAP (37◦ C, 16 hr). Hydrolysates were made from (A) a 20-mer synthesized using phosphoramidites S.3a-d and (B) a 20-mer synthesized using commercial 2-cyanoethyl deoxyribonucleoside phosphoramidites. Both were deprotected as described in Basic Protocol, steps 14 and 15. RP-HPLC analyses were performed as described in Basic Protocol, step 17. Identities of the RP-HPLC peaks are confirmed through co-injection with authentic 2 -deoxyribonucleosides. Adapted and reprinted from Cie´slak et al. (2004) with permission from the American Chemical Society.
Figure 3.11.5 121 MHz 31 P NMR spectrum of crude d(APS TPS CPS CPS GPS TPS APS GPS CPS TPS APS APS GPS GPS TPS CPS APS TPS GPS C) in NH4 OH. The crude 20-mer was prepared from S.3a-d and deprotected as described in Basic Protocol, steps 14 and 15. Adapted and reprinted from Cie´slak et al. (2004) with permission from the American Chemical Society.
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Analyze phosphorothioated oligonucleotide by 31 P NMR spectroscopy 29. Analyze the crude phosphorothioated oligonucleotide (synthesized using S.3a-d and deprotected as described in steps 14 and 15) in NH4 OH by 31 P NMR spectroscopy. A 31 P NMR spectrum of phosphorothioated d(ATCCGTAGCTAAGGTCATGC) is shown in Figure 3.11.5. The crude oligomer appears as a broad singlet at ∼52 ppm. SUPPORT PROTOCOL
PREPARATION OF O-(4-METHYLTHIO-1-BUTYL)-N,N,N ,N TETRAISOPROPYLPHOSPHORODIAMIDITE This protocol delineates the preparation of bis(N,N-diisopropylamino)chlorophosphine (S.4) in situ from the reaction of phosphorus trichloride and N,N-diisopropylamine, and its condensation with 4-methylthio-1-butanol (S.5) to produce the phosphorodiamidite S.2 (see Fig. 3.11.6).
Additional Materials (also see Basic Protocol) Phosphorus trichloride (Aldrich) N,N-Diisopropylamine (Aldrich) 4-Methylthio-1-butanol (Aldrich) Drierite, 8 mesh (Aldrich) 10-µm stainless-steel frit (20 × 10–mm o.d.; Kontes) 3-mm-i.d. Teflon canula 1. To 100 mL of dry benzene in an oven-dried 250-mL round-bottom flask, add by syringe through a rubber septum 1.75 mL (20 mmol) of freshly distilled phosphorus trichloride under a dry argon atmosphere. 2. Cool the stirred solution to 5◦ C using an ice bath. Add by syringe 17.6 mL (140 mmol) anhydrous N,N-diisopropylamine over a period of 30 min under an argon atmosphere. 3. Remove the ice bath and allow the stirred reaction to warm to 25◦ C under a positive pressure of argon until the formation of bis(N,N-diisopropylamino)chlorophosphine (S.4) is complete. The progress of the reaction is monitored by 31 P NMR spectroscopy. The formation of S.4 (singlet, 135.5 ppm, downfield relative to a phosphoric acid external standard) is essentially complete (>96%) within 72 hr.
4. Add by syringe 2.7 mL (22 mmol) of 4-methylthio-1-butanol (S.5) to the suspension. Stir for 2 hr at 25◦ C under a positive pressure of argon. 31
P NMR spectrum of reaction mixture shows a singlet at 118.5 ppm indicating the near quantitative formation of S.2.
5. Using argon pressure, filter the suspension through a 10-µm stainless steel frit (20 × 10–mm o.d.) coupled to a 3-mm-i.d. Teflon canula into a 250-mL round-bottom flask. Wash the salt with 40 mL of dry benzene and resume filtration. To release gas pressure, pierce a 20-G needle through the rubber septum of the receiving flask. 6. Rotoevaporate the filtrates under reduced pressure to afford S.2 as an oil.
4-Methylthio-1Butyl Group for Phosphate/ Thiophosphate Protection
Crude O-(4-methylthio-1-butyl)-N,N,N ,N -tetraisopropylphosphorodiamidite (S.2) can be used for the phosphinylation of S.1a-d without further purification. 1 H NMR (300 MHz, C6 D6 ): δ 3.56 (m, 2H), 3.53 (sept, J = 6.9 Hz, 2H), 3.49 (sept, J = 6.9 Hz, 2H), 2.30 (m, 2H), 1.80 (s, 3H), 1.63 (m, 4H), 1.23 (d, J = 6.9 Hz, 12H), 1.19 (d, J = 6.9 Hz, 12H). 13 C NMR (75 MHz, C6 D6 ): δ 15.2, 24.0, 24.1, 24.7, 24.8, 26.2, 31.1 (d, JPC = 9.6 Hz), 34.2, 44.6, 44.7, 64.1 (d, 2 JPC = 21.5 Hz). 31 P NMR (121 MHz, C6 D6 ): δ 118.5.
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Figure 3.11.6 Preparation of O-(4-methylthio-1-butyl)-N,N,N ,N -tetraisopropylphosphorodiamidite (S.2). Adapted and reprinted from Cie´slak et al. (2004) with permission from the American Chemical Society.
COMMENTARY Background Information The thermolabile 4-methylthio-1-butyl phosphate/thiophosphate-protecting group for DNA oligonucleotides has been investigated for its potential application to oligonucleotide synthesis on diagnostic microarrays and to the large-scale preparation of alkylation-free therapeutic oligonucleotides. With the advent of DNA microarrays as powerful diagnostic tools in clinical medicine (Rhodes and Chinnaiyan, 2002; Guo, 2003), the development of thermolytic groups for 5 -hydroxyl (Chmielewski et al., 2003) and phosphate (Grajkowski et al., 2001; Wilk et al., 2001, 2002; Cie´slak and Beaucage, 2003) protection toward the implementation of a “heat-driven” method (Grajkowski et al., 2003) for the synthesis of DNA oligonucleotides on planar glass surfaces is being investigated. Thermolabile 5 -/3 -hydroxyl- and phosphate-protecting groups are designed for rapid and efficient release from oligonucleotides when heated under neutral conditions. Such properties eliminate the use of harsh chemicals that are typically required for iterative oligonucleotide chain extension and final oligonucleotide deprotection. These chemicals, however, may damage the glass surface and result in oligonucleotide leaching, which would lead to a poorer performance of microarrays in terms of diagnostic sensitivity. In contrast, the large-scale preparation of oligonucleotides for therapeutic applications has challenged the suitability of 2-cyanoethyl deoxyribonucleoside phosphoramidites (Sinha et al., 1984) for such syntheses, given the potent DNA-alkylating properties of acrylonitrile (Wilk et al., 1999a,b) that is generated under the conditions used for nucleobase deprotection. DNA alkylation can permanently compromise the affinity of therapeutic oligonucleotides for their mRNA targets. To circumvent such a limitation, a number
of deoxyribonucleoside phosphoramidites functionalized with phosphorus-protecting groups that would prevent formation of DNAalkylating species during oligonucleotide deprotection have been reported. These groups include the 4-[(N-methyl,N-(2,2,2trifluoroacetyl)amino]butyl group (Wilk et al., 1999a,b; UNIT 2.7) and the 4-oxopentyl group (Wilk et al., 2001; UNIT 3.9), which can easily be removed from oligonucleotides by standard treatment with concentrated ammonium hydroxide or pressurized ammonia gas (Boal et al., 1996). The removal of these phosphate/thiophosphate-protecting groups occurred through an intramolecular cyclodeesterification reaction with the concomitant formation of side-products that were inoccuous to the nucleobases of DNA oligonucleotides (Wilk et al., 1999a,b; 2001). Deoxyribonucleoside phosphoramidites functionalized with phosphorus-protecting groups exhibiting thermolytic properties were also reported. Groups such as 2-(N-formyl,Nmethyl)aminoethyl (Grajkowski et al., 2001) and 3-[(N-tert-butyl)carboxamido]-1-propyl (Wilk et al., 2002) were cleaved from N-deprotected oligonucleotides upon heating at temperatures up to 90◦ C in an aqueous buffer at neutral pH. Under these conditions, the removal of these phosphate/thiophosphateprotecting groups was achieved within 3 hr and 1 hr, respectively. The thermolytic cleavage of the 3-[(N-tert-butyl)carboxamido]-1-propyl group is consistent with the participation of the amidic carbonyl group in an intramolecular cyclodeesterification reaction resulting in the release of a phosphate/thiophosphate diester as its 2-tert-butyliminotetrahydrofuran salt (UNIT 3.9; Wilk et al., 2002). The spatial arrangement of the amidic carbonyl group relative to the phosphate/thiophosphate leaving group is crucial in that it determines the thermolytic deprotection kinetics of the protecting group.
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Figure 3.11.7 Proposed mechanism for the cleavage of the 4-methylthio-1-butyl group under thermolytic conditions at neutral pH. Reagents and conditions: (i) 0.45 M 1H-tetrazole in MeCN, 1 min; (ii) 0.02 M I2 in THF/pyridine/water or 0.05 M 3H-benzodithiol-3-one-1,1-dioxide in MeCN, 3 min; (iii) 3% TCA in CH2 Cl2 , 1 min; (iv) MeNH2 gas (2.5 bar), 3 min; (v) 0.1 M TEAA, pH 7.0, 55◦ C, 30 min. LCAA-CPG, succinyl long-chain alkylamine controlled-pore glass; TCA, trichloroacetic acid; Thy, thymin-1-yl; TEAA, triethylammonium acetate; THF, tetrahydrofuran. Adapted and reprinted from Cie´slak et al. (2004) with permission from the American Chemical Society.
4-Methylthio-1Butyl Group for Phosphate/ Thiophosphate Protection
Indeed, the thermolytic deprotection of the homologous 3-[(N-tert-butyl)carboxamido]1-butyl and 2-(N-tert-butylcarboxamido)ethyl thiophosphate-protecting group is, respectively, 25-fold and >2000-fold slower than that of the 3-[(N-tert-butyl)carboxamido]1-propyl group under identical conditions (Wilk et al., 2002). Thus, it is critically important to consider the spatial relationship between the nucleophilic function and the phosphate/thiophosphate leaving group when designing novel thermolytic groups. These findings prompted the search for phosphate/thiophosphate-protecting groups that would be removed under milder thermolytic conditions in either the presence or absence of ammonium hydroxide. The selection of the 4-methythio-1-butyl group was based on its structural simplicity and its potential compliance with the spatial requirements for optimal thermolytic deprotection kinetics, assuming an intramolecular cyclodeesterification process. The preparation of deoxyribonucleoside phosphoramidites functionalized with the 4-methylthio-1-butyl group for P(III) protection (S.3a-d) began with the synthesis of the phosphinylating
reagent S.2, which was achieved upon reaction of the chlorophosphine S.4 with commercial 4-methylthio-1-butanol (S.5; see Support Protocol). Condensation of commercial nucleosides (S.1a-d) with crude S.2 in the presence of 1H-tetrazole afforded the deoxyribonucleosides phosphoramidites S.3a-d (see Basic Protocol). The thermolytic properties of the 4methylthio-1-butyl group were evaluated using a dinucleotide model. Specifically, the reaction of phosphoramidite S.3a with thymidine covalently attached to long-chain alkylamine controlled-pore glass (LCAA-CPG, S.6) and 1H-tetrazole gave the dinucleoside phosphite triester S.7 (Fig. 3.11.7). Exposure of S.7 to an iodine solution or to 3H-1,2benzodithiol-3-one-1,1-dioxide in MeCN produced the support-bound dinucleoside phosphotriester S.8 or S.9, respectively (Cie´slak et al., 2004). Removal of the 5 -O-DMTr group under acidic conditions followed by release of the dinucleotide from LCAA-CPG upon treatment with methylamine gas afforded S.10 or S.11 as the major product. RP-HPLC analysis of crude S.10 revealed the presence of the corresponding dinucleoside phosphodiester S.12
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to the extent of ∼10%, which is indicative of the facile removal of the 4-methylthio1-butyl phosphate protecting group. Heating either S.10 or S.11 in 0.1 M triethylammonium acetate (pH 7.0) for 30 min at 55◦ C, resulted in complete cleavage of the 4-methylthio-1-butyl group to give the dinucleoside phosphodiester S.12 or S.13 in near quantitative yield (>99%) on the basis of RP-HPLC analysis of the thermolytic deprotection reaction (Cie´slak et al., 2004). Alternatively, treatment of S.8 or S.9 with concentrated NH4 OH for 30 min at room temperature, followed by heating the ammoniacal solution for 2 hr at 55◦ C, produced comparable results. As shown in Figure 3.11.7, the thermolytic cleavage of the 4-methylthio-1-butyl group from S.10 or S.11 proceeds through an intramolecular cyclodeesterification reaction with the production of a sulfonium salt (S.14). To confirm the formation of S.14, the solidphase-linked dinucleoside phosphotriester S.8 was suspended in 0.1 M NaCl in D2 O and
heated for 30 min at 55◦ C. Analysis of the supernatant by 1 H NMR spectroscopy (Fig. 3.11.8A) revealed signals identical to those recorded for S.14 (Fig. 3.11.8B), which was also prepared as its chloride salt from the reaction of tetrahydrothiophene with methyl chloroformate (Byrne and Lafleur Lawter, 1986). These data convincingly demonstrate the participation of the 4-methylthio function in the thermolytic phosphate/thiophosphate deprotection of S.10 or S.11 (Cie´slak et al., 2004). The use of the 4-methylthio-1-butyl group for phosphate protection in solid-phase oligonucleotide synthesis was adequate only if the conventional 0.02 M iodine solution in THF/pyridine/H2 O was selected as the oxidant for iteratively converting phosphite to phosphate triester throughout oligonucleotide synthesis. The use of tert-butyl hydroperoxide or (1S)-(+)-(10-camphorsulfonyl)oxaziridine instead of iodine resulted in the partial conversion of the 4-methylthio-1-butyl group to its 4methylsulfinyl-1-butyl congener, which could
Figure 3.11.8 300 MHz 1 H NMR spectra of S.14 (as its chloride salt in D2 O). (A) S.14 produced by thermolytic deprotection of S.8 in D2 O containing 0.1 M NaCl. (B) S.14 prepared by reaction of tetrahydrothiophene with methyl chloroformate. Adapted and reprinted from Cie´slak et al. (2004) with permission from the American Chemical Society.
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not be thermolytically cleaved from the parent dinucleoside phosphotriester even upon heating for 3 hr at 90◦ C in a neutral buffer (Cie´slak et al., 2004). These results further validate the participation of the 4-methylthio function in the thermolytic phosphate/thiophosphate deprotection of S.10 or S.11 through an intramolecular cyclodeesterification process. The deoxyribonucleoside phosphoramidites S.3a-d were successfully used in the small scale (0.2-µmol) solid-phase synthesis of d(ATCCGTAGCTAAGGTCATGC) and its phosphorothioated analog. Following release from the CPG support, the thermolytic cleavage of the 4-methylthio-1-butyl protecting group from crude oligonucleotides was performed along with complete removal of nucleobase-protecting groups by treatment with concentrated NH4 OH for 10 hr at 55◦ C. Neither nucleobase alkylation (see Fig. 3.11.4) nor significant phosphorothioate desulfurization were detected under these conditions. To further substantiate the absence of nucleobase alkylation under conditions that would simulate millimole-scale oligonucleotide deprotection, deoxyribonucleosides and their Nprotected counterparts were heated in concentrated NH4 OH for 10 hr at 55◦ C in the presence or absence of a representative concentration of the sulfonium salt S.14 (Cie´slak et al., 2004). RP-HPLC analyses of each reaction mixture did not show significant nucleobase modifica-
4-Methylthio-1Butyl Group for Phosphate/ Thiophosphate Protection
tions under these conditions (Fig. 3.11.9). Exposure of O,O-diethyl thiophosphate to S.14 under concentration conditions similar to those prevailing in large-scale oligonucleotide synthesis did not show, on the basis of 31 P NMR data, desulfurization of the phosphorothioate model in concentrated NH4 OH over a period of 10 hr at 55◦ C. Typically, no conversion of O,Odiethyl thiophosphate (δP ∼55 ppm) to O,Odiethyl phosphate (δP ∼2 ppm) was detected (see Fig. 3.11.10). Altogether, the results reported therein indicate that the 4-methythio1-butyl group is suitable for the large-scale preparation of therapeutic oligonucleotides as well as for the microscale synthesis of oligonucleotides on diagnostic microarrays.
Critical Parameters and Troubleshooting The phosphorodiamidite S.2 is prepared using the bis(N,N-diisopropylamino)chlorophosphine intermediate S.4 (see Support Protocol, Fig. 3.11.6) in a manner similar to that reported for N,N,N ,N -tetraisopropyl-O[3-(N-tert-butylcarboxamino)-1-propyl]phosphorodiamidite (UNIT 3.9, Figure 3.9.5). Readers are referred to the Critical Parameters and Troubleshooting section of UNIT 2.7, which addresses issues pertaining to the preparation of phosphorodiamidites. The formation of S.4 must be carefully monitored by 31 P NMR spectroscopy to ensure complete exhaustion of dichloro(N,N-diisopropylamino)phosphine
Figure 3.11.9 RP-HPLC analysis of the DNA-modifying properties of S.14. (A) Chromatogram of the reaction of T, dC, dA, dG, N 4 -Bz-dC, N 6 -Bz-dA, and N 2 -i-Bu-dG with S.14 in NH4 OH under conditions simulating large-scale oligonucleotide deprotection. (B) Chromatogram of an identical reaction in the absence of S.14. Chromatographic conditions for RP-HPLC analyses are as described in UNIT 3.9. Adapted and reprinted from Cie´slak et al. (2004) with permission from the American Chemical Society.
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Figure 3.11.10 31 P NMR analysis (121 MHz) of the phosphorothioate desulfurization properties of S.14. (A) Reaction of O,O-diethyl thiophosphate (potassium salt) with S.14 under conditions simulating large-scale oligonucleotide deprotection in concentrated NH4 OH. (B) Potassium salt of O,O-diethyl thiophosphate in concentrated NH4 OH. Adapted and reprinted from Cie´slak et al. (2004) with permission from the American Chemical Society.
(δP ∼169 ppm in C6 D6 ) prior to adding 4-methylthio-1-butanol. It is recommended that the deoxyribonucleoside phosphoramidites S.3a-d be precipitated from cold hexane following silica gel purification. This precipitation step removes a hydrolyzed phosphinylating agent that often co-elutes with the desired phosphoramidite monomers. To ensure optimal coupling efficiency of individual deoxyribonucleoside phosphoramidites during solid-phase oligonucleotide synthesis, a number of parameters including exclusion of residual triethylamine and moisture must be taken into consideration. These issues are also addressed in UNIT 2.7. When using S.3a-d in solid-phase oligonucleotide synthesis, oxidants such as tert-butyl hydroperoxide and (1S)-(+)(10-camphorsulfonyl)oxaziridine should be avoided during chain assembly.
Anticipated Results The synthesis of deoxyribonucleoside phosphoramidites S.3a-d from phosphorodiamidite S.2 is performed essentially as recommended by Barone et al. (1984). The coupling efficiency of silica gel–purified and triethylamine-free phosphoramidites S.3a-d is comparable to that of commercial 2cyanoethyl deoxyribonucleoside phosphoramidites.
phosphoramidites S.3a-d require 1 to 2 days to complete.
Literature Cited Barone, A.D., Tang, J.-T., and Caruthers, M.H. 1984. In situ activation of bisdialkylaminophosphines—A new method for synthesizing deoxyoligonucleotides on polymer supports. Nucl. Acids Res. 12:4051-4061. Boal, J.H., Wilk, A., Harindranath, N., Max, E.E., Kempe, T., and Beaucage, S.L. 1996. Cleavage of oligodeoxyribonucleotides from controlledpore glass supports and their rapid deprotection by gaseous amines. Nucl. Acids Res. 24:31153117. Byrne, B. and Lafleur Lawter, L.M. 1986. The preparation of trimethylsulfonium chloride from methyl chloroformate and dimethyl sulfide. Tetrahedron Lett. 27:1233-1236. Chmielewski, M.K., March´an, V., Cie´slak, J., Grajkowski, A., Livengood, V., M¨unch, U., Wilk, A., and Beaucage, S.L. 2003. Thermolytic carbonates for potential 5 -hydroxyl protection of deoxyribonucleosides. J. Org. Chem. 68:10003-10012. Cie´slak, J. and Beaucage, S.L. 2003. Thermolytic properties of 3-(2-pyridyl)-1-propyl and 2-[N-methyl-N-(2-pyridyl)]aminoethyl phosphate/thiophosphate protecting groups in solid-phase synthesis of oligodeoxyribonucleotides. J. Org. Chem. 68:10123-10129.
Time Considerations
Cie´slak, J., Grajkowski, A., Livengood, V., and Beaucage, S.L. 2004. Thermolytic 4-methylthio-1-butyl group for phosphate/thiophosphate protection in solid-phase synthesis of DNA oligonucleotides. J. Org. Chem. 69:2509-2515.
The preparation of the phosphorodiamidite S.2 takes 3 to 4 days, whereas synthesis and purification of each of the deoxyribonucleoside
Grajkowski, A., Wilk, A., Chmielewski, M.K., Phillips, L.R., and Beaucage, S.L. 2001. The 2-(N-formyl,N-methyl)aminoethyl group as a
Synthesis of Unmodified Oligonucleotides
3.11.13 Current Protocols in Nucleic Acid Chemistry
Supplement 19
potential phosphate/thiophosphate protecting group in solid-phase oligodeoxyribonucleotide synthesis. Org. Lett. 3:1287-1290. Grajkowski, A., Cie´slak, J., Chmielewski, M.K., March´an, V., Phillips, L.R., Wilk, A., and Beaucage, S.L. 2003. Conceptual “heat-driven” approach to the synthesis of DNA oligonucleotides on microarrays. Ann. N.Y. Acad. Sci. 1002:1-11.
Wilk, A., Grajkowski, A., Srinivasachar, K., and Beaucage, S.L. 1999b. Improved chemistry for the production of synthetic oligodeoxyribonucleotides. Antisense Nucleic Acid Drug Dev. 9:361-366.
Guo, Q.M. 2003. DNA microarray and cancer. Curr. Opin. Oncol. 15:36-43.
Wilk, A., Chmielewski, M.K., Grajkowski, A., Phillips, L.R., and Beaucage, S.L. 2001. The 4-oxopentyl group as a labile phosphate/thiophosphate protecting group for synthetic oligodeoxyribonucleotides. Tetrahedron Lett. 42:5635-5639.
Iyer, R.P., Phillips, L.R., Egan, W., Regan, J.B., and Beaucage, S.L. 1990. The automated synthesis of sulfur-containing oligodeoxyribonucleotides using 3H-1,2-benzodithiol-3-one-1,1-dioxide as a sulfur-transfer reagent. J. Org. Chem. 55:46934699.
Wilk, A., Chmielewski, M.K., Grajkowski, A., Phillips, L.R., and Beaucage, S.L. 2002. The 3[(N-tert-butyl)carboxamido]-1-propyl group as an attractive phosphate/thiophosphate protecting group for solid-phase oligodeoxyribonucleotide synthesis. J. Org. Chem. 67:6430-6438.
Rhodes, D.R. and Chinnaiyan, A.M. 2002. DNA microarrays: Implications for clinical medicine. J. Invest. Surg. 15:275-279. Sinha, N.D., Biernat, J., McManus, J., and K¨oster, H. 1984. Polymer support oligonucleotide synthesis. 18. Use of β-cyanoethyl-N,N,dialkylamino/-N-morpholino phosphoramidite of deoxynucleosides for the synthesis of DNA fragments simplifying deprotection and isolation of the final product. Nucl. Acids Res. 12:4539-4557.
Contributed by Jacek Cie´slak, Andrzej Grajkowski, and Serge L. Beaucage Food and Drug Administration Bethesda, Maryland Victor Livengood National Institutes of Health Bethesda, Maryland
Wilk, A., Grajkowski, A., Phillips, L.R., and Beaucage, S.L. 1999a. The 4-[N-methyl-N(2,2,2-trifluoroacetyl)amino]butyl group as an alternative to the 2-cyanoethyl group for phosphate protection in the synthesis of oligodeoxyribonucleotides. J. Org. Chem. 64:75157522.
4-Methylthio-1Butyl Group for Phosphate/ Thiophosphate Protection
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Nucleoside Phosphoramidites Containing Cleavable Linkers
UNIT 3.12
The first step in any solid-phase oligonucleotide synthesis is attachment of the first nucleoside to the surface of the support (see UNIT 3.2). Typically, this attachment is made via an ester linkage connecting the 3 -OH position to one end of a dicarboxylic acid linker arm. The other end of the dicarboxylic acid is anchored to an amino- or hydroxyl-derivatized solid-phase support through an amide or ester bond, respectively. Since the chemistry for forming ester and amide linkages is not compatible with the phosphoramidite coupling chemistry used on automated synthesizers (see APPENDIX 3C), the nucleoside-derivatized support is generally prepared in a separate process, and an inventory of separate prederivatized supports (one for each base) must be prepared or purchased. For many nucleosides, derivatized supports are conveniently prepackaged into synthesis columns ready for use on column-based synthesizers. Multi-well plate-based synthesizers (Lashkari et al., 1995; Sindelar and Jaklevic, 1995; Brennan et al., 1998; Rayner et al., 1998; Cheng et al., 2002) present an added challenge because they require placement of the correct prederivatized supports into the correct wells according to each set of sequences being made. Because this can be a tedious and error-prone step, much effort has gone into developing “universal” supports that can be used in any well, regardless of the nucleoside required. Two strategies have been developed to allow this universal functionality. In the first strategy (Fig. 3.12.1A), universal supports with various diol linker structures S.2 are employed (Debear et al., 1987; Scott et al., 1994; Schwartz et al., 1995; Nelson et al., 1997; Scheuerlarsen et al., 1997; Azhayev, 1999; Lyttle et al., 1999; Azhayev and Antopolsky, 2001; Guzaev and Manoharan, 2003). The first nucleoside is added using a conventional nucleoside-3 O-phosphoramidite (S.1), which forms a phosphodiester linkage to the universal support (S.3). After synthesis, the relatively stable phosphate ester linkage connecting the oligonucleotide to the support must be selectively cleaved to generate a product with a 3 -OH and not a 3 -phosphate terminus. The difficulty of this selective cleavage is the main drawback to this approach. In this unit, a second approach to universal supports is described (Fig. 3.12.1B) that uses modified phosphoramidite reagents (linker phosphoramidites, S.4; Pon and Yu, 2001, 2004) instead of modified solid-phase supports. These modified phosphoramidites contain the same protected nucleosides and the same activatable phosphoramidite groups used in conventional synthesis. However, an easily cleavable linker is inserted between the 3 -OH of the nucleoside and the phosphoramidite group in such a manner that oligonucleotide products with the desired 3 -OH ends can be generated without any additional cleavage or deprotection steps. This method also allows much less expensive, underivatized amino supports S.5 to act as universal supports. Basic Protocol 1 describes the synthesis of the four linker phosphoramidite reagents S.4a-d starting from readily available protected nucleosides. The Alternate Protocol provides a method for preparing simpler and slightly less costly reagents in which the sulfonyldiethanol group is replaced by ethylene glycol. Although DNA synthesis requires a set of four linker phosphoramidites (one for each base), these reagents are useful in two different ways, as will be discussed below. Both methods for synthesis of linker phosphoramidites can be adapted for use with modified nucleosides; however, the cost and effort may outway the benefits if the modified linker phosphoramidites will not be used frequently.
Contributed by Richard T. Pon Current Protocols in Nucleic Acid Chemistry (2005) 3.12.1-3.12.24 C 2005 by John Wiley & Sons, Inc. Copyright
Synthesis of Unmodified Oligonucleotides
3.12.1 Supplement 23
Figure 3.12.1 Comparison of universal support strategies. In route (A), a conventional 3 -O-phosphoramidite reagent is coupled to a CPG support with a modified diol linker arm. After completion of the synthesis, a difficult-to-cleave phosphate P-O bond must be hydrolyzed to release an oligonucleotide with the desired 3 -OH terminus. In route (B), a modified linker phosphoramidite reagent is coupled to an underivatized amino CPG support. After completion of the synthesis, the desired oligonucleotide product can be released by hydrolysis of a more easily cleaved ester C-O bond.
Basic Protocol 2 describes the use of linker phosphoramidite reagents for oligonucleotide synthesis in multi-well plates using inexpensive underivatized amino supports (S.5) in a procedure that eliminates the problems associated with 3 -dephosphorylation.
Nucleoside Phosphoramidites Containing Cleavable Linkers
Finally, in Basic Protocol 3, the linker phosphoramidites containing a sulfonyldiethanol group (S.4) are used to create long strings of oligonucleotides linked end-to-end via tandem oligonucleotide synthesis (Pon and Yu, 2005). The ability to make sets of oligonucleotides in a single automated tandem synthesis with linker phosphoramidite reagents is a major advantage when large sets of oligonucleotides are required, i.e., for PCR primers or double-stranded sequences. However, tandem synthesis is best used for applications
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that are tolerant of minor impurities, since it is usually not practical to isolate and purify each component in the mixture. Fortunately, phosphoramidite coupling chemistry is efficient enough to allow most oligonucleotides to be used with only minimal purification (such as a desalting step), and the small amounts of truncated failure sequences present do not significantly affect their use in many applications.
SYNTHESIS OF LINKER PHOSPHORAMIDITES CONTAINING A CLEAVABLE SUCCINYL-SULFONYLDIETHANOL LINKER
BASIC PROTOCOL 1
This protocol describes the three synthetic steps (Fig. 3.12.2) required to convert protected 5 -O-dimethoxytrityl-N-acyl-2 -deoxyribonucleosides S.8a-d into linker phosphoramidite reagents S.4a-d. The first step involves reaction of the 3 -OH position with succinic anhydride S.9 to create nucleoside-3 -O-succinyl hemiesters S.10a-d. Then, the terminal carboxylic acid group is esterified with sulfonyldiethanol S.11 to create nucleosides with a 3 -O-linker ending in a hydroxyl group S.12a-d. Finally, nucleoside S.12 is converted to the desired linker phosphoramidite compound S.4a-d
Figure 3.12.2 Synthesis of linker phosphoramidites S.4a-d containing a 2,2 -sulfonyldiethanol spacer (see Basic Protocol 1). Synthesis of Unmodified Oligonucleotides
3.12.3 Current Protocols in Nucleic Acid Chemistry
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by phosphitylation of the terminal hydroxyl group with 2-cyanoethyl-N,N,N ,N tetraisopropylphosphorodiamidite S.13 and tetrazole S.14. The synthetic manipulations required for this procedure are quite straightforward. The conditions required for the succinylation and phosphitylation steps are widely used in the preparation of conventional solid-phase supports and 3 -O-phosphoramidite reagents. Only the addition of sulfonyldiethanol as a spacer between the succinic acid and phosphoramidite groups is unique to this protocol.
Materials Protected 2 -deoxyribonucleosides: 5 -O-(4,4 -Dimethoxytrityl)-N6 -benzoyl-2 -deoxyadenosine (S.8a) 5 -O-(4,4 -Dimethoxytrityl)-N4 -acetyl-2 -deoxycytidine or 5 -O-(4,4 -dimethoxytrityl)-N4 -benzoyl-2 -deoxycytidine (S.8b) 5 -O-(4,4 -Dimethoxytrityl)-N2 -isobutyryl-2 -deoxyguanosine (S.8c) 5 -O-(4,4 -Dimethoxytrityl)thymidine (S.8d) Succinic anhydride (S.9) Dichloromethane Triethylamine Methanol Chloroform (CHCl3 ) 0.5 M triethylammonium phosphate (TEAP) solution (see recipe) 60% to 65% (w/v) 2,2 -sulfonyldiethanol (S.11) solution in water Acetonitrile, anhydrous Pyridine, anhydrous 4-Dimethylaminopyridine (DMAP) O-Benzotriazol-1-yl-N,N,N ,N -tetramethyluronium hexafluorophosphate (HBTU) Diisopropylethylamine (DIPEA) Silica gel Diisopropylamine, anhydrous 2-Cyanoethyl-N,N,N ,N -tetraisopropylphosphorodiamidite (S.13) 0.45 M 1H-tetrazole (S.14) in anhydrous acetonitrile 5% (w/v) sodium bicarbonate in water Saturated aqueous sodium chloride Hexanes, distilled (remove dissolved oxygen by distilling under vacuum on a rotary evaporator just prior to use) 100-, 250-, and 500-mL round-bottom flasks Magnetic stirrer Fluorescent silica gel TLC plates, Merck 60 or similar 500-mL separatory funnels Rotary evaporator and warm water bath (60◦ C) Chromatography column, ∼3 in. (∼7.5 cm) diameter, with capacity for at least 9 in. (∼23 cm) silica gel Rubber septum Drying tube: an empty 10- or 20-mL syringe filled with Drierite (with indicator), with a small glass wool plug at each end Syringes Additional reagents and equipment for TLC (APPENDIX 3D) and flash chromatography (APPENDIX 3E) Nucleoside Phosphoramidites Containing Cleavable Linkers
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Couple nucleoside to succinic anhydride 1. For each protected nucleoside S.8a-d, combine 10 mmol nucleoside and 1.5 g (15 mmol) succinic anhydride (S.9) in a 250-mL round-bottom flask equipped with a magnetic stirrer. 2. Add 100 mL dichloromethane and then 4.2 mL (30 mmol) triethylamine. Stir at room temperature for 3 hr. 3. Monitor the reaction by TLC (APPENDIX 3D) using 5% and/or 10% (v/v) methanol/chloroform to verify the complete conversion of the starting nucleoside (faster moving) into the succinylated product (very slow moving). 4. If the starting nucleoside remains after 3 hr, continue stirring at room temperature. The reaction may take 5 hr or longer. The reaction can also be sped up by adding additional succinic anhydride (up to a total of 20 or 25 mmol).
5. When the reaction is complete, transfer the dichloromethane solution into a 500-mL separatory funnel, add 200 mL of 0.5 M aqueous TEAP solution, and mix. Aqueous TEAP is used to form the triethylammonium salt of the succinate hemiester S.10. It also helps to prevent an intractable emulsion from forming in the separatory funnel.
6. Remove the organic (lower) phase from the separatory funnel, and wash the aqueous TEAP phase two times with 100 mL dichloromethane. 7. Evaporate the combined dichloromethane solutions to furnish the nucleoside-3 -Osuccinate hemiester S.10 as a white foam. 8. Dry overnight under vacuum. The dried hemiester can be stored as a dry powder at room temperature, and can be used in the next step without further purification. Yield = 107%. Characterization data can be found in Gait et al. (1980) and Kumar et al. (1993).
Couple sulfonyldiethanol spacer to hemiester 9. Place 9.6 mL (50 mmol) of 65% (w/v) 2,2 -sulfonyldiethanol solution (S.11) and 50 mL anhydrous acetonitrile in a 500-mL round-bottom flask. 10. Using a rotary evaporator and a warm water bath (60◦ C), evaporate the solution until the solvent no longer comes off. Add another 50-mL portion of anhydrous acetonitrile and repeat the evaporation. 11. Repeat the evaporation step two more times using 30 mL (each) anhydrous pyridine. The purpose of the repeated co-evaporations is to remove all traces of water from the 2,2 -sulfonyldiethanol.
12. Redissolve the now anhydrous 2,2 -sulfonyldiethanol in 80 mL anhydrous pyridine and stir at room temperature to form a clear solution. 13. Add 10 mmol S.10, 1.59 g (13 mmol) DMAP, 4.93 g (13 mmol) HBTU, and 2.3 mL (13 mmol) DIPEA, and stir to dissolve. Continue stirring at room temperature for 10 min. 14. Check the reaction by TLC using 5% or 10% (v/v) methanol/CHCl3 . All of the starting material (very slow moving) should have been converted into a fastermoving spot (Rf ∼0.4 to 0.6).
15. When the reaction is complete (no starting material remains), concentrate the solution on a rotary evaporator until a slurry begins to form.
Synthesis of Unmodified Oligonucleotides
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Table 3.12.1
1
H NMR Chemical Shiftsa for Nucleoside Derivatives S.12a-d
S.12a (B = ABz )
S.12b (B = CBz )
S.12c (B = Gi-Bu )
S.12d (B = T)
6.54–6.50
6.29–6.25
6.22–6.15
6.40
H3 (br d, 1H)
5.58–5.56
5.44–5.42
5.55–5.53
5.49
-CO-O-CH2 -CH2 - SO2 - (t, 2H)
4.62–4.59
4.62–4.61
4.63
4.59
4.33
4.31–4.30 4.20–4.14
4.11
H1 (t, 1H)
H4 (br s, 1H)
H4 & -SO2 -CH2 -CH2 -OH (m, 3H) -SO2 -CH2 -CH2 -OH (br dd, 2H)
4.10–4.09
4.14–4.12
-OCH3 ×2 (s, 6H)
3.79
3.82–3.80
3.80–3.78
3.79
3.60–3.33
3.49
H5 , H5 & -CO-O-CH2 -CH2 -SO2 - (m, 4H)
3.50–3.46
3.55–3.48
-SO2 -CH2 -CH2 -OH (t, 2H)
3.28–3.25
3.31–3.28
-CH(CH3 )2 (m, 1H)
3.29 3.05–2.96
H2 (m, 1H)
3.08–3.00
2.79–2.77
2.65–2.59
2.49
H2 (m, 1H)
2.72–2.66
2.36–2.33
2.36–2.27
2.49
-CO-CH2 -CH2 -CO- (br s, 4H)
2.72
2.69
2.71
2.68
-CH3 (s, 3H)
1.37
CH3 -CH-CH3 (d, 3H)
1.13–1.10
CH3 -CH-CH3 (d, 3H)
1.05–1.03
a The internal reference for 1 H NMR spectra was tetramethylsilane at 0 ppm. The solvent was CDCl . All shifts are measured in ppm. 3
16. Remove the flask from the evaporator, redissolve the residue in 200 mL chloroform, and transfer to a 500-mL separatory funnel. Wash the chloroform solution four times with 200 mL water. 17. Evaporate the chloroform solution to dryness to yield the crude product S.12. 18. Purify the crude product by flash chromatography (APPENDIX 3E) on silica gel using a column of ∼3 in. (∼7.5 cm) diameter, containing ∼9 in. (∼23 cm) of silica gel, packed using a slurry in CHCl3 . Dissolve the crude product in 10 mL CHCl3 and apply to the column. Elute the column progressively with 1%, 2%, and 3% (v/v) MeOH/CHCl3 until the product elutes. 19. Monitor the fractions by TLC as above. Combine fractions containing the pure product S.12 and evaporate to dryness to yield a white or slightly yellowish foam. Expected yield: 70%–80%. For characterization data, see Table 3.12.1. Store as a dry powder at room temperature.
Synthesize linker phosphoramidite 20. Place 5 mmol nucleoside S.12 in a 100-mL round-bottom flask equipped with a magnetic stirrer. Seal the flask with a rubber septum and insert a drying tube filled with Drierite. It is important to exclude atmospheric moisture from this reaction. All liquid transfers are done using syringes.
Nucleoside Phosphoramidites Containing Cleavable Linkers
CAUTION: The drying tube filled with Drierite is essential to act as a vent for the flask. Do not forget to have this in place, especially when adding reagents.
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Table 3.12.2 TLC, UV, and 31 P NMR Chemical Shift Data for Phosphoramidites S.4a-d
S.4b (B = CBz )
S.4c (B = Gi-Bu )
S.4d (B = T)
EtOAc
10% TEA/CHCl3
EtOAc
0.50
0.67
0.75
0.25
UV (nm)
234.5, 280.5
236.5, 260.0, 305.5
237.0, 267.0, 276.0
267.0
31
150.85
150.88
150.97, 151.04
146.50
S.4a (B = ABz ) TLC solvent 10% TEA/CHCl3 Rf a
b
P NMR
a The solvent for UV spectra was 95% ethanol. b The solvent for 31 P NMR spectra was CDCl . Shifts are measured in ppm from H PO . 3 3 4
21. Add 60 mL anhydrous acetonitrile and stir at room temperature to dissolve. 22. While still stirring, add 0.82 mL (6 mmol) anhydrous diisopropylamine, 2.4 mL (7.5 mmol) 2-cyanoethyl-N,N,N ,N -tetraisopropylphosphorodiamidite (S.13), and finally 12.2 mL (5.5 mmol) 0.45 M 1H-tetrazole (S.14) in anhydrous acetonitrile. 23. Stir at room temperature for 2.5 hr and then remove an aliquot and analyze by TLC using 10% (v/v) triethylamine/chloroform. The starting nucleoside S.12 should be entirely consumed and converted into a slightly slower migrating product S.4.
24. When the reaction is complete, dilute with 150 mL dichloromethane and transfer to a 500-mL separatory funnel. 25. Wash the reaction mixture once with 200 mL of 5% (w/v) aqueous sodium bicarbonate and then twice with 200 mL saturated aqueous sodium chloride. 26. Concentrate the dichloromethane solution. 27. Purify by flash chromatography on silica gel. Use a solution of 42:52:6 (v/v/v) dichloromethane/hexanes/triethylamine to prepare the column and to load the crude sample. Elute the product with 5% to 6% (v/v) triethylamine/dichloromethane. It is important to pack the silica gel column using a solution containing triethylamine to neutralize the surface acidity of the silica gel before applying the crude phosphoramidite. The use of freshly degassed (distilled) hexanes is also recommended during this step to minimize accidental oxidation of the phosphoramidite.
28. Monitor fractions by TLC (Table 3.12.2). Combine the fractions containing the pure product S.4 and evaporate to dryness to produce a white foam. Expected yield: 70%–80%. For characterization data, see Table 3.12.2. Store as dry powders in screw-capped containers at –20◦ C for up to 1 year. The linker phosphoramidites (S.4 and S.16) have shorter solution stability than conventional phosphoramidites. The guanosine phosphoramidites (S.4c and S.16c) are the most sensitive, and should be used within 3 to 5 days of preparation. The other linker phosphoramidite solutions should be used within 1 to 2 weeks of preparation.
REPLACEMENT OF SULFONYLDIETHANOL WITH ETHYLENE GLYCOL 2,2 -sulfonyldiethanol
This alternative protocol replaces the labile spacer with a much simpler 1,2-ethanediol group (S.16; Fig. 3.12.3). Other similar alkyl diols such as 1,3propanediol or 1,4-butanediol can also be used. These alkyl diols are somewhat less expensive than sulfonyldiethanol and produce linker phosphoramidites with slightly greater stability. The resulting phosphoramidites are suitable for attachment to LCAACPG for standard oligonucleotide syntheses (see Basic Protocol 2), but should not be used for tandem syntheses (for further discussion, see Basic Protocol 3).
ALTERNATE PROTOCOL
Synthesis of Unmodified Oligonucleotides
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Supplement 23
Figure 3.12.3 Protocol).
Synthesis of linker phosphoramidites S.16a-d containing an ethylene glycol spacer (see Alternate
Additional Materials (also see Basic Protocol 1) Nucleoside-3 -O-succinate hemiester (S.10; see Basic Protocol 1) p-Toluenesulfonyl chloride (p-TsCl) 1-Methylimidazole, anhydrous (N-methylimidazole; NMI) Ethylene glycol, anhydrous 250-mL separatory funnel 1. In a 250-mL round-bottom flask equipped with a magnetic stirrer, dissolve 4.4 mmol nucleoside-3 -O-succinate hemiester S.10 in 50 mL anhydrous acetonitrile and 2.9 mL (35 mmol) pyridine. 2. Add 1.64 g (8.6 mmol) p-TsCl and then 1.3 mL (15.8 mmol) NMI. Stir at room temperature to dissolve completely. 3. Add 0.25 mL (4.4 mmol) anhydrous ethylene glycol and continue stirring another 20 min at room temperature. 4. Concentrate the reaction mixture by evaporating off the solvent. 5. Redissolve the residue in 50 mL chloroform and transfer to a 250-mL separatory funnel. Nucleoside Phosphoramidites Containing Cleavable Linkers
6. Wash the chloroform solution with 50 mL water, 50 mL saturated aqueous NaCl solution, and then 50 mL water again.
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7. Concentrate the crude material to a thick oil and then purify by flash chromatography on silica gel using 0% to 5% (v/v) methanol/chloroform (first 0%, then 2.5%, and finally 5%). 8. Monitor by TLC using 5% methanol/chloroform. Combine fractions containing the pure material S.15 (Rf = 0.40) and evaporate to dryness to form a foam. Expected yield: 70%–80%. Nucleosides S.15a-d are converted into phosphoramidites S.16a-d as described in Basic Protocol 1 for conversion of S.12 to S.4. TLC solvents and Rf values for S.16a-d are the same as for S.4a-d.
OLIGONUCLEOTIDE SYNTHESIS IN MULTI-WELL PLATES ON UNDERIVATIZED LCAA-CPG SUPPORTS
BASIC PROTOCOL 2
This protocol describes the synthesis of oligonucleotides in 96-well synthesis plates using an underivatized LCAA-CPG support instead of diol-based universal supports (S.2; Fig. 3.12.1). This protocol was developed using a MerMade 192 synthesizer (BioAutomation) equipped with ten phosphoramidite ports (A, C, G, T, and six additional positions). Other synthesizers can be similarly used, as long as there are sufficient phosphoramidite ports to accommodate four regular phosphoramidites and four linker phosphoramidites. A four-column ABI 394 synthesizer (Applied Biosystems) equipped with eight phosphoramidite ports has also been used successfully, especially when performing tandem oligonucleotide synthesis (see Basic Protocol 3). With this column-based instrument, empty plastic synthesis columns are used instead of 96-well plates to hold the LCAACPG support.
Materials Linker phosphoramidite reagents: S.4a-d (see Basic Protocol 1) or S.16a-d (see Alternate Protocol) Anhydrous acetonitrile Reagents for oligonucleotide synthesis: conventional 3 -O-phosphoramidites activator, deprotection, oxidizing, and capping solutions (APPENDIX 3C) Long-chain alkylamine controlled-pore glass (LCAA-CPG) support, ∼100 µmol/g ◦ amino group loading, 500 A pore size (S.5) 1:1 (v/v) ammonium hydroxide/40% (w/v) aqueous methylamine Automated DNA synthesizer for multi-well plates, with at least 8 monomer ports (e.g., MerMade 192 synthesizer, BioAutomation) 96-well Oro-Flex OF1100 polypropylene filter plate (0.7-mL wells; Orochem Technologies) with 96-well collection plate (1-mL wells) Centrifuge with 96-well plate adapters Clamping apparatus for multi-well plates (as recommended by synthesizer manufacturer) 65◦ C oven Centrifugal evaporator (e.g., Speedvac, Savant) that can accommodate 96-well plates Additional reagents and equipment for oligonucleotide synthesis (APPENDIX 3C) and for “dry” loading of the support (see Support Protocol) Set up and run synthesizer 1. Prepare solutions of the linker phosphoramidites S.4a-d or S.16a-d at the desired concentration (0.025 to 0.1 M) in anhydrous acetonitrile and install on unused phosphoramidite ports on the synthesizer (i.e., ports 5 to 8). See Commentary for guidelines in selecting the concentration to use.
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2. Install sufficient amounts of the four regular 3 -O-phosphoramidites S.1a-d (ports 1 to 4), and activator, deprotection, oxidizing, capping, and wash solutions for the number and length of oligonucleotides being made. These reagents are exactly the same as required for conventional synthesis.
3. Create an electronic file of the oligonucleotide sequences to be made according to the particular synthesizer’s specifications. For the MerMade 192 synthesizer, create a simple text file with each line containing a sequence name, a comma or tab delimiter, and the desired sequence in the 5 -to-3 orientation. Modify the 3 -terminal nucleoside in each sequence to reflect the phosphoramidite port used for the corresponding linker phosphoramidite. For example, if the adenosine linker phosphoramidite (S.4a or S.16a) is installed on port 5, modify all sequences ending in “A” to a text string ending with “5”: AAGGCCTT5 instead of AAGGCCTTA. The substitution of the last base by the linker phosphoramidite port position is best done by creating a software routine. This can be done through a short custom program or by using a combination of the MID, IF, and CONCATENATE (&) functions in an Excel spreadsheet as shown by the following formula. =MID(cell ref, 1, LEN(cell ref)-1) & IF(MID(cell ref, LEN(cell ref), 1)="A", "5", IF(MID(cell ref, LEN(cell ref), 1)="C", "6", IF(MID(cell ref, LEN(cell ref), 1)="G", "7", IF(MID(cell ref, LEN(cell ref), 1)="T", "8", "- Error")))) This formula uses the parameter cell ref to denote the Excel spreadsheet cell location of the oligonucleotide sequence. The expression "MID(cell ref, 1, LEN(cell ref)-1)" extracts all of the sequence except for the last character, and the "&" operator concatenates a new character representing the ports for the corresponding linker phosphoramidites. This is determined by using "MID(cell ref, LEN(cell ref), 1)" to extract the last character of the sequence and a series of embedded "IF" functions to see if the last character is A, C, G, or T (case insensitive). When a match is found, the expression substitutes a value of 5, 6, 7, or 8, respectively (these values may be changed as required) as the last character in the sequence. If the sequence does not end in one of the four standard bases, the suffix “-Error” will be attached.
4. Using one of the “dry” loading methods described below (see Support Protocol), load LCAA-CPG into an OF1100 filter plate at ∼2 to 10 mg/well (depending upon the synthesis scale). Gently tap or briefly centrifuge the plate to bring down any CPG particles sticking to the sides of the wells. Install the plate in the synthesizer. If recommended by the instrument’s manufacturer, seal any unused synthesis wells. 5. Select the appropriate software option for universal supports (first coupling cycle begins with the 3 -terminal base) instead of conventional or prederivatized supports (first coupling cycle begins with the penultimate base from the 3 -end). This option is available on the MerMade 192. On some older instruments, such as the ABI 394, an additional “fake” base must be added to the 3 -end of each oligonucleotide, since the instrument always assumes the last nucleoside is already attached to the support. Such instruments were either developed before universal supports were available or their manufacturer did not include support for universal supports.
6. Prime the reagents and start the synthesizer, as per the manufacturer’s instructions.
Cleave and deprotect oligonucleotides 7. Remove the OF1100 filter plate from the synthesizer and transfer to a fume hood. Nucleoside Phosphoramidites Containing Cleavable Linkers
8. Place a 96 × 1-mL multi-well collection plate under the OF1100 filter plate. Add 120 µL of 1:1 (v/v) ammonium hydroxide/40% (w/v) aqueous methylamine solution to each well of the filter plate. Wait ∼5 min.
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9. Add another 120-µL portion of ammonium hydroxide/methylamine solution to each well. Wait another 2 min. Deprotection with methylamine can only be performed if all of the cytosine bases are protected with N-acetyl protecting groups. If cytosine bases with N-benzoyl protecting groups are present, ammonium hydroxide without methylamine must be used (see APPENDIX 3C, deprotection of DNA oligonucleotides).
10. Transfer any remaining solution through the filter plate into the collection plate by briefly centrifuging (∼2 min at 12.6 × g or less) or applying vacuum (∼10 sec). When using a vacuum manifold, be careful not to remove too much of the volatile ammonia and methylamine, as complete deprotection may not occur.
11. Seal the plate tightly using the clamping apparatus recommended by the synthesizer manufacturer. Place in a preheated oven (65◦ C) and leave for 15 to 30 min. 12. Place the unopened plate assembly on ice until cooled to below room temperature (∼5 min). CAUTION: Do not attempt to open the collection plate while it is still warm. The wells are under pressure, and eye injury, cross-contamination, and loss of samples may result if the cover is removed without adequate cooling.
13. Place the collection plate in a centrifugal evaporator such as a Speedvac and evaporate off the methylamine and ammonia (∼1 hr). The residual aqueous solution containing the crude deprotected oligonucleotide product is ready for desalting or further purification (see Chapter 10).
DRY LOADING OF CPG SUPPORT INTO 96-WELL PLATES Adding the correct amount of each support to each synthesis well in a 96-well plate can be a tedious and error-prone task. This task is greatly simplified when a single universal support is used in every well. Originally, a suspension of CPG in a mixture of organic solvent blended to approximately the same density as the CPG beads (“wet” method) was used to transfer the CPG into each well (Lashkari et al., 1995; Cheng et al., 2002). However, this method is inconvenient and difficult to perform by hand, because the beads rapidly settle out of suspension and the solvent fumes are annoying. The wet method
Figure 3.12.4 Pipetter device for dry loading CPG into 96-well plates. An electric Pipet-Aid filler/dispenser is fitted with a 100-µL filter-type micropipet tip whose length is cut to deliver the required volume of CPG. A 1000-µL pipet tip is used to connect the 100-µL tip to the pipetter.
SUPPORT PROTOCOL
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Figure 3.12.5 Sliding dispenser for dry loading CPG into 96-well plates. This dispenser can be constructed from Plexiglas in any machine shop. The dimensions of the body are 45 mm × 55 mm × 18 mm (width by length by thickness) and the movable slide is 19 mm × 110 mm (width by length), with the portion between the stops machined to the exact thickness of the groove in which it slides. The distance between the fill and dispense holes is 20 mm. The amount (volume) of CPG dispensed depends upon the thickness of the slider and the diameter of the hole drilled into it. The bottom face of the dispenser contains nine wells drilled (∼1.5 mm deep) to align with the spacing of the synthesis plate being used. The authors thank Dr. John Hobbs, University of British Columbia, for suggesting the design of the sliding dispenser. Current Protocols in Nucleic Acid Chemistry
can be automated, but this requires a very expensive robotic or 96-tip multiple pipeting station (Rayner et al., 1998), which is difficult to justify when synthesizing only one or two plates per day. Instead, the CPG can be handled as a dry powder (“dry” method) using one of the two devices described below. In Figure 3.12.4, a simple and commonly available 100-µL pipet tip containing a filter (filter tip) is trimmed to hold a specific volume of CPG. The tip is then attached to an electric pipetting aid capable of applying either vacuum or mild air pressure. The pipet is inserted into a vial of CPG, the vacuum is applied to suck the beads into the tip, the tip is immediately placed in the synthesis well, and the beads are dispensed by switching from vacuum to air pressure. The amount of beads dispensed is determined by the length of the trimmed pipet tip. A tip of ∼0.5 cm will hold ∼2 to 3 mg support, which is suitable for ∼50 to 100 nmol-scale syntheses. Tips can be easily calibrated by dispensing the CPG onto an analytical balance. In Figure 3.12.5, a sliding dispenser for CPG is shown, which can be assembled from Plexiglas in most machine shops. This device has two advantages over the filter tip method. First, operation of the dispenser is faster because the dispenser has a built-in CPG reservoir and because the bottom is indexed for quick alignment with each well. Second, the dispenser can be built to dispense greater amounts of CPG than the filter tips. The dimensions of the loader and the slider are generally not critical, although the movable center “slider” should be machined to an exact thickness and flatness to prevent CPG beads from being jammed between the pieces. The volume of the hole drilled into the slider determines the amount of CPG to be dispensed. Either the thickness of the slider or the diameter of the hole can be adjusted. A slider that is 0.125 in. (∼3 mm) thick with a 5/32-in.-diameter (∼4 mm) hole will dispense ∼10 mg of CPG. To use the sliding dispenser, a polypropylene micro funnel (Aldrich) is inserted into the top port of the dispenser and filled with CPG. When the center slider is in the “fill” position, CPG will fall from the funnel into the slider and fill the cavity. The dispenser is then fitted over the well and the slider is pushed to the “dispense” position, which allows the CPG to fall into the well. The dispenser is moved to the next well and the slide is moved back and forth between the fill and dispense positions once more. By repeating this process, a 96-well plate can be filled with support quite quickly.
TANDEM SYNTHESIS OF MULTIPLE OLIGONUCLEOTIDES IN A SINGLE SYNTHESIS
BASIC PROTOCOL 3
This protocol describes how to attach an additional oligonucleotide sequence onto the 5 -end of another oligonucleotide by use of the S.4 linker phosphoramidites. This allows strings of oligonucleotides, linked end-to-end in tandem, to be synthesized in a single automated synthesis (Pon et al., 2002; Pon and Yu, 2005). The linkages between the oligonucleotides are simultaneously cleaved during cleavage from the support, so that the product is released as a mixture of oligonucleotides. This method eliminates the need to separately synthesize and deprotect each individual oligonucleotide before combining them into a desired mixture. It is useful for making pairs of PCR primers, sets of multiplexed primers, and double-stranded sequences, and for increasing the yield of a single sequence by making multiple copies simultaneously. It can also be combined with Basic Protocol 2 to perform tandem syntheses in a multi-well plate format. Tandem oligonucleotide synthesis should only be performed using the S.4 linker phosphoramidites, which contain the sulfonyldiethanol group. During cleavage, these residues undergo β-elimination to leave a monophosphate group on the support (S.20; Fig. 3.12.6B) and on the 5 -terminus of oligonucleotides synthesized in tandem
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3.12.13
Figure 3.12.6 Cleavage mechanisms for linker phosphoramidites. (A) The linkage S.17 formed by coupling S.16 to LCAA-CPG is hydrolyzed at the succinyl ester groups to yield oligonucleotide S.7 with a 3 -OH end and the support S.18 with a phosphoroamidate monoester. (B) The linkage S.19 formed by coupling S.4 to LCAA-CPG undergoes rapid β-elimination from the sulfonyldiethanol group to release a 3 -succinylated oligonucleotide from the support. During base deprotection, simultaneous hydrolysis at the 3 -ester position generates oligonucleotide S.7 with a 3 -OH end. A phosphoramidate group is left behind on the support. (C) The linkage S.21 connects two oligonucleotides synthesized in tandem using S.4. It undergoes β-elimination and ester hydrolysis to leave a 5 -monophosphate on oligonucleotide 1 (S.23) and a 3 -OH on oligonucleotide 2 (S.22). Nucleoside Phosphoramidites Containing Cleavable Linkers
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(S.23; Fig. 3.12.6C). In contrast, the alternative diol spacers in the S.16 phosphoramidites leave behind an unnatural 5 -alkyl phosphate group (S.18; Fig. 3.12.6A). These residues are harmless for attaching the first oligonucleotide to a support, because the alkyl phosphate group is discarded with the used support. In a tandem synthesis, however, the alkyl phosphate group will remain on the oligonucleotides.
Materials Linker phosphoramidites (S.4a-d; see Basic Protocol 1) Anhydrous acetonitrile Solid support: For column format: prepacked synthesis columns containing prederivatized solid-phase support For plate format: long-chain alkylamine controlled-pore glass (LCAA-CPG; ◦ S.5), ∼100 µmol/g amino group loading, 500 A pore size NAP-10 (prepacked Sephadex) column Additional reagents and equipment for oligonucleotide synthesis (APPENDIX 3C and Basic Protocol 2) 1. Prepare a 0.1 M solution of the required linker phosphoramidites S.4a-d in anhydrous acetonitrile. Install these on unused phosphoramidite ports on the synthesizer (i.e., ports 5 to 8). 2. Create an oligonucleotide sequence file by combining all of the sequences into a single text string. a. For underivatized LCAA-CPG supports: Modify the terminal 3 -position of each oligonucleotide sequence so that it corresponds to the desired linker phosphoramidite (see Basic Protocol 2, steps 3 to 5). b. For prepacked synthesis columns containing prederivatized supports: Substitute only the 3 -position of each additional oligonucleotide by a linker phosphoramidite reagent. For example, to synthesize the two 12-mer sequences 5 -AAA AAA AAA AAA-3 and 5 -TTT TTT TTT TTT-3 using a prepacked T-derivatized solid support, with linker phosphoramidite S.4a installed on phosphoramidite port 5, program the synthesizer to make AAA AAA AAA AA5 TTT TTT TTT TTT in a single automated synthesis. To perform the same synthesis on underivatized LCAA-CPG, with S.4d on port 6, program the synthesizer to make AAA AAA AAA AA5 TTT TTT TTT TT6.
3. Perform the automated synthesis, cleavage from the support, and deprotection steps in the usual fashion. Either ammonium hydroxide or ammonium hydroxide/methylamine deprotection protocols can be used. No other modifications to procedures are required.
4. Evaporate off the ammonium hydroxide or methylamine reagent, redissolve in 1 mL water, and desalt on a NAP-10 (prepacked Sephadex) column. The desalted product will be a mixture of each oligonucleotide sequence as well as all of the failure sequences normally produced (N–1, N–2, and so on). If the synthesis was successful and the high coupling yields typically expected (98% to >99%) were obtained, the product mixture is satisfactory for use in most applications. Purification of each individual component from this mixture is not recommended unless the tandem synthesis was performed to make multiple copies of a single sequence.
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Triethylammonium phosphate (TEAP) solution, 2 M In a chemical fume hood, set up a large (2- to 4-liter) beaker in an ice bath on top of a magnetic stirrer. Add 892 ml water and begin stirring. Carefully add 272 ml of 85% (w/v) aqueous phosphoric acid under continued stirring. Carefully add 836 ml triethylamine with continued stirring. The majority of the triethylamine will float on top of the aqueous solution. Stir the mixture in the ice bath until it forms a homogeneous solution (∼2 hr). Allow to warm to room temperature and then transfer to a sealed bottle for storage (stable for a year or more). Before use, dilute the 2 M stock solution to ∼0.5 M with distilled water. CAUTION: Eye and skin protection are required when handling 85% phosphoric acid, as it is a strong acid.
COMMENTARY Background Information The need for large numbers of oligonucleotides in a wide variety of applications has encouraged various means to reduce the cost of their production. In particular, development of highly parallel synthesis instruments using 96-, 384-, and 1536-well synthesis plates (Lashkari et al., 1995; Sindelar and Jaklevic, 1995; Brennan et al., 1998; Rayner et al., 1998; Cheng et al., 2002) has reduced the cost of labor, and reductions in synthesis scale have reduced reagent costs. Nonetheless, there is a continuing need to reduce costs by decreasing both the time and materials involved. This unit describes methods for doing so via two approaches.
Nucleoside Phosphoramidites Containing Cleavable Linkers
Universal supports Originally, solid-phase oligonucleotide synthesis began with the first nucleoside already attached to the solid-phase support, which is prepackaged into ready-to-use synthesis columns. Since there are only four common deoxyribonucleosides, it was not difficult to produce and maintain inventories of separate columns for each base. However, when multi-well plates are used, the process of placing the correct support into the correct position on each plate becomes quite tedious and errorprone. This difficulty spurred the search for a single universal support that could be used for any oligonucleotide sequence, so that all wells could be preloaded with a single material. The problem with this approach, however, is that the phosphoramidite reagents (S.1) used to synthesize the oligonucleotides would begin each oligonucleotide with a 3 -terminal phosphate group. This phosphate group inter-
feres with biological activity, since the natural configuration produced by enzymatic activity has 5 -phosphate and 3 -OH termini, not 5 -OH and 3 -phosphates. Fortunately, the linkage to a 3 -terminal monophosphate group can be cleaved if a cyclic phosphodiester configuration can be formed, as exemplified by RNA hydrolysis. A large number of different universal supports employing some type of diol-based configuration has been developed. Unfortunately, the dephosphorylation step required after synthesis on the first generation of these supports was slow, and prolonged conditions (longer times and higher temperatures) or additions of various metal salts were required (Debear et al., 1987; Scott et al., 1994; Schwartz et al., 1995; Nelson et al., 1997; Scheuerlarsen et al., 1997; Azhayev, 1999; Lyttle et al., 1999). Otherwise, the desired 3 -OH product was contaminated with the unwanted 3 -phosphorylated products. Recently, a second generation of improved universal supports has been developed (Azhayev and Antopolsky, 2001; Guzaev and Manoharan, 2003). These are superior to the first generation supports because the universal linker is designed to retain phosphorylated oligonucleotides and only release products with the desired 3 -OH ends. However, these supports still require some modification to the conventional cleavage and deprotection conditions, i.e., the use of dilute methanolic ammonia instead of concentrated aqueous ammonium hydroxide, and any material not released from the support reduces the recovered yield. To approach this problem from a different perspective, modified linker phosphoramidites
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Figure 3.12.7 Chemical 5 -phosphorylation using a commercially available 2,2 -sulfonyldiethanol-containing phosphoramidite S.25.
have been designed with an easily cleavable linker between the protected nucleoside and the phosphoramidite group. The first compounds used a succinyl-ethylene glycol combined linker (S.16; Fig. 3.12.3; Pon and Yu, 2001), but an improved reagent based on a succinyl-sulfonyldiethanol combination was eventually developed (S.4; Fig. 3.12.2; Pon and Yu, 2004). In both cases, the ester linkage on the 3 -position is identical to the succinyl ester attachment widely used to link nucleosides to prederivatized solid-phase supports. Thus, cleavage from the support can be done by standard methods, producing only S.7 with the desired 3 -OH groups (Fig. 3.12.6A and B). The phosphoramidate residues left behind (S.18 and S.20) remain attached to the support and are discarded with the used support. A major advantage to this approach is that underivatized amino supports, such as the widely used long-chain alkylamine controlledpore glass (LCAA-CPG) becomes a universal support (Fig. 3.12.1B; Pon et al., 1988). This material is much less expensive than other universal supports because a special linker arm does not have to be synthesized and attached to the support. Although, the phosphoramidate bond formed by attachment
of the linker phosphoramidite to the support in S.17 and S.19 is slightly different from the usual phosphate linkage, the P-N phosphoramidate linkage is completely stable to the conditions of oligonucleotide synthesis. Similar internucleotide phosphoramidate P-N bonds have been reported in the synthesis of various antisense oligonucleotides (Mag and Engels, 1989; Gryaznov and Letsinger, 1992). Substitution of the ethylene glycol group in S.16 with a 2,2 -sulfonyldiethanol group provides an improved linker phosphoramidite (S.4) because a second cleavage site is introduced. The sulfonyldiethanol group undergoes a β-elimination, very similar to the βelimination mechanism used to remove the 2cyanoethyl protecting groups from phosphate linkages. This facile deprotection to yield a free phosphoric acid group has been widely utilized to produce 5 -monophosphorylated oligonucleotides (Horn and Urdea, 1986) through use of the commercially available reagent S.25 (Fig. 3.12.7; Applied Biosystems, Glen Research, Chemgenes, or any supplier of phosphoramidites). Phosphoramidite S.25 has been used occasionally to initiate oligonucleotide synthesis on solid-phase supports (Gryaznov and Letsinger, 1993) and
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on glass (Glazer et al., 2001) or polypropylene (Shchepinov et al., 1997) microarrays when synthesis of 3 -terminally phosphorylated products is acceptable. The identical βelimination mechanism results in the release of the oligonucleotide from the support when either S.25 or S.4 are used. In linker phosphoramidite S.4, βelimination and hydrolysis under basic conditions provides two advantages. First, the rapid β-elimination releases the product from the support after only a short treatment (5 min) with aqueous ammonium hydroxide (Fig. 3.12.6B and C). This is much faster than the 1 to 2 hr required to cleave a succinic acid linkage (Pon and Yu, 1997). The faster release time is especially convenient when deprotection is performed manually (as is the case for most 96-well plate syntheses). The slower hydrolysis of the 3 -O-succinyl ester occurs off the support at the same time as base deprotection. Second, β-elimination of S.4 generates a phosphate group on whatever S.4 has been attached to (Fig. 3.12.6C). This formation of both 3 -OH and 5 -monophosphate groups at the junction where S.4 is used is critical when more than one oligonucleotide sequence or copy is produced through tandem synthesis.
Nucleoside Phosphoramidites Containing Cleavable Linkers
Tandem oligonucleotide synthesis Tandem oligonucleotide synthesis was originally reported in 1994 when a special phosphoramidite for preparing “two oligonucleotides per synthesis” (TOPS) was described (Hardy et al., 1994). This linking phosphoramidite contained a pair of 1,4anhydroerythritol rings that resembled the diol linker arm on a universal support. However, instead of being the starting point for a single oligonucleotide, it was used to synthesize a second oligonucleotide on the 5 -end of a preceding sequence. Unfortunately, just like the first generation universal supports, this reagent required harsher deprotection conditions to remove the 3 -phosphate group from the second oligonucleotide. Two improved schemes for tandem oligonucleotide synthesis have eliminated the difficult dephosphorylation step. In the first approach, the introduction of a 3 phosphate group was avoided by not using phosphoramidite coupling chemistry to begin the second or subsequent oligonucleotides. Instead, reagents such as HATU, HBTU, or HCTU are used in combination with 4-dimethylaminopyridine to couple the nucleoside-3 -O-hemiesters of either succinic
or hydroquinone-O,O -diacetic acid onto the 5 -end of a support-bound oligonucleotide (Pon et al., 2002). This coupling procedure was found to be very rapid and suitable for automation in the development of a method for reusing solid-phase supports (Pon et al., 2001). However, this method was difficult to implement because it required an automated synthesizer that had five additional reagent positions (for four nucleoside hemiesters and an additional coupling reagent). Because not many instruments at that time had such capacity, the tandem synthesis method using linker phosphoramidite reagents was developed. This method is described in this unit. The linker phosphoramidite S.4 has been quite suitable for tandem oligonucleotide synthesis. It has been used to produce a variety of oligonucleotides for situations in which it is advantageous to synthesize an oligonucleotide mixture in a single synthesis, rather than performing multiple syntheses (and their associated deprotection and workup steps) before combining the products for use. Figure 3.12.8 illustrates how a set of linker phosphoramidites can be used to assemble a string of oligonucleotides, linked end-to-end in tandem, on the surface of a support. Examples of these applications include pairs of PCR primers, complementary sequences that spontaneously base-pair into double-stranded sequences upon deprotection, and single oligonucleotides when synthesis of multiple tandem copies is used to increase the amount of sequence produced by any single synthesis column. This latter application is potentially useful for the large-scale synthesis of pharmaceutical oligonucleotides. What is unique about this approach is that the phosphate group arising from the linker phosphoramidite is transferred onto the 5 position of the preceding oligonucleotide. This is different from the 3 -phosphate that occurs if a conventional phosphoramidite (S.1) is used, in that no difficult 3 -dephosphorylation steps are needed. Although, 5 -phosphorylated oligonucleotides are not as widely used as 5 OH oligonucleotides, they more closely resemble the biological products created by enzymes. Thus, the 5 -phosphate groups do not interfere with the oligonucleotide’s use in the same way 3 -phosphate groups do.
Critical Parameters As with any oligonucleotide synthesis or organic synthesis involving moisture-sensitive reactions, good results depend upon the successful exclusion of moisture. Methods
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Figure 3.12.8 Synthetic scheme showing the tandem synthesis of multiple oligonucleotides in a single automated synthesis. Linker phosphoramidite reagent S.4 is used to begin each new oligonucleotide sequence on the 5 -OH end of a preceding sequence. After N rounds of linker phosphoramidite addition and oligonucleotide synthesis, a string of oligonucleotides linked end-to-end on the surface of the support is obtained (S.31). Treatment with ammonium hydroxide simultaneously cleaves the linker joining the string to the support and each linker joining two oligonucleotides together. The product released contains a mixture of the final oligonucleotide (S.32) with both 5 -OH and 3 -OH ends, and all preceding oligonucleotides (S.33 and S.34) with 5 -monophosphate and 3 -OH ends. If desired, product S.32 can also be obtained as a 5 -monophosphate by using the phosphorylating reagent S.25 in the last coupling cycle.
for maintaining these conditions and evaluating oligonucleotide quality are described in APPENDIX 3C. This section will concentrate on other parameters that are critical to oligonucleotide syntheses as outlined in Basic Protocols 2 and 3. When using linker phosphoramidites S.4 or S.16 to make oligonucleotides on LCAACPG, the two main parameters to consider are the age of the solutions and the amount of reagent to add. Due to the labile sulfonyldiethanol group, solutions of linker phosphoramidites should be used as quickly as possible. When the linker phosphoramidite decomposes, there is less phosphoramidite present to couple to the support, and a reduced nucleoside loading occurs. Since this is used at the start of synthesis, a coupling de-
crease in the first coupling cycle will reduce the amount of any subsequent oligonucleotide. Solutions of S.4a-b and S.4d lead to an ∼10% to 20% decrease in the amount oligonucleotide recovered when they have been in solution at room temperature for two weeks. Thus, it is recommended that these solutions be used within 1 to 2 weeks of preparation. The linker phosphoramidite reagents with the least stability are the guanine derivatives S.4c and S.16c. These solutions should be used within 3 to 5 days of preparation. The synthesis scale (amount of oligonucleotide produced) depends upon a number of factors. These are related to both the solidphase support used and the linker phosphoramidite. For the support, the key factor is the number of amino groups available to react with
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Nucleoside Phosphoramidites Containing Cleavable Linkers
the linker phosphoramidite. This is affected by the amino group loading, the pore size (which affects accessibility to the bulky phosphoramidite reagent), and the mass of support used. For the linker phosphoramidite, it is the concentration and amount of reagent added. The effect of these parameters is discussed in greater detail in Pon and Yu (2004). In summary, CPG supports with amino group loadings of ∼100 µmol/g are recommended, with ◦ an average pore size of not less than 500 A. Only about one-half of the amino groups specified by the manufacturer are actually accessible for oligonucleotide synthesis, but this is the case for both prederivatization protocols and this universal protocol. Presumably, this is because there are pores that are too small to be accessible to phosphoramidite reagents, but are still titratable in the manufacturer’s loading determination. The pore size of CPG particles also affects their density,◦ and CPG particles with an average 1000 A pore size are light enough to be knocked out of the wells in 96-well synthesis plates. For synthesis on the ◦ MerMade 192, the heavier 500 A CPG, which remains in the wells, is therefore preferred. However, because of the smaller pore volume, the maximum oligonucleotide length is limited to ∼50 bases. The mass of CPG beads in each synthesis is another obvious factor controlling the scale, and increasing the mass of beads is the easiest way to increase synthesis scale. The amount of CPG used in each well of a 96-well synthesis plate can be increased from ∼1 or 2 mg to ∼10 mg without producing any affect on oligonucleotide quality or increasing the consumption of reagents. This is because the minimum volume usually dispensed by these synthesizers (∼20 to 25 µL) is still sufficient to completely cover the increased mass of support. Generally, ∼2 mg of◦CPG (∼100 µmol/g of NH2 groups with 500 A pores) can be used for synthesis on an ∼40-nmol scale, and 10 mg of CPG can be used for a 200-nmol scale. Because linker phosphoramidites S.4 and S.16 behave very similarly to conventional phosphoramidites, the coupling time and activation do not need to be modified. Typical conditions are 0.25 M 5-ethylthiotetrazole in acetonitrile as the activating reagent (mixed 1:1 v/v with the phosphoramidites at the time of coupling) and a coupling time of 60 sec. However, unlike conventional phosphoramidites, which require the highest possible coupling yields, addition of linker phosphoramidites does not have to be quantitative, i.e., coupling to 100% of the amino groups on the CPG is not required. Instead, the amount of cou-
pling can be adjusted to provide the synthesis scale desired. Any unreacted amino groups are acylated in the next capping step. It is recommended that the volume of linker phosphoramidite be reduced to either the minimum amount required to just cover the CPG or the minimum amount that can be reliably dispensed, since adding additional volume has little benefit. The concentration of the linker phosphoramidite is a much more important parameter. A concentration of 0.1 M (the concentration at which S.1 is typically used) is probably the maximum that needs be considered. Concentrations of 0.15 and 0.2 M only produce slightly higher synthesis scales, and then only when supports with very high loadings (>100 µmol/g) are employed. Lower concentrations, such as 0.05 or 0.025 M, can be considered, especially when only small-scale syntheses are required. Thus, linker phosphoramidite consumption can be significantly reduced. Compared to synthesis with 0.1 M linker phosphoramidite, synthesis with 0.025 and 0.05 M solutions will yield approximately one-third to two-thirds the amount of crude product. The oligonucleotide quality is unaffected by the reduction in yield of the first coupling step, and the decreased amount of material is still more than adequate for most applications. The synthesis of multiple oligonucleotides by tandem synthesis is very similar to the synthesis of a single oligonucleotide, but there are additional factors that must be considered. These relate to the maximum length of the string of tandem-linked oligonucleotides, the difference between the 5 -OH end of the last sequence and the 5 -phosphorylated end(s) of the preceding sequence(s), and the yield reduction that occurs with each subsequent tandem oligonucleotide. First, the number of tandem oligonucleotides that can be prepared in a single synthesis is limited by the same factors limiting the length of any single oligonucleotide. Thus, the average coupling efficiency (Zhou et al., 2004), detritylation efficiency (Habus and Agrawal, 1994; Paul and Royappa, 1996), depurination during detritylation (Septak, 1996), and limitations of the support’s pore volume (Efcavitch et al., 1986) limit the total number of couplings that can be performed. With good reagents, properly functioning instrumentation, and long-mer oligonucleotide synthesis protocols, it is possible to routinely synthesize single oligonucleotides of up to 120 to 150 bases (for further guidelines on synthesis of long
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oligonucleotides, see discussion in APPENDIX 3C). This length is more than sufficient to accommodate the tandem synthesis of a pair of typical PCR primers. In Pon and Yu (2005), the tandem synthesis of a string of multiple oligonucleotides totaling up to 120 bases showed that oligonucleotide quality was very similar to that obtained by conventional syn◦ thesis. In that case, a wide-pore 1000 A CPG support with a reduced loading (10 µmol/g) was employed, and a milder 2% (v/v) solution of 1, 2-dichloroacetic acid was substituted for the 5% (v/v) solution regularly used. Second, as described in Background Information and illustrated in Figure 3.12.8, tandem oligonucleotide synthesis introduces a phosphate group onto the 5 -end of all but the final oligonucleotide. While this is not expected to have any adverse effects, it does limit the introduction of 5 -modifications such as fluorescent dyes or other labels. In this case, the 5 -modification can only be introduced onto the last oligonucleotide made, which has a 5 OH end. Finally, because automated oligonucleotide synthesis proceeds in a step-wise manner, the overall yield is highly dependent upon the coupling efficiency of every phosphoramidite addition. For the synthesis of single oligonucleotides, this dependence can be expressed by the equation:
where OY is overall yield, AY is average coupling yield, and N is oligonucleotide length. When tandem oligonucleotides are produced, the amount of each subsequent oligonucleotide can only be as much as the number of available 5 -ends of the preceding sequence (as defined by the above equation). Consequently, the amount of each oligonucleotide made in tandem will always be less than the amount of the preceding oligonucleotide. Fortunately, for most biological applications such as PCR primers, it is not critical to have both primers in an exact 1:1 proportion. Also, most applications are relatively insensitive to the small amounts of shorter failure sequences present in crude oligonucleotide preparations. Therefore, the crude products of tandem syntheses can be used without extensive purification or adjustment of the component proportions. Simple desalting of the crude tandem products on a Sephadex column is usually sufficient to remove low-molecular-weight impurities.
For large-scale oligonucleotide syntheses, tandem synthesis can be used to make multiple copies of the same sequence. This increases the amount of oligonucleotide that can be obtained from any individual synthesis column. Since the support is the single most expensive material in large-scale synthesis, strategies that either reuse the support or produce more material from a single synthesis are quite desirable. In the case of tandem synthesis, the amount of material produced is expressed by:
or
where M is the number of tandem oligonucleotides in the string being synthesized. Since the amount of material obtained decreases with each additional copy made in tandem, while the reagent consumption for each coupling step remains fixed, there is a diminishing economic return with each additional tandem synthesis. However, given the high cost of support and the high coupling efficiencies routinely obtained by commercial large-scale production facilities, making at least one extra copy in tandem is still beneficial. One possible difficulty with performing tandem oligonucleotide synthesis is the purification and analysis of the product mixture. Purification and isolation of individual products is not recommended when different sequences are made by tandem synthesis, especially when the products are of similar size, because similarly sized sequences may be impossible to resolve. However, purification and analysis of multiple copies of the same sequence is very similar to conventional analysis, especially if the last oligonucleotide in the tandem string is phosphorylated by reagent S.25 to match the other copies.
Troubleshooting Basic Protocol 1 and the Alternate Protocol both require some experience in organic synthesis. In particular, the reactions are sensitive to moisture contamination, and precaution must be taken to keep the anhydrous solvents and the sensitive reagents (i.e., HBTU, S.13, and S.14) dry. Each synthetic step should be monitored by TLC, including a sample of the starting material with the reaction mixture so
Synthesis of Unmodified Oligonucleotides
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Nucleoside Phosphoramidites Containing Cleavable Linkers
that their mobilities can be compared. Each step results in a significant mobility change, so identification of the reaction progress is straightforward. Basic Protocol 2 should also be straightforward to perform as long as the precautions for programming the synthesizer (as described in the protocol) and selecting the appropriate amounts of CPG and linker phosphoramidite (see Critical Parameters) are observed. It is essential that the correct parameter for the support type (universal support, not conventional support) be selected, and that the synthesis cycle not begin with a capping step. If an error is made in the first case, a linker phosphoramidite will not be used to start the synthesis and the resulting oligonucleotide will be one base short and irreversibly attached to the support through a phosphoramidate linkage. An error in the second case will cause the amino groups on the support surface to be acetylated before they can react with the linker phosphoramidite, so that no oligonucleotide synthesis can occur. If spare phosphoramidite ports are being used for the first time, one must also ensure that the injectors are properly calibrated and the synthesis scripts, cycles, and reagent files controlling the synthesizer are appropriately modified. Finally, all products should be quantitated by UV absorbance at 260 nm to ensure that the syntheses are proceeding satisfactorily and at the desired scale. Basic Protocol 3 is sensitive to the efficiency of the linker phosphoramidite coupling. If the amounts of additional oligonucleotides made in tandem seem lower than expected, the coupling efficiency of the linker phosphoramidite needs to be verified. The easiest way to do this is through collection of the trityl colors produced from coupling the 5 terminal base and the subsequent linker phosphoramidite (APPENDIX 3C for the trityl assay). The coupling yield of this step should be at least 75%, with more typical yields of 90% to 98%. Low yields may be caused by moisture contamination of the reagents, linker phosphoramidite decomposition (see Critical Parameters), or inadequate reagent delivery by the synthesizer. It is also difficult, if not impossible, to resolve oligonucleotides of the same length but different sequence. A test synthesis involving two products that differ in length by at least two bases, or where one product is labeled with a fluorescent dye, is recommended to make the products easily identifiable by HPLC or electrophoresis. This, along with trityl analysis, will confirm the success of the tandem synthesis.
Anticipated Results The yields recovered from Basic Protocol 1 are indicated in the protocol details. Experienced chemists may obtain greater yields, but generally a yield of 70% to 80% for each step is satisfactory. In Basic Protocol 2, the quality of the oligonucleotides should be identical to the quality obtained from conventional prederivatized supports. However, the synthesis scale can be varied considerably by alteration of the conditions chosen for the support and linker phosphoramidite. As described earlier, one can easily adjust the scale between 10 and 50 nmol. Synthesis at scales up to 200 nmol can be obtained by increasing the amount of CPG or by using CPG with a higher amino group loading. In Basic Protocol 3, the products obtained should be comparable to products made individually, when used in most applications. Thus, pairs of PCR primers made in tandem should give similar, if not identical, amplification to those made separately. Doublestranded fragments will spontaneously basepair with each other after deprotection, and the double-stranded nature of these products can be observed by comparing their mobility under denaturing versus non-denaturing conditions. Since at least one of the tandem products will be 5 -phosphorylated, the double-stranded fragment is also ready for enzymatic ligation.
Time Considerations The time required to prepare linker phosphoramidites will depend upon the experience of the researcher with chemical synthesis. However, because multiple steps are involved and the procedures must be repeated for each of the four common deoxynucleosides, preparation of the linker phosphoramidites is a major and labor-intensive task. This could easily take several months for a single person to complete. For this reason, the preparation of linker phosphoramidites from modified nucleosides is only practical if they will be used very frequently. Once the linker phosphoramidites are prepared, their use is quite straightforward and requires little extra preparation. Tandem oligonucleotide synthesis will require a much longer run time on an automated synthesizer because twice as many (if not more) automated couplings are required to make the multiple oligonucleotides. Since this can be scheduled to run overnight, the extra time on the synthesizer is not a significant drawback. However, when making large collections of multiple oligonucleotides, the time
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and effort saved by not having to individually set up, prepare, deprotect, purify, and package all of the required oligonucleotides will be substantial.
Literature Cited Azhayev, A.V. 1999. A new universal support for oligonucleotide synthesis. Tetrahedron 55:787800. Azhayev, A.V. and Antopolsky, M.L. 2001. Amide group assisted 3 -dephosphorylation of oligonucleotides synthesized on universal A-supports. Tetrahedron 57:4977-4986. Brennan, T., Biddison, G., Frauendorf, A., Schwarcz, L., Keen, B., Ecker, D.J., Davis, P.W., Tinder, R., and Swayze, E.E. 1998. Twodimensional parallel array technology as a new approach to automated combinatorial solidphase organic synthesis. Biotechnol. Bioeng. 61:33-45. Cheng, J.Y., Chen, H.H., Kao, Y.S., Kao, W.C., and Peck, K. 2002. High throughput parallel synthesis of oligonucleotides with 1536 channel synthesizer. Nucl. Acids Res. 30:e93. Debear, J.S., Hayes, J.A., Koleck, M.P., and Gough, G.R. 1987. A universal glass support for oligonucleotide synthesis. Nucleosides Nucleotides Nucleic Acids 6:821-830. Efcavitch, J.W., McBride, L.J., and Eadie, J.S. 1986. Effect of pore diameter on the support-bound synthesis of long oligodeoxynucleotides. In Biophosphates and Their Analogues—Synthesis, Structure, Metabolism and Activity (K.S. Bruzik and W.J. Stec, eds.) p. 6570. Elsevier Science Publishers, Amsterdam. Gait, M.J., Singh, M., Heathcliffe, G.R., Atkinson, T.C., Newton, C.R., and Markham, M.F. 1980. Rapid synthesis of oligodeoxyribonucleotides. IV. Improved solid phase synthesis of oligodeoxyribonucleotides through phosphotriester intermediates. Nucl. Acids Res. 8:10811096. Glazer, M., Fidanza, J., McGall, G., and Frank, C. 2001. Colloidal silica films for high-capacity DNA probe arrays. Chem. Mater. 13:4773-4782. Gryaznov, S.M. and Letsinger, R.L. 1992. Synthesis and properties of oligonucleotides containing aminodeoxythymidine units. Nucl. Acids Res. 20:3403-3409. Gryaznov, S.M. and Letsinger, R.L. 1993. Anchor for one step release of 3 -aminooligonucleotides from a solid support. Tetrahedron Lett. 34:12611264. Guzaev, A.P. and Manoharan, M. 2003. A conformationally preorganized universal solid support for efficient oligonucleotide synthesis. J. Am. Chem. Soc. 125:2380-2381. Habus, I. and Agrawal, S. 1994. Improvement in the synthesis of oligonucleotides of extended length by modification of detritylation step. Nucl. Acids Res. 22:4350-4351. Hardy, P.M., Holland, D., Scott, S., Garman, A.J., Newton, C.R., and McLean, M.J. 1994.
Reagents for the preparation of two oligonucleotides per synthesis (TOPSTM ). Nucl. Acids Res. 22:2998-3004. Horn, T. and Urdea, M.S. 1986. A chemical 5 phosphorylation of oligodeoxyribonucleotides that can be monitored by trityl cation release. Tetrahedron Lett. 27:4705-4708. Kumar, P., Ghosh, N.N., Sadana, K.L., Garg, B.S. and Gupta, K.C. 1993. Improved methods for 3 O-succinylation of 2 -deoxyribo- and ribonucleosides and their covalent anchoring on polymer supports for oligonucleotides synthesis. Nucleosides Nucleotides 12:565-584. Lashkari, D.A., Hunicke-Smith, S.P., Norgren, R.M., Davis, R.W., and Brennan, T. 1995. An automated multiplex oligonucleotide synthesizer: Development of high-throughput, lowcost DNA synthesis. Proc. Natl. Acad. Sci. U.S.A. 92:7912-7915. Lyttle, M.H., Dick, D.J., Hudson, D., and Cook, R.M. 1999. A phosphate bound universal linker for DNA synthesis. Nucleosides Nucleotides Nucleic Acids 18:1809-1824. Mag, M. and Engels, J.W. 1989. Synthesis and selective cleavage of oligodeoxyribonucleotides containing non-chiral internucleotide phosphoramidate linkages. Nucl. Acids Res. 17:59735988. Nelson, P.S., Muthini, S., Vierra, M., Acosta, L., and Smith, T.H. 1997. RainbowTM universal CPG: A versatile solid support for oligonucleotide synthesis. Biotechniques 22:752-756. Paul, C.H. and Royappa, A.T. 1996. Acid binding and detritylation during oligonucleotide synthesis. Nucl. Acids Res. 24:3048-3052. Pon, R.T. and Yu, S. 1997. Hydroquinone-O,O diacetic acid (‘Q-linker’) as a replacement for succinyl and oxalyl linker arms in solid phase oligonucleotide synthesis. Nucl. Acids Res. 25:3629-3635. Pon, R.T. and Yu, S. 2001. Linker phosphoramidite reagents for oligonucleotide synthesis on underivatized solid-phase supports. Tetrahedron Lett. 43:8943-8947. Pon, R.T. and Yu, S. 2004. Linker phosphoramidite reagents for the attachment of the first nucleoside to underivatized solid-phase supports. Nucl. Acids Res. 32:623-631. Pon, R.T. and Yu, S. 2005. Tandem oligonucleotide synthesis using linker phosphoramidites. Nucl. Acids Res. 33:1940-1948. Pon, R.T., Usman, N., and Ogilvie, K.K. 1988. Derivatization of controlled pore glass beads for solid phase oligonucleotide synthesis. Biotechniques 6:768-775. Pon, R.T., Yu, S., Guo, Z., Deshmukh, R., and Sanghvi, Y.S. 2001. Reusable solid-phase supports for oligonucleotides and antisense therapeutics. J. Chem. Soc., Perkin Trans. 1 26382643. Pon, R.T., Yu, S., and Sanghvi, Y.S. 2002. Tandem oligonucleotide synthesis on solid-phase supports for the production of multiple oligonucleotides. J. Org. Chem. 67:856-864.
Synthesis of Unmodified Oligonucleotides
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Rayner, S., Brignac, S., Bumeister, R., Belosludtsev, Y., Ward, T., Grant, O., O’Brien, K., Evans, G.A., and Garner, H.R. 1998. MerMade: An oligodeoxyribonucleotide synthesizer for high throughput oligonucleotide production in dual 96-well plates. Genome Res. 8:741747. Scheuerlarsen, C., Rosenbohm, C., Jorgensen, T.J.D., and Wengel, J. 1997. Introduction of a universal solid support for oligonucleotide synthesis. Nucleosides Nucleotides Nucleic Acids 16:67-80. Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., and Gough, G.R. 1995. A universal adapter for chemical synthesis of DNA or RNA on any single type of solid support. Tetrahedron Lett. 36:27-30. Scott, S., Hardy, P., Sheppard, R.C., and McLean, M.J. 1994. A universal support for oligonucleotide synthesis. In Innovation and Perspectives in Solid-Phase Synthesis. Peptides, Proteins, and Nucleic Acids, Biological and Biomedical Applications (R. Epton, ed.) pp. 115-124. Mayflower Worldwide Ltd., Birmingham, Ala.
Septak, M. 1996. Kinetic studies on depurination and detritylation of CPG-bound intermediates during oligonucleotide synthesis. Nucl. Acids Res. 24:3053-3058. Shchepinov, M.S., Case-Green, S.C., and Southern, E.M. 1997. Steric factors influencing hybridisation of nucleic acids to oligonucleotide arrays. Nucl. Acids Res. 25:1155-1161. Sindelar, L.E. and Jaklevic, J.M. 1995. Highthroughput DNA synthesis in a multichannel format. Nucl. Acids Res. 23:982-987. Zhou, X., Cai, S., Hong, A., You, Q., Yu, P., Sheng, N., Srivannavit, O., Muranjan, S., Rouillard, J.M., Xia, Y., Zhang, X., Xiang, Q., Ganesh, R., Zhu, Q., Matejko, A., Gulari, E., and Gao, X. 2004. Microfluidic PicoArray synthesis of oligodeoxynucleotides and simultaneous assembling of multiple DNA sequences. Nucl. Acids Res. 32:5409-5417.
Contributed by Richard T. Pon University of Calgary Calgary, Alberta, Canada
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Microwave-Assisted Functionalization of Solid Supports for Rapid Loading of Nucleosides
UNIT 3.13
The preparation and manufacture of macromolecules such as oligonucleotides, peptides, and carbohydrates are carried out using solid-phase synthesis (Beaucage and Iyer, 1992; Iyer and Beaucage, 1999). The assembly of these macromolecules involves step-wise coupling of individual monomer units to a leader monomer that is tethered to a functionalized solid support. Consequently, efficient approaches for functionalizing the solid support and tethering it to the leader monomer are critically important for successful solid-phase synthesis. Oligonucleotides are generally assembled on controlled-pore glass (CPG) or polystyrenebased supports that are tethered to the leader nucleoside via a linker and a spacer arm (Fig. 3.13.1). Typically, the underivatized solid supports are first derivatized with a reagent capable of incorporating an amino terminus on the support. Being highly versatile, the amino group can be reacted with a variety of reagents to provide supports carrying diverse functionalities such as hydroxyl, carboxyl, and so on. In turn, a carboxyl-terminated solid support can be linked to the 3 -hydroxy group of the leader nucleoside via an ester linkage. Such nucleoside-loaded supports, tethered via a succinic ester linkage to a solid support (Fig. 3.13.1), are most commonly employed in oligonucleotide synthesis. Commercially available nucleoside-loaded supports are very expensive, and many supports that carry modified nucleosides or specialized linkers are not commercially available. Consequently, many laboratories prefer to synthesize nucleoside-loaded supports from amino- or carboxyl-functionalized supports in-house. However, the reported protocols for functionalization and loading of supports are time-consuming, labor-intensive, and not very user-friendly. Clearly, it would be advantageous to have rapid and efficient protocols for the preparation of nucleoside-loaded supports starting from native, underivatized supports. This unit provides convenient and rapid protocols for the preparation of functionalized supports using microwave-assisted procedures starting from native, underivatized supports. The unit also describes improved protocols for the loading of nucleosides on functionalized supports. Thus, starting from underivatized CPG, protocols are described for: (1) the preparation of amino- and carboxyl-functionalized CPG, (2) the loading of nucleosides onto carboxyl-functionalized CPG, and (3) the recovery of excess
Figure 3.13.1 A leader nucleoside attached to a solid support via a succinyl linker and alkyl amino spacer arm.
Contributed by Radhakrishnan P. Iyer, Seetharamaiyer Padmanabhan, and John E. Coughlin Current Protocols in Nucleic Acid Chemistry (2005) 3.13.1-3.13.18 C 2005 by John Wiley and Sons, Inc. Copyright
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3.13.1 Supplement 23
nucleosides following loading of supports. With suitable modifications, these protocols can be applied towards the functionalization and loading of other solid supports. Basic Protocol 1 describes ultra-fast microwave-assisted amination (MAA) of CPG using (3-aminopropyl)triethoxysilane (APTES) without the aid of a solvent (Padmanabhan et al., 2005). Other amination reagents such as (3-aminopropyl)trimethoxysilane (APTMS), aminopropyldimethyl ethoxysilane (APDES), and N-[3(trimethoxysilyl)propyl]ethylenediamine (APTED) can also be used in place of APTES. Basic Protocol 2 describes ultra-fast microwave-assisted succinylation (MAS) of aminofunctionalized CPG. Nucleosides tethered to CPG using a succinyl linkage are almost universally employed in oligonucleotide synthesis. With appropriate modifications, these protocols can also be employed for the rapid attachment of other specialized linkers such as N-methyl sarcosine, hydroquinone-O,O -diacetic acid, and oxalic acid. Basic Protocol 3 deals with loading of nucleosides on succinylated CPG using a LOTUS workstation (Iyer et al., 2005). The Alternate Protocol describes nucleoside loading on CPG using an orbital shaker without the use of LOTUS. This procedure, though less convenient, is suited for smaller-scale operations and can be adopted by laboratories that do not have access to the LOTUS workstation. Basic Protocol 4 outlines a process for the recovery of 5 -O-DMTr-N-protected nucleosides, which are employed in large excess during loading reactions on solid supports, through a simple aqueous workup. BASIC PROTOCOL 1
MICROWAVE-ASSISTED AMINATION OF CONTROLLED-PORE GLASS This protocol is used to attach 3-aminopropyl moieties to commercially available, native controlled-pore glass (S.1; Fig. 3.13.2) using APTES to produce S.2. Previously reported literature methods for amination of CPG (Majors and Hopper, 1974; Tundo and Venturello, 1979; Matteucci and Caruthers, 1981) involve refluxing CPG with APTES in toluene for 48 to 72 hr followed by capping of unreacted hydroxyl groups with trimethylsilyl chloride. Such heterogeneous reactions are difficult to manage on a large scale, require more time for completion, and are not reproducible. The microwave-assisted protocol described here is ultra-fast, efficient, reproducible, and can be performed without the aid of a solvent. The protocol can be employed using other silylating reagents such as APTMS (which also yields the aminated support S.2) or APDES or APTED (to produce the corresponding amino-derivatized supports S.3 and S.4; Fig. 3.13.3). These reagents are available from Alpha Aesar, Acros Organics, or Oakwood Products. For additional information, see
Figure 3.13.2
Microwave-assisted amination (MAA) of CPG using APTES or APTMS.
MicrowaveAssisted Functionalization of Solid Supports
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Figure 3.13.3
Microwave-assisted amination (MAA) of CPG using different silyl reagents.
Background Information, discussion of MAA. Other silica supports such as wide-pore silica can also be employed. Although the procedures described here pertain to largescale work, with appropriate modifications, they can be adapted towards smaller-scale operations.
Materials ◦
Native controlled-pore glass (CPG; 500 A; Prime Synthesis). Various lots of CPG supplied by the manufacturer have the following range of physical properties: ◦ Pore size: 340 to 535 A Bulk density: 0.25 to 0.35 g/cm3 Solid surface area: 101 to 163 m2 /g Pore volume: 1.1 cm3 /g Particle size: 120 to 200 mesh Loading: 184 µmol/g (3-Aminopropyl)triethoxysilane (APTES) Toluene, reagent grade
Synthesis of Unmodified Oligonucleotides
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Methanol, reagent grade Dichloromethane, reagent grade Hexanes, reagent grade Ninhydrin solution, 0.2% (w/v) in ethanol 500-mL thick-walled glass reactor (Spring Bank) fitted with a Teflon screw-cap stopper and chemically resistant O-ring (Chemraz) Domestic microwave oven (800 watt, high power setting) Safety shield (3/4-in. polycarbonate; e.g., VWR) Temperature laser gun (Raytek minitemp) Fritted funnels, medium porosity, small (60 mL) and large (19-cm height × 16-cm diameter) Filter flask Vacuum pump Large glass dish (14 × 23 cm; Anchor Hocking Co.) Aluminum foil Master heat gun (Master Appliance Corp.) 8 × 35–mm flat-bottom sample tubes Vacuum manifold Additional reagents and equipment for amino group estimation by trityl analysis (UNIT 3.2) CAUTION: All microwave reactions should be carried out behind a safety shield, and in a specially fabricated thick-walled glass reactor fitted with a Teflon stopper and chemically resistant O-ring. The safety shield is used to protect against injuries from minor explosions.
Perform amino loading on CPG 1. In a 500-mL thick-walled pressure reactor fitted with a Teflon screw-cap stopper ◦ and a chemically resistant O-ring, combine 100 g native CPG (500 A) and 350 mL APTES. Quickly close the reactor vessel with the Teflon plug and gently mix the contents by shaking. 2. Place the reactor in a domestic microwave oven. 3. Place a safety shield to cover the area around the microwave. 4. Start to heat in 1-min cycles at high power. After each heat cycle, remove the reactor from the microwave oven, mix the contents well by shaking, and measure the temperature by directing the beam of a Raytek laser gun at the upper body of the reactor. Allow the reaction mixture to cool to ∼64◦ to 71◦ C before proceeding with the next heating cycle. CAUTION: The reactor and its contents will be hot. After two heating cycles in the microwave, the outside temperature of the reactor usually rises to ∼82o to 90◦ C. If the reaction mixture is not cooled after each cycle, the O-ring may melt, allowing the contents of the reactor to leak out. The laser gun is a convenient device for measuring the outside temperature of reaction vessels.
5. At the end of a total reaction cycle of ∼8 min, allow the reaction mixture to cool to RT. 6. Carefully open the reactor and filter the reaction mixture in vacuo using a large fritted funnel of medium porosity. MicrowaveAssisted Functionalization of Solid Supports
7. Wash the support twice each with 125 mL toluene, 250 mL methanol, 250 mL DCM, and finally 250 mL hexanes.
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8. Spread the resulting 3-aminopropyl-CPG support (S.2) in a thin layer in a large (14 × 23–cm) dish, cover the dish with perforated aluminum foil, and allow the support to dry for 2 days in a fume hood at ambient temperature. 9. Store the dried support in a closed container for up to 3 to 7 days at ambient temperature, or at 0◦ to 4◦ C for longer periods.
Perform qualitative analysis of amino loading 10. Treat 4 to 5 mg of 3-aminopropyl-CPG in a vial with a few drops of ninhydrin reagent and heat for a few seconds (to near boiling) using a heat gun. The presence of the amino group on the support is indicated by the formation of a strong purple color.
Perform quantitative analysis of amino loading 11. Place a sample (∼100 mg) of 3-aminopropyl-CPG in an 8 × 35–mm flat-bottom sample tube, cover the tube with a Kimwipe tissue and secure it around the neck of the tube, and place the tube in a jar attached to a vacuum manifold. Dry under high vacuum for 12 hr. Drying of the support should be done carefully in vacuo to remove traces of solvent. The use of a static gun (to prevent particles from flying) allows easier handling, weighing, and transferring of the support, which must be handled carefully since the particles are light and fluffy.
12. Determine amino loading by converting amino functions to dimethoxytrityl amino groups and performing trityl analysis (UNIT 3.2).
MICROWAVE-ASSISTED SUCCINYLATION OF 3-AMINOPROPYL-CPG This protocol describes microwave-assisted succinylation (Fig. 3.13.4) of 3-aminopropylCPG (S.2) to produce S.5 in an ultra-fast and efficient procedure. It is performed using reagent-grade dimethylformamide as solvent. The conventional procedure for preparation of succinylated LCAA-CPG (UNIT 3.2) requires overnight reaction in a shaker employing anhydrous pyridine as a solvent, and is less convenient.
BASIC PROTOCOL 2
Materials 3-Aminopropyl-CPG (S.2; see Basic Protocol 1) Succinic anhydride, reagent grade 4-Dimethylaminopyridine (DMAP), reagent grade N,N-Dimethylformamide (DMF), reagent grade, water content <15 ppm Methanol, reagent grade Dichloromethane (DCM), reagent grade Hexanes, reagent grade Ethyl acetate (EtOAc), reagent grade 500-mL thick-walled glass reactor (Spring Bank) fitted with Teflon screw-cap stopper and chemically resistant O-ring (Chemraz) Domestic microwave oven (800 watt, high power setting) Safety shield (3/4-in. polycarbonate; e.g., VWR) Temperature laser gun (Raytek minitemp) Fritted funnels, medium porosity, small (60 mL) and large (19-cm height, 16-cm diameter) Filtration flask 8 × 35–mm flat-bottom sample tubes Master heat gun (Master Appliance Corp.) Large glass dish (14 × 23 cm; Anchor Hocking Co.) Additional reagents and solutions for carboxyl group estimation (UNIT 3.2) Current Protocols in Nucleic Acid Chemistry
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Figure 3.13.4
Microwave-assisted succinylation (MAS) of 3-aminopropyl-CPG.
Perform succinylation 1. Place 150 g of 3-aminopropyl-CPG in a 500-mL thick-walled pressure reactor with a Teflon screw-cap stopper and O-ring. 2. Dissolve 60 g succinic anhydride and 7.2 g DMAP in 450 mL DMF (reagent grade, water content <15 ppm), and add this solution to the reactor containing the support. 3. Mix the contents well. Place the reactor in a microwave oven and set up the safety shield. Initiate eight 30-sec cycles of heating on high power (total time ∼5 min). Allow the dark-colored reaction mixture to cool in between heating cycles. CAUTION: Although temperature is not as critical as in the amination reaction, this microwave-assisted reaction is exothermic. Use shorter heating cycles if necessary.
Monitor succinylation 4. To test the support for the presence of unreacted amino groups, filter a small sample (∼5 mg) of the support under vacuum through a small fritted funnel of medium porosity, and then wash twice each with 10 mL methanol, 10 mL DCM, and 10 mL hexanes. 5. Place the washed succinylated support in a clean 8 × 35–mm flat-bottom sample tube and add 3 drops of ninhydrin solution. Heat the mixture with a heat gun to boiling. If any unreacted amino groups remain on the support, a purple color will develop. The absence of color indicates complete succinylation of the amino groups.
6. If any purple color appears, perform a few more heating cycles until the ninhydrin test is negative.
Filter product and analyze quantitatively 7. After completion of succinylation, let the reactor cool to room temperature. Carefully filter the purple-colored reaction mixture under vacuum through a large frit of medium porosity. 8. Wash the support three times each with 200 mL methanol, 200 mL DCM, 200 mL EtOAc, and finally 200 mL hexanes. 9. Spread the support in a thin layer in a 14 × 23–cm glass tray, cover with perforated aluminum foil, and dry for 48 hr at ambient temperature in a fume hood. 10. Determine the carboxyl loading of the support as described using 4-nitrophenol or N-monomethoxytrityl-6-amino-1-hexanol (see UNIT 3.2). MicrowaveAssisted Functionalization of Solid Supports
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LOADING OF NUCLEOSIDES ON SUCCINYLATED CPG USING LOTUS As is well known, the loading of nucleosides on a solid support is a multi-phase reaction that involves mixing of the nucleoside and various reagents with the solid support in an inert atmosphere (for further discussion, see Background Information). The established protocol (UNIT 3.2) requires the use of anhydrous pyridine as solvent in the presence of an excess of nucleoside (10 mmol per 100 g succinylated support; reviewed in UNIT 3.2). The excess nucleoside used in the reaction cannot be readily recovered afterward. This protocol describes an efficient approach for loading nucleosides on a solid support that involves the use of a specially fabricated reactor known as LOTUS (Fig. 3.13.5) in conjunction with dimethylformamide as a solvent. The LOTUS workstation is ideally suited to perform loading of nucleosides in scales higher than 20 to 50 g of support.
BASIC PROTOCOL 3
Three design elements make the LOTUS workstation unique for performing multi-phase reactions: (1) orbital shaking of the solid and liquid phases, (2) fluidization of phases induced by inert gas, and (c) active recycling of the liquid phase. The mixing process in LOTUS is designated as “orbital shaking coupled with active recycling” (OSCAR) of the liquid phase. Consequently, solid-phase reactions can be performed efficiently in LOTUS. LOTUS has been designed to incorporate pneumatic systems for better control of fluid flow while maintaining an inert atmosphere. LOTUS I has two pneumatically controlled solenoid valves (McMaster Carr) for delivery of reagents from solvent reservoirs under argon. Pressure release valves placed in the argon line provide outlet for release of excess pressure. The activation of the solenoid valves forces pressurized gas into the solvent and reactant chambers, thus allowing liquids to enter into the reactor. The advantage of LOTUS is that the entire process—including loading, capping, filtration, washing, and drying—is carried out using a single assembly in an inert atmosphere with minimal human exposure to toxic solvents and air-borne support particles. In addition, the use of dimethylformamide as a solvent provides several advantages: (1) it allows loading to be carried out with a smaller molar excess of nucleoside, (2) it permits facile monitoring of the loading reaction, and (3) it facilitates the recovery of excess nucleoside following completion of the loading reaction. This feature is particularly useful when expensive nucleosides are employed. The following protocol uses the loading of DMTr-dABz on CPG as an example to illustrate the procedure.
Materials Succinylated CPG (S.5; see Basic Protocol 2) 5 -O-(4,4 -Dimethoxytrityl)-N6 -benzoyl-2 -deoxyadenosine (DMTr-dABz ) or other 5 -O-DMTr-N-protected deoxyribonucleoside (Reliable Biopharmaceutical Corporation; ChemGenes) 4-Dimethylaminopyridine (DMAP), reagent grade Anhydrous N,N-dimethylformamide (DMF), freshly distilled over CaH2 Argon gas Anhydrous triethylamine (TEA), freshly distilled over CaH2 1-[3-(Dimethylamino)propyl]-3-ethylcarbodiimide hydrochloride (EDC), reagent grade Methanol, reagent grade Dichloromethane (DCM), reagent grade Hexanes, reagent grade CAP A and B capping solutions (American International Chemical; ChemGenes) LOTUS reactor workstation (Iyer et al., 2005; Fig. 3.13.5; Spring Bank Technologies) Solid addition funnels Current Protocols in Nucleic Acid Chemistry
Synthesis of Unmodified Oligonucleotides
3.13.7 Supplement 23
1-L round-bottom flask Rubber septa and syringe needles Fritted funnels, small size (60 ml), of medium porosity 8 × 35–mm flat-bottom sample tubes Vacuum manifold Additional reagents and equipment for trityl analysis (UNIT 3.2) CAUTION: DMTr-nucleosides and carbodiimide reagents such as EDC can cause severe skin and eye irritation and allergic reactions. Use appropriate skin and eye protection and wipe all work areas with a wet cloth immediately after use. NOTE: Anhydrous DMF and TEA are prepared by distilling reagent-grade solvents over ◦ calcium hydride and are stored over 4 A molecular sieves. They may also be purchased from commercial vendors (Aldrich, J.T. Baker). If procured from commercial sources, it is important to use certified solvents that have <5 ppm water.
Load nucleoside on CPG using LOTUS 1. Place 100 g succinylated CPG (S.5; 103 µmol carboxyl/g) into the LOTUS reactor through inlet d using a solid addition funnel. 2. Dissolve 19.7 g (3 eq, 30 mmol) DMTr-dABz and 3.66 g (3 eq, 30 mmol) DMAP in 400 mL anhydrous DMF in a 1-L round-bottom flask under argon atmosphere. If carbonyl determination has not been carried out, base reagent quantities on the amino loading of the CPG.
3. Introduce the solution into the LOTUS reactor through inlet port d under a blanket of argon. The inlet port d can be capped with a rubber septum to facilitate liquid transfer under argon. Carry out the transfer of all anhydrous solvents/reagents through a syringe needle or transfer tube (polypropylene) under a nitrogen/argon atmosphere.
4. Shake the contents using the orbital shaker at 100 rpm. 5. Introduce 4.2 mL (30 mmol) anhydrous triethylamine via a syringe needle through inlet port d. 6. Initiate “orbital shaking coupled with active recycling” (OSCAR) by turning on the pump and orbital shaker, and continue for 5-10 min. 7. Add 5.76 g (3 eq, 30 mmol) EDC through inlet port d. 8. Perform OSCAR by carrying out recycling in both the forward and reverse directions for 8 hr. Reverse recycling in an argon atmosphere helps to fluidize the reaction mixture and promote efficient mixing of the solid and liquid phases.
9. Stop recycling and continue orbital shaking alone for ∼8 hr. 10. Monitor the progress of the reaction by periodically removing an aliquot of recycling liquid through sample port o with a small spatula and performing trityl analysis (UNIT 3.2) to determine the amount of DMTr-dABz consumed.
MicrowaveAssisted Functionalization of Solid Supports
Typically a 10-µL aliquot, diluted to 10 mL with 5% DCA in DCM, is used to measure the absorbance of the trityl color and determine the concentration of unconsumed DMTr-dABz remaining in the reaction, using the formula: [DMTr-dABz ] (µmol/mL) = (A503 × D)/76 where A503 is absorbance at 503 nm and D is the dilution factor.
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Figure 3.13.5 Prototype LOTUS reactor and workstation. The LOTUS reaction vessel comprises an upper chamber (a) and a lower chamber (b) held together by a clamp (c). The upper chamber has a gas/solid/liquid inlet (d). Inlet d is used to introduce fresh chemical reactants into the reaction vessel under argon. A pressure release valve (e) is situated on the upper chamber. The lower chamber, equipped with a medium-porosity frit (f), is connected via a valve (s) and adapter (h) to the collection reservoir (i). The adapter also has a gas port (g), suitable for connecting to a vacuum line to draw excess reagents into the collection reservoir as desired. The contents of the collection reservoir can be drained through valve (r). The entire reaction vessel, including the collection reservoir, is mounted on an orbital shaker to ensure thorough and efficient mixing of the solid, liquid, and gas phases during the reaction. The reaction chamber and accessories are maintained under slightly positive inert atmosphere so that air- and moisture-sensitive reactions can be performed. The lower chamber comprises a sampling port (j) that allows a sample of solid matrix to be drawn for analysis during reactions and process operations. Liquid reactants can be independently sampled from an auxiliary port (k) in the recycle path. The online monitoring of both solid and liquid phases allows better process control during nucleoside loading. In order to ensure continuous and close contact between liquid and solid phases, the concept of active recycling is incorporated in the reactor design. Thus, the upper chamber is fitted with a recycled reagent inlet port (l) adapted for connection to flexible, chemically resistant tubing (m) that fits into the pump head of a peristaltic pump (n). The lower chamber also has a recycled reagent outlet port (o) that is adapted for connection to flexible tubing. Port o is connected to the recycled reagent inlet port (l) via tubing through the pump. The pump serves several functions: (1) it can be used to draw the reactants from a reservoir into the reaction vessel, (2) it helps recycle the liquid contents of the reaction vessel through recycled reagent ports o and l, (3) it helps modulate the delivery of reagents or the rate of recycling throughout the reaction via the flow controller (p). Recycling can be done in both forward and reverse directions and at different speeds. Forward recycling generates turbulent forces that assist in mixing of solid and liquid phases, and reverse recycling assists fluidization of solid and liquid phases, thereby aiding mixing. The combined fluid forces generated by forward and reverse recycling facilitates thorough mixing of the solid and liquid reaction phases, resulting in rapid reaction kinetics and efficient and reproducible loading without causing mechanical abrasion of the solid support. Reprinted from Iyer et al. (2005) with permission from Organic Preparations and Procedures International. Current Protocols in Nucleic Acid Chemistry
Synthesis of Unmodified Oligonucleotides
3.13.9 Supplement 23
11. After about 12 hr, remove a slurry sample (∼100 mg) of the support through port d, filter under vacuum through a small fritted funnel of medium porosity, and wash with 30 mL each methanol, DCM, and hexanes. Dry the support in vacuo using a vacuum pump, and perform trityl analysis on the dried support. Typically, loading of 60 to 70 µmol nucleoside/gram support is readily obtained after a 12-hr reaction.
12. If the loading is low, add (in order) 3.7 g DMAP (as a solution in 20 mL DMF), 5 mL triethylamine, and 2.95 g EDC, and continue OSCAR. It is critical that anhydrous solvents and reagents are employed in the reaction and solution. Also, reagents should be added sequentially as mentioned in order to obtain high loading within 24 hr.
Perform workup 13. At the end of the reaction, filter the contents of the reactor under vacuum through the medium-porosity frit f in the reactor, and collect the filtrate through port o. Set aside the filtrate for recovery of excess unreacted nucleoside (see Basic Protocol 4). 14. Wash the support (via port o) two times with 400 mL each methanol, then DCM, and finally hexanes. After each addition of solvent, initiate OSCAR for 10 min and perform filtration under vacuum. 15. Dry the support thoroughly under vacuum in the LOTUS apparatus. 16. Remove a sample of the solid support and perform trityl analysis (UNIT 3.2). Typically, nucleoside loadings of 75 to 80 µmol/g are obtained.
Cap support using LOTUS 17. Add 325 mL each of CAP A and CAP B to the nucleoside-loaded CPG through port d. 18. Perform OSCAR for 3 hr using both forward and reverse recycling. 19. Filter the capped support and then wash twice with 300 mL each methanol, then DCM, and then hexanes. 20. Place the support in an 8 × 35–mm flat-bottom sample tube, secure a Kimwipe tissue around the neck of the tube, and place it in a jar attached to a vacuum manifold. Dry the support in vacuo. Drying of support should be done carefully in vacuo to remove traces of solvent. Also, the use of a static gun allows easier handling and transferring of the support since the particles are light and fluffy.
21. Determine the nucleoside loading of dried support S.7 by trityl analysis. Typically, nucleoside loadings of 70 to 75 µmol/g are obtained.
22. Store the support in a closed container at ambient temperature for up to 3 to 7 days, or at 0◦ to 4◦ C for longer periods. ALTERNATE PROTOCOL
LOADING OF NUCLEOSIDES ON SUCCINYLATED CPG USING ORBITAL SHAKING This protocol deals with nucleoside loading on succinylated CPG using an orbital shaker instead of LOTUS. This procedure, though less convenient, can be followed in laboratories that do not have access to LOTUS.
MicrowaveAssisted Functionalization of Solid Supports
Additional Materials (also see Basic Protocol 3) 1-L single-neck round-bottom flasks Rubber septum
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Orbital shaker (e.g., Lab-Line Orbit Environ-Shaker) Fritted funnel, large size (19-cm height × 16-cm diameter), of medium porosity Load nucleoside on CPG 1. Load 65 g succinylated CPG (87 µmol carboxyl/g) in a 1-L single-neck round-bottom flask. 2. In a separate round-bottom flask, with stirring, dissolve 18.7 g (5 eq relative to carboxyl load of CPG) DMTr-dABz and 3.5 g DMAP in 260 mL anhydrous DMF (freshly distilled over CaH2 ). Add this solution to the support. Carry out the transfer of all anhydrous solvents/reagents through a syringe or transfer tube (polypropylene) under nitrogen/argon atmosphere.
3. Add 4 mL anhydrous triethylamine and then 5.47 g (5 eq) EDC to the above reaction mixture. 4. Seal the reaction flask with a rubber septum and mix under gentle orbital shaking (∼100 rpm) overnight. Set the orbital shaker at a low speed to enable a gentle rocking motion for mixing of the phases. Shaking at high speed can cause the fragile support particles to break up.
5. Monitor the coupling reaction by removing a small sample of reaction mixture (50 to 100 mg) using a disposable pipet. Filter under vacuum through a small fritted funnel of medium porosity and wash twice with 10 mL each methanol and then DCM. 6. Dry ∼20 mg sample and determine the loading by trityl analysis (UNIT 3.2). If the loading is not acceptable (>65 µmol/g), continue mixing for an extended period of time and, if necessary, add more DMAP (as solution in DMF) followed by TEA and then EDC to the reaction mixture.
Perform workup 7. When loading is complete, filter the reaction mixture through a large fritted funnel. 8. Wash the support twice with 300 mL each methanol, then DCM, and then hexanes. Dry in air overnight in the funnel.
Cap unreacted nucleophilic groups 9. Mix nucleoside-loaded CPG S.7 with 325 mL each of CAP A and CAP B mixtures in a 1-L single-neck round-bottom flask for 6 hr under orbital shaking. 10. Filter the capped support and wash with 300 mL each methanol, then DCM, and then hexanes twice. 11. Dry the support using a vacuum pump and determine the loading of dried support through trityl analysis (UNIT 3.2). Loading should typically be around 70 µmol/g.
12. Store the support in a closed container at ambient temperature for up to 3 to 7 days, or at 0◦ to 4◦ C for longer periods.
RECOVERY OF EXCESS UNREACTED NUCLEOSIDE 5 -O-DMTr-N-protected
A large excess of nucleosides are employed during loading reactions on solid supports. The use of DMF as a solvent for loading facilitates recovery of most of this excess nucleoside by simple aqueous workup. The following illustrative protocol deals with the recovery of excess DMTr-dABz from the filtrate obtained following the loading reaction in Basic Protocol 3 or the Alternate Protocol.
BASIC PROTOCOL 4
Synthesis of Unmodified Oligonucleotides
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Supplement 23
Materials Nucleoside-containing DMF filtrate (e.g., Basic Protocol 3, step 13) Sodium chloride Chloroform, reagent grade 5% (w/v) aqueous citric acid, reagent grade 5% (w/v) aqueous sodium carbonate, reagent grade Brine (saturated aqueous NaCl) Anhydrous sodium sulfate, reagent grade Separatory funnel with stopper 24-cm Buchner funnel with Whatman no. 1 filter paper Rotary evaporator Vacuum pump 1. Measure the volume of DMF filtrate obtained after isolation of DMTr-dABz -loaded CPG. 2. Slowly pour the filtrate, under stirring, into 10 vol ice-cold water containing 2 to 5 g NaCl. A white solid begins to separate.
3. Let stand for 1 hr at 4◦ C. 4. Filter the solid that separates under vacuum using a 24-cm Buchner funnel with Whatman no. 1 filter paper and wash the solid with water. 5. Take up the wet solid in chloroform and transfer the dissolved solution to a separatory funnel. 6. Separate the lower organic layer and wash it with 300 mL each of 5% aqueous citric acid (to remove any dissolved DMAP), followed by 5% aqueous sodium carbonate, and finally brine. 7. Dry the chloroform layer over 20 g anhydrous sodium sulfate and filter through a funnel. 8. Concentrate the chloroform solution on a rotary evaporator to give the nucleoside as a white residue. 9. Dry in vacuo overnight using a vacuum pump to obtain DMTr-dABz as a white solid of acceptable purity. The recovery of the excess nucleoside is >70%. The 1 H NMR spectrum of recovered DMTr-dABz is found to be identical to a standard sample obtained from Reliable Biopharmaceuticals.
COMMENTARY Background Information
MicrowaveAssisted Functionalization of Solid Supports
Synthetic oligonucleotides find extensive applications in target validation, functional genomics, diagnostics, and therapeutics, and as experimental tools in molecular biology (Sproat, 1995; Vlassov et al., 1997; Verma and Eckstein, 1998). As potential therapeutic agents, synthetic oligonucleotides have been designed to down-regulate intracellular RNA expression through RNA interference and ribozyme- and antisense-
based approaches, as well as to inhibit protein function through the aptamer approach (Sharp, 2001; Hannon, 2002; Zamore, 2002; Iyer et al., 2003; Manoharan, 2004). Two oligonucleotide drugs, Vitravene (an antisense oligonucleotide) and Macugen (an aptamer), have been approved for clinical use. The synthesis and manufacture of oligonucleotides for various applications are carried out using solid-phase phosphoramidite chemistry or H-phosphonate chemistry in
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Figure 3.13.6
Strategies for the functionalization and loading of solid supports.
automated synthesizers (Beaucage and Caruthers, 1981; Beaucage and Iyer, 1992; Iyer and Beaucage, 1999; UNITS 3.3–3.5). In solid-phase oligonucleotide assembly, monomeric units are sequentially coupled to a leader nucleoside bound to a solid support through a series of reaction cycles. Consequently, the availability of appropriate nucleoside-loaded supports is critical for the manufacture of oligonucleotides. Although several improvements in oligonucleotide synthesis have been reported over the past two decades (Iyer and Beaucage, 1999), only limited reports exist for the functionalization of solid supports or for the preparation of support-bound nucleosides (Majors and Hopper, 1974; Tundo and Venturello, 1979; Matteucci and Caruthers, 1981; Damha et al., 1990; UNIT 3.2). The functionalization and loading of supports are traditionally carried out according to path A in Figure 3.13.6, wherein native controlled-pore glass (CPG; S.1) is derivatized to give the aminopropylsilyl derivative S.2, is succinylated to produce the carboxy-tethered CPG derivative S.5, and is finally coupled at the carboxyl terminus to a 5 -DMTr-protected nucleoside (S.6) by esterification to give S.7. Path B is followed as an alternate approach where S.2 is coupled with the presynthesized nucleoside hemisuccinate derivative S.8 to produce S.7. Although these reported protocols for producing S.7 from S.1 have helped advance oligonucleotide synthesis
technology, the protocols are time-consuming, labor-intensive, and show high batch-to-batch variability in loading. There are some reports of improved procedures for loading of nucleosides on functionalized supports, but their utility for routine use has not been fully explored. Over the past two decades, microwaveassisted procedures have been successfully employed for the rapid and efficient synthesis of different classes of organic compounds (for reviews see Chatti et al., 2001; Listrom et al., 2001; Bose et al., 2002). Microwave-assisted organic synthesis has several advantages: (1) ultra-fast reaction kinetics, (2) cleaner reactions with improved yields and reduced formation of side products, (3) ability to effect chemo-, regio-, and stereoselective transformations, (4) flexibility to perform reactions with or without solvents, (5) economical and eco-friendly processes compared to conventional reactions, and (6) successful product formation in reactions that fail under conventional conditions. There is only one prior report of the application of microwave-assisted procedures in the solid-phase synthesis of nucleic acids, involving rapid deprotection and cleavage of oligonucleotides (Kumar and Gupta, 1997). Microwave-assisted amination (MAA) of native CPG Microwave-assisted procedures have recently been applied as an alternative approach for the functionalization of CPG using
Synthesis of Unmodified Oligonucleotides
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Supplement 23
Table 3.13.1 Influence of Solvents on Microwave-Assisted Amination of CPGa
Amination reagent
Solvent
Loading (µmol/g)
APTES
Neat
113
DMSO
116
DMF
86
a From Padmanabhan et al. (2005) with permission from Elsevier.
Table 3.13.2 Influence of Silyl Agents on Microwave-Assisted Amination of CPGa
Amination reagent
Reaction conditions
Loading (µmol/g)
APTES
Standard conditionsb
73
APTES
MAA, 5 min
98-113
APTMS
MAA, 5 min
110
APDES
MAA, 5 min
96
APTED
MAA, 5 min
79
a From Padmanabhan et al. (2005) with permission from Elsevier. b Loading with standard non-microwave methods (for comparison).
Table 3.13.3 Influence of Additives on MicrowaveAssisted Amination of CPGa
Reagent
Additiveb
APTES
DMAP
110
TFA
116
PTSA
128
BF3 ·Et2 O
45
Loading (µmol/g)
a From Padmanabhan et al. (2005) with permission form Elsevier. b Abbreviations: DMAP, 4-dimethylaminopyridine; PTSA, p-toluene-
sulfonic acid; TFA, trifluoroacetic acid.
MicrowaveAssisted Functionalization of Solid Supports
a domestic microwave oven (Padmanabhan et al., 2005). The microwave-assisted amination (MAA) protocol described in this unit is an optimized protocol that uses neat APTES without any solvent. The effect on loading of using different solvents in the MAA of CPG with APTES is given in Table 3.13.1. Although MAA can be conducted in DMF and DMSO, the use of CPG in neat APTES is more convenient and more eco-friendly. It is pertinent to mention that since an excess of APTES is used in this protocol, MAA of CPG has been attempted using APTES recovered from a previous MAA reaction. This procedure was unsuccessful, probably due to contamination of the recovered APTES with liberated ethanol. In attempts to see if higher amino loadings could be achieved with different amination reagents, MAA reactions were carried out (all neat) using (3-aminopropyl)trimethoxysilane
(APTMS) and N-[(3-trimethoxysilyl)propyl] ethylenediamine (APTED; Fig. 3.13.3). Although the corresponding aminated CPGs S.2 and S.4 were obtained, loading was not increased beyond that obtained with APTES. It is believed that amination involves the condensation of three hydroxyl groups on the CPG matrix with the three ethoxy groups of APTES to form S.2. It seemed reasonable that if each of the hydroxyl groups of CPG could be engaged in reaction with a single molecule of a monoethoxysilane derivative such as aminopropyldimethyl ethoxysilane (APDES), amino-CPG S.3 could be achieved with increased amino loading. However, although the MAA reaction of CPG with APDES did give the corresponding aminated product S.3, the loading was not increased beyond 96 µmol/g (Table 3.13.2). In another experiment, the amino-CPG obtained following the first MAA
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with APDES was subjected to a second MAA using APTES. There was no further improvement in loading, suggesting that all available sites on the CPG had been functionalized during the first reaction. As a safety precaution, it should also be noted that during MAA of CPG with APDES in the presence of tin(IV) chloride as a catalyst, an explosive reaction resulted. The MAA of CPG with APTES was also evaluated using different additives (Table 3.13.3). The best results were obtained with ptoluenesulfonic acid and TFA, giving S.2 with amino loadings of 128 and 116 µmol/g, respectively. With boron trifluoride etherate as a catalyst, the product S.2 had considerably reduced loading (∼40 µmol/g). These results suggest that additives and solvents influence the amino loading on CPG and hence the final loading of nucleosides. It is pertinent to mention that the MAA protocol can be modified to prepare amino-functionalized supports with a predetermined loading level (unpub. observ.). Microwave-assisted succinylation (MAS) of aminated CPG Following the success of MAA of CPG, the MAS of the functionalized support was attempted under microwave conditions. Microwave-assisted reaction of aminated CPG with succinic anhydride without the aid of any solvent resulted in a very dark yellow-colored support, probably due to the formation of an imide rather than the expected acid. Nevertheless, with DMF as a solvent, the MAS of the aminated CPG (50 to 100 g scale) was achieved successfully to give S.5 in the presence of a catalytic amount of DMAP in less than 5 min. Completion of the reaction was ascertained by testing for the absence of the amino group on a sample of S.2. The use of DMF in place of pyridine makes the MAS procedure highly attractive for preparation of succinylated CPG. In addition to DMF, the succinylation can also be carried out using DMSO, dimethylacetamide, or acetonitrile. Loading of nucleosides on succinylated CPG Following the success of microwaveassisted protocols for ultra-fast preparation of functionalized CPG, the loading of nucleosides on solid supports under microwave conditions was also evaluated. Preliminary studies of nucleoside loading of functionalized supports S.2 and S.5 (Fig. 3.13.6, paths A and B) using microwave conditions have been en-
couraging. The protocols are being optimized, and will be reported in due course. With the succinylated CPG S.5 at hand, an improved protocol for efficient loading of nucleosides on S.5 was developed using a specially fabricated reactor in conjunction with DMF as a solvent (Iyer et al., 2004a,b, 2005). The use of DMF facilitated efficient recovery of excess nucleoside, thereby resulting in higher economy of operations. Efficient mixing and contact of the functionalized solid support with the nucleoside and various reagents are necessary to achieve a high loading. The LOTUS workstation (Fig. 3.13.5) is ideally suited to accomplish this task. LOTUS enables the performance of all operations in a closed system, thereby minimizing user exposure to toxic solvents and air-borne support particles. Three LOTUS design elements facilitate efficient solid-liquid mixing: (1) orbital shaking of the solid and liquid phases, (2) fluidization of phases induced by inert gas, and (3) active recycling of the liquid phase. The mixing process in LOTUS is designated as “orbital shaking coupled with active recycling” (OSCAR). LOTUS consists of a reaction chamber equipped with accessories and a workstation to control operations by which both recycling and orbital shaking can be optionally and independently controlled. As shown in Figure 3.13.5, since all operations are performed in a closed system, human exposure to reagents, solvents, and supports is minimized while anhydrous conditions and an inert atmosphere are maintained. Basic Protocol 3 describes the use of LOTUS for loading 5 -O-(4,4 -dimethoxytrityl)N6 -benzoyl-2 -deoxyadenosine (DMTr-dABz ) on succinylated CPG. Using 100 g support in each batch, high nucleoside loadings of 70 to 80 µmol/g were achieved with only three equivalents of nucleoside in 12 to 14 hr. Following the reaction, both filtration and drying operations were performed in LOTUS. The nucleoside-loaded CPG was then treated with CAP A and CAP B in LOTUS. Thus, nucleoside loading as well as the required process operations could be conveniently performed using LOTUS as a multi-task workstation. Additionally, the use of DMF instead of pyridine prevented any unpleasant odor and also aided the facile recovery of excess nucleoside, which is especially desirable when using expensive nucleosides. Indeed, the recovery of excess nucleoside was carried out by simple aqueous workup of the DMF filtrate. Subsequently, the recovered nucleoside (DMTr-dABz ) was successfully loaded onto
Synthesis of Unmodified Oligonucleotides
3.13.15 Current Protocols in Nucleic Acid Chemistry
Supplement 23
succinylated CPG. The efficient intermixing of phases, induced by OSCAR, and on-line monitoring are key to faster reactions and dramatically improved nucleoside loading on a solid support using LOTUS. The nucleosideloaded support thus obtained has been successfully used for large-scale solid-phase synthesis of dinucleotide analogs as potential anti-HBV agents (Iyer et al., 2004a,b). Nucleoside loading on succinylated CPG can also be carried out by orbital shaking using DMF as a solvent, although the disadvantages of this protocol have been mentioned previously. Loading was carried out on succinylated CPG (100 g batches) using 5 equivalents of DMTr-dABz . Using this protocol, nucleoside-loaded CPG was obtained with loadings ranging from 60 to 70 µmol/g. Using similar procedures, nucleoside loadings on carboxy-terminated wide-pore silica, Tentagel, and aminomethyl polystyrene have also been performed. For comparison, the CPG-loaded nucleoside S.7 prepared as above and that prepared by the conventional method have been employed in a 10-µmol synthesis of dimer and 20-mer oligonucleotides (PO and PS) in an Expedite Synthesizer. In both instances, the stepwise coupling yield was >98% as ascertained by trityl analysis. Following the synthesis, each of the CPGs was treated with 28% NH4 OH at 55◦ C for 12 hr to isolate the fully deprotected di- and polynucleotides. RP-HPLC analysis of the crude mixtures showed that the profiles of compounds prepared using both supports were similar.
In conclusion, a microwave-assisted protocol for rapid and efficient functionalization of CPG is presented in which CPG S.5 carrying a carboxy terminus can be obtained from native CPG within a few hours, in contrast to conventional procedures that require several days. Efficient processes for loading of nucleosides on the resulting functionalized support are also described using anhydrous DMF as a solvent. The use of a novel reactor in conjunction with recycling technology enables efficient loading. The methodologies described here can be applied for functionalization of other solid supports and for loading of supports with nucleosides, amino acids, and small molecules for solid-phase synthesis of oligonucleotides, peptides, and ligands, respectively. The approach described here can potentially be employed for rapid functionalization of other solid matrices in the form of beads, slides, pins for application in microarrays, combinatorial chemistry including medical diagnostics, environmental clean-up (removal of toxic materials), radioimmunoassays, fluorescent immunoassays, ELISAs, and affinity chromatography.
Critical Parameters and Troubleshooting MAA of CPG MAA is an exothermic reaction and attention should be paid to ensure that reaction contents do not overheat. Accordingly, it is advisable that heat cycles be adjusted, as needed, after a pilot run on a small scale. Table 3.13.4 shows the outside temperature of the reactor
Table 3.13.4 Temperature Measurements During MicrowaveAssisted Amination of CPG with APTESa
MicrowaveAssisted Functionalization of Solid Supports
Cycle no.
Temperature after microwave exposure
Cooling temperature
1
45
—
2
82
70
3
82
68
4
88
68
5
86
68
6
90
65
7
88
71
8
84
68
9
91
64
10
83
r.t.
a Temperature measurements were made using a Raytek Minitemp laser gun. r.t.,
room temperature.
3.13.16 Supplement 23
Current Protocols in Nucleic Acid Chemistry
during heating and cooling cycles during MAA of 20 g CPG in the presence of 80 mL APTES. MAS of CPG It is important to ensure that reagents and catalysts are fresh or have been properly stored under anhydrous conditions after being opened. Succinic anhydride is hygroscopic and new containers should be stored in a desiccator after being opened. Reagent-grade DMF should be properly sealed after use and stored over molecular sieves to prevent moisture build-up. Like MAA, MAS is an exothermic reaction and attention should be paid to ensure that reaction contents do not overheat. Accordingly, it is advisable that heat cycles be suitably adjusted after a pilot run. Nucleoside loading It is important to ensure that coupling reagents and catalysts are anhydrous and fresh, or have been properly stored under anhydrous conditions after being opened. Often, lower loading yields or failure in loading reactions are a result of moisture in the reaction medium. DMTr-protected nucleosides should also be securely closed and stored at −20◦ C and thawed before opening the container. DMF and TEA should be freshly distilled from CaH2 , and dry solvents should be stored over molecular sieves. The transfer of all anhydrous solvents/reagents should be carried out through a syringe or polypropylene transfer tube under nitrogen/argon atmosphere. It is critical that anhydrous solvents and reagents are employed in the reaction and solutions. Reagents should be added sequentially as mentioned in order to obtain high loading within 24 hr. When an orbital shaker is used for loading, it should be set at a low speed to enable a gentle rocking motion for mixing of the phases. Shaking at high speed can cause the fragile particles to break up. Success in solid-support reactions is dependent on reagent concentrations. Reducing solvent volume, while ensuring that the solid support remains properly suspended in the solution, can increase the concentration of nucleoside and/or coupling reagents. Handling of solid support The use of a static gun allows easier handling and transferring of the support since the particles are light and fluffy. Prior to performing trityl assays, drying of support should be done carefully in vacuo to remove traces of
solvent. Place the support in a sample tube, secure a Kimwipe tissue around the neck, and place it in a jar attached to a vacuum manifold. Always release the vacuum slowly to minimize particle losses. As mentioned previously, although CPG particles are more rigid than silica particles, they are nevertheless fragile. It is recommended that orbital shaking be carried out at low speed for effective mixing. Use of magnetic stirrers or overhead stirrers should be completely avoided for mixing. Capping of unreacted amino groups on the support Although each amino group on the aminated CPG is expected to be coupled to succinic acid following MAS, there is a slight possibility that some unreacted amino groups will remain on the support. Uncapped amino groups on a nucleoside-loaded CPG could potentially consume phosphoramidite reagents and lower the synthesis yields if employed in oligonucleotide synthesis. An excellent discussion on the deleterious effects of uncapped amino groups on CPG can be found in UNIT 3.2. A capping step at the end of loading is essential to convert each amino functionality to an acetamido group and prevent its reaction with reagents employed in oligonucleotide synthesis.
Anticipated Results The protocols described in this unit are extremely useful for the rapid preparation of highly loaded CPG supports in a reproducible fashion. The microwave-assisted protocols for solid support functionalization are very reliable and easy to execute, even for an inexperienced investigator. It is important to use microwave-specific glassware along with all precautions mentioned in the protocol. It is possible to scale up these processes further using commercial microwave workstations using continuous flow or batch process technologies. The highly loaded supports obtained using these protocols are especially beneficial in synthesis and manufacture of 20- to 30-mer oligonucleotides. Since larger amounts of oligonucleotide can be produced in each synthesis, both scale and economy of operation are greatly enhanced. The protocols have been evaluated for the preparation of other supports such as Tentagel and aminomethyl polystyrene. The protocol also illustrates the utility of the LOTUS workstation in performing solid-phase synthesis.
Synthesis of Unmodified Oligonucleotides
3.13.17 Current Protocols in Nucleic Acid Chemistry
Supplement 23
Time Considerations Starting from native CPG, amino and carboxy functionalization reactions are carried out within minutes using these protocols. The slower steps in the process are the workup steps such as filtration, washing, and final drying, which depend on the scale of the operation.
Acknowledgments Support of this research from the National Institutes of Health, under Research Project Cooperative Agreement Grant Award 5 UO1 AI058270-02, is gratefully acknowledged.
Literature Cited Beaucage, S.L. and Caruthers, M.H. 1981. Deoxynucleoside phosphoramidites: A new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Lett. 22:1859-1862. Beaucage, S.L. and Iyer, R.P. 1992. Advances in the synthesis of oligonucleotides by the phosphoramidite approach. Tetrahedron 48:2223-2311. Bose, A.K., Manhas, M.S., Ganguly, S.N., Sharma, A.N., and Banik, B.K. 2002. MORE chemistry for less pollution: Applications for process development. Synthesis 11:1578-1591. Chatti, S., Bortolussi, M., and Loupy, A. 2001. Cation and leaving group effects in isosorbide alkylation under microwave in phase-transfer catalysis. Tetrahedron 57:4365-4370. Damha, M.J., Giannaris, P.A., and Zabarylo, S.V. 1990. An improved procedure for derivatization of controlled-pore glass beads for solidphase oligonucleotide synthesis. Nucl. Acids Res. 18:3813-3821. Hannon, G.J. 2002. RNA interference. Nature 418:244-251. Iyer, R.P. and Beaucage, S.L. 1999. Oligonucleotide synthesis. In Comprehensive Natural Products Chemistry, Vol. 7: DNA and Aspects of Molecular Biology (E.T. Kool, ed.) pp. 105-152. Elsevier Science, London. Iyer, R.P., Kuchimanchi, S.N., and Pandey, R.K. 2003. RNA interference: An exciting new approach for target validation, gene expression analysis and therapeutics. Drugs of the Future 28:51-59. Iyer, R.P., Jin, Y., Roland, A., Morrey, J.D., Mounir, S., and Korba, B. 2004a. Phosphorothioate Diand tri-nucleotides as a novel class of antiHBV agents. Antimicrob. Agents Chemother. 48:2199-2205.
MicrowaveAssisted Functionalization of Solid Supports
Iyer, R.P., Coughlin, J.E., and Padmanabhan, S. 2005. Efficient and rapid processes for loading of nucleosides on solid supports. Org. Prep. Proc. Intl. 37:205-212; Iyer, R.P., Coughlin, J.E., and Padmanabhan, S. Patent pending. Kumar, P. and Gupta, K.C. 1997. Microwaveassisted rapid deprotection of oligodeoxyribonucleotides. Nucl. Acids Res. 25:5127-5129. Listrom, P., Tierney, J., Wathey, B., and Westeman, J. 2001. Microwave assisted organic chemistry: A review. Tetrahedron 57:9225-9283. Majors, R.E. and Hopper, M.J. 1974. Studies of siloxane phases bonded to silica gel for use in high-performance liquid chromatography. J. Chrom. Sci. 12:767-778. Manoharan, M. 2004. RNA interference and chemicaly modified small interfering RNAs. Curr. Opin. Chem. Biol. 8:570-579. Matteucci, M.D. and Caruthers, M.H. 1981. The synthesis of deoxyoligonucleotides on a polymer support. J. Am. Chem. Soc. 103:3185-3190. Padmanabhan, S., Coughlin, J.E., and Iyer, R. P. 2005. Microwave-assisted rapid and efficient functionalization of solid supports. Tetrahedron Lett. 46:343-347; Padmanabhan, S., Iyer, R.P. Patent pending. Sharp, P.A. 2001. RNA interference. Genes Dev. 15:485-490. Sproat, B.S. 1995. Chemistry and applications of oligonucleotide analogues. J. Biotechnol. 41:221-238. Tundo, R.P. and Venturello, P. 1979. Synthesis, catalytic activity, and behavior of phase-transfer catalysts supported on silica gel. Strong influence of substrate adsorption on the polar polymeric matrix on the efficiency of the immobilized phosphonium salts. J. Am. Chem. Soc. 101:6606-6613. Vlassov, V.V., Vlassova, I.E., and Pautova, L.V. 1997. Oligonucleotides and polynucleotides as biologically active compounds. Prog. Nucleic Acids Res. Mol. Biol. 57:95-143. Verma, S. and Eckstein, F. 1998. Modified oligonucleotides: Synthesis and strategy for users. Annu. Rev. Biochem. 67:99-134. Zamore, P.D. 2002. Ancient pathways programmed by small RNAs. Science 296:1265-1269.
Contributed by Radhakrishnan P. Iyer, Seetharamaiyer Padmanabhan, and John E. Coughlin Spring Bank Technologies Milford, Massachusetts
Iyer, R.P., Roland, A., Jin, Y., Mounir, S., Korba, B., Julander, J.G., and Morrey, J.D. 2004b. Antihepatitis B virus activity of ORI-9020, a novel phosphorothioate dinucleotide, in a transgenic mouse model. Antimicrob. Agents Chemother. 48:2318-2320.
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Current Protocols in Nucleic Acid Chemistry
Solution-Phase Synthesis of Di- and Trinucleotides Using Polymer-Supported Reagents
UNIT 3.14
This unit describes an alternate methodology to chemically synthesize protected phosphodiester and phosphorothioate di- and trinucleotides in solution by means of solid-supported reagents (Dueymes et al., 2005). Such a synthesis uses commercially available 5 -O-dimethoxytrityl-3 -O-(2-cyanoethyl-N,N -diisopropyl)-N-protected2 -deoxynucleoside phosphoramidites and 3 -O-levulinyl-N-protected nucleosides in combination with polyvinyl pyridinium tosylate as the activator and polystyrene-bound trimethylammonium periodate or tetrathionate as the oxidizing or sulfurizing agent, respectively. The levulinyl group was chosen for 3 -O-protection because it is orthogonal to the other protecting groups borne by the dinucleotide, i.e., dimethoxytrityl, cyanoethyl, benzoyl, and isobutyryl. The coupling and oxidizing solid-supported reagents are commercially available or can be easily prepared from commercial ion-exchange resins, as described here. The original sulfurizing reagent is attached to Amberlyst A-26 anion-exchange resin by simple stirring of the resin in an aqueous solution of potassium tetrathionate. After completion of each reaction, the solid-supported reagents are removed by filtration. The fully protected building blocks are 5 -O-detritylated with 2% p-toluenesulfonic acid or 3 -O-delevulinylated by means of anionic-polymer-bound hydrazine. Reaction work-ups consist of filtrations, aqueous extractions, and precipitations. This approach avoids the use of any chromatography process. Generally, dimer synthesis is performed at up to a 10-mmol scale, requiring only the usual laboratory glassware.
SYNTHESIS OF 5 -OH-3 -O-LEVULINYL DINUCLEOSIDE CYANOETHYLPHOSPHOTRIESTERS
BASIC PROTOCOL
This protocol and Alternate Protocol 1 describe the solution-phase synthesis of 3 O-levulinylated dinucleotides with a phosphotriester or thionophosphotriester linkage, respectively, by means of the phosphoramidite method, according to the scheme shown in Figure 3.14.1. The procedures describe synthesis at a 10-mmol scale, although they can be scaled up. These protocols are also suitable for the synthesis of 3 -O-protected trinucleotides with phosphotriester or thionophosphotriester linkages. In this protocol, synthesis of a deoxyguanosine dinucleotide is presented as an example of a dinucleotide with a phosphotriester linkage.
Materials 3 -O-Levulinyl nucleosides (S.4; UNIT 2.11; Samchully Pharmaceuticals; Reese et al., 1998): 3 -O-Levulinyl-N 6 -benzoyl-2 -deoxyadenosine 3 -O-Levulinyl-N 4 -benzoyl-2 -deoxycytidine 3 -O-Levulinyl-N 2 -isobutyryl-2 -deoxyguanosine 3 -O-Levulinyl-thymidine Standard commercial 3 -phosphoramidites (S.3; e.g., Pierce, Cruachem): 5 -O-Dimethoxytrityl-3 -O-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite]N 6 -benzoyl-2 -deoxyadenosine 5 -O-Dimethoxytrityl-3 -O-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite]N 4 -benzoyl-2 -deoxycytidine 5 -O-Dimethoxytrityl-3 -O-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite]N2 -isobutyryl-2 -deoxyguanosine 5 -O-Dimethoxytrityl-3 -O-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite]thymidine Contributed by Franc¸ois Morvan and Jean-Jacques Vasseur Current Protocols in Nucleic Acid Chemistry (2006) 3.14.1-3.14.15 C 2006 by John Wiley & Sons, Inc. Copyright
Synthesis of Unmodified Oligonucleotides
3.14.1 Supplement 26
Acetonitrile (<10 ppm H2 O) Dichloromethane (<0.005% H2 O) Poly(4-vinylpyridinium-p-toluenesulfonate) resin (S.2; Aldrich; dry 10 hr at 80◦ C under 30 mbar vacuum over P2 O5 ; store in desiccator at room temperature up to several months under 10 mbar vacuum over P2 O5 ) Argon source 50 mM tetraethylammonium acetate (TEAA), pH 7.5 Amberlyst A-26 ion-exchange resin, periodate form (S.7; see Support Protocol 1) Methanol p-Toluenesulfonic acid Saturated aqueous NaHCO3 solution 0.2 M sodium bisulfite/0.5 M sodium chloride Na2 SO4 , anhydrous Diethyl ether P2 O5 Shaker (Edmund Buhler SM-30 or equivalent) ◦ HPLC column: Nucleosil C18, 5 µm, 100 A, 150 × 4.6 mm Fritted-glass funnel, 10 cm diameter, 15 cm height Rotary evaporator 2-L separatory funnel Vacuum desiccator Additional reagents and equipment for HPLC (UNIT 10.5), thin-layer chromatography (TLC; APPENDIX 3D), and MALDI-TOF mass spectrometry (UNIT 10.1) Perform coupling reaction 1. Azeotrope (coevaporate) 4.35 g (10.0 mmol) of 3 -O-levulinyl-N2 -isobutyryl-2 deoxyguanosine (S.4) and 10.74 g (13.0 mmol) of 5 -O-dimethoxytrityl-3 -[(2cyanoethyl)-N, N-diisopropylphosphoramidite]-N2 -isobutyryl-2 -deoxyguanosine (S.3) twice, each time with 100 mL acetonitrile, then dissolve in 100 mL dichloromethane. Before use, each nucleoside derivative is stored in a dessicator under reduced pressure (10 mbar) for 14 hr. For generation of other dimers, the phosphoramidite is generally used in an excess of 1.3 to 1.5 molar equivalents with respect to the 5 -OH nucleoside.
2. Add this solution to 30 g (100.0 mmol tosylate) of dry poly(4-vinylpyridinium-ptoluenesulfonate) (S.2) and shake with the shaker set at 150 rpm at room temperature under an argon atmosphere. Monitor the disappearance of the 5 -OH nucleoside unit (RT = 7.2 min) by HPLC (UNIT 10.5) using the following conditions: ◦
Column: Nucleosil C18, 5 µm, 100 A, 150 × 4.6 mm Flow rate: 1 mL/min Mobile phase A: acetonitrile Mobile phase B: 50 mM TEAA, pH 7.5 Linear gradient: 10% to 40% A in B over 5 min, then 40% to 80% A in B over 15 min.
Solution-Phase Synthesis Using PolymerSupported Reagents
For poly(4-vinylpyridinium-p-toluenesulfonate), Aldrich reports loading of the ptoluenesulfonate anion as determined by elemental analysis: N = 4.59%, S = 10.70%, corresponding to 3.34 mmol p-toluenesulfonate (tosyl) anion per g resin. Before use, the resin is dried over P2 O5 under vacuum (30 mbar) at 80◦ C for 10 hr and then stored in desiccator over P2 O5 under vacuum (10 mbar). The loading of each batch of resin must be checked by elemental analysis before use. The coupling reaction is performed under argon.
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Figure 3.14.1 Synthesis of dinucleotide 5 -OH-3 -O-Lev phosphotriesters and thionophosphotriesters using polymerbound reagents via the phosphoramidite approach. Symbols: B1 , B2 , bases; ABz , N 6 -benzoyladenin-9-yl; CBz , N 4 -benzoylcytosin-1-yl, GiBu , N 2 -isobutyrylguanin-9-yl, T, thymin-1-yl; CE, 2-cyanoethyl; DMTr, 4,4 -dimethoxytrityl; Lev, levulinyl; PS, polystyrene; PV, polyvinyl.
3.14.3 Current Protocols in Nucleic Acid Chemistry
Supplement 26
3. When the coupling is complete (after 1 hr to 3 hr), add 0.5 mL water and shake for an additional 15 min until the hydrolysis of any excess phosphoramidite is complete (monitored by HPLC). The phosphoramidite (S.3; RT = 22.1, 22.3 min) is hydrolyzed to the cyanoethyl Hphosphonate diester (S.6; RT = 15.0 min).
4. Filter the mixture under vacuum using a fritted-glass funnel and wash the resin three times, each time with 50 mL dichloromethane. 5. Concentrate the combined organic phase on a rotary evaporator to ∼100 mL. Proceed to the next reaction without further purification.
Perform oxidation reaction 6. Add 8.0 g (20.0 mmol IO4 – ) of Amberlyst A-26, periodate form (S.7), and shake at room temperature (generally for 2 hr) until there is complete conversion of the phosphite triester to the phosphate triester as monitored by 31 P NMR. Check the loading of the resin by elemental analysis before use. S.5: δ 140.4, 141.8 ppm; S.8: δ –1.9, –1.8 ppm. CAUTION: Special care must be exercised when using the polymer-supported periodate, which is a strong oxidizer presenting a danger of fire or explosion when in contact with other materials.
7. Filter the mixture using a fritted-glass funnel and wash the resin three times, each time with 50 mL dichloromethane.
Perform detritylation reaction 8. Add 107.0 mL of methanol to the filtrate and cool to 0◦ C in an ice bath. 9. Add 90 mL of 10% (w/v) p-toluenesulfonic acid in 7:3 (v/v) dichloromethane/ methanol prechilled to 0◦ C. Stir 30 min. 10. Perform TLC analysis (APPENDIX 3D) using 95:5 (v/v) dichloromethane/methanol as the solvent to confirm the completion of the reaction. Rf 0.48 for 5 -O-DMTr dimer and 0.30 for 5 -OH dimer.
11. Add 100 mL water and vigorously stir for 10 min at 0◦ C. During step 9, partial conversion of the levulinyl protection into its dimethylacetal derivative occurs. The reaction in this step restores the dimethylacetal to the levulinyl compound.
12. Add 100 mL saturated aqueous NaHCO3 solution and stir 10 min at 0◦ C. 13. Pour the solution into a 2-L separatory funnel and add 100 mL dichloromethane and 200 mL saturated aqueous NaHCO3 solution. Shake and isolate the organic layer. 14. Wash the organic layer with 300 mL of 0.2 M sodium bisulfite/0.5 M sodium chloride. Isolate the organic layer. 15. Dry the organic solution over 40 g of anhydrous Na2 SO4 , filter using a fritted-glass funnel, wash twice, each time with 25 mL dichloromethane, then concentrate the filtrate on a rotary evaporator. 16. Add 50 mL dichloromethane to the concentrated filtrate, then add the resulting solution dropwise to 1 L of vigorously stirring diethyl ether prechilled to 0◦ C. Solution-Phase Synthesis Using PolymerSupported Reagents
17. Filter the white precipitate using a fritted-glass funnel, then wash with 500 mL prechilled diethyl ether and dry in a vacuum desiccator over P2 O5 .
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18. Confirm the purity of the dimer by HPLC, (UNIT 10.1).
31 P
NMR, and MALDI-TOF-MS
5 -OH-dGiBu -dGiBu -3 -O-levulinyl cyanoethylphosphotriester (S.12): Yield of white amorphous solid, 7.5 g (76%). 31 P NMR: −1.82, −1.93 ppm. MALDI-TOF-MS (THAP matrix, + mode) m/z: exp. 888.3; calc. 888.3. RP-HPLC: purity at 260 nm, 95%; RT = 8.7 min.
SYNTHESIS OF 5 -OH-3 -O-LEVULINYL DINUCLEOSIDE CYANOETHYLTHIONOPHOSPHOTRIESTERS
ALTERNATE PROTOCOL 1
This protocol describes synthesis of thionophosphotriester-linked dinucleotides using the appropriate support-bound sulfurization reagent (S.10; Fig. 3.14.1). Synthesis of a 5 -OH, 3 -O-Lev dGiBu -T dinucleotide is presented as an example.
Additional Materials (also see Basic Protocol) Amberlyst A-26 ion-exchange resin, tetrathionate form (S.10; see Support Protocol 1) 2.5% (w/v) NaHCO3 2-L separatory funnel Perform coupling reaction 1. Perform coupling as in the Basic Protocol, steps 1 to 5, starting with 3.40 g (10.0 mmol) of 3 -O-levulinyl-thymidine and 12.40 g (15.0 mmol) of 5 -O-dimethoxytrityl-3 -O-[(2-cyanoethyl)-N, N-diisopropylphosphoramidite]N2 -isobutyryl-2 -deoxyguanosine. Perform sulfurization reaction 2. Add 26.6 g (50.0 mmol S4 O6 2– ) of Amberlyst A-26, tetrathionate form (S.10), and shake at room temperature (generally for 2 hr) until there is complete conversion of the phosphite triester to the thionophosphate triester as monitored by HPLC (see Basic Protocol, step 2) and 31 P NMR. S.5: 31 P NMR: δ 141.7, 143.5 ppm. RP-HPLC: RT = 16.0, 16.1 min. S.11: 31 P NMR: δ 68.2, 68.7 ppm. RP-HPLC: RT = 16.7, 17.0 min.
3. Filter the mixture using a fritted-glass funnel and wash the resin three times, each time with 50 mL dichloromethane.
Perform detritylation reaction 4. Carry out steps 8 to 12 of the Basic Protocol. Rf 0.55 for 5 -O-DMTr dimer and 0.37 for 5 -OH dimer.
5. Pour the solution into a 2-L separatory funnel and add 200 mL dichloromethane and 300 mL saturated aqueous NaHCO3 solution. Shake and isolate the organic layer. 6. Wash the organic layer three times, each time with 500 mL of 2.5% NaHCO3 . Extended washing is performed in order to remove the 5 -OH-3 -HP CE monomer.
7. Perform steps 15 to 18 of the Basic Protocol to complete the isolation. 5 -OH-dGiBu -T-3 -O-levulinyl cyanoethylthionophosphotriester (S.14): Yield of white amorphous solid, 7.37g (91%). 31 P NMR: 68.48 and 68.09 ppm. MALDI-TOF-MS (THAP matrix, + mode) m/z: exp. 809.7; calc. 809.8. RP-HPLC purity at 260 nm, 91%; RT = 9.7 min. Synthesis of Unmodified Oligonucleotides
3.14.5 Current Protocols in Nucleic Acid Chemistry
Supplement 26
ALTERNATE PROTOCOL 2
SYNTHESIS OF 3 -OH-5 -O-DIMETHOXYTRITYL DINUCLEOSIDE CYANOETHYLTHIONOPHOSPHOTRIESTERS This protocol describes the solution-phase synthesis of 3 -OH-5 -O-DMTr-N-protected dinucleotides with a thionophosphotriester linkage by means of phosphoramidite method according to the scheme shown in Figure 3.14.2. This protocol is also suitable for the synthesis of trinucleotides with thionophosphotriester linkages. Synthesis of a 3 -OH, 5 -O-DMTr dGiBu -dCBz dinucleotide is presented as an example.
Solution-Phase Synthesis Using PolymerSupported Reagents
Figure 3.14.2 Synthesis of dinucleotide 5 -O-DMTr-3 -OH thionophosphotriesters using polymerbound reagents via the phosphoramidite approach. Symbols: B1 , B2 , bases; ABz , N 6 benzoyladenin-9-yl; CBz , N 4 -benzoylcytosin-1-yl; GiBu , N 2 -isobutyrylguanin-9-yl; T, thymin-1-yl; CE, 2-cyanoethyl; DMTr, 4,4 -dimethoxytrityl; Lev, levulinyl; PS, polystyrene; PV, polyvinyl.
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Additional Materials (also see Basic Protocol) Amberlyst A-26 ion-exchange resin, tetrathionate form (S.10; see Support Protocol 1) Pyridine Silica gel Acetic acid Amberlyst A-15 ion-exchange resin, hydrazinium form (S.16; see Support Protocol 1) Fritted-glass column, 5 cm diameter, 60 cm height Perform coupling reaction 1. Perform coupling as in the Basic Protocol, steps 1 to 5, starting with 4.36 g (10.0 mmol) 3 -O-levulinyl-N2 -isobutyryl-2 -deoxyguanosine and 14.5 g (17.4 mmol) 5 -O-dimethoxytrityl-3 -[(2-cyanoethyl)-N, N -diisopropylphosphoramidite]N4 -benzoyl-2 -deoxycytidine. Perform sulfurization reaction 2. Add 10 mL of pyridine and 26.6 g (50.0 mmol, S4 O6 2− ) of Amberlyst A-26, tetrathionate form (S.10). Shake at room temperature (generally for 2 to 6 hr) until there is complete conversion of the phosphite triester to the thionophosphate triester as monitored by HPLC (see Basic Protocol, step 2) and 31 P NMR. S.5: 31 P NMR: δ 136.6 ppm. RP-HPLC: RT = 18.9 min. S.11: 31 P NMR: δ 64.7, 66.0 ppm. RP-HPLC: RT = 19.9 min. Pyridine is added to avoid partial detritylation
3. Filter the mixture using a fritted-glass funnel and wash the resin three times, each time with 50 mL of dichloromethane. 4. Evaporate the combined organic layers to an oil on a rotary evaporator. 5. Dissolve the oil in 44 mL of dichloromethane, add 20 g (50.0 mmol IO4 – ) of Amberlyst A-26, periodate form (S.7), and shake for 2 hr. 6. Filter the mixture using a fritted-glass funnel, then wash the resin twice, each time with 200 mL dichloromethane. 7. In a 2-L separatory funnel, wash the organic layer with 300 mL of 0.2 M sodium bisulfite/0.5 M sodium chloride. Isolate the organic layer. 8. Dry the organic solution over 40 g of anhydrous Na2 SO4 , vacuum filter using a fritted-glass funnel, and concentrate to an oil on a rotary evaporator. 9. Dissolve the oil in 60 mL dichloromethane and pour into a fritted-glass column (5 cm diameter, 60 cm height) containing 60 g silica gel. 10. Wash with 200 mL of 97:3 (v/v) dichloromethane/methanol. 11. Evaporate on a rotary evaporator. 12. Perform steps 16 to 18 of the Basic Protocol to complete the isolation. 3 -O-Levulinyl-dGiBu -dCBz -5 -O-DMTr cyanoethylthionophosphotriester (S.11): Yield of white amorphous solid, 10.32 g (861%).31 P NMR: 68.4 and 68.7 ppm. MALDI-TOF-MS (THAP matrix, + mode) m/z: exp. 1200.25; calc. 1201.24. RP-HPLC purity at 260 nm, 96%; RT = 19.0 min.
Perform delevulinylation reaction 13. Dissolve 3.5 g (2.9 mmol) of dimer S.11 in a solution consisting of 69 mL pyridine and 17 mL of acetic acid.
Synthesis of Unmodified Oligonucleotides
3.14.7 Current Protocols in Nucleic Acid Chemistry
Supplement 26
14. Add 11.8 g (43.7 mmol N2 H5 + ) of Amberlyst A-15, hydrazinium form (S.16), and shake at 150 rpm for 4 hr at room temperature. Monitor by TLC (APPENDIX 3D) using 95:5 (v/v) CH2 Cl2 /MeOH as the solvent. Rf 0.65 for 3 -O-Lev dimer and 0.51 for 3 -OH dimer.
15. Filter the resin using a fritted-glass funnel, and wash with 300 mL of dichloromethane. 16. Add 250 mL of saturated aqueous NaHCO3 to the filtrate and gently stir. 17. Pour the solution into a 2-L separatory funnel and shake to separate the organic from the aqueous layer. Retain both layers. 18. Extract the aqueous layer three times, each time with 150 mL dichloromethane, and pool all organic layers. 19. Add anhydrous Na2 SO4 to the combined organic phase, filter using a fritted-glass funnel, and concentrate on a rotary evaporator. 20. Add 30 mL of dichloromethane to the concentrated filtrate, then add the resulting solution dropwise with vigorous magnetic stirring to 300 mL of prechilled (0◦ C) diethyl ether. 21. Filter the white precipitate using a fritted-glass funnel, then wash it with 200 mL prechilled diethyl ether and dry in a vacuum desiccator over P2 O5 . 22. Confirm the purity of the dimer by HPLC, (UNIT 10.1).
31 P
NMR, and MALDI-TOF-MS
3 -OH-dGiBu -dCBz -5 -O-DMTr cyanoethylthionophosphotriester (S.17): Yield of white amorphous solid, 2.9 g (90%). 31 P NMR: 68.3 and 68.6 ppm. MALDI-TOF-MS (THAP matrix, + mode) m/z: exp. 1102.5; calc. 1103.1.RP-HPLC purity at 260 nm, 95%; RT = 17.9, 18.3 min. ALTERNATE PROTOCOL 3
ALTERNATE COUPLING PROTOCOL WHEN USING 3 -O-LEVULINYLN4 -BENZOYL 2 -DEOXYCYTIDINE Since 3 -O-levulinyl-N4 -benzoyl 2 -deoxycytidine is not soluble in dichloromethane, DMF is used as cosolvent. However the authors have noticed that traces of water trapped in the polyvinyl pyridinium resin is released due to an enhanced swelling of the resin in DMF. Hence, a preliminary drying step is needed, which is performed by means of molecular sieves shaken overnight with this mixture. All the other steps are the same as those described in the Basic Protocol.
Additional Materials (also see Basic Protocol) Dimethylformamide (DMF), anhydrous 4A molecular sieves 1. Dissolve 4.3 g (10.0 mmol) of 3 -O-levulinyl-N4 -benzoyl-2 -deoxycytidine in 60 mL hot (∼40◦ C) 1:1 (v/v) DMF/dichloromethane. 2. Add 10 g of 4A molecular sieves and 30 g (100.0 mmol tosylate) of poly(4vinylpyridinium-p-toluenesulfonate) (S.2), and shake overnight at 150 rpm, room temperature. Solution-Phase Synthesis Using PolymerSupported Reagents
3. Add 15 mmol of 5 -O-dimethoxytrityl-3 -O-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite] nucleoside in 45 mL of dichloromethane and shake at room temperature ∼2 hr.
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Current Protocols in Nucleic Acid Chemistry
4. Perform HPLC analysis (see Basic Protocol, step 2) to follow the disappearance of 3 -O-levulinyl-N4 -benzoyl-2 -deoxycytidine (RT = 8.6 min). 5. Continue with steps 3 to 5 of the Basic Protocol.
PREPARATION OF POLYMER-SUPPORTED REAGENTS Materials
SUPPORT PROTOCOL 1
Amberlyst A-26 ion-exchange resin, hydroxide (OH– ) form (Aldrich) Methanol Dichloromethane Sodium periodate P2 O5 Potassium tetrathionate (Aldrich) Amberlyst A-15 ion-exchange resin, H+ form (Aldrich) 2 M HCl Hydrazine monohydrate Absolute ethanol Fritted-glass filter funnel 4-L Erlenmeyer flask Shaker Vacuum desiccator To prepare oxidizing resin (periodate form) CAUTION: Special care must be exercised when preparing or using the polymersupported periodate, which is strong oxidizer presenting a danger of fire or explosion when in contact with other materials. 1a. On a fritted glass funnel, wash 10.0 g Amberlyst A-26 (OH– form) twice each with 25 mL methanol and then 25 mL dichloromethane. 2a. Pour the resin into a 4-L Erlenmeyer flask and add 10.0 g (43 mmol) sodium periodate in 200 mL deionized water. Shake 15 hr at 150 rpm. 3a. Filter using a fritted-glass funnel. Wash the resin with 1000 mL water once, followed by 30 mL of methanol twice, then 20 mL of dichloromethane twice. 4a. Dry in a desiccator under reduced pressure over P2 O5 . 5a. Determine the loading in periodate ion by elemental analysis. Store resin up to several years at room temperature in closed vials. Yield S.7: 8.5 g of dried resin. N = 3.40%, I = 27.16%, corresponding to 2.14 mmol of periodate ion per g of resin. Note that a similar oxidizing resin is available from Novabiochem.
To prepare sulfurizing resin (tetrathionate form) 1b. On a fritted glass funnel, wash 10.0 g Amberlyst A-26 (OH– form) twice each with 25 mL methanol and then 25 mL dichloromethane. 2b. Pour the resin into a 4-L Erlenmeyer flask and 30.35 g (100 mmol) potassium tetrathionate in 200 mL deionized water. Shake 20 hr at 150 rpm. 3b. Filter using a fritted-glass funnel. Wash the resin with 4000 mL of water once, followed by 100 mL of methanol twice, then 100 mL of dichloromethane twice.
Synthesis of Unmodified Oligonucleotides
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4b. Dry in a desiccator under reduced pressure over P2 O5 . 5b. Determine the loading in tetrathionate ion by elemental analysis. Store resin up to several years at room temperature in closed vials. Yield S.10: 8.5 g of dried resin. N = 4.24%, S = 23.25%, K+ <100 ppm, corresponding to 1.81 mmol of tetrathionate ion per g of resin.
To prepare delevulinylating resin (hydrazinium form) 1c. On a fritted glass funnel, wash 200.0 g Amberlyst A-15 (H+ form) twice each with 1000 mL water, twice each with 500 mL of 2 M HCl, then twelve times (or as necessary to bring the filtrate to pH 6) with 500 mL water. 2c. Pour the resin into a 4-L Erlenmeyer flask and add 2.3 L of a solution of 200 mL (4.12 mol) hydrazine monohydrate in 2.1 L deionized water. Shake 2.5 hr at 150 rpm. 3c. Filter using a fritted-glass funnel. Wash resin twelve times with 500 mL water, then three times with 500 mL ethanol, and finally two times with 500 mL dichloromethane. 4c. Dry in a desiccator under reduced pressure over P2 O5 . 5c. Determine the loading in hydrazinium ion by elemental analysis. Store resin up to several years at room temperature in closed vials. Yield S.16: 232 g of dried resin. N = 11.14%, S = 12.57%, corresponding to 3.98 mmol hydrazinium ion per g of resin. SUPPORT PROTOCOL 2
REGENERATION OF COUPLING RESIN Materials Used coupling resin S.2 [poly(4-vinylpyridinium-p-toluenesulfonate) resin] Acetonitrile Dichloromethane Methanol p-Toluenesulfonic acid P2 O5 2-L Erlenmeyer flask Fritted-glass funnel Platform shaker Vacuum desiccator 1. Pour 50 g of used S.2 in a 2-L Erlenmeyer flask with 500 mL of acetonitrile and shake 1 hr at 150 rpm on a platform shaker. 2. Filter using a fritted-glass funnel. 3. Pour the resin into a 2-L Erlenmeyer flask with 500 mL of dichloromethane and shake 1 hr at 150 rpm. 4. Filter using a fritted-glass funnel. 5. Pour the resin in a 2-L Erlenmeyer flask with 500 mL of methanol and shake 1 hr at 150 rpm. 6. Filter using a fritted-glass funnel.
Solution-Phase Synthesis Using PolymerSupported Reagents
7. Dissolve 200 g of p-toluenesulfonic acid in 200 mL of warm methanol (40◦ C) in a 2-L Erlenmeyer flask and add the resin. Shake 16 hr at 150 rpm on a platform rotator.
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8. Filter using a fritted-glass funnel and wash with 600 mL of methanol and 800 mL of dichloromethane. 9. Dry the resin under vacuum (∼30 mbar) over P2 O5 at 80◦ C for 10 hr, changing the hydrolyzed P2 O5 frequently.
COMMENTARY Background Information The studies that are reported here were originally driven to develop a convenient method for scaling up the synthesis of di- and trinucleosides with oxo- and thionophosphotriester internucleoside linkages without chromatographic purification. Synthesis of oligonucleotides is generally performed on a solid support according to the phosphoramidite approach (UNIT 3.3). Although very efficient, the cost of solid supports (Sanghvi et al., 2000) explains the quest for alternative synthetic methodologies, such as the HELP method described by Bonora (UNIT 4.27), which uses an MPEG soluble support, and the PASS methods (Mihaichuk et al., 2000), in which the growing oligonucleotide reacts with a phosphoramidite unit temporarily immobilized on a solid support. Compared to solid-phase synthesis, the scale-up of solution-phase synthesis is more conventional and economical, but involves purification and isolation procedures after each reaction step. A new approach where polymerbound reagents promote reactions, also called inverse solid-phase synthesis in combinatorial chemistry (Sherrington, 2001), prevails over these limitations, as the reagents are separated from the reaction mixtures by simple filtration (Ley et al., 2000). There are only a few examples of solutionphase synthesis of oligonucleotides involving solid-phase reagents. For instance, sulfonic acid resins were employed as substitutes for acetic acid/water mixtures to induce detritylation of 5 -O-DMTr phosphodiester oligonucleotides (Iyer et al., 1995). Polystyrenesulfonyl chloride was used as the coupling agent to create phosphodiester internucleoside linkages by the obsolete phosphodiester approach (Rubinstein and Patchornik, 1972, 1975). More recently, a polymer-supported xanthane hydride was used as a sulfur-transfer reagent for the solution-phase synthesis of phosphorothioate nucleotides (Zhang et al., 2002). It could be pointed out that the use of this reagent was not reported for oligonucleotide synthesis.
Coupling step Tetrazole, tetrazole salts, and heterocyclic analogs are commonly used as activators of the coupling reaction of 5 -OH nucleosides with nucleoside 3 -O-phosphoramidites (UNIT 3.3). Tetrazole, the most frequently employed activator, needs to be of high purity. Because of its predisposition to explosion (Eleuteri et al., 2000), especially during sublimation, tetrazole is no longer sold as a solid powder. Pyridinium acidic salts were occasionally used as safer and cheaper activators than tetrazole (Beier and Pfleiderer, 1999; Eleuteri et al., 2000). Their solid-supported analogs are readily accessible from commercial resins by simple ion-exchange washings. The pyridinium moiety could be ionically bound to polystyrene sulfonic acids such as Dowex 50W or Amberlyst 15 to give S.1, or could be covalently linked to the resin, as in polyvinylpyridinium tosylate (S.2). This last reagent was formerly used for the preparation of nucleoside 3 -Ophosphoramidites (Sanghvi et al., 2000). However, in the authors’ experience, this reagent has produced poor results with low conversion yields and extensive degradation. This unfavorable reactivity was explained by the low availability of the pyridinium moieties on the resin, but the residual water contained in the resin could explain, at least partially, the hydrolysis of the formed phosphoramidite in acidic medium. The coupling reactions between an excess of nucleoside 3 -phosphoramidite (S.3) and a 5 -OH nucleoside (S.4) are generally performed in dichloromethane to ensure adequate swelling of the polystyrene polymer. Reactions performed with insoluble 3 -Olevulinyl 5 -OH-N4 -benzoyl-2 -deoxycytosine require DMF as cosolvent. The reactions are followed by 31 P NMR and by reversedphase HPLC. 31 P NMR is used to follow the conversion of the phosphoramidites (δ 148 to 150 ppm) to dinucleotide 3 -5 phosphite triesters (S.5; δ 140 ppm), and HPLC is used to follow the disappearance of the 5 -OH nucleoside. Resins (S.1) gave unpredictable Synthesis of Unmodified Oligonucleotides
3.14.11 Current Protocols in Nucleic Acid Chemistry
Supplement 26
results in regard to the reaction times, while reliable results are observed with 10 molar equivalents of polymer (S.2). After 1 to 2 hr of coupling, the 5 -OH is totally consumed. Besides the signal of the phosphite triester, the 31 P NMR analysis of the medium shows the presence of an excess of starting phosphoramidites and of their cyanoethyl Hphosphonate diester analogs (S.6; δ 8 ppm) resulting from their partial hydrolysis by water held in the resin. Attempts to remove this water by washing, azeotropic coevaporation, or drying under high vacuum were ineffective. When DMF/dichloromethane is used (for 3 -O-levulinyl-5 -OH-N4 -benzoyl 2 deoxycytosine), extensive hydrolysis of the phosphoramidites is observed due to an additional release of trapped water from the resin. This results from a better swelling of the polymer in DMF compared to dichloromethane. To avoid this extensive hydrolysis, molecular sieves are added to the reaction mixtures before the addition of the coupling agent S.2. When the coupling is done, the excess phosphoramidite is totally converted to Hphosphonate (S.6) by addition of water in order to help the purification at the end of the overall process. The phosphite triesters (S.5) are not sensitive to this addition of water. At the end of the coupling, the resin is removed by filtration and the resulting mixtures are used directly for oxidation or sulfurization.
Solution-Phase Synthesis Using PolymerSupported Reagents
Oxidation or sulfurization step Oxidation and sulfurization of dinucleotide phosphite triesters give dinucleotide phosphotriesters and thionophosphotriesters, respectively. In solid-phase DNA synthesis, I2 /H2 O is typically used for oxidation, but other nonaqueous reagents such as tertbutylhydroperoxide are also employed successfully (for other examples, see UNIT 3.3). It was reported that the periodate anion bound to an anion-exchange resin is able to oxidize triphenylphosphane in triphenylphosphane oxide (Harrison and Hodge, 1982). This polymer-bound reagent could be used in a variety of solvents (Drewry et al., 1999), increasing the limited efficiency of periodate that results from its insolubility in nonpolar solvents. The oxidation of dinucleoside phosphite triesters (S.5) is carried out with polystyrene Amberlyst A-26, periodate form (S.7), in 2 hr, yielding dinucleoside phosphotriesters (S.8; δ −2 ppm). During the course of this reaction, the contaminating nucleoside H-phosphonate diesters (S.6) are oxidized to nucleoside cya-
noethyl phosphodiesters (S.9; δ 0 ppm). The resin is then filtered off. The ability of oligonucleotide phosphorothioates to act as inhibitors of gene expression is certainly the reason why many sulfur-transfer reagents have been employed in solid-phase oligonucleotide chemistry (UNIT 3.3) From these, the most extensively used is the commercially available 3-H-1,2-bendithiol-3-one 1,1-dioxide, referred to as Beaucage’s reagent (Iyer et al., 1990). There are far fewer examples of supported sulfur-transfer reagents reported for an inverse solid-phase synthesis. The 3-amino1,2,4-dithiazole-5-thione or xanthane hydride reported as a sulfur-transfer reagent for solid-phase synthesis of oligonucleotide phosphorothioates (Tang et al., 2000) was covalently linked to a methylacrylate-ethylene glycol copolymer to sulfurize nucleoside phosphite triesters (Zhang et al., 2002). Although efficient, this reagent is not regenerable. Consequently, its synthesis is costly and time-consuming compared to an ionically polymer-bound reagent. Attaching the tetrathionate ion to anionexchange polymer resins is an alternative to a covalently bound reagent. While sluggish, pyridinium tetrathionate was used as sulfurizing agent in oligonucleotide chemistry (Efimov et al., 1995). Because of its anionic character, the tetrathionate ion is easily bound to anion-exchange polymer resins through salt bridges with the pyridinium moieties of poly4-vinyl pyridine or with the quaternary ammonium cations of the polystyrenic Amberlyst A-26. This is performed by simple stirring of the resins in an aqueous solution of potassium tetrathionate. However, the ionic bond between tetrathionate and pyridinium ions is not sufficiently stable to consider the use of the polyvinyl pyridium resin. The loading of Amberlyst A-26 tetrathionate (S.10) determined by elemental analysis shows two ammonium cations on the resin bound to the dianion. The sulfurization of phosphite triesters is followed by 31 P NMR. In general, 5 molar equivalents of S.10 are able to convert dinucleotide phosphite triesters (S.5; δ 140 ppm) to dinucleotide thionophosphotriesters (S.11; δ 68 ppm) in 2 to 3 hr. The resin is then removed by filtration. Elemental analysis of the trithionate ion resulting from the reduction of the tetrathionate is bound to the resin. In contrast to the supported-periodate oxidation of phosphite triesters where the H-phosphonate diester side products (S.6) are oxidized to phosphodiesters (S.9), the
3.14.12 Supplement 26
Current Protocols in Nucleic Acid Chemistry
tetrathionate reaction is not able to sulfurize S.6. For the synthesis of 3 -OH dinucleotides, the sulfurization is followed with the oxidation of S.6 into the polar S.9 by the periodate resin S.7. This additional treatment does not alter the thionophosphotriester linkage of dinucleotide S.11. At this stage, nucleoside phosphodiesters (S.9) are removed from the reaction mixtures by filtration on a small pad of silica gel where they remain adsorbed. Detritylation or delevulinylation step As mentioned above, strong acid cationexchange resins have been employed for the detritylation of 5 -O-DMTr- oligonucleotide phosphodiesters in water (Iyer et al., 1995). The use of these resins in organic solvents has also been described for nucleoside chemistry (Patil et al., 1994). Their reactivity is considerably dependant on the nature of the polymers. Two main advantages have been pointed out for the use of these reagents in oligonucleotide chemistry. The trityl cation released during the acidic treatment is captured by the resin so that the oligonucleotide is obtained pure after filtration, and depurination is not observed during the course of the reaction. However, in the studies illustrated here, a large variety of sulfonic acid resins were tested, and extensive degradation of the dinucleotides was observed. Consequently, these solid-supported acids are not being investigated further. The crude mixtures obtained from oxidation or sulfurization are treated with 2% p-toluenesulfonic acid in a 7:3 (v/v) dichloromethane/methanol mixture at 0◦ C for 30 min to 1 hr. The course of the reactions is conventionally followed by TLC analysis. The detritylation of crude mixtures obtained after an oxidation step generates dinucleotide 5 -OH cyanoethyl phosphotriesters (S.12) contaminated with nucleoside 5 -OH3 -cyanoethyl phosphodiesters (S.13). The crude products from the sulfurization give dinucleotide 5 -OH thionophosphotriesters (S.14) with concomitant formation of nucleoside 5 OH-3 -H-phosphonate diesters (S.15). Standard aqueous NaHCO3 workup allows the removal of nucleotide contaminants (S.13 or S.15), which stay in solution in the aqueous phase. Finally, precipitation of the desired dinucleotides in diethyl ether allows the removal of the trityl residues. The results have been consistently good, as judged from the HPLC, 31 P NMR, and MALDI-TOF-MS analysis of the dinucleotides.
The 3 -O-levulinyl protection is generally removed with a solution of hydrazine in acetic acid/pyridine (van Boom and Burgers, 1976). In this approach, a polymer-supported hydrazine (S.16) is used to perform the deprotection. This reagent is prepared by washing the strong acid cation exchanger Amberlyst A-15 with an aqueous solution of hydrazine. Delevulinylation of 3 -O-Lev-5 -O-DMTr dinucleotides is directly performed from crude products containing oxo- or thionophosphotriesters (S.8 and S.11), respectively. The delevulinylation is complete in 3 hr and gives rise to dinucleotide 5 -O-DMTr-3 -OH cyanoethyl thionophosphotriesters (S.17). Standard aqueous NaHCO3 workup allows the removal of nucleotide (S.9) from the desired dinucleotides.
Critical Parameters and Troubleshooting The preparation of the polymer-supported reagents (Support Protocol 1) and the synthesis of the di- and trinucleotides (Basic and Alternate Protocols) are relatively simple and efficient. However, careful attention to details of basic organic procedures is required. Experience with chemical laboratory techniques such as extraction, filtration, evaporation, TLC, and HPLC is needed. Characterization of the products requires knowledge of elemental analysis, 31 P NMR, and MALDI-TOF mass spectrometry. General laboratory safety is of primary concern when hazardous compounds and solvents are involved. Strict anhydrous conditions are required for the coupling step, but not for the other steps involved in the process. Special attention must be paid to the shaking of the heterogeneous reaction in order to keep reaction times short. Stirring of the solution-phase reaction medium with polymer-supported reagents is best performed on a shaker at 150 rpm. Due to the low solubility of 3 -O-levulinyl4 N -benzoyl-2 -deoxycytidine in dichloromethane, coupling steps using this monomer are performed in DMF/dichloromethane (Alternate Protocol 3). Due to the presence of methanol during the detritylation, some dimethylacetalization of the keto group of the levulinyl protection is observed, with a higher retention time in HPLC and a higher mass (+46) in MALDITOF-MS. The addition of water at the end of the reaction (Basic Protocol, step 11) allows
Synthesis of Unmodified Oligonucleotides
3.14.13 Current Protocols in Nucleic Acid Chemistry
Supplement 26
conversion of this side product into the levulinyl derivative.
Anticipated Results The protocols described here for dimers can be successfully applied for the synthesis of trimers. Using these methods, di- and trinucleotides are obtained with good yields (76% to 96%) and high purities (90% to 96%) without chromatography. Synthesis of longer oligonucleotides such as hexamers is also possible, but increasing the length of the oligonucleotide decreases its solubility in dichloromethane and requires the addition of DMF to dissolve the intermediates during the coupling step, as well as addition of pyridine during the aqueous work-up after the detritylation step (Basic Protocol, step 12). Due to low yields and purities, the authors do not recommend use of the Basic Protocol for long oligonucleotides. However, the di- and trinucleotides can be converted into 3 -phosphoramidite or 3 -H-phosphonate monoester building blocks and used for solidphase synthesis on a DNA synthesizer. Since reagents are ionically attached to the polymers, the resins are easily regenerable (see Support Protocol 2). Therefore, their cost and their impact on the environment are moderate.
Time Considerations The synthesis of a dinucleotide using polymer-supported reagents can be accomplished in 1 to 2 days. The time for coupling comprises 1 to 3 hr, depending on the loading of the supported tosylate, and requires additional time for filtration, washing, and concentration. The time needed for oxidation is usually shorter (1 to 2 hr) than for sulfurization (2 to 6 hr). Use of 31 P NMR and HPLC to monitor the reactions could increase the time required for the overall process. Detritylation is usually fast (10 to 30 min), but work-up of the reaction, with extraction and precipitation, requires more time (∼2 hr depending on the scale). Delevulinylation requires about 4 hr, plus additional time for work-up and isolation (∼2 hr).
Literature Cited
Solution-Phase Synthesis Using PolymerSupported Reagents
Beier, M. and Pfleiderer, W. 1999. Pyridinium salts: An effective class of catalysts for oligonucleotide synthesis. Helv. Chim. Acta 82:879887. Drewry, D.H., Coe, D.M., and Poon, S. 1999. Solidsupported reagents in organic synthesis. Med. Res. Rev. 19:97-148.
Dueymes, C., Sch¨onberger, A., Adamo, I., Navarro, A.E., Meyer, A., Lange, M., Imbach, J.L., Link, F., Morvan, F., and Vasseur, J.J. 2005. Highyield solution-phase synthesis of di- and trinucleotide blocks assisted by polymer-supported reagents. Org. Lett. 7:3485-3488. Efimov, V.A., Kalinkina, A.L., Chakhmakhcheva, O.G., Hill, T.S., and Jayaraman, K. 1995. New efficient sulfurizing reagents for the preparation of oligodeoxyribonucleotides phosphorothioate analogs. Nucl. Acids Res. 23:4029-4033. Eleuteri, A., Capaldi, D.C., Krotz, A.H., Cole, D.L., and Ravikumar, V.T. 2000. Pyridinium trifluoroacetate/N-methylimidazole as an efficient activator for oligonucleotide synthesis via the phosphoramidite method. Org. Proc. Res. Dev. 4:182-189. Harrison, C.R. and Hodge, P. 1982. Polymersupported periodate and iodate as oxidizing agents. J. Chem. Soc. Perkin Trans. 1 509-511. Iyer, R.P., Phillips, L.R., Egan, W., Regan, J.B., and Beaucage, S.L. 1990. The automated synthesis of sulfur-containing oligodeoxyribonucleotides using 3H-1,2-benzodithiol-3-one 1,1-dioxide as a sulfur-transfer reagent. J. Org. Chem. 55:46934699. Iyer, R.P., Jiang, Z., Yu, D., Tan, W., and Agrawal, S. 1995. Improved procedure for the detritylation of DMT-oligonucleotides: Use of Dowex. Synth. Comm. 25:3611-3623. Ley, S.V., Baxendale, I.R., Bream, R.N., Jackson, P.S., Leach, A.G., Longbottom, D.A., Nesi, M., Scott, J.S., and Taylor, S.J. 2000. Muti-step organic synthesis using solid-supported reagents as scavengers: A new paradigm in chemical library generation. J. Chem. Soc. Perkin Trans. 1 3815-4195. Mihaichuk, J.C., Hurley, T.B., Vagle, K.E., Smith, R.S., Yegge, J.A., Pratt, G.M., Tompkins, C.J., Sebesta, D.P., and Pieken, W.A. 2000. The dimethoxytrityl resin product anchored sequential synthesis method (DMT PASS): A conceptually novel approach to oligonucleotide synthesis. Org. Proc. Res. Dev. 4:214224. Patil, S.V., Mane, R.B., and Salunkhe, M.M. 1994. A facile method for detritylation of 5 -Odimethoxytrityl-3 -O-tert-butyldimethylsilyl2 -deoxynucleosides. Synth. Comm. 24:24232428. Reese, C.B., Song, Q.L., Rao, M.V., and Beckett, I. 1998. Preparation of an octadeoxyribonucleoside heptaphosphorothioate by the phosphotriester approach in solution. Nucleosides Nucleotides Nucl. Acids 17:451-470. Rubinstein, M. and Patchornik, A. 1972. Polymers as chemical reagents: The use of poly(3,5 diethylstyrene)-sulfonyl chloride for the synthesis of internucleotide bonds. Tetrahedron Lett. 13:2881-2884. Rubinstein, M. and Patchornik, A. 1975. Poly(3,5diethylstyrene) sulfonyl chloride: A new reagent for internucleotide bond synthesis. Tetrahedron 31:1517-1519.
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Sanghvi, Y.S., Guo, Z., Pfundheller, H.M., and Converso, A. 2000. Improved process for the preparation of nucleosidic phosphoramidites using a safer and cheaper activator. Org. Proc. Res. Dev. 4:175-181. Sherrington, D.C. 2001. Polymer-supported reagents, catalysts, and sorbents: Evolution and exploitation–A personalized view. J. Polym. Sci.: Polym. Chem. 39:2364-2377. Tang, J.Y., Han, Y.X., Tang, J.X., and Zhang, Z.D. 2000. Large-scale synthesis of oligonucleotide phosphorothioates using 3-amino1,2,4-dithiazole-5-thione as an efficient sulfur-transfer reagent. Org. Proc. Res. Dev. 4:194-198. van Boom, J.H. and Burgers, P.M.J. 1976. Use of levulinic acid in the protection of oligonucleotides via the modified phosphotriester method: Synthesis of decaribonucleotide U-A-U-A-U-A-UA-A-A. Tetrahedron Lett. 52:4875-7878. Zhang, Z., Han, Y., Tang, J.X., and Tang, J.Y. 2002. A novel polymer-supported sulfurtransfer reagent for the synthesis of phosphorothioates. Tetrahedron Lett. 43:4347-4349.
Contributed by Franc¸ois Morvan and Jean-Jacques Vasseur Universit´e Montpellier II Montpellier, France
Synthesis of Unmodified Oligonucleotides
3.14.15 Current Protocols in Nucleic Acid Chemistry
Supplement 26
DNA Synthesis Without Base Protection Using the Phosphoramidite Approach
UNIT 3.15
The strategy for synthesis of oligodeoxynucleotides without the protection of base moieties has proven useful for the synthesis of oligodeoxynucleotide derivatives having base-labile functional groups (Wada et al., 2001; Kobori et al., 2002; Sekine et al., 2003; Ohkubo et al., 2004, 2005a). Elimination of the base-deprotecting step makes it possible for oligodeoxynucleotides on chips or microarrays to be prepared more rapidly. Recently, the high-throughput synthesis of DNA probes on glass plates has become more important (Shena, 1999; Gao et al., 2004). Several methods categorized as base-unprotected approaches are reviewed in UNIT 3.10. In the protocols that follow, detailed experimental procedures required for DNA synthesis without base protection are described. Basic Protocols 1 to 3 describe the synthesis of the 5 -O-(4,4 -dimethoxytrityl) 3 -phosphoramidites of dC, dA, and dG from the starting unprotected 2 -deoxynucleosides. Alternate Protocols 1 to 6 describe synthesis of the same phosphoramidites by removal of N-protecting groups from commercially available phosphoramidites. The typical procedure for the synthesis of oligodeoxyribonucleotides from N-unprotected phosphoramidites involves the use of benzimidazolium triflate in the presence of trifluoromethyl-1-hydroxylbenztriazole. The key details for automated oligodeoxyribonucleotide synthesis are described in Basic Protocol 4.
SYNTHESIS OF 5 -O-(4,4 -DIMETHOXYTRITYL)-2 -DEOXYCYTIDINE 3 -(2-CYANOETHYL-N,N-DIISOPROPYLPHOSPHORAMIDITE) FROM 2 -DEOXYCYTIDINE
BASIC PROTOCOL 1
The N-unprotected dC phosphoramidite is prepared by direct tritylation of deoxycytidine followed by phosphitylation, as illustrated in Figure 3.15.1.
Materials 2 -Deoxycytidine hydrochloride (S.1a; Yamasa Shoyu) ◦ Pyridine (anhydrous, dried over 4-A molecular sieves, distilled from CaH2 ) Argon source Triethylamine (Wako, distilled from CaH2 ) Dichloroacetic acid (Wako) 4,4 -Dimethoxytrityl chloride (DMTr-Cl) Chloroform (CHCl3 ) ◦ Methanol (MeOH; Wako, anhydrous, dried over 4-A molecular sieves) Brine (saturated aqueous NaCl) Sodium sulfate (Na2 SO4 , anhydrous) Hexane Silica gel (Wakogel C-200, 75 to 150 µm) ◦ Toluene (anhydrous, dried over 4-A molecular sieves) ◦ Dichloromethane (CH2 Cl2 , anhydrous, dried over 4-A molecular sieves) ◦ Tetrahydrofuran (THF, anhydrous, dried over 4-A molecular sieves, no stabilizer) Diisopropylethylamine (DIPEA, anhydrous, dried over CaH2 , distilled from CaH2 ) 2-Cyanoethoxy-(N, N-diisopropylamino)chlorophosphine (synthesized as described in Nagai et al., 1989; Tanimura et al., 1989) 200-mL round-bottom flasks with argon-filled balloon Rotary evaporator and vacuum pump Synthesis of Unmodified Oligonucleotides Contributed by Akihiro Ohkubo, Kohji Seio, and Mitsuo Sekine Current Protocols in Nucleic Acid Chemistry (2006) 3.15.1-3.15.22 C 2006 by John Wiley & Sons, Inc. Copyright
3.15.1 Supplement 26
Figure 3.15.1
Synthesis of N-unprotected 3 -phosphoramidites by direct tritylation and phosphitylation.
Thin-layer chromatography (TLC) silica plates (Kieselgel 60 F254 ; 0.2 mm thick; Merck) 500-mL separatory funnel No. 2 filter paper and glass funnel 5 × 6– and 5 × 7.5–cm chromatography columns Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare 5 -O-DMTr-2 -dC 1. Suspend 1.30 g (5 mmol) 2 -deoxycytidine hydrochloride (S.1a) in 3 mL pyridine in a 200-mL round-bottom flask and concentrate the suspension to dryness using a rotary evaporator and vacuum pump. Repeat this azeotropic drying process three times, each time with 3 mL pyridine. 2. Add 50 mL pyridine and a stir bar to the flask and attach a balloon filled with argon gas to create an argon atmosphere. 3. While stirring, add 699 µL (5 mmol) triethylamine and then slowly add 412 µL (5 mmol) dichloroacetic acid. 4. While stirring, add 1.86 g (5.5 mmol) DMTr-Cl. 5. Stir the mixture for 4 hr at room temperature and check the reaction by TLC (APPENDIX 3D) using 6:1 (v/v) CHCl3 /MeOH (Rf = 0.59). 6. Add 10 mL MeOH to quench the DMTr-Cl and stir 10 min. 7. Dissolve the mixture in 200 mL CHCl3 and wash with 200 mL brine in a 500-mL separatory funnel. 8. Dry the organic layer over anhydrous sodium sulfate, filter through no. 2 filter paper in a glass funnel, and evaporate using a rotary evaporator and vacuum pump. 9. Dissolve the crude product in a minimal amount of 7:3 (v/v) CHCl3 /hexane. 10. Carefully transfer the crude product to the top of a 5 × 6–cm silica gel column (APPENDIX 3E). Elute successively with the following eluents:
DNA Synthesis Without Base Protection Using Phosphoramidites
3.15.2 Supplement 26
a. 500 mL of 7:3 (v/v) CHCl3 /hexane containing 1% pyridine. b. 500 mL CHCl3 containing 1% (v/v) pyridine (to remove the residues of the reagents). c. 500 mL of 99:1 (v/v) CHCl3 /MeOH containing 1% (v/v) pyridine (to remove the byproducts). d. 2 L of 97:3 (v/v) CHCl3 /MeOH containing 1% (v/v) pyridine. Collect 100-mL fractions and analyze by TLC. Current Protocols in Nucleic Acid Chemistry
11. Combine fractions containing pure product. Evaporate under reduced pressure and remove last traces of pyridine by coevaporating three times each with 5 mL toluene and 5 mL CHCl3 . 12. Check the purity of the product. 5 -O-(4,4 -Dimethoxytrityl)-2 -deoxycytidine (S.4a): 1.88 g as a white foam (71% yield from S.1a). TLC (6:1 [v/v] CHCl3 /MeOH), 0.59. 1 H NMR (270 MHz, CDCl3 ): 7.75 (d, J = 7.32 Hz, H6), 7.24 to 7.05 (m, DMTr), 6.69 to 6.64 (m, DMTr), 6.10 (t, J = 5.9 Hz, H1 ), 5.41 (d, J = 6.8 Hz, H5), 4.38 to 4.32 (m, H3 ), 3.91 to 3.81 (m, H4 ), 3.63 (s, MeO-), 3.34 to 3.22 (m, H5 ), 2.48 to 2.39 (m, H2 ), 2.10 to 2.01 (m, H2 ). This material is used for preparation of the 3 -phosphoramidite without further purification.
Prepare 5 -O-DMTr-2 -dC-3 -phosphoramidite 13. Suspend 2.65 g (5 mmol) S.4a in 3 mL pyridine in a 200-mL round-bottom flask and concentrate to dryness using a rotary evaporator and vacuum pump. Repeat this azeotropic drying process three times each with 3-mL portions of pyridine, toluene, and CH2 Cl2 . 14. Add 50 mL THF and a stir bar to the flask and attach a balloon filled with argon gas to create an argon atmosphere. 15. While stirring, add 1.23 mL (7.5 mmol) DIPEA. 16. Cool the mixture to −78◦ C in a dry ice/acetone bath and slowly add 1.23 mL (5.5 mmol) of 2-cyanoethoxy-(N,N-diisopropylamino)chlorophosphine. Continue stirring for 10 min. 17. Warm the flask gradually to room temperature and stir 30 min. Check the reaction by TLC using 9:1 (v/v) CHCl3 /MeOH (Rf = 0.54). 18. Add 2 mL water to quench the 2-cyanoethoxy-(N,N-diisopropylamino)chlorophosphine and stir 5 min. 19. Dissolve the mixture in 200 mL CHCl3 and wash with 200 mL brine in a 500-mL separatory funnel. 20. Dry over anhydrous sodium sulfate, filter through a no. 2 filter paper in a glass funnel, then evaporate under reduced pressure. 21. Dissolve the crude product in a minimal amount of 5:5 (v/v) CHCl3 /hexane. 22. Carefully transfer the crude product to the top of a 5 × 7.5–cm silica gel column. Elute successively with the following eluents: a. 500 mL of 7:3 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. b. 500 mL of 5:5 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (to remove the residues of the reagents). c. 500 mL of 4:6 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (to remove the byproducts). d. 500 mL of 3:7 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. e. 500 mL of CHCl3 containing 1% (v/v) triethylamine. f. 500 mL of 98:2 (v/v) CHCl3 /MeOH containing 1% (v/v) triethylamine. Collect 100-mL fractions and analyze by TLC. 23. Combine fractions containing pure product and evaporate under reduced pressure. Remove the last traces of triethylamine by coevaporating three times each with 5 mL toluene and 5 mL CHCl3 .
Synthesis of Unmodified Oligonucleotides
3.15.3 Current Protocols in Nucleic Acid Chemistry
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24. Check the purity of the product. 5 -O-(4,4 -Dimethoxytrityl)-2 -deoxycytidine 3 -(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.5a): 3.02 g as a white foam (83% yield from S.4a). TLC (9:1 [v/v] CHCl3 /MeOH), 0.54. 1 H NMR (270 MHz, CDCl3 ): 7.90, 7.81 (2d, J = 7.6 Hz, H6), 7.38 to 7.34 (m, DMTr), 7.27 to 7.17 (m, DMTr), 6.79 (dd, J = 4.1, 8.9 Hz, DMTr), 6.26 (dd, J = 6.2, 12.4 Hz, H1 ), 5.37 (d, J = 7.6 Hz, H5), 4.65 to 4.48 (m, H3 ), 4.09 (d, J = 3.8 Hz, H4 ), 3.75 to 3.71 (m, DMTr, iPr2 CH), 3.58 to 3.32 (m, -CH2 CN, iPr2 CH, H5 ), 2.68 to 2.52 (m, -OCH2 CH2 CN, H2 ), 2.37 (t, J = 6.2 Hz, -OCH2 CH2 CN), 2.25 to 2.11 (m, H2 ), 1.13 to 0.99 (m, iPr). 13 C NMR (67.8 MHz, CDCl3 ): 165.6, 158.6, 155.8, 144.4, 141.2, 135.5, 135.4, 130.2, 130.1, 128.2, 127.9, 127.0, 117.5, 117.4, 113.2, 93.9, 86.7, 85.9, 85.1, 72.7, 72.0, 71..7, 62.6, 62.2, 58.4, 58.1, 55.2, 43.3, 43.2, 43.1, 41.0, 40.7, 24.6, 24.6, 24.5, 22.6, 20.4, 20.3, 20.2, 20.1. 31 P NMR (109.4 MHz, CDCl3 ): 149.7, 149.2. MS (ESI) m/z (M+Na) calcd. for C39 H48 N5 NaO7 P+ 752.3189, found 752.3194. ALTERNATE PROTOCOL 1
SYNTHESIS OF THE DEOXYCYTIDINE 3 -PHOSPHORAMIDITE BY REMOVAL OF BENZOYL PROTECTION This method is performed by starting with a commercially available base-protected phosphoramidite (S.6a) and is illustrated in Figure 3.15.2.
Additional Materials (also see Basic Protocol 1) 5 -O-(4,4 -Dimethoxytrityl)-4-N-benzoyl-2 -deoxycytidine 3 -(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.6a; Glen Research) 2 M methylamine (MeNH2 ; Aldrich) in THF 5 × 8–cm chromatography column 1. Dissolve 4.17 g (5 mmol) S.6a in 50 mL of 2 M MeNH2 /THF in a 200-mL roundbottom flask equipped with a stir bar. Seal flask with a glass stopper held in place by metal clamps (to avoid hazards posed by gas pressure), and keep in fume hood. 2. Stir for 12 hr at room temperature and check the reaction by TLC (APPENDIX 3D) using 9:1 (v/v) CHCl3 /MeOH (Rf = 0.54). 3. Evaporate the mixture under reduced pressure and dissolve the crude product in a minimal amount of 5:5 (v/v) CHCl3 /hexane.
BPro DMTrO
P
DMTrO
O
NH3
O N
BNH2
O
O OCH2CH2CN
6a, B = cytosin-1-yl, Pro = benzoyl 6b, B = adenin-9-yl, Pro = benzoyl 6c, B = guanin-9-yl, Pro = isobutyryl 7a, B = cytosin-1-yl, Pro = acetyl 8b, B = adenin-9-yl, Pro = phenoxyacetyl 9c, B = guanin-9-yl, Pro = 4-isopropylphenoxyacetyl
N
P
OCH2CH2CN
5a, B = cytosin-1-yl 5b, B = adenin-9-yl 5c, B = guanin-9-yl
Figure 3.15.2 Synthesis of N-unprotected 3 -phosphoramidites by removal of nucleobase protecting groups from commercial phosphoramidites.
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4. Carefully transfer the crude product to the top of a 5 × 8–cm silica gel column. Elute successively with the following eluents: a. 500 mL of 7:3 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. b. 500 mL of 5:5 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. c. 500 mL of 4:6 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (to remove the residues of the reagents). d. 500 mL of 3:7 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. e. 500 mL of CHCl3 containing 1% (v/v) triethylamine. f. 500 mL of 98:2 (v/v) CHCl3 /MeOH containing 1% (v/v) triethylamine. Collect 100-mL fractions and analyze by TLC. 5. Combine fractions containing pure product and evaporate under reduced pressure. Remove the last traces of triethylamine by coevaporating three times each with 5 mL toluene and 5 mL CHCl3 . 6. Check the purity of the product. 5 -O-(4,4 -Dimethoxytrityl)-2 -deoxycytidine 3 -(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.5a): 3.10 g as a white foam (85% yield from 6a). TLC (9:1 [v/v] CHCl3 / MeOH), 0.54.
SYNTHESIS OF THE DEOXYCYTIDINE 3 -PHOSPHORAMIDITE BY REMOVAL OF ACETYL PROTECTION
ALTERNATE PROTOCOL 2
This method is performed using the commercially available phosphoramidite S.7a and is illustrated in Figure 3.15.2.
Additional Materials (also see Basic Protocol 1) 5 -O-(4,4 -Dimethoxytrityl)-4-N-acetyl-2 -deoxycytidine 3 -(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.7a; Glen Research) 2 M NH3 in MeOH (Aldrich) 5 × 8–cm chromatography column 1. Dissolve 3.86 g (5 mmol) S.7a in 50 mL of 2 M NH3 /MeOH in a 200-mL roundbottom flask equipped with a stir bar. 2. Stir for 2 hr at room temperature and check the reaction by TLC (APPENDIX 3D) using 9:1 (v/v) CHCl3 /MeOH (Rf = 0.54). 3. Evaporate the mixture under reduced pressure and dissolve the crude product in a minimal amount of 5:5 (v/v) CHCl3 /hexane. 4. Carefully transfer the crude product to the top of a 5 × 8–cm silica gel column. Elute successively with the following eluents: a. 500 mL of 7:3 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. b. 500 mL of 5:5 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. c. 500 mL of 4:6 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (to remove the residues of the reagents). d. 500 mL of 3:7 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. e. 500 mL of CHCl3 containing 1% (v/v) triethylamine. f. 500 mL of 98:2 (v/v) CHCl3 /MeOH containing 1% (v/v) triethylamine. Collect 100-mL fractions and analyze by TLC.
Synthesis of Unmodified Oligonucleotides
3.15.5 Current Protocols in Nucleic Acid Chemistry
Supplement 26
5. Combine fractions containing pure product and evaporate under reduced pressure. Remove the last traces of triethylamine by coevaporating three times each with 5 mL toluene and 5 mL CHCl3 . 6. Check the purity of the product. 5 -O-(4,4 -Dimethoxytrityl)-2 -deoxycytidine 3 -(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.5a): 3.43 g as a white foam (94% yield from S.7a). TLC (9:1 [v/v] CHCl3 / MeOH), 0.54. BASIC PROTOCOL 2
SYNTHESIS OF 5 -O-(4,4 -DIMETHOXYTRITYL-2 -DEOXYADENOSINE 3 -(2-CYANOETHYL-N,N-DIISOPROPYLPHOSPHORAMIDITE) FROM 2 -DEOXYADENOSINE The N-unprotected dA phosphoramidite is prepared by direct tritylation of deoxyadenosine followed by phosphitylation, as illustrated in Figure 3.15.1.
Materials 2 -Deoxyadenosine (S.1b; Yamasa Shoyu) ◦ Pyridine (anhydrous, dried over 4-A molecular sieves, distilled from CaH2 ) Argon source Triethylamine (Wako, distilled from CaH2 ) Dichloroacetic acid (Wako) 4,4 -Dimethoxytrityl chloride (DMTr-Cl) Chloroform (CHCl3 ) ◦ Methanol (MeOH; Wako, anhydrous, dried over 4-A molecular sieves) Brine (saturated aqueous NaCl) Sodium sulfate (Na2 SO4 , anhydrous) Hexane Silica gel (Wakogel C-200, 75 to 150 µm) ◦ Toluene (anhydrous, dried over 4-A molecular sieves) ◦ Dichloromethane (CH2 Cl2 , anhydrous, dried over 4-A molecular sieves) ◦ Tetrahydrofuran (THF, anhydrous, dried over 4-A molecular sieves, no stabilizer) Diisopropylethylamine (DIPEA, anhydrous, dried over CaH2 , distilled from CaH2 ) 2-Cyanoethoxy-(N, N-diisopropylamino)chlorophosphine (synthesized as described in Nagai et al., 1989; Tanimura et al., 1989) 200-mL round-bottom flasks with argon-filled balloon Rotary evaporator and vacuum pump Thin-layer chromatography (TLC) silica plates (Kieselgel 60 F254 ; 0.2 mm thick; Merck) 500-mL separatory funnel No. 2 filter paper and glass funnel 5 × 6– and 5 × 7.5–cm chromatography columns Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare 5 -O-DMTr-2 -dA 1. Suspend 1.26 g (5 mmol) 2 -deoxyadenosine (S.1b) in 3 mL pyridine in a 200-mL round-bottom flask and concentrate the suspension to dryness using a rotary evaporator and vacuum pump. Repeat this azeotropic drying process three times, each time with 3 mL pyridine. DNA Synthesis Without Base Protection Using Phosphoramidites
2. Add 50 mL pyridine and a stir bar to the flask and attach a balloon filled with argon gas to create an argon atmosphere.
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3. While stirring, add 699 µL (5 mmol) triethylamine and then slowly add 412 µL (5 mmol) dichloroacetic acid. 4. While stirring, add 1.86 g (5.5 mmol) DMTr-Cl. 5. Stir the mixture for 4 hr at room temperature and check the reaction by TLC (APPENDIX 3D) using 6:1 (v/v) CHCl3 /MeOH (Rf = 0.68). 6. Add 10 mL MeOH to quench DMTr-Cl and stir 10 min. 7. Dissolve the mixture in 200 mL CHCl3 , wash with 200 mL brine in a 500-mL separatory funnel. 8. Dry organic layer over anhydrous sodium sulfate, filter through no. 2 filter paper in a glass funnel, and evaporate using a rotary evaporator and vacuum pump. 9. Dissolve the crude product in a minimal amount of 7:3 (v/v) CHCl3 /hexane. 10. Carefully transfer the crude product to the top of a 5 × 6–cm silica gel column (APPENDIX 3E). Elute successively with the following eluents: a. 500 mL of 7:3 (v/v) CHCl3 /hexane containing 1% (v/v) pyridine. b. 500 mL of CHCl3 containing 1% (v/v) pyridine (to remove the residues of the reagents). c. 500 mL of 99.5:0.5 (v/v) CHCl3 /MeOH containing 1% (v/v) pyridine. d. 500 mL of 99:1 (v/v) CHCl3 /MeOH containing 1% (v/v) pyridine (to remove the byproducts). e. 500 mL of 98.5:1.5 (v/v) CHCl3 /MeOH containing 1% (v/v) pyridine. f. 1 L of 97:3 (v/v) CHCl3 /MeOH containing 1% (v/v) pyridine. Collect 100-mL fractions and analyze by TLC. 11. Combine fractions containing pure product. Evaporate under reduced pressure and remove last traces of pyridine by coevaporating three times each with 5 mL toluene and 5 mL CHCl3 . 12. Check the purity of the product. 5 -O-(4,4 -Dimethoxytrityl)-2 -deoxyadenosine (S.4b): 2.04 g as a white foam (74% yield from S.1b). TLC (6:1 [v/v] CHCl3 /MeOH), 0.68. 1 H NMR (270 MHz, CDCl3 ): 8.25 (s, H2), 7.97(s, H8), 7.39 to 7.15 (m, DMTr), 6.79 to 6.75 (m, DMTr), 6.44 (t, J = 6.75 Hz, H1 ), 6.15 (brs, NH2 ), 4.70 to 4.65 (m, H3 ), 4.19 to 4.15 (m, H4 ), 3.75 (s, MeO-), 3.42 to 3.36 (m, H5 ), 2.81 to 2.74 (m, H2 ), 2.57 to 2.50 (m, H2 ). This material is used for preparation of the 3 -phosphoramidite without further purification.
Prepare 5 -O-DMTr-2 -dA-3 -phosphoramidite 13. Suspend 2.88 g (5 mmol) S.4b in 3 mL pyridine in a 200-mL round-bottom flask and concentrate to dryness using a rotary evaporator and vacuum pump. Repeat this azeotropic drying process three times each with 3-mL portions of pyridine, toluene and CH2 Cl2 . 14. Add 50 mL THF and a stir bar to the flask and attach a balloon filled with argon gas to create an argon atmosphere. 15. While stirring, add 1.23 mL (7.5 mmol) DIPEA. 16. Cool the mixture to −78◦ C in a dry ice/acetone bath and slowly add 1.23 mL (5.5 mmol) of 2-cyanoethoxy-(N, N-diisopropylamino)chlorophosphine. Continue stirring for 10 min.
Synthesis of Unmodified Oligonucleotides
3.15.7 Current Protocols in Nucleic Acid Chemistry
Supplement 26
17. Warm the flask gradually to room temperature and stir the mixture 30 min. Check the reaction by TLC using 9:1 (v/v) CHCl3 /MeOH (Rf = 0.63). 18. Add 2 mL water to quench 2-cyanoethoxy-(N, N-diisopropylamino)chlorophosphine, and stir 5 min. 19. Dissolve the mixture in 200 mL CHCl3 , wash with 200 mL brine in a 500-mL separatory funnel. 20. Dry over anhydrous sodium sulfate, filter through a no. 2 filter paper in a glass funnel, then evaporate under reduced pressure. 21. Dissolve the crude product in a minimal amount of 5:5 (v/v) CHCl3 /hexane. 22. Carefully transfer the crude product to the top of a 5 × 7.5–cm silica gel column. Elute successively with the following eluents: a. 500 mL of 7:3 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. b. 500 mL of 5:5 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (to remove the residues of the reagents). c. 500 mL of 4:6 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (to remove the byproducts). d. 500 mL of 3:7 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. e. 1.5 L of CHCl3 containing 1% (v/v) triethylamine. Collect 100-mL fractions and analyze by TLC. 23. Combine fractions containing pure product and evaporate under reduced pressure. Remove the last traces of triethylamine by coevaporating three times each with 5 mL toluene and 5 mL CHCl3 . 24. Check the purity of the product. 5-O-(4,4-Dimethoxytrityl)-2 -deoxyadenosine3 -(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.5b): 3.46 g as a white foam (92% yield from S.4b). TLC (9:1 [v/v] CHCl3 /MeOH), 0.63. 1 H NMR (270 MHz, CDCl3 ): 8.26 (s, H2), 7.99, 7.96 (2s, H7), 7.39 to 7.16 (m, DMTr), 6.76 (dd, J = 3.78, 8.91 Hz, DMTr), 6.42 (t, J = 6.35 Hz, H1 ), 5.79 (brs, NH2 ), 4.76 to 4.68 (m, H3 ), 4.28 to 4.24 (m, H4 ), 3.83 to 3.54 (m, DMTr, iPr2 CH, -CH2 CN), 3.40 to 3.31 (m, H5 ), 2.91 to 2.82 (m, H2 ), 2.65 to 2.55 (m, -OCH2 CH2 CN, H2 ), 2.44 (t, -OCH2 CH2 CN), 1.20 to 1.08 (m, iPr). 13 C NMR (67.8 MHz, CDCl3 ): 158.5, 155.4, 153.0, 149.7, 149.6, 144.5, 139.0, 135.7, 135.6, 130.0, 128.2, 128.1, 127.8, 126.8, 120.1, 117.5, 117.4, 113.1, 86.4, 85.6, 84.4, 74.2, 74.0, 63.5, 63.3, 58.4, 58.1, 55.2, 43.3, 43.1, 39.5, 24.6, 24.5, 24.4, 20.4, 20.3, 20.2, 20.1. 31 P NMR (109.4 MHz, CDCl3 ): 149.4, 149.3. MS (ESI) m/z (M+Na) calcd. for C40 H48 N7 NaO6 P+ 776.3301, found 776.3340. ALTERNATE PROTOCOL 3
SYNTHESIS OF THE DEOXYADENOSINE 3 -PHOSPHORAMIDITE BY REMOVAL OF BENZOYL PROTECTION This method is performed using the commercially available base-protected phosphoramidite 6b and is illustrated in Figure 3.15.2.
Additional Materials (also see Basic Protocol 2)
DNA Synthesis Without Base Protection Using Phosphoramidites
3.15.8 Supplement 26
5 -O-(4,4 -Dimethoxytrityl)-6-N-benzoyl-2 -deoxyadenosine 3 -(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.6b; Glen Research) 2 M methylamine (MeNH2 ; Aldrich) in THF 5 × 8–cm chromatography column 1. Dissolve 4.29 g (5 mmol) S.6b in 50 mL of 2 M MeNH2 /THF in a 200-mL roundbottom flask equipped with a stir bar. Seal flask with a glass stopper held in place by metal clamps (to avoid hazards posed by gas pressure), and keep in fume hood. Current Protocols in Nucleic Acid Chemistry
2. Stir for 2 hr at room temperature and check the reaction by TLC (APPENDIX 3D) using 9:1 (v/v) CHCl3 /MeOH (Rf = 0.63). 3. Evaporate the mixture under reduced pressure and dissolve the crude product in a minimal amount of 5:5 (v/v) CHCl3 /hexane. 4. Carefully transfer the crude product to the top of a 5 × 8–cm silica gel column. Elute successively with the following eluents: a. 500 mL of 7:3 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. b. 500 mL of 5:5 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. c. 500 mL of 4:6 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (to remove the byproducts). d. 500 mL of 3:7 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. e. 1.5 L of CHCl3 containing 1% (v/v) triethylamine. Collect 100-mL fractions and analyze by TLC. 5. Combine fractions containing pure product and evaporate under reduced pressure. Remove the last traces of triethylamine by coevaporating three times each with 5 mL toluene and 5 mL CHCl3 . 6. Check the purity of the product. 5-O-(4,4-Dimethoxytrityl)-2-deoxyadenosine3-(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.5b): 3.35 g as a white foam (89% yield from S.6b). TLC (9:1 [v/v] CHCl3 / MeOH), 0.63.
SYNTHESIS OF THE DEOXYADENOSINE 3 -PHOSPHORAMIDITE BY REMOVAL OF PHENOXYACETYL PROTECTION
ALTERNATE PROTOCOL 4
This method is performed using the commercially available base-protected phosphoramidite S.8b and is illustrated in Figure 3.15.2.
Additional Materials (also see Basic Protocol 2) 5 -O-(4,4 -Dimethoxytrityl)-6-N-phenoxyacetyl-2 -deoxyadenosine 3 -(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.8b; Glen Research) 2 M NH3 in MeOH (Aldrich) 1. Dissolve 4.43 g (5 mmol) S.8b in 50 mL of 2 M NH3 /MeOH in a 200-mL roundbottom flask equipped with a stir bar. 2. Stir for 1 hr at room temperature and check the reaction by TLC (APPENDIX 3D) using 9:1 (v/v) CHCl3 /MeOH (Rf = 0.63). 3. Evaporate the mixture under reduced pressure and dissolve the crude product in a minimal amount of 5:5 (v/v) CHCl3 /hexane. 4. Carefully transfer the crude product to the top of a 5 × 8–cm silica gel column. Elute successively with the following eluents: a. 500 mL of 7:3 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. b. 500 mL of 5:5 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. c. 500 mL of 4:6 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (to remove the byproducts). d. 500 mL of 3:7 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. e. 1.5 L of CHCl3 containing 1% (v/v) triethylamine. Collect 100-mL fractions and analyze by TLC.
Synthesis of Unmodified Oligonucleotides
3.15.9 Current Protocols in Nucleic Acid Chemistry
Supplement 26
5. Combine fractions containing pure product and evaporate under reduced pressure. Remove the last traces of triethylamine by coevaporating three times each with 5 mL toluene and 5 mL CHCl3 . 6. Check the purity of the product. 5-O-(4,4-Dimethoxytrityl)-2-deoxyadenosine3-(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.5b): 3.69 g as a white foam (98% yield from S.8b). TLC (9:1 [v/v] CHCl3 /MeOH), 0.63. BASIC PROTOCOL 3
SYNTHESIS OF 5 -O-(4,4 -DIMETHOXYTRITYL)-2 -DEOXYGUANOSINE 3 -(2-CYANOETHYL-N,N-DIISOPROPYLPHOSPHORAMIDITE) FROM 2 -DEOXYGUANOSINE In contrast to the N-unprotected dA and dC phosphoramidites, the dG derivative is prepared from deoxyguanosine by a method involving N-dimethylaminomethylene (dmf) protection followed by tritylation, removal of the dmf group, and then phosphitylation. The entire four-step procedure is illustrated in Figure 3.15.3.
Materials 2 -Deoxyguanosine (S.1c; Yamasa Shoyu) ◦ Pyridine (anhydrous, dried over 4-A molecular sieves, distilled from CaH2 ) ◦ Methanol (MeOH; Wako, anhydrous, dried over 4-A molecular sieves) Argon source N,N-Dimethylformamide dimethyl acetal (Tokyo Kasei Co. Ltd.; http://www.tokyokasei.co.jp/) 4,4 -Dimethoxytrityl chloride (DMTr-Cl) Chloroform (CHCl3 ) Brine (saturated aqueous NaCl) Sodium sulfate (Na2 SO4 , anhydrous) Diisopropyl ether Ammonium hydroxide Silica gel (Wakogel C-200, 75 to 150 µm) Hexane ◦ Toluene (anhydrous, dried over 4-A molecular sieves) ◦ Dichloromethane (CH2 Cl2 , anhydrous, dried over 4-A molecular sieves) Diisopropylethylamine (DIPEA, anhydrous, dried over CaH2 , distilled from CaH2 ) 2-Cyanoethoxy-(N,N-diisopropylamino)chlorophosphine (synthesized as described in Nagai et al., 1989; Tanimura et al., 1989) Triethylamine (Wako, distilled from CaH2 ) 100- and 200-mL round-bottom flasks with argon-filled balloon Rotary evaporator and vacuum pump Thin-layer chromatography (TLC) silica plates (Kieselgel 60 F254 ; 0.2 mm thick; Merck) 60-mm Buchner funnel with filter paper 500-ml separatory funnel No. 2 filter paper and glass funnel 5 × 6– and 5 × 7.5–cm chromatography columns Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E)
DNA Synthesis Without Base Protection Using Phosphoramidites
3.15.10 Supplement 26
Prepare 5 -O-DMTr-2 -dG 1. Suspend 1.33 g (5 mmol) 2 -deoxyguanosine (S.1c) in 3 mL pyridine in a 100-mL round-bottom flask and concentrate to dryness using a rotary evaporator and vacuum pump. Repeat this azeotropic drying process three times, each time with 3 mL pyridine. Current Protocols in Nucleic Acid Chemistry
Figure 3.15.3
Synthesis of N-unprotected 3 -phosphoramidites via transient N-dimethylaminomethylene protection.
2. Add 30 mL dry MeOH and a stir bar to the flask and attach a balloon filled with argon gas to create an argon atmosphere. 3. While stirring, add 1.81 mL (15 mmol) N,N-dimethylformamide dimethyl acetal. 4. Stir for 4 hr at 55◦ C and check the reaction by TLC (APPENDIX 3D) using 4:1 (v/v) CHCl3 /MeOH (Rf = 0.11). 5. Cool the mixture to room temperature, then collect the resulting colorless precipitates by filtration on a 60-mm Buchner funnel with filter paper and dry under suction. 6. Suspend the dried residue in 3 mL pyridine in a 200-mL round-bottom flask and concentrate to dryness using a rotary evaporator and vacuum pump. Repeat this azeotropic drying process three times, each time with 3 mL pyridine. 7. Add 50 mL pyridine and a stir bar to the flask and attach a balloon filled with argon gas to create an argon atmosphere. 8. While stirring, add 1.86 g (5.5 mmol) DMTr-Cl. 9. Stir the mixture for 4 hr at room temperature and check the reaction by TLC using 6:1 (v/v) CHCl3 /MeOH (Rf = 0.57). 10. Add 10 mL MeOH to quench DMTr-Cl and stir 10 min. 11. Dissolve the mixture in 200 mL CHCl3 and wash with 200 mL brine in a 500-mL separatory funnel. 12. Dry organic layer over anhydrous sodium sulfate, filter through no. 2 filter paper in a glass funnel, and evaporate using a rotary evaporator and vacuum pump. 13. Dissolve the crude product in a minimal amount of CHCl3 and add 200 mL diisopropyl ether. 14. Collect the resulting yellow precipitate by filtration on a 60-mm Buchner funnel with filter paper and dry under suction.
Synthesis of Unmodified Oligonucleotides
3.15.11 Current Protocols in Nucleic Acid Chemistry
Supplement 26
15. Dissolve the resulting material in 100 mL of 1:1 (v/v) pyridine/28% ammonium hydroxide. Stir 15 hr at room temperature and check the reaction by TLC using 6:1 (v/v) CHCl3 /MeOH (Rf = 0.52). 16. Evaporate the reaction mixture in a rotary evaporator and dissolve the crude product in a minimal amount of CHCl3 . 17. Carefully transfer the crude product to the top of a 5 × 6–cm silica gel column (APPENDIX 3E). Elute successively with the following eluents: a. 500 mL of 8:2 (v/v) hexane/CHCl3 containing 1% (v/v) pyridine. b. 500 mL of CHCl3 containing 1% (v/v) pyridine (to remove the residues of the reagents). c. 500 mL of 99:1 (v/v) CHCl3 /MeOH containing 1% (v/v) pyridine. d. 500 mL of 98:2 (v/v) CHCl3 /MeOH containing 1% (v/v) pyridine (to remove the byproducts). e. 1 L of 97:3 (v/v) CHCl3 /MeOH containing 1% (v/v) pyridine. f. 2 L of 95:5 (v/v) CHCl3 /MeOH containing 1% (v/v) pyridine. Collect 100-mL fractions and analyze by TLC. 18. Combine fractions containing pure product and evaporate under reduced pressure. Dissolve the mixture in 200 mL of 1:1 (v/v) pyridine/CHCl3 and wash with 200 mL brine in a 500-mL separatory funnel. 19. Dry organic layer over anhydrous sodium sulfate, filter through no. 2 filter paper in a glass funnel, and evaporate using a rotary evaporator and vacuum pump. 20. Coevaporate the resulting material three times each with 5 mL toluene and 5 mL CHCl3 . 21. Check the purity of the product. 5 -O-(4,4 -Dimethoxytrityl)-2 -deoxyguanosine (4c): 1.65 g as a white foam (58% yield from S.1c). TLC (6:1 [v/v] CHCl3 /MeOH), 0.52. 1 H NMR (270 MHz, CDCl3 ): 10.58 (brs, NH), 7.76 (s, H8), 7.34 to 7.17 (m, DMTr), 6.81 (dd, J = 6.3, 8.8 Hz, DMTr), 6.42 (brs, NH2 ), 6.12 (dd, J = 6.3, 6.8 Hz, H1 ), 4.35 to 4.32 (m, H3 ), 3.91 to 3.81 (m, H4 ), 3.71 (s, MeO-), 3.15 to 3.09 (m, H5 ), 2.67 to 2.57 (m, H2 ), 2.31 to 2.22 (m, H2 ). This material is used for preparation of the 3 -phosphoramidite without further purification.
Prepare 5 -O-DMTr-2 -dG-3 -phosphoramidite 22. Suspend 2.65 g (5 mmol) S.4c in 3 mL pyridine in a 200-mL round-bottom flask and concentrate to dryness using a rotary evaporator and vacuum pump. Repeat this azeotropic drying process three times each with 3-mL portions of pyridine, toluene, and CH2 Cl2 . 23. Add 50 mL CH2 Cl2 and a stir bar to the flask and attach a balloon filled with argon gas to create an argon atmosphere. 24. While stirring, add 1.23 mL (7.5 mmol) DIPEA. 25. Add 1.23 mL (5.5 mmol) 2-cyanoethoxy-(N,N-diisopropylamino)chlorophosphine. 26. Stir for 30 min and check the reaction by TLC using 9:1 (v/v) CHCl3 /MeOH (Rf = 0.48). DNA Synthesis Without Base Protection Using Phosphoramidites
27. Add 2 mL water to quench 2-cyanoethoxy-(N,N-diisopropylamino)chlorophosphine and stir 5 min.
3.15.12 Supplement 26
Current Protocols in Nucleic Acid Chemistry
28. Dissolve the mixture in 200 mL CHCl3 and wash with 200 mL brine in a 500-ml separatory funnel. 29. Dry organic layer over anhydrous sodium sulfate, filter through no. 2 filter paper in a glass funnel, and evaporate under reduced pressure. 30. Dissolve the crude product in a minimal amount of 5:5 (v/v) CHCl3 /hexane. 31. Carefully transfer the crude product to the top of a 5 × 7.5–cm silica gel column. Elute successively with the following eluents: a. 500 mL of 7:3 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. b. 500 mL of 5:5 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (to remove the residues of the reagents). c. 500 mL of 4:6 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (to remove the byproducts). d. 500 mL of 3:7 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (500 mL). e. 500 mL of CHCl3 containing 1% (v/v) triethylamine. f. 500 mL of 98.5:1.5 (v/v) CHCl3 /MeOH containing 1% (v/v) triethylamine. g. 500 mL of 97:3 (v/v) CHCl3 /MeOH containing 1% (v/v) triethylamine. Collect 100-mL fractions and analyze by TLC. 32. Combine fractions containing pure product and evaporate under reduced pressure. Remove the last traces of triethylamine by coevaporating three times each with 5 mL toluene and 5 mL CHCl3 . 33. Check the purity of the product. 5-O-(4,4-Dimethoxytrityl)-2-deoxyguanosine 3-(2 -cyanoethyl-N,N-diisopropylphosphoramidite) (S.5c): 3.08 g as a white foam (80% yield from S.4c). TLC (9:1 [v/v] CHCl3 /MeOH), 0.48. 1 H NMR (270 MHz, CDCl3 ): 7.66, 7.64 (2s, H8), 7.41 to 7.38 (m, DMTr), 7.30 to 7.14 (m, DMTr), 6.78 (dd, J = 2.70, 8.64 Hz, DMTr), 6.21 (t, J = 7.02 Hz, H1 ), 6.02 (brs, NH2 ), 4.73 to 4.70 (m, H3 ), 4.29 to 4.22 (m, H4 ), 3.90 to 3.53 (m, DMTr, iPr2 CH, -CH2 CN), 3.40 to 3.29 (m, H5 ), 2.85 to 2.72 (m, H2 ), 2.61 to 2.44 (m, H2 , -OCH2 CH2 CN), 1.19 to 1.04 (m, iPr). 13 C NMR (67.8 MHz, CDCl3 ):159.1, 158.5, 153.5, 151.52, 151.48, 144.5, 135.7, 130.10, 130.06, 129.0, 128.21, 128.17, 127.8, 126.9, 125.3, 117.6, 117.5, 127.4, 117.4, 113.1, 86.4, 85.7, 85.4, 84.0, 77.2, 74.4, 74.1, 73.7, 73.5, 63.4, 63.3, 58.43, 58.37, 58.2, 58.1, 55.2, 46.2, 43.3, 43.2, 39.1, 24.6, 24.50, 24.45, 20.4, 20.3, 20.2, 20.1. 31 P NMR (109.4 MHz, CDCl3 ): 149.5, 149.3. MS (ESI) m/z (M+H) calcd. for C40 H49 N7 O7 P+ 770.3431, found 770.3501.
SYNTHESIS OF THE DEOXYGUANOSINE 3 -PHOSPHORAMIDITE BY REMOVAL OF ISOBUTYRYL PROTECTION
ALTERNATE PROTOCOL 5
This method is performed using the commercially available base-protected phosphoramidite S.6c and is illustrated in Figure 3.15.2.
Additional Materials (also see Basic Protocol 2) 5 -O-(4,4 -Dimethoxytrityl)-2-N-isobutyryl-2 -deoxyguanosine 3 -(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.6c; Glen Research) 2 M methylamine (MeNH2 ; Aldrich) in THF 5 × 8–cm chromatography column 1. Dissolve 4.20 g (5 mmol) S.6c in 50 mL of 2 M MeNH2 /THF in a 200-mL roundbottom flask equipped with a stir bar. Seal flask with a glass stopper held in place by metal clamps (to avoid hazards posed by gas pressure), and keep in fume hood.
Synthesis of Unmodified Oligonucleotides
3.15.13 Current Protocols in Nucleic Acid Chemistry
Supplement 26
2. Stir for 14 hr at room temperature and check the reaction by TLC (APPENDIX 3D) using 9:1 (v/v) CHCl3 /MeOH (Rf = 0.48). 3. Evaporate the mixture under reduced pressure and dissolve the crude product in a minimal amount of 5:5 (v/v) CHCl3 /hexane. 4. Carefully transfer the crude product to the top of a 5 × 8–cm silica gel column. Elute successively with the following eluents: a. 500 mL of 7:3 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. b. 500 mL of 5:5 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. c. 500 mL of 4:6 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (to remove the byproducts). d. 500 mL of 3:7 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. e. 500 mL of CHCl3 containing 1% (v/) triethylamine. f. 500 mL of 98.5:1.5 (v/v) CHCl3 /MeOH containing 1% (v/v) triethylamine. g. 500 mL of 97:3 (v/v) CHCl3 /MeOH containing 1% (v/v) triethylamine. Collect 100-mL fractions and analyze by TLC. 5. Combine fractions containing pure product and evaporate under reduced pressure. Remove the last traces of triethylamine by coevaporating three times each with 5 mL toluene and 5 mL CHCl3 . 6. Check the purity of the product. 5-O-(4,4-Dimethoxytrityl)-2-deoxyguanosine 3-(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.5c): 2.88 g as a white foam (75% yield from S.6c). TLC (9:1 [v/v] CHCl3 /MeOH), 0.48. ALTERNATE PROTOCOL 6
SYNTHESIS OF THE DEOXYGUANOSINE 3 -PHOSPHORAMIDITE BY REMOVAL OF ISOPROPYLPHENOXYACETYL PROTECTION This method is performed using the commercially available base-protected phosphoramidite S.9c and is illustrated in Figure 3.15.2.
Additional Materials (also see Basic Protocol 1) 5 -O-(4,4 -Dimethoxytrityl)-2-N-(4-isopropylphenoxyacetyl)-2 -deoxyguanosine 3 -(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.9c; Glen Research) 2 M NH3 in MeOH (Aldrich) 5 × 8–cm chromatography column 1. Dissolve 4.80 g (5 mmol) S.9c in 50 mL of 2 M NH3 /MeOH in a 200-mL roundbottom flask equipped with a stir bar. 2. Stir for 2 hr at room temperature and check the reaction by TLC (APPENDIX 3D) using 9:1 (v/v) CHCl3 /MeOH (Rf = 0.54). 3. Evaporate the mixture under reduced pressure and dissolve the crude product in a minimal amount of 5:5 (v/v) CHCl3 /hexane. 4. Carefully transfer the crude product to the top of a 5 × 8–cm silica gel column. Elute successively with the following eluents: DNA Synthesis Without Base Protection Using Phosphoramidites
a. 500 mL of 7:3 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. b. 500 mL of 5:5 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. c. 500 mL of 4:6 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine (to remove the byproducts).
3.15.14 Supplement 26
Current Protocols in Nucleic Acid Chemistry
d. e. f. g.
500 mL of 3:7 (v/v) hexane/CHCl3 containing 1% (v/v) triethylamine. 500 mL of CHCl3 containing 1% (v/) triethylamine. 500 mL of 98.5:1.5 (v/v) CHCl3 /MeOH containing 1% (v/v) triethylamine. 500 mL of 97:3 (v/v) CHCl3 /MeOH containing 1% (v/v) triethylamine.
Collect 100-mL fractions and analyze by TLC. 5. Combine fractions containing pure product and evaporate under reduced pressure. Remove the last traces of triethylamine by coevaporating three times each with 5 mL toluene and 5 mL CHCl3 . 6. Check the purity of the product. 5-O-(4,4-Dimethoxytrityl)-2-deoxyguanosine 3-(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.5c): 3.46 g as a white foam (90% yield from S.9c). TLC (9:1 [v/v] CHCl3 /MeOH), 0.48.
TYPICAL PROCEDURE FOR SOLID-PHASE SYNTHESIS OF OLIGODEOXYRIBONUCLEOTIDES
BASIC PROTOCOL 4
Materials 5 -O-(4,4 -Dimethoxytrityl)-2 -deoxynucleoside 3 -(2-cyanoethylN,N-diisopropylphosphoramidite)s: C (S.5a), A (S.5b), G (S.5c), and T Thymidine-loaded HCP (28 µmol/g, succinate linker) 3% trichloroacetic acid in CH2 Cl2 (Glen Research) ◦ Dichloromethane (CH2 Cl2 , anhydrous, dried over 4-A molecular sieves) ◦ Acetonitrile (CH3 CN, anhydrous, dried over 3-A molecular sieves, distilled from CaH2 ) ◦ N-Methyl-2-pyrrolidone (NMP, anhydrous, Wako, dried over 4-A molecular sieves) 1-Hydroxybenzotriazole (HOBt, Dojindo Molecular Technologies) Oxidizing solution: 0.1 M I2 in 9:1 (v/v) pyridine/H2 O (for manual operation) or 0.1 M I2 in 2:1:7 (v/v/v) pyridine/H2 O/THF (Glen Research; for automated operation) ◦ Pyridine (anhydrous, dried over 4-A molecular sieves, distilled from CaH2 ) Concentrated NH3 , aqueous 1,8-Diazabicyclo[5.4.0]undec-7-ene (DBU, anhydrous, dried over CaH2 , distilled from CaH2 ) Tetrabutylammonium fluoride (TBAF) Acetic acid (AcOH; Wako) ◦ Tetrahydrofuran (THF, anhydrous, dried over 4-A molecular sieves, no stabilizer) 1-Hydroxy-6-(trifluoromethyl)benzotriazole (HOtf Bt, Aldrich) Benzimidazolium triflate (BIT; Hayakawa et al., 1996) ABI 392 DNA synthesizer Manual Operation A thymidine-loaded highly cross-linked polystyrene or HCP (1 µmol, 28 µmol/g, succinate linker; ABI) is used. Each cycle of chain elongation consists of: 1. detritylation: 3% trichloroacetic acid in CH2 Cl2 , 2 mL, 1 min 2. washing: CH2 Cl2 (three times, each with 1 mL), CH3 CN (three times, each with 1 mL) 3. first coupling: appropriate phosphoramidite unit (10 µmol) in CH3 CN/NMP (150 µL, 9:1 v/v), followed by HOBt (6.3 mg, 20 µmol) in CH3 CN/NMP (150 µL, 9:1 v/v), 1 min 4. washing: CH3 CN (three times, each with 1 ml)
Synthesis of Unmodified Oligonucleotides
3.15.15 Current Protocols in Nucleic Acid Chemistry
Supplement 26
Table 3.15.1 ABI 392 DNA Synthesizer Program Using N-Unprotected Phosphoramidites
Step
Operation
Reagent(s)
Time (min)
1
Washing
CH3 CN
0.2
2
Detritylation
3% Cl3 CCOOH/CH2 Cl2
1.5
3
Washing
CH3 CN
0.4
4
Coupling
0.1 M amidite + 0.2 M HOtf Bt in 15:1 (v/v) CH3 CN/NMP
1.0
5
Coupling
0.1 M amidite + 0.2 M HOtf Bt in 15:1 (v/v) CH3 CN/NMP
1.0
6
Washing
CH3 CN
0.2
7
Oxidation
0.1 M I2 in 2:1:7 (v/v/v) pyridine/H2 O/THF
0.5
8
Washing
CH3 CN
0.4
5. second coupling: appropriate phosphoramidite unit (10 µmol) in CH3 CN/NMP (150 µL, 9:1 v/v), followed by HOBt (6.3 mg, 20 µmol) in CH3 CN/NMP (150 µL, 9:1 v/v), 1 min 6. washing: CH3 CN (three times, each with 1 mL) 7. oxidation: 0.1 M I2 in pyridine/H2 O, 9:1 (v/v), 2 min 8. washing: pyridine (three times, each with 1 mL), CH3 CN (three times, each with 1 mL), CH2 Cl2 (three times, each with 1 mL). Generally, the average yield per cycle has been estimated to be 98% to 99% by the DMTr cation assay (UNIT 10.3). After chain elongation is finished, the DMTr group is removed by treatment with 3% trichloroacetic acid in CH2 Cl2 (2 mL) for 1 min, and the resin is washed with CH2 Cl2 (three times, each with 1 mL) and CH3 CN (three times, each with 1 mL). The oligomer is deprotected and released from the polymer support by treatment with concentrated aqueous NH3 (500 µL) for 40 min. The polymer support is removed by filtration and washed with CH3 CN (three times, each with 1 mL). The filtrate is evaporated and purified by reversed-phase or anion-exchange HPLC.
Automated Operation The synthesis of oligodeoxyribonucleotides by use of the ABI 392 DNA synthesizer is carried out on a polymer support with a succinate-type linker according to the reaction cycle shown in Table 3.15.1. The unmodified oligomer after chain elongation is deprotected and released from the polymer support by treatment with concentrated aqueous NH3 (500 µL) for 40 min. The polymer support is removed by filtration and washed with CH3 CN (three times, each with 1 mL). The filtrate is evaporated and purified by reversed-phase or anion-exchange HPLC.
DNA Synthesis Without Base Protection Using Phosphoramidites
On the other hand, synthesis of an oligomer having alkaline-labile functional groups is performed on a polymer support with a silyl-type linker using the ABI 392 DNA synthesizer program shown in Table 3.15.2. After chain elongation, the oligomer is deprotected by treatment with a 10% DBU solution in CH3 CN (500 µL) for 1 min. The oligodeoxyribonucleotide mixture is then released from the resin by treatment with 500 µl of a 1:1 mixture of 1 M TBAF in AcOH and THF for 1 hr. The polymer support is removed by filtration and washed with CH3 CN (three times, each with 1 mL). The filtrate is evaporated and purified by reversed-phase or anion-exchange HPLC.
3.15.16 Supplement 26
Current Protocols in Nucleic Acid Chemistry
Table 3.15.2 ABI 392 DNA Synthesizer Program for Oligomers with Alkaline-Labile Functional Groups
Step
Number
Operation Name
Time (sec)
1
106
Begin
2
64
18 to Waste
3.0
3
42
18 to Column
10.0
4
2
Reverse Flush
10.0
5
1
Block Flush
4.0
6
101
Phos Prep
3.0
7
140
Column 1 On
8
111
Block Vent
2.0
9
58
Tet to Waste
1.7
10
33
B + Tet to Column
2.5
11
34
Tet to Column
1.0
12
33
B + Tet to Column
2.5
13
43
Push to Column
14
141
Column 1 Off
15
142
Column 2 On
16
64
18 to Waste
4.0
17
1
Block Flush
3.0
18
111
Block Vent
2.0
19
58
Tet to Waste
1.7
20
33
B + Tet to Column
2.5
21
34
Tet to Column
1.0
22
33
B + Tet to Column
2.5
23
43
Push to Column
24
143
Column 2 Off
25
103
Wait
15.0
26
2
Reverse Flush
0.1
27
5
Flush to Collect
0.1
28
103
Wait
15.0
29
2
Reverse Flush
0.1
30
5
Flush to Collect
0.1
31
103
Wait
15.0
32
2
Reverse Flush
0.1
33
5
Flush to Collect
0.1
34
103
Wait
15.0
35
58
Tet to Waste
1.7
36
33
B + Tet to Column
2.5
37
34
Tet to Column
1.0
38
33
B + Tet to Column
2.5
39
43
Push to Column continued
Synthesis of Unmodified Oligonucleotides
3.15.17 Current Protocols in Nucleic Acid Chemistry
Supplement 26
Table 3.15.2 ABI 392 DNA Synthesizer Program for Oligomers with Alkaline-Labile Functional Groups, continued
Step
DNA Synthesis Without Base Protection Using Phosphoramidites
Number
Operation Name
Time (sec)
40
143
Column 2 Off
41
103
Wait
15.0
42
2
Reverse Flush
0.1
43
5
Flush to Collect
0.1
44
103
Wait
15.0
45
2
Reverse Flush
0.1
46
5
Flush to Collect
0.1
47
103
Wait
15.0
48
2
Reverse Flush
0.1
49
5
Flush to Collect
0.1
50
103
Wait
15.0
51
64
18 to Waste
4.0
52
42
18 to Column
10.0
53
2
Reverse Flush
7.0
54
1
Block Flush
3.0
55
64
18 to Waste
10.0
56
2
Reverse Flush
7.0
57
1
Block Flush
3.0
58
41
15 to Column
8.0
59
64
18 to Waste
4.0
60
1
Block Flush
3.0
61
103
Wait
30.0
62
42
18 to Column
10.0
63
4
Flush to Waste
6.0
64
42
18 to Column
10.0
65
2
Reverse Flush
7.0
66
1
Block Flush
3.0
67
105
Start Detrityl
68
64
18 to Waste
4.0
69
42
18 to Column
10.0
70
2
Reverse Flush
7.0
71
1
Block Flush
3.0
72
112
Trityl Advance
73
109
Waste – Port
74
120
Advance FC
75
120
Advance FC
76
120
Advance FC
77
120
Advance FC
78
113
End Advance continued
3.15.18 Supplement 26
Current Protocols in Nucleic Acid Chemistry
Table 3.15.2 ABI 392 DNA Synthesizer Program for Oligomers with Alkaline-Labile Functional Groups, continued
Step
Number
Operation Name
Time (sec)
79
40
14 to Column
15.0
80
3
Trityl Flush
5.0
81
40
14 to Column
15.0
82
103
Wait
15.0
83
3
Trityl Flush
5.0
84
40
14 to Column
10.0
85
103
Wait
10.0
86
3
Trityl Flush
5.0
87
40
14 to Column
6.0
88
103
Wait
10.0
89
3
Trityl Flush
5.0
90
42
18 to Column
10.0
91
3
Trityl Flush
8.0
92
110
Waste – Bottle
93
42
18 to Column
8.0
94
2
Reverse Flush
7.0
95
1
Block Flush
4.0
96
107
End
COMMENTARY Background Information
The synthesis of N-unprotected 2 deoxynucleoside-3 -phosphoramidite building blocks was first reported as a four-step reaction with N-dimethylaminomethylene2 -deoxyribonucleosides (Zemlicka and Holy, 1967; Holy and Zemlicka, 1969; McBride et al., 1986) as the key intermediates (Gryaznov and Letsinger, 1991, 1992). The N-dimethylaminomethylene-2 -deoxyribonucleoside derivatives S.2a-c (Fig. 3.15.3) can be readily synthesized by the reaction of 2 -deoxyribonucleosides S.1a-c with dimethylformamide dimethyl acetal, and are converted to the corresponding 5 -Odimethoxytritylated compounds S.3a-c by the usual tritylation procedure. Treatment of the 5 -O-DMTr products S.3a-c with ammonia gives the N-unprotected species S.4a-c, which are phosphitylated to give the N-unprotected 2 -deoxynucleoside 3 -phosphoramidite building blocks S.5a-c. Although this four-step procedure is somewhat time consuming, it has proven to be the most reliable method for obtaining the pure building blocks. This
method is presented for the preparation of the N-unprotected 2 -deoxyguanosine derivative (S.5c) in Basic Protocol 3. The N-unprotected 5 -O-DMTr 2 -deoxynucleoside derivatives S.4a-c have also been synthesized by the direct introduction of the DMTr group onto the 5 -hydroxyl group of 2 -deoxyribonucleosides (Fig. 3.15.1). Ishido first reported this straightforward method to obtain S.4a and S.4b (Ishido, 1989; Nishino et al., 1991). The reaction of S.1a and S.1b with DMTr-Cl in pyridine in the presence of dichloroacetic acid gave S.4a and S.4b in good yields. These reactions have proven to be practical. Later, Kataoka and Hayakawa (1999) reported an alternative to this method using DMTr-Cl/triethylamine/imidazole. In this paper, O-selective direct tritylation of 2 deoxyguanosine was first reported. The reproducibility of this process, however, is very low. A four-step procedure involving the initial protection of the nucleobase with the dimethylaminomethylene protecting group is more practical for the synthesis of 5 -O-DMTr-2 deoxyguanosine, as shown in Figure 3.15.1
Synthesis of Unmodified Oligonucleotides
3.15.19 Current Protocols in Nucleic Acid Chemistry
Supplement 26
DNA Synthesis Without Base Protection Using Phosphoramidites
(Hayakawa and Kataoka, 1998). The 5 O-DMTr-2 -deoxyribonucleoside derivatives S.4a-c thus obtained are readily converted to the corresponding 3 -phosphoramidite derivatives. This process can be successfully carried out for all of the N-unprotected species. Basic Protocols 1 and 2 describe direct tritylation of 2 -deoxycytidine and 2 -deoxyadenosine using the method of Kataoka and Hayakawa, followed by phosphitylation to give the Nunprotected phosphoramidites S.5a-b. Quite recently, a new method for preparation of N-unprotected 5 -O-DMTr2 -deoxyribonucleoside 3 -phosphoramidites has been reported (Ohkubo et al., 2005b). It was found that first-generation Nprotected 2 -deoxyribonucleoside 3 -phosphoramidite derivatives such as N-benzoyl2 -deoxycytidine 3 -phosphoramidite (S.6a), N-acetyl-2 -deoxycytidine 3 -phosphoramidite (S.7a), N-benzoyl-2 -deoxyadenosine 3 phosphoramidite (S.6b), and N-isobutyryl-2 deoxyguanosine 3 -phosphoramidite (S.6c), as well as currently used reagents such as N-phenoxyacetyl-2 -deoxyadenosine 3 -phosphoramidite (S.8b) and N-isopropylphenoxylacetyl-2 -deoxyguanosine 3 -phosphoramidite (S.9c), could be converted to the N-unprotected 3 -phosphoramidite derivatives S.5a-c in high yields by treatment with ammonia in methanol (Fig. 3.15.2). These methods are presented as Alternate Protocols. The first-generation 3 -phosphoramidite reagents S.6a-c have not been used with DNA synthesis strategies developed recently to address the strong demand for shorter total reaction times. These approaches require the more base-labile N-acetylated (S.7a) and N-phenoxyacetylated (S.8b, S.9c) species. The classically N-protected 5 -O-DMTr-2 deoxyribonucleoside derivatives have been synthesized as key intermediates for the synthesis of phosphoramidites by wellestablished procedures, and the cost of these old reagents is rather cheap compared with the currently used PAC-phosphoramidite building blocks. Thereby, the total cost of the N-unprotected 2 -deoxyribonucleoside 3 phosphoramidite building blocks becomes even cheaper than that of the PAC phosphoramidite units. The successful deacylation of the first-generation phosphoramidite building blocks has provided a new potential use for N-unprotected 2 -deoxyribonucleoside 3 phosphoramidites as versatile intermediates in the construction of oligodeoxyribonucleotides with new functional groups. For example, the
reaction of S.5a with phenylisocyanate gives new N-modified building blocks that are useful for synthesis of oligodeoxyribonucleotides having aromatic rings as substituents on the nucleobase moiety (Ohkubo et al., 2005b). Since the reaction of the free amino group on cytosine with isocyanate derivatives proceeds smoothly without damage to other functional groups such as the phosphoramidite linkage, this reaction provides a powerful new tool for synthesizing a wide variety of modified DNA monomers.
Critical Parameters and Troubleshooting In Basic Protocols 1 to 3, for synthesis of N-unprotected phosphoramidites from unprotected nucleosides, the reactions should be carried out using DMTr-Cl or 2-cyanoethoxy(N,N-diisopropylamino)chlorophosphine derivatives under anhydrous conditions. In particular, the O-selective phosphitylation should be performed using the chlorophosphine (not 2-cyanoethoxy-bis(diisopropylamino)phosphine) at a low temperature to avoid N-phosphitylation of the nucleobases. The reactions are checked by TLC prespotted with one drop of triethylamine to prevent degradation of the target phosphoramidite units. After the reactions are completed, H2 O (not MeOH or EtOH) should be added to the reaction mixture to quench the chlorophosphine. In the Alternate Protocols, N-protected phosphoramidite compounds should be treated with methylamine or ammonia under anhydrous conditions to avoid the hydrolysis of the phosphoramidite residues.
Anticipated Results The N-unprotected DNA synthesis is very attractive for high-throughput synthesis of DNA molecules and synthesis of alkali-labile modified DNAs such as oligonucleotides containing DNA lesions, because this strategy allows the omission of the nucleobase deprotection step usually performed by treatment with concentrated NH3 at room temperature for 3 to 8 hr. In this approach, after the chain elongation is carried out using the four N-unprotected monomer building blocks on an ABI 392 DNA synthesizer used for synthesis of unmodified DNAs, the desired crude mixture is obtained by selective cleavage of the succinate linker from the polymer support using concentrated NH3 at room temperature only for 40 min. As an example, the anion-exchange HPLC profile of a crude mixture of the unmodified DNA
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Figure 3.15.4 Anion-exchange HPLC profiles of crude mixtures obtained using a 0.2 M HOtf Bt + BIT solution under conditions used for the ABI 392 DNA synthesizer. (A) d(TTAAAAATTATTAAATTATT). Anion-exchange HPLC was done on a Waters Alliance system with a Waters 3D UV detector and a Gen-Pak FAX column (Waters, 4.6 × 100 mm). A linear gradient (10% to 80%) of solvent I (1 M NaCl in 25 mM phosphate buffer, pH 6.0) in solvent II (25 mM phosphate buffer, pH 6.0) was used at 50◦ C at a flow rate of 1.0 mL/min for 45 min. (B) d(TACCTGGiBuGTCCAT). A linear gradient (10% to 60%) of solvent I (1 M NaCl in 25 mM phosphate buffer, pH 6.0) in solvent II (25 mM phosphate buffer, pH 6.0) was used at 50◦ C at a flow rate of 1.0 mL/min for 45 min.
oligomer d[TAAAAATTATTAAATTATT] is shown in Figure 3.15.4A. Moreover, a method has been established for synthesizing oligonucleotides incorporating alkali-labile modified bases such as 4-Nacetylcytosine or 2-N-isobutyrylguanosine by the combined use of the N-unprotected approach and a new silyl-type linker that can be cleaved under neutral conditions. After the final condensation is finished, the cyanoethyl groups are removed by treatment with 10% DBU in MeCN for 1 min. The crude oligodeoxyribonucleotide is released from the resin by treatment with a 1 M TBAF/AcOH in THF and then analyzed by HPLC. As an example, the anion-exchange HPLC profile of a crude mixture of the modified DNA oligomer d[TACCTGGiBu GTCCAT] is shown in Figure 3.15.4B. In the near feature, the present Nunprotected strategy may make it possible to avoid the considerable and undesired release of DNA probes synthesized in situ on glass plates, which currently requires a final deprotection step using concentrated NH3 .
Time Considerations The time needed to synthesize an Nunprotected phosphoramidite unit is estimated to be 2 to 4 days. All N-unprotected phosphoramidite units are stable at the purification step. The synthesis and purification of oligonucleotides using the activated phosphite method
can be carried out in 2 to 4 days. The time needed for the Alternate Protocols is estimated to be 1 to 2 days.
Literature Cited Gao, X., Gulari, E., and Zhou, X. 2004. In situ synthesis of oligonucleotide microarrays. Biopolymers 73:579-596. Gryaznov, S.M. and Letsinger, R.L. 1991. Synthesis of oligonucleotides via monomers with unprotected bases. J. Am. Chem. Soc. 113:58765877. Gryaznov, S.M. and Letsinger, R.L. 1992. Selective O-phosphitylation with nucleoside phosphoramidite reagents. Nucl. Acids Res. 20:18791882. Hayakawa, Y., and Kataoka, M. 1998. Facile synthesis of oligodeoxyribonucleotides via the phosphoramidite method without nucleoside base protection. J. Am. Chem. Soc. 120:1239512401. Hayakawa, Y., Kataoka, M., and Noyori, R. 1996. Benzimidazolium triflate as an efficient promoter for nucleotide synthesis via the phosphoramidite method. J. Org. Chem. 61:79967997. Holy, A. and Zemlicka, J. 1969. Oligonucleotidic compounds. XXXIII. Hydrolysis of N-dimethyl-aminomethylenecytidine (DMC), N-dimethylaminomethyleneadenosine (DMA), and N-dimethylaminomethyleneguanosine (DMG) and related 2 deoxy compounds. Collect. Czech. Chem. Commun. 34:2449-2458. Ishido, R. 1989. Protection of 5 -hydroxy groups of cytidine derivatives. Jpn. Kokai Tokkyo Koho Jp Pat. 01308294.
Synthesis of Unmodified Oligonucleotides
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Kataoka, M. and Hayakawa, Y. 1999. A convenient method for the synthesis of N-free 5 -O-(p,p dimethoxytrityl)-2 -deoxyribonucleosides via 5 -O-selective tritylation of the parent substances. J. Org. Chem. 64:6087-6089. Kobori, A., Miyata, K., Ushioda, M., Seio, K., and Sekine, M. 2002. A new method for the synthesis of oligodeoxyribonucleotides containing 4N-alkoxycarbonyldeoxycytidine derivatives and their hybridization properties. J. Org. Chem. 67:476-485. McBride, L.J., Kierzek, R., Beaucage, S.L., and Caruthers, M.H. 1986. Nucleotide chemistry. 16. Amidine protecting groups for oligonucleotide synthesis. J. Am. Chem. Soc. 108:20402048. Nagai, H., Fujiwara, T., Fujii, M., Sekine, M., and Hata, T. 1989. Reinvestigation of deoxyribonucleoside phosphorothioites: Synthesis and properties of deoxyribonucleoside-3 dimethyl phosphites. Nucl. Acids Res. 17:8581-8593. Nishino, S., Nagano, Y., Hasegawa, Y., Yamamoto, H., Kamaike, K., and Ishido, Y. 1991. Efficient deanilidation of phosphoranilidates by the use of nitrites and acetic anhydride. Heteroatom Chem. 2:187-196. Ohkubo, A., Ezawa, Y., Seio, K., and Sekine, M. 2004. O-selectivity and utility of phosphorylation mediated by phosphite triester intermediates in the N-unprotected phosphoramidite method. J. Am. Chem. Soc. 126:10884-10896. Ohkubo, A., Aoki, K., Ezawa, Y., Sato, Y., Taguchi, H., Seio, K., and Sekine, M. 2005a. Synthesis of oligodeoxyribonucleotides containing hydroxymethylphosphonate bonds in the phosphoramidite method and their hybridization properties. Tetrahedron Lett. 46:8953-8957.
Ohkubo, A., Sakamoto, K., Miyata, K., Taguchi, H., Seio, K., and Sekine, M. 2005b. Convenient synthesis of N-unprotected deoxynucleoside 3 -phosphoramidite building blocks by selective deacylation of N-acylated species and their facile conversion to other N-functionalized derivatives. Org. Lett. 7:5389-5392. Sekine, M., Ohkubo, A., and Seio, K. 2003. Protonblock strategy for the synthesis of oligodeoxynucleotides without base protection, capping reaction, and P-N bond cleavage reaction. J. Org. Chem. 68:5478-5492. Shena, M. 1999. DNA Microarrays: A Practical Approach. Oxford University Press, London. Tanimura, H., Maeda, M., Fukuzawa, T., Sekine, M., and Hata, T. 1989. Chemical synthesis of the 24 RNA fragments corresponding to hop stunt viroid. Nucl. Acids Res. 17:8135-8147. Wada, T., Kobori, A., Kawahara, S., and Sekine, M. 2001. Synthesis and hybridization ability of oligodeoxyribonucleotides incorporating N-acyldeoxycytidine derivatives. Eur. J. Org. Chem. 4583-4593. Zemlicka, J. and Holy, A. 1967. Preparation of Ndimethylaminomethylene derivatives as a new method of a selective substitution of nucleoside amino groups. Collect. Czech. Chem. Commun. 32:3159-3168.
Contributed by Akihiro Ohkubo, Kohji Seio, and Mitsuo Sekine Tokyo Institute of Technology Tokyo, Japan
DNA Synthesis Without Base Protection Using Phosphoramidites
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A Universal and Recyclable Solid Support for Oligonucleotide Synthesis
UNIT 3.16
Franc¸ois Morvan,1 Albert Meyer,1 and Jean-Jacques Vasseur1 1
Universit´e Montpellier, Montpellier, France
ABSTRACT This unit provides a modified phosphoramidite method to synthesize oligodeoxyribonucleotides onto a universal and reusable hydroxyl solid support thanks to the use of deoxyribonucleoside tert-butyl and cyanoethyl phosphoramidites. The nucleoside tert-butyl phosphoramidite allows the introduction of an H-phosphonate diester linkage using the phosphoramidite method. After elongation, the H-phosphonate diester linker is cleaved by transesterification under mild basic conditions to yield an oligonucleotide with free 3 and 5 -hydroxyls and the starting solid support. Thus, the solid support is easily recycled and used for a subsequent synthesis. In addition, a nucleoside tert-butyl phosphoramidite could be introduced inside the oligonucleotide chain during the elongation to yield a second H-phosphonate diester linkage. After elongation, the two H-phosphonate diester linkages are cleaved, producing two oligonucleotides with free 3 - and 5 -hydroxyls. C 2007 by John Wiley & Sons, Inc. Curr. Protoc. Nucleic Acid Chem. 30:3.16.1-3.16.19. Keywords: phosphoramidite r H-phosphonate r automatized synthesis r tandem synthesis
INTRODUCTION This unit describes a modified phosphoramidite method to synthesize oligodeoxyribonucleotides onto a universal and reusable hydroxylated solid support through the use of deoxyribonucleoside tert-butyl- and 2-cyanoethyl-N,N-diisopropylphosphoramidites (Ferreira et al., 2005). Deoxyribonucleoside tert-butyl phosphoramidites are obtained from commercially available 5 -O, N-protected deoxyribonucleosides using tert-butyl tetraisopropylphosphorodiamidite. The coupling of a deoxyribonucleoside tert-butyl phosphoramidite with the hydroxylated solid support immediately followed by the detritylation step, without performing the oxidation step of the synthesis cycle, produces an H-phosphonate diester linkage. Then, standard deoxyribonucleoside 2cyanoethyl phosphoramidites are incorporated according to the phosphoramidite method (Beaucage and Caruthers, 1981; UNIT 3.3) with a few modifications that maintain the Hphosphonate diester linkage throughout the elongation of the oligonucleotide. Oxidations are done with tert-butyl hydroperoxide (Hayakawa et al., 1986) or 3H-1,2-benzodithiole3-one 1,1-dioxide (Iyer et al., 1990) to afford the phosphotriester or thionophosphorotriester, respectively, since these oxidizing reagents do not oxidize H-phosphonate diester linkages (Jung et al., 1994; Bologna et al., 1999). The capping step is performed with a phosphoramidite rather than the usual acetic anhydride to avoid P-acylation of the H-phosphonate linkage (UNIT 3.4), and dichloroacetic acid (DCA) is preferred for the detritylation step. At the end of chain elongation, a treatment with potassium carbonate in methanol releases the 3 -OH oligonucleotide from the starting hydroxylated solid support, which can be recycled and reused several times. Additionally, the nucleoside tert-butyl phosphoramidite can be introduced inside the oligonucleotide chain during elongation to yield a second H-phosphonate diester linkage and enable tandem oligonucleotide synthesis.
Current Protocols in Nucleic Acid Chemistry 3.16.1-3.16.19, September 2007 Published online September 2007 in Wiley Interscience (www.interscience.wiley.com). DOI: 10.1002/0471142700.nc0316s30 C 2007 John Wiley & Sons, Inc. Copyright
Synthesis of Unmodified Oligonucleotides
3.16.1 Supplement 30
CAUTION: All chemicals must be used in a chemical fume hood by qualified persons equipped with laboratory coats, safety glasses, and gloves. BASIC PROTOCOL 1
PREPARATION OF DEOXYRIBONUCLEOSIDE tert-BUTYL PHOSPHORAMIDITES Conversion of 5 -O- and N-protected deoxyribonucleosides into the corresponding tertbutyl phosphoramidites using tert-butyl tetraisopropylphosphorodiamidite is shown in Figure 3.16.1. Starting material, solvents, and glassware must be as dry as possible to obtain high yields.
Materials 5 -O-DMTr-N-protected nucleoside (Samchully Pharmaceuticals): 5 -O-(4,4 -Dimethoxytrityl)-N6 -benzoyl-2 -deoxyadenosine (S.1a) 5 -O-(4,4 -Dimethoxytrityl)-N4 -benzoyl-2 -deoxycytidine (S.1b) 5 -O-(4,4 -Dimethoxytrityl)-N2 -isobutyryl-2 -deoxyguanosine (S.1c) 5 -O-(4,4 -Dimethoxytrityl)-2 -deoxythymine (S.1d) Diisopropylammonium tetrazolide Acetonitrile (dried over CaH2 and distilled) Anhydrous methylene chloride (CH2 Cl2 ; freshly distilled over P2 O5 ) tert-Butyl tetraisopropylphosphorodiamidite (Aldrich) Argon Molybde stain (see recipe), optional Ethyl acetate Saturated aqueous sodium chloride solution (brine) Anhydrous sodium sulfate Cyclohexane Triethylamine (Et3 N) Silica gel (0.04 to 0.06 mm) Dioxane 50-mL round-bottom flask equipped with stir bar and CaCl2 guard TLC silica plates (Kieselgel 60 F-254; Merck) 254-nm UV lamp 250-mL separatory funnel 5-cm-diameter chromatography column
Universal and Recyclable Solid Support
Figure 3.16.1 Synthesis of the four protected deoxyribonucleoside tert-butyl phosphoramidites (S.2a-d). DMTr, dimethoxytrityl.
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Vacuum evaporator Lyophilizer Additional reagents and equipment thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare deoxyribonucleoside tert-butyl phosphoramidites 1. In a 50-mL round-bottom flask, coevaporate 1.84 mmol of 5 -O-DMTr-N-protected nucleoside (S.1a-d) and 0.157 g (0.92 mmol) diisopropylammonium tetrazolide three times, each with 10 mL dry acetonitrile. Dissolve in 15 mL anhydrous methylene chloride. 2. While stirring, add 0.671 g (2.20 mmol) of tert-butyl tetraisopropylphosphorodiamidite under an argon atmosphere. 3. Stir the mixture overnight. Monitor by TLC. Before spotting the reaction mixture, TLC plates must be neutralized by dipping in the eluting solution and air-dried. After elution, direct visualization using a 254-nm UV lamp reveals the starting and final compounds. In addition, staining the TLC plate with molybde stain solution reveals phosphorous compounds (i.e., starting phosphine and resulting phosphoramidite) as blue spots. The plate is dipped in the molybde solution, washed with water, stamped with absorbent paper to remove excess water, and then slowly heated to dry the plate. For A and T: 2:7:1 (v/v/v) cyclohexane/EtOAc/Et3 N; Rf1a = 0.05; Rf2a = 0.5; Rf1d = 0.1; Rf2d = 0.7. For C: 5:4:1 (v/v/v) cyclohexane/CH2 Cl2 /Et3 N; Rf1b = 0.1; Rf2b = 0.6. For G: 2:7:1 (v/v/v) CH3 CN/CH2 Cl2 /Et3 N; Rf1c = 0.3; Rf2c = 0.6.
4. Dilute the reaction mixture with 50 mL ethyl acetate, pour into a 250-mL separatory funnel, and wash two times with 100 mL brine. 5. Collect the organic layer, dry it by adding ∼10 g solid anhydrous sodium sulfate, filter on cotton in a funnel, and concentrate on a rotary evaporator under reduced pressure to obtain a foam.
Purify by flash column chromatography 6. Dissolve the crude S.2a, S.2b, or S.2d in a minimal volume of 75:20:5 (v/v/v) cyclohexane/CH2 Cl2 /Et3 N and apply the solution to a 5-cm-diameter chromatography column containing 40 g silica gel (0.04 to 0.06 mm) equilibrated in the same solvent. For S.2c, use 25:75:5 (v/v/v) cyclohexane/CH2 Cl2 /Et3 N to equilibrate the column and dissolve the crude product. It is compulsory to have triethylamine in the elution solvent to avoid activation (and degradation) of the phosphoramidite due to the acidity of silica.
7. Gradually increase the concentration of methylene chloride in the eluent, still containing 5% of Et3 N. 8. Monitor fractions by TLC and combine those containing the pure compound. 9. Evaporate to dryness on a vacuum evaporator, and coevaporate with 20 mL dioxane. 10. Dissolve the foam in 30 mL dioxane, freeze the solution as a film in the flask, and lyophilize to afford the resulting phosphoramidite as a white powder. The phosphoramidites S.2a-d can be stored for several months at −20◦ C without decomposition.
Synthesis of Unmodified Oligonucleotides
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11. Check phosphoramidite purity by 31 P NMR. Chemical shifts of the tert-butyl-N,N-diisopropylphosphoramidites are near 140 ppm (doublet) instead of 150 ppm for conventional phosphoramidites. Likewise, the chemical shift of hydrolyzed phosphine (tert-butyl H-phosphonate) when present is near 7 ppm (singlet) instead of 14 ppm for non-substituted alkyl H-phosphonates. 5 -O-(4,4 -Dimethoxytrityl)-N6 -benzoyl-2 -deoxyadenosine 3 -O-(tert-butyl)-N,N-diisopropylphosphoramidite (S.2a): yield 77%. TLC (cyclohexane/EtOAc/Et3 N 2/7/1 v/v/v) Rf : 0.50. 31 P NMR (CD3 CN): 139.3 and 139.5 ppm. 1 H NMR (CD3 CN): 1.08-1.36 (m, 21H), 2.52-2.69 (m, 1H), 2.99-3.14 (m, 1H), 3.27-3.41 (m, 2H), 3.56-3.69 (m, 2H), 3.75 (s, 6H), 4.21- 4.24 (m, 1H), 4.72-4.89 (m, 1H), 6.43-6.49 (m, 1H), 6.75-6.84 (m, 4H), 7.17-7.64 (m, 12H), 8.01 (m, 2H), 8.28 (d, J = 2.14 Hz, 1H), 8.59 (d, J = 1.35 Hz, 1H), 9.62 (br s, 1H). 5 -O-(4,4 -Dimethoxytrityl)-N6 -benzoyl-2 -deoxycytidine 3 -O-(tert-butyl)-N,N-diisopropylphosphoramidite (S.2b): yield 65%, TLC (cyclohexane/CH2 Cl2 /Et3 N 5/4/1 v/v/v) Rf : 0.60. 31 P NMR (CD3 CN): 139.7 and 139.6 ppm. 1 H NMR (CD3 CN): 1.07-1.34 (m, 21H), 2.25-2.36 (m, 1H), 2.53-2.62 (m, 1H), 3.39-3.46 (m, 2H), 3.56-3.67 (m, 2H), 3.79 (s, 6H), 4.16- 4.17 (m, 1H), 4.46-4.58 (m, 1H), 6.14-6.18 (m, 1H), 6.89-6.91 (m, 5H), 7.18-7.67 (m, 12H), 7.9 (pd, J = 8.6 Hz, 2H), 8.23 (d, J = 8.4 Hz, 1H), 8.34 (d, J = 8.6 Hz, 1H), 9.5 (br s, 1 H). 5-O-(4,4 -Dimethoxytrityl)-N2 -isobutyryl-2-deoxyguanosine 3 -O-(tert-butyl)-N,N-diisopropylphosphoramidite (S.2c): yield 79%. TLC (CH3 CN/CH2 Cl2 /Et3 N 2/7/1 v/v/v) Rf : 0.60. 31 P NMR (CD3 CN): 140.0 and 139.4 ppm. 1 H NMR (CD3 CN): 1.03-1.33 (m, 27H), 2.55-2.71 (m, 2H), 2.86-2.92 (m, 1H), 3.23-3.38 (m, 2H), 3.54-3.66 (m, 2H), 3.75 (s, 6H), 4.18- 4.21 (m, 1H), 4.52-4.99 (m, 1H), 6.23-6.26 (m, 1H), 6.75-6.82 (m, 4H), 7.19-7.42 (m, 9H), 7.85 (1d, 1H). 5 -O-(4,4 -Dimethoxytrityl)thymidine 3 -O-(tert-butyl)-N,N-diisopropylphosphoramidite (S.2d): yield 86%. TLC (cyclohexane/EtOAc/Et3 N 2/7/1 v/v/v) Rf : 0.70. 31 P NMR (CD3 CN): 139.8 and 139.0 ppm. 1 H NMR (CD3 CN): 1.06-1.52 (m, 21H), 2.21 (s, 3H), 2.34-2.42 (m, 2H), 3.31-3.38 (m, 2H), 3.55-3.67 (m, 2H), 3.8 (s, 6H), 4.09- 4.11 (m, 1H), 4.51-4.59 (m, 1H), 6.25-6.32 (m, 1H), 6.88-6.92 (m, 4H), 7.27-7.52 (m, 10H), 9.10 (br s, 1H). BASIC PROTOCOL 2
PREPARATION OF THE HEXANOL-LCAA-CPG SOLID SUPPORT The universal and reusable solid support S.5 is prepared from ◦commercially available long-chain alkylamine controlled-pore glass (LCAA-CPG; 500 A), which is converted ◦to a hydroxylated solid support (Fig. 3.16.2). A larger pore size (i.e., 1000 or 2000 A) is required for the synthesis of longer oligodeoxyribonucleotides (>50 mers). The hexanol-functionalized support is prepared from the LCAA-CPG according to the protocol described below. The loading of the alcohol function borne by the CPG solid support is determined by a trityl assay after coupling with a commercial deoxyribonucleoside 2-cyanoethyl phosphoramidite using a standard elongation cycle. The preparation of this universal solid support is easily scaled up to tens of grams, as long as there is efficient shaking during all the steps.
Materials ◦
Universal and Recyclable Solid Support
Long-chain alkylamine controlled-pore glass (LCAA-CPG, 500 A, 80 to 120 mesh, amino group, 80 to 90 µmol/g) 3% trichloroacetic acid (TCA) in methylene chloride (commercial deblocking solution) Triethylamine (Et3 N) Diisopropylethylamine (DIPEA) Anhydrous methylene chloride (CH2 Cl2 )
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Diethyl ether Phosphorous pentoxide (P2 O5 ) Anhydrous pyridine Succinic anhydride 4-Dimethylaminopyridine (DMAP) N-(3-Dimethylaminopropyl)-N -ethylcarbodiimide (DEC) N-Hydroxysuccinimide 6-Aminohexan-1-ol Anhydrous piperidine Methanol Deoxyribonucleoside phosphoramidite 0.1 M p-toluenesulfonic acid solution in acetonitrile 25-mL round-bottom flasks Frits (porosity 3) Vacuum desiccator DNA synthesis column 10-mL volumetric flasks Spectrophotometer Additional reagents and equipment for automated oligonucleotide synthesis and the trityl assay (APPENDIX 3C)
Figure 3.16.2 Preparation of the universal and reusable hexanol-LCA-CPG solid support (S.5). DEC, N-(3-dimethylaminopropyl)-N -ethylcarbodiimide; DMAP, 4-(dimethylamino)pyridine.
Synthesis of Unmodified Oligonucleotides
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Functionalize LCAA-CPG 1. In a 25-mL round-bottom flask, combine 1.0 g of LCAA-CPG and 5 mL of 3% trichloroacetic acid (TCA) in methylene chloride (commercial deblocking solution). Shake gently for 4 hr at room temperature. This step is performed to convert the resulting ammonium function, due to the acidic treatment, to the amino function, thereby activating the solid support.
2. Filter off the LCAA-CPG on a frit, and wash successively with 10 mL of 9:1 (v/v) Et3 N/DIPEA, 10 mL CH2 Cl2 , and 10 mL diethyl ether. 3. Dry in a vacuum desiccator over P2 O5 for 30 min at room temperature. 4. In a 25-mL round-bottom flask, combine successively 1.0 g of the activated LCAACPG, 6 mL anhydrous pyridine, 200 mg (2 mmol) succinic anhydride, and 40 mg (0.33 mmol) DMAP. Shake gently overnight at room temperature. 5. Filter off on a frit and wash with 3 mL pyridine followed by 5 mL CH2 Cl2 . 6. Dry in a vacuum desiccator over P2 O5 for 30 min at room temperature. 7. In a 25-mL round-bottom flask, combine successively the succinylated CPG (S.3), 5 mL CH2 Cl2 , 0.191 g (1 mmol) DEC, 60 mg (0.5 mmol) N-hydroxysuccinimide, and 40 µL Et3 N. Shake for 4 hr at room temperature. 8. Add 6 mg (0.05 mmol) of 6-aminohexan-1-ol and shake overnight at room temperature. 9. Add 5 mL piperidine and shake 15 min at room temperature. 10. Filter off on a frit, and wash successively with 3 mL pyridine, 5 mL methanol, and 5 mL CH2 Cl2 . 11. Dry the resulting hexanol-LCAA-CPG (S.5) in a vacuum desiccator over P2 O5 for 30 min at room temperature.
Determine loading 12. Weigh 20 mg of S.5 in an empty DNA synthesis column. 13. Couple a deoxyribonucleoside phosphoramidite to S.5 according to a standard elongation cycle in trityl-OFF mode, but omit the detritylation step that is usually performed before coupling. The first detritylation step actually can be performed, although the solid support has no trityl group.
14. Transfer the DMTr cation into a 10-mL volumetric flask and fill it up to 10 mL with a 0.1 M p-toluenesulfonic acid solution in acetonitrile. 15. Transfer 1 mL of the resulting solution to a 10-mL volumetric flask and add 9 mL of 0.1 M p-toluenesulfonic acid solution in acetonitrile. 16. Read the absorption at 498 nm (A498 ). 17. Calculate loading in micromole per gram CPG (µmol/g CPG) according to the following formula:
Universal and Recyclable Solid Support
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where m is mass of S.5 (step 1), 70,000 is the extinction coefficient of the cation DMTr, and the two 10 values correspond to the total volume of the solution (step 14) and the factor of dilution (step 15). Loading of 6-aminohexan-1-ol is typically between 40 and 50 µmol/g CPG.
OLIGONUCLEOTIDE SYNTHESIS This protocol describes the synthesis of oligonucleotides with phosphodiester or phosphorothioate linkages starting from the universal hexanol solid support S.5 and the deoxyribonucleoside tert-butyl phosphoramidite building blocks S.2a-d, and then using commercial nucleoside phosphoramidites with either conventional or ultra-fast-deprotecting nucleobase-protecting groups. Alternate Protocols 1 and 2 describe tandem synthesis to yield two oligonucleotides per synthesis.
BASIC PROTOCOL 3
The synthesis of oligodeoxyribonucleotides is performed using an ABI 381A DNA synthesizer according to the customized elongation cycle shown in Table 3.16.1 (Fig. 3.16.3). Since the solid support S.5 has no DMTr group, the first detritylation step can be omitted. Detritylation is performed with 3% DCA in CH2 Cl2 , and activation of deoxyribonucleoside phosphoramidites is effected by 5-(benzylthio)-1H-tetrazole (BMT, 0.3 M in dry acetonitrile). Capping is performed with BMT-activated bis(2-cyanoethyl)-N,Ndiisopropylphosphoramidite (0.03 M in dry acetonitrile) for 15 sec. Oxidation is performed with 1.1 M tert-butyl hydroperoxide in CH2 Cl2 or 0.05 M 3H-1,2-benzodithiol3-one 1,1-dioxide in dry acetonitrile for 60 sec. The average coupling yields per cycle are typically between 98% and 99%, as determined by the DMTr cation assay (UNIT 10.3 and APPENDIX 3C). After oligonucleotide chain assembly, the column is treated with a 0.01 M potassium carbonate solution in methanol for 10 min to release the oligonucleotide from the solid support S.5, which is then recycled.
Table 3.16.1 Customized Oligonucleotide Elongation Cycle for Deoxyribonucleoside 3 -O-(tertButyl)- and 3 -O-(2-Cyanoethyl)-N,N-diisopropylphosphoramidite Building Blocksa
Step
Reagents and solvents
Time (sec)
For deoxyribonucleoside 3 -O-(tert-butyl)-N,N-diisopropylphosphoramidites Wash
CH3 CN
10
Detritylation
3% DCA in CH2 Cl2
60 15
Wash
CH3 CN
Coupling
0.09 M tert-butyl amidite S.2a-d in CH3 CN + 0.3 M BMT in CH3 CN 45
Wash
CH3 CN
20
Detritylation
3% DCA in CH2 Cl2
60
Wash
CH3 CN
15
For deoxyribonucleoside 3 -O-(2-cyanoethyl)-N,N-diisopropylphosphoramidites Coupling
0.09 M 2-cyanoethyl amidite in CH3 CN + 0.3 M BMT in CH3 CN
Cappinga
0.03 M i-Pr2 NP(OCH2 CH2 CN)2 in CH3 CN + 0.3M BMT in CH3 CN 15
Oxidation
1.1 M tBuOOH in CH2 Cl2 or 0.05 M Beaucage reagent in CH3 CN
60
Wash
CH3 CN
15
Detritylation
3% DCA in CH2 Cl2
60
Wash
CH3 CN
15
15
Synthesis of Unmodified Oligonucleotides
a Capping step is optional.
3.16.7 Current Protocols in Nucleic Acid Chemistry
Supplement 30
Materials 5-Benzylthio-1H-tetrazole (BMT) Anhydrous acetonitrile (<10 ppm H2 O) Anhydrous tert-butyl hydroperoxide (tBuOOH; 5.5 M in decane; Fluka) Anhydrous methylene chloride (CH2 Cl2 ; freshly distilled over P2 O5 ) 3H-1,2-Benzodithiole-3-one 1,1-dioxide Bis(2-cyanoethyl)-N,N-diisopropylphosphoramidite 5 -O-(4,4 -Dimethoxytrityl)-N-protected-2 -deoxynucleoside 3 -O-(tert-butyl)-N,N-diisopropylphosphoramidite (S.2a-d; see Basic Protocol 1) Standard 3 -O-(2-cyanoethyl)-N,N-diisopropylphosphoramidites of 5 -O-(4,4 -dimethoxytrityl)-N-protected-2 -deoxyribonucleosides: 5 -O-DMTr-N6 -benzoyl-2 -deoxyadenosine (Pierce) 5 -O-DMTr-N4 -benzoyl-2 -deoxycytidine (Pierce) 5 -O-DMTr-N2 -isobutyryl-2 -deoxyguanosine (Pierce) 5 -O-DMTr-2 -deoxythymine (Pierce) 5 -O-DMTr-N6 -phenoxyacetyl-2 -deoxyadenosine (Glen Research) 5 -O-DMTr-N4 -acetyl-2 -deoxycytidine (Glen Research) 5 -O-DMTr-N2 -(isopropylphenoxyacetyl)-2 -deoxyguanosine (Glen Research) Hexanol-LCAA-CPG solid support (S.5; see Basic Protocol 2) 3% Dichloracetic acid (DCA) in CH2 Cl2 (Biosolve) 10 mM potassium carbonate (K2 CO3 ) in methanol (70 mg in 50 mL) Concentrated aqueous ammonia Methanol Phosphorous pentoxide (P2 O5 ) ABI 381A DNA synthesizer 1-mL syringes 2-mL microcentrifuge tubes Speed vacuum Sealed vial (vial with Teflon septum and screw top for HPLC sample preparations) 55◦ C dry bath Vacuum desiccator Additional reagents and equipment for automated oligonucleotide synthesis and the trityl assay (APPENDIX 3C) Synthesize oligonucleotide 1. Program the modified elongation cycle for an ABI 381A DNA synthesizer (Table 3.16.2). Create the user function USER F: 4,6,12,16,20,23 corresponding to Tet +12 TO COLM used for the capping step. 2. Prepare a 0.3 M solution of 5-benzylthio-1H-tetrazole in anhydrous acetonitrile (5.77 g in 100 mL). Place the bottle in position no. 9 using the change bottle procedure (see Table 3.16.3 for bottle positions). Place a solution of 3% DCA in CH2 Cl2 in position 14 for the detritylation step. 3. Prepare oxidizing solution and place the bottle in position no. 15 using the change bottle procedure. a. For phosphodiester oligonucleotides: Prepare a 1.1 M tert-butyl hydroperoxide solution by mixing 5 mL of anhydrous tert-butyl hydroperoxide (5.5 M in decane) with 20 mL methylene chloride. b. For phosphorothioate oligonucleotides: Prepare a 0.05 M solution of 3H-1, 2-benzodithiol-3-one 1,1-dioxide in dry acetonitrile (100 mg in 10 mL). Universal and Recyclable Solid Support
3.16.8
4. Prepare a 0.03 M solution of bis(2-cyanoethyl)-N,N-diisopropylphosphoramidite in anhydrous acetonitrile (81.3 mg in 10 mL). Place the bottle in position no. 12 using the change bottle procedure.
Supplement 30
Current Protocols in Nucleic Acid Chemistry
Figure 3.16.3 Oligonucleotide synthesis on the universal hydroxylated solid support. BMT, 5(benzylthio)-1H-tetrazole; CE, cyanoethyl; DCA, dichloroacetic acid; DMTr, dimethoxytrityl.
5. Prepare a 0.09 M solution (18 molar eq.) of S.2a, b, c, or d in dry acetonitrile and place the bottle in position no. 5 using the change bottle procedure.
Synthesis of Unmodified Oligonucleotides
3.16.9 Current Protocols in Nucleic Acid Chemistry
Supplement 30
Table 3.16.2 Customized Phosphoramidite Elongation Cycle for 381A ABI DNA Synthesizer
Active for bases Step no. Function no. Name
Universal and Recyclable Solid Support
Step time A
G
C
T
X
1
10
#18 TO WASTE
2
Yes
Yes
Yes
Yes
Yes
2
9
#18 TO COLM
10
Yes
Yes
Yes
Yes
Yes
3
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
Yes
4
1
BLOCK FLUSH
3
Yes
Yes
Yes
Yes
Yes
5
28
PHOS PREP
3
Yes
Yes
Yes
Yes
Yes
6
90
TET TO COLM
3
Yes
Yes
Yes
Yes
Yes
7
19
B + TET TO COLM
3
Yes
Yes
Yes
Yes
Yes
8
90
TET TO COLM
3
Yes
Yes
Yes
Yes
Yes
9
19
B + TET TO COLM
3
Yes
Yes
Yes
Yes
Yes
10
90
TET TO COLM
3
Yes
Yes
Yes
Yes
Yes
11
19
B + TET TO COLM
3
Yes
Yes
Yes
Yes
Yes
12
90
TET TO COLM
3
Yes
Yes
Yes
Yes
Yes
13
1
BLOCK FLUSH
3
Yes
Yes
Yes
Yes
Yes
14
10
#18 TO WASTE
2
Yes
Yes
Yes
Yes
Yes
15
4
WAIT
15
Yes
Yes
Yes
Yes
Yes
16
4
WAIT
30
No
No
No
No
Yes
17
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
Yes
18
1
BLOCK FLUSH
3
Yes
Yes
Yes
Yes
Yes
19
9
#18 TO COLM
10
Yes
Yes
Yes
Yes
Yes
20
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
Yes
21
9
#18 TO COLM
10
Yes
Yes
Yes
Yes
Yes
22
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
Yes
23
1
BLOCK FLUSH
3
Yes
Yes
Yes
Yes
Yes
24
28
PHOS PREP
3
Yes
Yes
Yes
Yes
No
25
90
TET TO COLM
3
Yes
Yes
Yes
Yes
No
a
26
97
USER F
3
Yes
Yes
Yes
Yes
No
27
90
TET TO COLM
3
Yes
Yes
Yes
Yes
No
a
28
97
USER F
3
Yes
Yes
Yes
Yes
No
29
90
TET TO COLM
3
Yes
Yes
Yes
Yes
No
a
30
97
USER F
3
Yes
Yes
Yes
Yes
No
31
90
TET TO COLM
3
Yes
Yes
Yes
Yes
No
32
4
WAIT
20
Yes
Yes
Yes
Yes
No
33
1
BLOCK FLUSH
3
Yes
Yes
Yes
Yes
No
34
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
No
35
10
#18 TO WASTE
2
Yes
Yes
Yes
Yes
No
36
9
#18 TO COLM
10
Yes
Yes
Yes
Yes
No
37
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
No
38
9
#18 TO COLM
10
Yes
Yes
Yes
Yes
No continued
3.16.10 Supplement 30
Current Protocols in Nucleic Acid Chemistry
Table 3.16.2 Customized Phosphoramidite Elongation Cycle for 381A ABI DNA Synthesizer, continued
Active for bases Step no. Function no. Name
Step time A
G
C
T
X
39
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
No
40
9
#18 TO COLM
10
Yes
Yes
Yes
Yes
No
41
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
No
42
1
BLOCK FLUSH
3
Yes
Yes
Yes
Yes
No
43
81
#15 TO WASTE
3
Yes
Yes
Yes
Yes
No
44
13
#15 TO COLM
10
Yes
Yes
Yes
Yes
No
45
10
#18 TO WASTE
2
Yes
Yes
Yes
Yes
No
46
4
WAIT
45
Yes
Yes
Yes
Yes
No
47
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
No
48
1
BLOCK FLUSH
3
Yes
Yes
Yes
Yes
No
49
10
#18 TO WASTE
2
Yes
Yes
Yes
Yes
No
50
9
#18 TO COLM
10
Yes
Yes
Yes
Yes
No
51
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
No
52
9
#18 TO COLM
10
Yes
Yes
Yes
Yes
No
53
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
No
54
9
#18 TO COLM
10
Yes
Yes
Yes
Yes
Yes
55
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
Yes
56
1
BLOCK FLUSH
3
Yes
Yes
Yes
Yes
Yes
57
33
CYC ENTRY
1
Yes
Yes
Yes
Yes
Yes
58
10
#18 TO WASTE
2
Yes
Yes
Yes
Yes
Yes
59
9
#18 TO COLM
10
Yes
Yes
Yes
Yes
Yes
60
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
Yes
61
1
BLOCK FLUSH
3
Yes
Yes
Yes
Yes
Yes
62
5
ADVANCE FC
1
Yes
Yes
Yes
Yes
Yes
63
6
WASTE PORT
1
Yes
Yes
Yes
Yes
Yes
64
82
#14 TO WASTE
3
Yes
Yes
Yes
Yes
Yes
65
14
#14 TO COLM
10
Yes
Yes
Yes
Yes
Yes
66
108
FLUSH TO TRIT
1
Yes
Yes
Yes
Yes
Yes
67
14
#14 TO COLM
10
Yes
Yes
Yes
Yes
Yes
68
108
FLUSH TO TRIT
1
Yes
Yes
Yes
Yes
Yes
69
14
#14 TO COLM
10
Yes
Yes
Yes
Yes
Yes
70
108
FLUSH TO TRIT
1
Yes
Yes
Yes
Yes
Yes
71
14
#14 TO COLM
10
Yes
Yes
Yes
Yes
Yes
72
108
FLUSH TO TRIT
1
Yes
Yes
Yes
Yes
Yes
73
14
#14 TO COLM
10
Yes
Yes
Yes
Yes
Yes
74
108
FLUSH TO TRIT
1
Yes
Yes
Yes
Yes
Yes
75
14
#14 TO COLM
10
Yes
Yes
Yes
Yes
Yes continued
Synthesis of Unmodified Oligonucleotides
3.16.11 Current Protocols in Nucleic Acid Chemistry
Supplement 30
Table 3.16.2 Customized Phosphoramidite Elongation Cycle for 381A ABI DNA Synthesizer, continued
Active for bases Step no. Function no. Name 76
108
77
9
78
Step time A
G
C
T
X
FLUSH TO TRIT
1
Yes
Yes
Yes
Yes
Yes
#18 TO COLM
10
Yes
Yes
Yes
Yes
Yes
108
FLUSH TO TRIT
1
Yes
Yes
Yes
Yes
Yes
79
7
WASTE BOTTLE
1
Yes
Yes
Yes
Yes
Yes
80
1
BLOCK FLUSH
3
Yes
Yes
Yes
Yes
Yes
81
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
Yes
82
9
#18 TO COLM
10
Yes
Yes
Yes
Yes
Yes
83
2
REVERSE FLUSH
5
Yes
Yes
Yes
Yes
Yes
84
1
BLOCK FLUSH
3
Yes
Yes
Yes
Yes
Yes
85
34
CYC END
1
Yes
Yes
Yes
Yes
Yes
a USER F = 4,6,12,16,20,23 Tet +12 TO COLM.
Table 3.16.3 Bottle Positions for 381A ABI DNA Synthesizer
Bottle
Reagent
5
5 -O, N-protected deoxyribonucleoside tert-butyl phosphoramidite
9
0.3 M 5-benzylthio-1H-tetrazole in CH3 CN
12
0.03 M bis(2-cyanoethyl)-N,N-diisopropylphosphoramidite in CH3 CN (capping solution)
14
3% DCA in CH2 Cl2 (detritylation solution)
15
1.1 M tBuOOH in CH2 Cl2 for phosphodiester (PO oxidizing solution)
15
0.05 M 3H-1,2-benzodithiol-3-one 1,1-dioxide (Beaucage reagent) in CH3 CN (PS oxidizing solution)
6. Prepare a 0.09 M solution of commercial deoxyribonucleoside 2-cyanoethyl phosphoramidites in dry acetonitrile and place the corresponding bottles (no. 1 to 4) on the synthesizer using the change bottle procedure. 7. Enter the sequence of the desired oligonucleotide. Add an extra “dummy” base 3 to the sequence to account for the solid support. For example, enter ACTGTGTXX to obtain ACTGTGTX, where X is the nucleoside originating from building block S.2a-d.
8. Pack a column with 1 µmol of solid support S.5 (e.g., 20 mg if loading is 50 µmol/g). Place it on the synthesizer. 9. Run the synthesis. To save some deblocking solution, press the JUMP button during step 60 and go to step 77, then resume the synthesis; otherwise, let the synthesis cycle perform the detritylation steps. A jump from step 60 to 77 allows for the column to be washed with dry acetonitrile removing any trace of water, if present. Universal and Recyclable Solid Support
3.16.12 Supplement 30
Synthesize oligonucleotide 10. When the synthesis is complete, remove the column. Use two syringes to push 1 mL of 10 mM K2 CO3 in methanol back and forth through the solid support over a period of 10 min. Current Protocols in Nucleic Acid Chemistry
11. Withdraw the solution and transfer to a 2-mL microcentrifuge tube. Evaporate off the methanol under speed vacuum. 12. Add 1 mL of concentrated aqueous ammonia and transfer to a sealed vial for HPLC. Heat 5 hr in a 55◦ C dry bath. 13. Cool to room temperature and then evaporate the ammonia and water under speed vacuum. 14. Redissolve the residue in 300 µL pure water for further analysis and characterization.
Recycle hexanol solid support 15. Wash the synthesis column successively with 2 mL water and 2 mL methanol. 16. Dry the synthesis column under vacuum in a desiccator over P2 O5 for 2 hr. The dried column can be stored for months at room temperature in a dry place. The same synthesis column has been used four times without a noticeable decrease in quantity or quality of the final oligodeoxyribonucleotides.
TANDEM OLIGONUCLEOTIDE SYNTHESIS Tandem oligonucleotide synthesis can be used to synthesize two oligonucleotides in a row, each starting with the incorporation of a tert-butyl phosphoramidite (S.2a-d). The overall procedure is illustrated in Figure 3.16.4. The synthesis is carried out using the same materials and procedures as in Basic Protocol 3, with the only difference being that the two desired oligonucleotide sequences are entered sequentially in step 7 of Basic Protocol 3. For example, AGTGTXTGACGXX would be entered to obtain
Figure 3.16.4 Tandem synthesis of oligodeoxyribonucleotides (ODNs). * indicates N-acylated nucleobases. BMT, 5-(benzylthio)-1H-tetrazole; DCA, dichloroacetic acid.
ALTERNATE PROTOCOL 1
Synthesis of Unmodified Oligonucleotides
3.16.13 Current Protocols in Nucleic Acid Chemistry
Supplement 30
oligonucleotides AGTGTX (ODN2 ) and TGACGX (ODN1 ), where X is the nucleoside originating from building block S.2a-d. After elongation, treatment with K2 CO3 in methanol releases the two oligonucleotides in solution. This strategy is useful for the synthesis of PCR primers or to double the yield of a single oligonucleotide. ALTERNATE PROTOCOL 2
TANDEM OLIGONUCLEOTIDE SYNTHESIS WITH SELECTIVE RELEASE OF EACH OLIGONUCLEOTIDE This protocol describes the synthesis of two oligonucleotides in a row starting from a commercially available solid support loaded with a deoxyribonucleoside linked through a 3 -O-succinyl linker. Upon complete synthesis of the first oligonucleotide, the synthesis of the second oligonucleotide begins with the addition of a deoxyribonucleoside tert-butyl phosphoramidite (S.2a-d) by transesterification of the H-phosphonate diester linkage as shown in Figure 3.16.5. A first treatment with methanolic K2 CO3 at 0◦ C releases only the second oligonucleotide (ODN*2 ), which is isolated and treated with concentrated ammonium hydroxide for full deprotection. Next, the solid support is treated with concentrated ammonium hydroxide (90 min at 20◦ C) to release the first oligonucleotide (ODN*1 ).
Universal and Recyclable Solid Support
Figure 3.16.5 Tandem oligonucleotide synthesis on a standard deoxyribonucleoside 3 O-succinylated solid support with selective release of oligodeoxyribonucleotides. BMT, 5(benzylthio)-1H-tetrazole; DCA, dichloroacetic acid; ODN, oligodeoxyribonucleotide. * indicates N-acylated nucleobases.
3.16.14 Supplement 30
Current Protocols in Nucleic Acid Chemistry
The ammoniacal oligonucleotide solution is then heated for 5 hr at 55◦ C to complete oligonucleotide deprotection. Although this approach does not permit recycling of the solid support, it allows the use of any commercial succinylated solid support for the tandem synthesis of two oligonucleotides that can be obtained individually.
Additional Materials (also see Basic Protocol 3) Commercial 3 -O-succinylated solid support Concentrated ammonium hydroxide 1. Prepare solutions as in Basic Protocol 3, steps 1 to 6. 2. Enter the sequence of the two oligonucleotides in a row. For example, enter AGTGTXTGACGA to obtain AGTGTX and TGACGA, where X is the nucleoside originating from building block S.2a-d.
3. Place the nucleoside 3 -O-succinylated solid support on the synthesizer and run the synthesis. 4. When the tandem oligonucleotide synthesis is complete, remove the synthesis column. 5. Release the second oligonucleotide from the solid support by using two syringes to push back and forth 1 mL of cold 10 mM K2 CO3 in methanol through the solid support over a period of 20 min at 0◦ C. Since the two oligonucleotides are linked together by an H-phosphonate diester linkage, a selective release of the second oligonucleotide is performed by transesterification. During this short treatment, the ester function between the first oligonucleotide and the solid support is not hydrolyzed. Therefore, the first oligonucleotide remains on the solid support.
6. Withdraw the solution and transfer to a microcentrifuge tube. Evaporate the methanol under speed vacuum. 7. Add 1 mL concentrated ammonium hydroxide to the solution and transfer to a sealed vial. Heat 5 hr in a 55◦ C dry bath. 8. Release the first oligonucleotide from the solid support by using two syringes to push back and forth 1 mL of concentrated ammonium hydroxide through the solid support over a period of 30 min. Repeat two times, for a total of three times. 9. Combine the ammoniacal solutions in a sealed vial and heat 5 hr in a 55◦ C dry bath. 10. Cool each of the oligonucleotide solutions (steps 7 and 9) and evaporate under speed vacuum. 11. Redissolve the residues in pure water for further analysis and characterization.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Molybde stain (Dittmer and Lester, 1964) Solution 1: Boil 40.11 g of MoO3 in 1 L of 25 N sulfuric acid until completely dissolved (3 to 4 hr). Allow the light yellow solution to cool slowly (overnight) to ambient temperature. Store up to several months in the dark. The solution will turn light blue. continued
Synthesis of Unmodified Oligonucleotides
3.16.15 Current Protocols in Nucleic Acid Chemistry
Supplement 30
Solution 2: Boil a suspension of 1.78 g molybdenum powder in 500 mL of solution 1 for 15 min. Cool and decant from the remaining residue. Store up to several months in the dark. Stain solution: Add one volume of each solution to 4.5 volumes of water. Prepare weekly and store in the dark. A dark green solution is formed.
COMMENTARY Background Information
Universal and Recyclable Solid Support
Oligonucleotides are widely used in research and as diagnostic probes. For many biological applications, the 3 -hydroxyl of oligonucleotides must be free. For convenience, universal solid supports are preferred since their use minimizes preparation and storage of solid supports functionalized with each of the four nucleobases, and also minimizes errors and manipulations. In addition, ∼40% of the cost of an oligonucleotide is from the solid support, which is wasted at the end of the synthesis. Thus, an obvious way to reduce the average cost of an oligonucleotide is to recycle the solid support. Furthermore, in the context of durable development, the amount of waste is reduced. It has been shown that the presence of an H-phosphonate diester linkage between an oligonucleotide and a universal hydroxylated solid support allows an efficient and clean release of the 3 -hydroxylated oligonucleotide under mild basic conditions (Ferreira et al., 2005). It has also been reported (Sobkowski et al., 1994) that ethanolamine and 3-aminopropanol cleave H-phosphonate diester linkages within 15 min. However, the removal of these aminoalcohols by evaporation is inconvenient due to their high boiling points. In contrast, methanolic K2 CO3 cleaves the H-phosphonate diester linkage with a better efficiency (10 min) and is easy to remove (Ferreira et al., 2005). The cleavage proceeds through a methanolmediated transesterification resulting in the formation of dimethylphosphite in addition to an oligonucleotide with a free 3 -hydroxyl group and a hydroxylated solid support (S.5, Fig. 3.16.3). This mild basic treatment does not damage the solid support, which can be reused after simple washing and drying steps. The H-phosphonate diester linkage can be formed using conventional nucleoside H-phosphonate monoesters, which require different reagents and solvents from those used in the phosphoramidite method. o-Methoxybenzyl (Meyer et al., 2004) or
tert-butyl (Ferreira et al., 2005) phosphite triesters produce H-phosphonate diester linkages through an Arbuzov-like mechanism under acidic conditions (Fig. 3.16.6). Thus, the advantage of using nucleoside tert-butyl phosphoramidites instead of classic nucleoside H-phosphonate monoesters in the formation of H-phosphonate diester linkages is that no reagents other than those used in the phosphoramidite chemistry are needed. The preparation of the deoxyribonucleoside tert-butyl phosphoramidite building blocks (S.2a-d) proceeds like that of conventional phosphoramidite derivatives starting from commercially available 5 -O, N-protected deoxyribonucleosides and tert-butyl tetraisopropylphosphorodiamidite. The coupling efficiency of these phosphoramidites is slightly lower due to the steric hindrance created by the tert-butyl group in the vicinity of the phosphorous atom. Thus, 5-methylthio-1H-tetrazole (pKa = 4.08; Wu and Pitsch, 1998), which is a more acidic activator than 1H-tetrazole (pKa = 4.89), is used for coupling reactions performed over a period of 45 sec. This strategy allows the synthesis of oligonucleotides (phosphodiester or phosphorothioate) that are comparable in terms of purity and quantity. Since the H-phosphonate diester linkage must be kept intact during the elongation cycle, the oxidation of phosphite triester linkages must be performed using tert-butyl hydroperoxide (Jung et al., 1994; Bologna et al., 1999) or the Beaucage reagent (Bologna et al., 1999), which are known to oxidize only phosphite triester linkages. The universal solid support can be reused several times after performing washes with water and methanol and subsequent drying under vacuum. The same oligonucleotide sequence has been synthesized four times on the same recycled solid support without any observed decrease in product quality and quantity from one synthesis to the next. Since methanolic K2 CO3 treatment removes the 2-cyanoethyl groups but only partially removes the nucleobase-protecting
3.16.16 Supplement 30
Current Protocols in Nucleic Acid Chemistry
Figure 3.16.6
H-Phosphonate diester linkage formation through an Arbuzov-like mechanism.
groups, it is necessary to complete the nucleobase deprotection of the released oligonucleotides through the use of concentrated ammonium hydroxide. Alternatively, a single treatment with methanolic K2 CO3 (10 min at room temperature and 20 min at 50◦ C) fully deprotects oligonucleotides bearing ultra-fastdeprotecting groups, such as acetyl on cytidine, phenoxyacetyl on adenosine, or (4isopropyl)phenoxyacetyl on guanosine. Multiple syntheses For production of PCR primers, it is more efficient to synthesize two oligonucleotide primers during the same run and obtain them together in solution. This approach improves productivity by reducing both handling and storage. The universal support approach allows the preparation of both primers in tandem using a deoxyribonucleoside tert-butyl phosphoramidite building block to initiate the synthesis of each sequence. Upon completion of the synthesis, the two oligonucleotides are released together and are fully deprotected by treatment with methanolic K2 CO3 and concentrated ammonium hydroxide, respectively. Tandem oligonucleotide synthesis can also be applied to the production of larger quantities of a single oligonucleotide, when required. In that case, the same sequence is synthesized two times in a row to double the amount of oligonucleotide product. This strategy could also be applied to the synthesis of siRNA where two complementary RNA strands are required. Two strategies are reported in the literature for tandem oligonucleotide synthesis; one approach involves a complex synthesis of building blocks and requires harsher basic conditions for oligonucleotide deprotection (Hardy et al., 1994). The other strategy requires specific reagents not usually used on a DNA synthesizer (Pon et al., 2002). In addition, UNIT 3.12 reports the use of deoxyribonucleoside phosphoramidites containing cleavable linkers that provide, after deprotection, one oligonucleotide with free 3 - and 5 hydroxyls and one oligonucleotide bearing a
5 -phosphate monoester and a free 3 -hydroxyl group. Finally, tandem oligonucleotides can be synthesized in which the first oligonucleotide is initiated from a commercially available succinylated solid support, and the second is initiated by the coupling of a deoxyribonucleoside tert-butyl phosphoramidite building block according to the customized elongation cycle to generate an H-phosphonate linkage. Subsequent treatment with K2 CO3 /methanol at 0◦ C for 20 min cleaves the H-phosphonate linkage and thus selectively releases the second oligonucleotide while the first remains linked to the succinylated CPG. A standard treatment of the CPG beads with concentrated ammonium hydroxide then releases the first oligonucleotide. In this way, two oligonucleotides are synthesized in a row, but their release is selective and each is obtained individually.
Critical Parameters The purification of deoxynucleoside 3 -O-(tert-butyl)-N,N-diisopropylphosphoramidites by flash chromatography on silica gel must be carried out with triethylamine in the eluent to avoid degradation due to the acidity of silica. If possible, the chromatography must be done within 2 hr. Automated synthesis of oligonucleotides according to the phosphoramidite method is well established. Due to the sensitivity of the method to moisture, the phosphoramidite and activator solutions must be kept dry to ensure high coupling efficiencies. This is achieved by ◦ adding a few beads of 4-A molecular sieves to the solutions. The deoxyribonucleoside tertbutyl phosphoramidite building blocks are less reactive than standard 2-cyanoethyl phosphoramidites. To increase their coupling efficiency, the use of 5-benzylthio-1H-tetrazole and an extended coupling reaction time of 45 sec are recommended. Capping is performed using bis-(2-cyanoethyl)-N,N-diisopropylphosphoramidite, since the presence of an H-phosphonate diester linkage precludes the use of acetic
Synthesis of Unmodified Oligonucleotides
3.16.17 Current Protocols in Nucleic Acid Chemistry
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anhydride. This reagent leads to shorter oligonucleotide 5 -phosphate monoesters with increased hydrophilicity compared to that of the expected full-length oligonucleotides, and thus facilitates oligonucleotide purification by reversed-phase chromatography. Other phosphoramidites can be used as capping reagents, including diethyl N,N-diisopropyl phosphoramidite (Yu et al., 1994; Alvarez et al., 1999) or O-(2-cyanoethyl)-O-(n-octyl)-N,N-diisopropylphosphoramidite (Natt and Haner, 1997).
Anticipated Results For investigators who are familiar with the preparation of phosphoramidites, the synthesis of deoxyribonucleoside 3 -O-(tert-butyl)N,N-diisopropylphosphoramidites is straightforward and yields ranging between 65% and 86% are expected. The preparation of the solid support is straightforward since all steps are performed on the support; only filtrations and washes are necessary for the work-up. Synthesis of oligonucleotides on the universal solid support, starting with one deoxyribonucleoside tert-butyl phosphoramidite building block and then using commercially available deoxyribonucleoside phosphoramidites, is as efficient as syntheses carried out on a succinylated solid support in terms of quantity and purity. After recycling, the universal solid support can be reused several times without losing its functional properties. Synthesis of tandem oligonucleotides by this strategy leads to products of sufficient purity to be used directly as PCR primers.
Time Considerations
Universal and Recyclable Solid Support
The synthesis of a deoxyribonucleoside 3 -O-(tert-butyl)-N,N-diisopropylphosphoramidite, including purification, requires ∼1 full day, with one overnight reaction. Preparation of the hydroxylated universal solid support requires ∼2 days. Since it is recycled, its preparation is not required each time. Typically, the duration of the 1-µmol customized oligodeoxyribonucleotide elongation cycle is ∼8 min. This cycle is slightly longer than the conventional cycle since the oxidation step using either tert-butyl hydroperoxide or 3H-1,2-benzodithiol-3-one 1,1-dioxide requires 60 sec instead of 10 sec with the iodine/ water solution, and the capping step using a phosphoramidite takes longer than capping with acetic anhydride. The synthesis of a 20-mer requires ∼2.5 hr.
Release of the oligonucleotide from the hydroxylated solid support is faster (only 10 min) than release from a succinylated solid support (90 min). The complete removal of conventional benzoyl and isobutyryl protecting groups is achieved within 5 hr at 55◦ C using concentrated ammonium hydroxide. However, when using ultra-fast deprotecting groups (i.e., acetyl, phenoxyacetyl, and isopropylphenoxyacetyl) for nucleobase protection, complete removal of these groups and release of the oligonucleotide from the support required only 20 min at 50◦ C when treated with 50 mM potassium carbonate in methanol. Recycling the hydroxylated solid support requires ∼10 min for the washes and 2 hr of drying time. Since the solid support is kept in the synthesis column during oligonucleotide release and recycling steps, no additional time is needed for repacking the column.
Literature Cited Alvarez, K., Vasseur, J.J., Beltran, T., and Imbach, J.L. 1999. Photocleavable protecting groups as nucleobase protections allowed the solid-phase synthesis of base-sensitive SATEprooligonucleotides. J. Org. Chem. 64:63196328. Beaucage, S.L. and Caruthers, M.H. 1981. Deoxynucleoside phosphoramidites—A new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Lett. 22:1859-1862. Bologna, J.C., Morvan, F., and Imbach, J.L. 1999. The prooligonucleotide approach: Synthesis of mixed phosphodiester and SATE phosphotriester prooligonucleotides using H-phosphonate and phosphoramidite chemistries. Eur. J. Org. Chem. 2353-2358. Dittmer, J.C. and Lester, R.L. 1964. Simple specific spray for detection of phospholipids on thinlayer chromatograms. J. Lipid Res. 5:126-127. Ferreira, F., Meyer, A., Vasseur, J.J., and Morvan, F. 2005. Universal solid supports for the synthesis of oligonucleotides via a transesterification of H-phosphonate diester linkage. J. Org. Chem. 70:9198-9206. Hardy, P.M., Holland, D., Scott, S., Garman, A.J., Newton, C.R., and Mclean, M.J. 1994. Reagents for the preparation of two oligonucleotides per synthesis (TOPS). Nucl. Acids Res. 22:29983004. Hayakawa, Y., Uchiyama, M., and Noyori, R. 1986. Nonaqueous oxidation of nucleoside phosphite to the phosphates. Tetrahedron Lett. 27:41914194. Iyer, R.P., Phillips, L.R., Egan, W., Regan, J.B., and Beaucage, S.L. 1990. The automated synthesis of sulfur-containing oligodeoxyribonucleotides using 3H-1,2-benzodithiol-3-one 1,1-dioxide as a sulfur-transfer reagent. J. Org. Chem. 55:46934699.
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Jung, P.M., Histand, G., and Letsinger, R.L. 1994. Hybridization of alternating cationic/anionic oligonucleotides to RNA segments. Nucleosides Nucleotides 13:1597-1605. Meyer, A., Morvan, F., and Vasseur, J.-J. 2004. H-Phosphonate oligonucleotides from phosphoramidite chemistry. Tetrahedron Lett. 45:37453748. Natt, F. and Haner, R. 1997. Lipocap—A lipophilic phosphoramidite-based capping reagent. Tetrahedron 53:9629-9636. Pon, R.T., Yu, S., and Sanghvi, Y.S. 2002. Tandem oligonucleotide synthesis on solid-phase supports for the production of multiple oligonucleotides. J. Org. Chem. 67:856-864.
Sobkowski, M., Stawinski, J., Sobkowska, A., and Kraszewski, A. 1994. Studies on reactions of nucleoside H-phosphonates with bifunctional reagents. 2. Stability of nucleoside H-phosphonate diesters in the presence of amino alcohols. J. Chem. Soc., Perkin Trans. 118031808. Wu, X.L. and Pitsch, S. 1998. Synthesis and pairing properties of oligoribonucleotide analogues containing a metal-binding site attached to beta-D-allofuranosyl cytosine. Nucl. Acids Res. 26:4315-4323. Yu, D., Tang, J.Y., Iyer, R.P., and Agrawal, S. 1994. Diethoxy N,N-diisopropyl phosphoramidite as an improved capping reagent in the synthesis of oligonucleotides using phosphoramidite chemistry. Tetrahedron Lett. 35:8565-8568.
Synthesis of Unmodified Oligonucleotides
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CHAPTER 4 Synthesis of Modified Oligonucleotides and Conjugates INTRODUCTION he genetic information encoded in either genomic DNA or transcribed mRNAs has become an enticing target for chemical modifications and potential therapeutic indications. Specifically, the search for methods that selectively prevent expression of unwanted genes has been actively pursued for more than a decade. Theoretically, it is possible to exploit the selective Watson-Crick hydrogen bond formation between complementary nucleic acid strands to control gene expression either by preventing transcription of genomic DNA (an antigene strategy) or by interfering with the translation of mRNAs (an antisense strategy) to gene products. With the advent of rapid and efficient methods for automated synthesis of oligonucleotides, it has indeed been possible to prepare modified oligonucleotides complementary to specific nucleotide sequences of mRNA targets. As a consequence of binding, and on the basis of particular cellular mechanisms, these oligonucleotides can disrupt the flow of genetic information from transcribed mRNAs to proteins. Thus, gene expression can be controlled at the mRNA level by modified oligonucleotides, and this antisense strategy has led, over the years, to the development of therapeutic oligonucleotides against cancer and various infectious diseases in humans. In this context, UNIT 4.1 reports recent advances in the application of oligonucleotides as drug candidates, describes the relationship between oligonucleotide modifications and their therapeutic profiles, and provides general guidelines for enhancing oligonucleotide drug properties.
T
Along similar lines, modern methods for the chemical synthesis of oligonucleotides have facilitated the covalent attachment of reporter and conjugate groups to DNA and RNA. Such modifications generate properties not normally present in native DNA or RNA oligonucleotides. For example, addition of a conjugate group to an oligonucleotide can be used to alter its solubility, charge, binding affinity, spectroscopic properties, and function. Furthermore, oligonucleotide conjugates can provide structural information on the nature of specific nucleic acid complexes such as protein-DNA(RNA) or ligand-DNA(RNA) complexes. Unlike enzymatic methods, chemical methods for the incorporation of modified nucleosides into oligonucleotides permit a precise control over the type, number, and placement of the reporter and conjugate groups within the sequence of interest. Typically, fluorescent markers, intercalators, cross-linkers, and DNA/RNA cleaving agents are added either to the 5 or 3 terminus of oligonucleotides, or inserted anywhere within an oligonucleotide sequence. These oligonucleotide conjugates can then be specifically dedicated to diagnostic or therapeutic applications. UNIT 4.2 presents an overview of the methods employed for the attachment of reporter and conjugate groups to the 5 terminus of oligonucleotides. The unit lists several reporter and conjugate groups that can be directly incorporated into oligonucleotides mainly via ligand phosphoramidite or H-phosphonate derivatives. This knowledge is reduced to practice in UNIT 4.3, which delineates step-by-step protocols for the synthesis of intercalating and photo-cross-linking ligand phosphoramidites as well as their addition to the 5 terminus of oligonucleotides. Reporter and conjugate groups are, however, not always stable to the conditions used during either oligonucleotide synthesis or the subsequent deprotection steps. Consequently,
Synthesis of Modified Oligonucleotides and Conjugates
Contributed by Serge L. Beaucage
4.0.1
Current Protocols in Nucleic Acid Chemistry (2007) 4.0.1-4.0.4 C 2007 by John Wiley & Sons, Inc. Copyright
Supplement 29
the incorporation of these groups into oligonucleotides must be accomplished according to an “indirect” approach. This method involves functionalization of the 5 terminus of oligonucleotides, mostly with nucleophilic groups. Although the indirect approach to the preparation of oligonucleotide conjugates is extensively reviewed in UNIT 4.2, procedures for the modification of oligonucleotides at the 5 terminus are outlined in UNIT 4.9. Following deprotection and purification, the functionalized oligonucleotides are then conjugated to selected electrophilic reporter or conjugate groups. Methods describing the incorporation of functional groups into ligands are reported in UNIT 4.8, and methods for conjugation of these ligands to the 5 -functionalized oligonucleotides are described in UNIT 4.10. Phosphoramidites tethered with single or multiple linkers denote a creative approach to the functionalization of oligonucleotides. These phosphoramidites are prepared from methoxyoxalamido (MOX)–derived secondary alcohol precursors, which are assembled upon reaction with amino alcohols or, repeatedly and sequentially, with aliphatic diamines and dimethyloxalate. This MOX phosphoramidite chemistry leads to oligonucleotides tethered with spacers varying in hydrophobicity, rigidity, and length, that are amenable to conjugation with diverse electrophilic ligands. This versatile functionalization method is featured in UNIT 4.29. An attractive method for the preparation of oligonucleotide conjugates consists of functionalizing the 5 -terminus of solid-phase-bound oligonucleotides with a diene phosphoramidite derivative followed by conjugation of the deprotected oligonucleotides with selected dienophile reporter groups via a Diels-Alder cycloaddition reaction. The details of this convenient conjugation method are provided in UNIT 4.18. For similar applications, a facile and efficient 5 -iodination of solid-phase-linked oligonucleotides is presented in UNIT 4.19. 5 -Iodo oligonucleotides allow conjugation with small molecules or macromolecules having a strongly nucleophilic group. While the functionalization of oligonucleotides at either the 5 - or 3 -terminus is invaluable in the preparation of oligonucleotide conjugates, the incorporation of uridine 2 -carbamates into oligonucleotides nicely complements these methods for direct or indirect attachment of various ligands to DNA or RNA. This versatile method is carefully detailed in UNIT 4.21. The purification of synthetic oligonucleotides from a mixture of shorter sequences is always a concern. Thus, purification methods that can easily provide oligonucleotides of high purity are of great value. UNIT 4.20 highlights a reversible biotinylation of the 5 -terminus of oligonucleotides, which allows simple and highly effective affinity purification of these biomolecules.
Introduction
This chapter is also intended to include overviews and step-by-step protocols addressing the attachment of reporter and conjugate groups to the nucleobase, the 3 terminus, and the carbohydrate portion of oligonucleotides for diagnostic applications. In this regard, UNIT 4.5 surveys a number of conjugation methods for oligonucleotides displaying various 3 functional groups, and UNIT 4.6 describes the step-by-step preparation of 3 -aminoalkylated or 3 -mercaptoalkylated oligonucleotides, and their respective conjugation to fluorescent reporter groups. UNIT 4.31 addresses the attachment of 1,4,7,10tetraazacyclododecane-1,4,7,10-tetracetic acid (DOTA) to deoxyuridine functionalized with an aminoalkylated linker at N3, and the conversion of the modified deoxyribonucleoside to its phosphoramidite derivative. This phosphoramidite is used in the preparation of a DOTA-labeled oligonucleotide and its gadolinium(III) chelate. This unit thus provides a straightforward method for site-specific incorporation of a magnetic resonance imaging contrast agent into oligonucleotides for target-specific applications. Other protocols describing the preparation of oligonucleotides with modified nucleobases, sugars, and internucleotide linkages for potential therapeutic applications will also be incorporated into this chapter to provide a comprehensive coverage of these cutting edge technologies. In
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this context, UNIT 4.4 delineates the solid-phase synthesis of chimeric 2-5A-DNA oligonucleotide conjugates along with the preparation of the phosphoramidite monomers used for such synthesis. In addition, the synthesis and purification of oligonucleotides composed of N3 →P5 phosphoramidate and either phosphodiester or phosphorothioate linkages, as presented in UNIT 4.7, is a relevant example of such protocols. UNIT 4.11 provides manual and automated methods for the synthesis of peptide nucleic acids (PNAs), whereas UNIT 4.12 details the synthesis of locked nucleic acids (LNAs). These modified nucleic acids exhibit high affinity for complementary RNA and DNA strands, and are thus amenable to a wide range of biomedical applications. In this regard, a procedure for the incorporation of LNAs into cells is reported in UNIT 4.13. The conjugation of peptides to oligonucleotides to generate nucleopeptides is the focus of intense scrutiny given the reported abilities of these conjugates at enhancing intracellular delivery of therapeutic oligonucleotides. However, the preparation of peptide-oligonucleotide conjugates is challenging. An inspiring method for the stepwise solid-phase synthesis of nucleopeptides is delineated in UNIT 4.22, thus making these bioconjugates more available for further cell delivery studies. Another innovative approach to the preparation of peptide-oligonucleotide conjugates is based on the respective functionalization of oligonucleotides and peptides with either a free or activated thiol group. Conjugation of the functionalized entities occurs rapidly and efficiently through formation of a disulfide link. The specifics of this method are featured in UNIT 4.28. A different strategy for internalization and cell-specific targeting of therapeutic oligonucleotides relates to the covalent attachment of these molecules to carbohydrates. UNIT 4.26 includes methods for the preparation of a mannose-derived ligand and a non-nucleosidic phosphoramidite functionalized with phthalimidooxy groups as building blocks in the solid-phase construction of a multiantennary oligonucleotide glycoconjugate. Such glycocluster structures have recently attracted considerable attention in terms of potential biomedical applications, as these are a prerequisite for high-affinity cellular binding and internalization of their conjugates by endocytosis. Synthetic nucleic acids containing vicinal 2 ,5 - and 3 ,5 -phosphodiester linkages have been useful to investigations related to pre-mRNA splicing events and to studies assessing the specificity of RNA lariat debranching enzymes. These branched nucleic acids may also find applications in the development of biosensors and diagnostics, or may serve as “molecular anchors” in the formation of triplex and tetraplex structures. Two synthetic strategies for the preparation of branched oligonucleotides are outlined and developed in UNIT 4.14. An important class of uncharged P-chiral analogs of nucleic acids includes morpholino oligonucleotides. These synthetic oligonucleotide analogs are typically used to block translation of mRNA, alter the splicing of pre-mRNA, or prevent other interactions between biological macromolecules and RNA through base-pairing events. Morpholino oligonucleotides combine efficacy, specificity, stability, lack of non-antisense effects, and exhibit good water-solubility properties. Exquisite technical information on how to design and deliver morpholino oligonucleotides to control gene expression in cell cultures is given in UNIT 4.30. Transfer RNAs (tRNAs) incorporate a variety of modified nucleobases, which are known to affect the biological activity of tRNAs through structural changes. To better understand the role of these modified nucleobases, UNIT 4.23 reports on the synthesis of ribonucleoside phosphoramidites functionalized with either 6-methylthio purine or 2methylthio-6-chloro purine for incorporation into relatively short RNA oligonucleotide models. Following oligonucleotide synthesis using these phosphoramidites, the modified nucleobases are chemically converted to a number of native tRNA nucleobase modifications to further studies in this research area.
Synthesis of Modified Oligonucleotides and Conjugates
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Modified nucleic acids such as 2 -deoxy-2 -fluoro-β-D-oligoarabinonucleotides are unique in that they form more stable duplex structures with RNA complements than identical unmodified DNA:RNA duplexes. In addition, the modified heteroduplexes are substrates for RNase H, an enzyme viewed by many as the major effector of antisense activity. Thus, given the promising antisense properties of 2 -fluoroarabinonucleic acids, methods for the synthesis of 2 -deoxy-2 -fluoro-β-D-arabinonucleoside phosphoramidites and their use in the automated synthesis of 2 -fluoroarabinonucleotides are included in UNIT 4.15. Substitution of native nucleobases with 7-deazapurines (pyrrolo[2,3-d]pyrimidines) functionalized with halogeno or alkynyl groups at C7 has been shown to stabilize DNA duplexes. This serves as a creative example in the design of therapeutic oligonucleotides with high-affinity binding for complementary nucleic acid targets. The preparation of these modified DNA oligonucleotides using phosphoramidite monomers is described in UNIT 4.25. Synthesis of the pyrrolo[2,3-d]pyrimidine 2 -deoxyribonucleoside precursors is provided in UNIT 1.10. An additional class of oligonucleotides for potential therapeutic applications includes phosphonoacetate and thiophosphonoacetate oligodeoxyribonucleotide analogs. These modified oligonucleotides are highly resilient to nucleases, and yet they are anionic, water soluble, and form hybrids with both native DNA and RNA complementary sequences. It is noteworthy that both oligonucleotide analogs stimulate RNase H–mediated hydrolysis of the complementary RNA strand of each DNA:RNA hybrid. UNIT 4.24 presents the detailed preparation of the deoxyribonucleoside phosphoramidites required for solidphase synthesis of phosphonoacetate and thiophosphonoacetate oligonucleotides, along with methods used for purification and characterization of these oligonucleotide analogs. Phosphorothioated oligonucleotides have also found therapeutic applications in antisense settings. Such modified oligonucleotides are chiral at phosphorous, and those exhibiting the Sp configuration are significantly more resistant to nucleases found in human plasma than P-diastereomeric or Rp -phosphorothioated oligonucleotides. Methods for the synthesis of oligonucleotides with stereodefined phosphorothioate linkages would therefore be valuable in the preparation of therapeutic oligonucleotides with optimal resistance to nucleases. UNIT 4.17 specifically describes the use of deoxyribonucleoside3 -O-(2-thio-1,3,2-oxathiaphospholane) derivatives in the P-stereocontrolled synthesis of phosphorothioated oligonucleotides. Another class of modified oligonucleotides with potential biomedical applications relates to phosphorothioated oligodeoxyribonucleotides expressing specific CpG motifs. These oligonucleotides strongly stimulate B cells, natural killers cells, and antigen-presenting cells to proliferate and/or secrete cytokines, chemokines, and immunoglobulins. Given that a number of structural factors influences the recognition of CpG-oligonucleotides and expression of their immunostimulatory properties, UNIT 4.16 provides an overview of the chemical strategies used to assess the significance of these factors toward the development of human therapies. With the incidence of therapeutic applications for phosphorothioated oligonucleotides, the large-scale preparation of these biopolymers through automated solid-phase techniques is increasingly important, but is challenging and costly in terms of reagent consumption. A new process whereby polyethylene glycol is employed as a soluble support for the large-scale synthesis of thioated oligonucleotides and selected conjugates is proposed in UNIT 4.27. Such a process offers several advantages over traditional solid-phase methods for oligonucleotide synthesis, and these are discussed in the unit. Introduction
Serge L. Beaucage
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A Brief History, Status, and Perspective of Modified Oligonucleotides for Chemotherapeutic Applications Modification of oligonucleotides for drug applications has a very short history (for highlights see Table 4.1.1 and Table 4.1.2). Intensive efforts to modify short strands of nucleic acids to enhance their properties as drug candidates began in earnest in the late 1980s. The notion that oligonucleotides could be made into drugs along with the discovery and development of two essential enabling technologies— automated DNA sequencing (Connell et al., 1987) and commercial automated oligonucleotide synthesis and purification in the early 1980s—set the stage for oligonucleotide drug discovery (reviewed by Zon, 1993). Subsequently, oligonucleotide medicinal chemistry and pharmacology, which form the basis of structure-activity relationship studies, have been highly developed in the process of converting oligonucleotides into drugs. Before this, little value was given to synthetically changing the structure of short strands of DNA or RNA except for some early modifications primarily directed to enhance structural studies and diagnostic applications. Now great interest in making drugs out of oligonucleotides has spurred intense structure-activity/property relationship (SAPR) studies to optimize drug properties (Cook, 1998a,b). Given that oligonucleotides are gene-based materials (or informational materials; Cohen, 1991) and considering the emergence of genomic target selection, which is also gene-
Table 4.1.1
UNIT 4.1
based, it is surprising that it has taken so long for the drug discovery community to recognize the potential value of modified oligonucleotides as drugs. However, when traditional medicinal chemists first consider modifying oligonucleotides, they are confronted with a number of daunting chemical, biophysical, and biochemical properties of oligonucleotides (Table 4.1.3), which has surely slowed progress in this area. In addition to the need to apply novel ligands to drug discovery, the nucleic acid targets that serve as receptors, particularly RNA molecules, are novel as well. In spite of the early difficulties in exploring the use of oligonucleotides as drugs, however, much progress has been made in recent years. The first antisense oligonucleotide drug, Vitravene, was recently approved by the Food and Drug Administration (FDA) for treating cytomegalovirus retinitis in AIDS patients, and several additional oligonucleotides are in late phase II and III clinical studies for anti-cancer and antiinflammatory indications (Table 4.1.4).
OVERVIEW OF BASE PAIRING AND GENE EXPRESSION A single strand of nucleic acid has a polymeric sugar-phosphate backbone with ribofuranosyl (in RNA) or deoxyribofuranosyl (in DNA) residues connected at the 3′ and 5′ hydroxyls by phosphoric acid to form phosphodiester linkages. Each sugar residue has one of
Historical Listing of Oligo- and Polynucleotides as Therapeutic Agents
Milestone
References
DNA alkylating agents Poly(rI) • poly(rC) Macromolecular antimetabolites “Antitemplates” Antisense oligonucleotides (targeting RNA) Heterocycle modified oligonucleotides: prodrugs for nucleosides Antigene oligonucleotides: triple strand formers Aptamers
Belikova et al. (1967); Summerton (1979) De Clercq et al. (1969) Chandra and Bardos (1972); reviewed by Bardos and Ho (1978) See Table 4.1.2 Alderfer et al. (1985)
Sense approach
Moser and Dervan (1987); LeDoan et al. (1987) Ellington and Szostak (1990); Tuerk and Gold (1990) Morishita et al. (1995)
Contributed by P. Dan Cook Current Protocols in Nucleic Acid Chemistry (2000) 4.1.1-4.1.17 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Modified Oligonucleotides and Conjugates
4.1.1
Table 4.1.2
Highlights of the Development of Chemotherapeutic Oligonucleotides
Milestone
References
Antisense experiments in cell culture
Stephenson and Zamecnik (1978); Zamecnik and Stephenson (1978) Merrifield, Letsinger, Caruthers (1962-1987) Ts’o et al. (1987) Eckstein, Stec, Zon (1967-1981) Cech (1986); Altman (1989) Isis Pharmaceuticals (1992-1994)
Automated synthesis, protocols, reagents Methylphosphonate oligonucleotides Phosphorothioates Ribozymes Initiation of first clinical trial; first report of clinical efficacy Twenty five oligonucleotides entered into clinical development First New Drug Application (NDA) for an antisense oligonucleotide First approved antisense oligonucleotide drug
five types of heterocycles (nucleobases) attached at the C1′ position of either the deoxyribofuranosyl or ribofuranosyl sugar. These heterocycles are the purines (guanine and adenine) and the pyrimidines (cytosine and thymine, with uracil replacing thymine in RNA). Complementary DNA describes a chemical complex of two strands of deoxyribonucleic acid that are bound together by WatsonCrick base-pair hydrogen bonding. This “essences of life” bonding specifies that guanine in one DNA strand specifically binds to cytosine in the other strand, and that adenine in one DNA strand specifically binds to thymine in the other strand (Fig. 4.1.1). In RNA, uracil specifically binds to adenine. A sequence (or a specific ordering of nucleobases) of an RNA or a DNA strand will bind specifically
(1992-present; see Table 4.1.4) Isis Pharmaceuticals (April 1998) Isis Pharmaceuticals (August 1998)
(hydrogen bond) to another sequence only if the nucleobases match up (base pair). The strands are then said to be complementary to one another. The complementary DNA strands twist into a helical motif with one strand having its bases ordered in the 3′ to 5′ direction and the other strand having its bases ordered in the opposite direction (5′ to 3′). This is referred to as antiparallel orientation (Fig. 4.1.2). The doublestranded DNA (dsDNA) serves as the template for synthesis of either complementary DNA molecules, to provide a process of self-replication, or complementary RNA molecules (from the sense strand) for the process of transcription. The complementary DNA strand, referred to as the antisense strand, is itself rarely transcribed to give an RNA molecule.
Table 4.1.3 Chemical, Biochemical, and Biophysical Properties of First-Generation Oligonucleotides
Parameter
Property
Molecular weight Charge Chirality in backbone Instabilities Solubility
High (20- to 25-mers, MW 7000 to 9000) Highly negative (19 to 24 charges) or neutral P=S, Me-P, N-Pa Biological and chemical (nucleases, pH, depurination, dethiation) Insoluble in common organic solvents depending on backbone modification (P=S versus Me-P)a No Complex synthesis of reactive monomers Unfamiliarity with instrument synthesis of oligomers Purification by ion exchange chromatography Analysis and characterization atypical
Crystalline? Ease of synthesis Modified Oligonucleotides for Chemotherapeutic Applications
aMe-P, methylphosphonate; N-P, phosphoramidate; P=S, phosphorothioate.
4.1.2 Current Protocols in Nucleic Acid Chemistry
Table 4.1.4
Oligonucleotides Entered into Clinical Developmenta
Disease indications
Oligonucleotide
Molecular target
ISIS 2105
HPV 6 and 11 E2 Genital warts genes product HCMV IE gene-2 CMV retinitis
ISIS 2922
Statusb
Sponsor
Terminated
Isis
NDA approved (August 1998) III, Crohn’s; II for others
Isis/Ciba Vision
Renal allograft, rheumatoid arthritis, Crohn’s disease, ulcerative colitis, psoriasis Cancer II
Isis/Boehringer Ingleheim
Cancer
II
Isis/Novartis
HCMV IE gene Gp120 Ha-ras gag gene gag gene UL36 and UL37 Protein kinase A p53 c-myc c-myb gag gene HIV integrase Bcl-2 gag gene
CMV retinitis AIDS Cancer AIDS AIDS CMV retinitis Cancer Cancer Restenosis Cancer AIDS AIDS Cancer AIDS
I/II Terminated I Terminated I I II Terminated Terminated Terminated Terminated I/II II Terminated
Cancer
I
MG-98 Resten-NG
Ribonucleotide reductase R2 subunit DNA methylase c-myc
Isis Isis Isis Hybridon Hybridon Hybridon Hybridon Lynx/Inex Lynx/Inex Lynx/Inex Chugai Aronex Genta Novopharm Biotech GeneSense
Heptazyme
IRES region
Angiozyme
Vascular endothelial growth factor receptor Immune system
ISIS 2302
Intercellular adhesion molecule (ICAM-1)
ISIS 3521/CPG 64128A ISIS 5132/CPG 69846A ISIS 13312 ISIS 5320 ISIS 2503 GEM-91 GEM-92 GEM-132 GEM-231 OL(1)p53 LR-3280
Protein kinase C-α c-raf kinase
GPs 0193 Zintevir G3139 GPI-2A GTI 2040
CPG-x
Solid tumors I Restenosis and I proliferative diseases Hepatitis C virus I
Isis/Novartis
MethylGene AVI BioPharma
Antiangiogensis (cancer)
I
Ribozyme Pharmaceuticals Ribozyme
Hepatitis B
I
CpG
aAbbreviations: E, early gene; HCMV, human cytomegalovirus; HPV, human papillomavirus; IE, immediate early gene;
IRES, internal ribosomal entry site; NDA, New Drug Application; UL, unique long. bI, II, and III indicate phase of clinical trials at time of printing.
Synthesis of Modified Oligonucleotides and Conjugates
4.1.3 Current Protocols in Nucleic Acid Chemistry
H O O
O O
O
Figure 4.1.1
C
P
N
H
O P HO
N
O
H
O O O P
N O O
N
O
O
−
O
−
O O
HO
N H
O
Watson-Crick hydrogen-bonding (base-pairing) rules (G to C, A to U or T).
The transcribed RNA molecules, in turn, serve as the templates that allow the ordering of the amino acids within the polypeptide chain of proteins, during the process of translation. Translation is an appropriate term because the nucleotide sequence information (language) of nucleic acids is translated into the amino acid sequence (language) of protein. The complete process—DNA makes RNA makes protein— has been referred to as the central dogma of molecular biology. A more common term for the process is gene expression. Thus, the two stages of gene expression are transcription (the conversion of DNA to RNA) and translation (the conversion of RNA to proteins). DNA also self-replicates to maintain an organism’s genome. Furthermore, regulation of gene expression in living organisms involves extensive recognition and binding to specific nucleic acid sequences by nucleic acid–binding proteins.
INFORMATIONAL DRUG DISCOVERY APPROACHES AND THEIR SUCCESS
Modified Oligonucleotides for Chemotherapeutic Applications
H(CH3)
N U/T N O
N G
N
O
O
O
A N H N H N H O
P
O
−
N
O
O
−
N H
N
Disease states (genetic, oncogenic, or infective) are typically a result of the production of abnormal proteins, the end-stage product of gene expression. Thus, it is not surprising that drug discovery has historically focused on interfering with the functions of the abnormal proteins rather than on preventing abnormal protein formation—treating the disease symptoms rather than the cause. In contrast, treatment approaches involving informational chemicals, such as oligonucleotides, attempt to prevent the formation (gene expression) of abnormal protein by targeting DNA, RNA, or regulatory proteins that are required for transcription and translation of the abnormal protein. Having a set of binding rules (Watson-Crick base-pairing rules) that allows one to rapidly and precisely select a molecule to synthesize that will inhibit gene
expression at the DNA or RNA level, and that offers exquisite specificity for its receptor, is unique in the history of drug research. Thus, the employment of informational drugs to complement genomic target validation in modern drug discovery is revolutionary. Several discovery approaches are being pursued that employ the encoded information of modified oligonucleotides as the drug agent (Tables 4.1.1 and 4.1.2). In the process of transcription, complementary RNA (message RNA or mRNA) is derived from the sense DNA strand by Watson-Crick base-pair recognition and binding. The antisense approach targets the initial sense complementary RNA strand (primary transcript) as well as many downstream sites that are available as RNA is metabolized in the process of protein production (e.g., splicing, transport, translation; Fig. 4.1.2). A strict definition of the antisense approach describes the inhibition of gene expression through targeting of a predetermined sequence in an RNA. There are several types of antisense approaches, which may be differentiated on the basis of their modes of action (Fig. 4.1.2). Antisense oligonucleotides primarily operate by sequence-specific binding to the targeted RNA. The resultant heteroduplex recruits a ubiquitous endonuclease, ribonuclease H (RNase H), to cleave the targeted RNA strand. Other potential antisense modes of action are by direct sequence-specific binding of the oligonucleotides to the targeted RNA (or merely occupying a site on the RNA) such that the function of essential RNA-binding proteins is disrupted or that secondary structure required for gene expression of the RNA is disrupted. A variety of distinct sites in the intermediary metabolism of RNA can potentially be targeted by antisense oligonucleotides with RNase H– dependent or –independent modes of action depicted in Figure 4.1.2.
4.1.4 Current Protocols in Nucleic Acid Chemistry
Figure 4.1.2
Informational drug-targeting sites. CAP, m7Gppp; W/C, Watson-Crick.
RNase L, like RNase H, is another cellular nuclease that cleaves RNA. Its endonucleolytic cleavage is triggered by binding to (2′-5′)linked polyadenylic sequences (Fig. 4.1.3). Torrence and co-workers have conjugated the RNase L activating moiety to oligonucleotides to provide sequence-specific cleavage of targeted RNA (Torrence et al., 1994). Ribozymes are RNA molecules with catalytic RNA-cleaving activity (Cech, 1986) and thus represent another type of antisense oli-
gonucleotide. These are typically much longer RNA molecules that contain both a catalytic region (which can cleave another RNA segment) and an adjacent sequence or two sequence arms that allow Watson-Crick base-pair binding to the target. A variety of ribozymes have been described that differ primarily in their threedimensional structures and lengths (reviewed by Rossi, 1998). The sequence-specific cleavage of a predetermined RNA target is accomplished without assistance from any protein.
NH2 N O O O HO P O P O P O O O O
O
N
N
N NH2
HO
O O P O O
N O
N
N N NH2
HO
O O P O O
HO
Figure 4.1.3
(2′-,5′)-Adenylic oligonucleotides.
N N
O O
N N
oligonucleotide
Synthesis of Modified Oligonucleotides and Conjugates
4.1.5 Current Protocols in Nucleic Acid Chemistry
A X = O,S,C 5' Heterocycle
A,C,G,T
X
Sugar
3' O Linkage
R 2'-Position
O P O O
O
4'
Connection sites
O
R
A,C,G,T Pendants
Replace sugar-phosphate (e.g., amide linkage, PNA) Subunits of oligonucleotide to modify
B Phosphorous modifications a. b. O O O P O O P S O O
O O P BH3 O
d.
e. O O O P NHR O P OR O O
O O P CH3 O h.
g.
f.
c.
O O P Se O
O S P S O
Linkage modifications i. j. Me N O
NH O P O O
HN
n.
m.
l.
k.
CH2 O P O
S
O
O
S
O
O
Sugar modifications p.
o.
r.
q. base
base O
s.
O
O OMe
F
base
base
O
O
O
OMe NH2
t. u.
base
O
v.
base
O
base
O
base O
O
O O
w.
O
NMe2
x. base
O
N NMe2 O P NHR
O
O NMe2 base
O O
Heterocycle modifications y. Me
z. O
NH2 Me
NH N cc. Me
aa.
O
O
N N
N
O
NH
NH2
NH2
Connection modification ff.
O
N
NH2
N
O
N
O
Sugar-phosphate replacement gg.
base N
O
Modified Oligonucleotides for Chemotherapeutic Applications
N
NH
N N
N NH2
N
ee.
dd.
NH2
I
N
N
O
N
bb.
NH2
N
N
O
O base HN
base N
O
4.1.6 Current Protocols in Nucleic Acid Chemistry
Oligonucleotides may also inhibit gene expression at the DNA level by several antigene approaches. The most often examined approach is triple strand formation by oligonucleotides binding in the major groove of dsDNA via several Hoogsteen base-pairing rules (Helene, 1993). A less often examined approach is the use of oligonucleotides to bind via Watson-Crick base-pairing rules to a single strand of DNA available from the formation of a transcription bubble or locally open-chain site of dsDNA. Utilizing nucleic acids as decoys to compete with natural cis-acting sites on dsDNA for essential regulatory proteins is referred to as the sense approach. In this case, synthetic oligonucleotides (typically dsDNA) are designed to bind to proteins in a sequence-specific manner. In another protein-binding approach, aptamers, derived from nucleic acid selection processes (see Chapter 9), can specifically target regulatory proteins. Common steps to all selection methods for nucleic acid–binding species are: (1) generation of a large pool of sequence diversity from chemically synthesized DNA pools, (2) transformation of the pools by enzymatic manipulations such as polymerase chain reaction (PCR) or in vitro transcription, (3) selection of functional shapes, and (4) amplification. As each RNA sequence folds into a distinct three-dimensional shape, and because of the large number of structures (millions) generated, complementary binding to a protein target may occur, providing useful biological properties (reviewed by Bacher and Ellington, 1998). Of those drug discovery approaches based on oligonucleotides, the antisense efforts are clearly the most advanced, in that twenty three first-generation phosphorothioate oligonucleotides have entered human clinical trials and one antisense oligonucleotide has achieved FDA approval (Table 4.1.4). Much less success has been accomplished by targeting DNA with
triple-strand-forming oligonucleotides, antisense ribozymes, and RNase L–modified oligonucleotides. From a chemical point of view, these and other less successful approaches listed in Tables 4.1.1 and 4.1.2 suffer from not being amenable to the readily available, firstgeneration backbone-modified oligonucleotides such as methylphosphonates, amidates, phosphorothioates, and α oligomers (Fig. 4.1.4B). Success in these drug discovery approaches, assuming that the biological rationale is valid, will require much more intensive chemical efforts.
MODIFICATIONS AND DRUG PROPERTY ALTERATIONS A diverse range of modifications at all possible modification sites of an oligonucleotide have been reported (reviewed in Cook, 1998a). Figure 4.1.4A illustrates a dimer of an oligonucleotide depicting possible modifications for enhancing oligonucleotide drug properties. These include alterations of heterocycles, carbohydrates, linkages (backbones), and connection and conjugation sites, as well as the complete removal of the sugar-phosphate backbone. Most of the positions available in a GC or AT dimer (∼26 positions for each dimer) that do not directly interfere with Watson-Crick base-pair hydrogen bonding have been modified at one time or another. The nucleobases or heterocycles of nucleic acids provide the recognition points for Watson-Crick base pairing and must maintain this specific interaction in any modification. Thus, the scope of heterocyclic modifications is quite limited; this is verified by the fact that only five or six modified heterocycles have demonstrated useful oligonucleotide drug properties. These can be characterized as lipophilic modifications at C5 of pyrimidines and C7 of 7deazapurines. Enhanced stacking has been suggested to be the reason for increased binding
Figure 4.1.4 Oligonucleotide modification. (A) Dimer structure showing potential modification sites. (B) Examples of oligonucleotide modifications: (a) natural phosphate diester; (b) chiral phosphorothioate; (c) chiral methyl phosphonate; (d) chiral phosphoramidate; (e) chiral phosphate triester; (f) chiral boranophosphate; (g) chiral phosphoroselenoate; (h) phosphorodithioate; (i) methylenemethylimino (MMI); (j) 3′-amide; (k) 3′ achiral phosphoramidate; (l) 3′ archiral methylene phosphonate; (m) thioformacetal; (n) thioethyl ether; (o) 2′-fluoro; (p) 2′-O-methyl; (q) 2′-O-(3amino)propyl; (r) 2′-O-(2-methoxy)ethyl; (s) 2′-O-2-(N,N-dimethylaminooxy)ethyl (DMAOE); (t) 2′O-2-[2-(N,N-dimethylamino)ethyloxy]ethyl (DMAEOE); (u) 2′-O-N,N-dimethylacetamidyl; (v) N-morpholinophosphordiamidate; (w) hexose nucleic acid; (x) locked nucleic acid (LNA); (y) 5-propynyluracil-1-yl; (z) 5-methylcytosin-1-yl; (aa) 2-aminoadenin-9-yl; (bb) 7-deaza-7-iodoadenin-9-yl; (cc) 7-deaza-7-propynyl-2-aminoadenin-9-yl; (dd) phenoxazinyl; (ee) phenoxazinylG-clamp; (ff) α−deoxyribofuranosyl; (gg) peptide nucleic acid (PNA). See Cook (1999). Current Protocols in Nucleic Acid Chemistry
Synthesis of Modified Oligonucleotides and Conjugates
4.1.7
Modified Oligonucleotides for Chemotherapeutic Applications
affinity with this type of heterocycle modification. 2-Aminoadenine represents another type of modification that provides enhanced binding affinity. In this case, an additional hydrogen bond is formed to uracil, providing significant increases in melting temperature (Tm). All of these modifications—positioned to lie in the major or minor groove of the heteroduplex—do not affect sugar conformation of the heteroduplex, and do not provide useful nuclease resistances, but will support an RNase H cleavage mechanism. Modifications in the ribofuranosyl moiety have provided the most value in the quest to enhance oligonucleotide drug properties. In particular, certain 2′-O modifications have greatly increased binding affinity, increased nuclease resistance, and altered pharmacokinetics, and are potentially less toxic than phosphorothioate oligonucleotides (Cook, 1998a). Preorganization of the sugar into a 3′-endo pucker conformation is responsible for the increased binding affinity. Unfortunately, no sugar modification has been reported that is useful in supporting RNase H cleavage. Linkage or backbone modifications involve changes to the nonbridging oxygen atoms, such as phosphorothioate (P=S), methylphosphonate (Me-P), phosphoramidate (N-P), and others shown in Figure 4.1.4B, as well as extensive changes in which the entire four-atom linkage between sugar moieties is replaced. The most useful linkage-replacement modification, MMI (Fig. 4.1.4B), provides greatly enhanced binding affinities, removes the nuclease-cleavable phosphodiester linkage and the chirality of the phosphorothioate linkage, and allows adjustments of oligonucleotide lipophilicity by controlling the negative charge. Of the linkage changes, only phosphorothioates induce RNase H cleavage. Several examples of connection modifications in oligonucleotides are α-nucleoside linkage (reverse connection of the heterocycle at C1′) and (2′-5′)-phosphodiester bonds (in place of the normal (3′-5′)-phosphodiester bond). Peptide nucleic acid (PNA) is an example of the complete removal of the sugar-phosphate backbone and replacement with a peptide linkage. In addition to the above modifications, pendants or conjugation groups can be attached at many positions in the various subunits to enhance drug properties. The antisense concept implies well-defined structural requirements for the oligonucleotide ligand that binds to a reasonably characterized RNA receptor. Although this knowledge of
rather precise binding is of great value and certainly sets this drug discovery approach apart from current approaches, additional modifications are limited to modifications that do not interfere with Watson-Crick basepairing rules. Modifications of oligonucleotides may be expected to address essentially every facet of antisense drug properties. Biophysical and biochemical properties that may be affected by modifications include binding affinity, basepair specificity, nuclease resistance, chemical stability, lipophilicity, solubility, endonucleolytic cleavage of the RNA of a heteroduplex, biological pharmacokinetic properties, toxicological properties, and pharmacological properties. Drug properties that will be affected by altering biophysical and biochemical parameters include the general areas of pharmacokinetics, pharmacodynamics, and toxicology. In addition, certain modifications may represent cost and proprietary patent advantages. Modifications of short strands of DNA and RNA for drug purposes should be rationally directed by drug property deficiencies of the parent or first-generation oligonucleotides (Table 4.1.5). Biological deficiencies are exposed by pharmacologic, pharmacokinetic, and toxicologic studies of the parent oligonucleotides at the biophysical, biochemical, in vitro, in vivo, and clinical levels. In addition, as modified oligonucleotides move from discovery into the development pipeline, chemical development deficiencies in, for example, larger-scale synthesis, purification, and analytical processes will emerge that must be resolved. Finally, in contemplating research in oligonucleotides, one should search the patent literature in addition to examining the traditional scientific literature, as considerable proprietary positions of various aspects of the technology have been achieved. Hopefully, the days of incorporating nucleosides, novel or known, into oligonucleotides based merely on availability are over. This nonrational approach was of value in the beginning of oligonucleotide structure-activity modifications simply to develop an information base. Now that more knowledge is available about what is required to achieve enhanced properties, current and future oligonucleotide modifications and studies of their structure-activity relationships should be soundly and rationally based on the need to resolve one or more of the deficiencies listed in Table 4.1.5, or to overcome emerging drug process or production problems.
4.1.8 Current Protocols in Nucleic Acid Chemistry
Table 4.1.5
Limitations of First-Generation P=Sa Oligonucleotides
Area
Limitation
Pharmacodynamics
Low affinity per nucleotide unit: low potency limited sites to target Inhibition of RNase H at high concentrations RNase H mechanism implicated Limited oral bioavailability Limited blood-brain barrier penetration Dose-dependent pharmacokinetics Release of cytokines Complement-associated effects on blood pressure? Clotting effects Accumulation in kidneys?
Pharmacokinetics
Toxicologics
aP=S, phosphorothioate.
OPTIMIZATION OF BINDING AFFINITY AND NUCLEASE RESISTANCE Oligonucleotide Binding Affinities Based on the ligand-receptor theory of pharmacological activity, increasing the affinity of an oligonucleotide for its RNA target should increase potency. A relatively simple physicochemical experiment is employed to determine the level of binding and specificity of a modified oligonucleotide. A complementary oligonucleotide is the simplest target to which an antisense oligonucleotide can hybridize. Thus, using length-matched complementary oligonucleotides at stoichiometric concentrations in a medium that mimics the intracellular environment as closely as possible (salt, pH, temperature, concentration), melting curves are employed to determine how tightly a modified oligonucleotide binds to its complement. This involves a spectrophotometric analysis to measure absorbance versus temperature. The melting temperature (Tm—the temperature at which the mixture is half duplex and half single-stranded) is determined (Freier et al., 1992). Oligonucleotide affinity, as measured by melting curves, increases with the length of the oligonucleotide-RNA heteroduplex. Thus, 15to 25-mers are typically used in antisense experiments rather than shorter oligonucleotides, which may have Tm values close to or below physiological temperature and, therefore, may only form low levels of the required heteroduplex. A few short oligomers (12-mers or less) have exhibited interesting biological activity, and these require modifications leading to high affinity per nucleotide unit. One interesting
example was the report of potent and selective inhibition of gene expression by phosphorothioates (7- and 8-mers) containing only 5-propynyl-substituted uracil and cytosine bases (Fig. 4.1.4B; Wagner et al., 1996). Furthermore, several recent studies correlate binding affinities of a series of 2′-O-modified oligonucleotides with increased in vitro and in vivo activity (Kawasaki et al., 1993; Morvan et al., 1993; Monia et al., 1993, 1996; Altmann et al, 1996a,b,c, 1997). An extensive comparative listing of binding affinities of 2′-O modifications was provided by Freier and Altmann (1997).
Oligonucleotide Nuclease Resistance A number of in vivo pharmacokinetic studies in several animal species indicate that P=S oligonucleotides are not as stable as initially thought (Zhang et al., 1996; Nicklin, 1998). Although the stability of P=S oligonucleotides may be sufficient for many drug applications, greater stability will be helpful in expanding dosage regimens (longer duration of action relates to less frequent dosing) and in developing oral bioavailability. Additionally, less degradation of modified oligomers will minimize metabolite toxicity. In summary, the preponderance of published antisense biological data suggests that oligonucleotides with higher binding affinities and greater stabilities towards nucleases are important medicinal chemistry objectives.
STANDARDS ESTABLISHED BY RECENT SAPR STUDIES The intense oligonucleotide research performed in the 1990s has provided a remarkable enhancement of several of the desired antisense
Synthesis of Modified Oligonucleotides and Conjugates
4.1.9 Current Protocols in Nucleic Acid Chemistry
Modified Oligonucleotides for Chemotherapeutic Applications
drug properties. The results achieved with modified oligonucleotides concerning binding affinity, base-pair specificity, nuclease resistance, and support of RNase H cleavage of the targeted RNA is impressive. One should consider the level of binding affinity (as represented by Tm), nuclease resistance (as represented by t1/2), and support of an RNase H mechanism that a new modification should possess in order to be of interest as a potential drug. A number of “winners” have been identified (Fig. 4.1.4B) and discussed in several reviews (Cook, 1998a,b). Certain modifications of oligonucleotides provide an increase in Tm of >1.5°C/modification (°C/mod) relative to a P=S oligonucleotide (∼1.0°C/mod relative to an unmodified P=O oligonucleotide), and nuclease resistance (t1/2) of >24 hr with snake venom phosphorodiesterase (SVPD; about the same as for P=S). In view of these values, the author suggests that a novel modification should exhibit a Tm >1.5°C/mod compared to its P=S oligonucleotide parent. In evaluation of binding properties, the modified oligonucleotides should be hybridized with an RNA complement, as this is the receptor required for the antisense approach. A clear correlation between Tm values derived from hybridization to a DNA complement or to an RNA has not been established. Also, correlations of Tm values have not been established for oligonucleotides having just one modification (point or pendent modification) or several modifications distributed throughout the sequence, or for those having a contiguous placement of the modification in the sequence (Cook, 1991; Lesnik et al., 1993; Buhr et al., 1996; Matteucci and Krosigk, 1996). Since the application of a modification will likely require its uniform placement in the sequence, or at least several contiguous bases in a row for a gapmer strategy (described below, Cook, 1993), binding affinity measurements (Tm) should be taken with these types of modified oligonucleotides. Furthermore, the modification must not compromise base-pair specificity. In this regard, information from a number of papers suggests that base-pair specificity actually increases as Tm values are increased. A specificity that is comparable to base mismatches of a P=S phosphorothioate would appear to be a useful standard. Nuclease resistance of a novel modification in a P=O oligonucleotide backbone should at least be at the level of uniformly modified P=S oligonucleotides. Given that measurements of t1/2 values, unlike Tm measurements, are deter-
mined using several procedures (e.g., by incubation with heat-inactivated fetal bovine serum, purified enzymes such as SVPD, cell or tissue extracts; or by in vivo dosing followed by extraction and analysis) under different conditions, the use of P=S oligonucleotide controls (standards) is necessary. In addition, several concentrations of nuclease should be employed to identify and minimize complications leading to enzyme inhibition (Cummins et al., 1995). Half-lives of ∼24 hr are often reported for P=S oligonucleotides in SVPD assays. When the 3′ end of an oligonucleotide is modified to have sufficient resistance to 3′-exonucleases, endonucleolytic cleavage becomes evident. Thus, modifications should also protect against endonucleases. In a gapmer strategy, this is accomplished by a phosphorothioate gap, which will also support an RNase H cleavage mechanism. Nuclease resistance of a modified oligomer, if not provided by the modification, may in many cases be enabled by employing a P=S oligonucleotide backbone. In considering the relative importance of the nuclease resistance of an antisense oligomer and its affinity level for its RNA target, recent biological results suggest that it may be more important to enhance the stability of an oligonucleotide than its binding affinity (Crooke et al., 1996a). Modifications that provide highbinding oligonucleotides with low nuclease resistance have not provided significant biological activity, whereas oligonucleotides such as phosphorothioates have. Although some modifications provide high binding affinities and high nuclease resistance, they may not exhibit useful antisense activities because they do not support an RNase H mechanism. A modification that supports an RNase H mode of action and provides high Tm and nuclease resistance t1/2 values has not been reported. Thus, an ideal oligonucleotide modification would provide an oligomer that hybridizes to target RNA with high binding affinity and specificity, would be stable to nucleolytic degradation, and would allow RNase H cleavage of the RNA target. This has led to the theory that to optimize the antisense activity of an oligomer, a combination of oligonucleotide modifications will be required (Cook, 1991, 1993). As noted above, after high-binding, nuclease-resistant 2′-O-modified oligonucleotides were developed, it was rather disappointing that oligomers uniformly modified were inactive or less active than their first-generation parent phosphorothioates. It is now well known that uniformly 2′-O-modified oligonucleotides do
4.1.10 Current Protocols in Nucleic Acid Chemistry
not support RNase H cleavage: the 2′-O-modified oligonucleotide-RNA heteroduplex presents a structural conformation that is recognized by the enzyme but is not cleaved (Crooke et al., 1995; Lima and Crooke, 1997). This lack of activity has led to the development of a chimeric strategy (gapmer technology; Cook, 1993; Monia et al., 1993; Yu et al., 1996). This approach focuses on the design of highbinding, nuclease-resistant antisense oligonucleotides that are “gapped” with a contiguous sequence of 2′-deoxyribonucleoside phosphorothioates (2′-deoxy/PS)(Fig. 4.1.5). On hybridization to target RNA, a heteroduplex is presented that supports RNase H cleavage of the RNA strand. The stretch of the modified oligonucleotide-RNA heteroduplex that is recognized by RNase H may be placed anywhere within the modified oligonucleotide. The modifications in the flanking regions of the gap should not only provide nuclease resistance to exo- and endonucleases, but should also not compromise binding affinity and base-pair specificity (Hoke et al., 1991). Modifications of the phosphorus atom of the natural phosphodiester linkage (producing methylphosphonates, phosphorothioates, and phosphoramidates) destabilize heteroduplexes −0.7° to −1.5°C per modification (Fig. 4.1.4B; Agrawal et al., 1990; Dagle et al., 1991; Guinosso et al., 1991). The decreased binding affinity of these modified oligonucleotides could be expected to reduce antisense effectiveness. In the case of chimeric 2′-O-methyl- or 2′fluoro-modified oligonucleotides, an enhancement in the binding affinity of ∼2.0° to 2.3°C (compared to P=S oligonucleotides) for each modification is obtained (Sproat et al., 1989; Guinosso et al., 1991; Miller et al., 1991; Kawasaki et al., 1993). However, it is now clear that 2′-O-methyl- and 2′-F-modified DNA are not sufficiently nuclease resistant to have antisense value as P=O backbones (Sproat et al., 1989; Morvan et al., 1993; Sproat and Lamond, 1993; Sands et al., 1995; Prasmickaite et al., 1998). The potential problem in this area can be circumvented by the use of 2′-O-methyl- or 2′-F-modified phosphorothioates in the flanking regions (doubly modified; Miller et al., 1991; Kawasaki et al., 1993). Flank
More recent research has focused on 2′-O modifications such as methoxyethyl (Altmann et al., 1996a,b,c) and aminopropyl (Griffey et al., 1996), which not only provide relatively high binding affinities but also a level of nuclease resistance that allows replacement of thiophosphates with natural phosphodiester linkages. With a favorable combination of Tm and t1/2, 2′-O modifications can be employed in the chimeric strategy (gapmer technology; Fig. 4.1.5), which allows a significant portion of P=S linkages to be replaced with P=O linkages. Just how many sulfurs can be replaced depends on the length of the oligomer and the gap size or RNase H cleavage site. Typically, a 21-mer with a 7-nucleotide gap has 65% of the P=S linkages replaced with P=O linkages. As noted in the discussion of limitations of P=S oligonucleotides, reduction of the sulfur content in a P=S oligonucleotide could have important implications in the pharmacokinetic and pharmacodynamic properties as well as the toxicity profiles of oligonucleotides (Altmann et al., 1996a). A very important aspect of gapmer technology is that the gap or RNase H cleavage site must be protected from endonucleolytic cleavage. Phosphodiester linkages and even an alternating P=S/P=O motif do not provide a useful level of nuclease resistance for biological activity (Dagle et al., 1991; Hoke et al., 1991). The recently reported lack of activity of a “gapped” 3′-amidate phosphodiester is likely due to endonuclease degradation (Heidenreich et al., 1997). Phosphorothioates are the only useful modification to allow a reasonable combination of binding affinity and nuclease resistance and also support an RNase H mechanism. Hence, as noted above, most antisense activities require an RNase H mechanism, which in turn requires sulfur in the form of thiophosphate somewhere in the chimeric molecule for nuclease resistance. An additional issue that should be kept in mind is the impact that oligonucleotide modifications may have on the cost of future antisense drugs. Again, as a standard, the cost of P=S oligonucleotides should be considered. The cost of phosphorothioates has been dramatically reduced due to improvements in the process, cost reduction of key reagents, and eco-
Gap
5'- 2'-Modifications/PS or PO
Flank 2'-Modifications/PS or PO
2'-deoxy/PS
Figure 4.1.5
Gapmer technology. PO, phosphodiester; PS, phosphorothioate.
-3'
Synthesis of Modified Oligonucleotides and Conjugates
4.1.11 Current Protocols in Nucleic Acid Chemistry
nomics of larger-scale syntheses. As of 1999, the cost of materials for a 20-mer P=S oligonucleotide was <$300/gram; it is projected to decline to <$50/gram as larger quantities are required. The 2′-O-methoxyethyl-modified oligonucleotides are derived from ribonucleosides and should eventually be substantially less expensive than modified oligonucleotides derived from deoxyribofuranosyl nucleosides. P=S oligonucleotide antisense drugs are expected to be cost competitive when considering parenteral (intravenous) treatment three times/week with a 1 mg/kg dose. Modified oligonucleotides must be cost effective and will need to be less expensive to synthesize (e.g., shorter), or provide a greater therapeutic index (increased potency) or other important advantages to offset an increase in the cost of synthesis. When considering research to modify oligonucleotides, the author believes it is important to know the extent to which proprietary protection of a modification can be obtained. Important patent positions for many types of modifications have been established during the 1990s as designing drugs from oligonucleotides has become of interest (Crooke et al, 1996b; Sheffery and Gordon, 1996; Craig et al., 1997).
GENERAL GUIDELINES FOR ENHANCING OLIGONUCLEOTIDE DRUG PROPERTIES
Modified Oligonucleotides for Chemotherapeutic Applications
The excitement roused by the prospect of using oligonucleotides as drugs stems from the fact that, unlike other drug discovery approaches, they are informational materials— i.e., chemicals having a specific set of rules (the Watson-Crick base-pairing rules) that clearly govern their binding to a specific nucleic acid receptor. In addition, the fact that DNA and RNA molecules are, compared to proteins, relatively new molecular targets is of great interest. The first test in a SAPR study to determine whether a modified oligonucleotide is worth pursuing is whether it maintains its sequence specificity according to Watson-Crick rules (Fig. 4.1.6). A newly modified oligonucleotide that does not possess an acceptable level of specificity (comparable to that of the P=S oligonucleotide) is an immediate failure. The next step is to determine how tightly the modified oligonucleotide binds to its target nucleic acid. As noted, the binding affinity of a modified oligomer is a physicochemical property, determined by measuring the oligomer’s sequence-
specific interaction with its length-matched RNA complement. The hybridization or melting process is performed under a rather standard set of conditions designed to mimic an intracellular environment (Freier et al., 1992). Because this calculation is done under artificial conditions, it may not accurately represent the binding of an oligomer to a native RNA inside a cell. The next essential property of an oligonucleotide is sufficient resistance to degradation by exo- and endonucleolytic plasma and tissue nucleases (Fig. 4.1.6). Nuclease resistance is determined in various assays, such as incubating the oligonucleotide in heat-inactivated fetal bovine serum, cellular extracts, or purified exonucleases or endonucleases (also see Standards Established by Recent SAPR Studies). These results are also likely to differ from the stability of an oligomer in vivo. However, these Tm and t1/2 assays do allow SAPR studies to proceed, and thus provide reasonable methods to compare various oligonucleotide modifications at an early developmental stage. This is quite different from traditional drug discovery approaches.
CONJUGATED OLIGONUCLEOTIDES The sugar, heterocycle, and backbone (linkage) subunit modifications as depicted in Figure 4.1.4A are core modifications that greatly enhance binding affinities and nuclease resistance of antisense oligonucleotides. To enhance other antisense drug properties of an optimized, core-modified oligonucleotide, a variety of molecules (pendants) have been attached (conjugated) in a point-modification motif (i.e., only one pendant in an antisense oligonucleotide). Pendant modifications have primarily been directed to enhancing oligonucleotide uptake. Other potential applications of pendants include increased solubility, lipophilicity, and means to attach synthetic cleaving agents, intercalators (for improvements in binding affinity), and cross-linking and alkylating groups (see UNITS 4.2 & 4.3). Several reviews have discussed oligonucleotide pendants (Cook, 1991, 1993; Manoharan, 1993; De Mesmaeker et al., 1995).
PERSPECTIVE One P=S oligonucleotide drug (Vitravene) is available as of 1999, and others will follow in the next several years. However, to continually improve this novel and exciting drug class, and to overcome certain limitations, structural changes are required. In the 1990s a diverse range of modifications, at all possible modifi-
4.1.12 Current Protocols in Nucleic Acid Chemistry
• Synthesis of novel or known nucleosides • Synthesis of novel amidites with appropriate protection • Development of oligomerization protocol process/analysis • Synthesis of modified oligonucleotide in standard sequences • Synthesis of length-matched complementry RNA sequences and single base mismatches and appropriate control sequences • Obtain binding affinities (change in Tm in degrees/modification) of matched and mismatched modified oligonucleotide
Sequence-specificity or binding affinity of modification below standards
Abandon modification
Sequence-specificity and binding affinity of modification improved compared to standards
• Prepare appropriate modified oligonucleotides for nuclease studies • Determine relative nuclease resistance
Nuclease-resistance of modified oligonucleotides less than standards
Consider additional modifications, e.g., combination of modifications, for enhancing nuclease resistance while maintaining binding affinity
Modified oligonucleotide possess greater nuclease-resistance than standards
Examine modified oligonucleotide with target sequence (tissue culture reduction of message and/or protein)
Advanced pharmacology, pharmacokinetics and toxicology studies
Figure 4.1.6
General pathway for structure-activity/property relationship studies.
cation sites of an oligonucleotide (Fig. 4.1.4A), has been reported. This application of traditional medicinal chemistry (SAPR studies) to drug discovery in antisense oligonucleotides, and in oligonucleotides in general, has answered many important questions. For example, as a result of this rather intense effort, modifications are now known that stabilize oligonucleotides towards nucleolytic degradation (e.g., 2′-O-methoxyethyl and 2′-O-aminopropyl), that greatly enhance binding affinities while maintaining base-pair specificity (e.g., 2′-O-methoxyethyl), and that support endonucleolytic cleavage by RNase H (e.g., 5propynyl pyrimidines). Although these are biochemical and biophysical properties, a large volume of cellular and animal studies support the notion that enhancing these properties correlates with enhanced antisense biological ac-
tivity in vivo. Unfortunately, a single modification that provides high binding affinity nuclease-resistance and support of an RNase H mechanism is not available. A modification of this nature is of current interest in the antisense approach. It is also known that changing the structure of phosphorothioate oligonucleotides provides an opportunity to alter their pharmacokinetic profile. Structural changes that remove sulfur (as thiophosphate) and/or change lipophilicity (e.g., by 2′-O modifications) have resulted in more favorable toxicity profiles (Altmann et al., 1996a). Although research has revealed these important antisense properties (and there may be many more to learn about), and has shown how to control them, it has not yet determined the optimum values at which these modifications should be aimed. In addition, the lack of
Synthesis of Modified Oligonucleotides and Conjugates
4.1.13 Current Protocols in Nucleic Acid Chemistry
Modified Oligonucleotides for Chemotherapeutic Applications
antisense oligonucleotides that are orally available and/or penetrate the blood-brain barrier represents the most important deficiency of antisense oligonucleotides. Recent reports of antisense P=S oligonucleotides doubly modified at the 3′ and 5′ ends with 2′-O-methyl or 2′-O-methoxyethyl—to provide a high level of nuclease resistance—have provided encouraging results suggesting that these pharmacokinetic deficiencies will soon be solved by appropriate chemical modifications (Agrawal et al., 1995). One should be aware of the level of accomplishments achieved in oligonucleotide medicinal chemistry research in the course of the 1990s. Binding affinities, nuclease resistance, support of RNase H, and cost of synthesis have been discussed in this unit, and they should be considered (as standards) before initiating or continuing oligonucleotide modification research. In addition, understanding the proprietary patent positions that have been established is an important research consideration. The author believes that, at this stage of oligonucleotide medicinal chemistry, it is highly unlikely that a single modification will be discovered that will significantly affect all of the important drug properties described above. The types of modified oligonucleotides currently being pursued (going beyond P=S oligonucleotides) possess a combination of modifications, and this trend will certainly continue as pendants will be conjugated to oligonucleotides with optimized core subunits to obtain a “completely” optimized oligonucleotide drug. The current “winners,” or the first modifications most likely to be incorporated into antisense oligonucleotides that will undergo clinical trials, are the RNA mimics 2′-Omethoxyethyl and 2′-O-aminopropyl, and the backbone modification MMI (Fig. 4.1.4B). These will likely be utilized in a gapmer strategy. However, efforts to prepare uniform modifications, such as RNA mimics (2′-O-modifications), MMI, and PNA are of considerable interest, in that reliance on RNase H for a mode of action would not be required. In addition to these modifications that act either by direct binding (RNase H independent) or via RNase H, the author believes that subunit pendant modifications (such as cholesterol conjugates and folic acid conjugates) will become increasingly important for optimizing multiply modified oligonucleotides.
LITERATURE CITED Agrawal, S., Mayrand, S.H., Zamecnik, P.C., and Pederson, T. 1990. Site-specific excision from RNA by RNase H and mixed-phosphate-backbone oligodeoxynucleotides. Proc. Natl. Acad. Sci. U.S.A. 87:1401-1405. Agrawal, S., Zhang, X., Lu, Z., Zhao, H., Tamburin, J.M., Yan, J., Cai, H., Diasio, R.B., Habus, I., Jiang, Z., Iyer, R.P., Yu, D., and Zhang, R. 1995. Absorption, tissue distribution and in vivo stability in rats of a hybrid antisense oligonucleotide following oral administration. Biochem. Pharmacol. 50:571-576. Alderfer, J.L., Loomis, R.E., Soni, S.D., Sharma, M., Bernacki, R., and Hughes, R. Jr. 1985. Halogenated nucleic acids: Biochemical and biological properties of fluorinated polynucleotides. Polymeric Mater. Med. 32:125-138. Altman, S. 1989. Ribonuclease P: An enzyme with a catalytic RNA subunit. Adv. Enzymol. 62:1-36. Altmann, K.H., Dean, N.M., Fabbro, D., Freier, S.M., Geiger, T., Häner, R., Hüsken, D., Martin, P., Monia, B.P., Müller, M., Natt, F., Nicklin, P., Phillips, J., Pieles, U., Sasmor, H., and Moser, H.E. 1996a. Second generation of antisense oligonucleotides: From nuclease resistance to biological efficacy in animals. Chimia 50:168-176. Altmann, K.H., Kesselring, R., and Pieles, U. 1996b. 6′-Carbon-substituted carbocyclic analogs of 2′deoxyribonucleosides: Synthesis and effect on DNA/RNA du plex stability. Tetrahedron 52:12699-12722. Altmann, K.H., Fabbrot, D., Dean, N.M., Geiger, T., Monia, B.P., Muller, M., and Nicklin, P. 1996c. Second-generation antisense oligonucleotides: Structure-activity/relationships and the design of improved signal-transduction inhibitors. Biochem. Soc. Trans. 24:630-637. Altmann, K.H., Martin, P., Dean, N.M., and Monia, B.P. 1997. Second generation antisense oligonucleotides—inhibition of pkc-α and c-raf kinase expression by chimeric oligonucleotides incorporating 6′-substituted carbocyclic nucleosides and 2′-O-ethylene glycol substituted ribonucleosides. Nucleosides Nucleotides 16:917. Bacher, J.M. and Ellington, A.D. 1998. Nucleic acid selection as a tool for drug discovery. Drug Discovery Today 3:265. Bardos, T.J. and Ho, Y.K. 1978. Chemical and Enzymatic Methods in the Synthesis of Modified Polynucleotides. In Symposium on the Chemistry and Biology of Nucleosides and Nucleotides (R.E. Harmon, R.K. Robins, and L. Townsend, eds.) pp. 55-68. Academic Press, Orlando, Fla. Belikova, A.M., Zarytova, V.F., and Grivneva, N.I. 1967. Synthesis of ribonucleosides and diribonucleoside phosphates containing 2-chloroethylamine and nitrogen mustard residues. Tetrahedron Lett. 7:3557-3562. Buhr, C.A., Wagner, R.W., Grant, D., and Froehler, B.C. 1996. Oligodeoxynucleotides containing C-7 propyne analogs of 7-deaza-2′-deoxyguanosine and 7-deaza-2′-deoxyadenosine. Nucl. Acids Res. 24:2974-2980.
4.1.14 Current Protocols in Nucleic Acid Chemistry
Cech, T.R. 1986. RNA as an enzyme. Sci. Am. 255:64-75. Chandra, P. and Bardos, T.J. 1972. Inhibition of DNA polymerases from RNA tumor viruses by novel template analogs. Partially thiolated polycytidylic acid. Res. Commun. Chem. Pathol. Pharmacol. 4:615-622. Cohen, J.S. 1991. Informational drugs: A new concept in pharmacology. Antisense Res. Dev. 1:191-193. Connell, C., Fung, S., Heiner, C., Bridgham, J. Chakarian, V., Heron, E., Jones, R., Menchen, S., Mordan, W., Raff, M., Recknor, M., Smith, L., Springer, J., Woo, S., and Hunkapiller, M. 1987. Automated DNA sequence analysis. BioTechniques 5:342. Cook, P.D. 1991. Medicinal chemistry of antisense oligonucleotides—future opportunities. AntiCancer Drug Design 6:585-607. Cook, P.D. 1993. Medicinal chemistry strategies for antisense research. In Antisense Research and Applications (S.T. Crooke and B. Lebleu, eds.) pp. 149-187. CRC Press, Boca Raton, Fla. Cook, P.D. 1998a. Antisense medicinal chemistry. In Handbook of Experimental Pharmacology (S.T. Crooke, ed.) pp. 51-101. Springer-Verlag, Heidelberg, Germany. Cook, P.D. 1998b. Second generation antisense oligonucleotides: 2′-Modifications. Annu. Rep. Med. Chem. 33:313. Cook, P.D. 1999. Making drugs out of oligonucleotides: A brief review and perspective. Nucleosides Nucleotides 18:1141-1162. Craig, A., Vanstone, D., and Sudhir, A. 1997. Patent strategies in the antisense oligonucleotide based therapeutic approach. Expert Opin. Ther. 7:1175. Crooke, S.T., Lemonidis, K.M., Neilson, L., Griffey, R., Lesnik, E.A., and Monia, B.P. 1995. Kinetic characteristics of Escherichia coli RNase H1: Cleavage of various antisense oligonucleotideRNA duplexes. J. Biochem. 312:599-608. Crooke, S.T., Graham, M.J., Zuckerman, J.E., Brooks, D., Conklin, B.S., Cummins, L.L., Greig, M.J., Guinosso, C.J., Kornbrust, D., Manoharan, M., Sasmor, H.M., Schleich, T., Tivel, K.L., and Griffey, R.H. 1996a. Pharmacokinetic properties of several novel oligonucleotide analogs in mice. J. Pharmacol. Exp. Ther. 277:923-937. Crooke, S.T., Bernstein, L.S., and Boswell, H. 1996b. Progress in the development and patenting of antisense drug discovery technology. Expert Opin. Ther. 6:855-870. Cummins, L.L., Owens, S.R., Risen, L.M., Lesnik, E.A., Freier, S.M, McGee, D., Guinosso, C.J., and Cook, P.D. 1995. Characterization of fully 2′-modified oligoribonucleotide hetero- and homoduplex hybridization and nuclease sensitivity. Nucl. Acids Res. 23:2019-2024. Dagle, J.M., Andracki, M.E., DeVine, R.J., and Walder, J.A. 1991. Physical properties of oligonucleotides containing phosphoramidate-modified internucleoside linkages. Nucl. Acids Res. 19:1805-1810.
De Clercq, E., Eckstein, F., and Merigan, T.C. 1969. Interferon induction increased through chemical modification of a synthetic polyribonucleotide. Science 165:1137-1139. DeMesmaeker, A., Haner, R., Martin, P., and Moser, H.E. 1995. Antisense oligonucleotides. Acc. Chem. Res. 28:366-374. Ellington, A.D. and Szostak, J.W. 1990. In vitro selection of RNA molecules that bind specific ligands. Nature 346:818-822. Freier, S.M. and Altmann, K.-H. 1997. The ups and downs of nucleic acid duplex stability: Structure-stability studies on chemically-modified DNA: RNA duplexes. Nucl. Acids Res. 25:4429-4443. Freier, S.M., Lima, W.F., Sanghvi, Y.S., Vickers, T., Zounes, M., Cook, P.D., and Ecker, D.J. 1992. Thermodynamics of antisense oligonucleotide hybridization. In Gene Regulation: Biology of Antisense RNA and DNA, Vol. 1 (Series: Molecular Cellular Biology) (R.P. Erickson and J.G. Izant, eds.) pp. 95-107. Raven Press, New York. Griffey, R.H., Monia, B.P., Cummins, L.L., Freier, S., Greig, M.J., Guinosso, C.J., Lesnik, E., Manalili, S.M., Mohan, V., Owens, S., Ross, B.R., Sasmor, H., Wancewicz, E., Weiler, K., Wheeler, P.D., and Cook, P.D. 1996. 2′-O-aminopropyl ribonucleotides: A zwitterionic modification that enhances the exonuclease resistance and biological activity of antisense oligonucleotides. J. Med. Chem. 39:5100-5109. Guinosso, C.J., Hoke, G.D., Freier, S.M., Martin, J.F., Ecker, D.J., Mirabelli, C.K., Crooke, S.T., and Cook, P.D. 1991. Synthesis and biophysical and biological evaluation of 2′-modified antisense oliogonucleotides. Nucleosides Nucleotides 10:259-262. Heidenreich, O., Gryaznov, S., and Nerenberg, M. 1997. RNase H-independent antisense activity of oligonucleotide N3′-P5′ phosphorothioates. Nucl. Acids Res. 25:776. Helene, C. 1993. Control of gene expression by triple-helix-forming oligonucleotides: The antigene strategy. In Antisense Research and Applications (S.T. Crooke and B. Lebleu, eds.) pp. 375-385. CRC Press, Boca Raton, Fla. Hoke, G.D., Draper, K., Freier, S.M., Gonzalez, C., Driver, V.B., and Zounes, M.C. 1991. Effects of phosphorothioate capping on antisense oligonucleotide stability, hybridization and antiviral efficacy versus herpes simplex virus infection. Nucl. Acids Res. 19:5743. Kawasaki, A.M., Casper, M.D., Freier, S.M., Lesnik, E.A., Zounes, M.C., Cummins, L.L., Gonzalez, C., and Cook P.D. 1993. Uniformly modified 2′-deoxy-2′- f l u o r o p h o sphorothioate oligonucleotides as nuclease-resistant antisense compounds with high affinity and specificity for RNA targets. J. Med. Chem. 36:831-841.
Synthesis of Modified Oligonucleotides and Conjugates
4.1.15 Current Protocols in Nucleic Acid Chemistry
LeDoan, T., Perrouault, L., Praseuth, D., Habhoub, N., Decout, J.L., Thuong, N.T., Lhomme, J., and Helene, C. 1987. Sequence-specific recognition, photocrosslinking and cleavage of the DNA double helix by an oligo-[α]-thymidylate covalently linked to an azidoproflavine derivative. Nucl. Acids Res. 15:7749.
Prasmickaite, L., Hogset, A., Maelandsmo, G., Berg, K., Goodchild, J., Perkins, T., Fodstad, O., and Hovig, E. 1998. Intracellular metabolism of a 2′-O-methyl-stabilized ribozyme after uptake by DOTAP transfection or as free ribozyme. A study by capillary electrophoresis. Nucl. Acids Res. 26:4241-4248.
Lesnik, EA., Guinosso, C.J., Kawasaki, A.M., Sasmor, H., Zounes, M., Cummins, L.L., Ecker, D.J., Cook, P.D., and Freier, S.M. 1993. Oligodeoxynucleotides containing 2′-O-modified adenosine: Synthesis and effects on stability of DNA:RNA duplexes. Biochemistry 32:78327838.
Rossi, J.J. 1998. Therapeutic ribozymes: Principles, applications, and problems. In Applied Antisense Oligonucleotide Technology (C.A. Stein and A.M. Krieg, eds.) pp. 511-525. Wiley-Liss, New York.
Lima, W.F. and Crooke, S.T. 1997. Binding affinity and specificity of Escherichia coli RNase H1: Impact on the kinetics of catalysis of antisense oligonucleotide-RNA hybrids. Biochemistry 36:390-398. Manoharan, M. 1993. Designer antisense oligonucleotides: Conjugation chemistry and functionality placement. In Antisense Research and Applications (S.T. Crooke and B. Lebleu, eds.) pp. 303-349. CRC Press, Boca Raton, Fla. Matteucci, M.D. and von Krosigk, U. 1996. Hybridization properties of oligonucleotides bearing a tricyclic 2′-deoxycytidine analog based on a carbazole ring system. Tetrahedron Lett. 37:5057-5060. Miller, P.S., Bhan, P., Cushman, C.D., Kean, J.M., and Levis, J.T. 1991. Antisense oligonucleotide methylphosphonates and their derivatives. Nucleosides Nucleotides 10:37-46. Monia, B.P., Lesnik, E.A., Gonzalez, C., Lima, W.F., McGee, D., Guinosso, C.J., Kawasaki, A.M., Cook, P.D., and Freier, S.M. 1993. Evaluation of 2′-modified oligonucleotides containing 2′-deoxy gaps as antisense inhibitors of gene expression. J. Biol. Chem. 268:14514-14522. Monia, B.P., Johnston, J.F., Sasmor, H., and Cummins, L.L. 1996. Nuclease resistance and antisense activity of modified oligonucleotides targeted to Ha-ras. J. Biol. Chem. 271:1453314540. Morishita, R., Givvons, G.H., Kaneda, Y., and Dzau, V.J. 1995. Pharmacokinetics of antisense oligodeoxynucleotides (cyclin B1 and cdc2 kinase) in the vessel wall in vivo: Enhanced therapeutic utility for restenosis by HVJ-liposome delivery. Gene 149:13-19. Morvan, F., Porumb, H., Degols, G., Lefebvre, I., Pompon, A., Sproat, S., Rayner, B., Malvy, C., Lebleu, B., and Imbach, J.-L. 1993. Comparative evaluation of seven oligonucleotide analogues as potential antisense agents. J. Med. Chem. 36:280-287. Moser, H.E. and Dervan, P.B. 1987. Sequence-specific cleavage of double helical DNA by triple helix formation. Science 238:645-650. Modified Oligonucleotides for Chemotherapeutic Applications
Nicklin, P. 1998. Pharmacokinetics properties of phosphorothioates in animals. In Handbook of Experimental Pharmacology (S.T. Crooke, ed.) pp. 141-168. Springer-Verlag, Heidelberg, Germany.
Sands, H., Gorey-Feret, L.J., Ho, S.P., Bao, Y., Cocuzza, A.J., Chidester, D., and Hobbs, F.W. 1995. Biodistribution and metabolism of internally 3H-labeled oligonucleotides. II. 3′,5′Blocked oligonucleotides. Therapeutics 47:636646. Sheffery, M. and Gordon, C.L. 1996. Leadership positions in antisense patents (company report). Mehta and Isaly Equity Research, New York. Sproat, B.S. and Lamond, A.I. 1993. 2′-O-Alkyloligoribonucleotides. In Antisense Research and Applications (S.T. Crooke and B. Lebleu, eds.) pp. 351-362. CRC Press, Boca Raton, Fla. Sproat, B.S., Lamond, A.I., Beijer, B., and Neuner, U. 1989. Highly efficient chemical synthesis of 2′-O-methyloligoribonucleotides and tetrabiotinylated derivatives; novel probes that are resistant to degradation by RNA or DNA specific nucleases. Nucl. Acids Res. 17:3373-3386. Stephenson, M.L. and Zamecnik, P.C. 1978. Inhibition of Rous sarcoma viral RNA translation by a specific oligodeoxyribonucleotide. Proc. Natl. Acad. Sci. U.S.A. 75:285-288. Summerton, J. 1979. Intracellular inactivation of specific nucleotide sequences: A general approach to the treatment of viral diseases and virally-mediated cancers. J. Ther. Biol. 78:77-99. Torrence, P.F., Xiao, W., Li, G., Lesnik, K., Khamnei, S., Maran, A., Maitra, R., Dong, B., and Silverman, R.H. 1994. 2′,5′-Oligoadenylate antisense chimeras for targeted ablation of RNA in carbohydrate modifications. In Antisense Research (Y.S. Sanghvi and P.D. Cook, eds.) pp. 118-132. American Chemical Society, Washington, D.C. Ts’o, P.O., Miller, P.S., Aurelian, L., Murakami, A., Agris, C., Blake, K.R., Lin, S.B., Lee, B.L., and Smith, C.C. 1987. An approach to chemotherapy based on base sequence information and nucleic acid chemistry. Matagen (masking tape for gene expression). Ann. N.Y. Acad. Sci. 507:220-241. Tuerk, C. and Gold, L. 1990. Systematic evolution of ligands by exponential enrichment: RNA ligands to bacteriophage T4 DNA polymerase. Science 249:505-510. Wagner, R.W., Matteucci, M.D., Grant, D., Huang, T., and Froehler, B.C. 1996. Potent and selective inhibition of gene expression by an antisense heptanucleotide. Nat. Biotechnol. 14:840-844.
4.1.16 Current Protocols in Nucleic Acid Chemistry
Yu, D., Iyer, R.P., Shaw, D.R., Lisziewicz, J., Li, Y., Jiang, Z., Roskey, A., and Agrawal, S. 1996. Hybrid oligonucleotides: Synthesis, biophysical properties, stability studies, and biological activity. Bioorg. Med. Chem. Lett. 4:1685-1692. Zamecnik, P.C. and Stephenson, M.L. 1978. Inhibition of Rous sarcoma virus replication and cell transformation by a specific oligodeoxynucleotide. Proc. Natl. Acad. Sci. U.S.A. 75:280284.
Zhang, R., Iyer, R.P., Yu, D., Tan, W., Zhang, X., Lu, Z., Zhao, H., and Agrawal, S. 1996. Pharmacokinetics and tissue disposition of a chimeric oligodeoxynucleoside phosphorothioate in rats after intravenous administration. J. Pharmacol. Exper. Ther. 278:971-979. Zon, G. 1993. History of antisense drug discovery. In Antisense Research and Applications (S.T. Crooke and B. Lebleu, eds.) pp. 1-5. CRC Press, Boca Raton, Fla.
Contributed by P. Dan Cook Isis Pharmaceuticals Carlsbad, California
Synthesis of Modified Oligonucleotides and Conjugates
4.1.17 Current Protocols in Nucleic Acid Chemistry
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups Because of the base-pairing properties of oligomeric strands of DNA, which allow specific recognition of a particular base sequence by a complementary one in a single-stranded DNA or RNA target, such strands are used as nucleic acid probes for molecular biology, diagnostics in medicine, and as artificial regulators of gene expression according to the “antisense” strategy (Cohen, 1989; Hélène and Saison-Behmoaras, 1994). Recently, synthetic oligonucleotides have been used to regulate gene expression via the “antigene” approach upon hybridization with double-stranded DNA through Hoogsteen hydrogen-bonding interactions of an oligonucleotide third strand with the purine bases of the double helix (Thuong and Hélène, 1993; Soyfer and Potoman, 1996). To improve the performance of oligonucleotides, various molecules have been attached to them to provide the following properties: 1. Easy detection with high sensitivity; 2. Increased affinity to complementary nucleic acids sequences; 3. Ability to induce irreversible modifications of the target sequences; 4. Capacity to recognize and permeate target cell membranes. The structural, chemical, and physicochemical properties of the ligands used in these applications vary widely, and may range from very simple to very complex when a low-stability molecule, such as a protein, is involved. These different parameters must be taken into account when choosing the positions within each molecule that will be involved in the chemical reactions needed for their covalent linking. Care is essential when determining the best conjugation method for a chosen ligand, in order to preserve its chemical, physicochemical, and biochemical properties, without impairing the oligonucleotide’s ability to bind specifically with the nucleic acid target. As a general rule, it is necessary to insert a linker (L) between the ligand and the oligonucleotide to provide enough conformational freedom for the desired complex to form between the oligonucleotide-ligand conjugate
UNIT 4.2
and the nucleic acid or molecular target. The performance of the oligonucleotide-ligand conjugate depends on the attachment sites of the two entities as well as on the linker’s nature and size. A ligand can be covalently attached to the oligonucleotide chain through the 5′ or 3′ end, the nucleobases, the sugars, or the internucleotide bridges. Using too short a linker may induce steric hindrance or prevent the ligand from adopting the most favorable functional conformation (Asseline et al., 1996); using too large a linker may impair cooperative binding between the oligonucleotide-ligand conjugate and the nucleic acid target. However, the nature and size of the linker are less important to small ligands and, in particular, to reporter groups. This unit reviews the covalent attachment of various ligands to the 5′ end of oligonucleotides and the properties of these molecules. The synthesis of oligonucleotide-ligand conjugates involving the coupling of ligands to other oligonucleotidic sites will be presented elsewhere.
SYNTHESIS The two major techniques used for the synthesis of oligodeoxynucleotides are the “phosphotriester” (Michelson and Todd, 1955; Reese and Shaffhill, 1968; Itakura et al., 1975) and “phosphite” (Letsinger and Lunsford, 1976; Beaucage and Caruthers, 1981; UNIT 3.3) methods. Although the first is more convenient for syntheses in solution, the second, using phosphoramidite intermediates, is the best one for solid-phase synthesis. Both methods involve very reactive intermediates requiring appropriate protection for the functional groups not engaged in the coupling reaction, and deprotection conditions that do not affect the integrity of the oligonucleotide chain. Because of these constraints, the phosphotriester and phosphite methods are essentially used to introduce simple and chemically stable functional groups or ligands to the 5′ end of oligonucleotides. In the case of ligands with poor chemical stability or complex structures, the best conjugation method involves coupling to unblocked oligomers that have 5′ ends functionalized with potent electrophilic or nucleophilic groups.
Contributed by Nguyen T. Thuong and Ulysse Asseline Current Protocols in Nucleic Acid Chemistry (2000) 4.2.1-4.2.33 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Modified Oligonucleotides and Conjugates
4.2.1
B'
HO
O
CN
O
H Z'LO P O O
Z'LO P R'O
N(i-Pr)2
1
2
3
O
O ZLO P O O
Z'LOH
B
N
O
4
6
5 O ZLNH2
N H
NC B'
O
7
DMTrO 9
ZL O O
B
n
O
CN
O H P O O
B'
DMTrO
HN O P O O
B' O
(i-Pr)2N
R'O
O
O P O
O
8
O P O O
O R'O
RO
ZL
B'
O
N
B' O
H2N
R'O
11
12
B
H2N 10
NC
O P O (i-Pr)2N
B' O N Tr H
13
Figure 4.2.1 Functionalization of oligodeoxynucleotides via phosphite and phosphotriester derivatives. Abbreviations used in figures: B, nucleic base; B′, protected base; DMF, dimethylformamide; DMSO, dimethyl sulfoxide; DMTr, 4,4′-dimethoxytrityl; Fmoc, 9-fluorenylmethyl; L, linker; MMTr, monomethoxytrityl; Px,9-phenylxanthen-9-yl(pixyl); R, oligodeoxynucleotide; R′, protected oligonucleotide; TCEP, tris-(2-carboxyethyl)phosphine; Tr, trityl; Z, functional group or ligand; Z′, protected functional group or ligand.
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
A protected oligonucleotide with a free hydroxyl group at the 5′ end immobilized on the support (S.1; Fig. 4.2.1) may easily be obtained by solid-phase synthesis using either nucleoside-3′-phosphoramidites or nucleoside-3′-Hphosphonates. The 5′-terminal hydroxyl can then be reacted with phosphoramidites (S.2), H-phosphonates (S.3), and 1,1′-carbonyldiimidazole.
Phosphoramidites and H-phosphonates often obtained from a molecule having a hydroxyl functionality (S.4), allow the direct introduction of a functional group or ligand (Z-L) via a phosphodiester bond (S.5) after oxidation and oligonucleotide deprotection. This synthetic route implies the preparation of “phophitylation reagents” and, if necessary, protection of the functional groups of the ligands not involved in the
4.2.2 Current Protocols in Nucleic Acid Chemistry
coupling reaction. When using 1,1′-carbonyldiimidazole, it is possible to introduce functional groups or ligands bearing a primary amino functionality to the 5′-terminus of oligonucleotides, without the need to prepare intermediate reagents or protecting the functional groups of the ligands not involved in the formation of the oligonucleotide-ligand conjugates, provided these are less reactive than the ligands’ primary amino group. The first step of this method is the formation of the carbonylimidazolide derivative (S.6) which then reacts with the primary amino functionality of ligand (S.7) to form the conjugate with a carbamate linkage (S.8) stable to the basic conditions used for oligonuleotide deprotection. An alternative to this conjugation method is to use modified supports derivatized with the ligand to assemble the oligonucleotide chain in the (5′→3′)-orientation via 5′-phosphoramidites (S.9; Horne and Dervan, 1990). This strategy is particularly well adapted for the modification of the 5′ end of oligodeoxyribonucleotide (N3′→P5′)-phosphoramidates (S.10), whose synthesis is achieved in the (5′→3′)-orientation by coupling the 3′-aminonucleoside (S.11) with 5′-H-phosphonate (S.12; Chen et al., 1995) or with the 5′-phosphoramidites (S.13; McCurdy et al., 1997).
Addition of Functional Groups to the 5′ End of Oligonucleotides Owing to the low reactivity of the 5′-hydroxyl functionality of oligonucleotides, its condensation with the functional group of ligands is not easily achieved without side reactions at the nucleic base level. Thus, the preparation of oligonucleotide-ligand conjugates from unblocked oligomers requires a reactive electrophilic or nucleophilic group at their 5′ end to react specifically, either directly or following activation, with a ligand bearing the appropriate functional group. Among the functional groups able to meet these requirements are the phosphoryl, thiophosphoryl, sufhydryl, amino, carboxylic, and cis-dihydroxyl groups. 5′-Terminal phosphate and phosphorothioate groups 5′-Phosphate (S.14; Fig. 4.2.2) or phosphorothioate (S.15) can be introduced either chemically during the oligonucleotide synthesis, or enzymatically on the 5′-hydroxyl group of an oligonucleotide chain through the use of T4 polynucleotide kinase, which transfers the γ-phosphoryl or thiophosphoryl of ATP (Eckstein, 1983) to the 5′-terminal hydroxy function
of oligodeoxyribonucleotides or oligoribonucleotides. The first chemical synthesis of dinucleotide 5′-phosphate (S.16) was achieved using the phosphotriester method in solution, by condensation of thymidine-5′-O-dibenzylphosphate (S.17) with 3′-acetyldeoxynucleotide (S.18) in the presence of dicyclohexylcarbodiimide followed by deblocking of the 3′-hydroxyl and the phosphate groups by alkaline hydrolysis and hydrogenolysis, respectively (Michelson and Todd, 1955). The 5′-phosphate-containing oligonucleotides are easily prepared on solid phase by condensation of the phosphoramidite S.19 (Uhlmann and Engels, 1986; Thuong and Chassignol, 1987), S.20 (Uhlmann and Engels, 1986; Schwarz and Pfleiderer, 1987), S.21 (Horn and Urdea, 1986), or S.22 (Guzaev et al., 1985) with the 5′-terminal hydroxyl of the oligonucleotide bound to the support, followed by oxidation and deprotection steps. The removal of the first ester group [2-cyanoethyl or 2-(4-nitrophenyl)ethyl] from the terminal phosphate triesters via β-elimination is achieved by treatment with concentrated aqueous ammonia, an approach similar to that used to deblock internucleotidic 2-cyanoethyl phosphate triesters. Deprotection of the second ester function of the terminal phosphate diesters, however, requires more drastic conditions, because of the increased electron density on the phosphodiester function making it a poorer leaving group. Although these phosphorylating methods are well adapted to oligodeoxyribonucleotides, they are difficult to use with oligoribonucleotides without inducing chain cleavage. Because of the inherent hydrophobicity of reagent S.22, it facilitated purification of the appended oligomers by reversed-phase chromatography. This protective group can be removed by detritylation followed by ammonium hydroxide treatment. It is worth noting that among these four phosphorylating reagents only S.19, which can be purified by “molecular distillation,” can be stored at −20°C for several months without decomposition (Thuong and Chassignol, 1987). The preparation of oligonucleotides containing a phosphorothioate group at the 5′ end is similarly achieved, by replacing the iodine oxidation step with a sulfurization one using a solution of elemental sulfur solution in CS2/pyridine (Horn and Urdea, 1986; Schwarz and Pfleiderer, 1987; Thuong and Chassignol, 1987). Since the original pioneering work, the sulfuriza-
Synthesis of Modified Oligonucleotides and Conjugates
4.2.3 Current Protocols in Nucleic Acid Chemistry
OH O P O O
OH S P O O
B O
OH O P O O
B O
RO
RO
14
15
Thy
O
O O P O O
O
B
HO OCH2Ph O P OCH2Ph Thy O O
16
OH O P O O
B O O
HO 17
O 18
O
NO2
CN
O (i-Pr)2N P
(i-Pr)2N P O
O
CN
NO2
19 20
DMTrO
O S O
O
CN
O P N(i-Pr)2 21
EtO2C CO2Et O DMTrO O P N(i-Pr)2
CN
22
Figure 4.2.2 Incorporation of 5′-phosphate and 5′-phosphorothioate groups into oligodeoxynucleotides. Abbreviations: Thy, thymin-l-yl. See Figure 4.2.1 for additional definitions of functional group abbreviations.
tion step has been carried out using easy-tohandle and more reactive reagents such as 3H1,2-benzodithiol-3-one 1,1-dioxide (Iyer et al., 1990), tetraethylthiuram disulfide (Vu and Hirschbein, 1991), or bis(O,O-diisopropoxyphosphinothioyl) disulfide (Stec et al., 1993).
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
5′-Thiol-containing oligonucleotides The introduction of a sulfhydryl group to the 5′ termini of oligonucleotides has been achieved using modified nucleoside derivatives S.23 (Fig. 4.2.3; Sproat et al., 1987a) or by using the nonnucleosidic derivatives S.24 (Connolly, 1985), S.25 (Kuijpers and van Boeckel, 1993), or S.26 (Gao et al., 1995).
The S-trityl protecting group can ultimately be removed by treatment with excess AgNO3, which must then be removed by precipitation with dithiothreitol (DTT). A simpler deblocking step can be accomplished by the use of the thioester derivative S.25 and the disulfide-containing compound S.26 upon incubation with aqueous ammonia and DTT, respectively. It is worth noting that the sulfhydryl group is easily oxidized by air at neutral or alkaline pH, so that oligomers obtained after purification are usually in the dimeric disulfide form. The free sulfhydryl must be liberated by reduction of the disulfide bridge just before conjugation. This reaction can be accomplished by treatment with DTT (Cleland, 1964) or tris-(2-carboxyethyl)phosphine (TCEP; Burns et al., 1991).
4.2.4 Current Protocols in Nucleic Acid Chemistry
B'
TrS
O
OMe O P N
TrS
(i-Pr)2N
O P
CN
O
O 24
23
S
O
O
O
S S
CN
O P N(i-Pr)2
O
CN
O
DMTrO
O P N(i-Pr)2
25 26 O S S N
N
N
B
S S L O P O O
27
O RO
28
Figure 4.2.3 Incorporation of 5′-thiol groups into oligodeoxynucleotides. See Figure 4.2.1 for definitions of functional group abbreviations.
An interesting approach is to deblock the protected sulfhydryl group in the presence of 2,2′-dithiodipyridine (S.27; Py-SS-Py), which reacts with a free sulfhydryl to form the pyridinyldisulfide derivative S.28 (Kuijpers and van Boeckel, 1993). This compound is air-stable and possesses an activated disulfide function that can either be used directly to achieve conjugation with thiol-containing compounds or easily reduced to the thiol form. During deprotection of oligonucleotides synthesized f r o m β-cyanoethylphosphoramidites, acrylonitrile released by β-elimination of cyanoethyl phosphotriesters reacts easily with a free sulfhydryl group to give the β-cyanoethylthioether derivative, greatly decreasing the synthesis yield of the 5′-thiol-containing oligonucleotides. This drawback can be circumvented by deblocking the oligonucleotide using a mixture of concentrated aqueous ammonia, 2,2′-dithiopyridine, and phenol (Asseline and colleagues, unpub. observ.). Using this method large amounts of the 5′-pyridinyldisulfide-containing oligonucleotides S.28 have been obtained both as phosphodiesters and phosphorothioates. 5′-Amino-containing oligonucleotides Addition of a primary amino group to the 5′ end of oligonucleotides has been performed via
5′-azido-2′,5′-dideoxyribonucleoside phosphorochloridites such as S.29 (Fig. 4.2.4; Mungall et al., 1975), 5′-masked amino-2′,5′-dideoxyribonucleoside phosphoramidites S.30 (Smith et al., 1985) and S.31 (Sproat et al., 1987b), or the non-nucleosidic phosphoramidite derivatives S.32 (Coull et al., 1986), S.33 (Connolly, 1987), and S.34 (Connel et al., 1987). The chlorophosphite S.29 was used for the synthesis of 5′-amino-oligothymidylate (P3′→N5′)-phosphoramidate (S.35) in solution. At the end of the oligonucleotide-chain assembly, the azido group is easily reduced to an amino function by triphenylphosphine in the presence of ammonium hydroxide. Alternatively, the phosphoramidites S.30, S.31, S.32, S.33, and S.34 are condensed with the 5′-terminal hydroxyl of oligonucleotides bound to the support using standard conditions. In the case of the N-trifluoroacetyl-containing compounds S.31, S.32, S.34, and the N-9-fluorenylmethoxycarbonyl (Fmoc)-containing compound S.30, release of the amino group is achieved during the deprotection of the oligonucleotide chain by concentrated aqueous ammonia treatment. Deprotection of the tritylated derivative S.33 requires acid treatment, which can be performed either while the oligonucleotide is still bound to the support or
Synthesis of Modified Oligonucleotides and Conjugates
4.2.5 Current Protocols in Nucleic Acid Chemistry
O
H Thy
N3
Cl
Thy
Fmoc N
O O
O
P
P
(i-Pr)2N
OPh
F3C
O
Thy
N H
O O
CN
O
P
(i-Pr)2N
30
29
H N
F3C
31
CN
O
MMtr
O P N(i-Pr)2
O
CN
O
32
OMe N H
O P N(i-Pr)2 33
H H N
O
O MeO P N O CF3 34
O
Thy H2N
O O P O H
H
n
N O
Thy
B'
O
nN
O R'O
36
HO 35
Figure 4.2.4 Incorporation of 5′-amino groups into oligodeoxynucleotides. See Figure 4.2.1 for definitions of functional group abbreviations.
after deblocking and purification of the tritylated oligonucleotide by reversed-phase chromatography. Another method consists of condensing the 5′-hydroxyl group of protected oligomers bound to the support with 1,1′-carbonyldiimidazole (similar to S.6) and then reacting the resulting imidazolide derivative with an alkylenediamine to afford the corresponding aminoalkylated oligonucleotides S.36 (Wachter et al., 1986).
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
5′-Carboxyl-containing oligonucleotides 5′-Carboxyl-containing oligonucleotides S.37 (Fig. 4.2.5) or S.38 were prepared, respectively, by condensation of the 5′-hydroxyl group of protected oligomers bound to a solid support with phosphoramidite (S.39; Kremsky et al., 1987), or by treatment with 1,1′-carbonyldiimidazole followed by reaction with an amino acid (Gotthikh et al., 1990). 5′-Diol-containing oligonucleotides By analogy with the specific activation of the 3′-terminal cis diol of RNA by periodate
oxidation, uridine was introduced at the 5′ end of oligonucleotides via a (5′-5′)-linkage (S.40; Fig. 4.2.6) generated from either the phosphodiester S.41 (Agrawal et al., 1986) or the phosphoramidites S.42 (Agrawal et al., 1986) and S.43 (Kuijpers et al., 1993). Conversion of one functional group to another Numerous processes allowing the conversion of one functional group to another are available. The 5′-phosphate linkers can be coupled with diamines and with aminothiols in the presence of condensing agents to give the 5′amino linker (Chu et al., 1983) and the 5′-sulfhydryl linkers (Chu and Orgel, 1988), respectively. The 5′-amino group reacts directly with iminothiolane (S.44; Fig. 4.2.7) to generate a sulfhydryl group via amidine bond formation (Jue et al., 1978). Heterobifunctional reagents such as succinimidyl 3-(2-pyridyldithio)propionate (SPDP; S.45; Cumber et al., 1985), succinimidyl trans-4-(N-maleimidylmethyl)cyclohexane-1-carboxylate (SMCC; S.46; Ghetie et al., 1990), succinimidyl (acetylthio)acetate (SATA; S.47; Julian et al., 1983;
4.2.6 Current Protocols in Nucleic Acid Chemistry
O
CO2H O O P O
B
HO2C
O
O
nN
B
O
H
RO
O RO
38
37
O
OMe O P
O
N(i-Pr)2 39
Figure 4.2.5 Incorporation of 5′-carboxyl groups into oligodeoxynucleotides. See Figure 4.2.1 for definitions of functional group abbreviations.
Ghetie et al., 1990), 4-[(succinimidyloxy)carboxyl]-α-methyl-α-(2-pyridyldithio)toluene (SMPT; S.48; Thorpe et al., 1987; Ghetie et al., 1990), succinimidyl 3-maleimidopropionate (S.49) succinimidyl 4-{[(iodoacetyl)amino]methyl}-cyclohexane-1-carboxylate (SIAC; S.50; Haugland, 1989), succinimidyl iodoacetate (S.51), and 4-nitrophenyl iodoacetate (S.52) can be used to introduce various activated functional groups. Activated esters S.45-S.52 react with primary amino groups to afford amide bonds. Alternatively, the pyridine disulfide derivatives S.45 and S.48 and the maleimido derivatives S.46 and S.49 react with the sulfhydryl groups to give disulfide and thioether derivatives, respectively. One of the two functional groups of these heterobifunctional reagents is used to couple the reagent to the oligonucleotide without affecting the other. The intact functional group is then reacted with selected ligands.
Phosphorothioate, amino, thiol, phosphate, carboxylic, and cis-diol groups possess very different reactivities. The first three are nucleophiles that can be used directly for the conjugation reaction. Conversely, the last three groups, being weakly nucleophilic, must be activated to enable their reaction with, for example, amino functions. In a general way, conjugation reactions are often performed in aqueous medium or in a mixture of water and organic solvents in which oligonucleotides are soluble. However, strong solvation of nucleophilic groups such as oxygenated anions and amines by water often led to slow and poor yield conjugation reactions; in such solvents. Generally, sulfhydryl and thiophosphoryl groups are only slightly deactivated in water, but the formation of amide and phosphoramidate is more strongly disfavored.
O
Ura
O B
O P O
Ura
O P O
O
O
O HO
Covalent Attachment of Ligands to Oligonucleotides Functionalized at the 5′ End
O
O Cl
OH
RO
PxO
40
41
H3CO
Ura
P O
NC
O
O
(i-Pr)2N
OPx
Ura
P O
O
(i-Pr)2N PxO
OPx AcO
42
OAc
43
Figure 4.2.6 Some structures involved in the incorporation of 5′-diol groups into oligodeoxynucleotides. See Figure 4.2.1 for definitions of functional group abbreviations.
Synthesis of Modified Oligonucleotides and Conjugates
4.2.7 Current Protocols in Nucleic Acid Chemistry
O NH2Cl
N
S S
O N
S O
44
O
SPDP 45
O
O
O
O
O
N O
O N
S
N O
O
O SMCC
SATA
46
47
O
O
O
O N O N S S
O O
O N
N O
O
SMPT
49
48
I
O
O NH
O
O
O
O
N O
O N I O
O
SIAC 50
51
O I
O
NO2 52
Figure 4.2.7 Heterobifunctional reagents for the conversion of one oligodeoxynucleotide 5′-functional group to another.
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
These drawbacks may be avoided by performing the coupling reactions in an organic medium. Solubilization of oligonucleotides in organic solvents can be achieved by exchange of phosphate counterions (NH4+, Et3NH+, Na+, K+) with lipophilic ammonium cations such as cetyltrimethylammonium (Zarytova et al., 1987), or by complexing the counterions (NH4+, Na+, or K+) with crown ethers (Odell, 1985; Thuong and Asseline, 1991) or Kryptofix (Dietrich et al., 1969). The latter method, easy to develop, generally gives good results. Possi-
ble organic solvents are methanol and dipolar aprotic solvents such as N,N-dimethylformamide (DMF) or dimethylsulfoxide (DMSO). It is worth noting that DMSO can oxidize thiols into disulfides (Tam et al., 1991) and hence is unsuitable for coupling reactions involving this functional group. Via a 5′-terminal phosphate Modification via the 5′-terminal phosphate is based on the specific activation of the phosphomonoester (which is much more nucleo-
4.2.8 Current Protocols in Nucleic Acid Chemistry
philic than internucleotidic phosphodiesters) to give an electrophilic intermediate that can then react with nucleophilic derivatives such as alcohols or amines to give esters and phosphoramidates, respectively. This method, initially described by Gilham and Khorana in 1958, consisted of the condensation of 5′-Otritylthymidine S.53 (Fig. 4.2.8) with 3′-Oacetylthymidine-5′-phosphate S.54 in the presence of dicyclohexylcarbodiimide (CDI) S.55a to give the dinucleoside monophosphate S.56. This method was then used to immobilize oligonucleotides on paper (Gilham, 1962). Due to the poor nucleophilicity of alcohols, this esterification reaction is not selective and leads to side reactions at the nucleic base level. Primary and secondary amines, which are much more nucleophilic than alcohols, selectively give phosphoramidate derivatives. More recently this method has been optimized by the use of a water-soluble carbodiimide S.55b in the presence of a catalyst such as imidazole, N-methylimidazole, or 4-dimethylaminopyridine. The use of these particular amines leads to the reactive intermediates S.57, S.58, and S.59, which will react in situ with primary and secondary aliphatic amines. (In these intermediates the corresponding phos-
OH O P O O
Thy
TrO
O HO
phate anion is not nucleophilic because one of its substituent group is electron withdrawing.) These two strategies can be used to couple ligands that are not sensitive to condensing agents to short nucleic acid segments. The conversion of the decanucleotide-5′-phosphate group S.60 (Fig. 4.2.9) to the aminated 5′-phosphoramidate S.61 was performed by direct coupling of ethylenediamine and S.60 in the presence of CDI, S.55b, and N-methylimidazole, or by reacting the phosphoimidazolide S.57 with the diamine (Chu et al., 1983) in aqueous medium. Using this method, cystamine S.62 (Chu and Orgel, 1988) and the orthophenanthroline (OP) derivative S.63 (Fig. 4.2.9; Chen and Sigman, 1986) have been linked to oligodeoxyribonucleotides via a phosphoramidate linkage. The same method was also applied to the conjugation of psoralen (Pso) to oligodeoxyribonucleotides S.64 and to oligodeoxyribonucleotide methylphosphonates S.65 (Lee et al., 1988; Bhan and Miller, 1990). In the case of longer nucleic acid chains or ligands involving functional groups able to react with CDI, such as proteins, it is necessary to prepare and isolate the phosphoimidazolide derivative S.57 prior to performing the coupling step (Chu et al., 1983; Chu and Orgel, 1988).
Thy O
55a, R1 = R2 =
AcO
55b, R1 = Et
54
53
R1 N C N R2
R2 =
TrO
O
O O P O O
HCl Me2N
Thy
O
N
Thy
O N P OR O
H3C
N
O N P OR O 58
57 AcO 56 O N P OR O
(Me)2N
59
Figure 4.2.8 Modification of oligonucleotides via 5′-terminal phosphate groups. See Figure 4.2.1 for definitions of functional group abbreviations.
Synthesis of Modified Oligonucleotides and Conjugates
4.2.9 Current Protocols in Nucleic Acid Chemistry
H2 N
OH O
S
O P O
O P O Thy
O
NH2
S
NH O
O
NH
Thy
O P O O
O
B
O
O
10
H
10
H
RO 62
61
60
N
O N H
N
H N
O NH H3 C
O P O O
B
O
O
B
CH3 RO
O 64
H N
O O H3 C
NH H3C P O O
O
NH O P O O
O
RO
63
O
O
B
CH3 O
O
H
n
65
Figure 4.2.9 Conjugation of ligands to oligonucleotides via 5′-terminal phosphate groups. See Figure 4.2.1 for definitions of functional group abbreviations.
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
Carbodiimide, which has a tendency to react with the nucleic bases and particularly with thymine (Chu et al., 1983), can advantageously be replaced by the triphenylphosphine/2,2′dipyridyldisulfide couple (Zarytova et al., 1987). Thus, when the cetyltrimethylammonium salt of oligonucleotides, N-methylimidazole or 4-dimethylaminopyridine (Zarytova et al., 1987) and the latter condensing reagent are mixed in DMF or DMSO, the zwitterionic derivatives S.58 and S.59, respectively, are easily generated. These react with amino-containing compounds to form phosphoramidates. This method has been used to conjugate the sulfhydryl group S.66 (Fig. 4.2.10; Boutorin et al., 1994), the coproporphyrin I chelate S.67 (Fedorova et al., 1990), the bleomycin A5 S.68 (Zarytova et al., 1993), and intercalating agents such as the phenazinium S.69 (Lokhov et al., 1992), 2methoxy-6-chloro-9-aminoacridine S.70 (Balbi et al., 1994), the ethidium S.71, the azidoethidium S.72 (Koshkin et al., 1994), the daunomycin S.73 (Dikalov et al., 1991), and the benzopyridoin-
dolium salt S.74 (Silver et al., 1997) to the 5′-terminus of oligonucleotides. Finally, it must be noted that ester bonds between 5′-phosphate-containing oligonucleotides and the free 3′-hydroxyl group of another oligonucleotide can easily be formed when using ligase and template oligonucleotide sequences.
Via a 5′-terminal phosphorothioate The nucleophilicity of the phosphorothioate group allows easy coupling with electrophilic reagents. Oligonucleotides bearing thiophosphate groups at the 5′ ends (S.15) react specifically with halogenoalkyl derivatives to afford the corresponding 5′-phosphothioloester. For example, various intercalating agents such as acridine (S.75; Fig. 4.2.11), proflavin (S.76; Praseuth et al., 1988a), daunomycin (S.77; Garbesi et al., 1997), thiazole orange (S.78; N.N.T. and U.A., unpub. observ.), azidophenacyl (S.79; Praseuth et al., 1988b), pyrene (S.80;
4.2.10 Current Protocols in Nucleic Acid Chemistry
H N O P O O
ZL
O
B
RO 66 to 74
ZL
=
HS 66
HO2C CO2H NH ZL
=
HO2C
N
N
HN O N H
67
Figure 4.2.10 Attachment of ligands to oligonucleotides via 5′-terminal phosphate groups. See Figure 4.2.1 for definitions of functional group abbreviations.
Ebata et al., 1995), and the cyclopropapyrroloindole subunit of the antitumor antibiotic CC-1065 (S.81; Lukhtanov et al., 1997a,b) have been linked to oligonucleotides in this manner. The kinetics and yields of these reactions depend on the nature of the ligands and the halogenoalkyl groups. In general, iodoacetamido derivatives [-NH-(CO)-CH2-I] are more reactive than the corresponding iodoalkyl derivatives. The weakly reactive chloroalkyl derivatives can be easily converted to the corresponding iodoalkyl derivatives via a halogen exchange reaction with sodium iodide in acetone or acetonitrile. It is noteworthy that the coupling reaction of orthophenanthroline to oligonucleotides sometimes gives very low yields. In one such case, the formation of a 5′-monophosphate (S.14) and a dimer with a disulfide bridge (S.82; Fig. 4.2.12) was observed (Thuong and Asseline, 1991). These side reactions are probably caused by superoxide anion O2− and H2O2 generated by the presence of OP, thiophosphate, oxygen, and traces
of divalent cations contained in the conjugation buffer. This is analogous to the mechanism proposed for the nucleolytic activity of the complex OP-Cu in the presence of thiol (Kuwabara et al., 1986; Hélène and Thuong, 1988). To prevent these side reactions it is necessary to eliminate traces of divalent cations contained in buffer solutions by chelation with, for example, Chelex 100 (Thuong and Asseline, 1991). Formation of the 5′-phosphothioloester (S.83) can also be obtained by reaction of the 5′-phosphorothioate oligomer with the maleimido derivative of OP (Shimizu et al., 1994). Another coupling reaction via the phosphorothioate group concerns the formation of the disulfides S.84a and S.84b, which is accomplished with ligands bearing a pyridinyldisulfide group (S.85; Chassignol and Thuong, 1998). One particularity of these conjugates is that oligonucleotide 5′-phosphorothioates can be regenerated by reduction of the disulfide linkages and, if necessary, transformed to the corresponding 5′-phosphate by oxidation.
Synthesis of Modified Oligonucleotides and Conjugates
4.2.11 Current Protocols in Nucleic Acid Chemistry
H2N
O
O
NH2
NH H2N ZL
N
N
N
=
O OH O
O
HO
H N
N H
O
O N
O HO
NH
HO
S
N
OH O
OH
O
HN
O HO
N H
H2N
NH
NH
Cu
S
OH O
O
NH2 68
OCH3 OH ZL
H N
N
=
ZL
= N
NH
69 Cl 70
ZL
=
H2N
NH
ZL
= N3
NH
N
N
72
71
O
OH
OH ZL
= H3CO
OH H
O
H3CO
O CH3
O
O
H3C
73
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
=
N N H CH3
OH
Figure 4.2.10
HN ZL
(L) = many different linkers 74
Continued
Via 5′-thiol oligonucleotides The sulfhydryl group, which is most widely used in the modification of proteins, is also often employed for the preparation of oligonucleotide conjugates. However, the tendency to form symmetrical dimers through disulfide bridge formation requires DTT or TCEP reduc-
tion. To improve coupling efficiency the reaction should be performed in oxygen-free solution. Conjugation via thiol groups can lead to the formation of either a chemically stable thioether linkage or a disulfide bridge that can be reduced to give the starting thiols.
4.2.12 Current Protocols in Nucleic Acid Chemistry
O ZL S P O
O
O
B
RO 75 to 81 OCH3
=
ZL
(CH2)n NH
N
(CH2)n R
N
NH
Cl 75 O
76
OH
O OH
H3CO
O
(CH2)n
CH3
N
O
OH H
S
O
H3C OH
HN
H3C
N
O 78 77
O HN
O N3 79
CH3 80
H3C N
O N
O O
N H
81
Figure 4.2.11 Conjugation of functonal groups to oligonucleotides via 5′-terminal phosphorothioate groups. See Figure 4.2.1 for definitions of functional group abbreviations.
The alkylation of thiol groups by iodoacetyl derivatives has been used to prepare the fluorescent conjugates derived from fluorescein (S.86a; Fig. 4.2.13; Ansorge et al., 1987) and N-(5-sulfo-1-naphtyl)ethylenediamine (S.86b; Connolly, 1985). This method has also been used to couple fullerene to the 5′-terminus of oligonucleotides (S.87; Boutorin et al., 1994). Alternatively, the pyrenyl S.88a (Kumar et al., 1991) and antibody-oligonucleotide conjugates S.88b (Kuijpers et al., 1993), have been formed by adding oligonucleotide 5′ thiols to the corresponding maleimido-ligands.
5′-Thiol-containing oligonucleotides also react easily with tresyl-activated or epoxy-activated Sepharose to form thioether linkages (Blanks and McLaughlin, 1988). Air oxidation of two sulfhydryl-containing compounds gives a mixture of homo- and heterodimers. The synthesis of conjugates via formation of cleavable disulfide bridges implies the reaction of one of the thiol groups with 2,2′-dithiodipyridine to form the R-S-SPyr intermediate, which exchanges with the other thiol group to complete conjugation. The reaction of peptide, peroxidase, and human IgG with iminothiolane afforded with thiol deriva-
Synthesis of Modified Oligonucleotides and Conjugates
4.2.13 Current Protocols in Nucleic Acid Chemistry
O O O P S S P O O O
B O OR
O
B
RO 82
N
O
N
O
H N
N H
O S P O O
N O O
O
B
RO
83 O ZL S S P O O
O
B
RO 84
O O ZL
= O O 84a
CH3 ZL
H N
= H2N
O
O
O
O N H
84b
N
S S L Z 85
Figure 4.2.12 Incorporation of functional groups into oligonucleotides via 5′-terminal phosphorothioate groups. See Figure 4.2.1 for definitions of functional group abbreviations.
tives S.89 (Fig. 4.2.14) condensation of these derivatives with the activated oligomers S.90 gave the conjugates S.91 (Chu and Orgel, 1988). Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
Via the 5′-amino group The alkylamino groups easily react at pH ≥ 8 with electrophilic reagents to form stable linkages. Thus, fluorescein isothiocyanate (S.92a; Fig. 4.2.15) and tetramethylrhodamine isothiocy-
anate (S.92b) have been coupled to 5′-aminooligonucleotides in aqueous solution to produce the thiourea derivatives S.93a and S.93b (Smith et al., 1985). The same reaction was used to link orthophenanthroline isothiocyanate S.92c to the 5′ end of oligonucleotides with (N3′→P5′)-linkages to afford S.93c (N.T.T. and U.A., unpub. observ.). Certain acyl chlorides or anhydrides such as dansyl chloride or ethylenediaminetraacetic di-
4.2.14 Current Protocols in Nucleic Acid Chemistry
O
HO
O
O
CO2
ZL =
ZL S (CH2)n O O P O
H
O
B
O
N
86a
HN
HO3S
RO ZL =
NH
86b
H O
O O
O
S
H N O P O O
B
O
O O P O O
O
B
87 RO O
S O
O
O
ZL N O
NH
N H
ZL N
OH O O P O O
S (CH2)2
O O
ZL = pyrenyl
N
B U
O O
RO
O P O O 88a
O
B
ZL = antibody RO 88b
Figure 4.2.13 Conjugation of ligands to oligonucleotides via 5′-thiol groups. See Figure 4.2.1 for definitions of functional group abbreviations.
anhydride (EDTA), and activated esters such as biotin-4-nitrophenylester, N-hydroxysuccinimido-biotin, and the N-hydroxysuccinimide ester of bathophenanthroline have been used to prepare oligonucleotide conjugates S.94 (Fig. 4.2.16), S.95a-c, S.96, and S.97 through the formation of either sulfonamide S.94 (Connolly, 1987) or amide bonds S.95a (Boutorin et
al., 1984; Chu et al., 1985), S.95b (Agrawal et al., 1986), S.95c (Schubert et al., 1995), S.96 (Wachter et al., 1986), and S.97 (Bannwarth et al., 1988). The 5′-arylazido-containing oligonucleotide S.98 (Fig. 4.2.17) has been obtained by arylation of the 5′-terminal amino group of oligonucleotides with 2,4-dinitro-5-az-
Synthesis of Modified Oligonucleotides and Conjugates
4.2.15 Current Protocols in Nucleic Acid Chemistry
Z
H N
SH NH 89
S S
H N
Z
N NH
O O P O O
O
O
S S
O
B
O P O O
R1
B
O
n
H
R1
O
Z = peptide or peroxidase or human IgG R1 = H or OH 91
n
H 90
Figure 4.2.14 Incorporation of conjugate groups into oligonucleotides via 5′-thiol groups. See Figure 4.2.1 for definitions of functional group abbreviations.
also enzymatically coupled to a 2-methy-9-hydroxyellipticinium salt in the presence of horseradish peroxidase to give the oxazolopyridocarbazole derivative S.100 (Mouscadet et al., 1994).
idofluorobenzene (Grimautdinova et al., 1984). In the same way the phenazinium derivatives S.99 were obtained by nucleophilic attack of 5′-amino alkylated oligonucleotides of the C2halogenated heterocycle (Lokhov et al., 1992). 5′-Amino-containing oligonucleotides were
Z N C S HO
O
O
(CH3)2N
N(CH3)2
O
N Z=
Z=
Z=
CO2
CO2
N
92c
92b
92a
Z
H N
H N
R
S
O
O (CH2)nO P O O
Thy
R1 =
HN
RO 93a, 93b
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
B
O
R1 =
O P O O
O
B
HN H
n
93c
Figure 4.2.15 Attachment of reporter and conjugate groups to oligonucleotides via 5′-amino groups. See Figure 4.2.1 for definitions of functional group abbreviations.
4.2.16 Current Protocols in Nucleic Acid Chemistry
Z
n O O P O
N H
O
B
O RO
94 to 95c
HO2C (Me)2N O S
Z=
HO2C
Z=
O N
N
HO2C
O
95a
94 O HN
N
NH
Z=
Z=
N
O
S
N
S O
95c
95b
O HN
NH O S
H N
H N 8
O O
O
B
O O P O O 96
O
B
RO O NH O
Thy
O O P O N
N
O
O
B
97 RO
Figure 4.2.16 Conjugation of functional groups to oligonucleotides via 5′-amino groups. See Figure 4.2.1 for definitions of functional group abbreviations.
Via a 5′-carboxyl group Like phosphates, carboxyl groups can be selectively activated and coupled to hydrazine or amine derivatives to give, for example, the biotin conjugate S.101 (Fig. 4.2.18; Kremsky et al., 1987) and the daunomycin conjugate S.102 (Gotthikh et al., 1990).
Via a 5′-terminal cis diol The 2′- and 3′-hydroxyl functionalities of the ribose-containing oligomer S.40, upon periodate oxidation, can easily be coupled with the hydrazino or primary amino groups of the ligands to give the unstable 3,5-dihydroxymorpholine derivative S.103 (Fig. 4.2.19). Sta-
Synthesis of Modified Oligonucleotides and Conjugates
4.2.17 Current Protocols in Nucleic Acid Chemistry
NO2 NO2
OH
NH
N
NH O P O
N3
O
O
NH
NH O P O O
B
O
B
RO
RO 99
98
H N H3C
N N
O
O O P O O
O
B
RO 100
Figure 4.2.17 Introduction of functional groups to oligonucleotides via 5′-amino groups. See Figure 4.2.1 for definitions of functional group abbreviations.
ble morpholine derivatives S.104 were obtained by reduction of S.103 with NaBH4 or NaBH3CN. Using this method, biotin was coupled via its hydrazino derivative to afford
S.104a (Agrawal et al., 1986). The 125I-labeled tyramine derivative S.104b (Kuijpers and van Boeckel, 1993) was also prepared in a similar manner. Furthermore, periodate oxida-
O HN
O
NH
OH
O CH3 OH
S H3CO
O
O
HO H O
H3C
HN NH
OH
O
O O
HN
nN
O
H
O
O
B
O O P O O
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
O
O O P O O
B
O
B
RO 101
102
RO
Figure 4.2.18 Conjugation of functional groups to oligonucleotides via 5′-carboxyl groups. See Figure 4.2.1 for definitions of functional group abbreviations.
4.2.18 Current Protocols in Nucleic Acid Chemistry
Ura HO Z
Ura
O
O
N
Z
N
OH O O P O O
O O P O O
B
O RO
O
B
RO 104a-b
103a-b O HN
NH
a, Z =
H N S O
(CH2)n
HO b, Z =
125
I
O
O HO OH
HN
N H
NH O
O O P O O
S
O
H N
H N
N H
O
B
RO 105
106
O O P O O
O
B
RO
Figure 4.2.19 Attachment of functional groups to oligonucleotides via 5′-terminal cis diols. See Figure 4.2.1 for definitions of functional group abbreviations.
tion of the 5′-vicinal diol–containing oligonucleotide S.105 (Fig. 4.2.16) followed by coupling with the hydrazinobiotin produced the conjugate S.106 (Kremsky et al., 1987).
Direct Addition of Ligands to the 5′ End of Oligonucleotides The synthesis of oligonucleotide conjugates derived from acridine, psoralen, and porphyrin was first achieved by the phosphotriester method in solution. Oligonucleotide-intercalator conjugates S.107 (Fig. 4.2.20; Thuong et al., 1984) and S.108 (Asseline and Thuong, 1988) were obtained by condensing oligonucleotides S.109 with the hydroxylated ligands
S.110 and S.111, respectively. Similarly, the porphyrin-oligonucleotide conjugate S.112 was synthesized from the protected oligomer S.113 and the hydroxylated porphyrin S.114 (Le Doan et al., 1986, 1987a) in the presence of a condensing reagent. Deprotection of the fully-protected oligonucleotides was achieved by successive treatment with the DBU salt of benzohydroxamic acid and concentrated ammonia or sodium hydroxide. Sodium hydroxide was used instead of aqueous ammonia for the N-deprotection of oligonucleotides covalently linked to acridine to avoid cleavage of the 9-aminoacridine linkage (Asseline et al., 1984; Thuong and Asseline, 1991).
Synthesis of Modified Oligonucleotides and Conjugates
4.2.19 Current Protocols in Nucleic Acid Chemistry
Oligonucleotides bearing a ligand at the 5′ end are easily prepared on solid-phase by condensation of the terminal 5′-hydroxyl group with those phosphoramidite or H-phosphonate derivatives of ligands that are stable to the conditions required for synthesis. Many functional groups have been coupled to the 5′ terminus of oligonucleotides using these methods. Among them are intercalating agents, photoreactive groups, and labels. The acridine and psoralen derivatives were first linked to the 5′ ends of oligonucleotides bound to a solid support by use of the phosphoramidites S.115 (Fig. 4.2.21; Thuong and Chassignol, 1988) and S.116 (Kurfürst et al., 1993; Dupret et al., 1994), respectively. The phosphoramidite derivatives of these compounds with different linkers S.117 and S.118 are now commercially available. The pyrene group was also conjugated to the 5′ terminus of oligonucleotides using its phosphoramidites derivatives S.119 and S.120 (Korskun et al., 1992); likewise, fagaronine was linked to the 5′ ends of oligonucleotides via the parent phosphoramidite S.121 (Chen et al., 1992). Anthraquinone and sapphyrin were similarly attached to oligonucleotides from their H-phosphonate derivatives S.122 (Lin and Matteucci, 1991) and S.123 (Sessler et al., 1996), respectively.
Oligonucleotide-biotin conjugates S.124a, S.124b, and S.124c (Fig. 4.2.22) were prepared via phosphoramidites S.125a, S.125b (Cocuzza and Zagorsky, 1991) and S.125c (Olejnik et al., 1996). Conjugate S.124c, which has a photocleavable linker to enable the release of the oligonucleotide from the conjugate under UV irradiation at a wavelength of 300 to 350 nm. Fluorescent oligonucleotide conjugates derived from fluorescein S.126a are easily obtained via phosphoramidite S.127. Replacing the hydrogen atoms of the fluorescein nucleus by chlorine leads to tetra- and hexachlorofluorescein derivatives S.126b and S.126c, the absorption and emission spectra of which are red shifted. Designed to exploit energy-transfer techniques, oligomers labeled at their 5′ ends with two chromophores were developed as either linear structures, S.128 (Fig. 4.2.23) and S.129 (Ju et al., 1995, 1996), or hairpin structures, S.130 (Fig. 4.2.24; Nazarenko et al., 1997). Fluorescence energy transfer is a dipole-dipole resonance interaction between two chromophores near to each other. One of these chromophores must be fluorescent and is called the “donor”, whereas the other identified as the “acceptor” may or may not be fluorescent. A spectral overlap between the acceptor’s excitation and donor’s emission wavelengths, and a
N
O
H3CO
O
O
HN
O (CH2)3 O O P O O
(CH2)5 O O P O O
O
B
O
(CH2)3 O O
107
B
n
O O P O O (CH2)5 HN OCH3
O Cl
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
O
n
O P O O
O
Cl
N 108
Figure 4.2.20 Direct addition of ligands to the 5′ ends of oligonucleotides by the phosphotriester coupling method. See Figure 4.2.1 for definitions of functional group abbreviations.
4.2.20 Current Protocols in Nucleic Acid Chemistry
O O
HO O
O O P OPhCl-4 B O O
O 110 Cl
n
O
HO
O P OPhCl-4 O
HN
N
109
H3CO 111
NH NH
N N
O
HN
HN HN (CH2)6 O OH
HN (CH2)6 O O O P O O
N
N
114
B O O P OPhCl-4 B O O
n
O O P O O H
N
O P OPhCL-4 O
OCH3 Cl
n
O
(CH2)5
H
N
N
(CH2)5
112
OCH3 Cl
N 113
Figure 4.2.20
Continued
proper distance between the two entities are necessary for the donor to transfer energy, upon light excitation, to the acceptor. The efficiency of energy transfer is inversely proportional to the sixth power of the distance between the two chromophores. In the case of the linear oligomers S.128 and S.129, the donor was 5-carboxyfluorescein
(FAM), whereas the acceptor was either FAM or its derivative 6-carboxy-4′,5′-dichloro-2′,7′dimethoxyfluorescein (JOE), or one of the two rhodamine derivatives 5-(and 6-)carboxyN,N,N′,N′-tetramethyl rhodamine (TAMRA) and 5-(and 6-)carboxy-X-rhodamine (ROX; all available from Molecular Probes). Upon irradiation at 488 nm, the energy emitted by FAM
Synthesis of Modified Oligonucleotides and Conjugates
4.2.21 Current Protocols in Nucleic Acid Chemistry
CN
O ZL O P N(i-Pr)2 OCH3
O O
ZL =
N
NH(CH2)5
O(H2C)6
ZL = O
Cl
116
115 OCH3
O
O
ZL =
N
H N
H3C ZL = CH3
O
DMTrO
O
Cl 117
118 H N
(CH2)4 O
ZL =
ZL =
ODMTr
120
119 (CH2)6 H3CO
OCH3
ZL =
N
H3CO
CH3
121
DMTrO
N H
N ODMTr
O N
O 122
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
HN NH
O O P O H 123
N
H O P O O
Figure 4.2.21 Direct addition of ligands to the 5′ end of oligonucleotides by the phosphoramidite coupling method. See Figure 4.2.1 for definitions of functional group abbreviations.
4.2.22 Current Protocols in Nucleic Acid Chemistry
O O
NH
HN
H N S O
NH
HN L
H N
O O P O
S
124a-c
O
B
O
H N
H N b
O
O P N(i-Pr)2
O 125a-c
RO
a
CN L
L
=
H N
L
=
H N H3C
H N
c
=
L
O
H N
NO2
N H
R2
R2 HO
O
O
R1 R1
CO2
C HN
R1
O
R1
(CH2)6
R2 OAc
O O
R1
O
O O P O B
O
R2 AcO
O
O
C HN (CH2)6
R1
O O P
126a R1 = R2 = H 126b R1 = Cl, R2 = H 126c R1 = R2 = Cl
CN
N(i-Pr)2
RO 127a 127b 127c
R1 = R2 = H R1 = Cl, R2 = H R1 = R2 = Cl
Figure 4.2.22 Additional examples of direct addition of ligands to the 5′ ends of oligonucleotides by the phosphoramidite coupling method. See Figure 4.2.1 for definitions of functional group abbreviations.
is transferred to the acceptors which in turn generated strong fluorescence at characteristic wavelengths. Typically, when FAM is separated from the acceptor by six consecutive 1′,2′-dideoxyribose phosphates, the emission spectra of S.128a-d showed a 2- to 12-fold enhancement in fluorescence intensity relative to that of the corresponding singly-labeled oligomers. The hairpin structure S.130 is functionalized with both the fluorescent dye FAM at the fluorescence quencher 4-[(4-(dimethylamino)-
phenyl] azobenzoic acid (DABCYL). The hairpin keeps both chromophores in close proximity to each other, causing the fluorescence of the fluorophore to be quenched by energy transfer to the quencher (the energy is dissipated as heat). When the probe S.130 encounters a complementary target molecule, it undergoes a spontaneous conformational change that causes the fluorophore and the quencher to move away from each other. Since the fluorophore is no longer in close proximity to the quencher, it fluoresces upon UV irradiation.
Synthesis of Modified Oligonucleotides and Conjugates
4.2.23 Current Protocols in Nucleic Acid Chemistry
FAM O O P O O
O
R1
n
O O P O O
A U
O
O O P O O
O
B
n1
O H
HO 128a
R1 = H
129a
R1 = B
O
A =
O
CO2
O HO
O
O
H3CO
128b
R1 = H
129b
R1 = B
OCH3 CO2
A =
O (CH3)2N
128c 129c
R1 = H
O
N(CH3)2
CO2
A =
R1 = B O
N 128d 129d
R1 = H
O
N _ SO3
A =
R1 = B
O
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
Figure 4.2.23 Direct addition of ligands to the 5′ terminus of oligonucleotides to permit sensitive fluorescence detection. Figure 4.2.1 for definitions of functional group abbreviations.
4.2.24 Current Protocols in Nucleic Acid Chemistry
DABCYL 3' 5' FAM 130
= (Me)2N
DABCYL
N
O N
Figure 4.2.24 An additional example of direct addition of ligands to the 5′ ends of oligonucleotides to provide fluorescence detection.
Lastly, the 5′ terminus of oligonucleotides can be modified to tether other oligonucleotides through a linker and enable formation of triplehelices upon recognition of an oligopurine stretch on each limb of the conjugates S.131 (Asseline and Thuong, 1994).
the conjugate. For oligonucleotide conjugates with the same number of negative charges, separation can be performed by reversed-phase chromatography. Retention times of the conjugates are increased with the introduction of hydrophobic groups. The length and nature of the linkers used to join oligonucleotides and ligands also affect the chromatographic mobility of the conjugates. Purification of the oligonucleotide conjugates can also be achieved by polyacrylamide gel electrophoresis (PAGE). After purification the nucleoside composition of each conjugate can be verified by hydrolysis with nucleases such as endonuclease P1 (from Penicillium citrinum), snake venom
CHARACTERIZATION Characterization of oligonucleotides bearing a ligand at the 5′ terminus can be achieved in different ways. The oligonucleotide-ligand conjugates can be purified by ion-exchange chromatography (Asseline et al., 1986; Thuong and Asseline, 1991); the retention time increases with the number of negative charges on
O O P O R O
O
N
O CH3
N H
N
O
O
N
H N
N
H3C
O
O O
R O P O O 131
Figure 4.2.25 Conjugation of two third oligonucleotide strands for triple helical formation with double-stranded DNA targets via alternate strand recognition. See Figure 4.2.1 for definitions of functional group abbreviations.
Synthesis of Modified Oligonucleotides and Conjugates
4.2.25 Current Protocols in Nucleic Acid Chemistry
phosphodiesterase (from Crotalus durissus), and alkaline phosphatase. The resulting hydrolysates are then analyzed by comparison with commercial nucleoside samples or previously characterized nucleosides. Such analysis is performed by TLC or preferably by reversed-phase HPLC. It should be pointed out that ligand attachment to the 5′ end of an oligonucleotide prevents hydrolysis of the conjugate by 5′-exonuclease. Thus, 5′-oligonucleotide conjugates should be resistant to digestion by calf spleen phosphodiesterase. Recent developments in mass spectrometry, namely matrix-assisted laser desorption/ionization time of flight (MALDI-TOF; UNIT 10.1) and electrospray ionization (ESI, UNIT 10.2) techniques have allowed molecular mass determination of derivatized oligonucleotides. Chromophore-oligonucleotide conjugates can also be characterized by UV/fluorescence spectroscopy and circular dichroism.
PROPERTIES Numerous physicochemical, biochemical, and biological studies devoted to oligonucleotide-ligand conjugates have been widely reviewed. The discussion below will cover just a few observations on the hybridization properties of conjugates, followed by a description of a few specific properties and applications of 5′-derivatized oligonucleotides.
Formation of Double-Stranded Duplexes
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
Hybrid stabilities The stability of hybrids formed between one oligonucleotide-ligand conjugate and its complementary nucleic acid sequence depends on factors, such as the nature of the ligand-linker combination and the site where the ligand is conjugated to the oligonucleotide chain. The importance of these parameters has been studied with oligonucleotide–intercalating agent conjugates. For example, phenazinium derivatives have been linked to the 5′ or 3′ end of oligonucleotides (Lokhov et al., 1992) as well as to the C-5 positions of deoxyuridine (Levina et al., 1993). Acridine has been conjugated to the 5′ and 3′ termini of oligonucleotides, the internucleotidic phosphodiesters, and the C-5 positions of deoxyuridine residues (Asseline et al., 1984, 1996). Ethidium derivatives have been attached to either the 5′ or 3′ ends of short oligonucleotide sequences (Koshkin et al., 1994). Results obtained with these different conjugates indicate that the attachment of the
intercalating agents to the 5′ or 3′ ends of oligonucleotides via a phosphodiester or a phosphoramide linkage generally leads to the formation of more stable hybrids. Specific properties 5′-Conjugated oligonucleotides with a free 3′-hydroxyl functionality are widely used as primers for polymerases. Thus, oligomers bearing a 5′-fluorescent label such as fluorescein or rhodamine were first used as primers for DNA sequencing by enzymatic methods to allow sensitive, fast fluorescence detection (Smith et al., 1985). The doubly labeled primers S.128 and S.129 recently introduced for sequencing not only increase detection sensitivity but allow detection at many wavelengths to simplify DNA sequencing readouts (Ju et al., 1995, 1996). The use of primers labeled with a fluorescent group and a quencher (like S.130) allows direct quantification of synthesized PCR fragments in which the fluorescence is restored during amplification (Nazarenko et al., 1997). DNA fragments obtained by PCR from primers containing two adenines and a psoralen derivative at their 5′ end easily give photocross-linking products under irradiation at 360 nm with the thymines of complementary sequences. This “Chemi-clamp” system (Appligene-Oncor), enables very sensitive detection of point mutations contained in genes (Costes et al., 1993; Dupret et al., 1994) by the use of denaturing gradient (denaturing gradient gel electrophoresis, or DGGE; Børresen-Dale et al., 1998).
Formation of Triple-Helical Complex Triple-helix stabilities Oligonucleotides can bind to the major groove of DNA polypurine⋅polypyrimidine sequences to form triple helices. Recognition involves the formation of Hoogsteen or reversed-Hoogsteen hydrogen bonds between bases of the oligonucleotide third strand and the purines of the target DNA. This sequencespecific recognition of double-helical DNA was first described for pyrimidine oligonucleotides (Le Doan et al., 1987b; Moser and Dervan, 1987), then for oligonucleotides containing G and T (Cooney et al., 1988), G and A (Beal and Dervan, 1991; Pilch et al., 1991), and T, C, and G (Giovannangeli et al., 1992a). The attachment of an acridine derivative (Collier et al., 1991; Sun et al., 1989) to the 5′ end of a pyrimidine oligonucleotide was shown to strongly stabilize the triple-helical complex
4.2.26 Current Protocols in Nucleic Acid Chemistry
formed with DNA by intercalation at the triplex-duplex junction. When attached to the 5′ end of a short oligopurine strand, oxazolopyridocarbazole was also shown to stabilize the triple-helical complex (Mouscadet et al., 1994). Another example of triple helix stabilization is demonstrated by conjugation of a daunorubicin derivative to the 5′ terminus of an oligopyrimidine oligonucleotide (Garbesi et al., 1997). Studies carried out with benzopyridoindole derivatives linked to C-5 of deoxyuridine located at either the 5′ or 3′ ends or at internal positions in the oligonucleotide chain, as well as on internucleotidic bridges, via phosphorothiolate or phosphoramidate linkages have shown that only the 5′ oligonucleotide conjugates induce triple-helical stabilization (unpub. observ.). Other results obtained with oligopyrimidine oligonucleotides linked to BePI, BgPI, BfPQ, and BhPQ derivatives through either the 5′ end or internal sites have shown that the 5′ end is the most favorable site for intercalator conjugation (Silver et al., 1997). Irreversible modification of nucleic acid target sequences The ability of oligonucleotides bearing an intercalating agent at their 5′ end to form stable triple-helical complexes with the DNA target by intercalation of the polycyclic ligand into the double-helix/triple-helix junction has been exploited to develop reactive oligomers able to induce irreversible localized modifications of the double-stranded DNA targets. Specifically, alkylating agents, cleaving reagents such as porphyrin or Cu-phenanthroline, and photocross-linking reagents have been conjugated to oligonucleotides to modify target nucleic acids. Cross-linking. Many of the intercalating agents attached to oligonucleotides have been shown to cross-link target nucleic acids upon UVA or visible light irradiation. These include proflavin and its azido derivative (Praseuth et al., 1988a,b), ethidium and its azido derivative (Koshkin et al., 1994), and psoralen (Takasugi et al., 1991; Giovannangeli et al., 1992b). In the first two cases, cross-links can be converted to cleavage products upon alkaline treatment. In addition, photooxidation of guanine bases by singlet oxygen production resulted from the reaction of an excited proflavin with molecular oxygen. In the case of psoralen, a photochemical [2+2]-cycloaddition reaction involves the furan or pyrone ring of psoralen and the 5,6double bond of thymine. The yield of crosslinked structures depends on the attachment site
and the linker used to tether the psoralen to the oligonucleotide. When the target is a DNA duplex, a psoralen derivative can be used to cross-link the two strands of the double helix when conjugated to the 5′ end of a pyrimidine triple-helix-forming oligonucleotide (Takasugi et al., 1991; Giovannangeli et al., 1992b). This reaction requires a 5′-TpA-3′ sequence at the junction between the duplex and triplex structures on the oligopurine-containing strand of the target duplex so that two thymines are properly located to form cyclobutane adducts with both the furan and the pyrone rings of a psoralen intercalated at the junction. Since the 5′-TpA-3′ sequence is self-complementary, one thymine is located in the duplex region on one strand and the other in the triplex region on the other duplex strand. The resulting product has psoralen linked to the three strands of the triplex structure (one covalent bond attaching psoralen to the oligonucleotide third strand and two photochemically generated bonds to thymines on the target DNA). Recently, N-5-methylcyclopropapyrroloindole—a structural analog of cyclopropapyrroloindole (CPI), the reactive subunit of antibiotic CC-1065, which alkylates adenines at N3—was conjugated to the 5′ end of an oligonucleotide (Lukhtanov et al., 1997b). This conjugate has proven capable of alkylating the target DNA duplex region immediately adjacent to its double-stranded complementary binding region with which it forms a triple helix. The remarkable stability of the CPI alkylating function to side reactions in physiological buffers allows the reactive oligonucleotide conjugate to rapidly and efficiently cross-link to the DNA target. Cleavage of the target DNA sequence. Cleaving reagents such as porphyrin, phenanthroline-Cu, or bleomycin-Cu have been linked to the 5′ ends of oligonucleotides in order to induce specific cuts on target nucleic acids. Among them orthophenanthroline (OP) used in the presence of Cu2+ and a reducing agent has proven to be the most efficient (Chen and Sigman, 1986, 1988; François et al., 1989). Using a 5′ oligonucleotide-OP conjugate bound to duplex DNA through a triple-helical structure, phenanthroline has been shown to intercalate at the triplex-duplex junction and induce cleavage of both DNA strands with high efficiency— up to 70% double-strand cleavage (François et al., 1989). Since OP is covalently linked to the oligonucleotide via its 5′ terminus, the oligonucleotide binds to the major groove of the DNA
Synthesis of Modified Oligonucleotides and Conjugates
4.2.27 Current Protocols in Nucleic Acid Chemistry
duplex. OP then intercalates at the duplex-triplex junction within the minor groove. Therefore, Cu2+ chelation and the cleavage reaction take place in the minor groove even though recognition of the duplex sequence occurs in the major groove. The cleavage reaction was shown to be sequence specific. Using SV40 circular DNA (5000 bp) as a target, a single double-stranded cut was induced by an OPoligonucleotide conjugate (François et al., 1989), providing a basis for sequence-specific artificial endonucleases. Inhibition of gene expression via triple-helix formation The synthesis of oligonucleotide-intercalator conjugates has paved the way for the development of new tools for molecular and cellular biology and for the design of new gene expression regulators. Oligonucleotides bearing an intercalator at their 5′ end have been used in the “antigene” strategy in which the target is double-helical DNA. The formation of a triplehelical complex within the promoter region of a selected gene may inhibit binding of transcription factors, thereby inhibiting transcription initiation (Grigoriev et al., 1992); this inhibition can be made irreversible when a psoralen conjugate is used in the presence of UV light (Grigoriev et al., 1993). Psoralen conjugates were also used as tools to demonstrate that DNA target sequences were accessible within the chromatin structure compressing genomic DNA in the cell nucleus (Giovannangeli et al., 1997). An acridine-oligonucleotide conjugate was recently shown to arrest the elongation of an RNA transcript when involved in a triplehelical complex formed downstream of the transcription initiation site (Escudé et al., 1996). Unconjugated oligomer did not exhibit any effect on transcription elongation and demonstrated that the gain in stability provided by the intercalator was required for a biological response to be observed.
CONCLUSION
Modification of the 5′ Terminus of Oligonucleotides for Attachment of Reporter and Conjugate Groups
Numerous ligands have been covalently linked to the 5′ end of oligonucleotides. Coupling at this location is among the easiest to perform and generally produces conjugates with good hybridization properties. These conjugates are very useful in the applications of oligonucleotides in various fields, as they possess specific properties. The presence of the free 3′-terminal hydroxyl group allows their use as primers for polymerases, and the strong
stabilization provided by intercalator ligands makes them efficient inhibitors of gene expression via triple-helix formation.
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Shimizu, M., Inoue, H., and Ohtsuka, E. 1994. Detailed study of sequence-specific DNA cleavage of triplex-forming oligonucleotides linked to 1,10-phenanthroline. Biochemistry 33:606613. Silver, G.C., Sun, J.S., Nguyen, C.H., Boutorin, A.S., Bisagni, E., and Hélène, C. 1997. Stable triple-helical DNA complexes formed by benzopyridoindole- and benzopyridoquinoxalineoligonucleotide conjugates. J. Am. Chem. Soc. 119:263-268. Smith, L.M., Fung, S., Hunkapiller, M.W., Hunkapiller, T., and Hood, L.E. 1985. The synthesis of oligonucleotides containing an aliphatic amino group at the 5′ terminus: Synthesis of fluorescent DNA primers for use in DNA sequence analysis. Nucl. Acids Res. 13:23992412. Sproat, B.S., Beijer, B., Rider, P., and Neumer, P. 1987a. The synthesis of protected 5′-mercapto2′,5′-dideoxyribonucleoside-3′-O-phosphoram idites; uses of 5′-mercapto-oligo-deoxyribonucleotides. Nucl. Acids Res. 15:4837-4848. Sproat, B.S., Beijer, B., and Rider, P. 1987b. The synthesis of protected 5′-amino-2′,5′-dideoxyribonucleoside-3′-O-phosphoramidites; applications of 5′-amino-oligodeoxyribonucleotides. Nucl. Acids Res. 15:6181-6196. Soyfer, V.N. and Potoman, V.N. 1996. Possible spheres of applications of intermolecular triplexes. In Triple-Helical Nucleic Acids, pp. 253284. Springer-Verlag, New York. Stec, W.J., Uznanski, B., and Wilk, A. 1993. Bis(O,O-diisopropoxy phosphinothioyl)-disulfide-A highly efficient sulfurizing reagent for cost-effective synthesis of oligo(nucleoside phosphorothioate)s. Tetrahedron Lett. 34:53175320. Sun, J.S., Francois, J.C., Montenay-Garestier, T., Saison-Behmoaras, T., Roig, V., Thuong, N.T., and Hélène, C. 1989. Sequence-specific intercalating agents. Intercalation at specific sequences on duplex DNA via major groove recognition by oligonucleotide-intercalator conjugates. Proc. Natl. Acad. Sci. U.S.A. 86:9198-9202. Sun, J.S., Giovannangeli, C., Francois, J.C., Kurfürst, R., Montenay-Garestier, T., Asseline, U., Saison-Behmoaras, T., Thuong, N.T., and Hélène, C. 1991. Triple-helix formation by α oligodeoxynucleotides and α oligodeoxynucleotide-intercalator conjugates. Proc. Natl. Acad. Sci. U.S.A. 88:6023-6027. Takasugi, M., Guendouz, A., Chassignol, M., Decout, J.L., Lhomme, J., Thuong, N.T., and Hélène, C. 1991. Sequence-specific photo-induced cross-linking of the two strands of doublehelical DNA by a psoralen covalently linked to a triple helic forming oligonucleotide. Proc. Natl. Acad. Sci. U.S.A. 88:5602-5606. Tam, J.P., Wu, C.-R., Liu, W., and Zhang, J.-W. 1991. Disulfide bond formation in peptides by dimethyl sulfoxide. Scope and applications. J. Am. Chem. Soc. 113:6657-6662.
4.2.32 Current Protocols in Nucleic Acid Chemistry
Thorpe, P.E., Wallace, P.M., Knowles, P.P., Relf, M.G., Brown, A.N.F., Watson, G.J., Knyba, R.E., Wawrzynczak, E.J., and Blakey, D.C. 1987. New coupling agents for synthesis of immunotoxins containing a hindered disulfide bond with improved stability in vivo. Cancer Res. 47:59245931. Thuong, N.T. and Asseline, U. 1991. Oligonucleotides attached to intercalators, photoreactive and cleavage agents. In Oligonucleotides and Analogues: A Practical Approach (F. Eckstein, ed.) pp. 283-308. IRL Press, Oxford. Thuong, N.T. and Chassignol, M. 1987. Synthèse et reactivité d’oligothymidylates substitués par un agent intercalant et un groupe thiophosphate. Tetrahedron Lett. 28:4157-4160. Thuong, N.T. and Chassignol, M. 1988. Solid phase synthesis of oligo-α- and oligo-β deoxynucleotides covalently linked to an acridine. Tetrahedron Lett. 29:5905-5908. Thuong, N.T. and Hélène, C. 1993. Sequencespecific recognition and modification of doublehelical DNA by oligonucleotides. Angew. Chem. Int. Ed. Engl. 32:666-690. Thuong, N.T., Hélène, C., and Asseline, U. 1984. European Patent, 84-400143-8. United States Patent, 1989, 4-835-263.
Vu, H. and Hirschbein, B.L. 1991. Internucleotide phosphite sulfurization with tetraethylthiuram disulfide phosphorothioate oligonucleotide synthesis via phosphoramidite chemistry. Tetrahedron Lett. 32:3005-3008. Wachter, L., Jablonski, J.-A., and Ramachandran, K.L. 1986. A simple and efficient procedure for the synthesis of 5′-aminoalkyl oligodeoxynucleotides. Nucl. Acids Res. 14:7985-7994. Zarytova, V.F., Godovikova, T.S., Kutyavin, I.V., and Khalimskaya L.M. 1987. Reactive oligonucleotide derivatives as affinity reagents and probes in molecular biology. In Biophosphates and Their Analogues. Synthesis, Structure, Metabolism and Activity (K.S. Brusik and W.J. Stec, eds.) pp. 149-164. Elsevier Science Publishers, New York. Zarytova, V.F., Sergeyev, D.S., and Godovikova, T.S. 1993. Synthesis of bleomycin A5 oligonucleotide derivatives and site-specific cleavage of the DNA target. Bioconjugate Chem. 4:189-193.
Contributed by Nguyen T. Thuong and Ulysse Asseline Centre de Biophysique Moléculaire, CNRS Orléans, France
Uhlmann, E. and Engels, J. 1986. Chemical 5′-phosphorylation of oligonucleotides valuable in automated DNA synthesis. Terahedron Lett. 27:1023-1026.
Synthesis of Modified Oligonucleotides and Conjugates
4.2.33 Current Protocols in Nucleic Acid Chemistry
Supplement 7
Direct Attachment of Conjugate Groups to the 5′ Terminus of Oligodeoxyribonucleotides
UNIT 4.3
The attachment of conjugate groups (intercalating and photoreactive groups) to the 5′ terminus of oligodeoxyribonucleotides can be achieved using either of two strategies. This unit presents one strategy, the direct incorporation of ligands during oligodeoxyribonucleotide synthesis using phosphoramidite derivatives. The second strategy, coupling of an unblocked oligomer with a ligand via a specific reaction between the reactive groups present in both entities, will be presented in future units. The protocols delineated below cover the direct addition of an intercalator, 2-methoxy-6-chloro-9-aminoacridine (see Basic Protocol 1), and a photo-cross-linking reagent, psoralen (see Basic Protocol 2), to the 5′ end of oligonucleotides via their phosphoramidite derivatives. These procedures require the following steps: the incorporation of hydroxylated linkers into ligands, the preparation of the phosphoramidite derivatives, and the addition of the latter to the 5 ′ ends of protected oligonucleotides bound to the support. After deprotection, the oligodeoxyribonucleotide conjugates are purified and characterized (see Support Protocol). Properties of a number of conjugates synthesized by these procedures are discussed in UNIT 4.2. CAUTION: All chemicals must be handled in a fume hood by individuals equipped with lab coats, glasses, and gloves. DIRECT ADDITION OF ACRIDINE DERIVATIVES TO THE 5′ END OF OLIGODEOXYRIBONUCLEOTIDES
BASIC PROTOCOL 1
This strategy (illustrated in Figs. 4.3.1, 4.3.2, and 4.3.3) involves the covalent attachment of a linker carrying a hydroxyl function to the acridine derivative S.1b, and the preparation of the acridinyl phosphoramidite derivative S.1c. The acridinyl phosphoramidite derivative can be coupled either automatically or manually to the 5′-terminal hydroxyl group of oligodeoxyribonucleotides bound to solid supports. After deprotection, the acridineoligodeoxyribonucleotide conjugate S.1d is purified and characterized as described in the Support Protocol.
OCH3
OCH3
H2N(CH2)6OH N
Cl
N
N H
6 OH
OH Cl
Cl 1a
1b
O
O
O
O
N HCl
O
O Br(CH2)6OH
OCH3 O
Figure 4.3.1 linkers.
K2CO3
6 OH
O
O 2a
O
OH
2b
2c
Synthesis of acridine and psoralen derivatives functionalized with hydroxylated
Contributed by Ulysse Asseline and Nguyen T. Thuong Current Protocols in Nucleic Acid Chemistry (2000) 4.3.1-4.3.16 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Modified Oligonucleotides and Conjugates
4.3.1
OCH3 Cl
OCH3
CN
O P
N
N H
6 OH
N
N H
DIEA
Cl
6O
P N(i-Pr)2
Cl 1b
1c
O O
Cl O
O
CN
O
O
P
6 OH
CN
O
N(i-Pr)2 O
6O
DIEA
O
P N(i-Pr)2
O 2c
Figure 4.3.2
CN
O
N(i-Pr)2
2d
Synthesis of the phosphoramidite derivatives of acridine and psoralen ligands.
HO
B'
O O
O P O O
CN O
B'
n
O P OCH3 O
1. N
O NH (CH2)6 O P N(i-Pr)2 1c
Cl
O
CN 1.
(CH2)6 O P O
2d 1H-tetrazole
1H-tetrazole
3. NH4OH
3. NaOH
N
O
Cl
H3CO HN
O
O
O (CH2)6
(CH2)6
O
O
O P O O
O P O O
O
O
B'
n
O
H 1d
Figure 4.3.3
N(i-Pr)2
2. oxidation
2. oxidation
Direct Attachment of Conjugate Groups to the 5′ Terminus of Oligodeoxyribonucleotides
CN
O
O
O
B'
n
H 2e
Direct addition of ligands to the 5′ terminus of oligodeoxyribonucleotides.
4.3.2 Current Protocols in Nucleic Acid Chemistry
Materials 6-Amino-1-hexanol 6,9-Dichloro-2-methoxyacridine Phenol Dichloromethane (CH2Cl2), distilled from P2O5 and passed over activated, basic aluminum oxide Distilled methanol (MeOH) 2 M sodium hydroxide (NaOH) Acetonitrile (CH3CN), HPLC grade 2,6-Dibromo-4-benzoquinone-N-chloroimine (DBPNC; Prolabo) Ethanol, distilled P2O5 Acetonitrile (CH3CN), DNA synthesis grade, anhydrous and stored over 3-Å molecular sieves N,N-Diisopropylethylamine (DIEA), distilled from KOH 2-Cyanoethyl-N,N-diisopropylchlorophosphite Ethyl acetate, distilled Hexane, distilled Triethylamine (Et3N), distilled from KOH 10% (w/v) Na2CO3 Saturated NaCl (brine) MgSO4 Oligodeoxyribonucleotide bound to solid supports, such as controlled-pore glass (CPG; UNIT 3.1), synthesized by classic phosphoramidite chemistry (1-µmol scale; UNIT 3.3 and APPENDIX 3C) DNA synthesis reagents recommended by the synthesizer’s manufacturer 0.4 M sodium hydroxide (NaOH) in 50:50 (v/v) MeOH/H 2O Dowex 50 resin, pyridinium form (Aldrich) Round-bottomed flasks (various sizes) with rubber septa and glass stoppers Drying tube containing CaCl2 Reflux condenser Analytical thin-layer chromatography (TLC) setup and silica-gel plates: e.g., Merck 5554 Kieselgel 60F plates, including UV lamp for detection Desiccator Beakers, various sizes Silica gel for column chromatography: Merck 9387 Kieselgel 60 or Merck 7734 Kieselgel 60 Chromatographic column (3 cm × 45 cm) Argon Nitrogen gas Spectrophotometer Rotary evaporator with water bath Chemically inert syringes with replaceable needles Funnel Separatory funnels (various sizes) Filter (porosity 4) Vials and Teflon-faced septa Disposable filters for syringes and for filtration of HPLC buffers Liquid chromatography apparatus equipped with multiwavelength detector Melting-point apparatus
Synthesis of Modified Oligonucleotides and Conjugates
4.3.3 Current Protocols in Nucleic Acid Chemistry
Synthesize 2-methoxy-6-chloro-9-(ω-hydroxyhexylamino)-acridine S.1b 1. Place 840 mg (7.18 mmol) 6-amino-1-hexanol, 1 g (3.59 mmol) 6,9-dichloro-2methoxyacridine, and 2.5 g phenol in a 25-mL round-bottomed flask equipped with a magnetic stir-bar, a reflux condenser, and a calcium chloride drying tube. Heat 80 to 90 min at 100°C. 2. Perform TLC analysis on a silica-gel plate using 90:10 (v/v) CH 2Cl2/MeOH as the eluent to check that the reaction is complete. Dilute an aliquot 10-fold with MeOH and load 1 µl on the TLC plate. This should show the disappearance of 6,9-dichloro-2-methoxyacridine (Rf = 0.9) and the formation of a new compound (Rf = 0.18).
3. Allow the mixture to cool, dilute it with 2.5 mL MeOH, and add the solution dropwise to a magnetically stirred 2 M NaOH solution (20 mL contained in a 100-mL beaker). Maintain stirring for 15 min. 4. Filter the yellow precipitate using a porosity 4 filter and wash it with water until neutral. Crystallize twice from 20:80 (v/v) H2O/MeOH. Expected yield: 80% (1.03 g, 2.87 mmol).
5. Analyze the product by TLC using silica-gel plates and 75:25 (v/v) CH2Cl2/MeOH as eluent. Dilute 1 to 2 mg in 100 µl MeOH and use 1 µl for spotting. In addition to 2-methoxy-6-chloro-9-(w-hydroxyhexylamino)-acridine (S.1b, Rf = 0.58),TLC analysis will show three side products, identified as the starting material 6,9-dichloro-2-methoxyacridine (Rf = 0.85), 2-methoxy-6-chloro-9-acridone (Rf = 0.81), and an unidentified third product (Rf = 0.39). These three products do not react during subsequent steps and are easily removed from the conjugates during purification. Reversed-phase HPLC as reported in Asseline et al., 1986 can also be used. Purity should be ∼95%. Characterize the product by other analytical methods as desired. Melting point: 152° to 153°C. H-NMR (DMSO-d6): δ 1.15-1.40 (m, 6H, CH2CH2CH2), 1.60-1.75 (m, 2H, CH2), 3.203.33 (m, 3H, CH2N + NH), 3.61-3.75 (m, 2H, CH2OH), 3.90 (s, 3H, OCH3), 4.20-4.28, (m, 1H), 7.25-7.88 (m, 5H, Acr), 8.25-8.38 (m, 1H, H8 Acr).
1
Mass analysis: I.S. Polarity positive. Calculated for C20H23ClN2O2: M+H= 359.18 and 361.18. Found 359.5 and 361.5.
6. Check for the absence of sodium phenolate by TLC analysis using the conditions described in step 5. Place plate in fume hood and dry using a hair dryer. Spray the TLC plate with a solution of 50 mg 2,6-dibromo-4-benzoquinone-N-chloroimine (DBPNC) in 20 mL ethanol and heat with a TLC drying plate or in an oven at 100°–130°C for several minutes. Phenol will give a blue-colored spot.
Direct Attachment of Conjugate Groups to the 5′ Terminus of Oligodeoxyribonucleotides
Prepare [2-methoxy-6-chloro-9-(ω-hexylamino)-acridinyl]-(2-cyanoethyl)-N,N-diisopropyl-phosphoramidite, S.1c 7. Place 300 mg (0.83 mmol) 2-methoxy-6-chloro-9-(ω-hydroxyhexylamino)acridine, S.1b, in a 25-mL round-bottomed flask and dry it by three rounds of coevaporation with anhydrous acetonitrile (3 × 5 mL). 8. Place a stir bar in the flask and stopper it with a septum. Push a needle through the septum and leave the flask in a P2O5-containing desiccator under vacuum overnight.
4.3.4 Current Protocols in Nucleic Acid Chemistry
9. Fill the desiccator with nitrogen before opening it. 10. Under argon atmosphere and magnetic stirring, add with a syringe 6 mL anhydrous CH3CN, 0.32 mL DIEA (235 mg, 1.82 mmol), and 0.2 mL 2-cyanoethyl-N,N-diisopropylchlorophosphite (216 mg, 0.91 mmol) at room temperature. 11. Let reaction proceed 30 min. Monitor its progress by TLC on a silica-gel plate using 50:50:4 (v/v/v) ethyl acetate:hexane:Et3N as eluent. Pre-elute the TLC plate once with the eluent, then load 1 µl of the reaction mixture and elute. After 30 min the starting material S.1b (Rf = 0.10) is transformed into two new compounds, a main product (≈80%; Rf = 0.50) and a side-product (≈15%; Rf = 0.65). Trace amounts of the starting material are also observed.
12. Add 20 mL CH2Cl2 and wash the organic layer with 3 mL of 10% (w/v) aqueous Na2CO3 and then 3 mL of saturated NaCl (brine). Decant the organic phase, dry it over MgSO4, and concentrate it to dryness. 13. Apply the residue, diluted with 2 to 3 mL anhydrous CH3CN to a silica-gel chromatography column (3 cm × 45 cm, containing 60 g silica gel) and elute with 500 mL of 50:50:4 (v/v/v) ethyl acetate:hexane:Et 3N. Monitor the fractions by TLC analysis using 50:50:4 (v/v/v) ethyl acetate:hexane:Et 3N as eluent, and collect the main product (Rf = 0.50). 14. Pool the fractions containing the pure product, and remove the solvent under reduced pressure. Coevaporate the residue twice using 8 mL anhydrous CH3CN each time. Expected yield: 65% (300 mg, 0.54 mmol) of a yellow oil. Purified acridinyl phosphoramidite S.1c can be stored at −20°C for a few months and should be reconstituted in anhydrous acetonitrile just prior to use (add a few pearls of 3-Å molecular sieves and wait ∼30 min before using the solution). P-NMR (CDCl3): δ 143.52 ppm.
31
H-NMR (CDCl3): δ 1.09-1.20 (m, 12H-[C(H)(CH3)2]2), 1.35-1.50 (m, 2H, CH2), 1.561.68 (m, 4H, CH2CH2), 1.70-1.83 (m, 2H, CH2), 2.61 (t, 2H, J = 6.34 Hz, CH2CN), 3.20-3.33 (m, 3H, CH2N + NH), 3.50-3.87 (m, 6H, CH2OP, (2 CH)), 3.97 (s, 3H, OCH3), 7.17-8.10 (m, 6H, Acr) 1
Couple the acridinyl phosphoramidite S.1c to 5′ end of oligodeoxyribonucleotide chain Automated coupling: 15a. When employing standard solid-phase DNA oligonucleotide synthesis cycles (1µmol scale) according to the classical phosphoramidite method, perform an additional detritylation step. 16a. Place a 0.1 M solution of the acridinyl phosphoramidite S.1c in anhydrous CH3CN on the synthesizer. Carry out coupling by recycling a mixture of 0.1 mL of the acridinyl phosphoramidite, 0.5 mL of 0.4 M 1H-tetrazole in anhydrous CH3CN, and 0.1 mL anhydrous CH3CN for 5 min. Repeat. For synthesizers without a recycle program, perform a double coupling step. For optimal coupling yields, changes in phosphoramidite concentration may be required depending on the synthesizer used.
17a. Remove the excess phosphoramidite and tetrazole, and perform an iodine oxidation reaction according to the standard procedure (UNIT 3.3, APPENDIX 3C).
Synthesis of Modified Oligonucleotides and Conjugates
4.3.5 Current Protocols in Nucleic Acid Chemistry
Manual coupling: 15b. When performing standard solid-phase oligodeoxyribonucleotide synthesis (1µmol scale) according to the classical phosphoramidite method, place the oligonucleotide bound to the support in a short (4-mL) vial stoppered with a septum. Push a needle through the septum and leave the vial in a desiccator containing P 2O5 under vacuum overnight. The oligonucleotide must have a free 5′-hydroxy group.
16b. Fill the desiccator with nitrogen before opening it. With a syringe, add a mixture of phosphoramidite S.1c (0.15 mL of a 0.15 M solution in anhydrous CH3CN) and 1H-tetrazole (0.5 mL of a 0.5 M solution in anhydrous CH 3CN) to the support-linked oligonucleotide. Leave the mixture for 10 min, shaking gently by hand from time to time. 17b. Remove the excess reagents with a syringe and perform an oxidation reaction by adding 1 mL of an aqueous iodine solution that is normally used on the synthesizer. Remove the iodine solution after 1 min and wash the support extensively with CH3CN. Deprotect the acridine-oligodeoxyribonucleotide conjugates 18. Treat the support bearing the acridine-oligodeoxyribonucleotide conjugates with 5 mL of 0.4 M NaOH in 50:50 (v/v) MeOH/H2O at room temperature. 19. After 1 to 2 hr, remove the solid support by filtration. Wash it twice with 0.5 mL water. Maintain the yellow solution at room temperature until deprotection is complete. The time required for oligonucleotide deprotection depends on the base composition of the sequence and on the nature of the protective groups used to protect the nucleic bases. Removal of the isobutyryl group from guanine residues requires ∼30 to 35 hr under these conditions.
20. Neutralize the solution (containing the oligonucleotide conjugate S.1d) by adding Dowex 50 ion-exchange resin until the pH of the solution reaches 6. Remove the resin by filtration and wash it with water until the resin becomes pale yellow in color. 21. Concentrate the solution under vacuum to remove MeOH. Add ∼10 mL water, then extract the aqueous solution three times with 3 mL ethyl acetate each time. Concentrate the aqueous solution to ∼3 to 4 mL. 22. Filter the solution through a 0.45-µm disposable filter to remove any particulates and prevent the clogging of HPLC columns. Purify and characterize the oligonucleotideacridine conjugate (see Support Protocol). SUPPORT PROTOCOL
Direct Attachment of Conjugate Groups to the 5′ Terminus of Oligodeoxyribonucleotides
PURIFICATION AND CHARACTERIZATION OF OLIGONUCLEOTIDE-ACRIDINE CONJUGATES The following general procedure is applicable to the purification of 10- to 20-mer oligodeoxyribonucleotides-acridine conjugates, assuming that the sequences under consideration are not likely to form self-associated complexes such as G-tetrads. Should this be the case, a specific purification procedure—not provided in this unit—must be used. Purification can by improved by performing ion-exchange chromatography at pH 12 (Raynaud et al., 1996). Another solution to prevent the formation of G tetrads is to partially replace 2’-deoxyguanosine with 7-deaza-2′deoxyguanosine (Raynaud et al., 1996). Generally, the crude oligodeoxyribonucleotide conjugates obtained from the protocol described above are purified by ion-exchange chromatography. After a desalting step, the
4.3.6 Current Protocols in Nucleic Acid Chemistry
purity of the conjugates is verified on a reversed-phase column; in most cases this will show that sufficient purity has been achieved. Alternatively, reversed-phase chromatography allows easy purification of oligodeoxyribonucleotide-acridine conjugates when acridine is bound to the 5′ end of oligonucleotides (making the conjugates more lipophilic than the underivatized oligonucleotides). Sometimes the conjugate can be insufficiently pure and a second purification is required. Usually it is performed using the system that was not used for the first purification (i.e., a second purification step can be made using reversed-phase after a first purification by ion-exchange chromatography and vice versa). In all cases, oligodeoxyribonucleotide-acridine conjugates are detected during analysis and purification by measuring UV absorption at 254 or 260 nm and at 425 nm (only the acridine moiety of the conjugate absorbs light at that wavelength). Materials 0.01 bis-Tris, pH 6 or 0.01 M NaH2PO4, pH 6.8 (APPENDIX 2A), each containing 10% or 10% HPLC-grade methanol (MeOH) Sodium chloride (NaCl) 0.025 M Tris⋅Cl, pH 7 (APPENDIX 2A) Acetonitrile (CH3CN) 1 M triethylammonium acetate (TEAA) buffer, pH 7 (stock solution; see recipe) 0.01 M Tris⋅Cl, pH 8 (APPENDIX 2A) 1 U snake venom phosphodiesterase (3′ exonuclease) 10 µg alkaline phosphatase Ion-exchange columns: Mono Q HR 5/5 or HR 10/10 (Amersham Pharmacia Biotech) or DEAE (Waters) Reversed-phase columns: e.g., Lichrospher 100 RP 18 (5 µm, 125 × 4 mm, or 10 µm, 250 ×10 mm; Merck) or CC Nucleosil 100-5 C18 (125/4) column (Macherey-Nagel) Desalting columns: HR 10/10 columns (Amersham Pharmacia Biotech) packed with Lichroprep RP 18 (Merck) or Sephadex G-10 or G-25 resin 1. Analyze the crude oligonucleotide-acridine conjugate solution via chromatography by one of the following procedures, generally loading 10 to 20 µl of the crude solution onto the column: a. Ion-exchange chromatography with Mono Q HR 5/5 or HR 10/10 column: Use a linear gradient of 0 to 100% 1 M NaCl in 0.01 M bis-Tris, pH 6, containing 10% or 20% MeOH, or 0 to 100% 1.5 M NaCl in 0.01 M NaH2PO4, pH 6.8, containing 10% or 20% MeOH, with a flow rate of 1 mL/min (HR 5/5) or 4 mL/min (HR 10/10). b. Ion-exchange chromatography with DEAE column (100 × 10 mm) : Use a linear gradient of 0 to 100% 1.5 M NaCl in 0.025 M Tris⋅Cl, pH 7, containing 10% MeOH, with a flow rate of 1 mL/min. c. Reversed-phase chromatography with Lichrospher 100 RP 18 column (5 µm, 125 × 4 mm; or 10 µm, 250 × 10 mm): Use a linear gradient of CH3CN (5% to 80%) in 0.1 M aqueous triethylammonium acetate, pH 7, with a flow rate of 1 mL/min (5-µm column) or 4 mL/min (10-µm column). d. Reversed-phase chromatography with CC Nucleosil 100-5 C18 (125/4) column: Use a linear gradient of CH3CN (5% to 80%) in 0.1 M aqueous triethylammonium acetate, pH 7, with a flow rate of 1 mL/min. The gradients used must be chosen so as to afford the best separation. Usually an increasing concentration of 1% of 1 M or 1.5 M NaCl per minute is used for ion-exchange chromatography and an increasing concentration of ∼1% CH3CN per minute is used for reversedphase chromatography.
Synthesis of Modified Oligonucleotides and Conjugates
4.3.7 Current Protocols in Nucleic Acid Chemistry
2. Purify the remaining oligonucleotide-acridine conjugate by preparative reversedphase or ion-exchange chromatography column. Use any type of column that was described in step 1, according to the recommended conditions. Usually 1 mmol of crude oligonucleotide-acridine conjugates can be purified in two or three runs on Mono Q HR 10/10 column. On preparative reversed-phase columns, reasonable purification can be achieved in a single run, but it is better to perform two. Alternatively, purification can be achieved on analytical columns by performing five or more runs.
3. Pool the fractions containing the conjugate and remove the organic solvents by evaporation under reduced pressure. 4. If ion-exchange purification was used in step 2, desalt the conjugate using appropriate desalting column. If reversed-phase purification was used, remove triethylammonium acetate by three consecutive lyophilizations. Resuspend the sample in 1 to 2 mL water between lyophilizations. 5. Analyze the oligonucleotide-acridine conjugate by ion-exchange chromatography when the purification was performed by reversed-phase, or by reversed-phase when the purification was achieved by ion-exchange, using the columns and parameters described in step 1. For example, analyze on a Lichrospher 100 RP 18 (5 mm; 125 mm × 4 mm) column from Merck using a linear gradient of CH3CN (5% to 35% over 40 min) in 0.1 M aqueous triethylammonium acetate, pH 7, with a flow rate of 1 mL/min. Expected retention time is 28 min 18 sec for the acridine-oligonucleotide conjugate Acr(CH2)6pd5[T3C2T C2TCT]. After purification, UV spectra of oligodeoxyribonucleotide-acridine conjugates show absorptions characteristic to the ligand. A typical spectrum of an oligonucleotide-acridine conjugate is shown in Figure 4.3.4 (left). In addition, nuclease digestion of the conjugates and reversed-phase HPLC analysis of the resulting monomers can be used to ascertain full removal of the protective groups and base composition.
Direct Attachment of Conjugate Groups to the 5′ Terminus of Oligodeoxyribonucleotides
Figure 4.3.4 Absorption spectra of Acr(CH2)6pd[T3C2T C2TCT] (left) and Pso(CH2)6pd[T4 5-Me C2T5-MeCT5-MeC3T5-MeCT] (right) recorded in water.
4.3.8 Current Protocols in Nucleic Acid Chemistry
6. Hydrolyze the oligonucleotide-acridine conjugate S.1d with nucleases by placing the following in a 1.5-mL microcentrifuge tube: ∼1 OD unit conjugate S.1d in 10 µl H2O 50 µl 0.01 M Tris⋅Cl, pH 8 1 U snake venom phosphodiesterase 10 µg alkaline phosphatase. Incubate 2 to 3 hr at 37°C. 7. Inactivate the enzyme by heating the mixture at 90°C for 2 min. 8. Analyse the hydrolysate by reversed-phase HPLC: e.g., using a Lichrospher 100 RP 18 column with a gradient of CH3CN (0% for 10 min, then 0 to 24% CH3CN over 30 min, then 24% to 56% CH3CN over 20 min) in 0.1 M aqueous triethylammonium acetate, pH 7, with a flow rate of 1 mL/min. These conditions permit the separation of natural nucleosides and a variety of modified nucleosides. Identification can be made by comparison with authentic nucleosides. For example: RtdC ≈ 5 min, Rt 5-MedC ≈ 10 min, RtT ≈ 13 min, RtdG ≈ 14 min 30 sec, RtdA ≈ 17 min 30 sec. Under these conditions the retention time of the acridine-linker derivative S.1b is ∼51 min. Prolonged incubation at pH 8 induces degradation of S.1b. Additional characterization can be made by MALDI and ESI mass spectrometry (UNITS 10.1 & 10.2). Acr(CH2)6pd[T3C2T C2TCT] has been characterized by MALDI-TOF using 2,4,6-trihydroxyacetophenone as matrix in the presence of diammonium L-tartrate. Quantification of oligonucleotide-acridine conjugates can be made by using e = 8850 M−1 cm−1 at l = 425 nm (Asseline and Thuong, 1988). Usually when the synthesis is performed at the micromole scale ∼15 to 20 OD units of pure oligonucleotide-acridine conjugate 15-mer with mixed bases is obtained.
DIRECT ADDITION OF PSORALEN DERIVATIVES TO THE 5′ END OF OLIGODEOXYRIBONUCLEOTIDES
BASIC PROTOCOL 2
This strategy (illustrated in Figs. 4.3.1, 4.3.2, and 4.3.3) involves the attachment of a hydroxylated linker to the psoralen ligand. Specifically, 5-methoxypsoralen S.2a is demethylated to give the 5-hydroxypsoralen S.2b. The latter is reacted with 6-bromohexanol in the presence of K2CO3 (following a procedure adapted from that previously reported by Kurfürst et al., 1993) to give the psoralen derivative S.2c. Phosphitylation of S.2c affords S.2d (in another procedure adapted from the same reference). The phosphoramidite derivative S.2d is then condensed with the 5′-terminal hydroxyl of oligodeoxyribonucleotides bound to solid supports. This conjugation is completed by the deprotection, purification, and characterization of the psoralenyl-oligonucleotide conjugates S.2e. Materials 5-Methoxypsoralen (S.2a) Pyridine hydrochloride CaCl2 Dichloromethane (CH2Cl2), distilled from P2O5 and passed over activated, basic aluminum oxide Methanol (MeOH), distilled P2O5 N,N-Dimethylformamide, redistilled under vacuum over ninhydrin and stored over 4-Å molecular sieves 6-Bromo-1-hexanol Potassium carbonate, anhydrous
Synthesis of Modified Oligonucleotides and Conjugates
4.3.9 Current Protocols in Nucleic Acid Chemistry
Ethyl acetate Pyridine, anhydrous N,N-Diisopropylethylamine (DIEA), distilled from KOH Sodium sulfate, anhydrous 2-Cyanoethyl-N,N-diisopropylchlorophosphite Triethylamine (Et3N) 10% (w/v) aqueous sodium carbonate (NaHCO3) Cold saturated aqueous NaCl Acetonitrile (CH3CN), DNA synthesis grade, anhydrous and stored over 3 Å-molecular sieves Oligodeoxyribonucleotides bound to solid supports, such as controlled-pore glass (CPG; UNIT 3.1), synthesized by classic phosphoramidite chemistry (1-µmol scale) DNA synthesis reagents recommended by synthesizer manufacturer Concentrated (25%) aqueous ammonia 25-mL round-bottomed flask Reflux condenser Drying tube Nitrogen gas Oven Analytical thin-layer chromatography (TLC) setup and silica-gel plates: e.g., Merck 5554 Kieselgel 60F plates, including UV lamp for detection Filter funnel, porosity 4 Desiccator Argon Silica gel for column chromatography: Merck 9387 Kieselgel 60 or Merck 7734 Kieselgel 60 Chromatographic columns: (3 cm × 50 cm) and (1.5 cm × 40 cm) Prepare 5-(hydroxypsoralen), S.2b 1. Place 1 g (4.64 mmol) 5-(methoxypsoralen) S.2a and 4.27 g (37 mmol) pyridine hydrochloride in a 25-mL round-bottomed flask equipped with a reflux condenser and a calcium chloride drying tube, under a nitrogen atmosphere. 2. Verify that pyridine hydrochloride is at the bottom of the flask and heat the mixture 90 min at 180°C. Monitor the reaction by TLC using 90:10 (v/v) CH2Cl2/MeOH as eluent. The starting material S.2a (Rf = 0.9) will be transformed into S.2b (Rf = 0.50).
3. Allow the mixture to cool to room temperature. Solubilize the reaction mixture with 3 mL MeOH and slowly pour the solution in 50 mL cold water while stirring magnetically to precipitate S.2b. Filter the solid over a filter funnel (porosity 4) and dry it over P2O5 in a desiccator for 2 days. Analyze S.2b again by TLC. The Rf value of S.2b is 0.50 when 90:10 (v/v) CH2Cl2/MeOH is used as eluent, and 0.65 when 50:50 (v/v) CH2Cl2/ethyl acetate is the eluent. Expected yield: 95% (885 mg, 4.38 mmol). Characterize the product by appropriate analytical methods. Melting point: 280°C. Direct Attachment of Conjugate Groups to the 5′ Terminus of Oligodeoxyribonucleotides
H-NMR (DMSO-d6): δ 3.34 (br s, 1H), 6.23 (d, 1H, J = 9.7 Hz, C3 Pso), 7.14 (S, 1H, C8 Pso), 7.18 (d, 1H, J = 2.2 Hz, C4’ Pso), 7.89 (d, 1H, J = 2.3 Hz, C5’ Pso), 8.23 (d, 1H, J = 9.5 Hz, C4 Pso). 1
Mass analysis. I.S. polarity positive. Calculated for C11H6O4: M+H=203.0. Found 203.0.
4.3.10 Current Protocols in Nucleic Acid Chemistry
Prepare 5-(6-hydroxyhexyloxy)-psoralen, S.2c 4. Place a solution of 5-hydroxypsoralen S.2b (0.6 g, 2.97 mmol) in 10 mL anhydrous N,N-dimethylformamide in a 25-mL round-bottomed flask equipped with a reflux condenser and a calcium chloride drying tube. Add successively 1.16 mL 6-bromo1-hexanol (1.61 g, 8.91 mmol) and 0.61 g anhydrous potassium carbonate (4.45 mmol). 5. Heat the flask at 65°C in the dark under an argon atmosphere and stir for 4 hr. 6. Allow the mixture to cool to room temperature and remove insoluble salts by filtration, then concentrate the filtrate to dryness under reduced pressure. 7. Purify the residue (2.5 g) on a silica-gel column (3 cm × 50 cm, 70 g silica) using increasing concentrations of MeOH in CH 2Cl2 (1:100 to 3:97, v/v): 200 mL of 1% MeOH in CH2Cl2; 200 mL of 2% MeOH in CH2Cl2; 200 mL of 2.5% MeOH in CH2Cl2; and 200 mL of 3% MeOH in CH2Cl2. 8. Monitor the fractions by silica-gel TLC using 90:10 (v/v) CH2Cl2/MeOH or 50:50 (v/v) CH2Cl2/ethyl acetate as eluent. Collect the fractions containing the pure product and remove the solvent under reduced pressure. The Rf value of S.2c is 0.43 when 90:10 (v/v) CH2Cl2/MeOH is the eluent, and 0.45 when 50:50 (v/v) CH2Cl2/ethyl acetate is the eluent. Expected yield: 75% (672 mg, 2.22 mmol). Melting point: 100°C. H-NMR (CDCl3): @ 1.38-1.60 (m, 6H), 1.82-1.95 (m, 2Η, ΧΗ2), 3.60-3.71 (m, 2H, CH2OH), 4.45 (t, 2H, J = 6.4 Hz, -CH2OAr), 6.27 (d, 1H, J = 9.7 Hz, C3 Pso), 6.94 (d, 1H, J = 2.3 Hz, C4’ Pso), 7.26 (s, 1H, C8), 7.58 (d, 1H, J = 2.4 Hz, C5’ Pso), 8.15 (d, 1H, J = 9.5 Hz, C4 Pso) 1
Mass analysis. I.S. polarity positive. Calculated for C17H18O5: M+H=303.0. Found 303.0.
Prepare [5-(ω-hexyloxy)-psoralenyl]-(2-cyanoethyl)-N,N-diisopropylphosphoramidite S.2d 9. Place 0.30 g 5-(6-hydroxyhexyloxy)-psoralen (1 mmol; S.2c) in a 25-mL round-bottomed flask, dry it by coevaporation with anhydrous pyridine and leave under vacuum for at least 12 hr. See Basic Protocol 1, steps 7, 8 and 9, for procedure.
10. Under a nitrogen atmosphere and magnetic stirring at room temperature add using a syringe 6 mL dichloromethane, 0.35 mL DIEA (0.258 g, 2 mmol) and then dropwise 0.29 mL 2-cyanoethyl-N,N-diisopropylchlorophosphite (0.307 mg, 3 mmol). Leave 20 min at room temperature. Monitor the reaction by silica-gel TLC using 95:5 (v/v) ethyl acetate/Et3N as eluent. Pre-elute the TLC plate with eluent before spotting the samples. 11. Shake the solution twice with 2 mL of 10% (w/v) aqueous Na2CO3 and once with 2 mL of a cold saturated aqueous NaCl solution. Dry the organic layer over anhydrous sodium sulfate and concentrate it to dryness. 12. Purify the residue (500 mg) on a silica-gel column (1.5 cm × 40 cm, 20 g silica) using 95:5 (v/v) ethyl acetate/Et3N as eluent. Monitor the fractions by silica-gel TLC using 95:5 (v/v) ethyl acetate/Et3N as eluent. The Rf value of S.2d is 0.87.
Synthesis of Modified Oligonucleotides and Conjugates
4.3.11 Current Protocols in Nucleic Acid Chemistry
13. Pool the fractions containing the pure product. Remove the solvents under reduced pressure and coevaporate with anhydrous CH3CN. Expected yield: 65% (326 mg, 0.65 mmol) of a colorless oil. Purified psoralenyl phosphoramidite S.2d can be stored at −20°C for a few months and should be reconstituted in anhydrous acetonitrile just prior to use (add a few pearls of 3-Å molecular sieves and wait ∼30 min before using the solution). P-NMR (CDCl3): d 149.47 ppm
31
H-NMR (CDCl3): d 1.12-1.23 (m, 12H -[C(H)(CH3)2]2), 1.38-1.70 (m, 6H, (CH2)3), 1.82-1.85 (m, 2H, CH2), 2.60 (t, 2H, J = 6. Hz, CH2CN), 3.52-3.73 (m, 4H, CH2O, [C(H)]2), 3.74-3.90 (m, 2H, CH2OP), 4.45 (t, 2H, J = 6.33 Hz, -CH2OAr), 6.27 (d, 1H, J = 9.8 Hz, C3 Pso), 6.94 (d, 1H, J = 2 Hz, C4’ Pso), 7.13 (s, 1H, C8), 7.58 (d, 1H, J = 2.3 Hz, C5’ Pso), 8.15 (d, 1H, J = 9.8 Hz, C4 Pso). 1
Couple the psoralenylphosphoramidite S.2d to the 5′-end of oligodeoxyribonucleotides Automated coupling: 14a. After standard solid-phase DNA oligonucleotide synthesis (1-µmol scale) according to the classical phosphoramide method, perform an additional detritylation step. 15a. Place a 0.1 M solution of the psoralenyl phosphoramidite S.2d in anhydrous CH3CN on the synthesizer. Carry out coupling by recycling a mixture of 0.1 mL of the psoralenyl phosphoramidite, 0.5 mL of 0.4 M 1H-tetrazole in anhydrous CH3CN, and 0.1 mL anhydrous CH3CN for 5 min. Repeat. For synthesizers without a “recycle” program, perform a double coupling step. For optimal coupling yields changes in amidite concentration may be required, depending on the synthesizers used.
16a. Remove the excess phosphoramidite and tetrazole, and perform an aqueous iodine oxidation reaction according to the standard procedure. Manual coupling: 14b. When performing standard solid-phase oligodeoxyribonucleotide synthesis (1µmol scale) according to the classical phosphoramidite method, place the oligonucleotide bound to the support in a short (4-mL) vial stoppered with a septum. Push a needle through the septum and leave the vial in a desiccator containing P2O5 under vacuum overnight. The oligonucleotide must have a 5′-hydroxy group.
15b. Fill the desiccator with nitrogen before opening it. Using a syringe, add a mixture of phosphoramidite S.2d (0.15 mL of a 0.15 M solution in anhydrous CH3CN) and 1H-tetrazole (0.5 mL of a 0.5 M solution in anhydrous CH3CN) to the support-linked oligonucleotide. Leave the mixture at room temperature for 10 min, shaking gently by hand from time to time. 16b. Remove the excess reagents with a syringe and perform an oxidation reaction by adding 1 mL iodine solution that is normally used on the synthesizer. Remove the iodine solution after 1 min and wash the support extensively with CH3CN. Direct Attachment of Conjugate Groups to the 5′ Terminus of Oligodeoxyribonucleotides
Deprotect oligodeoxyribonucleotide-psoralen conjugates 17. Add 5 mL concentrated 25% aqueous ammonia to a vial containing the psoralen-oligodeoxyribonucleotide conjugate bound to the support and leave it 1 hr at 55°C.
4.3.12 Current Protocols in Nucleic Acid Chemistry
18. Discard the support and leave the ammonium hydroxide solution an additional 6 hr at 55°C. 19. Evaporate the ammoniacal solution to dryness. 20. Solubilize the crude material S.2e with 12 mL water and extract three times with 3 mL ethyl acetate each time. Reduce the volume of the solution to ∼3 to 4 mL under reduced pressure. 21. Purify and characterize the oligodeoxyribonucleotide-psoralen conjugates (see Basic Protocol 1; the conjugates can be detected at 260 and 320 nm). For example analysis performed on a Lichrospher 100 RP 18 (5 mm; 125 mm × 4 mm) column using a linear gradient of CH3CN (5% to 35% over 40 min) in 0.1 M aqueous triethylammonium acetate, pH 7, at a flow rate of 1 mL/min indicates a retention time of 30 min 16 sec for the psoralen-oligonucleotide conjugate Pso(CH2)6pd[T45-MeC2T5-MeCT5-MeC3T5-MeCT]. As for oligonucleotide-acridine conjugates, base composition can be verified by nuclease degradation (same protocol). In this case reversed-phase analysis of hydrolysates performed under the conditions described for the oligonucleotide-acridine conjugates indicates a retention time of 53 min 36 sec for S.2c. Quantification of oligonucleotide- psoralen conjugates can be made by using e ≈ 10 000 M−1 cm−1 at l = 320 nm (unpub. observ.). When the synthesis is performed at the micromole scale, ∼15 to 20 OD units of pure oligonucleotide-psoralen conjugates 15-mer with mixed bases are obtained. The absorption spectrum of the psoralen-oligonucleotide conjugate Pso(CH2)6pd[T45-MeC2T5-MeC T5-MeC3T5-MeCT] is represented in Figure 4.3.4 (right).
REAGENTS AND SOLUTIONS 1M triethylammonium acetate (TEAA) buffer, pH 7 (stock solution) In a fume hood, add successively to a 1 liter beaker (equipped with a stirring bar) distilled water (810 mL), triethylamine (134 mL) and glacial acetic acid (57 mL). Mix thoroughly and adjust to pH 7 by adding either a few drops of triethylamine or a few drops of glacial acetic acid. COMMENTARY Background Information Two strategies can be used to attach ligands to the 5′ end of oligodeoxyribonucleotides. This unit describes one of these strategies, which involves the direct addition of a ligand to the 5′ end of an oligodeoxynucleotide during its initial synthesis via the phosphoramidite approach. Although methods have been developed for derivatizing the 5′ terminus of oligodeoxyribonucleotides in homogenous solution (Thuong et al., 1984, 1987; Asseline and Thuong, 1984; Asseline et al., 1988), only conjugation reactions on solid supports are reported in this unit (Thuong and Chassignol 1988, Dupret et al., 1994; Asseline et al., 1996). This strategy is applicable when the ligand is available in sufficient amounts, and its solubility and chemical stability are compatible with the conditions required for the preparation of its phosphoramidite or H-phosphonate deriva-
tives. In addition, the ligand should be stable under the conditions required for the coupling reaction and to those conditions used for complete deprotection of the oligodeoxynucleotide chain and its release from the support. The second strategy, not covered here, involves the incorporation of appropriate functional groups into two unprotected reactants. The coupling of these functionalized reactants results in the formation of oligodeoxyribonucleotide conjugates. Various oligodeoxyribonucleotide conjugates can be prepared using this strategy from only one oligodeoxyribonucleotide synthesis, provided that the required amount for each conjugate is low. Future units will describe the coupling of a number of ligands (intercalators, photoreactive and cleavage reagents, and labels) to oligodeoxyribonucleotides post-synthetically. These ligands are typically 2-methoxy-6-choro-9-
Synthesis of Modified Oligonucleotides and Conjugates
4.3.13 Current Protocols in Nucleic Acid Chemistry
Direct Attachment of Conjugate Groups to the 5′ Terminus of Oligodeoxyribonucleotides
aminoacridine as an intercalator, psoralen as a photo-cross-linking reagent, and phenanthroline-Cu as a cleavage reagent. More recently, the authors have used thiazole orange as a reporter group. This strategy involves the incorporation of linkers carrying suitable functional groups into ligands and the addition of amino, phosphorothioate, phosphate, or sulfhydryl functions to the 5′ end of oligonucleotides, as well as coupling methods for linking both entities. Alternatively, many reporter groups bearing functional groups that will react with 5′-thiol, 5′-terminal phosphorothioate, and 5′-amino groups are available from commercial sources. Heterobifunctional reagents, which allow reaction with a thiol function and with the primary amino function of various ligands, are also available commercially. These reagents, listed in UNIT 4.2, are very useful when there is no need for conjugates to have a well-defined linker between the oligodeoxyribonucleotide and the ligand. The coupling methods developed in this unit and future units can be used to prepare oligodeoxyribonucleotide conjugates composed of natural β-deoxyribonucleosides (Thuong and Chassignol 1988, Dupret et al., 1994, Asseline et al., 1996), unnatural α-D-deoxyribonucleosides (Kurfürst et al., 1993), or 2′-Omethylribonucleosides (unpub. observ.). When the chain assembly is performed in the usual (3′→5′) orientation, the modifying steps (i.e., direct incorporation of the ligand or the masked 5′-terminal functional group) are carried out at the end of the chain assembly just prior to the deprotection step. Given that oligodeoxyribonucleotides can be purchased fully protected and bound to the support, 5′-end modification can be performed manually. Procedures reported in this unit may be used to prepare oligodeoxyribonucleotides covalently linked via their 5′ end to molecules such as intercalating agents, and other reactive compounds the properties of which are described in UNIT 4.2. In many examples, oligodeoxyribonucleotide conjugates properties, such as strong complex stabilization when the ligand is an intercalator (Asseline et al., 1984, 1996, Sun et al., 1989; Giovannangeli et al., 1996) or best cleavage or cross-linking efficiency when the ligand is a reactive group, are largely dependent on the geometry of the complex formed between the ligand and its target (Takasugi et al., 1991; Giovannangeli et al., 1992; Costes et al., 1993, Grigoriev et al., 1993). Using these procedures linkage parameters between the oli-
godeoxyribonucleotide and the ligand, such as the size and nature of the linker connecting the two entities, may be varied to prepare oligodeoxyribonucleotide-ligand conjugates exhibiting optimal properties for specific experimental needs.
Critical Parameters and Troubleshooting In the case of acridine-containing oligodeoxyribonucleotides, the use of ammonia for the deprotection step must be avoided since it induces cleavage of the C9-N bond between the acridine and the linker. Therefore, a 0.4 N sodium hydroxide solution in 50:50 (v/v) H2O/MeOH is recommended. It is also recommended that the oligonucleotide-acridine conjugate solutions be stored between pH 6 and 7. Storage at pH values between 7 and 9 leads to considerable degradation. Protection of flasks and columns from light during the different steps of these protocols is recommended because acridine and psoralen derivatives are light-sensitive. In any case, purification steps must be as short as possible. It is always possible that the conditions required for deprotection of the conjugates may be incompatible with the stability of some modified nucleoside incorporated into the sequence of interest. To avoid obtaining a low yield of oligonucleotide-ligand conjugates, it is possible to deprotect an aliquot of the conjugates to verify the coupling efficiency of the amidite derivatives. If the coupling efficiency is insufficient, it is possible to perform additional coupling of the ligand. At the micromole scale it is usual to obtain about 20 OD units of pure conjugates when working with mixed DNA sequences. In some cases, when working with oligonucleotides containing sequences, inducing self associated structure, the purification step can be particularly challenging. In such cases, it is advisable to purify the acridine-oligonucleotide conjugates on a Mono-Q column at pH 12. In the case of sequences containing stretches of dG, the better way to solve the problem is to partially replace the dGs with 7-deaza-2′-deoxyguanosines (Raynaud et al., 1996).
Anticipated Results The yields reported for the various steps can be different when the syntheses are performed at scales other than those described. Using the protocols provided here and starting with 300 mg of the acridine-linker derivative S.1b, 300 mg of the purified acridine phosphoramidite S.1c can be obtained. This can be used to
4.3.14 Current Protocols in Nucleic Acid Chemistry
synthesize acridine-oligodeoxyribonucleotide conjugates S.1d. Usually 15 to 20 OD units of purified conjugates S.1d can be expected when the synthesis is performed at the µmol scale. In the case of the psoralen derivatives, starting with 310 mg of the psoralen-linker derivative S.2c it is possible to obtain 325 mg of the purified psoralen phosphoramidite derivative S.2d. The latter is then used to prepare psoralenoligodeoxyribonucleotide conjugates S.2e. When DNA synthesis is performed at the µmol scale, the yields obtained for conjugates S.2e (15 to 20 OD units) are similar to those obtained for the preparation of the conjugates S.1d.
Time Considerations Providing all reagents and materials required for each step are available, most of the reactions are simple and rapid. Preparation of compounds S.1b and S.2b requires 3 days including the drying step, while only two days are necessary to obtain compound S.2c. The time required for the preparation of phosphoramidite derivatives S.1c or S.2d is one day provided that the ligand-linker derivatives have been dried the previous day and left under vacuum in a desiccator overnight. Oligonucleotides chain assemblies and the addition of the ligand to the 5′-end of the oligonucleotide can be performed the same day when the ligand is added directly with the synthesizer. Otherwise the manual coupling requires leaving the support bearing the oligonucleotide under vacuum overnight. The deprotection step for psoralen-oligodeoxyribonucleotide conjugates S.2e takes a few hours while the duration of the deprotection step for acridine-oligodeoxyribonucleotide conjugates S.1d can be extended to almost two days. The time required for the purification of the ligand-oligonucleotide conjugates S.1d and S.2e can vary depending on the sequences. When two successive purification steps are necessary the time can be from five to ten days including rounds of lyophilization (overnight) or desalting and additional analyses. Characterization of the purified conjugates S.1d and S.2e can be achieved in three or four days.
Literature Cited Asseline, U., Thuong, N.T., and Hélène, C. 1986. Oligothymidylates substitutés en position 3′ par un dérivè de l’acridine. Nucleosides Nucleotides 5:45-63.
Asseline, U. and Thuong, N.T. 1988. Oligothymidylates substitués par un dérive de l’acridine en position 5′, à la fois en position 5′ et 3′ ou sur un phosphate internucleotidique. Nucleosides Nucleotides 7:431-455. Asseline, U., Toulmé, F., Thuong, N. T., Delarue, M., Montenay-Garestier, T., and Hélène, C. 1984. Oligodeoxynucleotides covalently linked to intercalating dyes as base sequence–specific ligands. Influence of dye attachment site. EMBO J. 3:795-800. Asseline, U., Bonfils, E., Dupret, D., and Thuong, N.T. 1996. Synthesis and binding properties of oligonucleotides covalently linked to an acridine derivative: A new study of the influence of the dye attachment site. Bioconjugate Chemistry 7:369-379. Costes, B., Girodon, E., Ghanem, N., Chassignol, M., Thuong, N.T., Dupret, D., and Goossens, M. 1993. Psoralen-modified oligonucleotide primers improve detection of mutations by denaturing gradient gel electrophoresis and provide an alternative to GC-clamping. Hum. Mol. Genet. 2:393-397. Dupret, D., Gossens, M., Chassignol, M., and Thuong, N.T. 1994. European Patent No. 0596028 A1,1994,05,11. Giovannangeli, C., Thuong, N.T., and Hélène, C. 1992. Oligodeoxynucleotide-directed photo-induced cross-linking of HIV proviral DNA via triple-helix formation. Nucleic Acids Res. 20:4275-4281. Giovannangeli, C., Perrouault, L., Escudé, C., Thuong, N.T., and Hélène, C. 1996. Specific inhibition of in vitro transcription elongation by triplex-forming oligonucleotide-intercalator conjugates targeted to HIV proviral DNA. Biochemistry 35:10539-10548. Grigoriev, M., Praseuth, D., Guieysse, A.L., Robin, P., Thuong, N.T., Hélène, C., and Harel-Bellan, A. 1993. Inhibition of gene expression by triple helix–directed DNA cross-linking at specific sites. Proc. Natl. Acad. Sci. USA. 90:35013505. Kurfürst, R., Roig, V., Chassignol, M., Asseline, U., and Thuong, N.T. 1993. Oligo-α-Deoxyribonucleotides with a modified nucleic base and covalently linked to reactive agents. Tetrahedron 49:6975-6990. Raynaud, F., Asseline, U., Roig, V., and Thuong, N.T. 1996. Synthesis and characterization of O6modified deoxyguanosine-containing oligodeoxyribonucleotides for triplex helix formation. Tetrahedron 52:2047-2064. Sun, J.S., François, J.C., Montenay-Garestier, T., Saison-Behmoaras, T., Roig, V., Thuong, N.T., and Hélène, C. 1989. Sequence-specific intercalating agents. Intercalation at specific sequences on duplex DNA via major groove recognition by oligonucleotide-intercalator conjugates. Proc. Natl. Acad. Sci. USA. 86:9198-9202.
Synthesis of Modified Oligonucleotides and Conjugates
4.3.15 Current Protocols in Nucleic Acid Chemistry
Takasugi, M., Guendouz, A., Chassignol, M., Decout, J.L., Lhomme, J., Thuong, N.T., and Hélène, C. 1991. Sequence-specific photo-induced cross-linking of the two strands of doublehelical DNA by a psoralen covalently linked to a triple helix forming oligonucleotide. Proc. Natl. Acad. Sci. USA. 88:5602-5606. Thuong, N.T. and Chassignol, M. 1988. Solid phase synthesis of oligo-α- and oligo-β-deoxynucleotides covalently linked to an acridine. Tetrahedron Lett. 29:5905-5908. Thuong, N.T., Hélène, C., and Asseline, U. 1984. European Patent No. 84-400143-8. 1989. U.S. Patent No. 4-835-263. Thuong, N.T., Asseline, U., Roig, V., Takasugi, M., an d Hélène, C. 1 98 7. Oligo(α-deoxynucleotide)s covalently linked to intercalating agents: Differential binding to ribo- and deoxyribopolynucleotides and stability towards nuclease digestion. Proc. Natl. Acad. Sci. USA. 84:5129-5133.
Contributed by Ulysse Asseline and Nguyen T. Thuong Centre de Biophysique Moléculaire, CNRS Orléans, France The authors would like to express their appreciation to their collaborators M. Chassignol, V. Roig, and Y. Aubert for their contribution to the development of varied oligonucleotide sequences linked to acridine and psoralen derivatives. This work was supported by Rhône-Poulenc, the Agence Nationale de Recherches sur le SIDA, and bio-Mérieux.
Direct Attachment of Conjugate Groups to the 5′ Terminus of Oligodeoxyribonucleotides
4.3.16 Supplement 7
Current Protocols in Nucleic Acid Chemistry
Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides
UNIT 4.4
This unit describes the synthesis of chimeric oligonucleotides in which the small (2′-5′)oligoriboadenylate activator of RNase L is covalently joined through a butanediol phosphate linker to a deoxyribooligonucleotide (Fig. 4.4.1). The products are termed “2-5A-antisense” when the deoxyribonucleotide portion of the chimera is targeted against a specific RNA. The overall synthetic strategy (see Basic Protocol 1) is based on the phosphite-triester approach to DNA/RNA synthesis (UNITS 3.3 & 3.5). Appropriately protected 2-cyanoethylphosphoramidite derivatives of adenosine, the butanediol linker, and the usual four deoxyribonucleosides are used for chain elongation, and the solid-phase methodology is employed with protected deoxyribonucleosides covalently linked to controlled-pore glass (CPG) through a long-chain alkyl amine (lcaa) and a succinyl moiety. 5′-Phosphorylation of the chimera completes the basic synthesis. After deprotection, the chimera is purified (see Basic Protocol 2) and characterized by a number of independent methods (see Basic Protocols 2 to 4). Two Support Protocols describe the synthesis of the central building block for the 2-5A domain of 2-5A-antisense oligonucleotide (see Support Protocol 1) and preparation of the synthon for the linker between 2-5A and the antisense oligonucleotide (see Support Protocol 2). CAUTION: When working with radioactivity, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following the guidelines provided by the local radiation safety officer.
5'-phosphomonoester
O HO P O − O
O HO
Ade
O − O P O O
O
Ade
2-5A domain HO
O − O P O O
HO
O
Ade
O − O P O O
HO
Ade
O
O − O P O O
O O P O O
−
linker domain
O O P O − O
O
Gua
O − O P O O
antisense domain
Thy
O
O − O P O 18-20
O
O
Ade
OH
Figure 4.4.1
General structure of 2-5A-antisense oligonucleotides.
Contributed by Mark R. Player and Paul F. Torrence Current Protocols in Nucleic Acid Chemistry (2000) 4.4.1-4.4.23 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Modified Oligonucleotides and Conjugates
4.4.1 Supplement 1
BASIC PROTOCOL 1
AUTOMATED SYNTHESIS OF CHIMERIC 2-5A-ANTISENSE OLIGONUCLEOTIDES ACCORDING TO SOLID-PHASE PHOSPHORAMIDITE CHEMISTRY The standard 2-5A-antisense chimera has four regions (Fig. 4.4.1): (1) antisense DNA, (2) linker dimer, (3) 2-5A tetramer, and (4) 5′-terminal phosphate monoester. This procedure describes construction of the antisense domain, coupling of the butanediol linkers, addition of the (2′-5′)-oligoriboadenylate domain, and 5′-phosphorylation. Details are given for two different automated ABI DNA synthesizers. 5′-O-Solid supports from ABI are used and are listed below; 3′-O-Solid supports from Glen Research are also listed, and are used to synthesize oligonucleotides with terminal (3′-3′)-phosphodiester bonds. Materials Reagents for oligonucleotide synthesis: Acetonitrile (CH3CN; diluent; Cruachem) Tetrazole/acetonitrile (activator/coupling solution; PE Biosystems) Acetic anhydride/lutidine/tetrahydrofuran (capping solution; PE Biosystems) N-Methylimidazole/tetrahydrofuran (capping solution; PE Biosystems) Trichloroacetic acid/CH2Cl2 (detritylation solution; PE Biosystems) Iodine/H2O/pyridine/tetrahydrofuran (oxidizer; PE Biosystems) Tetraethylthiuram disulfide (TETD)/CH3CN (PE Biosystems; for sulfurization) 0.1 M phosphoramidite solutions (see recipe): Linker CE phosphoramidite: 4-O-(4,4′-dimethoxytrityl)oxybutyl-1[(2-cyanoethyl)-N,N-diisopropylphosphoramidite] (see Support Protocol 2) RNA CE phosphoramidite (2-5A): N6-benzoyl-5′-O-(4,4′-dimethoxytrityl)3′-O-(tert-butyldimethylsilyl)adenosine-2′-(N,N-diisopropyl-2cyanoethyl)phosphoramidite (see Support Protocol 1) Phosphorylation reagent: 2-[2-(4,4′-Dimethoxytrityl)ethylsulfonyl]ethyl(2-cyanoethyl)-N,N-diisopropylphosphoramidite 5′-O-Dimethoxytrityl-N6-benzoyl-2′-deoxyadenosine-3′(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (dABz) 5′-O-Dimethoxytrityl-N2-isobutyryl-2′-deoxyguanosine-3′-(2-cyanoethylN,N-diisopropyl)phosphoramidite (dGi-Bu) 5′-O-Dimethoxytrityl-N4-benzoyl-2′-deoxycytidine-3′-(2-cyanoethyl-N,Ndiisopropyl)phosphoramidite (dCBz) 5′-O-Dimethoxytrityl-2′-deoxythymidine-3′-(2-cyanoethyl-N,Ndiisopropyl)phosphoramidite (T) 3:1 (v/v) concentrated NH4OH/ethanol 1 M n-tetrabutylammonium fluoride (TBAF)/tetrahydrofuran (THF; Aldrich) 10 mM n-tetrabutylammonium phosphate (TBAP), pH 7.5, in H2O
Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides
Automated DNA synthesizer (ABI Biotechnologies model 391 or 392) with 5′ or 3′ solid supports: 5′-O-Dimethoxytrityl-N6-benzoyl-2′-deoxyadenosine-3′-lcaa-CPG (1 µmol; dABz-lcaa-CPG; ABI) 5′-O-Dimethoxytrityl-N4-benzoyl-2′-deoxycytidine-3′-lcaa-CPG (1 µmol; dCBz-lcaa-CPG; ABI) 5′-O-Dimethoxytrityl-N2-isobutyryl-2′-deoxyguanosine-3′-lcaa-CPG (1 µmol; dGi-Bu-lcaa-CPG; ABI) 5′-O-Dimethoxytritylthymidine-3′-lcaa-CPG (1 µmol; T-lcaa-CPG; ABI) 3′-O-Dimethoxytrityl-N6-benzoyl-2′-deoxyadenosine-5′-lcaa-CPG (1 µmol; Glen Research)
4.4.2 Supplement 1
Current Protocols in Nucleic Acid Chemistry
3′-O-Dimethoxytrityl-N4-benzoyl-2′-deoxycytidine-5′-lcaa-CPG (1 µmol; Glen Research) 3′-O-Dimethoxytritylthymidine-5′-lcaa-CPG (1 µmol; Glen Research) 3′-O-Dimethoxytrityl-N2-isobutyryl-2′-deoxyguanosine-5′-lcaa-CPG (1 µmol; Glen Research). Water bath at 55°C Speedvac evaporator NOTE: The above-mentioned 5′ solid supports are used in the steps below. The 3′ solid supports are used in place of the 5′ supports when synthesizing oligonucleotides with terminal (3′-3′)-phosphodiester bonds. Synthesize 2-5A-antisense chimera For ABI 391 DNA synthesizer: 1a. Perform automated synthesis of 2-5A-antisense oligonucleotide on an ABI 391 DNA synthesizer according to manufacturer’s instructions. Use manufacturer’s 1 µmol scale synthetic cycles for the antisense region and modify step 13 of the manufacturer’s protocol for the remaining regions (see Table 4.4.1). 2a. Synthesize remaining regions of the chimera by sequentially adding 0.1 M solutions of linker CE phosphoramidite, RNA CE phosphoramidite (2-5A), and phosphorylation reagent. Use monomer CE phosphoramidites that are <1 week old. To change reagents, sequentially place on bottle position no. 5 as follows: a. Remove the previous reagent and replace with 5 mL CH3CN. b. Remove the column feed line and dry the column by turning valve no. 14 ON for 3 min and then OFF, then turning function no. 105 ON (4 mL CH3CN to column with the feed line unhooked) and then OFF. Replace column feed line. c. Change bottle no. 5 to the next region’s CE phosphoramidite. d. Repeat for synthesis of each chimera domain. For ABI 392 DNA synthesizer: 1b. Perform automated synthesis of 2-5A-antisense oligonucleotide on an ABI 392 DNA synthesizer according to manufacturer’s instructions. 2b. Synthesize remaining regions of the chimera by adding linker CE phosphoramidite, RNA CE phosphoramidite (2-5A), and phosphorylation reagent using monomer
Table 4.4.1 Synthesis Conditions for Different Domains of the 2-5A-Antisense Chimera
Chimera region DNA antisense Linker 2-5A Phosphate
Sequence no. and monomer sequence
Cycle no. and coupling time
Monomer CE phosphoramidite
Trityl mode
Sequence 2 5′-(Nx)-3′ Sequence 4 5′-(LL)(Nx)-3′ Sequence 3 5′-(AAAA)(LLNx)-3′ Sequence 1 5′-(p)(AAAALLNx)
Cycle 2 15 sec Cycle 4 300 sec Cycle 3 600 sec Cycle 1 60 sec
0.1 M DNA CE phosphoramidite 0.1 M linker CE phosphoramidite 0.1 M 2′,5′-ABz or PAC RNA CE phosphoramidite 0.2 M phosphorylation reagent
ON ON ON OFF
Synthesis of Modified Oligonucleotides and Conjugates
4.4.3 Current Protocols in Nucleic Acid Chemistry
Supplement 1
positions 5, 6, and 7, respectively. Use monomer CE phosphoramidites that are <1 week old. On the ABI 392, multiple cycle chimera sequences are created and saved on the computer’s hard disk. The linker CE phosphoramidite, 2-5A CE phosphoramidite, and phosphorylation reagent are kept on monomer positions 5, 6, and 7, and need not be removed until they expire. This minimizes exposure to air. Average stepwise yield (ASWY) and overall yield are available for each synthesis. Several cycles were created with the appropriate wait times by modification of the standard 1 mmol CE cycle (1mmol linker, 1 mmol phosphorylation). These changes are summarized in Table 4.4.2.
Cleave 2-5A-antisense oligonucleotide from CPG and deprotect it 3. Add 3 mL of 3:1 (v/v) concentrated NH4OH/ethanol to the CPG support and incubate 2 hr at room temperature. 4. Transfer to 55°C and incubate 8 to12 hr. Alternatively, in the case of all phenoxyacetyl (PAC)-protected bases, incubate at room temperature for 1 hr, followed by 55°C for 1 hr.
5. Transfer solution to a test tube, cool to 0°C, and evaporate solvent in a Speedvac evaporator. 6. Remove 3′-O-TBDMS group from the 2-5A region of the chimera by adding 1 mL of 1 M TBAF/THF and incubating 12 to 18 hr at room temperature. The chimera will normally remain in solution at the end of the incubation, and the solution will be homogenous. If it is not, briefly sonicate to achieve solution and incubate for another 4 hr.
7. Add 2 mL of 10 mM TBAP, pH 7.5, and remove THF in the Speedvac evaporator. 8. Dissolve crude chimera in distilled, deionized water to a total volume of 6 mL, and purify by HPLC (see Basic Protocol 2). If purification cannot be performed immediately, the chimera can be stored at –70°C in the dry state for up to 2 weeks.
Table 4.4.2 Automated Synthesizer Coupling Times and Conditions
Chimera region
Cycle name and coupling time
5′-(Nx)-3′
1 µmol CE; 15 sec
Monomer CE phosphoramidite
0.1 M DNA CE phosphoramidite 1 µmol linker; 300 sec 0.1 M linker CE 5′-(LL)(Nx) -3′ phosphoramidite 2-5A 5′-(AAAA)(LLNx)-3′ 1 µmol RNA; 600 sec 0.1 M 2′,5′-ABz RNA CE phosphoramidite Phosphate 5′-(P)(AAAALLNx)-3′ 1 µmol phosphorylation; 0.2 M 60 sec phosphorylation reagent Sulfurization Any linkage 1 µmol phosphothioate; TETD/CH3CN bottle 25 sec, 5 sec delivery no. 10 DNA antisense Linker
Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides
Monomer sequence
Trityl mode OFF
-
4.4.4 Supplement 1
Current Protocols in Nucleic Acid Chemistry
PREPARATION OF KEY INTERMEDIATE FOR SYNTHESIS OF THE (2′,5′)-OLIGOADENYLATE (2-5A) DOMAIN
SUPPORT PROTOCOL 1
This protocol describes the procedures used to generate the appropriately protected phosphoramidite, 5′-O-(4,4′-dimethoxytrityl)-3′-O-(tert-butyldimethylsilyl)-N6-benzoyladenosine-2′-(N,N-diisopropyl-2-cyanoethyl)phosphoramidite for adding adenosine through the formation of a 2′-5′ linkage to preformed antisense and linker domains. This phosphoramidite derivative may also be used to prepare unligated (2′-5′)-oligoadenylates using solid-phase synthesis. Materials Pyridine (Aldrich) dried over molecular sieves Adenosine (Aldrich) dried overnight in vacuo over P2O5 (Aldrich) 4-Dimethylaminopyridine (DMAP; Aldrich) Triethylamine (Aldrich) 4,4′-Dimethoxytrityl chloride (DMTr⋅Cl; Aldrich) Chloroform (CHCl3; Aldrich) Methanol (Aldrich) Trimethylsilyl chloride (Aldrich) Benzoyl chloride (Aldrich) Concentrated ammonium hydroxide (Aldrich) Diethyl ether (Aldrich) MgSO4 (anhydrous; Aldrich) Kieselgel 60 silica gel (220 to 440 mesh; Fluka) Ethyl acetate (Aldrich) Toluene (Aldrich) Imidazole (Aldrich) tert-Butyldimethylsilyl chloride (TBDMS⋅Cl; Aldrich) N,N-Dimethylformamide (DMF; anhydrous; Aldrich) Cyclohexane (Aldrich) 5% (w/v) aqueous Na2CO3 Methylene chloride (CH2Cl2; Aldrich) 1H-Tetrazole (Aldrich) P2O5 (Aldrich) Dry nitrogen 2-Cyanoethyl-(N,N,N′N′-tetraisopropyl)phosphoramidite (Aldrich) Benzene 250-mL flask with ground-glass stopper Rotary evaporator with vacuum pump 2 × 15-cm, 5 × 15-cm, and 5 × 20-cm chromatography columns 25-mL two-arm reaction flask fitted with a rubber septum 5-mL hypodermic syringe Additional reagents and equipment for thin-layer chromatography (TLC) and fast silica gel column (flash) chromatography Synthesize N6-benzoyl-5′-O-DMTr-adenosine 1. Mix the following in a 250-mL flask that can be sealed to the atmosphere: 100 mL dry pyridine 2.67 g (10 mmol) dry adenosine
Synthesis of Modified Oligonucleotides and Conjugates
4.4.5 Current Protocols in Nucleic Acid Chemistry
Supplement 1
61 mg (0.5 mmol) DMAP 1.9 mL (14 mmol) triethylamine 4.1 g (10 mmol) DMTr⋅Cl. Introduce a magnetic stirring bar, seal the flask with a ground-glass stopper, and stir 2 to 3 hr at room temperature. 2. Monitor the completeness of the reaction by analyzing an aliquot by silica gel TLC using 95:5 (v/v) CHCl3/CH3OH as the eluent. For comparison, run DMAP, adenosine, and DMTr⋅Cl as TLC standards. Use care to completely evaporate the pyridine from the TLC plate before development. 3. When the tritylation reaction is complete, place in an ice/water bath to maintain at 0°C, and slowly add 7.7 mL (60 mmol) trimethylsilyl chloride. Incubate 15 min. 4. Allow mixture to warm to room temperature, add 5.8 mL (50 mmol) benzoyl chloride, and stir ~2 hr at room temperature. 5. Cool mixture in the ice/water bath, add 20 mL cold water, and allow to stand for 5 min. 6. Add 20 mL concentrated ammonium hydroxide (final 2 M NH4OH), allow to warm to room temperature, and incubate 30 min at room temperature. 7. Concentrate to ~20 mL in a rotary evaporator in vacuo using a vacuum pump. 8. Add 700 mL diethyl ether, wash three times with 100 mL water, and dry the ether layer over 50 g anhydrous MgSO4. 9. Filter off MgSO4 and evaporate filtrate in a rotary evaporator. 10. Purify product by fast silica gel column (flash) chromatography using a 5 × 20-cm Kieselgel 60 (220 to 440 mesh) column and 2% methanol/0.2% pyridine/ethyl acetate as an eluent. Collect 5-mL fractions. 11. Analyze fractions by TLC as above, combine fractions that contain product (Rf = 0.6), and remove solvent in a rotary evaporator. 12. Remove last traces of residual pyridine by several additions and evaporations of toluene in 50-mL portions. Store, if necessary, dry at 4°C with a desiccant (stable at least 3 months). This is a critical step to obtain a solid product. The product should be obtained as a pale yellow solid foam (yield 50% to 70%): TLC Rf = 0.55 on silica gel (95:5 ethyl acetate/methanol); 1H NMR: (CDCl3) δ [ppm]: 3.34 (dd, 1H, H-5′ or 5′′), 3.49 (dd, 1H, H-5′ or 5′′), 4.39 (d, 1H, H-4′), 4.50 (dd, 1H, H-3′), 4.90 (t, 1H, H-2′), 6.13 (d, 1H, H-1′), 6.74-7.60 (m, aromatic protons), 8.02 and 8.04 (s, 1H, H-2 or H-8).
Synthesize N6-benzoyl-5′-O-DMTr-2′-O-TBDMS-adenosine and N6-benzoyl-5′-ODMTr-3′-O-TBDMS-adenosine 13. Dissolve the following and maintain at room temperature for 2.5 hr: 1.75 g (2.6 mmol) N6-benzoyl-5′-O-DMTr-adenosine Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides
0.544 g (8 mmol) imidazole 0.60 g (5 mmol) TBDMS⋅Cl 10 mL anhydrous DMF.
4.4.6 Supplement 1
Current Protocols in Nucleic Acid Chemistry
14. Verify that all N6-benzoyl-5′-O-DMTr-adenosine has reacted by running an aliquot on a silica gel TLC plate using 95:5 (v/v) ethyl acetate/methanol or 1:1 (v/v) ethyl acetate/cyclohexane as the eluent. 15. Add 1.5 mL of 5% aqueous Na2CO3 at 0°C to stop the reaction. Stir 15 min at <10°C. 16. Evaporate DMF in a rotary evaporator in vacuo, dissolve the resulting residue in 50 mL CH2Cl2, and wash three times with 50 mL water. 17. Dry CH2Cl2 layer over anhydrous MgSO4 and then evaporate solvent using a rotary evaporator. 18. Dissolve residue in 10 mL of 1:1 (v/v) ethyl acetate/cyclohexane containing 0.2% pyridine and apply to a 5 × 15-cm Kieselgel 60 silica gel column. Elute with 49:49:2 (v/v/v) ethyl acetate/cyclohexane/pyridine. Collect 5-mL fractions. 19. Monitor elution of reaction products using silica gel TLC with 1:1:0.004 (v/v/v) ethyl acetate/cyclohexane/pyridine. The first product to elute is the 2′-O-silylated isomer: N6-benzoyl-5′-O-DMTr-2′-OTBDMS-adenosine (Rf = 0.47). After complete elution of the 2′-O-silylated isomer, the concentration of ethyl acetate must be increased to elute the 3′-O-silylated isomer.
20. Switch eluent to ethyl acetate/cyclohexane/pyridine 66:32:2 (v/v/v) and continue TLC monitoring for elution of the 3′-O-silylated isomer. N6-Benzoyl-5′-O-DMTr-3′-O-TBDMS-adenosine has an Rf of 0.24.
21. Pool appropriate fractions containing each of the respective products and remove the solvent by rotary evaporation in vacuo. Subsequently add and evaporate 50-mL portions of toluene to remove traces of pyridine as judged by smell. Store, if necessary, dry at 4°C with a desiccant (stable at least 3 months). The products are obtained as colorless foams (typical yields: 480 mg 2′-O-silylated isomer; 660 mg 3′-O-silylated isomer). By silica gel TLC using 1:1:0.004 ethyl acetate/cyclohexane/pyridine, the 2′-isomer has a Rf = 0.47 and the 3′-isomer has a Rf = 0.24. The product used for further synthesis of 2-5A and 2-5A-antisense oligonucleotide is the 3′-O-silylated isomer: 1H NMR: (CDCl3), δ (ppm): 0.02 and 0.10 (ds, 6H, CH3Si), 0.98 (s, 9H, t-butyl), 3.27 and 3.54 (dd, 1H, H-5′ or H-5′′), 3.77 (s,3H, CH3O), 4.21 (m, 1H, H-4′), 4.60 (t, 1H, H-3′), 4.80 (t,1H, H-2′), 6.10 (d, 1H, H-1′), 8.03 and 8.05 (s, 1H, H-2 or H-8), 6.78-7.60, 8.28, 8.77 (m, aromatic protons). It is critical that complete separation of the silylated isomers be accomplished at this stage. Separation will be impossible after generation of the phosphoramidite. This is the most common cause of difficulty in this procedure.
Synthesize N6-benzoyl-5′-O-DMTr-3′-O-TBDMS-adenosine-2′-(N,N-diisopropyl-2cyanoethyl)phosphoramidite 22. Combine 48 mg (0.68 mmol) of 1H-tetrazole and 540 mg (0.68 mmol) of N6-benzoyl-5′-O-DMTr-3′-O-TBDMS-adenosine in a 25-mL two-arm reaction flask fitted with a rubber septum and a hypodermic needle. For this reaction, additional care must be taken to ensure anhydrous reaction conditions. The hypodermic needle is used for evacuation of air and admission of dry nitrogen and reagents.
23. Place the flask, fittings, and contents in a vacuum desiccator and dry overnight in vacuo in the presence of P2O5. 24. Admit dry nitrogen to the desiccator and thus to the reaction flask. Immediately remove the hypodermic needle from the rubber septum.
Synthesis of Modified Oligonucleotides and Conjugates
4.4.7 Current Protocols in Nucleic Acid Chemistry
Supplement 1
25. Add a solution of 5 mL CH2Cl2 and 206 mg (0.68 mmol) of 2-cyanoethyl-(N,N,N′,N′tetraisopropyl)phosphoramidite through the rubber septum using a 5-mL hypodermic syringe and needle. 26. Incubate 4 hr at room temperature and then overnight at 4°C. 27. Evaporate reaction mixture in vacuo on a rotary evaporator. 28. Add 2 mL of 6:3:1 (v/v/v) benzene/cyclohexane/triethylamine to the residue and apply to a 2 × 15-cm silica gel (Kieselgel 60) column. Use the same solvent for product elution. Collect 5-mL fractions. 29. Monitor fractions by silica gel TLC using 6:3:1 (v/v/v) benzene/cyclohexane/triethylamine as eluent. Two major products are generated in this reaction, namely, the two P-chiral diastereomers. These two stereoisomers have Rf values of 0.61 and 0.53 by silica gel TLC with this solvent.
30. Combine product-containing fractions, concentrate to dryness in vacuo, and dry by addition and evaporation of 50-mL portions of toluene. The product can be stored at 4°C. A partial separation of these two diastereomers can be achieved, but in practice this is not necessary. The product, consisting of a mixture of both P-chiral diastereoisomers, is obtained as a colorless foam (~500 mg); 31P NMR: (CDCl3, 1% C5D5N):δ (ppm) 150.73 and 150.38. SUPPORT PROTOCOL 2
PREPARATION OF A LINKER INTERMEDIATE THAT JOINS (2′-5′)-OLIGOADENYLATE (2-5A) TO ANTISENSE (3′-5′)-OLIGODEOXYRIBONUCLEOTIDES This protocol describes the synthesis of 4-O-(4,4′-dimethoxytrityl)oxybutyl-1-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite], which is the moiety used in the prototype 2-5A-antisense chimera to join the 2-5A domain to the antisense domain. Materials 1,4-Butanediol (Aldrich) Pyridine (anhydrous; Aldrich) Dry argon or nitrogen (Aldrich) 4,4′-Dimethoxytrityl chloride (DMTr⋅Cl; Aldrich) Chloroform (Aldrich) Methanol (Aldrich) Ethyl acetate (Aldrich) MgSO4 (anhydrous; Aldrich) Kieselgel 60 silica gel (Fluka) Methylene chloride (CH2Cl2; Aldrich) Ethyldiisopropylamine (Aldrich) 2-Cyanoethyl-N,N-diisopropylphosphoramidic chloride (Aldrich) Benzene (Aldrich) Petroleum ether (Aldrich) Triethylamine (Aldrich)
Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides
250-ml stoppered flask Rotary evaporator attached to a vacuum pump Hand-held UV lamp
4.4.8 Supplement 1
Current Protocols in Nucleic Acid Chemistry
Additional reagents and equipment for thin-layer chromatography (TLC) and fast silica gel column (flash) chromatography Synthesize 4-O-(4,4′-dimethoxytrityl)oxybutan-1-ol 1. Dissolve 9.0 g (100 mmol) of 1,4-butanediol in 50 mL anhydrous pyridine in a 250-ml stoppered flask. 2. Remove pyridine by evaporation at <40°C in a rotary evaporator attached to a vacuum pump. 3. Through the rotary evaporator stopcock, admit dry argon or nitrogen to the flask containing the butanediol, add an additional 50 mL dry pyridine, and repeat the evaporation procedure. 4. Repeat step 3 once more. Be certain to admit the dry nitrogen or argon after the final pyridine evaporation. A total of three evaporations will dry the butanediol sufficiently for the following tritylation reaction.
5. Dissolve dried 1,4-butanediol in 50 mL anhydrous pyridine. 6. Add 3.39 g (10 mmol) DMTr⋅Cl and allow the homogenous mixture to react 2 hr at room temperature in the stoppered flask. 7. Verify complete formation of the product by TLC using 99:1 (v/v) chloroform/methanol as developing solvent. 8. Pour the entire mixture onto 100 g ice in a beaker. Add a magnetic stirring bar and stir the mixure until the ice is completely melted. 9. Add 100 mL ethyl acetate and shake to extract the organic product. Separate the organic (top) and aqueous (bottom) layers. 10. Reextract the aqueous layer with an additional 50 mL ethyl acetate. Separate the organic layer again, and add the ethyl acetate layer to the one obtained from the previous extraction. 11. Add 10 g anhydrous MgSO4 to the combined organic layers to dry them. Filter off MgSO4 and evaporate all but ~10 mL ethyl acetate solution in a rotary evaporator at <40°C. 12. Add the concentrated ethyl acetate solution to the top of a column containing 250 g silica gel (Kieselgel 60). Elute with 99:1 (v/v) methylene chloride/methanol, collecting 10-mL fractions. 13. Check each fraction for product by spotting aliquots on a silica gel TLC plate and looking for the presence of UV-absorbing material with a hand-held UV lamp. If UV absorbance is detected, develop the plate using 99:1 (v/v) chloroform/methanol. Approximately 2.1 g (54% overall yield) of 4-O-(4,4′-dimethoxytrityl)oxybutan-1-ol can be obtained: 1H NMR (CDCl3, 1% deuteriopyridine):δ (ppm) 1.68(m, 4H, CH2); 3.10 (t, J = 5.7 Hz, 2H, CH2O); 3.62 (t, J = 5.8 Hz, 2H, CH2OH); 3.76 (s, 6H, CH3O); 6.79-7.46 (m, aromatic protons).
Synthesize 4-O-(4,4′-dimethoxytrityl)oxybutyl-1-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite] 14. Prepare a solution of the following reagents under anhydrous conditions: 390 mg (1 mmol) 4-O-(4,4′-dimethoxytrityl)oxybutan-1-ol
Synthesis of Modified Oligonucleotides and Conjugates
4.4.9 Current Protocols in Nucleic Acid Chemistry
Supplement 1
510 mg (4 mmol) ethyldiisopropylamine 3 mL dry methylene chloride. Cool in an ice bath. 15. Slowly add 237 mg (1 mmol) of 2-cyanoethyl-N,N-diisopropylphosphoramidic chloride under anhydrous conditions. Allow to warm to room temperature and to react for 1 hr. 16. Evaporate solvent using a rotary evaporator at <40°C. 17. Dissolve residue in 5 mL benzene and add mixture to the top of a 1.8 × 14-cm silica gel column. Elute the product with 6:3:1 (v/v/v) benzene/petroleum ether/triethylamine. 18. Check fractions for product by silica gel TLC using 6:3:1 (v/v/v) benzene/petroleum ether/triethylamine as the eluent. 19. Evaporate solvent from product-containing fractions. Yield: 580 mg (98%) 4-O-(4,4′-dimethoxytrityl)oxybutyl-1-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite]: 1H NMR (CDCl3, 1% deuteriopyridine):δ (ppm) 1.17 (t, J = 7.0 Hz, 12 H, CH3C); 1.70 (m, 4H, CH2C); 2.60 (t, J = 6.5 Hz, 2 H, CH2CN), 3.08 (t, J = 5.7 Hz, 2H, CH2O); 3.80 (m, 2H, CH); 6.80-7.49 (m, aromatic protons); 31P NMR (CDCl3, 1% deuteriopyridine):δ (ppm) 147.6. This phosphoramidite, like all phosphoramidites, is best stored in the presence of a desiccant at –20°C. BASIC PROTOCOL 2
PURIFICATION AND CHROMATOGRAPHIC CHARACTERIZATION OF CHIMERIC 2-5A ANTISENSE OLIGONUCLEOTIDES After deprotection, the chimera can be purified using either reversed-phase ion-pair chromatography or anion-exchange chromatography. The polystyrene reversed-phase ion-pair (PRP-1) column has a long lifetime, can tolerate extremes of pH and fluoride ions, and gives good recoveries of the 2-5A-antisense chimera. Desalting prior to ion-pair HPLC is not required, but is performed after. A typical chromatogram of a representative crude chimera is shown in Figure 4.4.2. The Nucleogen DEAE anion-exchange column has a shorter lifetime than the PRP-1 column, and cannot tolerate extremes of pH and fluoride ions. Preparative purification requires prior desalting. In addition, recoveries of chimera are poorer than those accomplished with the PRP-1 procedure. After purification by ion-pair or anion-exchange chromatography, the chimera is converted to a sodium salt by cation exchange, dialyzed, and filter sterilized. The purity of the chimera is estimated by capillary gel electrophoresis and ion-exchange HPLC.
Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides
Materials Crude 2-5A-antisense chimera (see Basic Protocol 1) Solvent A: 10 mM n-tetrabutylammonium phosphate (TBAP), pH 7.5, in H2O (for ion-pair chromatography) Solvent B: 10 mM TBAP, pH 7.5, in 8:2 (v/v) CH3CN/H2O (for ion-pair chromatography) Methanol Solvent C: 20 mM potassium phosphate, pH 7.0, in 8:2 (v/v) CH3CN/H2O (for anion-exchange chromatography) Solvent D: 20 mM potassium phosphate, pH 7.0, in aqueous 1 M KCl (for anion-exchange chromatography) Dowex 50W (Na+ form; Bio-Rad) Tris/methanol running buffer: 75 mM Tris phosphate, pH 7.6, in 9:1 (v/v) H2O/methanol
4.4.10 Supplement 1
Current Protocols in Nucleic Acid Chemistry
Figure 4.4.2 Polystyrene reversed-phase ion-pair chromatography of the crude (post-deprotection) 2-5A-DNA chimera: 5′-pA(2′,5′-pA)3-2′-p[(CH2)4p]2d[TCT CCG CTT CTT CCT GCC AT]. A 33 × 7-mm PRP-1 column was used and elution was accomplished with a convex gradient of 5% to 80% solvent B in solvent A over 50 min, where solvent A was 10 mM n-tetrabutylammonium phosphate (TBAP), pH 7.5, and solvent B was 10 mM TBAP in 8:2 (v/v) methanol/water, pH 7.5. The flow rate was 1.5 mL/min. Detection was at 270 mm. The major peak at a retention time of ~39 min was collected.
Solvent E: 25 mM Tris⋅Cl, pH 7.0 (APPENDIX 2A) in 1:200 (v/v) CH3CN/H2O Solvent F: 25 mM Tris⋅Cl, 1 M ammonium chloride, pH 7.0, in 1:200 (v/v) CH3CN/H2O High-performance liquid chromatograph (HPLC) with 270-nm UV detection 300 × 7-mm polystyrene reversed-phase column (PRP-1 semi-prep column; 10 µm, 100 Å; Hamilton; for ion-pair chromatography) Speedvac evaporator SepPak C18 cartridge 125 × 4–mm Nucleogen DEAE 60-7 column (7 µm, 60 Å; Macherey-Nagel; for anion-exchange chromatography) PolyPrep column Millex-GV 0.22-µm filter unit (Millipore) SpectraPor dialysis chamber (Vt = 5 mL) with a 3500 MWCO membrane (Spectrum) Capillary electrophoresis instrument (ABI model 270A-HT) with MICRO-GEL100 gel-filled capillaries (50-µm i.d., 27-cm effective length) and UV detection at 260 nm 250 × 4-mm Dionex PA-100 column (Dionex) Additional reagents and equipment for determining OD260 (UNIT 10.3)
Synthesis of Modified Oligonucleotides and Conjugates
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Purify 2-5A-antisense chimera For reversed-phase ion-pair chromatography: 1a. Inject 1 mL crude 2-5A-antisense chimera into a 300 × 7–mm polystyrene reversedphase column. 2a. Elute purified chimera using a convex gradient from 5:95 (v/v) solvents B/A to 80:20 solvents B/A over 45 min, at a flow rate of 1.5 mL/min. Collect 1-mL fractions and use UV detection at 270 nm. 3a. Combine fractions according to the UV profile, and evaporate to dryness in a Speedvac evaporator. 4a. Wash a SepPak C18 cartridge sequentially with 10 mL methanol and 10 mL H2O to remove shipping buffer. 5a. Dissolve chimera in 2 to 4 mL H2O and load onto the cartridge. 6a. Wash cartridge sequentially with 15 mL H2O, 10 mL of 5:95 (v/v) methanol/H2O, and 10 mL of 1:9 (v/v) methanol/H2O. 7a. Elute chimera with 10 mL of 1:1 (v/v) methanol/H2O and concentrate the eluate as needed in a Speedvac evaporator. Proceed to step 8. For anion-exchange chromatography: 1b. Desalt crude 2-5A-antisense chimera as described in steps 4a to 7a. 2b. Inject 1 mL chimera in water onto a 125 × 4–mm Nucleogen DEAE 60-7 column. 3b. Elute purified chimera using a linear gradient from 1:99 (v/v) solvents D/C to 100% solvent D over 30 min, followed by isocratic elution for 15 min, all at a flow rate of 1.0 mL/min. Collect 1-mL fractions and use UV detection at 260 nm. 4b. Combine fractions as indicated by the UV profile, and evaporate to dryness in a Speedvac evaporator. Proceed to step 8. Convert chimera to its sodium salt by cation exchange 8. Dissolve tetrabutylammonium salt (step 7a) or potassium salt (step 4b) of the chimera in 2 mL water. 9. Add 1 mL Dowex 50W slurry (Na + form) and stir 3 hr at 4°C. 10. Remove resin by passing the suspension through an empty PolyPrep column. 11. Elute chimera from the resin with 17 mL H 2O. Be sure to monitor the recovery by UV, as the oligonucleotide is retarded by the Dowex. Dialyze and sterilize chimera 12. Sterilize solution through a Millex-GV 0.22-µm filter unit. 13. Dialyze against water overnight at 4°C, using a SpectraPor dialysis chamber (Vt = 5 mL) with a 3500 MWCO membrane. 14. Repeat step 12. Determine OD260 (UNIT 10.3) to quantitate DNA. Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides
These steps are necessary for removal of low-molecular-weight impurities and to prepare the chimera for biological evaluation.
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Figure 4.4.3 Determination of 2-5A-antisense chimera purity by capillary gel electrophoresis on an ABI 270A-HT instrument using MICRO-GEL100 gel-filled capillaries (50-µm i.d., 27-cm effective length) with a running buffer of 75 mM Tris phosphate, pH 7.6, in 9:1 (v/v) H2O/methanol. The voltage was –14 kV and the operation T was 30°C. Detection was at 260 nm. The typical chromatogram shown is for the chimera: 5′-pA(2′,5′-pA)3-2′-p[(CH2)4p]2d[TCT CCG CTT CTT CCT GCC AT].
Estimate chimera purity by capillary gel electrophoresis 15. Determine oligonucleotide chimera purity on an ABI 270A-HT capillary electrophoresis instrument using MICRO-GEL100 gel-filled capillaries and Tris/methanol running buffer with UV detection at 260 nm. Run at –14 kV (17 µA) and at an operation temperature of 30°C. Typically, a chimera electropherogram can be obtained using a sample concentration of 0.06 OD260 and an electrokinetic injection of 2 s at –5 kV. A typical electropherogram is shown in Figure 4.4.3.
Estimate chimera purity by ion-exchange HPLC 16. Carry out ion-exchange HPLC using a 250 × 4–mm Dionex PA-100 column. Use an injection volume of 50 µL and ~0.1 OD260 of oligonucleotide. Employ a linear gradient from 1:9 (v/v) solvents F/E to 9:1 solvents F/E over 25 min, followed by isocratic elution for 25 min, all at a flow rate of 1.0 mL/min with UV detection at 270 nm. The Dionex PA-100 column gives better peak shape than many other ion-exchange columns.
COMPOSITION ANALYSIS OF CHIMERIC 2-5A-ANTISENSE OLIGONUCLEOTIDES USING SNAKE VENOM PHOSPHODIESTERASE DIGESTION AND REVERSED-PHASE HPLC When 2-5A-antisense chimeric oligonucleotides are digested by snake venom phosphodiesterase (SVPD), they yield 5′-AMP from the 2-5A domain, 5′-pA2′-p(CH2)4p(CH2)4OH from the terminal AMP of 2-5A and the linker moiety, and deoxyribonucleotide 5′-monophosphates from the antisense domain. This analysis provides a cornucopia of valuable information regarding the 2-5A-antisense chimera. It provides a compositional analysis (by providing the ratio of 5′-AMP to any or all of the constituent 5′-deoxyribonucleotides) and
BASIC PROTOCOL 3
Synthesis of Modified Oligonucleotides and Conjugates
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also ascertains the completeness of 5′-phosphorylation (through the appearance of adenosine as a product of 5′-unphosphorylated chimera), which is vital for full activation of RNase L. Materials Purified 2-5A-antisense chimera (see Basic Protocol 2) Snake venom phosphodiesterase (SVPD, Crotallus adamanteus; Amersham Pharmacia Biotech) 1 M Tris⋅Cl, pH 8.0 (APPENDIX 2A) 1 M MgCl2 Solvent A: aqueous 100 mM ammonium phosphate, pH 5.5 Solvent B: 1:1 (v/v) methanol/H2O Microcon-10 concentrator (Amicon) High-performance liquid chromatograph (HPLC) with 250 × 4.6–mm Ultrasphere C18 ODS column (5 µm, 80 Å; Thomson Instrument) 1. Analyze the nucleotide composition of purified 2-5A-antisense chimera by digestion with SVPD. Mix the following and incubate 3 hr at 37°C: 0.2 OD260 units chimera 0.15 units SVPD 50 mM Tris⋅Cl, pH 8.0 0.5 mM MgCl2 water to 100 µL. 2. Wash the membrane of a Microcon-10 concentrator with 100 µL water and then with 100 µL solvent A to remove shipping buffer. Centrifuge each wash 5 min at 700 × g, 4°C (see manufacturer’s instructions). Each spin-rinse takes ~15 min.
3. Discard filtrate, apply 100 µL enzyme digestion mixture, and spin-rinse for 15 min. 4. Wash membrane three more times with 100-µL portions of solvent A to ensure that all nucleotides have passed through the membrane. 5. Analyze the centrifugate by reversed-phase HPLC using a 250 × 4.6–mm Ultrasphere C18 ODS column. Inject 20 to 30 µL. Separate the digestion products using the following solvent program at a flow rate of 0.5 mL/min: a. isocratic elution with 1:99 (v/v) solvents B/A for 20 min b. linear gradient elution from 1:99 solvents B/A to 45:55 solvents B/A over 30 min c. isocratic elution with 45:55 solvents B/A for 20 min. Typical retention times for various nucleotides are: 5′-dCMP (ε260 = 7,610), 9.7 min; 5′-TMP (ε260 = 8,158), 27.3 min; 5′-dGMP (ε260 = 9,969), 29.6 min; 5′-AMP (ε260 = 12,300), 31.7 min; 5′-dAMP (ε260 = 14,300), 41.2 min; 5′-pA2′-p(CH2)4p(CH2)4OH [AMP(pBu)2] (ε260 = 12,300), 39.5 min. A typical chromatogram of a digestion is shown in Figure 4.4.4.
Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides
6. Normalize the absorbance of each peak by dividing by the extinction coefficient (ε260), and thereby ascertain the relative ratios of each nucleotide.
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Figure 4.4.4 Digestion of a representative 2-5A-antisense chimera with snake venom phosphodiesterase and analysis by HPLC. A snake venom phosphodiesterase digest of the chimera 5′-pA(2′,5′-pA)3-2′p[(CH2)4p]2d[TCT CCG CTT CTT CCT GCC AT] was injected into a 250 × 4.6-mm Beckman Ultrasphere C18 ODS column. Elution was with 2% solvent B in 98% solvent A isocratically for 20 min, then a linear gradient of 2% to 50% solvent B in solvent A for 15 min, then 50% solvent B isocratically for 20 min. Solvent A was 100 mM ammonium phosphate, pH 5.5, and solvent B was 1:1 (v/v) methanol/H2O. The flow rate was 0.5 mL/min and detection was at 260 nm.
SEQUENCING CHIMERIC 2-5A-ANTISENSE OLIGONUCLEOTIDES ACCORDING TO A MODIFIED MAXAM-GILBERT PROCEDURE
BASIC PROTOCOL 4
Although the size and unusual chemical structure of 2-5A-antisense chimeras do not permit enzymatic sequencing, the Maxam-Gilbert procedure can be employed if certain modifications are followed. In this procedure, the chimera is 3′-end labeled using [α-32P]ddATP and terminal nucleotide transferase. Materials Purified 2-5A-antisense chimera (see Basic Protocol 2) 5× TNT buffer (see recipe) [α-32P]ddATP (3000 Ci/mmol, 10 mCi/mL; Amersham) 15 U/µL terminal deoxyribonucleotide transferase (TNT) from calf thymus (Life Technologies) 0.5 M EDTA, pH 8 (APPENDIX 2A) 1 mg/mL calf thymus DNA (Clontech) 0.5 M sodium phosphate buffer, pH 6.8 (see recipe) Scintillation fluid Diethyl pyrocarbonate (DEPC; Sigma) DEPC buffer (see recipe) Ethanol 0.3 and 3 M sodium acetate (Quality Biological), pH 6, in water 4 mg/mL yeast tRNA (Clontech) Dimethyl sulfate (DMS; Aldrich) DMS buffer (see recipe) DMS stop buffer (see recipe)
Synthesis of Modified Oligonucleotides and Conjugates
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A>C buffer (see recipe) 1 N acetic acid Hydrazine (HZ; Aldrich) HZ stop buffer (see recipe) 5 M NaCl (Quality Biological) in water 1 M piperidine (Aldrich) in water, freshly prepared Denaturing gel-loading buffer (see recipe) 10× TBE electrophoresis buffer (APPENDIX 2A) Lyophilizer ChromaSpin-10 gel column (Clontech) Whatman DE81 filters Vacuum manifold Siliconized tubes Heating blocks at 25°, 60°, and 90°C Additional reagents and equipment for polyacrylamide gel electrophoresis (PAGE; APPENDIX 3B) 3′-End label 2-5A-antisense chimera 1. Lyophilize 10 pmol of purified 2-5A-antisense chimera. 2. Add the following in order: 10 µL 5× TNT buffer 32 µL ddH2O 6 µL [α-32P]ddATP (20 pmol) 2 µL 15 U/µL TNT. 3. Incubate 60 min at 37°C. 4. Stop reaction by adding 4 µL of 0.5 M EDTA, pH 8, and 6 µL of 1 mg/mL calf thymus DNA. Vortex gently. 5. Set aside 1 µL for determination of specific activity (steps 8 to 11). 6. Apply the remainder of the reaction to a precentrifuged ChromaSpin-10 gel column and elute with water by centrifuging 6 min at 700 × g, at 4°C. 7. Lyophilize eluted solution and store at –20°C until use (up to 2 weeks). Determine specific activity of labeled chimera 8. Dilute 1 µL labeling reaction (from step 5) in 99 µL of 0.2 M EDTA, pH 8. 9. Place quadruplicate 3-µL spots of diluted sample on separate Whatman DE81 filters. Dry at 50°C. 10. Wash two filters on a vacuum manifold with 30 mL of 0.5 M sodium phosphate buffer, pH 6.8, and dry them at 50°C. Washing will remove unincorporated nucleotide. The two unwashed filters are used to determine the total cpm in the sample. Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides
11. Add scintillation fluid to each filter and count them in a liquid scintillation counter. Use the cpm from the filters to determine the following: % incorporation = cpm incorporated/total cpm
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Current Protocols in Nucleic Acid Chemistry
% 3′ ends labeled = % incorporation × 2 specific activity = % incorporation × total cpm added to reaction/µg chimera. Expected values are 10% to 15% incorporation, with 20% to 30% of 3′ ends labeled, and specific activity of ~108 cpm/mg chimera.
Perform sequencing reactions 12. Set up rA reaction to cleave primarily at riboadenosine residues: a. Place a siliconized tube containing 1⁄6 of the labeled chimera on ice. b. Add 150 µL DEPC buffer followed by 2 µL of 1:9 (v/v) DEPC/ethanol. c. Vortex and incubate 5 min at 90°C. d. Add 15 µL of 3 M sodium acetate, pH 6. e. Add 5 µL of 4 mg/mL yeast tRNA and 800 µL ethanol. f. Vortex and bring to –70°C. 13. Set up G reaction for primary cleavage at deoxyguanosine and minor cleavage at riboadenosine. a. Place a siliconized tube containing 1⁄6 of the labeled chimera on ice. b. Add 200 µL DMS buffer followed by 1 µL dimethyl sulfate. c. Vortex and incubate 20 min at 25°C. d. Add 50 µL DMS stop buffer and 800 µL ethanol. e. Vortex and bring to –70°C. 14. Set up A>C reaction for cleavage primarily at deoxyadenosine, with secondary cleavage at deoxycytidine and riboadenosine. a. Place a siliconized tube containing 1⁄6 of the labeled chimera on ice. b. Add 100 µL A>C buffer. c. Vortex and incubate 15 min at 90°C. d. Add 150 µL of 1 N acetic acid, 5 µL of 4 mg/mL yeast tRNA, and 800 µL ethanol. e. Vortex and bring to –70°C. 15. Set up A+G reaction for cleavage at riboadenosine, deoxyadenosine, and deoxyguanosine. a. Place a siliconized tube containing 1⁄6 of the labeled chimera on ice. b. Add 150 µL DEPC buffer followed by 2 µL of 1:9 (v/v) DEPC/ethanol. c. Vortex and incubate 5 min at 90°C. d. Add 15 µL of 3 M sodium acetate, pH 6. e. Add 5 µL of 4 mg/mL yeast tRNA and 800 µL ethanol. f. Vortex and bring to –70°C. 16. Set up T+C reaction for cleavage at deoxythymidine, deoxycytidine, and riboadenosine. a. Place a siliconized tube containing 1⁄6 of the labeled chimera on ice. b. Add 15 µL water followed by 30 µL hydrazine. c. Vortex and incubate 60 min at 25°C. d. Add 400 µL HZ stop buffer and 800 µL ethanol. e. Vortex and bring to –70°C. 17. Set up C reaction for strong cleavage at deoxycytidine and riboadenosine.
Synthesis of Modified Oligonucleotides and Conjugates
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a. b. c. d. e.
Place a siliconized tube containing 1⁄6 of the labeled chimera on ice. Add 15 µL of 5M NaCl followed by 30 µL hydrazine. Vortex and incubate 60 min at 25°C. Add 400 µL HZ stop buffer and 800 µL ethanol. Vortex and bring to –70°C.
Wash all samples 18. Centrifuge 15 min at 12,000 × g, 4°C. 19. Withdraw and discard supernatant, checking with a survey meter that very little labeled chimera is removed. 20. Redissolve chimera in 400 µL of 0.3 M sodium acetate, pH 6. 21. Add 800 µL ethanol, vortex, bring to –70°C, and centrifuge 15 min at 12,000 × g, 4°C. 22. Repeat steps 19 and 21 (do not add sodium acetate). 23. Remove most of the supernatant and lyophilize pellet for 15 min. Cleave modified bases with piperidine 24. Add 100 µL of 1 M piperidine to DNA samples and vortex. 25. Make sure lids are securely shut and incubate 20 min at 60°C (for rA reaction) or 30 min at 90°C (for all other reactions). 26. Cool on ice and lyophilize for 15 min. 27. Add 15 µL water, vortex, and lyophilize again. Repeat. Perform polyacrylamide gel electrophoresis (PAGE) 28. Add 100 µL denaturing gel-loading buffer. 29. Denature samples 60 min at 90°C and then cool on ice. 30. Pour and set up 20% polyacrylamide/8 M urea sequencing gel (see APPENDIX 3B) using 1× TBE electrophoresis buffer and pre-run for 2 hr at 130 W constant power. Gel will heat to 55°C.
31. Load samples and run at 130 W constant power until the bromphenol blue has migrated ~19 cm down from the wells. Remove one or both plates. 32. Mark the upper left corner of the gel with radioactive ink and autoradiograph. A typical result is shown in Figure 4.4.5. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
A>C buffer 0.480 mg sodium hydroxide (1.2 N) 20 µL 1 mM EDTA H2O to 10 mL Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides
Aniline solution, 1 M, pH 4.5 931 µL aniline (Aldrich) 8.869 mL H2O
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Figure 4.4.5. Maxam-Gilbert sequencing of the 2-5A-antisense chimera 5′-pA(2′,5′-pA)3-2′p[(CH2)4p]2d[TCT CCG CTT CTT CCT GCC AT]. Lane 1, rA; lane 2, G; lane 3, A>C; lane 4, A+G; lane 5, T+C; lane 6, C.
Adjust pH to 4.5 with acetic acid Store in the dark at –20°C Precipitate will clear upon addition of acetic acid.
ATP/dTTP stock solution 4.8 mg dTTP (final 1.0 mM) 0.55 mg ATP (final 0.1 mM) Adjust pH to 7.0 Add H2O to 10.0 mL Denaturing gel-loading buffer 8 mL formamide (Fluka; 80% v/v) 1 g sucrose (10% w/v) 40 µL 2 mM EDTA 0.2% (w/v) bromphenol blue (Sigma) Add H2O to 10 mL Diethyl pyrocarbonate (DEPC) buffer 166 µL 3 M sodium acetate, pH 6 (final 50 mM, pH 5) 20 µL 0.5 M EDTA (final 1 mM) H2O to 10 mL
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Dimethyl sulfate (DMS) buffer 80 mg sodium cacodylate (Aldrich), pH 8.0 (final 50 mM) 20 µL 1 mM EDTA H2O to 10 mL DMS stop buffer 829 µL 1.5 M sodium acetate, pH 7.0 71 µL 1 M 2-mercaptoethanol 25 µL 100 µg/mL yeast tRNA 75 µL H2O Prepare fresh before use Hydrazine (HZ) stop buffer 400 µL 1.5 M sodium acetate, pH 7 (final 0.3 M) 200 µL 1 mM EDTA (final 0.1 mM) 15 µL 25 µg/mL yeast tRNA (final 0.2 µg/mL) H2O to 2 mL Prepare fresh before use Phosphoramidite solutions, 0.1 M Dry phosphoramidites over anhydrous P2O5 for ≥33 hr. Dilute in anhydrous acetonitrile to make each 0.1 M solution (see Table 4.4.3). Use within a week. Sodium phosphate buffer, 0.5 M, pH 6.8 4.73 g NaH2PO4 2.24 g Na2HPO4 H2O to 100 mL Terminal deoxyribonucleotide transferase (TNT) buffer, 5× 500 mM potassium cacodylate 1 mM dithiothreitol 10 mM CoCl2 Adjust pH to 7.2 COMMENTARY Background Information The 2-5A system (Johnston and Torrence, 1984; Player and Torrence, 1998) has been the basis of a targeted mRNA destruction method that derives from the covalent linkage of a (3′-5′)-antisense oligodeoxyribonucleotide
Table 4.4.3
Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides
and a (2′-5′)-oligoadenylate activator of RNase L, the 2-5A-dependent RNase (Lesiak et al., 1993; Torrence et al., 1993). This composite nucleic acid (Fig. 4.4.1; Xiao et al., 1996) could, through the antisense domain, target the chimera to a particular mRNA sequence, which
Preparation of Automated Synthesizer Reagents
CE phosphoramidite
CH3CN (mL)
Molarity (M)
Coupling cycles
Sources
500 mg dABz 500 mg dGi-Bu 500 mg dCBz 500 mg T 0.25 g butanediol linker 500 mg 2-5A 0.25 g phosphorylation reagent
5.6 5.8 5.9 6.6 3.75 5.0 3.8
0.1 0.1 0.1 0.1 0.1 0.1 0.1
25 24 24 27 10 24 7
PE Biosystems PE Biosystems PE Biosystems PE Biosystems Support Protocol 2 Support Protocol 1 Glen Research
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would then be targeted for destruction by the 2-5A component via a localized activation of the latent 2-5A-dependent RNase. The prototype 2-5A-antisense chimera consisted of an antisense domain made up of oligo(dT)18 connected to 2-5A through a linker (Lesiak et al., 1993; Torrence et al., 1993). The 2-5A and antisense moieties were joined together through phosphodiester bonds and two 1,4-butanediol molecules linked to each other by a phosphodiester function. Linkage to the 2-5A tetramer was at the 2′-terminal hydroxyl, and linkage to the antisense oligonucleotide was at the 5′-terminal hydroxyl. The mode of linkage to the 2-5A component was through the 2′ terminus of the oligomer, since a free 5′-monophosphate was required for maximal 2-5Adependent endonuclease activity (Johnston and Torrence, 1984; Player and Torrence, 1998). Linker elements were used to join 2-5A to the antisense DNA sequence rather than directly joining the terminal 2-5A adenosine nucleotide to the first nucleotide of the antisense sequence. Such a strategy was used because of the possibility that RNase L, once bound to the 2-5A component of the chimera, might disturb hybridization to target RNA, or conversely that the double helix generated by the antisense oligonucleotide and the sense RNA might interfere with binding to RNase L. The 2-5A antisense approach has led to sequence-specific cleavage of a modified human immunodeficiency virus RNA (Torrence et al., 1993) and of mRNA encoding the dsRNA-dependent protein kinase (PKR) in cell-free systems (Maitra et al., 1995), as well as ablation in intact HeLa cells of PKR mRNA, PKR protein, and the biological function of PKR (Maran et al., 1994). In addition, respiratory syncytial virus (RSV) replication has been inhibited by specific 2-5A-antisense targeting of the RSV M2 mRNA (Cirino et al., 1997). The actual chemical synthesis of 2-5A-antisense oligonucleotides involves the solid-support phosphite-triester approach to DNA/RNA synthesis (e.g., Beaucage and Caruthers, 1981). Appropriately protected 2-cyanoethylphosphoramidite derivatives of the riboadenosine, butanediol linker, and the usual four deoxyribonucleosides are used for chain elongation. Sugar protection consists of the usual 4,4′-dimethoxytrityl for 5′ hydroxyls and tert-butyldimethylsilyl for 2′ or 3′ hydroxyls. For base protection, ABz, CBz, and Gi-Bu have been used by this laboratory for some time with good success; however, the authors have used commercially available APAC, Ci-Bu, and GPAC,
which allow faster ammoniacal workup and yield slightly cleaner crude product. 2-Cyanoethylphosphoramidites are the functionality of choice for phosphodiester bond generation. The following generic oligonucleotide structural types are described in this unit: I: 5′-pA(2′,5′-pA)3-2′-p[(CH2)4p]2dN(3′,5′pdN)n II: 5′-pA(2′,5′-pA)3-2′-p[(CH2)4p]2dN(3′,5′pdN)m(3′,3′-pdN) Analytical results are somewhat different when the snake venom phosphodiesterase protocol is applied to a type II (Li et al., 1997) 2-5A-antisense chimera. Specifically, a (3′-3′)dinucleotide 5′-monophosphate derived from the last two nucleotides of the chimera’s 3′ terminus is produced. Structural information to corroborate the identity of the dinucleotide can be obtained by comparing their HPLC on-line UV spectra with the calculated spectra of the 1:1 summation of constituent mononucleotides. Finally, the structure of the (3′-3′)dinucleotide fragment can be corroborated by comparing the HPLC chromatogram of the enzymatically digested product with that of a synthetic dinucleotide produced by a DNA synthesizer using a nucleoside bound to CPG through the 5′-hydroxyl group. The retention times of the (3′-3′)-dinucleotide products vary depending upon their composition and, in some cases, can interfere with other peaks in the HPLC analysis of snake venom phosphodiesterase digests. Under these conditions, accurate analysis of these key digestion products can be problematic. In order to obviate this difficulty, digestion with snake venom phosphodiesterase can be carried out with the addition of bacterial alkaline phosphatase. This results in the removal of all noninternucleotide phosphates, so that digestion products consist of nucleosides and the (3′-3′)dinucleotide. Thus, this modified digestion procedure can result in a shift in retention times (Rt) such that each individual product is well separated from the others. This additional digestion procedure is not always necessary; its necessity depends on the composition of the dinucleotide product. Many (3′-3′)-dinucleotides can be resolved very well from all nucleotides, and can be easily identified by their on-line UV spectra and subsequently integrated. In general, the syntheses follow the strategies developed in Lesiak et al. (1993), Xiao et al. (1996), and Li et al. (1997). Type I 2-5A-antisense chimeras were the first structural type
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synthesized (Lesiak et al., 1993; Torrence et al., 1993) and were used with success in cell-free systems and in intact cells (Torrence et al., 1993; Maran et al., 1994; Maitra et al., 1995). Type II 2-5A-antisense chimeras with 3′-terminally inverted phosphodiester bonds have proven to be of greater stability toward exonuclease degradation and have been exploited to block respiratory syncytial virus replication (Cirino et al., 1997).
Critical Parameters In the prototypical 2-5A antisense chimera, 2-5A and antisense DNA were joined through two linkers of 1,4-butanediol phosphate that arise from the key intermediate 4-O-(4,4′dimethoxytrityl)oxybutyl-1-[(2-cyanoethyl)N,N-diisopropylphosphoramidite]. The yield of this intermediate is improved significantly when using 2:10:1 (v/v) ethyl acetate/hexane/triethylamine as the flash purification chromatography solvent. Triethylamine in the purification may prevent decomposition of this acid-sensitive compound during exposure to silica gel. Key considerations in the characterization of 2-5A-antisense chimeras include: (1) that (2′,5′)-phosphodiester bonds in the RNA portion of the chimera are not isomerized to (3′,5′)phosphodiesters during or after synthesis; (2) that 1,4-butanediol phosphate linker connects 2-5A and antisense DNA in the anticipated position; and (3) that phosphorylation does, in fact, occur at the 5′ terminus in high yield. Approaches to address these questions may be found in Xiao et al. (1996).
Anticipated Results
Synthesis and Characterization of Chimeric 2-5A-DNA Oligonucleotides
Under the conditions described herein, the coupling efficiencies for different regions of the 2-5A-antisense molecule should be: antisense DNA, 98% to 99%; butanediol linkers, 97% to 98%; (2′,5′)-RNA, 94% to 98 %; and phosphorylation, 98% to 99%. The average stepwise coupling yield for 2-5A-antisense chimera should be 96% to 98%. When 2-5A-antisense oligonucleotides are synthesized and purified according to the procedures outlined herein, products of purities approaching 95% are possible. Isolated yields of purified chimeric oligonucleotides of 70 A260 units are achievable. The digestion of a representative 2-5A-antisense chimera with snake venom phosphodiesterase is shown in Figure 4.4.4. This HPLC picture of the snake venom digestion differs when the substrate for digestion contains a
(3′-3′)-phosphodiester-linked dinucleotide. When a representative (3′-3′)-tailed chimera, 5′-pA(2′5′-pA)3-2′p(Bup)2-pd[AAT GGG ATC CAT TTT GTC C(3′-3′)C], is digested under standard conditions with snake venom phosphodiesterase, the HPLC of digestion products will reveal seven major peaks with retention times of 10.8, 29.2, 31.1, 32.59, 39.3, 40.2, and 41.5 min. The first five correspond to, respectively, dCMP, dTMP, dGMP, rAMP, and AMP(pBu)2. At 40.2 min, a new product appears that is not observed in a similar digestion of a standard first-generation 2-5A-antisense chimera with no 3′-3′ linkage. Lastly, dAMP is at 41.5 min. The overall ratio of above nucleotidic digestion products is 3:7:4:3:4 for dCMP/dTMP/dGMP/rAMP/dAMP, revealing an underabundance of dCMP from what would be expected from complete digestion. However, the peak with retention time of 40.2 min possessed a UV spectrum that could be generated by 1:1 addition of the spectra of two mononucleotides, 2 × dCMP. Thus, the structure pdC3′p3′dC can be assigned to this new peak. The structure of this product (3′-3′)-dinucleotide can be corroborated by comparison of the HPLC chromatogram of enzymatically digested product with synthetic dinucleotide. Similar HPLC digestion patterns were obtained when other (3′-3′)-tailed chimeras were digested with snake venom phosphodiesterase. For instance, pA4-[pBu]2-pd[GCC CAC CGG GTC CAC CAT(3′-3′)C] gave the dinucleotide pdT3′p3′dC (Rt = 46.0 min) and pA4-[pBu]2pd[TGG GAA GCT GTC ACT GTA GAG(3′3′)C] yielded pdG3′p3′dC (Rt = 44.4 min). The (3′-3′)-dinucleotides can be assigned structures based on a comparison of their UV spectra with the calculated spectra from 1:1 summation of constituent mononucleotides. The retention times of the (3′-3′)-dinucleotide products varied depending upon their composition. For instance, the retention time of pdC3′p3′dC is close to that of other digestion products such as dAMP (Rt = 41.5 min) or AMP(pBu)2 (Rt = 40.2 min). Under these conditions, accurate analysis of these key digestion products may be problematic. To deal with this difficulty, digestion with snake venom phosphodiesterase can be carried out with the addition of bacterial alkaline phosphatase. This results in the removal of all non-internucleotide phosphates, so that digestion products consist of nucleosides and the (3′-3′)-dinucleotide. Thus, for instance, when the chimera 5′pA(2′,5′-pA)3-2′-(pBu)2-pd[AAT GGG ATC CAT TTT GTC C(3′-3′)C] is digested with
4.4.22 Supplement 1
Current Protocols in Nucleic Acid Chemistry
snake venom phosphodiesterase and bacterial alkaline phosphatase, the following products are obtained: dC (Rt = 25.6 min), dG (Rt = 42.8 min), dT (Rt = 44.8 min), dC3′p3′dC (Rt = 47.8 min), rA (Rt = 50.1 min), dA (Rt = 51.8 min), and A2′pBupBu (Rt = 54.9). Thus, this modified digestion procedure resulted in a shift in Rt values such that each individual product was well separated from the others. This additional digestion procedure is not always necessary, and its use depends on the constitution of the dinucleotide product.
Time Considerations The following are estimations of the time needed for various stages of the synthesis, purification, and characterization of a chimeric 2-5A-antisense oligonucleotide. Support Protocol 1 requires 6 days for synthesis of N6-benzoyl-5′-O-dimethoxytrityl-3′O-tert-butyldimethylsilyladenosine-2′-N,Ndiisopropylphosphoramidite. Support Protocol 2 requires 2 days for synthesis of 4-O(4,4′-dimethoxytrityl)oxybutyl-1-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite. Basic Protocol 1 requires 6 to 8 hr for automated 2-5A-antisense chimera synthesis on an ABI 391 or 392 synthesizer, and 3 hr to overnight for cleavage and deprotection of the chimera. Basic Protocol 2 requires 6 hr for purification of the chimera by polystyrene reversedphase ion-pair HPLC; 6 hr for desalting and concentration of the chimera; 3 hr for cation exchange; 24 to 48 hr for dialysis and sterilization; 12 hr for capillary gel electrophosesis (a large number of samples may be run in this time when using an autosampler); and 4 hr for the Dionex HPLC purity check. Basic Protocol 3 requires 6 to 8 hr for snake venom digestion and HPLC analysis of three samples. Basic Protocol 4 requires 4 hr for labeling the chimera with terminal nucleotide transferase; 4 hr for determining the specific activity of the radioactive chimera; 6 to 8 hr for MaxamGilbert sequencing; plus the necessary time for autoradiograph exposure.
Literature Cited Beaucage, S.L. and Caruthers, M.H. 1981. Deoxyribonucleotide phosphoramidites—A new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Lett. 22:1859-1862. Cirino, N.M., Li, G., Xiao, W., Torrence. P.F., and Silverman, R.H. 1997. Targeting RNA for decay in respiratory syncytial virus infected cells with 2′→5′ oligoadenylate antisense. Proc. Natl. Acad. Sci. U.S.A. 94:1937-1942. Johnston, M.I. and Torrence, P.F. 1984. The role of interferon-induced proteins, double-stranded RNA and 2′,5′-oligoadenylate in the interferonmediated inhibition of viral translation. In Interferon: Mechanism of Production and Action (R.M. Freidman, ed.) pp. 189-298. Elsevier/ North-Holland, Amsterdam. Lesiak, K., Khamnei, S., and Torrence, P.F. 1993. 2′,5′-Oligoadenylate:antisense chimeras—Synthesis and properties. Bioconjugate Chem. 4:467-472. Li, G., Xiao, W., and Torrence, P.F. 1997. Synthesis and properties of second generation 2-5A-antisense chimeras with enhanced resistance to exonucleases. J. Med. Chem. 40:2959-2966. Maitra, R.K., Li, G., Xiao, W., Dong, B., Torrence, P., and Silverman, R.H. 1995. Catalytic cleavage of an RNA target by 2-5A antisense and RNase L. J. Biol. Chem. 270:15071-15075. Maran, A., Maitra, R.K., Kumar, A., Dong, B., Xiao, W., Li, G., Williams, B.R.G., Torrence, P.F., and Silverman, R.H. 1994. Blockage of NF-kB signaling by selective ablation of an mRNA target by 2-5A antisense chimeras. Science 265:789792. Player, M. and Torrence, P.F. 1998. The 2-5A system: Modulation of viral and cellular processes through acceleration of RNA degradation. Pharmacol. Therapeut. 78:55-113. Torrence, P.F., Maitra, R.K., Lesiak, K., Khamnei, S., Zhou, A., and Silverman, R.H. 1993. Targeting RNA for degradation with a (2′-5′)oligoadenylate-antisense chimera. Proc. Natl. Acad. Sci. U.S.A. 90:1300-1304. Xiao, W., Player, M.R., Li, G., Zhang, K., Lesiak, K., and Torrence, P.F. 1996. Synthesis and characterization of composite nucleic acids containing 2′,5′-oligoriboadenylate linked to antisense DNA. Antisense Nucleic Acid Drug Devel. 6:247-258.
Contributed by Mark R. Player and Paul F. Torrence Northern Arizona University Flagstaff, Arizona
Synthesis of Modified Oligonucleotides and Conjugates
4.4.23 Current Protocols in Nucleic Acid Chemistry
Supplement 1
Attachment of Reporter and Conjugate Groups to the 3′ Termini of Oligonucleotides Applications of oligonucleotide conjugates encompass mechanistic and hybridization probes, antisense agents, and sensors. Consequently, methods for their synthesis constitute an active area of research. This unit focuses on methods for the preparation of oligonucleotides conjugated at their 3′ termini (Fig. 4.5.1, S.1). Conjugation at the 3′ terminus of an oligonucleotide has been noted as being less common than at other sites within oligonucleotides, because this position commonly serves as the site for covalent linkage to the solid phase synthesis support (Goodchild, 1990); however, the valuable physicochemical properties of 3′-oligonucleotide conjugates, such as their ability to stabilize nucleic acid hybridization complexes and to retard the activity of exonucleases, provides strong incentive to develop methods for their preparation. The synthesis of 3′-oligonucleotide conjugates was included in previous reviews published during the past decade, and therefore this author will try to minimize redundancy. The reader is referred to these previous reviews for further information (Goodchild, 1990; Agrawal, 1994; Beaucage and Iyer, 1993; Fidanza et al., 1994). Methods for the synthesis of 3′-oligonucleotide bioconjugates will be divided into three general categories: preparation of modified supports and phosphoramidites, postsynthetic modification of deprotected oligonucleotides, and the recently developed method utilizing solution phase conjugation of protected oligonucleotides. The latter two methods require modified solid phase synthesis supports which release oligonucleotides containing the appropriate 3′-terminal functional group. Such supports are described briefly in this review. The variety of new supports that produce oligonucleotides containing 3′-hydroxyl termini are not included. Due to the growing importance of nucleopeptides and surface/polymer supported oligonucleotides, methods for the synthesis of these two general
5'-HO
oligo
O
conjugant
1
Figure 4.5.1 jugate.
Generic oligonucleotide 3′-con-
UNIT 4.5
families of conjugates are presented separately. A representative, but not exhaustive, set of examples of each method is presented, and apologies are extended to any authors whose work is not cited.
MODIFIED SOLID PHASE SYNTHESIS SUPPORTS AND PHOSPHORAMIDITES Modifying the 3′ termini of oligonucleotides with polymers such as poly-L-lysine or polyethylene glycol (PEG) is desirable in order to improve the pharmacological properties of antisense probes. 3′-PEG-derivatized oligonucleotides have been prepared from a specially designed solid phase synthesis support in which the PEG serves as a linker between the controlled-pore glass (CPG) support and the growing oligonucleotide (Fig. 4.5.2, S.2; Jaschke et al., 1993). By loading the PEG onto the solid support as its succinato ester, 3′-PEG-derivatized oligonucleotides are released under standard aqueous ammonium hydroxide deprotection conditions. Succinato derivatives of smaller molecules, such as cholesterol, have also been used to prepare 3′-oligonucleotide conjugates (Fig. 4.5.2, S.3; MacKellar et al., 1992). A variety of solid phase supports have been prepared from a common amino derivative of glycerol (Nelson et al., 1992). The masked nucleophile was originally employed as it N-9fluorenylmethoxycarbonyl (Fmoc)–protected derivatives as a means for introducing 3′-alkylamines into oligonucleotides (Nelson et al., 1989). In later studies, the support was modified to contain a longer butylamine linker in order to alleviate potential steric interactions between the conjugated moiety and the remainder of the oligonucleotide backbone (Fig. 4.5.3, S.4 and S.5; Nelson et al., 1989). Recently, this modification was used for preparing doubledye-labeled oligonucleotides containing tetramethylrhodamine at their 3′ termini (Mullah and Andrus, 1997). In each of the above examples a new solid phase synthesis support must be prepared for each 3′ reporter group or conjugate. Recently, a method that circumvents this procedure was reported (Nelson et al., 1997). The method employs a universal solid phase support (Fig. 4.5.4, S.6) which releases an intermediate 3′-
Contributed by Marc M. Greenberg Current Protocols in Nucleic Acid Chemistry (2000) 4.5.1-4.5.19 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Modified Oligonucleotides and Conjugates
4.5.1 Supplement 2
O O(CH2CH2O)nDMTr
(CH2)3NH O
O O
DMTrO O
N H
O DMTr, 4,4'-dimethoxytrityl
3
Figure 4.5.2 Solid phase supports for attaching polyethylene glycol (S.2) and cholesterol (S.3) to the 3′ termini of oligonucleotides.
tetrahydrofuranyl phosphodiester (Schwartz et al., 1995; Scheuer-Larsen et al., 1997). This intermediate undergoes subsequent cyclic phosphate formation and ultimately releases a 3′-hydroxyl oligonucleotide upon further alkaline treatment in a manner analogous to RNA hydrolysis. When a modified phosphoramidite is the first species coupled to the support, 3′modified oligonucleotides containing hydroxyl termini are produced upon cleavage. Although the method still requires the preparation of the appropriate modified phosphoramidite, it has the advantage that the same amidite can be used for introducing the respective modification at other sites within the oligonucleotide. Hence, one does not need to synthesize both a modified support and a modified phosphoramidite. Furthermore, when used in conjunction with a more labile diglycolate linker in lieu of the standard succinato moiety,
the oligonucleotides are cleaved using a mixture of t-butylamine, methanol, and water. The dye-labeled oligonucleotides prepared by the above methods have been used for detecting a polymerase chain reaction (PCR) product in real time (Mullah et al., 1998). In an earlier oligonucleotide conjugate synthesis method, attomolar oligonucleotide detection limits were achieved using multiple nonradioactive labels at the 3′ termini (Haralambidis et al., 1990a,b). The method relied upon linear synthesis of a polyamide on CPG, followed by incorporation of a linker molecule, which allows one to then carry out standard oligonucleotide synthesis. This strategy is useful for preparing nucleopeptides (see below), but more relevant to the present discussion is its utility for preparing oligonucleotides containing multiple labels via conjugation to alkylamine side chains of the polyamide portion of the mole-
O
O
O
O
NH
NH
HN
DMTrO
3
O
N H
2
S
4
O
DMTrO
O
O
NH
3
O
Attachment of Reporter and Conjugate Groups to the 3′ Termini of Oligonucleotides
N H
O
5
Figure 4.5.3 Solid support for conjugating biotin (S.4) or other conjugants (S.5) to oligonucleotides at their 3′ termini.
4.5.2 Supplement 2
Current Protocols in Nucleic Acid Chemistry
O
H N
O
ODMTr
O 6
O
H N std. oligo synthesis
O O P O X
O O
prot. oligo
OH-5'
OR
O
O aq. NH4OH, LiCl
−
−
O
O O O P O X
oligo
OH-5'
HO aq. NH4OH, LiCl
O O
oligo
O P O X O
OH-5'
–
O
oligo
HO X
Figure 4.5.4
OH-5'
X = modified phosphoramidite R = phosphate protecting group
A universal support for preparing 3′-modified oligonucleotides.
cule. The labels can be incorporated prior to or after oligonucleotide synthesis, while the nucleic acid component is on the solid support or in solution. In contrast, the thioester support (Fig. 4.5.5, S.7) enables introduction of conjugation as part of the main chain of the polymer (Hovinen et al., 1995). The thioester support was preceded by other versatile solid phase supports from which oligonucleotides containing a variety of 3′ termini could be prepared (Hovinen et al., 1993a,b, 1994); however, by utilizing a thioester linkage, more homogeneous products were obtained in good yields using a large excess of alkylamine nucleophiles (Hovinen et al., 1995).
SOLID PHASE SYNTHESIS SUPPORTS FOR PRODUCING 3′-FUNCTIONAL GROUPS SUITABLE FOR CONJUGATION In order to carry out postsynthetic conjugation of oligonucleotides, the biopolymers must contain a suitable functional group at their 3′ termini. Utilization of the nucleophilicity of sulfur in conjugation chemistry has resulted in the preparation of numerous solid phase supports that release oligonucleotides containing thiols at their 3′ termini (e.g., Fig. 4.5.6, S.8; Asseline et al., 1992; Bonfils and Thuong,
1991; Gottikh et al., 1990; Gupta et al., 1990, 1991; Kumar, 1993a,b; Zuckerman et al., 1987). In some instances the initially cleaved 3′-thiol group is transformed into other functional groups that are also useful for preparing bioconjugates. These groups include 3′-phosphates, 3′-phosphorothioates, 3′-alkylamines, and 3′-alkyl carboxylic acids (Asseline et al, 1992; Gottikh et al., 1990; Gupta et al., 1991). 3′-Phosphorylated oligonucleotides can be obtained by a variety of supports, some of which are designed to be compatible with chemically unstable biopolymers (Gryaznov and Letsinger, 1992; Guzaev and Lönnberg, 1997). Preparation of oligonucleotides containing 3′-dialdehydes are readily prepared by utilizing a solid phase support containing a ribonucleoside, but this limits the structure of the tether between the oligonucleotide and 3′-electrophile. Consequently, supports designed to release alkyl aldehydes have been developed (e.g., Fig. 4.5.6, S.9; Urata and Akagi, 1993). This glyceryl support is designed to be universal, and to release a terminal, vicinal diol upon cleavage
O O
NH
S
8
NH
OAc O
NH
S
ODMTr
O
ODMTr O
ODMTr
O
O
O
S
Ac, acetyl
9
7
Figure 4.5.5 Thioester support for the preparation of 3′-modified oligonucleotides.
Figure 4.5.6 Supports for the release of oligonucleotides containing 3′-thiols (S.8) and 3′aldehydes (S.9).
Synthesis of Modified Oligonucleotides and Conjugates
4.5.3 Current Protocols in Nucleic Acid Chemistry
Supplement 2
O O
NH
ODMTr O
10
O O
NH
O
O
ODMTr O
11
O N H
O
O O
O
N H
OH S
O 12
Figure 4.5.7
Attachment of Reporter and Conjugate Groups to the 3′ Termini of Oligonucleotides
O
ODMTr 4
13
Supports for the release of oligonucleotides containing various 3′-functional groups.
under standard concentrated aqueous ammonia conditions. Subsequent periodate oxidation generates the desired 3′-aldehyde. A versatile family of solid phase supports that enable one to introduce 3′-alkyl carboxylic acids, 3′-alkylamines, or 3′-alkyl thiols concomitantly with cleavage/deprotection of the oligonucleotides from the supports has also been reported (e.g., Fig. 4.5.7, S.10 and S.11; Hovinen et al., 1993a,b, 1994, 1995). In some instances differentiation of the cleavage and deprotection steps can be achieved by employing a thioester-based support (Fig. 4.5.5, S.7). An alternative variation that utilizes this concept appeared recently (Lyttle et al., 1997). In this instance, a common solid phase support (Fig. 4.5.7, S.12) is modified prior to oligonucleotide synthesis with an appropriate dimethoxytritylated tether to produce a support (e.g., Fig. 4.5.7, S.13) that will release oligonucleotides containing the desired 3′-terminal functional group upon cleavage/deprotection. In most instances, oligonucleotides are cleaved from the above supports under conditions that also result in cleavage of the exocyclic amines and phosphate diester protecting groups. In some instances, transamination is a competitive process (Hovinen et al., 1995). Orthogonal solid phase supports enable one to cleave oligonucleotides from the support without affecting the protecting groups throughout the biopolymer. These supports have proven invaluable in developing methods for bioconjugation using protected oligonu-
cleotides. The first reported series of orthogonal solid phase supports utilized UV-irradiation to induce cleavage. Oligonucleotides containing 3′-hydroxyl groups (Greenberg and Gilmore, 1994; Venkatesan and Greenberg, 1996), 3′-alkyl carboxylic acids (Yoo and Greenberg, 1995), 3′-alkylamines (McMinn and Greenberg, 1996), and 3′-phosphates (Avino et al., 1996; Dell ’Aquila et al., 1997; McMinn et al., 1998; Fig. 4.5.8, S.14 to S.17, respectively) were obtained using the o-nitrobenzyl photoredox reaction. More recently, Pd(0) labile supports that facilitate the release of 3′-phosphato, 3′-hydroxyl, and 3′-alkyl carboxylic acid containing oligonucleotides have been reported (Fig. 4.5.8, S.18, S.19, and S.20, respectively; Greenberg et al., 1998; Matray et al., 1997; Zhang et al., 1997).
POSTSYNTHETIC CONJUGATION Conjugation of 3′-Alkylamines The polyamide methods developed by Haralambidis (1990a,b) are compatible with conjugation to alkylamines in solution, following oligonucleotide deprotection. Common means of forming such conjugates involve in situ activation of carboxylic acids, or the use of the less reactive but more water tolerant N-hydroxysuccinimide esters (NHS-esters) or isothiocyanates. Using a support similar to those developed by the Clontech group (Fig. 4.5.9, S.21), Thaden and Miller (1993a,b) prepared an oli-
4.5.4 Supplement 2
Current Protocols in Nucleic Acid Chemistry
O H3C DMTrO
NH N
O O
O DMTrO
O
O n
O
NO2
O
CH3O
H3CO
O
NO2
n= 1-4
H N
O
O N H
O 14
DMTrO
H N
15
DMTrO
O
n
O
NO2
NO2
n= 4-6
CH3O
CH3O
H N
O
H N
O 2
2
O
O 17
16
O H3C DMTrO
O
NH N
O
O
DMTrO
O O
N H
3
O
O 3
O
7
O
N H
19
18
O DMTrO 3
O O
3
N H
20
Figure 4.5.8
Photolabile and Pd(0)-labile orthogonal oligonucleotide synthesis supports.
gonucleoside methylphosphonate containing a 3′-rhodamine reporter group. Coupling is achieved with ∼85% yield by reacting ten molar equivalents of rhodamine isothiocyanate with the biopolymer at room temperature for 18 hr (Thaden and Miller, 1993b). Oligonucleotides containing 3′-chlorin groups (Fig. 4.5.9, S.22) were prepared by activating the carboxylic acid on the chlorin component (Boutorine et al., 1996). The required alkylamine-containing oligonucleotide was prepared indirectly from the 3′-phosphorylated biopolymer via a redox con-
densation reaction with a diamine. The coupling proceeded in 30% to 40% conversion and, like the isothiocyanate example above, required several hours. In another example from the same contribution to the literature, the 3′phosphate of the oligonucleotide was activated with a symmetrical dihydrazide, and the free hydrazide was condensed with a chlorin containing an aldehyde (Boutorine et al., 1996). The resulting hydrazone also was obtained in modest overall yield.
Synthesis of Modified Oligonucleotides and Conjugates
4.5.5 Current Protocols in Nucleic Acid Chemistry
Supplement 2
O
DMTrO O
NH
O N 2
O
2
N H
O
21 H O HO H3C
CH3
N NH
HN N
H3C
CH3
HO2C O
N H
4
O N P O H − O
oligo
OH-5'
22
Figure 4.5.9
Introduction of dyes and porphyrins at the 3′ termini of oligonucleotides.
Conjugation of 3′-Alkyl Carboxylic Acids Oligonucleotides containing 3′-alkyl carboxylic acids can be conjugated to amines following activation in situ with standard peptide coupling agents (Gottikh et al., 1990); however, one should note that these conditions can give rise to significant amounts of nonspecific covalent modification. One can obtain the requisite 3′-alkyl carboxylic acid directly from designed supports, or this functional group can be introduced postsynthetically. For example, daunomycin was linked to the 3′ terminus of an oligonucleotide (Fig. 4.5.10, S.23) via amide bond formation. The multistep process started from a disulfide support which yields 3′-phosphorylated oligonucleotides (Fig. 4.5.10, S.8; Gottikh et al., 1990). Activation of the phosphate group was followed by reaction with an amino acid. In this particular instance, activation/coupling with carbodiimide took place over 5 hr at 4°C, and yields were not reported.
Conjugation of 3′-Alkyl Thiols Attachment of Reporter and Conjugate Groups to the 3′ Termini of Oligonucleotides
Significant advantage has been taken of the nucleophilicity of sulfur for preparing oligonucleotide conjugates. As mentioned above, a variety of supports have been developed that result in the release of oligonucleotides containing 3′-alkyl thiols. Cleavage of 3′-alkyl
thiolated oligonucleotides from these supports can be effected under conditions that do not remove the exocyclic amine and phosphate diester protecting groups; however, the thiolcontaining oligonucleotides are typically cleaved/deprotected under reductive conditions in aqueous ammonia, and conjugated in aqueous solutions using the fully deprotected biopolymers. Excellent yields of sulfide linked conjugates are obtained by reacting thiolated oligonucleotides with α-halocarbonyl-containing conjugants (e.g., Fig. 4.5.11, S.24; Gupta et al., 1991). 3′-Oligonucleotide conjugates can also be prepared by reacting 3′-thiolated oligonucleotides with activated disulfides or maleimides (Gupta et al., 1991; Harrison and Balasubramanian, 1997; Kumar, 1993b). Reaction with a maleimide proceeds via the formation of a stable Michael adduct. In one approach, a variety of biologically useful conjugates was prepared in which the Michael acceptor (maleimide) was covalently attached to the 3′ terminus of the oligonucleotide, and the thiol-containing conjugant of interest was loaded on a solid support (Harrison and Balasubramanian, 1997). When preparing oligonucleotide conjugates containing 3′-disulfides, either the 3′-thiolated oligonucleotide or the conjugating species can be activated. Disul-
4.5.6 Supplement 2
Current Protocols in Nucleic Acid Chemistry
O
O O
NH
S
S
oligo
5'-HO
ODMTr
O P OH O
O
−
8
O oligo
5'-HO
O P O HN
O
−
n
oligo
5'-HO
O P O N O
CO2H
O
O P O HN
−
n
23
O
OH
O
N N
H3C
O oligo
5'-HO
OH
−
HO
R=
R O
H3C
O
O
OCH3
HO
Figure 4.5.10
Postsynthetic modification of oligonucleotide 3′-carboxylic acids.
fide transfer has been used for conjugating oligonucleotides at their 3′ termini to small or large molecules and surfaces (Asseline et al., 1992; Gupta et al., 1991; Harrison and Balasubramanian, 1997; Kumar, 1993b; Zuckerman et al., 1987). For example, hybrid enzymes that hydrolyze RNA sequences specifically have also been prepared via disulfide exchange (Zuckerman et al., 1988; Zuckerman and Schultz, 1988).
Conjugation of 3′-Phosphorothioates Conjugations at phosphorous atoms near the 3′ termini of oligonucleotides have been carried out by selectively oxidizing an H-phosphonate linkage with an appropriately substituted amine (Letsinger et al., 1989). The resulting phosphoramidate is carried through a conventional
O oligo
5'-HO
O P O
solid phase synthesis protocol using either phosphoramidite or H-phosphonate chemistry. Ammonolysis and chromatographic purification give a product containing >50% of the respective label. A more common approach to synthesizing 3′-oligonucleotide conjugates takes advantage of the nucleophilicity of phosphorothioates. In one example, 3′-phosphorothioate-containing oligonucleotides were prepared on a solid phase synthesis support (Fig. 4.5.6, S.8; Asseline et al., 1992). Following standard oligonucleotide synthesis and ammoniacal deprotection/cleavage under reducing conditions, the fully deprotected oligonucleotides were conjugated to halogenated substrates (Fig. 4.5.12). Conjugation to derivatives of daunomycin, fluorescein, and 1,10phenanthroline (S.25) containing alkyl halide
−
O
−
SO3 Na
SH
2
O
+
−
SO3 Na
+
oligo
O P O
−
O
S 3
O N H
NH
24
O
I
5'-HO
+
N H
Figure 4.5.11
NH
Postsynthetic modification of oligonucleotide 3′-thiols.
Synthesis of Modified Oligonucleotides and Conjugates
4.5.7 Current Protocols in Nucleic Acid Chemistry
Supplement 2
O oligo
5'-HO
O P OH S
−
N O
+
5'-HO
oligo
O
N
O
O P S
−
N H
N N
O I
N H
Figure 4.5.12
Attachment of Reporter and Conjugate Groups to the 3′ Termini of Oligonucleotides
25
Postsynthetic modification of oligonucleotide 3′-phosphorothioates.
functional groups are believed to have proceeded in essentially quantitative yields over 24 hours in either protic organic solvents containing crown ethers to enhance biopolymer solubility, or mixtures of dimethylformamide and water. This strategy was utilized recently in studies on truncated derivatives of the potent antitumor antibiotics, CC-1065 and the duocarmycins, which alkylate deoxyadenosine at N-3 (Lukhtanov et al., 1996). The active pharmacophore, the cyclopropapyrroloindole, was covalently linked via an α-bromoacetamido linkage to a 3′-phosphorothioate in 50% to 60% yield. Several reports of template-mediated oligonucleotide ligation involving 3′-phosphorothioate DNA have also appeared. Oxidative coupling of a 3′-phosphorothioate oligonucleotide to a 5′-phosphorothioated biopolymer is achieved rapidly (5 min) under mild conditions (0°C) using 1 µM K3Fe(CN)6 (Gryaznov and Letsinger, 1993a). Little or no coupling is detected under these conditions in the absence of a template. In contrast, ligation of a 5′-phosphorothioated oligonucleotide to a 3′-bromoacetamide-containing oligonucleotide proceeded in ∼80% yield after 48 hr at 0°C in the absence of a template (Gryaznov and Letsinger, 1993b). As expected, the presence of a template, which increases the effective concentration of the reaction partners, significantly accelerated the reaction. Essentially quantitative yields of ligated product were obtained in only 20 minutes when a stoichiometric template was present. A distinct advantage of this latter coupling method is that the reaction does not require any exogenous condensing agents. In a subsequent investigation, autoligation was effectively carried out using reaction partners of the opposite polarity, 3′-phosphorothioate and 5′-bromoacetamide (Fig. 4.5.13; Gryaznov et al., 1994).
Conjugation to 3′-Aldehydes Template-mediated synthesis has also been very useful for the conjugation of oligonucleotides containing 3′-aldehydes (Goodwin and Lynn, 1992; Zhan and Lynn, 1997). Aldehydes are attractive electrophiles with which to form oligonucleotide conjugates. Condensation with primary amines under reductive conditions provides secondary amines which are stable to acid and base. Solid supports that produce oligonucleotides containing 3′-aldehydes directly are unknown. Consequently, 3′aldehydes are typically produced via periodate oxidation of a vicinal diol. A common practice for preparing 3′-oligonucleotide conjugates via aldehyde condensation takes advantage of incorporating a ribonucleoside at the 3′ terminus (Fig. 4.5.14; Leonetti et al., 1988). Periodate oxidation of the oligonucleotide containing a 3′-terminal ribonucleoside produces a dialdehyde which under reductive amination conditions generates a morpholine upon reaction with a primary amine. An often cited application of this method concerns the synthesis of poly-L-lysine conjugates of oligonucleotides (Leonetti et al., 1988, 1990). Haralambidis et al. (1994) have utilized this ability to introduce amino acids at the 3′ termini of oligonucleotides to enable attachment of a substituted benzaldehyde, which is then conjugated to an enzyme via reductive amination. Condensation of a dialdehyde with a primary amine was also used recently in the segmental synthesis of a biologically active hammerhead ribozyme (Bellon et al., 1996). Linkage of the two segments was carried out in a loop region of the hammerhead ribozyme which had been shown to not be crucial for catalytic activity. One segment was synthesized so as to incorporate a 5′-alkylamine, while the other half contained a 3′-terminal uridine which served as the source of the dialdehyde. Follow-
4.5.8 Supplement 2
Current Protocols in Nucleic Acid Chemistry
oligo
5'-HO
O
O − O P S OH
Br
oligo
N H
OH-3'
Template
5'-HO
oligo
O O P S O
Figure 4.5.13
O
−
oligo
N H
OH-3'
Template-mediated oligonucleotide coupling.
ing rapid and quantitative periodate oxidation, reductive condensation was carried out to 95% conversion over the course of seven days. In a more recent study, the electrophilic half of a ribozyme was synthesized on a glyceryl sup-
port (Fig. 4.5.6, S.9; Bellon et al., 1997; Urata and Akagi, 1993). In these experiments, conjugation of the two halves of a ribozyme within the loop II region of the unmodified hammerhead proceeded in as high as 81.2% yield
O 5'-DMTrO
prot. oligo
Prot.
O P O − O O
B
O
OTBDMS
O N H
O 1. deprotect 2. oxidize (NaIO4)
5'-HO
oligo
O O P O − O O
B
O H H
O
RNH2, NaCNBH3
5'-HO
oligo
O O P O − O
O
B
N R Prot., nucleobase protecting group TBDMS, tert-butyldimethylsilyl
Figure 4.5.14
Postsynthetic modification of oligonucleotides by reductive amination.
Synthesis of Modified Oligonucleotides and Conjugates
4.5.9 Current Protocols in Nucleic Acid Chemistry
Supplement 2
(48 hr) when borane pyridine was employed as a reducing agent.
Solution Phase Conjugation of Protected Oligonucleotides
Attachment of Reporter and Conjugate Groups to the 3′ Termini of Oligonucleotides
The need for orthogonal linkers that enable the removal of fully protected oligonucleotides from the support for further elaboration in solution was recognized by at least one leader in the field of oligonucleotide synthesis a number of years ago (Zon and Geiser, 1991). The first reports of conjugating protected oligonucleotides in solution appeared in 1997 (McMinn et al., 1997; also see McMinn and Greenberg, 1998, 1999). This new method for synthesizing oligonucleotide conjugates was made possible by the development of a family of orthogonal solid phase synthesis supports (Fig. 4.5.8, S.14 to S.20). Using light or Pd(0), these supports enable one to release oligonucleotides containing 3′-alkylamines, 3′-alkyl carboxylic acids, or 3′-phosphate diesters which retain their exocyclic amine, phosphate diester, and 5′-hydroxyl protecting groups. One advantage of this method is that potential deleterious side reactions are eliminated by utilizing protected oligonucleotides. Nonspecific covalent modifications of unprotected oligonucleotides can be a significant problem when conjugating biopolymers (Bischoff et al., 1987; Erout et al., 1996; Ghosh and Musso, 1987; Lund et al., 1988). In addition, the rates at which conjugation reactions proceed are considerably faster than similar bond-forming reactions using unprotected oligonucleotides. An explanation for this acceleration is uncertain at this time, but may be related to the solvent conditions. Conjugation reactions of protected oligonucleotides are carried out in aprotic organic solvents, whereas unprotected oligonucleotides are often conjugated in aqueous solvents. Amide bond formation in aqueous solvents may be adversely affected by stronger solvation (hydrogen bonding) of the reactants, as well as a lower effective molarity of amines due to protonation. The original report of solution phase conjugation of protected oligonucleotides utilized a redox condensation or a Mukaiyama reaction to activate carboxylic acids (Fig. 4.5.15; McMinn et al., 1997; Mukaiyama, 1976). The oligonucleotides contained 3′-alkylamines. During the course of developing the conjugation chemistry, it was discovered that the “fast deprotecting” amides used to protect deoxyadenosine and deoxyguanosine underwent transamidation with the 3′-terminal alkyl-
amines (McMinn and Greenberg, 1997). Thus, phenoxyacetyl-protected phosphoramidites should not be used in conjunction with alkylamine modifiers. The observed transamidation is of general importance, because of the commercial availability of alkylamine modifiers for oligonucleotide synthesis. This problem was overcome by using isobutyryl exocyclic amine protecting groups for deoxyadenosine, deoxycytidine, and deoxyguanosine. Subsequently, a variety of biologically relevant reporter groups such as biotin and acridine were conjugated in excellent yields (88%) under mild reaction conditions (2 hr at 55°C) using only ten molar equivalents of carboxylic acids and activating reagents relative to oligonucleotide substrate. Only cholesterol coupled in <88% yield (i.e., 83%), and this may be attributed to difficulties in isolation of the very nonpolar conjugate. Optimization of the reaction process revealed that yields were not diminished by using as few as five molar equivalents of reagents relative to 3′-functionalized oligonucleotide, and were unaffected by increasing the length of the biopolymer (McMinn and Greenberg, 1998). N-Protected tripeptides also served as suitable coupling partners. For example, conjugation of N-Fmoc-Gly-Gly-His proceeded in 89% yield (Fig. 4.5.16, S.26) and, in contrast to other methods, prior chemical elaboration of the tripeptide were not required in order to introduce this nuclease mimic (McMinn et al., 1997; Truffert et al., 1996). Coupling under the Mukaiyama conditions did result in some epimerization of the α-amino acid. This limitation was overcome using benzotriazol-1-yloxytripyrrolidinophosphonium hexafluorophosphate (PyBOP) or 2-(1H-benzotriazolyl)1,1,3,3-tetramethyluronium hexafluorophosphate (HBTU) as activating agents, with no detriment to the yields of the conjugates (McMinn and Greenberg, 1998). The greater ease of using PyBOP or HBTU also proved useful for the efficient synthesis of (5′,3′)-bisconjugates (Fig. 4.5.16, S.27; Kahl et al., 1998). Protected oligonucleotides containing 3′-alkylamines also proved to be suitable substrates for isocyanates. Nearly quantitative yields of alkyl aryl ureas (e.g., Fig. 4.5.17, S.28) were obtained upon reaction with ten molar equivalents of aryl isocyanates. Coupling of the less reactive alkyl isocyanate generated slightly lower yield (88%), despite using twice the number of equivalents of isocyanate. The ability to conjugate isocyanates to protected oligonucleotides containing 3′-alkylamines facilitated the synthesis of oligonucleotide-peptide conju-
4.5.10 Supplement 2
Current Protocols in Nucleic Acid Chemistry
DMTrO
H N
O
6
O
NO2
CH3O
H N
O 2
O 16 1. std. oligo synthesis 2. photolysis
O DMTrO
prot. oligo
O P O (CH2)6NH2 O
CN
RCO2H, (PyS)2, PPh3, DMAP O
DMTrO
H O P O (CH2)6 N
prot. oligo
O
CN
R O
deprotection
HO
oligo
O H O P O (CH2)6 N O
R
−
O
Py, pyridin-2-yl DMAP, 4-dimethylaminopyridine Figure 4.5.15
Solution phase conjugation of protected oligonucleotides.
gates in which the 3′ terminus of the oligonucleotide was coupled to the N terminus of a peptide (McMinn and Greenberg, 1998). Conjugate S.29 (Fig. 4.5.17) was obtained in a 70% yield upon reaction of twenty molar equivalents of the respective isocyanate for 4 hr at 55°C; however, the method was limited with respect to the sequences of peptides that can be employed. For instance, peptides containing bulky side chains at their amino termini coupled poorly. The resulting conjugates contain the opposite topology between the oligonucleotide and peptide as that obtained from the amide-forming reaction described in Figure 4.5.18.
Conjugation of the 3′ termini of oligonucleotides to the amine terminus of tripeptides was achieved in a more general manner by using protected oligonucleotides containing 3′alkyl carboxylic acids (Kahl and Greenberg, 1999). In general, conjugation of protected oligonucleotides containing 3′-alkyl carboxylic acids to primary alkyl amines proceeded equally efficiently as analogous reactions involving 3′-alkylamine-containing oligonucleotides. Furthermore, the amide-forming reaction was essentially unaffected by steric hindrance during coupling to the amino termini of peptides. Conjugation of 3′-alkyl-carboxylicacid-containing oligonucleotides to the amino termini of peptides produces conjugates con-
Synthesis of Modified Oligonucleotides and Conjugates
4.5.11 Current Protocols in Nucleic Acid Chemistry
Supplement 2
O
O d(oligonucleotide) O P O − O
N H N
5
O
H N
NH2
N H
O N H
26 O RNH(CH2)6O P O − O
O d(oligonucleotide) O P O(CH2)6NHR' − O
O R=
HN
3
H N NH
R' =
O
N N H
S O
O
H N
NH2 O
27
Figure 4.5.16 cleotides.
Oligonucleotide conjugates and bis-conjugates prepared from protected oligonu-
taining the same topology as the urea bondforming reaction (Fig. 4.5.18). In contrast to the urea bond-forming method, oligonucleotide-peptide conjugates synthesized in this manner proceeded efficiently even when the N-terminal amino acid was sterically hindered. For instance, S.30 (Fig. 4.5.18) was obtained in 95% yield via a PyBOP mediated coupling, whereas the analogous conjugate formed via coupling of an oligonucleotide amine and isocyanate proceeded in <20% yield (McMinn and Greenberg, 1998).
The Synthesis of Nucleopeptides The importance of nucleopeptides and protein-nucleic acid interactions has provided the driving force behind several methods for synthesizing these bioconjugates. Consequently, many of the methods for preparing 3′-oligonucleotide conjugates described above have been applied to the synthesis of nucleopeptides. Recently, the method involving the solution phase coupling of protected oligonucleotides containing 3′-alkylamines to carboxyl-protected peptides was applied to the synthesis of a conjugate consisting of the operator site of the λ
O
O 5'-d(oligonucleotide) O P O − O
5
N H
N H
NO2
28
O 5'-d(oligonucleotide) O P O − O Attachment of Reporter and Conjugate Groups to the 3′ Termini of Oligonucleotides
O
H N
N 5
N H
N H
O
O
O NH2 CH3
29 Figure 4.5.17 alkylamines.
Oligonucleotide conjugates prepared from protected substrates containing 3′-
4.5.12 Supplement 2
Current Protocols in Nucleic Acid Chemistry
O
O 5'-d(oligonucleotide) O P O O
(CH2)6 N H
−
R
O
H N
N H
O
N H
R'
O
+
R"
O
n
δ
O
(CH2)6 N H
−
R'
H N R
O
+
NH3
N H
O
n
−
R"
+
δ
δ
O 5'-d(oligonucleotide) O P O O
CH2Ph
O
H N
−
R'''
O
NH2
N H
3
3
+
O
−
δ
δ
30
−
δ O
O 5'-d(oligonucleotide) O P O
−
R'''= isobutyl
Figure 4.5.18
Topological control of nucleopeptide formation.
repressor helix-turn-helix protein and a modified version of the DNA-binding helix of λ repressor (McMinn and Greenberg, 1999). The desired bioconjugate (Fig. 4.5.19, S.31) was obtained in 72% yield using ten molar equivalents of peptide and PyBOP. Attempted synthesis of the related nucleopeptide (Fig. 4.5.19, S.32) containing the naturally occurring DNAbinding helix of λ repressor resulted in the isolation of S.33 (Fig. 4.5.19) in 96% yield. It was proposed that S.33 was formed via an N-
O
O 5'-d(CAG TGG TAT GAT A) O P O O
−
to O-transacylation during the concentrated aqueous ammonia deprotection of the initially formed conjugate. The fact that S.33 was not observed during the synthesis of S.31 suggests that even slight steric hindrance prevents the undesired transacylation process. Oligonucleotide conjugates of large proteins such as modified serum albumin have been prepared via disulfide exchange. In this study, a 3′-thiolated oligonucleotide obtained from a disulfide support was conjugated to a
(Gln-Ser-Ala-Val-Gly-Ala-Leu-Phe-Asn)NHCOCH3
N H
5
31 O
O 5'-d(CAG TGG TAT GAT A) O P O O
−
5
(Gln-Ser-Ala-Val-Gly-Ala-Leu-Phe-Asn)NHCOCH3
N H 32
O
O 5'-d(CAG TGG TAT GAT A) O P O O
−
5
N H
33
Figure 4.5.19
Solution phase synthesis of nucleopeptides.
(Gln-Ser)NH2
Synthesis of Modified Oligonucleotides and Conjugates
4.5.13 Current Protocols in Nucleic Acid Chemistry
Supplement 2
neoglycoprotein which was derivatized through an alkylamine on its surface with Nsuccinimidyl 3-(2-pyridyldithio)propionate (Bonfils et al., 1992). Alternatively, 3′-thiolated oligonucleotides were conjugated to peptides that were derivatized with maleimides (Arar et al., 1993; Soukchareun et al., 1998). Bioconjugates can be formed in high yield by this method; however, disadvantages include susceptibility of the thiolated oligonucleotides to oxidation, and difficulties in reactions with highly charged peptides due to aggregation (Soukchareun et al., 1998). Nucleopeptide conjugates have also been prepared via displacement of a 3′-iodoacetamide-substituted oligonucleotide by a cysteine-containing peptide (Tung et al., 1995). Two general methods have been developed for the synthesis of oligonucleotide-peptide conjugates on a single solid phase support. Bifunctional supports (e.g., Figs. 4.5.3 and 4.5.20, S.5 and S.34, respectively) can be employed for the synthesis of peptides (using N-Fmoc chemistry) and oligonucleotides sequentially (Basu and Wickstrom, 1995; Juby et al., 1991). This approach was also used for the synthesis of peptide libraries in which the oligonucleotide component on the bead served as a tag for identifying the peptide sequence (Nielsen et al., 1993). Alternatively, nucleopeptides can be synthesized in a linear fashion on a single support in which the peptide is synthesized first, followed by coupling of a transitional linking molecule, and then automated oligonucleotide synthesis (de la Torre, 1994; Soukchareun et al., 1995; Truffert et al., 1994, 1996). Typically, the linking molecule contains a protected primary alcohol and a carboxylic acid (or activated ester) at the other terminus. Finally, a convergent method for the synthesis of 3′-oligonucleotide-peptide conjugates was reported that takes advantage of a templatemediated transfer of a peptide from an intermediate thioester to a 3′-amine-containing oligonucleotide (Bruick et al., 1996).
Attachment of Reporter and Conjugate Groups to the 3′ Termini of Oligonucleotides
NC
O O P O O
ODMTr NHFmoc
34
Figure 4.5.20 Support for the linear synthesis of nucleopeptides.
Conjugation of Oligonucleotides to Surfaces Via their 3′ Termini Usage of surface-bound oligonucleotides in applications such as sequencing by hybridization and as sensors has resulted in a variety of methods for covalently linking the biopolymers to surfaces (Drmanac et al., 1993). Methods have also been developed for in situ synthesis of oligonucleotides on surfaces (Caviani Pease et al., 1994; Chee et al., 1996; Cohen et al., 1997; Maskos and Southern, 1992; McGall et al., 1996; Southern et al., 1994); however, only methods which involve conjugation of a previously synthesized oligonucleotide to a surface will be discussed in this commentary. Furthermore, in keeping with the theme of this commentary, only those methods that involve coupling to the 3′ terminus of an oligonucleotide will be presented. As is the case for the synthesis of nucleopeptides, many of the methods for synthesizing bioconjugates catalogued above have been applied to the problem of preparing surface-bound oligonucleotides. In one application directed towards detecting hybridization, silica surfaces are activated with terminal alkyl epoxides (Lamture et al., 1994). Oligonucleotides containing 3′-alkylamines are then conjugated to the surface via nucleophilic ring opening of the epoxide. In a related example, patterned DNA surfaces containing covalently bound oligonucleotides are also prepared via chemical modification of silica surfaces (Fig. 4.5.21; Chrisey et al., 1996a,b). The three step process for immobilizing the oligonucleotides begins with introducing an alkylamine by modifying the silica surface with the appropriate trialkoxysilane. Following derivatization of the alkylamines with a maleimide, 3′-thiolated oligonucleotides are conjugated to the surface. When combined with masks and existing photoresist technology, this chemistry is useful for preparing patterned surfaces on the micrometer scale. In another application directed towards developing sequencing by hybridization, oligonucleotides containing 3′-dialdehydes or 3′-alkylamines were coupled to acrylamide gels containing the complementary functionality via reductive amination (Timofeev et al., 1996). Finally, an extremely sensitive method for detecting oligonucleotide hybridization was recently reported utilizing thiolated oligonucleotides (5′ and 3′) bound to gold particles (Elghanian et al., 1997; Storhoff et al., 1998). Hybridization to target oligonucleotides in solution mediates the formation of networked gold particle aggregates. Aggregates of the gold
4.5.14 Supplement 2
Current Protocols in Nucleic Acid Chemistry
oligo
S O
H2N H2N H2N
H2N
HN
HN
HN
Si
Si
HN
Si
O
Si
O
Figure 4.5.21
O
O
O HN H2N
H2 N
N
HN
O
Si
HN
HN
Si
Si
O
OH-5'
O
N
O HN H2N
HN
O
Si
HN
O
Si
Modification of surfaces by conjugation of 3′-derivatized oligonucleotides.
particles result in readily measured color changes on as little as 10 femtomoles of material.
SUMMARY Although the scope of this review was limited to the synthesis of oligonucleotide conjugates through their 3′ termini, it is clear that there has been significant activity during the past decade in the development of methods for the synthesis of oligonucleotide conjugates in general. As more oligonucleotide-based therapeutics enter the clinic, and other applications such as genome sequencing and gene function develop, the impetus for the development of oligonucleotide conjugate synthesis methods will certainly continue.
LITERATURE CITED Agrawal, S. 1994. Functionalization of oligonucleotides with amino groups and attachment of amino specific reporter groups. In Protocols for Oligonucleotide Conjugates, Vol. 26: Synthesis and Analytical Techniques, pp. 93-120. Humana Press, Totowa, N.J. Arar, K., Monsigny, M., and Mayer, R. 1993. Synthesis of oligonucleotide-peptide conjugates containing a KDEL signal sequence. Tetrahedron Lett. 34:8087-8090. Asseline, U., Bonfils, E., Kurfurst, R., Chassignol, M., Roig, V., and Thuong, N. T. 1992. Solidphase preparation of 5′,3′-heterobifunctional oligodeoxyribonucleotides using modified solid supports. Tetrahedron 48:1233-1254.
Avino, A., Garcia, R.G., Diaz, A., Albericio, F., and Eritja, R. 1996. A comparative study of supports for the synthesis of oligonucleotides without using ammonia. Nucleosides Nucleotides 15:18711889. Basu, S. and Wickstrom, E. 1995. Solid phase synthesis of a D-peptide-phosphorothioate oligodeoxynucleotide conjugate from two arms of a polyethylene glycol-polystyrene support. Tetrahedron Lett. 36:4943-4946. Beaucage, S.L. and Iyer, R.P. 1993. The functionalization of oligonucleotides via phosphoramidite derivatives. Tetrahedron 49:1925-1963. Bellon, L., Workman, C., Scherrer, J., Usman, N., and Wincott, F. 1996. Morpholino-linked ribozymes: A convergent synthetic approach. J. Am. Chem. Soc. 118:3771-3772. Bellon, L., Workman, C., Jarvis, T.C., and Wincott, F.E. 1997. Post-synthetically ligated ribozymes: An alternative approach to iterative solid-phase synthesis. Bioconjugate Chem. 8:204-212. Bischoff, R., Coull, J.M., and Regnier, F.E. 1987. Introduction of 5′-terminal functional groups into synthetic oligonucleotides for selective immobilization. Anal. Biochem. 164:336-344. Bonfils, E. and Thuong, N.T. 1991. Solid phase synthesis of 5′, 3′-bifunctional oligodeoxyribonucleotides bearing a masked thiol group at the 3′-end. Tetrahedron Lett. 32:3053-3056. Bonfils, E., Depierreux, C., Midoux, P., Thuong, N.T., Monsigny, M., and Roche, A.C. 1992. Drug targeting: Synthesis and endocytosis of oligonucleotide-neoglycoprotein conjugates. Nucl. Acids Res. 20:4621-4629.
Synthesis of Modified Oligonucleotides and Conjugates
4.5.15 Current Protocols in Nucleic Acid Chemistry
Supplement 2
Boutorine, A.S., Brault, D., Takasugi, M., Delgado, O., and Helene, C. 1996. Chlorin-oligonucleotide conjugates: Synthesis, properties, and red light-induced photochemical sequence-specific DNA cleavage in duplexes and triplexes. J. Am. Chem. Soc. 118:9469-9476. Bruick, R.K., Dawson, P.E., Kent, S.B.H., Usman, N., and Joyce, G.F. 1996. Template-directed ligation of peptides to oligonucleotides. Chem. Biol. 3:49-56.
Goodchild, J. 1990. Conjugates of oligonucleotides and modified oligonucleotides: A review of their synthesis and properties. Bioconjugate Chem. 1:165-187. Goodwin, J.T. and Lynn, D.G. 1992. Template-directed synthesis: Use of a reversible reaction. J. Am. Chem. Soc. 114:9197-9198.
Caviani Pease, A., Solas, D., Sullivan, E.J., Cronin, M.T., Holmes, C.P., and Fodor, S.P.A. 1994. Light-generated oligonucleotide arrays for rapid DNA sequence analysis. Proc. Natl. Acad. Sci. U.S.A. 91:5022-5026.
Gottikh, M., Asseline, U., and Thuong, N.T. 1990. Synthesis of oligonucleotides containing a carboxyl group at either their 5′ end or their 3′ end and their subsequent derivatization by an intercalating agent. Tetrahedron Lett. 31:6657-6660.
Chee, M., Yang, R., Hubbell, E., Berno, A., Huang, X.C., Stern, D., Winkler, J., Lockhart, D.J., Morris, M.S., and Fodor, S.P.A. 1996. Accessing genetic information with high-density DNA arrays. Science 274:610-614.
Greenberg, M.M. and Gilmore, J.L. 1994. Cleavage of oligonucleotides from solid-phase supports using o-nitrobenzyl photochemistry. J. Org. Chem. 59:746-753.
Chrisey, L.A., Lee, G.U., and O’Ferrall, C.E. 1996a. Covalent attachment of synthetic DNA to selfassembled monolayer films. Nucl. Acids Res. 24:3031-3039.
Greenberg, M.M., Matray, T.J., Kahl, J.D., Dong, J.Y., and McMinn, D.L. 1998. Optimization and mechanistic analysis of oligonucleotide cleavage from palladium-labile solid-phase synthesis supports. J. Org. Chem. 63:4062-4068.
Chrisey, L.A., O’Ferrall, C.E., Spargo, B.J., Dulcey, C.S., and Calvert, J.M. 1996b. Fabrication of patterned DNA surfaces. Nucl. Acids Res. 24:3040-3047.
Gryaznov, S.M. and Letsinger, R.L. 1992. A new approach to synthesis of oligonucleotides with 3′ phosphoryl groups. Tetrahedron Lett. 33:41274128.
Cohen, G., Deutsch, J., Fineberg, J., and Levine, A. 1997. Covalent attachment of DNA oligonucleotides to glass. Nucl. Acids Res. 25:911-912.
Gryaznov, S.M. and Letsinger, R.L. 1993a. Template controlled coupling and recombination of oligonucleotide blocks containing thiophosphoryl groups. Nucl. Acids Res. 21:1403-1408.
de la Torre, B.G., Avino, A., Tarrason, G., Piulats, J., Albericio, F., and Eritja, R. 1994. Stepwise solidphase synthesis of oligonucleotide-peptide hybrids. Tetrahedron Lett. 35:2733-2736. Dell’Aquila, C., Imbach, J.-L., and Rayner, B. 1997. Photolabile linker for the solid-phase synthesis of base-sensitive oligonucleotides. Tetrahedron Lett. 38:5289-5292. Drmanac, R., Drmanac, S., Strezoska, Z., Paunesku, T., Labat, I., Zeremski, M., Snoddy, J., Funkhouser, W.K., Koop, B., Hood, L., and Crkvenjakov, R. 1993. DNA sequence determination by hybridization: A strategy for efficient large-scale sequencing. Science 260:1649-1652. Elghanian, R., Storhoff, J.J., Mucic, R.C., Letsinger, R.L., and Mirkin, C.A. 1997. Selective colorimetric detection of polynucleotides based on the distance-dependent optical properties of gold nanoparticles. Science 277:1078-1081. Erout, M.-N., Troesch, A., Pichot, C., and Cros, P. 1996. Preparation of conjugates between oligonucleotides and N-vinylpyrrolidine/N-acryloxysuccinimide copolymers and applications in nucleic acids assays to improve sensitivity. Bioconjugate Chem. 7:568-575.
Attachment of Reporter and Conjugate Groups to the 3′ Termini of Oligonucleotides
Ghosh, S.S. and Musso, G.F. 1987. Covalent attachment of oligonucleotides to solid supports. Nucl. Acids Res. 15:5353-5372.
Fidanza, J.A., Ozaki, H., and McLaughlin, L.W. 1994. Functionalization of oligonucleotides by the incorporation of thio-specific reporter groups. In Protocols for Oligonucleotide Conjugates, Vol. 26: Synthesis and Analytical Techniques (S. Agrawal, ed.) pp. 121-143. Humana Press, Totowa, N.J.
Gryaznov, S.M. and Letsinger, R.L. 1993b. Chemical ligation of oligonucleotides in the presence and absence of a template. J. Am. Chem. Soc. 115:3808-3809. Gryaznov, S.M., Schultz, R., Chaturvedi, S.K., and Letsinger, R.L. 1994. Enhancement of selectivity in recognition of nucleic acids via chemical autoligation. Nucl. Acids Res. 22:2366-2369. Gupta, K.C., Sharma, P., Sathyanarayana, S., and Kumar, P. 1990. A universal solid support for the synthesis of 3′-thiol group containing oligonucleotides. Tetrahedron Lett. 31:2471-2474. Gupta, K.C., Sharma, P., Kumar, P., and Sathyanarayana, S. 1991. A general method for the synthesis of 3′-sulfhydryl and phosphate group containing oligonucleotides. Nucl. Acid Res. 19:3019-3025. Guzaev, A. and Lönnberg, H. 1997. A novel solid sup por t f or sy nthesis 3′-phosphorylated chimeric oligonucleotides containing internucleosidic methyl phosphotriester and methylphosphonate linkages. Tetrahedron Lett. 38:3989-3992. Haralambidis, J., Duncan, L., Angus, K., and Tregear, G.W. 1990a. The synthesis of polyamide-oligonucleotide conjugate molecules. Nucl. Acids Res. 18:493-499.
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Current Protocols in Nucleic Acid Chemistry
Haralambidis, J., Angus, K., Pownall, S., Duncan, L., Chai, M., and Tregear, G.W. 1990b. The preparation of polyamide-oligonucleotide probes containing multiple non-radioactive labels. Nucl. Acids Res. 18:501-505.
Leonetti, J.P., Rayner, B., Lemaitre, M., Gagnor, C., Milhaud, P.G., Imbach, J.-L., and Lebleu, B. 1988. Antiviral activity of conjugates between poly(L-lysine) and synthetic oligodeoxyribonucleotides. Gene 72:323-332.
Haralambidis, J., Lagniton, L., and Tregear, G.W. 1994. The preparation of enzyme-labelled oligonucleotides by reductive amination. Bioorg. Med. Chem. Lett. 4:1005-1010.
Leonetti, J.-P., Degols, G., and Lebleu, B. 1990. Biological activity of oligonucleotide-poly(L-lysine) conjugates: Mechanism of cell uptake. Bioconjugate Chem. 1:149-153.
Harrison, J.G. and Balasubramanian, S. 1997. A convenient synthetic route to oligonucleotide conjugates. Bioorg. Med. Chem. Lett. 7:10411046.
Letsinger, R.L., Zhang, G., Sun, D.K., Ikeuchi, T., and Sarin, P.S. 1989. Cholesteryl-conjugated oligonucleotides: synthesis, properties, and activity as inhibitors of replication of human immunodeficieny virus in cell culture. Proc. Natl. Acad. Sci. U.S.A. 86:6553-6556.
Hovinen, J., Gouzaev, A.P., Azhayev, A.V., and Lönnberg, H. 1993a. A new method to prepare 3′-modified oligonucleotides. Tetrahedron Lett. 34:5163-5166. Hovinen, J., Guzaev, A., Azhayev, A., and Lönnberg, H. 1993b. Synthesis of 3′-functionalized oligonucleotides on a single solid support. Tetrahedron Lett. 34:8169-8172. Hovinen, J., Guzaev, A., Azhayev, A., and Lönnberg, H. 1994. Novel solid supports for the preparation of 3′-derivatized oligonucleotides: introduction of 3′-alkylphosphate tether groups bearing amino, carboxy, carboxamido, and mercapto functionalities. Tetrahedron 50:7203-7218. Hovinen, J., Guzaev, A., Azhayeva, E., Azhayev, A., and Lönnberg, H. 1995. Imidazole tethered oligodeoxyribonucleotides: Synthesis and RNA cleaving activity. J. Org. Chem. 60:2205-2209. Jaschke, A., Furste, J.P., Dieter, C., and Volker, A.E. 1993. Automated incorporation of polythylene glycol into synthetic oligonucleotides. Tetrahedron Lett. 34:301-304. Juby, C.D., Richardson, C.D., and Brousseau, R. 1991. Facile preparation of 3′ oligonucleotidepeptide conjugates. Tetrahedron Lett. 32:879882. Kahl, J.D. and Greenberg, M.M. 1999. Solution phase bioconjugate synthesis using protected oligonucleotides containing 3′-alkyl carboxylic acids. J. Org. Chem. 64:507-510. Kahl, J.D., McMinn, D.L., and Greenberg, M.M. 1998. High-yielding method for on-column derivatization of protected oligodeoxynucleotides and its application to the convergent synthesis of 5′,3′-bis-conjugates. J. Org. Chem. 63:48704871. Kumar, A. 1993a. A rapid solid phase method for the synthesis of 3′-thiol group containing oligonucleotides. Nucleosides Nucleotides 12:729-736. Kumar, A. 1993b. A versatile solid phase method for the synthesis of masked 3′-thiol group containing oligonucleotides. Nucleosides Nucleotides 12:1047-1059. Lamture, J.B., Beattie, K.L., Burke, B.E., Eggers, M.D., Ehrilch, D.J., Fowler, R., Hollis, M.A., Kosicki, B.B., Reich, R.K., Smith, S.R., Varma, R.S., and Hogan, M.E. 1994. Direct detection of nucleic acid hybridization on the surface of a charge coupled device. Nucl. Acids Res. 22:2121-2125.
Lukhtanov, E.A., Podyminogin, M.A., Kutyavin, I.V., Meyer, R.B. Jr., and Gamper, H.B. 1996. Rapid and efficient hybridization-triggered crosslinking within a DNA duplex by an oligodeoxyribonucleotide bearing a conjugated cyclopropapyrroloindole. Nucl. Acids Res. 24:683687. Lund, V., Schmid, R., Rickwood, D., and Hornes, E. 1988. Assessment of methods for covalent binding of nucleic acids to magnetic beads, Dynabeads, and the characteristics of the bound nucleic acids in hybridization reactions. Nucl. Acids Res. 16:10861-10880. Lyttle, M.H., Adams, H., Hudson, D., and Cook, R.M. 1997. Versatile linker chemistry for synthesis of 3′-modified DNA. Bioconjugate Chem. 8:193-198. MacKellar, C., Graham, D., Will, D.W., Burgess, S., and Brown, T. 1992. Synthesis and physical properties of anti-HIV antisense oligonucleotides bearing terminal lipophilic groups. Nucl. Acids Res. 20:3411-3417. Maskos, U. and Southern, E.M. 1992. Parallel analysis of oligodeoxyribonucleotide (oligonucleotide) interactions. I. Analysis of factors influencing oligonucleotide duplex formation. Nucl. Acids Res. 20:1675-1678. Matray, T.J., Dong, J.Y., McMinn, D.L., and Greenberg, M.M. 1997. Synthesis of oligonucleotides containing 3′-alkylcarboxylic acids using a palladium labile oligonucleotide solid phase synthesis support. Bioconjugate Chem. 8:99-102. McGall, G., Labadie, J., Brock, P., Wallraff, G., Nguyen, T., and Hinsberg, W. 1996. Light-directed synthesis of high-density oligonucleotide arrays using semiconductor photoresists. Proc. Natl. Acad. Sci. U.S.A. 93:13555-13560. McMinn, D.L. and Greenberg, M. M. 1996. Novel solid phase synthesis supports for the preparation of oligonucleotides containing 3′-alkyl amines. Tetrahedron 52:3827-3840. McMinn, D.L. and Greenberg, M.M. 1997. Synthesis of oligonucleotides containing 3′-alkyl amines using N-isobutyryl protected deoxyadenosine phosphoramidite. Tetrahedron Lett. 38:3123-3126.
Synthesis of Modified Oligonucleotides and Conjugates
4.5.17 Current Protocols in Nucleic Acid Chemistry
Supplement 2
McMinn, D.L. and Greenberg, M. M. 1998. Postsynthetic conjugation of protected oligonucleotides containing 3’-alkylamines. J. Am. Chem. Soc. 120:3289-3294. McMinn, D.L. and Greenberg, M.M. 1999. Convergent solution-phase synthesis of a nucleopeptide using a protected oligonucleotide. Bioorg. Med. Chem. Lett. 9:547-550. McMinn, D.L., Matray, T.J., and Greenberg, M.M. 1997. Efficient solution phase synthesis of oligonucleotide conjugates using protected biopolymers containing 3′-terminal alkyl amines. J. Org. Chem. 62:7074-7075. McMinn, D.L., Hirsch, R., and Greenberg, M. M. 1998. An orthogonal solid phase support for the synthesis of oligonucleotides containing 3′phosphates and its application in the preparation of photolabile hybridization probes. Tetrahedron Lett. 39:4155-4158. Mukaiyama, T. 1976. Oxidation-reduction condensation. Angew. Chem. Int. Ed. Engl. 15:94-103. Mullah, B. and Andrus, A. 1997. Automated synthesis of double dye-labeled oligonucleotides using tetramethylrhodamine (TAMRA) solid supports. Tetrahedron Lett. 38:5751-5754. Mullah, B., Livak, K., Andrus, A., and Kenney, P. 1998. Efficient synthesis of double dye-labeled oligodeoxyribonucleotide probes and their application in a real time PCR assay. Nucl. Acids Res. 26:1026-1031. Nelson, P., Frye, R.A., and Liu, E. 1989. Bifunctional oligonucleotide probes synthesized using a novel CPG support are able to detect single base pair mutations. Nucl. Acids Res. 17:71877194. Nelson, P.S., Kent, M., and Muthini, S. 1992. Oligonucleotide labeling methods. 3. Direct labeling of oligonucleotides employing a novel, nonnucleosidic, 2-aminobutyl-1,3-propanediol backbone. Nucl. Acids Res. 20:6253-6259. Nelson, P., Muthini, S., Kent, M.A., and Smith, T.H. 1997. 3′-Terminal modification of oligonucleotides using a universal solid support. Nucleosides Nucleotides 16:1951-1959. Nielsen, J., Brenner, S., and Janda, K.D. 1993. Synthetic methods for the implementation of encoded combinatorial chemistry. J. Am. Chem. Soc. 115:9812-9813. Scheuer-Larsen, C., Rosenbohm, C., Jorgensen, T.J.D, and Wengel, J. 1997. Introduction of a universal solid support for oligonucleotide synthesis. Nucleosides Nucleotides 16:67-80. Schwartz, M.E., Breaker, R.R., Asteriadis, G.T., and Gough, G.R. 1995. A universal adapter for chemical synthesis of DNA or RNA on any single type of solid support. Tetrahedron Lett. 36:27-30. Attachment of Reporter and Conjugate Groups to the 3′ Termini of Oligonucleotides
Soukchareun, S., Tregear, G.W., and Haralambidis, J. 1995. Preparation and characterization of antisense oligonucleotide-peptide hybrids containing viral fusion peptides. Bioconjugate Chem. 6:43-55.
Soukchareun, S., Haralambidis, J., and Tregear, G. 1998. Use of Nα-Fmoc-cysteine(S-thiobutyl) derivatized oligodeoxynucleotides for the preparation of oligodeoxynucleotide-peptide hybrid molecules. Bioconjugate Chem. 9:466-475. Southern, E.M., Case-Green, S.C., Elder, J.K., Johnson, M., Mir, K.U., Wang, L., and Williams, J.C. 1994. Arrays of complementary oligonucleotides for analysing the hybridisation behaviour of nucleic acids. Nucl. Acids Res. 22:13681373. Storhoff, J.J., Elghanian, R., Mucic, R.C., Mirkin, C.A., and Letsinger, R.L. 1998. One-pot colorimetric differentiation of polynucleotides with single base imperfections using gold nanoparticle probes. J. Am. Chem. Soc. 120:1959-1964. Thaden, J. and Miller, P.S. 1993a. Automated synthesis of oligodeoxyribonucleoside methylphosphonates having [N-(3-aminoprop-1-yl)-N-(2hydroxyethyl)-2-aminoethyl] phosphate or methylphosphonic acid at the 3′ end using a modified controlled pore glass support. Bioconjugate Chem. 4:395-401. Thaden, J. and Miller, P.S. 1993b. Photoaffinity behavior of a conjugate of oligonucleoside methylphosphonate, rhodamine, and psoralen in the presence of complementary oligonucleotides. Bioconjugate Chem. 4:386-394. Timofeev, E.N., Kochetkova, S.V., Mirzabekov, A.D., and Florentiev, V.L. 1996. Regioselective immobilization of short oligonucleotides to acrylic copolymer gels. Nucl. Acids Res. 24:3142-3148. Truffert, J.-C., Lorthioir, O., Asseline, U., Thuong, N.T., and Brack, A. 1994. On-line solid phase synthesis of oligonucleotide-peptide hybrids using silica supports. Tetrahedron Lett. 35:23532356. Truffert, J.-C., Asseline, U., Brack, A., and Thuong, N.T. 1996. Synthesis, purification and characterization of two peptide-oligonucleotide conjugates as potential artificial nucleases. Tetrahedron 52:3005-3016. Tung, C.-H., Wang, J., Leibowtiz, M.J., and Stein, S. 1995. Dual-specificity interaction of HIV-1 TAR RNA with tat peptide-oligonucleotide conjugates. Bioconjugate Chem. 6:292-295. Urata, H. and Akagi, M. 1993. A convenient synthesis of oligonucleotides with a 3′-phosphoglycolate and 3′-phosphoglycaldehyde terminus. Tetrahedron Lett. 34:4015-4018. Venkatesan, H. and Greenberg, M. M. 1996. Improved utility of photolabile solid phase synthesis supports for the synthesis of oligonucleotides containing 3′-hydroxyl termini. J. Org. Chem. 61:525-529. Yoo, D.J. and Greenberg, M. M. 1995. Synthesis of oligonucleotides containing 3′-alkyl carboxylic acids using universal, photolabile solid phase synthesis supports. J. Org. Chem. 60:3358-3364. Zhan, Z.J. and Lynn, D.G. 1997. Chemical amplification through template-directed synthesis. J. Am. Chem. Soc. 119:12420-12421.
4.5.18 Supplement 2
Current Protocols in Nucleic Acid Chemistry
Zhang, X., Gaffney, B.L., and Jones, R.A. 1997. RNA synthesis using a universal, base-stable alkyl linker. Nucl. Acids Res. 25:3980-3983.
Zuckerman, R.N. and Schultz, P.G. 1988. A hybrid sequence-selective ribonuclease S. J. Am. Chem. Soc. 110:6592-6594.
Zon, G. and Geiser, T.G. 1991. Phosphorothioate oligonucleotides: Chemistry, purification, analysis, scale-up and future directions. Anti-Cancer Drug Des. 6:539-568.
Zuckerman, R.N., Corey, D.R., and Schultz, P.G. 1988. Site-selective cleavage of RNA by a hybrid enzyme. J. Am. Chem. Soc. 110:1614-1615.
Zuckerman, R., Corey, D., and Schultz, P. 1987. Efficient methods for attachment of thiol specific probes to the 3′-ends of synthetic oligodeoxyribonucleotides. Nucl. Acids Res. 15:5305-5321.
Contributed by Marc M. Greenberg Colorado State University Fort Collins, Colorado
Financial support of this work from the National Science Foundation (CHE-9732843), the CATI Center for RNA Chemistry, Dupont, and the Alfred P. Sloan Foundation is greatly appreciated.
Synthesis of Modified Oligonucleotides and Conjugates
4.5.19 Current Protocols in Nucleic Acid Chemistry
Supplement 2
3′-Modified Oligonucleotides and their Conjugates
UNIT 4.6
This unit describes synthetic methods for the preparation of three types of oligonucleotide 3′-modifications. The first two, aminoalkyl and sulfhydrylalkyl, are important because these groups offer specific points of attachment to a large variety of dyes and reporter groups. Their preparation is described in the Basic Protocol and Alternate Protocol 1, respectively. Alternate Protocol 2 deals with the 3′-attachment of polyethylene glycol (PEG) or other diols to the synthetic oligo. PEG-functionalized oligos are of interest because of their enhanced cell membrane permeability, which is important for antisense drug development. Implementation of the following procedures requires working knowledge of automated DNA synthesizers, high-performance liquid chromatographs (HPLCs), and UV/visible spectrophotometers. The use of derivatized supports requires the ability to pack DNA synthesis columns. In addition, access to a fume hood is required, as many of the preparations use malodorous, toxic, or otherwise hazardous reagents. CAUTION: Safe conduct in the laboratory when handling common solvents and chemicals, and working knowledge of equipment such as vacuum lines and rotary evaporators, is essential. Safety glasses and gloves should be worn to minimize risk of exposure. CAUTION: Many of the steps involve pyridine, which is extremely toxic and has a foul odor. Dichloroacetic acid causes severe burns when spilled on the skin. These steps should be performed in a fume hood with gloves and eye protection. Do not use magnetic stir bars with controlled-pore glass—it generates fine particles that can clog frits or valves in DNA synthesizers. PREPARATION OF 3′-AMINOALKYL-FUNCTIONALIZED DNA OLIGONUCLEOTIDES
BASIC PROTOCOL
The synthesis of an anhydride-functionalized CPG that can be used in the preparation of 3′-aminoalkyl, 3′-thioalkyl, or 3′-hydroxyalkyl DNA is delineated. The preparation of a support for 3′-aminoalkyl DNA synthesis is then described, followed by the synthesis and purification of a 3′-fluorescein-DNA conjugate. Materials 1000 Å aminopropyl-conjugated controlled-pore glass (aminopropyl-CPG; Biosearch Technologies) Pyridine Methylene chloride Trimellitic anhydride chloride (Aldrich) 100% and 20% (v/v) acetonitrile 6-Amino-1-hexanol N,N-Dimethylformamide (DMF) 4,4′-Dimethoxytrityl chloride (DMTr chloride; Aldrich) 3% (v/v) dichloroacetic acid in methylene chloride 28% (v/v) aqueous ammonia 5- and 6-Carboxy fluorescein succinimide ester (Molecular Probes) Dimethyl sulfoxide (DMSO) 1 M sodium carbonate/1 M sodium bicarbonate solution 0.05 M aqueous ammonium acetate Contributed by Matthew H. Lyttle Current Protocols in Nucleic Acid Chemistry (2000) 4.6.1-4.6.8 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Modified Oligonucleotides and Conjugates
4.6.1 Supplement 3
Sephadex G-25 resin (Amersham Pharmacia Biotech) 1 M triethylammonium acetate (TEAA) 14% (v/v) aqueous ammonia (1 part concentrated ammonium hydroxide, 1 part water) 2.8% (v/v) aqueous ammonia (1 part concentrated ammonium hydroxide, 9 parts water) Buffer A: 0.025 M Tris⋅Cl and 0.01 M Tris base Buffer B: 0.025 M Tris⋅Cl, 0.01 M Tris base, and 1 M NaCl 125-mL Erlenmeyer flasks with stoppers 150-mL sintered glass funnels, coarse frit 1-liter side-arm filter flask with rubber gasket Water aspirator Heavy-walled vacuum tubing 3-way valve Speedvac (Savant) Spectrophotometer and 1-cm path length glass cuvettes (e.g., Beckman) Automated DNA synthesizer and reagents (e.g., PE Biosystems) 1.5-mL plastic screw-cap tubes 55°C water bath Reversed-phase DNA purification cartridges and reagents (e.g., Biosearch Technologies) 1 × 30–cm glass tube with a frit and valve on bottom 10-mL syringes High-performance liquid chromatograph (HPLC) with anion-exchange column Additional reagents and equipment for DNA purification with a reversed-phase cartridge (UNIT 10.7) Synthesize an anhydride-functionalized CPG 1. Weigh out 5 g aminopropyl-CPG and place in a 125-mL Erlenmeyer flask with stopper. 2. In a separate 125-mL Erlenmeyer flask, mix together 10 mL pyridine and 40 mL methylene chloride. Stopper the flask. 3. Weigh out 1 g trimellitic anhydride chloride and add it to the solution prepared in step 2. Swirl solution until the solid completely dissolves. 4. Pour solution into the flask containing CPG. Swirl and allow to stand 1 hr at ambient temperature. 5. In the meantime, assemble a 150-mL sintered glass funnel, rubber gasket, and a 1-liter side-arm filter flask, and connect these to a water aspirator with a piece of heavywalled vacuum tubing. Install a 3-way valve between the flask and aspirator to control the vacuum and prevent water from being drawn into the flask when the aspirator is turned off. 6. Pour the slurry containing the CPG into the sintered glass funnel and remove the solvent by suction through the frit. Wash the CPG with three 50-mL portions of methylene chloride followed by three 50-mL portions of acetonitrile. 3′-Modified Oligonucleotides and their Conjugates
For each wash, completely suspend the CPG in the solvent with a spatula before applying vacuum to ensure efficient washing.
7. Dry the CPG in a vacuum desiccator for several hours.
4.6.2 Supplement 3
Current Protocols in Nucleic Acid Chemistry
Add hydroxylated aminohexyl linker to the anhydride-functionalized CPG 8. Weigh out 2 g of 6-amino-1-hexanol and dissolve in 10 mL DMF. 9. Add the solution to 2 g of the support obtained in step 7 and swirl briefly. Allow the slurry to stand for 3 hr at room temperature. 10. Repeat steps 6 and 7. Protect the linker hydroxy function with a DMTr group 11. Add 20 mL dry pyridine to the CPG obtained in step 10 and then add 1 g DMTr chloride. Swirl the mixture until the DMTr chloride dissolves, and allow to stand for at least 18 hr at room temperature. 12. Repeat steps 6 and 7. 13. To measure the amount of available hydroxyls on the support, now DMTr protected, weigh out 20 mg of the dry CPG material into a 125-mL Erlenmeyer flask and add exactly 100 mL of 3% dichloroacetic acid in methylene chloride. 14. Measure the absorbance of the orange solution at 498 nm, and multiply this number by the abbreviated extinction coefficient of DMTr (71.2). The result is the loading of the support, in micromoles per gram. Loading should be ≥25 mmol/g.
Synthesize DNA on aminohexyl-linked CPG 15. Calculate how many milligrams of the support obtained in step 12 will be required for a 0.2-µmol synthesis. Pack the calculated amount into a DNA synthesis column. The required amount of support should be between 3 and 10 mg.
16. Using an automated DNA synthesizer, add the desired sequence of nucleobases to the support. Remember that the 3′-terminal nucleobase must be added as an amidite, so use a dummy nucleobase in the sequence entry at the 3′ terminus. Leave the 5′-DMTr on when oligonucleotide synthesis is complete.
17. After synthesis, place the CPG containing the DNA in a 1.5-mL screw-cap tube and remove the DNA from the CPG by heating in 1 mL of 28% aqueous ammonia for 5 hr at 55°C. 5′-Terminal dG sequences are prone to detritylate at 55°C; these sequences give better results when deprotected for 48 hr at room temperature.
18. Allow the tube to cool to room temperature, and purify the DNA with a reversedphase cartridge according to the manufacturer’s instructions (see UNIT 10.7). Evaporate the purified DNA solution and then measure the absorbance at 254 nm in water. Conjugate 3′-aminoalkyl DNA oligonucleotides to 5- and 6-carboxyfluorescein 19. For each 10 OD254 units of 3′-aminohexyl DNA oligonucleotide measured, weigh out 1 mg of 5- and 6-carboxyfluorescein succinimide ester and dissolve in 100 µL DMSO. For >50 OD254 units of DNA, increase DMSO by 10 mL per additional 10 OD units.
20. Add this solution to a 1.5-mL plastic screw-cap tube containing the dried DNA, followed by 500 µL of 1 M sodium carbonate/1 M sodium bicarbonate solution. Close the tube and allow to stand 18 to 24 hr at room temperature. For >50 OD254 units of DNA, increase buffer by 100 mL per additional 10 OD units.
Synthesis of Modified Oligonucleotides and Conjugates
4.6.3 Current Protocols in Nucleic Acid Chemistry
Supplement 3
21. Evaporate the liquid in a Speedvac evaporator, and dissolve the residue in 1.5 mL of 0.05 M aqueous ammonium acetate. 22. Mix 20 g Sephadex G-25 resin with 100 mL of 0.05 M aqueous ammonium acetate, and pour the slurry into a 1 × 30–cm glass tube with a frit and valve at the bottom. Add enough of the slurry so that the settled bed volume is 20 to 25 cm long. Allow the liquid to elute from the column until the liquid is level with the Sephadex slurry. Use fresh Sephadex for each purification.
23. Add the DNA solution (step 21) to the column, and allow the liquid to elute from the column until the liquid is level with the Sephadex column. 24. Elute the column with 0.05 M aqueous ammonium acetate at a flow rate of ∼2 mL/min. Collect the first colored band that elutes after the first 10 to 15 mL. Evaporate the collected fraction in 1.5-mL tubes in a Speedvac. For many applications, the fluorescein-DNA oligonucleotide conjugate is pure enough for good results. Analysis of purity can be performed by PAGE (UNIT 10.4) or HPLC (UNIT 10.5). If necessary, purification can be performed using a reversed-phase cartridge (see below).
Purify fluorescein-DNA oligonucleotide conjugate 25. Preequilibrate a reversed-phase DNA purification cartridge by eluting with 4 mL acetonitrile followed by 4 mL of 1 M TEAA at a flow rate of ∼1 mL/min using a 10-mL syringe. 26. Dissolve the sample obtained in step 24 in 1 mL of 14% aqueous ammonia. Apply the solution to the cartridge. Collect the effluent and reload it onto the cartridge two times. 27. Elute the cartridge with 4 mL of 2.8% aqueous ammonia followed by 4 mL water at ∼1 mL/min. Discard noncolored eluant. 28. Elute the cartridge with 3 mL of 20% acetonitrile in water at ∼1 mL/min, and collect the strongly green effluent. 29. Evaporate in a Speedvac and store the purified 3′-fluorescein DNA oligonucleotide conjugate at −20°C until needed. Samples retain fluorescence for 2 to 3 months if kept in the dark.
Analyze conjugate by HPLC 30. Dissolve the sample in 500 µL of 20% acetonitrile in water, and inject 2 to 20 µL, depending on the concentration, onto an anion-exchange column. Elute the conjugate with a linear gradient of 100% buffer A to 100% buffer B over 20 min at a flow rate of 1 mL/min. The fluorescein conjugate will elute 2 to 4 min later than the underivatized 3′-aminohexyl DNA oligonucleotide. ALTERNATE PROTOCOL 1
3′-Modified Oligonucleotides and their Conjugates
PREPARATION OF 3′-THIOALKYL-FUNCTIONALIZED DNA OLIGONUCLEOTIDES This procedure uses many of the same reagents and steps as the Basic Protocol. However, the oxygen of the sulfur-bearing spacer must be protected, or else mixtures of products will be obtained after DNA synthesis and cleavage. A synthesis of these O-DMTr-protected thioalkyl spacers has been previously reported (Gupta et al., 1991).
4.6.4 Supplement 3
Current Protocols in Nucleic Acid Chemistry
Additional Materials (also see Basic Protocol) 6-Mercapto-1-O-DMTr hexanol (Biosearch Technologies) Dithiothreitol (DTT) Triethylamine Fluorescein-5-maleimide (Molecular Probes) Sodium phosphate/sodium chloride buffer: 50 mM sodium phosphate and 150 mM sodium chloride, adjust to pH 7.2 (if necessary) Synthesize 3′-thiohexyl DNA oligonucleotides on CPG 1. Prepare anhydride-functionalized CPG (see Basic Protocol, steps 1 to 7) and place 2 g in a 125-mL Erlenmeyer flask with stopper. 2. In a separate 125-mL flask, dissolve 1 g of 6-mercapto-1-O-DMTr hexanol in 10 mL DMF, and add 1 mL of triethylamine. 3. Add this solution to the Erlenmeyer flask containing CPG. Stopper the flask and swirl the CPG until there is a uniform slurry in the flask. Allow the flask containing the CPG to stand for 3 hr at room temperature. 4. Perform washing and drying as described (see Basic Protocol, steps 6 and 7). 5. Assay the loading of the spacer on the CPG (see Basic Protocol, steps 13 and 14). Loading should be ≥20 mmol/g.
6. Synthesize DNA oligonucleotides on the CPG, and deprotect and purify the product (see Basic Protocol, steps 15 to 18), but add 2 to 3 mg DTT to the 28% aqueous ammonia solution (step 17) before heating to prevent oxidative dimerization of the thioakyl oligonucleotides. Conjugate 3′-thiohexyl DNA oligonucleotides to fluorescein-5-maleimide 7. Weigh out 2 mg fluorescein-5-maleimide for every 10 OD254 units of DNA oligonucleotide to be conjugated in a 1.5-mL plastic screw-cap tube, and dissolve in 100 µl DMF. 8. Add this solution to a 1.5-mL plastic screw-cap tube containing the thiohexyl DNA. 9. Add 500 µL sodium phosphate/sodium chloride buffer, pH 7.2, and allow to stand 18 to 24 hr in the dark at room temperature. 10. Perform Sephadex G-25 size exclusion (see Basic Protocol, steps 21 to 24). Reversed-phase cartridge purification does not work well for thiomaleimido fluoresceinDNA oligonucleotide conjugates. Purification of the conjugates can be accomplished by either preparative HPLC (UNIT 10.5) or preparative PAGE (UNIT 10.4).
PREPARATION OF 3′-POLYETHYLENE-GLYCOL-FUNCTIONALIZED DNA OLIGONUCLEOTIDES
ALTERNATE PROTOCOL 2
Dry solvents and equipment are essential for the key reaction in this sequence, which is the addition of a hydroxyl functionality to an anhydride. This procedure uses many of the same reagents and steps as previous protocols. Various polyethylene glycols are commercially available; selection depends on the experimental design. The example given below uses triethylene glycol. Additional Materials (also see Basic Protocol) Triethylene glycol (Aldrich) N-Methylimidazole (Aldrich)
Synthesis of Modified Oligonucleotides and Conjugates
4.6.5 Current Protocols in Nucleic Acid Chemistry
Supplement 3
1. Prepare anhydride-functionalized CPG (see Basic Protocol, steps 1 to 4), and place 2 g in a 125-mL Erlenmeyer flask with stopper. 2. In a separate 125-mL flask, dissolve 1 g triethylene glycol in 10 mL DMF, and add 500 µL N-methylimidazole. 3. Add this solution to the Erlenmeyer flask containing CPG. Stopper the flask and swirl the CPG until there is a uniform slurry in the flask. Allow the flask containing the CPG to stand for 18 to 24 hr at room temperature. 4. Perform washing and drying (see Basic Protocol, steps 6 and 7). 5. Assay the loading of the triethylene glycol spacer on the CPG (see Basic Protocol, steps 11 through 14). Loading should be ≥20 mmol/g.
6. Synthesize DNA oligonucleotides on the PEGylated CPG, and deprotect and purify the product (see Basic Protocol, steps 15 to 18). 7. Analyze 3′-PEGylated DNA oligonucleotides by ion-exchange HPLC (see Basic Protocol, step 30). The desired product will elute slightly later than the corresponding unmodified DNA.
COMMENTARY Background Information
3′-Modified Oligonucleotides and their Conjugates
The basic strategy employed in these protocols (Lyttle et al., 1997) calls for the synthesis of an anhydride-functionalized solid support as a common intermediate for each functional group attachment. A bifunctional molecule (spacer) that has an OH group at one end and an SH, NH2, or OH group at the other end is then added. The most nucleophilic functional group reacts with the anhydride to form a thioester, amide, or ester bond, respectively, while the OH group at the other end of the spacer is available to react with nucleoside phosphoramidites (see Figure 4.6.1). There is then an optional step of adding a 4,4′-dimethoxytrityl (DMTr) group to this alcohol group to spectrophotometrically gauge the amount of addition of the first phosphoramidite during automated oligonucleotide synthesis. In the case of the thiol functionality, this DMTr alcohol protection is mandatory, or else a mixture of products (resulting from SH and OH addition to the anhydride) will be obtained. The synthesis of the required O-DMTr-protected hydroxylalkylthiols is described in the literature (Gupta et al., 1991). 6-O-DMTr-hydroxyhexylthiol is available from Biosearch Technologies. Once the support is made, automated oligonucleotide synthesis is performed to construct the desired sequence, then the usual aqueous ammonia treatment is employed for DNA
deprotection and solid-support cleavage to provide the desired 3′-terminal functionality upon basic hydrolysis of the thioester, amide, or ester bonds. The product DNA is then purified with reversed-phase cartridges, and can be characterized by PAGE, HPLC, and matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS). In the case of 3′-NH2- or SHmodified oligos, a procedure for the attachment of fluorescein is described, including an example of a successful reversed-phase cartridge purification of a 3′-amino-modified DNA-fluorescein conjugate.
Critical Parameters and Troubleshooting These protocols describe three techniques for the 3′-terminal labeling of DNA. Successful execution of each protocol depends on the skill and patience of the researcher, as well as the quality of reagents and solvents. The approximate expected loading is mentioned in each protocol; if loading is lower than expected, there are several points to check. For steps that require the immobilized anhydride, use a new bottle of trimellitic anhydride chloride for the coupling reaction. A negative ninhydrin test (Stewart and Young, 1984) will assure that the anhydride was quantitatively linked to the resin prepared according to the Basic Protocol. DMF must be dry (i.e., <0.05% water) and not have a noticeably fishy aroma;
4.6.6 Supplement 3
Current Protocols in Nucleic Acid Chemistry
O
NH2 N H
+ O
O O
O O
Cl
O O
H2N HS
OH or ODMTr / Et3N
or PEG / 1-Methylimidazole
O
O X(CH2)nOR OH
N H O DMTr, 4,4'-dimethoxytrityl; PEG, polyethylene glycol
Figure 4.6.1
1: X = NH, R = H, n = 6 2: X = S, R = DMTr, n = 6 3: X = O, X(CH2)nOR = PEG, R = H
An anhydride-functionalized solid support for the synthesis of 3′-modified DNA.
DMF slowly hydrolyzes to formic acid and dimethyl amine. The latter component has a fishy smell, and will have deleterious effects on the chemistry. Temperature of reactions should be close to 20°C; even a decrease of 10°C will slow down reaction rates. The DMTr chloride addition reactions are also moisture sensitive; therefore, use a freshly opened pyridine bottle (<0.05% water) and new DMTr chloride to improve the results if they are low. Finally, for the novice unfamiliar with organic chemical manipulations, the advice of a more experienced colleague can be indispensable.
Anticipated Results DNA fragments have been synthesized on three 3′-modification supports and compared to those DNA oligonucleotides synthesized from a conventional hemisuccinate nucleosidederivatized CPG. Stepwise coupling yields were similar in all cases. Yields of reversedphase cartridge purified DNA compared favorably with the amount normally obtained for cartridge purification at the 200-nmol and 1µmol synthesis scales. Anion exchange (AX)HPLC of 3′-modified 14-mers (5′-CCGAGTACTATTCA-3′) synthesized from the three CPGs showed that good quality products (i.e., 85% to 95% pure by AX-HPLC integration)
were obtained in the case of the 3′-thiohexyland 3′-triethylene-glycol-functionalized oligomers. In the case of the 3′-aminohexyl-derivatized 14-mer, a substantially slower eluting contaminant (20% to 30%) was present. The amount of this impurity can be decreased by prolonged heating (i.e., 24 to 48 hr) with aqueous ammonia, suggesting that this contaminant is DNA containing an unhydrolyzed amidemetallic linker at the 3′-terminus. This slowereluting contaminant did not interfere in subsequent reactions involving conjugation of the 3′-aminohexyl-functionalized oligonucleotide and was separated from the desired fluorescein conjugate by a reversed-phase cartridge purification. Due to the high loadings of the 3′-aminoalkyl CPG obtained with this procedure (i.e., 50 to 80 µmol/g), ample reactive material is still produced for subsequent conjugation steps. When the 3′-aminohexyl-linked 14-mer DNA fragment was prepared at a 200-nmol scale, ~75% conversion to the desired conjugate was obtained upon reaction with 50 equivalents of a mixture of fluorescein 5- and 6-carboxyhydroxysuccinimide active esters. The fluorescein conjugates elute 2 to 3 min slower by AX-HPLC than the unreacted 3′aminohexyl or thiolalkyl-modified DNA. The crude fluorescein conjugate could be purified
Synthesis of Modified Oligonucleotides and Conjugates
4.6.7 Current Protocols in Nucleic Acid Chemistry
Supplement 3
to 95% purity by the reversed-phase cartridge method described above. Better amine-fluorescein conjugation efficiency (>99%) was obtained at a 1-µmol synthesis scale, perhaps because of concentration considerations. Purified yields of 3′-fluorescein DNA at the 1-µmol synthesis scale should be 15 to 25 OD units. The corresponding 3′-thiolalkyl-modified DNA gave lower conjugation efficiencies (19% to 49%) even when reacted with 100 equivalents of fluorescein-5-maleimide. Cartridge purification of the thiol-linked fluorescein-DNA conjugates was also less effective. A 20% crosslinked polyacrylamide gel of 3′-conjugated DNA fragments that were prepared from both 3′-thiolalkyl and aminohexyl terminal modifications showed strongly fluorescent products exhibiting about the same gel mobility as that of underivatized DNA of the same length and sequence. The gel lanes of the thiolalkyl-linked conjugates contain some underivatized material, in agreement with HPLC analysis.
Time Considerations Most of the steps require a few hours at most and can be accomplished during an 8-hr day. The Sephadex columns usually require ≥4 hr for setup, chromatography, and analysis of the products. In general, only the immobilized an-
hydride-functionalized CPG cannot be stored for extended periods of time, and should be utilized as soon as possible. In all other steps involving a solid support, reagents should be washed away and the support washed with a neutral, volatile solvent before storage. When 18 to 24 hr of reaction time is recommended, it is best to mix the reagents for the reaction in the early evening (∼5 p.m.) and then work up the reaction mixture the following day.
Literature Cited Gupta, K.C., Sharma, P., Kumar, P., and Sathyanarayana, S. 1991. A general method for the synthesis of 3′-sulfhydryl and phosphate group containing oligonucleotides. Nucl. Acids Res.19:3019-3025. Lyttle, M.H., Adams, H., Hudson, D., and Cook, R.M. 1997. Versatile linker chemistry for synthesis of 3′-modified DNA. Bioconjug. Chem. 8:193-198. Stewart, J. and Young, J. 1984. Laboratory techniques in solid phase peptide synthesis. In Solid Phase Peptide Synthesis, pp. 105-107. Pierce Chemical Company, Rockford, Ill.
Contributed by Matthew H. Lyttle Biosearch Technologies, Inc. Novato, California
3′-Modified Oligonucleotides and their Conjugates
4.6.8 Supplement 3
Current Protocols in Nucleic Acid Chemistry
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates and their Phosphodiester and Phosphorothioate Chimeras
UNIT 4.7
Oligonucleotide N3′→P5′ phosphoramidates (pnODNs), wherein each 3′-oxygen is replaced by a 3′-amine in the 2′-deoxyribose ring (Figure 4.7.1), have shown significant therapeutic potential as antisense and antigene agents via duplex and triplex formation, respectively. Their binding properties also make them good candidates for oligonucleotide-based diagnostics and research tools. This unit describes the solid-phase synthesis of pnODNs (see Basic Protocol 1 and Alternate Protocol 1) using a method based on a phosphoramidite amine-exchange reaction wherein the key step is an exchange of a solid support–bound 3′-amino oligonucleotide for the amino group of a phosphoramidite monomer. Purification of these analogs is performed by either ion-exchange chromatography (IEC; see Basic Protocol 2) or reversed-phase high-performance liquid chromatography (RP-HPLC; see Alternate Protocol 2). The unit also describes the synthesis of required monomers and application of the method to preparation of phosphodiester- and phosphorothioate-containing chimeras of pnODNs. SOLID-PHASE CHAIN ASSEMBLY OF OLIGONUCLEOTIDE N3′→P5′ PHOSPHORAMIDATES FOR IEC PURIFICATION
BASIC PROTOCOL 1
The pnODN syntheses are performed in the 5′→3′ direction using a 3′-tritylamino nucleoside bound to aminopropyl controlled-pore glass (CPG) by a succinyl linker as the solid support (Figure 4.7.2). The trityl group was chosen as the amino-protecting group because it provides the requisite stability to the coupling, oxidation, and capping reagents, and is quantitatively removed by a relatively short treatment with dichloroacetic acid in dichloromethane. The resulting 3′-ammonium dichloroacetate salt is then coupled t o 3′-tritylamino-5′-O-[(cis-2,6-dimethylpiperidino)(2-cyanoethoxyl)]phosphinyl2′,3′-dideoxynucleoside monomer (Figure 4.7.3; S.1) in the presence of 1H-tetrazole. After neutralization, the resulting internucleotide phosphoramidite is oxidized to the stable, protected phosphoramidate. If phosphodiester or phosphorothioate chimeras are desired, the 3′-O-(4,4′-dimethoxytrityl)-5′-O-[(cis-2,6-dimethylpiperidino)(2-cyanoethoxy)]phosphinyl-2′-deoxyribonucleoside monomers (Figure 4.7.3; S.2) are used for those linkages, followed by either oxidation or sulfurization, respectively. A mixture of isobutyric anhydride and N-methylimidazole is used to cap any unreacted 3′-amines
5'
Base
O O O P O
Figure 4.7.1
O P S Base O
Base
O O
−
O
5'
O P O Base
Base
O NH
−
O
5'
− Base
O
O
O
3'
3'
Phosphodiester (po)
Phosphorothioate (ps)
3'
N3'→P5' Phosphoramidate (pn)
Structures of phosphodiester, phosphorothioate, and phosphoramidate linkages.
Contributed by Karen L. Fearon and Jeffrey S. Nelson Current Protocols in Nucleic Acid Chemistry (2000) 4.7.1-4.7.40 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Modified Oligonucleotides and Conjugates
4.7.1 Supplement 3
CPG
O
H N
O
O
O
O
H N
CPG
B1
O
Cl2CHCO2H
O
O CH2Cl2
NHTr
Cl2CHCOO
CH3 OCH2CH2CN P N O O
1. Isobutyric anhydride/NMI/2,6-lutidine/THF 2. Repeat synthesis cycle
B1
− +NH3
1H-Tetrazole CH3CN
B2
CH3 NHTr 1
CPG
O
H N
O
B1
O
O
O
H N
CPG
O
O
O
NH
O
O
NH
1. 20% C5H5N/CH3CN
O P OCH2CH2CN
P NCCH2CH2O
2. H2O2/H2O/C5H5N/THF
B2
B1
O
O
B2
NHTr NHTr
Figure 4.7.2 Synthetic steps in the preparation of pnODNs by the amine-exchange method. CPG, controlled-pore glass; Tr, triphenylmethyl; NMI, N-methylimidazole; THF, tetrahydrofuran.
because less hindered electrophiles, such as acetic anhydride, react with the tritylamino group of the oligonucleotide product. The method of purification needs to be decided on and planned for during the setup of the solid-phase synthesis. RP-HPLC purification of pnODNs using the hydrophobicity of the trityl group (Tr) is problematic because once the cyanoethyl groups are removed during ammonolysis, the phosphoramidate linkages are no longer stable to the acidic, post-RP-HPLC detritylation conditions. A new purification method requiring nonacidic conditions was therefore developed; however, it is only applicable to compounds terminating with a 3′-hydroxyl group. If a 3′-amine is desired, then IEC purification must be used. Chimera-containing phosphorothioate linkages are best purified by RP-HPLC because the phosphorothioate groups cause a severe loss of resolution during IEC. This
CH3 OCH2CH2CN
CH3 OCH2CH2CN N
P
O
O
B
P
N
CH3
1a, 2a: 1c, 2c: 1g, 2g: 1t, 2t:
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
O
O
B
CH3 NHTr
ODMTr
1
2
B = N 6-benzoyladenin-9-yl 4 B = N -benzoylcytosin-1-yl B = N 2-isobutyryl-O 6-(N,N-diphenylcarbamoyl)guanin-9-yl B = Thymin-1-yl DMTr = 4,4'-dimethoxytrityl
Figure 4.7.3
Structures of key phosphoramidite monomers.
4.7.2 Supplement 3
Current Protocols in Nucleic Acid Chemistry
protocol describes synthesis requiring IEC purification, whereas Alternate Protocol 1 describes synthesis requiring RP-HPLC purification. Materials 3% (v/v) dichloroacetic acid (Cl2CHCO2H) in dichloromethane (CH2Cl2; see recipe) Acetonitrile (CH3CN; ≤0.001% H2O) 0.1 M phosphoramidite monomer solutions (see recipe): 0.1 M 3′-tritylamino-5′-O-[(cis-2,6-dimethylpiperidino)(2cyanoethoxy)]phosphinyl-2′,3′-dideoxynucleoside monomers (ABz, CBz, Gi-Bu,DPC, T; S.1; see Support Protocol 7) 0.1 M 3′-O-(4,4′-dimethoxytrityl)-5′-O-[(cis-2,6-dimethylpiperidino)(2cyanoethoxy)]phosphinyl-2′-deoxyribonucleoside monomers (ABz, CBz, Gi-Bu,DPC, T; S.2; see Support Protocol 7; for phosphodiester or phosphorothioate chimera) 0.167 M 1H-tetrazole in CH3CN (see recipe) 20% (v/v) pyridine in CH3CN (see recipe) 1.5:3.5:20:75 (v/v/v/v) H2O2/H2O/pyridine/THF (see recipe) 0.2 M S-Tetra (Stec et al., 1993) in pyridine, prepared under argon, or 3H-1,2-benzodithiol-3-one-1,1-dioxide in acetonitrile (Beaucage reagent; Glen Research) 1:1:8 (v/v/v) isobutyric anhydride/2,6-lutidine/THF (see recipe) 16.5% (v/v) N-methylimidazole (NMI)/THF (see recipe) 3′-Tritylamino-2′,3′-dideoxynucleoside-5′-O-hemisuccinate conjugated to aminopropyl-controlled-pore glass (CPG; ABz, CBz, Gi-Bu,DPC, or T; see Support Protocol 6) Concentrated aqueous ammonia Column-mode DNA synthesizer capable of 1-µmol-scale syntheses (e.g., 392 or 394, PE Biosystems) with at least four monomer positions (preferably eight for synthesis of chimeras) Empty 1-µmol synthesis columns Desiccator with vacuum 4-mL glass screw-cap vials Heat block or oven set at 58°C Additional reagents and equipment for automated DNA synthesis (see manufacturer’s instructions and APPENDIX 3C) NOTE: This chemistry is extremely water sensitive. Oven-dry all bottles and syringes used for transferring solvents and solutions. Dissolve solids using a manifold or firestone valve to maintain an argon atmosphere. Perform all transfers under argon. Input oligonucleotide sequence 1. Enter the desired sequence into the automated synthesizer in backwards order. The pnODN syntheses are performed in the 5′-to-3′ direction instead of the 3′-to-5′ direction, which is standard for commercially available synthesizers. For instance, if the desired sequence is 5′-GGACCCTCCTCCGGAGCCOH-3′, then the synthesizer is programmed as follows: 5′-cCGAGGCCTCCTCCCAGG-3′, where lowercase letters represent 3′-O-dimethoxytrityl (3′-O-DMTr) monomers and uppercase letters represent 3′-trityl-amino monomers. If a pn/po/pn chimera of sequence 5′GGACCCpTpCpCpTpCpCGGAGCCOH-3′ is desired, where p is a phosphodiester linkage, then the synthesizer should be programmed as follows: 5′-cCGAGGCctcctcCCAGG-3′.
Synthesis of Modified Oligonucleotides and Conjugates
4.7.3 Current Protocols in Nucleic Acid Chemistry
Supplement 3
2. For synthesis of a pn/ps/pn chimera, program each flank to end with the first residue of the subsequent flank. If this is overlooked, the attaching residues at the pn/ps and ps/pn junctions will contain a thioamidate and a phosphodiester linkage, respectively. For instance, if the desired sequence is 5′-GGACCsCsTsCsCsTsCsCsGGAGCCOH-3′, where s is a phosphorothioate linkage and the other linkages are phosphoramidate linkages, then the synthesizer should be programmed and run three times as follows: 5′-cCAGG-3′
with oxidation
5′-Gcctcctcc-3′
with sulfurization
5′-cCGAGG-3′
with oxidation.
This assumes that the synthesizer used requires the inclusion of the base already on the support when programming the sequence.
Program synthesizer 3. Program the synthesizer to perform the following cycle: a. 3% Cl2CHCO2H in CH2Cl2 for 60 sec for tritylamino or 90 sec for O-DMTr, then CH3CN wash (six times with 0.5 mL). b. Phosphoramidite monomer solution (S.1 or S.2; 0.1 M; 15 eq) + 1H-tetrazole (0.167 M; 65 eq) in CH3CN for 5 min. c. Neutralize and wash with 20% pyridine in CH3CN (six times with 0.4 mL). d. 1.5:3.5:20:75 H2O2/H2O/pyridine/THF (0.65 mL; 2 min) or, for a phosphorothioate linkage, 0.2 M (50 eq) S-Tetra in pyridine (220 sec), followed by CH3CN washes (six times with 0.5 mL). e. 1:1:8 isobutyric anhydride/2,6-lutidine/THF (0.5 mL) + 16.5% NMI/THF(0.5 mL) for 2 min, then CH3CN washes (six times with 0.5 mL). f. Repeat steps a to e. g. Repeat step a (“trityl-off”). The trityl-off step is not performed when RP-HPLC purification of ODN is required. Phosphoramidite/tetrazole deliveries should be modeled after the instrument manufacturer’s protocol for delivering these reagents during phosphodiester synthesis. Modifications may need to be made to accommodate differences between flow synthesizers that only deliver reagents (e.g., PE Biosystems) and those that can recirculate phosphoramidite monomers (e.g., Amersham Pharmacia Biotech). The above details refer to the former (PE-ABI) synthesizer. N,N-Diisopropylamino phosphoramidite monomers can be used in place of cis-2,6-dimethylpiperidino phosphoramidite monomers with the following changes. A coupleoxidize-couple-oxidize protocol (Nelson et al., 1997) must be used for the couplings to 3′-amino groups (i.e., repeat steps b through d before proceeding to step e for capping). Couplings to 3′-hydroxyls require only a single coupling. Also, the concentration of tetrazole in acetonitrile must be increased to 0.5 M because N,N-diisopropylamino phosphoramidites require more tetrazole for activation than cis-2,6-dimethylpiperidinophosphoramidites (Hirschbein et al., 1998). 3′-O-(4,4′-Dimethoxytrityl)-5′O[(N,N-diisopropylamino)(2-cyanoethoxy)]phosphinyl-2′-deoxyribonucleoside monomers (ABz, CBz, Gi-Bu, T) are available from Glen Research. Using a vortexing-mode synthesizer (e.g., PE Biosystems 390Z) on the 10-mmol scale, 3.6 eq of monomer and 9 eq of 1H-tetrazole is sufficient for complete coupling. Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
Commercially available 3H-1,2-benzodithiol-3-one-1,1-dioxide (Beaucage reagent) in acetonitrile can be used in place of S-Tetra/pyridine for sulfurization, if desired. Use the vendor’s protocol for this reagent.
4.7.4 Supplement 3
Current Protocols in Nucleic Acid Chemistry
4. Calculate the amount of monomer necessary for the desired syntheses by multiplying the number of couplings of each type of base by 15 µmol/coupling. Add 100 µmol to the total for each base in order to have enough solution to prime the lines and to cover the bottom of the bottle. Prepare and add reagents 5. Dissolve the monomers to a concentration of 0.1 M in CH3CN as described (see Reagents and Solutions). Due to the stability and expense of the monomers, dissolve only as much monomer as will be used in 1 week. If using N,N-diisopropylamino phosphoramidites, 30 mmol of monomer is used per coupling to 3′-amino groups because the couple-oxidize-couple-oxidize protocol must be performed.
6. Formulate a sufficient amount of each auxiliary reagent to complete the desired syntheses. Consumptions on a per-coupling basis are typically 0.4 mL for the tetrazole solution, 0.65 mL for the hydrogen peroxide formulation, 0.5 mL each for the capping reagents, and 2.4 mL for the 20% pyridine/CH3CN solution.
7. Load the reagents on the instrument, keeping them under argon at all times. a. Place the four 3′-tritylamino-5′-O-[(cis-2,6-dimethylpiperidino)(2-cyanoethoxyl)]phosphinyl-2′,3′-dideoxynucleoside monomers (ABz, CBz, Gi-Bu,DPC, T) (S.1) in monomer positions 1 to 4 and, for chimeras, place the four 3′-O-(4,4′dimethoxytrityl)-5′-O-[(cis-2,6-dimethylpiperidino)(2-cyanoethoxy)]phosphinyl2′-deoxyribonucleoside monomers (ABz, CBz, Gi-Bu,DPC, T) (S.2) in monomer positions 5 to 8. b. Place the hydrogen peroxide solution in the oxidation position. c. Place the S-Tetra or Beaucage reagent in the sulfurization position. d. Place the capping agents in their usual positions on the instrument. e. Place 20% pyridine/CH3CN in the position usually used for concentrated aqueous ammonia or, if available, an extra position on the synthesizer. 8. Weigh 1 µmol of 3′-tritylamino-2′,3′-dideoxynucleoside-5′-O-hemisuccinate-CPG corresponding to the 5′-terminal base. Place the CPG in an empty 1-µmol synthesis column and check the column thoroughly for leaks on the synthesizer by passing acetonitrile through it. Run synthesizer 9. Prime all of the lines with reagent and start the synthesizer according to the manufacturer’s guidelines. 10. At the end of the synthesis, remove the CPG column from the synthesizer and dry the CPG in a desiccator under vacuum for 15 min. 11. Carefully open the column and transfer the CPG to a 4-mL glass screw-cap vial. 12. Add 1 mL concentrated aqueous ammonia and deprotect the oligonucleotide for 8 to 12 hr at 58°C. 13. Store the deprotected pnODN in the ammonia solution at −20°C until purification (maximum of 4 weeks).
Synthesis of Modified Oligonucleotides and Conjugates
4.7.5 Current Protocols in Nucleic Acid Chemistry
Supplement 3
ALTERNATE PROTOCOL 1
SOLID-PHASE CHAIN ASSEMBLY FOR RP-HPLC PURIFICATION Additional Materials (also see Basic Protocol 1) 0.1 M 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-3′-O[(N,N-diisopropylamino)(2-cyanoethoxy)]phosphinyl uridine (see recipe) 0.5 M 1H-tetrazole in CH3CN 3:1 (v/v) concentrated aqueous ammonia/ethanol 1. Program, set up, and run the synthesizer as described (see Basic Protocol 1, steps 1 through 9), with the following modifications: a. Ensure that the terminal coupling is with a 3′-O-DMTr-protected deoxyribonucleoside (S.2) in order to ultimately have a terminal 3′-hydroxyl group on the oligonucleotide. Also, do not remove the DMTr group at the end of the synthesis (step 3g). b. Place 0.1 M 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-3′-O[(N,N-diisopropylamino)(2-cyanoethoxy)]phosphinyl uridine solution on the instrument in one of the monomer positions. Also, replace the 0.167 M 1H-tetrazole solution with 0.5 M 1H-tetrazole solution. It is important to use 0.5 M 1H-tetrazole for this coupling because the uridine monomer is an N,N-diisopropylamino phosphoramidite, which requires more tetrazole for activation than the cis-2,6-dimethylpiperidino phosphoramidites.
c. Program the synthesizer to couple the uridine monomer to the 3′ terminus of the pnODN or chimera (see Basic Protocol 1), but extend the coupling time to 10 min (step 3b). Once again, make sure not to remove the DMTr group at the end of the synthesis. 2. At the end of the synthesis, dry the CPG-column and deprotect the oligonucleotide as described (see Basic Protocol 1, steps 10 to 12), but use 1 mL of 3:1 (v/v) concentrated aqueous ammonia/ethanol in step 12. Do not heat >18 hr. The deprotected pnODN can be stored in the ammonia solution at −20°C until purification for a maximum of 1 week, although it is best to remove the ammonia and purify the crude oligonucleotide as soon as possible after synthesis. BASIC PROTOCOL 2
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
IEC PURIFICATION, ISOLATION, AND ANALYSIS The IEC purification method is used for pnODNs synthesized with a terminal 3′-amine or 3′-hydroxyl following removal of the trityl protecting group. Preparative IEC is able to separate failure sequences from the full-length product (n-mer). However, the resolution between the n-1 failure sequence and the product is not great; therefore, the main product peak must be fractioned and analyzed by analytical IEC or capillary gel electrophoresis (CGE) in order to decide which fractions to combine. Materials Deprotected oligonucleotide solution (see Basic Protocol 1), 4°C Buffer A: 0.01 M aqueous NaOH/0.01 M NaCl, pH 12 Buffer B: 0.01 M aqueous NaOH/1.5 M NaCl, pH 12 Concentrated aqueous ammonia, 4°C 0.5 M aqueous NaOH solution 100% ethanol UV/visible spectrometer
4.7.6 Supplement 3
Current Protocols in Nucleic Acid Chemistry
Analytical IEC column (preferably a 4 × 250–mm Dionex PA-100 NucleoPac column) HPLC or FPLC system compatible with high pH buffer systems equipped with a UV detector, data collection system, and a 1- or 2-mL sample injection loop 3-mL disposable syringe with luer lock 0.45-µm filter that fits the end of a luer lock syringe Speedvac evaporator (Savant) Preparative IEC column (preferably a Pharmacia MonoQ 10/10 column) Sample holder with 1.5-mL centrifuge tubes or fraction collector Sephadex G-25 column (e.g., Pharmacia NAP-10; optional) Analyze quality of crude oligonucleotide 1. Dilute 10 µL cold, deprotected oligonucleotide solution in 990 µL water and scan from 200 to 400 nm using a UV spectrometer. Determine the absorbance at 260 nm and multiply by 100 (dilution factor) to determine the concentration of crude oligonucleotide in the 1 mL sample. To prevent loss of sample and to improve the accuracy of the dilution, make sure the ammonia solution is cold before opening the vial.
2. Dilute 0.5 OD260 units oligonucleotide into 0.5 mL water to serve as an analytical IEC sample. 3. Preequilibrate the analytical IEC column for ≥10 min with buffer A at a flow rate of 1 mL/min. 4. Program an HPLC or FPLC to run a gradient of 0% to 50% buffer B versus buffer A over 40 min at a flow rate of 1 mL/min and inject the 0.5-mL ODN sample. Monitor the run at 260 nm. Reequilibrate the column for ≥10 min with buffer A before performing a second run.
Prepare sample and IEC purify 5. Filter the CPG away from the remaining cold ammonia solution using a 3-mL disposable syringe with an attached 0.45-µm filter. 6. Wash the CPG twice with 0.5 mL cold, concentrated aqueous ammonia. 7. Add 10 µL of 0.5 M aqueous NaOH solution, and concentrate in a Speedvac evaporator to ∼0.5 mL. Do not completely dry the sample.
8. Filter the concentrated oligonucleotide again using a new 3-mL disposable syringe with an attached 0.45-µm filter and wash with 0.3 mL water. 9. Preequilibrate the preparative IEC column for ≥15 min with buffer A. Run a “blank” gradient if the column has not been used recently, and between purifications of samples with different sequences.
10. Program the HPLC to run a gradient ramping at 1%/min of buffer B versus buffer A at a flow rate of 1 mL/min. Use the analytical IEC (step 4) to determine the approximate percent of buffer B that will be needed to elute the sample. 11. Prepare a sample holder with at least ten 1.5-mL centrifuge tubes or use a fraction collector. This is necessary to obtain the highest level of purity; preparative IEC does not have the resolution of analytical IEC.
Synthesis of Modified Oligonucleotides and Conjugates
4.7.7 Current Protocols in Nucleic Acid Chemistry
Supplement 3
12. Inject the entire crude ODN sample from the 1-µmol synthesis and monitor elution at 260 nm. Collect ∼0.5-mL fractions during the elution of the product peak. Store the fractions at 4°C until they have been analyzed and are ready for desalting. Analyze and desalt 13. Analyze a small amount (i.e., ∼0.1 OD260) of the fractions by analytical IEC. 14. Combine the fractions that are ≥85% pure and concentrate in a Speedvac evaporator to a volume of ~1 mL. Do not let the samples evaporate completely in the Speedvac evaporator; NaOH can potentially degrade pnODNs.
15. Precipitate the pnODN with 2.5 mL of 100% ethanol and cool for ≥30 min at −20°C. 16. Centrifuge for 2 min at 3000 × g, 4°C, and carefully remove the supernatant. 17. Dissolve the pellet in 1 mL of deionized water and repeat ethanol precipitation (steps 15 and 16) two more times to desalt the sample. Alternatively, desalt the sample on a Sephadex G-25 column using the manufacturer’s protocol.
18. Dissolve pellet in 1 mL of water and measure the OD260 as in step 1 to determine the yield of pnODN. 19. Inject a 0.2 OD260 sample on the analytical IEC column to determine the purity of the product. Alternatively, determine the purity using capillary gel electrophoresis (CGE) or polyacrylamide gel electrophoresis (PAGE; APPENDIX 3B).
20. Concentrate the pnODN to dryness and store up to 1 year at −20°C. ALTERNATE PROTOCOL 2
REVERSED-PHASE HPLC PURIFICATION, ISOLATION, AND ANALYSIS The RP-HPLC method developed for the pnODNs and their chimeras relies on the 3′-addition of a commercially available RNA monomer, 5′-O-(4,4′-dimethoxytrityl)-2′-Otert-butyldimethylsilyl-3′-O-[(N,N-diisopropylamino)(2-cyanoethoxy)]phosphinyl uridine, to the terminal 3′-OH via 1H-tetrazole activation, followed by oxidation to a 3′→3′ phosphodiester linkage (Figure 4.7.4). The DMTr group is retained at the end of the synthesis to enable hydrophobic purification. After the RP-HPLC purification, the 3′-terminal uridine phosphodiester is cleaved from the oligonucleotide product by treatment with fluoride and base. Additional Materials (also see Basic Protocol 2) Deprotected oligonucleotide solution (see Alternate Protocol 1), 4°C Buffer C: acetonitrile Buffer D (see recipe): 0.1 M TEAB/2% acetonitrile, pH 8 3:1 (v/v) concentrated aqueous ammonia/ethanol 1 M TEAB buffer, pH 8 (see recipe) Acetonitrile 1 M aqueous NaF (0.45-µm filtered)
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
Analytical RP-HPLC column (e.g., Polymer Laboratories 0.46 × 15–cm PLRP-S column) HPLC system compatible with reversed-phase buffers and solvents, equipped with a UV detector, data collection system, and a 2-mL sample injection loop
4.7.8 Supplement 3
Current Protocols in Nucleic Acid Chemistry
1. DCA/CH2Cl2 2. DMTrO
O
OTBDMS
O
i-Pr2N
Ura
P OCH2CH2CN
DMTrO
Ura
O
DMTrO
Ura
O
1H-tetrazole/CH3CN 3'-DMTrOpnDNA-5'-CPG
1. Purify by RP-HPLC 2. 1 M aq. NaF/NH4OH, o 58 C, 16 hr
O P O
3. Desalt
−
+
−
O-3'-pnDNA-5'
Ura
O
OH
O O P O
O-3'-pnDNA-5'
DMTrO 3'-HOpnDNA-5'
+
OTBDMS
O 3. H2O2/H2O/C5H5N/THF o 4. NH3/EtOH, 58 C, 8-12 hr
DMTrO
Ura
O
+ O O P O
OH
−
OH
HO
O O P O
−
OH
Figure 4.7.4 Method facilitating the purification of oligonucleotide phosphoramidates by RPHPLC. DCA, dichloroacetic acid; Ura, uracil-1-yl; TBDMS, tert-butyldimethylsilyl; i-Pr, isopropyl.
Semipreparative RP-HPLC column (e.g., Polymer Labs 0.8 × 30–cm PLRP-S column) Heat block or oven set at 58°C Analyze quality of crude oligonucleotide 1. Optional: Follow the procedure for measurement of the crude OD260 and analysis by analytical IEC (see Basic Protocol 2, steps 1 to 4). 2. Dilute a second 0.5 OD260 units of ODN sample into 0.5 mL water for analysis by analytical RP-HPLC. 3. Preequilibrate the analytical RP-HPLC column for ≥10 min with 5% buffer C versus buffer D at a flow rate of 1 mL/min. 4. Program the HPLC to run a gradient of 5% to 40% buffer C versus buffer D over 40 min, followed by holding at 40% buffer C for 10 min at 1 mL/min. 5. Inject the 0.5-mL ODN sample and monitor the run at 260 nm. Reequilibrate the column for ≥10 min with 5% buffer C versus buffer D before performing a second run. Typical analytical IEC and RP-HPLC chromatograms of a crude pnODN containing a hydrophobic 3′-terminal uridine phosphodiester are shown in Figure 4.7.5. There are two product peaks in the RP-HPLC chromatogram because of partial loss of the tert-butyldimethylsilyl (TBDMS) group from uridine. Add the two peaks together to determine the amount of ODN product present. The byproduct peak generated from diphenylcarbamoyl (DPC) deprotection of G is observed near the product at 260 nm.
Prepare ODN sample and RP-HPLC purify 6. Filter the CPG away from the remaining cold ammonia solution using a 3-mL syringe attached to a 0.45-µm filter. 7. Wash the CPG twice with 0.5 mL of 3:1 concentrated aqueous ammonia/ethanol. 8. Concentrate in a Speedvac evaporator to ~0.5 mL. Do not completely dry the sample.
Synthesis of Modified Oligonucleotides and Conjugates
4.7.9 Current Protocols in Nucleic Acid Chemistry
Supplement 3
Figure 4.7.5 (A) Analytical IEC chromatogram (40.1% pure) and (B) RP-HPLC chromatogram (42.9% pure) of the phosphoramidate oligonucleotide, 5′-CCCTCCTCCGGAGCCpUDMTr where p is a (3′,3′)-phosphodiester linkage. Two ODN product peaks are seen in the RP-HPLC because some of the TBDMS group on the 3′-terminal uridine is removed prematurely by ammonia treatment and/or subsequent workup. Peak 1, product containing uridine with 5′-O-DMTr but not 2′-O-TBDMS; peak 2, byproduct generated from diphenylcarbamoyl (DPC) deprotection of G; peak 3, product containing uridine with 5′-O-DMTr and 2′-O-TBDMS.
Do not add 0.5 M aqueous NaOH to this sample as the hydrophobic 3′-terminal uridine phosphodiester will cleave prematurely.
9. Filter the concentrated oligonucleotide again using a new 3-mL syringe attached to a 0.45-µm filter and wash twice with 0.3 mL of water. 10. Add 0.2 mL of 1 M TEAB buffer, pH 8, and 25 µL acetonitrile. The sample, once concentrated and buffered, should be purified within 12 hr. Occasionally the byproduct produced from DPC deprotection of G continues to precipitate out after filtration, especially if the sample is frozen; refilter the solution just prior to purification, if necessary, to prevent clogging of the column.
11. Preequilibrate a semipreparative RP-HPLC column for ≥15 min in 5% buffer C versus buffer D at a flow rate of 2 mL/min. Run a “blank” gradient if the column has not been used recently, and between purifications of samples with different sequences.
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
TEAB (buffer D) is used because, unlike the more commonly used triethylammonium acetate (TEAA), it remains basic during the post-RP-HPLC concentration and enables the isolation of pure pnODN without accompanying acid-mediated degradation.
12. Program the HPLC to run a gradient of 5% to 40% buffer C versus buffer D over 40 min, followed by a hold for 10 min at 40% buffer C at a flow rate of 2 mL/min.
4.7.10 Supplement 3
Current Protocols in Nucleic Acid Chemistry
13. Inject 75 to 120 OD260 crude pnODN and collect both product peaks. Monitor the chromatography at 296 nm for preparative runs. The byproduct peak generated from DPC deprotection of G is not observed at 296 nm; the higher wavelength is used to attenuate the peak height. In general, there are some impurities just prior to the products, as well as a backside shoulder; both of these should be avoided during collection of the major fractions. It is usually best to collect only to approximately half the highest UV reading on the backside of the peak because this region contains more short-mer impurities. An example of a semipreparative RP-HPLC chromatogram and the fractionation of the peaks is shown in Figure 4.7.6.
14. Combine the two product peaks, concentrate in the Speedvac evaporator until the sample can be transferred to a 4-mL screw-cap vial, and then concentrate the sample to dryness. It is not necessary to add NaOH to the fractions; the TEAB buffer will stay basic during the concentration.
Remove 3′-terminal uridine phosphodiester, desalt, and analyze 15. Add 200 µL concentrated aqueous ammonia and 200 µL of 1 M aqueous NaF to the dry pnODN, vortex the mixture until the pnODN is dissolved, and heat the sample for 12 to 16 hr at 58°C. The fluoride removes the TBDMS group and the base causes intramolecular cleavage of the uridine phosphodiester function.
16. Cool the pnODN solution and check a small aliquot (0.2 OD260) by analytical IEC for complete cleavage. The retention time of the cleaved product is shorter than that of the uridine-containing product, and the two early-eluting uridine byproducts (see Fig. 4.7.4) are present.
17. Concentrate the solution to ∼200 µL in the Speedvac evaporator and precipitate the pnODN with 0.6 mL of 100% ethanol to remove the bulk of NaF. Freeze the sample for ≥30 min at –20°C. 18. Centrifuge for 2 min at 10,000 × g, room temperature, and carefully remove the supernatant.
Figure 4.7.6 Semipreparative RP-HPLC chromatogram of a pnODN with the sequence 5′CCCTCCTCCGGAGCCpUDMTr where p is a (3′,3′)-phosphodiester linkage.
Synthesis of Modified Oligonucleotides and Conjugates
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19. Dissolve the product in 1 mL of water and desalt the sample on a Sephadex G-25 column using the manufacturer’s protocol. The Sephadex G-25 column is necessary in this case because ethanol precipitation does not remove the uridine byproducts.
20. Measure the OD units at 260 nm to determine the yield of pnODN. Inject 0.2 OD260 units on an analytical IEC column to determine the purity of the product. Alternatively, determine ODN purity by CGE or PAGE (APPENDIX 3B).
21. Concentrate the pnODN to dryness in the Speedvac evaporator and store up to 1 year at −20°C. SUPPORT PROTOCOL 1
SYNTHESIS OF 3′-TRITYLAMINO-2′,3′-DEOXYTHYMIDINE The synthesis of 3′-tritylamino-2′,3′-deoxythymidine (S.5) from thymidine is shown in Figure 4.7.7. The 3′-azido-5′-O-(4-methoxybenzoyl)-2′,3′-deoxythymidine (S.3) is prepared as previously reported (Czernecki and Valéry, 1991). Materials Thymidine N,N-Dimethylformamide (DMF) Triphenylphosphine p-Anisic acid Diisopropylazodicarboxylate (DIAD) Diethyl ether, 5°C Lithium azide (LiN3) Ethyl acetate Saturated aqueous NaCl Sodium sulfate (Na2SO4, anhydrous) 8:2 (v/v) ethyl acetate/hexane 1:1 (v/v) ethanol/dichloromethane (CH2Cl2) Hydrogen 10% Pd/C catalyst (Aldrich) Pyridine (anhydrous) Triethylamine Trityl chloride Chromatography-grade silica gel, 70-230 mesh 60 Å (Aldrich) 0.5% to 5% (v/v) triethylamine in 2% (v/v) methanol/CH2Cl2 2% to 5% (v/v) methanol/CH2Cl2 5:95, 1:9, and 2:8 (v/v) methanol/CH2Cl2 57:43 (v/v) 1,4-dioxane/methanol 2 M aqueous NaOH (APPENDIX 2A) Dowex 50W-X8 cation-exchange resin (pyridinium H+ form; see recipe) Saturated aqueous NaHCO3 1:1 (v/v) ethyl ether/hexane
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
Heating mantle, variac, and temperature controller Mechanical overhead stirrer (Fisher) TLC plates (e.g., 0.2-mm-thick precoated Merck silica gel 60 F254 plates) Rotary evaporator Parr shaker-type hydrogenator able to hold pressures up to 60 psi
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O HO
O
Thy
2. DIAD/Ph3P/DMF 3. LiN3/DMF, 110 oC
OH
O
1. DIAD/Ph3P/4-MeOC6H4CO2H
O
Thy
MeO
1. H2,10% Pd/C, EtOH/CH2Cl2 2. TrCl/Et3N/C5H5N
N3 3 (98%)
T
CH3 OCH2CH2CN O
N O
O
MeO
Thy
2 M aq. NaOH
HO
O
Thy
1,4-Dioxan/MeOH NHTr 4 (83%)
NHTr
P Cl CH3 1t
DBU/CH2Cl2
5 (90%)
Figure 4.7.7 Synthetic steps in the preparation of 3′-tritylamino-2′,3′-dideoxythymidine (S.5) from thymidine. Thy, thymin-1-yl; DIAD, diisopropylazodicarboxylate; DBU, 1,8-diazabicyclo[5.4.0]undec7-ene.
Additional reagents and equipment for thin-layer chromatography (TLC, APPENDIX 3D) and column chromatography (APPENDIX 3E) Synthesize 3′-azido-5′-O-(4-methoxybenzoyl)-2′,3′-deoxythymidine (S.3) 1. Dissolve 141.0 g (582.2 mmol) thymidine in 1125 mL DMF in a 3-liter round-bottom flask. 2. Add 183.2 g (698.4 mmol) triphenylphosphine and 106.3 g (698.4 mmol) p-anisic acid. 3. Add 137.5 mL (698.4 mmol) diisopropylazodicarboxylate, diluted in 150 mL DMF, over 35 min using an additional funnel. 4. After 40 min, add another 183.2 g (698.4 mmol) triphenylphosphine all at once, and another 141.3 g (698.4 mmol) diisopropylazodicarboxylate, dissolved in 150 mL DMF, over a 40-min period using an additional funnel. 5. Stir the resultant mixture for an additional 65 min. 6. Quench the reaction with 10 mL of water and concentrate the solution to a volume of ∼800 mL using a rotary evaporator equipped with a vacuum pump. 7. Precipitate the product by pouring it into 6000 mL cold diethyl ether, 5°C, with rapid stirring. 8. Filter the solid using a Büchner funnel and house vacuum, wash with 2000 mL cold diethyl ether, and dry in vacuo to give semipure 2,3′-anhydro-5′-O-(4-methoxybenzoyl)-2′,3′-deoxythymidine. Crude yield = 110% (230.2 g).
9. Using a mechanical stirrer, dissolve 230.0 g (642.5 mmol) of 2,3′-anhydro-5′-O-(4methoxybenzoyl)-2′,3′-deoxythymidine in 1200 mL DMF, add 47.2 g (963.8 mmol) lithium azide, and heat the mixture for 48 hr at 100° to 110°C. 10. Concentrate the solution on the rotary evaporator using a vacuum pump, dissolve the residue in 3000 mL ethyl acetate, and wash two times with 1000 mL water and three times with 600 mL saturated aqueous NaCl.
Synthesis of Modified Oligonucleotides and Conjugates
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11. Dry the organic solution over anhydrous Na2SO4, filter, and concentrate on the rotary evaporator with a vacuum line to afford 3′-azido-5′-O-(4-methoxybenzoyl)-2′,3′-deoxythymidine (S.3) as an amber foam. 12. Perform TLC analysis (APPENDIX 3D) on 0.2-mm-thick precoated Merck silica gel 60 F254 plates to confirm the purity of the product. Elute with 8:2 (v/v) ethyl acetate/hexane. Crude yield = 97.9% (228.9 g, 570.0 mmol). Rf (8:2 ethyl acetate/hexane) = 0.51. 1H NMR (CDCl3 /TMS): δ 9.36 (1H, br s), 7.98 (2H, d, J = 8.7 Hz), 7.22 (1H, s), 6.95 (2H, d, J = 8.7 Hz), 6.18 (1H, t, J = 6.5 Hz), 4.65 (1H, dd, J = 12.3, 3.3 Hz), 4.53 (1H, dd, J = 12.3, 3.6 Hz), 4.35 (1H, m), 4.21 (1H, dt, J = 3.6, 2.5 Hz), 3.87 (3H, s), 2.54 (1H, m), 2.35 (1H, m), 1.71 (3H, s).
Synthesize 5′-O-(4-methoxybenzoyl)-3′-tritylamino-2′,3′-deoxythymidine (S.4) 13. Dissolve 10.0 g (24.9 mmol) of S.3 in 500 mL of 1:1 (v/v) ethanol/CH2Cl2 and reduce via hydrogenation (60 psi H2) in the presence of 1 g of 10% Pd/C catalyst for 16 hr. 14. Remove the catalyst by vacuum filtration and evaporate the solvent in vacuo. Yield = 92% (8.6 g, 22.9 mmol) of the corresponding 3′-amine, which is taken directly to the next reaction.
15. Dry 8.6 g (22.9 mmol) of 3′-amino-5′-O-(4-methoxybenzoyl)-2′,3′-deoxythymidine by azeotropic removal of water two times with 50 mL pyridine, evaporating to dryness on a rotary evaporator each time. Dissolve in 50 mL anhydrous pyridine. 16. Add 6.17 mL (48.1 mmol) triethylamine and 7.0 g (25.2 mmol) trityl chloride. Stir this mixture for 2 hr at room temperature. Perform TLC analysis (see step 12), eluting with 5:95 (v/v) methanol/CH2Cl2, to determine if the reaction is complete 17. If the reaction is complete, go to step 18, otherwise add an additional 1.9 g (6.9 mmol) trityl chloride, if necessary, and continue stirring for an additional 2 hr. 18. Remove the solvents using a rotary evaporator equipped with a vacuum pump and purify the crude product (S.4) by gravity on a silica gel column (APPENDIX 3E) preequilibrated with 0.5% triethylamine in 2% methanol/CH2Cl2. Obtain product by eluting with 2% to 5% methanol/CH2Cl2. For acid-sensitive (trityl-containing and phosphoramidite) compounds, the column should be packed and equilibrated with 0.5% triethylamine.
19. Perform TLC analysis (step 12) to confirm the purity of the product. Elute with 5:95 (v/v) methanol/CH2Cl2. Yield = 90% (12.7 g, 20.6 mmol). Rf (5:95 methanol/CH2Cl2) = 0.45. 1H NMR (CDCl3 /TMS): δ 8.26 (1H, br s, exchanges with D2O), 7.84, 6.90 (4H, AB, J = 8.86 Hz), 7.55 (6H, d, J = 7.45 Hz), 7.29 (6H, t, J = 7.56 Hz), 7.21 (3H, t, J = 7.27 Hz), 7.02 (1H, s), 6.08 (1H, t, J = 6.19 Hz), 4.59 (1H, dd, J = 12.35, 2.35 Hz), 4.29 (1H, dd, J = 12.43, 3.94 Hz), 3.98 (1H, m), 3.88 (3H, s), 3.41 (1H, m), 1.97 (1H, br, exchanges with D2O), 1.65-1.75 (1H, m), 1.58 (3H, s), 1.30-1.40 (1H, m). HRMS (FAB+): calcd for [M + Cs]+, 750.1580, observed 750.1559.
Synthesize 3′-tritylamino-2′,3′-deoxythymidine (S.5) 20. Remove the 5′-O-anisoyl protecting group by dissolving 30.1 g (48.7 mmol) of S.4 in 150 mL of 57:43 (v/v) 1,4-dioxane/methanol, and then adding 73.1 mL (146.2 mmol) of 2 M aqueous NaOH. Stir the solution for 1.5 hr at room temperature. Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
21. Neutralize with ~150 g of Dowex 50W-X8 cation-exchange resin (dry pyridinium H+ form, 1.6 meq/g).
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22. Once the pH is neutral (∼10 min), filter the resin, wash three times with 40 mL of 2:8 methanol/CH2Cl2, and concentrate the crude product on the rotary evaporator using a vacuum line. 23. Dissolve the residue in 500 mL ethyl acetate and extract two times with 250 mL saturated aqueous NaHCO3, once with 250 mL water, and once with 250 mL saturated aqueous NaCl. 24. Dry the organic phase over anhydrous Na2SO4 and filter. 25. Remove the solvents using a rotary evaporator with a vacuum line and dissolve the resulting foam in 300 mL of 5:95 methanol/CH2Cl2. 26. Add this solution slowly to 1250 mL of a rapidly stirring mixture of 1:1 (v/v) diethyl ether/hexane to precipitate the pure 3′-tritylamino-2′,3′-deoxythymidine (S.5). 27. Perform TLC analysis (step 12) to confirm the purity of the product. Elute with 1:9 (v/v) methanol/CH2Cl2. Yield = 90% (21.2 g, 43.8 mmol). Rf (1:9 methanol/CH2Cl2) = 0.50. 1H NMR (CDCl3 /TMS): δ 8.30 (1H, br s, exchanges with D2O), 7.52 (6H, d, J = 7.46 Hz), 7.29 (6H, t, J = 7.55 Hz), 7.21 (3H, t, J = 7.25 Hz), 7.16 (1H, s), 6.01 (1H, t, J = 6.38 Hz), 3.85 (1H, d, J = 11.71 Hz), 3.74 (1H, m), 3.65 (1H, dd, J = 11.99, 2.59), 3.34 (1H, q, J = 6.54 Hz), 1.80-2.00 (1H, br, exchanges with D2O), 1.83 (3H, s), 1.45-1.55 (1H, m), 1.30-1.40 (1H, m). HRMS (FAB+): calcd for [M + Cs]+, 616.1212, observed 616.1226.
SYNTHESIS OF N4-BENZOYL-3′-TRITYLAMINO-2′,3′-DIDEOXYCYTIDINE The synthesis of N4-benzoyl-3′-tritylamino-2′,3′-deoxycytidine (S.9) is shown in Figure 4.7.8. The C monomer is more readily and efficiently synthesized by the dU→dC route, rather than by lithium azide ring opening of a 2,3′-anhydro-2′-deoxycytidine derivative (Reese and Skone, 1984; Nelson et al., 1997).
SUPPORT PROTOCOL 2
Additional Materials (also see Support Protocol 1) 2′-Deoxyuridine 4-Dimethylaminopyridine tert-Butyldimethylsilyl (TBDMS) chloride 2:1 (v/v) ethanol/CH2Cl2 1,2,4-Triazole Phosphorus oxychloride (POCl3) 1,4-Dioxane Benzoyl chloride Concentrated aqueous ammonia (28%), 4°C Tetrahydrofuran (THF) 1 M tetra-n-butylammonium fluoride (TBAF) in THF 1% (v/v) triethylamine in 30% to 50% (v/v) ethyl acetate/hexane Synthesize 3′-azido-5′-O-(tert-butyldimethylsilyl)-2′,3′-dideoxyuridine (S.6) 1. Dry 11.4 g (50 mmol) of 2′-deoxyuridine two times thoroughly via co-evaporation with 100 mL anhydrous DMF in vacuo. 2. Add 100 mL DMF followed by 8.36 mL (60 mmol) triethylamine, 0.31 g (2.5 mmol) of 4-dimethylaminopyridine, and 8.29 g (55.0 mmol) tert-butyldimethylsilyl chloride. Stir the reaction mixture for 1 hr at room temperature. 3. Dilute with 600 mL CH2Cl2 and extract three times with 200 mL water and once with 200 mL saturated aqueous NaCl.
Synthesis of Modified Oligonucleotides and Conjugates
4.7.15 Current Protocols in Nucleic Acid Chemistry
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O
O
NH
NH HO
O
N
O
1. TBDMS-Cl/cat. DMAP/Et3N/DMF
TBDMSO
2. DIAD/Ph3P/DMF 3. LiN3/ DMF/∆
OH
O
N
O
1. H2,10% Pd/C, EtOH/CH2Cl2 2. TrCl/Et3N/C5H5N
N3 6 (63%)
dU
NHBz
O
N
NH TBDMSO
O
N
O
1. POCl3/Et3N/1,2,4-triazole/MeCN 2. NH4OH/1,4-dioxan
TBDMSO
3. BzCl/C5H5N
NHTr
O
N
O TBAF/THF
NHTr
7 (85%)
8 (92%) NHBz N HO
O
N
O
CH3 OCH2CH2CN P N Cl CH3 1c DBU/CH2Cl2
NHTr 9 (88%)
Figure 4.7.8 Synthetic steps in the preparation of N4-benzoyl-3′-tritylamino-2′,3′-dideoxycytidine (S.9) from 2′-deoxyuridine. DMAP, 4-dimethylaminopyridine; Bz, benzoyl; TBAF, tetra-n-butylammonium fluoride.
4. Dry the organic layer over anhydrous Na2SO4, vacuum filter, and concentrate on a rotary evaporator using a vacuum pump. 5. Purify the resulting residue by gravity column chromatography (APPENDIX 3E) on silica gel with 2% to 10% methanol/CH2Cl2 to afford 5′-O-(tert-butyldimethylsilyl)-2′-deoxyuridine. Yield = 80% (13.7 g, 40.0 mmol).
6. Dissolve 13.7 g (40.0 mmol) of 5′-protected nucleoside and 16.8 g (64.0 mmol) triphenylphosphine in 100 mL DMF. While stirring, add 12.6 mL (64.0 mmol) diisopropylazodicarboxylate in 20 mL DMF. 7. Stir 2 hr at room temperature, concentrate the reaction mixture on a rotary evaporator using a vacuum pump to ∼30 mL, and pour into 1200 mL diethyl ether. The desired product precipitates out after ∼10 min of rapid stirring.
8. Place the resulting mixture overnight at 4°C. 9. Collect the precipitate by vacuum filtration, wash two times with 300 mL cold diethyl ether, and dry in vacuo to afford 2,3′-anhydro-5′-O-(tert-butyldimethylsilyl)-2′-deoxyuridine as a white solid. Yield = 90% (11.7 g, 36.0 mmol).
10. React 33.8 g (104.2 mmol) of 2,3′-anhydro-5′-O-(tert-butyldimethylsilyl)-2′-deoxyuridine with 7.65 g (156.3 mmol) LiN3 in 300 mL DMF for 48 hr at 95° to 100°C. Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
11. Cool the resulting brown homogeneous mixture to room temperature, and concentrate to an oil using a rotary evaporator and vacuum pump. 12. Dissolve the residue in 800 mL ethyl acetate and extract with 200 mL water.
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13. Extract the aqueous layer twice more with 75 mL ethyl acetate and combine the organics. Wash three times with 250 mL water and once with 250 mL saturated aqueous NaCl. 14. Dry the ethyl acetate solution over anhydrous Na2SO4, vacuum filter, and concentrate on a rotary evaporator with a vacuum line to afford 3′-azido-5′-O-(tert-butyldimethylsilyl)-2′,3′-dideoxyuridine (S.6) as a brownish foam. Proceed directly to hydrogenation. 15. Perform TLC analysis (APPENDIX 3D) on 0.2-mm-thick precoated Merck silica gel 60 F254 plates to confirm the purity of the product. Elute with 8:92 (v/v) methanol/CH2Cl2. Yield = 87% (33.2 g, 90.3 mmol). Rf (8:92 methanol/CH2Cl2) = 0.57. 1H NMR (CDCl3 /TMS): δ 8.87 (1H, br s, exchanges with D2O), 7.91 (1H, d, J = 8.10 Hz), 6.23 (1H, t, J = 5.88 Hz), 5.71 (1H, d, J = 8.18 Hz), 4.25 (1H, q, J = 5.91 Hz), 3.95-4.05 (2H, m), 3.83 (1H, dd, J = 11.40, 1.68 Hz), 2.45-2.55 (1H, m), 2.25-2.35 (1H, m), 0.95 (9H, s), 0.15 (3H, s), 0.14 (3H, s). HRMS (FAB+): calcd for [M + H]+, 368.1754, observed 368.1747.
Synthesize 5′-O-(tert-butyldimethylsilyl)-3′-tritylamino-2′,3′-dideoxyuridine (S.7) 16. Dissolve 33.2 g (90.3 mmol) of crude S.6 in 300 mL of 2:1 (v/v) ethanol/CH2Cl2 and reduce via hydrogenation (60 psi H2) in the presence of 3.0 g of 10% Pd/C catalyst for 18 hr. 17. Remove the catalyst by vacuum filtration and evaporate the solvent on a rotary evaporator using a vacuum line to afford the corresponding 3′-amine. Proceed directly to the next reaction. Yield = 99.4% (30.4 g, 89.8 mmol).
18. Azeotrope 30.4 g (89.8 mmol) of 3′-amino-5′-O-(tert-butyldimethylsilyl)-2′,3′ -dideoxyuridine two times with 300 mL pyridine and dissolve the solid in a mixture of 600 mL CH2Cl2 and 70 mL anhydrous pyridine. 19. Add 25.0 mL (179.6 mmol) triethylamine and 30.0 g (125.7 mmol) trityl chloride to this solution and stir for 2 hr at room temperature. 20. Purify (see Support Protocol 1, step 18) to afford 5′-O-(tert-butyldimethylsilyl)-3′tritylamino-2′,3′-dideoxyuridine (S.7). 21. Perform TLC analysis (step 15) to confirm the purity of the product. Elute with 8:2 (v/v) ethyl acetate/hexane. Yield = 85% (44.3 g, 75.9 mmol). Rf (8:2 ethyl acetate/hexane) = 0.58. 1H NMR (CDCl3/TMS): δ 8.24 (1H, br s, exchanges with D2O), 7.73 (1H, d, J = 8.25 Hz), 7.52 (6H, d, J = 7.78 Hz), 7.31 (6H, m), 7.23 (3H, t, J = 7.23 Hz), 6.21 (1H, t, J = 6.69 Hz), 5.60 (1H, d, J = 8.17 Hz), 3.84 (1H, m), 3.76 (1H, dd, J = 11.34, 2.00 Hz), 3.48 (1H, dd, J = 11.37, 2.27 Hz), 3.32 (1H, m), 2.07 (1H, br, exchanges with D2O), 1.60-1.70 (1H, m), 1.45-1.55 (1H, m), 0.84 (9H, s), 0.01 (3H, s), −0.05 (3H, s). HRMS (FAB+): calcd for [M + Na]+, 606.2764, observed 606.2751.
Synthesize N4-benzoyl-5′-O-(tert-butyldimethylsilyl)-3′-tritylamino-2′,3′-dideoxycytidine (S.8) 22. Add 22.5 mL (161.1 mmol) triethylamine dropwise over a period of 10 min to a stirring mixture of 11.1 g (161.1 mmol) of 1,2,4-triazole and 3.5 mL (37.1 mmol) phosphorus oxychloride in 125 mL anhydrous CH3CN at 0°C. 23. To this cold stirring mixture, add 9.4 g (16.1 mmol) of S.7 as a solution in 50 mL acetonitrile. Stir 2 hr at room temperature.
Synthesis of Modified Oligonucleotides and Conjugates
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24. Quench the reaction with 30 mL triethylamine and 10 mL water. 25. Remove the solvents using a rotary evaporator and vacuum line. 26. Dissolve the resulting brown solid in 250 mL CH2Cl2. Extract three times with 150 mL saturated aqueous NaHCO3 and once with 150 mL saturated aqueous NaCl. 27. Dry the organic solution over anhydrous Na2SO4, vacuum filter, and concentrate on a rotary evaporator using a vacuum line to afford 4-(1,2,4-triazol-1-yl)-5′-O-(tertbutyldimethylsilyl)-3′-tritylamino-2′,3′-dideoxyuridine as an orange solid. Crude yield = 100% (10.2 g, 16.1 mmol).
28. Dissolve this crude material in 200 mL of 1,4-dioxane and add 50 mL concentrated NH4OH, 4°C. 29. Stir the reaction mixture for 4 hr at room temperature and concentrate on a rotary evaporator using a vacuum pump to afford 5′-O-(tert-butyldimethylsilyl)-3′-tritylamino-2′,3′-dideoxycytidine as a beige solid. Crude yield = 100% (9.4 g, 16.1 mmol).
30. Azeotrope the crude material two times with 200 mL anhydrous pyridine. 31. Dissolve in 200 mL pyridine, cool externally in a 0°C ice bath, and add 2.2 mL (19.3 mmol) benzoyl chloride. 32. Allow the reaction to slowly warm to room temperature and stir an additional 16 hr at room temperature. 33. Cool the reaction mixture externally to 0°C and quench with 40 mL water. Stir 5 min. 34. Add 40 mL concentrated aqueous ammonia, 4°C, and stir the reaction mixture for an additional 15 min at 0°C. 35. Remove the solvents using a rotary evaporator and vacuum pump, dissolve the residue in 125 mL CH2Cl2, and extract three times with 75 mL saturated aqueous NaHCO3. 36. Dry the organic phase over Na2SO4, vacuum filter, and evaporate the solvents using a rotary evaporator and vacuum line. 37. Purify the crude material (see Support Protocol 1, step 18) to afford N4-benzoyl-5′O-(tert-butyldimethylsilyl)-3′-tritylamino-2′,3′-dideoxycytidine (S.8). 38. Perform TLC analysis (step 15) to confirm the purity of the product. Elute with 5:95 (v/v) methanol/CH2Cl2. Yield = 92% (10.2 g, 14.8 mmol). Rf (5:95 methanol/CH2Cl2) = 0.71. 1H NMR (CDCl3 / TMS): δ 8.70 (1H, d, J = 7.36 Hz, exchanges with D2O), 8.27 (1H, d, J = 7.36 Hz), 7.91 (2H, d, J = 7.46 Hz), 7.62 (1H, t, J = 7.20 Hz), 7.50-7.60 (8H, m; with 6H, d, J = 7.74 Hz at 7.52), 7.42 (1H, br d, J = 7.41 Hz), 7.30 (6H, t, J = 7.39 Hz), 7.22 (3H, t, J = 7.39 Hz), 6.26 (1H, t, J = 6.26 Hz), 3.80 (1H, br m), 3.77 (1H, br d, J = 11.39 Hz), 3.49 (1H, dd, J = 11.24, 2.33 Hz), 3.30 (1H, m), 1.90-2.10 (2H, br m; 1H exchanges in D2O), 1.52 (1H, dt, J = 13.57, 6.76 Hz), 0.86 (9H, s), 0.04 (3H, s), −0.01 (3H, s). HRMS (FAB+): calcd for [M + Cs]+, 819.2343, observed 819.2366.
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
4.7.18 Supplement 3
Current Protocols in Nucleic Acid Chemistry
Synthesize N4-benzoyl-3′-tritylamino-2′,3′-dideoxycytidine (S.9) 39. Remove the 5′-TBDMS protecting group by dissolving 2.0 g (2.85 mmol) S.8 in 15 mL THF and reacting it with 15 mL of 1 M TBAF in THF for 16 hr. 40. Concentrate the reaction mixture to a syrup on the rotary evaporator using a vacuum line and dissolve the residue in 25 mL CH2Cl2. Extract four times with 25 mL water and once with 25 mL saturated aqueous NaCl. 41. Dry the organic layer over Na2SO4, vacuum filter, and remove the solvent using the rotary evaporator and vacuum line. 42. Purify the crude product on a silica gel column preequilibrated with 1% triethylamine in 30% ethyl acetate/hexane, and elute with 30% to 50% ethyl acetate/hexane to afford N4-benzoyl-3′-tritylamino-2′,3′-dideoxycytidine (S.9). 43. Perform TLC analysis (step 15) to confirm the purity of the product. Elute with 5:95 (v/v) methanol/CH2Cl2. Yield = 88% (1.4 g, 2.50 mmol). Rf (5:95 methanol/CH2Cl2) = 0.55. 1H NMR (CDCl3 / TMS): δ 8.65 (1H, br s, exchanges with D2O), 8.19 (1H, d, J = 7.36 Hz), 7.87 (2H, d, J = 7.57 Hz), 7.62 (1H, t, J = 7.37 Hz), 7.47-7.57 (9H, m), 7.30 (6H, t, J = 7.50 Hz), 7.23 (3H, t, J = 7.24 Hz), 6.07 (1H, dd, J = 6.66, 4.31), 3.91 (1H, d, J = 12.00 Hz), 3.79 (1H, m), 3.73 (1H, d, J = 12.10 Hz), 3.30 (1H, q, J = 6.38 Hz), 1.80-2.00 (2H, m, 1 br H exchanges in D2O), 1.40 (1H, ddd, J = 13.89, 7.03, 4.41 Hz). HRMS (FAB+): calcd for [M + Na]+, 595.2321, observed 595.2310.
SYNTHESIS OF N2-ISOBUTYRYL-O6-(N,N-DIPHENYLCARBAMOYL)3′-TRITYLAMINO-2′,3′-DIDEOXYGUANOSINE
SUPPORT PROTOCOL 3
The synthesis of N2-isobutyryl-O6-(N,N-diphenylcarbamoyl)-3′-tritylamino-2′,3′-dideoxyguanosine (S.13) is depicted in Figure 4.7.9. The 3′-O-benzoyl-N2-isobutyryl- 2′-deoxyxyloguanosine (S.10) is synthesized as previously reported (Nishino et al., 1986; Herdewijn and Van Aerschot, 1989). Additional Materials (also see Support Protocol 1) N2-Isobutyryl-2′-deoxyguanosine Benzoyl chloride Trifluoromethanesulfonic anhydride 4-Dimethylaminopyridine tert-Butyldimethylsilyl chloride 1:1 (v/v) methanol/1,4-dioxane 1 M aqueous HCl Diethylazodicarboxylate Argon N,N-Diisopropylethylamine N,N-Diphenylcarbamyl chloride Triethylamine trihydrofluoride Toluene 7:3 and 6:4 (v/v) ethyl acetate/hexane 2-liter large-mouth Erlenmeyer flask
Synthesis of Modified Oligonucleotides and Conjugates
4.7.19 Current Protocols in Nucleic Acid Chemistry
Supplement 4
O N
O
NH N
HO
N
O
N
NHiBu
HO 2. (CF3SO2)2O/C5H5N/CH2Cl2, -15 oC 3. H2O
OH
NH
1. BzCl/C5H5N R O
dGiBu
N
N
NHiBu
10 R = OBz (35%) O N
1. TBDMS-Cl/DMAP/Et3N/DMF 2. 2 M NaOH, MeOH/dioxan (1:1 v/v), 5 oC
TBDMSO
3. LiN3/DEAD/Ph3P/DMF 4. H2,10% Pd/C, EtOH/CH2Cl2
O
N
NH N
NHiBu
TrCl/Et3N/C5H5N
NPh2
CH3 OCH2CH2CN P N Cl
NH2 11 (49%) O
O N R1O
O
N
O N
NH N
NHiBu 1. DPC-Cl/i-Pr NEt/C H N 2 5 5
HO
O
N
N N
NHiBu
CH3 1g
NHTr
2. Et3N•3HF/C5H5N/CH2Cl2
12 R1 = TBDMS (100%)
DBU/CH2Cl2
NHTr 13 (82%)
Figure 4.7.9 Synthetic steps in the preparation of N2-isobutyryl-O6-(N,N-diphenylcarbamoyl)-3′tritylamino-2′,3′-dideoxyguanosine (S.13) from N2-isobutyryl-2′-deoxyguanosine. DEAD, diethylazodicarboxylate; iBu, isobutyryl; DPC-Cl, N,N-diphenylcarbamyl chloride.
Synthesize 3′-O-benzoyl-N2-isobutyryl-2′-deoxyxyloguanosine (S.10) 1. Azeotrope 119.1 g (353.1 mmol) N2-isobutyryl-2′-deoxyguanosine two times with 600 mL pyridine and then add 1500 mL pyridine. The solid does not completely dissolve.
2. Add 45.1 mL (388.4 mmol) benzoyl chloride, dissolved in 250 mL pyridine, dropwise over 5 hr using an additional funnel, and then continue stirring for another 11 to 16 hr. 3. Quench the reaction with 20 mL water, then remove the solvent using a rotary evaporator and vacuum pump. 4. Dissolve the solid in 900 mL CH2Cl2, wash with 400 mL water, and then transfer the organic phase to a 2-liter large-mouth Erlenmeyer flask. 5. Add 400 mL water and stir vigorously to precipitate the white solid product. 6. Vacuum filter the solid and wash three times with 100 mL of water. 7. Dry the solid using a vacuum pump to obtain the 5′-O-benzoyl-N2-isobutyryl-2′-deoxyguanosine. Yield = 91.6% (142.7 g, 323.4 mmol). Rf (1:9 methanol/CH2Cl2) = 0.27.
8. Azeotrope 142.7 g (323.4 mmol) of 5′-O-benzoyl-N2-isobutyryl-2′-deoxyguanosine three times with 400 mL pyridine, and then add 240 mL pyridine and 2200 mL CH2Cl2. Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
9. Cool the mixture to −15°C and add, with stirring, 92.5 mL (549.7 mmol) trifluoromethanesulfonic anhydride dissolved in 500 mL CH2Cl2, dropwise over 1 hr, while maintaining the temperature between −10° and −15°C.
4.7.20 Supplement 4
Current Protocols in Nucleic Acid Chemistry
10. Perform TLC analysis (APPENDIX 3D) on 0.2-mm-thick precoated Merck silica gel 60 F254 plates to confirm that the reaction is complete. Elute with 1:9 (v/v) methanol/CH2Cl2 (Rf = 0.6). 11. When the reaction is complete (after ∼1.5 hr), slowly quench the reaction with 150 mL water, maintaining the reaction temperature below 10°C. 12. Allow the solution to warm to room temperature and continue stirring for an additional 16 hr to complete the inversion. 13. Wash the organic layer two times with 1000 mL water and once with 1000 mL saturated aqueous NaCl. 14. Dry the organic solution over anhydrous Na2SO4, vacuum filter, and concentrate on the rotary evaporator and vacuum line to a brown solid. 15. Dissolve the solid in 550 mL CH2Cl2, stir for 1 hr, and then cool overnight at 4°C. 16. Vacuum filter the white solid, wash with 750 mL cold CH 2Cl2, and then dry using a vacuum pump to afford 3′-O-benzoyl-N2-isobutyryl-2′-deoxyxyloguanosine (S.10). 17. Concentrate the mother liquor using a rotary evaporator and vacuum line and repeat the recrystallization to obtain a second crop of pure product. 18. Perform TLC analysis (step 10) to confirm the purity of the product. Combined yield = 38.6% (55.1 g, 124.9 mmol). Rf (1:9 methanol/CH2Cl2) = 0.27. 1H NMR (DMSO-d6): δ 12.04 (1H, s), 11.70 (1H, s), 8.16 (1H, s), 7.84 (2H, d, J = 7.79 Hz), 7.67 (1H, t, J = 7.54 Hz), 7.52 (2H, t, J = 7.69 Hz), 6.25 (1H, dd, J = 7.55, 2.00 Hz), 5.69 (1H, t, J = 4.3 Hz), 4.96 (1H, t, J = 5.49 Hz), 4.32 (1H, dt, J = 7.79, 5.88 Hz), 3.77 (2H, m), 3.00 (1H, m), 2.76 (2H, m), 1.11 (6H, d, J = 6.76 Hz). The product (S.10) coelutes by TLC with the 5′-O-benzoyl-N2-isobutyryl-2′-deoxyguanosine but can be distinguished by 1H NMR spectroscopy.
Synthesize 3′-amino-5′-O-(tert-butyldimethylsilyl)-N2-isobutyryl-2′,3′-dideoxyguanosine (S.11) 19. To a stirring solution of 4.86 g (11.0 mmol) S.10 in 20 mL DMF, add 3.4 mL (24.2 mmol) triethylamine, 54 mg (0.44 mmol) 4-dimethylaminopyridine, and 3.31 g (22.0 mmol) tert-butyldimethylsilyl chloride. 20. Stir the reaction for 2 hr at room temperature. 21. Add 10 mL methanol and, after stirring an additional 5 min, concentrate the reaction mixture on a rotary evaporator using a high vacuum. 22. Dissolve the residue in 150 mL CH2Cl2 and wash three times with 40 mL water and once with 60 mL saturated aqueous NaCl. 23. Dry the organic layer over Na2SO4, vacuum filter, and concentrate with a rotary evaporator using a vacuum line to afford 6.40 g (>100% crude yield) of 3′-O-benzoyl-5′-O-(tert-butyldimethylsilyl)-N2-isobutyryl-2′-deoxyxyloguanosine. 24. Dissolve the crude material in 100 mL 1:1 methanol/1,4-dioxane and cool to 5°C. 25. Add 44.0 mL (87.9 mmol) prechilled (5°C) 2 M aqueous NaOH and stir the reaction mixture for 15 to 20 min in an ice bath. Monitor this reaction carefully and neutralize the hydroxide as soon as possible in order to avoid loss of the isobutyryl group.
26. Neutralize the reaction with 97 mL of 1 M aqueous HCl to pH 6 to 7.
Synthesis of Modified Oligonucleotides and Conjugates
4.7.21 Current Protocols in Nucleic Acid Chemistry
Supplement 3
27. Remove the ice bath and concentrate the reaction mixture to ∼50 mL on the rotary evaporator with a high vacuum. 28. Extract three times with 75 mL CH2Cl2. 29. Wash the combined organics three times with 50 mL saturated aqueous NaHCO3 and two times with 50 mL saturated aqueous NaCl. 30. Dry the organic solution over Na2SO4, vacuum filter, and concentrate with a rotary evaporator using a vacuum line to afford 5′-O-(tert-butyldimethylsilyl)-N2-isobutyryl-2′-deoxyxyloguanosine. Proceed to the next reaction without further purification. Yield = 82% (4.1 g, 9.1 mmol).
31. To 47.3 g (104.7 mmol) crude 5′-O-(tert-butyldimethylsilyl)-N2-isobutyryl-2′-deoxyxyloguanosine dissolved in 1000 mL anhydrous DMF, add 15.4 g (314.1 mmol) LiN3 and 41.2 g (157.1 mmol) triphenylphosphine. 32. Add 24.7 mL (157.1 mmol) diethylazodicarboxylate and stir the reaction mixture for 5 hr at room temperature under argon. 33. Quench the reaction with 20 mL water and concentrate the reaction mixture on the rotary evaporator using a vacuum pump. 34. Dissolve the residue in 1500 mL ethyl acetate. 35. Wash three times with 1000 mL water and once with 1000 mL saturated aqueous NaCl. 36. Dry the organic solution over Na2SO4, vacuum filter, and concentrate using a rotary evaporator and vacuum line. Proceed directly to hydrogenation and purification of the 3′-amine. 37. Dissolve ≤104.7 mmol crude azide in 1600 mL of 1:1 (v/v) ethanol/CH2Cl2 and hydrogenate (60 psi H2) in the presence of 2.5 g of 10% Pd/C catalyst for 16 hr at room temperature. 38. Remove the catalyst by vacuum filtration and evaporate the solvent using a rotary evaporator and vacuum line to afford the crude 3′-amine. 39. Purify by gravity on a silica gel column (APPENDIX 3E) using 2% to 6% methanol/CH2Cl2 and then 1% triethylamine/6% methanol/CH2Cl2 to afford 3′-amino-5′O-(tert-butyldimethylsilyl)-N2-isobutyryl-2′,3′-dideoxyguanosine (S.11) as an off-white foam. 40. Perform TLC analysis (step 10) to confirm the purity of the product. Yield = 60% (28.2 g, 63.2 mmol). Rf (1:9 methanol/CH2Cl2) = 0.14. 1H NMR (CDCl3/TMS): δ 8.01 (1H, s), 6.17 (1H, dd, J = 6.77, 3.98 Hz), 3.80-3.90 (4H, mm), 2.83 (1H, septet, J = 6.80 Hz), 2.59 (1H, ddd, J = 13.26, 6.16, 4.03 Hz), 2.33 (1H, dt, J = 13.19, 6.79 Hz), 1.26 (6H, dd, J = 6.86, 2.79 Hz), 0.88 (9H, s), 0.07 (3H, s), 0.06 (3H, s). HRMS (FAB+): calcd for [M + H]+, 451.2489, observed 451.2480.
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
Synthesize 5′-O-(tert-butyldimethylsilyl)-N2-isobutyryl-3′-tritylamino-2′,3′-dideoxyguanosine (S.12) 41. Dissolve 28.5 g (63.2 mmol) S.11 in 500 mL pyridine, add 17.6 mL (126.4 mmol) triethylamine and 28.2 g (101.1 mmol) trityl chloride, and stir for 16 hr at room temperature.
4.7.22 Supplement 3
Current Protocols in Nucleic Acid Chemistry
42. Concentrate the reaction product to a solid using a rotary evaporator and vacuum pump. 43. Purify crude product (see Support Protocol 1, step 18) to afford 5′-O-(tert-butyldimethylsilyl)-N2-isobutyryl-3′-tritylamino-2′,3′-dideoxyguanosine (S.12). 44. Perform TLC analysis (step 10) to confirm the purity of the product. Yield = 100% (43.8 g, 63.2 mmol). Rf (1:9 methanol/CH2Cl2) = 0.72. 1H NMR (CDCl3 / TMS): δ 11.90 (1H, br s, exchanges with D2O), 8.01 (1H, br s, exchanges with D2O), 7.58 (1H, s), 7.56 (6H, d, J = 7.39 Hz), 7.31 (6H, t, J = 7.58 Hz), 7.23 (3H, t, J = 7.28 Hz), 6.00 (1H, dd, J = 6.86, 4.63 Hz), 3.88 (1H, dt, J = 5.91, 3.01 Hz), 3.75 (2H, ABX, JAB = 11.25 Hz), 3.52 (1H, m), 2.57 (1H, septet, J = 6.91 Hz), 2.00- 2.10 (1H, br s, exchanges with D2O), 1.72 (1H, dt, J = 13.63, 6.89 Hz), 1.59 (1H, ddd, J = 13.73, 6.69, 4.87 Hz), 1.28 (6H, dd, J = 6.90, 3.22 Hz), 0.81 (9H, s), −0.03 (3H, s), −0.04 (3H, s). HRMS (FAB+): calcd for [M + Cs]+, 825.2561, observed 825.2540.
Synthesize O6-(N,N-diphenylcarbamoyl)-N2-isobutyryl-3′-tritylamino-2′,3′-dideoxyguanosine (S.13) 45. Dissolve 30.3 g (43.7 mmol) S.12 in 90 mL anhydrous pyridine. 46. Add 11.4 mL (65.6 mmol) N,N-diisopropylethylamine and 11.1 g (48.1 mmol) N,N-diphenylcarbamyl chloride under argon and stir for 1.5 hr at room temperature. 47. Concentrate the intensely red/purple reaction mixture in vacuo (see step 42). 48. Dissolve the residue in 600 mL CH2Cl2, extract two times with 400 mL water and once with 400 mL saturated aqueous NaCl. 49. Dry the CH2Cl2 solution over Na2SO4, vacuum filter, and concentrate in vacuo (see step 36) to afford impure 5′-O-(tert-butyldimethylsilyl)-O6-(N,N-diphenylcarbamoyl)-N2-isobutyryl-3′-tritylamino-2′,3′-dideoxyguanosine. This product is generally taken on directly to desilylation, although it can also be purified on silica.
50. Perform TLC analysis (step 10) using 6:4 (v/v) ethyl acetate/hexane. Crude yield > 100% (43.8 g). Rf (6:4 ethyl acetate/hexane) = 0.58. 1H NMR (CDCl3 /TMS): δ 8.03 (1H, s), 7.90 (1H, br s, exchanges with D2O), 7.55 (6H, d, J = 7.63 Hz), 7.24-7.50 (16H, mm), 7.21 (3H, t, J = 7.22 Hz), 6.29 (1H, t, J = 6.07 Hz), 3.89 (1H, m), 3.75 (2H, ABX, JAB = 11.25 Hz), 3.49 (1H, br m), 3.01 (1H, br m), 2.77 (1H, septet, J = 6.78 Hz), 2.00-2.10 (br s, exchanges with D2O), 1.65-1.75 (2H, m), 1.28 (6H, d, J = 6.64 Hz), 0.83 (9H, s), −0.01 (3H, s), −0.02 (3H, s). HRMS (FAB+): calcd for [M + Cs]+, 1020.3245, observed 1020.3281.
51. Dissolve ∼43.7 mmol crude 5′-O-(tert-butyldimethylsilyl)-O6-(N,N-diphenylcarbamoyl)-N2-isobutyryl-3′-tritylamino-2′,3′-dideoxyguanosine in 200 mL CH2Cl2 and 25 mL pyridine. 52. Add 49.8 mL (305.8 mmol) triethylamine trihydrofluoride, followed by a 25-mL CH2Cl2 rinse, and stir the reaction mixture for 20 hr under argon at room temperature. 53. Dilute the reaction mixture with 600 mL CH2Cl2 and extract two times with 400 mL water. 54. Back-extract the first aqueous layer with 50 mL CH2Cl2. 55. Dry the combined organics over Na2SO4, vacuum filter, and concentrate in vacuo (see step 36).
Synthesis of Modified Oligonucleotides and Conjugates
4.7.23 Current Protocols in Nucleic Acid Chemistry
Supplement 3
56. Dissolve the residue in 100 mL CH2Cl2 and azeotrope three times with 50 mL toluene to remove traces of pyridine using a rotary evaporator and vacuum pump. 57. Purify on silica gel, using a column packed in 2% triethylamine in 7:3 (v/v) ethyl acetate/hexane and eluting with 7:3 ethyl acetate/hexane, to afford O6-(N,N-diphenylcarbamoyl)-N 2-isobutyryl-3′-tritylamino-2′,3′-dideoxyguanosine (S.13). 58. Confirm the purity of the product by TLC analysis (step 10), eluting with 6:4 (v/v) ethyl acetate/hexane. Yield = 82% (27.6 g, 35.7 mmol). Rf (6:4 ethyl acetate/hexane) = 0.20. 1H NMR (CDCl3 / TMS): δ 7.95 (1H, s), 7.84 (1H, br s, exchanges with D2O), 7.55 (6H, d, J = 7.85 Hz), 7.25-7.45 (16H, mm), 7.21 (3H, t, J = 7.25 Hz), 6.15 (1H, t, J = 6.31 Hz), 3.77-3.87 (2H, br m), 3.69 (1H, m), 3.62 (1H, m), 3.19 (1H, m), 2.80 (1H, septet, J = 6.86 Hz), 1.92-2.05 (2H, mm, 1H exchanges with D2O), 1.65 (1H, m), 1.24 (6H, d, J = 6.86 Hz). HRMS (FAB+): calcd for [M + Cs]+, 906.2380, observed 906.2350. Do not use 1 M tetra-n-butylammonium fluoride in THF to remove the TBDMS group because the O6-(N,N-diphenylcarbamoyl) group is not stable to this reagent. SUPPORT PROTOCOL 4
SYNTHESIS OF N6-BENZOYL-3′-TRITYLAMINO- 2′,3′-DIDEOXYADENOSINE The synthesis of N6-benzoyl-3′-tritylamino-2′,3′-dideoxyadenosine (S.18) from adenosine is illustrated in Figure 4.7.10. The 5′-O-(tert-butyldimethylsilyl)-2′-deoxyxyloadenosine is synthesized by slightly modified literature procedures (Wagner et al., 1974; Hansske and Robins, 1983), which is more efficient than the inversion route that is used for the pyroration of S.10. Additional Materials (also see Support Protocol 1) (−)-Adenosine Dibutyltin oxide p-Toluenesulfonyl chloride tert-Butyldimethylsilyl chloride 1 M lithium triethyl borohydride in THF Ammonium chloride Benzoyl chloride 7:10 (v/v) methanol/1,4-dioxane Pyridinium hydrochloride Diethylazodicarboxylate Argon 4:6 (v/v) ethyl acetate/hexane Tetrahydrofuran (THF) 1.0 M tetrabutylammonium fluoride (TBAF) in THF Synthesize 2′-O-(p-toluenesulfonyl)adenosine (S.14) 1. Gently reflux 25.0 g (93.5 mmol) (−)-adenosine and 23.3 g (93.5 mmol) dibutyltin oxide in 1000 mL methanol for 2 hr, until the cloudy suspension becomes clear. 2. Cool the solution to 4°C, then cautiously add 53.5 g (281 mmol) p-toluenesulfonyl chloride and 39.1 mL (281 mmol) triethylamine, keeping the reaction temperature at 4°C. 3. Stir the resulting cloudy suspension overnight at room temperature.
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
4. Vacuum filter the white solid, wash two times with 100 mL cold methanol, and dry using a vacuum pump to afford 2′-O-p-toluenesulfonyladenosine (S.14).
4.7.24 Supplement 3
Current Protocols in Nucleic Acid Chemistry
NH2 N HO
N
N
O
NH2 N
N
1. Bu2SnO/MeOH/reflux
HO
N
N
O
1. TBDMS-Cl/C5H5N
N
2. 1M LiEt3BH/THF, 4 oC 3. NH4Cl
o
HO
2. Ts-Cl/Et3N, 4 C
OH
HO
OTs
14 (100% crude)
A NH2 N R1O
R O
NHBz N
N
N
1. BzCl/C5H5N
N
R1O
R O
N
N
N
o
2. 2 M NaOH, 4 C MeOH/dioxan (7:10 v/v) 16 R = OH (84%) R1 = TBDMS
15 R = OH (59%) R1 = TBDMS
NHBz
NHBz N TBDMSO
O
N
1. LiN3/DEAD/Ph3P/DMF 2. H2,10% Pd/C, EtOH/CH2Cl2 3. TrCl/Et3N/C5H5N
N
N HO
N 1 M TBAF/THF
NHTr 17 (67%)
O
N
N N
CH3 OCH2CH2CN P N Cl CH3 1a
NHTr
DBU/CH2Cl2
18 (94%)
Figure 4.7.10 Synthetic steps in the preparation of N6-benzoyl-3′-tritylamino-2′,3′-dideoxyadenosine (S.18) from adenosine. Bu, butyl; Ts, p-toluenesulfonyl.
Crude yield = 114% (45 g). 1H NMR (DMSO-d6): δ 8.19 (1H, s), 8.02 (1H, s), 7.37 (2H, s), 7.42 (2H, d), 7.03 (2H, d), 6.11 (1H, d), 6.03 (1H, d), 5.75 (1H, t), 5.49 (1H, dd), 4.39 (1H, ddd), 4.07 (1H, br d), 3.62 (2H, m), 2.25 (3 H, s).
Synthesize 5′-O-(tert-butyldimethylsilyl)-2′-deoxyxyloadenosine (S.15) 5. Dissolve 45 g (< 93.5 mmol) S.14 in 1000 mL pyridine, add 28.2 g (187 mmol) tert-butyldimethylsilyl chloride, and stir overnight at room temperature. 6. Quench the reaction with 100 mL methanol, concentrate on a rotary evaporator using a vacuum pump, and azeotrope three times with 25 mL toluene. 7. Dissolve the solid in 200 mL methanol, stir for 2 hr to desilylate the N6-amino group, and concentrate with a rotary evaporator and vacuum line. 8. Purify the reaction product by gravity on a silica gel column (APPENDIX 3E) using 3% to 4% methanol/CH2Cl2 to give 5′-O-(tert-butyldimethylsilyl)-2′-O-(p-toluenesulfonyl)adenosine. Yield = 75.6% (37.9 g, 70.7 mmol). TLC (1:9 methanol/CH2Cl2) Rf = 0.51.
9. Dissolve 37.9 g (70.7 mmol) 5′-O-(tert-butyldimethylsilyl)-2′-O-(p-toluenesulfonyl)adenosine in 100 mL anhydrous THF and cool to 4°C. 10. Add dropwise 283 mL (283 mmol) prechilled 1.0 M lithium triethyl borohydride in THF. 11. Stir the solution for 30 min at 4°C and then overnight at room temperature. 12. Cool the solution to 4°C, carefully quench with 11.3 g (212 mmol) NH4Cl, and concentrate on the rotary evaporator with a vacuum line. 13. Dissolve the residue first in 50 mL methanol and then 500 mL diethyl ether. 14. Wash with 100 mL saturated aqueous NaCl, and concentrate to a foam (see step 12).
Synthesis of Modified Oligonucleotides and Conjugates
4.7.25 Current Protocols in Nucleic Acid Chemistry
Supplement 3
15. Purify on silica (2% to 4% methanol/CH2Cl2) to afford 5′-O-(tert-butyldimethylsilyl)-2′-deoxyxyloadenosine (S.15). Yield = 77.5% (20.1 g, 54.8 mmol). TLC (1:9 methanol/CH2Cl2) Rf = 0.43. 1H NMR (CDCl3 / TMS): δ 8.35 (1H, s), 7.98 (1H, s), 7.08 (1H, d, J = 9.34 Hz), 6.14 (1H, dd, J = 9.36, 2.68 Hz), 5.98 (2H, br s), 4.47 (1H, m), 4.10 (1H, m ), 3.97 (2H, m), 2.88 (1H, ddd, J = 15.47, 9.40, 6.32 Hz), 2.56 (1H, dd, J = 15.35, 2.76 Hz), 0.88 (9H, s), 0.063 (3H, s), 0.060 (3H, s). Make sure the reaction is fully quenched before concentrating the solution because lithium triethyl borohydride is pyrophoric. Other solvents, such as ethyl acetate or CH2Cl2, should not be used for the extraction as they lead to severe emulsions.
Synthesize N6-benzoyl-5′-O-(tert-butyldimethylsilyl)-2′-deoxyxyloadenosine (S.16) 16. Dissolve 5.0 g (13.6 mmol) S.15 in 25 mL pyridine, add 3.28 mL (27.4 mmol) benzoyl chloride, and stir for 2 hr at room temperature. 17. Quench the reaction with 1 mL water and concentrate on a rotary evaporator with vacuum pump. 18. Dissolve the residue in 80 mL of 7:10 (v/v) methanol/1,4-dioxane, cool to 4 °C. 19. Add 34 mL (68 mmol) prechilled 2.0 M aqueous NaOH and stir for 5 min to selectively remove the 3′-benzoyl group. Monitor the hydrolysis carefully and neutralize the hydroxide as soon as possible in order to avoid loss of the N6-benzoyl group.
20. Neutralize the solution to pH 7 with 4.0 g (35 mmol) pyridinium hydrochloride and concentrate on a rotary evaporator with vacuum pump. 21. Dissolve the residue in 100 mL CH2Cl2, extract two times with 50 mL saturated aqueous NaHCO3 and two times with 50 mL saturated aqueous NaCl, and concentrate in vacuo (see step 7). 22. Purify on silica (2% methanol/CH2Cl2) to give N6-benzoyl-5′-O-(tert-butyldimethylsilyl)-2′-deoxyxyloadenosine (S.16). Yield = 84.6% (5.4 g, 11.5 mmol). TLC (5:95 methanol/CH2Cl2) Rf = 0.38. 1H NMR (CDCl3 / TMS): δ 9.00 (1H, br s), 8.82 (1H, s), 8.32 (1H, s), 8.02 (2H, d, J = 7.26 Hz), 7.63 (1H, t, J = 7.41 Hz), 7.54 (2H, t, J = 7.56 Hz), 6.31 (1H, dd, J = 9.06, 2.44 Hz), 5.95 (1H, d, J = 7.63 Hz), 4.55 (1H, m), 4.11 (1H, pseudo q, J = 6.21 Hz), 4.02 (2H, dd, J = 8.08, 2.68 Hz), 2.90 (1H, ddd, J = 15.22, 9.12, 5.93 Hz), 2.58 (1H, dd, J = 15.33, 2.47 Hz), 0.88 (9H, s), 0.07 (3H, s), 0.06 (3H, s).
Synthesize N6-benzoyl-5′-O-(tert-butyldimethylsilyl)-3′-tritylamino-2′,3′-dideoxyadenosine (S.17) 23. Dissolve 17.6 g (37.5 mmol) S.16 in 375 mL anhydrous DMF and add 5.5 g (113.0 mmol) LiN3 and 14.8 g (56.3 mmol) triphenylphosphine. 24. Add 8.9 mL (56.3 mmol) diethylazodicarboxylate and stir the reaction mixture for 6 hr under argon at room temperature. 25. Quench the reaction with 10 mL water and concentrate in a rotary evaporator with a vacuum pump. Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
26. Dissolve the residue in 500 mL ethyl acetate, and wash three times with 300 mL water and once with 300 mL saturated aqueous NaCl.
4.7.26 Supplement 3
Current Protocols in Nucleic Acid Chemistry
27. Dry the ethyl acetate solution over Na2SO4, filter, and concentrate with a rotary evaporator and vacuum line. This crude (triphenylphosphine oxide–contaminated) 3′-azido-N6-benzoyl-5′-O-(tertbutyldimethylsilyl)-2′,3′-dideoxyadenosine (18.8 g) is taken on directly to hydrogenation and purified as the 3′-amine.
28. Dissolve 18.8 g of the crude azide in 250 mL of 1:1 (v/v) ethanol/CH 2Cl2 and reduce by hydrogenation (60 psi H2) in the presence of 1.0 g of 10% Pd/C catalyst for 16 hr at room temperature. 29. Remove the catalyst by vacuum filtration, and evaporate the solvent in a rotary evaporator with vacuum line to afford the crude 3′-amine. 30. Purify on silica (preequilibrate with 2% methanol/CH2Cl2 and elute with 2% to 6% methanol/CH2Cl2 and then 1% triethylamine/6% methanol/CH2Cl2) to afford 3′amino-N6-benzoyl-5′-O-(tert-butyldimethylsilyl)-2′,3′-dideoxyadenosine as an offwhite foam. 31. Perform TLC analysis (APPENDIX 3D) on 0.2-mm-thick precoated Merck silica gel 60 F254 plates to confirm the purity of the product. Elute with 8:92 (v/v) methanol/CH2Cl2. Yield = 68% (12.0 g, 25.6 mmol). Rf (8:92 methanol/CH2Cl2) = 0.30. 1H NMR (CDCl3 / TMS): δ 8.95 (1H, br s, exchanges with D2O), 8.81 (1H, s), 8.40 (1H, s), 8.02 (2H, d, J = 7.23 Hz), 7.62 (1H, t, J = 7.43 Hz), 7.54 (2H, t, J = 7.48), 6.49 (1H, dd, J = 6.81, 3.68 Hz), 3.80-3.98 (4H, mm), 2.76 (1H, ddd, J = 13.27, 6.42, 3.69), 2.39 (1H, dt, J = 13.43, 6.93), 0.92 (9H, s), 0.11 (3H, s), 0.00 (3H, s). HRMS (FAB+): calcd for [M + Cs]+, 601.1360, observed 601.1373.
32. Protect 29.5 g (63.0 mmol) of 3′-amino-N6-benzoyl-5′-O-(tert-butyldimethylsilyl)2′,3′-dideoxyadenosine by reacting with 12.9 mL (94.5 mmol) triethylamine and 21.1 g (75.6 mmol) trityl chloride in 350 mL CH2Cl2 for 16 hr at room temperature. 33. Dilute the reaction mixture with an additional 150 mL CH2Cl2. 34. Extract once with 400 mL water, three times with 300 mL saturated aqueous NaHCO3, and two times with 300 mL saturated aqueous NaCl. 35. Concentrate to a glassy foam with a rotary evaporator and vacuum line. 36. Purify the crude product on a silica gel column preequilibrated with 1% triethylamine in 4:6 (v/v) ethyl acetate/hexane, and elute with 4:6 ethyl acetate/hexane to afford N6-benzoyl-5′-O-(tert-butyldimethylsilyl)-3′-tritylamino-2′,3′-dideoxyadenosine (S.17). 37. Perform TLC analysis (step 33) using 5:95 (v/v) methanol/CH2Cl2 as the eluent. Yield = 98% (44.1 g, 62.1 mmol). Rf (5:95 methanol/CH2Cl2) = 0.48. 1H NMR (CDCl3 / TMS): δ 8.99 (1H, br s, exchanges with D2O), 8.76 (1H, s), 8.12 (1H, s), 8.00 (2H, d, J = 7.29 Hz), 7.60 (1H, t, J = 7.41 Hz), 7.54 (6H, d, J = 7.44 Hz), 7.51 (2H, t, J = 7.28 Hz), 7.28 (6H, t, J = 7.56 Hz), 7.20 (2H, t, J = 7.22 Hz), 6.36 (1H, t, J = 5.96 Hz), 3.90 (1H, m), 3.82 (1H, dd, J = 11.28, 2.74 Hz), 3.67 (1H, dd, J = 11.26, 3.10 Hz), 3.50 (1H, br m), 2.00-2.10 (1H, br s, exchanges with D2O), 1.68-1.83 (2H, mm), 0.82 (9H, s), −0.02 (3H, s), −0.03 (3H, s). HRMS (FAB+): calcd for [M + Cs]+, 843.2455, observed 843.2477.
Synthesize N6-benzoyl-3′-tritylamino-2′,3′-dideoxyadenosine (S.18) 38. Remove the 5′-TBDMS protecting group by dissolving 43.7 g (61.5 mmol) S.17 in 123.0 mL THF and reacting with 123.0 mL (123.0 mmol) of 1.0 M TBAF in THF for 24 hr, room temperature.
Synthesis of Modified Oligonucleotides and Conjugates
4.7.27 Current Protocols in Nucleic Acid Chemistry
Supplement 3
39. Concentrate the reaction mixture with vacuum line and dissolve the residue in 400 mL ethyl acetate. 40. Extract three times with 250 mL water and two times with 250 mL saturated aqueous NaCl. 41. Dry the organic layer over Na2SO4, vacuum filter, and remove the solvent with a rotary evaporator and vacuum line. 42. Purify the crude product on a silica gel column preequilibrated with 2% triethylamine in 8:2 (v/v) ethyl acetate/hexane, and elute with 8:2 ethyl acetate/hexane to 100% ethyl acetate to afford N6-benzoyl-3′-tritylamino-2′,3′-dideoxyadenosine (S.18). 43. Perform TLC analysis (step 33) using 5:95 (v/v) methanol/CH2Cl2 as the eluent. Yield = 94% (34.5 g, 57.9 mmol). Rf (5:95 methanol/CH2Cl2) = 0.40. 1H NMR (CDCl3 / TMS): δ 9.06 (1H, br s, exchanges with D2O), 8.68 (1H, s), 8.02 (1H, s), 8.01 (2H, d, J = 7.33 Hz), 7.60 (1H, t, J = 7.47 Hz), 7.53 (6H, d, J = 7.38 Hz), 7.51 (2H, t, J = 7.20 Hz), 7.29 (6H, t, J = 7.58 Hz), 7.21 (3H, t, J = 7.25 Hz), 6.24 (1H, dd, J = 7.56, 6.30 Hz), 4.85 (1H, dd, J = 9.87, 3.19 Hz, exchanges with D2O), 3 65-3.82 (3H, mm), 3.37 (1H, t, J = 10.16 Hz), 2.38 (1H, dt, J = 13.46, 7.02 Hz), 2.00-2.20 (1H, br s, exchanges with D2O), 1.75 (1H, ddd, J = 13.28, 5.96, 2.97 Hz). HRMS (FAB+): calcd for [M + Na]+, 619.2434, observed 619.2421. SUPPORT PROTOCOL 5
SYNTHESIS OF 3′-O-(4,4′-DIMETHOXYTRITYL)-PROTECTED DEOXYRIBONUCLEOSIDES An approach to the synthesis of 3′-O-(4,4′-dimethoxytrityl)-protected deoxyribonucleosides (S.20) is presented in Figure 4.7.11. These nucleosides are necessary for the synthesis of the phosphodiester or phosphorothioate portion of chimeric oligonucleotides. Additional Materials (also see Support Protocol 1) N6-Benzoyl-2′-deoxyadenosine N4-Benzoyl-2′-deoxycytidine N2-Isobutyryl-2′-deoxyguanosine Thymidine 4-Dimethylaminopyridine tert-Butyldimethylsilyl chloride Tetrahydrofuran (THF) 1 M tetrabutylammonium fluoride (TBAF) in THF N,N-Diisopropylethylamine
HO
O
1. TBDMS-Cl/DMAP/Et3N/C5H5N 2. DMTr-Cl/C5H5N
B
3. DPC-Cl/i-Pr2NEt/C5H5N iBu (only when B = G )
OH Bz
TBDMSO
Bz
B
DMTrO
iBu
B=A ,C ,G
O
, or T
19
CH3 OCH2CH2CN P N Cl HO
TBAF/THF (when B = ABz, CBz, T)
O
iBu,DPC
Et3N•3HF/C5H5N/CH2Cl2 (when B = G
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
)
B
CH3 2 DBU/CH2Cl2
DMTrO 20
Figure 4.7.11 General approach to the synthesis of 3′-O-(4,4′-dimethoxytrityl)-2′-deoxyribonucleosides (S.20) from N-protected 2′-deoxyribonucleosides.
4.7.28 Supplement 3
Current Protocols in Nucleic Acid Chemistry
N,N-Diphenylcarbamyl chloride 9:1 (v/v) CH2Cl2/pyridine Triethylamine trihydrofluoride 50% to 70% ethyl acetate/hexane Prepare 5′-O-(tert-butyldimethylsilyl)-2′-deoxyribonucleosides (S.19) 1. Azeotrope the N-protected 2′-deoxyribonucleoside (dABz, dCBz, dGi-Bu, or T) two times from 10 mL/g pyridine and suspend in pyridine at 10 mL/g. 2. To this stirring mixture, add sequentially 0.1 eq 4-dimethylaminopyridine, 1.2 eq triethylamine, and 1.05 to 1.2 eq tert-butyldimethylsilyl chloride. Stir for 8 to 24 hr at room temperature. 3. Remove the pyridine using a rotary evaporator and vacuum pump. 4. Dissolve the residue in 15 mL/g CH2Cl2 and extract two times with 10 mL/g water and one time with 10 mL/g saturated aqueous NaCl. 5. Dry the organic solution over anhydrous Na2SO4, vacuum filter, and concentrate under reduced pressure to a solid that is used in the next reaction without further purification. Prepare 5′-O-(tert-butyldimethylsilyl)-3′-O-(4,4′-dimethoxytrityl)-2′-deoxyribonucleosides 6. Azeotrope the 5′-O-(tert-butyldimethylsilyl)-protected 2′-deoxyribonucleoside two times from 10 mL/g pyridine and dissolve in pyridine at 10 mL/g. 7. While stirring this solution, add 1.2 to 1.3 eq 4,4′-dimethoxytrityl chloride. Stir the solution for 16 to 24 hr at room temperature. 8. Concentrate on a rotary evaporator using a vacuum pump. 9. Dissolve the residue in 15 mL/g CH2Cl2 and extract once each with 10 mL/g water, 10 mL/g saturated aqueous NaHCO3, and 10 mL/g saturated aqueous NaCl. 10. Dry the CH2Cl2 solution over Na2SO4, vacuum filter, and concentrate under reduced pressure to a foam. The product can be used directly in the next reaction (T) or purified on silica (dABz, dCBz, and dGi-Bu) using a gradient of 1% to 5% methanol in CH2Cl2.
Prepare 3′-O-(4,4′-dimethoxytrityl)-2′-deoxyribonucleosides (S.20) For dABz, dCBz, and T: 11a. Remove the 5′-TBDMS protecting group by dissolving the 5′-(tert-butyldimethylsilyl)-3′-O-(4,4′-dimethoxytrityl)-2′-deoxyribonucleoside in THF at 3 mL/g and reacting it with 1 M (2.0 eq) TBAF in THF for 16 to 24 hr, room temperature. 12a. Concentrate the solution under reduced pressure. 13a. Dissolve the residue in 15 mL/g CH2Cl2, and extract two times with 10 mL/g water and one time with 10 mL/g saturated aqueous NaCl. 14a. Dry the organic layer over Na2SO4, vacuum filter, and evaporate on a rotary evaporator with pump. 15a. Purify the crude 3′-O-(4,4′-dimethoxytrityl)-2′-deoxyribonucleosides (S.20) (see Support Protocol 1, step 18).
Synthesis of Modified Oligonucleotides and Conjugates
4.7.29 Current Protocols in Nucleic Acid Chemistry
Supplement 3
16a. Perform TLC analysis (APPENDIX 3D) on 0.2-mm-thick precoated Merck silica gel 60 F254 plates to confirm the purity of the product. Elute with 1:9 (v/v) methanol/CH2Cl2. N6-Benzoyl-3′-O-(4,4′-dimethoxytrityl)-2′-deoxyadenosine (S.20). Yield from dABz = 56.7% (55.4 g). Rf (1:9 methanol/CH2Cl2) = 0.68. 1H NMR (CDCl3/TMS): δ 9.14 (1H, br s, exchanges with D2O), 8.72 (1H, s), 8.06 (1H, s), 8.02 (2H, d, J = 7.43 Hz), 7.62 (1H, t, J = 7.33 Hz), 7.53 (2H, d, J = 7.74 Hz), 7.50 (2H, d, J = 7.54 Hz), 7.39 (4H, d, J = 8.81 Hz), 7.34 (2H, t, J = 7.54 Hz), 7.26 (2H, t, J = 7.95 Hz), 6.87 (4H, dd, J = 8.91, 2.43 Hz), 6.37 (1H, dd, J = 9.95, 5.26 Hz), 5.79 (1H, br d, J = 10.38 Hz, exchanges with D2O), 4.66 (1H, d, J = 5.32 Hz), 4.08 (1H, s), 3.81 (6H, s), 3.76 (1H, d, J = 12.78 Hz), 3.35 (1H, t, J = 11.86 Hz), 2.73 (1H, ddd, J = 13.21, 10.10, 7.99 Hz), 1.76 (1H, dd, J = 13.31, 5.30 Hz). HRMS (FAB+): calcd for [M + H] +, 658.2666; found, 658.2666. N4-Benzoyl-3′-O-(4,4′-dimethoxytrityl)-2′-deoxycytidine (S.20). Yield from dCBz = 74.7% (70.0 g) including additional mixed fractions that were purified further by precipitation from CH2Cl2 into a 20× volume of 3:1 hexane/diethyl ether over 1.5 hr. Rf (1:9 methanol/CH2Cl2) = 0.66. 1H NMR (CDCl3/TMS): δ 8.66 (1H, br s, exchanges with D2O), 8.09 (1H, d, J = 7.35 Hz), 7.87 (2H, d, J = 7.43 Hz), 7.62 (1H, t, J = 7.36 Hz), 7.53 (2H, d, J = 7.80 Hz), 7.48 (2H, d, J = 7.63 Hz), 7.37 (4H, d, J = 8.86 Hz), 7.32 (2H, t, J = 7.53 Hz), 7.25 (1H, t, J = 7.17 Hz), 6.85 (4H, d, J = 8.76 Hz), 6.25 (1H, dd, J = 7.63, 6.12 Hz), 4.36-4.43 (1H, br m), 3.94 (1H, d, J = 2.19 Hz), 3.81 (6H, s), 3.66 (1H, br d, J = 11.86 Hz), 3.26 (1H, br d, J = 11.90 Hz), 2.48 (1H, br s, exchanges with D2O), 2.22 (1H, dd, J = 13.13, 5.20 Hz), 2.08 (1H, quintet, J = 6.94 Hz). HRMS (FAB+): calcd for [M +Na]+, 656.2373; found, 656.2383. 3′-O-(4,4′-Dimethoxytrityl)-thymidine (S.20). Yield from T = 81.8% (45.2 g). Rf (1:9 methanol/CH2Cl2) = 0.56. 1H NMR (CDCl3 /TMS): δ 8.61 (1H, br s, exchanges with D2O), 7.46 (2H, d, J = 7.47 Hz), 7.36 (4H, d, J = 8.83 Hz), 7.32 (2H, t, J = 7.94 Hz), 7.25 (1H, t, J = 7.43 Hz), 6.86 (4H, d, J = 7.39 Hz), 6.15 (1H, dd, J = 8.87, 5.76 Hz), 4.38 (1H, d, J = 6.20 Hz), 3.99 (1H, d, J = 2.13 Hz), 3.81 (6H, s), 3.68 (1H, br d, J = 11.79 Hz), 3.30-3.37 (1H, br m), 2.47-2.55 (1H, br m, exchanges with D2O), 1.95 (1H, ddd, J = 13.98, 8.42, 6.00 Hz), 1.87 (3H, s), 1.67-1.74 (1H, m). HRMS (FAB+): calcd for [M + Na] +, 567.2107; found, 567.2111.
For dGi-Bu,DPC: 11b. Prepare a stirring solution of 101.9 g (135.2 mmol) 5′-O-(tert-butyldimethylsilyl)3′-O-(4,4′-dimethoxytrityl)-N2-isobutyryl-2′-deoxyguanosine in 300 mL pyridine, add 26.6 g (206.1 mmol) N,N-diisopropylethylamine and 41.4 g (178.7 mmol) N,N-diphenylcarbamyl chloride. 12b. Stir the dark solution for 2 hr and then concentrate on a rotary evaporator with a vacuum pump. 13b. Dissolve the residue in 600 mL CH2Cl2 and extract two times with 250 mL water and once with 250 mL saturated aqueous NaCl. 14b. Dry the organic solution over Na2SO4, vacuum filter, and concentrate to a purplecolored foam with a vacuum line on a rotary evaporator. 15b. Dissolve the crude nucleoside in 800 mL of a 9:1 (v/v) CH2Cl2/pyridine solution, then add 155.0 g (961.5 mmol) triethylamine trihydrofluoride and react for 16 hr at room temperature.
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
16b. Remove the solvents with a rotary evaporator and vacuum pump, dissolve the residue in 600 mL CH2Cl2, and wash two times with 250 mL water and 250 mL saturated aqueous NaCl. 17b. Dry the organic solution over Na2SO4, vacuum filter, and concentrate to a dark red foam with a vacuum line on a rotary evaporator.
4.7.30 Supplement 3
Current Protocols in Nucleic Acid Chemistry
18b. Purify the crude product by gravity on a silica gel column (APPENDIX 3E) preequilibrated with 1% Et3N/50% ethyl acetate/hexane and elute with 50% to 70% ethyl acetate/hexane to afford the protected 2′-deoxyguanosine (S.20). 19b. Perform TLC analysis (step 16a), eluting with 75:25 (v/v) ethyl acetate/hexane. Yield from dGi-Bu = 35.6% (43.3 g). Rf (75:25 ethyl acetate/hexane) = 0.38. 1H NMR (CDCl3/TMS): δ 8.00 (1H, s), 7.91 (1H, br s, exchanges with D2O), 7.49 (2H, d, J = 7.75 Hz), 7.20-7.45 (19H, mm with 4H, d, J = 8.93 Hz at 7.38), 6.86 (4H, dd, J = 8.82, 2.09 Hz), 6.26 (1H, dd, J = 9.80, 5.12 Hz), 4.65 (1H, d, J = 5.25 Hz), 4.35 (1H, dd, J = 10.35, 3.26 Hz, exchanges with D2O), 4.04 (1H, s), 3.80 (6H, s), 3.73 (1H, br d, J = 11.48 Hz), 3.39 (1H, br t, J = 11.54 Hz), 2.64-2.80 (2H, m), 1.68 (1H, dd, J = 13.18, 5.16 Hz), 1.24 (6H, d, J = 6.93 Hz). HRMS (FAB+): calcd for [M + Na]+, 857.3275; found, 857.3270. Do not use 1 M tetra-n-butylammonium fluoride in THF to remove the TBDMS group because the O6-(N,N-diphenylcarbamoyl) group is not stable to this reagent.
SYNTHESIS OF 3′-AMINONUCLEOSIDE-CONTAINING SOLID SUPPORT
SUPPORT PROTOCOL 6
Additional Materials (also see Support Protocol 1) 3′-Tritylamino-2′,3′-dideoxynucleosides (S.5, S.9, S.13, or S.18; see Support Protocols 1 to 4) 4-Dimethylaminopyridine Succinic anhydride 10% (v/v) aqueous citric acid, cold 1-Hydroxybenzotriazole 1:1 (v/v) 1-methyl-2-pyrrolidinone (anhydrous)/dimethyl sulfoxide (anhydrous) N,N-Diisopropylethylamine 2-(1H-Benzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate Aminopropyl-conjugated controlled-pore glass (aminopropyl-CPG; Sigma) 1:1:8 (v/v/v) acetic anhydride/2,6-lutidine/THF 16.5% (v/v) N-methylimidazole in THF (see recipe) Mechanical shaker Synthesize 3′-tritylamino-2′,3′-dideoxynucleoside-5′-O-hemisuccinates 1. To a solution of 1.5 mmol of 3′-tritylamino-2′,3′-dideoxynucleoside (S.5, S.9, S.13, or S.18;) in 5 mL CH2Cl2, add 0.22 g (1.8 mmol) 4-dimethylaminopyridine and 0.18 g (1.8 mmol) succinic anhydride. Stir for 1 hr at room temperature. 2. Quench the reaction with 0.6 mL methanol and dilute with 30 mL CH2Cl2. 3. Extract once each with 20 mL cold 10% aqueous citric acid, 20 mL water, and 20 mL saturated aqueous NaCl. 4. Dry the organic layer over Na2SO4, vacuum filter, and concentrate the product (S.21; Figure 4.7.12) to a foam on a rotary evaporator using a vacuum line. N6-Benzoyl-3′-tritylamino-2′,3′-dideoxyadenosine-5′-O-hemisuccinate: 100% yield (1.15 g). N4-Benzoyl-3′-tritylamino-2′,3′-dideoxycytidine-5′-O-hemisuccinate: 76% yield (0.77 g). 3′-Tritylamino-2′,3′-deoxythymidine-5′-O-hemisuccinate: 94% yield (0.82 g). O6-(N,N-Diphenylcarbamoyl)-N2-isobutyryl-3′-tritylamino-2′,3′-dideoxyguanosine5′-O-hemisuccinate: 78% yield (1.02 g).
Synthesis of Modified Oligonucleotides and Conjugates
4.7.31 Current Protocols in Nucleic Acid Chemistry
Supplement 9
O O
HO
O
B
O NHTr 21a 21c 21g 21t
Bz
B=A Bz B=C iBu,DPC B=G B= T
Figure 4.7.12 Structures of N-protected-3′-tritylamino-2′,3′-dideoxyribonucleoside-5′-O-hemisuccinates.
Conjugate 3′-tritylamino-2′,3′-dideoxynucleoside-5′-O-hemisuccinates to CPG 5. To a solution of 1 mmol 3′-tritylamino-2′,3′-dideoxynucleoside-5′-O-hemisuccinate and 0.13 g (0.95 mmol) of 1-hydroxybenzotriazole in 10 mL of 1:1 (v/v) 1-methyl2-pyrrolidinone/DMSO, add 0.35 mL (2.0 mmol) N,N-diisopropylethylamine and 0.36 g (0.95 mmol) 2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate. 6. Stir the solution 5 min, add 10.0 g aminopropyl-CPG, and put on a shaker for 6 hr, room temperature. 7. Vacuum filter the CPG and wash successively with 20 mL each of DMF, methanol, and ethyl ether. 8. Prepare a 1:1 (v/v) mixture of 1:1:8 acetic anhydride/2,6-lutidine/THF and 16.5% N-methylimidazole/THF, add this mixture to the support on the funnel, and let stand for 30 min to cap any unreacted amino groups on the CPG. 9. Vacuum filter the CPG and wash successively with 20 mL each of acetonitrile, methanol, and diethyl ether. 10. Dry the CPG using a vacuum pump. The nucleoside loadings, determined by trityl assay at 432 nm in 20% TFA/CHCl3 using a molar extinction coefficient of 40.7 ìmol−1 cm−1, were 38.6 ìmol/g for A, 33.6 ìmol/g for C, 29.0 ìmol/g for T, and 39.0 ìmol/g for G. For larger scales, use an overhead stirrer instead of a shaker. Do not use a magnetic stir bar because the CPG will be crushed. SUPPORT PROTOCOL 7
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
PHOSPHORAMIDITE SYNTHESIS The preparation of nucleoside 5′-O-cis-(2,6-dimethylpiperidinyl)-2-cyanoethylphosphoramidite monomers is described below. While either cis-2,6-dimethylpiperidino phosphoramidites or N,N-diisopropylamino phosphoramidites can be prepared and used for the preparation of pnODNs, the former are preferred because they allow the use of significantly lower equivalents per coupling (Fearon et al., 1998). Additional Materials (also see Support Protocols 1 to 5) 3′-Tritylamino-2′,3′-dideoxynucleosides (see Support Protocols 1 to 4) or 3′-O-(4,4′-dimethoxytrityl)-2′-deoxyribonucleosides (see Support Protocol 5) Phosphorus trichloride 3-Hydroxypropionitrile 10% (w/v) aqueous KOH 1:4 (v/v) toluene/hexane
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cis-2,6-Dimethylpiperidine 1,8-Diazabicyclo[5.4.0]undec-7-ene (DBU), distilled from CaH2 before use Synthesize 2-cyanoethylphosphorodichloridite 1. To a solution of 500 mL (5.73 mol) phosphorus trichloride in 250 mL acetonitrile, add dropwise at room temperature, with stirring and bubbling argon, a solution of 47 mL (0.69 mol) 3-hydroxypropionitrile in 250 mL acetonitrile. 2. Stir the solution 15 min at room temperature, with absorption of evolving HCl into a solution of 10% aqueous KOH. 3. Concentrate on a rotary evaporator using a pump and vacuum filter, under argon, into a distillation flask. 4. Distill the product under reduced pressure. The 2-cyanoethylphosphorodichloridite distills as a colorless liquid at 78° to 80°C at 1.0 mmHg. Yield = 75.7% (88.5 g). 31P NMR (CDCl3): δ 180.3.
Synthesize 2-cyanoethyl-cis-(2,6-dimethylpiperidinyl)chlorophosphoramidite 5. To a solution of 35.0 g (203.6 mmol) 2-cyanoethylphosphorodichloridite in 300 mL of 1:4 toluene/hexane, add 55 mL (408.1 mmol) cis-2,6-dimethylpiperidine dropwise, with stirring at 4°C. 6. Stir the reaction for 2 hr at room temperature. 7. Vacuum filter and wash the solid with 40 mL of 1:4 toluene/hexane under argon. 8. Concentrate the filtrate using a rotary evaporator and vacuum pump. 9. To the resultant oil, add 5 mL CH2Cl2 and 300 mL hexane, and crystallize the product overnight at 4°C. 10. Filter the 2-cyanoethyl-cis-(2,6-dimethylpiperidinyl)chlorophosphoramidite under argon. 11. Crush with a spatula, wash with 100 mL of 100:3 (v/v) hexane/CH2Cl2, and dry using a rotary evaporator and vacuum pump. 12. Concentrate the mother liquor (step 3) and recrystallize again (step 9) to obtain a second crop of pale yellow 2-cyanoethyl-cis-(2,6-dimethylpiperidinyl)chlorophosphoramidite. Combined yield = 76.5% (38.8 g). 31P NMR (CDCl3): δ 172.7.
Synthesize nucleoside 5′-O-[cis-(2,6-dimethylpiperidinyl)(2-cyanoethyl)]phosphoramidite monomers (S.1 and S.2) 13. Azeotrope 10.0 mmol of 3′-tritylamino-2′,3′-dideoxynucleoside (S.5, S.9, S.13, or S.18) or 3′-O-(4,4′-dimethoxytrityl)-2′-deoxyribonucleoside (S.20) two times from 50 mL CH3CN. 14. Dissolve azeotroped nucleoside in 30 mL CH2Cl2 and then add 3.0 mL (20.0 mmol) DBU. 15. With stirring, add a solution of 3.0 g (12.0 mmol) 2-cyanoethyl-cis-(2,6-dimethylpiperidinyl)chlorophosphoramidite in 8 mL CH2Cl2, under an argon atmosphere. Stir the reaction mixture for 15 min at ambient temperature. 16. Check the reaction by TLC.
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In order to obtain accurate TLC of the product, pre-wet the TLC plate by immersing it in 10% triethylamine/CH2Cl2, quickly let it dry, then immediately spot the sample, and elute with 5:70:25 Et3N/ethyl acetate/hexane.
17. To avoid decomposition, desalt the crude reaction by loading the mixture directly onto a silica gel column (APPENDIX 3E) preequilibrated in 5% triethylamine/CH2Cl2 and quickly elute it in the same solvent system. These phosphoramidites are not stable to an aqueous workup after the reaction.
18. Remove solvents under reduced pressure and purify the crude product on a silica gel column preequilibrated with 0.5% to 5% triethylamine in 2% methanol/CH2Cl2. Elute as indicated below. N6-Benzoyl-3′-tritylamino-5′-O-[(cis-2,6-dimethylpiperidino)(2-cyanoethoxy)]phosphinyl-2′,3′-dideoxyadenosine (S.1). Purify on silica (60% to 70% ethyl acetate/hexane containing 3% triethylamine). Yield = 83.1% (6.72 g). 31P NMR (CD3CN): δ 148.82, 149.16. Rf = 0.45. N4-Benzoyl-3′-tritylamino-5′-O-[(cis-2,6-dimethylpiperidino)(2-cyanoethoxy)]phosphinyl2′,3′-dideoxycytidine (S.1). Purify on silica (70% ethyl acetate/hexane containing 3% triethylamine). Yield = 82.7% (6.50 g). 31P NMR (CD3CN): δ 149.31, 149.68. Rf = 0.43. O6-(N,N-Diphenylcarbamoyl)-N2-isobutyryl-3′-tritylamino-5′-O-[(cis-2,6-dimethylpiperidino)(2-cyanoethoxy)]phosphinyl-2′,3′-dideoxyguanosine (S.1). Purify compound on silica (60% ethyl acetate/hexane containing 3% triethylamine). Yield = 76.9% (7.58 g). 31P NMR (CD3CN): δ 148.93, 149.50. Rf = 0.57. 3′-Tritylamino-5′-O-[(cis-2,6-dimethylpiperidino)(2-cyanoethoxy)]phosphinyl-2′,3′-dideoxythymidine (S.1). Purify compound on silica (50% ethyl acetate/hexane containing 3% triethylamine). Yield = 79.4% (5.52 g). 31P NMR (CD3CN): δ 149.13, 149.49. Rf = 0.60. N6-Benzoyl-3′-O-(4,4′-dimethoxytrityl)-5′-O-[(cis-2,6-dimethylpiperidino)(2-cyanoethoxy)]phosphinyl-2′-deoxyadenosine (S.2). Purify compound on silica (60% to 70% ethyl acetate/hexane containing 3% triethylamine). Yield= 76.7% (6.67 g). 31P NMR (CD3CN): δ 149.26, 149.39. Rf = 0.47. N4-Benzoyl-3′-O-(4,4′-dimethoxytrityl)-5′-O-[(cis-2,6-dimethylpiperidino)(2-cyanoethoxy)] phosphinyl-2′-deoxycytidine (S.2). Purify compound on silica (60% to 75% ethyl acetate/hexane containing 3% triethylamine). Yield = 74.4% (6.29 g). 31P NMR (CD3CN): δ 149.37, 149.76. Rf = 0.43. O6-(N,N-diphenylcarbamoyl)-N2-isobutyryl-3′-O-(4,4′-Dimethoxytrityl)-5′-O-[(cis2,6-dimethylpiperidino)(2-cyanoethoxy)]phosphinyl-2′-deoxyguanosine (S.2). Purify compound on silica (50% ethyl acetate/hexane containing 3% triethylamine). Yield = 71.0% (7.43 g). 31P NMR (CD3CN): δ 149.32, 149.51. Rf = 0.60. 3′-O-(4,4′-Dimethoxytrityl)-5′-O-[(cis-2,6-dimethylpiperidino)(2-cyanoethoxy)]phosphinyl2′-deoxythymidine (S.2). Purify compound on silica (60% ethyl acetate/hexane containing 3% triethylamine). Yield = 74.1% (5.61 g). 31P NMR (CD3CN): δ 149.24, 149.65. Rf = 0.60. Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. The acetonitrile used for all formulations must contain ≤0.001% water. Oven-dry all bottles and syringes. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Buffer D: 0.1 M TEAB/2% acetonitrile, pH 8 Dilute 100 mL of 1 M TEAB buffer, pH 8 (see recipe), in 880 mL water and add 20 mL acetonitrile. Check that the pH is 8.0 and correct with triethylamine or dry ice, if necessary. Filter through a 0.2-µm filter before use on the HPLC. Store up to 6 months at 4°C. Dichloroacetic acid in CH2Cl2, 3% (v/v) Dissolve 12 mL of dichloroacetic acid in 388 mL of CH2Cl2. Store up to 1 week at room temperature. This solution should only be kept for about 1 week due to its potential to generate HCl, which is extremely detrimental to the pnODN synthesis. Also, only use bottles of DCA <6 months old.
5′-O-(4,4′-Dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-3′-O-[(N,N-diisopropylamino)(2-cyanoethoxy)]phosphinyl uridine, 0.1 M Dissolve 86 mg of uridine phosphoramidite (Glen Research) per milliliter of acetonitrile in an oven-dried bottle under argon. This solution can be used on the synthesizer for 1 week if kept under argon. Also remember that this monomer requires 0.5 M 1H-tetrazole in acetonitrile for activation.
cis-2,6-Dimethylpiperidinyl phosphoramidite monomer solutions, 0.1 M Weigh solid nucleoside 5′-O-[cis-(2,6-dimethylpiperidinyl)(2-cyanoethyl)]phosphoramidite monomer (see Support Protocol 7) and transfer it to a monomer bottle. Cap the bottle with a rubber septum, use vacuum to evacuate the bottle, and refill it with argon. Using a syringe, dissolve the monomer in the appropriate amount of acetonitrile under argon. Use the following amounts of monomer per milliliter acetonitrile for 0.1 M solutions: S.1: 80.8 mg A; 78.4 mg C; 98.5 mg G; 69.5 mg T; S.2: 85.7 mg A; 83.3 mg C; 104.6 mg G; 75.6 mg T. Store monomer solutions up to 5 days at room temperature under argon. Store the solid monomers in desiccated bags at −20°C and allow to warm up to room temperature before opening.
Dowex 50W-X8 cation-exchange resin (pyridinium H+ form) Suspend Dowex 50W-X8 cation-exchange resin (sulfonic acid form; Janssen Pharmaceutica) in pyridine and let cool to room temperature. Filter the resin and wash three times with methanol. Store the filtered resin in a sealed bottle until use. H2O2/H2O/pyridine/THF, 1.5:3.5:20:75 (v/v/v/v) Dissolve 5 mL of 30% aqueous H2O2 (v/v; Aldrich) in 20 mL pyridine and 75 mL tetrahydrofuran. The water in the formulation comes from the aqueous hydrogen peroxide solution. This solution is stable for 3 weeks on the synthesizer, but should be disposed of within a few days of removing it from the synthesizer due to its potential to form explosive peroxides.
Isobutyric anhydride/2,6-lutidine/THF, 1:1:8 (v/v/v) Dissolve 20 mL isobutyric anhydride and 20 mL of 2,6-lutidine in 160 mL anhydrous tetrahydrofuran under argon. This solution is stable on the instrument for 2 months as long as it is kept dry and under argon.
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N-Methylimidazole in THF, 16.5% (v/v) This solution can be bought directly from PE Biosystems or made by dissolving 33 mL of N-methylimidazole in 167 mL anhydrous tetrahydrofuran. This solution is stable on the instrument for 2 months as long as it is kept dry and under argon.
Pyridine in CH3CN, 20% (v/v) Dissolve 40 mL pyridine, previously dried over chunks of CaH2 for at least 24 hr, in 160 mL acetonitrile under argon. This formulation must be kept dry in order to prevent hydrolysis of the internucleotide phosphoramidite during synthesis; it can be kept on the instrument for 1 month under argon.
Tetrazole in CH3CN, 0.167 M Under argon, add 50 mL of a 0.5 M solution of tetrazole in acetonitrile (PE Biosystems) to 100 mL acetonitrile. This solution is stable on the instrument for 2 months as long as it is kept dry and under argon.
Triethylammonium bicarbonate (TEAB) buffer, 1 M, pH 8 Add 139 mL of triethylamine to ~800 mL of water. In a hood, slowly add dry ice to the stirring solution until the pH measures 8.0. Add enough water to make the final volume 1 L. Correct the pH to 8.0 with either triethylamine or dry ice, if necessary. Store the solution up to 6 months at 4°C and check the pH before use. Triethylamine will be immiscible in water at first. The solution gets very cold and CO2 is given off during the dissolution of dry ice.
COMMENTARY Background Information
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
Oligonucleotides are widely used in research and diagnostics and are rapidly gaining acceptance as potential therapeutic agents. Antisense oligonucleotides, which are complementary to selected sequences of mRNA associated with a disease, control gene expression either by sterically blocking translation or processing of the RNA or by the irreversible cleavage of the target RNA by endogenous RNase H (Uhlman and Peyman, 1990). Alternatively, antigene therapeutics are oligonucleotides that prevent gene transcription by forming a triple-helical structure with targeted polypurine:polypyrimidine sites in double-stranded (ds) DNA (Thuong and Hélène, 1993). Many oligonucleotide analogs have been investigated in an attempt to improve properties such as resistance to nuclease degradation, binding strength to RNA and/or dsDNA, cellular uptake, and pharmacokinetic parameters (Uhlman and Peyman, 1990). Unfortunately, phosphorothioates, and the majority of other firstgeneration analogs that possess increased resistance to nuclease degradation, have a decreased binding affinity for their RNA or dsDNA targets. Additionally, phosphorothioate
ODNs may exhibit nonspecific effects, presumably due to adventitious protein binding (Stein and Cheng, 1993). The fully modified oligonucleotide N3′→P5′ phosphoramidates (pnODNs), on the other hand, possess an increased stability to nucleases relative to both phosphodiester and phosphorothioate ODNs, the ability to tightly and sequence-specifically bind to both RNA and dsDNA, and have shown minimal sequence-independent protein binding (Gryaznov and Chen, 1994; Chen et al., 1995; Gryaznov et al., 1995). These improved properties have translated well in both cell culture and in vivo antisense studies where pnODNs have demonstrated efficacy at 10% of the comparably effective dose of phosphorothioate ODNs (Gryaznov et al., 1996; Heidenreich et al., 1997; Skorski et al., 1997). In addition, a pnODN sequence showed no toxicity in preliminary in vivo studies in mice up to dosages of 150 mg/kg administered intravenously six times over a period of two weeks, whereas kidney and liver toxicity were seen for the isosequential phosphorothioate ODN at the same (and lower) dose (Zon, pers. comm.). Consistent with most oligonucleotide analogs other than phosphorothioates, uniformly
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modified pnODNs do not activate RNase H (DeDionisio and Gryaznov, 1995; Heidenreich et al., 1997). However, in many systems, RNase H activity has been necessary in order to obtain an “antisense” effect and is generally thought to increase the activity of such compounds. Although pnODNs have demonstrated significant RNase H–independent activity, the efficacy of chimeric ODNs that possess an RNase H-active core of 6 to 8 phosphorothioate linkages flanked by pnODN linkages is under investigation. The significantly increased and sequencespecific binding affinity of uniformly modified pnODNs for dsDNA renders these “antigene” compounds potentially useful for inhibition of gene expression via triplex helix formation. Hélène and coworkers (Escudé et al., 1996; Giovannangeli et al., 1997) have demonstrated that pnODNs work best when they are designed to form triple-helical DNA by Hoogsteen hydrogen bonding to the duplex with the third strand in a parallel orientation with respect to the polypurine target sequence. Using a eukaryotic transcription assay, they showed that a pnODN can arrest transcription at a specific triplex site in the type 1 human immunodeficiency virus nef gene under the control of a cytomegalovirus promoter. The synthetic method described herein (McCurdy et al., 1997; Nelson et al., 1997; Fearon et al., 1998), based on a phosphoramidite amine-exchange reaction, is 3 to 6 times more efficient than the oxidative phosphorylation-based method previously reported by Gryzanov and Chen (1994) and Chen et al. (1995). An advantage of the described method is that the monomer is activated and used in excess; thus, a small amount of inadvertent hydrolysis only wastes monomer and does not necessarily decrease the step yield of the coupling. Also, because the method is based on phosphoramidite chemistry, the synthesis of chimeras possessing any combination of phosphoramidate, phosphodiester, and/or phosphorothioate linkages at predefined positions is easily performed on a commercially available synthesizer with no instrument modifications. Fluorescent and other specialty phosphoramidites are also easily used in conjunction with this chemistry. This amine-exchange method was modified for use on a vortexing-mode synthesizer (e.g., PE Biosystems 390Z; 10 µmol-scale) and only ≤3.6 eq of the cis-2,6-dimethylpiperidinyl phosphoramidite monomers and 9 eq of 1Htetrazole were necessary to achieve optimal
coupling yields (Fearon et al., 1998). Even lower stoichiometric requirements may be possible using an activator other than 1H-tetrazole.
Compound Characterization Analytical IEC and/or capillary gel electrophoresis (CGE) is generally used to determine the chain-length purity of pnODNs and their chimeras (see Fig. 4.7.13). CGE was performed on a Beckman P/ACE 5510 system with 10% Microgel Capillaries (0.1 × 500 mm) in 35 mM Tris-borate buffer, pH 9.0, in the presence of 15% ethylene glycol, with a 5 sec injection at 10 kV and a running voltage of 25 kV. Alternatively, 20% polyacrylamide gel electrophoresis followed by visualization with Stains-All and densitometric scanning can also be used. If further characterization is desired, electrospray ionization mass spectrometry can be used to determine the molecular weight of the oligonucleotide synthesized; however, sodium counterions must first be exchanged for ammonium ions (Stultz and Marsters, 1991). In the case of chimeras, the number of phosphoramidate, phosphodiester, and/or phosphorothioate linkages can be determined by 31P-NMR spectroscopy in D2O.
Critical Parameters and Troubleshooting The phosphoramidite monomer, 1H-tetrazole, and 20% pyridine in acetonitrile solutions all must be kept rigorously anhydrous; the method is extremely sensitive to water. Additionally, the 5′-phosphoramidite monomers must have a purity of ≥99% by 31P-NMR spectroscopy in order to afford optimal coupling efficiency. The 2-cyanoethyl-protected phosphoramidate linkages are reasonably stable to acidic conditions; however, older formulations of 3% dichloroacetic acid in CH2Cl2 can degrade the protected phosphoramidate linkages and cause an increase in failure sequences. In general, only enough 3% dichloroacetic acid solution is formulated for a one week period. For the same reason, bottles of dichloroacetic acid from the vendor are generally used or discarded within 6 months of purchase. RP-HPLC using either the 3′-terminal tritylamino group or 3′-terminal O-DMTr group alone must be avoided because the use of acetic acid during post-HPLC detritylation severely degrades the pnODN. For this reason, the DMTr uridine incorporation method should be used for RP-HPLC purification and subsequent deprotection or IEC of the “trityl-off” pnODN should be performed. Do not add any base other
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Figure 4.7.13 Analytical CGE of a crude pnODN with the sequence 5′-ATGACTGAGTACAAACOH-3′.
than concentrated aqueous ammonia to the uridine-containing pnODNs to prevent premature cleavage of the phosphodiester function. It is best to remove the ammonia and RP-HPLC purify the pnODN as soon as possible. The uridine RP-HPLC purification method should be used for pnODN/phosphorothioate ODN chimeras because the phosphorothioate portion of the molecule greatly reduces the resolution of the IEC. Additionally, avoid exposure of these chimeras to aqueous NaOH solutions, which can result in small amounts of inadvertent conversion of the phosphorothioate linkages to phosphodiester linkages. Also avoid concentrating the uridine-containing oligonucleotide to dryness before purification. Isolated pnODNs and their chimeras should be stored frozen (−20°C) as lyophilized powders or aqueous solutions in deionized water or buffer at pH ≥ 7.4. During purification and analysis, working solutions of pnODNs should be stored at 4°C. If the pH of the solution is <7.0, there is potential for the pnODN to degrade.
Anticipated Results
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
The isolated yield of pnODNs and their chimeras depends on both length, sequence, and number of phosphoramidate, phosphorothioate, and phosphodiester linkages, but is generally 15 to 30 OD260 units for pnODNs and 20 to 35 OD260 units for chimeras 12 to 25
nt in length with a purity of ≥85%. Occasionally, certain sequences require a second IEC purification after RP-HPLC to obtain purities ≥85%. Figure 4.7.14 shows a representative IEC and CE for the purified pnODN with the sequence 5′-CCCTCCTCCGGAGCCOH-3′. Table 4.7.1 shows the yields and purities of several pnODNs and chimeras synthesized by this method.
Time Considerations The formulation of reagents and set-up of the synthesizer generally take about 3 to 4 hr. The cycle time to introduce each nucleotide is 22 min, which is multiplied by the length of the oligonucleotide being synthesized to determine the overall synthesis time. Deprotection is usually performed overnight in an oven equipped with a timer set to turn off after 8 to 12 hr. Analysis of the quality of the synthesis, sample preparation, and either IEC or RP-HPLC purification take about 3 hr per sample; however for IEC purification, the analysis of the resulting fractions can take an additional 6 to 10 hr. For RP-HPLC, there is an additional 16 hr treatment with fluoride and base to cleave the uridine phosphodiester linkage. The work-up and analysis steps take another 4 to 8 hr, depending on the method used for desalting. Thus, a sample can be synthesized, purified, workedup, and analyzed in 4 to 5 days. Using the advantages of automation and a multi-column
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A
B
instrument, the rather long turn-around time can be overcome by multiplexing the synthesis and purification steps. Generally, about 3 to 4 pnODNs can be synthesized and purified per week by one chemist, assuming an autosampler is utilized for fraction analysis. The synthesis of the monomers is a challenging task that will take a skilled organic chemist approximately 2 to 3 months for the four 3′tritylamino nucleoside monomers and 1 month for the four 3′-O-DMTr nucleoside monomers. Alternatively, the 3′-O-DMTr-nucleoside-5′O-(N,N,-diisopropylamino)phosphoramidite monomers and the 3′-tritylamino-nucleoside5′-O-(N,N-diisopropylamino)phosphoramidite monomers are available from Glen Research and Annovis, respectively, and can be used in place of the cis-(2,6-dimethylpiperidino) phosphoramidite monomers, as long as the coupleoxidize-couple-oxidize protocol is used for the couplings to 3′-amines. These monomers also require 0.5 M 1H-tetrazole in acetonitrile for activation.
Literature Cited Chen, J.-K., Schultz, R.G., Lloyd, D.H., and Gryaznov, S.M. 1995. Synthesis of oligodeoxyribonucleotide N3′→P5′ phosphoramidates. Nucl. Acids Res. 23:2661-2668. Czernecki, S. and Valéry, J.M. 1991. An efficient synthesis of 3′-azido-3′-deoxythymidine (AZT). Synthesis 1991:329-240. Figure 4.7.14 (A) Analytical IEC chromatogram and (B) CGE electropherogram of the RP-HPLC purified and deprotected phosphoramidate o l i g onuc leotide, 5′CCCTCCTCCGGAGCCOH-3′.
DeDionisio, L. and Gryaznov, S.M. 1995. Analysis of a ribonuclease H digestion of N3′→P5′ phosphoramidate-RNA duplexes by capillary gel electrophoresis. J. Chromatogr. B Biomed. Appl. 669:125-131.
Table 4.7.1 Yields and Purities of Phosphoramidate Oligonucleotides and Chimera Synthesized by the Amine-Exchange Method
Sequencea
Purification method
OD260 % Purity by CGE
CCCTCCTCCGGAGCCOH AGAGATTTTTACACCOH CCAGAGTCACACAACAOH CAGATCGTCCATGGTCOH CAGATpCpGpTpCpCpApTGGTCOH GGACCsCsTsCsCsTsCsCsGGAGCCOH TTGCCCACAsCsCsGsAsCsGsGsCGCCCACCAFAM
RP-HPLC IEC IEC IEC IEC RP-HPLC RP-HPLC
20 29 15 17 28 26 29
95% 98% 93% 99+% 85% 97% 70%b
a
The sequences are reported in the 5′→3′ direction. The linkages are phosphoramidate unless noted by “s” or “p,” which are phosphorothioate or phosphodiester linkages, respectively. CGE, capillary gel electrophoresis; FAM, fluorescein; IEC, ion-exchange chromatography; RP-HPLC, reversed-phase high-pressure liquid chromatography. b Polyacrylamide gel electrophoresis of this compound, followed by staining with Stains-All and densitometric scanning, indicated a purity of 90%.
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Escudé, C., Giovannangeli, C., Sun, J.-S., Lloyd, D.H., Chen, J.-K., Gryaznov, S.M., Garestier, T., and Hélène, C. 1996. Stable triple helices formed by oligonucleotide N3′→P5′ phosphoramidates inhibit transcription elongation. Proc. Natl. Acad. Sci. U.S.A. 93:4365-4369. Fearon, K.L., Nelson, J.S., Hirschbein, B.L., Foy, M.F., Nguyen, M.Q., McCurdy, S.N., Frediani, J.E., Okruszek, A., DeDionisio, L.A., Raible, A.M., and Boyd, V. 1998. An improved synthesis of oligonucleotide N3′→P5′ phosphoramidates and their chimera using hindered phosphoramidite monomers and a novel handle for reverse phase purification. Nucl. Acids Res. 26:3813-3824. Giovannangeli, C., Diviacco, S., Labrousse, V., Gryaznov, S., Charneau, P., and Hélène, C. 1997. Accessibility of nuclear DNA to triplex-forming oligonucleotides: The integrated HIV-1 provirus as a target. Proc. Natl. Acad. Sci. U.S.A. 94:7984. Gryaznov, S. and Chen, J.-K. 1994. Oligodeoxyribonucleotide N3′→P5′ phosphoramidates: Synthesis and hybridization properties. J. Am. Chem. Soc. 116:3143-3144. Gryaznov, S.M., Lloyd, D.H., Chen, J.-K., Schultz, R.G., DeDionisio, L.A., Ratmeyer, L., and Wilson, W.D. 1995. Oligonucleotide N3′→P5′ phosphoramidates. Proc. Natl. Acad. Sci. U.S.A. 92:5798-5802.
Skorski, T., Perrotti, D., Nieborowska-Skorska, M., Gryaznov, S., and Calabretta, B. 1997. Antileukemia effect of c-myc N3′→P5′ phosphoramidate antisense oligonucleotides in vivo. Proc. Natl. Acad. Sci. U.S.A. 94:3966-3971. Stec, W.J., Uznanski, B., Wilk, A., Hirschbein, B.L., Fearon, K.L., and Bergot, B.J. 1993. Bis(O,O-diisopropoxy phosphinothioyl) disulfide: A highly efficient sulfurizing reagent for cost-effective synthesis of oligo(nucleoside phosphorothiaote)s. Tetrahedron Lett. 34:5317-5320. Stein, C.A. and Cheng, Y.-C. 1993. Antisense oligonucleotides as therapeutic agents: Is the bullet really magical? Science 261:1004-1012. Stultz, J.T. and Marsters, J.C. 1991. Improved electrospray ionization of synthetic oligonucleotides. Rapid Commun. Mass. Spectrom. 5:359-363. Thuong, N.T. and Hélène, C. 1993. Sequence-specific recognition and modification of doublehelical DNA by oligonucleotides. Angew. Chem. Int. Ed. Engl. 32:666-690. Uhlman, E. and Peyman, A. 1990. Antisense oligonucleotides: A new therapeutic principle. Chem. Rev. 90:544-584.
Gryaznov, S., Skorski, T., Cucco, C., NieborowskaSkorska, M., Chiu, C.Y., Lloyd, D., Chen, J.-K., Koziolkiewicz, M., and Calabretta, B. 1996. Oligonucleotide N3′→P5′ phosphoramidates as antisense agents. Nucl. Acids Res. 24:1508-1514.
Wagner, D., Verheyden, J.P.H., and Moffatt, J.G. 1974. Preparation and synthetic utility of some organotin derivatives of nucleosides. J. Org. Chem. 39:24-30.
Hansske, F. and Robins, M.J. 1983. A deoxygenative [1,2]-hydride shift rearrangement converting cyclic cis-diol monotosylates to inverted secondary alcohols. J. Am. Chem. Soc. 105:6736-6737.
Key References
Heidenreich, O., Gryaznov, S., and Nerenberg, M. 1997. RNase H-independent antisense activity of oligonucleotide N3′→P5′ phosphoramidates. Nucl. Acids Res. 25:776-780. Herdewijn, P. and Van Aerschot, A. 1989. Synthesis of 9-(3-azido-2,3-dideoxy-β-D-erythro-pentofuranosyl)- 2,6-diaminopurine (AzddDPA). Tetrahedron Lett. 30:855-858. McCurdy, S.N., Nelson, J.S., Hirschbein, B.L., and Fearon, K.L. 1997. An improved method for the synthesis of N3′→P5′ phosphoramidate oligonucleotides. Tetrahedron Lett. 38:207-210. Nelson, J.S., Fearon, K.L., Nguyen, M.Q., McCurdy, S.N., Frediani, J.E., Foy, M.F., and Hirschbein, B.L. 1997. N3′→P5′ Oligodeoxyribonucleotide phosphoramidates: A new method of synthesis based on a phosphoramidite amineexchange reaction. J. Org. Chem. 62:7278-7287.
Synthesis and Purification of Oligonucleotide N3′→P5′ Phosphoramidates
Reese, C.B. and Skone, P.A. 1984. The protection of thymine and guanine residues in oligodeoxyribonucleotide synthesis. J. Chem. Soc. Perkin Trans. I 1263-1271.
Nishino, S., Yamamoto, H., Nagato, Y., and Ishido, Y. 1986. Partial protection of carbohydrate derivatives. Part 19. Highly regioselective 5′-Oaroylation of 2′-deoxyribonucleosides in terms of dilution-drop-by-drop-addition procedure. Nucleosides Nucleotides 5:159-168.
Nelson et al., 1997. See above. This reference details the couple-oxidize-coupleoxidize amine-exchange method for synthesis of pnODNs, as well as the synthesis of the 3′-tritylamino protected nucleosides. Fearon et al., 1998. See above. This reference describes the optimized amine-exchange method for the synthesis of pnODNs and the synthesis of the 3′-O-DMTr-protected nucleosides and the cis-2,6-dimethylpiperidinyl phosphoramidite monomers. It also demonstrates the synthesis of pnODNs using low equivalents of monomer and 1H-tetrazole and describes the equilibrium and oxidation steps in detail.
Contributed by Karen L. Fearon and Jeffrey S. Nelson Lynx Therapeutics Hayward, California
4.7.40 Supplement 3
Current Protocols in Nucleic Acid Chemistry
Incorporation of Halogenoalkyl, 2-Pyridyldithioalkyl, or Isothiocyanate Linkers into Ligands
UNIT 4.8
In UNIT 4.3, the direct incorporation of ligands to the 5′ ends of oligodeoxyribonucleotides via their phosphoramidite derivatives is described. A second strategy for the addition of ligands to the 5′ ends of oligodeoxyribonucleotides involves the introduction of appropriate functional groups to two unprotected reactants. Specific coupling of these reactants results in the formation of oligodeoxyribonucleotide conjugates. Many different oligodeoxyribonucleotide conjugates can be prepared starting with only one oligodeoxyribonucleotide synthesis, provided that the required amount for each conjugate is low. This method is particularly useful when (1) a low amount of the ligand is available, (2) the ligand is unstable under the conditions required for oligonucleotide deprotection, or (3) the poor solubility of the ligand in solvents usually used for oligonucleotide synthesis does not allow the preparation of its phosphoramidite or H-phosphonate derivatives. In this approach, functional groups such as amino, phosphate, phosphorothioate, thiol, and carboxyl, that are capable of reacting with functionalized ligands, are attached to the 5′ ends of oligodeoxyribonucleotides. This unit describes the incorporation of linkers containing reactive groups such as halogenoalkyl, 2-pyridyldithioalkyl, or isothiocyanate into ligands. Halogenoalkyl (Asseline et al., 1992, 1996) and 2-pyridyldithioalkyl (Chassignol and Thuong, 1998) linkers can be reacted with either a phosphorothioate or a thiol group incorporated into the oligodeoxyribonucleotide, while the isothiocyanate can react with aminoalkylated oligodeoxyribonucleotides. These ligands are typically 2-methoxy-6-chloro-9-amino-acridine as an intercalator (see Basic Protocol 1), psoralen as a photo-cross-linking reagent (see Basic Protocol 2), phenanthroline-Cu as a cleaving reagent (see Basic Protocol 3), and thiazole orange as a label (see Basic Protocol 4). The addition of carboxyl, amino, phosporothioate, phosphate, and sulfhydryl functions to the 5′ ends of oligonucleotides, as well as methods for linking these functionalized oligonucleotides and ligands, are reported in UNITS 4.9 & 4.10. Alternatively, many labels carrying functional groups that react with 5′-thiol, 5′-terminal phosphorothioate, and 5′-amino groups are commercially available. Heterobifunctional reagents, which allow coupling between two compounds following two successive specific reactions, are also available from commercial sources. The latter compounds, listed in UNIT 4.2, are very useful when conjugates with a well-defined linker between the oligodeoxyribonucleotide and the ligand are not required. CAUTION: All chemicals must be handled in a fume hood. Investigators should be equipped with a laboratory coat, glasses, and gloves. FUNCTIONALIZATION OF AN ACRIDINE DERIVATIVE WITH A BROMOALKYL LINKER
BASIC PROTOCOL 1
The synthesis of 2-methoxy-6-chloro-9-(ω-bromohexylamino)acridine S.1e (Fig. 4.8.1) is achieved by bromination of 2-methoxy-6-chloro-9-(ω-hydroxyhexylamino)acridine S.1b (discussed in UNIT 4.3; Fig. 4.3.1). Synthesis of Modified Oligonucleotides and Conjugates Contributed by Ulysse Asseline and Nguyen T. Thuong Current Protocols in Nucleic Acid Chemistry (2001) 4.8.1-4.8.15 Copyright © 2001 by John Wiley & Sons, Inc.
4.8.1 Supplement 5
OCH3
N
NH(CH2)6OH
OCH3
CBr4, (C6H5)3P
N
NH(CH2)6Br
DMF
Cl
Cl 1b
Figure 4.8.1
1e
Bromination of an acridine derivative. DMF, N,N-dimethylformamide.
Materials 2-Methoxy-6-chloro-9-(ω-hydroxyhexylamino)acridine S.1b (UNIT 4.3) Acetonitrile distilled from P2O5, stored over 3A molecular sieves N,N-Dimethylformamide (DMF) redistilled in vacuo over ninhydrin, stored over 4A molecular sieves Triphenylphosphine (Aldrich) CBr4 (Aldrich) Dichloromethane (CH2Cl2) distilled from P2O5 and passed over basic aluminum oxide Methanol, distilled or synthesis grade Nitrogen source 10-mL round-bottom flask Magnetic stirrer with Teflon stir bars Vacuum pump (oil pump) capable of creating <0.1 mmHg pressure, with manifold and cold trap Preparative 20 × 20–cm glass-backed silica TLC plates (2-mm thickness; Merck) Short-wave UV light box Mortar and pestle 1.5 × 15–cm chromatography column (empty) 2.5 × 7–cm Kieselgel 60F analytical TLC plates (Merck) Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) 1. Place 138 mg (0.38 mmol) of 2-methoxy-6-chloro-9-(ω-hydroxyhexylamino)acridine S.1b into a 10-mL round-bottom flask, and dry by coevaporating three times with 5 mL acetonitrile. 2. Add 5 mL anhydrous DMF and stir with a Teflon stir bar and magnetic stirrer. 3. Add 105 mg (0.40 mmol) triphenylphosphine and 132 mg (0.40 mmol) CBr4 to the acridine solution, and stir the reaction mixture overnight at room temperature. 4. Concentrate the solution to dryness using a vacuum pump. 5. Purify 450 mg of residue by preparative TLC (APPENDIX 3D) using 20 × 20–cm glass-backed silica TLC plates and 85:15 (v/v) CH2Cl2/methanol as the eluent. Dry the plate in a fume hood and visualize by UV shadowing. Incorporation of Halogenoalkyl, 2Pyridyldithioalkyl, or Isothiocyanate Linkers into Ligands
Two distinct yellow bands should be visible. The upper one (which will also be the larger one if the yield is good) corresponds to S.1e; the band just below it corresponds to the residual starting material.
4.8.2 Supplement 5
Current Protocols in Nucleic Acid Chemistry
CAUTION: It is preferable to carry out steps 5 to 8 in a fume hood to avoid breathing solvent vapors and silica powder.
6. Scrape off the silica gel band corresponding to S.1e. Grind the silica gel to a fine powder using a mortar and pestle. 7. Transfer the silica to an empty 1.5 × 15–cm chromatography column and elute with 100 mL of 65:35 (v/v) CH2Cl2/methanol under slight pressure of nitrogen until the yellow color almost disappears. The pressure should be adjusted so that the solvent elutes from the bottom of the column at ~3 to 5 drops/sec.
8. Perform analytical TLC on a 2.5 × 7–cm Kieselgel 60F plate using 85:15 (v/v) CH2Cl2/methanol (Rf S1.b = 0.49, Rf S.1e = 0.54). When 8:2 (v/v) ethyl acetate/hexane is used as the eluent, the starting material S.1b and the product S.1e show an Rf of 0.1 and 0.38, respectively. The yield of S.1e is 60% (97 mg, 0.23 mmol). mp = 108°-110°C. 1H-NMR (CDCl3): δ: 1.55-1.56 (m, 4H, CH2CH2), 1.90 (m, 2H, CH2), 1.99 (m, 2H, CH2), 3.43 (t, 2H, J = 6.6 Hz, CH2Br), 3.94-3.98 (m, 3H, CH2N + NH), 4.0 (s, 3H, OCH3), 7.13-7.86 (m, 5H, Acr), 8.00-8.04 (m, 1H, H-8 Acr). 13C-NMR (DMSO-d6): δ: 31.50, 33.32, 35.61, 38.22, 41.22, 55.14, 62.30, 109.04, 118.14, 121.42, 127.50, 129.96, 132.30, 134.15, 143.06, 145.80, 149.44, 152.30, 160.04, 161.75. Mass analysis. Electrospray ionization mass spectrometry (ESI-MS) polarity positive. Calcd. for C20H22ClBrN2O: 420, 422, and 424; found: 421, 423, and 425 (M+H).
FUNCTIONALIZATION OF A PSORALEN DERIVATIVE WITH AN IODOALKYL OR 2-PYRIDYLDITHIOALKYL LINKER
BASIC PROTOCOL 2
These syntheses are illustrated in Figure 4.8.2. The 5-(6-iodohexyloxy) derivative of psoralen S.2f is obtained by condensation of the hydroxyl derivative S.2b (UNIT 4.3, Fig. 4.3.1) with 1,6-diiodohexane in the presence of K2CO3, following a procedure adapted from the authors’ previously published work (Takasugi et al., 1991). Replacement of the iodine atom of the 5-(6-iodohexyloxy) derivative S.2f by a 2-pyridyldithio group is achieved by a two-step procedure that differs from the one reported by Chassignol and Thuong (1998). First, a thioacetyl derivative S.2g is obtained by the reaction of S.2f with
O
O
O
O
I(CH2)6I, K2CO3 OH
O(CH2)6I
DMF, 65 oC
O
O 2b
2f
CH3COSK
acetone 50 oC
O
O
O
O
N O(CH2)6SS
S S
N O(CH2)6SCOCH3
N CH3CN/CH3OH, NH4OH O
O 2h
2g
Figure 4.8.2 Functionalization of 5-hydroxypsoralen (S.2b) with an iodoalkyl linker (S.2f) or a 2-pyridyldithioalkyl linker (S.2h). DMF, N,N-dimethylformamide.
Synthesis of Modified Oligonucleotides and Conjugates
4.8.3 Current Protocols in Nucleic Acid Chemistry
Supplement 5
potassium thioacetate. The product S.2g is then hydrolyzed in situ by ammonia treatment and the resulting thiol derivative is reacted with 2,2′-dipyridyl disulfide to give the psoralen derivative S.2h. Materials 5-Hydroxypsoralen S.2b (UNIT 4.3) N,N-Dimethylformamide redistilled in vacuo over ninhydrin, stored over 4A molecular sieves 1,6-Diiodohexane (Aldrich) Anhydrous potassium carbonate Argon source CH2Cl2 Distilled pentane (CE Instruments) Distilled methanol Nitrogen source Potassium thioacetate (Aldrich) Distilled acetone Distilled ethyl acetate Distilled hexane (CE Instruments) Sodium sulfate (Aldrich) 2,2′-Dipyridyl disulfide (Aldrich) Acetonitrile distilled from P2O5, stored over 3A molecular sieves 29% (v/v) aqueous ammonium hydroxide 2.5 mg/mL 2,6-dibromo-4-benzoquinone-N-chloroimine (DBPNC; Merck) in ethanol 100-mL and 25-mL round-bottom flasks and stoppers Reflux condenser CaCl2 drying tube Magnetic stirrer with temperature-controlled oil bath and Teflon stir bars 5-cm glass filter funnel (porosity 4) Vacuum pump (oil pump) capable of creating <0.1 mmHg pressure, with manifold and cold trap Kieselgel 60F chromatography column (e.g., 3-cm diameter, 50-cm height, 50 g silica gel; Merck) 2.5 × 7–cm analytical TLC plates (e.g., Kieselgel 60F plates, Merck) Short-wave UV light box Rotary evaporator with a water aspirator Desiccator containing P2O5 7-cm filter funnel with filter paper 25-mL flask with stopper Preparative 20 × 20–cm glass-backed silica gel TLC plates (2-mm thickness; e.g. Merck) Mortar and pestle 1.5 × 15–cm chromatography column (empty) Additional reagents and equipment for thin-layer chromatography (TLC, APPENDIX 3D) and column chromatography (APPENDIX 3E) Incorporation of Halogenoalkyl, 2Pyridyldithioalkyl, or Isothiocyanate Linkers into Ligands
Prepare 5-(6-iodohexyloxy)psoralen S.2f 1. Prepare a solution of 0.6 g (2.97 mmol) 5-hydroxypsoralen S.2b in 15 mL anhydrous N,N-dimethylformamide in a 100-mL round-bottom flask equipped with a reflux
4.8.4 Supplement 5
Current Protocols in Nucleic Acid Chemistry
condenser and a CaCl2 drying tube. Stir reaction with a Teflon stir bar on a magnetic stirrer. 2. Add successively 5.4 mL (11.15 g, 33 mmol) 1,6-diiodohexane and 0.6 g (4.35 mmol) anhydrous potassium carbonate. Heat the stirred reaction mixture at 65°C for 4 hr in the dark under an argon atmosphere. 3. Allow the mixture to cool to room temperature and filter off the insoluble mineral salts by suction using a 5-cm glass filter funnel (porosity 4). Concentrate the filtrate to dryness using a vacuum pump. 4. Dissolve 14 g residue in 7 mL CH2Cl2 and purify on a 50-g Kieselgel 60F chromatography column eluting first with 200 mL of 50:50 (v/v) CH2Cl2/pentane to eliminate the excess diiodohexane, and then with increasing concentrations (0:100 to 4:96, v/v) of methanol in CH2Cl2. Perform chromatography under slight pressure of nitrogen. The pressure should be adjusted so that the solvent elutes from the bottom of the column at ~3 to 5 drops/sec.
5. Monitor fractions by analytical TLC using 2.5 × 7–cm Kieselgel 60F plates and 9:1 (v/v) CH2Cl2/methanol as the eluent. Visualize by UV shadowing and collect fractions containing pure product (Rf = 0.85). 6. Remove the solvent in a rotary evaporator with a water aspirator and wash the white solid S.2f with 2 mL pentane. Dry in a desiccator containing P2O5 for ≥3 to 4 hr. Yield 78% (0.96 g, 2.33 mmol). m.p = 90°-95°C. 1H-NMR (CDCl3): d: 1.45-1.62 [m, 4H, (CH2)2], 1.81-1.94 [m, 4H, (CH2)2], 3.21 (t, 2H, J = 6.8 Hz,CH2I), 4.45( t, 2H, J = 6.4 Hz, -CH2OAr), 6.28 (d, 1H, J = 9.6 Hz, H-3 Pso), 6.94 (d, 1H, J = 2.45 Hz, H′-4 Pso), 7.1 (s, 1H, H-8 Pso), 7.58 (d, 1H, J = 2.5 Hz, H′-5 Pso), 8.16 (d, 1H, J = 9.8 Hz, H-4 Pso). 13 C-NMR (DMSO-d6): δ: 15.13, 30.50, 35.36, 35.78, 38.97, 78.70, 99.44, 111.80, 112.29, 118.64, 119.22, 145.70 (d, J = 95 Hz), 152.15 (d, J = 145.5 Hz), 154.98, 158.34, 163.84, 166.32, 152.30, 160.04, 161.75.
Prepare 5-[6-(acetylthio)hexyloxy]psoralen S.2g 7. Place the following in a 25-mL round-bottom flask equipped with a reflux condenser: 200 mg (0.48 mmol) 5-(6-iodohexyloxy)-psoralen S.2f 65.8 mg (0.57 mmol) potassium thioacetate 10 mL anhydrous acetone Heat at 50°C under magnetic stirring. 8. Monitor the reaction by analytical TLC and UV shadowing, using 3:1 (v/v) ethyl acetate/hexane as the eluent. After 2 hr, the iodinated compound S.2f (Rf = 0.45) is completely transformed into the thiol ester S.2g (Rf = 0.34). The reaction can also be monitored using 96:4 (v/v) CH2Cl2/acetone (Rf S.2f = 0.72, Rf S.2g = 0.60).
9. Remove the solid by gravity filtration through a 7-cm filter funnel fitted with filter paper. 10. Dilute the solution with 50 mL CH2Cl2 and wash the organic solution three times with 5 mL water. 11. Dry the organic phase over sodium sulfate and concentrate to dryness using a rotary evaporator with a water aspirator. 12. Stir residue with 2 mL hexane to obtain a solid.
Synthesis of Modified Oligonucleotides and Conjugates
4.8.5 Current Protocols in Nucleic Acid Chemistry
Supplement 5
Yield 93% (163 mg, 0.44 mmol). mp = 68°-70°C. 1H-NMR (CDCl3): d: 1.42-1.70 (m, 6H, (CH2)3), 1.82-1.93 (m, 2H, CH2), 2.34 (s, 3H, CH3), 2.89 (t, 2H, J = 7.1 Hz, -CH2S), 4.34 (t, 2H, J = 6.4 Hz, -CH2OAr), 6.28 (d, 1H, J = 9.6 Hz, H-3 Pso), 6.94 (d, 1H, J = 2.4 Hz, H′-4 Pso), 7.13 (s, 1H, H-8 Pso), 7.58 (d, 1H, J = 2.3 Hz, H′-5 Pso), 8.15 (d, 1H, J = 9.6 Hz, H-4 Pso). 13C-NMR (DMSO-d6): d: 31.07, 34.01, 34.50, 35.30, 36.76, 61.10, 78.70, 99.40, 111.80, 112.26, 118.60, 119.20, 145.67 (d, J = 87.5 Hz), 152.11 (d, J = 138 Hz), 154.98, 158.34, 163.86, 166.32, 201.51. Mass analysis. ESI-MS polarity positive. Calcd. for C19H20O5S: 360; found: 361 (M+H).
Prepare 5-[6-(2-pyridyldithio)hexyloxy]psoralen S.2h 13. Place the following (in the order listed) in a 25-mL stoppered flask: 150 mg (0.41 mmol) 5-[(6-acetylthio)hexyloxy]psoralen S.2g 0.458 g (2.05 mmol) 2,2′-dipyridyl disulfide 10 mL 50:50 (v/v) acetonitrile/methanol 0.1 mL 29% aqueous ammonium hydroxide. Leave the mixture to react over 2 to 3 days at room temperature. 14. Monitor the reaction by analytical TLC and UV shadowing, using 3:7 (v/v) ethyl acetate/hexane as the eluent. Spray the TLC plate with 2.5 mg/mL DBPNC in ethanol and heat until color appears. After 60 hr, the starting material S.2g (Rf = 0.48) is totally transformed into a new product (Rf = 0.36). 5-[6-(2-Pyridyldithio)hexyloxy]psoralen S.2h appears as a bright yellow-colored spot.
15. Concentrate the solution under reduced pressure using a rotary evaporator with a water aspirator. 16. Purify 650 mg of residue by preparative TLC using 20 × 20–cm glass-backed silica gel TLC plates and 96:4 (v/v) CH2Cl2/acetone as the eluent. Elute plate two times. Visualize by UV shadowing at 365 nm. S.2h can be seen as a pale blue band. CAUTION: It is preferable to carry out steps 16 to 18 in a fume hood to avoid breathing solvent vapors and silica powder.
17. Scrape off the silica gel band corresponding to S.2h. Grind the silica gel to a fine powder using a mortar and pestle. 18. Transfer the silica to an empty 1.5 × 15–cm chromatography column and elute with 100 mL of 85:15 (v/v) CH2Cl2/acetone under slight pressure of nitrogen until the product is eluted (as monitored by TLC and UV shadowing). The pressure should be adjusted so that the solvent elutes from the bottom of the column at ~3 to 5 drops/sec.
Incorporation of Halogenoalkyl, 2Pyridyldithioalkyl, or Isothiocyanate Linkers into Ligands
Yield = 86% (140 mg, 0.35 mmol). 1H-NMR (CDCl3): d: 1.50-1.58 (m, 4H, (CH2)2), 1.72-1.78 (m, 2H, CH2), 1.81-1.89 (m, 2H, CH2), 2.81(t, 2H, J = 7.2 Hz, -CH2-S-S), 4.43 (t, 2H, J = 6.4 Hz, -CH2OAr), 6.27 (d, 1H, J = 9.8 Hz, H-3 Pso), 6.93 (d, 1H, J = 2.1 Hz, H′-4 Pso), 7.05-7.09 (m, 1H, Ar), 7.18 (s, 1H, H-8 Pso), 7.58 (d, 1H, J = 2.3 Hz, H′-5 Pso), 7.60-7.72 (m, 2H, Ar), 8.14 (d, 1H, J = 9.8 Hz, H-4 Pso), 8.45-8.47 (m, 1H, Ar). 13C-NMR (DMSO-d6): d: 31.14, 33.59, 34.42, 35.36, 44.16 (t, J = 153 Hz), 78.70, 99.44, 111.82, 112.27, 118.62, 119.21, 125.43 (d, J = 116.5 Hz), 127.25, 143.92 (d, J = 116.5 Hz), 145.67 (d, J = 94.5 Hz), 152.11 (d, J = 138.5 Hz), 154.99, 155.68, 158.3, 163.85, 165.61, 166.32. Mass analysis. ESI-MS polarity positive. Calcd. for C22H21NO4S2: 427; found: 429 (M+H).
4.8.6 Supplement 5
Current Protocols in Nucleic Acid Chemistry
FUNCTIONALIZATION OF AN ORTHOPHENANTHROLINE DERIVATIVE WITH A BROMOALKYL LINKER OR AN ISOTHIOCYANATE GROUP
BASIC PROTOCOL 3
These syntheses are illustrated in Figure 4.8.3. The preparation of 5-(ω-bromohexanoamido)-1,10-phenanthroline S.3c was adapted from a previously reported two-step procedure (Thuong and Asseline, 1991). Reduction of 5-nitro-1,10-phenanthroline S.3a by ammonium sulfide leads to the 5-amino derivative S.3b, and acylation of the latter with 6-bromohexanoyl chloride gives the bromoalkyl derivative S.3c. Alternatively, the isothiocyanate derivative S.3d is obtained by treatment of the amino derivative S.3b with carbon disulfide in the presence of dicyclohexylcarbodiimide in pyridine. Materials 20% (w/v) aqueous ammonium sulfide Nitrogen source 5-Nitro-1,10-phenanthroline (S.3a; Aldrich) Absolute ethanol, stored over 4A molecular sieves Chloroform Sodium sulfate, anhydrous (Aldrich) Acetonitrile distilled from P2O5, stored over 3A molecular sieves (for S.3c only) N,N-Diisopropylethylamine (Aldrich; for S.3c only) 6-Bromohexanoyl chloride (Aldrich; for S.3c only) 5% (v/v) aqueous NaHCO3 (for S.3c only) Dichloromethane (CH2Cl2) distilled from P2O5 and passed over basic aluminum oxide
N
NHCO(CH2)5Br
N
3c
Br(CH2)5COCl DIPEA, CH3CN
N
NO2
N
(NH4)2S
NH2
o
ethanol/water, 80 C
N
N
3a
3b
CS2, DCC C6H5N
N
N
C
S
N
3d
Figure 4.8.3 Derivatization of orthophenanthroline (S.3a) with a bromoalkyl linker (S.3c) or an isothiocyanate group (S.3d). DCC, 1,3-dicyclohexylcarbodiimide; DIPEA, N,N-diisopropylethylamine.
Synthesis of Modified Oligonucleotides and Conjugates
4.8.7 Current Protocols in Nucleic Acid Chemistry
Supplement 5
Methanol Pyridine redistilled from p-toluenesulfonylchloride, stored over 3A molecular sieves (for S.3d only) 1,3-Dicyclohexylcarbodiimide (Aldrich; DCC; for S.3d only) CS2 (Merck; for S.3d only) Dioxane, freshly distilled (Aldrich; for S.3d only) 500-mL three-necked flask Reflux condenser 100-mL dropping funnel 0.5-cm nitrogen inlet tube Magnetic stirrer with temperature-controlled oil bath and Teflon stir bars 500-mL separatory funnel 7-cm filter funnel and filter paper Rotary evaporator with water aspirator 5-cm glass filter funnel (porosity 4) Desiccator containing P2O5 25-mL flask with rubber stopper Neutral-activated chromatography column, 1.6-cm diameter, 50-cm height (e.g., 40 g of Kieselgel 60 or Aluminumoxid 90; Merck; for S.3c only) 2.5 × 7–cm analytical TLC plates (e.g., Kieselgel 60F or Aluminumoxid 60F254 neutral, Type E; Merck) Short-wave UV light box 10-mL round-bottom flask with rubber septa and glass stoppers (for S.3d only) Silica gel column, 1.5-cm diameter, 45-cm height (17 g Kieselgel 60; Merck; for S.3d only) Additional reagents and equipment for thin-layer chromatography (TLC, APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare 5-amino-1,10-phenanthroline S.3b 1. Under an efficient fume hood, place 60 mL (0.17 mol) of 20% aqueous ammonium sulfide in a 500-mL three-necked flask equipped with a reflux condenser, a 100-mL dropping funnel, and a 0.5-cm nitrogen inlet tube. Heat the flask to 65°C under a nitrogen atmosphere. 2. Dissolve 1 g (4.48 mmol) of 5-nitro-1,10-phenanthroline (S.3a) in 40 mL boiling absolute ethanol and transfer this solution to the dropping funnel. Add this solution dropwise over 1 hr to the magnetically stirred 20% aqueous ammonium sulfide solution at 80°C. 3. Add another 25 mL of 20% aqueous ammonium sulfide and reflux for an additional hour. 4. Allow the stirred solution to cool to room temperature and extract four times with 70 mL chloroform. 5. Pool the chloroform extracts and back-extract two times with 20 mL water using a 500-mL separatory funnel. Incorporation of Halogenoalkyl, 2Pyridyldithioalkyl, or Isothiocyanate Linkers into Ligands
6. Dry the organic solution over anhydrous sodium sulfate and remove the drying agent by gravity filtration through a 7-cm funnel fitted with filter paper. Concentrate the filtrate in a rotary evaporator with a water aspirator. 7. Dissolve 680 mg yellow residue in 20 mL boiling absolute ethanol, filter, and add 15 mL water to the filtrate. Stopper the flask and let stand 2 days at 2° to 5°C.
4.8.8 Supplement 5
Current Protocols in Nucleic Acid Chemistry
8. Collect the yellow crystals by suction using a 5-cm glass filter funnel (porosity 4) and dry in a desiccator containing P2O5 for at least one night. Yield, 60% (520mg, 2.66 mmol). mp = 250°C (with decomposition at 255°-260°C). 1 H-NMR (DMSO-d6): d: ppm, 6.10 (b s, 2H, NH2), 6.85 (s, 1H), 7.47-7.50 (m, 1H), 7.70-7.73 (m, 1H), 8.02-8.04 (m, 1H), 8.65-8.67 (m, 2H), 9.03-9.05 (m, 1H). 13C-NMR (DMSO-d6): δ: 107.98 (d, J = 153 Hz), 128.09 (d, J = 58.5 Hz), 128.38, 129.40 (d, J = 65.5 Hz), 136.79, 137.01, 138.95 (d, J = 109 Hz), 146.67, 148.89, 150.96, 152.37, 155.54 (d, J = 73 Hz). Mass analysis. ESI-MS polarity positive. Calcd. for C12H9N3: 195; found: 197 (M+H).
Prepare 5-(w-bromohexanoamido)-1,10-phenanthroline S.3c 9a. Dry 5-amino-1,10-phenanthroline S.3b (step 8) by coevaporating three times with 10 mL anhydrous acetonitrile. 10a. Combine the following with magnetic stirring in a 25-mL stoppered flask: 0.195g (1 mmol) 5-amino-1,10-phenanthroline S.3b 0.297 g (0.4 mL, 2.30 mmol) N,N-diisopropylethylamine 15 mL acetonitrile. 11a. Add dropwise, at room temperature, a solution of 0.245 g (0.176 mL, 1.15 mmol) of 6-bromohexanoyl chloride in 0.3 mL anhydrous acetonitrile using a syringe and needle inserted through the rubber stopper. The mixture becomes homogeneous during the course of the addition.
12a. Add 5 mL of 5% aqueous NaHCO3 and extract three times with 20 mL dichloromethane. 13a. Pool the organic extracts and back-extract three times with 8 mL water. 14a. Dry the organic phase over anhydrous sodium sulfate and remove the drying agent by gravity filtration through a funnel fitted with filter paper. Evaporate in a rotary evaporator with a water aspirator. 15a. Dissolve 750 mg yellow residue in 1 mL CH2Cl2 and purify on a 40-g neutral-activated chromatography column, eluting with (in order) 200 mL each 99:1, 98:2, and 97:3 (v/v) CH2Cl2/methanol. Perform chromatography under slight pressure of nitrogen. The pressure should be adjusted so that the solvent elutes from the bottom of the column at ~3 to 5 drops/sec.
16a. Monitor by analytical TLC and UV shadowing, using Aluminumoxid 60F254 plates and 97:3 (v/v) CH2Cl2/methanol. Collect fractions containing pure product (Rf S.3c = 0.3). Since the compound decomposes after a few weeks of storage at –20°C, it should be stored at –70°C. Yield = 65% (0.230 g, 0.62 mmol). mp= 90°-95°C (decomposition). 1H-NMR (CDCl3) d ppm, 1.60-1.92 (m, 6H, (CH2)3), 2.58 (m, 2H, CH2C(O)), 3.43 (t, 2H, J = 6.4 Hz, CH2Br), 7.24 (s, 1H), 7.58-7.59 (m, 2H), 7.90 (bs, 1H), 8.15-8.17 (m, 1H), 8.29-8.31 (m, 1H), 9.08-9.13 (m, 2H). 13C-NMR (DMSO-d6): d: 30.58, 33.51, 38.26, 41.32, 41.97, 126.15, 129.10, 129.75, 130.86, 134.31, 137.88, 142.03, 149.94, 152.02, 155.43, 156.10, 178.48. Mass analysis. ESI-MS polarity positive. Calcd. for C18H18 N3BrO: 371 and 373; found: 372 and 374 (M+H). Synthesis of Modified Oligonucleotides and Conjugates
4.8.9 Current Protocols in Nucleic Acid Chemistry
Supplement 5
Prepare 5-(1,10-phenanthroline) isothiocyanate S.3d 9b. Dry 5-amino-1,10-phenanthroline S.3b (step 8) by coevaporating two times with 5 mL anhydrous pyridine. 10b. Combine the following in a 10-mL round-bottom flask equipped with a stir bar: 97.6 mg (0.5 mmol) 5-amino-1,10-phenanthroline S.3b 412.6 mg (2 mmol) 1,3-dicyclohexylcarbodiimide 2.5 mL anhydrous pyridine 152.28 mg (126 µL, 2 mmol) CS2. 11b. Stir the mixture at room temperature and monitor the reaction by analytical TLC on a Kieselgel 60F plate using 9:1 (v/v) CH2Cl2/methanol as the eluent. Visualize by UV shadowing. The starting material S.3b (Rf = 0.23; yellow spot by UV shadowing at 254 nm) is transformed into the isothiocyanate derivative S.3d (Rf = 0.54; blue spot). S.3b and S.3d are poorly resolved on TLC plates.
12b. When the reaction is complete (usually 24 hr), remove pyridine and excess CS2 by evaporation. 13b. Take up the residue with 3 mL freshly distilled dioxane. Stir the mixture for 5 min and filter off the insoluble material by gravity using a funnel fitted with filter paper. Concentrate the filtrate using a rotary evaporator with a water aspirator. Dioxane should be freshly distilled to avoid peroxides.
14b. Purify 250 mg residue on a 17-g Kieselgel 60 column using the following eluents (in order): 100 mL CH2Cl2, 50 mL of 98:2 (v/v) CH2Cl2/methanol, and 100 mL of 95:5 (v/v) CH2Cl2/methanol. Perform chromatography under slight pressure (see step 15a). Yield 76% (90 mg, 0.38 mmol). mp = 91°-93°C. 1H-NMR (CDCl3): d: ppm, 7.27 (s, 1H), 7.54-7.57 (m, 1H), 7.63-7.66 (m, 1H), 8.11-8.13 (m, 1H), 8.57-8.59 (m, 1H), 9.06-9.07 (m, 1H), 9.18-9.19 (m, 1H). 13C-NMR (DMSO-d6): d: 119.06, 129.68, 131.47, 134.79, 138.20, 141.84, 144.07, 148.14, 150.08, 151.45, 152.19, 155.11, 156.61. Mass analysis. ESI-MS polarity positive. Calcd. for C13H7N3S: 237; found: 239 (M+H). BASIC PROTOCOL 4
FUNCTIONALIZATION OF A THIAZOLE ORANGE DERIVATIVE WITH AN IODOALKYL LINKER The preparation of iodooctylthiazole orange, illustrated in Figure 4.8.4, is achieved by a procedure adapted from the literature (Brooker et al., 1942; Benson et al., 1993). 3-Methyl-2-(methylthio)benzothiazolium iodide S.4b reacts with N-(8-iodooctyl)-4methylquinolium iodide S.4d to give iodooctylthiazole orange S.4e. Materials
Incorporation of Halogenoalkyl, 2Pyridyldithioalkyl, or Isothiocyanate Linkers into Ligands
3-Methyl benzothiazole-2-thione S.4a Methyl iodide (Fluka) Distilled methanol Diethyl ether, anhydrous 1,8-Diiodooctane (Aldrich) Dioxane, freshly distilled (Aldrich) Lepidine (Aldrich) CH2Cl2 Nitrogen source
4.8.10 Supplement 5
Current Protocols in Nucleic Acid Chemistry
CH3 N S S
CH3I
I
−
SCH3
CH3OH
4a
CH3 N+ (CH2)8I
S
I
− + N
4b TEA ethanol
S
(CH2)8I
N
I(CH2)8I
I
− + N
4e
dioxan, reflux CH3 4c
N H3C
CH3 4d
Figure 4.8.4 Thiazole orange derivative functionalized with an iodoalkyl linker (S.4e). TEA, triethylamine.
Ethanol, prewarmed (55°C) Triethylamine (Merck) 100-mL and 10-mL round-bottom flask and rubber septa/glass stoppers Magnetic stirrer with temperature-controlled oil bath and Teflon stir bars Reflux condenser 5-cm glass filter funnel (porosity 4) Desiccator containing P2O5 Two-necked round-bottom flask 10-mL dropping funnel Silica gel chromatography columns: 35 g, 2.5-cm diameter, 45-cm height; and 25 g, 1.2-cm diameter, 40-cm height (e.g., Kieselgel 60, Merck) 2.5 × 7–cm analytical TLC plates (e.g., Kieselgel 60F, Merck) Short-wave UV light box Additional reagents and equipment for thin-layer chromatography (TLC, APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare 3-methyl-2-(methylthio)benzothiazolium iodide S.4b 1. Mix the following in a 100-mL round-bottom flask equipped a Teflon stir bar, magnetic stirrer, and reflux condenser: 2.40 g (13.24 mmol) 3-methyl benzothiazole-2-thione S.4a 4.25 g (1.86 mL, 30 mmol) methyl iodide 30 mL distilled methanol. 2. Reflux the magnetically stirred reaction mixture for 3 hr. When S.4a dissolves, a yellowish precipitate appears.
3. Cool the reaction mixture in an ice bath and filter the precipitate by suction using a 5-cm glass filter funnel (porosity 4). 4. Wash the precipitate twice with 10 mL dry diethyl ether. Remove solvent using a rotary evaporator with a water aspirator, and then dry in a desiccator containing P2O5. CAUTION: Diethyl ether can form peroxides. It is a highly flammable solvent that should be handled with care.
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4.8.11 Current Protocols in Nucleic Acid Chemistry
Supplement 5
Yield 95% (4.04 g, 12.5 mmol). mp = 152°-154°C (148°-149°C). Rf = 0.20 by TLC and UV shadowing using CH2Cl2/methanol (9:1, v/v) as eluent. 1H-NMR (DMSO-d6): d ppm, 3.12 (s, 3H, SCH3), 4.10 (s, 3H, NCH3), 7.71 (t, 1H, J = 7.8 Hz), 7.82 (t, 1H, J = 7.8 Hz), 8.18 (d, J = 8 Hz, 1H), 8.40 (d, J = 8 Hz, 1H). 13C-NMR (DMSO-d6): d: 24.37 (d, J = 116.5 Hz), 42.76 (d, J = 87.5 Hz), 121.96 (d, J =87.5 Hz), 130.21 (d, J = 58 Hz), 133.24 (d, J = 109 Hz), 134.48, 135.40 (d, J = 116.5 Hz), 148.77, 187.48. Mass analysis. ESI-MS polarity positive. Calcd. for C9H10NS2: 196; found: 197 (M+H).
Prepare N-(8-iodooctyl)-4-methylquinolium iodide S.4d 5. Stir a solution of 9.15 g (4.97 mL, 25 mmol) of 1,8-diiodooctane in 10 mL of refluxing dioxane in a two-necked round-bottom flask equipped with a reflux condenser and a 10-mL dropping funnel. Add dropwise 0.69 mL (0.746 g, 5 mmol) lepidine S.4c over 30 min with continued heating, and maintaining heating (reflux) for an additional 3 hr. 6. Stop heating and stir mixture for 16 hr at room temperature. 7. Decant the resulting brown oil, wash with anhydrous diethyl ether, and dry under vacuum. 8. Purify on a 35-g silica gel chromatography column using 200 mL of 99:1 (v/v) CH2Cl2/methanol and then 400 mL of 98:2 (v/v) CH2Cl2/methanol as the eluents. Perform chromatography under slight pressure of nitrogen. The pressure should be adjusted so that the solvent elutes from the bottom of the column at ~3 to 5 drops/sec. Yield 53% (1.35 g, 2.6 mol). Brown oil Rf S.4d = 0.35 by TLC and UV shadowing using 9:1 (v/v) CH2Cl2/methanol as eluent. 1H-NMR (DMSO-d6) d ppm, 1.17-1.36 (m, 8H, (CH2)4), 1.68-1.73 (m, 2H, CH2), 1.90-1.93 (m, 2H, CH2), 2.99 (s, 3H, CH3), 3.24 (t, 2H, J = 7 Hz, CH2I), 4.99 (t, 2H, J = 7.8 Hz, NCH2), 8.03-8.07 (m, 2H, Ar), 8.23-8.26 (m, 1H, Ar), 8.53-8.60 (m, 2H, Ar), 9.41-9.42 (m, 1H). 13C-NMR (DMSO-d6): δ: 7.36, 15.31, 25.96 (d, J = 153 Hz), 31.85, 33.83, 34.46, 35.60, 38.94, 63.02 (d, J = 131.5 Hz), 125.54, 128.73 (d, J = 116.5 Hz), 133.40, 135.20, 135.80 (d, J = 109.5 Hz), 141.5 (d, J = 80 Hz), 142.96, 154.53 (d, J = 138.5 Hz), 164.76. Mass analysis. ESI-MS polarity positive. Calcd. for C18H25IN: 382; found: 382 (M+H).
Prepare 8-iodooctylthiazole orange S.4e 9. In a 10-mL round-bottom flask equipped with magnetic stirrer, dissolve 0.156 g (0.48 mmol) S.4b and 0.246 g (0.48 mmol) S.4d in 3 mL warm (55°C) absolute ethanol. 10. Add 20 µL (0.15 mmol) triethylamine and leave the red-colored mixture with magnetic stirring at room temperature. Monitor the reaction by analytical TLC using 9:1 (v/v) CH2Cl2/methanol as the eluent. Visualize by UV shadowing. The spots corresponding to S.4b (Rf = 0.20) and S.4d (Rf = 0.31) disappear while a new red-colored spot appears (Rf = 0.46). The reaction is complete after ~15 min.
11. Remove solvent using a rotary evaporator with a water aspirator, and dry compound in a desiccator containing P2O5. 12. Purify 450 mg residue on a 25-g silica gel chromatography column using 600 mL of 99:1 (v/v) CH2Cl2/methanol as the eluent. Perform chromatography under slight pressure of nitrogen. Incorporation of Halogenoalkyl, 2Pyridyldithioalkyl, or Isothiocyanate Linkers into Ligands
Yield 62% (0.208 g, 0.29 mmol). mp = 123°-124°C. Rf = 0.46 using 9:1 (v/v) CH2Cl2/methanol as the eluent. 1H-NMR (DMSO-d6) d ppm, 1.28-1.45 (m, 8H, (CH2)4), 1.75-1.78 (m, 2H, CH2), 1.85-1.88 (m, 2H, CH2), 3.15 (t, 2H, J = 7.3 Hz, CH2I), 3.99 (s, 3H, NCH3), 4.99 (t, 2H, J = 7.5 Hz, NCH2), 6.70 (s,1H), 7.24-7.25 (m, 1H), 7.27-7.32 (m,1H), 7.44-7.52 (m, 1H), 7.52 (d, J = 8Hz 1H), 7.62 (d, J = 8Hz 1H), 7.66-7.73 (m, 2H), 8.81(d, J = 8Hz, 1H),
4.8.12 Supplement 5
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8.90 (d, J = 8Hz, 1H). 13C-NMR (DMSO-d6): δ: 15.34, 31.95, 33.88, 34.52, 34.96, 35.88, 38.94, 40.10, 60.35, 94.33 (d, J = 145.5 Hz), 114.05, 119.21 (d, J = 116.5 Hz), 124.25, 129.11 (d, J = 116.5 Hz), 130.07, 130.47, 130.71, 132.07, 133.01 (d, J = 131 Hz), 134.47, 139.40, 143.24, 146.70, 150.55, 154.77, 166.26. Mass analysis. ESI-MS polarity positive. Calcd. for C26H30IN2S: 529; found: 529 (M+H).
COMMENTARY Background Information The attachment of conjugate groups to the 5′ terminus of oligodeoxyribonucleotides can be achieved following two strategies. The first strategy is direct incorporation using phosphoramidites of ligands. The second strategy involves the coupling of an unblocked oligomer with the ligand via a specific reaction between the reactive groups present in both entities. This requires the functionalization of ligands with halogenoalkyl, 2-pyridyldithioalkyl, or isothiocyanate groups (described here) and the addition of amino, carboxyl, thiophosphate, phosphate, and thiol groups to the 5′ ends of oligodeoxyribonucleotides (UNIT 4.9). The preparation oligonucleotide-ligand conjugates by coupling a functionalized ligand to an unprotected oligonucleotide carrying a suitable functional group offers many advantages over the direct incorporation of the ligands during the oligonucleotide synthesis. In particular, this method is very useful when only a small amount of ligand is available, when the ligand does not resist the chemical conditions required for oligonucleotide deprotection, or when its poor solubility in the solvents typically used for oligonucleotide synthesis does not allow preparation of phosphoramidite or Hphosphonate derivatives. Furthermore, provided that the required amount for each conjugate is low, this method is more convenient method than the direct method for obtaining many different oligodeoxyribonucleotide-ligand conjugates starting from only one oligodeoxyribonucleotide synthesis. This unit describes the attachment of a halogenoalkyl and 2-pyridyldithioalkyl linker to a ligand for coupling with an oligodeoxyribonucleotide functionalized with either a 5′-thiophosphate or a 5′-pyridyldisulfide group. The use of a halogenoalkylated ligand yields a conjugate with a stable phosphothioloester or thioether linkage. The use of a 2-pyridyldithioalkylated ligand yields a conjugate with a disulfide bond that can be cleaved by the use of a reducing agent. The third option reported in this unit is the attachment of an isothiocyanate group to a ligand for coupling with an oligodeoxyribonucleotide that carries an aminoalkylated linker.
Procedures reported in this unit with acridine, psoralen, orthophenanthroline, and thiazole orange may be used to prepare other families of ligands. Using these procedures, the parameters of linkage between the oligodeoxyribonucleotide and the ligand—such as the size and nature of the linker used to connect the two entities—may be varied to prepare oligodeoxyribonucleotide-ligand conjugates with optimal properties. For many examples reported, the properties of the oligodeoxyribonucleotideligand conjugates are largely dependent on the geometry of the complex formed between the ligand and its target (Takasugi et al., 1991; Giovannangeli et al., 1992; Costes et al., 1993; Grigoriev et al., 1993). For example, the strongest stabilization is achieved when the ligand is an intercalator (Sun et al., 1989; Asseline et al., 1996; Giovannangeli et al., 1996), the best efficiency of cleavage when the ligand is a cleaving agent (François et al. 1989a,b), or effective cross-linking when the ligand is a reactive group (Takasugi et al., 1991).
Critical Parameters and Troubleshooting Protection of flasks and columns from light throughout the steps of these protocols is recommended since acridine, psoralen, and thiazole orange derivatives are sensitive to light. This can be achieved by wrapping flasks and columns with aluminum foil. Purification steps must also be kept as short as possible. In order to avoid low yields during purification, it is important to completely remove solvents such as dimethylformamide, pyridine, and ethanol, and to pack the columns with extreme care. The use of nonanhydrous solvents can affect the yields obtained. Storing the products in hermetically sealed vials at –20°C in the dark is recommended. Under these conditions, good reactivity of these compounds is usually observed after months or even years of storage. The bromoalkylated phenanthroline derivative, however, may show significant degradation after a few months at –20°C. Flasks and vials containing functionalized ligands should always be allowed to reach room temperature before being opened in order to
Synthesis of Modified Oligonucleotides and Conjugates
4.8.13 Current Protocols in Nucleic Acid Chemistry
Supplement 5
avoid moisture, which can cause sticky compounds (instead of solids) to be obtained. These sticky compounds are difficult to handle and have reduced stability over time. Rf values given in this unit correspond to TLC analyses performed on 2.5 × 7–cm aluminum-backed sheets developed in jars (diameter 4.5 cm, height 8 cm) with caps, using the appropriate mixtures of solvents. The use of jars and plates with different sizes can give different Rf values.
Anticipated Results The yields reported for the various steps can be different when the syntheses are performed at scales other than those described. Using the protocols provided here and starting with 138 mg of the hydroxyalkylated acridine derivative S.1b, 97 mg of the purified bromoalkylated derivative S.1e can be obtained. In the case of the psoralen derivatives, starting with 600 mg of the 5-hydroxypsoralen S.2b, it is possible to obtain 970 mg of the iodoalkylated psoralen derivative S.2f. The latter (200 mg) is then used to prepare the acetylthio derivative S.2g (163 mg), which is then transformed into the 2-pyridyldithioalkylated compound S.2h (140 mg). Starting with 1 g of 5-nitro-1,10-phenanthroline S.3a, 520 mg of the amino derivative S.3b can be obtained. Using 195 mg of the latter, it is possible to obtain 245 mg of the bromoalkylated derivative S.3c. Alternatively, starting with 97 mg of the amino derivative S.3b, 90 mg of the isothiocyanate derivative S.3d can be obtained. In the case of the thiazole orange derivative preparation, starting with 2.4 g thione S.4a, 4.04 g of S.4b can be obtained; using 0.746 g of lepidine S.4c, it is possible to prepare 1.35 g of the iodoalkylated lepidine derivative S.4d. Reaction between 0.156 g of S.4b and 0.246 g of S.4d can give 0.208 g of thiazole orange S.4e.
Time Considerations
Incorporation of Halogenoalkyl, 2Pyridyldithioalkyl, or Isothiocyanate Linkers into Ligands
Provided that all reagents and materials required for each step are available, most of the procedures are simple and rapid. Preparation of compound S.1e requires 2 to 3 days while only 2 days are necessary for compound S.2f. Compound S.2g can be obtained, starting from derivative S.2f, in 1 day; its transformation into S.2g requires 3 to 4 days. The time required for the preparation of compound S.3b is usually 3 days (including 2 days for the crystallization step), and the time required for the synthesis of S.3c and S.3d is 1 and 2 days,
respectively. The synthesis of compounds S.4b and S.4d requires 1 and 2 days, respectively. Starting from S.4b and S.4d, compound S.4e is obtained in 1 day.
Literature Cited Asseline, U., Bonfils, E., Kurfürst, R., Chassignol, M., Roig, V., and Thuong, N.T. 1992. Solidphase preparation of 5′,3′-heterobifunctional oligodeoxyribonucleotides using modified solid supports. Tetrahedron 48:1233-1254. Asseline, U., Bonfils, E., Dupret, D., and Thuong, N.T. 1996. Synthesis and binding properties of oligonucleotides covalently linked to an acridine derivative. A new study of the influence of the dye attachment site. Bioconjugate Chem. 7:369379. Benson, S.C., Singh, P., and Glazer, A.N. 1993. Heterodimeric DNA-binding dyes designed for energy transfer: Synthesis and spectroscopic properties. Nucl. Acids Res. 21:5727-5735. Brooker, L.G., Keyer, G.H., and Williams, W.W. 1942. The absorption of unsymmetrical cyanines. Resonance as a basis for classification of dyes. J. Am. Chem. Soc. 64:199-210. Chassignol, M. and Thuong, N.T. 1998. Phosphodisulfide bond: A new linker for the oligonucleotide conjugation. Tetrahedron Lett. 39:82718274. Costes, B., Girodon, E., Ghanem, N., Chassignol, M., Thuong, N.T., Dupret, D., and Goossens, M. 1993. Psoralen-modified oligonucleotide primers improve detection of mutations by denaturing gradient gel electrophoresis and provide an alternative to GC-clamping. Hum. Mol. Genet. 2:393397. François, J.-C., Saison-Behmoaras, T., Barbier, C., Chassignol, M., Thuong, N.T., and Hélène, C. 1989a. Sequence-specific recognition and cleavage of duplex DNA via triple-helix formation by oligonucleotides covalently linked to a phenanthroline-copper chelate. Proc. Natl. Acad. Sci. U.S.A. 86:9702-9706. François, J.-C., Saison-Behmoaras, T., Chassignol, M., Thuong, N.T., and Hélène, C. 1989b. Sequence-targeted cleavage of single- and doublestranded DNA by oligothymidylates covalently linked to 1,10-phenanthroline. J. Biol. Chem. 264:5891-5898. Giovannangeli, C., Thuong, N.T., and Hélène, C., 1992. Oligodeoxynucleotide-directed photo-induced cross-linking of HIV proviral DNA via triple-helix formation. Nucl. Acids Res. 20:42754281. Giovannangeli, C., Perrouault, L., Escudé C., Thuong, N.T., and Hélène, C. 1996. Specific inhibition of in vitro transcription elongation by triplex-forming oligonucleotide-intercalator conjugates targeted to HIV proviral DNA. Biochemistry 35:10539-10548.
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Current Protocols in Nucleic Acid Chemistry
Grigoriev, M., Praseuth, D., Guieysse, A.L., Robin, P., Thuong, N.T., Hélène, C., and Harel-Bellan, A. 1993. Inhibition of gene expression by triple helix-directed DNA cross-linking at specific sites. Proc. Natl. Acad. Sci. U.S.A. 90:35013505. Sun, J.S., François, J.C., Montenay-Garestier, T., Saison-Behmoaras, T., Roig, V., Thuong, N.T., and Hélène, C. 1989. Sequence-specific intercalating agents. Intercalation at specific sequences on duplex DNA via major groove recognition by oligonucleotide-intercalator conjugates. Proc. Natl. Acad. Sci. U.S.A. 86:9198-9202.
Takasugi, M., Guendouz, A., Chassignol, M., Decout, J.L., Lhomme, J., Thuong, N.T., and Hélène, C., 1991. Sequence-specific photo-induced cross-linking of the two strands of doublehelical DNA by a psoralen covalently linked to a triple helix forming oligonucleotide. Proc. Natl. Acad. Sci. U.S.A. 88:5602-5606. Thuong, N.T. and Asseline, U. 1991. Oligonucleotides attached to intercalators, photoreactive and cleavage agents. In Oligonucleotides and Analogues: A Practical Approach (F. Eckstein, ed.) pp. 283-308. IRL Press, Oxford.
Contributed by Ulysse Asseline and Nguyen T. Thuong Centre de Biophysique Moléculaire, CNRS Orléans, France
The authors would like to express their appreciation to their past and present collaborators for their contribution to the development of various families of oligonucleotide-ligand conjugates. This work was supported by Rhône-Poulenc, the Agence Nationale de Recherches sur le SIDA and bio-Mérieux.
Synthesis of Modified Oligonucleotides and Conjugates
4.8.15 Current Protocols in Nucleic Acid Chemistry
Supplement 5
Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
UNIT 4.9
The chemical attachment of reporter and various conjugate groups (e.g., intercalators as well as photoreactive and cleaving agents) to the 5′ terminus of oligodeoxyribonucleotides can be achieved following two strategies. The first involves the direct addition of a ligand to the 5′ end of the oligodeoxyribonucleotide via its phosphoramidite or H-phosphonate derivative. This strategy has been described in UNIT 4.3. The second strategy involves the incorporation of appropriate functional groups into both the ligand and the 5′ terminus of an oligonucleotide. Specific coupling of these reactants results in the formation of oligodeoxyribonucleotide conjugates. Using the second strategy, many different oligodeoxyribonucleotide conjugates can be prepared starting from only one oligodeoxyribonucleotide, provided that the amount required for each conjugate is low. In this approach, functional groups (such as amino, phosphate, phosphorothioate, thiol, carboxyl, and cis-diol) that are capable of undergoing specific reactions with selected ligand functional groups are added to the 5′ end of oligodeoxyribonucleotides. Examples of incorporation of halogenoalkyl, 2-pyridyldithioalkyl, or isothiocyanate linkers into ligands have been reported in UNIT 4.8. This unit describes methods used in the authors’ laboratory for the preparation of oligodeoxyribonucleotides carrying the following 5′-terminal functional groups: carboxyl and amino (see Basic Protocol 1), phosphorothioate and phosphate (see Basic Protocol 2), and masked thiol (see Basic Protocol 3; see Alternate Protocols 1 and 2). The functional groups are incorporated upon completion of oligonucleotide chain assembly prior to deprotection and purification. Another strategy—adding functional groups after deprotection and purification—has been described elsewhere (Grimm et al., 2000). The coupling of these functionalized oligonucleotides with the reactive functions incorporated into the ligands reported in UNIT 4.8, and finally the purification and characterization of the oligodeoxyribonucleotide-ligand conjugates, is described in UNIT 4.10. CAUTION: All chemicals must be handled in a fume hood by personnel equipped with a laboratory coat, glasses, and gloves. NOTE: For each family of 5′-oligonucleotide modifications, a sample of detritylated oligodeoxyribonucleotide bound to the support is deprotected and purified separately using the general conditions described in APPENDIX 3C. These samples are used as references for chromatography and polyacrylamide gel electrophoresis analyses. ADDITION OF A CARBOXYLATED OR AMINOALKYLATED LINKER TO THE 5′ END OF OLIGODEOXYRIBONUCLEOTIDES
BASIC PROTOCOL 1
The addition of carboxylated or aminoalkylated linkers at the 5′ end of oligodeoxyribonucleotides is achieved via a two-step procedure adapted from Gottikh et al. (1990) and Wachter et al. (1986), respectively. The different steps involved are illustrated in Figure 4.9.1. In these procedures, the 5′-hydroxyl group of an oligonucleotide bound to a support (S.1) is activated by treatment with carbonyldiimidazole followed by reaction with either an amino acid (aminovaleric acid or aminocaproic acid) or a bis amine (1,5-diaminopentane or 1,6-diaminohexane). After deprotection and cleavage from the support, the oligodeoxyribonucleotides containing carboxylated or aminoalkylated linkers (S.3 and S.2, respectively) are purified by chromatography. Synthesis of Modified Oligonucleotides and Conjugates Contributed by Ulysse Asseline and Nguyen T. Thuong Current Protocols in Nucleic Acid Chemistry (2001) 4.9.1-4.9.28 Copyright © 2001 by John Wiley & Sons, Inc.
4.9.1 Supplement 5
H2N(CH2)mNHC(O)O
B
O O
1. Carbonyldiimidazole
O P O 2. H2N(CH2)mNH2, m = 5, 6 3. Deprotection
−
O
O
B
O HO
B'
O
n
H 2
O O P OCH2CH2CN O
B'
O O
n
P
−
1
O2C(CH2)mNHC(O)O
1. Carbonyldiimidazole
−
2. H2N(CH2)mCO2 DBU salt, m = 4, 5
B
O O
O P O
−
O
O
3. Deprotection
B
O H
n
3
Figure 4.9.1 Addition of a carboxyl or amino function to the 5′ end of an oligodeoxyribonucleotide. B, adenine, cytosine, guanine, or thymine; B′, thymine or any N-protected nucleobase; DBU, 1,8-diazabicyclo[5.4.0]undec-7-ene; P, controlled-pore glass.
Materials 5′-Detritylated oligodeoxyribonucleotide bound to a controlled-pore glass (CPG) support Nitrogen gas 40 mg/mL anhydrous 1,1′-carbonyldiimidazole in anhydrous dioxane Anhydrous dioxane Amino acid or bis amine (select one): 10 mg/mL 5-aminovaleric acid or 6-aminocaproic acid, 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) salt (Aldrich), in pyridine 12 mg/mL 1,5-diaminopentane or 1,6-diaminopentane in pyridine Pyridine, redistilled from p-toluenesulfonyl chloride, stored over 3A molecular sieves Acetonitrile (CH3CN), DNA synthesis grade, stored over 3A molecular sieves, and HPLC grade Concentrated ammonium hydroxide (25%) Ethyl acetate, distilled 1.5 M NaCl 25 mM Tris⋅Cl, pH 7 (APPENDIX 2A), containing 10% (v/v) distilled HPLC-grade methanol 1 M triethylammonium acetate (TEAA) buffer, pH 7 (stock solution) Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
8-mL vial with septum and screw cap 22-G hypodermic needle Desiccator containing P2O5 and KOH 50° and 55°C ovens or water baths
4.9.2 Supplement 5
Current Protocols in Nucleic Acid Chemistry
0.45-µm filter attached to a disposable syringe UV spectrophotometer Rotary evaporator with water bath and a water aspirator Ion-exchange chromatography system and column (select one): Mono Q HR 5/5 or HR 10/10 column (Amersham Pharmacia Biotech) DEAE column (8 µm, 100 × 10 mm; Waters) High-performance liquid chromatograph (HPLC) equipped with multiwavelength detector and reversed-phase column (select one): Lichrospher 100 RP 18 column (5 µm; 125 mm × 4 mm; Merck) Lichrospher 100 RP 18 column (10 µm; 250 mm × 10 mm; Merck) Delta-Pak C4 column (5 µm, 100 Å; 150 × 3.9 mm; Waters) Lyophilizer Additional reagents and equipment for analytical and preparative ion-exchange chromatography and reversed-phase HPLC (RP-HPLC), and for sample purification (UNIT 4.3) Activate 5′-hydroxyl group 1. Place 1 µmol of 5′-detritylated oligodeoxyribonucleotide bound to a CPG support in an 8-mL vial with a septum. Push a 22-G hypodermic needle through the septum and place the vial in a desiccator containing P2O5 and KOH. Keep under vacuum overnight. 2. Fill the desiccator with nitrogen before opening it. 3. Add 500 µL of 40 mg/mL anhydrous 1,1′-carbonyldiimidazole in anhydrous dioxane under nitrogen and allow to react for 25 min with occasional shaking. 4. Remove excess 1,1′-carbonyldiimidazole solution with a syringe and wash the support with 1 mL anhydrous dioxane. Add linker 5. Add 500 µL of one of the following solutions and incubate 3 to 4 hr with occasional shaking by hand. 10 mg/mL 5-aminovaleric acid or 6-aminocaproic acid (DBU salt) in pyridine (for S.3) 12 mg/mL 1,5-diaminopentane or 1,6-diaminopentane in pyridine (for S.2). Deprotect and remove from support 6. Remove liquid with a syringe. Wash support twice with 1 mL of 2:1 (v/v) pyridine in DNA synthesis–grade CH3CN, and twice with 1 mL CH3CN. 7. Add 5 mL concentrated ammonium hydroxide, seal the vial with a screw cap, and heat 1 hr at 50°C. 8. Cool the ammoniacal solution to room temperature. The solution should be cooled before opening the vial for safety and to prevent spillage and loss of product.
9. Discard the support, transfer the ammoniacal solution to a new vial, and incubate for an additional 6 hr at 55°C. 10. Cool solution to room temperature and evaporate to dryness under reduced pressure using a rotary evaporator with a water aspirator.
Synthesis of Modified Oligonucleotides and Conjugates
4.9.3 Current Protocols in Nucleic Acid Chemistry
Supplement 5
11. Solubilize crude material with 6 mL water and extract three times with 4 mL ethyl acetate. Discard the organic phase. 12. Filter solution using a 0.45-µm filter attached to a disposable syringe. Wash filter with 0.3 mL water and add wash to filtrate. 13. Dilute 25 µL crude deprotected oligonucleotide solution with 975 µL water and record the UV spectrum between 220 and 400 nm using a UV spectrophotometer. Determine absorbance at 260 nm and multiply by 40 (the dilution factor) to determine the concentration of the crude oligonucleotide solution. Analyze quality of synthesis 14. Analyze by ion-exchange chromatography and RP-HPLC. Set the instrument for detection at 260 nm. Preequilibrate columns with starting buffer for ≥15 min. Run a blank gradient before analysis and purification, and between analyses and purifications of samples with different sequences or different modifications. Analyze the unmodified oligonucleotide sequence for comparison. a. For ion-exchange chromatography: Use a Mono Q HR 5/5 or HR 10/10 column or a DEAE column with a linear gradient of 1.5 M NaCl (0% to 100%) in 25 mM Tris⋅Cl, pH 7, containing 10% (v/v) methanol. Elute at a flow rate of 1 mL/min with an HR 5/5 column or DEAE column, or at 4 mL/min with an HR 10/10 column. b. For RP-HPLC: Use a Lichrospher 100 RP 18 column with a linear gradient of HPLC-grade CH3CN (5% to 80%) in 0.1 M TEAA buffer, pH 7, at a flow rate of 1 mL/min. Alternatively, use a semipreparative Lichrospher 100 RP 18 column at a flow rate of 4 mL/min, or a Delta-Pak C4 column at a flow rate of 0.8 mL/min. The gradients must be chosen to afford the best separation. Usually 1 M NaCl is increased at 1% per minute for ion-exchange chromatography, and CH3CN is increased at 0.5% to 1% per minute for RP-HPLC. Characterization data can be found following step 17.
Purify sample 15. Using the system determined to give the best separation, purify the remaining crude 5′-modified oligonucleotide by preparative reversed-phase or ion-exchange chromatography. Because preparative chromatography columns do not have the resolution of analytical columns, the product peak should be fractionated to obtain good purification.
16. Proceed with sample purification as reported in UNIT 4.3 (steps 3 to 8 of the Support Protocol). Measure the OD260 to determine the yield of 5′-modified oligonucleotide. 17. Lyophilize the modified oligonucleotide and store up to two years at –20°C in a tightly sealed vial. Characterization of carboxylated oligodeoxyribonucleotides: Yield: 25 OD260 units.
Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
The best separation was obtained by ion-exchange chromatography on a Mono Q column (HR 50 × 5 mm) using a linear gradient of NaCl (0.2 to 0.5 M over 40 min) in 25 mM Tris⋅Cl, pH 7, containing 10% methanol at a flow rate of 1 mL/min. Retention times were 22 min for –OOC(CH2)4-NH-CO-d[CTCTCGCACCCATCTCTC] and –OOC(CH2)5-NHCO-d[CTCTCGCACCCATCTCTC], and 21 min for d[CTCTCGCACCCATCTCTC]. Ionexchange analysis of a crude oligonucleotide derivatized with aminovaleric acid is shown in Figure 4.9.2.
4.9.4 Supplement 5
Current Protocols in Nucleic Acid Chemistry
Figure 4.9.2 Ion-exchange chromatography analysis of crude HOOC(CH2)4-NH-COd[CTCTCGCACCCATCTCTC] (see Basic Protocol 1) on a Mono Q column.
RP-HPLC of carboxylated oligonucleotides on a Lichrospher 100 RP 18 (5 mm; 125 × 4 mm) column using a gradient of CH3CN (10% to 30% over 50 min) in 0.1 M TEAA buffer, pH 7, at 1 mL/min gives retention times of 11 min 25 sec for –OOC(CH2)4-NH-COd[CTCTC GC AC CCATCTCTC], 13 min 13 sec for –OOC(CH2)5-NH-COd[CTCTCGCACCCATCTCTC], and 9 min 40 sec for d[CTCTCGCACCCATCTCTC]. RP-HPLC performed on a Delta-Pak C4 (5 mm, 100 Å, 150 × 3.9 mm) column with a linear gradient of CH3CN (5% to 35% over 60 min) in 0.1 M TEAA buffer, pH 7, at 0.8 mL/min gives retention times of 12 min 10 sec for –OOC(CH2)5-NH-COd[CTCTCGCACCCATCTCTC], and 11 min 8 sec for d[CTCTCGCACCCATCTCTC]. Figure 4.9.3 shows a chromatogram obtained for a mixture of purified –OOC(CH2)5-NHCO-d[CTCTCGCACCCATCTCTC] and d[CTCTCGCACCCATCTCTC].
Figure 4.9.3 RP-HPLC analysis on a Delta Pak C4 column of a purified mixture of HOOC(CH2)5NH-CO-d[CTCTCGCACCCATCTCTC] and d[CTCTCGCACCCATCTCTC] (see Basic Protocol 1).
Synthesis of Modified Oligonucleotides and Conjugates
4.9.5 Current Protocols in Nucleic Acid Chemistry
Supplement 5
Mass analysis. ESI-MS polarity negative. HOOC(CH2)5-NH-COd[CTCTCGCACCCATCTCTC]. Calcd. for C177H233N56O112P17: 5461 Da; found: 5460 ± 3 Da (M-H). HOOC(CH2)4-NH-CO-d[CTCTCGCACCCATCTCTC] Calcd. for C176H231N56O112P17: 5447 Da; found: 5446 Da (M-H). Denaturing 20% PAGE does not allow easy separation of 5′-modified oligonucleotides with carboxylated linkers from the corresponding unmodified oligonucleotide. Characterization of aminoalkylated oligodeoxyribonucleotides: Yield: 25-30 OD260 units. Ion-exchange chromatography of 5′-aminoalkylated oligonucleotides on a DEAE column (8 mm, 100 × 10 mm) using a linear gradient of NaCl (0 to 0.6 M over 60 min) in 25 mM Tris⋅Cl, pH 7, containing 10% methanol at 1 mL/min gives retention times of 44 min 54 sec for H2N(CH2)5-NH-CO-d[CCGCTTAATACTGA] and H2N(CH2)6-NH-COd[CCGCTTAATACTGA], and 45 min 48 sec for d[CCGCTTAATACTGA]. Figure 4.9.4 shows a chromatogram obtained for crude H2N(CH2)5-NH-CO-d[CCGCTTAATACTGA]. RP-HPLC on a Lichrospher 100 RP 18 column using the conditions described above gives retention times of 10 min 25 sec for H2N(CH2)5-NH-CO-d[CCGCTTAATACTGA], 10 min 54 sec for H2N(CH2)6-NH-CO-d[CCGCTTAATACTGA], and 9 min 42 sec for d[CCGCTTAATACTGA]. RP-HPLC on a Delta-Pak C4 column (5 mm, 100 Å, 150 × 3.9 mm) with a linear gradient of CH3CN (5% to 35% over 60 min) in 0.1 M TEAA buffer, pH 7, at 0.8 mL/min gives retention times of 11 min 40 sec for H2N(CH2)5-NH-CO-d[CCGCTTAATACTGA], 12 min 21 sec for H2N(CH2)6-NH-CO-d[CCGCTTAATACTGA], and 11 min 15 sec for d[CCGCTTAATACTGA]. Figure 4.9.5 shows a chromatogram for a mixture of purified H2N(CH2)6-NH-CO-d[CCGCTTAATACTGA] and d[CCGCTTAATACTGA]. Denaturing 20% PAGE (APPENDIX 3B) allows easy separation of 5′-modified oligonucleotides with aminoalkylated linkers from the corresponding unmodified oligonucleotide (Fig. 4.9.6). Mass analysis. ESI-MS polarity negative. H2N(CH2)5-NH-CO-d[CCGCTTAATACTGA]. Calcd. for C142H185N52O83P13: 4351 Da; found: 4351 ± 1 Da (M-H). H2N(CH2)6-NH-COd[CCGCTTAATACTGA]. Calcd. for C143H187N52O83P13: 4365 Da; found: 4365 ± 1 Da (M-H).
Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
Figure 4.9.4 Ion-exchange chromatography on a DEAE column of crude H2N-(CH2)5-NH-COd[CCGCTTAATACTGA] (see Basic Protocol 1).
4.9.6 Supplement 5
Current Protocols in Nucleic Acid Chemistry
Figure 4.9.5 RP-HPLC on a Delta Pak C4 column of a purified mixture of H2N-(CH2)6-NH-COd[CCGCTTAATACTGA] and d[CCGCTTAATACTGA] (see Basic Protocol 1).
ADDITION OF A PHOSPHOROTHIOATE OR PHOSPHATE GROUP TO THE 5′ TERMINUS OF OLIGODEOXYRIBONUCLEOTIDES
BASIC PROTOCOL 2
The phosphorothioate group is incorporated at the 5′ end of oligodeoxyribonucleotides via a two-step procedure involving the coupling of bis-(2-cyanoethyl)-diisopropylamidophosphite with the 5′-terminal hydroxyl function of an oligodeoxyribonucleotide bound to a support (S.4), followed by a sulfurization step, as seen in the bottom of Figure 4.9.7. The phosphate group is obtained by replacing the sulfurization step with a standard aqueous iodine oxidization (top). After deprotection and cleavage from the support, the 5′-phosphorothioate-containing (S.6) or 5′-phosphate-containing (S.5) oligodeoxyribonucleotide is purified by liquid chromatography. The procedure was adapted from a previously described procedure (Thuong and Asseline, 1991).
Figure 4.9.6 PAGE analysis of d[CCGCTTAATACTGA] (lane 1), H2N-(CH2)5-NH-COd[CCGCTTAATACTGA] (lane 2), and H2N-(CH2)6-NH-CO-d[CCGCTTAATACTGA] (lane 3; see Basic Protocol 1).
Synthesis of Modified Oligonucleotides and Conjugates
4.9.7 Current Protocols in Nucleic Acid Chemistry
Supplement 5
O HO P O O
−
B
O O
1. i-Pr2NP(OCH2CH2CN)2,1H-Tetrazole, MeCN
O P O 2. Oxidation 3. Deprotection
−
O
O
B
O HO
B'
O
n
H 5
O O P OCH2CH2CN O
B'
O O
n
P 4
O HO P O S
−
B
O O
1. i-Pr2NP(OCH2CH2CN)2,1H-Tetrazole, MeCN
O P O
2. Sulfurization 3. Deprotection
−
O
O
B
O H
n
6
Figure 4.9.7 Addition of a phosphorothioate or phosphate group to the 5′ end of an oligodeoxyribonucleotide.
Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
Materials Diisopropylethylamine, distilled from KOH 2-Cyanoethanol Diethyl ether dried over sodium wires Nitrogen source N,N′-Diisopropylphosphoramidous dichloride (Aldrich) Argon atmosphere 5′-Detritylated oligodeoxyribonucleotide bound to a controlled-pore glass (CPG) support 0.5 M tetrazole in anhydrous CH3CN Anhydrous CH3CN 10 mg/mL Beaucage reagent in anhydrous CH3CN or 100 mg/mL tetraethylthiuram disulfide in anhydrous CH3CN (optional; for sulfurization) Iodine solution (same composition as for DNA synthesis; optional; for oxidation) Concentrated ammonium hydroxide (NH4OH; 25%) Dithiothreitol (DTT; optional) Ethyl acetate, distilled Isopropanol 2.5 mg/mL 2,6-dibromo-4-benzoquinone-N-chloroimine (DBPNC; Prolabo) in ethanol 1-liter three-neck round-bottom flask Dropping funnel Reflux condenser with a calcium chloride drying tube
4.9.8 Supplement 5
Current Protocols in Nucleic Acid Chemistry
Gas inlet adapter Glass filter (porosity 4) Rotary evaporator with a water aspirator and a water bath Falling-film distillation head (Aldrich) or other vacuum distillation apparatus 50° and 55°C ovens or water baths Kieselgel 60F plates for analytical TLC (Merck) Additional reagents and equipment for automated oligonucleotide synthesis (APPENDIX 3C), thin-layer chromatography (TLC; APPENDIX 3D), and purification and characterization of the product (see Basic Protocol 1) Prepare bis-(2-cyanoethyl)diisopropylamidophosphite 1. Place 70 mL (51.6 g, 0.40 mol) diisopropylethylamine and 25.6 g (0.36 mol) 2-cyanoethanol in 300 mL dry diethyl ether in a three-neck 1-liter round-bottom flask equipped with a dropping funnel, a reflux condenser with a calcium chloride drying tube, and a gas inlet adapter connected to a nitrogen source. Cool to 0°C while stirring vigorously with a magnetic stir bar or by mechanical stirring. 2. Add 36 g (33.18 mL, 0.18 mol) N,N-diisopropylphosphoramidous dichloride to 50 mL dry diethyl ether. Add dropwise, under an argon atmosphere, to the solution in step 1 using a dropping funnel. 3. Allow the stirred mixture to warm to room temperature for 1 hr. 4. Remove the precipitated salt by vacuum filtration with a glass filter and wash it with 100 mL dry diethyl ether. Concentrate filtrate by evaporation under reduced pressure using a rotary evaporator with a water aspirator. 5. Purify the residue by vacuum distillation. An apparatus can be fashioned in the laboratory, or a falling-film distillation head can be used. Yield 60% (29.3 g, 0.108 mol). 1H-NMR (CDCl3), δ: 1.19 [d, 12H, J = 6.8 Hz, (CH3)2CH]; 2.64 [t, J = 6.2 Hz, (OCH2CH2CN)]; 3.55-3.68 [m, 2H, (CH3)2CH]; 3.78-3.94 [4H, m, OCH2CH2CN)]. 31P-NMR (CDCl3), δ: 150.2 ppm. 13C-NMR (DMSO-d6), δ: 25.25, 29.78, 48.06 (d, J = 36.5 Hz), 63.84 (d, J = 73 Hz), 124.36. Mass analysis. ESI-MS, polarity positive. Calcd. for C12H22N3O2P: 271; found: 272 (M+H). The purified product can be stored up to 1 year at –20°C without loss of coupling efficiency. However, when the reaction is performed on a small scale, bis-(2-cyanoethyl)diisopropylamidophosphite is purified by chromatography on a silica gel column using 4:1 (v/v) diethyl ether/hexane containing 2% to 3% triethylamine as the eluent, and the fractions are monitored by TLC using the same eluent (pre-elute the plate before analysis). After treating the plate with DBPNC (step 13), a pale-colored spot appears (Rf = 0.6). When purified this way, the product shows loss of coupling efficiency after 3 to 4 months of storage. Bis-(2-cyanoethyl)diisopropylamidophosphite is now commercially available from Chemgenes.
Couple to oligonucleotide 6. Place 0.1 M bis-(2-cyanoethyl)diisopropylamidophosphite in anhydrous CH3CN on the synthesizer. React a fully 5′-detritylated oligodeoxyribonucleotide bound to a CPG support for 5 min at a flow rate of 1 mL/min by recycling a mixture containing: 0.1 mL 0.1 M phosphoramidite in anhydrous CH3CN 0.5 mL 0.5 M tetrazole in anhydrous CH3CN 0.1 mL CH3CN.
Synthesis of Modified Oligonucleotides and Conjugates
4.9.9 Current Protocols in Nucleic Acid Chemistry
Supplement 7
Perform this reaction twice. For synthesizers without a recycle program, perform a double coupling step. Alternatively, bis-(2-cyanoethyl)diisopropylamidophosphite can be manually added to the 5′ terminus of oligonucleotides bound to a solid support according to the method described for the phosphoramidites 1c and 2d in UNIT 4.3.
Perform sulfurization/oxidation 7a. For sulfurization: Treat product with 1 mL of 10 mg/mL Beaucage reagent in anhydrous CH3CN added at a rate of 1 mL/min, or with 15 mL of 100 mg/mL tetraethylthiuram disulfide in CH3CN added at 1 mL/min. 7b. For oxidation: Perform automated iodine oxidation step. Either the sulfurization or the oxidation step can be performed manually.
Deprotect and remove from support 8. Add 5 mL concentrated ammonium hydroxide and 100 eq (15 mg) DTT to the oligodeoxyribonucleotide 5′-bis-(2-cyanoethyl)phosphorothioate and heat 1 hr at 55°C. For the oligodeoxyribonucleotide 5′-bis-(2-cyanoethyl)phosphate, omit the DTT. DTT is used to prevent formation of an acrylonitrile adduct on the thiophosphate group.
9. Cool the ammoniacal solution to room temperature. The solution should be cooled before opening the vial for safety and to prevent spillage and loss of product.
10. Discard the support, transfer the ammoniacal solution to a new vial, and incubate for an additional 7 hr at 50°C. 11. Cool the ammoniacal solution and evaporate it to dryness under reduced pressure using a rotary evaporator and a water aspirator. 12. Solubilize the crude material with 6 mL water and extract four times with 4 mL ethyl acetate for the 5′-phosphorothioate, or three times for the 5′-phosphate. 13. For medium-sized oligodeoxyribonucleotides (10- to 25-mers), verify the presence of the phosphorothioate group by analyzing the crude deprotected oligodeoxyribonucleotide by analytical TLC (APPENDIX 3D) using isopropanol/concentrated NH4OH/H2O as eluent in varying proportions depending on the length and base composition of the oligodeoxyribonucleotide. Spray the plate with 2.5 mg/mL DBPNC in ethanol and heat until color appears. Typically, 65:9:15 (v/v/v) isopropanol/NH4OH/H2O is suitable for ≤10-mers, and 55:10:25 or 55:10:35 is suitable for 10- to 20-mers. Phosphorothioate-containing oligodeoxyribonucleotides give a pink-colored spot on the TLC plate after treatment with DBPNC and heating. (Eluent mixture must be prepared at least a half-day before use.)
Purify and characterize 14. Purify and characterize the 5′-phosphorothioate- or 5′-phosphate-containing oligodeoxyribonucleotide as described (see Basic Protocol 1, steps 12 to 17).
Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
Ion-exchange chromatography of 5′-phosphorothioate- and 5′-phosphate-containing oligodeoxyribonucleotides using the conditions described above with a DEAE column gives retention times of 49 min 24 sec for sp-d[CCGCTTAATACTGA] (53 min 36 sec for oligonucleotide disulfide), 47 min 36 sec for p-d[TTCTCCCCCGCTTA], 46 min 12 sec for d[TTCTCCCCCGCTTA], and 45 min 48 sec for d[CCGCTTAATACTGA]. Figure 4.9.8
4.9.10 Supplement 7
Current Protocols in Nucleic Acid Chemistry
Figure 4.9.8 Ion-exchange chromatography on a DEAE column of crude sp-d[CCGCTTAATACTGA] (see Basic Protocol 2).
shows the chromatogram of crude sp-d[CCGCTTAATACTGA], and Figure 4.9.9 shows the chromatogram of crude p-d[TTCTCCCCCGCTTA]. Denaturing 20% PAGE does not allow easy separation of oligonucleotide 5′-thiophosphates and 5′-phosphates from the corresponding unmodified oligonucleotide. In the case of 5′-thiophosphate oligonucleotides, however, the presence of a weak band with a mobility corresponding to that of a 30-mer confirmed that the peak observed at 53 min 36 sec is probably the chimeric oligonucleotide disulfide 3′-[AGTCATAATTCGCC]-ps-spd[CCGCTTAATACTGA]-3′. The 5′-thiophosphate oligonucleotide can be recovered by treatment of the chimera with DTT. RP-HPLC on a Lichrospher RP 18 column using the conditions described above gives retention times of 9 min for sp-d[CCGCTTAATACTGA], 9 min 42 sec for d[CCGCTTAATACTGA], 10 min 30 sec for p-d[TTCTCCCCCGCTTA], and 10 min 50 sec for d[TTCTCCCCCGCTTA].
Figure 4.9.9 Ion-exchange c hromatography on a DEAE column of cr ude pd[TTCTCCCCCGCTTA] (see Basic Protocol 2).
Synthesis of Modified Oligonucleotides and Conjugates
4.9.11 Current Protocols in Nucleic Acid Chemistry
Supplement 5
Figure 4.9.10 RP - HP LC on a Delta Pak C4 column of a pur ified mixture of pd[TTCTCCCCCGCTTA] and d[TTCTCCCCCGCTTA] (see Basic Protocol 2).
RP-HPLC on a Delta-Pak C4 column (5 mm, 100 Å, 150 × 3.9 mm) using a linear gradient of CH3CN (5% to 16 % by volume over 30 min) in 0.1 M TEAA buffer, pH 7, at 0.8 mL/min gives retention times of 10 min 40 sec for sp-d[CCGCTTAATACTGA], 11 min 15 sec for d[CCGCTTAATACTGA], 11 min 36 sec for p-d[TTCTCCCCCGCTTA], and 12 min 26 sec for d[TTCTCCCCCGCTTA]. Figure 4.9.10 shows a chromatogram obtained for a mixture of purified p-d[TTCTCCCCCGCTTA] and d[TTCTCCCCCGCTTA]. Mass analysis. ESI-MS polarity negative. sp-d[CCGCTTAATACTGA]. Calcd. for C136H175N50O84P14S: 4319 Da; found: 4318 Da (M-H). Yield: 35-40 OD260 units. Mass analysis. ESI-MS polarity negative. p-d[TTCTCCCCCGCTTA]. Calcd. for C133H176N41O89P14: 4206 Da; found: 4205 ± 1 Da (M-H). Yield: 35-40 OD260 units. BASIC PROTOCOL 3
ADDITION OF A MASKED THIOL GROUP TO THE 5′ TERMINUS OF OLIGODEOXYRIBONUCLEOTIDES USING AN S-DIPHENYLPHOSPHINATE PHOSPHORAMIDITE Two strategies can be used to add a masked thiol group to the 5′ end of an oligodeoxyribonucleotide. This protocol presents the first strategy, which is illustrated on the left of Figure 4.9.11. In this approach, an S-diphenylphosphinate phosphoramidite (S.7a) is added to the 5′ terminus of a support-bound oligodeoxyribonucleotide. This gives a 5′ thiol group masked with an alkali-labile protecting group (S.7b). Addition of 2,2′-dithiodipyridine during oligonucleotide deprotection converts the released thiol function to a 2-pyridyldisulfide group (S.10). This strategy can also be performed using an S-acetyl phosphoramidite (S.8a) as described in Alternate Protocol 1. The second strategy (right side of Fig. 4.9.11) uses a tritylated alkyl disulfide-containing linker (S.9a) and is described in Alternate Protocol 2.
Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
The preparation of the S-diphenylphosphinate derivative S.7a is included in the steps below and is illustrated in Figure 4.9.12. The synthesis of S.7a is achieved in four steps from the commercial methyldiphenylphosphinite S.11. Sulfurization of S.11 gives the methyldiphenylthionophosphinate S.12, which is then demethylated by treatment with trimethylamine. The resulting tetramethylammonium salt S.13 reacts with 2-[2-(2-io-
4.9.12 Supplement 5
Current Protocols in Nucleic Acid Chemistry
i-Pr2N P 1.
L
HO
OCH2CH2CN
B'
O
i-Pr2N
O
RS
1. DMTrO 7a R = Ph2P(O) 8a R = CH3CO
L
O
L
O
P
OCH2CH2CN
9a
O P OCH2CH2CN
1H-tetrazole 2. Oxidation
S S
O
1H-tetrazole 2. Oxidation
B'
O O
n
P
O
O RS
L
O P OCH2CH2CN O
B'
O
DMTrO
O DMTrO
L
S S
O P O L
−
S S
B
O
O
L
O
O
−
O
NH4OH O
B
O P OCH2CH2CN O
B'
O
n
H
n
7b R = Ph2P(O) 8b R = CH3CO
B'
O
O
O P
O
O
O P O
B'
O P OCH2CH2CN O
O
O O P OCH2CH2CN
L
P
n
9b
9c
DTT or TCEP, PySSPy
O N
S S
NH4OH Phenol PySSPy
O P O L
−
B
O
O
O O P O
−
O
O
B
O H
n
10
Figure 4.9.11 Addition of a masked thiol group to the 5′ end of an oligodeoxyribonucleotide. DTT, dithiothreitol; L, CH2CH2OCH2CH2OCH2CH2; PySSPy, 2,2′-dipyridyldisulfide; TCEP, tris(2-carboxyethyl)phosphine hydrochloride.
doethoxy)ethoxy]ethanol to give 2-{2-[2-(S-diphenylphosphinate)ethoxy]ethoxy}ethanol S.14. 2-[2-(2-Iodoethoxy)ethoxy]ethanol is obtained by treatment of 2-[2-(2-chloroethoxy)ethoxy]ethanol (S.15; Fig. 4.9.16) with NaI in acetone in the presence of NaHCO3. The alcohol S.14 is then treated with diisopropylethylamine and 2-cyanoethylN,N-diisopropylchlorophosphoramidite to give the phosphoramidite S.7a. Materials Methyldiphenylphosphinite (S.11; Aldrich) Toluene Elemental sulfur (S8) Hexane, distilled Ethyl acetate, distilled Triethylamine, distilled from KOH 2.5 mg/mL 2,6-dibromo-4-benzoquinone-N-chloroimine (DBPNC; Prolabo) in ethanol
Synthesis of Modified Oligonucleotides and Conjugates
4.9.13 Current Protocols in Nucleic Acid Chemistry
Supplement 5
O
S S8
P OMe
P
11
OMe
NMe3
P
− + NMe4 S
13
12 OH I
L
i-Pr2N P
O P
L S
OCH2CH2CN
O
O
i-Pr2NP(Cl)OCH2CH2CN
P
L
OH
S
DIPEA
7a
14
Figure 4.9.12 Preparation of the S-diphenylphosphinate phosphoramidite derivative S.7a. DIPEA, diisopropylethylamine; L, CH2CH2OCH2CH2OCH2CH2.
Celite 521 (Aldrich) 20% (w/v) trimethylamine/acetonitrile Diethyl ether, dried over sodium wires, ice cold and room temperature Dichloromethane (CH2Cl2) distilled over P2O5 and passed through a column of basic alumina Methanol, distilled 2-[2-(2-Chloroethoxy)ethoxy]ethanol (S.15; Fig. 4.9.16; Aldrich) NaI Sodium bicarbonate (NaHCO3) Acetone, anhydrous Anhydrous acetonitrile (CH3CN) Diisopropylethylamine, distilled from KOH 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite (Aldrich) Nitrogen gas 10% (w/v) aqueous sodium carbonate (Na2CO3) Saturated aqueous NaCl, ice cold Sodium sulfate, anhydrous 5′-Detritylated oligodeoxyribonucleotide bound to a controlled-pore glass (CPG) support 2,2′-Dithiodipyridine Phenol Concentrated ammonium hydroxide 1 M triethylammonium acetate (TEAA) buffer, pH 7 (stock solution) Tris-(2-carboxyethyl)phosphine (TCEP), hydrochloride
Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
250-mL three-necked round-bottom flask Reflux condenser Magnetic stirrer with heating element Kieselgel 60F plates for analytical TLC (Merck) Rotary evaporator with a water aspirator Calcium chloride drying tube
4.9.14 Supplement 5
Current Protocols in Nucleic Acid Chemistry
3 × 50–cm chromatography column containing 40 g silica gel (e.g., Kieselgel 60; Merck) and 1.6 × 45–cm column containing 25 g silica gel 10-mL vial and stoppers Separatory funnel Spectrophotometer Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D); column chromatography (APPENDIX 3E); direct addition of an acridinyl phosphoramidite (UNIT 4.3); and analysis, purification, and characterization of product (see Basic Protocol 1) Prepare diphenylthiophosphinate salt S.13 1. Place 3 g (13.8 mmol) methyldiphenylphosphinite S.11 and 60 mL toluene in a 250-mL three-necked round-bottom flask equipped with a reflux condenser and a magnetic stir bar. Add 883 mg (27.6 mmol) S8 slowly to the stirring solution at room temperature. The reaction is slightly exothermic.
2. Heat suspension at 50°C and monitor reaction by analytical TLC (APPENDIX 3D) and UV shadowing using 90:10:4 (v/v/v) hexane/ethyl acetate/triethylamine as eluent. Spray the plate with 2.5 mg/mL DBPNC in ethanol and heat until color appears. The sulfur-containing compound S.12 (Rf = 0.43) gives a pink-colored spot on the TLC plate after treatment with DBPNC. The starting material S.11 (Rf = 0.70) appears as a white spot.
3. After the reaction is complete (1 to 2 hr), allow reaction mixture to cool to room temperature, filter through celite 521, and concentrate under reduced pressure using a rotary evaporator with a water aspirator. Yield 85% (2.92 g, 11.78 mmol). White solid. 31P-NMR (CDCl3), δ: 85.85 ppm. 1H-NMR (CDCl3), δ: 3.75 (d, 3H, J = 12.2 Hz, OCH3), 7.41-7.53 (m, 6H, Ar), 7.81-7.91 (m, 4H, Ar). 13C-NMR (DMSO-d6), δ: 56.81, 134.44, 136.21, 137.70. Mass analysis. ESI-MS polarity positive. Calcd. for C13H13OSP: 248; found: 249 (M+H).
4. Add 2 g (8.05 mmol) methyldiphenylthionophosphinate S.12 to 50 mL of 20% trimethylamine/acetonitrile in a 100-mL flask equipped with a stirring bar. Stopper the flask and stir 2 to 3 days at room temperature, monitoring the reaction by analytical TLC and UV shadowing, with 9:1 (v/v) hexane/ethyl acetate as eluent. A white precipitate appears (Rf = 0) and the starting material (Rf = 0.40) disappears. The reaction can be faster if performed at higher temperature (50°C for 10 hr) in a sealed reactor.
5. Filter precipitate by vacuum filtration and wash with 5 mL ice-cold diethyl ether. Analyze S.13 by TLC using 85:10:5 (v/v/v) CH2Cl2/methanol/triethylamine as eluent (Rf = 0.72). Yield 95% (2.35g, 7.65 mmol). 31P-NMR (DMSO-d6), δ: 52.36 ppm. 1H-NMR (DMSO-d6), δ: 3.06 (s, 12H, NMe4), 7.16-7.20 (m, 6H, Ar), 7.70-7.76 (m, 4H, Ar). 13C-NMR (DMSO-d6), δ: 59.85, 132.34, 133.42, 135.97. Mass analysis. ESI-MS polarity positive. Calcd. for C12H10OSP: 233; found: 235 (M+H).
Prepare 2-[2-(2-iodoethoxy)ethoxy]ethanol 6. Place the following in a 250-mL round-bottom flask equipped with a reflux condenser and magnetic stirring bar:
Synthesis of Modified Oligonucleotides and Conjugates
4.9.15 Current Protocols in Nucleic Acid Chemistry
Supplement 7
4 g (24 mmol) 2-[2-(2-choroethoxy)ethoxy]ethanol S.15 14.96 g (100 mmol) NaI 8.36 g (100 mmol) NaHCO3 140 mL anhydrous acetone. 7. Heat at 60°C until the reaction is complete. Monitor reaction by analytical TLC using 9:1 (v/v) CH2Cl2/methanol as eluent. After one night (5 to 6 hr) the starting chlorinated material (Rf = 42) is transformed into the iodinated compound (Rf = 0.56).
8. Remove solid by filtration and evaporate solvent under reduced pressure using a rotary evaporator with a water aspirator. 9. Precipitate excess NaI by adding 8 mL CH2Cl2. Remove solid by filtration and evaporate solvent under reduced pressure. Repeat for a total of four times with 8 mL CH2Cl2 and two times with 10 mL diethyl ether. Take care to use diethyl ether free of peroxides. Yield 83% (4.94 g, 19.9 mmol). Mass analysis ESI-MS polarity positive. Calcd. for C6H3IO3: 260; found: 261 (M+H). This compound can be prepared in advance and stored for months at –20°C.
Prepare S-diphenylphosphinate derivative 7a 10. Place 1 g (3.25 mmol) diphenylthiophosphinate salt S.13, 1.01 g (3.9 mmol) 2-[2-(2iodoethoxy)ethoxy]ethanol, and 30 mL anhydrous CH3CN in a 50-mL flask equipped with a magnetic stirring bar, reflux condenser, and a calcium chloride drying tube. Allow reaction to proceed 15 hr at 50°C. 11. Allow the mixture to cool, remove solid by filtration, and concentrate the solution to dryness. 12. Purify the residue S.14 (2 g) on a 3 × 50–cm column containing 40 g silica gel (APPENDIX 3E), using 100 mL each of 0% to 5% (v/v) methanol in CH2Cl2 in 1% increments. Elute under slight pressure of nitrogen. The pressure must be adjusted until the solvent elutes from the bottom of the column at ~3 drops/sec. Yield 70% (0.834g, 2.27 mmol). 1H-NMR (CDCl3), δ: 2.95-3.03 (2H, m, CH2S), 3.58-3.63 (6H, m, CH2O), 3.68-3.74 (4H, m, CH2O), 7.45-7.58 (6H, m, Ar), 7.85-7.92 (4H, m, Ar). 31 P-NMR (CDCl3), δ: 45.40 ppm. 13C-NMR (DMSO-d6), δ: 33.77, 65.64, 75.05, 77.78, 134.42, 136.35, 138.02. Mass analysis. ESI-MS polarity positive. Calcd. for C18H23O4SP: 366; found: 367 (M+H).
13. Dry S.14 by three rounds of coevaporation with 10 mL anhydrous CH3CN and leave under vacuum overnight. 14. Add 0.317 g (0.426 mL, 2.45 mmol) diisopropylethylamine to a solution of 0.30 g (0.82 mmol) S.14 in 5 mL CH2Cl2. Cool in an ice-water bath. 15. Add 0.23 mL (1.02 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite dropwise to the cold solution via a syringe under a nitrogen atmosphere. Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
16. Monitor reaction by analytical TLC and by UV shadowing, using 70:30:4 (v/v/v) ethyl acetate/hexane/triethylamine as eluent. Pre-elute the plate once with eluent, and then load 1 to 2 µl reaction mixture and elute.
4.9.16 Supplement 7
Current Protocols in Nucleic Acid Chemistry
After 15 to 20 min the starting material (Rf = 0.15) is completely transformed into compound S.7a (Rf = 0.31). TLC using 95:5 (v/v) ethyl acetate/triethylamine gives Rf = 0.33 for the starting material and Rf = 0.70 for the phosphoramidite derivative.
17. When the reaction is complete (15 to 20 min), dilute the reaction mixture with 45 mL ethyl acetate and wash the organic phase twice with 3 mL of 10% aqueous Na2CO3, and twice with 3 mL ice-cold saturated aqueous NaCl solution, using a separatory funnel. 18. Dry organic phase over 15 g sodium sulfate and evaporate to dryness under reduced pressure. 19. Purify the oily residue S.7a (0.8 g) on a 1.6 × 45–cm column containing 25 g silica gel, using 95:5 (v/v) ethyl acetate/triethylamine as eluent. Elute under slight pressure of nitrogen. Store up to two years (or more) at −20°C under argon. The pressure must be adjusted until the solvent elutes from the bottom of the column at ~2 drops/sec. Yield 75% (0.50 g, 0.61 mmol). 31P-NMR (CDCl3), δ: 39.13 and 144.11 ppm. 1H-NMR (CDCl3), δ: 1.13-1.16 (m, 12H, [CH(CH3)2]2), 2.59-2.62 (m, 2H, CH2CN), 2.93-2.98 (m, 2H, CH2SP), 3.51-3.65 (m, 8H, CH2O + CH2OP), 3.65-3.69 (m, 1H), 3.70-3.83 (m, 2H, CH2OP), 3.66-3.70 (m, 1H, C(H)), 7.43-7.47 (m, 4H, Ar), 7.50-7.52 (m, 2H, Ar), 7.83-7.88 (m, 4H, Ar). 13C-NMR (DMSO-d6), δ: 25.25, 29.79, 33.77, 47.87, 63.64, 67.80, 75.00, 75.92, 134.42, 135.51, 138.00. Mass analysis. ESI-MS polarity positive. Calcd. for C27H40N2O5SP2: 566; found: 568 (M+H).
Add masked thiol function to oligonucleotide 20. Couple the phosphoramidite S.7a to the 5′-terminal hydroxyl of a fully protected oligodeoxyribonucleotide bound to a CPG support according to the method described for direct addition of an acridinyl phosphoramidite (UNIT 4.3, steps 15a to 17b of Basic Protocol 1). Deprotect and remove from support 21. Place 1 µmol of the oligodeoxyribonucleotide bound to the support in a 10-mL vial and add successively: 26 mg (0.118 mmol, 120 eq) 2,2′-dithiodipyridine 17 mg (0.179 mmol, 10 eq/phosphate) phenol 3 mL methanol 3.5 mL concentrated ammonium hydroxide. Allow to stand at room temperature for 60 hr. 22. Remove support by vacuum filtration. Wash support successively with 1 mL concentrated ammonium hydroxide, 1 mL water, 1 mL methanol. Pool the original filtrate and the three washes and concentrate them under reduced pressure. 23. Add 6 mL water to the residue and extract four times with 4 mL ethyl acetate to remove organic contaminants. 24. Analyze, purify and characterize the product S.10 as described (see Basic Protocol 1, steps 12 to 17). Store up to two years (or more) at −20°C in a tightly sealed container. Yield: 35-40 OD260 units. Ion-exchange chromatography on a DEAE column using conditions described in Basic Protocol 1 gives retention times of 52 min for C5H5N-S-S-CH2CH2-(OCH2CH2)2-pd[CTCTCGCACCCATCTCTC], and 50 min 36 sec for d[CTCTCGCACCCATCTCTC]. Figure 4.9.13 shows a chromatogram obtained for crude C5H5N-S-S-CH2CH2(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC].
Synthesis of Modified Oligonucleotides and Conjugates
4.9.17 Current Protocols in Nucleic Acid Chemistry
Supplement 5
Figure 4.9.13 Ion-exchange chromatography on a DEAE column of crude C5H5N-S-S-CH2CH2(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC] (see Basic Protocol 3).
RP-HPLC on a Lichrospher RP 18 column using conditions described in Basic Protocol 1 gives retention times of 15 min 14 sec for C5H5N-S-S-CH2CH2-(OCH2CH2)2-pd[CTCTCGCACCCATCTCTC], and 9 min 40 sec for d[CTCTCGCACCCATCTCTC]. See Figure 4.9.14. Mass analysis. ESI-MS polarity negative. Calcd. for C182H238N56O114P18S2: 5652 Da; found: 5651 Da (M-H). The presence of the pyridyldisulfide group at the 5′ end of the oligomer can be detected by reductive cleavage of the disulfide bridge with dithiothreitol or tris-(2-carboxyethyl)phosphine, as described below. The 2-pyridinethione that is released can be detected spectrophotometrically at 343 nm (Carlsson et al., 1978).
Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
Figure 4.9.14 RP-HPLC on a Lichrospher RP 18 column of crude C5H5N-S-S-CH2CH2(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC] (see Basic Protocol 3).
4.9.18 Supplement 5
Current Protocols in Nucleic Acid Chemistry
Figure 4.9.15 Absorption spectra of a solution of C5H5N-S-S-CH2CH2-(OCH2CH2)2-pd[CTCTCGCACCCATCTCTC] before (broken line) and after (solid line) reduction of the disulfide bridge (see Basic Protocol 3).
Verify presence of pyridyldisulfide group 25. Prepare an oligonucleotide solution in 0.1 M TEAA buffer, pH 7, that gives an absorbance of ∼0.5 or 1 OD260. Record the absorption spectrum of this solution between 220 and 450 nm using the same buffer as a reference. 26. Add 10 eq TCEP (dissolved in 5 µl water) to both the sample and reference. Wait 30 min and record the absorption spectrum again. A new absorption band is observed, corresponding to the 2-pyridinethione released (λmax = 343 nm). Knowing the ε value of the oligonucleotide at 260 nm, the absorbance ratio at 260 and 343 nm allows one to determine the quantity of 2-pyridyldisulfide group present in oligonucleotide S.10. Figure 4.9.15 illustrates results from this procedure applied to the oligonucleotide C5H5N-S-S-CH2CH2-(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC].
ADDITION OF A MASKED THIOL GROUP TO THE 5′ TERMINUS OF OLIGODEOXYRIBONUCLEOTIDES USING AN S-ACETYL PHOSPHORAMIDITE
ALTERNATE PROTOCOL 1
This protocol is a modification of the strategy in Basic Protocol 3 (Fig. 4.9.11, left side) that adds an S-acetyl phosphoramidite S.8a to the 5′ terminus of the support-bound oligodeoxyribonucleotide. The alcohol S.16, which is required for the preparation of S.8a (Fig. 4.9.16), is obtained in one step by reaction of 2-[2-(2-chloroethoxy)ethoxy]ethanol S.15 with potassium thioacetate. This procedure is a shortened version of that described by Kuijpers and Van Boeckel (1993). The synthesis can also be started with 2-[2-(2-iodoethoxy)ethoxy]ethanol, which is prepared as described in Basic Protocol 3. However, S.15 is commercially available and thus more convenient. Additional Materials (also see Basic Protocol 3) Potassium thioacetate 1.6 × 45–cm column containing 20 g silica gel (e.g., Kieselgel 60; Merck)
Synthesis of Modified Oligonucleotides and Conjugates
4.9.19 Current Protocols in Nucleic Acid Chemistry
Supplement 7
OH Cl
L
O
+
CH3C(O)SK
H3C
L
S
OH
16
15
i-Pr2NP(Cl)OCH2CH2CN
i-Pr2N
O H3C
DIPEA
S
L
O
P
OCH2CH2CN
8a
Figure 4.9.16 Preparation of the S-acetyl phosphoramidite derivative S.8a. DIPEA, diisopropylethylamine; L, CH2CH2OCH2CH2OCH2CH2.
Synthesize 2-[2-(2-acetylthioethoxy)ethoxy]ethanol S.16 1. Add 2 g (1.72 mL, 11.86 mmol) 2-[2-(2-chloroethoxy)ethoxy]ethanol S.15 to 2 g (17.5 mmol) potassium thioacetate and 35 mL anhydrous acetone in a 25-mL flask equipped with a reflux condenser and a magnetic stirring bar. 2. Heat the stirred mixture 12 hr at 50°C. 3. Allow the mixture to cool to room temperature. 4. Filter off the white precipitate and concentrate the filtrate under reduced pressure. 5. Purify the residue S.16 (2.4 g) on a 3 × 50–cm column containing 40 g silica gel (APPENDIX 3E) using a gradient of 98:2 to 9:1 (v/v) ethyl acetate/methanol. Elute under slight pressure of nitrogen. The pressure must be adjusted until the solvent elutes from the bottom of the column at ~2 to 3 drops/sec.
6. Perform analytical TLC (APPENDIX 3D) using 8:2 (v/v) ethyl acetate/hexane as eluent. Spray plate with 2.5 mg/mL DBPNC in ethanol and heat until the color appears. A pale yellow spot (Rf = 0.41) appears on the TLC plate after treatment with DBPNC. Yield 80% (1.97g, 9.48 mmol). 1H-NMR (CDCl3): δ: 3.73 (2H, m, CH2OH), 3.70-3.55 (8H, m, 4(CH2O)), 3.11 (2H, t, CH2S). 13C-NMR (DMSO-d6), δ: 34.49, 35.80, 36.72, 66.43, 75.12, 75.88, 78.56, 201.33. Mass analysis. ESI-MS polarity positive. Calcd. for C8H16O4S: 208; found: 209 (M+H).
Prepare S-acetyl phosphoramidite derivative S.8a 7. Dry 0.208 g (1 mmol) S.16 by three rounds of coevaporation with 5 mL anhydrous CH3CN and leave under vacuum overnight. 8. Add 0.387 g (0.522 mL, 3 mmol) diisopropylethylamine to a solution of 0.208 g (1 mmol) S.16 in 5 mL CH2Cl2. Cool in an ice-water bath. 9. Add 0.30 mL (1.3 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite dropwise to the cold solution via syringe under a nitrogen atmosphere.
Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
10. Monitor reaction by analytical TLC using 80:20:4 (v/v/v) hexane/ethyl acetate/triethylamine as eluent. Pre-elute the TLC plate once with the eluent, then load 1 to 2 µl reaction mixture and elute. After 15 min, the starting material (Rf = 0.38) is completely transformed into compound S.8a (Rf = 0.70).
4.9.20 Supplement 7
Current Protocols in Nucleic Acid Chemistry
11. When the reaction is complete, dilute the reaction mixture with 45 mL ethyl acetate and wash the organic phase twice with 3 mL of 10% aqueous Na2CO3 and then twice with ice-cold saturated aqueous NaCl using a separatory funnel. 12. Dry the organic phase over 15 g sodium sulfate and evaporate it to dryness under reduced pressure. 13. Purify the residue S.8a (0.45g) on a 1.6 × 45–cm column containing 20 g silica gel using 80:15:5 (v/v/v) hexane/ethyl acetate/triethylamine as eluent. Elute under slight pressure of nitrogen. Store up to two years (or more) at −20°C under argon. The pressure must be adjusted until the solvent elutes from the bottom of the column at ~2 to 3 drops/sec. Yield 75% (0.306 g, 0.75 mmol). 31P-NMR (CDCl3), δ: 150.63 ppm. 1H-NMR (CDCl3), δ: 1.15-1.19 (m, 12H, [CH(CH3)2]2), 2.33 (s, 3H, CH3), 2.64 (t, 2H, J = 6.56 Hz, CH2CN), 3.08 (t, 2H, J = 6.40 Hz, CH2S) 3.57-3.72 (m, 12H, CH2O), 3.74-3.84 (m, 2 CH). Mass analysis. ESI-MS polarity positive. Calcd. for C17H33O5SN2P: 408; found: 409 (M+H).
Add masked thiol function to oligonucleotide 14. Couple the phosphoramidite S.8a to the oligonucleotide and then deprotect, purify, and characterize the product S.10 as described (see Basic Protocol 3, steps 20 to 26). Yield: 40-50 OD260 units. ALTERNATE PROTOCOL 2
ADDITION OF A MASKED THIOL GROUP TO THE 5′ TERMINUS OF OLIGODEOXYRIBONUCLEOTIDES USING A DISULFIDE PHOSPHORAMIDITE DERIVATIVE The second strategy for adding a masked thiol group to the 5′ end of an oligodeoxyribonucleotide is shown in the right side of Figure 4.9.11. The masked thiol is incorporated using a tritylated alkyl disulfide–containing linker (S.9a), which allows full deblocking of the oligodeoxyribonucleotide but not the thiol function. The 4,4′-dimethoxytrityl (DMTr) group makes it possible to monitor the incorporation yield of the linker, and facilitates the purification of the 5′-derivatized oligodeoxyribonucleotide. The phosphoramidite S.9a is obtained via a four-step procedure (Fig. 4.9.17) starting from of 2-[2-(2-acetylthioethoxy)ethoxy]ethanol S.16, which was prepared in Alternate Protocol 1. First, alkali treatment removes the acetyl group to release a thiol-containing linker (S.17), which dimerizes in contact with oxygen in the air (S.18). The dimerized linker is
O H3C
S
L
NH4OH OH
16
OH HS L
O2
HO L
S
17
S
L
OH
18 DMTrCl
i-Pr2N DMTrO
S S
L
O
L
P
i-Pr2NP(Cl)OCH2CH2CN OCH2CH2CN
9a
DIPEA
DMTrO L
S
S
L
OH
19
Figure 4.9.17 Preparation of the tritylated disulfide phosphoramidite derivative S.9a. DIPEA, diisopropylethylamine; DMTr, 4,4′-dimethoxytrityl; L, CH2CH2OCH2CH2OCH2CH2.
Synthesis of Modified Oligonucleotides and Conjugates
4.9.21 Current Protocols in Nucleic Acid Chemistry
Supplement 5
monotritylated (S.19) and then phosphinylated to give the phosphoramidite S.9a. This procedure is a shortened version of that described by Bonfils and Thuong (1991). Additional Materials (also see Basic Protocols 1, 2, and 3) 2-[2-(2-Acetylthioethoxy)ethoxy]ethanol S.16 (see Alternate Protocol 1) 5% ammonium hydroxide Anhydrous pyridine Dimethoxytrityl chloride (DMTr⋅Cl) 10% perchloric acid MgSO4 80% (v/v) acetic acid Ethanol 1 M triethylammonium acetate (TEAA) buffer, pH 6 (stock solution) Dithiothreitol (DTT) Methanol, HPLC grade 1.6 × 45–cm chromatography column containing 20 g silica gel (e.g., Kieselgel 60; Merck) 10-mL gel-filtration column Lyophilizer Additional reagents and equipment for gel-filtration chromatography Prepare dihydroxylated disulfide S.18 1. Solubilize 2-[2-(2-acetylthioethoxy)ethoxy]ethanol S.16 (1 g, 4.80 mmol) in 2 mL of 5% ammonium hydroxide. Magnetically stir the resulting thiol S.17 and allow to dimerize to the disulfide S.18 upon contact with air. 2. Monitor reaction by analytical TLC (APPENDIX 3D) using 9:1 (v/v) CH2Cl2/methanol as eluent. Spray with 2.5 mg/mL DBPNC in ethanol and heat until the color appears. The starting material (Rf = 0.54) appears as a pale-yellow colored spot on the TLC plate after treatment with DBPNC. The thiol derivative S.17 (Rf = 0.50) appears as a brown-colored spot, and its disulfide S.18 (Rf = 0.34) as an intense yellow spot.
3. When the reaction is complete (usually 24 hr), concentrate the mixture and purify the residue (1 g) by flash chromatography (APPENDIX 3E) on a 3 × 50–cm column containing 40 g silica gel using increasing concentrations of methanol in CH2Cl2 (0% to 6%) as eluent. Elute under slight pressure of nitrogen. Monitor elution by TLC. The pressure must be adjusted until the solvent elutes from the bottom of the column at ~2 to 3 drops/sec. Yield: 40% (0.633 g, 1.92 mmol).
Prepare tritylated disulfide S.19 4. Dry 0.7 g (2.09 mmol) S.18 by three rounds of coevaporation with 10 mL anhydrous pyridine and keep it stirring in 5 mL anhydrous pyridine under a nitrogen atmosphere. 5. Cool solution to 5° to 10°C and add 0.18 g (0.53 mmol) DMTr⋅Cl. 6. Allow solution to warm to room temperature. Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
7. Monitor reaction by analytical TLC using 9:1 (v/v) CH2Cl2/methanol as eluent. Spray the plate with 10% perchloric acid.
4.9.22 Supplement 5
Current Protocols in Nucleic Acid Chemistry
Upon spraying the plate with perchloric acid, two orange-colored spots are obtained, corresponding to the monodimethoxytritylated compound S.19 (Rf = 0.65) and the bis-dimethoxytritylated compound (Rf = 0.95).
8. When the reaction is complete (6 to 8 hr), remove the solvent by evaporation under reduced pressure. 9. Dilute the residue with 10 mL CH2Cl2 and wash twice with 3 mL saturated aqueous NaCl and then twice with 3 mL water using a separatory funnel. 10. Dry the organic layer over MgSO4 and concentrate it to dryness. 11. Purify the residue S.19 (0.85 g) by chromatography on a 3 × 50–cm column containing 40 g silica gel using increasing concentrations of methanol in CH2Cl2 (0% to 4%) as eluent. Elute under slight pressure of nitrogen. Monitor elution by TLC. The pressure must be adjusted until the solvent elutes from the bottom of the column at ~2 to 3 drops/sec. Yield 75% (0.250 g, 0.397 mmol). 1H-NMR (CDCl3), δ: 2.80-2.92 (m, 4H, CH2S), 3.23 (t, J = 5.18 Hz, 2H), 3.58-3.80 (m, 18H, CH2O), 3.82 (s, 6H, OCH3), 6.80-7.47 (m, 13H, Ar). 13 C-NMR (DMSO-d6): δ: 35.14, 40.79, 43.40, 54.07, 60.48, 65.70, 68.35, 74.13, 75.20, 75.51, 77.83, 90.79, 118.61, 132.08, 133.23, 135.15, 141.28, 150.49, 163.51. Mass analysis. ESI-MS polarity positive. Calcd. for C33H44O8S2: 632; found 633 (M+H).
Prepare tritylated disulfide phosphoramidite derivative S.9a 12. Dry 0.30 g (0.47 mmol) S.19 by three rounds of coevaporation with 6 mL anhydrous CH3CN and leave under vacuum overnight. 13. Add 0.183 g (0.248 mL, 1.42 mmol) diisopropylethylamine to a solution of 0.30 g (0.47 mmol) S.19 in 4 mL CH2Cl2. Cool in an ice-water bath. 14. Add 0.137 mL (0.61 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite dropwise to the cold solution via a syringe under a nitrogen atmosphere. 15. Monitor reaction by analytical TLC using 50:50:4 (v/v/v) ethyl acetate/hexane/triethylamine as eluent. Pre-elute the TLC plate once with the eluent, then load 1 µl reaction mixture and elute. Visualize by UV shadowing and trityl detection. After 20 min the starting material (Rf = 0.15) is completely tranformed into the phosphoramidite S.9a (Rf = 0.59).
16. When the reaction is complete, dilute reaction mixture with 30 mL ethyl acetate and wash the organic phase twice with 3 mL of 10% aqueous Na2CO3 and then twice with 3 mL ice-cold saturated aqueous NaCl using a separatory funnel. 17. Dry the organic phase over sodium sulfate and concentrate it to dryness under reduced pressure. 18. Purify the oily residue S.9a (0.4 g) on a 1.6 × 45–cm column containing 20 g silica gel using 50:50:4 (v/v/v) ethyl acetate/hexane/triethylamine as eluent. Elute under slight pressure of nitrogen. Store up to two years (or more) at −20°C under argon. Yield 75% (0.293 g, 0.352 mmol). 31P-NMR (CDCl3), δ: 150.48 ppm. 1H-NMR (CDCl3), δ: 1.16-1.24 (m, 12H, [CH(CH3)2]2), 2.80 (t, 2H, J = 6 Hz, CH2CN), 3.20-3.24 (m, 4H, CH2S), 3.58-3.83 (m, 20H, CH2O, CH2OP +2 CH), 3.75 (s, 6H, OCH3), 6.80-7.47 (m, 13H, Ar). Mass analysis. ESI-MS polarity positive. Calcd. for C42H62O9S2N2P: 831; found: 832 (M+H).
Couple masked thiol function to oligonucleotide 19. Couple tritylated disulfide phosphoramidite S.9a to the 5′-terminal hydroxyl of a fully protected oligodeoxyribonucleotide bound to a CPG support according to the
Synthesis of Modified Oligonucleotides and Conjugates
4.9.23 Current Protocols in Nucleic Acid Chemistry
Supplement 7
method described for direct addition of an acridinyl phosphoramidite (UNIT 4.3, steps 15a to 17b of Basic Protocol 1). Detritylating a small amount of the support-bound oligonucleotide allows one to verify the coupling efficiency and, if necessary, to perform a second coupling to improve the yield.
20. Deprotect the oligodeoxyribonucleotide S.9b (Fig. 4.9.11) as described for the deprotection of the oligodeoxyribonucleotides with 5′-bis-(2-cyanoethyl)phosphate groups (see Basic Protocol 2, steps 8 to 11, of this unit) but without DTT in step 8. 21. Analyze, purify, and characterize the tritylated disulfide-containing oligodeoxyribonucleotide S.9c as described (see Basic Protocol 1, steps 11 to 17). Yield: 40-50 OD260 units. Ion-exchange chromatography on DEAE using conditions described in Basic Protocol 1 gives retention times of 52 min 12 sec for DMTrOCH2CH2-(OCH2CH2)2-S-S-CH2CH2(OCH2CH2)2- p- d[C TCTC GCAC CCATCCTC] and 50 min 36 sec for d[CTCTCGCACCCATCTCTC]. Figure 4.9.18 shows a chromatogram obtained for crude DMTrOCH2CH2-(OCH2CH2)2-S-S-CH2 CH2-(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC]. RP-HPLC on a Lichrospher RP 18 column using a linear gradient of CH3CN (5% to 50% over 60 min) in 0.1 M TEAA buffer, pH 7, at 1 mL/min gives retention times of 32 m i n 2 6 sec for DMTrOCH2CH2-(OCH2CH2)2-S-S-CH2CH2-(OCH2CH2)2-pd[CTCTCGCACCCATCTCTC] and 12 min for d[CTCTCGCACCCATCTCTC]. Figure 4.9.19 shows a chromatogram obtained for crude DMTrOCH2CH2-(OCH2CH2)2-S-SCH2CH2-(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC]. An additional peak with a retention time of 13 min 40 sec has been identified as the detritylated oligonucleotide HOCH2CH2-(OCH2CH2)2-S-S-CH2CH2-(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC]. Mass analysis. ESI-MS polarity negative. HOCH2CH2-(OCH2CH2)2-S-S-CH2CH2(OCH2CH2)2- p-d[CTCTCGCACCCATCTCTC]. Calcd. for C182H247N55O117S2P18: 5699; found: 5697 ± 4 (M-H). Sometimes a significant amount of detritylation occurs when handling the functionalized oligonucleotide. In this case, it is best to detritylate prior to purification by adding 4 mL of 80% acetic acid to the tritylated oligonucleotide. After 20 to 30 min incubation at room temperature, the acetic acid is removed by evaporation under reduced pressure. The residue
Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
Figure 4.9.18 Ion-exchange chromatography on a DEAE column of crude DMTrO-CH2CH2(OCH2CH2)2-S-S-CH2CH2-(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC] (see Alternate Protocol 2).
4.9.24 Supplement 7
Current Protocols in Nucleic Acid Chemistry
Figure 4.9.19 RP-HPLC on a Lichrospher RP 18 column of crude DMTrO-CH2CH2-(OCH2CH2)2S-S-CH2CH2-(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC] (see Alternate Protocol 2).
is then coevaporated three times with 4 mL ethanol, dissolved in 4 mL water, and extracted three times with 4 mL ethyl acetate. The purity of the detritylated oligonucleotide is verified by RP-HPLC using a linear gradient of CH3CN (0% to 30% over 60 min) in 0.1 M TEAA, pH 7, at 1 mL/min. If required, further purification is performed by RP-HPLC. When the modified oligodeoxyribonucleotide is purified at the tritylated step, detritylation should be performed before storage to avoid loss of trityl, resulting in a mixture of products. When transformation into oligomer S.10 is performed immediately after purification, the tritylated compound can be used. The chromatography data given above is for the tritylated compound; mass data is for detritylated product.
Transform disulfide-containing oligodeoxyribonucleotide into oligomer S.10 22. Prepare a solution of 10 OD260 units S.9c in 1 mL of 0.05 M TEAA buffer, pH 6. 23. Add 30 eq DTT or 3 eq TCEP and monitor cleavage of the disulfide bridge by RP-HPLC on an RP 18 column using the conditions described above (see Basic Protocol 1, step 14b). The thiol-containing oligodeoxyribonucleotide HS-CH2CH2-(OCH2CH2)2-pd[CTCTCGCACCCATCTCTC] is eluted first (11 min 30 sec) while the oligonucleotide containing the disulfide linker, HOCH2CH2-(OCH2CH2)2-S-S-CH2CH2-(OCH2CH2)2-pd[CTCTCGCACCCATCTCTC], is eluted ~2 min later (13 min 25 sec).
24. When the reaction is complete (overnight), extract excess DTT five times using 1 mL ethyl acetate. It is not necessary to remove the excess TCEP.
25. Load aqueous solution on a 10-mL gel-filtration column and elute with 0.05 M TEAA buffer, pH 6. Collect oligodeoxyribonucleotide in a solution containing 20 eq of 2,2′-dithiodipyridine in 2 mL HPLC-grade methanol. Monitor the elution of the oligonucleotide spectrophotometrically at 260 nm. The choice of Sephadex G-10 or G-25 depends on the molecular weight of the oligodeoxyribonucleotide. The oligonucleotide is usually eluted in the void volume (∼3 to 4 mL).
26. Allow eluent/dithiodipyridine/methanol mixture to incubate for 5 to 6 hr. 27. Remove methanol by evaporation, and extract excess 2,2′-dithiodipyridine four times using 2 mL ethyl acetate.
Synthesis of Modified Oligonucleotides and Conjugates
4.9.25 Current Protocols in Nucleic Acid Chemistry
Supplement 5
28. Verify formation of 2-pyridyldisulfide by RP-HPLC on RP 18 column using the conditions described (see Basic Protocol 1, step 14b). The 2-pyridyldithioalkylated oligodeoxyribonucleotide S.10 has a longer retention time (15 min 14 sec) than that of the mercaptoalkylated oligonucleotide (11 min 30 sec).
29. Purify the 5′-terminal 2-pyridyldithioalkylated oligodeoxyribonucleotide S.10 by RP-HPLC using the same column and conditions used for analysis. Purification can be achieved in three or four runs.
30. Remove organic solvent by evaporation and remove buffer by three consecutive lyophilizations. Store up to two years at −20°C in a tightly sealed container. Yield 60%. Mass analysis. ESI-MS polarity negative. Calcd. for C182H238N56O114P18S2: 5652 Da; found: 5651 Da (M-H).
COMMENTARY Background Information
Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
This unit reports methods for the preparation of modified oligodeoxyribonucleotides containing 5′-terminal carboxyl, amino, phosphorothioate, phosphate, or masked thiol groups. These 5′-modified oligonucleotides may be used to prepare oligodeoxyribonucleotides covalently linked to a wide variety of molecules such as labels, intercalating agents, and reactive compounds, a number of which are reported in UNIT 4.10. The methods developed in this unit can be used to prepare 5′-modified oligodeoxyribonucleotides built with natural β-deoxyribonucleosides, phosphorothioate phosphodiesters (Aubert et al., 2000), unnatural α-D-deoxyribonucleosides (Kurfürst et al., 1993), or 2′-O-methyldeoxyribonucleosides. Given that oligodeoxyribonucleotides can be purchased fully protected and bound to a support, 5′-end modification can be performed manually. When oligonucleotide synthesis is performed in the 5′→ 3′ orientation as in the case of N3′→P5′ phosphoramidates, 5′-derivatization can be achieved using a support bearing either the ligand or the masked functional group. These methods can also be adapted to oligonucleotides containing modified nucleosides, but it must be verified that the conditions required for deprotection are compatible with the stability of the modified nucleosides. For oligoribonucleotides and modified oligonucleotides that are sensitive to basic conditions, it is recommended to introduce 5′-phosphate and 5′-thiophosphate groups enzymatically (Eckstein, 1983). Alternate methods reported by Grimm et al. (2000) use the properties of a terminal phosphomonoester group for the introduction of linkers ending with amino, carboxyl, or sulfhydryl functions into unprotected oligonu-
cleotides. These methods are mainly limited to the introduction of either one functional group at one end or two identical functional groups at both ends of the oligonucleotide. In contrast, the methods reported in this unit, including the incorporation of masked functional groups prior to deprotection, allow the incorporation of a wide combination of functional groups into oligonucleotides for specific subsequent coupling of various ligands. Masked amine, sulfhydryl, and carboxyl can be introduced via a linker to various sites of the oligonucleotide chain: base, sugar, internucleotidic phosphate, and 3′ terminus (UNIT 4.5), as well as phosphate and thiophosphate at the 3′ terminus or thiophosphate at one or many internucleotidic positions during the assembly steps; these groups are released during deprotection. Amine and masked thiol or thiophosphate are compatible as well as masked thiol and amine in the phosphorothioate series (Aubert et al., 2000). The procedure for incorporation of a thiophosphate at the 5′ terminus can be modified to introduce a masked thiol group at the 3′ terminus by using a modified support (Bonfils and Thuong, 1991). Using the S-acetyl-containing linker S.16, synthesis of 25-mer oligodeoxyribonucleotide in the phosphorothioate series has been performed at a 10-µmol scale with very good yields (70 OD260 units per µmol).
Critical Parameters and Troubleshooting The success of these procedures is dependent on a number of critical parameters. The methods reported include either carbonyldiimidazole activation or the use of phosphoramidite chemistry, both of which are highly water sensitive. For these reasons, strictly anhydrous
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Current Protocols in Nucleic Acid Chemistry
solvents and solutions should be used as well as dry starting material. Precautions should be taken to exclude moisture, as well as oxygen in the case of the phosphoramidite derivatives. Phosphoramidites such as S.7a, S.8a, S.9a, and bis-(2-cyanoethyl)diisopropylamidophosphite, as well as most of the intermediate compounds used for their preparation, should be stored in tightly closed flasks or vials at –20°C under argon atmosphere. Cold flasks and vials should always be allowed to reach room temperature before they are opened, and vials should remain open as little as possible. To avoid low yields of oligonucleotides containing 5′-terminal functional groups, extreme care should be taken when using a syringe to remove solutions from vials containing the support-bound oligonucleotide in order to avoid unnecessary loss. After modification of the 5′ termini, it is recommended to deprotect an aliquot of the oligonucleotide conjugates to verify the efficiency of the reaction. In the case of the addition of the tritylated disulfide phosphoramidite S.9a, the coupling yield can be evaluated by quantitative trityl analysis performed as reported in UNIT 3.2. If the yield of 5′-modification is not sufficient, it is possible to perform an additional coupling step with the appropriate phosphoramidite derivatives. After 26 months storage of the tritylated disulfide phosphoramidite S.9a under the conditions described above, 5′-modification of an oligonucleotide has been performed on a µmol scale (by using 20 eq of S.9a) with 90% to 95% yield as determined by trityl quantification. This result indicates that, although the compound contains both a disulfide linkage and a tricoordinated phosphorus, it is unlikely to self-destruct to a large extent. This behavior differs from that of TCEP used to reduce the disulfide bond, probably for the following reasons: (1) the reducing properties of the phosphoramidite derivatives are weaker than those of the TCEP derivative; (2) the reductive cleavage of the disulfide linkage by TCEP requires the presence of one water molecule (Burns et al., 1991) and it is unlikely that 1 eq of water can be present in the stock of phosphoramidite S.9a; (3) the oligoethylene glycol linker can adopt a crown ether–like structure that prevents close proximity of the disulfide linkage and the tricoordinated phosphorus. After deprotection and before loading chromatography columns, organic impurities are extracted from crude oligodeoxyribonucleotide solutions with organic solvents. Oligodeoxyribonucleotide solutions must be fil-
tered through a 0.45-µm disposable filter to remove any particulates and prevent the clogging of analytical chromatography columns. In order to prevent chelation of phosphorothioate oligodeoxyribonucleotides, all solvents and buffer solutions used for their purification must be passed through a column of Chelex 100 resin (Bio-Rad) to remove divalent cations. The crude oligodeoxyribonucleotides obtained from these protocols are purified by ion-exchange chromatography on Mono Q (Amersham Pharmacia Biotech) and DEAE (Waters) columns. After a desalting step, the purity of these oligonucleotides is verified on a Lichrospher 100 RP column. When purified by ion-exchange chromatography, oligodeoxyribonucleotides containing 5′-functional groups are, in most cases, sufficiently pure for subsequent coupling reactions with selected ligands. These purification procedures are suitable for 10- to 25-mers. They are not suitable for oligonucleotides that can form self-associated complexes such as G-tetrads. Purification of tritylated compounds such as the 5′-tritylated disulfide-containing oligonucleotide S.9c or the 2-pyridyldithioalkylated oligonucleotide S.10 can easily be carried out not only with a reversed-phase column but also with an ion-exchange column such as DEAE (Waters). In the latter case, the presence of a lipophilic group allows good separation between the full-length 5′-modified and 5′-unmodified oligodeoxyribonucleotides. For 5′sulfur-containing oligodeoxyribonucleotides, purification by RP-HPLC on a Lichrospher RP 18 column using a low gradient of CH3CN in TEAA buffer, pH 7, allows separation of 5′mercaptoalkylated and 5′-(2-pyridyldithioalkylated) oligodeoxyribonucleotides. For 5′carboxyalkylated or 5′-aminoalkylated oligodeoxyribonucleotides, the presence of the 5′-functional group can be verified by RPHPLC by comparison with unmodified oligodeoxyribonucleotides, which are eluted more quickly than their 5′-derivatized counterparts. The presence of the amino function on 5′-aminoalkylated oligodeoxyribonucleotides can be verified by TLC using ninhydrin. For oligodeoxyribonucleotide 5′-phosphates, the use of a Delta-Pak C4 or RP 18 HPLC column allows separation of unmodified oligodeoxyribonucleotides from 5′-phosphate-containing oligomers. Purification can be difficult when working with oligonucleotides containing particular sequences, e.g., stretches of G. The best solution may be to replace the difficult se-
Synthesis of Modified Oligonucleotides and Conjugates
4.9.27 Current Protocols in Nucleic Acid Chemistry
Supplement 5
quences, e.g., by replacing dG with 7-deaza-2′dG (Raynaud et al., 1996). Nuclease digestion of the conjugates and RP-HPLC analysis of the resulting monomers ascertain full removal of the protecting groups and base composition. Additional characterization can be made by matrix-assisted laser desorption/ionization MS (MALDI-MS) and electrospray ionization MS (ESI-MS). After purification and characterization, oligonucleotides should be lyophilized and stored in tightly closed vials at −20°C. Under these conditions, they are stable up to two years (or more).
Anticipated Results Protocols reported in this unit are given for syntheses at a 1-µmol scale. When purification requires only ion-exchange chromatography or RP-HPLC, yields of 25 to 45 OD260 units can be expected for oligonucleotides of medium size (12- to 16-mers). The 5′-modifications of oligonucleotides reported in this unit have also been performed at a 10-µmol scale with similar yields.
Time Considerations Provided that all reagents and materials required for each step are available, most of the procedures are simple and rapid. When all the phosphoramidite derivatives are ready to use, the time required to prepare one oligonucleotide modified at its 5′ terminus with a carboxylated or aminoalkylated linker, a phosphate, a thiophosphate, or a masked thiol group (including purification and characterization) is ∼1 week. The time required for the preparation of the S-acetyl phosphoramidite derivative S.8a and the tritylated disulfide phosphoramidite derivative S.9a is ∼3 days. The preparation of the S-diphenylphosphinate phosphoramidite derivative S.7a requires 1 week or more.
Literature Cited Aubert, Y., Bourgerie, S., Meunier, L., Mayer, R., Roche, A.-C., Monsigny, M., Thuong, N.T., and Asseline, U. 2000. Optimized synthesis of phosphorothioate oligodeoxyribonucleotides substituted with a 5′-protected thiol function and a 3′-amino group. Nucleic Acids Res. 28:818-825.
Modification of the 5′ Terminus of Oligodeoxyribonucleotides for Conjugation with Ligands
Bonfils, E. and Thuong, N.T. 1991. Solid-phase synthesis of 5′-3′-bifunctional oligodeoxyribonucleotides bearing a masked thiol group at their 3′-end. Tetrahedron Lett. 35:3053-3056. Burns, J.A., Butler, J.C., Moran, J., and Whitesides, G.M. 1991. Selective reduction of disulfides by tris(2-carboxyethyl)phosphine. J. Org. Chem. 56:2648-2650. Carlsson, J., Drevin, H., and Axen, R. 1978. Proteinthiolation and reversible protein-protein conjugation. Biochem. J. 173:723-737. Eckstein, F. 1983. Phosphorothioate analogues of nucleotides. Tools for investigation of biochemical processes. Angew. Chem. Int. Ed. Engl. 22:423-506. Gottikh, M., Asseline, U., and Thuong, N.T. 1990. Synthesis of oligonucleotides containing a carboxyl group at either their 5′-end or their 3′-end and their subsequent derivatization by an intercalating agent. Tetrahedron Lett. 31:6657-6660. Grimm, G.N., Boutorine, A.S., and Hélène, C. 2000. Rapid routes of synthesis of oligonucleotide conjugates from non-protected oligonucleotides and ligands possessing different nucleophilic or electrophilic functional groups. Nucleosides Nucleotides and Nucleic Acids 19:1943-1965. Kuijpers, W.H. and Van Boeckel, C.A. 1993. A new strategy for the solid-phase synthesis of 5′-thiolated oligodeoxyribonucleotides. Tetrahedron 49:10944-10944. Kurfürst, R., Roig, V., Chassignol, M., Asseline, U., and Thuong, N.T. 1993. Oligo-α-deoxyribonucleotides with a modified nucleic acid base and covalently linked to reactive agent. Tetrahedron 32:6975-6990. Raynaud, F., Asseline, U., Roig, V., and Thuong, N.T. 1996. Synthesis and characterization of O6modified deoxyguanosine-containing oligodeoxyribonucleotides for triple-helix formation. Tetrahedron 52:2047-2064. Thuong, N.T. and Asseline, U. 1991. Oligodeoxyribonucleotides attached to intercalators, photoreactive and cleavage agents. In Oligodeoxyribonucleotides and Analogues: A practical approach (F. Eckstein, ed.) pp. 283-308. IRL Press, Oxford. Wachter, L., Jablonski, J.A., and Ramachandran, K.L. 1986. A simple and efficient procedure for the synthesis of 5′-aminoalkyl oligonucleotides. Nucleic Acids Res. 14:7985-7994.
Contributed by Ulysse Asseline and Nguyen T. Thuong Centre de Biophysique Moléculaire, CNRS Orléans, France
We would like to express our appreciation to our past and present collaborators for their contribution to the development of varied oligonucleotide families over the past years. The work was supported by Rhône-Poulenc, Agence Nationale de Recherche contre le SIDA and Bio-Mérieux.
4.9.28 Supplement 5
Current Protocols in Nucleic Acid Chemistry
Conjugation of 5′-Functionalized Oligodeoxyribonucleotides with Properly Functionalized Ligands
UNIT 4.10
This unit describes a collection of methods for the chemical attachment of reporter groups and various conjugate groups (e.g., intercalators, photoreactive agents, cleaving agents) to the 5′ terminus of oligodeoxyribonucleotides. Two strategies can be used. The first (described in UNIT 4.3) involves the direct addition of the ligand to the 5′ end of the oligodeoxyribonucleotide via its phosphoramidite or H-phosphonate derivative. This approach works well when there is a sufficient amount of the ligand and when its solubility and chemical stability are compatible with the conditions required for the preparation of the phosphoramidite or H-phosphonate derivative. In addition, the ligand should be stable under the conditions used for the coupling step and for the complete deprotection of the oligodeoxyribonucleotide, including its release from the support. The second strategy involves the incorporation of appropriate functional groups into both the ligand and the 5′ terminus of the oligodeoxyribonucleotide. Specific coupling of these reactants results in the formation of oligodeoxyribonucleotide conjugates. Using this strategy, several oligodeoxyribonucleotide conjugates can be prepared from only one oligodeoxyribonucleotide synthesis provided that the required amount for each conjugate is low. In this approach, appropriate functional groups on the ligand and oligonucleotide must be matched so that a conjugation reaction can be performed. This unit delineates methods used for the following coupling reactions: 5′-carboxylated oligonucleotides with aminoalkylated ligands (see Basic Protocol 1); 5′-aminoalkylated oligonucleotides with ligands that have been functionalized with an isothiocyanate or N-hydroxysuccinimidyl group (see Basic Protocol 2); oligonucleotide 5′-phosphorothioates with ligands functionalized with halogenoalkyl, 2-pyridyldithio, or iodoacetamidyl groups (see Basic Protocol 3); oligonucleotide 5′-phosphates with ligands functionalized with amino groups (see Basic Protocol 4); and 5′-mercaptoalkylated oligonucleotides with ligands functionalized with halogenoalkyl, 2-pyridiyldithio, or iodoacetamidyl groups (see Basic Protocol 5). The initial incorporation of these functional groups into the ligand and oligonucleotide has been described in UNITS 4.8 & 4.9, respectively. The ligands described in this unit are the intercalator 2-methoxy-6-chloro-9-aminoacridine, the photoreactive group psoralen, the cleaving reagent phenanthroline-Cu, and the label thiazole orange (UNIT 4.8). Alternatively, many labels carrying functional groups that react with 5′-thiol, 5′-terminal phosphorothioate, and 5′-amino groups are commercially available. Heterobifunctional reagents, which allow reactions with a thiol function and a primary amine, are also available from commercial sources. The latter compounds, listed in UNIT 4.2, are very useful when conjugates between the oligodeoxyribonucleotide and the ligand do not require a well-defined linker. The procedures reported in this unit are for 10- to 25-mer oligodeoxyribonucleotides derivatized at their 5′ termini by various ligands, and require that the sequences under consideration will not form self-associated complexes such as G-tetrads. Such oligonucleotide structures require a specific procedure that is not discussed in this unit. Purification can be improved by performing ion-exchange chromatography at pH 12 when the obligonucleotide-ligand conjugate is stable at the pH. Another solution to prevent the formation of G-tetrads is to partially replace 2′-deoxyguanosine with 7-deaza-2′-deoxyguanosine (Raymond et al., 1996) or with 7-chloro-7-deaza-2′-deoxyguanosine (Aubert et al., 2001). Contributed by Ulysse Asseline and Nguyen T. Thuong Current Protocols in Nucleic Acid Chemistry (2001) 4.10.1-4.10.23 Copyright © 2001 by John Wiley & Sons, Inc.
Synthesis of Modified Oligonucleotides and Conjugates
4.10.1 Supplement 7
CAUTION: All chemicals must be handled in a fume hood by personnel equipped with a laboratory coat, glasses, and gloves. NOTE: When purifying and handling phosphorothioate- or phenanthroline-containing oligodeoxyribonucleotides, it is necessary to mix solutions, buffers, and solvents with Chelex 100 resin in order to remove divalent cations. BASIC PROTOCOL 1
CONJUGATION OF 5′-CARBOXYLATED OLIGODEOXYRIBONUCLEOTIDES TO AMINOALKYLATED LIGANDS The conjugation of oligodeoxyribonucleotides containing a carboxyl group at the 5′ terminus with the primary amino function of a ligand (or with an amine-like function such as a hydrazine derivative) is accomplished in the presence of carbodiimide. It generally requires a large excess of both ligand and carbodiimide (100 to 500 eq). Usually the oligodeoxyribonucleotide is solubilized in buffer and the ligand is added to the solution in amine-free dimethylformamide (DMF) dimethylsulfoxide (DMSO). The procedure is illustrated in Figure 4.10.1, using the hydrazine derivative of Lucifer Yellow (LY) as an example. The coupling reaction is performed at pH 5.5 to 6 in order to prevent the modification of the nucleobase. At this pH, the hydrazine derivative remains reactive. When the coupling reaction is complete, as determined by thin-layer chromatography (TLC; APPENDIX 3D), the conjugate is isolated from the reaction mixture. Different strategies can be used depending on the solubility of the ligand in the solvent or mixture of solvents used to perform the conjugation reactions. A general procedure for analysis, purification, and characterization of conjugates is given for 10- to 25-mer non-self-associating oligodeoxyribonucleotides derivatized at the 5′ end by a variety of ligands.
−O C(CH ) NHC(O)O 2 2 m
B
O
RNHCO(CH2)mNHC(O)O
B
O
m = 4, 5 O
O RNH2
O P O− O
O
O P O−
EDC
B
O
B
O
O H
O
n
H
n
−O S 3 O
O
N
N NH H
R = LY = H2N −O 3S
Conjugation of Functionalized Oligodeoxyribonucleotides and Ligands
O
Figure 4.10.1 Conjugation of 5′-carboxylated oligodeoxyribonucleotides with aminated ligands. Abbreviations: B, base (i.e., thymine, cytosine, adenine, or guanine); EDC, 1-[3-(dimethylamino)propyl]-3-ethylcarbodiimide hydrochloride; LY, Lucifer Yellow (R).
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Current Protocols in Nucleic Acid Chemistry
Materials Purified 5′-carboxylated oligodeoxyribonucleotide (UNIT 4.9) 0.5 M pyridine buffer, pH 5.5 to 6 (use pyridine redistilled from p-toluenesulfonyl chloride and stored over 3A molecular sieves; adjust pH with HCl) 10 to 20 mg/mL aminoalkylated ligand (UNIT 4.8; e.g., Lucifer Yellow hydrazine derivative) dissolved in water, dimethylsulfoxide (DMSO), or N,N-dimethylformamide (DMF; redistilled in vacuo over ninhydrin) 1-[3-(Dimethylamino)propyl]-3-ethylcarbodiimide hydrochloride (EDC) 80:20 or 50:50 (v/v) dichloromethane (CH2Cl2)/methanol 55:10:20 or 55:10:30 (v/v/v) isopropanol/NH4OH/H2O Sephadex G-10, G-15, or G-25 resin (Pharmacia; for H2O-soluble ligands) 0.5 M triethylammonium acetate (TEAA), pH 7 Dichloromethane or ethyl acetate (for very lipophilic ligands) 4% (w/v) LiClO4 in distilled acetone, or n-butanol (for ligands of medium lipophilicity) Mobile phase A: 5% (v/v) acetonitrile in 0.1 M aqueous TEAA, pH 7, all HPLC grade Mobile phase B: 80% (v/v) acetonitrile in 0.1 M aqueous TEAA, pH 7, all HPLC grade 2-mL vial equipped with Teflon-faced septum Analytical silica gel TLC plates (e.g., 5554 Kieselgel 60F plates; Merck) 5-mL column UV lamp and viewing box Rotary evaporator with water bath and vacuum pump High-performance liquid chromatography (HPLC) system, including: Chemically inert syringes with replaceable needles Reversed-phase column: 125-mm × 4-mm, 5-µm Lichrospher 100 RP18 (Merck), CC Nucleosil 100-5 C18 (125/4; Macherey-Nagel), or polystyrene reversed-phase (PRP-1; Hamilton) Multiwavelength detector capable of measuring UV-Vis absorption between 230 and 600 nm Spectrophotometer Lyophilizer Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and purification and characterization of oligonucleotide-acridine conjugates (UNIT 4.3) NOTE: All buffered solutions used for HPLC purifications should be filtered through a 0.22- or 0.45-µm disposable filter. Couple ligand to oligonucleotide 1. In a 2-mL vial equipped with a Teflon-faced septum and magnetic stir bar, dissolve 5 OD260 units of purified 5′-carboxylated oligodeoxyribonucleotide in 0.1 mL of 0.5 M pyridine/HCl buffer, pH 5.5 to 6. 2. Add 0.1 mL of 10 to 20 mg/mL aminoalkylated ligand and 10 mg EDC. Stir 3 to 4 hr at room temperature. Water-soluble ligands should be prepared in water. Non-water-soluble ligands should be prepared in amine-free DMF or DMSO. EDC is a water soluble form of carbodiimide and is used to form amide bonds. Synthesis of Modified Oligonucleotides and Conjugates
4.10.3 Current Protocols in Nucleic Acid Chemistry
Supplement 7
Evaluate coupling efficiency 3. Monitor the coupling reaction by thin-layer chromatography (TLC; APPENDIX 3D) on analytical silica gel plates, eluting first with either 80:20 or 50:50 (v/v) dichloromethane/methanol, and then with 55:10:20 or 55:10:30 (v/v/v) isopropanol/NH4OH/H2O. The Lucifer yellow conjugate is directly visible as a yellow spot. In most cases, the main excess chromophore travels to the top of the TLC plate while the conjugated oligodeoxyribonucleotide moves slightly faster (i.e., higher Rf) than the starting oligomer. When monitoring the coupling of Lucifer Yellow with the oligonucleotide HOOC(CH2)5NH-CO-d[CTCTCGCACCCATCTCTC], elute the TLC plate directly with 55:10:20 (v/v/v) isopropanol/NH4OH/H2O (not with dichloromethane/methanol). The Lucifer Yellow excess has an Rf of 0.62, while the starting oligodeoxyribonucleotide and the conjugated product have Rf values of 0.31 and 0.34, respectively. The coupling reaction can also be monitored by reversed-phase HPLC using the procedures described below. In most cases, the oligodeoxyribonucleotide-ligand conjugate exhibits a longer retention time than that of the unconjugated oligomer.
Recover oligodeoxyribonucleotide-ligand conjugate 4a. For water-soluble ligand (e.g., Lucifer Yellow): Remove excess ligand by gel filtration using a 5-mL column of Sephadex G-10, G-15, or G-25 resin, depending on the size of the oligodeoxyribonucleotide. Collect 0.5-mL fractions and monitor for presence of conjugate by spotting 3 µL on TLC plates. Visualize by UV shadowing at 254 nm. 4b. For very lipophilic ligand: Concentrate the reaction mixture to dryness under reduced pressure using a rotary evaporator with a water bath and a vacuum pump. Dissolve the residue in water or 0.5 M TEAA, pH 7. Remove excess ligand by extraction with organic solvents such as dichloromethane or ethyl acetate. 4c. For medium-lipophilicity ligand: Precipitate conjugate by adding 10 vol of 4% LiClO4 in distilled acetone or n-butanol. Vortex 1 min and centrifuge 5 min at 2000 × g, room temperature. Decant supernatant and resuspend pellet in 200 µL water. Perform gel filtration as in step 4a. Precipitation can also be performed using n-butanol prechilled at –30°C.
Purify and characterize conjugate 5. Preequilibrate a 125-mm × 4-mm, 5-µm Lichrospher 100 RP18 RP-HPLC column for at least 15 min with mobile phase A. 6. Before analysis and purification, run a blank linear gradient from 100% mobile phase A to 100% mobile phase B at a flow rate of 1 mL/min over 80 min. Repeat this step between analyses and purifications of samples having different DNA sequences or modifications.
7. Analyze sample by RP-HPLC using the flow rate and gradient in step 6. Detect the presence of oligodeoxyribonucleotide-ligand conjugates by measuring UV absorption at 254 or 260 nm.
Conjugation of Functionalized Oligodeoxyribonucleotides and Ligands
This method allows easy separation of underivatized or 5′-functionalized oligodeoxyribonucleotides from oligodeoxyribonucleotide-ligand conjugates, which are, in most instances, more lipophilic. In most cases, RP-HPLC using a Lichrospher 100 RP18 column or a CC Nucleosil 100-5 C18 column is efficient for purifying the conjugate, and these columns give nearly equivalent results. A PRP-1 column can be used when these columns are not efficient, as in the case of oligonucleotide-orthophenanthroline conjugates. Detection is usually performed where the oligonucleotide absorbs light (254 or 260 nm) and at a second wavelength where the ligand absorbs light (usually in the visual range). It is also possible to record the UV-Vis spectrum corresponding to each peak of the chromatogram between 230 and 600 nm.
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Current Protocols in Nucleic Acid Chemistry
Purification of conjugates can also be performed by ion-exchange chromatography (e.g., for oligonucleotide-orthophenanthroline conjugates; see Basic Protocol 2). Ion-exchange chromatography that requires a desalting step should only be used when RP-HPLC is not successful. Conjugates can also be purified by preparative polyacrylamide gel electrophoresis (PAGE; APPENDIX 3B). The conjugates have migrate more slowly through the gel than the starting oligonucleotides. For all compounds described in this unit, except for oligodeoxyribonucleotide-phenanthroline conjugates, UV detection can be achieved at any wavelength at which the ligand absorbs UV light. The steepness of the gradient used must be chosen empirically to afford the best chromatographic separation between the oligonucleotide conjugate and the 5′-functionalized oligonucleotide precursor. Usually a rate of increase of 1% acetonitrile per minute is used for RP-HPLC.
8. After determining the system that gives the best separation, purify the remaining crude 5′-modified oligonucleotide by preparative reversed-phase or ion-exchange chromatography. Alternatively, purify directly on an analytical column when the coupling reaction is achieved on a small scale (i.e., ∼5 OD units).
9. Purify and characterize as reported in UNIT 4.3 (steps 3 to 8 of Support Protocol), but analyze the purified conjugate by RP-HPLC. After purification, UV absorption spectra of purified oligodeoxyribonucleotide-ligand conjugates show absorptions that are characteristic to the ligand (acridine, psoralen, pyrene, fluorescein, thiazole orange).
10. Determine the extinction coefficient (ε) of the oligodeoxyribonucleotide-ligand conjugate. It is generally accepted that the ε value of the oligodeoxyribonucleotide-ligand conjugate at a given wavelength is the sum of the ε values of the oligodeoxyribonucleotide and the ligand at the same wavelength.
11. Measure the OD260 to determine the yield of the oligodeoxyribonucleotide-ligand conjugate.
Figure 4.10.2 UV-visible spectrum of the Lucifer Yellow–oligodeoxyribonucleotide conjugate LY-NH-CO(CH2)5-NH-CO-d[CTCTCGCACCCATCTCTC] recorded in water between 230 and 520 nm.
Synthesis of Modified Oligonucleotides and Conjugates
4.10.5 Current Protocols in Nucleic Acid Chemistry
Supplement 7
12. Lyophilize the conjugate and store at −20°C (stable for one to many years). Nuclease digestion of the conjugates and RP-HPLC analysis of the resulting monomers can also be performed to verify that base modifications did not occur during coupling. Moreover, these methods confirm the structure of oligodeoxyribonucleotide-ligand conjugates. Additional characterization can be made by matrix-assisted laser desorption/ionization (MALDI) and electrospray ionization (ESI) mass spectrometry. Fo llow ing co nju ga tion of Lu cifer Yellow with HOOC(CH2)5-NH-COd[CTCTCGCACCCATCTCTC], analyses performed on a 125-mm × 4-mm, 5-ìm Lichrospher 100 RP18 column using a linear gradient of 5% to 42.5% acetonitrile in 0.1 M TEAA (100% to 50% mobile phase A versus B) over 50 min at a flow rate of 1 mL/min give a retention time of 15 min 59 sec for the product LY-NH-CO(CH2)5-NH-COd[CTCTCGCACCCATCTCTC], compared to 15 min 25 sec for the starting oligonucleotide. Detection was achieved at both 260 and 420 nm. Figure 4.10.2 shows the UV-visible spectrum of the LY-oligodeoxyribonucleotide conjugate in water. Ma ss an alysis: ESI po larity n egative, LY-NH-CO(CH2)5-NH-COd[CTCTCGCACCCATCTCTC]. Calcd. for C189H240N61O120P17S2: 5875 Da; found 5874 ± 2 Da (M–H). BASIC PROTOCOL 2
CONJUGATION OF 5′-AMINOALKYLATED OLIGODEOXYRIBONUCLEOTIDES TO LIGANDS FUNCTIONALIZED WITH AN ISOTHIOCYANATE OR N-HYDROXYSUCCINIMIDYL GROUP Whereas 5′-aminoalkylated oligodeoxyribonucleotides are conjugated with carboxylated ligands in the presence of a water-soluble carbodiimide, activated ligands such as pentafluorophenyl or N-hydroxysuccinimidyl esters or isothiocyanate derivatives are often used in a wide variety of solvents. The coupling step can be performed in a mixture of aqueous buffer and organic solvents (e.g., DMF, DMSO), or in pure organic solvents. In the latter case, oligodeoxyribonucleotides with lipophilic counter-ions such as cetyltrimethylammonium or triethylammonium are used to facilitate solubilization. The procedure is illustrated in Figure 4.10.3. When the coupling reaction is performed in organic solvent, a tertiary amine is added to ensure that the amino function on the oligodeoxyribonucleotide is unprotonated. This protocol describes the conjugation of 5′-aminoalkylated oligodeoxyribonucleotides with orthophenanthroline (OP) isothiocyanate, Oregon Green (OR) isothiocyanate, and the N-hydroxysuccinimidyl ester of 1-pyrene butanoic acid. Purification procedures are similar to those described in Basic Protocol 1. The optimal purification procedure must be determined empirically. Materials Purified 5′-aminoalkylated oligodeoxyribonucleotide (UNIT 4.9) 0.5 M sodium bicarbonate (NaHCO3) buffer, pH 9.5 5 to 10 mg/mL functionalized ligand (select one) in N,N-dimethylformamide (DMF): Orthophenanthroline (OP) isothiocyanate (UNIT 4.8, structure 3d) N-Succinimidyl ester of 1-pyrene butanoic acid (Molecular Probes) Oregon Green (OR) 5- and 6-isothiocyanate (mixed isomers; Molecular Probes) Ninhydrin 25 mM Tris⋅Cl, pH 7 (APPENDIX 2A), containing 10% (v/v) methanol: without NaCl and with 1 M NaCl (for ion-exchange with a Mono Q column) or with 1.5 M NaCl (for ion-exchange with a DEAE column)
Conjugation of Functionalized Oligodeoxyribonucleotides and Ligands
2-mL vial equipped with Teflon-faced septum Mono Q HR 5/5 or HR 10/10 (Pharmacia; for OP isothiocyanate ion-exchange HPLC)
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Current Protocols in Nucleic Acid Chemistry
100-mm × 10-mm, 8-µm DEAE (Waters; for OP isothiocyante ion-exchange HPLC) HR 10/10 column packed with Lichroprep PR 18 (Art 13900, Merck) or Sephadex G-10 or G-25 (for desalting) Additional reagents and equipment for TLC (see Basic Protocol 1 and APPENDIX 3D) and for isolating, purifying, and characterizing the final conjugate (see Basic Protocol 1) Perform and monitor coupling reaction 1. In a 2-mL vial equipped with a Teflon-faced septum and magnetic stir bar, dissolve 5 OD260 units of purified 5′-aminoalkylated oligodeoxyribonucleotide in 0.2 mL of 0.5 M sodium bicarbonate buffer, pH 9.5. Oligonucleotides are typically purified as triethylammonium salts, in which case 10- to 12-mers are soluble in pure DMSO. Otherwise, counter-ion exchange can be performed to
R1NHC(S)NH(CH2)mNHC(O)O
B
O O
R1N C S
O P O−
NaHCO3 H2N(CH2)mNHC(O)O
O
O
B
B
O
O
m = 5, 6
n
H
O O P O− O
O
B
O H
n R2C(O)NH(CH2)mNHC(O)O
O
B
O
O R2
N O
NaHCO3
O O P O−
O
O
O
B
O n
H N R1 = OP = N HO
O
O
1
R = OR =
F
F COO−
R2 = PY =
Figure 4.10.3 Conjugation of 5′-aminoalkylated oligodeoxyribonucleotides with ligands functionalized with isothiocyanate or N-hydroxysuccinimidyl groups. Abbreviations: OP, orthophenanthroline; OR, Oregon Green; PY, 1-pyrenepropyl.
Synthesis of Modified Oligonucleotides and Conjugates
4.10.7 Current Protocols in Nucleic Acid Chemistry
Supplement 7
prepare cetyltrimethylammonium salts using a procedure adapted from Zarytova et al. (1987). Start with 5 to 10 OD260 units of oligonucleotide in a 100-ìL volume and add 10 vol of 4% LiClO4 in acetone. Vortex 2 min and then centrifuge 5 min at 2000 × g, room temperature. Wash pellet two to three times with distilled acetone to remove the LiClO4. Dry pellet and dissolve in 20 to 30 ìL deionized water. Add 3 ìL of 8% hexadecyltrimethylammonium bromide, vortex, and centrifuge again. Dry the pellet.
2. Add 0.2 mL of 5 to 10 mg/mL functionalized ligand in DMF. Stir the mixture at room temperature overnight. 3. Monitor the reaction by TLC (APPENDIX 3D) using the conditions described above (see Basic Protocol 1, step 3). The presence of OP, pyrene, and their corresponding oligodeoxyribonucleotide conjugates can be easily visualized on a TLC plate by irradiation at 254 or 365 nm. At 254 nm, unmodified and conjugated oligonucleotides can be visualized as grey or black spots, depending on their concentration. At 365 nm, OP-oligonucleotide conjugates appear as slightly fluorescent spots, and pyrene-oligonucleotide conjugates appear as gold fluorescent spots. OR and its oligodeoxyribonucleotide conjugates can be directly visualized on the TLC plate as yellow-green spots. After elution and drying of the TLC plate, the presence of the remaining 5′-amino-containing oligodeoxyribonucleotide can be detected as a grey or black spot by irradiation at 254 nm, or as a pink spot by spraying with ninhydrin and heating the plate.
Recover, purify, and characterize conjugate 4a. For orthophenanthroline: Recover, purify, and characterize conjugate as described above (see Basic Protocol 1, steps 5 to 12) using the appropriate modifications below. a. Reversed-phase HPLC: Use a PRP-1 column with a linear gradient from 100% to 50% mobile phase A versus B over 50 min, and a flow rate of 1 mL/min for both purification and evaluation of efficiency.
Conjugation of Functionalized Oligodeoxyribonucleotides and Ligands
Figure 4.10.4 UV spectrum of the orthophenanthroline-oligodeoxyribonucleotide conjugate OP-NH-C(S)-NH(CH2)5-NH-CO-d[CCGCTTAATACTGA] recorded in water between 230 and 350 nm.
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Current Protocols in Nucleic Acid Chemistry
Figure 4.10.5 UV-visible spectrum of the Oregon Green–oligodeoxyribonucleotide conjugate OR-NH-C(S)-NH(CH2)5-NH-CO-d[CCGCTTAATACTGA] recorded in water between 230 and 600 nm.
b. Ion-exchange HPLC using a Mono Q column: Use a Mono Q HR 5/5 or 10/10 column, a linear gradient of 1 M NaCl (0% to 60%) in 25 mM Tris⋅Cl, pH 7, containing 10% methanol over 40 min, and a flow rate of 1 mL/min (HR 5/5) or 4 mL/min (HR 10/10). After purification, desalt the conjugate using an HR 10/10 column packed with Lichroprep RP18, Sephadex G-10, or G-25, and eluting with water. c. Ion-exchange HPLC using a DEAE column: Use a 100-mm × 10-mm, 8-µm DEAE column, a linear gradient of 1.5 M NaCl (0% to 60%) in 25 mM Tris⋅Cl, pH 7, containing 10% methanol over 60 min, and a flow rate of 1 mL/min. After purification, desalt the conjugate using an HR 10/10 column packed with Lichroprep RP18, Sephadex G-10, or G-25, and eluting with water. For RP-HPLC of OP isothiocyanate coupled to the oligonucleotide H2N-(CH2)5-NH-COd[CCGCTTAATACTGA], analysis on an RP18 column using the conditions described above (see Basic Protocol 1) yields a higher retention time than that expected for oligomer conjugates exhibiting similar lipophilicity; however, peaks are poorly resolved. Therefore, analysis and purification of these conjugates are performed on a PRP-1 column. A retention time of 16 min 10 sec is observed for the conjugate OP-NH-C(S)-NH(CH2)5-NH-CO-
Figure 4.10.6 UV-visible spectrum of the pyrene-oligodeoxyribonucleotide conjugate PYC(O)-NH-(CH2)6-NH-CO-d[CCGCTTAATACTGA] recorded in water between 230 and 400 nm. Current Protocols in Nucleic Acid Chemistry
Synthesis of Modified Oligonucleotides and Conjugates
4.10.9 Supplement 7
d[CCGCTTAATACTGA] compared to a retention time of 12 min 30 sec for the starting oligonucleotide H2N(CH2)5-NH-CO-d[CCGCTTAATACTGA]. When the same conjugate and starting oligonucleotide are analyzed by ion exchange on a Mono Q HR 5/5 column using a linear gradient of 1 M NaCl (20% to 60%) in 25 mM Tris⋅Cl, pH 7, with 10% methanol over 40 min at a flow rate of 1 mL/min, a retention time of 19 min 30 sec is obtained for the conjugate compared to 17 min 30 sec for the starting oligodeoxyribonucleotide. The coupling of the OP derivative to the oligodeoxyribonucleotide does not induce a significant change in the absorption spectrum. Figure 4.10.4 shows the UV spectrum of OP-NH-C(S)-NH(CH2)5-NH-CO-d[CCGCTTAATACTGA] in water. Denaturing polyacrylamide gel electrophoresis allows resolution of the coupling product from the unreacted oligodeoxyribonucleotide. The mobility of the conjugate in the gel is more retarded than that of the starting oligodeoxyribonucleotide. The difference between the electrophoretic mobilities is similar to that observed between H2N(CH2)5-NH-COd[CCGCTTAATACTGA] and d[CCGCTTAATACTGA], shown in Figure 4.9.6.
S− O P O− O
B
O O
R1X
R2COCH2I
n
H R3SSPy
R1S
O
O P O−
O P O− O
B
O
O
O
B
O
n
H
N
HO
R1 =
B
n
H
O
O
O
O
R2 =
OCH3
O O
O n
H
Cl
R2COCH2S
R3SS
O P O−
O
R1 = R3 = COO−
NH
O
(CH2)6
(CH2)6 NH
Acr-NH-(CH2)6-
Pso-(CH2)6FLU O (CH2)8 − I N+
HN R1 =
R1 =
N S
N
N H3C TO-(CH 2)8-
Conjugation of Functionalized Oligodeoxyribonucleotides and Ligands
OP-NHCO(CH2)5-
Figure 4.10.7 Conjugation of oligodeoxyribonucleotide 5′-phosphorothioates with halogenoalkylated ligands (left), an iodoacetamidylated ligand (right), and a 2-pyridyldithioalkylated ligand (center). For halogenoalkylated ligands, R1 = Acr-NH-(CH2)6- (UNIT 4.8, structure 1e); Pso(CH2)6- (UNIT 4.8, structure 2f), OP-C(O)-(CH2)5- (UNIT 4.8, structure 3c); and TO-(CH2)8- (UNIT 4.8, structure 4e). For the iodoacetamidylated ligand, R2 = FLU. For the 2-pyridyldithioalkylated ligand, R3 = P50-(CH2)6- (UNIT 4.8, structure 2h). Abbreviations: Acr, acridine; FLU, fluorescein; OP, orthophenanthroline; Pso, psoralen; PY, 2-pyridyl; TO, thiazole orange.
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Current Protocols in Nucleic Acid Chemistry
4b. For Oregon Green and N-hydroxysuccinimidyl ester of pyrene: Recover, purify, and characterize conjugate as described above (see Basic Protocol 1, steps 5 to 12). For coupling of Oregon Green (OR) 5- or 6-isothiocyanate with oligonucleotide H2N(CH2)5-NH-CO-d[CCGCTTAATACTGA], detection at both 260 and 500 nm yields two peaks for the conjugate OR-NH-C(S)-NH(CH2)5-NH-CO-d[CCGCTTAATACTGA], with retention times of 19 min 5 sec and 20 min 46 sec, corresponding to the two geometrical isomers of OR. The starting H2N-(CH2)6-NH-CO-d[CCGCTTAATACTGA] is eluted with a retention time of 14 min. Figure 4.10.5 shows the UV-visible spectrum of the OR-oligodeoxyribonucleotide conjugate in water. Ma ss an alysis: ESI polarity negative, OR-NH-C(S)-NH-(CH2)5-NH-COd[CCGCTTAATACTGA]. Calcd. for C163H194N53O88P13SF2: 4776 Da; found 4477 ± 1 Da (M–H). For coupling of the N-hydroxysuccinimidyl ester of pyrene (PY) with oligonucleotide H2N(CH2)6-NH-CO-d[CCGCTTAATACTGA], detection at both 260 and 350 nm gives a retention time of 35 min 42 sec for the conjugate PY-NH-C(O)-NH(CH2)6-NH-COd[CCGCTTAATACTGA], compared to 14 min for the starting oligonucleotide. Figure 4.10.6 shows the UV-visible spectrum of the PY-oligodeoxyribonucleotide conjugate in water. Mass analysis: ESI polarity negative, PY-C(O)-NH-(CH2)6-NH-CO-d[CCGCTTAATACTGA]. Calcd. for C163H201N52O84P13: 4636 Da; found 4636 Da (M–H).
CONJUGATION OF OLIGODEOXYRIBONUCLEOTIDE 5′-PHOSPHOROTHIOATES TO LIGANDS FUNCTIONALIZED WITH HALOGENOALKYL, 2-PYRIDYLDITHIO, OR IODOACETAMIDYL GROUPS
BASIC PROTOCOL 3
The derivatization of oligodeoxyribonucleotide 5′-phosphorothioates, illustrated in Figure 4.10.7, is achieved over a wide pH range (5 to 8) with ligands carrying iodoacetamidyl, halogenoalkyl, or 2-pyridyldithio groups. The conjugation reaction can be performed in water, aqueous DMF, or aqueous methanol. The reaction can also be performed in pure methanol in the presence of crown ethers to facilitate solubilization of the oligodeoxyribonucleotide salts (i.e., 18-crown-6 for potassium salts and 15-crown-5 for sodium salts). Procedures are presented for (1) halogenoalkyl derivatives of acridine, psoralen, orthophenanthroline, and thiazole orange; (2) a 2-pyridyldithio derivative of psoralen; and (3) an iodoacetamido derivative of fluorescein. The halogenoalkyl derivatives are reacted with oligodeoxyribonucleotide 5′-phosphorothioates in methanol in the presence of crown ether. The 2-pyridyldithio derivative of psoralen requires minor changes to the conjugation procedure. The protocol for the iodoacetamido derivative is identical to that for the halogenoalkyl-containing ligands; however, since iodoacetamido derivatives are, in general, more reactive than the halogenoalkyl containing ligands, the coupling times are often shorter. Iodoacetamido derivatives of numerous ligands are commercially available. The reader can also refer to Asseline et al. (1996) for preparation of an iodoacetamido derivative of acridine.
Synthesis of Modified Oligonucleotides and Conjugates
4.10.11 Current Protocols in Nucleic Acid Chemistry
Supplement 7
Materials Purified lyophilized oligodeoxyribonucleotide 5′-phosphorothioates (Na+ or K+ salt; UNIT 4.9) 12.5 mg/mL 15-crown-5 or 18-crown-6 (Aldrich) in methanol Functionalized ligand (select one): Halogenoalkylated acridine, psoralen, orthophenanthroline, or thiazole orange (UNIT 4.8, structures 1e, 2f, 3c, and 4e, respectively) 2-Pyridyldisulfide psoralen (UNIT 4.8, structure 2h) Iodoacetamidylated fluorescein (Aldrich) Methanol (for halogenoalkylated ligands and iodoacetamidylated fluorescein) Dichloromethane 0.5 M sodium or potassium phosphate buffer, pH 7 (APPENDIX 2A) Sephadex G-10 or G-15 column 0.1 M triethylammonium acetate (TEAA) buffer, pH 7 2-mL vial equipped with a Teflon-faced septum 30° to 35°C water bath (optional; for halogenoalkylated ligands) UV-Vis spectrophotometer Additional reagents and equipment for TLC (see Basic Protocol 1and APPENDIX 3D) and for isolating, purifying, and characterizing the final conjugate (see Basic Protocol 1) NOTE: In order to prevent chelation of phosphorothioates and phenanthrolines attached to oligodeoxyribonucleotides, all solvents and buffer solutions used for their purification must be passed through a column of Chelex 100 resin to remove divalent cations. Prepare oligodeoxyribonucleotide 5′-phosphorothioate 1. In a 2-mL vial equipped with a Teflon-faced septum and magnetic stir bar, dissolve 5 OD260 units of purified lyophilized oligodeoxyribonucleotide 5′-phosphorothioate (sodium salt or potassium salt) in 0.4 mL of 12.5 mg/mL 15-crown-5 (for sodium salt) or 18-crown-6 (for potassium salt) in methanol. Perform conjugation For halogenalkylated derivatives: 2a. Dissolve 10 eq halogenoalkylated acridine, psoralen, orthophenanthroline, or thiazole orange in a minimal volume of methanol (enough to dissolve the ligand). Add to the oligodeoxyribonucleotide and seal the vial. For the psoralen derivative, also add 0.2 mL dichloromethane and vortex 20 or 30 sec to fully dissolve. 3a. Stir the mixture at room temperature or at 30° to 35°C. The conjugation is usually completed after a few hours (5 to 8 hr), but a longer reaction time is sometimes required (e.g., 24 hr).
4a. Monitor the reaction by TLC using the conditions described above (see Basic Protocol 1, step 3).
Conjugation of Functionalized Oligodeoxyribonucleotides and Ligands
Acridine- and thiazole orange–oligodeoxyribonucleotide conjugates can be easily visualized on TLC plates as yellow and red spots, respectively. Psoralen and orthophenanthroline conjugates can be visualized by irradiation at 254 nm (grey or black spots) or 365 nm (pale fluorescent spots). The presence of unreacted oligodeoxyribonucleotide 5′-thiophosphates can be detected by spraying with DBPNC and heating; the 5′-thiophosphate-containing oligodeoxyribonucleotides give a pink spot.
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Current Protocols in Nucleic Acid Chemistry
Figure 4.10.8 UV-visible spectrum of the acridine-oligodeoxyribonucleotide conjugate AcrNH(CH2)6-S-p-d[CCGCTTAATACTGA] recorded in water between 230 and 530 nm.
For 2-pyridydisulfide psoralen derivative: 2b. Dissolve 5 eq 2-pyridyldisulfide psoralen derivative in 0.2 mL dichloromethane. Add to the oligodeoxyribonucleotide and seal the vial. Vortex for 20 or 30 sec to completely solubilize the mixture. 3b. Stir the mixture at room temperature for 3 to 4 hr. 4b. Monitor the reaction by TLC (step 4a).
Figure 4.10.9 UV-visible spectrum of the psoralen-oligodeoxyribonucleotide conjugate Pso(CH2)6-S-p-d[CCGCTTAATACTGA] recorded in water between 220 and 400 nm.
Synthesis of Modified Oligonucleotides and Conjugates
4.10.13 Current Protocols in Nucleic Acid Chemistry
Supplement 7
For iodoacetamido fluorescein derivative: 2c. Dissolve 10 eq iodoacetamidylated fluorescein in a minimal volume of methanol (enough to dissolve the ligand). Add to the oligodeoxyribonucleotide and seal the vial. 3c. Stir the mixture at room temperature or at 30° to 35°C. The conjugation is usually completed after a few hours (5 to 8 hr), but a longer reaction time is sometimes required (e.g., 24 hr).
4c. Monitor the reaction by TLC (step 4a). Fluorescein conjugates can be visualized directly on the TLC plate as yellow spots.
Analyze, purify, and characterize conjugate 5. When the reaction is complete or nearly complete, add 0.4 mL of 0.5 M sodium or potassium phosphate buffer, pH 7. 6. Extract three times with 1 mL dichloromethane each to remove the bulk of the excess ligand. 7. Remove all remaining ligand by gel filtration on a Sephadex G-10 or G-15 column, eluting with water or 0.1 M TEAA buffer, pH 7. Monitor elution by detection at 260 nm using a UV-Vis spectrophotometer. The size of the column depends on the scale of the synthesis. Usually a 5-mL column is used.
8. Analyze, purify, and characterize the oligodeoxyribonucleotide-ligand conjugates as described above (see Basic Protocol 1, steps 5 to 12). For halogenoalkylated acridine derivative: The conjugate has a retention time of 19 min 32 sec, compared to 13 min 7 sec for the unreacted oligodeoxyribonucleotide 5′-thiophosphate. In addition to detection at 260 nm, a second detection at 425 nm confirms the presence of acridine. Figure 4.10.8 shows the UV-visible spectrum of the acridine-oligodeoxyribonucleotide conjugate in water. Mass analysis: ESI polarity negative, AcrNH(CH2)6-S-p-d[CCGCTTAATACTGA]. Calcd. for C156ClH197N52O85P14S: 4659 Da; found 4657 ± 2 Da (M–H).
Conjugation of Functionalized Oligodeoxyribonucleotides and Ligands
Figure 4.10.10 UV-visible spectrum of the thiazole orange–oligodeoxyribonucleotide conjugate TO-(CH2)8-S-p-d[CCGCTTAATACTGA] recorded in water between 230 and 600 nm.
4.10.14 Supplement 7
Current Protocols in Nucleic Acid Chemistry
Figure 4.10.11 UV-visible spectrum of the psoralen-oligodeoxyribonucleotide conjugate Pso(CH2)6-S-S-p-d[CCGCTTAATACTGA] recorded in water between 220 and 400 nm.
For halogenoalkylated psoralen derivative: The conjugate has a retention time of 25 min 19 sec. In addition to detection at 260 nm, a second detection at 340 nm confirms the presence of psoralen. Figure 4.10.9 shows the UV-visible spectrum of the psoralen-oligodeoxyribonucleotide conjugate recorded in water. Mass analysis: ESI polarity negative, Pso-(CH2)6-S-p-d[CCGCTTAATACTGA]. Calcd. for C153H192N50O88P14S: 4602 Da; found 4603 ± 1 Da (M–H). For halogenoalkylated thiazole orange derivative: The conjugate has a retention time of 26 min 19 sec. In addition to detection at 260 nm, a second detection at 510 nm confirms the presence of thiazole orange. Figure 4.10.10 shows the UV-visible spectrum of the thiazole orange–oligodeoxyribonucleotide conjugate in water. Mass analysis: ESI polarity negative, TO-(CH2)8-S-p-d[CCGCTTAATACTGA]. Calcd. for C162H205N52O84P14S: 4719 Da; found 4720 ± 1 Da (M–H). For halogenoalkylated orthophenanthroline derivative: As previously reported (see Basic Protocol 2), the phenanthroline-oligonucleotide conjugates are poorly eluted on an RP18 reversed-phase HPLC column. Thus, this conjugate is analyzed and purified as described in Basic Protocol 2 (step 4a, substep a) for the coupling of the orthophenanthroline isothiocyanate derivative with a 5′-aminoalkylated oligodeoxyribonucleotide. Analysis on
Figure 4.10.12 UV-visible spectrum of the fluorescein-oligodeoxyribonucleotide conjugate FLU-NH-C(O)-CH2-S-p-d[CCGCTTAATACTGA] recorded in 0.1 M sodium bicarbonate buffer, pH 9, between 230 and 600 nm.
Synthesis of Modified Oligonucleotides and Conjugates
4.10.15 Current Protocols in Nucleic Acid Chemistry
Supplement 7
a PRP-1 column using a gradient from 5% to 42.5% CH3CN in 0.1 M TEAA, pH 7, over 50 min at a flow rate of 1 mL/min gives a retention time of 15 min 24 sec for the conjugate OP-NH-C(O)-(CH2)5-S-p-d[CCGCTTAATACTGA], compared to 11 min 21 sec for the starting oligonucleotide 5′-thiophosphate. As in Basic Protocol 2, this coupling of orthophenanthroline halogenoalkyl derivative with the oligodeoxyribonucleotide 5′-thiophosphate does not induce significant changes in the absorption spectrum. For 2-pyridyldisulfide psoralen: Analysis under these conditions gives a retention time of 28 min 7 sec for the conjugate. As in the case of the halogenoalkyl derivative of psoralen, additional detection at 340 nm confirms the presence of psoralen. Figure 4.10.11 shows the UV-visible spectrum of the psoralen-oligodeoxyribonucleotide conjugate (via a disulfide bond) in water. Mass analysis: ESI polarity negative, Pso-(CH2)6-S-S-pd[CCGCTTAATACTGA]. Calcd. for C153H192N50O88P14SS: 4634 Da; found 4633 ± 1 Da (M–H). For 5′-iodoacetamidylated fluorescein: The conjugate of the 5′-iodoacetamidylated fluorescein (isomer 5; FLU) with S-p-d[CCGCTTAATACTGA] has a retention time of 17 min 18 sec using an RP18 reversed-phase HPLC column under the conditions described above. Figure 4.10.12 shows the UV-visible spectrum of the fluorescein-oligodeoxyribonucleotide conjugate in 0.1 M sodium hydrogen carbonate buffer, pH 7. Mass analysis: ESI polarity n ega tive, FLU-NH-C(O)-CH2- S- p- d[CCGCTTAATACTGA]. Calcd. for C158H189N51O90P14S: 4707 Da; found 4708 Da (M–H). Ion-exchange for halogenoalkylated ligand-oligodeoxyribonucleotide conjugates: Analysis and purification of these conjugates can also be performed on a Mono Q or DEAE ion-exchange column using the eluents and conditions described above (see Basic Protocol 2, step 4a, substeps b and c). The conjugate is retained more than the oligonucleotide 5′-thiophosphate because of the hydrophobicity of the ligand. For example, using a Mono Q column under the conditions described in Basic Protocol 2, the conjugate OP-NH-C(O)(CH2)5-S-p-d[CCGCTTAATACTGA] is eluted at 28 min 30 sec, compared to 26 min for the starting oligodeoxyribonucleotide 5′-phosphorothioate. PAGE of halogenoalkylated ligand-oligodeoxyribonucleotide conjugates: Denaturing polyacrylamide gel electrophoresis (UNIT 10.4) allows the resolution of the conjugate OP-NH-C(O)-(CH2)5-S -p -d[CCGCTTAATACTGA] from unreacted S-pd[CCGCTTAATACTGA]. The electrophoretic mobility of the conjugate on the gel is retarded when compared to that of the oligodeoxyribonucleotide 5′-thiophosphate. A third band with weak intensity and lower electrophoretic mobility relative to the other two bands is also observed. This band may correspond to the dimer of the oligodeoxyribonucleotide 5′-phosphorothioate. Mass analysis: ESI polarity negative, OP-NH-C(O)-(CH2)5-S-pd[CCGCTTAATACTGA]. Calcd. for C154H193N53O85P14S: 4609 Da; found 4610 ± 1 Da (M-H).
R OH
NH −
O P O−
O P O O
O
RNH2
B
O
O
B
EDC O
Conjugation of Functionalized Oligodeoxyribonucleotides and Ligands
H
O n
H
n
Figure 4.10.13 Condensation reaction of oligodeoxyribonucleotide 5′-phosphates with aminated compounds. R, H2N(CH2)5CH2-.
4.10.16 Supplement 7
Current Protocols in Nucleic Acid Chemistry
CONJUGATION OF OLIGODEOXYRIBONUCLEOTIDE 5′-PHOSPHATES TO LIGANDS FUNCTIONALIZED WITH AMINO GROUPS
BASIC PROTOCOL 4
The conjugation of oligodeoxyribonucleotides carrying a phosphate group at the 5′ terminus with the primary amino function of a ligand is accomplished in the presence of carbodiimide. It generally requires a large excess of both ligand and carbodiimide (100 to 500 eq). Usually the oligodeoxyribonucleotide is solubilized in a buffer and the ligand is added to the solution in amine-free DMF. The procedure is illustrated in Figure 4.10.13 and exemplified by the coupling of 1,5-diaminopentane with oligodeoxyribonucleotide 5′-phosphates. Materials Purified oligodeoxyribonucleotide 5′-phosphate (UNIT 4.9) 0.5 M aqueous 1,5-diaminopentane, pH 4.5 1-[3-(Dimethylamino)propyl]-3-ethylcarbodiimide hydrochloride (EDC) 4% (w/v) LiClO4 in acetone Mobile phase A: 0.1 M triethylammonium acetate buffer pH 7, containing 5% CH3CN Mobile phase B: 0.1 M triethylammonium acetate (TEAA) buffer, pH 7 containing 80% CH3CN 2-mL vial equipped with a Teflon-faced septum HPLC system (also see Basic Protocol 1) Column: 150-mm × 3-mm, 5-µm RP8 glass cartridge system (Merck) Additional reagents and equipment for TLC (see Basic Protocol 1 and APPENDIX 3D) and for isolating, purifying, and characterizing the final conjugate (see Basic Protocol 1) 1. In a 2-mL vial equipped with a Teflon-faced septum and magnetic stir bar, dissolve 5 OD260 units of purified oligodeoxyribonucleotide 5′-phosphate in 0.2 mL of 0.5 M aqueous 1,5-diaminopentane, pH 4.5. 2. Add 10 mg water-soluble EDC and incubate with stirring at room temperature for 4 to 5 hr. 3. Divide the reaction mixture between two microcentrifuge tubes. 4. Add 1 mL of 4% (w/v) LiClO4 in acetone to each tube. Vortex for at least 1 min and centrifuge for 5 min at 2000 × g, room temperature. Discard supernatant and dissolve the residue in 0.1 mL water. Repeat three times. 5. Analyze condensation product by reversed-phase HPLC using a 150-mm × 3-mm, 5-µm RP8 glass cartridge system with a gradient from 100% to 50% mobile phase A versus B, over 50 min at a flow rate of 0.5 mL/min. Usin g these co nd ition s, the co nd en sation product H2N-(CH2)5-NH-pd[TTCTCCCCCGCTTA] shows one peak with a retention time of 15 min 34 sec, compared to 14 min 45 sec for the oligodeoxyribonucleotide 5′-phosphate. Mass analysis: ESI p ola rity neg ative, H2N-(CH2)5-NH-p-d[TTCTCCCCCGCTTA]. Calcd. for C138H189N43O88P14: 4289 Da; found 4289 ± 1 Da (M–H). The presence of the primary amino function at the end of the linker can be verified after TLC elution by spraying the plate with ninhydrin solution and heating.
Synthesis of Modified Oligonucleotides and Conjugates
4.10.17 Current Protocols in Nucleic Acid Chemistry
Supplement 7
BASIC PROTOCOL 5
CONJUGATION OF 5′-MERCAPTOALKYLATED OLIGODEOXYRIBONUCLEOTIDES TO LIGANDS FUNCTIONALIZED WITH HALOGENOALKYL, IODOACETAMIDYL, OR 2-PYRIDYLDITHIO GROUPS This coupling reaction can be performed in buffer (pH 7 to 9), in a mixture of buffer and DMF, or directly in DMF using oligodeoxyribonucleotides converted to their cetyltrimethylammonium salts. Solutions must be degassed by bubbling with argon or nitrogen. Halogenoalkylated, iodoacetamidylated, and 2-pyridyldithioalkylated derivatives of various ligands are conjugated with an oligodeoxyribonucleotide bearing a masked mercaptoalkylated linker at its 5′ end. The procedures are illustrated in Figure 4.10.14. The coupling of halogenoalkylated thiazole orange with HS-CH2CH2-(OCH2CH2)2-pd[CTCTCGCACCCATCTCTC] is performed in situ by treatment of C5H5N-S-SCH2CH2-(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC] with the reducing agent Tris-(2-carboxyethyl)phosphine (TCEP). The coupling of iodoacetamidyl ligands with 5′-mercaptoalkylated oligodeoxyribonucleotides is performed using the same procedure.
O R1S
O P O L O− O
1. TCEP
O P O−
2. R 1X
O
O
S S
B
O
O N
B
O
O P O L O−
O
n
H
B
O O P O− O
B
O O
n
H
O R2S
S
O P O L O−
B
O O
1. TCEP
O P O−
2. R 2SSPy
O
O
B
O H
(CH2)8 − I N+
n
(CH2)6
R1 =
R2 =
O
S N
O
O
O
H3C TO-(CH 2)8-
Conjugation of Functionalized Oligodeoxyribonucleotides and Ligands
Pso-(CH2)6-
Figure 4.10.14 Conjugation of oligodeoxyribonucleotides bearing a 5′-(2-pyridyldithioalkylated) linker with a halogenoalkylated ligand (top) or a 2-pyridyldithioalkylated ligand (bottom). R1 = TO-(CH2)8- (UNIT 4.8, structure 4e); R2 = Pso-(CH2)6- (UNIT 4.8, structure 2h). Abbreviations: T C E P, Tr i s - ( 2 - c a r box yet hy l ) ph os ph in e hy dr o c h lo r id e; X , I; P y, 1-pyr idyl; L, CH2CH2OCH2CH2OCH2CH2.
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Current Protocols in Nucleic Acid Chemistry
The coupling of 2-pyridyldithioalkylated ligands with 5′-(2-pyridyldithioalkylated)-oligodeoxyribonucleotides is quite similar, but in this case the amount of TCEP must be strictly controlled in order to prevent the cleavage of the product, which also contains a disulfide bridge. This protocol describes the condensation of C5H5N-S-S-CH2CH2(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC] with 2-pyridyldithioalkylated psoralen. Conjugate Halogenoalkylated and Iodoacetamidylated Ligands to 5′-Mercaptoalkylated Oligodeoxyribonucleotides Materials Purified 5′-(2-pyridyldithioalkylated) oligodeoxyribonucleotide (UNIT 4.9) 30 mM sodium bicarbonate buffer, pH 9 Dimethylformamide (DMF) Nitrogen or argon gas Tris-(2-carboxyethyl)phosphine (TCEP) Halogenoalkylated or iodoacetamidylated ligand (e.g., halogenoalkylated thiazole orange; UNIT 4.8, structure 4e) Diisopropylethylamine (optional) 2-mL vial equipped with a Teflon-faced septum Additional reagents and equipment for TLC (see Basic Protocol 1 and APPENDIX 3D) and for isolating, purifying, and characterizing the final conjugate (see Basic Protocol 1) 1a. In a 2-mL vial equipped with a Teflon-faced septum and magnetic stir bar, dissolve 10 OD260 units of purified 5′-(2-pyridyldithioalkylated) oligodeoxyribonucleotide in 0.15 to 0.3 mL of the desired solvent—i.e., 30 mM sodium bicarbonate buffer, pH 9, buffer/DMF, or DMF. The solvent must be chosen empirically. The resulting reaction mixture must be homogeneous.
2a. Degas the solution by bubbling argon or nitrogen gas through it for 10 min. 3a. Dissolve 2 or 3 eq TCEP in a minimal volume (2 to 3 µl) of water and add to the degassed oligonucleotide solution. Leave 30 min at room temperature. 4a. Dissolve 5 to 20 eq halogenoalkylated or iodoacetamidylated ligand in 0.15 to 0.3 mL DMF and add to the degassed oligonucleotide solution. If the conjugation reaction is performed in DMF without buffer, also add 5 µl diisopropylethylamine. 5a. Stir the mixture at room temperature or at 30° to 35°C for 3 to 4 hr. Monitor reaction by TLC (see Basic Protocol 1, step 3). The thiazole orange conjugate is readily visible on the TLC plate as a red spot.
6a. Analyze, purify, and characterize the conjugate as described for condensation of halogenoalkylated ligands with oligodeoxyribonucleotide 5′-thiophosphates (see Basic Protocol 1, steps 5 to 12). Analysis of the coupling reaction between the thiazole orange (TO) derivative and C5H5NS-S-CH2CH2-(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC] is performed on an RP18 (5% to 42.5% of CH3CN over 50 min in 0.1 M TEAA, pH 7 at a flow rate of 1 mL/min) reversed-phase HPLC column using the conditions described in Basic Protocol 3. The conjugate TO-(CH2)8-S-CH2CH2-(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC] elutes with a retention time of 24 min 25 sec. A second peak at 18 min 11 sec corresponds to either th e th iol-co nta inin g o ligo deoxyrib on ucleotide HS-CH2CH2-(OCH2CH2)2-pd [CTCTCGCACCCATCTCTC] or its (5′-5′)- dimer d erivative 3′d[CTCTCTACCCACGCTCTC]-p-CH2CH2-(OCH 2CH2)2-S-CH2CH2-(OCH2CH2)2-pd[CTCTCGCACCCATCTCTC]. The presence of thiazole orange in the conjugate is
Synthesis of Modified Oligonucleotides and Conjugates
4.10.19 Current Protocols in Nucleic Acid Chemistry
Supplement 7
Figure 4.10.15 UV-visible spectrum of the psoralen-oligodeoxyribonucleotide conjugate Pso(CH2)6-S-S-CH2CH2-(OCH2CH2 )2-p-d[CTCTCGCACCCATCTCTC] recorded in water between 230 and 400 nm.
detected at 260 and 510 nm. Mass analysis: ESI polarity negative, TO-(CH2)8-S-CH2CH2(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC]. Calcd. for C203H264N57O114S2P18: 5948 Da; found 5955 ± 3 Da (M–H).
Conjugate 2-Pyridyldisulfide-Containing Ligands With 5′-Mercaptoalkylated Oligodeoxyribonucleotides Materials Purified 5′-(2-pyridyldithioalkylated) oligodeoxyribonucleotides (UNIT 4.9) 30 mM sodium bicarbonate buffer, pH 9 Nitrogen or argon gas Tris-(2-carboxyethyl)phosphine (TCEP) 2-Pyridyldithioalkylated ligand (e.g., 2-pyridyldisulfide derivative of psoralen; UNIT 4.8, structure 2f) Dimethylformamide (DMF) 2-mL vial equipped with a Teflon-faced septum Additional reagents and equipment for TLC (see Basic Protocol 1 and APPENDIX 3D) and for isolating, purifying, and characterizing the final conjugate (see Basic Protocol 1) 1b. In a 2-mL vial equipped with a Teflon-faced septum and magnetic stir bar, dissolve 10 OD260 units of purified 5′-(2-pyridyldithioalkylated) oligodeoxyribonucleotide in 0.2 mL of 30 mM sodium bicarbonate buffer, pH 9. 2b. Degas the solution by bubbling argon or nitrogen through it for 10 min. 3b. Dissolve ≤2 eq TCEP in a minimal volume (2 to 3 µl) of water and add to the degassed oligonucleotide solution. Leave 90 min at room temperature.
Conjugation of Functionalized Oligodeoxyribonucleotides and Ligands
4b. Dissolve 1.2 eq 2-pyridyldithioalkylated ligand in 360 µl DMF and add to the degassed oligonucleotide solution. 5b. Stir the mixture at room temperature for 4 hr. Monitor reaction by TLC (see Basic Protocol 1, step 3).
4.10.20 Supplement 7
Current Protocols in Nucleic Acid Chemistry
The psoralen conjugate can be visualized as a pale blue fluorescent spot by irradiation at 365 nm.
6b. Analyze, purify, and characterize the conjugate as described for condensation of halogenoalkylated ligands with oligodeoxyribonucleotide 5′-thiophosphates (see Basic Protocol 1, steps 5 to 12). Analysis of the coupling reaction between the psoralen derivative and the oligonucleotide HS-CH2CH2-(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC] is performed on an RP18 reversed-phase HPLC column using the conditions described above (see Basic Protocol 1). The conjugate Pso-(CH2)6-S-S-CH2CH2-(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC] has a retention time of 35 min 38 sec. A second peak with a retention time of 18 min 11 sec corresponds to either the thiol-containing oligodeoxyribonucleotide HS-CH2CH2(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC] or its (5′-5′)-dimer derivative 3′[CTCTCTACCCACGCTCTC]-p-CH2CH2-(OCH2CH2)2-S-S-CH2CH2-(OCH2CH2)2-pd[CTCTCGCACCCATCTCTC]. The psoralen-containing conjugate can be detected at 260 and 320 nm. The UV-visible spectrum of the psoralen-oligodeoxyribonucleotide conjugate in water is shown in Figure 4.10.15. Mass analysis: ESI polarity negative, Pso-(CH2)6-SCH2CH2-(OCH2CH2)2-p-d[CTCTCGCACCCATCTCTC]. Calcd. for C194H249N55O118S2P18: 5852 Da; found 5852 ± 2 Da (M–H).
COMMENTARY Background Information The methods reported in this unit describe the conjugation of ligands containing halog en oalkyl, iodoacetamidyl, and 2pyridylthioalkyl linkers with oligodeoxyribonucleotides functionalized with either a 5′thiophosphate or a 5′-pyridyldithio group. When using halogenoalkylated or iodoacetamidylated ligands, the conjugates obtained have irreversible phosphothioester or thioether linkages. In contrast, the conjugates obtained from 2-pyridylthioalkylated ligands have phosphodisulfide or disulfide bonds that can be cleaved by the use of a reducing agent such as TCEP or dithiothreitol (DTT). These reactions, which proceed without activation, can be performed either in organic solvents (using a crown ether to solubilize the oligonucleotides) or in a mixture of organic solvents and aqueous buffer. The conjugation of oligodeoxyribonucleotides carrying aminoalkylated linker is illustrated by examples using isothiocyanate or N-hydroxysuccinimidyl derivatives of ligands. In these cases, the amino function must be unprotonated and the reaction is carried out in the presence of buffer. The coupling of 5′-modified oligonucleotides with phosphate and carboxylated linker with nucleophilic ligands such as primary amine or hydrazino derivatives requires the use of a coupling reagent. Procedures reported in this unit may be used to prepare oligodeoxyribonucleotides covalently linked via their 5′ ends to a wide variety of molecules such as labels, intercalating
agents, and reactive compounds, the properties of which are reported in UNIT 4.2. In many cases, the properties of the oligodeoxyribonucleotideligand conjugates are largely dependent on the geometry of the complex formed between the ligand and its target. The linkage between the oligodeoxyribonucleotide and the ligand (i.e., the size and nature of the linker) may be varied to optimize the properties of oligodeoxyribonucleotide-ligand conjugates. The conjugation methods presented in this unit can be used to prepare oligonucleotide conjugates composed of natural β-deoxyribonucleosides, phosphorothioate oligodeo xy rib on ucleo tides, α-D-deoxyribonucleosides or 2′-O-methylribonucleosides, and many other modified oligonucleotides. The methods reported for conjugating 5′-modified oligodeoxyribonucleotides with appropriately functionalized ligands are also valid for coupling these ligands with the same functional groups incorporated into other positions of the oligonucleotides (i.e., phosphate thiophosphate, thiolcarboxyl, and amino at the 3′ end); or thiol carboxyl, and amino attached via a linker to the 2′ position of the sugar, nucleobase, or internucleotide phosphate. Ligands functionalized with an iodoacetamidyl group can also be reacted with internucleotide thiophosphate groups (Asseline et al., 1996). The methods reported in UNIT 4.9 for introducing functional groups at the 5′ ends of oligodeoxyribonucleotides are compatible in most cases with the presence of a second functional group at another position in the oligodeoxyri-
Synthesis of Modified Oligonucleotides and Conjugates
4.10.21 Current Protocols in Nucleic Acid Chemistry
Supplement 7
bonucleotide. It is thus possible to prepare bisderivatized oligonucleotides with two different ligands (Aubert et al., 2000). The conjugation of functionalized ligands to unprotected oligonucleotides carrying suited functional groups offers many advantages over the direct incorporation of the ligands during oligonucleotide synthesis (UNIT 4.3). In particular, this method is very useful when a limited amount of the ligand is available, when the ligand does not resist the chemical conditions required for oligonucleotide deprotection, or when it is only weakly soluble in the solvents needed for oligonucleotide synthesis, thus preventing the preparation of a phosphoramidite or H-phosphonate derivative. Furthermore, the purification of oligonucleotide-ligand conjugates, and particularly those involving two ligands, can be easier using this method, because the oligonucleotides are purified prior to their coupling with ligands, which is not the case when ligands are incorporated during oligonucleotide chain elongation before deprotection. Lastly, as long as only small amounts of conjugate are required, this method is the most convenient one for obtaining many different oligodeoxyribonucleotide-ligand conjugates starting from only one oligodeoxyribonucleotide synthesis.
Critical Parameters
Conjugation of Functionalized Oligodeoxyribonucleotides and Ligands
For TLC analyses of conjugation reactions, the reservoir containing the solvent mixture (including concentrated ammonia) must be well-saturated. It is better to prepare it at least half a day before use. Before loading HPLC columns, organic impurities, such as excess ligands and coupling reagents, should be extracted from crude oligodeoxyribonucleotide solutions with organic solvents or removed by gel filtration. The oligodeoxyribonucleotide solutions must then be filtered through a 0.45-µm disposable filter to remove any particulates and prevent the clogging of columns. An important parameter is the choice of the solvents used to perform the conjugation reactions since oligodeoxyribonucleotides are soluble in water and in buffer solutions, while most ligands are not. A second important parameter is the choice of the method used to recover the oligonucleotide-ligand conjugate prior to its purification by chromatography. When working with a new ligand, it is critical to experiment with different methods on an analytical scale in order to chose the best one.
When working with acridine-containing oligodeoxyribonucleotides, the pH of all solutions must be kept below 7 or above 9.5 in order to prevent the cleavage of the bond between the acridine ring and the linker. The coupling of halogenoalkylated orthophenanthroline derivatives with oligodeoxyribonucleotide 5′-thiophosphates sometimes gives low product yields. Liquid chromatography analysis (Thuong and Asseline, 1991) shows, in addition to the starting oligodeoxyribonucleotide 5′-thiophosphate and the expected oligodeoxyribonucleotide-phenanthroline conjugate, the presence of two other products identified as the (5′-5′)-oligodeoxyribonucleotide phosphorothioate dimer obtained by disulfide bond formation, and the corresponding oligodeoxyribonucleotide 5′-phosphate. The formation of these side-products can be explained by an oxidoreduction reaction triggered by the simultaneous presence of thiophosphate, oxygen, orthophenanthroline, and divalent cations. It is important to note that the characterization of orthophenanthroline-oligonucleotide conjugates by mass spectrometry is often difficult. Oligodeoxyribonucleotides containing a thiol function require degassing of solvents and buffer solutions in order to remove oxygen and any oxidizing reagents, which might lead to the formation of a dimer via disulfide bond formation. Under the conditions described in these protocols, reactivity of oligonucleotide and ligand functional groups with nucleobases should not be problematic. For 5′-carboxylated oligonucleotides, amide linkages should not form with the exocyclic amino function of the nucleobase at the recommended pH (5.5 to 6), as it has been reported that, under these conditions, nucleobases are not modified at pH ≤6 (Ivanovskaya et al., 1987). Ligands functionalized with isothiocyanate or N-hydroxysuccinimide ester groups should also not react with the primary amino function of the nucleobase, which is less reactive than the primary aliphatic amino function at the linker extremity. Succinimidyl esters have a high selectivity for reaction with aliphatic amines. Aromatic isothiocyanates have also been reported to selectively react with aliphatic primary amines (Smith et al., 1985). Finally, aminoalkylated compounds should not form phosphoramidate linkages with internucleotidic phosphodiesters, which are less reactive than the terminal phosphomonoester group. This modification has been reported to be highly selective, and internucleotidic phos-
4.10.22 Supplement 7
Current Protocols in Nucleic Acid Chemistry
phates are not affected (Ivanovskaya et al., 1987). Oligonucleotide-ligand conjugates should be lyophilized and stored in tightly closed vials at −20°C. The ligands should be protected from light at all times, since they are sensitive to light (UNIT 4.8). Under these conditions, they are stable for a year or more. RP-HPLC analyses of four oligonucleotide-ligand conjugates (Figs. 4.10.2, 4.10.8, 4.10.9, 4.10.10) that were stored as aqueous solutions for 18 months in the dark at –20°C showed no detectable degradation.
Anticipated Results The yields obtained for the conjugates are good, typically exceeding 80% for reactions involving 5′-thiophosphate-, 5′-amino-, and 5′thiol-containing oligonucleotides. The yields for 5′-carboxylated and 5′-phosphate oligonucleotides are more variable, ranging from 50% to 80%. The yield of recovered oligonucleotide-ligand conjugate after purification and desalting depends on the nature of the ligand and on the scale of the synthesis. When working on a micromolar scale, it is possible to obtain 20 to 25 OD260 units of the acridine- or psoralen-oligonucleotide conjugate. Purification of 5′-oligonucleotide conjugates by ionexchange chromatography leads to lower recovery yields because of the requirement for an additional desalting step. The thiazole orange– oligodeoxyribonucleotide conjugate is also obtained in good yield (6 OD260 units are recovered from 10 OD260 units of the 5′-modified oligodeoxyribonucleotide starting material); however, when reactions are performed with only 4 to 5 OD260 units of 5′-modified oligodeoxyribonucleotides, the amount of purified oligonucleotide-ligand conjugate obtained is ∼1.5 to 2 OD260 units.
Time Considerations Provided that all reagents and materials required for each step are available, most of the procedures described herein are simple and rapid. The time required to perform each protocol—including purification, characterization, and lyophylization—is ∼1 week.
Literature Cited Asseline, U., Bonfils, E., Dupret, D., and Thuong, N.T. 1996. Synthesis and binding properties of oligonucleotides covalently linked to an acridine derivative: New study of the influence of the dye attachment site. Bioconjugate Chem. 7:369-379. Aubert, Y., Bourgerie, S., Meunier, L., Mayer, R., Roche, A.-C., Monsigny, M., Thuong, N.T., and Asseline, U. 2000. Optimized synthesis of phosphorothioate oligodeoxyribonucleotides substituted with a 5′-protected thiol function and a 3′-amino group. Nucl. Acids Res. 28:818-825. Aubert, Y., Perrouault, L., Hélene, C., Giovannangeli, C., and Asseline, U. 2001. Synthesis and properties of triple helix-forming oligodeoxyribonucleotides containing 7-chloro-7-deaza-2′deoxyguanosine. Bioorg. Med. Chem. 9:16171624. Ivanovskaya, M.G., Gottihk, M.B., and Shabarova, Z.A. 1987. Modification of Oligo(poly)nucleotide Phosphomonoester groups in aqueous solutions. Nucleosides-Nucleotides 6:913-934. Raymond, F., Asseline, U., Roig, V., and Thuong, N.T. 1996. Synthesis and characterization of O6 modified deoxyguanosine-containing oligodeoxyribonucleotides for triple helix formation. Tetrahedon 52:2047-2064. Smith, L.M., Fung, S., Hunkapiller, M.W., Hunkapiller, T.J., and Hood, L.E. 1985. The synthesis of oligonucleotides containing an aliphatic amino group at the 5′ terminus: Synthesis of fluorescent DNA primers for use in DNA analysis. Nucl. Acids. Res. 13:2399-2412. Thuong, N.T. and Asseline, U. 1991. Oligodeoxyribonucleotides attached to intercalators, photoreactive and cleavage agents. In Oligodeoxyribonucleotides and Analogues: A Practical Approach (F. Eckstein, ed.) pp. 283-308. IRL Press, Oxford.
Contributed by Ulysse Asseline and Nguyen T. Thuong Centre de Biophysique Moleculaire, CNRS Orléans, France The authors would like to express their appreciation to past and present collaborators for their contribution to the development of varied families of oligonucleotide conjugates over the past years. This work was supported by Rhône-Poulenc, Agence Nationale de Recherche contre le SIDA, and bioMérieux.
Synthesis of Modified Oligonucleotides and Conjugates
4.10.23 Current Protocols in Nucleic Acid Chemistry
Supplement 7
Synthesis and Purification of Peptide Nucleic Acids
UNIT 4.11
Peptide nucleic acids (PNAs) are DNA analogs in which the normal phosphodiester backbone is replaced by 2-aminoethyl glycine linkages (Fig. 4.11.1). In spite of this significant chemical modification, the hybridization of PNAs with DNA or RNA follows normal rules for Watson-Crick pairing and occurs with high affinity. The fact that PNAs possess a dramatically different backbone and bind efficiently has created enormous interest in their application. This unit describes the synthesis and purification of PNAs. PNAs can be synthesized using either automated procedures (see Basic Protocol 1; Mayfield and Corey, 1999) or manual procedures (see Basic Protocol 2; Norton et al., 1995). The goal here is to provide the reader with the basic information necessary to initiate experiments with PNAs. NOTE: It is essential that the reagents used for PNA synthesis, whether automated or manual, be kept as anhydrous as possible, because contamination with water will result in incomplete coupling of monomers during synthesis. AUTOMATED SYNTHESIS OF PEPTIDE NUCLEIC ACIDS The automated synthesis of PNAs is illustrated in Figure 4.11.2. Automated synthesis is a convenient strategy for obtaining PNAs, and is performed using the Expedite synthesizer (Applied Biosystems). This instrument has been specially adapted for PNA synthesis, with proprietary software for FMOC chemistry instead of the phosphoramidite chemistry used for typical oligonucleotide synthesis. It is probably possible to adapt other synthesizers for PNA synthesis; however, the investigator will need to consider whether the number of PNAs needed justifies the time and expense required to adapt the instrumentation.
BASIC PROTOCOL 1
The reagents used are available from Applied Biosystems separately or as an Expedite PNA kit. Two resins are available: a xanthen alkonic acid (XAL) resin and a peptide amide linker (PAL) resin. Although either is adequate for automated synthesis, the XAL resin is preferred because the cleavage step is considerably shorter (5 min according to Applied Biosystems, compared to 90 min for the PAL resin).
N 2H
base N O NH O base N O OH O
Figure 4.11.1 Structure of a peptide nucleic acid. Contributed by Dwaine A. Braasch, Christopher J. Nulf, and David R. Corey Current Protocols in Nucleic Acid Chemistry (2002) 4.11.1-4.11.18 Copyright © 2002 by John Wiley & Sons, Inc.
Synthesis of Modified Oligonucleotides and Conjugates
4.11.1 Supplement 9
coupling step base*
NHFMOC
P
O
deprotection 20% piperidine in DMF
base*
N
O N
HO
NH2
P
NHFMOC
O
HATU, 0.2 M DIPEA 0.3 M 2, 6-lutidine, DMF, NMP
N H
P
1. capping step 5% Ac2O, 6% 2, 6-lutidine in DMF 2. deprotection
n=n+1
n
base*
P
N H
coupling step
N
N H
O
N H
P
NHFMOC
O
trace of "capped" unreacted oligomer
+
N
NH2
O
1. deprotection 2. cleavage from the support 20% m -cresol in TFA
when n = n max
base
n max
base O
O
N
HO
O
O
N
n
base*
base*
O
NHFMOC
O
N
N H
O
NH2
O
PNA monomers NHBHOC
O
NH
N
N
N
N
O
O
N
O
O
HO
N
N
N
N
O
NHFMOC
HO
NH
N
NHBHOC
O N
N
NHFMOC
O
O
O N
HO
NHBHOC
O
N
NHFMOC
HO
O
NHFMOC
Figure 4.11.2 Automated PNA synthesis as described in Basic Protocol 1. Abbreviations: Ac2O, acetic anhydride; base*, N-protected nucleobase (see PNA monomers); BHOC, benzhydryloxycarbonyl; DIPEA, diisopropylethylamine; DMF, N,Ndimethylformamide; FMOC, 9-fluorenylmethoxycarbonyl; HATU, O-(7-azabenzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate; NMP, N-methylpyrrolidone; P, FMOC-XAL-PEG-polystyrene; TFA, trifluoroacetic acid.
2-Aminoethoxy-2-ethoxy acetic acid (AEEA) is used as a linker molecule for PNA synthesis. A linker molecule is used to introduce a controlled space between PNAs or between a PNA and some other functional or spatially addressable large group, such as rhodamine or biotin. Linked PNAs (bis-PNAs) possess an enhanced ability to bind by strand invasion and are an important tool for recognition of duplex DNA. Linker coupling is often inefficient, so it is always done two times.
Synthesis and Purification of Peptide Nucleic Acids
Although the synthesis described here is an automated process, this does not excuse the experimenter from closely monitoring the various steps throughout the synthesis. Careful observation to ensure proper machine function can result in catching errors as they occur and allow manual intervention that can rescue the synthesis and prevent loss of reagents and time (see Critical Parameters).
4.11.2 Supplement 9
Current Protocols in Nucleic Acid Chemistry
Materials PNA Expedite reagents (Applied Biosystems) FMOC-PNA monomers (Fig. 4.11.2): 9-fluorenylmethoxycarbonyl-protected peptide nucleic acid monomers (A, T, C, and G), base protected with benzhydryloxycarbonyl (BHOC) Diluent: N-methylpyrrolidone (NMP) Activator: 7-aza-1-hydroxybenzotriazole (HOAt) or O-(7-azabenzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HATU) Linker: 2-aminoethoxy-2-ethoxy acetic acid (AEEA) Base solution: 0.2 M diisopropylethylamine (DIPEA)/0.3 M 2,6-lutidine Deblocking solution: 20% (v/v) piperidine in N,N-dimethylformamide (DMF) Capping solution: 5% (v/v) acetic anhydride/6% (v/v) 2,6-lutidine in DMF N,N-Dimethylformamide (DMF), anhydrous (Burdick Jackson; wash A is Opti-Dry DMF from Fisher; wash B is anhydrous DMF from Applied Biosystems) Amino acids (Novabiochem, Advanced Chemtech) Isopropyl alcohol (optional) Cleavage cocktail: 20% (v/v) m-cresol (Sigma-Aldrich) in trifluoroacetic acid (TFA; Burdick Jackson) Diethyl ether, –20°C Expedite 8909 synthesizer (Applied Biosystems) FMOC-XAL-PEG-PS synthesis column (0.2 µmol prepacked; Applied Biosystems) 10-mL syringe 1.5-mL, 0.2-µm polytetrafluoroethylene (PTFE) or regenerated cellulose spin column (Millipore) Additional reagents and equipment for automated synthesis (see manufacturer’s instructions) and for purification and analysis of PNAs (see Basic Protocol 3) NOTE: Powdered reagents such as monomers, activator, and linker (AEEA) should be unpacked on arrival and stored at –20°C in a sealed container containing Drierite desiccant. Monomers should be inspected upon arrival. Clumps of reagent may indicate that water has been introduced during shipping. NOTE: The authors use three sources of DMF because the bottles from Fisher (Opti-Dry) and Applied Biosystems fit onto the input fitting and reagent port on the synthesizer, and the third can be used following synthesis. All DMF must be anhydrous and should have a low amine content to reduce the likelihood of side reactions. DMF should be purchased in 100-mL volumes to ensure that it is used quickly, minimizing the likelihood that contaminating water will interfere with synthesis. Prepare reagents 1. Dry all powdered reagents in vacuo overnight prior to solubilizing. Although the authors have found that drying in vacuo is most necessary during humid weather, it is always a worthwhile precaution.
2. Warm amber bottles in which FMOC-PNA monomers were shipped to room temperature prior to solubilization to prevent water from condensing inside the container. 3. Solubilize FMOC-PNA monomers by adding 3.25 mL NMP diluent directly to each amber bottle (final 216 mM monomer). Allow the mixture to sit undisturbed for 30 min at room temperature.
Synthesis of Modified Oligonucleotides and Conjugates
4.11.3 Current Protocols in Nucleic Acid Chemistry
Supplement 8
Premature vortexing of the monomer at this point will cause partially dissolved clumps of monomer to stick to the side of the bottle. If this occurs, extra time should be taken to bring the clumps to the bottom of the bottle. Typically, monomers A, T, and G do not require vortexing, and a gentle swirling of the bottle’s contents will suffice. Solubilization of monomer C often requires some assistance. Intermittent heating of monomer C in a shallow water bath at 37°C can assist in solubilization, as can occasional vortexing or sonication. The authors have also diluted the C monomer in an additional 0.5 mL of NMP and have observed that this facilitates dissolving the reagent without decreasing synthesis yield. Experimenters should note that solubility can be dependent on the lot number of PNA monomer; some lots can be more or less difficult to dissolve than others. The volumes indicated for reagent solubilization (steps 3 to 5) are adequate for 37 cycles, but the volumes prepared must ultimately accommodate the specific sequences and coupling events required for a given synthesis. Each cycle requires 90 ìL monomer, 90 ìL activator, 90 ìL base, 1.2 mL deblocking solution, 2.39 mL DMF wash A, 36 ìL DMF wash B, and 300 ìL capping solution. These volumes are estimates. The investigator should always load significantly more reagent on the synthesizer to ensure that no reagent runs out during the final coupling steps.
4. Dissolve PNA activator (HOAt or HATU) in 13.5 mL anhydrous DMF (final 182 mM). Although HOAt is generally less expensive, it is also less efficient as a coupling reagent than HATU, which was developed specifically for PNA synthesis. HOAt should only be used for shorter sequences (≤12 couplings). HATU has been used for syntheses requiring >40 couplings.
5. Dissolve AEEA linker in 2.4 mL NMP diluent (final 209 mM). Perform PNA synthesis 6. Load the above reagents as well as base solution, amino acids, deblocking solution, and capping solution onto an Expedite 8909 synthesizer with an FMOC-XAL-PEGPS column. Monomers, activator, and linker should not remain on the machine for >2 weeks. Amino acids should not be left on the machine for >2 days due to their tendency to crystallize. Amino acids are the least expensive reagent and it is cost effective to replace these solutions frequently rather than risk failed syntheses or damage to the instrument.
7. Perform synthesis according to manufacturer’s programs and specifications, stopping before removal of the final FMOC group. It is critical that the final FMOC be left on the PNA until a decision has been made to cleave the PNA from the resin or add another group. If synthesis is complete and the PNA is to be cleaved from the resin, proceed to step 9.
8. Optional: Perform any desired manual additions (e.g., see Support Protocols 1 and 2). 9. Implement the final deblock option in the prime menu of the Expedite PNA software. Cycle the final deblock procedure one time to complete the double deblock procedure. Wash and prepare PNA-bound resin 10. Remove the column from the synthesizer and wash four times with 10 mL DMF, reversing the direction of flow through the column each time. Synthesis and Purification of Peptide Nucleic Acids
11. Wash the column four times with 10 mL isopropyl alcohol, reversing the direction of flow through the column each time to facilitate drying of the resin.
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12. Dry the resin in vacuo for a minimum of 30 min or by blowing filtered house air across the column for 3 to 5 min, reversing the ends frequently. Resin is sufficiently dry when the resin plug slides easily from end to end as the air input is reversed.
Cleave PNA from solid-phase resin 13. Transfer the dried, resin-bound, deprotected PNA to a 1.5-mL, 0.2-µm PTFE or regenerated cellulose spin column. 14. Add 250 µL cleavage cocktail and incubate 90 min at room temperature. Although cleavage of PNAs from the FMOC-XAL-PEG-PS resin occurs in 5 min, cleavage of the protecting groups on the PNA side-chains requires at least 90 min. Complete cleavage of the BHOC side chains can be verified by MALDI-TOF mass spectrometry. Incomplete cleavage will result in a mass that exceeds the expected value by increments of the BHOC mass (100.1). Also, in the authors’ experience, products that retain one or more protecting groups will have longer retention times by RP-HPLC (the protecting groups are hydrophobic). Uniform cleavage is dependent on the freshness of the TFA, while the m-cresol serves as a molecular scavenger. The quality of the TFA can be checked by observing the color of the solution. It should be clear to slightly yellow. If the color progresses to a more orangebrown color, it should not be used for cleavage, but may still be used in preparing RP-HPLC buffer A (see Basic Protocol 3). In addition to solution color, TFA should release a small amount of fume (smoke) when the container is opened. If this is not observed, it should not be used for any procedure involving PNAs. CAUTION: TFA is caustic and should be dispensed using only glass pipets or pipet tips containing a charcoal filter (Intermountain Scientific). The charcoal filter serves as a barrier that protects the pipettor seals from the TFA fumes.
15. Centrifuge 2 min at 1300 × g for a PTFE filter or 2 min at 8400 × g for a regenerated cellulose filter. 16. Repeat steps 14 and 15, but reduce cleavage time to 5 min. 17. Collect the cleavage filtrate, remove the filter unit, and precipitate the PNA by adding 1 mL cold (–20°C) diethyl ether. Invert the tube several times to ensure complete precipitation. If the synthesis has been successful, the precipitated PNA should be obvious.
18. Centrifuge precipitated PNA 2 min at 1300 × g. Discard the supernatant. 19. Wash the pellet three times with 1 mL diethyl ether, vortexing to suspend the pellet. 20. Centrifuge 2 min at 8400 × g to repack the pellet. Centrifugation speeds of more than 8400 × g should be avoided because tight packing of PNA makes it difficult to dissolve the pellet.
21. Remove as much of the supernatant as possible by aspiration and then air dry the pellet for 5 to 10 min in a chemical fume hood. 22. Hydrate the pellet with 200 µL sterile water (for a 2-µmol synthesis) and allow the tube to remain undisturbed for 10 to 15 min at 65°C. Only slight vortexing should be required to complete PNA solubilization.
23. Purify and analyze PNA (see Basic Protocol 3).
Synthesis of Modified Oligonucleotides and Conjugates
4.11.5 Current Protocols in Nucleic Acid Chemistry
Supplement 8
SUPPORT PROTOCOL 1
ADDING PEPTIDES TO PNAS The addition of peptide sequences to PNAs is a convenient method for obtaining conjugates in which the peptide domain enhances hybridization (Zhang et al., 2000) or cell uptake (Simmons et al., 1997) of the attached PNA . Peptides can be added to a PNA in a number of ways. If the peptide contains three or fewer different amino acids, it can be conveniently added immediately before or after automated synthesis using the three open ports on the Expedite 8909 synthesizer in addition to the four dedicated to PNA monomers. All amino acids should be double coupled since amino acid coupling is sometimes inefficient. Also, since amino acids are inexpensive relative to PNA monomers, generous use of amino acids during coupling is a cost-effective strategy for optimizing synthesis yields. If more than three different amino acids need to be added, it is often more convenient to contract a dedicated peptide synthesis facility to add the completed peptide. In this case, it is recommended that the first amino acid of the peptide be coupled to the newly synthesized PNA prior to shipping the resin. The authors have found that this procedure reduces the likelihood of the N terminus becoming blocked during shipping and storage. The synthesis of the PNA should be coordinated with the facility that will add the peptide, so that delays between syntheses are avoided. Delay between syntheses can result in the spontaneous loss of FMOC groups, exposing the N terminus to modification and preventing its extension. The facility adding the peptide should be instructed to double or triple couple the first amino acid that is added at their facility. Prior to shipping, the column should be washed with DMF and dried in vacuo overnight. The thiol group of cysteine provides a convenient reactive group for PNA modification. Additional care is required to adequately cleave PNAs or PNA conjugates containing sulfhydryl groups, as the authors and others (Goodwin et al., 1998) have noted that the sulfhydryl can be modified during deprotection and purification. To avoid modification of cysteine, the cleavage cocktail should be supplemented with 7.5 mg pure crystalline phenol (Fisher) and 250 µL ethanedithiol (Sigma-Aldrich) per 1 mL of cleavage cocktail. Cysteine-containing PNAs should be neutralized immediately after purification to avoid the formation of TFA adducts. Neutralization can be achieved by adding 0.5 mL of an aqueous solution of 0.1 M ammonium acetate to the RP-HPLC fraction collection tube prior to collecting the PNA fraction. Incomplete neutralization will result in a product that is 97 mass units higher than expected, corresponding to an adduct with TFA.
SUPPORT PROTOCOL 2
ADDITION OF BIOTIN Once a PNA is synthesized, fluorescent groups, biotin, or other labels can be added to the free N terminus prior to deprotection. The labeling of PNAs with biotin is described as an example. Additional Materials (also see Basic Protocol 1) Biotin (Sigma) 42°C water bath 1-mL syringe 1. Perform a normal automated PNA synthesis (see Basic Protocol 1, steps 1 to 8). 2. Dissolve 20 mg biotin, 6.1 mg HATU, and 1.4 mg HOAt in 800 µl DMF. Warm the solution to 42°C and vortex intermittently until completely dissolved.
Synthesis and Purification of Peptide Nucleic Acids
Biotin is not readily soluble and will require warming and vortexing. Due to its poor solubility, biotin solutions should never be put on the synthesizer, as it will clog the lines.
3. Add 200 µL PNA base solution and mix well. Allow components to activate for 5 min at 42°C.
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4. Remove N-terminal FMOC group by performing the final two deblock cycles to remove the N-terminal cap of the resin-bound PNA (see Basic Protocol 1, step 9). 5. Wash the column two times from each end with 10 mL DMF. 6. Draw up the biotin solution into a 1-mL syringe and place into one end of the column. 7. Place a second syringe into the other end of the column and push the solution back and forth for 30 min. 8. Repeat steps 5 to 7 with another 1-mL preparation of biotin solution. Addition of biotin is repeated because of the inherently poor coupling efficiency of biotin.
9. Proceed to the procedure for cleavage of PNAs from solid-phase resin (see Basic Protocol 1, steps 10 to 23). MANUAL SYNTHESIS OF PEPTIDE NUCLEIC ACIDS The manual synthesis of PNAs is illustrated in Figure 4.11.3. Manual synthesis of PNAs is advantageous because PNAs can be obtained in larger amounts (>2 µmol) than on the Expedite synthesizer. By changing the amount of resin used, one can prepare as much or as little PNA as needed for each experiment. Manual synthesis also avoids the need for a dedicated automated synthesizer. As with automated synthesis, it is important to keep reagents and materials as anhydrous as possible. However, the coupling reactions are more exposed to atmospheric water. The authors have found that syntheses that use tert-butyloxycarbonyl (BOC) monomers generally produce better yields than syntheses using FMOC monomers.
BASIC PROTOCOL 2
Manual PNA synthesis is often so efficient that a capping step can be dispensed with. It is sometimes advisable, however, to include a capping step after coupling of the monomer to simplify the HPLC purification. The individual experimenter will have to determine the necessity for capping. It is typically necessary for long syntheses or for syntheses that have failed in the past. Many different types of apparatus can be used for manual peptide synthesis, and it is likely that these can be adapted for PNA synthesis. The apparatus described here (Fig. 4.11.4; Norton et al., 1995) uses common laboratory glassware and offers robust performance. While the details of manual synthesis will vary with apparatus, the outline of the procedure and the precautions that need to be taken will remain the same. Because a single missed step can ruin a labor-intensive synthesis, the authors follow a detailed checklist for each step. The checklist provides a written record that all steps were performed. A sample spreadsheet detailing the amount of reagent needed for synthesis is shown in Figure 4.11.5. Materials Nitrogen source N,N-Dimethylformamide (DMF; OptiDry; Fisher) 4-Hydroxymethylphenylamidomethyl (PAM) resin protected with tert-butyloxycarbonyl (BOC; Applied Biosystems) Carrier resin: PAM resin capped with an acetyl group (see Support Protocol 3) BOC-PNA monomers (Fig. 4.11.3; Applied Biosystems): tert-butyloxycarbonyl-protected peptide nucleic acid monomers (A, C, G, and T), base protected with benzyloxycarbonyl 2-(1H-Benzotriazol-1-yl)-1,1,3,3-tetramethyl uronium hexafluorophosphate (HBTU) and 1-hydroxybenzotriazole (HOBt) activators (Applied Biosystems) Fresh dichloromethane (DCM; Fisher) m-Cresol Trifluoroacetic acid (TFA; Burdick Jackson)
Synthesis of Modified Oligonucleotides and Conjugates
4.11.7 Current Protocols in Nucleic Acid Chemistry
Supplement 9
coupling step base*
NHBOC
P
O
deprotection 5% m -cresol in TFA P
base*
N
O N
HO
NH2
NHBOC
O
HBTU/HOBT/DIPEA in DMF
N H
P
n=n+1
n
base* O
P
N H
O
O
N H
n
base*
coupling step
N
O
N H
P
NHBOC
O
N
NH2
O
1. deprotection 2. cleavage from the support TFA/m -cresol/thioanisole/TFMSA
when n = n max
base
n max
base O
O
N
HO
deprotection
base*
N
NHBOC
O
N
N H
O
NH2
O
PNA monomers NHCO2Bn
O
NH
N
N
N
N
O
O
N
O
O
NHBOC
HO
N
N
N
O
NHBOC
HO
NH
N
NHCO2Bn
O N
N
O
N
O
O N
HO
NHCO2Bn
O
N
NHBOC
HO
O
NHBOC
Figure 4.11.3 Manual PNA synthesis as described in Basic Protocol 2. Although a capping step can be added, it is often not required and is not shown here. Abbreviations: base*, N-protected nucleobase (see PNA monomers); BOC, tert-butyloxycarbonyl; Bn, benzyl; DIPEA, diisopropylethylamine; DMF, dimethylformamide; HBTU, 2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyl uronium hexafluorophospate; HOBt, 1hydroxybenzotriazole; P, PAM resin; TFA, trifluoroacetic acid; TFMSA, trifluoromethanesulfonic acid.
Pyridine Diisopropylethylamine (DIPEA) Methanol Thioanisole Trifluoromethanesulfonic acid (TFMSA; Aldrich) Diethyl ether, ice cold
Synthesis and Purification of Peptide Nucleic Acids
250°C oven 125-mL vacuum filtration side-arm flasks 24/40 rubber septa 15-mL medium (C) fritted Pyrex funnel Vacuum tubing 3-way valves 250-mL Wheaton bottles with caps
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fritted filter
side-arm flask
coupling flask
deprotection flask
3-way valve
vacuum vacuum release valve N2
Figure 4.11.4 Apparatus for manual synthesis. Reprinted from Norton et al. (1995) with permission from Elsevier Science.
C-Terminus 1 2 3 4 5 6 7 8 9 10 11 12 13 14 N-Terminus
Monomer terminal amide lysine A A C A G A T T G G G A T
Molecular Weight g/mol 17.00 128.00 275.32 275.32 251.30 275.32 291.32 275.32 266.28 266.28 291.32 291.32 291.32 275.32 266.28
PNA mass after each coupling g/mol – 145.00 420.32 695.64 946.94 1222.26 1513.58 1788.90 2055.18 2321.46 2612.78 2904.10 3195.42 3470.74 3737.02 Total Mass
A Monomer G Monomer C Monomer T Monomer HBTU HOBt DIPEA DMF
Amount of Resin Used Substitution Number for Resin Total Active Sites PNA Mass
10 mg 0.66 mmol/g 0.0066 mmol 3737.02 mg/mmol
Amount of PNA to be Synthesized
24.6 mg
Equivalents 5.0 5.0 5.0 5.0 4.5 5.0 10.0 –
Amount Per Coupling 17.41 mg 17.94 mg 16.62 mg 12.68 mg 11.26 mg 4.46 mg 8.54 µL Up to 1 mL
3737.02
Figure 4.11.5 Example of a spreadsheet for the manual synthesis of a 14-base PNA including lysine at the C terminus. The resin used produces a theoretical yield of 24.6 mg of PNA upon cleavage.
Synthesis of Modified Oligonucleotides and Conjugates
4.11.9 Current Protocols in Nucleic Acid Chemistry
Supplement 8
Lyophilizer 10-mL flask with a ground glass joint Desiccator Additional reagents and equipment for purification and analysis of PNAs (see Basic Protocol 3) NOTE: All powdered reagents such as monomers and activator should be unpacked on arrival and stored at –20°C in a sealed container containing Drierite desiccant. Monomers should be inspected upon arrival. Clumps of reagent may indicate that water has been introduced during shipping. DMF should have a low amine content to reduce the likelihood of side reactions. Dry reagents and supplies 1. Dry all monomers and coupling reagents overnight in vacuo before beginning synthesis. To prevent accumulation of moisture, BOC-PNA monomers that have been refrigerated should be warmed to room temperature before opening.
2. Dry all glassware, pipet tips, and 1.5-mL microcentrifuge tubes in a 250°C oven overnight prior to use. Flush pipet tips and microcentrifuge tubes with nitrogen to ensure they are dry prior to synthesis. Set up apparatus 3. Clear a work area in a chemical hood. 4. Fit rubber septa over the tops of two 125-mL side-arm flasks and make a single hole in the middle of each septum to allow a 15-mL fritted funnel to be inserted without too much force. 5. Assemble the manual synthesis apparatus as in Figure 4.11.4 using vacuum tubing and 3-way valves such that the vacuum and nitrogen bubbling can be easily manipulated on and off. 6. Ensure that the 15-mL fritted funnel is clean and unclogged. Pull 5 mL DMF through the funnel to make sure it drains quickly. Prepare reagents 7. In a 1.5-mL microcentrifuge tube, weigh out the appropriate amount of BOC-protected PAM resin and add enough carrier resin to give ∼50 mg total resin weight. The purpose of the carrier resin is to facilitate handling of the resin by allowing researchers to handle larger volumes. The amount of PNA to be synthesized is dependent on the amount of resin used and the number of active sites on the resin. The number of active sites is based on the substitution number (Fig. 4.11.5), which should be listed on the resin bottle when purchased. The total number of active sites is based only on the amount the BOC-protected (noncarrier) PAM resin, as the carrier resin has been capped and contains no active sites. PAM resin can also be purchased with the first amino acid already attached (Advanced Chemtech).
8. Load 1.5-mL microcentrifuge tubes with the proper amount of dry monomer for each coupling reaction. Use a 5-fold excess of monomer over the resin active sites. Synthesis and Purification of Peptide Nucleic Acids
It is easiest to set up a spreadsheet (Fig. 4.11.5) with quantities of monomers, HBTU, HOBt, and DIPEA used for each activation/coupling reaction, checking them off during synthesis of the PNA.
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9. For each coupling reaction, measure out a 5-fold excess of HOBt (relative to resin active sites), a 4.5-fold excess of HBTU, 10 eq DIPEA, and 1 mL DMF. Draw DMF from the bottle using a syringe under nitrogen or argon gas to keep it as anhydrous as possible. HOBt, HBTU, DIPEA, and DMF will be added to the monomer in each microcentrifuge tube just minutes before starting the coupling reaction (step 17). This method is the most efficient way of activating the monomers, and thus preventing deletions in the final product. Aldrich sells DMF in Sure/Seal bottles, which have a rubber opening on a crimped lid so that a syringe can be inserted without uncapping the bottle and exposing the contents to air. A nitrogen-filled balloon may also be inserted through the rubber opening to continually flush the bottle with nitrogen during the synthesis.
10. Prepare 200 mL fresh 50:50 (v/v) DMF/DCM for washing. Also prepare 100 mL of 5% (v/v) m-cresol/TFA and 100 mL pyridine in separate clean, dry 250-mL Wheaton bottles with caps. The pyridine is used to wash the resin with the base prior to coupling. CAUTION: TFA is caustic and should be dispensed using only glass pipets or pipet tips containing a charcoal filter (Intermountain Scientific). The charcoal filter serves as a barrier that protects the pipettor seals from the TFA fumes.
11. Assemble disposable glass pipets and bulbs, one for each reagent, and place near or on each reagent bottle. For convenience and to prevent contamination, the authors usually tape a test tube onto each bottle as a pipet holder.
12. Place the 15-mL fritted funnel into the deblocking flask. Wet the rubber septum and the funnel with a little methanol to help it slide in. 13. Add the dry resin to the 15-mL fritted funnel without solubilizing it in DMF. It is easier to get it all in this way.
Perform synthesis 14. Swell resin in 1 mL DMF for 1 hr with nitrogen bubbling. 15. Close and bleed vacuum line. 16. Add 1 mL of 5% m-cresol/TFA to begin deprotection. Stir 3 min with nitrogen bubbling. 17. While the BOC group is being removed, activate (esterify) the first monomer to be added by adding HOBt, HBTU, DMF, and DIPEA (from step 9) to the monomer tube (step 8). Vortex until everything has gone into solution. Typically, these solutions turn a tan color after vortexing. Failure to change color can sometimes be an indication that the activation step is not proceeding properly. As the synthesis progresses, liquid will accumulate in the flasks. It is important to watch the level of liquids in the coupling and deprotection flasks and empty them when the volumes approach the fritted funnel. The deprotection flask can be emptied during a coupling step and vice versa.
18. Vacuum off deblocking solution from the resin. 19. Wash resin two times with 1 mL of 50:50 DMF/DCM. 20. Close and bleed vacuum line.
Synthesis of Modified Oligonucleotides and Conjugates
4.11.11 Current Protocols in Nucleic Acid Chemistry
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21. Remove funnel from the deprotection flask and insert into the rubber septum over the coupling flask. 22. Add activated monomer to resin and mix 20 min, with nitrogen bubbling, to couple the monomer to the resin (or growing PNA). 23. Vacuum off solution. 24. Wash six times with 1 mL of 50:50 DMF/DCM. 25. Wash with 1 mL pyridine then with 1 mL dry DMF. If capping is necessary, add 1 mL of 1:1 (v/v) acetic anhydride/DMF to the resin and mix for 20 min. Wash well with DMF and proceed to the next step.
26. Vacuum off solution. 27. Close and bleed vacuum line. 28. Remove funnel from the coupling flask and insert into the septum over the deblocking flask. 29. Repeat steps 16 to 28 for remaining monomer additions. 30. Remove the final protecting group as in steps 15 and 16. 31. Vacuum off deblocking solution. 32. Wash six times with 1 mL of 50:50 DMF/DCM. 33. Wash six times with 1 mL methanol. 34. Lyophilize overnight. When the synthesis is complete, it is customary to wash the resin with methanol and to dry in vacuo overnight before the cleavage is carried out. If the synthesis takes >1 day, it will be necessary to store the resin. Stop the synthesis just prior to deblocking the PNA, leaving the protecting group on. Wash the resin several times with HPLC-grade methanol. Remove the funnel with the resin from the coupling flask, put a Kimwipe over the top, and secure it with a rubber band. Label the funnel and put it in a desiccator. Resume the synthesis the following day by starting with swelling of the resin in NMP and deblocking the PNA (steps 14 to 16).
Cleave PNA from resin 35. Connect a clean 125-mL vacuum flask to the synthesis apparatus in place of the coupling flask. 36. Attach a rubber septum over the top of the flask and insert the funnel that contains the resin. 37. Prepare 4 mL cleavage cocktail in a clean, dry 10-mL flask that has a ground glass joint. Mix 1 part m-cresol, 1 part thioanisole, and 6 parts TFA. Cap the flask, put on gloves and a shield, crack open a vial of TFMSA, and add 2 parts TFMSA. Typically, the cleavage cocktail turns brown at this point. CAUTION: TFMSA is corrosive and extremely destructive to mucous membranes, the upper respiratory tract, eyes, and skin. Avoid skin contact and inhalation. Always use suitable protection.
38. Using a pipettor, add 1 mL cleavage cocktail to the resin. Using a very small flow of nitrogen into the flask, allow solution to barely bubble for ∼1 hr. Synthesis and Purification of Peptide Nucleic Acids
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39. Turn off the nitrogen and apply a vacuum to pull the cleavage solution into the clean flask. Remove as much of the cleavage solution as possible before proceeding to the precipitation. Excess TFA will make it more difficult to precipitate the PNA. Excess TFA can be removed from the solution by blowing a steady stream of nitrogen into the flask until most of the solution is gone, or by applying a vacuum to the flask for several minutes.
40. With most of the cleavage solution gone, add ≥45 mL ice-cold diethyl ether to the flask and set it on ice. 41. Take a spatula and scrape the sides of the flask to loosen all of the PNA into solution. There should be a white precipitate in the solution.
42. Pour the PNA/diethyl ether solution into a 50-mL conical centrifuge tube, cap tightly, and centrifuge 3 min at 1040 × g (e.g., 2500 rpm in a Beckman S4180 swinging bucket rotor), 5°C, pelleting the PNA to the bottom of the tube. 43. Carefully decant or aspirate off the diethyl ether. 44. Wash pellet three times by adding 50 mL ice-cold diethyl ether to the tube, vortexing, centrifuging, and removing the supernatant. 45. After decanting the last time, place a Kimwipe over the tube containing the wellwashed pellet and place it in a desiccator. Attach the house vacuum and allow PNA to dry overnight. 46. Purify and analyze PNA (see Basic Protocol 3). The PNA can be stored at −20°C as a lyophilized product or as an aqueous stock solution (in deionized water). Traces of TFA will slightly acidify the solution and readily dissolve the PNA.
PREPARATION OF CARRIER RESIN FOR MANUAL PNA SYNTHESIS Carrier resin is prepared by deprotecting a simple BOC-protected PAM resin and performing a capping step using acetic anhydride.
SUPPORT PROTOCOL 3
Additional Materials (also see Basic Protocol 2) tert-Butyloxycarbonyl-protected 4-hydroxymethylphenylamidomethyl resin (e.g., BOC-Ala-PAM, BOC-Val-PAM, BOC-Ile-PAM; Applied Biosystems) Acetic anhydride HPLC-grade dichloromethane HPLC-grade methanol 1. Set up the manual synthesis apparatus (Fig. 4.11.4) with a single flask. 2. Place ∼2 g BOC-protected PAM resin in the fritted funnel and swell with 1 mL DMF for 1 hr. 3. Vacuum off DMF. 4. Add a large excess (e.g., 1 mL) of 5% (v/v) m-cresol/TFA and deprotect the resin for 10 min with bubbling. 5. Wash two times with 1 mL DMF and repeat deprotection two more times. The resin should be fully deprotected at this point.
6. Wash five times with 1 mL DMF and once with 1 mL pyridine. 7. Add 1 mL of 1:1: (v/v) acetic anhydride/DMF and bubble for 20 min. 8. Wash once with 1 mL DMF and repeat step 7.
Synthesis of Modified Oligonucleotides and Conjugates
4.11.13 Current Protocols in Nucleic Acid Chemistry
Supplement 8
9. Wash once each with 1 mL DMF, 1 mL dichloromethane, and 1 mL methanol. 10. Dry overnight in a desiccator. The resin can be prepared in bulk and stored at 4°C in the desiccator. BASIC PROTOCOL 3
PURIFICATION AND ANALYSIS OF PEPTIDE NUCLEIC ACIDS PNAs can be purified by reversed-phase HPLC (RP-HPLC) followed by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF-MS) (Fig. 4.11.6). PNAs are not, however, purified or analyzed by standard procedures used for oligonucleotides. PNA purification and analysis is more similar to that for peptides. Materials PNA sample solution (see Basic Protocols 1 and 2) RP-HPLC buffer A: 0.1% (v/v) trifluoroacetic acid (TFA; Burdick Jackson) in water, passed through a 47-mm, 0.4-µm nylon membrane (Whatman) RP-HPLC buffer B: 0.1% (v/v) TFA in acetonitrile (Optima grade; Fisher), filtered through an Anodisc 47 filter (0.22-µm; Whatman) α-Cyano-4-hydroxycinnamic acid (Sigma) Isopropanol High-performance liquid chromatograph (HPLC) with C18 reversed-phase column (300-Å Microsorb-MV column; Varian Analytical Instruments) Matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrometer (Voyager-DE workstation; Applied Biosystems) Lyophilizer UV spectrophotometer Additional reagents and equipment for HPLC and MALDI-TOF-MS
A 12.358
B m.w. 5144.58
%B 100
60 40 20
11.153
0
5
10.073
9.261 9.910
4.898
3.171
10.181 10.418
80
Retention time (min)
Synthesis and Purification of Peptide Nucleic Acids
Mass (amu)
Figure 4.11.6 Typical (A) HPLC and (B) MALDI-TOF mass spectrometry data.
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Purify by HPLC 1. Centrifuge a PNA solution 3 min at 12,000 × g, room temperature, to remove particulate. 2. Heat a C18 reversed-phase HPLC column to 55°C. PNAs tend to form internal structure or higher order aggregates; sharper peaks will often be obtained if the column is maintained at 55°C using a heated water jacket.
3. Set up a gradient of 0% to 5% (v/v) RP-HPLC buffer B in buffer A for 6 min followed by 5% to 100% buffer B in buffer A for 24 min. 4. Inject sample. 5. Collect fractions corresponding to major peaks. Analyze by MALDI-TOF-MS 6. Spot a 1-µL aliquot of each HPLC fraction on the laser target of a MALDI-TOF mass spectrometer. 7. Overlay the sample with 1 µL matrix consisting of 10 mg/mL α-cyano-4-hydroxycinnamic acid in 25:75 (v/v) RP-HPLC buffers A/B. 8. Activate the laser and collect data. Resolution is best when lower laser energies are used (∼1400 mV) but higher energies are sometimes necessary, especially with long PNAs and PNA-peptide conjugates. Analytical scale analysis by HPLC or mass spectral analysis of the crude product often reveals that only one product has been formed. If this is the case, the PNA can be conveniently purified by Sep-Pak Vac 6-mL (1 g) C18 cartridge (Waters Chromatography). Alternatively, PNAs can be purified by preparative C18 HPLC (Baker Bond, J.T. Baker). PNAs typically elute between 32% and 37% RP-HPLC buffer B and are routinely >80% full-length material.
Solubilize and store PNA 9. Pool collected fractions containing the PNA with the appropriate mass. Freeze in an isopropanol/dry ice bath and lyophilize. PNAs that are not to be used immediately can be stored indefinitely in a lyophilized form at –20°C in a sealed storage box containing desiccant.
10. Dissolve the resulting pellet in 200 µL sterile water and allow the tube to sit 5 to 10 min undisturbed at room temperature. PNAs are readily soluble in aqueous solutions at pH 5.0 to 6.0 at high concentrations (millimolar) but are less soluble at higher pH’s. Solubility at neutral pH can be enhanced by the incorporation of charged amino acid residues at the termini of PNAs during synthesis. Heating solutions containing PNAs can also enhance solubility, and the authors recommend always heating PNAs to >50°C immediately prior to use to ensure that aggregation is minimized. The authors have repeatedly frozen and thawed solubilized PNAs without observing diminished hybridization, although for long-term storage it is best to store PNAs in lyophilized form.
Analyze by UV spectrophotometry 11. Heat PNA stock solutions 5 min at 65°C to reduce aggregation prior to dilution and measurement. PNAs readily aggregate because they are relatively hydrophobic. Heating is a convenient way to ensure that the measured concentration reflects the total concentration of PNA in the sample. Repeated heating of PNA samples does not affect their activity.
12. Observe the optical density at 260 nm (typically a 1:500 dilution).
Synthesis of Modified Oligonucleotides and Conjugates
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13. Determine the concentration using the following equation: c (mM) = (A260 × 30 ng/µL × 500)/mol. wt. where 30 ng/µL is the extinction coefficient, 500 is the dilution factor, and mol. wt. is the molecular weight of the PNA. Alternatively, the concentration can be determined as a cumulative function of the extinction coefficients of the PNA monomers. While more time-consuming, the latter method is more accurate since it is based on the extinction coefficient for a specific PNA sequence. The authors have found that it yields concentrations that vary by as much as 50% to 60% from those derived from use of the standard conversion factor. To determine the concentration using extinction coefficients, add up the extinction coefficients for the PNA sequence [A =13.7, C = 6.06, G = 11.7, T= 8.6, mL/ìmol(cm)]. Calculate the OD260/mL of crude PNA and divide by the total extinction coefficient to obtain the millimolar concentration. Determination of melting temperature provides a useful functional analysis for PNAs. A clean melting curve ensures that the PNA is present in solution at the anticipated concentration and can hybridize to its target sequences. The Tm value also provides important information for optimizing annealing conditions and interpreting results.
COMMENTARY Background Information
Synthesis and Purification of Peptide Nucleic Acids
Peptide nucleic acids (PNAs; Nielsen et al., 1991) are DNA analogs that have a neutral amide backbone (Fig. 4.11.1) and possess physical properties that differ from those possessed by nucleic acids with traditional phosphodiester or phosphorothioate backbones. Although they hybridize with high affinity to DNA and RNA according to normal WatsonCrick base-pairing rules (Egholm et al., 1993), the neutral backbone eliminates the electrostatic repulsion that characterizes the hybridization of DNA and RNA strands. PNA hybridization to single-stranded DNA or RNA occurs with high affinity, and hybridization to duplex DNA is characterized by an outstanding potential for strand invasion (Smulevitch et al., 1996; Lohse et al., 1999; Nielsen, 2001). The absence of a negatively charged backbone also reduces the likelihood that PNAs will associate with cellular proteins (Hamilton et al., 1996) and generate misleading phenotypes. Another difference relative to DNA or RNA is that the strength of PNA hybridization is independent of salt concentration. Given the many nucleic acid derivatives available, why should researchers consider using PNAs? PNAs possess distinctive chemical properties that confer numerous favorable properties, including high-affinity binding, rapid rates of hybridization, efficient strand invasion, resistance to digestion by nucleases and proteases, and low propensity to bind to proteins. PNAs do not spontaneously enter cultured cells, but can be introduced through simple transfection protocols (Hamilton et al.,
1999; Herbert et al., 1999; Braasch and Corey, 2001; Doyle et al., 2001). PNA synthesis is efficient and versatile and PNAs can be readily purified by HPLC and characterized by mass spectral analysis. Most PNAs will be less soluble than DNA or RNA, but lower solubility can be overcome by adjusting the pH of stock solutions or by heating PNA solutions prior to use. Learning how to obtain and work with PNAs is not trivial, but the power of PNA recognition amply justifies the effort for many applications. Advantages and disadvantages of PNAs are summarized in Table 4.11.1. PNAs can also be purchased directly from Applied Biosystems. Other vendors throughout the world have been licensed to sell PNAs. Currently, international vendors include Nippon Flour Mill, Omgen, Sawady Technologies, and OSWEL-University of Southhampton. Applied Biosystems can supply information for the vendor most convenient to a particular laboratory. Obtaining PNAs from commercial sources will probably be a less expensive option for laboratories that require a limited number of PNAs on a 2-µmol scale, especially if the laboratories do not have experience making peptides.
Critical Parameters The solid-phase synthesis of PNAs uses protocols similar to those developed for peptide synthesis and the physical properties of PNAs are more similar to hydrophobic peptides than to DNA or RNA. PNA solubility at neutral pH is relatively low, and PNAs tend to aggregate upon storage. These properties can pose prob-
4.11.16 Supplement 8
Current Protocols in Nucleic Acid Chemistry
Table 4.11.1 Advantages and Disadvantages of Peptide Nucleic Acids
Advantages
Disadvantages
Synthesis by standard peptide synthesis protocols
Less soluble than DNA or RNA
Easy to derivatize at N or C terminus High-affinity hybridization
Tendency for some sequences to aggregate Requires carrier DNA and lipid for delivery into mammalian cells or attachment to import peptides
High rates of hybridization Exceptional potential for strand invasion Low potential for binding to proteins that bind negatively charged polymers
lems for researchers unfamiliar with PNAs, and methods for obtaining useful concentrations of soluble PNA will be discussed below. The authors have not found poor solubility to be a major barrier to experiments with PNAs since the solubility of PNAs is a pH-dependent property. PNAs are soluble to high concentrations at pH ≤5.0, and usually remain soluble at neutral pH when diluted to lower concentrations or when pH is slowly adjusted upward. A substantial advantage is that PNAs can be readily derivatized by peptides or other groups that can form a covalent linkage with the free amino terminus. For automated synthesis, proper maintenance of the instrument is essential. One common problem is clogged lines that can lead to reduced flow rate. Line-filters should be replaced monthly on all internal bottles to ensure that the flow of reagents is consistent for all syntheses. When bottles are removed or placed on the instrument, the open lip of each bottle should be wiped with a clean Kimwipe to prevent cross-contamination of bottles and reduce wear on the O-ring seals. The O-ring seals should be replaced every 3 months regardless of any appearance of physical stress.
Troubleshooting A PNA that has been correctly synthesized will usually give rise to predominantly one peak by HPLC purification. However, PNAs that possess a high likelihood for forming internal structure may give rise to multiple peaks. The potential for multiple peaks can be lessened by heating the HPLC column at 55°C. If mass spectral analysis shows that the major peaks share a single predominant product, multiple peaks may not be a cause for concern. The most
common problem with PNA synthesis is formation of truncated products. One cause of failed syntheses is use of reagents that are contaminated with water, leading to a reduction in coupling efficiency. PNA monomer and activator should be routinely dried in vacuo prior to synthesis. The authors also use fresh bottles of solvent, ≤14 days old. Alternatively, failed sequences could arise due to problems with automated synthesis or human error during manual synthesis. If truncated products are formed, the HPLC will show a series of peaks with shorter retention times than would be expected for full-length product. Another problem is failure to fully cleave protecting groups after completion of synthesis. Since the protecting groups are hydrophobic, a failure to cleave them results in products with longer retention times after RP-HPLC. PNAs that have this problem will appear to have at least two peaks upon HPLC analysis and can be salvaged by a second treatment with TFA and m-cresol. Retention of FMOC groups can also be confirmed by observation of a mass of 222 atomic mass units (amu) greater than expected. To avoid this problem, the authors use TFA within 1 month of first opening bottles. The solubility properties of PNA are discussed above. PNA oligomers have different solubility properties than analogous DNA oligonucleotides and will often be less soluble. Dissolving PNA stocks at pH ≤5.0 and heating solutions prior to use should alleviated this. The authors routinely check the concentration of PNA stocks by monitoring their absorbance at 260 nm to ensure that the expected concentration is being maintained.
Synthesis of Modified Oligonucleotides and Conjugates
4.11.17 Current Protocols in Nucleic Acid Chemistry
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Anticipated Results The authors’ laboratory has made several hundred PNAs. When the Expedite synthesizer is working properly and when all reagents are fresh and dry, an excellent yield of the desired product is almost invariably achieved. Crude material before purification contains ≥50% desired product, while HPLC-purified product contains >90%. PNAs and PNA-peptide conjugates that require as many as 40 coupling steps can be synthesized in good yield. It is possible to synthesize PNAs with long runs of purines or PNAs that are self-complementary as long as critical coupling steps are repeated. PNA synthesis is actually easier than peptide synthesis because the “personalities” of only four, rather than twenty, monomers are involved in determining coupling efficiency. With care, almost any PNA can be obtained.
Time Considerations Preparing the reagents for automated or manual PNA synthesis requires 1 to 2 hr. Automated PNA synthesis can be completed within hours, depending on the length of the PNA. Manual synthesis should take no more than 1 hr per cycle depending on whether a capping step is performed. Deprotection and purification can be accomplished in 1 day. HPLC analysis and purification should take 2 to 3 hr combined. MALDI-TOF-MS should require no more than 30 min.
Literature Cited Braasch, D.A. and Corey, D.R. 2001. Synthesis, analysis, purification, and intracellular delivery of peptide nucleic acids. Methods 23:97-107. Doyle, D.F., Braasch, D.A, Simmons, C.G., Janowski, B.A., and Corey, D.R. 2001. Intracellular delivery and inhibition of gene expression by peptide nucleic acids. Biochemistry 40:53-64.
Synthesis and Purification of Peptide Nucleic Acids
Hamilton, S.E., Simmons, C.G., Kathriya, I., and Corey, D.R. 1999. Cellular delivery of peptide nucleic acids and inhibition of human telomerase. Chem. Biol. 6:343-351. Herbert, B.-S., Pitts, A.E., Baker, S.I., Hamilton, S.E., Wright, W.E., Shay, J.W., and Corey, D.R. 1999. Inhibition of telomerase in immortal human cells leads to progressive telomere shortening and cell death. Proc. Nat. Acad. Sci. U.S.A. 96:14726-14281. Lohse, J., Dahl, O., and Nielsen, P.E. 1999. Doubleduplex invasion by peptide nucleic acid: A general principle for sequence-specific targeting of double-stranded DNA. Proc. Natl. Acad. Sci. U.S.A. 96:11804-11808. Mayfield, L.D. and Corey, D.R. 1999. Automated synthesis of peptide nucleic acids (PNAs) and peptide nucleic acid-peptide conjugates. Anal. Biochem. 268:401-404. Nielsen, P.E. 2001. Targeting double-stranded DNA with PNA. Curr. Med. Chem. 8:545-550. Nielsen, P.E., Egholm, M., Berg, R.H., and Buchardt, O. 1991. Sequence-selective recognition of double stranded DNA by a thymine-substituted polyamide. Science 254:1497-1500. Norton, J.C., Waggenspack, J.J., Varnum, E., and Corey, D.R. 1995. Targeting peptide nucleic acid protein conjugates to structural features within duplex DNA. Bioorg. Med. Chem. 3:437-445. Simmons, C.G., Pitts, A.E., Mayfield, L.D., Shay, J.W., and Corey, D.R. 1997. Synthesis and membrane permeability of PNA-peptide conjugates. Bioorg. Med. Chem. Lett. 7:3001-3007. Smulevitch, S.V., Simmons, C.G., Norton, J.C., Wise, T.W., and Corey, D.R. 1996. Enhanced strand invasion by oligonucleotides through manipulation of backbone charge. Nature Biotech. 14:1700-1704. Zhang, X., Ishihara, T., and Corey, D.R. 2000. Strand invasion by mixed base PNAs and PNApeptide chimera. Nucl. Acids Res. 28:3332-3338.
Internet Resources http://www.appliedbiosystems.com/ds/pna.taf
Egholm, M., Buchardt, O., Christensen, L., Behrens, C., Freier, S.M., Driver, D.A., Berg, R.H., Kim, S.K., Norden, B., and Nielsen, P.E. 1993. PNA hybridizes to complementary oligonucleotides obeying the Watson-Crick hydrogenbonding rules. Nature 365:566-568.
Ordering information and bibliography.
Goodwin, T.E., Holland, R.D., Lay, J.O., and Raney, K.D. 1998. A simple procedure for solid-phase synthesis of peptide nucleic acids with N-terminal cysteine. Bioorg. Med. Chem. Lett. 8:22312234.
PNA synthesis provider in The Netherlands.
Hamilton, S.E., Iyer., M., Norton, J.C., and Corey, D.R. 1996. Specific and nonspecific inhibition of RNA synthesis by DNA, PNA and phosphorothioate promoter analog duplexes. Bioorg. Med. Chem. Lett. 6:2897-2900.
http://www.horizonpress.com/gateway/pna.html Links to PNA-related sites. http://www.isogen.nl/pna.html
http://www.bostonprobes.com Supplier of PNA diagnostic probes.
Contributed by Dwaine A. Braasch, Christopher J. Nulf, and David R. Corey University of Texas Southwestern Medical Center at Dallas Dallas, Texas
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Current Protocols in Nucleic Acid Chemistry
Locked Nucleic Acids: Synthesis and Characterization of LNA-T Diol
UNIT 4.12
This unit describes the eleven-step convergent synthesis of the thymidine analog of locked nucleic acids (LNAs) starting from a commercially available sugar. The first five protocols describe the synthesis of a glycosyl donor suitable for synthesis of several LNA monomers; the overall procedure is illustrated in Figure 4.12.1. First, the starting sugar, 1,2:5,6-di-O-isopropylidene-α-D-allofuranose (S.1), is benzylated (S.2; see Basic Protocol 1 and Alternate Protocol). In the next set of procedures (see Basic Protocol 2), the 5,6-O-isopropylidene group is then selectively removed, the product (S.3) is treated with sodium periodate to give the aldehyde derivative (S.4), and a 4-C-hydroxymethyl group is introduced (S.5). The product is then mesylated (S.6; see Basic Protocol 3) and acetylated (see Basic Protocol 4), yielding the glycosyl donor, 1,2-di-O-acetyl-3-O-benzyl-5-O-methanesulfonyl-4-C-methanesulfonyloxymethyl-D-erythropentofuranose (S.7). Finally, the last protocol describes the synthesis of LNA-T diol from this sugar precursor (see Basic Protocol 5; Fig. 4.12.2). All procedures are experimentally simple and use readily available standard reagents. The synthesis of the other LNA monomers follows the same general pathway. Glycosylation reactions of adenine and guanine are known to give isomeric mixtures. Thus, chromatographic purification is to some extent necessary to obtain pure compounds. Furthermore, these nucleosides contain exocyclic amino groups that need protection. CAUTION: All the reactions should be carried out in a fume hood and contact with chemicals should be avoided. SYNTHESIS OF 3-O-BENZYL-1,2:5,6-DI-O-ISOPROPYLIDENE-α-D-ALLOFURANOSE USING BENZYL BROMIDE IN TETRAHYDROFURAN This protocol describes the benzylation of 1,2:5,6-di-O-isopropylidene-α-D-allofuranose (S.1) by treatment with sodium hydride and benzyl bromide resulting in 3-O-benzyl1,2:5,6-di-O-isopropylidene-α-D-allofuranose (S.2; Fig. 4.12.1). Purification is performed by a combination of extraction and crystallization.
BASIC PROTOCOL 1
CAUTION: Hydrogen gas, which can be explosive, is evolved in steps 5 and 9. Perform steps 2 through 9 under a nitrogen stream and use extreme care. Materials 60% (v/v) sodium hydride in mineral oil Hexane (stored over 3A molecular sieves) Nitrogen (N2) stream (or argon stream) Tetrahydrofuran (THF; stored over 3A molecular sieves) Dimethylformamide (DMF; stored over 3A molecular sieves) 1,2:5,6-Di-O-isopropylidene-α-D-allofuranose (Pfanstiehl Laboratories) Benzyl bromide Brine (saturated aqueous NaCl) MgSO4 500-mL three-neck round-bottom flask 250-mL dropping funnel Nitrogen inlet 20-mL syringe Sintered glass funnel, pore size 3 Contributed by Henrik M. Pfundheller and Christian Lomholt Current Protocols in Nucleic Acid Chemistry (2002) 4.12.1-4.12.16 Copyright © 2002 by John Wiley & Sons, Inc.
Synthesis of Modified Oligonucleotides and Conjugates
4.12.1 Supplement 8
Rotary evaporator connected to vacuum pump Generate 3-alkoxyde 1. Equip a 500-mL three-neck round-bottom flask with a 250-mL dropping funnel, nitrogen inlet, and a magnetic stir bar. 2. Place 11.2 g of 60% sodium hydride in the flask, add 30 mL hexane, and stir for a few minutes under a stream of nitrogen. 3. Stop stirring and allow sodium hydride to settle. Remove the excess hexane carefully with a 20-mL syringe and discard. 4. Add 25 mL THF and 5 mL DMF to the sodium hydride and cool the suspension in an ice bath. 5. Place 52 g (0.20 mol) of 1,2:5,6-di-O-isopropylidene-α-D-allofuranose dissolved in 100 mL THF in the 250-mL dropping funnel and add the solution dropwise over a 30-min period to the sodium hydride suspension. 6. Remove from the ice bath and stir for 1.5 hr at room temperature. Benzylate 3-alkoxyde 7. Over a 20-min period, add 25 mL benzyl bromide (in small portions) to the solution and then stir the mixture for 16 hr. 8. Cool the mixture in an ice bath for 15 min. 9. Destroy excess sodium hydride by careful addition of 100 mL water.
O O
O O O
O
1. NaH O
OH
HO HO O
80% AcOH
2. BnBr
O
O
OBn O
O
1
OBn O
2
3
O
HO
O
NalO4
H2CO, NaOH HO
O OBn O
MsO
Ac2O, AcOH, conc. H2SO4 O
OBn O
6
Locked Nucleic Acids: Synthesis and Characterization of LNA-T Diol
5
O
MsO
O OBn O
4
MsCl, Pyridine
O
MsO
O
OAc
MsO OBn OAc
7
Figure 4.12.1 Synthesis of the universal glycosyl donor. Bn, benzyl; Ms, methanesulfonyl.
4.12.2 Supplement 8
Current Protocols in Nucleic Acid Chemistry
Purify and isolate product 10. Separate phases and extract the aqueous phase (lower phase) with 75 mL THF. 11. Combine the two organic phases and wash two times with 100 mL brine. 12. Dry over MgSO4, remove the MgSO4 by vacuum filtration through a sintered glass funnel, and wash the solid with 50 mL THF. 13. Concentrate the filtrate in a rotary evaporator attached to a vacuum pump. 14. Dissolve the residue in 100 mL boiling hexane and place solution overnight at 5°C. 15. Isolate the formed crystals of 3-O-benzyl-1,2:5,6-di-O-isopropylidene-α-D-allofuranose (S.2) by filtration and dry under vacuum. Yield: 64 g (91%). An additional 4 g of product can be obtained from the mother liquor.
SYNTHESIS OF 3-O-BENZYL-1,2:5,6-DI-O-ISOPROPYLIDENE-α-D-ALLOFURANOSE USING BENZYL BROMIDE IN DIMETHYLFORMAMIDE
ALTERNATE PROTOCOL
This procedure describes the benzylation of 1,2:5,6-di-O-isopropylidene-α-D-allofuranose (S.1) using DMF as solvent. Using DMF instead of THF makes the reaction mixture less flammable, although DMF is toxic. Purification of the reaction mixture is obtained through precipitation of the product from the reaction mixture followed by crystallization. All reagents and materials are listed above (see Basic Protocol 1). CAUTION: Hydrogen gas, which can be explosive, is evolved in steps 5 and 9. Perform steps 2 through 9 under a nitrogen stream and use extreme care. Generate 3-alkoxyde 1. Equip a 500-mL three-neck round-bottom flask with a 250-mL dropping funnel, nitrogen inlet, and a magnetic stir bar. 2. Place 4.8 g of 60% sodium hydride in the flask, add 20 mL hexane, and stir for a few minutes under a stream of nitrogen. 3. Stop stirring and let the sodium hydride settle, then remove the excess hexane carefully with a 20-mL syringe and discard. 4. Add 20 mL DMF to the sodium hydride and cool the suspension in an ice bath. 5. Place 26 g (0.10 mol) of 1,2:5,6-di-O-isopropylidene-α-D-allofuranose dissolved in 30 mL DMF in the 250-mL dropping funnel and add solution dropwise over a 30-min period to the sodium hydride suspension. 6. Remove from the ice bath and stir for 1.5 hr at room temperature. Benzylate 3-alkoxyde 7. Over a 15-min period, add 14.3 mL benzyl bromide (in small portions) to the solution and then stir mixture for 1 hr at room temperature. 8. Cool the mixture in an ice bath. 9. Destroy excess sodium hydride by careful addition of 5 mL water. Purify and isolate product 10. Pour the reaction mixture into 175 mL ice water with stirring.
Synthesis of Modified Oligonucleotides and Conjugates
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11. When the ice has melted, filter off the precipitate by vacuum filtration through a sintered glass funnel, and wash the solid three times with 100 mL ice-cold water. 12. Dry the solid using a rotary evaporator connected to a vacuum pump. 13. Add 150 mL boiling hexane and keep overnight at 5°C to allow crystal formation. 14. Isolate the formed crystals of 3-O-benzyl-1,2:5,6-di-O-isopropylidene-α-D-allofuranose (S.2) by filtration and dry under vacuum. Yield: 31 g (89%). BASIC PROTOCOL 2
SYNTHESIS OF 3-O-BENZYL-4-C-HYDROXYMETHYL-1,2-O-ISOPROPYLIDENE-α-DERYTHROPENTOFURANOSE In this protocol, selective removal of the 5,6-O-isopropylidene group is performed by treatment of S.2 with 80% acetic acid (steps 1 to 5) to give 3-O-benzyl-1,2-O-isopropylidene-α-D-allofuranose (S.3). The formed 5,6-diol is then oxidatively cleaved with sodium periodate (steps 6 to 11) to give the corresponding 5-aldehyde derivative (S.4). Treatment of S.4 with formaldehyde and aqueous sodium hydroxide in a mixed aldol condensation/Cannizzaro reaction (steps 12 to 19) introduces a 4-C-hydroxymethyl group, giving 3-O-benzyl-4-C-hydroxymethyl-1,2-O-isopropylidene-α-D-erythropentofuranose (S.5). Materials 3-O-Benzyl-1,2:5,6-di-O-isopropylidene-α-D-allofuranose (S.2; see Basic Protocol 1 or Alternate Protocol) Acetic acid Dichloromethane Ethyl acetate Toluene Tetrahydrofuran (THF) NaIO4 Brine (saturated aqueous NaCl) 1,4-Dioxane 37% (w/v) aqueous formaldehyde 4 M NaOH MgSO4 Hexane 1-L round-bottom flask Rotary evaporator connected to vacuum pump Sintered glass funnel, pore size 3 Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) Deprotect 5,6-O-isopropylidene group 1. Place 63.1 g (0.18 mol) of 3-O-benzyl-1,2:5,6-di-O-isopropylidene-α-D-allofuranose (S.2) into a 1-L round-bottom flask containing a magnetic stir bar.
Locked Nucleic Acids: Synthesis and Characterization of LNA-T Diol
2. Add 70 mL water and 290 mL acetic acid and stir at room temperature until no starting material can be observed (∼25 to 35 hr) as monitored by TLC (APPENDIX 3D) using 9:1 (v/v) dichloromethane/ethyl acetate.
4.12.4 Supplement 8
Current Protocols in Nucleic Acid Chemistry
3. Remove the magnetic stir bar and concentrate the solution in a rotary evaporator connected to a vacuum pump. Avoid temperatures that are >45°C in steps 3 and 4, since this will result in removal of the 1,2-O-isopropylidene group.
4. Co-evaporate the residue (S.3) under vacuum two times with 100 mL toluene. Perform periodate oxidation of vicinal 5,6-diol 5. Add 100 mL THF, 100 mL water, and a large magnetic stir bar to the residue. 6. Over a 30-min period, add 43 g NaIO4 in small portions under vigorous stirring. 7. Stir the mixture for 3 hr. 8. Filter off the solid by vacuum filtration through a sintered glass funnel, and wash two times with 50 mL ethyl acetate. 9. Separate the phases and extract the aqueous phase (lower layer) two times with 50 mL ethyl acetate. 10. Combine the two organic phases, wash with 50 mL brine, and concentrate the organic phase (containing S.4) under vacuum in the rotary evaporator. Perform mixed aldol condensation/Cannizzaro reaction 11. Dissolve the residue in 120 mL of 1,4-dioxane. Add a magnetic stir bar and 40 mL aqueous 37% formaldehyde. 12. Over a 40-min period, add dropwise 90 mL of 4 M NaOH and then stir the mixture overnight. 13. Separate the phases and extract the aqueous phase (lower layer) one time with 100 mL ethyl acetate and then two times with 50 mL ethyl acetate. 14. Combine the two organic phases, wash with 100 mL brine, and extract the brine with 50 mL ethyl acetate. 15. Dry the organic phase over MgSO4 and remove the MgSO4 by filtration. 16. Wash the solid two times with 50 mL ethyl acetate and concentrate the combined filtrates in the rotary evaporator. 17. Dissolve the residue in 50 mL dichloromethane and add this solution to 350 mL hexane under vigorous stirring. 18. Isolate the precipitated 3-O-benzyl-4-C-hydroxymethyl-1,2-O-isopropylidene-α-Derythropentofuranose (S.5) by filtration and dry under vacuum. Yield: 45.7 g (82%) of a white solid.
Synthesis of Modified Oligonucleotides and Conjugates
4.12.5 Current Protocols in Nucleic Acid Chemistry
Supplement 8
BASIC PROTOCOL 3
SYNTHESIS OF 3-O-BENZYL-1,2-O-ISOPROPYLIDENE-5-O-METHANESULFONYL-4-CMETHANESULFONYLOXYMETHYL-α-D-ERYTHROPENTOFURANOSE This protocol describes the mesylation of the two hydroxyl groups in 3-O-benzyl-4-Chydroxymethyl-1,2-O-isopropylidene-α-D-erythropentofuranose (S.5) with methanesulonylchloride, giving 3-O-benzyl-1,2-O-isopropylidene-5-O-methanesulfonyl4-C-methanesulfonyloxymethyl-α-D-erythropentofuranose (S.6). The resulting mesyl groups are used in later steps for both protection and functionalization. Materials 3-O-Benzyl-4-C-hydroxymethyl-1,2-O-isopropylidene-α-D-erythropentofuranose (S.5; see Basic Protocol 2) Dichloromethane (stored over 3A molecular sieves) Pyridine (stored over 3A molecular sieves) Methanesulfonylchloride (e.g., Aldrich) Brine (saturated aqueous NaCl) 1 M HCl Saturated aqueous NaHCO3 MgSO4 Methanol 500-mL round-bottom flask 250-mL dropping funnel Guard tube/nitrogen inlet Sintered glass funnel, pore size 3 Rotary evaporator connected to vacuum pump Perform mesylation 1. Place 43.5 g (0.14 mol) of 3-O-benzyl-4-C-hydroxymethyl-1,2-O-isopropylidene-αD-erythropentofuranose (S.5) in a 500-mL round-bottom flask equipped with a magnetic stir bar, 250-mL dropping funnel, and a guard tube/nitrogen inlet. This reaction must be performed under a dry atmosphere.
2. Add 75 mL dichloromethane and 55 mL pyridine and cool mixture in an ice bath. 3. Place 24 mL (0.31 mol) methanesulfonylchloride in the dropping funnel and add dropwise over a 45-min period. 4. Remove from the ice bath and stir 3 hr at room temperature. 5. Add 100 mL water and 100 mL brine and stir 45 min at room temperature. Purify and isolate product 6. Separate the phases and extract the aqueous phase (top layer) three times with 75 mL dichloromethane. 7. Combine the two organic phases and wash three times with 150 mL of 1 M HCl, one time with 150 mL of saturated aqueous NaHCO3, and one time with 150 mL brine. 8. Dry the organic phase over MgSO4 and remove the MgSO4 by vacuum filtration through a sintered glass funnel. Locked Nucleic Acids: Synthesis and Characterization of LNA-T Diol
9. Wash the solid two times with 25 mL dichloromethane and concentrate the combined filtrates in a rotary evaporator with a vacuum pump.
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10. Dissolve the residue in 50 mL boiling methanol. Cool to room temperature and then let sit overnight at 5°C. 11. Isolate the formed crystals of 3-O-benzyl-1,2-O-isopropylidene-5-O-methanesulfonyl-4-C-methanesulfonyloxymethyl-α-D-erythropentofuranose (S.6) by filtration and dry under vacuum. Yield: 56.7 g (90%) of colorless crystals.
SYNTHESIS OF 1,2-DI-O-ACETYL-3-O-BENZYL-5-O-METHANESULFONYL-4-CMETHANESULFONYLOXYMETHYL-D-ERYTHROPENTOFURANOSE
BASIC PROTOCOL 4
This protocol describes the deprotection of the 1,2-O-isopropylidene group in 3-Obenzyl-1,2-O-isopropylidene-5-O-methanesulfonyl-4-C-methanesulfonyloxymethyl-αD-erythropentofuranose (S.6) followed by acetylation, giving the universal glycosyl donor 1,2-di-O-acetyl-3-O-benzyl-5-O-methanesulfonyl-4-C-methanesulfonyloxymethylD-erythropentofuranose (S.7) as a mixture of anomers. Deprotection and acetylation are performed in a one-pot procedure by treatment of S.6 with a mixture of acetic acid, acetic anhydride, and catalytic amounts of concentrated sulfuric acid. Materials 3-O-Benzyl-1,2-O-isopropylidene-5-O-methanesulfonyl-4-C-methanesulfonyloxymethyl-α-D-erythropentofuranose (S.6; see Basic Protocol 3) Acetic acid Acetic anhydride Concentrated H2SO4 Dichloromethane Saturated aqueous Na2CO3 Saturated aqueous NaHCO3 MgSO4, anhydrous 1-L round-bottom flask Guard tube/nitrogen inlet Sintered glass funnel, pore size 3 Rotary evaporator connected to vacuum pump Perform 1,2-O-isopropylidene acetolysis and acetylation 1. Place 46.6 g (0.10 mol) 3-O-benzyl-1,2-O-isopropylidene-5-O-methanesulfonyl-4C-methanesulfonyloxymethyl-α-D-erythropentofuranose (S.6) in a 1-L round-bottom flask equipped with a guard tube/nitrogen inlet and a magnetic stir bar. 2. Add 150 mL acetic acid and 30 mL acetic anhydride, and cool mixture in an ice bath. 3. Add 0.2 mL concentrated H2SO4 and let sit 5 min. 4. Remove from the ice bath and stir mixture for 18 hr at room temperature. Purify and isolate product 5. Add 150 mL dichloromethane to the reaction mixture followed by slow addition of 500 mL saturated aqueous Na2CO3. CAUTION: A large amount of CO2 gas is evolved in this step.
6. Stir the mixture for 3 hr at room temperature.
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7. Separate phases and extract the aqueous phase (top layer) two times with 150 mL dichloromethane. 8. Combine the two organic phases, add 300 mL saturated aqueous NaHCO3, and stir 2 hr at room temperature. 9. Separate phases and extract the aqueous phase (top layer) two times with 75 mL dichloromethane. 10. Dry the combined organic phases over MgSO4 and remove the MgSO4 by vacuum filtration through a sintered glass funnel. 11. Wash the solid two times with 25 mL dichloromethane. 12. Concentrate the combined filtrates to an oil in a rotary evaporator connected to a vacuum pump. The product, 1,2-di-O-acetyl-3-O-benzyl-5-O-methanesulfonyl-4-C-methanesulfonyloxymethyl-D-erythropentofuranose (S.7; mixture of anomers), is obtained as a yellow oil. Yield: 51.0 g (100%). BASIC PROTOCOL 5
SYNTHESIS OF (1S,3R,4R,7S)-7-HYDROXY-1-HYDROXYMETHYL-3-(THYMIN-1-YL)-2,5DIOXABICYCLO[2.2.1]HEPTANE (LNA-T DIOL) This protocol describes the synthesis of LNA-T diol from the universal glycosyl donor (S.7); the procedure is illustrated in Figure 4.12.2. The coupling of S.7 with silylated thymine (steps 1 to 9) gives 1-(2-O-acetyl-3-O-benzyl-5-O-methanesulfonyl-4-Cmethanesulfonyloxymethyl-β-D-erythropentofuranosyl)thymine (S.8). Basic hydrolysis of the 2′-O-acetyl group in S.8 (steps 10 to 16) liberates the free 2′-OH group, which instantly attacks the 4′-C-methanesulfonyloxy group, thereby causing ring formation between 2′-O and 4′-C. The resulting (1S,3R,4R,7S)-7-benzyloxy-1-methanesulfonyloxymethyl-3-(thymin-1-yl)-2,5-dioxabicyclo[2.2.1]heptane (S.9) constitutes the bicyclic LNA skeleton. Substitution of the methanesulfonyloxy group with benzoate (steps 17 to 22) and basic hydrolysis of the benzoate (steps 23 to 31) result in (1S,3R,4R,7S)-7-benzyloxy-1-hydroxymethyl-3-(thymin-1-yl)-2,5-dioxabicyclo[2.2.1]heptane (S.11) . Finally, reductive removal of the benzyl group in S.11 using Pd(OH)2/C-ammonium formate yields LNA-T diol (S.12; steps 32 to 38).
Locked Nucleic Acids: Synthesis and Characterization of LNA-T Diol
Materials 1,2-Di-O-acetyl-3-O-benzyl-5-O-methanesulfonyl-4-C-methanesulfonyloxymethylD-erythropentofuranose (S.7; see Basic Protocol 4) Acetonitrile (stored over 3A molecular sieves) Thymine N,O-Bis(trimethylsilyl)acetamide (Fluka) Trimethylsilyl trifluoromethanesulfonate (Fluka) Saturated aqueous NaHCO3 Dichloromethane Brine (saturated aqueous NaCl) Tetrahydrofuran (THF) LiOH⋅H2O Acetic acid Ethyl acetate MgSO4, anhydrous Anhydrous dimethylformamide (DMF; stored over 3A molecular sieves) Sodium benzoate
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Hexane Methanol 20% Pd(OH)2/C (palladium hydroxide catalyst on carbon; Fluka) Ammonium formate 2-cm-thick Celite pad 250-mL, 500-mL, and 1-L round-bottom flasks Rotary evaporator connected to vacuum pump Condensers Guard tubes/nitrogen inlets Sintered glass funnel, pore size 3 Glycosylate thymine 1. Place 25.5 g (50 mmol) of 1,2-di-O-acetyl-3-O-benzyl-5-O-methanesulfonyl-4-Cmethanesulfonyloxymethyl-D-erythropentofuranose (S.7) in a 500-mL round-bottom flask. Add 100 mL acetonitrile and concentrate the mixture using a rotary evaporator connected to a vacuum pump. It has been observed that skipping this evaporation step, which is performed to remove traces of water, can lead to lower yields from the reaction.
O
NH MsO
1. Thymine/BSA
O
MsO
OAc 2. TMS-OTf
MsO OBn OAc
N
O
O
MsO OBn OAc
7
8 O
O
NH LiOH MsO
N
NH
O
BzONa BzO
O
OBn O
N
OBn O
10
9 O
O
NH LiOH HO
N O
OBn O
11
O
O
NH
O
Pd(OH)2 /C NH4HCO2
HO
N
O
O
OH
O
12
Figure 4.12.2 Synthesis of LNA-T diol. Bn, benzyl; BSA, bis(trimethylsilyl)acetamide; Bz, benzoyl; Ms, methanesulfonyl; TMS-OTf, trimethylsilyl trifluoromethanesulfonate.
Synthesis of Modified Oligonucleotides and Conjugates
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2. Dissolve in 100 mL acetonitrile, and add 7.0 g (55 mmol) thymine, 24.5 mL (10 mmol) N,O-bis(trimethylsilyl)acetamide, and a magnetic stir bar. 3. Equip the 500-mL round-bottom flask with a condenser and a guard tube/nitrogen inlet, and heat the mixture to reflux (∼90°C) for 45 min. 4. Cool the reaction mixture to room temperature and add 10 mL trimethylsilyl trifluoromethanesulfonate. 5. Heat the mixture to reflux (∼90°C) overnight. 6. Cool the mixture to room temperature and pour it into 200 mL saturated aqueous NaHCO3. Stir the mixture for 5 min. 7. Extract three times with 200 mL dichloromethane. 8. Combine the organic phases and wash with 200 mL saturated aqueous NaHCO3 and then 100 mL brine. 9. Concentrate the organic phase (containing S.8) in a 1-L round-bottom flask in the rotary evaporator. Form 2′-O, 4′-C ring 10. Dissolve the residue in 100 mL THF and add 200 mL water and 10 g LiOH⋅H2O. Stir the mixture for 1.5 hr at room temperature. 11. Neutralize to pH ∼8 with acetic acid (∼10 mL). 12. Extract the mixture three times with 200 mL ethyl acetate. 13. Combine the organic phases and wash with 100 mL brine. 14. Dry the organic phase over MgSO4 and remove the MgSO4 by vacuum filtration through a sintered glass funnel. 15. Wash the solid two times with 25 mL ethyl acetate and concentrate in the rotary evaporator. 16. Dissolve the residue (S.9) in 200 mL acetonitrile and concentrate two times in a 500-mL round-bottom flask. Substitute methanesulfonyloxy group with benzoyloxy group 17. Dissolve the residue in 250 mL anhydrous DMF and add 20 g sodium benzoate. 18. Equip the flask with a guard tube/nitrogen inlet and heat the mixture for 16 hr at 90°C. The reaction results in a gummy mixture. If effective stirring cannot be achieved, an additional 250 mL anhydrous DMF should be added.
19. Cool the reaction mixture to room temperature. 20. Add water until the formed precipitate dissolves. 21. Pour the reaction mixture into 800 mL of stirring ice water (1600 mL of ice water if 500 mL of DMF has been used as solvent). 22. Isolate the precipitated product (S.10) by filtration and wash the product two times with 100 mL water. Locked Nucleic Acids: Synthesis and Characterization of LNA-T Diol
4.12.10 Supplement 8
Current Protocols in Nucleic Acid Chemistry
Hydrolyze benzoyl ester 23. Suspend the solid in 200 mL THF and 300 mL water in a 1-L round-bottom flask containing a magnetic stir bar. 24. Add 10 g LiOH⋅H2O and stir the mixture for 5 hr at room temperature. 25. Neutralize to pH ∼8 with acetic acid (∼10 mL) and concentrate to half volume in the rotary evaporator. 26. Extract three times with 200 mL dichloromethane 27. Combine the organic phases and wash with 200 mL saturated aqueous NaHCO3 followed by 200 mL brine. 28. Dry over MgSO4 and remove MgSO4 by filtration. 29. Wash the solid two times with 25 mL dichloromethane and concentrate the combined filtrates in the rotary evaporator. 30. Dissolve the residue in a minimum of ethyl acetate (∼50 mL) and pour this solution slowly into 600 mL hexane under vigorous stirring. 31. Collect the precipitated product (S.11) by filtration. Yield: 14.2 g (79%) of a white solid.
Remove 3′-O-benzyl group 32. Dissolve 14.2 g of S.11 in 100 mL methanol in a 250-mL round-bottom flask equipped with magnetic stir bar and condenser. 33. Add 1.5 g of 20% Pd(OH)2/C and 8.0 g ammonium formate. 34. Heat the mixture for 1 hr at 60°C. 35. Vacuum filter the hot solution through a 2-cm-thick Celite pad. 36. Wash the Celite with 200 mL methanol. Combine the filtrates and concentrate in the rotary evaporator. 37. Dissolve the residue in 50 mL boiling methanol and add 200 mL ethyl acetate followed by 600 mL hexane. 38. Isolate the precipitated product (1S,3R,4R,7S)-7-hydroxy-1-hydroxymethyl-3(thymin-1-yl)-2,5-dioxabicyclo[2.2.1]heptane (LNA-T diol; S.12) by filtration and dry under vacuum. Yield: 10.2 g (95%) as white precipitate.
COMMENTARY Background Information LNA oligonucleotides were introduced several years ago and were shown to have very high binding affinity and selectivity towards complementary nucleic acids (Koshkin et al., 1998b; Obika et al., 1998; Singh et al., 1998). These interesting properties have been evaluated in different diagnostic and therapeutic settings. LNA oligonucleotides have been used in diagnostic genotyping for the detection of the prothrombic mutations factor V Leiden (Ørum
et al., 1999) and have also been successfully evaluated in living rats for control of gene expression (Wahlestedt et al., 2000). Today, much effort is being put into the development of simple and reliable microarrays for multiplex genotyping in combination with multiplex target amplification using LNA oligonucleotides (Choleva et al., 2001). Furthermore, LNA oligonucleotides are successfully being investigated with respect to recognition of double-stranded DNA, as shown by inhibition of
Synthesis of Modified Oligonucleotides and Conjugates
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the NF-κB transcription factor p50 (Obika et al., 2001). The key steps in the synthesis of LNA nucleosides are the introduction of the 4-C-hydroxymethyl functionality followed by selective derivatization of the resulting two diastereoto pic h yd ro xy methy l groups. 4′-C-Hydroxymethyl nucleosides have been synthesized using both linear (Youssefyeh et al., 1977; Jones et al., 1979) and convergent (Leland and Kotick, 1974; Youssefyeh et al., 1977, 1979) strategies based on aldol condensation of 5(′)-aldehydes with formaldehyde followed by a Cannizzaro reaction or sodium borohydride reduction. LNA nucleosides were previously synthesized following linear (Obika et al., 1997; Koshkin et al., 1998a) as well as convergent (Singh et al., 1998; Koshkin et al., 1998a,b) strategies. Due to the reported problems (i.e., low yields, workup difficulties) for th e p reparation of 5′-aldehydo/4′-C-hydroxymethyl nucleosides from natural ribonucleosides, the authors decided to focus on the convergent strategy for the large-scale synthesis of LNA nucleosides further stimulated by the possibilities of introducing a variety of different nucleobases. This unit describes the authors’ efforts to develop a simplified and efficient synthesis of a sugar intermediate (S.7; Figure 4.12.1; see Basic Protocols 1 to 4), which can be used as a glycosyl donor in the coupling reactions with different nucleobases. This is exemplified by the synthesis of LNA-T diol (S.12; Figure 4.12.2; see Basic Protocol 5). These protocols give high yields of the desired products (>79% for each step), are experimentally simple, and avoid the use of time-consuming column chromatography. The synthesis of the adenine, cytosine, and guanine derivatives of LNA follow the same strategy and are described elsewhere (Koshkin et al., 2001).
Locked Nucleic Acids: Synthesis and Characterization of LNA-T Diol
Synthesis of the universal glycosyl donor S.7 Standard benzylation (see Basic Protocol 1) of the commercially available 1,2:5,6-di-O-isopropylidene-α-D-allofuranose with THF as solvent results in the corresponding 3-O-benzylated derivative S.2 (Horton and Tindall, 1970) in almost quantitative yield after crystallization from hexanes. In order to simplify the workup, the mineral oil from the sodium hydride suspension is initially removed with a syringe after addition of hexanes. As an alternative to THF, DMF can be used as the solvent (see Alternate Protocol) as reported earlier (Brimacombe and Ching, 1968). The starting sugar is commer-
cially available but can also be synthesized from the gluco-epimer via oxidation and selective reduction (Sowa and Thomas, 1966). Using 80% acetic acid, the 5,6-O-isopropylidene protecting group is selectively removed (see Basic Protocol 2) to give S.3 as reported (Horton and Tindall, 1970). It is important to keep the reaction at a low temperature to avoid removal of the 1,2-O-isopropylidene protecting group. After evaporation of the reaction mixture, the 5,6-glycol is oxidatively cleaved by periodate, giving the 5-aldehydo derivative after a simple work-up procedure. Finally, the aldehyde is condensed with formaldehyde followed by in situ crossed Cannizzaro reaction with excess formaldehyde, giving the desired 4-C-hydroxymethyl derivative S.5 (Youssefyeh et al., 1977, 1979) in 82% yield (3 steps) after crystallization from hexanes. In the first publications describing the synthesis of LNA nucleosides using the convergent strategy (Koshkin et al., 1998b; Singh et al., 1998), 3-O-benzyl-4-C-hydroxymethyl-1,2O-isopropylidene-α-D-ribofuranose S.5 was regioselectively 5-O-benzylated (Waga et al., 1993) in 71% yield, taking advantage of the different positioning of the two diastereotopic hydroxymethyl groups on each face of the bicyclo[3.3.0]octane system. However, chromatographic purification of the reaction mixture was necessary to separate the two isomers. Acetylation (Koshkin et al., 1998b; Singh et al., 1998) or tosylation (Koshkin et al., 1998a) of the 4-C-hydroxymethyl group, acetolysis, and subsequent 1,2-di-O-acetylation was followed by nucleobase-coupling and ring-closing reactions to give the desired LNA nucleoside derivative. Based on these results, the authors decided to synthesize 1,2-di-O-acetyl-3-Obenzyl-4-C-methanesulfonoxymethyl-5-Omethanesulfonyl-D-erythropentofuranose S.7 to overcome the early chromatographic step and increase the overall yield of the synthesis. The diol S.5 is permesylated using standard conditions (see Basic Protocol 3) to give the desired compound in 90% yield after crystallization from methanol. Subsequent standard acetolysis and basic diacetylation (see Basic Protocol 4) quantitatively yield S.7 as an anomeric mixture (∼1:5) that can be used as a glycosyl donor in coupling reactions with different nucleobases. The overall yield from S.1 is 67%. Synthesis of LNA-T diol S.12 Following the method of Vorbrüggen (Vorbrüggen et al., 1981; Vorbrüggen and Höfle,
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Table 4.12.1
Selected Rf Values and Melting Points
Compound number
Rf (90:10 CH2Cl2/ethyl acetate)a
1 2 3 4 5 6 7 8 9 10 11 12
0.30 0.70
0.55 0.40 + 0.35 0.05
Rf (95:5 CH2Cl2/methanol)a 0.78 0.14 0.40 0.16 0.70 0.60 + 0.55 0.25 0.22 0.45 0.13 0.05
Melting point (°C)b
65c
99d 109
169 196e
aSolvent ratios are given in v/v. TLC plates: Merck silica gel 60 F . 254 bUncorrected, measured on a Büchi melting point B-540. c64°C to 65°C (Brimacombe, 1968). d101°C to 102°C (Youssefyeh, 1979). e204°C to 205°C (Obika, 2001).
1981), S.7 is stereoselectively coupled (see Basic Protocol 5) with silylated thymine followed by a one-pot deacetylation and intramolecular ring-closing reaction upon treatment with LiOH, giving the LNA derivative S.9. This two-step one-pot procedure is fast and smooth compared to the two-step procedure originally described (Koshkin et al., 1998a,b). The methanesulfonyloxy group is substituted with benzoyl upon reaction of S.9 with sodium benzoate in hot DMF (Codington et al., 1960), followed by hydrolysis of the benzoate ester using LiOH to give S.11 in 79% yield (four steps) after crystallization from hexanes. Direct basic hydrolysis of the 5′-O-methanesulfonyl group was unsuccessful (Koshkin et al., 2001). Debenzylation with Pd(OH)2/C and ammonium formate as the H donor is fast and efficient, giving the desired LNA-T diol S.12 in 95% yield (50% from S.1) after a simple workup procedure and crystallization from ethyl acetate/hexanes. The corresponding phosphoramidite for use in automated oligonucleotide synthesis (Caruthers, 1991) can be synthesized via 5′-O-dimethoxytritylation and 3′-O-phosphitylation (Sinha et al., 1983; Koshkin et al., 1998b).
Compound Characterization All isolated compounds appear as one spot on a TLC plate except S.7, which exists as a mixture of anomers (∼1:5). Rf values and ap-
propriate solvent systems are given in Table 4.12.1. For bicyclic LNA structures, 1H-NMR shows three singlet signals for H1′, H2′, and H3′. This can be taken as verification of the formation of the rigid LNA bicyclic structure, as small coupling constants are indicative of H1′-C1′-C2′-H2′ dihedral angles close to 90°, which are characteristic of the N conformation (Altona and Sundaralingam, 1973; Obika et al., 1997; Koshkin et al., 1998b). Data for selected compounds S.2: 1H-NMR (CDCl3) δ 7.39-7.26 (m, 5H), 5.75 (d, J = 3.6 Hz, 1H), 4.78 (d, J = 12.2 Hz, 2H), 4.61-4.56 (m, 2H), 4.36 (m, 1H), 4.164.12 (m, 1H), 4.01-3.95 (m, 2H), 3.91-2.85 (m, 1H), 1.60 (s, 3H), 1.40 + 1.38 + 1.36 (3 × s, 9H). 13C-NMR (CDCl3) δ 137.5, 128.5, 128.3, 128.0, 112.9, 109.7, 103.9, 78.0, 77.8, 77.4, 74.7, 72.1, 64.9, 26.7, 26.4, 26.0, 25.0. MALDI-MS m/z: 351.2 [M+H]+. S.5: 1H-NMR (CDCl3) δ 7.37-7.26 (m, 5H), 5.76 (d, J = 3.9 Hz, 1H), 4.80 (d, J = 11.7 Hz, 1H), 4.63 (t, J = 4.0 Hz, 1H), 4.56 (d, J = 11.7 Hz, 1H), 4.20 (d, J = 5.2 Hz, 1H), 3.93 (d, J = 12.0 Hz, 1H), 3.85 (d, J = 12.0 Hz, 1H), 3.78 (d, J = 11.9 Hz, 1H), 3.54 (d, J = 12.0 Hz, 1H), 2.50 (br, 1H), 2.32 (br, 1H), 1.63 (s, 3H), 1.33 (s, 3H). 13C-NMR (CDCl3) δ 137.2, 128.4, 128.1, 127.7, 113.4, 104.3, 86.3, 78.3, 78.2,
Synthesis of Modified Oligonucleotides and Conjugates
4.12.13 Current Protocols in Nucleic Acid Chemistry
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72.6, 64.0, 63.0, 26.5, 25.8. MALDI-MS m/z: 311.1 [M+H]+. S.6: 1H-NMR (CDCl3) δ 7.38-7.26 (m, 5H), 5.73 (d, J = 3.8 Hz, 1H), 4.85 (d, J = 12.0 Hz, 1H), 4.75 (d, J = 11.6 Hz, 1H), 4.64 (t, J = 3.8 Hz, 1H), 4.55 (d, J = 11.6 Hz, 1H), 4.39 (d, J = 12.0 Hz, 1H), 4.30 (d, J = 11.0 Hz, 1H), 4.17 (d, J = 5.2 Hz, 1H), 4.12 (d, J = 11.0 Hz, 1H), 3.05 (s, 3H), 2.95 (s, 3H), 1.66 (s, 3H), 1.32 (s, 3H). 13C-NMR (CDCl3) δ 136.5, 128.4, 128.2, 127.9, 113.8, 104.3, 83.0, 78.2, 77.6, 72.6, 69.3, 68.5, 37.8, 37.2, 26.0, 25.4. MALDI-MS m/z: 467.1 [M+H]+. S.7 (main isomer): 1H-NMR (CDCl3) δ 7.37-7.28 (m, 5H), 6.17 (s, 1H), 5.37 (d, J = 4.8 Hz, 1H), 4.62 (d, J = 11.1 Hz, 1H), 4.52 (d, J = 11.1 Hz, 1H), 4.50 (d, J = 11.6 Hz, 1H), 4.42 (d, J = 4.8 Hz, 1H), 4.37 (d, J = 11.6 Hz, 1H), 4.30 (d, J = 10.6 Hz, 1H), 4.19 (d, J = 10.6 Hz, 1H), 3.01 (s, 6H), 2.14 (s, 3H), 2.10 (s, 3H). 13C-NMR (CDCl ) δ 169.1, 168.7, 136.3, 3 128.5, 128.3, 128.1, 97.2, 82.7, 78.6, 73.9, 73.1, 68.8, 68.4, 37.5, 37.3, 20.9, 20.5. MALDI-MS m/z: 511.1 [M+H]+. S.11: 1H-NMR (CDCl3) δ 9.28 (br, 1H), 7.45(d, J = 1.1 Hz, 1H), 7.38-7.22 (m, 5H), 5.66 (s, 1H), 4.67 (d, J = 11.6 Hz, 1H), 4.56 (d, J = 11.7 Hz, 1H), 4.54 (s, 1H), 4.05 (d, J = 7.9 Hz, 1H), 4.01 (d, J = 12.5 Hz, 1H), 3.96 (s, 1H), 3.95 (d, J = 12.6 Hz, 1H), 3.83 (d, J = 7.9 Hz, 1H), 1.88 (s, 3H). 13C-NMR (CDCl3) δ 163.9, 149.8, 137.0, 134.7, 128.5, 128.2, 127.8, 110.3, 88.2, 87.3, 76.9, 75.9, 72.3, 72.0, 57.6, 12.7. MALDI-MS m/z: 360.1 [M+H]+. S.12: 1H-NMR (DMSO-d6) δ 11.33 (br, 1H), 7.60 (d, J = 1.1 Hz, 1H), 5.68 (d, J = 4.1 Hz, 1H), 5.38 (s, 1H), 5.20 (br t, J = 5.6 Hz, 1H), 4.09 (s, 1H), 3.89 (d, J = 4.0 Hz, 1H), 3.80 (s, J = 7.8 Hz, 1H), 3.74 (d, J = 5.5 Hz, 2H), 3.61 (d, J = 7.8 Hz, 1H), 1.75 (d, J = 1.1 Hz, 3H). 13C-NMR (DMSO-d6) δ 164.1, 150.1, 135.1, 108.6, 89.0, 86.5, 79.1, 71.2, 68.9, 56.2, 12.6. Anal. calcd. for C11H14N2O6⋅2⁄3H2O: C 46.81, H 5.48, N 9.92; found: C 46.64, H 5.22, N 10.05. MALDI-MS m/z: 270.9 [M+H]+.
Critical Parameters and Troubleshooting
Locked Nucleic Acids: Synthesis and Characterization of LNA-T Diol
In general, the presented synthesis of LNAT has been developed to be tolerant to small changes in reaction conditions. Nonetheless, in some steps it is necessary to take precautions to ensure anhydrous conditions. When necessary, it was found sufficient to use solvents that were stored over molecular sieves. It is advisable to check all reactions by TLC (APPENDIX 3D; see Table 4.12.1 for Rf values)
before workup to ensure that complete conversion of starting material has been obtained. If the reaction has not gone to completion, a prolonged reaction time and/or additional reagent (typically 10% excess) will typically bring the reaction to completion. In Basic Protocol 1, the development of hydrogen gas demands a steady flow of inert atmosphere through the reaction flask. If such precaution is not taken, build up of hydrogen gas could potentially result in explosion of the reaction mixture. In Basic Protocol 2, steps 3 and 4, it is important to avoid temperatures higher than 45°C during the evaporation of acetic acid. If the temperature gets too high, the 1,2-O-isopropylidene group will be lost. A small amount of product that has lost the 1,2-O-isopropylidene group is acceptable in the following steps, but will lower the overall yield of the reaction. In steps 5 to 7, all the starting 5,6-diol S.4 must be consumed since unreacted starting material will be unaffected in subsequent reactions and is difficult to separate from the product during isolation of S.6. In Basic Protocols 3 and 4, the use of anhydrous reaction conditions is necessary to obtain a complete reaction. The use of anhydrous reaction conditions is essential during the glycosylation of S.7 with silylated nucleobases (see Basic Protocol 5, steps 1 to 5), since water will h yd ro lyze th e tr imethy lsilyl trifluoromethanesulfonate. It is also crucial to have anhydrous conditions during the substitution of the methanesulfonyloxy group with sodium benzoate (Basic Protocol 5, steps 17 to 18). It has been observed that even small amounts of water in the reaction mixture can not only slow down the reaction, but result in hydrolysis of the formed benzoate giving S.11, which will not be recovered during the precipitation step in the work-up procedure. If hydrolysis has occurred during the reaction, it is possible to extract S.11 from the aqueous phase by extraction with ethyl acetate. In Basic Protocol 5, step 34, it is advisable to keep the reaction at a temperature just below the boiling point. It has been observed that when the reaction mixture reaches the boiling point, large amounts of salt precipitate in the condenser.
Anticipated Results The overall yield for synthesis of LNA-T diol is 50% (eleven steps). Expected yields for isolated intermediates are S.2, 90%; S.5, 82%; S.6, 90%; S.7, 100%; S.11, 79%; and S.12,
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95%. The scale of the synthesis can be increased to 1 mol for Basic Protocols 1 to 3 without any major problems. The synthesis was developed to avoid chromatographic purification. The yields can be increased slightly by purifying the mother liquors (e.g., by column chromatography); however, this is time consuming.
Time Considerations The time needed to complete the synthesis of LNA-T diol is estimated to be 10 to 14 days. Most of the reactions are not affected by a prolonged reaction time. Thus, one can leave the reaction mixtures overnight at room temperature. An exception is the removal of the 5,6-O-isopropylidene group from S.2 (see Basic Protocol 2, step 2), where extended exposure to acetic acid will result in loss of the 1,2-O-isopropylidene group. All isolated compounds are stable and can be stored without special precautions.
Literature Cited Altona, C. and Sundaralingam, M. 1973. Conformational analysis of the sugar ring nucleosides and nucleotides. Improved method for the interpretation of proton magnetic resonance coupling constants. J. Am. Chem. Soc. 95:2333-2344. Brimacombe, J.S. and Ching, O.A. 1968. Nucleophilic displacement reactions in carbohydrates. Carbohydr. Res. 8:82-88. Caruthers, M.H. 1991. Chemical synthesis of DNA and DNA analogues. Acc. Chem. Res. 24:278284. Choleva, Y., Nørholm, M., Pedersen, S., Mouritzen, P., Høiby, P.E., Nielsen, A.T., Møller, S., Jakobsen, M.H., and Kongsbak, L. 2001. Multiplex SNP genotyping using locked nucleic acids and microfluidics. J. Assoc. Lab. Automation 6:9297. Codington, J.F., Fecher, R., and Fox, J.J. 1960. Pyrimidine nucleosides. VII. Reactions of 2′,3′,5′-trimesyloxyuridine. J. Am. Chem. Soc. 82:2794-2803. Horton, D. and Tindall, C.G. Jr. 1970. Methyleneinsertion reactions with unsaturated sugars, synthesis of 4-C-cyclopropyl-D-ribo-tetrafuranose derivatives. Carbohydr. Res. 15:215-232. Jones, G.H., Taniguchi, M., Tegg, D., and Moffat, J.G. 1979. 4′-Substituted nucleosides. 5. Hydroxymethylation of nucleoside 5′-aldehydes. J. Org. Chem. 44:1309-1317. Koshkin, A.A., Rajwanshi, V.K., and Wengel, J. 1998a. Novel convenient syntheses of LNA [2.2.1]bicyclo nucleosides. Tetrahedron Lett. 39:4381-4384.
Koshkin, A.A., Singh, S.K., Nielsen, P., Rajwanshi, V.K., Kumar, R., Meldgaard, M., Olsen, C.E., and Wengel, J. 1998b. LNA (locked nucleic acids): Synthesis of the adenine, cytosine, guanine, 5-methylcytosine, thymine and uracil bicyclonucleoside monomers, oligomerisation, and unprecedented nucleic acid recognition. Tetrahedron 54:3607-3630. Koshkin, A.A., Fensholdt, J., Pfundheller, H.M., and Lomholt, C. 2001. A simplified and efficient route to 2′-O, 4′-C-methylene-linked bicyclic ribonucleosides (LNA). J. Org. Chem. 66:85048512. Leland, D.L. and Kotick, M.P. 1974. Studies on 4-C-(hydroxymethyl)pentofuranoses. Synthesis of 9-[4-C-(hydroxymethyl)-α-L-threo-pentofuranosyl]adenine. Carbohydr. Res. 38:C9-C11. Obika, S., Nanbu, D., Hari, Y., Morio, K., In, Y., Ishida, T., and Imanishi, T. 1997. Synthesis of 2′- O, 4′-C-methyleneuridine and -cytidine. Novel bicyclic nucleosides having a fixed C3′endo sugar puckering. Tetrahedron Lett. 38:8735-8738. Obika, S., Nanbu, D., Hari, Y., Andoh, J., Morio, K., Doi, T., and Imanishi, T. 1998. Stability and structural features of the duplexes containing nucleoside analogues with a fixed N-type conf o rm ation, 2′-O,4′-C-methyleneribonucleosides. Tetrahedron Lett. 39:5401-5404. Obika, S., Uneda, T., Sugimoto, T., Nanbu, D., Minami, T., Doi, T., and Imanishi, T. 2001. 2′-O,4′C-Methylene bridged nucleic acid (2′,4′-BNA): Synthesis and triplex-forming properties. Bioorg. Med. Chem. 9:1001-1011. Ørum, H., Jakobsen, M.H., Koch, T., Vuust, J., and Borre, M.B. 1999. Detection of the factor V Leiden mutation by direct allele-specific hybridization of PCR amplicons to photoimmobilized locked nucleic acids. Clin. Chem. 45:1898-1905. Singh, S.K., Nielsen, P., Koshkin, A.A., and Wengel, J. 1998. LNA (locked nucleic acids): Synthesis and high-affinity nucleic acid recognition. Chem. Commun. (1998):455-456. Sinha, N.D., Biernat, J., and Köster, H. 1983. β-Cyanoethyl N,N-dialkylamino/N-morpholinomonochloro phosphoramidites, new phosphitylating agents facilitating ease of deprotection and work-up of synthesized oligonucleotides. Tetrahedron Lett. 24:5843-5846. Sowa, W. and Thomas, G.H.S. 1966. The oxidation of 1,2:5,6-di-O-isopropylidene-D-glucose by dimethylsulfoxide-acetic anhydride. Can. J. Chem. 44:836-838. Vorbrüggen, H. and Höfle, G. 1981. On the mechanism of nucleoside synthesis. Chem. Ber. 114:1256-1268. Vorbrüggen, H., Krolikiewicz, K., and Bennua, B. 1981. Nucleoside synthesis with trimethylsilyl triflate and perchlorate as catalysts. Chem. Ber. 114:1234-1255.
Synthesis of Modified Oligonucleotides and Conjugates
4.12.15 Current Protocols in Nucleic Acid Chemistry
Supplement 8
Waga, T., Nishizaki, T., Miyakawa, I., Ohrui, H., and Meguro, H. 1993. Synthesis of 4′-C-methylnucleosides. Biosci. Biotech. Biochem. 57:14331438. Wahlestedt, C., Salmi, P., Good, L., Kela, J., Johnsson, T., Høkfelt, T., Broberger, C., Porreca, F., Lai, J., Ren, K., Ossipov, M., Koshkin, A., Jakobsen, N., Skouv, J., Oerum, H., Jacobsen, M.H., and Wengel, J. 2000. Potent and nontoxic antisense oligonucleotides containing locked nucleic acids. Proc. Natl. Acad. Sci. U.S.A. 97:5633-5638. Youssefyeh, R., Tegg, D., Verheyden, J.P.H., Jones, G.H., and Moffat, J.G. 1977. Synthetic routes to 4′-hydroxymethylnucleosides. Tetrahedron Lett. 5:435-438. Youssefyeh, R.D, Verheyden, J.P.H., and Moffat, J.G. 1979. 4′-Substituted nucleosides. 4. Synthesis of some 4′-hydroxymethyl nucleosides. J. Org. Chem. 44:1301-1309.
Key References Koshkin et al., 1998a,b. See above. This unit’s protocols were developed based on the synthetic results described in these papers. Wengel, J. 1999. Development of locked nucleic acid. Acc. Chem. Res. 32:301-310. This article describes the development of LNA.
Contributed by Henrik M. Pfundheller and Christian Lomholt Exiqon A/S Vedbaek, Denmark
Locked Nucleic Acids: Synthesis and Characterization of LNA-T Diol
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Cellular Delivery of Locked Nucleic Acids (LNAs)
UNIT 4.13
This unit describes the introduction of locked nucleic acid (LNA) oligomers (Fig. 4.13.1; Koshkin et al., 1998; Obika et al., 1998; Wang et al., 1999; reviewed in Braasch and Corey, 2001) into cells. It is intended to extend the discussion of the synthesis and characterization of LNA that is found in UNIT 4.12. INTRODUCTION OF LNA OLIGOMERS INTO CELLS As is the case for most other types of oligonucleotide, there is little reason to believe that LNA oligomers will be able to spontaneously enter most types of cultured cells and locate a cellular target. Since LNAs possess a negatively charged backbone, one simple method for promoting uptake is the use of cationic lipid. A step-wise sample procedure for a 48-well plate format is given below and a schematic summary is provided in Figure 4.13.2.
BASIC PROTOCOL
NOTE: The conditions required for successful transfections will vary from one cell line to the next. It is impossible to predict which combination of lipid and oligonucleotide will be most effective; this must be determined empirically for each cell line. Materials Cells grown to confluence in 75-cm2 tissue culture flasks Complete growth medium (see recipe) 100 µM LNA stock solution (see Support Protocol 1) Opti-MEM I (Invitrogen Life Technologies; reduced-serum medium, containing L-glutamine and no phenol red) LipofectAMINE (Invitrogen Life Technologies) 48-well tissue culture plate (Costar) Repeating pipettor (e.g., Eppendorf) 12.5-mL Combitips (Eppendorf) 65° and 37°C water baths or a thermal cycler 12 × 75–mm round-bottom tubes 37°C, 5% CO2 incubator
base
RO O
O O
P
O O– base
O O
Figure 4.13.1 Structure of LNA.
Synthesis of Modified Oligonucleotides and Conjugates
Contributed by Dwaine A. Braasch and David R. Corey
4.13.1
R′O
Current Protocols in Nucleic Acid Chemistry (2002) 4.13.1-4.13.9 Copyright © 2002 by John Wiley & Sons, Inc.
O
Supplement 9
Additional reagents and equipment for trypsinizing and counting cells (e.g., CPMB APPENDIX 3F) NOTE: LipofectAMINE and Opti-MEM I are important to the success of the experiment and should not be substituted. Prepare cells 1. Beginning with a 75-cm2 tissue culture flask of cells grown to confluence, trypsinize the cells according to standard procedures (e.g., CPMB APPENDIX 3F). The total number of cells (and thus the number of flasks) will depend on the number of LNAs being transfected. For transfection of a single LNA, 33,000 to 39,000 cells are needed (11,000 to 13,000 cells per well at three different concentrations of LNA-LipofectAMINE complex).
2. Suspend the cells in fresh complete growth medium.
inhibitor (100 µM LNA) 6.4 µL
Opti-MEM 143.6 µL
step 9
1.9 µL
Opti-MEM 148.1 µL
step 10 step 11
LipofectAMINE
total volume =
300 µL
15 min
200 nM, V t = 3200 µL
add 2900 µL Opti-MEM
step 12 complex formation
step 14
100 nM, V t = 3000 µL 1500 µL 200 nM solution step 15 1500 µL Opti-MEM 25 nM, V t = 2000 µL 500 µL 100 nM solution 1500 µL Opti-MEM
dispense 200 µL/well
step 15
step 16
step 17 transfection overnight
Cellular Delivery of Locked Nucleic Acids (LNAs)
Figure 4.13.2 Preparation of LipofectAMINE-LNA complexes for cellular transfection, including subsequent dilutions of stock complexes. Vt = total volume. Volumes are sufficient for dispensing six replicates and have been optimized for COS-7 cells.
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3. Draw up the entire volume of cell suspension into a 10-mL pipet and dispense back into the flask with the tip of the pipet pressed lightly against the bottom of the flask. Aggregates of cells will be disrupted, yielding a single-cell suspension that gives more accurate cell counts.
4. Perform a cell count in triplicate using a Coulter counter or hemacytometer (e.g., CPMB APPENDIX 3F) and average the results. 5. Prepare cell suspension in complete growth medium at a density of 44,000 to 52,000 cells/mL. 6. Plate cells 4 to 6 hr prior to transfection. For each LNA to be transfected, plate 250 µL cell suspension (11,000 to 13,000 cells) in three wells of a 48-well tissue culture plate. For accuracy with larger numbers of LNAs, use a repeating pipettor and a 12.5-mL Combitip.
7. Replace cover on plate and disperse cells evenly within the wells by sliding the plate back and forth, gently bumping it against the front lip of a laminar flow hood work surface. Keep cells 4 to 6 hr at 37°C in a 5% CO2 incubator for cells to attach to the surface. Prepare LNA-LipofectAMINE complexes 8. Warm 100 µM LNA stock solution 5 min at 65°C for a 15-mer or 95°C for a 25-mer to disrupt aggregates, and then maintain at 37°C until transfection. Also warm Opti-MEM I to 37°C. 9. In a 12 × 75–mm round-bottom tube, dilute 6.4 µL of 100 µM LNA with 143.6 µL Opti-MEM I. 10. In a separate 12 × 75–mm round-bottom tube, dilute 1.9 µL LipofectAMINE with 148.1 µL Opti-MEM I. This solution can be scaled up in a single tube depending on how many unique conditions are being tested. If, for example, five inhibitors (LNAs) are to be tested, then it would be advantageous to prepare sufficient lipid mix for seven conditions.
11. Add 150 µL LipofectAMINE from step 10 to each LNA in step 9 (total 300 µL) and tap the tube briskly 15 times to mix the reagents and initiate the formation of LNA-LipofectAMINE complexes. 12. Allow the tube to sit 15 min at room temperature in the dark. Opti-MEM I is light sensitive.
Transfect cells 13. While waiting for complexes to form, aspirate off the complete growth medium in each well of the tissue culture plate and replace with 250 µL Opti-MEM I per well. Also set up two tubes for a dilution series of LNA-LipofectAMINE complexes and add 1.5 mL Opti-MEM I to each. 14. When the incubation (step 12) is complete, add 2.9 mL Opti-MEM I to the LNALipofectAMINE complexes and mix well. This gives 3.2 mL at 200 nM LNA for the starting concentration for the serial dilution. The authors typically use 200 nM or 500 nM as the starting concentration.
Synthesis of Modified Oligonucleotides and Conjugates
4.13.3 Current Protocols in Nucleic Acid Chemistry
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15. Transfer 1.5 mL stock to one of the tubes in step 13 (final 3 mL at 100 nM) and mix well. Transfer 500 µL of this solution to the second tube (final 2 mL at 25 mM) and mix well. If 500 nM is used as the starting concentration, a 200 nM dilution should be included.
16. Aspirate the Opti-MEM I wash from the cells and immediately dispense 200 µL of each LNA-LipofectAMINE dilution to the appropriate wells, working backward through the dilution scheme for a given LNA. 17. Allow cells to incubate overnight at 37°C. 18. Aspirate off transfection solution and replace with 250 µL complete growth medium. 19. Incubate >24 hr at 37°C prior to conducting an assay for the effects of the LNA. When developing a protocol for delivering LNAs into cells, it is useful to obtain a fluorophore-labeled LNA. Delivery of the LNA can be visualized by microscopy, facilitating the evaluation and subsequent optimization of delivery conditions. SUPPORT PROTOCOL 1
PREPARATION OF LNA OLIGOMER STOCK SOLUTIONS LNA oligomers arrive lyophilized and should be handled like DNA or RNA oligomers. Materials Locked nucleic acid oligomers (LNAs; Proligo) DNase/RNase-free water (Life Technologies) Spectrophotometer 1. If LNAs have been refrigerated, allow them to equilibrate to room temperature. 2. Centrifuge the samples 2 min at 14,000 × g, room temperature, to collect LNA at the bottom of the tube. 3. Add DNase/RNase-free water to give a stock solution of ∼1 mM. Allow the oligomer to sit undisturbed for 10 to 15 min at room temperature. The estimated concentration is based on the volume, on the mass reported by the manufacturer, and on the molecular weight of a given LNA.
4. Vortex in 5-sec bursts several times. Heat to 65°C for up to 15 min and cool to room temperature. 5. Allow tubes to sit undisturbed for 5 min, room temperature. 6. Centrifuge 2 min at 14,000 × g, room temperature, to pellet any remaining undissolved material. Observe the tube contents carefully at this point, as occasionally there are insoluble materials that can interfere with cellular assays. It is best not to proceed with cellular assays if an LNA exhibits this behavior unless one has significant experience in desalting and purifying oligomers. Consult the manufacturer if solubility properties are not satisfactory.
7. Remove a 1-µL aliquot and dilute it with 144 µL distilled water. Ascertain the absorbance at 260 nm. 8. Calculate the concentration of the LNA using the following equation, where 33 ng/µL is the extinction coefficient of the LNA and 145 is the dilution factor: Cellular Delivery of Locked Nucleic Acids (LNAs)
c (mM) = (A260 × 33 ng/µL × 145)/mol. wt. of LNA
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The value 33 ng/ìL assumes an average extinction coefficient based on an equal population of all bases. If the LNA contains a preponderance of one or two bases, the equation can be modified (e.g., see CPMB APPENDIX 3D) using the appropriate extinction coefficients. The extinction coefficients of the LNA nucleotide analogs are not available to the authors, but it may be reasonable to assume that they are the same as for DNA nucleotides (Proligo, pers. comm.).
9. Adjust concentration to 100 µM and store the stock solution for up to 1 year at 4°C. DETERMINATION OF Tm FOR LNA OLIGOMERS To understand the potential for LNA oligonucleotides to recognize intracellular targets, it is useful to determine Tm values for LNAs with complementary RNA or DNA oligomers.
SUPPORT PROTOCOL 2
Materials LNA oligonucleotides DNA or RNA oligomers 10× Ca2+- and Mg2+-free phosphate-buffered saline (CMF-PBS; Invitrogen Life Technologies or see recipe) 0.1 M Na2HPO4 buffer, pH 7.5 (Fisher) Mineral oil (Sigma) Stoppered cuvette (1-cm pathlength and 1.5-cm Z dimension; Spectrosil Far UV Quartz, Uvonic Instruments) Spectrophotometer with temperature-controlled cuvette holder 1. Calculate the concentration of each of the single-stranded components (LNA and either DNA or RNA). The concentration of LNAs can be determined from the A260 value of a diluted aliquot (see Support Protocol 1, step 8). It is sometimes useful to heat the LNAs to 65°C or higher, depending on the oligomer length and the number of LNA bases, to break up intra- and intermolecular hydrogen bonding and aggregation. When preparing oligonucleotide pairs for Tm analysis or tranfection, it is prudent to heat the samples and ensure that the concentrations are accurate.
2. Prepare a small volume (∼20 to 30 µL) of 100 µM heteroduplex nucleic acid in a solution containing 2.5× CMF-PBS final concentration. 3. Dilute a 5-µL aliquot of the 100 µM heteroduplex mixture with 145 µL of 0.1 M Na2HPO4 buffer, pH 7.5, in a 1-cm-pathlength stoppered cuvette. 4. Overlay this solution with 145 µL mineral oil to minimize evaporation. 5. Monitor the change in the absorbance at 260 nm every 5°C as the temperature is ramped from 100° to 12°C and also back up to 100°C. 6. Fit the data collected from these analyses using van’t Hoff thermal denaturation/renaturation curve analysis to determine the Tm values from the denaturation and renaturation curves (UNIT 7.3). It is not uncommon for entirely LNA oligomers with >11 bases to possess Tm values >95°C.
Synthesis of Modified Oligonucleotides and Conjugates
4.13.5 Current Protocols in Nucleic Acid Chemistry
Supplement 8
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
CMF-PBS (Ca2+- and Mg2+-free phosphate-buffered saline), 10× 10.4 mM KH2PO4 1551.7 mM NaCl 29.6 mM Na2HPO4⋅7H2O, pH 7.4 Store up to 1 year at 4°C Complete growth medium Dulbecco’s modified Eagle’s medium (DMEM) high-glucose without L-glutamine (e.g., Mediatech Cellgro, Fisher) containing: 20 mM HEPES buffer, pH 7.4 (cell-culture grade, Sigma) 10% (w/v) FBS (Atlanta Biologicals) 1× PSF (see recipe) 0.7 mg/mL tylosin (Sigma) 2 mM L-glutamine (Invitrogen Life Technologies) Store up to 4 months at 4°C PSF (penicillin/streptomycin/Fungizone), 100× 10,000 U penicillin 10 mg streptomycin 25 µg/mL amphotericin B (e.g., Fungizone, Invitrogen Life Technologies) Store up to 1 year at −20°C COMMENTARY Background Information LNAs offer several advantages for nucleic acid recognition (Table 4.13.1), which should encourage investigators to consider their use. Currently, oligonucleotides that contain LNA bases can be obtained commercially from Proligo (http://www.proligo.com). Synthesis is based on phosphoramidite chemistry and employs LNA monomers of A, T, G, and 5-methylC. The most striking advantage conferred by use of LNA bases is a dramatic increase in the
Table 4.13.1
Cellular Delivery of Locked Nucleic Acids (LNAs)
affinity of binding to complementary sequences (Table 4.13.2). A single LNA base can increase the melting temperature (Tm) of binding by 10°C, and oligomers that contain several strategically positioned LNA bases can bind with even higher affinity than analogous peptide nucleic acid (PNA) oligomers (Braasch and Elayadi, unpub. observ.). LNA bases are introduced into oligonucleotides by standard synthesis methods (UNIT 4.12), allowing LNA bases to be interspersed among DNA or RNA bases. As a result, important properties such as
Advantages of Locked Nucleic Acids
High affinity hybridization
Tight binding by short LNAs or at high temperatures
Synthesized like DNA/RNA
Enables ready adaptation of existing synthesizers. Simple to intersperse DNA or RNA bases to modulate Tm values or RNase H sensitivity.
Negatively charged backbone
Good solubility; investigators who work with DNA or RNA will find LNA easy to work with. Ability to incorporate phosphorothioate linkages to improve stability or pharmacokinetic properties.
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Table 4.13.2 Melting Temperature (Tm) Values for LNAs and LNA-DNA Hybrids
LNA or LNA-DNAa
Tm
Tm(ref)b ∆Tm
∆Tm/LNA Reference base
GTGTTTTGC GTGTCCGAGACGTTG
52 72
28 59
24 13
5 1.5
Kumar et al. (1998) Wahlstedt et al. (2000)
GTGTCCGAGACGTTG GTGTCCGAGACGTTG
83 >90
59 59
24 >31
3 2
Wahlstedt et al. (2000) Wahlstedt et al. (2000)
CACTATACG CTGATATGC
40 36.8
29 27.2
11 9.6
3.3 9.6
Koshkin et al. (1998) Bondensgaard et al. (2000)
CTGATATGC AGGGTCGCTmeCGGTGT
51.6 >96
27.2 53
24.4 43
8.1 3
Bondensgaard et al. (2000) Braasch (unpub. observ.)
AGGGTCGCTmeCAATGT meCAGTTAGGGTTAG
83 81
NPc 50
— 31
— 3.1
Braasch (unpub. observ.) Braasch (unpub. observ.)
meCAGTTAGAATTAG
TAGGGT
65 56
NPc NDc
— —
— —
Braasch (unpub. observ.) Braasch (unpub. observ.)
TAGGGTTA AGGATmeCTAGGTGAA
74 >96
22 53
52 25
6.5 2.9
Braasch (unpub. observ.) Braasch (unpub. observ.)
AGGATmeCTAGG AGGATmeCTAGGTGAA
73 59
39 53
34 6
3.4 0.6
Braasch (unpub. observ.) Braasch (unpub. observ.)
aUnderlined bases are LNA. meC, 5-methylcytosine. All oligomers are shown from 5′ to 3′. bReference T values are for analogous DNA oligonucleotides. m cND, melting temperature not detected; NP, analysis not performed. ∆T and ∆T /LNA base could thus not be calculated. m m
Tm or RNase H activation can be tailored to meet the specifications of individual applications. This combination of high-affinity hybridization with standard synthesis protocols is powerful because it encourages adapting LNAs to existing protocols that use oligonucleotides.
Critical Parameters When designing oligonucleotides that are intended to function inside cells, the ability of the oligonucleotides to form duplexes that act as substrates for RNase H is a primary consideration (Crooke, 1999). RNase H degrades RNA-DNA hybrids, allowing oligonucleotides that contain DNA to promote the cleavage of mRNA. If antisense inhibition of gene expression is desired, this can be an advantage since the target mRNA is permanently inactivated, and the antisense oligomer can then move on to inactivate additional mRNA molecules. Oligonucleotides that cannot recruit RNase H (i.e., that do not contain DNA portions) can block the binding of the translation apparatus when targeted to the 5′ terminus of the untranslated region (Baker et al., 1997; Doyle et al., 2001). Oligomers that do not activate RNase H can also redirect splicing when targeted to splice
sites (Kang et al., 1998), but when they are targeted to other mRNA sequences, they are likely to be displaced by the ribosome. Oligomers that contain only LNA bases activate RNase H poorly (Wahlestedt et al., 2000). However, because LNAs are made using protocols similar to those used for DNA synthesis, it is straightforward to incorporate DNA bases into LNA-DNA chimera. This provides the experimenter with the choice of whether or not to incorporate RNase H sensitivity into oligonucleotide design by including a contiguous run of at least six DNA bases. If antisense gene inhibition is desired, this strategy may allow a wider range of sequences to be targeted. If simple steric blocking of the RNA target is required, the potential to direct RNase H cleavage is unnecessary and even counterproductive because it might lead to unintentional destruction of nontargeted RNA substrates. LNA bases confer some increase in the stability of oligomers to degradation by nucleases. However, to achieve maximal stability in animal studies, it is likely that one or two phosphorothioate (PS) linkages will need to be substituted at both the 3′- and 5′-terminal linkages. As noted above, LNA synthesis is similar to the
Synthesis of Modified Oligonucleotides and Conjugates
4.13.7 Current Protocols in Nucleic Acid Chemistry
Supplement 8
synthesis of DNA or RNA, allowing PS linkages to be added routinely (Kumar et al., 1998). Complete substitution of phosphodiester linkages with PS linkages has also been noted to improve the pharmacokinetic properties of antisense oligonucleotides (Geary et al., 2001). It is reasonable to believe that LNA-containing oligomers will also need to be modified with PS linkages to achieve in vivo efficacy, though animal studies will be necessary to establish if this truly is the case for LNA-containing chimeric oligonucleotides. The choice of whether or not to exploit the potential for RNase H activation influences how antisense activity will be examined on a case-by-case basis. LNA oligomers that contain DNA segments that can activate RNase H will cause RNA to be degraded, allowing efficacy to be judged by northern analysis. LNA oligomers that cannot activate RNase H can be evaluated by examining the expression of the protein target or by measuring its activity. As with any antisense experiment, use of control oligonucleotides that contain mismatched bases is necessary to support the belief that an effect is specific, i.e., due to binding to the intended mRNA target.
Troubleshooting
Cellular Delivery of Locked Nucleic Acids (LNAs)
When oligonucleotides are introduced into cells, they may prove to be toxic. This toxicity could be due to successful inhibition of the target gene function. However, toxicity could also be due to (1) binding to one or more nontarget proteins, (2) hybridization to one or more nontarget nucleic acid sequences, or (3) poisoning of the cells by small molecule impurities or endotoxins. The primary consideration is that the observed effects should be consistent with the biology of the system being examined. If cell death is observed, LNAs can be purified by desalting to remove small molecule contaminants. Alternatively, a fresh synthesis of LNA can be performed to determine if newly made material behaves similarly. Toxicity could also be caused by improper choice of transfection conditions, reagent concentrations, cell line, or lipids. The window between the conditions that produce optimal LNA delivery and those that cause cells to die is likely to be small. Use of fluorophore-labeled LNAs provides a convenient method for evaluating the success of a given protocol. The authors have also observed that some syntheses of LNAs are relatively insoluble. If this occurs, the experimenter should consult with the manufacturer.
Anticipated Results There are many effective antisense oligonucleotides that do not contain LNA bases. Why use LNA? Why not stay with standard oligonucleotide designs? The use of LNAs is only three years old, and there are no definitive answers to these questions. However, it is reasonable to speculate that the ability of LNA bases to dramatically improve the affinity of binding might increase the potency, specificity, and predictability of antisense action. The resulting increase in efficacy would allow “knock down” phenotypes to be generated more easily, and the fact that LNA bases can be easily incorporated into oligonucleotides allows this hypothesis to be readily tested. Microscopic examination of the delivery of a fluorophore-labeled LNA is a useful demonstration that transfection conditions are promoting LNA uptake by cells. In the authors’ experience, micrographs show that LNA is distributed throughout the cytoplasm and nucleus, with some punctate staining indicating areas of high concentration.
Time Considerations LNAs normally can be obtained within 2 weeks of placing an order with Proligo. LNAs should completely dissolve in water within 20 min. Quantification of LNA concentration by UV spectrophotometry and determination of a Tm value should take ∼2 hr. Transfection of LNA into cells should require an additional 2 to 4 hr. The time required for observation of a phenotype will vary from hours to days, depending on the gene being targeted.
LITERATURE CITED Baker, B.F., Lot, S.S., Condon, T.P., Cheng-Flournoy, S., Lesnik, E.A., Sasmor, H.M., and Bennett, C.F. 1997. 2′-O-(2-Methoxy)ethyl-modified anti-intercellular adhesion molecule 1 (ICAM-1) oligonucleotides selectively increase the ICAM-1 mRNA level and inhibit formation of the ICAM-1 translation initiation complex in human umbilical vein endothelial cells. J. Biol. Chem. 272:11994-12000. Bondensgaard, K., Petersen, M., Singh, S.K., Rajwanshi, V.K., Kumar, R., Wengel, J., and Jacobsen, J.P. 2000. Structural studies of LNA:RNA duplexes by NMR: Conformations and RNase H activity. Chem. Eur. J. 6:2687-2695. Braasch, D.A. and Corey, D.R. 2001. Locked nucleic acids: Fine-tuning nucleic acid recognition. Chem. Biol. 8:1-7. Crooke, S.T. 1999. Molecular mechanisms of antisense drugs: Human RNase H. Antisense Nucl. Acid Drug Devel. 9:377-379.
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Current Protocols in Nucleic Acid Chemistry
Doyle, D.F., Braasch, D.A., Simmons, C.G., Janowski, B.A., and Corey, D.R. 2001. Inhibition of gene expression inside cells by peptide nucleic acids: Effect of mRNA target sequence, mismatched bases, and PNA length. Biochemistry 40:53-64.
Obika, S., Nanbu, D., Hari, Y., Andoh, J., Morio, K., Doi, T., and Imanishi, T. 1998. Stability and structural features of the duplexes containing the nucleoside analogues with a fixed N-type conf o rmation, 2′-O,4′-C-methyleneribonucleosides. Tetrahedron Lett. 39:5401-5404.
Geary, R.S., Yu, R.Z., and Levin, A.A. 2001. Pharmacokinetics of phosphorothioate antisense oligonucleotides. Curr. Opin. Investigational New Drugs 2:562-573.
Wahlestedt, C., Salmi, P., Good, L., Kela, J., Johnsson, T., Hokfelt, T., Broberger, C., Porreca, F., Lai, J., Ren, K., Ossipov, M., Koshkin, A., Jakobsen, N., Skouv, J., Oerum, H., Havsteen Jacobsen, M., and Wengel, J. 2000. Potent and nontoxic antisense oligonucleotides containing locked nucleic acids. Proc. Natl. Acad. Sci. U.S.A. 97:5633-5638.
Kang, S.H., Cho, M.J., and Kole, R. 1998. Up-regulation of luciferase gene expression with antisense oligonucleotides—Implications and applications in functional assay developments. Biochemistry 37:6235-6239. Koshkin, A.A., Singh, S.K., Nielsen, P., Rajwanshi, V.K., Kumar, R., Meldgaard, M., Olsen, C.E., and Wengel, J. 1998. LNA (locked nucleic acids): Synthesis of the adenine, cytosine, guanine, 5-methylcytosine, thymine, and uracil bicyclonucleoside monomers, oligomerisation and unprecedented nucleic acid recognition. Tetrahedon 54:3607-3630. Kumar, R., Singh, S., Koshkin, A.A., Rajwanshi, V.K., Meldgaard, M. and Wengel, J. 1998. The first analogues of LNA (locked nucleic acids): Pho sphorothioate-LNA and 2′-thio-LNA. Bioorg. Med. Chem. Lett. 8:2219-2222.
Wang, G., Gunic, E., Girardet, J-L., and Stoisavljevic, V. 1999. Conformationally locked nucleosides. Synthesis and hybridization properties of oligodeoxynucleotides containing 2′4′-Cbridged 2′-deoxynucleosides. Bioorg. Med. Chem. Lett. 9:1147-1150.
Contributed by Dwaine A. Braasch and David R. Corey University of Texas Southwestern Medical Center at Dallas Dallas, Texas
Synthesis of Modified Oligonucleotides and Conjugates
4.13.9 Current Protocols in Nucleic Acid Chemistry
Supplement 8
Solid-Phase Synthesis of Branched Oligonucleotides
UNIT 4.14
This unit describes the synthesis of nucleic acids containing vicinal 2′,5′- and 3′,5′-phosphodiester bonds. These molecules occur in the cell nucleus, and are formed during the splicing of precursor messenger RNA (pre-mRNA). As such they have many potential applications in nucleic acid biochemistry, particularly as tools for probing the substrate specificity of lariat debranching enzymes, and as tools for studying pre-mRNA splicing (e.g., Nam et al., 1994; Carriero et al., 2001). The assembly of these branched nucleic acids (bNAs) on a solid support can be achieved by following two strategies (Damha and Zabarylo, 1989; Braich and Damha, 1997). The first, referred to as the convergent strategy, is based on well-established automated phosphoramidite chemistry (UNITS 3.3 & 3.5). This method uses a ribonucleoside bisphosphoramidite as the branch-introduction synthon (see Basic Protocol 1). The branching reagent serves to couple together solid-support-bound chains, thus forming a branch juncture with the desired vicinal 2′,5′- and 3′,5′-phosphodiester bonds (see Basic Protocol 2). For efficient branching to occur, CPG supports with high nucleoside loadings are used (see Support Protocol 1). With this approach, Y- and V-shaped molecules having identical 2′ and 3′ chains are readily assembled. The second method is a divergent approach that permits the regiospecific synthesis of bNAs using readily available phosphoramidite reagents (see Basic Protocol 3). An important feature of this method is the assembly of a linear DNA:RNA chimera containing a single 2′-O-silylribonucleoside residue in the middle of the chain. Subsequent removal of the 2-cyanoethyl and silyl protecting groups without detaching the nascent oligonucleotide from the solid support is another salient feature of this approach. This releases an internal 2′-OH group from which orthogonal synthesis of a branch can be carried out. This unit also describes methods used in the authors’ laboratory for the deprotection (see Support Protocol 2), purification, and characterization of branched oligonucleotides. Preferred methods for purification of bNAs are anion-exchange HPLC (see Support Protocol 3) and polyacrylamide gel electrophoresis (see Support Protocol 4). The branched nature of the molecule is confirmed by enzymatic hydrolysis of the bNA to its constituent nucleosides using nuclease P1 (see Support Protocol 3). Further characterization may be conducted via nucleoside composition analysis using snake venom phosphodiesterase (UNIT 10.6) or MALDI-TOF-MS (UNIT 10.1). SYNTHESIS AND CHARACTERIZATION OF THE ADENOSINE BRANCHING SYNTHON N6-BENZOYL-5′-O-(4,4′-DIMETHOXYTRITYL)ADENOSINE-2′,3′-BIS-O-(2-CYANOETHYL-N,N-DIISOPROPYL) PHOSPHORAMIDITE The authors’ group has been predominantly interested in the synthesis of branched RNA fragments related to the lariat intermediates formed during pre-mRNA splicing. Such intermediates contain almost exclusively adenosine at the branch point; therefore, the protocol given below describes the synthesis of the adenosine branching phosphoramidite synthon (BIS-A; S.3; Fig. 4.14.1) used for the synthesis of symmetrical, branched DNA and RNA oligonucleotides (Damha and Ogilvie, 1988). The same protocol may be adapted to the synthesis of the corresponding U, C, and G bisphosphoramidites. The starting protected nucleoside, N6-benzoyl-5′-O-(4,4′-dimethoxytrityl)adenosine (S.1), is commercially available (ChemGenes). Alternatively, it may be synthesized from adenosine using the transient benzoylation procedure of Ti et al. (1982), followed by dimethoxytritylation of the 5′-hydroxyl group (Hakimelahi et al., 1982; Wu et al., 1989). Contributed by Sandra Carriero and Masad J. Damha Current Protocols in Nucleic Acid Chemistry (2002) 4.14.1-4.14.32 Copyright © 2002 by John Wiley & Sons, Inc.
BASIC PROTOCOL 1
Synthesis of Modified Oligonucleotides and Conjugates
4.14.1 Supplement 9
O
[ABz] [DMTr]
Ph
HN O
N
N
OMe Ph
HN N N O
O
N N
N
i -Pr2N
DMTrO
N
O
P OCH2CH2CN
Cl 2
NCCH2CH2O
O
O
P
P
i -Pr2N
i -Pr2N
OCH2CH2CN
DMAP, DIPEA, THF OH
HO OMe
1
3 (4 diastereomers)
Figure 4.14.1 Reaction scheme demonstrating the synthesis of N6-benzoyl-5′-O-(4,4′-dimethoxytrityl)adenosine-2′,3′-Obis-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (S.3) from the starting protected nucleoside N6-benzoyl-5′-O-(4,4′-dimethoxytrityl)adenosine (S.1). The structure of the chlorophosphoramidite (S.2) is also shown. Abbreviations: Bz, benzoyl; DIPEA, N,N-diisopropylethylamine; DMAP, 4-dimethylaminopyridine; DMTr, 4,4′-dimethoxytrityl.
The synthesis of the branching synthon involves the phosphitylation of the 2′ and 3′ secondary hydroxyls of the ribose sugar (Fig. 4.14.1) using an excess of 2-cyanoethylN,N-diisopropylchlorophosphoramidite (S.2). Reaction conditions, workup, chromatographic purification, and characterization of the product N6-benzoyl-5′-O-(4,4′-dimethoxytrityl)adenosine-2′,3′-bis-O-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (S.3) are described. Materials N6-Benzoyl-5′-O-(4,4′-dimethoxytrityl)adenosine (S.1; ChemGenes) 4-Dimethylaminopyridine (DMAP; 99%; Aldrich) Nitrogen or argon gas, dry Anhydrous THF (see recipe) in a septum-sealed distillation collection bulb Anhydrous DIPEA (see recipe) 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite (S.2; ChemGenes) 1:1 (v/v) dichloromethane/diethyl ether 20% (v/v) sulfuric acid (optional) Ethyl acetate prewashed with 5% (w/v) NaHCO3 NaCl solution, saturated Sodium sulfate (Na2SO4), anhydrous 50:47:3 (v/v/v) CH2Cl2/hexanes/triethylamine Silica gel (230- to 400-mesh) in 50:47:3 CH2Cl2/hexanes/triethylamine 95% (v/v) ethanol Diethyl ether
Solid-Phase Synthesis of Branched Oligonucleotides
50-mL oven- or flame-dried round-bottom flask with rubber septum Glass syringe and needle, oven dried 2 × 5 cm silica-coated thin-layer chromatography (TLC) plate with fluorescent indicator (e.g., Kieselgel 60 F254 aluminum sheets) 254-nm UV light source 500-mL separatory funnel Gravity filtration device and filter paper
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Current Protocols in Nucleic Acid Chemistry
250- and 500-mL round-bottom flasks Rotary evaporator with a water aspirator 5 × 25–cm glass chromatography column with solvent reservoir bulb Additional reagents and materials for thin-layer chromatography (TLC; APPENDIX 31 3D), column chromatography (APPENDIX 3E), P-NMR (UNIT 7.2), and mass spectrometry (UNITS 10.1 & 10.2) Phosphitylate 2′- and 3′-OH 1. Place 1.3 g (2.0 mmol) N6-benzoyl-5′-O-(4,4′-dimethoxytrityl)adenosine (S.1) and 84 mg (0.7 mmol) DMAP into a 50-mL oven- or flame-dried round-bottom flask. 2. Cap the flask with a rubber septum and purge the flask with dry nitrogen or argon. Care must be taken to avoid the presence of moisture throughout the entire reaction.
3. Withdraw anhydrous THF from a septum-sealed distillation collection bulb using an oven-dried glass syringe and needle. Add 6.0 mL THF to the purged flask with stirring until the starting material is completely dissolved. 4. Add 3.6 mL (21 mmol) anhydrous DIPEA and stir the mixture. 5. Slowly add 1.8 mL (8.3 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite (S.2). If the THF solution is sufficiently dry, a white precipitate should form after ∼1 min. This is the diisopropylethylammonium hydrochloride salt that forms during the reaction.
6. Stir the mixture 1 hr at room temperature or until the reaction is complete. Reaction of the secondary hydroxyls of the starting material with S.2 is fast. If TLC analysis (see below) reveals that the reaction is not complete after 1 hr, a further 0.5 mmol S.2 should be added dropwise and stirring continued until all of the starting material is consumed.
Monitor reaction by TLC 7. Spot the reaction mixture onto a precut 2 × 5–cm silica-coated TLC plate with fluorescent indicator and develop using 1:1 (v/v) dichloromethane/diethyl ether (APPENDIX 3D). Addition of 1% to 3% trietylamine may help prevent detritylation.
8. Visualize heterocyclic bases under a 254-nm UV light source. If desired, spray the plate with 20% sulfuric acid in order to visualize the dimethoxytrityl-bearing species. CAUTION: Wear protective eyewear. TLC analysis should indicate complete conversion to products, which exhibit larger Rf values than the starting protected nucleoside. Since the phosphitylation reaction gives rise to two new chiral centers (2′- and 3′-P), the product (S.3) consists of a mixture of four diastereomers. The products appear as two spots (Rf = 0.51 and 0.40) or as one dumbbellshaped spot, because the solvent system partially resolves the four diastereomeric products. A minor side product forms, which migrates between the product (S.3) and starting material (S.1). This is likely the nucleoside-2′,3′-H-bis-phosphonate (Rf = 0.25) that forms via hydrolysis of S.3.
Work up reaction 9. Transfer the reaction mixture to a 500-mL separatory funnel and add 100 mL prewashed ethyl acetate. The ethyl acetate is prewashed with 5% NaHCO3 in order to prevent detritylation and/or activation of the phosphoramidite moiety.
10. Wash the ethyl acetate layer five times each with 100 mL saturated NaCl solution. The diisopropylammonium hydrochloride salt dissolves.
Synthesis of Modified Oligonucleotides and Conjugates
4.14.3 Current Protocols in Nucleic Acid Chemistry
Supplement 9
11. Dry the organic layer over anhydrous Na2SO4. Add more Na2SO4 if the salt crystals clump together upon swirling. When the solution is dry, the nonhydrated crystals will float in solution upon swirling.
12. Gravity filter the resulting solution through filter paper into a 250-mL round-bottom flask. Rinse the Na2SO4 crystals with 10 to 20 mL ethyl acetate. 13. Remove the solvent under reduced pressure (i.e., in a rotary evaporator with a water aspirator) to yield the crude product as a yellow oil. Isolate and characterize product 14. Prepare a 5 × 25–cm glass chromatography column by adding a slurry of 40 g silica gel in 50:47:3 CH2Cl2/hexanes/triethylamine. Precondition with the same solvent. 15. Dissolve the crude material in a minimum amount of 50:47:3 CH2Cl2/hexanes/triethylamine and load on column. Perform chromatography at a rate of ∼1 in. solvent/min using a small amount of air pressure (APPENDIX 3E). Collect product in 10-ml fractions in small test tubes. 16. Combine product-containing fractions into a 500-mL round-bottom flask and concentrate to an oil on a rotary evaporator. 17. Remove residual triethylamine by co-evaporating the oil first with 50 mL of 95% ethanol followed by 50 mL diethyl ether, to provide the pure product as a pale yellow foam. 18. Store bisphosphoramidite at –20°C under an inert atmosphere protected from light. Phosphoramidites are particularly sensitive to UV light; therefore, it is best to store the bisphosphoramidite in a dark bottle (or a bottle covered with aluminum foil) in a –20°C freezer. Under these conditions, the phosphoramidite may be stored for an indefinite period of time. Prior to use, its purity may be verified via TLC analysis. If partial decomposition has occurred, or the coupling reactions with S.3 are poor, the compound should be subjected to chromatography again as described above.
19. Characterize by TLC (APPENDIX 3D), 31P-NMR (UNIT 7.2), and mass spectrometry (UNITS 10.1 & 10.2). 31
P-NMR spectra (400 MHz) of S.3 were measured on a Varian XL-400 spectrometer using CD3CN as the solvent (Fig. 4.14.2). Chemical shifts are reported in parts per million (ppm) and are downfield (positive value) from 85% H3PO4 (external standard). Fast atom bombardment mass spectrometry (FAB-MS) analysis was conducted on a Kratos MS25RFA high-resolution mass spectrometer using a p-nitrobenzyl alcohol (NBA) matrix. N6-Benzoyl-5′-O-(4,4′-dimethoxytrityl)adenosine-2′,3′-bis-O-(2-cyanoethyl-N,N-diisopropylphosphoramidite) (S.3): yield 52% (560 mg); Rf (1:1 CH2Cl2/diethyl ether): 0.51 and 0.40; 31P-NMR (400 MHz, CD3CN): diastereomer 1, 152.3 and 150.7 ppm (5JP-P = 10.1 Hz); diastereomer 2, 151.9 and 150.5 ppm (5JP-P = 6.6 Hz); diastereomer 3, 151.8 and 151.0 ppm (5JP-P = 4.6 Hz); diastereomer 4, 151.3 and 151.2 ppm (5JP-P = 8.9 Hz); FAB-MS anal. calc’d.: 1074.17; observed: 1074.57; [M]+. BASIC PROTOCOL 2
Solid-Phase Synthesis of Branched Oligonucleotides
CONVERGENT SYNTHESIS OF SYMMETRICAL BRANCHED NUCLEIC ACIDS The procedure described below for the synthesis of bNAs is carried out on a 1-µmol scale using an ABI 381A DNA synthesizer (Damha and Zabarylo, 1989; Damha et al., 1992). The condensation of two adjacent linear oligonucleotides (prepared from standard RNA or DNA phosphoramidites; Fig. 4.14.3) with the adenosine bisphosphoramidite synthon (S.3; Fig. 4.14.1) produces bNAs that contain identical branches connected via vicinal 2′,5′- and 3′,5′-phosphodiester linkages (Fig. 4.14.4). The same protocol may be utilized for the synthesis of bNAs containing D-xylose or D-arabinose instead of D-ribose at the
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Current Protocols in Nucleic Acid Chemistry
branchpoint, using the appropriate bisphosphoramidite synthons (Damba and Ogilvie, 1988; Noronha, Carriero, Agha, and Damha, unpub. observ.). Branched nucleic acid synthesis works very well on commercially available solid supports (i.e., LCAA-CPG) containing 20 to 40 µmol nucleoside per gram support; however, even better yields are attainable on CPG supports with higher loadings (e.g., 90 µmol/g; Fig. 4.14.5). Such supports can be prepared using HATU/DMAP as the coupling reagents (see Support Protocol 1), and are ideal for the synthesis of short-length bNAs (e.g., trimers) since, in this case, high loadings ensure proper distance between the neighboring CPGbound nucleosides (Damha and Zabarylo, 1989). Materials 5′-O-(4,4′-Dimethoxytrityl)-N-protected-2′-deoxyribonucleoside- or -ribonucleoside-derivatized succinyl-LCAA-CPG (ChemGenes; also see Support Protocol 1) Cap A and B capping reagents (see recipes) DNA and/or RNA 3′-phosphoramidites (S.4a-d and S.5a-d; Fig. 4.14.3)
2 2 1 1
3
3 4
152.5
152.0
151.5
4
151.0
150.5 PPM
Figure 4.14.2 31P-NMR of N6-benzoyl-5′-O-DMTr-adenosine-2′,3′-bis-O-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (S.3). Due to the two chiral phosphorus centers, the compound exists as a mixture of four diastereomers (22 = 4; Fig. 4.14.1). Each diastereomer displays two sets of phosphorus signals (4 isomers × 2 31P signals = 8 signals). The doubling or splitting of each signal (8 × 2 = 16) is due to long-range coupling between the two chiral 31P atoms (nuclear spin = 1⁄2) of the bisphosphoramidite. The spectrum was recorded on a Varian XL-400 spectrometer using CD3CN as the solvent. The chemical shift and coupling constants for the four diastereomers (shown as numbers near peaks) are as follows: diastereomer 1, 152.3 and 150.7 ppm (5JP-P = 10.1 Hz); diastereomer 2, 151.9 and 150.5 ppm (5JP-P = 6.6 Hz); diastereomer 3, 151.8 and 151.0 ppm (5JP-P = 4.6 Hz); diastereomer 4, 151.3 and 151.2 ppm (5JP-P = 8.9 Hz).
Synthesis of Modified Oligonucleotides and Conjugates
4.14.5 Current Protocols in Nucleic Acid Chemistry
Supplement 9
DNA 3′-phosphoramidites B
DMTrO
RNA 3′-phosphoramidites B
DMTrO
O O
i -Pr2N P
OCH2CH2CN 4a-d
DNA 5′-phosphoramidites OCH2CH2CN
O NCCH2CH2O
O
i -Pr2N
P
B
O
O
O Si
P
DMTrO
i -Pr2N
6a-d
5a-d
4-6a B = N 6-benzoyladenin-9-yl 4-6b B = N 2-isobutyrylguanin-9-yl 4-6c B = N 4-benzoylcytosin-1-yl 4d, 6d B = thymin-1-yl 5d B = uracil
Figure 4.14.3 Chemical structures of DNA and RNA phosphoramidites used for bNA synthesis: 5′-O-DMTr-N-protected-2′-deoxyribonucleoside-3′-O-(2-cyanoethyl-N,N-diisopropylamino)phosphoramidites (S.4a-d), 5′-O-DMTr-N-protected-2′-O-TBDMS-ribonucleoside-3′-O-(2-cyanoethylN,N-diisopropylamino)phosphoramidites (S.5a-d), and the inverted phosphoramidites 3′-O-DMTrN-protected-2′-deoxyribonucleoside-5′-O-(2-cyanoethyl-N,N-diisopropylamino)phosphoramidites (S.6a-d).
N6-Benzoyl-5′-O-(4,4′-dimethoxytrityl)adenosine-2′,3′-bis-O-(2-cyanoethyl-N,Ndiisopropyl)phosphoramidite (BIS-A; S.3; see Basic Protocol 1) Anhydrous acetonitrile (see recipe) Activator solution: 0.5 M 1H-tetrazole (sublimed) in anhydrous acetonitrile Oxidant solution (see recipe) Detritylation solution (see recipe) Nitrogen or argon gas (optional) Synthesis columns for 1-µmol scale synthesis, with seals and filters (PE Biosystems) and 13-mm aluminum seals (Chromatographic Specialties) ABI 381A automated DNA synthesizer (PE Biosystems) Synthesizer bottles for phosphoramidites, oven dried Fraction collector and 15-mL test tubes 50-mL buret Quartz cuvettes UV-Vis spectrophotometer Additional reagents and equipment for oligonucleotide synthesis (APPENDIX 3C), cleaving and deprotecting oligonucleotides (see Support Protocol 2), and anion-exchange HPLC (see Support Protocol 3) or denaturing PAGE (see Support Protocol 4) NOTE: 1H-Tetrazole is no longer commercially available in crystalline form. Solutions of 0.45 M 1H-tetrazole in anhydrous acetonitrile may be purchased from ChemGenes. For a list of supplementary phosphoramidite activating reagents, see UNIT 3.5. Solid-Phase Synthesis of Branched Oligonucleotides
CAUTION: All solutions for the DNA/RNA synthesizer should be prepared in a well-ventilated fume hood.
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Current Protocols in Nucleic Acid Chemistry
Prepare columns for synthesis 1. Transfer an accurately weighed amount of 5′-O-(4,4′-dimethoxytrityl)-N-protected2′-deoxyribonucleoside- or -ribonucleoside-derivatized succinyl-LCAA-CPG for a 1-µmol synthesis to an assembled synthesis column. The support-bound nucleoside represents the first nucleotide at the 3′-end of the oligonucleotide to be synthesized. See Support Protocol 1 for high-loading CPG supports.
NNNNNN 7 1. NH4OH/EtOH 2. TREAT-HF (RNA only)
DMTrON
DMTrON
DMTrON
DNA/RNA synthesis (3′→5′)
DMTrON
HON
HON
HON
HON
N N N N N
N N N N N
N N N N N
N N N N N
trityl off succinyl-LCAA-CPG 3
failure sequences N N N N N N
N N N N N N
HOABz 2′ 3′
HOABz 3′ 2′
N N N N N N
N N N N N N
N N N N N N
trityl off
failure sequences HOABz 3′ 2′
P*
AcON
DNA/RNA synthesis (3′→5′)
N N N N N
trityl off
HOABz 3′ 2′
N N N N N N
N N N N N N
N N N N N N
P*
N N N N N
1. NH4OH/EtOH 2. TREAT-HF (RNA only)
2′
NNNNNN
3′
1. NH4OH/EtOH 2. TREAT-HF (RNA only)
NNNNNN
A
2′
Y-shaped
NNNNNN
A 3′
NNNNNN
V-shaped NNNNNN
10 2′
NNNNNN
A 3′
8
P*
2′
+
AcON
NNNNNN
3′
NNNNNN
11a
11b
NNNNNN
A
2′
P*
2′
+
A P*
3′
NNNNNN
A 3′
NNNNNN 9a
P* 9b
NNNNNN
NNNNNN
7
7
Figure 4.14.4 Schematic representation demonstrating the convergent synthesis of V- and Yshaped bDNA or bRNA oligonucleotides on a solid support. The method can also be used for the synthesis of short branched sequences—e.g., tetranucleoside triphosphates, NpA(2′pN)3′pN— particularly when the nucleoside loading on the solid support approaches 90 µmol/g. Abbreviations: A, bisphosphoramidite; Bz, benzoyl; DMTr, 4,4′-dimethoxytrityl; N, any nucleotide (RNA or DNA); TREAT-HF, triethylammonium trihydrofluoride; S.7, linear oligonucleotide; S.8 and S.10, full-length bNAs (V- and Y-shaped); S.9a-b and S.11a-b, unbranched linear failure sequences. P* indicates the presence of a 2′- or 3′-linked phosphate to the adenosine branch point, which is formed by the hydrolysis and oxidation of the residual phosphoramidite during solid-phase synthesis.
Synthesis of Modified Oligonucleotides and Conjugates
4.14.7 Current Protocols in Nucleic Acid Chemistry
Supplement 9
(i) T10
A
T10
A
XC +
(ii) T10 (iii) T10
BPB
A
T10 T10 P* T10 T10 P*
Relative % oligonucleotide
CPG loading (µmol/g)
80 70 60 50 40 30 20 10 0 20
30
40
50
60
70
80
CPG loading (µmol/g) i: Branched DNA
ii: Extended isomeric DNA
Figure 4.14.5 Effect of CPG loading on yield of the Y-shaped bDNA, T10A2′,5′-T10 3′,5′-T10 synthesized via the convergent strategy (see Basic Protocol 2). (A) PAGE analysis of the amount of Y-shaped product (i; S.10) and failure sequences (ii; S.11a-b) formed as a function of CPG loading on a 20% denaturing polyacrylamide gel. P* indicates the presence of a 2′or 3′-linked phosphate. (B) Chart demonstrating the increase in the amount of S.10 (i) with increasing nucleoside-CPG loading. The chart also demonstrates the inverse relationship between the amount of extended isomeric linear failures (ii; S.11a-b) and nucleoside-CPG loading. The percentage oligonucleotide was determined by integration of the HPLC peak areas of the compounds in question (see Support Protocol 3) using gradient 3 (see Table 4.14.4).
Table 4.14.1 Automated Cycle on an ABI 381A DNA Synthesizer for Capping of Free CPG-Bound Amino and Hydroxyl Groupsa
Synthesis step
Function
Time (sec)
Column washing steps:
Solid-Phase Synthesis of Branched Oligonucleotides
1 2 3 4
Acetonitrile to waste Acetonitrile to column Argon reverse flush Argon block flush
Column capping steps: 5 6 7
Cap A + cap B to column Wait Repeat steps 5 and 6
Column washing steps: 8 9 10 11 12
Argon reverse flush Acetonitrile to column Repeat steps 8 and 9 Argon reverse flush Argon block flush
5 60 5 5 15 300
5 30 5 5
aAlternatively, CPG may be capped manually using acetic anhydride (see Support Protocol 1).
4.14.8 Supplement 9
Current Protocols in Nucleic Acid Chemistry
2. Acetylate any underivatized amino and hydroxyl groups on the solid support using an ABI 381A automated DNA synthesizer and the capping cycle given in Table 4.14.1 (APPENDIX 3C). This step also removes traces of water from the solid support. Alternatively, see Support Protocol 1 for the manual capping procedure on nucleosideloaded CPG.
Synthesize branched oligonucleotides 3. Weigh out the appropriate amount of DNA (S.4a-d) and/or RNA (S.5a-d) 3′-phosphoramidites (Fig. 4.14.3) and dilute to the appropriate concentration with anhydrous acetonitrile as indicated in Table 4.14.2. 4. Transfer 100 mg BIS-A (S.3) to an oven-dried synthesizer bottle and dilute to 0.03 M with anhydrous acetonitrile. Low concentrations of BIS-A should be employed in the branching reaction, as high concentrations minimize the yield of fully branched product (S.8 and S.10) and favor the extended isomeric side products (S.9a-b and S.11a-b). It is important to prepare the BIS-A stock solution using ≥100 mg material to ensure that (unavoidable) traces of moisture do not consume significant amounts of BIS-A during the coupling (branching) step and reduce the overall yield of bNA synthesis (Fig. 4.14.6E). When the ABI 381A DNA synthesizer is used, this stock solution can be used for as many as 18 branching reactions (170 ìL/addition). Once bNA synthesis is complete, the bottle containing the BIS-A reagent can be removed from the synthesizer, sealed, purged under an inert atmosphere, and left in a freezer (−20°C) for ∼2 weeks. Alternatively, the stock solution can be evaporated under vacuum, and the solid bisamidite recovered for future use.
5. Place all synthesizer reagents (i.e., activator, capping, oxidant, and detritylation solutions, and acetonitrile) and diluted phosphoramidites (step 3) on the appropriate ports of the synthesizer. 6. Place the BIS-A phosphoramidite bottle on the spare phosphoramidite port (the “X” port on the 381A synthesizer).
Table 4.14.2 Concentrations and Optimal Coupling Times of Phosphoramidites in Synthesis of bNAs on an ABI 381A DNA Synthesizer
Phosphoramiditea
Mol. wt. (g/mol)
Concentration (g/mL)
Coupling time (sec)
DNA phosphoramidites (0.1 M in CH3CN b): 857.7 S.4a, S.6a
86
90
S.4b, S.6b S.4c, S.6c S.4d, S.6d
839.7 833.7 744.6
84 83 74
120 90 90
RNA phosphoramidites (0.15 M in CH3CN): S.5a 988.2 S.5b 970.2 S.5c 964.2 S.5d 861.0
148 146 145 129
600 900 600 600
Bisphosphoramidite (0.03 M in CH3CN): S.3 1074.2
32.2
1800
aPhosphoramidite structures are shown in Figure 4.14.3. bFinal concentration of the first inverted DNA phosphoramidite coupled to the 2′-OH of the rA in divergent and
regiospecific synthesis is 0.3 M (see 5′-pD′′ in Fig. 4.14.8).
Synthesis of Modified Oligonucleotides and Conjugates
4.14.9 Current Protocols in Nucleic Acid Chemistry
Supplement 9
Table 4.14.3 Automated 1-µmol Synthesis Cycle for bDNA and bRNA on an ABI 381A DNA Synthesizer
Synthesis step
Function
Time (sec)
Detritylation of support-bound nucleoside: 1 Acetonitrile to waste 2 Acetonitrile to column 3 Argon reverse flush 4 Argon block flush 5 Advance fraction collector 6 3% TCA to waste 7 3% TCA to column 8 Acetonitrile to column 9 3% TCA to column 10 Argon block flush Column washing steps: 11 Acetonitrile to waste 12 Acetonitrile to column 13 Argon reverse flush 14 Argon block flush 15 Acetonitrile to waste 16 Acetonitrile to column 17 Argon reverse flush 18 Argon block flush Phosphoramidite coupling steps: 19 Phosphoramidite preparation 20 Activator to column 21 Phosphoramidite + activator to column 22 Repeat steps 20 and 21 two times 23 Activator to column 24 Waita 25 Argon reverse flush 26 Argon block flush Column capping steps: 27 Cap A + cap B to column 28 Wait 29 Repeat steps 27 and 28 30 Acetonitrile to waste 31 Argon block flush 32 Acetonitrile to waste 33 Argon reverse flush 34 Argon block flush Oxidation steps: 35 Oxidant to waste 36 Oxidant to column 37 Acetonitrile to waste 38 Argon block flush 39 Wait Column washing steps: 40 Acetonitrile to waste 41 Argon reverse flush 42 Argon block flush 43 Acetonitrile to waste 44 Acetonitrile to column 45 Argon reverse flush 46 Repeat steps 44 and 45 six times 47 Argon block flush Solid-Phase Synthesis of Branched Oligonucleotides
5 45 5 5 1 10 140 30 80 10 5 120 5 5 5 60 5 5 3 5 5 3 5 5 17 45 5 5 5 5 5 5 20 5 5 20 5 10 5 5 18 5 5
aSee Table 4.14.2 for coupling times of various phosphoramidites.
4.14.10 Supplement 9
Current Protocols in Nucleic Acid Chemistry
A
B (i) T10 A
GT9 GT9
i
ii
AU T10
(iii) linear failure sequences
0
10
+
(ii) T10
20
A
30
A
P* GT9 GT9
iii
P*
40 Time (min)
50
60
70
80
D
C (i) CCCUACUAA
(iii) linear failure sequences
GUAUGCCC GUAUGCCC
P* CCCUACUAA GUAUGCCC + (ii) GUAUGCCC CCCUACUAA P*
AU
0
10
20
30
40 Time (min)
E i
i
ii
ii
iii
50
60
70
iii
80
Figure 4.14.6 Analysis of bDNA and bRNA molecules synthesized using the convergent strategy (see Basic Protocol 2). (A-B) Analysis of a successful synthesis of the Y-DNA 5′-T10A2′,5′-GT9 3′,5′-GT9 by (A) anion-exchange HPLC (see Support Protocol 3) using gradient 3 (Table 4.14.4) and (B) 20% denaturing PAGE (see Support Protocol 4). (C-E) Analysis of a successful (C-D) and unsuccessful (E) synthesis of the mixed base Y-RNA 5′-CCCUACUAA2′,5′-GUAUGCCC3′,5′-GUAUGCCC by (C) anion-exchange HPLC (see A for conditions) and (D-E) 20% denaturing PAGE. The regioisomeric extended failure sequences (ii) are resolved into two peaks by HPLC (A and C), but appear as one band by gel analysis (B, D, and E). In panel E, the major product is the unbranched 8-mer 5′-GUAUGCCC-3′, which accumulates due to the unsuccessful branching of the bisphosphoramidite. P* indicates the presence of a 2′- or 3′-linked phosphate.
7. Enter the sequence to be synthesized in the 5′-to-3′ direction, where the 3′-nucleotide corresponds to the nucleoside bound to the CPG. For example, to synthesize the hypothetical V-shaped branched oligonucleotide (S.8) shown in Figure 4.14.4, enter the sequence 5′-XNNNNNN-3′, where X corresponds to the bisphosphoramidite and N is any base (phosphoramidite) of choice. If the hypothetical Y-shaped branched oligonucleotide (S.10) is desired, enter the sequence 5′NNNNNNXNNNNNN-3′.
8. Perform synthesis in the trityl-off mode according to the synthesis cycle outlined in Table 4.14.3 and utilizing the coupling times recommended in Table 4.14.2. Collect dimethoxytrityl solutions in 15-mL test tubes using an external fraction collector. Turning the trityl mode off ensures that the last nucleotide at the 5′ end has a free hydroxyl group, which is desirable for purification using anion-exchange HPLC (see Support Protocol 3).
Synthesis of Modified Oligonucleotides and Conjugates
4.14.11 Current Protocols in Nucleic Acid Chemistry
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9. Upon completion of the synthesis, dry the CPG by manually conducting an argon reverse flush operation on the synthesizer for 10 min. Alternatively, dry the CPG under a stream of nitrogen or argon, or in a vacuum desiccator for 30 min. 10. Cleave the oligonucleotides from the support and deprotect the exocyclic amino, phosphate, and 2′-silyl (RNA only) protecting groups (see Support Protocol 2). 11. Purify the bNAs from failure sequences by anion-exchange HPLC (see Support Protocol 3) or denaturing PAGE (see Support Protocol 4). Typical HPLC and PAGE profiles for the synthesis of branched V-shaped and Y-shaped DNA and RNA molecules are demonstrated in Figures 4.14.6 and 4.14.7.
Measure branching efficiency by trityl color analysis 12. Dilute the dimethoxytrityl solutions collected after each successive coupling (step 8) with 10 mL detritylation solution using a 50-mL buret. 13. Aliquot 100 µL into a quartz cuvette and dilute with 2 mL detritylation solution. Use only quartz cuvettes, as the 1,2-dichloroethane will dissolve disposable polystyrene cuvettes.
14. Measure the absorbance of the solution on a UV-Vis spectrophotometer between 450 and 550 nm, and record the absorbance peak at ∼505 nm for the dimethoxytrityl cation.
XC
d c a
a′ b
b
BPB
1
Solid-Phase Synthesis of Branched Oligonucleotides
2
3
4
Figure 4.14.7 PAGE analysis of bDNA molecules synthesized via the convergent strategy (see Basic Protocol 2), demonstrating the mobilities of bNAs through a cross-linked 20% polyacrylamide gel. Lane 1, crude linear DNA 12-mer 5′-TACTAAGTATGT-3′ (a); lane 2, crude V-DNA 13-mer 5′-A2′,5′-GTATGT3′,5′-GTATGT (c); lane 3, crude Y-DNA 18-mer TACTAA2′,5′-GTATGT3′,5′-GTATGT (d); lane 4, running dyes (xylene cyanol and bromphenol blue). The fastest-migrating sequence in lanes 2 and 3 is the 6-mer 5′-GTATGT-3′ (b), which is the immediate precursor of the V-shaped molecule (c). Notice the large gap between the failure sequence (b) and product V-DNA (c) in lane 2, owing to more than doubling the molecular weight upon branching. In lane 3, extension of the failure sequence yields a separate band (a′).
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15. Determine the efficiency of each coupling step using the equation (Ax/Ax−1) × 100%, where Ax is the absorbance of the trityl cation released at any given step and Ax−1 is the release in the previous step. For efficient coupling, these values should be close to 100% for linear portions of the oligonucleotide (also see UNIT 10.3). For efficient branching, the value for the coupling of S.3 should be 50%, since one tritylated bisphosphoramidite is coupled to two nucleotide chains. For this step, trityl yields that are significantly greater than or less than 50% indicate little branch formation (e.g., 80% indicates the formation of mainly the extended isomeric compounds S.9a-d and S.11a-d).
REGIOSPECIFIC SYNTHESIS OF BRANCHED NUCLEIC ACIDS The protocol described below outlines the regiospecific and divergent synthesis of bNAs via phosphoramidite chemistry according to the method of Braich and Damha (1997; Fig. 4.14.8). This protocol allows for the synthesis of bNA molecules with different DNA sequences surrounding the branchpoint nucleotide. The methodology requires the use of standard DNA and RNA 3′-phosphoramidites (S.4a-d and S.5a-d) for the synthesis of a linear DNA strand incorporating a single ribonucleotide unit. Once this sequence is assembled, all of the 2-cyanoethyl phosphate-protecting groups are selectively removed by treatment with triethylamine. This step is necessary as phosphotriesters are susceptible to cleavage/modification by the ensuing fluoride treatment. The CPG-bound oligomer is then treated with fluoride ions to cleave the tert-butyldimethylsilyl group at the 2′ position of the ribose unit, from which another chain (2′-branch) can be synthesized. This is accomplished using “inverted” DNA phosphoramidites (deoxyribonucleoside 5′-phosphoramidites; S.6a-d), allowing branch synthesis to occur in the opposite (5′-to-3′) direction. With the exception of the decyanoethylation and desilylation steps, the entire process is conducted using an ABI 381A DNA synthesizer.
BASIC PROTOCOL 3
Materials 5′-O-(4,4′-Dimethoxytrityl)-N-protected-2′-deoxyribonucleoside-derivatized succinyl-LCAA-CPG (ChemGenes; also see Support Protocol 1) Cap A and B capping reagents (see recipes) DNA 3′-phosphoramidites (S.4a-d; Chem Genes; Fig. 4.14.3) Anhydrous acetonitrile (see recipe) RNA 3′-phosphoramidite (S.5a-d; ChemGenes; Fig. 4.14.3) Activator solution: 0.5 M 1H-tetrazole (sublimed) in anhydrous acetonitrile Oxidant solution (see recipe) Detritylation solution (see recipe) 4:6 (v/v) triethylamine/acetonitrile (see recipe) Anhydrous THF (see recipe) 1 M tetra-n-butylammonium fluoride (TBAF; Aldrich) in THF, fresh Inverted DNA 5′-phosphoramidites (S.6a-d; ChemGenes; Fig. 4.14.3) Argon or nitrogen gas (optional) Synthesis columns for 1 µmol scale synthesis, with seals and filters (PE Biosystems) and 13-mm aluminum seals (Chromatographic Specialties) ABI 381A automated DNA synthesizer (PE Biosystems) External fraction collector and 15-mL test tubes Empty DNA synthesizer bottles, oven-dried 10- and 1-mL disposable syringes 25-mL glass syringe Additional reagents and equipment for cleaving and deprotecting the oligonucleotide (see Support Protocol 2), anion-exchange HPLC (see Support
Synthesis of Modified Oligonucleotides and Conjugates
4.14.13 Current Protocols in Nucleic Acid Chemistry
Supplement 9
B
DMTrO
O OTBDMS
O
O P OCH2CH2CN O HOD
D D D D D D
D D D D D
DMTrOD
DNA synthesis (3′→5′)
3′-pR amidites
3′-pD amidites
succinylLCAA-CPG
1. DNA synthesis (3′→5′) 2. cap with Ac2O
AcO
AcOD
AcO
D D D D D D O
D D D D D D O
O P O− O
O
O P OCH2CH2CN
O P O−
extend from 2′-OH in 5′→3′ direction with 5′-pD′ amidites
B
D D D D D O B
O
O
O O
O P
O P O− O D D D D D D
OCH2CH2CN
O
OH
O P O−
O D″ D′ D′ D′ D′ D′
D D D D D D
1. cleave from support and deprotect 2. purify (3′-5′)
R
1. remove all 2cyanoethyl groups 2. remove 2′-OTBDMS group
O
B O O
OTBDMS
O P OCH2CH2CN
O
O
D D D D D D
D D D D D D
12
(2′-5′)
D″ D′ D′ D′ D′ D′
D D D D D D 13
Solid-Phase Synthesis of Branched Oligonucleotides
Figure 4.14.8 Schematic representation for the divergent and regiospecific synthesis of branched DNA. The branching synthon is a standard RNA phosphoramidite (S.5a-d; Fig. 4.14.3). Abbreviations: 3′-pD, DNA 3′-phosphoramidites (S.4a-d); 5′-pD′, inverted DNA 5′-phosphoramidites (S.6ad); 5′-pD′′, higher concentration (0.3M) of inverted DNA 5′−phosphoramidite as first nucleotide coupled to the ribose branch point; 3′-pR, RNA 3′-phosphoramidite (S.5a-d); DMTr, 4,4′-dimethoxytrityl; TBDMS, tert-butyldimethysilyl.
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Protocol 3) or denaturing PAGE (see Support Protocol 4), and measuring coupling efficiency by trityl color analysis (see Basic Protocol 2) CAUTION: All solutions required for bNA solid-phase synthesis should be prepared in a well-ventilated fume hood. Synthesize linear DNA (S.12) 1. Prepare synthesis column (1 µmol) with the appropriate 5′-O-dimethoxytrityl-2′-deoxyribonucleoside-derivatized succinyl-LCAA-CPG (see Basic Protocol 2, steps 1 and 2). 2. Weigh out the proper amount of DNA 3′-phosphoramidites (S.4a-d; Fig. 4.14.3) and dilute to 0.1 M with anhydrous acetonitrile (see Table 4.14.2). For synthesis on the ABI 381A DNA synthesizer, the volume of each phosphoramidite addition to the column is 170 ìL.
3. Weigh out the appropriate amount of 3′-RNA phosphoramidite (S.5a-d; Fig. 4.14.3) and dilute to 0.15 M with anhydrous acetonitrile (see Table 4.14.2). The RNA 3′-phosphoramidite is the branching synthon. Any of the four standard RNA 3′-phosphoramidites (A, G, C, or U) may be used depending on the specific branch point to be introduced.
4. Install all reagents (i.e., activator, capping, oxidant, and detritylation solutions, and acetronitrile) and phosphoramidite solutions on the synthesizer, placing the RNA 3′-phosphoramidite on the spare port (the “X” port on the 381A synthesizer). 5. Enter the base sequence of the linear oligonucleotide to be synthesized in the 5′-to-3′ direction, where the last entry (3′ nucleotide) corresponds to the nucleoside bound to the CPG. For example, to synthesize the hypothetical linear oligonucleotide (S.12) shown in Figure 4.14.8, enter the sequence 5′-NNNNNNXNNNNNN-3′, where N is any deoxyribonucleoside phosphoramidite and X represents the branch point of the RNA.
6. Perform synthesis in the trityl off mode according to the synthesis cycle outlined in Table 4.14.3 and utilizing the coupling times shown in Table 4.14.2. Collect the dimethoxytrityl solutions in 15-mL test tubes in an external fraction collector. Turning the trityl mode off ensures that the last nucleotide at the 5′ end has a free hydroxyl group, which is desirable for purification using anion-exchange HPLC (see Support Protocol 3).
7. Acetylate the hydroxyl group at the 5′ terminus by running the automated capping cycle (Table 4.14.1). Capping the free 5′-OH is necessary as it ensures that extension from this functional group will not occur during the synthesis of the “’orthogonal” 2′-branch.
Cleave 2-cyanoethyl protecting group 8. Dry the CPG by manually conducting an argon reverse flush operation on the synthesizer for 10 min. 9. Remove the synthesis column from the synthesizer and connect it to a 10-mL disposable syringe filled with 4:6 (v/v) triethylamine/acetonitrile. Slowly push the deprotection solution through the column over a 90-min period. Deprotection of the 2-cyanoethyl phosphate-protecting group converts the phosphotriester to the more stable phosphodiester, which withstands the conditions required for desilylation in the ensuing step. To ensure complete decyanoethylation, push the solution slowly through the column and then pull in on the syringe slightly in order to displace the CPG beads from the base of the column.
Synthesis of Modified Oligonucleotides and Conjugates
4.14.15 Current Protocols in Nucleic Acid Chemistry
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10. Wash the CPG beads extensively with 30 mL acetonitrile followed by 30 mL THF using a 25-ml glass syringe attached to the column via the syringe adapter. Cleave 2′-O-TBDMS group 11. Push 1 mL of 1 M TBAF in THF through the column over a period of 10 min using a 1-mL disposable syringe. It is essential to use fresh TBAF. In order to ensure complete desilylation, push the solution slowly through the column and then pull in on the syringe slightly in order to displace the CPG beads from the base of the column. Prolonged treatment with TBAF results in cleavage of the oligonucleotide chain from the solid support (Braich and Damha, 1997). Incomplete desilylation results in the accumulation of silylated linear DNA (S.12; Fig. 4.15.8), which does not allow branch extension from the 2′ position of the branch point (see Fig. 4.14.9B).
12. Wash CPG beads with 50 mL THF followed by 50 mL acetonitrile. 13. Reinstall the column on the synthesizer. Synthesize 2′,5′-linked branch (S.13) 14. Modify the DNA synthesis cycle such that synthesis step 15 becomes the first step in the cycle. Steps 1 to 14 (TCA treatment) may be disregarded since the assembled chain lacks a dimethoxytrityl group.
15. Weigh out the appropriate amounts of inverted DNA 5′-phosphoramidites (S.6a-d; Fig. 4.14.3) into the amidite bottles. Add acetonitrile to the first DNA phophoramidite to prepare a 0.3 M solution. Add acetonitrile to make 0.1 M solutions of each of the remaining monomers. The first inverted DNA 5′-phosphoramidite is added as a 0.3 M solution due to steric hindrance around the ribose 2′-hydroxyl group.
16. Install all the inverted phosphoramidites on the synthesizer and place the first inverted phosphoramidite (0.3 M concentration) on the spare port (the “X” port on the 381A). 17. Enter the linear sequence to be synthesized in the 5′-to-3′ direction, where the last entry corresponds to the first phosphoramidite to be coupled to the 2′-hydroxyl of the branch point. For example, to synthesize the hypothetical branched DNA oligonucleotide (S.13) shown in Figure 4.14.8, enter the sequence 5′-NNNNNN-3′, where N is the first DNA 5′-phosphoramidite to be coupled to the 2′-hydroxyl group of ribose.
18. Synthesize the 2′-branch in the trityl off mode using the modified synthesis cycle starting from step 15 of Table 4.14.3, and utilizing the coupling times shown in Table 4.14.2, except for the first phophoramidite (0.3 M), which should have a coupling time of 30 min. 19. Upon completion of the synthesis, dry the CPG by reverse flushing the column with argon for 10 min. Alternatively, dry the CPG under a stream of nitrogen or argon, or in a vacuum desiccator for 30 min. 20. Cleave the oligonucleotides from the support and deprotect the amino- and phosphate-protecting groups (see Support Protocol 2). Solid-Phase Synthesis of Branched Oligonucleotides
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21. Purify the bNAs from failure sequences by anion-exchange HPLC (see Support Protocol 3) or denaturing PAGE (see Support Protocol 4) and measure coupling efficiency by trityl color assay (see Basic Protocol 2, steps 12 to 15). Typical PAGE profiles for the successful and unsuccessful regiospecific synthesis of a branched Y-shaped DNA molecule are demonstrated in Figure 4.14.9.
PREPARATION OF LCAA-CPG SUPPORTS WITH HIGH NUCLEOSIDE LOADINGS
SUPPORT PROTOCOL 1
The method described allows for the rapid derivatization of LCAA-CPG having nucleoside loadings up to ∼90 µmol/g, which is 3 to 4 times the loading found in commercially available solid supports. While commercial samples provide more than adequate yields of bNAs (Damha et al., 1992), those with higher loadings (50 to 90 µmol/g) provide the best results (Fig. 4.14.5). For example, the synthesis of small bNAs (i.e., trimers and tetramers) requires that the CPG be densely loaded so that efficient branching may occur. The protocol below, adapted from the work of Pon et al. (1999) and Damha et al. (1990), allows for the rapid esterification of 5′-O-protected ribonucleosides and deoxyribonucleosides to succinyl-LCAA-CPG. The key condensing reagent is a mixture of HATU and 4-DMAP. Materials 5′-O-(4,4′-Dimethoxytrityl)-N-protected-ribonucleoside or -2′-deoxyribonucleoside (ChemGenes) O-(7-Azabenzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HATU; PE Biosystems) 4-Dimethylaminopyridine (DMAP; 99%, Aldrich)
A (i) A
B
T10 T10
XC
XC (iii) TACTAA (iv) TACTAA
(ii) A
GTATGT CAAGTT Si CAAGTT
Si T10 BPB BPB 1
2
3
4
Figure 4.14.9 20% denaturing PAGE analysis of a successful (A) and a less successful (B) synthesis of bNAs. (A) Lane 1, pure bNA (i) prepared by the divergent synthesis method (see Basic Protocol 3); lane 2, pure bNA (i) prepared by the convergent synthesis method (see Basic Protocol 2); lane 3, crude bNA (i) prepared by the divergent synthesis method; lane 4, running dyes (xylene cyanol and bromphenol blue). (B) Preparative PAGE purification of bNA (iii) prepared via the divergent (regiospecific) approach. The diminutive amount of fully branched product (iii) is due to the incomplete desilylation of the 2′-O-TBDMS group (Si) on the RNA branching synthon, as seen by the significant amount of (iv) that remains.
Synthesis of Modified Oligonucleotides and Conjugates
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Supplement 9
Succinylated long-chain-aminoalkyl controlled-pore glass (succinyl-LCAA-CPG; UNIT 3.2) Anhydrous acetonitrile (see recipe) Dichloromethane, reagent grade (Fisher) Methanol, reagent grade (Fisher) Cap A and B capping reagents (see recipes) 15-mL glass bottle with septum 1-mL syringe and needle Wrist-action shaker Sintered glass funnel or Buchner funnel with filter paper Additional reagents and equipment for quantitation of released trityl groups (UNIT 3.2) and acetylation of support (UNIT 3.2; optional) 1. In a 15-mL glass bottle, combine the following: 0.1 mmol 5′-O-(4,4′-dimethoxytrityl)-N-protected-ribonucleoside or -2′-deoxyribonucleoside 0.1 mmol HATU 12 mg DMAP 250 mg succinyl-LCAA-CPG. 2. Cap the bottle with a septum and add 1 mL anhydrous acetonitrile via a 1-mL syringe and needle. 3. Shake 2 to 4 hr on a wrist-action shaker at room temperature. Do not stir the slurry with a magnetic stir bar as this will break up the glass beads into fine particles that may clog the frit on the DNA synthesizer.
4. Vacuum filter succinyl-LCAA-CPG into a side-arm filter flask through a sintered glass funnel or Buchner funnel with filter paper. 5. Wash the CPG sequentially with 25 mL reagent-grade dichloromethane, 25 mL methanol, and 25 mL dichloromethane. It has been reported that the free carboxylic groups are inconsequential and do not react during phosphoramidite synthesis (Lyttle et al., 1997).
6. Transfer the CPG to a glass vial and dry under vacuum using in a desiccator attached to a vacuum pump. The derivatized CPG may be stored indefinitely at room temperature, preferably in a vacuum desiccator.
7. Determine nucleoside loading through the quantitation of the released trityl groups from the support-bound nucleoside (UNIT 3.2). 8. Acetylate the solid support with cap A and B capping reagents on a DNA synthesizer using the capping cycle outlined in Table 4.14.1. Alternatively, perform capping as described in UNIT 3.2.
Solid-Phase Synthesis of Branched Oligonucleotides
4.14.18 Supplement 9
Current Protocols in Nucleic Acid Chemistry
COMPLETE DEPROTECTION OF BRANCHED OLIGONUCLEOTIDES (DNA AND RNA)
SUPPORT PROTOCOL 2
This protocol describes the steps necessary for cleaving the bNA from the solid support and removing the protecting groups from the heterocyclic bases and sugar-phosphate backbone. The first step is treatment of the solid support with concentrated aqueous ammonia to concomitantly cleave both the bNA from the support and the N-acyl and 2-cyanoethyl phosphate-protecting groups. A subsequent deprotection step with triethylammonium trihydrofluoride (TREAT-HF) cleaves the 2′-O-tert-butyldimethylsilyl (TBDMS) protecting groups from branched oligoribonucleotides (Gasparutto et al., 1992). The desilylated material is then precipitated directly using 1-butanol (Sproat et al., 1995). A procedure for the quantitation of oligonucleotides is also described. Materials Branched oligonucleotide attached to CPG (bNA-CPG; see Basic Protocols 2 and 3) 29% ammonium hydroxide, 4°C (store up to 1 month at 4°C) 70% and 100% (v/v) ethanol, former at 4°C DEPC-treated water (optional; APPENDIX 2A) Autoclaved water Triethylammonium trihydrofluoride (TREAT-HF; 98%; Aldrich) 3 M sodium acetate, pH 5.5 (APPENDIX 2A) 1-Butanol, analytical grade, 4°C 1.5-mL screw-cap microcentrifuge tubes with O-ring seals (preferred) Wrist-action shaker Speedvac evaporator (Savant) Double-beam UV spectrophotometer, calibrated Cleave and deprotect oligonucleotide 1. Transfer bNA-CPG to a 1.5-mL screw-cap microcentrifuge tube, preferably with an O-ring seal. 2. Add 750 µL cold 29% ammonium hydroxide and 250 µL of 100% ethanol, screw the cap on tightly, and incubate 24 to 48 hr at room temperature on a wrist-action shaker. If the branched oligonucleotide sequence contains isobutyryl-protected guanosine nucleotides, then deprotection must proceed for ≥48 hr, room temperature. The ammonium hydroxide should be relatively fresh (<1 month old). Seal the cap on the ammonium hydroxide solution tightly and store at 4°C.
3. Microcentrifuge 1 min at maximum speed, room temperature, and cool 30 min on dry ice or 1 to 2 hr at −20°C. The contents must be cooled before the screw cap is released to prevent volatile ammonium hydroxide from boiling over, which could result in loss of product.
4. Remove supernatant and transfer to a fresh 1.5-mL microcentrifuge tube. 5. Wash CPG pellet with 500 µL of 100% ethanol, microcentrifuge to settle the CPG, and transfer the supernatant wash to a second 1.5-ml tube. Repeat two more times (total four tubes). 6. Cool all four tubes for 1 hr on dry ice, then dry in a Speedvac evaporator. Introducing the vacuum too quickly will lead to bumping of the ammonia solution and sample loss.
Synthesis of Modified Oligonucleotides and Conjugates
4.14.19 Current Protocols in Nucleic Acid Chemistry
Supplement 9
7. Pool in a total of 1 mL water. For branched RNAs, use DEPC-treated water that has been autoclaved to remove residual DEPC. The product is a crude deprotected branched DNA or partially deprotected branched RNA. For RNA, DEPC-treated water should be used in all subsequent steps.
Quantitate oligonucleotide 8. Dilute crude bNA 10 or 100 fold with autoclaved water and measure the absorbance at 260 nm (A260) on a calibrated, double-beam, UV spectrophotometer using autoclaved water as a reference. If using a single-beam spectrophotometer, a baseline should be run with a sample containing autoclaved water (blank).
9. Determine the concentration of the stock solution using the dilution factor and Beer’s Law (A260 = εbc), where ε is the molar extinction coefficient, b is the UV cell pathlength (typically 1 cm), and c is the concentration of oligonucleotide present. The molar extinction coefficients (liter/mol cm) and molecular weights of the oligonucleotides can be obtained from http://paris.chem.yale.edu/extinct.html and http://medstat.med.utah.edu/masspec/oligoii.htm.
Cleave 2′-O-TBDMS group 10. Evaporate bRNA samples (step 7) using a Speedvac evaporator and resuspend in 5 µL TREAT-HF per A260 unit (step 8). 11. Cover the microcentrifuge tubes with aluminum foil and incubate 24 to 48 hr at room temperature on a wrist-action shaker. Alternatively, the 2′-O-TBDMS group may be removed using a mixture of TREAT-HF and N-methylpyrrolidinone according to the method of Wincott et al. (1995).
For large bRNAs (≥10-mers): 12a. Precipitate the oligonucleotides directly from the desilylation reaction by adding 25 µL of 3 M sodium acetate, pH 5.5, followed by 1 mL cold analytical-grade 1-butanol. Vortex 1 min and cool 3 to 6 hr at −20°C. A white precipitate should be clearly visible after addition of 1-butanol.
13a. Microcentrifuge 10 min at maximum speed, 4°C, carefully remove the supernatant, and wash the white pellet twice with 500 µL cold 70% ethanol each. Disruption of the pellet during washing steps could result in loss of sample.
14a. Dry the pellet in a Speedvac evaporator, resuspend in 1 ml autoclaved water, and requantitate the amount of crude oligonucleotide (steps 8 and 9). Once the 2′-O-TBDMS group has been removed, the fully deprotected RNA is sensitive to nucleases and base hydrolysis, which will cause strand cleavage. In order to prevent this, special considerations for working with RNA (APPENDIX 2A) must be observed. Water used for RNA dissolution and buffer preparation should be of the highest quality available (Milli-Q) and should be treated with DEPC.
For small bNAs (<10-mers): 12b. Quench the desilylation reaction with an equal volume of autoclaved water and evaporate in a Speedvac evaporator. Resuspend in 1 mL autoclaved water and requantitate the amount of crude oligonculeotide (steps 8 and 9). Solid-Phase Synthesis of Branched Oligonucleotides
Shorter sequences (i.e., trimers and tetramers) do not precipitate efficiently.
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ANALYSIS AND PURIFICATION OF BRANCHED OLIGONUCLEOTIDES USING ANION-EXCHANGE HPLC
SUPPORT PROTOCOL 3
Branched oligonucleotides may be easily and efficiently analyzed and purified from the crude mixture by anion-exchange HPLC as described below (Fig. 4.14.6). bNAs of high purity are attainable (>95%; Fig. 4.14.10). The bNA of interest can be readily isolated from the reaction mixture failure sequences. The resultant product is precipitated directly by the addition of 1-propanol (Sproat et al., 1995). This direct precipitation method works very well for the direct isolation of large bNAs (>10 nt). Smaller bNAs do not precipitate out efficiently and must be further purified by size-exclusion chromatography (or reversed-phase Sep-Pak cartridges) subsequent to HPLC separation. As an alternative to HPLC purification, the bNAs may be purified by denaturing PAGE (see Support Protocol 4). Characterization of the bNAs is conveniently done via MALDI-TOF-MS as described in UNIT 10.1. The matrix and co-matrix typically used are 6-aza-2-thiothymine (ATT) and dibasic ammonium citrate, respectively (Lecchi et al., 1995). The branched nature of the molecules may also be confirmed via the yeast debranching enzyme (yDBR), a phosphodiesterase specific to hydrolysis of the 2′,5′-phosphodiester bond of oligonucleotides that contain vicinal 2′,5′- and 3′,5′-phosphodiester linkages (Nam et al., 1994; Ooi et al., 2001). Nucleoside composition analysis of bNA is carried out using snake venom phosphodiesterase (SVPD) according to the method of Eadie et al. (1987; UNIT 10.6). This enzyme cleaves bDNA or bRNA from the 3′ termini yielding 5′-monophosphates, which can then be converted to their constituent nucleosides by in situ treatment with alkaline phosphatase (AP). The resulting nucleoside mixture is analyzed by reversed-phase HPLC as described in UNIT 10.5. Alternatively, bNA can be digested with nuclease P1 from Penicillium citrinum, an endonuclease that cleaves bNA to produce the constituent nucleoside 5′-monophosphates and its branch core trinucleoside diphosphate—i.e., A(2′p5′N)3′p5′N (Damha et al., 1992). The released branched trinucleoside diphosphate structure can be readily synthesized (see Basic Protocol 2) and used as a standard during HPLC analysis of the enzyme digest (UNIT 10.6). NOTE: For branced RNAs, use DEPC-treated water throughout (APPENDIX 2A).
UACUAA
GUAUGU GUAUGU
95% purity AU
0.00
10.00
20.00
30.00
40.00 Time (min)
50.00
60.00
70.00
80.00
Figure 4.14.10 Analysis of the purity of a bNA synthesized via the convergent strategy. Purification was conducted using anion-exchange HPLC (see Support Protocol 3) with linear gradient 2 (Table 4.14.4). The chromatogram was obtained using the same conditions.
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Table 4.14.4
Conditions for Separation of bNAs by Anion-Exchange HPLCa
Length of bNA (nt) <5 6-15 >15
Gradient
% Buffer A (H2O)
% Buffer B (1 M LiClO4)
Run time (min)
1
100-90
0-10
60
2 3
100-80 90-80
0-20 10-20
60 60
aPerformed on a Waters 7.5 × 75–mm Protein Pak DEAE-5PW column.
Materials Deprotected branched oligonucleotide (see Support Protocol 2) Autoclaved water 1 M LiClO4 (see recipe) Reagent-grade 1-propanol, 4°C (>5-mers; Fisher) Sephadex G-25 columns (Amersham Pharmacia Biotech) or Sep-Pak cartridges (Waters Chromatography) for <5-mers Nuclease P1 buffer (see recipe) 0.3 U/µL Penicillium citrinum nuclease P1 (NP1) 9 U/µL alkaline phosphatase (AP) 0.1 M triethylammonium acetate (TEAA), pH 7.0 Acetonitrile, HPLC grade 50°C water bath or heating block High-performance liquid chromatograph (HPLC) with: Anion-exchange column (7.5 × 75–mm Waters Protein-Pak DEAE-5PW) UV detector with adjustable range or dual-wavelength detection Sample loop Column heater Reversed-phase C18 column (e.g., Whatman Partisil ODS-2, 10-ìm, 4.6 × 250–mm; Chromatographic Specialties) Syringe Speedvac evaporator (Savant) Additional reagents and equipment for reversed-phase chromatography (UNIT 10.1 & 10.6) and quantitation of bNAs by UV spectrophotometry (see Support Protocol 2) Analyze bNAs by anion-exchange HPLC 1. Aliquot 0.3 to 0.6 A260 units of deprotected branched oligonucleotide (25 to 100 µL volume) into a sterile 1.5-mL microcentrifuge tube. 2. Heat the sample 1 min in a 50°C water bath or heating block. Heating the sample disrupts any intramolecular secondary structures.
3. To settle any particulates, microcentrifuge 2 min at maximum speed, room temperature. It is important to avoid loading any small particulates into the injector of the HPLC as they may clog the injection loop.
4. Preequilibrate HPLC anion-exchange column with initial buffer conditions (Table 4.14.4) and set the UV detector wavelength to 260 nm. Solid-Phase Synthesis of Branched Oligonucleotides
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5. Load the sample into the sample loop using an appropriate syringe and inject. Elute bNA at 50°C with a flow rate of 1 mL/min and the gradient and run time specified in Table 4.14.4. Record the chromatogram. The branched DNA or RNA product should be very well resolved from the extended linear failure sequences and other failure sequences present in the crude mixture. Full-length bNAs elute last (i.e., highest retention time). Typical chromatograms for a 31-mer bDNA and 25-mer bRNA are demonstrated in Figure 4.14.6A and C. Note the excellent separation between the bNA of interest and the failure sequences.
Purify bNAs by anion-exchange HPLC 6. Dissolve 40 to 60 A260 units crude bNA mixture in 1 mL autoclaved water. Heat and microcentrifuge as in steps 2 to 3. Loading >60 A260 units may overload the column and compromise the separation of the bNA from the linear failure sequences.
7. Run the sample as described (steps 4 and 5), but set the detector to 290 nm and collect 1-ml fractions from the peaks of interest in sterile 1.5-mL microcentrifuge tubes. The detector wavelength is set to 290 nm in order to avoid saturation of the detector signal. If the HPLC is equipped with a detector capable of monitoring dual wavelengths, monitor both the 260- and 290-nm profiles. The anticipated retention time should be very similar to that obtained during routine HPLC analysis.
8. Pool peak fractions and dry them in a Speedvac evaporator. Add 250 µL autoclaved water. For large bRNAs (≥5-mers): 9a. Precipitate from perchlorate salts (in sample) by adding 4 vol (1 mL) reagent-grade cold 1-propanol and cooling 4 to 6 hr at −20°C. Lithium perchlorate (LiClO4) is much more soluble in organic solvents than other perchlorate salts, making precipitation easy and efficient, and thus preventing a final desalting step. The DNA or RNA isolated is in its lithium salt form.
10a. Microcentrifuge 10 min at maximum speed, room temperature, carefully remove the supernatant, and wash the white pellet twice with 500 µL cold 1-propanol. Disrupting the pellet during washing steps can result in loss of sample.
11a. Dry in a Speedvac evaporator, resuspend in 1 mL autoclaved water, and proceed to step 12. For small bNAs (<5-mers): 9b. Desalt by size-exclusion chromatography on Sephadex G-25 columns according to manufacturer’s instructions or by reversed-phase chromatography on Sep-Pak cartridges (UNIT 10.1). Proceed to step 12. Small bNAs do not precipitate out of solution efficiently.
Characterize bNAs 12. Quantitate the bNA sample by UV spectrophotometry (see Support Protocol 2, steps 8 and 9). 13. Dissolve 0.5 A260 units bNA in 20 mL of nuclease P1 buffer. 14. Add 3 µL (0.9 U) NP1 and 1 µL (9 U) AP. Mix well and incubate 24 hr at 37°C. The alkaline phosphatase may be contaminated with adenosine deaminase, which converts adenosine into inosine. The retention time of an appropriate inosine nucleoside control should be obtained prior to HPLC analysis of the digestion mixture (UNIT 10.6).
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15. Dry the sample in a Speedvac evaporator and dissolve residue in 15 µL autoclaved water. 16. Analyze the mixture by reversed-phase HPLC on a C18 column using the mobile phases 0.1 M TEAA, pH 7.0, and acetonitrile with the gradient described in UNIT 10.6. 17. Calculate the relative ratios of nucleoside to branched trinucleotide diphosphate by dividing the area of each peak by the extinction coefficients specified in UNIT 10.6. The extinction coefficient for the branched trinucleotide diphosphate may be calculated using the oligonucleotide extinction coefficient calculator at http://paris.chem.yale.edu/ extinct.html. The extinction coefficients for bDNA and bRNA are assumed to be the same as their linear counterparts. SUPPORT PROTOCOL 4
ANALYSIS AND PURIFICATION OF BRANCHED OLIGONUCLEOTIDES BY DENATURING PAGE A method for the analysis and purification of bNAs by denaturing polyacrylamide gel electrophoresis (PAGE) is described (also see UNIT 10.4 and APPENDIX 3B). PAGE is a very convenient way to assess efficiency of bNA synthesis since the molecular weight of oligonucleotides bound on the solid support more than doubles after a convergent branching reaction—e.g., reaction of neighboring decathymidylic acid chains with bisphosphoramidite synthon S.3 to give a 21-unit-long bNA molecule (Fig. 4.14.4 and product ii in Fig. 4.14.6B). This generates a gap between the desired bNA product and its precursor molecules, greatly facilitating the separation process. Any type of standard laboratory electrophoresis equipment may be utilized. Most bNAs of >10 nucleotides in length are very well resolved on a 20% denaturing polyacrylamide gel. If shorter sequences must be purified, better resolution is achieved with a 24% denaturing polyacrylamide gel. Setup and polymerization of the gel along with electrophoretic separation conditions are described. The resultant bands may be visualized by UV shadowing and photographed. A technique for the rapid extraction of oligonucleotides from the gel matrix is also described (Chen and Ruffner, 1996). Materials 20% (w/v) denaturing acrylamide gel solution (see recipe) Deprotected branched oligonucleotide sample, dry (see Support Protocol 2) Formamide loading buffer (see recipe) Gel extraction buffer (see recipe) Running dye (see recipe)
Solid-Phase Synthesis of Branched Oligonucleotides
Gel electrophoresis equipment (APPENDIX 3B) with: 16 × 18–cm glass plates Spacers: 0.75 mm (analysis) or 1.5 mm (purification) Gel combs: 0.75 mm thick with 12 to 20 wells (analysis) or 1.5 mm thick with one to three large wells (purification) Sonicator or ∼50°C water bath (optional) 20 × 20–cm silica-coated thin-layer chromatography (TLC) plate with fluorescent indicator (e.g., Kieselgel 60 F254 aluminum sheets) Handheld UV lamp (254 nm) Camera equipped with UV filter (optional) 90°C water bath or heating block (optional) UV shadow box Wrist-action shaker (optional) Speedvac evaporator (Savant) Sephadex G-25 (Amersham Pharmacia Biotech) or reversed-phase cartridges (e.g., Sep-Pak cartridges; Waters Chromatography)
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Additional reagents and equipment for denaturing PAGE (UNIT 10.4 and APPENDIX 3B), reversed-phase chromatography using Sep-Pak cartridges (UNIT 10.1), UV spectrophotometry (see Support Protocol 2), MALDI-TOF-MS (UNIT 10.1), and enzymatic digestion (see Support Protocol 3) CAUTION: Acrylamide and N,N′-methylenebisacrylamide are neurotoxins. Prepare all solutions containing these two reagents in a fume hood. Minimize exposure and contact to both crystalline forms and solutions by conducting all handling (including weighing) in a well-ventilated area and wearing disposable gloves at all times. Analyze bNAs by denaturing PAGE 1. Assemble a gel sandwich using 16 × 18–cm glass plates separated by 0.75-mm spacers. See APPENDIX 3B and UNIT 10.4 for a thorough description of denaturing PAGE.
2. Transfer 30 mL of 20% denaturing acrylamide gel solution to an Erlenmeyer flask and add 200 µL fresh 10% (w/v) APS immediately followed by 20 µL TEMED. Swirl and degas the solution by attaching the Erlenmeyer flask to a house vacuum line. Maintain rapid stirring to avoid bumping. The gel solution may also be degassed prior to APS and TEMED addition by placing the 20% acrylamide gel solution on a sonicator for 5 to 10 min. If using larger glass plates, adjust the volumes of acrylamide, APS, and TEMED accordingly.
3. Pour the solution between the plates, insert a 0.75-mm-thick comb, and allow gel to polymerize (30 to 45 min). 4. Place gel in electrophoresis apparatus, removing the comb and bottom spacer, rinse the wells, and prerun the gel 30 min at 500 V, room temperature. 5. Dissolve 0.6 A260 units deprotected bNA sample in 10 µL formamide loading buffer. If the dissolution process is not immediate, place the sample in a sonicator bath 1 to 2 min or heat the samples briefly at ∼50°C. 6. Load the samples into the wells. Load an equal amount of running dye in the first and last well as an external reference marker. Run the gel at 500 V until the bromphenol blue dye is 3⁄4 of the way down the gel. 7. Disassemble the glass plates and wrap the gel in plastic wrap. Place the wrapped gel over a 20 × 20–cm silica-coated TLC plate with fluorescent indicator and visualize the bands by shining a handheld UV lamp over the gel. Take a picture of the gel using a camera equipped with a UV filter. CAUTION: Wear safety glasses to avoid eye burn. A typical crude bNA reaction mixture will consist of at least three bands as shown in Figures 4.14.6 and 4.14.7. The fastest-migrating band is the linear precursor sequence (S.7; Fig. 4.14.4), followed by the extended isomeric failure sequences (S.9a-b for the synthesis of V-shaped bNAs and S.11a-b for the synthesis of Y-shaped bNAs), and finally the products (S.8 and S.10). If the coupling efficiencies between successive nucleotides is less than optimal, a ladder of failure sequences will be evident below the extended branch failure and linear sequences. Note that the extended isomeric failure sequences are a mixture of regioisomers (S.9a-b and S.11a-b) that are sometimes resolved into two close-moving bands (this is also evident by HPLC analyses). The slowest of the predominant bands is the bNA of interest. If linear markers are run alongside the purified bNA, one observes that bNA has retarded mobility relative to a linear oligonucleotide of identical composition (length and sequence composition). This is due to the increased frictional effects of the bNA relative to the linear sequences as they move through the highly cross-linked gel environment.
Synthesis of Modified Oligonucleotides and Conjugates
4.14.25 Current Protocols in Nucleic Acid Chemistry
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Purify bNAs by denaturing PAGE 8. Assemble and run a preparative gel as for the analytical gel (steps 1 to 6) with the following modifications: a. Increase gel thickness (spacers and comb) to 1.5 cm and scale up volume of gel to 50 mL gel solution, 350 µL APS, and 35 µL TEMED. b. Use up to 100 A260 units crude bNA dissolved in 100 µL formamide loading buffer. c. Use a comb with one large well to purify 60 to 100 A260 units, one well of a two-well comb for 30 to 60 units, and one well of a three-well comb for <30 units. 9. Disassemble the glass plates, remove the gel, and wrap it in plastic wrap. Place the wrapped gel over a silica-coated TLC plate and visualize the bands by shining UV light over the gel. If desired, photograph the gel using a camera equipped with a UV filter. CAUTION: Wear safety glasses to avoid eye burn.
10. Excise the slowest moving band and place the gel piece into a sterile 15-mL tube. Crush the gel piece using a sterile spatula and soak in 3 mL gel extraction buffer. 11. Heat 5 min in a 90°C water bath or heating block and rapidly freeze 5 min in dry ice. Thaw contents rapidly at 90°C and centrifuge 10 min at maximum speed × g in a tabletop centrifuge to settle the crushed gel pieces. Alternatively, after extracting in 3 mL gel extraction buffer (step 10) or water, shake the slurry 12 to 16 hr at room temperature on a wrist-action shaker. The rapid “crush and soak” method described has been outlined in a paper by Chen and Ruffner (1996).
12. Extract the supernatant and dry in a Speedvac evaporator. Desalt on a Sephadex G-25 column according to manufacturer’s instructions, or via reversed-phase chromatography using Sep-Pak cartridges according to UNIT 10.1. 13. Quantitate the amount of bNA recovered via UV spectrophotometry (see Support Protocol 2, steps 8 and 9). 14. Characterize bNAs by MALDI-TOF-MS (UNIT 10.1) and/or enzymatic digestion of the constituent nucleotides with SVPD/AP and NP1/AP (see Support Protocol 3). REAGENTS AND SOLUTIONS Use deionized, double-distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Acetonitrile, anhydrous Predry by refluxing over reagent-grade phosphorus pentoxide (Fisher) followed by refluxing and distillation over calcium hydride (Aldrich) under inert atmosphere. Prepare fresh before each use. Alternatively, low-water acetonitrile suitable for use on a DNA synthesizer can be purchased (VWR Canlabs or BDH) and used as such, or refluxed and distilled from calcium hydride under inert atmosphere.
Cap A capping reagent Prepare anhydrous 2,4,6-collidine (Aldrich) by refluxing and distillation over calcium hydride. Store up to 6 months at room temperature under inert atmosphere over activated 4A molecular sieves. Before use mix 1 part dry 2,4,6-collidine with 1 part acetic anhydride and 8 parts anhydrous THF (see recipe). Solid-Phase Synthesis of Branched Oligonucleotides
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Cap B capping reagent Prepare anhydrous N-methylimidazole (Aldrich) by stirring over calcium hydride for 24 hr, followed by vacuum distillation. Dilute to 16% (v/v) in anhydrous THF (see recipe). Store up to 6 months at room temperature under inert atmosphere over activated 4A molecular sieves. Denaturing acrylamide gel solution, 20% (w/v) 20 g acrylamide 1 g N,N′-methylenebisacrylamide 21.02 g electrophoresis-grade urea (final 7 M) 5 mL 10× TBE electrophoresis buffer (APPENDIX 2A) H2O to 50 mL Store up to 1 to 2 months at 4°C CAUTION: Acrylamide and N,N′-methylenebisacrylamide are neurotoxins. Prepare all solutions in a well-ventilated fume hood, and take precautions to minimize exposure. Wear gloves at all times. To aid in dissolution, the acrylamide solution can be heated in a warm (<50°C) water bath with constant stirring. Do not overheat as this will cause the acrylamide to polymerize.
Detritylation solution 3% (w/v) trichloroacetic acid (TCA, 99%, Aldrich) 1,2-Dichloroethane (analytical grade, Fisher) Store up to 6 months at room temperature Diisopropylethylamine (DIPEA), anhydrous Dry by mild heating (50° to 60°C) and stirring 16 hr over calcium hydride followed by distillation under vacuum. Store up to 2 to 4 months at 4°C over 4A activated molecular sieves in septum-sealed bottles. Alternatively, 99.5% redistilled DIPEA can be purchased (Aldrich) and kept anhydrous by storage over activated 4A molecular sieves up to 1 year at 4°C.
Formamide loading buffer Deionize 10 mL formamide by adding 1 g mixed bed resin (Bio-Rad) and stirring 30 min. Filter through Whatman filter paper. Mix 4 vol deionized formamide with 1 vol of 10× TBE electrophoresis buffer (APPENDIX 2A). Store up to 1 year at −20°C. Gel extraction buffer 363 mg Tris base (final 30 mM) 1.75 g NaCl (final 300 mM) 112 mg disodium EDTA (final 3 mM) H2O to 100 mL Store up to 6 months at 4°C pH to 7.5 Lithium perchlorate (LiClO4), 1 M Dissolve 106.4 g LiClO4 (99%, Aldrich) in 1 liter water. Store up to 1 month at room temperature. Filter through a 0.45-µm filter before use. Nuclease P1 buffer 1.21 g Tris base (final 0.1 M Tris⋅Cl) 13.6 mg ZnCl2 (final 1 mM) H2O to 100 mL Adjust pH to 7.2 with HCl Store up to 6 months at −20°C
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Oxidant solution 1.23 g iodine crystals (0.1 M) 25:20:2 (v/v/v) tetrahydrofuran/pyridine/water Store up to 6 months at room temperature Running dye 9.8 mL deionized formamide 0.2 mL 10× TBE electrophoresis buffer (APPENDIX 2A) 10 mg xylene cyanol (final 0.1%) 10 mg bromphenol blue (final 0.1%) Store up to 1 year at −20°C Tetrahydrofuran (THF), anhydrous Dry by continuous reflux and distillation over sodium metal and benzophenone under an inert atmosphere until a purple color persists. Prepare fresh before each use. Triethylamine/acetonitrile, 4:6 (v/v) Dry triethylamine by refluxing and distillation over calcium hydride. Store up to 1 year at 4°C over activated 4A molecular sieves. Mix 8 mL anhydrous triethylamine and 12 mL anhydrous acetonitrile (see recipe) in a septum bottle and seal. Prepare fresh. To use, withdraw via syringe through the septum. COMMENTARY Background Information
Solid-Phase Synthesis of Branched Oligonucleotides
Discovery and applications of bNAs The body of work devoted to branched nucleic acids (bNAs) indicates that they are of current interest in chemistry and biochemistry. Branched ribonucleic acids (bRNAs) were first detected in nuclear RNA in 1983 (Wallace and Edmonds, 1983). It was later discovered that an RNA “lariat” is the first biochemical intermediate in the splicing of precursor messenger RNA (Ruskin et al., 1984). Several other bRNA structures such as Y-shaped trans-splicing intermediates and multicopy single-stranded DNAs (msDNA) were subsequently discovered in eukaryotes and prokaryotes, respectively (Yee et al., 1984; Inouye et al. 1987). These branched molecules all contain a common structural feature, namely vicinal 2′,5′and 3′,5′-phosphodiester bonds at a branch point nucleotide. A novel polypeptide termed the RNA lariat debranching enzyme selectively hydrolyzes the 2′,5′-phosphodiester bond at the branch point of excised intron lariats, thus converting them into linear molecules (Ruskin and Green, 1985; Chapman and Boeke, 1991; Nam et al., 1994). The novelty of bRNAs has raised interest as to their possible role in regulating RNA processing. Recently, there has been increasing interest in synthetic bDNA and bRNA for use in
diagnostic applications (Urdea et al., 1991) and biosensor development (Uddin et al., 1997), as tools for studying mRNA splicing (Carriero et al., 2001) and debranching (Nam et al., 1994; Ooi et al., 2001), and as “molecular anchors” for inducing the formation of novel triplex (e.g., Hudson et al., 1995) and tetraplex structures (Robidoux et al., 1997). Chemical synthesis of bRNA The stepwise assembly of the vicinal 2′,5′and 3′,5′-internucleotide linkages present in bRNA via traditional phosphotriester methods can be difficult, as cyclic phosphate may be formed (see Damha et al., 1992, and references therein). This problem was circumvented by using phosphodiester intermediates, which are more stable to nucleophilic attack by vicinal 2′-OH groups relative to phosphotriesters. Short bRNA molecules were first prepared by solution-phase methods. Ogilvie and coworkers originally prepared branched tri- and tetranucleotides having identical nucleotides at the branch point 2′ and 3′ positions (symmetric bRNA; Damha et al., 1985; Damha and Ogilvie, 1988). Their method was also adapted to the synthesis of bRNA isomers having different nucleotide units at the 2′ and 3′ positions (Damha and Ogilvie, 1988). Regiospecific synthesis of a tetranucleotide of natural branch point sequence was performed by Kierzek et al.
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(1986) by solution-phase phosphoramidite chemistry. Chattopadhyaya and co-workers have focused on several strategies for the synthesis of small bRNA and RNA lariats with self-cleavage properties (Rousse et al., 1994). The authors’ group has been describing in detail a methodology for the solid-phase synthesis of Y-shaped RNAs and DNAs, in which the branch point adenosine is linked to identical oligonucleotide chains via vicinal 2′,5′- and 3′,5′-phosphodiester linkages. Hyperbranched or “dendritic” structures have also been synthesized from this laboratory (Hudson and Damha, 1993; Hudson et al., 1998). The synthetic convergent strategy is based on the discovery that nucleoside 2′,3′-O-bisphosphoramidite synthons react with adjacent solid-support-bound chains yielding symmetric V-like molecules. Synthesis is then continued in the 3′-to-5′ direction from the apex of the V to yield Y-shaped structures. This method was adapted to the synthesis of bRNAs with 2′ and 3′ chains of different base composition (Ganeshan et al., 1995). All other synthetic strategies reported so far for the regiospecific assembly of branched oligonucleotides mimicking the natural lariat structures are based upon solution-phase phosphotriester methods, except for the recent work by Sproat et al. (1994), who use a divergent solid-phase phosphoramidite strategy. This strategy has permitted the synthesis of mediumsized branched oligoribonucleotides; however, it requires the use of nonstandard monomers and branch-point synthons. Given this limitation, and those of the authors’ current methods, alternative strategies for the regiospecific synthesis of branched oligonucleotides were sought. This led ultimately to the synthesis of multi-copy single-stranded DNA, a peculiar nucleic acid that is produced in E. coli and M. xantus (Damha and Braich, 1998). In this unit, general procedures for both convergent and divergent solid-phase synthesis of branched oligonucleotides are presented. The conditions of the branching reactions are crucial to the ultimate success of the overall procedure, and the protocols described in this unit should be followed carefully. In spite of these areas of caution, bNA sequences can now be prepared routinely using, for the most part, commercially available reagents and standard laboratory equipment. The novice should have no hesitation in setting forth!
Compound Characterization Analysis of crude and purified bNAs is conveniently carried out via anion-exchange
HPLC (see Support Protocol 3). The buffer concentration required for the elution of bNA is dependent upon the molecular weight and charge of the molecule being purified (Table 4.14.4). Separation of bNA from failure sequences is greatly facilitated by the relatively large molecular weight of the bNA species. Gel electrophoresis is another convenient way to isolate/purify bNA fragments (see Support Protocol 4). In this case, the bNA of interest exhibits significant retarded mobility relative to the failure sequences as a result of frictional effects as the bNA molecules migrate through the cross-linked gel environment. Further characterization is conducted by MALDI-TOF-MS (UNIT 10.1) using 6-aza-2-thiothymine (ATT) and dibasic ammonium citrate as matrices (Lecchi et al., 1995). Nucleoside composition analysis is conducted via enzymatic hydrolysis with the exonuclease snake venom phosphodiesterase (Eadie et al., 1987; UNIT 10.6) and the endonuclease nuclease P1 (see Support Protocol 3). The yeast debranching enzyme (yDBR), a phosphodiesterase specific to the 2′,5′-phosphodiester bond of oligonucleotides containing vicinal 2′,5′- and 3′,5′-phosphodiester linkages, may also be used to confirm the branched nature of the molecules according to the method of Ooi et al. (2001).
Critical Parameters and Troubleshooting Basic Protocol 1 The synthesis of the branch synthon (S.3) from 5′-O-DMTr-N6-benzoyl-riboadenosine is extremely moisture sensitive. Thus, it is imperative that all solvents, reagents, and apparatuses (e.g., syringes, needles, reaction vessel) be anhydrous. The phosphitylating reagent (stored at −20°C) should be warmed to room temperature in a desiccator prior to use. If the isolated yield of S.3 is significantly lower than expected, verify that the phosphitylating chlorophosphoramidite is a clear viscous liquid, and that no crystalline material is present (this residue is indicative of hydrolysis). Basic Protocol 2 There are two important considerations when synthesizing bNA by the convergent approach. Firstly, the yield of bNA is dependent upon the molar concentration of the branching bisphosphoramidite synthon S.3. Dilute solutions (0.03 M) of S.3 must be employed to maximize coupling with adjacent CPG-bound oligonucleotide chains (branching). If higher
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concentrations are used, yields of branched products are reduced, and mainly extended isomeric side products are formed. The efficiency of the branching reaction can be assessed during synthesis by quantitation of the trityl cations released immediately before and after the branching reaction. Theoretically, 100% branch formation should give an apparent coupling yield of 50% since the trityl absorption following branching should be half of the previous absorption (during branching two nucleotide chains become joined to a single bisphosphoramidite synthon). Thus, trityl yields significantly greater or less than 50% indicate little branch formation (e.g., 80% apparent trityl yield indicates the formation of mainly the extended isomeric compounds). It is important to prepare the bisphosphoramidite stock solution (0.03 M) using at least 100 mg S.3. This ensures that (unavoidable) traces of moisture will not use up significant amounts of bisphosphoramidite during the branching step and reduce the overall yield of synthesis. The purity of the bisphosphoramidite can be assessed prior to use via TLC analysis or 31PNMR. If partial decomposition has occurred, or if the branching reactions during synthesis are poor, the compound should be rechromatographed as described (see Basic Protocol 1). Secondly, solid supports having a high degree of derivatization (>50 µmol/g) provide the best results for the synthesis of short bNA fragments (<7 nt). This is because highly substituted supports ensure the appropriate distance between the reactive 5′-OH end groups of the immobilized oligonucleotide chains. Supports with a nucleoside loading of 30 to 50 µmol/g give very good results for the synthesis of medium-size branched oligonucleotides.
Solid-Phase Synthesis of Branched Oligonucleotides
Basic Protocol 3 The divergent approach requires synthesis of a linear oligonucleotide chain followed by backward synthesis from the 2′-hydroxyl of an internal ribonucleotide residue. This is carried out with commercially available inverted nucleoside 5′-phosphoramidites. Key to the success of this synthesis is the ability to cleave the 2′-O-silyl protecting group of the internal ribonucleotide residue and the use of an excess of the first inverted 5′-phosphoramidite reagent to force the branching reaction to a maximal extent. This is in sharp contrast to the convergent strategy, where branching requires a delicate control of reaction conditions. Incomplete removal of the 2′-O-TBDMS group from the RNA branch point (Fig. 4.14.8)
results in the accumulation of the linear DNARNA-DNA precursor (S.12), which is easily detected by HPLC and PAGE. To ensure complete 2′-O-desilylation of the CPG-bound oligonucleotide, it is essential to use fresh TBAF reagent and to introduce the TBAF solution slowly to the synthesis column (via syringe), pushing and pulling the piston slightly to displace the CPG beads stuck at the base of the column. This allows all of the CPG-bound oligonucleotide to come in contact with the TBAF reagent. It is important not to expose the CPG-bound bNA to the TBAF reagent for >10 min, as prolonged treatment results in significant cleavage of the oligonucleotide from the solid support. After washing steps, the first inverted phosphoramidite is introduced as a 0.3 M acetonitrile solution (as opposed to 0.1 M for all other DNA 3′- and 5′-phosphoramidite couplings) and allowed to react with the CPGbound oligomer for 30 min (as opposed to 90 and 120 sec for all other DNA phosphoramidites).
Anticipated Results The anticipated results of a convergent bNA synthesis are provided in Figures 4.14.6 and 4.14.7. Syntheses carried out on a 1-µmol scale will characteristically yield 50 to 80 A260 units crude material, while the isolated yields (bNA) generally fall in the range of 5 to 20 A260 units. bNAs that have been purified by anion-exchange HPLC are of high purity, usually >95% (Fig. 4.14.10). In the case of divergent bNA synthesis, typical crude yields are in the range of 50 to 100 A260 units. A standard gel analysis of bNA prepared by the divergent method is shown in Fig. 4.14.9. After HPLC or gel purification, 5 to 20 A260 units of bNA are typically recovered from divergent syntheses (slightly higher yields are obtained with HPLC). As for convergent synthesis, purity is >95%.
Time Considerations Provided that all reagents and materials required for each step are available, most of the procedures are simple and rapid. The synthesis of S.3 from 5′-DMTr-N6-benzoyl-riboadenosine requires 2 to 4 hr to complete, including the workup. The reaction should not be allowed to proceed overnight, as decomposition may occur. Column chromatography requires ∼1 to 2 hr including setup. Ideally, column purification should be conducted immediately following the phosphitylation reaction. When the branching synthon S.3 and all the other phosphoramidite derivatives are ready to
4.14.30 Supplement 9
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use, the time required to prepare, purify, and isolate a Y-shaped RNA or DNA via the convergent approach is 3 days and 4 to 6 days, respectively. The preparation of bNA via the divergent approach requires 3 days or more.
Literature Cited Braich, R.S. and Damha, M.J. 1997. Regiospecific solid-phase synthesis of branched oligonucleotides. Effect of vicinal 2′,5′- (or 2′,3′-) and 3′,5′-phosphosdiester linkages on the formation of hairpin DNA. Bioconjugate Chem. 8:370377. Carriero, S., Braich, R.S., Hudson, R.H.E., Anglin, D., Friesen, J.D., and Damha, M.J. 2001. Inhibition of in vitro pre-mRNA splicing in S. cerevisiae by branched oligonucleotides. Nucleosides, Nucleotides, and Nucl. Acids. 20:873-877. Chapman, K.B. and Boeke, J.D. 1991 Isolation and characterization of the gene encoding yeast debranching enzyme. Cell 65:483-492. Chen, Z. and Ruffner, D.E. 1996. Modified crushand-soak method for recovering oligodeoxynucleotides from polyacrylamide gel. BioTechniques 21:820-822. Damha, M.J., and Braich, R.S. 1998. Synthesis of branched DNA/RNA chimera similar to the msDNA molecule of Myxoccus xanthus. Tetrahedron lett. 39:3907-3910. Damha, M.J. and Ogilvie, K.K. 1988. Synthesis and spectroscopic analysis of branched RNA fragments: Messenger RNA splicing intermediates. J. Org. Chem. 53:3710-3722. Damha, M.J. and Zabarylo, S.V. 1989. Automated solid-phase synthesis of branched oligonucleotides. Tetrahedron Lett. 30:6295-6298. Damha, M.J., Pon, R.T., and Ogilvie, K.K. 1985. Chemical synthesis of branched RNA: Novel trinucleoside diphosphates containing vicinal 2′5′ and 3′-5′ phosphodiester linkages. Tetrahedron Lett. 26:4839-4842. Damha, M.J., Giannaris, P.A., and Zabarylo, S.V. 1990. An improved procedure for derivatization of controlled-pore glass beads for solid-phase oligonucleotide synthesis. Nucl. Acids Res. 18:3813-3821. Damha, M.J., Ganeshan, K., Hudson, R.H.E., and Zabarylo, S.V. 1992. Solid-phase synthesis of branched oligoribonucleotides related to messenger RNA splicing intermediates. Nucl. Acids Res. 20:6565-6573. Eadie, J.S., McBride, L.J., Efcavitch, J.W., Hoff, L.B., and Cathcart, R. 1987. High-performance liquid chromatographic analysis of oligodeoxyribonucleotide base composition. Anal. Biochem. 165:442-447. Ganeshan, K., Tadey, T., Nam, K., Braich, R., Purdy, W.C., Boeke, J.D., and Damha, M.J. 1995. Novel approaches to the synthesis and analysis of branched RNA. Nucleosides & Nucleotides 14:1009-1013.
Gasparutto, D., Livache, T., Bazin, H., Duplaa, A.M., Guy, A., Khorlin, A., Molko, D., Roget, A., and Teoule, R. 1992. Chemical synthesis of a biologically active natural tRNA with its minor bases. Nucl. Acids Res. 20:5159-5166. Hakimelahi, G.H., Proba, Z.A., and Ogilvie, K.K. 1982. New catalysts and procedures for the dimethoxytritylation and selective silylation of ribonucleosides. Can. J. Chem. 60:1106-1113. Hudson, R.H.E. and Damha, M.J. 1993. Nucleic acid dendrimers: Novel biopolymer structures. J. Am. Chem. Soc. 115:2119-2124. Hudson, R.H.E., Uddin, A.H., and Damha, M.J. 1995. Association of branched nucleic acids: Structural and physicochemical analysis of antiparallel TAT triple-helical DNA. J. Am. Chem. Soc. 117:12470-12477. Hudson, R.H.E., Robidoux, S., and Damha, M.J. 1998. Divergent synthesis of nucleic acid dendrimers. Tetrahedron Lett. 32:1299-1302. Inouye, S., Furuichi, T., Dhundle, A., and Inouye, M. 1987. Molecular Biology of RNA: New Perspectives (M. Inouye and B. S. Dudock, eds.) pp. 271. Academic Press, San Diego. Kierzek, R., Kopp, D.W., Edmonds, M., and Caruthers, M.H. 1986. Chemical synthesis of branched RNA. Nucl. Acids Res. 14:4751-4764. Lecchi, P., Le, H.M.T., and Pannell, L.K. 1995. 6-Aza-2-thiothymine: A matrix for MALDI spectra of oligonucleotides. Nucl. Acids Res. 23:1276-1277. Lyttle, M.H., Adams, H., Hudson, D., and Cook, R.M. 1997. Versatile linker chemistry for synthesis of 3′-modified DNA. Bioconjugate Chem. 8:193-198. Nam, K., Hudson, R.H.E., Chapman, K.B., Ganeshan, K., Damha, M.J., and Boeke, J.D. 1994. Yeast lariat debranching enzyme: Substrate and sequence specificity. J. Biol. Chem. 269:20613-20621. Ooi, S.L., Dann, C. III, Nam, K., Leahy, D., Damha, M.J., and Boeke, J.D. 2001. Ribonucleases part A: Functional roles and mechanisms. Methods Enzymol. 342:233-250. Pon, R.T., Yu, S., and Sanghvi, Y.S. 1999. Rapid esterification of nucleosides to solid-phase supports for oligonucleotide synthesis using uronium and phosphonium coupling reagents. Bioconjugate Chem. 10:1051-1057. Robidoux, S., Klinck, R., Gehring, K., and Damha, M.J. 1997. Association of branched oligonucleotides into the i-motif. J. Biomol. Struct. Dyn. 15:517-527. Rousse, B., Puri, N., Viswanadham, G., Agback, P., Glemarec, C., Sandstroem, A., Sund, C., and Chattopadhyaya, J. 1994. Solution conformation of hexameric and heptameric lariat-RNAs and their self-cleavage reactions which give products mimicking those from some catalytic RNAs (ribozymes). Tetrahedron 50:1777-1810. Ruskin, B. and Green, M. 1985. An RNA processing activity that debranches RNA lariats. Science 229:135-140.
Synthesis of Modified Oligonucleotides and Conjugates
4.14.31 Current Protocols in Nucleic Acid Chemistry
Supplement 9
Ruskin, B., Krainer, A.R., Maniatis, T., and Green, M.R. 1984. Excision of an intact intron as a novel lariat structure during pre-mRNA spicing in vitro. Cell 38:317-331. Sproat, B.S., Beijer, B., Grotli, M., Ryder, U., Morand, K.L., and Lamond, A.I. 1994. Novel solidphase synthesis of branched oligoribonucleotides including a substrate for the RNA debranching enzyme. J. Am. Chem. Soc. Perkin. Trans. 1:419-431. Sproat, B., Colonna, F., Mullah, B., Tsou, D., Andrus, A., Hampel, A., and Vinayak, R. 1995. An efficient method for the isolation and purification of oligoribonucleotides. Nucleosides & Nucleotides 14:255-273. Still, W.C., Kahn, M., and Mitra, A. 1978. Rapid chromatographic technique for preparative separations with moderate resolution. J. Org. Chem. 43:2923-2925. Ti, G.S., Gaffney, B.L., and Jones, R.A. 1982. Transient protection: Efficient one-flask syntheses of protected deoxynucleosides. J. Am. Chem. Soc. 104:1316-1319. Uddin, A.H., Piunno, P.A.E., Hudson, R.H.E., Damha, M.J., and Krull, U.J. 1997. A fiber optic biosensor for fluorimetric detection of triplehelical DNA. Nucl. Acids Res. 25:4139-4146. Urdea, M.S., Horn, T., Fultz, T.J., Anderson, M., Running, J.A., Hamren, S., Ahle, D., and Chang, C.A. 1991. Branched DNA amplification multimers for the sensitive, direct detection of human hepatitis viruses. Nucl. Acids Symp. Series. 24:197-200. Wallace, J.C. and Edmonds, M. 1983. Polyadenylated nuclear RNA contains branches. Proc. Natl. Acad. Sci. U.S.A. 80:950-954. Wincott, F., Di Renzo, A., Shaffer, C., Grimm, S., Tracz, D., Workman, C., Sweedler, D., Gonzalez, C., Scaringe, S., and Usman, N. 1995. Synthesis, deprotection, analysis and purification of RNA and ribozymes. Nucl. Acids. Res. 23:2677-2684. Wu, T., Ogilvie, K.K., and Pon, R.T. 1989. Prevention of chain cleavage in the chemical synthesis of 2′-silylated oligoribonucleotides. Nucl. Acids Res. 17:3501-3517. Yee, T., Furuichi, T., Inouye, S., and Inouye, M. 1984. Multicopy single-stranded DNA isolated from a Gram-negative bacterium, Myxococcus xanthus. Cell 38:203-209.
Key References Damha and Zabarylo, 1989. See above.
Reports on the first general procedure for the convergent solid-phase synthesis of branched oligonucleotides via an adenosine bisphosphoramidite. Damha et al., 1992. See above. Reports on the convergent synthesis of branched RNA oligonucleotides using the standard silyl-phosphoramidite RNA synthesis methodology. The bNAs synthesized are related to the splicing intermediates derived from S. cerevisiae. Nam et al., 1994. See above. Reports on the substrate and sequence specificity of the yeast lariat debranching enzyme (yDBR), a unique 2′,5′-phosphodiesterase. The enzyme accepts a variety of substrates including group II intron lariats, msDNA, and synthetic bNAs. Padgett, R.A., Konarska, M.M., Grabowski, P.J., Hardy, S.F., and Sharp, P.A. 1984. Lariat RNA’s as intermediates and products in the splicing of messenger precursors. Science 225:898-903. This report provides evidence that the branched lariat structure is an intermediate of splicing of an adenovirus ML2 RNA transcript. Specifically demonstrated is that the excised intron contains an unusual nuclease-resistant core consisting of a branched trinucleotide structure with vicinal 2′,5′and 3′,5′-phosphodiester linkages. Sharp, P.A. 1994. Split genes and RNA splicing. Cell 77:805-815. A Nobel lecture. A paramount review describing the splicing of introns from nascent RNA, the evolutionary significance of introns, and the plethora of factors involved in post-transcriptional processing. Wallace and Edmonds, 1983. See above. A first account demonstrating the occurrence of a branched nuclear polyadenylated RNA containing vicinal 2′,5′- and 3′,5′-phosphodiester bonds. Such molecules were absent from cytoplasmic polyadenylated RNA, implicating these structures as intermediates during mRNA processing.
Internet Resources http://paris.chem.yale.edu/extinct.html A useful site for the calculation of molecular weights of oligonucleotides and peptides as well as the determination of extinction coefficients (ε).
Contributed by Sandra Carriero and Masad J. Damha McGill University Montreal, Canada
Solid-Phase Synthesis of Branched Oligonucleotides
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Solid-Phase Synthesis of 2′-Deoxy-2′-fluoroβ-D-Oligoarabinonucleotides (2′F-ANA) and Their Phosphorothioate Derivatives
UNIT 4.15
This unit describes the chemical synthesis of 2′-deoxy-2′-fluoro-β-D-oligoarabinonucleotides (2′F-ANA), both with phosphodiester and phosphorothioate linkages. The protocols described herein include araF phosphoramidite preparation (see Basic Protocol 1), assembly on DNA synthesizers (see Basic Protocol 2), and final deprotection and purification of oligonucleotides (see Basic Protocol 3). The preparation of araF phosphoramidite building blocks is carried out by introducing a 2-cyanoethyl-N,N-diisopropylaminophosphinyl group at the 3′-O-position of conveniently protected araF nucleosides. The preparation of araF-protected nucleosides is described in UNIT 1.7. Assembly of 2′F-ANA sequences can be carried out under similar conditions as for DNA sequences, but longer coupling times for ara F phosphoramidite monomers are required. For phosphorothioate analogs, longer sulfurization times are also required, as compared to S-DNA synthesis. Assembly of the oligonucleotides is carried out by the stepwise addition of phosphoramidite building blocks to nucleoside or nucleotide hydroxyl termini preimmobilized on a solid support until the desired sequence is obtained (APPENDIX 3C). Each addition of new building block requires five steps: detritylation, coupling, capping, oxidation or sulfurization, and capping. The last capping reaction also serves to dry the column prior to the next coupling cycle. Cleavage of the sequence from the solid support and the removal of the nucleobase- and phosphodiester-protecting groups is carried out using an aqueous ammonia/ethanol mixture. Crude oligonucleotides obtained in this way can be purified by ion-exchange high-performance liquid chromatography (HPLC; UNIT 10.5) or denaturing polyacrylamide gel electrophoresis (PAGE; UNIT 10.4). NOTE: All glassware should be oven dried.
MMTrO
O F OH
MMTrO
B
B
i -Pr2NP(Cl)OCH2CH2CN O
i -Pr2NEt, THF i -Pr2N
1a b c d
O F
B = N 6-benzoyladenin-9-yl B = N 2-isobutyrylguanin-9-yl B = N 4-benzoylcytosin-1-yl B = thymin-1-yl
P
2a b c d
OCH2CH2CN (75%) (70%) (80%) (95%)
Figure 4.15.1 Synthesis of the four araF-protected nucleoside phosphoramidites (S.2a-d). MMTr, 4-monomethoxytrityl; i-Pr2NEt, N-ethyl-N,N-diisopropylamine; i-Pr2-NP(Cl)OCH2CH2CN, 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite.
Synthesis of Modified Oligonucleotides and Conjugates
Contributed by Ekaterina Viazovkina, Maria M. Mangos, Mohamed I. Elzagheid, and Masad J. Damha
4.15.1
Current Protocols in Nucleic Acid Chemistry (2002) 4.15.1-4.15.22 Copyright © 2002 by John Wiley & Sons, Inc.
Supplement 10
BASIC PROTOCOL 1
PREPARATION OF araF PHOSPHORAMIDITES Conversion of protected araF nucleosides into the corresponding phosphoramidite building blocks is shown in Figure 4.15.1 (Wilds and Damha, 2000). In order to obtain phosphoramidites in sufficiently high yields, the starting materials and reaction solvents should be as dry as possible and all glassware should be oven dried. Likewise, to simplify purification, the reaction should be completed using a minimal amount of phosphitylation reagent. Materials Protected araF nucleosides (UNIT 1.7): N6-Benzoyl-9-[2-deoxy-2-fluoro-5-O-(4-methoxytrityl)-β-D-arabinofuranosyl] adenine (S.1a) N2-Isobutyryl-9-[2-deoxy-2-fluoro-5-O-(4-methoxytrityl)-β-D-arabinofuranosyl] guanine (S.1b) 4 N -Benzoyl-1-[2-deoxy-2-fluoro-5-O-(4-methoxytrityl)-β-D-arabinofuranosyl] cytosine (S.1c) 1-[2-Deoxy-2-fluoro-5-O- (4-methoxytrityl)-β-D-arabinofuranosyl]thymine (S.1d) THF, anhydrous (see recipe) N-Ethyl-N,N-diisopropylamine (DIPEA; Aldrich), double distilled 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite (Chem Genes) Dichloromethane 1:9 and 1:19 (v/v) methanol/dichloromethane Saturated sodium bicarbonate solution Magnesium sulfate, anhydrous Solvent system: 1:99 (v/v) triethylamine/chloroform (for araF-T) 1:9:10 (v/v) triethylamine/dichloromethane/hexanes (for araF-A and araF-C) 1:99 (v/v) triethylamine/dichloromethane (for araF-G) Silica gel (230 to 400 mesh) Diethyl ether 50-mL round-bottom flask equipped with stir bar and rubber septum Syringe TLC Merck silica plates (Kieselgel 60 F-254) 254-nm UV lamp 500-mL separatory funnel Vacuum evaporator (e.g., Savant Speedvac) 3-cm-diameter chromatography column Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare araF phosphoramidite monomers 1. In a 50-mL round bottom flask equipped with a stir bar and rubber septum, dissolve 1 mmol of each protected araF nucleoside (S.1a-d) in 3 to 5 mL of anhydrous THF. 2. While stirring, add 0.63 mL (3.6 mmol) double distilled N-ethyl-N,N-diisopropylamine (DIPEA) followed by the slow addition of 0.24 mL (1.1 mmol) 2-cyanoethylN,N-diisopropylchlorophosphoramidite.
Synthesis of 2′-Deoxy-2′-fluoroβ-D-oligoarabinonucleotides (2′F-ANA)
A white precipitate of diisopropylethylammonium hydrochloride salts forms after 10 min and is indicative of sufficiently anhydrous conditions for a successful reaction.
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3. Stir the reaction mixture for an additional 2 hr or until complete consumption of starting material is observed. For TLC analysis (APPENDIX 3D), remove 10 to 20 µL reaction mixture by syringe and dilute with 100 µL dichloromethane in a small tube. Analyze on a TLC Merck silica plate using 1:19 (v/v) methanol/dichloromethane for araF-T, -C, and -A, and 1:9 (v/v) methanol/dichloromethane for araF-G. Co-spot the starting material for comparison. Visualize by exposure with a 254-nm UV lamp. TLC solvents in this step do not include TEA. However, TLC solvents listed under step 11, can be substituted for those listed here. The phosphoramidites migrate faster than the starting material. In this reaction, two stereoisomeric products are formed, but they are not separable in the methanol/dichloromethane system.
4. If, as demonstrated by TLC, the reaction is not complete (i.e., more than ∼5% of starting material remaining), add an additional portion (∼0.2 eq.) of 2-cyanoethylN,N-diisopropylchlorophosphoramidite, stir for 2 hr more, and repeat TLC analysis (step 3). If the reaction looks complete by TLC analysis, start workup of the phosphoramidite. It is important to minimize the amount of phosphitylating reagent used, otherwise the excess is hydrolyzed during the workup to form H-phosphonate impurities (31P NMR: singlet, ∼14 ppm), which can cause difficulties during subsequent purification of the amidites.
5. Dilute the reaction mixture with 150 mL dichloromethane, transfer it to a 500-mL separatory funnel, and wash with 150 mL saturated sodium bicarbonate solution. 6. Collect the organic layer, dry it by adding solid anhydrous magnesium sulfate until it no longer clumps, and filter and concentrate under reduced pressure to obtain a crude product. Chromatographic purification of the crude amidite is recommended, but if the reaction mixture looks pure by TLC and evaporates easily to give a stable foam, this step may be omitted. In case of a second day purification, it is strongly recommended to evaporate the crude compound to a foam, dry it under high vacuum, and keep the product in a vacuum desiccator. If the product persists as an oil after these steps are taken, co-evaporate it with diethyl ether. A foam can usually be obtained after co-evaporation.
Purify phosphoramidite monomers by flash column chromatography 7. Redissolve the crude material in a minimal amount of one of the following solvent systems: 1:99 (v/v) triethylamine/chloroform for araF-T 1:9:10 (v/v) triethylamine/dichloromethane/hexanes for araF-A and araF-C 1:99 (v/v) triethylamine/dichloromethane for araF-G. 8. Apply sample to a 3-cm-diameter chromatography column containing 50 g 230- to 400-mesh silica gel, preconditioned with the appropriate chromatography solvent (step 7). Column chromatography is performed according to the method of Still et al. (1978) using a small amount of air pressure at a rate of ∼1 inch of solvent per minute (also see APPENDIX 3E).
Synthesis of Modified Oligonucleotides and Conjugates
4.15.3 Current Protocols in Nucleic Acid Chemistry
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9. Apply the sample to the column and begin eluting with the preconditioning solvent. a. For araF-C and araF-A: Elute in 1:9:10 (v/v) triethylamine/dichloromethane/hexanes. b. For araF-T: Follow a stepwise gradient of triethylamine/chloroform until a 1:31 ratio is reached. For 1 mmol crude sample, collect 100 mL eluate at 1% TEA, 100 mL at 2% TEA, and so on. c. For araF-G: Follow a step gradient of methanol/triethylamine/dichloromethane from 0:1:99 to 5:1:94 (v/v). For 1 mmol crude sample, collect 100 mL at 0% methanol, 100 mL at 1% methanol, and so on. Always add at least 0.5% of triethylamine to the chromatographic solvent mixtures used for elution and preconditioning of the column in order to avoid hydrolysis of the amidites to the H-phosphonates during purification. Step gradients indicated are only intended as guidelines and should be adjusted according to various factors including amount of crude product applied and flow rate of mobile phase.
10. Monitor fractions by TLC (APPENDIX phosphoramidite.
3D;
step 3) and pool those containing pure
11. Evaporate the phosphoramidite on a vacuum evaporator and dry under high vacuum to obtain a stable foam. If a foam does not form, try co-evaporation of the pure product with diethyl ether. Typically, a colorless or pale yellow foamy product forms easily. This can be stored in a vacuum desiccator over phosphorus pentoxide or at −20°C in a freezer for several months without decomposition. For araF-A (S.1a), the typical yield after chromatographic purification is ∼70% to 75%. TLC using 5:45:50 (v/v/v) triethylamine/hexanes/dichloromethane results in two spots corresponding to two isomers at Rf 0.22 and 0.33. 31P NMR (202.3 MHz, acetone-d6): 150.9 and 151.1, 19F NMR (470.27 MHz, acetone-d6, without external reference): yields −197.3 and –197.7 (ddd, J1′-F = 19 Hz, J′2′-F = 52 Hz, J3′-F = 19 Hz), and FAB-MS using an NBA matrix yields 846.34 [M+]. For araF-G (S.1b), the typical yield after chromatographic purification is ∼70% to 75%. TLC using 20:1 (v/v) chloroform/methanol yields a spot of Rf 0.54; 31P NMR (200.06 MHz, acetone-d6, 85% ortho-phosphoric acid as external reference): 151.8 and 151.0; 19F-NMR (282.32 MHz, acetone-d6, 99% trifluoroacetic acid as external reference): −119.7 and –119.4; and FAB-MS using an NBA matrix yields 822 [M+]. For araF-C (S.1c), the typical yield after chromatographic purification is ∼80% to 85%. TLC using 5:45:50 (v/v/v) triethylamine/hexanes/dichloromethane yields two spots, corresponding to two isomers, at Rf 0.26 and 0.34; 31P NMR (500 MHz, acetone-d6): 151.3 and 150.8; 19F NMR (470.27 MHz, acetone-d6, without external reference): −199.0 and –199.2 (ddd, J1′-F = 18 Hz, J′2′-F = 52 Hz, J3′-F = 18 Hz); and FAB-MS using an NBA matrix yields 828 [M+]. For araF-T (S.1d), typical yield after chromatographic purification is ∼90% to 95%. TLC using 10:9:1 (v/v/v) chloroform/ethyl acetate/ethanol yields two spots, corresponding to two isomers, at Rf 0.76 and 0.83; 31P NMR (500 MHz, acetone-d6): 151.3 and 150.8; 19F NMR (470.27 MHz, acetone-d6, without external reference): −199.0, –199.2 (ddd, J1′-F = 18 Hz, J′2′-F = 52 Hz, J3′-F = 18 Hz); and FAB-MS using an NBA matrix yields 822 [M+].
Synthesis of 2′-Deoxy-2′-fluoroβ-D-oligoarabinonucleotides (2′F-ANA)
4.15.4 Supplement 10
Current Protocols in Nucleic Acid Chemistry
SOLID-PHASE ASSEMBLY OF PROTECTED araF PHOSPHORAMIDITES This protocol describes the setup and step-by-step synthesis of araF oligonucleotides. The methodology has been optimized for use with the Expedite 8909 DNA synthesizer equipped with a workstation (PerSeptive Biosystems), but can easily be adapted to other automated DNA synthesizers. All syntheses have been performed on 1-µmol scales, both for phosphodiester- and phosphorothioate-containing araF oligonucleotides. The coupling time for araF nucleosides requires a 15-min “wait” step, as compared to 1.5 min for deoxynucleosides on the Expedite instrument. Likewise, the detritylation time has been extended to 150 sec to allow for effective removal of the monomethoxytrityl (MMTr) protection as opposed to the more labile dimethoxytrityl (DMTr) groups that are commonly used with standard DNA monomers. For thio-araF-oligonucleotide synthesis, the overall success of oligomer synthesis was evaluated with different sulfur-transfer reagents, including 3H-1,2-benzodithiol-3-one-1,1-dioxide (Beaucage reagent; Iyer et al., 1990), 3-ethoxy-1,2,4-dithiazoline-5-one (EDITH; Xu et al., 1996), and 3-amino-1,2,4dithiazoline-5-thione (ADTT; Tang et al., 2000). In all cases, the concentration of sulfurization reagent was increased ∼5 fold as compared to customized S-DNA synthesis, and the time required for the oxidation was prolonged to 10 min as opposed to 1 to 2 min for S-DNA synthesis. Current procedures can easily be adapted to the synthesis of molecules with mixed nucleotide composition (e.g., DNA/araF chimeras of variable gap sequences).
BASIC PROTOCOL 2
Materials AraF phosphoramidites (see Basic Protocol 1) Argon gas Acetonitrile, anhydrous (see recipe) Liquid Reagent kit for the Expedite 8909 instrument (PerSeptive Biosystems): Acetonitrile wash and amidite diluent: anhydrous acetonitrile Activator solution: dissolve 1.8 g sublimed tetrazole (0.5 M) in 50 mL acetonitrile; store up to 2 wks at room temperature. Cap A (see recipe) Cap B (see recipe) Oxidizer solution (see recipe) Deblock solution: 15 g trichloroacetic acid in 500 mL dichloromethane; store up to 1 yr at room temperature Sulfurization reagent (see recipe) Amidite column (see recipe) Synthesizer vials with caps Vacuum desiccator containing phosphorus pentoxide Syringe Automated DNA synthesizer (e.g., Expedite 8909, Perseptive Biosystems) with trityl monitor Additional reagents and equipment for oligonucleotide synthesis (APPENDIX 3C) Synthesize phosphodiester and phosphorothioate araF oligonucleotides 1. Calculate the amount of araF phosphoramidites required for the synthesis of a desired sequence. The typical concentration of amidites used with the Expedite synthesizer corresponds to 50 mg/mL. Note that the volume of amidite solution delivered to the column during each coupling step is ∼240 ìL. However, an excess of 50 mg amidite (i.e., 1 mL solution) should be included when preparing each amidite solution in order to purge the lines of the synthesizer.
Synthesis of Modified Oligonucleotides and Conjugates
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Table 4.15.1 Automated 1 µmol Synthesis Cycle for the Synthesis of Phosphodiester araF Oligonucleotides on the Expedite 8909 Instrumenta
Functionb Amountc Time (sec) Description Detritylation of the support bound nucleoside: Trityl monitor on/off
—
1
Deblock
10
0
Deblock
70
150
Diverted wash A
40
0
Trityl mon. on/off Diverted wash A
— 40
1 0
Coupling of phosphoramidite: Wash 5 Activator 5
0 0
A + activator
6
0
A + activator
9
500
Wash
8
400
Wash
7
0
Wash A
20
0
Cap A and B
7
0
Cap A and B
6
15
Wash A
6
15
Wash A
14
0
Trityl analysis monitor is turned on for data collection Detritylation solution is rapidly delivered to column Slow, prolonged delivery of detritylation solution to column Lines are flushed with anhydrous acetonitrile (wash A) Trityl quantitation is stopped Lines are flushed with anhydrous acetonitrile Lines are flushed with acetonitrile Lines are flushed with tetrazole solution Rapid addition of the phosphoramidite (A) and tetrazole solutions to the column Coupling of the free activated phosphoramidite to the support-bound terminal nucleoside Slow delivery of acetonitrile to the column to purge remaining phosphoramidite solution Lines and column are washed with acetonitrile
Capping of the column: System is flushed with anhydrous acetonitrile Equal volumes of cap A and cap B solutions are rapidly delivered to column Slower delivery of cap A and cap B to column (to maximize capping efficiency) Slow delivery of acetonitrile to column to purge remaining cap A and B solutions Lines and column are flushed with anhydrous acetonitrile
Oxidation of phosphoramidite: Oxidizer
20
0
Wash A
15
0
Cap A and B
7
0
Wash A
30
0
Oxidant is delivered to the column to oxidize the newly formed phosphite triester linkages Lines and column are flushed with anhydrous acetonitrile
Capping of the column:
Synthesis of 2′-Deoxy-2′-fluoroβ-D-oligoarabinonucleotides (2′F-ANA)
Equal volumes of cap A and B solutions are rapidly delivered to column Lines and column are flushed with anhydrous acetonitrile
aStandard DNA coupling cycles supplied by the manufacturer have been optimized for 2′-araF oligonucleotide assembly. bAll entries are as described in a typical cycle sequence for the Expedite instrument; see manufacturer’s directions for
further information.
cRepresents the number of pulses required for the corresponding step; each pulse has approximately 16 µL volume.
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Table 4.15.2 Automated 1-µmol Synthesis Cycle for the Synthesis of Phosphorothioate araF Oligonucleotides on Expedite 8909 Instrumenta
Function
Amountb
Time (sec)
Description
Detritylation of the support-bound nucleoside: Trityl monitor on/off Deblock
1 10
1 0
Trityl analysis monitor is turned on for data collection Detritylation solution is rapidly delivered to column
Deblock
70
150
Slow, prolonged delivery of detritylation solution to column
Diverted wash A
40
0
Lines are flushed with anhydrous acetonitrile (wash A)
Trityl monitor on/off
0
1
Trityl quantitation is stopped
Diverted wash A
40
0
Lines are flushed with anhydrous acetonitrile
Coupling of phosphoramidite: Wash
5
0
Lines are flushed with acetonitrile
Activator
5
0
Lines are flushed with tetrazole solution
A + activator
6
0
Rapid addition of the phosphoramidite (A) and tetrazole solutions to the column
A + activator
9
500
Coupling of the free activated phosphoramidite to the support-bound terminal nucleoside
Wash
8
400
Slow delivery of acetonitrile to the line to purge remaining phosphoramidite solution
Wash
7
0
Lines and column are rapidly washed with acetonitrile
Wash A
20
0
System is flushed with anhydrous acetonitrile
Cap A and B
7
0
Equal volumes of cap A and cap B solutions are rapidly delivered to column
Cap A and B
6
15
Slower delivery of cap A and cap B to column (to maximize capping efficiency)
Wash A
6
15
Slow delivery of acetonitrile to column to purge remaining cap A and B solutions
Wash A
14
0
Lines and column are rapidly flushed with anhydrous acetonitrile
Capping of the column:
Sulfurization of phosphoramidite: Sox
15
0
Sulfurizing reagent is rapidly pulsed through column to sulfurize the newly formed phosphite triester linkages
Sox
25
400
Slower delivery of sulfurizing reagent through column (to maximize sulfur transfer efficiency)
Wash A
15
200
Slow delivery of acetonitrile to column to purge remaining sulfurizing solution from system
Wash A
15
0
System is rapidly flushed with acetonitrile
Cap A and B
7
0
Equal volumes of cap A and B solutions are rapidly delivered to column
Wash A
30
0
Lines and column are flushed with anhydrous acetonitrile
Capping of the column:
aStandard DNA coupling cycles supplied by the manufacturer have been optimized for 2′-araF oligonucleotide assembly. bRepresents the number of pulses required for the corresponding step; each pulse has approximately 16 µL volume.
4.15.7 Current Protocols in Nucleic Acid Chemistry
Supplement 10
2. Weigh out the calculated amounts of phosphoramidites into the appropriate synthesizer vials and leave these to dry in a vacuum desiccator over phosphorus pentoxide overnight. 3. Flush the desiccator with argon gas before opening, and carefully remove the preweighed amidites. 4. Dissolve the phosphoramidites in anhydrous acetonitrile to a final concentration of 50 mg/mL. Use of syringe to transfer the acetonitrile through the rubber septum of each capped synthesizer vial to ensure the reagents are protected from humidity. Do not open the phosphoramidite vials until they are ready to be placed directly on the synthesizer.
5. Connect the solutions from the Liquid Reagents kit for the Expedite 8909 instrument, the phosphoramidite solutions, and the sulfurization reagent to the automated DNA synthesizer according to the manufacturer’s directions. The Expedite 8909 synthesizer is equipped with an extra position for the sulfur transfer reagent. If a synthesizer without this option is to be used, the sulfurization reagent may be placed at the position of the oxidizer, provided that the line has previously been washed with acetonitrile to avoid cross-contamination of the reagents.
6. Purge the lines of the synthesizer with all the solutions and solvents. 7. Install the appropriate amidite column. 8. Modify the synthetic cycles for araF phosphoramidites. Examples of synthesis cycles for phosphodiester and phosphorothioate araF oligonucleotides are given in Table 4.15.1 and Table 4.15.2. 9. Enter the sequence to be synthesized. 10. Carry out the assembly on the instrument according to the manufacturer’s instructions, choosing the “DMTr-off” (trityl-off) option. 11. Check the coupling efficiency periodically using the trityl monitor equipped with the instrument (also see APPENDIX 3C). 12. When the synthesis is complete, remove the column from the synthesizer and dry it under vacuum. 13. Deprotect and purify to obtain the desired oligonucleotide (see Basic Protocol 3). BASIC PROTOCOL 3
Synthesis of 2′-Deoxy-2′-fluoroβ-D-oligoarabinonucleotides (2′F-ANA)
DEPROTECTION AND PURIFICATION OF araF OLIGONUCLEOTIDES This protocol describes the steps necessary for cleavage of araF sequences, assembled as described above (see Basic Protocol 2), from the solid support and removal of the protecting groups from the heterocyclic bases and phosphates. The procedure for the quantitation of isolated oligonucleotides is also described. AraF oligonucleotides can be successfully analyzed and purified by anion-exchange HPLC or denaturing gel electrophoresis (PAGE; UNIT 10.4 and APPENDIX 3B), as described below. Analytical amounts of material can easily be isolated by gel electrophoresis, while large amounts require HPLC purification. Desalting of the oligomers after gel or chromatographic isolation completes the purification procedure. There are two different methods that may be employed to accomplish this, either size-exclusion chromatography or reversed-phase SepPak C18 cartridge purification.
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Materials Fully protected oligonucleotides, attached to the solid support of a synthesis column (see Basic Protocol 2) Ethanol, anhydrous 29% ammonium hydroxide Denaturing acrylamide gel stock solution (see recipe) Loading buffer (e.g., formamide/dye mix, UNIT 10.4) Sephadex G-25 column (Amersham Pharmacia Biotech) 1 M NaClO4 Anhydrous and 25% or 50% (v/v) acetonitrile 1.5 mL microcentrifuge tubes or screw-cap microcentrifuge tubes with O-ring seal Platform shaker 55°C heating block or water bath (optional) Vacuum evaporator (e.g., Savant Speedvac) with low and high vacuum sources UV spectrophotometer and cuvette 0.75- and 1.5-mm-thick gel plates TLC Merck silica plate (Kieselgel 60 F-254) Hand-held 254-nm UV lamp Camera with UV filter (optional) 0.45 µm hydrophilic fluid filter (Creative Medical) 0.22 µm membrane filter (Millipore; optional) Anion-exchange high-performance liquid chromatograph (HPLC) with: Gradient maker 0.5 mm × 7.5 cm Protein Pak DEAE 5PW column (Waters) Column heater Sep-Pak C18 cartridges (Waters Chromatography) 10 mL syringe Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (APPENDIX 3B and UNIT 10.4) Deprotect oligonucleotides 1. Remove the fully protected oligonucleotides attached to the solid support of the synthesis column, and place in a 1.5 mL microcentrifuge tube or a screw-cap microcentrifuge tube with O-ring seal (seal tightly) if deprotecting at increased speed and temperature (step 3). 2. Add 0.25 mL anhydrous ethanol and 0.75 mL of 29% ammonium hydroxide. 3. Place the tube on a platform shaker for 48 hr at room temperature. Alternatively, deprotect by incubating in a 55°C heating block or waterbath for 15 hr. 4. Once the deprotection is complete, cool the microcentrifuge tube ∼1 hr in the freezer (i.e., −20°C) before opening. 5. Microcentrifuge the samples briefly so that the CPG beads are allowed to settle, then transfer the supernatant to another microcentrifuge tube. Wash the CPG with ethanol twice and place the supernatant from each wash in additional microcentrifuge tubes. 6. Dry the samples in a vacuum evaporator, applying low vacuum first to remove all traces of ammonia. Introducing high vacuum to the ammonia/ethanol solution may cause bumping of the liquid and loss of the sample. Synthesis of Modified Oligonucleotides and Conjugates
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7. When the ammonia has been evaporated and sample volumes are decreased enough, combine the supernatants from each microcentrifuge tube. Evaporate the final sample to dryness under high vacuum. Quantitate oligonucleotide 8. Dissolve the oligonucleotide from the previous step in 1 mL deionized water. Remove a 10-µL aliquot and dilute this to 1 mL with deionized water in a UV cuvette. 9. Measure the absorption of the sample at 260 nm. Calculate the amount of crude oligonucleotide obtained in A260 units. Typically, a 1-ìmol synthesis may give yields of 100 to 180 A260 units of crude material, depending on the sequence length and composition.
Analyze crude oligonucleotide by denaturing PAGE 10. Prepare a 24% denaturing polyacrylamide gel on a 0.75-mm-thick analytical gel plate for gel electrophoresis (APPENDIX 3B and UNIT 10.4). 11. Prepare samples of oligonucleotides to be analyzed (e.g., 0.5 to 1 A260 units per analysis). Dissolve each of these in 10 µL loading buffer. 12. Load the samples into wells and run these alongside loading buffer alone, which is placed in the first and last lanes of the gel. Connect the gel to the power supply and start running at 200 V until the dye has fully diffused into the gel. Increase the voltage to 500 V and maintain at this setting until the faster running dye marker has traveled 3⁄ down the length of the gel. 4 13. Turn off the current and dismantle the gel apparatus. Put the gel in plastic wrap. Place a fluorescing TLC Merck silica plate under the gel and examine by illumination with a hand-held 254-nm UV lamp. If necessary, photograph the gel using a camera with a UV filter. Oligonucleotides absorb UV light and appear as dark bands in the gel against a fluorescent background. Typically, crude FANA oligonucleotides, both diester and thio, consist of one intense band. The presence of multiple bands below the desired product usually indicates poor coupling efficiency.
Purify oligonucleotides by denaturing PAGE and desalt using Sephadex G-25 14. Prepare a 24% to 16% denaturing polyacrylamide gel on a 1.5-mm-thick gel plate for preparative analysis (APPENDIX 3B and UNIT 10.4). Compared to analytical separations, a lower percentage of acrylamide may be used in preparative analyses (from 16% to 24%), depending on the quality of the synthesis. For example, a crude oligonucleotide which migrates as one main band in analytical compositions can be sufficiently purified using 16% acrylamide in the preparative run. When multiple bands corresponding to failed sequences are present in the gel, a higher percentage of acrylamide solution should be used.
15. Prepare and dry the oligonucleotide sample to be purified (∼20 to 30 A260 units). Dissolve the sample in 100 µL loading buffer. Sample loading to the gel may vary, depending on the quality of the synthesis.
16. Load the sample on the gel and repeat steps 12 and 13 above. 17. Using a razor blade, cut out the segment of gel containing the band of interest. Synthesis of 2′-Deoxy-2′-fluoroβ-D-oligoarabinonucleotides (2′F-ANA)
18. Transfer the excised band to a 15-mL conical plastic tube with screw cap and crush with a spatula. Add 10 mL water and leave the tube on a platform shaker overnight to extract the oligonucleotide from the gel.
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Current Protocols in Nucleic Acid Chemistry
19. Filter the supernatant through a 0.45-µm hydrophilic fluid filter and concentrate on a vacuum evaporator. 20. Desalt the sample using a Sephadex G-25 column according to the manufacturer’s directions. 21. Quantitate the amount of oligonucleotide isolated via UV spectroscopy. 22. Lyophilize the oligonucleotide. The purified, dry oligonucleotide can be stored for long periods (e.g., >1 yr) at −20°C. If the oligonucleotide is to be used in combination with microbially sensitive materials (e.g., cell culture), filtration through a 0.22-ìm filter is recommended. Collect the filtrate in previously autoclaved tubes and lyophilize.
Analyze and purify oligonucleotide by anion-exchange HPLC and desalt on SepPak C18 cartridge 23. Prepare the anion-exchange HPLC by setting the detector wavelength to 260 nm, heating the column to 50°C, and equilibrating with buffer A (deionized water) using a flow rate of 1 mL/min or as specified by the manufacturer. See UNIT 10.5 and legends to Figures 4.15.2 to 4.15.4 for more details.
A
0.35 0.30
AU
0.25 0.20 0.15 0.10 0.05 0.00 0.00
10.00
20.00
30.00
40.00
50.00
60.00
70.00
80.00
90.00
60.00
70.00
80.00
90.00
Minutes
B 0.50 0.40
AU
0.30 0.20 0.10 0.00 0.00
10.00
20.00
30.00
40.00
50.00
Minutes
Figure 4.15.2 Ion-exchange HPLC analysis of crude S-oligonucleotide ATA TCC TTg TCg TAT CCC (cap letters represent araF nucleotides, DNA residues in lowercase). (A) Sulfurization with Beaucage reagent. The small peak at ∼58 min represents S-oligonucleotides with one P-O insertion. The main peak at ∼62 min represents full S-FANA. (B) Sulfurization with ADTT. System: Waters 600E Multisolvent Delivery System with Waters 486 Tunable Absorbance detector and oven, driven by Millennium (V 3.20) software; column: Waters 0.5-mm × 7.5-cm Protein Pak DEAE 5PW, 50°C; solvent A: H2O; solvent B: 1 M NaClO4; flow rate: 1 mL/min; detection: 260 nm; gradient conditions: 0% B for 9 min, 0% to 15% B in 3 min, 15% to 50% B in 60 min, 50% to 80% B in 2 min, hold 80% B 10 min, 80% to 0% B in 2 min, hold 0% B 10 min.
Synthesis of Modified Oligonucleotides and Conjugates
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24. Load ∼1 A260 unit crude oligonucleotide sample (steps 8 and 9) to the column. Wash 5 min with buffer A, then apply a linear gradient of 0% to 25% buffer B (1M NaClO4) in 60 min (i.e., 0.42%/min) to elute diester araF oligonucleotides, or of 0% to 50% buffer B in 60 min (i.e., 0.83%/min) for S-FANA oligonucleotides. Wash the column and re-equilibrate. The desired fraction should elute close to the end of the applied gradient.
25. Optimize the conditions for the next run. Examples of suitable chromatographic gradients that may be used are given in the legends to Figures 4.15.2 to 4.15.4. 26. Continue purification of the next portions of oligonucleotide using the previously optimized conditions. Increase the amount of oligonucleotide loaded to the column and set the detector at 280 to 290 nm depending on the amount of the sample to be loaded. Use higher wavelengths for larger samples to avoid saturating the UV detector. Sample loading to the column depends on the quality of the reaction mixture and type of oligonucleotide. Typical loading amounts for phosphodiester FANA oligonucleotides range from 10 to 30 A260 units of crude reaction mixture using the conditions described above. Loading for S-FANA oligonucleotides can be increased considerably as a result of their stronger adsorption on the ion-exchange column. The authors have been able to purify ∼100 A260 units of crude mixture in one loading.
27. Quantitate the total amount of oligonucleotide obtained.
A 0.060 0.050
AU
0.040 0.030 0.020 0.010 0.000 0.00
10.00
20.00
30.00
40.00
50.00
60.00
70.00
80.00
50.00
60.00
70.00
80.00
Minutes
B 1.00
AU
0.80 0.60 0.40 0.20 0.00 0.00
10.00
20.00
30.00
40.00 Minutes
Synthesis of 2′-Deoxy-2′-fluoroβ-D-oligoarabinonucleotides (2′F-ANA)
Figure 4.15.3 Ion-exchange HPLC analysis of (A) crude and (B) purified S-oligonucleotide CTC TAg cgt ctT AAA (cap letters represent araF nucleotides, lowercase letters represent DNA residues). System: Waters Binary HPLC Pump with Waters 2487 Dual λ Absorbance detector, equipped with in-line degasser and oven, driven by Breeze (V 3.20) software; column: Waters 0.5-mm × 7.5-cm Protein Pak DEAE 5PW, 50°C; solvent A: H2O; solvent B: 1 M NaClO4; flow rate: 1 mL/min; detection: 260 nm; gradient conditions: 0% B for 9 min, 0% to 15% B in 3 min, 15% to 50% B in 45 min, 50% to 80% B in 2 min, hold 80% B 10 min, 80% to 0% B in 2 min, hold 0% B 10 min. Peaks at <20 min correspond to failure sequences and loss of protecting groups (e.g., benzoyl from A or C).
4.15.12 Supplement 10
Current Protocols in Nucleic Acid Chemistry
28. For the desalting step, prepare SepPak C18 cartridges first by washing them with 10 mL (each) water, then anhydrous acetonitrile, then water, using a 10-mL syringe. Make sure that the cartridges are well equilibrated with water; traces of acetonitrile will prevent adsorption of the oligonucleotide and may lead to substantial loss of material.
29. Load ∼20 to 30 A260 units of oligonucleotide per cartridge, wash the cartridge with 10 mL water, and then elute the pure desalted oligonucleotide with 5 mL of 25% or 50% acetonitrile in water for phosphodiester and phosphorothioate-oligonucleotides, respectively. Quantitate the amount of oligonucleotide obtained. Always collect water fractions from all desalting steps. If the cartridge was not equilibrated properly or was overloaded, the oligonucleotide may elute with the water fraction. If this occurs, load the water fraction with oligonucleotide to a new cartridge and repeat the desalting procedure.
30. Evaporate the oligonucleotide on a vacuum evaporator using low vacuum first to remove acetonitrile, followed by complete drying of the sample under high vacuum. 31. If necessary, filter the oligonucleotide through a 0.22 µm membrane filter (see comments in step 22). Desalting after HPLC purification can be done on Sephadex G-25 as well.
A 1.20 1.00
AU
0.80 0.60 0.40 0.20 0.00 0.00
10.00
20.00
30.00 Minutes
40.00
50.00
60.00
10.00
20.00
30.00 Minutes
40.00
50.00
60.00
B 0.70 0.60
AU
0.50 0.40 0.30 0.20 0.10 0.00 0.00
Figure 4.15.4 Ion-exchange HPLC analysis of (A) crude and (B) purified phosphodiester oligonucleotide ATg TCC TTg TCg gTg Agg TTA GG (cap letters represent araF nucleotides, lowercase for DNA residues). System: Waters Binary HPLC Pump with Waters 2487 Dual λ Absorbance detector, equipped with in-line degasser and oven, driven by Breeze (V 3.20) software; column: Waters 0.5-mm × 7.5-cm Protein Pak DEAE 5PW, 50°C; solvent A: H2O; solvent B: 1 M NaClO4; flow rate: 1 mL/min; detection: 260 nm; gradient conditions: hold 0% B 2 min, from 0% to 30% B in 40 min, from 30% to 45% B in 2 min, hold 45% B 10 min, 45% to 0% B in 2 min, hold 0% B 10 min.
Synthesis of Modified Oligonucleotides and Conjugates
4.15.13 Current Protocols in Nucleic Acid Chemistry
Supplement 10
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Acetonitrile, anhydrous Predry acetonitrile by refluxing and distilling over phosphorus pentoxide overnight. Reflux and distill over calcium hydroxide. Store sealed or on the synthesizer up to 1 month at room temperature. Amidite column Prepare a succinyl-LCAA-CPG support derivatized with 5′-(4-monomethoxytrityl) and the base-protected araF nucleoside of choice (S.1a-d; UNIT 3.2). Average nucleoside loadings are 20 to 40 µmol/g, according to trityl analysis in APPENDIX 3C. Cap A 10 mL acetic anhydride 10 mL 2,6-lutidine or 2,4,6-collidine 80 mL THF (see recipe) Store sealed or on synthesizer up to 1 month at room temperature Cap B 16 mL N-methylimidazole 84 mL THF (see recipe) Store sealed or on synthesizer up to 1 month at room temperature Denaturing acrylamide gel stock solution For 24% analytical gel: 120 g acrylamide 6 g N, N′-methylene-bisacrylamide 210 g urea 50 mL 10× TBE electrophoresis buffer (APPENDIX 2A) Water to 500 mL Store protected from light for up to 1 year at 4°C. Immediately before pouring, degas 50 mL stock solution for 10 min and add 20 µL TEMED and 200 µL freshly prepared 10% (w/v) ammonium persulfate. Allow gel to polymerize overnight at room temperature (also see APPENDIX 3B and UNIT 10.4 for pouring acrylamide gels). For 16% to 24% preparative gel: Prepare stock solution and gel as above, but use 80 g acrylaminde and 4 g N,N′-methylenebisacrylamide for 16% gel, or scale as appropriate for up to 24%. Immediately before pouring, degas 65 mL stock solution for 10 min and add 35 µL TEMED and 350 µL freshly prepared 10% (w/v) ammonium persulfate. Oxidizer solution 1.23 g iodine 25 mL THF (see recipe) 20 mL pyridine 2 mL H2O Store sealed up to 6 months at room temperature or on synthesizer up to 2 wks at room temperature. Synthesis of 2′-Deoxy-2′-fluoroβ-D-oligoarabinonucleotides (2′F-ANA)
4.15.14 Supplement 10
Current Protocols in Nucleic Acid Chemistry
Sulfurization reagent Any of the reagents listed below can be used to sulfurize the phosphite triester linkage. Beaucage reagent: 2 g 3H-1,2-benzodithiol-3-one 1,1-dioxide (0.2 M) 50 mL anhydrous acetonitrile (see recipe) Store on the synthesizer up to 2 or 3 days at room temperature. Discard if precipitate is observed. EDITH reagent: 1 g 3-ethoxy-1,2,4-dithiazoline-5-one (0.125 M) 100 mL anhydrous acetonitrile (see recipe) Store up to 1 month at room temperature ADTT reagent: 1.5 g 3-amino-1,2,4-dithiazoline-5-thione (0.1 M) 50 mL anhydrous acetonitrile (see recipe) 50 mL pyridine Store up to 1 month at room temperature Tetrahydrofuran (THF), anhydrous Dry tetrahydrofuran by distillation over sodium and benzophenone. Storage of anhydrous THF is not recommended as peroxides may be formed. Always use freshly distilled. COMMENTARY Background Information Antisense oligonucleotides (AONs) represent an extensive class of biologically relevant, nonnatural polymeric nucleic acids that have potential utility as novel pharmaceuticals or in molecular diagnostics applications as probes of gene function. These analogs are specifically designed to induce selective gene interference via tight and specific associations with their intended genetic targets. For example, such a “bottleneck” approach can target the mRNA directly to inhibit the expression of a dysfunctional gene, and thereby transcend traditional approaches that control a disease phenotype by targeting the malformed protein (for review, see Myers and Dean, 2000, and references therein). Furthermore, with the complete sequence of the human genome now at hand, the future design of antisense-based therapeutics may be feasible without prior knowledge of the protein structure. Indeed, adequate optimization of the biological properties of chemically modified AONs is paramount to the success of this and related technologies. The most efficacious AON members are those that ideally possess remarkable stabilities against both general serum and cellular exo- and endonucleolytic activities, and adequate lipophilicity to ensure efficient cell permeation. They should also ex-
hibit high discrimination toward fully complementary host cell sequences and allow minimal residual expression of the target gene (see Lebedeva and Stein, 2001, and references therein). Additionally, suppression of gene activity in eukaryotic organisms may proceed catalytically with the assistance of intracellular ribonuclease H (RNase H), a ubiquitous enzyme implicated in DNA replication and repair processes (Walder and Walder, 1988; Crouch and Toulmé, 1998). In fact, the extent to which protein synthesis is inhibited via antisense action in vivo seems to correlate with the propensity of the preformed AON:RNA substrate to elicit RNase H destruction of the mRNA component. Thus, a single enzyme-active AON can be recycled by this pathway to silence multiple copies of an RNA transcript, and display superior inhibition to those that do not activate the enzyme (Uhlmann and Peyman, 1990). Importantly, these events hinge upon presentation of the correct hybrid shape by the substrate duplex to the enzyme for subsequent hydrolysis of the targeted species (Nakamura et al., 1991; Oda et al., 1993). However, most AONs lack the appropriate structural attributes for enzyme elicitation and typically display less than optimal cellular potencies (see for example, Lima and Crooke, 1997).
Synthesis of Modified Oligonucleotides and Conjugates
4.15.15 Current Protocols in Nucleic Acid Chemistry
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Table 4.15.3 Thermal and Biological Properties of Oligonucleotide Analogsa
Oligonucleotide analogue
∆Tm/mod. RNase H activation (°C)
Reference
thioate-LNA LNA α-L-LNA 2′F-RNA N-methanocarba-NA MMI-2′OMe-RNA 2′-MOE-RNA 2′OMe-RNA RNA CeNA 2′F-ANA thioate-2′F-ANA DNA thioate-DNA ANA S-methanocarba-NA 2′β-Me-ANA [3.3.0]bc-ANA
+10.0 +5.6 +4.6 +3.0 +2.6 +2.0 +2.0 +1.7 +0.7 +2 +1.2 +0.5 +0.5 0 –0.5 –0.6 –2.8 –3.5c
Wengel et al., 1999 Wengel et al., 1999 Sørensen et al., 2002 Manoharan, 1999 Altmann et al., 1994a Sangvhi, 1998 Manoharan, 1999 Manoharan, 1999 Wilds and Damha, 2000 Wang et al., 2000 Wilds and Damha, 2000 Lok et al., 2002 Wilds and Damha, 2000 Wilds and Damha, 2000 Noronha et al., 2000 Altmann et al., 1994b Schmit et al., 1994 Christensen et al., 1998
No No Yes No n/ab No No No No Yes Yes Yes Yes Yes Yes n/ab No No
aAbbreviations: α-L-LNA, α-L-ribo-configured locked nucleic acid; 2′β-Me-ANA, 2′-deoxy-2′-O-
methylarabinonucleic acid; ANA, arabinonucleic acid; [3.3.0]bc-ANA, 2′,3′-[3.3.0]-bicycloarabinonucleic acid; CeNA, cyclohexene nucleic acid; 2′F-ANA, 2′-deoxy-2′fluoroarabinonucleic acid; 2′F-RNA, 2′-deoxy-2′fluororibonucleic acid; LNA, locked nucleic acid; N-methanocarba-NA, Northern-locked 4′,6′-methanocarbocyclic nucleic acid; S-methanocarba-NA, Southern-locked 1′,6′-methanocarbocyclic nucleic acid; MMI-2′OMe-RNA, methylene(methylimino)-2′-OMe-RNA; 2′-MOE-RNA, 2-O-(2-methoxyethyl) RNA; 2′OMe-RNA, 2′methoxyribonucleic acid; thioate-DNA, phosphorothioate DNA; thioate-2′F-ANA, phosphorothioate 2′-deoxy2′fluoroarabinonucleic acid; DNA, deoxyribonucleic acid; thioate-LNA, phosphorothioate locked nucleic acid. bData not available. cReported for the mixed base deoxy 9mer with 3 dispersed bc-ANA inserts; note that contiguous bc-ANA insertion enhances duplex stability up to +1.7°C / mod. relative to thioate-DNA.
Synthesis of 2′-Deoxy-2′-fluoroβ-D-oligoarabinonucleotides (2′F-ANA)
An important parameter that governs overall duplex geometry is the conformations adopted by the individual sugars in the AON. To this end, modifications at the 2′-position have proven most valuable as tools for influencing ring conformational equilibria. Additionally, much of the literature devoted to antisense modifications has shown that crude predictions of sugar gauche and anomeric effects can be made with some accuracy as to the effect a particular 2′-modification exerts on the pseudosugar equilibrium (e.g., Plavec et al., 1994; Thibaudeau et al., 1994; Damha et al., 1995; Thibaudeau and Chattopadhyaya, 1997; Noronha and Damha, 1998; Wilds and Damha, 1999). The authors’ evaluation of these effects indicated to them that a switch in chirality of the α-2′-OH group in ribose residues to the β-configuration in the arabinose epimers should drive
the N-to-S equilibrium to the southern hemisphere and enforce a noncanonical A:B topology in hybrids with RNA, in common with the native RNA:DNA hybrids. It is noteworthy that most 2′-modifications do not activate RNase H, as their double-stranded helices with RNA are tailored to adopt a pure A form—i.e., RNA:RNA-like (for reviews, see Manoharan, 1999, and references therein). In fact, arabinonucleic acids (ANAs) represent the first examples of sugar-modified antisense agents capable of eliciting RNase H–mediated hydrolysis of target RNA. Additionally, ANAs exhibit adequate hydrolytic stability as a result of the trans relationship of the 2′- and 3′-OH groups, which effectively retards internal nucleophilic participation of the 2′-OH group in chain isomerization and/or phosphodiester cleavage. Despite these appealing properties, oligoarabinonucleotides of
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A MMTrO
O
B
MMTrO
O F
B
O
O O P O
O
X−
B
O P O
O F
X−
O
B
O
X = O, DNA X = S, S-DNA
X = O, 2’F-ANA X = S, S-2’F-ANA
B enzyme-active AON segment
HO O− O P O O
B O
O O P O O−
enzyme-active AON segment
3
enzyme-active AON segment
O O P O O− 4
O O P O O−
enzyme-active AON segment
Figure 4.15.5 (A) DNA and 2′F-ANA primary structures and backbone chemistries. (B) Illustration of linker-modified DNA or 2′F-ANA antisense constructs. Panel B reprinted from Mangos and Damha (2002) with permission from Bentham Science Publishers.
mixed base composition display weaker binding affinities relative to the native DNA:RNA heteroduplexes (Giannaris and Damha, 1994; Damha et al., 1998; Noronha et al., 2000). The observed thermal destabilization may arise from cis-placement of the sterically demanding β-C2′-OH group with respect to the base in the major groove of the helix, which purportedly weakens local π-stacking and/or normal Watson-Crick pairing by distorting the N-glycosyl orientation (Noronha et al., 2000). Alternatively, molecular dynamics simulations have suggested the arabinose sugars to be held in C2′-endo geometries by probable intrasugar hydrogen bonding between the 2′-α-substituent and the C5′-oxygen (Venkateswarlu and Ferguson, 1999). The strand topology is consequently forced to adopt a B-like conformation, which is disfavored for binding with RNA. This, however, contradicts recent NMR analyses (Trempe et al., 2001), which show a clear bias of the sugar conformation toward the eastern (O4′-endo) hemisphere and a lack of C2′OH/O5′ interactions in the arabinose sugars (Damha et al., 2001). Interestingly, the CD spectral profiles of DNA:RNA and ANA:RNA complexes reveal similar duplex helicities; however, the ability of the latter to direct RNase H–mediated phosphodiester hydrolysis of the target occurs with comparably lower efficiencies. This lower
processivity of the enzyme toward ANA:RNA hybrids may arise from a weaker tenacity of ANA for RNA targets and a correspondingly lower effective concentration of substrate duplex. Accordingly, the extent of cleavage increases significantly when substrate and enzyme are incubated at ambient rather than physiological temperatures (Noronha et al., 2000). Apart from conferring greater glycosyl stability towards potential depurination, a β-F atom in place of the ara-2′-OH group (i.e., 2′F-ANA) markedly increases the duplex melting temperature (Damha et al., 1998), imparting higher stability than hybrids of RNA and ANA, thioate-DNA or even DNA (Table 4.15.3). The origin of these effects likely stems from greater strand preorganization of the fluorinated strands relative to their singlestranded DNA counterparts. In fact, simple substitution with the more electronegative F atom almost exclusively forces the arabinosugar equilibrium to adopt an O4′-endo (or eastern) conformation (Berger et al., 1998; Trempe et al., 2001). Like the arabinose compounds, this predominance over a C2′-endo geometry is presumably forced to occur in order to minimize steric conflicts between the F atom and the proximal heterocycle (Berger et al., 1998). Rather, the 2′-substituent is conveniently accommodated in the major groove
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Synthesis of 2′-Deoxy-2′-fluoroβ-D-oligoarabinonucleotides (2′F-ANA)
by assuming a more eastern geometry, which displaces it sufficiently far from the enzyme’s locus of action while retaining other geometrical elements necessary for activity. In fact, 2′F-ANA, exhibits superior RNase H competency relative to ANA, probably by virtue of the sterically innocuous properties of fluorine and the higher thermal stability of FANA/RNA hybrids (Damha et al., 2001). Although much conformational work has been directed to the phosphodiester analogs (vide supra), many of these observations can be extended to 2′F-ANA oligomers with phosphorothioate (S) scaffolds. The dominant influence of fluorine on sugar-pucker profiles and subsequent enzyme hydrolytic susceptibility is retained in S-FANA (Lok et al., 2002), in which additional nuclease resistance is provided to the 2′-modified oligomers by the thioate backbone (reviewed by Cook, 1998). However, uniformly modified S-FANA containing hybrids are suboptimally recognized by RNase H as compared to their phosphodiester derivatives, but can greatly be improved with deoxy incorporation (Lok et al., 2002). Likewise, other oligonucleotide modifications that combine intervening enzyme-active AON segments with conformationally restricted “flanking” residues usually support cleavage of their intended targets. The peripheral segments also serve to provide added nuclease resistance to the core segment, which is usually a stretch of nuclease-labile deoxynucleotides. Although the entropic consequences imposed by the flanking segments on the antisense strand endow it with favorable duplexation properties, RNase-H-induction is invariably compromised (Shen et al., 1998). Accordingly, non-RNase H competent flanking backbone segments (e.g., 2′OMe-RNA) show strong dependencies on gap size, with a minimal requirement of six to eight contiguous deoxynucleotides within the core to elicit observable activities. These requirements are significantly relaxed for SFANA in which insertion of even a single deoxynucleotide as the “gap” restores high in vitro activities. In fact, fluoroarabino-deoxy chimeras comprising four or more neighboring deoxynucleotides exhibit remarkably greater potencies than uniform S-DNA sequences and show greater inhibition of luciferase expression in HeLa cells without affecting cell viability (Lok et al., 2002). Furthermore, cleavage occurs throughout the entire RNA chain, rather than specifically within the gap as for the methylated ribosyl congeners, which adopt a characteristic C3′-endo pucker and likely induce
local helical deformations spanning the methoxy regions of the AON:RNA chimera hybrids. These results further highlight the pivotal importance of the O4′-endo antipodal conformation as a major determinant for RNase H recognition. Consequently, the most promising antisense agents are hypothesized to be those in which the arabinose configuration at C2′ is retained (Venkateswarlu and Ferguson, 1999). However, other carbohydrate modified ANA derivatives with this same pseudosugar disposition have displayed poor RNase H–associated hydrolysis of target RNA. Significantly, various bicyclic ANAs possess an O4′-endo bias and could hypothetically induce RNase H at high concentrations, but generally show abolished activities under conditions representative of the intracellular environment (Minasov et al., 2000). Interestingly, the inherent flexibility of the sugar conformations strongly correlate with the relative processivity of the enzyme toward hybrids of ANA-derived AON and RNA according to the following: DNA (flexible) > 2′F-ANA/ANA >> bc-ANA (rigid). These trends, together with the activity enhancements observed upon introducing relatively flexible deoxy residues in the more rigid FANA strand (vide supra), prompted the authors to investigate the potential synergistic attributes of intermingling sites of flexibility within the 2′FANA polymers. Indeed, the incorporation of acyclic linkers with high degrees of flexibility in the AON strand, either a 2′,5′-linked secouridine residue S.3 or a butanediol linker S.4 (Figure 4.15.5), appear to better accommodate the stringent conformational requirements of the enzyme without disrupting other fundamental recognition elements within the enclosing AON segments (Mangos et al., unpub. observ.). Sizable increases in enzyme-mediated scission of target RNA occur upon substituting an acyclic linker in the middle of a known RNase H–competent analog (i.e., DNA or 2′F-ANA). Moreover, the susceptibility of the hybrid to RNA hydrolysis remains fully operative in other sequence contexts with up to eight-fold enhancements over strands lacking the acyclic insertion. For example, in sequences wherein the acyclic inserts are moved along the 2′F-ANA scaffold, a significant change in activity is observed that depends specifically on the insertion site. The greatest activity occurs in constructs with a centrally placed acyclic linker, although differences between 5′- versus 3′-end insertion are also observed, with a greater amount of enzyme proc-
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essing occurring when the linker is placed near the 3′-end of the AON. The authors therefore speculate that RNase H induction can remarkably be improved by introducing subtle changes in local 2′F-ANA strand dynamics that enable better adherence and/or processing of the hybrid substrate by the enzyme. These and future studies with 2′F-ANA and their constructs should provide considerable insights as to the structural factors that comprise the “optimal” AON/RNA substrate. For these reasons, the protocols of this unit provide a method of quickly and conveniently preparing 2′-fluoroarabino oligonucleotides with any one or all of the structural modifications described herein. Although linker technologies are not discussed, these are quite simple to implement and exemplify the diversity with which the protocols can be applied to new AON designs that further exploit the interesting properties of 2′F-ANA.
Critical Parameters The synthesis of araF-phosphoramidites, as for other phosphoramidites, is very moisture and acid sensitive. Therefore, all reagents and apparatus should be anhydrous. If the product has to be isolated by flash column chromatography (APPENDIX 3E), it is absolutely necessary to include triethylamine in all solvent systems used to equilibrate the column and for subsequent sample elution. Heating the product above 40°C is not recommended during evaporation. If the phosphoramidite has been stored for a long period of time prior to oligomer synthesis, its purity should be verified by TLC (APPENDIX 3D) or 31P NMR. If necessary, the phosphoramidite can be repurified by flash chromatography. Phosphoramidite oligonucleotide chemistry is extremely water sensitive. All coupling reagents should be absolutely dry, and fresh reagents should be prepared as these are critical to the success of the synthesis. It is not recommended to keep the reagents on the synthesizer for longer than 1 week. Furthermore, the synthesizer should be in good working condition. All lines should be purged with each reagent and solvent prior to starting a synthesis and cleaned with acetonitrile afterward to prevent crystallization and blocking of the lines. If synthesis is performed on the Expedite instrument, a relatively low concentration of phosphoramidites can be used without loss of coupling efficiency. For example, commercial RNA protocols require 50 mg/mL phosphoramidite concentration, which is almost 3-
fold lower than for RNA synthesis on older instruments (e.g., 1.5 M amidite solutions are required for use with the ABI 381A DNA synthesizer). Phosphoramidite and activator solutions are continuously delivered to the column during the coupling step. Such permanent delivery is performed by slow pulses of reagents to the column, which permanently renew them and thereby increase the efficiency of the synthesis. If a synthesizer with stationary delivery of reagents is to be used, the amidite concentration should be increased to that used for RNA synthesis (or slightly lower). The authors were able to successfully synthesize diester araF oligonucleotides on the ABI 381A DNA instrument using 0.1 to 0.15 M concentrations of phosphoramidites. Many different sulfur-transfer reagents for S-DNA have been documented in the literature over the last several years. However, none of these reagents are able to give 100% P-S oxidation. As a consequence, a small percentage of phosphodiester bonds are inevitably present in S-oligonucleotides. The diester fragments are much less stable to hydrolysis, making them more susceptible to nucleolytic cleavage in cell culture, which limits certain biological applications of these antisense oligonucleotides. Fortunately, pure S-oligonucleotides can easily be analyzed and purified from P-O containing S-oligonucleotides by anion-exchange HPLC (Bergot and Egan, 1992). This method enables the isolation of S-oligonucleotides that contain even a single P-O linkage within the strand. The percentage of P-O bonds in the product is dependent on the source of sulfurizing reagent used, as different sources may lead to different percentages in the fully synthesized oligomers. Although undesired oxidation is not very problematic in S-DNA synthesis, 2′F-ANA oligonucleotides are intrinsically more difficult to sulfurize as a consequence of the electron-withdrawing effects of the fluorine substituent in the sugars. This limitation may be overcome by increasing the concentration of the sulfur-transfer reagent, accompanied by extended reaction times in order to obtain S-oligonucleotides of better quality. The authors of the current protocol applied several sulfurization reagents to S-araF synthesis, namely the Beaucage reagent, EDITH, and ADTT, and have found that the former normally gives a higher percentage of P-O bonds relative to the latter two reagents. EDITH and ADTT work very similarly and give oligomers of low P-O compositions. Figure 4.15.2 shows a chromatographic profile for a crude S-DNA/S-araF chimera synthesized
Synthesis of Modified Oligonucleotides and Conjugates
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using either the Beaucage reagent or ADTT. As is shown, the S-FANA oligonucleotide synthesized with ADTT displays more uniform P-S incorporation (i.e., P-O insertions are minimal in this oligomer as detected by chromatography). However, EDITH and ADTT reagents are very expensive and not widely available commercially, which makes the Beaucage reagent a suitable alternative. It should also be emphasized that the limitations observed with the Beaucage reagent are reserved only for 2′FANA polymers; experimentally, sulfurization with this reagent proceeds much more efficiently with other types of modified oligonucleotides (e.g., S-DNA), thereby making it the reagent of choice for those applications. The amount of contaminating P-O linkages within S-FANA oligonucleotides does not typically exceed 1% per phosphate, even when using the Beaucage reagent, but may sometimes be significantly larger. Unfortunately, PO insertions cannot be detected by PAGE. Oligonucleotides with multiple phosphodiester bonds will migrate as a single narrow band through the gel, but present multiple peaks upon anion-exchange chromatographic analysis. Consequently, chromatographic purification of S-FANA oligonucleotides is strongly recommended. In general, both HPLC and PAGE purification can be applied to araF oligonucleotides. Sequences that consist entirely of phosphodiester backbones may be purified either by HPLC or PAGE, depending upon the scale used in the synthesis. If 1 to 20 A260 units are required, PAGE purification is sufficient; for preparative amounts ranging from 100 to 200 A260 units of purified oligomer, HPLC is recommended. For S-FANA oligonucleotides, ion-exchange purification is most convenient, regardless of the amount to be purified.
Troubleshooting
Synthesis of 2′-Deoxy-2′-fluoroβ-D-oligoarabinonucleotides (2′F-ANA)
Low coupling efficiency. Several conditions can cause poor monomer coupling yields. (1) The reagents and acetonitrile may contain water. This is minimized by drying the phosphoramidites and tetrazole in a vacuum desiccator over phosphorus pentoxide for one or two days prior to use. (2) Lines in the synthesizer may be partially blocked. Check delivery of each reagent to the lines by monitoring flow rates. Perform routine DNA synthesis first and monitor coupling efficiencies by trityl analysis. HPLC analysis of S-araF oligonucleotide gives multiple product peaks. This problem is usually caused by incomplete sulfurization, es-
pecially if the Beaucage reagent has been used. If so, the product may still migrate as one band under PAGE conditions. This can be circumvented by increasing the concentration of the Beaucage reagent or using another sulfurizing reagent. Beaucage reagent precipitates. The Beaucage reagent is known to be extremely sensitive to the quality of solvent and is decomposed rather easily. If this process is rapid, a switch to an acetonitrile source of better quality is recommended. Furthermore, the glass bottle for Beaucage reagent should be silanized by the investigator (APPENDIX 2A; Iyer et al., 1990) or obtained in silanized form from various commercial sources. Alternatively, acetonitrile solutions of Beaucage reagent can be placed in a 15-mL plastic conical tube, which is then deposited within an appropriate glass bottle and attached to the gene-machine to prevent direct contact of the reagent with the glass.
Anticipated Results Using araF phosphoramidites as building blocks in conjunction with the methods presented here, it should be possible to routinely obtain araF oligonucleotides with yields ranging from 120 to 180 A260 units of crude material for 1-µmol scale syntheses, and isolated yields of ∼30 to 100 A260 units. The sulfurization step with EDITH or ADTT reagents helps to minimize the extent of P-O insertions in S-oligonucleotides. These considerations, together with the optimized ion-exchange HPLC chromatographic procedures, should enable the facile isolation of araF oligonucleotides of high purity.
Time Considerations Preparation of araF phosphoramidites from the appropriately protected nucleosides usually takes ∼1 day per monomer. Depending on the type of chemistry (diester- versus thio-araF-oligonucleotides, or DNA/araF oligonucleotide chimeras), the time required for assembly of a 20-mer oligonucleotide (typical for antisense applications) on the Expedite instrument is from 6 to 12 hr. An additional 2 days are required to perform deprotection, followed by 3 to 5 days for isolation, desalting, and final analysis as described (see Basic Protocol 3), if purification is required.
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Literature Cited Altmann, K.-H., Kesselring, R., Francotte, E., and Rihs, G. 1994a. 4′,6′-Methano carbocyclic thymidine: A conformationally constrained building block for oligonucleotides. Tetrahedron Lett. 35:2331-2334. Altmann, K.-H., Imwinkelried, M., Kesselring, R., and Rihs, G. 1994b. 1′,6′-Methano carbocyclic thymidine: Synthesis, X-ray crystal structure, and effect on nucleic acid duplex stability. Tetrahedron Lett. 35:7625-7628. Berger, I., Tereshko, V., Ikeda, H., Marquez, V.E., and Egli, M. 1998. Crystal structures of B-DNA with incorporated 2′-deoxy-2′-fluoro-arabinofuranosyl thymines: Implications of conformational preorganization for duplex stability. Nucl. Acids Res. 26:2473-2480. Bergot, B.J. and Egan, W. 1992. Separation of synthetic phosphorothioate oligonucleotides from their oxygenated (phosphodiester) defect species by strong-anion-exchange high-performance liquid chromatography. J. Chromatogr. 599:35-42. Christensen, N.K., Petersen, M., Nielson, P., Jacobsen, J.P., Olsen, C.E., and Wengel, J. 1998. A novel class of oligonucleotide analogues containing 2′-O,3′-C-linked [3.2.0]bicycloarabinonucleoside monomers: Synthesis, thermal affinity studies and molecular modeling. J. Am. Chem. Soc. 120:5458-5463. Cook, P.D. 1998. Second generation antisense oligonucleotides: 2′-modifications. Annu. Rep. Med. Chem. 33:313-325. Crouch, R.J. and Toulmé, J.J. (eds.) 1998. Ribonucleases H. INSERM, Paris. Damha, M.J., Meng, B., Yannopoulos, C.G., Wang, D., and Just, G. 1995. Structural basis for the RNA selectivity of oligonucleotides containing alkylsulfide internucleoside linkages and 2′-Osubstituted 3′-deoxyribose. Nucl. Acids Res. 19:3967-3973. Damha, M.J., Wilds, C.J., Noronha, A., Brukner, I., Borkow, G., Arion, D., and Parniak, M.A. 1998. Hybrids of RNA and arabinonucleic acids (ANA and 2′F-ANA) are substrates of ribonuclease H. J. Am. Chem. Soc. 120:12976-12977. Damha, M.J., Noronha, A.M., Wilds, C.J., Trempe, J.-F., Denisov, A., and Gehring, K. 2001. Properties of arabinonucleic acids (ANA & 2′FANA): Implications for the design of antisense therapeutics that invoke RNase H cleavage of RNA. Nucleosides Nucleotides 20:429-440. Giannaris, P.A. and Damha, M.J. 1994. Hybridization properties of oligoarabinonucleotides. Can. J. Chem. 72:909-918. Iyer, R.P., Phillips, L.R., Egan, W., Regan, J.B., and Beaucage, S.L. 1990. The automated synthesis of sulfur-containing oligodeoxyribonucleotides using 3H-1,2-benzodithiol-3-one 1,1-dioxide as a sulfur-transfer reagent. J. Org. Chem. 55:46934699.
Lebedeva, I. and Stein, C.A. 2001. Antisense oligonucleotides: Promise and reality. Annu. Rev. Pharmacol. Toxicol. 41:403-419. Lima, W.F. and Crooke, S.T. 1997. Binding affinity and specificity of Escherichia coli RNase H1: Impact on the kinetics of catalysis of antisense oligonucleotide-RNA hybrids. Biochemistry 36:390-398. Lok, C.-N., Viazovkina, E., Min, K-L., Nagy, E., Wilds, C.J., Damha, M.J., and Parniak, M.A. 2002. Potent gene-specific inhibitory properties of mixed-backbone antisense oligonucleotides comprised of 2′-deoxy-2′-fluoro-D-arabinose and 2′-deoxyribose nucleotides. Biochemistry 41:3457-3467. Mangos, M.M. and Damha, M.J. 2002. Flexible and frozen sugar-modified nucleic acids: modulation of biological activity through furanose ring dynamics in the antisense strands. Curr. Topics Med. Chem. 2:1145-1169. Manoharan, M. 1999. 2′-Carbohydrate modifications in antisense oligonucleotide therapy: Importance of conformation, configuration and conjugation. Biochim. Biophys. Acta 1489:117130. Minasov, G., Teplova, M., Nielsen, P., Wengel, J., and Egli, M. 2000. Structural basis of cleavage by RNase H of hybrids of arabinonucleic acids and RNA. Biochemistry 39:3525-3532. Myers, N.M. and Dean, K.J. 2000. Sensible use of antisense: How to use oligonucleotides as research tools. TIPS 21:19-23. Nakamura, H., Oda, Y., Iwai, S., Inoue, H., Ohtsuka, E., Kanaya, S., Kimura, S., Katsuda, C., Katayanagi, K., Morikawa, K., Miyashiro, H., and Ikehara, M. 1991. How does RNase H recognize a DNA:RNA hybrid? Proc. Natl. Acad. Sci. U.S.A. 88:11535-11539. Noronha, A. and Damha, M.J. 1998. Triple helices containing arabinonucleotides in the third (Hoogsteen) strand: Effects of inverted stereochemistry at the 2′-position of the sugar moiety. Nucl. Acids Res. 26:2665-2671. Noronha, A.M., Wilds, C.J., Lok, C.-N., Viazovkina, K., Arion, D., Parniak, M.A., and Damha, M.J. 2000. Synthesis and biophysical properties of arabinonucleic acids (ANA): Circular dichroic spectra, melting temperatures and ribonuclease H susceptibility of ANA:RNA hybrid duplexes. Biochemistry 39:7050-7062. Oda, Y., Iwai, S., Ohtsuka, E., Ishikawa, M., Ikehara, M., and Nakamura, H. 1993. Binding of nucleic acids to E. coli RNase HI observed by NMR and CD spectroscopy. Nucl. Acids Res. 21:46904695. Plavec, J., Thibaudeau, C., and Chattopadhyaya, J. 1994. How does the 2′-hydroxy group drive the pseudorotational equilibrium in nucleoside and nucleotide by the tuning of the 3′-gauche effect? J. Am. Chem. Soc. 116:6558-6560. Synthesis of Modified Oligonucleotides and Conjugates
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Sangvhi, Y.S. 1998. Synthesis of nitrogen containing linkers for antisense oligonucleotides. In Carbohydrate Mimics (Y. Chapleur, ed.) pp. 523536. Wiley-VCH, Germany. Schmit, C., Bèvierre, M-O., De Mesmaeker, A., and Altmann, K.-H. 1994. The effects of 2′- and 3′-alkyl substituents on oligonucleotide hybridization and stability. Bioorg. Med. Chem. Lett. 4:1969-1974. Shen, L.X., Kandimalla, E.R., and Agrawal, S. 1998. Impact of mixed-backbone oligonucleotides on target binding affinity and target cleaving specificity and selectivity by E. coli RNase H. Bioorg. Med. Chem. 6:1695-1705. Sørensen, M.D., Kvaernø, L., Bryld, T., Håkansson, A.E., Verbeure, B., Gaubert, G., Herdewijn, P., and Wengel, J. 2002. Alpha-L-ribo-configured locked nucleic acid (alpha-L-LNA): Synthesis and properties. J. Am. Chem. Soc. 124:21642176. Still, W.C., Kahn, M., and Mitra, A. 1978. Rapid chromatographic technique for preparative separation with moderate resolution. J. Org. Chem. 43:2923-2925. Tang, J.-Y., Han Y., Tang, J.X., and Zhang, Z. 2000. Large scale synthesis of oligonucleotide phosphorothioates using amino-1,2,4-dithiazoline-5thione as an efficient sulfur-transfer reagent. Org. Proc. Dev. 4:194-198.
Wilds, C.J. and Damha, M.J. 1999. Duplex recognition by oligonucleotides containing 2′-Deoxy2′-fluoro-D-arabinose and 2′-deoxy-2′-fluoroD-ribose. Intermolecular contacts versus sugar puckering in the stabilization of triple helical complexes. Bioconjug. Chem. 10:299-305. Wilds, C.J., and Damha M.J. 2000. 2′-deoxy-2′fluoro-β-D-arabinonucleosides and oligonucleotides (2′F-ANA): Synthesis and physicochemical studies. Nucl. Acids Res. 28:36253635. Xu, Q., Musier-Forsyth, K., Hammer, R.P., and Barany, G. 1996. Use of 1,2,4-dithiazolidine3,5-dione (DtsNH) and 3-ethoxy-1,2,4-dithiazoline-5-one (EDITH) for synthesis of phosphorothioate-containing oligodeoxyribonucleotides. Nucl. Acids Res. 24:1602-1607.
Key References Crouch and Toulmé, 1998. See above A comprehensive book on ribonucleases H, their sources, properties, biological utility, and antisense applications.
Thibaudeau, C. and Chattopadhyaya, J. 1997. The discovery of intramolecular stereoelectronic forces that drive the sugar conformation in nucleosides and nucleotides. Nucleosides Nucleotides 16:523-529.
Damha et al., 2001. See above.
Thibaudeau, C., Plavec, J., Garg, N., Papchikhin, A., and Chattopadhyaya, J. 1994. How does the electronegativity of the substituent dictate the strength of the gauche effect? J. Am. Chem. Soc. 116:4038-4043.
Freier, S.M. and Altmann, K.-H. 1997. The ups and downs of nucleic acid duplex stability: Structurestability studies on chemically-modified DNA:RNA duplexes. Nucl. Acids Res. 25:44294443.
Trempe, J.F., Wilds, C.J., Denisov, A.Y., Pon, R.T., Damha, M.J., and Gehring, K. 2001. NMR solution structure of an oligonucleotide hairpin with a 2′F-ANA/RNA stem: Implications for RNase H specificity toward DNA/RNA hybrid duplexes. J. Am. Chem. Soc. 123:4896-4903.
An extensive resource that relates duplex stabilities to nucleotide structure with 197 examples of oligonucleotide modifications.
Uhlmann, E. and Peyman, A. 1990. Antisense oligonucleotides: A new therapeutic principle. Chem. Rev. 90:543-584. Venkateswarlu, D. and Ferguson, D.M. 1999. Effects of C2′-substitution on arabinonucleic acid structure and conformation. J. Am. Chem. Soc. 121:5609-5610. Walder, R.Y. and Walder, J.A. 1988. Role of RNase H in hybrid-arrested translation by antisense oligonucleotides. Proc. Natl. Acad. Sci. U.S.A. 85:5011-5015.
Synthesis of 2′-Deoxy-2′-fluoroβ-D-oligoarabinonucleotides (2′F-ANA)
Wengel, J., Koshkin, A., Singh, S.K., Nielsen, P., Meldgaard, M., Rajwanshi, V.K., Kumar, R., Skouv, J., Nielsen, C.B., Jacobsen, J.P., Jacobsen, N., and Olsen, C.E. 1999. LNA (Locked nucleic acid). Nucleosides Nucleotides 18:13651370.
Wang, J., Verbeure, B., Luyten, I., Lescrinier, E., Froeyen, M., Hendrix, C., Rosemeyer, H., Seela, F., Aerschot, A.V., and Herdewijn, P. 2000. Cyclohexene nucleic acids (CeNA): Serum stable oligonucleotides that activate RNase H and increase duplex stability with complementary RNA. J. Am. Chem. Soc. 122:8595-8602.
Highlights the role of the 2′-sugar position in ANA and 2′F-ANA conformations, and the origins of RNase H activity.
Kvaernø, L. and Wengel, J. 2001. Antisense molecules and furanose conformations—is it really that simple? Chem. Commun. 1419-1424. A concise review briefly comparing RNA binding versus cleavage with promising antisense candidates. Lebedeva and Stein, 2001. See above. Insightful discussion of antisense design that focuses on in vivo toxicity from nonspecific interactions and potential irrelevant cleavage of nontargeted RNA.
Contributed by Ekaterina Viazovkina, Maria M. Mangos, Mohamed I. Elzagheid, and Masad J. Damha McGill University Montreal, Canada
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Chemistry of CpG DNA The innate immune system of vertebrates has evolved to recognize specific pathogenassociated molecular patterns (PAMPs) present in invading microorganisms through pattern recognition receptors (PRRs; Lien and Ingalls, 2002). One of several such PAMPs is the unmethylated CpG dinucleotide present in specific sequence contexts (CpG motifs) in pathogenic microorganisms (Hemmi et al., 2000). On sensing a CpG motif, PRRs trigger complex signal transduction pathways that ultimately activate a number of transcription factors, including NF-κB and AP-1. These induce specific patterns of gene expression associated with the development and maintenance of immune responses. The immune responses to bacterial and synthetic oligonucleotides containing CpG motifs include proliferation of B cells, production of cytokines IL-12, γ-IFN, IL-6, and TNF-α (Klinman et al., 1996; Zhao et al., 1997), and production of costimulatory molecules by monocytes/macrophages, B cells, dendritic cells (DCs), and natural killer (NK) cells.
HISTORY Tokunaga and coworkers were the first to report that DNA from Mycobacterium bovis induces production of interferons α, β, and γ (IFN-α, -β, and -γ), augments NK cell activity, and shows antitumor activity (Tokunaga et al., 1984). The same authors, using short synthetic DNA fragments, showed that palindromic sequences containing CpG dinucleotides efficiently induce NK cell activity and induce IFNs (Yamamoto et al., 1992a). Subsequent studies demonstrated that bacterial DNA, but not mammalian DNA, induces murine B cell proliferation and cytokine secretion (Messina et al, 1991; Yamamoto et al., 1992b). Later studies showed that unmethylated CpG dinucleotides in specific sequence contexts present in bacterial DNA, synthetic oligodeoxyribonucleotides, and DNA vaccines are responsible for the observed immune responses (Krieg et al., 1995; Sato et al., 1996). Compared to bacterial DNA, the occurrence of CpG motifs in vertebrate DNA is sparse. In addition, the vertebrate C residues usually carry a 5-methyl substituent (Bird, 1986), which enables the immune system to distinguish between its own DNA and bacterial DNA that signals an infection.
UNIT 4.16
RECEPTORS The recognition of CpG DNA has been shown to occur through Toll-like receptor 9 (TLR9), which belongs to a family of proteins called TLRs (Hemmi et al., 2000). The TLRs function as PRRs to initiate the innate immune response against invading microorganisms. However, direct evidence for binding of CpG DNA to TLR9 has not yet been documented. Although most other TLRs are membrane receptors, growing evidence suggests that TLR9 is localized in the cytoplasm (Takeshita et al., 2001; Ahmad-Nejad et al., 2002). Therefore, cellular uptake and endosomal localization seem to be prerequisites for CpG DNA recognition (Stacey et al., 2000).
Sequence and Structural Specificity of TLR9 TLR9 proteins from different vertebrates vary in the CpG DNA sequences they recognize (Bauer et al., 2001a). Therefore, a CpG motif that is active in one species may not be in another. For example, mouse TLR9 prefers an unmethylated CpG dinucleotide flanked by two purine bases on the 5′ side and two pyrimidine bases on the 3′ side, such as GACGTT (Krieg et al., 1995). CpG dinucleotides preceded by C or followed by G are generally less active in mice. Human immune cells respond optimally to GTCGTT or TTCGTT motifs (Hartmann et al., 2000). Certain other sequences, such as a palindromic AACGTT sequence, are known to induce immune responses in both mouse and human systems (Yamamoto et al., 1992a; Van Uden and Raz, 2000). Thus, TLR9 variants from different species recognize CpG dinucleotides flanked by a variety of sequences, though to different extents (Rankin et al., 2001; Zhang et al., 2001; Wernette et al., 2002).
Signaling Events Though the evidence suggests that TLR9 is the receptor for phosphorothioate CpG DNAs, the role of TLR9 in mediating the observed effects of phosphodiester CpG oligonucleotides and bacterial DNA is not yet clear. The signaling events initiated by TLR9 in response to CpG DNA include recruitment of MyD88, IRAK, and TRAF6, which leads to activation of IκB kinase, MAP kinase, the stress kinases JNK-1 and -2, and p38, which in turn causes activation of transcription factors ATF-
Contributed by Ekambar R. Kandimalla and Sudhir Agrawal Current Protocols in Nucleic Acid Chemistry (2003) 4.16.1-4.16.14 Copyright © 2003 by John Wiley & Sons, Inc.
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2, AP-1, and NF-κB (Fig. 4.16.1). The activated transcription factors induce the synthesis of several regulatory cytokines and costimulatory molecules.
SIGNIFICANCE OF CpG DINUCLEOTIDES AND CHEMISTRY OF CpG DNA A CpG dinucleotide present in specific sequence contexts is essential for immunostimulatory activity, whereas the inverted dimer, GpC, is inactive. As discussed above, the flanking sequences play an important role in determining activity of CpG DNA (Yamamoto et al., 1992a; Krieg et al., 1995; Pisetsky, 1999). Recently, other structural features that influence the activity of synthetic oligonucleotides have been reported, including the nature of the internucleotide linkage, the nature and conformation of the sugar ring, modification or removal of nucleobases, accessibility of the 5′ end, and the nature and size of any 5′-terminal blocking group.
DNA Backbone Shorter CpG DNA molecules with unmodified phosphodiester backbones may elicit potent immune stimulation in vitro (Sonehara et al., 1996; Iho et al., 1999). However, phosphorothioate-modified oligonucleotides are used more commonly to prevent rapid degradation by nucleases present in cells. Although phosphodiester and phosphorothioate CpG DNAs elicit superficially similar immune re-
sponses, recent studies showed certain distinct differences in the actions of these two backbones (Ballas et al., 2001; Rothenfusser et al., 2001; Verthelyi et al., 2001, 2002; Dalpke et al., 2002; Gursel et al., 2002), which are not well understood (Fig. 4.16.2 and Table 4.16.1). Phosphodiester backbone Phosphodiester CpG DNAs containing palindromic structures and/or poly(dG) sequences effectively activate NK cells (Yamamoto et al., 1992a, 2000; Iho et al., 1999) and induce IFN-α/β production from plasmacytoid DCs (Bauer et al., 2001b; Kadowaki et al., 2001; Krug et al., 2001a,b). The effects of phosphodiester CpG DNAs are generally similar to those of bacterial DNA, and the induction of type I IFN is far greater than with phosphorothioate CpG DNAs. However, B cell activation by phosphodiester CpG DNA appears minimal compared with bacterial and phosphorothioate CpG DNAs. Phosphodiester CpG DNAs induce high levels of IFN-γ and IL-12, but produce IL-6 only minimally. Phosphorothioate substitutions on either end of the phosphodiester CpG DNA improve resistance to nucleases (Dalpke et al., 2002). Incorporation of poly(dG) sequences at the ends of the phosphodiester DNA has been reported to enhance nuclease stability and increase cellular uptake through scavenger receptors (Pearson et al., 1993; Kimura et al., 1994; Agrawal et al., 1 996). CpG DNAs containing phosphorothioate poly(dG) sequences at the 5′ and
cytosol
CpG DNA bacterial DNA (CpG) CpG oligonucleotides plasmid DNA (CpG)
TIR domain
MAPK (p38, ERK, JNK)
TLR9
AP-1
TRAF6 IRAK ds viral RNA [poly(I⋅C)]
nucleus
TIR domain TLR3 MyD88
IκB NF-κB
Chemistry of CpG DNA
Figure 4.16.1 The two members of the Toll-like receptor (TLR) family that recognize bacterial CpG DNA and double-stranded (ds) viral RNA. Key components involved in the signaling pathways are shown. The activated transcription factors ultimately upregulate the expression of a number of cytokines and costimulatory molecules. Abbreviations: AP-1, activator protein 1; ERK, extracellular-signal-regulated protein kinase; IκB, inhibitor of κB; IRAK, IL-1 receptor associated kinase; JNK, c-jun N-terminal kinase; MAPK, mitogen activated protein kinase; MyD88, myeloid differentiation factor 88; NF-κB, nuclear factor κB; TIR, Toll/IL-1 receptor; TLR, Toll-like receptor.
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R = O− phosphodiester
O
B
O O
R P O O
R = S− O
B
phosphorothioate
O
R = CH3 methylphosphonate
Figure 4.16.2 Chemical structure of a typical dinucleotide showing the internucleotide phosphate linkage (boxed) and the three backbone chemistries studied in CpG DNA for immunostimulatory activity. Some of the important immunostimulatory effects observed with each backbone modification are given in Table 4.16.2.
3′ ends and a palindromic phosphodiester in the middle are potent adjuvants in vitro and in vivo (Dalpke et al., 2002; Verthelyi et al., 2002). However, it is unclear whether activity is due to the phosphodiester backbone, the poly(dG) sequences, or secondary structures formed by palindromic and poly(dG) sequences. Recently, for the first time, the authors showed the use of phosphodiester CpG DNA without requiring palindromic structures and poly(dG) sequences for immunostimulatory activity (Yu et al., 2002a). For this study, two phosphodiester CpG oligodeoxynucleotides were attached via a glyceryl linker through their 3′ ends. These 3′-3′-linked CpG DNAs are referred to as immunomers. Phosphodiester immunomers showed remarkable stability against nucleases in medium containing 10% FBS that was not heat inactivated. Surprisingly, phosphodiester immunomers induced increased IL-12 secretion and minimal amounts of IL-6 secretion in mouse spleen cell cultures (Yu et al., 2002a). These studies suggest that it would be possible to modulate cytokine secretion profiles induced by CpG DNAs by using different backbone chemistries. In J774 cell cultures they activated NF-κB and induced cytokine secretion comparable to that of an unmodified 18-mer phosphorothioate CpG DNA containing the same CpG motif. Moreover,
phosphodiester immunomers showed antitumor activity in nude mice bearing human breast (MCF-7) and prostate (DU145) cancer xenografts, suggesting that single-stranded phosphodiester CpG DNA can be a potent pharmacological agent. Phosphorothioate backbone In contrast, phosphorothioate CpG DNAs do not require palindromic sequences to induce im mu ne respo nses. I n g en er al, phosphorothioate CpG oligonucleotides strongly stimulate B cell proliferation, as well as activate monocytes/macrophages, DCs, and B cells to produce cytokines and immunoglobulins (Branda et al., 1993; Krieg et al., 1995; Zhao et al., 1997; Stacey et al., 2000). Phosphorothioate CpG DNAs are the most extensively studied to date, and several are in clinical trials. Stereoenriched phosphorothioate backbone Substitution of a nonbridging oxygen with sulfur creates a chiral phosphorus center with Rp- and Sp-diastereomers. If there are n phosphorothioate linkages in the backbone, conventional automated synthesis yields 2n stereoisomers. Studies using stereoenriched all-Rp and all-Sp phosphorothioate CpG DNAs showed that the all-Rp analog induced lower cell pro-
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liferation than all-Sp or racemic CpG DNA analogs (Yu et al., 2000a). However, it is not clear if this is because of their differential susceptibility to nucleases (Tang et al., 1995; Yu et al., 2000a) or because one of the diastereomers is preferentially recognized by the receptor. Role of backbone charge and nonionic methylphosphonate linkages Substitution of a nonbridging oxygen on the internucleotide phosphate with a methyl group produces a nonionic phosphorus center (Fig. 4.16.2). Recent studies show that an uncharged
Table 4.16.1
methylphosphonate internucleotide linkage between C and G of the CpG dinucleotide diminishes the immune response (Zhao et al., 1996). Further, methylphosphonate substitutions within the three internucleotide linkages to the 5′-side of the CpG dinucleotide also suppress activity (Yu et al., 2001b). It appears that a negative charge on these internucleotide linkages is important for recognition and/or interaction between the CpG DNA and its receptor. Surprisingly, substituting the fifth or sixth linkage on the 5′ side of the CpG dinucleotide significantly enhances immunostimu-
Properties of DNA Backbones with Different Linkagesa
Linkage
Property
Phosphodiester
Usually require longer length palindromic sequences to be stable against nucleases May require higher concentrations to compensate for nuclease degradation Often require phosphorothioate-end modifications Poly(dG) end modifications are extensively used Produce high levels of IL-12, IFN-γ, and IFN-α/β, and induce minimal IL-6 Potent activators of NK and dendritic cells, but weak activators of B cells Produce TH1-type immune responses. Potent adjuvants, and antitumor and antiasthmatic agents Stable against nucleases Strong activators of B cells. Also directly activate macrophages/monocytes and DCs, but do not directly activate NK and T cells. Produce a number of cytokines, including IL-12 Produce Ig Produce TH1 immune responses. Currently investigated for their potential in clinical trials as anticancer, antiallergic, and antiinfectious agents Only site-specific modifications have been studied CH3 between C and G of the CpG dinucleotide suppresses the immune response CH3 within three internucleotide linkages to 5′ of CpG suppresses the response CH3 at five or six linkages from 5′ of CpG enhances the response CH3 in 3′ flanking sequences has minimal effect May be possible to alter cytokine secretion by site-specific incorporation of one or two linkages in CpG DNA
Phosphorothioate
Methylphosphonate
aRefer to Figure 4.16.2 for graphical depiction of the linkages.
Chemistry of CpG DNA
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latory activity, which may reflect tighter binding to the receptor. In contrast, the presence of nonionic internucleotide linkages to the 3′ side of the CpG dinucleotide has an insignificant effect on activity.
2′-Sugar modifications Replacing either nucleoside in the CpG dinucleotide with one that is modified at the 2′-hydroxyl group (R in Fig. 4.16.3A) impedes immunostimulatory activity (Zhao et al., 1996). However, 2′-O-methyl- or 2′-O-methoxyethylribonucleosides in the sequences flanking the CpG dinucleotide have different effects depending on the position of substitution (Zhao et al., 1999; 2000). In general, 2′-O-alkylribonucleosides adjacent to the CpG dinucleotide on the 5′ side impair activity, while the same substitution on the 3′ side has minimal effect. Importantly, activity increases when the substitutions are incorporated distal to the CpG dinucleotide on either side (Zhao et al., 1999, 2000; Agrawal and Kandimalla, 2001). Hence, the effects on receptor binding due to conformational changes within ribose are strongly dependent on the position of substitution relative to the CpG dinucleotide.
Sugar Modifications While TLR9 has been shown to be involved in CpG DNA immune activation, a closely related family member, TLR3, has been reported to specifically recognize viral and synthetic double-stranded RNA and induce immune responses (Alexopoulou et al., 2001). Ribose and deoxyribose sugar moieties adopt 2′- and 3′-endo conformations (Fig. 4.16.3) that attribute distinct structural, physicochemical, and biological properties to RNA and DNA, respectively. It appears that the vertebrate innate immune system has evolved to recognize both of these nucleic acid structures from invading microorganisms, but through different receptors (Fig. 4.16.1). The authors of this unit have extensively studied the effects of sugar modifications in CpG DNA, as discussed below.
A
3′-Sugar modifications The incorporation of unnatural 3′-deoxynucleosides results in the formation of 2′-5′-inter-
B 5′
O
5′
B
O
O
O
O O
P
S−
O
O
B
O
O O
B
O
P
S−
O
R
O2′ O
O
B
B
O
R
S−
O
P
P O
O3′ O
P O
S− O
B
O3′ S−
O
P
S−
O
Figure 4.16.3 (A) Chemical structure of a DNA chain containing a single ribonucleoside (boxed). R is –OH (RNA), –OCH3 (2′-O-methyl RNA), or –OCH2CH2OCH3 (2′-O-methoxyethyl RNA). (B) Chemical structure of a DNA chain with a 3′-deoxyribonucleoside (R = –H) or 3′-O-methylribonucleoside (R = –OCH3) (boxed). Note that the incorporation of a 3′-deoxy- or 3′-O-methylribonucleoside results in a 2′-5′-linkage in an otherwise 3′-5′-linked DNA.
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3000
IL-12
2000
Cytokine (pg/mL)
1000
0 15000
IL-6
10000
5000 0 225
IL-10
150 75 0 2
1
3
M
5
4
6
CpG DNA
Figure 4.16.4 Effect of site-specific incorporation of a 3′-deoxyribonucleoside in different CpG DNA molecules (1 and 4) either in the 5′-flanking (2 and 5) or 3′-flanking sequence (3 and 6) on induction of cytokine (IL-12, -6, and -10) secretion in BALB/c mouse spleen cell cultures. Data for specific compounds is taken from Yu et al. (2002c). BALB/c mouse spleen cell cultures were incubated for 24 hrs with 1.0 µg/mL of CpG DNA. IL-12 secretion is not affected by either 5′ or 3′ modification. IL-6 and -10 secretion is increased when the modification is in the 5′-flanking sequence and either decreased or unaffected when the modification is in the 3′-flanking sequence. M is medium alone. Sequences of CpG DNA are: (1) 5′-d(CCTACTAGCGTTCTCATC)-3′; (2) 5′d(CCTAC*TAGCGTTCTCATC)-3′; (3) 5′-d(CCTACTAGCGTTCTC*ATC)-3′; (4) 5′-d(CTATCTGACG TTCTCTGT)-3′; (5) 5′-d(CTATC*TGACGTTCTCTGT)-3′; (6) 5′-d(CTATCTGACGTTCTC*TGT)-3′. CpG motifs are underlined. Italics indicate the 3′-deoxyribonucleosides. Asterisks indicate the position of the 2′-5′-linkages.
Chemistry of CpG DNA
nucleotide linkages in an otherwise 3′-5′-linked DNA (Fig. 4.16.3B). The presence of a 3′-deoxynucleoside either within the CpG dinucleotide or adjacent to it abrogates immunostimulatory activity (Yu et al., 2002b). However, the same modification distal to the CpG dinucleotide in the 5′-flanking sequence potentiates activity. In the 3′-flanking sequence, 3′deoxynucleosides have an insignificant effect on immunostimulation. Interestingly, 3′-deoxynucleosides in either the 5′- or the 3′-flanking sequence distal to the CpG dinucleotide result in different cytokine secretion profiles compared with unmodified CpG DNA (Fig. 4.16.4). It seems that changes in recognition
and/or interaction with the receptor that are brought about by introducing chemical changes in CpG DNA are reflected in downstream events such as cytokine secretion. However, it is not clear yet if these distinct effects are a consequence of the differences in structural and/or kinetic interactions between the modified CpG DNA and its receptor. Nonetheless, by incorporating appropriate chemical modifications into CpG DNA, it may be possible to modulate cytokine secretion in a desirable fashion for specific disease indications. Similar effects have been reported with 3′-O-methylribonucleosides (Zhao et al., 2000; Agrawal and Kandimalla, 2001).
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Significance of Deoxycytidine and Deoxyguanosine in CpG Dinucleotides: Synthetic Nucleosides Although flanking sequences strongly influence the interaction of CpG DNA and its receptors, the principal determinant in receptor recognition of a CpG DNA molecule (single- or double-stranded) is the unmethylated CpG dinucleotide itself. Any chemical modification introduced within the CpG dinucleotide that changes the DNA conformation—such as substitution of deoxyribose with ribose, 2′-O-substituted ribose, or 3′-deoxyribose, neutralization of the anionic phosphate charge between C and G, or deletion of the C or G nucleobase— completely abolishes recognition by the receptor and the subsequent immune responses. In addition, a methyl substitution at the 5 position of cytosine results in loss of activity (Zhao et al., 1996). Synthetic pyrimidines By replacing cytosine with synthetic pyrimidines, the authors have carried out an extensive study to delineate the importance for immunostimulation of each functional group on the pyrimidine ring of a CpG dinucleotide (Fig. 4.16.5). This study showed that deletion of any of the functional groups of cytosine (i.e., 2-keto, 3-imino, and 4-amino) resulted in loss of activity, suggesting that all three groups are
important for recognition and/or interaction with the receptor (Kandimalla et al., 2001). Unlike a hydrophobic methyl group, a hydroxyl substituent at the 5 position of cytosine in CpG does not suppress immunostimulatory activity (Kandimalla et al., 2001). In addition, while the 4-amino group of cytosine is absolutely required, an alkyl substitution on the amino group does not interfere with recognition and subsequent activity (Kandimalla et al., 2001). Synthetic purines Similarly, a number of synthetic purine analogs substituted for guanine in a CpG dinucleotide have been studied (Fig. 4.16.6). Deletion or modification of functional groups at the 1, 2, and 6 positions result in the loss of immunostimulatory activity (Kandimalla et al., 2001). The deletion of nitrogen at the 7 position, however, does not, suggesting that N7 is not required for receptor recognition. These studies have provided important clues regarding the functional groups of guanine in a CpG dinucleotide that are required for recognition and/or binding to the receptor (Kandimalla et al., 2001). Importantly, these studies identified new, trademarked, synthetic dinucleotides (i.e., YpG, CpR, and YpR, where Y and R are the synthetic pyrimidine and purine analogs in Figs. 4.16.5 and 4.16.6, respectively) that alter cytokine secretion profiles compared with the
NH2 H3 C NH2 4
O
O
N
NH2
N
O
N
N
1 2
N
O
5-methylcytosine
O
O O P
O
HO
N
3
5 6
N N
NH2
O H 3C
S−
O
5-methylisocytosine
NHCH2CH3
uracil
O
O
N
N
NH N
5-hydroxycytosine
N
NH O
4-N -ethylcytosine
N
O
P-base
Figure 4.16.5 Structure of 2′-deoxycytidine showing hydrogen bond acceptor (inward arrows) and donor (outward arrow) groups on cytosine. Chemical structures of some of the synthetic pyrimidine analogs studied are shown.
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O
O N
N
NH
N
N
N
hypoxanthine
NH
N N
NH2
N
7-deazaguanine
2-aminopurine
NH2
NH2 N N
O 7N 5 8
O
O
6
N 4 N 3
O
N
N
N
N
N
O
N H
N NH2
N H
1
NH purine
2
9
N
N
NH2
N
2,6-diaminopurine
isoguanine
NH2 CH3O
O
N
N N
N
NH2
N
O
N
NH
NH
O P S−
O
N H
N
O
7-deazaxanthine
K-base
H N N
N
NH2
8-bromoguanine O
O NH
O
NH
Br
N
8-oxoguanine
NH
N NH2
N
N
NH2
7-deaza-8-azaguanine
Figure 4.16.6 Structure of 2′-deoxyguanosine showing hydrogen bond acceptor (inward arrows) and donor (outward arrows) groups on guanine. Chemical structures of some of the synthetic purine analogs studied are shown.
natural CpG dinucleotide. Additionally, these studies indicate divergent synthetic nucleotide motif recognition patterns of the receptor and the possibility of modulating downstream cytokine secretion profiles using synthetic motifs placed appropriately in oligonucleotide sequences.
Role of Nucleobases
Chemistry of CpG DNA
Recently, the need for each nucleobase in a CpG DNA for immune stimulation in mice by using abasic or 1′,2′-dideoxynucleosides (Fig. 4.16.7A) has been reported. The presence of a nucleobase is absolutely required in both C and G positions of the CpG dinucleotide for activity. However, deletion of a nucleobase in the 5′-flanking region at a distance of three or more nucleosides from the CpG dinucleotide increases immunostimulatory activity (Yu et al.,
2001a). A similar deletion in the 3′-flanking sequence does not significantly affect activity, suggesting that not all nucleobases are involved in recognition. Possibly, nucleobases in certain positions cause steric hindrance when binding to the receptor. Deletion of one or two of them might relieve this strain, improving recognition and/or binding to the receptor. Moreover, nucleobase deletions in the 5′-flanking sequence increased IL-6 production compared with parent CpG DNA, while those in the 3′-flanking sequence had the opposite effect (Fig. 4.16.7B).
Requirement of Nucleosides Recently it has been shown that non-nucleoside linkers (Fig. 4.16.8) could replace certain nucleosides in the 5′- and 3′-flanking sequences (Yu et al., 2002c). An alkyl linker in the flanking sequence 5′ to the CpG dinucleotide potenti-
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A
B 3000
5′
O
B
O
IL-12 2250
O P
1500
S−
O
Cytokine (pg/mL)
O
O
O O
P
0 4000
IL-6
3000
S−
O
750
O
B
2000 1000
O3′ O
P
0
S−
M
7
1
8
CpG DNA
O
Figure 4.16.7 (A) Chemical structure of a DNA chain with a 1′,2′-dideoxyribonucleoside (boxed). B indicates base. (B) Effect of site-specific incorporation of a 1′,2′-dideoxyribonucleoside in a CpG DNA molecule (1) in either in the 5′-flanking (7) or 3′-flanking (8) sequence on induction of cytokine (IL-12 and IL-6) secretion in BALB/c mouse spleen cell cultures. Data for specific compounds are taken from Yu et al. (2001a). BALB/c mouse spleen cell cultures were incubated for 24 hr with 1.0 µg/mL of CpG DNA. Note that IL-12 secretion is not affected by either 5′ or 3′ modification. IL-6 secretion is increased when the modification is in the 5′-flanking sequence and decreased when the modification is in the 3′-flanking sequence. M is medium alone. Sequences of CpG DNAs are: (1) 5′-d(CCTACTAGCGTTCTCATC)-3′; (7) 5′-d(CCTXCTAGCGTTCTCATC)-3′; (8) 5′-d(CCTACTAGCGTTCXCATC)-3′. CpG motifs are underlined. X indicates the position of 1′,2′-dideoxyribonucleoside.
ated immunostimulatory activity. Interestingly, the same substitution in the 3′-flanking sequence did not affect immunostimulatory activity compared with parent CpG DNA. While a C3-linker optimally improved activity, longer ethyleneglycol and branched alkyl linkers (Fig. 4.16.8) were also beneficial. A linker in the 5′-flanking sequence increased IL-6 secretion several fold (Yu et al., 2002c). However, it is not clear whether the differences observed resulted from altered recognition/binding events with the receptor or initiation of different downstream signaling and transcriptional events compared with the parent CpG DNA.
Accessibility to the 5′ End of CpG DNA is Required for Immunostimulatory Activity The findings discussed above clearly suggest that the sequence 5′ to the CpG dinucleotide plays a major role in immune stimulation, while that on the 3′ side has an insignificant effect. The authors’ recent studies using 3′-3′- and 5′-5′-linked CpG DNAs suggest that an accessible 5′ end of CpG DNA is required for immunostimulatory activity (Yu et al., 2000b). Increased activity is observed when two CpG DNAs are tethered through their 3′ ends, while little or none is seen when they are tethered through their 5′ ends (Yu et al., 2000b). Synthesis of Modified Oligonucleotides and Conjugates
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O
B
O
alkyl linkers O(CH2)n O
O O
S−
P
n = 2, 3, 4, 6, or 9
O linker
ethylene glycol linkers
O O
P
S−
O
O(CH2CH2O)n CH2CH2O O
branched alkyl linkers
O O
P
n = 2 or 5
B
S−
NH2
OH
O O
O
or
O
O
Figure 4.16.8 Chemical structure of a DNA chain containing an alkyl linker. Structures of various linkers studied are shown. B indicates base.
Size of the 5′- or 3′-Attached Ligand Influences CpG DNA Activity
Chemistry of CpG DNA
Subsequent studies showed that accessibility at the 5′ end depends on the size of the ligand or moiety conjugated to this end of CpG DNA (Kandimalla et al., 2002). Conjugation of a small residue, such as a phosphorothioate group, at the 5′ end has an insignificant effect on immunostimulatory activity. However, conjugation of larger groups—including fluorescein, a mononucleotide, a tetramer, or a longer oligonucleotide (5′-5′-linked)—significantly interferes with activity. Surprisingly, conjugation of an oligonucleotide or a ligand to the 3′ end of CpG DNA (3′-3′-linked) has either an insignificant effect on activity or increases activity. Studies of cellular uptake and activation of transcription factor NF-κB in J774 cells using fluorescein-conjugated CpG DNAs suggest that both 5′- and 3′-conjugates have similar cellular uptake, but only the 3′-conjugate activates NF-κB, not the 5′-conjugate. CpG DNA has been shown to efficiently mediate antigen uptake and presentation by DCs only when antigen or allergen is conjugated to the CpG DNA (Shirota et al., 2000, 2001; Tighe et al., 2000a). CpG DNA conju-
gates are currently in clinical trials for allergies (Tighe et al., 2000a,b; Horner et al., 2002). Routinely, ligands are conjugated to oligonucleotides at the 5′ end because of convenient synthetic protocols. However, the conjugation of a macromolecule or incorporation of G-rich sequences at the 5′ end could interfere with the recognition of the CpG DNA by its receptor, thereby reducing activity. The authors’ studies suggest that the conjugation of functional ligands (e.g., antigen, antibody, allergen, another CpG DNA) at the 3′ end of CpG DNA not only contributes to increased stability against nuclease digestion but also increased immunostimulatory potency of CpG DNA in vivo.
CONCLUSIONS CpG DNA is a powerful tool that can be used to modulate the immune system for treatment of a wide variety of disease indications. Several first-generation CpG DNA molecules are in clinical trials for a number of diseases either as monotherapies, in combination with antigens, vaccines, or monoclonal antibodies, or as conjugates with antigens (Gurunathan et al., 2000; Agrawal and Kandimalla, 2002; Kandimalla and Agrawal, 2002; Krieg, 2002). Extensive
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safety data is available from clinical studies of antisense oligonucleotides. Antisense oligonucleotides, which are used at several-fold higher concentrations than CpG DNA, have been administered to humans without serious adverse safety concerns. To date, several hundred people have been treated for up to two years without any evidence of anti-DNA antibody formation. Many of these molecules that are in clinical trials contain CpG dinucleotides. The presence of CpG dinucleotides in antisense oligonucleotides induces immune responses (Agrawal and Kandimalla, 2000; Lewis et al., 2000; Agrawal and Kandimalla, 2001; Jahrsdorfer et al., 2002), sometimes resulting in uncontrolled cytokine secretion, causing toxicity concerns (Agrawal and Zhao, 1998a,b; Agrawal, 1999a,b). A number of second-generation chemical modifications of CpG motifs in antisense oligonucleotides have been reported to suppress immune-related side effects (Agrawal and Zhao, 1998a,b; Agrawal, 1999a,b; Agrawal and Kandimalla, 2000). The study of CpG DNA chemistry is a step closer towards understanding the biological effects and development of CpG DNA for human therapies. The chemical studies discussed in this review suggest that, in addition to the CpG dinucleotide, a number of other factors in CpG DNA molecules influence recognition by receptors, secretion of cytokines, and the resulting immunological effects. The ability to regulate cytokine induction through the use of second-generation CpG DNA modifications discussed in this review is an important factor in advancing the use of CpG DNA for specific disease indications with reduced toxicity concerns. Further understanding of the biological effects of second-generation CpG DNAs will lead to the design of more potent CpG DNA pharmacological agents.
LITERATURE CITED Agrawal, S. 1999a. Factors affecting the specificity and mechanism of action of antisense oligonucleotides. Antisense Nucleic Drug Dev. 9:371375. Agrawal, S. 1999b. Importance of nucleotide sequence and chemical modifications of antisense oligonucleotides. Biochim. Biophys. Acta 1489:53-68.
Agrawal, S. and Kandimalla, E.R. 2002. Medicinal chemistry and therapeutic potential of CpG DNA. Trends Mol. Med. 8:114-121. Agrawal, S. and Zhao, Q. 1998a. Antisense therapeutics. Curr. Opin. Chem. Biol. 2:519-528. Agrawal, S. and Zhao, Q. 1998b. Mixed backbone oligonucleotides: Improvement oligonucleotide-induced toxicity in vivo. Antisense Nucleic Acid Drug Dev. 8:135-139. Agrawal, S., Iadarola, P.L., Temsamani, J., Zhao, Q., and Shaw, D. 1996. Effect of G-rich sequences on the synthesis, purification, binding, cell uptake, and hemolytic activity of oligonucleotides. Bioorg. Med. Chem. Let. 6:2219-2224. Ahmad-Nejad, P., Hacker, H., Rutz, M., Bauer, S., Vabulas, R.M., and Wagner, H. 2002. Bacterial CpG-DNA and lipopolysaccharides activate Toll-like receptors at distinct cellular compartments. Eur. J. Immunol. 32:1958-1968. Alexopoulou, L., Holt, A.C., Medzhitov, R., and Flavell, R.A. 2001. Recognition of doublestranded RNA and activation of NF-κB by Tolllike receptor 3. Nature 413:732-738. Ballas, Z.K., Krieg, A.M., Warren, T., Rasmussen, W., Davis, H.L., Waldschmidt, M., and Weiner, G.J. 2001. Divergent therapeutic and immunologic effects of oligodeoxynucleotides with distinct CpG motifs. J. Immunol. 167:48784886. Bauer, S., Kirschning, C.J., Hacker, H., Redecke, V., Hausmann, S., Akira, S., Wagner, H., and Lipford, G.B. 2001a. Human TLR9 confers responsiveness to bacterial DNA via species-specific CpG motif recognition. Proc. Natl. Acad. Sci. U.S.A. 98:9237-9242. Bauer, M., Redecke, V., Ellwart, J.W., Scherer, B., Kremer, J.P., Wagner, H., and Lipford, G.B. 2001b. Bacterial CpG-DNA triggers activation and maturation of human CD11c-, CD123+ dendritic cells. J. Immunol. 166:5000-5007. Bird, A.P. 1986. CpG-rich islands and the function of DNA methylation. Nature 321:209-213. Branda, R.F., Moore, A.L., Mathews, L., McCormack, J.J., and Zon, G. 1993. Immune stimulation by an antisense oligomer complementary to the rev gene of HIV-1. Biochem. Pharmacol. 45:2037-2043. Dalpke, A.H., Zimmermann, S., Albrecht, I., and Heeg, K. 2002. Phosphodiester CpG oligonucleotides as adjuvants: Polyguanosine runs enhance cellular uptake and improve immunostimulative activity of phosphodiester CpG oligonucleotides in vitro and in vivo. Immunology 106:102-112.
Agrawal, S. and Kandimalla, E.R. 2000. Antisense therapeutics: Is it as simple as complementary base recognition? Mol. Med. Today 6:72-81.
Gursel, M., Verthelyi, D., Gursel, I., Ishii, K.J., and Klinman, D.M. 2002. Differential and competitive activation of human immune cells by distinct classes of CpG oligodeoxynucleotides. J. Leukoc. Biol. 71:813-820.
Agrawal, S. and Kandimalla, E.R. 2001. Antisense and/or immunostimulatory oligonucleotide therapeutics. Current Cancer Drug Targets 1:197-209.
Gurunathan, S., Klinman, D.M., and Seder, R.A. 2000. DNA vaccines: Immunology, application, and optimization. Annu. Rev. Immunol. 18:927974.
Synthesis of Modified Oligonucleotides and Conjugates
4.16.11 Current Protocols in Nucleic Acid Chemistry
Supplement 12
Hartmann, G., Weeratna, R.D., Ballas, Z.K., Payette, P., Blackwell, S., Suparto, I., Rasmussen, W.L., Waldschmidt, M., Sajuthi, D., Purcell, R.H., Davis, H.L., and Krieg, A.M. 2000. Delineation of a CpG phosphorothioate oligodeoxynucleotide for activating primate immune responses in vitro and in vivo. J. Immunol. 164:1617-1624. Hemmi, H., Takeuchi, O., Kawai, T., Kaisho, T., Sato, S., Sanjo, H., Matsumoto, M., Hoshino, K., Wagner, H., Takeda, K., and Akira, S. 2000. A Toll-like receptor recognizes bacterial DNA. Nature 408:740-745. Horner, A.A., Takabaysahi, K., Zubeldia, J.M., and Raz, E. 2002. Immunostimulatory DNA-based therapeutics for experimental and clinical allergy. Allergy 57:24-29. Iho, S., Yamamoto, T., Takahashi, T., and Yamamoto, S. 1999. Oligodeoxynucleotides containing palindrome sequences with internal 5′-CpG-3′ act directly on human NK and activated T cells to induce IFN-γ production in vitro. J. Immunol. 163:3642-3652. Jahrsdorfer, B., Jox, R., Muhlenhoff, L., Tschoep, K., Krug, A., Rothenfusser, S., Meinhardt, G., Emmerich, B., Endres, S., and Hartmann, G. 2002. Modulation of malignant B cell activation and apoptosis by bcl-2 antisense ODN and immunostimulatory CpG ODN. J. Leukoc. Biol. 72:83-92. Kadowaki, N., Antonenko, S., and Liu, Y.J. 2001. Distinct CpG DNA and polyinosinic-polycytidylic acid double-stranded RNA, respectively, stimulate CD11c- type 2 dendritic cell precursors and CD11c+ dendritic cells to produce type I IFN. J. Immunol. 166:2291-2295. Kandimalla, E.R. and Agrawal, S. 2002. Towards optimal design of second-generation immunomodulatory oligonucleotides. Curr. Op. Mol. Ther. 4:122-129. Kandimalla, E.R., Yu, D., Zhao, Q., and Agrawal, S. 2001. Effect of chemical modifications of cytosine and guanine in a CpG-motif of oligonucleotides: Structure-immunostimulatory activity relationships. Bioorg. Med. Chem. 9:807-813. Kandimalla, E.R., Bhagat, L., Yu, D., Cong, Y., Tang, J., and Agrawal, S. 2002. Conjugation of ligands at the 5′-end of CpG DNA affects immunostimulatory activity. Bioconj. Chem. 13:966-974. Kimura, Y., Sonehara, K., Kuramoto, E., Makino, T., Yamamoto, S., Yamamoto, T., Kataoka, T., and Tokunaga, T. 1994. Binding of oligoguanylate to scavenger receptors is required for oligonucleotides to augment NK cell activity and induce IFN. J. Biochem. 116:991-994. Klinman, D.M., Yi, A.K., Beaucage, S.L., Conover, J., and Krieg, A.M. 1996. CpG motifs present in bacterial DNA rapidly induce lymphocytes to secrete interleukin 6, interleukin 12, and interferon γ. Proc. Natl. Acad. Sci. U.S.A. 93:28792883. Chemistry of CpG DNA
Krieg, A.M., Yi, A.K., Matson, S., Waldschmidt, T.J., Bishop, G.A., Teasdale, R., Koretzky, G.A., and Klinman, D.M. 1995. CpG motifs in bacterial DNA trigger direct B-cell activation. Nature 374:546-549. Krug, A., Rothenfusser, S., Hornung, V., Jahrsdorfer, B., Blackwell, S., Ballas, Z.K., Endres, S., Krieg, A.M., and Hartmann, G. 2001a. Identification of CpG oligonucleotide sequences with high induction of IFN-α/β in plasmacytoid dendritic cells. Eur. J. Immunol. 31:2154-2163. Krug, A., Towarowski, A., Britsch, S., Rothenfusser, S., Hornung, V., Bals, R., Giese, T., Engelmann, H., Endres, S., Krieg, A.M., and Hartmann, G. 2001b. Toll-like receptor expression reveals CpG DNA as a unique microbial stimulus for plasmacytoid dendritic cells which synergizes with CD40 ligand to induce high amounts of IL-12. Eur. J. Immunol. 31:3026-3037. Lewis, E.J., Agrawal, S., Bishop, J., Chadwick, J., Cristensen, N.D., Cuthill, S., Dunford, P., Field, A.K., Francis, J., Gibson, V., Greenham, A.K., Kelly, F., Kilkuskie, R., Kreider, J.W., Mills, J.S., Mulqueen, M., Roberts, N.A., Roberts, P., and Szymkowski, D.E. 2000. Non-specific antiviral activity of antisense molecules targeted to the E1 region of human papillomavirus. Antiviral Res. 48:187-196. Lien, E., and Ingalls, R.R. 2002. Toll-like receptors. Crit. Care Med. 30:S1-S11. Messina, J.P., Gilkeson, G.S., and Pisetsky, D.S. 1991. Stimulation of in vitro murine lymphocyte proliferation by bacterial DNA. J. Immunol. 147:1759-1764. Pearson, A.M., Rich, A., and Krieger, M. 1993. Polynucleotide binding to macrophage scavenger receptors depends on the formation of basequartet-stabilized four-stranded helices. J. Biol. Chem. 268:3546-3554. Pisetsky, D.S. 1999. The influence of base sequence on the immunostimulatory properties of DNA. Immunol. Res. 19:35-46. Rankin, R., Pontarollo, R., Ioannou, X., Krieg, A.M., Hecker, R., Babiuk, L.A., and van den Hurk, S.v.d.L. 2001. CpG motif identification for veterinary and laboratory species demonstrates that sequence recognition is highly conserved. Antisense Nucleic Acid Drug Dev. 11:333-340. Ro thenfusser, S., Hornung, V., Krug, A., Towarowski, A., Krieg, A.M., Endres, S., and Hartmann, G. 2001. Distinct CpG oligonucleotide sequences activate human gamma delta T cells via interferon-α/-β. Eur. J. Immunol. 31:3525-3534. Sato, Y., Roman, M., Tighe, H., Lee, D., Corr, M., Nguyen, M.D., Silverman, G.J., Lotz, M., Carson, D.A., and Raz, E. 1996. Immunostimulatory DNA sequences necessary for effective intradermal gene immunization. Science 273:352-354.
Krieg, A.M. 2002. CpG motifs in bacterial DNA and their immune effects. Annu. Rev. Immunol. 20:709-760.
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Shirota, H., Sano, K., Kikuchi, T., Tamura, G., and Shirato, K. 2000. Regulation of murine airway eosinophilia and Th2 cells by antigen-conjugated CpG oligodeoxynucleotides as a novel antigen-specific immunomodulator. J. Immunol. 164:5575-5582. Shirota, H., Sano, K., Hirasawa, N., Terui, T., Ohuchi, K., Hattori, T., Shirato, K., and Tamura, G. 2001. Novel roles of CpG oligodeoxynucleotides as a leader for the sampling and presentation of CpG-tagged antigen by dendritic cells. J. Immunol. 167:66-74. Sonehara, K., Saito, H., Kuramoto, E., Yamamoto, S., Yamamoto, T., and Tokunaga, T. 1996. Hexamer palindromic oligonucleotides with 5′-CG3′ motif(s) induce production of interferon. J. Interferon Cytokine Res. 16:799-803. Stacey, K.J., Sester, D.P., Sweet, M.J., and Hume, D.A. 2000. Macrophage activation by immunostimulatory DNA. Curr. Top. Microbiol. Immunol. 247:41-58. Takeshita, F., Leifer, C.A., Gursel, I., Ishii, K.J., Takeshita, S., Gursel, M., and Klinman, D.M. 2001. Role of Toll-like receptor 9 in CpG DNAinduced activation of human cells. J. Immunol. 167:3555-3558. Tang, J.Y., Roskey, A.R., Li, Y., and Agrawal, S. 1995. Enzymatic synthesis of stereoregular (all Rp) oligonucleotide phosphorothioate and its properties. Nucleosides Nucleotides 14:985989. Tighe, H., Takabayashi, K., Schwartz, D., Van Nest, G., Tuck, S., Eiden, J.J., Kagey-Sobotka, A., Creticos, P.S., Lichtenstein, L.M., Spiegelberg, H.L., and Raz, E. 2000a. Conjugation of immunostimulatory DNA to the short ragweed allergen amb a 1 enhances its immunogenicity and reduces its allergenicity. J. Allergy Clin. Immunol. 106:124-134. Tighe, H., Takabayashi, K., Schwartz, D., Marsden, R., Beck, L., Corbeil, J., Richman, D.D., Eiden, J.J. Jr., Spiegelberg, H.L., and Raz, E. 2000b. Conjugation of protein to immunostimulatory DNA results in a rapid, long-lasting and potent induction of cell-mediated and humoral immunity. Eur. J. Immunol. 30:1939-1947. Tokunaga, T., Yamamoto, H., Shimada, S., Abe, H., Fukuda, T., Fujisawa, Y., Furutani, Y., Yano, O., Kataoka, T., Sudo, T., Makiguchi, N., and Suganuma, T. 1984. Antitumor activity of deoxyribonucleic acid fraction from Mycobacterium bovis BCG. I. Isolation, physicochemical characterization, and antitumor activity. J. Natl. Cancer Inst. 72:955-962. Van Uden, J., and Raz, E. 2000. Introduction to immunostimulatory DNA sequences. Springer Semin. Immunopathol. 22:1-9. Verthelyi, D., Ishii, K., Gursel, M., Takeshita, F., and Klinman, D. 2001. Human peripheral blood cells differentially recognize and respond to two distinct CpG motifs. J. Immunol. 166:2372-2377.
Verthelyi, D., Kenney, R.T., Seder, R.A., Gam, A.A., Friedag, B., and Klinman, D.M. 2002. CpG oligodeoxynucleotides as vaccine adjuvants in primates. J. Immunol. 168:1659-1663. Wernette, C.M., Smith, B.F., Barksdale, Z.L., Hecker, R., and Baker, H.J. 2002. CpG oligodeoxynucleotides stimulate canine and feline immune cell proliferation. Vet. Immunol. Immunopathol. 84:223-236. Yamamoto, S., Yamamoto, T., Kataoka, T., Kuramoto, E., Yano, O., and Tokunaga, T. 1992a. Unique palindromic sequences in synthetic oligonucleotides are required to induce INF and augment INF-mediated natural killer activity. J. Immunol. 148:4072-4076. Yamamoto, S., Yamamoto, T., Shimada, S., Kuramoto, E., Yano, O., Kataoka, T., and Tokunaga, T. 1992b. DNA from bacteria, but not from vertebrates, induces interferons, activates natural killer cells and inhibits tumor growth. Microbiol. Immunol. 36:983-997. Yamamoto, S., Yamamoto, T., Iho, S., and Tokunaga, T. 2000. Activation of NK cell (human and mouse) by immunostimulatory DNA sequence. Springer Semin. Immunopathol. 22:35-43. Yu, D., Kandimalla, E.R., Roskey, A., Zhao, Q., Chen, L., Chen, J., and Agrawal, S. 2000a. Stereo-enriched phosphorothioate oligodeoxynucleotides: Synthesis, biophysical and biological properties. Bioorg. Med. Chem. 8:275-284. Yu, D., Zhao, Q., Kandimalla, E.R., and Agrawal, S. 2000b. Accessible 5′-end of CpG-containing phosphorothioate oligodeoxynucleotides is essential for immunostimulatory activity. Bioorg. Med. Chem. Lett. 10:2585-2588. Yu, D., Kandimalla, E.R., Zhao, Q., Cong, Y., and Agrawal, S. 2001a. Modulation of immunostimulatory activity of CpG oligonucleotides by site-specific deletion of nucleobases. Bioorg. Med. Chem. Lett. 11:2263-2267. Yu, D., Kandimalla, E.R., Zhao, Q., Cong, Y., and Agrawal, S. 2001b. Immunostimulatory activity of CpG oligonucleotides containing non-ionic methylphosphonate linkages. Bioorg. Med. Chem. 9:2803-2808. Yu, D., Zhu, F.G., Bhagat, L., Wang, H., Kandimalla, E.R., Zhang, R., and Agrawal, S. 2002a. Potent CpG oligonucleotides containing phosphodiester linkages: In vitro and in vivo immunostimulatory properties. Biochem. Biophys. Res. Commun. 297:83-90. Yu, D., Kandimalla, E.R., Zhao, Q., Cong, Y., and Agrawal, S. 2002b. Immunostimulatory properties of phosphorothioate CpG DNA containing both 3′-5′- and 2′-5′-internucleotide linkages. Nucl. Acids Res. 30:1613-1619. Yu, D., Kandimalla, E.R., Cong, Y., Tang, J., Tang, J.Y., Zhao, Q., and Agrawal, S. 2002c. Design, synthesis, and immunostimulatory properties of CpG DNAs containing alkyl-linker substitutions: Role of nucleosides in the flanking sequences. J. Med. Chem. 45:4540-4548.
Synthesis of Modified Oligonucleotides and Conjugates
4.16.13 Current Protocols in Nucleic Acid Chemistry
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Zhang, Y., Shoda, L.K., Brayton, K.A., Estes, D.M., Palmer, G.H., and Brown, W.C. 2001. Induction of interleukin-6 and interleukin-12 in bovine B lymphocytes, monocytes, and macrophages by a CpG oligodeoxynucleotide (ODN 2059) containing the GTCGTT motif. J. Interferon Cytokine Res. 21:871-881. Zhao, Q., Temsamani, J., Iadarola, P.L., Jiang, Z., and Agrawal, S. 1996. Effect of different chemically modified oligodeoxynucleotides on immune stimulation. Biochem. Pharmacol. 51:173-182. Zhao, Q., Temsamani, J., Zhou, R.Z., and Agrawal, S. 1997. Pattern and kinetics of cytokine productio n fo llowing administration of phosphorothioate oligonucleotides in mice. Antisense Nucleic Acid Drug. Dev. 7:495-502.
Zhao, Q., Yu, D., and Agrawal, S. 1999. Site of chemical modifications in CpG containing phosphorothioate oligodeoxynucleotide modulates its immunostimulatory activity. Bioorg. Med. Chem. Lett. 9:3453-3458. Zhao, Q., Yu, D., and Agrawal, S. 2000. Immunostimulatory activity of CpG containing phosphorothioate oligodeoxynucleotide is modulated by modification of a single deoxynucleoside. Bioorg. Med. Chem. Lett. 10:1051-1054.
Contributed by Ekambar R. Kandimalla and Sudhir Agrawal Hybridon, Inc. Cambridge, Massachusetts
Chemistry of CpG DNA
4.16.14 Supplement 12
Current Protocols in Nucleic Acid Chemistry
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Phosphorothioate Linkages
UNIT 4.17
Phosphorothioate analogs of oligonucleotides (PS-oligos) constitute an important tool for studying the metabolism of nucleic acids (Eckstein, 2000, and references therein) and have been evaluated as potential therapeutics in the so-called “antisense” (Stein and Krieg, 1998) and “antigene” strategies (Thuong and Helene, 1993). In 1998, the U.S. Food and Drug Administration (FDA) approved the first PS-oligo, Fomirvirsen (trade name, Vitravene), for therapeutic application against cytomegalovirus (CMV) retinitis (Manoharan, 1999). Most of the second-generation antisense compounds that are currently undergoing clinical trials are PS-oligos (e.g., Isis Pharmaceuticals, Hybridon; Maier et al., 2000). PS-oligos are isoelectronic with natural oligonucleotides and, importantly, they are much more resistant towards intra- and extracellular nucleases (Wickstrom, 1986). These features are important with respect to their therapeutic applications. However, substitution of sulfur for one nonbridging oxygen in the internucleotide phosphate group induces asymmetry at the phosphorus atom, and standard chemical methods for the synthesis of oligo(deoxyribonucleoside phosphorothioate)s provide a mixture of 2n diastereomers, where n is the number of phosphorothioate linkages (Wilk and Stec, 1995). Therefore, even for relatively short PS-oligos (10- to 15-mers), thousands of diastereomers would be involved in interactions with other chiral biomolecules (e.g., DNA, RNA, or proteins) and, in principle, each diastereomer might interact in a slightly different manner. The enzymatic synthesis of PS-oligos allows for the preparation of PS-oligonucleotides of RP-configuration at each phosphorothioate linkage (all-RP-PS-oligos) due to the stereoselectivity of all DNA and RNA polymerases identified to date (Hacia et al., 1994; Lackey and Patel, 1997; Tang et al., 1995). The first method for stereocontrolled chemical synthesis of PS-oligos, which was elaborated in the authors’ laboratory (Stec et al., 1991), is based on a new chemistry employing P-diastereomerically pure nucleoside monomers possessing the 2-thio-1,3,2-oxathiaphospholane moiety attached to appropriately protected nucleosides at the 3′-O position (S.1; Fig. 4.17.1). Further studies resulted in the synthesis of monomers with the oxathiaphospholane ring substituted at position 4 with either two methyl groups (S.2; Stec et al., 1995) or a spiro pentamethylene ring (S.3; Stec et al., 1998). These substituents enhance a differentiation in chromatographic mobility of diastereomers, rendering their separation less laborious. The oxathiaphospholane monomers react in the presence of 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) with the 5′-OH group of a nucleoside (or growing oligonucleotide attached at the 3′ end to a DBU-resistant solid support) to yield a dinucleotide (or an elongated oligomer) with an internucleotide phosphorothioate diester bond, as depicted in Figure 4.17.2. The process is fully stereospecific and occurs with retention of configuration at the phosphorus atom. The chemical yield of the condensation process is not as efficient as that of the phosphoramidite or H-phosphonate methods (UNITS 3.3 & 3.4), but repetitive yields of 92% to 94% allow syntheses of medium-sized oligomers (up to 15-mers). Longer oligonucleotides were obtained in poor yields and the syntheses were not reproducible. Oxathiaphospholanes that are 18O-labeled at the endocyclic position allowed for the synthesis of PS-oligos with internucleotide PS[18O]-phosphorothioate moieties of predetermined chirality (Guga et al., 2001). Synthesis of Modified Oligonucleotides and Conjugates Contributed by Piotr Guga and Wojciech J. Stec Current Protocols in Nucleic Acid Chemistry (2003) 4.17.1-4.17.28 Copyright © 2003 by John Wiley & Sons, Inc.
4.17.1 Supplement 14
For the synthesis of stereodefined PS-oligos via the oxathiaphospholane methodology presented in this unit, pure P-diastereomers of nucleoside oxathiaphospholane monomers are required. They are not commercially available, but can be efficiently obtained by phosphitylation of widely available 5′-O-DMTr-N-protected deoxyribonucleosides with oxathiaphospholane phosphitylating reagent followed by sulfurization. The methodology of their synthesis and use in solid-phase synthesis of PS-oligos is presented in consecutive protocols. Basic Protocol 1 describes a detailed procedure for the synthesis of the phosphitylating reagent 2-chloro-spiro-4,4-pentamethylene-1,3,2-oxathiaphospholane. The procedure is general and may be applied to other analogs, depending on the aldehyde (or mercaptoalcohol) used. Alternate Protocol 1 describes a procedure for synthesis of 18O-labeled mercaptoalcohol, which is used to synthesize labeled phosphitylating reagent and, subsequently, 18O-labeled nucleoside monomers. These can be used for synthesis of stereodefined PS[18O]oligos, which are useful compounds in studying the mechanism(s) of enzymatic reactions. Support Protocol 1 describes a method for transfer of dry solvents, required for this procedure. Basic Protocol 2 outlines the synthesis of 5′-O-DMTr-N-protected-deoxyribonucleoside3′-O-(2-thio-spiro-4,4-pentamethylene-1,3,2-oxathiaphospholane)s (S.3) and their chromatographic separation into P-diastereomers. This method, although described for dA, dC, dG, and T derivatives, can be also used for derivatizing other appropriately protected nucleosides. For example, in the authors’ laboratory, N6-benzoyl-7-deaza-5′-O(4,4′-dimethoxytrityl)-3′-O-(2-thio-spiro-4,4-pentamethylene-1,3,2-oxathiaphospholane)2′-deoxyadenosine was obtained and separated into diastereomers (unpub.). Similarly, this method is suitable for phosphitylation of protected nucleosides with other oxathiaphospholane reagents containing different substituents, although the separation of diastereo-mers may be very difficult. Alternate Protocol 2 describes the conversion of 5′-O-DMTr-N-protected-deoxyribonucleoside-3′-O-(2-thio-spiro-4,4-pentamethylene-1,3,2-oxathiaphospholane)s to their 2oxo-analogs with selenium dioxide. These monomers can be used to elongate stereodefined PS-oligos and generate segments of unmodified nucleotide units possessing phosphate internucleotide linkages. This goal cannot be achieved with the phosphoramidite or H-phosphonate methods, because the phosphorothioate linkages already
DMTrO
O
B
O X P 1
O 1 2 3 4 Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
X = S, X = S, X = S, X = O,
S
3
R
5
R
R=H R = CH 3 R,R = −(CH 2)5− R,R = −(CH 2)5−
Figure 4.17.1 Structural features of deoxyribonucleoside oxathiaphospholane derivatives. Abbreviations: B, thymin-1-yl or N-protected nucleobase; DMTr, 4,4′-dimethoxytrityl. Adapted from Stec et al. (1998) with permission from the American Chemical Society.
4.17.2 Supplement 14
Current Protocols in Nucleic Acid Chemistry
−
S O
OR′
P OR S DBU
O S
OR
P
S
ψ O
−
S
OR′
P
S
−
S
R′O RO
OR′
OR
DMTrO R=
O
P
−
S
O
B
O
−
O
S R′O RO
P
−
S
B
R′ = O Sar
Figure 4.17.2 Mechanism of base-promoted oxathiaphospholane ring-opening condensation. Abbreviations: B, thymin-1yl or N-protected nucleobase; DBU, 1,8-diazabicyclo[5.4.0]undec-7-ene; DMTr, 4,4′-dimethoxytrityl; ψ, pseudorotation; Sar, sarcosinylated or DBU-resistant solid support.
generated by the oxathiaphospholane method are diesters and would be oxidized in the I2/water/pyridine routinely used for conversion of phosphites to phosphates. Basic Protocol 3 outlines details of manual solid-phase synthesis of PS-oligos using oxathiaphospholane monomers. In principle, this synthesis can be performed on an automatic synthesizer, but the necessary modification of the manufacturer’s protocols is impossible for the majority of synthesizers. The protocol for 1-µmol-scale automated solid-phase synthesis using an ABI 391 synthesizer (Applied Biosystems) has been published (Stec et al., 1998), but in many instances the software does not allow for any changes in the protocol. Also, the instrument should be able to deliver an additional solvent (methylene chloride) to the column in order to wash delivery lines after coupling. This is necessary to avoid formation of deposits inside the tubing and valves, which may lead to major failure of the instrument and expensive replacement of the clogged valve blocks. One also has to consider that, in using an automated synthesizer, significant amounts of monomer solutions are wasted during optimization of the protocol, and due to the dead volumes of the system. Therefore, manual synthesis of a limited number of oligomers may be economically more justified. Support Protocol 2 describes preparation of solid supports for the synthesis of PS-oligos, which must be DBU-resistant because this strong base is necessary for the coupling step. This requirement is fulfilled by the use of Brown’s sarcosinyl-succinoyl linker (Brown et al., 1989).
Synthesis of Modified Oligonucleotides and Conjugates
4.17.3 Current Protocols in Nucleic Acid Chemistry
Supplement 14
H
O
O S2Cl2
H
H
S
S
O
−HCl
HO NaBH4
S
propan-2-ol 6 (70%)
5
OH S
OH SH
7 (85%)
O
LiAlH4
PCl3
diethyl ether
pyridine
P Cl S
8 (70%)
9 (70%)
Figure 4.17.3 Synthesis of 2-chloro-spiro-4,4-pentamethylene-1,3,2-oxathiaphospholane (S.9) starting from cyclohexanecarboxaldehyde (S.5). Adapted from Stec et al. (1998) with permission from the American Chemical Society.
CAUTION: It is imperative that all reactions be run in a suitable fume hood with efficient ventilation. Many of the reactions in this unit are highly exothermic; safety glasses and reagent-impermeable protective gloves should be worn. BASIC PROTOCOL 1
SYNTHESIS OF PHOSPHITYLATING REAGENT: 2-CHLORO-spiro-4,4-PENTAMETHYLENE-1,3,2-OXATHIAPHOSPHOLANE The most simple oxathiaphosphitylating reagent, 2-chloro-1,3,2-oxathiaphospholane, can be obtained from the reaction of 2-mercaptoethanol with phosphorus trichloride in the presence of two molar equivalents of triethylamine (Martynov et al., 1969; Willson et al., 1975; Stec et al., 1991). Condensation of an appropriately protected 3′-OH-nucleoside with 2-chloro-1,3,2-oxathiaphospholane in pyridine solution, performed in the presence of dry elemental sulfur, provides nucleoside 3′-O-(2-thiono-1,3,2-oxathiaphospholane)s (S.1; Fig. 4.17.1). However, their separation as P-diastereomers is very laborious and requires several consecutive silica gel chromatographic runs of partially enriched fractions. Therefore, it is recommended to synthesize 2-chloro-spiro-4,4-pentamethylene1,3,2-oxathiaphospholane (S.9) starting from cyclohexanecarboxaldehyde (S.5), as depicted in Figure 4.17.3. Using isobutyraldehyde in the same sequence of reactions, 2-chloro-4,4-dimethyl-1,3,2-oxathiaphospholane can be obtained. However, the phosphitylating reagent S.9, when used for synthesis of nucleotide monomers, provides much more useful compounds in terms of chromatographic separability as pure P-diastereomers. It is important to note that the fast-eluting isomers of 4,4-dimethyl- and 4,4-pentamethylene-oxathiaphospholane monomers are precursors of RP internucleotidic phosphorothioate bonds. Conversely, analog internucleotide phosphorothioate bonds of RP configuration are formed from the slow-eluting isomer of S.1.
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
Using this methodology, different analogs can be synthesized. However, it is important to obtain a phosphitylating reagent without additional centers of asymmetry, as the number of diastereomers will double with each new center, rendering separation of P-diastereomers very difficult or impossible. The relationship between chromatographic mobility and absolute configuration of the monomers must be checked for each new analog.
4.17.4 Supplement 14
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Materials 4 to 5 M and 1.5 M sodium hydroxide (NaOH) Sulfur monochloride (S2Cl2), freshly distilled over 2 g elemental sulfur (S8; dried ≥12 hr under vacuum) per 50 mL S2Cl2 Argon (or, optionally, nitrogen), dry Cyclohexanecarboxaldehyde (Fluka) Methylene chloride Diethyl ether, anhydrous Sodium borohydride (NaBH4) Isopropyl alcohol Anti-bumping granules 20% (w/v) hydrochloric acid Chloroform Magnesium sulfate, anhydrous Hexane Lithium aluminum hydride Ethyl acetate, dry Tetrahydrofuran (THF), with traces of added moisture 10% (v/v) H2O/THF Phosphorus trichloride (PCl3) Benzene, anhydrous Pyridine Dry molecular sieves (4A, 4- to 6-µm-o.d. beads, Aldrich) 250-mL absorber with safety flask (see Fig. 4.17.4) 250-mL four-neck round-bottom flask Heated oil bath capable of magnetic stirring Thermometer (capable of reading 150°C) 100-mL dropping funnel Reflux condensers Glass gas inlet adapter (preferred) or syringe needle and rubber septum Rotary evaporator with a water aspirator and a diaphragm vacuum pump (10 to 15 mmHg; optional) 500-mL Erlenmeyer flask (29/42 joint) 1-L two-neck round-bottom flask (two 29/42 joints) Stopcock, 29/42 Flexible adapter (glass M/F joints, 29/42, on corrugated Teflon tubing; optional) 500-mL separatory funnel Filter funnel and Whatman no.1 filter paper (or equivalent) High-vacuum fractional distillation apparatus High-vacuum oil pump (0.01 mmHg) NOTE: Upon storage, cyclohexanecarboxaldehyde undergoes polymerization. Order only the amount required for use within 2 to 3 weeks. NOTE: Within this unit, evaporation of solvents is performed using a rotary evaporator connected to a water aspirator, unless otherwise specified.
Synthesis of Modified Oligonucleotides and Conjugates
4.17.5 Current Protocols in Nucleic Acid Chemistry
Supplement 14
to the hood
4 M NaOH safety flask
absorber
thermometer NOTE: glass gas adapter not shown
aldehyde sulfur monochloride
magnetic stir bar
oil bath 567 4 8 3 2 1 119
4 567 3 8 2 9 1 1
magnetic stirrer
Figure 4.17.4 System assembly for synthesis of S.6. The glass gas adapter (or septum and needle) for delivery of dry argon should be mounted in the fourth neck of the flask (not shown).
Synthesize 2,2′-dithiobis(cyclohexanecarbaldehyde) (S.6) 1. Prepare a 250-mL absorber containing ∼150 mL of 4 to 5 M NaOH (see Fig. 4.17.4). 2. Assemble a reactor consisting of a 250-mL four-neck, round-bottom flask (to be heated in an oil bath with magnetic stirring) equipped with a thermometer, a 100-mL dropping funnel, a condenser, and a magnetic stir bar. Connect the outlet of the condenser via a safety flask to the absorber and be sure that the absorber vents into the hood. The capacity of the safety flask must be sufficient to accommodate the solution of sodium hydroxide from the absorber.
3. Add 30 g (17.8 mL, 0.22 mol) of freshly distilled sulfur monochloride to the flask. Make sure that the thermometer is in contact with the liquid. Deliver dry argon close to the bottom of the flask through the fourth joint either with a glass gas inlet adapter or with a syringe needle inserted through a rubber septum. Since gaseous hydrogen chloride is liberated from the reaction, the use of a glass inlet adapter, rather than a syringe needle, is recommended.
4. Apply heating until sulfur monochloride reaches 60°C. Stabilize the temperature. Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
5. Add, dropwise through the dropping funnel, 50 g (56 mL, 0.44 mol) of cyclohexanecarboxaldehyde over a 60-min period with stirring, keeping the temperature at 60°C. Remove the oil bath and allow the mixture to continue stirring another 10 min.
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6. Stop stirring and remove the stir bar. Leave the reaction mixture until it reaches room temperature and solidifies (∼1 hr). 7. Dissolve the solid residue in 150 mL of methylene chloride, then evaporate to dryness using a rotary evaporator connected to a water aspirator. CAUTION: During evaporation gaseous hydrogen chloride is liberated.
8. Add 150 mL of diethyl ether and gently reflux until the solid residue is dissolved. Transfer the solution into a 500-mL Erlenmeyer flask and close the stopcock. Cool down the mixture (containing the synthesized S.6) and keep overnight in a refrigerator (4°C) to allow crystallization. Collect crystalline S.6 by filtration. Approximately 35 g of 2,2′-dithiobis(cyclohexanecarbaldehyde) (S.6) should be collected as a white solid (∼70% yield, m.p. 88° to 89°C). 1H NMR (CDCl3, δ): 8.98 ppm (s, 1H, CHO), 1.2-2.1 ppm (m, 10H). FAB MS (positive mode, Cs+, 13 keV, matrix NBA) m/z 286, [M]+, 25%; m/z 111, [C6H10CHO]+, 100%.
Synthesize 2,2′-dithiobis(cyclohexanemethanol) (S.7) 9. In a 1-L two-neck flask (two 29/42 joints), equipped with a reflux condenser and a stopcock, suspend 5.67 g (0.15 mol) of NaBH4 in 500 mL isopropyl alcohol. Add anti-bumping granules and heat to boiling. 10. While gently refluxing, remove the stopcock momentarily and add, with a chemical spoon, ∼2 g of S.6 every 3 to 5 min in 10 to 12 portions for a total of 21.5 g (0.075 mol). CAUTION: The addition of each portion of 2,2′-dithiobis(cyclohexanecarboxaldehyde) results in enhanced boiling and emission of vapors of isopropyl alcohol through the open neck. The stopcock should thus be closed as soon and possible. Alternatively, one can use a flexible adapter (glass M/F joints on corrugated Teflon tubing) for stepwise delivery of 2,2′-dithiobis(cyclohexanecarboxaldehyde) without opening the reactor.
11. Reflux the mixture for 1 hr, then evaporate to dryness and add 200 mL of 1.5 M sodium hydroxide. 12. Cautiously neutralize the mixture with 20% hydrochloric acid, checking pH with indicator strips. 13. Transfer the mixture to a 500-mL separatory funnel and extract the solution twice, each time with 150 mL chloroform. Dry the organic layer with anhydrous magnesium sulfate and evaporate the solvent. 14. Dissolve the product in 200 mL of diethyl ether and add, with stirring, a few 3- to 5-mL aliquots of hexane until the mixture becomes translucent. Leave in a refrigerator (4°C) overnight for crystallization. Collect crystalline S.7 by filtration. Approximately 19 g of 2,2′-dithiobis(cyclohexanemethanol) (S.7) should be collected as a colorless crystalline material (85% to 88% yield, m.p. 49° to 50°C). 1H NMR (CDCl3, δ): 3.54 ppm (s, 2H, CH2OH), 2.23 ppm (s, 1H, CH2OH), 1.2-1.8 ppm (m, 10H). 13C NMR (CDCl3, δ): 21.9, 25.73, 32.45, 56.18, 67.99 ppm. FAB MS (positive mode, Cs+, 13 keV, matrix NBA) m/z 290, [M]+, 45%; m/z 273, [M-OH]+, 25%; m/z 113, [C6H10CH2OH]+, 100%.
Synthesize 2-mercaptocyclohexanemethanol (S.8) 15. In a 1-L two-neck flask equipped with a reflux condenser and a dropping funnel (atmosphere of dry argon or nitrogen) suspend 5.9 g (0.16 mol) lithium aluminum hydride in 500 mL of dry diethyl ether. CAUTION: The suspension of lithium aluminum hydride in diethyl ether is highly flammable. Advise coworkers of the hazard and keep an appropriate fire extinguisher at hand.
Synthesis of Modified Oligonucleotides and Conjugates
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16. Add dropwise through the dropping funnel a solution of 4.6 g (0.16 mol) S.7 in 150 mL of diethyl ether over 60 min with magnetic stirring. Continue stirring for an additional 60 min. The reaction is exothermic and mild reflux occurs.
17. Cautiously decompose excess reducing agent by adding dropwise through the funnel 3 mL of dry ethyl acetate, followed by 10 mL THF containing traces of moisture, and then ∼10 mL of 10% water/THF, until the solid suspension becomes gray and finally white. 18. Filter off inorganic salts using a filter funnel and Whatman no. 1 filter paper, and dry the filtrate over anhydrous magnesium sulfate. Filter off the drying agent and evaporate the filtrate to dryness. 19. Distill the residue in a high-vacuum fractional distillation apparatus under reduced pressure (0.05 mmHg, provided by high-vacuum oil pump). Collect the fraction boiling between 74° and 76°C, which contains S.8. CAUTION: Avoid overheating the vessel. Keep pressure below 0.1 mmHg. Approximately 3.2 g of 2-mercaptocyclohexanemethanol (S.8) should be collected as a colorless oil (70% yield, nD20 = 1.5188). 1H NMR (CDCl3, δ): 3.49 ppm (s, 2H, CH2OH), 2.15 ppm (bs, 1H, CH2OH), 1.31 ppm (s, 1H, CHSH), 1.15-1.85 ppm (m, 10H). 13C NMR (CDCl3, δ): 22.0, 26.07, 36.06, 52.34, 73.12 ppm. FAB MS (negative mode, Cs+, 13 keV, matrix GLY) m/z 145, [M]–, 100. FAB MS (positive mode, Cs+, 13 keV, matrix GLY) m/z 113, [C6H10CH2OH]+, 45%; m/z 129, [C6H10SHCH2]+, 20%.
Synthesize 2-chloro-spiro-4,4-pentamethylene-1,3,2-oxathiaphospholane (S.9) 20. In a 1-L two-neck flask equipped with a thermometer and a dropping funnel, add 28.2 g (0.21 mol) PCl3 to 500 mL of dry benzene under an argon (optionally nitrogen) atmosphere. Cool the flask to 5°C with an ice bath. Add, dropwise through the funnel, a solution of 20 g (0.14 mol) S.8 and 22 mL (0.27 mol) pyridine in 35 mL dry benzene over a 15-min period with magnetic stirring. Keep the temperature of the reaction mixture below 10°C. 21. Continue stirring at room temperature for 30 min and filter off pyridine hydrochloride with exclusion of moisture. Load the reaction mixture in a filter funnel inside a bag filled with dry argon (or nitrogen) and gently apply suction to keep the bag slightly inflated with continuous delivery of dry gas. 22. Evaporate the solvent under reduced pressure with exclusion of moisture (preferably in a rotary evaporator equipped with a diaphragm vacuum). If a water aspirator must be used, insert a drying tube filled with blue indicator silica gel between the rotary evaporator and the aspirator to reduce the risk of hydrolysis of the product. Apply vacuum gently.
23. Distill the product in a high-vacuum fractional distillation apparatus under reduced pressure (0.01 mmHg, provided by a high-vacuum oil pump). Collect the fraction boiling between 82° and 84°C, which contains S.9. CAUTION: Avoid overheating the vessel. Keep pressure at 0.01 mmHg. When overheated, spontaneous decomposition of the crude product may occur, leading to destruction of the apparatus.
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
Approximately 20 g of 2-chloro-spiro-4,4-pentamethylene-1,3,2-oxathiaphospholane (S.9) should be collected as a colorless liquid (70% to 75% yield). 31P NMR (C6D6; δ) 217.7 ppm; EI (electron impact) MS: (70 eV) m/z 210, [M]+, 12%; m/z 175, [M-Cl]+, 5.8%; m/z 90, 100%.
24. Store S.9 in a tightly closed vessel inside another tightly closed container filled with several grams of dry 4A molecular sieves at –20°C (stable for at least 1 year).
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Current Protocols in Nucleic Acid Chemistry
SYNTHESIS OF 2,2′-DITHIOBIS([18O]CYCLOHEXANECARBOXALDEHYDE) The 18O-labeled 2,2′-dithiobis(cyclohexanecarboxaldehyde), which can be further transformed into the corresponding phosphitylating reagent as described in Basic Protocol 1, is obtained by hydrolysis of the N-phenylimine derivative of 2,2′-dithiobis(cyclohexanecarboxaldehyde) with H2[18O], catalyzed with gaseous hydrogen chloride (Fig. 4.17.5). The N-phenylimine derivative is obtained from 2,2′-dithiobis(cyclohexanecarboxaldehyde) (S.6; see Basic Protocol 1, step 8) upon treatment with aniline.
ALTERNATE PROTOCOL 1
Additional Materials (also see Basic Protocol 1) 2,2′-Dithiobis(cyclohexanecarboxaldehyde) (S.6; see Basic Protocol 1, step 8) Aniline, freshly distilled in inert atmosphere 95:5 (v/v) chloroform/hexane H2[18O] (95 atom%) Hydrogen chloride, anhydrous Tetrahydrofuran (THF), dried over sodium hydride 250-mL two-neck round-bottom flasks Azeotropic trap (e.g., mini Dean-Stark trap, Aldrich) 8 × 40–cm chromatography column packed with silica gel 60, 230 to 400 mesh (Merck) Rubber septum TLC silica gel plates with UV indicator (Merck; also see APPENDIX 3D) High vacuum valve (e.g., Rotaflo, Quickfit) Drying tube (8 × 5⁄8 in. with connectors, Aldrich) 2- to 5-mL gas-tight syringe Rotary evaporator with water aspirator or membrane pump Buchner funnel with glass frit Additional reagents and equipment for column chromatography (APPENDIX 3E), thin-layer chromatography (TLC; APPENDIX 3D), and high-vacuum transfer of solvent (see Support Protocol 1)
O
H S
H S
O
PhN
NPh S
PhNH2
S
−H2O 6
10 18O
18
[ O ]-H2O
H S
H S
HCl
18O
18O
1. NaBH 4
P Cl S
2. LiAlH 4 3. PCl 3 11
12
Figure 4.17.5 Synthesis of 2-chloro-spiro-4,4-pentamethylene-1,3,2-[18O]oxathiaphospholane (S.12) starting from 2,2′-dithiobis(cyclohexanecarboxaldehyde) (S.6).
Synthesis of Modified Oligonucleotides and Conjugates
4.17.9 Current Protocols in Nucleic Acid Chemistry
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Synthesize N-phenylimine derivative of 2,2′-dithiobis(cyclohexanecarboxaldehyde) (S.10) 1. In a 250-mL two-neck flask equipped with an azeotropic trap, reflux condenser, and dropping funnel, dissolve 14.3 g (0.05 mol) of 2,2′-dithiobis(cyclohexanecarboxaldehyde) (S.6) in 150 mL benzene. Add anti-bumping granules and heat to boiling. 2. Add, dropwise through the dropping funnel, a solution of 10.0 mL (10.2 g, 0.11 mol) aniline in 25 mL benzene over a 30-min period. Continue the reaction for 30 min, keeping gently boiling with azeotropic removal of liberated water. The end of the reaction is confirmed by disappearance of a resonance line of the aldehyde proton in the 1H NMR spectrum.
3. Cool the reaction mixture to room temperature and evaporate the solvent under reduced pressure using a rotary evaporator with a water aspirator. 4. Dissolve the residue in 15 to 20 mL of benzene and apply to an 8 × 40–cm chromatography column packed with ∼200 g of 230 to 400 mesh silica gel. 5. Elute the column with chloroform and collect the eluate in 12- to 15-mL fractions. 6. Analyze fractions by TLC on silica gel plates (APPENDIX 3D). Develop TLC plates with 95:5 (v/v) chloroform/hexane. 7. Combine all fractions that contain the desired product (S.10; Rf = 0.55). Evaporate the solvent under reduced pressure. Typically 18 g (80% yield) of the N-phenylimine derivative (S.10; see Fig. 4.17.5) should be obtained. 1H NMR (CDCl3, δ): 6.67-6.80 ppm (m, 1H), 7.07-7.55 ppm (m, 4H), 3.6 ppm (very broad singlet, 1H, CH=NPh), 1.29-2.18 ppm (m, 10H). 13C NMR (CDCl3, δ): 22.8, 24.88, 25.22, 30.05, 30.15, 33.23, 56.13, 56.793, 60.13, 60.62, 76.38, 77.01, 77.65, 114.84, 118.14, 120.63, 120.69, 125.42, 125.69, 128.11, 128.78, 129.00, 146.28, 150.99, 151.33, 164.78, 165.62, 194.09, 194.77. This number of resonances in the 13C NMR spectrum reflects the presence of cis- and trans-isomers of the N-phenylimine derivative.
Synthesize labeled 2,2′-dithiobis([18O]cyclohexanecarboxaldehyde) (S.11) 8. Place 16 g (0.037 mol) S.10 in a 250-mL two-neck round-bottom flask with a magnetic stir bar inside, with a high-vacuum valve in one joint and a rubber septum in the other, and dry overnight at high vacuum (0.01 mmHg; provided by high-vacuum oil pump). At the end of drying, deliver dry argon gas to the flask through the septum. 9. Prepare an absorber containing ∼150 mL of 4 to 5 M NaOH (see Basic Protocol 1, step 1). 10. Using the vacuum line technique (see Support Protocol 1), transfer ∼100 mL of dry THF to the flask containing S.10. 11. Connect the vacuum valve with the absorber through an 8 × 5⁄8–in. drying tube (filled with anhydrous magnesium sulfate) and a safety flask (Fig. 4.17.4). 12. Add 1.8 mL (0.9 mol) H2[18O] (20% excess) with a 2- to 5-mL gas-tight syringe. 13. Flush the apparatus continously with dry argon and cool the mixture to <5°C in an ice bath.
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
14. Remove the septum and quickly install a glass gas inlet adapter for delivery of hydrogen chloride. Adjust the adapter so that its end is ∼1.5 to 2 cm above the level of the liquid. 15. Slowly deliver anhydrous hydrogen chloride with magnetic stirring of the mixture. Precipitation of anilinium hydrochloride should be observed during this process.
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16. Continue hydrolysis until the reaction mixture becomes pale green-yellow. Remove ice bath and allow the mixture to reach room temperature. 17. Filter off the precipitate using a Buchner funnel. Wash twice with 20 to 30 mL THF. 18. Combine filtrates and evaporate solvent under reduced pressure. CAUTION: The THF vapor will be strongly acidic, as it is saturated with hydrogen chloride.
19. Dissolve the residue in 100 mL of THF and evaporate the solvent again. 20. Dissolve the residue in diethyl ether and crystallize by cooling (see Basic Protocol 1, step 8). 21. Analyze the product using electron impact (EI) mass spectrometry. Each aldehyde group of the final product is isotopically labeled to the extent of 87%. The spectrum should contain three peaks at m/z 286, 288, and 290, corresponding to the unlabeled, singly labeled, and doubly labeled compounds, respectively. The intensities of the ions are 1.7%, 22.6%, and 75.7%, respectively. The isotope content remains unchanged during subsequent reactions leading to the final phosphitylating reagent.
22. Continue the synthesis (see Basic Protocol 1, starting from step 9). HIGH-VACUUM TECHNIQUE FOR TRANSFER OF DRY SOLVENTS This technique allows for transfer of dry solvents from a reservoir (where the solvent is stored over a drying reagent such as sodium hydride) into a reaction flask with exclusion of moisture. For successful transfer, the quality of vacuum valves is absolutely essential. Figure 4.17.6 shows principal construction of the apparatus, which can be assembled from generic glassware. All components must be tested to make sure that they are safe for high-vacuum usage (i.e., at <0.5 mmHg).
SUPPORT PROTOCOL 1
1. Set up the apparatus as shown in Fig. 4.17.6, but leave the flask containing THF over NaH not yet immersed in the warm water bath and the receiving flask not yet immersed in the dry ice/isopropanol bath. 2. Close high-vacuum valves A and B. 3. Attach the apparatus to high vacuum (<0.5 mmHg). 4. Open valve B for several seconds. 5. Close valve B. 6. Open valve A for several seconds. If gentle boiling of THF is observed, go to step 9. 7. Close valve A. 8. Decrease the pressure in the system by repeating steps 4 to 7. 9. While valve B is closed, immerse the receiving flask in the dry ice/isopropanol bath. 10. Immerse the reservoir in the warm water bath (∼50° to 55°C). Alternatively, use a hair dryer to provide heat to the reservoir. 11. After the required volume of solvent has been transferred, stop heating and remove the dry ice bath. Allow both flasks to reach ambient temperature. 12. Disconnect the apparatus from the high vacuum and attach the outlet to a line with dry argon.
Synthesis of Modified Oligonucleotides and Conjugates
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to high vacuum
valve B
valve A
dry ice/isopropanol bath THF over NaH warm water bath Figure 4.17.6 Apparatus for high-vacuum transfer of dry solvents.
13. Open valve B and fill the system with argon. 14. Close valves A and B. 15. Disconnect the flask with transferred solvent and close immediately with a stopcock or septum. BASIC PROTOCOL 2
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
SYNTHESIS OF 5′-O-DMTr-DEOXYRIBONUCLEOSIDE-3′-O-(2-THIO-4,4spiro-PENTAMETHYLENE-1,3,2-OXATHIAPHOSPHOLANE)S AND THEIR SEPARATION INTO P-DIASTEREOMERS The oxathiaphospholane method of stereocontrolled synthesis of PS-oligos, depicted in Figure 4.17.2, is based on availability of diastereomerically pure oxathiaphospholane monomers S.1-S.3. Separation of monomers S.1 and S.2 as pure P-diastereomers requires tedious silica gel column chromatography or costly preparative HPLC. Monomer S.3, obtained by introducing a pentamethylene substituent at position 4 of the oxathiaphospholane ring, possesses a satisfactory separability of diastereomers. Appropriate 5′-ODMTr-N-protected deoxyribonucleosides are phosphitylated at room temperature with S.9 or S.12 in acetonitrile, in the presence of diisopropylethylamine, to yield the corresponding phosphites, which are further sulfurized with elemental sulfur. Silica gel column chromatography affords S.3 (or its 18O-labeled analog) as a diastereomeric mixture in satisfactory yield (75% to 85%). The deoxyguanosine derivative of S.3 should be additionally protected at O6 with diphenylcarbamoyl chloride to improve the yield of the condensation step. Without this protection, the repetitive yield of condensation of the deoxyguanosine oxathiaphospholane monomer drops below 90%. The S.3 monomers (B = T, ABz, CBz, Gi-Bu,DPC) are separated by column chromatography into fast- and sloweluting species. The stereochemistry of the coupling has been checked for each of 32 combinations of diastereomeric dinucleotides NPSN (N = dG, dA, dC, T). All four fast-eluting diastereomers of S.3 are precursors of the dinucleoside 3′,5′-phosphorothioates of RP configuration. Slow-eluting isomers of S.3 yield phosphorothioate linkages of SP configuration.
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Current Protocols in Nucleic Acid Chemistry
NOTE: It is important to use acid-free eluents for all silica gel chromatography of the monomers to avoid partial detritylation of the products. This can be done by distillation of chloroform with 1 mL pyridine, and then by adding 2 to 3 mL pyridine per liter of all eluents (chloroform and mixtures of ethyl acetate, butyl acetate, and benzene). Materials 5′-O-DMTr-N-protected deoxyribonucleosides (Chemgenes): N6-Benzoyl-5′-O-(4,4′-dimethoxytrityl)-2′-deoxyadenosine (5′-O-DMTr-dABz) N2-Isobutyryl-5′-O-(4,4′-dimethoxytrityl)-2′-deoxyguanosine (5′-O-DMTr-dGi-Bu) N4-Benzoyl-5′-O-(4,4′-dimethoxytrityl)-2′-deoxycytidine (5′-O-DMTr-dCBz) 5′-O-(4,4′-Dimethoxytrityl)-2′-deoxythymidine (5′-O-DMTr-T) Argon (or, optionally, nitrogen), dry Acetonitrile, anhydrous 2-Chloro-spiro-4,4-pentamethylene-1,3,2-oxathiaphospholane (S.9 or labeled S.12; see Basic Protocol 1 or Alternate Protocol 1) Elemental sulfur, anhydrous (dried overnight at high vacuum) Chloroform (distilled with 1 mL pyridine per L) Toluene, dry Pyridine, anhydrous Diisopropylethylamine (Aldrich), anhydrous Diphenylcarbamoyl chloride (Aldrich) 9:1 (v/v) chloroform/methanol Merck 60H silica gel, particle size 5 to 40 µm Ethyl acetate Butyl acetate Benzene 25-mL two-neck round-bottom flasks High-vacuum valve (Rotaflo, Quickfit) Rubber septum High-vacuum oil pump (0.01 mmHg) 2-mL and 10-mL gas-tight syringes Buchner funnel 25 × 3–cm chromatography column packed with 20 g of 230 to 400 mesh silica gel TLC silica gel plates with UV indicator (Merck; also see APPENDIX 3D) Constant temperature water bath 30 × 2–cm chromatography column Filter paper (Whatman no. 1) High-performance TLC (HP-TLC) plates (silica gel 60 F254) with UV indicator (Merck; also see APPENDIX 3D) Additional reagents and equipment for column chromatography (APPENDIX 3E) and thin-layer chromatography (TLC; APPENDIX 3D) Phosphitylate protected deoxyribonucleosides 1. Place 10 mmol of 5′-O-DMTr-N-protected deoxyribonucleoside (dABz, dGi-Bu, T, or dCBz) in a 25-mL two-neck round-bottom flask containing a magnetic stir bar, with a high-vacuum valve in one joint and a rubber septum in the other, and dry overnight at high vacuum (0.01 mmHg; provided by high-vacuum oil pump). 2. Close the vacuum valve and fill the flask with dry argon (optionally nitrogen). 3. Using a gas-tight syringe, add 10 mL of anhydrous acetonitrile through the septum. 4. Using a gas-tight syringe, add 1.91 mL (11 mmol) of anhydrous diisopropylethylamine.
Synthesis of Modified Oligonucleotides and Conjugates
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5. To the magnetically stirred solution, using a gas-tight syringe, add dropwise at room temperature 1.63 mL (2.32 g, 11 mmol) of 2-chloro-spiro-4,4-pentamethylene-1,3,2oxathiaphospholane (S.9) over a 5-min period. 6. Stir an additional 5 min and add ∼0.5 g (∼15 mmol) dry elemental sulfur. Continue stirring for 12 hr. 7. Filter off excess sulfur using a Buchner funnel. Evaporate the solvent and dissolve the residue in 4 mL of chloroform (distilled with pyridine). 8. Apply crude product to a 25 × 3–cm column packed with 20 g of 230 to 400 mesh silica gel. Elute the column with chloroform, collecting 8- to 10-mL fractions. Identify the appropriate fractions by TLC (APPENDIX 3D) using standard silica gel plates and 95:5 (v/v) CHCl3/methanol (Rf = 0.6575). Combine fractions and evaporate the solvents under reduced pressure (15 to 20 mmHg) with a water bath temperature not exceeding 30°C. 9. Add 5 to 6 mL of dry toluene and evaporate to dryness with exclusion of moisture using a membrane pump. Repeat this step twice. After the solvent is evaporated, apply high vacuum to the flask for 2 hr. Close the flask with a septum and pierce the septum with a needle. Store the flask in a desiccator and apply high vacuum for ≥12 hr. The monomers can be stored at room temperature in a desiccator for a month. Th e desired 5′-O-DMTr-deoxyribonucleoside-3′-O-(2-thio-spiro-4,4-pentamethylene1,3,2-oxathiaphospholane)s are obtained in 75% to 85% yield. For the guanosine derivative, diphenylcarbamoyl protection at O6 is required (steps 10 to 15). The diastereomeric composition, 31P NMR chemical shifts, and TLC parameters (HP-TLC plates) of compounds S.3a to c are given in Table 4.17.1. Elemental analysis (found/calculated): S.3a (B = T) C 61.67%/60.79%, H 6.15%/5.77%, N 3.73%/3.73%, P 4.06%/4.13%, S 8.00%/8.54%; S.3b (B = ABz) C 62.84%/62.56%, H 5.48%/5.37%, N 7.73%/8.11%, P 3.41%/3.58%, S 6.74%/7.42%; S.3c (B = CBz) C 62.53%/62.92%, H 5.47%/5.52%, N 5.14%/5.00%, P 3.65%/3.69%, S 7.26%/7.63%.
Table 4.17.1 Characteristics of the 5′-O-DMTr-deoxyribonucleoside-3′-O-(2-thio-spiro-4,4pentamethylene-1,3,2-oxathiaphospholane)sa
Base
Yield (%)
Composition (fast:slow)
δP (ppm, CD3CN)
Rf (TLC)b
T (3a)
84
50:50
ABz (3b)
84
49:51
CBz (3c)
86
48:52
Gi-Bu,DPC (3d)
78
52:48
105.3 (fast) 105.6 (slow) 104.7 (fast) 105.1 (slow) 105.3 (fast) 105.6 (slow) 106.2 (fast)d 106.9 (slow)d
0.61 (fast) 0.54 (slow) 0.54 (fast) 0.46 (slow) 0.60 (fast) 0.40 (slow) 0.37 (fast)c 0.26 (slow)c
aTable adapted from Stec et al. (1998) with permission from the American Chemical Society. bTLC performed on HP-TLC plates with UV indicator (Merck) and a developing system of 1:1 (v/v) butyl acetate/benzene
(T and Gi-Bu) or 1:2 (v/v) ethyl acetate/butyl acetate (ABz and CBz).
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
cR values reported for monomers before O6-protection with diphenylcarbamoyl chloride (DPC). After protection, R f f
values are 0.74 and 0.63 using the same solvent system. dδ values reported for monomers after O6-protection with DPC. P
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Perform O6 protection of deoxyguanosine derivative 10. Place 0.85 g (1 mmol) of N2-isobutyryl-5′-O-(4,4′-dimethoxytrityl)-3′-O-(2-thiospiro-4,4-pentamethylene-1,3,2-oxathiaphospholanyl)-2′-deoxyguanosine (from step 9) in a two-neck 25-mL round-bottom flask with a magnetic stir bar, with a high-vacuum valve in one neck and a rubber septum in the other, and dry overnight at high vacuum (0.01 mmHg). 11. Using a 10-mL gas-tight syringe, add 5 mL of anhydrous pyridine. 12. Using a 2-mL gas-tight syringe, add 0.26 mL (1.5 mmol) diisopropylethylamine and 0.46 g (2.0 mmol) diphenylcarbamoyl chloride, with stirring, at room temperature. Continue stirring mixture 1 hr. 13. Concentrate mixture to dryness, dissolve in 1.5 mL of chloroform, and apply on a 25 × 3–cm column packed with 20 g of 230 to 400 mesh silica gel. Elute the column with 300 mL chloroform, collecting 10- to 12-mL fractions. 14. Analyze fractions by TLC (APPENDIX 3D) on standard silica gel plates. Develop TLC plates with 9:1 (v/v) chloroform/methanol. 15. Combine all fractions that contain the desired product (Rf = 0.79). Evaporate the solvents under reduced pressure (15 to 20 mmHg) with the temperature of the water bath not exceeding 35°C. Dissolve the residue in dry toluene and evaporate the solvent. Store the pure product S.3d (a pale yellow oil) in a tightly closed vessel. Approximately 0.95 g (90% to 95% yield) should be isolated. MS. (+FAB) m/z 1041.6 (M+, 1%), m/z 1042.6 (M++1, 0.6%), m/z 303.2 (DMTr+, 100%). Elemental analysis(found/calculated): C 63.60%/63.44%, H 5.78%/5.52%, N 7.77%/8.08%, P 2.72%/2.98%, S 5.78%/6.15%. The diastereomeric composition, 31P NMR chemical shifts, and TLC parameters (HP-TLC plates) of S.3d are given in Table 4.17.1.
Separate diastereomers of S.3a to d 16. The day before separation, load a 30 × 2–cm column with a degassed suspension of ∼20 g of silica gel (Merck 60H, particle size 5 to 40 µm) in ∼100 mL of the appropriate mixture of solvents: 2:1:0.003 (v/v/v) ethyl acetate/butyl acetate/pyridine for dA and dC derivatives 1:2:0.003 (v/v/v) ethyl acetate/butyl acetate/pyridine for dG derivative 1:1:0.002 (v/v/v) butyl acetate/benzene/pyridine for T monomer. Gently cover the top surface of the gel with a disc of Whatman no.1 filter paper of diameter close to the inside diameter of the column. Flush the column with 150 mL of the eluant. Maintain a 2- to 3-mm layer of eluant over the gel. IMPORTANT NOTE: Because the differences in chromatographic mobilities of P-diastereomers are very small, the glass frit must be mounted within the cylindrical part of the column to assure undisturbed, laminar flow of the eluant. To achieve good resolution, the column should be packed with the silica gel suspension at least 24 hr before chromatography. Since isocratic elution is used for the separation, the same column may be used for two to three consecutive separations of a given monomer.
17. Dissolve ∼300 mg of a monomer (mixture of diastereomers) in 1.5 mL of the appropriate eluant (see step 16) and apply gently on the gel. 18. Elute the column with 300 mL of appropriate eluant and collect 10- to 12-mL fractions. Analyze fractions by TLC on HP-TLC plates (Table 4.17.1). Combine appropriate fractions and concentrate to dryness under reduced pressure (15 to 20 mmHg) with the temperature of the water bath not exceeding 35°C.
Synthesis of Modified Oligonucleotides and Conjugates
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19. Evaporate the pure diastereomers twice with anhydrous toluene, with exclusion of moisture, and store in a tightly closed vessel for up to 1 month at room temperature. Typically, for dA, dT, and dC monomers, one passage gives 75% to 80% separated diastereomers of 96% to 100% diastereomeric purity, as assessed by 31P NMR. For the dG derivative, the “fast” isomer is usually obtained in lower yield (28% to 30%, 100% diastereomeric purity) while the “slow” isomer is obtained in ∼50% yield, but only of 90% diastereomeric purity, and must be rechromatographed. ALTERNATE PROTOCOL 2
SYNTHESIS OF 5′-O-DMTr-DEOXYRIBONUCLEOSIDE-3′-O-(2-OXO-spiro4,4-PENTAMETHYLENE-1,3,2-OXATHIAPHOSPHOLANE)S 5′-O-DMTr-deoxyribonucleoside-3′-O-(2-oxo-spiro-4,4-pentamethylene-1,3,2-oxathiaphospholane)s (S.4) are synthesized from their corresponding 2-thio monomers (S.3; see Basic Protocol 2) and can be used to elongate PS-oligos obtained via the oxathiaphospholane approach and thereby generate short unmodified oligonucleotide segments. This goal cannot be achieved using standard phosphoramidite chemistry because, during the routine oxidation step by means of I2/water/pyridine, the diester phosphorothioate linkages already present in the oligomer would undergo PS-to-PO conversion. The oligonucleotide synthesis protocol differs from that used for the synthesis of PS-oligos in the following ways: (1) because of their relatively low stability, the 2-oxo-monomers should not be repurified before synthesis (see Basic Protocol 3, step 1); and (2) the amount of DBU necessary for condensation (step 16) may be reduced by 50%. Materials 5′-O-DMTr-deoxyribonucleoside-3′-O-(2-thio-spiro-4,4-pentamethylene-1,3,2oxathiaphospholane)s (S.3; see Basic Protocol 2) Silica gel 60, 230 to 400 mesh Acetonitrile, anhydrous Argon (or, optionally, nitrogen), dry Selenium dioxide, anhydrous (dried overnight at high vacuum) 95:5 (v/v) chloroform/methanol (distill chloroform with 1 mL pyridine per L) Two-neck 10-mL round-bottom flask High-vacuum valve (e.g., Rotaflo, Quickfit) Rubber septum High-vacuum oil pump 10-µmol-scale DNA synthesis column 1- to 2-mL polypropylene syringes with luer ends 5-mL gas-tight syringe TLC silica gel plates with UV indicator (Merck; also see APPENDIX 3D) 10-mL gas-tight syringe Luer male-to-male adapter Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) CAUTION: Selenium dioxide is toxic.
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
1. Place ∼300 mg of 5′-O-DMTr-deoxyribonucleoside-3′-O-(2-thio-spiro-4,4-pentamethylene-1,3,2-oxathiaphospholane) (S.3) in a two-neck 10-mL round-bottom flask containing a magnetic stir bar, with a high-vacuum valve in one joint and a rubber septum in the other, and dry overnight at high vacuum (0.01 mmHg; provided by high-vacuum oil pump).
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2. Assemble a 10-µmol-scale DNA synthesis column filled to 80% of its volume with silica gel 60 (230 to 400 mesh). Wash the gel three times, each time with 5 mL anhydrous acetonitrile. Apply a stream of dry argon and continue drying for 15 to 20 min after the gel loosens. Close both ends of the column with two 1- to 2-mL luer syringes to eliminate contact with atmospheric moisture. The dryness of the gel in the column is crucial for stability of the product. Every effort should be made to protect the dried gel from moisture.
3. Close the vacuum valve on the flask containing dried S.3 and fill the flask with dry argon. 4. Using a 5-mL gas-tight syringe, add 4 mL of anhydrous acetonitrile. 5. With continuous flow of dry argon, remove the vacuum valve and add, in several portions, ∼80 mg selenium dioxide. 6. Monitor the progress of the reaction by TLC (APPENDIX 3D) on silica gel plates. Develop TLC plates with 95:5 (v/v) chloroform/methanol (see Table 4.17.2 for Rf values of S.4). Because the differences in chromatographic mobilities are small, always apply the starting material on an adjacent lane as a reference.
7. After the reaction is complete, withdraw the supernatant with a 10-mL gas-tight syringe and remove the needle. 8. Attach a luer male-to-male adapter to the bottom of the silica gel column and gently load the withdrawn supernatant at the top. Collect the effluent in a dry flask with continuous flow of dry argon. Wash the silica gel with 1 to 2 mL of dry acetonitrile. As filtration progresses, an orange layer of selenium compounds moves through the gel to the bottom of the column. Stop the filtration when the layer reaches two-thirds of the column length from the top.
9. Replace the high-vacuum valve on the flask and evaporate acetonitrile at high vacuum. Apply the vacuum slowly while shaking the flask continuously until the oil residue makes a foam. 5′-O-DMTr-deoxyribonucleoside-3′-O-(2-oxo-spiro-4,4-pentamethylene-1,3,2-oxathiaphospholane)s (S.4) are unstable and should be used for synthesis within a few hours after preparation. They are typically isolated at 40% to 55% yield. Their 31P NMR chemical shifts and TLC parameters are reported in Table 4.17.2.
Table 4.17.2 Characteristics of the 5′-O-DMTr-deoxyribonucleoside-3′-O-(2-oxo-spiro-4,4pentamethylene-1,3,2-oxathiaphospholane)sa
Base T (4a) ABz (4b) CBz (4c) Gi-Bu,DPC (4d)
Yield (%)
δP (ppm, CD3CN)
Rf (TLC)b
55 41 45 54
44.7, 44.3 45.1, 44.9 44.6, 44.1 45.3, 44.5
0.71 0.71 0.70 0.74
aTable adapted from Stec et al. (1998) with permission from the American Chemical Society. bTLC performed on silica gel plates with UV indicator (Merck) and a developing system of 95:5 (v/v) chloroform/meth-
anol. The 2-oxo-monomers migrate slightly more slowly than their 2-thio-precursors.
Synthesis of Modified Oligonucleotides and Conjugates
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BASIC PROTOCOL 3
MANUAL SOLID-PHASE SYNTHESIS OF STEREODEFINED OLIGO(NUCLEOSIDE PHOSPHOROTHIOATE)S Synthesis of oligo(nucleoside phosphorothioate)s using the oxathiaphospholane method can be performed either in a standard 1-µmol column (1-µmol-scale synthesis) or in a reassembled OPC column (2-µmol scale; see below). Because of the strong base used for the condensation step (1,4-diazabicyclo[5.4.0]undec-7-ene), a sarcosinylated solid support must be used (see Support Protocol 2). Typically, supports are functionalized with 25 to 40 µmol nucleoside per gram. A lower nucleoside concentration is not recommended, because a larger amount of support will be necessary to achieve the synthesis at the recommended scale, and may not leave enough space for efficient mixing with incoming reagents. The condensation step is extremely sensitive even to trace amounts of moisture. Therefore, the acetonitrile to be used as a solvent for DBU, and deoxyribonucleoside oxathiaphospholane monomers should be dried over P2O5 (5 g/L) and distilled under reduced pressure (∼200 mmHg) through a 20-cm Vigreux column with exclusion of moisture under an atmosphere of dry argon. At least one-third of the initial volume must remain in the flask. Acetonitrile dried in this way must be transferred using a gas-tight syringe under an atmosphere of dry argon, or by the vacuum line technique (see Support Protocol 1). This protocol describes a 2-µmol-scale synthesis. The synthesis proceeds from the 3 ′-end to the 5 ′-end of the sequence. The first nucleoside from the 3′-end is attached to the solid support. To avoid mistakes, the investigator is recommended to have a synthesis step check list to check off each of the executed steps during consecutive synthetic cycles, as illustrated in Table 4.17.3 for synthesis of the sequence 5′-TGACTGCA-3′. Notably, after the last condensation step, the capping procedure is not executed. Materials Deoxyribonucleoside oxathiaphospholane monomers (see Basic Protocol 2 and/or Alternate Protocol 2) Chloroform (optional), distilled with 1 mL pyridine per L Toluene, anhydrous Low-pressure argon or nitrogen, dried (see recipe) Sarcosinylated solid support functionalized with a nucleoside (first from the 3′ end of sequence to be synthesized) at a concentration ranging from 20 to 30 µmol/g support (see Support Protocol 2) Capping reagent A: 1:1:8 (v/v/v) acetic anhydride/pyridine/tetrahydrofuran (THF) Capping reagent B: 7 g 4-dimethylaminopyridine/93 mL THF Anhydrous acetonitrile (H2O < 20 ppm; see recipe) in bottle with rubber septum, with dry gas delivered inside through a line ending in a needle
Table 4.17.3 Sample Check List for Oligonucleotide Synthesis (Sequence 5′-TGACTGCA-3′)
Nucleoside monomer
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
— C-OTP monomer G-OTP monomer T-OTP monomer C-OTP monomer A-OTP monomer G-OTP monomer T-OTP monomer
Detritylation
Coupling
Capping
— √
— √
√
—
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Detritylating reagent: 3.5% (w/v) dichloroacetic acid in methylene chloride Acetonitrile, HPLC grade (Baker) 1:4.5 (v/v) 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) in anhydrous acetonitrile (H2O <20 ppm) Methylene chloride Aqueous ammonia, concentrated (Baker) 10 × 3–cm chromatography column packed with 10 g 230- to 400-mesh silica gel 60 (optional) 4-mL sample vials with open-top screw caps with Teflon-faced rubber septa 19- to 22-G, 1- to 1.5-in. luer-lock needles Vacuum desiccator (≤0.05 mmHg; provided by high-vacuum oil pump) with condenser cooled by liquid N2 1-, 2-, and 5-mL all-polypropylene luer-lock syringes 19- to 22-G, 2- to 3-in. luer-lock needles with blunt 90° tips Columns for DNA synthesis: For 1-µmol scale: Applied Biosystems DNA synthesis column (cat. no. 400407), empty, 1.0 µmol crimp-style For 2-µmol scale: emptied and reassembled Applied Biosystems oligonucleotide purification cartridge (OPC; cat. no. 400771) Column filters (two for each column; Applied Biosystems, cat. no. 400059) Aluminum seals (caps; two for each column; Aldrich cat. no. Z11413-8; Wheaton aluminum cap, 13 mm, tear-off) Crimper for aluminum seals (Aldrich, cat. no. z 11423) Polypropylene (or other chemically inert) luer male-to-male adapter 100- and 500-µL gas-tight syringes (Hamilton) Glass drying tube (∼2-cm i.d., 25-cm length), with three-way valve (or two independent valves) at the top and a rubber septum at the opposite side High-vacuum (0.05 mmHg) oil pump 500-mL filtering flask capped with a rubber septum pierced with a 3- to 4-mm i.d. hole Water aspirator with a manostat Two waste bottles: one for chlorinated waste (methylene chloride, detritylating reagent) and another for water-miscible wastes (acetonitrile, capping reagents A and B) 60°C water bath (optional) Speedvac concentrator with vacuum provided by water aspirator 19- to 22-G, 4- to 5-in. luer-lock needles with blunt 90° tips Additional reagents and equipment for column chromatography (optional; APPENDIX 3E) and purification of oligonucleotides (UNITS 10.3-10.5) CAUTION: The drying tube must be tested to make sure that it is safe for high-vacuum usage, i.e., at a pressure lower than 0.05 mmHg. Prepare monomers for synthesis 1. If the deoxyribonucleoside oxathiaphospholane monomers (S.3 only) have been stored for more than one week, repurify them by flash chromatography (APPENDIX 3E) on a 10 × 3–cm silica gel column using chloroform (distilled with pyridine) as eluant. Concentrate appropriate fractions with exclusion of moisture. Coevaporate the residue twice with anhydrous toluene and apply high vacuum to generate a foam. Repurification is necessary to keep repetitive yield at 92% to 94%. The 2-oxo-monomers (S.4) should not be repurified in this manner, and should be used within a few hours of synthesis (see Alternate Protocol 2).
Synthesis of Modified Oligonucleotides and Conjugates
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2. If necessary, use a spatula to disaggregate the foam of oxathiaphospholane monomers. 3. Put ∼30 mg of given monomer (S.3 or S.4) in an appropriately marked 4-mL sample vial (one vial for each condensation step to be performed) and cover tightly with an open-top screw cap with a Teflon-faced rubber septum. Pierce each septum with a 19- to 22-G, 1- to 1.5-in. luer-lock needle. Before the vials are stored in the desiccator (step 4), be sure that none of the needles became clogged during piercing. This can be done by temporarily inserting a second needle to deliver a stream of dry argon to the vial. The unrestricted flow of argon from the first needle should be easily sensed.
4. Dry monomers in a vacuum desiccator at ≤0.05 mmHg for at least 12 hr. After the drying is complete, fill the desiccator with dry argon, open it cautiously, and immediately remove the needles from the vials. The condenser should be free of any solvents before the drying starts. It is recommended that the condenser be cooled with liquid nitrogen. It is imperative that the oil pump and the whole vacuum system intended for drying monomers be free of acidic impurities such as acetic or hydrochloric acid. Traces of acids would partially detritylate the monomers.
Perform initial setup procedures 5. Prepare and place a permanent mark upon all-polypropylene syringes for capping (2 mL), detritylation (5 mL), washing with acetonitrile (5 mL), and washing with methylene chloride (5 mL). Fit each with a 19- to 22-G, 2- to 3-in. needle with a blunt 90° tip. Use the syringes only for designated purposes, as cross-contamination may lead to poor synthesis yields. The needles should be blunted for safety reasons.
6. Fill a column with an appropriate support for synthesis and assemble it. Be sure to insert column filters at both ends of the column. Secure the aluminum caps with a crimper. Remove the middle part of each aluminum cap. Insert a luer male-to-male adapter into one outlet of the column and attach the outlet needle (19- to 22-G, 4- to 5-in. with a blunt 90° tip). Do not hesitate to apply significant force to secure aluminum caps. This is crucial for avoiding leakage of chemicals during synthesis. The long outlet needle (4- to 5-in.) is more convenient for swirling of the column during consecutive synthetic steps.
7. Set the manostat for the filtering flask at 400 to 450 mmHg. Start synthesis cycle 8. Mix 1 mL capping reagent A and 1 mL capping reagent B in the 2-mL dedicated syringe for capping (see step 5), remove the needle, and gently load the mixture into the column. Continue capping for 2 min with intermittent swirling of the column. Expel reagent from the column into a proper waste bottle. CAUTION: Avoid undue force when pushing the plunger. Excessively fast loading or expelling of the reagents may damage the column filters.
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
9. Fill the dedicated 5-mL syringe for acetonitrile washing (see step 5) with HPLC-grade acetonitrile. Remove the needle and flush the column with intermittent swirling. Collect the effluent in the proper waste bottle. Expel the remaining liquid, then insert the outlet needle of the column through the septum on the top of the filtering flask, deliver a stream of argon from the top of the column, and gently apply suction to the filtering flask. Continue drying until the support in the column becomes loose.
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10. Fill the dedicated 5-mL syringe for detritylation (see step 5) with detritylating reagent. Remove the needle and flush the column with intermittent swirling. Collect the effluent in the proper waste bottle or in a test tube for quantitative cationic DMTr analyses. Expel the remaining liquid. IMPORTANT NOTE: Upon contact with the support, the detritylation solution becomes red due to the presence of DMTr cations removed from the nucleoside. If at the end of delivery the effluent is still colored, continue detritylation with an additional volume of fresh reagent to complete the detritylation process.
11. Wash the support with 5 mL of acetonitrile and dry it as described in step 9. 12. Put the column and gas-tight syringes (500 µL and 100 µL) in the drying tube, gently apply high vacuum, and continue drying for 10 min. Close the vacuum valve on the drying tube and deliver dry gas to the tube. Take the syringes out of the tube (leave the column inside), close the tube with the septum, close the gas valve, and apply vacuum again. Use the syringes only for designated purposes. CAUTION: Do not allow the rubber septum to be expelled from the bottom of the tube by excessively high gas pressure.
13. Withdraw 300 µL of dry acetonitrile (H2O <20 ppm) using the 500-µL gas-tight syringe and add the solvent to the vial containing the appropriate oxathiaphospholane monomer. 14. Close the vacuum valve on the drying tube and deliver dry gas to the tube. Take the column out of the tube, close the tube with the septum, and close the gas valve. Insert a dry 1-mL syringe into the inlet of the column. 15. Withdraw 90 µL of 1:4.5 (v/v) DBU/acetonitrile using the 100-µL gas-tight syringe and add the reagent to the vial containing the dissolved oxathiaphospholane monomer. Mix the contents of the vial for a few seconds. 16. Pierce the septum of the vial with the needle of the column and, using the syringe attached to the column, suck up the contents of the vial to fill the column. Swirl the column intermittently for 10 min. Expel the liquid to the waste bottle. 17. Wash the support with 5 mL of methylene chloride, followed by 5 mL of HPLC-grade acetonitrile, and dry as described in step 9. Complete synthesis cycle 18. Repeat steps 8 to 17 with each successive monomer until elongation is complete. 19. Execute a routine cleavage of the product from the support using a few milliliters of concentrated aqueous ammonia for 2 hr, followed (if necessary) by heating of the ammoniacal solution in a tightly closed vessel at 55°C for 12 hr to remove nucleobase-protecting groups. 20. Concentrate the sample under reduced pressure in a Speedvac concentrator using a water aspirator. 21. Purify the oligonucleotide according to published protocols (UNITS 10.3-10.5).
Synthesis of Modified Oligonucleotides and Conjugates
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SUPPORT PROTOCOL 2
ATTACHMENT OF NUCLEOSIDE 3′-O-SUCCINYL HEMIESTERS TO SARCOSINYLATED SOLID-PHASE SUPPORT This protocol attaches 5′-O-DMTr-N-protected nucleoside-3′-O-succinyl hemiesters to long-chain alkylamine controlled-pore glass (LCAA-CPG) through a sarcosinyl linker. Since 5′-O-DMTr-N-protected nucleoside-3′-O-succinate hemiesters are commercially available, their syntheses are not described. Alternatively, one may prepare them by coupling succinic anhydride with a protected nucleoside according to published procedures (see UNIT 3.2 and references therein). Materials Long-chain alkylamine controlled-pore glass (LCAA-CPG) beads (80 to 120 mesh, 500 Å; Sigma) 9-Fluorenylmethoxycarbonyl (Fmoc)–sarcosine monohydrate (Fluka) Argon (or, optionally, nitrogen), dry 1,3-Dicyclohexylcarbodiimide (DCC; Aldrich) Dimethylformamide (DMF), anhydrous Pyridine, anhydrous 1:1:1 (v/v/v) acetonitrile/methanol (reagent grade)/pyridine Acetonitrile, anhydrous 10% (v/v) piperidine in pyridine 5′-O-DMTr-N-protected nucleoside-3′-O-succinyl hemiester (Sigma) 25-mL two-neck round-bottom flasks High-vacuum valve (e.g., Rotaflo, Quickfit) Rubber septum High-vacuum oil pump 1-mL and 10-mL gas-tight syringes Buchner funnel with glass frit Filter flask Vacuum source (e.g., water aspirator) 50-mL Erlenmeyer flask with stopcock CAUTION: 1,3-Dicyclohexylcarbodiimide can cause skin or eye irritation and allergic reactions. Use appropriate protection. Couple Fmoc-sarcosine to LCAA-CPG 1. Place 2 g LCAA-CPG beads and 0.5 g (1.6 mmol) Fmoc-sarcosine in a 25-mL two-neck round-bottom flask containing a magnetic stir bar, with a high-vacuum valve in one joint and a rubber septum in the other, and dry overnight at high vacuum (0.01 mmHg; provided by a high-vacuum oil pump). At the end of drying, deliver dry argon (or nitrogen) to the flask through the septum. 2. Open the flask, quickly add 0.5 g (2.4 mmol) DCC, and close the flask. 3. Using gas-tight syringes, add 5 mL anhydrous DMF followed by 0.5 mL anhydrous pyridine. 4. Stir the mixture at room temperature for 12 hr. 5. Transfer the suspension to a fritted-glass Buchner funnel and filter off the solution.
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
6. Wash support on the funnel three times, each time with 50 mL of 1:1:1 (v/v/v) acetonitrile/methanol/pyridine with gentle suction. 7. Wash with 50 mL acetonitrile and continue suction for 2 to 3 min.
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8. Transfer the support to a 50-mL Erlenmeyer flask, add 20 mL of 10% piperidine in pyridine, and seal the flask. Shake occasionally over a 30-min period. Decant the liquid. 9. Filter and wash as in steps 5 to 7. 10. Allow the support to dry at room temperature in a fume hood. Couple nucleoside-3′-O-succinyl hemiesters to LCAA-CPG-Sar 12. Place 2 g LCAA-CPG-Sar and ∼0.35 g (∼0.5 mmol) 5′-O-DMTr-N-protected nucleoside-3′-O-succinyl hemiester in a 25-mL two-neck round-bottom flask containing a magnetic stir bar, with a high-vacuum valve in one joint and a rubber septum in the other, and dry overnight at high vacuum (0.01 mmHg; provided by a high-vacuum oil pump). At the end of drying, deliver dry argon (or nitrogen) to the flask through the septum. 13. Open the flask, quickly add 0.2 g (1 mmol) DCC, and close the flask. 14. Using gas-tight syringes, add 10 mL anhydrous DMF, followed by 0.8 mL anhydrous pyridine. 15. Stir the mixture at room temperature for 24 hr. 16. Filter and wash as in steps 5 to 7. 17. Allow the support to dry at room temperature in a fume hood. Typically, using long-chain alkylamine controlled-pore glass, 80 to 120 mesh, 500 Å (Sigma) a concentration of 25 to 40 ìmol of nucleoside per gram of support is obtained. The support should be stored in a tightly closed vessel in the dark. It is stable for several years at room temperature. IMPORTANT NOTE: The support should be prevented from contact with acidic vapors to avoid detritylation.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Acetonitrile, anhydrous (H2O <20 ppm) Dry acetonitrile over 5 g P2O5 per L for at least 24 hr, then distill under reduced pressure through a 20-cm Vigreux column with exclusion of moisture under an atmosphere of dry argon; at least one-third of the initial volume must remain in the flask. Transfer to reaction vessels using gas-tight syringes under dry argon, or by vacuum line technique (see Support Protocol 1). IMPORTANT NOTE: For condensation steps in Basic Protocol 3, the water content of acetonitrile must be <20 ppm as measured by the Karl Fischer technique. “DNA/RNA synthesis grade” acetonitrile supplied by leading manufacturers is usually not suitable for this purpose unless dried as above.
Low-pressure argon or nitrogen, dried Dry the gas by passing it through a 0.5- to 0.7-m-long column (3- to 4-cm i.d.) filled with blue indicator silica gel (Aldrich cat. no. 336815), then through a similar column filled with granular molecular sieves covered with P2O5. Delivery of inert gas under a slight positive pressure is accomplished by attaching a source of gas via tubing to a T-shaped glass connector. One arm of the connector is attached to the inlet of the first drying column; the other arm is connected to a rubber balloon. After the balloon is inflated up to a diameter of 40 to 50 cm, the valve on the tank regulator is closed, and the internal pressure of the balloon will be sufficient to assure appropriate flow of the gas. The balloon needs to be reinflated from time to time.
Synthesis of Modified Oligonucleotides and Conjugates
4.17.23 Current Protocols in Nucleic Acid Chemistry
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The outlet of the second drying column should be attached through a bubbler (partially filled with mineral oil) to a manifold with 2 to 3 delivery lines, each ending with a valve and a male luer adapter. Typically, argon has lower water content than nitrogen, therefore the drying columns have a longer life-time if argon gas is used.
COMMENTARY Background Information
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
Oligo(nucleoside phosphorothioate)s are congeners of natural oligonucleotides where one of the two nonbridging oxygens in each internucleotide phosphate is replaced by sulfur. By virtue of asymmetry at the phosphorus atom, these constitute a mixture of P-diastereomers, and syntheses of such molecules with predetermined P-chirality at each internucleotide bond are challenging. While the stereocontrolled synthesis of PS-oligos is an art, even more challenging are studies on the properties of such P-stereodefined congeners, mainly with respect to their interactions with other biomolecules such as natural DNA, RNA, and proteins. The affinity of PS-oligos towards DNA and RNA seems to be dictated mainly by base pairing, but is also affected, to a certain extent, by their P-chirality. Notably, intuitive speculations (Zon et al., 1987) or molecular mechanics calculations (Jaroszewski et al., 1992; Hartmann et al., 1999) have led to conclusions contradicting the results of melting experiments performed with heteroduplexes involving P-stereodefined PS-oligos (Boczkowska et al., 2002). This underscores the importance of experimental verification of such hypotheses, which is possible only through access to stereodefined PS-oligos. Physicochemical studies demonstrated that stereoregular PS-oligos of the sequence d(CG)4 of opposite configurations at phosphorus differ significantly in their ability to adopt the Z-conformation in high concentrations of sodium chloride (Boczkowska et al., 2000). Preliminary studies demonstrated that PS-oligos of RP-configuration containing polyadenylate sequences are able to form unusually stable triplexes with two antiparallel complementary RNA strands. The molecular basis of this phenomenon is still unknown, but the role of sulfur in the Rp configuration is essential, as neither [all-SP]-PS-oligomers nor unmodified DNA oligomers are able to form corresponding triplexes of comparable stability (Stec, unpub. observ.) P-stereodefined (PS) oligos have been used for studying the mode of action of several bacterial and human enzymes (Koziolkiewicz
et al., 1997, 2001, 2002). The observation that plasma 3′-exonucleases—a class of proteins responsible for degradation of oligonucleotides in blood—are RP-selective enzymes provided the invaluable information that this enzymatic activity, detrimental toward antisense therapeutics, can be stopped by a single SP-phosphorothioate at the 3′ end of PS-oligonucleotides. Besides this practical aspect, elucidation of the mechanism of the nucleolytic cleavage by 3′-exonuclease(s) provided, for the first time, the information necessary for the classification of this family of proteins (Koziolkiewicz et al., 2002). The interactions of phosphate groups with proteins is well documented. Therefore, the presented method for stereodefined labeling of internucleotide phosphate groups with sulfur and stable oxygen isotopes opens a new avenue for mechanistic studies on DNA/protein interactions at atomic resolution. In this unit, experimental details of the oxathiaphospholane method, developed for stereocontrolled synthesis of oligo(nucleoside phosphorothioate)s, are presented. The method is based upon the synthesis of appropriately protected nucleoside 3′-O-(2-thiono-1,3,2oxathiaphospholane)s and their separation into diastereomerically pure species. It has been demonstrated that DBU-assisted condensation of these monomers with the 5′-OH group of a nucleoside or growing oligonucleotide in the iterative process of chain elongation on a solid support is stereospecific. Stereopurity of [allRP]- and [all-SP]-oligonucleotides has been validated via degradation with stereoselective nucleases such as snake venom phosphodiesterase (Eckstein et al., 1979) and nuclease P1 (Eckstein et al., 1983), respectively. Additional studies have also demonstrated that episulfide, which is released from the ring-opening condensation process, does not modify growing oligonucleotides. One has to realize that this methodology is laborious and costly, and suffers from numerous imperfections which do not allow for the preparation of long PS-oligonucleotides (>15mers). Nonetheless, the oxathiaphospholane
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method has provided many stereodefined PSoligonucleotides suitable for many applications, including NMR studies (Kanehara et al., 1996; Furrer et al., 1999) and biological evaluation in cell cultures, although newcomers to the field may have found it difficult to use. It is clear to the authors of this unit that new modified oxathiaphospholane monomers are highly desirable, as they would not only allow improvement in the repetitive yield of a single condensation step, but would also provide a new and efficient method for protecting internucleotide phosphorothioate diesters against oxidation, necessary for effective combination of the oxathiaphospholane method with phosphoramidite chemistry. Such a combination may provide an effective access to so-called gap-mer or chimeric constructs (Metelev et al., 1994; Pickering et al., 1996; Maier et al., 2000) consisting of oligo(nucleoside phosphate)s and P-stereodefined oligo(nucleoside phosphorothioate) segments. Work on these extensions to the presented method is in progress. Various attempts at stereocontrolled synthesis of PS-oligos undertaken by other research establishments have been reported, among them efforts directed towards stereospecific sy nthesis of dinucleoside 3′,5′-phosphorothioates (Jin et al., 1996; Jin and Just, 1998; Wada et al., 1998; Wang and Just, 1999; Lu and Just, 2000; Oka et al., 2002), Agrawal’s work on the synthesis of nearly stereopure PS-oligos (Iyer et al., 1995, 1998), and the successful stereocontrolled synthesis of longmers reported by Beaucage’s laboratory (Wilk et al., 2000). These groups utilized the ingenious phosphoramidite methodology originally developed by Beaucage and Caruthers (1981). The availability of short P-stereodefined PSoligos (up to pentamers) following separation of diastereomers by means of RP-HPLC should also be noted (Murakami et al., 1994; Tamura, 1998).
neous decomposition of the material inside the vessels. One should frequently monitor the pressure in the system during distillation and keep temperature within the indicated range. For the synthesis of final phosphitylating reagent as well as for phosphitylation of nucleosides, the dryness of solvents and glassware is extremely important, because P(III) chlorides react instantly with traces of water. Like oligonucleotide synthesis via the phosphoramidite or H-phosphonate approach, the dryness of monomers, solvents, and equipment used for coupling is essential for good yields in the oxathiaphospholane approach. Here, the importance of this factor is further stressed because, even with all precautions taken, the repetitive yield of coupling is between 92% and 94%, and any further decrease in coupling yields due to the presence of traces of water may render oligonucleotide synthesis impractical. It is recommended that the cationic DMTr release be quantitated after each condensation step to check if the actual repetitive yield is acceptable.
Anticipated Results Synthesis of phosphitylating reagent, although not trivial, should furnish the consecutive products with reasonable yield (60% to 70%). The reagent is stable and can be stored for months. The same applies to the deoxyribonucleoside 2-thio-oxathiaphospholane monomers. They can be stored in a desiccator at room temperature for several months. Following flash chromatography, the material is recovered at >95% yield, and can then be used for oligonucleotide synthesis. Synthesis of a 15-mer PS-oligo at a 2-µmol scale usually provides 8 to 10 OD units of pure material. This is ∼5% of theoretical yield. It is important that during HPLC purification only the upper part of the oligonucleotide peak be collected (>40% of peak height) to obtain a more homogenous oligomer.
Critical Parameters and Troubleshooting
Time Considerations
Chemical synthesis of the phosphitylating reagent requires some skills to prevent release of the unpleasant odor characteristic of organosulfur compounds. It is recommended that glassware be cleaned using an oxidant, like sodium hypochlorite or hydrogen peroxide, to convert sulfhydryl or disulfide-containing compounds into sulfones. As mentioned in relevant protocols, overheating of the vessels during high-vacuum distillation is dangerous, as it may lead to sponta-
Since neither the oxathiaphosphitylating reagent nor the oxathiaphospholane monomers are commercially available, their preparation is necessary and takes a considerable amount of time. It is reasonable to assume that the synthesis o f 2 -chloro-spiro-4,4-pentamethylene1,3,2-oxathiaphospholane starting from cyclohexanecarboxaldehyde can be accomplished in 10 working days, provided that the technical staff prepares, simultaneously, the necessary anhydrous solvents.
Synthesis of Modified Oligonucleotides and Conjugates
4.17.25 Current Protocols in Nucleic Acid Chemistry
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Phosphitylation/sulfurization of commercially available, appropriately protected nucleosides at 10 mmol scale, as well as preparation of sarcosinylated supports, require 10 additional days. Separation of nucleoside monomers into P-diastereomerically pure species is a very important and rather difficult step. Undoubtedly, some experience is necessary to obtain good results. It should be emphasized that proper loading of the chromatographic column, as well as very careful elution and TLC analysis of effluent, are crucial for yield and diastereomeric purity of the resolved monomers. It must also be taken into account that separation of diastereomers is a time-consuming step, as only ∼300 mg of monomer can be applied on a single silica gel column. The 100 to 120 mg of each pure diastereomer recovered from that amount is sufficient for 3 to 4 coupling steps. To avoid delay resulting from laborious chromatographic purifications, it is good practice to stockpile pure diastereomers, as they are chemically stable and can be stored for long periods of time. Once the monomers are made, the synthesis of PS-oligos proceeds much more rapidly, and typically one oligomer can be made and purified within 4 to 5 days. Usually, during manual synthesis, one can accomplish 8 to 10 couplings daily. If the synthesis is to be continued on the next day, it should be interrupted after the capping step, followed by washing and drying of the support. The synthetic column should then be stored in a desiccator.
Literature Cited Beaucage, S.L. and Caruthers, M.H. 1981. Deoxynucleoside phosphoramidites: A new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Lett. 22:1859-1862. Boczkowska, M., Guga, P., Karwowski, B., and Maciaszek, A. 2000. Effect of P-chirality of internucleotide bonds on B-Z conversion of stereodefined self-complementary phosphorothioate oligonucleotides of [PS]-d(CG)4 and [PS]-d(GC)4 series. Biochemistry 39:1105711064. Boczkowska, M., Guga, P., and Stec, W.J. 2002. Stereodefined phosphorothioate analogues of DNA: Relative thermodynamic stability of model PS-DNA/DNA and PS-DNA/RNA complexes. Biochemistry 41:12483-12487.
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
Brown, T., Pritchard, C.E., Turner, G., and Salisbury, S.A. 1989. A new base-stable linker for solidphase oligonucleotide synthesis. J. Chem. Soc. Chem. Commun. 891-893.
Eckstein, F. 2000. Phosphorothioate oligodeoxynucleotides: What is their origin and what is unique about them? Antisense Nucleic Acid Drug Dev. 10:117-121. Eckstein, F., Burgers, P.M.J., Sathyanarayana, B.K., and Saenger, W. 1979. Crystal and molecular structure of adenosine 5′-O-phosphorothioate OP-nitrophenyl ester (Sp diastereomer). Substrate stereospecificity of snake venom phosphodiesterase. Eur. J. Biochem. 100:585-591. Eckstein, F., Potter, B.V.L., and Connolly, B. 1983. Synthesis and configurational analysis of a dinucleoside phosphate isotopically chiral at phosphorus. Stereochemical course of Penicillium citrum nuclease P1 reaction. Biochemistry 22:1369-1377. Furrer, P., Billeci, T.M., Donati, A., Kojima, C., Karwowski, B., Sierzchala, A., Stec, W.J., and James, T.L. 1999. Structural effect of complete [RP]-phosphorothioate and phosphorodithioate substitution in the DNA strand of a model antisense inhibitor-target RNA complex. J. Mol. Biol. 285:1609-1621. Guga, P., Domanski, K., and Stec, W.J. 2001. Oxathiaphospholane approach to the synthesis of P-chiral, isotopomeric deoxy(ribonucleoside phosphorothioate)s and phosphates labeled with oxygen isotope. Angew. Chem. Int. Ed. Engl. 40:610-613. Hacia, J.G.,Wold, B.J., and Dervan, P.B. 1994. Phosphorothioate oligonucleotide-directed triple helix formation. Biochemistry 33:5367-5369. Hartmann, B., Bertrand, H.-O., and Fermandjian, S. 1999. Sequence effects on energetic and structural properties of phosphorothioate DNA: A molecular modelling study. Nucl. Acids Res. 27:3342-3347. Iyer, R.P., Yu, D., Ho, N.-H., Tan, W., and Agrawal, S. 1995. A novel nucleoside phosphoramidite synthon derived from 1R,2S-ephedrine. Tetrahedron: Asymmetry 6:1051-1054. Iyer, R.P., Guo, M.J., Yu, D., and Agrawal, S. 1998. Solid-phase stereoselective synthesis of oligonucleoside phosphorothioates: The nucleoside bicyclic oxazaphospholidines as novel synthons. Tetrahedron Lett. 39:2491-2494. Jaroszewski, J.W., Syi, J.-L., Maizel, J., and Cohen, J.S. 1992. Towards rational design of antisense DNA: Molecular modeling of phosphorothioate DNA analogues. Anticancer Drug Des. 7:253262. Jin, Y. and Just, G. 1998. Stereoselective synthesis of dithymidine phosphorothioates using xylose derivatives as chiral auxiliaries. J. Org. Chem. 63:3647-3654. Jin, Y., Biancotto, G., and Just, G. 1996. A stereoselective synthesis of dinucleotide phosphorothioates using chiral phosphoramidites as intermediates. Tetrahedron Lett. 37:973-976.
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Kanehara, H., Wada, T., Mizuguachi, M., and Makino, K. 1996. Influence of a thiophosphate linkage on the duplex stability: Does Sp configuration always lead to higher stability than Rp? Nucleosides Nucleotides 15:1169-1178.
Oka, N., Wada, T., and Saigo, K. 2002. Diastereocontrolled synthesis of dinucleoside phosphorothioates using a novel class of activators, dialkyl(cyanomethyl)ammonium tetrafluoroborates. J. Am. Chem. Soc. 124:4962-4963.
Koziolkiewicz, M., Wójcik, M., Kobylanska, A., Karwowski, B., Rebowska, B., Guga, P., and Stec, W.J. 1997. Stability of stereoregular oligo(nucleoside phosphorothioate)s in human plasma: Diastereoselectivity of plasma 3′-exonuclease. Antisense Nucleic Acids Drug Dev. 7:43-48.
Pickering, J.G., Isner, J.M., Ford, C.M., Weir, L., Lazarovits, A., Rocnik, E.F., and Chow, L.H. 1996. Processing of chimeric antisense oligonucleotides by human vascular smooth muscle cells and human atherosclerotic plaque. Implications for antisense therapy of restenosis after angioplasty. Circulation 93:772-780.
Koziolkiewicz, M., Owczarek, A., Domanski, K., Nowak, M., Guga, P., and Stec, W.J. 2001. Stereochemistry of cleavage of internucleotide bond by Serratia marcescens endonuclease. Bioorg. Med. Chem. 9:2403-2409.
Stec, W.J., Grajkowski, A., Koziolkiewicz, M., and Uznanski, B. 1991. Novel route to oligo(deoxyribonucleoside phosphorothioates): Stereocontrolled synthesis of P-chiral oligo(deoxyribonucleoside phosphorothioates). Nucl. Acids Res. 19:5883-5888.
Koziolkiewicz, M., Owczarek, A., Wójcik, M., Domanski, K., Guga, P., and Stec, W.J. 2002. Retention of the configuration in the action of human plasma 3′-exonuclease on oligo(deoxynucleoside phosphorothioate): A new method for assignment of absolute configuration at phosphorus in isotopomeric deoxyadenosine 5′-O[18O]phosphorothioate. J. Am. Chem. Soc. 124:4623-4627. Lackey, D.B. and Patel, J. 1997. Biochemical synthesis of chirally pure RP oligonucleotide phosphorothioates. Biotechnol. Lett. 19:475-478. Lu, Y. and Just, G. 2000. Stereoselective synthesis of RP- and SP-dithymidine phosphorothioates via chiral indolooxazaphosphorine intermediates derived from tryptophan. Angew. Chem. Int. Ed. Engl. 39:4521-4524. Maier, M.A., Guzaev, A.P., and Manoharan, M. 2000. Synthesis of chimeric oligonucleotides containing phosphodiester, phosphorothioate, and phosphoramidate linkages. Org. Lett. 2:1819-1822. Manoharan, M. 1999. 2′-Carbohydrate modifications in antisense oligonucleotide therapy: Importance of conformation, configuration and conjugation. Biochim. Biophys. Acta 1489:117130. Martynov, I.V., Kruglyak, Y.L., Leibovskaya, G.A., Khromova, Z.I., and Stukov, O.G. 1969. Phosphonylated oximes: IV. Reactions of 1,3,2-dioxa- and 1,3,2-oxathiaphospholanes with αhalogen nitroso alkanes. Zh. Obsch. Khim 39:996. Metelev, V., Lisziewicz, J., and Agrawal, S. 1994. Study of antisense oligonucleotide phosphorothioates containing segments of oligodeoxynucleotides and 2′-O-methyloligoribonucleotides. Bioorg. Med. Chem. Lett. 4:29292934. Murakami, A., Tamura, Y., Wada, H., and Makino, K. 1994. Separation and characterization of diastereoisomers of antisense oligodeoxyribonucleoside phosphorothioates. Anal. Biochem. 233:285-290.
Stec, W.J., Grajkowski, A., Karwowski, B., Kobylanska, A., Koziolkiewicz, M., Misiura, K., Okruszek, A., Wilk, A., Guga, P., and Boczkowska, M. 1995. Diastereomers of nucleoside 3′-O-(2thio-1,3,2-oxathia(selena)phospholanes): Building blocks for stereocontrolled synthesis of oligo(nucleoside phosphorothioate)s. J. Am. Chem. Soc. 117:12019-12029. Stec, W.J., Karwowski, B., Boczkowska, M., Guga, P., Koziolkiewicz, M., Sochacki, M., Wieczorek, M., and Blaszczyk, J. 1998. Deoxyribonucleoside 3′-O-(2-thio- and 2-oxo-spiro-4,4-pentamethylene-1,3,2-oxathiaphospholane)s: Monomers for stereocontrolled synthesis of oligo(deoxyribonucleoside phosphorothioate)s and chimeric PS/PO oligonucleotides. J. Am. Chem. Soc. 120:7156-7167. Stein, C.A. and Krieg, A.M. (eds.) 1988. Applied Antisense Oligonucleotide Technology. WileyLiss, New York. Tamura, Y., Miyoshi, L.H., Yokota, T., and Makino, K. 1998. Preparation of stereoregulated antis ens e oligode oxyribonucleoside phosphorothioate and interaction with its complementary DNA and RNA. Nucleosides Nucleotides 17:269-282. Tang, J., Roskey, A., Li, Y., and Agrawal, S. 1995. Enzymatic synthesis of stereoregular [all RP]oligonucleotide phosphorothioate and its properties. Nucleosides Nucleotides 14:985-990. Thuong, N.T. and Helene, C. 1993. Sequence-specific recognition and modification of doublehelical DNA by oligonucleotides. Angew. Chem. Int. Ed. Engl. 32:666-690. Wada, T., Kobayashi, N., Mori, T., and Sekine, M. 1998. Stereocontrolled synthesis of dithymidine phosphorothioates by use of a functionalized 5′-protecting group bearing an imidazole residue. Nucleosides Nucleotides 17:351-364. Wang, J.C. and Just, G. 1999. Indol-oxazaphosphorine precursors for stereoselective synthesis of dinucleotide phosphorothioates. J. Org. Chem. 64:8090-8097. Wickstrom, E. 1986. Oligodeoxynucleotide stability in subcellular extracts and culture media. J. Biochem. Biophys. Methods 13:97-102.
Synthesis of Modified Oligonucleotides and Conjugates
4.17.27 Current Protocols in Nucleic Acid Chemistry
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Wilk, A. and Stec, W.J. 1995. Analysis of oligo(deoxynucleoside phosphorothioate)s and their diastereomeric composition. Nucl. Acids Res. 23:530-534. Wilk, A., Grajkowski, A., Phillips, L.R., and Beaucage, S.L. 2000. Deoxyribonucleoside cyclic Nacylphosphoramidites as a new class of monomers for the stereocontrolled synthesis of oligothymidylyl- and oligodeoxycytidylylphosphorothioates. J. Am. Chem. Soc. 122:21492156. Willson, M., Goncalves, H., Boudjebel, H., and Burgada, R. 1975. Étude du méchanisme des réactions d’acidalyse, d’alcoolyse et d’aminolyse des oxathiaphosphalanes-1,3,2. Bull. Soc. Chim. Fr. Part 2, 615-620.
Zon, G., Summers, M.F., Gallo, K.A., Shao, K.-L., Koziolkiewicz, M., Uznanski, B., and Stec, W.J. 1987. Stereochemistry of oligodeoxyribonucleotide phosphate triesters. In Biophosphates and Their Analogues: Synthesis, Structure, Metabolism and Activity (K.S. Bruzik and W.J. Stec, eds.) pp. 165-178. Elsevier, Amsterdam.
Contributed by Piotr Guga and Wojciech J. Stec Polish Academy of Sciences Lodz, Poland
Synthesis of Phosphorothioate Oligonucleotides with Stereodefined Linkages
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Synthesis of Oligonucleotide Conjugates via Aqueous Diels-Alder Cycloaddition
UNIT 4.18
This unit describes in detail the preparation of 5′-labeled oligonucleotides via the aqueous Diels-Alder conjugation method. The unit is divided into three protocols: the first describing the preparation of the diene-amidite (see Basic Protocol 1); the second presenting the modification of oligonucleotides with the diene-amidite (see Basic Protocol 2); and the third describing the labeling of 5′-diene-modified oligonucleotides via Diels-Alder conjugation with TAMRA-5-maleimide (see Basic Protocol 3). All procedures are simple to perform with standard equipment and reagents. CAUTION: All reactions must be performed in a well-ventilated fume hood. The use of proper laboratory precautions when handling all chemicals is mandatory. In particular, exposure of personnel to tert-butyldimethylsilyl chloride, bromine, 1,1′-carbonyldiimidazole, 2-cyanoethyl diisopropylchlorophosphoramidite, and N,N-diisopropylethylamine must be avoided. SYNTHESIS OF THE DIENE-AMIDITE The five-step synthesis route for the diene-amidite (S.7) employs commercially available 3-cyclohexene-1-methanol (S.1) as the starting material (Fig. 4.18.1). In the first step, the hydroxyl group of S.1 is protected by silylation with tert-butyldimethylsilyl chloride (TBDMS⋅Cl) to give the alkene S.2. In the next step, bromine is added. The resulting dibromide (S.3) is subjected to a bis-dehydrohalogenation reaction with potassium tert-butoxide to form the diene S.4. Removal of the TBDMS protecting group with strongly acidic Dowex resin and subsequent distillation of the crude product leads to pure 2,4-cyclohexadiene-1-methanol (S.5). Conjugation of S.5 to 6-amino-1-hexanol is accomplished in reaction with 1,1′-carbonyldiimidazole (CDI). The crude carbamate S.6 is purified by crystallization. Finally, S.6 is converted into the phosphoramidite S.7 by treatment with 2-cyanoethyl diisopropylchlorophosphoramidite and N,N-diisopropylethylamine (DIPEA). Purification of the crude product is performed by flash chromatography. Materials 3-Cyclohexene-1-methanol (S.1; Aldrich) Imidazole Argon source N,N-Dimethylformamide (DMF) tert-Butyldimethylsilyl chloride (TBDMS⋅Cl; Aldrich) Hexanes Ethyl acetate (EtOAc) TLC stain (see recipe) Brine (saturated aqueous NaCl) Magnesium sulfate (MgSO4) Bromine Dichloromethane (CH2Cl2) 10% (w/v) sodium thiosulfate (Na2S2O3) Aliquat 336 (tricaprylylmethylammonium chloride; Aldrich) Tetrahydrofuran (THF) Potassium tert-butoxide (Aldrich) Saturated ammonium chloride (NH4Cl) Dowex 50WX4-50 strongly acidic ion-exchange resin (Aldrich) Contributed by Michael Leuck and Andreas Wolter Current Protocols in Nucleic Acid Chemistry (2003) 4.18.1-4.18.14 Copyright © 2003 by John Wiley & Sons, Inc.
BASIC PROTOCOL 1
Synthesis of Modified Oligonucleotides and Conjugates
4.18.1 Supplement 14
TBDMS⋅Cl/imidazole
OH
Br2, CH2Cl2
OTBDMS
1 hr, 96%
DMF, 20 hr, quant. 1
2
OTBDMS
KOt Bu/Aliquat 336
OTBDMS
THF, 2 hr
Br Br
4 3
O
Dowex-H +/MeOH
CDI
OH
2 hr, 84% (2 steps)
O
DMF, 1 hr
N
N
5
O 6-amino-1-hexanol, 16 hr
O
OH
N H
50%
6
O
i -Pr2NP(Cl)OCH2CH2CN/DIPEA THF, 1 hr, 75%
O
O
N H
P
O
CN
i-Pr2N 7
Figure 4.18.1 Preparation of diene-amidite (S.7). Abbreviations: Aliquat 336, tricaprylylmethylammonium chloride; CDI, 1,1′-carbonyldiimidazole; DIPEA, N,N-diisopropylethylamine; TBDMS⋅Cl, tert-butyldimethylsilyl chloride.
Methanol 1,1′-Carbonyldiimidazole (CDI; Aldrich) 6-Amino-1-hexanol (Aldrich) N,N-Diisopropylethylamine (DIPEA; Aldrich) 2-Cyanoethyl diisopropylchlorophosphoramidite (Aldrich) Saturated sodium bicarbonate (NaHCO3)
Synthesis of Oligonucleotide Conjugates via Diels-Alder Cycloaddition
100- and 250-mL round-bottom flasks with outlet stopcock adapters for gas line TLC plates: 7.5 × 5–mm silica gel 60 F254 precoated aluminum-backed TLC sheets (EM Science) Heat gun 1-L and 500-mL separatory funnels Rotary evaporator equipped with a vacuum pump and vacuum controller Rubber septa
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Current Protocols in Nucleic Acid Chemistry
10- and 50-mL vials 3- and 5-mL syringes with 2-in., 20-G stainless steel needles Shaker Short-path distillation apparatus (Aldrich) Biotage Flash 40 chromatography system with Flash 40M silica cartridge Additional reagents and equipment for TLC (APPENDIX 3D) and flash chromatography (APPENDIX 3E) Perform silylation 1. Weigh 2.78 g (24.8 mmol) of 3-cyclohexene-1-methanol (S.1) and 3.38 g (49.6 mmol) of imidazole into a 100-mL round-bottom flask equipped with a magnetic stir bar and an argon inlet. Apply an atmosphere of argon. 2. Add 25 mL of DMF and stir until all solids are dissolved. 3. Add 4.49 g (29.8 mmol) TBDMS⋅Cl to the reaction mixture and stir for 20 hr at room temperature. Monitor reaction by TLC (APPENDIX 3D) using 100:1 (v/v) hexanes/ethyl acetate. Visualize spots by treating plate with TLC stain and heating with a heat gun. TLC (hexanes/EtOAc 100:1): Rf = 0.05 for starting material (S.1); 0.29 for TBDMS-ether (S.2).
Perform aqueous workup of S.2 4. Pour the reaction mixture into a 1-L separatory funnel. Add 250 mL of brine and extract the product with 150 mL of hexanes. 5. Repeat the extraction twice with 150 mL each of hexanes. 6. Combine the hexanes fractions and wash with 250 mL of brine. 7. After separation dry the organic phase over MgSO4. 8. Remove the MgSO4 by filtration and wash it twice, each time with 20 mL of hexanes. 9. Concentrate the combined filtrates under vacuum using a rotary evaporator equipped with a vacuum pump. Typical yield of 4-(tert-butyldimethylsilyloxymethyl)cyclohex-1-ene (S.2): 5.62 g (100%) as colorless liquid. TLC (hexanes): Rf = 0.29; 13C NMR (CDCl3): δ 127.0, 126.2, 68.0, 36.3, 28.2, 26.0, 25.7, 25.3, 24.8, 18.4, –5.3.
Brominate alkene 10. Weigh 5.62 g (24.8 mmol) of S.2 into a 250-mL round-bottom flask equipped with a magnetic stir bar. 11. Add 50 mL of CH2Cl2, seal the flask with a rubber septum, and apply an atmosphere of argon. Dissolve the silyl ether (S.2) under stirring, then cool the solution with an ice bath to 0°C. 12. In a separate 10-mL vial, dissolve 1.3 mL (25.2 mmol) of bromine in 3 mL of CH2Cl2. Fill a 5-mL syringe with the bromine/CH2Cl2 solution. 13. Attach the syringe to the flask (step 11) using a 20-G stainless steel needle to pierce the septum, and add the bromine solution dropwise under cooling. At the beginning, the red color disappears immediately after addition. Finally, an orangered color remains.
14. After complete addition of the bromine solution, remove the ice bath and let the mixture stir for another 60 min. Monitor by TLC (see step 3). TLC (hexanes/EtOAc 100:1): Rf = 0.29 for TBDMS-ether (S.2); 0.15 for dibromide (S.3).
Synthesis of Modified Oligonucleotides and Conjugates
4.18.3 Current Protocols in Nucleic Acid Chemistry
Supplement 14
Perform aqueous workup of S.3 15. Pour the reaction mixture into a 500-mL separatory funnel. 16. Rinse the flask with 150 mL of CH2Cl2 and add to the separatory funnel. 17. Wash the CH2Cl2 layer with 150 mL of 10% Na2S2O3, then with 150 mL of water. 18. Dry the organic phase over MgSO4. 19. Remove the MgSO4 by filtration and wash twice with 20 mL of CH2Cl2. 20. Concentrate the combined filtrates under vacuum in a rotary evaporator. Typical yield of trans-1,2-dibromo-4-tert-butyldimethylsilyloxymethylcyclohexane (S.3): 9.19 g (96%) as yellow oil. TLC (hexanes/EtOAc 100:1): Rf = 0.15. 13C NMR (CDCl3): δ 67.3, 53.6, 53.5, 34.2, 31.41, 3.12, 3.88, 2.25, 2.3, –5.4.
Perform bis-dehydrohalogenation 21. Weigh 9.19 g (23.8 mmol) of S.3 and 0.20 g (0.5 mmol) of Aliquat 336 into a 250-mL round-bottom flask equipped with a magnetic stir bar. 22. Add 70 mL of THF, seal the flask with an outlet stopcock adapter attached to an argon line, and apply an atmosphere of argon. Dissolve the dibromide (S.3) under stirring, then cool the solution with an ice bath to 0°C. 23. Open the reaction flask by removing the adapter, add 5.88 g (52.4 mmol) of potassium tert-butoxide in one portion, seal the flask again with the adapter, and apply an atmosphere of argon (a yellow precipitate is immediately formed). Remove the ice bath after 5 min and continue stirring for 2 hr. Monitor by TLC. TLC (hexanes/EtOAc 100:1): Rf = 0.15 for dibromide (S.3); 0.25 for diene (S.4).
Perform aqueous workup of S.4 24. Pour the reaction mixture into a 500-mL separatory funnel. 25. Rinse the flask with 100 mL of hexanes and add to the separatory funnel. 26. Wash the hexanes layer with 100 mL saturated NH4Cl. 27. After separation, extract the aqueous phase with 50 mL of hexanes and wash the combined hexanes phases with 100 mL of water. 28. Dry the organic phase over MgSO4. 29. Remove the MgSO4 by filtration and wash it twice, each time with 20 mL of hexanes. 30. Concentrate the combined filtrates under vacuum at 40°C, with pressure ≥180 mbar, in a rotary evaporator. Perform desilylation 31. Weigh 8.60 g of Dowex 50WX4-50 into a 50-mL vial and wash it carefully three times, each time with 20 mL methanol. After the third washing, the supernatant should be colorless.
32. Dissolve the S.4 residue from step 30 in 80 mL methanol, transfer it into a 250-mL round-bottom flask, and add the washed Dowex 50WX4-50 resin. 33. Attach the flask to a shaker and shake the reaction mixture for 2 hr at 200 rpm. Monitor by TLC using 3:1 (v/v) hexanes/ethyl acetate. Synthesis of Oligonucleotide Conjugates via Diels-Alder Cycloaddition
TLC (hexanes/EtOAc 3:1): Rf = 0.80 for diene (S.4); 0.20 for diene-alcohol (S.5).
4.18.4 Supplement 14
Current Protocols in Nucleic Acid Chemistry
Purify diene-alcohol S.5 34. Remove the ion-exchange resin by filtration and wash it twice with 20 mL methanol. 35. Concentrate the combined filtrates under vacuum at 40°C, with pressure ≥180 mbar, in a rotary evaporator. 36. Transfer the residue into a round-bottom flask of appropriate size (5 or 10 mL) and attach it to a short-path distillation apparatus. Distill the product under vacuum, collecting the fraction ranging from b.p. 47°C (2.2 mbar) to 50°C (1.7 mbar). Typical yield of 2,4-cyclohexadiene-1-methanol (S.5): 2.20 g (84%) as colorless liquid. TLC (hexanes/EtOAc 3:1): Rf = 0.20. 1H NMR (CDCl3): δ 5.98 - 5.62 (m, 4 H), 3.58 (d, J = 5.9 Hz, 2 H), 2.56 - 2.42 (m, 1 H), 2.37 - 2.05 (m, 2 H), 1.50 (s, 1 H). 13C NMR (CDCl3): δ 126.9, 125.4, 125.0, 123.6, 64.6, 35.5, 24.9.
Conjugate diene-alcohol to 6-amino-1-hexanol 37. Weigh 2.20 g (20.0 mmol) of S.5 into a 100-mL round-bottom flask equipped with a magnetic stir bar. 38. Add 20 mL of DMF, seal the flask with an outlet stopcock adapter attached to an argon line, and apply an atmosphere of argon. Dissolve the diene (S.5) under stirring. 39. Open the reaction flask by removing the adapter, add 3.40 g (21.0 mmol) of CDI in one portion, seal the flask again with the adapter, and apply an atmosphere of argon. Continue stirring for 1 hr. Monitor by TLC using 1:1 (v/v) hexanes/ethyl acetate. TLC (hexanes/EtOAc 1:1): Rf = 0.32 for alcohol (S.5); 0.20 for the imidazolyl carbamate.
40. Open the reaction flask by removing the adapter, add 2.34 g (20.0 mmol) of 6-amino-1-hexanol in one portion, seal the flask again with the adapter, and apply an atmosphere of argon. Continue stirring overnight. Monitor by TLC using 1:2 (v/v) hexanes/ethyl acetate. TLC (hexanes/EtOAc 1:2): Rf = 0.56 for the imidazolyl carbamate; 0.44 for carbamate (S.6).
Perform aqueous workup of carbamate S.6 41. Concentrate the reaction mixture under vacuum in a rotary evaporator. 42. Dissolve the residue in 100 mL of CH2Cl2 and transfer the solution into a 250-mL separatory funnel. 43. Wash the CH2Cl2 layer twice, each time with 70 mL of water. 44. After separation, dry the organic phase over MgSO4. 45. Remove the MgSO4 by filtration and wash it twice, each time with 10 mL of CH2Cl2. 46. Concentrate the combined filtrates under vacuum in a rotary evaporator. 47. Recrystallize the crude product from CH2Cl2. Typical yield of 6-(cyclohexa-1,3-dien-5-yl)methylcarbamoylhexan-1-ol (S.6): 2.53 g (50%) as colorless crystals, m.p. 65°C. TLC (hexanes/EtOAc 1:2): Rf = 0.44. 1H NMR (300 MHz, CD3CN): δ 6.01-5.96 (m, 1H), 5.94-5.89 (m, 1H), 5.86-5.78 (m, 1H), 5.72-5.67 (m, 1H), 5.51 (bs, 1H), 3.96-3.89 (m, 2H), 3.49 (q, J = 5.6 Hz, 2H), 3.08 (q, J = 6.6 Hz, 2H), 2.68-2.50 (m, 1H), 2.49 (t, J = 5.6 Hz, 1H), 2.31-2.21 (m, 1H), 2.15-2.02 (m, 1H), 1.54-1.45 (m, 4H), 1.40-1.26 (m, 4H). 13C NMR (75 MHz, CD3 CN): δ 156.8, 127.0, 126.0, 125.6, 125.0, 61.7, 61.5, 40.7, 33.0, 32.8, 29.9, 26.5, 25.5, 25.0. MS (FAB): m/z 254 [M+H]+. Synthesis of Modified Oligonucleotides and Conjugates
4.18.5 Current Protocols in Nucleic Acid Chemistry
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Perform phosphitylation 48. Weigh 2.53 g (10.0 mmol) of S.6 into an oven-dried 100-mL round-bottom flask equipped with a magnetic stir bar. 49. Add 25 mL of dry THF and 2.6 mL (15 mmol) of DIPEA, seal the flask with a septum, and apply an atmosphere of argon. Dissolve the alcohol (S.6) under stirring. 50. Weigh 2.49 g (10.5 mmol) of 2-cyanoethyl diisopropylchlorophosphoramidite into a 3-mL syringe. 51. Attach the syringe to the flask via the septum and add the chlorophosphoramidite dropwise. Rinse the syringe with 3 mL of dry THF and add this solution to the reaction mixture. Continue stirring for 1 hr. Monitor by TLC using ethyl acetate. TLC (EtOAc): Rf = 0.50 for carbamate (S.6); 0.83 for amidite (S.7).
Perform aqueous work-up and purification of S.7 52. Pour the reaction mixture into a 250-mL separatory funnel. 53. Add 75 mL of CH2Cl2 and wash the organic layer three times with 75 mL saturated NaHCO3. 54. Dry the organic phase over MgSO4. 55. Remove the MgSO4 by filtration and wash it twice with 10 mL CH2Cl2. 56. Concentrate the combined filtrates under vacuum in a rotary evaporator. 57. Dissolve the residue in 4 mL ethyl acetate and inject it on a Biotage Flash 40 chromatography system employing a Biotage Flash 40M silica gel cartridge equilibrated with ethyl acetate. Elute the product with ethyl acetate. 58. Concentrate the product-containing fractions under vacuum in a rotary evaporator. Dry the residue under high vacuum. Typical yield of 1-[(2-cyanoethoxy)diisopropylaminophosphinyloxy]-6-(cyclohexa-1,3dien-5-yl)methylcarbamoylhexane (S.7): 3.40 g (75%) as colorless oil. TLC (EtOAc): Rf = 0.83. 1H NMR (CD3CN): δ 6.01-5.87 (m, 2H), 5.83-5.77 (m, 1H), 5.71 (m, 1H), 5.52 (bs, 1H), 3.97-3.89 (m, 2H), 3.89-3.70 (m, 2H), 3.70-3.56 (m, 4H), 3.07 (q, J = 6.6 Hz, 2H), 2.70-2.53 (m, 1 H), 2.66 (t, J = 5.9 Hz, 2H), 2.31-2.15 (m, 1H), 2.15-2.01 (m, 1H), 1.64-1.26 (m, 8H), 1.20 (s, 3H), 1.19, (s, 3H), 1.18 (s, 3H), 1.17 (s, 3H). 13C NMR (CD3 CN): δ 156.7, 127.0, 125.8, 125.3, 124.0, 118.9, 65.7, 63.6, 63.4, 58.6, 58.4, 43.1, 42.9, 40.7, 33.2, 31.2, 31.1, 29.9, 26.3, 25.6, 25.0, 24.2, 24.1, 20.4, 20.3. 31P NMR (CD3 CN): δ 148.0. MS (FAB): m/z 455 [M+H]+. Anal. calcd. for C23H40N3O4P: C, 60.91; H, 8.89; N, 9.26; found: C, 60.55; H, 9.23; N, 9.10. RP-HPLC: Rt = 12.67 min; Waters Nova-Pak C18 column (4 ìm, 3.9 × 150 mm) eluting with a linear gradient from 40% to 100% A (solvent A, acetonitrile; solvent B, 250 mM triethylammonium acetate, pH 6.5) in 15 min with a flow rate of 1 mL/min; UV detection performed at λ = 254 nm.
Synthesis of Oligonucleotide Conjugates via Diels-Alder Cycloaddition
4.18.6 Supplement 14
Current Protocols in Nucleic Acid Chemistry
CPG
N1
n −1 standard synthesis cycles
standard synthesis cycle
N1.....Nn
O CPG
diene-amidite
N1.....Nn
P O
O
cleavage/deprotection conc. ammonia
CPG
N-amidite
3′ N1.....Nn
P
O −
O
+
H N
O
O O
CN
H N
O O
NH4
Figure 4.18.2 Synthesis of diene-modified oligonucleotides. Standard synthesis cycle: deblocking (deblock solution); coupling (N-amidite/DCI); capping (Cap A and B solutions); oxidation (oxidizer solution). N-amidite: standard dT, dC(bz), dA(bz), dG(ib) phosphoramidites.
SYNTHESIS OF 5′-DIENE-MODIFIED OLIGONUCLEOTIDES This protocol describes the setup and steps required for automated assembly of 5′-dienemodified oligonucleotides on a 1-µmol scale. The protocol was developed for the Expedite 8909 DNA synthesizer, but can easily be adapted to other automated DNA synthesizers. The reactions carried out in each cycle are shown in Figure 4.18.2. For further details on automated oligonucleotide synthesis, see APPENDIX 3C.
BASIC PROTOCOL 2
The instrument manufacturer’s synthesis protocol in DMTr-ON mode is applied for the assembly of the oligonucleotide with standard reagents for solid-phase oligonucleotide synthesis. The diene-amidite (S.7) is employed at a concentration of 50 mg/mL in acetonitrile in the last coupling cycle. After completion of the synthesis, the oligonucleotide is cleaved from the CPG support and subsequently deprotected by treatment with concentrated ammonium hydroxide. Desalting of the crude oligonucleotide is accomplished by alcohol precipitation. Materials Dry acetonitrile (DNA synthesis grade; H2O content <30 µg/mL) Deblock solution: trichloroacetic acid (TCA) in dichloromethane (Proligo) Cap A solution: acetic anhydride in tetrahydrofuran (Proligo) Cap B solution: 1-methylimidazole in tetrahydrofuran/pyridine (Proligo) Oxidizer solution: iodine in tetrahydrofuran/water/pyridine (Proligo) Activator: 0.25 M 4,5-dicyanoimidazole (DCI) in acetonitrile (Proligo) 50 mg/mL standard phosphoramidites [dT, dC(bz), dA(bz), and dG(ib); Proligo] in dry acetonitrile 50 mg/mL diene-amidite (S.7; see Basic Protocol 1) in dry acetonitrile Helium source Concentrated ammonium hydroxide (NH4OH) 3 M sodium acetate (APPENDIX 2A) Isopropanol 70% (v/v) ethanol
Synthesis of Modified Oligonucleotides and Conjugates
4.18.7 Current Protocols in Nucleic Acid Chemistry
Supplement 15
Automated DNA synthesizer (e.g., Expedite 8909, PerSeptive Biosystems) Synthesis column for 1-µmol scale (Proligo): controlled-pore glass (CPG; 500 Å for <50-mers, 1000 Å for ≥50-mers) Heating block Speedvac evaporator Additional reagents and equipment for quantitating oligonucleotides by OD260 measurement (UNIT 10.3) Perform automated synthesis 1. Install the bottles with all reagents (acetonitrile, deblock solution, cap A and B solutions, oxidizer solution, activator solution), phosphoramidite solutions [dT, dC(bz), dA(bz), and dG(ib)], and diene-amidite S.7 in an automated DNA sythesizer according to the manufacturer’s instructions (also see APPENDIX 3C). Flush with helium and prime the fluidics system three times. 2. Use the sequence editor to enter a new sequence or edit an existing sequence for synthesis. 3. Load the appropriate synthesis column for 1-µmol scale. 4. Choose the DMTr-ON option in the setup menu and perform the synthesis. The standard synthesis cycle is applied in the last cycle for the diene-amidite (S.7) coupling. Cycle steps are listed in Table 4.18.1.
5. Remove the synthesis column from the instrument. 6. Cleave the oligonucleotide from the solid support in the column by treatment with 1 mL concentrated ammonium hydroxide for 30 min at room temperature. 7. Collect the ammonia solution containing the oligonucleotide into a screw-cap tube and deprotect the oligonucleotide by heating in a heating block at 55°C for 8 hr or overnight. 8. Dry the deprotected oligonucleotide in a Speedvac evaporator. Desalt oligonucleotides 9. Dissolve ∼100 OD260 units of crude oligonucleotide in 100 µL of deionized water. 10. Add 11 µL of 3 M sodium acetate solution and mix. 11. Add 333 µL of isopropanol, then vortex. 12. Incubate overnight at room temperature. 13. Microcentrifuge 10 min at maximum speed (≥10,000 × g), room temperature. 14. Remove the supernatant and discard; avoid disturbing the pellet. 15. Add 300 µL of 70% ethanol, mix briefly, and place in a freezer at −20°C for 2 hr. 16. Microcentrifuge 5 min at maximum speed (≥10,000 × g), room temperature. 17. Remove the supernatant and discard. 18. Evaporate pellet to dryness in a Speedvac evaporator. 19. Quantify the oligonucleotide by OD260 measurement (UNIT 10.3). Synthesis of Oligonucleotide Conjugates via Diels-Alder Cycloaddition
4.18.8 Supplement 15
Current Protocols in Nucleic Acid Chemistry
Table 4.18.1 Standard Expedite 1-µmol DNA Synthesis Protocol Applied for Synthesis of Diene-Modified Oligonucleotides using Diene-Amidite (S.7).
Operation
Function
Mode
Amount
Time (sec) Description
Index Fract. Coll. Default Trityl Mon. On/Off Dblk Dblk Diverted Wsh A Trityl Mon. On/Off Diverted Wsh A Index Fract. Coll.
NA
1
0
Event out ON
WAIT NA
0 1
1.5 1
PULSE PULSE PULSE
10 50 40
0 49 0
NA
0
1
Wait START data collection Dblk to column Deblock Flush system with Wsh A STOP data collection
PULSE
40
0
NA
2
0
Deblocking 144 0 141 16 16 38 141 38 144
Flush system with Wsh A Event out OFF
Coupling 1
Wsh
PULSE
5
0
Flush system with Wsh Flush system with Act Monomer + Act to column Couple monomer Couple monomer Couple monomer Flush system with Wsh
2
Act
PULSE
5
0
22
S.7 + Act
PULSE
6
0
22 2 1 1
S.7 + Act Act Wsh Wsh
PULSE PULSE PULSE PULSE
1 4 7 8
8 32 56 0
12
Wsh A
PULSE
20
0
13 12 12
Caps Wsh A Wsh A
PULSE PULSE PULSE
8 6 14
0 15 0
Ox Wsh A
PULSE PULSE
15 15
0 0
Ox to column Flush system with Wsh A
Caps Wsh A
PULSE PULSE
7 30
0 0
Caps to column End of cycle wash
Capping Flush system with Wsh A Caps to column Cap Flush system with Wsh A
Oxidizing 15 12 Capping 13 12
aNA, not applicable.
Synthesis of Modified Oligonucleotides and Conjugates
4.18.9 Current Protocols in Nucleic Acid Chemistry
Supplement 14
BASIC PROTOCOL 3
LABELING OF A 5′-DIENE-MODIFIED OLIGONUCLEOTIDE WITH TAMRA-5-MALEIMIDE This protocol describes the conjugation of TAMRA-5-maleimide to a diene-modified oligonucleotide via aqueous Diels-Alder cycloaddition (Fig. 4.18.3). The crude 5′-dienemodified oligonucleotide is dissolved in sodium acetate buffer, pH 4.5, and treated with a 10-fold excess of TAMRA-5-maleimide. The mixture is heated at 60°C for 4 hr. Excess dye is removed by size-exclusion chromatography employing NAP-5 columns. The labeled oligonucleotides obtained can be purified by high-performance liquid chromatography (HPLC; UNIT 10.5) or polyacrylamide gel electrophoresis (PAGE; UNIT 10.4). Materials Diene-modified oligonucleotide (see Basic Protocol 2) 100 mM sodium acetate buffer, pH 4.5 (APPENDIX 2A) 40 mM TAMRA-5-maleimide solution: 5.0 mg TAMRA-5-maleimide (Molecular Probes) in 268 µL dimethylformamide (DMF) Heating block NAP-5 columns (Amersham Pharmacia Biotech) Speedvac evaporator Additional reagents and equipment for quantitating oligonucleotides by OD260 measurement (UNIT 10.3) Label diene-modified oligonucleotide 1. Prepare a 1 mM stock solution of the diene-modified oligonucleotide in deionized water. Calculations are based on the OD260 readings obtained (see Basic Protocol 2, step 19) and the calculated extinction coefficient of the oligonucleotide (UNIT 10.3).
O O 3′ N1.....Nn P
−
H N
O
+
O
N
O
O
NaOAc buffer pH 4.5
TAMRA
O
4 hr, 60 °C
O O 3′ N1.....Nn
P
−
O
Synthesis of Oligonucleotide Conjugates via Diels-Alder Cycloaddition
O
H N
O O
N
TAMRA
O
Figure 4.18.3 Labeling of diene-modified oligonucleotide with TAMRA-5-maleimide via aqueous Diels-Alder cycloaddition.
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2. In a 1.5-mL microcentrifuge tube, prepare a mixture of: 20 µL 1 mM diene-modified oligonucleotide (20 nmol) 50 µL 100 mM sodium acetate buffer, pH 4.5 25 µL deionized water 5 µL 40 mM TAMRA-5-maleimide solution. 3. Incubate the mixture for 4 hr at 60°C in a heating block. If desired, store mixture up to 2 to 4 weeks at −20°C. Remove excess dye by size-exclusion chromatography 4. Remove the top of the NAP-5 column and discard the preservative solution into a waste container. Remove the bottom cap. 5. Equilibrate the NAP-5 column with 10 mL of deionized water. 6. Load 100 µL of the labeling solution and allow it to enter the NAP-5 gel bed completely. 7. Load 400 µL of deionized water; discard the eluate. 8. Elute twice, each time with 500 µL of deionized water, and collect 500-µL fractions in separate vials. 9. Identify product-containing fractions by OD260 measurement (UNIT 10.3). 10. Combine the product-containing fractions and evaporate to dryness in a Speedvac evaporator. 11. Optional: Purify crude oligonucleotide by RP-HPLC (UNIT 10.5) or PAGE (UNIT 10.4). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
TLC stain Solution A: Weigh 0.2 g of 3-methoxyphenol (Aldrich) into a 250-mL beaker and add absolute ethanol to make 100 g of solution (0.2% w/w final). Solution B: Add 5.3 mL concentrated sulfuric acid (H2SO4) to a 250-mL beaker containing ∼50 mL of absolute ethanol. Add ethanol to make 100 mL (1 M H2SO4 final). TLC stain: In a 250-mL screw-cap vial, mix equal volumes of solutions A and B (usually 50 mL). Prepare fresh. CAUTION: Sulfuric acid is corrosive. Avoid skin and eye contact. Addition of the acid to ethanol is exothermic.
COMMENTARY Background Information The chemical synthesis of modified oligonucleotides is of increasing interest for studying biological processes in vivo and in vitro, in particular for diagnostic applications. Oligonucleotides are modified to prepare conjugates with reporter groups such as fluorophores or biotin, ligands for radioactive labeling, peptides, enzymes, antibodies, and carbohy-
drates (Goodchild, 1990). Other applications require the attachment of oligonucleotides to surfaces, e.g., to produce microarrays (Bowtell and Sambrook, 2002) or to conjugate oligonucleotides to nanoparticles (Letsinger et al., 2000). A number of strategies have been applied to chemically modify oligonucleotides at the 3′ and/or 5′ terminus (UNITS 4.2, 4.3, 4.5, & 4.6) or in the middle of the sequence. For 3′-modi-
Synthesis of Modified Oligonucleotides and Conjugates
4.18.11 Current Protocols in Nucleic Acid Chemistry
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fication, a branched linker is attached to the solid support employed for oligonucleotide synthesis. The linker either comprises a functional group for postsynthetic modification reactions or an attached reporter group or ligand that alleviates the need for postsynthetic modifications (UNITS 4.5 & 4.6). Functionalized nucleoside building blocks are used to incorporate a modification in the middle of the sequence (Telser et al., 1989) or at the 5′ terminus (Smith et al., 1985). For 5′-modification, a non-nucleosidic phosphoramidite such as fluorochrome amidite or a biotin-functionalized amidite can be coupled to the oligonucleotide in the last synthesis cycle (UNITS 4.2 & 4.3). Many fluorochrome-phosphoramidites and other amidites containing reporter groups of haptens are commercially available to allow on-support labeling. As an alternative method, the phosphoramidite employed in the last synthesis cy-
Protected FG
cle can contain a protected functional group. After its deprotection, the functional group is available for postsynthetic conjugation to a modifier containing the corresponding functionality for attachment (UNIT 4.2). Examples of standard conjugation methods are given in Figure 4.18.4. The Diels-Alder conjugation method (Hill et al., 2001) extends the portfolio of conjugation chemistries, as it contains some unique features. (1) The method is applicable for conjugation reactions in aqueous media and therefore allows postsynthetic labeling of oligonucleotides. (2) Protection of the functional group is not necessary, as the diene moiety is stable under the standard synthesis and deprotection conditions employed for solid-phase oligonucleotide synthesis.
Incoming FG
FG
O
Covalent linkage
O O
NHMMTr
NH2
N
O
NHTFA
NH O amine
NHS-ester
amide
O
O S
S
SR
SH
N
N
O disulfide
thiol
O
O
O P OR OR phosphate triester
O
maleimide
HO
O P
−
O
O
terminal phosphate
alcohol
phosphate diester O
N
N O
O diene
O −
O
Synthesis of Oligonucleotide Conjugates via Diels-Alder Cycloaddition
sulfide
O OH
P
O
maleimide
Diels-Alder adduct
Figure 4.18.4 Common oligonucleotide conjugation methods. Abbreviations: FG, functional group; MMTr, 4-monomethoxytrityl; TFA, 2,2,2-trifluoroacetyl.
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(3) The Diels-Alder conjugation reaction is fast and selective for dienophiles, such as maleimides. (4) The method is orthogonal to other common methods (see Fig. 4.18.4) and therefore allows combination with other functionalities (e.g., amines) to prepare dual-labeled probes in one pot. Basic Protocol 3 allows for the conjugation of diene-modified oligonucleotides to any substrate containing maleimides or other suitable dienophiles and allows the preparation of modified oligonucleotides used in the spectrum of biological applications.
Critical Parameters Synthesis of diene-alcohol The synthesis of the diene-alcohol S.5 is straightforward and can be conveniently performed on a larger scale. It is not necessary to purify intermediates S.2 to S.4, as compound S.5 can be satisfactorily purified by fractional distillation. For successful performance of the bis-dehydrohalogenation, it is crucial to use fresh potassium tert-butoxide and to use anhydrous conditions; otherwise, prolonged reaction times or incomplete conversion will be observed. If the reaction is not complete after the time given in the protocol, an additional portion of potassium tert-butoxide (∼10% of the initial amount) should be added and the reaction mixture stirred for another hour. Compounds S.4 and S.5 appear to be volatile liquids. After evaporation of the organic phases containing these volatile intermediates, drying under high vacuum should be avoided. Purification and storage of diene-amidite If wet silica gel is used for the flash chromatography of diene-amidite S.7, the formation of byproducts may be observed. Equilibration of the column employing a 1% solution of triethylamine in EtOAc, instead of EtOAc alone, helps to buffer the acidic silica gel. Subsequently, the column should be treated with EtOAc to replace the triethylamine solution. After purification, the amidite (S.7) should be dried under high vacuum and stored under dry argon in a refrigerator at 4°C. It is advantageous to distribute the amidite to appropriate clean, dry synthesizer bottles immediately after preparation in order to avoid repeated opening of the amidite container. The bottles should be flushed with argon and sealed.
Oligonucleotide synthesis The synthesizer used for oligonucleotide synthesis should be operated and maintained according to the manufacturer’s instructions. It is crucial to use appropriate solvents and reagents and to prime all the lines carefully prior to synthesis. A freshly prepared solution of diene-amidite (S.7) should be used. It is not necessary to prolong the coupling time for the final diene-amidite coupling compared to coupling of standard amidites. Diels-Alder cycloaddition reaction The reaction conditions for the Diels-Alder conjugation reaction have given the best labeling results. For sensitive substrates, it could be desirable to carry out the Diels-Alder conjugation under different reaction conditions, such as lower temperature or higher pH. Successful conjugation may only be possible in exchange for a lower yield. Limitations are: (1) fast hydrolysis of maleimides at pH >8, (2) decreased reaction rates at temperatures <35°C, and (3) the requirement that DMF content not exceed 20%. Purification of crude oligonucleotide conjugates can be performed by standard methods such as reversed-phase high-performance liquid chromatography (RP-HPLC; UNIT 10.5) or polyacrylamide gel elctrophoresis (PAGE; UNIT 10.4).
Anticipated Results By applying the methods in this unit and observing the critical parameters, it is possible to prepare the diene-amidite (S.7) at a 30% overall yield, with a purity of ≥98%, determined by RP-HPLC and 31P NMR analysis (see Basic Protocol 2). Diene-functionalization of oligonucleotides employing pure phosphoramidite (S.7) usually provides coupling efficiencies >99%. The conjugation of dienefunctionalized oligonucleotides to maleimides typically proceeds with efficiencies >80%.
Time Considerations The preparation of diene-amidite (S.7) takes ∼2 weeks, if the scale given in the protocol is applied. The time for the assembly of a 5′-dienemodified oligonucleotide (synthesis of a 20mer on 1-µmol scale), including deprotection and desalting, takes ~3 days. The conjugation reaction, including size-exclusion chromatography, typically requires 2 days. Synthesis of Modified Oligonucleotides and Conjugates
4.18.13 Current Protocols in Nucleic Acid Chemistry
Supplement 14
Literature Cited Bowtell, D. and Sambrook, J. (eds.) 2002. DNA Microarrays: A Molecular Cloning Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Goodchild, J. 1990. Conjugates of oligonucleotides and modified oligonucleotides: A review of their synthesis and properties. Bioconjugate Chem. 1:165-187. Hill, K.W., Taunton-Rigby, J., Carter, J.D., Kropp, E., Vagle, K., Pieken, W., McGee, D.P.C., Husar, G., Leuck, M., Anziano, D., and Sebesta, D.P. 2001. Diels-Alder bioconjugation of dienemodified oligonucleotides. J. Org. Chem. 66:5352-5358. Letsinger, R.L., Elghanian, R., Biswanadham, G., and Mirkin, C. 2000. Use of a steroid cyclic disulfide anchor in constructing gold nanoparticle-oligonucleotide conjugates. Bioconjugate Chem. 11:289-291.
Smith, L.M., Fung, S., Hunkapiller, M.W., Hunkapiller, T.J., and Hoo, L.E. 1985. The synthesis of oligonucleotides containing an aliphatic amino group at the 5′ terminus: Synthesis of fluorescent DNA primers for use in DNA sequence analysis. Nucl. Acids Res. 13:2399-2412. Telser, J., Cruickshank, K.A., Morrison, L.E., and Netzel, T.L. 1989. Synthesis and characterization of DNA oligomers and duplexes containing covalently attached molecular labels: Comparison of biotin, fluorescein, and pyrene labels by thermodynamic and optical spectroscopic measurements. J. Am. Chem. Soc. 111:6966-6976.
Contributed by Michael Leuck and Andreas Wolter Proligo LLC Boulder, Colorado
Synthesis of Oligonucleotide Conjugates via Diels-Alder Cycloaddition
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Current Protocols in Nucleic Acid Chemistry
5′-Iodination of Solid-Phase-Linked Oligodeoxyribonucleotides
UNIT 4.19
This unit describes a method for converting the 5′-hydroxyl of CPG-bound oligodeoxyribonucleotides to an iodo group. This procedure is useful when an electrophilic group is desired in the terminal 5′-position of an oligodeoxyribonucleotide. Because the iodo group is an efficient leaving group in SN2 displacement reactions, its introduction at the 5′-position allows for facile ligation to small molecules or macromolecules containing a strongly nucleophilic group. Additionally, 5′-iodo oligonucleotides can easily be converted to a wide variety of other functional groups. Two protocols are described in this unit. For both procedures, an oligonucleotide is prepared on an automated DNA synthesizer (see APPENDIX 3C) and is then iodinated in-column (i.e., while still protected and immobilized on the CPG resin in the column). The first method (see Basic Protocol) describes a manual procedure performed by adding reagents to the column after it is removed from the synthesizer. The second method (see Alternate Protocol) provides an automated method performed by programming the synthesizer to perform the same iodination reaction. The manual procedure is suitable for most applications, while the automated method may be preferable for carrying out multiple iodination reactions. CAUTION: All chemicals must be used in a fume hood by qualified individuals equipped with laboratory coats, safety glasses, and gloves. MANUAL PROCEDURE FOR 5′-IODINATION OF OLIGODEOXYRIBONUCLEOTIDES ON A SOLID SUPPORT This protocol outlines a method to convert the 5′-hydroxyl on protected CPG-bound oligodeoxyribonucleotides to an iodo group (Fig. 4.19.1). In this procedure, an iodination solution is passed between two syringes fitted to the ends of a DNA synthesis column containing the attached oligodeoxyribonucleotide, converting the 5′-hydroxyl to an iodo group. The 5′-iodo-modified oligodeoxyribonucleotide can subsequently be cleaved and deprotected using 28% aqueous NH4OH.
BASIC PROTOCOL
Materials Anhydrous N,N-dimethylformamide (DMF) Iodination solution (see recipe) Methylene chloride (CH2Cl2) 28% (v/v) ammonium hydroxide (NH4OH) DNA synthesizer (e.g., ABI; see APPENDIX 3C) 0.2 to 10 µmol DNA synthesis column with long-chain alkylamine controlled-pore glass (CPG) support 1- and 10-mL syringes Shaker, rocker, or other agitating device C18 reversed-phase HPLC column (UNIT 10.5) Speedvac evaporator Additional reagents and equipment for oligonucleotide synthesis (Chapters 3 and 4 and APPENDIX 3C) and reversed-phase HPLC (UNIT 10.5) Synthesis of Modified Oligonucleotides and Conjugates Contributed by Eric T. Kool and Gregory P. Miller Current Protocols in Nucleic Acid Chemistry (2003) 4.19.1-4.19.8 Copyright © 2003 by John Wiley & Sons, Inc.
4.19.1 Supplement 14
HO
I
B1
O O
O P
O
(PhO)3PCH3I OCH2CH2CN
O
O
O P dry DM F
CPG
OCH2CH2CN
O
B2
O
O
O N H
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O
O O
n CPG
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O
Figure 4.19.1 Iodination of solid-phase-linked oligodeoxyribonucleotides. Abbreviations: B1 and B2, thymine and/or N-protected nucleobases; CPG, long-chain alkylamine controlled-pore glass (resin).
Synthesize oligonucleotide and obtain control sample 1. Using a DNA synthesizer, synthesize 0.2 to 10 µmol of CPG-bound protected oligodeoxyribonucleotide with the 5′-O-(4,4′-dimethoxytrityl) group removed (DMTr-OFF; see APPENDIX 3C). 2. Open DNA synthesis column, remove and set aside ∼1 mg of resin, then close column. This sample of the resin will be cleaved and deprotected without undergoing the iodination reaction and thus can be used as an HPLC reference to determine the retention time of the starting material and estimate the retention time of the 5′-iodinated product.
Dehydrate resin 3. Attach an empty 1.0-mL syringe to one end of the DNA synthesis column. To the other end, attach a 1.0-mL syringe filled with anhydrous DMF. 4. Slowly push DMF through column to the other syringe. This step should remove any water that may be adsorbed to the resin that would otherwise react with the water-sensitive iodination reagent.
Iodinate oligodeoxyribonucleotide 5. Remove the DMF-filled syringe and quickly replace with a 1.0-mL syringe filled with iodination solution. 6. Push iodination solution back and forth between the two syringes several times. 7. Seal the junctions between the column and the syringes with Parafilm to prevent leakage. 8. Put on shaker, rocker, or other agitating device for 15 to 20 min at room temperature. If the 5′-end of the oligonucleotide contains a bulky group, it may be desirable to increase the reaction time (see Critical Parameters and Troubleshooting for details).
9. Unwrap the column and use the syringes to remove the iodination solution from the column. 5′-Iodination of Solid-Phase-Linked Oligodeoxyribonucleotides
10. Using a 10-mL syringe, push 5 to 10 mL DMF through the column, followed by a few milliliters of CH2Cl2. Blow-dry beads using air. The CH2Cl2 is not mandatory, but it facilitates drying of the CPG beads, making it easier to remove them from the column.
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Cleave and deprotect iodinated deoxyribonucleotide 11. Open column and transfer beads to a sealed vial for deprotection. 12. Cleave and deprotect the iodinated product and the sample of starting material (from step 2) by soaking each resin in 28% NH4OH (∼1 mL for 1-µmol column) at room temperature for 18 to 24 hr. 13. Remove ammoniacal solution, rinse beads twice with small portions (e.g., 0.25 mL for 1-µmol column) of deionized water, combine portions, and evaporate liquids in a Speedvac evaporator. 14. Analyze purity by HPLC using a C18 reversed-phase column (UNIT 10.5). AUTOMATED PROCEDURE FOR 5′-IODINATION OF OLIGODEOXYRIBONUCLEOTIDES ON A SOLID SUPPORT
ALTERNATE PROTOCOL
This protocol involves the use of a DNA synthesizer to perform the same iodination reaction described in the Basic Protocol. The advantage of this automated procedure is that the iodination reaction can be conveniently run on the DNA synthesizer as a 15-min procedure following DNA synthesis. Depending on the number of columns on the particular synthesizer, multiple reactions may be run simultaneously. Although the iodination reagent is not harmful to the synthesizer, the reagent must be filtered prior to use to ensure that no particulates clog the lines. Because of this additional step, it is often more convenient to use the manual procedure if only one or two iodination reactions will be performed. If multiple reactions are to be performed, the automated procedure is generally faster and more convenient. Additional Materials (also see Basic Protocol) Glass wool or line filter (e.g., preparation and delivery line filter from ABI) Empty DNA synthesis column (e.g., 1000-Å CPG column; ABI) 10-mL syringes Clean, oven-dried phosphoramidite bottle compatible with synthesizer Reagent bottle compatible with synthesizer, filled with anhydrous DMF Synthesize oligonucleotide 1. Using a DNA synthesizer, synthesize 0.2 to 10 µmol CPG-bound protected oligodeoxyribonucleotide with the 5′-O-DMTr group removed (see APPENDIX 3C). Some resin (≤1 mg) may be removed at this point to use as a reference in HPLC.
Filter iodination solution for use in synthesizer 2. Filter iodination solution under anhydrous conditions as follows. Insert a small filter or glass wool into an empty DNA synthesis column. Take up iodination solution in a 5- or 10-mL syringe, and attach it to the column/filter. Attach another syringe to the opposite end of the column. Push solution through column/filter into the opposite syringe. Generally, each reaction requires ≤1 mL of iodination solution. The Preparation and Delivery Line Filter from ABI is the ideal size and shape to fit into a 1000-Å ABI CPG column. If the solution contains a large quantity of particulates, the filter may become clogged. If this happens, it may be necessary to remove the syringes (while making sure to minimize exposure to the air) and wash the filter. To wash it, simply push dry DMF through the column filter in the opposite direction from that in which the iodination solution was flowing. The above filtration system is only one of a variety of ways that the filtration can be performed.
Synthesis of Modified Oligonucleotides and Conjugates
4.19.3 Current Protocols in Nucleic Acid Chemistry
Supplement 14
Table 4.19.1 IodoCycle Program for ABI 392 and 394 Synthesizersa
Step
Function
Time (sec)
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31
Begin Prep DMF DMF to waste DMF to column Reverse flush Block flush Phos prep Iodo to column DMF to waste Block flush Wait Reverse flush Iodo to column DMF to waste Block flush Wait DMF to column Flush to waste DMF to column Flush to waste DMF to column Reverse flush Block flush CH3CN to waste CH3CN to column Reverse flush CH2Cl2 to waste CH2Cl2 to column Reverse flush Block flush End
15 4 15 10 4 3 20 3 4 300 10 20 3 4 300 15 10 20 10 20 10 4 3 20 10 3 20 10 4
aModified from Miller and Kool (2002) with permission from the American
Chemical Society.
3. Attach a needle to the syringe containing the iodination solution. 4. Inject the iodination solution into an oven-dried and septum-sealed phosphoramidite bottle. Prepare synthesizer for iodination reaction 5. Write an iodination procedure for the DNA synthesizer containing the following steps:
5′-Iodination of Solid-Phase-Linked Oligodeoxyribonucleotides
a. b. c. d. e.
Rinse beads with anhydrous DMF. Send iodination solution to column. Wait (5 min). Flush iodination solution to waste. Send iodination solution to column.
4.19.4 Supplement 14
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f. Wait (5 min). g. Rinse with anhydrous DMF. An example of a functional procedure for ABI 392 and 394 synthesizers is shown in Table 4.19.1.
6. Install the iodination reagent into one of the phosphoramidite positions and install a bottle of anhydrous DMF into one of the reagent positions on the DNA synthesizer. 7. Edit sequence: enter a single base as the sequence. If this is not done, the iodination cycle will repeat itself for each base in the sequence.
Iodinate oligodeoxyribonucleotide 8. Start synthesis. Due to differences between synthesizers, it is recommended that the column be observed while the cycle is run the first few times to make sure the “iodination solution to column” and “wash” sequences are of the correct length of time. A period of 20 to 30 sec for “iodination solution to column” should be sufficient to completely fill the columns on an ABI 392 or 394 synthesizer.
Cleave and deprotect iodinated deoxyribonucleotide 9. After the synthesis is complete, transfer the CPG resin to a sealed vial. Cleave and deprotect the iodinated product by soaking the resin in 28% NH4OH (∼1 mL for 1-µmol column) at room temperature for 18 to 24 hr. 10. Remove iodination solution and DMF from synthesizer. To ensure that the iodination reagent does not damage or contaminate the synthesizer, replace the iodination reagent bottle with one containing anhydrous DMF and use the manual functions to briefly rinse the lines. The unused iodination solution can be stored in the phosphoramidite bottle (see recipe for storage conditions).
11. Analyze purity using HPLC with a C18 reversed-phase column (UNIT 10.5). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Iodination solution 0.226 g (0.5 mmol) methyltriphenoxyphosphonium iodide [(PhO)3PCH3I; Aldrich; 0.5 M final] 0.85 ml anhydrous N,N-dimethylformamide (DMF) Prepare fresh or store up to 3 months at –70°C If the solution will be used on a DNA synthesizer, it should be filtered before use (see Alternate Protocol). (PhO)3PCH3I reacts quickly with water and is light sensitive. Minimize exposure of the reagent to the atmosphere while weighing, and promptly flush the vial and the original reagent bottle with N2 or argon when finished. It is convenient to prepare the iodination solution during oligonucleotide synthesis. If the solution will be used within a few hours of its preparation, there is no need to protect it from light. If the iodination solution has been frozen, always refilter (see Alternate Protocol, step 2) after thawing. (PhO)3PCH3I is often the consistency of mud or a thick oil. Although this makes it more difficult to manipulate, it generally still works well.
Synthesis of Modified Oligonucleotides and Conjugates
4.19.5 Current Protocols in Nucleic Acid Chemistry
Supplement 14
COMMENTARY Background Information
5′-Iodination of Solid-Phase-Linked Oligodeoxyribonucleotides
In 1997, Xu and co-workers described a DNA-ligation method which allowed for the efficient ligation of a 5′-iodo-modified oligodeoxyribonucleotide with a 3′-phosphorothioated oligonucleotide (Xu and Kool, 1997). In the presence of a complementary template, ligation proceeded efficiently under mild conditions. The product of this ligation method is an oligonucleotide identical to natural DNA with the exception that one oxygen atom in the phosphodiester backbone is replaced by a sulfur atom; it therefore retains most of the chemical properties of natural DNA (Xu and Kool, 1998). The 5′-iodo-modified oligomers were prepared on a DNA synthesizer using a prev iou sly synthesized 5′-iodo-dT phosphoramidite. Unfortunately, 5′-iodo-modified phosphoramidites are time-consuming and expensive to synthesize, so the utility of this ligation system was significantly limited. It was subsequently found that the 5′-iodooligodeoxyribonucleotides are in fact accessible via a much simpler, more direct route (Miller and Kool, 2002). The protocols described in this unit are based on this new approach, in which 5′-iodination is carried out directly on the DNA column, before deprotection and cleavage. The protecting groups used on the DNA bases and backbone effectively prevent the very reactive (PhO)3PCH3I from reacting at any site other than the 5′-hydroxyl. This is true when using both Glen Research’s Sterling 2-cyanoethyl (CE) phosphoramidites (5′-DMTr-dABz, 5′-DMTr-dCBz, and 5′-DMTrdGi-Bu) or their Ultramild CE phosphoramidites (5′-DMTr-dAi-Bu or 5′-DMTr-dAAcOPh, 5′DMTr-dGAcOPh-i-Pr, and 5′-DMTr-dCAc). Also, it was found that when deprotection and cleavage are carried out at room temperature for 24 hr, very little hydrolysis of the iodo group occurs. This new method of iodinating the 5′end of oligodeoxyribonucleotides effectively makes the 5′-iodo/3′-phosphorothioate ligation system accessible to anyone with access to synthetic DNA. Unfortunately, it was found th at b oth 2′-O-tert-butyldimethylsilyl (TBDMS) and 2′-O-triisopropylsilyloxymethyl (TOM) protecting groups on protected oligoribonucleotides are cleaved by both (PhO)3PCH3I and another iodinating system, PPh3/I2/imidazole in DMF (Kool and Miller, unpub. observ.). In addition to their usefulness in DNA ligation reactions, the 5′-iodo oligomers are also
good intermediates for making other types of 5′-modifications. The 5′-iodo group is easily converted to thiols, amines, azides, thioethers, thiocyanates, and selenides while the oligomer is still attached to resin (Kool and Miller, unpub. observ.). 5′-Thiols and amines are convenient handles for attaching isothiocyanates, succinimidyl esters, maleimides, iodoacetamides, and other commercially available labels. Alternatively, thiol-containing moieties will react directly with 5′-iodo oligomers, forming very stable thioethers.
Critical Parameters and Troubleshooting The most important key to success in running this reaction is to keep the iodination reagent anhydrous. The large excess of reagent called for in this protocol should temper this potential problem. Care should nevertheless be taken to minimize the amount of time the reagent is exposed to the air, and the reagent and iodination solution should always be stored under an inert gas. Another important point is that the reaction time may need to be increased when bulkier groups are in the 5′-position. For example, when pyrene is in the 5′-position, the iodination reaction requires ∼25 min for completion. Additionally, if there is a modified base containing a strong nucleophile near the 5′-end, it may displace the iodo group, cyclizing the oligomer either during synthesis or during deprotection. This is often observed when the 5′-base is a purine, in which the N3 nitrogen attacks the 5′-carbon, resulting in some cyclized material (Dimitrijevich et al., 1979). This cyclized byproduct may be hydrolyzed back to the 5′hydroxy starting material upon treatment with NH4OH.
Anticipated Results The hydrophobicity of the iodo group allows for easy separation of 5′-modified oligodeoxyribonucleotides from their 5′-hydroxy starting materials via reversed-phase HPLC (UNIT 10.5). For example, using a gradient of 0% to 20% CH3CN in 50 mM triethylammonium acetate (TEAA), pH 7.0, over a 20-min period to elute the crude reaction mixture from an analytical C18 reversed-phase column, iodinated trimers elute 5 min later than their corresponding starting materials, while iodinated 13-mers elute ∼2 min after their respective starting materials (see Fig. 4.19.2). A chromatogram of the de-
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protected 5′-hydroxyl starting material should therefore give an indication of when the iodinated product will come off of the column. Also, knowledge of the starting material retention time allows for calculation of percent conversion to the iodinated product. Electrospray ionization (ESI) mass spectrometry (UNIT 10.2) should be used to verify the identity of the product. The success of this iodination reaction appears to depend on the identity of the 5′-base,
3000
13T-I
12
but not the overall base composition, of a particular oligomer. Generally, 5′-pyrimidines afford the highest yields because they do not cyclize as easily as purines. Yields of 70% to 90% are typical for pyrimidines and 50% to 80% for purines, based on total integration of HPLC chromatograms (Miller and Kool, 2002, and unpub. observ.). Because the starting material will contain failure sequences from incomplete phosphoramidite coupling during oligomer synthesis, HPLC chromatograms will
13T-OH
13
14
15
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13C-I
13C-OH
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0 11
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13
13A-I
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17
13A-OH
2000
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0 12
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14
13G-I
13G-OH
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0 12
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14 Minutes
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Figure 4.19.2 RP-HPLC profiles of 5′-d(NGTAGGCAAGAGT) before and after 5′-iodination. From top to bottom, N = T, C, A, and G, respectively. Reprinted from Miller and Kool (2002) with permission from the American Chemical Society.
Synthesis of Modified Oligonucleotides and Conjugates
4.19.7 Current Protocols in Nucleic Acid Chemistry
Supplement 14
contain multiple peaks. However, a single predominant product peak with a longer retention time should be observed when using a reversedphase column.
Time Considerations The manual procedure described in the Basic Protocol can be completed in less than an hour. The automated procedure requires more time to set up the reactions, but once the reagent and synthesizer are prepared, many reactions can be performed quickly. For example, if a 4-column synthesizer is used with a 15-min iodination cycle, 16 reactions can be performed per hour.
Miller, G.P. and Kool, E.T. 2002. A simple method for electrophilic functionalization of DNA. Org. Lett. 4:3599-3601. Xu, Y. and Kool, E.T. 1997. A novel 5′-iodonucleoside allows efficient non-enzymatic ligation of single-stranded and duplex DNAs. Tetrahedron Lett. 38:5595-5598. Xu, Y. and Kool, E.T. 1998. Chemical and enzymatic properties of bridging 5′-S-phosphorothioester linkages in DNA. Nucl. Acids Res. 26:31593164.
Key References Miller and Kool, 2002. See above. Reports a method to iodinate the 5′-carbon of oligodeoxyribonucleotides on a solid support.
Literature Cited Dimitrijevich, S.D., Verheyden, J.P.H., and Moffatt, J.G. 1979. Halo sugar nucleosides. 6. Synthesis of some 5′-deoxy-5′-iodo and 4′,5′-unsaturated purine nucleosides. J. Org. Chem. 44:400-406.
Contributed by Eric T. Kool and Gregory P. Miller Stanford University Stanford, California
5′-Iodination of Solid-Phase-Linked Oligodeoxyribonucleotides
4.19.8 Supplement 14
Current Protocols in Nucleic Acid Chemistry
Reversible Biotinylation of the 5′-Terminus of Oligodeoxyribonucleotides and Its Application in Affinity Purification
UNIT 4.20
In UNIT 4.2, several methods for biotinylation of DNA are described (see structures 95b, 96, 101, 104a, 106, and 124a-c in that unit). Except for structure 124c, in which the biotin moiety can be removed nondestructively by UV irradiation, other biotinylation methods are irreversible. UNIT 4.3 presents protocols for direct attachment of acridine and psoralen derivatives to the 5′-terminus of DNA via their phosphoramidites. In this unit, protocols are given for the preparation of two reversible biotinylation phosphoramidites for direct labeling of the 5′-terminus of DNA (see Basic Protocols 1 and 2 and Support Protocol 2) and their applications in NeutrAvidin-coated microsphere-mediated affinity purification of synthetic oligodeoxyribonucleotides (see Support Protocol 1). The DNAs synthesized using these two phosphoramidites feature a diisopropyl silyl acetal linkage between biotin and their 5′-termini; these linkages can be readily broken by fluoride ions. The first of the two phosphoramidites yields 5′-OH unmodified DNA upon treatment with HF/pyridine, while the second phosphoramidite yields 5′-phosphate DNA on treatment with HF/pyridine followed by aqueous methylamine. CAUTION: All chemicals must be handled in a well-ventilated fume hood by individuals equipped with laboratory coats, safety glasses, and gloves. REVERSIBLE BIOTINYLATION VIA A DIISOPROPYL SILYL ACETAL LINKER YIELDING 5′-OH DNA This protocol describes the preparation of the reversible biotinylation phosphoramidite S.3 (see Fig. 4.20.1), its coupling to the 5′-end of DNA on an automatic solid-phase synthesizer, and postsynthetic cleavage/deprotection to afford the biotinylated DNA S.4 (see Fig. 4.20.1). A sample sequence is shown in S.5. Other appropriately protected deoxyribonucleosides, such as N6-benzoyldeoxyadenosine, N2-isobutyryldeoxyguanosine, and N4-acetyldeoxycytosine, should be able to be biotinylated using the same procedure. Materials Biotinyl alcohol S.1 (see Support Protocol 2 for preparation) Imidazole (99%) Argon and nitrogen gas N,N-Dimethylformamide (DMF, anhydrous) Diisopropylethylamine (DIEA) Diisopropyldichlorosilane (Fluka) Thymidine (99%+) Ethyl acetate 5% (w/v) sodium bicarbonate (NaHCO3) Sodium sulfate (Na2SO4, anhydrous) Chloroform (CHCl3) Methanol 40-µm silica gel (Baker) TLC plates: silica gel on aluminum (60F-254, 200-µm thickness) Methylene chloride (CH2Cl2, anhydrous) 2-Cyanoethyl-N,N,N′,N′-tetraisopropylphosphoramidite (2-cyanoethyltetraisopropylphosphorodiamidite, 97%) Contributed by Shiyue Fang and Donald E. Bergstrom Current Protocols in Nucleic Acid Chemistry (2003) 4.20.1-4.20.17 Copyright © 2003 by John Wiley & Sons, Inc.
BASIC PROTOCOL 1
Synthesis of Modified Oligonucleotides and Conjugates
4.20.1 Supplement 14
1H-Tetrazole (99%+, sublimed) Tetrahydrofuran (THF, 99%+) 5′-DMTr, 2-cyanoethyl phosphoramidite monomers: 5′-O-dimethoxytrityl-N6-benzoyl-2′-deoxyadenosine-3′-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (dABz) 5′-O-dimethoxytrityl-N2-isobutyryl-2′-deoxyguanosine-3′-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (dGi-Bu) 5′-O-dimethoxytrityl-N4-acetyl-2′-deoxycytidine-3′-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (dCAc) 5′-O-dimethoxytrityl-2′-deoxythymidine-3′-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite (T) Acetonitrile (CH3CN, anhydrous)
O S H
H NH
N O
O
N H
O
H N
OH O
1
O
1. i-Pr2SiCl2, imidazole, DIEA 2. thymidine, imidazole O S H N O
N H
H NH
O
NH
H N
O
O
O Si O
O
O
2 (92%)
O
O
N
OH
(i-Pr2N)2POCH2CH2CN, 1H-tetrazole O O
S H N O
N H
H NH
O
NH
H N
O
O Si O
N
O
O
O
O O
3 (85%)
NC
O
P
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O O S H HN
N H
H NH
O
NH
H N
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O Si O
O
O
N
O
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O
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O−
O
O S H HN
H NH
N H
O
O
H N
O Si O TACAGTGACT O
5′
3′
OH
5
O
Reversible Biotinylation of the 5′-Terminus of Oligodeoxyribonucleotides
Figure 4.20.1 Preparation of reversible biotinylation phosphoramidite S.3 and structures of biotinylated 5′-OH DNAs S.4 and S.5.
4.20.2 Supplement 14
Current Protocols in Nucleic Acid Chemistry
∼29% (v/v) ammonium hydroxide (NH4OH) ∼40% (v/v) methylamine (CH3NH2) 50-mL one-neck round-bottomed flasks (oven dried) Vacuum pump 10-, 5-, 1-, 0.25-mL graduated glass syringes 20-G, 5-cm stainless steel needles Septa (14/20, 24/40) 250-mL separatory funnel Filter funnel and Whatman no. 1 filter paper Rotary evaporator equipped with a water aspirator Flash chromatography columns (4.5 × 12–cm, 3.0 × 12–cm) UV lamp 5-mL screw-cap vials Speedvac evaporator Additional reagents and equipment for thin-layer chromatography (TLC, APPENDIX 3D), column chromatography (APPENDIX 3E), solid-phase automatic DNA synthesis (APPENDIX 3C) using classical phosphoramidite method (UNIT 3.3), and purification of oligonucleotides by SDS-PAGE (UNIT 10.4) and reversed-phase HPLC (UNIT 10.5) Prepare biotinyl thymidine S.2 1. Dry 1.7 g (2.62 mmol) biotinyl alcohol S.1 in a 50-mL one-neck round-bottomed flask (equipped with a magnetic stir bar) under vacuum overnight at room temperature. 2. Add 211 mg (3.1 mmol) imidazole and then flush the flask with an argon flow for 10 min. 3. Add 5 mL of dry DMF and 2.7 mL (15.5 mmol) DIEA with 10-mL and 5-mL glass syringes attached to 20-G stainless steel needles, respectively, and cool the mixture to 0°C. 4. Add 839 µL (4.65 mmol) diisopropyldichlorosilane with a 1-mL glass syringe in one portion. Stir the mixture at 0°C for 1 hr, then at room temperature for 4 hr on a magnetic stir plate. The tertiary hydroxyl group in S.1 substitutes one of the two chlorine atoms in diisopropyldichlorosilane; the other chlorine atom will be substituted by the primary hydroxyl group of thymidine in the next step.
5. In another 50-mL oven-dried, one-neck round-bottomed flask, place 1.5 g (6.2 mmol) thymidine and 422 mg (6.2 mmol) imidazole. Seal the flask with a septum, flush with argon gas for ∼10 min, and then add 5 mL dry DMF with a 20-G needle attached to a 10-mL glass syringe. Shake and dissolve the solids. 6. Cool the solution prepared in step 4 to 0°C. Add the solution prepared in step 5 with a 10-mL glass syringe and stir the mixture at 0°C for 4 hr. 7. Add 50 mL ethyl acetate and 50 mL of 5% NaHCO3, transfer the mixture to a 250-mL separatory funnel, and separate the organic phase. 8. Extract the aqueous phase four times with 50 mL ethyl acetate. 9. Combine the organic phases, dry over ∼2 g anhydrous Na2SO4, and filter off the solids using a filter funnel and Whatman no. 1 filter paper. Concentrate the filtrate to dryness on a rotary evaporator with a water aspirator.
Synthesis of Modified Oligonucleotides and Conjugates
4.20.3 Current Protocols in Nucleic Acid Chemistry
Supplement 14
10. Dissolve the residue in ∼10 mL of 9:1 (v/v) CHCl3/methanol and apply to the top of a 4.5 × 12–cm flash chromatography column (APPENDIX 3E) prepared with a slurry of 40-µm silica gel in the same solvent system. Elute the column with the same solvent system. Monitor the fractions by TLC (APPENDIX 3D). 11. Pool the fractions that contain the UV-active spot with an Rf = 0.4 (9:1 CHCl3/methanol), concentrate to dryness on a rotary evaporator, and then dry under vacuum. S.2 is obtained with an expected yield of 92% (2.49 g). 1H NMR (CD3OD, 500 MHz) δ 0.91-1.05 (m, 14H), 1.19 (s, 6H), 1.28 (s, 9H), 1.28-1.29 (m, 1H), 1.40-1.43 (m, 2H), 1.58-1.83 (m, 5H), 1.81 (s, 3H), 2.12-2.30 (m, 4H), 2.94-3.05 (m, 2H), 3.25-3.33 (m, 5H), 3.46-3.51 (m, 4H), 3.56-3.57 (m, 4H), 3.90-4.00 (m, 3H), 4.23 (dd, 1H, J = 4.5, 7.7 Hz), 4.38-4.40 (m, 1H), 5.13 (dd, 1H, J = 4.8, 7.7 Hz), 7.37 (d, 2H, J = 8.1 Hz), 7.44 (d, 2H, J = 8.4 Hz), 7.47 (s, 1H); 13C NMR (CD3OD, 500 MHz) δ 12.7, 14.5, 14.7, 18.3, 18.3, 18.3, 18.4, 26.8, 29.4, 29.5, 29.8, 30.2, 31.6, 32.1, 32.5, 35.8, 36.7, 38.8, 40.2, 40.3, 41.0, 41.5, 56.0, 56.6, 59.2, 64.0, 64.1, 70.5, 70.6, 71.3, 71.9, 74.8, 86.0, 88.5, 111.5, 125.5, 129.7, 133.6, 137.5, 152.2, 156.0, 158.0, 166.2, 171.7, 176.0, 176.2, 211.4.
Prepare biotinyl thymidine phosphoramidite S.3 12. Dry 220 mg (0.22 mmol) S.2 in a 50-mL oven-dried, one-neck round-bottomed flask (equipped with a magnetic stir bar) under vacuum overnight at room temperature. 13. Flush the flask with an argon flow for ∼5 min, then add (in order) 5 mL dry CH2Cl2 and 76 µL (0.23 mmol) of 2-cyanoethyl-N,N,N′,N′-tetraisopropylphosphoramidite with 10-mL and 250-µL, respectively, glass syringes. Stir the mixture at room temperature until it becomes a clear solution. 14. Add 15.4 mg (0.22 mmol) of 1H-tetrazole in three portions over a period of 1 hr. For each addition, open the neck slightly, quickly add with a spatula, and then stop the flask immediately to minimize the introduction of air into the flask. 15. Stir the reaction mixture for an additional 4 hr, and then concentrate to ∼2 mL on a rotary evaporator. 16. Apply the suspension to the top of a 3.0 × 12–cm flash chromatography column prepared with a slurry of silica gel in 1:1 (v/v) CHCl3/THF. Elute the column with the same solvent system and monitor the fractions by TLC. 17. Pool the fractions that contain the UV-active spot with an Rf = 0.3 (1:1 CHCl3/THF), remove solvents on a rotary evaporator, and dry the product under vacuum. S.3 is obtained as a white foam with an expected yield of 85% (225 mg). 1H NMR (CDCl3, 250 MHz) δ 0.99–1.13 (m, 14H), 1.24 (d, 12H, J = 6.8 Hz), 1.28 (s, 6H), 1.36 (s, 9H), 1.36–1.90 (m, 8H), 1.90 (s, 3H), 2.21–2.39 (m, 6H), 2.75 (dt, 2H, J = 2.0, 5.8 Hz), 3.07–3.10 (m, 2H), 3.32–3.42 (m, 5H), 3.55–3.60 (m, 4H), 3.65 (s, 4H), 3.62–3.92 (m, 4H), 4.01–4.09 (m, 3H), 4.28–4.33 (m, 1H), 4.68–4.72 (m, 1H), 5.20–5.24 (m, 1H), 6.30 (t, 1H, J = 7.5 Hz), 7.49 (d, 4H, J = 7.8 Hz), 7.59 (s, 1H); 31P NMR (CDCl3, 250 MHz) δ 149.7, 149.8.
Couple phosphoramidite S.3 to 5′-end of oligodeoxyribonucleotide 18. Using the 5′-DMTr, 2-cyanoethyl phosphoramidites and following the classical phosphoramidite method (UNIT 3.3, APPENDIX 3C), synthesize the desired oligodeoxyribonucleotide on a DNA synthesizer. After completion, perform an additional detritylation step. Reversible Biotinylation of the 5′-Terminus of Oligodeoxyribonucleotides
For the example in Figure 4.20.1 (S.5), the sequence 3′-TCAGTGACA-5′ was synthesized on a 1-ìmol scale.
4.20.4 Supplement 14
Current Protocols in Nucleic Acid Chemistry
19. Place a solution of phosphoramidite S.3 (0.1 M in dry acetonitrile) on the synthesizer. Carry out coupling by recycling a mixture of 0.1 mL phosphoramidite solution and 0.5 mL of 1H-tetrazole solution (0.45 M in dry acetonitrile) for 15 min. 20. Remove the excess coupling reagents and perform iodine oxidation according to the standard procedure (UNIT 3.3, APPENDIX 3C). Dry the product on CPG under nitrogen flow. Cleave and deprotect biotinylated oligonucleotide 21. Place the oligonucleotide-bound CPG in a 5-mL screw-cap vial, add 500 µL of ∼29% NH4OH and 500 µL of ∼40% methylamine, and heat the suspension to 65°C for 30 min. 22. Cool the vial to –20°C, remove the supernatant, and save. Wash the CPG three times with 500 µL water, keeping all washes. Combine the supernatant and water washes, and dry on a Speedvac evaporator. 23. Purify the biotinylated DNA by gel electrophoresis or reversed-phase HPLC according to procedures in UNIT 10.4 and UNIT 10.5, respectively. The HPLC profile of crude biotinylated DNA S.5 (see Fig. 4.20.1) is illustrated in Figure 4.20.2.
REVERSIBLE BIOTINYLATION VIA A DIISOPROPYL SILYL ACETAL LINKER YIELDING 5′-PHOSPHATE DNA
BASIC PROTOCOL 2
Absorbance (260 nm)
This protocol describes the preparation of the reversible biotinylation phosphoramidites S.7 (see Fig. 4.20.3), its coupling to the 5′-end of DNA on a solid-phase synthesizer, and postsynthetic cleavage/deprotection to afford the biotinylated DNA S.8 (Fig. 4.20.3). A sample sequence is shown in S.9.
a b c d 0
5
10
15
20 25 30 Retention time (min)
35
40
45
50
Figure 4.20.2 HPLC profiles of DNAs S.5 (trace a), 5′-TACAGTGACT-3′ (trace b), S.9 (trace c), and 5′-H2O3PO-ACAGTGACT-3′ (trace d) generated on a C18 reversed-phase column (100 Å, 250 × 4.6 mm, Varian Analytical Instruments), using a linear gradient of 0% to 45% solvent B (90% acetonitrile) in solvent A (0.1 M triethylammonium acetate, 5% acetonitrile) over 60 min at a flow rate of 1 mL/min by detecting the absorbance of DNA at 260 nm. For a detailed protocol for performing reversed-phase HPLC, refer to UNIT 10.5. Synthesis of Modified Oligonucleotides and Conjugates
4.20.5 Current Protocols in Nucleic Acid Chemistry
Supplement 14
Materials Biotinyl alcohol S.1 (see Support Protocol 2 for preparation) Imidazole (99%) Argon and nitrogen gas N,N-Dimethylformamide (DMF, anhydrous) Diisopropylethylamine (DIEA) Diisopropyldichlorosilane (Fluka) Diethyl bis(hydroxymethyl)malonate (97%) Methylene chloride (CH2Cl2) 5% (v/v) citric acid
O S H
H NH
N O
O
N H
H N
O
OH O
1
O
1. i-Pr2SiCl2, imidazole, DIEA 2. diethyl bis(hydroxymethyl)malonate, DIEA O S H
H NH
N O
O
N H
OH CO2Et
CN O
O
N H
H NH
CO2Et
H N
O
O
Si
O
O
O
O
P
N
CO2Et
7 (61%)
O S H HN
O
(i-Pr2N)2POCH2CH2 CN, 1H-tetrazole O
N
Si
6 (92%)
S
O
O O
O
H
CO2Et
H N
O
O
N H
H NH
CO2Et
H N
O
O
Si
O
O
O CO2Et
O O
P
O
−
B
O
O
O
8
O P
O−
O
O S H HN
H NH
N H
O
O
CO2Et
H N
O O
Si
O
O CO2Et
O P O−
O
ACAGTGACT 5′
3′
OH
O
9
Reversible Biotinylation of the 5′-Terminus of Oligodeoxyribonucleotides
Figure 4.20.3 Preparation of reversible biotinylation phosphoramidite S.7 and structures of biotinylated 5′-phosphate DNAs S.8 and S.9.
4.20.6 Supplement 14
Current Protocols in Nucleic Acid Chemistry
Sodium sulfate (Na2SO4, anhydrous) Chloroform (CHCl3) Methanol 40-µm silica gel (Baker) TLC plates: silica gel on aluminum (60F-254, 200-µm thickness) Acetonitrile (distilled over CaH2) 2-Cyanoethyl-N,N,N′,N′-tetraisopropylphosphoramidite (2-cyanoethyltetraisopropylphosphorodiamidite, 97%) 0.45 M 1H-tetrazole (99%+, sublimed) in acetonitrile 5% (w/v) sodium bicarbonate (NaHCO3) Tetrahydrofuran (THF, 99%+) Triethylamine (TEA) 50-mL one-neck, round-bottomed flasks (oven dried) Vacuum pump 20-G, 5-cm stainless steel needles 5-, 1-, and 0.5-mL graduated glass syringes Septa (14/20, 24/40) 250-mL separatory funnels Filter funnel and Whatman no. 1 filter paper Rotary evaporator equipped with a water aspirator Flash chromatography columns (3.0 × 12–cm, 3.0 × 10–cm) UV lamp Additional reagents and equipment for thin-layer chromatography (TLC, APPENDIX 3D), column chromatography (APPENDIX 3E), and incorporation of biotinylated phosphoramidite into oligonucleotide (see Basic Protocol 1) Prepare biotin diethyl bis(hydroxymethyl)malonate conjugate S.6 1. Dry 605 mg (0.93 mmol) biotinyl alcohol S.1 in a 50-mL oven-dried, one-neck, round-bottomed flask (equipped with a magnetic stir bar) under vacuum overnight at room temperature. 2. Add 63 mg (0.93 mmol) imidazole and then flush the flask with an argon flow for 10 min. 3. Add 2 mL of dry DMF and 498 µL (2.79 mmol) DIEA with a 20-G needle attached to a 5-mL and 1-mL glass syringe, respectively, and cool the mixture to 0°C. 4. Add 252 µL (1.40 mmol) diisopropyldichlorosilane with a 500-µL glass syringe in one portion. Stir the mixture at 0°C for 1 hr, then at room temperature for 4 hr on a magnetic stir plate. 5. In another 50-mL oven-dried, one-neck, round-bottomed flask, place 500 mg (2.20 mmol) diethyl bis(hydroxymethyl)malonate and 126 mg (1.86 mmol) imidazole. Seal the flask with a septum, flush with an argon flow for ∼10 min, and then add 2 mL of dry DMF with a 5-mL glass syringe. Shake to dissolve the solids. 6. Cool the solution prepared in step 5 to 0°C. Add the reaction mixture obtained in step 4 with a 5-mL glass syringe very slowly (over at least a 30-min period) and stir the resulting solution at 0°C for 5 hr. 7. Add 30 mL CH2Cl2 and 50 mL of 5% citric acid, transfer the mixture to a 250-mL separatory funnel, and separate the organic phase. 8. Extract the aqueous phase four times with 30 mL CH2Cl2. 9. Combine the organic phases, dry over ∼2 g anhydrous Na2SO4, and filter off the solids using a filter funnel and Whatman no. 1 filter paper. Concentrate the filtrate to dryness on a rotary evaporator with a water aspirator.
Synthesis of Modified Oligonucleotides and Conjugates
4.20.7 Current Protocols in Nucleic Acid Chemistry
Supplement 14
10. Dissolve the residue in ∼8 mL of 19:1 (v/v) CHCl3/methanol and apply to the top of a 3.0 × 12–cm flash chromatography column (APPENDIX 3E) prepared with a slurry of 40-µm silica gel in the same solvent system. Elute the column with the same solvent system. Monitor the fractions by TLC (APPENDIX 3D) using 9:1 (v/v) CHCl3/methanol. 11. Pool the fractions that contain the UV-active spot with an Rf = 0.5 (9:1 CHCl3/methanol), concentrate to dryness on a rotary evaporator, and then dry under vacuum. The product, (S.6), has the highest Rf value, and is the major UV-active spot in the reaction mixture under these TLC conditions. The expected yield is 92% (839 mg). 1H NMR (CDCl3, 500 MHz) δ 0.88-1.01 (m, 14H), 1.24-1.27 (m, 12H), 1.32 (s, 9H), 1.47-1.53 (m, 2H), 1.65-1.85 (m, 6H), 2.18-2.31 (m, 4H), 3.03-3.10 (m, 2H), 3.25 (dt, 1H, J = 7.4, 4.7 Hz), 3.38-3.50 (m, 4H), 3.54-3.56 (m, 4H), 3.60 (s, 4H), 4.14-4.27 (m, 9H), 5.22-5.24 (m, 1H), 6.37 (t, 1H, J = 5.5 Hz), 6.58 (t, 1H, J = 5.3 Hz), 7.39 (d, 2H, J = 8.5 Hz), 7.57 (d, 2H, J = 8.5 Hz); 13C NMR (CDCl3, 500 MHz) δ 13.4, 14.2, 17.7, 27.8, 29.8, 30.5, 31.3, 32.0, 35.1, 35.6, 38.4, 39.2, 39.5, 40.0, 55.2, 57.5, 61.6, 61.7, 61.8, 62.7,70.0, 70.1 (X 2), 70.3, 73.8, 124.7, 129.1, 131.9, 155.1, 156.3, 169.2, 170.2, 173.3, 174.2.
Prepare biotin diethyl bis(hydroxymethyl)malonate conjugate phosphoramidite S.7 12. Dry 715 mg (0.72 mmol) S.6 in a 50-mL oven-dried, one-neck round-bottomed flask (equipped with a magnetic stir bar) under vacuum overnight at room temperature. 13. Flush the flask with an argon flow for ∼5 min, then sequentially add 2 mL dry acetonitrile and 264 µL (0.80 mmol) 2-cyanoethyl-N,N,N′,N′-tetraisopropylphosphoramidite with a 5-mL and 500-µL glass syringe, respectively. 14. Add 1.68 mL (0.76 mmol) of 0.45 M 1H-tetrazole (in acetonitrile) with a 5-mL glass syringe, stir the mixture at room temperature for 3 hr, and then quench the reaction with 50 mL of 5% NaHCO3. 15. Transfer the reaction mixture to a 250-mL separatory funnel, extract the mixture five times with 30 mL CH2Cl2, and combine the organic phases. 16. Dry over ∼2 g anhydrous Na2SO4, filter off the solids, and concentrate the filtrate to dryness on a rotary evaporator. 17. Dissolve the residue in ∼5 mL of 7:3:0.5 (v/v/v) CHCl3/THF/TEA, and apply to the top of a 3.0 × 10–cm flash chromatography column prepared with a slurry of silica gel in the same solvent. Elute the column with the same solvent system and monitor the fractions by TLC. 18. Pool the fractions that contain the UV-active spot with an Rf = 0.5 (7:3:0.5 CHCl3/THF/TEA), remove solvents on a rotary evaporator, and dry under vacuum. S.7 is obtained as a white foam with an expected yield of 61% (520 mg). 1H NMR (CDCl3, 250 MHz) δ 0.91-1.07 (m, 14H), 1.14-1.18 (m, 12H), 1.22-1.29 (m, 12H), 1.32 (s, 9H), 1.41-1.51 (m, 2H), 1.65-1.82 (m, 4H), 2.16-2.34 (m, 4H), 2.60 (t, 2H, J = 6.8 Hz), 3.04-3.06 (m, 2H), 3.21-3.26 (m, 1H), 3.40-3.48 (m, 4H), 3.52-3.61 (m, 8H), 3.76-3.82 (m, 2H), 4.05-4.25 (m, 11H), 5.19-5.22 (m, 1H), 6.28 (t, 1H, J = 5.5 Hz), 6.39 (t, 1H, J = 5.3 Hz), 7.39 (d, 2H, J = 8.0 Hz), 7.57 (d, 2H, J = 8.3 Hz); 31P NMR (CDCl3, 250 MHz) δ 166.36.
Synthesize biotinylated DNA 19. Perform oligonucleotide synthesis, coupling of the biotinylated phosphoramidite S.7, and cleavage and deprotection of the biotinylated DNA (S.8) as described (see Basic Protocol 1, steps 18 to 23). Reversible Biotinylation of the 5′-Terminus of Oligodeoxyribonucleotides
For the example in Figure 4.20.3 (S.9), the sequence 3′-TCAGTGACA-5′ was synthesized on a 1-ìmol scale prior to biotinylation. The HPLC profile of crude biotinylated S.9 is illustrated in Figure 4.20.2.
4.20.8 Supplement 14
Current Protocols in Nucleic Acid Chemistry
APPLICATION OF THE REVERSIBLE BIOTINYLATION METHOD IN AFFINITY PURIFICATION USING AVIDIN-COATED MICROSPHERES
SUPPORT PROTOCOL 1
This protocol describes the attachment of the reversibly biotinylated full-length DNAs S.4 and S.8 (see Figs. 4.20.1 and 4.20.3, respectively) to NeutrAvidin-coated microspheres, the removal of nonbiotinylated failure sequences resulting from inefficient coupling and other impurities, and the recovery of pure, unmodified DNAs S.10 and S.12 (see Fig. 4.20.4) from the solid microspheres. Materials UltraLink Immobilized NeutrAvidin (1000-Å pore size; 50- to 80-µm particle size; 0.08 µmol biotin/ml gel binding capacity; Pierce), prepared as 50% (v/v) slurry containing 0.02% (w/v) sodium azide PBS (APPENDIX 2A) Biotinylated DNA (see Basic Protocol 1 for S.4 or Basic Protocol 2 for S.8) Acetone (dried over anhydrous sodium sulfate) Tetrahydrofuran (THF, distilled over sodium/benzophenone ketyl) 70:30 (v/v) hydrogen fluoride in pyridine (HF/pyridine) Methoxytrimethylsilane ∼40% (v/v) methylamine (CH3NH2; for 5′-phosphate DNA S.8 only) 10- and 1.5-mL centrifuge tubes Lyophilizer or Speedvac evaporator 1. Transfer 5 mL (for 1-µmol scale DNA synthesis) UltraLink Immobilized NeutrAvidin gel (i.e., 10 mL of 50% gel slurry) into a 10-mL centrifuge tube, centrifuge 30 sec at 2100 × g, room temperature, and remove the supernatant. 2. Resuspend the gel in 5 mL PBS, centrifuge 30 sec at 2100 × g, room temperature, and remove the supernatant. Repeat wash two additional times. 3. Dissolve 1 µmol biotinylated DNA in 2 mL PBS and transfer the solution to the centrifuge tube containing the NeutrAvidin gel. Wash the DNA tube two times with 0.5 mL PBS and add to the gel suspension. Incubate at room temperature for 1 hr with occasional gentle shaking. 4. Centrifuge the suspension 30 sec at 2100 × g, room temperature, and remove the supernatant. Wash and then dry the gel as follows: at least three times with 3 mL PBS three times with 3 mL water two times with 5 mL acetone three times with 5 mL THF. 5. Suspend the gel in 5 mL THF, add 300 µL of 70:30 (v/v) HF/pyridine with a 500-µL pipet, and incubate the suspension 1 hr at room temperature with occasional gentle shaking. The fluoride ion cleaves the two Si-O bonds in S.4 and S.8 (see Fig. 4.20.4). CAUTION: HF reacts with glass and is toxic. The reaction must be performed in a plastic centrifuge tube in a well-ventilated fume hood.
6. Add 3 mL methoxytrimethylsilane with a 1-mL pipet, incubate at room temperature for 10 min, and then centrifuge the mixture 30 sec at 2100 × g and discard the supernatant. Methoxytrimethylsilane quenches the excess HF, giving the volatile side products fluorotrimethylsilane and methanol (see Fig. 4.20.4).
Synthesis of Modified Oligonucleotides and Conjugates
4.20.9 Current Protocols in Nucleic Acid Chemistry
Supplement 14
7a. For 5′-OH DNA (from S.4): Collect pure DNA S.10 by washing the gel six times with 0.5 mL water, saving each wash. Combine water washes and dry in a Speedvac evaporator or lyophilizer. Expected recovery yield is >80%. The purity of the DNA can be checked by reversed-phase HPLC (UNIT 10.5). A typical profile for the short sequence 5′-TACAGTGACT-3′ is illustrated in Figure 4.20.2.
7b. For 5′-phosphate DNA (from S.8): Wash the gel with 1 mL THF, then add 3 mL of ∼40% methylamine and incubate at room temperature for 30 min. Centrifuge the mixture 30 sec at 2100 × g, remove the supernatant, and wash the gel seven times with 1 mL water, saving each wash. Combine the supernatant and water washes, and dry the pure DNA S.12 in a Speedvac evaporator or lyophilizer.
O O
A
S H HN
O
N H
H NH
NH
H N
O
O
Si
O
N
O
O
O
O
O
4
O P
O−
O
1. HF/pyridine 2. Me 3SiOMe
O NH HO
O S H HN
O
N H
H NH
H N
O
OH O
+
F
Si
F
+
Me3SiF
+
O
O O−
O P
O
N
O
O
10
O
B
S H HN
O
N H
H NH
CO2Et
H N
O
O
Si
O
O
O
O
O
P
O
O
8
O P
CO2Et HO O H HN
O
N H
H NH
H N
O
OH
+
F
O−
O
1. HF/pyridine 2. Me 3SiOMe
S
B
O
O−
CO2Et
Si
F
O P
O
+
CO2Et
O−
Me3SiF
+
O
O
O
B
O O P
O−
O O CO2Et HO
O CO2Et
O P
O O
O
O−
−O
B
~40% MeNH2
P
O O P
11
Reversible Biotinylation of the 5′-Terminus of Oligodeoxyribonucleotides
O
O
O
O−
B
O O−
O P
12
O−
O
Figure 4.20.4 Removal of biotin from biotinylated DNAs S.4 and S.8 to generate unmodified 5′-OH DNA S.10 and unmodified 5′-phosphate DNA S.12, respectively.
4.20.10 Supplement 14
Current Protocols in Nucleic Acid Chemistry
O
2,2 -(ethylenedioxy)bis(ethylamine)
1. CH3MgBr
OH
O
O
2. AcOH
O levulinic acid
14 (75%)
O
tert-butylchlorodiphenylsilane
H NH
OH O
O OH
H N
O
13 (61%)
S H HN
O
H 2N
S H HN
DMAP
Ph O
H NH
Si
CMe3
1. 4-(tert-butyl)benzoyl chloride
Ph
2. K2CO3
O
O
biotin
15
O S H
OH
isobutyl chloroformate
H NH
N O
O S H
DIEA
O
O
16 (76%)
17
O S
14 , DMAP
O H NH
N O
O
O
H
H NH
N O
O
N H
O
O
H N
OH O
1 (66%)
Figure 4.20.5 Preparation of the biotinyl alcohol S.1.
Aqueous methylamine removes the 5′-tag in S.11 to generate the 5′-phosphate DNA S.12 (see Fig. 4.20.4). Expected recovery yield is >70%. The purity of the DNA can be checked by reversed-phase HPLC (UNIT 10.5). A typical profile for the short sequence 5′-H2O3POACAGTGACT-3′ is illustrated in Figure 4.20.2.
SYNTHESIS OF BIOTINYL ALCOHOL The biotinyl alcohol S.1 is required for the preparation of the reversible biotinylation phosphoramidites S.3 and S.7. This protocol describes its preparation, which is shown in Figure 4.20.5. Additional Materials (also see Basic Protocol 1) Levulinic acid (98%) Dry ice/acetone 3.0 M methyl magnesium bromide in ether Acetic acid (99.7%+) Magnesium sulfate (MgSO4, anhydrous)
SUPPORT PROTOCOL 2
Synthesis of Modified Oligonucleotides and Conjugates
4.20.11 Current Protocols in Nucleic Acid Chemistry
Supplement 14
2,2′-(Ethylenedioxy)bis(ethylamine) (98%) Ether Triethylamine Ninhydrin Ethanol (anhydrous) Biotin (AnaSpec) 4-(Dimethylamino)pyridine (DMAP) Pyridine (distilled from CaH2) t-Butylchlorodiphenylsilane (98%) 4-t-Butylbenzoylchloride (98%) Potassium carbonate (K2CO3) 5% citric acid Isobutylchloroformate (98%) 100-, 20-, 10-, 5-, and 1-mL graduated glass syringes 1000-mL two-neck round-bottomed flask, oven dried Condenser 500- and 250-mL separatory funnels Aldrich short-path distillation apparatus 50-, 100- and 250-mL one-neck round-bottomed flasks, oven dried Flash chromatography columns (4.5 × 30–cm and 4.5 × 10–cm) Heat gun Rubber septum 2- and 6-cm stainless steel needles Prepare lactone S.13 1. Add 14 mL (300 mmol) levulinic acid and 300 mL THF with a 20-mL and 100-mL glass syringe, respectively, to a 1000-mL oven-dried, two-neck, round-bottomed flask equipped with a magnetic stir bar, connected to an argon flow through a condenser. 2. Cool the flask to −78°C with a slurry of dry ice in acetone. Slowly add 100 mL (300 mmol) of 3.0 M methyl magnesium bromide in ether with a 100-mL glass syringe. Stir the mixture for 3 hr while gradually warming to room temperature, then heat to 50°C overnight. This should form a light yellow solution.
3. Cool the solution to room temperature, add ∼100 mL acetic acid with a disposable pipet, and stir the mixture for ∼12 hr. At this stage, maintaining an anhydrous atmosphere is unnecessary.
4. Add 40 mL water and then remove volatile components (ether and THF) on a rotary evaporator. 5. Transfer the red residue to a 500-mL separatory funnel and extract three times with 100 mL (each extraction) CH2Cl2. 6. Combine the organic phases, dry over ∼5 g anhydrous MgSO4, and filter off the solids. Concentrate the filtrate on a rotary evaporator. 7. Distill the residue by means of a short-path distillation apparatus using a vacuum of ∼0.6 mmHg. Collect the second fraction, which typically distills at 70°C (∼0.6 mmHg). Reversible Biotinylation of the 5′-Terminus of Oligodeoxyribonucleotides
The product 5,5-dimethyl-dihydrofuran-2-one (S.13) should be a colorless oil with an expected yield of 61% (9.5 g). 1H NMR (CDCl3, 250 MHz): δ 1.44 (s, 6H), 2.06 (t, 2H, J = 8.23 Hz), 2.63 (t, 2H, J = 8.3 Hz).
4.20.12 Supplement 14
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Prepare amino alcohol S.14 8. Place 24.3 mL (166 mmol) 2,2′-(ethylenedioxy)bis(ethylamine), 9.5 g (83.2 mmol) S.13, and 10 mL water in a 100-mL one-neck, round-bottomed flask equipped with a condenser and a magnetic stir bar. Stir the mixture overnight at 90°C. This should give a light yellow solution.
9. Remove water and excess 2,2′-(ethylenedioxy)bis(ethylamine) by distilling at ∼70°C at 0.6 mmHg under vacuum as in step 7. 10. Dissolve the light yellow oily residue in 15 mL of 5:2:2:1 (v/v/v/v) ether/methanol/acetonitrile/triethylamine and apply the solution to the top of a 4.5 × 30–cm flash chromatography column prepared using a slurry of silica gel in the same solvent mixture. Elute the column and monitor the fractions by TLC using the same solvent mixture. Detect the product by rinsing the TLC plate briefly in a solution of 0.5 g ninhydrin in 200 mL ethanol and heat with a heat gun until purple spots appear (∼45 sec). The product should appear as a purple spot with an Rf = 0.8.
11. Pool the fractions containing the pure product, evaporate solvents on a rotary evaporator, and dry the light yellow oily product under vacuum. The expected yield of the product (S.14) is 75% (16.3 g). 1H NMR (CD3OD, 500 MHz): δ 1.20 (s, 6H), 1.78 (t, 2H, J = 8.1 Hz), 2.35 (t, 2H, J = 7.5 Hz), 2.88 (t, 2H, J = 4.5 Hz), 3.89-3.42 (m, 2H), 3.54-3.57 (m, 4H), 3.62 (s, 4H); 13C NMR (CD3OD, 500 MHz): δ 28.8, 31.0, 38.5, 38.8, 40.8, 69.0, 69.3, 69.6, 69.7, 72.0, 174.0.
Prepare N-t-butylbenzoylbiotin S.16 12. Place 2.44 g (10.0 mmol) biotin and 0.61 g (5.0 mmol) DMAP in a 250-mL oven-dried, one-neck, round-bottomed flask equipped with a magnetic stir bar and capped with a rubber septum. Connect the flask to an argon flow with a 2-cm stainless steel needle and flush the flask with argon for ∼5 min. 13. Sequentially add 15 mL dry pyridine and 3.0 mL (15 mmol) t-butylchlorodiphenylsilane with a 20- and 5-mL glass syringe, repectively, and stir the mixture overnight at room temperature. t-Butylchlorodiphenylsilane is used for temporary protection of the carboxylic acid group to form the intermediate S.15.
14. Add 3.0 mL (15 mmol) 4-t-butylbenzoylchloride with a 5-mL glass syringe, stir the mixture for 3 hr, and then quench the reaction by adding 2 mL methanol. 15. Remove volatile components on a rotary evaporator. Dissolve the residue in a mixture of the following: 12 mL THF 6 mL methanol 6 mL water 6.9 g (50 mmol) K2CO3. Stir 30 min at room temperature. This mild basic condition removes the t-butyldiphenylsilane group without affecting the 4-t-butylbenzoyl protecting group, to give S.16.
16. Quench the reaction with 50 mL of 5% citric acid. Extract the mixture six times with 20 mL ethyl acetate. Combine the organic extracts, dry over ∼2 g anhydrous MgSO4, filter off the solids, and concentrate the filtrate to dryness on a rotary evaporator.
Synthesis of Modified Oligonucleotides and Conjugates
4.20.13 Current Protocols in Nucleic Acid Chemistry
Supplement 14
17. Dissolve the residue in ∼4 mL CHCl3, place the solution on top of a 4.5 × 10–cm flash chromatography column, and elute first with ∼350 mL CHCl3 and then with 19:1 (v/v) CHCl3/methanol. Monitor the fractions by TLC using 9:1:10 (v/v/v) CHCl3/methanol/ether as the solvent system. 18. Pool fractions that contain the UV-active spot with an Rf = 0.5 (9:1:10 CHCl3/methanol/ether), concentrate to dryness on a rotary evaporator, and dry under vacuum. The expected yield of the product 1-(4-tert-butylbenzoyl)-biotin (S.16) is 76% (3.07 g). 1H NMR (CD3OD, 500 MHz) δ 1.15 (s, 9H), 1.27-1.63 (m, 6H), 2.14 (t, 2H, J = 7.4 Hz), 2.79-2.91 (m, 2H), 3.11-3.15 (m, 1H), 4.07 (dd, 1H, J = 4.6, 7.8 Hz), 4.98-5.00 (m, 1H), 7.25 (d, 2H, J = 8.5 Hz), 7.31 (d, 2H, J = 8.6 Hz); 13C NMR (CD3OD, 500 MHz) δ 26.1, 29.6, 29.9, 31.7, 34.8, 35.9, 38.9, 56.7, 59.3, 64.1, 125.7, 129.8, 133.8, 156.1, 158.2, 171.9, 177.6.
19. Place 1.9 g (4.8 mmol) S.16 in a 100-mL one-neck, round-bottomed flask equipped with a magnetic stir bar. Then place 292 mg (2.39 mmol) DMAP and 656 mg (2.5 mmol) S.14 in a separate 50-mL one-neck, round-bottomed flask equipped with a stir bar. Dry both on a vacuum pump overnight at room temperature. Prepare biotinyl alcohol S.1 20. Stop the neck of the flask containing S.16 with a rubber septum. Flush the flask with an argon flow with two 6-cm stainless steel needles (one inlet and one outlet) for ∼10 min and then remove the outlet needle. 21. Place the flask in an ice bath and sequentially add 4 mL dry DMF and 920 µL (5.28 mmol) DIEA with a 5-mL and 1-mL glass syringe, respectively. Let sit 10 min. 22. Add 684 µL (5.27 mmol) isobutylchloroformate with a 1-mL glass syringe and stir the mixture 1 hr at 0°C. This forms the intermediate S.17.
23. Flush the flask containing DMAP and S.14 (step 19) with an argon flow for 10 min, then add 4 mL dry DMF with a 5-mL glass syringe. 24. Very slowly add the solution of DMAP and S.14 to the solution of intermediate S.17 with a 10-mL glass syringe (at least over a 15-min period). After addition, stir the mixture 10 min at 0°C. 25. Quench the reaction with 2 mL water. Add 20 mL of 5% citric acid, transfer the mixture to a 250-mL separatory funnel, and extract six times with 30 mL (each extraction) ethyl acetate. 26. Pool the extracts, dry over ∼2 g anhydrous Na2SO4, filter off the solids, and evaporate the filtrate on a rotary evaporator. 27. Dissolve the residue in ∼10 mL of 19:1 (v/v) CH2Cl2/methanol and apply to the top of a 4.5 × 10–cm flash chromatography column prepared with a slurry of silica gel in the same solvent. Elute the column with the same solvent and monitor the fractions by TLC using a solvent system of 9:1 (v/v) CH2Cl2/methanol. 28. Collect the fractions that contain a UV-active spot with an Rf = 0.5 (9:1 CH2Cl2/methanol), pool these fractions, and evaporate to dryness on a rotary evaporator to give S.1.
Reversible Biotinylation of the 5′-Terminus of Oligodeoxyribonucleotides
Under vacuum, biotinyl alcohol (S.1) can form a white foam. Expected yield is 66% (2.1 g). 1H NMR (CD3OD, 500MHz) δ 1.24 (s, 6H), 1.38 (s, 9H), 1.46-1.55 (m, 2H), 1.65-1.85 (m, 6H), 2.28 (t, 2H, J = 7.3 Hz), 2.34 (t, 2H, J = 7.7 Hz), 3.06-3.13 (m, 2H), 3.36-3.43 (m, 5H), 3.55-3.66 (m, 8H), 4.31 (t, 1H, J = 6.9 Hz), 5.22 (t, 1H, J = 6.6 Hz), 7.48 (d, 2H, J = 8.2 Hz), 7.54 (d, 2H, J = 8.1 Hz); 13C NMR (CD3OD, 500 MHz) δ 26.9, 29.3, 29.5, 29.9, 31.8, 32.4, 35.9, 36.8, 38.9, 40.3, 40.4, 40.4, 56.7, 59.3, 64.0, 70.7, 70.7, 70.9, 71.4, 79.6, 125.6, 129.8, 133.7, 156.1, 158.1, 171.8, 176.1, 176.6.
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COMMENTARY Background Information The strong noncovalent highly specific interaction between biotin and streptavidin or avidin (association constant = 1015/M) has found numerous applications in areas such as immunology, cell biology, and molecular biology (McInnes and Symons, 1989). Consequently, many methods have been developed for biotinylation of various target molecules. Biotinylation of DNA has been achieved either enzymatically (Langer et al., 1981) or chemically. Site-specific chemical biotinylation of DNA can be performed manually after solidphase synthesis, cleavage, and complete deprotection (Agrawal et al., 1986; Urdea et al., 1988; Gildea et al., 1990; De Vos et al., 1994) or automatically by the synthesizer during solidphase synthesis (Pon, 1991; Neuner, 1996; Olejnik et al., 1996). For the latter method, which is more convenient to perform, biotinylation reagents that are compatible with DNA synthesis, cleavage, and deprotection conditions are required. For some applications, releasing the target DNA from streptavidin or avidin may be necessary. For this purpose, cleavable linkers have been incorporated in between biotin and target DNA to avoid the harsh conditions required for breaking the biotin-streptavidin/avidin bonding. For example, a disulfide bond has been incorporated into the linker, which can be broken by reduction with dithiothreitol (Shimkus et al., 1985; Dawson et al., 1991). The drawback of this method is that part of the linker is left on the oligonucleotide, therefore modifying the oligonucleotide, which may not be further processed under biological and biochemical conditions. Another method is to incorporate an acid-labile linker in between biotin and DNA (Gildea et al., 1990). Although the DNA is not modified after cleavage from biotin, the synthesis of the reagent for biotinylation is complicated, and biotinylation is performed manually. A third method uses a photocleavable linkage between biotin and DNA (Olejnik et al., 1996). Using this method, biotinylation is performed automatically by the synthesizer during solid-phase synthesis and, after cleavage, unmodified DNA is obtained. The unsatisfactory feature of this method is that target DNA may be damaged by UV irradiation during photocleavage (Cadet and Vigny, 1990; Greenberg and Gilmore, 1994; Greenberg, 1995).
The biotinylation methods described in this unit use a fluoride-cleavable diisopropyl silyl acetal linkage between biotin and DNA (Fang and Bergstrom, 2003a,b). This method has several advantages over known methods. The biotinylation is automatically carried out by the synthesizer during solid-phase synthesis, no damage of target DNA during cleavage should occur, and, after cleavage, both unmodified 5′-OH (Fang and Bergstrom, 2003b) and unmodified 5′-phosphate DNA (Fang and Bergstrom, 2003a) can be obtained depending on which of the two phosphoramidites is used (see Figs. 4.20.1, 4.20.3, and 4.20.4). One of the potential applications of this reversible biotinylation method is affinity purification of synthetic oligonucleotides mediated by biotin-avidin/streptavidin binding, as demonstrated in Support Protocol 1. The biotinylated full-length DNA can be readily attached to the NeutrAvidin-coated microspheres, nonbiotinylated failure sequences and other impurities can be removed by simply washing with buffer and water, and high-quality unmodified DNA can be obtained by simple incubation with fluoride ions (followed by aqueous amine treatment in the case of 5′-phosphorylated DNA; Guzaev et al., 1995) under mild conditions, followed by a water wash (see Fig. 4.20.4). Currently, synthetic DNA is generally purified by HPLC and/or gel electrophoresis, which are considered time consuming. DMTr-selective cartridge purification is convenient, but the quality of DNA thus obtained is less satisfactory (UNIT 10.7). The affinity purification method described in this unit is convenient, and high quality DNA can be obtained. It is anticipated that the advantages of this method will be more obvious when it is required for purification of very long DNA sequences and/or isolation of target DNA from very complex mixtures.
Critical Parameters and Troubleshooting For the preparation of the two phosphoramidites (S.3 and S.7, see Basic Protocols 1 and 2 and Support Protocol 2), many steps must be carried out under strict anhydrous conditions. When this is the case, make certain that glassware and reagents, especially solvents, are dry. Although some reactions can be performed in open air, the authors recommend all transformations be performed under an argon or nitrogen atmosphere.
Synthesis of Modified Oligonucleotides and Conjugates
4.20.15 Current Protocols in Nucleic Acid Chemistry
Supplement 14
The oligonucleotide to be biotinylated must be synthesized using acetyl-protected C rather than benzoyl-protected C; otherwise, transamination may occur during the aqueous CH3NH2/NH4OH cleavage/deprotection (Reddy et al., 1994; Wincott et al., 1995). When using the biotinylation method for affinity purification of synthetic oligonucleotides, make sure to handle the NeutrAvidincoated microspheres gently. Vortexing of the gel should be avoided, as this may result in leaking of gel material and low recovery yield of full-length DNA.
Anticipated Results Yields reported for each step during the preparation of the phosphoramidites S.3 and S.7 may vary depending on how carefully the reaction, work-up, and purification are performed. Since these reactions can be easily performed on a gram scale, it is not difficult to obtain sufficient amounts of S.3 and S.7 for biotinylating several batches of DNA. The yield for biotinylation of the 5′-end of DNA on solid support is >99% in each case. For affinity purification by a NeutrAvidin-coated gel, a recovery yield of full-length unmodified DNA can be >70% in both cases.
Time Considerations The preparation of the phosphoramidites S.3 or S.7 requires ∼13 days including drying glassware and reagents, setting up and working up reactions, and purifying and drying products. DNA synthesis, 5′-biotinylation, and postsynthetic cleavage/deprotection can be achieved in 1 or 2 days depending on the length of the sequence. Affinity purification can be carried out within 6 hr, but drying the product may require ∼12 hr.
Literature Cited Agrawal, S., Christodoulou, C., and Gait, M.J. 1986. Efficient methods for attaching non-radioactive labels to the 5′ ends of synthetic oligodeoxyribonucleotides. Nucl. Acids Res. 14:6227-6245. Cadet, J. and Vigny, P. 1990. The photochemistry of nucleic acids. In Bioorganic Photochemistry (H. Morrison, ed.) Vol. 1, pp. 170-184. John Wiley & Sons, New York.
Reversible Biotinylation of the 5′-Terminus of Oligodeoxyribonucleotides
Dawson, B.A., Herman, T., Haas, A.L., and Lough, J. 1991. Affinity isolation of active murine erythroleukemia cell chromatin: Uniform distribution of ubiquitinated histone H2A between active and inactive fractions. J. Cell. Biochem. 46:166-173.
De Vos, M.J., Van Elsen, A., and Bollen, A. 1994. New non-nucleosidic phosphoramidites for the solid phase multi-labeling of oligonucleotides: Comb- and multifork-like structure. Nucleosides Nucleotides 13:2245-2265. Fang, S. and Bergstrom, D.E. 2003a. Reversible biotinylation phosphoramidite for 5′-end-labeling, phosphorylation and affinity purification of synthetic oligonucleotides. Bioconjugate Chem. 14:80-85. Fang, S. and Bergstrom, D.E. 2003b. Fluoridecleavable biotinylation phosphoramidite for 5′end-labeling, affinity purification of synthetic oligonucleotides. Nucl. Acids Res. 31:708-715. Gildea, B.D., Coull, J.M., and Koster, H. 1990. A versatile acid-labile linker for modification of synthetic biomolecules. Tetrahedron Lett. 31:7095-7098. Greenberg, M.M. 1995. Photochemical release of protected oligodeoxyribonucleotides containing 3′-glycolate termini. Tetrahedron 51:29-38. Greenberg, M.M. and Gilmore, J.L. 1994. Cleavage of oligonucleotides from solid-phase supports using o-nitrobenzyl photochemistry. J. Org. Chem. 59:746-753. Guzaev, A., Salo, H., Azhayev, A., and Lönnberg, H. 1995. A new approach for chemical phosphorylation of oligonucleotides at the 5′-terminus. Tetrahedron 51:9375-9384. Langer, P.R., Waldrop, A.A., and Ward, D.C. 1981. Enzymatic synthesis of biotin-labeled polynucleotides: Novel nucleic acid affinity probes. Proc. Natl. Acad. Sci. U.S.A. 78:6633-6637. McInnes, J.L. and Symons, R.H. 1989. Preparation and detection of nonradioactive nucleic acid and oligonucleotide probes. In Nucleic Acid Probes (R.H. Symons, ed.) pp. 33-80. CRC Press, Boca Raton, Fla. Neuner, P. 1996. New non nucleosidic phosphoramidite reagent for solid phase synthesis of biotinylated oligonucleotides. Bioorg. Med. Chem. Lett. 6:147-152. Olejnik, J., Krzymanska-Olejnik, E., and Rothschild, K.J. 1996. Photocleavable biotin phosphoramidite for 5′-end-labelling, affinity purification and phosphorylation of synthetic oligonucleotides. Nucl. Acids Res. 24:361-366. Pon, R.T. 1991. A long chain biotin phosphoramidite reagent for the automated synthesis of 5′-biotinylated oligonucleotides. Tetrahedron Lett. 32:1715-1718. Reddy, M.P., Hanna, N.B., and Farooqui, F. 1994. Fast cleavage and deprotection of oligonucleotides. Tetrahedron Lett. 35:4311-4314. Shimkus, M., Levy, J., and Herman, T. 1985. A chemically cleavable biotinylated nucleotide: Usefulness in the recovery of protein-DNA complexes from avidin affinity columns. Proc. Natl. Acad. Sci. U.S.A. 82:2593-2597.
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Urdea, M.S., Warner, B.D., Running, J.A., Stempien, M., Clyne, J., and Horn, T. 1988. A comparison of non-radioisotopic hybridization assay methods using fluorescent, chemiluminescent and enzyme labeled synthetic oligodeoxyribonucleotide probes. Nucl. Acids Res. 16:49374956. Wincott, F., DiRenzo, A., Shaffer, C., Grimm, S., Tracz, D., Workman, C., Sweedler, D., Gonzalez, C., Scaringe, S., and Usman, N. 1995. Synthesis, deprotection, analysis and purification of RNA and ribozymes. Nucl. Acids Res. 23:2677-2684.
Contributed by Shiyue Fang and Donald E. Bergstrom Purdue University West Lafayette, Indiana Walther Cancer Institute Indianapolis, Indiana
Synthesis of Modified Oligonucleotides and Conjugates
4.20.17 Current Protocols in Nucleic Acid Chemistry
Supplement 14
Uridine 2′-Carbamates: Facile Tools for Oligonucleotide 2′-Functionalization
UNIT 4.21
This unit contains procedures for synthesis of uridine 2′-carbamate phosphoramidites and oligonucleotides thereof. 3′,5′-Silyl-diprotected uridine can be converted into the corresponding 2′-carbamate by reaction with 1,1′-carbonyldiimidazole followed by treatment with an aliphatic amine. The 2′-carbamate can then be converted in several steps into a 3′-phosphoramidite suitable for machine-assisted oligonucleotide synthesis. The preparation of eleven different uridine 2′-carbamates and their phosphoramidites from several primary and secondary amines is described in the first two methods (see Basic Protocol 1 and Alternate Protocol). Their use in oligonucleotide synthesis is then described (see Basic Protocol 2). 2′-Carbamate modification is stable to conditions of standard phosphoramidite oligonucleotide synthesis. Although 2′-carbamate modification is somewhat destabilizing for DNA-DNA and DNA-RNA duplexes, it is suitable for the direction of ligands into the minor groove or into non-base-paired sites (e.g., loops, bulges) of oligo- and polynucleotides. Pyrene-modified oligonucleotide 2′-carbamates show a considerable increase in fluorescence intensity upon hybridization to a complementary RNA, and have interesting binding properties when hybridized to a mismatched DNA. CAUTION: Carry out all operations involving organic solvents and reagents in a wellventilated fume cupboard, and wear gloves and protective glasses. PREPARATION OF URIDINE 2′-CARBAMATE PHOSPHORAMIDITES FROM PRIMARY AND SECONDARY AMINES Nucleoside 2′-carbamates have been used previously in the syntheses of various modified nucleosides (McGee et al., 1996; Zhang et al., 2003) and oligonucleotides (Freier and Altmann, 1997; Seio et al., 1998; Dubey et al., 2000; Prhavc et al., 2001). The present protocol is based primarily on Korshun et al. (2002) and is illustrated in Figure 4.21.1. St able 2′-O-(imidazol-1-ylcarbonyl)-3′,5′-O-(tetraisopropyldisiloxan-1,3-diyl)uridine (S.3) is first prepared from uridine (S.1). It is then treated with primary or secondary aliphatic amines to give uridine 2′-carbamates (S.4) in high yield. Preparation of 2′-carbamates bearing a variety of N-substituents is described (S.4a-f; see Fig. 4.21.2). After 3′,5′-O-deprotection with triethylamine trihydrofluoride, 5′-O-dimethoxytritylation, and 3′-O-phosphinylation with bis(N,N-diisopropylamino)-2-cyanoethoxyphosphine, the corresponding phosphoramidites (S.7a-f) are obtained. They are used in machine-assisted synthesis of modified oligodeoxynucleotides containing uridine-2′-carbamate residues bearing these N-substituents. Strategic planning Formation of 2′-carbamates. In many cases, the imidazolide S.3 need not be isolated, but instead can be reacted in situ with an excess of the appropriate amine in dry dichloromethane (CH2Cl2). In the case of an amine hydrochloride (e.g., for S.4d), 1.1 eq triethylamine (TEA) or N,N-diisopropylethylamine (DIPEA) is added to the reaction mixture to liberate the free base. Reaction times differ vastly from one amine to another. Whilst propargylamine reacts rapidly in CH2Cl2 (<1 hr), N-methylpropargylamine, a secondary amine, requires overnight reaction. 2-Aminomethyl-15-crown-5 reacts very slowly in CH2Cl2. To accelerate the reaction, a change of solvent to acetonitrile (CH3CN) and a temperature increase to 55°C is advised. Thus, two general procedures for the preparation of carbamates S.4a-d versus S.4e-f are presented below. 3′,5′-O-Silyl-protected uridine 2′-carbamates are then isolated by column chromatography. Contributed by Vladimir A. Korshun, Dmitry A. Stetsenko, and Michael J. Gait Current Protocols in Nucleic Acid Chemistry (2003) 4.21.1-4.21.26 Copyright © 2003 by John Wiley & Sons, Inc.
BASIC PROTOCOL 1
Synthesis of Modified Oligonucleotides and Conjugates
4.21.1 Supplement 15
O NH HO
O OH
O
O
N
NH
NH
O
O O
OH
N
O
Si O
Si
O
O O
OH
Si
O
N
O 2
3
O
O NH
O
N
O
Si O
O
HO
N
O
NH
O
DMTrO
N
O
R
N
O
O NH
R Si
O
N
O
1
O
N
O
Si
OH
O
R
N
R1
R1
O
OH
5a-f
O
N R1
O
4a-f
O
O 6a-f
O NH DMTrO
N
O
O R
i-Pr2N
P
O
O
N R1 O
NCCH2CH2O 7a-f
Figure 4.21.1 General scheme for conversion of uridine into 2′-carbamate 3′-phosphoramidites (see Basic Protocol 1). For R and R1 groups, see Figure 4.21.2.
Desilylation. Tetrabutylammonium fluoride trihydrate (TBAF) in tetrahydrofuran (THF) was found unsuitable for removal of the Markiewicz protecting group from S.4, because of the observed 2′-3′ migration of the carbamate group under alkaline conditions (Korshun et al., 2002). Triethylamine trihydrofluoride, a nonbasic fluoride source, gave smooth silyl deprotection without any migration. Aromatic uridine 2′-carbamates S.5c and S.5d are crystalline solids. The other S.5 compounds are viscous oils, but are nevertheless suitable for 5′-dimethoxytritylation without further purification.
Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
Dimethoxytritylation. No 2′-3′ migration is detected when pyridine is used as a solvent for 5′-dimethoxytritylation or as an additive during column chromatography instead of TEA. The pyrene derivative S.6d and the dipeptide derivative S.6f show the highest propensity towards isomerization during column chromatography in the presence of TEA. Generally, 5′-O-DMTr-2′-carbamates S.6 show fast 2′-3′ migration under strong basic conditions, but slow migration rates in the presence of protic solvents. The 2′- and 3′-isomers are usually easily distinguishable by TLC. Column-purified 5′-O-DMTr-derivatives (S.6) are stable as dry amorphous solids for at least 12 months. Usually,
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Current Protocols in Nucleic Acid Chemistry
co-evaporation with dry CH2Cl2 is a convenient way to remove traces of other solvents; however, it must be thoroughly vented off to give S.6 as a foam. Phosphinylation. 5′-O-DMTr-uridine 2′-carbamates S.6 are phosphinylated with bis(N,Ndiisopropylamino)-2-cyanoethoxyphosphine in dry CH2Cl2 in the presence of diisopropylammonium tetrazolide to give phosphoramidites S.7, which are then isolated by column chromatography. None of the starting carbamates gave rise to isomerization products during the phosphinylation reaction. Compound characterization. Chemical characterization data are provided for all compounds. NMR spectra were recorded on a 300-MHz Bruker DRX300 NMR spectrometer in DMSO-d6 unless otherwise stated. Chemical shifts (δ) are given in ppm and referenced to tetramethylsilane as an internal standard and 85% phosphoric acid (H3PO4) as an external standard. Coupling constants (J) are given in Hertz and refer to apparent multiplicities. D2O exchange was carried out on all samples.
Compound
R
4a-7a
H
R2
4b-7b
CH3
R2
4c-7c
H
4d-7d
H
R1
I R2
R2
O
O
4e-7e
H
O
R2
O
O
O
4f-7f
4g-7g
H
4h-7h
H
4i-7i
H
4j-7j
R2
H
H
COCF3 N
R2
R2
O
COCF3 N
R2
R2
NH2
N H
O
O
NHCOCF3
N COCF3
O
O
NHCOCF3
NHFmoc S S
H N
O
O
NHCOCF 3
O
4l-7l
H
R2
O
O O
Figure 4.21.2 Side-chain substituents of uridine 2′-carbamates. R2 stands for the rest of the molecule; see Figures 4.21.1 and 4.21.3.
Synthesis of Modified Oligonucleotides and Conjugates
4.21.3 Current Protocols in Nucleic Acid Chemistry
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Materials Uridine (S.1), 99% pure Anhydrous pyridine, 99.8% pure (Aldrich) Nitrogen (or argon) gas Markiewicz reagent: 1,3-dichloro-1,1,3,3-tetraisopropyldisiloxane, 96% pure (Lancaster) Ethyl acetate (EtOAc), HPLC grade 5% (w/v) sodium hydrogencarbonate (NaHCO3) Sodium sulfate (Na2SO4), anhydrous Toluene, analytical grade Silica gel: 0.040- to 0.063-mm Macherey-Nagel Kieselgel 60 Chloroform (CHCl3), HPLC grade, ethanol free Anhydrous dichloromethane (CH2Cl2), distilled from powdered CaH2 (Fisher) Hexane, HPLC grade 1,1′-Carbonyldiimidazole, 95% pure (Sigma) Amine for S.3 conversion (select one): Propargylamine (for S.4a) N-Methylpropargylamine (for S.4b) 4-Iodobenzylamine (Lancaster; for S.4c) 1-Pyrenemethylamine hydrochloride (Aldrich; for S.4d) 2-Aminomethyl-15-crown-5 (for S.4e) H-Leu-Phe-NH2 hydrochloride (for S.4f) Triethylamine (TEA), ≥99% pure Acetonitrile (CH3CN), HPLC grade (optional; for S.4e and S.4f) N,N-Diisopropylethylamine (DIPEA), ≥99% pure (Aldrich; optional; for S.4f) 5% (w/v) citric acid Methanol (MeOH), analytical grade Acetone, analytical grade Dry tetrahydrofuran (THF), freshly distilled from LiAlH4 (store over 4A molecular sieves under nitrogen) Triethylamine trihydrofluoride Absolute ethanol, analytical grade (optional; for S.5c and S.5d) Diethyl ether, analytical grade (optional; for S.5c and S.5d) 4,4′-Dimethoxytrityl chloride (DMTr⋅Cl, 95% pure; Avocado Research Chemicals) Diisopropylammonium tetrazolide Bis(N,N-diisopropylamino)-2-cyanoethoxyphosphine, 98% pure (Fluka) 20% (w/v) sodium chloride (NaCl) Rotary evaporator equipped with a water aspirator 4 × 20–cm sintered glass chromatography column, porosity 3 Vacuum oil pump TLC plate: silica-coated aluminum plate with fluorescent indicator (Merck silica gel 60 F254) 254-nm UV lamp 30-mL screw-top Teflon bottle (Nalgene) Additional reagents and equipment for thin-layer chromatography (TLC, APPENDIX 3D) and column chromatography (APPENDIX 3E)
Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
4.21.4 Supplement 15
Current Protocols in Nucleic Acid Chemistry
Protect 3′- and 5′-hydroxy groups of uridine 1. Co-evaporate 3.663 g (15 mmol) uridine (S.1) twice with 30 mL anhydrous pyridine using a rotary evaporator equipped with a water aspirator, and then apply a dry nitrogen (or argon) atmosphere. 2. Dissolve the residue in 60 mL pyridine with magnetic stirring and rapidly add 5.0 g (15.8 mmol) Markiewicz reagent in one portion. Stopper the flask and continue stirring overnight. 3. Dilute the mixture with 300 mL EtOAc. Wash twice with 150 mL water and then once with 150 mL of 5% (w/v) NaHCO3. 4. Dry over anhydrous Na2SO4 and filter off the drying agent. Evaporate the solution to dryness and then co-evaporate three times with 40 mL toluene. 5. Pack a 4 × 20–cm sintered glass column with silica gel in CHCl3. Load the sample and elute minor side products with 1:9 (v/v) EtOAc/CHCl3. Elute desired compound (S.2) with 1:2 (v/v) EtOAc/CHCl3. 6. Evaporate fractions containing the product and co-evaporate three times with 40 mL anhydrous CH2Cl2. 7. Dry the resulting white foam 2 hr in vacuo (0.05 to 0.5 Torr). 8. Grind the foam with a stainless steel spatula and dry the resulting powder 12 to 16 hr using a vacuum oil pump. Make sure traces of chlorinated solvent are completely removed from the compound, otherwise some decomposition may occur, caused by the HCl traces generated.
9. Characterize the product by TLC (on silica gel; APPENDIX 3D) and 1H NMR. The compound is stable at least 12 months during storage at ambient temperature. 3′,5′-O-(Tetraisopropyldisiloxan-1,3-diyl)uridine (S.2). Yield of white amorphous solid 6.897 g (94%). Rf: 0.35 (1:1 v/v EtOAc/hexane). 1H NMR: 11.33 (s, 1H, H-3, exchangeable with D2O), 7.68 (d, 1H, J5,6 = 8.1 Hz, H-6), 5.55–5.50 (m, 3H, H-5, H-1′, 2′-OH), 4.16 (m, 1H, 2J = 8.6 Hz, J4,5′a = 4.9 Hz, H-5′a), 4.12 (m, 2H, H-4′, H-5′b), 3.97 (m, 1H, H-2′), 3.92 (dd, 1H, J2′,3′ = 2.5 Hz, J3′,4′ = 13.1 Hz, H-3′), 1.09–0.90 (m, 28H, Pri).
Activate 2′-hydroxy group 10. Dissolve 2.434 g (5.0 mmol) S.2 in 50 mL dry CH2Cl2 and add 852 mg (5.25 mmol) 1,1′-carbonyldiimidazole in one portion. This solution can be used directly for unhindered primary and secondary amines (step 16a). For hindered primary amines (step 16b), the product should be isolated first (steps 11 to 14).
11. Monitor reaction by TLC in 1:1 (v/v) EtOAc/hexane. The starting compound S.2 (Rf = 0.35) usually disappears after 0.5 to 2 hr.
12. Wash the solution twice with 50 mL water and dry over Na2SO4. 13. Filter off the Na2SO4, evaporate the solvent in vacuo, and co-evaporate the residue three times with 40 mL dry CH2Cl2 using a rotary evaporator. 14. Dry the residue in vacuo overnight using an oil pump. Synthesis of Modified Oligonucleotides and Conjugates
4.21.5 Current Protocols in Nucleic Acid Chemistry
Supplement 15
15. Characterize the compound by TLC and 1H NMR. 2′-O-(Imidazolylcarbonyl)-3′,5′-(tetraisopropyldisiloxan-1,3-diyl)uridine (S.3) is obtained as a white crystalline powder (2.872 g, 98.9%) that is pure according to TLC and 1 H NMR. Rf 0.18 (1:1 v/v EtOAc/hexane), m.p. 193°–195°C (recrystallized from EtOAc/CHCl3). 1H NMR: 11.44 (s, 1H, H-3), 8.39 (s, 1H, imidazole), 7.69 (s, 1H, imidazole), 7.64 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.09 (s, 1H, imidazole), 5.85 (s, 1H, H-1′), 5.79 (d, 1H, J2′,3′ = 5.4 Hz, H-2′), 5.61 (d, 1H, J5,6 = 8.0 Hz, H-5), 4.74 (m, 1H, H-3′), 4.13–3.92 (m, 3H, H-4′, H-5′), 1.10–0.80 (m, 28H, Pri). Imidazolide S.3 is stable during aqueous extraction, but hydrolyzed under both acidic and alkaline conditions. Considerable decomposition was also observed during silica gel chromatography (1:1:1 to 1:1:2 v/v/v CHCl3/EtOAc/acetone, 66% yield of impure product).
Treat S.3 with primary or secondary aliphatic amine For unhindered primary and secondary amines 16a. To the solution of S.3 (prepared from 5 mmol S.2 in step 10), add the appropriate amine and allow reaction to proceed for the appropriate amount of time at 25°C: For S.4a: 0.514 mL (7.5 mmol) propargylamine for 1 hr For S.4b: 0.633 mL (7.5 mmol) N-methylpropargylamine overnight For S.4c: 1.630 g (7.0 mmol) 4-iodobenzylamine for 30 hr For S.4d: 1.874 g (7.0 mmol) 1-pyrenemethylamine hydrochloride and 1.0 mL (7.2 mmol) TEA for 72 hr. Monitor the completion of the reaction by TLC using EtOAc to monitor the disappearance of the imidazolide, or the appropriate solvents in step 23 to monitor the accumulation of product. 17a. Dilute the reaction mixture with 50 mL CH2Cl2. Proceed to step 18. For hindered primary amines 16b. Prepare a solution of 1.162 g (2.0 mmol) imidazolide S.3 in 25 mL CH3CN. Add the appropriate amine and incubate at 55°C until all of the starting compound is consumed: For S.4e: 514 mg (2.0 mmol) 2-aminomethyl-15-crown-5 for 96 hr For S.4f: 942 mg (3.0 mmol) H-Leu-Phe-NH2 hydrochloride and 0.61 mL (3.5 mmol) DIPEA for 72 hr. Monitor progress by TLC using EtOAc to monitor the disappearance of the imidazolide, or the appropriate solvents in step 23 to monitor the accumulation of product. 17b. Cool the mixture to ambient temperature and evaporate to dryness. Dilute with 100 mL EtOAc. Proceed to step 18. Work up S.4 18. Wash with 100 mL water, followed by 100 mL of 5% (w/v) citric acid, and then another 100 mL water. 19. Dry over Na2SO4, filter off the Na2SO4, and evaporate the solution.
Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
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20. Purify the residue by silica gel column chromatography (APPENDIX 3E) in the appropriate solvent system: For S.4a: 33% to 40% (v/v) EtOAc in CHCl3 For S.4b: 1:2 (v/v) EtOAc/CHCl3 For S.4c: 25% to 33% (v/v) EtOAc in CHCl3 For S.4d: 25% to 33% (v/v) EtOAc in CHCl3 For S.4e: 2% to 10% (v/v) MeOH in CHCl3 For S.4f: stepwise gradient of 33% to 50% (v/v) EtOAc in CHCl3, then 33% to 50% acetone in 1:1 (v/v) CHCl3/EtOAc. Typical volumes are 1 to 1.5 L for S.4a-e and ∼4 L for S.4f.
21. Monitor fractions by TLC, combine fractions containing product, and evaporate. 22. Co-evaporate the residue three times with 20 to 50 mL CH2Cl2, dry in vacuo for 2 hr, and grind the foam as described above (step 8). Dry in vacuo again to afford the compounds listed below as white amorphous solids. 23. Characterize the compound by TLC and 1H NMR. 2′-O-(Propargylaminocarbonyl)-3′,5′-O-(tetraisopropyldisiloxan-1,3-diyl)uridine (S.4a). Yield 2.780 g (97.9%). Rf: 0.49 (1:1 v/v CHCl3/EtOAc). 1H NMR: 11.42 (s, 1H, H-3), 7.84 (t, 1H, J = 5.4 Hz, OCONH), 7.69 (d, 1H, J5,6 = 8.0 Hz, H-6), 5.64 (s, 1H, H-1′), 5.59 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.34 (d, 1H, J2′,3′ = 4.9 Hz, H-2′), 4.50 (m, 1H, H-3′), 4.34–3.88 (m, 2H, H-5′), 3.85–3.69 (m, 3H, CH2N, H-4′), 3.06 (s, 1H, CH), 1.10–0.80 (m, 28H, Pri). 2′-O-(N-Methyl-N-propargylaminocarbonyl)-3′,5′-O-(tetraisopropyldisiloxan-1,3-diyl)uridine (S.4b). Yield 2.827 g (97.2%). Rf: 0.56 (1:1 v/v CHCl3/EtOAc). 1H NMR: 11.41(s, 1H, H-3), 7.66 (d, 1H, J5,6 = 8.1 Hz, H-6), 5.67 (s, 1H, H-1′), 5.59 (d, 1H, J5,6 = 8.1 Hz, H-5), 5.35 (m, 1H, H-2′), 4.53 (dd, 1H, J2′,3′ = 6.0 Hz, J3′,4′ = 7.9 Hz, H-3′), 4.17–3.83 (m, 5H, H-4′, H-5′, CH2N), 3.21 (s, 1H, CH), 2.93 (s, 1.8H), 2.84 (s, 1.2H) (NCH3, rotamers), 1.07–0.82 (m, 28H, Pri). 2′-O-(4-Iodobenzylaminocarbonyl)-3′,5′-O-(tetraisopropyldisiloxan-1,3-diyl)uridine (S.4c). Yield 3.491 g (93.6%). Rf: 0.50 (1:1 v/v CHCl3/EtOAc). 1H NMR: 11.41 (s, 1H, H-3), 7.96 (br t, 1H, OCONH), 7.68 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.62 (d, 2H, J = 7.6 Hz, ArH), 7.05 (d, 2H, J = 7.6 Hz, ArH), 5.65 (s, 1H, H-1′), 5.58 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.35 (d, 1H, J2′,3′ = 5.4 Hz, H-2′), 4.48 (m, 1H, H-3′), 4.20–3.80 (m, 5H, CH2Ar, H-4′, H-5′), 1.08–0.88 (m, 28H, Pri). 2′-O-(Pyrene-1-ylmethylaminocarbonyl)-3′,5′-O-(tetraisopropyldisiloxan-1,3-diyl)uridi ne (S.4d). Yield 3.492 g (93.9%). Rf: 0.54 (1:1 v/v CHCl3/EtOAc). 1H NMR: 11.43 (s, 1H, H-3), 8.42 (d, 1H, J9′,10′ = 9.3 Hz, pyrene H-10′), 8.31–7.97 (m, 9H, ArH, OCONH), 7.69 (d, 1H, J5,6 = 8.1 Hz, H-6), 5.68 (s, 1H, H-1′), 5.59 (d, 1H, J5,6 = 8.1 Hz, H-5), 5.40 (d, 1H, J2′,3′ = 5.8 Hz, H-2′), 4.91 (m, 2H, CH2Ar), 4.66 (m, 1H, H-3′), 4.04 (m, 1H, 2J5′a,5′b = 12.6 Hz, J4′,5′a = 3.2 Hz, H-5′a), 3.93–3.77 (m, 2H, H-4′, H-5′b), 1.04–0.67 (m, 28H, Pri). 2′-O-(1,4,7,10,13-Pentaoxacyclopentadecan-2-ylmethyl)-aminocarbonyl-3′,5′-O-(tetraisopropyldisiloxan-1,3-diyl)uridine (S.4e). Yield 1.165 g (76.4%). Rf: 0.32 (17:3 v/v CHCl3/MeOH). 1H NMR: 11.41 (s, 1H, H-3), 7.68 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.40 (t, 1H, J = 5.7 Hz, OCONH), 5.64 (s, 1H, H-1′), 5.59 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.33 (d, 1H, J2′,3′ = 5.7 Hz, H-2′), 4.48 (dd, 1H, J2′,3′ = 5.7 Hz, J3′,4′ = 8.2 Hz, H-3′), 3.99 (m, 2H, 2J5′a,5′b = 12.9 Hz, J4′,5′a = 2.7 Hz, J4′,5′b = 4.1 Hz, H-5′), 3.84 (m, 1H, H-4′), 3.64–3.36 (m, 19H#, CH(CH2OCH2)4CH2), 3.00 (m, 2H, CH2N), 1.06–0.82 (m, 28H, Pri). (#Calculated value; the signal of water is also present in the region).
Synthesis of Modified Oligonucleotides and Conjugates
4.21.7 Current Protocols in Nucleic Acid Chemistry
Supplement 15
Nα-[3′,5′-O-(Tetraisopropyldisiloxan-1,3-diyl)uridin-2′-O-ylcarbonyl]-L-leucyl-L-phenyl alaninamide (S.4f). Yield 610 mg (38.6%). Rf: 0.45 (EtOAc). ESI-MS: [M+H]+ calcd. 790.39, found 790.43. 1H NMR: 11.43 (s, 1H, H-3), 7.87 (d, 1H, J = 8.2 Hz, NHCHCH2Ph), 7.70 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.44 (d, 1H, J = 8.4 Hz, OCONH), 7.39 (br s, 1H, NH2), 7.19 (m, 5H, Ph), 7.06 (br s, 1H, NH2), 5.64 (s, 1H, H-1′), 5.59 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.35 (d, 1H, J2′,3′ = 5.1 Hz, H-2′), 4.47 (m, 2H, H-3′, CHCH2Ph), 4.15–3.83 (m, 4H, H-4′, H-5′, CHBui), 3.03–2.73 (m, 2H, 2J = 13.4 Hz, JAX = 4.7 Hz, JBX = 9.1 Hz, CH2Ph), 1.49 (m, 1H, CH2CHMe2), 1.29 (m, 2H, CH2Pri), 1.10–0.72 (m, 34H, SiCH(CH3)2, CH2CH(CH3)2). When S.4e and S.4f are prepared according to steps 16a and 17a, yields are 44% and 23%, respectively, after 90 days.
3′,5′-O-Desilylate uridine 2′-carbamates 24. Dissolve 2 mmol S.4 in 5 mL freshly distilled THF in a 30-mL screw-top Teflon bottle. Add 0.814 mL (5 mmol) triethylamine trihydrofluoride and magnetically stir the mixture 6 hr or overnight at ambient temperature. Monitor deprotection by TLC using 15% (v/v) MeOH/CHCl3. 25. Dilute the mixture with 25 mL hexane, shake the bottle well, and allow the two phases to separate. 26. Discard the upper layer and wash the residue three times by decantation with 25 mL of 1:1 (v/v) toluene/hexane. For uridine 2′-carbamates S.5a, S.5b, S.5e, and S.5f, the oily residues are co-evaporated first with toluene (step 27a) and then with pyridine (step 28) before dimethoxytritylation. For S.5c and S.5d, crystalline uridine 2′-carbamates are isolated by ethanol trituration (step 27b), and these are co-evaporated by pyridine only (step 28).
27a. For S.5a, S.5b, S.5e, and S5.f: Co-evaporate the crude oily uridine-2′-carbamates three times with 20 mL toluene. 27b. For S.5c and S.5d: Triturate the residue in 5 mL absolute ethanol, filter off the crystalline product, and wash it with 5 mL ethanol followed by 10 mL diethyl ether. Dry in vacuo. Characterize by TLC and 1H NMR. 2′-O-(4-Iodobenzylaminocarbonyl)uridine (S.5c). Yield 0.993 g (98.6%). Rf: 0.34 (17:3 v/v CHCl3/MeOH), m.p. 192°–197°C (EtOH). 1H NMR: 11.36 (s, 1H, H-3), 7.90 (m, 2H, H-6, OCONH), 7.65 (d, 2H, J = 7.4 Hz, ArH), 7.02 (d, 2H, J = 7.4 Hz, ArH), 5.99 (m, 1H, H-1′), 5.66 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.52 (d, 1H, J = 4.4 Hz, 3′-OH), 5.18 (m, 1H, 5′-OH), 5.05 (m, 1H, H-2′), 4.25–4.05 (m, 3H, H-3′, CH2N), 3.88 (m, 1H, H-4′), 3.59 (m, 2H, H-5′). 2′-O-(Pyren-1-ylmethylaminocarbonyl)uridine (S.5d). Yield 0.467 g (93.5%). Rf: 0.37 (17:3 v/v CHCl3/MeOH), m.p. 200°–208°C (EtOH). 1H NMR: 11.38 (s, 1H, H-3), 8.42– 7.90 (m, 11H, H-6, OCONH, ArH), 6.03 (d, 1H, J1′,2′ = 5.7 Hz, H-1′), 5.68 (d, 1H, J5,6 = 7.8 Hz, H-5), 5.57 (m, 1H, 3′-OH), 5.21 (m, 1H, 5′-OH), 5.14 (m, 1H, H-2′), 4.91 (m, 2H, CH2Ar), 4.24 (m, 1H, H-3′), 3.91 (m, 1H, H-4′), 3.60 (m, 2H, H-5′).
5′-O-Dimethoxytritylate uridine 2′-carbamates 28. Co-evaporate residue from step 27a or 27b three times with 20 mL pyridine. Dissolve in 30 mL pyridine and half-evaporate, then apply a dry nitrogen atmosphere.
Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
29. Cool the flask in an ice bath and add 0.85 g (2.5 mmol) DMTr⋅Cl in one portion. Monitor the reaction by TLC using 17:3 (v/v) CHCl3/MeOH for S.6e and EtOAC for all others. Add 0.17-g (0.5-mmol) portions of DMTr⋅Cl every 4 hr until the starting nucleoside S.5 disappears completely. The total amount of DMTr⋅Cl needed is dependent on the residual fluoride content and usually varies from 3 to 4 mmol.
4.21.8 Supplement 15
Current Protocols in Nucleic Acid Chemistry
30. After completion of the reaction, quench excess DMTr⋅Cl with 1 mL MeOH, wait 10 min, and dilute the mixture with 100 mL CHCl3. 31. Wash with 100 mL water, 100 mL of 5% NaHCO3, and again with 100 mL water. 32. Dry over Na2SO4, filter off the Na2SO4, and then evaporate the mixture. 33. Co-evaporate three times with 25 mL toluene and purify the residue on a silica gel column in the appropriate solvent system: For S.6a: 1% to 5% (v/v) MeOH/0.5% (v/v) TEA in CHCl3 For S.6b: 1% to 2% (v/v) MeOH/0.5% (v/v) TEA in 1:1 (v/v) CHCl3/EtOAc For S.6c: 0.5% to 2% (v/v) MeOH/0.5% (v/v) TEA in 1:1 (v/v) CHCl3/EtOAc For S.6d: 0.5% to 1.5% (v/v) MeOH/0.5% (v/v) pyridine in CHCl3 For S.6e: 3% to 9% (v/v) MeOH/0.5% (v/v) TEA in 2:1 (v/v) CHCl3/EtOAc For S.6f: 1% to 3% (v/v) MeOH/0.5% (v/v) pyridine in 2:1 (v/v) CHCl3/EtOAc. 34. Combine the fractions containing product S.6, evaporate, and then co-evaporate three times with 25 mL CH2Cl2. Dry the residue in vacuo to afford the compounds as amorphous solids. 35. Characterize the compounds by TLC and 1H NMR. 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(propargylaminocarbonyl)uridine (S.6a). Yield 1.156 g (92.1%). Rf: 0.60 (EtOAc). 1H NMR: 11.38 (br s, 1H, H-3), 7.85 (t, 1H, J = 5.7 Hz, OCONH), 7.66 (m, 1H, H-6), 7.40–7.19 (m, 9H, ArH), 6.89 (d, 4H, J = 8.7 Hz, ArH), 5.79 (d, 1H, J = 5.7 Hz, 3′-OH), 5.75 (s, 1H, H-1′), 5.39 (d, 1H, J5,6 = 8.1 Hz, H-5), 5.11 (m, 1H, H-2′), 4.35 (m, 1H, H-3′), 4.06 (m, 1H, H-4′), 3.79 (m, 2H, CH2N), 3.73 (s, 6H, OCH3), 3.40–3.15 (m, 3H#, H-5′, CH). 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(N-methyl-N-propargylaminocarbonyl)uridine (S.6b). Yield 1.097 g (85.5%). Rf: 0.73 (EtOAc). 1H NMR: 11.39 (s, 1H, H-3), 7.70 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.41–7.19 (m, 9H, ArH), 6.89 (d, 4H, J = 8.6 Hz, ArH), 5.89 (d, 1H, J1′,2′ = 3.3 Hz, H-1′), 5.54 (m, 1H, 3′-OH), 5.39 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.15 (m, 1H, H-2′), 4.32 (m, 1H, H-3′), 4.12–3.93 (m, 3H, H-4′, CH2N), 3.73 (s, 6H, OCH3), 3.35–3.16 (m, 3H#, H-5′, CH), 2.92 (s, 1.8H), 2.85 (s, 1.2H), (NCH3, rotamers). 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(4-iodobenzylaminocarbonyl)uridine (S.6c). Yield 1.378 g (85.5%). Rf: 0.75 (EtOAc). 1H NMR: 11.42 (s, 1H, H-3), 7.95 (t, 1H, J = 6.0 Hz, OCONH), 7.70 (d, 1H, J5,6 = 8.1 Hz, H-6), 7.66 (d, 1H, J = 8.2 Hz, ArH), 7.40–7.18 (m, 9 H, ArH), 7.06 (d, 2H, J = 8.2 Hz, ArH), 6.88 (d, 4H, J = 8.8 Hz, ArH), 5.92 (d, 1H, J1′,2′ = 4.8 Hz, H-1′), 5.61 (d, 1H, J3′,OH = 5.6 Hz, 3′-OH), 5.38 (d, 1H, J5,6 = 8.1 Hz, H-5), 5.17 (apparent t, 1H, J1′,2′ = J2′,3′ = 4.8 Hz, H-2′), 4.33 (m, 1H, H-3′), 4.14 (d, 2H, J = 6.0 Hz, NCH2), 3.98 (m, 1H, H-4′), 3.73 (s, 6H, OCH3), 3.35–3.16 (m, 2H#, H-5′). 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(pyren-1-ylmethylaminocarbonyl)uridine (S.6d). Yield 1.482 g (92.2%). Rf: 0.75 (EtOAc). 1H NMR: 11.44 (s, 1H, H-3), 8.41 (d, 1H, J9′,10′ = 9.3 Hz, H-10′), 8.32–8.00 (m, 9H, ArH, OCONH), 7.73 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.42–7.10 (m, 9H, ArH), 6.88 (d, 4H, J = 8.6 Hz, ArH), 5.97 (d, 1H, J1′,2′ = 4.6 Hz, H-1′), 5.65 (d, 1H, J3′,OH = 5.6 Hz, 3′-OH), 5.39 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.26 (m, 1H, H-2′), 4.95 (d, 2H, J = 5.3 Hz, NCH2), 4.38 (m, 1H, H-3′), 3.99 (m, 1H, H-4′), 3.72 (s, 6H, OCH3), 3.35–3.15 (m, 2H#, H-5′). 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(1,4,7,10,13-pentaoxacyclopentadecan-2-ylmethylaminocarbonyl)uridine (S.6e). Yield 1.248 g (85.5%). Rf 0.29 (17:3 v/v CHCl3/MeOH). 1H NMR: 11.39 (s, 1H, H-3), 7.69 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.41–7.12 (m, 10H, ArH, OCONH), 6.89 (d, 4H, J = 8.8 Hz, ArH), 5.90 (d, 1H, J1′,2′ = 4.7 Hz, H-1′), 5.53 (m, 1H, 3′-OH), 5.38 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.15 (m, 1H, H-2′), 4.32 (m, 1H, H-3′), 3.97 (m, 1H, H-4′), 3.73 (s, 6H, OCH3), 3.69–3.16 (m, 21H#, H-5′, CH(CH2OCH2)4CH2), 3.02 (m, 2H, CH2N).
Synthesis of Modified Oligonucleotides and Conjugates
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Nα-[5′-O-(4,4′-Dimethoxytrityl)uridin-2′-yloxycarbonyl]-L-leucyl-L-phenylalaninamide (S.6f). Yield 0.706 g (83.1%). Rf: 0.45 (EtOAc). 1H NMR: 11.45 (s, 1H, H-3), 7.90 (d, 1H, J = 8.4 Hz, NHCHCH2Ph), 7.72 (d, 1H, J5,6 = 8.1 Hz, H-6), 7.48–7.10 (m, 12H, ArH, NH2,OCONH), 6.89 (d, 4H, J = 8.8 Hz, ArH), 5.90 (d, 1H, J1′,2′ = 4.4 Hz, H-1′), 5.52 (d, 1H, J3′,OH = 5.9 Hz, 3′-OH), 5.43 (d, 1H, J5,6 = 8.1 Hz, H-5), 5.13 (m, 1H, H-2′), 4.47 (m, 2H, H-3′, CHCH2Ph), 4.37 (m, 1H, H-3′), 4.17 (m, 1H, CHBui), 3.98 (m, 1H, H-4′), 3.73 (s, 6H, OCH3), 3.52–3.10 (m, 2H#, H-5′), 3.05–2.70 (m, 2H, CH2Ph), 1.57–1.22 (m, 3H, CH2CHMe2), 0.86–0.69 (m, 6H, CH(CH3)2). (#Calculated value; the signal of water is also present in the region).
Phosphinylate 5′-O-DMTr-uridine 2′-carbamates 36. Co-evaporate 1.0 mmol S.6 two times with 20 mL dry CH2Cl2, dissolve in 40 mL dry CH2Cl2, and then apply a dry nitrogen atmosphere. 37. Add 171 mg (1.0 mmol) diisopropylammonium tetrazolide and 0.38 mL (1.2 mmol) bis(N,N-diisopropylamino)-2-cyanoethoxyphosphine and half-evaporate on a rotary evaporator. Stir the mixture 2 hr under nitrogen. Monitor by TLC using 25% acetone/1% TEA in CHCl3 (pair of diastereomers). 38. When conversion of S.6 is complete, dilute the mixture with 100 mL CHCl3 and wash with 100 mL of 5% NaHCO3 followed by 100 mL of 20% (w/v) NaCl. 39. Dry over Na2SO4, filter off the Na2SO4, and evaporate the solution to dryness. 40. Purify the residue on silica gel in the appropriate solvent system. For S.7a: 33% to 100% (v/v) EtOAc/1% (v/v) TEA in CHCl3 For S.7b: 30% to 70% (v/v) EtOAc/1% (v/v) TEA in CHCl3 For S.7c: 20% to 25% (v/v) acetone/1% (v/v) TEA in CHCl3 For S.7d: 5% to 25% (v/v) acetone/1% (v/v) TEA in CHCl3 For S.7e: 20% to 66% (v/v) acetone/1% (v/v) TEA in CHCl3 For S.7f: 50% to 100% (v/v) acetone/1% (v/v) TEA in 1:1 (v/v) CHCl3/EtOAc. 41. Combine fractions containing product S.7, evaporate, and dry in vacuo to afford the compounds as white amorphous solids. 42. Characterize the compounds by TLC and 1H and 31P NMR. 3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-O-(4,4′-dimethoxytrityl)-2′O-(propargylaminocarbonyl)uridine (S.7a). Yield 1.029 g (82.8%). Faster moving diastereomer: Rf: 0.56 (EtOAc). 1H NMR: 11.45 (s, 1H, H-3), 7.95 (m, 1H, OCONH), 7.72 (d, 1H, J5,6 = 8.1 Hz, H-6), 7.41–7.18 (m, 9H, ArH), 6.88 (d, 4H, J = 8.5 Hz, ArH), 5.89 (d, 1H, J1′,2′ = 5.0 Hz, H-1′), 5.43 (d, 1H, J5,6 = 8.1 Hz, H-5), 5.33 (apparent t, 1H, J1′,2′ = J2′,3′ = 5.0 Hz, H-2′), 4.56 (m, 1H, H-3′), 4.15 (m, 1H, H-4′), 3.85–3.68 (m, 8H, OCH3, CH2N), 3.63–3.44 (m, 4H, POCH2, CHCH3), 3.34–3.22 (m, 2H#, H-5′), 3.09 (t, 1H, 4J = 2.3 Hz, CH), 2.58 (t, 2H, J = 6.1 Hz, CH2CN), 1.19–0.95 (m, 12H, CHCH3). 31P NMR (CD3CN): 150.8. Slower moving diastereomer: Rf: 0.41 (EtOAc). 1H NMR: 11.44 (s, 1H, H-3), 7.90 (t, 1H, J = 5.7 Hz, OCONH), 7.73 (d, 1H, J5,6 = 8.1 Hz, H-6), 7.41–7.18 (m, 9H, ArH), 6.87 (m, 4H, ArH), 5.91 (d, 1H, J1′,2′ = 5.1 Hz, H-1′), 5.42 (d, 1H, J5,6 = 8.1 Hz, H-5), 5.36 (apparent t, 1H, J1′,2′ = J2′,3′ = 5.1 Hz, H-2′), 4.55 (m, 1H, H-3′), 4.10 (m, 1H, H-4′), 3.82–3.68 (m, 10H, OCH3, CH2N, POCH2), 3.46 (m, 2H, CHCH3), 3.35–3.21 (m, 2H#, H-5′), 3.12 (m, 1H, CH), 2.74 (t, 2H, J = 5.8 Hz, CH2CN), 1.19–0.89 (m, 12H, CHCH3). 31 P NMR (CD3CN): 150.7. Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
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3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-O-(4,4′-dimethoxytrityl)-2′O-(N-methyl-N-propargylaminocarbonyl)uridine (S.7b). Yield 1.004 g (79.5%). Rf: 0.58, 0.69 (EtOAc). 1H NMR: 11.42 (s, 1H, H-3), 7.70 (m, 1H, H-6), 7.42–7.16 (m, 9H, ArH), 6.87 (m, 4H, ArH), 5.90 (m, 1H, H-1′), 5.47–5.18 (m, 2H, H-5, H-2′), 4.55 (m, 1H, H-3′), 4.20–3.92 (m, 3H, H-4′, CH2N), 3.73 (s, 6H, OCH3), 3.62–3.18 (m, 7H#, POCH2, CHCH3, H-5′, CH), 2.94–2.78 (m, 3H, NCH3), 2.74, 2.56 (2m, 2H, CH2CN), 1.20–0.88 (m, 12H, CHCH3). 31P NMR (CD3CN): 150.9 (26%), 150.8 (30%), 150.4 (44%), diastereomers and rotamers. 3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-O-(4,4′-dimethoxytrityl)-2′O-(4-iodobenzylaminocarbonyl)uridine (S.7c). Yield 0.674 g (67%). Rf: 0.47, 0.62 (EtOAc). 1H NMR: 11.46 (s, 1H, H-3), 8.04 (m, 1H, OCONH), 7.73–7.61 (m, 3H, ArH, H-6), 7.42–7.18 (m, 9H, ArH), 7.07 (m, 2H, ArH), 6.87 (m, 4H, ArH), 5.93 (m, 1H, H-1′), 5.46–5.28 (m, 2H, H-5, H-2′), 4.53 (m, 1H, H-3′), 4.12 (m, 3H, H-4′, NCH2), 3.78–3.19 (m, 12H#, POCH2, CHN, H-5′, OCH3), 2.68, 2.58 (2m, 2H, CH2CN), 1.23–0.89 (m, 12H, CHCH3). 31P NMR (CD3CN): 149.3. 3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-O-(4,4′-dimethoxytrityl)-2′O-(pyren-1-ylmethylaminocarbonyl)uridine (S.7d). Yield 0.978 g (97.4%). Rf: 0.55, 0.68 (EtOAc). 1H NMR: 11.47 (s, 1H, H-3), 8.41 (d, 1H, J9′,10′ = 9.3 Hz, H-10′), 8.36–7.98 (m, 9H, ArH, OCONH), 7.73 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.43–7.17 (m, 9H, ArH), 6.87 (d, 4H, J = 8.5 Hz, ArH), 5.95 (d, 1H, J1′,2′ = 5.1 Hz, H-1′), 5.42 (m, 2H, H-5, H-2′), 4.91 (d, 2H, J = 5.3 Hz, NCH2), 4.57 (m, 1H, H-3′), 4.17 (m, 1H, H-4′), 3.71 (s, 6H, OCH3), 3.42–3.35 (m, 6H, POCH2, CHN, H-5′), 2.88, 2.58 (2t, 2H, J = 5.8 Hz, CH2CN, diastereo- mers), 1.17, 1.02–0.89 (2m, 12H, CHCH3). 31P NMR (CD3CN): 149.3. 3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-O-(4,4′-dimethoxytrityl)-2′O-(1,4,7,10,13-pentaoxacyclopentadecyl-2-methylaminocarbonyl)uridine (S.7e). Yield 0.955 g (93.4%). Rf: 0.11, 0.24 (EtOAc). 1H NMR: 11.44 (br s, 1H, H-3), 7.70 (m, 1H, H-6), 7.49–7.15 (m, 10H, ArH, OCONH), 6.87 (m, 4H, ArH), 5.91 (m, 1H, H-1′), 5.46–5.25 (m, 2H, H-5, H-2′), 4.53 (m, 1H, H-3′), 4.11 (m, 1H, H-4′), 3.80–3.19 (m, 31H#, POCH2, CHN, H-5′, CH(CH2OCH2)4CH2, OCH3), 3.00 (m, 2H, CH2N), 2.75, 2.59 (2m, 2H, CH2CN), 1.20–0.90 (m, 12H, CHCH3). 31P NMR (CD3CN): 149.28, 149.27 (diastereomers). (#Calculated value; the signal of water is also present in the region). Nα-[3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-O-(4,4′-dimethoxytrityl) uridin-2′-yloxycarbonyl]-L-leucyl-L-phenylalaninamide (S.7f). Yield 0.271 g (51.6%). Rf: 0.13, 0.20 (EtOAc). 1H NMR: 11.47 (s, 1H, H-3), 8.08 (d, 2H, J = 11.7 Hz, NH2), 7.83 (d, 1H, J = 8.6 Hz, NHCHCH2Ph), 7.70 (d, 1H, J5,6 = 8.1 Hz, H-6), 7.43–7.07 (m, 15H, ArH, OCONH), 6.88 (d, 4H, J = 8.8 Hz, ArH), 5.86 (d, 1H, J1′,2′ = 4.8 Hz, H-1′), 5.46 (d, 1H, J5,6 = 7.8 Hz, H-5), 5.32 (t, 1H, J = 5.3 Hz, H-2′), 4.64 (m, 1H, CHCH2Ph), 4.46 (m, 1H, H-3′), 4.17 (m, 1H, CHBui), 3.95 (m, 1H, H-4′), 3.73 (s, 6H, OCH3), 3.58–3.46 (m, 4H, POCH2, CHN), 3.04–2.99 (m, 2H, H-5′), 2.87–2.81 (m, 2H, CH2Ph), 2.59 (m, 2H, CH2CN), 1.23–0.98 (m, 12H, NCHCH3), 0.83–0.72 (m, 3H, CH2CHMe2), 0.65–0.59 (m, 6H, CH2CH(CH3)2). 31P NMR (DMSO-d6): 148.9.
PREPARATION OF URIDINE 2′-CARBAMATE PHOSPHORAMIDITES FROM PRIMARY AMINES THAT REQUIRE ADDITIONAL SIDE-CHAIN PROTECTION With an excess of di- or polyamines, imidazolide S.3 reacts mostly in a 1:1 ratio. The reaction with N-(3-aminopropyl)-1,3-propanediamine and spermine affords exclusively primary amino-substituted products (see Figure 4.21.3). This observation is in agreement with the recently reported good selectivity of imidazolecarbonyl derivatives towards primary versus secondary amines (Rannard and Davis, 2000). The remaining amino groups are either protected by trifluoroacetylation or used for further reaction (e.g., with protected cysteine derivative in S.4j). Trifluoroacetyl (Tfa) and 9-fluorenylmethoxycarbonyl (Fmoc) are the most common amino-protecting groups used in oligonucleotide synthesis. Aminoalcohols also show high selectivity towards N-substitution. The primary hydroxyl group of carbamate S.4k is further protected by a trimethylacetyl group to give
ALTERNATE PROTOCOL
Synthesis of Modified Oligonucleotides and Conjugates
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O NH O O Si
N
O
Si O
O COCF3
H N
O
N
NHCOCF3
O O
4g
NH O
N
O
Si
O COCF3
O
O
Si
H N
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N
N
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NHCOCF3
COCF3
4h
O
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NH
NH O
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Si O Si
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O
O
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Si O
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N
O
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Si
O
N H N
O
O
O
O
NHCOCF3
O
O 3
4i O NH O
N
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O NHFmoc
O Si
O
H N
O
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H N
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S S O
O 4j O
O
NH
NH O O Si
N
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Si O
O
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H N
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OH
O Si
O
Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
H N
O O
O 4k
O
N
O
Si
O
O O
4l
Figure 4.21.3 Preparation of side-chain-protected 3′,5′-O-silylated uridine 2′-carbamates (see Alternate Protocol).
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S.4l. This protecting group was found to be rather stable under oligonucleotide deprotection conditions (see Basic Protocol 2). NOTE: For Tfa 2′-carbamates (S.4g-S.4i), attempts to reduce the amine-to-imidazolide ratio to 1:1 give a decreased yield of the target compound and increased amounts of disubstituted byproducts. NOTE: Although Tfa derivatives are stable in the presence of TEA, a premature Fmoc release may occur from Fmoc compounds. Therefore, use of pyridine is advised as an additive in the column chromatography of Fmoc-containing intermediates. Additional Materials (also see Basic Protocol 1) 3′,5′-O-(Tetraisopropyldisiloxan-1,3-diyl)uridine (S.2; see Basic Protocol 1, step 9) Amine for S.3 conversion (select one): N-(3-Aminopropyl)-1,3-propanediamine, 98% pure (Aldrich; for S.4g) Spermine, 99% pure (Aldrich; for S.4h) 4,7,10-Trioxa-1,13-tridecanediamine, 97% pure (Sigma; for S.4i-S.4j) 2-(2-Aminoethoxy)ethanol (for S.4k) Reagent for amine protection (select one): S-Ethyl trifluorothioacetate, 97% pure (Aldrich; for S.4g-S.4i) Nα-Fmoc-S-tert-butylthio-L-cysteine pentafluorophenyl ester, 99% pure (Novabiochem; for S.4j) Trimethylacetyl chloride, 99% pure (Aldrich; for S.4l) Prepare di- and polyamine 2′-carbamates (S.4g-i) and protect by trifluoroacetylation For S.4g 1a. Prepare a solution of S.3 from 2.434 g (5.0 mmol) S.2 and 852 mg (5.25 mmol) 1,1′-carbonyldiimidazole in 50 mL dry CH2Cl2. Monitor the reaction by TLC (APPENDIX 3D) in 1:1 (v/v) EtOAc/hexane. 2a. Prepare a magnetically stirred, ice-cooled solution of 5.6 mL (40 mmol) N-(3-aminopropyl)-1,3-propanediamine in 100 mL CH2Cl2. 3a. Add the solution from step 1a dropwise to the solution in step 2a and incubate 2 hr. 4a. Remove the cooling bath and incubate the mixture overnight at room temperature. 5a. Wash the mixture with 200 mL water, 200 mL of 5% NaHCO3, and then another 200 mL water. 6a. Dry over Na2SO4, filter to remove the solid, and evaporate the solution to dryness. 7a. Co-evaporate the residue with 30 mL THF, dissolve in 10 mL dry THF, and add 5.0 mL (39 mmol) S-ethyl trifluorothioacetate in one portion. Incubate 20 hr. CAUTION: S-Ethyl trifluorothioacetate generates ethanethiol, which has a strong stench.
8a. Evaporate the mixture to dryness, co-evaporate the residue with CHCl3, and purify on a silica gel column using 1:1 to 1:3 (v/v) CHCl3/EtOAc (APPENDIX 3E). The product is obtained as a white amorphous solid. Trituration of S.4g in CH2Cl2/hexane gives colorless crystals.
Synthesis of Modified Oligonucleotides and Conjugates
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9a. Analyze the product S.4g by TLC and 1H NMR. Proceed to step 12. 2′-O-[4-Trifluoroacetyl-7-(trifluoroacetylamino)-4-azaheptan-1-ylaminocarbonyl]-3′,5′O-(tetraisopropyldisiloxan-1,3-diyl)uridine (S.4g). Yield 2.558 g (61.2%), m.p. 74°–77°C. Rf: 0.69 (EtOAc). 1H NMR: 11.42 (s, 1H, H-3), 9.51, 9.43 (2 br t, 1H, NHTFA, rotamers), 7.69 (d, 1H, J5,6 = 8.1 Hz, H-6), 7.51, 7.45 (2m, 1H, OCONH, rotamers), 5.65 (d, 1H, J1′,2′ = 1.8 Hz, H-1′), 5.59 (d, 1H, J5,6 = 8.1 Hz, H-5), 5.32 (m, 1H, H-2′), 4.50 (m, 1H, H-3′), 3.97 (m, 2H, 2J5′a,5′b = 12.5 Hz, J4′,5′a = 4.2 Hz, J4′,5′b = 2.4 Hz, H-5′), 3.83 (m, 1H, H-4′), 3.33 (m, 4H#, CH2NCH2), 3.19 (m, 2H, CH2NHTFA), 2.98 (m, 2H, OCONHCH2), 1.74 (m, 4H, CH2), 1.10–0.85 (m, 28H, Pri). (#Calculated value; the signal of water is also present in the region).
For S.4h 1b. Prepare a solution of S.3 as in step 1a. 2b. Prepare a magnetically stirred, ice-cooled solution of 2.63 g (13 mmol) spermine in 200 mL CH2Cl2. 3b. Add the solution from step 1b dropwise to the solution in step 2b and incubate 2 hr. 4b. Remove the cooling bath and stir the mixture overnight at room temperature. 5b. Dilute with 150 mL CHCl3 and wash two times with 200 mL of 20% NaCl. 6b. Dry over Na2SO4, filter, and evaporate the solution to dryness. 7b. Trifluoroacetylate the residue as in step 7a. 8b. Purify as in step 8a using 50% to 0% (v/v) CHCl3 in EtOAc. 9b. Analyze the product S.4h by TLC and 1H NMR. Proceed to step 12. 2′-O-[4,9-bis(Trifluoroacetyl)-12-(trifluoroacetylamino)-4,9-diazadodecan-1-ylaminocarbonyl]-3′,5′-O-(tetraisopropyldisiloxan-1,3-diyl)uridine (S.4h). Yield 2.457 g (49.0%). Rf: 0.67 (EtOAc). 1H NMR: 11.42 (s, 1H, H-3), 9.51, 9.44 (2 br t, 1H, NHTFA, rotamers), 7.69 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.51, 7.45 (2t, 1H, J = 5.2 Hz, OCONH, rotamers), 5.65 (s, 1H, H-1′), 5.59 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.32 (m, 1H, H-2′), 4.49 (m, 1H, H-3′), 3.98 (m, 2H, 2J5′a,5′b = 12.6 Hz, J4′,5′a = 4.2 Hz, H-5′), 3.83 (m, 1H, H-4′), 3.40–3.28 (m, 8H#, CH2N(TFA)CH2), 3.20 (m, 2H, CH2NHTFA), 2.98 (m, 2H, OCONHCH2), 1.86–1.60 (m, 4H, CH2CH2NH), 1.51 (m, 4H, CH2CH2CH2CH2), 1.06–0.80 (m, 28H, Pri). (#Calculated value; the signal of water is also present in the region).
For S.4i 1c. Prepare a solution of S.3 from 1.703 g (3.5 mmol) S.2 and 592 mg (3.65 mmol) 1,1′-carbonyldiimidazole in 30 mL dry CH2Cl2. Monitor the reaction by TLC in EtOAc. 2c. Prepare a magnetically stirred, ice-cooled solution of 3.9 mL (17.5 mmol) 4,7,10-trioxa-1,13-tridecanediamine in 100 mL CH2Cl2. 3c. Add the solution from step 1c dropwise to the solution in step 2c and incubate 2 hr. 4c. Remove the cooling bath and incubate the mixture overnight at room temperature. 5c. Wash the mixture with 150 mL water, 150 mL of 5% NaHCO3, and then another 150 mL water. 6c. Dry over Na2SO4, filter, and evaporate the solution to dryness. Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
7c. Dissolve the residue in 10 mL dry CH2Cl2 and add 3.0 mL (24 mmol) S-ethyl trifluorothioacetate in one portion. Incubate the mixture overnight. CAUTION: S-Ethyl trifluorothioacetate generates ethanethiol, which has a strong stench.
4.21.14 Supplement 15
Current Protocols in Nucleic Acid Chemistry
8c. Evaporate the mixture to dryness and purify on a silica gel column using 1:1 to 1:4 (v/v) CHCl3/EtOAc. 9c. Analyze the product S.4i by TLC and 1H NMR. Proceed to step 12. 2′-O-[13-(Trifluoroacetylamino)-4,7,10-trioxatridecan-1-ylaminocarbonyl]-3′,5′-O-(tetraisopropyldisiloxan-1,3-diyl)uridine (S.4i). Yield 1.888 g (65.1%) as colorless oil. Rf: 0.42 (EtOAc). 1H NMR: 11.41 (s, 1H, H-3), 9.36 (br s, 1H, NHTFA), 7.69 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.36 (t, 1H, J = 5.7 Hz, OCONH), 5.64 (d, 1H, J1′,2′ = 1.8 Hz, H-1′), 5.59 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.31 (m, 1H, H-2′), 4.49 (dd, 1H, J2′,3′ = 5.0 Hz, J3′,4′ = 7.9 Hz, H-3′), 3.99 (m, 2H, 2J5′a,5′b = 13.0 Hz, J4′,5′a = 2.7 Hz, J4′,5′b = 4.4 Hz, H-5′), 3.82 (m, 1H, H-4′), 3.55–3.30 (m, 12H#, (CH2OCH2)3), 3.22 (apparent q, 2H, J = 6.5 Hz, CH2NHTFA), 3.00 (m, 2H, OCONHCH2), 1.69 (apparent quintet, 2H, J = 6.8 Hz, CH2), 1.60 (apparent quintet, 2H, J = 6.6 Hz, CH2), 1.06–0.81 (m, 28H, Pri). (#Calculated value; the signal of water is also present in the region).
Prepare functionalized 2′-carbamate (S.4j) by acylation 1d. Prepare a solution of S.3 as in step 1a. 2d. Prepare a magnetically stirred, ice-cooled solution of 5.5 mL (25 mmol) 4,7,10-trioxa-1,13-tridecanediamine in 100 mL CH2Cl2. 3d. Add the solution from step 1d dropwise to the solution in step 2d and incubate 2 hr. 4d. Remove the cooling bath and incubate the mixture overnight at room temperature. 5d. Wash the mixture with 200 mL water, 200 mL of 5% NaHCO3, and then another 200 mL water. 6d. Dry over Na2SO4, filter, and evaporate the solution to dryness. 7d. Dissolve the crude amine in 20 mL dry CH2Cl2 and add 2.988 g (5.0 mmol) Nα-Fmoc-S-tert-butylthio-L-cysteine pentafluorophenyl ester. Incubate 2 hr. 8d. Dilute the mixture with 200 mL CHCl3 and wash with 150 mL of 5% NaHCO3, 150 mL of 5% citric acid, and 150 mL water. 9d. Dry as in step 6d and then purify the residue on a silica gel column using 1:1:0 to 1:3:0 to 1:2:1 (v/v/v) CHCl3/EtOAc/acetone. 10d. Pool the appropriate fractions, evaporate, and co-evaporate three times with 20 mL CH2Cl2. Dry in vacuo. 11d. Analyze the product S.4j by TLC and 1H NMR. Proceed to step 12. 2′-{O-13-[S-(tert-Butylthio)-N-(9-fluorenylmethoxycarbonyl)-L-cysteinylamino]-4,7,10trioxatridecan-1-ylaminocarbonyl}-3′,5′-O-(tetraisopropyldisiloxan-1,3-diyl)uridine (S.4j). Yield 3.732 g (65.2%), white amorphous solid. Rf: 0.41 (EtOAc). 1H NMR: 11.41 (s, 1H, H-3), 8.01 (t, 1H, J = 5.2 Hz, NHCOCH), 7.88 (d, 2H, J = 7.4 Hz, ArH (fluorene)), 7.75–7.64 (m, 4H, ArH (fluorene H-1,4,5,8), H-6, NHFmoc), 7.43–7.27 (m, 5 H, ArH (fluorene H-2,3,6,7), OCONHCH2), 5.64 (s, 1H, H-1′), 5.59 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.31 (d, 1H, J2′,3′ = 5.8 Hz, H-2′), 4.49 (m, 1H, H-3′), 4.32–4.13 (m, 4H, CHCH2O (Fmoc), COCHN), 3.98 (m, 2H, 2J5′a,5′b = 13.0 Hz, J4′,5′a = 4.2 Hz, J4′,5′b = 2.5 Hz, H-5′), 3.82 (m, 1H, H-4′), 3.50–3.26 (m, 12H#, (CH2OCH2)3), 3.15–2.82 (m, 6H, CH2N, CH2S), 1.60 (m, 4H, CH2CH2CH2), 1.28 (s, 9H, But), 1.06–0.81 (m, 28H, Pri). (#Calculated value; the signal of water is also present in the region). Synthesis of Modified Oligonucleotides and Conjugates
4.21.15 Current Protocols in Nucleic Acid Chemistry
Supplement 15
Prepare hydroxyalkyl 2′-carbamate (S.4k) and trimethylacetylate to give S.4l 1e. To obtain 2-(2-hydroxyethoxy)ethyl-3′,5′-silyl-2′-carbamate (S.4k), apply the procedure for unhindered primary amines (see Basic Protocol 1, steps 16a to 17a and 18 to 22), using 1.50 mL (15.0 mmol) 2-(2-aminoethoxy)ethanol in step 16a (∼3 hr) and 2:1:1 to 1:1:1 to 1:1:2 (v/v/v) CHCl3/EtOAc/acetone for chromatography in step 20. 2′-O-[2-(2-Hydroxyethoxy)ethylaminocarbonyl]-3′,5′-O-(tetraisopropyldisiloxan-1,3diyl)uridine (S.4k). Yield 2.269 g (73.5%). Rf: 0.28 (EtOAc). 1H NMR: 11.41 (s, 1H, H-3), 7.68 (d, 1H, J5,6 = 7.9 Hz, H-6), 7.40 (br s, 1H, OCONH), 5.64 (s, 1H, H-1′), 5.59 (d, 1H, J5,6 = 7.9 Hz, H-5), 5.31 (d, 1H, J2′,3′ = 5.4 Hz, H-2′), 4.55 (m, 2H, H-3′, OH), 4.12–3.79 (m, 3H, H-4′, H-5′), 3.53–3.27 (m, 6H#, CH2OCH2CH2), 3.12 (m, 2H, CH2N), 1.10–0.82 (m, 28H, Pri). (#Calculated value; the signal of water is also present in the region).
2e. To a stirred solution of 2.162 g (3.5 mmol) S.4k in 20 mL dry CH2Cl2, add 1.0 mL (12.5 mmol) pyridine and 0.615 mL (5 mmol) trimethylacetyl chloride. 3e. Keep the reaction at ambient temperature until the starting compound disappears, as monitored by TLC using EtOAc (∼20 hr). 4e. Dilute the mixture with 100 mL CH2Cl2. 5e. Wash with 100 mL water, 100 mL of 5% NaHCO3, and another 100 mL water. 6e. Dry over Na2SO4, filter, and evaporate to dryness. 7e. Purify the residue on silica gel using 10% to 50% (v/v) EtOAc/CHCl3. 8e. Combine the fractions, evaporate, and co-evaporate three times with 20 mL CH2Cl2. Dry in vacuo. 9e. Analyze the product S.4l by TLC and 1H NMR. Proceed to step 12. 2′-O-[2-(2-Pivaloyloxyethoxy)ethylaminocarbonyl]-3′,5′-O-(tetraisopropyldisiloxan-1,3diyl)uridine (S.4l). Yield 1.977 g (80.5%). Rf: 0.34 (1:1 v/v CHCl3/EtOAc). 1H NMR: 11.41 (s, 1H, H-3), 7.69 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.37 (br t, 1H, J = 4.6 Hz, OCONH), 5.64 (s, 1H, H-1′), 5.59 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.31 (d, 1H, J2′,3′ = 4.9 Hz, H-2′), 4.50 (m, 1H, H-3′), 4.17–3.78 (m, 5H, H-4′, H-5′, CH2OPiv), 3.59 (m, 2H, CH2OCH2), 3.40 (m, 2H#, CH2N), 3.10 (m, 2H, CH2OCH2), 1.12 (s, 9H, But), 1.07–0.83 (m, 28H, Pri). (#Calculated value; the signal of water is also present in the region).
Prepare 5′-O-dimethoxytritylated side-chain-protected 2′-carbamates (S.6g-j and S.6l) 12. Deprotect the 3′- and 5′-hydroxyl groups of side-chain-protected 2′-carbamates (S.4g-j and S.4l) by treatment with triethylamine trihydrofluoride in THF (see Basic Protocol, steps 24 to 26). Coevaporate with toluene (step 27a). 13. 5′-O-Dimethoxytritylate crude 3′,5′-unprotected side-chain-protected 2′-carbamates (S.5g-j and S.5l) by treatment with DMTr⋅Cl in pyridine, and purify S.6g-j and S.6l by column chromatography (see Basic Protocol 1, steps 28 to 34).
Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
For S.6g: stepwise gradient of 1% to 4% (v/v) MeOH/0.5% (v/v) TEA in 2:1 (v/v) CHCl3/EtOAc For S.6h: 1% to 5% (v/v) MeOH/0.5% (v/v) TEA in 2:1 (v/v) CHCl3/EtOAc For S.6i: 1% to 3.5% (v/v) MeOH/0.5% (v/v) TEA in 2:1 (v/v) CHCl3/EtOAc For S.6j: 0% to 6% (v/v) MeOH/1% (v/v) pyridine in 1:1 (v/v) CHCl3/EtOAc For S.6l: 0.5% to 2% (v/v) MeOH/0.5% (v/v) TEA in 2:1 (v/v) CHCl3/EtOAc.
4.21.16 Supplement 15
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14. Analyze the products by TLC and 1H NMR. 5′-O-(4,4′-Dimethoxytrityl)-2′-O-[4-trifluoroacetyl-7-(trifluoroacetylamino)-4-azaheptan-1ylaminocarbonyl]uridine (S.6g). Yield 1.556 g (86.8%), white amorphous solid. Rf: 0.49 (EtOAc). 1H NMR: 11.37 (br s, 1 H, H-3) (exchangeable with D2O), 9.51, 9.46 (2 br s, 1H, NHTFA, rotamers), 7.69 (m, 1H, H-6), 7.53–7.19 (m, 10H, ArH (DMTr), OCONH), 6.89 (m, 4H, ArH (DMTr)), 5.92 (m, 1H, H-1′), 5.56 (m, 1H, 3′-OH) (exchangeable with D2O), 5.37 (m, 1H, H-5), 5.15 (m, 1H, H-2′), 4.34 (m, 1H, H-3′), 3.97 (m, 1H, H-4′), 3.73 (s, 6H, OCH3), 3.40–3.14 (m, 8H#, H-5′, CH2NTFA), 3.00 (m, 2H, OCONHCH2), 1.73 (m, 4H, CH2CH2CH2). 5′-O-(4,4′-Dimethoxytrityl)-2′-O-[4,9-bis(trifluoroacetyl)-12-(trifluoroacetylamino)- 4,9diazadodecan-1-aminocarbonyl]uridine (S.6h). Yield 1.773 g (83.4%). Rf: 0.50 (EtOAc). 1 H NMR: 11.40 (s, 1H, H-3), 9.51, 9.44 (2m, 1H, NHTFA rotamers), 7.69 (m, 1H, H-6), 7.51–7.18 (m, 10H, ArH, OCONH), 6.88 (d, 4H, J = 8.8 Hz, ArH), 5.92 (m, 1H, H-1′), 5.56 (m, 1H, 3′-OH), 5.37 (m, 1H, H-5), 5.15 (m, 1H, H-2′), 4.34 (m, 1H, H-3′), 3.96 (m, 1H, H-4′), 3.73 (s, 6H, OCH3), 3.44–3.14 (m, 12H#, H-5′, CH2NTFA), 3.01 (m, 2H, OCONHCH2), 1.73 (m, 4H, CH2CH2NH), 1.52 (m, 4H, CH2CH2CH2CH2). (#Calculated value; the signal of water is also present in the region). 5′-O-(4,4′-Dimethoxytrityl)-2′-O-[13-(trifluoroacetylamino)-4,7,10-trioxatridecan-1ylaminocarbonyl]uridine (S.6i). Yield 1.427 g (80.3%). Rf: 0.22 (EtOAc). 1H NMR: 11.39 (s, 1H, H-3), 9.36 (br s, 1H, NHTFA), 7.70 (d, 1H, J5,6 = 8.1 Hz, H-6), 7.41–7.19 (m, 10H, ArH, OCONH), 6.89 (d, 4H, J = 8.8 Hz, ArH), 5.89 (d, 1H, J1′,2′ = 5.0 Hz, H-1′), 5.54 (d, 1H, J3′,OH = 5.6 Hz, 3′-OH), 5.38 (d, 1H, J5,6 = 8.1 Hz, H-5), 5.13 (apparent t, 1H, J1′,2′ = J2′,3′ = 5.0 Hz, H-2′), 4.32 (m, 1H, H-3′), 3.96 (m, 1H, H-4′), 3.73 (s, 6H, OCH3), 3.53–3.36 (m, 12H, (CH2OCH2)3), 3.28–3.16 (m, 4H, H-5′, CH2NHTFA), 3.02 (m, 2H, OCONHCH2), 1.75–1.56 (m, 4H, CH2CH2CH2). 5′-O-(4,4′-Dimethoxytrityl)-2′-O-{13-[S-(tert-butylthio)-N-(9-fluorenylmethyloxycarbonyl) cysteinylamino]-4,7,10-trioxatridecane-1-ylaminocarbonyl}uridine (S.6j). Yield 2.304 g (95.5%). Rf: 0.21 (EtOAc). 1H NMR: 11.39 (s, 1H, H-3), 8.02 (t, 1H, J = 5.5 Hz, NHCOCH), 7.88 (d, 2H, J = 7.5 Hz, ArH (fluorene)), 7.70 (m, 4H, ArH (fluorene), H-6, NHFmoc), 7.44–7.10 (m, 14H, ArH (fluorene H-2,3,6,7, DMTr), OCONHCH2), 6.88 (d, 4H, J = 8.9 Hz, ArH (DMTr)), 5.90 (d, 1H, J1′,2′ = 5.0 Hz, H-1′), 5.54 (d, 1H, J3′,OH = 5.6 Hz, 3′-OH), 5.38 (d, 1H, J5,6 = 8.1 Hz, H-5), 5.14 (apparent t, 1H, J1′,2′ = J2′,3′ = 5.0 Hz, H-2′), 4.37–4.15 (m, 5H, CHCH2O (Fmoc), COCHN, H-3′), 3.96 (m, 1H, H-4′), 3.72 (s, 6H, OCH3), 3.50–3.33 (m, 12H, (CH2OCH2)3), 3.28–2.88 (m, 8H, CH2N, CH2S, H-5′), 1.61 (apparent quintet, 4H, J = 6.6 Hz, CH2CH2CH2), 1.28 (s, 9H, But). 5′-O-(4,4′-Dimethoxytrityl)-2′-O-[2-(2-trimethylacetoxyethoxy)ethylaminocarbonyl] uridine (S.6l). Yield 1.377 g (90.4%). Rf: 0.60 (EtOAc). 1H NMR: 11.40 (s, 1H, H-3), 7.69 (d, 1H, J5,6 = 8.1 Hz, H-6), 7.41–7.19 (m, 10H, ArH, OCONH), 6.89 (d, 4H, J = 8.8 Hz, ArH), 5.90 (d, 1H, J1′,2′ = 5.0 Hz, H-1′), 5.53 (d, 1H, J3′,OH = 5.7 Hz, 3′-OH), 5.38 (d, 1H, J5,6 = 8.1 Hz, H-5), 5.12 (apparent t, 1H, J1′,2′ = J2′,3′ = 5.0 Hz, H-2′), 4.32 (m, 1H, H-3′), 4.11 (m, 2H, CH2OPiv), 3.97 (m, 1H, H-4′), 3.73 (s, 6H, OCH3), 3.57, 3.43 (2m, 4H, CH2OCH2), 3.27–3.02 (m, 4H, H-5′, CH2N), 1.11 (s, 9H, But).
Prepare phosphoramidites S.7g-j and S.7l 15. Phosphinylate S.6 compounds by treatment with bis(N,N-diisopropylamino)-2-cyanoethoxyphosphine and diisopropylammonium tetrazolide in dry CH2Cl2, and purify S.7 by column chromatography (see Basic Protocol 1, steps 36 to 41). For S.7g: 20% to 66% (v/v) acetone/1% (v/v) TEA in CHCl3 For S.7h: 20% to 50% (v/v) acetone/1% (v/v) TEA in CHCl3 For S.7i: 20% to 66% (v/v) acetone/1% (v/v) TEA in CHCl3 For S.7j: 20% to 40% (v/v) acetone/1% (v/v) pyridine in CHCl3 For S.7l: 1% (v/v) TEA in 1:1 (v/v) acetone/CHCl3.
Synthesis of Modified Oligonucleotides and Conjugates
4.21.17 Current Protocols in Nucleic Acid Chemistry
Supplement 15
16. Analyze the products by TLC and 1H and 31P NMR. 3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-O-(4,4′-dimethoxytrityl)-2′O-[4-trifluoroacetyl-7-(trifluoroacetylamino)-4-azaheptan-1-ylaminocarbonyl]uridine (S.7g). Yield 1.396 g (93.8%). Rf: 0.14, 0.25 (EtOAc). 1H NMR: 11.40 (br s, 1H, H-3), 9.52, 9.44 (2 br s, 1H, NHTFA, rotamers), 7.70 (m, 1H, H-6), 7.53 (m, 1H, OCONH), 7.41–7.15 (m, 9H, ArH), 6.87 (m, 4H, ArH), 5.92 (m, 1H, H-1′), 5.45–5.22 (m, 2H, H-5, H-2′), 4.54 (m, 1H, H-3′), 4.12 (m, 1H, H-4′), 3.80–3.09 (m, 18H#, OCH3, POCH2, CHN, H-5′, CH2NTFA), 2.99 (m, 2H, OCONHCH2), 2.74, 2.57 (2m, 2H, CH2CN), 1.73 (m, 4H, CH2CH2CH2), 1.20–0.87 (m, 12H, CHCH3). 31P NMR (CD3CN): 150.74 (54%), 150.66 (22%), 150.58 (24%), diastereomers and rotamers. 3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-O-(4,4′-dimethoxytrityl)-2′O-[4,9-bis(trifluoroacetyl)-12-(trifluoroacetylamino)-4,9-diazadodecyl-1-aminocarbonyl] uridine (S.7h). Yield 1.242 g (98.3%). Rf: 0.18, 0.32 (EtOAc). 1H NMR: 11.36 (br s, 1H, H-3), 9.52, 9.45 (2 br s, 1H, NHTFA, rotamers), 7.69 (m, 1H, H-6), 7.53 (m, 1H, OCONH), 7.41–7.16 (m, 9H, ArH), 6.87 (m, 4H, ArH), 5.92 (m, 1H, H-1′), 5.44–5.21 (m, 2H, H-5, H-2′), 4.53 (m, 1H, H-3′), 4.12 (m, 1H, H-4′), 3.80–3.12 (m, 22H#, OCH3, POCH2, CHN, H-5′, CH2NTFA), 2.99 (m, 2H, OCONHCH2), 2.74, 2.57 (2m, 2H, CH2CN), 1.74 (m, 4H, CH2CH2NH), 1.51 (m, 4H, CH2CH2CH2CH2), 1.19–0.86 (m, 12H, CHCH3). 31P NMR (CD3CN): 150.8 (56%), 150.7 (20%), 150.6 (24%), diastereomers and rotamers. 3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-O-(4,4′-dimethoxytrityl)-2′O-[13-(trifluoroacetylamino)-4,7,10-trioxatridecan-1-ylaminocarbonyl]uridine (S.7i). Yield 1.069 g (98.2%). Rf: 0.20, 0.34 (EtOAc). 1H NMR: 11.44 (s, 1H, H-3), 9.37 (br s, 1H, NHTFA), 7.71 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.47–7.19 (m, 10H, ArH, OCONH), 6.87 (m, 4H, ArH), 5.91 (m, 1H, H-1′), 5.42 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.31 (m, 1H, H-2′), 4.55 (m, 1H, H-3′), 4.12 (m, 1H, H-4′), 3.80–3.16 (m, 24H#, (CH2OCH2)3, POCH2, CHN, H-5′, OCH3), 3.01 (m, 2H, OCONHCH2), 2.74, 2.57 (2m, 2H, CH2CN), 1.74–1.55 (m, 4H, CH2CH2CH2), 1.18–0.92 (m, 12H, CHCH3). 31P NMR (CD3CN): 149.35, 149.28 (diastereomers). 3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-O-(4,4′-dimethoxytrityl)-2′O-{13-[S-(tert-butylthio)-N-(9-fluorenylmethyloxycarbonyl)cysteinylamino]-4,7,10- trioxatridecane-1-ylaminocarbonyl}uridine (S.7j). Yield 1.130 g (81.4%). Rf: 0.30, 0.53 (EtOAc). Faster moving diastereomer: 1H NMR: 11.44 (s, 1H, H-3), 8.02 (m, 1H, NHCOCH), 7.88 (d, 2H, J = 7.4 Hz, ArH (fluorene)), 7.70 (m, 4H, ArH (fluorene), H-6, NHFmoc), 7.47–7.08 (m, 14H, ArH (fluorene H-2,3,6,7, DMTr), OCONHCH2), 6.88 (d, 4H, J = 8.6 Hz, ArH (DMTr)), 5.89 (d, 1H, J1′,2′ = 5.1 Hz, H-1′), 5.42 (d, 1H, J5,6 = 7.9 Hz, H-5), 5.30 (m, 1H, H-2′), 4.55 (m, 1H, H-3′), 4.35–4.07 (m, 5H, CHCH2O (Fmoc), COCHN, H-4′), 3.72 (s, 6H, OCH3), 3.60–3.21 (m, 24H#, (CH2OCH2)3, POCH2, NCHCH3, H-5′), 3.15–2.85 (m, 6H, CH2N, CH2S), 2.57 (m, 2H, CH2CN), 1.59 (m, 4H, CH2CH2CH2), 1.28 (s, 9H, But), 1.13–0.93 (m, 12H, CHCH3). Slower moving diastereomer: 1H NMR: 11.44 (s, 1H, H-3), 8.02 (m, 1H, NHCOCH), 7.88 (d, 2H, J = 7.4 Hz, ArH (fluorene)), 7.70 (m, 4H, ArH (fluorene), H-6, NHFmoc), 7.42–7.12 (m, 14H, ArH (fluorene H-2,3,6,7, DMTr), OCONHCH2), 6.87 (d, 4H, J = 8.2 Hz, ArH (DMTr)), 5.91 (m, 1H, H-1′), 5.42 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.34 (m, 1H, H-2′), 4.53 (m, 1H, H-3′), 4.33–4.03 (m, 5H, CHCH2O (Fmoc), COCHN, H-4′), 3.80–3.20 (m, 24H#, (CH2OCH2)3, POCH2, NCHCH3, H-5′, OCH3), 3.12–2.87 (m, 6H, CH2N, CH2S), 2.74 (t, 2H, J = 5.7 Hz, CH2CN), 1.61 (m, 4H, CH2CH2CH2), 1.27 (s, 9H, But), 1.12–0.88 (m, 12H, CHCH3).
Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-O-(4,4′-dimethoxytrityl)-2′O-[2-(2-trimethylacetoxyethoxy)ethylaminocarbonyl]uridine (S.7l). Yield 0.947g (98.4%). Rf: 0.43, 0.57 (EtOAc). 1H NMR: 11.44 (s, 1H, H-3), 7.70 (d, 1H, J5,6 = 8.0 Hz, H-6), 7.47–7.19 (m, 10H, ArH, OCONH), 6.88 (m, 4H, ArH), 5.90 (m, 1H, H-1′), 5.43 (d, 1H, J5,6 = 8.0 Hz, H-5), 5.30 (m, 1H, H-2′), 4.55 (m, 1H, H-3′), 4.10 (m, 3H, H-4′, CH2OCO), 3.72 (s, 6H, OCH3), 3.62–3.01 (m, 12H#, POCH2, CHN, H-5′, NCH2CH2OCH2), 2.74, 2.57 (2m, 2H, CH2CN), 1.23–0.90 (m, 21H, CHCH3, But). 31P NMR (CD3CN): 149.5, 149.4, 149.3 (diastereomers and rotamers). (#Calculated value; the signal of water is also present in the region).
4.21.18 Supplement 15
Current Protocols in Nucleic Acid Chemistry
SYNTHESIS, ISOLATION, AND CHARACTERIZATION OF OLIGONUCLEOTIDES CONTAINING URIDINE 2′-CARBAMATES
BASIC PROTOCOL 2
3′-Phosphoramidites of 5′-dimethoxytritylated uridine 2′-carbamates are stable during prolonged storage at –20°C. The compounds are less reactive compared to standard 2′-deoxyribonucleoside phosphoramidites due to the steric bulk of the 2′-carbamate group. Therefore, the coupling time should be increased to 10 to 15 min to achieve a 97% to 98% yield, which is acceptable for introduction of one or a few modifications into an oligonucleotide chain. In all cases, a 2′-carbamate function is completely stable to the conditions of oligonucleotide synthesis as well as to final deprotection with concentrated aqueous ammonia (55°C, 8 to 16 hr). In general, an increased coupling time is the only change in oligonucleotide synthesis and isolation that is necessary for preparation of uridine-2′-carbamate-containing oligonucleotides. However, there are two important details that should be noted. First, the 2′-trimethylacetate ester obtained from phosphoramidite S.7l is unexpectedly stable to ammonia treatment. Only after 96 hr of standard deprotection does the MALDI-TOF spectrum show complete ester hydrolysis in a doubly modified oligo-2′-deoxyribonucleotide. Second, the phosphoramidite S.7f is sparingly soluble in acetonitrile and N,N-dimethylformamide and is highly prone to gel formation, which makes it difficult to use for standard oligonucleotide synthesis. All the other phosphoramidites can be used as 0.1 M solutions in anhydrous acetonitrile. Oligonucleotide synthesis can be carried out on any automated DNA/RNA synthesizer. The authors have used Applied Biosystems 380B and 394 on both a 0.2- and 1-µmol scale using 2′-deoxyribonucleoside and 2′-O-methyl-ribonucleoside phosphoramidites (Transgenomics), as well as 2′-TOM-ribonucleoside phosphoramidites (Glen Research; also see UNITS 2.9 & 3.8). A number of techniques may be used for isolation and purification of oligonucleotides with uridine 2′-carbamate modifications, such as reversed-phase or ion-exchange HPLC, on cartridges (e.g., PolyPak, Glen Research), or by denaturing PAGE. A standard vertical gel electrophoresis apparatus is suitable for PAGE of modified oligonucleotides. The molecular mass of the oligonucleotides was checked by MALDITOF mass spectrometry on an Applied Biosystems Voyager DE workstation in positive ion mode. Thermal denaturation experiments with oligonucleotide duplexes were performed on a Perkin Elmer Lambda 40 UV/Vis spectrophotometer with Peltier temperature programmer in the hybridization buffer. Fluorescence spectra were recorded using a Perkin Elmer LS 50B Luminescence Spectrometer in the same buffer. For evaporation of small volumes, a Speedvac evaporator is ideal. Materials Uridine 2′-carbamate phosphoramidite(s) (S.7a-j,l; see Basic Protocol 1 and Alternate Protocol) Acetonitrile, anhydrous Phosphoramidites: 2′-Deoxyribonucleoside phosphoramidites (Transgenomics) 2′-O-Methyl-ribonucleoside phosphoramidites (2′-OMe; Transgenomics) 2′-O-[(Triisopropylsilyl)oxy]methyl-ribonucleoside phosphoramidites (2′-O-TOM; Glen Research; also see UNITS 2.9 & 3.8) 30% ammonia Nitrogen Mobile phase A: 5% CH3CN in 0.1 M triethylammonium acetate (TEAA), pH 7.0 (DMTr-ON, HPLC) Mobile phase B: 100% CH3CN (DMTr-ON, HPLC) 1 to 400 mM sodium perchlorate in 20 mM Tris⋅Cl (pH 6.8; APPENDIX 2A)/25% formamide (DMTr-OFF, HPLC)
Synthesis of Modified Oligonucleotides and Conjugates
4.21.19 Current Protocols in Nucleic Acid Chemistry
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15% (w/v) polyacrylamide gel (APPENDIX 3D) containing 2 M urea in 0.5× TBE electrophoresis buffer (APPENDIX 2A) (DMTr-OFF, PAGE) 0.5 M LiClO4 (DMTr-OFF, PAGE) Matrix solution I: 40 mg/mL 2,6-dihydroxyacetophenone in methanol Matrix solution II: 80 mg/mL diammonium hydrogen citrate in water Water aspirator Screw-capped tube (Sarstedt) or vial Speedvac evaporator (Savant) Spin-X tube (Costar) Reversed-phase cartridges for DNA isolation (e.g., PolyPak, Glen Research; DMTr-ON) HPLC system (optional) with: Column: 3.9 × 300–mm Phenomenex Bondclone 10 C18 column (DMTr-ON) or 9 × 250–mm Dionex NucleoPac PA-100 column (DMTr-OFF) Detector: 254 nm (DMTr-ON) or 280 nm (DMTr-OFF) Lyophilizer Microcon tube (Millipore) or NAP-10 column (DMTr-OFF, PAGE) Additional reagents and equipment for automated solid-phase oligonucleotide synthesis (APPENDIX 3C) and purification of oligonucleotides (UNITS 10.1, 10.4, 10.5, 10.7 & APPENDIX 3B) 1. Dissolve the appropriate uridine 2′-carbamate phosphoramidite(s) in anhydrous CH3CN at up to 1.0 M. 2. Start the automated solid-phase oligonucleotide synthesis (APPENDIX 3C) from an appropriate solid support–filled column (e.g., on a 0.2- or 1-µmol scale). 3. Elongate the desired oligonucleotide chain using modified 2′-carbamate phosphoramidites (S.7a-j and S.7l) and standard 2′-deoxy-, 2′-OMe-, or 2′-O-TOM-ribonucleoside phosphoramidites in either DMTr-ON or -OFF mode. Allow at least 10 to 15 min for each 2′-carbamate phosphoramidite coupling. 4. After completion of the assembly, remove the column, wash with CH3CN (e.g., two times with 10 mL for 1-µmol synthesis), and quickly dry using a water aspirator. 5. Transfer the support into a screw-capped tube or vial and add 1 mL of 30% ammonia. 6a. For oligonucleotides prepared from phosphoramidites S.7a-j: Vortex the mixture and incubate at least 8 hr at 55°C or 24 hr at ambient temperature. 6b. For oligonucleotides prepared from phosphoramidite S.7l: Vortex the mixture and incubate at least 48 hr at 55°C per modified residue to ensure complete trimethylacetate ester deprotection. 7. Allow the solution to cool to ambient temperature, evaporate most of the ammonia under a stream of nitrogen, and dry the rest in vacuo using a Speedvac evaporator. For DMTr-ON oligonucleotides, it may be necessary to add 5 ìL TEA to completely preserve the DMTr group.
8. Add 0.5 mL deionized water, transfer the solution to the upper part of a Spin-X tube, and microcentrifuge 5 min at 13,000 rpm.
Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
9. Add 0.25 mL deionized water to the upper part of the tube, centrifuge again to wash, and then repeat. Concentrate the combined filtrates (sample and two washes) in vacuo.
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Current Protocols in Nucleic Acid Chemistry
10a. For DMTr-ON oligonucleotides: Purify on a reversed-phase cartridge following manufacturer’s recommendations (also see UNIT 10.7) or by RP-HPLC (UNIT 10.5) using the following recommended conditions: 3.9 × 300–mm Phenomenex Bondclone 10 C18 column Buffer A: 5% CH3CN in 0.1 M TEAA, pH 7.0 Buffer B: 100% CH3CN Gradient: 0% to 60% buffer B in 45 min Flow rate: 1 mL/min Detection: 254 nm. Pool the appropriate fractions, re-evaporate twice with water, and lyophilize. 10b. For HPLC of DMTr-OFF oligonucleotides: Purify by ion-exchange HPLC (UNIT 10.5) using the following recommended conditions: 9 × 250–mm Dionex NucleoPac PA-100 column Gradient: 1 to 400 mM sodium perchlorate in 20 mM Tris⋅Cl (pH 6.8)/25% formamide Flow rate: 1 mL/min Detection: 280 nm. Pool the appropriate fractions, desalt on a NAP-10 column, and lyophilize. 10c. For PAGE of DMTr-OFF oligonucleotides: Use denaturing gel electrophoresis (UNIT 10.4 & APPENDIX 3B) in 15% polyacrylamide containing 2 M urea in 0.5× TBE electrophoresis buffer. Cut the band(s) and elute the product with 0.5 M LiClO4. Desalt using a Microcon tube or a NAP-10 column, and lyophilize. 11. Dissolve the oligonucleotide in deionized water and quantitate by measuring UV absorbance at 260 nm. Store the solution up to 2 years frozen (i.e., −20°C). 12. Check the molecular mass of the oligonucleotides by MALDI-TOF-MS (UNIT 10.1) using a 1:1 (v/v) mixture of matrix solutions I and II. Prepare matrix just before loading the samples onto a plate. For a discussion of hybridization (duplex stability) studies and fluorescence measurements using 2′-carbamate-modified oligonucleotides, see Background Information. The optimal hybridization buffer for these experiments is 10 mM sodium phosphate, pH 7.0 (APPENDIX 2A) containing 0.1 mM EDTA and 100 mM NaCl. See Figure 4.21.4 for sample spectra and Table 4.21.1 for melting temperatures in a duplex stability study.
COMMENTARY Background Information Application of nucleoside carbamates The studies that are described here were originally driven by a desire to develop a convenient method for the introduction of various ligands into ribozymes. Use of ligands such as polyamines and amino acids together with the hairpin ribozyme was investigated as a possible way to enhance the ribozyme’s RNA cleavage potential under low metal ion concentration (Stolze et al., 2001). Since nucleobases in the core region are often involved in noncovalent
interactions important for maintaining structure and function, the sugar moiety of nucleosides, particularly the 2′-hydroxyl groups, seem to be attractive for ligand attachment, especially because 2′-functionalization already has proven value in various applications (Zatsepin et al., 2002). There are numerous examples of 2′-O-alkylnucleoside derivatives and their introduction into oligonucleotides. An irritating shortcoming is a sometimes poor yield of 2′-O-alkylation. An interesting opportunity for 2′-functionalization is via a carbamate function, easily
Synthesis of Modified Oligonucleotides and Conjugates
4.21.21 Current Protocols in Nucleic Acid Chemistry
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5015.60
A 30000
CUCCCAGGCUCAAAT O
2′ O
O
2′ O
HN
HN
Counts
20000
10000
0 3000
5000 Mass (m/z )
6000
7000
CUCCCAGGCUCAAAT 40000
2′ O O HN
5050.15
B
4000
2′ O
O
HN
Counts
30000
20000
10000
0 3000
4000
C
7000
CUCCCAGGCUCAAAT 20000
2′ O
4958.24 Counts
6000
5000 Mass (m/z )
O
O
2′ O HN
HN
HN
HN
NH
NH
10000 NH2
0 3000
Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
4000
5000 Mass (m/z )
6000
NH2
7000
Figure 4.21.4 MALDI-TOF mass spectra of 2′-carbamate oligodeoxyribonucleotides containing two modified uridines CU*CCCAGGCU*CAAAT, where U* is from phosphoramidites S.7d (A), S.7e (B), and S.7h (C) (see Figures 4.21.1 and 4.21.2 and Table 4.21.1).
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Table 4.21.1 and RNA
Melting Temperatures of Duplexes of 2′-Carbamate Oligonucleotides with Complementary DNA
Sequence, 5′ to 3′
U* = U-2′-OCONHCH2R where R =
CTCCCAGGCU*C- Control, U* = 2′-deoxyuridine AAAT C≡CH 4-Iodophenyl 1-Pyrenyl 1,4,7,10,13-Pentaoxacyclopentadecan-2-yl CH2OCH2CH2OH (CH2)2[O(CH2)2]2O(CH2)3NH2 (CH2)2NH(CH2)3NH2 (CH2)2NH(CH2)4NH(CH2)3NH2 CU*CCCAGGCTC- Control, U* = 2′-deoxyuridine AAAT 4-Iodophenyl 1-Pyrenyl 1,4,7,10,13-Pentaoxacyclopentadecan-2-yl CH2OCH2CH2OH (CH2)2[O(CH2)2]2O(CH2)3NH2 (CH2)2NH(CH2)3NH2 (CH2)2NH(CH2)4NH(CH2)3NH2
generated from primary or secondary alcohols by successive treatment with 1,1′-carbonyldiimidazole (CDI) and an aliphatic amine. This chemistry has been used successfully for 5′modification of oligonucleotides (Wachter et al., 1986). Aliphatic 5′-carbamates are known to be stable to ammonia deprotection at 55°C, in contrast to aromatic nucleoside 3′-carbamates (Sproat and Brown, 1983). Oligonucleotides containing nucleoside 2′carbamates have been reported previously. Cytidine 2′-carbamate was obtained by CDI activation followed by amine treatment (Dubey et al., 2000). In general, 2′-carbamate modifications are detrimental to the stability of DNARNA duplexes (Freier and Altmann, 1997; Prhavc et al., 2001). Recently we have described the preparation of a number of uridine 2′-carbamates and their introduction into oligo2′-deoxy- and 2′-OMe-ribonucleotides, and studied their hybridization with complementary DNA and RNA. It was found that 2′-pyrene carbamate-modified oligonucleotides show interesting binding properties and remarkable
DNA
RNA
Tm (°C)
∆Tm/mod Tm (°C) (°C)
∆Tm/mod (°C)
56.4
—
59.1
—
50.0 49.9 54.7 49.8
−6.4 −6.5 −1.7 −6.6
54.5 54.7 57.8 53.9
−4.7 −4.5 −1.4 −5.3
50.1 49.8 51.5 51.4 56.2
−6.3 −6.6 −4.9 −5.0 —
53.7 53.9 53.4 55.1 58.8
−5.5 −5.3 −5.8 −4.0 —
53.4 57.5 53.5
−2.8 +1.3 −2.8
54.4 55.5 54.8
−4.4 −3.3 −4.0
52.8 52.6 53.1 54.1
−3.4 −3.6 −3.1 −2.3
56.2 55.5 55.9 59.0
−2.6 −3.3 −2.9 +0.2
DNA mismatch affinity (Korshun et al., 2002), and exhibit enhanced fluorescence when bound to complementary RNA, but not DNA (see Fig. 4.21.5). Synthesis of uridine 2′-carbamates and their phosphoramidites The authors developed a general approach for the preparation of uridine 2′-carbamates using 1,1′-carbonyldiimidazole (CDI) activation of the 2′-hydroxy group of 3′,5′Markiewicz-protected uridine (S.2; see Basic Protocol 1). The reaction with CDI in dry dichloromethane gives the corresponding activated imidazolide in nearly quantitative yield. Dichloromethane proved to be the best solvent for the transformation, whereas use of the more polar tetrahydrofuran and acetonitrile led to a much longer reaction time. Other investigators have described a successful preparation of 2′carbamates using disuccinimidyl carbonate (DSC) and triethylamine in acetonitrile (Prhavc et al., 2001). In the authors’ hands, however,
Synthesis of Modified Oligonucleotides and Conjugates
4.21.23 Current Protocols in Nucleic Acid Chemistry
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Fluorescence (arb. units)
25 20 15 10
3 5
2 1
0 350
400
nm
450
Figure 4.21.5 Fluorescence spectra of 2′-pyrene carbamate oligodeoxyribonucleotide CTCCCAGGCU*CAAAT (1) and its duplex with complementary DNA (2) and RNA (3) (see Table 4.21.1).
Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
DSC gave poorer results, requiring a higher excess of reagent and longer reaction times, while giving lower conversion yields. An additional advantage of CDI is that in many cases the imidazolide S.3 need not be isolated, but can be reacted in situ with an excess of amine. The corresponding Markiewicz-protected 2′-carbamates (S.4) are stable compounds that can be easily purified by column chromatography. A facile technique was developed to protect side-chain amino groups of di- and polyamine 2′-carbamate derivatives, which involved trifluoroacetylation without isolation of the unprotected intermediate (see Alternate Protocol). Initial attempts to remove the Markiewicz group from 2′-carbamates by tetrabutylammonium fluoride trihydrate (TBAF) in THF showed smooth silyl deprotection, but also the appearance of a by-product with similar mobility. After several hours of treatment, an equilibrium was reached where an ∼1:1 ratio of the two products was obtained. The side-product proved to be the corresponding 3′-carbamate, generated by carbamoyl migration in the 1,2diol system (Korshun et al., 2002). The possibility of carbamoyl migration has been discussed in a similar case, but the 3′-carbamate itself was neither detected nor isolated (Seio et al., 1998). Interestingly, those authors used a safer reagent by buffering TBAF solution with
acetic acid. In other cases, 0.1 M TBAF in THF was used (Dubey et al., 2000; Prhavc et al., 2001). In our hands, however, TBAF in THF always gave a mixture of regioisomers. However, using the alternative desilylating reagent triethylamine trihydrofluoride, only pure 2′carbamate product S.5 was obtained, even after overnight treatment. The following step of 5′-dimethoxytritylation required some modification. Although care was taken to get rid of most of the fluoride by repeated toluene/hexane washings, some of the latter apparently remained in the crude mixture, which thus consumed additional amounts of 4,4′-dimethoxytrityl chloride (DMTr⋅Cl; up to 3 eq). In one case (S.6e), this led to an increased formation of 3′,5′-ditritylated product and lower yield of the target compound. A new work-up procedure was recently adopted to obviate the need for higher excess of DMTr⋅Cl. It uses an excess of inexpensive ethoxytrimethylsilane to scavenge the remaining fluoride, with corresponding formation of only volatile products (Stetsenko, pers. comm.). Use of pyridine (0.5% v/v) instead of TEA as an additive for column chromatography of 5′DMTr-protected 2′-carbamates is strongly advised to prevent 2′,3′-carbamate migration on silica gel. The final 3′-phosphinylation went smoothly and uneventfully according to the published procedure (Caruthers et al., 1987).
4.21.24 Supplement 15
Current Protocols in Nucleic Acid Chemistry
Synthesis of oligonucleotides containing uridine 2′-carbamate residues Automated solid-phase oligonucleotide synthesis was accomplished using a manufacturer’s protocol that has been modified only by increasing the coupling time of the 2′-carbamate phosphoramidites to 10 to 15 min (see Basic Protocol 2). Uridine-2′-carbamate-modified oligonucleotides were routinely synthesized in DMTr-ON mode and isolated on a reversed-phase cartridge (PolyPak) following the supplier’s standard technique. The results were consistently good as judged from the MALDI-TOF mass spectra of doubly modified oligomers (see Fig. 4.21.4). Effect of 2′-carbamate modifications on duplex stability Melting studies with uridine-2′-carbamatemodified oligonucleotides (see Table 4.21.1) confirmed the overall destabilizing effect of the 2′-carbamate group in both the DNA-DNA and DNA-RNA series. Most carbamate modifications result in a considerable decrease in Tm values of the duplexes, but there was almost no correlation between the steric bulk of the carbamate group and the Tm of the corresponding duplexes. The least destabilizing is pyrene carbamate, especially in the case of DNA-DNA duplexes. As expected, a single modification close to the 5′-terminal position of the oligonucleotide caused less destabilization than one in the middle. A number of studies of pyrene attached to the 2′-position of individual nucleosides within duplexes have been published recently (Silverman and Cech, 1999; Yamana et al., 2001). Pyrene as well as other planar polyaromatic hydrocarbons often show a greater stabilizing effect in DNA-DNA duplexes than in DNA-RNA duplexes (Yamana et al., 2001). The pyrene 2′-carbamate modification is not an exception, giving a clear stabilizing effect of pyrene superimposed on the destabilizing influence of the 2′-carbamate group, while even the spermine carbamate, which should carry three positive charges under physiological pH, is only negligibly stabilizing in just one most favorable situation. That pyrene is more stabilizing in DNADNA duplexes than DNA-RNA duplexes might suggest the better fit of pyrene into the shallower DNA-DNA minor groove. This notion is strengthened by interesting fluorescent properties of pyrene 2′-carbamate oligonucleotides. The total fluorescence intensity upon duplex formation of single pyrene-labeled oli-
gomers with a complementary DNA does not change very much (see Fig. 4.21.5). By contrast, the binding to a complementary RNA gives a 5- to 30-fold increase in fluorescence for single-pyrene oligonucleotides and a 30fold increase for double-pyrene oligonucleotides. This gives evidence that the excited pyrene 2′-carbamate is effectively quenched in single-stranded oligodeoxyribonucleotides and in DNA-DNA duplexes, but not in DNARNA duplexes. Similar examples of fluorescence increases upon hybridization with a complementary RNA are demonstrated for probes containing 2′-attached pyrene (Yamana et al., 2001). Thus, uridine 2′-carbamate modification in a DNA strand is usually destabilizing for both DNA-DNA and DNA-RNA duplexes, and therefore might be less suitable for those cases where higher stability of a duplex is preferential (e.g., antisense applications). However, the 2′carbamate modification affords a convenient way to place various functional groups into the minor groove of a DNA-DNA duplex (e.g., crown ether for metal ion binding, or an aliphatic amino group for postsynthetic labeling with activated derivatives of fluorescent dyes). Other possible applications include the specific minor groove delivery of reactive functionalities (e.g., for site-directed protein cross-linking or conjugation with peptides or other biomolecules), as well as modifications of flexible parts of highly structured RNA molecules for studies of ribozyme catalysis or RNA folding.
Critical Parameters and Troubleshooting The synthesis of 2′-carbamate compounds (see Basic Protocol 1 and Alternate Protocol) is relatively short, straightforward, and fairly efficient. However, careful attention to details of basic organic synthesis procedures is required. Preparation of the various compounds requires prior experience with routine chemical laboratory techniques such as solvent evaporation, extraction, TLC, and column chromatography. HPLC and gel electrophoresis are the most widely used methods of modified oligonucleotide isolation. Characterization of the products demands knowledge of 1H and 31P NMR, UV and MALDI-TOF mass spectroscopy. General laboratory safety is also of primary concern when hazardous materials are involved. Strict adherence to the outlined methods is therefore highly recommended.
Synthesis of Modified Oligonucleotides and Conjugates
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Anticipated Results Good to moderate yields of the final uridine 2′-carbamate 3′-phosphoramidites (S.7) from the parent Markiewicz-protected nucleoside (S.2) are expected following these procedures (see Basic Protocol 1 and Alternate Protocol). Analogously, others have reported good yields of 2′-carbamate nucleosides obtained by both similar (Dubey et al., 2000) and different chemistries (Prhavc et al., 2001). Basic Protocol 2 is intended to make use of the 2′-carbamate phosphoramidites to synthesize, purify, and characterize 2′-carbamate-containing oligonucleotides.
Time Considerations The synthesis of the 2′-carbamate 3′-phosphoramidites starting from Markiewicz-protected uridine (S.2) can be accomplished in 2 to 3 weeks per amine derivative, depending on the nature of the side chain and whether an additional protecting group is needed. The authors suggest, therefore, the preparation of a large batch of S.2 (from ∼30 mmol upwards) to use in portions as needed. The time for oligonucleotide synthesis varies from standard phosphoramidite methods only by the increased 2′-carbamate coupling time of 10 to 15 min per residue. The times needed for deprotection, isolation, purification, and analysis of modified oligonucleotides are the same as with standard techniques.
Literature Cited Caruthers, M.H., Barone, A.D., Beaucage, S.L., Dodds, D.R., Fisher, E.F., McBride, L.J., Matteucci, M., Stabinsky, Z., and Tang J.-Y. 1987. Chemical synthesis of deoxyoligonucleotides by the phosphoramidite method. Methods Enzymol. 154:287-313. Dubey, I., Pratviel, G., and Meunier, B. 2000. Synthesis and DNA cleavage of 2′-O-amino-linked metalloporphyrin-oligonucleotide conjugates. J. Chem. Soc., Perkin Trans. 1:3088-3095. Freier, S.M. and Altmann, K.-H. 1997. The ups and downs of nucleic acid duplex stability: Structurestability studies on chemically-modified DNA:RNA duplexes. Nucl. Acids Res. 25:44294443. Korshun, V.A., Stetsenko, D.A., and Gait, M.J. 2002. Novel uridin-2′-yl carbamates: Synthesis, incorporation into oligodeoxyribonucleotides, and remarkable fluorescence properties of 2′-pyren-1-ylmethylcarbamate. J. Chem. Soc., Perkin Trans. 1:1092-1104. Uridine 2′-Carbamates: Facile Tools For Oligonucleotide 2′-Functionalization
McGee, D.P.C., Sebesta D.P., O’Rourke, S.S., Martinez, R.L., Jung, M.E., and Pieken, W.A. 1996. Novel nucleosides via intramolecular functionalization of 2,2′-anhydrouridine derivatives. Tetrahedron Lett. 37:1995-1998. Prhavc, M., Lesnik, E.A., Mohan, V., and Manoharan, M. 2001. 2′-O-Carbamate-containing oligonucleotides: Synthesis and properties. Tetrahedron Lett. 42:8777-8780. Rannard, S.P. and Davis, N.P. 2000. The selective reaction of primary amines with carbonyl imidazole containing compounds: Selective amide and carbamate synthesis. Org. Lett. 2:2117-2120. Seio, K., Wada, T., Sakamoto, K., Yokoyama, S., and Sekine, M. 1998. Chemical synthesis and properties of conformationally fixed diuridine monophosphates as building blocks of the RNA turn motif. J. Org. Chem. 63:1429-1443. Silverman, S.K. and Cech, T.R. 1999. RNA tertiary folding monitored by fluorescence of covalently attached pyrene. Biochemistry 38:14224-14237. Sproat, B.S. and Brown, D.M. 1985. A new linkage for solid phase synthesis of oligodeoxyribonucleotides. Nucl. Acids Res. 13:2979-2987. Stolze, K., Holmes, S.C., Earnshow, D.J., Singh, M., Stetsenko, D.A., Williams, D., and Gait, M.J. 2001. Novel spermine-amino acid conjugates and basic tripeptides enhance cleavage of the hairpin ribozyme at low magnesium ion concentration. Bioorg. Med. Chem. Lett. 11:3007-3010. Wachter, A., Jablonski, J.-A., and Ramachandran, K.L. 1986. A simple and efficient procedure for the synthesis of 5′-aminoalkyl oligodeoxynucleotides. Nucl. Acids Res. 14:7985-7994. Yamana, K., Zako, H., Asazuma, K., Iwase, R., Nakano, H., and Murakami, A. 2001. Fluorescence detection of specific RNA sequences using 2′-pyrene-modified oligoribonucleotides. Angew. Chem. Int. Ed. Engl. 40:1104-1106. Zatsepin, T.S., Stetsenko, D.A., Arzumanov, A.A., Romanova, E.A., Gait, M.J., and Oretskaya, T.S. 2002. Synthesis of peptide-oligonucleotide conjugates with single and multiple peptides attached to 2′-aldehyde through thiazolidine, oxime and hydrazine linkages. Bioconjug. Chem. 13:822-830. Zhang, L., Cui, Z., and Zhang, B. 2003. An efficient synthesis of 3′-amino-3′-deoxyguanosine from guanosine. Helv. Chim. Acta 86:703-710.
Contributed by Vladimir A. Korshun Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry Moscow, Russia Dmitry A. Stetsenko and Michael J. Gait Medical Research Council Laboratory of Molecular Biology Cambridge, United Kingdom
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Current Protocols in Nucleic Acid Chemistry
Stepwise Solid-Phase Synthesis of Nucleopeptides
UNIT 4.22
Modification of oligonucleotides by attachment of peptide chains has attracted considerable interest during the last few years. The covalent union of a peptide chain may have beneficial effects on synthetic oligonucleotides (Tung and Stein, 2000), such as (1) increasing their stability to exonucleases, (2) accelerating or enhancing hybridization to their target sequences, and (3) facilitating their transport through cell membranes, which is particularly important for antisense (or antigene) applications (Gait, 2003). No rules are established to name and describe the different types of peptide-oligonucleotide conjugates. The word nucleoprotein is used to describe both covalently and noncovalently linked nucleic acid–protein complexes. Since in the former a phosphate group links the 5′- or 3′-terminal hydroxyl group of a nucleic acid with the side-chain hydroxyl group of serine, threonine, or tyrosine of a protein, phosphodiester-linked peptide-oligonucleotide conjugates have often been denoted as nucleopeptides (Schattenkerk et al., 1984; Robles et al., 1991; Waldmann and Gabold, 1997). Some authors, however, have used the term nucleopeptide to refer to other types of conjugates (UNIT 4.5; McMinn and Greenberg, 1999), and still others have referred to nucleopeptides as peptide-oligonucleotide conjugates (see below; Stetsenko et al., 2002). This unit describes stepwise solid-phase methods to obtain nucleopeptides containing 2′-deoxynucleosides and any of the proteinogenic amino acids, including the trifunctional ones. The procedures reported here are also suitable for the preparation of other peptideoligonucleotide conjugates, since most of the critical points of the procedures are related to the presence of purine nucleosides and some trifunctional amino acids in the target molecule. Methods for synthesis of the desired support-bound peptide (see Basic Protocol 1) are followed by methods for preparing the solid support (see Support Protocol 1) and several protected amino acids that are not commercially available (see Support Protocols 2 to 5). This is followed by extension of the oligonucleotide and cleavage and deprotection of the nucleopeptide (see Basic Protocol 2), and analysis, purification, and characterization of the target molecule (see Basic Protocols 3 and 4). Additional methods are given for reduction of nucleopeptides containing sulfoxide-protected methionine (see Support Protocol 6) and deprotection and purification of cysteine-containing nucleopeptides (see Support Protocol 7). STRATEGIC PLANNING Synthesis Strategy The synthesis of any type of peptide-oligonucleotide conjugate is a chemical challenge, because two completely different chemical entities must be assembled in the same molecule. Since the beginning of this work, the aim of the authors was to develop the simplest possible methodology for the preparation of nucleopeptides. In this respect, it was clear that the use of solid-phase chemistry would be one of the keys to simplifying the overall procedure. Both peptides and oligonucleotides are usually obtained by well-established solid-phase procedures, but the two methodologies differ considerably in many aspects. First, the coupling step is obviously different. In peptide synthesis, carboxylic acids have to be activated to form amide bonds with the free amine groups immobilized on the solid support, while elongation of oligonucleotide chains requires a phosphoric ester to be formed between two hydroxyl groups. Second, the protecting groups required at every
Contributed by Anna Grandas, Vicente Marchán, Laurent Debéthune and Enrique Pedroso Current Protocols in Nucleic Acid Chemistry (2004) 4.22.1-4.22.54 Copyright © 2004 by John Wiley & Sons, Inc.
Synthesis of Modified Oligonucleotides and Conjugates
4.22.1 Supplement 16
NH C CO
NH CH CO CHR O −
+
O P
O
CHR
base
O
O
HO
B
−
O P
O
O
O
B
O R = H : Ser R = CH3 : Thr
O
Figure 4.22.1 Base-promoted cleavage of nucleopeptides with serine- or threonine-nucleoside phosphodiester bonds.
step to mask the appropriate functionalities—and, consequently, the treatments necessary for their removal—are also quite different. Peptide chains are commonly obtained either by using groups that are labile to acids of different strength (the so-called Boc/benzyl strategy) or by using a combination of base-labile temporary α-amine-protecting groups and acid-labile permanent side-chainprotecting groups (Fmoc/tBu strategy). As most readers know, oligonucleotide synthesis combines the use of very mild acidic conditions to eliminate the temporary protecting groups (in order to prevent depurination) with that of a basic ammonia treatment to remove the permanent protecting groups (see UNIT 2.1). This scenario illustrates that the two methodologies cannot easily be made compatible, as the standard final treatment in peptide synthesis is a reaction in acidic conditions, under which an oligonucleotide chain would not survive. Another point of concern is the stability of the [hydroxylated amino acid]-nucleoside phosphodiester linkage, which can undergo a β-elimination process that degrades the target molecule (Shabarova, 1970). Figure 4.22.1 shows the β-elimination side reaction that may affect nucleopeptides (depurination is shown in Figure 2.1.8). The synthesis protocol described here is a stepwise approach (Fig. 4.22.2). It has been optimized so that the methodology is accessible to groups of nonspecialists in the field and, whenever possible, is compatible with commercially available synthons. Taking into account that peptide chains are normally elongated from the C to N terminus and oligonucleotides from the 3′ to 5′ terminus (although reverse elongation is also possible), elongation of the oligonucleotide chain at the side-chain hydroxyl of a resinlinked peptide seemed the best alternative. First, this allows greater stability, as the peptide is more robust than the oligonucleotide, particularly to acidic treatments. Second, this allows the linking amino acid to be placed at any position in the peptide chain. Stepwise assembly of the peptide moiety on a resin-linked oligonucleotide would only yield nucleopeptides with the linking amino acid at the C terminus. Moreover, difficulties in coupling amino acids onto resin-linked oligonucleotides have in some cases been reported (Bergmann and Bannwarth, 1995; Marchán et al., 2000). Stepwise Solid-Phase Synthesis of Nucleopeptides
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Current Protocols in Nucleic Acid Chemistry
A
−
+
Cl H3N CH
PS MBHA resin
CH3 1. 5% DIPEA/DCM 2. Fmoc-Phe-OH, DCC 3. Ac 2O Fmoc NH CH CO NH CH
PS
CH2 Ph IRAA
CH3 1. 20% piperidine/DMF 2. H-HMFS-OH, DCC
H-HMFS-
NH CO (CH2)2
CO NH CH CO NH CH
PS
CH2 Ph H O 1. Boc-Ala-OH, DCC, DMAP 2. Ac 2O, pyr
CH3
NH CO (CH2)2 CO NH CH CO NH CH
PS
CH2 Ph Boc
NH CH CO O CH3
CH3 Boc-Ala-HMFS-Phe-MBHA = Boc-Ala
Figure 4.22.2 (continues on next page) Stepwise solid-phase synthesis of nucleopeptide Ac-LysTrp-Lys-Hse(p3édGCATCG)-Ala-OH. (A) Preparation of the [C-terminal amino acid]-resin includes coupling of an IRAA and an HMFS linker (handle) to the resin (see Support Protocol 1), followed by coupling of the C-terminal amino acid (see Basic Protocol 1). (B) Peptide assembly (see Basic Protocol 1) followed by oligonucleotide assembly and final deprotection (see Basic Protocol 2) yield the crude target nucleopeptide. 1HOBt must be added to couple homoserine. Bases: B1 and B6, N2-isobutyrylguanin-9-yl; B2 and B5, N4-benzoylcytosin-1-yl; B3, N6-benzoyladenin-9-yl; B4, thymin1-yl. Other abbreviations: aa, amino acid; Boc, tert-butoxycarbonyl; CNE, 2-cyanoethyl; DCC, N,N′-dicyclohexylcarbodiimide; DCM, dichloromethane; DIPEA, N,N-diisopropylethylamine; DMAP, 4-(N,N-dimethylamino)pyridine; DMF, N,N-dimethylformamide; DMTr, 4,4′-dimethoxytrityl; dN, 2′deoxyribonucleoside; Fmoc, 9-fluorenylmethoxycarbonyl; For, formyl; HOBt, 1-hydroxybenzotriazole; Hse, homoserine; H-HMFS-OH, N-[9-(hydroxymethyl)-2-fluorenyl]succinamic acid; IRAA, internal reference amino acid; MBHA, p-methylbenzhydrylamine; NMI, N-methylimidazole; PS, polystyrene-co-1%-divinylbenzene; pyr, pyridine; TCA, trichloroacetic acid; Tfa, trifluoroacetyl; TFA, trifluoroacetic acid.
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B 1. 30% TFA/DCM 2. 5% DIPEA/DCM 4× 3. Boc-aa-OH, DCC 1 Boc Ala
Hse Ac Lys Trp
4. Ac 2O
Tfa For
Lys NH CH CO Ala CH2
Tfa
CH2 OH
H 1. 2. 3. 4.
DMTr-dN-O-P(OCNE)Ni Pr 2, tetrazole capping (Ac 2O, NMI) oxidation (1 M t -BuOOH) 3% TCA/DCM
n = 1–6
O
O
Bn
6× O O P OCH2CH2CN O Ac Lys Trp Tfa For
1:1 conc. NH 3 /dioxane overnight, 55°C
Lys Hse Ala Tfa
Ac-Lys-Trp-Lys-Hse(p 3′dGCATCG)-Ala-OH
Figure 4.22.2 continued.
Oligonucleotide assembly onto a resin-linked peptide is usually carried out with 3′-phosphoramidite nucleoside derivatives, which links the peptide to the 3′ end of the oligonucleotide. Substitution of 5′-phosphoramidites for the 3′-derivatives allows peptide-5′-oligonucleotides to be obtained.
Stepwise Solid-Phase Synthesis of Nucleopeptides
Choice of Protecting Groups The main concern in the overall process is the choice of protecting groups. They must be chosen so that the integrity of the target molecule is not affected by the final deprotection conditions. This means, as previously stated, avoiding the use of harsh acidic and basic treatments. Peptide assembly is compatible with the use of Nα-Boc-protected amino acids, and, although they are often commercially available with acid-labile side-chain-protecting groups, some derivatives with base-labile groups at the side chain are available. The only problem, then, is the peptide-oligonucleotide phosphate union, which in some cases can be particularly labile to bases. Since the data available at the outset of this work suggested that basic treatments could to some extent (though not completely) degrade nucleopeptides in which serine or threonine are the linking amino acids, it seemed possible to choose permanent protecting groups such that their removal under basic conditions would keep the side reaction to acceptable levels (<30%).
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Table 4.22.1
Recommended Permanent Protection Schemea,b All nucleopeptides
For base-stable nucleopeptides onlyc
Bz Ac Dmf —
Bz Bz i-Bu —
OFm — Fmoc (Fmoc)2 H, Tos (Dnp) Ac For (O)f S-tBug,h
OFm —c Tfa (Fmoc)2 Tos (Dnp) Ac For (O) —c
HMFS NH3/dioxane, room temperatureh
NPE NH3/dioxane, 55°C or (i) TBAF, (ii) NH3/dioxane, 55°C
Nucleobases A C G T Amino acids Asp, Glu Asn, Gln Lys Arg Hisd Ser, Thr, Tyre Trp Met Cys Other parameters Handle Cleavage conditions
aRefer to Figure 4.22.3 for abbreviations and structures of the amino acid protecting groups. Dash
indicates no protecting group needed. bAbbreviations for base protection: Ac, acetyl; Bz, benzoyl; Dmf, dimethylaminomethylene; i-Bu,
isobutyryl. cHomoserine or tyrosine nucleopeptides that do not contain Asn, Gln, or Cys. dHistidine is introduced protected with the tosyl (Tos) group, which is removed prior to oligonucleotide elongation. Dnp-protected histidine must be used if His is placed at the C-terminal position. Dnp removal requires overnight ammonia/dioxane treatment at room temperature. For further comments on histidine protection, see Strategic Planning. eThese residues are incorporated unprotected when they are the linking amino acid and protected as indicated when they are not. fThe crude nucleopeptide must be treated with N-methylmercaptoacetamide to reduce the sulfoxide to a thioether. gDTT must be added to the ammonia/dioxane solution to simultaneously remove the thiol protecting group. hThese very mild deprotection conditions must be used if serine or threonine are the linking residues or if the nucleopeptide contains asparagine and cysteine. In other circumstances, strongest deprotection treatments may be allowed (see Strategic Planning, discussion of final deprotection conditions).
All of the permanent protecting groups used in this procedure are shown in Table 4.22.1 and Fig. 4.22.3, and are discussed further below. They are chosen to be labile to the standard oligonucleotide-deprotecting reagent, aqueous ammonia. This is fully compatible with tyrosine nucleopeptides. Furthermore, if the nucleopeptide is not to reproduce the linking site of a naturally occurring nucleoprotein, the natural nonproteinogenic amino acid homoserine can be used as the linking residue (Fig. 4.22.2B), as first described in 1997 (Beltrán et al., 1997; Robles et al., 1997). No β-elimination can take place in this case, since the phosphate group is linked to the γ-carbon of the amino acid. Note, however, that the great tendency of homoserine to form γ-lactones precludes its use as the C-terminal amino acid. For the synthesis of serine and threonine nucleopeptides, the mildest final deprotection conditions must be used (see below).
Synthesis of Modified Oligonucleotides and Conjugates
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Amino acid
Side chain
Protected side chain
Structure of the protecting groups
Asp, Glu
(CH2)1-2 CO OH
(CH2)1-2 CO O Fm
Asn, Gln
(CH2)1-2 CO NH2
—
Lys
(CH2)4 NH Fmoc
Arg
Fm
(CH2)4 NH Tfa
(CH2)4 NH2
NH (CH2)3 NH
CO CF3
(CH2)3 NH NH2
NH Fmoc
N
His
CH2
N
N
CH2 NH
N
Ser
CH2 OH
CH2 O Ac
Thr
CH OH CH3
CH O Ac CH3
CH2
OH
CH2
Tos
CO
Dnp
SO2
O
Fmoc
N CH2
Tyr
Tfa
N Fmoc
NH
CH2
CH3 Tos NO2 Dnp
O2N O Ac
CO CH3
CH2
N For
Ac
O
Trp
For H
Met
CH2
CH2
Cys
CH2
SH
S CH3
O S CH3
CH2
CH2
CH2
S S t Bu
Figure 4.22.3 Structures of the unprotected and protected side chains of the trifunctional amino acids, and structures of the permanent protecting groups suitable for nucleopeptide synthesis. Abbreviations: Ac, acetyl; Dnp, 2,4-dinitrophenyl; Fm, 9-fluorenylmethyl; Fmoc, 9-fluorenylmethoxycarbonyl; For, formyl; Tfa, trifluoroacetyl; Tos, tosyl.
Solid Support Copolymers of styrene and 1% divinylbenzene are classic supports for peptide synthesis, while controlled-pore glass beads are typically used for the preparation of oligonucleotides. Polyethyleneglycol-polystyrene copolymers have been used in the synthesis of both types of molecules. In the authors’ hands, the best results have been achieved using the polystyrene-1%-divinylbenzene copolymer (Robles et al., 1999). The authors use the p-methylbenzhydrylamine resin (see Fig. 4.22.2A), but aminomethylpolystyrene should also work.
Stepwise Solid-Phase Synthesis of Nucleopeptides
Internal Reference Amino Acid Incorporation of an internal reference amino acid (IRAA; Fig. 4.22.2A) has two benefits. First, it allows one to adjust the substitution degree of the solid support to the desired level. Previous work (Montserrat et al., 1994) showed that substitution degrees of >0.20 mmol/g are not suitable for oligonucleotide synthesis, and that amine-functionalized polystyrene resins generally have higher loadings. The loading can be reduced by incompletely incorporating one amino acid and capping the unreacted amines. Use of Nα-Fmoc derivatives has the advantage that deprotection of an aliquot with piperidine and spectrophotometric quantitation of the resulting N-(9-fluorenylmethyl)piperidine easily allows the degree of substitution to be calculated.
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The second benefit of incorporation of an IRAA is that it allows some reaction yields to be determined. On one hand, the yield of incorporation of the C-terminal amino acid can be assessed by the ratio of C-terminal amino acid to IRAA obtained after acid hydrolysis and amino acid analysis of an aliquot of resin. On the other hand, the comparison of the peptide/IRAA ratio before and after cleavage allows one to determine the cleavage yield. Obviously, the IRAA must be different from the nucleopeptide amino acids. Handle The handle permanently protects the carboxyl group of the C-terminal amino acid (Fig. 4.22.2A). It is a bifunctional molecule with one carboxyl and one hydroxyl group. The former forms a stable amide bond with the IRAA and the latter an ester with the C-terminal amino acid. Cleavage of the nucleopeptide-resin ester linkage requires basic conditions to which the nucleopeptide remains stable. The authors have tested two different handles: 2-nitrophenyl ester (NPE; Eritja et al., 1991) and fluorenylmethyl ester (HMFS; Albericio et al., 2001). The fluorenylmethyl ester is more labile than the 2-nitrophenyl ester, and is therefore the most adequate to synthesize serine and threonine nucleopeptides. Since mild cleavage conditions are required to prevent peptide-related side reactions (as well as to prevent base-promoted peptide-oligonucleotide cleavage, see below), the HMFS handle is now routinely used in the authors’ laboratory and is the recommended option. Protecting Groups for the Peptide Chain Peptide assembly is carried out using the tert-butoxycarbonyl (Boc) group to temporarily protect the α-amino function. Removal of this protecting group requires treatment with 30% to 40% trifluoroacetic acid (TFA) in dichloromethane, which is fully compatible with the base-labile ester that links the C-terminal amino acid to the resin. Protection of the trifunctional amino acid side chains is discussed for each type of residue (see Table 4.22.1 and Fig. 4.22.3). Aspartic and glutamic acids The β- and γ-carboxyl groups of these residues are protected as 9-fluorenylmethyl (Fm) esters, which have the same lability as the nucleopeptide-HMFS linkage. Boc-Asp(OFm)OH and Boc-Glu(OFm)-OH are commercially available. Side reactions associated with aspartimide formation in some peptide sequences (most notably, Asp-Gly) may take place during the final deprotection treatment (Jeyaraj et al., 2002). Asparagine and glutamine The primary carboxamides of these residues’ side chains can be left unprotected. Phosphitylation of these groups by tetrazole-activated phosphoramidites has been shown to be virtually nil (Garcia de la Torre et al., 1999; Debéthune et al., 2002a), so there is very little risk of obtaining branched nucleopeptides. Final deprotection must be carried out using the mildest conditions (see below) to prevent side reactions associated with the 55°C ammonia treatment of asparagine-containing nucleopeptides (Debéthune et al., 2002a). Lysine The ε-amino function can be protected with either the trifluoroacetyl (Tfa) or 9-fluorenylmethoxycarbonyl (Fmoc) groups (Robles et al., 1999). Tfa can only be removed by ammonia treatment at 55°C, while milder basic treatments eliminate the Fmoc groups. Both derivatives are commercially available.
Synthesis of Modified Oligonucleotides and Conjugates
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Arginine The easiest way to prevent reactions at the guanidinium group is to keep it protonated. Washing with solutions of weak proton donors (e.g., 1-hydroxybenzotriazole, tetrazole) prior to amino acid or nucleoside coupling is required to ensure that the strongly basic guanidinium group is not free. Nevertheless, optimal results were achieved using the Boc-Arg(Fmoc)2-OH derivative (Debéthune et al., 2002a). Boc-Arg-OH⋅HCl can be purchased from commercial suppliers and the synthesis of Boc-Arg(Fmoc)2-OH is described in this unit (see Support Protocol 2). Histidine The imidazole ring must be protected during histidine incorporation to prevent racemization at the α-carbon. Two commercially available Boc-histidine derivatives can be used for nucleopeptide synthesis; they are imidazole-protected using either the tosyl (Tos) group or the 2,4-dinitrophenyl (Dnp) group (Beltrán et al., 1998). The tosyl group is labile to nucleopeptide deprotection conditions and can also be removed by treatment with 1-hydroxybenzotriazole (HOBt; Fujii and Sakakibara, 1974). Since the best-quality crude nucleopeptides were obtained when phosphoramidites were coupled onto histidine-unprotected peptide-resins, it is recommended that the experimenter (1) use Boc-His(Tos)OH for peptide synthesis, (2) treat with HOBt to deprotect the imidazole ring (if no HOBt has been used at any amino acid coupling step), and (3) proceed with the oligonucleotide assembly. In nucleopeptides with histidine at the C terminus, the imidazole ring should be kept protected throughout the entire synthesis process. This is best achieved using the Dnp group. Undesired detachment of the growing nucleopeptide chains from the resin, which can slowly take place if the imidazole ring is unprotected, can thus be avoided. The Dnp group is labile to fluoride treatment (see below) as well as ammonia treatment at either room temperature (overnight) or 55°C (6 hr). Serine, threonine, and tyrosine The side-chain hydroxyl of these amino acids (when they are not used as the linking residue) are protected with the acetyl (Ac) group (Fig. 4.22.3), which can be removed in the mildest final deprotection conditions. Boc-Ser(Ac)-OH and Boc-Tyr(Ac)-OH can be purchased from commercial suppliers. The preparation of Boc-Thr(Ac)-OH is described in this unit (see Support Protocol 5). Tryptophan Best results are achieved using the commercially available indole-protected derivative Boc-Trp(For)-OH, in which the indole ring is N-formylated (Robles et al., 1999). The formyl group can be removed by ammonia treatment at room temperature. Methionine The sulfide function of the methionine side chain can be alkylated during peptide elongation, and is oxidized at the nucleotide incorporation cycles. To prevent side reactions, the commercially available sulfoxide derivative Boc-Met(O)-OH is recommended (Marchán et al., 2000). After nucleopeptide synthesis and deprotection, the sulfoxide is reduced to a sulfide by reaction with N-methylmercaptoacetamide (see Support Protocol 6).
Stepwise Solid-Phase Synthesis of Nucleopeptides
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Cysteine Boc-Cys(S-tBu)-OH (where S-tBu is tert-butylthio), in which the thiol group has been transformed into a sterically hindered disulfide, allows cysteine-containing nucleopeptides to be obtained (Debéthune et al., 2002a). Care must be taken during the final deprotection step to avoid loss of HS-S-tBu by β-elimination. The target product is safely obtained if the final deprotection is carried out by treating the nucleopeptide-resin with ammonia at room temperature in the presence of 1,4-dithiothreitol (see Support Protocol 7). Boc-Cys(S-tBu)-OH can be easily obtained from the commercially available H-Cys(StBu)-OH derivative (see Support Protocol 3). Linking Amino Acid Serine, threonine, and tyrosine are introduced as Nα-Boc derivatives with the side-chain hydroxyl unprotected. No acylation of the free hydroxyl has been detected if the subsequent amino acids are incorporated using N,N′-dicyclohexylcarbodiimide (DCC) to activate the carboxyl group. Nα-Boc-protected homoserine derivatives are not available from commercial suppliers. The synthesis of the required compound is described here (see Support Protocol 4; Beltrán et al., 1997). This derivative, which is obtained as the triethylammonium salt Boc-Hse(DMTr)-O– HTEA+, will not couple onto the growing peptide chain unless a proton donor is added to the carbodiimide activator; therefore, this coupling is performed in the presence of HOBt. Peptide Assembly Peptide synthesis is carried out manually in polypropylene syringes fitted with polyethylene disks. An automatic synthesizer can also be used. As stated above, carbodiimides (N,N′-dicyclohexylcarbodiimide or N,N′-diisopropylcarbodiimide) are used to activate the carboxyl group of the incoming amino acid. 4-Dimethylaminopyridine must be added during the incorporation of the first (C-terminal) amino acid to achieve good esterification yields. Addition of HOBt is required to prevent side reactions during the activation and incorporation of asparagine, glutamine, and unprotected arginine, and to couple the homoserine derivative (HOBt can be replaced by tetrazole in this case). Some peptide sequences are particularly prone to form diketopiperazines (Pro-aa or aa-Pro, where aa is any amino acid, are typical examples; Gisin and Merrifield, 1972). This means that the free amine group of a dipeptide reacts with the resin-linked terminal carboxyl, releasing a cyclic dipeptide (diketopiperazine) and liberating free hydroxyl groups on the solid support. This unwanted side reaction can be minimized (Gairí et al., 1990) by reacting the third amino acid with the trifluoroacetate salt of the dipeptide-resin in the presence of a strong carboxyl activator reagent (PyAOP, 7-aza-benzotriazol-1-yloxytripyrrolidinephosphonium hexafluorophosphate) and an excess of N,N-diisopropylethylamine. Protecting Groups for the Oligonucleotide Moiety Standard DMTr and 2-cyanoethyl groups are used for temporary protection of the 5′-hydroxyl (or 3′-hydroxyl, if 5′-phosphoramidites are used) and for permanent protection of the phosphates, respectively. For the preparation of tyrosine and homoserine nucleopeptides, and if no side reaction–originating amino acids (Asn, Asp, or Cys) are present, standard nucleobase-protecting groups can be used: benzoyl (Bz) for A and C, and isobutyryl (i-Bu) for G. If base-labile nucleopeptides are to be obtained and/or the hybrid contains “delicate” amino acids, more labile nucleobase-protecting groups must be used. C is then protected with the acetyl (Ac) group, and the exocyclic amine of G is protected as formamidine with the dimethylaminomethylene group (Dmf). The structures of all of these protecting groups appear in Figures 2.1.7 and 2.1.10.
Synthesis of Modified Oligonucleotides and Conjugates
4.22.9 Current Protocols in Nucleic Acid Chemistry
Supplement 16
Oligonucleotide Elongation The oligonucleotide is assembled on an automatic DNA synthesizer (ABI 380B). Small modifications of the standard synthesis cycle are introduced to optimize performance with polystyrene resins (Bardella et al., 1990). The authors usually replace the standard iodine oxidation reagent with a 1 M solution of t-BuOOH, which facilitates washing of the resin after the oxidation step. Use of this reagent is absolutely mandatory when the nucleopeptide contains tyrosine or unprotected tryptophan, to prevent iodination of their aromatic side chains. Nucleopeptides with phosphorothioate internucleoside linkages have been obtained using the Beaucage reagent for the sulfurization step (Robles et al., 1999). Final Deprotection Conditions The polystyrene-divinylbenzene copolymer is hydrophobic and does not swell in aqueous medium. Consequently, treatments of nucleopeptide-resins with ammonia are not carried out using a 33% concentrated aqueous solution, but with a 1:1 (v/v) mixture of concentrated aqueous ammonia/dioxane. To cleave [base-stable nucleopeptide]-resin bonds, a two-step procedure is recommended: (1) treatment with tetrabutylammonium fluoride in anhydrous tetrahydrofuran, which detaches the nucleopeptide from the resin and removes most amino acid–protecting groups, and (2) treatment of the resulting filtrate with concentrated ammonia (or 1:1 ammonia/dioxane) to eliminate other remaining protecting groups. The authors have obtained better cleavage yields with this procedure than with an overnight treatment of the nucleopeptide-resin with ammonia at 55°C. The reaction with fluorides is a β-elimination process that yields the nucleopeptide with a C-terminal carboxylic acid function (Fig. 4.22.4A). The tyrosine-nucleoside phosphodiester bond remains unaffected by fluoride treatment.
A
O
H
NH CH C O R
base
+
O
NH CH C O − R
B
O
HO
NH CH C O R
NH3
+
O
NH CH C NH2 R
Stepwise Solid-Phase Synthesis of Nucleopeptides
Figure 4.22.4 Mechanisms of cleavage of the [C-terminal amino acid]-resin ester linkage. (A) Base-promoted β-elimination yields a C-terminal carboxylic acid function; (B) nucleophile-promoted cleavage yields a C-terminal carboxamide.
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Current Protocols in Nucleic Acid Chemistry
The nucleopeptide-HMFS linkage can also be cleaved under milder conditions by treatment with 1:1 ammonia/dioxane at room temperature. Although cleavage yields are lower than if the reaction is carried out at 55°C, this is the only alternative if the target nucleopeptide has serine or threonine as the linking residue, or if it contains amino acids (or peptide sequences) which are prone to give side reactions under these conditions (see above). An additional point of concern when the nucleopeptide-resin is treated directly with ammonia is that cleavage can take place either by β-elimination (Fig. 4.22.4A), which affords the nucleopeptide acid, or by nucleophilic attack of ammonia at the ester bond between the C-terminal amino acid and the HMFS handle (Fig. 4.22.4B). Nucleopeptides are then obtained as C-terminal carboxamides. Acid/carboxamide mixtures (in which the acid is the main component) are always obtained when glycine is the C-terminal amino acid (Debéthune et al., 2002a). The amount of amide is virtually nil when other residues occupy the C-terminal position. Summary As a general rule, the authors recommend assembling the nucleopeptide using a set of permanent protecting groups that can be removed using the mildest deprotection conditions, but little or no care has to be taken if tyrosine or homoserine is the linking amino acid and the nucleopeptide does not include “delicate” amino acids. However, even under the mildest possible final deprotection conditions, degradation of serine nucleopeptides by base-mediated cleavage of the serine-nucleoside phosphodiester bond can take place in some cases (Debéthune et al., 2002a). The data available indicate that this is a sequence-dependent reaction whose extent cannot as yet be predicted. From an operational point of view, the preparation of a nucleopeptide requires an adequately functionalized solid support and suitably protected amino acid and nucleoside derivatives. Access to an HPLC, a UV spectrophotometer, an electrospray or MALDITOF mass spectrometer, and an amino acid analyzer is required. Denaturing polyacrylamide gel electrophoresis (PAGE) facilities may also be necessary. STEPWISE MANUAL SOLID-PHASE SYNTHESIS OF THE PEPTIDE In this protocol, the C-terminal amino acid is first coupled to a solid support that has already been coupled with the internal reference amino acid (IRAA) and the HMFS linker (see Support Protocol 1 and Fig. 4.22.2A). Incorporation of the terminal amino acid is compared to the IRAA by amino acid analysis. After acylation of unreacted HMFS hydroxyl groups, the subsequent amino acids are added until the final support-bound peptide has been synthesized (Fig. 4.22.2B).
BASIC PROTOCOL 1
During synthesis, each coupling is monitored by a ninhydrin or chloranil test. The ninhydrin test (Kaiser et al., 1970) is only reliable for primary amines. For a qualitative coupling assay in the case of proline, the chloranil test (Christensen, 1979) should be used to assay for secondary amines. In general, the support-bound peptide (step 28) can be used directly for elongation of the oligonucleotide chain (see Basic Protocol 2). In some cases, especially with long peptides, analysis of the deprotected crude peptide by HPLC and mass spectrometry techniques (steps 29 to 41) is recommended to ensure peptide homogeneity before oligonucleotide elongation.
Synthesis of Modified Oligonucleotides and Conjugates
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Materials Derivatized p-methylbenzhydrylamine solid support (H-HMFS-IRAA-MBHA-PS; see Support Protocol 1) Dichloromethane (DCM; peptide synthesis grade) N,N-Dimethylformamide (DMF; peptide synthesis grade) t-Butoxycarbonyl-protected amino acids (Boc-aa-OH): Ala, Arg⋅HCl, Asn, Asp(OFm), Gln, Glu(OFm), Gly, His(Dnp), His(Tos), Ile, Leu, Lys(Fmoc), Lys(Tfa), Met(O), Phe, Pro, Ser, Ser(Ac), Thr, Trp(For), Tyr, Tyr(Ac), and Val (Novabiochem, Bachem, Neosystem) Boc-Arg(Fmoc)2-OH (see Support Protocol 2) Boc-Cys(S-tBu)-OH (see Support Protocol 3) Boc-Hse(DMTr)-O–HNEt3+ (see Support Protocol 4) Boc-Thr(Ac)-OH (see Support Protocol 5) N,N′-Dicyclohexylcarbodiimide (DCC) or N,N′-diisopropylcarbodiimide (DiPC) 4-Dimethylaminopyridine (DMAP) Methanol (MeOH; HPLC grade) Concentrated HCl, analytical grade Propionic acid Phenol, crystalline 3 M aqueous p-toluenesulfonic acid containing 0.2% tryptamine 0.06 M citrate buffer, pH 2 Acetic anhydride Benzoyl chloride (optional) Pyridine, synthesis grade 30% (v/v) trifluoroacetic acid (TFA; peptide synthesis grade) in DCM 5% (v/v) N,N-diisopropylethylamine (DIPEA) in DCM 1-Hydroxybentotriazole (HOBt; for coupling unprotected Arg, Asn, Gln, Hse) 0.5 M HOBt in DCM (for coupling unprotected Arg) (Azabenzotriazole-1-yl-oxy-tris-pyrrolidino)phosphonium hexafluorophosphate (PyAOP; for coupling any aa onto Pro) N,N-Diisopropylethylamine (DIPEA) Ethyl acetate (EtOAc) 1 M sulfuric acid Sodium sulfate (Na2SO4), anhydrous Ninhydrin reagents A and B (see recipes) Saturated chloranil solution: ∼0.75 g of 2,3,5,6-tetrachloro-1,4-benzoquinone in 25 mL toluene Acetone Peroxide-free dioxane (see recipe) 1 M 1,4-dithiothreitol (DTT) in peroxide-free dioxane (for S-tBu-protected Cys) 32% (v/v) ammonium hydroxide (store at 4°C) HPLC mobile phase A: 0.045% TFA in water (HPLC grade) HPLC mobile phase B: 0.036% TFA in acetonitrile (HPLC grade) α-Cyano-4-hydroxycinnamic acid (CHA) or dihydroxybenzoic acid (DHB) matrix 0.1% TFA in 1:1 (v/v) acetonitrile/water (HPLC grade)
Stepwise Solid-Phase Synthesis of Nucleopeptides
2- to 10-mL disposable polypropylene syringes with porous polyethylene discs and Teflon two-way stop cocks (RTV SF2, Shimadzu Scientific Research) Vacuum filtration system Teflon stir rod Desiccator Pyrex tubes for amino acid analyses Methane/oxygen flame
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110° and 155°C heating block Rotary evaporator equipped with a water aspirator or vacuum pump (use aspirator unless pump is indicated) 0.45-µm nylon filters Automatic amino acid analyzer (Beckman System 6300) 25-mL separatory funnel Filter paper 25- and 50-mL round-bottom flasks Small glass tubes for ninhydrin or chloranil tests 4-mL screw-cap pressure tubes, O-ring seal preferred Additional reagents for HPLC and MALDI-TOF-MS (see Basic Protocol 3 and Chapter 10) CAUTION: Trifluoroacetic acid is a very corrosive acid. Breathing the vapors is very harmful and it is possible to be quickly overcome by them. Always manipulate with extreme caution in a well-ventilated fume hood and wear appropriate protective clothing. Prepare syringe with solid support for peptide synthesis 1. Transfer an accurately weighed amount of H-HMFS-IRAA-MBHA-PS solid support for a 50-µmol synthesis into a 5- to 10-mL polypropylene syringe fitted with a porous polyethylene disc and equipped with a Teflon two-way stop cock. 2. Place the syringe on a vacuum filtration system. Incorporate C-terminal amino acid onto support 3. Wash the solid support using three 30-sec washes each of DCM, DMF, and then DCM again. For washes and for deprotection and neutralization treatments, a solvent volume of one or two times the volume of resin is recommended. For coupling, it is important to keep the reaction mixture as concentrated as possible. Coupling reactions must therefore be carried out in the minimum amount of solvent required to swell the resin and allow stirring. It is important to handle the polystyrene resin carefully to avoid mechanical degradation of the solid matrix. In all steps that require mixing, the resin should be stirred very carefully with a Teflon stir rod.
4. Weigh out the required amount of Boc-aa-OH (10 eq), DCC (10 eq), and DMAP (0.5 eq), and dissolve the mixture in a minimal volume of 1:1 (v/v) DCM/DMF. DiPC can replace DCC in all reactions described in this protocol and in the Support Protocols. The urea derivative formed after activation of the carboxyl group is more soluble in organic solvents. The esterification conditions employed for the incorporation of the C-terminal amino acid may cause some racemization, especially with amino acids such as unprotected histidine. In order to avoid this side reaction, glycine has been selected in most cases as the C-terminal amino acid. However, some problems may arise during the ammonium hydroxide deprotection (see Strategic Planning for more details).
5. Add the reaction mixture to the solid support and let stand 1.5 hr, occasionally stirring with a Teflon stir rod. 6. Wash the solid support successively with three 30-sec washes of DMF and three 30-sec washes of DCM. 7. Repeat steps 4 to 6 using 5 eq Boc-aa-OH, 5 eq DCC, and 0.25 eq DMAP and a reaction time of 1 hr. 8. Wash the solid support three times for 30 sec with MeOH and dry in a desiccator.
Synthesis of Modified Oligonucleotides and Conjugates
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Determine C-terminal amino acid incorporation: General procedure for amino acid analysis 9. Withdraw a 5- to 10-mg aliquot of dry resin and place it in a Pyrex tube. 10. Add 300 µl of 1:1 (v/v) concentrated HCl/propionic acid to the resin. When tyrosine is present, add a phenol crystal to the mixture to prevent decomposition of this amino acid during acid hydrolysis. Some amino acids, such as cysteine, homoserine, serine, threonine, and tryptophan, are partially or completely destroyed during acid hydrolysis. Arginine is partially converted to ornithine, and asparagine and glutamine to aspartic and glutamic acids, respectively. When the determination of some of these particular residues is necessary, enzymatic digestion must be carried out (see Basic Protocol 3, step 51). When determining tryptophan incorporation, hydrolysis must be carried out with a 3 M p-toluenesulfonic acid solution containing 0.2% of 3-(2-aminoethyl)indole (tryptamine), for 12 hr at 110°C in a vacuumsealed Pyrex tube.
11. Close the Pyrex tube hermetically using a methane/oxygen flame and heat 1.5 hr in a 155°C heating block. 12. Cool to room temperature, open the Pyrex tube, and evaporate the acidic mixture to dryness in a rotary evaporator equipped with a vacuum pump. CAUTION: Always wear gloves and glasses and work in a well-ventilated fume hood when carrying out these operations. Be careful when closing and opening the tube.
13. Redissolve the dry residue in 500 µL of 0.06 M citrate buffer, pH 2, and filter the resulting solution through a 0.45-µm nylon filter. Inject in an automatic amino acid analyzer. The amount of sample necessary for carrying out the amino acid analysis depends on the analyzer. For some analyzers, 5 nmol amino acid is optimal. In that case, injection of 50 ìL of the resulting solution will give 2 to 10 nmol of each residue. Prior to analyzing the sample, a standard that contains all 20 natural amino acids is analyzed to calculate a response factor for each residue under the analysis conditions employed. The response factors are used to calculate the amount of each residue in the sample.
14. Compare incorporation of the C-terminal amino acid to incorporation of the IRAA. If (C-terminal amino acid/IRAA) < 0.9, repeat steps 7 to 13. If 0.9 ≤ (C-terminal amino acid/IRAA) ≤ 1, proceed with step 15. In general, a nearly quantitative incorporation is achieved.
Acylate unreacted hydroxyl groups of HMFS linker 15. Add the required amount of acetic anhydride or benzoyl chloride (10 eq), pyridine (10 eq), and DCM to the solid support and let stand 10 min with occasional stirring using a Teflon rod. As for the coupling steps, a minimum volume of DCM is used for this step. The volume should be sufficient to swell the resin and allow stirring with the Teflon rod.
16. Wash the resin three times for 30 sec with DCM and repeat steps 15 and 16 to ensure that all unreacted hydroxyl groups are capped. 17. Wash the resin successively with three 30-sec washes each of DMF, DCM, and MeOH. Dry directly on a vacuum filtration system.
Stepwise Solid-Phase Synthesis of Nucleopeptides
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Incorporate subsequent amino acids 18. Add the required volume of 30% (v/v) TFA in DCM (one or two times the resin volume) and let stand 5 min with occasional stirring using a Teflon rod. 19. Wash with DCM, add an equal volume of 30% TFA in DCM, and let stand 25 min with occasional stirring. 20. Wash thoroughly with DCM, then wash four times for 60 sec with 5% (v/v) DIPEA in DCM, and five times for 30 sec with DCM. 21. Weigh out the required amount of Boc-aa-OH (3 eq) and DCC (3 eq), and dissolve the mixture in the minimum quantity of 1:1 (v/v) DCM/DMF. Add to the solid support and let stand 1.5 hr with occasional stirring using a Teflon rod. Modify as necessary for the following amino acid–specific conditions. When coupling arginine with the side chain protonated, weigh out the required amount of Boc-Arg-OH⋅HCl⋅2H2O (3 eq), DCC (3 eq), and HOBt (6 eq), and dissolve the mixture in the minimum amount of 1:1 (v/v) DCM/DMF. To avoid undesired side reactions, wash the resin four times for 60 sec with 0.5 M HOBt in DCM between steps 20 and 21. For asparagine or glutamine, weigh out the required amount of Boc-Asn-OH or Boc-GlnOH (3 eq), DCC (3 eq), and HOBt (3 eq), dissolve in the minimum quantity of 1:1 (v/v) DCM/DMF, and let stand 10 min at 0°C before adding it to the resin. When coupling the second amino acid onto Pro-resin, skip the neutralization step with 5% (v/v) DIPEA in DCM (step 20), weigh out the required amount of Boc-aa-OH (5 eq) and PyAOP (5 eq), dissolve the mixture in the minimum quantity of 1:1 (v/v) DCM/DMF, and add it to the resin. Then add 10 eq DIPEA to the resin and let stand for 1.5 hr, stirring occasionally with a Teflon rod. An acidic additive is necessary for the incorporation of the triethylammonium salt of the homoserine derivative. Weigh out the required amount of Boc-Hse(DMTr)-O–HNEt3+ (3 eq), DCC (3 eq), and HOBt (3 eq), and dissolve the mixture in the minimum quantity of 1:1 (v/v) DCM/DMF. After incorporation onto the support, elimination of the tert-butoxycarbonyl group with trifluoroacetic acid removes the DMTr group, so it is possible to quantify the coupling yield by quantitating the released DMTr cation (see Basic Protocol 2, step 11). When using Boc-Thr(Ac)-O–DCHA+, a workup is necessary to isolate the protected amino acid in the neutral species. Suspend Boc-Thr(Ac)-O–DCHA+ in EtOAc (∼5 mL/mmol amino acid) and transfer to a 25-mL separatory funnel. Add the same volume of 1 M sulfuric acid. Shake to achieve complete dissolution of the amino acid derivative. Separate the organic layer, and extract the aqueous phase with more EtOAc (twice with 5 mL). Dry the organic layer over anhydrous Na2SO4. Add more Na2SO4 if the salt crystals clump together upon swirling. Gravity filter the resulting solution through filter paper into a 50-mL round-bottom flask. Rinse the Na2SO4 crystals with 5 to 10 mL EtOAc. Remove the solvent under reduced pressure (i.e., in a rotary evaporator) to yield a white solid. Perform several co-evaporations with DCM to eliminate traces of EtOAc.
22. Wash the resin successively with three 30-sec washes of DMF, DCM, and MeOH. Dry directly on a vacuum filtration system. Assess amino acid coupling 23. Withdraw a 0.5- to 1-mg aliquot of dry resin and place it in a small glass tube. 24a. For primary amines: Add 3 drops of ninhydrin reagent A and 1 drop of ninhydrin reagent B. Heat the tube 3 min in a 110°C heating block. Allow to cool to room temperature. Assess color and proceed to step 25. It is important to have good quality ninhydrin reagents in order to obtain a reliable result. See Reagents and Solutions for more specific details. This test was developed for the primary amines of amino acids and may not be reliable for other primary amines.
Synthesis of Modified Oligonucleotides and Conjugates
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24b. For proline: Add 5 drops saturated chloranil solution and 20 drops acetone. Stir 5 min at room temperature. Assess color and proceed to step 25. 25a. For negative tests: If the reaction mixture is yellow (indicative of a successful amino acid coupling), proceed with the incorporation of the next amino acid (step 26). 25b. For positive tests: If the reaction mixture is blue for ninhydrin or green-blue for chloranil (indicative of the presence of free primary or secondary amines, respectively), repeat the coupling of the amino acid using the same amounts and conditions (steps 18 to 22) and then repeat the ninhydrin or chloranil test. 25c. For repeated positive tests: If a blue or green-blue color persists after a second coupling of the amino acid, cap the unreacted amines by adding acetic anhydride (10 eq), DIPEA (10 eq), and DMF to the solid support. Let stand 10 min, stirring occasionally with a Teflon rod. Wash the resin sequentially using three 30-sec washes each of DMF followed by DCM. Repeat the ninhydrin or chloranil test to confirm that all unreacted amines are capped before proceeding with the incorporation of the next amino acid (step 26). 26. Perform incorporation of the second and subsequent amino acids (steps 18 to 22), confirming the success of each coupling (steps 23 to 25) before proceeding to the next. Some minor modifications have to be introduced depending on the desired functional group at the N-terminal position of the peptide. If the amino group has to be acylated, remove the tert-butoxycarbonyl group and acylate by treating with acetic anhydride (3 eq) in DMF for 15 to 25 min. A longer treatment could cause undesired acetylation of the free side chain of the hydroxylated amino acid. When a free N-terminal position is desired, incorporate the last residue as an Fmoc-protected amino acid instead of a Boc derivative. The fluorenylmethoxycarbonyl group will be removed during the final ammonium hydroxide treatment. In some cases the tert-butoxycarbonyl group can be used to block the N-terminal position of the peptide. If that is the case, no further modification is necessary to the standard procedure because the Boc group is stable to the final ammonium hydroxide treatment.
27. Once the final (N-terminal) amino acid has been successfully coupled, proceed with the amino acid analysis as described in steps 9 to 13. When an Fmoc-amino acid derivative is introduced at the N-terminal position of the peptide, it is recommended to eliminate the Fmoc group before carrying out acid hydrolysis. This is because the Fmoc group may not be completely removed during the acid hydrolysis treatment, especially when it protects some sterically hindered residues such as isoleucine. For removal of the Fmoc group, treat the resin aliquot with 20% (v/v) piperidine in DMF as described below (see Support Protocol 1).
28. Calculate the relative ratio of the amino acids according to their peak areas and response factors (see step 13). If the peptide was correctly elongated, the ratio of residues will be close to the theoretical value.
Perform cleavage and deprotection (optional) 29. Take a 5- to 10-mg aliquot of dry peptide-resin and place it in a 4-mL screw-cap pressure tube, preferably with an O-ring seal. 30. Add 1 mL peroxide-free dioxane and 1 mL of 32% ammonium hydroxide solution, screw the cap on tightly, and leave 6 hr at room temperature. Stepwise Solid-Phase Synthesis of Nucleopeptides
If the peptide sequence contains trifluoroacetyl-protected lysine, perform the deprotection at 55°C for 12 to 15 hr. If it contains tert-butylthio-protected cysteine, use 1 mL of 1 M 1,4-dithiothreitol in dioxane and 1 mL of 32% ammonium hydroxide.
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Table 4.22.2 Typical Gradient and Mobile Phase for RP-HPLC of Deprotected Crude Peptidea
Elapsed time (min)
Percent mobile phase Bb
0 30 31 35 36 45
10 50 100 100 10 10
aGradient conditions are based on a flow of 1 mL/min using a Kromasil reversed-phase
C18 column (5-µm-diameter spherical silica, 4.0 × 250 mm) at room temperature with a 45-min overall cycle. Mobile phase A: 0.045% trifluoroacetic acid (TFA) in water; mobile phase B: 0.036% TFA in acetonitrile. bPercentage is at elapsed time.
31. Filter through a 2-mL polypropylene disposable syringe with a polyethylene or cotton filter and collect the solution in a 25-mL round-bottom flask. 32. Wash the pressure tube several times with MeOH and pass through the syringe. With tert-butylthio-protected cysteine-containing peptides, removal of 1,4-dithiothreitol is required prior to HPLC analysis. Dilute the crude sample to 2 mL by adding water and wash three times with 2 mL EtOAc. Proceed with the aqueous phase.
33. Concentrate the combined solution under reduced pressure to a volume of ∼0.5 mL. Dilute the deprotected crude peptide with 0.5 mL HPLC mobile phase B and take an aliquot (10 to 50 µL) for HPLC analysis. Analyze crude peptide by HPLC (optional) 34. Program the gradient system to start with 10% mobile phase B, increasing the percentage of mobile phase B with time (Table 4.22.2). Ensure that there is sufficient mobile phase to keep intakes covered throughout the run. For more specific details about HPLC analysis, see Basic Protocol 3 and UNIT 10.5.
35. Set the UV detector wavelength at 220 nm. Equilibrate the HPLC system with the starting mobile phase composition (10% mobile phase B) until a flat baseline is achieved at this detection wavelength. 36. Inject the sample by programming the autosampler or by loading it into the sample loop using the appropriate syringe. Elute the sample by starting the gradient. Record the chromatogram. Typically, small peptides will elute at ∼10 to 15 min under the recommended conditions (Table 4.22.2). When long peptides are prepared, it is recommended to use a gradient from 10% to 70% or 80% mobile phase B over 30 min and then wash as usual. Ammonium hydroxide treatment gives a mixture of the C-terminal carboxylic acid and carboxamide peptides, which usually elute in this order. Some UV-absorbing compounds are eluted at high acetonitrile percentages. These peaks are associated with handle-derived products formed during ammonium hydroxide treatment.
Synthesis of Modified Oligonucleotides and Conjugates
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Analyze crude peptide by MALDI-TOF-MS (optional) 37. Weigh 10 mg matrix (CHA or DHB) and place in a 1.5-mL microcentrifuge tube. A CHA matrix is recommended for small peptides (mol. wt. <500) and DHB for a wide range of masses.
38. Add 1 mL of 0.1% TFA in 1:1 (v/v) acetonitrile/water to the matrix. Vortex 30 sec to allow complete dissolution. Optimal results are obtained when the highest-purity matrices are used. Matrix solutions should be made fresh prior to use. Storage for more than 3 days is not recommended.
39. To a 0.5-mL microcentrifuge tube, add 1 µL peptide sample solution and 1 µL matrix solution. Mix by withdrawing and expelling the solution ten times with the pipet. 40. Spot 1 µL solution on a MALDI-TOF sample plate and dry. To allow homogeneous crystallization, do not disturb the spotted samples after they start to crystallize. 41. Perform peptide MALDI-TOF-MS analysis in positive mode. SUPPORT PROTOCOL 1
PREPARATION OF THE SOLID SUPPORT (H-HMFS-IRAA-MBHA-PS) This protocol describes the coupling of the internal reference amino acid and handle to the solid support used for nucleopeptide synthesis (see Fig. 4.22.2A). Additional Materials (also see Basic Protocol 1) p-Methylbenzhydrylamine resin: (α-amino-α-xylyl)-polystyrene reticulated with 1% divinylbenzene (f = 0.5 to 0.6 mmol/g; Novabiochem) 9-Fluorenylmethoxycarbonyl-protected internal reference amino acid (Fmoc-aa; Novabiochem, Bachem, Neosystem) 20% (v/v) piperidine in DMF N-[9-(Hydroxymethyl)-2-fluorenyl]succinamic acid (H-HMFS-OH; Alberico et al., 2001) 2- and 10-mL polypropylene syringes fitted with polyethylene discs Teflon two-way stopcocks 50-mL volumetric flasks Double-beam UV spectrophotometer, calibrated, and quartz cuvettes Wash solid support 1. Transfer an accurately weighed amount of p-methylbenzhydrylamine resin into a polypropylene syringe fitted with a polyethylene disc (i.e., 250 mg of resin into a 10-mL syringe). 2. Place the syringe on a vacuum filtration system. 3. Wash the solid support with three 30-sec washes of DCM. For general guidelines on volumes and handling resin, see Basic Protocol 1, step 3.
4. Add the required volume of 30% (v/v) trifluoroacetic acid in DCM and let stand 5 min, occasionally stirring with a Teflon rod. 5. Wash with DCM, add an equal volume of 30% (v/v) trifluoroacetic acid in DCM, and let stand 25 min. 6. Wash sequentially as follows:
Stepwise Solid-Phase Synthesis of Nucleopeptides
Three times for 30 sec with DCM Four times for 60 sec with 5% (v/v) DIPEA in DCM Five times for 30 sec with DCM.
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Incorporate internal reference amino acid 7. Weigh out the required amount of Fmoc-aa (0.5 eq) and DCC (0.5 eq), and dissolve the mixture in the minimum quantity of 1:1 (v/v) DCM/DMF. The internal reference amino acid must not be present in the target nucleopeptide sequence.
8. Add the reaction mixture to the solid support and let stand 25 min, stirring occasionally with a Teflon rod. Coupling time and amount of reactants are optimized to obtain a desired loading of ≤0.25 mmol/g. An increased coupling time will yield higher substitution degrees that are not adequate for oligonucleotide elongation.
9. Wash the solid support sequentially using three 30-sec washes each of DMF followed by DCM. Determine loading 10. Withdraw a 10-mg aliquot of dry resin and place it in a 2-mL polypropylene syringe fitted with a polyethylene disc and stopcock. 11. Add 1 mL of 20% (v/v) piperidine in DMF and let stand 3 min, occasionally stirring with a Teflon rod. 12. Open the stopcock and collect the filtrate in a 50-mL volumetric flask. Wash three times with 2 mL DCM and collect in the same flask. 13. Repeat the piperidine treatment for 7 min and wash again three times with DCM. Collect these filtrates together with the previous ones, and dilute with DCM to the 50-mL mark. Piperidine treatments quantitatively remove the fluorenylmethoxycarbonyl protecting group. An N-(9-fluorenylmethyl)piperidine (Fmp) adduct is formed during this treatment. Its molar extinction coefficient at 300 nm (ε300) is 7800. Its quantitation allows the loading of the solid support to be determined.
14. Prepare a reference solution by adding 2 mL of 20% (v/v) piperidine in DMF to a 50-mL volumetric flask and diluting to the mark with DCM. 15. Measure the sample absorbance at 300 nm (A300) on a calibrated, double-beam UV spectrophotometer using the above solution as a reference. If A300 >1, dilute sample as necessary to achieve a value <1. If using a single-beam spectrophotometer, a baseline should be run with a sample containing the reference solution (blank).
16. Calculate the solid support loading (f) as: f (mmol/g) =
A ×V ×1000 ε× l × m
Equation 4.22.1
where V is the total volume of the solution (in mL), ε is the molar extinction coefficient of the Fmp adduct at 300 nm (7800 L/mol/cm), l the absorbance path of the cuvette (in cm), and m is the amount of solid support (in mg). 17. If f ≈ 0.15 to 0.25 mmol/g, proceed according to step 18 with the entire support batch. If it is lower, repeat steps 7 to 9 with shorter coupling times.
Synthesis of Modified Oligonucleotides and Conjugates
4.22.19 Current Protocols in Nucleic Acid Chemistry
Supplement 16
Acylate unreacted amine groups and remove Fmoc 18. Add the required amount of acetic anhydride (10 eq), DIPEA (10 eq), and DMF to the solid support and let stand 10 min, stirring occasionally with a Teflon rod. 19. Wash the resin three times for 30 sec each with DCM and then repeat step 18. Perform the ninhydrin test with 0.5 to 1 mg of resin (see Basic Protocol 1, step 24a) to be sure that all unreacted amino groups are capped (solution should be yellow). 20. Wash the resin three times for 30 sec each with DMF, then DCM, and then MeOH. Dry directly on the vacuum filtration system. 21. Add the required volume of 20% (v/v) piperidine in DMF and leave 3 min, stirring occasionally with a Teflon rod. 22. Wash five times for 30 sec each with DCM and once again add the required volume of 20% (v/v) piperidine in DMF. Let stand 10 min, stirring occasionally with a Teflon rod. 23. Wash thoroughly three times for 30 sec each with DMF and then DCM. Dry directly on the vacuum filtration system. Incorporate handle 24. Weigh out the required amount of H-HMFS-OH (3 eq) and DCC (3 eq) and dissolve the mixture in the minimum quantity of 1:1 (v/v) DCM/DMF. The synthesis of N-[9-(hydroxymethyl)-2-fluorenyl]succinamic acid is carefully described elsewhere (see Albericio et al., 2001).
25. Add the reaction mixture to the solid support and let stand for 4 to 5 hr, stirring occasionally with a Teflon rod. 26. Wash the solid support successively (three times for 30 sec each) with DMF, then DCM, and finally MeOH. Dry directly on the vacuum filtration system. 27. Withdraw a 0.5- to 1-mg aliquot of dry resin and carry out the ninhydrin test (see Basic Protocol, step 24a). If the coupling is not complete (i.e., solution is blue, not yellow), repeat the handle incorporation using 1.5 eq for 1 to 2 hr. SUPPORT PROTOCOL 2
Stepwise Solid-Phase Synthesis of Nucleopeptides
PREPARATION OF Boc-Arg(Fmoc)2-OH Fmoc protection of Boc-Arg-OH is shown in Figure 4.22.5. Materials N-(tert-Butoxycarbonyl)-L-arginine hydrochloride (Novabiochem) Acetonitrile, anhydrous (see recipe) 9-Fluorenylmethyl chloroformate (Fmoc⋅Cl) Argon (Ar), dry Dichloromethane (DCM, anhydrous; see recipe) in a septum-sealed distillation collection bulb N,N-Diisopropylethylamine (DIPEA), anhydrous (see recipe) Chlorotrimethylsilane Dichloromethane (DCM; synthesis grade) Sodium sulfate (Na2SO4), anhydrous Methanol (MeOH) Acetic acid (AcOH) Hexanes 230- to 400-mesh silica gel
4.22.20 Supplement 16
Current Protocols in Nucleic Acid Chemistry
Boc NH CH COOH (CH2)3 NH H 2N
1. TMS⋅Cl, DIPEA 2. Fmoc-Cl
Boc NH CH COOH (CH2)3 NH
DCM +
NH2− Cl
Fmoc NH
N Fmoc
Figure 4.22.5 Protection of the guanidinium group of Boc-arginine. Abbreviations: Boc, tert-butoxycarbonyl; DCM, dichloromethane; DIPEA, N,N-diisopropylethylamine; Fmoc, 9-fluorenylmethoxycarbonyl; TMS, trimethylsilyl.
250-mL oven-dried round-bottom flask and rubber septum Desiccator containing P2O5 Glass syringe and needle, oven dried 40°C heat block 250-mL separatory funnel Gravity and vacuum filtration devices and filter paper 500-mL round-bottom flasks 2 × 5–cm silica-coated thin-layer chromatography (TLC) plates 254-nm UV light source Rotary evaporator with a water aspirator 5 × 25–cm glass chromatography column with solvent reservoir bulb Additional reagents and equipment for TLC (APPENDIX 3D), column chromatography (APPENDIX 3E), NMR, IR, and mass spectrometry Fmoc-protect Boc-Arg-OH 1. Place 1.0 g (3.04 mmol) of N-(tert-butoxycarbonyl)-L-arginine hydrochloride in a 250-mL oven-dried round-bottom flask. 2. Co-evaporate three times with anhydrous acetonitrile. 3. Weigh 2.4 g (9.28 mmol) of Fmoc⋅Cl and dry overnight under vacuum in a desiccator containing P2O5. 4. Cap the flask with a rubber septum and purge the flask with dry Ar. 5. Withdraw anhydrous DCM from a septum-sealed distillation collection bulb using an oven-dried glass syringe and needle. Add 25 mL DCM to the purged flask with magnetic stirring. 6. Add 1.6 mL (9.4 mmol) anhydrous DIPEA and slowly add 1.5 mL (11.9 mmol) chlorotrimethylsilane. 7. Stir the reaction mixture 90 min at 40°C. The original suspension slowly becomes a solution.
8. Cool in an ice bath. Dissolve 2.4 g (9.28 mmol) Fmoc⋅Cl in a minimal amount of dry DCM. Add 1.6 mL DIPEA and the Fmoc⋅Cl to the cooled reaction. 9. Stir the mixture 20 min in the ice bath and then 6.5 hr at room temperature. Synthesis of Modified Oligonucleotides and Conjugates
4.22.21 Current Protocols in Nucleic Acid Chemistry
Supplement 16
Work up reaction 10. Transfer the reaction mixture into a 250-mL separatory funnel and wash with 50 mL water. Wash the aqueous layer three times each with 50 mL DCM. 11. Dry the combined organic layer over anhydrous Na2SO4. Add more Na2SO4 if the salt crystals clump together upon swirling. When the solution is dry, the nonhydrated crystals will float in solution upon swirling.
12. Gravity filter the resulting solution through filter paper into a 500-mL round-bottom flask. Rinse the Na2SO4 crystals with 10 to 20 mL DCM. 13. Remove the solvent under reduced pressure (i.e., in a rotary evaporator with water aspirator) to yield an orange oil. Monitor reaction by TLC 14. Dissolve an aliquot of oil in a minimal amount of DCM, spot the solution onto a precut 2 × 5–cm TLC plate, and develop using 93:5:2 (v/v/v) DCM/MeOH/acetic acid (APPENDIX 3D). CAUTION: Wear protective eyewear.
15. Visualize the spots under a 254-nm UV light source. TLC analysis should indicate a main product with Rf = 0.54 that corresponds to the desired amino acid derivative. Minor amounts of fast-moving products are usually present in the mixture.
Isolate and characterize product 16. Dissolve the orange oil in 1.5 to 2 mL DCM and pour the solution onto 30 mL hexanes. A yellowish precipitate is obtained. This procedure partially eliminates the less polar contaminants.
17. Filter the precipitate under vacuum. 18. Prepare a 5 × 25–cm glass chromatography column with solvent reservoir bulb by adding a slurry of 50 g of 230- to 400-mesh silica gel in 1:1 (v/v) hexanes/DCM. Precondition with the same solvent. 19. Dissolve the crude material in a minimum amount of 1:1 hexanes/DCM and load onto the column. Perform chromatography using a small amount of air pressure (APPENDIX 3E) and eluting with solvent mixtures of increasing polarity: 1:1 (v/v) hexanes/DCM 25:75 (v/v) hexanes/DCM DCM 99:1, 98:2, 97:3, 96:4, and 95:5 (v/v) DCM/MeOH. 20. Combine the product-containing fractions into a 500-mL round-bottom flask and concentrate to a white solid (60% yield) on a rotary evaporator. 21. Determine the melting temperature. Characterize by TLC, 1H and 13C NMR, IR, and mass spectrometry. Boc-Arg(Fmoc)2-OH: Rf 0.54 (93:5:2 DCM/MeOH/AcOH); m.p. 106°-109°C; 1H NMR (300 MHz, CDCl3): δ 7.81-7.26 (m, 16H), 5.14 (m, 1H), 4.75 (m, 2H), 4.30 (m, 5H), 3.52 (m, 2H), 1.43 (s, 9H), 1.26 (m, 4H) ppm; 13C NMR (75 MHz, CDCl3): δ 161, 159, 157, 128, 120, 78, 68, 47, 44, 29, 28, 25 ppm; IR (film): 3800, 1720, 1620, 1515, 1460, 1260, 1110 cm–1; FAB-MS (positive mode, magic bullet, 3:1 dithiothreitol/dithioerythritol): m/z 719.4 [(Boc-Arg(Fmoc)2-OH)+H]+, calcd. mass for C41H42N4O8: 718.8. Stepwise Solid-Phase Synthesis of Nucleopeptides
4.22.22 Supplement 16
Current Protocols in Nucleic Acid Chemistry
H2N CH COOH CH2
t Bu S
S
(Boc)2O, NaOH
Boc NH CH COOH
dioxane, H 2O
CH2
t Bu S
S
Figure 4.22.6 Protection of the α-amine of H-Cys(S-tBu)-OH. Boc, tert-butoxycarbonyl.
PREPARATION OF Boc-Cys(S-tBu)-OH Boc-protection of cysteine is shown in Figure 4.22.6.
SUPPORT PROTOCOL 3
Materials S-(tert-Butylthio)-L-cysteine (H-Cys(S-tBu)-OH; Novabiochem) 2:1 (v/v) dioxane/water 1 M NaOH Di-tert-butyldicarbonate (Boc2O) Hexanes 1 M HCl Ethyl acetate (EtOAc) Sodium sulfate (Na2SO4), anhydrous 50-mL and 250-mL round-bottom flasks 100-mL separatory funnel Gravity filtration device and filter paper Rotary evaporator equipped with water aspirator Additional reagents and equipment for TLC (APPENDIX 3D), 1H NMR, and mass spectrometry Introduce Boc protecting group 1. Place 0.5 g (2.4 mmol) S-(tert-butylthio)-L-cysteine in a 50-mL round-bottom flask. 2. Suspend in 15 mL of 2:1 (v/v) dioxane/water and adjust the pH to 9 by addition of 1 M NaOH. 3. Add 0.60 g (2.74 mmol) di-tert-butyldicarbonate and keep the reaction overnight at room temperature with magnetic stirring. To avoid decomposition of di-tert-butyldicarbonate, store at –20°C.
Work up reaction 4. Transfer the reaction mixture to a 100-mL separatory funnel and wash twice with 50 mL hexanes. 5. Cool the aqueous phase in an ice bath and add 1 M HCl dropwise with manual stirring until the pH is 2 to 3. 6. Transfer the acidified aqueous phase to the 100-mL separatory funnel and extract three times with 50 mL EtOAc. 7. Wash the combined organic phases two times with 25 mL water. 8. Dry the organic layer over anhydrous Na2SO4. Add more Na2SO4 if the salt crystals clump together upon swirling.
Synthesis of Modified Oligonucleotides and Conjugates
4.22.23 Current Protocols in Nucleic Acid Chemistry
Supplement 16
9. Gravity filter the resulting solution through filter paper into a 250-mL round-bottom flask. Rinse the Na2SO4 crystals with 10 to 20 mL EtOAc. 10. Remove the solvent under reduced pressure (i.e., in a rotary evaporator with a water aspirator) to yield a white solid (80% yield). Use without further purification. 11. Determine the melting temperature. Characterize by TLC, 1H NMR, and mass spectrometry. Boc-Cys(S-tBu)-OH: Rf 0.47 (95:5 DCM/MeOH); m.p. 119°-120°C; 1H NMR (300 MHz, CDCl3): δ 5.40 (d, 1H, J = 7.8 Hz), 4.61 (m, 1H), 3.20 (m, 2H), 1.47 (s, 9H), 1.34 (s, 9H) ppm; FAB-MS (positive mode, 4-nitrobenzyl alcohol): m/z 309.4 [(Boc-Cys(S-tBu)OH)+H]+ and 332.4 [(Boc-Cys(S-tBu)OH)+Na]+, calcd. mass for C12H23NO4S2: 309.5. SUPPORT PROTOCOL 4
PREPARATION OF Boc-Hse(DMTr)-O–HTEA+ Boc protection and tritylation of homoserine are shown in Figure 4.22.7. Materials L-Homoserine (Novabiochem) Pyridine, anhydrous (see recipe) Argon (Ar), dry Chlorotrimethylsilane Triethylamine Di-tert-butyldicarbonate (Boc2O) 32% ammonium hydroxide 4,4′-Dimethoxytrityl chloride (DMTr⋅Cl) Dichloromethane (DCM), neutralized (see recipe) Methanol (MeOH) Triethylamine Acetonitrile, anhydrous (see recipe) 250-mL oven-dried round-bottom flask with rubber septum Oven-dried glass syringes and needles Rotary evaporator equipped with a water aspirator 500-mL round-bottom flask Additional reagents and equipment for column chromatography (see Support Protocol 2 and APPENDIX 3E), TLC (APPENDIX 3D), 1H and 13C NMR, and mass spectrometry Introduce Boc protecting group 1. Place 0.5 g (4.20 mmol) of L-homoserine in a 250-mL oven-dried round-bottom flask. 2. Dry by co-evaporating twice with 25 mL anhydrous pyridine. 3. Cap the flask with a rubber septum and purge the flask with dry Ar. 4. Add 10 mL anhydrous pyridine and 3.50 mL (25.2 mmol) chlorotrimethylsilane with oven-dried glass syringes and needles. 5. Let the reaction stand 2 hr under an Ar atmosphere at room temperature. 6. Add 1.1 g (5 mmol) di-tert-butyldicarbonate and 0.7 mL (5 mmol) triethylamine, and stir continuously 20 hr at room temperature under an Ar atmosphere. 7. Slowly add 10 mL cold water and 7 mL concentrated ammonium hydroxide (32% v/v), and stir 30 min at room temperature.
Stepwise Solid-Phase Synthesis of Nucleopeptides
The final ammonium hydroxide concentration achieved (∼ 2 M) will ensure complete removal of silyl groups.
4.22.24 Supplement 16
Current Protocols in Nucleic Acid Chemistry
H2N CH COOH CH2
1. TMS⋅Cl 2. (Boc) 2O, TEA
CH2
− +
Boc NH CH COO NH4 aq NH3
CH2 CH2
Pyr
OH
OH −
+
Boc NH CH COO HNEt3 CH2
DMTr⋅Cl, Pyr (silica gel chromatography with 2% TEA)
CH2 DMTr
O
Figure 4.22.7 Synthesis of the homoserine derivative. Abbreviations: Boc, tert-butoxycarbonyl; DMTr, 4,4′-dimethoxytrityl; Pyr, pyridine; TEA, triethylamine; TMS, trimethylsilyl.
8. Evaporate to dryness under reduced pressure (i.e., in a rotary evaporator) to yield an oil. 9. Dry the remaining oil by co-evaporating twice with 25 mL anhydrous pyridine. 10. Dissolve in 10 mL anhydrous pyridine and add 1.75 g (5.2 mmol) DMTr⋅Cl. 11. Stir 24 hr at room temperature under an Ar atmosphere. 12. Evaporate to dryness under reduced pressure. Isolate and characterize the product 13. Perform silica gel column chromatography (see Support Protocol 2, steps 18 to 19, and APPENDIX 3E), eluting with neutralized DCM and increasing amounts of MeOH (0% to 10%) in the presence of 2% triethylamine. 14. Combine the product-containing fractions in a 500-mL round-bottom flask and concentrate to an oil in a rotary evaporator. 15. Remove residual triethylamine by co-evaporating the oil with anhydrous acetonitrile to provide the pure product as a white solid (35% yield). 16. Determine the melting temperature. Characterize by TLC, 1H and 13C NMR, and mass spectrometry. Boc-Hse(DMTr)-O–HNEt3+: Rf 0.62 (9:1 DCM/MeOH); m.p. 67°-68°C; [α]D = 9.93 (c 0.62, MeOH); 1H NMR (300 MHz, CDCl3): δ 7.44-7.15 (m, 9H), 6.82-6.78 (m, 4H), 5.55 (d, 1H, J = 7.2 Hz), 4.22 (m, 1H), 3.77 (s, 6H), 3.30-3.10 (m, 2H), 2.98 (q, 6H, J = 7.3 Hz), 2.17 (m, 1H), 1.99 (m, 1H), 1.39 (s, 9H), 1.19 (t, 9H, J = 7.3 Hz) ppm; 13C NMR (75 MHz, CDCl3): δ 176.1, 158.3, 155.2, 145.1, 135.4, 130.0-126.5, 113.0, 86.1, 60.8, 55.1, 52.8, 45.0, 32.7, 28.3, 8.4 ppm; FAB-MS (positive mode, Xe, magic bullet, 3:1 dithiothreitol/dithioerythritol): m/z 623.1 [(Boc-Hse(DMTr)-O–HNEt3+)+H]+ and 522.1 [(BocHse(DMTr)-OH)+H]+, calcd. mass for C38H50N2O7 (Boc-Hse(DMTr)-O–HNEt3+): 622.8, calcd. mass for C30H35NO7 (Boc-Hse(DMTr)-OH): 521.6.
Synthesis of Modified Oligonucleotides and Conjugates
4.22.25 Current Protocols in Nucleic Acid Chemistry
Supplement 16
Boc NH CH COOH
NO2 , TEA
BrCH2
H3C CH
Boc NH CH COOCH2
NO2
H3C CH
EtOAc
OH
OH Ac2O, DMAP EtOAc
Boc NH CH COOH
H2, 5% Pd/C
H3C CH
EtOAc/MeOH
O Ac
Boc NH CH COOCH2
NO2
H3C CH O Ac
NH
(DCHA)
hexanes/ether +
Boc NH CH COO − HDCHA H3C CH O Ac
Figure 4.22.8 Protection of the threonine hydroxyl group and preparation of its solid dicyclohexylammonium salt. Abbreviations: Boc, tert-butoxycarbonyl; DCHA, N,N-dicyclohexylamine; DMAP, 4-(N,N-dimethylamino)pyridine; TEA, triethylamine.
SUPPORT PROTOCOL 5
PREPARATION OF Boc-Thr(Ac)-OH Preparation of the protected threonine residue is shown in Figure 4.22.8. Materials N-(tert-Butoxycarbonyl)-L-threonine (Novabiochem) Ethyl acetate (EtOAc) 4-Nitrobenzyl bromide Triethylamine 1 M HCl 5% (w/v) aqueous sodium hydrogencarbonate (NaHCO3) solution Sodium sulfate (Na2SO4), anhydrous Dichloromethane (DCM) Methanol (MeOH) Acetic anhydride 4-Dimethylaminopyridine (DMAP) Argon (Ar), dry 5% (dry basis) palladium on activated carbon Hydrogen gas Hexanes Diethyl ether Dicyclohexylamine
Stepwise Solid-Phase Synthesis of Nucleopeptides
4.22.26 Supplement 16
Current Protocols in Nucleic Acid Chemistry
100-, 250-, and 500-mL round-bottom flasks Reflux condenser Gravity filtration device and filter paper 250- and 500-mL separatory funnels Rotary evaporator with a water aspirator 25-mL oven-dried round-bottom flask with rubber septum Additional reagents and equipment for column chromatography (see Support Protocol 2 and APPENDIX 3E), TLC (APPENDIX 3E), 1H and 13C NMR, IR, and mass spectrometry Prepare Boc-Thr-ONbn 1. Place 2.5 g (12.5 mmol) of N-(tert-butoxycarbonyl)-L-threonine in a 100-mL roundbottom flask. Add 20 mL EtOAc. 2. Add 3.4 g (15.7 mmol) 4-nitrobenzyl bromide and 1.6 g (15.7 mmol) triethylamine and reflux 6 hr. 3. Cool the resulting solution and gravity filter through filter paper. 4. Transfer the filtrate to a 500-mL separatory funnel and wash once with 50 mL water, once with 50 mL of 1 M HCl, and three times with 50 mL of 5% aqueous NaHCO3 solution. 5. Dry the organic layer over anhydrous Na2SO4. Add more Na2SO4 if the salt crystals clump together upon swirling. 6. Gravity filter the resulting solution through filter paper into a 250-mL round-bottom flask. Rinse the Na2SO4 crystals with 10 to 20 mL EtOAc. 7. Remove the solvent under reduced pressure (i.e., in a rotary evaporator with a water aspirator). 8. Perform silica gel column chromatography (see Support Protocol 2, steps 18 and 19, and APPENDIX 3E), eluting with DCM and increasing amounts of MeOH (0% to 5%). 9. Combine product-containing fractions into a 500-mL round-bottom flask and concentrate to a white solid (70% yield) on a rotary evaporator. 10. Determine the melting temperature. Characterize by TLC (APPENDIX 3D), 1H and 13C NMR, IR, and mass spectrometry. Boc-Thr-ONbn: Rf 0.60 (95:5 CHCl3/MeOH); m.p. 95°-98°C; [α]D = –9.8 (c 0.62, MeOH); 1H NMR (300 MHz, CDCl3): δ 8.2 (d, 2H, J = 8.8 Hz), 7.5 (d, 2H, J = 8.8 Hz), 5.4 (m, 1H), 5.3 (d, 1H, J = 3.2 Hz), 4.4 (m, 2H), 1.5 (s, 9H), 1.3 (d, 3H, J = 6.6 Hz) ppm; 13 C NMR (75 MHz, CDCl3): δ 171, 156, 152, 143, 128, 124, 80, 68, 66, 59, 28, 20 ppm; IR (KBr): 3450, 1760, 1670, 1520, 1350 cm–1; FAB-MS (positive mode, Xe, magic bullet): m/z 355.1 [(Boc-Thr-ONbn)+H]+, 377.0 [(Boc-Thr-ONbn)+Na]+, calcd. mass for C16H22N2O7: 354.36.
Prepare Boc-Thr(Ac)-ONbn 11. Place 3.2 g (9 mmol) of Boc-Thr-ONbn in a 100-mL round-bottom flask. Dissolve in 20 mL EtOAc. 12. Add 4.3 mL (45 mmol) acetic anhydride and 0.55 g (4.5 mmol) DMAP and stir for 2 hr at room temperature. 13. Add 10 mL MeOH and stir 10 min. 14. Concentrate to an oil in a rotary evaporator. 15. Perform silica gel column chromatography as in step 8. 16. Combine the product-containing fractions in a 500-mL round-bottom flask and concentrate to an oil (95% yield) in a rotary evaporator.
Synthesis of Modified Oligonucleotides and Conjugates
4.22.27 Current Protocols in Nucleic Acid Chemistry
Supplement 16
17. Characterize by TLC, 1H and 13C NMR, IR, and mass spectrometry. Boc-Thr(Ac)-ONbn: Rf 0.84 (95:5 CHCl3/MeOH); [α]D = −7.2 (c 0.23, MeOH); 1H NMR (300 MHz, CDCl3): δ 8.2 (d, 2H, J = 8.8 Hz), 7.5 (d, 2H, J = 8.8 Hz), 5.4 (m, 1H), 5.2 (d,1H, J = 3.6 Hz), 4.5 (dd, 1H, J = 2.2, 9.4 Hz), 2.1 (s, 3H), 1.5 (s, 9H), 1.3 (d, 3H, J = 6.2 Hz), ppm; 13C NMR (75 MHz, CDCl3): δ 177, 170, 156, 148, 142, 129, 124, 81, 70, 66, 57, 28, 21 ppm; IR (film): 1750, 1710, 1610, 1530, 1350 cm−1; FAB-MS (positive mode, Xe, nitrobenzyl alcohol): m/z 397.0 [(Boc-Thr(Ac)-ONbn)+H]+, calcd. mass for C18H24N2O8: 396.40.
Prepare Boc-Thr(Ac)-OH (oil) 18. Place 0.95 g (2.4 mmol) Boc-Thr(Ac)-ONbn into a 25-mL oven-dried round-bottom flask. Cap the flask with a rubber septum and purge with dry Ar. 19. Dissolve in 10 mL of 1:1 (v/v) EtOAc/MeOH. 20. Add 0.18 mg of 5% palladium on activated carbon. Bubble hydrogen gas into the reaction mixture for 3 hr. CAUTION: Hydrogen is an extremely flammable gas. Keep the container in a well-ventilated place away from sources of ignition. Take precautionary measures against static discharges. Always manipulate in a well-ventilated fume hood and wear appropriate protective laboratory equipment.
21. Eliminate the catalyst by gravity filtration through filter paper. Rinse with 10 to 20 mL MeOH. 22. Concentrate the filtrate to dryness in a rotary evaporator. 23. Dissolve the resulting oil in 100 mL EtOAc and transfer to a 250-mL separatory funnel. 24. Wash the organic layer six times with 50 mL of 1 M HCl and then three times with 50 mL water. 25. Dry the organic layer over anhydrous Na2SO4. Add more Na2SO4 if the salt crystals clump together upon swirling. 26. Gravity filter the resulting solution through filter paper into a 250-mL round-bottom flask. Rinse the Na2SO4 crystals with 10 to 20 mL EtOAc. 27. Remove the solvent under reduced pressure (i.e., in a rotary evaporator). 28. Perform silica gel column chromatography as in step 8. 29. Combine the product-containing fractions into a 500-mL round-bottom flask and concentrate to an oil (55% yield) in a rotary evaporator. 30. Characterize by TLC, 1H and 13C NMR, and IR. Boc-Thr(Ac)-OH: Rf 0.71 (95:5 CHCl3/MeOH); 1H NMR (300 MHz, CDCl3): δ 5.75 (m, 1H), 4.39 (m, 1H), 4.26 (d, 1H, J = 8.6 Hz), 4.15 (m, 1H), 1.45 (s, 9H), 1.25 (d, 3H, J = 5.6 Hz) ppm; 13C NMR (75 MHz, CDCl3): 174, 170, 156, 81, 71, 57, 28, 21, 17 ppm; IR (film): 3400, 1750, 1710, 1520 cm–1.
Prepare Boc-Thr(Ac)-O–DCHA+ (solid) 31. Dissolve Boc-Thr(Ac)-OH in 1:1 (v/v) hexanes/diethyl ether and add an equimolar amount of dicyclohexylamine. Stir overnight at 0°C. Stepwise Solid-Phase Synthesis of Nucleopeptides
32. Filter and then wash with prechilled 1:1 (v/v) hexanes/diethyl ether to obtain a white solid (93% yield).
4.22.28 Supplement 16
Current Protocols in Nucleic Acid Chemistry
33. Characterize by TLC, 13C NMR, and mass spectrometry. Boc-Thr(Ac)-O–DCHA+: Rf 0.47 (85:10:5 CHCl3/MeOH/AcOH); m.p. 172°-173°C; 13C NMR (75 MHz, CDCl3): δ 174, 170, 156, 78, 73, 59, 53, 29, 28, 25, 24, 21, 17 ppm; FAB-MS (positive mode, Xe, magic bullet, 3:1 dithiothreitol/dithioerythritol): m/z 443.0 [(BocThr(Ac)-O–DCHA+)+H]+ and 465.0 [(Boc-Thr(Ac)-O–DCHA+)+Na]+, calcd. mass for C23H42N2O6: 442.60.
ELONGATION OF THE OLIGONUCLEOTIDE CHAIN AND CLEAVAGE AND DEPROTECTION OF THE NUCLEOPEPTIDE In this protocol, the peptide-bound support is added to an oligonucleotide synthesis column and the nucleotide is extended from the appropriate amino acid hydroxyl function (homoserine in Fig. 4.22.2B). In general, oligonucleotide synthesis follows standard automated DNA methods (see APPENDIX 3C); however, some minor modifications have to be introduced in the oligonucleotide synthesis cycle, since a polystyrene support is used instead of controlled-pore glass (CPG). The main washes are made with neutralized DCM rather than acetonitrile, and THF washes are used after certain steps. A 2% N,N-diisopropylethylamine solution is used to wash the support after detritylation. Washes with anhydrous acetonitrile are employed before phosphoramidite couplings. The synthesis cycle is outlined in Table 4.22.3. Oligonucleotide synthesis is monitored by the trityl assay, and the yield of the first incorporation is also determined.
BASIC PROTOCOL 2
Complete deprotection of the nucleopeptide can be carried out using ammonium hydroxide alone or using tetra-n-butylammonium fluoride (TBAF) followed by ammonium hydroxide. Ammonia treatment may give (especially with glycine at the C terminus) a mixture of the C-terminal carboxylic acid and carboxamide nucleopeptides. The TBAF procedure gives exclusively the C-terminal carboxylic acid nucleopeptide irrespective of the bifunctional spacer used. The TBAF treatment is recommended if nucleopeptides are sufficiently stable, but may not be compatible with nucleopeptides in which serine or threonine are the linking residues (see Strategic Planning). In some cases, presence of tetra-n-butylammonium counterions renders the reversed-phase HPLC analysis (see Basic Protocol 3) difficult. In those cases, it is recommended to obtain the sodium salt of the nucleopeptide as described below. PAGE analysis (see Basic Protocol 4) is also more reliable with desalted products. For additional requirements for deprotection, analysis, and purification, see Support Protocol 7. Materials Peptide-derivatized support (see Basic Protocol 1) DNA phosphoramidites (Glen Research): 5′-O-(4,4′-dimethoxytrityl)-N-protected2′-deoxyribonucleoside-3′-O-(2-cyanoethyl-N,N-diisopropyl)-phosphoramidites, where the nucleobases are: N6-benzoyladenin-9-yl N2-isobutyrylguanin-9-yl N2-dimethylaminomethyleneguanin-9-yl N4-benzoylcytosin-1-yl N4-acetylcytosin-1-yl thymin-1-yl Argon (Ar) and nitrogen (N2, optional) gas, dry Anhydrous acetonitrile (see recipe) in a septum-sealed distillation collection bulb Activator solution (see recipe) Cap A and B capping reagents (Glen Research) Oxidizer solution (see recipe) Peroxide-free THF (see recipe)
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Table 4.22.3 Automated 5-µmol Synthesis Cycle for the Oligonucleotide Chain Elongation on an ABI 380B DNA Synthesizer
Synthesis step
Function
Time (sec)
Detritylation 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22
DCM to waste DCM to column Ar reverse flush Ar block flush Waste port Advance fraction collector 3% TCA to waste 3% TCA to column Ar column flush 3% TCA to column Ar column flush DCM to waste DCM to column Ar column flush Ar block flush Waste to bottle 2% DIPEA to waste 2% DIPEA to column Ar reverse flush DCM to waste DCM to column Ar reverse flush
5 20 10 5 1 1 5 60 10 60 10 3 180 10 5 1 3 30 10 3 20 10
ACN to waste ACN to column Ar reverse flush Ar block flush ACN to waste ACN to column Ar reverse flush Ar block flush
5 45 30 5 5 45 45 5
Phosphoramidite preparation Activator to column Phosphoramidite + activator to column Activator to column THF to waste Wait THF to column Ar reverse flush Ar block flush
3 3 25 3 5 300 30 10 5
Column washing 23 24 25 26 27 28 29 30 Phosphoramidite coupling 30 31 32 33 34 35 36 37 38
continued Stepwise Solid-Phase Synthesis of Nucleopeptides
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Table 4.22.3 Automated 5-µmol Synthesis Cycle for the Oligonucleotide Chain Elongation on an ABI 380B DNA Synthesizer, continued
Synthesis step
Function
Time (sec)
Capping preparation Cap A + cap B to column Wait DCM to waste DCM to column Ar reverse flush Ar block flush
10 30 120 5 45 10 5
Column capping 39 40 41 42 43 44 45
Oxidation and column washing 46 47 48 49 50 51 52 53 54 55 56 57 58
Oxidizer to waste Oxidizer to column DCM to waste Wait THF to waste THF to column Ar reverse flush DCM to waste DCM to column Ar reverse flush Ar block flush Repeat steps 54 and 55 three times Ar block flush
7 30 5 60 5 60 10 5 30 10 5 5
Detritylation solution: 3% (w/v) trichloroacetic acid (TCA, 99%, Glen Research) in neutralized DCM 2% (v/v) N,N-diisopropylethylamine (DIPEA) in neutralized DCM Neutralized DCM (see recipe) 0.1 M p-toluenesulfonic acid monohydrate in acetonitrile 70% (v/v) perchloric acid Absolute ethanol (EtOH) Peroxide-free dioxane (see recipe) 32% ammonium hydroxide (store at 4°C) 1 M dithiothreitol (DTT) in peroxide-free dioxane Tetra-n-butylammonium fluoride (TBAF, Aldrich) Anhydrous tetrahydrafuran (see recipe) in a septum-sealed distillation collection bulb Methanol (MeOH), HPLC grade Glacial acetic acid 200- to 400-mesh Dowex 50WX4-400 ion-exchange resin (4.8 meq Na+/g; Fluka Chemic) 1 M NaOH Synthesis column for 5-µmol-scale synthesis: old empty OPC cartridge (ABI) with a body and two caps (13-mm aluminum seals; Chromatographic Specialties)
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Empty DNA synthesizer bottles, oven dried, with rubber septa Vacuum desiccator containing P2O5 Glass syringes and needles, oven dried ABI 380B automated DNA synthesizer External fraction collector and 15-mL test tubes Double-beam UV spectrophotometer, calibrated, and quartz cuvettes 20-mL screw-cap pressure tubes, O-ring seal preferred 55°C oven Disposable polypropylene syringes and polyethylene filters (cotton optional in some steps) 50-mL round-bottom flasks Teflon two-way stopcocks Rotary evaporator with a water aspirator Lyophilizer Vacuum filtration system Additional reagents and equipment for automated oligonucleotide synthesis (APPENDIX 3C) and amino acid analysis (see Basic Protocol 1) CAUTION: All solutions and reagents required for the DNA synthesizer should be manipulated and prepared in a well-ventilated fume hood. NOTE: Oxidizer solution, DIPEA solution, neutralized DCM, and peroxide-free THF should be filtered (Pro-XR nylon filters, 0.25 µm, 25 mm) before they are added to the synthesizer bottles. Prepare column 1. Transfer an accurately weighed amount of peptide-derivatized support for a 5-µmol synthesis (i.e., 100 mg peptide-support with a loading of 0.20 µmol/mg) to an assembled oligonucleotide synthesis column. The peptide-resin contains the hydroxyl function where the first nucleotide will be anchored and the oligonucleotide chain subsequently elongated. Standard 1-ìmol scale columns may fit enough a resin for a 1 to 2-ìmol-scale synthesis on polystyrene.
Perform oligonucleotide synthesis 2. Weigh the appropriate amount of DNA phosphoramidites in oven-dried synthesizer bottles, cap with rubber septa, insert a disposable needle, and dry them in a P2O5-containing desiccator under vacuum (6 hr minimum). 3. Open the desiccator under an Ar atmosphere. Using oven-dried glass syringes and needles, dissolve the compounds in sufficient anhydrous acetonitrile (from a septumsealed distillation collection bulb) to yield a 0.2 M concentration. Care must be taken to avoid the presence of moisture throughout the entire process.
4. Place the synthesizer reagents in the following ports of an ABI 380B synthesizer:
Stepwise Solid-Phase Synthesis of Nucleopeptides
Activator: port 9 Cap A and B: ports 11 and 12 Oxidizer: port 15 Peroxide-free THF: port 16 Detritylation solution: port 14 2% DIPEA in neutralized DCM: port 17 Neutralized DCM: port 18 Anhydrous acetonitrile: port 13 DNA phosphoramidite solutions (step 3): ports 1 to 4.
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5. Enter the sequence to be synthesized. Introduce an additional nucleotide at the 3′ end that will correspond to the peptide-resin containing the hydroxyl function where the oligonucleotide will be anchored. For example, to synthesize the tryptophan-containing nucleopeptide shown in Figure 4.22.2B, enter the sequence 5′-GCTACGT-3′, where the 3′-T corresponds to the peptidelinked resin.
6. Perform synthesis in the DMTr-off mode using a synthesis cycle modified as outlined in Table 4.22.3. Collect DMTr cation solutions in 15-mL test tubes using an external fraction collector. When nucleopeptides with >10 nucleobases are prepared, synthesis in DMTr-on mode is recommended in order to facilitate the purification.
7. Upon completion of the synthesis, dry the resin by manually conducting an Ar reverse flush operation on the synthesizer for 2 to 3 min. Alternatively, dry the support under a stream of N2 or Ar. Monitor DNA synthesis by DMTr assay 8. Dilute the first three and last three DMTr cation solutions to 10 mL with 0.1 M p-toluenesulfonic acid monohydrate in acetonitrile. Mix thoroughly. CAUTION: Handle the solution containing trichloroacetic acid in DCM and acetonitrile with gloves, because it is corrosive as well as toxic. Always manipulate in a well-ventilated fume hood. The volume of DMTr solutions may change during the course of a synthesis because DCM is a volatile solvent. Fractions may sit for several days before being assayed without affecting the results. Samples that have gone to dryness must be thoroughly redissolved. Addition of the p-toluenesulfonic acid ensures ionization of the DMTr group, making the solutions more strongly colored.
9. Dilute each sample 20- to 50-fold with the same solution. Measure the absorbance at 498 nm (A498) versus the 0.1 M p-toluenesulfonic acid monohydrate solution in acetonitrile. These dilutions are necessary because high absorbance measurements are not reliable.
10. Calculate the rough stepwise coupling efficiency for the synthesis as a whole, using the following equation: (stepwise efficiency) n =
avg. absorbance last three fractions avg. absorbance first three fractions Equation 4.22.2
where n is the number of DMTr nucleotides in the oligonucleotide (equal to the length of the oligonucleotide for DMTr-off synthesis). Quantify incorporation yield of the first nucleotide 11. Evaporate the first DMTr cation solution to dryness and redissolve the remaining residue in a known volume of 3:2 (v/v) 70% perchloric acid/EtOH. CAUTION: Handle 3:2 (v/v) 70% perchloric acid/EtOH with gloves in a well-ventilated fume hood. Perchloric acid is very corrosive and can be explosive when it contacts organic materials. Perchloric-containing residues must be appropriately neutralized prior to disposal.
12. Dilute the sample 20- to 50-fold with the same solution in order to obtain an A498 <1. Measure the A498 of the sample versus the 3:2 (v/v) 70% perchloric acid/EtOH solution.
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13. Calculate loading as: loading =
A498 × DF × 1000 ε × l × m
Equation 4.22.3
where DF is the dilution factor, ε is the molar extinction coefficient of the DMTr cation at 498 nm (71,700 L/mol⋅cm), l is the absorbance path of the cuvette (in cm), and m is the amount of peptide-support introduced in the column (in mg). 14. Determine the incorporation yield of the first nucleotide onto the peptide-support as the ratio between the obtained loading value and the peptide-support loading determined by amino acid analysis. Perform complete deprotection of nucleopeptide With ammonium hydroxide treatment 15a. Open the column and transfer the nucleopeptide-support to a 20-mL screw-cap pressure tube, preferably with an O-ring seal. 16a. Add 5 mL peroxide-free dioxane and 5 mL of 32% ammonium hydroxide. Screw the cap on tightly, and incubate 15 hr at 55°C in an oven or 6 to 8 hr at room temperature (see Strategic Planning for more details). When the nucleopeptide contains Cys(S-tBu), use 5 mL of 1 M DTT in dioxane and 5 mL of 32% ammonium hydroxide and treat for 6 to 8 hr at room temperature (see Support Protocol 7). CAUTION: The 32% ammonium hydroxide solution must be kept at 4°C with the cap tightly sealed. Concentrated ammonium hydroxide is extremely caustic. Breathing the vapors is harmful. Always wear glasses and gloves when using this compound and work in a well-ventilated fume hood, as it is possible to be quickly overcome by ammonia fumes and be blinded.
17a. Cool the mixture to room temperature. Filter through a disposable polypropylene syringe with a polyethylene filter or cotton, and collect in a 50-mL round-bottom flask. 18a. Wash the pressure tube several times with MeOH and pass the washes through the syringe. Concentrate the resulting filtrate under reduced pressure to a volume of ∼2 mL. To prevent inadvertent detritylation of dimethoxytritylated nucleopeptides, avoid heat and acid during the evaporation step. Elimination of dioxane and ammonium hydroxide should be done at room temperature to preserve the 5′-DMTr on the oligonucleotide chain. Addition of triethylamine is recommended to ensure no loss of the DMTr group.
With TBAF and ammonium hydroxide treatment 15b. Open the column and transfer the nucleopeptide-support into a polypropylene disposable syringe with a polyethylene filter closed with a stopcock. 16b. Add 5 mL of 0.05 M TBAF in anhydrous THF and let stand 30 min, stirring occasionally. Stepwise Solid-Phase Synthesis of Nucleopeptides
17b. Collect the filtrate in a 50-mL round-bottom flask. Wash three times with 2 mL MeOH followed by 5 mL THF and add to the filtrate. Add a few drops of glacial acetic acid and evaporate to dryness using a rotary evaporator.
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18b. Redissolve the residue in 10 mL of 1:1 (v/v) peroxide-free dioxane/32% ammonium hydroxide and place in a 55oC oven for 15 hr, or at room temperature for 6 to 8 hr, depending on the stability expected for the target nucleopeptides. Concentrate the resulting filtrate under reduced pressure to a volume of ~2 mL. Obtain sodium salt of nucleopeptide (optional; for TBAF only) 19. Add 200- to 400-mesh Dowex 50WX4-400 ion-exchange resin to a polypropylene syringe fitted with a polyethylene disc. 20. Wash with 200 mL water. 21. Wash with 200 mL of 1 M NaOH, added dropwise. 22. Wash with water until the solution has the pH of deionized water. 23. Dissolve deprotected nucleopeptide residue (step 18b) in a minimum amount of water (1 to 2 mL) and load onto the equilibrated Dowex resin. Elute with water until no UV-absorbing filtrate is obtained (check by measuring the absorbance at 260 nm versus water). Collect 1 to 2 mL fractions in disposable tubes. 24. Combine the product-containing fractions and lyophilize. Calculate cleavage yield 25. Wash the resin (resulting from ammonia or TBAF treatment; step 17a or 17b) with MeOH and dry directly on a vacuum filtration system. 26. Proceed with amino acid analysis as described (see Basic Protocol 1, steps 9 to 13). 27. Calculate the cleavage yield as the ratio between any amino acid in the nucleopeptide still attached to the solid support and the internal reference amino acid. Choose residues that are not destroyed or modified during the hydrolysis process, such as amino acids with aliphatic side chains, phenylalanine, or aspartic and glutamic acids.
ANALYSIS, PURIFICATION, AND CHARACTERIZATION OF NUCLEOPEPTIDES Crude nucleopeptides are first analyzed and then purified by liquid chromatography techniques. Analysis is routinely carried out using reversed-phase HPLC (RP-HPLC), although analytical PAGE (see Basic Protocol 4) can provide additional information, or may be required if the sample is highly impure. Purification is also generally performed by RP-HPLC. Medium-pressure systems (MPLC) may be the best option for relatively homogeneous crude nucleopeptides, since relatively large amounts of product (30 to 1000 OD units) can be purified in a single run. When this technique is inefficient, or if complicated crude nucleopeptides are obtained, the higher resolving power of high-pressure systems is required. Either analytical or semipreparative HPLC columns can be used, depending on the amount of pure product to be isolated. Preparative gel electrophoresis (see Basic Protocol 4) provides very good resolution and may allow pure nucleopeptides to be obtained when RP-HPLC fails. Its main disadvantage is low recovery of product from the gel. After purification, the overall synthesis yield is determined and the nucleopeptide is quantified by spectrophotometry at 260 nm. The nucleopeptide is then analyzed by MALDI-TOF or ESI mass spectrometry. Oligonucleotide analysis, purification, and characterization procedures are described in UNITS 10.1-10.7. Base composition is determined by digestion using a combination of alkaline phosphatase (AP) and either snake venom phosphodiesterase (SVPD) or calf spleen phosphodiesterase (SpPD). The assessment of base composition is more often carried out using SVPD than SpPD. Degradation with SVPD and AP can be performed in a single reaction mixture, because both enzymes work at the same slightly basic pH. Degradation with SpPD and AP requires two separate
BASIC PROTOCOL 3
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treatments under different conditions. The same information is generally obtained using either protocol, and consistent results from both reinforce that the nucleoside composition is as expected. Finally, the product is analyzed by amino acid analysis after acid hydrolysis or enzymatic digestion of the peptide moiety. Enzymatic digestion is carried out in a sequential mode using three enzymes: papaine, microsomal leucin-aminopeptidase, and prolidase. (1) Papaine is a nonspecific enzyme that cleaves at -X-↑-Y-, where Y is not specific and X can be any residue, but is preferably arginine, lysine, or residues consecutive to phenylalanine. (2) Microsomal leucin-aminopeptidase is an exopeptidase that hydrolyzes peptide bonds adjacent to a free α-amine group (H2N-X-↑-Y-, where X is not proline and Y is not specific). It does not hydrolyze γ-amino groups. (3) Prolidase is a dipeptidase that hydrolyzes dipeptides with a proline or hydroxyproline residue at the C terminus (H2NX-↑-Y-, where X is not proline, but Y is proline or hydroxyproline). When coupled with amino acid analysis, this protocol allows the content of asparagine and aspartic acid to be determined, as well as other acid-labile amino acids (e.g., methionine, tryptophan). Materials Crude deprotected nucleopeptide (see Basic Protocol 2 or Support Protocol 7) 0.01 M ammonium acetate, pH 7.0 HPLC mobile phase A: 0.01 M ammonium acetate, pH 7.0 HPLC mobile phase B: 1:1 (v/v) acetonitrile/water Methanol (MeOH), HPLC grade MPLC mobile phase A: 0.05 M ammonium acetate, pH 7.0 MPLC mobile phase B: 70% (v/v) 0.05 M ammonium acetate solution, pH 7.0, in 1:1 (v/v) acetonitrile/water Matrix: 2′,4′,6′-trihydroxyacetophenone (THAP) or 3-hydroxypicolinic acid (3-HPA) Ammonium citrate Acetonitrile, HPLC grade 0.1% (v/v) triethylamine in water Isopropanol (optional) 0.1 M Tris⋅Cl, pH 8.0 (APPENDIX 2A) 0.1 M MgCl2 1.5 U/500 µL snake venom phosphodiesterase (SVDP; Boehringer-Mannheim; EC 3.1.4.1) 0.0337 U/µL bacterial alkaline phosphatase (AP; Sigma; EC 3.1.16.1) 0.2 M ammonium acetate, pH 5.4 and 8.3 0.23 U/µL calf spleen phosphodiesterase (SpPD; Sigma; EC 3.1.3.1) Concentrated HCl 0.1 M ammonium acetate, pH 5.3 1:32 (v/v) 2-mercaptoethanol/water 20 mg/mL (286 U/mL) papaine (from Papaya latex; Sigma; EC 3.4.22.2) Glacial acetic acid 3 mg/mL (72 U/mL) microsomal leucin aminopeptidase (mLAP type VI-S from kidney pork microsomes; Sigma; EC 3.4.11.2) 0.025 M MnCl2 buffer solution, pH 8.3 5 mg/mL (875 U/mL) prolidase (from pork kidney; Sigma; EC 3.4.13.9)
Stepwise Solid-Phase Synthesis of Nucleopeptides
High-performance liquid chromatograph (HPLC) with: Injector (autosampler preferred), sample loop, and syringe (for manual loading) 0.1 to 5 mL/min pumping system (binary) UV/Vis detector, variable wavelength between 190 and 600 nm (preferred) or dual-wavelength detection
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Data integrating system Gradient system: displays and stores for redisplay and reformatting (preferred) or programmable Analytical column: reversed-phase C18 column (i.e., Kromasil, 10 µm, 4.0 × 250 mm) Semipreparative column: reversed-phase C18 column (i.e., Kromasil, 10 µm, 10.0 × 250 mm) Automatic fraction collector (optional) Lyophilizer Double-beam UV spectrophotometer, calibrated Quartz cuvettes Preparative chromatographic system (medium-pressure liquid chromatography, MPLC): Piston pump Reversed-phase C18-filled glass column (i.e., Vydac 15 to 20-µm i.d., 300-Å porosity, 22 × 2 cm) Automatic fraction collector UV/Vis detector with fixed-wavelength detection Teflon tubing connectors and adapters Gradient-forming device with two equal-diameter cylinders and a Teflon stopcock between them Chart recorder Pyrex tubes 37°C water bath or heating block Benchtop centrifuge Additional reagents and equipment for reduction of sulfoxide-protected methionine (see Support Protocol 6; optional), deprotection of tert-butylthio-protected cysteine (see Support Protocol 7; optional), MALDI-TOF-MS (UNIT 10.1), ESI-MS (UNIT 10.2), and amino acid analysis (see Basic Protocol 1) Analyze crude nucleopeptide by analytical RP-HPLC 1. Take 0.05 to 0.10 OD units of crude deprotected nucleopeptide and dilute with 0.01 M ammonium acetate buffer, pH 7.0, to a volume of 25 to 100 µL. Typical injection volumes for analysis are 10 to 100 ìL depending on the sample loop size of the injector. For nucleopeptides containing tert-butylthio-protected cysteine, perform deprotection (see Support Protocol 7) prior to the analysis of the crude product.
2. Program the HPLC gradient system to start with 5% HPLC mobile phase B, increasing the percentage of HPLC mobile phase B with time (Table 4.22.4). Ensure that there is sufficient mobile phase to keep intakes covered throughout the run. For 5′-dimethoxytritylated nucleopeptides 5% to 60% mobile phase B in 30 min is the gradient recommended. For post-HPLC detritylation, see UNIT 10.5.
3. Set the UV detector wavelength at 260 nm. Using an analytical column, equilibrate the HPLC system with the starting mobile phase composition until a flat baseline is achieved at the desired detection wavelength. 4. Inject the sample by programming the autosampler or by loading it into the sample loop using an appropriate syringe. Elute the sample by starting the gradient. Record the chromatogram.
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Table 4.22.4 Gradient and Mobile Phase for RP-HPLC of Crude Nucleopeptidea
Elapsed time (min)
Percent mobile phase Bb
0 30 31 35 36 45
5 35 100 100 5 5
aGradient conditions are based on a flow of 1 mL/min using an analytical
Kromasil reversed-phase C18 column (10-µm diameter spherical silica, 4.0 × 250 mm) at ambient temperature with a 45-min injection cycle. Mobile phase A: 0.01 M ammonium acetate, pH 7.0; mobile phase B: 1:1 (v/v) acetonitrile/water. bPercentage is at elapsed time.
Typically, nucleopeptides will elute at ∼12 to 18 min under the recommended conditions (Table 4.22.4). For 5′-dimethoxytritylated nucleopeptides, the elution time will be longer (∼18 to 25 min). In some cases, when the nucleopeptide is small (e.g., <8 nucleobases) and has been cleaved with ammonium hydroxide, it is possible to differentiate the nucleopeptides with C-terminal carboxylic acid and carboxamide groups. When the nucleopeptide contains sulfoxide-protected methionine or proline, it is possible to obtain two or more peaks (depending on the number of these residues) due to sulfoxide diastereomers or proline conformations (cis or trans).
5. For nucleopeptides containing sulfoxide-protected methionine, proceed with reduction (see Support Protocol 6) prior to purification. Purify crude nucleopeptide by analytical or semipreparative RP-HPLC 6a. For small amounts of nucleopeptide (1 to 10 OD units): Perform HPLC as in steps 1 to 4. Load the analytical column with ∼1 to 2 OD units at every injection and elute at a flow rate of 1 mL/min. Collect the desired fractions either with an automated fraction collector or by observing the chromatogram in real time and manually collecting the eluate. 6b. For larger amounts of nucleopeptide (>10 OD units): Use a semipreparative column under the described conditions. Elute at a flow rate of 3 mL/min. 7. Lyophilize the collected solution and quantify the amount of product obtained (steps 8 to 11). Quantify nucleopeptide 8. Dissolve the lyophilized nucleopeptide in 5 mL water. Take 20 µL and dilute to a volume of 1 mL with water. An adequate dilution factor has to be chosen to keep absorbance <1. Modify the dilution depending on the synthesis scale and expected yield.
9. Measure the absorbance of the diluted sample in a quartz cuvette at 260 nm (A260) on a calibrated, double-beam, UV spectrophotometer using water as a reference. Stepwise Solid-Phase Synthesis of Nucleopeptides
If using a single-beam spectrophotometer, a blank should be run with a sample containing water (blank).
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Table 4.22.5
Molar Extinction Coefficients of Nucleobases
Nucleobase (ni)
εi at 260 nm
T dC dG dA
8830 7700 11500 15200
10. Determine the amount of product (in OD260 units) and the concentration of the stock solution using the dilution factor and Beer’s Law (A260 = εlc), where ε is the molar extinction coefficient (see below), l is the path length of the UV cell (typically 1 cm), and c is the molar concentration of oligonucleotide. Lyophilize the nucleopeptide solution. In a nucleopeptide, the contribution of aromatic amino acids (tryptophan, phenylalanine, and tyrosine) to the global oligonucleotide absorbance at 260 nm is considered negligible in comparison with the absorption of the nucleobases. The molar extinction coefficient of the nucleopeptide is given by ε = εi × F, where εi represents the molar extinction coefficient of each nucleobase (see Table 4.22.5) and F is a correction factor (0.9 when the oligonucleotide is single-stranded and 0.8 when it is double-stranded or self-complementary).
11. Calculate the overall synthesis yield. From the ratio of product obtained (in OD units) to the theoretical amount (calculated from the loading of the C-terminal amino acid resin). Average yields for nucleopeptide synthesis, cleavage, and purification are ∼10% to 30%.
Purify crude nucleopeptide by preparative MPLC 12. Prepare a preparative chromatographic system as follows: a. Connect a piston pump to the top of a preparative reversed-phase C18-filled glass column. b. Connect the bottom to an automatic fraction collector through a UV/Vis detector. c. Connect the pump to the mixing chamber of a gradient-forming device. Connect the detector to a chart recorder using the appropriate ports. 13. Wash the column with 100 mL MeOH and 200 mL water by placing the pump entrance line in the appropriate bottles. 14. Set the UV detector wavelength at 260 nm with the appropriate lamp and filter. Turn on the detector and chart recorder. 15. Equilibrate the column with 200 mL MPLC mobile phase A at 3 to 4 mL/min. Stop the pump. 16. Place 600 mL MPLC mobile phase A and 600 mL MPLC mobile phase B in the appropriate compartments of the gradient-forming device. Place a magnetic stir bar in the mixing chamber (initially MPLC mobile phase A) and a magnetic stirrer below. 17. Place the pump solvent line inside the mixing chamber. 18. Dissolve the crude nucleopeptide sample in MPLC mobile phase A to a volume of 5 mL. 19. Remove the supernatant solution from the column cap position. Introduce the nucleopeptide sample solution onto the column and close.
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20. Turn on the pump (3 to 4 mL/min) and the magnetic stirrer. Open the stopcock of the gradient-forming device to generate the gradient. Turn on the automatic fraction collector. Preparative chromatographic elution profiles are very similar to analytical ones. The gradient conditions described above extrapolate the typical analytical gradient (from 5% to 35% HPLC mobile phase B in mobile phase A). When DMTr-on nucleopeptides are purified, MPLC mobile phase B should be changed to 40% 0.05 M ammonium acetate solution, pH 7.0, in 1:1 (v/v) acetonitrile/water.
21. Select the tubes containing the desired product. Reanalyze those solutions by analytical HPLC (steps 1 to 4). In general, the desired nucleopeptide (main chromatographic peak) is eluted within four or five tubes and has a high level of purity (>95% to 99% as assessed by analytical HPLC).
22. Pool the fractions with the same chromatographic profile and purity requirements. Lyophilize the collected solution and quantify (steps 8 to 11). Analyze nucleopeptide by MALDI-TOF-MS 23. Weigh 10 mg THAP matrix or 50 mg 3-HPA matrix into a 1.5-mL microcentrifuge tube. In a separate tube, weigh 50 mg ammonium citrate. Optimal results are obtained when the highest-purity matrices are used. Matrix and ammonium citrate solutions should be made fresh prior to use. Storage for >3 days is not recommended. In general, MALDI-TOF analysis of nucleopeptides is carried out following the same protocols as for simple oligonucleotides (UNIT 10.1). The standard matrices are used (THAP and 3-HPA) and negative-ionization mode is recommended because of the global negative charge of the nucleopeptides. For routine nucleopeptide analysis (up to 8 to 10 nucleobases), THAP is recommended. 3-HPA should be used when THAP does not give positive results.
24. Add 1 mL of 1:1 (v/v) acetonitrile/water to the THAP or 3-HPA matrix, and 1 mL water to the ammonium citrate salt. Vortex for 30 sec to allow complete dissolution. 25. Prepare a 50 to 100 µM nucleopeptide solution in water. 26. To a 0.5-mL microcentrifuge tube, add 1 µL nucleopeptide sample solution, 1 µL ammonium citrate solution, and 1 µL matrix solution. Mix by withdrawing and expelling the solution ten times with a pipet. 27. Spot 1 µL on the MALDI-TOF sample plate and dry. To allow for homogeneous crystallization, do not disturb the spotted sample after it starts to crystallize. Do not load the sample plate into the mass spectrometer before the plate is dry. Good signal can be obtained from anywhere around the edges of the crystallized spot.
28. Analyze by MALDI-TOF-MS (see UNIT 10.1). Analyze nucleopeptide by ESI-MS 29. Prepare a 100 to 200 µM stock nucleopeptide solution in water. The best concentration for carrying out ESI-MS analysis depends on the instrument used. In general, nucleopeptides are analyzed by ESI-MS in the negative mode, as with MALDITOF-MS. See UNIT 10.2 for more specific details.
30. Add 5 µL of 0.1% triethylamine to 100 µL nucleopeptide solution. Stepwise Solid-Phase Synthesis of Nucleopeptides
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31. Mix 100 µL of the above solution with an equal volume of acetonitrile (preferred), isopropanol, or MeOH. In general, the aqueous nucleopeptide solution is mixed in a 1:1 (v/v) ratio with an organic solvent such as acetonitrile, MeOH, or isopropanol, because the solvent is readily evaporated and facilitates the transfer of the ion from the liquid to the gas phase. The solvent also allows for the generation of a large number of ions. Another issue is that, as the pH increases, more negative ions are produced and the signal intensity rises. Addition of an organic base such as triethylamine to the nucleopeptide solution increases the pH and also reduces the extent of cation adducts observed in the mass spectra.
32. Analyze by ESI-MS (UNIT 10.2). Analyze base composition by SVPD/AP digestion 33. Evaporate 0.5 to 1.0 OD unit of nucleopeptide sample to dryness under vacuum in an appropriate vessel (i.e., 1.5-mL microcentrifuge tube). 34. Add the following: 34 µL freshly deionized water 50 µL 0.1 M Tris⋅Cl, pH 8.0 10 µL 0.1 M MgCl2 1 µL 1.5 U/500 µL SVPD 5 µL 0.0337 U/µL bacterial AP. 35. Vortex the sample and centrifuge briefly to collect the liquid at the bottom of the tube. Incubate 8 to 15 hr at 37°C. 36. Vortex the sample and chill with dry ice at least 10 min. 37. Centrifuge 5 min in a benchtop centrifuge at maximum speed, room temperature. Carefully remove the supernatant with a pipet and transfer to a new labeled tube. Discard the original tube containing the pellet. 38. Evaporate the sample to complete dryness under vacuum. 39. Dissolve the dried sample in 50 µL water and vortex at least 30 sec.
Table 4.22.6 Digestsa
Gradient and Mobile Phase for RP-HPLC Analysis of Nucleopeptide
Elapsed time (min)
Percent mobile phase Bb
0 5 20 25 30 31 40
10 10 30 100 100 10 10
aGradient conditions are based on a flow of 1 mL/min using a Kromasil reversed-phase C18 column (10-µm
diameter spherical silica, 4.0 × 250 mm) at ambient temperature with a 40-min injection cycle. Mobile phase A: 0.01 M ammonium acetate, pH 7.0; mobile phase B: 1:1 (v/v) acetonitrile/water. bPercentage is at elapsed time
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40. Perform reversed-phase HPLC of the digested sample on a C18 column. Load ∼0.2 OD unit per injection and elute with an acetonitrile/ammonium acetate/water gradient as specified in Table 4.22.6. Set the detector to 260 nm. See UNIT 10.6 for more specific details on RP-HPLC.
41. Collect data for 30 min. The nucleoside elution order depends on the column and HPLC system used. Approximate elution times are 5.0 min for dC, 6.0 min for dG, 6.7 min for T, and 13.1 min for dA.
42. To calculate the base composition, use the molar extinction coefficients from Table 4.22.5 and the integrated area of each peak. First calculate the average value of the ratio between the integrated area and the molar extinction coefficient (X): X = ∑ (peak area/εi ) / N Equation 4.22.4
where εi is the molar extinction coefficient of each nucleoside (Table 4.22.5) and N the number of every particular nucleobase in the sequence. Then calculate the composition for each nucleoside: n i = (peak area/εi ) / X Equation 4.22.5
where ni represents the experimental calculated value for the nucleoside i. Although SVPD is a 3′-exonuclease, it has been shown to degrade nucleopeptides where the 3′-hydroxyl function of the oligonucleotide chain is blocked by linkage to a peptide chain. This modification would prevent degradation during a short incubation time, but not after long treatments as described above. Also, SVPD contains some endonuclease contamination that favors complete degradation of the oligonucleotide chain. In general, the nucleoside composition calculated after enzymatic digestion is very similar to the expected composition. In some cases, the value of the 3′-nucleoside is low depending on the amino acid that links the peptide and oligonucleotide chain. When homoserine serine, or threonine is used, an extra peak, with higher retention time than that of simple nucleosides, may be observed in the RP-HPLC profile that corresponds to the 3′-nucleoside-peptide.
Analyze base composition by SpPD/AP digestion 43. Evaporate 0.5 to 1.0 OD unit of nucleopeptide sample to dryness under vacuum in an appropriate vessel (i.e., 1.5-mL microcentrifuge tube). 44. Add 30 µL of 0.2 M ammonium acetate buffer, pH 5.4, and 10 µL of 0.23 U/µL SpPD. 45. Vortex the sample and centrifuge briefly to collect the liquid at the bottom of the tube. Incubate 12 to 15 hr at 37°C. 46. Lyophilize and then add the following:
Stepwise Solid-Phase Synthesis of Nucleopeptides
34 µL freshly deionized water 50 µL 0.1 M Tris⋅Cl, pH 8.0 10 µL 0.1 M MgCl2 5 µL 0.0337 U/µL bacterial AP.
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47. Continue as in steps 35 to 42. SpPD is a 5′-exonuclease enzyme, and degrades the oligonucleotide chain of the nucleopeptide starting from the 5′-terminal hydroxyl group. SpPD also degrades the 3′-nucleoside-peptide linkage, independently of the linking amino acid. In general, nucleoside composition after enzymatic digestion is very similar to the expected composition. In some cases, however, the deoxyadenosine value is lower than expected. This is because of deaminase contamination, and depends on the enzyme batch employed. In these cases, an extra peak appears in the HPLC trace, which is attributed to a modified deoxyadenosine nucleoside.
Perform amino acid analysis of nucleopeptide under acidic conditions 48. Place 5 OD260 units of purified nucleopeptide in a Pyrex tube. 49. Add 300 µL water and 300 µL concentrated HCl. 50. Determine amino acid content as described (see Basic Protocol 1, steps 11 to 13). When quantifying amino acids from peptide-oligonucleotide conjugates, high values of glycine are obtained due to co-elution with oligonucleotide hydrolysis side-products.
Perform enzymatic analysis of peptides and nucleopeptides 51. Evaporate 3 to 5 nmol peptide or nucleopeptide sample to dryness under vacuum in an appropriate vessel (i.e., a 1.5-mL microcentrifuge tube). 52. Dissolve the sample in 15 µL of 0.1 M ammonium acetate buffer, pH 5.3. Add 1 µL of 1:32 (v/v) 2-mercaptoethanol/water. 53. Add 3 µL of 20 mg/mL (286 U/mL) papaine solution and incubate 2 hr at 37°C. 54. Add one drop of glacial acetic acid and lyophilize. 55. Redissolve the sample in 15 µL of 0.2 M ammonium acetate buffer, pH 8.3, and 1 µL of 1:32 (v/v) 2-mercaptoethanol/water. 56. Add 3 µL of 3 mg/mL (72 U/mL) microsomal leucin aminopeptidase solution and incubate 3 hr at 37°C. 57. Repeat the addition of 3 µL enzyme solution and incubate another 12 hr at 37°C. 58. Add one drop of concentrated acetic acid and lyophilize. 59. Redissolve the sample in 15 µL of 0.1 M ammonium acetate and 0.025 M MnCl2 buffer solution, pH 8.3. 60. Add 3 µL of 5 mg/mL (875 U/mL) prolidase solution and incubate 3 hr at 37°C. 61. Add one drop of concentrated acetic acid and lyophilize. 62. Perform amino acid analysis as described (see Basic Protocol 1, step 13). REDUCTION OF SULFOXIDE-PROTECTED METHIONINE-CONTAINING NUCLEOPEPTIDES
SUPPORT PROTOCOL 6
Additional Materials (also see Basic Protocol 3) Deprotected crude nucleopeptide (see Basic Protocol 2) N-Methylmercaptoacetamide 1. Quantitate crude nucleopeptide as described (see Basic Protocol 3, steps 8 to 10). 2. Dissolve 50 OD260 units lyophilized crude nucleopeptide in 1 mL water. 3. Place at 37°C and add a 20- to 100-fold excess (2 to 10 µL) N-methylmercaptoacetamide. Stir occasionally.
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4. After 12 hr, monitor the reaction by reversed-phase C18 HPLC (see Basic Protocol 3, steps 1 to 4). In general, the methionine-reduced nucleopeptide is eluted after the sulfoxide-containing nucleopeptide (∼1 to 2 min).
5. If necessary, add more N-methylmercaptoacetamide to achieve complete reduction within 24 to 36 hr. 6. When complete reduction is achieved, lyophilize and proceed to isolate the desired reduced nucleopeptide (see Basic Protocol 3, step 6 or 12). SUPPORT PROTOCOL 7
DEPROTECTION AND PURIFICATION OF CYSTEINE-CONTAINING NUCLEOPEPTIDES Additional Materials (also see Basic Protocol 3) Nucleopeptide-support (see Basic Protocol 2, step 7) 1 M 1,4-dithiotreitol (DTT) in peroxide-free dioxane (see recipe) 32% ammonium hydroxide 0.05 M ammonium acetate buffer, pH 7.0 20-mL screw-cap pressure tube, O-ring seal preferred Disposable polypropylene syringe with polyethylene filter (or cotton) 50-mL round-bottom flask Peristaltic pump Sephadex G-10 column Deprotect cysteine-containing nucleopeptide 1. Open the synthesis column and transfer the nucleopeptide-support to a 20-mL screw-cap pressure tube, preferably with an O-ring seal. 2. Add 5 mL of 1 M DTT in peroxide-free dioxane and 5 mL of 32% ammonium hydroxide, screw the cap on tightly, and incubate 6 to 8 hr at room temperature. It is necessary to have a large excess of DTT during the ammonia deprotection step to remove the tert-butylthio protecting group from cysteine and to avoid secondary reactions such as β-elimination of HS-S-tBu. Dimethylaminomethylene-protected guanosine and acetyl-protected cytosine phosphoramidites must be used in combination with tertbutylthio-protected cysteine residues, and cleavage and deprotection must be performed at room temperature for 6 to 8 hr. Refer to safety recommendations and other notes above (see Basic Protocol 2).
3. Filter the solution through a disposable polypropylene disposable syringe with a polyethylene filter or cotton and collect the filtrate in a 50-mL round-bottom flask. 4. Wash the pressure tube three times with MeOH and pass the washes through the syringe into the same flask. 5. Concentrate the combined solution under reduced pressure (in a rotary evaporator with a water aspirator) to a volume of ∼2 mL. Perform gel filtration 6. Set up a purification system as described for MPLC purification (see Basic Protocol 3, step 12), but with a peristaltic pump. 7. Equilibrate a Sephadex G-10 chromatography column with 200 mL of 0.05 M ammonium acetate buffer, pH 7.0. Stepwise Solid-Phase Synthesis of Nucleopeptides
8. Dilute the crude nucleopeptide sample with 0.05 M ammonium acetate buffer, pH 7.0, to a volume of 5 mL.
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9. Remove the supernatant solution from the column cap position. Introduce the nucleopeptide sample solution in the column and close. 10. Set the UV detector wavelength at 260 nm with the appropriate lamp and filter. Turn on the detector and chart recorder. Turn on the peristaltic pump and elute slowly (~1 mL/min) with 0.05 M ammonium acetate buffer, pH 7.0. Under the conditions described, the nucleopeptide-containing peak is eluted before the DTT-containing peak. This is because separation is based on molecular weight differences, with the larger compounds being eluted faster.
11. Observing the chromatogram, select the tubes where the nucleopeptide-containing peak is eluted and combine eluents. Lyophilize, analyze by HPLC, and purify as described (see Basic Protocol 3). It is important to point out that cysteine-containing nucleopeptides may dimerize very quickly because of oxidation by atmospheric oxygen, yielding a dimer in which two nucleopeptides are bonded through a disulfide bridge between the peptide chains. When two or more cysteine residues are present in the nucleopeptide, polymers can be formed. To prevent oxidation, avoid having samples in solution at room temperature. Keep frozen.
ANALYSIS AND PURIFICATION OF NUCLEOPEPTIDES BY POLYACRYLAMIDE GEL ELECTROPHORESIS
BASIC PROTOCOL 4
For more details on oligonucleotide polyacrylamide gel electrophoresis, see UNIT 10.4 and APPENDIX 3B. Materials Urea, ultrapure 38% (w/v) acrylamide/2% (w/v) bisacrylamide (see recipe) 10× TBE buffer (1.3 M Tris, 0.45 M boric acid, 25 mM EDTA) 10% (w/v) ammonium persulfate (store up to 1 week at 4°C) N,N,N′,N′-Tetramethylethylenediamine (TEMED) Bromphenol blue Xylene cyanol Glycerol Deprotected crude or purified nucleopeptide (see Basic Protocol 2 or 3) 50% (v/v) aqueous formamide Stains-all (Aldrich) Formamide Concentrated HCl Isopropanol 3 M Tris⋅Cl, pH 8.8 (APPENDIX 2A) 2 M ammonium acetate buffer, pH 7.0 Acetonitrile, HPLC grade 50% (v/v) aqueous methanol (MeOH) 250-mL Erlenmeyer flask 20- and 60-mL syringes 95° and 37°C water baths or heating blocks Sep-Pak cartridges (Waters) with clamps Polyethylene disc Disposable tubes Lyophilizer Additional reagents and equipment for polyacrylamide electrophoresis (UNIT 10.4 and APPENDIX 3B)
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Set up gel 1. Assemble gel plates, spacers, and combs as described in manufacturer’s instructions.
APPENDIX 3B
or following
2. In a 250-mL Erlenmeyer flask, combine the following: 12.6 g urea 15 mL 38% (w/v) acrylamide/2% (w/v) bisacrylamide (20% acrylamide) 1.5 mL 10× TBE buffer 30 mL water. CAUTION: Acrylamide and bisacrylamide are hazardous. Solutions of acrylamide deteriorate quickly, especially when exposed to light or left at room temperature (see Reagents and Solutions). For oligonucleotides with <25 bases, 20% is the optimum acrylamide concentration (UNIT 10.4). To speed dissolution of urea, the gel mixture can be heated before adding TEMED and ammonium persulfate; however, to prevent degradation of acrylamide, do not heat above 55°C. Allow to cool to room temperature before adding TEMED and ammonium persulfate to prevent polymerization while pouring the gel.
3. Add 300 µL of 10% (w/v) ammonium persulfate and 30 µL TEMED. Mix thoroughly and wait 30 sec. 4. Gently pull acrylamide solution into a 60-mL syringe, avoiding bubbles, and pour the gel, allowing it to flow slowly down between the gel plates (see APPENDIX 3B). 5. Insert the comb. Dislodge any trapped air bubbles, especially in the wells, by tapping gently on the glass plates. Set up electrophoresis apparatus 6. After the gel has polymerized (usually 30 to 60 min), remove bottom spacer of gel sandwich and remove extraneous polyacrylamide from around the combs. 7. Fill bottom reservoir of gel apparatus with 0.5× TBE buffer. Place the gel in the electrophoresis apparatus and clamp plates to support. 8. Pour 0.5× TBE buffer into the upper reservoir to ∼3 cm above the top of the gel. Using a Pasteur pipet, rinse wells thoroughly with 0.5× TBE buffer to remove stray fragments of polyacrylamide. 9. Preheat gel ∼1 hr by running at a constant 750 V. The ideal wattage of the gel should generate enough heat so that the gel plates are warm, but not too hot to touch.
Load and run the gel 10. To remove urea that has leached into them, flush the sample wells with 0.5× TBE buffer just prior to loading the gel. 11. Prepare a 5× stock solution with marker dyes: 0.25% (w/v) bromphenol blue 0.25% (w/v) xylene cyanol 30% (w/v) glycerol. Dilute to 1× with water before loading onto the gel. Stepwise Solid-Phase Synthesis of Nucleopeptides
In a 20% acrylamide gel, migration of bromphenol blue and xylene cyanol will be similar to 6-nt and 22-nt linear single-stranded oligonucleotides, respectively.
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12. Dissolve each sample of crude or purified nucleopeptide (0.1 to 0.2 OD260 units) in 20 µL of 50% aqueous formamide by vortexing. Microcentrifuge the dissolved samples briefly to collect them at the bottom of the tube. 13. Heat samples at 95°C in covered microcentrifuge tubes. Place on ice. 14. Load 10 µL sample per well under the surface of the buffer and just above the surface of the well. 15. Load 5 µL of 1× marker dye solution (step 11). 16. Run gel immediately at 750 V constant voltage. Maintain a constant gel temperature of ∼65°C. Observe migration of marker dyes to determine electrophoresis time. Temperatures >65°C can result in cracked plates or smeared bands; temperatures that are too low can lead to incomplete denaturation.
Process gel 17. Prepare a 10% stock dye solution by dissolving 100 mg Stains-all in 100 mL formamide. Adjust pH to 7.3 to 7.4 by adding concentrated HCl. Store at 4°C, protected from light. 18. Mix the following: 10 mL 10% Stains-all solution 10 mL formamide 50 mL isopropanol 1 mL 3 M Tris⋅Cl, pH 8.8 129 mL water. 19. Remove the gel sandwich from the apparatus and carefully place the gel in a shallow pan containing enough staining solution to cover the gel. 20. Let stand 6 hr to overnight, then decant the stain and gently rinse the gel in water. If the background is too high, soaking the gel in water and exposing to infrared light can effect some destaining.
21. Photograph the gel under ambient light against a white background. Alternatively, dry using a gel dryer (APPENDIX 3B) and scan or photograph. Purify nucleopeptide by preparative gel electrophoresis 22. Prepare a denaturing polyacrylamide gel as described above and in APPENDIX 3B. Prepare twice the amount of reagents as required for analytical gel electrophoresis. 23. Load 2 to 8 OD260 units of nucleopeptide in 40 µL of 50% aqueous formamide and perform electrophoresis. 24. Remove the gel and visualize the nucleopeptide bands using a fluorescent TLC plate and UV lamp (APPENDIX 3B). 25. Slice the gel on the perimeter of the band with a clean razor blade, and transfer the fragment to a disposable tube. Crush the fragment with a spatula. 26. Add 3 to 4 mL of 2 M ammonium acetate buffer, pH 7.0, and vortex briefly. Let stand with vigorous stirring for 12 to 24 hr in a 37°C water bath. Alternatively, freeze and thaw at least three times to enhance extraction of the product from gel. 27. Mount a Sep-Pak cartridge with a clamp and insert an empty 20-mL syringe on top of the cartridge. 28. Fill the syringe barrel with 10 mL acetonitrile. Insert and depress the plunger to pass the solution through the Sep-Pak cartridge over an elapsed time of 15 to 30 sec.
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29. Repeat step 28 with 10 mL water and 10 mL of 2 M ammonium acetate buffer, pH 7.0. 30. Remove the syringe and insert a polypropylene syringe fitted with a polyethylene disc onto the top of Sep-Pak cartridge. Add the 3 to 4 mL buffer and gel (step 26) and let fluid pass through the column by gravity. 31. Wash the disposable tube (from the gel suspension) with 3 to 4 mL water and add to the syringe. Repeat wash three more times. 32. Insert a new disposable syringe and elute nucleopeptide with 5 to 7 mL of 50% MeOH. Collect 1-mL fractions in disposable tubes. 33. Use UV absorbance to identify the fractions where the nucleopeptide has eluted. Lyophilize and quantify. REAGENTS AND SOLUTIONS Use deionized, Milli-Q-purified water (Millipore, 18 mΩ × cm−1) in all recipes and protocol steps, unless otherwise noted. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Acetonitrile, anhydrous Reflux and distill over calcium hydride powder under an inert (i.e., N2 or Ar) atmosphere. Store over calcium hydride lumps under an Ar atmosphere. Prepare fresh before each use. Alternatively, DNA-synthesis-grade low-water-content acetonitrile can be purchased and used as such, or refluxed and distilled from calcium hydride under an inert atmosphere.
Acrylamide/bisacrylamide solution, 38%/2% (w/v) Dissolve 380 g acrylamide and 20 g bisacrylamide in water to a volume of 1 liter. Filter through a 0.5-µM membrane. Store 2 to 4 weeks at 4°C, protected from light. Activator solution Prepare 0.8 M sublimed 1H-tetrazole (see recipe) in anhydrous THF (see recipe) using a septum-sealed distillation collection bulb. For a complete dissolution of tetrazole, use an ultrasonic bath. Prepare fresh before each use. Dichloromethane (DCM), anhydrous Reflux and distill dichloromethane over reagent-grade phosphorus pentoxide under an inert (N2 or Ar) atmosphere. Store over calcium hydride lumps under Ar atmosphere. Prepare fresh before each use. DCM, neutralized Pass DCM through a basic alumina column in order to eliminate acid traces. Store up to one week at 20°C. N,N-Diisopropylethylamine (DIPEA), anhydrous Place under calcium hydride lumps under an Ar atmosphere for at least one night before use. Prepare fresh before each use. Dioxane, peroxide free Pass dioxane through a basic alumina column in order to eliminate all traces of peroxide. Store until a positive peroxide test is obtained (mix equal volumes of 5% (w/v) KI aqueous solution and dioxane). Stepwise Solid-Phase Synthesis of Nucleopeptides
A yellowish color indicates the presence of peroxides. No yellowish color should be observed if the solvent is peroxide-free.
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Ninhydrin reagent A Dissolve 40 g phenol in 10 mL absolute EtOH. Mix 2 mL of 2 mM potassium cyanide and 100 mL pyridine freshly distilled over ninhydrin. Mix each solution separately with 4 g Amberlite MB-3 resin (Merck) for 45 min. Filter and mix both solutions together to obtain the desired reagent A. Store up to 1 to 2 years at 4°C. Ninhydrin reagent B Dissolve 2.5 g ninhydrin in 50 mL absolute EtOH. Keep the solution protected from light in an amber glass container (up to 1 to 2 years at 4°C). Oxidizer solution Dilute 5.0 to 6.0 M anhydrous tert-butylhydroperoxide in decane (Aldrich or Fluka) to a final concentration of 1 M in anhydrous DCM (see recipe). Prepare fresh before each use. Pyridine, anhydrous Reflux and distill pyridine over reagent-ninhydrin powder under an inert (i.e., N2 or Ar) atmosphere. Reflux and distill again over calcium hydride powder. Place over calcium hydride lumps under an Ar atmosphere. Prepare fresh before each use. Tetrahydrofuran (THF), anhydrous Dry peroxide-free tetrahydrofuran (see recipe) by continuous reflux and distillation over sodium metal and benzophenone under inert atmosphere until purple color persists. Prepare fresh before each use. THF, peroxide free Pass THF through a basic alumina column in order to eliminate peroxides. Store until a positive test is obtained (see peroxide-free dioxane). Protect from light. 1H-Tetrazole, sublimed Transfer a commercial solution of 1H-tetrazole in acetonitrile into a round-bottom flask (this can be mixed with unused wet solutions of tetrazole from the DNA synthesizer). Evaporate to dryness under reduced pressure using a rotary evaporator with a water aspirator. Use a sublimation apparatus to sublime the remaining solid to obtain pure crystalline 1H-tetrazole. Store under Ar up to 5 years at ambient temperature. CAUTION: 1H-Tetrazole is an explosive compound. Never heat near the melting point (157°-159°C). Wear safety glasses and gloves during sublimation procedure. Manipulate with caution in a well-ventilated fume hood. 1H-Tetrazole is no longer commercially available in solid form. Solutions of 1H-tetrazole in acetonitrile may be purchased from typical organic chemical suppliers such as Aldrich.
COMMENTARY Background Information The methodologies for the preparation of various kinds of peptide-oligonucleotide conjugates have been reviewed by different authors (Tung and Stein, 2000; Zubin et al., 2002; UNIT 4.5). The research published since the last review report on adaptations, modifications, problems, or improvements of previously described methods (e.g., Chen et al., 2002, 2003; Drioli et al., 2002; Kachalova et al., 2002; Viladkar, 2002; Zatsepin et al., 2002), but no new synthesis design has been described. For these reasons,
as well as for space limitations, only a few comments on the preparation of nucleopeptides and on the use of stepwise solid-phase methodology to obtain peptide-oligonucleotide conjugates will be made here (i.e., only representative examples will be cited). All the methodologies used to prepare hybrids in which the two components are directly linked through phosphate diesters (here referred to as nucleopeptides) share a common outline: the key linking phosphodiester bond is
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Stepwise Solid-Phase Synthesis of Nucleopeptides
created having all the nonparticipating functional groups protected. This key phosphate has been formed using different chemistries that yield phosphodiester bonds, namely the phosphate triester (Kuyl-Yeheskiely et al., 1989; Ueno et al., 1993), the H-phosphonate (Kuyl-Yeheskiely et al., 1987), and the phosphite triester (Dreef-Tromp et al., 1992a; Sakakura and Hayakawa, 2000; Debéthune et al., 2002a; Jeyaraj et al., 2002) approaches. Synthesis of nucleopeptides has been carried out in solution and on solid supports, both by convergent (Dreef-Tromp et al., 1992a; Robles et al., 1995; Sakakura and Hayakawa, 2000) and stepwise strategies (Debéthune et al., 2002a). Protection schemes have included a large variety of permanent protecting groups that are either chemically or enzymatically (Flohr et al., 1999) removable. Many nucleopeptides have been prepared since the pioneering work of Z.A. Shabarova in the sixties (Shabarova, 1970). In 1992, van Boom’s group was able to obtain a nucleopeptide with up to ten nucleosides (Dreef-Tromp et al., 1992a) and two trifunctional amino acids besides the linking residue. The methodology described here has allowed the synthesis of nucleopeptides containing up to five amino acids with functionalized side chains (besides the linking residue), and to obtain a nucleopeptide with thirteen amino acids and fifteen nucleosides. In the authors’ opinion, one of the main advantages of this stepwise solid-phase methodology is that it circumvents the need to manipulate protected peptides. These are often highly insoluble in most organic solvents, which renders both the purification and the phosphitylation (or phosphorylation) difficult. Moreover, most of the monomers required can be purchased from commercial suppliers. Besides the trifunctional amino acid derivatives whose synthesis is described here, the only nucleoside synthons that should have to be synthesized in the laboratory are the 5′-phosphoramidites of 2′-deoxycytidine and 2′-deoxyguanosine. The commercially available 5′phosphoramidites of these nucleosides are only suitable for the preparation of nucleopeptides with 5′-oligonucleotide-homoserine or tyrosine linkages, and their standard nucleobaseprotecting groups are not compatible with baselabile nucleopeptides. Finally, it is worth emphasizing that compared to other methods the use of an amino acid to attach the two moieties has the advantage of allowing one to place the linking unit at any
position in the peptide chain, with no need to prepare a specially designed support. The only exception to this assertion, as previously mentioned, is that homoserine cannot be placed at the C-terminal position. The acid treatment that removes the Boc group simultaneously eliminates its DMTr hydroxyl-protecting group, and the propensity for homoserine to lactonize would easily provoke cleavage of the peptideresin bond, resulting in loss of peptide chains. As previously stated, peptide-oligonucleotide conjugates that are not directly linked or are linked through amide bonds have also been synthesized using stepwise procedures. In most cases, hybrids have been assembled onto insoluble matrices, but the modifiable solubility of polyethylene glycol has also been exploited (Drioli et al., 2002). Peptide assembly is more often carried out first, but synthesis starting with oligonucleotide elongation has also been reported (Bergmann and Bannwarth, 1995; Sarracino et al., 1998). In general, there is no need for concern about the stability of the linkage between the two moieties, but some degradation of the target molecule during the final deprotection treatment has been found to be associated with the use of immobilized trifunctional linkers (Basu and Wicsktrom, 1995). It is important to note that the protection scheme proposed here for the preparation of nucleopeptides is of general application and can be extended to the synthesis of any peptideoligonucleotide conjugate. In addition, when the target molecule contains trifunctional amino acids that may suffer side reactions, and/or base-labile linkers are used for their assembly, the recommendation to use the most labile set of permanent protecting groups also applies. The most common applications of peptideoligonucleotide conjugates have been discussed earlier (see unit introduction). Such molecules can also find application in structural studies (Ho et al., 1999; Gómez-Pinto et al., 2003), in studies on the structural requirements of an enzyme (Debéthune et al., 2002b), or in studies on the behavior of metal complexes (Civitello et al., 2001; Marchán et al., 2001). Hopefully, access to user-friendly synthesis procedures and the future development of newer, more convenient procedures will inspire an increasing number of laboratories to prepare more conjugates, and thus enlarge their fields of application.
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A
0
B
35
0
C
35
0
D
35
0
35
Min.
Figure 4.22.9 Examples of HPLC profiles of nucleopeptides: (A) crude and (B) purified Ac-Lys-Trp-Lys-Hse(p3′dGCATCG)-Ala-OH, and (C) crude and (D) purified Ac-Tyr(p3′dTTTCAGAAAATCTAG)-Leu-Asp-Pro-Arg-Ile-Thr-Val-OH. See Basic Protocol 3 for gradient conditions, eluents, and column employed.
Critical Parameters As previously pointed out, the most critical point in nucleopeptide synthesis is the choice of protecting groups, which must be accompanied by appropriate selection of the final deprotection conditions (see Strategic Planning and Table 4.22.1). It is also important to ensure that oligonucleotide elongation is carried out on a homogeneous immobilized peptide. The amino acid composition should be checked before the oligonucleotide synthesis is begun. Depending on the peptide length and composition, HPLC and MS analysis of the crude product obtained after deprotection of an aliquot of peptide-resin may also be worthwhile. If large nucleopeptides or conjugates are to be prepared, leaving the DMTr group after incorporation of the last nucleoside synthon may facilitate the HPLC analysis and purification of the crude product. Two purification runs, both by HPLC or by combining HPLC and PAGE, may sometimes be required.
Troubleshooting Large nucleopeptides, especially depending on the amino acid composition, may show complex HPLC profiles. HPLC analysis at higher temperature (50° to 55°C) may reduce conformational equilibria, and in some cases simplify the HPLC profile of the crude product. As previously suggested, analysis by PAGE under denaturing conditions may also be a good alternative.
major peak in the crude nucleopeptide after cleavage and deprotection. The coupling yield of the first nucleoside phosphoramidite is variable, with an average value of 70%. Nearly quantitative yields, as in any other oligonucleotide synthesis, are obtained in the subsequent couplings. The cleavage yield with concentrated ammonia/dioxane treatment at 55°C usually ranges from 60% to 80%. An additional treatment with TBAF, if allowed by the nucleopeptide structure, leaves <5% of the nucleopeptide on the resin. The purity of the crude nucleopeptide varies depending on the nucleopeptide size and composition. In the HPLC profiles shown in Figure 4.22.9, the main peak (expected product) accounts for ∼75% of the total area in the case of the tryptophan-containing nucleopeptide, and ∼60% in the arginine-containing one. MALDITOF-MS analysis of the crude sample allows confirmation that it contains the target product. Purification can usually be accomplished in a single run by either reversed-phase MPLC or HPLC on semipreparative C18 columns. Overall yields (nucleopeptide assembly, deprotection, and purification) will also vary depending on the homogeneity of the crude product and whether purification requires one or two steps. The arginine-containing nucleopeptide, for instance, is an example of a nucleopeptide of medium difficulty, and was obtained in 17% yield.
Time Considerations Anticipated Results If peptide elongation has proceeded satisfactorily, the expected peptide should be the
The preparation of nucleopeptides using these procedures may be a time-demanding task if trifunctional amino acid derivatives, as well as
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the handle linking the hybrid molecule to the resin, have to be prepared. With all of the precursors, reagents, and solutions in hand, the synthesis and purification of a nucleopeptide containing ten amino acids and a 10-mer oligonucleotide can be accomplished in ∼3 weeks, depending, for instance, on the availability of techniques such as amino acid analysis to assess that correct yields have been obtained, or on whether peptide purity is checked before proceeding with oligonucleotide elongation. The time required to purify the target molecule may vary depending on the scale of synthesis and, obviously, the quality of the crude product. A tentative schedule would be the following: Week 1: Preparation of solid support, manual synthesis of peptide chain, and analysis of peptide-resin. Week 2: Oligonucleotide elongation, deprotection and cleavage, and analysis of the crude nucleopeptide to set up the purification conditions. Week 3: Purification of the nucleopeptide and full characterization of the product (amino acid analysis after acid hydrolysis or enzymatic digestion, determination of the relative proportion of nucleosides after enzymatic digestion, and mass spectrometric analysis).
Literature Cited Albericio, F., Cruz, M., Debéthune, L., Eritja, R., Giralt, E., Grandas, A., Marchán, V., Pastor, J.J., Pedroso, E., Rabanal, F., and Royo, M. 2001. An improved synthesis of N-[(9-hydroxymethyl)-2fluorenyl]succinamic acid (HMFS), a versatile handle for the solid-phase synthesis of biomolecules. Synthetic Commun. 31:225-232. Bardella, F., Giralt, E., and Pedroso, E. 1990. Polystyrene-supported synthesis by the phosphitetriester approach: An alternative for the large scale synthesis of small oligodeoxyribonucleotides. Tetrahedron Lett. 31:6231-6234.
Stepwise Solid-Phase Synthesis of Nucleopeptides
Chen, C.-P., Li, X.-X., Zhang, L.-R., Min, J.-M., Chan, J.Y.-W., Fung, K.-P., Wang, S.-Q., and Zhang, L.-H. 2002. Synthesis of antisense oligonucleotide-peptide conjugate targeting to GLUT-1 in HepG-2 and MCF-7 cells. Bioconjugate Chem. 13:525-529. Chen, C.-P., Zhang, L.,-R., Peng, Y,-F., Wang, X.,B., Wang, S.,-Q., and Zhang, L.H. 2003. A concise method for the preparation of peptide and arginine-rich peptide-conjugated antisense oligonucleotide. Bioconjugate Chem. 14:532-538. Christensen, T. 1979. A qualitative test for monitoring coupling completeness in solid phase peptide synthesis using chloranil. Acta Chem. Scand. B33:763-766. Civitello, E.R., Leniek, R.J., Hossler, K.A., Haebe, K., and Stearns, D.M. 2001. Synthesis of peptide-oligonucleotide conjugates for chromium coordination. Bioconjugate Chem. 12:459-463. Debéthune, L., Marchán, V., Fábregas, G., Pedroso, E., and Grandas, A. 2002a. Towards nucleopeptides containing any trifunctional amino acid (II). Tetrahedron 58:6965-6978. Debéthune, L., Kohlhagen, G., Grandas, A., and Pommier, Y. 2002b. Processing of nucleopeptides mimicking the topoisomerase I-DNA covalent complex by tyrosyl-DNA phosphodiesterase. Nucl. Acids Res. 30:1198-1204. Dreef-Tromp, C.M., van der Maarel, J.C.M., van den Elst, H., van der Marel, G.A., and van Boom, J.H. 1992a. Solid-phase synthesis of the nucleopeptide fragment H-Asp-Ser[pAAAGTAAGCC]-Glu-OH from the nucleoprotein of Bacillus subtilis phage φ29. Nucl. Acids Res. 20:4015-4020. Drioli, S., Adamo, I., Ballico, M., Morvan, F., and Bonora, G.M. 2002. Liquid-phase synthesis and characterization of a conjugated chimeric oligonucleotide-PEG-peptide. Eur. J. Org. Chem. 3473-3480. Eritja, R., Robles, J., Fernández-Forner, D., Albericio, F., Giralt, E., and Pedroso, E. 1991. NPE-resin, a new approach to the solid-phase synthesis of protected peptides and oligonucleotides I: Synthesis of the supports and their application to oligonucleotide synthesis. Tetrahedron Lett. 32:1511-1514.
Basu, S., and Wickstrom, E. 1995. Solid phase synthesis of a D-peptide-phosphorothioate oligodeoxynucleotide conjugate from two arms of a polyethylene glycol-polystyrene support. Tetrahedron Lett. 36:4943-4946.
Flohr, S., Jungmann, V., and Waldmann, H. 1999. Chemoenzymatic synthesis of nucleopeptides. Chem. Eur. J. 5:669-681.
Beltrán, M., Maseda, M., Robles, J., Pedroso, E., and Grandas, A. 1997. Homoserine derivatives for the preparation of base-stable nucleopeptide analogues. Lett. Pept. Sci. 4:147-155.
Fujii, T. and Sakakibara, S. 1974. Studies on the synthesis of histidine peptides. I. NIm-Tosylhistidine derivatives as starting materials. Bull. Chem. Soc. Jpn. 47:3146-3151.
Beltrán, M., Pedroso, E., and Grandas, A. 1998. A comparison of histidine protecting groups in the synthesis of peptide-oligonucleotide conjugates. Tetrahedron Lett. 39:4115-4118.
Gairí, M., Lloyd-Williams, P., Albericio, A., and Giralt, E. 1990. Use of BOP reagent for the suppression of diketopiperazine formation in boc/bzl solid-phase peptide synthesis. Tetrahedron Lett. 31:7363-7366.
Bergmann, F. and Bannwarth, W. 1995. Solid phase synthesis of directly linked peptide-oligodeoxynucleotide hybrids using standard synthesis protocols. Tetrahedron Lett. 36:1839-1842.
Gait, M.J. 2003. Peptide-mediated cellular delivery of antisense oligonucleotides and their analogues. Cell. Mol. Life Sci. 60:1-10.
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García de la Torre, B., Albericio, F., SaisonBehmoaras, E., Bachi, A., and Eritja, R. 1999. Synthesis and binding properties of oligonucleotides carrying nuclear localization sequences. Bioconjugate Chem. 10:1005-1012.
Ollivier, N., Olivier, C., Gouyette, C., Huynh-Dinh, T., Gras-Masse, H., and Melnyk, O. 2002. Synthesis of oligonucleotide-peptide conjugates using hydrazone chemical ligation. Tetrahedron Lett. 43:997-999.
Gisin, B.F. and Merrifield, R.B. 1972. Carboxylcatalyzed intramolecular aminolysis side reaction in solid-phase peptide synthesis. J. Am. Chem. Soc. 94:3102-3106.
Robles, J., Pedroso, E., and Grandas, A. 1991. Solid phase synthesis of a model nucleopeptide with a phosphodiester bond between the 5′ end of a trinucleotide and a serine residue. Tetrahedron Lett. 32:4389-4392.
Gómez-Pinto, I., Marchán, V., Gago, F., Grandas, A., and González, C. 2003. Solution structure and stability of tryptophan-containing nucleopeptide duplexes. ChemBiochem. 4:40-49. Ho, W.C., Steinbeck, C., and Richert, C. 1999. Solution structure of the aminoacyl-capped oligodeoxyribonucleotide du plex (WTGCGCAC)2. Biochemistry 38:12597-12606. Jeyaraj, D.A., Prinz, H.R., and Waldmann, H.R. 2002. Synthesis of nucleopeptides by employing an enzyme-labile urethane protecting group. Chem. Eur. J. 8:1879-1887. Kachalova, A.V., Stetsenko, D.A., Romanova, E.A., Tashlitsky, V.N., Gait, M.J., and Oretskaya, T.S. 2002. A new and efficient method for the synthesis of 5′-conjugates of oligonucleotides through amide-bond formation in solid phase. Helv. Chim. Acta 85:2409-2416. Kaiser, E., Colescott, R.L., Bossinger, C.D., and Cook, P.I. 1970. Color test for detection of free amino groups in the solid-phase synthesis of peptides. Anal. Biochem. 34:595-598. Kuyl-Yeheskiely, E., Tromp, C.M., Schaeffer, A.HR., van der Marel, G.A., and van Boom, J.H. 1987. A model study directed towards the preparation of nucleopeptides via H-phosphonate intermediates. Nucl. Acids Res. 15:1807-1818. Kuyl-Yeheskiely, E., Dreef-Tromp, C.M., Geluk, A., van der Marel, G.A., and van Boom, J.H. 1989. Synthesis of the nucleopeptide H-PheTyr(pGC)-NH2 and H-Phe-Ser(pGC)-Ala-OH via a phosphotriester approach. Nucl. Acids Res. 17:2897-2905. Marchán, V., Rodríguez-Tanty, C., Estrada, M., Pedroso, E., and Grandas, A. 2000. Alternative procedures for the synthesis of methionine-containing peptide-oligonucleotide hybrids. Eur. J. Org. Chem. 2495-2500. Marchán, V., Moreno, V., Pedroso, E., and Grandas, A. 2001. Towards a better understanding of the cisplatin mode of action. Chem. Eur. J. 7:808815. McMinn, D.L. and Greenberg, M.M. 1999. Convergent solution-phase synthesis of a nucleopeptide using a protected oligonucleotide. Bioorg. Med. Chem. Lett. 9:547-550. Montserrat, F.X., Grandas, A., Eritja, E., and Pedroso, E. 1994. Criteria for the economic large scale solid-phase synthesis of oligonucleotides. Tetrahedron 50:2617-2622.
Robles, J., Pedroso, E., and Grandas, A. 1995. Solidphase synthesis of a nucleopeptide from the linking site of adenovirus-2 nucleoprotein, -Ser(p5′CATCAT)-Gly-Asp-. Convergent versus stepwise strategy. Nucl. Acids Res. 23:4151-4161. Robles, J., Maseda, M., Beltrán, M., Cocernau, M., Pedroso, E., and Grandas, A. 1997. Synthesis and enzymatic stability of phosphodiester-linked peptide-oligonucleotide hybrids. Bioconjugate Chem. 8:785-788. Robles, J., Beltrán, M., Marchán, V., Pérez, Y., Travesset, I., Pedroso, E., and Grandas, A. 1999. Towards nucleopeptides containing any trifunctional amino acid. Tetrahedron 55:13251-13264. Sakakura, A. and Hayakawa, Y. 2000. A novel synthesis of oligonucleotide-peptide conjugates with a base-labile phosphate linker between the two components according to the allyl-protected phosphoramidite strategy. Tetrahedron 56:44274435. Sarracino, D.A., Steinberg, J.A., Vergo, M.T., Woodworth, G.F., Tetzlaff, C.N., and Richert, C. 1998. 5′-Peptidyl substituents allow a tuning of the affinity of oligodeoxyribonucleotides for RNA. Bioorg. Med. Chem. Lett. 8:2511-2516. Schattenkerk, C., Wreesmann, C.T.J., de Graaf, M.J., van der Marel, G.A., and van Boom, J.H. 1984. Synthesis of naturally occurring nucleopeptide fragment via a phosphotriester approach. Tetrahedron Lett. 25:5197-5200. Shabarova, Z.A. 1970. Synthetic nucleotide-peptides. In Progress in Nucleic Acid Research and Molecular Biology, Vol. 10 (J.N. Davidson and W.E. Colum, eds.) pp. 145-182. Academic Press, London. Stetsenko, D.A., Malakhov, A.D., and Gait, M.J. 2002. Total stepwise solid-phase synthesis of oligonucleotide-(3′→N)-peptide conjugates. Org. Lett. 2002:3259-3262. Tung, C.-H. and Stein, S. 2000. Preparation and applications of peptide-oligonucleotide conjugates. Bioconjugate Chem. 11:605-618. Ueno, Y., Saito, R., and Hata, T. 1993. Studies on the synthesis of nucleotidyl-peptides. II. The preparation of a nucleotidyl-peptide having a 5′-nucleotidyl-(P-O)-serine phosphodiester bond. Nucl. Acids Res. 21:4451-4457. Viladkar, S.M. 2002. Guanine rich oligonucleotideamino acid/peptide conjugates: Preparation and characterization. Tetrahedron 58:495-502. Waldmann, H. and Gabold, S. 1997. Chemoenzymatic synthesis of nucleopeptides. Chem. Commun. 1861-1862.
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Zatsepin, T.S., Stetsenko, D.A., Arzumanov, A.A., Romanova, E.A., Gait, M.J., and Oretskaya, T.S. 2002. Synthesis of peptide-oligonucleotide conjugates with single and multiple peptides attached to 2′-aldehydes through thiazolidine, oxime, and hydrazine linkages. Bioconjugate Chem. 13:822-830. Zubin, E.M., Romanova, E.A., and Oretskaya, T.S. 2002. Modern methods for the synthesis of peptide-oligonucleotide conjugates. Russ. Chem. Rev. 71:239-264.
Key References Dreef-Tromp, C.M., van den Elst, HR., van den Boogaart, J.E., van der Marel, G.A., and van Boom, J.H. 1992b. Solid-phase synthesis of an RNA nucleopeptide fragment from the nucleoprotein of poliovirus. Nucl. Acids Res. 20:24352439.
Largest phosphodiester-linked peptide-oligoribonucleotide hybrid synthesized to date. Tung and Stein, 2000. See above. Review covering synthesis and applications of peptide-oligonucleotide conjugates.
Contributed by Anna Grandas, Vicente Marchán, Laurent Debéthune, and Enrique Pedroso Universitat de Barcelona Barcelona, Spain
The authors wish to acknowledge the contribution of Dr. Jordi Robles, who set up the basic nucleopeptide stepwise solid-phase synthesis methodology, in addition to the contributions of many other people over the years.
Stepwise Solid-Phase Synthesis of Nucleopeptides
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Synthesis of Oligoribonucleotides Containing N6-Alkyladenosine and 2-Methylthio-N6-Alkyladenosine
UNIT 4.23
This unit describes synthesis of 5 -O-(4,4 -dimethoxytrityl)-2 -O-tert-butyldimethylsilyl3 -O-[(2-cyanoethoxy)-(N,N-diisopropylamino)]phosphinyl-6-methylthiopurine riboside (S.9, Fig. 4.23.1; see Basic Protocol 1) and 5 -O-(4,4 -dimethoxytrityl)-2 -Otert-butyldimethylsilyl-3 -O-[(2-cyanoethoxy)-(N,N-diisopropylamino)]phosphinyl-2methylthio-6-chloropurine riboside (S.18, Fig. 4.23.2; see Alternate Protocol). Both modified phosphoramidites and natural ribonucleoside phosphoramidites are used for the synthesis of precursor forms of oligoribonucleotides, which are then converted into oligoribonucleotides containing N6 -alkyladenosine and 2-methylthio-N6 -alkyladenosine by treatment with an appropriate primary amine (Kierzek and Kierzek, 2003a,b; see Basic Protocol 2).
SYNTHESIS OF THE 6-METHYLTHIOPURINE RIBOSIDE PHOSPHORAMIDITE
BASIC PROTOCOL 1
The synthesis of 5 -O-DMTr-2 -O-TBDMS-3 -O-(2-cyanoethyl-N,N-diisopropyl)-6methylthiopurine riboside phosphoramidite starts with complete conversion of inosine (S.1; Fig. 4.23.1) into 2 ,3 ,5 -tri-O-acetylinosine (S.2) by reaction with acetic anhydride in pyridine. S.2 is quantitatively transformed into 2 ,3 ,5 -tri-O-acetyl-6-thiopurine riboside (S.3) by treatment with Lawesson’s reagent for 1.5 hr at 100◦ C (Cava and Levinson, 1985; Kierzek and Kierzek, 2003a). This product is transformed into 2 ,3 ,5 -tri-O-acetyl6-methylthiopurine riboside (S.4) with methyl iodide in the presence of potassium carbonate in N,N-dimethylformamide (DMF) over 2 hr at 50◦ C in ∼81% yield (Wetzel and Eckstein, 1975). Acetyl groups are removed with ammonium hydroxide in pyridine, and the resulting 6-methylthiopurine riboside (S.5) is treated with 4,4 -dimethoxytrityl chloride (DMTr-Cl). After purification by short-column chromatography, the 5 -O-DMTr-6methylthiopurine riboside (S.6) is obtained in ∼74% yield (Smith et al., 1962). This product is reacted with tert-butyldimethylsilyl chloride (TBDMS-Cl) in pyridine in the presence of imidazole to give a mixture of 2 - (S.7), 3 - (S.8), and 2 ,3 -silylated derivatives, which after purification and subsequent 3 -O-TBDMS derivative isomerization gives 5 O-DMTr-2 -O-TBDMS-6-methylthiopurine riboside (S.7) in ∼66% yield (Ogilvie at al., 1978). The phosphoramidite S.9 is prepared by reaction of S.7 with 2-cyanoethyl diisopropylchlorophosphoramidite in anhydrous acetonitrile in the presence of diisopropylethylamine (Beaucage and Caruthers, 1981). The product is obtained in ∼85% yield after purification by short-column chromatography on silica gel.
Materials Inosine (S.1) Pyridine (reagent grade or better) Acetic anhydride Dichloromethane (anhydrous) Methanol (anhydrous) Saturated aqueous sodium bicarbonate solution Sodium sulfate (anhydrous) Toluene
Contributed by Elzbieta Kierzek and Ryszard Kierzek Current Protocols in Nucleic Acid Chemistry (2004) 4.23.1-4.23.21 C 2004 by John Wiley & Sons, Inc. Copyright
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Figure 4.23.1 Acetylation, thiolation, methylation, acetyl deprotection, dimethoxytritylation, silylation, and phosphinylation of purine riboside derivative. Abbreviations: Ac, acetyl; DMTr, 4,4 -dimethoxytrityl; i-Pr, isopropyl; TBDMS, tertbutyldimethylsilyl.
1,4-Dioxane (anhydrous) Lawesson’s reagent (Aldrich) N,N-Dimethylformamide (DMF; anhydrous) Potassium carbonate (anhydrous), ground Methyl iodide Celite 545 Silica gel 60H (Merck) Concentrated aqueous ammonium hydroxide (NH4 OH; 28% to 30%) 4,4 -Dimethoxytrityl chloride (DMTr-Cl) Imidazole tert-Butyldimethylsilyl chloride (TBDMS-Cl) Ethyl acetate Hexanes (anhydrous) 10% (w/v) aqueous sodium phosphate, monobasic (NaH2 PO4 ) Benzene (anhydrous) Dry ice/ethanol bath Acetonitrile (commercial with <20 ppm water or dried over 3Å molecular sieves) Diisopropylethylamine (DIPEA; anhydrous) 2-Cyanoethyl diisopropylchlorophosphoramidite Acetone Triethylamine (TEA; anhydrous) Oligoribonucleotides with N6 -Alkyladenosine
250- and 25-mL round-bottom flasks Silica gel 60F thin-layer chromatography (TLC) plates (Merck)
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250-mL and 1-L separatory funnels Rotary evaporator Vacuum pump and water aspirator Reflux condenser 100◦ and 50◦ C oil baths (silicone oil) 60- and 150-mL sintered glass funnels Rubber septa Vacuum adaptor for 250-mL round-bottom flask and appropriate traps Vacuum desiccator 1-mL and 10-mL glass syringes with needles Silanized silica gel 60F TLC plates (Merck), optional Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (UNIT 2.4 & APPENDIX 3E) Acetylate inosine 1. Suspend 8.01 g (30 mmol) inosine (S.1) in 75 mL anhydrous pyridine in a 250-mL round-bottom flask and begin stirring. Add 9.90 mL (105 mmol) acetic anhydride and leave 16 hr at room temperature. 2. Analyze the reaction mixture by TLC (APPENDIX 3D) on silica gel 60F TLC plates using 9:1 (v/v) dichloromethane/methanol (S.2 Rf = 0.54). The TLC analysis is performed with a short-wave length UV lamp at 254 nm.
3. Pour the reaction mixture carefully into a 1-L separatory funnel containing 400 mL saturated aqueous sodium bicarbonate solution and extract three times with 200 mL dichloromethane. 4. Dry combined organic layers with ∼10 g anhydrous sodium sulfate and evaporate on a rotary evaporator with a water aspirator. Coevaporate residue three times with ∼50 mL toluene. Dry under vacuum pump. 2 ,3 ,5 -Tri-O-acetylinosine (S.2): 11.89 g (100%). TLC (dichloromethane/methanol 9:1 [v/v]): Rf = 0.54. UV (MeOH): λmax = 224, 256 nm; λmin = 228 nm. 1 H NMR (CDCl3 ): δ 2.10 (s, 3H, CH3 ), 2.15 (s, 3H, CH3 ), 2.16 (s, 3H, CH3 ), 4.41–4.47 (m, 3H, H4 , H5 , H5 ), 5.61 (t, J = 5.8 Hz, 1H, H3 ), 5.88 (t, J = 6.5 Hz, 1H, H2 ), 6.19 (d, J = 5.1 Hz, 1H, H1 ), 8.11 (s, 1H, H2), 8.31 (s, 1H, H8). 13 C NMR (CDCl3 ): δ 20.3, 20.5, 20.7 (C(O)CH3 ), 63.0 (C5 ), 70.5 (C3 ), 73.3 (C2 ), 80.4 (C4 ), 86.6 (C1 ), 125.1 (C5), 138.6 (C8), 145.8 (C4), 148.7 (C2), 158.7 (C6), 169.3, 169.5, 170.3 (C(O)CH3 ).
Thionylate inosine derivative 5. Dissolve 11.83 g (30 mmol) S.2 in 100 mL 1,4-dioxane in a 250-mL round-bottom flask and add 7.28 g (18 mmol) Lawesson’s reagent. Attach a reflux condenser and reflux the reaction mixture in an oil bath for 1.5 hr at 100◦ C. CAUTION: Reaction must be run in an efficiently working hood.
6. Periodically analyze the reaction mixture by TLC on silica gel 60F plates using 9:1 (v/v) dichloromethane/methanol (S.3 Rf = 0.64). 7. On the rotary evaporator with water aspirator, remove approximately half the volume of dioxane. 8. Add 10 mL methanol followed by 200 mL saturated aqueous sodium bicarbonate solution. Cooling the reaction mixture and evaporation of dioxane result in gel formation, but addition of methanol dissolves the reaction mixture, thus allowing easy workup.
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9. Extract the reaction mixture three times with 150 mL dichloromethane. 10. Dry combined organic layers with ∼20 g anhydrous sodium sulfate and rotary evaporate to a solid foam. 2 ,3 ,5 -Tri-O-acetyl-6-thiopurine riboside (S.3): TLC (dichloromethane/methanol 9:1 [v/v]): Rf = 0.64. UV (MeOH): λmax = 233, 325 nm; λmin = 268 nm.
Methylate 6-thioinosine derivative 11. Coevaporate the reaction mixture with 50 mL DMF in a 250-mL round-bottom flask and dissolve the residue in 50 mL DMF. 12. Add 24.4 g (33 mmol) ground anhydrous potassium carbonate and 2.07 mL (33 mmol) methyl iodide. Heat the reaction mixture in the oil bath for 2 hr at 50◦ C with intensive stirring. Anhydrous potassium carbonate must be ground to a powder before addition to the reaction mixture.
13. Periodically analyze the reaction mixture by TLC on silica gel 60F plates using 9:1 (v/v) dichloromethane/methanol (S.4 Rf = 0.86). 14. Cool the reaction mixture to room temperature and filter off potassium carbonate by vacuum filtration through a Celite 545 cake on a 60-mL sintered glass funnel. Transfer the filtrate to a 1-L separatory funnel and add 150 mL saturated aqueous sodium bicarbonate solution. 15. Extract the reaction mixture three times with 150 mL dichloromethane. Dry combined organic layers with ∼20 g anhydrous sodium sulfate and rotary evaporate with water aspirator. 16. Purify the reaction mixture by short-column chromatography (UNIT 2.4 & APPENDIX 3E). Prepare the column (150-mL sintered glass funnel, ∼100 g silica gel 60H) in dichloromethane, and form the gradient by increasing the amount of methanol by 0.5% (v/v) for every 100 mL dichloromethane, up to 2.5% methanol. Analyze composition of the fractions (∼30 mL) on silica gel 60F TLC plates in 9:1 (v/v) dichloromethane/methanol. 17. Combine the fractions containing product (S.4) and rotary evaporate with water aspirator. 2 ,3 ,5 -Tri-O-acetyl-6-methylthiopurine riboside (S.4): 10.27 g (81% relative to S.2). TLC (dichloromethane/methanol 9:1 [v/v]): Rf = 0.86. UV (MeOH): λmax = 221, 284 nm; λmin = 241 nm. 1 H NMR (CDCl3 ): δ 2.09 (s, 3H, CH3 ), 2.13 (s, 3H, CH3 ), 2.16 (s, 3H, CH3 ), 2.75 (s, 3H, SCH3 ), 4.35–4.49 (m, 3H, H4 , H5 , H5 ), 5.67 (t, J = 5.6 Hz, 1H, H3 ), 5.96 (t, J = 5.4 Hz, 1H, H2 ), 6.22 (d, J = 5.4 Hz, 1H, H1 ), 8.15 (s, 1H, H2), 8.76 (s, 1H, H8).
Remove acetyl groups and perform dimethoxytritylation 18. To 10.27 g (24.19 mmol) S.4 in a 250-mL round-bottom flask, add 40 mL pyridine and 20 mL concentrated aqueous NH4 OH. Close the flask with a rubber septum and leave for 16 hr at room temperature.
Oligoribonucleotides with N6 -Alkyladenosine
19. Analyze reaction mixture by TLC on silica gel 60F plates using 9:1 (v/v) dichloromethane/methanol (S.5 Rf = 0.33). 20. Rotary evaporate the reaction mixture and coevaporate residue three times with anhydrous pyridine (∼50 mL each time).
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21. Dissolve the oily residue in 125 mL anhydrous pyridine and add 8.99 g (26.61 mmol) DMTr-Cl. Stir the reaction mixture for 1 to 1.5 hr at room temperature. 22. Periodically analyze reaction mixture by TLC on silica gel 60F plates using 9:1 (v/v) dichloromethane/methanol (S.6 Rf = 0.40). If reaction is not complete after 1.5 hr, add 5% to 10% (w/w) DMTr-Cl, based on the initial amount added, and stir 1 hr. 23. Carefully add the reaction mixture to 250 mL saturated aqueous sodium bicarbonate solution in a 1-L separatory funnel. Extract three times with 200 mL dichloromethane. 24. Dry combined organic layers with ∼20 g anhydrous sodium sulfate and rotary evaporate with water aspirator. Coevaporate residue three times with 50 mL toluene. 25. Purify the reaction mixture by short-column chromatography. Prepare the column (150-mL sintered glass funnel, ∼100 g silica gel 60H) in dichloromethane and make the gradient by increasing the amount of methanol by 0.5% (v/v) for every 100 mL dichloromethane, up to 2% methanol. 26. Analyze composition of fractions (∼30 mL) by TLC on silica gel 60F plates using 9:1 (v/v) dichloromethane/methanol. Combine fractions containing product (S.6) and rotary evaporate with water aspirator to a solid foam. 5 -O-(4,4 -Dimethoxytrityl)-6-methylthiopurine riboside (S.6): 10.73 g (74%). TLC (dichloromethane/methanol 9:1 [v/v]): Rf = 0.40. FAB-MS: m/z = 601.5. UV (MeOH): λmax = 215, 226, 280 nm; λmin = 220, 256 nm. 1 H NMR (CDCl3 ): δ 2.64 (s, 3H, SCH3 ), 3.31–3.36 (m, 1H, H5 ), 3.43–3.48 (m, 1H, H5 ), 3.72 (s, 6H, OCH3 ), 4.37 (d, J = 3.8 Hz, 1H, H4 ), 4.46–4.49 (m, 1H, H3 ), 4.85 (t, J = 7.0 Hz, 1H, H2 ), 6.06 (d, J = 6.0 Hz, 1H, H1 ), 6.73 (d, J = 9.0 Hz, 4H, DMTr), 7.14–7.28 (m, 9H, DMTr), 8.20 (s, 1H, H2), 8.54 (s, 1H, H8). 13 C NMR (CDCl3 ): δ 11.8 (SCH3 ), 55.2 (OCH3 ), 63.5 (C5 ), 72.9 (C3 ), 76.1 (C2 ), 86.3 (C4 ), 91.1 (C1 ), 113.2 (DMTr), 126.9 (C5), 127.8, 128.0, 129.1, 129.9, 135.4, 135.5 (DMTr), 140.9 (C8), 144.2 (C4), 151.2 (C2), 158.6 (C6).
Silylate 2 -OH 27. To a 250-mL round-bottom flask, add 10.56 g (17.59 mmol) S.6 and coevaporate twice with 50 mL anhydrous pyridine using the water aspirator. Dissolve the residue in 80 mL anhydrous pyridine. 28. Add 3.14 g (45.73 mmol) imidazole and 3.45 g (22.87 mmol) TBDMS-Cl. Stir the reaction mixture 2 hr at room temperature. 29. Periodically analyze reaction mixture by TLC on silica gel 60F plates using 1:1 (v/v) ethyl acetate/hexanes. If reaction is not complete after 2 hr, add 5% to 10% (w/w) TBDMS-Cl and 10% to 20% (w/w) imidazole, based on the original amounts added, and stir 1 hr. One should be able to observe the disappearance of substrate (S.6 Rf = 0.07) and appearance of the silylated derivatives. The reaction mixture contains the 2 -silylated isomer (S.7 Rf = 0.77) and 3 -silylated isomer (S.8 Rf = 0.58), as well as the 2 ,3 -disilylated derivative (Rf = 0.95).
30. Carefully add the reaction mixture to 250 mL saturated aqueous sodium bicarbonate solution in a 1-L separatory funnel and extract three times with 200 mL dichloromethane. 31. Extract combined organic layers twice with 200 mL of 10% aqueous NaH2 PO4 . During extraction with aqueous monobasic sodium phosphate, the reaction mixture forms an emulsion with relative ease. Vigorous shaking of the mixture should be avoided.
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32. Dry combined organic layers with ∼10 g anhydrous sodium sulfate and rotary evaporate. Coevaporate residue three times with ∼50 mL toluene. 33. Purify the reaction mixture by short-column chromatography. Prepare the column (150-mL sintered glass funnel, ∼100 g silica gel 60H) in dichloromethane and make the gradient by increasing the amount of methanol by 0.25% (v/v) for every 200 mL dichloromethane, up to 0.5% methanol. This column chromatography is used to separate the mixture of 2 -silylated and 3 -silylated isomers (which run together) from the 2 ,3 -disilylated derivative and other impurities.
34. Analyze the composition of the fractions (∼30 mL) by TLC on silica gel 60F plates using 9:1 (v/v) dichloromethane/methanol. Collect only the fractions containing 2 and 3 -silylated isomers (Rf = 0.61), and rotary evaporate to a solid foam. Dispose of fractions carrying 2 ,3 -disilylated derivative (Rf = 0.81). 35. To purify the mixture of 2 - and 3 -silylated isomers, prepare a column (150-mL sintered glass funnel, ∼100 g silica gel 60H) in toluene and make the gradient by increasing the amount of ethyl acetate by 0.5% (v/v) for every 100 mL toluene, up to 4% ethyl acetate. 36. Analyze composition of the fractions (∼30 mL) by TLC on silica gel 60F plates using 1:1 (v/v) ethyl acetate/hexanes. Collect separate fractions for pure 2 -isomer, mixed fractions containing mostly the 2 -isomer, and mixed fractions containing mostly the 3 -isomer. 37. Add ∼100 mL methanol to the fraction containing mostly 3 -isomer and leave at room temperature for ∼2 days. Isomerization of tert-butyldimethylsilyl groups usually takes ∼2 days and gives similar amounts of 2 - and 3 -isomers.
38. Rotary evaporate methanol with water aspirator and combine the residue with the mixed fractions containing mostly 2 -isomer. Purify this mixture as described in steps 35 and 36. If the synthesis is run on a large scale, a second isomerization of the fraction containing mostly 3 -isomer should be carried out, and the purification should be repeated.
39. Combine the fractions containing pure product, rotary evaporate, and coevaporate three times with ∼30 mL benzene in a 250-mL round-bottom flask. Dissolve in ∼10 mL benzene per 1 g product. 40. To lyophilize the product, freeze benzene solution in a dry ice/ethanol bath, rotating the flask in such a way that the freezing liquid evenly covers most of the surface of the round-bottom flask. Connect flask with a vacuum adaptor to a vacuum line and apply vacuum for 10 to 16 hr. Be sure that vacuum traps are large enough to condense all benzene and that solid benzene will not clog the vacuum line. Lyophilized product is very stable and can be stored safely for many years at 4◦ C.
Oligoribonucleotides with N6 -Alkyladenosine
5 -O-(4,4 -Dimethoxytrityl)-2 -O-tert-butyldimethylsilyl-6-methylthiopurine riboside (S.7): 8.29 g (66%). TLC (ethyl acetate/hexanes 1:1 [v/v]): Rf = 0.77. 1 H NMR (CDCl3 ): δ −0.20 (s, 3H, SiCH3 ), −0.04 (s, 3H, SiCH3 ), 0.76 (s, 9H, t-butyl), 2.67 (s, 3H, SCH3 ), 3.30–3.36 (m, 1H, H5 ), 3.43–3.48 (m, 1H, H5 ), 3.73 (s, 6H, OCH3 ), 4.37 (d, J = 3.5 Hz, 1H, H4 ), 4.45–4.50 (m, 1H, H3 ), 4.85 (t, J = 6.1 Hz, 1H, H2 ), 6.06 (d, J = 5.4 Hz, 1H, H1 ), 6.73 (d, J = 9.0 Hz, 4H, DMTr), 7.14–7.28 (m, 9H, DMTr), 8.20 (s, 1H, H2), 8.54 (s, 1H, H8). 13 C NMR (CDCl3 ): δ −5.2, −5.0 (SiCH3 ), 11.7 (SCH3 ), 17.8 (C(CH3 )), 25.5 (C(CH3 )3 ), 55.2 (OCH3 ), 63.4 (C5 ), 71.6 (C3 ), 75.6 (C2 ), 84.2 (C4 ), 88.2 (C1 ), 113.2 (DMTr), 126.9 (C5), 127.9, 128.1, 129.1, 130.0, 131.8, 135.6 (DMTr), 140.3 (C8), 144.5 (C4), 152.0 (C2), 158.5 (C6).
4.23.6 Supplement 17
Current Protocols in Nucleic Acid Chemistry
Convert to phosphoramidite 41. Place 1.10 g (1.54 mmol) S.7 in a 25-mL round-bottom flask closed with rubber septum equipped with a ventilation needle. Dry overnight under vacuum in a desiccator. 42. With 1- and 10-mL glass syringes and needles, add the following:
7.5 mL anhydrous acetonitrile (<20 ppm water) 0.40 mL (2.31 mmol) DIPEA 0.44 g (1.85 mmol, 0.41 mL) 2-cyanoethyl diisopropylchlorophosphoramidite. Stir reaction mixture 1 hr at room temperature. Because of its high viscosity, the phosphine should be added through the rubber septum into the flask as it sits on a balance to accurately measure the amount added by weight. This is especially important for small-scale syntheses.
43. Analyze reaction mixture by TLC on silica gel 60F plates using 45:45:10 (v/v/v) acetone/hexanes/TEA (S.9 Rf = 0.73) or on silanized silica gel 60F TLC plates using 7:3 (v/v) acetone/water (Rf = 0.28). If reaction is not complete after 1 hr, add 10% to 20% (w/w) DIPEA followed by 5% to 10% (w/w) 2-cyanoethyl diisopropylchlorophosphoramidite, based on the original amounts added, and stir 30 min. Silica gel TLC plates should be saturated with 45:45:10 (v/v/v) acetone/hexanes/TEA and dried before spotting the reaction mixture.
44. Add the reaction mixture to 50 mL saturated aqueous sodium bicarbonate in a 250-mL separatory funnel and extract three times with 50 mL dichloromethane containing 1% (v/v) TEA. 45. Dry combined organic layers with ∼5 g anhydrous sodium sulfate and rotary evaporate with water aspirator to a foam. 46. Purify the reaction mixture by short-column chromatography. Prepare the column (60-mL sintered glass funnel, ∼30 g silica gel 60H) in 90:10:1 (v/v/v) hexanes/ethyl acetate/TEA and make the gradient by increasing the amount of ethyl acetate by 5% (v/v) for every 100 mL mixture, up to 30% ethyl acetate. Keep amount of triethylamine constant (1%) during the entire purification. 47. Analyze composition of the fractions (∼20 mL) by TLC on silica gel 60F plates in 45:45:10 (v/v/v) acetone/hexanes/TEA. Collect separate fractions containing the phosphoramidite (S.9) and evaporate. 48. Lyophilize the product as described in steps 39 and 40. Lyophilized product is very stable and can be stored safely for at least 2 years at −20◦ C. 5 -O-(4,4 -Dimethoxytrityl)-2 -O-tert-butyldimethylsilyl-3 -O-[(2-cyanoethoxy)-(N,Ndiisopropylamino)]phosphinyl-6-methylthiopurine riboside (S.9): 1.20 g (85%). TLC (acetone/hexanes/triethylamine 45:45:10 [v/v/v]): Rf = 0.73; on silanized plates (acetone/water 7:3 [v/v]): Rf = 0.28. FAB-MS: m/z = 915.2. 1 H NMR (CDCl3 ): δ −0.22 (d, 3H, SiCH3 ), −0.04 (d, 3H, SiCH3 ), 0.75 (s, 9H, t-butyl), 1.04–1.21 (m, 12H, 2i-Pr), 2.73 (s, 3H, SCH3 ), 3.28–3.35 (m, 1H, H5 ), 3.49–3.66 (m, 5H, H5 , CH2 CH2 ), 3.78 (s, 6H, OCH3 ), 3.85–4.00 (m, 2H, 2CH), 4.34–4.44 (m, 2H, H4 , H3 ), 5.08 (t, J = 5.6 Hz, 1H, H2 ), 6.01 (d, J = 6.0 Hz, 0.5H, H1 ), 6.07 (d, J = 6.0 Hz, 0.5H, H1 ), 6.79–6.83 (m, 4H, DMTr), 7.21–7.48 (m, 9H, DMTr), 8.17 (s, 0.5H, H2), 8.20 (s, 0.5H, H2), 8.60 (s, 0.5H, H8), 8.62 (s, 0.5H, H8). 13 C NMR (CDCl3 ): δ −5.2, −4.7 (Si(CH3 )), 11.7 (SCH3 ), 17.9 (C(CH3 )), 20.0, 20.4, 24.6, (NC(CH3 )2 ), 25.6 (C(CH3 )3 ), 42.9, 43.5 (NC(CH3 )2 ), 55.2 (OCH3 ), 57.6, 58.8 (CH2 CH2 O-), 63.3 (C5 ), 72.7 (C3 ), 73.4, 74.5 (CH2 CH2 O-), 75.2 (C2 ), 83.9 (C4 ), 88.1 (C1 ), 113.2 (DMTr), 117.4 (CN), 126.9 (C5), 127.9, 128.1, 129.1, 130.0, 131.9, 135.6 (DMTr), 140.6 (C8), 144.5 (C4), 151.9 (C2), 158.5 (C6). 31 P NMR (CDCl3 ): δ 149.4, 150.8.
Synthesis of Modified Oligonucleotides and Conjugates
4.23.7 Current Protocols in Nucleic Acid Chemistry
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Figure 4.23.2 Acetylation, chlorination, introduction of methylthio group, acetyl deprotection, dimethoxytritylation, silylation, and phosphinylation of purine riboside derivative. Abbreviations: Ac, acetyl; DMAP, 4-dimethylaminopyridine; DMF, N,N-dimethylformamide; DMTr, 4,4 -dimethoxytrityl; Et, ethyl; i-Pr, isopropyl; TBDMS, tert-butyldimethylsilyl.
ALTERNATE PROTOCOL
Oligoribonucleotides with N6 -Alkyladenosine
SYNTHESIS OF THE 2-METHYLTHIO-6-CHLOROPURINE RIBOSIDE PHOSPHORAMIDITE Guanosine (S.10; Fig. 4.23.2) is O-peracetylated with acetic anhydride in acetonitrile (Matsuda, 1986) and the resulting 2 ,3 ,5 -tri-O-acetylguanosine (S.11) is converted to 2 ,3 ,5 -tri-O-acetyl-2-amino-6-chloropurine riboside (S.12) by phosphorus oxychloride treatment (Robins and Uzna˜nski, 1981). The 2-amino group of S.12 is transformed into 2-methylthio with dimethyl disulfide and isoamyl nitrite in acetonitrile using a 45-min treatment at 45◦ C (Nair and Fasbender, 1993). After column purification, 2 ,3 ,5 -tri-Oacetyl-2-methylthio-6-chloropurine riboside (S.13) is obtained in ∼53% yield. Acetyl groups are removed with triethylamine (TEA) in methanol. The product, 2-methylthio6-chloropurine riboside (S.14) is treated with 4,4 -dimethoxytrityl chloride (DMTr-Cl) in pyridine for 2 hr at room temperature. After column chromatography, 5 -O-DMTr-2methylthio-6-chloropurine riboside (S.15) is obtained in ∼75% yield (Smith et al., 1962). The reaction of S.15 with tert-butyldimethylsilyl chloride (TBDMS-Cl) gives a mixture of 2 - and 3 -isomers (S.16 and S.17; Ogilvie et al., 1978). The 2 -O-TBDMS isomer (S.16) is separated (∼57% yield) and treated with 2-cyanoethyl diisopropylchlorophosphoramidite in anhydrous acetonitrile in the presence of diisopropylethylamine. After purification by silica gel column chromatography, the phosphoramidite S.18 is obtained in ∼84% yield (Beaucage and Caruthers, 1981).
4.23.8 Supplement 17
Current Protocols in Nucleic Acid Chemistry
Additional Materials (also see Basic Protocol 1) Guanosine (S.10; anhydrous) 4-Dimethylaminopyridine (DMAP) Phosphorus oxychloride (POCl3 ), freshly distilled N,N-Dimethylaniline Dimethyl disulfide Isoamyl nitrite 500-mL and 1-L round-bottom flasks 140◦ and 60◦ C oil baths (silicone oil) 250-µL and 5-mL glass syringes with needles Acetylate guanosine 1. Add 8.49 g (30 mmol) anhydrous guanosine (S.10) and 0.64 g (2.25 mmol) DMAP to 375 mL anhydrous acetonitrile in a 1-L round-bottom flask and begin stirring. It is important to use anhydrous guanosine and not the hydrated form.
2. Add 16.7 mL (118 mmol) TEA and 10.2 mL (108 mmol) acetic anhydride and stir 30 min at room temperature. After a few minutes, solid guanosine dissolves.
3. Analyze reaction mixture by TLC (APPENDIX 3D) on a silica gel 60F TLC plate using 9:1 (v/v) dichloromethane/methanol (S.11 Rf = 0.31). The TLC analysis is performed with a short-wavelength UV lamp at 254 nm.
4. Concentrate to ∼50 mL on a rotary evaporator equipped with a water aspirator. 5. Add the reaction mixture carefully to 400 mL saturated aqueous sodium bicarbonate solution in a 1-L separatory funnel and extract three times with 150 mL dichloromethane. 6. Dry combined organic layers with ∼10 g anhydrous sodium sulfate and rotary evaporate. 2 ,3 ,5 -Tri-O-acetylguanosine (S.11): TLC (dichloromethane/methanol 9:1 [v/v]): Rf = 0.31.
Introduce 6-chloro group 7. To dried residue of S.11 (30 mmol) in a 500-mL round-bottom flask, add 82.5 mL (900 mmol) freshly distilled POCl3 and 5.24 mL (36 mmol) N,N-dimethylaniline. Close flask with a reflux condenser and place in a 140◦ C oil bath. Continue heating until 4 min from the moment it starts to reflux. CAUTION: Phosphorous oxychloride can be dangerous and should be handled carefully. It is recommended to run the reaction first on a small scale.
8. Cool flask in a wet ice bath (∼10 min). 9. Remove excess phosphorous oxychloride on the rotary evaporator. It is important to clean the rotary evaporator very well with dichloromethane before and after evaporation of phosphorus oxychloride.
10. Add 200 mL dichloromethane to the residue and transfer to a 1-L separatory funnel. 11. Wash four to six times with 500 mL water to remove most of the hydrochloric acid (until pH is ∼4). For the final wash, use 500 mL saturated aqueous sodium bicarbonate.
Synthesis of Modified Oligonucleotides and Conjugates
4.23.9 Current Protocols in Nucleic Acid Chemistry
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12. Dry the combined organic layers with ∼30 g anhydrous sodium sulfate and rotary evaporate with water aspirator. 13. Purify the reaction mixture by short-column chromatography (UNIT 2.4 & APPENDIX 3E). Prepare the column (150-mL sintered glass funnel; ∼100 g silica gel 60H) in dichloromethane and wash with dichloromethane only. 14. Analyze composition of the fractions (∼20 mL) by TLC on silica gel 60F plates using 9:1 (v/v) dichloromethane/methanol (S.12 Rf = 0.71). 15. Pool fractions containing product and rotary evaporate. 2 ,3 ,5 -Tri-O-acetyl-2-amino-6-chloropurine riboside (S.12): 9.49 g (74% relative to S.10). TLC (dichloromethane/methanol 9:1 [v/v]): Rf = 0.71. UV (MeOH): λmax = 224, 247, 310 nm; λmin = 234, 268 nm. 1 H NMR (CDCl3 ): δ 2.05 (s, 3H, CH3 ), 2.07 (s, 3H, CH3 ), 2.11 (s, 3H, CH3 ), 4.31–4.49 (m, 3H, H4 , H5 , H5 ), 5.71 (t, J = 5.6 Hz, 1H, H3 ), 5.93 (t, J = 5.6 Hz, 1H, H2 ), 5.98 (d, J = 4.8 Hz, 1H, H1 ), 7.86 (s, 1H, H8). 13 C NMR (CDCl3 ): δ 20.3, 20.4, 20.6 (C(O)CH3 ), 62.6 (C5 ), 70.1 (C3 ), 72.9 (C2 ), 80.0 (C4 ), 87.0 (C1 ), 120.3 (C5), 142.2 (C8), 151.1 (C4), 151.9 (C6), 167.3 (C2), 169.2, 169.4, 170.2 (C(O)CH3 ).
Introduce 2-methylthio group 16. In a 500-mL round-bottom flask, dissolve 7.23 g (16.9 mmol) S.12 in 140 mL anhydrous acetonitrile. 17. Add 15.2 mL (169 mmol) dimethyl disulfide and 4.53 mL (33.8 mmol) isoamyl nitrite. Heat the reaction mixture in a 60◦ C oil bath under refluxing condenser for 45 min. CAUTION: The reaction must be run in a well-ventilated fume hood.
18. Periodically analyze the reaction mixture by TLC on silica gel 60F plates using 9:1 (v/v) dichloromethane/methanol (S.13 Rf = 0.64) or on silanized silica gel 60F TLC plates in 7:3 (v/v) acetone/water (Rf = 0.70). 19. Concentrate to ∼30 mL on the rotary evaporator with water aspirator. Add the reaction mixture to 300 mL saturated aqueous sodium bicarbonate in a 1-L separatory funnel and extract three times with 100 mL dichloromethane. 20. Dry combined organic layers with ∼30 g anhydrous sodium sulfate and rotary evaporate. 21. Purify the reaction mixture by short-column chromatography. Prepare the column (150-mL sintered glass funnel; ∼100 g silica gel 60H) in dichloromethane and make the gradient by increasing the amount of methanol by 0.25% (v/v) for every 100 mL mixture, up to 2% methanol. 22. Analyze the composition of the fractions (∼30 mL) by TLC on silica gel plates using 9:1 (v/v) dichloromethane/methanol. 23. Collect fractions containing product (S.13) and rotary evaporate with water aspirator.
Oligoribonucleotides with N6 -Alkyladenosine
2 ,3 ,5 -Tri-O-acetyl-2-methylthio-6-chloropurine riboside (S.13): 4.13 g (53%). TLC (dichloromethane/methanol 9:1[v/v]): Rf = 0.64; on silanized TLC plates (acetone/water 7:3 [v/v]): Rf = 0.70. UV (MeOH): λmax = 233, 263, 306 nm; λmin = 248, 281 nm. 1 H NMR (CDCl3 ): δ 2.10 (s, 3H, CH3 ), 2.11 (s, 3H, CH3 ), 2.15 (s, 3H, CH3 ), 2.65 (s, 3H, SCH3 ), 4.29–4.48 (m, 3H, H4 , H5 , H5 ), 5.65 (t, J = 5.8 Hz, 1H, H3 ), 5.99 (t, J = 5.0 Hz, 1H, H2 ), 6.12 (d, J = 4.5 Hz, 1H, H1 ), 8.10 (s, 1H, H8). 13 C NMR (CDCl3 ): δ 14.8 (SCH3 ), 20.3, 20.4, 20.6 (C(O)CH3 ), 62.7 (C5 ), 70.1 (C3 ), 72.9 (C2 ), 80.0 (C4 ), 87.0 (C1 ), 120.3 (C5), 142.1 (C8), 151.2 (C4), 151.9 (C6), 167.3 (C2), 169.2, 169.4, 170.2 (C(O)CH3 ).
4.23.10 Supplement 17
Current Protocols in Nucleic Acid Chemistry
Remove acetyl groups and perform dimethoxytritylation 24. To 7.96 g (17.34 mmol) S.13 in 250-mL round-bottom flask, add 140 mL of 9:1 (v/v) methanol/TEA and leave for ∼48 hr at room temperature. 25. Periodically analyze reaction mixture by TLC on silica gel 60F plates using 9:1 (v/v) dichloromethane/methanol (S.14 Rf = 0.17). 26. When the deprotection of acetyl groups is completed, evaporate reaction mixture on the rotary evaporator and coevaporate the residue three times with ∼30 mL anhydrous pyridine. 27. Dissolve the residue in 75 mL anhydrous pyridine and add 6.45 g (19.07 mmol) DMTr-Cl. Stir reaction mixture for 1 to 1.5 hr at room temperature. 28. Periodically analyze reaction mixture by TLC on silica gel 60F plates using 9:1 (v/v) dichloromethane/methanol (S.15 Rf = 0.59). If reaction is not complete after 1.5 hr, add 5% to 10% (w/w) DMTr-Cl, based on initial amount added, and stir 1 hr. 29. Add the reaction mixture to 350 mL saturated aqueous sodium bicarbonate solution in a 1-L separatory funnel. Extract three times with 250 mL dichloromethane. 30. Dry combined organic layers with ∼30 g anhydrous sodium sulfate and rotary evaporate with water aspirator. Coevaporate the residue three times with ∼50 mL toluene. 31. Purify the reaction mixture by short-column chromatography. Prepare the column (150-mL sintered glass funnel; ∼100 g silica gel 60H) in dichloromethane and make the gradient by increasing the amount of methanol by 0.25% (v/v) for every 100 mL dichloromethane, up to 3% methanol. 32. Analyze composition of the fractions (∼30 mL) by TLC on silica gel 60F plates using 9:1 (v/v) dichloromethane/methanol. Combine the fractions containing product (S.15) and rotary evaporate to a solid foam. 5 -O-(4,4 -Dimethoxytrityl)-2-methylthio-6-chloropurine riboside (S.15): 8.26 g (75%). TLC (dichloromethane/methanol 9:1 [v/v]): Rf = 0.59. FAB-MS: m/z = 635.3. 1 H NMR (CDCl3 ): δ 2.49 (s, 3H, SCH3 ), 3.37–3.40 (m, 2H, H5 , H5 ), 3.74 (s, 6H, OCH3 ), 4.34 (q, J = 3.2 Hz, 1H, H4 ), 4.46–4.49 (m, 1H, H3 ), 4.85 (t, J = 5.9 Hz, 1H, H2 ), 6.01 (d, J = 5.4 Hz, 1H, H1 ), 6.74 (d, J = 9.0 Hz, 4H, DMTr), 7.17–7.24 (m, 9H, DMTr), 8.14 (s, 1H, H8). 13 C NMR (CDCl3 ): δ 14.8 (SCH3 ), 55.2 (OCH3 ), 63.5 (C5 ), 72.4 (C3 ), 75.6 (C2 ), 81.4 (C4 ), 86.8 (C1 ), 113.2 (DMTr), 127.1 (C5), 127.8, 129.1, 129.9, 135.3, 139.5 (DMTr), 142.4 (C8), 144.1 (C4), 147.3 (C6), 158.6 (C2).
Silylate 2 -OH 33. To a 250-mL round-bottom flask, add 5.44 g (8.57 mmol) S.15 and coevaporate with 30 mL anhydrous DMF. 34. Add 70 mL anhydrous DMF followed by 1.50 g (21.44 mmol) imidazole and 1.54 g (10.28 mmol) TBDMS-Cl. Stir the reaction mixture 5 hr at room temperature. 35. Periodically analyze reaction mixture by TLC on silica gel 60F plates using 4:6 (v/v) ethyl acetate/hexanes. If reaction is not complete after 5 hr, add 5% to 10% (w/w) TBDMS-Cl and 10% to 20% (w/w) imidazole, based on the original amounts added, and stir 2 hr. Silylated derivatives should appear in the reaction mixture. After reaction is complete, the mixture is composed of the 2 -silylated isomer (S.16 Rf = 0.70), the 3 -silylated isomer (S.17 Rf = 0.45), and the 2 ,3 -disilylated derivative (Rf = 0.86).
36. Add the reaction mixture to 250 mL saturated aqueous sodium bicarbonate solution in a 1-L separatory funnel and extract three times with 100 mL dichloromethane.
Synthesis of Modified Oligonucleotides and Conjugates
4.23.11 Current Protocols in Nucleic Acid Chemistry
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37. Extract combined organic layers twice with 100 mL of 10% aqueous NaH2 PO4 . During extraction with aqueous monobasic sodium phosphate, the reaction mixture forms an emulsion with relative ease. Vigorous shaking of the mixture should be avoided.
38. Dry combined organic layers with ∼20 g anhydrous sodium sulfate and rotary evaporate with water aspirator. Coevaporate residue three times with 30 mL toluene. 39. Purify the reaction mixture by short-column chromatography. Prepare the column (60-mL sintered glass funnel; ∼30 g silica gel 60H) in dichloromethane and elute products with dichloromethane. This column chromatography separates 2 -silylated and 3 -silylated isomers (which run together) from the 2 ,3 -disilylated derivative and other impurities.
40. Analyze composition of the fractions (∼20 mL) by TLC on silica gel 60F plates using 9:1 (v/v) dichloromethane/methanol. Collect only the fractions containing 2 and 3 -silylated isomers (Rf = 0.59), and rotary evaporate to a solid foam. Dispose of fractions carrying 2 ,3 -disilylated derivative (Rf = 0.82). 41. Purify the mixture of 2 - and 3 -silylated isomers again. Prepare the chromatography column (60-mL sintered glass funnel; ∼30 g silica gel 60H) in toluene and make the gradient by increasing the amount of ethyl acetate by 0.5% (v/v) for every 100 mL toluene, up to 5% ethyl acetate. 42. Analyze composition of the fractions (∼30 mL) by TLC on silica gel plates using 4:6 (v/v) ethyl acetate/hexanes. Collect separate fractions for pure 2 -isomer, mixed fractions containing mostly 2 -isomer, and mixed fractions containing mostly 3 isomer. 43. Add ∼70 mL methanol to the fraction containing mostly 3 -isomer and leave at room temperature for ∼2 days. Isomerization usually takes ∼2 days and gives equimolar amounts of 2 - and 3 -isomers.
44. Evaporate methanol and combine the residue with the mixed fractions containing mostly 2 -isomer. Purify this mixture as described in steps 41 and 42. If the synthesis is run on a large scale, a second isomerization of the fraction containing mostly 3 -isomer should be carried out, and the purification should be repeated.
45. Combine the fractions containing product, rotary evaporate, and coevaporate three times with ∼30 mL benzene in a 250-mL round-bottom flask. Dissolve in ∼10 mL benzene per 1 g product. 46. To lyophilize the product, freeze benzene solution in a dry ice/ethanol bath, rotating the flask in such a way that the freezing liquid evenly covers most of the surface of the round-bottom flask. Connect flask with a vacuum adaptor to a vacuum line and apply vacuum for 10 to 16 hr. Be sure that vacuum traps are large enough to condense all benzene and that solid benzene will not clog the vacuum line. Lyophilized product is very stable and can be stored safely for many years at 4◦ C.
Oligoribonucleotides with N6 -Alkyladenosine
5 -O-(4,4 -Dimethoxytrityl)-2 -O-tert-butyldimethylsilyl-2-methylthio-6-chloropurine riboside (S.16): 3.66 g (57%). TLC (ethyl acetate/hexanes 4:6 [v/v]): Rf = 0.70. 1 H NMR (CDCl3 ): δ −0.15 (s, 3H, SiCH3 ), 0.12 (s, 3H, SiCH3 ), 0.85 (s, 9H, t-butyl), 2.55 (s, 3H, SCH3 ), 3.41–3.52 (m, 2H, H5 , H5 ), 3.79 (s, 6H, OCH3 ), 4.27 (q, J = 3.5 Hz, 1H, H4 ), 4.37 (q, J = 4.2 Hz, 1H, H3 ), 4.85 (t, J = 7.2 Hz, 1H, H2 ), 6.08 (d, J = 5.4 Hz, 1H, H1 ), 6.80 (d, J = 9.0 Hz, 4H, DMTr), 7.22–7.43 (m, 9H, DMTr), 8.20 (s, 1H, H8). 13 C NMR (CDCl3 ): δ −5.1, −4.9 (Si(CH3 )), 14.7 (SCH3 ), 17.9 (C(CH3 )), 25.5 (C(CH3 )3 ), 55.2 (OCH3 ), 63.4 (C5 ), 71.6 (C3 ), 75.3 (C2 ), 84.3 (C4 ), 88.1 (C1 ), 113.3 (DMTr), 127.1 (C5), 127.97, 128.02, 130.0, 135.3, 135.4 (DMTr), 142.1 (C8), 150.9 (C4), 158.6 (C6), 166.9 (C2).
4.23.12 Supplement 17
Current Protocols in Nucleic Acid Chemistry
Convert to phosphoramidite 47. Place 0.51 g (0.68 mmol) S.16 in a 25-mL round-bottom flask closed with rubber septum equipped with a ventilation needle. Dry overnight under vacuum in a desiccator. 48. With 250-µL and 5-mL glass syringes and needles, add the following:
4.0 mL anhydrous acetonitrile (<20 ppm water) 0.18 mL (1.02 mmol) DIPEA 0.19 g (0.82 mmol, 0.18 mL) 2-cyanoethyl diisopropylchlorophosphoramidite. Stir reaction mixture 1 hr at room temperature. Because of its high viscosity, the phosphine should be added through the rubber septum into the flask as it sits on a balance to accurately measure the amount added by weight. This is especially important for small-scale syntheses.
49. Analyze reaction mixture by TLC on silica gel 60F plates using 45:45:10 (v/v/v) acetone/hexanes/TEA (S.18 Rf = 0.72) or on silanized 60F plates using 8:2 (v/v) acetone/water (Rf = 0.32). If reaction is not complete after 1 hr, add 10% to 20% (w/w) DIPEA followed by 5% to 10% (w/w) 2-cyanoethyl diisopropylchlorophosphoramidite, based on original amounts added, and stir for 0.5 hr. Silica gel TLC plates should be saturated with 45:45:10 (v/v/v) acetone/hexanes/TEA and dried before spotting the reaction mixture. For silanized plates for this derivative, 8:2 (v/v) acetone/water is preferred.
50. Add the reaction mixture to 50 mL saturated aqueous sodium bicarbonate solution in a 250-mL separatory funnel and extract three times with 50 mL dichloromethane containing 1% (v/v) TEA. 51. Dry combined organic layers with ∼5 g anhydrous sodium sulfate and rotary evaporate. 52. Purify the reaction mixture by short-column chromatography. Prepare the column (60-mL sintered glass funnel; ∼30 g silica gel 60H) in 90:10:1 (v/v/v) hexanes/ethyl acetate/TEA and make the gradient by increasing the amount of ethyl acetate by 5% (v/v) for every 100 mL mixture, up to 30% ethyl acetate. Keep the amount of TEA constant (1%) during the entire purification. 53. Analyze composition of the fractions by TLC on silica gel 60F plates in 45:45:10 (v/v/v) acetone/hexanes/TEA. Collect fractions containing phosphoramidite (S.18) and rotary evaporate. 54. Lyophilize the product as described in steps 45 and 46. Lyophilized product is very stable and can be stored safely for at least 2 years at −20◦ C. 5 -O-(4,4 -Dimethoxytrityl)-2 -O-tert-butyldimethylsilyl-3 -O-[(2-cyanoethoxy)-(N,Ndiisopropylamino)]phosphinyl-2-methylthio-6-chloropurine riboside (S.18): 0.54 g (84%). TLC (acetone/hexanes/triethylamine 45:45:10 [v/v/v]): Rf = 0.72; on silanized TLC plates (acetone/water = 8:2 [v/v]): Rf = 0.32. FAB-MS: m/z = 949.6. 1 H NMR (CDCl3 ): δ −0.20 (s, 3H, SiCH3 ), −0.04 (s, 3H, SiCH3 ), 0.78 (s, 9H, t-butyl), 1.16–1.22 (m, 12H, 2i-Pr), 2.55 (s, 3H, SCH3 ), 3.33–3.40 (m, 1H, H5 ), 3.45–3.50 (m, 1H, H5 ), 3.58–3.70 (m, 4H, -CH2 CH2 -), 3.78 (s, 3H, OCH3 ), 3.79 (s, 3H, OCH3 ), 3.84–4.02 (m, 2H, 2CH), 4.33–4.41 (m, 2H, H4 , H3 ), 4.87–4.91 (m, 1H, H2 ), 6.08 (d, J = 6.0 Hz, 0.5H, H1 ), 6.11 (d, J = 6.0 Hz, 0.5H, H1 ), 6.80–6.84 (m, 4H, DMTr), 7.22–7.45 (m, 9H, DMTr), 8.22 (s, 1H, H8). 13 C NMR (CDCl3 ): δ −5.1, −4.7 (Si(CH3 )), 14.6 (SCH3 ), 17.9 (C(CH3 )), 20.37, 20.45, 24.6 (NC(CH3 )2 ), 25.5 (C(CH3 )3 ), 42.9, 43.5 (NC(CH3 )2 ), 55.2 (OCH3 ), 57.5, 58.7 (CH2 CH2 O-), 63.4 (C5 ), 72.8 (C3 ), 73.6, 75.3 (CH2 CH2 O-), 76.1 (C2 ), 83.9 (C4 ), 87.7 (C1 ), 113.3 (DMTr), 117.4 (CN), 127.1 (C5), 128.0, 128.1, 130.0, 135.2, 135.5 (DMTr), 142.2 (C8), 150.8 (C4), 158.7 (C6), 166.7 (C2). 31 P NMR (CDCl3 ): δ 149.7, 150.7.
Synthesis of Modified Oligonucleotides and Conjugates
4.23.13 Current Protocols in Nucleic Acid Chemistry
Supplement 17
BASIC PROTOCOL 2
SYNTHESIS OF OLIGORIBONUCLEOTIDES CONTAINING N6 -ALKYLADENOSINE OR 2-METHYLTHIO-N6 -ALKYLADENOSINE The procedure starts with chemical synthesis of a precursor oligoribonucleotide carrying the 6-methylthiopurine riboside (S.19; Fig. 4.23.3) or 2-methylthio-6-chloropurine riboside (S.23; Fig. 4.23.4), which is then postsynthetically converted into N6 -alkyladenosine (S.22) or 2-methylthio-N6 -alkyladenosine (S.24), respectively. The synthesis was performed on an Applied Biosystems 392 synthesizer but could be done on any commercially available synthesizer (see APPENDIX 3C for additional details on automated synthesis). Commercially available 5 -O-DMTr-2 -O-TBDMS-3 -O-[(2-cyanoethoxy)(N,N-diisopropylamino)]phosphinyl derivatives of uridine and N-protected cytidine, adenosine, and guanosine are used in addition to the modified precursor phosphoramidite S.9 or S.18. In both cases, the coupling yield of modified phosphoramidite based on the trityl assay (APPENDIX 3C) is comparable to that of standard commercial 2 -O-silylated phosphoramidites. When the synthesis is completed (with DMTr-OFF), the solid support carrying the precursor oligoribonucleotide S.19 is oxidized with a 20 mM solution of magnesium monoperoxyphtalate in 9:1 (v/v) dioxane/water for 2.5 hr at room temperature to give a mixture of sulfoxide (S.20) and sulfone (S.21) derivatives of the precursor oligoribonucleotide. Under these conditions, oxidation of the thiomethyl group does
Oligoribonucleotides with N6 -Alkyladenosine
Figure 4.23.3 Oxidation and aminolysis of precursor oligoribonucleotide carrying 6methylthiopurine riboside to give the modified oligoribonucleotide containing N6 -alkyladenosine. Abbreviations: DMTr, 4,4 -dimethoxytrityl; i-Pr, isopropyl; R, alkyl chain of primary amine; TBDMS, tert-butyldimethylsilyl.
4.23.14 Supplement 17
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Figure 4.23.4 Aminolysis of precursor oligoribonucleotide carrying 2-methylthio-6-chloropurine riboside to give the modified oligoribonucleotide containing 2-methylthio-N6 -alkyladenosine. Abbreviations: DMTr, 4,4 -dimethoxytrityl; i-Pr, isopropyl; R, alkyl chain of primary amine; TBDMS, tert-butyldimethylsilyl.
not cause any side reactions (e.g., modification of native nucleobases). The support is washed with 9:1 (v/v) dioxane/water followed by acetonitrile and is then dried. The oxidation procedure is not required for the solid support carrying the precursor oligoribonucleotide S.23. Next, the solid support (with S.20/S.21 or S.23) is treated with the appropriate primary amine in acetonitrile. After amine treatment, the samples are evaporated, and a mixture of 3:1 (v/v) aqueous ammonia/ethanol is used (8 hr at 55◦ C) to deprotect and cleave the modified oligomer from the support. The support is filtered off and the solution dried down. The residue is coevaporated with anhydrous pyridine and treated with 1 M triethylammonium fluoride in pyridine for 48 hr at 55◦ C. The reaction mixture is evaporated and desalted on a Sep-Pak cartridge, and modified oligonucleotides are purified by TLC (APPENDIX 3D), high-performance liquid chromatography (HPLC; UNIT 10.5), or polyacrylamide gel electrophoresis (PAGE; UNIT 10.4).
Materials Commercial phosphoramidites: 5 -O-(4,4 -dimethoxytrityl)-2 -O-tertbutyldimethylsilyl-3 -O-[(2-cyanoethoxy)-(N,N-diisopropylamino)]phosphinyl uridine and N-protected cytidine, adenosine, and guanosine Modified phosphoramidite: 5 -O-(4,4 -dimethoxytrityl)-2 -O-tertbutyldimethylsilyl-3 -O-[(2-cyanoethoxy)-(N,N-diisopropyl)]phosphinyl-6methylthiopurine riboside (S.9; see Basic Protocol 1) or -2-thiomethyl-6-chloropurine riboside (S.18; see Alternate Protocol) Acetonitrile, anhydrous Magnesium monoperoxyphthalate hexahydrate (for oligomers containing S.9)
Synthesis of Modified Oligonucleotides and Conjugates
4.23.15 Current Protocols in Nucleic Acid Chemistry
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Dioxane (for oligomers containing S.9) Primary amine: e.g., isopentenylamine hydrochloride and triethylamine (TEA; for i6 A or ms2 i6 A) or methylamine (for m6 A or ms2 m6 A) Pyridine (reagent grade or better) 2 M methylamine in tetrahydrofuran (for m6 A or ms2 m6 A) Concentrated aqueous ammonia (28% to 32%) Ethanol 1 M triethylammonium fluoride in pyridine (or other desilylating agent) 10 mM ammonium acetate Automated oligonucleotide synthesizer (Applied Biosystems 392) 1.5- and 5-mL screw-top tubes 15-mL sintered glass funnels Water aspirator 55◦ C water bath Speedvac evaporator (Savant) Spin filters Sep-Pak cartridges (Waters) 10-mL disposable syringes Additional reagents and equipment for automated oligoribonucleotide synthesis (APPENDIX 3C) Synthesize desired oligoribonucleotide 1. Synthesize precursor oligoribonucleotide (S.19 or S.23) on a 1-µmol scale in DMTrOFF mode on an automated oligonucleotide synthesizer using the protocol provided by the manufacturer (also see APPENDIX 3C). Use phosphoramidites (commercial and modified) at 0.1 M in anhydrous acetonitrile. To achieve a 0.1 M solution of modified phosphoramidite, dissolve 0.25 g S.9 in 2.73 mL acetonitrile or 0.25 g S.18 in 2.64 mL acetonitrile. Oxidize thiomethyl group (for S.19 only) 2. Transfer the oligoribonucleotide-support from the synthesis column to a 1.5-mL screw-top microcentrifuge tube. 3. Dissolve 98.8 mg (0.2 mmol) magnesium monoperoxyphthalate hexahydrate in 10 mL of 9:1 (v/v) dioxane/water. Add 1 mL of this solution to the solid support and leave for 2.5 hr at room temperature with occasional shaking. 4. Filter off the oxidizing solution using 15-mL sintered glass funnel. 5. Wash the solid support three times with 5 mL of 9:1 (v/v) dioxane/water and three times with 5 mL anhydrous acetonitrile. Dry support with a water aspirator.
Treat with primary amine 6. Transfer oligoribonucleotide-support (mixture of S.20 and S.21 from step 5 or S.23 from step 1) to a 5-mL screw-top tube.
Oligoribonucleotides with N6 -Alkyladenosine
a. For general procedure, add 0.45 mL anhydrous acetonitrile and 0.05 mL of the appropriate primary amine. Seal tube tightly and leave for 12 hr at room temperature with shaking. b. For N6 -isopentenyladenosine or 2-methylthio-N6 -isopentenyladenosine (S.22 or S.24; R = isopentenyl), use 25 mg (0.21 mmol) isopentenylamine hydrochloride in 0.45 mL anhydrous pyridine and 0.05 mL (0.42 mmol) TEA, and incubate 12 hr at 55◦ C with shaking for S.22 or 2 to 5 hr at 55◦ C for S.24.
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Current Protocols in Nucleic Acid Chemistry
c. For N6 -methyladenosine or 2-methylthio-N6 -methyladenosine (S.22 or S.24; R = methyl) use 0.5 mL of 2 M methylamine in tetrahydrofuran and incubate 12 hr at 55◦ C with shaking for S.22 or 2 to 5 hr at 55◦ C for S.24. To convert precursor oligoribonucleotides into oligomers containing N6 neopentylyladenosine, N6 -(1-methylpropyl)adenosine, N6 -(1-methylbutyl)adenosine, N6 isopentyladenosine, N6 -propargyladenosine, and 2-methylthio-N6 -isopentyladenosine, the following primary amines are used: neopentylamine (2,2-dimethylpropylamine), 1-methylpropylamine (sec-butylamine), 1-methylbutylamine, isoamylamine (3methylbutylamine), propargylamine, and isoamylamine (3-methylbutylamine), respectively. When synthesizing these or any other modified adenosines, conversion yields will be affected by the choice and concentration of the primary amine used, as well as the temperature and time of the substitution reaction. When S.19 is used, the oxidative activation of the SCH3 is also critically important (Kierzek and Kierzek, 2003a).
7. Cool the reaction mixture at −20◦ C for 5 min, carefully open the sealed screw-top tube, and evaporate to dryness in a Speedvac evaporator.
Deprotect and cleave oligomer from support 8. Add 0.8 mL of 3:1 (v/v) concentrated aqueous ammonia/ethanol and leave tightly sealed for 8 hr at 55◦ C. 9. Cool at −20◦ C for 5 min and filter off the solid support using a spin filter. Wash the support twice with 1 mL water. 10. Transfer combined filtrates to a 5-mL screw-top tube and evaporate to dryness in the Speedvac evaporator.
Desilylate oligomer 11. Coevaporate the residue with 0.5 mL anhydrous pyridine in the Speedvac evaporator. 12. Add 0.5 mL of 1 M triethylammonium fluoride in pyridine, close tightly, and leave for 48 hr at 55◦ C. Removal of tert-butyldimethylsilyl groups can also be performed with tetra-nbutylammonium fluoride or triethylamine trihydrofluoride according to standard procedures.
13. Evaporate pyridine in the Speedvac evaporator.
Desalt product 14. Connect a Sep-Pak cartridge to the end of a 10-mL disposable syringe. Use the syringe to pass 10 mL acetonitrile followed by 10 mL of 10 mM ammonium acetate through the cartridge. The reaction mixture can also be desalted with a Sephadex G-10 or G-25 column (Amersham Biosciences).
15. Dissolve the oligonucleotide residue in 10 mL of 10 mM ammonium acetate and load on the cartridge. Wash column with 10 mL of 10 mM ammonium acetate. 16. Elute crude mixture of modified oligoribonucleotide with 5 mL of 3:7 (v/v) acetonitrile/water and evaporate with the Speedvac evaporator. 17. Purify the modified oligoribonucleotide using any standard method, such as TLC (APPENDIX 3D), HPLC (UNIT 10.5), or PAGE (UNIT 10.4). After purification, from a 1-µmol-scale synthesis, ∼10 to 30 OD260 units of modified oligoribonucleotide containing N6 -alkyladenosine (S.22) or 2-methylthio-N6 -alkyladenosine (S.24) are obtained (Table 4.23.1).
Synthesis of Modified Oligonucleotides and Conjugates
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Table 4.23.1 Retention Times, Synthesis Yields, and Spectrometric Mass Analysis of Oligoribonucleotides Obtained by Postsynthetic Modification
N6 -Alkyladenosine (A = R6 A) R6
Retention time (min)a
Yield (%)b
Massc
2-Methylthio-N6 -alkyladenosine (A = ms2 R6 A) Retention time (min)a
UACAUGUA H Methyl Neopentyl 1-Methylpropyl 1-Methylbutyl Isopentyl Isopentenyl Propargyl
24.0 24.2 33.8 30.4 34.3 34.6 33.0 25.8
64 40 47 42 61 52 41 40
24.0 24.5 32.6 29.1 33.2 33.2 31.4 25.2
64 47 51 72 71 45 56 64
2494.99 2508.93 2564.93 2550.90 2564.95 2564.78 2563.75 2532.64
26.9 30.0
49 35
2540.91 2554.79
44.7
24
2608.97
UACAUGUA 2494.99 2508.92 2564.87 2550.88 2565.01 2564.94 2562.92 2532.98
25.3 27.9
48 64
2541.02 2555.04
44.7
41
2608.93
UACAUGUA H Methyl Neopentyl 1-Methylpropyl 1-Methylbutyl Isopentyl Isopentenyl Propargyl
24.0 25.2 35.1 31.6 35.9 36.3 34.3 26.7
64 51 58 77 51 51 25 49
UACAUGUA 2494.99 2508.61 2564.90 2551.05 2564.84 2564.78 2563.30 2532.81
27.2 31.0
27 54
2541.01 2555.00
48.8
65
2608.57
ACAUGUA H Methyl Neopentyl 1-Methylpropyl 1-Methylbutyl Isopentyl Isopentenyl Propargyl Oligoribonucleotides with N6 -Alkyladenosine
26.0 27.1 38.5 34.6 39.2 39.7 38.0 26.0
74 49 69 75 91 55 65 42
Massc
UACAUGUA
UACAUGUA H Methyl Neopentyl 1-Methylpropyl 1-Methylbutyl Isopentyl Isopentenyl Propargyl
Yield (%)b
ACAUGUA 2188.35 2202.32 2258.49 2244.46 2258.52 2258.47 2256.11 2226.46
26.3 31.1
35 53
2234.65 2249.03
53.6
20
2302.92
continued
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Table 4.23.1 Retention Times, Synthesis Yields, and Spectrometric Mass Analysis of Oligoribonucleotides Obtained by Postsynthetic Modification, continued
N6 -Alkyladenosine (A = R6 A) R6
Retention time (min)a
Yield (%)b
Massc
2-Methylthio-N6 -alkyladenosine (A = ms2 R6 A) Retention time (min)a
Yield (%)b
Massc
CCGGUCmUCCAAAACCGG H Methyl
29.9 32.9
16 24
5445.18 5459.53
a HPLC: Supelco RP C18; eluent A = 100 mM triethylammonium acetate; eluent B = 1:1 (v/v) acetonitrile/A; gradient
= 1% B/min. b Overall yield refers to precursor oligomer synthesis, postsynthetic modification, and deprotection and is based on C18
HPLC analysis. c Molecular weights measured by atmospheric pressure ionization–electrospray liquid chromatography/mass spectroscopy
(API-ES LC-MS) in the negative ionization mode.
COMMENTARY Background Information There are 96 different natural modifications of nucleotides occurring in all types of RNA (Limbach et al., 1994; Bjork, 1995). Transfer RNAs are particularly rich in modified nucleotides, with 80 different modified nucleotides known. The modifications are mostly present in single-stranded regions of tRNA such as hairpin and multibranch loops. Positions 34 and 37 of the anticodon arm are particularly highly modified. The functions of modified nucleotides are not clearly defined, but it is known that they affect structure and biological activities of RNA, influence the secondary and tertiary interactions of tRNA, change hydrogen bond and stacking interactions as well as puckering of ribose residues, and influence binding of divalent cations (Persson, 1993; Grosjean et al., 1998). Among 19 modifications of adenosine, 15 have either a substitution on the N6 -exocyclic amine group or a 2-methylthio group with a substitution on the N6 -exocyclic amine group at the same time. Examples include N6 threonylcarbamoyladenosine (t6 A), N6 -isopentenyladenosine (i6 A), N6 -methyladenosine (m6 A), 2-methylthio-N6 -isopentenyladenosine (ms2 i6 A), and 2-methylthio-N6 methyladenosine (ms2 m6 A). Because these residues are present at position 37 of the tRNA anticodon loop, studies of their biological function and of their influence on RNA structure are most important. The syntheses of oligoribonucleotides containing the last four modifications are performed by the methods described in this unit.
Studies of modified RNA activities are significantly limited by access to modified oligoribonucleotides. In the literature, syntheses of RNA carrying simple modifications are well described. Introduction of nucleosides bearing complex modifications is still challenging, however. In 1978, Wiewi´orowski and co-workers described the first chemical synthesis of an anticodon loop heptamer containing t6 A (Adamiak et al., 1978). Recently, several chemical syntheses of anticodon loops and arms containing t6 A were published (Sochacka, 1999; Boudou et al., 2000; Stuart et al., 2000; Sundaram et al., 2000). The most important improvement in the synthesis of oligoribonucleotides containing N6 -alkyladenosines and 2-methylthioN6 -alkyladenosines is the application of a postsynthetic approach to introducing the modification, which is the approach described in this unit. This allows very simple simultaneous synthesis of several modified oligoribonucleotides from only one precursor oligoribonucleotide. The first crucial element of this approach is selection and development of the efficient synthesis of the modified phosphoramidites S.9 and S.18. The second crucial element is setting conditions for postsynthetic modification of the precursor form of the oligoribonucleotides to obtain a very high yield of modification and at the same time keep elements of the oligoribonucleotides intact during the entire modification process. The longest modified oligoribonucleotides that have been obtained by the described
Synthesis of Modified Oligonucleotides and Conjugates
4.23.19 Current Protocols in Nucleic Acid Chemistry
Supplement 17
postsynthetic modification of precursor oligonucleotides are octadecamers, which correspond to the length of the anticodon arm of tRNA. Because of the high yield of transformation of the precursor oligonucleotides, however, the final length of the modified oligoribonucleotides should be limited only by the general reliability of chemical RNA synthesis and purification methods. This high yield also should allow the introduction of several N6 -alkyladenosines and/or 2-methylthio-N6 -alkyladenosines into an oligoribonucleotide, if the N6 -alkyl substituent is the same.
cubation time is used for the precursor oligoribonucleotides (2.5 hr at room temperature) because of the heterogeneous conditions of oxidation and because no side products of oxidation have been observed. During modification of the precursor form of the oligoribonucleotide, some problems may arise with transformation into N6 isopentenyladenosine and 2-methylthio-N6 isopentenyladenosine oligonucleotides (S.22 and S.24, respectively; R = isopentenyl). This is due to the low solubility of isopentenylamine hydrochloride in acetonitrile or even in pyridine. This makes the deprotection process inconvenient. This salt is removed using a SepPak cartridge before purification, however.
Critical Parameters
Oligoribonucleotides with N6 -Alkyladenosine
The synthesis of modified phosphoramidites and transformation of precursor oligoribonucleotides into oligonucleotides containing N6 -alkyladenosine and 2methylthio-N6 -alkyladenosine do not present significant difficulties. Some prior experience in the chemistry of nucleosides and oligonucleotides is nonetheless very useful. It is important to be very careful during synthesis of 2 ,3 ,5 -tri-O-acetyl-2-amino6-chloropurine riboside (S.12) because the reaction, particularly when performed on a large scale, uses several hundred milliliters of phosphorus oxychloride that has to be refluxed at 140◦ C for a short period of time only. It is best to perform this reaction first on a small scale for practice. In the next reaction, during synthesis of 2 ,3 ,5 -tri-O-acetyl-2-methylthio-6chloropurine riboside (S.13), 10% to 20% 2 ,3 ,5 ,N2 -tetraacetyl-6-chloropurine riboside is also formed, and this side product must be separated by short-column chromatography. It is also very important to carefully separate the 2 - and 3 -O-TBDMS isomers during synthesis of 5 -O-DMTr-2 -O-TBDMS ribosides S.7 and S.16. A first prepurification column allows for a relatively easy separation of both isomers during the second silica gel column purification. The oxidation of the 6-methylthio group with magnesium monoperoxyphthalate in Basic Protocol 2 must be complete, because only the oxidation products (6-methylsulfone and 6-methylsulfoxide derivatives) can be converted into N6 -alkylated adenosine derivatives by treatment with the appropriate amine. Experiments carried out by the authors with protected 6-methylthiopurine riboside in this oxidation solution demonstrate that the oxidation of the 6-methylthio group was completed within 1 hr at room temperature. A longer in-
Anticipated Results Synthesis of precursor phosphoramidites is not trivial but can be achieved by someone with basic experience in organic chemistry. The yield of particular steps of the synthesis ranges between 50% and 100%. Some reactions have high yield and no side products, which allows subsequent reaction without any purification of the intermediate. Precursor oligoribonucleotides are synthesized on an Applied Biosystems 392 synthesizer using a standard procedure of RNA synthesis. On the basis of HPLC analysis of the crude reaction mixture, the transformation of precursor oligoribonucleotide into oligoribonucleotide containing N6 -alkyladenosine or 2-methylthio-N6 -alkyladenosine proceeds with an 80% to 90% yield. The efficiency of transformation is dependent on structure (mostly a steric hindrance effect of the alkyl chain) and nucleophilicity of the primary amine. The modified oligoribonucleotides are purified from the entire crude reaction mixture by TLC or HPLC with a 20% to 60% overall yield (Table 4.23.1).
Time Considerations Synthesis of modified phosphoramidites (S.9 and S.18) takes ∼2 to 3 weeks for each precursor. The transformation of precursor oligoribonucleotides (S.19 and S.23) and purification of modified oligoribonucleotides (S.22 and S.24) requires ∼1 week.
Literature Cited Adamiak, R.W., Biala, E., Grzeskowiak, K., Kierzek, R., Kraszewski, A., Markiewicz, W.T., Okupniak, J., Stawinski, J., and Wiewi´orowski, M. 1978. The chemical synthesis of the anticodon loop of an eukaryotic initiator tRNA containing the hypermodified nucleoside N6 -/
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Current Protocols in Nucleic Acid Chemistry
N-threonylcarbamoyl/-adenosine/t6 A/1 . Acids Res. 5:1889-1905.
Nucl.
Beaucage, S.L. and Caruthers, M.H. 1981. Deoxynucleoside phosphoramidites—a new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Lett. 22:1859-1862. Bj¨ork, G.R. 1995. tRNA: Structure, biosynthesis and function. In Biosynthesis and Function of Modified Nucleosides (D. S¨oll and U. RajBhandary, eds.) pp. 165-206. American Society for Microbiology Press, Washington, D.C. Boudou, V., Langridge, J., Van Aerchot, A., Hendrix, C., Millar, A., Weiss, P., and Herdewijn, P. 2000. Synthesis of the anticodon hairpin tRNAf Met containing N-{[9-(β-Dribofuranosyl)-9H-purin-6-yl]carbamoyl}L-threonine. Helv. Chim. Acta 83:152-161. Cava, M.P. and Levinson, M.J. 1985. Thionation reactions of Lawesson reagents. Tetrahedron 41:5061-5087. Grosjean, H., Houssier, C., Romby, P., and Marquet, R. 1998. Modulatory role of modified nucleotides in RNA loop-loop interaction. In Modification and Editing of RNA (H. Grosjean and R. Benne, eds.) pp. 113-133. American Society of Microbiology Press, Washington, D.C. Kierzek, E. and Kierzek, R. 2003a. The synthesis of oligoribonucleotides containing N6 -alkyladenosines and 2-methylthio-N6 alkyladenosines via post-synthetic modifications of precursor oligomers. Nucl. Acids Res. 31:4461-4471. Kierzek, E. and Kierzek, R. 2003b. The thermodynamic stability of RNA duplexes and hairpins containing N6 -alkyladenosines and 2-methylthio-N6 -alkyladenosines. Nucl. Acids Res. 31:4472-4480. Limbach, P.A., Crain, P.F., and McClosky, J.A. 1994. Summary: The modified nucleosides of RNA. Nucl. Acids Res. 22:2183-2196. Matsuda, A. 1986. A convenient method for the selective acylation of guanine nucleosides. Synthesis 5:385-386. Nair, V. and Fasbender, A.J. 1993. C-2 functionalized N6-cyclosubstituted adenosines—highly selective agonists for the adenosine-a1-receptor. Tetrahedron 49:2169-2184.
Ogilvie, K.K., Beaucage, S.L., Schifman, A.L., Theriault, N.Y., and Sadana, K.L. 1978. The synthesis of oligoribonucleotides. II. The use of silyl protecting groups in nucleoside and nucleotide chemistry. VIII. Can. J. Chem. 56:27682780. Persson, B.C. 1993. Modification of tRNA as a regulatory device. Mol. Microbiol. 8:1011-1016. Robins, M.J. and Uzna˜nski, B. 1981. Nucleic-acid related compounds. 33. Conversions of adenosine and guanosine to 2,6-dichloro, 2-amino-6chloro, and derived purine nucleosides. Can. J. Chem. 59:2601-2607. Smith, M., Rammler, D.H., Goldberg, I.H., and Khorana, H.G. 1962. Studies on polynucleotides. XVI. Specific synthesis of the C3-C5 internucleotide linkage. Synthesis of uridyl-(3 5 )-uridine and uridyl-(3 -5 )-adenosine. J. Am. Chem. Soc. 84:430-440. Sochacka, E. 1998. The chemical synthesis of E.coli tRNALys anticodon loop fragment and its analogues. Nucleosides Nucleotides 17:327338. Stuart, J.W., Gdaniec, Z., Guenther, R., Marszalek, M., Sochacka, E., Malkiewicz, A., and Agris, P.F. 2000. Functional anticodon architecture of human tRNALys3 includes disruption of intraloop hydrogen bonding by the naturally occurring amino acid modification, t6 A. Biochemistry 39:13396-13404. Sundaram, M., Crain, P.F., and Davis, D.R. 2000. Synthesis and characterization of the native anticodon domain of E. coli. Simultaneous incorporation of modified nucleosides mnm5 s2 U, t6 A, and pseudouridine using phosphoramidite chemistry. J. Org. Chem. 65:5609-5614. Wetzel, R. and Eckstein, F. 1975. Synthesis and reactions of 6-methylsulfonyl-9-β-Dribofuranosylpurine. J. Org. Chem. 40:658660.
Contributed by Elzbieta Kierzek and Ryszard Kierzek Institute of Bioorganic Chemistry, Polish Academy of Sciences Poznan, Poland
Synthesis of Modified Oligonucleotides and Conjugates
4.23.21 Current Protocols in Nucleic Acid Chemistry
Supplement 17
Oligodeoxyribonucleotide Analogs Functionalized with Phosphonoacetate and Thiophosphonoacetate Diesters
UNIT 4.24
Phosphonoacetate and thiophosphonoacetate oligodeoxyribonucleotides are analogs wherein the phosphodiester backbone is replaced with phosphonoacetate or thiophosphonoacetate internucleotide linkages (Fig. 4.24.1). The reactive N,N(diisopropylamino)phosphinyl acetate monomers are convenient to prepare from standard protected nucleosides and give high-yield synthesis of oligodeoxyribonucleotides on solid supports. Phosphonoacetate and thiophosphonoacetate oligodeoxyribonucleotides are highly stable to nucleases. They are water-soluble and anionic at neutral pH (pKa values of 3.8 and 3.9). They hybridize with complementary DNA or RNA to yield duplexes having stabilities comparable to those achieved with phosphorothioate DNA. Both analogs stimulate RNase H degradation of complementary RNA. Chimeric oligomers end-capped with phosphonoacetate or thiophosphonoacetate internucleotide linkages have been shown to be more active than natural DNA at stimulating RNase H in vitro.
Figure 4.24.1 Structures of phosphonoacetate, thiophosphonoacetate, phosphodiester, and phosphorothioate internucleotide linkages. B: thymin-1-yl, cytosin-1-yl, adenin-9-yl, or guanin-9-yl.
Contributed by Douglas J. Dellinger, Christina M. Yamada, and Marvin H. Caruthers Current Protocols in Nucleic Acid Chemistry (2004) 4.24.1-4.24.26 C 2004 by John Wiley & Sons, Inc. Copyright
Synthesis of Modified Oligonucleotides and Conjugates
4.24.1 Supplement 18
Figure 4.24.2
Synthesis of 1,1-dimethyl-2-cyanoethyl [bis(N,N-diisopropylamino)phosphinyl]acetate.
DMTrO
O OH
B
DMTrO
O
3 1H -tetrazole
4a B = thymin-1-yl b = 4-N -acetylcytosin-1-yl c = 6-N -benzoyladenin-9-yl d = 2-N -isobutyrylguanin-9-yl
B
O
O P
CH2
C
i -Pr2N
CH3 O
C
CH2CN
CH3 5a-d
Figure 4.24.3 Preparation of protected 2 -deoxynucleoside-3 -O-(N,N-diisopropylamino)phosphinyl acetates. DMTr, 4,4 dimethoxytrityl.
Modified oligodeoxyribonucleotides containing thiophosphonoacetate linkages with stable, intact esters are readily taken up by cells in the absence of cationic lipids. This unit describes the solid-phase synthesis and purification of modified oligodeoxyribonucleotides (ODNs) having phosphonoacetate (PACE) and thiophosphonoacetate (S-PACE) internucleotide linkages (Dellinger et al., 2003). The first protocol outlines the preparation of appropriate synthons (Figs. 4.24.2 and 4.24.3; see Basic Protocol 1). The next two protocols illustrate the synthesis of PACE or S-PACE modified DNA (Fig. 4.24.4; see Basic Protocol 2) and chimeric oligomers (Sheehan et al., 2003) having either of these analogs combined with natural or phosphorothioate internucleotide linkages (Fig. 4.24.5; see Basic Protocol 3). The final protocols describe the drying of monomers (see Support Protocol) in addition to the purification and characterization of ODNs (see Basic Protocol 4). CAUTION: All reactions should be carried out in a well-ventilated fume hood and contact with chemicals should be avoided. It is essential that reagents, solvents, and synthons be used under anhydrous conditions in order to maximize yield and success in preparing these oligomers. To avoid incompatibilities, a solvent-resistant Teflon head pump capable of 34 liter/min and a maximum vacuum of 1.5 torr is used for all solvent removals performed on a rotary evaporator. All intermediates and products are stored in desiccators at −20◦ C, which are allowed to warm to room temperature before they are opened. BASIC PROTOCOL 1 Oligonucleotides with Phosphonoacetate and Thiophosphonoacetate Diesters
SYNTHESIS OF PROTECTED 2 -DEOXYNUCLEOSIDE3 -O-(N,N-DIISOPROPYLAMINO)PHOSPHINYL ACETATES Generating protected 2 -deoxynucleoside-3 -O-(N,N-diisopropylamino)phosphinyl acetates requires three steps: (1) preparation of 1,1-dimethylcyanoethyl bromoacetate (S.2; Fig. 4.24.2), (2) condensation of S.2 with bis(diisopropylamino)chlorophosphine to yield the phosphinylating reagent 1,1-dimethyl-2-cyanoethyl [bis(N,Ndiisopropylamino)phosphinyl]acetate (S.3), and (3) synthesis and purification of
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protected 2 -deoxynucleoside-3 -O-(N,N-diisopropylamino)phosphinyl acetates (S.5; Fig. 4.24.3). The first step is the synthesis of an appropriately protected bromoacetate. The 1,1dimethyl-2-cyanoethyl ester is recommended, and is prepared from 3-hydroxy-3methylbutyronitrile and bromoacetyl bromide. However, the reaction conditions for generating this reagent are quite general. Any number of alcohols can be used with these protocols to produce the corresponding esters. The second step is synthesis of 1,1-dimethyl-2-cyanoethyl [bis(N,N-diisopropylamino)phosphinyl]acetate. This phosphinylating reagent is prepared by a Reformatsky reaction (Bayless and Hauser, 1954) using the zinc bromide derivative of an ester-protected acetic acid and bis(N,N-diisopropylamino)chlorophosphine. The third step, preparation of protected 2 -deoxynucleoside-3 -O-(N,N-diisopropylamino)phosphinyl acetates, can be conveniently performed starting with standard 5 dimethoxytrityl-protected 2 -deoxynucleosides. For cost and convenience, the use of Nacetyl, N-benzoyl, and N-isobutyryl protection are recommended for dC, dA, and dG, respectively. These reagents can be obtained from ChemGenes. Other fast-deprotecting exocyclic amine–blocking groups can be used as well. As described in the accompanying protocols, the use of exocyclic amine–blocking groups compatible with fast deprotection conditions (such as methylamine for 15 min) is important because exposure of the oligodeoxyribonucleotide products to highly basic solutions must be limited. Prolonged exposure to strong base can result in decarboxylation of PACE/S-PACE internucleotide linkages and chain cleavage. The synthesis chemistry is compatible with most modified 2 -deoxynucleosides. The final protected 2 -deoxynucleoside-3 -O-(N,Ndiisopropylamino)phosphinyl acetates are purified by silica-gel column chromatography. CAUTION: Many phosphines are highly toxic or neuroactive agents. The phosphines described in these protocols are new reagents. Additional safety precautions should always be taken when working with novel reagents of this type, as they are uncharacterized relative to toxicity. It is extremely important to prevent inhalation or skin contact with these reagents. As in any laboratory situation, proper safety equipment should include a certified working fume hood, safety glasses, a lab coat, and reagent-impenetrable gloves. All glassware used in the synthesis or purification of these reagents should be thoroughly decontaminated before removal from the fume hood. CAUTION: Prior to initiating synthesis, prepare the fume hood to handle large amounts of gaseous HBr. It is important to know the local regulations regarding such hazardous material and the proper way to trap or vent this acid. When trapping HBr, be sure to carefully calculate the mole equivalents of generated acid and ensure that the trap has enough capacity. A variety of effective procedures for working with mineral acids can be found in common laboratory manuals; the method of choice will depend upon local regulations.
Materials Argon, dry Toluene, anhydrous (Aldrich) Bromoacetyl bromide (S.1, Fig. 4.24.2; Aldrich) Nitrogen, dry (optional) 50-g bottle of 3-hydroxy-3-methylbutyronitrile (Fluka) Bis(diisopropylamino)chlorophosphine (Digital Specialty Chemicals) Tetrahydrofuran, anhydrous (THF; Aldrich) Zinc metal, granular (Aldrich)
Synthesis of Modified Oligonucleotides and Conjugates
4.24.3 Current Protocols in Nucleic Acid Chemistry
Supplement 18
Acetonitrile, anhydrous Phosporic acid/CD3 CN NMR standard Reagent-grade hexanes, anhydrous 5 -O-(4,4 -Dimethoxytrityl)-protected 2 -deoxynucleoside (S.4; ChemGenes): 5 -O-DMTr-N6 -benzoyl-2 -deoxyadenosine 5 -O-DMTr-N4 -acetyl-2 -deoxycytidine 5 -O-DMTr-N2 -isobutyryl-2 -deoxyguanosine 5 -O-DMTr-2 -deoxythymidine Dichloromethane, anhydrous (Aldrich) 0.45 M tetrazole in acetonitrile (Glen Research) Ethyl acetate, reagent grade Diisopropylethylamine, anhydrous (DIPEA; Aldrich) Silica gel 60 for medium-pressure liquid chromatography (MPLC; 230 to 400 mesh; Aldrich) 120◦ C oven 500-mL and 1-liter round-bottom flasks with 24/40 joints Drying tubes containing calcium sulfate (Drierite) and equipped with a 24/40 joint PTFE-coated stir bar 125-mL, 250-mL, and 2-L Erlenmeyer flasks Top-loading balance Addition funnels with 24/40 joints, pressure-equalization arms (100- and 500-mL capacity), and PTFE stopcocks 325-mm Friedrich’s condensers with 24/40 joints (Labglass) Acid vapor trap and acid-resistant tubing Heating mantle to fit a 1-liter round-bottom flask, with controller Rotary evaporator with solvent-resistant Teflon head pump 1- and 2-liter recovery flasks (Labglass) Rotary evaporator trap (Labglass) Short-path distillation apparatus (Labglass) Three-flask distribution receiver Vacuum pump with an inlet acid vapor trap Capillary bleed tube (Kontes Glass) or needle valve attached to a Y-connector Powder funnels 24/40 rubber septa Hand-held heat gun (Aldrich) 500-mL separatory funnel Ground-glass 24/40 joint 50-mL glass pipet Glass-backed silica-gel TLC plates with fluorescent indicator (Aldrich) TLC developing tank (Aldrich) UV viewing cabinet (Aldrich) Flash-chromatography column (Aldrich cat) and flow controller Sea sand 350-mL fritted glass Buchner funnel, medium porosity, fitted with a 24/40 vacuum adapter (Labglass) Vacuum desiccator and solid PTFE vacuum pump with maximum vacuum of 1.5 torr (Aldrich) Additional reagents and equipment for 31 P NMR, thin-layer chromatography (TLC; APPENDIX 3D), column chromatography (APPENDIX 3E), and fast atom bombardment mass spectrometry (FAB-MS) Oligonucleotides with Phosphonoacetate and Thiophosphonoacetate Diesters
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Synthesize 1,1-dimethylcyanoethyl bromoacetate 1. Dry all glassware overnight in an oven at 120◦ C. 2. Remove the 1-liter round-bottom flask with 24/40 joint from the oven and cool under a stream of dry argon. Place the flask in a fume hood and secure it on a magnetic stir plate. 3. Add 500 mL anhydrous toluene and a PTFE-coated stir bar to the flask. Cap with a drying tube. Secure the entire apparatus to the fume hood’s lattice support or a support stand. 4. Remove the 125-mL Erlenmeyer flask from the oven and cool to room temperature under a stream of dry argon. Tare the flask in the hood using a top-loading balance. 5. Weigh out 108 g (600 mmol) bromoacetyl bromide (S.1) into the flask and then quickly transfer to the toluene-containing flask by temporarily removing the drying tube, adding the bromoacetyl bromide, and then reattaching the drying tube. 6. Rinse the Erlenmeyer flask with anhydrous toluene and add the rinse to the solution. Stir on the magnetic stir plate to completely dissolve the bromoacetyl bromide. 7. Remove the 100-mL addition funnel from the oven and cool to room temperature using a stream of dry nitrogen or argon. Make sure that the PTFE stopcock is closed and tightened to prevent leaks. 8. Transfer the entire contents of a 50-g (500 mmol) bottle of 3-hydroxy-3methylbutyronitrile to the addition funnel. Rinse the reagent bottle with anhydrous toluene and add to the funnel. 9. Remove the drying tube from the 1-liter round-bottom flask and attach the addition funnel. Place the drying tube on the addition funnel, open the stopcock, and add 3-hydroxy-3-methylbutyronitrile dropwise to the stirred solution of bromoacetyl bromide. 3-Hydroxy-3-methylbutyronitrile is a tertiary alcohol that reacts very slowly with bromoacetyl bromide. This slow reactivity allows one to add the two components rapidly without cooling the flask. If another alcohol is used, especially a primary alcohol, the reaction can be highly exothermic at room temperature. When using another alcohol, it is important to first perform a test reaction on a small scale: cool the flask containing bromoacetyl bromide in an ice bath and add the alcohol dropwise to the stirred reaction mixture. This test reaction is used to determine how exothermic the reaction will become and how much cooling will be required for a larger-scale reaction. It is also important to remember that the efficiency of cooling the contents of a larger-volume flask with an external cooling bath decreases significantly as the ratio of surface area to volume decreases.
10. Remove the funnel and fit a 325-mm Friedrich’s condenser with attached drying tube to the flask. Fix an acid vapor trap to the drying tube using acid-resistant tubing. Place the flask in a mantle and heat the reaction mixture to reflux (∼110◦ C) overnight. 11. Cool the flask to room temperature and remove the toluene using a rotary evaporator. In order to minimize loss or contamination, transfer the solution to a 2-liter recovery flask and attach a rotary evaporator trap between the recovery flask and the evaporator. Toluene is often considered a difficult solvent to remove on a rotary evaporator due to its boiling point and tendency to “bump.”
12. Purify the product as an oil by vacuum distillation using a short-path distillation apparatus, three-flask distribution receiver, and vacuum pump with inlet acid vapor trap. Perform distillation at ∼0.1 mmHg using a capillary bleed tube inserted into the distillation flask or a needle valve attached to a Y-connector on the vacuum line.
Synthesis of Modified Oligonucleotides and Conjugates
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Distill the oil into two components: an initial minor fraction with a wide boiling range and the major fraction at 120◦ to 122◦ C. Store the major fraction up to 6 months at −20◦ C. The product is black before distillation, colorless after, and yellow upon storage. It is important to place an acid vapor trap in the vacuum line in order to trap residual HBr, and to change the pump oil immediately following distillation, to prevent corrosion and seizure of the vacuum pump. The needle valve and Y-connector give better control over the distillation pressure. The constant boiling product (1,1-dimethylcyanoethyl bromoacetate; S.2) gave 97.3 g of clear, colorless liquid (88% yield). 1 H NMR (CDCl3 ): δ 3.75 (s, 2H), 2.86 (s, 2H), 1.53 (s, 6H). Electron impact mass spectrometry (EI-MS) generated molecular ions of 205 m/e and 207 m/e with isotopic abundances of 51% and 49%.
Synthesize phosphinylating reagent 13. Remove the 500-mL round-bottom flask with 24/40 joint from the oven. Cool under a stream of dry argon, place in the fume hood, and fit with a drying tube. 14. With a balance placed in the hood, weigh 20 g (75 mmol) bis(diisopropylamino)chlorophosphine into a plastic weighing boat and then quickly transfer the phosphine to the cool, dry 500-mL flask by temporarily removing the drying tube from the flask and adding the phosphine via a powder funnel. This reagent is moisture sensitive. It is therefore important to minimize its exposure to the atmosphere by working quickly.
15. Partially fill the flask with 300 mL anhydrous THF, add a stir bar, and place the flask on a magnetic stir plate. Cap the flask with the drying tube and secure to the fume hood’s lattice support or a support stand. 16. Stir the solution vigorously for 1 hr or until the phosphine has dissolved. The bis(diisopropylamino)chlorophosphine is often contaminated with small amounts of diisopropylamine hydrochloride and H-phosphonate hydrolysis products. These contaminants are less soluble in THF than the desired phosphine and can form an insoluble precipitate. To ensure effective conversion to the product, it is recommended that the bis(diisopropylamino)chlorophosphine be evaluated by 31 P NMR prior to use. The chlorophosphine should appear as a singlet in anhydrous CD3 CN with a chemical shift of 134.4 ppm. Peaks between 0 and 20 ppm represent various hydrolyzed or oxidized products. If peaks other than these appear in the spectrum, the reagent should be rejected or repurified (purity should be >90%). Purity of the chlorophosphine can be roughly estimated by integrating various peaks and using the results to determine the amount of impure phosphine to use in the reaction. The chlorophosphine can be synthesized and purified by recrystallization using the procedures described in Dellinger et al. (2003).
17. While the phosphine is being dissolved, remove the 1-liter three-neck round-bottom flask from the oven and cool under a stream of dry argon. Place it on a magnetic stir plate and secure it to the fume hood’s lattice support or a support stand. Fit a drying tube to the center joint and rubber septa to the side joints. 18. Weigh 7.2 g (110 mmol) granular zinc metal into a plastic weigh boat. Temporarily remove the drying tube from the flask, immediately transfer the zinc to the flask using a powder funnel, and replace the drying tube. Oligonucleotides with Phosphonoacetate and Thiophosphonoacetate Diesters
It is important to keep the zinc metal dry and under inert atmosphere, as the presence of large amounts of zinc oxide can inhibit the initiation of the Reformatsky coupling reaction.
19. Remove two 500-mL addition funnels, a 250-mL Erlenmeyer flask, and the 325-mm Friedrich’s condenser from the oven and cool to room temperature under a stream of
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dry argon. Make sure that the PTFE stopcocks are closed and tightened to prevent leaks. 20. Working quickly, decant the THF/phosphine solution into one of the addition funnels while leaving any insoluble material in the flask. Cap the funnel with a rubber septum and place on one of the side joints of the three-neck flask. Slurry the residual insoluble precipitate into 50 mL anhydrous THF and pour onto the zinc in the three-neck roundbottom flask. It is important that this step be performed quickly, as the phosphine solution is moisture sensitive.
21. Place the Erlenmeyer flask on the top-loading balance and tare. Carefully weigh 18.2 g (82.5 mmol) 1,1-dimethylcyanoethyl bromoacetate (S.2) into the dry Erlenmeyer flask. Add 150 mL anhydrous THF and swirl the flask to dissolve the reagent. 22. Pour this solution into the second addition funnel, cap the funnel with a rubber septum, and place it on the third neck of the flask. 23. Place the Friedrich’s condenser in the center neck of the three-neck flask and insert a drying tube on top of the condenser. Connect the condenser to a water source and drain using appropriate tubing. Add a PTFE-coated magnetic stir bar to the three-neck flask. 24. Add one third of the phosphine (100 mL) and bromoacetate (50 mL) solutions to the flask and heat this mixture using a hand-held heat gun focused on the zinc, with the magnetic stir plate turned off, until boiling begins. It is important to focus the heat from the heat gun onto the zinc metal.
25. Once the THF boils, remove the heat gun and observe the reaction mixture. Assessment that the Reformatsky reaction has initiated is quite obvious as it will be noticeably exothermic, and the cloudy colorless mixture will become clear and slightly yellow. If the reaction does not initiate, activate the heat gun and locally heat the zinc until the THF once again begins to boil. Once the reaction has initiated, the solution will reflux without the use of a heat gun.
26. Continue the reaction at reflux by slowly adding the remainder of the phosphine and bromoacetate solutions. If the reaction becomes too vigorous, engage the magnetic stir plate, as stirring usually controls the rate. If it is still too vigorous, cool the flask using an ice bath. 27. Once addition is complete, keep the solution at reflux (which is lost as the reaction slows) for 30 min by using a heating mantle. Cool to room temperature. 28. Analyze by 31 P NMR for completeness by dissolving a few drops of the reaction mixture in an NMR tube containing anhydrous acetonitrile. First, lock the NMR instrument and shim the magnet on a phosphoric acid/CD3 CN standard. Then remove the standard, turn off the lock, insert the NMR tube with the crude sample into the instrument, and acquire experimental results. 31
P NMR samples of these crude mixtures are typically evaluated in anhydrous, nondeuterated acetonitrile with the deuterium lock turned off due to the difficulty and expense of obtaining absolutely anhydrous deuterated solvents. Commercial “anhydrous” deuterated solvents falsely indicate that a significant amount of the reaction products are hydrolyzed. With the same anhydrous, nondeuterated acetonitrile commonly used for bulk reactions, a more accurate evaluation of hydrolysis products has been obtained. The chlorophosphine (δ P = 135 ppm) should be completely consumed and converted to a single product (S.3; δ P = 48 ppm).
Synthesis of Modified Oligonucleotides and Conjugates
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29. Cool reaction mixture and decant into a 1-liter recovery flask, leaving behind the unreacted metal. Remove THF using a rotary evaporator to yield a yellow/brown viscous oil.
Isolate phosphinylating reagent 30. To isolate the product by trituration, add 250 mL anhydrous reagent-grade hexanes, place a rubber septum in the flask’s 24/40 joint, and shake vigorously to dissolve the phosphine. Allow the oil and hexanes to settle and decant the hexane layer into a 2-liter recovery flask. Repeat for a total of three successive extractions. The oil will be converted to a solid.
31. Dissolve the resulting solid in 150 mL anhydrous acetonitrile, place in a 500-mL separatory funnel, and extract twice with 150-mL portions of anhydrous hexanes. 32. Analyze the acetonitrile solution by 48 ppm) and discard.
31 P
NMR for absence of the product (δP =
33. Combine all hexane fractions in the 2-liter recovery flask and concentrate the product on a rotary evaporator to a slightly yellow oil. Redissolve in 300 mL anhydrous hexanes and place the flask in a −20◦ C freezer overnight. 34. Decant the hexane solution into a clean, dry, 500-mL round-bottom flask and remove solvent using a rotary evaporator to give 16.4 g (88% yield) of a slightly yellow oil. Store up to 6 months at −20◦ C. The product, 1,1-dimethyl-2-cyanoethyl [bis(N,N-diisopropylamino)phospinyl]acetate (S.3), can be further purified by recrystallization in anhydrous pentane. 1 H NMR (CDCl3 ): δ 3.48 (m, 4H), 2.99 (s, 2H), 2.79 (d, J = 3.12 Hz, 2H), 1.30 (m, 24H). EI-MS generated a molecular ion of 371 m/e with fragmentation loss of CH2 COOC(CH3 )2 CH2 CN at 231 m/e. 31 P NMR (CD3 CN): δ 48.1 ppm. The authors typically only recrystalize with pentane to achieve good data from analysis by 1 H NMR.
Synthesize (N,N-diisopropylamino)phosphinyl acetate monomers 35. Remove a 500-mL round-bottom flask from the oven and cool under a stream of dry argon. Place the flask on a magnetic stir plate in the fume hood, add a stir bar, and attach a ground-glass joint capped with a drying tube. Fasten to the fume hood’s lattice support or a support stand. 36. Weigh 10.0 g DMTr-protected 2 -deoxynucleoside (S.4) into a plastic weighing boat. To avoid absorption of moisture, temporarily remove the drying tube from the flask, quickly transfer the nucleoside using a powder funnel, and return the drying tube to the flask. Add 250 mL anhydrous dichloromethane and begin stirring.
Oligonucleotides with Phosphonoacetate and Thiophosphonoacetate Diesters
Occasionally the protected 2 -deoxynucleosides from commercial suppliers contain a significant amount of absorbed water. Any water in the solution will preferentially react with the phosphinylating agent and reduce the final yield of product. It is therefore prudent to test the protected 2 -deoxynucleoside/dichloromethane solution for the amount of water present using a Karl-Fisher titration prior to performing the phosphinylation reaction. (Once the shipment has been shown to be anhydrous, it can be assumed subsequent solutions made from the same batch will be anhydrous as long as care has been taken in handling and storage.) The authors use an Aquastar V-200 Karl-Fischer titrator (VWR ). A small amount of water in the protected 2 -deoxynucleoside can be neutralized by using a molar equivalent excess of the phosphinylating agent as an internal desiccant. Larger amounts of water require that the 2 -deoxynucleoside solution be evaporated to dryness on a rotary evaporator, redissolved in anhydrous pyridine, and evaporated to dryness again. The excess water is thus removed as a pyridine/water azeotrope. This procedure should be repeated until the water content of the pyridine solution is similar to the anhydrous solvent alone. It is subsequently important to completely remove all the pyridine,
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Table 4.24.1 Reactants Used to Prepare 2 -Deoxynucleoside-3 -O-(N,Ndiisopropylamino)phosphinyl Acetates
5 -DMTr-dABz
5 -DMTr-dC
5
Ac
-DMTr-dGi-Bu
5 -DMTr-T
Protected 2 -deoxynucleoside
Phosphinylating reagent
0.45 M tetrazole
10 g
6.8 g
27.2 mL
10 g
7.8 g
31.1 mL
10 g
7.0 g
27.9 mL
10 g
8.2 g
32.7 mL
as it will act as a base and neutralize the tetrazole acid catalyst. This is accomplished by using a similar azeotropic evaporation process with toluene. Once the protected 2 deoxynucleoside is free of pyridine, it is redissolved in anhydrous dichloromethane and the synthesis continued.
37. Place a top-loading balance in the hood and weigh the appropriate amount of phosphinylating reagent (S.3) into a plastic weighing boat (Table 4.24.1). Temporarily remove the drying tube, quickly transfer the phosphine to the 500-mL flask using a powder funnel, and return the drying tube to the flask. Continue stirring to dissolve completely (∼10 to 30 min). The phosphine is moisture sensitive, so it is important to minimize its exposure to the atmosphere.
38. Add the volume of 0.45 M tetrazole in anhydrous acetonitrile shown in Table 4.24.1 using a dry 50-mL glass pipet. Stir 24 hr at room temperature. The volume given provides 0.8 molar equivalents tetrazole to the reaction.
39. Spot the reaction mixture on a glass-backed silica-gel TLC plate with fluorescent indicator, alongside a dichloromethane solution of the appropriate starting material. Air dry the spots. 40. Carefully place the TLC plate in a developing tank pre-equilibrated with reagentgrade ethyl acetate and allow the solvent elution front to reach the top of the plate. Air dry the plate, place in a UV viewing cabinet, and illuminate using shortwave irradiation. The reaction is determined complete by spot-to-spot conversion to a faster eluting product. For additional details on TLC, see APPENDIX 3D.
41. Confirm conversion to product by loss of the 31 P NMR signal corresponding to S.3 (δ = 48.1 ppm). 42. If conversion is incomplete by these assays, allow the reaction mixture to stir another 24 hr. 43. Quench the reaction by adding 0.8 molar equiv anhydrous DIPEA and stirring 5 min. 44. Pour the solution into a 1-liter recovery flask and concentrate to a viscous oil using a rotary evaporator to remove solvent. Proceed to purification immediately.
Purify product by column chromatography 45. For each preparation, place ∼700 mL dry silica gel 60 (230 to 400 mesh) and 700 mL solvent (Table 4.24.2) in a 2-liter Erlenmeyer flask and swirl to equilibrate. If needed, use additional solvent to assure that the silica gel is completely solvated and easy to pour. Pour ∼16 cm solvated silica gel into a flash chromatography column and use solvent to wash any silica from the sides of the column. For additional details on column chromatography, see APPENDIX 3E.
Synthesis of Modified Oligonucleotides and Conjugates
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Table 4.24.2 Column Elution Solvents for Various Synthons
2 -Deoxynucleoside base
% Ethyl acetate (v/v)
% Hexanes (v/v)
Adenine
100
0
Cytosine
50
50
Guanine
100
0
Thymine
100
0
46. Assemble the column in the fume hood by tightly clamping it to the fume hood’s lattice support or a support stand. Attach the flow controller. Push solvent through the column at a flow rate of ∼300 mL/min using air or nitrogen to compress the silica gel bed. Remove the pressure and flow controller when the solvent level reaches the compressed silica. Sprinkle sea sand on the silica gel bed to a depth of ∼2 cm. 47. Carefully pour an additional 500 mL solvent into the column without disturbing the compressed silica gel bed. Reattach the flow controller and push the liquid through the column at the same rate until it reaches the silica gel bed. Because most silica gels are acidic, it is typical to add a small percentage of triethylamine to the eluting solvent when purifying protected 2 -deoxynucleoside phosphoramidites. This prevents loss of the DMTr protecting group and acid-catalyzed hydrolysis of the phosphoramidite. However, the presence of triethylamine in the eluting solvent has a deleterious effect on the stability of the 2 -deoxynucleoside-3 -O-(N,N-diisopropylamino)phosphinyl acetates . As a result, losses in recovered yield are greater in the presence of triethylamine than when this base is not used. Therefore, addition of triethylamine to these solvents is not recommended. Typically, there is a considerable amount of diisopropylammonium tetrazolide in the crude 2 -deoxynucleoside-3 -O-(N,N-diisopropylamino)phosphinyl acetate products to help neutralize excess acid on the silica gel. However, detritylation has still been observed infrequently during column chromatography. Detritylation on the column is easily detectable as a bright orange band on the silica column. To eliminate this detritylation, a small amount of TEA may be used to neutralize excess acid on silica gel during column packing and equilibration, but excess amine should be thoroughly flushed from the column prior to adding the crude material to the top of the column. Under either of these conditions, it is important to run the column at a rapid flow rate and not leave the product in contact with the silica gel for prolonged periods of time. The DMTrprotected 2 -deoxythymidine-3 -O-(N,N-diisopropylamino)phosphinyl acetates seem to be most susceptible to degradation during prolonged exposure to silica gel. When purifying these synthons, it is important to elute the column rapidly but carefully and not pause during purification.
48. Dissolve the crude product (step 43) in a minimum volume of ethyl acetate and add to the column by slowly dropping the mixture onto the sand layer using a Pasteur pipet. Use a small volume of solvent (50 mL) to wash the flask and add this solution dropwise to the sand layer. 49. Push the reaction mixture and solvent onto the silica gel by briefly pressurizing the column until the solvent level once again reaches the top of the compressed bed. Carefully add an additional 50 mL solvent to the sand layer and once again push it onto the silica gel by briefly pressurizing the column until the liquid level has reached the compressed bed.
Oligonucleotides with Phosphonoacetate and Thiophosphonoacetate Diesters
50. Carefully fill the column with solvent without disturbing the compressed silica gel bed. Attach the flow controller and elute the reaction product at ∼300 mL/min. Collect 50-mL fractions. 51. Check for UV-absorbing material by spotting a small amount of eluted solvent onto a glass-backed TLC plate containing a fluorescent indicator. Air dry and view the plate
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in the UV viewing cabinet using shortwave irradiation. Keep samples containing UV-absorbing material and discard those without. 52. For detailed analysis, spot TLC plates with column fractions that contain UV-absorbing material along with a dichloromethane solution of the appropriate starting material and the crude reaction mixture. Air dry the spots, place the plate in a developing tank pre-equilibrated with ethyl acetate, and elute until the solvent front reaches the top of the plate. Air dry and view in the UV viewing cabinet using shortwave irradiation. 53. Identify column fractions containing pure material suspected of being the product. Combine in a recovery flask and remove solvent on a rotary evaporator. The resulting products are usually glassy or foamed solids. The authors frequently elect to convert them to granular powders by precipitation into anhydrous hexanes (step 54), because powders are easier to manipulate for solid-phase DNA synthesis.
Work up final product 54. For each preparation, place 1 liter anhydrous hexanes in a 2-liter recovery flask. Redissolve the purified solid in a minimal amount of anhydrous dichloromethane and add dropwise to the hexanes using a Pasteur pipet. Add the DCM/product mixture a few drops at a time, rapidly swirling the solution. A white solid, corresponding to the product, precipitates from the hexanes.
55. Once addition is complete, isolate the solid by filtration using a 350-mL mediumporosity, fritted glass Buchner funnel fitted with a 24/40 vacuum adapter. During filtration, keep the solid submersed in solvent to prevent the granular precipitate from converting to a gum. 56. When filtration is complete, immediately place the funnel in a vacuum desiccator and attach to a two-stage vacuum pump or high-vacuum line. Make sure that the vacuum line or vacuum pump is “trapped” so that there is no chance that oil will back up into the vacuum desiccator. Allow the precipitate to be evacuated in the vacuum desiccator for at least 12 hr in order to remove all residual solvent. Rapid evacuation of solvent from the powder can often lead to an “explosion” of the product such that it covers the inside walls of the desiccator. This can be prevented by slowly evacuating the desiccator in stages. Additionally, covering the funnel with aluminum foil containing a few small holes can help prevent this unfortunate loss of product.
57. Once the precipitate has been dried under vacuum, close the stopcock to isolate the vacuum desiccator, remove the high-vacuum line, and bleed the desiccator slowly with dry argon or nitrogen. The resulting white powder is easily partitioned into DNA synthesizer reagent vials.
58. Identify and analyze each product for purity by 31 P NMR and FAB-MS. Store up to 6 months at −20◦ C. 1,1-Dimethyl-2-cyanoethyl 5 -O-(4,4 -dimethoxytrityl)-2 -deoxythymidine-3 -O-(N,Ndiisopropylamino)phosphinyl acetate (S.5a) is isolated with an average yield of 84%. 31 P NMR (CD3 CN): δ 120.3, 120.8 ppm. FAB-HRMS: calcd. for C44 H55 N4 O9 P (M+H)+ 815.3785, found 815.3775. 1,1-Dimethyl-2-cyanoethyl 5 -O-(4,4 -dimethoxytrityl)-N4 -acetyl-2 -deoxycytidine-3 -O(N,N-diisopropylamino)phosphinyl acetate (S.5b) is isolated with an average yield of 78%. 31 P NMR (CD3 CN): δ 121.3, 121.8 ppm. FAB-HRMS: calcd. for C45 H56 N5 O9 P (M+H)+ 842.3894, found 842.3914. 1,1-Dimethyl-2-cyanoethyl 5 -O-(4,4 -dimethoxytrityl)-N6 -benzoyl-2 -deoxyadenosine3 -O-(N,N-diisopropylamino)phosphinyl acetate (S.5c) is isolated with an average yield
Synthesis of Modified Oligonucleotides and Conjugates
4.24.11 Current Protocols in Nucleic Acid Chemistry
Supplement 18
of 75%. 31 P NMR (CD3 CN): δ 120.8, 121.6 ppm. FAB-HRMS: calcd. for C51 H58 N7 O8 P (M+H)+ 928.4163, found 928.4159. 1,1-Dimethyl-2-cyanoethyl 5-O-(4,4-dimethoxytrityl)-N2 -isobutyryl-2-deoxyguanosine3 -O-(N,N-diisopropylamino)phosphinyl acetate (S.5d) is isolated with an average yield of 89%. 31 P NMR (CD3 CN): δ 121.7, 122.0 ppm. FAB-HRMS: calcd. for C48 H60 N7 O9 P(M+H)+ 910.4268, found 910.4235.
BASIC PROTOCOL 2
AUTOMATED SYNTHESIS OF PHOSPHONOACETATE AND THIOPHOSPHONOACETATE DNA Phosphonoacetate DNA Synthesis The chemical synthesis of PACE ODNs (Fig. 4.24.4) can be accomplished manually, but the use of an automated DNA synthesizer is preferred. This protocol describes the synthesis as performed on an ABI model 394 automated DNA synthesizer. The chemistry is completed on a solid support with standard controlled-pore glass (CPG) DNA synthesis columns with long-chain alkylamino (LCAA) linker (1 µmol dA, Ac-dC, dG, dT; 500-Å glass pore size; Glen Research). When using a dC-containing CPG column, it is important that the dC be protected with an acetyl rather than benzoyl group. The use of the acetyl protecting group on dC allows for the rapid deprotection of the ODNs using methylamine at 55◦ C for 15 min without methylation of the N4 position of the cytosine residues. The use of the more typical benzoyl-protected dC monomers and CPG columns results in a small but detectable amount of methylation of the N4 position during ODN deprotection using these rapid conditions. The use of the typical benzoyl-protected dA and isobutyrylprotected dG is compatible with rapid ODN deprotection using methylamine and does not result in detectable methylation of these heterobases. The recommended synthesis cycle is adapted from a standard 1-µmol 2-cyanoethylphosphoramidite DNA cycle (Caruthers et al., 1987). The coupling wait-time is increased to 1998 sec by using two successive 999-sec wait steps. The oxidation step is accomplished prior to the capping step; the oxidation wait time is 180 sec. Table 4.24.3 summarizes the synthesis cycle as applied to the 394 automated synthesizer. A general description of automated DNA synthesis can be found in APPENDIX 3C. The monomers used for PACE ODN synthesis are the 1,1-dimethyl-2-cyanoethyl 5 O-(4,4 -dimethoxytrityl)-2 -deoxynucleoside-3 -O-(N,N-diisopropylamino)phosphinyl acetates described above (see Basic Protocol 1). They are also commercially available from MetaSense Technologies (see http://www.Metasensetechnologies.com for ordering and helpful information). It is important to dry the protected 2 -deoxynucleoside3 -O-(N,N-diisopropylamino)phosphinyl acetates overnight to remove any possible moisture before use (see Support Protocol). Indeed, prior to dissolving the synthons, researchers should familiarize themselves with techniques used for handling air-sensitive or moisture-sensitive reagents. The most common failure modes for DNA synthesis result from moisture contamination of amidites or activators. A good treatise on this subject, Aldrich Technical Bulletin: AL-134 Handling AirSensitive Reagents, can be obtained from Aldrich Chemical in PDF format at http://www.sigmaaldrich.com/aldrich/bulletin/al techbull al134.pdf.
Oligonucleotides with Phosphonoacetate and Thiophosphonoacetate Diesters
Freshly sublimed tetrazole dissolved in anhydrous acetonitrile at a concentration of 0.45 M is used as an activator. A prepared solution of 0.45 M tetrazole in acetonitrile can be purchased from Glen Research. Trichloroacetic acid (3% w/v in anhydrous dichloromethane; Glen Research) is used to deprotect the 5 -DMTr group prior to each round of coupling. Capping is accomplished using a two-part capping solution modified from Hogrefe et al. (1993). Cap A consists of 10% acetic anhydride in anhydrous tetrahydrofuran (Glen Research), and Cap B consists of 0.625% (w/v)
4.24.12 Supplement 18
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Figure 4.24.4 Synthesis scheme for preparing phosphonoacetate and thiophosphonoacetate oligodeoxyribonucleotides. Circled P is the LCAA-CPG support (long-chain alkylamino controlled-pore glass hemisuccinate). Abbreviations: B, thymin1-yl, 4-N-acetylcytosin-1-yl, 6-N-benzoyladenin-9-yl, or 2-N-isobutyrylguanin-9-yl; BDT, 3H-1,2-benzodithiol-3-one 1,1dioxide; CSO, (1S)-(+)-(10-camphorsulfonyl)oxazaridine; DMAP, 4-dimethylaminopyridine; DMTr, 4,4 -dimethoxytrityl; Py, pyridine; TCA, trichloroacetic acid. Table 4.24.3 Synthesis Cycle for Phosphonoacetate or Thiophosphonoacetate DNA
Step
Delivery time (sec)
Wait time (sec)
Port
1. Dichloromethane
35
—
19
2. Detritylation
85
—
14
3. Acetonitrile
20
—
18
2.5 × 2
999 × 2
5-8, 9
10
—
18
8
180
15
8
60
10
7. Acetonitrile
40
—
18
8. Capping
10
10
11 and 12
9. Acetonitrile
30
—
18
4. Coupling
a
5. Acetonitrile 6. Oxidation or b
sulfurization
a Coupling involves simultaneous delivery of an appropriately protected 2 -deoxynucleoside-3 -O-(N,N-
diisopropylamino)phosphinyl acetate (ports 5 to 8) and tetrazole (port 9). b Synthesis of phosphonoacetate DNA requires placement of CSO at port 15, whereas synthesis of thio-
phosphonoacetate DNA requires BDT at port 10. The appropriate wait time is then programmed into the cycle.
4-N,N-dimethylaminopyridine (DMAP, Aldrich) in anhydrous pyridine (Aldrich). Oxidation of the nascent internucleotide phosphonite to the phosphonate is carried out with 0.1 M (1S)-(+)(10-camphorsulfonyl)oxaziridine (Aldrich) in anhydrous acetonitrile (1 g CSO in 43.6 mL CH3 CN). The acetonitrile used should be anhydrous synthesis grade, and dichloromethane should be certified ACS grade (both available from Fisher Biotech).
Thiophosphonoacetate DNA Synthesis With the exception of the sulfurization step, the synthesis of S-PACE ODNs is analogous to the procedure used for PACE DNA. Because the same synthons are used to prepare both
Synthesis of Modified Oligonucleotides and Conjugates
4.24.13 Current Protocols in Nucleic Acid Chemistry
Supplement 18
PACE and S-PACE ODNs, the synthesis cycle in Table 4.24.3 can be followed, except that 3H-1,2-benzodithiol-3-one-1,1-dioxide (BDT) is used as a sulfurization reagent in place of the CSO used for oxidation in PACE DNA synthesis. The sulfurization wait time is 60 sec, and the sulfurization solution is 0.05 M BDT (Glen Research) in anhydrous acetonitrile (1 g BDT in 100 mL CH3 CN). SUPPORT PROTOCOL
DRYING OF PHOSPHONOACETATE AND THIOPHOSPHONOACETATE MONOMERS Steps are provided to describe the drying and preparation of the starting monomers prior to ODN synthesis.
Materials 1,1-Dimethyl-2-cyanoethyl 5 -O-(4,4 -dimethoxytrityl)-2 -deoxynucleoside-3 -O(N,N-diisopropylamino)phosphinyl acetates (S.5a-d; see Basic Protocol 1; also available from MetaSense Technologies) Argon or nitrogen, anhydrous Anhydrous acetonitrile, synthesis grade (Fisher Biotech) Amber serum vials with rubber septa 18-G × 1-in. needle Vacuum desiccator Two-stage vacuum pump or high vacuum line, with trap Syringe, dry 1. Weigh the appropriate amount of each synthon into separate amber serum vials and place a rubber septum on each. Press an 18-G × 1-in. needle through each septum until it extends about half-way. Make sure that the needles do not contact any powder. 2. Place the vials in a vacuum desiccator attached to a two-stage vacuum pump or a high vacuum line, with a trap, and evacuate for at least 12 hr prior to use. 3. Close the stopcock to isolate the vacuum desiccator, remove the high vacuum line, and bleed the desiccator slowly with dry argon or nitrogen. Open the desiccator and quickly remove the needles from the rubber septa. 4. Dissolve dried synthons in synthesis-grade anhydrous acetonitrile at a concentration of 0.1 M by filling a dry syringe with the required amount of solvent (Table 4.24.4) and then transferring the solvent to the amber serum vial under dry nitrogen or argon. Store up to 6 months at −20◦ C. For the synthesis of an average 20-mer having five of each base (dC, dG, dT, dA) and on a 1-µmol CPG column, less than 250 mg of each synthon is required.
Table 4.24.4 Acetonitrile Solvent Requirements for 2 -Deoxynucleoside-3 -O(N,N-diisopropylamino)phosphinyl Acetates
Volume acetonitrile
Oligonucleotides with Phosphonoacetate and Thiophosphonoacetate Diesters
2 -Deoxynucleoside base
250 mg
1g
Adenine
2.7 mL
11.0 mL
Cytosine
3.0 mL
11.9 mL
Guanine
2.8 mL
11.0 mL
Thymine
3.1 mL
12.3 mL
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Current Protocols in Nucleic Acid Chemistry
AUTOMATED SYNTHESIS OF MODIFIED DNA CHIMERAS The protocol outlined in this section can be used to synthesize modified DNA having either PACE or S-PACE internucleotide linkages combined with phosphate and thiophosphate backbones (Fig. 4.24.5). Many oligomers having these four linkages in various combinations have been synthesized, including those presented in Table 4.24.5 (also see BioSpring GmbH, http://www.biospring.de).
BASIC PROTOCOL 3
Preparation for Synthesis of DNA Chimeras The reagents and preparations for synthesis of chimeras having either PACE or S-PACE internucleotide linkages in combination with phosphate or thiophosphate backbones are analogous to those outlined in Basic Protocol 2. In order to generate S-PACE and thiophosphate linkages, the sulfurization solution used is BDT in anhydrous acetonitrile. The concentration of BDT for sulfurization remains the same for synthesizing thiophosphate and S-PACE internucleotide linkages. Oxidation to PACE and phosphate backbones is carried out with CSO; however, the concentration of CSO required to oxidize a phosphite triester to a phosphotriester (producing a normal phosphate backbone) is significantly greater than the concentration required to oxidize the phosphonite diester to the phosphonate diester (producing the PACE backbone). Effective oxidization of phosphite triesters to phosphotriesters requires a 0.5 M solution of CSO and exposure for at least 3 min. This higher concentration of CSO is used to oxidize both phosphite and phosphonite internucleotide linkages during the preparation of chimeras. The use of CSO for the nonaqueous oxidation of phosphite triesters is described in a Glen Report (Glen Research, 1996), which can be downloaded as a PDF file from the Glen Research Web site (http://www.glenres.com). Synthons used to prepare PACE and S-PACE linkages (see Basic Protocol 1 or MetaSense Technologies) are dried as described in the Support Protocol, dissolved in anhydrous acetonitrile (see Table 4.24.4 for appropriate concentrations), and then transferred to ports 5 to 8 on the automated DNA synthesizer. For preparing phosphate and thiophosphate internucleotide linkages, standard base-protected synthons, 5 -O-(4,4 -dimethoxytrityl)-3 -O-[(N,N-diisopropylamino)(2-cyanoethoxy)phosphinyl]2 -deoxyribonucleoside monomers, are used according to published procedures (Caruthers et al., 1987). They are available from Glen Research (dT-CE, Ac-dC-CE, dG-CE, and dA-CE phosphoramidites). These synthons are dissolved as 0.1 M solutions in anhydrous acetonitrile and transferred to ports 1 to 4 on the DNA synthesizer. The capping solution is described in Basic Protocol 2.
Table 4.24.5 MALDI-TOF Mass Spectrometry of ODNsa
Abbreviation
Structure
Calculated mass
Observed mass
PACE
d(CaTaCaAaAaGaTaGaGaGaCaTaGaGaTaGaAaC)
6270.2
6271.3
S-PACE
d(CsTsCsAsAsGsTsGsGsGsCsTsGsGsTsGsAsC)
6542.1
6540.5
PACE EO
d(CaTCaAAaGTaGGaGCaTGaGTaGAaC)
5933.8
5933.1
S-PACE EO
d(CsT-CsA-AsG-TsG-GsG-CsT-GsG-TsG-AsC)
6205.8
6208.2
PACE Cap
d(CaTaCaAaAaGaT-G-G-G-C-TaGaGaTaGaAaC)
6139.9
6137.2
S-PACE Cap
d(CsTsCsAsA-G-T-G-G-G-C-T-G-GsTsGsAsC)
6163.8
6165.5
a Nomenclature: a, phosphonoacetate; s, thiophosphonoacetate; -, phosphorothioate; the absence of a symbol between
nucleoside letters corresponds to a normal phosphate linkage (e.g., CT).
Synthesis of Modified Oligonucleotides and Conjugates
4.24.15 Current Protocols in Nucleic Acid Chemistry
Supplement 18
Figure 4.24.5 Synthesis scheme for preparing oligodeoxyribonucleotide chimeras having phosphonoacetate or thiophosphonoacetate internucleotide linkages combined with phosphodiester or phosphorothioate backbones. See Figure 4.24.4 for abbreviations.
Synthesis of Oligodeoxyribonucleotide Chimeras A general synthesis plan for various chimeras is outlined in Table 4.24.6. This table includes two series of synthons—i.e., the standard 2 -deoxynucleoside-3 phosphoramidites and the 2 -deoxynucleoside-3 -O-(N,N-diisopropylamino)phosphinyl acetates—and two choices for the P(III) to P(V) oxidation of the internucleotide linkage—i.e., CSO and BDT. Each cycle, which outlines the steps for synthesizing one internucleotide linkage, uses only one synthon and oxidation condition. In some cases, chimeras having all four bases can be synthesized using one cycle. For example, phosphodiester/PACE or phosphorothioate/S-PACE chimeras are prepared using only one oxidant (CSO or BDT, respectively). Chimeric syntheses of phosphorothioate/PACE or phosphodiester/S-PACE ODNs require two separate cycles: one for the phosphorothioate and S-PACE linkages (using the sulfurizing reagent BDT) and another to generate phosphodiester and PACE internucleotide backbones (using the oxidant CSO). Oligonucleotides with Phosphonoacetate and Thiophosphonoacetate Diesters
A chimera requiring two cycles is illustrated by the PACE Cap ODN shown in Table 4.24.5. This oligomer has phosphorothioate linkages flanked by PACE at the 3 and 5 ends. In this case, a heptamer having a PACE backbone is first synthesized using appropriate 2 -deoxynucleoside-3 -O-(N,N-diisopropylamino)phosphinyl acetates as synthons and CSO as oxidant. At this point, BDT as a sulfurizing agent is substituted for CSO. This
4.24.16 Supplement 18
Current Protocols in Nucleic Acid Chemistry
Table 4.24.6 Synthesis Cycle for ODN Chimeras
Step
Delivery time (sec)
Wait time (sec)
Port
1. Dichloromethane
35
—
19
2. Detritylation
85
—
14
3. Acetonitrile
20
—
18
standard phosphoramidites
2.5 × 2
25
1-4, 9
phosphinoamidites
2.5 × 2
999 × 2
5-8, 9
10
—
18
8
180
15
8
60
10
7. Acetonitrile
40
—
18
8. Capping
10
10
11 and 12
9. Acetonitrile
30
—
18
4. Couplinga :
5. Acetonitrile 6. Oxidation or b
sulfurization
a For each cycle, coupling involves simultaneous delivery of a standard protected 2 -deoxynucleoside-3 -
phosphoramidite (ports 1 to 4) and tetrazole (port 9) or an appropriately protected 2 -deoxynucleoside-3 -O-(N,Ndiisopropylamino)phosphinyl acetate (ports 5 to 8) and tetrazole (port 9). b The choice of oxidation (CSO) or sulfurization (BDT) depends upon the synthesis plan.
is accomplished by reprogramming the cycle to use port 10 containing the BDT reagent in place of CSO at port 15. Synthesis then continues using appropriately protected standard 2 -deoxynucleoside-3 -phosphoramidites in order to prepare a 12-mer having five phosphorothioate and six PACE linkages. Once again the cycle is changed so that CSO (port 15) replaces BDT (port 10) in the synthesis program. The final PACE internucleotide linkages are added using 2 -deoxynucleoside-3 -O-(N,N-diisopropylamino)phosphinyl acetates to generate the PACE Cap ODN. Automated synthesis for certain types of chimeras that require two oxidants is therefore limited by the programming capabilities of the ABI 394 instrument. For preparing ODNs such as PACE Cap, this reprogramming problem is minor, as it must be done only twice.
DEPROTECTION, PURIFICATION, AND CHARACTERIZATION OF MODIFIED DNA
BASIC PROTOCOL 4
The following steps are used for PACE, S-PACE, and chimeric DNA. After partial deprotection and cleavage from the solid support, the ODNs are easily purified by reversedphase HPLC (RP-HPLC) using the 5 -DMTr protecting group for hydrophobic affinity (trityl-on purification). After detritylation and further RP-HPLC purification, they are analyzed by ion-exchange HPLC to determine yield and by matrix-assisted laser desorption/ionization time-of-flight mass spectometry (MALDI-TOF-MS) to obtain ODN mass information.
Materials DNA synthesis column containing oligodeoxyribonucleotides (ODN) linked to controlled-pore glass (CPG) Acetonitrile, anhydrous Argon or nitrogen, anhydrous 1.5% (v/v) 1,8-diazabicyclo-[5.4.0]undec-7-ene (DBU; Aldrich) in anhydrous acetonitrile 40% methylamine, aqueous (Aldrich)
Synthesis of Modified Oligonucleotides and Conjugates
4.24.17 Current Protocols in Nucleic Acid Chemistry
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RP-HPLC mobile phases: A: 100 mM triethylammonium acetate, pH 8.0 B: acetonitrile 10 mM Tris·Cl, pH 8.0 (APPENDIX 2A) 80% acetic acid 50 mM triethylammonium acetate, pH 8 TE buffer, pH 8.0 (APPENDIX 2A) IEX-HPLC mobile phases: A: 10 mM NaOH/80 mM NaBr B: 10 mM NaOH/1.5 M NaBr 2,4,6-Trihydroxyacetophenone monohydrate (THAP) matrix Ammonium citrate 1:1 (v/v) acetonitrile/H2 O 10-mL Luer-tip syringe Male Luer-to-tubing connector (Aldrich) Conical vial: 3-mL Reacti-Vial sealed with Teflon-bonded silicon Tuf-Bond discs (Pierce), preferred 55◦ C heating block (e.g., Reacti-Block Aluminum Heat-Block, Pierce) or oven Speedvac evaporator (Savant) RP-HPLC apparatus with 25-cm × 9.4-mm-i.d. Zorbax 300SB-C18 column (Agilent Technologies) Ion-exchange (IEX)-HPLC apparatus with 1-mL RESOURCE Q column (Amersham Pharmacia Biotech) 100-well plate, gold plated Voyager-DE STR Biospectrometry Workstation mass analyzer (PerSeptive Biosystems) Recover CPG-bound ODN 1. Remove the DNA synthesis column containing the ODN linked to CPG from the synthesizer. Wash with 10 mL anhydrous acetonitrile using a 10-mL Luer-tip syringe to push the solvent through the column. 2. Flush with a low-pressure stream of dry argon or nitrogen using a male Luer-totubing connector fitted onto the inert gas line and placing the Luer-tip snugly into the DNA synthesis column. Blow a mild, low-pressure stream through the column until the CPG flows freely as the column is rotated. 3. Open the column and pour the contents into a conical glass vial (preferably a 3-mL Reacti-Vial).
Remove dimethylcyanoethyl group 4. Place 1 mL freshly prepared 1.5% DBU solution into the vial containing CPG and tighten the cap containing the Teflon-bonded silicon disc onto the vial. Vortex to stir the contents and incubate 60 min at room temperature. The vial can be vortexed occasionally during this deprotection reaction.
5. Vortex again and allow the CPG to settle into the conical portion of the vial. Remove the DBU solution using a clean Pasteur pipet and discard. Repeat three times using 1 mL acetonitrile each time to ensure complete removal of the DBU.
Oligonucleotides with Phosphonoacetate and Thiophosphonoacetate Diesters
Cleave ODN and remove exocyclic amine–protecting group 6. Add 1 mL aqueous 40% methylamine to the CPG and place the Teflon cap snugly on the vial. Heat the solution 15 min at 55◦ C by placing the vial in a preheated heating block or oven. 7. Remove the vial from the heat source, vortex, and place in an ice bucket.
4.24.18 Supplement 18
Current Protocols in Nucleic Acid Chemistry
8. Once cooled, remove the top of the vial and transfer the methylamine solution to a microcentrifuge tube using a glass Pasteur pipet. Be careful to allow the CPG to settle to the bottom of the glass vial before transferring the methylamine solution. The conical shape of the Reacti-Vial aids this process and prevents transfer of CPG. The residual CPG can be washed with an additional volume of cold methylamine solution to optimize recovery. Be careful, however, not to over-fill the microcentrifuge tube.
9. Evaporate solution to dryness, preferably in a centrifugal evaporator/concentrator (e.g., SpeedVac) without heating. Store dry mixture of oligodeoxyribonucleotide products in the sealed tube up to 6 months at −20◦ C.
Purify ODN by RP-HPLC and remove dimethoxytrityl group 10. Dissolve the crude oligodeoxyribonucleotide mixture in 0.5 mL water. Perform tritylon purification by RP-HPLC on a 25-cm Zorbax 300SB-C18 column. Elute with RP-HPLC mobile phases using the following gradient at a flow rate of 1.2 mL/min: Duration 0-2 min 2-27 min 27-52 min
Mobile phase B 8% 8% to 20% 20% to 80%.
Collect fractions containing the full-length product. 11. Concentrate under vacuum (e.g., SpeedVac) in a microcentrifuge tube and redissolve in 100 µL of 10 mM Tris·Cl, pH 8. It is important to use a buffered solution at pH 7 to 8 for dissolving these ODNs. At acidic pH, PACE and S-PACE ODNs are less soluble and can stick to the tube, which reduces recovery.
12. Remove the trityl group by treatment for 1 hr with 1 mL of 80% acetic acid. 13. Concentrate to dryness (e.g., in a SpeedVac) and dissolve in 0.5 mL of 50 mM triethylammonium acetate, pH 8. 14. Repeat preparative HPLC with the 25-cm × 9.4-mm-i.d. Zorbax 300SB-C18 column to isolate the fully deprotected ODNs. Use the same gradient conditions as for tritylon isolation of ODNs (step 10). Concentrate the resulting products under vacuum (e.g., Speedvac) and store up to 6 months at −20◦ C in ∼100 µl TE buffer, pH 8.0.
Characterize ODNs by IEX-HPLC 15. To determine synthetic yield of full-length product, analyze by ion-exchange HPLC using a 1-mL RESOURCE Q column. Elute with IEX-HPLC mobile phases using a gradient of 0% to 100% IEX-HPLC mobile phase B over 45 min at a flow rate of 1.5 mL/min. Analyze ODNs by mass spectrometry 16. Dissolve the modified ODN in water at a concentration of 100 µM. 17. Add 45 mg (0.2 mmol) THAP matrix and 2 mg (8.2 µmol) ammonium citrate to 500 µL of 1:1 (v/v) acetonitrile/water to form a supersaturated solution (a cloudy suspension). Allow suspended particles to settle to the bottom of the tube, forming a clear supernatant. 18. Pipet 1 µL THAP matrix supernatant onto a gold-plated 100-well plate. Pipet 1 µL ODN solution onto the same location. Allow spot to dry. There is no need for a desalting step because the ODN was purified by RP-HPLC.
Synthesis of Modified Oligonucleotides and Conjugates
4.24.19 Current Protocols in Nucleic Acid Chemistry
Supplement 18
19. Insert the plate into a Voyager Biospectrometry Workstation mass analyzer. Perform all MALDI-TOF-MS measurements in the positive ion mode. Table 4.24.5 lists the mass spectrometry results from a series of phosphonoacetate, thiophosphonoacetate, and chimeric oligomers.
COMMENTARY Background Information For many years now, oligodeoxyribonucleotides (ODNs) and their analogs have proven to be indispensable for most research in biology and biochemistry. They are used to study various biological processes through sequencing, PCR applications, and directed mutagenesis, as well as to diagnose diseases and study gene expression (DNA chips), identify single nucleotide polymorphisms, and modulate or control gene expression. The latter application has broad implications from basic research on gene function to the use of ODNs as therapeutic agents. As one would expect, success in these areas has been possible because of extensive research using a large number of ODNs. As each analog investigated thus far has a specific repertoire of advantages and disadvantages, it is worthwhile to continue developing new derivatives useful for biological research. This objective led to the PACE and SPACE oligomers whose synthesis chemistries are outlined in this manuscript.
Oligonucleotides with Phosphonoacetate and Thiophosphonoacetate Diesters
Synthesis using phosphonoacetic acid Phosphonoacetic acid has been known as an effective mimic of phosphoric acid and its derivatives for many decades. At neutral to basic pH, the three-dimensional structure and charge density of phosphonoacetate is remarkably similar to phosphate. The ability to mimic phosphate was demonstrated by the use of phosphonoacetic acid as an antiviral agent (Overby et al., 1977). Most methods for the synthesis of phosphonoacetic acid proceed through an oxidative transformation like an Arbuzov reaction (Arbuzov and Dunin, 1927), resulting in a phosphorus molecule in the oxidation state P(V). Once the carboxylic acid group is attached, the molecule becomes quite inactive to methods of coupling by activated transesterification. Even with these difficulties, phosphonoacetic acid was successfully coupled to a variety of nucleosides in an attempt to mimic 5 - and 3 -phosphorylated nucleosides; these molecules also demonstrated antiviral activity (Griengl et al., 1988; Lambert et al., 1989). The authors initially attempted to develop a method for synthesizing PACE ODNs using methyl 5 -O-(4,4 -dimethoxytrityl)-2 -
deoxythymidine-3 -O-phosphonoacetate prepared by the same methods developed for making phosphonoacetic acid–derived mononucleosides. This synthon was activated with various arylsulfonyl chlorides or 1-(2mesitylenesulfonyl)-3-nitro-1,2,4-triazole and condensed with 2 -deoxythymidine linked to a controlled-pore glass support. Unfortunately, coupling yields for these reactions never exceeded 5%, even after 24 hr (similar results were obtained with the acetic acid ethyl ester). A further problem with this approach was then discovered, namely that the alkaline conditions previously described to remove either the methyl or ethyl ester resulted in partial to complete destruction of the phosphonate internucleotide linkage. Due to these problems, it was concluded that P(V) coupling would never produce a useful method for PACE DNA synthesis. Rudolph et al. (1996) subsequently reported the ability to synthesize a 13-mer chimera having alternating phosphate and phosphonoacetate internucleotide linkages. Using the identical techniques previously described for making phosphonoacetic acid–derived mononucleosides, they were able to prepare a solutionphase 2 -deoxythymidine phosphonoacetate dimer as the acetic acid methyl ester and convert it to the 3 -phosphoramidite. Joining these dimers using classical phosphoramidite chemistry then produced the reported 13-mer chimera having alternating phosphonoacetate and phosphodiester linkages. Because the suspected 13-mer products were only characterized by relative HPLC retention times, it is unclear whether the methyl phosphonoacetate ester was completely removed from this molecule by extended base hydrolysis; however, this noteworthy manuscript reaffirmed the confidence that the successful synthesis of PACE oligodeoxyribonucleotides would require the development of novel P(III) methods. Synthesis using phosphinylacetic acid Although the literature is somewhat sparse, phosphinylacetic acids have been known for quite some time (Matrosov et al., 1972). These molecules were most often evaluated as potential solvents for charged heavy metal ions and used for applications such as the
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purification of uranium by selective solvolysis. As a result, these types of compounds have been characterized for their chemical and physical properties, but have never been described as reactive synthons in organic chemistry or used to prepare analogs of peptides, oligosaccharides, oligodeoxyribonucleotides, or lipids. Based upon the literature and the authors’ experience, a potentially important reason for this lack of interest was the difficult challenge inherent in synthesizing these derivatives and the inaccurate characterization data based upon the early literature. A successful new approach was therefore developed as outlined in Figures 4.24.2 and 4.24.3. Initially, methyl 2 -deoxythymidine-3 -O-(N,Ndiisopropylamino)phosphinyl acetate was synthesized and used to prepare a dimer having a PACE internucleotide linkage. The high isolated yield of this dimer (81%) was extremely encouraging. Further studies, however, confirmed that removal of the methyl ester with various alkaline solutions led to degradation of the internucleotide linkage. As a consequence, the acetic acid 2-cyanoethyl ester was next investigated, as there was an expectation that it could be removed using a non-nucleophilic base such as DBU. Although this type of ester protection proved promising, it was not entirely satisfactory, as ∼8% of the PACE internucleotide linkages were further modified to phosphate under DBU treatment. These observations led the group to the sterically hindered acetic acid 1,1-dimethyl-2-cyanoethyl ester, which could be removed essentially quantitatively (<1% hydrolysis) with non-nucleophilic base. Removal of protecting groups A related early concern during the development of this chemistry was potential problems associated with removal of the amine-protecting groups commonly used on cytosine, adenine, and guanine. These blocking groups are routinely eliminated with mild base at elevated temperatures, conditions that are known to degrade alkyl phosphonate internucleotide linkages. An alternative was found in the method developed by Reddy et al. (1994), which uses 40% methylamine in water to rapidly remove exocyclic amine from ODNs containing N-acetyl-2 -deoxycytidine residues without methylation of the exocyclic amine groups. By slightly modifying these conditions, it was possible to achieve complete removal of the heterobase-protecting groups in 15 min at 55◦ C. The results showed that,
under these conditions, both PACE and SPACE ODNs were relatively stable, as 1.4% and 1.2%, respectively, of the internucleotide linkages were degraded in 15 min. Mass spectroscopy of the resulting ODN products also demonstrated no methylation of cytidine residues protected by acetyl groups. Although not entirely satisfactory (the problem is under further investigation), this procedure is currently used in the authors’ laboratory to remove the base-protecting groups.
Compound Characterization Both 1 H and 31 P NMR are indispensable for analyzing precursors and synthons used to prepare these ODNs. 1 H NMR spectra are recorded on Varian VXR-300 and Bruker 400 MHz spectrometers with tetramethylsilane as an internal standard. A Bruker 400 MHz spectrophotometer and an external 85% H3 PO4 capillary are used for recording oligomer 31 P NMR in D2 O. In the case of chimeras, the number of PACE, S-PACE, phosphate, and thiophosphate internucleotide linkages can be determined, as they have different 31 P NMR spectra. MALDI-TOF mass spectrometry (UNIT 10.1) should be used to analyze the molecular weight of these oligomers and to confirm that the correct base composition has been achieved. This is important because PACE and S-PACE ODNs are resistant toward degradation by nucleases. As a consequence, classical enzymatic methods cannot be used to characterize these ODNs. Further analysis of these ODNs is possible using standard 20% polyacrylamide gel electrophoresis (UNIT 10.4) followed by visualization and densitometric scanning. It is also possible to derivatize these oligomers with fluorescein and use this label to visualize ODNs on polyacrylamide gels (Dellinger et al., 2003). In all cases, PACE and S-PACE ODNs exhibit electrophoretic mobilities similar to natural DNA of the same length.
Critical Parameters and Troubleshooting Reagent quality All nucleic acid chemists know that water (preferably the lack thereof) is the most critical parameter for synthesizing DNA, RNA, and their analogs. Certainly this rule applies to the synthesis of PACE and S-PACE ODNs. Among the various sources of water contamination, the most significant is acetonitrile, which must be anhydrous for several key synthesis steps. These include its use as a solvent for the synthons and tetrazole. Acetonitrile
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used to wash the CPG just prior to each condensation step should also be anhydrous. Currently, commercial synthesis-grade acetonitrile is usually sufficiently anhydrous for these steps. It should, however, be checked with a Karl Fisher titrator before use. In order to be absolutely sure that the acetonitrile is anhydrous, distillation from calcium hydride immediately before use is recommended. Another source of water is its adsorption on the synthons during storage. This can be minimized by storing the synthons at −20◦ C under dry argon in a desiccator equipped with Drierite. Prior to opening the desiccator, it should be allowed to attain room temperature. Following transfer of the synthons to vials for use on the synthesizer, the samples should be dried under vacuum overnight to remove final traces of water. In addition to water, there are other precautions that should be followed in order to ensure high repetitive yields during ODN synthesis. One is the quality of tetrazole used to activate the synthons. Commercially available material, if prepared specifically for oligodeoxyribonucleotide synthesis (0.45 M acetonitrile solution), is usually sufficiently pure for direct use; however, even reagent-grade tetrazole as obtained from various suppliers should be purified by sublimation before use in ODN synthesis. Another potential problem relates to the preparation of various synthons. During their synthesis, diisopropylammonium tetrazolide is generated. This salt must be completely removed during flash chromatography; otherwise, it will depress the overall ODN synthesis rate and yield by buffering the condensation reaction. Excessive amounts of this salt can be detected in the 1 H NMR spectra of the purified synthons.
Oligonucleotides with Phosphonoacetate and Thiophosphonoacetate Diesters
P(III) to P(V) oxidation Another potential problem that can be encountered during ODN synthesis is incomplete sulfurization or oxidation of P(III) to P(V). This leads to degradation of the oligomer, because trivalent phosphorus derivatives are known to be unstable to acid (a key step in the synthesis cycle is acid detritylation). It is difficult to monitor this problem during synthesis because the consequences (loss of stepwise trityl yields and overall deterioration of oligomer quality as observed by HPLC of the products) are common to several unrelated problems in DNA synthesis. If lack of complete oxidation is suspected, this step can be checked by 31 P gel-phase NMR (Greef et al., 1996); however, a more com-
mon solution is to simply replace the oxidation and sulfurization reagents with fresh samples. Once again, it is important to use the correct concentration of the oxidizing agent. The phosphonite internucleotide linkage produced after the phosphonamidite coupling reaction is more easily oxidized than the phosphite triester internucleotide linkage produced after a standard phosphoramidite coupling reaction. The phosphonite internucleotide linkage can be quantitatively oxidized by a 3-min oxidation using 0.1 M CSO in anhydrous acetonitrile. However, the phosphite triester internucleotide linkage requires 0.5 M CSO for at least 3 min. This is an important note for the synthesis of chimeric DNA sequences. Degradation It is well known that alkyl phosphonate internucleotide linkages are unstable toward basic conditions (Hogrefe et al., 1993). Phosphonoacetate and thiophosphonoacetate ODNs suffer from the same problem. For example, with aqueous 40% methylamine at 55◦ C, 1.4% and 1.2%, respectively, of the internucleotide linkages in these oligomers are degraded after 15 min. It is therefore important to limit the deprotection time with this reagent to the minimum required for effective removal of the exocyclic amine–protecting group on cytosine, guanine, and adenine bases. A related potential problem is the use of DBU to remove the dimethylcyanoethyl protecting group. These conditions must be anhydrous. It is recommended that the DBU be dried over 4-Å molecular sieves prior to use and careful attention be given to the concentration of DBU used for removal of the dimethylcyanoethyl protecting groups. The presence of water in the DBU solution will generate hydroxide anions, resulting in degradation of PACE and S-PACE internucleotide linkages, and can also lead to the premature cleavage of the ODN from the solid support. It is important that the concentration of DBU (1.5%) be accurately measured and that exposure of the CPG to this solution be limited to 60 min. Significantly higher concentrations of DBU (even in the absence of water) or very long exposure times can result in premature cleavage of the ODN from the CPG, reducing the recovery yields. Due to the deleterious effect of hydroxide ion on PACE and S-PACE internucleotide linkages, any base should be excluded from the chromatography solvents used to purify these synthons. As a consequence, chromatography must be carried out rapidly in order to minimize any detritylation, which can occur due to the acidity of the
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silica gel. The acidity of silica gel can vary greatly from batch to batch and from manufacturer to manufacturer. In the authors’ experience, most batches of silica gel are not acidic enough to give any significant amount of detritylation. Furthermore, there is a considerable amount of diisopropylammonium tetrazolide in the crude 2 -deoxynucleoside-3 O-(N,N-diisopropylamino)phosphinyl acetate products to help neutralize some excess acid on the silica gel. Nonetheless, detritylation does occur, albeit infrequently, during column chromatography. Detritylation on the column is easily detectable as a bright orange band on the silica column. To eliminate this, a small amount of TEA may be used to neutralize excess acid on silica gel during column packing and equilibration, but excess amine should be thoroughly flushed from the column prior to adding the crude material to the top of the column. Deprotection of internucleotide linkages The use of anhydrous DBU deprotection of cyanoethyl protecting groups on ODNs can result in the undesired side product of N3 alkylation of thymidine residues from the production of acrylonitrile under these conditions. These products can easily be detected by mass spectroscopy (both MALDI-MS and LC/MS). This is one of the reasons that the use of the dimethylcyanoethyl protecting group is recommended over the more typical cyanoethyl protecting group. When protecting groups for the PACE and S-PACE internucleotide linkages were first evaluated, it was discovered that removal of the cyanoethyl protecting group using anhydrous DBU resulted in small amounts of these N3 alkylation products. By switching from cyanoethyl to dimethylcyanoethyl protection of the PACE and S-PACE internucleotide linkages, it was found that detectable amounts of N3 alkylation of the thymidine residues on the ODN products were eliminated, even as evaluated by highly sensitive LC/MS/MS techniques. Handling modified ODNs Once the PACE oligodeoxyribonucleotides are produced, it is important to realize that the internucleotide bond is a weaker acid (pKa ∼ = 3.8) than the phosphodiester bond of native DNA (pKa ∼ = 1.8). The implications in handling these modified oligodeoxyribonucleotides have to do with their solubility in water under acidic conditions. Phosphonoacetate internucleotide bonds can be protonated with weak acids such as acetic acid. Once proto-
nated, the oligodeoxyribonucleotide is significantly less soluble in aqueous solutions. It is important that working solutions are buffered at neutral to slightly higher pH in order to prevent loss by precipitation and adherence to surfaces. This is especially important during trityl-on purification of the oligodeoxyribonucleotide. The final step is removal of the DMTr group using 80% glacial acetic acid followed by desalting on a reversed-phase column. The acetic acid is removed in a vacuum evaporator, leaving behind the modified ODN in its protonated form. In order to completely dissolve the ODN, it is important to use a buffer with enough capacity to neutralize the acidic sites on the backbone. This is accomplished by vortexing the sample vigorously during the initial resuspension of the ODN in a pH 8.0 triethylammonium acetate (TEAA) buffer, evaporating to dryness, and then redissolving the sample once again in TEAA prior to desalting.
Anticipated Results
1,1-Dimethyl-2-cyanoethyl 5 -O-(4,4 dimethoxytrityl)-2 -deoxynucleoside-3 -O(N,N-diisopropylamino)phosphinyl acetates have been used to prepare ODNs up to 25 nt in length. These oligomers contain PACE, S-PACE, phosphorothioate, and phosphate internucleotide linkages in various combinations and have isolated yields of ∼30%. Several examples of those that have been synthesized, including their mass spectral analysis, are included in Table 4.24.5. A typical HPLC profile as obtained by reversed-phase column chromatography (trityl-on) is shown in Figure 4.24.6. Although the peaks are broad, as expected due to the large number of diastereomers, it is clear that the expected product dominates the HPLC profile of the crude reaction mixture. When this oligomer is repurified by column chromatography after removal of the 4,4 dimethoxytrityl group, very few additional side products appear. At this stage of purification, these oligomers are essentially homogeneous (confirmed by analytical ion-exchange HPLC; Dellinger et al., 2003) and can be used for various biochemical experiments. A typical mass spectrum for the PACE Cap ODN (Table 4.24.5) is shown in Figure 4.24.7. Oligomers with stable acetic acid esters (methyl and butyl) have also been prepared and tested for uptake by HeLa and SKBR3 cells. The results (Sheehan et al., 2003) show that these ODNs reversibly accumulate in the cytoplasm of cells (presumably
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Figure 4.24.6 Reversed-phase column chromatography (trityl on) of the total reaction mixture obtained during the preparation of the PACE Cap ODN (see Table 4.24.5 for sequence). Gradient conditions and the column are described in the text.
Figure 4.24.7 MALDI-TOF-MS analysis of PACE Cap ODN. The mass spectrum was determined using conditions described in the text.
Oligonucleotides with Phosphonoacetate and Thiophosphonoacetate Diesters
by pinocytosis followed by ester hydrolysis). Current research in this area is focused upon enhancing in vivo acetic acid ester hydrolysis by using biologically relevant esters and studying these oligomers for antisense activity. The isolated yields of PACE and S-PACE ODNs depend upon the length, sequence, and number of analog internucleotide linkages. This is because both PACE and S-PACE internucleotide linkages are somewhat labile toward aqueous 40% methylamine. These yields for 18-mers, based upon the amount of
2 -deoxynucleoside attached to CPG, range from 20% to 30% following two reversedphase HPLC column purification steps (tritylon followed by trityl-off). The resulting oligomers are at least 95% homogeneous when evaluated by ion-exchange analytical column chromatography and are ready to be used for various biochemical experiments.
Time Considerations Synthesis of 1,1-dimethyl-2-cyanoethyl [bis(N,N-diisopropylamino)phosphinyl]acetate can take up to four days. The procedure begins
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with the preparation of 1,1-dimethyl-2cyanoethyl bromoacetate, which requires an overnight reflux of bromoacetyl bromide with 3-hydroxy-3-methylbutyronitrile in toluene. The next morning the reflux is stopped and the reaction mixture evaporated to an oil. The 1,1-dimethyl-2-cyanoethyl bromoacetate is purified by distillation, which requires 3 to 4 hr, and the day ends with 1 H NMR analysis of the bromoacetate distillate and 31 P NMR examination of the bis(N,Ndiisopropylamino)chlorophosphine. Clean up of the reflux equipment and set up of the distillation apparatus can usually be accomplished during the evaporation process. The Reformatsky coupling reaction to yield 1,1-dimethyl-2-cyanoethyl [bis(N,Ndiisopropylamino)phosphinyl]acetate will require 3 to 5 hr to set up and complete. Initial isolation of the product by trituration and liquid/liquid extraction will take an additional 3 to 4 hr, including combining the hexane fractions and evaporating the hexane solutions to an oil. The product is then redissolved in hexanes and allowed to cool overnight to precipitate additional impurities. This solution is decanted, concentrated to an oil using a rotary evaporator, and characterized by NMR and mass spectroscopy. This final process can take at least 3 to 4 hr. If optional crystallization of the product from pentane is carried out, the total procedure will be extended by several hours to one day. Preparation of the protected 2 deoxynucleoside-3 -O-(N,N-diisopropylamino)phosphinyl acetates takes ∼2 days. Phosphinylation of the protected 2 -deoxynucleoside is generally completed overnight, and the reaction mixture is evaluated the next morning by TLC and 31 P NMR. If the reaction is complete, triethylamine is added to quench the reaction and the product is purified by medium-pressure silica-gel column chromatography. This purification and isolation procedure usually requires 3 to 4 hr. For ease of handling, the product should be converted to a powder by precipitation into hexanes. This process usually requires only 30 min, but the powder must be dried in a vacuum desiccator overnight in order to remove residual hexanes. Set up for DNA synthesis requires attachment of various reagents and solvents to the DNA synthesizer. These include dichloromethane, acetonitrile, tetrazole (0.45 M in acetonitrile) the detritylation mixture, the two-part capping solution, and both oxidation and sulfurization reagents. This
process requires ∼1 hr. The four appropriately protected 2 -deoxynucleoside-3 -O-(N,Ndiisopropylamino)phosphinyl acetates are dissolved in anhydrous acetonitrile and the bottles placed on the DNA synthesizer. The DNA synthesis cycle for adding one 2 -deoxynucleotide requires ∼45 min. Thus, the automated synthesis of a 20-mer requires ∼15 hr. Deprotection and cleavage of the ODN from CPG takes ∼2 hr. Major steps are removal of the dimethylcyanoethyl group with anhydrous DBU (60 min) and treatment with aqueous methylamine (15 min, 55◦ C), which cleaves the oligomer from CPG and removes exocyclic amine–protecting groups. Following these steps, the reaction mixture is cooled and transferred to a microcentrifuge tube for concentration to dryness (1 to 2 hr). The ODNs are purified by reversed-phase HPLC using the 4,4 -dimethoxytrityl group for hydrophobic affinity. The entire procedure requires ∼12 hr. The first step is trityl-on RPHPLC (50 min) followed by collection of appropriate product fractions and concentration to dryness (2 to 3 hr). Next, the 4,4 dimethoxytrityl group is removed with aqueous 80% glacial acetic acid (90 min, room temperature) followed by concentration to dryness in a centrifugal concentrator. The ODN is then repurified and desalted by RP-HPLC. The same column and gradient can be used for both purifications. Appropriate product fractions are collected, combined, and concentrated to dryness. The resulting products are usually stored up to 6 months at −20◦ C in TE buffer, pH 8.0.
Literature Cited
¨ Arbuzov, A.E. and Dunin, A.A. 1927. Uber phosphon-carbons¨auren. Ber. 60B:291-295. Bayless, P.L. and Hauser, C.R. 1954. A Reformatskii type condensation of aroyl chlorides with ethyl 2-bromoisobutyrate by means of zinc to form β-keto esters. J. Am. Chem. Soc. 76:2306-2308. Caruthers, M.H., Barone, A.D., Beaucage, S.L., Dodds, D.R., Fisher, E.F., McBride, L.J., Matteucci, M., Stabinsky, Z., and J.-Y. Tang. 1987. Chemical synthesis of deoxyoligonucleotides by the phosphoramidite method. Methods Enzymol. 154:287-313. Dellinger, D.J., Sheehan, D.M., Christensen, N.K., Lindberg, J.G., and Caruthers, M.H. 2003. Solid phase chemical synthesis of phosphonoacetate and thiophosphonoacetate oligodeoxyribonucleotides. J. Am. Chem. Soc. 125:940-950. Glen Research. 1996. Non-aqueous oxidation with 10-camphorsulfonyl-oxziridine. Glen Research Corporation Technical Report 9:8-9.
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Greef, C.H., Seeberger, P.H., Caruthers, M.H., Beaton, G., and Bankaitis-Davis, D. 1996. Synthesis of phosphorodithioate RNA by the H-phosphonothioate method. Tetrahedron Lett. 37:4451-4454. Griengl, H., Hayden, W., Penn, G., Declercq, E., and Rosenwirth, B. 1988. Phosphonoformate and phosphonoacetate derivatives of 5-substituted 2 -deoxyuridines—synthesis and antiviral activity. J. Med. Chem. 31:1831-1839. Hogrefe, R.I., Reynolds, M.A., Vaghefi, M.M., Young, K.M., Riley, T.A., Klem, R.E., and Arnold, L.J. Jr. 1993. An improved method for the synthesis and deprotection of methylphosphonate oligonucleotides. In Protocols for Oligonucleotides and Analogs, Vol. 20 (S. Agrawal, ed.) pp. 143-164. Humana Press, Totowa, New Jersey. Lambert, R.W., Martin, J.A., Thomas, G.J., Duncan, I.B., Hall, M.J., and Heimer, E.P. 1989. Synthesis and antiviral activity of phosphonoacetic and phosphonoformic acid-esters of 5-bromo2 -deoxyuridine and related pyrimidine nucleosides and acyclonucleosides. J. Med. Chem. 32:367-374. Matrosov, E.I., Tsvetlsov, E.N., Malevannaya, R.A., and Kabachnik, M.I. 1972. Infrared spectra and association of phosphinylacetic acid. Zh. Obshch. Khim. 42:1695-1700. Overby, L.R., Duff, R.G., and Mao, J.C. 1977. Antiviral potential of phosphonoacetic acid. Ann. N.Y. Acad. Sci. 284:310-320.
Rudolph, M.J., Reitman, M.S., MacMillan, E.W., and Cook. A.F. 1996. Phosphonoacetate derivatives of oligodeoxyribonucleotides. Nucleosides Nucleotides 15:1725-1739. Sheehan, D., Lunstad, B., Yamada, C.M., Stell, B., Caruthers, M.H., and Dellinger, D.J. 2003. Biochemical properties of phosphonoacetate and thiophosphonoacetate oligodeoxyribonucleotides. Nucl. Acids Res. 31:4109-4118.
Key References Dellinger et al., 2003. See above. This reference outlines the chemistry used to prepare phosphonoacetate and thiophosphonoacetate ODNs from protected 2 -deoxynucleoside-3 O-(N,N-diisopropylamino)phosphinyl acetate synthons. Sheehan et al., 2003. See above. This reference presents initial biochemical and biophysical results with phosphonoacetate and thiophosphonoacetate ODNs.
Contributed by Douglas J. Dellinger Agilent Laboratories Boulder, Colorado Christina M. Yamada and Marvin H. Caruthers University of Colorado Boulder, Colorado
Reddy, M.P., Hanna, N.B., and Farooqui, F. 1994. Fast cleavage and deprotection of oligonucleotides. Tetrahedron Lett. 35:4311-4314.
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Base-Modified Oligodeoxyribonucleotides: Using Pyrrolo[2,3-d]pyrimidines to Replace Purines
UNIT 4.25
This unit describes the preparation of phosphoramidites and oligonucleotides derived from nucleosides related to the canonical DNA constituents 2 -deoxyadenosine (dA), 2 -deoxyguanosine (dG), and 2 -deoxyisoguanosine (iGd ). Because the 7-deazapurines (pyrrolo[2,3-d]pyrimidines) closely resemble purines in structure, these compounds are ideal shape mimics to be incorporated into DNA or RNA. Nucleosides may also have a halogen or alkynyl substituent at the 7 position of the 7-deazapurine moiety. Figure 4.25.1 shows a series of typical phosphoramidites used for oligonucleotide synthesis. The figure also compares systematic numbering, which is used for compound names in each protocol, with purine numbering, which is used elsewhere throughout the unit. Basic Protocols 1, 2, and 3 describe the preparation of the phosphoramidites of 7-deaza-7-iodo-2 -deoxyadenosine (I7 c7 Ad ; S.2d; Seela and Zulauf, 1998), 7-deaza7-iodo-2 -deoxyguanosine (I7 c7 Gd ; S.3d; Ramzaeva and Seela, 1996), and 7-deaza-2 deoxyisoguanosine (c7 iGd ; S.6a; Seela and Wei, 1999), respectively. These phosphoramidites are then employed in the solid-phase synthesis of 7-deazapurine-modified oligonucleotides (see Basic Protocol 4). The duplex stabilities of the modified oligonucleotides are subsequently measured (see Basic Protocol 5) and compared with the stabilities of oligonucleotide duplexes containing only the canonical DNA fragments. NOTE: Preparation of the starting nucleosides S.9d and S.12d is described in UNIT 1.10. The structure numbers in that unit do not correspond to the structure numbers used here. NOTE: Unless otherwise noted, use reagent-grade chemicals in all protocol steps. Store all phosphoramidites and oligonucleotides at –18◦ C.
SYNTHESIS OF THE PHOSPHORAMIDITE OF 7-DEAZA-7-IODO-2 -DEOXYADENOSINE
BASIC PROTOCOL 1
The synthesis of the phosphoramidite (S.2d) of 7-deaza-7-iodo-2 -deoxyadenosine (I7 c7 Ad ) is shown in Figure 4.25.2. The unprotected nucleoside S.9d serves as starting material; the preparation of this iodinated nucleoside is described in UNIT 1.10 (also see Seela and Zulauf, 1996). The preparation of the intermediates is straightforward, and the reaction steps presented result in high yields. Directions for preparing the phosphoramidites S.1a, S.1j, S.2a, and S.2f-S.2h (7-deazapurine derivatives of 2 -deoxyadenosine) are similar to those for preparing compound S.2d, with modifications that can be found by referring to the appropriate references (see Fig. 4.25.1). Synthesis of S.8 is similar to S.2 (also see Balow et al., 1998; Okamoto et al., 2002).
Materials 4-Amino-7-(2-deoxy-β-D-erythro-pentofuranosyl)-5-iodo-7H-pyrrolo[2,3d]pyrimidine (systematic numbering; S.9d; see UNIT 1.10), also known as 7-deaza-7-iodo-2 -deoxyadenosine (purine numbering) Methanol (MeOH) N,N-Dimethylformamide dimethylacetal Silica gel 60 for flash chromatography (particle size, <0.063 mm; Merck) Anhydrous dichloromethane (CH2 Cl2 ) Anhydrous pyridine
Contributed by Frank Seela and Xiaohua Peng Current Protocols in Nucleic Acid Chemistry (2005) 4.25.1-4.25.25 C 2005 by John Wiley & Sons, Inc. Copyright
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Figure 4.25.1
Legend at right.
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Figure 4.25.2 Synthesis of the phosphoramidite derivative of 7-deaza-7-iodo-2 -deoxyadenosine. DMTr, 4,4 -dimethoxytrityl.
4,4 -Dimethoxytrityl chloride (DMTr-Cl) 5% (w/v) sodium bicarbonate (NaHCO3 ) Brine (i.e., saturated aqueous NaCl) Anhydrous sodium sulfate (Na2 SO4 ) Anhydrous N,N-diisopropylethylamine (DIPEA; 99.5% pure) 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite Anhydrous triethylamine (Et3 N) Acetone 10-, 25-, 50-, and 100-mL round-bottom flasks Rotary evaporator connected to a vacuum pump 4 × 70–cm chromatography columns 0.2-mm-thick silica gel 60 F254 aluminum TLC plates (Merck) 254-nm UV lamp High-vacuum pump (final pressure, <1 mmHg) 100-mL separatory funnels 5-cm-diameter funnel with folded 10-cm-diameter Whatman no. 1 filter Argon source Additional reagents and equipment for flash chromatography (APPENDIX 3E) and thin-layer chromatography (TLC; APPENDIX 3D)
Figure 4.25.1 (at left) Selected phosphoramidites used for the synthesis of 7-deazapurinecontaining oligonucleotides. DMTr, 4,4 -dimethoxytrityl. References: S.1a (Seela and Kehne, 1985); S.1j (Seela et al., 2005a); S.2a (Seela et al., 1989); S.2d,f-h (Seela and Zulauf, 1998; Seela and Zulauf, 1999); S.3a (Seela and Driller, 1989); S.3c,d (Ramzaeva and Seela, 1996); S.3e (Seela and Shaikh, 2005), S.3f-h (Ramzaeva et al., 1997); S.3i (Seela and Chen, 1997); S.4a (Seela and Kaiser, 1986); S.5a (Seela et al., 2003); S.6a (Seela and Wei, 1999); S.7b,c and S.8a-d (Seela et al., 2005b).
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Protect exocyclic 4-amine 1. In a 100-mL round-bottom flask equipped with a Teflon stir bar, suspend (with stirring) 400 mg (1.06 mmol) 4-amino-7-(2-deoxy-β-D-erythro-pentofuranosyl)-5iodo-7H-pyrrolo[2,3-d]pyrimidine (S.9d) in 20 mL MeOH at room temperature. 2. Add 1.34 mL (10.0 mmol) N,N-dimethylformamide dimethylacetal and stir the resulting mixture 10 hr at 40◦ C, using an oil bath and contact thermometer to maintain the temperature. 3. Adsorb the reaction mixture onto ∼2 g silica gel 60. Remove the solvent from the adsorbed sample using a rotary evaporator connected to a vacuum pump. 4. Load the dry material onto a 4 × 70–cm column packed with silica gel 60 (bed height, 10 cm). Perform flash chromatography (APPENDIX 3E) using 0.4 bar pressure and an elution gradient from 100% CH2 Cl2 (200 mL) to 90:10 (v/v) CH2 Cl2 /MeOH (400 mL). Collect 10-mL fractions. 5. Monitor fractions by TLC (APPENDIX 3D) on silica gel 60 F254 plates using 90:10 (v/v) CH2 Cl2 /MeOH as the eluent. Visualize products under 254-nm light using a UV lamp. 6. Pool all product fractions, evaporate the solvent using a rotary evaporator, and dry the resulting sample overnight under high vacuum. 7-(2-Deoxy-β-D-erythro-pentofuranosyl)-4-{[(dimethylamino)methylidene]amino}-5iodo-7H-pyrrolo[2,3-d]pyrimidine (S.10) is obtained as a colorless solid (389 mg, 85% yield). TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.5. UV (MeOH): λmax (ε) = 277 (10,400), 323 nm (19,000). 1 H NMR (DMSO-d6 ): δ 2.18 (m, 1H, 2 -Hα ), 2.47 (m, 1H, 2 -Hβ ), 3.18 and 3.22 (2 × s, 6H, Me2 N), 3.54 (m, 2H, 5 -H), 3.81 (m, 1H, 4 -H), 4.32 (m, 1H, 3 -H), 5.00 (t, J = 5.4 Hz, 1H, 5 -OH), 5.23 (d, J = 3.9 Hz, 1H, 3 -OH), 6.52 (t, J = 7.0 Hz, 1H, 1 -H), 7.70 (s, 1H, 6-H), 8.30 (s, 1H, 2-H), 8.82 (s, 1H, N=CH). Anal. calcd. for C14 H18 IN5 O3 : C, 38.99; H, 4.21; N, 16.24; found: C, 39.09; H, 4.27; N, 16.10.
Protect 5 -OH group 7. In a 25-mL round-bottom flask equipped with a Teflon stir bar, prepare a solution of 300 mg (0.70 mmol) compound S.10 in 3.0 mL anhydrous pyridine. 8. Add 256 mg (0.76 mmol) DMTr-Cl in a single portion (with stirring) and stir the resulting mixture 1 hr at 50◦ C, using an oil bath and contact thermometer to maintain the temperature. 9. Pour the reaction mixture into a 100-mL separatory funnel containing 20 mL of 5% NaHCO3 . 10. Extract the aqueous solution twice with 30 mL CH2 Cl2 . Combine the resulting organic layers, wash with brine, and then dry over 2 g anhydrous Na2 SO4 . 11. Filter off the Na2 SO4 by gravity filtration through a folded 10-cm-diameter filter. Evaporate the filtrate to dryness using a rotary evaporator. 12. Dissolve the crude product in ∼2 mL CH2 Cl2 , load it onto a 4 × 70–cm column packed with silica gel 60, and perform flash chromatography (step 4) using an elution gradient from 100% CH2 Cl2 (200 mL) to 90:10 (v/v) CH2 Cl2 /acetone (300 mL). Collect 10-mL fractions. 13. Monitor fractions by TLC (step 5) using 90:10 (v/v) CH2 Cl2 /MeOH as the eluent. Using Pyrrolo[2,3d]pyrimidines to Replace Purines
14. Pool all product fractions, evaporate the solvent using a rotary evaporator, and dry the resulting sample overnight under high vacuum.
4.25.4 Supplement 20
Current Protocols in Nucleic Acid Chemistry
7-[2-Deoxy-5-O-(4,4 -dimethoxytriphenylmethyl)-β-D-erythro-pentofuranosyl]-4{[(dimethylamino)methylidene]amino}-5-iodo-7H-pyrrolo[2,3-d]pyrimidine (S.11) is obtained as a colorless foam (360 mg, 71% yield). TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.6. UV (MeOH): λmax (ε) = 236 (29,200), 275 (12,200), 322 nm (19,600). 1 H NMR (DMSO-d6 ): δ 2.24 (m, 1H, 2 -Hα ), 2.57 (m, 1H, 2 -Hβ ), 3.18 (m, 2H, 5 -H), 3.18 and 3.22 (2 × s, 6H, Me2 N), 3.72 (2 × s, 6H, 2 × MeO), 3.92 (m, 1H, 4 -H), 4.37 (m, 1H, 3 -H), 5.30 (d, J = 4.0 Hz, 1H, 3 -OH), 6.54 (t, J = 6.6 Hz, 1H, 1 -H), 6.84 (m, 4H, aromatic H), 7.22-7.38 (m, 9H, aromatic H), 7.56 (s, 1H, 6-H), 8.31 (s, 1H, 2-H), 8.82 (s, 1H, N=CH). Anal. calcd. for C35 H36 IN5 O5 : C, 57.30; H, 4.95; N, 9.55; found: C, 57.48; H, 5.12; N, 9.44.
Prepare phosphoramidite (S.2d) 15. In a 10-mL round-bottom flask equipped with a Teflon stir bar and kept under argon, prepare a solution of 300 mg (0.41 mmol) compound S.11 in 5 mL anhydrous CH2 Cl2 . 16. Add 104 µL (0.60 mmol) anhydrous N,N-diisopropylethylamine and 118 µL (0.53 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite and stir the resulting mixture 15 min at room temperature under argon. 17. Transfer the reaction mixture to a 100-mL separatory funnel and dilute with 30 mL CH2 Cl2 . Wash twice, each time with 20 mL of 5% NaHCO3 , wash once with 20 mL brine, and then dry over 3 g Na2 SO4 . 18. Filter off the Na2 SO4 and evaporate the filtrate to dryness (step 11). 19. Prewash a 4 × 70–cm silica gel 60 column (bed height, 8 cm) with 200 mL CH2 Cl2 containing 1% (v/v) anhydrous Et3 N and then with 100 mL of 100% CH2 Cl2 . 20. Dissolve the crude product in ∼2 mL CH2 Cl2 , load it onto the column, and perform flash chromatography (step 4) using 95:5 (v/v) CH2 Cl2 /acetone (400 mL) as the eluent. Collect 10-mL fractions. 21. Monitor fractions by TLC (step 5) using 90:10 (v/v) CH2 Cl2 /acetone as the eluent. 22. Pool all product fractions and evaporate the solvent using a rotary evaporator. 23. Dissolve in a small volume (∼2 mL) of CH2 Cl2 in a 25-mL round-bottom flask, evaporate the solvent using a rotary evaporator, and dry the resulting sample under high vacuum for 24 hr at 40◦ C. 7-[2-Deoxy-5-O-(4,4 -dimethoxytriphenylmethyl)-β-D-erythro-pentofuranosyl]-4{[(dimethylamino)methylidene]amino}-5-iodo-7H-pyrrolo[2,3-d]pyrimidine-3 -[(2cyanoethyl)-N,N-diisopropylphosphoramidite] (S.2d) is obtained as a colorless foam (222 mg, 58% yield). TLC (CH2 Cl2 /acetone, 9:1): Rf = 0.24 and 0.32. 31 P NMR (CDCl3 ): δ 149.0, 149.2.
24. If the synthesized phosphoramidite still contains impurities, remove those impurities in the following way. a. In a 100-mL round-bottom flask equipped with a Teflon stir bar, dissolve 0.3 to 1.0 g of the synthesized phosphoramidite in 1 to 2 mL CH2 Cl2 . b. Add ∼90 mL cyclohexane dropwise, with continuous shaking or stirring to dislodge any solid stuck to the walls of the flask. c. Refrigerate overnight. d. Remove the supernatant from the flask and dissolve the residue in 5 mL CH2 Cl2 . e. Transfer the contents of the flask to a 25-mL round-bottom flask, evaporate the solvent using a rotary evaporator, and dry the resulting sample under high vacuum for 24 hr at 40◦ C.
Synthesis of Modified Oligonucleotides and Conjugates
4.25.5 Current Protocols in Nucleic Acid Chemistry
Supplement 20
Figure 4.25.3 Synthesis of the phosphoramidite derivative of 7-deaza-7-iodo-2 -deoxyguanosine. DMTr, 4,4 dimethoxytrityl; i-Bu2 O, isobutyric anhydride.
BASIC PROTOCOL 2
SYNTHESIS OF THE PHOSPHORAMIDITE OF 7-DEAZA-7-IODO-2 -DEOXYGUANOSINE The synthesis of the phosphoramidite (S.3d) of 7-deaza-7-iodo-2 -deoxyguanosine (I7 c7 Gd ; Fig. 4.25.3) is performed as described for the phosphoramidite S.2d (see Basic Protocol 1), with the exception that protection of the 2-amino group is transient (Ti et al., 1982). The preparation of the iodinated nucleoside S.12d is described in UNIT 1.10 (also see Ramzaeva and Seela, 1995). Directions for preparing the phosphoramidites S.3a, S.3c, and S.3e-S.3i (7-deazapurine derivatives of 2 -deoxyguanosine) are similar to those for preparing compound S.3d, with modifications that can be found by referring to the appropriate references (see Fig. 4.25.1).
Materials
Using Pyrrolo[2,3d]pyrimidines to Replace Purines
4-Amino-7-(2-deoxy-β-D-erythro-pentofuranosyl)-5-iodo-7H-pyrrolo[2,3d]pyrimidin-2-one (systematic numbering; S.12d; see UNIT 1.10), also known as 7-deaza-7-iodo-2 -deoxyguanosine (purine numbering) Anhydrous pyridine Chlorotrimethylsilane (Me3 SiCl) Isobutyric anhydride 25% (v/v) ammonium hydroxide (NH4 OH) 4,4 -Dimethoxytrityl chloride (DMTr-Cl) 5% (w/v) sodium bicarbonate (NaHCO3 ) Anhydrous dichloromethane (CH2 Cl2 ) Anhydrous sodium sulfate (Na2 SO4 ) Silica gel 60 for flash chromatography (particle size, <0.063 mm; Merck) Acetone Methanol (MeOH) Anhydrous N,N-diisopropylethylamine (DIPEA) 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite Anhydrous triethylamine (Et3 N) Ethyl acetate (EtOAc)
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10-, 25-, 50-, and 250-mL round-bottom flasks Rotary evaporator connected to a vacuum pump 3-cm-diameter Buchner funnel with filter paper circles High-vacuum pump (final pressure, <1 mmHg) 100-mL separatory funnels 5-cm-diameter funnel with folded 10-cm-diameter Whatman no. 1 filter 4 × 70–cm chromatography columns 0.2-mm-thick silica gel 60 F254 aluminum TLC plates (Merck) 254-nm UV lamp Argon source Additional reagents and equipment for flash chromatography (APPENDIX 3E) and thin-layer chromatography (TLC; APPENDIX 3D) Protect exocyclic 2-amine 1. In a 25-mL round-bottom flask equipped with a stir bar, prepare a solution of 300 mg (0.77 mmol) 4-amino-7-(2-deoxy-β-D-erythro-pentofuranosyl)-5-iodo-7Hpyrrolo[2,3-d]pyrimidin-2-one (S.12d) in 4 mL anhydrous pyridine. 2. Add 0.48 mL (3.78 mmol) Me3 SiCl and stir the resulting mixture 15 min at room temperature. 3. Add 0.62 mL (3.74 mmol) isobutyric anhydride and stir the resulting mixture 3 hr at room temperature. 4. Cool the reaction mixture in an ice bath. After cooling, add 1 mL cold H2 O and stir 5 min on ice. 5. Add 1 mL of 25% NH4 OH and stir the resulting mixture 15 min at room temperature. 6. Evaporate the contents of the round-bottom flask down to a small volume using a rotary evaporator. 7. Crystallize the product (S.13) by adding ∼10 ml H2 O, which should yield colorless crystals. Collect the crystals by vacuum filtration with a 3-cm Buchner funnel, wash twice with 2 mL H2 O, and then dry overnight under high vacuum. 7-(2-Deoxy-β-D-erythro-pentofuranosyl)-3,7-dihydro-5-iodo-2-(isobutyrylamino)-4Hpyrrolo[2,3-d]pyrimidin-4-one (S.13) is obtained as colorless crystals (312 mg, 88% yield). TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.4. m.p. 188◦ C. 1 H NMR (DMSO-d6 ): δ 1.01 (s, 6H, 2 Me), 2.12 (m, 1H, 2 -Hα ), 2.37 (m, 1H, 2 -Hβ ), 2.73 (m, 1H, CH), 3.50 (m, 2H, 5 -H), 3.78 (m, 1H, 4 -H), 4.30 (m, 1H, 3 -H), 4.89 (t, J = 5.7 Hz, 1H, 5 -OH), 5.20 (d, J = 4.3 Hz, 1H, 3 -OH), 6.35 (t, J = 6.5 Hz, 1H, 1 -H), 7.43 (s, 1H, 6-H), 11.49 (s, 1H, NH), 11.76 (s, 1H, NH). Anal. calcd. for C15 H19 IN4 O5 : C, 38.98; H, 4.14; N, 12.12; found: C, 39.11; H, 4.37; N, 11.96.
Protect 5 -OH group 8. In a 25-mL round-bottom flask equipped with a Teflon stir bar, prepare a solution of 400 mg (0.87 mmol) compound 13 in 3.0 mL anhydrous pyridine. 9. Add 328 mg (0.97 mmol) DMTr-Cl and stir the resulting mixture 12 hr at room temperature. 10. Pour into a 100-mL separatory funnel containing 30 mL of 5% NaHCO3 . 11. Extract the aqueous solution twice, each time with 50 mL CH2 Cl2 . Combine the resulting organic layers and dry them over 5 g Na2 SO4 . 12. Filter off the Na2 SO4 by gravity filtration through a folded 10-cm-diameter filter. Evaporate the filtrate to dryness using a rotary evaporator.
Synthesis of Modified Oligonucleotides and Conjugates
4.25.7 Current Protocols in Nucleic Acid Chemistry
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13. Dissolve the crude product in ∼2 mL CH2 Cl2 and load it onto a 4 × 70–cm column packed with silica gel 60 (bed height, 10 cm). Perform flash chromatography (APPENDIX 3E) using 0.4 bar pressure and an elution gradient from 100% CH2 Cl2 (200 mL) to 85:15 (v/v) CH2 Cl2 /acetone (400 mL). Collect 10-mL fractions. 14. Monitor fractions by TLC (APPENDIX 3D) on silica gel 60 F254 plates using 5:1 (v/v) CH2 Cl2 /MeOH as the eluent. Visualize products under 254-nm light using a UV lamp. 15. Pool all product fractions, evaporate the solvent using a rotary evaporator, and dry the resulting sample overnight under high vacuum. 7-[2-Deoxy-5-O-(4,4 -dimethoxytriphenylmethyl)-β-D-erythro-pentofuranosyl]-3,7dihydro-5-iodo-2-(isobutyrylamino)-4H-pyrrolo[2,3-d]pyrimidin-4-one (S.14) is obtained as a colorless, amorphous solid (600 mg, 91% yield). TLC (CH2 Cl2 /MeOH, 5:1): Rf = 0.5. 1 H NMR (DMSO-d6 ): δ 1.13 (m, 12H, 4 Me), 2.24 (m, 2H, 2 -H), 2.77 (m, 1H, CH), 3.12 (m, 2H, 5 -H), 3.75 (s, 6H, 2 MeO), 3.93 (m, 1H, 4 -H), 4.35 (m, 1H, 3 -H), 5.30 (d, J = 4.5 Hz, 1H, 3 -OH), 6.39 (t, J = 6.5 Hz, 1H, 1 -H), 6.86-7.39 (m, 13H, aromatic H, 6-H), 11.54 (s, 1H, NH), 11.82 (s, 1H, NH). Anal. calcd. for C36 H37 IN4 O7 : C, 56.55; H, 4.88; N, 7.33; found: C, 56.42; H, 4.82; N, 7.30.
Prepare phosphoramidite (S.3d) 16. To a 25-mL round-bottom flask equipped with a Teflon stir bar, add 230 mg (0.30 mmol) compound S.14. Purge with argon, and then add 5.0 mL anhydrous CH2 Cl2 . 17. Add 100 µL (0.58 mmol) anhydrous N,N-diisopropylethylamine and 100 µL (0.45 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite and stir the resulting mixture 30 min at room temperature under argon. 18. Transfer the reaction mixture to a 100-mL separatory funnel and dilute with 30 mL CH2 Cl2 . Wash twice, each time with 20 mL of 5% NaHCO3 , and then dry over Na2 SO4 . 19. Filter off the Na2 SO4 and evaporate the filtrate to dryness using a rotary evaporator. 20. Prewash a 4 × 70–cm silica gel 60 column (bed height, 10 cm) with 200 mL CH2 Cl2 containing 1% (v/v) anhydrous Et3 N and then with 100 mL of 100% CH2 Cl2 . 21. Dissolve the crude product in ∼2 mL CH2 Cl2 , load it onto the column, and perform flash chromatography (step 13) using 95:5 (v/v) CH2 Cl2 /acetone (400 mL) as the eluent. Collect 10-mL fractions. 22. Monitor fractions by TLC (step 14) using 69:30:1 (v/v/v) CH2 Cl2 /EtOAc/Et3 N as the eluent. 23. Pool the S.3d-containing fractions and evaporate the solvent using a rotary evaporator. 24. Dissolve in a small volume (∼2 mL) of CH2 Cl2 . Transfer this solution to a 10-mL round-bottom flask, evaporate the solvent using a rotary evaporator, and dry the resulting sample under high vacuum for 24 hr at 40◦ C. 7-[(2-Deoxy-5-O-(4,4 -dimethoxytriphenylmethyl)-β-D-erythro-pentofuranosyl]3,7-dihydro-5-iodo-2-(isobutyrylamino)-4H-pyrrolo[2,3-d]pyrimidin-4-one-3 -[(2cyanoethyl)-N,N-diisopropylphosphoramidite] (S.3d) is obtained as a colorless foam (125 mg, 54% yield). TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.8. 31 P NMR (CDCl3 ): δ 148.0, 148.7. Using Pyrrolo[2,3d]pyrimidines to Replace Purines
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25. If the synthesized phosphoramidite still contains impurities, remove those impurities in the following way. a. In a 100-mL round-bottom flask equipped with a Teflon stir bar, dissolve 0.3 to 1.0 g of the synthesized phosphoramidite in 1 to 2 mL CH2 Cl2 . b. Add ∼90 mL cyclohexane dropwise, with continuous shaking or stirring to dislodge any solid stuck to the walls of the flask. c. Refrigerate overnight. d. Remove the supernatant from the flask and dissolve the residue in 5 mL CH2 Cl2 . e. Transfer the contents of the flask to a 25-mL round-bottom flask, evaporate the solvent using a rotary evaporator, and dry the resulting sample under high vacuum for 24 hr at 40◦ C.
SYNTHESIS OF THE PHOSPHORAMIDITE OF 7-DEAZA-2 -DEOXYISOGUANOSINE
BASIC PROTOCOL 3
The synthesis of the phosphoramidite S.6a is performed as described for the phosphoramidite S.3d (see Basic Protocol 2), with the exception that the 2-oxo group of 7deaza-2 -deoxyisoguanosine (S.15a) must be protected with a diphenylcarbamoyl group (Fig. 4.25.4). To avoid the formation of the 2,5 -bis(diphenylcarbamoyl) derivative, the ratio of diphenylcarbamoyl chloride to compound S.15a should not exceed 3:2. The preparation of the starting nucleoside S.15a is described in Seela and Wei (1999). Directions for preparing the phosphoramidites S.7b and S.7c (7-deazapurine derivatives of 2 -deoxyisoguanosine) are similar to those for preparing compound S.6a, with modifications that can be found by referring to the appropriate references (see Fig. 4.25.1).
Materials 4-Amino-7-(2-deoxy-β-D-erythro-pentofuranosyl)-1,7-dihydro-2H-pyrrolo[2,3d]pyrimidin-2-one (systematic numbering; S.15a; see Seela and Wei, 1999), also known as 7-deaza-2 -deoxyisoguanosine (purine numbering) Anhydrous pyridine Diphenylcarbamoyl chloride Anhydrous N,N-diisopropylethylamine (DIPEA) Crushed ice 5% (w/v) sodium bicarbonate (NaHCO3 ) Anhydrous dichloromethane (CH2 Cl2 ) Anhydrous sodium sulfate (Na2 SO4 ) Silica gel 60 for flash chromatography (particle size, <0.063 mm; Merck) Methanol (MeOH) Chlorotrimethylsilane (Me3 SiCl) Isobutyryl chloride 25% (v/v) ammonium hydroxide (NH4 OH) 4,4 -Dimethoxtrityl chloride (DMTr-Cl) Acetone 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite Brine (i.e., saturated aqueous NaCl) Anhydrous triethylamine (Et3 N) 10-, 50-, 100-, and 250-mL round-bottom flasks 50- and 100-mL separatory funnels 5-cm-diameter funnel with folded 10-cm-diameter Whatman no. 1 filter Rotary evaporator connected to a vacuum pump 4 × 70–cm chromatography columns
Synthesis of Modified Oligonucleotides and Conjugates
4.25.9 Current Protocols in Nucleic Acid Chemistry
Supplement 20
Figure 4.25.4 Synthesis of the phosphoramidite derivative of 7-deaza-2 -deoxyisoguanosine. DMTr, 4,4 -dimethoxytrityl; i-Bu-Cl, isobutyryl chloride.
0.2-mm-thick silica gel 60 F254 aluminum TLC plates (Merck) 254-nm UV lamp High-vacuum pump (final pressure, <1 mmHg) Argon source Additional reagents and equipment for flash chromatography (APPENDIX 3E) and thin-layer chromatography (TLC; APPENDIX 3D)
Using Pyrrolo[2,3d]pyrimidines to Replace Purines
Protect exocyclic 2-oxo group 1. In a 100-mL round-bottom flask equipped with a Teflon stir bar, suspend (with stirring) 1.15 g (4.32 mmol) 4-amino-7-(2-deoxy-β-D-erythro-pentofuranosyl)-1,7dihydro-2H-pyrrolo[2,3-d]pyrimidin-2-one (S.15a) in 25 mL anhydrous pyridine.
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2. Add 1.54 g (6.65 mmol) diphenylcarbamoyl chloride and 1.1 mL (6.32 mmol) anhydrous N,N-diisopropylethylamine and stir the resulting mixture 2 hr at room temperature. 3. Add ∼5 g crushed ice to the reaction mixture and stir 5 to 10 min. 4. Pour into a 100-mL separatory funnel containing 30 mL of 5% NaHCO3 . 5. Extract the aqueous solution twice, each time with 40 mL CH2 Cl2 . Combine the resulting organic layers and dry them over 5 g Na2 SO4 . 6. Filter off the Na2 SO4 by gravity filtration through a folded 10-cm-diameter filter. Evaporate the filtrate to dryness using a rotary evaporator. 7. Dissolve the crude product in ∼5 mL of 97:3 (v/v) CH2 Cl2 /MeOH and load it onto a 4 × 70–cm column packed with silica gel 60 (bed height, 10 cm). Perform flash chromatography (APPENDIX 3E) using 0.4 bar pressure and an elution gradient from 97:3 (v/v) CH2 Cl2 /MeOH (300 mL) to 90:10 (v/v) CH2 Cl2 /MeOH (500 mL). Collect 10-mL fractions. 8. Monitor fractions by TLC (APPENDIX 3D) on silica gel 60 F254 plates using 90:10 (v/v) CH2 Cl2 /MeOH as the eluent. Visualize products under 254-nm light using a UV lamp. 9. Pool all product fractions, evaporate the solvent using a rotary evaporator, and dry the resulting sample overnight under high vacuum. 4-Amino-7-(2-deoxy-β-D-erythro-pentofuranosyl)-7H-pyrrolo[2,3-d]pyrimidin-2-yl diphenylcarbamate (S.16) is obtained as a colorless foam (1.58 g, 79% yield). TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.40. UV (MeOH): λmax (ε) = 269 nm (12900). 1 H NMR (DMSO-d6 ): δ 2.15 (m, 1H, 2 -Hα ), 2.42 (m, 1H, 2 -Hβ ), 3.53 (m, 2H, 5 -H), 3.80 (m, 1H, 4 -H), 4.33 (m, 1H, 3 -H), 4.96 (t, J = 5.5 Hz, 1H, 5 -OH), 5.25 (d, J = 4.0 Hz, 1H, 3 -OH), 6.39 (t, J = 6.8 Hz, 1H, 1 -H), 6.59 (d, J = 3.5 Hz, 1H, 5-H), 7.25-7.48 (m; 13H, 6-H; NH2 and aromatic H). Anal. calcd. for C24 H23 N5 O5 : C, 62.46; H, 5.02; N, 15.18; found: C, 62.63; H, 5.17; N, 15.08.
Protect exocyclic 4-amine 10. In a 100-mL round-bottom flask equipped with a Teflon stir bar, prepare a solution of 1.03 g (2.23 mmol) compound 16 in 20 mL anhydrous pyridine. 11. Add 2.0 mL (15.76 mmol) Me3 SiCl and stir the resulting mixture 25 min at room temperature. 12. Add 0.80 mL (7.64 mmol) isobutyryl chloride and stir the resulting mixture 2 hr at room temperature. 13. Cool the reaction mixture in an ice bath. After cooling, add 5.0 mL prechilled H2 O and stir 5 to 10 min on ice. 14. Add 12 mL of 25% NH4 OH and stir the resulting mixture 2.5 hr at room temperature. 15. Pour the mixture into a 100-mL separatory funnel containing 30 mL of 5% NaHCO3 . 16. Extract the aqueous solution four times, each time with 40 mL CH2 Cl2 . Combine the resulting organic layers and dry them over 5 g anhydrous Na2 SO4 . 17. Filter off the Na2 SO4 by gravity filtration through a folded 10-cm-diameter filter. Evaporate the filtrate to dryness using a rotary evaporator. 18. Dissolve the crude product in ∼5 mL of 98:2 (v/v) CH2 Cl2 /MeOH and perform flash chromatography (step 7) using an elution gradient from 98:2 (v/v) CH2 Cl2 /MeOH (300 mL) to 95:5 (v/v) CH2 Cl2 /MeOH (500 mL). Collect 10-mL fractions.
Synthesis of Modified Oligonucleotides and Conjugates
4.25.11 Current Protocols in Nucleic Acid Chemistry
Supplement 20
19. Monitor fractions by TLC (step 8) using 90:10 (v/v) CH2 Cl2 /MeOH as the eluent. 20. Pool all product fractions, evaporate the solvent using a rotary evaporator, and dry the resulting sample overnight under high vacuum. 7-(2-Deoxy-β-D-erythro-pentofuranosyl)-4-[(2-methylpropanoyl)amino]-7Hpyrrolo[2,3-d]pyrimidin-2-yl diphenylcarbamate (S.17) is obtained as a colorless foam (0.95 g, 80% yield). TLC (CH2 Cl2 /MeOH, 9:1): Rf = 0.52. UV (MeOH): λmax (ε) = 221 (40,000), 233 (36,500), 294 nm (7,200). 1 H NMR (DMSO-d6 ): δ 1.11 and 1.14 (2 × s, 6H, 2 Me), 2.21 (m, 1H, 2 -Hα ), 2.42 (m, 1H, 2 -Hβ ), 2.87 (m, 1H, CH), 3.54 (m, 2H, 5 -H), 3.83 (m, 1H, 4 -H), 4.36 (m, 1H, 3 -H), 4.92 (t, J = 5.3 Hz, 1H, 5 -OH), 5.31 (d, J = 4.2 Hz, 1H, 3 -OH), 6.55 (t, J = 6.5 Hz, 1H, 1 -H), 6.84 (d, J = 4.0 Hz, 1H, 5-H), 7.30-7.63 (m, 11H, 6-H and aromatic H), 11.0 (s, 1H, NH). Anal. calcd. for C28 H29 N5 O6 : C, 63.27; H, 5.50; N, 13.18; found: C, 63.19; H, 5.60; N, 13.07.
Protect 5 -OH group 21. In a 50-mL round-bottom flask equipped with a Teflon stir bar, prepare a solution of 531 mg (1.0 mmol) compound 17 in 10.0 mL anhydrous pyridine. 22. Add 373 mg (1.10 mmol) DMTr-Cl and stir the resulting mixture 3.5 hr at room temperature. 23. Add 5 mL MeOH and stir the resulting mixture 15 min at room temperature. 24. Pour into a 50-mL separatory funnel containing 15 mL of 5% NaHCO3 . 25. Extract the aqueous solution three times, each time with 15 mL CH2 Cl2 . Combine the resulting organic layers and dry them over Na2 SO4 . 26. Filter off the Na2 SO4 by gravity filtration through a folded 10-cm-diameter filter. Evaporate the filtrate to dryness using a rotary evaporator. 27. Dissolve the crude product in ∼2 mL CH2 Cl2 and load it onto a 4 × 70–cm column packed with silica gel 60 (bed height, 8 cm). Perform flash chromatography (step 7) using an elution gradient from 100% CH2 Cl2 (300 mL) to 90:10 (v/v) CH2 Cl2 /acetone (300 mL). Collect 10-mL fractions. 28. Monitor fractions by TLC (step 8) using 90:10 (v/v) CH2 Cl2 /acetone as the eluent. 29. Pool all product fractions, evaporate the solvent using a rotary evaporator, and dry the resulting sample overnight under high vacuum. 7-[2-Deoxy-5-O-(4,4 -dimethoxytriphenylmethyl)-β-D-erythro-pentofuranosyl]-4-[(2methylpropanoyl)amino]-7H-pyrrolo[2,3-d]pyrimidin-2-yl diphenylcarbamate (S.18) is obtained as a colorless foam (622 mg, 75% yield). TLC (CH2 Cl2 /acetone, 9:1): Rf = 0.70. UV (MeOH): λmax (ε) = 221 (55,700), 233 (56,100), 294 nm (7,600). 1 H NMR (DMSO-d6 ): δ 1.13 and 1.15 (2 × s, 6H, 2 Me), 2.30 (m, 2H, 2 -H), 2.88 (m, 1H, CH), 3.16 (m, 2H, 5 -H), 3.72 (s, 6H, 2 MeO), 3.95 (m, 1H, 4 -H), 4.38 (m, 1H, 3 -H), 5.37 (d, J = 4.1 Hz, 1H, 3 -OH), 6.58 (t, J = 6.3 Hz, 1H, 1 -H), 6.58 (d, J = 4.0 Hz, 1H, 5-H), 7.22-7.46 (m, 24H, 6-H and aromatic H), 11.0 (s, 1H, NH). Anal. calcd. for C28 H29 N5 O6 : C, 70.57; H, 5.68; N, 8.40; found: C, 70.29; H, 5.65; N, 8.33.
Prepare phosphoramidite (S.6a) 30. In a 50-mL round-bottom flask equipped with a Teflon stir bar and kept under argon, prepare a solution of 320 mg (0.38 mmol) compound 18 in 11.0 mL anhydrous CH2 Cl2 .
Using Pyrrolo[2,3d]pyrimidines to Replace Purines
31. Add 104 µL (0.60 mmol) anhydrous N,N-diisopropylethylamine and 120 µL (0.54 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite and stir the resulting mixture 30 min at room temperature under argon.
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32. Transfer the reaction mixture to a 100-mL separatory funnel and dilute with 30 mL CH2 Cl2 . Wash twice, each time with 20 mL of 5% NaHCO3 , wash once with 20 mL brine, and then dry over 3 g Na2 SO4 . 33. Filter off the Na2 SO4 by gravity filtration through a folded 10-cm-diameter filter. Evaporate the filtrate to dryness using a rotary evaporator. 34. Prewash a 4 × 70–cm silica gel 60 column (bed height, 10 cm) with 200 mL CH2 Cl2 containing 1% (v/v) anhydrous Et3 N and then with 100 mL of 100% CH2 Cl2 . 35. Dissolve the crude product in ∼2 mL CH2 Cl2 , load it onto the column, and perform flash chromatography (step 7) using 93:7 (v/v) CH2 Cl2 /acetone (400 mL) as the eluent. Collect 10-mL fractions. 36. Monitor fractions by TLC (step 8) using 93:7 (v/v) CH2 Cl2 /acetone as the eluent. 37. Pool all product fractions and evaporate the solvent using a rotary evaporator. 38. Dissolve in a small volume (∼2 mL) of CH2 Cl2 . Transfer this solution to a 25-mL round-bottom flask, evaporate the solvent using a rotary evaporator, and dry the resulting sample under high vacuum for 24 hr at 40◦ C. 7-[2-Deoxy-5-O-(4,4 -dimethoxytriphenylmethyl)-β-D-erythro-pentofuranosyl]-4-[(2methylpropanoyl)amino]-7H-pyrrolo[2,3-d]pyrimidin-2-yl diphenylcarbamate 3 -[(2cyanoethyl)-N,N-diisopropylphosphoramidite] (S.6a) is obtained as a colorless foam (337 mg, 84% yield). TLC (CH2 Cl2 /acetone, 9:1): Rf = 0.8. 31 P NMR (CDCl3 ): δ 149.7, 149.9.
39. If the synthesized phosphoramidite still contains impurities, remove those impurities in the following way. a. In a 100-mL round-bottom flask equipped with a Teflon stir bar, dissolve 0.3 to 1.0 g of the synthesized phosphoramidite in 1 to 2 mL CH2 Cl2 . b. Add ∼90 mL cyclohexane dropwise, with continuous shaking or stirring to dislodge any solid stuck to the walls of the flask. c. Refrigerate overnight. d. Remove the supernatant from the flask and dissolve the residue in 5 mL CH2 Cl2 . e. Transfer the contents of the flask to a 25-mL round-bottom flask, evaporate the solvent using a rotary evaporator, and dry the resulting sample under high vacuum for 24 hr at 40◦ C.
SYNTHESIS AND PURIFICATION OF OLIGONUCLEOTIDES CONTAINING 7-DEAZA-2 -DEOXYRIBONUCLEOSIDES
BASIC PROTOCOL 4
This protocol describes the preparation and purification of oligonucleotides containing the 7-deaza-2 -deoxyribonucleosides shown in Figure 4.25.5. The oligonucleotides 20 to 58 (shown in Tables 4.25.1, 4.25.2, 4.25.3, 4.25.4, and 4.25.5) are prepared via phosphoramidite chemistry on a 1-µmol scale using standard reaction cycles on an automated DNA synthesizer (ABI 392-08; Applied Biosystems) operating in DMTr-on mode. The phosphoramidites S.1 to S.8 (Fig. 4.25.1) and other standard phosphoramidites are employed. The average coupling yield of the base-modified phosphoramidites is >95% in all cases. Oligonucleotides synthesized using this protocol are purified by reversed-phase HPLC (RP-HPLC), and composition analysis is subsequently performed. Product masses are determined by matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrometry (UNIT 10.1). The preparation of the oligonucleotide 5 -d(ATiC iCA15a
Synthesis of Modified Oligonucleotides and Conjugates
4.25.13 Current Protocols in Nucleic Acid Chemistry
Supplement 20
Figure 4.25.5 Base-modified nucleosides incorporated into the oligonucleotides shown in Tables 4.25.1, 4.25.2, 4.25.3, 4.25.4, and 4.25.5.
TTA T15aA)-3 (53; where iC = m5 iCd = 5-methyl-2 -deoxyisocytidine) is described below as a typical example (Table 4.25.5). The phosphoramidite S.6a was employed for the incorporation of S.15a. The synthesis and purification of oligonucleotides 20 through 52 and 54 through 58 are performed in a similar manner using the appropriate phosphoramidites. After standard cleavage from the solid support and deprotection of the cyanoethyl groups in 25% NH4 OH for 1 hr at room temperature, the oligonucleotides 53, 54, 55, 57, and 58 (all containing 2 -deoxyisoguanosine analogs) are base-deprotected by heating the ammonia solution to 60◦ C for 20 to 24 hr. (The standard base deprotection procedure calls for heating of the solution to 60◦ C for 16 hr.) The authors confirmed the homogeneity of each synthesized oligonucleotide by ion-exchange chromatography on a NucleoPac-PA-100 column (4 × 50 cm; Dionex).
Materials
Using Pyrrolo[2,3d]pyrimidines to Replace Purines
Phosphoramidite solutions (0.1 M in MeCN): 2 -deoxyadenylate, 2 -deoxythymidylate, 5-methyl-2 -deoxyisocytidylate, and 7-deaza-2 -deoxyisoguanylate (S.6a; see Basic Protocol 3) 25% (v/v) aqueous ammonium hydroxide (NH4 OH) Mobile phase A: acetonitrile (MeCN) Mobile phase B: 95:5 (v/v) 0.1 M TEAA buffer (pH 7.0)/acetonitrile (see recipe) 2.5% (v/v) dichloroacetic acid (Cl2 CHCOOH) in anhydrous dichloromethane (CH2 Cl2 ) Anhydrous triethylamine (Et3 N)
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Current Protocols in Nucleic Acid Chemistry
Methanol (MeOH) 0.1 M Tris·Cl, pH 8.9 (see recipe) 36.2 U/mL snake venom phosphodiesterase I (SVPD; EC 3.1.4.1; Crotalus adamanteus; Amersham) 1 U/µL alkaline phosphatase (EC 3.1.3.1; calf intestine; Roche Diagnostics) ABI 392-08 automated DNA synthesizer (Applied Biosystems) SpeedVac evaporator (Savant) HPLC apparatus, including: 4 × 125–mm and 4 × 250–mm RP-18 LiChrospher columns (Merck) 655A-12 HPLC pump (Merck/Hitachi) 655A variable-wavelength monitor (Merck/Hitachi) L-5000 controller unit (Merck/Hitachi) D-2000 chromatogram peak integrator (Merck/Hitachi) 5-mL polypropylene tubes 1-mL quartz cuvette (1-cm path length) UV-Vis spectrophotometer Additional reagents and equipment for automated oligonucleotide synthesis (APPENDIX 3C) using phosphoramidite chemistry (UNIT 3.3), reversed-phase high-performance liquid chromatography (RP-HPLC; UNIT 10.5), and matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF-MS; UNIT 10.1) NOTE: Phosphoramidites should be dried under high vacuum (<1 mmHg) for 20 to 24 hr at 40◦ C before being applied to the DNA synthesizer.
Synthesize and purify 5 -DMTr-protected oligonucleotide 1. Using phosphoramidite chemistry, prepare the oligonucleotide 5 -DMTr-53 on a 1-µmol scale in an ABI 392-08 synthesizer operating in DMTr-on mode (also see UNIT 3.3 and APPENDIX 3C). The oligonucleotide is automatically cleaved from the solid support by treatment for 1 hr with 25% (v/v) NH4 OH, and the cyanoethyl groups are removed from the oligonucleotide (at room temperature) as well.
2. Base-deprotect the oligonucleotide by heating to 60◦ C in 2 mL of 25% NH4 OH for 20 to 24 hr. 3. Evaporate the 5 -DMTr-53 solution to dryness in a SpeedVac evaporator. Dissolve the resulting sample in 500 µL H2 O. 4. Purify by RP-HPLC (UNIT 10.5) on a 4 × 250–mm RP-18 column with 260-nm detection, using the following elution gradient at a flow rate of 1.0 mL/min: 0-3 min 3-15 min 15-20 min
10% to 15% mobile phase B 15% to 50% mobile phase B 50% to 10% mobile phase B
Collect the desired fraction (retention time, ∼12 min) in a 5-mL polypropylene tube. 5. Evaporate the collected fraction to dryness in a SpeedVac evaporator.
Detritylate and purify 5 -OH oligonucleotide 6. Treat purified 5 -DMTr-53 with 200 µL of 2.5% Cl2 CHCOOH in CH2 Cl2 for 5 min at room temperature. 7. Neutralize the sample with 100 µL Et3 N and evaporate to dryness in a SpeedVac evaporator. Dissolve in 500 µL H2 O.
Synthesis of Modified Oligonucleotides and Conjugates
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8. Purify by RP-HPLC (step 4) using the following gradient: 0-25 min 25-30 min 30-35 min
0% to 20% mobile phase B 20% mobile phase B 20% to 0% mobile phase B
Collect the desired fraction (retention time, ∼15 min) in a 5-mL polypropylene tube. 9. Evaporate the collected fraction to dryness in a SpeedVac evaporator. Dissolve the resulting sample in 500 µL H2 O.
Desalt purified oligonucleotide 10. Load the sample onto a 4 × 125–mm RP-18 column and desalt by performing RP-HPLC using H2 O to elute the salt (1 hr at 0.7 mL/min) followed by 3:2 (v/v) MeOH/H2 O to elute the oligonucleotide (∼3 to 5 min at 0.7 mL/min). Collect the oligonucleotide fraction in a 5-mL polypropylene tube. 11. Evaporate to dryness in a SpeedVac evaporator. Dissolve the resulting sample in 100 µL H2 O and store frozen at –18◦ C. As an alternative to steps 6 to 11, the oligonucleotide can be obtained as a sodium salt by (1) treating the evaporated fraction from step 5 with 250 µL of 80% (v/v) acetic acid for 30 min at room temperature; (2) evaporating the resulting solution to near-dryness using a SpeedVac evaporator (without heating); (3) adding 350 µL of 1 M NaCl; and (4) adding 1.1 mL ethanol and cooling in an ice bath for 15 min to precipitate the product. The precipitated oligonucleotide can then be collected by centrifugation for 15 min at 12,000 rpm, ∼10◦ C, with the supernatant being saved until an optical density measurement can be made.
Determine product yield 12. Add 2 µL of the product solution to 1 mL H2 O in a 1-mL quartz cuvette. Measure the optical density of the sample at 260 nm (OD260 ) in a UV-Vis spectrophotometer, using H2 O as a reference. Calculate the total amount of oligonucleotide synthesized by multiplying the measured OD260 /mL by the dilution factor (500) and the sample volume (0.1 mL). For 2 µL product in 1 mL H2 O, the authors measured an OD260 of 0.64, corresponding to a total yield of 0.64 OD260 /mL × 500 × 0.1 mL = 32 OD260 units.
13. Convert the calculated yield from OD units to mmol using the equation: yield (mmol) = yield (OD260 units)/[0.9 × ε260 (M−1 · cm−1 )]. The molar extinction coefficients (ε 260 ) of the nucleosides used to make oligonucleotide 53 are as follows: c7 iGd (S.15a), 7,400; dA, 15,400; dT, 8,800; m5 iCd , 6,300. Each coefficient is multiplied by the number of residues in the sequence. Thus, the total amount of oligonucleotide 53 obtained in this example is 32/[0.9 × (2 × 7400 + 2 × 6300 + 4 × 15400 + 4 × 8800)] = 0.29 × 10−3 mmol, corresponding to a yield of 29%.
Analyze oligonucleotide composition 14. Dissolve 0.5 OD260 units (4.5 nmol) of product in 300 µL of 0.1 M Tris·Cl, pH 8.9. 15. Add 5 µL SVPD and incubate 1 hr at 37◦ C. Snake venom phosphodiesterase I cleaves the phosphodiester bonds of the oligonucleotide from the 3 direction to give 5 monophosphates.
16. Add 5 µL alkaline phosphatase and incubate 1 hr at 37◦ C. Using Pyrrolo[2,3d]pyrimidines to Replace Purines
Alkaline phosphatase cleaves the 5 monophosphates to give free nucleosides.
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Current Protocols in Nucleic Acid Chemistry
17. Load the mixture onto a 4 × 250–mm RP-18 column and perform RP-HPLC using the following gradient at a flow rate of 0.7 mL/min: 0-20 min 20-40 min 40-60 min
100% mobile phase B 100% to 35% mobile phase B 35% to 100% mobile phase B.
To provide a reference for identifying peaks in the chromatogram yielded by the experimental mixture, chromatograms of the purified monomeric nucleosides are also obtained.
18. Calculate the relative amount of each constituent in the product solution by dividing the peak area by the molar extinction coefficient. Relevant molar extinction coefficients (λ = 260 nm) for analyzing product composition include the following: c7 iGd , 7,400; Cl7 c7 iGd , 6,350; Br7 c7 iGd , 5,840; iGd , 4,300; dA, 15,400; dT, 8,800; dG, 11,700; dC, 7,600; m5 iCd , 6,300.
19. Determine the molecular mass of the purified product by MALDI-TOF-MS (e.g., on a Bruker Biflex-III spectrometer operating in reflector mode; see UNIT 10.1). The observed mass of oligonucleotide 53 (3670.3 Da) is in good agreement with the calculated mass (3669.5 Da).
DETERMINATION OF TRANSITION (MELTING) TEMPERATURE AND THERMODYNAMIC DATA
BASIC PROTOCOL 5
The Tm values of duplexes containing the base-modified nucleosides S.9a (Seela and Kehne, 1983), S.9b,c,i (Seela and Thomas, 1994), S.9d,f (Seela and Zulauf, 1996), S.9j (F. Seela and K. Xu, unpub. observ.), S.12a-d,f,i (Winkeler and Seela, 1983; Ramzaeva and Seela, 1995; Seela and Chen, 1997), S.12e (Buhr et al., 1996), S.15a-c (Seela and Peng, 2004), and S.19a-d (Seela et al., 1987; Seela and Peng, 2004) were determined from absorbance-temperature profiles. Tm values (Tables 4.25.1, 4.25.2, 4.25.3, 4.25.4, and 4.25.5) were measured on a Cary-1/3 UV-Vis spectrophotometer (Varian) equipped with a thermoelectric controller. The temperature was increased by 1.0◦ C/min from 10◦ to 95◦ C, and the melting profile was monitored at 260 nm. UV absorbance was recorded every 30 sec, and Tm values were determined based on polynomial fits of the resulting UV absorbance curves. The temperature at which the first derivative of the UV melting curve was at a maximum was taken to be the Tm . Thermodynamic data were calculated using the program Meltwin 3.0 (McDowell and Turner, 1996). All experiments were performed in low-salt or high-salt buffer, pH 7.0 (see recipes).
Oligonucleotides Containing the 2 -Deoxyadenosine Analogs S.9a-d,f,i From Table 4.25.1, it is apparent that the replacement of the standard dA residue with the 7-deaza compound S.9a has only a minor influence on duplex stability (Tm = 0.25◦ C per modified base for duplex 21·21 versus 20·20). Incorporation of the 7-methylated nucleoside S.9i results in a slight increase in stability (Tm = 0.65◦ C per modified base for 25·25). However, when nucleosides containing 7-halogeno substituents (S.9b-d) or a 7-hexynyl group (S.9f) are introduced, Tm values increase significantly (Tm = 2◦ C per modified base for S.9b-c, 2.15◦ C for S.9d, and 1.55◦ C for S.9f). Oligonucleotides Containing the 2,6-Diaminopurine Derivatives S.19a-d A very similar situation is observed in the case of the 2,6-diamino derivatives S.19ad (Table 4.25.2). The replacement of the dA residue with the nucleoside S.19a (R = H) has no influence on duplex stability (duplex 29·30), while the incorporation of the 7-halogenated derivatives S.19b-d significantly increases the Tm value (Tm = 2.3◦ to 3.0◦ C per modified base; see duplexes 31·32, 33·34, and 35·36). Furthermore, compounds S.19a and S.19d result in higher Tm values compared with the
Synthesis of Modified Oligonucleotides and Conjugates
4.25.17 Current Protocols in Nucleic Acid Chemistry
Supplement 20
Table 4.25.1 Tm Values and Thermodynamic Data for Self-Complementary Oligonucleotides Containing 2 -Deoxyadenosine Analogs S.9a-d,f,ia
Duplex
Tm (◦ C)
[d(A-T)6 ]2 (20·20)
33
Tm b (◦ C)
H◦ (kcal/mol)
S◦ (cal/mol · K)
G◦ 310 (kcal/mol)
−45
−125
−6.3
36
+0.25
−65
−186
−6.8
c
57
+2
−60
−159
−10.6
c
57
+2
−62
−165
−10.9
[d(9d-T)6 ]2 (24·24)
59
+2.15
−65
−174
−11.4
c
41
+0.65
−45
−142
−8.0
52
+1.55
−47
−123
−9.3
[d(9a-T)6 ]2 (21·21) [d(9b-T)6 ]2 (22·22) [d(9c-T)6 ]2 (23·23) [d(9i-T)6 ]2 (25·25)
[d(9f-T)6 ]2 (26·26)
a Thermodynamic parameters are derived from the fitting of melting curves measured at 260 nm (see Basic Protocol 5).
In all assays, the concentration of each oligonucleotide strand was 5 µM in high-salt buffer (1.0 M NaCl), pH 7.0 (see recipe). b Change (per modified base) relative to duplex 20·20. c See Seela and Thomas (1995).
Table 4.25.2 Tm Values and Thermodynamic Data for Oligonucleotide Duplexes Containing S.19a-d or S.9a,d,ja
H◦ (kcal/mol)
S◦ (cal/mol · K)
G◦ 310 (kcal/mol)
−90
−257
−10.9
0
−80
−225
−10.4
55
+2.7
−92
−256
−12.9
5 -d(TAG GTC 19cAT ACT)-3 (33) 3 -d(ATC C19cG TT19c TGA)-5 (34)
56
+3.0
−95
−265
−13.4
5 -d(TAG GTC 19dAT ACT)-3 (35) 3 -d(ATC C19dG TT19d TGA)-5 (36)
54
+2.3
−83
−227
−12.3
5 -d(TAG GTC 9aAT ACT)-3 (37) 3 -d(ATC C9aG TT9a TGA)-5 (38)
44
−1.0
−81
−229
−9.9
5 -d(TAG GTC 9dAT ACT)-3 (39) 3 -d(ATC C9dG TT9d TGA)-5 (40)
51
+1.3
−89
−249
−11.7
5 -d(TAG GTC 9jAT ACT)-3 (39a) 3 -d(ATC C9jG TT9j TGA)-5 (40a)
49
+0.7
−81
−227
−11.1
Duplex
Tm (◦ C)
5 -d(TAG GTC AAT ACT)-3 (27) 3 -d(ATC CAG TTA TGA)-5 (28)
47
5 -d(TAG GTC 19aAT ACT)-3 (29) 3 -d(ATC C19aG TT19a TGA)-5 (30)
47
5 -d(TAG GTC 19bAT ACT)-3 (31) 3 -d(ATC C19bG TT19b TGA)-5 (32)
Tm b (◦ C)
a Thermodynamic parameters are derived from the fitting of melting curves measured at 260 nm (see Basic Protocol 5). In all assays, the concentration
of each oligonucleotide strand was 5 µM in low-salt buffer (0.1 M NaCl), pH 7.0 (see recipe). b Change (per modified base) relative to duplex 27·28.
corresponding nucleosides lacking the 2-amino group (S.9a and S.9d, respectively). In contrast to other halogenated derivatives, the incorporation of 7-deaza-2 -deoxyadenosine carrying a fluorine substituent at the 7 position (S.9j) stabilizes oligonucleotide duplexes only to a small extent.
Using Pyrrolo[2,3d]pyrimidines to Replace Purines
Hybridization experiments involving the pairing of S.9a or S.19a with each of the four canonical nucleosides (Table 4.25.3) show that nucleoside S.19a forms a rather stable base pair with dC (duplex 44·41; Okamoto et al., 2002), while S.9a does not (duplex 44·45). This indicates that the additional 2-amino group of S.19a plays a crucial role in
4.25.18 Supplement 20
Current Protocols in Nucleic Acid Chemistry
Table 4.25.3 Tm Values and Thermodynamic Data for Oligonucleotide Duplexes Containing S.9a or S.19a Opposite a Canonical Nucleosidea
H◦ (kcal/mol)
S◦ (cal/mol · K)
G◦ 310 (kcal/mol)
−90
−257
−10.9
0
−89
−252
−10.8
36
−11
−65
−185
−7.67
5 -d(TAG GGC AAT ACT)-3 (43) 3 -d(ATC C19aG TTA TGA)-5 (41)
45
−2
−87
−249
−10.1
5 -d(TAG GCC AAT ACT)-3 (44) 3 -d(ATC C19aG TTA TGA)-5 (41)
46
−1
−93
−266
−10.5
5 -d(TAG GTC AAT ACT)-3 (27) 3 -d(ATC C9aG TTA TGA)-5 (45)
46
−1
−86
−245
−10.4
5 -d(TAG GAC AAT ACT)-3 (41) 3 -d(ATC C9aG TTA TGA)-5 (45)
33
−14
−68
−196
−7.1
5 -d(TAG GGC AAT ACT)-3 (43) 3 -d(ATC C9aG TTA TGA)-5 (45)
45
−2
−85
−241
−10.1
5 -d(TAG GCC AAT ACT)-3 (44) 3 -d(ATC C9aG TTA TGA)-5 (45)
43
−4
−85
−243
−9.6
Duplex
Tm (◦ C)
5 -d(TAG GTC AAT ACT)-3 (27) 3 -d(ATC CAG TTA TGA)-5 (28)
47
5 -d(TAG GTC AAT ACT)-3 (27) 3 -d(ATC C19aG TTA TGA)-5 (41)
47
5 -d(TAG GAC AAT ACT)-3 (42) 3 -d(ATC C19aG TTA TGA)-5 (41)
Tm b (◦ C)
a Thermodynamic parameters are derived from the fitting of melting curves measured at 260 nm (see Basic Protocol 5). In all assays, the concentration
of each oligonucleotide strand was 5 µM in low-salt buffer (0.1 M NaCl), pH 7.0 (see recipe). b Change (per modified base) relative to duplex 27·28.
the observed pairing with dC; a bidentate base pair is suggested as the interaction between S.19a and dC (see Fig. 4.25.6, motif IIIa).
Oligonucleotides Containing the 2 -Deoxyguanosine Analogs S.12a,c-f,i Compared with dA analogs and 2,6-diaminopurine derivatives, the situation is quite different for the 7-deaza analogs of dG (Table 4.25.4). The incorporation of 7-deaza-2 deoxyguanosine (S.12a) in place of dG results in duplex destabilization compared with d(G-C)4 (46). The 7-methyl derivative S.12i and the 7-hexynyl nucleoside S.12f lead to duplex stabilization compared with the parent nucleoside S.12a (further destabilization compared with dG). When 7-bromo or 7-iodo derivatives (S.12c or S.12d) are incorporated, a further increase in the Tm value is observed. The increase in Tm amounts to ∼1◦ C per modified base in each case (octamers 48 and 49). Like the 7-halogeno derivatives, the 7-propynylated nucleoside S.12e also leads to duplex stabilization (27b·28b), as the small propynyl group is well accommodated in the major groove of DNA. Oligonucleotides Containing the 2 -Deoxyisoguanosine Analogs S.15a-c The influence of the 7-deazaisoguanosine series (S.15a-c) on oligonucleotide stability was studied with a parallel duplex (52·27) and an antiparallel duplex (56·52; Table 4.25.5). The 7-deazaisoguanine-containing parallel duplex (53·27) shows almost identical base pair stability compared with the duplex containing the parent isoguanine. However, the incorporation of chlorinated (S.15b) or brominated (S.15c) derivatives increases Tm by ∼2.0◦ C per modified base for both parallel and antiparallel duplexes. Motifs for the base-modified base pairs related to dG-dC, c7 iGd -iCd , and c7 iGd -dC are depicted in Figure 4.25.6.
Synthesis of Modified Oligonucleotides and Conjugates
4.25.19 Current Protocols in Nucleic Acid Chemistry
Supplement 20
Figure 4.25.6
Base pair motifs for 7-substituted 7-deazapurines paired with 2 -deoxythymidine or 2 -deoxycytidine.
Table 4.25.4 Tm Values and Thermodynamic Data for Self-Complementary Oligonucleotides Containing 2 -Deoxyguanosine Analogs S.12a,c-f,ia
H◦ (kcal/mol)
S◦ (cal/mol · K)
−81
−244
−0.88
−62
−190
67
+0.88
−72
−212
[d(12d-C)4 ]2 (49·49)
70
+1.25
−91
−266
[d(12i-C)4 ]2 (50·50)
58
−0.25
−82
−250
59
−0.13
−62
−189
−90
−257
Duplex
Tm (◦ C)
[d(G-C)4 ]2 (46·46)
60
[d(12a-C)4 ]2 (47·47)
53
[d(12c-C)4 ]2 (48·48)
[d(12f-C)4 ]2 (51·51)
Tm b (◦ C)
5 -d(TAG GTC AAT ACT)-3 (27) 3 -d(ATC CAG TTA TGA)-5 (28)
47
5 -d(TA12a 12aTC AAT ACT)-3 (27a) 3 -d(ATC CA12a TT12a TGA)-5 (28a)
44
−0.38
−104
−328
5 -d(TA12e 12eTC AAT ACT)-3 (27b) 3 -d(ATC CA12e TTA T12eA)-5 (28b)
55
+1.0
−91
−251
a Thermodynamic parameters are derived from the fitting of melting curves measured at 260 nm (see Basic Protocol 5).
Using Pyrrolo[2,3d]pyrimidines to Replace Purines
In all assays, the concentration of each oligonucleotide strand was 7.5 µM in low-salt buffer (0.1 M NaCl), pH 7.0 (see recipe). G data are unavailable. b Change (per modified base) relative to duplex 46·46 or 27·28.
4.25.20 Supplement 20
Current Protocols in Nucleic Acid Chemistry
Table 4.25.5 Tm Values and Thermodynamic Data for Duplexes Containing 2 -Deoxyisoguanosine Analogs S.15a-ca
H◦ (kcal/mol)
S◦ (cal/mol · K)
G◦ 310 (kcal/mol)
−74
−211
−8.8
0
−51
−135
−8.7
43
+2.0
−79
−225
−9.5
5 -d(ATiC iCA15c TTA T15cA)-3 (55) 5 -d(TAG GTC AAT ACT)-3 (27)
44
+2.5
−74
−208
−9.5
3 -d(TAiG iGTiC AAT AiCT)-5 (56) 5 -d(ATiC iCAiG TTA TiGA)-3 (52)
60
−94
−257
−14.8
3 -d(TA15b 15bTiC AAT AiCT)-5 (57) 5 -d(ATiC iCA15b TTA T15bA)-3 (54)
67
+1.8
−96
−255
−16.4
3 -d(TA15c 15cTiC AAT AiCT)-5 (58) 5 -d(ATiC iCA15c TTA T15cA)-3 (55)
68
+2.0
−83
−175
−14
Duplexb
Tm (◦ C)
5 -d(ATiC iCAiG TTA TiGA)-3 (52) 5 -d(TAG GTC AAT ACT)-3 (27)
39
5 -d(ATiC iCA15a TTA T15aA)-3 (53) 5 -d(TAG GTC AAT ACT)-3 (27)
39
5 -d(ATiC iCA15b TTA T15bA)-3 (54) 5 -d(TAG GTC AAT ACT)-3 (27)
Tm c (◦ C)
a Thermodynamic parameters are derived from the fitting of melting curves measured at 260 nm (see Basic Protocol 5). In all assays, the concentration
of each oligonucleotide strand was 5 µM in high-salt buffer (1.0 M NaCl), pH 7.0 (see recipe). b iC, 5-methyl-2 -deoxyisocytidine; iG, 2 -deoxyisoguanosine. c Change (per modified base) relative to duplex 52·27 or duplex 56·52.
Conclusion From the data shown above, it is apparent that the incorporation of 7-substituted 7deazapurine nucleosides into oligonucleotide duplexes leads to an increase in duplex stability, as indicated by the observed Tm values. Base pair stabilization was found to result from the increased stacking of the nucleobases, the hydrophobic character of the substituents at the 7 position, and the increased hydrogen bond strength, which is related to the pKa values of the nucleosides (see UNIT 1.10). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
High-salt buffer (1.0 M NaCl), pH 7.0 To 900 mL H2 O, add: 58.5 g NaCl (1 M final) 9.5 g MgCl2 (100 mM final) 12.8 g (CH3 )2 AsO2 Na · H2 O (60 mM final) Adjust pH to 7.0 with 2 M HCl Add water to 1 L Store tightly capped for up to 2 years at room temperature Low-salt buffer (0.1 M NaCl), pH 7.0 To 900 mL H2 O, add: 5.85 g NaCl (0.1 M final) 0.95 g MgCl2 (10 mM final) 2.14 g (CH3 )2 AsO2 Na · H2 O (10 mM final) Adjust pH to 7.0 with 2 M HCl Add water to 1 L Store tightly capped for up to 2 years at room temperature
Synthesis of Modified Oligonucleotides and Conjugates
4.25.21 Current Protocols in Nucleic Acid Chemistry
Supplement 20
Triethylammonium acetate (TEAA) buffer, 0.1 M (pH 7.0)/acetonitrile, 95:5 (v/v) To 900 mL H2 O, add 14.0 mL triethylamine. Adjust the pH to between 7 and 7.5 with 90% (v/v) acetic acid, add water to 1.0 L, and then filter through RC membrane filter paper (pore size, 0.45 µm; diameter, 0.47 mm; Schleicher & Schuell). Add 50 mL acetonitrile to 950 mL of the filtrate and store tightly capped for up to 4 months at room temperature. Tris·Cl, pH 8.9 To 90 mL H2 O, add 1.21 g tris(hydroxymethyl)methylamine [(HOCH2 )3 CNH2 ] (0.1 M final). Adjust pH to 8.9 with 20% (v/v) HCl, and then add water to 100 mL. Store tightly capped for up to 2 years at 5◦ C. COMMENTARY Background Information
Using Pyrrolo[2,3d]pyrimidines to Replace Purines
7-Deazapurine-containing oligonucleotides have been used to inhibit gene expression (antisense oligonucleotides; Balow et al., 1998), and 7-deazapurine nucleosides have been used in DNA and RNA sequencing with labeled 2 -dNTPs and in 2 ,3 -ddNTPmediated termination of chain extension (Mizusawa et al., 1986). Moreover, 7 deazapurine nucleoside residues have been introduced into nucleic acids for studies of electron transfer (Latimer and Lee, 1991; Li et al., 2004). As N7 is absent in these basemodified oligonucleotides, neither dG quartet formation (Seela and Mersmann, 1993) nor the generation of mini-hairpins occurs, thereby preventing band compression during DNA or RNA sequencing (Barr et al., 1986). On the other hand, special supramolecular assemblies (Seela and Wei, 1997) and nanostructures (Seela et al., 2002) can be constructed with the help of 7-dezazapurine-containing DNA, with these structures showing superior properties compared with those containing regular purine bases. The 7 position of a 7-deazapurine is an ideal site for introducing reporter groups or (to enhance DNA duplex stability) introducing electron-withdrawing substituents of medium size (Ramzaeva and Seela, 1996; Seela and Zulauf, 1998). Such substituents (in particular, halogen or alkynyl substituents) are well accommodated in the major groove of duplex DNA, and some have the potential to be used for further chemical and functional manipulations of the oligonucleotides. The 7-iodo-substituted derivatives of 2 -deoxyadenosine (S.2d and S.9d) and 2 -deoxyguanosine (S.3d and S.12d) are particularly valuable, since they are readily alkynylated via Pd-catalyzed cross-coupling reactions. Alternatively, DNA modifications can be performed using
solid-phase methods that are applied during or after automated solid-phase synthesis. Over the past decade, efforts have also been made to modify canonical DNA by replacing purine bases with 7-deazapurines to improve binding of oligonucleotides to their DNA and RNA targets. The design and synthesis of modified oligonucleotides with increased duplex stability are of particular interest since improvements in antisense effects have been correlated with increases in Tm values. Efforts to investigate oligonucleotides containing 7-deazapurines, including the design and synthesis of appropriately protected phosphoramidites, have been undertaken in the authors’ laboratory for a number of years. Access to these systems became realistic with the development of stereoselective nucleobase anion glycosylation, which was first realized using 7-deazapurine-2 -deoxyribonucleosides (Winkeler and Seela, 1983; Seela et al., 1988). Because 7-deazapurines (pyrrolo[2,3d]pyrimidines) closely resemble purines in structure, they are ideal shape mimics of the canonical purine bases in nucleic acids. 7Deazapurine nucleosides show extraordinary stability in their N-glycosylic bonds, which makes them stable against acid- or enzymecatalyzed hydrolysis. On the other hand, 7deazpurines are more easily oxidized than purines. The syntheses of the phosphoramidites S.1-S.8 (Fig. 4.25.1) use well-established chemistry. Dialkylaminoalkylidene protection of the exocyclic amino groups is readily achieved by reaction with the corresponding acetal reagents, while acyl protection of the exocyclic amino groups is achieved using acyl chlorides or acid anhydrides. Protection of the 2-oxo group of 7-deaza-2 -deoxyisoguanosine is performed by diphenylcarbamoylation of S.15a-c with
4.25.22 Supplement 20
Current Protocols in Nucleic Acid Chemistry
Table 4.25.6
13
C NMR Chemical Shifts (δ) for 7-Deazapurine-2 -Deoxyribonucleosidesa,b
Sys Pur
C(2)c C(2)
9d
152.0 157.3 103.2
10
C(4)c C(6)
C(4a) C(5)
151.5 160.2 110.3
11
151.2 160.2 110.3
12d
152.7 157.7
C(5) C(7)
C(6) C(7a)c C(1 ) C(8) C(4)
51.9 126.9 149.8 53.6 128.6 150.9
C(2 )
C(3 )
C(4 ) C(5 )
83.0
—d
71.0
87.5
62.0
83.0
d
71.1
87.5
61.9
d
—
53.9 128.3 151.0
82.8
—
70.9
85.6
64.3
99.8 102.2 121.5 150.5
82.2
38.5
70.9
87.0
61.9
d
13
147.5 156.1 103.9
55.5 124.3 147.0
82.6
—
70.8
87.3
61.7
14
147.6 156.1 104.2
55.0 124.1 147.1
82.7
—d
70.6
85.5
64.1
15a
153.9 156.3
92.6 100.8 118.9 152.6
83.4
—d
71.1
87.2
62.0
16
158.8 155.8 100.9 100.1 121.6 150.7
82.7
—d
71.0
87.2
62.0
17
154.3 153.6 106.3 104.4 119.5 152.5
82.6
—d
70.9
87.4
61.9
18
158.0 153.6 106.2 104.6 123.8 151.4
82.2
—d
70.6
85.5
64.0
a Sys, carbon position in systematic numbering scheme; Pur, carbon position in purine numbering scheme. b Measured in ppm downfield from DMSO-d . 6 c Peak assignments are tentative. d Superimposed upon DMSO-d peak. 6
diphenylcarbamoyl chloride in pyridine. Dimethoxytritylation of 5 -hydroxy groups is achieved using a standard protocol, which always results in high yields. Protected nucleosides are easily converted to the desired phosphoramidites by treatment with 2-cyanoethylN,N-diisopropylchlorophosphoramidite. DNA syntheses using phosphoramidites S.1-S.8 employ conventional phosphoramidite chemistry. The coupling yield is always >95%. Purification of the resulting oligonucleotides is performed in DMTr-on and/or DMTr-off mode using an RP-18 column. The Tm values of oligonucleotide duplexes are determined by UV spectrophotometry, and thermodynamic data are determined by curve shape analysis. The duplex stability of 7-deazapurinecontaining DNA can then be compared with that of the corresponding unmodified DNA duplex. 7-Halogenated and 7-alkynylated 7-deazapurines stabilize DNA duplexes that have antiparallel or parallel chain orientation.
Compound Characterization 1
H, 13 C, and 31 P NMR spectra were obtained on AC-250 and AMX-500 spectrometers (Bruker), with δ values given in ppm downfield from internal tetramethylsilane (SiMe4 ) or external 85% (v/v) H3 PO4 solution (for 31 P NMR). UV spectra were measured on a U-3200 spectrometer (Hitachi; λmax in nm, ε in M−1 · cm−1 ). Melting points were determined using a Linstr¨om apparatus and are
not corrected. Elemental analyses were performed by the Mikroanalytisches Laboratorium Beller (G¨ottingen, Germany). Elemental analyses, UV data, 1 H and 31 P NMR data, and melting points are shown in the appropriate protocol steps. 13 C NMR data are shown in Table 4.25.6.
Critical Parameters Phosphoramidites are sensitive to hydrolysis when acid is present, but they are stable in neutral solution. Their purity can be checked in neutral aqueous buffer by HPLC. The 7deazapurine nucleosides, including their phosphoramidites, are more easily oxidized than are their parent purine compounds. However, all oligonucleotide syntheses have been performed using a standard oxidation reagent (0.02 or 0.1 M iodine in 90:5:5 [v/v/v] THF/pyridine/H2 O), and significant degradation has not been observed. The success of the DNA synthesis depends on the quality of the reagents; only reagent-grade chemicals should be used to synthesize the phosphoramidites described here. All phosphoramidite reagents should be stored under anhydrous conditions and should be dried under high vacuum at 40◦ C for 24 hr before use. Additional care must be taken with the phosphoramidites of 7-deazaguanosines and isoguanosines, which easily absorb water and are sensitive to oxidation upon exposure to light.
Synthesis of Modified Oligonucleotides and Conjugates
4.25.23 Current Protocols in Nucleic Acid Chemistry
Supplement 20
Troubleshooting If the phosphoramidite coupling yield is <90%, the following measures are strongly recommended: (1) repurify the phosphoramidites; (2) dry the phosphoramidites under high vacuum for >40 hr at 40◦ to 50◦ C; (3) increase the concentrations of the phosphoramidite solutions used; (4) dry all chemical reagents with 3-Å molecular sieves for 2 days prior to use; and (5) prolong the coupling time for modified phosphoramidites. High coupling yields can give confidence that the reagents used are of good quality.
Anticipated Results Using the procedures described in this unit, phosphoramidites of 7-substituted 7deazapurines can be prepared on a small or large scale. Oligonucleotide duplexes with increased stability can be obtained by incorporating 7-substituted 7-deazapurine2 -deoxyribonucleosides. With high-quality phosphoramidites, DNA synthesis is routine. Upon DMTr-on and DMTr-off purification by reversed-phase or ion-exchange HPLC, homogenous products are obtained.
Time Considerations The time needed for phosphoramidite synthesis depends on the time required for reaction and purification, and on the technical expertise of the investigator; the synthesis of phosphoramidites as described in these protocols takes ∼2 weeks for an experienced chemist. Oligonucleotide synthesis using these basemodified phosphoramidite reagents requires more time than does oligonucleotide synthesis using standard phosphoramidites.
Literature Cited Balow, G., Mohan, V., Lesnik, E.A., Johnston, J.F., Monia, B.P., and Acevedo, O.L. 1998. Biophysical and antisense properties of oligodeoxynucleotides containing 7-propynyl-, 7-iodo- and 7-cyano-7-deaza-2-amino-2 deoxyadenosines. Nucl. Acids Res. 26:33503357. Barr, P.J., Thayer, R.M., Laybourn, P., Najarian, R.C., Seela, F., and Tolan, D.R. 1986. 7Deaza-2 -deoxyguanosine-5 -triphosphate: Enhanced resolution in M13 dideoxy sequencing. Biotechniques 4:428-432.
Using Pyrrolo[2,3d]pyrimidines to Replace Purines
Buhr, C.A., Wagner, R.W., Grant, D., and Froehler, B.C. 1996. Oligodeoxynucleotides containing C-7 propyne analogs of 7deaza-2 -deoxyguanosine and 7-deaza-2 deoxyadenosine. Nucl. Acids Res. 24:29742980.
Latimer, L.J. and Lee, J.S. 1991. Ethidium-bromide does not fluoresce when intercalated adjacent to 7-deazaguanine in duplex DNA. J. Biol. Chem. 266:13849-13851. Li, H., Peng, X., and Seela, F. 2004. Fluorescence quenching of parallel-stranded DNA bound ethidium bromide: The effect of 7-deaza-2 deoxyisoguanosine and 7-halogenated derivatives. Bioorg. Med. Chem. Lett. 14:6031–6034. McDowell, J.A. and Turner, D.H. 1996. Investigation of the structural basis for thermodynamic stabilities of tandem GU mismatches: Solution structure of (rGAG GU CUC)2 by two-dimensional NMR and simulated annealing. Biochemistry 35:14077-14089. Mizusawa, S., Nishimura, S., and Seela, F. 1986. Improvement of the dideoxy chain termination method of DNA sequencing by use of deoxy-7deazaguanosine triphosphate in place of dGTP. Nucl. Acids Res. 14:1319-1324. Okamoto, A., Tanaka, K., and Saito, I. 2002. 2Amino-7-deazaadenine forms stable base pairs with cytosine and thymine. Bioorg. Med. Chem. Lett. 12:97-99. Ramzaeva, N. and Seela, F. 1995. 7-Substituted 7-deaza-2 -deoxyguanosines: Regioselective halogenation of pyrrolo[2,3-d]pyrimidine nucleosides. Helv. Chim. Acta 78:1083-1090. Ramzaeva, N. and Seela, F. 1996. Duplex stability of 7-deazapurine DNA: Oligonucleotides containing 7-bromo or 7-iodo-7-deazaguanine. Helv. Chim. Acta 79:1549-1558. Ramzaeva, N., Mittelbach, C., and Seela, F. 1997. 7Deazaguanine DNA: Oligonucleotides with hydrophobic or cationic side chains. Helv. Chim. Acta 80:1809-1822. Seela, F. and Chen, Y. 1997. Methylated DNA: The influence of 7-deaza-7-methylguanine on the structure and stability of oligonucleotides. Helv. Chim. Acta 80:1073-1086. Seela, F. and Driller, H. 1989. Alternating d(GC)3 and d(C-G)3 hexanucleotides containing 7deaza-2 -deoxyguanosine or 8-aza-7-deaza-2 deoxyguanosine in place of dG. Nucl. Acids Res. 17:901-910. Seela, F. and Kaiser, K. 1986. Phosphoramidites of base-modified 2 -deoxyinosine isosteres and solid-phase synthesis of d(GCI∗CGC) oligomers containing an ambiguous base. Nucl. Acids Res. 14:1825-1844. Seela, F. and Kehne, A. 1983. 2 -Desoxytubercidin: Synthese eines 2 -desoxyadenosin-isosteren durch phasentransferglycosylierung. Liebigs Ann. Chem. 876-884. Seela, F. and Kehne, A. 1985. 2 -Desoxytubercidin: Synthese des O-3 -phosphoramidites und kondensation zu 2 -desoxytubercidylyl(3 →5 )-2 desoxytubercidin. Tetrahedron 41:5387-5392. Seela, F. and Mersmann, K. 1993. 7Deazaguanosine: Synthesis of an oligoribonucleotide building block and disaggregation of the UGGGGU G4 structure by the modified base. Helv. Chim. Acta 76:1435-1449.
4.25.24 Supplement 20
Current Protocols in Nucleic Acid Chemistry
Seela, F. and Peng, X. 2004. Regioselective syntheses of 7-halogenated 7-deazapurine nucleosides related to 2-amino-7-deaza-2 -deoxyadenosine and 7-deaza-2 -deoxyisoguanosine. Synthesis 8:1203-1210.
Seela, F., Berg, H., and Rosemeyer, H. 1989. Bending of oligonucleotides containing an isosteric nucleobase: 7-Deaza-2 -deoxyadenosine replacing dA within d(A)6 tracts. Biochemistry 28:6193-6198.
Seela, F. and Shaikh, K.I. 2005. Oligonucleotides containing 7-propynyl-7-deazaguanine: Synthesis and base pair stability. Tetrahedron In press.
Seela, F., Wiglenda, T., Rosemeyer, H., Eickmeier, H., and Reuter, H. 2002. 7-Deaza-2 - deoxyxanthosine dihydrate forms water-filled nanotubes with C-H···O hydrogen bonds. Angew. Chem. Int. Ed. Engl. 41:603-605.
Seela, F. and Thomas, H. 1994. Synthesis of certain 5-substituted 2 -deoxytubercidin derivatives. Helv. Chim. Acta 77:897-903. Seela, F. and Thomas, H. 1995. Duplex stabilization of DNA: Oligonucleotides containing 7substituted 7-deazaadenines. Helv. Chim. Acta 78:94-108. Seela, F. and Wei, C. 1997. 7-Deazaisoguanine quartets: Self-assembled oligonucleotides lacking the Hoogsteen motif. Chem. Commun. 18691870. Seela, F. and Wei, C. 1999. The base-pairing properties of 7-deaza-2 -deoxyisoguanosine and 2 deoxyisoguanosine in oligonucleotide duplexes with parallel and antiparallel chain orientation. Helv. Chim. Acta 82:726-745. Seela, F. and Zulauf, M. 1996. Palladium-catalyzed cross coupling of 7-iodo-2 -deoxytubercidin with terminal alkynes. Synthesis 726-730. Seela, F. and Zulauf, M. 1998. 7-DeazaadenineDNA: Bulky 7-iodo substituents or hydrophobic 7-hexynyl chains are well accommodated in the major groove of oligonucleotide duplexes. Chem. Eur. J. 4:1781-1790. Seela, F. and Zulauf, M. 1999. Oligonucleotides containing 7-deazaadenines: The influence of the 7-substituent chain length and charge on the duplex stability. Helv. Chim. Acta 82:18781898. Seela, F., Steker, H., Driller, H., and Bindig, U. 1987. 2-Amino-2 -desoxytubercidin und verwandte pyrrolo[2,3-d]pyrimidinyl-2 desoxyribofuranoside. Liebigs Ann. Chem. 15-19. Seela, F., Westermann, B., and Bindig, U. 1988. Liquid-liquid and solid-liquid phase-transfer glycosylation of pyrrolo[2,3-d]pyrimidines: Stereospecific synthesis of 2-deoxy-β-Dribofuranosides related to 2 -deoxy-7carbaguanosine. J. Chem. Soc. Perkin Trans. I:697-702.
Seela, F., Shaikh, K., and Wiglenda, T. 2003. Synthesis and properties of halogenated 7-deaza2 - deoxyxanthosine and protected derivatives for oligonucleotides synthesis. Nucleosides Nucleotides Nucl. Acids 22:1239-1241. Seela, F., Chittepu, P., He, Y., He, J., and Xu, K. 2005a. 6-Azapyrimidine and 7-deazapurine 2 -deoxy-2 -fluoroarabinonucleosides: Synthesis, conformation and properties of oligonucleotides. Nucleosides Nucleotides Nucleic Acids In press. Seela, F., Peng, X., and Ming, X. 2005b. 7Deazapurine-2,6-diamine and 7-deazaguanine: Synthesis and property of 7-substituted nucleosides and oligonucleotides. Nucleosides Nucleotides Nucleic Acids In press. Ti, G.S., Gaffney, B.L., and Jones, R.A. 1982. Transient protection: Efficient one-flask syntheses of protected deoxynucleosides. J. Am. Chem. Soc. 104:1316-1319. Winkeler, H.D. and Seela, F. 1983. Synthesis of 2-amino-7-(2 -deoxy-β-D-erythropentofuranosyl)-3,7-dihydro-4H-pyrrolo[2,3d]pyrimidin-4-one, a new isostere of 2 deoxyguanosine. J. Org. Chem. 48:3119-3122.
Internet Resources http://www.seela.net Research web site of Dr. Frank Seela, author of this unit.
Contributed by Frank Seela and Xiaohua Peng Universit¨at Osnabr¨uck Osnabr¨uck, Germany and Center for Nanotechnology (CeNTech) M¨unster, Germany
Synthesis of Modified Oligonucleotides and Conjugates
4.25.25 Current Protocols in Nucleic Acid Chemistry
Supplement 20
An Aminooxy-Functionalized Non-Nucleosidic Phosphoramidite for the Construction of Multiantennary Oligonucleotide Glycoconjugates on a Solid Support
UNIT 4.26
This unit contains procedures for the synthesis of an aminooxy-functionalized nonnucleosidic phosphoramidite and fully acetylated 4-oxobutyl α-D-mannopyranoside, and a description of their use in the preparation of multivalent oligonucleotide glycoconjugates. Commercially available diethyl 2,2-bis(hydroxymethyl)malonate can be converted in six steps into a phosphoramidite that bears two phthaloyl-masked aminooxy groups in addition to a 4,4 -dimethoxytritylated hydroxyl group, and allows normal oligonucleotide chain elongation. After conventional chain assembly, the phthaloyl protecting groups can be removed by a hydrazinium acetate treatment and subsequently oximated with a carbohydrate building block carrying an anomeric aldehyde tether. Synthesis of the non-nucleosidic phosphoramidite and the carbohydrate building block are described in Basic Protocols 1 and 2, respectively. The use of these reagents for the preparation of oligonucleotide glycoconjugates is then presented as Basic Protocol 3. The phthaloyl protecting groups may be removed with hydrazinium acetate in pyridine, i.e., under conditions that cleave neither the commonly used succinyl linker nor the 2-cyanoethyl phosphate-protecting groups. Glycosidic aldehydes react readily with the exposed aminooxy groups, affording fully protected oxime conjugates that have been shown to withstand standard ammonolytic deprotection and cleavage from the support. Accordingly, a phthaloyl protection strategy may well be adopted for the construction of glycooligonucleotide conjugates on a solid support. CAUTION: Perform all operations involving organic solvents and reagents in a wellventilated fume hood, and wear gloves and protective glasses.
PREPARATION OF THE AMINOOXY-FUNCTIONALIZED PHOSPHORAMIDITE FROM DIETHYL 2,2-BIS(HYDROXYMETHYL)MALONATE Aminooxy-functionalized building blocks have been previously used for the solidphase 5 -conjugation of oligonucleotides. A 5 -terminal aminooxy group has been introduced as a 2-[(2-ureido)-4-(2-phthalimidooxyethoxy)quinoline]ethyl phosphoramidite (Hamma and Miller, 2003), an 11-phthalimidoxy-3,6,9-trioxaundecyl phosphoramidite (Salo et al., 1999), and an N-trityl-6-aminooxyhexyl phosphoramidite (Defrancq and Lhomme, 2001). The present protocol is based on Katajisto et al. (2004). The synthesis of the aminooxy-functionalized phosphoramidite (S.6) is outlined in Figure 4.26.1. Diethyl 2,2-bis(hydroxymethyl)malonate (S.1) is first converted to its di-O-methoxymethylene derivative (S.2) and the latter is aminolyzed with 3-aminopropanol to obtain S.3. The free hydroxy functions are then displaced with N-phthaloyl-protected aminooxy groups by a Mitsunobu reaction using N-hydroxyphthalimide as the nucleophile and diethyl azodicarboxylate (DEAD) and triphenylphosphine as activators. The crude product is subjected to acid-catalyzed hydrolysis to remove the methoxymethylene protection (S.4). 4,4 -Dimethoxytritylation of one of the exposed hydroxyl groups (S.5) followed by phosphitylation of the other with 2-cyanoethyl N,N-diisopropylphosphonamidic chloride completes the synthesis of the phosphoramidite (S.6). This building block is used Contributed by Johanna Katajisto, Pasi Virta, and Harri L¨onnberg Current Protocols in Nucleic Acid Chemistry (2005) 4.26.1-4.26.16 C 2005 by John Wiley & Sons, Inc. Copyright
BASIC PROTOCOL 1
Modified Oligonucleotides and Conjugates
4.26.1 Supplement 21
Figure 4.26.1 Synthesis of the non-nucleosidic phosphoramidite S.6 (see Basic Protocol 1). DMTr, 4,4 -dimethoxytrityl. Reprinted from Katajisto et al. (2004) with permission from the American Chemical Society.
in machine-assisted oligonucleotide synthesis to construct conjugates bearing protected aminooxy functions (see Basic Protocol 3).
Materials
An AminooxyFunctionalized Non-Nucleosidic Phosphoramidite for Oligonucleotide Glycoconjugates
Diethyl 2,2-bis(hydroxymethyl)malonate (S.1.), 95% pure (Acros) Dry tetrahydrofuran (THF), freshly distilled from sodium (store over 4-Å molecular sieves) Trimethyl orthoformate, 98% pure (Aldrich) p-Toluenesulfonic acid monohydrate, ≥98.5% pure (Aldrich) 5% (w/v) aqueous sodium hydrogen carbonate (NaHCO3 ) Diethyl ether Saturated aqueous sodium chloride (NaCl) Sodium sulfate (Na2 SO4 ), anhydrous Silica gel: 0.040- to 0.063-mm Fluka Kieselgel 60 (dry overnight in 150◦ C oven) Dichloromethane (CH2 Cl2 ), ≥99% pure Bromocresol green indicator: dissolve 0.04 g bromocresol green (Merck) in 100 mL ethanol and add 0.1 M aqueous NaOH until the blue color appears (store up to 1 month at room temperature) 3-Aminopropanol, 99% pure (Aldrich)
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Current Protocols in Nucleic Acid Chemistry
Methanol (MeOH), ≥99.8% pure Dry benzene, distilled from powdered CaH2 (store over 4-Å molecular sieves) Triphenylphosphine, 99% pure (Aldrich) N-Hydroxyphthalimide, 97% pure (Aldrich) Diethyl azodicarboxylate (DEAD), ≥97% (Fluka) 2-Propanol (i-PrOH), Baker analyzed 80% (v/v) aqueous acetic acid (AcOH) Dry pyridine, distilled from powdered CaH2 (store over 4-Å molecular sieves) 4,4 -Dimethoxytrityl chloride (DMTr-Cl), 97% (Aldrich) Dry toluene, distilled from powdered CaH2 (store over 4-Å molecular sieves) Dry acetonitrile, HPLC grade (store over 4-Å molecular sieves) Phosphorus pentoxide (P2 O5 ), 97% pure (Aldrich) Dry nitrogen (or argon) Anhydrous triethylamine, distilled from powdered CaH2 (store over 4-Å molecular sieves) 2-Cyanoethyl N,N-diisopropylphosphonamidic chloride (TRC) Ethyl acetate (EtOAc), analytical grade Hexane, HPLC grade Separatory funnels Rotary evaporator equipped with a water aspirator 5 × 35–cm sintered glass chromatography column, porosity 2 TLC plate: silica-coated aluminium plate with fluorescent indicator (Merck silica gel 60 F254 ) 3 × 20–cm sintered glass chromatography column, porosity 3 254-nm UV lamp Vacuum desiccator Additional reagents and equipment for column chromatography (APPENDIX 3E) and thin-layer chromatography (TLC, APPENDIX 3D) Protect hydroxy groups of diethyl 2,2-bis(hydroxymethyl)malonate 1. Dissolve 10.0 g (45.4 mmol) diethyl 2,2-bis(hydroxymethyl)malonate (S.1) in 60 mL THF with magnetic stirring. Add 6.5 mL (59.0 mmol) trimethyl orthoformate and a catalytic amount (10 mg) of p-toluenesulfonic acid monohydrate to the reaction mixture. Stopper the flask and continue stirring overnight. 2. Pour the reaction mixture into 100 mL of magnetically stirred ice-cold 5% (w/v) aqueous NaHCO3 . Extract the mixture twice with 100 mL diethyl ether. Combine the organic phases and wash them in a separatory funnel with 100 mL saturated aqueous NaCl. 3. Dry the pooled organic phase over 15 g anhydrous Na2 SO4 and filter off the drying agent by gravity using filter paper. Evaporate the solution to dryness using a rotary evaporator equipped with a water aspirator. 4. Purify the residue by silica gel chromatography as follows. Pack a 5 × 35–cm glass column with 400 mL silica gel in CH2 Cl2 . Redissolve the residue from step 3 in 5 to 10 mL CH2 Cl2 , apply this sample to the column, and elute the column with CH2 Cl2 (typically 1.5 to 2 L). Collect fractions of ∼50 mL. Basic column chromatography techniques are described in APPENDIX 3E.
5. Monitor fractions by TLC (APPENDIX 3D) on a fluorescent indicator plate using CH2 Cl2 as solvent. Visualize the desired product by spraying plate with bromocresol green indicator and then warming it gently on a hot plate. The product (S.2) can be seen as an yellow spot on a blue background (Rf = 0.66).
Modified Oligonucleotides and Conjugates
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6. Combine the fractions containing the product and evaporate to dryness using a rotary evaporator. 7. Characterize the product by 1 H and 13 C NMR and mass spectrometry. Diethyl 2-methoxy-1,3-dioxane-5,5-dicarboxylate (S.2): Yield of colorless oil: 11.2 g (94%). 1 H NMR: 5.28 (s, 1H), 4.54 (d, 2H, 2 JAB = 10.5 Hz), 4.04-4.52 (m, 6H), 3.38 (s, 3H), 1.23-1.34 (m, 6H). 13 C NMR: 169.5, 167.4, 108.9, 62.8, 62.1, 52.8, 13.9. HRMS (ESI): anal. calcd. for C11 H18 O7 [M + Na]+ 285.0945, found 285.0950.
Perform aminolysis with 3-aminopropanol 8. Reflux a mixture of 7.00 g (26.7 mmol) S.2 with 13.5 mL (160 mmol) 3aminopropanol for 48 hr. Monitor progress by TLC as in step 5 using 1:9 (v/v) MeOH/CH2 Cl2 as an eluent. By TLC analysis with bromocresol green, the unreacted starting material (S.2; Rf = 0.92), a monoamide intermediate (Rf = 0.52), and the desired product (S.3; Rf = 0.32) can all be detected.
9. Isolate and purify the desired diamide S.3 from the excess of 3-aminopropanol by silica gel column chromatography as follows. Pack a 5 × 35–cm glass column with 400 mL silica gel in 5% (v/v) MeOH/CH2 Cl2 . Apply the solution from step 8 directly onto the column and elute with 5% (v/v) MeOH/CH2 Cl2 (typically 1.5 to 2 L). Collect fractions of ∼50 mL. 10. Monitor fractions by TLC as in step 8. Combine the fractions containing the product and evaporate in vacuo with a rotary evaporator to yield S.3 as a clear oil. 11. Characterize the product by 1 H and 13 C NMR and mass spectrometry. N,N-Bis(3-hydroxypropyl)-2-methoxy-1,3-dioxane-5,5-dicarboxamide (S.3): Yield of a clear oil: 6.5 g (76% relative to S.2). 1 H NMR: 5.29 (s, 1H), 4.50 (d, 2H, 2 JAB = 13.6 Hz), 4.18 (d, 2H, 2 JAB = 13.6 Hz), 3.61-3.73 (m, 4H), 3.38-3.55 (m, 7H), 1.73 (m, 4H). 13 C NMR: 169.6, 169.6, 109.8, 65.1, 59.6, 58.3, 53.2, 36.9, 18.8. HRMS (ESI): anal. calcd. for C13 H24 N2 O7 [M + Na]+ 343.1476, found 343.1469.
Introduce phthaloyl-protected aminooxy groups and remove methoxymethylene protection 12. Coevaporate 1.10 g (3.43 mmol) S.3 twice with 20 mL dry benzene using a rotary evaporator equipped with a water aspirator. 13. Dissolve the residue in 20 mL anhydrous THF together with 1.98 g (7.55 mmol) triphenylphosphine and 1.23 g (7.55 mmol) N-hydroxyphthalimide. Add 1.17 mL (7.55 mmol) DEAD dropwise to the reaction mixture and continue stirring overnight. 14. Remove the solvent in vacuo with a rotary evaporator. Pack a 3 × 20–cm column with 100 mL silica gel in CH2 Cl2 . Redissolve the residue in a minimal amount (5 to 10 mL) of CH2 Cl2 , apply the residue to the column, then elute with a step gradient from neat CH2 Cl2 to 97:3 (v/v) CH2 Cl2 /i-PrOH (typically 300 to 400 mL) to separate the product from excess reagents and most of the triphenylphosphine oxide formed during the reaction. Collect fractions of ∼10 mL. 15. Monitor fractions by TLC on fluorescent indicator plates using 1:9 (v/v) MeOH/CH2 Cl2 and UV detection (Rf = 0.52). Combine the fractions containing the product and evaporate to dryness using a rotary evaporator. An AminooxyFunctionalized Non-Nucleosidic Phosphoramidite for Oligonucleotide Glycoconjugates
Purification of crude 2-methoxy-N,N-bis(3-phthalimidooxypropyl)-1,3-dioxane-5,5dicarboxamide was found to be seriously hampered by the presence of triphenylphosphine oxide formed during the Mitsunobu reaction. To avoid loss of the material by repeated purifications, the crude product is subjected to acid-catalyzed hydrolysis of the methoxymethylene protecting group (step 16).
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The product, still contaminated by some triphenylphosphine oxide, exhibited the following NMR signals: 1 H NMR: 7.45-7.84 (m, 8H), 5.27 (s, 1H), 4.56 (d, 2H, 2 JAB = 10.3 Hz), 4.20-4.31 (m, 6H), 3.58 (m, 4H), 3.36 (s, 3H), 2.01 (m, 4H). 13 C NMR: 169.4, 168.9, 163.7, 134.5, 132.0, 123.0, 109.7, 76.1, 64.9, 62.1, 53.1, 36.5, 27.5. Identity of the compound was verified by HRMS (ESI): anal. calcd. for C29 H30 N4 O11 [M + Na]+ 633.1776, found 633.1803.
16. Dissolve the crude product in 20 mL of 80% (v/v) aqueous acetic acid and let stand 2 hr at room temperature. 17. Evaporate the solution in vacuo using a rotary evaporator. Coevaporate the residue three times with 50 mL water using a rotary evaporator. 18. Purify the compound (S.4) by silica gel chromatography as follows. Pack a 3 × 20–cm glass column with 100 mL silica gel in CH2 Cl2 . Redissolve the residue from step 17 in 2 to 3 mL CH2 Cl2 , load onto the column, then elute using a step gradient from neat CH2 Cl2 to 5% (v/v) MeOH/CH2 Cl2 (typically 300 to 400 mL). Collect fractions of ∼10 mL. 19. Monitor by TLC as in step 15 (Rf of S.4 = 0.37). Combine fractions containing pure product and evaporate to dryness using a rotary evaporator. 20. Characterize compound S.4 by 1 H and 13 C NMR and mass spectrometry. 2,2-Bis(hydroxymethyl)-N,N-bis(3-phthalimidooxypropyl)malondiamide (S.4): Yield of a colorless oil: 300 mg (15% starting from S.3). TLC (1:9 v/v MeOH/CH2 Cl2 ): Rf = 0.37. 1 H NMR: 7.72-7.91 (m, 8H), 4.26 (t, 4H, J = 5.7 Hz), 3.94 (broad s, 4H), 3.62 (dd, 4H, J = 7.5 and 11.3 Hz), 2.01 (m, 4H). 13 C NMR: 169.5, 168.9, 163.6, 134.6, 131.8, 123.5, 65.0, 55.0, 36.4, 27.8. HRMS (ESI): anal. calcd. for C27 H28 N4 O10 [M + H]+ 569.1878, found 569.1896. Compound S.4 contains four exchangable protons (two NH, two OH) that are not visible on the 1 H NMR spectrum.
Dimethoxytritylate S.4 21. Coevaporate 700 mg (1.23 mmol) S.4 twice with 10 mL anhydrous pyridine using a rotary evaporator, then dissolve residue in 10 mL dry pyridine. 22. Add 417 mg (1.23 mmol) DMTr-Cl, and continue stirring the mixture overnight at room temperature. 23. Evaporate the mixture in vacuo using a rotary evaporator, dilute the residue with 30 mL CH2 Cl2 , and wash twice in a separatory funnel, each time with 15 mL of 5% NaHCO3 . 24. Dry over 10 g anhydrous Na2 SO4 and filter off the drying agent by gravity using filter paper. Evaporate the solution to dryness, and then coevaporate three times with 20 mL dry toluene using a rotary evaporator. 25. Pack a 3 × 20–cm glass column with 100 mL silica gel in CH2 Cl2 . Redissolve the crude oily residue in 3 mL CH2 Cl2 , apply to the column, then elute using a step gradient from neat CH2 Cl2 to 3% MeOH/CH2 Cl2 (typically 300 to 400 mL). Collect fractions of ∼10 mL. 26. Monitor fractions by TLC on fluorescent indicator plates using 1:9 (v/v) MeOH/CH2 Cl2 . Visualize the product bearing the DMTr group, in addition to normal UV detection, by heating the TLC plate on a hot plate. Combine fractions containing the product and evaporate in vacuo using a rotary evaporator to yield the product as a white amorphous solid. Upon heating, the product (S.5) can be seen as a yellow spot on a white background (Rf = 0.41).
Modified Oligonucleotides and Conjugates
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27. Characterize the compound (S.5) by 1 H and 13 C NMR and mass spectrometry. 2-(4,4 -Dimethoxytrityloxymethyl)-2-(hydroxymethyl)-N,N-bis(3-phthalimidooxy propyl)malonamide (S.5): Yield of white solid foam: 440 mg (40% relative to S.4). 1 H NMR: 7.78-7.83 (m, 4H), 7.71-7.76 (m, 4H), 7.66 (t, 2H, J = 6.4 Hz), 7.11-7.38 (m, 9H), 6.80 (m, 4H), 4.18 (m, 4H), 4.10 (d, 2H, J = 6.4 Hz), 3.81 (t, 1H, J = 6.4 Hz), 3.75 (s, 6H), 3.46-3.62 (m, 6H), 1.96 (m, 4H). 13 C NMR: 170.9, 163.5, 158.5, 144.3, 135.3, 134.5, 130.1, 128.9, 128.1, 127.9, 126.9, 123.6, 113.2, 86.5, 76.0, 65.1, 64.0, 58.7, 55.2, 36.4, 27.9. HRMS (ESI): anal. calcd. for C48 H46 N4 O12 [M + Na]+ 893.3004, found 893.3046.
Phosphitylate S.5 28. Dry 220 mg (0.25 mmol) S.5 overnight over P2 O5 in a vacuum desiccator. Dissolve this in 2 mL dry acetonitrile, and apply a dry nitrogen atmosphere. 29. Add 175 µL (1.26 mmol) triethylamine and 62 µL (0.28 mmol) 2-cyanoethyl N,Ndiisopropylphosphonamidic chloride to the reaction solution. Stir the mixture 45 min under nitrogen. Monitor the reaction by TLC in 60% (v/v) EtOAc/hexane, visualizing the product by heating the TLC plate on a hot plate. The product (S.6) can be seen as a yellow spot on a white background (Rf = 0.82).
30. Apply the reaction mixture to a 3 × 20–cm column containing 50 mL dried silica gel and isolate the pure compound by eluting with 60:39:1 (v/v/v) dry ethyl acetate/hexane/triethylamine (typically 150 to 200 mL). Collect fractions of ∼5 mL. 31. Monitor fractions by TLC as in step 26 (Rf for S.6 = 0.83). Combine fractions containing the phosphoramidite, and evaporate and dry in vacuo using a rotary evaporator to yield S.6 as a white solid foam. 32. Characterize the compound by 1 H and 13 C NMR and mass spectrometry. 2-Cyanoethyl 3-[(4,4 -dimethoxytrityl)oxy]-2,2-bis[2-aza-1-oxo-5-(phthalimidooxy) pentyl]propyl-N,N-diisopropylphosphoramidite (S.6) is stable during prolonged storage at −20◦ C. Yield of solid white foam: 230 mg (85%). 1 H NMR: 7.71-7.84 (m, 8H), 7.61 (t, 2H, J = 5.1 Hz), 7.10-7.45 (m, 9H), 6.80 (m, 4H), 4.03-4.45 (m, 6H), 3.69-3.78 (m, 10H), 3.25-3.57 (m, 6H), 2.56 (m, 2H, J = 7.2 Hz), 1.91-1.99 (m, 4H), 1.12 (d, 12H, J = 7.2 Hz). 13 C NMR: 170.7, 163.3, 159.5, 144.1, 135.1, 134.3, 129.9, 128.7, 127.9, 127.7, 123.3, 113.1, 86.4, 75.9, 64.8, 64.5, 53.5, 59.4, 58.7, 58.3, 55.0, 43.0, 36.2, 27.9, 24.3, 18.7. 31 P NMR: 146.6. MS (ESI): anal. calcd. for C57 H63 N6 O13 P [M + Na]+ 1093.4, found 1094.1.
BASIC PROTOCOL 2
An AminooxyFunctionalized Non-Nucleosidic Phosphoramidite for Oligonucleotide Glycoconjugates
PREPARATION OF THE MANNOSYL ALDEHYDE LIGAND The carbonyl group is known to be exceptionally susceptible to attack by an α-nucleophile such as an aminooxy group. Previous studies have shown that the oxime conjugates obtained withstand the standard ammonolysis performed upon completion of the oligonucleotide chain elongation to deprotect and cleave the product from the support (Salo et al., 1999). In the present protocol, based primarily on Katajisto et al. (2004), fully acetylated 4-oxobutyl α-D-mannopyranoside (S.9) is used as an example of a glycosyl aldehyde ligand with which the solid-supported oximation is carried out to obtain the desired oligonucleotide glycoconjugate. This compound can be easily prepared by a two-step procedure, as depicted in Figure 4.26.2. Boron trifluoride etherate–promoted glycosidation of commercially available peracetylated α-D-mannopyranose (S.7) with 1,4-butanediol yields a 4-hydroxybutyl mannoside peracetate (S.8), which is then subjected to Swern oxidation to yield S.9. This protocol enables easy preparation of the desired target compound in spite of the relatively low yield obtained in converting S.7 to S.8.
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Figure 4.26.2 Synthesis of the sugar ligand S.9 (see Basic Protocol 2). Ac, acetyl. Reprinted from Katajisto et al. (2004) with permission from the American Chemical Society.
Materials α-D-Mannose pentaacetate (S.7; Sigma) Dry benzene, distilled from powdered CaH2 (store over 4-Å molecular sieves) Dry nitrogen (or argon) Dry acetonitrile (MeCN), HPLC grade (store over 4-Å molecular sieves) 1,4-Butanediol, 99% pure (Aldrich) Boron trifluoride etherate (Merck) Dichloromethane (CH2 Cl2 ), ≥ 99% pure Sodium sulfate (Na2 SO4 ), anhydrous Silica gel: 0.040- to 0.063-mm Fluka Kieselgel 60 (dry overnight in oven at 150◦ C) Ethyl acetate (EtOAc), analytical reagent Hexane, HPLC grade 10% (v/v) H2 SO4 Oxalyl chloride, 99% pure (Aldrich) Dry argon Dry ice/isopropanol bath Dry dimethyl sulfoxide (DMSO; store over 4-Å molecular sieves) Triethylamine (TEA), freshly distilled from powdered CaH2 Diethyl ether 1 M aqueous HCl, ice cold Saturated aqueous sodium hydrogen carbonate (NaHCO3 ) Saturated aqueous sodium chloride (NaCl) Methanol (MeOH), ≥99.8% pure Separatory funnels Rotary evaporator equipped with a water aspirator 3 × 20–cm sintered glass chromatography column, porosity 3 TLC plate: silica-coated aluminium plate with fluorescent indicator (Merck silica gel 60 F254 ) 254-nm UV lamp Additional reagents and equipment for column chromatography (APPENDIX 3E) and thin-layer chromatography (TLC, APPENDIX 3D)
Modified Oligonucleotides and Conjugates
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Glycosidate S.7 with 1,4-butanediol 1. Coevaporate 5.00 g (12.8 mmol) α-D-mannose pentaacetate (S.7) twice with dry benzene using a rotary evaporator equipped with a water aspirator. Apply a dry nitrogen (or argon) atmosphere. 2. Dissolve the residue in 30 mL dry MeCN. Add 1.10 mL (12.8 mmol) 1,4-butanediol and then 4.82 mL (38.4 mmol) boron trifluoride etherate, and allow the reaction to proceed at room temperature for 2 hr. Extension of the reaction time gives a decreased yield of the target compound and increased amount of monoacetylated 1,4-butanediol as a byproduct. For further discussion, see Background Information.
3. Dilute the reaction mixture with 70 mL CH2 Cl2 and wash with 50 mL water in a separatory funnel. Dry over 10 g anhydrous Na2 SO4 , filter by gravity using filter paper to remove the solid, then evaporate the solution to dryness using a rotary evaporator with a water aspirator. 4. Separate the product from the unreacted starting material S.7 by silica gel chromatography as follows. Pack a 3 × 20–cm glass column with 100 mL silica gel in 2:1 (v/v) EtOAc/hexane. Redissolve sample in a minimal amount (∼5 mL) of 2:1 (v/v) EtOAc/hexane, apply to column, and elute with 2:1 (v/v) EtOAc/hexane (typically 300 to 400 mL). Collect fractions of ∼10 mL. Basic column chromatography techniques are described in APPENDIX 3E.
5. Monitor the fractions by TLC (APPENDIX 3D) on fluorescent indicator plates using 1:9 (v/v) MeOH/CH2 Cl2 . Visualize the desired product by dipping the TLC plate in 10% aqueous H2 SO4 and then heating it on a hot plate. The unreacted starting material and the desired product are both charred and can be visualized as black spots on the plate (S.7, Rf = 0.81; S.8, Rf = 0.55).
6. Combine the fractions containing the product (S.8) and evaporate to dryness using a rotary evaporator. 7. Characterize S.8 by 1 H and 13 C NMR and high-resolution mass spectrometry. 4-Hydroxybutyl 2,3,4,6-tetra-O-acetyl-α-D-mannopyranoside (S.8): Yield of a colorless oil: 1.05 g (19%). 1 H NMR: 7.26 (6s, 1H) 5.33 (dd, 1H, J = 4.1 and 10.8 Hz), 5.29 (t, 1H, J = 10.8 Hz), 5.22 (dd, 1H, J = 4.3 and 1.6 Hz), 4.81 (d, 1H, J = 1.4 Hz), 4.28 (dd, 1H, J = 5.4 and 12.7 Hz), 4.10 (dd, 1H, J = 2.9 and 12.7 Hz), 3.98 (m, 1H), 3.73 (m, 1H), 3.68 (t, 2H, J = 5.4 Hz), 3.50 (m, 1H), 2.14, 2.10, 2.05, and 1.99 (each s, 3H), 1.72-1.63 (m, 4H). 13 C NMR: 170.7, 170.1, 169.9, 169.8, 97.6, 69.7, 69.1, 68.4, 68.3, 66.2, 62.5, 29.4, 25.7, 20.9, 20.8, 20.7. HRMS (ESI): anal. calcd. for C18 H28 O11 [M + Na]+ 443.1524, found 443.1516.
Perform Swern oxidation of S.8 8. Predry 1.00 g (2.38 mmol) S.8 overnight over P2 O5 in a vacuum desiccator. Dissolve in 10 mL dry CH2 Cl2 . 9. Prepare a magnetically stirred solution of 1.67 mL (19.1 mmol) oxalyl chloride in 5 mL dry CH2 Cl2 . Apply a dry argon atmosphere and cool the reaction mixture to −60◦ C using a dry ice/isopropanol bath. Add 2.70 mL (38.1 mmol) DMSO dropwise to the solution and allow the mixture to react under vigorous stirring 30 min at −60◦ C. An AminooxyFunctionalized Non-Nucleosidic Phosphoramidite for Oligonucleotide Glycoconjugates
10. Slowly add S.8 to the stirring solution and continue stirring at −60◦ C for 40 min. CAUTION: The reaction is violent and exothermic at ambient temperature. The reaction proceeds via an intermediate complex that is unstable at temperatures higher than −60◦ C. CAUTION: Dimethyl sulfide, which has a strong stench, is generated during the reaction; the reaction must therefore be carried out in a well-ventilted fume hood.
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11. Add 7.91 mL (57.1 mmol) TEA, remove the cooling bath, and allow the solution to warm to 0◦ C. 12. Pour the mixture into 100 mL magnetically stirred, ice-cold water, and extract three times with 100 mL diethyl ether. 13. Combine the organic phases and wash in a separatory funnel successively with 50 mL ice-cold 1 M aqueous HCl, 50 mL saturated aqueous NaHCO3 , and finally with saturated aqueous NaCl. 14. Dry over 10 g anhydrous Na2 SO4 , filter by using filter paper to remove the solid, then evaporate in vacuo using a rotary evaporator. 15. Apply the reaction mixture to a 3 × 20–cm column containing 100 mL silica gel and isolate the pure compound by eluting with 2:1 (v/v) EtOAc/hexane (typically 300 to 400 mL). Collect fractions of ∼10 mL. 16. Monitor fractions by TLC in 1:9 (v/v) MeOH/CH2 Cl2 and visualize the product by charring with 10% H2 SO4 (see step 5). Combine the fractions containing the product (S.9) and evaporate to dryness using a rotary evaporator. The desired product is seen as a black spot (Rf = 0.53).
17. Characterize S.9 by 1 H and 13 C NMR and high-resolution mass spectrometry. 4-Oxobutyl 2,3,4,6-tetra-O-acetyl-α-D-mannopyranoside (S.9): Yield of a clear oil: 595 mg (60%). 1 H NMR: 9.82 (s, 1H), 5.12-5.31 (m, 3H), 4.80 (s, 1H), 4.28 (dd, 1H, J = 5.6 and 12.9 Hz), 4.12 (m, 1H), 3.95 (m, 1H), 3.75 (m, 1H), 3.50 (m, 1H), 2.58 (m, 2H), 2.16, 2.11 and 2.05 (each s, 3H), 2.00-1.77 (m, 5H). 13 C NMR: 201.3, 170.6, 170.1, 169.9, 169.7, 97.5, 69.6, 68.9, 68.3, 67.9, 66.1, 62.4, 40.5, 22.3, 21.9, 20.9, 20.7, 20.4, 20.2. HRMS (ESI): anal. calcd. for C18 H26 O11 [M + Na]+ 441.13673, found 441.13667.
SYNTHESIS OF OLIGONUCLEOTIDE GLYCOCONJUGATES BY ON-SUPPORT OXIMATION
BASIC PROTOCOL 3
The standard phosphoramidite protocol can be applied to incorporate S.6 units into oligonucleotides, except that a prolonged coupling time (600 sec) should be used with S.6 to obtain an acceptable 95% coupling efficiency. For example, a protected heterosequence 5 -d(TXTXTGACGATCTCAT)-3 , where X stands for the non-nucleosidic residue S.6, may be synthesized as outlined in Figure 4.26.3. The support-bound oligonucleotide (S.10) is treated with 0.5 M hydrazinium acetate in pyridine for 30 min to remove the phthaloyl groups, and the exposed aminooxy groups are converted to stable oxime conjugates (S.11) upon reaction with the mannopyranosyl aldehyde building block (S.9). Final global deprotection and cleavage of the oxime conjugate S.12 from the support is accomplished by standard ammonolysis (33% aqueous NH3 , 7 hr at 55◦ C). The oligodeoxyribonucleotides can be assembled on any automated DNA/RNA synthesizer. The authors have used an Applied Biosystems 392 DNA synthesizer on 1.0-µmol scale with a commercial 1000-Å CPG-succinyl-thymidine support and standard phosphoramidite chemistry. Phosphoramidite S.6 was used as a 0.15 M solution in dry acetonitrile, the coupling time being 600 sec. After cleavage from the support, the oxime conjugates were evaporated, dissolved in water, purified by RP-HPLC, desalted, and characterized by ESI-MS.
Modified Oligonucleotides and Conjugates
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An AminooxyFunctionalized Non-Nucleosidic Phosphoramidite for Oligonucleotide Glycoconjugates
Figure 4.26.3 Synthesis of the oligonucleotide glycoconjugate S.12 utilizing on-support oximation. Ac, acetyl; Pht, phthaloyl.
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Materials Phosphoramidite S.6 (see Basic Protocol 1) Dry acetonitrile (MeCN), HPLC grade (store over 4-Å molecular sieves) 2 -Deoxyribonucleoside phosphoramidites (Glen Research) 0.5 M hydrazinium acetate solution: 0.124:4:1 (v/v/v) mixture of hydrazine monohydrate (>98% pure, Aldrich), analytical-grade pyridine, and glacial acetic acid Pyridine, analytical grade Acetonitrile (MeCN), analytical grade Mannosyl aldehyde ligand S.9 (see Basic Protocol 2) 33% aqueous ammonia Mobile phase A: 0.05 M ammonium acetate, pH 7.0, in H2 O Mobile phase B: 0.05 M ammonium acetate in 65% (v/v) aqueous MeCN 55◦ C water bath Speedvac evaporator HPLC system with 4.6 × 150–mm analytical ThermoHypersil C18 column and 260-nm detector Desalting column: 7.5 mm × 30 cm TSKgel G 2000 size-exclusion chromatography column, particle size 10 µm (Toso-Haas) UV spectrophotometer Additional reagents and equipment for automated solid-phase oligonucleotide synthesis (APPENDIX 3C) and for purification (UNIT 10.5) and characterization (UNIT 10.2) of oligonucleotides Incorporate S.6 into oligodeoxyribonucleotide chain 1. Prepare a 0.15 M solution of phosphoramidite S.6 in anhydrous MeCN. 2. Start the automated solid-phase oligonucleotide synthesis (APPENDIX 3C) on a 1000-Å CPG-succinyl-thymidine support (1-µmol scale). 3. Elongate the desired oligonucleotide chain using standard 2 -deoxyribonucleoside phosphoramidites and phosphoramidite S.6 in DMTr-OFF mode (APPENDIX 3C). Extend the coupling time to 600 sec for every S.6 coupling. After coupling of S.6, the detritylation is carried out by using two consecutive detritylation steps (60 sec each) separated by a trityl flush step (5 sec). Otherwise stardard protocols are employed.
4. After completion of the assembly, remove the column and place under vacuum to dry the support.
Remove phthaloyl groups 5. Transfer the support to a 1.5-ml polypropylene microcentrifuge tube, add 1 mL of 0.5 M hydrazinium acetate solution, and leave 30 min at room temperature. 6. Vacuum filter the support into a DNA synthesizer column. Wash five times, each time with 10 mL pyridine, then wash an additional five times, each time with 10 mL MeCN. Dry in vacuo using an oil pump. Owing to the high nucleophilicity of the aminooxy functions, solvents of analytical grade, which do not contain any aldehydes or ketones as impurities, should be used for the washings to avoid premature oximation.
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Perform on-support oximation 7. Transfer the support to a 1.5-ml polypropylene microcentrifuge tube and add a solution of the aldehyde-tethered sugar ligand S.9 (100 µmol in 200 µL dry MeCN). Leave the mixture at room temperature for 16 hr to ensure complete oximation. 8. Vacuum filter the support into a DNA synthesizer column. Wash five times, each time with 20 mL MeCN. Dry in vacuo using an oil pump.
Cleave and isolate oligonucleotide glycoconjugate 9. Transfer the support to a 1.5-ml polypropylene microcentrifuge tube and add 1 mL of 33% aqueous ammonia. Vortex and then incubate ≥7 hr at 55◦ C. 10. Evaporate the ammonia and dry in vacuo using a Speedvac evaporator. 11. Add 200 µL deionized water and microcentrifuge 5 min at 12,000 rpm. 12. Purify the filtrate by RP-HPLC (UNIT conditions:
10.5)
using the following recommended
4.6 × 150–mm analytical ThermoHypersil C-18 column Mobile phase A: 0.05 M ammonium acetate in H2 O Mobile phase B: 0.05 M ammonium acetate in 65% MeCN Gradient: 0% to 100% B over 30 min Flow rate: 1 mL/min Detection: 260 nm. These conditions are suitable for the purification of oligos synthesized in DMTr-OFF mode.
13. Pool the appropriate fractions, desalt on a TSKgel G 2000 size-exclusion chromatography column, and evaporate the conjugate to dryness using a Speedvac evaporator. 14. Dissolve the oligonucleotide glycoconjugate in deionized water and quantify by measuring the UV absorbance at 260 nm. Store the solution frozen. 15. Characterize S.12 by ESI-MS (UNIT 10.2). MS (ESI): anal. calcd. 5898.4, found 5897.3. See Figure 4.26.4 for an RP-HPLC profile of crude S.12 (retention time, 15.8 min). Purity: 54% (percentage of S.12 in crude reaction mixture on the basis of HPLC signal areas). For a discussion of melting studies of S.12, see Background Information. The experiments were performed in 10 mM sodium phosphate, pH 7.0 (APPENDIX 2A), containing 100 mM NaCl. The concentration of the duplex of S.12 and its complementary sequence was 2µM.
COMMENTARY Background Information
An AminooxyFunctionalized Non-Nucleosidic Phosphoramidite for Oligonucleotide Glycoconjugates
The aim of the studies described in this unit has been to develop a strategy for the solidphase synthesis of structurally diverse oligonucleotide glycoconjugates. Interest in covalent conjugation of sugar ligands to oligonucleotides has increased during recent years, as it is expected that applications will be found in cell-specific targeting of antisense oligonucleotides or small interfering ribonucleic acids (siRNA; Wang et al., 2003). Recognition of various cell-surface lectins, and hence the cell type, is based on simultaneous interaction with several sugar ligands. Accordingly, a glycocluster structure is a prerequisite
for high-affinity binding and subsequent internalization of the conjugate by endocytosis. The number of protocols described for solidphase synthesis of covalent oligonucleotide glycoconjugates is still limited. The examples available include on-support glycosylation with trichloroacetimidates (Adinolfi et al., 1999) and use of base-glycosylated nucleoside phosphoramidites (De Kort et al., 1999; Hunziker, 1999; Matsuura et al., 2001) and glycoside phosphoramidites (Akhtar et al., 1995; Sheppard et al., 2000) as building blocks for the chain assembly. In a more recent study, a method for solid-phase synthesis of oligonucleotide conjugates bearing a tetraantennary
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Figure 4.26.4 RP-HPLC profile of crude S.12 at 260 nm. Reprinted from Katajisto et al. (2004) with permission from the American Chemical Society.
glycocluster at the 5 terminus has been described (Dubber and Fr´echet, 2003). Building blocks derived from pentaerythritol were used as branching units and glycoside phosphoramidites were used for insertion of the 5 terminal glycosyl groups. In contrast, the protocols in this unit are based on solution-phase synthesis of a nonnucleosidic phosphoramidite building block bearing two phthaloyl-masked aminooxy groups, the incorporation of this building block into oligonucleotides by conventional phosphoramidite chemistry, and solid-phase oximation of the deblocked aminooxy groups with glycosyl aldehydes. The final deprotection and cleavage from the solid support is achieved by conventional ammonolysis. This methodology allows variation of the number, site, and identity of the sugar ligands, and hence is suitable for the creation of conjugate libraries. Because the synthesized building block allows normal
oligonucleotide chain assembly, the distance between the sites of glycosyl attachment may be tuned with additional nucleosidic and nonnucleosidic phosphoramidite building blocks. Conversion of various sugars to peracetates and their subsequent glycosidation with a diol and oxidation to an aldehyde offers a straightforward method for conjugation of any desired sugar by simple and efficient oxime formation. The oxime conjugates are highly water soluble and stable at physiological pH, making them applicable to biological studies. Synthesis of the non-nucleosidic phosphoramidite building block Building block S.6 is synthesized from commercially available diethyl 2,2bis(hydroxymethyl)malonate (S.1). It has been shown previously (Guzaev et al., 1996) that S.1 and its mono-4,4 -dimethoxytrityl (DMTr) derivative are decomposed by release
Modified Oligonucleotides and Conjugates
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of formaldehyde when treated with primary amines. Accordingly, it is necessary to protect both hydroxyl groups of S.1 prior to aminolysis with 3-aminopropanol to obtain S.3. Use of an orthoester protection for this purpose ensures easy deblocking by acid-catalyzed hydrolysis under mild conditions. Using an excess of 3-aminopropanol, the aminolysis of S.2 to the desired diol is almost quantitative. The Mitsunobu reaction (Mitsunobu, 1981) used for introduction of the phthaloylprotected aminooxy groups is the difficult step of the synthesis. Treatment of S.3 with a mixture of N-hydroxyphthalimide, triphenylphosphine, and diethyl azodicarboxylate over a period of 24 hr results in complete disappearance of the starting material without any marked formation of side products. The purification of the di-O-phthalimidooxy product is, however, problematic. Traces of triphenylphosphine oxide formed during the reaction tend to comigrate with the product even after repeated purifications by column chromatography on silica gel. To avoid extensive loss of the target compound, the crude product should be directly subjected to acid-catalyzed hydrolysis to remove the methoxymethylene protecting group. The resulting bis(hydroxymethyl) derivative S.4 can then be more easily separated from impurities by column chromatography on silica gel, although in a low yield. One of the hydroxy groups of S.4 is finally selectively protected as a monoDMTr ether (S.5) using an equivalent amount of DMTr-Cl. The remaining hydroxyl is phosphitylated with 2-cyanoethyl N,N-diisopropylphosphonamidic chloride to obtain S.6. The yield of dimethoxytritylation cannot be increased by using excess DMTr-Cl, since this leads to increased formation of the bis(dimethoxytritylated) product.
An AminooxyFunctionalized Non-Nucleosidic Phosphoramidite for Oligonucleotide Glycoconjugates
Synthesis of the glycosyl aldehyde ligand The glycosyl aldehyde ligand used for the solid-phase oximation is obtained by an easy two-step procedure. A fully acetylated sugar ligand is first subjected to conventional boron trifluoride etherate–promoted glycosidation (Salvador et al., 1995) with an α,ωalkanediol. MeCN appears to be the solvent of choice for this transformation, since less polar CH2 Cl2 or CHCl3 dissolve the diol poorly, leading to extensive formation of monoacylated diol as a byproduct whose mobility is similar to the desired product. The same side reaction takes place when either the reaction time is prolonged or the diol is used in excess. Despite the relatively low yield of this reac-
tion, the use of an unprotected diol shortens the route to the desired target compound, and hence increases the overall yield. In addition, the unreacted sugar peracetate can easily be recycled. The second step, a Swern oxidation of the hydroxy function to an aldehyde group, is a smooth and high-yielding reaction, as expected (Mancuso et al., 1978). Synthesis of the oligonucleotide conjugate and its hybridization properties The present protocol describes preparation of a multivalent oligonucleotide glycoconjugate containing four α-Dmannopyranosyl units. The key steps are introduction of non-nucleosidic bis(hydroxymethyl)malondiamide-derived phosphoramidites, each bearing two phthaloyl-masked aminooxy groups, into the oligonucleotide chain, and oximation of their deblocked aminooxy groups with a glycosyl aldehyde on the support. A standard phosphoramidite protocol can be applied to the coupling of the non-nucleosidic units, although a prolonged coupling time (600 sec) is needed to obtain a sufficiently high coupling yield (>95%). The aminooxy functions may be deblocked without difficulty by a hydrazinium acetate treatment. This treatment does not cleave the succinyl linker (Salo et al., 1999). Oximation of the exposed aminooxy groups with a large excess of peracetylated sugar aldehyde building block in MeCN proceeds essentially to completion within 16 hr at ambient temperature. The oxime conjugate may then be released in solution by normal ammonolysis, which simultaneously deprotects the base, sugar, and phosphate moieties of the conjugate. No signs of hydrolysis or degradation products can be observed by RP-HPLC analysis of the crude product (see Fig. 4.26.4). After normal RP-HPLC purification, the identity of the conjugate may be further verified by ESI-MS analysis. It should be noted that oxidation of the primary hydroxyl group of an otherwise protected sugar to an aldehyde group may appear to be an attractive alternative, but this is not recommended. Although such an aldehyde is easily obtained and solid-supported oximation using the aldehyde is high-yielding, upon standard ammonolytic cleavage, the sugar ligand evidently undergoes dehydration to a 4,5-ene derivative, in all likehood due to conjugation of the 4,5 double bond with the oxime group. Melting studies show that the synthesized oligonucleotide glycoconjugate forms a
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stable duplex with a complementary 12-mer oligodeoxyribonucleotide when the glycosylated 5 terminus falls outside the duplex. The thermal stability of this duplex is only moderately lower than that of the respective fully complementary 12-mer duplex. The melting points were observed to be 44.9◦ C and 48.0◦ C, respectively. More marked destabilization was observed, as expected, on using a complementary 18-mer oligodeoxyribonucleotide carrying an A7 sequence at the 3 terminus, i.e., opposite the glycosylated part of the conjugate. A melting point of 41.9◦ C was observed for hybridization with the glycosylated conjugate.
Compound Characterization Basic Protocols 1 and 2. Chemical characterization data are provided for all compounds in the appropriate protocol steps. The NMR spectra were recorded on a Bruker 200 or JEOL JNM-GX 400- or 500-MHz spectrometer in CDCl3 . The chemical shifts (δ) are given in ppm from internal tetramethylsilane, and the coupling constants (J) are given in hertz (Hz). Mass spectra were recorded on an Applied Biosystems Mariner System 5272 using electrospray ionization (ESI). Basic Protocol 3. Oxime conjugates were characterized by ESI-MS on an Applied Biosystems Mariner System 5272. The melting curve (absorbance versus temperature) for S.12 in hybridization buffer was measured at 260 nm on a Perkin-Elmer Lambda 2 UV-Vis spectrometer equipped with Peltier temperature controller.
Critical Parameters and Troubleshooting The phosphoramidite S.6 and the sugar aldehyde ligand S.9 may be prepared starting from simple and inexpensive starting materials via a fairly straightforward selective transformation of functional groups. The individual steps are based on rather conventional chemistry. Nevertheless, most of the reactions are conducted under anhydrous conditions and, hence, careful attention to proper drying of the solvents and glassware is essential. Preparation of the compounds requires experience with routine chemical laboratory techniques such as extraction, TLC (APPENDIX 3D), and silica gel column chromatography (APPENDIX 3E). Knowledge of machine-assisted DNA synthesis (APPENDIX 3C) and isolation of the products by HPLC (UNIT 10.5) is needed. Characterization of the compounds demands
knowledge of 1 H, 13 C, and 31 P NMR spectroscopies and electrospray ionization mass spectrometry (ESI-MS; UNIT 10.2).
Anticipated Results Good to moderate yields of the individual steps of the total synthesis of building blocks S.6 and S.9 are expected. The total yield of S.6 may also be increased to some extent by avoiding repetitive column chromatographic purifications after the Mitsunobu reaction. These building blocks allow quite convenient synthesis of multivalent oligonucleotide glycoconjugates on a solid support as described in Basic Protocol 3. The target compound is expected to be clearly the main product, and its isolation by RP-HPLC is therefore fairly straightforward.
Time Considerations The synthesis of the phosphoramidite S.6 starting from diethyl 2,2bis(hydroxymethyl)malonate (S.1) can be accomplished in 2 weeks (including five overnight incubations) and the synthesis of the sugar ligand S.9 within 2 to 3 days. The time needed for machine-assisted oligonucleotide chain assembly differs from the standard protocol only by the extended coupling time of 600 sec per non-nucleosidic phosphoramidite unit. The phthaloyl deprotection and subsequent oximation by overnight treatment increase the total time of the conjugate synthesis by 24 hr. The times needed for deprotection and cleavage from the support, RP-HPLC purification, and characterization are the same as with standard methods.
Literature Cited Adinolfi, M., Barone, G., De Napoli, L., Guariniello, L., Iadonisi, A., and Piccialli, G. 1999. Solid-phase glycosidation of oligonucleotides. Tetrahedron Lett. 40:2607-2610. Akhtar, S., Routledge, A., and Patel, R. 1995. Synthesis of mono- and dimannoside phosphoramidate derivatives for the solid-phase conjugation to oligonucleotides. Tetrahedron Lett. 36:73337336. De Kort, M., Ebrahimi, E., Wijsman, E.R., Van der Marel, G.A., and Van Boom, J.H. 1999. Synthesis of oligodeoxynucleotides containing 5-(β-D-glycopyranosyloxymethyl)-2 deoxyuridine, a modified nucleoside in the DNA of Trypanosoma Brucei. Eur. J. Chem. 9: 2337-2344. Defrancq, E. and Lhomme, J. 2001. Use of an aminooxy linker for the functionalization of oligodeoxyribonucleotides. Bioorg. Med. Chem. Lett. 11:931-933.
Modified Oligonucleotides and Conjugates
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Dubber, M. and Fr´echet, J.M.J. 2003. Solid-phase synthesis of multivalent glycoconjugates on a DNA synthesizer. Bioconjugate Chem. 14:239246.
Mitsunobu, O. 1981. The use of diethyl azodicarboxylate and triphenylphosphine in synthesis and transformation of natural products. Synthesis 1-28.
Guzaev, A., Salo, H., Azhayev, A., and L¨onnberg, H. 1996. Novel nonnucleosidic building blocks for the preparation of multilabeled oligonucleotides. Bioconjugate Chem. 7:240-248.
Salo, H., Virta P., Hakala, H., Prakash, T.P., Kawasaki, A.M., Manoharan, M., and L¨onnberg, H. 1999. Aminoxy functionalized oligonucleotides: Preparation, on-support derivatization, and post-synthetic attachment to polymer support. Bioconjugate Chem. 10:815823.
Hamma, T. and Miller, P.S. 2003. 4-(2Aminoethoxy)-2-ethylureido)quinolineoligonucleotide conjugates: Synthesis, binding interactions, and derivatization with peptides. Bioconjugate Chem. 14:320-330. Hunziker, J. 1999. Synthesis of 5-(2-amino2-deoxy-D-glucopyranosyloxymethyl)-2 deoxyuridine and its incorporation into oligothymidylates. Bioorg. Med. Chem. Lett. 9:201-204. Katajisto, J., Virta, P., and L¨onnberg, H. 2004. Solidphase synthesis of multiantennary oligonucleotide glycoconjugates utilizing on-support oximation. Bioconjugate Chem. 15:890-896.
Salvador, L.A., Elofsson, M., and Kihlberg, J. 1995. Preparation of building blocks for glycopeptide synthesis by glycosylation of Fmoc amino acids having unprotected carboxyl groups. Tetrahedron 51:5643-5656. Sheppard, T.L., Wong, C., and Joyce, G.F. 2000. Neoglycoconjugates: Design and synthesis of a new class of DNA-carbohydrate conjugates. Angew. Chem. Int. Ed. Engl. 39:3660-3663.
Mancuso, A.J., Huang, S., and Swern, D. 1978. Oxidation of long-chain and related alcohols to carbonyls by dimethyl sulfoxide “activated” by oxalyl chloride. J. Org. Chem. 43:2480-2482.
Wang, L., Prakash, R.K., Stein, C.A., Koehn, R.K., and Ruffner, D.E. 2003. Progress in the delivery of therapeutic oligonucleotides: Organ/cellular distribution and targeted delivery of oligonucleotides in vivo. Antisense Nucleic Acid Drug Dev. 13:169-189.
Matsuura, K., Hibino, M., Yamada, Y., and Kobayashi, K. 2001. Construction of glycoclusters by self-organization of site-specifically glycosylated oligonucleotides and their cooperative amplification of lectin-recognition. J. Am. Chem. Soc. 123:357-358.
Contributed by Johanna Katajisto, Pasi Virta, and Harri L¨onnberg University of Turku Turku, Finland
An AminooxyFunctionalized Non-Nucleosidic Phosphoramidite for Oligonucleotide Glycoconjugates
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Large-Scale Preparation of Conjugated Oligonucleoside Phosphorothioates by the High-Efficiency Liquid-Phase (HELP) Method
UNIT 4.27
This unit describes the soluble-polymer-supported synthesis of phosphorothioated oligonucleotides and their postsynthetic conjugation with a new chain of high-molecularweight polyethylene glycol (PEG) or other polymers. The protocols give all the details of synthesis, deprotection, isolation, and modification of the oligonucleotides. Basic Protocol 1 outlines the step-by-step procedure of the liquid-phase process used for assembling the thioated oligonucleotide chain. The procedure is similar to classical solidphase synthesis, with the exception that all the reactions are performed in homogeneous solutions. Purification of intermediates, however, is very similar to methods performed in processes conducted on an insoluble support; the insolubility of PEG in diethyl ether or ethanol allows the selective precipitation of the supported nucleic acid product and the effective removal of all soluble byproducts. Alternate Protocol 1 describes the assembly of the chain using preassembled dimers to allow production of the oligonucleotides on a larger scale. The reduced number of procedural steps provides an increase in the yield of the final product. Basic Protocol 2 describes the introduction of a linker with a reactive amino-terminal function on the PEG-supported oligonucleotides and the final detachment of the modified oligonucleotides from the support. This is required because the terminal hydroxyl moiety of the oligonucleotide does not guarantee a successful condensation reaction with the large polymeric chain, whose reactivity generally decreases with increasing molecular weight. Basic Protocol 3 describes the conjugation of the modified oligonucleotide with a properly activated high-molecular-weight PEG in solution. The most convenient procedures for activation of the terminal hydroxyl of PEG are outlined. Alternate Protocol 2 describes a different method for activation of the PEG moiety to enable conjugation. This process, although less effective, can be used for conjugation should the one in Basic Protocol 3 not be implemented. ◦
NOTE: Store all anhydrous solvents under argon, over activated 4-A molecular sieves (<20 ppm water). Dry glassware to be used for oligonucleotide synthesis in an oven at 110◦ C and store in a desiccator over KOH pellets prior to use. Store all amidite solution bottles over desiccant in polypropylene containers. Allow all cold bottles to warm to room temperature before use.
PREPARATION OF PHOSPHOROTHIOATED OLIGONUCLEOTIDES VIA HELP This process, called high-efficiency liquid phase, or HELP, consists of three main steps: (1) functionalization of a soluble support with a 5 -DMTr-nucleoside via a succinyl linkage, (2) elongation of the oligonucleotide, and (3) cleavage of the succinyl linkage and removal of all protecting groups. Many advantages arise from this synthetic process. The
Contributed by Gian Maria Bonora Current Protocols in Nucleic Acid Chemistry (2005) 4.27.1-4.27.26 C 2005 by John Wiley & Sons, Inc. Copyright
BASIC PROTOCOL 1
Synthesis of Modified Oligonucleotides and Conjugates
4.27.1 Supplement 22
Figure 4.27.1 Preparation of the 5 -O-DMTr-nucleoside-3 -hemisuccinate and subsequent functionalization of MPEG.
oligonucleotide synthesis is carried out in homogeneous medium on a soluble polymeric support. The polymer-bound product is recovered from the reaction mixture by a simple precipitation that allows rapid and easy elimination of excess reagents and soluble byproducts. Preparation of the nucleoside succinate and functionalization of the support are illustrated in Figure 4.27.1. Polyethylene glycol (PEG) is used as a soluble support for oligonucleotide synthesis. The molecular weight of the PEG ranges from 5 to 20 kDa, depending on the final size of the desired oligonucleotide. In the process described herein, a 10kDa monomethoxy-PEG (MPEG) is employed because the monofunctional derivative is easier to handle during the synthetic procedures. A commercial bifunctional PEG may be used to increase production of the final product from the same molar amount of PEG support; however, this may lead to unpredictable results as a consequence of interference between two oligonucleotide chains growing on the same support. The well-established phosphoramidite-based chemistry is successfully employed in this procedure. The iterative coupling process is illustrated in Figure 4.27.2. As a general observation, the phase homogeneity allows for easy scaling up of the process and, in principle, requires smaller amounts of reagents because of the absence of the inherent diffusion problems that occur with insoluble resin beads. It is also worth noting that the spectral transparency and high solubility of the conjugate allow for a rapid and nondestructive spectrophotometric analysis of any synthetic step. The ammonia treatment at the end of this protocol cleaves the succinyl linkage with MPEG, removes cyanoethyl protecting groups from the phosphorothioate backbone, and removes any standard base-protecting groups, yielding a free, crude oligonucleotide. Should an additional conjugation reaction be performed on the support-bound oligonucleotide (see Basic Protocol 2 and 3), the cleavage process will be performed at the end of that reaction.
Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
Purification of the conjugated oligonucleotide is achieved by ion-exchange chromatography because the separation of the final conjugated oligonucleotide from the shorter failure sequences cannot be accomplished be reversed-phase chromatography. A broad, single peak is commonly observed due to the presence of the large PEG chain, which dominates the chromatographic behavior of the supported products.
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Figure 4.27.2 HELP synthesis cycle for phosphorothioated oligonucleotides. Abbreviations: B, base (thymin-1-yl, N 4 benzoylcytosin-1-yl, N 6 -benzoyladenin-9-yl, N 2 -isobutyrylguanin-9-yl); DDD, diethoxydithiocarbonate disulfide; DMTr, 4,4 dimethoxytrityl; DNA, synthetic oligonucleoside phosphorothioate triester; MPEG, polyethylene glycol monomethyl ether; NMI, N-methylimidazole; TCA, trichloroacetic acid; DCE, 1,2-dichloroethane.
Materials 5 -O-(4,4 -Dimethoxytrityl)-protected nucleoside (for attachment to support; Fig. 4.27.1; Glen Research) ◦ Pyridine, dried using 4-A molecular sieves 4-Dimethylaminopyridine (DMAP) Succinic anhydride, recrystallized from chloroform Argon source Chloroform Ethanol (EtOH) 60% perchloric acid Toluene, anhydrous Dichloromethane, anhydrous 10% (w/v) citric acid, ice cold Na2 SO4 , anhydrous n-Hexane Diethyl ether, anhydrous KOH pellets Kieselgel 60, 70-230 mesh (Merck) Methanol (MeOH) ◦ 1,2-Dichloroethane (DCE), dried using 4-A molecular sieves
Synthesis of Modified Oligonucleotides and Conjugates
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N,N -Dicyclohexylcarbodiimide (DCC) Polyethylene glycol monomethyl ether 10,000 (MPEG10000 ; Nektar Therapeutics) N-Methylimidazole (NMI) Acetone Ethyl acetate Iodine (I2 ), crystalline Methyl tert-butyl ether (MTBE) ◦ Acetonitrile (MeCN), dried using 4-A molecular sieves 2,6-Lutidine Acetic anhydride, distilled over anhydrous sodium acetate 6% (w/v) trichloroacetic acid (TCA) in dry DCE (store at 4◦ C in a dark bottle) 0.2 M base-protected 5 -O-(4,4 -dimethoxytrityl)-3 -O-(2-cyanoethyl-N,Ndiisopropyl) nucleoside phosphoramidites (dABz -CE, dCBz -CE, dGiBu -CE and dT-CE; Glen Research) in MeCN (see recipe for phosphoramidites) 0.5 M 1H-tetrazole in MeCN (see recipe) 0.5 M DDD in MeCN (see recipe) Concentrated (∼30%) ammonium hydroxide 80% (v/v) acetic acid 250-mL three-neck round-bottom flasks Rubber septa Calcium chloride drying tubes Precoated silica gel 60 TLC sheets (5 × 10 cm) with fluorescent indicator (F254 ; Merck) UV lamp (UVP Mineralight lamp UVG-11, 254 nm) 25- and 250-mL separatory funnels Filter paper Rotary evaporator with water aspirator as vacuum source Sintered glass filters (Gooch G-3 and G-4) Vacuum desiccator 100 × 35–mm column for flash chromatography 100-mL and 250-mL round-bottom flasks Spectrophotometer Three-way stopcock High-vacuum oil pump Syringes and needles Glass container with tight seal 60◦ C water bath Lyophilizer Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare 5 -DMTr-nucleoside-3 -hemisuccinate 1. Coevaporate 5 mmol of the appropriate 5 -DMTr-protected nucleoside twice with 10 mL dry pyridine in a 250-mL three-neck round-bottom flask sealed with rubber septa. Attach a calcium chloride drying tube and dissolve the final residue in 20 mL dry pyridine. 2. Add 0.5 eq DMAP and 1 eq succinic anhydride slowly over a 30-min period, with stirring. Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
3. Protect the solution against light and stir under argon atmosphere for 24 hr at room temperature.
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4. Monitor the reaction by TLC (APPENDIX 3D) on precoated silica gel 60 F254 sheets using 90:10:0.05 (v/v/v) chloroform/ethanol/pyridine as eluent. Visualize spots using a 254-nm UV lamp and also by spraying plates with 3:2 (v/v) 60% perchloric acid/ethanol. Pyridine is included in the eluent to avoid detritylation. After spraying with acid solution, DMTr-bearing compounds appear as orange-colored spots, with the product running more slowly than the starting material. An additional orange spot running with the solvent front is due to some DMTr-OH byproduct originating from the hydrolysis of the starting DMTr-Cl.
5. Using a rotary evaporator and a water aspirator, evaporate the solution to dryness and then coevaporate twice with 50 mL toluene to remove traces of pyridine. Dissolve the thick oil in 20 mL dichloromethane and wash it in a 250-mL separatory funnel with 40 mL ice-cold 10% (w/v) citric acid. 6. Extract the aqueous solution twice with 50 mL dichloromethane, pool the organic layers, and wash twice with 50 mL water. Dry the organic layer over 5 to 10 g anhydrous Na2 SO4 , filter off the drying agent under gravity using filter paper, and evaporate to dryness using a rotary evaporator. 7. Dissolve the residue in 20 mL of 0.5% (v/v) pyridine/dichloromethane and add it slowly, with stirring, to 300 mL of a 1:1 (v/v) mixture of n-hexane and anhydrous diethyl ether cooled in an ice bath. Filter the white powder through a Gooch G-4 sintered glass filter and dry under vacuum in a vacuum desiccator over KOH pellets. 8. Purify the crude product by flash chromatography (APPENDIX 3E) using a 100 × 35– mm silica gel (Kieselgel 60, 70-230 mesh) column and a stepwise gradient of 0% to 10% methanol in chloroform containing 0.5% pyridine. Check each fraction by TLC (see step 4). Store the purified product (5 -DMTr-nucleoside-3 -hemisuccinate) at 4◦ C.
Functionalize MPEG with 5 -DMTr-nucleoside-3 -hemisuccinate 9. Coevaporate the 5 -DMTr-nucleoside-3 -hemisuccinate (0.3 mmol) twice with 10 mL dry pyridine, and dissolve the residue in 5 mL dry 0.5% (v/v) pyridine/DCE in a 100-mL round-bottom flask equipped with a calcium chloride drying tube. Always coevaporate the starting materials completely when reactions are performed with MPEG or MPEG-supported derivatives. MPEG is very hygroscopic!
10. Cool the solution in an ice bath, then add 0.15 mmol DCC with stirring. Continue stirring for 15 min. 11. While stirring, prepare a 100-mL round-bottom flask containing 1.0 g (0.1 mmol) MPEG10000 , previously coevaporated twice with 10 mL dry pyridine and dissolved in 5 mL dry 0.5% (v/v) pyridine/DCE. After stirring, filter the solution from step 10 through a G-3 sintered glass filter into this MPEG10000 solution. 12. Add 0.15 mmol NMI with stirring, protect the solution against light, and stir under an argon atmosphere overnight at room temperature. Monitor the reaction by TLC on precoated silica gel 60 F254 sheets using 10:5:1 (v/v/v) acetone/ethyl acetate/water as eluent. Visualize spots using a 254-nm UV lamp and by exposure to iodine, introduced as solid crystals at the bottom of the developing chamber, to detect MPEG-bearing compounds as a brown spot. Alternatively, visualize the DMTrbearing compounds with perchloric acid solution (see step 4). MPEG-bound derivatives always show an Rf = 0.
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13. Filter the reaction mixture through a G-3 sintered glass filter into a 250-mL roundbottom flask and wash the solid with 5 mL of dry 0.5% (v/v) pyridine/DCE. Cool the solution in an ice bath and add 70 mL MTBE dropwise over a period of 30 min with vigorous stirring. 14. Filter the white powder through a G-3 glass filter. Wash the solid extensively with anhydrous diethyl ether (usually three times using 10 mL each time). Dry under vacuum in a vacuum desiccator over KOH pellets. 15. Suspend the crude product in 10 mL dry MeCN, filter through a G-4 glass filter, then wash the solid residue with 5 mL dry MeCN. Reduce the volume of the clear solution to 5 mL under vacuum, then cool the concentrate in an ice bath. Add 70 mL MTBE dropwise over 30 min with vigorous stirring. 16. Filter the white powder through a G-3 glass filter, wash the solid extensively with anhydrous diethyl ether, and dry it under vacuum in a vacuum desiccator over KOH pellets. Store the product (5 -DMTr-nucleoside-3 -succ-MPEG) at 4◦ C. The precipitation and filtration of all MPEG derivatives must be executed slowly and accurately to avoid any loss of material.
17. To measure the degree of functionalization, dissolve a 5-mg aliquot of product in 10 mL of 3:2 (v/v) 60% perchloric acid/ethanol, dilute the resulting solution 10fold with this same acid/ethanol solution, and read the absorbance at 498 nm in a spectrophotometer. Calculate the loading of MPEG from the equation: loading (µmol/g) = [A498 × 10 × 14.3]/mg weighed support where 10 is the cell path length (in mm) and 14.3 is the extinction coefficient at 498 nm (on a mg scale).
Acetylate functionalized MPEG (capping step) 18. Dissolve the 5 -DMTr-nucleoside-3 -succ-MPEG in 5 mL dry MeCN in a 100-mL round-bottom flask equipped with a calcium chloride drying tube. All volumes in this and subsequent steps are based on the starting amount of MPEG (1.0 g) in step 11. To proceed with different amounts of MPEG, all volumes should be scaled appropriately.
19. Cool the solution in an ice bath and add, with stirring, 0.5 mL 2,6-lutidine, 0.5 mL NMI, and 0.5 mL acetic anhydride. Let stand 3 min at 0◦ C. 20. Add 70 mL MTBE dropwise over a period of 10 to 15 min. Filter the reaction mixture through a G-3 glass filter and wash the precipitate extensively with anhydrous diethyl ether. Dry the solid under vacuum in a vacuum desiccator over KOH pellets. Store the product at 4◦ C. Capping is required to irreversibly block any unreacted MPEG or MPEG-nucleoside conjugates.
Detritylate 5 -DMTr-nucleoside-3 -succ-MPEG 21. Dissolve the 5 -DMTr-nucleoside-3 -succ-MPEG in 10 mL dry DCE in a 250-mL round-bottom flask equipped with a calcium chloride drying tube. Add 10 mL of 6% (w/v) TCA in DCE with vigorous stirring. Continuing stirring 15 min. Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
If the 5 -terminal residue is a purine (dG or dA), cool the solution on ice before adding the acidic solution, to avoid any depurination.
22. Precipitate the deprotected nucleoside-3 -succ-MPEG by dropwise addition of 70 mL MTBE at 0◦ C. Filter the solid product through a G-3 glass filter and wash it extensively with anhydrous diethyl ether.
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23. To ensure complete detritylation, repeat TCA treatment (steps 21 to 22). Since incomplete (97% to 98%) detritylation sometimes occurs, the TCA treatment is always repeated.
24. Dry the solid under vacuum in a vacuum desiccator over KOH pellets. Store the product at 4◦ C.
Couple with 5 -DMTr-nucleoside-3 -phosphoramidite 25. Place the nucleoside-3 -succ-MPEG in a 250-mL three-neck round-bottom flask and seal the flask with rubber septa and a three-way stopcock. Coevaporate three times with 10 mL anhydrous MeCN and dry the material under vacuum using a highvacuum oil pump. Flush with argon for 4 to 5 min, then dissolve the dried residue in the minimum amount of MeCN required (usually <2 to 3 mL). 26. While stirring, add 3 eq (0.3 mmol) of the 5 -DMTr-protected nucleoside phosphoramidite to be coupled (from a 0.2 M solution in MeCN) and 10 eq (1.0 mmol) 1H-tetrazole (from a 0.5 M solution in MeCN) simultaneously via syringes through the septum. Stir the solution under an argon atmosphere for 5 min at room temperature. 27. Cool the reaction mixture in an ice bath and add 70 mL MTBE dropwise over a period of 10 to 15 min with vigorous stirring. Filter the white powder through a G-3 glass filter, rapidly wash it with ether, and dry it under vacuum in a vacuum desiccator over KOH pellets. Store the product at 4◦ C. The use of freshly prepared amidite solution (maximum 1 to 2 weeks old and stored in a freezer over molecular sieves) is a critical step for a good yield of the condensation reaction.
Sulfurize the phosphate triester group 28. Dissolve the 5 -DMTr-oligonucleotide-3 -succ-MPEG in 5 mL dry MeCN in a 100-mL round-bottom flask equipped with a calcium chloride drying tube. 29. While stirring, add 10 eq (1 mmol) DDD from a 0.5 M solution in MeCN. Continue stirring for 15 min at room temperature. 30. Add 70 mL ice-cold MTBE dropwise over a period of 10 to 15 min with vigorous stirring. Filter the white precipitate through a G-3 glass filter, wash it with anhydrous diethyl ether, and redissolve it in 5 mL dry MeCN. 31. Repeat precipitation by adding 70 mL ice-cold MTBE. Filter the precipitate through a G-3 glass filter, rapidly wash with anhydrous diethyl ether, and dry under vacuum in a vacuum desiccator over KOH pellets. 32. Dissolve the crude product in 100 mL ethanol, warming the solution up to a maximum of 38◦ C until the solid is completely dissolved. Recrystallize by letting the solution cool down in a 4◦ C refrigerator for 1 hr. 33. Filter the resulting white powder through a G-3 glass filter under vacuum and repeat crystallization until TLC analysis using 90:10:0.05 (v/v/v) chloroform/ethanol/pyridine does not reveal any orange spot at Rf > 0, indicating that all excess reagents have been removed. 34. Filter again, dry under vacuum in a vacuum desiccator over KOH pellets, and store the product at 4◦ C. Synthesis of Modified Oligonucleotides and Conjugates
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Supplement 22
Acetylate 5 -DMTr-oligonucleotide-3 -succ-MPEG (capping) 35. Dissolve the sulfurized 5 -DMTr-oligonucleotide-3 -succ-MPEG in 5 mL dry MeCN in a 100-mL round-bottom flask and cool it in an ice bath. Add, with stirring, 0.5 mL 2,6-lutidine, 0.5 mL NMI, and 0.5 mL acetic anhydride. Leave the solution at 0◦ C for 3 min. 36. Precipitate the capped product from its ice-cold solution by dropwise addition of 70 mL MTBE. 37. Filter the mixture through a G-3 glass filter and wash the precipitate extensively with anhydrous diethyl ether. 38. Dry the product under vacuum in a vacuum desiccator over KOH pellets. Store at 4◦ C. 39. Repeat steps 21 to 38 for each additional nucleotide residue until the desired sequence is completed.
Cleave succinyl linkage and remove all protecting groups 40. Dissolve the completed 5 -DMTr-oligonucleotide-3 -succ-MPEG in 5 mL of 30% ammonium hydroxide for each 20 mg product in a tightly closed glass container and leave the solution without stirring for 10 hr at 60◦ C. The reaction time in the ammonia solution is reduced from the usual 24 hr to only 10 hr, as a better-quality product is thus obtained.
41. Evaporate the solution to dryness, rinse with water, and evaporate again. Repeat the procedure until the material is free of any ammonia odor. 42. Dissolve the residue in 5 mL water and extract four times, each time with 5 mL anhydrous diethyl ether in a 25-mL separatory funnel. 43. Lyophilize the aqueous layer and dissolve the residue in 5 mL of 80% acetic acid for each 20 mg product in a 100-mL round-bottom flask. Stir for 30 min at room temperature. 44. Extract the resulting yellow solution five times with 5 mL anhydrous diethyl ether in a 25-mL separatory funnel, and evaporate the aqueous layer to dryness using a high-vacuum oil pump. 45. Rinse the residue with water, evaporate, and repeat the rinsing and evaporation until the material is free of acetic acid odor. 46. Dissolve the residue in water and lyophilize. Store at −15◦ C. 47. To utilize the crude oligonucleotide, separate it from the MPEG by suspending 1 g of the lyophilized residue in 10 mL acetone or MeOH and filtering the oligonucleotide on a G-4 glass filter under vacuum. MPEG is highly soluble in acetone and methanol. If further purification is desired, see Support Protocol. ALTERNATE PROTOCOL 1
Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
PREPARATION OF PHOSPHOROTHIOATED OLIGONUCLEOTIDES VIA HELP USING DIMER BLOCKS One of the main features of this protocol is the use of a “blockmer” strategy. The use of dimers may reduce the levels of shorter deletion sequences (failures) that are produced during synthesis. The (n–1)mer is especially difficult to purify using standard reversed-phase HPLC. Since incomplete coupling and sulfurization may be the reasons for its formation, one way to reduce this and similar byproducts is to use preformed dimeric units in the stepwise synthesis of the oligomer. An important advantage in the
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use of dimeric units is the reduction of the number of steps required for oligonucleotide synthesis. When performing the liquid-phase procedure, all the intermediate purifications are achieved through a precipitation and filtration procedure that leads to a small but unavoidable loss of product. This loss becomes quite important when several operations are repeated, as in the synthesis of longer sequences. The synthesis of protected dimers is performed in a classical solution procedure using the standard phosphoramidite method (Beaucage and Caruthers, 1981). Dimer phosphoramidites used by the author were prepared by Isis Pharmaceuticals as part of a collaborative project and are not, to the author’s knowledge, commercially available. This protocol describes the detailed procedure for the synthesis of d(GPS GPS TPS T) starting from a cyanoethyl-protected dimer. Continued elongation of the chain follows the same sequence of reactions up to the desired sequence length.
Additional Materials (also see Basic Protocol 1) 5 -DMTr-thymidylyl-(3 →5 )-thymidine (2-cyanoethyl)thiophosphate (5 -DMTr-d(TPS(CE) T) dimer) 0.2 M 5 -DMTr-d(GiBu PS(CE) GiBu )-3 -phosphoramidite in MeCN Polyethylene glycol monomethyl ether 5000 (MPEG5000 ; Nektar Therapeutics) 250-mL and 1-L round-bottom flasks 1-L three-neck round-bottom flasks Synthesize 5 -DMTr-d(TPS(CE) T)-3 -succ 1. Coevaporate 3 g (3.3 mmol) 5 -DMTr-d(TPS(CE) T) dimer twice with 20 mL dry pyridine in a 250-mL three-neck round-bottom flask sealed with rubber septa. Attach a calcium chloride drying tube and dissolve the residue in 14 mL dry pyridine. 2. Add 0.202 g (1.65 mmol) DMAP and 0.33 g (3.3 mmol) succinic anhydride slowly over a 30-min period, with stirring. 3. Protect the solution against light and stir under argon atmosphere for 24 hr at room temperature. 4. Monitor the reaction by TLC (APPENDIX 3D) on precoated silica gel 60 F254 sheets using 90:10:0.05 (v/v/v) chloroform/ethanol/pyridine as eluent. Visualize spots using a 254-nm UV lamp and also by spraying plates with 3:2 (v/v) 60% perchloric acid/ethanol. Pyridine is included in the eluent to avoid detritylation. After spraying with acid solution, DMTr-bearing compounds appear as orange-colored spots, with the product running more slowly than the starting material. An additional orange spot running with the solvent front is due to some DMTr-OH byproduct originating from the hydrolyisis of the starting DMTr-Cl.
5. Using a rotary evaporator and a water aspirator, evaporate the solution to dryness and then coevaporate three times with 20 mL toluene to remove traces of pyridine. Dissolve the thick oil in 30 mL dichloromethane and wash it in a 250-mL separatory funnel with 60 mL ice-cold 10% (w/v) citric acid. 6. Extract the aqueous solution twice with 20 mL dichloromethane, pool the organic layers, and wash twice with 50 mL water. Dry the organic layer over 5 to 10 g anhydrous Na2 SO4 , filter off the drying agent under gravity using filter paper, and evaporate the filtrate to dryness using a rotary evaporator. 7. Dissolve the thick oil in 14 mL of 0.5% (v/v) pyridine/dichloromethane and add it slowly, with stirring, to 140 mL of a 1:1 (v/v) mixture of n-hexane and anhydrous diethyl ether cooled in an ice bath. Filter the white powder through a Gooch G-3 glass sintered funnel and dry under vacuum in a vacuum desiccator over KOH pellets. Store the product at 4◦ C. Current Protocols in Nucleic Acid Chemistry
Synthesis of Modified Oligonucleotides and Conjugates
4.27.9 Supplement 22
8. Purify the crude product by flash chromatography (APPENDIX 3E) using a 100 × 35– mm silica gel (Kieselgel 60, 70-230 mesh) column and a stepwise gradient of 0% to 10% methanol in chloroform containing 0.5% pyridine. Check each fraction by TLC (see step 4). 9. Pool product-containing fractions, evaporate to dryness using a high-vacuum oil pump, and dissolve in 10 mL dichloromethane. 10. Add dropwise, with stirring, to 100 mL cold (5◦ C) 1:1 n-hexane/anhydrous diethyl ether over a period of 5 to 10 min. 11. Filter the white powder through a G-4 sintered glass funnel and dry under vacuum in a vacuum desiccator over KOH pellets. Store the product at 4◦ C. From 3.0 g of starting dimer, 1.2 g of purified 5 -DMTr-d(TPS(CE) T)-3 -succ is obtained.
Functionalize MPEG with 5 -DMTr-d(TPS(CE) T)-3 -succ 12. Coevaporate the 5 -DMTr-d(TPS(CE) T)-3 -succ (0.9 mmol) twice with 10 mL dry MeCN, and dissolve the residue in 3 mL dry 0.5% (v/v) pyridine/DCE in a 100-mL round-bottom flask equipped with a calcium chloride drying tube. Always coevaporate the starting materials completely when reactions are performed with MPEG or MPEG-supported derivatives. MPEG is very hygroscopic!
13. Cool the solution in an ice bath, then add 0.5 mmol DCC with stirring. Continue stirring for 15 min. 14. While stirring, prepare a 250-mL round-bottom flask containing 1.5 g (0.3 mmol) MPEG5000 previously coevaporated twice with 10 mL dry MeCN and dissolved in 7.5 mL dry 0.5% (v/v) pyridine/DCE. After stirring, filter the solution from step 13 through a G-3 sintered glass filter into this MPEG5000 . 15. Add 0.6 mmol NMI with stirring, protect the solution against light, and stir overnight under argon atmosphere at room temperature. Monitor the reaction by TLC on precoated silica gel 60 F254 sheets using 10:5:1 (v/v/v) acetone/ethyl acetate/water as eluent. Visualize spots using a 254-nm UV lamp and by exposure to iodine, introduced as solid crystals at the bottom of the developing chamber, to detect MPEG-bearing compounds as a brown spot. Alternatively, visualize the DMTrbearing compounds with perchloric acid solution (see step 4). MPEG-bound derivatives always show an Rf = 0.
16. Filter the solution through a G-4 sintered glass funnel into a 250-mL round-bottom flask, and reduce the reaction volume to 1/3 of its original volume using a rotary evaporator and a water aspirator. 17. Cool the solution in an ice bath and add 70 mL MTBE dropwise with vigorous stirring to the cold solution over a period of 20 min. Filter the white powder through a G-4 sintered glass funnel and wash the solid extensively with anhydrous diethyl ether (usually three times with 10 mL each time). Dry under vacuum in a vacuum desiccator over KOH pellets. 18. Evaluate the complete removal of unreacted excess reagents by TLC as described in step 15. Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
5 -DMTr-d(TPS(CE) T)-3 -succ-MPEG has an Rf = 0.
19. Suspend the crude product in 100 mL EtOH at 38◦ C. Remove any undissolved white residue (unreacted dinucleotide succinate) by filtration through a G-3 sintered glass filter under vacuum, and place the filtrate in a refrigerator at 4◦ C for 2 hr.
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20. Filter the white powder through a G-4 sintered glass funnel. Store the product at 4◦ C. From 1.5 g of starting MPEG, 1.6 g of functionalized polymer is collected.
Acetylate 5 -DMTr-d(TPS(CE) T)-3 -succ-MPEG (capping) 21. Dissolve the 5 -DMTr-d(TPS(CE) T)-3 -succ-MPEG in 8 mL dry MeCN in a 250-mL round-bottom flask and cool in an ice bath. 22. Add 0.8 mL 2,6-lutidine, 0.8 mL NMI, and 0.8 mL acetic anhydride with stirring. Leave the solution for 3 min at 0◦ C. 23. Add 70 mL MTBE dropwise to the cold and vigorously stirred solution. 24. Filter the white powder through a G-4 sintered glass funnel, wash the product extensively with anhydrous diethyl ether, and dry under vacuum in a vacuum desiccator over KOH pellets.
Detritylate 5 -DMTr-d(TPS(CE) T)-3 -succ-MPEG 25. Dissolve 5 -DMTr-d(TPS(CE) T)-3 -succ-MPEG (0.3 mmol) in 50 mL dry DCE in a 1-L round-bottom flask equipped with a calcium chloride drying tube. 26. Add 50 mL of 6% TCA solution in DCE dropwise with vigorous stirring over a period of 15 min. 27. Cool the solution in an ice bath, and precipitate the d(TPS(CE) T)-3 -succ-MPEG by dropwise addition of 350 mL MTBE over 10 to 15 min. Filter the mixture through a G-4 sintered glass funnel and wash the precipitate extensively with anhydrous diethyl ether. 28. Assess completion of detritylation by following the absorbance at 498 nm of the DMTr group (see Basic Protocol 1, step 17) and by observing any orange color after spraying a spot on a TLC sheet with acid/ethanol solution (step 4). If detritylation is incomplete, repeat the TCA treatment (steps 26 to 27). 29. Dry the white powder in a vacuum desiccator over KOH pellets. Eliminate any TCA residue in the fully detritylated product by recrystallization from DCE solution (50 mL) using MBTE (350 mL) as precipitating solvent. Filter product and dry in a vacuum desiccator over KOH pellets. Store product at 4◦ C.
Couple d(TPS(CE) T)-3 -succ-MPEG with dinucleotide phosphoramidite derivative 30. Place d(TPS(CE) T)-3 -succ-MPEG) (0.3 mmol) in a 1-L three-neck round-bottom flask sealed with rubber septa and equipped with a three-way stopcock. Coevaporate three times with 20 mL dry MeCN and dry under vacuum using a high-vacuum oil pump. Flush with argon for 4 to 5 min. 31. Dissolve the dried residue in 13.5 mL MeCN injected with a syringe through the septum. 32. While stirring, add 3 eq (0.9 mmol) 5 -DMTr-d(GiBu PS(CE) GiBu )-3 -phosphoramidite (from a 0.2 M solution in MeCN) and 10 eq (3 mmol) 1H-tetrazole (from a 0.5 M solution in MeCN) simultaneously via syringes through the septum. Stir the solution under argon atmosphere for 5 min at room temperature. 33. Cool the solution in an ice bath and add 350 mL MTBE dropwise with vigorous stirring over 10 to 15 min. Filter the white powder through a G-4 sintered glass funnel, rapidly wash with anhydrous diethyl ether, and dry under vacuum in a vacuum desiccator over KOH pellets. 34. Dissolve in 300 mL of warm (40◦ C) EtOH and leave in a refrigerator for 2 hr. Filter the crystals. Repeat this recrystallization process until TLC with 90:10:0.05 (v/v/v)
Synthesis of Modified Oligonucleotides and Conjugates
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chloroform/ethanol/pyridine does not reveal any orange spot at Rf > 0, indicating that all excess reagent has been removed. 35. Filter and dry the resulting white powder under vacuum in a vacuum desiccator over KOH pellets. Store the product at 4◦ C.
Sulfurize the condensation product 36. Place the condensation product (0.3 mmol) in a 1-L round-bottom flask and dissolve it in 31 mL dry MeCN. While stirring, add 10 eq (3 mmol) DDD from a 0.5 M solution in MeCN. Stir the solution at room temperature for 15 min. 37. Add 400 mL ice-cold MTBE dropwise, with vigorous stirring. 38. Filter the white precipitate through a G-4 sintered glass funnel, wash with anhydrous diethyl ether, and dry under vacuum in a vacuum desiccator over KOH pellets. Store the product (5 -DMTr-d(GPS(CE) GPS(CE) TPS(CE) T)-3 -succ-MPEG) at 4◦ C. Starting from 5.0 g of fully protected MPEG-dimer, 5.6 g of fully protected MPEG-tetramer is obtained. To perform additional couplings, acetylate the MPEG-bound tetramer as in steps 39 to 42, then repeat steps 25 to 38 until the final sequence is obtained.
Acetylate 5 -DMTr-d(GPS GPS TPS T)-3 -succ-MPEG (capping) 39. Dissolve 5 -DMTr-d(GPS(CE) GPS(CE) TPS(CE) T)-3 -succ-MPEG in 30 mL dry MeCN in a 1-L round-bottom flask and cool in an ice bath. 40. Add 3 mL 2,6-lutidine, 3 mL NMI, and 3 mL acetic anhydride with stirring. Leave the solution for 3 min at 0◦ C. 41. Precipitate the product from its ice-cold solution by adding 400 mL MTBE dropwise over 10 to 15 min. 42. Filter the mixture through a G-4 sintered glass funnel, wash the precipitate extensively with anhydrous diethyl ether, and dry under vacuum in a vacuum desiccator over KOH pellets. Store the product at 4◦ C. A chromatogram of an octomer synthesized using this approach is shown in Figure 4.27.3. If further purification is desired, see Support Protocol.
Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
Figure 4.27.3 Analytical IE-FPLC of a fully deprotected octamer synthesized by the dimer blocks procedure (Alternate Protocol 1). (A) crude, (B) FPLC purified. Reprinted from Bonora et al. (2000) with permission from the American Chemical Society.
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PREPARATION OF 5 -AMINOALKYLATED PHOSPHOROTHIOATE OLIGONUCLEOTIDES
BASIC PROTOCOL 2
To overcome the low nucleophilicity of the terminal 5 -OH group of oligonucleotides, and to increase the efficiency of a postsynthetic conjugation with these molecules, a linker carrying a primary amino group is required. This functional group is added to the HELP-synthesized oligonucleotide using a reagent that allows a standard amidite-based procedure and ensures high condensation yields and fast reaction times. It is important to underline that the same conjugation reaction between a high-molecular-weight polymer such as PEG and a biomolecule can, in principle, be extended to peptides or other biologically active molecules with a proper amino function. As described below and illustrated in Figure 4.27.4, a primary amino group is added to the 5 terminus of a fully thioated 15-mer obtained by liquid-phase synthesis on MPEG (HELP technique) following standard detritylation of the crude product. This reaction benefits from the advantages given by the soluble support polymer. As previously described, it is possible to drive the reaction to high yields using an excess of reagents; any unreacted material is easily removed by the usual precipitation and filtration procedures at the end of the reaction. Subsequently, the hydrolysis of the MPEG-oligonucleotide linker will permit the recovery of free aminoalkylated oligonucleotide.
Materials 5 -DMTr-oligonucleotide-3 -succ-MPEG (see Basic Protocol 1) 80% (v/v) acetic acid Diethyl ether, anhydrous ◦ Acetonitrile (MeCN), dried using 4-A molecular sieves 0.5 M 2-[2-(4-monomethoxytrityl)aminoethoxy]ethyl-(2-cyanoethyl)-N,Ndiisopropylphosphoramidite (Glen Research in MeCN) 0.5 M 1H-tetrazole in MeCN (see recipe) Argon source Methyl tert-butyl ether (MTBE) Ethanol (EtOH) KOH pellets TBHP solution: 80% tert-butyl hydroperoxide in 3:2 (v/v) di-tert-butylperoxide/water (Fluka Sigma-Aldrich) Concentrated NH4 OH Acetone 0.03 M 2,4,6-trinitrobenzenesulfonic acid (TNBS) in borate buffer, pH 9.3 Borate buffer, pH 9.3 100-mL round-bottom flask High-vacuum oil pump Lyophilizer 100-ml three-neck round-bottom flask and rubber septa Syringes and needles Sintered glass filters (Gooch G-3 and G-4) Vacuum desiccator Vessel with tight seal 50◦ C oven Refrigerated centrifuge Spectrophotometer Deprotect terminal 5 -hydroxyl group 1. Dissolve the 5 -DMTr-oligonucleotide-3 -succ-MPEG in 80% acetic acid at 5 mL/ 20 mg in a 100-mL round-bottom flask and stir 30 min at room temperature.
Synthesis of Modified Oligonucleotides and Conjugates
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Supplement 22
Figure 4.27.4 Functionalization of a thioated 15-mer DNA oligonucleotide with an aminoalkylated linker at the 5 terminus. Abbreviations: MMTr, 4-monomethoxytrityl; MPEG, polyethylene glycol monomethyl ether; MTBE, methyl tert-butyl ether.
2. Extract the resulting yellow mixture five times with 40 mL anhydrous diethyl ether, and evaporate the aqueous layer to dryness using a high-vacuum oil pump. 3. Dissolve the residue in 25 mL water and evaporate under vacuum using a rotary evaporator and water aspirator. 4. Repeat step 3 until the material is free of acetic acid odor. 5. Dissolve the residue in water and lyophilize.
Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
Conjugate with 5 -amino-modifier phosphoroamidite reagent 6. Coevaporate 1 g oligonucleotide-3 -MPEG (15.6 µmol of 5 -OH groups) three times with 5 mL anhydrous MeCN in a 100-mL three-neck round-bottom flask. 7. Dissolve the residue in 5.0 mL anhydrous MeCN by injecting with a syringe through a rubber septum.
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8. Inject, simultaneously through the septa, 100 µmol of 2-[2-(4-monomethoxytrityl)aminoethoxy]ethyl-(2-cyanoethyl)-N,N-diisopropylphosphoramidite (as a 5 -amino-modifier phosphoramidite) and 400 µmol of 1H-tetrazole (from a 0.5 M solution in MeCN). 9. Stir the mixture under an argon atmosphere 5 min at room temperature. 10. Precipitate the MPEG-bound product from its cold solution by adding 70 mL MTBE with vigorous stirring. Filter the white solid through a G-4 sintered glass filter and wash it with 100 mL anhydrous diethyl ether. 11. Recrystallize the solid from EtOH to remove any excess reagents from the product (see Alternate Protocol 1, step 34). 12. Collect the final product and dry it under vacuum in a vacuum desiccator over KOH pellets. Based on monomethoxytrityl (MMTr) absorbance at 472 nm [ε = 51900 in 70% HClO4 /EtOH 3/2 (v/v)], a functionalization degree of 67% is measured. The product can be reacted again with the 5 -amino-modifier phosphoramidite as described above, with the exception of a prolonged reaction time (15 min). A final 86.5% degree of functionalization is achieved and the product is recovered quantitatively. This procedure is recommended only when a higher yield is needed.
Oxidize phosphite triester function 13. Dissolve the tritylated 5 -aminoalkylated oligonucleotide-3 -MPEG in 7 mL anhydrous MeCN in an ice bath and stir the solution. 14. Add 0.6 mL TBHP solution and stir the reaction 15 min at 0◦ C. 15. Precipitate the product by adding 70 mL MTBE. Collect the white powder, wash it with anhydrous diethyl ether, and dry under vacuum over KOH pellets.
Deprotect phosphate and detach oligonucleotide from MPEG 16. Dissolve oxidized 5 -aminoalkylated oligonucleotide-3 -MPEG in 200 mL concentrated NH4 OH and place the solution in a tightly closed vessel in an oven at 50◦ C for 18 hr. 17. Coevaporate the solution repeatedly with 25 mL distilled water until the material is completely free of ammonia (approximately three times). Dissolve the residue in 100 mL distilled water and extract with four times with 100 mL anhydrous diethyl ether. 18. Lyophilize the aqueous solution and collect the white solid. 19. Remove MPEG by dissolving the solid in 10 mL of 0.5% NH4 OH and precipitating the oligonucleotide by addition of 150 mL acetone with vigorous stirring in an ice bath. 20. Centrifuge 20 min at 4000 rpm (3000 to 4000 × g should be sufficient), 0◦ C. Recover the precipitate containing the ammonium salt of the tritylated 5 -aminoalkylated oligonucleotide (15-mer), and lyophilize it. More than 70% of MPEG is recovered from the acetone solution after a single treatment. From 1.0 g of starting MPEG-oligonucleotide, 0.3 g of modified oligonucleotide can be obtained.
Deprotect terminal amino group 21. Dissolve the tritylated 5 -aminoalkyl-oligonucleotide in 80% acetic acid at 100 mL/g and stir 4 hr at room temperature. A pale yellow color should develop.
Synthesis of Modified Oligonucleotides and Conjugates
4.27.15 Current Protocols in Nucleic Acid Chemistry
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AU
A
26.7 min
13.3
40.0
AU
B
0
5
min
10
15
Figure 4.27.5 RP-HPLC (A) and GPC (B) of the aminoalkylated oligonucleotide. GPC was performed on a PL Aquagel-OH 30 (8 µm, 30 × 0.75–cm) eluted with Milli-Q purified water. The two peaks in panel A are due to intermolecular associations, as suggested by analyses performed under more diluted conditions. Adapted from Ballico et al. (2003) with permission from the American Chemical Society.
22. Add 10 mL distilled water and extract the solution five times with 25 mL anhydrous diethyl ether. Evaporate the aqueous phase to dryness under reduced pressure using a high-vacuum oil pump. Coevaporate with 10 mL distilled water until the material is free of acetic acid. 23. Dissolve in 10 mL distilled water per g residue and lyophilize. A white product should be quantitatively collected. Figure 4.27.5 shows chromatograms of an aminoalkylated oligonucleotide.
Quantitate free NH2 by TNBS test 24. Add 250 µL of 0.03 M TNBS in borate buffer, pH 9.3, to 1 mg of sample. Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
25. Dilute to 10 mL with borate buffer, pH 9.3, stir, and allow to stand 30 min at room temperature.
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26. Read the absorbance at 421 nm (ε = 12860). This test reaction is very useful to determine if the starting MPEG-oligonucleotide is fully modified. The same reaction can be advantageously used to measure any further degree of modification of the starting amino groups.
5 -CONJUGATION OF PHOSPHOROTHIOATED OLIGONUCLEOTIDES WITH HIGH-MOLECULAR-WEIGHT POLYETHYLENE GLYCOLS: ACTIVATION WITH N,N -DISUCCINIMIDYL CARBONATE
BASIC PROTOCOL 3
To obtain a stable linkage between MPEG and an aminoalkylated oligonucleotide, the terminal hydroxyl of the polymer must be activated. As illustrated in Figure 4.27.6, two common procedures involve reaction of MPEG with N,N -disuccinimidyl carbonate (presented here) or with p-nitrophenyl chloroformate (see Alternate Protocol 2). Both procedures produce a reactive intermediate leading to the formation of a stable urethane linkage with the aminoalkylated biomolecule. If performed in organic solution, the conjugation is achieved under heterogeneous conditions, but solubilization of the MPEG-conjugated oligonucleotides can be expected owing to the amphiphilic nature of the polymer. On the other hand, the reaction performed in aqueous solution is characterized by a complete homogeneity, which offers better reactivity. The latter procedure is preferred. The presence of MPEG on the final product offers all the advantages given by the soluble polymer–supported procedures, in particular during the intermediate purifications, as reported in the experimental conditions.
Materials MPEG5000 (Nektar Therapeutics) Toluene, anhydrous Pyridine, anhydrous Dichloromethane (CH2 Cl2 ), anhydrous Acetonitrile (MeCN), anhydrous N,N -Disuccinimidyl carbonate Methyl tert-butyl ether (MTBE) Isopropanol, cold Diethyl ether, anhydrous, cold KOH pellets Ethanol (EtOH) Fully deprotected 5 -aminoalkylated oligonucleotide (see Basic Protocol 2) Na2 CO3 /NaHCO3 buffer, pH 9 Chloroform (CHCl3 ), anhydrous Rotary evaporator Vacuum desiccator G-3 and G-4 sintered glass filters High-vacuum oil pump Prepare MPEG-OSu carbonate 1. Coevaporate 5.0 g (1.0 mmol) of MPEG5000 twice with 15 mL anhydrous toluene per g, and leave the vessel on a rotary evaporator for 30 min to complete dehydration. 2. Add 1.0 mL anhydrous pyridine and dissolve the slurry by adding 5 mL anhydrous CH2 Cl2 and 2 mL anhydrous MeCN.
Synthesis of Modified Oligonucleotides and Conjugates
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Figure 4.27.6 Conjugation of a 5 -aminoalkylated oligodeoxyribonucleoside phosphorothioate (15-mer) with a highmolecular-weight polyethylene glycol. Abbreviations: MPEG, polyethylene glycol monomethyl ether; MTBE, methyl tertbutyl ether.
3. While stirring at room temperature, add 0.13 g of N,N -disuccinimidyl carbonate for each gram of MPEG (2.5 molar equiv). Leave the reaction overnight. 4. Precipitate the product by adding 200 mL MTBE while stirring in an ice bath. Filter on a G-3 glass filter, wash the solid with 25 mL each of cold isopropanol and cold anhydrous diethyl ether, and dry under vacuum in a vacuum desiccator over KOH pellets. 5. Recrystallize MPEG-OSu carbonate from EtOH (see Alternate Protocol 1, step 34). Store product at 4◦ C. A quantitative recovery of MPEG-OSu carbonate is achieved.
Perform conjugation reaction 6. Dissolve 0.1 g (19 µmol) of fully deprotected 5 -aminoalkylated oligonucleotide in 1.0 mL of Na2 CO3 /NaHCO3 buffer, pH 9.0. 7. Add 0.11 g (21 µmol) MPEG-OSu carbonate and leave stirring at room temperature for 72 hr. For oligos of different lengths, the molar amount of oligo in step 6 should be held contast. A 10% molar excess of MPG over oligo should be used.
8. Evaporate the solution to dryness using a high-vacuum oil pump, and dissolve the residue in 20 mL CHCl3 .
Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
9. Filter the insoluble material through a G-4 sintered glass filter and precipitate the product from the filtrate by adding 100 mL MTBE while stirring vigorously in an ice bath. 10. Filter the white powder on a G-4 glass filter, wash it with 25 mL each of cold isopropanol and anhydrous diethyl ether, collect the powder, and dry it under vacuum in a vacuum desiccator over KOH pellets.
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A yield of 0.12 g (0.015 mmol) crude MPEG-conjugated oligonucleotide is usually obtained. This activation gives the best condensation yields with high-molecular-weight PEGs. However, with PEGs larger than 5 kDa, the conjugation requires a large excess of activated PEG (e.g., 2- to 3-fold) to obtain a satisfactory yield.
5 -CONJUGATION OF PHOSPHOROTHIOATED OLIGONUCLEOTIDES WITH HIGH-MOLECULAR-WEIGHT POLYETHYLENE GLYCOLS: ACTIVATION WITH p-NITROPHENYL CHLOROFORMATE
ALTERNATE PROTOCOL 2
This method is an alternative to Basic Protocol 3 that uses the other common activating agent. Both procedures give a reactive intermediate that will form a stable urethane linkage with the NH2 group of the biomolecule. The activation used here appears less efficient than that in Basic Protocol 3, but can be useful as an option.
Additional Materials (also see Basic Protocol 3) p-Nitrophenyl chloroformate Triethylamine (TEA) Ethanol (EtOH) Prepare MPEG-p-nitrophenyl carbonate 1. Coevaporate 5.0 g (1.0 mmol) of MPEG5000 twice with 15 mL anhydrous toluene per g, and leave the vessel on a rotary evaporator for 30 min to complete dehydration. 2. Dissolve the solid residue in 20 mL anhydrous CH2 Cl2 . 3. While stirring, add 0.08 g p-nitrophenyl chloroformate per g MPEG (2 molar equiv) and 0.28 mL TEA per g MPEG (1 molar equiv). 4. Leave the reaction stirring for 24 hr at room temperature. Maintain the pH around 8 by adding TEA. 5. Precipitate the product by adding 200 mL MTBE while stirring in an ice-bath. Filter on a G-3 glass filter, wash the precipitate with 25 mL each of cold isopropanol and diethyl ether, and dry it under vacuum in a vacuum desiccator over KOH pellets. 6. Recrystallize the collected product from EtOH. A quantitative recovery of MPEG-p-nitrophenyl carbonate is achieved.
Perform conjugation reaction 7. Dissolve 0.1 g (19 µmol) of fully deprotected 5 -aminoalkylated oligonucleotide in 1.0 mL Na2 CO3 /NaHCO3 buffer, pH 9.0. 8. Add 0.11 g (21 µmol) of MPEG-p-nitrophenyl carbonate and leave the solution stirring at room temperature for 72 hr. 9. Evaporate the solution to dryness using a high-vacuum oil pump. Suspend the residue in 20 mL CHCl3 and stir for 10 min. 10. Filter the insoluble material through a G-4 glass filter and precipitate the product from the filtrate by additing 100 mL MTBE while stirring vigorously in an ice bath. 11. Filter the white powder on a G-4 glass filter, wash it with 25 mL each of cold isopropanol and anhydrous diethyl ether, collect the powder, and dry it under vacuum in a vacuum desiccator over KOH pellets. Synthesis of Modified Oligonucleotides and Conjugates
4.27.19 Current Protocols in Nucleic Acid Chemistry
Supplement 22
SUPPORT PROTOCOL
PURIFICATION AND ANALYSIS OF OLIGONUCLEOTIDES These methods are used to purify the final product at the end of Basic Protocol 1 and Alternate Protocol 1. NOTE: Prepare all HPLC solvents and buffers with double-distilled water filtered through a Millipore GS 0.22-µm sterile filter.
Materials 0.1 and 3.0 M NaCl, pH 11.5 (see recipe) Methanol 0.05 M triethylammonium acetate (TEAA), pH 7.0 (see recipe) 20:80 (v/v) TEAA/MeCN (see recipe) 0.22-µm sterile membrane filters Sonicator Centrifuge Chromatography system and column(s): Amersham Biosciences FPLC system with UV detector, MonoQ HR 5/5 ion-exchange column, and PepRPC HR 5/5 reversed-phase column Gilson HPLC system with UV detector and 15 × 0.46–cm Progel-TSK Oligo-DNA-RP column (Supelco) Hewlett Packard series 1100 HPLC system with Lambda-Max model 481 UV/vis detector and MEGA 2 integrator (Carlo Erba) and 15 × 0.46–cm Progel-TSK Oligo-DNA-RP column (Supelco) For preparative IE-FPLC 1a. Dissolve 2 mg of the completely deprotected 15-mer in 1.5 mL start buffer (mobile phase A; 0.1 M NaCl, pH 11.5), filter through a sterile 0.22-µm membrane filter, and degas by sonication. 2a. Perform ion-exchange purification using a MonoQ HR 5/5 column on an Amersham Biosciences FPLC system with the following conditions:
Mobile phase A: 0.1 M NaCl, pH 11.5 Mobile phase B: 3.0 M NaCl, pH 11.5 Elution gradient: 20% B for 3 min (hold) 20% to 80% B in 22 min Flow rate: 1.0 mL/min Temperature: room temperature UV detector: 260 nm. 3a. Collect the material of the peak corresponding to the 15-mer and desalt the product by reversed-phase chromatography using a PepRPC HR 5/5 column on an Amersham Biosciences FPLC system with the following conditions:
Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
Mobile phase A: water Mobile phase B: methanol Elution gradient: 100% A for 15 min (hold) 0% to 100% B in 15 min Flow rate: 0.7 mL/min UV detector: 260 nm.
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For analytical IE-FPLC 1b. Dissolve 2 mg sample in 1.5 mL water, filter through a sterile 0.22-µm membrane filter, dilute the filtrate 10-fold with start buffer (mobile phase A; 0.1 M NaCl, pH 11.5), and degas the solution by sonication. 2b. Inject 100 µL of the degassed solution and perform analytical ion-exchange FPLC using a MonoQ HR 5/5 column on an Amersham Biosciences FPLC system using the following conditions: Mobile phase A: 0.1 M NaCl, pH 11.5 Mobile phase B: 3.0 M NaCl, pH 11.5 Elution gradient: 0% B for 5 min (hold) 0% to 100% B in 35 min Flow rate: 1.0 mL/min Temperature: room temperature UV detector: 260 nm.
For analytical RP-HPLC: Method 1 1c. Dissolve 2 mg sample in 1.5 mL water, filter through a sterile 0.22-µm membrane filter, dilute the filtrate 10-fold with start buffer (mobile phase A; 0.05 M TEAA, pH 7.0), and degas the solution by sonication. 2c. Inject 100 µL of the degassed solution and perform reversed-phase HPLC using a Progel-TSK Oligo-DNA-RP column (15 × 0.46 cm) on a Gilson HPLC system with the following conditions: Mobile phase A: 0.05 M TEAA, pH 7.0 Mobile phase B: 20:80 (v/v) TEAA/MeCN Elution gradient: 10% to 40% B in 40 min Flow rate: 1.0 mL/min Temperature: room temperature UV detector: 260 nm.
For analytical RP-HPLC: Method 2 1d. Prepare sample as in step 1c. 2d. Inject 100 µL of the degassed solution and perform reversed-phase HPLC using a Progel-TSK Oligo-DNA-RP column (15 × 0.46 cm) on a Hewlett Packard series 1100 system with the following conditions: Mobile phase A: 0.05 M TEAA, pH 7.0 Mobile phase B: 20:80 (v/v) TEAA/MeCN Elution gradient: 10% B for 2 min (hold) 10% to 40% B in 38 min Flow rate: 1.0 mL/min Temperature: room temperature UV detector: 260 nm.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
DDD in MeCN, 0.5 M Dissolve 1.21 g diethoxydithiocarbonate disulfide (DDD; Isis Pharmaceutical; store ◦ at –20◦ C) in 10 mL MeCN (dried using 4-A molecular sieve) in a dark bottle. Store unused solution at 4◦ C.
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NaCl, pH 11.5, 0.1 and 3.0 M 3.0 M solution: Dissolve 350.64 g (6 mol) NaCl in 2 L double-distilled water and adjust pH to 11.5 with 5 N NaOH. Filter through a Millipore GS 0.22-µm sterile filter and degas by sonication (30 min) prior to use. Store at 4◦ C. 0.1 M solution: Dilute 3 M NaCl with distilled water, readjust pH, and filter and degas as above. Store at 4◦ C.
Phosphoramidites in MeCN 0.1 M DMTr-protected nucleoside phosphoramidites: In a dark bottle sealed with◦ a rubber cap, dissolve 1 g phosphoramidite in 13.4 mL MeCN (dried using 4-A molecular sieves), injected with a syringe. Prepare immediately before use and keep under argon at –20◦ C. 0.2 M dimer-phosphoramidites: Prepare as above, but adjust volume of MeCN to give a final concentration of 0.2 M. As an example, 1.0 g of dT dimer is dissolved in 4.45 mL of dry MeCN.
TEAA, pH 7.0, 0.05 M Prepare a stock solution of 0.5 M triethylammonium acetate (TEAA) by adding 34.8 mL (0.25 mol) triethylamine (TEA) to 14.3 mL (0.25 mol) glacial acetic acid in 100 mL of double-distilled water. Dilute the mixture to 500 mL with water and adjust the pH to 7.0 using triethanolamine (TEA) or glacial acetic acid. Store at 4◦ C. Prepare 0.05 M working solution by diluting 100 mL of the 0.5 M stock solution to 1 L with double-distilled water, filtering through a Millipore GS 0.22-µm sterile membrane filter, and degassing by sonication for 30 min before use.
TEAA/MeCN, 20:80 (v/v) Mix 200 mL 0.05 M TEAA, pH 7.0 (see recipe), with 800 mL MeCN. Store at 4◦ C. Degas the solution by sonication for 30 min before use. 1H-Tetrazole in MeCN, 0.5 M In a bottle sealed with a rubber cap, dissolve 2 g of 1H-tetrazole in 57.1 mL dry MeCN (injected with a syringe). Store the solution under argon at room temperature. COMMENTARY Background Information
Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
The investigation of therapeutic properties of synthetic oligonucleotides and their chemically modified analogs has reached different stages of the clinical trial process, including the approval of the first antisense drug, Vitravene, a registered trademark of Isis Pharmaceuticals (Grillone and Lanz, 2001). The current studies, together with the prospect of successful commercialization, have triggered a demand for large-scale oligonucleotide synthesis. High coupling efficiency, low reagent consumption, and simple and cost-efficient purification are crucial factors
for successful production of oligonucleotidebased drugs (Seliger, 1993; Andrade et al., 1994). Solid-phase automated synthesis has been the method of choice due to ease of operation (Padmapriya et al., 1994; Ravikumar et al., 1995; Tsou et al., 1995). However, despite the development of new high-capacity synthesizers, the application of solid-phase automated synthesis for production of large quantities of oligonucleotides still remains problematic. The limitation is mainly due to the heterogeneity of the reaction, which leads to the use of quite a large excess of high-cost monomers in order to achieve high coupling
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efficiencies. The liquid-phase synthesis was proposed as an answer to the abovementioned limitation (Bayer and Mutter, 1972). In this technique, sequential synthesis is performed with the growing oligonucleotide chain attached to a soluble polymer support, and chain elongation is then carried out in homogeneous medium. The polymer-bound oligomer is usually recovered from the reaction mixture by precipitation, thus allowing rapid elimination of excess reagent and soluble byproducts. In a new liquid-phase method called highefficiency liquid-phase (HELP), polyethylene glycol (PEG; mol. wt. 5 to 20 kDa) is used as a soluble support polymer for oligonucleotide synthesis (Bonora, 1995). The oligonucleotide chain is joined through a succinate linkage. Once completed, the oligonucleotide is released from the support under standard conditions and purified by chromatographic methods. Oligomers of up to 20 nucleotides in length have been synthesized by following this approach. This method has been applied for synthesis of phosphorothioate analogs (Scremin and Bonora, 1993) in which a nonbridging oxygen is formally replaced by a sulfur atom to overcome the instability of natural DNA toward degradative enzymes (Cohen, 1993). Usually, sulfurization of the phosphite backbone is carried out as a repeating step during each elongation cycle. Thus, conversion of the phosphite moiety to phosphorothioate needs to be performed at the highest efficiency in order to achieve high purity of oligonucleotide-based drugs. As a consequence, the improvement of the thiolation reaction by the use of a proper sulfurizing reagent represents one of the main aspects of research in this field. Alternatively, the introduction of fully protected phosphorothioate dimers instead of monomers during the coupling step reduces the number of coupling and thiolation steps, and consequently reduces the number of unconverted phosphite linkages. This will certainly result in higher homogeneity of the final phosphorothioate backbone, as recently demonstrated (Krotz et al., 1997). The dimeric synthons are fully protected 3 →5 phosphorothiate dinucleotides, with a phosphoroamidite group at the 3 position as a reacting moiety for coupling with the 5 -hydroxyl of the growing oligonucleotide chain. Sulfurization of the phosphite triester moiety is carried out during each synthetic cycle with the recently developed diethoxydithiocarbonate disulfide (DDD), which seems to offer advantages over traditional sulfurizing
reagents (Eleuteri et al., 1999). A higher sulfurization yield is expected. The pharmacological properties of synthetic oligonucleotides can be modulated by their conjugation with proper molecules (Manoharan, 2001). The rationale of these modifications is dictated by the necessity for better cellular uptake, biostability, and pharmacokinetic properties. Among the molecules described in the literature, it is possible to distinguish between low- and high-molecularmass units. Within the first group, lipophilic conjugates such as cholesterol and other steroids, vitamins, and folic acid have been widely employed (MacKellar et al., 1992). Oligosaccharides and peptides have been considered for their ability to deliver oligonucleotides specifically to the targeted cells (Garcia de la Torre et al., 1999). Cleaving and cross-linking agents have also been proposed to improve the overall biological performances (Magda et al., 1997). Furthermore, large conjugating molecules such as proteins and antibodies have been linked, and great attention has been turned to polyamines as polycationic carriers (Markiewicz et al., 1998). Among the different biocompatible polymers, polyethylene glycol (PEG) was extensively investigated on the basis of previous success achieved with the PEGylation of protein (Kodera et al., 1998). In fact, this procedure is well on its way to becoming a standard component of the pharmaceutical tool box, since PEG possesses a unique set of properties, including analytical methods for conjugate characterization; absence of toxicity, immunogenicity and antigenicity; low massdependent elimination via the kidney; and high solubility in water and other organic media. At the oligonucleotide level, the presence of highmolecular-weight PEGs has showed a minimal effect on hybridization behavior, while a clear enhancement of in vivo stability and cellular permeation has been observed without any adverse toxic effect (Pang, 1993). Additionally, taking advantage of the recently developed procedure for producing pure, selectively and reversibly protected, bifunctional PEGs (Drioli et al., 2001), it is easy to imagine the production of PEG conjugates carrying both an oligonucleotide and an additional molecule such as a peptide, steroid, intercalator, or whatever can be devised to further improve their pharmacological properties. The coupling of PEG to a bioactive molecule is achieved by a polymeric derivative having an activated functional group at
Synthesis of Modified Oligonucleotides and Conjugates
4.27.23 Current Protocols in Nucleic Acid Chemistry
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one or both termini (Zalipsky, 1995), chosen on the basis of the reactive groups of the molecule to be PEG-conjugated. In the case of oligonucleotides, which offer only low nucleophilic functionalities such as primary hydroxyl groups, it is quite difficult to attain an extensive modification through a direct reaction with PEG. Moreover, the introduction of a large PEG chain by classical solid-phase procedures, commonly used for many postsynthetic oligonucleotide modifications, suffers from the phase heterogeneicity of the process, which implies poor reactivity and unpredictable kinetic effects. An acceptable yield has been reported in the literature only for PEGs of lower molecular weight following standard phosphoramidite conditions (Tarasow et al., 1997). However, the adverse effects are enhanced with increasing mass of the conjugating polymer (J¨aschke et al., 1994), with better biological performances being observed with sizes up to 20 to 40 kDa. An oligonucleotide conjugation using a larger PEG molecule in a classical solution reaction has been reported; a very large excess of the ester-activated form of polymer was employed, but the yield of condensation was not discussed (Wlotzka et al., 2002). Very recently, a conjugation procedure similar to the one described here has been reported (Jeong et al., 2003). For all these reasons, the postsynthetic solution-phase PEG conjugation of synthetic oligonucleotides can be properly optimized to achieve an optimum level of modification. To avoid the low reactivity of the terminal hydroxyl group, and to extend this procedure, possibly, to the peptide level, a terminal amino function was introduced on the oligonucleotides.
Critical Parameters and Troubleshooting
Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
The favorable aspects of the HELP technique are: 1. The synthesis may easily be scaled at the bench over a range between 0.2 and 1.0 mmol. The upper limit for the synthesis scale may be larger than 1.0 mmol, since all steps are carried out in a classical solution approach. 2. The functionalization of the supporting polymer can be efficiently achieved both with a monomer and a protected dimeric unit. 3. The deblocking process has been efficiently completed following standard oligonucleotide deprotection conditions.
4. The fully deblocked oligomers are efficiently separated using standard ion-exchange procedures. 5. The overall and average yields, as judged from the final DMTr assay, are satisfactory and comparable regardless of the synthetic scales investigated thus far. 6. The overall recovery after all the dissolution and precipitation steps is acceptable. A higher amount of product should be expected when similar processes are carried out on a larger scale. The unfavorable aspects are: 1. The intermediate purification steps demand a relatively large volume of precipitating solvent. The solvent could, however, be fully recovered by devising a proper recycling process. 2. A reduced efficiency of the coupling reaction is commonly observed with increasing chain length. As a consequence, the repetition of a coupling step is often required in the preparation of 8-mer or longer oligonucleotides. A third coupling step has even been necessary for the final condensation leading to a 15-mer. 3. The desalting procedure demands a standard molecular sieving method. Selective precipitation does not work efficiently, likely due to the reduced length of the oligomer and hence its unfavorable solubility properties. 4. The molecular weight of the PEG used in the post-conjugation process is strictly related to the final condensation yield. A molecular weight higher than 5 kDa reduces the efficiency of this process. 5. The final postsynthetic conjugation works quite efficiently if a proper excess of activated PEG is employed. Subsequent separation of the conjugate from unreacted PEG is obviously necessary and demands a careful chromatographic process. The following precautions must be considered: 1. The size of the supporting MPEG must be commensurate with the size of the desired oligomers. An MPEG that has a molecular weight 30% higher than that of the desired oligonucleotide is acceptable (for example, a 10-kDa MPEG is recommended for the preparation of a 15-mer oligonucleotide). 2. The synthetic process works efficiently if absolute care is taken in drying all the materials employed—reagents and support—to avoid any possible water contamination. 3. The overall process is fully effective only if careful DMTr analysis of all the
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Table 4.27.1 Expected Time for One Cycle of Thioate Oligonucleotide Synthesis via HELP Procedurea
Reaction step
Time (min)
Recovery step
Time (min)
Detritylation
15 + 15b
Precipitation
20 + 20c
Coupling
5 (15)d
Precipitation
20
Sulfurization
15
Precipitation
20 + 20c
Recrystallizatione
60
Capping
3
Precipitation
20
Total reaction time
53 (63)
Total recovery time
180
a Standard condensation cycle: detritylation-coupling-sulfurization-capping. b This process is usually repeated to ensure complete removal of the DMTr group. c Precipitations with ether are repeated to ensure complete removal of the reagents. d Because of solubility problems, coupling time can be extended up to 15 min when necessary. e Recrystallization from EtOH is suggested only for the complete removal of soluble intermediates.
intermediate coupling yields is done on strictly purified products to avoid any DMTr contribution from non-MPEG-bound contaminants that may still be present. 4. When the coupling yield is lower than near-quantitative, it is absolutely mandatory to repeat the coupling step before proceeding with the subsequent capping, oxidation/sulfurization, and detritylation steps. 5. A possible solution to the problem of low coupling yield could be the use of a higher excess of reagent. However, a more difficult intermediate purification can be expected in this case, owing to the need to remove a higher amount of non-MPEG-bound reagents. 6. A key step for a successful preparation of the desired oligonucleotide is the precipitation and filtration step; hence, great care is required for this operation. If conducted appropriately, <0.2% of product will be lost after each precipitation with ether. The recrystallization from ethanol has a higher expected loss (1% to 2%). For this reason, it must be executed only when essential for the overall purification.
Anticipated Results The efficiency of the reported HELP process is at least comparable to alternative standard procedures, if not higher. The possibility of increasing oligonucleotide synthesis to a scale that is unaffordable through solid-phase procedures can now be envisaged. A largerscale HELP-based production will be more efficiently evaluated in a pilot-scale study, since in that case, the simultaneous investigation of the recovery of support, unreacted reagents, and solvents can be better conceptualized. Future development could address the production of next-generation oligonucleotides of
different structures demanding new kinds of protected monomers. In that case, it is possible to foresee new, as yet unexpected, solubility and reactivity problems when these new reagents are applied to solid-phase procedures.
Time Considerations Preparation of the starting succinate usually takes 1 day, while the following derivatization of the supporting MPEG is achieved after an overnight reaction. The time demanded for a single cycle of the HELP synthesis of oligonucleotides is summarized in Table 4.27.1. The final conjugation procedure is quite time-consuming, since large PEG molecules are not very reactive. Consequently, to ensure the best condensation yield, a reaction time spanning a whole weekend is suggested. The deprotection and purification processes are the bottleneck of the overall procedure, since at least 1 week must be planned for the preparative purification.
Literature Cited Andrade, M., Scozzari, A.N., Cole, D.L., and Ravikumar, V.T. 1994. Efficient synthesis of antisense oligodeoxyribonucleotide phosphorothioates. Bioorg. Med. Chem. Lett. 4:2017-2022. Ballico, M., Cogoi, S., Drioli, S., and Bonora, G.M. 2003. Postsynthetic conjugation of biopolymers with high molecular mass poly(ethylene glycol): Optimization of a solution process tested on synthetic oligonucleotides. Bioconjugate Chem. 14:1038-1043. Bayer, E. and Mutter, M. 1972. Liquid-phase synthesis of peptides. Nature 237:513-514. Beaucage, S.L. and Caruthers, M.H. 1981. Deoxynucleoside phosphoramidites: A new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Lett. 22:1859-1862.
Synthesis of Modified Oligonucleotides and Conjugates
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Bonora, G.M. 1995. Polyethylene-glycol: A highefficiency liquid-phase (help) for the largescale synthesis of the oligonucleotides. Appl. Biochem. Biotechnol. 54:3-17.
Manoharan, M. 2001. Oligonucleotide conjugates in antisense technology. In Antisense Drug Technology (S. Crooke, ed.) pp. 391-469. Marcel Dekker, New York.
Bonora, G.M., Rossin, R., Zaramella, S., Cole, D.L., Eleuteri, A., and Ravikumar, V.T. 2000. A liquid-phase process suitable for large-scale synthesis of phosphorothioate oligonucleotides. Org. Proc. Res. Dev. 14:225-231.
Markiewicz, W.T., Godzina, P., Markievicz, M., and Astriab, A. 1998. Synthesis of a polyamino-oligodeoxyribonucleotide combinatorial library. Nucleosides Nucleotides Nucleic Acids 17:1871-1880.
Cohen, J.S. 1993. Phosphorothioate oligodeoxynucleotides. In Antisense Research and Applications (S.T. Crooke and B. Lebleu, eds.) pp. 205222. CRC Press, Boca Raton, Fla.
Padmapriya, A.A., Tang, J.-Y., and Agrawal, S. 1994. Large-scale synthesis, purification, and analysis of oligodeoxynucleotide phosphorothioates. Antisense Res. Dev. 4:185-199.
Drioli, S., Benedetti, F., and Bonora, G.M. 2001. Pure, homo-bifunctional poly(ethylene glycol) orthogonally protected: Synthesis and characterization. Reactive and Functional Polymers 48:119-128.
Pang, S.N.J. 1993. Final report on the safety assessment of polyethylene glycols. J. Am. Coll. Toxicol. 12:429-456.
Eleuteri, A., Cheruvallath, Z.S., Capaldi, D.C., Cole, D.L., and Ravikumar, V.T. 1999. Oligodeoxyribonucleotide phosphorothioates: Substantial reduction of (N-1)-mer content through the use of blockmer phosphoramidite synthons. Nucleosides Nucleotides Nucleic Acids 18:1803-1807. Garcia de la Torre, B., Albericio, F., SaisonBehmoaras, E., Bachi, A., and Eritja, R. 1999. Synthesis and binding properties of oligonucleotides carrying nuclear localization sequences. Bioconjugate Chem. 10:1005-1012. Grillone, L.R. and Lanz, R. 2001. Formivirsen. Drugs Today 37:245-255. J¨aschke, A., F¨urste, J.P., Nordhoff, E., Hillenkamp, F., Cech, D., and Erdmann, V.A. 1994. Synthesis and properties of oligodeoxyribonucleotidepolyethylene glycol conjugates. Nucl. Acids Res. 22:4810-4017. Jeong, J.H, Kim, S.W., and Park, T.G. 2003. A new antisense oligonucleotide delivery system based on self-assembled ODN-PEG hybrid conjugate micelles. J. Control. Release 93:183-191. Kodera, Y., Matsushima, A., Hiroto, M., Nishimura, H., Ishii, A., Ueno, T., and Inada, Y., 1998. PEGylation of proteins and bioactive substances for medical and technical applications. Prog. Polymer Sci. 23:1233-1271.
Conjugated Oligonucleoside Phosphorothioates Using the HELP Method
Ravikumar, V.T., Andrade, M., Wyrzykiewicz, T.K., Scozzari, A.N., and Cole, D.L. 1995. Large-scale synthesis of oligodeoxyribonucleotide phosphorothioate using controlled-pore glass as support. Nucleosides Nucleotides Nucleic Acids 14:1219-1226. Scremin, C.L. and Bonora, G.M. 1993. Liquidphase synthesis of phosphorothioate oligonucleotides on polyethylene-glycol support. Tetrahedron Lett. 34:4663-4666. Seliger, H. 1993. Protocols for Oligonucleotides and Analogs. In Methods in Molecular Biology, Vol. 20 (S. Agrawal ed.) pp. 391-435. Humana Press, Totowa, N.J. Tarasow, T.M., Tinnermeier, D., and Zyzniewski, C. 1997. Characterization of oligodeoxyribonucleotide-polyethylene glycol conjugates by electrospray mass spectrometry. Bioconjugate Chem. 8:89-93. Tsou, D., Hampel, A., Andrus, A., and Vinayak, P. 1995. Large-scale synthesize of oligoribonucleotides on high-loaded polystyrene (HLP) support. Nucleosides Nucleotides Nucleic Acids 14:1481-1492. Wlotzka, B., Leva, S., Eschgfaller, B., Burmeister, J., Kleinjung, F., Kaduk, C., Muhn, P., HessStumpp, H., and Klussmann, S. 2002. In vivo properties of an anti GnRG Spiegelmer: An example of an oligonucleotide-based therapeutic substance class. Proc. Natl. Acad. Sci. U.S.A. 99:8899-8902.
Krotz, A.H., Klopchin, P., Cole, D.L., and Ravikumar, V.T. 1997. Improved impurity profile of phosphorothioate oligonucleotides through the use of dimeric phosphoramidite synthons. Bioorg. Med. Chem. Lett. 7:73-78.
Zalipsky, S. 1995. Chemistry of polyethylene glycol conjugates with biologically active molecules. Adv. Drug Deliv. Rev. 16:157-182.
MacKellar, C., Grahham, D., Will, D.W., Burgess, S., and Brown, T. 1992. Synthesis and physical properties of anti-HIV antisense oligonucleotides bearing terminal lipophilic groups. Nucl. Acids Res. 20:3411-3417.
Contributed by Gian Maria Bonora University of Trieste Trieste, Italy
Magda, D., Wright, M., Crofts, S., Lin, A., and Sessler, J.L. 1997. Metal complex conjugate of antisense DNA which displays ribozyme-like activity. J. Am. Chem. Soc. 119:6947-6948.
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Disulfide Conjugation of Peptides to Oligonucleotides and Their Analogs
UNIT 4.28
This unit contains detailed procedures for the conjugation of peptides to oligonucleotides and their analogs via a disulfide linkage. The first section discusses the strategic planning for disulfide conjugation and provides protocols for the preparation of various types of peptide and oligonucleotide analog materials to be conjugated. Then protocols are provided for the conjugation of well-known cell-penetrating peptides to various oligonucleotide analogs that maintain negatively charged phosphate groups (e.g., 2 -Omethyloligoribonucleotides, locked nucleic acids). The final protocol focuses on peptide conjugation to peptide nucleic acids (PNA), which do not contain negatively charged phosphates. The differences between these conjugation protocols, as well as the reasons for the differences, are highlighted.
STRATEGIC PLANNING Peptide Synthesis Peptides used for conjugation can be synthesized by the user or obtained from a commercial supplier. A requirement for the peptide is that it should contain a thiol group, either free or activated (if conjugating to a free thiol on the oligonucleotide cargo). This is almost always in the form of a cysteine residue that is attached to one of the peptide termini. Any peptide synthesis machine can be used for the assembly steps, but the authors have used Fmoc/tert-butyl solid-phase peptide chemistry on either an Applied Biosystems Pioneer peptide synthesizer (100-µmol scale) or an Advanced ChemTech APEX 396 robotic synthesizer (5- to 10-µmol scale). C-terminal carboxylic acid peptides were synthesized with the C-terminal amino acid attached to the solid support. For peptides where the cysteine is on the N-terminus, Boc-Cys(Npys)-OH can be used instead of standard Fmoc-Cys(Tr)-OH so that the peptide is activated on-resin without the need for activation after solid-phase synthesis and purification. With peptides where the thiol is not on the N-terminus, thiol activation can only be carried out after complete synthesis of the peptide due to the instability of the activated thiol to piperidine and other strong nucleophiles. Oligonucleotide Synthesis Oligonucleotides suitable for disulfide conjugation can have the thiol linker modification at either the 5 - or the 3 -end. These modified oligonucleotides can often be obtained from a commercial supplier or otherwise synthesized by the user. For 3 modifications, a modified solid support is used. 5 -Thiol modification is achieved using an appropriate commercial thiol-linker phosphoramidite. The authors found difficulties in removing the protecting group of a trityl thiol modifier, due to the need to use silver nitrate. Yields of the thiol product were much lower than when a disulfide-protected thiol was liberated. Furthermore, there were reproducibility problems with the conjugation reactions, which were often low yielding, presumably due to the presence of residual silver salts. Complete reproducibility was observed with a thiol derived from a disulfide-protected linker. Thiol deprotection should not be carried out at the same time as cleavage of the oligonuclotide from the solid support and removal of other protecting groups, since this will lead to significant amounts of acrylonitrile addition to the thiol moiety, forming an unreactive thioether. Furthermore, such action unnecessarily complicates the purification and can result in disulfide-linked dimer formation during the ensuing slow desalting process.
Contributed by John J. Turner, Donna Williams, David Owen, and Michael J. Gait Current Protocols in Nucleic Acid Chemistry (2006) 4.28.1-4.28.21 C 2006 by John Wiley & Sons, Inc. Copyright
Synthesis of Modified Oligonucleotides and Conjugates
4.28.1 Supplement 24
Figure 4.28.1
Formation of disulfide-linked conjugates of peptides and antisense cargoes.
Disulfide Bond Formation Originally, asymmetric disulfide conjugation between two different biomolecules was achieved by the oxidation of alkyl thiols by atmospheric oxygen or by a mild oxidizing reagent (e.g., “diamide”). However, this method produces mixtures of unwanted homodimers as well as the desired asymmetric disulfide. By pre-activation of one of the thiols, the requisite asymmetric disulfide can be obtained as the sole product. Thiol activation is typically achieved using either the pyridylsulfenyl (pys) or 3-nitropyridylsulfenyl (Npys) group (Fig. 4.28.1). Although a stoichiometric amount of both reagents is theoretically sufficient to produce the desired disulfide product, usually the more expendable of the two reagents (often the peptide) is used in excess to drive the conjugation reaction to completion. Thiol activation of peptides is initially the most obvious choice, since these are synthesized on a larger scale and the loss of material due to the extra step is less important. However, when reacting several different peptides with one antisense cargo, it is more appropriate to activate the oligonucleotide. Another advantage of this strategy is that thiol-containing peptides that are also highly basic (e.g., contain many Arg or Lys residues) do not readily form homodimers, presumably due to electrostatic repulsion. By contrast, oligonucleotide cargoes are susceptible to homodimer formation when stored as the free thiol. Activation with the pys group can prevent such homodimerization. However, even this may not be sufficient in all cases to prevent homodimer formation, especially when storing solutions for extended periods of time (>3 months).
Disulfide Conjugation of Peptides to Oligonucleotides and Their Analogs
Purification of Conjugates Crucial to successful synthesis of peptide conjugates of oligonucleotides and their analogs is separation of the desired conjugate from the excess of components (usually the peptide). This is particularly difficult when a highly cationic peptide is conjugated to a negatively charged oligonucleotide because the excess peptide may bind to the conjugate, causing aggregation and precipitation from aqueous solution. A fluorophore on the oligonucleotide greatly assists in the monitoring of precipitation during the reaction. It is necessary to use a strong denaturing agent during conjugation and also during subsequent purification by anion-exchange chromatography. Furthermore, it is important to choose an ion-exchange column for purification that has a short path length and large pores, such
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as Resource Q (Amersham Biosciences). Because of precipitation, it is not advisable to purify oligonucleotides and conjugates from peptides having strong cationic charge using reversed-phase purification techniques. CAUTION: Wear gloves and protective glasses and carry out all operations involving organic solvents and reagents in a well-ventilated chemical fume hood.
CONJUGATION OF PEPTIDES WITH OLIGONUCLEOTIDE ANALOGS CONTAINING NEGATIVELY CHARGED PHOSPHATES
BASIC PROTOCOL 1
The synthesis of peptides conjugated by disulfide bond to oligonucleotides and their analogs has been known for some years (Eritja et al., 1991; Bongartz et al., 1994; Corey, 1995; Viv`es and Lebleu, 1997; Antopolsky et al., 1999; Astriab-Fisher et al., 2000). The protocols shown here are based on the authors’ own synthetic methods (Turner et al., 2005a) that involve improvements in conjugation and purification conditions as well as analysis. Each of the precursor biomolecules is prepared separately by solid-phase synthesis and purified prior to the conjugation steps. The synthetic conjugate product must be carefully purified from the conjugation reaction and characterized. Alternate steps are given for the peptide synthesis, using two different kinds of synthesizer, and depending on whether the activated Cys is at the N- or C-terminus. Oligonucleotide assembly can be carried out on any automated DNA/RNA synthesizer. The authors used an Applied Biosystems 394 or 3400 four-column synthesizer on a 1-µmol scale. The molecular mass of the oligonucleotides should be checked by MALDI-TOF-MS, for example on an Applied Biosystems Voyager DE-PRO workstation in positive ion mode. For evaporations of small volumes, use of a Speedvac concentrator is ideal.
Materials Fmoc-protected amino acid monomers (Novabiochem) including: Fmoc-Arg(Pbf)-OH Fmoc-Asn(Tr)-OH Fmoc-Cys(Tr)-OH Fmoc-Gln(Tr)-OH Fmoc-Glu(OtBu)-OH Fmoc-His(Tr)-OH Fmoc-Lys(Boc)-OH Fmoc-Trp(Boc)-OH N,N-Dimethylformamide (DMF, AnalaR-grade, BDH Chemicals), freshly distilled PyBop (Novabiochem) N,N-Diisopropylethylamine (DIPEA, 99+%, Applied Biosystems) Piperidine (>99.5%, Romil) NovaSyn TGR resin (for C-terminal amide synthesis, Novabiochem) PEG-PS resin (for C-terminal carboxylic acid synthesis, Applied Biosystems) Boc-Cys(Npys)-OH (for N-terminal cysteine; Bachem Bioscience) Isopropanol Trifluoroacetic acid (TFA, >99.9%, Romil) Triisopropylsilane (TIS, >99%, Aldrich) Diethyl ether, 4◦ C Acetonitrile (MeCN, HPLC-grade, Fisher Scientific) Millipore water or double-distilled deionized water 1,2-Ethanedithiol (EDT, >98%, Fluka) Water (HPLC-grade) 1.0 M NH4 HCO3 solution (aq.) 10 mg/mL 2-aldrithiol (Aldrich) in DMF
Synthesis of Modified Oligonucleotides and Conjugates
4.28.3 Current Protocols in Nucleic Acid Chemistry
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Nucleoside phosphoramidites (as needed): 2 -Deoxyribonucleoside phosphoramidites (Glen Research) 2 -O-Me-ribonucleoside phosphoramidites (Transgenomics) Locked nucleic acid (LNA) phosphoramidites (Link Technologies) Anhydrous acetonitrile 3 -(6-Fluorescein)-CPG (for 3 -fluorescent oligonucleotides, Glen Research) Thiol modifier C6-S-S (for 5 -thiol modification, Glen Research) 0.02 M iodine solution in 78:2:20 (v/v/v) THF/pyridine/water (Proligo) 5-Ethylthio-1H-tetrazole (a 0.25 M solution in MeCN, Link Technologies) 30% (v/v) aq. ammonia Sodium perchlorate (AnalaR, BDH Chemicals) 2.0 M Tris·Cl, pH 6.8 Formamide (p.a. ≥99.0%, Fluka) Sterilized water 2.0 M triethylammonium acetate, pH 7 (TEAA, Glen Research) 1.0 M aqueous D/L-dithiothreitol (DTT, ≥99%, Aldrich) Triethylamine (TEA, ≥99.5%, Fluka) Peptide synthesizer (e.g., APEX 396 or Pioneer peptide synthesizer) Desiccator attached to vacuum 15-mL polyethylene syringe (IST empty reservoir, Kinesis) 20-µm polyethylene frit (Kinesis) Speedvac concentrator Benchtop centrifuge 15- and 50-mL centrifuge tubes (Falcon) 0.22- (water) and 0.45-µm (water and organic) filters (Millipore) HPLC system, chemically inert, suitable for ion-exchange chromatography, with: PEEK tubing throughout Injector, sample loop, and syringe (manual loading) UV/Vis detector, variable wavelength 190 and 500 nm (preferable) or dual-wavelength detection Helium for degassing Jupiter reversed-phase HPLC column with guard (analytical and semi-prep, Phenomenex) DNAPac PA-100 (9 × 250–mm) Dionex column (semi-prep) with guard attached Resource Q column (1 mL/min analytical or 6 mL/min semi-prep, Amersham Biosciences) Lyophilizer Rotary evaporator and water pump 1.5-mL screw-cap tube (Sarstedt) or vial 55◦ C bath or temperature block (optional) Spin-X tube (Costar) Dialysis tubing, 3500 MWCO (Medicell International Ltd.) UV spectrometer Sephadex NAP-25 column (Amersham Biosciences) 0.5-mL microcentrifuge tube Slide-a-lyzer (0.5- to 3-mL capacity, 3500 MWCO, Pierce)
Disulfide Conjugation of Peptides to Oligonucleotides and Their Analogs
Additional reagents and equipment for peptide synthesis, MALDI-TOF-MS (see Support Protocol 1), Ellman’s test (see Support Protocol 2), oligonucleotide synthesis (APPENDIX 3C), and amino acid analysis (optional; see UNIT 4.22) NOTE: Amino acid protecting groups are tBu, tert-butyl; Boc, tert-butyloxycarbonyl; Fmoc, 9-fluorenylmethoxycarbonyl; Pbf, 2,2,4,6,7-pentamethyldihydrobenzofuran-5sulfonyl; pys, pyridylsulfenyl; Npys, 3-nitropyridylsulfenyl; and Tr, trityl.
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Table 4.28.1 Synthesis Cycle for Peptide Synthesis on the APEX 396 Synthesizer
Synthesis step
Reagents and volumes (µL)
Time (min)
Deprotection
20% piperidine solution (1000)
3
20% piperidine solution (1000)
12
Wash (five times)
DMF (1000)
1
Couple (two times)
Amino acid (300)
25
PyBop (300) DIPEA (300) Wash (five times)
DMF (1000)
1
Synthesize peptide with an activated cysteine For an APEX 396 peptide synthesizer (N-terminal cysteine) 1a. Dissolve the amino acid derivatives in DMF to give 0.5 M solutions (allow 600 µL per amino acid residue). 2a. Dissolve PyBop in DMF to give a 0.5 M solution (allow 600 µL per amino acid residue). 3a. Make a 1 M DIPEA solution in DMF (allow 600 µL per amino acid residue). 4a. Make a 20% piperidine solution in DMF (allow 2 mL per amino acid residue). 5a. Weigh out 15 µmol resin, put in a reactor well of the peptide synthesizer, and swell the resin two times with 1 mL DMF (5 min each). Drain off the DMF from the well. 6a. Program the synthesizer for the desired sequence. 7a. Start the synthesis beginning with Fmoc deprotection and first coupling and continue until the sequence is completed. See Table 4.28.1. Ensure that there is sufficient DMF and nitrogen gas present for the whole synthesis.
8a. Carry out the final peptide coupling with Boc-Cys(Npys)-OH. Do not carry out final Fmoc deprotection. 9a. Wash the resin with 3 mL DMF, then 3 mL isopropanol, and dry the resin under vacuum in a desiccator for 4 hr. 10a. To simultaneously cleave the peptide from the support and deprotect, treat with 1 mL of 95% TFA/2.5% water/2.5% TIS for 4 to 6 hr (using the longer time with high Arg content). 11a. Filter off the resin using a 15-mL polyethylene syringe fitted with a 20-µm frit and wash with an additional 1 mL TFA. Concentrate the filtrate to ∼10% of the original volume using a rotary evaporator, and precipitate the peptide with 25 mL cold (4◦ C) diethyl ether. 12a. Vortex the mixture and compact the precipitate by centrifuging 5 min at 1666 × g (2500 rpm), room temperature. Carefully decant the diethyl ether solution and wash the precipitate with 25 mL cold diethyl ether three additional times, centrifuging the residue and decanting the supernatant each time. CAUTION: It is necessary to use sealed centrifuge buckets when centrifuging solutions of flammable liquids such as ether.
Synthesis of Modified Oligonucleotides and Conjugates
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13a. Analyze the crude product and purify by HPLC on a Phenomenex Jupiter reversedphase column (analytical or semi-prep as appropriate) using the following conditions:
Buffer A: 0.1% TFA (aq.) Buffer B: 90:10 (v/v) acetonitrile/buffer A Gradient: 0% to 90% buffer B initially, optimizing according to peptide. All buffered solutions used for HPLC purifications should be filtered through a 0.45-µm disposable filter prior to use (see also UNIT 10.5).
14a. Collect appropriate fractions, lyophilize, redissolve in 1 mL water or buffer A, and analyze by HPLC and MALDI-TOF-MS (see Support Protocol 1). Continue with step 15.
For a Pioneer peptide synthesizer (N-terminal cysteine) 1b. Weigh 0.4 mmol of the appropiate amount of amino acid derivatives into separate tubes (4 equivalent excess per coupling). 2b. Dissolve PyBop in DMF to give a 0.395 M solution (calculated for 3.95 equivalents excess per coupling). 3b. Make a 0.79 M DIPEA solution in DMF (calculated for 7.9 equivalents excess per coupling). 4b. Make a 20% piperidine solution in DMF (calculated for 50 mL per amino acid residue). 5b. Weigh out 100 µmol resin (amino acid loading typically 0.2 mmol/g, thus 0.5 g resin). Place resin in the synthesis column and connect to the synthesizer. Swell the resin in 1 mL DMF for 2 min. 6b. Program synthesizer for the desired sequence. 7b. Start the synthesis beginning with Fmoc deprotection and the first coupling and continue, using manufacturer’s programs and specifications. Ensure that there is sufficient DMF and nitrogen gas present for the whole synthesis.
8b. Carry out steps 8a to 14a as above. Continue with step 15.
For a C-terminal cysteine 1c. Repeat steps 1a to 6a or 1b to 6b as above depending on the synthesizer type. 2c. Start the synthesis beginning with Fmoc deprotection and couple first with FmocCys(Tr)-OH in the same way as in 7a or 7b, depending on the synthesizer type. 3c. Do not add Boc-Cys(NPys)-OH as the final amino acid; do carry out the final deprotection of Fmoc. 4c. Wash the resin with 3 mL DMF, then 3 mL isopropanol, and dry the resin under vacuum in a desiccator. 5c. To simultaneously cleave the peptide from the support and deprotect, treat with 1 mL of 94% TFA/2.5% water/2.5% EDT/1% TIS for 4 to 6 hr (depending on Arg content). Disulfide Conjugation of Peptides to Oligonucleotides and Their Analogs
6c. Carry out steps 11a to 14a. 7c. Quantify the peptide using the Ellman’s test (see Support Protocol 2).
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8c. Activate the C-terminal cysteine-containing peptide as synthesized by dissolving in HPLC-grade water at a concentration between 1 and 5 mM. In the case of hydrophobic peptides, the solution should contain 10% to 30% acetonitrile to maintain solubility.
9c. Add 1 M aqueous NH4 HCO3 solution to make an overall concentration of 0.1 M buffer. 10c. Add 2-aldrithiol solution (10 equivalents), mix thoroughly by vortexing the solution, centrifuge briefly (5 sec) at 7412 × g (6000 rpm), and allow to stand for 2 to 3 hr at room temperature. 11c. Dilute with an equal volume of 0.1% aq. TFA (mobile phase A). 12c. Vortex thoroughly and centrifuge 3 min at 16,060 × g (13,000 rpm), room temperature. 13c. Purify the supernatant by RP-HPLC. Collect the product and lyophilize. Analyze by MALDI-TOF-MS (see Support Protocol 1) and by HPLC. 14c. Quantify either by weighing the product or, if the amount is too small, by using amino acid analysis (see general procedure for amino acid analysis in UNIT 4.22). Continue with step 15.
Synthesize oligonucleotide containing a thiol linker 15. Dissolve the appropriate phosphoramidites (standard 2 -deoxyribonucleoside, 2 -O-Me-ribonucleoside, LNA, or others) in anhydrous MeCN as 100 mM solutions. 16. Start the automated solid-phase oligonucleotide synthesis from an appropriate solid support-filled column, e.g., on a 1-µmol scale. See APPENDIX 3C for details on automated oligonucleotide synthesis. For 3 -fluorescent oligonucleotides, use 3 -(6-fluorescein)-CPG (Glen Research) for the support.
17. Elongate the oligonucleotide chain in DMTr-OFF mode following the standard operating procedures for the machine as directed by the manufacturer. Use 0.25 M 5-ethylthio-1H-tetrazole as the coupling agent. For LNA monomers, allow quadruple the time for iodine oxidation (60 sec cf. 15 sec for other monomers) and 15 min per coupling. Use the thiol modifier in the final coupling step. 18. After completion of the assembly, remove the column, wash with MeCN (two times with 10 mL for 1-µmol synthesis) and quickly dry on a water pump. 19. Transfer the support into a 1.5-mL screw-cap tube or vial and add 1 mL of 30% aq. ammonia. On some DNA synthesizers, such as the ABI 394 or 3400 machines, steps 18 and 19 are carried out automatically on the synthesizer and the product oligonucleotide is collected in a vial.
20. Vortex the mixture and incubate for at least 8 hr at 55◦ C or for 24 hr at ambient temperature. 21. Cool the solution in a −20◦ C freezer, and evaporate off the ammonia and dry in vacuo using a Speedvac concentrator. 22. Add 0.5 mL of deionized water, transfer the solution to a Spin-X tube, and centrifuge 5 min at 16,060 × g (13,000 rpm), room temperature. Wash the filter with deionized water (two times with 0.25 mL) and concentrate in vacuo. Current Protocols in Nucleic Acid Chemistry
Synthesis of Modified Oligonucleotides and Conjugates
4.28.7 Supplement 24
23. Purify DMTr-OFF oligonucleotides using an ion-exchange (IE) HPLC with the following conditions:
Column: 9 × 250–mm DNAPac PA-100 Buffer A: 1 mM sodium perchlorate, 20 mM Tris·Cl pH, 6.8, and 25% formamide Buffer B: 400 mM sodium perchlorate, 20 mM Tris·Cl, pH 6.8, and 25% formamide Gradient: 15% to 55% buffer B in 20 min Flow rate: 3 mL/min Detection at 260 nm wavelength. For oligonucleotides with fluorescein as the fluorophore, measurement at 480 nm is also possible. All buffered solutions used for HPLC purifications should be filtered through a 0.45-µm disposable filter prior to use (see also UNIT 10.5).
24. Pool the appropriate HPLC fractions. Desalt by dialysis against 2500 mL water at 4◦ C for 24 hr using a 3500 MWCO membrane, and changing the water three times. Lyophilize the dialysate. 25. Dissolve the oligonucleotide in deionized water and quantify by measuring the UV absorbance at 260 nm. Store the solution frozen at −20◦ C (stable at least 6 months). 26. Check the molecular mass of the oligonucleotide by MALDI-TOF-MS (see Support Protocol 1).
Liberate thiol on the oligonucleotide 27. Dissolve 300 nmol purified oligonucleotide in 292.5 µL sterilized water. 28. Add 7.5 µL of 2.0 M TEAA buffer, followed by 3 µL of 1 M DTT and 3 µL triethylamine. 29. Mix thoroughly by vortexing and allow to stand for 1.5 hr at room temperature. 30. Monitor thiol formation by HPLC (see step 23) and MALDI-TOF-MS (see Support Protocol 1). 31. Equilibriate a NAP-25 column with sterilized water. Load the sample, collect the desalted product, and lyophilize. 32. Quantify the absorbance value at 260 nm and carry out an Ellman’s test (see Support Protocol 2) to determine if there is any DTT impurity present.
Conjugate oligonucleotide thiol to activated peptide 33. Place 20 µL of a 1 mM aqueous solution of thiol oligonucleotide in a 0.5-mL microcentrifuge tube. 34. Add 70 µL formamide and 5 µL of 2 M TEAA buffer. Vortex and then microcentrifuge briefly. 35. Add the activated peptide (80 nmol in 8 µL) to the oligonucleotide, vortex, and allow to stand for 30 min.
Disulfide Conjugation of Peptides to Oligonucleotides and Their Analogs
For conjugations to highly basic peptides, precipitation can occur. The supernatant should be removed and the residue redissolved in a mixture of formamide and 2 M TEAA buffer before being combined with the supernatant and purified.
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36. Purify the conjugate using an IE-HPLC in one injection with the following conditions:
Column: Resource Q 1 mL/min Buffer A: 20 mM Tris·Cl, pH 6.8, and 50% formamide Buffer B: 400 mM sodium perchlorate, 20 mM Tris·Cl, pH 6.8, 50% formamide Gradient: 0% to 100% buffer B in 20 min Flow rate: 1 mL/min Detection: 280 nm wavelength. For oligonucleotides with fluorescein as fluorophore, simultaneous measurement at 480 nm can be done. Buffered solutions used for these HPLC purifications should be filtered through a 0.22µm disposable filter prior to use (see also UNIT 10.5). A fine filter is required because a guard column is not used.
37. Collect purified product, load in Slide-a-lyzer as directed, and dialyze for 24 hr against water at 4◦ C. 38. Recover sample and lyophilize. 39. Redissolve the conjugate in water or an appropriate buffer to analyze by HPLC and MALDI-TOF-MS (see Support Protocol 1). 40. Determine the yield by absorbance calculation at 260 nm.
CONJUGATION OF C-TERMINAL CYS-CONTAINING PEPTIDES TO OLIGONUCLEOTIDES VIA ACTIVATION OF THE OLIGONUCLEOTIDE
ALTERNATE PROTOCOL
In this protocol, the oligonucleotide is the activated species. Use the peptide thiol synthesized in Basic Protocol 1. A peptide with a C-terminal thiol is obtained from Basic Protocol 1, steps 1c to 7c. For peptides with an N-terminal thiol, again follow Basic Protocol 1, steps 1c to 7c, but in step 2c add the Fmoc-Cys(Tr)-OH in the last coupling step instead of the first.
Additional Materials (also see Basic Protocol 1) Sephadex NAP-10 column (Amersham Biosciences) Activate oligonucleotide 1. Dissolve 200 nmol oligonucleotide obtained from Basic Protocol 1, step 25, in 195 µL sterilized water. 2. Add 5 µL of 2 M TEAA buffer and 44 µL aldrithiol solution (2 µmol, 10 eq.). 3. Vortex and allow to stand 1 hr at room temperature. 4. Desalt using a Sephadex NAP-10 column. 5. Analyze and quantify (see Basic Protocol 1, steps 25 and 26).
Conjugate activated oligonucleotide to peptide thiol 6. Place 20 µL of a 1 mM aqueous solution of activated oligonucleotide in a 0.5-mL microcentrifuge tube. 7. Add 70 µL formamide and 5 µL of 2 M TEAA buffer. Vortex and then briefly centrifuge. 8. Add 50 nmol peptide thiol (see Basic Protocol 1, step 7c) in 5 µL to the oligonucleotide, vortex, and allow to stand for 30 min at room temperature.
Synthesis of Modified Oligonucleotides and Conjugates
4.28.9 Current Protocols in Nucleic Acid Chemistry
Supplement 24
For conjugations to highly basic peptides, precipitation can occur. The supernatant should be removed and the residue redissolved in a mixture of formamide and 2 M TEAA buffer before being combined with the supernatant and purified.
9. Purify and quantify as in Basic Protocol 1, steps 36 to 40. BASIC PROTOCOL 2
CONJUGATION OF PEPTIDES WITH PEPTIDE NUCLEIC ACIDS The conjugation of PNA oligomers to peptides using the disulfide route is more straightforward than with the oligonucleotide counterpart, since the likelihood of precipitation due to the addition of highly basic peptides is minimal. There have been a number of reports previously describing disulfide conjugation of PNA to peptides (Pooga et al., 1998; Braun et al., 2002; Kaushik et al., 2002; Koppelhus et al., 2002; Kilk et al., 2004), but this protocol is based on the authors’ own procedures (Turner et al., 2005b) that involve improvements in purification conditions and analysis. Synthesis of the activated PNA oligomer is achieved using the APEX 396 Robotic peptide synthesizer described in Basic Protocol 1. Oligomers synthesized should be analyzed and purified using a reversed-phase HPLC column heated to 45◦ C. Molecular mass can be verified by MALDI-TOF-MS using the CHCA matrix (see Support Protocol 1). For further information on the synthesis of PNA oligomers, see UNIT 4.11. It is well documented that a PNA oligomer can be sparingly soluble in aqueous solution, and that its conjugation to a sparingly soluble peptide (e.g., Transportan) may exacerbate this problem. Therefore, such behavior needs to be taken into consideration in PNA synthesis design. In order to assist in the solubility of the PNA oligomer, lysine residues are often added (typically three or four) to the C- and/or N-terminus (see UNIT 4.11).
Materials
Disulfide Conjugation of Peptides to Oligonucleotides and Their Analogs
Fmoc (Bhoc) PNA monomers (Applied Biosystems) N-Methylpyrrolidinone (NMP, ≥99.5%, Fluka) PyBop (Novabiochem) N,N-Dimethylformamide (DMF, AnalaR-grade, BDH Chemicals), freshly distilled N,N-Diisopropylethylamine (DIPEA, 99+%, Applied Biosystems) 2,6-Lutidine (≥99%, Aldrich) Piperidine (>99.5%, Romil) Fmoc-PAL-PEG-PS amide resin (Applied Biosystems) Isopropanol Trifluoroacetic acid (TFA, >99.9%, Romil) Triisopropylsilane (TIS, >99%, Aldrich) Phenol Diethyl ether, 4◦ C 5% acetic anhydride/6% 2,6-lutidine solution in DMF (PNA capping solution, Applied Biosystems) 1.0 M aq. NH4 OAc (AnalaR-grade, BDH Chemicals) APEX 396 Robotic peptide synthesizer 1-mL polyethylene syringe (IST empty reservoir, Kinesis) 10-µm polyethylene frit (Kinesis) Plastic tap Filtration unit 15-mL centrifuge tube (Falcon) Heating jacket for HPLC column UV spectrometer Lyophilizer Additional reagents and equipment for reversed-phase HPLC (see Basic Protocol 1)
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Table 4.28.2 Synthesis Cycle for PNA Synthesis on the APEX 396 Synthesizer
Synthesis step
Reagents and volumes (µL)
Time (min)
Deprotection
20% piperidine solution (800)
1
20% piperidine solution (800)
4
Wash (five times)
DMF (1000)
1
Couple (two times)
PNA or amino acid (100) PyBop (100) DIPEA/lutidine (100) DMF (100)
30
Wash (five times)
DMF (1000)
1
Cap (two times)
PNA capping solution (1000)
5
Wash (five times)
DMF (1000)
1
Synthesize PNA oligomer containing an activated (Npys) cysteine thiol 1. Dissolve PNA monomers in NMP to give 0.2 M solutions (allow 200 µL per PNA or amino acid residue). 2. Dissolve PyBop in DMF to give a 0.2 M solution (allow 200 µL per PNA or amino acid residue). 3. Make a 0.2 M DIPEA/0.2 M 2,6-lutidine solution in DMF (allow 200 µL per PNA or amino acid residue). 4. Make a 20% piperidine solution in DMF (allow 1.6 mL per PNA or amino acid residue). 5. Weigh out 5 µmol resin, place in a reactor well, and swell the resin two times with 1 mL DMF (5 min each). 6. Program the synthesizer for the desired sequence. Typically three lysine residues are added to the C-terminus and one is added to the N-terminus followed by the activated cysteine for the final coupling.
7. Start the synthesis beginning with Fmoc deprotection and the first coupling, and continue until the sequence is completed. See Table 4.28.2. Ensure that there is sufficient DMF and nitrogen gas present for the whole synthesis.
8. Carry out the final coupling with Boc-Cys(Npys)-OH. Do not carry out final Fmoc deprotection. 9. Wash the resin with 3 mL DMF, then 3 mL isopropanol, and dry the resin under vacuum in a desiccator for 4 hr. 10. Place resin in a 1-mL polyethylene syringe fitted with 10-µm frit and plastic tap. 11. To simultaneously cleave the PNA oligomer from the support and deprotect it, add 1 mL of a 95% TFA/2.5% water/2.5% triisopropylsilane solution containing 10% phenol for 4 hr. 12. Filter into a 15-mL centrifuge tube, washing the resin with an additional 0.5 mL TFA. Concentrate to ∼200 µL using a rotary evaporator and water pump, and precipitate the PNA oligomer with 10 mL cold (4◦ C) diethyl ether. 13. Vortex the mixture and compact the precipitate by centrifuging 5 min at 1666 × g (2500 rpm), room temperature. Carefully decant the diethyl ether solution and wash the precipitate with 10 mL diethyl ether three additional times, centrifuging the residue and carefully decanting the supernatant each time.
Synthesis of Modified Oligonucleotides and Conjugates
4.28.11 Current Protocols in Nucleic Acid Chemistry
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CAUTION: It is necessary to use sealed centrifuge buckets when centrifuging solutions of flammable liquids such as ether.
14. Analyze the crude product and purify by use of a Phenomenex Jupiter reversedphase HPLC column (analytical or semi-prep as appropriate) heated to 45◦ C. Monitor by UV at 260 nm. Use the same buffers as in Basic Protocol 1, step 13a. A typical gradient for a 16- to 18-mer with four lysines is 5% to 30% buffer B in 25 min.
15. Collect appropriate fractions, lyophilize, redissolve in water/acetonitrile as required, and analyze by HPLC and MALDI-TOF-MS (see Support Protocol 1). 16. Quantify at 260 nm. The extinction coefficient for individual nucleobases is different than for the corresponding oligonucleotides (see UNIT 4.11).
Conjugate PNA oligomer to thiol peptide 17. Place 20 µL of a 1 mM aqueous solution of oligonucleotide in a microcentrifuge tube. 18. Add 10 µL of 1.0 M NH4 OAc and then the thiol peptide to be conjugated (40 nmol, 4 µL of a 10 mM stock solution; see Basic Protocol 1 for synthesis of the thiol peptide). In the case of lipophilic peptides that are sparingly soluble in water at neutral pH (e.g., Transportan), add 50 µL of acetonitrile prior to peptide addition.
19. Mix the solution by vortexing, centrifuge briefly (5 sec at 7412 × g), and allow to stand for 30 min at room temperature. 20. Purify the resulting conjugate in one injection by reversed-phase HPLC using a Phenomenex Jupiter column as described in step 14, using a flow rate of 1.5 mL/min and a gradient of 5% to 30% buffer B in 25 min when conjugating to highly basic peptides, or a gradient of 5% to 50% buffer B when conjugating to more lipophilic peptides. Buffers are the same as for peptides (see Basic Protocol 1, step 13a).
21. Collect the product and lyophilize. 22. Redissolve in sterile water, analyze by HPLC and MALDI-TOF-MS (see Support Protocol 1), and quantify by measuring absorbance at 260 nm (see UNIT 4.11). SUPPORT PROTOCOL 1
DETERMINATION OF MOLECULAR MASS BY MALDI-TOF MASS SPECTROMETRY Although mass determination through MALDI-TOF analysis is dealt with in several existing units (UNIT 4.11 for PNA, UNIT 4.22 for peptides and oligonucleotides, and UNIT 10.1 for general protocols for machine use), a number of variations in the matrices have been found to work well for the various constructs described here. General operational procedures can be found in the manufacturer’s manual and in UNIT 10.1.
Materials Disulfide Conjugation of Peptides to Oligonucleotides and Their Analogs
α-Cyano-4-hydroxycinnamic acid (CHCA, ≥99.0%, Aldrich) Acetonitrile 3% (v/v) aq. trifluoroacetic acid (TFA, >99.9%, Romil) 2,6-Dihydroxyacetophenone (DHAP, ≥99.0%, Fluka) Methanol Diammonium hydrogen citrate (≥99.0%, Fluka)
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Millipore water or double-distilled deionized water 2,4,6-Trihydroxyacetophenone (THAP, ≥99%, Fluka) 1.5-mL solvent-resistant microcentrifuge tubes MALDI-TOF mass spectrometer Prepare matrix For peptides, PNA oligomers, and peptide-PNA conjugates 1a. Dissolve 10 mg CHCA in 0.5 mL acetonitrile in a 1.5-mL solvent-resistant microcentrifuge tube. 2a. Add 0.5 mL of 3% aq. TFA solution. 3a. Mix thoroughly. Continue with step 4.
For oligonucleotides and peptide-oligonucleotide conjugates (low salt content in sample) 1b. Dissolve 20 mg DHAP in 0.5 mL methanol in a 1.5-mL solvent-resistant mirocentrifuge tube. 2b. Dissolve 40 mg diammonium hydrogen citrate in 0.5 mL water in a separate tube. 3b. Mix the two solutions together thoroughly. Continue with step 4.
For oligonucleotides and peptide-oligonucleotide conjugates (high salt content in sample; also used to check for contamination by peptides) 1c. Add 0.5 mL acetonitrile to 60 mg THAP in a 1.5-mL solvent-resistant microcentrifuge tube and vortex for 1 min. 2c. Dissolve 5.7 mg diammonium hydrogen citrate in 0.5 mL water and add to THAP/acetonitrile mixture. 3c. Vortex for 30 sec and centrifuge 3 min at 16,060 × g (13,000 rpm), room temperature. Use the supernatant and continue with step 4. Excess THAP will be in pellet form at the bottom of the tube.
Prepare sample 4. Spot 1 µL of the matrix onto the plate. 5. Immediately (within 5 sec) add 0.1 µL of the sample and mix by pipetting the solution up and down five times with a pipet. The concentration of the sample is critical for good mass acquisition (see UNIT 10.1). Sample concentrations exceeding 1 mM can give inferior spectra. For THAP matrix (which is saturated), immediate crystallization can occur upon dispensing the sample. In this case, do not mix. In general, cease mixing the sample/matrix mixture if crystallization commences.
6. Allow to crystallize and dry completely. 7. Acquire spectra in positive ion mode (see UNIT 10.1).
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SUPPORT PROTOCOL 2
DETERMINATION OF THIOL CONTENT BY THE ELLMAN’S TEST The Ellman’s test is a widely used and established technique for the quantitation of thiol content in a given solution.
Materials Ellman’s reagent: 2 mM dithio-bis-2-nitrobenzoic acid (DTNB) in 50 mM sodium acetate (NaOAc) 2.0 M Tris·Cl, pH 8.0 UV spectrometer with 1-mL, 1-cm path length quartz glass cuvette (Suprasil, Hellma) 1. Add 50 µL Ellman’s reagent, 100 µL of 2.0 M Tris·Cl, and water (850 µL minus the sample volume) to a quartz cuvette. 2. Mix thoroughly by carefully pipetting 200 µL of solution up and down ten times with a pipet. 3. Zero the UV spectrometer at 412 nm. 4. Repeat step 2 and check that the UV spectrometer still reads zero. If not, repeat steps 2 and 3 until it does. 5. Add the sample to the cuvette, mix carefully ten times, and take the reading. 6. Calculate absorbance as follows:
A(sample) = [total volume (µL)/sample volume (µL)] × A412 7. Repeat steps 1 through 6 two times, varying the amount of sample used (e.g., 10, 15, 20 µL), and take the average. 8. Calculate the thiol content as follows:
thiol (M) = average A(sample)/13,600 where 13,600 M−1 cm−1 is the extinction coefficient of the reagent. Due to the sensitivity of the test, when fluorophores are present (e.g., fluorescein, 492 nm maximum) ensure that a background check at pH 8.0 (i.e., no Ellman’s reagent present) at 412 nm is carried out so that the absorbance value reading can be adjusted if necessary.
COMMENTARY Background Information
Disulfide Conjugation of Peptides to Oligonucleotides and Their Analogs
Antisense oligonucleotides and siRNA The antisense oligonucleotide field emerged some 25 years ago with the pioneering work of Zamecnik (Zamecnik and Stephenson, 1978). However, a severe limitation to biological activity of oligonucleotides and their analogs has been the poor cellular uptake and delivery into the right cell compartment. Most oligonucleotides and analogs are not taken up into cells in culture without use of an additional carrier, of which cationic lipids (such as Lipofectamine 2000) are the most popular (Bennett et al., 1992). The same problems of poor cell uptake extend
to siRNA and PNA. For therapeutic use, cationic lipids have disadvantages because of the need for careful formulation and difficulties in maintaining stability. Major efforts have been made to find alternative methods of oligonucleotide and siRNA drug delivery that do not require formulation, but merely involve chemical modification of the oligonucleotide itself. Peptide conjugates of oligonucleotides to enhance cell delivery One idea that has been pursued for several years is the covalent conjugation of cellpenetrating peptides (CPP), also known as protein transduction domains (PTD) (Lindgren
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et al., 2000; Lindsay, 2002; Wadia and Dowdy, 2002; Lochmann et al., 2004; Zorko and Langel, 2005). Such peptides have been shown to translocate into mammalian cells in culture, but it has been controversial as to what extent such peptides are taken up by endocytotic pathways or by non-energy-dependent processes. It appears that some cationic peptides, such as Tat, are predominantly taken up by endocytosis (Richard et al., 2003), but even now the uptake mechanism is disputed (Potocky et al., 2003; Kaplan et al., 2005; Ziegler et al., 2005). It is also less clear for more hydrophobic peptides such as Transportan (Zorko and Langel, 2005). Methods of synthesis of peptide conjugates of oligonucleotides and analogs have been reviewed (Zubin et al., 2002; Zatsepin et al., 2005) and their applications for cell delivery covered extensively (Stetsenko et al., 2000; Gait, 2003; Thierry et al., 2003; Shi and Hoekstra, 2004; Juliano, 2005). Very recently there have been a couple of papers suggesting that peptide conjugates of siRNA are also useful for carrier-free delivery (Chiu et al., 2004; Muratovska and Eccles, 2004). Such peptide carrier–oligonucleotide or siRNA cargo conjugates can be synthesized either with a stable linkage (such as amide, thioether, oxime, or hydrazine) or with an unstable linkage (such as disulfide). Disulfide-linked peptide-oligonucleotide and peptide-PNA conjugates Very little information had been available as to whether unstable disulfide linkages may have any advantage over more stably linked conjugates. Recent evidence from the PNA-peptide field suggests, at least for this class of conjugates, that disulfide-linked conjugates may have higher activity (Koppelhus et al., 2002; Turner et al., 2005b). However, this has not been proven in the case of peptide conjugates of negatively charged oligonucleotides, where sufficient biological activity has been hard to achieve (Antopolsky et al., 1999; Astriab-Fisher et al., 2000; Turner et al., 2005a). Potentially, the disufide bond may be cleaved by reduction within a mammalian cell to allow cargo release linkage, but the rates of such intracellular cleavage are unknown and may vary depending on the cargo as well as the peptide type. Disulfide conjugates have become popular with laboratories studying the uptake and activity of peptideantisense cargo conjugates because the chemistry is simple and reasonably specific, and the thiol functional groups needed on each component are readily introduced. However,
with regards to the synthesis of highly cationic peptide-oligonucleotide conjugates, reliable protocols have been missing, and there have even been reports suggesting that such conjugations are impossible to achieve (Prater and Miller, 2004). The protocols provided here for synthesis, purification, and analysis of peptide-oligonucleotide (Turner et al., 2005a) and peptide-PNA (Turner et al., 2005b) conjugates should fill this gap and encourage others to explore a wider variety of peptides and oligonucleotides for conjugation, toward the goal of improved cell delivery. The authors’ results in this area have shown that conjugation by several types of CPP does indeed enhance cellular uptake into HeLa cells and fibroblasts in cell culture, but that biological activity levels in a model system involving steric block of HIV-1 Tat-dependent transactivation in the nucleus are limited in most cases by slow release from endosomal compartments (Turner et al., 2005a,b). However, recent exciting results suggest that disulfidelinked PNA-peptides targeting the HIV-1 TAR region have great potential as antiviral and virucidal agents (Tripathi et al., 2005) and also in the targeting of neuropeptide receptor mRNAs (Kilk et al., 2004). Disulfidelinked peptide-oligonucleotide conjugates are also useful for delivery into living cells for the detection of mRNA by a molecular beacons approach (Nitin et al., 2004).
Critical Parameters and Troubleshooting For the synthesis of peptides using the APEX or Pioneer machines, problematic sequences (often indicated by repeating motifs or large sections of hydrophobicity) should be identified and the use of double couplings or double-double couplings employed. Although the APEX machine can in theory synthesize 96 peptides in parallel, it is best to operate with smaller numbers due to the variation in exposure time to reagents in the different steps (e.g., piperidine). The authors have found that groups of four to six peptides at a time are manageable. Although the machine is by definition automated, it should be monitored as much as possible to catch any errors that may occur. Manual intervention at this stage can rescue the synthesis and prevent loss of reagents and time. The programming of the machine should also be double-checked before commencing the synthesis. The Pioneer peptide synthesizer, which is a flow-through instrument where one peptide is synthesized at a time, is less
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prone to error, but is no longer commercially available. Reagent solutions should not be used if older than 3 days. This is especially true for PyBop. Any solutions exhibiting discoloration should be discarded. Masses of +170 and +50 are occasionally observed as a result of degraded resin. Replacing the resin solves this. For PNA synthesis, much of what has been described for the synthesis of peptides on the APEX machine also holds true (i.e., programming and monitoring the machine during synthesis). Several additional factors are also critical to the synthesis of the PNA oligomers. (1) A short piperidine treatment should be used for cleavage of the Fmoc protecting group. The Bhoc nucleobase protecting group is susceptible to cleavage when longer (standard peptide) reaction times are used. (2) The solubility of the PNA monomers and, in general, the growing chain can be maintained by the addition of N-methylpyrrolidone (NMP) to the reaction mixture, if necessary. (3) The quality of the coupling reagent is of greater importance in PNA synthesis than in peptide synthesis. Although an amide bond is formed in both cases, peptide synthesis and solutions are more tolerant in general to the presence of small amounts of water than PNA synthesis. Poor yields can in some cases be rectified by adding molecular sieves to the solutions for PNA synthe-
Disulfide Conjugation of Peptides to Oligonucleotides and Their Analogs
sis. Stock solutions of thiol-containing peptides that are not highly cationic and of PNA oligomers should be stored in 0.1% aqueous TFA to inhibit dimerization. For both, acidic media may be required for solvation. Oligonucleotide synthesis is generally straightforward. With LNA, longer coupling times and a stronger activator (e.g., 5-ethylthiotetrazole) are used. For conjugations, it is important that the quality of both starting materials be high. Pure oligonucleotide or PNA and peptide suitable for conjugation will simplify the purification of the conjugate. It has been observed occasionally that the desalting procedure after the liberation of the oligonucleotide thiol with DTT can be incomplete. Traces of DTT in the final solution can diminish the coupling efficiency with activated peptides. Desalting again will resolve this problem. Alternatively, begin with a bigger Sephadex NAP column. It is not advisable to dialyze the thiol oligonucleotide, as dimer formation will occur. Maintaining solubility is vital for the smooth formation and purification of conjugates. The addition of a highly cationic peptide to a solution of oligonucleotide without formamide present will result in immediate aggregation. This is best visualized with fluorescently labeled materials. A high percentage of the solution should be formamide to ensure solubility. This percentage can rise to >90%
Figure 4.28.2 HPLC chromatogram of conjugate formation from peptide RQIKIWFQNRRMKWKKGGC with (pys)S-(CH2 )6 -5 -2 -O-Me/LNA[CUC CCA GGC UCA]-3 -fluorescein; peaks (i) salts and formamide, (ii) excess peptide, (iii) conjugate product, (iv) unconjugated oligo(pys). The solid trace is at 280 nm and the dashed trace is at 480 nm, which identifies the fluorescein label on the oligonucleotide.
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Figure 4.28.3 MALDI-TOF mass spectra of GRKKKRRQRRRPC(S-)-S-(CH2 )6 -5 -2 -O-Me/LNA[CUC CCA GGC UCA]-3 -fluorescein conjugate: (A) purified and (B) pure conjugate with 0.5 eq. peptide added.
for reactions involving highly cationic peptides. It is best to keep highly concentrated stocks of peptides and oligonucleotides, so that the amount of formamide needed does not prevent single injection purification by HPLC. A high concentration is advisable to ensure quick reaction times. When purifying the conjugate, care must be taken when formamide and 2 M TEAA are used to resolubilize aggregated material. It has been observed that HPLC runthrough can occur when large amounts have been used. This material can be collected and re-injected in the following run.
Care must be taken when loading Slidea-Lyzer cassettes so as not to puncture the dialysis membrane. In addition, for highly basic peptide-oligonucleotide conjugates, there is a risk of self-aggregation onto the membrane during dialysis, resulting in a reduction in yield. The Slide-a Lyzer cassette can be washed with appropriate buffer (0.1 to 2 M TEAA) to solubilize the conjugate. This can then be lyophilized/concentrated or transferred into the appropriate buffer with a NAP column. Alternatively, dialysis can be carried out using an appropriate buffer instead
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Figure 4.28.4 RP-HPLC of (A) the synthesis of RRRRRRRQIKIWFQNRRMKWKKGGCCK[CTCCCAGGCTCAGATC]PNA KKK and (B) analysis of the purified product.
Disulfide Conjugation of Peptides to Oligonucleotides and Their Analogs
of water. Lyophilized conjugates can require a modest amount of salt to aid their solution. Often, adding a small amount of 2 M TEAA is sufficient—typically, enough to make a 10 mM solution, but in some cases up to 1 M TEAA is needed. For these latter, difficult cases, it is necessary to add concentrated
salt solution first and then dilute the solution with water. MALDI-TOF mass spectral analysis of conjugates dissolved in high salt may be problematic due to poor crystallization. Use THAP matrix (see Support Protocol 1), diluting a small amount of the sample, if necessary.
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Figure 4.28.5
MALDI-TOF mass spectrum of the conjugate described in Figure 4.28.4.
The conjugation reaction and purification of PNA-peptide conjugates is less problematic, since aggregation with cationic peptides does not occur. However, the overall solubility of conjugates should be considered (e.g., the conjugation of hydrophobic peptides to PNA). Self-aggregation of PNA oligomers can occur at neutral pH. This is circumvented by the addition of formamide. A heated HPLC column is advisable to obtain good resolution with all conjugates synthesized. Provided that the oligonucleotide (or analog) and peptide remain soluble at the concentrations described, disulfide formation should be complete within 30 min. Reaction times significantly greater than this can cause scrambling of the disulfide bond and hence a reduction in yield. Due to the short reaction times, it is not necessary to remove oxygen from the aqueous solution.
purity ≥90%. Oligonucleotide-peptide conjugations proceed >90% according to HPLC (see Fig. 4.28.2 for an example). Due to the self-aggregation properties of some conjugates (discussed above), yields are often lower. Yields may vary between 12% for selfaggregating conjugates and 75% for minimally self-aggregating conjugates. The presence of excess peptide can be verified by MALDI-TOF mass spectral analysis using the THAP matrix (see Fig. 4.28.3). Note that mass M/2 is often observed with THAP matrix. Conversion of a PNA oligomer into a conjugate is typically 80% to 90% (see Fig. 4.28.4). Yields obtained are typically 45% to 70%. Purity of all conjugates is >90%. MALDITOF mass spectra of PNA conjugates typically show molecular mass of the conjugate, molecular mass of conjugate/2, PNA thiol, and peptide thiol (see Fig. 4.28.5).
Time Considerations Anticipated Results Good yields and good purity can be expected for both peptides and oligonucleotides. Activation of peptide or oligonucleotide thiol groups proceeds near quantitatively (as judged by HPLC analysis). However, after isolation of the desired activated material, a typical yield is ≥85%. Liberation of the thiol from oligonucleotides typically proceeds >90%, sometimes quantitatively. For PNA synthesis, yields of ∼20% are expected after purification, depending on the sequence and length, with
The assembly of peptides and PNA can be completed within hours depending on the length of the oligomer. Allow 1 to 3 hr to program and set up the machine as well as prepare reagents, and then ∼1 hr per cycle. Oligonucleotide synthesis is more rapid, with cycles of 15 to 30 min. Conjugations are typically finished within 30 min. HPLC purification for oligo-peptide conjugates is 30 min per run. Dialysis is usually achieved with one change of water allowing at least 6 hr between (resulting in
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≥1 × 106 dilution of salt content). Purification of PNA-peptide conjuates takes 45 min. Starting with synthesis of the peptide and oligonucleotide (or analog), allow 2 to 3 weeks for the synthesis of conjugates altogether. Allow longer if experience is short in these areas and/or more than one peptide or oligonucleotide is being synthesized. If the oligonucleotide and peptide are obtained commercially, allow at least 1 week for the synthesis and analysis of conjugates.
Literature Cited Antopolsky, M., Azhayeva, E., Tengvall, U., Auriola, S., J¨aa¨ skel¨ainen, I., R¨onkk¨o, S., Honkakoski, P., Urtti, A., L¨onnberg, H., and Azhayev, A. 1999. Peptide-oligonucleotide phosphorothioate conjugates with membrane translocation and nuclear localization properties. Bioconjug. Chem. 10:598-606. Astriab-Fisher, A., Sergueev, D.S., Fisher, M., Shaw, B.R., and Juliano, R.L. 2000. Antisense inhibition of P-glycoprotein expression using peptide-oligonucleotide conjugates. Biochem. Pharmacol. 60:83-90. Bennett, C.F., Chiang, M.-Y., Chan, H., Shoemaker, J.E.E., and Mirabelli, C.K. 1992. Cationic lipids enhance cellular uptake and activity of phosphorothioate antisense oligonucleotides. Mol. Pharmacol. 41:1023-1033. Bongartz, J.P., Aubertin, A.M., Milhaud, P.G., and Lebleu, B. 1994. Improved biological activity of antisense oligonucleotides conjugated to a fusogenic peptide. Nucl. Acids Res. 22:4681-4688. Braun, K., Peschke, P., Pipkorn, R., Lampel, S., Wachsmuth, M., Waldeck, W., Friedrich, E., and Debus, J. 2002. A biological transporter for the delivery of peptide nucleic acids (PNAs) to the nuclear compartment of living cells. J. Mol. Biol. 318:237-243. Chiu, Y.-L., Ali, A., Chu, C., Cao, H., and Rana, T.M. 2004. Visualizing a correlation between siRNA, localization, cellular uptake and RNAi in living cells. Chem. Biol. 11:1165-1175. Corey, C.R. 1995. 48000-fold acceleration of hybridisation by chemically modified oligonucleotides. J. Amer. Chem. Soc. 117:93739374. Eritja, R., Pons, A., Escarceller, M., Giralt, E., and Albericio, F. 1991. Synthesis of defined peptide-oligonucleotide hybrids containing a nuclear transport signal sequence. Tetrahedron 47:4113-4120.
Disulfide Conjugation of Peptides to Oligonucleotides and Their Analogs
Gait, M.J. 2003. Peptide-mediated cellular delivery of antisense oligonucleotides and their analogues. Cell. Mol. Life Sci. 60:1-10.
enters cells by macropinocytosis. J. Control Release 102:247-253. Kaushik, N., Basu, A., Palumbo, P., Nyers, R.L., and Pandey, V.N. 2002. Anti-TAR polyamide nucleotide analog conjugated with a membranepermeating peptide inhibits Human Immunodeficiency Virus Type I production. J. Virol. 76:3881-3891. Kilk, K., Elmquist, A., Saar, K., Pooga, M., Land, T., Bartfai, T., Soomets, U., and Langel, U. 2004. Targeting of antisense PNA oligomers to human galanin receptor type 1 mRNA. Neuropeptides 38:316-324. Koppelhus, U., Awasthi, S.K., Zachar, V., Holst, H.U., Ebbeson, P., and Nielsen, P.E. 2002. Celldependent differential cellular uptake of PNA, peptides and PNA-peptide conjugates. Antisense & Nucleic Acid Drug Dev. 12:51-63. Lindgren, M., H¨allbrink, M., Prochiantz, A., and Langel, U. 2000. Cell-penetrating peptides. Trends Pharmacol. Sci. 21:99-103. Lindsay, M.A. 2002. Peptide-mediated cell delivery: Application in protein target validation. Curr Opin Pharmacol 2:587-594. Lochmann, D., Jauk, E., and Zimmer, A. 2004. Drug delivery of oligonucleotides by peptides. Eur. J Pharm. Biopharm. 58:237-251. Muratovska, A. and Eccles, M.R. 2004. Conjugate for efficient delivery of short interfering RNA (siRNA) into mamalian cells. FEBS Lett. 558:63-68. Nitin, N., Santangelo, P.J., Kim, G., Nie, S., and Bao, G. 2004. Peptide-linked molecular beacons for efficient delivery and rapid mRNA detection in living cells. Nucl. Acids Res. 32:e58. Pooga, M., Soomets, U., H¨allbrink, M., Valkna, A., Saar, K., Rezaei, K., Kahl, U., Hao, J.-X., Xu, X.-J., Wiesenfeld-Hallin, Z., H¨okfelt, T., Bart¨ 1998. Cell penetrating fai, T., and Langel, U. PNA constructs regulate galanin receptor levels and modify pain transmission in vivo. Nat. Biotechnol. 16:857-861. Potocky, T.B., Menon, A.K., and Gellman, S.H. 2003. Cytoplasmic and nuclear delivery of a TAT-derived peptide and a β-peptide after endocytic uptake into HeLa cells. J. Biol. Chem. 278:50188-50194. Prater, C.E. and Miller, P. 2004. 3 Methylphosphonate-modified oligo-2 -Omethytlribonucleotides and their Tat peptide conjugates: Uptake and stability in mouse fibroblasts in culture. Bioconjug. Chem. 15:498-507. Richard, J.-P., Melikov, K., Viv`es, E., Ramos, C., Verbeure, B., Gait, M.J., Chernomordik, L.V., and Lebleu, B. 2003. Cell-penetrating peptides. A re-evaluation of the mechanism of cellular uptake. J. Biol. Chem. 278:585-590.
Juliano, R.L. 2005. Peptide-oligonucleotide conjugates for the delivery of antisense and siRNA. Curr. Opin. Mol. Ther. 7:132-138.
Shi, F. and Hoekstra, D. 2004. Effective intracellular delivery of oligonucleotides in order to make sense of antisense. J. Control Release 97:189-209.
Kaplan, I.M., Wadia, J.S., and Dowdy, S.F. 2005. Cationic TAT peptide transduction domain
Stetsenko, D.A., Arzumanov, A.A., Korshun, V.A., and Gait, M.J. 2000. Peptide conjugates of
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oligonucleotides as enhanced antisense agents: A review. Mol. Biol. (Russ.) 34:852-859. Thierry, A.R., Viv`es, E., Richard, J.-P., Prevot, P., Martinand-Mari, C., Robbins, I., and Lebleu, B. 2003. Cellular uptake and intracellular fate of antisense oligonucleotides. Curr. Opin. Mol. Ther. 5:133-138. Tripathi, S., Chaubey, B., Ganguly, S., Harris, D., Casale, R.A., and Pandey, P.K. 2005. Anti-HIV1 activity of anti-TAR polyamide nucleic acid conjugated with various membrane transducing peptides. Nucl. Acids Res. 33:4345-4356. Turner, J.J., Arzumanov, A.A., and Gait, M.J. 2005a. Synthesis, cellular uptake and HIV1 Tat-dependent trans-activation inhibition activity of oligonucleotide analogues disulphide-conjugated to cell-penetrating peptides. Nucl. Acids Res. 33:27-42. Turner, J.J., Ivanova, G.D., Verbeure, B., Williams, D., Arzumanov, A., Abes, S., Lebleu, B., and Gait, M.J. 2005b. Cell-penetrating peptide conjugates of peptide nucleic acids (PNA) as inhibitors of HIV-1 Tat-dependent trans-activation in cells. Nucl. Acids Res. 33:6837-6849. Viv`es, E. and Lebleu, B. 1997. Selective coupling of a highly basic peptide to an oligonucleotide. Tetrahedron Lett. 38:1183-1186. Wadia, J.S. and Dowdy, S.F. 2002. Protein transduction technology. Curr. Opin. Biotechnol. 13:52-56.
Zamecnik, P.C. and Stephenson, M.L. 1978. Inhibition of Rous sarcoma virus replication and cell transformation by a specific oligodeoxynucleotide. Proc. Natl. Acad. Sci. U.S.A. 75:280-284. Zatsepin, T.S., Turner, J.J., Oretskaya, T.S., and Gait, M.J. 2005. Conjugates of oligonucleotides and analogues with cell penetrating peptides as gene silencing agents. Curr. Pharm. Des. 11:3639-3654. Ziegler, A., Nervi, P., D¨urrenberger, M., and Seelig, J. 2005. The cationic cell-penetrating peptide CPPTat derived from the HIV-1 protein TAT is rapidly transported into living fibroblasts: Optical, biphysical, and metabolic evidence. Biochemistry 44:138-148. Zorko, M. and Langel, U. 2005. Cell-penetrating peptides: Mechanism and kinetics of cargo delivery. Adv. Drug Deliv. Rev. 57:529-545. Zubin, E.M., Romanova, E.A., and Oretskaya, T.S. 2002. Modern methods for the synthesis of peptide-oligonucleotide conjugates. Russ. Chem. Rev. 71:239-264.
Contributed by John J. Turner, Donna Williams, David Owen, and Michael J. Gait Medical Research Council, Laboratory of Molecular Biology Cambridge, United Kingdom
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Methoxyoxalamido Chemistry in the Synthesis of Tethered Phosphoramidites and Functionalized Oligonucleotides
UNIT 4.29
This unit elaborates on the synthesis of specialty phosphoramidites tethered with single or multiple linkers through the use of methoxyoxalamido (MOX) precursors. The strategy is based on the reaction of dimethyl oxalate with a proper aliphatic amine (phosphoramidite base) to form a MOX precursor that, in turn, is reacted with a selected primary aliphatic amine (linker) to form a simple tethered synthon for phosphoramidite preparation (Fig. 4.29.1). The strategy is quite general since a vast number of commercially available primary aliphatic amines can be employed. Basic Protocol 1 describes syntheses of exemplary phosphoramidites tethered with single linkers. Additional flexibility of the MOX strategy comes from the possibility of reiterating the dimethyl oxalate/primary diamine treatments leading to synthons with multiple linkers (Fig. 4.29.2). Synthesis of selected phosphoramidites with tethers is outlined in the Alternate Protocol. Finally, Basic Protocol 2 describes the use of tethered phosphoramidites for 5 -derivatization of oligonucleotides.
Figure 4.29.1 Synthetic pathway for phosphoramidites tethered with a single linker. MOX, methoxyoxalamido; DMTr, dimethoxytrityl; MMTr, monomethoxytrityl.
Synthesis of Modified Oligonucleotides and Conjugates
Contributed by Alan M. Morocho and Nikolai N. Polushin
4.29.1
Current Protocols in Nucleic Acid Chemistry (2006) 4.29.1-4.29.19 C 2006 by John Wiley & Sons, Inc. Copyright
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Figure 4.29.2 Synthetic pathway to phosphoramidites tethered with multiple linkers. MOX, methoxyoxalamido; DMTr, dimethoxytrityl; MMTr, monomethoxytrityl; n = number of cycles.
BASIC PROTOCOL 1
PREPARATION OF PHOSPHORAMIDITES TETHERED WITH SINGLE LINKERS Synthetic procedures for preparation of specialty phosphoramidites using MOX chemistry are all straightforward and robust. All necessary chemicals are commercially available and generally inexpensive. One of the starting reagents, 5 -amino-5 deoxythymidine, is somewhat expensive but can be prepared from inexpensive thymidine as described by Bannwarth (1988). There are four major steps in the synthesis of a phosphoramidite tethered with a single linker. First, the MOX derivative is prepared from a proper amino alcohol (phosphoramidite base). Second, the MOX precursor is treated with a primary aliphatic diamine or a primary amino alcohol to form an aminated or hydroxylated tether synthon. Next, the primary amino or hydroxyl group is protected with a monomethoxytrityl (MMTr) or dimethoxytrityl (DMTr) group, respectively. Lastly, the secondary hydroxyl of the MMTr/DMTr-protected intermediate is phosphinylated to give the final phosphoramidite.
Methoxyoxalamido Chemistry for Tethered Phosphoramidites
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Figure 4.29.3 depicts structures of the initial MOX precursors S.1 and S.2. Figure 4.29.4 shows phosphoramidites tethered with single aminated linkers (S.3d-S.6d) and their intermediates (S.3a,c-S.6a,c). Figure 4.29.5 shows phosphoramidites tethered with single hydroxylated linkers (S.10d and S.11d) and their precursors (S.10a,c and S.11a,c). Note that the b intermediate for S.5, S.6, and S.8 is used only for the formation of the tethered compounds described in the Alternate Protocol. The following considerations are important in strategic planning for the syntheses described in this unit: Choice of starting material (phosphoramidite base). The choice of starting materials, trans-4-aminocyclohexanol hydrochloride and 5 -amino-5 -deoxythymidine, is based on their availability and the presence of secondary hydroxyl groups, which imparts stability to the corresponding phosphoramidites. Formation of MOX precursors. At least a 2-fold excess of dimethyl oxalate over the amino alcohol should be used to suppress formation of a dimer byproduct. Addition of triethylamine, although not necessary in the case of free amines, does accelerate the reaction. In the case of trans-4-aminocyclohexanol hydrochloride, at least one equivalent of triethylamine must be added to free the protonated amino group. Reaction of MOX precursor with an aliphatic primary diamine. To suppress dimer formation, at least a 4-fold molar excess of diamine over the MOX precursor (S.1 or S.2) should be used. Typically, simple precipitation of the product (S.3a-S.6a) in ether gives purity acceptable to perform the next step. If the diamine used is of a substantial value, it can be recovered from the supernatant by distillation or crystallization. Reaction of MOX precursor with an aliphatic primary amino alcohol. Since only one amino group is available to react with a MOX precursor, only one equivalent of an amino alcohol is needed. Again, the product (S.10a, S.11a) is purified simply by precipitation in ether. Protection of primary amino/hydroxyl group. Due to poor solubility of amino alcohols S.3a-S.6a in pyridine, reaction with MMTr-Cl is carried out in 1:1 (v/v) dimethylformamide (DMF)/pyridine. Even in this solvent composition, the starting materials are not fully soluble, and the reaction mixture is stirred for 2 to 3 days to ensure complete transformation. Diols S.10a and S.11a are soluble in 1:1 DMF/pyridine, and the reaction with DMTr-Cl is generally complete within 12 hr. After extraction and concentration, the product is precipitated into 3:2 (v/v) ether/pentane and is normally used in the next step without further purification. However, if the purity of the MMTr/DMTr-protected intermediate is <85% (based on NMR and TLC analyses), column purification is recommended.
Figure 4.29.3
Initial MOX precursors.
Synthesis of Modified Oligonucleotides and Conjugates
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Figure 4.29.4
Phosphoramidites and precursors with simple or multiple aminated linkers. MMTr, monomethoxytrityl.
Phosphinylation. MMTr-protected amino alcohols S.3c-S.6c and DMTr-protected diols S.10c-S.11c are phosphinylated with 2-cyanoethyl-N,N,N ,N -tetraisopropylphosphorodiamidite in dry CH2 Cl2 in the presence of tetrazole. The corresponding phosphoramidites (S.3d-S.6d and S.10d-S.11d) are then purified by column chromatography on silica gel. Compound characterization 1 H and 31 P NMR spectra were recorded on a Varian XL-300 spectrometer at 300 and 121 MHz, respectively, in DMSO-d6 unless otherwise noted. Chemical shifts (δ) are reported in ppm downfield from tetramethylsilate (TMS) for 1 H NMR and from H3 PO4 for 31 P NMR. Electrospray ionization mass spectra (ESI-MS) were recorded on a Mariner spectrometry system (PerSeptive Biosystems) with samples dissolved in 1:1 (v/v) methanol/water.
Materials
Methoxyoxalamido Chemistry for Tethered Phosphoramidites
trans-4-Amino-1-cyclohexanol hydrochloride (97%; Aldrich) Triethylamine (Et3 N), ≥99% Methanol (MeOH), HPLC grade Dimethyl oxalate, 99% (Aldrich) Diethyl ether, anhydrous Chloroform (CHCl3 ), HPLC grade
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◦
60 A silica gel, 200 to 400 mesh (EM Science) 5 -Amino-5 -deoxythymidine (Berry & Associates, http://www.berryassoc.com or Fidelity Systems, http://www.fidelitysystems.com) Aliphatic primary diamines Ethylenediamine (EDA), ≥99.5% (Aldrich) for S.3a 2,4,8,10-Tetraoxaspiro[5.5]undecane-3,9-dipropanamine (TUDA), 97% (Aldrich) for S.4a 4,7,10-Trioxa-1,13-tridecanediamine (TTDD), ≥98% (Aldrich) for S.5a and S.6a Aliphatic primary amino alcohols 6-Amino-1-hexanol (AH), 97% (Aldrich) for S.11a 2-(2-Aminoethoxy)ethanol (AEE), 98% (Aldrich) for S.10a Dichloromethane (CH2 Cl2 ), HPLC grade N,N-Dimethylformamide (DMF), anhydrous Pyridine, anhydrous, 99.8% (Aldrich) 4-Monomethoxytrityl chloride (MMTr-Cl; ChemGenes) Saline solution: 25% to 30% (w/v) aqueous NaCl Na2 SO4 , reagent grade, anhydrous Pentane, HPLC grade 4,4 -Dimethoxytrityl chloride (DMTr-Cl; ChemGenes) Tetrazole, dried Argon gas 2-Cyanoethyl-N,N,N ,N -tetraisopropylphosphorodiamidite (ChemGenes) 10% (w/v) aqueous sodium hydrogen carbonate (NaHCO3 ) Dichloromethane (CH2 Cl2 ), anhydrous Ethyl acetate (EtOAc), HPLC grade Toluene, anhydrous Phosphorus pentoxide (P2 O5 ) Rotary evaporator equipped with a vacuum pump or water aspirator (5 to 10 Torr) Buchner funnel, 140-mL capacity with glass frit (porosity 4 to 8 µm) Paper filters (coarse porosity) 4 × 40–cm sintered glass columns Vacuum oil pump (0.05 to 0.5 Torr) Separatory funnel Thin-layer chromatography (TLC) Kieselgel 60 F254 plates (EM Science) I2 -silica chamber 254-nm UV lamp Heat gun Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare MOX precursors For compound S.1 from trans-4-aminocyclohexanol 1a. Prepare a solution of 3.03 g (20 mmol) trans-4-amino-1-cyclohexanol and 5.6 mL (40 mmol) Et3 N in 40 mL MeOH. 2a. In a separate flask dissolve 4.72 g (40 mmol) dimethyl oxalate in 20 mL MeOH. 3a. Add the solution of trans-4-amino-1-cyclohexanol and Et3 N dropwise over 2 hr to the stirring solution of dimethyl oxalate. 4a. Concentrate the reaction mixture on a rotary evaporator to ∼20 mL. Add 200 mL diethyl ether while stirring.
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Figure 4.29.5
Phosphoramidites and precursors with simple or multiple hydroxylated linkers. DMTr, dimethoxytrityl.
5a. Filter the precipitate under vacuum using a Buchner funnel, and wash twice with 30 mL diethyl ether. Dissolve the precipitate in 60 mL CHCl3 . ◦
6a. Pack a 4 × 40–cm sintered glass column with 100 g of 60 A silica gel in CHCl3 . Load the sample and run a gradient of 0% to 10% MeOH in CHCl3 . Collect and pool fractions containing the desired compound. Identify the correct fractions by TLC using UV or I2 staining for visualization. Evaporate solvent on a rotary evaporator. The large sample volume does not require concentration before loading onto the column because the loading solvent is nonpolar and will not push the compound through the column.
7a. Dry the resulting white crystals overnight in vacuo (vacuum pump, 0.05 to 0.5 Torr) to yield 3.27 g (81%) of S.1 as a white solid. 8a. Characterize the product by TLC, 1 H NMR, and ESI-MS.
Methoxyoxalamido Chemistry for Tethered Phosphoramidites
S.1: TLC (9:1 CHCl3 /MeOH): Rf = 0.3. 1 H NMR (DMSO-d6 ): δ 8.67-8.82 (d, 1H, NH), 4.55-4.62 (d, 1H, OH), 3.73-3.80 (s, 3H, OCH3 ), 3.44-3.66 (m, 1H, H4), 3.25-3.44 (m, 1H, H1), 1.62-1.91 (m, 4H, 2CH2 ), 1.08-1.50 (m, 4H, 2CH2 ). ESI-MS: m/z 201.8 (M + H+ ), 223.7 (M + Na+ ), 239.8 (M + K+ ); calcd. for C5 H9 NO4 : 201.2.
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For compound S.2 from 5 -amino-5 -deoxythymidine 1b. Prepare a solution of 4.82 g (20 mmol) 5 -amino-5 -deoxythymidine and 2.8 mL (20 mmol) Et3 N in 40 mL MeOH. 2b. In a separate flask dissolve 4.72 g (40 mmol) dimethyl oxalate in 20 mL MeOH. 3b. Add the solution of 5 -amino-5 -deoxythymidine and Et3 N dropwise over 2 hr to the stirring solution of dimethyl oxalate. 4b. Concentrate the reaction mixture on a rotary evaporator to ∼20 mL. Add 200 mL diethyl ether while stirring. 5b. Filter the precipitate under vacuum using a Buchner funnel, and wash twice with 30 mL diethyl ether. Dissolve the precipitate in 80 mL CHCl3 . ◦
6b. Pack a 4 × 40–cm sintered glass column with 100 g of 60 A silica gel in CHCl3 . Load the sample and run a gradient of 0% to 10% MeOH in CHCl3 . Collect and pool fractions containing the desired compound. Identify the correct fractions by TLC using UV or I2 staining for visualization. Evaporate solvent on a rotary evaporator. 7b. Dry the resulting white crystals overnight in vacuo (vacuum pump, 0.05 to 0.5 Torr) to yield 6.15 g (94%) of S.2 as a white solid. 8b. Characterize the product by TLC, 1 H NMR, and ESI-MS. S.2: TLC (8:2 CHCl3 /MeOH): Rf = 0.32. 1 H NMR (DMSO-d6 ): δ 11.27-11.37 (s, 1H, HN-3), 9.03-9.19 (t, 1H, COCONH), 7.49-7.54 (s, 1H, H6), 6.09-6.21 (t, 1H, H1 ), 5.305.38 (d, 1H, OH-3 ), 4.12-4.26 (m, 1H, H3 ), 3.74-3.93 (m, 4H, H4 +OCH3 ), 3.28-3.50 (m, 2H, CH2 -5 ), 1.95-2.23 (m, 2H, H2 ,2 ), 1.73-1.90 (s, 3H, CH3 -5). ESI-MS: m/z 349.9 (M + Na+ ); calcd. for C13 H17 N3 O7 : 327.29.
Derivatize MOX precursor 9. Dissolve an aliphatic primary diamine (40 mmol) or an aliphatic primary amino alcohol (12 mmol) in 50 mL MeOH and stir vigorously. 10. Dissolve 10 mmol of MOX precursor (S.1 or S.2) in 25 mL of 1:1 (v/v) MeOH/CH2 Cl2 (HPLC grade) and add to the diamine or amino alcohol solution in portions over 30 min. 11. Stir the reaction mixture for 2 hr. During this time a partial precipitation of product is typically observed.
12. Add 300 mL diethyl ether. Cool the suspension to 4◦ C for 2 hr to complete the precipitation. 13. Filter the precipitate under vacuum using a Buchner funnel. Wash the solid twice with 30 mL diethyl ether and dry overnight in vacuo (vacuum pump, 0.05 to 0.5 Torr). 14. Characterize the compound by TLC, 1 H NMR, and ESI-MS. From aliphatic primary diamine: S.3a from S.1 and EDA (95%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0. 1 H NMR (DMSOd6 ): δ 8.56-8.67 (t, 1H, CH2 -NH-COCO), 8.37-8.48 (d, 1H, COCO-NH-ACH), 4.48-4.58 (d, 1H, OH), 3.43-3.58 (m, 1H, H4 (ACH)), 3.25-3.40 (m, 1H, H1 (ACH)), 3.04-3.13 (q, 2H, CH 2 -NH-COCO), 2.55-2.63 (t, 2H, NH2 -CH 2 ), 1.72-1.85 (br d, 2H, 2 × CH (ACH)), 1.58-1.70 (br d, 2H, 2 × CH (ACH)), 1.29-1.48 (m, 2H, 2 × CH (ACH)), 1.081.26 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 230.4 (M + H+ ); calcd. for C10 H19 N3 O3 : 229.1.
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S.4a from S.1 and TUDA (77%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0. 1 H NMR (DMSO-d6 ): δ 8.69-8.79 (t, 1H, CH2 -NH-COCO), 8.38-8.49 (d, 1H, COCO-NH-ACH), 4.40-4.53 (m, 2H, 2 × CH2 -CH-(O-)2 (spiro ring)), 4.19-4.31 (br d, 2H, 2 × CH (spiro ring)), 3.44-3.62 (m, 5H, 4 × CH (spiro ring) + H4 (ACH)), 3.27-3.40 (m, 3H, 2 × CH (spiro ring) + H1 (ACH)), 3.03-3.18 (q, 2H, CH 2 -NH-COCO), 2.46-2.52 (m, H2 NCH 2 ), 1.74-1.88 (m, 2H, 2 × CH (ACH)), 1.60-1.73 (m, 2H, 2 × CH (ACH)), 1.31-1.59 (m, 10H, 4 × CH2 + 2 × CH (ACH)), 1.09-1.29 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 444.1 (M + H+ ); calcd. for C21 H37 N3 O7 : 443.3. S.5a from S.1 and TTDD (85%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0. 1 H NMR (DMSO-d6 ): δ 8.67-8.80 (t, 1H, CH2 -NH-COCO), 8.40-8.55 (d, 1H, COCO-NH-ACH), 4.35-4.80 (br s, 1H, OH), 3.11-3.58 (m, 16H, 6 × CH2 -CH 2 -O + H4 (ACH) + H1 (ACH) + CH 2 -NH-COCO), 2.53-2.62 (t, 2H, NH2 -CH 2 ), 1.32-1.87 (m, 10H, 6 × CH (ACH) + 2 × CH2 -CH 2 -CH2 ), 1.10-1.28 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 390.0 (M + H+ ), 412.0 (M + Na+ ); calcd. for C18 H35 N3 O6 : 389.2. S.6a from S.2 and TTDD (82%). TLC (8:2 CHCl3 /MeOH, UV): Rf = 0. 1 H NMR (DMSOd6 ): δ 8.70-8.97 (m, 2H, (CH2 )3 -NH-COCO + COCO-NH-5 ), 7.42-7.51 (s, 1H, H6), 6.08-6.18 (t, 1H, H1 ), 4.15-4.23 (m, 1H, H3 ), 3.81-3.90 (m, 1H, H4 ), 3.28-3.56 (m, 14H, 6 × CH2 -CH 2 -O + H5 ,5 ), 3.13-3.26 (m, 2H, (CH2 )2 -CH 2 -NH-COCO), 2.522.62 (m, 2H, NH2 -CH 2 ), 1.97-2.15 (m, 2H, H2 ,2 ), 1.77-1.85 (s, 3H, CH3 -5), 1.49-1.75 (m, 4H, CH2 -CH 2 -CH2 ). ESI-MS: m/z 517.0 (M + H+ ); calcd. for C22 H37 N5 O9 : 515.3. From aliphatic primary amino alcohol: S.10a from S.1 and AEE (77%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.16. 1 H NMR (DMSO-d6 ): δ 8.58-8.65 (t, 1H, CH2 -NH-COCO), 8.45-8.52 (d, 1H, COCO-NH-ACH), 4.53-4.62 (m, 2H, HO-CH2 + ACH-OH), 3.23-3.59 (m, 10H, 3 × CH2 -CH 2 -O + H4 (ACH) + CH 2 -NH-COCO + H1 (ACH)), 1.75-1.86 (br d, 2H, 2 × CH (ACH)), 1.601.71 (br d, 2H, 2 × CH (ACH)), 1.32-1.49 (m, 2H, 2 × CH (ACH)), 1.10-1.27 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 275.6 (M + H+ ); calcd. for C12 H22 N2 O5 : 274.1. S.11a from S.1 and AH (60%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.18. 1 H NMR (DMSO-d6 ): δ 8.66-8.74 (t, 1H, CH2 -NH-COCO), 8.39-8.47 (d, 1H, COCO-NH-ACH), 4.52-4.57 (d, 1H, ACH-OH), 4.29-4.35 (t, 1H, HO-CH2 ), 3.43-3.59 (m, 1H, H4 (ACH)), 3.26-3.40 (m, 3H, CH 2 -OH + H1 (ACH)), 3.04-3.14 (q, 2H, CH 2 -NH-COCO), 1.741.85 (br d, 2H, 2 × CH (ACH)), 1.59-1.71 (br d, 2H, 2 × CH (ACH)), 1.09-1.50 (m, 12H, CH2 -(CH 2 )4 -CH2 + 4 × CH (ACH)). ESI-MS: m/z 287.7 (M + H+ ); calcd. for C14 H26 N2 O4 : 286.2.
Perform tritylation With MMTr-Cl for primary amine (S.3a-S.6a) 15a. Prepare a 10 mmol slurry of the amino alcohol in 100 mL of 1:1 (v/v) anhydrous DMF/pyridine and begin stirring. 16a. Add 11 mmol MMTr-Cl and stir at room temperature for 2 to 3 days. 17a. Dilute◦the mixture with 400 mL CHCl3 and filter through a thin (0.5 to 1 cm) layer of 60 A silica gel, under vacuum using a Buchner funnel. 18a. Transfer the filtrate to a separatory funnel and extract with 400 mL saline solution. 19a. Separate the organic layer and dry with anhydrous Na2 SO4 . Filter through a paper filter and concentrate the filtrate to ∼50 mL on a rotary evaporator. 20a. Add 300 mL diethyl ether and then 200 mL pentane with vigorous stirring. 21a. Cool the suspension to 4◦ C and maintain at this temperature for 2 hr. Methoxyoxalamido Chemistry for Tethered Phosphoramidites
22a. Filter the precipitate under vacuum using a Buchner funnel. Wash the solid twice with 30 mL of 1:1 (v/v) diethyl ether/pentane and dry overnight in vacuo (vacuum pump, 0.05 to 0.5 Torr).
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23a. Characterize the compound by TLC, 1 H NMR, and MS. S.3c from S.3a (56%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.42. 1 H NMR (DMSO-d6 ): δ 8.75-8.84 (t, 1H, CH2 -NH-COCO), 8.40-8.48 (d, 1H, COCO-NH-ACH), 7.11-7.41 (m, 12H, MMTr), 6.78-6.86 (d, 2H, MMTr), 4.52-4.58 (d, 1H, OH), 3.69-3.74 (s, 3H, OCH3 ), 3.43-3.60 (m, 1H, H4 (ACH)), 3.20-3.40 (m, 3H, CH 2 -NH-COCO + H1 (ACH)), 2.86-2.95 (m, 1H, MMTr-NH), 2.03-2.15 (m, 2H, MMTr-NH-CH 2 ), 1.74-1.86 (br d, 2H, 2 × CH (ACH)), 1.60-1.72 (br d, 2H, 2 × CH (ACH)), 1.32-1.50 (m, 2H, 2 × CH (ACH)), 1.10-1.28 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 502.6 (M + H+ ); calcd. for C30 H35 N3 O4 : 501.3. S.4c from S.4a (67%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.45. 1 H NMR (DMSO-d6 ): δ 8.70-8.79 (t, 1H, CH2 -NH-COCO), 8.40-8.48 (d, 1H, COCO-NH-ACH), 7.11-7.41 (m, 12H, MMTr), 6.80-6.89 (d, 2H, MMTr), 4.53-4.58 (d, 1H, OH), 4.43-4.49 (t, 1H, CH2 CH-(O-)2 (spiro ring)), 4.35-4.42 (t, 1H, CH2 -CH-(O-)2 (spiro ring)), 4.19-4.28 (br d, 2H, 2 × CH (spiro ring)), 3.68-3.74 (s, 3H, OCH3 ), 3.44-3.60 (m, 5H, 4 × CH (spiro ring) + H4 (ACH)), 3.23-3.42 (m, 3H, 2 × CH (spiro ring) + H1 (ACH)), 3.05-3.16 (q, 2H, CH 2 -NH-COCO), 1.88-2.00 (m, 2H, MMTr-NH-CH 2 ), 1.75-1.86 (br d, 2H, 2 × CH (ACH)), 1.60-1.72 (br d, 2H, 2 × CH (ACH)), 1.32-1.58 (m, 10H, 4 × CH2 + 2 × CH (ACH)), 2H, 1.11-1.27 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 716.2 (M + H+ ), 738.2 (M + Na+ ); calcd. for C41 H53 N3 O8 : 715.4. S.5c from S.5a (65%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.43. 1 H NMR (DMSO-d6 ): δ 8.64-8.75 (t, 1H, CH2 -NH-COCO), 8.41-8.48 (d, 1H, COCO-NH-ACH), 7.11-7.41 (m, 12H, MMTr), 6.80-6.87 (d, 2H, MMTr), 4.53-4.58 (d, 1H, OH), 3.69-3.73 (s, 3H, OCH3 ), 3.27-3.56 (m, 14H, 6 × CH2 -CH 2 -O + H4 (ACH) + H1 (ACH)), 3.10-3.23 (q, 2H, CH 2 -NH-COCO), 2.53-2.62 (t, 1H, MMTr-NH), 1.97-2.08 (q, 2H, MMTr-NHCH 2 ), 1.74-1.86 (br d, 2H, 2 × CH (ACH)), 1.59-1.73 (m, 6H, 2 × CH2 -CH 2 -CH2 + 2 × CH (ACH)), 1.31-1.50 (m, 2H, 2 × CH (ACH)), 1.10-1.28 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 662.6 (M + H+ ), 684.6 (M + Na+ ); calcd. for C38 H51 N3 O7 : 661.4. S.6c from S.6a (83%). TLC (8:2 CHCl3 /MeOH, UV): Rf = 0.61. 1 H NMR (DMSO-d6 ): δ 11.25-11.33 (s, 1H, HN-3), 8.71-8.98 (m, 2H, (CH2 )3 -NH-COCO + COCO-NH-5 ), 7.46-7.51 (s, 1H, H6), 7.11-7.43 (m, 12H, MMTr), 6.79-6.88 (d, 2H, MMTr), 6.09-6.19 (t, 1H, H1 ), 5.29-5.36 (d, 1H, OH-3 ), 4.12-4.23 (m, 1H, H3 ), 3.82-3.91 (m, 1H, H4 ), 3.68-3.77 (s, 3H, OCH3 ), 3.28-3.59 (m, 14H, 6 × CH2 -CH 2 -O + H5 ,5 ), 3.11-3.24 (q, 2H, CH 2 -NH-COCO), 2.53-2.61 (t, 2H, MMTr-NH-CH 2 ), 1.98-2.10 (m, 2H, H2 ,2 ), 1.78-1.84 (s, 3H, CH3 -5), 1.61-1.76 (m, 4H, CH2 -CH 2 -CH2 ). ESI-MS: m/z 787.9 (M + H+ ), 810.9 (M + Na+ ); 826.9 (M + K+ ); calcd. for C42 H53 N5 O10 : 787.4.
With DMTr-Cl for primary hydroxyl (S.10a-S.11a) 15b. Dissolve 10 mmol of the alcohol in 100 mL of 1:1 (v/v) anhydrous DMF/pyridine and begin stirring. 16b. Add 3.73 g (11 mmol) DMTr-Cl and stir overnight at room temperature. Check the course of the reaction by TLC using 1:9 (v/v) MeOH/CHCl3 . Visualize TLC plates under UV or by I2 staining. If the reaction is incomplete, add an additional 0.2 equiv of DMTr-Cl and continue stirring for 5 hr. The starting compound has lower mobility and does not become orange upon heating the TLC plate with a heat gun.
17b. Dilute the mixture with 400 mL CHCl3 . 18b. Transfer the solution to a separatory funnel and extract with 400 mL saline solution. 19b. Separate the organic layer and dry with anhydrous Na2 SO4 . Filter through a paper filter and concentrate the filtrate to ∼50 mL on a rotary evaporator. 20b. Add 300 mL diethyl ether and then 200 mL pentane with vigorous stirring. 21b. Cool the suspension to 4◦ C and maintain at this temperature for 2 hr.
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22b. Filter the precipitate under vacuum using a Buchner funnel. Wash the solid twice with 30 mL of 1:1 (v/v) diethyl ether/pentane and dry overnight in vacuo (vacuum pump, 0.05 to 0.5 Torr). 23b. Characterize the compound by TLC, 1 H NMR, and ESI-MS. S.10c from S.10a (78%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.54. 1 H NMR (DMSO-d6 ): δ 8.58-8.66 (t, 1H, CH2 -NH-COCO), 8.43-8.49 (d, 1H, COCO-NH-ACH), 7.17-7.42 (m, 9H, DMTr), 6.84-6.93 (d, 4H, DMTr), 4.52-4.56 (d, 1H, ACH-OH), 3.69-3.77 (s, 6H, OCH3 ), 3.43-3.60 (m, 5H, 2 × CH2 -CH 2 -O + H4 (ACH)), 3.25-3.39 (m, 3H, CH 2 -NHCOCO + H1 (ACH)), 2.99-3.07 (t, 2H, DMTr-O-CH 2 ), 1.73-1.85 (br d, 2H, 2 × CH (ACH)), 1.57-1.68 (br d, 2H, 2 × CH (ACH)), 1.30-1.47 (m, 2H, 2 × CH (ACH)), 1.091.26 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 599.7 (M + Na+ ); calcd. for C33 H40 N2 O7 : 576.3. S.11c from S.11a (85%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.56. 1 H NMR (DMSO-d6 ): δ 8.65-8.73 (t, 1H, CH2 -NH-COCO), 8.38-8.44 (d, 1H, COCO-NH-ACH), 7.17-7.39 (m, 9H, DMTr), 6.84-6.92 (d, 4H, DMTr), 4.52-4.57 (d, 1H, ACH-OH), 3.70-3.75 (s, 6H, 2 × OCH3 ), 3.42-3.59 (m, 1H, H4 (ACH)), 3.26-3.40 (m, 1H, H1 (ACH)), 3.02-3.13 (q, 2H, CH 2 -NH-COCO), 2.88-2.96 (t, 2H, DMTr-O-CH 2 ), 1.74-1.85 (br d, 2H, 2 × CH (ACH)), 1.60-1.70 (br d, 2H, 2 × CH (ACH)), 1.10-1.58 (m, 12H, CH2 -(CH 2 )4 -CH2 + 4 × CH (ACH)). ESI-MS: m/z 588.7 (M + H+ ); calcd. for C35 H44 N2 O6 : 588.3.
Phosphinylate secondary hydroxyl group 24. Prepare a mixture of 5 mmol dry secondary alcohol (from step 23) and 330 mg (4.75 mmol) tetrazole in 50 mL anhydrous CH2 Cl2 under a dry argon atmosphere. 25. Add 2.15 mL (6.5 mmol) 2-cyanoethyl-N,N,N,N -tetraisopropylphosphorodiamidite and stir the reaction mixture for at least 1 hr or until complete disappearance of the starting material as indicated by TLC analysis. 26. Dilute the reaction with 200 mL CH2 Cl2 (HPLC grade) and extract with 200 mL of 10% aqueous NaHCO3 . 27. Separate the organic layer, dry over anhydrous Na2 SO4 , filter off the drying agent, and concentrate to ∼20 mL on a rotary evaporator. 28. Precipitate the residue into 500 mL of 1:1 (v/v) diethyl ether/pentane and allow the product to settle. 29. Decant the supernatant and dissolve the residue in 50 mL CH2 Cl2 (anhydrous) containing 4% Et3 N. Purify the phosphoramidite by flash chromatography on 40 g silica gel (see APPENDIX 3E) using the appropriate elution solvent system:
20% to 60% (v/v) EtOAc/4% (v/v) Et3 N in pentane (for S.3d-S.5d and S.10d-S.11d) 5% to 50% (v/v) pyridine in EtOAc (for S.6d). 30. Collect and pool fractions containing the desired compound. Identify the correct fractions by TLC using UV or I2 staining for visualization. Evaporate solvent on a rotary evaporator and then co-evaporate twice with ∼50 mL toluene. 31. Dry the residue overnight in vacuo (vacuum pump, 0.05 to 0.5 Torr) over P2 O5 . 32. Characterize the compounds by TLC and 31 P NMR. S.3d from S.3c (85%). TLC (4% Et3 N in 1:1 ethyl acetate/pentane, UV): Rf = 0.50. 31 P NMR (DMSO-d6 ): δ 145.20. Methoxyoxalamido Chemistry for Tethered Phosphoramidites
S.4d from S.4c (87%). TLC (4% Et3 N in 1:1 ethyl acetate/pentane, UV): Rf = 0.35. 31 P NMR (DMSO-d6 ): δ 145.10.
4.29.10 Supplement 25
Current Protocols in Nucleic Acid Chemistry
S.5d from S.5c (91%). TLC (4% Et3 N in 1:1 ethyl acetate/pentane, UV): Rf = 0.39. 31 P NMR (DMSO-d6 ): δ 144.78. S.6d from S.6c (72%). TLC (2% Et3 N in 9:1 ethyl acetate/pyridine, UV): Rf = 0.34. 31 P NMR (DMSO-d6 ): δ 147.13, 147.09. S.10d from S.10c (85%). TLC (4% Et3 N in 1:1 ethyl acetate/pentane, UV): Rf = 0.53. 31 P NMR (DMSO-d6 ): δ 144.74. S.11d from S.11c (84%). TLC (4% Et3 N in 1:1 ethyl acetate/pentane, UV): Rf = 0.62. 31 P NMR (DMSO-d6 ): δ 145.20.
PREPARATION OF PHOSPHORAMIDITES TETHERED WITH MULTIPLE LINKERS
ALTERNATE PROTOCOL
Iterative treatment of the initial MOX precursor S.1 or S.2 with a diamine and dimethyl oxalate leads to MOX derivatives that can also be converted to phosphoramidites (Fig. 4.29.2). This protocol describes the synthesis of a number of tethered phosphoramidites when utilizing 4,7,10-trioxa-1,13-tridecanediamine (TTDD) as the linker. All of these compounds are prepared starting from S.5a and S.6a (see Basic Protocol 1) by performing a second MOX reaction to give the tethered MOX precursors S.5b and S.6b. The final iteration can be performed with a diamine to give a primary amino group (S.7a-S.9a; Fig. 4.29.4) or with an amino alcohol to give a primary hydroxyl group (S.12a-S.14a; Fig. 4.29.5). Tritylation and phosphinylation are then performed as in Basic Protocol 1. When an iteratively tethered MOX intermediate is prepared, a 4-fold excess of dimethyl oxalate is used (as opposed to a 2-fold excess in the case of MOX precursors S.1 and S.2) to minimize formation of a dimer byproduct. If needed, the excess dimethyl oxalate can be easily recovered from the reaction mixture by sublimation.
Perform second MOX reaction (prepare MOX precursors S.5b and S.6b) 1. Prepare a stirring solution of 40 mmol dimethyl oxalate and 20 mmol Et3 N in 50 mL MeOH. 2. Add 10 mmol amino alcohol S.5a or S.6a dissolved in 50 mL of 1:1 (v/v) MeOH/CH2 Cl2 to the stirring solution, portion-wise, over a 2-hr period. Portions should not exceed 5 mL. The amount of a dimer byproduct increases if the amino alcohol solution is added too quickly.
3. Stir the reaction mixture for an additional 1 hr and concentrate to dryness on a rotary evaporator. 4. Sublime the excess dimethyl oxalate at ∼5 Torr and 70◦ C. 5. Dissolve the residue in 50 mL CH2 Cl2 and purify by flash chromatography on a 4 × 40–cm column containing 60 g silica gel (APPENDIX 3E) using a 0% to 30% gradient of MeOH in CHCl3 for elution. 6. Collect and pool fractions containing the desired compound. Identify the correct fractions by TLC using UV or I2 staining for visualization. Evaporate solvent on a rotary evaporator. 7. Dry the residue overnight in vacuo (vacuum pump, 0.05 to 0.5 Torr). 8. Characterize the product by TLC, 1 H NMR, and ESI-MS. S.5b from S.5a (87%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.49. 1 H NMR (DMSO-d6 ): δ 8.83-8.93 (t, 1H, COCONH, MOX), 8.65-8.73 (t, 1H, NHCOCO), 8.38-8.46 (d, 1H, COCONH, ACH), 4.53-4.58 (d, 1H, OH, ACH), 3.73-3.78 (s, 3H, OCH3 , MOX), 3.433.57 (m, 9H, 4 × CH2 + CH), 3.25-3.43 (m, 5H, 2 × CH2 + CH), 3.10-3.24 (q, 8H, 2
Synthesis of Modified Oligonucleotides and Conjugates
4.29.11 Current Protocols in Nucleic Acid Chemistry
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× CH2 ), 1.58-1.84 (overlapping multiplets, 8H, + 2 × CH2 (ACH) + 2 × CH2 (ACH)), 1.30-1.49 (m, 2H, CH2 , ACH), 1.07-1.26 (m, 2H, CH2 , ACH). ESI-MS: m/z 476.3 (M + H+ ), 498.3 (M + Na+ ); calcd. for C21 H37 N3 O9 : 475.5. S.6b from S.6a (90%). TLC (8:2 CHCl3 /MeOH, UV): Rf = 0.54. 1 H NMR (DMSO-d6 ): δ 11.26-11.32 (s, 1H, HN-3), 8.81-8.94 (m, 2H, 2 × (CH2 )3 -NH-COCO), 8.69-8.80 (t, 1H, COCO-NH-5 ), 7.43-7.50 (s, 1H, H6), 6.09-6.18 (t, 1H, H1 ), 5.27-5.38 (br s, 1H, OH-3 ), 4.14-4.23 (br s, 1H, H3 ), 3.82-3.90 (m, 1H, H4 ), 3.73-3.79 (s, 3H, OCH3 ), 3.28-3.57 (m, 14H, 6 × CH2 -CH 2 -O + H5, 5 ), 3.13-3.25 (m, 4H, 2 × (CH2 )2 -CH 2 NH-COCO), 1.98-2.18 (m, 2H, H2 ,2 ), 1.78-1.86 (s, 3H, CH3 -5), 1.62-1.77 (m, 4H, CH2 -CH 2 -CH2 ). ESI-MS: m/z 625.0 (M + Na+ ); calcd. for C25 H39 N5 O12 : 601.3.
Perform second derivatization 9. Follow steps 9 to 14 of Basic Protocol 1 using TTDD to prepare amino alcohols (S.7a-S.8a) or using AEE to prepare diols (S.12a or S.13a) from S6.b and S.5b. S.7a from S.6b and TTDD (83%). TLC (8:2 CHCl3 /MeOH, UV): Rf = 0. 1 H NMR (DMSO-d6 ): δ 8.65-8.92 (m, 4H, 3 × NH-COCO + COCO-NH-5 ), 7.44-7.50 (s, 1H, H6), 6.07-6.17 (t, 1H, H1 ), 5.20-5.45 (br s, 1H, OH-3 ), 4.14-4.23 (m, 1H, H3 ), 3.813.90 (m, 1H, H4 ), 3.25-3.58 (m, 26H, 12 × CH2 -CH 2 -O + H5 ,5 ), 3.07-3.24 (m, 8H, 3 × (CH2 )2 -CH 2 -NH-COCO + NH2 -CH 2 ), 1.98-2.17 (m, 2H, H2 ,2 ), 1.77-1.85 (s, 3H, CH3 -5), 1.59-1.76 (m, 8H, CH2 -CH 2 -CH2 ). ESI-MS: m/z 790.7 (M + H+ ); calcd. for C34 H59 N7 O14 : 789.4. S.8a from S.5b and TTDD (85%). TLC (8:2 CHCl3 /MeOH, UV): Rf = 0. 1 H NMR (DMSO-d6 ): δ 8.66-8.76 (t, 3H, 3 × CH2 -NH-COCO), 8.40-8.47 (d, 1H, COCO-NHACH), 4.44-4.68 (br s, 1H, OH), 3.27-3.56 (m, 26H, 12 × CH2 -CH 2 -O + H4 (ACH) + H1 (ACH)), 3.12-3.24 (q, 6H, 3 × CH 2 -NH-COCO), 2.60-2.71 (t, 2H, NH2 -CH 2 ), 1.75-1.86 (m, 2H, 2 × CH (ACH)), 1.55-1.74 (m, 10H, 4 × CH2 -CH 2 -CH2 + 2 × CH (ACH)), 1.32-1.49 (m, 2H, 2 × CH (ACH)), 1.10-1.27 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 664.3 (M + H+ ); calcd. for C30 H57 N5 O11 : 663.4. S.12a from S.6b and AEE (89%). TLC (8:2 CHCl3 /MeOH, UV): Rf = 0.41. 1 H NMR (DMSO-d6 ): δ 11.25-11.32 (s, 1H, HN-3), 8.81-8.90 (t, 1H, (CH2 )3 -NH-COCO), 8.688.79 (m, 2H, COCO-NH-5 + NH-COCO), 8.56-8.65 (t, 1H, NH-COCO), 7.42-7.51 (s, 1H, H6), 6.08-6.18 (t, 1H, H1 ), 5.28-5.36 (d, 1H, OH-3 ), 4.55-4.64 (t, 1H, HO-CH2 ), 4.13-4.24 (m, 1H, H3 ), 3.80-3.90 (m, 1H, H4 ), 2.97-3.58 (m, 26H, 8 × CH2 -CH 2 -O + H5 ,5 + O-CH2 -CH 2 -NH-COCO + HO-CH 2 + 2 × O-CH2 -CH2 -CH 2 -N), 1.98-2.16 (m, 2H, H2 ,2 ), 1.78-1.85 (s, 3H, CH3 -5), 1.62-1.76 (m, 4H, CH2 -CH 2 -CH2 ). ESI-MS: m/z 675.8 (M + H+ ), 697.8 (M + Na+ ); calcd. for C28 H46 N6 O13 : 674.3. S.13a from S.5b and AEE (90%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.20. 1 H NMR (DMSO-d6 ): δ 8.67-8.79 (m, 2H, 2 × O-(CH2 )3 -NH-COCO), 8.58-8.65 (t, 1H, O(CH2 )2 -NH-COCO), 8.40-8.48 (d, 1H, COCO-NH-ACH), 4.53-4.63 (m, 2H, ACH-OH + HO-CH 2 ), 3.25-3.56 (m, 20H, H4 (ACH) + 8 × O-CH 2 + H1 (ACH) + O-CH2 CH 2 NHCOCO), 3.13-3.23 (m, 4H, 2 × (CH2 )2 CH 2 -NHCOCO), 3.01-3.12 (t, 2H, HO-CH 2 ), 1.60-1.86 (m, 8H, 4 × CH (ACH) + 2 × CH2 CH 2 CH2 ), 1.32-1.49 (m, 2H, 2 × CH (ACH)), 1.11-1.26 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 549.8 (M + H+ ), 571.8 (M + Na+ ); calcd. for C24 H44 N4 O10 : 548.3.
Perform third MOX reaction (prepare MOX precursor S.8b) 10. Follow steps 1 to 8 of this protocol to prepare S.8b from S.8a.
Methoxyoxalamido Chemistry for Tethered Phosphoramidites
S.8b from S.8a (71%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.48. 1 H NMR (DMSO-d6 ): δ 8.82-8.93 (t, 1H CH3 O-COCO-NH), 8.66-8.75 (t, 3H, 3 × CH2 -NH-COCO), 8.41-8.48 (d, 1H, COCO-NH-ACH), 4.53-4.57 (d, 1H, OH), 3.73-3.76 (s, 3H, CH3 O-COCO), 3.25-3.55 (m, 26H, 12 × CH2 -CH 2 -O + H4 (ACH) + H1 (ACH)), 3.11-3.21 (m, 8H, 3 × CH 2 -NH-COCO), 1.75-1.86 (m, 2H, 2 × CH (ACH)), 1.60-1.74 (m, 10H, 4 × CH2 -CH 2 -CH2 + 2 × CH (ACH)), 1.33-1.49 (m, 2H, 2 × CH (ACH)), 1.11-1.26 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 750.3 (M + H+ ); calcd. for C33 H59 N5 O14 : 749.4.
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Perform third derivatization 11. Follow steps 9 to 14 of Basic Protocol 1 using TTDD to prepare the amino alcohol (S.9a) or using AEE to prepare the diol (S.14a) from S.8b. S.9a from S.8b and TTDD (87%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0. 1 H NMR (DMSO-d6 ): δ 8.67-8.75 (t, 5H, 5 × CH2 -NH-COCO), 8.40-8.47 (d, 1H, COCO-NHACH), 4.53-4.57 (d, 1H, OH), 3.25-3.55 (m, 38H, 18 × CH2 -CH 2 -O + H4 (ACH) + H1 (ACH)), 3.12-3.23 (q, 10H, 5 × CH 2 -NH-COCO), 2.55-2.62 (t, 2H, NH2 -CH 2 ), 1.31-1.86 (m, 18H, 6 × CH (ACH) + 6 × CH2 -CH 2 -CH2 ), 1.07-1.28 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 938.3 (M + H+ ); calcd. for C42 H79 N7 O16 : 937.6. S.14a from S.8b and AEE (91%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.16. 1 H NMR (DMSO-d6 ): δ 8.66-8.78 (m, 4H, O-(CH2 )3 -NH-COCO), 8.57-8.65 (t, 1H, O-(CH2 )2 NH-COCO), 8.40-8.47 (d, 1H, COCO-NH-ACH), 4.51-4.63 (m, 2H, HO-CH2 + ACHOH), 3.25-3.57 (m, 34H, H4 (ACH) + 16 × CH 2 + H1 (ACH)), 3.11-3.24 (q, 8H, 4 × (CH2 )2 CH 2 NHCOCO), 1.60-1.86 (m, 12H, 4 × CH (ACH) + 4 × CH2 CH 2 CH2 ), 1.30-1.49 (m, 2H, 2 × CH (ACH)), 1.09-1.27 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 845.3 (M + Na+ ); calcd. for C36 H66 N6 O15 : 822.5.
Tritylate primary amino or hydroxyl group 12. Follow steps 15a to 23a of Basic Protocol 1 to protect primary amino groups with MMTr-Cl, or steps 15b to 23b to protect primary hydroxyl groups with DMTr-Cl. S.7c from S.7a (72%). TLC (8:2 CHCl3 /MeOH, UV): Rf = 0.56. 1 H NMR (DMSO-d6 ): δ 11.25-11.33 (s, 1H, HN-3), 8.82-8.91 (t, 1H, (CH2 )3 -NH-COCO), 8.62-8.80 (m, 3H, COCO-NH-5 + 2 × NH-COCO), 7.45-7.52 (s, 1H, H6), 7.11-7.43 (m, 12H, MMTr), 6.80-6.88 (d, 2H, MMTr), 6.09-6.18 (t, 1H, H1 ), 5.29-5.36 (d, 1H, OH-3 ), 4.13-4.23 (m, 1H, H3 ), 3.82-3.90 (m, 1H, H4 ), 3.68-3.77 (s, 3H, OCH3 ), 3.28-3.59 (m, 26H, 12 × CH2 -CH 2 -O + H5 ,5 ), 3.12-3.26 (m, 6H, 3 × (CH2 )2 -CH 2 -NH-COCO), 2.53-2.61 (t, 2H, MMTr-NH-CH 2 ), 1.98-2.11 (m, 2H, H2 ,2 ), 1.78-1.84 (s, 3H, CH3 -5), 1.61-1.76 (m, 8H, CH2 -CH 2 -CH2 ). ESI-MS: m/z 1062.8 (M + H+ ); calcd. for C54 H75 N7 O15 : 1061.5. S.8c from S.8a (61%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.42. 1 H NMR (DMSO-d6 ): δ 8.66-8.75 (t, 3H, CH2 -NH-COCO), 8.41-8.48 (d, 1H, COCO-NH-ACH), 7.11-7.42 (m, 12H, MMTr), 6.80-6.88 (d, 2H, MMTr), 4.53-4.57 (d, 1H, OH), 3.69-3.74 (s, 3H, OCH3 ), 3.27-3.56 (m, 26H, 12 × CH2 -CH 2 -O + H4 (ACH) + H1 (ACH)), 3.12-3.23 (q, 6H, 3 × CH 2 -NH-COCO), 2.53-2.61 (t, 1H, MMTr-NH), 1.97-2.08 (q, 2H, MMTr-NH-CH 2 ), 1.75-1.86 (br d, 2H, 2 × CH (ACH)), 1.60-1.74 (m, 10H, 4 × CH2 -CH 2 -CH2 + 2 × CH (ACH)), 1.32-1.50 (m, 2H, 2 × CH (ACH)), 1.12-1.27 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 936.4 (M + H+ ); calcd. for C50 H73 N5 O12 : 935.5. S.9c from S.9a (66%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.40. 1 H NMR (DMSO-d6 ): δ 8.67-8.75 (t, 5H, CH2 -NH-COCO), 8.41-8.48 (d, 1H, COCO-NH-ACH), 7.11-7.42 (m, 12H, MMTr), 6.80-6.88 (d, 2H, MMTr), 4.52-4.57 (d, 1H, OH), 3.68-3.74 (s, 3H, OCH3 ), 3.25-3.56 (m, 38H, 18 × CH2 -CH 2 -O + H4 (ACH) + H1 (ACH)), 3.11-3.23 (q, 10H, 5 × CH 2 -NH-COCO), 2.53-2.60 (t, 1H, MMTr-NH), 1.97-2.08 (q, 2H, MMTr-NH-CH 2 ), 1.75-1.86 (br d, 2H, 2 × CH (ACH)), 1.60-1.74 (m, 14H, 6 × CH2 -CH 2 -CH2 + 2 × CH (ACH)), 1.30-1.50 (m, 2H, 2 × CH (ACH)), 1.12-1.27 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 1210.3 (M + H+ ); calcd. for C62 H95 N7 O17 : 1209.7. S.12c from S.12a (81%). TLC (8:2 CHCl3 /MeOH, UV): Rf = 0.66. 1 H NMR (DMSOd6 ): δ 11.27-11.30 (s, 1H, HN-3), 8.82-8.89 (t, 1H, (CH2 )3 -NH-COCO), 8.71-8.78 (m, 2H, COCO-NH-5 + NH-COCO), 8.58-8.65 (t, 1H, NH-COCO), 7.45-7.48 (s, 1H, H6), 7.19-7.41 (m, 9H, DMTr), 6.85-6.91 (d, 4H, DMTr), 6.09-6.15 (t, 1H, H1 ), 5.32-5.35 (d, 1H, OH-3 ), 4.15-4.22 (m, 1H, H3 ), 3.82-3.89 (m, 1H, H4 ), 3.71-3.75 (s, 6H, OCH3 ), 3.29-3.59 (m, 20H, 8 × CH2 -CH 2 -O + H5 ,5 + O-CH2 -CH 2 -NH-COCO), 3.14-3.23 (m, 4H, 2 × (CH2 )2 -CH 2 -NH-COCO), 3.01-3.11 (t, 2H, DMTr-O-CH 2 ), 1.99-2.15 (m, 2H, H2 ,2 ), 1.78-1.82 (s, 3H, CH3 -5), 1.62-1.74 (m, 4H, CH2 -CH 2 -CH2 ). ESI-MS: m/z 697.8 (M – DMTr + Na+ ), 999.9 (M + Na+ ); calcd. for C49 H64 N6 O15 : 976.4.
Synthesis of Modified Oligonucleotides and Conjugates
4.29.13 Current Protocols in Nucleic Acid Chemistry
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S.13c from S.13a (75%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.42. 1 H NMR (DMSO-d6 ): δ 8.59-8.79 (m, 3H, 2 × O-(CH2 )3 -NH-COCO + O-(CH2 )2 -NH-COCO), 8.41-8.47 (d, 1H, COCO-NH-ACH), 7.17-7.42 (m, 9H, DMTr), 6.85-6.91 (d, 4H, DMTr), 4.53-4.57 (d, 1H, ACH-OH), 3.70-3.77 (s, 6H, 2 × OCH3 ), 3.28-3.60 (m, 20H, H4 (ACH) + 8 × O-CH 2 + H1 (ACH) + O-CH2 CH 2 -NHCOCO), 3.12-3.23 (m, 4H, 2 × (CH2 )2 CH 2 NHCOCO), 3.00-3.07 (t, 2H, DMTr-O-CH 2 ), 1.75-1.87 (m, 2H, 2 × CH (ACH)), 1.611.74 (m, 6H, 2 × CH2 CH 2 CH2 + 2 × CH (ACH)), 1.32-1.49 (m, 2H, 2 × CH (ACH)), 1.10-1.27 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 571.6 (M – DMTr + Na+ ), 873.7 (M + Na+ ); calcd. for C45 H62 N4 O12 : 850.4. S.14c from S.14a (86%). TLC (9:1 CHCl3 /MeOH, UV): Rf = 0.41. 1 H NMR (DMSO-d6 ): δ 8.66-8.77 (m, 4H, 4 × O-(CH2 )3 -NH-COCO), 8.58-8.64 (t, 1H, O-(CH2 )2 -NH-COCO), 8.40-8.46 (d, 1H, COCO-NH-ACH), 7.17-7.42 (m, 9H, DMTr), 6.85-6.92 (d, 4H, DMTr), 4.52-4.56 (d, 1H, ACH-OH), 3.70-3.78 (s, 6H, 2 × OCH3 ), 3.28-3.60 (m, 32H, H4 (ACH) + 15 × CH 2 + H1 (ACH)), 3.14-3.28 (q, 8H, 4 × (CH2 )2 CH 2 NHCOCO), 3.02-3.08 (t, 2H, DMTr-O-CH 2 ), 1.75-1.86 (m, 2H, 2 × CH (ACH)), 1.61-1.74 (m, 10H, 4 × CH2 CH 2 CH2 + 2 × CH (ACH)), 1.32-1.50 (m, 2H, 2 × CH (ACH)), 1.09-1.28 (m, 2H, 2 × CH (ACH)). ESI-MS: m/z 845.3 (M – DMTr + Na+ ); calcd. for C57 H84 N6 O17 : 1124.6.
Phosphinylate secondary hydroxyl group 13. Follow steps 24 to 32 of Basic Protocol 1 to phosphinylate the secondary hydroxyl of the MMTr/DMTr-protected intermediates. To purify the phosphoramidites by flash chromatography, use the following elution solvent systems: 5% to 50% (v/v) pyridine in EtOAc (for S.7d and S.12d) 40% to 100% (v/v) EtOAc/4% (v/v) Et3 N in pentane (for S.8d, S.9d, S.13d, and S.14d). 7d from 7c (80%). TLC (2% Et3 N in 9:1 ethyl acetate/pyridine, UV): Rf = 0.35. 31 P NMR (DMSO-d6 ): δ 147.12, 147.10. 8d from 8c (83%). TLC (4% Et3 N in 1:1 ethyl acetate/pentane, UV): Rf = 0.37. 31 P NMR (DMSO-d6 ): δ 144.77. 9d from 9c (70%). TLC (4% Et3 N in 1:1 ethyl acetate/pentane, UV): Rf = 0.31. 31 P NMR (DMSO-d6 ): δ 144.78. 12d from 12c (68%). TLC (2% Et3 N in 9:1 ethyl acetate/pyridine, UV): Rf = 0.36. 31 P NMR (DMSO-d6 ): δ 147.12, 147.04. 13d from 13c (65%). TLC (4% Et3 N in 1:1 ethyl acetate/pentane, UV): Rf = 0.39. 31 P NMR (DMSO-d6 ): δ 144.74. 14d from 14c (69%). TLC (4% Et3 N in 1:1 ethyl acetate/pentane, UV): Rf = 0.36. 31 P NMR (DMSO-d6 ): δ 144.75. BASIC PROTOCOL 2
Methoxyoxalamido Chemistry for Tethered Phosphoramidites
SYNTHESIS, DEPROTECTION, AND PURIFICATION OF OLIGONUCLEOTIDES DERIVATIZED WITH TETHERED PHOSPHORAMIDITES All tethered phosphoramidites described in this unit are based on cyclic carbon scaffolds with a secondary hydroxyl, and are therefore as stable as standard phosphoramidites. They can be stored for years at −20◦ C. After phosphoramidite installation on the synthesizer, the coupling yields do not deteriorate for at least 2 weeks. The described phosphoramidites are generally as reactive as standard phosphoramidites. However, if tetrazole is used as a catalyst, it is recommended to increase coupling time to 3 to 5 min to ensure complete incorporation. If the more potent 5-ethyl-thio tetrazole (ETT) is used, coupling time could be reduced to 1 to 3 min. For higher-molecular-weight phosphoramidites tethered to multiple linkers (S.9d and S.14d) the coupling time should be set to 10 min with tetrazole and 5 min with ETT.
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All tethers, including those with multiple oxalamido groups, are stable to final deprotection with neat ethanolamine (70◦ C, 15 to 30 min) or concentrated aqueous ammonia (70◦ C, 8 to 16 hr). Because the oxalamido group is not very stable in aqueous alkaline solutions, deprotection procedures involving NaOH treatment must be avoided. The ethanolamine-based deprotection procedure detailed below is fast, reliable, and does not necessitate tight sealing of the reaction vial. It should be stressed, however, that N4 -acetyl-protected deoxycytidine phosphoramidites must be used to prevent N4 -sideproduct formation (Polushin et al., 1994; Reddy et al., 1994). Oligonucleotide synthesis can be carried out on any automated DNA/RNA synthesizer. The authors have used the Biosset ASM-800 DNA/RNA synthesizer. The signature feature of this synthesizer is low reagent consumption, which becomes essential when valuable phosphoramidites are used. To purify crude 5 -modified oligonucleotides, a number of standard techniques can be employed, e.g., reversed-phase or ion-exchange HPLC, cartridge purification, or denaturing PAGE. Of these, PAGE is easy to implement and generally gives oligonucleotides of higher purity. For PAGE purification, the last MMTr/DMTr protecting group (from 5 -functionalization with tethered phosphoramidites) should be removed on the synthesizer (DMTr-OFF mode). The MMTr group is more stable, so the deprotecting time should be set to 3 to 5 min. The molecular mass of a number of functionalized oligonucleotides was assessed by MALDI-TOF-MS on a Voyager-DE mass spectrometer.
Materials Tethered phosphoramidites (S.3d-S.14d; see Basic Protocol 1 and Alternate Protocol) Acetonitrile (CH3 CN), anhydrous, DNA synthesis grade Standard 2 -deoxyribonucleoside phosphoramidites (Transgenomic) 0.5 M tetrazole in CH3 CN (Glen Research) or 0.25 M 5-ethylthio-1H-tetrazole (ETT, Glen Research) Ethanolamine (EA), ≥99% (Aldrich) 10% (w/v) LiClO4 in ethanol (EtOH) Ethanol (EtOH), 200 proof 7 M urea 15% (w/v) polyacrylamide gel (APPENDIX 3B) containing 7 M urea in 0.5× TBE electrophoresis buffer (APPENDIX 2A) 0.25 M triethyl ammonium bicarbonate (TEAB), aqueous solution 1.7-mL microcentrifuge tubes 70◦ C incubator or water bath Microcentrifuge Speedvac evaporator (Savant) Sephadex G-25 NAP-10 columns (Pharmacia) Lyophilizer Water aspirator or vacuum pump (5 to 10 Torr) Additional reagents and equipment for automated solid-phase oligonucleotide synthesis (APPENDIX 3C), purification of oligonucleotides (UNITS 10.1, 10.4, 10.5, 10.7 & APPENDIX 3B), and determination of molecular mass (UNIT 10.1) 1. Dissolve the desired tethered phosphoramidite in anhydrous CH3 CN at 0.1 M in an appropriate vial. Install the vial onto a DNA/RNA synthesizer. 2. Start the automated solid-phase oligonucleotide synthesis (APPENDIX 3C) from an appropriate solid-support-filled column.
Synthesis of Modified Oligonucleotides and Conjugates
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3. Elongate the desired oligonucleotide chain using standard 2 -deoxyribonucleoside phosphoramidites. Use the selected specialty phosphoramidite for the last coupling. If tetrazole is used as a catalyst, allow 3 to 5 min for coupling phosphoramidites S.3d-S.8d and S.10d-S.13d, and 10 min for coupling phosphoramidites S.9d and S.14d. In the case of ETT, reduce the coupling times to 1 to 3 min and 5 min, respectively. 4. Remove the last MMTr/DMTr protecting group manually or by engaging the DMTrOFF mode. Allow 3 to 5 min for removal of the MMTr group. 5. Remove the column from the synthesizer and dry for 15 to 20 min in vacuo using a water aspirator or vacuum pump. 6. Remove the column filter and transfer the support into a 1.7-mL microcentrifuge tube (up to 10 mg per tube) and add 50 µL EA. Vortex briefly to make sure that all support is covered with EA and then microcentrifuge briefly at 13,000 rpm. 7. Incubate at 70◦ C for 20 to 25 min. For oligonucleotides >50 nt, increase the incubation time to 30 to 35 min. 8. Add 20 µl of 10% (w/v) LiClO4 in EtOH and vortex. Add 700 µL of EtOH, vortex, and leave at room temperature for at least 15 to 20 min to ensure full precipitation of the modified oligonucleotide. 9. Microcentrifuge 5 min at 13,000 rpm. Discard the supernatant. Wash the residue with 700 µL EtOH, microcentrifuge 5 min at 13,000 rpm, and discard the wash. 10. Add 200 µL deionized water, vortex, and microcentrifuge briefly at 13,000 rpm. Transfer the solution to a new 1.7-mL tube. Wash the remaining solid support with 200 µL deionized water, microcentrifuge, and transfer the wash to the main solution. 11. Concentrate the solution to dryness in vacuo on a Speedvac evaporator and dissolve the residue in 7 M urea. 12. Purify by denaturing polyacrylamide gel electrophoresis (PAGE; UNIT 10.4 & APPENDIX 3B) using 15% polyacrylamide and 7 M urea in 0.5× TBE electrophoresis buffer. 13. Cut out the band(s) and elute the product with 0.25 M TEAB. Desalt using a Sephadex G-25 NAP-10 column and lyophilize. 14. Dissolve the oligonucleotide in deionized water and quantitate by measuring UV absorbance at 260 nm. Store the solution at –20◦ C. 15. Analyze the products by MALDI-TOF-MS (UNIT mass.
10.1)
to determine molecular
COMMENTARY Background Information
Methoxyoxalamido Chemistry for Tethered Phosphoramidites
Use of specialty phosphoramidites, such as amino modifiers and spacers, is widespread in modern oligonucleotide synthesis. A number of phosphoramidites with aminated and hydroxylated linkers are commercially available (Glen Research), but they generally possess at least three limitations. First, these phosphoramidites are mainly derived from aliphatic primary alcohols and are thus not
very stable even at low temperatures due to susceptibility to Arbuzov rearrangement (Polushin, 2000). This instability leads to preparation and storage complications and also necessitates fairly rapid phosphoramidite consumption on automated DNA synthesizers, thus making the commercially available amino modifiers and spacers less than ideal for reliable large-scale oligonucleotide production. Second, the available amino modifier
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and spacer phosphoramidites are quite limited in variety. They are primarily flexible in nature, since they contain methylene or oxyethylene units within the tether chain, and are rather short, not exceeding 19 atoms in length. Third, the production chemistry for these phosphoramidites is not amenable to constructing a vast number of distinctly different monomers, each possessing varying degrees of flexibility, hydrophobicity, and length. This unit elaborates on a general synthetic route toward phosphoramidite linkers. The method is based on methoxyoxalamido (MOX) chemistry, which is very robust in the synthesis of modified oligonucleotides (Polushin, 2000). MOX chemistry is easy to implement and offers great control over tether composition, rigidity, and length. All βcyanoethyl phosphoramidites in this unit are derived from secondary alcohols and can be used on the synthesizer for at least two weeks without any loss in coupling efficiency. Phosphoramidite synthesis The general synthetic routes towards novel phosphoramidites tethered with single or multiple linkers are outlined in Figures 4.29.1 and 4.29.2, respectively. Phosphoramidite preparations proceed by a single linker addition (for a single tether) or by serial linker additions (for a compound tether). In either case, the reactions proceed in a straightforward manner and the synthesis proves to be robust. The stepwise yields are high, and crystallization or precipitation effectively cleans up the majority of the reactions leading up to the phosphoramidite. Column chromatography is used only at the final phosphinylation step and to purify the MOX intermediates (S.5b, S.6b, and S.8b). The choice of starting compound (phosphoramidite base) for the synthesis of tethered phosphoramidites is based on two major considerations. First, it must have a primary amino group as a starting point for MOX chemistry. Second, the hydroxyl group for attachment of the phosphoramidite moiety should be secondary to impart greater stability to the phosphoramidite (Polushin, 2000). It is also essential that the starting material be commercially available and inexpensive, or can be easily prepared from an inexpensive reagent. The trans-4-aminocyclohexanol hydrochloride and 5 -amino-5 -deoxythymidine used as starting compounds in this unit both satisfy all the above demands.
Structural versatility The practical advantage of the described synthetic approach lies in its amenability to structural versatility by utilizing an arsenal of distinctly different, commercially available diamines and amino alcohols. It is possible to tailor the synthesis to suit practically any requirement in chemical or physical properties of the tethers. The selection of tethers for this unit encompasses only a short range of hydrophobicity, rigidity, and length, but the synthetic strategy can easily produce a much wider variety of chain features. Further flexibility comes from the possibility of assembling compound tethers through the use of oxalamido bonding. This modular assembly allows different combinations of available diamines to be used, and thus allows selective modification of the physical nature of the tether at each step. For example, if more rigidity is needed at the end of the tether, a diamine with a constrained configuration can be used at the last step of tether construction. By exploiting the serial addition strategy and a fairly long diamine such as the 15atom TTDD, one can easily prepare specialty phosphoramidites with extraordinarily long tethers. Through three iterations of MOX chemistry, phosphoramidites with a 56- or a 48-atom tether were synthesized (S.9d or S.14d, respectively). Theoretically, there is no limit to the length that can be achieved using this strategy. Recently biotin amino phosphoramidites with tethers up to 106 atoms in length have been synthesized (Polushin, unpub. observ.). It is essential to stress that all tethered compounds prepared through the MOX strategy are monodispersed, unlike previously described polydispersed tethers prepared from low-molecular-weight polyethylene glycol (PEG) polymer (Jaschke et al., 1994). Synthesis of modified oligonucleotides: Coupling and deprotection Most of the described phosphoramidites are solids and thus easier to handle. All are freely soluble in acetonitrile. The average coupling efficiency of the phosphoramidites at 0.1 M, when using ETT as a catalyst over a 3 to 5 min coupling time, was greater than 95%. The tethered phosphoramidites presented here have been used to synthesize a number of 5 -modified oligonucleotides. The modified oligonucleotides were observed as distinct bands on polyacrylamide gels, with mobilities
Synthesis of Modified Oligonucleotides and Conjugates
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Figure 4.29.6 PAGE analysis of crude 5 -modified T10 oligonucleotides. Lane 1 of each gel corresponds to the unmodified T10 oligonucleotide. The other lanes correspond to oligonucleotides modified at the 5 terminus with the following phosphoramidites: S.3d (A2), S.4d (A3), S.5d (A4), S.6d (A5), S.8d (A6), S.7d (A7), S.9d (A8), S.10d (B2), S.11d (B3), S.13d (B4), S.12d (B5), S.14d (B6).
Table 4.29.1 MALDI-TOF-MS Data for Modified T10 Oligonucleotides
Starting oligonucleotide
Modifier
Lanea
MW (calculated)
MW (experimental)
(Tp)9 T
none
A1, B1
2978
2980
(Tp)9 T
3d
A2
3271
3274
(Tp)9 T
4d
A3
3485
3488
(Tp)9 T
5d
A4
3431
3433
(Tp)9 T
6d
A5
3557
3560
(Tp)9 T
8d
A6
3706
3709
(Tp)9 T
7d
A7
3832
3834
(Tp)9 T
9d
A8
3980
3983
(Tp)9 T
10d
B2
3316
3318
(Tp)9 T
11d
B3
3328
3330
(Tp)9 T
13d
B4
3590
3592
(Tp)9 T
12d
B5
3717
3719
(Tp)9 T
14d
B6
3865
3867
a Lane numbers correspond to Figure 4.29.6.
Methoxyoxalamido Chemistry for Tethered Phosphoramidites
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less than those of unmodified counterparts. As an example, Figure 4.29.6 shows a series of crude T10 oligonucleotides modified by the addition of tethered phosphoramidites. MS data for these oligomers are presented in Table 4.29.1. To determine the effect of oligonucleotide deprotection conditions on tethered compounds, several MMTr-protected intermediate amino alcohols and DMTr-protected intermediate diols were subjected to neat ethanolamine, ammonia/methylamine (30% aq. ammonium hydroxide/2 M methanolic methylamine, 1:1, v/v), and aqueous NaOH (0.2 M NaOH in water/methanol, 1:1, v/v). Exposure to NaOH for even 1 hr at ambient temperature was too harsh, and some degradation of all tethers was detected under these conditions. However, the tethers did hold up to ammonia, methylamine, and ethanolamine, and they were stable at the temperature and duration required for complete deprotection of the modified oligonucleotides.
Critical Parameters and Troubleshooting The synthesis of tethered phosphoramidites is straightforward and efficient. However, familiarity with general chemical laboratory techniques such as filtration, extraction, concentration, TLC, and column chromatography is a must. General knowledge of standard analytical techniques (1 H NMR, 31 P NMR, UV, and mass spectroscopies) is necessary for characterization of products. The efficiency of specialty phosphoramidite incorporation may be influenced by the particular model of the DNA/RNA synthesizer; thus, some optimization of critical parameters (e.g., coupling time, delivery time) might be needed to maximize the coupling yields. Practical knowledge of gel electrophoresis is required for oligonucleotide purification. Laboratory safety must always be a primary concern. The amidites are quite stable in the presence of triethylamine and no special precautions are used prior to removal of triethylamine. Although triethylamine is known to reduce phosphoramidite coupling efficiency, the final toluene co-evaporations are sufficient to remove triethylamine completely.
prepared in 40% to 50% yields starting from the MOX precursors. Coupling efficiency of all phosphoramidites should be greater than 95%.
Time Considerations Each of the initial MOX precursors S.1 and S.2 can be prepared in 2 days. Since these precursors are the starting materials for all other compounds, at least 10-g preparations are recommended. The synthesis of a phosphoramidite with a single hydroxylated linker (S.10d-S.11d) generally takes 3 to 5 days, while synthesis of a phosphoramidite with a single aminated linker (S.3d-S.6d) takes 2 days longer. One round of tether elongation (diamine treatment followed by dimethyl oxalate treatment) can be accomplished in 2 to 3 days. Typically, only one or a few incorporations of specialty phosphoramidites are required in the synthesis of a modified oligonucleotide. Thus, the length of the oligonucleotide and the synthesizer throughput dictate the time needed for synthesis. On the ASM-800 DNA/RNA synthesizer, eight 5 -modified 20-mers can be synthesized in ∼2 hr. EA deprotection and desalting (by precipitation) takes less than an hour. PAGE purification and desalting requires 1 to 2 days.
Literature Cited Bannwarth, W. 1988. Solid phase synthesis of oligonucleotides containing phosphoramidate internucleotide linkages and their specific chemical cleavage. Helv. Chim. Acta 71:1517-1527. Jaschke, A., Furste, J.P., Nordhoff, E., Hillenkamp, F., Cech, D., and Erdmann, V.A. 1994. Synthesis and properties of oligodeoxyribonucleotidepolyethylene glycol conjugates. Nucl. Acids Res. 22:4810-4817. Polushin, N.N. 2000. The precursor strategy: Terminus methoxyoxalamido modifiers for single and multiple functionalization of oligodeoxyribonucleotides. Nucl. Acids Res. 28:31253133. Polushin, N.N., Morocho, A.M., Chen, B.C., and Cohen, J.S. 1994. On the rapid deprotection of synthetic oligonucleotides and analogs. Nucl. Acids Res. 22:639-645. Reddy, M.P., Hanna, N.B., and Farooqui, F. 1994. Fast cleavage and deprotection of oligonucleotides. Tetrahedron Lett. 35:4311-4314.
Anticipated Results Good to moderate yields of the final specialty phosphoramidites from the corresponding MOX precursors are expected. Phosphoramidites tethered with single linkers (S.3d-S.6d, S.10d, and S.11d) have been
Contributed by Alan M. Morocho and Nikolai N. Polushin Fidelity Systems, Inc. Gaithersburg, Maryland
Synthesis of Modified Oligonucleotides and Conjugates
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Supplement 25
Using Morpholinos to Control Gene Expression
UNIT 4.30
Morpholino oligos (Morpholinos) are synthetic uncharged P-chiral analogs of nucleic acids. They are typically constructed by linking together 25 subunits, each bearing one of the four nucleic acid bases. Figure 4.30.1 illustrates the structure of three Morpholino subunits joined by inter-subunit linkages. The morpholino phosphorodiamidate backbone of Morpholinos consists of morpholine rings that bear methylene groups and are bound through modified phosphates in which the anionic oxygen is replaced by a nonionic dimethylamino group. The substituted phosphate is bound through an oxygen atom to the morpholine’s exocyclic methylene group, and through a phosphorous-nitrogen bond to the nitrogen atom of another morpholine ring. One standard DNA nucleobase (adenine, guanine, cytosine, or thymine) is bound to each morpholine ring. The ends of Morpholinos are conventionally named 3 and 5 by analogy with the nomenclature for nucleic acids (though if one were to number the atoms of a morpholino oligonucleotide backbone by IUPAC rules, the numbers assigned to the ends would be different). The secondary amine of the morpholine ring at the end of an unmodified morpholino oligonucleotide is called the 3 end of the oligo, whereas the 5 end terminates with a chiral carboxamidated phosphorodiamidate group (Fig. 4.30.1). Antisense Morpholinos block the interactions of macromolecules with mRNA by base pairing with the targeted mRNA in a complementary fashion, thus preventing initiation
Figure 4.30.1
Structure of a Morpholino 3-mer.
Contributed by Jon D. Moulton Current Protocols in Nucleic Acid Chemistry (2006) 4.30.1-4.30.24 C 2006 by John Wiley & Sons, Inc. Copyright
Synthesis of Modified Oligonucleotides and Conjugates
4.30.1 Supplement 27
complex read-through or modifying splicing in cells ranging from bacterial (Geller et al., 2005) to human (Suwanmanee et al., 2002). In particular, antisense Morpholinos have become a standard tool for developmental biologists to manipulate gene expression in embryos such as zebrafish and Xenopus sp. (Ekker and Larson, 2001). These modified oligonucleotides combine efficacy, specificity, stability, lack of non-antisense effects, and good water-solubility properties. This unit presents three protocols for the design of a knockdown experiment using Morpholinos (Basic Protocol 1), preparation of Morpholino solutions (Basic Protocol 2), and introduction of Morpholinos into cells by endocytosis in the presence of an amphiphilic peptide (Basic Protocol 3). The Commentary provides a thorough discussion of conditions and considerations for the application of Morpholinos. BASIC PROTOCOL 1
DESIGN OF A MORPHOLINO KNOCKDOWN EXPERIMENT This protocol outlines the choices commonly encountered while designing a Morpholino knockdown experiment. Considerations for the steps are addressed in the Commentary. 1. Choose the target gene. 2. Choose the cells or organism into which the oligo will be delivered. 3. Choose between splice blocking or translation blocking, which determines the molecular assays available for measuring antisense activity. 4. Obtain the sequence of the target RNA. Use the mRNA 5 -UTR and the first 25 coding bases for translation blockers, or pre-mRNA with introns and exons defined for splice blockers. 5. Choose a delivery method. 6. Select control oligos. 7. Decide whether end-modification of any oligos is necessary. 8. For blocking splicing, select which pre-mRNA splice boundary (intron-exon) to block. 9. Select the oligo target (following the targeting rules described in the Commentary) and determine the Morpholino sequence (the inverse complement of the target). 10. Use a transcript database and a homology search tool such as BLAST to test the selected target for homologies with other RNAs. If the selected target is too homologous with a region of an off-target mRNA, a partially complementary Morpholino might affect the expression of that mRNA. Another target on the desired mRNA must be selected to prevent off-site Morpholino interaction.
11. Order the synthesis of the selected Morpholinos. BASIC PROTOCOL 2
PREPARATION AND VERIFICATION OF MORPHOLINO STOCK SOLUTIONS This protocol describes the preparation of stock aqueous solutions of Morpholinos at concentrations of 1 mM or 500 µM, if necessary.
Materials Using Morpholinos to Control Gene Expression
Lyophilized Morpholino oligonucleotide (Gene Tools) Distilled autoclaved water without DEPC, sterile 0.1 M HCl
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Glass or polypropylene/polyethylene tubes with labels 65◦ C water bath Quartz spectrophotometer cell (1 cm path length) Parafilm Lint-free lab tissues UV spectrophotometer (or UV colorimeter) capable of measurements at 265 nm Morpholino product information sheet Prepare Morpholino solution 1. Read the amount of Morpholino given on the vial label and, using sterile technique, add the appropriate volume of distilled sterile water to make a 1 mM stock solution (i.e., 0.1 mL water for a vial containing 100 nmol Morpholino). The aqueous solubility of Morpholinos is sequence-dependent, but most Morpholino sequences with G content below 36% will dissolve in water at the recommended stock concentration of 1 mM. Do not keep Morpholino solutions of <1 µM because submicromolar concentrations can lose significant activity by binding to glass and plastic surfaces. It is strongly recommended that Morpholino stock solutions be made with distilled water, but isotonic buffers (e.g., Ringer’s solution, Danieau buffer) can also be used. The use of distilled water facilitates the process of concentrating Morpholinos, should that be required. If water must be treated with diethylpyrocarbonate (DEPC), it is very important to autoclave the treated water to destroy residual DEPC before using it to dissolve Morpholinos. Otherwise, DEPC reacts with adenines and compromises the ability of Morpholinos to bind to their targets (Henderson et al., 1973).
2. Cap the vial, shake it, and wait 5 min. 3. Swirl and inspect the solution to see whether the oligo has dissolved. If it has not dissolved, warm the vial for 5 min in a water bath at 65◦ C. 4. If desired, dispense into several tubes. Label tube(s) with the concentration and oligo name, and store any tubes that will not be used immediately. Scrupulously avoid microbial contamination of the stock solutions. Store fluorescenttagged Morpholinos in a closed box so that light will not bleach fluorescent moieties. Morpholinos are stable in stock solutions stored at either 25◦ C or 4◦ C. Morpholinos can also be stored frozen. However, ice crystal formation during slow freezing can cause the concentration of the oligos in the bulk solution phase to increase until the Morpholinos precipitate. Thus, after thawing, Morpholino solutions should be heated for 10 min in a water bath at 65◦ C to ensure complete dissolution prior to use.
Check Morpholino concentration by UV absorbance 5. Turn on the UV spectrophotometer and let it warm up for a few minutes. Set the spectrophotometer to report absorbance at 265 nm. 6. Clean the quartz spectrophotometer cell, if needed, and rinse the inside twice with 0.1 M HCl. Carefully shake excess liquid from cell. Do not touch the outside of the quartz spectrophotometer cell on the surfaces where light will pass through, as skin oils can skew the measurements.
7. Pipet 995 µL of 0.1 M HCl into the quartz cell and place the cell in the spectrophotometer. Blank the spectrophotometer at 265 nm. Synthesis of Modified Oligonucleotides and Conjugates
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Supplement 27
8. Remove the cell from the spectrophotometer and pipet 5 µL aqueous Morpholino solution into the quartz cell. Like natural nucleic acids, the nucleobases of a Morpholino are stacked and produce a hypochromic effect. Without unstacking the bases, the use of the molar absorptivity of an individual nucleobase to calculate the concentration of the oligo would lead to an erroneously low value. Oligos with A, C, and G bases can be unstacked by dissolving the oligos in 0.1 M HCl. Under these conditions, A, C, and G bases are protonated and are out of the stacked state due to electrostatic repulsion. When the nucleobases of the oligo are unstacked, the molar absorptivity of each nucleobase can be applied to determine the concentration of the oligo.
9. Place a piece of Parafilm over the open end of the cell, placing a thumb over the Parafilm to seal the cell, and invert several times to mix. 10. Remove the Parafilm. Wipe the outside of the cell with a lint-free tissue, if needed. 11. Place the cell in the spectrophotometer and read the absorbance at 265 nm (A265 ). 12. Calculate the molar concentration (C) of the original Morpholino solution as: C = (A265 × 200)/(ε b) where 200 is the factor for dilution in HCl, ε is the molar absorptivity, and b is the path length of the cell (1 cm). The molar absorptivity (ε) of the Morpholino is provided on the product information sheet. Alternatively, ε can be calculated by multiplying the molar absorptivity of each nucleobase (A, C, G, and T) by the number of instances that the nucleobase is present in the oligo, and adding these products. This Beer’s law calculation works when absorbance ≤2, where the relationship of absorbance to concentration is linear. If the measured absorbance is >2, the sample should be diluted and remeasured. BASIC PROTOCOL 3
DELIVERY OF MORPHOLINOS INTO CELLS USING ENDO-PORTER Endo-Porter is an amphiphilic peptide. After co-endocytosis with Morpholinos, EndoPorter permeabilizes endosomal membranes, releasing the Morpholino from the endosomes to the cytosol (Summerton, 2005). Endo-Porter was optimized using a HeLa cell line. Because tolerance of other cell types toward Endo-Porter often varies, a range of Endo-Porter concentrations should be tested before beginning knockdown experiments.
Materials 1 mM Endo-Porter solution (aqueous or DMSO formulation; Gene Tools) Cell cultures in plates or flasks at 80% to 100% confluence 1 mM Morpholino stock solution (Gene Tools) 1 mM fluoresceinated dextran, 10 kDa Cell culture medium with 10% serum Fluorescence microscope Select amount of Endo-Porter for cell type 1. Prepare concentrations of 2, 4, 6, and 8 µM Endo-Porter by pipetting 2, 4, 6, and 8 µL of a 1 mM Endo-Porter solution into 1-mL aliquots of cell culture.
Using Morpholinos to Control Gene Expression
2. Add 10 µM fluorescently labeled Morpholino (10 µL of 1 mM stock per 990 µL cell culture) or 10 kDa fluoresceinated dextran (10 µL of 1 mM stock per 990 µL cell culture). 3. Allow endocytotic uptake to proceed over a period of 24 hr.
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4. Observe intracellular fluorescence using a fluorescence microscope. See discussion on assessing delivery in the Commentary section.
5. Observe cells 72 hr after delivery to determine any cellular toxicity. For subsequent Morpholino delivery to the selected cell type, use the concentration of Endo-Porter that gave the best delivery without toxicity.
Deliver Morpholinos to cells 6. Using a cell culture not previously exposed to Endo-Porter, replace spent culture medium with fresh medium (with up to 10% serum). 7. Add the Morpholino stock solution to produce the desired concentration and swirl well to mix. For functional experiments (e.g., gene knockdown, splice blocking), Morpholinos are typically effective at concentrations as low as 1 µM. However, it is recommended that a range of concentrations be tested (such as 1, 4, and 10 µM Morpholino) to define optimal conditions.
8. Add Endo-Porter to produce the optimized concentration for the cell type and immediately swirl to mix. 9. Place the plates or flasks in the incubator. Wait at least 16 hr before assessing uptake by fluorescence, and at least 24 hr before measuring knockdowns by molecular assays. The delay needed to assay a knockdown depends on the stability of any preexisting protein encoded by the targeted mRNA; a protein with a long half-life will take longer to disappear from the cells.
COMMENTARY Background Information The morpholino phosphorodiamidate backbone of a Morpholino oligonucleotide has no significant ionic charge at neutral pH, in contrast with the polyanionic phosphodiester backbone of a natural nucleic acid. This favors the interaction of Morpholinos with nucleic acids, since there is no repulsion between anionic backbones as there is in duplexes of natural nucleic acids. Dissolved in pure water, nucleic acids lose their ability to form stable Watson-Crick bonds due to anionic repulsion between strands, whereas Morpholinos will still bind to complementary sequences (Summerton, 2004). Because Morpholinos are uncharged, they have no strong electrostatic interactions with proteins. Unmodified Morpholinos have little or no affinity for bovine or human serum albumin (H.M. Moulton, unpub. observ.). In contrast, interactions of anionic phosphorothioate oligos with proteins cause multiple physiological, non-antisense effects (Lebedeva and Stein, 2001). Proteins that bind nucleic acids generally interact electrostatically with the anionic phosphates of nucleic acids, stabilizing binding. Morpholinos appear to have little or no interaction with nucleic acid–binding proteins (Hudziak et al., 1996).
Morpholinos are very stable to nucleolytic enzymes. There are no known enzymes that degrade Morpholinos. Specifically, Morpholinos have been exposed to a range of nucleases (e.g., DNase I, DNase II, Benzonase, S1 nuclease, mung bean nuclease, Bal 31 nuclease, RNase A, RNase T1, phosphodiesterase I, and phosphodiesterase II) and proteases (e.g., pronase E, proteinase K, and pig liver esterase) under conditions where lytic enzymes would degrade their substrates. In no case was degradation of the Morpholinos detected (Hudziak et al., 1996). Morpholinos were incubated in serum and in liver homogenate without degradation (Summerton and Weller, 1997). When peptide-Morpholino conjugates were extracted from cells and analyzed by MALDITOF mass spectrometry, the Morpholino oligo entity was not degraded in the cells (Nelson et al., 2005). No crystal structure or high-resolution NMR structural analysis of phosphorodiamidate Morpholinos has been published. However, the study of a morpholino phosphorodiamidate ApA dimer using circular dichroic spectroscopy showed stacking of bases in aqueous phosphate buffer (Kang et al., 1992). On the basis of molecular modeling, the bases
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Table 4.30.1 RNA Binding Affinity of Various Oligo Types Ranked by Dissociation Temperature in Physiological Isotonic Buffers
Figure 4.30.2
Using Morpholinos to Control Gene Expression
Affinity
Type of oligo
Strongest
RNA:RNA, PNA:RNA, 2 -O-methyl-RNA:RNA (all very similar)
Strong
Morpholino:RNA
Medium
DNA:RNA
Weakest
Phosphorothioate:RNA
Comparison of RNase H–dependant, RISC-dependant, and steric blocking oligos.
of Morpholinos should stack in a fashion analogous to those of natural nucleic acids, allowing strong interactions with complementary nucleic acid sequences by Watson-Crick base pairing. A 400 MHz 1 H NMR analysis of a carbamate-linked Morpholino found the morpholine ring in the chair conformation (Stirchak et al., 1989). Molecular modeling of a Morpholino with the morpholine rings in the chair conformation suggests that a Morpholino and an RNA form an A-form heteroduplex with a helical pitch similar to that of an A-form RNA-RNA duplex (J.E. Summerton, unpub. observ.). Various types of antisense oligos are ranked by their affinity for binding to single strands of sense RNA based on their dissociation temperatures in physiological salt buffers (Table 4.30.1; Stein et al., 1997). The affinity of RNA for RNA is greater than the affinity of Morpholinos for RNA. However, single strands of mRNA folded into secondary structures contain single-stranded regions, such as the loops of stem-loops, with which Morpholinos can readily hybridize. Given that double-stranded regions of most RNA secondary structures are shorter than 25 base pairs, the overall binding affinity of Morpholinos for RNA is suffi-
cient to invade and displace those short doublestranded regions (Summerton, 1999). Antisense oligos such as DNA, RNA, and phosphorothioate (S-DNA) oligos recruit RNase H to degrade their mRNA targets (Summerton, 1999). RNAi and siRNA also employ an antisense mechanism to recognize a sense mRNA through interaction with a RISC complex, which leads to enzymatic degradation of complementary mRNA and translational blocking of partially complementary mRNA (Scacheri et al., 2004). In contrast, instead of degrading mRNA, antisense Morpholinos were designed to block the translation of mRNA into protein (Summerton and Weller, 1997). Figure 4.30.2 compares steric blocking, RNase H–dependant, and RISCdependant oligos. When comparing an RNase H– dependant oligo (a methylphosphonate diester/phosphodiester chimera) with a Morpholino, a CpGNNN motif was shown to induce apoptosis and cell cycle arrest when present in the RNase H–dependent oligo but not when present in the Morpholino (Tidd et al., 2001). There have been no reports of Morpholinos inducing either interferon production or induction of NF-κB
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mediated inflammation, and Morpholinos containing CpG motifs do not stimulate immune responses (J.E. Summerton and A. Krieg, unpub. observ.), suggesting that Morpholino-RNA heteroduplexes do not stimulate Toll-like receptors. Morpholinos complementary to sequences in the 5 -UTR and the first 25 coding bases of an mRNA can halt the progression of the initiation complex toward the start codon, preventing assembly of the entire ribosome. This inhibits the translation of the mRNA sequence into a polypeptide. Morpholinos targeted downstream of the start codon are usually ineffective for blocking translation (Summerton, 1999). In addition to their application to knock down gene expression, because stericblocking oligos do not trigger degradation of RNA, Morpholinos are also widely used to block splicing of pre-mRNA. Splicing in eukaryotes is directed by snRNPs that bind to introns and mark the intron-exon boundaries. Morpholinos targeted to these snRNP-binding sites can modify splicing (Sazani et al., 2001), either preventing splicing and causing an intron inclusion (Giles et al., 1999) or redirecting splicing and causing an exon excision (Draper et al., 2001). Blocking a splice site can cause activation of a cryptic splice site, complicating interpretation of the splice modification by producing partial deletions of exons (Draper et al., 2001) or partial inclusions of introns. Morpholinos stimulate site-specific ribosome frameshifting when bound just downstream of a shift site on an mRNA, and they do so with far higher efficiency than RNA, phosphorothioate oligos, or 2 -O-methyl RNA oligos (Howard et al., 2004). Although Morpholinos are most often used to block the translation initiation complex or the snRNPs that direct splicing, there are other mRNA sequences that are attractive targets for steric blocking. Specifically, Morpholinos can block miRNA activity by binding to the miRNA and preventing it from binding its mRNA target (Kloosterman et al., 2004), or by binding to the site on the mRNA where the miRNA would otherwise bind. Along similar lines, Morpholinos targeted across the cleavage site of a hammerhead ribozyme inhibited auto-cleavage, leading to over two orders of magnitude increase in the expression of a downstream reporter gene (Yen et al., 2004). While Morpholinos have also been shown to block intronic splice silencers (Bruno et al., 2004) and exonic splice enhancers (McClorey
et al., 2006), no publications have yet explored other potential regulatory targets such as zipcode binding sites, riboswitches, or binding sites for elements of the nonsense-mediated mRNA decay pathway. Morpholinos are commonly microinjected into embryos at the single-cell or few-cell stages to block genes involved in development (Heasman et al., 2000; Nasevicius and Ekker, 2000; Nutt et al., 2001). Morpholinos are also commonly used in cell cultures (Tyson-Capper and Europe-Finner, 2006). Applications in intact adult organisms have until recently been limited by poor in vivo delivery into the cytosol of cells (Summerton, 1999; Sazani et al., 2002). However, very recent advances in conjugating Morpholinos to cell-penetrating peptides (Nelson et al., 2005) now allow effective systemic delivery into adult organisms (Alonso et al., 2005; Kinney et al., 2005; Neuman et al., 2005; Enterlein et al., 2006). Combinations of several oligonucleotide sequences can bind to several different RNA targets simultaneously if introduced together into embryos (Ekker, 2000) or cell cultures (Summerton, 2005), allowing multiple knockdowns or synergistic targeting of a single messenger. Targeting of viral RNA with Morpholinos has been reported for hepatitis C (Jubin et al., 2000; McCaffrey et al., 2003), dengue virus (Kinney et al., 2005), ebola virus (Enterlein et al., 2006; Warfield et al., 2006), SARS virus (Neuman et al., 2005), West Nile virus (Deas et al., 2005), equine arterivirus (van den Born et al., 2005), mouse hepatitis virus (Neuman et al., 2004), novirhabdovirus (Alonso et al., 2005), and vesivirus (Stein et al., 2001). In addition to translation start sites, successful targets for inhibition of viral replication include cyclization sequences (Deas et al., 2005), terminal stem loops (Deas et al., 2005), and internal ribosomal entry sites (IRES; Jubin et al., 2000). Radioisotope delivery into organisms can be pretargeted using Morpholinos (Mang’era et al., 2001). Practitioners of nuclear medicine strive to minimize radiation exposure of a patient while delivering radionuclides to target tissues for imaging or for therapeutic applications. By attaching radioisotopes to antibodies that are specific for target tissues, the antibodies can anchor isotopes on these tissues. Because the large antibody molecules diffuse slowly, the isotopes must be maintained in the plasma at high concentrations or for long durations to achieve good delivery of radioisotope-linked antibodies to their targets.
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Pretargeting with Morpholinos involves introducing an antibody-Morpholino conjugate into the bloodstream; this can be done using high concentrations or re-dosing to saturate the target without exposing the patient to radiation during this pretargeting stage. Next, a conjugate of a radioisotope (possibly chelated) with a complementary Morpholino is added to the blood. Because the Morpholino has a much smaller molecular mass than an antibody, the radionuclide-Morpholino conjugate diffuses relatively quickly and is captured at the target tissue more rapidly through MorpholinoMorpholino pairing. Unbound radionuclideMorpholino conjugate is rapidly eliminated through the kidneys. This technique allows delivery of radioisotopes to the targeted tissue while exposing the organism to lower doses of radiation away from the targeted region. In the process of developing these techniques, pharmacokinetics of Morpholino-radionuclide conjugates have been studied in vivo (Liu et al., 2002a,b; He et al., 2003). In a recent modification, signals are amplified by binding a polymer bearing many complementary Morpholinos to each Morpholino-conjugated antibody fragment, followed by delivering radioisotopelabeled Morpholino complementary to the polymer-linked Morpholinos (He et al., 2003, 2004).
Critical Parameters Choosing Morpholino sequences The parameters considered when selecting oligonucleotide target sequences include CG%, G%, self-complementarity, tetra-G
moieties, length of the oligo, and the intended temperature at which the oligo will be used. The targeting recommendations are summarized below and in Table 4.30.2. CG range. A range of 40% to 60% CG is considered ideal for 25-base Morpholinos in 37◦ C systems. Oligos with <40% CG may lack the affinity needed for effective steric blocking, while oligos with >60% CG are more likely to interact with off-target messengers through high-affinity subsequences. G content. G content affects aqueous solubility of an oligo, with higher G contents being less soluble, particularly when the oligo is dissolved in isotonic salt solutions. Oligos with G contents up to 36% should be soluble in the millimolar range in pure water or aqueous buffer. However, freeze-thaw cycles are likely to cause high-G oligos to precipitate and the oligos must be heated to redissolve (see Basic Protocol 2). Self-complementarity. Self-complementary sequences can cause either intramolecular interactions, forming stem-loops, or intermolecular interactions, forming dimeric Morpholinos. When a short sequence of one part of an oligo is complementary to another short sequence separated by an intervening sequence, stem-loops can form. If small selfcomplementary sequences are separated by zero to a few bases, formation of a stable stemloop is unlikely because a hairpin with a small loop is not energetically favored. To prevent loss of oligo activity through competition between self-pairing and target binding, it is prudent to limit self-complementary sequences in oligo designs to 16 contiguous hydrogen bonds
Table 4.30.2 Summary of Targeting Recommendations for 37◦ C Systems
Using Morpholinos to Control Gene Expression
Parameter
Recommendation
Comments
CG range
40%-60%
At lower GC, affinity may be too low to block processes; higher GC favors nonspecific binding of subsequences.
G content
Up to 36% G
Higher G causes loss of water solubility; avoid upper end of acceptable range, if possible.
Selfcomplementarity
16 contiguous H-bonds maximum
For intermolecular (complementary palindrome) and intramolecular (stem loop) binding. Example: AGCGCT has 16 H-bonds (2+3+3+3+3+2 = 16). Check for non-Watson-Crick G-T pairing, which can participate in self-complementarities.
Consecutive G
3 consecutive Gs maximum
Runs of ≥4 G can associate through Hoogsteen bonding to form oligo tetramers.
Oligo length
25 bases or shorter by only a few bases
Using shorter oligos can decrease the chance of off-target interaction for high CG oligos.
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or less, where CG pairs contribute 3 hydrogen bonds and AT pairs contribute 2 hydrogen bonds. For instance, the short sequences ATGGC and GCCAT can form 13 contiguous hydrogen bonds (2+2+3+3+3 = 13). When analyzing sequences for self-complementarity, check for both Watson-Crick base-pairing and for GT base-pairing. Like an AT pair, a GT pair also forms two hydrogen bonds. However, because the overall stability of the GT pair is far lower than an AT pair, a GT pair can be scored as a single hydrogen bond when calculating its contribution to the stability of a self-complementary moiety (Aboul-ela et al., 1985). An oligo containing a self-complimentary sequence can form dimers. To prevent loss of oligo activity through competition between dimer formation and target binding, it is prudent to limit complimentary palindromes to 16 contiguous hydrogen bonds or less. For instance, if two oligos bearing the self-complimentary sequence ATGCATGCGT encounter each other, they can form 22 contiguous hydrogen bonds (2+1+3+3+2+2+3+3+1+2 = 22, taking into account the GT pairs) and would likely have poor antisense activity. G tetrads. Nucleic acids containing GGGG moieties can interact through Hoogsteen bonding to form oligo tetramers (Cheong and Moore, 1992). Morpholinos containing G tetrads have reduced activity, likely through the same mechanism. Because of this, contiguous stretches of four or more G bases should be avoided when designing Morpholinos. MIL and oligo length. The minimum inhibitory length (MIL) of an antisense oligo is the length needed to achieve 50% reduction in translation of a targeted gene at a concentration typically achieved in cells. The MIL of Morpholinos varies somewhat between targets, but averages about 14 bases for 37◦ C cell cultures (Summerton, 1999). To ensure good affinity between Morpholinos and their RNA targets, the oligos are usually synthesized as 25-mers. CG content can influence the MIL of an oligo, with a higher CG oligo having a shorter MIL. Oligos with high CG content might interact with off-target RNA; these oligos can be shortened by a few bases to lessen the likelihood of off-target interactions. The marginal loss of affinity resulting from shortening a high-CG oligo will not ruin activity but will slightly improve specificity. A more effective way to improve specificity is to choose a target with a lower CG content.
Temperature and oligo selection The targeting guidelines were developed for oligos to be used at 37◦ C. Many embryos are grown at lower temperatures. When temperatures are decreased appreciably, stability of base-pairing increases. The ideal CG content for oligos designed for use at lower temperatures is lower than the 40% to 60% CG recommended for 37◦ C systems. The ideal CG content for colder systems (e.g., fish and frogs) must be determined experimentally. Similarly, the allowable number of base pairs in self-complementary sequences should be reduced for colder systems. Solubility is also decreased at lower temperatures, so it is prudent to select oligos with lower G contents for use in colder systems. Targetable region for translation blockers To block translation, a 25-mer Morpholino can target anywhere between the 5 cap to 25 nucleotides into the coding sequence. The target can extend downstream into the coding sequence as long as the translational start codon is covered. In the first steps of translation, the initiation complex forms at the 5 cap and then scans through the UTR to the start codon (Fig. 4.30.3A). At the start codon, the large ribosomal subunit binds, the initiation factors dissociate, and translation proceeds through the coding region. If a Morpholino gets in the way of the initiation complex before the initiation complex reaches the start codon, it prevents assembly of the ribosome and translation of the mRNA. Nonetheless, it is preferable to target the start codon instead of upstream for two reasons. First, the quality of sequence deposited in public databases is often poor in the UTR, especially for older sequence records. Second, though rare in vertebrate genomes, internal ribosome entry sites (IRES) do exist and can allow a ribosome to enter and assemble downstream of a Morpholino bound in the 5 -UTR. Targetable region for splice blockers To block splicing, Morpholinos are targeted to pre-mRNA across or near the boundaries between exons and introns. A pre-mRNA that undergoes splicing has two flanking exons (the first and last exon) and an arbitrary number of internal exons. The first exon has a single splice site, a splice donor, where it contacts intron 1. The internal exons have two junctions each, a splice acceptor at the upstream end and a splice donor at the downstream end. The last exon has only a splice acceptor at its upstream end. Targeting the splice sites of the internal exons usually causes exon excision, resulting in an mRNA missing the exon with
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Figure 4.30.3
Using Morpholinos to Control Gene Expression
Targetable regions for translation blocking (A) and splice blocking (B).
the blocked splice site (Fig. 4.30.3B). Targeting splice sites of the flanking (first or last) exons usually causes intron inclusion, resulting in an mRNA containing the first or last intron. Sometimes blocking a splice site activates a cryptic splice site, resulting in an mRNA with an unexpected mass. The snRNPs that direct splicing bind at the intronic sides of the splice junctions, so Morpholinos are chosen that are complementary to more intronic sequence than exonic sequence. Morpholinos can have good activity if targeted entirely to intronic sequence near the splice junction, but activity decreases as the target is moved farther into the intron (P.A. Morcos, unpub. observ.). Splicing can also be modified by preventing excision of an arbitrary intron by blocking the nucleophilic adenosine that closes the splicing lariat (P.A. Morcos, unpub. observ.) or by targeting splice-regulatory sequences. It is often the goal of a splice-blocking experiment to eliminate activity of a protein. If the active site of the protein is known, a straightforward strategy is to target a Morpholino to the exon encoding the active site, causing the loss of that exon and of the active site. When the active site is not known, other useful strategies are available. One is to eliminate an upstream exon that has a number of nucleotides not evenly divisible by three, causing downstream translation to be frameshifted. Another is to trigger inclusion of the first intron, especially useful if it contains an in-frame stop codon or if its number of nucleotides is not evenly divisible by three. Sometimes causing a random exon exclusion or intron inclusion is
sufficient to eliminate activity of a protein, perhaps due to a resulting change in the protein’s tertiary structure. Quality of sequence Since a few mismatches can seriously decrease the activity of a Morpholino, the quality of the target sequence is an important consideration when designing Morpholinos. There are sometimes errors in sequence database files. Variations in sequence between strains of an organism can also present a problem. The most definitive way to ensure the correct target sequence is to sequence the targeted gene in the strain that will be used in the experiments. Mismatched unintentional targets and Morpholinos When a 25-base Morpholino is used near its lowest effective concentration, its effects are very specific. Under such conditions the oligo can also interact with sequences containing one or two mismatches when compared to the oligo’s perfectly complementary target, though even a single mismatch can decrease activity (Khokha et al., 2002). However, few to no such sequences are expected to occur randomly in a base pool the size of the Morpholino-targetable sites in the human transcriptome (Summerton, 1999). Effect of concentration on specificity When the concentration of any antisense oligo is increased well above its minimum effective concentration, it can interact with targets containing more mismatches; at some concentration a Morpholino will begin knocking down expression of off-target mRNAs.
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Therefore it is important that the oligo concentration be kept as low as practicable while still eliciting the desired targeted knockdown. The concentration at which off-target effects occur, the concentration at which targeted knockdown occurs, and the ratio of these concentrations are all sequence-specific and therefore unknown for each new oligo sequence. In most cases, an effective and specific concentration window exists such that, for complementary mRNA and off-target mispaired mRNA at similar concentrations, the onset of the targeted knockdown will occur at a lower concentration than the onset of the offtarget knockdown. However, knocking down high-copy-number mRNAs requires higher oligo concentrations, increasing the probability of knocking down low-copy-number offtarget mRNAs; such a situation can narrow or even close the effective and specific concentration window. Acceptable off-target homology A single mismatch in a Morpholino 25-mer may cause a significant decrease in antisense activity (Khokha et al., 2002), though many single-mismatched oligos have retained good activity. When used near the concentration at which a perfectly complementary oligo elicits a knockdown, five mismatches distributed throughout a 25-mer usually decreases activity of the mismatched oligo to near undetectable levels (S.T. Knuth, unpub. observ.). It is prudent to check the target sequence of a proposed oligo against a nucleotide sequence database in order to identify regions where the Morpholino might bind to off-target mRNA. When searching for homologous targets, keep in mind that 25-base Morpholinos will only block translation when targeted to the 5 -UTR and first 25 bases of coding sequence. Morpholinos can modify splicing if targeted in introns near intron-exon boundaries. If the Morpholino has homology to an off-target mRNA outside of these limited regions, binding of the oligo to the mRNA is not likely to affect expression of the off-target mRNA (though blocking miRNA targets or regulatory sequences such as exonic splice enhancers may affect expression). When comparing a 25-base Morpholino against an off-target sequence in a region where a Morpholino might have a biological effect, the fraction of homologous bases should always be below 80%. However, that percentage ignores important considerations about the distribution of the mismatches throughout the oligo. About 14 contiguous
bases of homology is the minimum inactivating length for a Morpholino (Summerton, 1999). However, if 10 bases of perfect homology are flanked with a mismatch at either side and some runs of homologous bases are just beyond the flanking single mismatches, the oligo may still bind sufficiently to block translation or splicing. High CG content can make shorter homologous sequences active, since CG pairs are more stable than AT pairs. Distributing five mismatches throughout a 25mer almost always results in loss of knockdown at low concentrations, so 5-mispair oligos are commonly used as specificity controls. If all five mismatches are at one end of the oligo, there are still 20 contiguous complementary bases in a 25-mer, and those 20 bases would retain considerable antisense activity. When checking a Morpholino target against a sequence database and finding a partially homologous region, following a rule of thumb like “<80% homology won’t cause off-target knockdown” can lead to trouble; it is important to consider the distribution of the mismatches. Additional factors to consider when analyzing partially homologous targets are that losing a CG pair due to a mismatch impacts the oligo activity more than losing an AT pair (three Hbonds compared to two), and that mismatches sometimes form GT pairs, which contribute about half the stability of an AT pair (Aboulela et al., 1985). Delivery of Morpholinos to the cytosol/nuclear compartment of cells Unmodified Morpholinos. Since unmodified Morpholinos diffuse between the cytosol and the nucleus, delivery of Morpholinos to the cytosol is sufficient to ensure entry into the nucleus (Morcos, 2001). However, unmodified Morpholinos do not readily diffuse across the plasma membrane of most cell types. If unmodified Morpholinos are added to cell cultures without delivery reagents, high concentrations and long exposure times must be used to achieve minimal delivery (Sazani et al., 2001). Further demonstrating plasma membrane impermeability, when Morpholinos are microinjected into one blastomere of a Xenopus laevis embryo at the two-cell stage, daughter cells of the injected cell will contain Morpholino activity while daughter cells of the uninjected cell contain no detectable Morpholino activity (Nutt et al., 2001). There have been some reports of particular cell types in tissue explants that take up experimentally useful concentrations of unmodified Morpholinos. These cell types include epithelial cells in
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mouse embryo pancreatic explant cultures during E11 through 13 (Prasadan et al., 2002) and liver cells in mouse embryo E10 liver explants (Monga et al., 2003). Using an engineered mouse with a stably integrated green fluorescent protein (GFP) up-regulation splicecorrection reporter system (see Up-regulation system, below), Sazani showed that there is scant uptake of unmodified Morpholinos into most tissues from the blood of adult mice (Sazani et al., 2002). Scrape loading. Scrape loading of Morpholinos into adherent cell cultures was an early method for introducing Morpholinos into cultured cells (Partridge et al., 1996). When adherent cells are gently lifted from the bottom of a well using a soft rubber scraper, the cells become transiently permeable, allowing Morpholinos to diffuse into the cytosol from the medium. This technique will not deliver oligos to all cells in a culture, and reproducibility depends on the technique of the experimenter. This method has fallen out of favor as more reproducible techniques producing more homogeneous delivery have been developed. Microinjection. Microinjection of Morpholinos into early embryos is a widely used technique for knocking down gene expression. Microinjection introduces Morpholinos directly into the cytosol. As the cytoplasm is apportioned into daughter cells at cell division, both daughter cells will contain Morpholinos. Some embryos, such as Xenopus sp., have strong permeability barriers that prevent appreciable leakage to the daughters of uninjected cells (Nutt et al., 2001). Other embryos such as the zebrafish, Danio rerio, allow diffusion of Morpholinos between cells through the first few cell divisions (for a good model of Morpholino diffusion in zebrafish embryos, see Kimmel and Law, 1985a,b). Electroporation. Electroporation has become a standard method for delivery of Morpholinos into chick embryos (Kos et al., 2003), especially for studies of neural tube development (Tucker, 2004). Electroporation has also been used to deliver Morpholinos into other embryos including mice (Mellitzer et al., 2002), into brains of developing rats (Takahashi et al., 2002), into zebrafish (Cerda et al., 2006), into clipped fins of zebrafish (Thummel et al., 2005), and into cell cultures (Jubin, 2005). Uncharged Morpholinos can be electroporated; the electroporation procedure makes cells transiently permeable so that Morpholinos can diffuse across the plasma membrane.
Endo-Porter. Endo-Porter is a reagent developed to deliver Morpholino oligos conveniently and reproducibly to the cytosol of cultured cells through an endocytotic pathway. Endo-Porter is an amphiphilic peptide that becomes cationic at low pH. In culture medium, Endo-Porter is uncharged but sticks to the surface of cells. Upon endocytosis, Endo-Porter is protonated in the acidic endosome and permeabilizes the endosomal membrane, releasing the endosomal contents into the cytoplasm. Morpholinos co-endocytosed with membraneassociated Endo-Porter are released into the cytoplasm when the endosome is permeabilized (Summerton, 2005). Endo-Porter allows simultaneous delivery of multiple Morpholinos. The concentration of Morpholinos can be varied independently of the Endo-Porter concentration, allowing dose-response antisense studies while holding the delivery reagent concentration constant. Cells treated with a 5 µM carboxyfluoresceinated Morpholino and 8 µM Endo-Porter gave transfection efficiencies of 82% for human amnion-derived WISH cells and 78% for human myometrial cells when assayed by confocal microscopy (TysonCapper and Europe-Finner, 2006), though concentrations too low to be detected by fluorescence might still be sufficient to have measurable antisense activity. Endo-Porter has been used successfully with traditionally hard-to-transfect cells such as cardiomyocytes (Masaki et al., 2005). It works well with unmodified Morpholinos or carboxyfluoresceinated Morpholinos, but best delivery is achieved with lissaminated Morpholinos (S.T. Knuth, unpub. observ.). Endo-Porter is commercially available in neat DMSO or in a less-effective aqueous formulation for cells sensitive to DMSO. The recommended concentration of EndoPorter is 6 µM, achieved by using 6 µL of a 1 mM Endo-Porter solution per milliliter of cell culture; this concentration gives good delivery without toxicity to many cell types. However, cell types vary in their tolerance to Endo-Porter, with some cells tolerating higher exposures while other cells are harmed by a 6 µM Endo-Porter solution. When trying Endo-Porter with a new cell type, it is prudent initially to test a range of Endo-Porter concentrations (e.g., 2, 4, 6, and 8 µM) to assess delivery and to check the tolerance of the cells for the reagent. Special Delivery. Morpholinos are sometimes delivered using cationic delivery reagents, such as ethoxylated polyethylenimine (EPEI) or Lipofectamine. However,
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since Morpholinos are not charged they will not form electrostatic complexes with cationic delivery reagents. Without such complexation, the Morpholinos are poorly delivered to the cytosol of treated cell cultures. To overcome this limitation, Morpholinos can be annealed to complementary or partially complementary strands of anionic nucleic acids. Special Delivery oligos are heteroduplexes of Morpholinos and partially complementary DNA, and are delivered after complexation with EPEI (Morcos, 2001). Special Delivery oligos were designed as a replacement for scrape loading of adherent cells, but can also be used with cells in suspension. Special Delivery oligos provide a more homogeneous delivery than scrape loading, and many studies have been published using them. However, several problems are inherent in the system: (1) EPEI is somewhat toxic to cells; (2) the concentration ratio of heteroduplex to EPEI is fixed; (3) only a single oligo sequence can be delivered at an effective concentration at any one time; and (4) the complexation procedure, which must be done prior to each delivery, adds complexity and variability to the experiment. While Special Delivery oligos can be made by following a fairly simple protocol, they are no longer available commercially as paired heteroduplexes. This approach has mostly been supplanted by Endo-Porter, which is simpler to use, more versatile, more effective, and less toxic in most cell types. Peptide conjugates. Cell-penetrating peptides covalently conjugated to Morpholinos are in development to enhance cytosolic delivery of Morpholinos in cell culture (Neuman et al., 2005) and in vivo (Kinney et al., 2005; Neuman et al., 2005). Most published research describing Morpholino-peptide conjugates has used arginine-rich peptides (Moulton et al., 2004; Neuman et al., 2004; Deas et al., 2005; Kinney et al., 2005; Nelson et al., 2005; McClorey et al., 2006). Conjugation with arginine-rich peptides alters the specificity, target affinity, and toxicity of Morpholinos (Nelson et al., 2005). Due to the high density of cationic charges on the peptide moiety, Morpholinos conjugated with arginine-rich peptides associate with subcellular structures and with outer cell surfaces. This property might lead to falsepositive artifacts when assessing delivery of arginine-conjugated peptides by fluorescencebased methods, such as fluorescence microscopy, fluorometry, or flow cytometry. To determine the concentration of an internalized conjugate using fluorescence-based methods,
the membrane-associated conjugate should be removed in order to avoid overestimation. Trypsin treatment has been effective for eliminating binding of Morpholino-peptide conjugates to the outside of cells (Moulton and Moulton, 2003). Minimum effective Morpholino concentration To avoid off-target knockdowns, the lowest concentration of Morpholino producing the desired knockdown should be determined. When delivering Morpholinos to cell cultures using Endo-Porter, starting with a 10 µM Morpholino concentration for both fluorescent delivery assays and functional experiments increases the chances that the fluorescence will be visible in the cytosol and that the first functional experiment will produce measurable results. Because a Morpholino concentration of 10 µM might cause nonspecific effects due to interaction with nontarget genes, functional assays should be performed using a range of Morpholino concentrations. Determining the minimum Morpholino concentration that produces measurable results allows one, subsequently, to avoid off-target knockdowns and to conserve oligo. Effective Morpholino concentrations in culture medium for knockdown experiments are typically in the 1 to 10 µM range. Simultaneous oligo strategy Oligos can sometimes be delivered together to enhance their effects. Pairs of nonoverlapping translation-blocking Morpholinos targeting the same mRNA can be used simultaneously in order to decrease the concentration required for a knockdown (Ekker and Larson, 2001). If the paired oligos are simultaneously introduced into the same cells, they are sometimes effective at much lower concentrations than for either oligo alone. If oligos are individually toxic in zebrafish, their use in combination at concentrations below their toxicity thresholds might elicit the desired phenotype without toxicity. Efficiency of splice-blocking can be increased by blocking both donor and acceptor splice sites flanking a single exon (P.A. Morcos, unpub. observ.). Targeting several exons simultaneously is an effective way to deplete a wild-spliced mRNA (Draper et al. 2001). When designing oligos intended for co-delivery, check for complementarity between oligos that may cause them to form Morpholino heterodimers and lose activity (see Troubleshooting: Oligo activity decreases with pairs of oligos).
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Assessing oligo delivery It is best to begin a set of Morpholino experiments in a cell line by confirming and optimizing delivery. Most experimental problems involving Morpholinos in cell culture are due to insufficient delivery of oligo and can be solved by optimizing delivery to the particular type of cells used. Checking whether good cytosolic delivery can be achieved before starting to use custom-made Morpholinos is usually the most efficient use of time and resources. By fluorescence. Fluorescence can be measured by fluorescence microscopy, flow cytometry, or fluorometry. Only fluorescence microscopy can distinguish cytosolic and nuclear fluorescence (indicating successful delivery of a fluorescent Morpholino) from endosomal or surface-bound fluorescence (which does not contribute to antisense activity). A fluorescence microscope and a fluorescently labeled marker such as a Morpholino or a 10-kDa dextran are required for a reliable delivery assay. Using a 10-kDa fluoresceinated dextran or a carboxyfluoresceinated standard Morpholino control before using a more expensive, custom-made Morpholino produces reliable uptake assays at reduced cost. After delivery, live cells may be conveniently observed using an inverted epifluorescence microscope. Fixing cells can lead to false positives for delivery due to permeabilization of the plasma membrane and release of the oligo from endosomes during fixation. Using an objective with a higher numerical aperture increases the amount of light gathered from a cell and helps reveal dim fluorescence. If diffuse fluorescence is seen throughout the cytosol of the cells, the Morpholino has been delivered successfully. Bright punctate spots are likely labeled oligos trapped in endosomes. Punctate fluorescence does not indicate delivery, but it does not preclude it either. For delivery with Endo-Porter, start by assaying a range of Endo-Porter concentrations for delivery efficacy and cell tolerance (see Basic Protocol 3) or by trying a concentration of 6 µM Endo-Porter in the selected cell culture. After Endo-Porter delivery, antisense activity can be detected using as little as 1 µM Morpholino. However, although antisense activity can be achieved at Morpholino concentrations that do not produce detectable fluorescence, proof-of-delivery experiments do require detectable fluorescence. To accumulate enough fluorescence for microscopy, a concentration of about 10 µM Morpholino is needed. The Endo-Porter and labeled Morpholino should be left on the cells overnight
to allow time for endocytotic uptake and accumulation. By measuring antisense activity. If delivery is successful and a Morpholino targeting translation or splicing works as designed, a decrease in protein concentration or a shift in RT-PCR product mass (respectively) can be measured. Successful delivery might also be indicated by phenotypic effects, such as a decrease in targeted enzyme activity (Hayashi et al., 2005) or a change in morphology (Ekker, 2000). However, assaying only for a phenotypic effect becomes problematic if the expected change in phenotype does not occur; if antisense activity is not separately assessed at the level of protein concentration or mRNA mass, the experimenter will not be able to discern whether (1) the oligo failed to reach and interact with its target mRNA to produce the knockdown or splice-block, or (2) the knockdown or spliceblock was successful but did not cause the expected phenotypic change. Assaying translation blocking activity Activity of translation-blocking Morpholinos can be assayed using immunoblots. However, while Morpholinos can halt new translation, they do not cause degradation of existing protein; it therefore takes some time after Morpholino treatment before immunoblots will show evidence of a knockdown. The time required will vary with the half-life of the protein. If no antibody is available for the protein product when targeting an mRNA for translation blocking, then indirect assays such as the change in phenotype of an embryo must sometimes be used to assess the effectiveness of translation blocking. Morpholinos can phenocopy (mimic the phenotype of) many known mutations that affect morphology during development; embryos with phenotypes modified by Morpholino treatment are known as morphants (Ekker, 2000). In some cases, the enzymatic activity of a target protein can be assayed (Hayashi et al., 2005). An enzyme activity assay may serve as an assay for Morpholino activity, though there must be a delay between application of the Morpholino and the enzyme activity assay to allow for degradation of preexisting protein (see also the discussion of complementation in Troubleshooting). The effect of a Morpholino on target RNA stability varies with the sequence. Target mRNA concentrations in Morpholino-treated cells may be decreased, unchanged, or increased relative to untreated cells. Changes
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in mRNA concentrations may be due to changes in the secondary structure of the mRNA on binding a Morpholino, thereby altering the availability of the mRNA for nucleolytic degradation. Consequently, mRNA assays such as Northern blots or RT-PCR are not suitable for assaying the activity of a translation-blocking Morpholino. Assaying splice blocking activity Because blocking splicing changes the mass of the mRNA produced, RT-PCR with appropriate choice of primers is a good molecular assay for detecting the activity of spliceblocking Morpholinos. However, it is important to keep in mind that it cannot be predicted with certainty whether a splice-blocking Morpholino will cause an exon deletion (most common), an intron insertion, or activation of a cryptic splice site (which can cause a partial insertion or deletion). Because cryptic sites often redirect splicing of only a fraction of the targeted pre-mRNA population, splice blocking might produce a mixture of RT-PCR product masses (Draper et al., 2001). To detect any of these changes, it is best to use primers targeted to the two exons flanking and closest to, but not including, the Morpholino’s splice junction. Targeting a splice junction on an internal exon is likely to cause exon deletion. Primers should be chosen so that, if an exon deletion occurs, the RT-PCR product will be large enough to detect easily on a gel (one hundred to several hundred bases). That means for the system exon1–intron1–(splice blocker target)–exon2– intron2–exon3, the RT-PCR primers should be targeted to exon 1 and exon 3 in order to detect either intron 1 insertions (unusual) or exon 2 deletions (common). If the first (most 5 ) or last (most 3 ) splice junction in an mRNA is targeted, the usual result is an intron insertion instead of an exon deletion. However, targeting the first splice junction might activate a cryptic splice site in the first exon, resulting in deletion of the 3 end of the first exon or inclusion of a 5 fragment of the first intron. When targeting the last exon, an intron insertion is a more likely outcome. This is because consensus sequences of splice acceptors are more complex than those of splice donors, so it is less likely that the last exon will contain a near-consensus cryptic splice acceptor. When assaying the activity of a spliceblocking Morpholino at the molecular level using RT-PCR, it is important to compare the expected size of the RT-PCR product after the splice modification with the size of the
RT-PCR product produced by an untreated cell or organism. For easiest detection, spliceblocked RT-PCR products would be about half or twice the size of the native-spliced product (for exon deletion or intron insertion, respectively). A real system usually won’t allow such a tidy result, but it is necessary for the change in mass to be clearly visible on the gel (e.g., a 5% change in mass can be difficult to detect). Fidelity of replication can be a problem in RT-PCR, so it is best to design shorter RTPCR products if all else is equal. This means for the knockdown of exon 2, targeting primers in exon 1 near the junction with intron 1 and in exon 3 near the junction with intron 2 would be the best choice. However, all else is not equal; since fragments should be large enough that they are clearly visible, it is prudent to move the primers farther into the flanking exons. When possible, primers should amplify RT-PCR products with lengths of hundreds of bases to ensure full-length replication and visible bands. Splice modifications can cause downstream frameshifts or inclusion of intronic sequence in the mature messenger. Either of these results can cause a range of complicating effects, including truncation of the protein product by appearance of in-frame stop codons, translation suppression by appearance of a miRNA target site, degradation or suppression of the messenger through siRNA or miRNA activity, and nonsense-mediated mRNA decay. Splice modification may or may not cause a change detectable by an immunochemical assay such as an ELISA or immunoblot, since the conformation of the modified protein around the antigenic site may or may not be changed by a splice modification. A large insertion or deletion might result in loss of antibody binding or at least a significant shift in the band position on a western blot, but a small insertion or deletion could be difficult to detect. It is possible that inserting an intron or deleting an exon will cause the protein product to lose function, but it is far from certain. If the active site of the protein is known and the exon encoding the active site is targeted, a loss of function is likely. However, if the active site is not known, then splice-blocking might not change the protein’s activity. A protein might be made that retains the conformation of its active site even though it has an inserted or deleted polypeptide moiety at a different part of the protein. This means that looking for a phenotypic change in an embryo or assaying enzyme activity is often inadequate for
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assessing the splice-blocking activity of a Morpholino. This also means that while RT-PCR is a useful tool to confirm splice blocking activity, one should independently assay for protein function before concluding that a targeted gene is not required for a biological process, because successful splice-blocking may not alter the activity of the protein in the process. Up-regulation system Assaying antisense activity by knocking down a protein can lead to false positives, because toxicity can cause a decrease in gene expression and this can be misinterpreted as targeted gene knockdown unless careful controls are used. To address this problem and to provide an increased signal-to-noise ratio for antisense activity assays, Ryzard Kole’s group developed a set of signal up-regulation reporter systems based on splice modification. These systems use a mutation in human β-globin that creates a new splice site and causes thalessemia. The splice-mutant has a stop codon in-frame in the mRNA as well as a frameshift in the downstream coding region; blocking the mutant site splices out the stop and restores the correct reading frame. Constructs coupling this mutation to luciferase or GFP have been engineered. Of particular interest are the pLuc705 HeLa cell line (Schmajuk et al., 1999), which expresses luciferase when the mutant splice site is spliceblocked with control oligo, and the Sazani mouse (Sazani et al., 2002), which expresses GFP when splice-blocked with the appropriate oligo.
Using Morpholinos to Control Gene Expression
Controls When an oligo is used to target an mRNA, a parallel experiment should be done using a negative control oligo. Negative control oligos include the standard control oligo and an invert oligo. A 5-mispair specificity control oligo is also sometimes used as a negative control. The negative control shows that the effects observed during the antisense experiment are due to the sequence of the targeting oligo and not to the backbone chemistry of the Morpholino or the cytosolic delivery method used. Standard control oligo. A standard control Morpholino with the sequence CCTCCTACCTCAGTTACAATTTATA has been used in many organisms as a negative control sequence without triggering off-target or non-antisense effects. This negative control produced no toxic or teratogenic effects even when administered at considerably higher concentrations
than typically used for specific knockdown experiments. Any custom-sequence control oligo has some risk of interacting with off-target RNA; in contrast, the standard control has an established history of inactivity and is a reliable choice for a negative control oligo. The standard control Morpholino is designed to splice-block the mutant splice site used in the pLuc705 up-regulation reporter system. Invert control. If a negative control oligo needs to be related to the custom-made targeting oligo in terms of base composition, the invert oligo is a good choice of sequence. The invert has the same base sequence as the targeting oligo, but the sequence is reversed in the 5 -to-3 orientation (i.e., 5 -ACGGTGC would become 5 -CGTGGCA). The advantage of an invert over a scrambled sequence is that the invert sequence can be conveniently generated by a simple algorithm and will have the same CG content, G content, and selfcomplementarities as the targeting oligo. However, there is always a risk with any custommade oligo that the oligo may interact with unintended RNAs. Sense control. Sense Morpholinos have sometimes caused an increase in concentration of the mRNA targeted by an antisense oligo (P.A. Morcos, unpub. observ.). Thus, a sense sequence is not a good choice for a negative control Morpholino. 5-Mispair oligo. Off-target knockdown by any antisense molecule increases with increasing concentration. A 5-mispair oligo is used to define the effective and specific concentration window for a targeting oligo. Using the targeting oligo within its effective and specific concentration range decreases the chance of causing experimental artifacts by interaction with off-target RNA. A 5-mispair control oligo has nearly the same sequence as a targeting oligo, but has five mismatched bases distributed through the sequence. The mismatches should be distributed fairly evenly through a 25-mer oligo, starting a few bases in from each end. Ideally, the mismatches should be formed by exchanging C for G and G for C, since these mismatches disrupt the formation of three hydrogen bonds per base pair. Ideally, when a targeting oligo is used near the lowest concentration that produces a discernable effect, the 5-mispair oligo used at the same concentration will not produce the effect. However, as with any custom-sequence control oligo, there is a possibility that the 5-mispair oligo will interact with an untargeted RNA, triggering an off-target effect. If an
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oligo targets a high-copy-number transcript, requiring relatively high Morpholino concentration for knockdown, and the mispair oligo interacts with a low-copy-number transcript, the mispair oligo might cause effects even at concentrations below the concentration at which the targeting oligo becomes effective. The 5-mispair oligo can be used in an experiment that determines the effective and specific window of concentrations for a targeting oligo, which is the concentration range between the onset of measurable activity for the targeting oligo and the onset of measurable activity for its 5-mispair oligo. The definition of the effective and specific concentration range based on a 5-mispair oligo evolved through trial and error. Originally a 4-mispair specificity control was recommended, but a 4-mispair oligo sometimes measurably decreased the target protein concentration at concentrations low enough that the corresponding targeting oligo was just becoming effective, so there was not a wide enough effective and specific concentration range to be consistently useful. Many investigators use the 5-mispair oligo as a negative control, but that was not its intended purpose. Assuming that adding the mismatches does not create too much complementarity to an important off-target mRNA and trigger an off-target knockdown, the 5-mispair oligo usually behaves as a negative control when used at concentrations low enough that the targeting oligo is just becoming effective. However, the 5-mispair oligo is intended as a specificity control that shows the targeting oligo is being used in its effective and specific range. To demonstrate specificity of the targeting oligo, the targeting oligo, a 5-mispair control oligo, and a true negative control oligo (such as the standard control) are used at the same concentration in parallel treatments. If the 5-mispair and negative control oligos produce the same results, and the targeting oligo produces a different result, the experiment indicates that the targeting oligo has been used within its effective and specific concentration range. Appearance of an effect due to interaction of a target RNA with the 5-mispair control oligo suggests that, at that same concentration, the targeting oligo might also interact with offtarget RNAs. Two nonoverlapping translation blockers. Another strategy for showing that the effect of a translation-blocking Morpholino is due to the knockdown of its targeted mRNA is to use a second oligo targeted to a different and nonoverlapping site in the 5 -UTR of the tar-
geted mRNA. If the second oligo has the same effect on the cells (or organism) as the first, this supports the hypothesis that the effect observed is due to the knockdown of the targeted gene. Two splice blockers targeting one internal exon. If a Morpholino targeting a splice donor site produces the same result as a Morpholino targeting the splice acceptor of the same exon, this supports the hypothesis that the effect observed is due to the excision of the targeted exon. However, failure of the two oligos to produce the same result may be due to activation of a cryptic splice site(s) by one or both of the oligos. mRNA rescue. A very strong proof of specificity involves the use of a rescue mRNA. A rescue mRNA codes for the same protein targeted by the Morpholino knockdown, but has a modified 5 -UTR that is not targeted by the Morpholino. For this experiment, the rescue mRNA and Morpholino are delivered to the cytosol together. If the co-delivered rescue mRNA and Morpholino produce the same wild-type phenotype as untreated cells or organisms, this supports the hypothesis that the morphant phenotype elicited by the Morpholino alone is due to interaction with the targeted RNA. Unfortunately, the mRNA rescue experiment cannot work for some genes when used in embryos. The timing of the onset of translation is crucial for some developmental genes, and the early onset of translation resulting from co-injection of Morpholino and rescue mRNA in the early zygote may alter the developmental process so that these embryos never recapitulate the wild-type phenotype. Furthermore, the location of gene expression is often crucial for development, but oocyte microinjection causes rescue mRNAs to be present in all cells of the embryo until degraded or diluted by growth. End modifications Several optional modifications attached to the ends of Morpholinos are commercially available. Carboxyfluorescein, lissamine, and primary amines (Fig. 4.30.4) are the most commonly used. Optional groups are usually added to the secondary amine on the 3 -end of the oligo, and are assumed to be 3 modifications unless explicitly declared to be 5 modifications. Fluorophores and biotin are attached to Morpholinos through flexible polyethyleneglycol spacers. The length of the spacers was chosen based on antisense activity studies to ensure that the fluorophores would not interfere with binding of the Morpholinos to their
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Figure 4.30.4
Using Morpholinos to Control Gene Expression
Commercially available 3 -end modifications of Morpholino oligos.
target RNA sequences. The primary amine modification includes a short spacer of two methylenes. Carboxyfluorescein. Carboxyfluorescein is a green-emitting fluorophore that was chosen from among the fluoresceins for its good chemical stability. While its photostability is better than that of many of the fluoresceins, all of the fluoresceins are subject to photobleaching, so carboxyfluorescein should not be exposed to intense light unnecessarily. The exci-
tation wavelength of a carboxyfluoresceinated Morpholino in water is 502 nm and its emission wavelength is 525 nm. Carboxyfluorescein has two negative charges at neutral pH. Lissamine. Lissamine is a red-emitting sulforhodamine B. The excitation wavelength of a lissaminated Morpholino in water is 575 nm, and the emission wavelength is 593 nm. Lissamine is a zwitterion at neutral pH, with one positive and one negative charge. Adding a lissamine to a Morpholino increases its delivery
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efficiency with Endo-Porter, but adding lissamine to a Morpholino can decrease its aqueous solubility. It is therefore recommended to use a carboxyfluorescein tag when a fluorochrome is needed, especially for Morpholino sequences with relatively high G contents (>30% G). Primary amine. Morpholinos may be modified with a primary amine to provide a reactive site for attachment of other moieties to the oligo. An unmodified Morpholino has a secondary amine on the 3 end, the pKa of which is 6.5. The primary amine, with its pKa of 10.2, provides a more reactive site. When a primary amine is attached to the 3 end of the oligo, this converts the 3 -secondary amine of the morpholine ring to a tertiary amine as a consequence of the attachment of a short spacer tethering the new primary amine. When a primary amine is attached to the 5 end of the oligo, the 3 -secondary amine is acetylated so that a reagent added to react with the primary amine will not react with the 3 end of the oligo. When reacting a primary amine with a derivatizing reagent, it is prudent to include an additional short spacer to prevent steric hindrance between the moiety being added and the Morpholino. Morpholino stock solutions and reconcentrating Morpholinos Morpholino stock solutions in distilled water should be kept sterile and can be autoclaved. Do not use water containing diethylpyrocarbonate. Morpholino stock solutions can be dissolved in buffers such as Ringer’s solution or Danieau buffer, but this can cause problems later if the stock solution must be reconcentrated, since lyophilization can be more difficult from a buffer. Also, Morpholinos are substantially more soluble in distilled water than in isotonic salt solutions. A solution of Morpholino in water can be concentrated by using a Speedvac or by lyophilization (freeze-drying). Lyophilizing Morpholinos from water produces a fluffy solid that dissolves fairly readily if the sequence has good solubility properties. However, dissolution of Morpholinos concentrated with a Speedvac may be more difficult and will likely require patience and heating to 65◦ C. Temperature during handling Morpholinos are not degraded by nucleolytic enzymes. Solutions of DNA and RNA are normally kept on ice during experiments to prevent nucleolytic degradation, but this is not
a concern with Morpholinos. However, some Morpholino solutions have low enough solubility that icing a solution may cause a loss of activity, due to the oligo coming out of solution. Therefore, icing Morpholino solutions is not only unnecessary but can cause problems; Morpholinos should be kept at room temperature during experiments. Material affinity Morpholinos have some affinity for plastics, so passaging very dilute (submicromolar) solutions through plastic containers may cause appreciable decreases in activity. Similarly, filter sterilization may cause Morpholino solutions to lose some activity as some oligo binds to the filter. When put through the same procedures with the same exposure to plastic surfaces, high-concentration Morpholino solutions have a smaller fractional decrease in concentration than low-concentration Morpholino solutions. Therefore, if exposure to plastic surfaces is required, it is best to do the procedures with Morpholinos in a relatively concentrated state (>1 µM). Similarly, if Morpholinos are to be stored in solution for more than a few days, it is best to store them at high concentration. Since solutions of Morpholinos at very low concentrations (<1 µM) may lose activity over a time scale of minutes to hours, dilutions should be made just before use. If Morpholino solutions of less than about 100 µM are filtered, the concentration may be affected appreciably given the large surface area of filters. As the oligo bound is proportional to the surface it is exposed to, a small-diameter filter should be used to minimize oligo losses. Pall Acrodisc HT Tuffryn 0.2-µm membrane filters were found to bind less Morpholino per area than other filters tested (J.E. Summerton, unpub. observ.). The concentration of oligo in a solution can be measured spectrophotometrically just before and after performing a procedure to determine the loss of oligo caused by the procedure.
Troubleshooting Loss of antisense activity over time Morpholinos can be safely stored at temperatures ranging from room temperature to −80◦ C. Some Morpholino solutions lose activity when stored frozen, not due to degradation of the oligos but simply to aggregation. The activity can be recovered by heating the solution to 65◦ C for 10 min prior to use.
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Loss of fluorescence over time Fluorescent tags can be photobleached by exposure to bright light or prolonged exposure to dim light. Always store fluorescent materials in the dark. Wrapping aluminum foil around tubes containing fluorescent materials is an easy and prudent method for protecting fluorophores. Fluorescent materials can autoquench at high concentrations and decrease their light emission, so do not check for fluorescence at very high concentration. Labeled Morpholinos, as 10 µM solutions, are well below the concentration at which their fluorophores autoquench. No apparent activity If a Morpholino does not produce the anticipated result, there are several possibilities to consider. Has delivery been confirmed? If the oligo is not reaching the cytosol of the cells, no antisense activity will be observed. Is the activity checked by a molecular assay? If the selected activity assay determines a phenotype, such as a change in embryo morphology or in enzyme activity, the oligo may be successfully knocking down translation or modifying splicing, but a second protein may be complementing the lost activity of the target protein, thereby confounding the assay. Assaying translation blocking by immunoblot and splice blocking by RT-PCR can help determine whether the oligo is not interacting with its target or has not been delivered, or whether there is a more subtle reason for the failure to produce the expected phenotype, such as complementation by another protein. Feedback upregulation can also cause a knockdown to fail; greatly increased transcription of the targeted mRNA in response to an attempted knockdown can overwhelm the ability of the oligo to block all of the targeted messengers. Oligo activity decreases with pairs of oligos When two or more oligos are together in a cell, they may hybridize with each other if they share complementary sequences. If a pair of oligos has less activity than each individual oligo, check the sequences for complementarities. Sixteen contiguous hydrogen bonds of complementarity is the maximum recommended for oligos used together in cells or organisms at 37◦ C.
Using Morpholinos to Control Gene Expression
Clogging microinjectors If a Morpholino solution causes a microinjector to clog, one can: (1) heat the solution to disrupt tiny clumps (65◦ C for 10 min), (2) filter-sterilize the solution (although some oligo may be lost on the filter), or (3) try in-
jecting a higher volume of a less concentrated solution.
Anticipated Results Translation blockers If a translation-blocking Morpholino knocks down expression of a protein, this activity can be revealed by a delayed decrease in the protein signal on an immunoblot using an antibody to the protein. The successful knockdown should also decrease the activity of the targeted protein, though an assay for the activity of that protein can be confounded by complementation by another protein. Splice blockers If a splice-blocking Morpholino changes the mass of an mRNA, this activity can be revealed soon after delivery by a change in the mass of an RT-PCR product produced using appropriately chosen primers. A successful exon excision should also result in a delayed decrease of the activity encoded by the deleted exon as preexisting protein degrades. Spliceblocking may also decrease activities encoded on untargeted exons of a target RNA due to frameshifts, inserted stop codons, or changes in tertiary structure.
Time Considerations After delivery, wait for antisense effects to be measurable When a Morpholino is delivered into the cytosol of a cell, preexisting protein is not altered by the Morpholino. For a translation blocking oligo, this means that even if translation of a protein is immediately and completely halted on Morpholino delivery, an assay for protein concentration will not immediately reveal a successful knockdown. At least part of the existing protein must be degraded before the knockdown will be evident on an immunoblot. Similarly, although a splice-blocking oligo may cause a rapid change in the mass of an RT-PCR product, the protein produced prior to splice blocking will persist in the cell until degraded. Delivery systems using endocytotic uptake, such as Endo-Porter or Special Delivery, increase the lag between the start of delivery and the appearance of a knockdown or spliceblock signal. An overnight wait is generally sufficient to allow for endocytotic uptake. For embryonic studies, the presence of maternal transcripts in a zygote may delay the loss of protein activity when splice-blocking Morpholinos are used. Though a splice blocker
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can modify splicing of pre-mRNA transcribed in the zygote, maternal transcripts are already spliced before the onset of zygotic transcription and will be expressed in their unmodified form. Translation-blockers can block both maternal and zygotic transcripts, and can thus provide a more rapid knockdown than splice blockers. Redelivery When translation of a protein is blocked by a Morpholino, existing protein in the cell persists until broken down. After delivery, as cells grow and divide, the concentration of Morpholino oligos in their cytoplasm decreases due to dilution. Because of these two processes, when Morpholinos are used to knock down genes that code for unusually abundant or stable proteins, redelivery of the oligos may be required before a significant decrease in protein levels can be detected by immunoblotting. After an initial treatment with a Morpholino at the start of day 1, redelivery on day 4 usually suffices to produce a clear knockdown of stable and abundant proteins by day 6. However, attempts to block translation of actin have so far failed to produce a decrease in actin levels on immunoblots, suggesting that Morpholinos cannot knockdown some very abundant proteins (P.A. Morcos, unpub. observ.).
Acknowledgements Thanks to Dr. Shannon T. Knuth for editing and for information from protocols she has written, and to Conrad Shultz and Drs. James E. Summerton, Hong M. Moulton, Yongfu Li, and Paul A. Morcos for their editing and suggestions.
Literature Cited Aboul-ela, F., Koh, D., Tinoco, I. Jr., and Martin, F.H. 1985. Base-base mismatches. Thermodynamics of double helix formation for dCA3XA3G + dCT3YT3G (X, Y = A,C,G,T). Nucl. Acids Res. 13:4811-4824. Alonso, M., Stein, D.A., Thomann, E., Moulton, H.M., Leong, J.C., Iversen, P., and Mourich, D.V. 2005. Inhibition of infectious haematopoietic necrosis virus in cell cultures with peptideconjugated morpholino oligomers. J. Fish Dis. 28:399-410. Bruno, I.G., Jin, W., and Cote, G.J. 2004. Correction of aberrant FGFR1 alternative RNA splicing through targeting of intronic regulatory elements. Hum. Mol. Genet. 13:2409-2420. Cerda, G.A., Thomas, J.E., Allende, M.L., Karlstrom, R.O., and Palma, V. 2006. Electroporation of DNA, RNA, and morpholinos into zebrafish embryos. Methods 39:207-211.
Cheong, C. and Moore, P.B. 1992. Solution structure of an unusually stable RNA tetraplex containing G- and U-quartet structures. Biochemistry 31:8406-8414. Deas, T.S., Binduga-Gajewska, I., Tilgner, M., Ren, P., Stein, D.A., Moulton, H.M., Iversen, P.L., Kauffman, E.B., Kramer, L.D., and Shi, P.Y. 2005. Inhibition of flavivirus infections by antisense oligomers specifically suppressing viral translation and RNA replication. J. Virol. 79:4599-4609. Draper, B.W., Morcos, P.A., and Kimmel, C.B. 2001. Inhibition of zebrafish fgf8 pre-mRNA splicing with Morpholino oligos: A quantifiable method for gene knockdown. Genesis 30:154156. Ekker, S.C. 2000. Morphants: A new systematic vertebrate functional genomics approach. Yeast 17:302-306. Ekker, S.C. and Larson, J.D. 2001. Morphant technology in model developmental systems. Genesis 30:89-93. Enterlein, S., Warfield, K.L., Swenson, D.L., Stein, D.A., Smith, J.L., Gamble, C.S., Kroeker, A.D., Iversen, P.L., Bavari, S., and Muhlberger, E. 2006. VP35 knockdown inhibits ebola virus amplification and protects against lethal infection in mice. Antimicrob. Agents Chemother. 50:984993. Geller, B.L., Deere, J., Tilley, L., and Iversen, P.L. 2005. Antisense phosphorodiamidate morpholino oligomer inhibits viability of Escherichia coli in pure culture and in mouse peritonitis. J. Antimicrob. Chemother. 55:983988. Giles, R.V., Spiller, D.G., Clark, R.E., and Tidd, D.M. 1999. Antisense Morpholino oligonucleotide analog induces missplicing of c-myc mRNA. Antisense Nucleic Acid Drug Dev. 9:213-220. Hayashi, Y., Horibata, Y., Sakaguchi, K., Okino, N., and Ito, M. 2005. A sensitive and reproducible assay to measure the activity of glucosylceramide synthase and lactosylceramide synthase using HPLC and fluorescent substrates. Anal. Biochem. 345:181-186. He, J., Liu, G., Zhang, S., Rusckowski, M., and Hnatowich, D.J. 2003. Pharmacokinetics in mice of four oligomer-conjugated polymers for amplification targeting. Cancer Biother. Radiopharm. 18:941-947. He, J., Liu, G., Gupta, S., Zhang, Y., Rusckowski, M., and Hnatowich, D.J. 2004. Amplification targeting: A modified pretargeting approach with potential for signal amplification-proof of a concept. J. Nucl. Med. 45:1087-1095. Heasman, J., Kofron, M., and Wylie, C. 2000. Betacatenin signaling activity dissected in the early Xenopus embryo: A novel antisense approach. Dev. Biol. 222:124-134. Henderson, R.E., Kirkegaard, L.H., and Leonard, N.J. 1973. Reaction of diethylpyrocarbonate
Synthesis of Modified Oligonucleotides and Conjugates
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with nucleic acid components. Adenosinecontaining nucleotides and dinucleoside phosphates. Biochim. Biophys. Acta 294:356-364. Howard, M.T., Gesteland, R.F., and Atkins, J.F. 2004. Efficient stimulation of site-specific ribosome frameshifting by antisense oligonucleotides. RNA 10:1653-1661. Hudziak, R.M., Barofsky, E., Barofsky, D.F., Weller, D.L., Huang, S.B., and Weller, D.D. 1996. Resistance of Morpholino phosphorodiamidate oligomers to enzymatic degradation. Antisense Nucleic Acid Drug. Dev. 6:267-272. Jubin, R. 2005. Optimizing electroporation conditions for intracellular delivery of Morpholino antisense oligonucleotides directed against the hepatitis C virus internal ribosome entry site. Methods Mol. Med. 106:309-322. Jubin, R., Vantuno, N.E., Kieft, J.S., Murray, M.G., Doudna, J.A., Lau, J.Y., and Baroudy, B.M. 2000. Hepatitis C virus internal ribosome entry site (IRES) stem loop IIId contains a phylogenetically conserved GGG triplet essential for translation and IRES folding. J. Virol. 74:1043010437. Kang, H., Chou, P.J., Johnson, W.C. Jr., Weller, D., Huang, S.B., and Summerton, J.E. 1992. Stacking interactions of ApA analogues with modified backbones. Biopolymers 32:1351-1363. Khokha, M.K., Chung, C., Bustamante, E.L., Gaw, L.W., Trott, K.A., Yeh, J., Lim, N., Lin, J.C., Taverner, N., Amaya, E., Papalopulu, N., Smith, J.C., Zorn, A.M., Harland, R.M., and Grammer, T.C. 2002. Techniques and probes for the study of Xenopus tropicalis development. Dev. Dyn. 225:499-510. Kimmel, C.B. and Law, R.D. 1985a. Cell lineage of zebrafish blastomeres. II. Formation of the yolk syncytial layer. Dev. Biol. 108:86-93. Kimmel, C.B. and Law, R.D. 1985b. Cell lineage of zebrafish blastomeres. III. Clonal analyses of the blastula and gastrula stages. Dev. Biol. 108:94101. Kinney, R.M., Huang, C.Y., Rose, B.C., Kroeker, A.D., Dreher, T.W., Iversen, P.L., and Stein, D.A. 2005. Inhibition of dengue virus serotypes 1 to 4 in vero cell cultures with Morpholino oligomers. J. Virol. 79:5116-5128. Kloosterman, W.P., Wienholds, E., Ketting, R.F., and Plasterk, R.H. 2004. Substrate requirements for let-7 function in the developing zebrafish embryo. Nucl. Acids Res. 32:6284-6291. Kos, R., Tucker, R.P., Hall, R., Duong, T.D., and Erickson, C.A. 2003. Methods for introducing Morpholinos into the chicken embryo. Dev. Dyn. 226:470-477. Lebedeva, I. and Stein, C.A. 2001. Antisense oligonucleotides: Promise and reality. Annu. Rev. Pharmacol. Toxicol. 41:403-419.
Using Morpholinos to Control Gene Expression
Liu, G., He, J., Zhang, S., Liu, C., Rusckowski, M., and Hnatowich, D.J. 2002a. Cytosine residues influence kidney accumulations of 99mTclabeled Morpholino oligomers. Antisense Nucleic Acid Drug. Dev. 12:393-398.
Liu, G., Zhang, S., He, J., Liu, N., Gupta, S., Rusckowski, M., and Hnatowich, D.J. 2002b. The influence of chain length and base sequence on the pharmacokinetic behavior of 99mTcMorpholinos in mice. Q. J. Nucl. Med. 46:233243. Mang’era, K.O., Liu, G., Yi, W., Zhang, Y., Liu, N., Gupta, S., Rusckowski, M., and Hnatowich, D.J. 2001. Initial investigations of 99mTc-labeled Morpholinos for radiopharmaceutical applications. Eur. J. Nucl. Med. 28:1682-1689. Masaki, M., Izumi, M., Oshima, Y., Nakaoka, Y., Kuroda, T., Kimura, R., Sugiyama, S., Terai, K., Kitakaze, M., Yamauchi-Takihara, K., Kawase, I., and Hirota, H. 2005. Smad1 protects cardiomyocytes from ischemia-reperfusion injury. Circulation 111:2752-2759. McCaffrey, A.P., Meuse, L., Karimi, M., Contag, C.H., and Kay, M.A. 2003. A potent and specific Morpholino antisense inhibitor of hepatitis C translation in mice. Hepatology 38:503-508. McClorey, G., Moulton, H.M., Iversen, P.L., Fletcher, S., and Wilton, S.D. 2006. Antisense oligonucleotide-induced exon skipping restores dystrophin expression in vitro in a canine model of DMD. Gene Ther. 13:1373-1381. Mellitzer, G., Hallonet, M., Chen, L., and Ang, S.L. 2002. Spatial and temporal ‘knock down’ of gene expression by electroporation of doublestranded RNA and Morpholinos into early postimplantation mouse embryos. Mech. Dev. 118:57-63. Monga, S.P., Monga, H.K., Tan, X., Mule, K., Pediaditakis, P., and Michalopoulos, G.K. 2003. Beta-catenin antisense studies in embryonic liver cultures: Role in proliferation, apoptosis, and lineage specification. Gastroenterology 124:202-216. Morcos, P.A. 2001. Achieving efficient delivery of Morpholino oligos in cultured cells. Genesis 30:94-102. Moulton, H.M. and Moulton, J.D. 2003. Peptideassisted delivery of steric-blocking antisense oligomers. Curr. Opin. Mol. Ther. 5:123-132. Moulton, H.M., Nelson, M.H., Hatlevig, S.A., Reddy, M.T., and Iversen, P.L. 2004. Cellular uptake of antisense Morpholino oligomers conjugated to arginine-rich peptides. Bioconjug. Chem. 15:290-299. Nasevicius, A. and Ekker, S.C. 2000. Effective targeted gene ‘knockdown’ in zebrafish. Nat. Genet. 26:216-220. Nelson, M.H., Stein, D.A., Kroeker, A.D., Hatlevig, S.A., Iversen, P.L., and Moulton, H.M. 2005. Arginine-rich peptide conjugation to Morpholino oligomers: Effects on antisense activity and specificity. Bioconjug. Chem. 16:959-966. Neuman, B.W., Stein, D.A., Kroeker, A.D., Paulino, A.D., Moulton, H.M., Iversen, P.L., and Buchmeier, M.J. 2004. Antisense Morpholinooligomers directed against the 5 end of the genome inhibit coronavirus proliferation and growth. J. Virol. 78:5891-5899.
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Neuman, B.W., Stein, D.A., Kroeker, A.D., Churchill, M.J., Kim, A.M., Kuhn, P., Dawson, P., Moulton, H.M., Bestwick, R.K., Iversen, P.L., and Buchmeier, M.J. 2005. Inhibition, escape, and attenuated growth of severe acute respiratory syndrome coronavirus treated with antisense Morpholino oligomers. J. Virol. 79:9665-9676. Nutt, S.L., Bronchain, O.J., Hartley, K.O., and Amaya, E. 2001. Comparison of morpholino based translational inhibition during the development of Xenopus laevis and Xenopus tropicalis. Genesis 30:110-113. Partridge, M., Vincent, A., Matthews, P., Puma, J., Stein, D., and Summerton, J. 1996. A simple method for delivering Morpholino antisense oligos into the cytoplasm of cells. Antisense Nucleic Acid Drug Dev. 6:169-175. Prasadan, K., Daume, E., Preuett, B., Spilde, T., Bhatia, A., Kobayashi, H., Hembree, M., Manna, P., and Gittes, G.K. 2002. Glucagon is required for early insulin-positive differentiation in the developing mouse pancreas. Diabetes 51:3229-3236. Sazani, P., Kang, S.H., Maier, M.A., Wei, C., Dillman, J., Summerton, J., Manoharan, M., and Kole, R. 2001. Nuclear antisense effects of neutral, anionic and cationic oligonucleotide analogs. Nucl. Acids Res. 29:3965-3974. Sazani, P., Gemignani, F., Kang, S.H., Maier, M.A., Manoharan, M., Persmark, M., Bortner, D., and Kole, R. 2002. Systemically delivered antisense oligomers upregulate gene expression in mouse tissues. Nat. Biotechnol. 20:1228-1233. Scacheri, P.C., Rozenblatt-Rosen, O., Caplen, N.J., Wolfsberg, T.G., Umayam, L., Lee, J.C., Hughes, C.M., Shanmugam, K.S., Bhattacharjee, A., Meyerson, M., and Collins, F.S. 2004. Short interfering RNAs can induce unexpected and divergent changes in the levels of untargeted proteins in mammalian cells. Proc. Natl. Acad. Sci. U.S.A. 101:1892-1897. Schmajuk, G., Sierakowska, H., and Kole, R. 1999. Antisense oligonucleotides with different backbones. Modification of splicing pathways and efficacy of uptake. J. Biol. Chem. 274:2178321789. Stein, D., Foster, E., Huang, S.B., Weller, D., and Summerton, J. 1997. A specificity comparison of four antisense types: Morpholino, 2 -Omethyl RNA, DNA, and phosphorothioate DNA. Antisense Nucleic Acid Drug Dev. 7:151-157. Stein, D.A., Skilling, D.E., Iversen, P.L., and Smith, A.W. 2001. Inhibition of Vesivirus infections in mammalian tissue culture with antisense Morpholino oligomers. Antisense Nucleic Acid Drug Dev. 11:317-325. Stirchak, E.P., Summerton, J.E., and Weller, D.D. 1989. Uncharged stereoregular nucleic acid analogs: 2. Morpholino nucleoside oligomers with carbamate internucleoside linkages. Nucl. Acids Res. 17:6129-6141.
Summerton, J. 1999. Morpholino antisense oligomers: The case for an RNase Hindependent structural type. Biochim. Biophys. Acta 1489:141-158. Summerton, J. 2004. Morpholinos and PNAs compared. Lett. Pept. Sci. 10:215-236. Summerton, J. 2005. Endo-Porter: A novel reagent for safe, effective delivery of substances into cells. Ann. N.Y. Acad. Sci. 1058:1-14. Summerton, J. and Weller, D. 1997. Morpholino antisense oligomers: Design, preparation, and properties. Antisense Nucleic Acid Drug Dev. 7:187-195. Suwanmanee, T., Sierakowska, H., Fucharoen, S., and Kole, R. 2002. Repair of a splicing defect in erythroid cells from patients with betathalassemia/HbE disorder. Mol. Ther. 6:718726. Takahashi, M., Sato, K., Nomura, T., and Osumi, N. 2002. Manipulating gene expressions by electroporation in the developing brain of mammalian embryos. Differentiation 70:155-162. Thummel, R., Bai, S., Sarras, M.P. Jr, Song, P., McDermott, J., Brewer, J., Perry, M., Zhang, X., Hyde, D.R., and Godwin, A.R. 2005. Inhibition of zebrafish fin regeneration using in vivo electroporation of Morpholinos against fgfr1 and msxb. Dev Dyn. 235:336-346. Tidd, D.M., Giles, R.V., Broughton, C.M., and Clark, R.E. 2001. Expression of c-myc is not critical for cell proliferation in established human leukemia lines. BMC Mol. Biol. 2:13. Tucker, R.P. 2004. Antisense knockdown of the beta1 integrin subunit in the chicken embryo results in abnormal neural crest cell development. Int. J. Biochem. Cell Biol. 36:1135-1139. Tyson-Capper, A.J. and Europe-Finner, G.N. 2006. Novel targeting of cyclooxygenase-2 (COX-2) pre-mRNA using antisense morpholino oligonucleotides directed to the 3 acceptor and 5 donor splice sites of exon 4: Suppression of COX-2 activity in human amnion-derived WISH and myometrial cells. Mol Pharmacol. 69:796804. van den Born, E., Stein, D.A., Iversen, P.L., and Snijder, E.J. 2005. Antiviral activity of Morpholino oligomers designed to block various aspects of equine arteritis virus amplification in cell culture. J. Gen. Virol. 86:3081-3090. Warfield, K.L., Swenson, D.L., Olinger, G.G., Nichols, D.K., Pratt, W.D., Blouch, R., Stein, D.A., Aman, M.J., Iversen, P.L., and Bavari, S. 2006. Gene-specific countermeasures against ebola virus based on antisense phosphorodiamidate morpholino oligomers. PLoS Pathog. 2:e1. Yen, L., Svendsen, J., Lee, J.S., Gray, J.T., Magnier, M., Baba, T., D’Amato, R.J., and Mulligan, R.C. 2004. Exogenous control of mammalian gene expression through modulation of RNA selfcleavage. Nature 431:471-476. Synthesis of Modified Oligonucleotides and Conjugates
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Key References Draper et al., 2001. See above. First description of splice blocking in a zebrafish, including an analysis of a cryptic splice site. Nelson et al., 2005. See above. Description of peptide-Morpholino conjugates now in use for in vivo experiments. Summerton, 1999. See above. Review article presenting data determining the effective region for targeting translation blocking oligos and presenting a detailed discussion of Morpholino specificity and minimum inhibitory length. Summerton and Weller, 1997. See above. Structure and early synthetic scheme for Morpholino oligos.
Internet Resources http://www.gene-tools.com Commercial source for Morpholinos.
http://p196.ezboard.com/bmorpholinos Discussion board for Morpholino users. http://www.zfin.org Zebrafish Information Network. References related to Morpholino use in zebrafish are searchable in an annotated database. http://zfin.org/cgi-bin/webdriver?MIval=aanewmrkrselect.apg Annotated database of zebrafish Morpholino sequences by gene name. http://www.avibio.com AVI BioPharma, Inc., Morpholino therapeutics company.
Contributed by Jon D. Moulton Gene Tools, LLC Philomath, Oregon
http://pubs.gene-tools.com Morpholino publication database. As of printing, >1500 publications have reported experiments with Morpholino oligos in a broad range of systems. Citations and many abstracts are searchable here.
Using Morpholinos to Control Gene Expression
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Current Protocols in Nucleic Acid Chemistry
Solid-Phase Oligonucleotide Labeling with DOTA
UNIT 4.31
This unit contains protocols for the synthesis of a nucleosidic phosphoramidite tethered to a protected DOTA ligand and a description of its use in the preparation of oligonucleotide conjugates. Commercial 5 -O-(4,4 -dimethoxytrityl)-2 -deoxyuridine is covalently bound to N-protected 1,4,7,10-tetraazacyclododecane by an N3 -aminoalkyl linker and then converted to its phosphoramidite derivative, allowing normal oligonucleotide chain elongation. Upon completion of the oligonucleotide syntheses, the conjugates are deprotected and converted to the corresponding gadolinium(III) chelates by treatment with gadolinium(III) citrate, and purified by polyacrylamide gel electrophoresis (PAGE). The Gd-DOTA-labeled oligonucleotides can be used in applications based on magnetic resonance imaging. Synthesis of the nucleosidic phosphoramidite building block is described in Basic Protocol 1, and its use in the preparation of oligonucleotide conjugates is detailed in Basic Protocol 2. CAUTION: Perform all operations involving organic solvents and reagents in a wellventilated chemical fume hood, and wear gloves and protective glasses.
PREPARATION OF THE NUCLEOSIDIC PHOSPHORAMIDITE TETHERED TO DOTA
BASIC PROTOCOL 1
The synthesis of the DOTA-labeled phosphoramidite building block S.8 is based on the procedure described in Jaakkola et al. (2006) and is outlined in Figure 4.31.1. First, S.1 (prepared as described in Heppeler et al., 1999) is converted to the tetraester S.2 by reaction with bromomethyl acetate. Subsequent hydrogenolysis gives S.3. Synthesis of the nucleoside S.6 starts from commercially available 5 -O-(4,4 -dimethoxytrityl)2 -deoxyuridine (S.4), which is initially converted to S.5 by Mitsunobu reaction with N-trifluoroacetyl-6-aminohexan-1-ol (Hovinen and Hakala, 2001). Ammonolysis of the trifluoroacetate S.5 gives rise to S.6. Condensation of S.3 and S.6 in the presence of O(7-azabenzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HATU) and N,N-diisopropylethylamine gives the nucleoside derivative S.7. Finally, conventional phosphitylation yields the phosphoramidite S.8.
Materials 1,4,7,10-Tetraazacyclododecane-1-carboxymethylbenzyl ester (S.1; Heppeler et al., 1999) Dry acetonitrile, ≥99.5% pure, ≤0.005% H2 O (Merck) Anhydrous potassium carbonate, p.a. (Merck) Methyl bromoacetate, 99% pure (Acros Organics) Silica gel 60, 0.063 to 0.200 nm (Merck) Methanol (MeOH), ≥99.9% pure (Merck) Dichloromethane (CH2 Cl2 ), 99% pure (Lab-Scan) 10% palladium on activated carbon (Pd/C; Aldrich) Hydrogen gas Celite 521 (Aldrich) 5 -O-(4,4 -Dimethoxytrityl)-2 -deoxyuridine (S.4; Sigma) N-Trifluoroacetyl-6-aminohexan-1-ol (Sinha and Striepeke, 1991) Triphenylphosphine, 99% pure (Aldrich) Dry tetrahydrofuran (THF), ≥99.5% pure, ≤0.0075% H2 O (Merck)
Contributed by Lassi Jaakkola, Alice Ylikoski, and Jari Hovinen Current Protocols in Nucleic Acid Chemistry (2007) 4.31.1-4.31.11 C 2007 by John Wiley & Sons, Inc. Copyright
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4.31.1 Supplement 29
Figure 4.31.1 Synthesis of the DOTA-labeled nucleosidic phosphoramidite (see Basic Protocol 1). Bn, benzyl; DIAD, diisopropyl azodicarboxylate; DIPEA, N,N-diisopropylethylamine; DMF, N,N-dimethylformamide; DMTr, 4,4 -dimethoxytrityl; HATU, O-(7-azabenzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate. Modified from Jaakkola et al. (2006) with permission from the American Chemical Society.
Diisopropyl azodicarboxylate (DIAD), 95% pure (Aldrich) Diethyl ether, ≥99.7% pure (Merck) Aqueous ammonia, p.a. 28% to 30% (Merck) Sodium sulfate (Na2 SO4 ), anhydrous, ≥99.0% pure (Merck) N,N-Diisopropylethylamine (DIPEA), ≥99.5% pure (Aldrich) N,N-Dimethylformamide (DMF), dry, 99.8% pure, ≤0.01% H2 O (Lab-Scan) O-(7-Azabenzotriazol-1-yl)-N,N,N ,N -tetramethyluronium hexafluorophosphate (HATU; Applied Biosystems) Sodium hydrogen carbonate (NaHCO3 ) solutions, saturated and 5% (w/v) 2-Cyanoethyl-N,N,N ,N -tetraisopropylphosphordiamidite, 97% pure (Aldrich) 0.45 M 1H-tetrazole in acetonitrile (Applied Biosystems) Triethylamine (TEA), ≥99% pure (Merck) Glass filters (3-µm pore size) Rotary evaporator equipped with an oil pump Chromatography columns: 18 × 4 cm, 5 × 30 cm, 4 × 15 cm, and 4 × 2.5 cm TLC plate: silica-coated glass plate with fluorescent indicator (Merck silica gel 60 F254 ) Hydrogenation apparatus (Parr Instruments Company) Ultrasonic bath 254-nm UV lamp Reflux condenser Solid-Phase Oligonucleotide Labeling with DOTA
Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E)
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Current Protocols in Nucleic Acid Chemistry
Alkylate S.1 1. Dissolve 0.45 g (1.4 mmol) S.1 in 8 mL dry acetonitrile with stirring. 2. Add 0.79 g (5.7 mmol) anhydrous potassium carbonate. 3. Prepare a solution of 0.54 mL (5.7 mmol) methyl bromoacetate in 2 mL dry acetonitrile. Add this to the stirring S.1 solution over a 30-min period. 4. Stir for 2 hr at room temperature (∼25◦ C). 5. Filter off all solid material using a glass filter with a 3-µm pore size. 6. Concentrate the filtrate in vacuo on a rotary evaporator equipped with an oil pump. 7. Apply the residue onto a silica gel column (18 × 4–cm column) and elute with 1:9 (v/v) methanol/CH2 Cl2 . The typical volume for elution is 1 L and the size of the fractions is ∼20 mL.
8. Monitor fractions by TLC with 1:9 (v/v) MeOH/CH2 Cl2 . Visualize the desired product by staining the plates in an iodine chamber. The product S.2 can be seen as a dark spot with Rf = 0.41.
9. Combine the fractions containing the product and evaporate to dryness using a rotary evaporator connected to an oil pump. 10. Characterize by 1 H NMR spectroscopy and mass spectrometry. 1,4,7,10-Tetraazacyclododecane-4,7,10-tricarboxymethylmethyl ester 1-carboxymethylbenzyl ester (S.2): Yield of pale yellow oil: 0.45 g (60%). 1 H NMR (CDCl3 ): 7.35 (5H, m); 5.20 (2H, s); 3.76 (6H, s); 3.74 (3H, s); 3.49-2.35 (24 H). +ESI-TOF-MS: calcd. for C26 H40 N4 NaO8 + (M+Na)+ , 559.27; found, 559.27.
Reduce S.2 11. Dissolve 0.35g (0.64 mmol) S.2 in 50 mL methanol. 12. Add 55 mg of 10% Pd/C. 13. Hydrogenate in a hydrogenation apparatus overnight at atmospheric pressure. 14. Filter through Celite 521 and concentrate the filtrate in vacuo on a rotary evaporator. The product can be used for the next step without further purification.
15. Monitor the product by TLC in 2:8 (v/v) MeOH/CH2 Cl2 . Visualize by staining the plates in an iodine chamber. The product S.3 can be seen as a dark spot with Rf = 0.52.
16. Characterize the product by mass spectrometry. 1,4,7,10-Tetraazacyclododecane-1,4,7,10-tetraacetic acid trismethyl ester (S.3): +ESITOF-MS: calcd. for C19 H34 N4 NaO8 + (M+Na)+ , 469.23; found, 469.22.
Synthesize S.5 17. Dissolve 5.0 g (9.51 mmol) S.4, 2.61 g (12.2 mmol) N-trifluoroacetyl-6-aminohexan1-ol, and 2.98 g (11.46 mmol) triphenylphosphine in 50 mL dry THF. 18. Add 2.22 mL (11.2 mmol) DIAD to the stirring solution over a 15-min period. Continue stirring for 2.5 hr at room temperature. 19. Concentrate in vacuo using a rotary evaporator to obtain an oil. 20. Add 50 mL diethyl ether and keep the reaction mixture in an ultrasonic bath until all oil has been dissolved.
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21. Allow to stand 1 hr at room temperature. 22. Remove the triphenylphosphine oxide that forms by filtration using a glass filter with a 3-µm pore size. 23. Concentrate the filtrate to half volume using a rotary evaporator connected to an oil pump. 24. Remove any remaining triphenylphosphine oxide by repeating the filtration. 25. Concentrate the filtrate to an oil in vacuo on a rotary evaporator. 26. Purify the oily crude residue on a silica gel column (5 × 30–cm column), eluting with 1:9 (v/v) MeOH/CH2 Cl2 . The typical volume for elution is 1 to 2 L, and the size of the fractions is ∼25 mL.
27. Monitor fractions by TLC in 1:9 (v/v) MeOH/CH2 Cl2 . The product bearing the DMTr group can be visualized, in addition to normal UV detection, by heating up the fluorescent indicator plate on a hot plate. The product can be seen as an orange spot on a white background (Rf = 0.49).
28. Combine fractions containing the product and evaporate in vacuo using a rotary evaporator to afford S.4 as a white amorphous solid. 29. Characterize the compound by 1 H and 13 C NMR spectroscopy and mass spectrometry. N3 -(6-Trifluoroacetamidohex-1-yl)-5 -O-(4,4 -dimethoxytrityl)-2 -deoxyuridine (S.5): Yield: 6.2 g (91%). 1 H NMR (DMSO-d6 ): 9.36 (1H, br, NH); 7.68 (1H, d, J = 8.1 Hz, H6); 7.35 (2H, DMTr); 7.25 (7H, DMTr); 6.87 (4H, d, DMTr); 6.17 (1H, t, J = 6.2 Hz, H1 ), 5.47 (1H, d, J = 8.1 Hz, H5), 5.38 (1H, d, J = 4.7 Hz, 3 -OH), 4.31 (1H, m, H3 ), 3.90 (1H, m, H4 ), 3.5 (1H, dd, H5 ), 3.80 (2H, t, NCH2 ), 3.71 (6H, s, 2 × OMe), 3.25 (1H, m, H5 ), 3.20 (1H, dd, H5 ); 3.15 (2H, q, CH2 NH), 2.23 (2H, m, H2 , H2 ), 1.47 (4H, m); 1.25 (4H, m). 13 C NMR (DMSO-d6 ): 161.7 (C4), 158.0 (C=O), 156.5 (q, CF3 ); 150.3 (C2); 144.8 (DMTr); 138.8 (C6); 129.7, 127.8, 127.7, 126.7, 113.1 (DMTr); 100.7 (C5), 85.7 (DMTr); 85.5 (C4 ); 85.2 (C1 ); 69.8 (C3 ); 63.3 (C5 ); 55.5 (2 × OMe); 40.1 (NCH2 ); 39.7 (C2 ); 39.0 (CH2 NHCO); 28.0, 25.9, 25.8 (CH2 ). +ESI-TOF-MS: calcd. for C38 H42 F3 N3 NaO8 + (M+Na)+ , 748.28; found, 748.27.
Ammonolyze S.5 30. Suspend 1.41 g (1.95 mmol) S.5 in a mixture of 50 mL methanol and 50 mL aqueous ammonia. Heat overnight at reflux at 80◦ C with stirring. 31. Allow to cool to room temperature and then remove all volatiles in vacuo using a rotary evaporator connected to an oil pump. 32. Add 100 mL water and 100 mL CH2 Cl2 . Shake vigorously and isolate the organic phase. 33. Dry over Na2 SO4 , filter off the drying agent, and concentrate the filtrate in vacuo using a rotary evaporator connected to an oil pump. The product can be used for the next step without further purification.
34. Characterize the product by 1 H NMR spectroscopy and mass spectrometry.
Solid-Phase Oligonucleotide Labeling with DOTA
N3 -(6-Aminohex-1-yl)-5 -O-(4,4 -dimethoxytrityl)-2 -deoxyuridine (S.6): 1 H NMR (CDCl3 ): 7.75 (1H, d, J = 8.3 Hz, H6); 7.40-7.23 (9H, DMTr); 6.84 (4H, d, J = 8.9 Hz, DMTr); 6.31 (1H, t, J = 6.2 Hz, H1 ); 5.45 (1H, d, J = 8.3 Hz, H5); 4.53 (1H, m, H3 ); 4.00 (1H, m, H4 ); 3.89 (2H, m); 3.78 (6H, s, OMe); 3.49 (1H, dd, J = 3.0, 10.6 Hz, H5 ); 3.41 (1H, dd, J = 3.3, 10.6 Hz, H5 ); 2.64 (2H, t, J = 6.5 Hz); 2.42 (1H, m, H2 ); 2.24 (1H, m, H2 ); 2.19 (3H, br); 1.62 (2H, qv, J = 6.4 Hz); 1.40 (2H, qv, J = 6.7 Hz); 1.34 (4H, m). +ESI-TOF-MS: calcd. for C36 H44 N3 O7 + (M+H)+ , 630.31; found, 630.34.
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Introduce DOTA to S.6 35. Dissolve 0.26 g (0.58 mmol) S.3 and 100 µL DIPEA in 9 mL dry DMF. 36. Add 220 mg (0.58 mmol) HATU and stir the mixture for 30 min at room temperature. 37. Add 0.37 g (0.58 mmol) S.6 and stir the mixture for 4 hr at room temperature. 38. Concentrate the filtrate in vacuo using a rotary evaporator connected to an oil pump. 39. Dissolve the residue in 50 mL CH2 Cl2 and wash two times with 25 mL sat. NaHCO3 . 40. Dry over Na2 SO4 , filter off the drying agent, and concentrate the filtrate in vacuo. 41. Purify the oily crude residue on a silica gel column (4 × 15–cm column), eluting with 1:9 (v/v) MeOH/CH2 Cl2 . The typical volume for elution is 300 to 400 mL, and the size of the fractions is ∼15 mL.
42. Monitor fractions by TLC in 1:9 (v/v) MeOH/CH2 Cl2 . The product bearing the DMTr group can be visualized, in addition to normal UV detection, by heating up the fluorescent indicator plate on a hot plate. The product can be seen as an orange spot on a white background (Rf = 0.51).
43. Combine fractions containing the product and evaporate in vacuo using a rotary evaporator to afford S.6 as a white amorphous solid. 44. Characterize the compound by 1 H NMR spectroscopy and mass spectrometry. DOTA-labeled nucleoside (S.7): Yield 0.43 g (82%). 1 H NMR (CDCl3 ): 7.75 (1H, d, J = 8.3 Hz, H6); 7.40-7.22 (9H, DMTr); 6.84 (4H, d, J = 8.8 Hz); 6.47 (1H, br t, J = 4.7 Hz, NH); 6.32 (1H, t, J = 6.3 Hz, H1 ); 5.43 (1H, d, J = 8.0 Hz, H5); 4.59 (1H, m, H3 ); 4.05 (1H, m, H4 ); 3.89 (2H, m); 3.79 (6H, s, OMe); 3.74 (6H, s); 3.73 (3H, s); 3.42 (2H, d, J = 2.9 Hz, H5 and H5 ); 3.20 (8H); 2.60 (16H); 2.46 (1H, m, H2 ); 2.27 (1H, m, H2 ); 1.70 (1H, br); 1.62 (2H, m); 1.50 (2H, m); 1.35 (4H, m). +ESI-TOF-MS: calcd. for C55 H76 N7 O14 + (M+H)+ , 1058.54; found, 1058.54.
Phosphitylate S.7 45. Dry 300 mg (0.28 mmol) S.7 by coevaporating two times with 5 mL dry acetonitrile, and dissolve in 10 mL dry acetonitrile. 46. Add 127 mg (0.42 mmol) 2-cyanoethyl-N,N,N ,N -tetraisopropylphosphorodiamidite and 0.63 mL of 0.45 M 1H-tetrazole in dry acetonitrile. Stir the mixture for 30 min at room temperature. 47. Pour the reaction mixture into 25 mL of 5% NaHCO3 with vigorous stirring. 48. Extract twice with 25 mL CH2 Cl2 . 49. Dry the combined organic layers over Na2 SO4 , filter off the drying agent, and concentrate the filtrate in vacuo. 50. Purify the residue on a silica gel column (4 × 2.5–cm column), eluting with 5:95 (v/v) MeOH/CH2 Cl2 containing 1% (v/v) TEA as the eluent. Typical volume for elution is 300 to 400 mL and the size of the fractions is ∼10 mL.
51. Monitor fractions by TLC in 5:95 (v/v) MeOH/CH2 Cl2 containing 1% (v/v) TEA. The product bearing the DMTr group can be visualized, in addition to normal UV detection, by heating up the fluorescent indicator plate on a hot plate. The product can be seen as an orange spot on a white background (Rf = 0.32).
52. Combine fractions containing the phosphoramidite, evaporate and dry in vacuo using a rotary evaporator to afford S.8 as a white solid foam.
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53. Characterize the compound by 31 P NMR spectroscopy and mass spectrometry. Phosphoramidite S.8 is stable for prolonged storage (at least 1 year) at −20◦ C. DOTA-labeled phosphoramidite (S.8): Yield of solid white foam: 230 mg (85%). 31 P NMR (CDCl3 ): 149.60; 149.20. +ESI-TOF-MS: calcd. for C64 H93 N9 O15 P [M + H]+ , 1258.65; found 1258.66. BASIC PROTOCOL 2
SYNTHESIS OF OLIGONUCLEOTIDE-DOTA CONJUGATES The standard phosphoramidite method can be applied to incorporate S.8 into oligonucleotides, except that a prolonged coupling time (10 min) should be used to achieve an acceptable 98% coupling efficiency. The oligodeoxyribonucleotides can be assembled on any automated DNA/RNA synthesizer. The overall strategy is outlined in Figure 4.31.2, and the synthesis of 5 -d(XTAATGTAGCCCCTGAA)-3 , where X stands for S.7, is presented as an example. This sequence was assembled on a 0.2-µmol scale using recommended protocols and DMTr-off synthesis. After chain assembly, the oligonucleotide conjugate is treated as described previously (Hovinen and Hakala, 2001). The support-bound oligonucleotide S.9 is treated with 0.1 M sodium hydroxide for 4 hr at room temperature to ensure complete hydrolysis of the ester protecting groups and to cleave the oligonucleotide conjugate from the support. After evaporation in the presence of ammonium chloride and treatment with aqueous ammonia for the final global deprotection, the oligonucleotide conjugate is treated with gadolinium(III) citrate. The chelate is then desalted, concentrated in vacuo, and purified by urea-PAGE. The purified product S.11 is concentrated using butanol, desalted, and characterized by UV spectroscopy and ESI-TOF-MS. Familiarity with automated DNA synthesis (APPENDIX 3C) and purification of oligonucleotides by urea-PAGE (APPENDIX 3B) is required to carry out this protocol. Characterization of the oligonucleotide conjugate requires knowledge of ESI-TOF-MS techniques (UNIT 10.2).
Figure 4.31.2 Synthesis of the oligonucleotide conjugate tethered to DOTA (see Basic Protocol 2). CPG, controlled-pore glass; DMTr, 4,4 -dimethoxytrityl. Modified from Jaakkola et al. (2006) with permission from the American Chemical Society. Solid-Phase Oligonucleotide Labeling with DOTA
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Materials DOTA-labeled phosphoramidite (S.8; see Basic Protocol 1) Dry acetonitrile, ≥99.5%, ≤0.005% H2 O (Merck) Deoxyribonucleoside phosphoramidites (e.g., Proligo) 0.1 M NaOH 1 M NH4 Cl Aqueous ammonia, p.a. 28% to 30% (Merck) Gadolinium(III) citrate: 0.1 M Gd(III)Cl3 /0.2 M citric acid 20% polyacrylamide gel containing 7 M urea (APPENDIX 3B) 1 M Na2 CO3 , pH 9.8 1-Butanol, ≥99.5% pure (Merck) Sterile, nuclease-free, deionized water Automated DNA/RNA synthesizer (e.g., Applied Biosystems) ◦ 0.2-µmol DNA synthesis column (1000 A CPG; Applied Biosystems) 1-mL syringes 2-mL vials Speedvac evaporator (e.g., SPD121P, Savant) NAP5 gel filtration columns (GE Healthcare) Centrifuge (e.g., BR4i, JOUAN) Additional reagents and equipment for automated DNA synthesis (APPENDIX 3C) and urea-PAGE (APPENDIX 3B) Incorporate S.7 into oligodeoxyribonucleotide chain 1. Prepare a 0.2 M solution of S.8 in anhydrous acetonitrile. Dissolve the standard deoxyribonucleoside phosphoramidites in anhydrous acetonitrile according to the manufacturer’s instructions. 2. Start the automated solid-phase oligonucleotide synthesis according to the manufacturer’s instructions. Typically, use a 0.2-µmol synthesis column packed with a 1000 ◦ A CPG support, the recommended synthesis protocol for a 0.2-µmol scale, and the DMTr-off mode. Adjust the coupling time to 600 sec for S.8. 3. Upon completion of chain assembly (S.10), remove the synthesis column.
Cleave oligonucleotide conjugate from support and hydrolyze methyl ester protecting groups 4. Attach a 1 mL-syringe to one end of the synthesis column. Take up 1 mL of 0.1 M NaOH using another syringe and attach it to the other end of the synthesis column. Transfer the 0.1 M NaOH solution back and forth through the column every 30 min for 4 hr at ambient temperature. 5. Collect the NaOH solution containing the oligonucleotide conjugate into a clean 2-mL vial, add 100 µL of 1 M NH4 Cl, and evaporate to dryness using a Speedvac evaporator.
Ammonolyse and introduce gadolinium(III) 6. Dissolve the oligonucleotide conjugate in 1 mL of concentrated aqueous ammonia and incubate 16 hr at 55◦ C. Ammonium hydroxide is used for final global deprotection.
7. Add 15 mol equiv per ligand of gadolinium(III) citrate and keep at least overnight at ambient temperature.
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Purify oligonucleotide conjugate S.11 8. Desalt the oligonucleotide conjugate containing gadolinium(III) using a NAP5 gel filtration column and sterile deionized water according to manufacturer’s instructions. 9. Concentrate the oligonucleotide conjugate in vacuo using a Speedvac evaporator. 10. Purify the oligonucleotide conjugate using a 20% polyacrylamide gel containing 7 M urea. 11. Inspect the gel under UV illumination and collect the area of the gel that contains the gadolinium(III) oligonucleotide conjugate. The main product with highest molecular weight is collected.
12. Elute the oligonucleotide conjugate passively from the gel pieces by soaking overnight in 10 mM aqueous Na2 CO3 , pH 9.8, at ambient temperature. 13. Collect the aqueous solution containing the oligonucleotide conjugate and concentrate the solution by adding 2 vol of 1-butanol per aqueous volume. Mix well and centrifuge for 2 min at 90 × g (700 rpm), room temperature. Discard the butanol layer and repeat the butanol concentration until the volume of the aqueous phase is <500 µL. 14. Desalt the gadolinium(III) oligonucleotide conjugate using a NAP5 gel filtration column and sterile deionized water according to the manufacturer’s instructions. Store the oligonucleotide solution frozen at −20◦ C (stable at least 2 years).
Characterize oligonucleotide conjugate 15. Quantify the amount of DNA by measuring its UV absorbance at 260 nm. The OD260 is 4.04 and the total yield of S.11 is 200 µg (19% relative to the 0.2-µmol scale of oligonucleotide synthesis).
16. Characterize the purified oligonucleotide conjugate by –ESI-TOF-MS. DOTA-labeled oligonucleotide S.11: –ESI-TOF MS: m/z 566.55 [M–10H]10– , 629.96 [M–9H]9– , 708.71 [M–8H]8– , 810.22 [M–7H]7– , 945.28 [M–6H]6– , 1134.32 [M–5H]5– , 1418.64 [M–4H]4– . Reconstitution of the data gives M = 5674.05, which is in accordance with the proposed structure.
COMMENTARY Background Information
Solid-Phase Oligonucleotide Labeling with DOTA
Because of its excellent metal-chelating properties and high in vivo and in vitro stability, 1,4,7,10-tetraazacyclododecane-1,4,7,10tetraacetic acid (DOTA) is widely used as an organic ligand in magnetic resonance imaging (MRI) and positron emission tomography (PET) (Aime et al., 1999; Caravan et al., 1999; Woods et al., 2002). Gd3+ -DOTA is one of the most commonly used clinical MRI contrast agents (Anderson and Welch, 1999; Runge, 2000). Bioactive molecules labeled with DOTA, in turn, have found applications as target-specific radiopharmaceuticals (Volkert and Hoffman, 1999). In several applications, covalent conjugation of DOTA to oligonucleotides is required. Although several bifunctional DOTA deriva-
tives permitting oligonucleotide conjugation are commercially available, oligonucleotide labeling is a multi-step procedure involving synthesis of an oligonucleotide bearing a tether functionalized with a nucleophilic ω-group, and its reaction with an activated DOTA derivative. Since the labeling reaction is performed in solution in the presence of an excess of an activated label, laborious purification procedures cannot be avoided. When attachment of several label molecules is necessary, the purification and characterization of the desired oligonucleotide conjugate may be extremely difficult. These problems can be avoided by performing the labeling reaction by oligonucleotide synthesis on a solid support. In the present method, described by Jaakkola et al. (2006), the DOTA-labeled
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uridine phosphoramidite S.8 is synthesized from 1,4,7,10-tetraazacyclododecane-1-carboxymethylbenzyl ester (S.1; Heppeler et al., 1999) and N3 -(6-trifluoroacetamidohex-1-yl)5 -O-(4,4 -dimethoxytritly)-2 -deoxyuridine (S.5; Hovinen and Hakala, 2001). Subsequently, the phosphoramidite is incorporated into an oligonucleotide chain using a modified deprotection protocol, and the desired metal ion is introduced as a citrate salt. Although Basic Protocol 1 presents only the synthesis of the DOTA-uridine phosphoramidite, the method can also be used to prepare the corresponding thymidine derivative using N3 -(6-trifluoroacetamidohex-1-yl)-5 -O-(4,4 dimethoxytrityl)thymidine (Hovinen and Takalo, 2005) instead of S.5. The present method provides a number of significant advantages. (1) The phosphoramidite S.8 can be incorporated into the oligonucleotide structure using an oligonucleotide synthesizer and standard procedures. (2) The position of the label in the oligonucleotide chain is not restricted. (3) The method allows multilabeling. This feature is particularly advantageous in applications where high detection sensitivity is required. (4) Since the metal is introduced after completion of oligonucleotide chain assembly, the DOTA conjugate can be used in various applications by simply changing the metal. (5) Because oligonucleotide labeling is performed using a solid-phase synthesis strategy, the oligonucleotide conjugate is always free from unconjugated chelate. This is extremely important for in vivo applications. Compound characterization Phosphoramidite synthesis. Chemical characterization data are provided for all compounds in the appropriate protocol steps. 1 H NMR spectra were recorded in CDCl3 or DMSO-d6 on a JEOL JNM-GX spectrometer at 500 or 600 MHz. 13 C NMR spectra were recorded at 125.65 MHz. 31 P NMR spectra were recorded on a JEOL LA 400 spectrometer at 161.9 MHz. Chemical shifts (δ) are given in ppm from internal tetramethylsilane (1 H, 13 C) or external H3 PO4 (31 P), and coupling constants (J) are given in hertz. Mass spectral analyses were performed by generating positive ions using an electrospray ionization (ESI) time-of-flight (TOF) spectrometer (Applied Biosystems Mariner). Oligonucleotide synthesis. Oligonucleotide conjugates were characterized by mass spectrometry. Mass spectral analyses were per-
formed by generating negative ions using an ESI-TOF spectrometer (Applied Biosystems Mariner System 5272).
Critical Parameters and Troubleshooting Synthesis of nucleosidic phosphoramidite Although several strategies for the preparation of selectively alkylated cyclen derivatives have been published, only the method of Heppeler et al. (1999) was reproducible in the authors’ hands. For example, introduction of a single t-butyl carboxymethyl group in various solvents in the presence or absence of organic or inorganic bases was unsuccessful, and synthesis of a tris-carboxymethylated cyclen gave rise to a complex mixture of products. Even in those cases where the desired product was obtained as the major component, its purification by silica gel column chromatography was extremely difficult. Eventually, benzyl bromoacetate was identified as the alkylating agent of choice, and S.1 was prepared in 60% yield by following Heppeler’s procedure. Removal of the benzyl group of S.2 by hydrogenolysis gives the corresponding carboxylic acid derivative S.3 in high yield. It is essential to use methanol as the solvent to avoid transesterification. Furthermore, high pressures should not be used, as they may cause cleavage of the NCH2 COOR methylene groups. Since S.3 adsorbs tightly on silica gel, purification by column chromatography is not recommended. Fortunately, it is pure enough to be used for the next reaction without further purification. The nucleoside S.5 was prepared by a Mitsunobu reaction (Mitsunobu, 1981) between 5 -O-(4,4 -dimethoxytrityl)-2 -deoxyuridine and N-trifluoroacetyl-6-aminohexan-1-ol (Hovinen and Hakala, 2001). Here, diethyl azodicarboxylate (DEAD) was substituted with diisopropyl azodicarboxylate (DIAD) due to the limited availability of the former compound. Since the alkylation occurs exclusively at N3, protection of the 3 -hydroxy group is not needed, and the desired product can be obtained in good yield. However, purification of S.5 by column chromatography is problematic, since the triphenylphosphine oxide formed during the reaction tends to co-migrate with S.5 on silica gel. Accordingly, it is highly recommended to remove the major amount of triphenylphosphine oxide by precipitation from diethyl ether prior to chromatographic purification of the product.
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S.5 can easily be converted to S.6 by treatment with methanolic ammonia. Due to the low solubility of S.5 in polar solvents, an overnight reaction under reflux conditions is required. Since the reaction is quantitative and clean, chromatographic purification of S.6 is not required. Reaction of S.3 with S.6 in the presence of HATU and DIPEA gives the DOTAnucleoside conjugate S.7 in high yield. Purification is easily performed by silica gel column chromatography. The reaction does not proceed with less reactive activators such as dicyclohexylcarbodiimide (DCC), in all likelihood due to the low reactivity of the carboxylic acid function of S.3. Furthermore, it is essential to preactivate S.3 with HATU for at least 10 min to prevent S.6 from forming guanidine derivatives. If the preactivation time of S.3 is too short, the separation of S.7 from the reaction mixture is difficult. The phosphoramidite S.8 is prepared in a straightforward manner. The individual steps are based on conventional chemistry. Since most of the reactions are conducted under anhydrous conditions, careful attention to proper drying of solvents and glassware is essential. Preparation of the compounds requires experience with routine chemical laboratory techniques such as extraction, TLC (APPENDIX 3D), and silica gel column chromatography (APPENDIX 3E). Synthesis of oligonucleotide conjugate The present protocol describes the preparation of an oligonucleotide conjugate containing a DOTA unit at the 5 -terminus. A standard phosphoramidite protocol can be applied to the coupling of S.8, although a prolonged coupling time (10 min) is needed to obtain a sufficiently high coupling yield (>98%). Labels attached at the N3 position of uracil residues naturally weaken hydrogen bonds in a duplex. Thus, these labels should only be used upstream or downstream of the coding sequence. Knowledge of automated DNA synthesis (APPENDIX 3C) and isolation of the products by gel electrophoresis (UNIT 10.4, APPENDIX 3B) is needed. Characterization of the compounds demands knowledge of 1 H, 13 C, and 31 P NMR spectroscopies and electrospray ionization mass spectrometry (ESI-MS; UNIT 10.2). Solid-Phase Oligonucleotide Labeling with DOTA
Metal ion chelation It is essential to treat the protected oligonucleotide with aqueous sodium hydroxide for at least 4 hr at room temperature (∼25◦ C) prior to
conventional ammonolysis to avoid formation of carboxamides that will result from reaction of ammonia with the DOTA ester groups. Although DOTA amides do chelate various metal ions, the amide chelates are known to be less stable than the corresponding carboxylic acid– derived chelates (Paul-Roth and Raymond, 1995). Furthermore, if saponification is incomplete, ammonolysis will result in the formation of a mixture containing mono-, di-, and triamides, which are complicated to separate by gel electrophoresis. The metal must be introduced as a citrate. Citric acid forms rather stable and soluble complexes with several metal ions, which do not dissociate even under the basic conditions needed for oligonucleotide deprotection. This is extremely important with lanthanide ions, since trivalent lanthanide(III) ions are powerful catalysts in the hydrolysis of phosphate esters (Butcher and Westheimer, 1955). Furthermore, a prolonged reaction time (overnight at room temperature) and an excess of gadolinium(III) citrate are required to ensure complete chelate formation. According to gel electrophoresis, the crude oligomer bearing the tethered DOTA is completely stable to the reaction conditions required for deprotection and chelate formation. The desired oligonucleotide conjugate (>95% of the crude reaction mixture) is easily isolated from the gel and characterized by mass spectrometry. The observed molecular weight is in accordance with the proposed structure. As shown here, the phosphoramidite S.8 is suitable for site-specific incorporation of MRI contrast agents into oligonucleotides. Although only the preparation of Gd3+ chelates is presented here, the method could also be used in applications based on PET and SPECT (single photon emission tomography) using, for example, 67/68 Ga3+ and 111 In3+ as central ions, respectively. Naturally, for PET and SPECT applications, the metal should be introduced just prior to analysis. The slow kinetics of DOTA chelate formation should be taken into account when short-lived radioisotopes must be used. However, as proposed by Velikyan et al. (2004), the complexation can be accelerated using microwave radiation.
Anticipated Results Good to moderate yields of the individual steps of the total synthesis of the DOTAphosphoramidite are expected. The building block allows convenient synthesis of oligonucleotide conjugates on a solid support. The
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target compound is expected to be clearly the main product, and its isolation by gel electrophoresis is therefore fairly straightforward.
Time Considerations The synthesis of phosphoramidite S.8 starting from the cyclen S.3 and 5 -O-(4,4 dimethoxytriyl)-2 -deoxyuridine (S.4) can be accomplished in ∼10 days (including two overnight reactions). The time needed for automated oligonucleotide chain assembly differs from the standard protocol only by the extended coupling time of 600 sec per DOTA phosphoramidite unit. The deprotection and subsequent conversion of the oligonucleotide to the corresponding metal chelate increases the total time of the conjugate synthesis by 24 hr. The times needed for purification and characterization are the same as with standard methods.
Literature Cited Aime, S., Botta, M., Fasano, M., and Terreno, E. 1999. Lanthanide(III) chelates for NMR biomedical applications. Chem. Soc. Rev. 27:1929. Anderson, C.J. and Welch, M.J. 1999. Radiometallabeled agents (non-technetium) for diagnostic imaging. Chem. Rev. 99:2219-2234. Butcher, W.W. and Westheimer, F.H. 1955. The lanthanum hydroxide gel promoted hydrolysis of phosphate esters. J. Am. Chem. Soc. 77:24202424. Caravan, P., Ellison, J.J., McMurry, T.J., and Lauffer, R.B. 1999. Gadolinium(III) chelates as MRI contrast agents: Structure, dynamics, and applications. Chem. Rev. 99:2293-2352. Heppeler, A., Froilevaux, S., M¨acke, H.R., Jermann, H.E., B´eh´e, M., Powell, P., and Hennig, M. 1999. Radiometal-labeled macrocyclic chelator-derivatised somatostatin analogue with superb tumor-targeting properties and potential for receptor-mediated tumor therapy. Chem. Eur. J. 5:1974-1981.
Hovinen, J. and Takalo, H. 2005. Oligonucleotide labeling reactants and their use. US Pat. 6,949,639. Jaakkola, L., Ylikoski, A., and Hovinen, J. 2006. Simple synthesis of a building block for solid phase labeling of oligonucleotides with 1,4,7,10-tetraazacyclododecane-1,4,7,10tetraacetic acid (DOTA). Bioconjug. Chem. 17:1105-1107. Mitsunobu, O. 1981. The use of diethyl azodicarboxylate and triphenylphosphine in synthesis and transformation of natural products. Synthesis 1-28. Paul-Roth, C. and Raymond, K.N. 1995. Amide functional group contribution to the stability of gadolinium(III) complexes: DTPA derivatives. Inorg. Chem. 34:1408-1412. Runge, V.M. 2000. Safety of approved MR contrast media for intravenous injection. J. Magn. Reson. Imaging 12:205-213. Sinha, N.D. and Striepeke, S. 1991. Oligonucleotides with reporter groups attached to the 5 terminus. In Oligonucleotides and Analogues. A Practical Approach (F. Eckstein, ed.) p. 189. Oxford University Press, Oxford. Velikyan, I., Lendvai, G., V¨alil¨a, M., Roivainen, A., Yngve, U., Bergstr¨om, M., and L˚angstr¨om, B. 2004. Microwave-accelerated 68 Ga-labelling of oligonucleotides. J. Labeled Comp. Radiopharm. 47:79-89. Volkert, W.A. and Hoffman, T.J. 1999. Therapeutic radiopharmaceuticals. Chem. Rev. 99:22692292. Woods, M., Kovacs, Z., and Sherry, A.D. 2002. Targeted complexes of lanthanide(III) ions as therapeutic and diagnostic pharmaceuticals. J. Supramol. Chem. 2:1-15.
Contributed by Lassi Jaakkola, Alice Ylikoski, and Jari Hovinen PerkinElmer Life and Analytical Sciences Turku, Finland
Hovinen, J. and Hakala, H. 2001. Versatile strategy for oligonucleotide derivatization. Introduction of lanthanide(III) chelates to oligonucleotides. Org. Lett. 3:2474-2476.
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CHAPTER 5 Methods for Cross-Linking Nucleic Acids INTRODUCTION
C
haracterization of synthetic oligonucleotides shapes much of our understanding of native, higher-molecular-weight DNA and RNA molecules. Although of immense utility, oligonucleotides usually possess lower structural and thermal stability and have greater end effects than the larger nucleic acid constructs they are intended to model. Hence, the physiochemical and biological properties of oligonucleotides may not always compare favorably to those of larger nucleic acids. One of the most successful strategies to stabilize oligonucleotides is to connect the strands that comprise elements of helical structure with an interstrand cross-link. Cross-linked oligonucleotides are often very resistant to denaturation (induced thermally or by changes in pH, salt, or oligonucleotide concentration) relative to their unmodified constructs. A wide variety of novel chemistries exist to introduce cross-links into nucleic acids. In nearly all of these methods, solid-phase synthesis of oligomers site-specifically labeled with modified nucleosides bearing the appropriate reactive functional groups is used to generate cross-links in high yield. In addition to providing increased structural stability, cross-links have been used to probe the geometry and conformational dynamics of both medium- and large-sized nucleic acids and have been used to explore the topology of protein-ligand complexes. In the units presented in this chapter, along with those in forthcoming supplements, the reader will be provided with state-of-the art protocols to form cross-links within nucleic acids and nucleic acid–ligand complexes. To provide the reader with a comprehensive array of techniques, the chapter will present the chemistry to synthesize both interstrand and intrastrand cross-links, cyclic nucleic acids, and nucleic acid–ligand complexes.
UNIT 5.1 presents protocols to postsynthetically modify 2-amino-containing oligoribonucleotides with either an alkyl-phenyl disulfide or an alkyl thiol group. These groups react under mild conditions to form disulfide cross-links by thiol-disulfide interchange. When incorporated on opposite faces of a short, continuous RNA helix, these reactants, as expected, do not form a disulfide bond. In contrast, when these reactive groups are placed in proximity, disulfide cross-links form rapidly. In addition, by incorporating these groups at various positions of large RNAs through semisynthesis, the dynamics of thermal motions can be detected. Such motions are believed to be linked to biological function, and the protocols presented are among the few simple ways to assess such dynamics.
Methods to synthesize small circular oligonucleotides for use in diagnostic, therapeutic, and laboratory operations are presented in UNIT 5.2. These systems have gained considerable attention in recent years because they form unusually strong and specific complexes with RNA and DNA strands. In addition to their properties as molecular recognition agents, synthetic circular DNAs 20 to 200 nucleotides in size can also serve as catalysts for the amplified synthesis of DNA and RNA, a process termed “rolling circle synthesis.” One of the most convenient methods for generating oligonucleotides possessing either intrastrand or interstrand cross-links is through incorporation of oligo(ethylene glycol) bridges by solid-phase synthesis as described in UNIT 5.3. Many of the reagents needed are either commercially available or can be prepared in a few easy synthetic steps. Unlike many other DNA and RNA cross-links, aspects of the structural and thermodynamic impact of modifying nucleic acids with oligo(ethylene glycol) has been studied. Contributed by Gary D. Glick Current Protocols in Nucleic Acid Chemistry (2003) 5.0.1-5.0.2 Copyright © 2003 by John Wiley & Sons, Inc.
Methods for Cross-Linking Nucleic Acids
5.0.1 Supplement 13
In UNIT 5.4, a second group of methods for incorporating disulfide cross-links within RNA structure is presented. These protocols describe methods for the synthesis of alkylthiolmodified ribonucleosides, their incorporation into synthetic RNA, and the formation of intramolecular disulfide bonds in RNA by air oxidation. The disulfide bonds can be formed in quantitative yields between thiols positioned in close proximity in either RNA secondary or tertiary structure. Disulfide cross-links are useful tools to probe solution structures of RNA, to monitor dynamic motions, to stabilize folded RNAs, and to study the process of tertiary structure folding. In UNIT 5.5, site-specific cross-links are introduced into oligodeoxyribonucleotides by electrophilic substitution. A nucleophilic base (deoxythiouridine) is incorporated into an oligodeoxyribonucleotide, and an electrophilic tether is used to cross-link that base to a complementary DNA strand. A variety of different electrophilic DNA strands can be generated from the same deoxythiouridine-containing oligodeoxyribonucleotide by changing the nature of the electrophilic tether. In UNIT 5.6, the preparation of short endcapped DNA duplexes is presented. Using this approach, the 5′ terminus of one strand of a duplex is cross-linked to the complementary 3′ strand with a hydrophobic or hydrophilic linker. Several different linkers are presented along with methods for their incorporation into DNA during solid-phase synthesis. Finally, in UNIT 5.7, terminal disulfide cross-links are generated by substituting the terminal bases of oligodeoxyribonucleotides with a modified thymidine residue. The usefulness of this approach is discussed, with a demonstration that the cross-links do not modify the structure of the oligodeoxyribonucleotides, and examples of applications. Gary D. Glick
Introduction
5.0.2 Supplement 13
Current Protocols in Nucleic Acid Chemistry
Engineering Disulfide Cross-Links in RNA Using Thiol-Disulfide Interchange Chemistry
UNIT 5.1
This unit presents methods for incorporating disulfide cross-links within RNA structures using thiol-disulfide interchange chemistry (Fig. 5.1.1). The alkyl phenyl disulfide S.1 and the alkyl thiol S.2 are incorporated at specific ribose 2′ positions within RNA, and then react through thiol-disulfide interchange to form the more stable dialkyl disulfide (see Basic Protocol 4). Such disulfide cross-linking can be used to prepare a simple covalent conjugate of two RNA molecules or, in more complex systems, to exert a specific conformational constraint to a dynamic RNA molecule. Compared to the approach previously used to obtain disulfide cross-linking of RNA—oxidation of two thiols (Goodwin et al., 1996; Sigurdsson et al., 1995)—thiol-disulfide interchange has the advantages that it proceeds under mild conditions without an oxidative catalyst and can be kinetically characterized. Preparation of disulfide cross-linking precursors S.1 and S.2 begins with a site-specific incorporation of a 2′-amino-2′-deoxy residue within each of the two RNA species that are to be cross-linked, achieved through standard solid-phase RNA synthesis with a protected 2′-amino-2′-deoxy nucleotide phosphoramidite (see Basic Protocol 1). After deprotection of the synthetic RNA, the 2′ amine is modified with N-succinimidyl-3-(2pyridyldithio) propionate (S.3), affording the alkyl pyridyl disulfide S.4 (Fig. 5.1.2). Thiol-disulfide interchange of S.4 with thiophenol affords the alkyl phenyl disulfide S.1 (see Basic Protocol 2); reduction of S.4 with dithiothreitol yields the alkyl thiol S.2 (Fig. 5.1.2; see Basic Protocol 3). The RNA system to be cross-linked by thiol-disulfide interchange is limited to a two-piece system. Because both cross-linking precursors arise from the same intermediate (S.4, Fig. 5.1.2), S.1 and S.2 cannot be differentiated synthetically on the same RNA molecule.
O
O
O
O
B HS
O N H O P O O O
H
N
O P O O
+
S S
O
B
O
2
1
O
O
B O
O
O N H O P O O O
S S
N
B
H
O
O P O O
O
HS
Figure 5.1.1 Disulfide cross-linking of RNA through thiol-disulfide interchange. B, nitrogenous base.
Methods For Cross-Linking Nucleic Acids
Contributed by Scott B. Cohen and Thomas R. Cech
5.1.1
Current Protocols in Nucleic Acid Chemistry (2000) 5.1.1-5.1.10 Copyright © 2000 by John Wiley & Sons, Inc.
O
O
B
O +
NH2 O O P O O
O
O
N O
S S
O
O
N
3
B
N O H O P O O O
S S
N
4 DTT HS
O
O
B
O
N O H O P O O O
S S
B
O
N O H O P O O O
SH
2
1 S
N H
Figure 5.1.2 Preparation of the alkyl phenyl disulfide S.1 and the alkyl thiol S.2 as precursors for thiol-disulfide interchange. B, nitrogenous base.
Therefore, the cross-linking system must be designed such that two components of the RNA molecule (two complementary strands, a ribozyme-substrate complex, etc.), containing either S.1 or S.2 separately, can be associated through base pairing or other noncovalent interaction. After the full RNA molecule is associated, initiation of crosslinking and the rate at which it proceeds are controlled by manipulating the pH of the reaction mixture. The RNA component containing the alkyl thiol S.2 is present in saturating excess over the RNA component containing the alkyl phenyl disulfide S.1 to discourage spurious cross-linking resulting from the presence of unassociated S.1. NOTE: Experiments involving RNA require careful precautions to prevent contamination and RNA degradation; see APPENDIX 2A (do not use DEPC; this should be unnecessary, and is inadvisable with 2′ amine chemistry). BASIC PROTOCOL 1
PREPARATION OF RNA OLIGONUCLEOTIDES CONTAINING A SITE-SPECIFIC 2′ AMINE GROUP
RNA Cross-Linking Using Thiol-Disulfide Interchange Chemistry
Solid-phase synthesis of the two RNA oligonucleotides containing a unique 2′ amine employs the same procedures used for standard RNA phosphoramidites with the inclusion of a 2′-amino-2′-deoxy nucleotide phosphoramidite. The 2′-amino-2′-deoxy C-, U-, and G-phosphoramidites are synthesized according to the procedures of Verheyden et al. (1971), Imazawa and Eckstein (1979), and Benseler et al. (1992) and are produced commercially by Nexstar Pharmaceuticals. The following protocol describes deprotection and purification procedures for the solid support–bound product. Treatment with ammonium hydroxide to remove the exocyclic and 2′ amine protecting groups remaining from the synthesis procedure is followed by removal of the 2′-O-silyl protecting groups with fluoride to liberate the 2′ hydroxyl. One RNA will then be radiolabeled and modified with
5.1.2 Current Protocols in Nucleic Acid Chemistry
an alkyl phenyl disulfide group (S.1) as described in Basic Protocol 2. The other RNA will be modified to contain an alkyl thiol group (S.2; Basic Protocol 3). For a general overview of oligonucleotide synthesis, see APPENDIX 3C. Materials Solid support–bound product of automated RNA synthesis (1-µmol synthesis scale) 3:1 (v/v) concentrated ammonium hydroxide (NH4OH)/absolute ethanol 24:46:30 (v/v/v) triethylamine/1-methyl-2-pyrrolidinone/triethylamine trihydrofluoride (see recipe) TE buffer, pH 7.5 (APPENDIX 2A) NAP-25 Sephadex column (Amersham Pharmacia Biotech) 1 M sodium chloride Absolute ethanol TBE buffer (APPENDIX 2A) 80% formamide/TBE solution (see recipe) Denaturing polyacrylamide gel: 20% polyacrylamide/8 M urea in TBE buffer, dimensions 20 cm long × 26 cm wide × 0.3 cm thick (see APPENDIX 3B, CPMB UNIT 7.6 or Sambrook et al., 1989) TEN buffer (see recipe) 4-mL screw-cap vial Teflon tape Water baths, 55° and 65°C Rotary drying system for microcentrifuge tubes (Savant) 40-mL Oak Ridge centrifuge tube Preparative centrifuge (Sorvall or Beckman) UV lamp (hand held) 50-mL polypropylene centrifuge tube 0.45-µm cellulose acetate filter Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis and UV shadowing (APPENDIX 3B, CPMB UNIT 7.6 or Sambrook et al., 1989) Remove the RNA protecting groups 1. Transfer the solid support–bound RNA product into a 4-mL screw-cap vial. 2. Add 1.5 mL of 3:1 (v/v) NH4OH/ethanol. Seal the vial with Teflon tape and incubate 8 hr at 55°C. 3. Cool the vial to –20°C and decant the supernatant into a 1.5-mL microcentrifuge tube. Concentrate the supernatant under vacuum with a rotary drying system. 4. Dissolve the dry residue in 400 µL of 24:46:30 (v/v/v) triethylamine/1-methyl-2pyrrolidinone/triethylamine trihydrofluoride. Incubate 1.5 hr at 65°C. Heating at 65°C (as suggested by Wincott et al., 1995) will help dissolve the dry residue. For RNAs shorter than 10 nt, reduce the amount of fluoride solution to 100 mL and skip step 5.
Purify the crude RNA 5. Transfer the solution to 5 mL TE buffer, pH 7.5, and mix thoroughly. 6. Load 2.5 mL of the resulting solution onto each of two NAP-25 Sephadex columns. Elute the RNA from each column with 3.5 mL water, collecting the first 3.5 mL of eluate from each (7 mL total product solution).
Methods For Cross-Linking Nucleic Acids
5.1.3 Current Protocols in Nucleic Acid Chemistry
7. Add 1 mL of 1 M sodium chloride to the product solution and transfer to a 40-mL Oak Ridge centrifuge tube. 8. Precipitate the RNA by adding 24 mL absolute ethanol and then centrifuging 30 min at 10,000 × g, 2°C. Decant the supernatant and allow the RNA pellet to air dry. 9. Dissolve the RNA pellet in 150 µL TE buffer and 150 µL of 80% formamide/TBE (300 µL total). 10. Purify the RNA to single-nucleotide resolution by denaturing polyacrylamide gel electrophoresis at 25 W until the full-length product has migrated about two-thirds of the way down the gel, as indicated by the dye markers (see APPENDIX 3B, CPMB UNIT 7.6 or Sambrook et al., 1989, for standard procedures). A gel 20 cm long × 26 cm wide × 0.3 cm thick is sufficient for three separate RNA oligonucleotide preparations.
Isolate the pure RNA 11. Identify the full-length RNA band by UV shadowing and excise the gel slice. 12. Place the gel slice in a 50-mL polypropylene centrifuge tube, crush thoroughly, and add 15 mL TEN buffer. Place the suspension on a shaker for 24 hr at 2°C. 13. Pellet the polyacrylamide by centrifugation for 10 min at 2000 × g, 2°C. 14. Filter the supernatant through a 0.45-µm cellulose acetate filter. 15. Precipitate the RNA by adding 3 vol absolute ethanol and then centrifuging 30 min at 10,000 × g, 2°C. 16. Dissolve the RNA in 100 µL TE buffer, pH 7.5. Determine RNA concentration spectrophotometrically by measuring A260, and adjust to 100 µM. Store up to 6 months at −20°C or indefinitely at –80°C. BASIC PROTOCOL 2
PREPARATION OF 32P-LABELED RNA CONTAINING AN ALKYL PHENYL DISULFIDE GROUP The RNA component of the cross-linking system containing the alkyl phenyl disulfide S.1 is prepared as a 32P-labeled reagent to allow monitoring of the cross-linking reaction, indicated by a shift to a product that migrates more slowly under denaturing electrophoresis conditions. Labeling of the RNA at the 5′ end with 32P is followed by chemical modification of the 2′ amine with N-succinimidyl-3-(2-pyridyldithio) propionate (S.3), yielding the corresponding amide S.4. Thiol-disulfide interchange of the pyridyl disulfide moiety of S.4 with thiophenol yields the alkyl phenyl disulfide S.1 (Fig. 5.1.2).
RNA Cross-Linking Using Thiol-Disulfide Interchange Chemistry
Materials RNA oligonucleotide with 2′ amine group (100 µM in TE buffer; see Basic Protocol 1) ≥0.1 Ci/µL [γ-32P]ATP (6000 Ci/mmol) 10 U/µL T4 polynucleotide kinase and 10× buffer (New England Biolabs) 1 M sodium chloride Absolute ethanol 1 M sodium borate buffer, pH 8 500 mM N-succinimidyl-3-(2-pyridyldithio) propionate (S.3; Pierce Chemicals) in N,N-dimethylformamide (prepare solution just before use) 70 mM thiophenol in absolute ethanol TE buffer, pH 7.5 (APPENDIX 2A)
5.1.4 Current Protocols in Nucleic Acid Chemistry
1× TBE buffer (APPENDIX 2A) 80% formamide/TBE solution (see recipe) Denaturing polyacrylamide gel: 20% polyacrylamide/8 M urea in TBE buffer, dimensions 20 cm long × 10 cm wide × 0.05 cm thick (see APPENDIX 3B, CPMB UNIT 7.6 or Sambrook et al., 1989) TEN buffer (see recipe) 10 mM sodium acetate buffer, pH 4.5 (APPENDIX 2A) Preparative centrifuge (Sorvall or Beckman) X-ray film for autoradiography Water bath, 37°C Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (APPENDIX 3B, CPMB UNIT 7.6 or Sambrook et al., 1989) CAUTION: Step 8 should be performed in a fume hood because of the stench of thiophenol. Any labware that comes in contact with thiophenol should be soaked in bleach solution. Label the RNA 1. Mix the following in a 1.5-mL microcentrifuge tube (9 µL total volume): 1 µL 100 µM RNA oligonucleotide with 2′ amine 5 µL ≥0.1 Ci/µL [γ-32P]ATP (6000 Ci/mmol; ≥0.5 mCi total) 1 µL 10× T4 polynucleotide kinase buffer 2 µL H2O. Warm the solution to 37°C. 2. Add 1 µL of 10 U/µL T4 polynucleotide kinase and incubate 30 min at 37°C. 3. Add 200 µL water and 50 µL of 1 M sodium chloride; mix well. 4. Precipitate the labeled RNA by adding 900 µL absolute ethanol and centrifuging 20 min at 16,000 × g, 2°C. 5. Decant the supernatant and allow the RNA pellet to air dry. Modify the 2′ amine 6. Dissolve the labeled RNA in: 140 µL water 20 µL 1 M sodium borate buffer, pH 8 20 µL 1 M sodium chloride. Warm the solution to 37°C. 7. Add 20 µL freshly dissolved 500 mM N-succinimidyl-3-(2-pyridyldithio) propionate (S.3) in N,N-dimethylformamide and mix well. Incubate 20 min at 37°C. Upon addition of S.3, the solution will become cloudy because of the compound’s limited water solubility but will clear during the course of the reaction as a result of hydrolysis.
8. In a fume hood, add 200 µL of 70 mM thiophenol in ethanol. Incubate 2 min at 23°C. The reaction should display a light yellow color from formation of pyridine-2-thione.
9. Precipitate the RNA by adding 700 µL absolute ethanol and centrifuging 20 min at 16,000 × g, 2°C. 10. Decant the supernatant and allow the RNA pellet to air dry.
Methods For Cross-Linking Nucleic Acids
5.1.5 Current Protocols in Nucleic Acid Chemistry
Purify the modified RNA 11. Dissolve the crude product in 10 µL TE buffer and 10 µL of 80% formamide/TBE solution (20 µL total). 12. Purify the RNA by denaturing polyacrylamide gel electrophoresis at 15 W (see APPENDIX 3B, CPMB UNIT 7.6 or Sambrook et al., 1989, for standard procedures). A gel 20 cm long is sufficient to provide clean separation from the faster-migrating RNA that remains unmodified after treatment with S.3 (∼25%).
13. Identify the band by autoradiography and excise the gel slice. 14. Crush the gel slice thoroughly in a 1.5-mL microcentrifuge tube and add 500 µL TEN buffer. Incubate the suspension 10 min on ice, vortexing occasionally. 15. Pellet the polyacrylamide by centrifugation for 2 min at 16,000 × g, 2°C, and carefully withdraw the supernatant into a fresh 1.5-mL microcentrifuge tube. 16. Precipitate the RNA by adding 4 vol absolute ethanol and centrifuging 30 min at 16,000 × g, 2°C. 17. Dissolve the modified RNA in 100 µL 10 mM sodium acetate buffer, pH 4.5. Determine radioactivity level to confirm that it is ≥105 cpm/µL. Store in 20-µL aliquots at −80°C. BASIC PROTOCOL 3
PREPARATION OF RNA CONTAINING ALKYL THIOL GROUP The RNA component of the cross-linking system containing the alkyl thiol S.2 is prepared using similar chemistry as for S.1. The 2′ amine is modified with N-succinimidyl-3-(2pyridyldithio) propionate (S.3), yielding the corresponding amide S.4. Reduction of the pyridyl disulfide moiety of S.4 with dithiothreitol yields the alkyl thiol S.2 (Fig. 5.1.2). Materials 20 nmol RNA oligonucleotide with 2′ amine group (see Basic Protocol 1) 1 M sodium borate buffer, pH 8 1 M sodium chloride 500 mM N-succinimidyl-3-(2-pyridyldithio) propionate (S.3; Pierce Chemicals) in N,N-dimethylformamide (prepare just before use) Absolute ethanol 1 M dithiothreitol (DTT) in water (APPENDIX 2A) TEN buffer (see recipe) TE buffer, pH 7.5 (APPENDIX 2A) Water bath, 37°C Modify the 2′ amine 1. In a 1.5-mL microcentrifuge tube, precipitate 20 nmol of the RNA to be modified by mixing 200 µL of the 100 µM 2′-amine-modified RNA with 50 µL of 1 M sodium chloride followed by 750 µL absolute ethanol, then centrifuging 20 min at 16,000 × g, 2°C. 2. Dissolve the RNA pellet in:
RNA Cross-Linking Using Thiol-Disulfide Interchange Chemistry
120 µL H2O 40 µL 1 M sodium borate buffer, pH 8 20 µL 1 M sodium chloride. Warm the solution to 37°C.
5.1.6 Current Protocols in Nucleic Acid Chemistry
3. Add 20 µL freshly dissolved 500 mM S.3 in N,N-dimethylformamide and mix well. Incubate 20 min at 37°C. This reaction may be scaled up as needed, maintaining the following final concentrations: 200 mM NaB(OH)3, 100 mM NaCl, 100 mM RNA, and 50 mM S.3.
4. Precipitate the RNA by adding 600 µL absolute ethanol and centrifuging 20 min at 16,000 × g, 2°C. 5. Repeat steps 2 to 4. Double treatment with S.3 should modify ≥95% of the 2′ amines.
Liberate the thiol 6. Dissolve the RNA pellet in 140 µL water and 40 µL of 1 M sodium borate buffer, pH 8. Liberate the thiol by adding 20 µL of 1 M DTT. Incubate 30 min at 37°C. 7. Precipitate the RNA by adding 50 µL of 1 M sodium chloride and 750 µL absolute ethanol, and centrifuging 20 min at 16,000 × g, 2°C. 8. Dissolve the modified RNA in 300 µL TEN buffer. Precipitate the RNA by adding 900 µL absolute ethanol and then centrifuging again as in step 7. 9. Dissolve the modified RNA in 100 µL TE buffer. Determine RNA concentration spectrophotometrically by measuring A260, and adjust to 100 µM. Store up to 6 months at −20°C or indefinitely at –80°C. CROSS-LINKING OF RNA THROUGH THIOL-DISULFIDE INTERCHANGE The two RNA components containing S.1 and S.2 are associated at pH 4.5, where the nucleophilicity of the alkyl thiol S.2 is attenuated such that cross-linking before association is discouraged. Cross-linking is then initiated by increasing the pH, usually to the range of pH 7 to 8. The cross-linking reaction is conducted with the thiol component S.2 in saturating excess (up to 10 times the equilibrium dissociation constant) over the phenyl disulfide component S.1 to discourage spurious cross-linking resulting from the presence of unassociated S.1.
BASIC PROTOCOL 4
The following protocol is from original ribozyme cross-linking experiments by Cohen and Cech (1997). The concentrations of RNA components, cross-linking pH, and other experimental conditions can be adjusted to accommodate other experimental systems.
Materials 32 P-labeled RNA modified with phenyl disulfide S.1 (≥105 cpm/µL in 10 mM sodium acetate buffer, pH 4.5; see Basic Protocol 2) RNA modified with alkyl thiol S.2 (1 µM in TE buffer; see Basic Protocol 3) 100 mM sodium acetate buffer, pH 4.5 (APPENDIX 2A) 100 mM magnesium chloride 5 M sodium chloride Formamide quenching solution (see recipe) 1 M sodium HEPES buffer, pH 7.5 (APPENDIX 2A) 0.65-mL microcentrifuge tube Water bath, 30°C Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (APPENDIX 3B, CPMB UNIT 7.6 or Sambrook et al., 1989)
Methods For Cross-Linking Nucleic Acids
5.1.7 Current Protocols in Nucleic Acid Chemistry
1. In a 0.65-mL microcentrifuge tube, mix (90 µL total volume): 10 µL 1 µM S.2-modified RNA 20 µL 100 mM sodium acetate buffer, pH 4.5 10 µL 100 mM magnesium chloride 20 µL 5 M sodium chloride 30 µL H2O. Warm the solution to 30°C. 2. Add 10 µL of ≥105 cpm/µL 32P-labeled S.1-modified RNA. Incubate 30 min at 30°C. 3. Remove 5 µL of the reaction and transfer to a fresh microcentrifuge tube containing 25 µL formamide quenching solution. Store frozen on dry ice. This aliquot serves to measure the small amount of cross-linking (if any) that occurs during the association incubation at pH 4.5.
4. Initiate cross-linking by adding 10 µL of 1 M sodium HEPES buffer, pH 7.5, to attain a final reaction pH of ∼7.2. The absolute rates of cross-linking can be controlled by manipulating the pH of the cross-linking reaction (∆log k/∆pH ∼1 in the pH range of 4.5 to 8.0).
5. At the desired time intervals, transfer 5 µL of the reaction to separate tubes containing 25 µL formamide quenching solution. Store the time aliquots frozen on dry ice until analysis by denaturing polyacrylamide gel electrophoresis (see APPENDIX 3B, CPMB UNIT 7.6 or Sambrook et al., 1989). For example, in the original ribozyme cross-linking experiments, 3- to 5-min intervals were used.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Formamide, 80%, in TBE To 40 mL formamide add 25 mg bromphenol blue and 25 mg xylene cyanol; mix well. Add 10 mL of 5× TBE buffer (APPENDIX 2A). Store up to 6 months at 23°C. The resulting 50 mL contains 80% formamide, 0.05% bromphenol blue, 0.05% xylene cyanol, 90 mM Tris-borate, and 1 mM EDTA.
Formamide quenching solution 800 µL formamide 0.5 mg bromphenol blue 0.5 mg xylene cyanol 100 µL 1 M sodium acetate buffer, pH 4.5 100 µL 0.5 M disodium EDTA Prepare fresh This must be prepared fresh on the day of use because EDTA will precipitate during extended storage at pH 4.5. The resulting 1.0 mL contains 80% formamide, 0.05% bromphenol blue, 0.05% xylene cyanol, 100 mM sodium acetate buffer (pH 4.5), and 50 mM EDTA. RNA Cross-Linking Using Thiol-Disulfide Interchange Chemistry
5.1.8
TEN buffer 10 mM Tris⋅Cl, pH 7.5 (APPENDIX 2A) 1 mM disodium EDTA (APPENDIX 2A) 250 mM sodium chloride Store indefinitely at room temperature (e.g., 23°C). Current Protocols in Nucleic Acid Chemistry
Triethylamine/1-methyl-2-pyrrolidinone/triethylamine trihydrofluoride, 24/46/30 (v/v/v) To a stirred solution of 12 mL triethylamine and 23 mL 1-methyl-2-pyrrolidinone (Aldrich), add dropwise 15 mL triethylamine trihydrofluoride (Aldrich). Continue stirring until the solution is homogeneous (1.4 M fluoride ion). Store in 1-mL aliquots up to 6 months at −20°C. CAUTION: Wear gloves when working with fluoride solutions. This solution is derived from Wincott et al. (1995).
COMMENTARY Background Information The authors’ goal was to use site-specific disulfide cross-linking to measure long-range conformational dynamics within a large (>300nt) catalytic RNA molecule (ribozyme). Existing methods of forming disulfide bonds had relied on chemical oxidation of two thiols, a reaction that is prohibitively slow in the absence of a catalyst. Redox-active metal complexes, such as copper(II) phenanthroline, are effective oxidants (Patai, 1974) but are incompatible with RNA because they promote oxidative cleavage of the ribose backbone (Chen and Sigman, 1988). In addition, large structured RNAs often contain binding sites for multivalent metal ions, further complicating analysis. Sulfoxide catalysis has been used to form disulfides in RNA (Sigurdsson et al., 1995), but the experimental conditions required (≥50% dimethyl sulfoxide) are not compatible with a folded and active ribozyme structure. The new disulfide cross-linking procedure (Cohen and Cech, 1997) that is presented in this unit can be applied to large RNAs and allows kinetic characterization of the cross-linking reaction. Incorporation of cross-linking precursors S.1 and S.2 within RNA relies on the presence of a 2′ amine, incorporated by use of the corresponding phosphoramidite during solid-phase RNA synthesis (Verheyden et al., 1971; Imazawa and Eckstein, 1979; Benseler et al., 1992). Modification of the 2′ amine is then performed after the RNA is deprotected, labeled with 32P, or otherwise manipulated. For example, a 2′ amine is compatible with T4 DNA ligase; ligation of a 41-nt synthetic RNA containing a 2′ amine to a 269-nt transcribed RNA as described by Moore and Sharp (1992) provided a route for preparing a 310-nt ribozyme containing the alkyl thiol S.2 (Cohen and Cech, 1997). Thiol-disulfide interchange chemistry (Fig. 5.1.1) proceeds under mild conditions (aqueous
solution, pH ∼7) without an oxidative catalyst and has been used to measure conformational dynamics between helical domains within a 310-nt ribozyme (Cohen and Cech, 1997). Association of the substrate domain containing S.1 with a set of ribozymes, each containing S.2 at a different position, afforded substrateribozyme cross-linking representing interhelical displacements of at least 50 Å. The kinetic profile of the cross-linking revealed the distribution of motions between the two domains. Cross-linking was achieved under a variety of experimental conditions (temperature 30° to 50°C; pH 6 to 8; NaCl concentration 0 to 1.0 M; MgCl2 concentration 0 to 100 mM) and allowed preparation of a series of conformationally constrained substrate-ribozyme complexes.
Critical Parameters Optimization of the modification reaction of the 2′ amine with S.3 revealed a strong temperature dependence: the efficiency of the reaction was significantly lower at 23°C than at 37°C. Allow ample time for the solution to warm to 37°C before addition of S.3. The absolute rates of cross-linking can be controlled by manipulating the pH of the crosslinking reaction (∆log k/∆pH ~1). The pH may require optimization. For example, if the crosslinking precursors S.1 and S.2 are in close proximity within the associated RNA molecule, then a reaction pH of 8 may result in cross-linking that is too fast to measure (<1 min). Lowering the pH from 8 to 7 will slow the reaction. Careful purification of the 32P-labeled RNA containing S.1 will enhance the yield of the cross-linking reaction. In the preparation of S.1, the concentration of RNA in the modification reaction with S.3 is relatively low (∼1 µM, compared to 100 µM when preparing S.2); as a consequence, the reaction is less efficient. With careful electrophoresis, it is possible to
Methods For Cross-Linking Nucleic Acids
5.1.9 Current Protocols in Nucleic Acid Chemistry
isolate the RNA containing the S.1 modification as a distinct band because it will migrate more slowly than the RNA left with a free 2′ amine. Complete separation has been achieved for RNAs up to 30 nt.
Anticipated Results The yield of cross-link is defined as the percentage of 32P-labeled RNA containing S.1 that becomes cross-linked to the RNA modified with S.2. Typically, cross-linking yields are ∼70%; yields as low as 40% and as high as 90% have been observed. Factors that may affect the yield of cross-link include the temperature (experiments at 2°C afforded lower yields than those at 30°C), the proximity of S.1 and S.2 within the RNA molecule (which influences the reaction rate), and the purity of the RNA components, particularly S.1.
Time Considerations Preparation of the RNA oligonucleotide containing a 2′ amine will take 2 to 3 days. Preparation of RNA modified with S.1 or S.2 can usually be done within a single day for each. The length of time required for cross-linking will depend on the experimental system; crosslinking reactions have been observed to be complete within minutes to several hours.
Literature Cited Benseler, F., Williams, D.M., and Eckstein, F. 1992. Synthesis of suitably-protected phosphoramidites of 2′-fluoro-2′-deoxyguanosine and 2′amino-2′-deoxyguanosine for incorporation into oligoribonucleotides. Nucleosides Nucleotides 11:1333-1351. Chen, C.B. and Sigman, D.S. 1988. Sequence-specific scission of RNA by 1,10-phenanthrolinecopper linked to deoxyoligonucleotides. J. Am. Chem. Soc. 110:6570-6572. Cohen, S.B. and Cech, T.R. 1997. Dynamics of thermal motions within a large catalytic RNA investigated by cross-linking with thiol-disulfide interchange. J. Am. Chem. Soc. 119:6259-6268.
Goodwin, J.T., Osborne, S.E., Scholle, E.J., and Glick, G.D. 1996. Design, synthesis, and analysis of yeast tRNAPhe analogs possessing intraand interhelical disulfide cross-links. J. Am. Chem. Soc. 118:5207-5215. Imazawa, M. and Eckstein, F. 1979. Facile synthesis of 2′-amino-2′-deoxyribo-furanosyl purines. J. Org. Chem. 44:2039-2041. Moore, M.J. and Sharp, P.A. 1992. Site-specific modification of pre-mRNA: the 2′-hydroxyl groups at the splice sites. Science 256:992-997. Patai, S. 1974. The Chemistry of the Thiol Group. John Wiley & Sons, New York, N.Y. Sambrook, J., Fritsch, E.F., and Maniatis, T. 1989. Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. Sigurdsson, S.T., Tuschl, T., and Eckstein, F. 1995. Probing RNA tertiary structure: Interhelical cross-linking of the hammerhead ribozyme. RNA 1:575-583. Verheyden, J.P.H., Wagner, D., and Moffatt, J.G. 1971. Synthesis of some pyrimidine 2′-amino2′-deoxynucleosides. J. Org. Chem. 36:250-254. Wincott, F., DiRenzo, A., Shaffer, C., Grimm, S., Tracz, D., Workman, C., Sweedler, D., Gonzalez, C., Scaringe, S., and Usman, N. 1995. Synthesis, deprotection, analysis, and purification of RNA and ribozymes. Nucl. Acids Res. 23:2677-2684.
Key References Cohen and Cech, 1997. See above. Describes the initial development of thiol-disulfide interchange chemistry and its application to measuring conformational dynamics within a 310-nt catalytic RNA. Sigurdsson et al., 1995. See above. Presents the first example of the use of chemistry derived from a 2′ amine to tag the RNA backbone in a sequence-specific fashion.
Contributed by Scott B. Cohen and Thomas R. Cech University of Colorado Boulder, Colorado
RNA Cross-Linking Using Thiol-Disulfide Interchange Chemistry
5.1.10 Current Protocols in Nucleic Acid Chemistry
Chemical and Enzymatic Methods for Preparing Circular Single-Stranded DNAs
UNIT 5.2
Each of the following three protocols for preparing single-stranded DNA (ssDNA) circles from 28 to at least 188 nucleotides (nt) in length involves a “one-pot” cyclization scheme using various chemical ligation reagents or enzymes and short synthetic DNA splints. Each ligation can proceed from two or more short linear DNAs (precursor segments), which are intermolecularly ligated to give a linear precircle. This DNA is then intramolecularly ligated to form a circular DNA. Alternatively, a single full-length cyclization starting material can be used, again using intramolecular ligation conditions to close it. Enzymatic ligation (see Basic Protocol) is based on the use of a double-helical splint complex with a complementary ssDNA splint (Fig. 5.2.1). This complex assists enzymatic ligation by juxtaposing reactive ends of precursor segments for subsequent ligation to a ssDNA circle using T4 DNA ligase. Chemical ligation with cyanogen bromide (see Alternate Protocol 1) involves an additional, alternative method of assistance through a triple-helical splint complex (Fig. 5.2.1). Chemical ligation occurs by simultaneous dimeric ligation of linear DNAs to circular form upon addition of cyanogen bromide. The final procedure (see Alternate Protocol 2) is a novel chemical autoligation method that also uses a double-helical splint complex. The reaction proceeds via nucleophilic displacement of iodide from 5′-iodothymidine by the 3′-phosphorothioate group to create a 5′-bridging phosphorothioate linkage. A series of five support protocols cover the preparation and purification (optional) of the end-modified DNAs required for all ligation methods.
Figure 5.2.1 Cyclization of ssDNAs requires a splint to assist in end joining. Representation of two types of splint complexes used for ligation: double-helical splint complex using conventional Watson-Crick hydrogen bonding, and triple-helical splint complex using both Watson-Crick and Hoogsteen hydrogen bonding.
Methods for Cross-Linking Nucleic Acids
Contributed by Amy M. Diegelman and Eric T. Kool
5.2.1
Current Protocols in Nucleic Acid Chemistry (2000) 5.2.1-5.2.27 Copyright © 2000 by John Wiley & Sons, Inc.
STRATEGIC PLANNING Each protocol can be used for the synthesis of single-stranded circles ranging in size from 28 to at least 188 nt. The enzymatic ligation method has no inherent sequence restrictions; however, the autoligation method requires a 5′-iodothymidine and 3′-phosphorothioate group at the ligation junction, which in turn requires preparation of this DNA using a modified phosphoramidite, 5-iodothymidine, as well as a sulfurizing reagent. The chemical ligation method using cyanogen bromide has been very effective for the synthesis of variously sized circles by dimeric ligation of triple-helical splint complexes (Ruben et al., 1995). It is possible to prepare circles by any of the three ligation methods; however, one method may be more suitable for a given circle based on certain considerations. Figure 5.2.2 is a decision tree designed to help in choosing a protocol. The first decision involves the size of the circle. Circles <28 nt are not covered by these protocols, and can best be prepared using methodology described by Alazzouzi et al. (1997). Small circles, 28 to 50 nt, are most easily synthesized using a single precursor segment and any of the three basic protocols, with particular steps omitted as noted (Fig. 5.2.2). Medium-sized circles, 50 to 140 nt, should be synthesized from two smaller precursor segments, as preparative yields for convergent or dimeric synthesis of this sort are usually significantly better than those obtained using single precursors. Any of the three protocols can be used for the synthesis of medium-sized circles—be sure to note the specific sequence and size considerations for each protocol, however. Large circles, >140 nt, are best prepared from at least three precursor segments. The basic two-step ligation protocol is the same for the synthesis of these large circles; however, all precursor segments and all but one splint are included in the first ligation and the last splint is incorporated in the second ligation. Along with the
Chemical and Enzymatic Methods
Figure 5.2.2 Strategic planning for DNA circle synthesis using a decision tree. First decision is circle size. This decision will determine the number of precursor segments to use. The next decision involves analysis of sequence requirements and choice of a ligation method. AP, Alternate Protocol; BP, Basic Protocol; THSC, triple-helical splint complex.
5.2.2 Current Protocols in Nucleic Acid Chemistry
Figure 5.2.3 Schematic representation of the steps in “one-pot” ligation for the enzymatic cyclization of ssDNAs by T4 DNA ligase. (A) Complementary splints 20-nt long are used to form double-helical splint complexes to juxtapose reactive 3′-hydroxyl and 5′-phosphate ends of linear ssDNAs for enzymatic ligation to a phosphodiester linkage (B) using T4 DNA ligase.
Methods for Cross-Linking Nucleic Acids
5.2.3 Current Protocols in Nucleic Acid Chemistry
specific considerations for each of the three protocols, it must be kept in mind that synthesis of large circles from multiple precursor segments proceeds with low efficiency; the enzymatic method gives the lowest yields, behind the chemical and autoligation methods, whose yields are somewhat better and fairly comparable to one another. Both the enzymatic and reagent-free ligation protocols operate on a 10- to 15-nmol scale and can easily be scaled up to 100 to 150 nmol. However, scales >150 nmol involve rather large volumes for lyophilization and can become quite costly. NOTE: If the cyclized oligonucleotides are to be transcribed, all solutions and equipment coming into contact with DNA must be autoclaved to be free of RNase contaminants. All water used should be ultrapure (at least distilled and deionized). BASIC PROTOCOL
DOUBLE-HELICAL SPLINT COMPLEX–ASSISTED ENZYMATIC CYCLIZATION OF OLIGONUCLEOTIDES USING T4 DNA LIGASE The basic protocol describes synthesis and purification of ssDNA circles using a complementary short ssDNA template (a splint) to juxtapose the reactive 3′-hydroxyl and 5′-phosphate groups of precursor segments for ligation by T4 DNA ligase. The protocol is applicable to circles in the size range of 28 to at least 188 nucleotides. This “one-pot” synthesis consists of a two-step ligation of two approximately equal-length precursor segments, which are first enzymatically ligated into a full-length precircle, without isolation, and subsequently enzymatically cyclized into a ssDNA circle (Fig 5.2.3). Circles <50 nt may be synthesized from a single precursor segment by a single enzymatic ligation and circles >140 nt by ligation of more than two precursor segments; see Strategic Planning for more information. The products are separated by preparative denaturing polyacrylamide gel electrophoresis (PAGE), and circularity is confirmed by endonuclease cleavage. CAUTION: Acrylamide and N,N′-methylenebisacrylamide are neurotoxins. Preparation of solutions with these compounds should be performed in a well-ventilated fume hood, and extreme precautions taken to minimize contact with solids or solutions. NOTE: To synthesize larger quantities, the first ligation reaction can be scaled up. The second ligation reaction can also be scaled up, but multiple reaction tubes should be used rather than a larger-scale one-pot synthesis. NOTE: For cyclization using a single precursor segment, skip steps 4 to 7 (i.e., omit the first ligation and proceed directly from DNA quantitation to the second ligation described).
Chemical and Enzymatic Methods
Materials 20 nt splint ssDNA(s) Precursor segments containing 5′-phosphate and 3′-hydroxyl groups Ultrapure (e.g., distilled, deionized) water 2× ligation buffer: 20 mM MgCl2/100 mM Tris⋅Cl (pH 7.5) 1 M dithiothreitol (DTT; APPENDIX 2A) 25 mM and 100 mM adenosine triphosphate (ATP) 400 U/µL T4 DNA ligase (New England Biolabs) 1× TBE (APPENDIX 2A) 10% denaturing polyacrylamide gel mix (see recipe or purchase) 10% (w/v) ammonium persulfate (APS) TEMED or TMEDA (Life Technologies) Formamide loading buffer (see recipe)
5.2.4 Current Protocols in Nucleic Acid Chemistry
0.2 N NaCl 5× S1 nuclease buffer: 50 mM NaCl/50 mM NaOAc/5 mM ZnCl2, pH 4.6 332 U/µL S1 nuclease (from Aspergillus oryzae; Amersham Pharmacia Biotech) Stop solution (see recipe) Stains-all dye solution (see recipe) UV spectrophotometer 1.5-mL microcentrifuge tubes and cap locks, autoclaved 15-mL screw-top centrifuge tubes, autoclaved 90°C heat block or thermal cycler Glass wool Dialysis tubing, MWCO 1000 (e.g., SpectraPor) Gel electrophoresis equipment: Vertical (sequencing) gel stand 2000-V power supply Glass plates: 13 × 15.5 cm, 13 × 16.5 cm, 6.5 × 15 cm, and 6.5 × 16 cm Gel combs: 1.5-mm-thick with 1.5-cm-wide wells and 0.4 mm thick with 5-mm-wide wells Spacers: 1.5 mm thick and 0.4 mm thick UV shadow box or light source Lyophilizer or SpeedVac Saran Wrap (or other UV-transparent plastic wrap) Razor blades, sterile Glass stir rod 50-mL filter tubes with 0.45-µm cellulose acetate filter (e.g., Spin-X II, Corning Costar) Obtain splint DNAs 1. Construct or purchase ssDNAs of the desired splint and precursor-segment sequences. Splint sequences should bridge, by 10 nt on each side, the gap between precursor segments for ligation (Fig. 5.2.3A). For ssDNAs purchased commercially, splints should be fully deprotected and precursor segments should be fully deprotected and 5′-phosphorylated. Alternatively, ssDNAs may be constructed on a DNA synthesizer using the standard DNA cycle and a 0.2-µmol scale; see Support Protocol 1 for synthesis of modified precursor segments, see Support Protocol 4 for deprotection, and see Support Protocol 5 for (optional) purification.
Quantitate DNA 2. Redissolve each DNA in 1.00 mL water. Prepare a 100× dilution sample by combining 10.00 µL of this stock solution with 990 µL water. Synthesized DNA after deprotection and lyophilization should be powdery and white and should readily redissolve in water. If the solution does not appear homogeneous, mild heating (∼60°C) and/or brief sonication (1 min) can be used to help dissolve the DNA. Insoluble material may cause some turbidity but is not cause for concern.
3. Use a properly calibrated UV spectrophotometer to obtain a measurement of the absorbance at 260 nm (A260) of the 100× dilution sample prepared in step 2. Calculate the concentration (c) of DNA for the stock sample prepared in step 2 in moles per milliliter (recalling the dilution factor from step 2 of 102) using Beer’s Law: A260 = ε × b × c b is generally 1, so c (in mol/mL) = (A260 × 102)/ε
Methods for Cross-Linking Nucleic Acids
5.2.5 Current Protocols in Nucleic Acid Chemistry
Extinction coefficients (ε) for each sequence can be calculated using the nearest-neighbor method (Borer, 1985). See also UNIT 7.3. IMPORTANT NOTE: If a single precursor segment is to be used (i.e., synthesis of a ssDNA circle <50 nt), omit the first ligation (steps 4 to 7) and begin with step 8.
Perform first ligation 4. Combine 125.0 µL of 2× ligation buffer with 15 nmol of each precursor segment and 18.3 nmol of one of the splints (splint 1) in an autoclaved 1.5-mL microcentrifuge tube. Final concentrations: 10 mM MgCl2, 50 mM Tris⋅Cl, 60 mM each precursor segment, 73 mM splint 1. IMPORTANT NOTE: For large DNA circles (>140 nt), use all precursor segments and all but one splint in step 4. It may not matter which splint is chosen for the first ligation here; however, more is required for the second ligation, so if one is limiting, it should be used here. If the combined volume of DNA (two precursor segments and splint) exceeds 119 mL, one or more DNAs must be lyophilized to reduce the volume.
5. Calculate the volume of water to add according to the following equation: volume water = 250 µL − (125.0 µL + volume DNA [step 4] + 5.40 µL) Add the calculated amount of water to the microcentrifuge tube, vortex briefly, and secure closed with a cap lock. The 250 mL represents the total reaction volume, 125.0 mL represents the 2× ligation buffer, volume DNA represents the combined volumes of precursor segments and splint, and 5.40 mL represents volumes of reagents to be added in step 7.
6. Incubate the capped tube 10 min in a 90°C heat block. After 10 min, turn the heat block off and allow it to slowly cool, with its contents, to room temperature (∼25°C). Step 6 takes ∼2 to 2.5 hr to complete. Alternatively, a PCR thermal cycler can be used. Set the cycler to heat for 10 min at 90°C, then slowly cool (0.5°C/min) to room temperature. The slow cooling allows the splints to bind their complementary targets efficiently, aligning reactive ends for ligation by the enzyme.
7. When the block and its contents have reached room temperature (∼25°C), add 2.50 µL of 1 M DTT, 1.00 µL of 25 mM ATP, and 1.88 µL of 400 U/µL T4 DNA ligase to the tube. Gently invert the tube and let stand 4 to 6 hr at room temperature. Final concentrations: 10 mM DTT, 100 mM ATP, 3 U/mL T4 DNA ligase. Perform second ligation
8. In autoclaved 15-mL centrifuge tube, combine 5.0 mL of 2× ligation buffer with either 167.0 µL of reaction from step 7 (2⁄3 vol) or 10 nmol full-length synthesized precursor segment (if cyclization is from a single precursor segment), plus 30 nmol of the remaining splint (splint 2). Final concentrations: 10 mM MgCl2, 50 mM Tris⋅Cl, 1 mM precircle, 3 mM splint 2, 1.2 mM splint 1. IMPORTANT NOTE: For circles >140 nt, the final splint is added in this step.
Chemical and Enzymatic Methods
The 167.0 mL of reaction from step 7 represents, in theory, a maximum yield of 10 nmol precircle from ligated precursor segments. Alternatively, if the circle size is <50 nt, 10 nmol of full-length synthesized precursor segment can be used. Note that DNA concentrations are much less for this second ligation (1 mM precircle) than for the first ligation (60 µM
5.2.6 Current Protocols in Nucleic Acid Chemistry
each precursor segment). This favors the intramolecular cyclization reaction over an intermolecular reaction that produces undesired multimeric linear species.
9. Calculate the volume of water to add according to the following equation: volume water = 10,000 µL − (5000 µL + volume DNA [step 8] + 118.3 µL) Add this volume of water to the 15-mL tube(s), vortex briefly, and secure closed. The 10,000 mL represents the total reaction volume; 5000 mL represents the 2× ligation buffer; volume DNA represents the combined volumes of either 167.0 mL of step 7 reaction or the synthesized precursor segment, plus splint; and 118.3 mL represents volumes of reagents to be added later. Normally it will not be necessary to lyophilize any of the components at this step, as the calculated volume of water often exceeds 4.5 mL.
10. Wrap the tube(s) thoroughly with glass wool and incubate 10 min in a 90°C heat block, inverting them every 2 min. After 10 min, turn the heat block off and allow it to cool, with its contents, to room temperature (∼25°C). The glass wool will allow even heating and cooling of the reagents to ensure an efficient slow annealing of splint and precircle. Inverting the tubes also ensures thorough heating of the contents. As before, this slow annealing takes ∼2 to 2.5 hr to complete.
11. When the block and its contents have reached room temperature (∼25°C), add 100.0 µL 1 M DTT, 10.00 µL of 100 mM ATP, and 8.33 µL of 400 U/µL T4 DNA ligase to each tube. Gently invert the tube and allow reaction to proceed 12 to 16 hr at room temperature. Final concentrations: 10 mM DTT, 100 mM ATP, 0.33 U/mL T4 DNA ligase. Note the different concentrations of enzyme and DNA used here compared to the first ligation. While the final ATP and DTT concentrations remain the same, the enzyme and DNA concentrations are lowered to favor the intramolecular cyclization reaction and to conserve enzyme and thus decrease cost.
Dialyze reaction products 12. Transfer the entire reaction volume from step 11 to MWCO 1000 dialysis tubing clamped at one end to make a bag. Secure the other end of the dialysis bag closed. Dialyze against 2 L ultrapure water for 3 hr. Repeat. 13. Lyophilize the dialysate to dryness. The dried residue should look off-white in color.
Gel purify reaction products 14. Prepare a vertical, 1.5-mm-thick (preparative), denaturing 10% polyacrylamide gel using a comb that produces 1.5-cm-wide wells. A gel prepared between plates used for sequencing (13 × 16.5 inch) allows for ample vertical loading of reaction mixture and sufficient migration length for separation and resolution of bands. For these conditions, prepare gel solution from 250 mL of 10% gel mix, 1.67 mL of 10% APS, and 53.3 mL TEMED. The gel should contain ∼12 wells for reaction solution plus a few for DNA markers. For more detailed description of procedures for denaturing PAGE, see APPENDIX 3B.
15. Place the gel in a gel electrophoresis apparatus, using 1× TBE in both top and bottom buffer reservoirs, and prerun 30 min at 30 W constant power, room temperature. 16. Redissolve each 10 mL lyophilized sample (from second ligation, step 13) in 60 µL water and 60 µL formamide loading buffer by brief vortexing.
Methods for Cross-Linking Nucleic Acids
5.2.7 Current Protocols in Nucleic Acid Chemistry
If the solution does not appear homogeneous, mild heating (∼60°C) and/or brief sonication (1 min) can be used to help dissolve the DNA mixture.
17. Load 10 µL of the solution from step 16 in each of the 1.5-cm-wide wells. Load DNA markers, step 2 solutions, and first ligation-reaction mixture (step 7) in separate adjacent wells (use ∼1 nmol of each to permit later visualization by UV shadowing). Use of DNA markers allows for confirmation of product lengths. Precursor segments (step 2) and solution from step 7 (first ligation) can be used to identify precursor segments and precircles in the reaction lanes.
18. Run the gel at 30 W constant power, room temperature, to the desired length. On a 10% gel bromphenol blue migrates similarly to a 26-nt linear DNA, so this band is usually run off, since most circles will be larger. Xylene cyanol runs like a 55-nt linear DNA, and for larger circles (>70 nt) should be run almost off the gel.
19. Disassemble the gel apparatus. Sandwich the prep gel between two pieces of Saran Wrap. View the gel under ultraviolet light with fluorescent white background (glass silica plates work well) to visualize the bands. The reaction lanes should show several bands. Figure 5.2.4 is a representation of a typical gel, with lane 6 containing second ligation reaction mixture. Depending on how far the gel is run, the splints (lane 1) will probably be run off, but the precursor segments may be visible and their identity can be confirmed by their co-migration with the precursor-segment markers (step 2 solutions) run alongside (lanes 2 and 3). The next band up (migrating more slowly) from the precursor segments should be the precircle band; this is confirmed by its co-migration with the largest band from the first ligation reaction (step 7) run alongside (lane 4). Although the secondary structure present in many DNA circles may cause them to migrate more slowly than their linear counterparts (Serwer and Allen, 1984), identification of the circle (lane 8) can be difficult, as there may be one or more products in between the linear precircle and monomer circle. These are likely to be ligation of an odd number of precursor segments (lane 7), and results are especially complex if precursor segments are of different sizes. Migration of circles is not always consistent and can be temperature dependent. Therefore, most bands migrating more slowly than confirmed precircle should be cut out for analysis.
Isolate purified DNAs after gel electrophoresis 20. Cut out the desired band with a clean, sterile razor blade and place the pieces in a 15-mL centrifuge tube. Using a glass stir rod, crush the gel pieces thoroughly. Combine crushed pieces with ∼5 to 10 mL of 0.2 N NaCl to make an easily shaken slurry. Repeat, in separate tubes, for each band of interest isolated. Shake each slurry for ∼12 hr. Precircle and higher bands should be isolated for reasons stated in step 19. Be sure all crushed gel pieces are transferred to the slurry. Conversion of precircle to circle may appear high by UV shadowing (step 19), but isolated yields after step 22 are often <50%; it is therefore important to ensure quantitative transfer of all materials.
21. Filter the slurry by brief (1- to 5-min) centrifugation through 50-mL filter tubes. Resuspend the gel pieces in 5 mL water, centrifuge again, and combine the filtrates. It is advisable to save the filtered gel pieces until after quantitation, as additional extractions with 0.2 N NaCl may be desired to increase isolated product yield, although only minimal (<10%) additional recoveries should be expected. Chemical and Enzymatic Methods
22. Dialyze the filtrate from step 21 as described in step 12, but using 2 L ultrapure water for 6 hr and repeating three times. 23. Quantitate as described in step 3 using the dialysate from step 22 with no dilution.
5.2.8 Current Protocols in Nucleic Acid Chemistry
Figure 5.2.4 Representation of a typical cyclization gel. Lanes 1 to 3 represent crude synthesized DNAs used for ligation; banding underneath attests to their impurity. Lane 4 represents the first ligation-reaction mixture; the largest (slowest-migrating) species is the precircle, shown after isolation in lane 5. Lane 6 represents the second ligation (cyclization) and the possible products as confirmed by their migration with isolated species [lanes 7 to 9: 1.5× precircle (from ligation of 3 precursor segments in a linear fashion), circle and dimer]. Note the migration of the bands isolated in lanes 7 to 9 is typical but may differ slightly with different methods and sequences. Note: While the ligation mechanism selects for correct sequence precursor segments, it is not unusual to see less than full-length impurities (n − 1, n − 2, etc.), and these impurities are often eliminated by careful isolation of the desired band.
Incorrect identification of bands may result in erroneous concentration calculations from the use of the wrong extinction coefficient. Confirmation of identification will be discussed in the next section. If a misidentification is discovered, simple recalculation with a different extinction coefficient is all that is required to correct the calculation error, as the absorbance is the same regardless of identification.
Confirm circularity of product 24. Place ∼0.2 nmol of the suspected circular DNA in each of two tubes and ∼0.2 nmol of isolated precircle in each of two tubes (four tubes total). Label one of each pair “control” and one “S1 nuclease reaction,” and lyophilize. 25. To each tube from step 24, add 0.60 µL of 5× S1 nuclease buffer. To the control tubes, add 2.50 µL water; to the reaction tubes, add 2.00 µL water. To the reaction tubes, add 0.50 µL diluted S1 nuclease (prepared from 0.50 µL of 332 U/µL enzyme stock +
Methods for Cross-Linking Nucleic Acids
5.2.9 Current Protocols in Nucleic Acid Chemistry
Figure 5.2.5 Representation of a typical characterization gel. Lanes 1 and 3 represent isolated precircle and circle controls. Lanes 2 and 4 represent precircle and circle cleavage by S1 nuclease, an endonuclease. As would be expected for endonuclease cleavage of a circle (lane 4), the initial cleavage produces a species that exhibits the same migration as the full-length cyclization precursor (precircle). Initial endonuclease cleavage of the linear precircle (see in lane 2 and also lane 4), by contrast, produces a continuous banding pattern.
373 µL water). Incubate all tubes 10 min at 37°C. Stop reactions by adding 1 vol (3.10 µL) stop solution to each. 26. Prepare a vertical, 0.4-mm-thick (analytical), denaturing 10% polyacrylamide gel using a comb that produces ∼5-mm-wide wells. Prerun the gel as in step 15. Using half-width sequencing plates (6.5 × 16 in) and 5-mm well width comb, the entire reaction volume (6.20 mL) can be analyzed. For these conditions, prepare gel solution from 50 mL of 10% gel mix, 333 mL of 10% APS, and 10.67 mL TEMED.
27. Load the entire reaction volume, 6.20 µL, on the gel and run dyes to the same distance as for the second ligation purification gel (step 18). 28. When electrophoresis is complete, disassemble the gel apparatus and transfer the analytical gel to a piece of Saran Wrap. Transfer the gel to an ultrapure water bath and soak 10 min to remove urea. Chemical and Enzymatic Methods
5.2.10 Current Protocols in Nucleic Acid Chemistry
29. Remove the water soak and add 200 mL of Stains-all dye solution. Wait 10 min for staining of bands. Background staining can be lightened by exposing the gel to incandescent light (100 W, held at 1 to 2 feet) while it is immersed in a fresh water bath.
30. Assess circularity of the DNA: this is confirmed by the absence of banding between the circle band and the precircle band (each confirmed by appropriate control). Initial cleavage of a circle produces a single band with the mobility of the full-length precursor, the precircle, Figure 5.2.5 (compare lanes 1 and 4). If the putative circular DNA is actually linear, as is the case for the precircle reaction lane (lane 2), banding will be seen directly under the suspected band (compare lanes 1 and 2).
TRIPLE-HELICAL SPLINT COMPLEX–ASSISTED CHEMICAL CYCLIZATION OF OLIGONUCLEOTIDES USING CYANOGEN BROMIDE
ALTERNATE PROTOCOL 1
The protocol for chemical ligation using cyanogen bromide involves synthesis and purification of ssDNA circles using a pyrimidine-rich triple-helical-forming ssDNA template (a splint) to juxtapose reactive 3′-hydroxyl and 5′-phosphate (or 5′-hydroxyl and 3′-phosphate) ends of linear ssDNAs for chemical ligation using cyanogen bromide. It is noteworthy that unlike in enzymatic ligation (see Basic Protocol), either orientation of phosphate or hydroxyl is usable (i.e., the precursor segments can be either 3′- or 5′-phosphorylated). The protocol is applicable for triplex-forming circles (TFCs) in the size range of 28 to ∼150 nt. The synthesis of TFCs begins with two approximately equal-length precursor segments that are highly pyrimidine rich at the ligation site, allowing for formation of a triple-helical splint complex with a purine-rich splint for simultaneous dimeric ligation into a ssDNA circle (see Fig. 5.2.6A). The basic chemistry of ligation can be either that depicted in Figure 5.2.3B or Figure 5.2.6B. The products are separated by preparative denaturing PAGE, and circularity confirmed by endonuclease cleavage. CAUTION: Cyanogen bromide is highly toxic and volatile; all work with this compound should be performed in a well-ventilated fume hood. Additional Materials (also see Basic Protocol) One purine-rich, triple-helical-forming splint ssDNA Two triplex-forming precursor segments containing either 3′- or 5′-phosphates 2× ligation buffer: 200 mM NiCl2/400 mM imidazole⋅HCl (pH 7.0) Cyanogen bromide (BrCN; solid) 1. Construct or purchase ssDNAs of the desired splint and precursor-segment sequences. Splint sequence should bridge, by 10 nt on each side, the gap between precursor segments for ligation (see Fig. 5.2.6A). For ssDNAs purchased commercially, splints should be fully deprotected and precursor segments should be fully deprotected and either 5′- or 3′-phosphorylated. For use of commercially purchased DNAs, proceed to step 2. Alternatively, ssDNAs may be constructed on a DNA synthesizer using the standard DNA cycle and a 0.2-mmol scale; see Support Protocol 2 for 3′-phosphate- or Support Protocol 1 for 5′-phosphate-containing precursor segments, Support Protocol 4 for deprotection, and Support Protocol 5 for (optional) purification.
Quantitate DNA 2. Quantitate DNA (see Basic Protocol, steps 2 and 3).
Methods for Cross-Linking Nucleic Acids
5.2.11 Current Protocols in Nucleic Acid Chemistry
Figure 5.2.6 Schematic representation of ligation steps for the chemical cyclization of ssDNAs using cyanogen bromide, BrCN. (A) A purine-rich ssDNA splint is used to form a triple-helical splint complex to juxtapose reactive 3′-phosphate and 5′-hydroxyl ends of linear DNAs for a simultaneous dimeric ligation to a phosphodiester linkage, (B), using BrCN. Note: For chemical ligation using BrCN, a 5′-phosphate and 3′-hydroxyl such as are used in Figure 5.2.3B are also suitable.
Ligation 3. Combine 750 µL of 2× ligation buffer with 75 nmol of each precursor segment and 82.5 nmol of the splint in an autoclaved 1.5-mL microcentrifuge tube Chemical and Enzymatic Methods
Final concentrations: 100 mM NiCl2, 200 mM imidazole⋅HCl, 50 mM each precursor segment or precircle, 55 mM splint.
5.2.12 Current Protocols in Nucleic Acid Chemistry
If combined volume of DNA (two precursor segments and splint) exceeds 750 µL, one or more DNAs must be lyophilized to reduce the volume.
4. Calculate the volume of water to add according to the following equation: volume water = 1500 µL − (750 µL + volume DNA [step 3]) Add the calculated amount of water to the microcentrifuge tube and vortex thoroughly. The 1500 mL represents the total reaction volume, 750 mL represents the 2× ligation buffer, and DNA represents the combined volumes of both precursor segments and the splint.
5. Weigh out 19.9 mg cyanogen bromide (125 mM final concentration) and add to the microcentrifuge tube. Thoroughly vortex to completely dissolve cyanogen bromide and allow to react at 23°C (room temperature) for 12 hr. CAUTION: Cyanogen bromide is highly toxic; all work with this compound should be performed in a well-ventilated fume hood. Do not let the reaction proceed beyond 12 hr, as degradation of DNA may become problematic.
Dialyze reaction after ligation 6. Dialyze the ligation-reaction mixture (see Basic Protocol, steps 12 and 13). Gel purify DNA products 7. Gel purify the dialyzed mixture (see Basic Protocol, steps 14 to 19). The reaction lanes should show several bands. Figure 5.2.4 is a representation of a typical gel, with lane 6 containing the ligation-reaction mixture. Depending on how far the gel is run, the splint will probably be run off, but the precursor segments or precircle may be visible and their identity can be confirmed by their co-migration with appropriate DNA markers from step 2 (lanes 1 to 3) run alongside. Although DNA circles usually migrate more slowly than their linear counterparts (Serwer and Allen, 1984), identification of circles is not trivial. Circle migration is not always consistent and can be temperature dependent. Triple-helical splint complexes can yield multimers from ligation of an odd number of precursor segments (lane 7), although the dimeric mechanism for ligation reduces this tendency. However, most bands migrating more slowly than precursor segments or precircle should be cut out for analysis.
Isolate purified DNAs 8. Isolate purified DNAs from the electrophoretic gel (see Basic Protocol, steps 20 to 23). Precircle and higher products should be isolated for reasons stated in step 7.
Confirm circularity 9. Confirm circularity of the desired DNA product (see Basic Protocol, steps 24 to 30). DOUBLE-HELICAL SPLINT COMPLEX–ASSISTED REAGENT-FREE CYCLIZATION OF OLIGONUCLEOTIDES USING A 3′-PHOSPHOROTHIOATE GROUP AND 5′-IODOTHYMIDINE The protocol describes synthesis and purification of ssDNA circles using a complementary ssDNA template (a splint) to create a double-helical splint complex to juxtapose reactive 3′-phosphorothioate group and 5′-iodothymidine of ssDNAs for autoligation by nucleophilic displacement. The protocol is applicable to circles in the size range of 28 to at least 188 nucleotides. The “one-pot” synthesis consists of two approximately equal-
ALTERNATE PROTOCOL 2
Methods for Cross-Linking Nucleic Acids
5.2.13 Current Protocols in Nucleic Acid Chemistry
Chemical and Enzymatic Methods
Figure 5.2.7 Schematic representation of “one-pot” autoligation steps for the reagent-free cyclization of ssDNAs. (A) Complementary splints are used to form double-helical splint complexes to juxtapose reactive 3′-phosphorothioate and 5′-iodothymidine ends of linear ssDNAs for autoligation via nucleophilic attack by phosphorothioate on iodothymidine. (B) Basic chemistry of autoligation.
5.2.14 Current Protocols in Nucleic Acid Chemistry
length linear precursor segments, which are ligated to form a linear precircle and subsequently cyclized into a ssDNA circle (see Fig. 5.2.7). Circles <50 nt may be synthesized from a single precursor segment by a single ligation and circles >140 nt may be synthesized by ligation of more than two precursor segments; see Strategic Planning for more information. The products are separated by preparative denaturing PAGE and circularity confirmed by endonuclease cleavage. NOTE: To synthesize larger quantities, the first ligation reaction can be scaled up. The second ligation reaction can also be scaled up, but multiple reaction tubes should be used rather than a larger-scale one-pot synthesis. NOTE: If a single precursor segment is to be used (i.e., synthesis of a ssDNA circle <50 nt), skip steps 3 to 5 (i.e., omit the first ligation and proceed directly from quantitation to the second ligation described). Additional Materials (also see Basic Protocol) 20 nt splint ssDNAs Precursor segments containing a 3′-phosphorothioate and 5′-iodothymidine 2× ligation buffer: 20 mM MgCl2/20 mM Tris⋅acetate, pH 7.00 Obtain the ssDNAs 1. Construct or purchase ssDNAs of the desired splint and precursor-segment sequences. Precursor segment oligos must be designed such that the chosen ligation junction contains a thymine on the 5′ side; this allows incorporation of the modified base, 5′-iodothymidine, for autoligation. Splint DNAs should bridge, by 10 nt on each side, the gap between precursor segments for ligation (see Fig. 5.2.7A). For ssDNAs purchased commercially, be sure splints are fully deprotected and precursor segments are fully deprotected and synthesized with a 5′-iodothymidine and 3′-phosphorothioate group. Alternatively, ssDNAs may be synthesized on a DNA synthesizer using the standard 0.2-mmol DNA cycle for splints and a slightly modified protocol for precursor segments and precircles; see Support Protocol 3 for 3′-phosphorothioate- and 5′-iodothymidine-containing precursor segments, and see Support Protocol 4 for deprotection.
Quantitate DNA 2. Quantitate DNA (see Basic Protocol, step 2). Perform first ligation
3. Combine 125.0 µL of 2× ligation buffer with 15 nmol of each precursor segment and 18.3 nmol of one of the splints (splint 1) in an autoclaved 1.5-mL microcentrifuge tube. IMPORTANT NOTE: For large DNA circles, >140 nt, use all precursor segments and all but one splint in step 3. Final concentrations: 10 mM MgCl2, 10 mM Tris⋅acetate, 60 mM each precursor segment, 73 mM splint 1. It may not matter which splint is chosen for the first ligation here; however, more is required for the second ligation, so if one is limiting, it should be used here. If combined volume of DNA (two precursor segments and splint) exceeds 125.0 mL, one or more DNAs must be lyophilized to reduce the volume.
4. Calculate the volume of water to add according to the following equation: volume water = 250 µL − (125.0 µL + volume DNA [step 3]) Add the calculated amount of water to the 1.5-mL tube, vortex briefly, and secure closed with a cap lock.
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5.2.15 Current Protocols in Nucleic Acid Chemistry
The 250 mL represents the total reaction volume, 125.0 mL represents the 2× ligation buffer, and volume DNA represents the combined volumes of either precursor segments or precircle, plus the splint.
5. Incubate the capped tubes 18 hr at room temperature. Perform second ligation 6. In an autoclaved 15-mL centrifuge tube, combine 5 mL of 2× ligation buffer with either 167.0 µL (2⁄3 volume) of the reaction from step 5 or 10 nmol full-length synthesized precursor segment if cyclization is from a single precursor segment, and 30 nmol of remaining splint (splint 2). Final concentrations: 10 mM MgCl2, 10 mM Tris⋅acetate, 1 mM precircle, 3 mM splint 2, 1.2 mM splint 1. IMPORTANT NOTE: For circles >140 nt, the final splint is added in this step. The 167.0 mL of reaction from step 5 represents, in theory, a maximum yield of 10 nmol of precircle from ligated precursor segments. Alternatively, 10 nmol of synthesized full-length precursor segment can be used if circle size is <50 nt. Note that DNA concentrations are much lower for this second ligation (1 mM precircle) than for the first (60 mM each precursor segment). This is meant to favor the intramolecular cyclization reaction versus an intermolecular reaction that produces multimeric linear species.
7. Calculate the volume of water to add according to the following equation: volume water = 10,000 µL − (5000 µL + volume DNA [step 6]) Add this volume of water to the 15-mL tube(s), vortex briefly, and secure closed. The 10,000 mL represents the total reaction volume, 5000 mL represents the 2× ligation buffer, and volume DNA represents the combined volumes of either 167 mL step 5 reaction or synthesized precircle, plus the splint. Normally it will not be necessary to be lyophilized any of the components at this step, as the calculated volume of water often exceeds 4.5 mL.
8. Incubate the tubes 18 hr at room temperature. Dialyze reaction 9. Dialyze the reaction mixture from the second ligation (see Basic Protocol, steps 12 and 13). Gel purify DNA products 10. Gel purify the dialyzed mixture (see Basic Protocol, steps 14 to 19).
Chemical and Enzymatic Methods
The reaction lanes should show several bands. Figure 5.2.4 is a representation of a typical gel, with lane 6 containing cyclization mixture. Depending on how far the gel is run, the splints will probably be run off, but precursor segments may be visible and their identity can be confirmed by their co-migration with the splints and precursor-segment markers (step 2) run alongside (lanes 1 to 3). The next band up from precursor segments should be the precircle band and can be confirmed by co-migration with the largest band from the ligation reaction (step 5) run alongside (lane 4). There is the potential for non-unit-length linear multimers from the linear ligation of an odd number of precursor segments leading to products in-between precircle and circle (lane 7). Although DNA circles usually migrate more slowly than their linear counterparts (Serwer and Allen, 1984), identification of circles is not trivial. Circle migration is not always consistent and may be temperature dependent. Therefore, most bands migrating more slowly than confirmed precircle should be cut out for analysis.
5.2.16 Current Protocols in Nucleic Acid Chemistry
Isolate purified DNAs 11. Isolate purified DNAs from the electrophoretic gel (see Basic Protocol, steps 20 to 24). Precircle and higher products should be isolated for reasons stated in step 10.
Confirm circularity 12. Confirm circularity of the desired DNA product (see Basic Protocol, steps 24 to 30). AUTOMATED SYNTHESIS OF PRECURSOR SEGMENTS CONTAINING 5′-PHOSPHATE FOR LIGATION
SUPPORT PROTOCOL 1
The support protocol describes incorporation of 5′-phosphate end groups on precursor segments necessary for their use in the enzymatic ligation methodology presented in the Basic Protocol. While DNAs can be purchased containing the desired end groups, modification during automatic DNA synthesis is straightforward. Introduction of a 5′-phosphate is described. NOTE: Precursor segments will contain a 3′-hydroxyl and 5′-phosphate after deprotection (see Support Protocol 4). Materials DNA synthesis reagents for 0.2-µmol synthesis Phosphorylating reagent (Chemical Phosphorylating Reagent, Glen Research) DNA synthesizer with phosphoramidite chemistry 1. Synthesize all natural bases for precursor segments, with completion of the synthesis cycle retaining the 5′-DMT protecting group. 2. Place phosphorylating reagent in a synthesizer port. 3. Set up the synthesis cycle for normal DNA, using the DNA column from step 1 as the starting column. Enter the sequence with the first (3′) base as the last base added to the column in step 1 and the next added “nucleotide” as the port containing the phosphorylating reagent. Complete the synthesis cycle with DMT removal. This adds a 5′ phosphate to each of the precursor segments.
4. Proceed to deprotection (see Support Protocol 4). AUTOMATED SYNTHESIS OF PRECURSOR SEGMENTS CONTAINING 3′-PHOSPHATE FOR LIGATION
SUPPORT PROTOCOL 2
The support protocol describes incorporation of 3′-phosphate end groups on precursor segments necessary for their use in the chemical ligation methodology presented in Alternate Protocol 1. While DNAs can be purchased containing the desired end groups, modification during automatic DNA synthesis is straightforward. Introduction of a 3′-phosphate is described. Materials DNA synthesis reagents for 0.2-µmol synthesis Phosphorylating reagent (Chemical Phosphorylating Reagent, Glen Research) DNA synthesizer with phosphoramidite chemistry NOTE: This is analogous to addition of a 3′ phosphorothioate in Support Protocol 3, but standard oxidizing (rather than sulfurizing) conditions are used to afford a 5′-hydroxyl
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5.2.17 Current Protocols in Nucleic Acid Chemistry
and 3′-phosphate, rather than a 3′-phosphorothioate, after deprotection (Support Protocol 4). 1. Begin synthesis of the precursor segment using any of the four base columns. Set up the synthesis cycle for normal DNA. 2. Place phosphorylating reagent in a synthesizer port. 3. Set up the synthesis cycle for normal DNA. Enter the sequence with the first (3′) base as the base located on the column and the next added “nucleotide” as the port containing phosphorylating reagent. Complete the cycle with DMT left on. This adds a 5′-phosphate to the column. Any normal base column can be used here, as deprotection of the finished product will cleave to the 5′ side of this initially added phosphate group, thus cleaving the finished oligo from the 3′ base present on the CPG bead. The DMT group should be left on for the next base to be added, which is the 3′ base in the sequence.
4. Add the first (3′) nucleotide for the desired sequence to the column. Complete synthesis of the remaining bases on the oligo with removal of the DMT protecting group. 5. Proceed to deprotection (see Support Protocol 4). SUPPORT PROTOCOL 3
AUTOMATED SYNTHESIS OF PRECURSOR SEGMENTS CONTAINING 3′-PHOSPHOROTHIOATE AND 5′-IODOTHYMIDINE FOR LIGATION This support protocol describes incorporation of 3′-phosphorothioate and 5′-iodothymidine end groups on precursor segments necessary for their use in the autoligation chemistry presented in Alternate Protocol 2. While DNAs can be purchased containing the desired end groups, modification during automatic DNA synthesis is straightforward. Phosphorylation at the 3′ end, subsequent modification of this end group to a 3′-phosphorothioate, and introduction of a 5′-iodothymidine are described. Materials DNA synthesizer with phosphoramidite chemistry DNA synthesis reagents for 0.2-µmol synthesis Phosphorylating reagent (S.1 in Fig. 5.2.8; Chemical Phosphorylating Reagent, Glen Research) Sulfurizing reagent (S.3; Applied Biosystems) Iodothymidine phosphoramidite (5′-I-dT-CE phosphoramidite, Glen Research) NOTE: Precursor segments will contain a 5′-iodothymidine and a 3′-phosphorothioate rather than 5′-hydroxyl and 3′-hydroxyl after deprotection (see Support Protocol 4). 1. Begin synthesis of the precursor segment using any of the four normal base columns. Set up the synthesis cycle for normal DNA. 2. Place phosphorylating reagent in a synthesizer port. 3. Enter the sequence with the first (3′) base as the base located on the column and the next added “nucleotide” as the port containing phosphorylating reagent. Complete the cycle with DMT left on. This adds a 5′-phosphate to the 3′ base of the column.
Chemical and Enzymatic Methods
Any base column can be used here, as deprotection of the finished product will cleave to the 5′ side of this initially added phosphate group (S.4), thus cleaving the finished oligo
5.2.18 Current Protocols in Nucleic Acid Chemistry
Figure 5.2.8 Schematic representation of steps involved in incorporation of a 3′-phosphorothioate group during DNA synthesis. First, a 3′-phosphate is coupled to the base on the CPG bead, S.1. Addition of the next base, S.2, the 3′ base of the desired sequence, proceeds with the trivalent phosphorus of this internucleotide bond converted to a phosphorothioate linkage using a sulfurizing reagent, S.3. Standard deprotection cleaves 3′ to this phosphorothioate linkage, S.4, creating a 3′-phosphorothioate group for coupling, S.5.
from the 3′-nucleotide present on the CPG bead (S.5). The DMT group should be left on for the next base to be added, which is the 3′ base in the sequence of interest. Alternatively, commercially available phosphate base columns can be used.
4. Set up the synthesizer to use a sulfurizing reagent (S.3) rather than an oxidizing reagent for this coupling only. 5. Proceed to add this next base and stop the synthesis after completion of this coupling. This modified synthesis cycle (involving S.2 and S.3) adds the first (3′) nucleotide for the sequence of interest to the column from step 3. The sulfurizing reagent converts the 3′ trivalent phosphorus linkage to a phosphorothioate linkage, S.4, rather than oxidized to a phosphodiester linkage as occurs during the standard DNA synthesis cycle. Subsequent
Methods for Cross-Linking Nucleic Acids
5.2.19 Current Protocols in Nucleic Acid Chemistry
deprotection will cleave 3′ to this linkage, leaving a 3′-phosphorothioate group as the nucleophile for autoligation (S.5).
6. Change the synthesis cycle back to standard (oxidizing agent instead of sulfurizing) and complete the synthesis of the desired precursor segments to the point at which synthesis of the natural bases (i.e., without iodothymidine) is complete, retaining the 5′-DMT protecting group. IMPORTANT NOTE: Do not forget to change the synthesis cycle back to oxidizing from sulfurizing.
7. Place the modified phosphoramidite (5′-I-dT-CE phosphoramidite) in a synthesizer port. 8. Set up the synthesis cycle for normal DNA using the DNA column from step 5 as the starting column. Enter the sequence with the first (3′) base as the last base added to the column from step 5, and the next added “nucleotide” as the port containing the modified phosphoramidite. Complete the synthesis cycle without DMT removal. This adds the modified phosphoramidite 5′-iodothymidine to each of the precursor segments. A DMT removal step is unnecessary because this phosphoramidite does not have a DMT protecting group.
9. Proceed to deprotection (see Support Protocol 4). SUPPORT PROTOCOL 4
DEPROTECTION OF PRECURSOR SEGMENTS After completion of synthesis, the oligonucleotides must be cleaved from the solid support and protecting groups which are needed during synthesis must be removed. The procedure is different for precursor segments that do and do not contain 3′-phosphorothioate and 5′-iodothymidine end groups in that gentler incubation conditions are used for the former. Materials Concentrated ammonium hydroxide Pasteur pipets Glass wool 1.5-mL screw-cap vial 1. Place DNA-loaded CPG beads from the column with 1 mL concentrated ammonium hydroxide in a 1.5-mL screw-cap vial. Incubate 24 hr at room temperature for precursors containing 3′-phosphorothioate and 5′-iodothymidine end groups, or 14 to 16 hr at at 55°C for precursors not containing those groups. 2. Filter the solution through a glass wool–plugged pipet to separate CPG beads. Use a second Pasteur pipet to transfer the solution.
3. Lyophilize the filtrate to remove ammonia and water. The crude DNA product should be powdery and white. SUPPORT PROTOCOL 5
Chemical and Enzymatic Methods
PURIFICATION OF DEPROTECTED PRECURSOR SEGMENTS (OPTIONAL) Slightly increased product yields may be possible through the use of purified products. However, this will result in some product loss and is not needed, as each splint complex– assisted ligation reaction mechanism selects for correct, full-length sequences, and the final product is gel purified to eliminate undesired DNAs. This support protocol is not recommended for precursor segments containing 3′-phosphorothioate or 5′-iodothymid-
5.2.20 Current Protocols in Nucleic Acid Chemistry
ine end groups. The protocol describes a standard denaturing acrylamide gel method for purification of deprotected oligonucleotides. CAUTION: Acrylamide and N,N′-methylenebisacrylamide are neurotoxins. Preparation of solutions containing these compounds should be performed in a well-ventilated fume hood, and extreme precautions taken to minimize contact with solids or solutions. Materials 10% denaturing polyacrylamide gel mix (see recipe or purchase) 10% (w/v) ammonium persulfate (APS) in water TEMED or TMEDA (Life Technologies) 1× TBE (APPENDIX 2A) Ultrapure (e.g., distilled, deionized) water Formamide loading buffer (recipe) 0.2 N NaCl Dialysis tubing, MWCO 1000 (e.g., SpectraPor) 50-mL filter tubes with 0.45-µm cellulose acetate filter (e.g., Spin-X II, Corning Costar) Gel electrophoresis equipment: Vertical (sequencing) gel stand 2000-V power supply Glass plates: 13 × 15.5 cm and 13 × 16.5 cm 2.5-mm-thick gel combs with 2.5-cm-wide wells 2.5-mm-thick spacers UV shadow box or light source Lyophilizer or SpeedVac Saran Wrap (or other UV-transparent plastic wrap) Sterile razor blades Glass stir rod 1. Prepare a vertical, 2.5-mm-thick (preparative), denaturing 10% polyacrylamide gel using a comb producing 2.5-cm-wide wells. The well size is not critical, and the walls can even be removed to create one large well. A gel prepared between plates used for sequencing (13 × 16.5 in.) allows for ample vertical loading of reaction mixture and sufficient migration length for separation and resolution of bands. For these conditions, prepare gel solution from 300 mL of 10% gel mix, 2 mL of 10% APS, and 64 mL TEMED. For more detailed description of procedures for denaturing PAGE, see APPENDIX 3B.
2. Place the gel in a gel electrophoresis apparatus, using 1× TBE in both top and bottom buffer reservoirs, and prerun 30 min at 30 W constant power, room temperature. 3. Redissolve each crude DNA (after deprotection and lyophilization) in 100 µL water and 100 µL formamide loading buffer by brief vortexing. Synthesized DNA after deprotection and lyophilization should be powdery and white and readily redissolve in water. If the solution does not appear homogeneous, mild heating (∼60°C) and/or brief sonication (1 min) can be used to help dissolve the DNA. Insoluble materials may cause some turbidity but are not cause for concern.
4. Load 25 µL of solution from step 3 in each of the 2.5-cm-wide wells; or, alternatively, remove well walls and load the entire sample in the large well created. 5. Run the gel at 30 W constant power, room temperature, to the desired length.
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5.2.21 Current Protocols in Nucleic Acid Chemistry
Bromphenol blue migrates similarly to a 26-nt linear DNA on a 10% gel, and xylene cyanol runs like a 55-nt linear DNA.
6. Disassemble the gel apparatus, separate the plates, and sandwich the gel between two pieces of Saran Wrap. View the gel under ultraviolet light with fluorescent white background (glass silica plates work well) to visualize the DNA. There should be an intense, slow-migrating band, which is the desired sequence, as well as faster-migrating shorter sequences below.
7. Cut out the desired band with a clean, sterile razor blade and place the cut up pieces in three or four 15-mL centrifuge tubes. Using a glass stir rod, crush the gel pieces thoroughly. Combine crushed gel pieces in each tube with ∼5 to 10 mL of 0.2 N NaCl to make an easily shaken slurry. Repeat, in separate tubes, for each band isolated. Shake each slurry for ∼12 hr. Note that high-purity DNAs can be obtained by individually crushing thin vertical portions of the large piece isolated above. Be sure all crushed gel pieces are transferred to the slurry, as isolated yields are often <50%, and it is therefore important to ensure a quantitative transfer of all materials.
8. Filter the slurry by brief centrifuging 1 to 5 min through 50-mL filter tubes. Resuspend the gel pieces in 5 mL water, centrifuge again, and combine the filtrates. It is advisable to save the filtered gel pieces until after quantitation, as additional extractions with 0.2 N NaCl may be desired to increase isolated product yield, although only minimal (<10%) further recoveries should be expected.
9. Transfer the entire reaction volume from step 8 to MWCO 1000 dialysis tubing clamped at one end to make a bag. Secure the other end of the dialysis bag closed. Dialyze against 2 L of ultrapure water for 3 hr. Repeat. 10. Lyophilize the dialysate to dryness. The dried residue should powdery and white.
REAGENTS AND SOLUTIONS Use distilled, deionized water or other ultrapure water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX. Most of these reagents can be purchased premade from a biological supply company.
Denaturing polyacrylamide gel mix, 10% Dissolve 450 g urea, 95.0 g acrylamide, and 5.00 g N,N′-methylenebisacrylamide in 100.0 mL of 10× TBE (APPENDIX 2A) diluted to a final volume of 1.00 L with water. Filter using 0.45-µm nylon filter paper. CAUTION: Acrylamide and N,N′-methylenebisacrylamide are neurotoxins. Preparation of solutions with these compounds should be performed in a well-ventilated fume hood, and extreme precautions taken to minimize contact with compounds or solutions.
Formamide loading buffer Dissolve 10.0 mL of 10 mM EDTA, pH 8.0 (APPENDIX 2A), in 40 mL formamide. Add 5.00 mg each bromophenol blue and xylene cyanol dyes. Chemical and Enzymatic Methods
Stains-all dye solution Dissolve 20.0 mg stains-all (3,3′-diethyl-9-methyl-4,5,4′,5′-debenzothiacarbocyanine; Sigma) in 25.0 mL formamide and dilute to 200 mL with water.
5.2.22 Current Protocols in Nucleic Acid Chemistry
Stop solution Dissolve 5.77 g urea, 1.2 mg each bromophenol blue and xylene cyanol, and 720 µL of a stock (500 mM) EDTA solution in water to a final volume of 12 mL. Final concentrations are 8 M urea, 30 mM EDTA, and 0.1 mg/mL of each gel running dye.
COMMENTARY Background Information Circularized single-stranded deoxyribonucleotides (ssDNAs) are more resistant to nuclease activity than their linear counterparts (Rumney and Kool, 1992), a property that may be important for in vivo studies. Circular DNAs have been investigated for their unique DNA binding properties (Wang and Kool, 1994; Kool, 1996) and also as useful models in studying DNA structures such as hairpin motifs by NMR (Ippel et al., 1995). DNA circles are also currently being used for more diagnostic applications, such as the creation of padlock probes (Nilsson et al., 1994, 1997). Circular oligonucleotide templates have also been used for the synthesis of concatemeric polypeptides (Brown, 1997). Additionally, circular ssDNAs offer a novel means of both DNA and RNA amplification as these molecules are accepted as templates by both DNA and RNA polymerases (Fire and Xu, 1995; Liu et al., 1996; Daubendiek and Kool, 1997). Circular ssDNAs have been constructed using a range of chemical, enzymatic, and reagent-free ligation methods. Chemical ligation using cyanogen bromide has been effective for the synthesis of circles from triple-helical (Ruben et al., 1995) as well as double-helical splint complexes (Dolinnaya et al., 1993). Chemical ligation using carbodiimide chemistry (Ashley and Kushlan, 1991) is becoming a less common method of ligation despite reported higher yields as compared to cyanogen bromide (Dolinnaya et al., 1991), possibly because of the long reaction times required. Enzymatic ligation, which gives comparable yields albeit with higher cost, eliminates the need for either of these chemicals, which may be damaging to DNA. Novel approaches to the synthesis of small circular DNAs are becoming more common as solid-phase methodology improves (De Napoli et al., 1995; Alazzouzi et al., 1997). Of great interest are new autoligation, or reagent-free, methods (Herrlein and Letsinger, 1994; Herrlein et al., 1995), since these eliminate the need for a separate ligation reagent. The finite stability of the tosyl-modified DNAs required for these couplings limits the methods’ practical applications. A recent report
by Xu and Kool (1997) describes a novel 5′-iodonucleoside that eliminates this stability issue. While this method produces a non-natural bridging phosphorothioate, the transcription and replication ability of the DNAs does not appear to be affected, as the circles were transcribed and replicated successfully in a rollingcircle fashion (Y. Xu and E.T. Kool, unpub.observ.). It is noteworthy that this bridging 5′phosphorothioate linkage in DNA is stable for at least several months in the pH range 5 to 9. Each of the three protocols in this unit is potentially applicable to the synthesis of circles ranging in size from 28 nt to at least 188 nt; however, sequence and size may render one or another method more suitable in specific cases. The enzymatic ligation method (see Basic Protocol) has few sequence or size restrictions, though small circles, and larger circles that may have sequence-induced secondary structure, may not be good substrates for the enzyme. The chemical ligation approach using cyanogen bromide (see Alternate Protocol 1) has been very effective in the creation of pyrimidine-rich circles by ligation of triple-helical splint complexes (Ruben et al., 1995). It has not, however, proven effective to the authors’ satisfaction in creating DNA circles from a double-helical splint complex using the methodology described by Dolinnaya et al. (1993). The phosphorothioate autoligation method (see Alternate Protocol 2) requires segment precursors to contain a 5′-thymidine (from iodothymidine) and a 3′-phosphorothioate group at the ligation junction. For larger circles, another issue arises: the tradeoff between optimized ligation yields and DNA synthesis yields. Longer synthesized DNAs will be produced in lower yields than shorter ones, but ligation of shorter oligos may proceed in higher yields than their longer counterparts. Consider the synthesis of a 100-mer ssDNA circle. While an average stepwise coupling yield of 98% on the synthesizer results in 37% overall yield of each 50-mer precursor segment, the same coupling yields give only 14% of the full-length 100-mer precircle. If the single ligation of precircle to circle proceeds in a 40% overall yield, and the double ligation
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5.2.23 Current Protocols in Nucleic Acid Chemistry
only in 30% overall yield, the overall yield of circle from this “convergent” (double-ligation) approach is still 2-fold greater. So in planning strategically for cyclization, one should take into account circle size and optimization of both DNA synthesis and ligation when designing starting materials. Enzymatic ligation proceeds via a 3′-hydroxyl and 5′-phosphate, affording a “natural” phosphodiester backbone. However, this “natural” method comes with an expensive price-tag and, with larger circles (>100 nt), potentially low yields. While the cyanogen bromide method uses the same starting materials for ligation, also yielding the desired “natural” linkage, the reagents for ligation, while substantially cheaper, may be moderately caustic to DNA. So although yields may be comparable to those from enzymatic ligation, the benefit of cheaper reagents may not outweigh the cost of possible product damage. The autoligation method has a cost advantage, since it eliminates the need for a separate ligation reagent, but creates a non-natural phosphorothioate linkage.
Compound Characterization As described in each individual protocol, confirmation of product circularity is quite rapid and simple and may be of critical importance when considering future applications. The analysis of this characterization is quite unambiguous, as described in the sections within each protocol on confirmation of circularity. Initial cleavage of a circle produces a single band with the mobility of the full-length cyclization precursor (Fig. 5.2.5). If the putative circular DNA is actually linear, banding will be seen directly under the suspected band. This type of banding should also be seen for the precircle reaction lane. This reaction, and/or one analogous to it, must be completed to ensure product quality.
Critical Parameters
Chemical and Enzymatic Methods
There are few complicating factors with these reactions, although several potential problems will be addressed in the troubleshooting section. However, there are a few “critical parameters” worth mentioning. For all syntheses, dialysis of the final ligation-reaction mixture to remove all salts prior to lyophilization and gel purification is critical, as excess salts can cause problems with gel migration. It is worthwhile to note that for all reactions done via a double ligation (i.e., with two precursor segments), the concentration of precircle in the
second step is purposely lowered to favor this intramolecular ligation producing circles as compared to the intermolecular ligation creating linear multimers. Another point to note is that while both enzymatic and autoligation occur via the same double-helical splint complex, only the enzymatic ligation protocol calls for slow annealing of precursor DNAs and splints prior to addition of ligation reagents (steps 6 and 10 of the Basic Protocol). This simply represents different methodologies consistently used in the authors’ lab, and while enzymatic ligation may proceed just as efficiently without these steps, this asyet-unexplored option has not been described in the protocol. For enzymatic ligation, the enzyme should be stored at −20°C prior to its use, and reaction temperature should be maintained consistently at ≤37°C, as higher temperatures may cause enzyme denaturation and inactivation. The only special precaution when using cyanogen bromide ligation is that, since this chemical is quite toxic and volatile, appropriate care must be taken when using it. By much the same token, the reaction should be stopped promptly when instructed, as prolonged exposure may increase the DNA’s susceptibility to damage. The only specific considerations for the phosphorothioate method are in the synthesis of the modified precursor segments. It is important to follow the directions for synthesis exactly, first phosphorylating the initial base column with subsequent addition of the desired 3′ base and sulfurization of only this internucleotide linkage. As for the 5′-iodothymidine phosphoramidite, this particular compound is unique—compared to other similar compounds designed to ligate ssDNAs via an SN2 mechanism—in having greatly enhanced stability, which significantly extends the shelf lives of both the phosphoramidite and the modified precursor-segment DNAs (Xu and Kool, 1997). For a discussion of common problems arising with these procedures, including their diagnosis and possible solutions, see Table 5.2.1.
Anticipated Results For all three methods, optimized yields after purification are modest, and often less than the observed qualitative conversion. For circles of ∼65 nt, yields of 33% for enzymatic ligation (Diegelman, pers. comm.) and 20% for autoligation (Xu et al., 1997) have been reported along with a comparable yield of 30% for
5.2.24 Current Protocols in Nucleic Acid Chemistry
Table 5.2.1 Troubleshooting Guide for Synthesis of Circular DNAs
Problem
Possible cause
Possible solution
Poor DNA synthesis yields
Reagents contaminated with water
Ensure reagents are anhydrous (distill solvents fresh) Boil 15 min in water and sonicate 30 min in methanol Use new phosphoramidites Distill fresh CH3CN to prepare phosphorylating reagent Purchase fresh reagents Briefly heat at 60°C and sonicate to dissolve DNA or briefly centrifuge crude DNA and work with supernatant Set absorbance at 260 nm; check that UV lamp is working properly Ensure cuvette is clean and unscratched and light passes through its clear sides Ensure instrument is properly zeroed with a cuvette containing only water Consult spectrophotometer manual on ensuring unimpeded light path See “Crude DNA not dissolving in water,” above If quantitating gel-purified products, try re-extracting the slurry mixture with 0.2 N NaCl Rerun reaction, ensuring addition of all DNAs needed Rerun reaction using a fresh batch of enzyme, and ensure proper storage (–20°C) after use Rerun reaction using fresh DTT or BrCN stock Rerun reaction for shorter period
Glass frits in synthesizer dirty Phosphoramidites degraded Poor phosphorylation yields Reagents contaminated with water
Crude DNA not dissolving in water
Phosphoramidites degraded Insoluble material found after deprotection
No/low absorbance seen for Measurement not at correct wavelength DNA quantitation Cuvette dirty or not inserted properly Instrument not properly zeroed Instrument not properly aligned No DNA in sample DNA concentration low after isolation of products from gel pieces No ligation seen by UV shadowing
One or more DNAs (especially splint) omitted from reaction mixture Enzyme old or inactive (enzymatic ligation) DTT or BrCN degraded (enzymatic ligation) Reaction allowed to proceed too long (chemical ligation) DNA modification (phosphorylation) poor, leading to reduced ligation yield (enzymatic and chemical ligation) Poor incorporation of the modified base, 5′-iodothymidine (autoligation)
Poor splint hybridization
Isolated product not behaving as expected for circular DNA
Isolated product is not a circular DNA
Isolated product is circular but has degraded significantly (less likely; check for circularity first)a
See “Poor phosphorylation yields,” above
Ensure coupling of modified phosphoramidite and sulfurization were effectiveb; rerun reaction, ensuring DNA synthesizer cycle is set up properly Rerun reaction; possibly redesign new splint junction (sequence context problem may exist) Rerun Sl nuclease reaction on other isolated bands Resynthesize DNA, taking greater precautions to avoid degradation
aCommon forms of damage are formation of abasic sites from depurination during synthesis, and degradation by reagents (e.g., BrCN) or heat or acid
treatments. bSulfurization can be examined by electrospray or MALDI-TOF MS (UNITS 10.1 & 10.2). Iodothymidine incorporation can be checked by comparing the
oligonucleotide to one without iodo-T added using either high-resolution sequencing PAGE (APPENDIX 3B) or C18-reversed-phase HPLC.
5.2.25 Current Protocols in Nucleic Acid Chemistry
Supplement 9
cyanogen bromide ligation of a 42-nt triplexforming circle (Ruben et al., 1995). For all syntheses involving more than one precursor segment and splint, and methodology involving a double ligation (two step, one pot) rather than a dimerization approach (triple-helical splintcomplex ligation with cyanogen bromide), intermediate, non-unit-length products (besides the desired circles) are often seen. In general, the observed yield—as determined by UV shadowing to estimate product conversion—is often twice the isolated yield. This apparently represents the inability to completely extract product from gel slurry; reextraction with additional eluent (0.2 N NaCl) may increase product yields somewhat, although only minimal (<10%) recoveries should be expected.
Time Considerations Completion of any of the above three protocols should take no longer than 7 days, and most can be completed in less time either by purchasing synthesized DNAs or by using fast-deprotection phosphoramidites. Actual hands-on work occupies much less total time, but several waiting periods are required. Synthesis of oligos, with overnight deprotection, followed by lyophilization and quantitation the next day should take ∼1.5 days. A double ligation set up to run the remainder of day 2 with the second ligation running overnight should take another 1.5 days, resulting in completion of the ligations by the middle of day 3. Dialysis for 6 hr and lyophilization overnight allows gel purification on day 4 and elution from gel pieces overnight to the morning of day 5. Following a 24-hr dialysis, the circular products should be ready partway through day 6 for quantitation and structural confirmation experiments.
Literature Cited Alazzouzi, E., Escaja, N., Grandas, A., and Pedroso, E. 1997. A straightforward solid-phase synthesis of cyclic oligodeoxyribonucleotides. Angew. Chem. Int. Ed. Engl. 36:1506-1508. Ashley, G.W. and Kushlan, D.M. 1991. Chemical synthesis of oligodeoxynucleotide dumbbells. Biochemistry 30:2927-2933.
Chemical and Enzymatic Methods
Borer, P.N. 1975. Optical properties of nucleic acids. In Handbook of Biochemistry and Molecular Biology, Vol. I, 3rd ed. (G.D. Fasman, ed.) p. 589. CRC Press, Boca Raton, Fla. Brown, S. 1997. Metal-recognition by repeating polypeptides. Nature Biotechnol. 15:269-272.
Daubendiek, S.L. and Kool, E.T. 1997. Generation of catalytic RNAs by rolling transcription of synthetic DNA nanocircles. Nature Biotechnol. 15:273-277. De Napoli, L., Galeone, A., Mayol, L., Messere, A., Montesarchio, D., and Piccialli, G. 1995. Automatic solid phase synthesis of cyclic oligonucleotides: A further improvement. Bioorg. Med. Chem. 3:1325-1329. Dolinnaya, N.G., Sokolova, N.I., Ashirbekova, D.T., and Shabarova, Z.A. 1991. The use of BrCN for assembling modified DNA duplexes and DNARNA hybrids: Comparison with water-soluble carbodiimide. Nucl. Acids Res. 9:3067-3072. Dolinnaya, N.G., Blumenfeld, M., Merenkova, I.N., O re tsk a y a, T.S., Kryn etsk aya, N.F., Ivanovskaya, M.G., Vasseur, M., and Shabarova, Z.A. 1993. Oligonucleotide circularization by splint-directed chemical ligation. Nucl. Acids Res. 21:5403-5407. Fire, A. and Xu, S.Q. 1995. Rolling replication of short DNA circles. Proc. Natl. Acad. Sci. U.S.A. 92:4641-4645. Herrlein, M.K. and Letsinger, R.L. 1994. Selective chemical autoligation on a double-stranded DNA splint. Nucl. Acids Res. 22:5076-5078. Herrlein, M.K., Nelson, J.S., and Letsinger, R.L. 1995. A covalent lock for self-assembled oligonucleotide conjugates. J. Am. Chem. Soc. 117:10151-10152. Ippel, J.H., Lanzotti, V., Galeone, A., Mayol, L., Van den Boogaart J.E., Pikkemaat, J.A., and Altona, C. 1995. Slow conformational exchange in DNA minihairpin loops: A conformational study of the circular dumbbell d
. Biopolymers 36:681-694. Kool, E.T. 1996. Circular oligonucleotides: New concepts in oligonucleotide design. Annu. Rev. Biophys. Biomol. Struct. 25:1-28. Liu, D., Daubendiek, S.L., Zillman, M.A., Ryan, K., and Kool, E.T. 1996. Rolling circle DNA synthesis: Small circular oligonucleotides as efficient templates for DNA polymerases. J. Am. Chem. Soc. 118:1587-1594. Nilsson, M., Malmgren, H., Samiotaki, M., Kwiatkowski, M., Chowdhary, B.P., and Landegren, U. 1994. Padlock probes: Circularizing oligonucleotides for localized DNA detection. Science 265:2085-2088. Nilsson, M., Krejci, K., Koch, J., Kwiatkowski, M., Gustavsson, P., and Landegren, U. 1997. Padlock probes reveal single-nucleotide differences, parent of origin and in situ distribution of centromeric sequences in human chromosomes 13 and 21. Nature Genet. 16:252255. Ruben, E., Rumney, S. IV, Wang, S., and Kool, E.T. 1995. Convergent DNA synthesis: A non-enzymatic dimerization approach to circular oligodeoxynucleotides. Nucl. Acids Res. 23:3547-3553.
5.2.26 Supplement 9
Current Protocols in Nucleic Acid Chemistry
Rumney, S. IV and Kool, E.T. 1992. DNA recognition by hybrid oligoether-oligodeoxynucleotide macrocycles. Angew. Chem. Int. Ed. Engl. 31:1617-1619. Serwer, P. and Allen, J.L. 1984. Conformation of double-stranded DNA during agarose gel electrophoresis: Fractionation of linear and circular molecules with molecular weights between 3 × 106 and 26 × 106. Biochemistry 23:922-927. Wang, S. and Kool, E.T. 1994. Circular RNA oligonucleotides. Synthesis, nucleic acid binding properties, and a comparison with circular DNAs. Nucl. Acids Res. 22:2326-2333. Xu, Y. and Kool, E.T. 1997. A novel 5′-iodonucleoside allows efficient nonenzymatic ligation of single-stranded and duplex DNAs. Tetrahedron Lett. 38:5595-5598.
Dolinnaya et al., 1993. See above. Reports the synthesis of medium-sized (∼40 nt) ssDNA circles from a double-helical splint complex with cyanogen bromide. Certain sequence requirements for successful ligation are described. Ruben et al., 1995. See above. Reports the cyanogen bromide–mediated synthesis of medium-sized (34- to 72-nt) ssDNA circles from two short segments using a triple-helical splint complex. Xu and Kool, 1997. See above. Reports an autoligation method incorporating a novel 5′-iodothymidine phosphoramidite (now commercially available) which possesses increased stability as compared to one previously described (Herrlein et al., 1995).
Key References Alazzouzi et al., 1997. See above. Reports a unique methodology for the solid-phase synthesis of small (<32 nt) ssDNA circles. No splint is needed, but yields are low for the larger circles in this size range.
Contributed by Amy M. Diegelman and Eric T. Kool University of Rochester Rochester, New York
Methods for Cross-Linking Nucleic Acids
5.2.27 Current Protocols in Nucleic Acid Chemistry
Engineering Specific Cross-Links in Nucleic Acids Using Glycol Linkers
UNIT 5.3
Simple glycol linkers can be used to cross-link nucleic acid sequences. In the most straightforward approach, such cross-links can be used in place of nucleotide sequences to bridge two domains of higher-order nucleic acid structures. Such linkers can also be viewed as tethers between two independently hybridizing nucleic acid sequences, or between a nucleic acid and some other ligand or reporter group. Although most any carbon chain can be employed to introduce cross-links in nucleic acids, the hydrophilic nature of the ethylene glycol chain gives it one particular advantage. Whereas simple carbon chains may tend to collapse on themselves as the result of the hydrophobic effect, the glycol chains’ alternating ethyl and oxygen ether subunits are more likely to be hydrated in aqueous solutions and thus maintain a more extended conformation, which permits them to easily bridge two different sites within the macromolecule. Additionally, a variety of ethylene glycol–based linkers are readily available (Fig. 5.3.1) and only require simple protection reactions in order to be used as cross-linking agents. Oligo(ethylene glycol) linkers have been used most commonly to replace a portion (Williams and Hall, 1996) or the entirety of the loop structure at the end of DNA (Durand et al., 1990; Altmann et al., 1995) or RNA helices (Benseler et al., 1993; Ma et al., 1993; Thomson et al., 1993; Fu et al., 1994; Hendry et al., 1994; Komatsu et al., 1996), essentially to achieve cross-linking of the terminal residues of the double-stranded helix. However, in some cases ethylene glycol linkers have been used to tether different strands of nucleic acids (Cload and Schepartz, 1991; Amaratunga and Lohman, 1993; Moses and Schepartz, 1996) or even to tether minor groove-binding ligands to the nucleic acid (Robles et al., 1996; Rajur et al., 1997; Robles and McLaughlin, 1997). In most cases, the glycol linker is incorporated as part of the nucleic acid backbone, such that at each terminus the linker is incorporated into a phosphodiester linkage that also incorporates either the 3′ or 5′ hydroxyl of the adjacent nucleoside residue. It is also possible to incorporate more than a single linker at the same site. Thus, two residues of tri(ethylene glycol) could be used instead of hexa(ethylene glycol) (Benseler et al., 1993; Fu et al., 1994)—in the former case a negatively charged phosphodiester would bridge the two linkers. This approach can be used to generate structures with varying linker lengths via the preparation of only a single linker building block. In the most common protocol, the linker is protected at one terminus as the 4,4′-dimethoxytrityl derivative (see Basic Protocol 1), and is converted to a phosphoramidite at the second terminus (see Basic Protocol 2). With such derivatives, the linker is simply incorporated into the DNA or RNA sequence by the same procedures as are used for common nucleoside phosphoramidites (see Basic Protocol 3). Preparation of the pro-
O O O O O O
Figure 5.3.1
O O O O O O
O O O O O
O O O O
O O O
O O
O
Varying lengths for readily available ethylene glycol–based linkers.
Contributed by Timothy O’Dea and Larry W. McLaughlin Current Protocols in Nucleic Acid Chemistry (2000) 5.3.1-5.3.8 Copyright © 2000 by John Wiley & Sons, Inc.
Methods for Cross-Linking Nucleic Acids
5.3.1
tected linker-phosphoramidites follows a common procedure regardless of length; protocols for the hexa(ethylene glycol) linker are presented here. BASIC PROTOCOL 1
PROTECTION OF THE GLYCOL CHAIN WITH A TRITYL GROUP The following protocol outlines the protection of one terminus of an ethylene glycol chain with a trityl group. The first reaction, illustrated in Figure 5.3.2, promotes monoprotection of the ethylene glycol chain with 4,4′-dimethoxytrityl chloride. Although the specific protocol for hexa(ethylene glycol) follows, this protocol has also been successful with glycol chains of various lengths: 1,3-propanediol, tri(ethylene glycol), the tetra- and penta-compounds, and so on. The monoprotected ethylene glycol product can be purified by silica-gel column chromatography. Materials Hexa(ethylene glycol) (HEG) Anhydrous pyridine (preferably freshly distilled) Nitrogen or argon gas 4,4′-Dimethoxytrityl chloride (DMT-Cl) 5% (v/v) methanol in dichloromethane 10% (v/v) aqueous sulfuric acid (H2SO4; Table A.2A.1) Triethylamine (Et3N, TEA) Dichloromethane (CH2Cl2, DCM; preferably freshly distilled) 5% (w/v) aqueous sodium hydrogen carbonate (NaHCO3) Sodium sulfate (Na2SO4) Methanol (CH3OH, MeOH) HO
O
O
O
O
OH
O
OCH3
O
O
O
O
O
OH
O
OCH3 OCH3
O
O
O
O
O
O
O
P OCH2CH2CN N(i-Pr)2
OCH3
O O
O
O
Engineering Specific Cross-Links in Nucleic Acids Using Glycol Linkers
G
G
C
C
C
C
G
G
O
O O
Figure 5.3.2 Reaction pathway for the preparation of a glycol linker and a sample nucleic acid sequence containing the linker.
5.3.2 Current Protocols in Nucleic Acid Chemistry
Non-acid-generating desiccant: e.g., sodium hydroxide or calcium carbonate 100-mL round-bottom flask and rubber stopper Device for maintaining nitrogen or argon atmosphere (e.g., balloon, syringe, and rubber stopper; see step 2) Needle and syringe Separatory funnel Silica gel Column for chromatography Rotary evaporator Thin-layer chromatography (TLC) apparatus (see APPENDIX 3D) CAUTION: Pyridine and its vapors are toxic; exposure to pyridine must be minimal. The reaction should be performed in a fume hood. Monoprotect ethylene glycol 1. Coevaporate 1.25 g (5 eq, 4.43 mmol) HEG twice with ∼10 mL anhydrous pyridine in a 100-mL flask. 2. Under an anhydrous nitrogen or argon atmosphere, add 10 mL anhydrous pyridine and a dry stir bar, and seal the 100 mL-flask with a rubber stopper. The easiest means to create a nitrogen or argon atmosphere is via a balloon sealed to a syringe with a needle. To construct: Remove plunger from syringe, cut off the now opened end, slip a balloon onto this end, and seal well with parafilm. Fill balloon with gas, attach needle, and punch needle through rubber stopper.
3. Begin stirring at ambient temperature. 4. In a separate flask under nitrogen, dissolve 300 mg (1 eq, 0.885 mmol) DMT-Cl in ∼3 mL anhydrous pyridine. 5. Using a syringe, puncture the rubber stopper and gradually add the DMT-Cl solution to the reaction flask. Useful increments are 0.5 mL every 5 min over a 30-min period. The reaction can be monitored by TLC (silica gel, 60 Å, see APPENDIX 3D) using 5% methanol in DCM as eluant. The Rf is 0.45. The product is visible under UV and turns orange when reacted with 10% aqueous H2SO4.
6. After 2 hr, add 2 mL TEA and dilute with ∼25 mL DCM. TEA neutralizes the acid that has been generated, which otherwise will cleave the mono-DMT derivative of the ethylene glycol linker.
7. Extract the organic layer twice with 5% NaHCO3 (∼40 mL) and once with distilled water (∼40 mL) using a separatory funnel. 8. Dry the organic layer over Na2SO4 and remove solvent with a rotary evaporator. The product remains as a clear or slightly colored oil.
Purify mono-DMT–ethylene glycol product 9. Pack a silica-gel column (~15 g, roughly 10× expected solute amount), using 0.5% TEA in DCM as eluant. Again, TEA reduces the acidic nature of the silica gel, thus reducing decomposition of the mono-DMT–ethylene glycol during chromatography.
10. Dissolve the mono-DMT–ethylene glycol product (from step 8) in a minimum quantity of DCM/TEA and pour onto the column. Elute with at least 400 mL of 0.5%
Methods for Cross-Linking Nucleic Acids
5.3.3 Current Protocols in Nucleic Acid Chemistry
TEA in DCM, followed by a step gradient using 400-mL aliquots of 0.5% TEA/DCM containing from 0.5% to 3% MeOH. The product will elute in <3% MeOH.
11. Test fractions by TLC (APPENDIX 3D; Rf = 0.45) using 5% MeOH in DCM as the eluant. 12. Combine fractions containing the correct product and remove solvent by rotary evaporation (high vacuum is needed to remove excess TEA in product). 13. Store in a sealed vial at ambient temperature over a desiccant. The 4,4′-dimethoxytrityl-protected hexa(ethylene glycol) product (DMT-HEG) is stable for several months with minimal decomposition provided it is not stored over a desiccant that liberates acid (e.g., P2O5). BASIC PROTOCOL 2
PHOSPHITYLATION OF THE MONOPROTECTED GLYCOL LINKER The following protocol details the phosphitylation of a 4,4′-dimethoxytrityl-protected glycol linker with 2-(cyanoethyl)-N,N-diisopropylchlorophosphoramidite. For an efficient reaction with high yield, conditions must be kept scrupulously anhydrous. While the following procedure outlines the use of a monoprotected hexa(ethylene glycol) linker, the protocol has been successful with monoprotected glycol compounds of various lengths. The reaction is illustrated in Figure 5.3.2. Materials 4,4′-Dimethoxytrityl-protected hexa(ethylene glycol) (DMT-HEG; see Basic Protocol 1) Anhydrous pyridine (preferably freshly distilled; UNIT 3.2) Non-acid-generating desiccant: e.g., sodium hydroxide or calcium carbonate Nitrogen or argon gas Anhydrous dichloromethane (CH 2Cl2, DCM; preferably freshly distilled) Diisopropylethylamine 2-(Cyanoethyl)-N,N-diisopropylchlorophosphoramidite Ethyl acetate 10% (v/v) triethylamine (Et3N, TEA) in ethyl acetate 5% (w/v) aqueous NaHCO3 Saturated aqueous NaCl Sodium sulfate (Na2SO4) 25-mL round-bottom flask and rubber stopper CAUTION: Pyridine and its vapors are toxic; exposure to pyridine must be minimal. The reaction should be performed in a fume hood. Phosphitylate DMT–ethylene glycol 1. Coevaporate 300 mg (1 eq, 0.51 mmol) of DMT-HEG twice with ∼10 mL anhydrous pyridine. Place under high vacuum over a non-acid-generating desiccant and leave overnight. 2. In a rubber-stoppered 25-mL round-bottom flask with a dry stir bar under an anhydrous nitrogen or argon atmosphere, dissolve the DMT-HEG in 1 mL anhydrous DCM and 0.22 mL (3 eq, 1.54 mmol, 157 mg) anhydrous diisopropylethylamine.
Engineering Specific Cross-Links in Nucleic Acids Using Glycol Linkers
A balloon sealed to a syringe provides an easy means to create a nitrogen or argon atmosphere (see Basic Protocol 1, step 2, for details). TEA may be used here as an alternative to diisopropylamine, if preferred.
5.3.4 Current Protocols in Nucleic Acid Chemistry
3. While stirring, add 0.115 mL (1 eq, 0.51 mmol, 121 mg) 2-(cyanoethyl)-N,N-diisopropylchlorophosphoramidite to the reaction flask using a syringe. The reaction can be monitored via TLC using 9:1 (v/v) ethyl acetate/TEA as the eluant. The Rf of DMT-HEG is 0.50 and that of DMT-HEG-phosphoramidite is 0.80. The product is visible under UV and turns orange when treated with 10% H2SO4.
4. After 25 min, dilute reaction with ∼20 mL ethyl acetate. 5. Extract the organic layer twice with 5% aqueous NaHCO3 and once with saturated aqueous NaCl. 6. Filter organic layer over Na2SO4 and evaporate solvent with rotary evaporator. Purify DMT–ethylene glycol–phosporamidite 7. Pack a silica-gel TLC column with 1% (v/v) TEA in ethyl acetate. 8. Elute the product with increasing percentages of TEA (1% to 5%) in ethyl acetate. 9. Test fractions by TLC (Rf = 0.80) using 10% TEA in ethyl acetate as the eluant. 10. Combine fractions containing the correct product and remove solvent using rotary evaporator (high vacuum is needed to remove the TEA). 11. Store in a sealed vial at −20°C The DMT-HEG-P will remain stable for several weeks.
PREPARATION OF ETHYLENE GLYCOL LINKERS FOR INCORPORATION INTO OLIGONUCLEOTIDES
BASIC PROTOCOL 3
The DMT-protected and phosphitylated glycol linkers can be inserted into DNA sequences using standard automated phosphoramidite synthesis. Since the glycol linker is an oil, several preparative steps facilitate its incorporation using an automated synthesizer. For an overview of oligonucleotide synthesis, see APPENDIX 3C. Materials Dimethoxytrityl-protected hexa(ethylene glycol) phosphoramidite (DMT-HEG-P) (see Basic Protocol 2) Anhydrous dichloromethane (CH2Cl2, DCM; preferably freshly distilled) Anhydrous acetonitrile (preferably freshly distilled) Bottle from DNA synthesizer, tared 1. Dissolve 266 mg DMT-HEG-P (0.34 mmol) in 1.00 mL anhydrous CH2Cl2 under an anhydrous nitrogen or argon atmosphere. A balloon sealed to a syringe provides an easy means to create a nitrogen or argon atmosphere (see Basic Protocol 1, step 2, for details).
2. Using a syringe, transfer 0.100 mL DMT-HEG-P solution to a suitable tared DNA synthesizer bottle. 3. Remove solvent on rotary evaporator and dry under high vacuum overnight. 4. Weigh DNA synthesis bottle to determine exact amount of DMT-HEG-P. 5. Dissolve 24 mg DMT-HEG-P (∼30 µmol) in 250 µL anhydrous acetonitrile. Care must be taken to ensure DMT-HEG-P is dissolved completely.
Methods for Cross-Linking Nucleic Acids
5.3.5 Current Protocols in Nucleic Acid Chemistry
6. Place the bottle on the automated DNA synthesizer and purge as recommended by the manufacturer. 7. The ethylene glycol linker can now be incorporated into oligonucleotides by solidsupport synthesis using standard phosphoramidite protocols. The oligonucleotide can be synthesized with the DMT group on and then purified by HPLC analysis, or with the DMT group off and then purified by gel electrophoresis. If poor coupling occurs, see Critical Parameters for possible solutions.
COMMENTARY Background Information
Engineering Specific Cross-Links in Nucleic Acids Using Glycol Linkers
The cross-linking agents described in this unit are those that are employed during the assembly of the DNA sequences—in this respect they are introduced at very specific sites. Other simple carbon-based linkers can also be employed in a similar manner, but as noted earlier, simple carbon chains may tend to collapse on themselves in an aqueous environment, while the glycol chains are more likely to be hydrated and thus maintain a more extended conformation. Other approaches to cross-linking are also available, most notably the introduction of thiol-based linkers, which upon oxidation form a disulfide cross-link between two sites within a higher-order nucleic acid complex (Ferentz and Verdine, 1991; Wolfe and Verdine, 1993; Goodwin and Glick, 1994; Cain and Glick, 1998; UNITS 5.1 & 5.4). It has been difficult to design effective protocols to confirm the presence of the linker within the nucleic acid sequence. With other types of modified sequences, DNA digests can often be used to confirm the presence of the modification. In the present case, the linkers are not easily identifiable, and such digests only confirm the presence of the nucleoside components. However, when the linker is present in RNA, it is possible to treat a small quantity of the nucleic acid fragment with T2 RNase and an alkaline phosphatase, which results in cleavage of all linkages save that between the linker and the 5′ terminus of the nucleotide (there is no requisite 2′-OH at this linkage). The resulting nucleoside attached to the linker can then be identified after the appropriate standard is prepared (Fu et al., 1994). Careful analysis of the digest by HPLC, with the use of an appropriate standard, can confirm the presence of the linker in the sequence of interest (Fu et al., 1994). However, this procedure can be tedious, and requires the preparation of the necessary standard(s). Recent work in the authors’ lab (D.J. Fu, G. Xiang, and L. McLaughlin, unpub. observ.) has indicated that MALDI-
TOF (UNIT 10.1) analyses of such nucleic acid analogues are much simpler and are effective in providing evidence for the presence of the linker in both DNA and RNA target sequences.
Critical Parameters The synthesis of these linkers should not present significant problems for anyone with even a moderate level of laboratory experience. As can be noted with the protocol, the glycol linkers tend to be quite inexpensive and are used in excess over the DMT-Cl reagent to ensure that only the mono-protected product results. These reactions can be performed with stoichiometries of 1:1, but usually some of the bis-protected DMT-linker results. This unwanted product can be removed during the purification step. For these protocols to succeed, the reactions must be performed under anhydrous conditions. Because of DMT-Cl’s sensitivity to water, great care must be taken to maintain a dry environment for the reaction. All ethylene glycol reagents should be coevaporated with pyridine and kept under vacuum before use. A nitrogen or argon atmosphere for the reaction helps maintain the anhydrous conditions while the reaction is being run. The simplest apparatus is a balloon fixed to a syringe body and filled with dry argon/nitrogen. The reaction flask is sealed with a rubber septum and a needle affixed to the syringe is pushed through the septum. This simple apparatus keeps the reaction mixture under a slight positive pressure with an anhydrous inert gas. High yields require that one be aware of the acid lability of the DMT-protecting group. To limit decomposition as a result of trace quantities of acid, a small amount of the organic base triethylamine (TEA) is added during work-up. It is also critical to have some TEA present (∼0.5%) during the chromatographic purification step employing a silica-gel column. TEA neutralizes the slight acidity of the silicic acid, promoting greater stability of the product when
5.3.6 Current Protocols in Nucleic Acid Chemistry
it is adsorbed on the column support. The ∼0.5% TEA does not significantly alter chromatographic mobility, and its presence results in a greater overall yield of recovered product. For the phosphitylation protocol, taking into consideration the lability of the phosphoramidite is critical for a successful experiment. Once synthesized, effort must be taken to minimize the exposure of the phosphoramidite to air and acidic compounds of any kind. Both the reagent and the isolated product should be stored in parafilm-sealed vials at −20°C. When the phosphoramidite reagent loses its pale-yellow color and becomes a deeper yellowish-orange color, the reagent has typically degraded and should not be used. Preparation of the linker for use with a DNA synthesizer is only complicated by the fact that it is an oil rather than a solid. The authors have found the simplest procedure is one in which some of the oil is transferred to a suitable flask, weighed, and dissolved in sufficient anhydrous solvent. From this solution, an aliquot corresponding to ∼30 µmol of linker (per coupling) is transferred to the DNA synthesis bottle. The solvent is then removed from the bottle under vacuum and the residue is kept under high vacuum overnight. The requisite amount of acetonitrile can then be added to the bottle before the latter is attached to the DNA synthesis machine—ensure that the residue in the bottle completely dissolves first. To obtain efficient coupling of the linker to a DNA strand, the synthesizer programming need not to be altered. However, if efficient coupling is not achieved, several parameters can be changed to attain better coupling. First, extended “wait” periods can be added to the cycle—these are typically the time periods during which the coupling reaction takes place. A second option is to perform two coupling steps in sequence without any intervening capping or oxidation steps.
Anticipated Results Yields for the protection of the glycol linker with DMT should be >70%. When isolated by column chromatography the DMT-ethylene glycol product is a pale-orange oil with Rf = 0.45 (5% MeOH in DCM). The yields expected for the phosphitylation protocol should be >70% when isolated from the column as a clear oil (Rf = 0.8, 1:9 TEA/ethyl acetate). Successful phosphitylation can be achieved without the column chromatography step. In this case, simply perform the aqueous work-up, dry the solution, and evaporate to an oil. 31P NMR will
confirm the ratio of the phosphitylated product to any phosphorus contaminants. So long as the latter are minimal in quantity, effective incorporation of the linker can be obtained with material prepared in this manner.
Time Considerations Monoprotection of ethylene glycol and its isolation can be accomplished in <5 hr. The phosphitylation protocol can be done in <2 hr when the DMT-ethylene glycol product is prepared ahead of time (see Basic Protocol 1). Incorporation of the ethylene glycol linker into the oligonucleotide will not require more than a half-hour beyond the normal coupling time required of a standard phosphoramidite.
LITERATURE CITED Altmann, S., Labhardt, A.M., Bur, D., Lehmann, C., Bannwarth, W., Billeter, M., Wuthrich, K., and Leupin, W. 1995. NMR studies of DNA duplexes singly cross-linked by different synthetic linkers. Nucl. Acids Res. 23:4827-4835. Amaratunga, M. and Lohman, T.M. 1993. Escherichia coli Rep helicase unwinds DNA by an active mechanism. Biochemistry 32:6815-6820. Benseler, F., Fu, D.J., Ludwig, J., and McLaughlin, L.W. 1993. Hammerhead-like molecules containing non-nucleoside linkers are active RNA catalysts. J. Am. Chem. Soc. 115:8483-8484. Cain, R.J. and Glick, G.D. 1998. Use of cross-links to study the conformational dynamics of triplex DNA. Biochemistry 37:1456-1464. Cload, S.T. and Schepartz, A. 1991. Polyether tethered oligonucleotide probes. J. Am. Chem. Soc. 113:6324-6326. Durand, M., Chevrie, K., Chassignol, M., and Thuong, N.T. 1990. Circular dichroism studies of an oligodeoxyribonucleotide containing a hairpin loop made of a hexaethylene glycol chain—conformation and stability. Nucl. Acids Res. 18:6353-6359. Ferentz, A.E. and Verdine, G.L. 1991. Disulfide cross-linked oligonucleotides. J. Am. Chem. Soc. 113:4000-4002. Fu, D.J., Benseler, F., and McLaughlin, L.W. 1994. Hammerhead ribozymes containing non-nucleoside linkers are active RNA catalysts. J. Am. Chem. Soc. 116:4591-4598. Goodwin, J.T. and Glick, G.D. 1994. Synthesis of a disulfide stabilized RNA hairpin. Tetrahedron Lett. 35:1647-1650. Hendry, P., Moghaddam, M.J., McCall, M.J., Jennings, P.A., Ebel, S., and Brown, T. 1994. Using linkers to investigate the spatial separation of the conserved nucleotides A9 and G12 in the hammerhead ribozyme. Biochim. Biophys. Acta 1219:405-412. Komatsu, Y., Kanzaki, I., and Ohtsuka, E. 1996. Enhanced folding of hairpin ribozymes with replaced domains. Biochemistry 35:9815-9820.
Methods for Cross-Linking Nucleic Acids
5.3.7 Current Protocols in Nucleic Acid Chemistry
Ma, M.Y.X., McCallum, K., Climie, S.C., Kuperman, R., Lin, W.C., Sumner-Smith, M., and Barnett, R.W. 1993. Design and synthesis of RNA miniduplexes via a synthetic linker approach. 2. Generation of covalently closed, double-stranded cyclic HIV-1 TAR RNA analogs with high Tat-binding affinity. Nucl. Acids Res. 21:2585-9. Moses, A.C., and Schepartz, A. 1996. Triplex tethered oligonucleotide probes. J. Am. Chem. Soc. 118:10896-10897. Rajur, S.B., Robles, J., Wiederholt, K., Kuimelis, R.W., and McLaughlin, L.W. 1997. Hoechst 33258 tethered by a hexa(ethylene glycol) linker to the 5′-termini of oligodeoxynucleotide 15mers: Duplex stabilization and fluorescence properties. J. Org. Chem. 62:523-529. Robles, J. and McLaughlin, L.W. 1997. DNA triplex stabilization using a tethered minor-groove binding Hoechst 33258 analogue. J. Am. Chem. Soc. 119:6014-6021.
Robles, J., Rajur, S.B., and McLaughlin, L.W. 1996. A parallel-stranded DNA triplex tethering a Hoechst 33258 analogue results in complex stabilization by simultaneous major groove and minor groove binding. J. Am. Chem. Soc. 118:58205821. Thomson, J.B., Tuschl, T., and Eckstein, F. 1993. Activity of hammerhead ribozymes containing non-nucleotidic linkers. Nucl. Acids Res. 21:5600-5603. Williams, D.J. and Hall, K.B. 1996. Thermodynamic comparison of the salt dependence of natural RNA hairpins and RNA hairpins with non-nucleotide spacers. Biochemistry 35:14665-14670. Wolfe, S.A. and Verdine, G.L. 1993. Ratcheting torsional stress in duplex DNA. J. Am. Chem. Soc. 115:12585-12586.
Contributed by Timothy O’Dea and Larry W. McLaughlin Boston College Chestnut Hill, Massachusetts
Engineering Specific Cross-Links in Nucleic Acids Using Glycol Linkers
5.3.8 Current Protocols in Nucleic Acid Chemistry
Engineering Disulfide Cross-Links in RNA Via Air Oxidation
UNIT 5.4
This unit describes methods for the synthesis of thiol-modified ribonucleosides, their incorporation into synthetic RNA, and the formation of intramolecular disulfide bonds in RNA by air oxidation. The disulfide bonds can be formed in quantitative yields between thiols positioned in close proximity in either RNA secondary or tertiary structure. Disulfide cross-links are useful tools to probe solution structures of RNA, monitor dynamic motions, and stabilize folded RNAs. Several steps are involved in the successful formation of a disulfide-cross-linked RNA. First, a location for incorporation of the disulfide bond is selected (Glick, 1998). For best results, this should be based on the highest-resolution structural data available for the particular RNA. In most cases, it is desirable to avoid interfering with interactions that stabilize structure, such as hydrogen bonding. Once the site for the disulfide bond has been chosen, the necessary thiol-modified nucleosides must be chemically synthesized. In most cases, the authors use alkylthiol linkers on the 2′ hydroxyl and the N3 position of pyrimidine residues, although linkers can be positioned at nearly any position of a nucleoside. The thiol-modified nucleoside phosphoramidites are synthesized with the thiol functionality protected as a tert-butyl disulfide, a protecting group that is stable under all conditions of solid-phase synthesis and subsequent manipulations. Incorporation of the thiol-modified nucleosides is accomplished by solid-phase chemical synthesis of the RNA. Following removal of both the exocyclic amine-protecting groups and the tert-butyldimethyl silyl groups used to protect the 2′ hydroxyls, the full-length oligoribonucleotide is purified by denaturing polyacrylamide gel electrophoresis (PAGE). Reduction of the mixed disulfide with dithiothreitol (DTT) liberates the free thiols. These are then oxidized by vigorous stirring in air to form an intramolecular disulfide bond in quantitative yield. The Basic Protocol describes the procedures for formation of intramolecular disulfide bonds by air oxidation of thiol-mediated RNA. Optimized protocols for solid-phase synthesis and deprotection of tRNAs (tRNA is used here as a model RNA) containing thiol-modified residues are presented in Support Protocol 1. Support Protocol 2 describes a method for purification of tRNAs containing thiol-modified residues. Support Protocol 3 describes a fluorescence assay to quantify the concentration of free thiol remaining during cross-link formation. Support Protocols 4, 7, and 8 describe the synthesis of three different thiol-containing nucleoside phosphoramidites, while Support Protocol 6 describes the synthesis of an intermediate compound needed for Support Protocols 7 and 8. Support Protocol 5 details the preparation of a thiol-modified controlled-pore glass (CPG) support used in the solid-phase synthesis of the modified RNA. NOTE: All procedures should be conducted using sterile techniques. When handling the RNA, suitable precautions should be taken to avoid RNase contamination. Gloves must be worn during handling of all equipment. Treatment of solutions with DEPC to inactivate RNases is not necessary, but could be done if preferred. CAUTION: Acrylamide is a neurotoxin, minimize inhalation and skin exposure. The reagents and solvents used in both solid-phase RNA synthesis and the synthesis of the thiol-modified phosphoramidites should be handled according to the manufacturer’s safety data sheet for each reagent.
Methods for Cross-Linking Nucleic Acids
Contributed by Emily J. Maglott and Gary D. Glick
5.4.1
Current Protocols in Nucleic Acid Chemistry (2000) 5.4.1-5.4.34 Copyright © 2000 by John Wiley & Sons, Inc.
BASIC PROTOCOL
FORMATION OF INTRAMOLECULAR DISULFIDE CROSS-LINKS IN RNA The RNA used in this protocol has been synthesized with the thiol groups protected as tert-butyl disulfides. Liberation of the free thiols is achieved by reduction of the mixed disulfides with DTT or other reducing agents, such as glutathione. Following removal of the DTT by continuous flow dialysis, the intramolecular disulfide bond is formed in the dilute RNA solution by air oxidation in a mildly basic solution (e.g., pH 8.0). Materials tert-Butyl-disulfide-modified RNA (see Support Protocol 1) 100 mM sodium phosphate buffer, pH 8.3 (see recipe) Dithiothreitol (DTT) Sodium phosphate buffer/NaCl, pH 7.0 (see recipe) 5000-molecular-weight-cutoff (MWCO) cellulose ester membrane 200 mM MgCl2 (see recipe) 0.1 N NaOH TE buffer (APPENDIX 2A) 1.5 M sodium acetate (NaOAc), pH 5.5 Absolute ethanol 10-well (500 µL) microdialyzer (e.g., Spectrum) 0.2-µm bottle-top filter Argon (Ar) tank Peristaltic pump 70°C water bath pH meter with microelectrode Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (e.g., CPMB UNIT 7.6 and APPENDIX 3B of this manual) or for derivatization and fluorescence spectroscopy for quantitation of thiols (see Support Protocol 3) Remove tert-butyl disulfide protecting groups to liberate free thiols 1. In a 0.5-mL microcentrifuge tube, dissolve the tert-butyl-disulfide-modified RNA (0.5 OD260 U) to a concentration of 50 µM in 16 µL of 100 mM sodium phosphate buffer, pH 8.3. 2. Add 0.054 mg DTT (200 eq per disulfide), close the tube, and incubate the mixture at 25°C for 12 hr. The authors have found that between 12 and 14 hr of incubation effects complete reduction of the tert-butyl disulfide protecting groups. This can be conveniently conducted overnight.
3. Open the tube and dilute the reaction mixture with 100 µL sodium phosphate buffer/NaCl, pH 7.0. 4. Assemble a 10-well continuous-flow microdialyzer with a 5000-MWCO cellulose ester membrane following the manufacturer’s instructions. Other types of dialysis assemblies should work as well.
Engineering Disulfide Cross-Links in RNA via Air Oxidation
5. Filter 3 L sodium phosphate buffer/NaCl through a 0.2-µm bottle-top filter and sparge the filtered buffer with Ar for ≥30 min before use. 6. Fill the dialysis chamber with sodium phosphate buffer/NaCl, removing all air bubbles from under the wells by forcing them out one of the flow ports as the chamber fills.
5.4.2 Current Protocols in Nucleic Acid Chemistry
7. Transfer the reduced RNA to a well of the microdialyzer and rinse the reaction tube with phosphate buffer/NaCl, adding the rinsings to the dialyzer well. Dialyze the RNA against the same buffer at a flow rate of 5.0 mL/min for 10 hr at room temperature. Control experiments show that this method completely removes the DTT.
8. Transfer the dialyzed RNA sample from the dialyzer well to a 1.5-mL microcentrifuge tube. Rinse the well of the dialyzer with 100 to 200 µL of sodium phosphate buffer/NaCl and add to the RNA sample. Form intramolecular disulfide bonds via air oxidation 9. Dilute the reduced, dialyzed tRNA sample to a final RNA concentration of 1 to 4 µM with sodium phosphate buffer/NaCl to prevent intermolecular disulfide bonds from forming. 10. Add 200 mM MgCl2 stock solution to 5 mM final concentration. Fold the RNA by heating the RNA sample to 70°C for 2 min in a water bath and then allowing the sample to cool to room temperature (15 to 30 min). This procedure has been optimized for yeast tRNAPhe samples; details of the folding protocol (i.e., time and temperature for refolding, with Mg2+ concentration) will be different for other RNAs.
11. Calibrate a microelectrode at pH 7.0. Immerse the electrode in the RNA sample. Adjust the pH of the sample to 8.0 with 0.1 N NaOH (typically 3 to 5 µL for a 400-µL sample). 12. Stir the RNA solution at room temperature exposed to air, loosely covering the tubes with a paper towel to minimize dust contamination. 13. Monitor disulfide bond formation by removing 5-µL aliquots of the reaction mixture and assessing them either by denaturing PAGE (e.g., CPMB UNIT 7.6) or by fluorescence spectroscopy after derivatization with 7-diethylamino-3-(4′-maleimidylphenyl)-4methylcoumarin (see Support Protocol 3). Disulfide bond formation is usually complete in ≤12 hr. Progress of cross-linking can be monitored throughout; however frequent monitoring is time-consuming and will decrease final yields. Cross-linked RNAs have slower gel mobility than the non-cross-linked RNAs (Sigurdsson and Eckstein, 1996). The fluorescence assay using 7-diethylamino-3-(4′-maleimidylphenyl)-4-methylcoumarin (Parvari et al., 1983) is more sensitive than with Ellman’s reagent.
14. Conduct native gel electrophoresis (e.g., CPMB UNIT 2.7) to verify formation of intramolecular cross-links. 15. Dialyze the cross-linked RNA against 1 L TE buffer using continuous-flow microdialysis as described in steps 4 to 8. Do not precipitate directly from the phosphate buffer, as recovered yields will be poor. Disulfide cross-link formation is usually quantitative, as judged by ethidium bromide staining of a sample analyzed by PAGE, and purification to remove non-cross-linked RNA is not necessary.
Methods for Cross-Linking Nucleic Acids
5.4.3 Current Protocols in Nucleic Acid Chemistry
Purify RNA from eluate 16. Add 1⁄5 vol of 1.5 M NaOAc, pH 5.5, and 3 vol absolute ethanol. Mix thoroughly and place at –20°C overnight. 17. Centrifuge 1 hr at 16,000 × g, 4°C. 18. Decant the solution and dry the pellet of RNA under vacuum. 19. Combine the RNA into one tube by dissolving the pellets in water and transferring to one tube. Rinse the tubes at least 3 times with water to ensure transfer of all RNA. Evaporate the RNA solutions dryness under vacuum and store at –20°C until use. 20. Store the precipitated RNA at −20°C (stable at least 1 year). SUPPORT PROTOCOL 1
SYNTHESIS OF RNA CONTAINING tert-BUTYLDISULFIDE-MODIFIED NUCLEOSIDES BY SOLID-PHASE METHODS Incorporation of modified nucleosides into an RNA can be accomplished with the necessary phosphoramidites and solid-phase synthesis technology. The following procedure has been optimized to use standard, commercially available ribonucleoside phosphoramidites and reagents combined with thiol-modified nucleoside phosphoramidites to allow total chemical synthesis of RNAs at least 76 nucleotides long. Careful handling to minimize exposure of the phosphoramidites and reagents to moisture throughout the procedure enables milligram quantities of 76-mer RNAs containing modified nucleosides to be obtained from a single 1-µmol-scale synthesis. Materials Methylene chloride (CH2Cl2) in CaH2 Acetonitrile (CH3CN) in CaH2 Thiol-modified nucleoside phosphoramidites (see Support Protocols 4, 7, and 8) Nucleoside phosphoramidites Argon (Ar) Anhydrous ethanol Absolute ethanol, room temperature and –20°C Brine (saturated aqueous NaCl) Nitrogen (N2) stream Ammonia gas (NH3) 1.0 M tetrabutylammonium fluoride (TBAF) in tetrahydrofuran (THF) 1.5 M sodium acetate (NaOAc), pH 5.5 (see recipe) Ethyl acetate (EtOAc)
Engineering Disulfide Cross-Links in RNA via Air Oxidation
Rotoevaporator Automated nucleic acid synthesizer Trap-Pak molecular sieve bags (Perseptive Biosystems) Rubber septa Cannula CPG column loaded with appropriate nucleoside (purchased, or see Support Protocol 5) Desiccator 1- and 2-dram glass vials (oven dried >8 hr at 180°C) Teflon tape Dry bath, 55°C Speedvac evaporator or equivalent Spatula, RNase-free
5.4.4 Current Protocols in Nucleic Acid Chemistry
5-in. (12.5-cm) glass pipets Rotary shaker Prepare thiol-modified phosphoramidites for automated synthesis 1. At least 24 hr before beginning the RNA synthesis, distill 1 L CH2Cl2 and 100 mL CH3CN from CaH2. 2. Dry the necessary thiol-modified phosphoramidites by coevaporation with distilled CH3CN (~1 mL/50 mg) three times, then under high vacuum overnight. The quantity of phosphoramidite depends on both the RNA sequence (if the same or different nucleosides are used) and the number of sequences being synthesized at one time. One coupling typically takes 5 mg dissolved in 5 mL on a Perkin-Elmer Expedite 8909 synthesizer.
3. Prepare the automated nucleic acid synthesizer for RNA synthesis following the manufacturer’s suggested protocols. Replacing all reagent solutions on the synthesizer with newly opened bottles of reagents immediately prior to use significantly improves both the quality and the yield of the synthesis. Although commercial preparations of 3% TCA can be used, it is recommended that solutions of 3% TCA be made with freshly distilled CH2Cl2 immediately prior to synthesis. The use of Trap-Pak molecular sieve bags to remove residual water in the solvent in CH3CN solutions (one Trap-Pak per liter) is also recommended. The protocols described here have been conducted with both AN-Bz, CN-Bz, GN-iBu, and U 5′-O-dimethoxytrityl, 2′-O-tert-butyldimethylsilyl, 3′-O-diisopropyl-β-cyanoethyl phosphoramidites as well as AN-Pac and GN-Pac 5′-O-dimethoxytrityl, 2′-O-tert-butyldimethylsilyl, and 3′-O-diisopropyl-β-cyanoethyl phosphoramidites. Other exocyclic amine-protecting groups should also be suitable for these protocols.
4. Backfill the flask(s) containing the thiol-modified phosphoramidite(s) with Ar and quickly seal with a rubber septum. 5. Working under Ar, dilute the thiol-modified phosphoramidite(s) with distilled CH3CN to 50 mg/mL. Using a cannula, place the solution into an oven-dried bottle of appropriate size for the synthesizer and place on the synthesizer. This amidite concentration is recommended for an Expedite 8909 synthesizer (Perseptive Biosystems). Other synthesizers will also work; amidites should be diluted according to the manufacturer’s protocols.
6. Prime the delivery lines with all ancillary reagents twice, and then prime the delivery lines with the amidite solutions twice, to ensure that the flow lines from each bottle of reagent and from the amidite bottle are filled. 7. Place a CPG column loaded with the appropriate nucleoside onto the synthesizer. See Glick (1991) for details on the loading of thiol-modified supports.
8. Begin the RNA synthesis following the manufacturer’s protocols. Addition of CH2Cl2 wash steps both before and after TCA treatment improves coupling efficiencies (see Goodwin et al., 1996). RNA synthesis is very sensitive to water. Eliminate water from all reagents and handling procedures as much as possible. The use of freshly distilled CH3CN and CH2Cl2 and newly opened dry reagents improves coupling efficiencies and allows the synthesis of RNAs exceeding 75 residues. It is also recommended that phosphoramidites be handled under an Ar atmosphere in a glove bag, especially in humid conditions. While solutions of A, C, and U phosphoramidites are stable for at least 5 days, the coupling efficiency of G phosphoramidites is increased with fresh dilution on a daily basis. A typical RNA coupling cycle
Methods for Cross-Linking Nucleic Acids
5.4.5 Current Protocols in Nucleic Acid Chemistry
takes 15 min (Gait et al., 1991; Goodwin et al., 1994). The synthesis of one 76-mer tRNA sequence takes ∼19.5 hr.
Deprotect synthetic RNA 9. Cool a 2-dram oven-dried vial under N2. 10. Add 2 to 3 mL anhydrous ethanol to the vial and cool in a brine/ice water bath under N2 for 2 to 5 min. 11. Remove the N2 line and insert a needle attached to a lecture bottle of NH3 and a vent. Saturate the ethanol by constant bubbling with NH 3 for at least 30 min. This procedure should be conducted in a fume hood.
12. Open the synthesis column and pour the solid support into a 1-dram oven-dried vial. 13. Quickly pour ∼1 mL of the anhydrous ethanolic ammonia solution into the vial. Seal threads of the vial with Teflon tape and cap tightly. Seal the outside of the cap with Teflon tape and parafilm to prevent evaporation. The inner Teflon tape can be put on the vial before pouring the solution, but be careful not to pour the ethanolic ammonia onto the tape or the seal will not be adequate. Usually bubbling occurs as the solution is poured onto the resin.
14. Place the sealed vial in a 55°C dry bath for 18 hr. 15. Remove the vial from the dry bath and allow to cool to room temperature before opening. Carefully open the vial and pass a gentle stream of N2 over the solution for 5 to 15 min to remove the ammonia. When the cap is removed, there should be a small release of pressure; this indicates that the ethanolic ammonia solution was thoroughly saturated during deprotection. A significant portion of the ethanol may evaporate during this procedure. If the solution becomes viscous (typically, with less than ∼300 mL ethanol remaining), add 500 mL of anhydrous ethanol to the vial before proceeding to step 16.
16. Transfer the ethanol solution from the vial to an autoclaved 1.5-mL microcentrifuge tube and evaporate under vacuum to ≤400 µL total volume. Transfer any remaining solution to the microcentrifuge tube and then rinse the solid support with 500 µL anhydrous ethanol. Add the rinsing to the microcentrifuge tube and evaporate the solution under vacuum to ≤400 µL. 17. Thoroughly rinse the solid support with anhydrous ethanol five more times using 100 to 200 µL ethanol each time, adding the rinsings to the microcentrifuge tube. Evaporate the solution to dryness under vacuum. Coevaporate with 500 µL anhydrous ethanol. Repeat the coevaporation once more using 200 µL ethanol. The residue resulting from step 17 may be stored for a few days at −20°C before proceeding with the remainder of this protocol.
Desilylate synthetic RNA 18. Cool a 1-dram oven-dried vial in a desiccator.
Engineering Disulfide Cross-Links in RNA via Air Oxidation
19. Gently loosen the residue obtained after step 17 from the microcentrifuge tube with the tip of an RNase-free spatula. Carefully transfer the residue to the dried vial. 20. Add ∼400 µL of 1.0 M TBAF in THF to the microcentrifuge tube with a glass pipet. Vortex, centrifuge, and transfer the solution to the vial with a glass pipet. Repeat this rinsing procedure twice more using ∼300 µL TBAF each time.
5.4.6 Current Protocols in Nucleic Acid Chemistry
TBAF solutions should be of the highest purity and lowest water content possible. For optimal results, bottles of TBAF should only be used once. TBAF is light sensitive.
21. Seal the vial with Teflon tape, parafilm, and cover in foil. Place the vial on a rotary shaker and shake at low speed for 24 hr. 22. Open the vial and quench the reaction with 1 mL of 1.5 M NaOAc, pH 5.5. Transfer the solution in two approximately equal aliquots into two 1.5-mL microcentrifuge tubes. Rinse the vial 5 times with 150 µL water and add the rinsings to the tubes. 23. Evaporate the solutions to ∼600 µL total volume in each tube. Add 600 to 700 µL EtOAc to each tube and homogenize as thoroughly as possible. Centrifuge to separate the layers. Remove the upper layer (EtOAc) with a pipet and repeat the extraction a second time. Evaporate the solution under vacuum for ∼10 min to remove residual EtOAc. 24. Transfer about half of the solution in each tube to another 1.5-mL microcentrifuge tube, resulting in four tubes containing ∼300 µL solution each. 25. Add 1 mL absolute ethanol to each tube, mix thoroughly, and place at −20°C overnight. 26. Centrifuge all tubes for at least 1 hr at 16,000 × g, 4°C. Decant the supernatant from the pellet and then wash the pellet with 200 µL of –20°C absolute ethanol. Dry the pellet under vacuum. 27. Quantify the crude RNA by dissolving the pellets in water and measuring the absorbance of a 1-µL aliquot of the solution at 260 nm. Estimate the molar extinction coefficient (ε260) using nearest-neighbor calculations. The ε260 calculation is done as described by Breslauer et al. (1986); see UNIT 7.3. This assumes that the modified residues behave like their unmodified counterparts. Both crude and purified RNA samples should be stored either pelleted from ethanol precipitation or dried down from aqueous solutions at −20°C.
28. Purify the crude RNA to single-nucleotide resolution (see Support Protocol 2). PURIFICATION OF SYNTHETIC RNA CONTAINING tert-BUTYLDISULFIDE PROTECTED THIOL-MODIFIED NUCLEOSIDES
SUPPORT PROTOCOL 2
Single-nucleotide resolution of the products resulting from solid-phase synthesis, the fulllength oligomer as well as failed sequences, can be achieved using denaturing PAGE (also see APPENDIX 3B). The procedures that are outlined have been optimized for the purification of a 76-mer tRNA sequence. These protocols can be modified for sequences of other lengths by adjusting the percentage of acrylamide in the gel matrix until both the full-length RNA and the sequence resulting from incomplete synthesis one residue from the end are resolved. Materials Crude synthetic tRNA (see Support Protocol 1) 80% formamide containing 0.05% xylene cyanol (XC) tracking dye 8% denaturing polyacrylamide solution (see recipe) 1× TBE electrophoresis buffer ( APPENDIX 2A) 1× and 4× TAE electrophoresis buffer (APPENDIX 2A) 1.5 M sodium acetate (NaOAc), pH 5.5 Absolute ethanol Power supply Silica-gel plate
Methods for Cross-Linking Nucleic Acids
5.4.7 Current Protocols in Nucleic Acid Chemistry
UVG-11 Mineralight lamp (254 nm, 115 V) or equivalent Razor blade, RNase free Hoefer Six-Pac electroeluter or equivalent Inner elution tubes Porous polyethylene plugs Blotter-paper discs Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (e.g., see CPMB UNIT 7.6 and APPENDIX 3B of this manual) Separate RNA species by PAGE 1. Dissolve 20 to 40 OD260 U of crude synthetic tRNA in 60 µL water, mix thoroughly, and allow to stand for ∼1 hr to thoroughly dissolve the RNA. There will probably also be insoluble material present.
2. Add 60 µL of 80% formamide/0.05% XC and centrifuge briefly to separate the insoluble matter. 3. Prepare an 8% denaturing polyacrylamide gel 31.0 cm × 38.5 cm × 0.8 mm, with 14.6-mm-wide wells (e.g., see CPMB UNIT 7.6 and APPENDIX 3B). Electrophoresis conditions are outlined for the purification of a 76-nt tRNA sequence. For sequences of different lengths, adjust the percentage acrylamide and electrophoresis time accordingly.
4. Load 20 µL of the RNA sample into each of six wells, being careful not to load any insoluble material onto the gel. 5. Electrophorese the gel in 1× TBE at 55 W until the XC tracking dye has migrated ∼11 inches (∼4 to 4.5 hr). Under these conditions 76-mer RNA migrates with the XC—be careful not to run the RNA off the gel.
Electroelute full-length synthetic RNA from gel slice 6. Place the gel over a silica-gel plate in a dark location. Briefly shine 254-nm light on the gel to locate the RNA. With a clean RNase-free razor blade, excise the full-length RNA from the gel. RNA is photoreactive; minimize exposure of RNA to UV light.
7. Pack the RNA slices into three or four inner elution tubes containing 300 µL of 1× TAE using blotter-paper discs and porous polyethylene plugs according to manufacturer’s directions. Cut off the bottom tip of the inner elution tube and insert into a 1.5-mL microcentrifuge tube containing 200 µL of 4× TAE. Electroelute in a Hoefer Six-Pac eluter for 90 min at +50 V. For efficient electroelution, remove all air bubbles from the inner elution tubes. Also, the elutions work better if the inner elution tubes are packed tightly with gel slices, but it is important not to crush the slices. If the amperage does not decrease below 0.2 mA during the course of electroelution, the gel slices can be recovered from the inner elution tube and soaked in 400 mL of 4× TAE at room temperature overnight. The eluted RNA should then be ethanol precipitated as described in the next step and combined with the RNA eluted during electroelution. Engineering Disulfide Cross-Links in RNA via Air Oxidation
8. Remove the inner elution tube carefully, rinse the electrode with 20 µL water, and add to the eluted RNA. Remove the electrode and pipet all buffer remaining above the porous plug into the eluted RNA. The total volume should be ∼350 mL.
5.4.8 Current Protocols in Nucleic Acid Chemistry
Purify RNA from eluate 9. Add 70 µL of 1.5 M NaOAc, pH 5.5, and 1 mL absolute ethanol. Mix thoroughly and place at −20°C overnight. 10. Centrifuge 1 hr at 16,000 × g, 4°C. 11. Decant the solution and dry the pellet of RNA under vacuum. 12. Combine the RNA into one tube by dissolving the pellets in water and transferring to one tube. Rinse the tubes at least 3 times with water to ensure transfer of all RNA. Evaporate the RNA solutions dryness under vacuum and store at −20°C until use (stable at least 1 year). QUANTIFICATION OF THIOLS IN RNA USING 7-DIETHYLAMINO-3-(4′-MALEIMIDYLPHENYL)-4-METHYLCOUMARIN
SUPPORT PROTOCOL 3
Reaction of free thiols with 7-diethylamino-3-(4′-maleimidylphenyl)-4-methylcoumarin produces a fluorescent adduct that can be quantified spectroscopically (Parvari et al., 1983). This provides a convenient method to determine when the cross-linking reaction is complete. The method can be used to accurately determine thiol concentrations as low as 0.5 µM. Materials RNA solution containing free thiols (see Basic Protocol) 5× CPM buffer (see recipe) 0.4 mM CPM in isopropyl alcohol (see recipe) 1% Triton X-100 Sodium phosphate buffer/NaCl, pH 7.0 (see recipe) Fluorimeter and cuvette 1. To a 5-µL aliquot of an RNA solution containing free thiols, add 1 µL of 5× CPM buffer and 5 µL of 0.4 mM CPM in isopropyl alcohol. Incubate the sample 10 min at room temperature. 2. Dilute the reaction with 489 µL of 1% Triton X-100. 3. Prepare a blank consisting of: 5 µL sodium phosphate buffer/NaCl, pH 7.0 1 µL 5× CPM buffer 5 µL 0.4 mM CPM in isopropyl alcohol 489 µL Triton X-100. 4. Transfer blank solution to a fluorimeter cuvette and measure the fluorescence intensity at 480 nm (λex = 390 nm). 5. In the same way, measure the fluorescence intensity of the RNA solution from step 2, and correcting for background fluorescence. A calibration curve can be constructed using DTT as a standard (0.5 to 5 mM is a useful range).
Methods for Cross-Linking Nucleic Acids
5.4.9 Current Protocols in Nucleic Acid Chemistry
SUPPORT PROTOCOL 4
SYNTHESIS OF 5′-O-(4,4′-DIMETHOXYTRITYL)-2′-O-(tertBUTYLDIMETHYLSILYL)-N3-(ETHYL)URIDINE-3′-O-(N,N-DIISOPROPYLβ-CYANOETHYLPHOSPHORAMIDITE) tert-BUTYL DISULFIDE (S.5) The preparation of a thiol-containing nucleoside phosphoramidite modified at the N3 position of uridine is described. The synthesis of 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tertbutyldimethylsilyl)-N3-(ethyl)uridine-3′-O-(N,N-diisopropyl-β-cyanoethylphosphoram idite) tert-butyl disulfide (S.5) proceeds in four sets of steps from uridine (Fig. 5.4.1). The 3′-O-tert-butyldimethylsilyl-protected isomer (S.4) obtained as a side product of the third set of steps is used to make a thiol-modified controlled-pore glass (CPG) support (Fig. 5.4.2; see Support Protocol 5). Materials Uridine Distilled acetonitrile (CH3CN) Distilled triethylamine (Et3N) N,N-Dimethylformamide (DMF) Chlorotrimethylsilane (TMSCl) N2 Petroleum ether Diethyl ether (Et2O) Sodium hydride (NaH) 1-Tosyl-2-benzoylmercaptoethanol (see Glick et al., 1991) 48% (w/v) aqueous HF Methylene chloride (CH2Cl2) Brine (saturated aqueous NaCl) Sodium sulfate (Na2SO4) O
O NH N
HO
a-d
O
N
HO
O HO
O SBz
N O
N
DMTrO
O HO
OH
SSt-Bu
N
e-f
OH
O
O HO
OH
1
2 O
O N
DMTrO
O
O
O Ot-BuDMS
HO
g
h
O
N
DMTrO
SSt-Bu
N
SSt-Bu
N
NC
3
t-BuDMSO
Ot-BuDMS N(i-Pr)2
SSt-Bu
N N
O P
5
O
DMTrO
O
O
O OH 4
Engineering Disulfide Cross-Links in RNA via Air Oxidation
Figure 5.4.1 Synthesis of 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N3-(ethyl)uridine-3′-O-(N,N-diisopropyl-β-cyanoethylphosphoramidite) tert-butyl disulfide. (a) TMSCl, Et3N, DMF; (b) NaH, DMF; (c) p-TsOCH2CH2SBz, DMF; (d) HF (aq); (e) DMTrCl, pyridine; (f) 1-(tert-butylthio)1,2-hydrazinedicarboxmorpholide, LiOH, CH3OH; (g) TBDMSCl, imidazole, DMF; (h) chloro-N,Ndiisopropylamine-β-cyanoethyl phosphine, 2,4,6-collidine, N-methylimidazole, THF. Abbreviations: Bz, benzoyl; DMF, dimethylformamide; DMTr, 4,4′-dimethoxytrityl; TBDMS, tert-butyldimethylsilyl; THF, tetrahydrofuran; Ts, tosyl.
5.4.10 Current Protocols in Nucleic Acid Chemistry
O
O SSt-Bu
N DMTrO
t-BuDMSO
N
O
a,b
O
O SSt-Bu
N DMTrO
OH
N
O
O
t-BuDMSO
O
O
SSt-Bu
N c DMTrO
N
O
O
t-BuDMSO
O
O
4 O
OC6Cl5 6
O
NH
CPG
7
Figure 5.4.2 Synthesis of 5′-O-(4,4′-dimethoxytrityl)-3′-O-(tert-butyldimethylsilyl)-2′-O-succinylCPG N3-(ethyl)uridine tert-butyl disulfide. (a) succinic anhydride, DMAP, pyridine; (b) pentachlorophenol, DCC, DMAP, CH2Cl2; (c) CPG (1000 Å), Et3N, DMF. Abbreviations: DCC, dicyclohexyl-carbodiimide; DMAP, 4-dimethylaminopyridine; DMF, dimethylformamide; DMSO, dimethyl sulfoxide.
Methanol (CH3OH) Pyridine 4,4′-Dimethoxytrityl chloride (DMTrCl) 1-(tert-Butylthio)-1,2-hydrazine carboxmorpholide (see Wünsch et al., 1982) LiOH⋅H2O Ethyl acetate (EtOAc) Imidazole tert-Butyldimethylsilyl chloride (TBDMSCl) Tetrahydrofuran (THF) 2,4,6-Collidine N-Methylimidazole Chloro-N,N-diisopropylamine-β-cyanoethyl phosphine Additional reagents and equipment for flash chromatography ( APPENDIX 3E) Prepare N3-(2-thiobenzoylethyl)uridine (S.1) 1. Dry 24.42 g uridine (100 mmol) by coevaporation with distilled CH3CN. 2. Dissolve the dried uridine in 83.6 mL freshly distilled Et3N (600 mmol, 6 eq) and 250 mL DMF. 3. Cool the solution to 4°C. 4. Slowly add 42 mL chlorotrimethylsilane (330 mmol, 3.3 eq) and stir the reaction under N2 for 2 hr. 5. Remove salts that precipitate during the course of the reaction by filtration under N2. 6. Triturate the residual salts in the filtrate with 1:1 (v/v) petroleum ether/diethyl ether. 7. Dissolve the residue in 400 mL DMF and cool to 4°C. 8. Add 4.40 g NaH (110 mmol, 1.1 eq) to the reaction mixture with stirring under N2. 9. After hydrogen evolution subsides, add 37 g 1-tosyl-2-benzoylmercaptoethanol (110 mmol, 1.1 eq) and stir the reaction overnight at 45°C. This step is based on the procedure detailed by Glick (1991).
10. Cool the solution to room temperature and remove the silyl groups by the addition of 5 mL of 48% aqueous HF.
Methods for Cross-Linking Nucleic Acids
5.4.11 Current Protocols in Nucleic Acid Chemistry
11. After 1 hr, dilute the reaction mixture with CH2Cl2 and wash successively with water and brine. 12. Dry the organic layer over Na2SO4 and concentrate under vacuum. 13. Purify the oily residue by flash chromatography ( APPENDIX 3E) using 19:1 (v/v) CH2Cl2/CH3OH to obtain S.1 as a white foam (26 g, 63% yield). Prepare 5′-O-(4,4′-dimethoxytrityl)-N3-(ethyl)uridine tert-butyl disulfide (S.2) 14. Coevaporate 15.0 g compound S.1 (37 mmol) once from 100 mL of 9:1 (v/v) CH3CN/pyridine, then dissolve in 185 mL pyridine. 15. Add 15.0 g 4,4′-dimethoxytrityl chloride (44 mmol, 1.2 eq) in 2.5-g portions over a 6-hr period at 4°C with stirring. 16. Allow the reaction to warm to room temperature overnight while stirring under N2. 17. Add 5 mL CH3OH and stir the mixture an additional 10 min. 18. Remove the solvents under vacuum, and coevaporate the residue with 100 mL CH3CN to yield a light yellow-orange foam. 19. Dissolve the crude tritylated product (14 g, 20 mmol) in 100 mL CH3OH and add 8.31 g 1-(tert-butylthio)-1,2-hydrazinedicarboxmorpholide (Wünsch et al., 1982) (24 mmol, 1.2 eq) and 2.52 g LiOH⋅H2O (60 mmol, 3.0 eq). 20. Stir the reaction under N2 for 12 hr and then concentrate the mixture under vacuum. 21. Dissolve the residue in EtOAc and wash with brine. 22. Dry the organic layer over Na2SO4 and concentrate under vacuum. 23. Purify the residue by flash chromatography ( APPENDIX 3E) using a step gradient of 3:2 to 2:3 (v/v) petroleum ether/EtOAc to obtain S.2 as a white foam (9.7 g, 70% yield). Prepare 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N3-(ethyl)uridine tert-butyl disulfide (S.3) and 5′-O-(4,4′-dimethoxytrityl)-3′-O-(tert-butyldimethylsilyl)-N3-(ethyl)uridine tert-butyl disulfide (S.4) 24. Dissolve 9.5 g compound S.2 (13.6 mmol) in 50 mL DMF. 25. Add 2.31 g imidazole (34 mmol, 2.5 eq) and 2.58 g tert-butyldimethylsilyl chloride (17.1 mmol, 1.25 eq). 26. Stir the mixture overnight under N2. 27. Dilute the reaction with EtOAc, wash the mixture with brine, dry over Na2SO4, and evaporate under vacuum. 28. Purify the residue by flash chromatography (APPENDIX 3E) using 9:1 (v/v) petroleum ether/EtOAc to obtain S.3 and S.4 as white foams (S.3: 3.9 g, 40% yield; S.4: 2.1 g, 25% yield).
Engineering Disulfide Cross-Links in RNA via Air Oxidation
Prepare 5′-O-(4,4′-dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N3-(ethyl)uridine3′-O-(N,N-diisopropyl-β-cyanoethylphosphoramidite) tert-butyl disulfide (S.5) 29. Dissolve 0.81 g compound S.3 (1.0 mmol) in 3.0 mL THF. 30. Add 1.0 mL 2,4,6-collidine (7.5 mmol, 7.5 eq) and 40 µL N-methylimidazole (0.5 mmol, 0.5 eq).
5.4.12 Current Protocols in Nucleic Acid Chemistry
31. Add 0.56 mL chloro-N,N-diisopropylamine-β-cyanoethyl phosphine (2.5 mmol, 2.5 eq) dropwise while stirring under N 2. 32. After 2 hr, dilute the reaction with EtOAc and wash the mixture with NaHCO3 and brine. 33. Dry the organic layer over Na2SO4, and concentrate under vacuum. 34. Purify the residue by flash chromatography (APPENDIX 3E) using 80:15:5 (v/v/v) petroleum ether/EtOAc/Et3N to obtain S.5 as a brittle white foam (0.82 g, 82% yield). SYNTHESIS OF 5′-O-(4,4′-DIMETHOXYTRITYL)-3′-O-(tert-BUTYLDIMETHYLSILYL)-2′-O-SUCCINYL-CPG N3-(ETHYL)URIDINE tert-BUTYL DISULFIDE CONTROLLED-PORE GLASS SUPPORT
SUPPORT PROTOCOL 5
5′-O-(4,4′-dimethoxytrityl)-3′-O-(tert-butyldimethylsilyl)-2′-O-succinyl-CPG N3-(ethyl)uridine tert-butyl disulfide (Fig. 5.4.2), a thiol-modified controlled-pore glass (CPG) support, is prepared using a side product (S.4) obtained in Support Protocol 4. This nucleoside support can be used to introduce a thiol modification at the 3′ terminus of an RNA species. The nucleoside loading concentration is usually 32 µmol/g (Schaller et al., 1963). Materials Compound S.4 (see Support Protocol 4) Pyridine Succinic anhydride 4-Dimethylaminopyridine (DMAP) Methylene chloride (CH2Cl2) Brine (saturated aqueous NaCl) Sodium sulfate (Na2SO4) Pentachlorophenol Dicyclohexylcarbodiimide (DCC) Petroleum ether Ethyl acetate (EtOAc) Long-chain alkyl amino controlled-pore glass (CPG), 1000-Å pore size, 100 µmol amino groups/g, 120/200 mesh N,N-Dimethylformamide (DMF) Triethylamine (Et3N) Methanol (CH3OH) Diethyl ether (Et2O) Acetic anhydride Additional reagents and equipment for flash chromatography (APPENDIX 3E) Prepare 5′-O-(4,4′-dimethoxytrityl)-3′-O-(tert-butyldimethylsilyl)-2′-O-pentachlorophenylsuccinate N3-(ethyl)uridine tert-butyl disulfide (S.6) 1. Dissolve 1.77 g compound S.4 (2.2 mmol) in 12.0 mL pyridine under N2. 2. Add 0.69 g succinic anhydride (6.6 mmol, 3.0 eq) and 0.133 g DMAP (1.1 mmol, 0.5 eq) and incubate 12 hr. 3. Concentrate the reaction mixture under vacuum. 4. Dissolve the residue in CH2Cl2 and wash with brine. 5. Dry the organic layer over Na2SO4 and concentrate under vacuum.
Methods for Cross-Linking Nucleic Acids
5.4.13 Current Protocols in Nucleic Acid Chemistry
6. Dissolve the crude succinate in 25 mL CH2Cl2 and add 0.88 g pentachlorophenol (3.3 mmol, 1.5 eq), 67 mg DMAP (0.55 mmol, 0.25 eq), and 0.91 g DCC (4.4 mmol, 2.0 eq). 7. After 8 hr, add petroleum ether to precipitate dicyclohexylurea. 8. Gravity filter the reaction mixture and concentrate under vacuum. 9. Purify the residue by flash chromatography (APPENDIX 3E) using 4:1 (v/v) petroleum ether/EtOAc to obtain S.6 as a white foam (2.3 g, 90% yield). Prepare 5′-O-(4,4′-dimethoxytrityl)-3′-O-(tert-butyldimethylsilyl)-2′-O-succinyl-CPG N3-(ethyl)uridine tert-butyl disulfide (S.7) 10. Suspend long-chain alkyl amino CPG in 4.0 mL DMF with 0.58 g compound S.6 (0.5 mmol, 5 eq) and 0.14 mL Et3N (1.0 mmol, 10 eq). 11. Gently swirl the mixture in the dark for 2 days. 12. Vacuum filter the support and rinse successively with 15 mL DMF, 50 mL CH3OH, and 50 mL Et2O. 13. Remove the residual solvents under vacuum. 14. Acetylate the unreacted amino groups by swirling the support for 1 hr with 0.70 mL acetic anhydride (7.0 mmol, 100 eq) and 10 mg DMAP (70 µmol, 1 eq) in 4 mL pyridine. 15. Rinse the support successively with 30 mL pyridine, 90 mL CH3OH, and 90 mL Et2O. 16. Remove the residual solvents under vacuum. SUPPORT PROTOCOL 6
SYNTHESIS OF 3′,5′-O-(TETRAISOPROPYLDISILOXANE-1,3-DIYL)-2′-OALLYL-O4-(2-NITROPHENYL) URIDINE (S.10) INTERMEDIATE Preparation of thiol-modified nucleoside phosphoramidite modified at the 2′-hydroxyl position of pyrimidines proceeds from preparation of 3′,5′-O-(tetraisopropyldisiloxane1,3-diyl)-2′-O-allyl-O4-(2-nitrophenyl)uridine. Preparation of this compound is described. First, uridine is converted to 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)O4-(2-nitrophenyl) uridine (S.9) in two sets of steps as described by Sproat and Lamond (1991). This compound is then allylated using the procedure described by Sproat et al. (1991).
Engineering Disulfide Cross-Links in RNA via Air Oxidation
Materials Uridine Pyridine 1,3-Dichloro-1,1,3,3-tetraisopropyldisiloxane Methylene chloride (CH2Cl2) Methanol (CH3OH) Saturated aqueous sodium bicarbonate (NaHCO3) Sodium sulfate (Na2SO4) Triethylamine (Et3N) Chlorotrimethylsilane Brine (saturated aqueous NaCl) 2-Mesitylenesulfonyl chloride 4-Dimethylaminopyridine (DMAP) 2-Nitrophenol 1,4-Diazabicyclo[2.2.2]octane (DABCO)
5.4.14 Current Protocols in Nucleic Acid Chemistry
p-Toluenesulfonic acid monohydrate p-Dioxane Ethyl acetate (EtOAc) Petroleum ether Tetrahydrofuran (THF) Triphenylphosphine Tris(dibenzylideneacetone) dipalladium(0) Allyl ethyl carbonate Additional reagents and equipment for flash chromatography (APPENDIX 3E) Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-uridine (S.8) 1. Dissolve 499 mg uridine (2.04 mmol) in 5.1 mL pyridine. 2. Cool to 0°C under N2. 3. Dissolve 732 µL 1,3-dichloro-1,1,3,3-tetraisopropyldisiloxane (2.29 mmol, 1.1 eq) in 0.5 mL CH2Cl2 and add dropwise to uridine solution. 4. Stir the reaction, allowing it to warm to room temperature, for 4 hr. 5. Dilute the reaction by adding 0.4 mL CH 3OH, and evaporate under vacuum. 6. Dissolve the residue in 15 mL CH2Cl2 and wash three times with saturated aqueous NaHCO3. 7. Dry the organic layer over Na2SO4 and evaporate under vacuum. 8. Dissolve the residue in toluene and evaporate under vacuum to obtain S.8 as a white foam (1.0 g, 100%). Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-O4-(2-nitrophenyl) uridine (S.9) 9. Dissolve 3.4 g compound S.8 (6.9 mmol) in 35 mL CH2Cl2 and cool to 0°C. It may be necessary to repeat the synthesis of S.8 to obtain the necessary quantity for the synthesis of S.9.
10. Add 9 mL Et3N (64.6 mmol, 9.4 eq) followed by 6.4 mL chlorotrimethylsilane (50.4 mmol, 7.3 eq) and stir the reaction 4 hr under N2. 11. Pour the reaction onto 100 mL saturated aqueous NaHCO3 and stir for 10 min. 12. Wash the organic layer with brine, dry over Na 2SO4, and concentrate under vacuum to a peach foam (3.8 g, 100%). 13. Dissolve the residue in 36 mL CH2Cl2 and 4.8 mL Et3N (34.6 mmol, 5 eq). 14. Add 2.25 g 2-mesitylenesulfonyl chloride (10.3 mmol, 1.5 eq) and 0.5 g DMAP (3.5 mmol, 0.5 eq). Stir for 15 min. 15. Add 2.0 g 2-nitrophenol (14.4 mmol, 2.1 eq) and 0.42 g DABCO (3.7 mmol, 0.55 eq) and continue stirring for 3 hr. 16. Dilute the reaction with 40 mL CH2Cl2 and wash with saturated aqueous NaHCO3. 17. Backwash the aqueous layer twice with CH2Cl2. 18. Dry the combined organic layers over Na2SO4 and concentrate under vacuum. 19. Dissolve the residue in 21 mL CH 2Cl2 and add a solution of 2.70 g p-toluenesulfonic acid monohydrate in 21 mL p-dioxane.
Methods for Cross-Linking Nucleic Acids
5.4.15 Current Protocols in Nucleic Acid Chemistry
20. Stir the reaction for 2 min and then quench by adding 2.5 mL Et3N. 21. Pour the reaction onto saturated aqueous NaHCO3 and backwash the aqueous layer with CH2Cl2 twice. 22. Dry the combined organic layers over Na2SO4 and concentrate under vacuum. 23. Purify the residue by flash chromatography (APPENDIX petroleum ether to obtain S.9 (4.5 g, 6.9 mmol, 100%).
3E)
using 55% EtOAc in
Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-2′-O-allyl-O4-(2-nitrophenyl) uridine (S.10) 24. Dissolve the 4.5 g compound S.9 in 37 mL THF (37 mL). 25. Add 371 mg triphenylphosphine (1.4 mmol, 0.2 eq) and 168 mg tris(dibenzylideneacetone) dipalladium(0) (0.2 mmol, 0.026 eq). 26. Add 1.9 mL allyl ethyl carbonate (14.0 mmol, 2 eq) dropwise. 27. Heat the reaction to reflux for 2 hr. 28. Cool the reaction to room temperature and concentrate under vacuum. 29. Purify the residue by flash chromatography (APPENDIX petroleum ether to obtain S.10 (3.3 g, 72%). SUPPORT PROTOCOL 7
3E)
using 18% EtOAc in
SYNTHESIS OF 5′-O-(4,4′-DIMETHOXYTRITYL)-N4-(BENZOYL)-2′-O(ETHYL)CYTIDINE-3′O-(N,N-DIISOPROPYL->-CYANOETHYLPHOSPHORAMIDITE) tert-BUTYL DISULFIDE (S.20) The intermediate prepared in Support Protocol 6 is converted to 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-N4-(benzoyl)-2′-O-allyl cytidine (S.11) using the method of Sproat and Lamond (1991) replacing isobutyryl chloride with benzoyl chloride. The synthesis of the fully protected phosphoramidite can be accomplished in nine sets of steps (Fig. 5.4.3).
Engineering Disulfide Cross-Links in RNA via Air Oxidation
Materials Compound S.10 (see Support Protocol 6) Tetrahydrofuran (THF) Ammonia (NH3) N2 Methanol (CH3OH) Petroleum ether Ethyl acetate (EtOAc) Pyridine N,N-Dimethylformamide (DMF) Benzoic anhydride Ammonium hydroxide (NH4OH) N-Methylmorpholine-N-oxide Acetone Osmium tetroxide (OsO4) Saturated aqueous sodium bisulfite Diethyl ether (Et2O) Saturated aqueous sodium bicarbonate (NaHCO3) Brine (saturated aqueous NaCl) Sodium sulfate (Na2SO4)
5.4.16 Current Protocols in Nucleic Acid Chemistry
O2N O O NH N
HO
N
O
a
O
N
O
O
(i-Pr)2Si
O HO
O
NH
O (i-Pr)2Si O
b-e
O (i-Pr)2Si O (i-Pr)2Si
OH
OH O2N
N
O
O O
OH
8 NHBz
9
O N N f
N
O (i-Pr)2Si O (i-Pr)2Si
O
N
O
g,h
O
O (i-Pr)2Si
O
O
i
O
(i-Pr)2Si
O
O
O
11 10 NHBz
NHBz
N
N
N
O
O
O O (i-Pr)2Si O (i-Pr)2Si O O
j
O
(i-Pr)2Si
N
OH
O (i-Pr)2Si
O
O
OH
12
H
NHBz
NHBz
N
N O O (i-Pr)2Si O (i-Pr)2Si O O
k
O
13
NHBz
N
O
O
N
O O (i-Pr)2Si O (i-Pr)2Si O O
l
OH
N O
N
O O (i-Pr)2Si O (i-Pr)2Si O O
m
OMs
15
14
NHBz
N N
HO
N O
o
O HO
O
SBz
16
NHBz
n
O
N
HO
HO
SBz
O
O O
SSt-Bu
18
17
NHBz NHBz N N p
N
DMTrO
O
O HO
O
SSt-Bu
N
DMTrO
NC
O
O
O
q O P
O
SSt-Bu
N(i-Pr)2
19 20
Figure 5.4.3 Synthesis of 5′-O-(4,4′-dimethoxytrityl)-N4-(benzoyl)-2′-O-(ethyl)cytidine tert-butyl disulfide 3′-O-(N,N-diisopropyl-β-cyanoethylphosphoramidite). (a) pyridine, 1,3-dichloro-1,1,3,3tetraisopropyldisiloxane, CH2Cl2; (b) chlorotrimethylsilane, Et3N, CH2Cl2; (c) 2-mesitylenesulfonyl chloride, DMAP, Et3N, CH2Cl2; (d) 2-nitrophenol, DABCO; (e) p-toluenesulfonic acid monohydrate, p-dioxane, CH2Cl2; (f) triphenylphosphine, tris(dibenzylideneacetone) dipalladium(0), allyl ethyl carbonate, THF; (g) NH3, THF; (h) pyridine, benzoic anhydride, DMF; (i) OsO4, N-methylmorpholine-N-oxide, acetone, H2O; (j) NaIO4, p-dioxane, H2O; (k) NaBH4, CH3OH; (l) methanesulfonyl chloride, pyridine; (m) thiobenzoic acid, Et3N, DMF; (n) HF (aq), CH3CN; (o) 1-(tert-butylthio)-1,2hydrazinedicarboxmorpholide, LiOH, CH3OH, THF; (p) DMTrCl, Et3N, DMF, DMAP; (q) chloro-N,Ndiisopropylamine-β-cyanoethyl phosphine, N,N-diisopropylethylamine, CH2Cl2. Abbreviations: Bz, benzoyl; DMAP, 4-dimethylaminopyridine; DMF, dimethylformamide; DMTr, 4,4′-dimethoxytrityl; Ms, methanesulfonyl; THF, tetrahydrofuran.
Methods for Cross-Linking Nucleic Acids
5.4.17 Current Protocols in Nucleic Acid Chemistry
Methylene chloride (CH2Cl2) p-Dioxane Sodium periodate (NaIO4) Sodium borohydrate (NaBH4) Methanesulfonyl chloride Triethylamine (Et3N) Thiobenzoic acid Acetonitrile (CH3CN) 48% (w/v) aqueous HF 1-(tert-Butylthio)-1,2-hydrazinedicarboxmorpholide (Wünsch et al., 1982) LiOH⋅H2O 1 N aqueous sodium citrate 4-Dimethylaminopyridine (DMAP) 4,4′-Dimethoxytrityl chloride (DMTrCl) N,N-Diisopropylethylamine Chloro-N,N-diisopropylamine-β-cyanoethyl phosphine Pressure tube (−78°C) CO2/isopropyl alcohol (i-PrOH) bath Additional reagents and equipment for flash chromatography (APPENDIX 3E) Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)- N4-(benzoyl)-2′-O-allyl cytidine (S.11) 1. Dissolve 1.94 g compound S.10 (2.99 mmol) in 8.5 mL THF in a pressure tube. 2. Cool the tube to −78°C using a CO2/i-PrOH bath. 3. Add 5 g NH3 (299 mmol, 100 eq). 4. Seal the reaction vessel and stir while allowing it to warm to room temperature for 62 hr. 5. Cool the reaction mixture to −78°C using CO2/i-PrOH bath and open the pressure tube. 6. Allow the mixture to warm under N 2 to room temperature. 7. Concentrate under vacuum. 8. Purify the residue by flash chromatography (APPENDIX 3E) using a step gradient of 0 to 10% CH3OH in 7:3 (v/v) petroleum ether/EtOAc. 9. Dissolve the purified residue in 14 mL pyridine and 6.5 mL DMF. 10. Add 0.72 g benzoic anhydride (3.20 mmol, 1.5 eq) and stir the reaction under N2 for 12 hr at room temperature. 11. Cool the reaction to 0°C and add 0.4 mL water. 12. Stir the reaction 5 min at 0°C, then add 0.3 mL NH4OH. Engineering Disulfide Cross-Links in RNA via Air Oxidation
13. Continue stirring the reaction 10 min at 0°C, then concentrate the mixture under vacuum. 14. Purify the residue by flash chromatography (APPENDIX 3E) using 3:2 (v/v) petroleum ether/EtOAc to obtain S.11 (1.42 g, 75% yield) as a white foam.
5.4.18 Current Protocols in Nucleic Acid Chemistry
Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)- 2′-O-(2,3-dihydroxypropyl)cytidine (S.12) 15. Dissolve 1.34 g compound S.11 (2.30 mmol) and 0.27 g N-methylmorpholine-N-oxide (2.34 mmol, 1.1 eq) in 21 mL of 6:1 (v/v) acetone/water. 16. Add 5 mg OsO4 (21 µmol, 0.01 eq) and stir the reaction mixture in the dark for 2.5 hr. 17. Add 1 mL saturated aqueous sodium bisulfite to precipitate osmium salts. 18. Decant the solution and dissolve the light-brown residue in Et 2O. 19. Wash the solution with saturated aqueous NaHCO3 and brine. 20. Dry the combined organic layers over Na2SO4 and concentrate under vacuum. 21. Purify the yellow residue by flash chromatography (APPENDIX 3E) using a step gradient of 19:1 to 37:3 CH2Cl2/CH3OH to obtain S.12 (1.27 g, 90% yield). Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-N4-(benzoyl)-2′-O-(ethanal)cytidine (S.13) 22. Dissolve the 1.27 g compound S.12 (1.91 mmol) in 20 mL of 3:1 (v/v) p-dioxane/water. 23. Add 0.49 g NaIO4 (2.29 mmol, 1.2 eq) and stir the reaction mixture in the dark for 4.5 hr. 24. Dilute the reaction with Et2O and wash with water. 25. Dry the organic layer over Na2SO4 and concentrate under vacuum. 26. Purify the residue by flash chromatography (APPENDIX 3E) using 19:1 (v/v) CH2Cl2/CH3OH to obtain S.13 as a white foam (2.94 g, 97% yield). Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-N4-(benzoyl)-2′-O(2-hydroxyethyl)cytidine (S.14) 27. Dissolve 1.17 g compound S.13 (1.85 mmol) in 19 mL CH3OH. 28. Add 21 mg NaBH4 (0.56 mmol, 0.3 eq) and stir the mixture in the dark under N2 for 90 min. 29. Dilute the solution with Et 2O and wash with saturated aqueous NaHCO3 and brine. 30. Wash the aqueous layer once with Et2O. 31. Dry the combined organic layers over Na2SO4 and concentrate under vacuum. 32. Purify the residue by flash chromatography (APPENDIX 3E) using 19:1 (v/v) CH2Cl2/CH3OH to obtain S.14 as a white foam (1.09 g, 93% yield). Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-N4-(benzoyl)-2′-O-(ethyl-2methylsulfonate)cytidine (S.15) 33. Dissolve the 1.09 g compound S.14 (1.72 mmol) in 17 mL CH2Cl2 and 1.4 mL pyridine (17.2 mmol, 10 eq). 34. Cool under N2 to 0°C. 35. Add 0.19 mL methanesulfonyl chloride (2.41 mmol, 1.2 eq) dropwise. 36. Stir the mixture under N2 while gradually warming to room temperature overnight.
Methods for Cross-Linking Nucleic Acids
5.4.19 Current Protocols in Nucleic Acid Chemistry
37. Dilute the reaction with Et2O and wash with saturated aqueous NaHCO3 and brine. 38. Dry the organic layer over Na2SO4 and concentrate under vacuum. 39. Purify the residue by flash chromatography (APPENDIX 3E) using 24:1 (v/v) CH2Cl2/CH3OH to obtain S.15 as a white foam (1.13 g, 92% yield). Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-N4-(benzoyl)-2′-O-(thiobenzoylethyl) cytidine (S.16) 40. Dissolve 1.04 g compound S.15 (1.46 mmol) in 5.8 mL DMF. 41. Add 2.0 mL Et3N (14.6 mmol, 10 eq) and 0.34 mL thiobenzoic acid (2.91 mmol, 2.0 eq). 42. Stir the mixture under N2 in the dark overnight. 43. Dilute the solution with Et2O and wash with saturated aqueous NaHCO3 and brine. 44. Dry the organic layer over Na2SO4 and concentrate under vacuum to a dark orangebrown solid. 45. Purify the residue by flash chromatography (APPENDIX 3E) using a step gradient of 0 to 10% CH3CN in 2:1 (v/v) petroleum ether/EtOAc to obtain S.16 as a white foam (0.84 g, 76% yield). Prepare N4-(benzoyl)-2′-O-(thiobenzoylethyl)cytidine (S.17) 46. Dissolve 0.37 g compound S.16 (0.49 mmol) in 4.3 mL CH3CN. 47. Add 0.5 mL of 48% aqueous HF and stir the reaction mixture for 7 hr. 48. Dilute the solution with Et2O and wash with H2O. 49. Dry the organic layer over Na2SO4 and concentrate under vacuum. 50. Purify the pink residue by flash chromatography (APPENDIX 3E) using 19:1 (v/v) CH2Cl2/CH3OH to obtain S.17 as a white foam (0.25 g, 100% yield). Prepare N4-(benzoyl)-2′-O-(ethyl)cytidine tert-butyl disulfide (S.18) 51. Dissolve the 0.25 g compound S.17 (0.49 mmol) in 3.8 mL of 1:1 (v/v) CH3OH/THF. 52. Add 0.20 g 1-(tert-butylthio)-1,2-hydrazinedicarboxmorpholide (Wünsch et al., 1982) (0.58 mmol, 1.2 eq) and 41 mg LiOH⋅H2O (0.97 mmol, 2.0 eq). 53. Stir the reaction under N2 at 0°C for 45 min. 54. Dilute the reaction with EtOAc, and then wash with 1 N aqueous sodium citrate and the saturated aqueous NaHCO3. 55. Dry the organic layer over Na2SO4 and concentrate under vacuum. 56. Purify the residue by flash chromatography (APPENDIX 3E) using 24:1 (v/v) CH2Cl2/CH3OH to obtain S.18 as a pink foam (0.18 g, 74% yield).
Engineering Disulfide Cross-Links in RNA via Air Oxidation
Prepare 5′-O-(4,4′-dimethoxytrityl)-N4-(benzoyl)-2′-O-(ethyl)cytidine tert-butyl disulfide (S.19) 57. Dissolve 0.10 g compound S.18 (0.21 mmol) and 13 mg DMAP (0.10 mmol, 0.5 eq) in 0.8 mL DMF and 26 µL pyridine (0.31 mmol, 1.5 eq). 58. Add 85 mg DMTrCl (0.25 mmol, 1.2 eq) and stir the reaction under N2 for 6 hr.
5.4.20 Current Protocols in Nucleic Acid Chemistry
59. Dilute the reaction with CH2Cl2 and wash with saturated aqueous NaHCO3 and brine. 60. Dry the organic layer over Na2SO4 and concentrate under vacuum. 61. Purify the residue by flash chromatography (APPENDIX 3E) using 1:1 (v/v) petroleum ether/acetone to obtain S.19 as a tan foam (0.13 g, 77% yield). Prepare 5′-O-(4,4′-dimethoxytrityl)-N4-(benzoyl)-2′-O-(ethyl)cytidine 3′-O-(N,N-diisopropyl->-cyanoethylphosphoramidite) tert-butyl disulfide (S.20) 62. Dissolve 55 mg compound S.19 (0.07 mmol) in 0.3 mL CH2Cl2 containing 60 µL N,N-diisopropylethylamine (0.35 mmol, 5 eq). 63. Cool under N2 to 0°C. 64. Add 23 µL chloro-N,N-diisopropylamine-β-cyanoethyl phosphine (0.10 mmol, 1.5 eq) dropwise and stir the reaction under N2 while allowing it to warm to room temperature. 65. After 2 hr, quench the excess chloridate with 0.3 mL CH3OH and concentrate the mixture under vacuum. 66. Purify the residue by flash chromatography (APPENDIX 3E) using 2:1 (v/v) petroleum ether/acetone to obtain S.20 as a white foam (58 mg, 84% yield). SYNTHESIS OF 5′-O-(4,4′-DIMETHOXYTRITYL)-2′-O-(ETHYL)URIDINE-3′O-(N,N-DIISOPROPYL->-CYANOETHYLPHOSPHORAMIDITE) tert-BUTYL DISULFIDE
SUPPORT PROTOCOL 8
The intermediate S.10 prepared in Support Protocol 6 can be converted to 3′,5′-O(tetraisopropyldisiloxane-1,3-diyl)-2′-O-allyl uridine (S.21) using nitrobenzaldoxime and 1,1,3,3-tetramethylguanidine as described by Sproat and Lamond (1991). The synthesis of the fully protected phosphoramidites can be accomplished in nine sets of steps (Fig. 5.4.4). Materials 2-Nitrobenzaldoxime 1,1,3,3-Tetramethylguanidine Acetonitrile (CH3CN) Compound S.10 (see Support Protocol 6) Ethyl acetate (EtOAc) Sodium sulfate (Na2SO4) Methanol (CH3OH) Methylene chloride (CH2Cl2) Acetone N-Methylmorpholine-N-oxide Osmium tetroxide (OsO4) Saturated aqueous sodium bisulfite Celite 1,4-Dioxane Sodium periodate (NaIO4) Diethyl ether (Et2O) Sodium bicarbonate (NaHCO3) Sodium borohydride (NaBH4) Brine (saturated aqueous NaCl) Pyridine
Methods for Cross-Linking Nucleic Acids
5.4.21 Current Protocols in Nucleic Acid Chemistry
O
O
a
10
N
O
b
O
O (i-Pr)2Si O
N
O
O
(i-Pr)2Si
N O O (i-Pr)2Si O (i-Pr)2Si O O
O
OH
HO
O
O
OMs
h
SBz
DMTrO
27
O 28
O NH
NH
O
O HO
SBz
O NH
N
SBz
i
g
O
N O O (i-Pr)2Si O (i-Pr)2Si O O 26
O NH
O
NH f
25
O
H
O
N O O (i-Pr)2Si O (i-Pr)2Si O O
24
HO
O O 23
NH e
O
O
O (i-Pr)2Si O
OH
O NH
O
O
22
O
N
O
N
(i-Pr)2Si
OH
O (i-Pr)2Si O
O
NH c
O
O
(i-Pr)2Si
21
d
O NH
NH
N
DMTrO
O
O HO
O
DMTrO
j
SSt-Bu
29
N
NC
O
O
O O P
O
SSt-Bu
N(i-Pr)2 30
Figure 5.4.4 Synthesis of 3′-O-(N,N-diisopropyl-β-cyanoethyl-phosphoramidite)-5′-O-(4,4′-dimethoxytrityl)-2′-O-(ethyl)uridine tert-butyl disulfide. (a) 2-Nitrobenzaldoxime, 1,1,3,3-tetramethylguanidine, CH3CN; (b) OsO4, N-methylmorpholine-N-oxide, acetone, H2O; (c) NaIO4, p-dioxane, H2O; (d) NaBH4, CH3OH; (e) methanesulfonyl chloride, pyridine; (f) thiobenzoic acid, Et3N, DMF; (g) HF (aq), CH3CN; (h) DMTrCl, Et3N, DMF, DMAP; (i) 1-(tert-butylthio)-1,2-hydrazine-dicarboxmorpholide, LiOH, CH3OH, THF; (j) chloro-N,N-diisopropylamine-β-cyanoethyl phosphine, N,Ndiisopropylethylamine, CH2Cl2. Abbreviations: Bz, benzoyl; DMAP, 4-dimethylaminopyridine; DMF, dimethylformamide; DMTr, 4,4′-dimethoxytrityl; Ms, methanesulfonyl.
Methanesulfonyl chloride N,N-Dimethylformamide (DMF) Triethylamine (Et3N) Thiobenzoic acid 48% (w/v) aqueous HF 4,4′-Dimethoxytrityl chloride (DMTrCl) 4-Dimethylaminopyridine (DMAP) Tetrahydrofuran (THF) LiOH⋅H2O 1-(tert-Butylthio)-1,2-hydrazinedicarboxmorpholide 1 N aqueous sodium citrate N,N-Diisopropylethylamine Chloro-N,N-diisopropylamine-β-cyanoethyl phosphine Petroleum ether Additional reagents and equipment for flash chromatography (APPENDIX 3E) Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-2′-O-allyluridine (S.21) 1. Dissolve 2.1 g 2-nitrobenzaldoxime (12.3 mmol, 4 eq) and 1.4 mL 1,1,3,3-tetramethylguanidine (11.1 mmol, 3.6 eq) in 28 mL CH3CN. Engineering Disulfide Cross-Links in RNA via Air Oxidation
2. Stir this solution under N2. 3. Add the solution to 2.0 g compound S.10 (3.1 mmol).
5.4.22 Current Protocols in Nucleic Acid Chemistry
4. Stir the reaction for 30 min at room temperature and then concentrate under vacuum. 5. Dissolve the residue in EtOAc and wash with water. 6. Backwash the water layer 3 times with EtOAc. 7. Combine the organic layers and dry over Na2SO4. 8. Concentrate under vacuum. 9. Purify the residue by flash chromatography (APPENDIX 3E) using a step gradient of 0 to 5% CH3OH in CH2Cl2 to obtain S.21 (1.61 g, 99%). Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-2′-O-(2,3-dihydroxypropyl)uridine (S.22) 10. Dissolve 1.61 g compound S.21 (3.06 mmol) in 30 mL of 6:1 (v/v) acetone/water. 11. Add 394 mg N-methylmorpholine-N-oxide (3.4 mmol, 1.1 eq) and 7.7 mg OsO4 (0.03 mmol, 0.01 eq) and stir in the dark for 3 hr. 12. Precipitate the osmium salts with 2 mL saturated aqueous sodium bisulfite and pour the mixture through a funnel containing Celite. 13. Evaporate the solvent under vacuum to obtain S.22 as a white foam (1.77 g, 100% yield). Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-2′-O-(ethanal)uridine (S.23) 14. Dissolve 437 mg compound S.22 (0.8 mmol) in 7.8 mL of 3:1 (v/v) 1,4-dioxane/H2O. 15. Add 209 mg NaIO4 (1.0 mmol, 1.25 eq) and stir the reaction in the dark for 6 hr. 16. Dilute the solution with Et 2O and filter. 17. Wash the filtrate with saturated aqueous NaHCO 3, dry over Na2SO4, and concentrate under vacuum to obtain S.23 as a pale-yellow foam (400 mg, 97% yield). Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-2′-O-(2-hydroxyethyl)uridine (S.24) 18. Dissolve 470 mg compound S.23 (0.9 mmol) in 7.4 mL CH3OH. It may be necessary to repeat the synthesis of S.22 to obtain the necessary quantity for the synthesis of S.23.
19. Add 16 mg NaBH4 (0.4 mmol, 0.48 eq) and stir the mixture overnight under N2. 20. Dilute the mixture with Et2O, then wash once with saturated aqueous NaHCO3 and twice with brine. 21. Dry the combined organic layers over Na2SO4, filter, and evaporate under vacuum to obtain S.24 as a white foam (464 mg, 98% yield). Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-2′-O-(ethyl-2-methylsulfonate)uridine (S.25) 22. Dissolve 138 mg compound S.24 (0.3 mmol) in a mixture of 2.6 mL CH2Cl2 and 0.21 mL pyridine. 23. Cool to 0°C. 24. Add 28 µL methanesulfonyl chloride (0.4 mmol, 14 eq) dropwise while stirring under N2.
Methods for Cross-Linking Nucleic Acids
5.4.23 Current Protocols in Nucleic Acid Chemistry
25. Allow the reaction to warm to room temperature over 18 hr. 26. Dilute the mixture with CH2Cl2 and wash once with saturated aqueous NaHCO3. 27. Dry the organic layer over Na2SO4, filter, and evaporate under vacuum. 28. Purify the residue by flash chromatography (APPENDIX 3E) using 17:3 (v/v) CH2Cl2/acetone to obtain S.25 as a white foam (133 mg, 84% yield). Prepare 3′,5′-O-(tetraisopropyldisiloxane-1,3-diyl)-2′-O-(thiobenzoylethyl)uridine (S.26) 29. Dissolve 552 mg compound S.25 (0.9 mmol) in a mixture of 3.6 mL DMF and 1.3 mL Et3N (9.1 mmol, 10.0 eq). It may be necessary to repeat the synthesis of S.25 to obtain the necessary quantity for the synthesis of S.26.
30. Add 0.21 mL thiobenzoic acid (1.8 mmol, 2 eq) and stir overnight under N2. 31. Dilute the mixture with Et2O, then wash once with saturated aqueous NaHCO3 and twice with brine. 32. Dry the combined organic extracts over Na2SO4, filter, and concentrate under vacuum. 34. Purify the residue by flash chromatography (APPENDIX 3E) using a step gradient of 47:3 to 4:1 (v/v) CH2Cl2/acetone to obtain S.26 as a white foam (472 mg, 80% yield). Prepare 2′-O-(thiobenzoylethyl)uridine (S.27) 35. Dissolve the 472 mg compound S.26 (0.7 mmol) in 5.8 mL CH3CN. 36. Add 1.3 mL of 48% aqueous HF and stir for 4 hr. 37. Dilute the mixture with EtOAc and wash once with H 2O. 38. Dry the organic layer over Na2SO4 and filter. 39. Evaporate under vacuum to obtain S.27 as a white foam (327 mg, 100% yield). Prepare 5′-O-(4,4′-dimethoxytrityl)-2′-O-(thiobenzoylethyl)uridine (S.28) 40. Dissolve 277 mg compound S.27 (0.7 mmol) in 2.7 mL DMF and 82 µL pyridine (1.0 mmol, 1.5 eq). 41. Add 280 mg DMTrCl (0.8 mmol, 1.2 eq) and 54 mg DMAP (0.4 mmol, 0.65 eq). 42. Stir the reaction overnight under N2. 43. Dilute the reaction with CH 2Cl2 and wash the solution once with saturated aqueous NaHCO3 and twice with brine. 44. Dry the combined organic extracts over Na2SO4, filter, and concentrate under vacuum. 45. Purify the residue by flash chromatography (APPENDIX 3E) using 48:1 (v/v) CH2Cl2/CH3OH) to obtain S.28 as a yellow foam (302 mg, 63% yield). Engineering Disulfide Cross-Links in RNA via Air Oxidation
Prepare 5′-O-(4,4′-dimethoxytrityl)-2′-O-(ethyl)uridine tert-butyl disulfide (S.29) 46. Dissolve 293 mg compound S.28 (0.4 mmol) in 3.3 mL of 1:1 (v/v) THF/CH3OH. 47. Add 26 mg LiOH⋅H2O (0.6 mmol, 1.5 eq).
5.4.24 Current Protocols in Nucleic Acid Chemistry
48. After stirring for 2 min, add 164 mg 1-(tert-butylthio)-1,2-hydrazinedicarboxmorpholide (0.5 mmol, 1.2 eq) (Wünsch et al., 1982). 49. Stir the mixture for an additional 10 min under N2. 50. Dilute the solution with CH2Cl2. 51. Wash the solution with 1 N aqueous sodium citrate, saturated aqueous NaHCO 3, and brine. 52. Dry the organic layer over Na2SO4. 53. Filter the solution and evaporate under vacuum. 54. Purify the residue by flash chromatography (APPENDIX 3E) using 49:1 (v/v) CH2Cl2/CH3OH to obtain S.29 as a white foam (285 mg, 100% yield). Prepare 3′-O-(N,N-diisopropyl->-cyanoethyl-phosphoramidite)-5′-O-(4,4′dimethoxytrityl)-2′-O-(ethyl)uridine tert-butyl disulfide (S.30) 55. Dissolve 49 mg compound S.29 (0.07 mmol) in 0.28 mL CH2Cl2 containing 62 µL N,N-diisopropylethylamine (0.4 mmol, 5 eq) and cool to 0°C. 56. Add 24 µL chloro-N,N-diisopropylamine-β-cyanoethyl phosphine (0.1 mmol, 1.5 eq) dropwise. 57. Stir the mixture for 2 hr under N2. 58. Dilute the solution with CH3OH. 59. Concentrate the solution under vacuum. 60. Purify the residue by flash chromatography (APPENDIX 3E) using 19:6 (v/v) petroleum ether/acetone to obtain S.30 as a white foam (49 mg, 78% yield). REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
CPM buffer, 5× 7.6 g Tris⋅Cl (250 mM) 38 g NaCl (2.6 mM) 46.5 mg Na2EDTA (0.5 mM) 0.38 mL Triton X-100 (0.15%, v/v) H2O to 200 mL Adjust pH to 7.5 with 1 N NaOH or HCl Add H2O to 250 mL Store up to 1 month at 25°C CPM in isopropanol, 0.4 mM Dissolve 1.6 mg of 7-diethylamino-3-(4′-maleimidylphenyl)-4-methylcoumarin (Molecular Probes) in 10 mL HPLC-grade isopropanol. Store up to 1 week at 25°C. Denaturing polyacrylamide gel solution, 8% 0.32 g bisacrylamide 9.28 g acrylamide 57.7 g urea (8 M final) 24 mL 5× TBE buffer (APPENDIX 2A) H2O to 120 mL Prepare fresh daily Current Protocols in Nucleic Acid Chemistry
Methods for Cross-Linking Nucleic Acids
5.4.25
MgCl2, 200 mM Dissolve 10.2 g magnesium chloride hexahydrate in 250 mL water. Store at room temperature (stable for several months). Sodium acetate, 1.5 M, pH 5.5 Dissolve 12.3 g anhydrous sodium acetate (NaOAc) in 100 mL water. Adjust pH to 5.5 with glacial acetic acid. Store up to 6 months at 25°C. Sodium phosphate buffer, 100 mM, pH 8.3 17.8 g Na2HPO4 H2O to 900 mL Adjust pH to 8.3 with 1 N H3PO4 or NaOH Add H2O to 1 L Store up to 6 months at 25°C Sodium phosphate buffer/NaCl, pH 7.0 100 mM sodium phosphate buffer, pH 7.0 (APPENDIX 2A) 5 mM NaCl (APPENDIX 2A) Check pH and adjust to 7.0 with 1 N H3PO4 or NaOH, if necessary Store up to 6 months at 25°C COMMENTARY Background Information
Engineering Disulfide Cross-Links in RNA via Air Oxidation
Generation of cysteine mutants in proteins capable of forming disulfide bonds is a useful technique to stabilize structure as well as to probe structure, folding, and dynamics. Disulfide cross-links have also been incorporated into DNA to examine ground-state structure, trap non-ground-state structures, and examine protein-DNA and DNA-DNA interactions (Glick, 1998). Disulfide cross-links have more recently been incorporated into RNA sequences to stabilize secondary structure (Goodwin and Glick, 1994; Allerson and Verdine, 1995), to examine dynamic motion between helices in the Tetrahymena ribozyme (Cohen and Cech, 1997), and to probe solution conformation of a tRNA, the hammerhead ribozyme, and the hairpin ribozyme (Sigurdsson et al., 1995; Goodwin et al., 1996; Earnshaw et al., 1997; Maglott and Glick, 1998). The methods that have been used to incorporate disulfide bonds into RNA have relied on either air oxidation or thiol-disulfide exchange. Although air oxidation is slower than thiol-disulfide exchange, using the methodology presented here it is possible to achieve quantitative conversion of the purified thiolmodified RNA to disulfide-cross-linked RNA. The ability to form intramolecular disulfide cross-links in quantitative yields makes it possible to produce milligram quantities of these constructs, which can then be used in a wide variety of biophysical and
biochemical experiments. In addition, attachment of the alkylthiol tethers at positions other than the 5′ or 3′ hydroxyls allows for radiolabeling of constructs modified at the termini of the RNA. Disulfide cross-links can be incorporated into RNA secondary and tertiary structures at many different locations. The specific thiolmodified nucleosides that are needed to create a particular cross-link depend on the location of the cross-links within the RNA. It is possible to form cross-links between two of the same thiol-modified nucleosides or between two different thiol-modified nucleosides. The preparation of nucleosides with thiol modifications at the N3 position of uridine as well as at the 2′ hydroxyl of both cytidine and uridine is described in this unit. Preparation of many other thiol-modified nucleosides has been reported, including those with alkylthiols on both G and A (Glick, 1998). Use of particular bases depends on the specific application (see Glick, 1998). The thiol group in each of these nucleosides is protected as a tert-butyl disulfide, which is stable to all conditions of solid-phase synthesis. These protocols involve relatively simple synthetic organic chemistry techniques and standard glassware and equipment. In addition to the thiol-modified nucleosides described here, synthesis of both adenosine and guanosine derivatized with 2′-O-alkyl linkers has been achieved (Manoharan et al., 1993; Douglas et al., 1994; Gundlach et al., 1997).
5.4.26 Current Protocols in Nucleic Acid Chemistry
Alkylthiol-modified nucleosides have also been described with linkers at, among others, the C5 position of pyrimidines (Sun et al., 1996), the C8 position of purines (Gundlach et al., 1997), and all exocyclic amine groups on both purines and pyrimidines (Allerson et al., 1997).
Compound Characterization N3-(2-Thiobenzoylethyl)uridine (S.1). TLC (9:1 CH2Cl2/CH3OH) Rf = 0.41. 1H NMR (300 MHz, CD3CN) δ 3.30 (2 H, t, J = 5 Hz, CH2SC(O)Ar), 3.65-3.81 (2H, 2 dd, J = 2.5, 10 Hz, 5′, 5′′), 3.96 (1H, m, 4′), 4.16-4.19 (4H, m, 2′, 3′, NCH2), 5.66 (1H, d, J = 8.1 Hz, 6), 5.80 (1H, d, J = 3.8 Hz, 1′), 7.43-7.62 (3H, m, Ar), 7.81 (1H, d, J = 8.1 Hz, 5), 7.88 (2H, d, J = 7.3 Hz, Ar). 13C NMR (75 MHz, CD3CN) δ 27.4 (CH2SC(O)Ar), 40.9 (NCH2), 62.1 (5′), 70.8 (3′), 75.6 (2′), 85.9 (4′), 91.8 (1′), 102.1 (5), 128.0, 129.9, 134.7, 138.0 (Ar), 140.3 (6), 152.4 (2), 163.9 (4), 192.4 (SC(O)Ar). IR (KBr) ν 3548, 3448, 3086, 3060, 2944, 2901, 1706, 1673, 1655, 1462, 1388, 1286, 1211, 1124, 1079, 919, 813, 779, 695, 647, 569 cm−1. MS (FAB, 3-NBA/trifluoroacetic acid) m/z 409 (M+ + 1). 5′-O-(4,4′-Dimethoxytrityl)-N3-(ethyl)uridine tert-butyl disulfide (S.2). 1H NMR (360 MHz, CD3CN) δ 1.31 (9H, s, SS(CH3)3), 2.90 (2H, t, J = 6.0 Hz, CH2SS), 3.36 (2H, m, 5′,5′′), 3.75 (6H, s, 2 OCH3), 4.02 (1H, m, 4′), 4.11 (2H, m, NCH2), 4.18 (1H, m, 2′), 4.32 (1H, m, 3′), 5.38 (1H, d, J = 8.1 Hz, 6), 5.78 (1H, d, J = 3.2 Hz, 1′), 6.85-7.44 (13H, m, Ar), 7.73 (1H, d, J = 8.1 Hz, 5). 13C NMR (90 MHz, CD3CN) δ 30.1 (SSC(CH3)3), 37.2 (CH2SS), 41.0 (NCH2), 48.5 (SSC(CH3)3), 56.0 (OCH3), 63.4 (5′), 70.5 (3′), 75.5 (2′), 83.8 (4′), 87.5 (OC(Ph)3), 91.4 (1′), 101.9 (5), 114.2 (Ar), 128.0, 129.0, 131.1, 136.5, 136.7 (Ar), 139.7 (6), 145.9 (Ar), 152.0 (2), 159.8 (Ar), 163.3 (4). IR (film; NaCl) ν 3452, 2959, 2940, 1708, 1665, 1608, 1509, 1457, 1252, 1177, 1103, 1035, 829, 810, 702 cm−1. MS (FAB, 3-NBA) m/z 695 (M+ + 1). 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N3-(ethyl)uridine tert-butyl disulfide (S.3) TLC (4:1 petroleum ether/EtOAc) Rf = 0.30. 1H NMR (300 MHz, CD3CN) δ 0.12 (6H, s, Si(CH3)2), 0.90 (9H, s, SiC(CH3)3), 1.30 (9H, s, SSC(CH3)3), 2.88 (2H, t, J = 8.0 Hz, CH2HS), 3.38 (2H, 2 dd, J = 2.4, 11.0 Hz, 5′,5′′), 3.74 (6H, s, 2 OCH3), 4.05 (3H, m, 4′, NCH2), 4.24 (1H, m, 2′), 4.31 (1H, m, 3′), 5.35 (1H, d, J = 8.2 Hz, 6), 5.83 (1H, d, J = 4.0 Hz, 1′), 6.84-7.43 (13H, m, Ar), 7.74 (1H, d, J = 8.1
Hz, 5). 13C NMR (75 MHz, CD3CN) δ –4.4 (Si(CH3)2), 18.8 (SiC(CH3)3), 26.3 (SiC(CH3)3), 30.2 (SSC(CH3)3), 37.5 (CH2SS), 41.0 (NCH2), 48.4 (SSC(CH3)3), 55.9 (OCH3), 63.7 (5′), 71.2 (3′), 76.9 (2′), 84.1 (4′), 87.7 (OC(Ph)3), 90.6 (1′), 102.1 (5), 114.1 (Ar), 127.8, 128.8, 128.9, 130.9, 136.3, 136.5 (Ar), 139.3 (6), 145.6 (Ar), 151.7 (2), 159.7 (Ar), 162.8 (4). IR (film; NaCl) ν 3856, 3546, 2955, 2930, 2857, 2361, 2334, 1710, 1668, 1608, 1509, 1456, 1253, 1177, 1122, 1036, 836 cm−1. MS (FAB, 3-NBA) m/z 809 (M+ + 1). 5′-O-(4,4′-Dimethoxytrityl)-3′-O-(tert-butyldimethylsilyl)-N3-(ethyl)uridine tert-butyl disulfide (S.4) TLC (4:1 petroleum ether/EtOAc) Rf = 0.14. 1H NMR (300 MHz, CD3CN) δ 0.04, 0.05 (6H, 2 s, Si(CH3)2), 0.81 (9H, s, SiC(CH3)3), 1.30 (9H, s, SSC(CH3)3), 2.88 (1H, dd, J = 3.8, 11.1 Hz, 5′), 2.89 (2H, t, J = 7.6 Hz, CH2SS), 3.46 (1H, dd, J = 2.4, 11.0 Hz, 5′′), 3.74 (6H, s, 2 OCH3), 4.00 (1H, m, 4′), 4.10 (3H, m, 2′, NCH2), 4.33 (1H, m, 3′), 5.38 (1H, d, J = 8.1 Hz, 6), 5.80 (1H, d, J = 3.0 Hz, 1′), 6.84-7.42 (13H, m, Ar), 7.74 (1H, d, J = 8.1 Hz, 5). 13C NMR (75 MHz, CD3CN) δ –4.4, –4.1 (Si(CH3)2), 18.7 (SiC(CH3)3), 26.3 (SiC(CH3)3), 30.3 (SSC(CH3)3), 37.5 (CH2SS), 41.0 (NCH2), 48.3 (SSC(CH3)3, 56.0 (OCH3), 63.3 (5′), 72.0 (3′), 75.4 (2′), 84.1 (4′), 87.7 (OC(Ph)3), 92.0 (1′), 102.1 (5), 114.1 (Ar), 127.7, 128.7, 129.0, 131.0, 136.4 (Ar), 139.6 (6), 145.5 (Ar), 151.7 (2), 159.7 (Ar), 162.9 (4). IR (film; NaCl) ν 3869, 2955, 2929, 2858, 2364, 2334, 1710, 1670, 1509, 1456, 1252, 1176, 1116, 1035, 836 cm−1. MS (FAB, 3NBA) m/z 809 (M+ + 1). 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(tert-butyldimethylsilyl)-N3-(ethyl)uridine-3′-O-(N,N-diisopropyl-β-cyanoethylphosphoramidite) tertbutyl disulfide (S.5). TLC (80:15:5 petroleum ether/EtOAc/Et3N) Rf = 0.28. 1H NMR (300 MHz, CD3CN) δ (two diastereomers) 0.09, 0.12 (6H, 2 s, Si(CH3)2), 0.83, 0.85 (9H, 2 s, SiC(CH3)3), 1.01, 1.14 (12H, d, J = 9 Hz, 2 NCH(CH3)2), 1.31 (9H, s, SSC(CH3)3), 2.45 (2H, m, OCH2CH2CN), 2.88 (2H, m, CH2SS), 3.40 (2H, m, 5′, 5′′), 3.50-3.90 (4H, m, OCH2CH2CN, 2 NCH(CH3)2), 3.75 (6H, s, 2 OCH3), 4.10 (2H, m, NCH2CH2SS), 4.20-4.42 (3H, m, 2′, 3′, 4′), 5.37, 5.39 (1H, d, J = 8.1 Hz, 6), 5.84, 5.89 (1H, d, J = 6.6 Hz, 1′), 6.83-7.46 (13H, m, Ar), 7.72, 7.77 (1H, d, J = 8.1 Hz, 5). 13C NMR (75 MHz, CD CN) δ (two dias3 tereomers) –4.2 (Si(CH3)2), 18.9 (SiC(CH3)3), 21.1 (OCH2CH2CN), 25.0, 25.1, 25.2, 25.3 (NCH(CH3)2), 26.4 (SiC(CH3)3), 30.4 (SC(CH3)3), 37.8 (CH2SS), 41.2 (NCH2
Methods for Cross-Linking Nucleic Acids
5.4.27 Current Protocols in Nucleic Acid Chemistry
Engineering Disulfide Cross-Links in RNA via Air Oxidation
CH2SS), 44.1, 44.3, 44.5, 44.6 (NCH(CH3)2), 48.5 (SSC(CH3)3), 56.2 (OCH3), 59.4, 60.0 (POCH2CH2CN), 64.0, 64.2 (5′), 73.5, 73.7 (3′), 76.1, 76.5 (2′), 83.9, 84.0 (4′), 88.1 (OC(Ph)3), 90.3, 90.4 (1′), 102.4, 102.5 (5), 114.4 (Ar), 128.1, 129.0, 129.2, 129.3, 131.2, 136.4, 136.5, 136.6 (Ar), 139.3, 139.4 (6), 145.7, 145.8 (Ar), 152.1 (2), 160.0 (Ar), 162.9, 163.0 (4). 31P NMR (202 MHz, CD3CN) δ (two diastereomers) 147.4, 146.8. IR (KBr) ν 2965, 2931, 2859, 2254, 1712, 1671, 1611, 1509, 1456, 1391, 1365, 1253, 1179, 1037, 979, 836 cm−1. MS (FAB, 3-NBA) m/z 1009.5 (M+ + 1). Anal. Calcd for C51H73N4O9PS2Si: C, 60.70; H, 7.28; N, 5.55. Found: C, 60.77; H, 7.24; N, 5.47. 5′-O-(4,4′-Dimethoxytrityl)-3′-O-(tert-butyldimethylsilyl)-2′-O-pentachlorophenylsuccinate N3-(ethyl)uridine tert-butyl disulfide (S.6). TLC (4:1 petroleum ether/EtOAc) Rf = 0.68. 1H NMR (360 MHz, CD3CN) δ –0.07, 0.01 (6H, s, Si(CH3)2), 0.79 (9H, s, SiC(CH3)3), 1.29 (9H, s, SSC(CH3)3), 2.79 (4H, m, succinate CH2), 3.03 (2H, m, CH2SS), 3.29-3.47 (2H, m, 5′, 5′′), 3.76 (6H, s, 2 OCH3), 3.96-4.12 (3H, m, 4′, NCH2), 4.38 (1H, m, 3′), 5.43 (2H, m, 6, 2′), 5.97 (1H, d, J = 4.8 Hz, 1′), 6.21-7.45 (13H, m, Ar), 7.65 (1H, d, J = 8.1 Hz, 5). 13C NMR (90 MHz, CD3CN) δ –4.4, –4.1 (Si(CH3)2), 18.6 (SiC(CH3)3), 26.1 (SiC(CH3)3), 29.4, 29.5 (succinate CH2), 30.2 (SSC(CH3)3), 37.0 (CH2SS), 41.1 (NCH2CH2SS), 48.5 (SSC(CH3)3), 56.0 (OCH3), 63.6 (5′), 71.4 (3′), 76.1 (2′), 85.3 (4′), 88.0 (OC(Ph)3), 88.7 (1′), 102.5 (5), 114.2 (Ar), 128.1, 129.0, 129.1, 131.2, 136.4 (Ar), 139.4 (6), 145.7 (Ar), 152.0 (2), 159.9 (Ar), 163.1 (4), 169.6, 171.8 (succinate CO2). IR (film; NaCl) ν 2956, 2931, 1786, 1751, 1713, 1672, 1608, 1509, 1455, 1390, 1363, 1253, 1229, 1177, 1154, 1107, 1036, 837 cm−1. MS (FAB, 3-NBA) m/z 1173 (M+ + 1). Anal. Calcd for C52H59N2O11S2SiCl5: C, 53.96; H, 5.10; N, 2.42. Found C, 54.03; H, 5.15; N, 2.40. 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)uridine (S.8). TLC (1:1 petroleum ether/ EtOAc) Rf 0.28; 1H NMR (300 MHz, CDCl3) δ 1.03-1.11 (28H, m, isopropyl), 3.56-4.40 (5H, m, 2′,3′,4′,5′,5′′), 5.70 (1H, dd, 5), 5.74 (1H, s, 1′), 7.71 (1H, d, 6), 8.83 (1H, s, NH); 13C NMR (90 MHz, CDCl ) δ 12.63-13.55 3 (isopropyl CHs), 16.98-17.65 (isopropyl CH3s), 60.29 (5′), 68.85 (3′), 75.32 (2′), 82.05 (4′), 91.15 (1′), 102.18 (5), 140.10 (6), 150.48 (2), 163.84 (4); IR (KBr pellet) ν 3434, 3200, 3063, 2946, 2868, 1703, 1465, 1388, 1270, 1211, 1123, 1091, 1060, 1038, 993, 907, 885,
861, 809, 774, 705, 553 cm−1; MS (CI, CH4) m/z 487 (MH+). 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)O4-(2-nitrophenyl) uridine (S.9). TLC (1:1 petroleum ether/EtOAc) Rf 0.27; 1H NMR (300 MHz, CDCl3) δ 1.00-1.11 (28H, m, isopropyl), 3.99-4.37 (5H, m, 2′,3′,4′,5′,5′′), 5.74 (1H, s, 1′), 6.21 (1H, d, 5), 7.32 (1H, d, Ar), 7.41 (1H, ddd, Ar), 7.67 (1H, ddd, Ar), 8.13 (1H, dd, Ar), 8.23 (1H, d, 6); 13C NMR (75 MHz, CDCl3) δ 12.89-13.70 (isopropyl CHs), 17.14-17.71 (isopropyl CH3s), 60.71 (5′), 69.28 (3′), 75.29 (2′), 82.34 (4′), 92.46 (1′), 94.78 (5), 125.76, 126.14, 126.81, 134.88, 141.90, 145.08 (Ar), 145.19 (6), 154.68 (2), 171.10 (4). 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)2′-O-allyl-O4-(2-nitrophenyl) uridine (S.10). TLC (1:1 petroleum ether/EtOAc) Rf 0.68. 1H NMR (300 MHz, CDCl3) δ 1.00-1.12 (28H, m, isopropyl), 3.93-4.44 (7H, m, 2′,3,4′,5′,5′′, allyl CH2), 5.16 (1H, ddd, =CH2a), 5.39 (1H, m, =CH2b), 5.76 (1H, s, 1′), 5.92 (1H, m, CH=), 6.19 (1H, d, 5), 7.33 (1H, dd,Ar), 7.41 (1H, ddd, Ar), 7.67 (1H, ddd, Ar), 8.13 (1H, dd, Ar), 8.38 (1H, d, 6); 13C NMR (75 MHz, CDCl3) δ 12.92-13.79 (isopropyl CHs), 17.16-17.79 (isopropyl CH3s), 59.85 (5′), 68.04 (3′), 71.51 (OCH2), 81.02 (2′), 82.16 (4′), 90.46 (1′), 94.50 (5), 117.29 (=CH2), 125.80, 126.13, 126.76, 134.73 (Ar), 134.82 (CH=), 142.00 (Ar), 144.77 (6), 145.26 (Ar) 154.72 (2), 171.06 (4); IR (KBr pellet) ν 3105, 2947, 2869, 1683, 1633, 1603,1549, 1531, 1454, 1365, 1351, 1288, 1220, 1166, 1126, 1073, 1041, 1014, 991, 886, 858, 780, 698 cm−1; MS (CI, CH4) m/z 648 (M+). 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)N4-(benzoyl)-2′-O-allyl cytidine (S.11). TLC (2:3 EtOAc/petroleum ether) Rf = 0.32. 1H NMR (300 MHz, CDCl3) δ 0.96-1.11 (28H, m, 4 (CH3)2HSi, 4 (CH3)2HSi), 3.95-4.02 (2H, m, 2′-H, 5′-Ha), 4.16 (1H, dd, J = 4.1, 9.6 Hz, 3′-H), 4.24-4.32 (2H, m, 4′-H, 5′-Hb), 4.39-4.52 (2H, m, 7-Ha,b), 5.19 (1H, d, J = 10.5 Hz, 9-Htrans), 5.42 (1H, d, J = 16.0 Hz, 9-Hcis), 5.84 (1H, s, 1′-H), 5.87-6.00 (1H, m, 8-H), 7.47-7.62 (4H, m, 5-H, ArH), 7.90-7.92 (2H, m, ArH), 8.39 (1H, d, J = 7.5 Hz, 6-H). 13C NMR (75 MHz, CDCl3) δ 12.6, 13.0, 13.2, 13.5 ((CH3)2)CHSi), 16.9, 17.0, 17.1, 17.4, 17.5 ((CH3)2)CHSi), 59.5 (5′), 67.8 (3′), 71.3 (7), 80.8 (2′), 81.9 (4′), 90.1 (1′), 96.1 (5), 117.2 (9), 127.5, 128.9 (Ar.), 133.0 (8, Ar.), 134.4 (6), 144.4 (2), 162.2 (4). IR (film; NaCl) ν 2946, 2868, 1699, 1667, 1620, 1553, 1488, 1264, 1126, 1040, 886 cm−1. FAB MS (3-NBA) m/z 630 (M+ + 1).
5.4.28 Current Protocols in Nucleic Acid Chemistry
3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)2′-O-(2,3-dihydroxypropyl)cytidine (S.12). TLC (19:1 CH2Cl2/CH3OH) Rf = 0.23; 1H NMR (360 MHz, CDCl3) δ (two diastereomers) 0.98-1.11 (28H, m, 4 (CH3)2)CHSi, 4 (CH3)2)CHSi), 3.66 (1H, dd, J = 5.2, 10.7 Hz, CH(OH)CH2aOH), 3.72-3.76 (2H, m, CH2CH(OH)CH2, CH(OH)CH2bOH), 3.884.06 (4H, m, 2′, 5′, OCH2CH(OH)), 4.15-4.28 (2H, m, 3′, 4′), 4.30 (1H, d, J = 13.5 Hz, 5′′), 5.83 (1H, s, 1′), 7.49-7.63 (4H, m, 5, Ar), 7.90-7.92 (2H, m, Ar), 8.31-8.34 (1H, m, 6), 8.94 (1H, br s, NH). 13C NMR (90 MHz, CDCl3) δ (two diastereomers) 12.5, 12.6, 12.9, 13.0 ((CH 3) 2CHSi), 16.8, 16.9, 17.0, 17.3, 17.4, 17.4, 17.6 ((CH2CHSi), 59.2 (5′), 63.6, 63.9 (CH(OH)CH2OH), 67.8 (3′), 70.4, 70.5 (OCH 2CH(OH)), 73.4, 74.3 (CH3)2CH(OH)CH2), 82.0 (2′), 83.1, 83.3 (4′), 89.9, 90.2 (1′), 96.3 (5), 127.6, 129.0, 132.8 (Ar), 133.3 (6), 144.3 (2), 162.5 (4). IR (film; NaCl) ν 3367, 2946, 2868, 1700, 1655, 1617, 1486, 1257, 1126, 1040, 886, 704 cm −1. MS (FAB, 3-NBA) m/z 664 (M + + 1). 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)N4-(benzoyl)-2′-O-(ethanal)cytidine (S.13). TLC (37:3 CH2Cl2/CH3OH) Rf = 0.50; 1H NMR (360 MHz, CDCl3) δ 0.98-1.11 (28H, m, 4 (CH3)2CHSi, 4 (CH3)2CHSi), 3.98-4.07 (2H, m, 2′, 5′), 4.18-4.31 (3H, m, 3′, 4′ ,5′′), 4.474.59 (2H, m, OCH2CHO), 5.86 (1H, s, 1′), 7.49-7.63 (4H, m, 5, Ar), 7.89-7.91 (2H, m, Ar), 8.37 (1H, d, J = 7.5 Hz, 6), 8.81 (1H, br s, NH), 9.81 (1H, s, CH2CHO). 13C NMR (90 MHz, CDCl3) δ 12.3, 12.9, 13.0, 13.4 ((CH3)2CHSi), 16.7, 16.9, 17.3, 17.4 ((CH3)2CHSi), 59.2 (5′), 68.1 (3′), 76.2 (OCH2CHO), 81.8 (2′), 83.0 (4′), 89.7 (1′), 96.2 (5), 127.5, 129.0, 132.8 (Ar), 133.2 (6), 144.3 (2), 162.4 (4), 200.7 (CH2CHO). IR (film; NaCl) ν 2946, 2868, 1700, 1667, 1619, 1484, 1253, 1130, 1064, 1040, 886, 703 cm−1. MS (FAB, 3-NBA) m/z 632 (M+ + 1). 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)N 4-(benzoyl)-2′-O-(2-hydroxyethyl)cytidine (S.14). TLC (37:3 CH2Cl2/CH3OH) Rf = 0.56. 1HNMR (360 MHz, CDCl ) δ 0.99-1.10 (28H, 3 m, 4 (CH3)2CHSi, 4 (CH3)2CHSi), 3.17 (1H, br s, CH2CH2OH), 3.72-3.77 (2H, m, CH2CH2OH), 3.93-4.03 (4H, m, 2′, 5′, OCH2CH2OH), 4.15-4.25 (2H, m, 3′, 4′), 4.30 (1H, d, J = 13.6 Hz, 5′′), 5.83 (1H, s, 1′), 7.49-7.63 (4H, m, 5, Ar), 7.89-7.92 (2H, m, Ar), 8.35 (1H, d, J = 7.5 Hz, 6), 8.92 (1H, br s, NH). 13C NMR (90 MHz, CDCl3) δ 12.6, 12.9, 13.0, 13.4 ((CH3)2CHSi), 16.8, 16.9,
17.0, 17.3, 17.4, 17.4 ((CH3)2CHSi), 59.3 (5′), 61.7 (CH2CH2OH), 68.0 (3′) , 7 3 .1 (OCH2CH2OH), 82.0 (2′), 82.3 (4′), 90.5 (1′), 96.2 (5), 127.5, 129.0, 132.9 (Ar), 133.2 (6), 144.3 (2), 162.4 (4). IR (film; NaCl) ν 2946, 2868, 1699, 1664, 1619, 1485, 1263, 1128, 1074, 1063, 1040, 886, 703 cm−1. MS (FAB, 3-NBA) m/z 634 (M+ + 1). 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)N4-(benzoyl)-2′-O-(ethyl-2-methylsul- fonate)cytidine (S.15). TLC (19:1 CH2Cl2/ CH3OH) Rf = 0.35. 1H NMR (360 MHz, CDCl3) δ 0.97-1.11 (28H, m, 4 (CH3)2CHSi, 4 (CH3)2CHSi), 3.14 (3H, s, CH3SO2), 3.97-4.01 (2H, m, 2′, 5′), 4.17-4.21 (4H, m, 3′, 4′, OCH2HH2OSO2), 4.30 (1H, d, J = 13.7 Hz, 5′′), 4.46-4.49 (2H, m, CH2CH2OSO2), 5.82 (1H, s, 1′), 7.50-7.62 (4H, m, 5, Ar), 7.89-7.92 (2H, m, Ar), 8.38 (1H, d, J = 7.5 Hz, 6). 13C NMR (90 MHz, CDCl3) δ 12.4, 12.9, 13.1, 13.4 ((CH3)2CHSi), 16.8, 16.9, 17.1, 17.3, 17.4, 17.4 ((CH3)2CHSi), 37.80 (CH3SO2), 59.3 (5′), 67.9 (3′), 69.0 (CH2CH2OSO2), 69.5 (OCH2 CH2OSO2), 81.9 (2′), 82.4 (4′), 89.5 (1′), 96.2 (5), 127.6, 129.1, 132.8 (Ar), 133.3 (6), 144.5 (2), 162.4 (4). IR (film; NaCl) ν 2945, 2868, 1664, 1484, 1170, 1128, 1074, 1063, 1040, 885, 704 cm−1 MS (FAB, 3-NBA) m/z 712 (M+ + 1). 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)N4-(benzoyl)-2′-O-(thiobenzoylethyl) cytidine (S.16). TLC (6:3:1 petroleum ether/EtOAc/CH3CN) Rf = 0.57. 1H NMR (360 MHz, CDCl3) δ 0.98-1.11 (28H, m, 4 (CH3)2CHSi, 4 (CH3)2CHSi), 3.35-3.43 (2H, m, CH2CH2SBz), 3.97-4.02 (2H, m, 2′, 5′), 4.09-4.13 (2H, m, OCH2CH2SBz), 4.18 (1H, dd, J = 3.9, 9.6 Hz, 3′), 4.25 (1H, dd, J = 2.0, 9.6 Hz, 4′), 4.30 (1H, d, J = 13.5 Hz, 5′′), 5.84 (1H, s, 1′), 7.40-7.62 (7H, m, 5, Ar), 7.91-7.99 (4H, m, Ar), 8.37 (1H, d, J = 7.4 Hz, 6), 8.97 (1H, br s, NH). 13C NMR (90 MHz, CDCl3) δ 12.5, 12.8, 13.1, 13.4 ((CH3)2CHSi), 16.8, 16.9, 17.1, 17.3, 17.4, 17.5 ((CH3)2CHSi), 29.1 (CH2CH2SBz), 59.4 (5′), 67.9 (3′), 69.9 (OCH2CH2SBz), 81.8 (2′), 81.9 (4′), 90.0 (1′), 96.0 (5), 127.2, 127.6, 128.5, 129.0, 133.0 (Ar), 133.2 (6), 137.1 (Ar), 144.6 (2), 162.3 (4), 191.5 (Ar). IR (film; NaCl) ν 2945, 2868, 1699, 1667, 1620, 1489, 1373, 1265, 1126, 1075, 1063, 1040, 691 cm−1. EI MS m/z 754 (M+ + 1). N4-(Benzoyl)-2′-O-(thiobenzoylethyl)cytidine (S.17). TLC (19:1 CH2Cl2/CH3OH) Rf = 0.21. 1H NMR (360 MHz, CDCl ) δ 3.25-3.42 (2H, 3 m, CH2SBz), 3.91-3.96 (2H, m, 2′, 5′), 4.054.19 (3H, m, 5′′, OCH2CH2SBz), 4.20-4.25
Methods for Cross-Linking Nucleic Acids
5.4.29 Current Protocols in Nucleic Acid Chemistry
Engineering Disulfide Cross-Links in RNA via Air Oxidation
(1H, m, 4′), 4.35-4.37 (1H, m, 3′), 5.87 (1H, s, 1′), 7.33-7.54 (7H, m, 5, Ar), 7.82-7.94 (4H, m, Ar), 8.57 (1H, d, J = 7.5 Hz, 6), 9.22 (1H, br s, NH). 13C NMR (90 MHz, CDCl3) δ 28.8 (CH2SBz), 59.8 (5′), 67.4 (3′), 69.4 (OCH2CH2SBz), 82.0 (2′), 84.6 (4′), 90.1 (1′), 96.7 (5), 126.8, 127.2, 127.6, 128.3, 128.6, 128.8, 132.9, 133.1 (Ar), 133.5 (6), 136.7 (Ar), 146.0 (2), 162.6 (4), 191.4 (Ar). IR (film; NaCl) ν 3345, 2928, 1699, 1658, 1617, 1558, 1485, 1379, 1258, 1111, 1067, 913, 705, 689 cm−1. MS (FAB, 3-NBA) m/z 512 (M+ + 1). N4-(Benzoyl)-2′-O-(ethyl)cytidine tert-butyl disulfide (S.18). TLC (24:1 CH2Cl2/ CH3OH) Rf = 0.29; 1H NMR (360 MHz, CDCl3) δ 1.32 (9H, s, C(CH3)3), 2.93 (2H, t, J = 5.8 Hz, CH2SS), 3.93-3.99 (2H, m, 2′, 5′), 4.09-4.18 (3H, m, 5′′, OCH2CH2SS), 4.24-4.30 (1H, m, 4′), 4.33-4.41 (1H, m, 3′), 5.84 (1H, d, J = 1.7 Hz, 1′), 7.48-7.62 (4H, m, 5, Ar), 7.87-7.89 (2H, m, Ar), 8.48 (1H, d, J = 7.5 Hz, 6), 9.01 (1H, br s, NH). 13C NMR (90 MHz, CDCl3) δ 29.8 (C(CH3)3), 40.3 (CH2SS), 48.0 (C(CH3)3), 60.3 (5′), 67.8 (3′), 69.0 (OCH2CH2SS), 81.7 (2′), 85.0 (4′), 90.9 (1′), 96.7 (5), 127.6, 129.0, 132.9 (Ar), 133.3 (6), 146.4 (2), 162.5 (4). IR (film; NaCl) ν 3374, 2960, 2922, 1699, 1648, 1617, 1558, 1487, 1379, 1260, 1110 cm−1. FAB MS (3-NBA) m/z 496 (M+ + 1). 5′-O-(4, 4′-Dimethoxytrityl)-N4)-benzoyl)2′-O-(ethyl)cytidine tert-butyl disulfide (S.19). TLC (1:1 acetone/petroleum ether) Rf = 0.23; 1H NMR (360 MHz, CD CN) δ 1.30 (9H, s, 3 C(CH3)3), 2.95 (2H, t, J = 6.3 Hz, CH2SS), 3.39-3.44 (2H, m, OCH2CH2SS), 3.76 (6H, s, 2 OCH3), 3.90-4.03 (3H, m, 2′, 5′, 5′′), 4.124.19 (1H, m, 4′), 4.39-4.47 (1H, m, 3′), 5.84 (1H, d, J = 1.7 Hz, 1′), 6.87-6.89 (4H, m, Ar), 7.15-7.61 (13H, m, 5, Ar), 7.92-7.95 (2H, m, Ar), 8.46 (1H, d, J = 7.6 Hz, 6). 13C NMR (90 MHz, CD3CN) δ 30.2 (C(CH3)3), 41.0 (CH2SS), 48.5 (C(CH3)3), 56.0 (OCH3), 62.1 (5′), 68.9 (3′), 70.1 (OCH2CH2SS), 83.3 (2′), 83.6 (4′), 87.7 (OC(Ph)3), 90.2 (1′), 97.2 (5), 114.3, 128.1, 129.1, 129.1, 129.2, 129.7, 131.0, 131.2, (Ar), 133.9 (6), 134.5, 136.6, 137.0, 145.5 (Ar), 145.9 (2), 155.7, 159.8 (Ar), 163.9 (4), 168.2 (Ar). IR (KBr) ν 3392, 2959, 2924, 1700, 1667, 1610, 1553, 1510, 1482, 1377, 1252, 1113, 1033, 704 cm−1. MS (FAB, 3NBA) m/z 798 (M+ + 1). 5′-O-(4, 4′-Dimethoxytrityl)-N4)-benzoyl)2′-O-(ethyl)cytidine tert-butyl disulfide 3′-O(N,N-diisopropyl β cyanoethylphosphoramidite) (S.20). TLC (2:1 petroleum ether/acetone) Rf = 0.40; 1H NMR (300 MHz,
CD3CN) δ (two diastereomers) 1.05-1.20 (12H, m, 2 NCH(CH3)2), 1.30 (9H, s, C(CH3)3), 2.50-2.67 (2H, m, OCH2CH2CN), 2.94-3.01 (2H, m, CH2SS), 3.41-3.71 (6H, m, OCH2CH2SS, OCH2CH2CN, 2 NCH(CH3)2), 3.79 (6H, s, 2 OCH3), 3.80-4.22 (4H, m, 2′, 4′, 5′, 5′′), 4.43-4.63 (1H, m, 3), 5.88 (1H, s, 1′), 6.87-6.92 (4H, m, Ar), 7.03-7.66 (13H, m, 5, Ar), 7.91-7.97 (2H, m, Ar), 8.42-8.55 (1H, 2 d, J = 7.6 Hz, 6). 13C NMR (75 MHz, CD3CN) δ (two diastereomers) 21.2, 21.3 (OCH2 CH2CN), 24.9, 25.0, 25.2, 25.3 (NCH(CH3)2), 30.3 (C(CH3)3), 41.4 (CH2SS), 44.1, 44.3 (NCH(CH3)2), 48.5 (C(CH3)3), 56.0 (OCH3), 59.3, 59.6 (OCH2CH2CN), 61.8, 62.2 (5′), 70.2, 70.4 (3′), 70.8 (OCH2CH2SS), 82.2 (2′), 83.1 (4′), 87.8 (OC(Ph)3), 90.8, 91.1 (1′), 97.3 (5), 114.2, 128.1, 129.0, 129.0, 129.3, 129.6, 131.2 (Ar), 133.8 (6), 134.5, 136.6, 136.7, 145.4 (Ar), 145.6 (2), 155.4, 159.8 (Ar), 163.6 (4). 31P NMR (202 MHz; CD3CN) δ 147.57, 146.51. IR (film; NaCl) ν 2967, 2934, 1708, 1686, 1509, 1510, 1462, 1251, 1180, 1035, 979 cm−1. MS (FAB, 3-NBA) m/z 998 (M+ + 1). Anal. Calcd for C52H64N5O9PS2: C, 62.56; H, 6.48; N, 7.02. Found: C, 62.41; H, 6.41; N, 6.92. 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)2′-O-allyluridine (S.21). TLC (1:1 petroleum ether/EtOAc) Rf 0.59; 1H NMR (360 MHz, CDCl3) δ 1.01-1.11 (28H, m, isopropyl), 3.88 (1H, m, 4′), 3.98 (1H, dd, OCH2a), 4.18 (2H, m, 2′,3′), 4.26 (1H, dd, OCH2b), 4.39 (2H, m, 5′,5′′), 5.21 (1H, ddd, =CH2a), 5.40 (1H, ddd, =CH2b), 5.68 (1H, dd, 5), 5.76 (1H, s, 1′), 5.93 (1H, m, CH=), 7.93 (1H, d, 6), 9.14 (1H, s, NH); 13C NMR (90 MHz, CDCl ) δ 12.68-13.63 3 (isopropyl CHs), 17.00-17.70 (isopropyl CH3s), 59.62 (5′), 68.25 (3′), 71.46 (OCH2), 81.27 (2′), 82.00 (4′), 89.29 (1′), 101.69 (5), 117.59 (=CH2), 134.51 (CH=), 139.77 (6), 150.06 (2), 163.61 (4); MS (DCI, CH4) m/z 527 (M+). 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)2′-O-(2,3-dihydroxypropyl)uridine (S.22). TLC (51:4 CH2Cl2/CH3OH) Rf = 0.22; 1H NMR (360 MHz, CDCl3) δ 1.02-1.10 (28H, m, 4 SiCH(CH3)2), 3.65-3.75 (3H, m, CH2CH(OH)CH2OH, CH2CH(OH)CH2OH), 3.86-4.04 (4H, m, 2′, 5′′, CH2CH(OH) CH2OH), 4.12-4.17 (2H, m, 3′, 4′), 4.25 (1H, d, J = 13.6 Hz, 5′), 5.71 (1H, dd, J = 1.4, 8.1 Hz, 5), 5.74 (1H, d, J = 1.6 Hz, 1′), 7.88 (1H, dd, J = 1.9, 8.1 Hz, 6). 13C NMR (75 MHz, CDCl3) δ 13.07-13.90 (SiCH(CH3)2), 17.1817.82 (SiCH(CH3)2), 59.66 (5′), 68.58 (3′), 70.94 (CH2CH(OH)CH2OH), 74.20 (CH2CH (OH)CH2OH), 82.22 (4′), 83.99 (CH2CH
5.4.30 Current Protocols in Nucleic Acid Chemistry
(OH)CH2OH), 89.43 (1′), 89.67 (2′), 102.33 (5), 139.24 (6), 150.82 (2), 163.33 (4). IR (Nujol) ν 3424, 3160, 3092, 2951, 2946, 2935, 2925, 2918, 2909, 2896, 2891, 2873, 2867, 2863, 2856, 2850, 2726, 1693, 1685, 1462, 1378, 1272, 1163, 1125, 1062, 1035, 1010, 991, 884, 703 cm−1. 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)2′-O-(ethanal)uridine (S.23) TLC (51:5 CH2Cl2/CH3OH) Rf = 0.48; 1H NMR (300 MHz, CDCl3) δ 1.03-1.12 (28 H, m, 4 SiCH(CH3)2), 3.71 (2 H, s, OCH2CHO), 3.954.01 (2 H, m, 2′, 5′′), 4.23-4.30 (2 H, m, 3′,4′), 4.45-4.47 (1 H, m, 5′ ), 5.70 (1 H, dd, J = 2.2, 8.1 Hz, 5), 5.78 (1 H, s, 1′), 7.90 (1 H, d, J = 8.1 Hz, 6), 8.93 (1 H, s, NH). 13C NMR (75 MHz, CDCl3) δ 12.87-13.82 (SiCH(CH3)2), 17.11-17.78 (SiCH(CH3)2), 59.64 (5′), 68.95 (3′), 76.62 (OCH2CHO), 82.00 (4′), 83.85 (2′), 89.32 (1′), 102.03 (5), 139.41 (6), 150.18 (2), 163.21 (4), 200.35 (OCH2CHO). IR (KBr) ν 3482, 2947, 2868, 1700, 1685, 1464, 1386, 1272, 1256, 1166, 1131, 1096, 1063, 1056, 1038, 1012, 991, 886, 704 cm−1. 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)2′-O-(2-hydroxyethyl)uridine (S.24). TLC (6:3:1 petroleum ether/EtOAc/CH3CN) Rf = 0.34; 1H NMR (360 MHz, CDCl3) δ 1.01-1.10 (28H, m, 4 SiCH(CH3)2), 3.74-3.76 (2H, m, CH2CH2OH), 3.92-4.00 (4H, m, 2′, 5′, CH2CH2OH), 4.17 (2H, m, 3′, 4′), 4.26 (1H, d, J = 13.7 Hz, 5′), 5.70 (1H, d, J = 8.1 Hz, 5), 5.75 (1H, s, 1′), 7.90 (1H, d, J = 8.1 Hz, 6), 9.96 (1H, s, NH). 13C NMR (90 MHz, CDCl3) δ 12.73-13.62 (SiCH(CH3)2), 16.97-17.65 (SiCH(CH3)2), 59.46 (5′), 61.78 (OCH2 CH2OH), 68.37 (3′), 73.20 (OCH2CH2OH), 81.99 (4′), 83.04 (2′), 89.62 (1′), 101.99 (5), 139.38 (6), 150.58 (2), 163.80 (4). IR (KBr) ν 2946, 2869, 1701, 1464, 1387, 1271, 1165, 1129, 1094, 1062, 1039, 992, 885, 705 cm−1. MS (CI, CH4) m/z 531 (M+). 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)2′-O-(ethyl-2-methylsulfonate)uridine (S.25) TLC (17:3 CH2Cl2/acetone) Rf = 0.48; 1H NMR (360 MHz, CDCl3) δ 1.03-1.12 (28H, m, 4 SiCH(CH3)2), 3.12 (3H, s, SCH3), 3.89 (1H, d, J = 4.1 Hz, 2′), 3.97 (1H, dd, J = 2.2, 13.6 Hz, 5′), 4.10-4.15 (3H, m, 4′, OCH2CH2S), 4.204.23 (1H, m, 3′), 4.26 (1H, d, J = 13.6 Hz, 5′′), 4.44-4.46 (2H, m, OCH2CH2S), 5.70 (1H, dd, J = 2.1, 8.2 Hz, 5), 5.73 (1H, s, 1′), 7.91 (1H, d, J = 8.2 Hz, 6), 8.95 (1H, s, NH). 13C NMR δ 12.85-13.74 (75 M Hz, CDCl3) (SiCH(CH3)2), 17.07-17.73 (SiCH(CH3)2), 38.00 (SCH3), 59.59 (5′), 68.63 (3′), 69.29 (OCH2CH2OMs), 69.44 (OCH2CH2OMs),
81.93 (4′), 83.27 (2′), 88.96 (1′), 101.97 (5), 139.38 (6), 150.33 (2), 163.60 (4). IR (KBr) ν 3213, 3065, 2947, 2869, 1699, 1465, 1355, 1272, 1179, 1129, 1095, 1065, 1040, 993, 885, 705 cm−1. 3′,5′-O-(Tetraisopropyldisiloxane-1,3-diyl)2′-O-(thiobenzoylethyl)uridine (S.26). TLC (4:1 CH2Cl2/acetone) Rf = 0.68; 1H NMR (360 MHz, CDCl3) δ 1.00-1.10 (28H, m, 4 SiCH(CH3)2), 3.36 (2H, m, OCH2CH2S), 3.91 (1H, d, J = 3.3 Hz, 2′), 3.99 (1H, dd, J = 1.5, 13.6 Hz, 5′), 4.02-4.06 (2H, m, OCH2CH2S), 4.18 (2H, m, 3′, 4′), 4.25 (1H, d, J = 13.6 Hz, 5′′), 5.68 (1H, dd, J = 1.7, 8.2 Hz, 5), 5.73 (1H, s, 1′), 7.44 (2H, m, Ar), 7.56 (1H, m, Ar), 7.90 (1H, d, J = 8.2 Hz, 6), 7.96 (1H, m, Ar), 9.47 (1H, s, NH). 13C NMR (90 MHz, CDCl3) δ 12.71-13.62 (SiCH(CH3)2), 17.00-17.70 (SiCH(CH3)2), 29.16 (OCH2CH2S), 59.57 (5′), 68.45 (3′), 70.12 (OCH2CH2S), 81.88 (4′), 82.60 (2′), 89.34 (1′), 101.71 (5), 127.45, 128.74, 133.53, 137.18 (Ar), 139.73 (6), 150.14 (2), 163.92 (4), 191.67 (CO). IR (KBr) ν 3205, 3058, 2945, 2868, 1699, 1463, 1387, 1271, 1207, 1164, 1126, 1093, 1064, 1040, 914, 885, 705, 690 cm−1. 2′-O-(Thiobenzoylethyl)uridine (S.27). TLC (9:1 CH2Cl2/CH3OH) Rf = 0.14; 1H NMR (360 MHz, CD3OD) δ 3.27-3.38 (2H, m, OCH2CH2S), 3.75 (1H, dd, J = 3.0, 12.4 Hz, 5′), 3.82-3.96 (2H, m, 2′, 5′′), 3.99-4.02 (1H, m, 4′), 4.05-4.09 (2H, m, OCH2CH2S), 4.24 (1H, m, 3′), 5.67 (1H, d, J = 8.1 Hz, 5), 5.95 (1H, d, J = 3.7 Hz, 1′), 7.49 (2H, m, ArH), 7.62 (1H, m, ArH), 7.93 (2H, m, ArH), 8.06 (1H, d, J = 8.1 Hz, 6). 13C NMR (90 MHz, CD3OD) δ 29.70 (ΟΧΗ2CH2S), 61.77 (5′), 70.01 (3′), 70.65 (OCH2CH2S), 83.80 (4′), 86.26 (2′), 89.31 (1′), 102.74 (5), 128.27, 130.04, 134.94, 138.30 (Ar), 142.58 (6), 152.35 (2), 166.36 (4), 193.18 (CO). IR (KBr) ν 3473, 3422, 3168, 3107, 3051, 2954, 2916, 2874, 2860, 2572, 2540, 1693, 1672, 1647, 1615, 1446, 1388, 1267, 1209, 1143, 1111, 1098, 915, 770, 687 cm−1. MS (CI, CH4) m/z 409 (M+ + 1). 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(thiobenzoylethyl)uridine (S.28). TLC (19:1 CH2Cl2/CH3OH) Rf = 0.59. 1H NMR (360 MHz, CD3CN) δ 3.30-3.41 (5H, m, 2′, 5′, 5′′, OCH2CH2S), 3.75 (6H, s, 2 OCH3), 3.87-3.99 (3H, m, 4′, OCH2CH2S) 4.01-4.04 (1H, m, 3′), 5.24 (1H, d, J = 8.2 Hz, 5), 5.83 (1H, d, J = 1.7 Hz, 1′), 6.87 (4H, m, ArH), 7.21-7.33 (9H, m, ArH), 7.41-7.52 (5H, m, ArH), 7.61-7.66 (1H, m, ArH), 7.74 (1H, d, J = 8.2 Hz, 6), 7.93-7.96 (4H, m, ArH). 13C NMR (90 MHz, CD3CN) δ
Methods for Cross-Linking Nucleic Acids
5.4.31 Current Protocols in Nucleic Acid Chemistry
Engineering Disulfide Cross-Links in RNA via Air Oxidation
17.03 (OCH2CH2S), 56.02 (OCH3), 63.11 (5′), 69.86 (3′), 70.28 (OCH2CH2S), 83.21 (2′), 84.04 (4′), 87.65 (OC(Ph)3), 88.68 (1′), 102.55 (5), 114.24 (Ar), 128.06, 129.02, 129.14, 129,98, 131.17, 131.64, 134.81, 136.49, 136.73, 137.89 (Ar), 141.17 (6), 145.87 (Ar), 151.41 (2), 159.86 (Ar), 161.07 (4), 192.35 (SC(O)Ph). MS (CI, CH4) m/z 710 (M+). 5′-O-(4,4′-Dimethoxytrityl)-2′-O-(ethyl)uridine tert-butyl disulfide (S.29) TLC (49:1 CH2Cl2/CH3OH) Rf = 0.12; 1H NMR (360 MHz, acetone-d6) δ 1.32 (9H, s, C(CH3)3) 3.00 (2H, t, J = 6.5 Hz, OCH2CH2S), 3.48 (2H, m, 5′, 5′′), 3.80 (6 H, s, OCH3), 3.94-4.07 (4H, m, 2′, 3′, OCH2CH2S), 4.15-4.17 (1H, m 4′), 5.29 (1H, d, J = 8.1 Hz, 5), 5.94 (1H, d, J = 2.5 Hz, 1′), 6.90-6.93 (4H, m, ArH), 7.32-7.37 (7H, m, ArH), 7.47-7.50 (2H, m, ArH), 7.92 (1H, d, J = 8.1 Hz, 6). 13C NMR (90 MHz, acetone-d6) δ 29.28-30.56 (acetone, C(CH3)3), 40.79 (OCH2CH2S), 48.32 (C (CH3)3), 55.61 (OCH3), 63.15 (5′), 69.94 (3′), 70.16 (OCH2CH2S), 83.30 (2′), 84.03 (4′), 87.58 (Ar), 88.75 (1′), 102.41 (5), 114.09, 127.83, 128.82, 129.13, 131.14, 136.37, 136.67 (Ar), 140.88 (6), 145.93 (Ar), 151.29 (2), 159.80 (Ar), 163.58 (4). MS (FAB, 3-NBA) m/z 695 (M+ + 1). 3′-O-(N,N-Diisopropyl-β-cyanoethylphosphoramidite)-5′-O-(4,4′-dimethoxytrityl)-2O-(ethyl)uridine tert-butyl disulfide (S.30). TLC (2:1 petroleum ether/acetone) Rf = 0.34; 1H NMR (360 MHz, CD CN) δ (two dias3 tereomers) 1.04-1.06 (4H, m, 2 NCH(CH3)2), 1.14-1.18 (12H, m, 2 NCH(CH3)2), 1.30, 1.31 (9H, 2 s, SSC(CH3)3), 2.51-2.69 (2H, m, OCH2CH2C N ) , 2 . 8 9 - 2 . 9 4 ( 2 H , m , 3.37-3.44 (2H, m, OCH2CH2S ) , OCH2CH2CN), 3.58-3.66 (2H, m, 5′, 5′′), 3.76, 3.77 (6H, 2 s, OCH3), 3.80-3.95 (2H, m, OCH2CH2S), 4.06-4.12 (2H, m, 2′,4′), 4.40-4.53 (1H, m, 3′), 5.20, 5.21 (1H, 2 d, J = 8.1 Hz, 5), 5.85 (1H, m, 1′), 6.86-6.90 (4H, m, ArH), 7.25-7.36 (7H, m, ArH), 7.42-7.47 (2H, m ArH), 7.73,7.82 (1H, 2 d, J = 8.1 Hz, 6). 13C NMR (75 MHz, acetone-d6) δ (two diastereomers) 20.96 (NCH(CH3)2), 25.0225.32 (NCH(CH3)2), 29.14-30.68 (acetone, SSC(CH3)3), 41.17 (OCH2CH2S), 44.07, 44.24 (CH2CN), 48.27 (SSC(CH3)3), 55.68 (OCH3), 59.25-59.92 (POCH2), 62.72, 63.00 (5′), 70.27, 70.41 (OCH2CH2S), 71.30, 71.48 (3′), 82.11, 82.81 (2′), 83.14-83.41 (4′), 87.78 (Ar), 89.04, 89.29 (1′), 102.53, 102.59 (5), 114.10 (Ar), 118.81, 118.96 (CN), 127.80, 128.72, 129.19, 131.15, 131.53, 136.27, 136.35, 136.45 (Ar), 140.65, 140.72 (6),
145.68, 145.75 (Ar), 151.21 (2), 159.80 (Ar), 163.37 (4). 31P NMR (202 MHz, CD3CN) δ 147.44, 146.90. IR (KBr) ν 3199, 3059, 2966, 2932, 2871, 2837, 2248, 1692, 1608, 1615, 1510, 1458, 1364, 1253, 1179, 1119, 1085, 1070, 1035, 980, 828, 810 cm−1. MS (FAB, 3-NBA) m/z 895 (M+ +1). Anal. Calcd for C45H59N4O9PS2: C, 60.38. H, 6.64. N, 6.26. Found: C, 60.59. H, 6.59. N, 5.95.
Critical Parameters There are three critical parameters for the formation of intramolecular disulfide bonds in RNA. First, prior to oxidation the reducing agent used to remove the thiol protecting groups, DTT, must be entirely removed from sample. Second, the concentration of the RNA must be very low (1 to 4 µM) to avoid intermolecular cross-link formation. Third, for efficient cross-link formation the pH of the solution must be maintained between 8.0 and 8.4. At pH values <8.0 the concentration of thiolate ion will be reduced and the rate of cross-link formation will be reduced. However, prolonged exposure of the RNA to pH values >8.4 may lead to degradation.
Troubleshooting See Table 5.4.1 for discussion of common problems encountered with these procedures and methods to identify, solve, and avoid them.
Anticipated Results The authors have achieved 100% conversion of tert-butyl disulfide–protected tRNAs to intramolecularly disulfide-cross-linked tRNAs at three different sites of disulfide incorporation. Similar results should be obtained for thiols positioned at proximal locations either in a folded tertiary structure or within secondary structure. The solid-phase synthesis of tRNAs containing the thiol-modified nucleosides described here results in yields of 0.5 mg tRNA after purification to single-nucleotide resolution. Each thiol-modified nucleoside is incorporated into the RNA oligomer as efficiently as commercially available phosphoramidites.
Time Considerations Conversion of the tert-butyl disulfide–protected RNA to disulfide-cross-linked RNA takes 36 to 48 hr; both the reduction of the tert-butyl disulfide protecting groups and the air oxidation can be done as overnight reactions. To ensure complete reduction of the tert-butyl disulfide protecting groups the
5.4.32 Current Protocols in Nucleic Acid Chemistry
Table 5.4.1
Troubleshooting Guide for Disulfide Cross-Linking
Problem
Possible causes
RNA does not cross-link or does not cross-link quantitatively
Protecting groups were not completely Add more DTT and/or incubate longer. reduced After DTT removal, thiols on the RNA can be quantified using the fluorescent assay. DTT was not completely removed Dialyze at a slower rate; use more before oxidation commenced buffer; monitor the dialysate with the fluorescence assay to determine when all DTT has been removed pH of the reaction was <8.0 Adjust pH to 8.0 RNA was not properly folded before Fold the RNA oxidation commenced RNA was not pure Repurify the starting thiol-modified RNA Thiols are not positioned close enough Select alternate positions for to react modification Resulting cross-linked RNA would be Select alternate positions for a strained, high-energy structure modification Moisture was present in Dry amidites by evaporation from phosphoramidite freshly distilled CH3CN several times Handle amidites under Ar Repurify the phosphoramidite Dilute the amidite with CH3CN immediately before coupling instead of at the beginning of the synthesis
Modified nucleosides incorporate poorly during RNA synthesis
RNA should be incubated with DTT for ≥12 hr, while >14 hr appears unnecessary. The slow rate of dialysis (5 mL/min), using 3 L of buffer over 10 hr, ensures that the DTT is completely removed and that the buffer exchange into the lower-salt buffer is complete. Cross-link formation between thiols positioned at locations that are proximal in the folded RNA generally occurs in <12 hr. It has taken up to 48 hr for completion of some cross-linking reactions; in such cases the pH is monitored periodically and adjusted as necessary, and water is also added to compensate for evaporation. Synthesis and purification of thiol-modified tRNA takes between 1 and 2 weeks depending on the amount of RNA to be purified before proceeding with the cross-linking reaction. This includes 1 day to synthesize the necessary phosphoramidites before commencing the solid-phase synthesis.
Solution
Literature Cited Allerson, C.R., Chen, S.L., and Verdine, G.L. 1997. A chemical method for site-specific modification of RNA: The convertible nucleoside approach. J. Am. Chem. Soc. 119:7423-7433. Allerson, C.R. and Verdine, G.L. 1995. Synthesis and biochemical evaluation of RNA containing an intrahelical disulfide crosslink. Chem. Biol. 2:667-675. Breslauer, K.J., Frank, R., Blöcker, H., and Markey, L.A. 1986. Predicting DNA duplex stability from the base sequence. Proc. Natl. Acad. Sci. U.S.A. 83:3746-3750. Cohen, S.B. and Cech, T.R. 1997. Dynamics of thermal motions within a large catalytic RNA investigated by crosslinking with thiol-disulfide interchange. J. Am. Chem. Soc. 119:6259-6268. Douglas, M.E., Beijer, B., and Sproat, B.S. 1994. An approach towards thiol mediated labelling in the minor groove of oligonucleotides. Bioorg. Med. Chem. Lett. 4:995-1000. Earnshaw, D.J., Masquida, B., Müller, S., Sigurdsson, S.T., Eckstein, F., Westhof, E., and Gait, M.J. 1997. Inter-domain cross-linking and molecular modelling of the hairpin ribozyme. J. Mol. Biol. 274:197-212.
Methods for Cross-Linking Nucleic Acids
5.4.33 Current Protocols in Nucleic Acid Chemistry
Gait, M.J., Pritchard, C., and Slim, G. 1991. Oligoribonucleotide synthesis. In Oligonucleotides and Analogues. A Practical Approach (F. Eckstein, ed.) pp. 25-48. Oxford University Press, Oxford. Glick, G.D. 1991. Synthesis of a conformationally restricted DNA hairpin. J. Org. Chem. 56:67466747. Glick, G.D. 1998. Design, synthesis, and analysis of conformationally restricted nucleic acids. Biopolymers 48:83-96. Goodwin, J.T., Osborne, S.E., Scholle, E.J., and Glick, G.D. 1996. Design, synthesis, and analysis of yeast tRNAPhe analogs possessing intraand interhelical disulfide cross-links. J. Am. Chem. Soc. 118:5207-5214.
Schaller, H., Weimann, G., Lerch, B., and Khorana, H.G. 1963. Studies on polynucleotides. XXIV. The stepwise synthesis of specific deoxyribopolynucleotides (4). Protected derivatives of deoxyribonucleosides and new syntheses of deoxyribonucleoside-3′ phosphates. J. Am. Chem. Soc. 85:3821-3827. Sigurdsson, S.T., Tuschl, T., and Eckstein, F. 1995. Probing RNA tertiary structure: Interhelical crosslinking of the hammerhead ribozyme. RNA 1:575-583. Sigurdsson, S.T. and Eckstein, F. 1996. Isolation of oligoribonucleotides containing intramolecular cross-links. Anal. Biochem. 235:241-242.
Goodwin, J.T., Stanick, W.A., and Glick, G.D. 1994. Improved solid-phase synthesis of long oligoribonucleotides: Application to tRNAPhe and tRNAGly. J. Org. Chem. 59:7941-7943.
Sproat, B.S. and Lamond, A.I. 1991. 2′-O-Methyloligoribonucleotides: Synthesis and application. In Oligonucleotides and Analogues. A Practical Approach (F. Eckstein, ed.) pp. 49-86. Oxford University Press, Oxford.
Gundlach, C.W. IV, Ryder, T.R., and Glick, G.D. 1997. Synthesis of guanosine analogs bearing pendant alkylthiol tethers. Tetrahedron Lett. 38:4039-4042.
Sproat, B.S., Iribarren, A., Beijer, B., Pieles, U., and Lamond, A.I. 1991. 2′-O-alkyloligoribonucleotides: Synthesis and applications in studying RNA splicing. Nucleosides Nucleotides 10:25-36.
Maglott, E.J. and Glick, G.D. 1998. Probing structural elements in RNA using engineered disulfide bonds. Nucl. Acids Res. 26:1301-1308.
Sun, S., Tang, X.-Q., Merchant, A., Anjaneyulu, P.S.R., and Piccirilli, J.A. 1996. Efficient synthesis of 5-(thioalkyl)uridines via ring opening of α-ureidomethylene thiolactones. J. Org. Chem. 61:5708-5709.
Manoharan, M., Johnson, L.K., Tivel, K.L., Springer, R.H., and Cook, P.D. 1993. Introduction of a lipophilic thioether tether in the minor groove of nucleic acids for antisense applications. Bioorg. Med. Chem. Lett. 3:2765-2770. Parvari, R., Pecht, I., and Soreq, H. 1983. A microfluorimetric assay for cholinesterases, suitable for multiple kinetic determinations of picomoles of released thiocholine. Anal. Biochem. 133:450-456.
Wünsch, E., Moroder, L., and Romani, S. 1982. 1-(tert-Butylthio)-1,2-hydrazinedicarboxoylic acid derivatives. New reagents for the introduction of the S-tert-butylthio group into cysteine and cysteine derivatives. Hoppe-Seylers Z. Physiol. Chem. 363:1461-1464.
Contributed by Emily J. Maglott and Gary D. Glick University of Michigan Ann Arbor, Michigan
Engineering Disulfide Cross-Links in RNA via Air Oxidation
5.4.34 Current Protocols in Nucleic Acid Chemistry
Use of Electrophilic Substitution to Form Site-Specific Cross-Links in DNA
UNIT 5.5
This unit describes the incorporation of a nucleophilic base into an oligodeoxyribonucleotide (ODN), and the use of electrophilic tethers to convert this ODN into an electrophilic probe capable of cross-linking a complimentary DNA strand (see Basic Protocol). The postsynthetic method that generates the electrophilic ODN allows the introduction of functionality not compatible with standard solid-phase synthetic conditions. It also allows the generation of a variety of different electrophilic DNA strands from the same deoxythiouridine-containing ODN by changing the nature of the electrophilic tether. In addition, the site of modification places the tether in intimate contact with the major groove, allowing structural features of the groove to aid in directing the electrophilic attack. The synthesis of the electrophile, N,N′-bis-bromoacetyl-1,2-diaminobenzene, is also described in this unit (see Support Protocol). FORMATION OF SITE-SPECIFIC CROSS-LINKS IN DNA The nucleophilic base used in these studies, 4-thio-2′-deoxyuridine (dS4U), is incorporated into an ODN using an S-(2-cyanoethyl)-protected phosphoramidite and standard solidphase synthesis conditions and reagents (Coleman and Siedlecki, 1991). The DNA is left on the column, and the S-cyanoethyl group is removed by treatment with 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) in dry acetonitrile. The ODN is then cleaved from the column and purified by a two-step process involving an initial trityl-on purification followed by a trityl-off purification. Cross-linking is performed by derivatizing the modified ODN with the desired electrophilic tether and incubating this probe with a 32 P-labeled target (Fig. 5.5.1; Coleman and Kesicki, 1995; Coleman and Pires, 1997).
BASIC PROTOCOL
Materials S-(2-Cyanoethyl)-protected 4-thio-2′-deoxyuridine (Glen Research) Dry solvent-grade acetonitrile 1.0 M DBU solution (see recipe) 50 mM NaSH/NH4OH solution (see recipe) Nitrogen gas (optional) HPLC buffer A (see recipe) HPLC buffer B: HPLC-grade acetonitrile 80% (v/v) distilled glacial acetic acid Dry ice
S
O NH
1) O
HO Nx O P O O–
N
O
NH HN Br
Br
NH
O Br
O
N
NH
O 2) complimentary DNA
S H
–
N
N
O O P O Ny OH
N
O
H N
dna
N N
H
O
O
dna
Figure 5.5.1 Cross-linking between 4-thio-2′-deoxyuridine and N,N′-bis-bromoacetyl-1,2diaminobenzene.
Methods for Cross-Linking Nucleic Acids
Contributed by Richard M. Pires and Robert S. Coleman
5.5.1
Current Protocols in Nucleic Acid Chemistry (2001) 5.5.1-5.5.6 Copyright © 2001 by John Wiley & Sons, Inc.
Supplement 6
Double-deionized water 2 mg/mL (1.5 U/mg) snake venom phosphodiesterase (SVP) from Crotalus durissus 1 U/µL alkaline phosphatase (AP) from calf intestine 0.1 M potassium phosphate buffer, pH 8.0 (APPENDIX 2A) Electrophile: N,N′-bis-bromoacetyl-1,2-diaminobenzene (see Support Protocol) Dimethylformamide (DMF) n-Butanol Radiolabeled complimentary DNA solution (1 to 5 OD/mL) DNA synthesizer with bottle (and septum) that fits a phosphoramidite inlet port Vacuum pump Desiccator Centrifugal evaporator Binary high-performance liquid chromatograph (HPLC) equipped with UV detector and recorder Polystyrene reversed-phase (PRP-1) HPLC column (Hamilton) Spectrophotometer Small spin column containing 0.3 ml Sephadex G-25 equilibrated with double-deionized water Additional reagents and equipment for DNA synthesis (APPENDIX 3C), SVP/AP digestion (UNIT 10.6), and denaturing polyacrylamide gel electrophoresis (PAGE; APPENDIX 3B) Synthesize dS4U-containing ODN 1. Dissolve 50 mg S-(2-cyanoethyl)-protected 4-thio-2′-deoxyuridine in 2 mL dry solventgrade acetonitrile in a bottle that fits a phosphoramidite inlet port of a DNA synthesizer. Use an empty bottle that previously contained a phosphoramidite standard. Wash and dry bottle thoroughly before reuse.
2. Place a septum on the bottle, and evaporate sample to dryness using a vacuum pump. Repeat this azeotropic drying procedure two times. 3. Add enough dry solvent-grade acetonitrile to prepare a 0.1 M phosphoramidite solution, and place the bottle on phosphoramidite port no. 5 of the synthesizer. Set the synthesizer to perform a standard synthesis (APPENDIX 3C), except do not remove the trityl group of the last base, and leave the DNA attached to the column. A 200-nmol or 100-mmol synthesis is appropriate.
4. After synthesis is complete, evaporate remaining phosphoramidite solution to dryness as in step 2, and store at −20°C in a desiccator (stable indefinitely). This storage method substantially increases the shelf life of the phosphoramidite.
5. Remove the column containing DNA from the synthesizer, and cleave the S-cyanoethyl group by passing 2 mL of 1.0 M DBU solution through the column over 2 hr using the double-syringe technique. The double-syringe technique is performed by placing a syringe at each end of the column and passing the reagent back and forth slowly. Use of Electrophilic Substitution to Form Site-Specific Cross-Links in DNA
6. Flush column two times with 3 mL dry solvent-grade acetonitrile. Residual DBU interferes with the purification of DNA.
7. Cleave DNA from column with 5 mL of 50 mM NaSH/NH4OH solution over 3 hr at room temperature using the double-syringe technique.
5.5.2 Supplement 6
Current Protocols in Nucleic Acid Chemistry
8. Allow sample to sit 16 hr at room temperature in 50 mM NaSH/NH4OH solution to remove the base-protecting groups. 9. Evaporate sample to dryness, either with a stream of nitrogen gas or under vacuum using a centrifugal evaporator. Purify dS4U-containing ODN 10. Redissolve sample in 100 µL HPLC buffer A and inject onto a PRP-1 column. Perform reversed-phase HPLC using 95% HPLC buffer A to 50% HPLC buffer A over 30 min at a flow rate of 1 mL/min. NaSH elutes from the column in ∼3 min and has a horrible odor. The untritylated failure sequences elute at ∼10 min, whereas the full-length tritylated product elutes at ∼20 min.
11. Collect purified product and evaporate to dryness using a centrifugal evaporator. Redissolve in 1 mL of 80% distilled glacial acetic acid. Allow sample to sit 20 min at room temperature. 12. Freeze sample using dry ice and evaporate to dryness under vacuum using a centrifugal evaporator. 13. Redissolve sample in 100 µL HPLC buffer A. The solution may turn cloudy, because the trityl alcohol produced is relatively insoluble in HPLC buffer A. If it turns cloudy, centrifuge to remove solid material.
14. Inject sample onto the PRP-1 column. Perform reversed-phase HPLC using 92% HPLC buffer A to 75% HPLC buffer A over 30 min at a flow rate of 1 mL/min. The purified ODN elutes at ∼12 min.
15. Evaporate sample to dryness with a centrifugal evaporator, redissolve in doubledeionized water, and determine the concentration by measuring A260. 16. Check sample for correct base composition by SVP/AP digestion (UNIT 10.6). 17. Perform HPLC using 95% HPLC buffer A to 90% HPLC buffer A over 12 min at a flow rate of 1 mL/min. Monitor dA, dG, dC, and dT at A254, and monitor dS4U at A332. Derivatize ODN with N,N′-bis-bromoacetyl-1,2-diaminobenzene 18. Dilute 5 µL dS4U-containing ODN stock solution obtained in step 17 to 95 µL with 0.1 M potassium phosphate buffer, pH 8.0. The ODN stock solution is usually close to 50 OD/mL.
19. Mix ∼1 mg electrophile in 30 µL DMF. 20. Add electrophilic solution to ODN stock solution and allow mixture to sit 1.5 hr at room temperature. 21. Terminate reaction by passing solution through a small spin column containing 0.3 mL Sephadex G-25 equilibrated with double-deionized water. 22. Concentrate sample to dryness by precipitating with n-butanol. 23. Place sample briefly under vacuum to remove any trace of n-butanol. Use derivatized ODN immediately after preparation.
Cross-link target DNA using electrophilic ODN 24. Dissolve electrophilic ODN in 4 µL of 0.1 M potassium phosphate buffer, pH 8.0.
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25. Add 2.0 µL radiolabeled complimentary DNA solution. The radiolabeled ODN stock solution is usually between 1 and 5 OD/mL.
26. Incubate sample 15 min to 16 hr at room temperature. Maximum cross-linking usually requires an overnight incubation.
27. Analyze cross-linking using denaturing polyacrylamide gel electrophoresis (APPENDIX 3B). SUPPORT PROTOCOL
SYNTHESIS OF N,N′-BIS-BROMOACETYL-1,2-DIAMINOBENZENE To synthesize the electrophilic tether, the appropriate diamine reacts with bromoacetyl bromide (Skinner et al., 1967). Materials 1,2-Diaminobenzene Nitrogen gas Tetrahydrofuran (THF), freshly distilled Triethylamine (TEA), freshly distilled Bromoacetyl bromide Saturated aqueous NaHCO3 Ethyl acetate Saturated NaCl 5% (v/v) HCl MgSO4 Silica 9:1 (v/v) dichloromethane/methanol (CH2Cl2/CH3OH) Flame-dried 100-mL flask 3 × 15–cm glass column Additional reagents and equipment for flash column chromatography (APPENDIX 3E) 1. Place 500 mg 1,2-diaminobenzene in a flame-dried 100-mL flask under nitrogen gas. 2. Dissolve diamine in 50 mL freshly distilled THF. 3. Add 1.6 mL freshly distilled TEA. 4. Place flask under nitrogen gas and cool sample 5 min at −78°C. 5. Add 1 mL bromoacetyl bromide with a syringe over a 10-min period. 6. Stir mixture 30 min at −78°C, then 90 min at room temperature. 7. Add 50 mL saturated aqueous NaHCO3 to stop the reaction. 8. Dilute sample with 50 mL ethyl acetate. Extract organic layer with saturated NaCl, then with 5% HCl. 9. Dry organic phase over MgSO4.
Use of Electrophilic Substitution to Form Site-Specific Cross-Links in DNA
10. Evaporate sample onto 1 to 2 g silica for loading, and purify in a 3 × 15–cm glass column by flash column chromatography (APPENDIX 3E) using 9:1 CH2Cl2/CH3OH. Column-purified samples result in a higher percentage of cross-linking than recrystallized samples. The product should be used immediately.
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
1,8-Diazabicyclo[5.4.0]undec-7-ene (DBU) solution, 1.0 M Dissolve 15.2 g DBU in 75 mL solvent-grade acetonitrile that has been freshly distilled from CaH2. Allow DBU to dissolve, then add acetonitrile to 100 mL final volume. Store up to 12 months at 25°C in a desiccator. HPLC buffer A Dissolve 69.7 mL triethylamine that has been freshly distilled from CaH2 in 850 mL double-deionized H2O. Adjust to pH 6.5 using distilled acetic acid. Adjust volume to 1 L. Then dilute 100 mL to 500 mL (final 0.1 M triethylamine). Store up to 6 months at 5°C. If acetic acid is not distilled, impurities may appear in the HPLC.
NaSH/NH4OH solution, 50 mM Add 0.28 g NaSH⋅H2O to 75 mL concentrated NH4OH from a freshly opened bottle. Allow NaSH to dissolve, then add NH4OH to 100 mL final volume. Store up to 6 months at 4°C. COMMENTARY Background Information
dS4U is a versatile modified DNA base that can be used to accomplish a variety of postsynthetic transformations in synthetic ODNs. However, if the 4-thio group is left unprotected during solid-phase synthesis of dS4U-containing ODNs, the yield of product is typically near 20% per step. The 2-cyanoethyl group is a convenient protecting group for sulfur that can be readily removed by treatment with a nonnucleophilic base such as DBU. Yields from ODN syntheses utilizing the 2-cyanoethyl protecting group are equivalent to those containing only the four naturally occurring bases (Coleman and Kesicki, 1994). In addition to its reaction with electrophiles, as described above, oxidation-reduction chemistry can be used to form disulfide-constrained hairpins and duplexes. dS4U can also be modified by strong nucleophiles.
Critical Parameters The location of the 4-thio group near the base-pairing atoms of the duplex places the electrophilic tether in intimate contact with the major groove. Because of this positioning of the tether, the efficiency of cross-linking depends not only on the nucleophilicity of the target base but also on the local steric and electronic environment of the duplex. The steric environment can significantly influence the accessibility of the target to the nucleophile. For instance, when deoxythiouridine is placed
in the sequence 5′-d(TAATACGACCXACTATA)-3′ (X = dS4U), lower than expected levels of cross-linking to the complimentary target occur (≤35%). Modeling studies indicate that the methyl group of the T that is base paired to the dA in the 3′ direction blocks the approach of electrophiles to N7 of dG in this duplex. This is supported by studies where this T was replaced by dU, resulting in higher levels of cross-linking (Coleman and Kesicki, 1994). The cross-linking efficiency also seems to mirror the local charge density of the major groove. Guanines that are expected to be in a more negative environment based on calculations also appear to cross-link more readily (Coleman and Pires, 1997). In addition, the linker design is of critical importance in cross-linking efficiency. Considerations important for linker design include the tether geometry, H-bonding ability, and reactivity. The most effective electrophile found has been N,N′-bis-bromoacetyl-1,2-diaminobenzene. Using the optimal sequence with this electrophile, >90% cross-linking of a duplex can be achieved (Coleman and Pires, 1997). One final consideration is duplex stability. As the cross-linking occurs in the context of the duplex, factors that destabilize the duplex are expected to reduce the efficiency of cross-linking. Melting studies have shown that substitution of dC with dS4U destabilizes a 17-mer duplex by 9°C. Melting studies on the same 17-mer containing a non-cross-linking model
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of the electrophilic tether show a transition temperature that is an additional 3°C lower. Interpretation of data from melting experiments performed on the cross-linked duplex are complicated by the rapid depurination of the target strand that occurs as the temperature is raised, although the duplex appears to be intact up to ≥10°C above the transition temperature for the non-modified duplex (Coleman and Pires, 1997).
Anticipated Results The percent cross-linking obtained depends on the parameters discussed above. With the appropriate sequence and the most effective bis-electrophile, cross-linking efficiencies of between 80% and 95% are routinely obtained. Cross-linking levels of 30% to 80% have been obtained with less effective bis-electrophiles (e.g., N,N′-bis-bromoacetyl-1,3-diaminobenzene).
Time Considerations Synthesis of DNA, including cleavage and deprotection, generally takes 1 day. Purification is limited by the time required to remove solvent from samples, and generally requires 1.5 to 2 days. The cross-linking reaction takes ∼2 hr to set up for an overnight reaction. To prepare the electrophilic tether, 3 hr are re-
quired to set up and run the reaction, and another 2 hr are needed to purify the product.
Literature Cited Coleman, R.S. and Kesicki, E.A. 1994. Synthesis and postsynthetic modification of oligodeoxynucleotides containing 4-thio-2′-deoxyuridine (dS4U). J. Am. Chem. Soc. 116:11636-11642. Coleman, R.S. and Kesicki, E.A. 1995. Template-directed cross-linking of oligonucleotides: Site specific covalent modification of dG-N7 within duplex DNA. J. Org. Chem. 60:6252-6253. Coleman, R.S. and Pires, R.M. 1997. Covalent cross-linking of duplex DNA using 4-thio-2′-deoxyuridine as a readily modifiable platform for introduction of reactive functionalities into oligonucleotides. Nucl. Acids Res. 25:4771-4777. Coleman, R.S. and Siedlecki, J.M. 1991. Synthesis and stability of S-cyanoethyl protected thiouridine and 2′-deoxy-4-thiouridine. Tetrahedron Lett. 32:3033-3034. Skinner, W.A., Cory, M., Shellenberger, T.E., and Degraw, J.I. 1967. Effect of organic compounds on reproductive processes. V: Alkylating agents derived from aryl-, aralkyl-, and cyclohexylmethylenediamines. J. Med. Chem. 10:120-121.
Contributed by Richard M. Pires and Robert S. Coleman The Ohio State University Columbus, Ohio
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Synthesis of Endcap Dimethoxytrityl Phosphoramidites for Endcapped Oligonucleotides
UNIT 5.6
This unit describes the preparation of short endcapped DNA duplexes. Endcaps may be either aromatic (hydrophobic) or aliphatic (hydrophilic or hydrophobic) molecules that specifically cross-link the 5′ end of one strand with the 3′ end of the complementary strand in a DNA duplex. Endcaps may be viewed as a replacement of the loop region nucleotides of a DNA hairpin, with the added advantage of increased thermal stability. Specific cross-links at the terminus of a DNA duplex can be engineered through thiol-disulfide exchange with thiol-modified nucleosides incorporated at the end of the oligonucleotide strands (UNIT 5.1). The endcap approach, instead, requires the incorporation of the endcap into the sequence during oligonucleotide synthesis. Many different types of molecules may be used as endcaps, including oligo(ethylene glycol) linkers (UNIT 5.3), unsubstituted alkyl chains (Altmann et al., 1995), and aromatic molecules such as stilbene diethers (Lewis et al., 1999), carboxamides (Letsinger and Wu, 1995; Lewis et al., 2000), azobenzene (Yamana et al., 1996, 1998), and naphthalene diimides (Bevers et al., 1998, 2000). Basic Protocol 1 describes the synthesis of a naphthalene diimide hydrophobic endcap that prefers to base stack with GC base pairs. The phosphoramidite derivative of the endcap can be readily synthesized in three short steps for incorporation into an oligonucleotide sequence. The preparation of a terthiophene hydrophobic endcap is described in Basic Protocol 2, again requiring just three synthetic steps to obtain the phosphoramidite derivative. The terthiophene endcap has higher lipophilicity than the naphthalene diimide endcap and provides higher stability when stacked over an AT base pair. Finally, Basic Protocol 3 outlines the preparation of a hydrophilic 2,2′-oxydiacetamide endcap, which provides lower enhancement in stability as compared to either of the hydrophobic endcaps, but a more rigid and well-defined structure than the oligo(ethylene glycol) endcaps. Synthesis of endcapped oligonucleotides can be carried out using standard automated synthesis protocols with only minor modifications. The naphthalene diimide endcap is base sensitive and hence exposure to strong base such as ammonia must be avoided. Decomposition of the naphthalene diimide endcap can be avoided by using ultramild phosphoramidites (Glen Research) during oligonucleotide synthesis. The resulting oligonucleotide can be cleaved from the support as well as deprotected in a 0.05 M solution of potassium carbonate in methanol. The other requirement is to extend the time of the coupling cycle during the incorporation of the endcap phosphoramidite to 15 min during oligonucleotide synthesis. The endcapped oligonucleotides can be purified by denaturing PAGE (UNIT 10.4) or by reversed-phase HPLC (UNIT 10.5). MALDI-TOF mass spectrometry (UNIT 10.1) can be effectively used to characterize the endcapped oligonucleotides. SYNTHESIS OF N-[3-O-(2-CYANOETHYL-N,N-DIISOPROPYLPHOSHORAMIDITE)PROPYL]-N′-[3-(4,4′-DIMETHOXYTRITYLOXY)PROPYL]-NAPHTHALENE-1,4,5,8-TETRACARBOXYLIC DIIMIDE The sequence of reactions outlined in this protocol can be seen in Figure 5.6.1. The alkyl substituents of the naphthalene diimide central core are derived from 3-aminopropanol. This can be easily substituted by another aminoalcohol to provide an endcap that is either Contributed by Maneesh R. Pingle, Pei-Sze Ng, Xiaolin Xu, and Donald E. Bergstrom Current Protocols in Nucleic Acid Chemistry (2003) 5.6.1-5.6.15 Copyright © 2003 by John Wiley & Sons, Inc.
BASIC PROTOCOL 1
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5.6.1 Supplement 12
O
O
O
O
O
O
3-aminopropanol HO
O
O
N
N
O
O
OH
2
1
DMTr ⋅ Cl HO
O
O
N
N
O
O
ODMTr
3
O
O
N
N
O
O
i -Pr2NP(Cl)OCH2CH2CN O N
P
O
ODMTr
CN
4
Figure 5.6.1 Synthetic scheme for the preparation of the phosphoramidite derivative of the naphthalene diimide endcap. Abbreviations: DMTr⋅Cl, 4,4′-dimethoxytrityl chloride; i-Pr2NP(Cl)OCH2CH2CN, 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite.
shorter (2-aminoethanol) or longer (4-aminobutanol), but this will yield slightly less stable duplexes.
Synthesis of Endcap DMTr Phosphoramidites for Endcapped Oligonucleotides
Materials Napthalene-1,4,5,8-tetracarboxylic dianhydride (S.1; Figure 5.6.1) 3-Aminopropanol 2 M sodium carbonate Chloroform, reagent grade Activated charcoal Methanol, reagent grade Dichloromethane, reagent grade Pyridine, anhydrous 4,4′-Dimethoxytrityl chloride (DMTr⋅Cl) 5% (w/v) sodium bicarbonate Sodium sulfate, anhydrous Silica gel (230 to 400 mesh, 60 Å, E Merck) Triethylamine, reagent grade Hexanes, reagent grade Ethyl acetate, reagent grade Dichloromethane, anhydrous Diisopropylethylamine
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2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite Ethyl acetate, prewashed with 5% (w/v) sodium bicarbonate 25-, 50-, 100-, and 250-mL round-bottom flasks Buchner funnel Filtration flask Whatman no. 1 filter paper Vacuum oven Water-cooled reflux condenser Oil bath Filter funnel, prewarmed (∼50° to 60°C) Rotary evaporator equipped with a water aspirator and vacuum pump Nitrogen atmosphere (see inert atmosphere/vacuum manifold in UNIT 1.1, Fig. 1.1.3) 125-mL Erlenmeyer flasks 125-mL separatory funnels Glass funnel Glass wool 2 × 20–cm and 3 × 18–cm glass chromatography columns 25-mL pear-shaped flask 1-mL syringe and stainless steel needle Additional reagents and solutions for thin-layer chromatography (TLC; APPENDIX 3D) and flash chromatography (APPENDIX 3E) CAUTION: Exposure to pyridine and its vapors should be minimized. All reactions should be performed in a fume hood. The reactions for dimethoxytrityl protection and the phosphitylation of the dimethoxytrityl-protected endcap are sensitive to moisture and the glassware used for the reactions must be scrupulously dry. NOTE: All listed reagents are available from Sigma-Aldrich. Synthesize S.2 1. Suspend 2.14 g (8 mmol) naphthalene-1,4,5,8-tetracarboxylic dianhydride (S.1) in 150 mL water in a 250-mL round-bottom flask containing a stir bar. 2. Add 1.5 g (20 mmol) 3-aminopropanol drop-wise over 2 min while stirring the contents of the flask. Warm to 70°C and stir on a magnetic stirrer for 2 hr. 3. Cool the flask to room temperature and filter the solution using a Buchner funnel fitted with Whatman no. 1 filter paper attached to a filtration flask. Wash the residue on the filter paper with 50 mL of 2 M sodium carbonate and then with 200 mL of water. Allow the residue to dry on the filter paper. 4. Transfer the dry residue to a 100-mL round-bottom flask and dry overnight under vacuum (in a vacuum oven at <1 Torr and room temperature) to obtain a beige solid. 5. Dissolve the crude product in 25 mL chloroform and reflux the solution in the flask by attaching a water-cooled reflux condenser to the flask and heating the flask in an oil bath. When the solution begins refluxing (61°C), add 50 mg activated charcoal and reflux the solution for an additional 10 min. 6. Quickly filter the solution through a warm filter funnel (∼50° to 60°C) fitted with Whatman no. 1 paper. 7. Remove the solvent under reduced pressure on a rotary evaporator equipped with a water aspirator to yield N,N′-bis(3-hydroxypropyl)-naphthalene-1,4,5,8-tetracarboxylic diimide (S.2) as an off-white powder.
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8. Analyze the product by TLC (APPENDIX 3D) on silica gel, developing the plate with 10% (v/v) methanol in reagent-grade dichloromethane (Rf = 0.4). Tritylate to give S.3 9. Coevaporate 382 mg (1 mmol) S.2 three times with 20-mL portions of anhydrous pyridine in a 50-mL round-bottom flask using a rotary evaporator connected to a vacuum pump. 10. Dissolve the contents of the flask in 20 mL anhydrous pyridine and add 406 mg (1.2 mmol) DMTr⋅Cl while maintaining the flask under a dry nitrogen atmosphere. Stir the contents of the flask for 2 hr under nitrogen. 11. Remove the solvent on a rotary evaporator connected to a vacuum pump and dissolve the mixture of crude products in 50 mL reagent-grade dichloromethane. 12. Transfer the solution to a 125-mL separatory funnel and wash with 50 mL of 5% sodium bicarbonate solution and then with three 50-mL portions of water. 13. Transfer the dichloromethane solution to a 125-mL Erlenmeyer flask and add 1 g anhydrous sodium sulfate. Swirl the solution for a few minutes and allow to stand for 10 min. 14. Remove the drying agent by filtering the solution through a glass funnel with a glass wool plug. 15. Remove the solvent under reduced pressure using a rotary evaporator and water aspirator. 16. Purify the product (S.3) by flash chromatography (APPENDIX 3E) on a 3 × 18–cm column of silica gel neutralized by pretreatment with 1% (v/v) triethylamine in hexanes. Elute with hexanes and ethyl acetate, starting with 100% hexanes and increasing the amount of ethyl acetate to 70% in 10% increments every 100-mL fraction. Although chromatography of the DMTr derivative can be performed without using any triethylamine in the mobile phase, it is recommended that between 0.5% and 1% triethylamine be incorporated in the elution solvent. For more details on the purification of DMTr derivatives see UNIT 2.3.
17. Analyze fractions by TLC on silica gel, developing the plates with 50% (v/v) ethyl acetate in hexanes. Pool all fractions containing the product (Rf = 0.3) in a 250-mL round-bottom flask and remove the solvent under reduced pressure (see step 15). 18. Dry the purified N-(3-hydroxypropyl)-N′-[3-(4,4′-dimethoxytrityloxy)propyl]-naphthalene-1,4,5,8-tetracarboxylic diimide (S.3) overnight under vacuum (in a vacuum oven at <1 Torr and room temperature) to produce a yellow foam. Synthesize phosphoramidite S.4 19. Dissolve 267 mg (0.379 mmol) S.3 in 5 mL anhydrous dichloromethane in a 25-mL pear-shaped flask under a stream of dry nitrogen. 20. Add 234 µL (1.346 mmol) diisopropylethylamine. Then add 100 µL (0.448 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite drop-wise through a 1-mL syringe while gently swirling the flask. Synthesis of Endcap DMTr Phosphoramidites for Endcapped Oligonucleotides
21. Stir the reaction gently under nitrogen at room temperature for 1 hr.
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22. Pour the contents of the flask into 20 mL ethyl acetate (previously washed with 5% sodium bicarbonate solution) in a 125-mL separatory funnel. Wash the ethyl acetate solution with three 20-mL portions of 5% sodium bicarbonate solution and then with three 10-mL portions of water. 23. Dry the ethyl acetate solution over anhydrous sodium sulfate (steps 13 and 14). 24. Remove the solvent under reduced pressure (step 15). 25. Purify the product (S.4) by flash chromatography on a 2 × 20–cm silica gel column pretreated with 1% (v/v) triethylamine in hexanes, eluting with 40% ethyl acetate and 1% triethylamine in hexanes. Collect the eluate in 10-mL fractions. 26. Analyze by TLC on silica gel, developing the plates with 40% ethyl acetate and 1% triethylamine in hexanes. Pool all fractions containing the product (Rf = 0.85) in a 250-mL round-bottom flask. 27. Remove the solvent under reduced pressure on a rotary evaporator connected to a water aspirator to produce N-[3-O-(2-cyanoethyl-N,N-diisopropylphosphoramidite)propyl]-N′-[3-(4,4′-dimethoxytrityloxy)propyl]-naphthalene-1,4,5,8-tetracarboxylic diimide (S.4) as an oily residue. 28. Dissolve the residue in 2 mL anhydrous dichloromethane and evaporate under vacuum on a rotary evaporator connected to a water aspirator. Repeat this step until a yellow foam is obtained and dry the foam overnight under vacuum using a vacuum pump at <1 Torr. The phosphoramidite may be stored for short periods of time (<10 days) at 4°C under anhydrous conditions, but best results are obtained if it is incorporated into an oligonucleotide sequence within a few days of its preparation.
SYNTHESIS OF 5-[3-O-(2-CYANOETHYL-N,N-DIISOPROPYLPHOSPHORAMIDITE)PROPYL]-5′′-[3-(4,4′-DIMETHOXYTRITYLOXY)PROPYL]2,2′:5′,2′′-TERTHIOPHENE
BASIC PROTOCOL 2
The scheme in Figure 5.6.2 outlines the synthesis of the phosphoramidite derivative of a terthiophene endcap. Materials 2,2′:5′,2′′-Terthiophene (S.5; Fig. 5.6.2) Tetrahydrofuran (THF), anhydrous Dry ice/acetone freezing bath n-Butyllithium Boron trifluoride diethyl etherate Trimethylene oxide Saturated sodium bicarbonate Diethyl ether, reagent grade Brine solution (saturated aqueous NaCl) Sodium sulfate, anhydrous Silica gel (230 to 400 mesh, 60 Å, E Merck) Hexanes, reagent grade Ethyl acetate, reagent grade Pyridine, anhydrous 4,4′-Dimethoxytrityl chloride (DMTr⋅Cl) Dichloromethane, anhydrous 5% (w/v) sodium bicarbonate
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5
S
2
2″ 2′
S
S
5″
S
1) n -butyllithium
5′
2) trimethylene oxide
HO
S OH
S
5
6
S DMTr ⋅ Cl
HO
S ODMTr
S 7
S
i -Pr2NP(Cl)OCH2CH2CN N
O
S ODMTr
S
P O CN 8
Figure 5.6.2 Synthetic scheme for the preparation of the phosphoramidite derivative of the terthiophene endcap. Abbreviations: DMTr⋅Cl, 4,4′-dimethoxytrityl chloride; i-Pr2NP(Cl)OCH2CH2CN, 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite.
Triethylamine Diisopropylethylamine 2-Cyanoethyl-N,N-diisopropylchlorophosphoramidite 25-mL round-bottom flasks Nitrogen atmosphere (see inert atmosphere/vacuum manifold in UNIT 1.1, Fig 1.1.3) 1-mL syringe and stainless steel needle 125-mL separatory funnels 125-mL Erlenmeyer flasks Glass funnel Glass wool Rotary evaporator with water aspirator and vacuum pump 2 × 20–cm and 4 × 20–cm glass chromatography columns Additional reagents and equipment for column chromatography (APPENDIX 3E) and thin-layer chromatography (TLC; APPENDIX 3D) NOTE: All listed reagents are available from Sigma-Aldrich. Synthesis of Endcap DMTr Phosphoramidites for Endcapped Oligonucleotides
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Synthesize S.6 1. Dissolve 333 mg (1.3 mmol) 2,2′:5′,2′′-terthiophene (S.5) in 15 mL anhydrous THF in a 25-mL round-bottom flask containing a Teflon-coated 0.5-in. (1.3-cm) magnetic stir bar under a stream of nitrogen. 2. Cool the solution to −78°C by immersing the flask in a dry ice/acetone freezing bath. 3. Add 1.8 mL (1.3 mmol) n-butyllithium and warm the reaction to 0°C by transferring the flask to an ice bath. 4. Stir the reaction for 2 hr on a magnetic stirrer. 5. Add 384 µL (3 mmol) boron trifluoride diethyl etherate and then add 198 µL (3 mmol) trimethylene oxide with a 1-mL syringe. 6. Stir the reaction for 40 min on a magnetic stirrer. 7. Quench the reaction by addition of 10 mL saturated sodium bicarbonate solution and then allow the reaction to warm to room temperature. 8. Transfer the contents of the flask to a 125-mL separatory funnel and extract the aqueous layer with 30 mL diethyl ether. 9. Wash the ether layer with 25 mL brine solution. 10. Transfer the ether layer to a 125-mL Erlenmeyer flask and add 0.5 g anhydrous sodium sulfate. 11. Remove the drying reagent by filtering through a glass funnel fitted with a glass wool plug. 12. Remove the solvent under reduced pressure using a rotary evaporator and water aspirator. 13. Purify the product (S.6) by flash chromatography (APPENDIX 3E) on a 4 × 20–cm silica gel column, eluting first with hexanes, then with 35% (v/v) ethyl acetate in hexanes, and finally with 65% ethyl acetate in hexanes. 14. Analyze the fractions by TLC (APPENDIX 3D) on silica gel, developing the plates in 50% (v/v) ethyl acetate in hexanes. Combine all fractions containing the product (Rf = 0.13) and remove the solvent under reduced pressure (step 12). 15. Dry 5,5′′-bis(3-hydroxypropyl)-2,2′:5′,2′′-terthiophene (S.6) overnight under vacuum using a rotary evaporator with a water aspirator. Tritylate to give S.7 16. Coevaporate 138 mg (0.38 mmol) S.6 with 10 mL anhydrous pyridine in a 25-mL round-bottom flask using a rotary evaporator connected to a vacuum pump. 17. Dissolve the contents of the flask in 5 mL anhydrous pyridine and add 167 mg (0.49 mmol) DMTr⋅Cl while maintaining the flask under a dry nitrogen atmosphere. Stir the contents of the flask for 3 hr under nitrogen. 18. Remove the solvent on a rotary evaporator connected to a vacuum pump and dissolve the mixture of crude products in 25 mL anhydrous dichloromethane. 19. Transfer the solution to a 125-mL separatory funnel and wash with 25 mL of 5% sodium bicarbonate solution followed by three 20-mL portions of water. 20. Dry the solution with anhydrous sodium sulfate (steps 10 and 11).
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21. Remove the solvent under reduced pressure (step 12). 22. Purify the product (S.7) by flash chromatography on a 2 × 20–cm silica gel column, neutralized by pretreatment with 1% (v/v) triethylamine in hexanes. Elute with 70% (v/v) ethyl acetate in hexanes. Although chromatography of the DMTr derivative can be performed without using any triethylamine in the mobile phase, it is recommended that between 0.5% and 1% triethylamine be incorporated in the elution solvent. For more details on the purification of DMTr derivatives see UNIT 2.3.
23. Analyze fractions by TLC using 50% (v/v) ethyl acetate in hexanes and combine all fractions containing the product (Rf = 0.31). Remove the solvent under reduced pressure (step 12). 24. Dissolve the residue in 2 mL anhydrous dichloromethane and evaporate under vacuum on a rotary evaporator connected to a water aspirator. Repeat this step until the product 5-(3-hydroxypropyl)-5′′-[3-(4,4′-dimethoxytrityloxy)propyl]-2,2′:5′,2′′terthiophene (S.7) solidifies into a foam. Synthesize phosphoramidite S.8 25. Dissolve 83 mg (0.125 mmol) S.7 in 5 mL anhydrous dichloromethane in a 25-mL round-bottom flask under a stream of nitrogen. 26. Add 78 µL (0.446 mmol) diisopropylethylamine to the flask and then add 50 µL (0.225 mmol) 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite drop-wise through a 1-mL syringe while gently swirling the flask. 27. Stir the reaction gently under nitrogen for 1 hr. 28. Pour the contents of the flask into 20 mL ethyl acetate in a 125-mL separatory funnel. Wash the ethyl acetate solution with three 20-mL portions of 5% sodium bicarbonate solution followed by three 10-mL portions of water. 29. Dry the ethyl acetate solution over anhydrous sodium sulfate (see steps 10 and 11). 30. Remove the solvent under reduced pressure (step 12). 31. Purify the product S.8 by flash chromatography on a 2 × 20–cm silica gel column, eluting with 75% (v/v) ethyl acetate in hexanes. 32. Analyze by TLC using 25% ethyl acetate and 1% triethylamine in hexanes and combine all fractions containing the product (Rf = 0.5). Remove the solvent under reduced pressure (step 12). 33. Dissolve the residue in 2 mL anhydrous dichloromethane and evaporate under vacuum on a rotary evaporator connected to a water aspirator. Repeat this step until the product 5-[3-O-(2-cyanoethyl-N,N-diisopropylphosphoramidite)propyl]-5′′-[3(4,4′-dimethoxytrityloxy)propyl]-2,2′:5′,2′′-terthiophene (S.8) solidifies into a foam. The phosphoramidite may be stored for short periods of time (<10 days) at 4°C under anhydrous conditions, but best results are obtained if it is incorporated into an oligonucleotide sequence within a few days of its preparation.
Synthesis of Endcap DMTr Phosphoramidites for Endcapped Oligonucleotides
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Current Protocols in Nucleic Acid Chemistry
H2N
TBDMS ⋅ Cl
OH
H 2N
3-aminopropanol
TBDMSO
diglycolyl chloride
OTBDMS 9
H N
H N
O
OTBDMS
2.9% HCl in 95% ethanol
H N
HO
O
10
DMTr ⋅ Cl
OH
O
O
O
O
H N
11
HO
H N
H N
O O
ODMTr
O 12
(i -Pr2N)2OCH2CH2CN
N
H N
O P O
H N
O
ODMTr
O
O CN 13
Figure 5.6.3 Synthetic scheme for the preparation of the phosphoramidite derivative of the 2,2′-oxydiacetamide endcap. Abbreviations: DMTr⋅Cl, 4,4′-dimethoxytrityl chloride; (i-Pr2N)2OCH2CH2CN, 2-cyanoethyl-N,N,N′,N′-tetraisopropylphosphorodiamidite; TBDMS⋅Cl, tert-butyldimethylsilyl chloride.
SYNTHESIS OF N-[3-O-(2-CYANOETHYL-N,N-DIISOPROPYLPHOSPHORAMIDITE)PROPYL]-N′-[3-(4,4′-DIMETHOXYTRITYLOXY)PROPYL]-2,2′OXYDIACETAMIDE
BASIC PROTOCOL 3
The sequence of reactions in this protocol describes the synthesis of the phosphoramidite derivative of an aliphatic hydrophilic endcap. The synthetic scheme is shown in Figure 5.6.3. All intermediate compounds up to the trityl-protected endcap are stable for storage purposes and may be prepared in larger quantities. It is advisable to prepare the final phosphoramidite derivative in the required quantity only when needed for oligonucleotide synthesis. Materials 3-Aminopropanol Dichloromethane, anhydrous Triethylamine (TEA), anhydrous (preferably freshly distilled) tert-Butyldimethylsilyl chloride (TBDMS⋅Cl) 4,4-Dimethylaminopyridine Brine solution (saturated NaCl) Sodium sulfate, anhydrous Diglycolyl chloride 5% (v/v) acetic acid 5% (w/v) sodium bicarbonate
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Concentrated HCl 95% (v/v) ethanol Methanol, reagent grade Hexanes, reagent grade Pyridine, anhydrous (preferably freshly distilled) 4,4′-Dimethoxytrityl chloride (DMTr⋅Cl) Silica gel (230 to 400 mesh, 60 Å, E Merck) Dichloromethane, reagent grade 1% (w/v) sodium hydroxide (optional) Acetonitrile, anhydrous 2-Cyanoethyl-N,N,N′,N′-tetraisopropylphosphorodiamidite 1H-Tetrazole 25-, 50-, 100- and 250-mL round-bottom flasks Nitrogen atmosphere (see inert atmosphere/vacuum manifold in UNIT 1.1, Fig. 1.1.3) 1-mL syringe and stainless steel needles 125- and 250-mL separatory funnels 125- and 250-mL Erlenmeyer flasks Glass funnels Glass wool Rotary evaporator with a water aspirator and vacuum pump 2 × 20–cm and 4 × 25–cm glass chromatography columns Additional reagents and equipment for column chromatography (APPENDIX 3E) and thin-layer chromatography (TLC; APPENDIX 3E) NOTE: All listed reagents are available from Sigma-Aldrich. Silylate 3-aminopropanol to give S.9 1. Dissolve 7 g (1 eq, 0.09 mol) 3-aminopropanol in 90 mL anhydrous dichloromethane in a dry 250-mL round-bottom flask containing a magnetic stir bar, under a stream of nitrogen. 2. Cool the flask in an ice bath and add 14.37 ml (1.1 eq, 0.1 mol) anhydrous triethylamine, 15.93 g (1.1 eq, 0.1 mol) TBDMS⋅Cl, and 0.5 g 4,4-dimethylaminopyridine to the flask. Stir the mixture overnight under nitrogen. Addition of TBDMS⋅Cl results in a vigorous reaction that generates heat and results in the formation of a white precipitate. It is important to cool the reaction flask in an ice bath. The contents of the reaction flask will slowly return to room temperature when the ice in the bath has melted, and the reaction will proceed at room temperature for the duration.
3. Add 50 mL water to the reaction mixture and stir vigorously for 10 min to dissolve the white precipitate. 4. Transfer the contents of the flask to a 250-mL separatory funnel and discard the aqueous layer. Wash the organic fraction with three 30-mL fractions of water followed by three 30-mL fractions of brine solution. 5. Transfer the dichloromethane solution to a 250-mL Erlenmeyer flask and add 1 g anhydrous sodium sulfate. Swirl the solution for a few minutes and then allow to stand for 10 min. 6. Remove the drying agent by filtering the solution through a glass funnel with a glass wool plug. Synthesis of Endcap DMTr Phosphoramidites for Endcapped Oligonucleotides
7. Remove the solvent under reduced pressure using a rotary evaporator and water aspirator at room temperature to obtain the product 3-tert-butyldimethylsilyloxypropylamine (S.9) as a colorless oil.
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Synthesize diacetamide S.10 8. Dissolve 8.17 g (2 eq, 0.04 mol) S.9 in 30 mL anhydrous dichloromethane in a 100-mL round-bottom flask containing a dry stir bar. Cool the flask in an ice bath. 9. Add 7.6 mL (3 eq, 0.05 mol) anhydrous triethylamine to the flask and then add 2.26 mL (1 eq, 0.018 mol) diglycolyl chloride drop-wise while stirring the reaction mixture on a magnetic stirrer. The reaction mixture will turn brown on addition of the diglycolyl chloride.
10. After addition of diglycolyl chloride is complete, remove the flask from the ice bath and stir the contents overnight at room temperature. 11. Add 40 mL water to the reaction mixture and stir vigorously for 10 min. 12. Transfer to a 125-mL separatory funnel and discard the aqueous layer. Wash the organic layer with two 30-mL portions of 5% acetic acid solution, then with two 30-mL portions of 5% sodium bicarbonate solution, and finally with two 60-mL portions of brine solution. 13. Transfer the dichloromethane solution to a 125-mL Erlenmeyer flask containing 0.5 g anhydrous sodium sulfate, swirl the solution for a few minutes, and allow the solution to stand for 10 min. 14. Remove the drying agent by filtering the solution through a glass funnel with a glass wool plug. 15. Remove the dichloromethane under reduced pressure using a rotary evaporator and water aspirator at room temperature to obtain the product N,N′-bis(3-tert-butyldimethylsilyloxypropyl)-2,2′-oxydiacetamide (S.10) as a brown oil. Dry the oil overnight in a 25-mL round-bottom flask under high vacuum (in a vacuum pump at <1 Torr). Desilylate to give S.11 16. Prepare a 2.9% (w/w) solution of concentrated HCl in 95% ethanol. 17. Dissolve 7.63 g (0.016 mol) S.10 in 15 mL of 95% ethanol in a 50-mL round-bottom flask containing a stir bar. 18. Add 15 mL of the HCl solution to the flask and stir the contents of the flask for 20 min on a magnetic stirrer. 19. Remove the solvent under reduced pressure using a rotary evaporator and water aspirator at room temperature to produce a brown oil. 20. Dissolve the brown oil in 15 mL methanol and transfer to a 125-mL separatory funnel. Wash the methanol layer with three 20-mL portions of hexanes. 21. Remove the solvent under reduced pressure using a rotary evaporator and water aspirator at room temperature to obtain the product N,N′-bis(3-hydroxypropyl)-2,2′oxydiacetamide (S.11) as a brown oil. Tritylate to give S.12 22. Coevaporate 7.63 g (0.016 mol) S.11 three times with 20-mL portions of anhydrous pyridine in a 100-mL round-bottom flask using a rotary evaporator connected to a vacuum pump. Dissolve the contents of the flask in 30 mL anhydrous pyridine. 23. Maintain the solution under a nitrogen atmosphere and add 5.707 g (0.016 mol) DMTr⋅Cl. Stir the contents of the flask overnight under a nitrogen atmosphere.
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24. Remove the solvent under reduced pressure using a rotary evaporator connected to a vacuum pump and purify the compound S.12 by flash chromatography (APPENDIX 3E) on a 4 × 25–cm silica gel column, eluting with 0% to 1% (v/v) methanol in dichloromethane containing 1% (v/v) triethylamine. Collect the effluent in 10-mL fractions. 25. Analyze by TLC (APPENDIX 3E) on silica gel, developing with 1% (v/v) methanol and 1% (v/v) triethylamine in dichloromethane. Combine the fractions that contain the desired product (Rf = 0.27) and evaporate the solvent under reduced pressure on a rotary evaporator connected to a water aspirator to obtain the product as a foam. 26. Optional: If the product does not foam, dissolve the oily residue in 50 mL dichloromethane and transfer to a 125-mL separatory funnel. Wash the dichloromethane solution with 25 mL of 1% sodium hydroxide. Dry the dichloromethane solution over anhydrous sodium sulfate and remove the solvent under reduced pressure (steps 13 to 15). 27. Dry the product N-(3-hydroxypropyl)-N′-[3-(4,4′-dimethoxytrityloxy)propyl]-2,2′oxydiacetamide (S.12) overnight in vacuo (in a vacuum pump at <1 Torr) to obtain a light brown foam. Synthesize phosphoramidite S.13 28. Coevaporate 123.5 mg (0.224 mmol) S.12 with anhydrous acetonitrile and then with anhydrous dichloromethane in a 25-mL round-bottom flask on a rotary evaporator connected to a water aspirator. Dry the material overnight under vacuum (in a vacuum pump at <1 Torr). 29. Dissolve the contents of the flask in 2 mL anhydrous dichloromethane and then add 0.078 mL (0.234 mmol) 2-cyanoethyl-N,N,N′,N′-tetraisopropylphosphorodiamidite (using a 1-mL syringe) and 15.9 mg (0.224 mmol) 1H-tetrazole over 15 min while maintaining the flask under nitrogen. Stir the contents of the flask under nitrogen for 2 hr. 30. Analyze the reaction by TLC on silica gel, developing the plate with 5:4:1 (v/v/v) hexanes/dichloromethane/triethylamine. 31. Puri fy the product N-[3-O-(2-cyanoethyl-N,N-diisopropylphosphoramidite)propyl]-N′-[3-(4,4′ -dimethoxytrityloxy)propyl]-2,2′-oxydiacetamide (S.13) on a 2 × 20–cm column of silica gel, eluting with 5:4:1 hexanes/dichloromethane/triethylamine. 32. Combine the fractions containing the product (Rf = 0.2) and remove the solvent under reduced pressure on a rotary evaporator connected to a water aspirator. Dry the product overnight under vacuum by connecting the flask to a vacuum pump (<1 Torr). The phosphoramidite may be stored for short periods of (<10 days) at 4°C under anhydrous conditions, but best results are obtained if it is incorporated into an oligonucleotide sequence within a few days of its preparation.
COMMENTARY Background Information
Synthesis of Endcap DMTr Phosphoramidites for Endcapped Oligonucleotides
Oligonucleotides bearing specific crosslinks generally display greater stability than their uncross-linked counterparts. This can be most easily observed in their melting behavior, with cross-linked oligonucleotides having higher melting temperature (Tm) values. The
authors’ goal was to develop both hydrophobic and hydrophilic endcaps that not only stabilize very short oligonucleotide duplexes (e.g., 4 bp) but also offer a variety of environments that may be appropriate for different applications. Hydrophobic endcaps such as stilbene diether and stilbene dicarboxamide (Letsinger
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whereas the naphthalene diimide endcap shows the greatest enhancement in melting temperature when stacked over a GC base pair. In both cases, the endcapped duplexes show significantly higher melting temperatures than their hairpin counterparts. The hydrophilic 2,2′-oxydiacetamide linker displays lower Tm values than either of the aromatic endcaps or the hexa(ethylene glycol) linker. However, the 2,2′-oxydiacetamide linker may offer some advantage when it is important to have a very hydrophilic moiety of minimal size with greater rigidity than the hexa(ethylene glycol) linker. The length spanned by these endcaps can be tuned by changing the chain length of the hydroxyalkyl portion of the endcaps. However, the authors have found that the length provided by the aminopropyl unit is optimal for obtaining the highest Tm values.
and Wu, 1995; Lewis et al., 1995) have been used to both stabilize DNA hairpins as well as study photo-induced charge separation and charge recombination (Lewis et al., 2000) and charge transfer (Lewis et al., 1997, 2001). Endcapped oligonucleotides are capable of specific protein binding as demonstrated by a stilbene dicarboxamide–endcapped oligonucleotide that mimics the REV responsive element and binds with equal affinity to the REV protein of HIV-1 (Nelson et al., 1996). Stilbene diether– endcapped duplexes have been shown to crystallize in a pinwheel arrangement with four oligonucleotide duplexes per asymmetric subunit (Lewis et al., 1999, 2000). Naphthalene and perylene diimide endcaps have been used to stabilize both duplex and triplex oligonucleotides (Bevers et al., 1998, 2000). An azobenzene endcap has also been used to stabilize short duplexes (Yamana et al., 1996, 1998). Applications of aliphatic hydrophilic endcaps are discussed in UNIT 5.3. Table 5.6.1 shows the melting temperature values for hairpin sequences with 4-bp stems and 4-nt loops. Replacement of the nucleotide loop by an endcap gives the results shown in Table 5.6.2. Both aromatic hydrophobic endcaps provide an overall increase in Tm values over the aliphatic endcaps. The more lipophilic terthiophene endcap provides the highest stability when it is stacked over an AT base pair,
Compound Characterization N,N′-Bis(3-hydroxypropyl)-naphthalene1,4,5,8-tetracarboxylic diimide (S.2). 1H NMR (250 MHz, CDCl3, δ): 8.8 (s, 4H, ar), 4.38 (t, J = 6.3 Hz, 3H, CH2), 3.65 (t, J = 5.7 Hz, 3H, CH2), 2.02 (m, 3H, CH2). 13C NMR (250 MHz, DMSO, δ): 162.0 130.7 129.4 125.6 59.9 38.0 30.7. MS: (+ve ESI) (M+H) 383, (–ve ESI) (M) 382. HRMS (FAB): calc. 383.1243; actual 383.1249.
Table 5.6.1
Melting Temperatures for 4-bp Stem Hairpin Oligonucleotides
Sequencea
Tm
ATGCTTTTGCAT ATGCAAAAGCAT GCTATTTTTAGC GCTAAAAATAGC
58.4°C 62.0°C 38.5°C 50.9°C
aThe underlined nucleotides form the loop of the hairpin oligonucleotide.
Table 5.6.2
Melting Temperatures for Endcapped Oligonucleotides
Sequencea
Type of endcap
Tm
ATGC-X-GCAT ATGC-X-GCAT GCTA-X-TAGC ATGC-X-GCAT GCTA-X-TAGC ATGC-X-GCAT GCTA-X-TAGC
Hexa(ethylene glycol) Naphthalene diimide Naphthalene diimide Terthiophene Terthiophene 2,2′-Oxydiacetamide 2,2′-Oxydiacetamide
61.4°C 74.6°C 61.6°C 62.3°C 65.8°C 51°C 41.7°C
aX refers to the location of the endcap.
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Synthesis of Endcap DMTr Phosphoramidites for Endcapped Oligonucleotides
N-(3-Hydroxypropyl)-N′-[3-(4,4′-dimethoxytrityloxy)propyl]-naphthalene-1,4,5,8-tetracarboxylic diimide (S.3). 1H NMR (250 MHz, CD2Cl2, δ): 8.69 (s, 4H. naphthalene), 7.43 – 7.18 (m, 9H, DMTr), 6.8 – 7.9 (m, 4H, DMTr), 4.2 (m, 4H, CH2), 3.74 (s, 6H, OCH3), 3.68 (m, 2H, CH2), 3.09 (t, J = 6.1 Hz, 2H, CH2), 2.1 (m, 4H, CH2). HRMS (FAB): calc. 684.2471; actual 684.2464. N-[3-O-(2-Cyanoethyl-N,N-diisopropylphosphoramidite)propyl]-N′-[3-(4,4′-dimethoxy trityloxypropyl]-naphthalene-1,4,5,8-tetracarb oxylic diimide (S.4). 31P NMR: 147.8 (referenced to external 85% H3PO4). 5,5′′-Bis(3-hydroxypropyl)-2,2′:5′,2′′-terthiophene (S.6). 1H NMR (250 MHz, CDCl3, δ): 6.90 (d, 4H), 6.70 (d, 2H), 3.75 (t, 4 H), 2.90 (t, 4H), 1.95 (m, 4H). 5-(3-Hydroxypropyl)-5′′-[3-(4,4′-dimethoxytrityloxy)propyl]-2,2′:5′,2′′-terthiophene (S.7). 1H NMR (250 MHz, CDCl3, δ): 6.6-7.5 (m, 19H), 3.8 (s, 6H), 3.7 (t, 2 H), 3.2 (t, 2H), 2.9 (m, 4H), 1.95 (m, 4H). 5-[3-O-(2-Cyanoethyl-N,N-diisopropylphosphoramidite)propyl]-5′′-[3-(4,4′-dimethoxytrityloxy)propyl]-2,2′:5′,2′′-terthiophene (S.8). 1H NMR: 6.6-7.2 (m, 19H), 2.63.9 (1s, 1m, 12H), 3.2 (t, 2H), 2.9 (m, 4H), 2.7 (t, 2H), 2.5 (m, 2H), 2.0 (m, 4H), 1.2 (d, 12H), 1.0 (t, 4H). 31P NMR: 170. 3-tert-Butyldimethylsilyloxypropylamine (S.9). 1H NMR (250 MHz, CDCl3, δ): –0.010 (s, 6H), 0.831 (s, 9H), 1.595 (p, 2H), 2.737 (t, 2H), 3.637 (t, 2H). 13C NMR (300 MHz, CDCl3, δ): -5.4173, 18.2349, 25.8746, 36.2497, 39.3576, 61.1958. HRMS (ESI, +): calc. 190.1627; actual 190.1630. N,N′-Bis(3-tert-butyldimethylsilyloxypropyl)-2,2′-oxydiacetamide (S.10). 1H NMR (250 MHz, CDCl3, δ): 2.019 (s, 10H), 2.854 (s, 18H), 3.709 (p, 4H), 5.380 (q, 4H), 5.686 (t, 4H), 5.976 (s, 4H). 13C NMR (300 MHz, CDCl3, δ): –5.476, 18.244, 25.848, 31.676, 37.352, 61.907, 71.270. HRMS (ESI, +): calc. 476.3102; actual 476.3091. N,N′-Bis(3-hydroxypropyl)-2,2′-oxydiacetamide (S.11). 1H NMR (250 MHz, CD3OD, δ): 744 (q, 4H), 3.37 (t, 4H), 3.60 (t, 4H), 4.039 (s, 4H). 13C NMR (250 MHz, CD3OD, δ): 33.098, 37.290, 60.547, 71.409, 171.729. HRMS (ESI, +): calc. 249.1450; actual 249.1446. N-(3-Hydroxypropyl)-N′-[3-(4,4′-dimethoxytrityloxy)propyl]-2,2′-oxydiacetamide (S.12). 1H NMR (250 MHz, CD2Cl2, δ): 1.602 (p, 2H), 1.805 (p, 2H), 3.164 (t, 2H), 3.283-
3.429 (m, 4H), 3.543 (t, 2H), 3.779 (s, 6H), 3.914, 3.943 (d, 4H), 6.817-7.365 (m, 13H). N-[3-O-(2-Cyanoethyl-N,N-diisopropylphosphoramidite)propyl]-N′-[3-(4,4′-dimethoxytrityloxy)propyl]-2,2′-oxydiacetamide (S.13). 31P NMR: 148 (referenced to external 85% H3PO4).
Critical Parameters Anyone with a moderate amount of experience in chemical synthesis should not find any difficulties in the synthesis of any of these endcaps. The reactions used to generate the DMTr and phosphoramidite derivatives are extremely sensitive to moisture and must be performed under anhydrous conditions. Additionally, the DMTr group is acid labile, and thus care must be taken to avoid any exposure to acid. As a precaution, small amounts of triethylamine can be used during chromatography. The tritylation reaction in each protocol yields a mixture of unreacted, monotritylated, and detritylated products. The ratio of reactants was chosen to maximize the yield of the monotritylated product. The reactant (untritylated), monotritylated, and ditritylated products can be distinguished by TLC. The ditrylated product runs at higher Rf, while the reactant migrates more slowly than the monotritylated products. The phosphoramidite derivative also requires careful handling, and exposure to air or acids must be avoided. It is preferable to synthesize the phosphoramidite derivative of the required endcap just before oligonucleotide synthesis.
Anticipated Results Yields of endcapped oligonucleotides should generally be >90% using standard automated synthesis procedures (see, e.g., APPENDIX 3C). The coupling time for the endcap phosphoramidite, however, should be increased to 15 min.
Time Considerations The phosphoramidite derivatives of the naphthalene diimide and terthiophene hydrophobic endcaps can each be prepared in 3 days including the time required for purification. The hydrophilic oxydiacetamide endcap synthesis requires significantly longer times and can be completed in 7 to 8 days.
5.6.14 Supplement 12
Current Protocols in Nucleic Acid Chemistry
Literature Cited Altmann, S., Labhardt, A.M., Bur, D., Lehmann, C., Bannworth, W., Billeter, M., Wuthrich, K., and Leupin, W. 1995. NMR studies of DNA duplexes singly cross-linked by different synthetic linkers. Nucl. Acids Res. 23:4827-4835.
Lewis, F.D., Letsinger, R.L., Wasielewski, M.R., and Egli, M. 2000a. Structure and electron transfer in synthetic DNA hairpins. Biophys. J. 78:817Symp.
Bevers, S., O’Dea, T.P., and McLaughlin, L.W. 1998. Perylene- and naphthalene-based linkers for duplex and triplex stabilization. J. Am. Chem. Soc. 120:11004-11005.
Lewis, F.D., Wu, T.F., Liu, X.Y., Letsinger, R.L., Greenfield, S.R., Miller, S.E., and Wasielewski, M.R. 2000b. Dynamics of photoinduced charge separation and charge recombination in synthetic DNA hairpins with stilbenedicarboxamide linkers. J. Am. Chem. Soc. 122:2889-2902.
Bevers, S., Schutte, S., and McLaughlin, L.W. 2000. Naphthalene- and perylene-based linkers for the stabilization of hairpin triplexes. J. Am. Chem. Soc. 122:5905-5915.
Lewis, F.D., Letsinger, R.L., and Wasielewski, M.R. 2001. Dynamics of photoinduced charge transfer and hole transport in synthetic DNA hairpins. Accounts Chem. Res. 34:159-170.
Letsinger, R.L. and Wu, T.F. 1995. Use of a stilbenedicarboxamide bridge in stabilizing, monitoring, and photochemically altering folded conformations of oligonucleotides. J. Am. Chem. Soc. 117:7323-7328.
Nelson, J.S., Giver, L., Ellington, A.D., and Letsinger, R.L. 1996. Incorporation of a non-nucleotide bridge into hairpin oligonucleotides capable of high-affinity binding to the Rev protein of HIV-1. Biochemistry 35:5339-5344.
Lewis, F.D., Wu, T.F., Burch, E.L., Bassani, D.M., Yang, J.S., Schneider, S., Jager, W., and Letsinger, R.L. 1995. Hybrid oligonucleotides containing stilbene units—Excimer fluorescence and photodimerization. J. Am. Chem. Soc. 117:87858792.
Yamana, K., Yoshikawa, A., and Nakano, H. 1996. Synthesis of a new photoisomerizable linker for connecting two oligonucleotide segments. Tetrahedron Lett. 37:637-640.
Lewis, F.D., Wu, T.F., Zhang, Y.F., Letsinger, R.L., Greenfield, S.R., and Wasielewski, M.R. 1997. Distance-dependent electron transfer in DNA hairpins. Science 277:673-676. Lewis, F.D., Liu, X.Y., Wu, Y., Miller, S.E., Wasielewski, M.R., Letsinger, R.L., Sanishvili, R., Joachimiak, A., Tereshko, V., and Egli, M. 1999. Structure and photoinduced electron transfer in exceptionally stable synthetic DNA hairpins with stilbenediether linkers. J. Am. Chem. Soc. 121:9905-9906.
Yamana, K., Yoshikawa, A. Noda, R., and Nakano, H. 1998. Synthesis and binding properties of oligonucleotides containing an azobenzene linker. Nucleosides Nucleotides 17:233-242.
Contributed by Maneesh R. Pingle, Pei-Sze Ng, Xiaolin Xu, and Donald E. Bergstrom Purdue University West Lafayette, Indiana
Methods for Cross-Linking Nucleic Acids
5.6.15 Current Protocols in Nucleic Acid Chemistry
Supplement 12
Engineering Terminal Disulfide Bonds Into DNA Characterization of synthetic oligonucleotides shapes much of the understanding of “native,” higher-molecular-weight DNA and RNA molecules. Although of immense utility, short oligonucleotides usually possess lower structural and thermal stability and have greater end effects than the larger nucleic acid constructs they are intended to model. Hence, the physiochemical properties of oligonucleotides may not always compare favorably to those of larger nucleic acids (Elson et al., 1970; Scheffler et al., 1970; Baldwin, 1971; Record and Lohman, 1978; Breslauer, 1986; Breslauer et al., 1986; Olmsted et al., 1991; MacGregor, 1996; SantaLucia et al., 1996). Arguably, the most successful approach to stabilize oligonucleotides is to connect the strands that comprise a helical structure with an interstrand cross-link (Pinto and Lippard, 1985; Borowy-Borowski et al., 1990; Kirchner and Hopkins, 1991; Boger et al., 1991; Sigurdsson et al., 1993; Willis et al., 1993). Methods to cross-link nucleic acids can generally be divided into two catagories. In the first category, cross-links are formed using an exogenous reagent, most commonly a (bis)electrophile. Because (bis)electrophiles can react with nearly all of the nucleophilic sites on the bases, these agents often have little or no sequence specificity and form complex mixtures of cross-linked aducts (Millard et al., 1990, 1991; Kirchner et al., 1992). Although some natural products (like mitomycin C) and synthetic compounds (such as cisplatin and psoralen) can form lesions at unique sites, these reagents require the presence of specific recognition sites within a target sequence to generate the cross-link, which limits their utility (Sherman et al., 1985; Tomasz et al., 1987; Teng et al., 1989; Lemaire et al., 1991). In some cases, cross-link formation by alkylating agents can also have undesirable effects such as disrupting base stacking (Sherman et al., 1985; Millard et al., 1990). In the second category, cross-links can be introduced into oligonucleotides through solidphase synthesis of oligomers site-specifically labeled with modified nucleosides that present reactive groups. Positioning the reactive groups in proximity on opposing strands of a helix allows for the formation of a cross-link. This general strategy requires appropriate selection of both loci to be bridged, as well as a chemistry
UNIT 5.7
to form the cross-link. In one of the most effective examples of this approach, Webb and Matteucci (1986) demonstrated that DNA oligomers containing cytosines bearing an aziridine group on the N4 position form cross-links after annealing to a complementary oligomer, followed by selective opening of the aziridine by the exocyclic amine of an opposing dC or dA. These cross-links have proved useful for some experiments, but this chemistry in particular, and related methods in general, have several limitations. For example, ethanobridged cross-links are not formed in high yield, the internal mismatch required to form the cross-link can give rise to (local) disruption of helical geometry, and the resulting crosslinks can potentially interfere with protein and drug binding (Catalano and Benkovic, 1989; Cowart et al., 1989; Cowart and Benkovic, 1991). An alternate approach involves incorporating an intrastrand cross-link at the terminus of a helical structure. For example, one end of a duplex can be covalently linked by bridging the 3′-and 5′-terminal hydroxyl groups with either oligonucleotide, oligoglycol, or alkyl linkers to form stem-loop structures or “hairpins” (Durand et al., 1990; Kool, 1991; Rumney and Kool, 1992; Salunkhe et al., 1992; Bannwarth et al., 1994; Gao et al., 1994, 1995; Williams and Hall, 1996). Both ends of a duplex can also be covalently capped with linkers to generate double hairpins or “dumbbells” (Germann et al., 1985; Wemmer and Benight, 1985; Erie et al., 1987, 1989; Benight et al., 1988; Ashley and Kushlan, 1991; Amaratunga et al., 1992; Doktycz et al., 1992; Paner et al., 1992). Because dumbbells denature in a monomolecular fashion and do not suffer from end effects, they are particularly useful in thermodynamic experiments (Erie et al., 1987, 1989). However, the synthesis of DNA dumbbells has in some cases proved to be quite challenging, thus limiting their utility (Erie et al., 1987). In addition, because the linkers are tethered to the terminal hydroxyl groups, some standard enzymatic manipulations such as end-labeling are not possible.
Contributed by Gary D. Glick Current Protocols in Nucleic Acid Chemistry (2003) 5.7.1-5.7.13 Copyright © 2003 by John Wiley & Sons, Inc.
Methods for Cross-Linking Nucleic Acids
5.7.1 Supplement 13
DESIGN OF AN IDEAL CROSS-LINK To develop a cross-linking approach that would be simple and general and would overcome the drawbacks of other methods, the author of this unit, and colleagues, chose eight important criteria that should be met (Table 5.7.1). It was clearly desirable to incorporate the cross-links by solid-phase synthesis to permit the modifications to be placed site-specifically and in close proximity within any given target sequence. Yet it was critical to find chemistry that would be specific and efficient for cross-link formation. Thiols were selected as the reactive functional groups because the mild Table 5.7.1
Requirements for an “Ideal” Nucleic Acid Cross-Linka
Requirementsb
Rationale
Potential solution
Increase the conformational stability towards thermal-, ionic-, pH-, and concentration-induced conformational changes Form interstrand cross-link site-specifically
Conformationally stable nucleic acids that are structurally homogeneous will facilitate their structural and thermodynamic analysis Limits undesired adduct formation resulting in high yields of cross-linked product Necessary if the structural aspects of cross-linked constructs are to be compared to their unmodified nucleic acid precursors
Covalent interstand cross-links confer a large degree of conformational stability
Allows for protein and ligand-nucleic acid interactions to be studied; maximizes (thermal) stability
Linkers at terminus of helix
Cross-link does not alter native geometry
Grooves of the helices, counter-ion binding, and hydration must remain unaltered
Engineering Terminal Disulfide Bonds Into DNA
redox chemistry used to form disulfide bonds from thiols is specific for sulfur. In addition, disulfide bonds are formed in high yield, often quantitatively, and are stable to a wide variety of solvents and reagents, yet are cleaved by reduction. Finally, the use of disulfide bonds to constrain macromolecular architecture has already been demonstrated in the peptide/protein literature (Clarke and Fersht, 1993, and references therein). The next step in developing this chemistry was to select appropriate loci for modification. Because cross-links residing in either the major or minor groove could interfere with ligand recognition and hydration, cross-links were de-
Position functional groups proximal to one another on opposing strands of a helix Position cross-link within a sterically accessible space
3′- and 5′-hydroxyl groups are Needed for wide range of free to [32P]-end-label biochemical assays (e.g., footprinting/sequencing) Prepared and isolated in large Allows for high-resolution quantities physical measurements such as NMR, DSCc, and X-ray diffraction studies
Cross-link located on the base rather than terminal hydroxyl groups Efficient cross-linking chemistry
Reversible
Assess structural and thermodynamic effects of base modification before and after cross-linking
Reversible disulfide cross-link
Flexible to a wide variety of nucleic acid structures
Broad applicability will be necessary if this methodology is to be used uniformly in nucleic acids
Position functional tether at any base or sugar site; cross-link within a sterically accessible space
aFrom Glick (1998). bRelative to the unmodified nucleic acid. cDSC, differential scanning calorimetry.
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Current Protocols in Nucleic Acid Chemistry
signed at the terminus of the helical structure (Figure 5.7.1). To avoid using the 3′- and 5′-hydroxyl groups, the terminal bases themselves were modified. This strategy exploited the fact that the terminal residues in duplexes have a reduced stability due to end effects, often referred to as “end fraying” (Patel and Hilbers, 1975). Therefore, altering the hydrogen bonding groups of the two opposing bases at the terminus of a duplex should not adversely affect helical stability, provided the thiol modifications do not disrupt base stacking. The free energy of a dT-dT mismatch located at the terminus of a duplex is ∼1.0 kcal/mol more stable than a dT-dT mismatch within a duplex (Aboul-ela et al., 1985; Freier et al., 1986). While this energetic penalty could be compensated for upon formation of the cross-link, it necessarily limits the overall (thermodynamic) stabilization the cross-link can provide. When two thymidine residues directly oppose each other in duplex DNA, their N3 positions project toward the center of the helix and converge to ∼4.5 Å. Based on this observation, replacing the terminal bases on one or both ends of a duplex with N3-(alkylthiol)thymidine should be an ideal way to form interstrand disulfide cross-links. Molecular modeling studies suggest that disulfide cross-links can form when the terminal bases of a duplex are
A
replaced with either N3-(methylthiol)thymidine or N3-(ethylthiol)thymidine. Since N3(methylthiol)thymidine is expected to decompose under the acidic conditions used to detritylate synthetic oligonucleotides, N3-(ethyl thiol)thymidine (T*SH) was chosen (Figure 5.7.1B). The following sections describe key aspects pertaining to the synthesis, structure, dynamics, thermodynamic stability, and in vitro biological properties of these disulfide crosslinked DNA oligonucleotides. Two applications of this cross-linking chemistry are then discussed, as are recent experiments using different thiol-modified nucleosides to generate disulfide cross-links within RNA secondary and tertiary structure. Lastly, other methods that have been developed to incorporate disulfide cross-links into nucleic acids are briefly outlined.
DISULFIDE CROSS-LINKED DNA DUPLEXES Synthesis The properties of disulfide cross-linked DNA duplexes were explored by synthesizing analogs of two previously well-studied oligonucleotides. The first is a hairpin whose sequence is derived from the ColE1 cruciform
B O SR N N
O
dRib s s
T*SH; R = H T*tBu; R = StBu T*2 ; R = ) 2
Figure 5.7.1 ((A) Synthetic strategy. To place cross-links within a duplex requires synthesizing thiol-modified nucleosides, incorporating the modified bases within a target by solid-phase synthesis, removal of all protecting groups, and air oxidation to form the cross-link. (B) Chemical structure of modified thymidine and cross-link. During synthesis, the thiol group on T*SH is protected as a tert-butyl mixed disulfide (T*tBu) and this protecting group is stable to all conditions of solid-phase synthesis, deprotection, and purification. The coupling efficiency of this base is indistinguishable from thymidine (>99% per cycle). To place T*tBu at the 3′ terminus, the 3′-hydroxyl group is attached to controlled-pore glass. For T*2, R is the disulfide cross-link. From Glick (1998).
Methods for Cross-Linking Nucleic Acids
5.7.3 Current Protocols in Nucleic Acid Chemistry
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5′ GGC A A T CC CCG T T AGG
1
C
A
CGCGA A T T CGC G T T
C
A C G C G A A T T C G C T*tBu
T
2
4a
5a
A T
T
T*tBu G C G A A T T C G C T*tBu
G C G C T T A A G C G T*tBu
T
T* G C A A T C C T* C G T T A G G
2b
3
T
C S S
GCGC T T A AGCG C
T
T*tBu G C A A T C C T*tBu C G T T A G G
2a
5′
C G C G A A T T C G C T* G C G C T T A A G C G T*
T
S S
S S
4b
T* G C G A A T T C G C T* T* C G C T T A A G C G T*
S S
5b
Figure 5.7.2 Duplex sequences and cross-linked analogs. From Glick (1998).
(S.1; Figure 5.7.2; Blatt et al., 1993), and the second is the Dickerson/Drew dodecamer (S.3; Wing et al., 1980). Synthesis of S.2a, S.4a, and S.5a is conducted using standard phosphoramidite chemistry using N-benzoyl- and N-isobutyryl-protected bases (Glick et al., 1992; Cain et al., 1995; Osborne et al., 1996). After deprotection and purification of each thiol-modified DNA by reversed-phase HPLC, the t-butyl mixed disulfide protecting groups on each sequence are cleaved with DTT (∼20 equiv per thiol, pH 8, overnight at 4°C). Reduced DNA samples are separated from the DTT by reversed-phase HPLC. Disulfide bond formation is performed by dissolving each sample (DNA concentration of ∼2 µM) in phosphate buffer (pH 8.3) and stirring vigorously at 25°C with exposure to air. The reactions are usually complete in 8 hr, as determined by a negative Ellman’s test. The cross-linked DNAs (S.2b, S.4b, and S.5b) are then isolated by either PAGE or HPLC, and incorporation of the modified bases is confirmed by enzymatic digestion. Importantly, disulfide bond formation is quantitative.
Structural Studies
Engineering Terminal Disulfide Bonds Into DNA
The effects of introducing the thiol modifications were first assessed by comparing UV thermal denaturation profiles of S.2a to that of the corresponding wild-type sequence (S.1). The melting temperatures (Tm) values are within 1°C and the transitions are nearly superimposable, which suggests that the thiol linkers
do not alter the stability or thermal denaturation pathways of the modified hairpin relative to its unmodified counterpart (Glick, 1991). These data also suggest that all of the stability conferred by the disulfide bond will be reflected in the cross-linked structure, since there is not a significant energetic penalty for replacing the terminal bases with the N3-thiol-modified thymidines. Introducing a disulfide cross-link in S.1 to generate S.2b increases the Tm from 65° to 81°C in low-ionic-strength buffers. If the concentration of Na+ is >100 mM, the crosslinked hairpin denatures above 96°C and Tm values cannot be measured by UV spectroscopy (Glick, 1991). To further investigate the structural effects of the disulfide modification, the solutionphase conformations of S.1 and S.2b were determined by NMR spectroscopy (Cain et al., 1995). The 8-bp-long stem of these sequences adopts a B-form helix, whereas the 5-bp-long single-stranded loop appears to be flexible and cannot be represented by a unique static conformation. NOESY cross-peak volumes, proton (both labile and non-labile) and phosphorus chemical shifts, as well as both homo- and heteronuclear coupling constants for the crosslinked hairpin are virtually identical to those measured for the unmodified sequence, even for the residues that are proximal to the crosslink. Thus, within the resolution of NMR spectroscopy, the two hairpins are structurally isomorphous.
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Similar to the results described above for the ColE1 hairpin, replacing one or both of the term inal dG-dC base pairs in d(CGCGAATTCGCG)2 with the modified thymidine does not significantly alter the thermal stability of these duplexes relative to the parent sequence (Osborne, 1996; Osborne et al., 1996). When the exchangeable and non-exchangeable protons and the 31P resonances of S.5b are assigned and compared to those for the parent duplex, S.3, the spectra are virtually identical, which strongly indicates that these sequences also adopt very similar structures. (Because S.4b is not symmetric, a direct comparison with either S.3 or S.5b is not possible.) To provide further evidence of this point, 2-D NOESY spectra of S.5b as a function of mixing time can be measured and interproton distances obtained from initial build-up plots. These data for S.5b are nearly identical to those for the wild-type dodecamer. Based on these findings, along with the results of CD and nuclease-susceptibility experiments, these alkylthiol modifications, in either the disulfide cross-linked or reduced and protected forms, do not alter native structure (Osborne et al., 1996).
Thermodynamic Measurements To elucidate the thermal and thermodynamic consequences of modifying and constraining DNA duplexes via the disulfide chemistry, differential scanning calorimetry (DSC) measurements can be conducted to characterize their thermally induced denaturation. DSC measurements on duplexes S.3 to S.5 (Osborne et al., 1996) represent the first use of calorimetry to investigate the thermodynamic consequences of constraining DNA with disulfide cross-links. The use of calorimetry is critical here because S.3 does not denature in a twostate fashion and cannot be analyzed using a van’t Hoff treatment. Consistent with the NMR data, DSC data show that a large energetic penalty is not incurred by replacing the terminal base pair(s) with N3-(ethylthiol)thymidine: the Tm values of S.3, S.4a, and S.5a are within 11°C of each other. The overall free energies for S.3, S.4a, and S.5a are comparable; this similarity arises due to compensations in both enthalpy and entropy (Osborne, 1996). The DSC measurements suggest that introducing cross-link(s) into S.3 results in two changes. First, constraining the dodecamer results in a significant increase in thermal stability. A single disulfide cross-link changes the molecularity of the complex from bimolecular to monomolecular (S.4b). However, a second
disulfide cross-link (S.5b) results in a constrained conformation with a reduced entropy compared to S.4b. From the DSC data, the cross-link imparts ∼3 kcal/mol of stabilization. The change in entropy certainly reflects differences in the native and/or denatured states of S.5b compared to S.4b. On first inspection, it would appear that the most likely source for the observed decrease in entropy is the denatured state, since the conformational freedom of the denatured state of S.5b is less than that of S.4b. However, based on the data, differential entropic contributions from the native states, as well as influences from differential solvation in both the initial and the final states, cannot be excluded. Notwithstanding, the data show that the entropy value is responsible for the increase in thermal stability of S.5b relative to S.4b. This is the first conformationally constrained nucleic acid system where the increase in melting temperature is predominantly due to a decrease in entropy (Erie et al., 1987, 1989).
Dynamics Measurements The static structure of DNA can explain many aspects of its function and properties. However, a local base-pair opening is implicated in a number of important chemical, biological, and mechanical processes involving DNA (Frank-Kamenetskii, 1985; Ramstein and Lavery, 1988, 1990; Guéron and Leroy, 1995; Tari and Secco, 1995). Hence, defining the dynamics of base-pair opening is necessary to fully understand the physiochemical and biological properties of DNA. Although the opening kinetics for several different constructs have been measured, the opening kinetics for oligonucleotides constrained with cross-links (e.g., hairpin loops, glycol bridges) have not been reported (Gueron and Leroy, 1995). To address this, the base-pair lifetimes and apparent dissociation constants of S.1 and S.2b were measured (Cain and Glick, 1997). Comparison of the lifetimes and apparent dissociation constants for corresponding base-pairs of the two hairpins indicates that the cross-link neither increases the number of base-pairs involved in fraying nor alters the lifetime, dissociation constant, or the opened structure from which exchange occurs for the base-pairs that are not frayed. The cross-link does, however, stabilize the frayed penultimate base-pair of the stem duplex by increasing the closing rate of this base-pair. Significantly, the disulfide cross-link is more effective at preventing fraying than the 5-bp-long hairpin loop.
Methods for Cross-Linking Nucleic Acids
5.7.5 Current Protocols in Nucleic Acid Chemistry
Supplement 13
5′
5′
S S 3′
3′
Figure 5.7.3 A triplex schematic. From Glick (1998).
DISULFIDE CROSS-LINKED DNA TRIPLEXES Design
Engineering Terminal Disulfide Bonds Into DNA
Folding of DNA triple helices based on the pyrimidine⋅purine-pyrimidine motif (py⋅pupy; dot = Hoogsteen and dash = Watson-Crick base-pairing) is affected by a number of factors including sequence length (i.e., the number of triplets formed), composition, the presence of base-pair mismatches, as well as solution conditions including pH and monovalent and divalent counterion concentrations (Moser and Dervan, 1987; Pilch et al., 1990; Kiessling et al., 1992; Plum et al., 1995). For example, a major factor in the stability of triplexes that contain C+⋅G-C triplets is the necessity for protonation of the N3 position of cytosine, which generally limits the stability of small triple helices of this type to pH below 7 (Callahan et al., 1991; Singleton and Dervan, 1992). The narrow pH range required for folding of these triplexes has hampered efforts to assess the physical properties of this motif under physiological conditions. Such data are clearly needed to design triplex sequences of higher affinity and specificity for diagnostic purposes in vitro and for use in vivo. While methods exist to stabilize py⋅pu-py triple helices, they rely on modifications that may alter the native triplex structure (Sun and Hélène, 1993; Thuong and Hélène, 1993). Therefore, it became important to determine if covalently locking the third strand of a triplex to the major groove through a structurally nonperturbing disulfide cross-link would afford constructs that are stable under physiological conditions. In other words, can the constraint provided by a disulfide cross-link alter the apparent pKa of a triplex?
This question was tested by using an intramolecular triplex rather than an intermolecular construct because the former should require only one cross-link to link the Hoogsteen strand to the Watson-Crick duplex (Figure 5.7.3; Goodwin et al., 1994; Osborne et al., 1997). To place a cross-link between the Hoogsteen and Watson-Crick strands, sites of chemical modification on the terminal bases must first be identified. In a T⋅T-A triplet, which is analogous to the base-pair substitution used to crosslink the terminus of duplexes (see Figure 5.7.1), the N3 position on the Hoogsteen thymidine and the C5 position of the Watson-Crick thymidine converge. Since the cross-link used for B-form duplexes is inappropriate here, a triplex cross-link was designed using a C5alkylthiol-modified thymidine (C53S, where three is the number of atoms in the linker, including thiol) and N3-(ethylthiol)thymidine (Figure 5.7.4; Goodwin and Glick, 1993). The optimal pH for air oxidation of thiols to form disulfide bonds is generally above the pKa of the thiol of interest (∼8.5). At the pKa of many thiols, triple helices containing C+⋅G-C base triplets are unfolded because the N3 position of cytosine is not protonated. To form a disulfide cross-link in a py⋅pu-py triplet, it is necessary to identify a sequence that (partly) folds into a triplex at a pH where cross-link formation readily proceeds. Recently, Häner and Dervan (1990) reported a 34-bp-long intramolecular triple helix that remains partially folded at up to pH 7.5 at 24°C in a buffer containing 25 mM Mg2+. In principle, therefore, a disulfide crosslink can form under the conditions needed to fold a suitably modified variant of this oligonucleotide (Figure 5.7.5).
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dR
N
O N
O
O H
H N
H T
N
dR
N
dR O T
N A
O
H
N O
N
N
N
dR
N H
H
O
H
N
N
N
T H
N
dR
N N
N
A
O
dR T dR
N
O N
H
O O
S T*S
H
N
S H
N N O
dR
N
N
dR
N N A
C53S
Figure 5.7.4 Chemical structure of triplex cross-link. From Glick (1998).
Preparation of triplex sequences containing T*tBu and C53StBu can be achieved via automated solid-phase DNA synthesis using standard protocols with an average stepwise coupling efficiency of >98.5%. Removal of the phosphate- and base-protecting groups followed by reversed-phase HPLC purification provides a ∼42% yield of S.7a (based on a 1-µmol synthesis). To form the cross-link, the tert-butylthiol protecting groups are removed with DTT and the reduced sample is vigorously stirred at room temperature in PBS (pH 7.25, 5 mM MgCl2) while exposed to air. After 24 hr, no starting material is observed by HPLC, and aliquots from the reaction mixtures test negative with Ellman’s reagent. Cross-linked DNA is purified by reversed-phase HPLC to give S.7b in ∼28% isolated overall yield (based on
a 1-µmol synthesis; ∼75% yield from S.7a). Enzymatic nucleoside composition analysis confirms incorporation of the modified bases and formation of the cross-link, and native and denaturing PAGE indicates that the structure formed is monomeric.
Structural Studies When imino proton spectra of S.6 and S.7b are assigned using standard 2-D techniques under conditions that favor triplex formation (pH 6, 0.5 mM Mg2+), the imino proton spectra of both sequences are virtually identical, which indicates that the modifications do not disrupt native triplex structure (Osborne et al., 1997). When S.6 or S.7a is titrated from pH 6 to 8, dissociation of the third strand is observed. By contrast, the cross-linked triplex S.7b is con-
Methods for Cross-Linking Nucleic Acids
5.7.7 Current Protocols in Nucleic Acid Chemistry
Supplement 13
+
+
+
T27 C28 T29 C30 T31 T32 C33 X34 T21-26 5′- A1 G2 A3 G4 A5 A6 G7 Y8 T20 C19 T18 C17 T16 T15 C14 A13
T9-12
6: X, Y = T 7a: X = T*tBu 7b: X = T*S disulfide cross-link Y = C53S-StBu Y = C 5 3S
Figure 5.7.5 Triplex sequence. From Glick (1998).
formationally stable over this pH range and only begins to unfold at pH ∼9. Moreover, S.7b is stable from pH 6 to 8 in the absence of Mg2+. Because of line broadening above 5°C, NMR cannot be used to study these triplexes. Therefore, CD spectroscopy is used. Monitoring the triplex CD signature band at 215 nm in the presence of Mg2+, S.7b does not begin to melt until ∼60°C at pH 6 (30°C for S.6 and S.7a) and 40°C at pH 8 (<20°C for S.6 and S.7a). Constructing titration curves from the CD data reveals that the apparent pKa for S.7b is ∼8.6, which is at least 1.5 pKa units greater than S.6 and S.7a. These results clearly demonstrate that the presented cross-link can be used to stabilize higher-order DNA structures.
Thermodynamic Measurements
Engineering Terminal Disulfide Bonds Into DNA
DSC measurements can be conducted to elucidate the thermodynamic consequences of modifying and constraining DNA triplexes with the disulfide chemistry (Völker et al., 1997). Cross-linked triplexes S.6 and S.7a melt in a biphasic manner above pH 6, with the initial triplex-to-duplex transition (Hoogsteen strand release) occurring at lower temperatures than melting of the hairpin. In contrast, cross-linking increases the thermal stability of the Hoogsteen transition such that the triplex and hairpin duplex denature simultaneously. Model-independent thermodynamic data obtained by DSC reveal that the cross-link-induced increase in triplex thermal stability corresponds to a free energy stabilization of ∼3 kcal/mol, with this stabilization being entirely entropic in origin. In other words, the cross-link is enthalpically neutral, but nevertheless induces a triplex stabilization of 3 kcal/mol due to a reduction in the entropy change associated with triplex melting. To deduce the origin(s) of this entropic impact, the pH and ionic strength dependence
of the melting transitions were measured. From a comparison of the melting transitions at different pH values and ionic strengths, it can be estimated that 0.4 more protons are associated with the cross-linked triplex state than with the uncross-linked triplex, and 1.3 fewer counterions are released on melting the cross-linked triplex. Thus, the entropic stabilization is not solely a result of a reduction in conformational entropy.
Dynamics Measurements At the start of this project, it was not known whether the base-pairs (Hoogsteen and Watson-Crick) that comprise triple-helical DNA open to any significant extent within a stable triplex. To address this question, the conformational dynamics of S.6 and S.7b were studied by 2-D exchange and NOE spectroscopy, and by measuring base-catalyzed imino-proton exchange rates (Cain and Glick, 1998). Under conditions that promote triplex formation (pH 6.0, 1°C), S.6 and S.7b exhibit a small and identical degree of conformational heterogeneity. However, at higher temperatures (pH 6.0, 37°C), S.6 exhibits more extensive heterogeneity than S.7b. The exchange times for WatsonCrick imino protons are ∼1 hr for both triplexes. However, the Hoogsteen base-pair lifetimes of S.6 could not be measured because this sequence is conformationally labile under the alkaline conditions necessary to conduct the exchange experiments. Due to the extraordinary pH stability conferred by the cross-link, it is possible to measure the Hoogsteen lifetimes for S.7b. The lifetimes of the these base-pairs range from ∼3 to 370 msec. As for Watson-Crick base-pairs, the Hoogsteen lifetimes are highest at the central region within the triplex and taper off towards the termini, which is suggestive of imino ex-
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Current Protocols in Nucleic Acid Chemistry
5′ GGC A A T CC CCG T T AGG
N
N N
N
N = A, 8 T, 9 G, 10 C, 11
N
Figure 5.7.6 Hairpin ligands for the binding study. From Glick (1998).
change mechanisms involving end effects. In all cases, the lifetime of a Hoogsteen base-pair is shorter than that of the Watson-Crick basepair in the same triplet, which is consistent with the greater stability of Watson-Crick over Hoogsteen base-pairs. The rate of triplex-toduplex conversion for S.7b at 1°C and pH 6.9 is low, with an upper bound of 3.2 × 10−4 sec−1. The imino protons of S.7b exchange slowly in PBS with exchange times as long as 1 hr, but the base-pair lifetimes are all <3 min, reflecting the fact that imino proton exchange is not opening-rate limited. Because the disulfide crosslink effectively prevents conformational heterogeneity associated with py⋅pu-py triple helices containing C+⋅G-C base triplets at neutral pH, constructs possessing this modification can serve as model systems to examine the structural and thermodynamic aspects of triplex formation in vitro, and to develop sequences that bind DNA with higher affinity and specificity.
APPLICATIONS Defining the biochemical, structural, dynamic, and thermodynamic impacts of constraining DNA with the disulfide chemistry described above opens the way for applying these cross-links in a host of different experiments. Uses of this chemistry include, among others, the joining of large pieces of DNA or RNA through disulfide bonds for the construction of nano-scale architectures, the design of redox-activated switches/devices, and the probing of secondary and tertiary structure. Below is a description of two representative applications of this chemistry. In the first, a disulfide bond was employed to stabilize or “trap” a DNA structure that would not be stable in its absence; in the second, a disulfide bond was used to report on conformational transitions in a protein binding study.
Stabilizing Non-Ground-State Structures In the absence of disulfide cross-links, the molecules described in the above sections are
themselves relatively stable with respect to conformational isomerization. In principle, however, it should also be possible to trap higherenergy non-ground-state structures with disulfide cross-links. Synthetic access to such constructs should be important in areas such as targeted drug delivery, protein-DNA recognition, and biophysical studies of alternate DNA geometries. The practicality of using disulfide cross-links in such endeavors can be demonstrated from the example of trapping the Dickerson/Drew dodecamer premelting intermediate with a disulfide cross-link. Both S.3 and S.5a (Figure 5.7.2) thermally denature in a biphasic manner in buffers containing 10 mM Na+ and 50 µM DNA (Marky et al., 1983). The first transition defines melting of the duplex to a hairpin structure, while the second transition represents denaturation of the hairpin to a random coil. The premelting intermediate is cross-linked by air oxidation of the sulfhydryl groups in S.5b at 50°C in diluted solution with low-salt buffer (Glick et al., 1992). Unlike the stem-loop intermediate produced by initial melting of the parent dodecamer, the hairpin does not denature as a result of increasing temperature, Na+ concentration, or DNA concentration (Glick et al., 1992; Wang et al., 1994). To address the structural consequences of the cross-link, the solution-phase conformation of the hairpin was determined by NMR spectroscopy (Wang et al., 1994, 1995). The stem region of this hairpin forms a B-form DNA duplex with a helical rise of 3.5 Å and a helical twist of 34.6°. The first three nucleotides in the loop stack over the 5′ end of the helix and are followed by a sharp turn at residue T8, which acts to close the loop. The conformation agrees with the “loop folding principle” advanced by Haasnoot, which predicts extension of the helix by 3 bases followed by a sharp bend in the loop (Haasnoot et al., 1986). The aromatic bases face into the major groove while the negatively charged backbone is in contact with solution.
Methods for Cross-Linking Nucleic Acids
5.7.9 Current Protocols in Nucleic Acid Chemistry
Supplement 13
Significantly, the cross-link does not alter the geometry of the stem duplex.
SYNTHESIS OF DISULFIDE CROSS-LINKED DNAs USING CONVERTIBLE NUCLEOSIDES
Probes in Molecular Recognition Experiments
The chemistry developed and advanced by Verdine and co-workers stands as the best method to place cross-links within the helical regions of DNA (Ferentz et al., 1991, 1993). Using convertible nucleoside chemistry, they showed that placing N6-thioalkyl derivatives of 2′-deoxyadenosine in consecutive base pairs on opposite strands of a duplex afforded disulfide cross-links upon air oxidation. These crosslinks reside in the major groove and impart increased stability with minimal distortion of native DNA geometry. This chemistry was extended several years latter to the synthesis of minor groove cross-links using modified dC residues. These major and minor groove crosslinks have been used with excellent effect in a variety of ways, such as studying protein-DNA interactions (Erlanson et al., 1993, 1997), stabilizing intrinsically bent DNA (Wolfe and Verdine, 1993), inducing torsional stress in short oligonucleotides (Wolfe et al., 1995), and cross-linking Z-DNA (Wolfe and Verdine, 1993).
DNA ligands often undergo conformational changes upon binding to proteins. In the absence of X-ray data, this “induced fit” can be difficult to observe. In principle, the constraint imposed by the disulfide bond should provide a way to investigate conformational changes in DNA structure that occur upon protein recognition. This point can be illustrated by the binding of monoclonal antibody BV04-01 to disulfide-cross-linked analogs of hairpins S.8 through S.11 (Figure 5.7.6; Stevens et al., 1993). This anti-DNA autoantibody was isolated from an autoimmune mouse that develops a syndrome related to human lupus, and BV0401 may be involved in the pathogenesis of this disorder in mice. BV04-01 binds only ssDNA, which can be modeled by the loop region of DNA hairpins (Stevens et al., 1993, and references therein). If conformational reorganization of the hairpin ligands is required for binding, then BV0401 should possess a lower affinity for the crosslinked sequences, since the disulfide bond renders them resistant to structural changes. However, if preorganization is important for the formation of complexes, then the more rigid oligomers should bind with equal or greater affinity than the unmodified ligands. Measurements of BV04-01 binding to cross-linked hairpins reveals a nearly 100-fold increase in Kd relative to S.8 through S.11. If the weaker affinity of BV04-01 for the cross-linked molecules results from the constraint imposed by the cross-link rather than as a result of a structural perturbation introduced by the alkylthiol linkers, then removing this constraint by reduction of the disulfide bond should afford a set of ligands that bind with roughly the same affinity as the unmodified hairpins. Indeed, BV04-01 recognition of the reduced hairpins is indistinguishable from binding to S.8 through S.11. DNA footprinting experiments show that, upon binding, residues within the duplex are recognized by single-strand-specific reagents, which provides further evidence that the stem duplex of the hairpin ligands is partially denatured (Swanson et al., 1994).
Engineering Terminal Disulfide Bonds Into DNA
Literature Cited Aboul-ela, F., Koh, D., Tinoco, I. Jr., and Martin, F.H. 1985. Base-base mismatches. Thermodynamics of double- helix f ormation for dCA3XA3G + dCT3YT3G (X, Y = A,C,G,T). Nucl. Acids Res. 13:4811-4824. Amaratunga, M., Snowden-Ifft, E., Wemmer, D.E., and Benight, A.S. 1992. Studies of DNA dumbbells. II. Construction and characterization of DNA dumbbells with a 16 base-pair duplex stem and Tn end loops (n = 2, 3, 4, 6, 8, 10, 14). Biopolymers 32:865-879. Ashley, G.W. and Kushlan, D.M. 1991. Chemical synthesis of oligodeoxynucleotide dumbbells. Biochemistry 30:2927-2933. Baldwin, R.L. 1971. Experimental tests of the theory of deoxyribonucleic acid melting with d(TA) oligomers. Acc. Chem. Res. 4:265-272. Bannwarth, W., Dorn, A., Iaiza, P., and Pannekouke, X. 1994. Short optimally capped duplex DNA as a conformationally restricted analog of B-DNA. Helv. Chim. Acta 77:182-193. Benight, A.S., Schurr, J.M., Flynn, P.F., Reid, B.R., and Wemmer, D.E. 1988. Melting of a self-complementary minicircle. Comparison of optical melting theory with exchange broadening of the nuclear magnetic resonance spectrum. J. Mol. Biol. 200:377-399. Blatt, N.B., Osborne, S.E., Cain, R.J., and Glick, G.D. 1993. Conformational studies from the ColE1 cruciform. Biochimie 75:433.
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Current Protocols in Nucleic Acid Chemistry
Boger, D.L., Munk, S.A., and Ishizaki, T. 1991. (+)-CC-1065 DNA alkylation: Observation of an unexpected relationship between cyclopropane electrophile reactivity and the intensity of DNA alkylation. J. Am. Chem. Soc. 113:2779-2780. Borowy-Borowski, H., Lipman, R., and Tomasz, M. 1990. Recognition between mitomycin C and specific DNA sequences for cross-link formation. Biochemistry 29:2999-3006.
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Cain, R.J. and Glick, G.D. 1998. Use of cross-links to study the conformational dynamics of triplex DNA. Biochemistry 37:1456-1464. Cain, R.J., Zuiderweg, E.R.P., and Glick, G.D. 1995. Solution structure of a DNA hairpin and its disulfide cross-linked analog. Nucl. Acids Res. 23:2153-2160. Callahan, D.E., Trapane, T.L., Miller, P.S., Ts’o, P.O.P., and Kan, L.-S. 1991. Comparative circular dichroism and fluorescence studies of oligodeoxyribonucleotide and oligodeoxyribonucleoside methylphosphonate pyrimidine strands in duplex and triplex formation. Biochemistry 30:1650-1655. Catalano, C.E. and Benkovic, S.J. 1989. Inactivation of DNA polymerase I (Klenow fragment) by adenosine 2′,3′-epoxide 5′-triphosphate: Evidence for the formation of a tight-binding inhibitor. Biochemistry 28:4374-4382. Clarke, J. and Fersht, A.R. 1993. Engineered disulfide bonds as probes of the folding pathway of barnase: Increasing the stability of proteins against the rate of denaturation. Biochemistry 32:4322-4329. Cowart, M. and Benkovic, S.J. 1991. A novel combined ch emical-enzymic synthesis of crosslinked DNA using a nucleoside triphosphate analog. Biochemistry 30:788-796. Cowart, M., Gibson, K.J., Allen, D.J., and Benkovic, S.J. 1989. DNA substrate structural requirements for the exonuclease and polymerase activities of prokaryotic and phage DNA polymerases. Biochemistry 28:1975-1983. Doktycz, M.J., Goldstein, R.F., Paner, T.M., Gallo, F.J., and Benight, A.S. 1992. Studies of DNA dumbbells. I. Melting curves of 17 DNA dumbbells with different duplex stem sequences linked by T4 endloops: Evaluation of the nearest-neighbor stacking interactions in DNA. Biopolymers 32:849-864.
Erlanson, D.A., Wolfe, S.A., Chen, L., and Verdine, G.L. 1997. Selective base-pair destabilization enhances binding of a DNA methyltransferase. Tetrahedron 53:12041-12056. Ferentz, A.E. and Verdine, G.L. 1991. Disulfidecross-linked oligonucleotides. J. Am. Chem. Soc. 113:4000-4002. Ferentz, A.E., Keating, T.A., and Verdine, G.L. 1993. Synthesis and characterization of disulfide cross-linked oligonucleotides. J. Am. Chem. Soc. 115:9006-9014. Frank-Kamenetskii, M.D. 1985. Flunctuational motility of DNA. In Structure & Motion: Membranes, Nucleic Acids, and Proteins (E. Clementi, G. Corongiu, M.H. Sarma, and R.H. Sarma, eds.) pp. 417-432. Adenine Press, Guilderland, N.Y. Freier, S.M., Kierzek, R., Caruthers, M.H., Neilson, T., and Turner, D.H. 1986. Free energy contributions of G⋅U and other terminal mismatches to helix stability. Biochemistry 25:3209-3213. Gao, H., Chidambaram, N., Chen, B.C., Pelham, D.E., Patel, R., Yang, M., Zhou, L., Cook, A., and Cohen, J.S. 1994. Double-stranded cyclic oligonucleotides with non-nucleotide bridges. Bioconjugate Chem. 5:445-453. Gao, H., Yang, M., and Cook, A.F. 1995. Stabilization of double-stranded oligonucleotides using backbone-linked disulfide bridges. Nucl. Acids Res. 23:285-292. Germann, M.W., Schoenwaelder, K.-H., and van de Sande, J.H. 1985. Right- and left-handed (Z) helical conformations of the hairpin d(CG)5T4(C-G)5 monomer and dimer. Biochemistry 24:5698-5702. Glick, G.D. 1991. Synthesis of a conformationally restricted DNA hairpin. J. Org. Chem. 56:67466747. Glick, G.D., 1998. Design, synthesis, and analysis of conformationally constrained nucleic acids. Biopolymers 48:83-96.
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Glick, G.D., Osborne, S.E., Knitt, D.S., and Marino, J.P. Jr. 1992. Trapping and isolation of an alternate DNA conformation. J. Am. Chem. Soc. 114:5447-5448. Goodwin, J.T. and Glick, G.D. 1993. Incorporation of alkylthiol chains at C-5 of deoxyuridine Tetrahedron Lett. 34:5549-5552. Goodwin, J.T., Osborne, S.E., Swanson, P.C., and Glick, G.D. 1994. Synthesis of a disulfide crosslinked DNA triple helix. Tetrahedron Lett. 35:4527-4531. Guéron, M. and Leroy, J.L. 1995. Studies of base pair kinetics by NMR measurement of proton exchange. Methods Enzymol. 261:383-413. Haasnoot, C.A.G., Hilbers, C.W., van der Marel, G.A., van Boom, J.H., Singh, U.C., Pattabiraman, N., and Kollman, P.A. 1986. On loop folding in nucleic acid hairpin-type structures. J. Biomol. Struct. Dyn. 3:843-857.
Moser, H.E. and Dervan, P.B. 1987. Sequence-specific cleavage of double helical DNA by triple helix formation. Science 238:645-650. Olmsted, M.C., Anderson, C.F., and Record, M.T. Jr. 1991. Importance of oligoelectrolyte end effects for the thermodynamics of conformational transitions of nucleic acid oligomers: A grand canonical Monte Carlo analysis. Biopolymers 31:1593-1604. Osborne, S.E. 1996. Ph.D. Thesis, University of Michigan.
Häner, R. and Dervan, P.B. 1990. Single-stranded DNA triple-helix formation. Biochemistry 29:9761-9765.
Osborne, S.E., Völker, J., Stevens, S.Y., Breslauer, K.J., and Glick, G.D. 1996. Design, synthesis, and analysis of disulfide cross-linked DNA duplexes. J. Am. Chem. Soc. 118:11993-12003.
Kiessling, L.L., Griffin, L.C., and Dervan, P.B. 1992. Flanking sequence effects within the pyrimidine triple-helix motif characterized by affinity cleaving. Biochemistry 31:2829-2834.
Osborne, S.E., Cain, R.J., and Glick, G.D. 1997. Structure and dynamics of disulfide cross-linked DNA triple helices. J. Am. Chem. Soc. 119:11711182.
Kirchner, J.J. and Hopkins, P.B. 1991. Nitrous acid cross-links duplex DNA fragments through deoxyguanosine residues at the sequence 5′-CG. J. Am. Chem. Soc. 113:4681-4682.
Paner, T.M., Amaratunga, M., and Benight, A.S. 1992. Studies of DNA dumbbells. III. Theoretical analysis of optical melting curves of dumbbells with a 16 base-pair duplex stem and Tn end loops (n = 2, 3, 4, 6, 8, 10, 14). Biopolymers 32:881-892.
Kirchner, J.J., Sigurdsson, S.T., and Hopkins, P.B. 1992. Interstrand cross-linking of duplex DNA by nitrous acid: Covalent structure of the dG-todG cross-link at the sequence 5′-CG. J. Am. Chem. Soc. 114:4021-4027. Kool, E.T. 1991. Molecular recognition by circular oligonucleotides: Increasing the selectivity of DNA binding. J. Am. Chem. Soc. 113:62656266. Lemaire, M.-A., Schwartz, A., Rahmouni, A.R., and Leng, M. 1991. Interstrand cross-links are preferentially formed at the d(GC) sites in the reaction between cis-diamminedichloroplatinum(II) and DNA. Proc. Natl. Acad. Sci. U.S.A. 88:19821985. MacGregor, R.B. Jr. 1996. Chain length and oligonucleotide stability at high pressure. Biopolymers 38:321-327. Marky, L.A., Blumenfeld, K.S., Kozlowski, S., and Breslauer, K.J. 1983. Salt-dependent conformational transitions in the self-complementary deoxydodecanucleotide d(CGCAATTCGCG): Evidence for hairpin formation. Biopolymers 22:1247-1257. Millard, J.T., Raucher, S., and Hopkins, P.B. 1990. Mechlorethamine cross-links deoxyguanosine residues at 5′-GNC sequences in duplex DNA fragments. J. Am. Chem. Soc. 112:2459-2460. Engineering Terminal Disulfide Bonds Into DNA
Millard, J.T., Weidner, M.F., Kirchner, J.J., Ribeiro, S., and Hopkins, P.B. 1991. Sequence preferences of DNA interstrand crosslinking agents: Quantitation of interstrand crosslink locations in DNA duplex fragments containing multiple crosslinkable sites. Nucl. Acids Res. 19:18851891.
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Salunkhe, M., Wu, T., and Lestinger, R.L. 1992. Control of folding and binding of oligonucleotides by use of a nonnucleotide linker. J. Am. Chem. Soc. 114:8768-8772. SantaLucia, J. Jr., Allawi, H.T., and Seneviratne, P.A. 1996. Improved nearest-neighbor parameters for predicting DNA duplex stability. Biochemistry 35:3555-3562. Scheffler, I.E., Elson, E.L., and Baldwin, R.L. 1970. Helix formation by d(TA) oligomers. II. Analysis of the helix-coil transitions of linear and circular oligomers. J. Mol. Biol. 48:145-171.
Tomasz, M., Lipman, R., Chowdary, D., Pawlak, J., Verdine, G.L., and Nakanishi, K. 1987. Isolation and structure of a covalent cross-link adduct between mitomycin C and DNA. Science 235:1204-1208. Völker, J., Osborne, S.E., Glick, G.D., and Breslauer, K.J. 1997. Thermodynamic properties of a conformationally constrained intramolecular DNA triple helix. Biochemistry 36:756-767. Wang, H., Osborne, S.E., Zuiderweg, E.R.P., and Glick, G.D. 1994. Three-dimensional structure of a disulfide-stabilized non-ground-state DNA hairpin. J. Am. Chem. Soc. 116:5021-5022.
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Swanson, P.C., Cooper, B.C., and Glick, G.D. 1994. High-resolution epitope mapping of an antiDNA autoantibody using model DNA ligands. J. Immunol. 152:2601-2612.
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Tari, L.W. and Secco, A.S. 1995. Base-pair opening and spermine binding—B-DNA features displayed in the crystal structure of a gal operon fragment: Implications for protein-DNA recognition. Nucl. Acids Res. 23:2065-2073.
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Teng, S.P., Woodson, S.A., and Crothers, D.M. 1989. DNA sequence specificity of mitomycin cross-linking. Biochemistry 28:3901-3907. Thuong, N.T. and Hélène, C. 1993. Sequence-specific recognition and modification of doublehelical DNA by oligonucleotides. Angew. Chem. Int. Ed. Engl. 32:666-690.
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Contributed by Gary D. Glick University of Michigan Ann Arbor, Michigan
The author wishes to thank the talented group of associates in his laboratory who have so skillfully conducted the research described in this unit. He also thanks K.J. Breslauer, Jens Völker, and E.R.P. Zuiderweg, whose expertise and insight into nucleic acids has formed the basis for outstanding collaborations. The worked described here was supported by NIH Grants GM-52831 and GM43861.
Methods for Cross-Linking Nucleic Acids
5.7.13 Current Protocols in Nucleic Acid Chemistry
Supplement 13
CHAPTER 6 Chemical and Enzymatic Probes for Nucleic Acid Structure INTRODUCTION ver the past 10 years, significant progress has been made in elucidating the structure of nucleic acids to atomic resolution. By comparison with proteins, however, relatively few high-resolution nucleic structures have been determined. One main reason why crystallographic/NMR studies of nucleic acids have proven difficult is because oligonucleotides often self-associate and/or equilibrate between several different conformations at the concentrations required for many high-resolution biophysical measurements. To help circumvent these problems, an array of chemical and biochemical approaches have been developed to explore the conformation dynamics of oligonucleotides. Perhaps the most common way to assess structure is by chemical and enzymatic footprinting. In footprinting, the accessibility and/or reactivity of particular functional groups in either a secondary or tertiary structure is examined and compared to the same positions in the denatured state. Footprinting experiments often employ base- and phosphate-specific modifying reagents, hydroxyl radical and transition metal complexes, and a broad spectrum of nucleases. Based on the cleavage properties of these reagents, as determined in control experiments, local structure can be studied. The most critical point to bear in mind when designing footprinting experiments is that since chemical and enzymatic structure probing is indirect, several different reagents should be employed to examine any given region of structure.
O
This chapter will provide the reader with a comprehensive collection of protocols to examine the solution structures of both DNA and RNA by footprinting. The initial units focus more heavily on methods to study RNA structure, while units appearing in future supplements will integrate more DNA protocols. UNIT 6.1 provides an extremely thorough coverage of the most useful chemical and enzymatic probes to examine RNA secondary and tertiary structure. For the chemical probes, both strand scission and primer extension detection protocols are provided. Oftentimes, when RNA molecules adopt tertiary structure, distinct metal-binding pockets are created. Under appropriate conditions, this metal binding can be exploited to induce strand cleavage in the RNA backbone. This type of strand cleavage can be an exquisitely sensitive probe of local tertiary structure. UNITS 6.2-6.4 & 6.8 describe six of the most useful metal complexes for probing nucleic acid structure. describe the application of hydroxyl radicals to investigate RNA and DNA tertiary structure, respectively. Unlike nearly every other footprinting reagent, hydroxyl radicals cleave the sugar-phosphate backbone at every residue. This reagent possesses little or no sequence selectivity and will provide uniform cleavage at all positions in a given RNA or DNA secondary structure. Because some positions often become protected from cleavage upon tertiary folding, this reagent is useful to monitor global folding at equilibrium. Moreover, as will be described in Chapter 11, hydroxyl radical cleavage can also be used to follow the kinetics of RNA folding.
UNITS 6.5 & 6.7
Contributed by Gary D. Glick Current Protocols in Nucleic Acid Chemistry (2004) 6.0.1-6.0.2 C 2004 by John Wiley & Sons, Inc. Copyright
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.0.1 Supplement 17
presents a discussion of chemical reagents that can be used specifically to study the major groove of DNA. The unit focuses on the most commonly used and readily available reagents: dimethyl sulfate, diethylpyrocarbonate, potassium permanganate, osmium tetroxide, and bromine formed in situ from bromide and monoperoxysulfate. Characteristics of each reagent are reviewed, as are their applications to the study of DNA, DNA-protein complexes, and DNA-drug complexes.
UNIT 6.6
provides a method for analyzing RNA activity at the level of specific functional groups. In nucleotide analog interference mapping (NAIM), oligonucleotides with random phosphorothioate-tagged nucleotide analogs are assayed for an activity of interest (e.g., binding or folding). Inactive and active species are identified, and iodine cleavage of the phosphorothioate tag is used to identify the site of substitution. Modifications of the method, such as nucleotide interference analog suppression (NAIS), are also described. UNIT 6.9
Gary D. Glick
Introduction
6.0.2 Supplement 17
Current Protocols in Nucleic Acid Chemistry
Probing RNA Structure with Chemical Reagents and Enzymes
UNIT 6.1
The function of an RNA molecule in vivo and in vitro will often depend on its tertiary structure as well as the information encoded in its Watson-Crick base-pairing potential. There are now a number of powerful methods that can be used simultaneously to determine structural properties of small and large RNAs. This unit and subsequent units provide methods for RNA footprinting, detecting positions sensitive to chemical and enzymatic attack in the presence or absence of bound proteins. RNA footprinting differs profoundly from DNA footprinting in that the starting structure of the RNA cannot be taken for granted even in general terms, and the footprinting reactions are often carried out with structure-sensitive reagents precisely for the purpose of elucidating the RNA solution structure. When interpreting these structures, however, it is necessary to always consider whether the functional form of the RNA molecule is being examined. The first four protocols in this unit provide methods for modifying and cleaving RNA for the purpose of probing structure. Three of these protocols use structure-sensitive chemicals—dimethyl sulfate (DMS; see Basic Protocol 1), diethylpyrocarbonate (DEPC; see Alternate Protocol 1), and ethylnitrosourea (ENU; see Alternate Protocol 2). The fourth describes the use of various nucleases (see Table 6.1.1) to probe RNA structure (see Basic Protocol 2). Three other protocols can be used together to analyze the base-pairing status of RNA. The reagents modify Watson-Crick positions of nucleotides not involved in base pairing or tertiary hydrogen bonding. Dimethyl sulfate (see Basic Protocol 3) is used to probe unpaired adenines and cytidines, while kethoxal (see Alternate Protocol 3) and 1-cyclohexyl-3-(2-morpholinoethyl)carbodiimide metho-p-toluenesulfonate (CMCT; see Alternate Protocol 4) are used to investigate unpaired guanosines and uridines, respectively. RNA cleavages and base modifications can be detected by primer extension (see Support Protocol 4), or cleavage sites can be detected by using RNA that has been labeled at the 5′ terminus with T4 polynucleotide kinase (see Support Protocol 1) or the 3′ terminus by T4 RNA ligase (see Support Protocol 2). Support Protocol 3 details the preparation of electrophoresis standards for use with end-labeled RNA. CAUTION: When working with radioactivity, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following the guidelines provided by the local radiation safety officer. NOTE: Experiments involving RNA require careful precautions to prevent contamination and RNA degradation (see APPENDIX 2A). STRATEGIC PLANNING Footprinting with an Appropriate RNA Before undertaking these often labor-intensive footprinting experiments it is worthwhile to evaluate the extent to which all or most of the RNA sample to be examined is in an appropriate folded state (see Critical Parameters; for general guidelines for RNA folding, see UNIT 6.3). There are a number of ways of determining this when the RNA is to be probed in vitro; the method used depends on which experimental system is available to assess the structure of the RNA. In the simplest case, the RNA might have an assayable function that can be demonstrated in vitro. An example of this is an artificial RNA ligand selected for binding to a protein target in vitro. RNAs isolated from cells are generally more
Chemical and Enzymatic Probes for Nucleic Acid Structure
Contributed by William A. Ziehler and David R. Engelke
6.1.1
Current Protocols in Nucleic Acid Chemistry (2000) 6.1.1-6.1.21 Copyright © 2000 by John Wiley & Sons, Inc.
Table 6.1.1 RNA Structure-Probing Reagents and Their Specificities
Reagenta For cleavage: DMSc DEPCc ENUc Hydroxyl radicals Pb2+ Mung bean nucleased RNase A RNase CL3 RNase I(ONE) RNase Phy M RNase T1 RNase T2 RNase U2 RNase V1d S1 nucleased For modification: CMCT DMS Kethoxal
Targetb
Where discussed
ss or ds G (N7) ss or ds A (N7)
Basic Protocol 1 Alternate Protocol 1 Phosphate oxygens Alternate Protocol 2 Ribose sugar backbone UNIT 6.5 Phosphates of ss UNIT 6.3 nucleotides ss N/pN Basic Protocol 2 ss Cp/N and Up/N Basic Protocol 2 ss Cp/N Basic Protocol 2 ss Np/N Basic Protocol 2 ss Ap/N and Up/N Basic Protocol 2 ss Gp/N Basic Protocol 2 ss Np/N Basic Protocol 2 ss Ap/N Basic Protocol 2 ds N/pN Basic Protocol 2 ss N/pN Basic Protocol 2 Primarily U (N3) and possibly G (N1) A (N1) and C (N3) G (N1, 2-NH2)
Supplier Aldrich Sigma Sigma
Roche or Promega Roche USB Promega (RNase ONE) Pharmacia Life Technologies Life Technologies Pharmacia Pharmacia Roche or Promega
Alternate Protocol 4
Aldrich
Basic Protocol 3 Alternate Protocol 3
Aldrich ICN or Research Organics
aAbbreviations: CMCT, 1-cyclohexyl-3-(2-morpholinoethyl)carbodiimide metho-p-toluenesulfonate; DEPC, diethylpyrocarbonate; DMS, dimethyl sulfate; ds, double-stranded; ENU, ethylnitrosourea; ss, single-stranded. bCleavage site indicated by shill. Specific position of cleavage or modification are indicated in parentheses. cThis reagent requires further chemical processing to cleave the RNA. dNuclease requires divalent cations for activity. For RNase V1, MgCl (10 mM final) is recommended, mung bean nuclease 2 and S1 nuclease require zinc acetate as per supplier’s instructions.
difficult to interpret. If an RNA is stripped of its native protein structure in the process of being isolated from cells, or if an RNA is made synthetically, it should be assumed that the folding of that RNA is suspect until proven otherwise. Small RNAs that are tightly structured, such as tRNAs, can often refold in functional form, but this cannot be assumed. The chances of misfolding increase dramatically for larger RNAs where there are more potential isoforms. An RNA that is isolated as part of a ribonucleoprotein (RNP) complex, such as a ribosomal subunit, is generally assumed to be correctly folded, but even that assumption should be viewed with caution. Choice of Structure-Sensitive Reagents
Probing RNA Structure with Chemical Reagents and Enzymes
The object in RNA footprinting is to determine which positions in the RNA are accessible to specific types of attack by chemical reagents or nucleases. Cleavage or modification by these reagents is normally performed under conditions where the target RNA is cleaved at most once per molecule, thus lowering the chance that the first cleavage/modification causes an RNA rearrangement that alters access for a second attack. The cleavage or modification specificities of individual nucleases and commonly used chemicals are listed in Table 6.1.1. It is worth considering the general advantages and disadvantages of different classes of reagents. Nucleases have the lowest resolution because their steric radii do not allow access to many sites that are solvated; however, they are easily used in a wide range of physiological buffers. Most of the nucleases described below are specific
6.1.2 Current Protocols in Nucleic Acid Chemistry
for nucleotides not involved in Watson-Crick pairing, with variable sensitivity to other types of structure and base specificity. The exception is cobra venom ribonuclease (V1), which is the only reagent described here that is specific for double-stranded regions. Even V1 is not exceptionally helpful in identifying double-stranded structures, however, because the ability of V1 to cleave helices is variable. Cleavage by V1 is infrequent, and V1 can cleave adjacent to rather than within helices. The available chemical reagents discussed in this and subsequent units allow better access to all regions of the RNA tertiary structure and cumulatively attack a wide range of nucleotide positions. In general, using a larger number of different reagents to probe allows a more detailed view of solvent-exposed positions. Exposure of phosphates, sugars, and aromatic ring positions can be probed. Reagents specific for Watson-Crick positions are often used as diagnostics for the existence of standard base pairing at contiguous nucleotides. Some of the protocols described below are based on methods described previously (Peattie and Gilbert, 1980; Knapp, 1989; Krol and Carbon, 1989). Location of chemical and nuclease cleavages can be determined using end-labeled RNA or primer extension. Hydroxyl radicals (UNIT 6.5) and Pb2+ (UNIT 6.3) will directly hydrolyze the phosphodiester backbone, as will the various nucleases according to their sequence and structural preferences. Other reagents, including DMS, DEPC, and ENU, require additional chemical steps, ultimately resulting in strand scission. Detection of Modifications or Cleavage Sites The method used to detect cleavages or modified positions varies with both the nature of the RNA being probed and the reagents being used. Short RNAs (<200 nucleotides) can be labeled at either the 5′ or 3′ end (Support Protocols 1 and 2), then folded and subjected to reagents that result in RNA strand cleavage. Separation of the cleaved RNA on denaturing polyacrylamide gels and determination of fragment sizes identifies the position of cleavage sites. Alternatively, a labeled DNA oligomer primer can be annealed at any point along the length of an unlabeled RNA and extended to a point of cleavage or Watson-Crick base modification (Support Protocol 4). Either case terminates extension and produces a labeled DNA fragment corresponding to the length from the label to the termination site. There are a variety of labels that can be used with these two general detection schemes. The most common label is 32P, which can be detected with either X-ray film or a phosphorimager. The question of whether to use end labeling or primer extension for detection depends on the RNA to be analyzed and how many reagents are needed. Primer extension is more often useful for longer RNAs, although end labeling is still widely used to probe small RNAs. The advantage of end labeling is that it requires fewer manipulations (i.e., no primer-extension reactions) after the cleavage reactions. Its disadvantages include that it can only be used for relatively short RNAs that can be labeled before cleavage reagent treatment and that it requires cleavage of the RNA strand for detection. Transfer RNAs, 5S rRNAs, and other small, structured RNAs have typically been investigated by end labeling. Primer extension has four main disadvantages. The first is that the extreme 3′ end of the RNA cannot be probed because the primer must anneal to the 3′ side of the region to be extended across. The second disadvantage is that sequence-dependent pausing and termination by the reverse transcriptase, even on intact RNA, tends to give a high background at some positions that obscures the true signal from cleavage or modification. This problem can be reduced by using different reverse transcriptases and/or extension conditions, but is never completely eliminated with structured RNAs. The third disadvan-
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.1.3 Current Protocols in Nucleic Acid Chemistry
tage is that additional manipulations are required to perform the primer-extension reactions following the cleavage or modification reactions. Finally, cellular RNAs may contain posttranscriptional modifications that will terminate the extension reaction. However, primer-extension RNA footprinting is potentially useful in many situations. For example, RNA of any size can be probed because a labeled primer can be placed anywhere along the length of RNA, and multiple primers can be used in separate reactions to detect cleavages or modifications along the entire length. The RNA also does not need to be purified or labeled in advance, allowing probing of preformed RNA-protein complexes even in crude cell lysates. Following the cleavage/modification reactions, protein is removed by organic extraction and the purified RNA is subjected to primer extension. Lastly, the extension reactions can be used to detect several chemical modifications at Watson-Crick positions in addition to any form of RNA strand cleavage. BASIC PROTOCOL 1
MODIFICATION AND CLEAVAGE OF RNA USING DIMETHYL SULFATE Dimethyl sulfate (DMS) alkylates the N7 cyclic amine of guanosine. The methylated nucleoside is reduced, opening the imidizole ring and weakening the glycosidic bond. Aniline treatment catalyzes the β-elimination of the ribose sugar, leaving the nucleotide 5′ of the displaced guanosine with a 3′ phosphate and the nucleotide 3′ of the guanosine with a 5′ phosphate. The N7 position is pointed into the major groove of an A-form RNA helix and is accessible to modification. Therefore, the N7 of double-stranded and single-stranded guanosines should be detected unless participating in tertiary structure or non-Watson-Crick base-pair interactions. See Strategic Planning for a discussion of choice of RNA, folding, and detection methods. Materials 1 M HEPES, pH 7.8 1 M KCl 0.1 M MgCl2 1 µg/µL carrier RNA (see recipe) Sample RNA, end labeled (see Support Protocol 1 or Support Protocol 2) or unlabeled RNase-free water (see recipe) ≥99% dimethyl sulfate (DMS) (Aldrich) 3 M sodium acetate, pH 5.2 (APPENDIX 2A) 100% and 70% (v/v) ethanol 1 M Tris⋅Cl, pH 8.0 (APPENDIX 2A) 0.2 M NaBH4, prepared fresh 1 M aniline acetate buffer (see recipe) FEXS solution (see recipe; for end-labeled RNA) Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (e.g., APPENDIX 3B or CPMB UNIT 2.12) or primer extension (see Support Protocol 4) NOTE: DMS is highly toxic and a suspected carcinogen; use appropriate precautions for handling, storage, and disposal. Methylate guanosine N7 1. Prepare reaction mix as follows:
Probing RNA Structure with Chemical Reagents and Enzymes
8 µL 1 M HEPES, pH 7.8 (final 200 mM) 4 µL 1 M KCl (100 mM) 4 µL 0.1 M MgCl2 (10 mM) 8 µL 1 µg/µL carrier RNA (0.2 µg/µL).
6.1.4 Current Protocols in Nucleic Acid Chemistry
2. Combine 50,000 cpm end-labeled sample RNA or 0.5 µg unlabeled sample RNA (for primer extension) with RNase-free water to make 15 µL total and add to reaction mix. 3. Add 1 µL of 99+% DMS, mix, and incubate 2 min at room temperature. 4. Stop the reaction by precipitating with 4 µL of 3 M sodium acetate and 120 µL of 100% ethanol on dry ice. Microcentrifuge 30 min at maximum speed (10,000 rpm), 4°C. Remove supernatant, rinse with 70% ethanol, and dry the pellet in a speedvac. Reduce methylated RNA and perform aniline cleavage 5. Resuspend pellet with 20 µL of 1 M Tris⋅Cl and 20 µL of 0.2 M NaBH4. 6. Incubate 30 min on ice in the dark. 7. Precipitate as in step 4. 8. Resuspend pellet in 40 µL of 1 M aniline acetate buffer and incubate 15 min in the dark at 60°C. 9. Precipitate as in step 4. 10. If end-labeled RNA was used, resuspend in 12 µL FEXS solution and proceed with direct gel electrophoresis (e.g., APPENDIX 3B). If RNA sample was unlabeled, see Support Protocol 4 and perform primer extension. See Support Protocol 3 for preparation of appropriate end-labeled RNA standards for electrophoresis.
MODIFICATION AND CLEAVAGE OF RNA USING DIETHYLPYROCARBONATE
ALTERNATE PROTOCOL 1
Diethylpyrocarbonate (DEPC) alkylates the N7 cyclic amine of adenosine. The mechanism and procedure are essentially identical to those described for DMS (see Basic Protocol 1), except that DEPC is used and the incubation time is increased (replace step 3 in Basic Protocol 1 with the following step). See Strategic Planning for discussions of choice of RNA, folding, and detection methods. Additional Materials (also see Basic Protocol 1) ≥97% diethylpyrocarbonate (DEPC) (store up to 1 year at 2° to 8°C) NOTE: DEPC is toxic. Handle with appropriate care. 3b. Add 1 µL DEPC, mix, and incubate 45 min at room temperature.
MODIFICATION AND CLEAVAGE OF RNA USING ETHYLNITROSOUREA
ALTERNATE PROTOCOL 2
Ethylnitrosourea (ENU) alkylates phosphate oxygens that are not involved in tertiary structure interactions. These include phosphates of both single-stranded and doublestranded nucleotides not participating in higher-ordered structure. Following alkaline treatment, the phosphotriester hydrolyzes resulting in RNA strand scission. See Strategic Planning for discussions of choice of RNA, folding, and detection methods. Additional Materials (also see Basic Protocol 1) ENU/ethanol solution (see recipe) 0.1 M Tris⋅Cl, pH 9.0 (APPENDIX 2A)
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.1.5 Current Protocols in Nucleic Acid Chemistry
Alkylate phosphates 1. Prepare reaction mix as follows: 2 µL 1 M HEPES, pH 7.8 (final 200 mM) 1 µL 1 M KCl (100 mM) 1 µL 0.1 M MgCl2 (10 mM) 2 µL 1 µg/µL carrier RNA (0.2 µg/µL) 2. Combine 50,000 cpm end-labeled sample RNA or 0.5 µg unlabeled sample RNA (for primer extension) with RNase-free water to make 3 µL total and add to reaction mix. 3. Add 1 µL of ENU/ethanol solution, mix, and incubate 30 min at room temperature. 4. Stop the reaction by precipitating with 1 µL of 3 M sodium acetate and 30 µL of 100% ethanol on dry ice. Microcentrifuge 30 min at maximum speed (10,000 rpm), 4°C. Remove supernatant, rinse with 70% ethanol, and dry the pellet in a speedvac. Hydrolyze phosphotriesters 5. Resuspend pellet in 10 µL of 0.1 M Tris⋅Cl, pH 9.0, and incubate 5 min at 50°C. 6. Precipitate as in step 4. 7. If RNA has been end-labeled, resuspend in 12 µL FEXS solution and proceed with gel electrophoresis (e.g., APPENDIX 3B or CPMB UNIT 2.12). If RNA sample was not labeled, perform primer extension (see Support Protocol 4). See Support Protocol 3 for preparation of appropriate end-labeled RNA standards for electrophoresis. BASIC PROTOCOL 2
CLEAVAGE OF RNA USING NUCLEASES Nucleases hydrolyze the RNA phosphodiester backbone resulting in strand scission. Cleavages can be identified using primer extension (see Support Protocol 4) or end-labeled RNA (see Support Protocol 1 and Support Protocol 2). Table 6.1.1 lists nuclease specificities and cleavage locations. Some nucleases use divalent metal cofactors to catalyze the reaction. Divalent-independent nucleases can be used to probe RNA in the presence and absence of divalent-dependent tertiary structures. Most nucleases are specific for single-stranded bases and require access to the base determinants for cleavage site recognition. Uncleaved nucleotide targets may be obscured from cleavage by tertiary structure or local steric hindrance. Being relatively large, nucleases may not penetrate some RNA structures completely, and it is common for some potential targets to remain uncleaved. The data obtained from one nuclease will identify general structural trends (e.g., single- or double-stranded character), but often needs to be corroborated using additional nucleases or chemical probes. See Strategic Planning for discussions of choice of RNA, folding, and detection methods.
Probing RNA Structure with Chemical Reagents and Enzymes
Materials One of the following nucleases (see Table 6.1.1): Mung bean nuclease RNase A RNase CL3 RNase I(ONE) RNase Phy M RNase T1
6.1.6 Current Protocols in Nucleic Acid Chemistry
RNase T2 RNase U2 RNase V1 S1 nuclease 0.1 M Tris⋅Cl, pH 7.5 (APPENDIX 2A) 1 M KCl (APPENDIX 2A) 0.1 M MgCl2 1 µg/µL carrier RNA (see recipe) Sample RNA, end labeled (see Support Protocol 1 or Support Protocol 2) or unlabeled RNase-free water (see recipe) 3 M sodium acetate, pH 5.2 (APPENDIX 2A) 100% and 70% (v/v) ethanol FEXS solution (see recipe) Additional reagents and equipment for direct gel electrophoresis (e.g., APPENDIX 3B or CPMB UNIT 2.12) or primer extension (see Support Protocol 4) 1. Prepare six reaction tubes, each containing: 1 µL 0.1 M Tris⋅Cl, pH 7.5 (final 10 mM) 1 µL 1 M KCl (100 mM) 1 µL 0.1 M MgCl2 (10 mM) 2 µL 1 µg/µL carrier RNA (0.2 µg/µL). 2. For each reaction mix, combine 50,000 cpm end-labeled sample RNA or 0.5 µg unlabeled sample RNA (for primer extension) with RNase-free water to make 4 µL total. Add to each reaction mix. 3. Dilute nuclease at six 5-fold serial dilutions (covering four orders of magnitude) with 10 mM Tris⋅Cl to give the appropriate amounts in a volume of 1 µL. Dilute RNase V1 with 10 mM Tris⋅Cl and 10 mM MgCl2. Nuclease activity varies by lot and by manufacturer. Thus, it is necessary to titrate the amount of nuclease necessary to cleave ∼10% of the RNA of interest. A second titration consisting of narrower increments may be necessary once the desired activity is found. Table 6.1.2 lists activity units that have produced 10% cleavage using the reaction conditions described above. These should be used as guidelines only. Table 6.1.2 Quantities of Nuclease That Achieve ~10% Cleavage in 10 min
Nuclease Mung bean nuclease RNase A RNase CL3 RNase I(ONE) RNase Phy M RNase T1 RNase T2 RNase U2 RNase V1 S1 nuclease aND, not determined
Quantitya ND
0.2 ng 0.025-0.100 U 0.012 U 1.0 U 0.138 U ND
1.0 U 0.047 U ND
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.1.7 Current Protocols in Nucleic Acid Chemistry
Fresh nuclease dilutions should be used each time, although dilutions of RNases T1, A, and I(ONE) can be frozen with minimal loss of activity.
4. Add 1 µL of each nuclease dilution to a reaction mix, mix well, and incubate 10 min at room temperature. 5. Add 1 µL of 3 M sodium acetate and 30 µL of 100% ethanol and precipitate on dry ice. Microcentrifuge 30 min at maximum speed (10,000 rpm), 4°C. Remove supernatant, rinse with 70% ethanol, and dry the pellet in a speedvac. 6. If the RNA sample was end-labeled, resuspend in 12 µL of FEXS solution and proceed with gel electrophoresis (e.g., APPENDIX 3B or CPMB UNIT 2.12). If sample RNA was not labeled, see Support Protocol 4 and perform primer extension. See Support Protocol 3 for preparation of appropriate end-labeled RNA standards for electrophoresis. SUPPORT PROTOCOL 1
LABELING THE 5′ RNA TERMINUS USING T4 POLYNUCLEOTIDE KINASE AND [C-32P]ATP Bacteriophage T4 polynucleotide kinase will transfer the γ phosphate of [γ-32P]ATP to an RNA substrate containing a 5′ hydroxyl group, or it will exchange the radiolabeled γ phosphate with a single 5′ phosphate present on the RNA substrate. The 5′ end of cellular RNAs may contain methyl guanosine cap structures that prohibit labeling by this method. Chemical solid-phase synthesis produces an RNA with a 5′ hydroxyl available for radiolabeling using [γ-32P]ATP and polynucleotide kinase, whereas RNA transcribed in vitro has a 5′ triphosphate that must first be removed with a phosphatase. Dephosphorylation is accomplished with bacterial alkaline phosphatase and its accompanying buffer. An alternate method, not described here, of generating 5′-labeled RNA by enzymatic synthesis is to transcribe in vitro with T7 RNA polymerase and [γ-32P]GTP, incorporating 32 P at only the 5′ terminal triphosphate (Milligan and Uhlenbeck, 1989). This protocol can also be used to produce 5′ end-labeled DNA oligomers to be used as primers in primer extension (see Support Protocol 4). This is done by performing the labeling reaction below (steps 8 to 11) with the appropriate DNA oligomer. Materials RNA of interest 150 U/µL bacterial alkaline phosphatase (BAP; Life Technologies) RNase-free water (see recipe) 10× dephosphorylation buffer: 100 mM Tris⋅Cl, pH 8.0 (APPENDIX 2A) 25:24:1 (v/v/v) phenol/chloroform/isoamyl alcohol (APPENDIX 2A) 3 M sodium acetate, pH 5.2 (APPENDIX 2A) 100% and 70% (v/v) ethanol 10 mM EDTA (APPENDIX 2A) 10× kinase buffer (see recipe) 0.1 M dithiothreitol (DTT) 150 µCi/µL [γ-32P]ATP (6000 Ci/mmol; NEN Life Sciences) 10,000 U/mL T4 polynucleotide kinase Stop mix (see recipe) Water baths, 37°C and 90° to 100°C
Probing RNA Structure with Chemical Reagents and Enzymes
Additional reagents and equipment for phenol/chloroform/isoamyl alcohol extraction (APPENDIX 2A)
6.1.8 Current Protocols in Nucleic Acid Chemistry
Dephosphorylate RNA (optional) 1. Calculate picomoles of RNA in 0.2 µg and set up the following reaction mix: 0.2 µg RNA of interest 10 µL of 10× dephosphorylation buffer RNase-free water to give 100 µL final volume. 70 U BAP per picomole RNA(dilute with 1x dephosphorylation buffer if necessary) Phosphate treatment is necessary if the RNA sample was generated by in vitro transcription. See also supplier’s technical bulletin for recommended reaction conditions.
2. Incubate 1 hr at 65°C. 3. Extract with 100 µL of 25:24:1 phenol/chloroform/isoamyl alcohol to stop the reaction. 4. Remove the aqueous phase to a new tube. Add 10 µL of 3 M sodium acetate and 300 µL of 100% ethanol. Precipitate on dry ice and then microcentrifuge 30 min at maximum speed (10,000 rpm), 4°C. Remove supernatant, rinse with 70% ethanol, and dry the pellet in a speedvac. 5. Bring 0.2 µg RNA to 18 µL with RNase-free water. Denature RNA sample (optional) 6. Add 2 µL of 10 mM EDTA. Heat 2 min at 90° to 100°C, then quickly transfer to an ice water bath and cool 5 min. Denaturation will increase labeling efficiency of base-paired or recessed 5′ termini; however, some RNA samples may be labeled more efficiently when folded.
7. Briefly microcentrifuge at maximum speed to collect any condensation. Label RNA 8. Combine the following and incubate 1 hr at 37°C. 13 µL denatured RNA (0.13 µg) 2 µL 10× kinase buffer 2 µL 0.1 M DTT 2 µL [γ-32P]ATP 1 µL T4 polynucleotide kinase (10 U). A crude preparation of [γ-32P]ATP is sufficient for this reaction and is less expensive. To label DNA primers, replace RNA with ~1 mg DNA oligomer and proceed through step 11.
9. Add 4 µL stop mix and 100 µL of 100% ethanol (5 vol). 10. Precipitate on dry ice, then microcentrifuge 30 min at maximum speed (10,000 rpm), 4°C. 11. Carefully remove supernatant (very radioactive) and dry the pellet in a speedvac. If the RNA is not going to be used immediately, resuspend the pellet in the appropriate buffer or water to reduce radiation-induced damage and freeze at −80°C. The length of time the RNA can be stored is limited by its activity; the half life of 32P is 14 days.
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.1.9 Current Protocols in Nucleic Acid Chemistry
12. If a heterogeneous population exists, purify the radiolabeled RNA by denaturing gel electrophoresis (APPENDIX 3B). The RNA must have a discrete length to accurately identify nucleotide cleavages/modifications in the subsequent structure probing experiments. SUPPORT PROTOCOL 2
LABELING THE 3′ RNA TERMINUS USING T4 RNA LIGASE AND [32P]pCp Bacteriophage T4 RNA ligase will catalyze phosphodiester formation between a nucleotide 3′ hydroxyl and a nucleotide 5′ phosphate. The RNA of interest requires a 3′ hydroxyl that will be coupled to 5′-[32P]cytidine-3′,5′-bisphosphate ([32P]pCp). [32P]pCp can be purchased or generated using 3′-cytosine-monophosphate, [γ-32P]ATP, and T4 polynucleotide kinase (England et al., 1980; UNIT 6.3). The latter method is recommended because it generates the concentrated form needed at a reasonable cost. Enzymatically synthesized RNAs have a 3′ hydroxyl that can be directly ligated with radiolabeled [32P]pCp. Breakdown of RNA by nucleases, metal or alkaline hydrolysis, and cleavage by ribozymes (excluding RNase P) produces 2′-3′ cyclic phosphates, which subsequently hydrolyze to 3′ phosphates and cannot be directly 3′ end labeled. Dephosphorylation of 3′ phosphates is accomplished with bacterial alkaline phosphatase and accompanying buffer (see Support Protocol 1). Additional Materials (also see Support Protocol 1) 0.5 M HEPES, pH 7.9 at 50 mM 0.1 M MgCl2 0.1 mg/mL bovine serum albumin (BSA) 1 mM ATP Dimethyl sulfoxide (DMSO) 500 µM 5′-[32P]cytidine-3′,5′-bisphosphate ([32P]pCp; UNIT 6. 3 or Amersham) 20 to 25 U/µL T4 RNA ligase Dephosphorylate RNA (optional) 1. If necessary, dephosphorylate RNA as described (see Support Protocol 1, steps 1 to 4), but increase RNA to 2 µg and increase BAP accordingly. 2. Resuspend pellet in 10 µL RNase-free water to make 0.2 µg/µL (∼6 µM for a 100-nucleotide RNA). Label RNA 3. Mix the following (10 µL total volume): 1 µL 10 µM RNA of interest (final 1 µM) 1 µL 0.5 M HEPES, pH 7.9 at 50 mM (50 mM) 2 µL 0.1 M MgCl2 (20 mM) 1 µL 0.1 mg/mL BSA (10 µg/mL) 1 µL 30 mM DTT (3 mM) 1 µL 1 mM ATP (0.1 mM) 1 µL DMSO (10% v/v)
Probing RNA Structure with Chemical Reagents and Enzymes
1 µL 500 µM [32P]pCp (50 µM) 1 µL 20 to 25 U/µL T4 RNA ligase (2 to 2.5 U/ µL) 4. Incubate ≥12 hr at 4°C (typically overnight).
6.1.10 Current Protocols in Nucleic Acid Chemistry
5. Add 90 µL RNase-free water, mix, and extract with 100 µL of 25:24:1 phenol/chloroform/isoamyl alcohol. 6. Add 10 µL of 3 M sodium acetate and 300 µL of 100% ethanol. 7. Precipitate and purify as described (see Support Protocol 1, steps 10 to 12). PREPARING STANDARDS FOR ELECTROPHORESIS OF END-LABELED RNA CLEAVAGE PRODUCTS
SUPPORT PROTOCOL 3
Limited alkaline hydrolysis and digestion with RNase T1 under denaturing conditions are used to prepare two separate sets of electrophoresis standards in order to determine sites of end-labeled RNA cleavage. Alkaline hydrolysis results in a relatively even ladder of truncated RNAs. A denaturing RNase T1 digest cleaves the RNA 3′ of each guanosine. Comparison of the two products with the modified/cleaved end-labeled RNA after electrophoresis identifies guanosines along the alkaline hydrolysis ladder, and therefore the intervening products present in the alkaline hydrolysis reaction can be determined from the RNA sequence. End-labeled RNA cleavage products from structure-probing reactions can thus be identified. Materials RNA of interest, 32P-end-labeled to 50,000 cpm (see Support Protocol 1 or Support Protocol 2) 100% (v/v) ethanol 3 M sodium acetate, pH 5.2 (APPENDIX 2A) Na2CO3/EDTA solution (see recipe) CU buffer (see recipe) RNase T1 (Life Technologies) 2 µg/µL tRNA carrier (see recipe for carrier RNA) CEU buffer (see recipe) FEXS solution (see recipe) 95°C water bath Alkaline Hydrolysis of RNA 1. Ethanol precipitate 50,000 cpm of end-labeled RNA in 1/10 vol of 3M sodium acetate, pH 5.2 and 3 vol of 100% ethanol, microcentrifuge 30 min at maximum speed, 4°C (10,000 rpm), and dry the pellet in a speedvac. 2. Resuspend with 1.2 µL Na2CO3/EDTA solution and mix with a pipet. 3. Incubate between 90 and 120 sec at 95°C, then 1 min on ice. Initially, vary incubation time at 95°C to achieve a relatively even ladder of products following electrophoresis, thereafter use the most successful incubation time.
4. Add 2.8 µL of CU buffer and mix to neutralize. 5. Add 8 µL FEXS solution. Store frozen at −80°C, if necessary. The time the RNA can be stored is dependent on the signal detection and the 32P half life (14 days).
RNase T1 Digest of RNA 6. Add 5 µL of 2 µg/µL carrier tRNA to a second aliquot of 50,000 cpm end-labeled RNA. Ethanol precipitate with 1/10th vol 3 M sodium acetate, and 3 vol 100%
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.1.11 Current Protocols in Nucleic Acid Chemistry
ethanol, microcentrifuge 30 min at maximum speed (10,000 rpm), 4°C, and dry the pellet in a speedvac. 7. Resuspend with 3 µL CEU buffer and mix with a pipet. 8. Incubate 5 min at 50°C. 9. Add 1 µL RNase T1 diluted in CEU buffer and incubate 15 min at 50°C. The appropriate RNase T1 dilution will need to be determined by titration, but 1 to 3 U typically produces good results. Alternatively, other urea-tolerant nucleases, such as RNase A (C and U specific), can be used.
10. Add 8 µL FEXS solution. Store frozen at −80°C, if necessary. BASIC PROTOCOL 3
MODIFICATION OF RNA AT WATSON-CRICK POSITIONS USING DIMETHYL SULFATE Watson-Crick positions of adenosines (N1) and cytidines (N3) not involved in base pairing or tertiary-structure hydrogen bonding are alkylated by DMS. Primer extension (see Support Protocol 4) of modified RNA and comparison with dideoxynucleotide sequencing reactions identifies the modified bases. Used together, dimethyl sulfate (for A and C), kethoxal (for G; see Alternate Protocol 3), and CMCT (for U; see Alternate Protocol 4) provide a complete analysis of the base-pairing status (i.e., secondary structure) of the RNA of interest. See Strategic Planning for discussions of RNA sample source and folding. Materials 0.5 M HEPES, pH 7.8 1 M KCl (APPENDIX 2A) 0.1 M MgCl2 1 µg/µl RNA of interest RNase-free water (see recipe) 10.56 M dimethyl sulfate (DMS; 99+%) 100% and 70% (v/v) ethanol 3 M sodium acetate, pH 5.2 (APPENDIX 2A) Additional reagents and equipment for primer extension (see Support Protocol 4) CAUTION: DMS is highly toxic and a suspected carcinogen; appropriate precautions should be taken for handling, storage, and disposal. 1. Prepare four reaction tubes, each containing: 1 µL 0.5 M HEPES, pH 7.8 (final 50 mM) 1 µL 1 M KCl (100 mM) 1 µL 0.1 M MgCl2 (10 mM) 2 µL 1 µg/µL RNA of interest (0.2 µg/µL)
Probing RNA Structure with Chemical Reagents and Enzymes
4 µL RNase-free water 2. Just prior to use, dilute DMS in four 2-fold serial dilutions in 100% ethanol to give the appropriate amounts in a volume of 1 µL.
6.1.12 Current Protocols in Nucleic Acid Chemistry
DMS must initially be titrated to attain the desired RNA modification typically ~10%. The following dilutions are provided as examples only. Dilute 1 mL of 10.56 M DMS stock in 16.6 mL of 100% ethanol to give 600 mM. Further dilute 2-, 4-, and 8-fold with 100% ethanol to make 300, 150, and 75 mM dilutions, respectively. The final reaction concentrations will be 60, 30, 15, and 7.5 mM.
3. Add 1 µL of each DMS dilution to a reaction mix, mix well, and incubate 20 min at room temperature. 4. Add 1 µL of 3 M sodium acetate and 30 µL of 100% ethanol and precipitate on dry ice. Microcentrifuge 30 min at maximum speed (10,000 rpm), 4°C. Remove supernatant, rinse with 70% ethanol, and dry the pellet in a speedvac. 5. Resuspend pellet in 40 µL RNase-free water and proceed with primer extension (see Support Protocol 4) using 4 µL (0.2 µg) per reaction. MODIFICATION OF RNA AT WATSON-CRICK POSITIONS USING KETHOXAL
ALTERNATE PROTOCOL 3
Kethoxal alkylates the N1 and 2-NH2 Watson-Crick positions of guanosines not involved in base pairing or tertiary-structure hydrogen bonding. The procedure is the same as for modification with DMS (see Basic Protocol 3), except that step 2 is replaced with the following step. Used together, dimethyl sulfate (A & C; Basic Protocol 3), kethoxal (G), and CMCT (U; Alternate Protocol 4) provide a complete analysis of the base-pairing status (i.e., secondary structure) of the RNA of interest. See Strategic Planning for discussion of RNA sample source and folding. Additional Materials (also see Basic Protocol 3) 4.27 M kethoxal stock (ICN or Research Organics) 2b. Just prior to use, dilute 4.27 M kethoxal in four 2-fold serial dilutions in RNase-free water to give the appropriate amounts in a volume of 1 µL. Kethoxal must be titrated to attain the desired RNA modification. The following dilutions are provided as examples only. Dilute 1 mL of 4.27 M kethoxal stock in 20.35 mL water to give 200 mM. Further dilute 2-, 4-, and 8-fold with water to make 100, 50, and 25 mM dilutions, respectively. Final reaction concentrations will be 20, 10, 5, and 2.5 mM.
MODIFICATION OF RNA AT WATSON-CRICK POSITIONS USING CMCT 1-Cyclohexyl-3-(2-morpholinoethyl)carbodiimide metho-p-toluenesulfonate (CMCT) alkylates the N3 Watson-Crick position of uridines not involved in base pairing or tertiary-structure hydrogen bonding. The approach is essentially as described for modification with DMS (see Basic Protocol 3), except that the reactions are performed as outlined below. Used together, dimethyl sulfate (A & C; Basic Protocol 3), kethoxal (G; Alternate Protocol 3), and CMCT (U) provide a complete analysis of the base-pairing status (i.e., secondary structure) of the RNA of interest. See Strategic Planning for discussion of RNA sample source and folding. Additional Materials (also see Basic Protocol 3) 0.5 M potassium borate, pH 8.0 0.5 M 1-cyclohexyl-3-(2-morpholinoethyl)carbodiimide metho-p-toluenesulfonate (CMCT) in RNase-free H2O (stable for several weeks at –20°C; Aldrich) 37°C water bath
ALTERNATE PROTOCOL 4
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.1.13 Current Protocols in Nucleic Acid Chemistry
1. Prepare three reaction tubes, each containing: 1 µL 0.5 M potassium borate, pH 8.0 (final 50 mM) 1 µL 1 M KCl (100 mM) 1 µL 0.1 M MgCl2 (10 mM) 2 µL 1 µg/µL RNA of interest (0.2 µg/µL) 2. Just prior to use, heat 0.5 M CMCT at 37°C and vortex to ensure the reagent is completely solubilized. Immediately dilute 0.5 M CMCT solution to 200 mM and 50 mM in RNase-free water. CMCT must be titrated to attain the desired RNA modification. The above dilutions can be used for the initial titration. Final reaction concentrations using the stock solution and the two dilutions will be 250, 100, and 25 mM, respectively.
3. Add 5 µL of each CMCT dilution to a reaction mix, mix well, and incubate 20 min at room temperature. 4. Precipitate and proceed to primer extension as described (see Basic Protocol 3, steps 4 and 5). SUPPORT PROTOCOL 4
DETECTION OF RNA CLEAVAGE OR MODIFICATION BY PRIMER EXTENSION Primer extension of a radiolabeled DNA oligomer will synthesize a complementary DNA strand to the RNA of interest. Reverse transcriptase will stop upon encountering alkylated Watson-Crick determinants, resulting in a product one base short of the modification. The enzyme will also run off RNA cleavage products, producing a DNA corresponding to the site of cleavage. Strong secondary structure, such as hairpin loops within the RNA, can cause the enzyme to pause, creating a prematurely terminated product. It is therefore essential to perform the reaction on unmodified/uncleaved RNA as a control. This enables pause sites from secondary structure to be subtracted from the modification/cleavage analysis. Because carrier RNA can affect the reverse transcription reaction, control reactions should include the carrier RNA as well. The reaction requires a DNA oligomer complementary to the RNA of interest, 3′ of the region to be examined. It might be necessary to use several DNA oligomers to examine an RNA several hundred nucleotides long. The extension products are resolved on standard DNA sequencing gels, allowing ∼200 nucleotides to be examined per DNA primer. Location of primer-extension terminations is determined by comparison to standard dideoxynucleotide sequencing reactions using the same primer and a template of the same sequence as the RNA being examined. The primer-extension reaction requires a 32P-labeled DNA oligomer that is generated using bacteriophage T4 polynucleotide kinase. Reaction conditions for labeling the primer are identical to those used for end labeling the 5′ RNA terminus (see Support Protocol 1), but substituting the DNA oligomer for the RNA.
Probing RNA Structure with Chemical Reagents and Enzymes
Materials [5′-32P]DNA oligomer (see Support Protocol 1) complementary to 3′ end of the sample RNA Sample RNA, cleaved or modified (see Basic Protocols 1 to 3 and Alternate Protocols 1 to 4) RNase-free water (see recipe) Sample RNA, uncleaved and unmodified 25 mM 4dNTP mix (see dNTPs in APPENDIX 2A) 0.1 M dithiothreitol (DTT)
6.1.14 Current Protocols in Nucleic Acid Chemistry
5× first strand buffer (see recipe) 200 U/µL Superscript II RNase H(−) Moloney murine leukemia virus reverse transcriptase (MMLV RT; Life Technologies; see Troubleshooting for alternative RTs) 3 M sodium acetate, pH 5.2 (APPENDIX 2A) 100% and 70% (v/v) ethanol FENXB solution (see recipe) Dideoxynucleotide sequencing reactions (e.g., CPMB UNIT 7.4A) of cDNA for sample RNA Sequencing gel: 6% (w/v) acrylamide (20:1 mono-/bis-), 8 M urea, 1× TBE buffer (APPENDIX 2A for TBE buffer) Water bath or heating block, 42° and 70°C Additional reagents and equipment for running a sequencing gel (APPENDIX 3B) Anneal primer 1. Mix the following: 100,000 cpm [5′-32P]DNA oligomer 0.2 µg sample RNA, cleaved or modified RNase-free water to 18.5 µL. It may be necessary to titrate the [32P]DNA oligomer and RNA concentrations to optimize the extension reaction conditions.
2. Set up a similar control reaction using unmodified, uncleaved RNA. This control should be performed to identify structure-induced terminations and to ensure that the majority of the DNA oligomer is being fully extended through the region of interest.
3. Incubate 10 min at 70°C, followed by 30 min at 42°C. 4. Briefly microcentrifuge down any condensation that forms. Extend DNA 5. To the primer/RNA mix add: 3 µL 25 mM 4dNTP mix 2 µL 0.1 M DTT 6 µL 5× first strand buffer. Total volume is now 29.5 µL.
6. Add 0.5 µL (100 U) Superscript II and mix. 7. Incubate 45 min at 42°C. 8. Add 3 µL of 3 M sodium acetate, and 90 µL of 100% ethanol. Precipitate on dry ice and microcentrifuge 30 min at maximum speed (10,000 rpm), 4°C. 9. Remove supernatant, rinse pellet with 70% ethanol, and dry the pellet in a speedvac. 10. Resuspend in 6 µL FENXB solution for resolution on a 6% acrylamide sequencing gel (APPENDIX 3B). Use dideoxynucleotide sequencing reactions as markers (e.g., CPMB Unit 7.4A). Typically a cDNA template, rather than an RNA template, is used for the sequencing reactions, allowing them to be performed with a DNA-dependent DNA polymerase (e.g., Sequenase; United States Biochemical).
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.1.15 Current Protocols in Nucleic Acid Chemistry
REAGENTS AND SOLUTIONS Use deionized water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Aniline acetate buffer, 1 M 10 µL of 99.5% aniline 93 µL H2O 6 µL glacial acetic acid Make fresh CAUTION: Aniline is highly toxic.
Carrier RNA, 1 mg/mL or 2 mg/mL Carrier RNA can be unlabeled RNA of interest or total yeast tRNA. Extract yeast tRNA with 25:24:1 (v/v/v) phenol/chloroform/isoamyl alcohol (APPENDIX 2A) and ethanol precipitate to remove any protein contaminants. Store at –20°C. Citrate/EDTA/urea (CEU) buffer For 100 mL: 10.25 mL 0.1 M citric acid 14.75 mL 0.1 M sodium citrate (final 25 mM citrate) 0.2 mL 0.5 M EDTA (APPENDIX 2A; final 1 mM) ∼30 mL H2O 42.04 g urea (final 7 M) Adjust to pH 4.7 to 5.0 with HCl Stable indefinitely at –20°C Citrate/urea (CU) buffer For 100 mL: 10.25 mL 0.1 M citric acid 14.75 mL 0.1 M sodium citrate (final 25 mM citrate) ∼30 mL of H2O 60.06 g urea (10 M final) Adjust to pH 4.7 to 5.0 with HCl Stable indefinitely at –20°C Ethylnitrosourea (ENU)/ethanol solution Add N-nitroso-N-ethylurea (ENU) to 100 µL of 100% ethanol until solution is saturated. Centrifuge to pellet insoluble reagent. Make fresh. CAUTION: ENU is highly toxic and a suspected carcinogen. follow appropriate precautions for handling, storage, and disposal.
FENXB solution 95% formamide, redistilled 20 mM EDTA (APPENDIX 2A) 2 mM NaOH 0.05% xylene cyanol 0.05% bromphenol blue Stable indefinitely at –5 to –20°C
Probing RNA Structure with Chemical Reagents and Enzymes
FEXS solution 95% formamide, redistilled 10 mM EDTA (APPENDIX 2A) 0.05% xylene cyanol 0.1% SDS Stable indefinitely at –5 to –20°C
6.1.16 Current Protocols in Nucleic Acid Chemistry
First strand buffer, 5× 250 mM Tris⋅Cl , pH 8.3 (APPENDIX 2A) 375 mM KCl 15 mM MgCl2 Stable indefinitely at –20°C Kinase buffer, 10× 700 mM Tris⋅Cl, pH 7.6 (APPENDIX 2A) 100 mM MgCl2 50 mM dithiothreitol (DTT) Stable indefinitely at –20°C Na2CO3/EDTA solution For 1 mL: 50 µL 1 M Na2CO3, pH 11.7 (final 50 mM) 2 µL 0.5 M EDTA (APPENDIX 2A; final 1 mM) 948 µL H2O Stable indefinitely at –20°C Stop mix 20 mM EDTA (APPENDIX 2A) 0.3 M NaCl 1 µg/µL glycogen Stable indefinitely at –20°C Water, RNase free Test water source for contaminating nuclease activity by incubating labeled RNA in water and buffers for intended reaction time. Ascertain integrity of sample by electrophoresis. Often deionized and glass-distilled water will be free of nucleases. However, if water and/or buffer reagents contain nuclease activity, treat with DEPC to inactivate (APPENDIX 2A). Autoclaved solutions are sometimes allowed to stand at room temperature, with loose caps, for several days to remove all DEPC. Trace DEPC can inhibit reverse transcriptase and other enzymes.
COMMENTARY Background Information Sensitivity to chemical and enzymatic reagents can often be used to test hypotheses regarding solution exposure and singlestranded versus double-stranded character along the folded RNA chain; however, the data generated in these experiments is most effective when used in reference to a secondary structure model for the RNA generated by other approaches. Computer-based energy-minimization algorithms have proven extremely useful in predicting simple RNA structures or in providing multiple local folding possibilities in larger RNAs (Zuker, 1989; Gautheret et al., 1990; Major et al., 1991; Malhotra et al., 1993; Walter et al., 1994; Gorodkin et al., 1997). These folding algorithms suffer from several
severe limitations in predicting longer RNAs, however, tending to miss long-distance interactions and lacking sufficient predictions of nonstandard interactions among nucleotides. In some instances, the physiologically relevant RNAs will also contain nucleoside modifications that are not necessarily known or taken into account. Perhaps most importantly, the functional form of the molecules might not be the most stable folded form under the conditions used to fold the RNA in vitro. It is possible that manipulation of the folding conditions might identify methods that drive the majority of a purified RNA into an appropriate structure, assuming that there is some method to determine what constitutes an appropriate structure.
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.1.17 Current Protocols in Nucleic Acid Chemistry
Manipulation of folding conditions for RNAs synthesized in vitro will be discussed below. Structural analysis of RNAs and ribonucleoproteins (RNPs) can be used to determine secondary structure, facilitate modeling of tertiary structure, and identify RNA-protein contacts. The source of the material to be probed will often dictate which labeling and probing techniques are necessary. Primer extension is the preferred method for analysis of large or cellularly derived RNAs. The native RNA or RNP complex can be assayed without manipulations that may disrupt the integrity of the sample. Postmodification or cleavage products are assayed by extension of labeled DNA oligomers with reverse transcriptase. End labeling techniques can also sometimes be successful. However, in vivo–derived RNAs often possess 5′ methyl cap structures preventing labeling at that terminus, and one or both termini in the native RNA can be recessed or obscured by protein in RNP structures. Chemical or enzymatic synthesis of RNA permits incorporation of labeling groups during synthesis, or if necessary, labeling sites can be engineered in the synthetic RNAs. Studies involving novel sequences, subregions of larger RNAs, or RNP reconstitution experiments sometimes require the use of synthetic RNA. Chemical and enzymatic synthesis of RNA also provides an opportunity to incorporate nucleosides modified with fluorescent, antigenic, or crosslinking groups during synthesis.
Critical Parameters
Probing RNA Structure with Chemical Reagents and Enzymes
The first principle in RNA structure analysis is to begin with an RNA properly folded into its physiologically relevant structure. Synthetic RNA may or may not be in its properly folded state. Gentle purification techniques should retain the in vivo conformation of cellularly derived RNAs and RNPs. However, folding of many RNAs in vivo is not readily reproduced in vitro, in many instances because the RNAs require association of proteins to assume physiologically appropriate structures. In these cases it might be necessary to either isolate the RNAs from cells or to associate the appropriate proteins in vitro in a manner that can be demonstrated to cause the RNA to assume a functional structure. The best insurance that the RNA or RNP of interest is correctly folded is to test its functional or enzymatic activity. Other folding assays will be necessary if no functional assay exists. The number of folding isoforms of an RNA sample can sometimes be determined using
native polyacrylamide gel electrophoresis. RNAs with extensive self-complementarity often form multimers, which can be detected on the native gel. Heat denaturation can be used to melt present structures. Cooling rate, RNA concentration, and the presence of mono- and divalent cations all affect the subsequent folding isoform populations. In general, a faster cooling rate, the absence of cations, and low RNA concentrations favor intramolecular folding. Likewise, slow cooling in high salt with high RNA concentrations favors intermolecular folding and the formation of multimers. It should be noted that the RNA might need to adopt more than one structure to carry out its function(s) in vivo. Driving all of the RNA sample (or protein-RNA complex) into a single form might reflect only one aspect of the functional RNA structure. Once the RNA is properly folded, the structure-probing experiments can begin. There are four factors that are key to experiment reproducibility. The first is a consistent reaction mix. Make a batch of all ingredients minus the reagent, mix, then aliquot to the necessary number of tubes. Always make enough for the desired number of reactions plus one. This will ensure that the amount of each component, especially radiolabeled RNA, is identical between reactions. The second factor is reaction time. Many of the probing reactions are short (e.g., 10 min) to minimize the effect of any contaminating nuclease activity. The number of reactions attempted at once should not compromise strict adherence to the reaction time. Longer reaction times may be used effectively, but short times have been found to reduce secondary or nonspecific activities. The third factor is the use of unlabeled carrier RNA. Carrier RNA provides a constant substrate concentration for the probing reagents. The concentration of radiolabeled RNA is much less than the carrier and is negligible in most reactions. Therefore, any changes in labeling efficiency between RNA preparations will have little effect. Lastly, it is necessary to titrate the amount of reagent for each particular RNA of interest so that ∼10% of the RNA is cleaved. The likelihood of multiple cleavages per molecule is thereby reduced. Typically, the concentration of RNA is held constant and the amount of reagent is titrated until the desired extent of modification is achieved. If the RNA sample of interest is extremely limited, reagents are titrated using a different RNA with the identical unlabeled carrier RNA and buffer conditions.
6.1.18 Current Protocols in Nucleic Acid Chemistry
The presence of excess carrier RNA facilitates reproducibility of cleavage conditions.
Troubleshooting Primer-extension analysis of probing reactions can encounter several pitfalls. For example, breakdown in the sample results in termination products not produced by the probing reagents. Extension of unmodified, uncleaved RNA should identify these sites and allow for their subtraction from the probing data. However, extensive degradation or strong secondary structure–induced terminations may obscure cleavages or modifications at several nucleotides of interest. Synthetic RNA sample integrity can be improved by changing the original purification strategy or by further purification to ensure that a discrete population exists. RNA samples in crude cellular fractions should be prepared as rapidly as possible at 0° to 4°C and stored frozen (−80°C) when not in use to minimize degradation by endogenous nucleases. To alleviate secondary structure–induced terminations, the primer-extension reaction conditions and/or the reverse transcriptase can be changed. Superscript II has demonstrated an ability to proceed quite well through secondary structure, but other reverse transcriptases, such as AMV and Retrotherm, may function better on a particular RNA. The heat-stable Retrotherm can also be used at elevated temperatures to minimize RNA secondary structure. Achieving an interpretable amount of modification or cleavage is another problem often encountered. The desired modification or cleavage goal is 10% of the total RNA, in the realm of single-hit kinetics. This rule minimizes the presence of products resulting from multiple modifications or cleavages. Typically, the concentration of reagent is adjusted until the 10% goal is reached for a batch of RNA, carrier RNA, and reagent. Single cleavages or modifications per RNA molecule usually allow for detection of all available targets within the region of interest. However, a hyper-reactive site may exist that prevents further primer extension or observation of end-labeled RNA products larger than the hyper-reactive nucleotide. In this scenario, a new primer complementary to the region 5′ of the RNA modification will be needed, or in the case of end-labeled RNA, labeling the opposite end will allow analysis of the remaining RNA region of interest. All of the probing reactions are purposefully designed to have nearly identical buffer and salt conditions. RNA structure can change consid-
erably based on the amount and nature of counterions and pH. Consistency in reaction conditions between different probing reagents helps to ensure the same structure is being examined. The buffers and pH are changed only when they may interfere with the chemical nature of the reagent. It should be noted that the maximal activities of several reagents and most of the nucleases occur under different conditions than those listed, but all have been used successfully as described. The standard salt conditions can be altered as desired but consistency is important. Also, formation of RNA tertiary structures is often very dependent on the amount and type of counter-ions present. Uncharacteristic modifications or cleavages from probing reagents are sometimes observed. Interpretation of atypical activities should be regarded with appropriate caution. For example, reagents used outside their optimal reaction conditions or the presence of contaminating activities could be responsible. It is also possible to observe inconsistencies between reagents with the same targets. Thus, using two reagents that should cleave or modify an unpaired nucleotide might give the result that the nucleotide is sensitive to one but not the other. In all probability, the reagents are functioning properly, but subtle differences in substrate binding and recognition can favor one over the other.
Anticipated Results There are several cautionary notes that should be considered when interpreting RNA footprinting data. First, it should always be remembered that both purified RNAs and RNA-protein complexes can exist in multiple isoforms, whether folded and bound in vitro or isolated intact from cells. As far as possible, the heterogeneity of RNA sample of interest should be investigated and taken into account when interpreting sensitivity. Second, sensitivity to cleavage or modification reagents is far more useful in showing solution exposure than “protection” from reagents. Inaccessibility of even the same region of a base to an individual enzyme or chemical can vary from reagent to reagent, and can have a number of causes including involvement in hydrogen bonding, folding into the interior of the RNA where it is less exposed to the solvent, inappropriate chemical states, and direct blockage by interacting proteins. Thus, if protection is observed in a particular region when a protein is added, the protection might result from either direct coverage by the protein or indirect effects of
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.1.19 Current Protocols in Nucleic Acid Chemistry
the bound protein acting at a distance. It should be reemphasized that even detailed footprinting results are often consistent with multiple hypothetical structures, and for any complex RNA it is likely that additional, independent types of evidence will be needed to sort out the possibilities. Results of structure-probing experiments are useful to qualitatively model RNA structure. The more reagents employed, the more descriptive the analysis. Suspected anomalous modifications or cleavages by a single reagent can therefore be subtracted. Modification/cleavage products can be quantitated by phosphorimager analysis and analyzed using a computer (e.g., ImageQuant from Molecular Dynamics; visual inspection of autoradiograms is fine for qualitative analysis). The accessibility of an RNA target is directly proportional to the modification/cleavage product band intensity. End-labeled RNA or labeled primer-extension products have one label per product molecule. However, it is not simple to interpret the reason that one target is preferred over another. Targets on the surface of a molecule are likely to be more accessible than those buried within the interior of a tertiary structure, although local steric effects may have as much impact on reagent reactivity as global structure. Data obtained using multiple reagents will better uncover the true solution structure of a target, and the trend of several reagents should be regarded as the rule, rather than a single hit by one reagent. One should also keep in mind that solution structure is dynamic and that RNA molecular motion is dependent on temperature and ionic environment.
Time Considerations
Probing RNA Structure with Chemical Reagents and Enzymes
The design of structure-probing experiments is dictated by the stability of the RNA sample of interest and the half-life of radiolabeled material. Cellularly derived RNAs or RNPs often contain contaminating nuclease activities, and these can be considerable depending on the purification scheme employed. It is therefore imperative to preserve samples frozen at −80°C and to use them immediately upon thawing. Synthetic RNAs are often free of contaminating nucleases, depending on the handling and manipulations the sample has received. The structure-probing reactions are usually short (e.g., 10 min) to reduce the effects of any contaminating activities. Control reactions should identify extraneous cleavages, provided that the majority of the sample is not degraded in the reaction interval. A time course
incubation of the RNA of interest in the desired reaction buffer may prove beneficial in identifying sample integrity versus incubation time. Longer reaction times, allowing for more samples, might then be used. Modified/cleaved RNA intended for primerextension analysis can be stored frozen at −80°C while the radiolabeled DNA primer is made. End-labeled RNA should be resolved by electrophoresis promptly after modification/cleavage reactions are complete. Note that only 10% of the RNA sample should be modified/cleaved; therefore, the detectable radiolabel signal from the modified/cleaved products will only be 10% of the sample’s specific activity. Experiments should proceed readily to maximize signal from a batch of radiolabeled RNA. Once all reaction components are in hand, setting up the probing reactions, incubating, and terminating the reactions for a single reagent should take ∼1 to 3 hr. Primer extension typically takes 2 to 3 hr, and gel electrophoresis on standard sequencing (6%) polyacrylamide gels requires 2 to 3 hr.
Literature Cited England, T., Bruce, A., and Uhlenbeck, O. 1980. Specific labeling of 3′ termini of RNA with T4 RNA ligase. Methods Enzymol. 65:65-74. Gautheret, D., Major, F., and Cedergren, R. 1990. Computer modeling and display of RNA secondary and tertiary structures. Methods Enzymol. 183:318-330. Gorodkin, J., Heyer, L., and Stormo, G. 1997. Finding the most significant common sequence and structure motifs in a set of RNA sequences. Nucl. Acids Res. 25:3724-3732. Knapp, G. 1989. Enzymatic approaches to probing of RNA secondary and tertiary structure. Methods Enzymol. 180:192-212. Krol, A. and Carbon, P. 1989. A guide for probing native small nuclear RNA and ribonucleoprotein structures. Methods Enzymol. 180:212-227. Major, F., Turcotte, M., Gautheret, D., Lapalme, G., Fillion, E., and Cedergren, R. 1991. The combination of symbolic and numerical computation for three-dimensional modeling of RNA. Science 253:1255-1260. Malhotra, A., Gabb, H., and Harvey, S. 1993. Modeling large nucleic acids. Curr. Opin. Struct. Biol. 3:241-246. Milligan, J.F. and Uhlenbeck, O.C. 1989. Synthesis of small RNAs using T7 RNA polymerase. Methods Enzymol. 180:51-62. Peattie, D.A. and Gilbert, W. 1980. Chemical probes for higher-order structure in RNA. Proc. Natl. Acad. Sci. U.S.A. 77:4679-4682.
6.1.20 Current Protocols in Nucleic Acid Chemistry
Walter, A.E., Turner, D.H., Kim, J., Lyttle, M.H., Muller, P., Matthews, D.H., and Zuker, M. 1994. Coaxial stacking of helixes enhances binding of oligoribonucleotides and improves predictions of RNA folding. Proc. Natl. Acad. Sci. U.S.A. 91:9218-9222.
Contributed by William A. Ziehler and David R. Engelke University of Michigan Ann Arbor, Michigan
Zuker, M. 1989. On finding all suboptimal foldings of an RNA molecule. Science 244:48-52.
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.1.21 Current Protocols in Nucleic Acid Chemistry
Probing Nucleic Acid Structure with Shape-Selective Rhodium and Ruthenium Complexes
UNIT 6.2
This unit describes the use of transition metal complexes, specifically rhodium and ruthenium compounds, as photochemical probes of the structure of DNA and RNA. Such complexes have proven to provide a versatile platform for the design of chemical reagents to probe nucleic acid structure. The central transition metal ion provides both a rigid, substitutionally inert framework and an octahedral geometry for ligand coordination. The metal complexes can be constructed to define shapes, symmetries, and functionalities that complement those of the nucleic acid target. In addition, the rich photochemistry of rhodium and ruthenium allows easy identification of sites of complex binding by lightinduced nucleic acid cleavage. The modular construction of such complexes, allowing use of a variety of commercially available or custom-synthesized ligands, makes it possible to generate many different probes to examine various subtle and not-so-subtle characteristics of nucleic acids. In the procedures that are described, end-labeled DNA or RNA is incubated with a single transition metal reagent and then irradiated with visible or ultraviolet light to promote cleavage of the nucleic acid polymer strand. The resulting cleavage products are then observed by polyacrylamide gel electrophoresis (PAGE). These common procedures are used with different metal complexes to probe various characteristics of nucleic acid systems. For rhodium compounds, strand cleavage is affected by direct metal complex photochemistry; for ruthenium compounds, cleavage is induced by singlet oxygen-induced damage followed by treatment with piperidine. Metal complexes of varied structure are utilized to explore different characteristics of the nucleic acids. Depending on the binding mode and specificity of the complexes, it is possible to explore characteristics such as groove width and depth (see Basic Protocol 1), the presence of unusual structures like cruciforms or Z DNA (see Basic Protocol 2), mismatches or abasic sites (see Basic Protocol 3), and tertiary folding of nucleic acids (see Basic Protocol 5). In addition, such complexes can be used as photofootprinting agents to study other molecules binding to nucleic acids (see Basic Protocol 4) and as singlet oxygen sensitizers for specific reaction with guanine bases in DNA or RNA (see Basic Protocol 6). Inorganic syntheses of the required metal complexes are included for each procedure (see Support Protocols 1 to 6). General guidelines for preparing labeled nucleic acids, for photolysis, and for mapping cleavage sites are also presented (see Support Protocols 7 to 9). CAUTION: The procedures described in this unit utilize strong acids, flammable liquids, glassware under vacuum, and cryogens. All reagents should be used with care, using appropriate personal and laboratory safety equipment, and in well-ventilated fume hoods. The syntheses of the transition metal reagents involve polycyclic aromatic compounds, most of which are either suspected or known carcinogens. All precautions recommended by suppliers should be taken in order to prevent undue exposure to dusts or solutions. The metal complexes made in these protocols bind to nucleic acids with high affinity and promote strand cleavage upon exposure to light. As a result, they should be handled with care and contact with them should be avoided. The light sources used in DNA photocleavage experiments produce high-intensity ultraviolet light. Focused light beams can cause skin burns and severe eye damage. The radioactivity used in nucleic acid labeling procedures must be used in compliance with all appropriate regulations and guidelines. When performing electrophoresis portions of the protocols, it must be remembered that unpolymerized acrylamide is a potent, cumulative neurotoxin and all contact with it must
Chemical and Enzymatic Probes for Nucleic Acid Structure
Contributed by Brian A. Jackson and Jacqueline K. Barton
6.2.1
Current Protocols in Nucleic Acid Chemistry (2000) 6.2.1-6.2.39 Copyright © 2000 by John Wiley & Sons, Inc.
be avoided. In all cases, the waste products produced by these syntheses and biochemical experiments should be disposed of according to good laboratory practices. BASIC PROTOCOL 1
MAPPING DNA MAJOR AND MINOR GROOVE CHARACTERISTICS Two metal complexes of differing structural characteristics, bis(1,10-phenanthroline)(9,10-phenanthrenequinone diimine)rhodium(III), [Rh(phen)2(phi)]3+, and tris(3,4,7,8-tetramethyl-1,10-phenanthroline)ruthenium(II), [Ru(TMP)3]2+, can be used to examine variations in structure and local conformation in stretches of putatively B-form DNA. For reference, the structures of the complexes are included as Figure 6.2.1. The two coordination compounds bind to DNA by different modes and, as a result, recognize different structural characteristics of the nucleic acid. [Rh(phen)2(phi)]3+ binds in the major groove by intercalation of its phi ligand, and thus recognizes sites on the DNA where the geometry of the double helix can accommodate its bulky phenanthroline ligands. Upon irradiation with UV light, it directly cleaves the DNA at its site of binding. Conversely, [Ru(TMP)3]2+ binds to the surface of the DNA minor groove and shows a preference for A-form helices. The ruthenium complex cleaves DNA by sensitizing singlet oxygen upon irradiation. When taken together, DNA cleavage patterns generated by both of these complexes serve as a straightforward method to map out the geometrical characteristics of a long DNA sequence. For the mapping of an isolated DNA sequence, end-labeled DNA is incubated with each metal complex and photolysed to promote strand cleavage. Because [Rh(phen)2(phi)]3+ directly cleaves the DNA, no subsequent treatment of those samples is required before electrophoresis. For [Ru(TMP)3]2+, samples must be subsequently treated with hot piperidine to convert the singlet oxygen damage to strand breaks. In addition, due to the diffusable nature of the singlet oxygen, [Ru(TMP)3]2+ cleavage must also be compared with cleavage by [Ru(phen)3]2+, which binds nonspecifically to DNA. The difference between the cleavage patterns of the two complexes reveals the sequence-specific recognition effect. Specific details for these rhodium and ruthenium photolysis reactions are given in this protocol. A more general yet thorough discussion of photolysis is given in Support Protocol 8, which should be consulted before carrying out this procedure. Materials Labeled DNA solution (see Support Protocol 7): for cleavage with [Rh(phen)2(phi)]3+, typically 100 µM base pairs in 2× buffer for cleavage with [Ru(TMP)3]2+ or [Ru(TMP)3]2+, typically 200 µM base pairs in 2× Tris/NaCl/imidazole buffer (see recipe)
CH3 3+
H N N H
CH3
N
Rh N
N
N
Ru
N
H3C
CH3
N
H3C
N
2+
H3C
N CH3
N
CH3
N
CH3
CH3
H3C CH3
Probing Nucleic Acid Structure
Figure 6.2.1
[Rh(phen)2(phi)]3+ and [Ru(TMP)3]2+.
6.2.2 Current Protocols in Nucleic Acid Chemistry
Metal complex solution (see recipe for preparation of stock solutions): rac- or ∆-[Rh(phen)2(phi)]3+ (typically 10 µM; see Support Protocol 1 for synthesis; see Support Protocol 2 for separation of enantiomers) rac-[Ru(TMP)3]2+ (typically 60 µM; see Support Protocol 3) rac-[Ru(phen)3]2+ (typically 60 µM; see Support Protocol 3) 1 M (10% v/v) piperidine solution in deionized water Dry ice Heating block or bath set to 90°C Speedvac (Savant) or lyophilizer Additional reagents and equipment for photolysing metal complexes (see Support Protocol 8) and mapping cleavage sites (see Support Protocol 9) Cleave DNA with [Rh(phen)2(phi)]3+ 1. Photolyse DNA/metal complexes as described (see Support Protocol 8, steps 1 to 4), mixing equal volumes of labeled DNA solution and rac- or ∆-[Rh(phen)2(phi)]3+ metal complex solution for a total volume of 10 to 50 µL. Include dark and light controls for each DNA of interest. Irradiate either 1 to 5 min at 313 nm or 5 to 20 min at 365 nm. It is best to separate the enantiomers and use only the ∆ enantiomer because it has better DNA-binding characteristics. If both are used, separate dark controls are required to ensure that one of the two enantiomers has not become contaminated in any way. The choice of irradiation paradigm is dependent on the light source available. The small differences in the results obtained at these two wavelengths are not relevant for these structure determinations.
2. Dry samples in a Speedvac evaporator or precipitate samples as described (see Support Protocol 8, steps 5 to 7). 3. Count samples and add loading buffer (see Support Protocol 8, step 8). 4. Map cleavage sites as described (see Support Protocol 9). Identify the specific bases of cleavage by comparing to Maxam-Gilbert sequencing reactions as described. Sites of [Rh(phen)2(phi)]3+ binding are indicated by cleavage bands in the experimental lanes. Although cleavage band intensity is related to the affinity of the complex for individual DNA sites, possible differences in photocleavage efficiency from site to site make direct comparisons difficult.
Cleave DNA with [Ru(TMP)3]2+ and [Ru(phen)3]2+ 5. Photolyse parallel samples using both rac-[Ru(TMP)3]2+ and rac-[Ru(phen)3]2+ complexes as described (see Support Protocol 8, steps 1 to 4), mixing equal volumes of metal complex solution and labeled DNA solution for a total volume of 10 to 50 µL. Include dark and light controls for each DNA and for each metal complex. Irradiate 20 to 40 min at 442 nm. Imidazole is used in DNA solutions for these reactions to help solubilize the ruthenium probe, which is extremely hydrophobic. The metal complex stock solutions prepared for this experiment can be close to saturation and may require a small amount of a co-solvent, such as ethanol, to keep the complexes in solution. Complex precipitation can be a source of experimental error and irreproducibility.
6. Dry samples in a Speedvac evaporator or precipitate samples as described (see Support Protocol 8, steps 5 to 7).
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.3 Current Protocols in Nucleic Acid Chemistry
7. Add 100 µL of piperidine solution to each of the dried samples and heat 30 min at 90°C. Treatment with hot piperidine causes strand scission at the sites of singlet oxygen damage on the DNA bases. CAUTION: Piperidine is a highly toxic liquid of high vapor pressure and is readily absorbed through the skin. Contact should be avoided and, if possible, manipulations should be made in a fume hood.
8. Freeze samples on dry ice and lyophilize off the piperidine solution. 9. Count samples and add loading buffer (see Support Protocol 8, step 8). 10. Map cleavage sites as described (see Support Protocol 9). 11. Subtract histograms of [Ru(TMP)3]2+ and [Ru(phen)3]2+ cleavage patterns to remove the guanine bias associated with singlet oxygen reactivity. Identify the specific bases of cleavage by comparison to Maxam-Gilbert sequencing reactions as described in Support Protocol 9. Cleavage by [Ru(phen)3]2+ is the same at all guanine residues due to its nonspecific binding behavior. Cleavage by the methylated complex is nonuniform as a result of preferential binding at individual sites. Cleavage by [Ru(TMP)3]2+is also observed at nonguanine residues near its binding sites. The difference between the cleavage patterns shows characteristic “bell-shaped” cleavage distributions as a result of high local concentrations of the diffusing singlet oxygen near the sites of [Ru(TMP)3]2+ binding. SUPPORT PROTOCOL 1
SYNTHESIS OF BIS(1,10-PHENANTHROLINE) (9,10-PHENANTHRENEQUINONE DIIMINE)RHODIUM(III) TRICHLORIDE Although several syntheses of [Rh(phen)2(phi)]3+ have been reported, the most reliable and highest-yielding procedure involves a three-step process. The synthetic scheme is summarized in Figure 6.2.2. In this preparation, the two ancillary phenanthroline ligands are added to the complex first (modified from Gillard et al., 1965). The resulting singly charged [Rh(phen)2(Cl)2]1+ is then converted to [Rh(phen)2(NH3)2]3+ by treatment with
1+
RhCl3•H2O
N
N
reflux
Cl
reflux
N
H3N
Rh
Cl
EtOH/H2O
N
Rh
H3N
N
N N
N
Hz•HCl N
3+
N
3+ O
O N H N
3:1 CH3CN/H2O 0.1 M NaOH
Probing Nucleic Acid Structure
Figure 6.2.2
N H
N
Rh N N
Synthesis of [Rh(phen)2(phi)]3+. Hz⋅HCl, hydrazine monohydrochloride.
6.2.4 Current Protocols in Nucleic Acid Chemistry
ammonium hydroxide (modified from Gidney et al., 1972). The bis-ammine complex is subsequently condensed with phenanthrene quinone to form the 9,10-phenanthrenequinone diimine (phi) intercalating ligand (modified from Mürner et al., 1998). All steps are carried out under ambient atmosphere and few extra precautions must be taken. Further details on the synthesis are available in Mürner et al. (1998). Materials Rhodium trichloride monohydrate 1,10-Phenanthroline Hydrazine monohydrochloride 95% (v/v) ethanol, denatured Concentrated (28% to 30%) ammonium hydroxide solution Phenanthrene quinone 3:1 (v/v) acetonitrile (reagent grade or better)/0.4 M NaOH solution in distilled water 1 M HCl solution (APPENDIX 2A) Sephadex SP-C25 cation-exchange resin 1 M MgCl2 solution (APPENDIX 2A) in distilled water HPLC-grade acetonitrile 0.1% (v/v) trifluoroacetic acid (TFA) in distilled water (for Sep-Pak elution) and in deionized, filtered water (for HPLC) Reflux condenser Temperature-controlled magnetic stir plate Oil bath or heating mantle 50- and 100-mL round-bottom flasks 60-mL medium-frit glass funnel 250-mL filter flasks Filter adapter Aspirator or vacuum pump Rotary evaporator Chromatography column: 1 to 2 feet (30 to 61 cm) long, 1 to 1.5 in. (2.5 to 3.8 cm) diameter 500-mL Erlenmeyer flasks Waters Sep-Pak 5-g C18 cartridges 15-mL plastic centrifuge tube or 25- to 50-mL round-bottom flask Lyophilizer (optional) High-pressure liquid chromatography (HPLC) system with C18 reversed-phase column Additional reagents and equipment for proton nuclear magnetic resonance (1H NMR), UV/visible spectrometry, and mass spectrometry Prepare [Rh(phen)2(Cl)2]Cl 1. Set up a reflux condenser with a temperature-controlled magnetic stir plate, an oil bath or heating mantle, and a 100-mL round-bottom flask containing a stir bar. 2. Combine 1 g rhodium trichloride monohydrate and 1.6 g of 1,10-phenanthroline (2 eq) in the 100-mL round-bottom flask. 3. Add 25 mg hydrazine monohydrochloride and 75 mL of a 1:1 (v/v) mixture of water and 95% ethanol.
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.5 Current Protocols in Nucleic Acid Chemistry
4. Stir and bring the mixture to reflux. Continue refluxing until all material has gone into solution and the color has changed from violet to yellow (∼4 to 6 hr). Hydrazine monohydrochloride acts as a reductive catalyst. By reducing the Rh(III) to Rh(II), ligand substitution is facilitated.
5. Remove residual unreacted ligand by filtering the hot solution through a 60-mL medium-frit glass funnel using an aspirator or a vacuum pump. Discard the ligand and collect filtrate in a 250-mL filter flask. 6. Refrigerate the filtrate overnight. The product will precipitate as yellow, powdery crystals. The solubility of the singly charged dichloro product in water is quite low. Any traces of [Rh(phen)3]3+ that are formed in the course of the reaction remain in solution and are eliminated by filtration.
7. Save the crystal product by filtration as in step 5. Collect the mother liquor in a 250-mL filter flask and store at 4°C in case it is needed to form a second crop of crystals. 8. Dry the first crop of product under vacuum in a sealed flask, using an aspirator or vacuum pump. Typical yields for the first crop are in the range of 60%.
9. Characterize the product by 1H NMR and mass spectrometry. Spectroscopic data are included below (see Compound Characterization).
Prepare [Rh(phen)2(NH3)2]Cl3 10. Combine 0.5 to 1.0 g [Rh(phen)2(Cl2)]Cl and 30 to 50 mL concentrated ammonium hydroxide solution in a 100-mL round-bottom flask. 11. With rapid stirring, bring the solution to reflux. Continue heating until all the yellow solid has dissolved (∼15 to 30 min). Remove the reaction from the heat and allow to cool. As the complex is converted from the singly charged dichloride to the triply charged diammine, solubility in ammonium hydroxide increases. As a result, solubility can serve as a rough measure of the reaction’s progress.
12. Remove the solvent from the reaction mixture using a rotary evaporator with a vacuum pump. The diammine product, which appears as a translucent yellow scale in the flask, is typically recovered in 90% to 100% yield.
13. Characterize the product by 1H NMR, UV/visible spectrometry, and mass spectrometry. Spectroscopic data are included below (see Compound Characterization).
Prepare [Rh(phen)2(phi)]Cl3 14. Combine 50 mg [Rh(phen)2(NH3)2]Cl3 and 25 mg phenanthrene quinone (an excess) in a 50-mL round-bottom flask. 15. Add 30 to 40 mL of 3:1 acetonitrile/0.4 M NaOH solution and stir briskly. Probing Nucleic Acid Structure
Sometimes the two solvent components initially form immiscible layers, but with rapid stirring there is sufficient mixing for the reaction to progress.
6.2.6 Current Protocols in Nucleic Acid Chemistry
16. Stir at room temperature for 6 to 12 hr. During this time the solution will go from yellow to a deep orange. The reaction progresses by base deprotonation of the coordinated ammine ligands, which add to the quinone with subsequent loss of water. Although extreme precautions are not necessary, it is recommended that the reaction flask be covered with aluminum foil to avoid unnecessary exposure to light.
17. Stop the reaction by neutralizing the NaOH with an equimolar amount of 1 M HCl solution. Purify by cation exchange 18. Swell 10 to 25 g dry Sephadex SP-C25 resin with 100 to 200 mL of 0.05 M MgCl2 solution for ∼15 min. The resin should swell completely but not consume all the free MgCl2 solution. A volume of extra solution (~1 cm above the level of the swelled resin) is necessary for proper pouring of the column. If all the solution is consumed, more should be added to the mixture.
19. Swirl the slurry and prepare a cation-exchange column by pouring the slurry into a 1- to 2-foot-long chromatography column and allowing it to settle as the excess solvent flows through. Stop the flow from the bottom of the column when the MgCl2 solution level reaches the top of the packed column resin. For adequate purification, the column should be ≥6 inches (∼15 cm) high.
20. Dilute the reaction mixture into a large volume (300 to 400 mL) of distilled water. Add the solution slowly to the top of the column, taking care not to unduly disturb the resin surface. The metal complex will adhere to the resin at the very top of the packed column. Unreacted quinone will flow through immediately.
21. Elute metal complexes with increasing concentrations of MgCl2. Begin with 2 to 3 column vol of 0.05 M MgCl2 and increase steadily to 0.5 M. Collect all eluted bands in individual 500-mL Erlenmeyer flasks. At the lowest concentrations, any additional unreacted quinone will elute off the column. This will be followed, as the concentration increases, by any unreacted diammine complex. The orange product band will elute between 0.2 and 0.5 M MgCl2.
22. For each column fraction containing [Rh(phen)2(phi)]Cl3, wash a Waters Sep-Pak 5-g C18 cartridge with 1 vol HPLC-grade acetonitrile followed by 1 vol water. Isolation of the complex on the reversed-phase cartridge will remove excess MgCl2 from the desired product.
23. Load the product fractions onto the cartridges either by gravity or under suction. Allow the entire product fraction to be adsorbed onto the resin. 24. Wash with 2 to 3 vol distilled water. 25. Elute the desalted product band with 1:1 (v/v) acetonitrile/0.1% TFA. Collect the material in either a 15-mL plastic centrifuge tube or a 25- to 50-mL round-bottom flask, depending on which technique will be used to remove the solvent. 26. Isolate the metal complex from this solution by rotary evaporation or lyophilization. The complex purified by cation-exchange chromatography has a purity >95%. Nonetheless, in the authors’ laboratory it is customary to further purify the material by reversed-phase HPLC (RP-HPLC), as described below. If an HPLC apparatus is unavailable, other methods of verifying the purity of the metal complex can be used
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.7 Current Protocols in Nucleic Acid Chemistry
(e.g., thin-layer chromatography using silica gel and 4:1:1:0.1 M CH3CN/water/n-butanol/saturated KNO3).
Purify by RP-HPLC 27. Dissolve a small amount of the material (e.g., 1 to 2 mg) in a volume of water appropriate for the injection loop volume of the HPLC apparatus. Inject it onto a C18 reversed-phase HPLC column. Run a gradient from 0.1% TFA to a 1:1 (v/v) mixture of 0.1 % TFA and acetonitrile over 30 min. The metal complex elutes between 20% and 30% acetonitrile.
28. Collect the product peak and evaporate or lyophilize the solvent. 29. Characterize the product by 1H NMR, UV/visible spectrometry, and mass spectrometry. Spectroscopic data are included below (see Compound Characterization). SUPPORT PROTOCOL 2
SEPARATION OF ENANTIOMERS OF BIS(1,10-PHENANTHROLINE) (9,10-PHENANTHRNENEQUINONE DIIMINE)RHODIUM(III) TRICHLORIDE The rac- and ∆-enantiomers of [Rh(phen)2(phi)]Cl3 are separated by ion-exchange chromatography with a chiral eluent. Typically, the procedure calls for a solution of cobalt L-cysteine sulfinate, Co(lcysu)3, which is made by procedures described by Dollimore and Gillard (1973) and presented in this protocol. Although procedures for both separating the enantiomers and making the cobalt complex are summarized here, the reader is directed to the original papers (Yoshikawa and Yamasaki, 1979; Cartwright et al., 1987) for more comprehensive details. Alternatively, small amounts of the enantiomers of [Rh(phen)2(phi)]3+ can be separated using a commercially available chiral HPLC column. Although chiral HPLC cannot produce the same amount of material as the chromatographic separation described here, it is useful for small-scale preparations or analysis of materials resolved by other means. Additional Materials (also see Support Protocol 1) Hexamminecobalt(III) chloride Nitrogen or argon gas L-Cysteine Potassium hydroxide 30% (v/v) hydrogen peroxide [Rh(phen)2(phi)]Cl3 (see Support Protocol 1) Sephadex CM-C25 ion-exchange resin 0.1 M KCl solution in deionized water
Probing Nucleic Acid Structure
Thermometer 500-mL, 2- or 3-neck round-bottom flasks with appropriate stoppers Bubbler for nitrogen or argon gas, equipped with a long pipet pH paper 60-mL coarse-frit glass funnels 500-mL filter flasks 1000-mL Erlenmeyer flask Chromatography columns: 2 to 4 feet (61 to 122 cm) long, 1 to 1.5 in. (2.5 to 3.8 cm) diameter 0.5 to 1 feet (15 to 30 cm) long, 1 to 1.5 in. (2.5 to 3.8 cm) diameter Solvent bottles for reservoirs
6.2.8 Current Protocols in Nucleic Acid Chemistry
Recirculating column pump (optional) Column end caps and tygon tubing for connecting recirculating pump and solvent reservoirs Additional reagents and equipment for circular dichroism (CD) spectroscopy Prepare K[Co(lcysu)3]⋅6H2O 1. Set up a reflux condenser with an oil bath or heating mantle and a thermometer. 2. Dissolve 5.34 g hexamminecobalt(III) chloride in 140 to 150 mL distilled water in a 500-mL round-bottom flask. 3. Deoxygenate the solution by bubbling with inert gas (nitrogen or argon) for 15 to 30 min. 4. Add 12.12 g L-cysteine and 16.84 g potassium hydroxide and heat in the oil bath or heating mantle to 70°C. Monitor the presence of ammonia by holding moistened pH paper over the outlet of the reflux condenser, and maintain the temperature until no further evolution of gaseous ammonia can be detected. The total reaction time is typically 24 hr.
5. Add 140 to 150 mL of 95% ethanol to precipitate the green potassium (+)-tris(L-cysteinato-SN)cobaltate(III). 6. Cool the mixture in an ice bath, vacuum filter through a 60-mL coarse-frit funnel using a 500-mL filter flask, and wash the resulting solid with 95% ethanol. 7. Add 100 mL of 30% hydrogen peroxide to another 500-mL round-bottom flask, and slowly add the collected precipitate while rigorously maintaining the solution’s temperature below 10°C in an ice bath. The material reacts rapidly, producing a bright yellow solution.
8. Precipitate the potassium (+)-tris(lcysu)cobaltate(III)⋅6H2O product by adding 200 mL of 95% ethanol. 9. Collect the product by vacuum filtration using another 60-mL coarse frit, wash the solid with 95% ethanol, and dry under vacuum. The K[Co(lcysu)3]⋅6H2O complex is light sensitive and should be stored in the dark for up to 1 to 2 months at 4°C. In addition, columns run with solutions of the complex should be protected from light by aluminum foil to minimize breakdown of the eluent.
Perform ion exchange 10. Prepare a 150 mM K[Co(lcysu)3]3− stock solution by dissolving the dry solid at 110.7 g/L in deionized water. The solution should be used as soon as possible. However, if storage is necessary, it can be stored for up to 1 week at 4°C in the dark.
11. Dissolve 30 to 50 mg [Rh(phen)2(phi)]Cl3 in a small volume of water (e.g., ~10 mL or more, depending on solubility). 12. Swell 50 to 100 g Sephadex CM-C25 ion-exchange resin with 400 to 600 mL of 0.1 M KCl solution in a 1000-mL Erlenmeyer flask for ∼30 min. 13. Pour the resulting loose suspension into a 2- to 4-foot chromatography column and allow to settle. If an insufficient amount of resin has been added for the desired column length (2 to 4 feet), add additional resin before the column has packed down.
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.9 Current Protocols in Nucleic Acid Chemistry
For enantiomer separation, a long column is required. If the column has settled completely and additional resin is added, an interface in the resin will be created that will affect band mobility. If the addition of more resin is necessary after complete column packing, add the resin and then mix the entire column by inversion before allowing it to pack again. This provides a more uniform column and better separation.
14. Wash the column thoroughly with several volumes of deionized water to remove any excess KCl solution. Drain excess water (i.e., to the level of the packed resin). If the potassium solution is not completely removed from the column, the cobalt eluent will precipitate on the resin.
15. Carefully load the rhodium solution (step 11) onto the top of the column, taking care not to disturb the resin surface. The colored complex should stick to the first 0.25 to 0.5 cm of column packing. If desired, an additional amount of swelled resin can be carefully layered over the product band with a long pipet to serve as a protective layer between the material and added solvent.
16. Run the column using a recirculating pump that circulates a small volume of cobalt eluent through the column at a flow rate of ∼1 mL/min. Begin eluting with a three-fold dilution of the 150 mM [Co(lcysu)3]3− stock solution (i.e., 50 mM), taking care not to disturb the resin bed. If a recirculating pump is unavailable, it is equally effective to prepare a larger volume of cobalt solution and recirculate it manually over time from a collection vessel at the base of the column to a reservoir above.
17. When the dilute cobalt solution begins to elute through the base of the column, increase the concentration of the eluent to 100 mM for an additional column volume, and then to 150 mM for an additional column volume. The stepwise equilibration of the column is necessary because the column resin will not contract uniformly if the ionic strength is increased too rapidly. Such nonuniform shrinkage will adversely affect the separation of the metal complex enantiomers.
18. Continue elution until two distinct, separated enantiomer bands are visible (usually a period of several days to two weeks, depending on the elution rate). Although the yellow color of the cobalt eluent makes it difficult to monitor the progress of the separation, the distinctively darker rhodium bands can usually be discerned.
19. When sufficient separation is obtained, stop elution and rinse the column thoroughly with several column volumes of distilled water to remove as much of the cobalt as possible. Once the excess eluent is washed out, the orange product bands should be easily located.
Isolate enantiomers 20. Allow the column to run dry and then expel the resin with air pressure onto a clean, flat surface.
Probing Nucleic Acid Structure
The column should come out as an intact cylinder with the product separation intact. If necessary, compressed air can be forced through the top of the column to remove all excess solvent before expelling the resin. In addition, it is often useful to mark the positions of the column bands on the outside of the column barrel before expelling the resin. The bands are often more difficult to locate in the dry resin, and comparison to the marked column can be very useful.
6.2.10 Current Protocols in Nucleic Acid Chemistry
21. Excise the bands of the separated enantiomers from the column with a laboratory spatula and place them into two 0.5- to 1-foot chromatography columns. With [Co(lcysu)3]3− as the eluent, the delta enantiomer of the rhodium complex elutes faster.
22. Wash with 0.1 M MgCl2 to remove any residual cobalt complex and ensure that the rhodium will have chloride counter ions when eluted from the resin. 23. Elute the separated enantiomers with 1 M MgCl2 and collect in 250-mL Erlenmeyer flasks. 24. Concentrate and desalt the products on Waters Sep-Pak C18 cartridges as described above (see Support Protocol 1, steps 22 to 26). 25. Characterize the product by CD spectroscopy to ensure the success of the enantiomeric separation. In addition, showing that the CD spectra of the two enantiomers are exact mirror images of one another demonstrates that no contaminating cobalt eluent is present. Spectroscopic data are included below (see Compound Characterization).
SYNTHESIS OF TRIS(PHENANTHROLINE) COMPLEXES OF RUTHENIUM(II)
SUPPORT PROTOCOL 3
In this protocol, modified from Lin et al. (1976), two tris(phenanthroline) complexes, [Ru(TMP)3]2+ and [Ru(phen)3]2+, are synthesized by analogous methods in a single step. The ruthenium trichloride starting material is combined with the appropriate phenanthroline ligands and refluxed to allow formation of complexes. The resulting materials are isolated by precipitation, purified by ion-exchange chromatography, and desalted on C18 reversed-phase cartridges. Additional Materials (also see Support Protocol 1) Ruthenium trichloride hydrate 6 M HCl solution (see Table A.2A.1) 3,4,7,8-Tetramethyl-1,10-phenanthroline (TMP) or 1,10-phenanthroline (phen) 30% (w/v) hypophosphorus acid solution 2 M NaOH solution 30-mL medium-frit glass funnels Prepare ruthenium complex 1. Set up a reflux condenser with a temperature-controlled magnetic stir plate, an oil bath or heating mantle, and a 100- to 200-mL round-bottom flask. Heat 60 to 75 mL distilled water in the flask to 90°C with stirring. 2. Add the following: 0.71 g ruthenium trichloride hydrate 2 drops 6 M HCl solution 2.1 g TMP or 1.7 g phen. NOTE: Stainless steel spatulas should not be used during the synthesis of this material as the phenanthroline ligands will complex with any dissolved iron impurities.
3. Continue heating for 10 to 20 min. During this time, the dark black solution will change to a dark green color.
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.11 Current Protocols in Nucleic Acid Chemistry
4. Neutralize 1.7 mL hypophosphorus acid solution to pH 7 with ∼5 mL of 2 M NaOH solution and add to the reaction mixture. Continue to reflux for an additional 60 min. The color will change to orange-red.
5. Remove from heat and filter hot through a 30-mL medium-frit funnel. Discard the precipitate. 6. While stirring, slowly add 20 mL of 6 M HCl solution to the filtrate. Collect the orange precipitate that forms by suction filtration. 7. Dissolve the isolated precipitate in a small volume of water and acetonitrile. The exact mixture depends on solubility. A good procedure is to start with 10 mL water and add acetonitrile until all material is dissolved (typically 15 to 25 mL total volume).
Purify product 8. Prepare a cation-exchange column as described (see Support Protocol 1, steps 18 and 19). For adequate purification, this column should be ≥10 cm high.
9. Load the entire product solution onto the top of the column with a long pipet, taking care not to unduly disturb the resin surface. The metal complex will adhere to the resin at the very top of the packed column. Unreacted phenanthroline ligands and any uncharged or anionic side products will flow through immediately.
10. Begin to elute with distilled water. After two to three column volumes of water have passed through and most of the yellow ligand has eluted, change to 0.05 M MgCl2. Increase the salt concentration over time to 0.2 M. The product should come off as a single broad orange band.
11. Collect the fraction in a 500-mL Erlenmeyer flask and isolate the product on a Sep-Pak reversed-phase cartridge as described (see Support Protocol 1, steps 22 to 26). 12. Characterize the products by 1H NMR, UV/visible spectrometry, and mass spectrometry. Spectroscopic data are included below (see Compound Characterization). BASIC PROTOCOL 2
SHAPE-SELECTIVE CLEAVAGE OF UNUSUAL STRUCTURES IN NUCLEIC ACIDS
Probing Nucleic Acid Structure
Tris(4,7-diphenyl-1,10-phenanthroline)rhodium(III), [Rh(DIP)3]3+, is a shape-selective structure probe of nucleic acids. Because of the sterically bulky phenyl substituents on the phenanthroline ligands (Fig. 6.2.3), this complex, like [Ru(TMP)3]2+, cannot intercalate easily into B-form DNA. Instead, the hydrophobic complex specifically targets unusual, non-B-form structures. Like other rhodium complexes, the complex promotes direct, photo-induced strand cleavage at sites where it is intimately bound. With much lower efficiency, the complex also damages DNA oxidatively, yielding damage at 5′-GG3′ and 5′-GA-3′ sites analogous to that observed by DNA-mediated electron transfer chemistry (Hall et al., 1996). The unusual structures targeted by the complex have been shown to include cruciforms, Z DNA, and other tertiary sites whose structures are not yet well characterized (Müller et al., 1987; Kirshenbaum et al., 1988; Lee and Barton, 1993). As a result, cleavage at a particular site by [Rh(DIP)3]3+ does not establish the presence of a particular DNA structure, but rather that the conformation of the nucleic acid at that
6.2.12 Current Protocols in Nucleic Acid Chemistry
3+
N N
N
Rh
N
N N
Figure 6.2.3
[Rh(DIP)3]3+.
site is an unusual one that is neither B nor A form and is not single stranded. For a discussion of nucleic acid structure, see APPENDIX 1B. Because cleavage with [Rh(DIP)3]3+ is very specific, experiments mapping a large piece of DNA for unusual structural forms are usually performed at low resolution to first identify candidate sites before moving to single-nucleotide resolution. Such experiments can scan an entire plasmid that contains sequences of interest, using agarose gel electrophoresis rather than PAGE to determine whether a [Rh(DIP)3]3+ cleavage site exists and approximately where it is located. Such low-resolution experiments can be done more quickly than experiments at single-nucleotide resolution, and thus ensure that experimental effort is not dissipated scrupulously screening a piece of DNA with no [Rh(DIP)3]3+ cleavage sites. Furthermore, this two-stage methodology provides an opportunity to determine whether the rhodium cleavage occurs on both DNA strands or only one and whether it depends on DNA superhelicity. Once sites have been identified at low resolution, restriction fragments of the plasmid or synthetic oligonucleotides containing the identified sites can be photocleaved and assayed by PAGE to determine the individual bases where cleavage takes place. Materials Supercoiled plasmid DNA (either commercial or containing a specific site of interest) Appropriate restriction enzymes (see steps for details) and buffer systems (according to manufacturer’s specifications) 1 µg/mL ethidium bromide solution (APPENDIX 2A) in water or 1× TBE electrophoresis buffer 7.5 M ammonium acetate solution in deionized water Absolute ethanol (200 proof, dehydrated) Dry ice 10× Tris/acetate buffer, pH 7.0 (see recipe) Metal complex solution (see recipe): 20 µM rac-[Rh(DIP)3]Cl3 (for synthesis, see Support Protocol 4) S1 single-strand-specific nuclease and appropriate buffer systems (according to manufacturer’s specifications) 9 mM base pairs of calf thymus DNA solution in deionized water, buffered to pH 7 to 9 DNA molecular weight standards (e.g., commercially available 100-base ladder) 1× TBE electrophoresis buffer (APPENDIX 2A) Loading buffer (e.g., formamide loading buffer or urea loading buffer; see recipes)
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.13 Current Protocols in Nucleic Acid Chemistry
Maxam-Gilbert sequencing reactions on labeled, unirradiated samples of 250- to 500-bp DNA fragment of interest (see Support Protocol 9) UV transilluminator Speedvac evaporator (Savant) or lyophilizer UV/visible spectrometer 90°C heating block Phosphorimager (optional) Additional reagents and equipment for restriction digests (e.g., CPMB UNIT 3.1), agarose gel electrophoresis (e.g., CPMB UNIT 2.5), photolysis (see Support Protocol 8), radiolabeling DNA (see Support Protocol 7), nondenaturing PAGE (e.g., CPMB UNIT 2.7A), denaturing PAGE (sequencing gels; APPENDIX 3B), autoradiography (optional; e.g., CPMB APPENDIX 3A), and mapping cleavage sites (see Support Protocol 9) NOTE: Because of the low solubility of rac-[Rh(DIP)3]Cl3 in water, a more concentrated solution of the material may need to be made in ethanol (or an ethanol/water mixture) and diluted to the desired stock concentration (20 µM) with water. Low-resolution cleavage Linearize plasmid DNA 1. Linearize 20 to 30 µg supercoiled plasmid DNA with the appropriate single-cutting restriction enzyme according to manufacturer’s instructions (e.g., see CPMB UNIT 3.1). This amount of DNA should be sufficient for one analysis (e.g., eight sample and control tubes, as described in step 7 below). The enzyme chosen must have only a single cut site on the plasmid DNA of interest.
2. Verify that the plasmid sample is completely linearized by analyzing ∼1 µg on a 1% (w/v) agarose gel. Stain the gel in 1 µg/mL ethidium bromide solution and visualize the stained DNA with a UV transilluminator. If the sample is not completely digested, retreat with enzyme until it is linearized.
3. Precipitate the remaining DNA by adding 1⁄5th to 1⁄4th vol of 7.5 M ammonium acetate solution and 4 to 5 vol absolute ethanol, mixing well, and chilling 1 hr on dry ice. 4. Microcentrifuge 12 min at maximum speed (14,000 rpm), at either room temperature or 4°C. 5. Decant the supernatant carefully and dry the pellet 15 to 20 min in a Speedvac evaporator or a lyophilizer to remove residual ethanol. Avoid overdrying the pellet or it may adhere to the walls of the microcentrifuge tube. 6. Resuspend DNA in deionized water, determine its concentration (and the concentration of the supercoiled plasmid stock) by UV/visible spectrometry (ε260 = 6600 M−1 cm−1 per nucleotide), and dilute both to 400 µM nucleotides in 2× Tris/acetate buffer.
Probing Nucleic Acid Structure
Irradiation experiments with both the supercoiled and liner plasmid samples will immediately assess any supercoiling dependence of a structure recognized by [Rh(DIP)3]Cl3. This is particularly relevant in the case of some cruciform structures that are only extruded from the double-stranded form in the presence of supercoiling strain.
6.2.14 Current Protocols in Nucleic Acid Chemistry
Cleave DNA 7. Combine equal volumes of 20 µM rac-[Rh(DIP)3]Cl3 and 400 µM DNA stock solutions as described (see Support Protocol 8, steps 1 to 3), but mix samples only by gentle tapping of the tube. For each plasmid of interest, prepare a light and a dark control and two experimental samples for both the linear and supercoiled forms (total eight tubes). One half of the experimental samples will be treated with S1 nuclease (see step 11 below). If desired, the dark control can be performed with either linear or supercoiled DNA; both are not required, as they assess contaminants in the metal solution. Centrifugation of irradiation samples should be avoided because it may result in metal complex precipitation.
8. Irradiate 20 to 60 sec at 330 nm with a Hg-Xe arc lamp or 10 min with a transilluminator (see Support Protocol 8, step 4) and precipitate DNA (steps 3 to 5 above). Under these conditions, photolysis leads to partial or complete conversion of supercoiled samples to nicked or linear plasmid DNA.
9. Resuspend the supercoiled irradiation samples in the buffer appropriate for the enzyme used above and treat with the same restriction enzyme for the same total incubation time as used in step 1. 10. Precipitate samples as in steps 3 to 5. 11. Treat half of the experimental samples (including both supercoiled and linearized plasmid irradiations) with S1 nuclease according to manufacturer’s instructions. In the authors’ experiments this involves buffering DNA solutions to pH 6, adding ?? zinc sulfate, and treating with 10 to 20 units of nuclease for 20 min at 37°C. This enzymatic treatment converts any single-strand breaks caused by the complex into double-strand breaks that can be observed in a low-resolution agarose gel experiment. It also provides the opportunity to assess whether there is a supercoiling dependence to the type of strand break induced by the complex.
12. Precipitate samples as in steps 3 to 5. Analyze samples 13. Electrophorese 1 µg DNA/lane on a 1% (w/v) agarose gel. Run DNA molecular weight standards alongside the experimental samples to better assess the approximate position of any breaks vis-a-vis the restriction enzyme site used to linearize the plasmid. 14. Stain the gel in 1 µg/mL ethidium bromide solution and visualize using a transilluminator. If desired, record data photographically using a Polaroid camera. Negatives of the photographs can be quantitated using a densitometer to determine the exact amounts of conversion of various forms of the plasmid.
High-resolution cleavage Linearize plasmid DNA 15. Linearize DNA as described above (steps 1 to 6), but use an enzyme that has a single cut site within 100 to 300 bp of any interesting structures revealed in the low-resolution experiment. This will generate pieces that are short enough for adequate resolution.
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.15 Current Protocols in Nucleic Acid Chemistry
Cleave DNA 16. Irradiate samples as described in steps 7 and 8 above. Irradiation times for high-resolution experiments must be optimized for a particular system to ensure that the amount of photocleavage is sufficient for PAGE analysis, but that the end-labeled DNA is not overcleaved. To determine an acceptable irradiation time, prepare a set of identical samples and irradiate each for steadily increasing times. A good starting point is ∼5 min at 330 nm. Select an irradiation time where the cleavage bands are easily observable on the gel but where most of the full-length DNA band (≥90%) remains uncleaved. At long irradiation times, in the absence of strong binding sites for the complex, non-structure-specific damage is observed at 5′-GG-3′ and 5′-GA-3′ sites. This should not be confused with structure-specific cleavage.
17. Precipitate all samples by adding 1 to 5 µL of 9 mM calf thymus DNA, 1⁄6th to 1⁄4th vol of 7.5 M ammonium acetate, and 5 vol absolute ethanol. Mix well and heat to 90°C for 5 min. Mix again and incubate on dry ice for 30 min. Heating ensures complete dissociation of the metal-DNA complex prior to precipitation. It is important to remove the metal complex as it can affect the electrophoretic mobility of the DNA products.
18. Microcentrifuge 12 min at maximum speed (14,000 rpm). 19. Carefully remove the supernatants, resuspend each pellet in 50 to 100 µL water, and repeat steps 17 and 18, but without heating. 20. Remove the supernatants, rinse the resulting pellets twice with cold 80% (v/v) ethanol, and dry under vacuum. 21. Treat supercoiled samples with restriction enzyme (as in step 15) and precipitate as described in steps 17, 18 and 20, but without heating. Analyze samples 22. Resuspend all samples in deionized water and label them with [32P]phosphate or [32P]nucleotides (see Support Protocol 7). Labeling of the 5′ end of the plasmid will require dephosphorylation with a phosphatase and then rephosphorylation with [g-32P]ATP. Labeling at the 3′ end is routinely done by filling in the overhang left by the restriction enzyme with an appropriate polymerase and radiolabeled nucleoside triphosphates.
23. Cut DNA with an additional restriction enzyme selected to excise the region of interest in a segment that is 250 to 500 bp long. 24. Isolate the desired fragment on a 5% to 10% nondenaturing polyacrylamide gel. 25. Identify the position of the band of interest by autoradiography, excise the gel band, and isolate the labeled DNA by soaking the gel slice 2 to 4 hr in 1× TBE electrophoresis buffer. The progress of elution can be monitored by comparing the amount of radiation in the supernatant to that remaining in the gel.
26. Precipitate the isolated DNA (steps 17 to 20).
Probing Nucleic Acid Structure
27. Resuspend in denaturing loading buffer, heat 2.5 to 5 min at 90°C, and resolve on a 6% to 8% denaturing polyacrylamide sequencing gel. Run with Maxam-Gilbert sequencing reactions on labeled, unirradiated samples of the same DNA fragment (see Support Protocol 9). Visualize results by autoradiography or by phosphorimagery.
6.2.16 Current Protocols in Nucleic Acid Chemistry
SYNTHESIS OF TRIS(4,7-DIPHENYL-1,10-PHENANTHROLINE) RHODIUM(III) TRICHLORIDE
SUPPORT PROTOCOL 4
In this protocol, which is modified from Gidney et al. (1972), [Rh(DIP)3]3+ is synthesized in a single step from rhodium trichloride and 4,7-diphenyl-1,10-phenathroline (DIP). The product is isolated by precipitation. Because of the low solubility of the complex in all solvents applicable for ion-exchange chromatography, the product is purified by recrystallization. Additional Materials (also see Support Protocol 1) 4,7-Diphenyl-1,10-phenanthroline (DIP) 2.5 mg/mL hydrazine monohydrochloride in water Saturated NaCl solution Acetone 100-mL Erlenmeyer flask 30-mL medium-frit glass funnel 100-mL filter flask 1. Set up a reflux condenser with a temperature-controlled magnetic stir plate and an oil bath or heating mantle. 2. Dissolve 50 mg rhodium trichloride monohydrate in 10 mL deionized water in a 50-mL round-bottom flask. 3. Dissolve 200 mg of 4,7-diphenyl-1,10-phenanthroline in 10 mL of hot (80°C) 95% ethanol in another 50-mL round-bottom flask. While stirring, add the ligand solution to the rhodium solution. Some precipitation may occur. NOTE: No stainless steel spatulas should be used during the synthesis of this material as the phenanthroline ligands will complex with any dissolved iron impurities.
4. Add 1 mL of 2.5 mg/mL hydrazine monohydrochloride to the reaction mixture. 5. Connect the reaction flask to the condenser and bring to reflux. Continue to reflux the mixture for 72 hr. After 30 min the solution should be clear yellow. Over the 72-hr period, it should become bright orange.
6. Allow the reaction to cool and remove the solvent using a rotary evaporator with a vacuum pump. This should yield a yellow-orange solid. A yellow side product of the reaction is often produced along with the desired complex. It has been assigned, by NMR and mass spectrometry, as [Rh(DIP)2Cl2]Cl. It is much less soluble in ethanol than the desired trischelate complex and can be separated on that basis in the following recrystallization step. Presence of red or pink coloration indicates contamination by iron complexes.
7. Transfer the solid to a 100-mL Erlenmeyer flask and recrystallize from 50 mL of hot 1:1 (v/v) ethanol/water containing 1 mL saturated NaCl solution. 8. Quickly decant the complex solution from any residual solid and allow to cool to room temperature. Chill at 4°C for several hours to ensure that crystallization is as complete as possible.
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.17 Current Protocols in Nucleic Acid Chemistry
Because of the low solubility of the material, the crystals will come out very quickly once the solution begins to cool.
9. Collect the product by vacuum filtration using a 30-mL medium-frit glass funnel. This should yield peach-colored crystals. Impurities (e.g., red iron complexes) cause significant changes in color. If necessary, the resulting solid can be recrystallized by dissolving in a minimum volume of ethanol and pouring the resulting solution into a large volume (200 to 250 mL) of acetone. Because of the low solubility of this compound in all solvents applicable for ion-exchange chromatography, purification of the material using that methodology is not practical. Spectroscopic data are included below (see Compound Characterization). BASIC PROTOCOL 3
RECOGNITION OF MISMATCHES AND ABASIC SITES IN DNA In addition to the use of shape-selective ancillary ligands to probe the characteristics of the grooves or structures of DNA as discussed with [Ru(TMP)3]2+ and [Rh(DIP)3]3+, shape-selective intercalating ligands can be used to probe differences in base pairing at the core of the DNA helix. Bis(2,2′-bipyridine)(5,6-chrysenequinone diimine)rhodium(III), [Rh(bpy)2(chrysi)]3+ (see Fig. 6.2.4), is such a complex that utilizes a broad, four-ring chrysenequinone diimine intercalating ligand. Like [Rh(DIP)3]3+, where the size of substituents on the ancillary ligands only allows interaction at sites of tertiary structure, the steric bulk of the chrysi ligand prevents it from intercalating into DNA except at sites where disrupted base pairing opens up the structure of the helix. This selectivity has been applied to the recognition of the destabilized helical structures characteristic of both mismatched base pairs in DNA (Jackson and Barton, 1997) and abasic sites (B.A. Jackson and J.K. Barton, unpub. observ.). Although the recognition properties of the two enantiomers of the complex are different, racemic material is sufficient for routine experiments. Photocleavage experiments with [Rh(bpy)2(chrysi)]3+ are performed as described (see Support Protocol 8). Since the structures that are probed with this molecule (i.e., mismatches and abasic sites) are not generally found at specific sites of plasmid restriction fragments, experiments are routinely done on synthetic oligonucleotides. As a result, DNA concentrations are expressed in terms of polymers rather than concentration of base pairs or nucleotides. Materials Metal complex solution (see recipe): rac-[Rh(bpy) 2(chrysi)]Cl3 solution (typically 2 µM; for synthesis, see Support Protocol 5) Labeled DNA solution (see Support Protocol 7): typically 20 µM polymers in 2× buffer
H
N N H
Probing Nucleic Acid Structure
Figure 6.2.4
N
Rh
N N
N
[Rh(bpy)2(chrysi)]3+.
6.2.18 Current Protocols in Nucleic Acid Chemistry
Additional reagents and equipment for photolysis (see Support Protocol 8) and for mapping cleavage sites (see Support Protocol 9) 1. Mix equal volumes of rac-[Rh(bpy)2(chrysi)]Cl3 solution and labeled DNA solution for a total volume of 10 to 50 µL as described (see Support Protocol 8, steps 1 to 3). Set up dark and light controls for each DNA of interest. 2. Irradiate 5 min at 313 nm or 15 to 30 min at 365 nm and treat as described (see Support Protocol 8, steps 4 to 8). 3. Perform electrophoresis and autoradiography as described (see Support Protocol 9). Sites of [Rh(bpy)2(chrysi)]3+ binding are indicated by cleavage bands in the experimental lanes.
4. Identify specific sites of cleavage by comparing to Maxam-Gilbert sequencing reactions as described in Support Protocol 9. Although cleavage band intensity is related to the affinity of the complex for individual DNA sites, differences in photocleavage efficiency from site to site make direct comparisons difficult.
SYNTHESIS OF BIS(2,2′-BIPYRIDINE)(5,6-CHRYSENEQUINONE DIIMINE)RHODIUM(III) TRICHLORIDE
SUPPORT PROTOCOL 5
[Rh(bpy)2(chrysi)]3+ is synthesized using the same condensation chemistry described for the synthesis of [Rh(phen)2(phi)]3+ (see Support Protocol 1) and is modified from Mürner et al. (1998). The [Rh(bpy)2Cl2]Cl intermediate is made using the same protocol, except that bipyridine is substituted for the phenanthroline ancillary ligands. It is then converted to the diammine complex and condensed with chrysenequinone. Because chrysenequinone is not commercially available, its one-step synthesis from the parent aromatic hydrocarbon is also described (Mürner et al., 1998; originally from Greabe and Hönigsberger, 1900). Additional Materials (also see Support Protocol 1) Chrysene Sodium dichromate Glacial acetic acid 2,2′-Bipyridine Mortar and pestle 250- and 1000- to 2000-mL round-bottom flasks 250-mL beaker 60-mL coarse-frit glass funnels 250- and 1000-mL filter flasks 2000-mL Erlenmeyer flask Synthesize 5,6-chrysenequinone 1. Set up a reflux condenser with a temperature-controlled magnetic stir plate and an oil bath or heating mantle. 2. Grind 5 g chrysene into a fine powder with a mortar and pestle. This procedure ensures that the material will be oxidized efficiently, and is necessary when the chrysene is in the form of crystalline “chunks.” If it can be obtained as a fine powder, further grinding is not required.
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.19 Current Protocols in Nucleic Acid Chemistry
CAUTION: To prevent exposure to the fine dust produced in this procedure, the grinding should be performed in a fume hood. In addition, appropriate personal safety devices should be used, including gloves, a laboratory coat, and a face mask.
3. Add the 5 g chrysene and 20 g sodium dichromate to a 250-mL round-bottom flask. 4. Add 50 g glacial acetic acid, connect the flask to a condenser, and bring the mixture to reflux with stirring. Continue refluxing for a total of 9 hr. As heating continues, the solution will change from the orange-yellow of the dichromate/chrysene to a deep green with visible orange-red crystals of the chrysenequinone product.
5. Remove the reaction mixture from heat and immediately add to a 250-mL beaker containing 75 to 100 mL boiling distilled water. The product will crash out of solution while the green chromium by-products remain soluble.
6. Collect the product by filtering the solution through a 60-mL coarse-frit glass funnel using a 250-mL filter flask and a water aspirator or vacuum pump. 7. Wash crystals thoroughly with 250 to 500 mL water. Recoveries at this step are typically 80% to 90%.
Recrystallize 5,6-chrysenequinone 8. Place ∼0.8 to 0.9 g product into a 1000-mL round-bottom flask and add ∼800 to 900 mL of 95% ethanol (∼1 liter per gram of solid). 9. Connect to the reflux condenser and bring to a boil with stirring. The solution should become a clear orange-red color. If there are still orange crystals that remain undissolved, more ethanol should be added. However, any yellow chrysene starting material should not dissolve.
10. Decant the hot solution away from any undissolved material into a 2000-mL Erlenmeyer flask. Allow the solution to cool to room temperature, then chill to 4°C for several hours to promote further crystallization. 11. Collect the product by filtration with a 60-mL coarse-frit glass funnel and a 1000-mL filter flask. 12. Rinse with a small volume of ethanol (e.g., 50 to 100 mL) and dry the isolated product under vacuum. 13. Characterize the product by 1H NMR and mass spectrometry. Spectroscopic data are included below (see Compound Characterization).
Synthesize [Rh(bpy)2(chrysi)]3+ 14. Prepare [Rh(bpy)2(NH3)2]Cl3 intermediate as described for [Rh(phen)2(NH3)2]Cl3 (see Support Protocol 1, steps 1 to 13), but substitute 2,2′-bipyridine for the phenanthroline ancillary ligands. Spectroscopic data are included below (see Compound Characterization).
Probing Nucleic Acid Structure
15. Prepare [Rh(bpy)2(chrysi)]3+ by condensation of [Rh(bpy)2(NH3)2]Cl3 with purified 5,6-chrysenequinone as described for the synthesis of [Rh(phen)2(phi)]Cl3 (see Support Protocol 1, steps 14 to 29), adjusting the amounts of materials commensurate with their different molecular weights.
6.2.20 Current Protocols in Nucleic Acid Chemistry
5,6-Chrysenequinone is less soluble in the acetonitrile/water solvent system but, as the reaction progresses, undissolved starting material goes into solution as the dissolved quinone is consumed. Unlike the previous synthesis, which went from yellow to orange, the color change here is more subtle. The formation of the diimine product gradually changes the red-orange of the quinone solution to a dark brown. After the reaction is completed, ion-exchange purification is done identically as described above. The products should be characterized by 1H NMR, UV/visible spectrometry, and mass spectrometry. Spectroscopic data are included below (see Compound Characterization).
PHOTOFOOTPRINTING OF DNA-BINDING MOLECULES The complex bis(9,10-phenanthrenequinone diimine)(2,2′-bipyridyl)rhodium(III), [Rh(phi)2(bpy)]3+, is a strong, sequence-neutral DNA-binding molecule. As with the complexes discussed above, it also cleaves DNA upon photoactivation. This neutral binding behavior and photocleavage can be applied to the footprinting of both DNA-binding proteins and small DNA-binding molecules. Because the cleavage chemistry involves direct reaction rather than a diffusible species, high-resolution footprints are obtained. Since the intercalative interaction of the molecule “senses” both grooves of the DNA helix, these molecules can be used to footprint molecules that bind in either the major or the minor groove (Uchida et al., 1989).
BASIC PROTOCOL 4
Materials Metal complex solution (see recipe): [Rh(phi)2(bpy)]Cl3 solution (typically 10 µM; for synthesis, see Support Protocol 6) Labeled DNA solution (see Support Protocol 7): typically 10 µM base pairs in a 2× buffer appropriate to the DNA binding molecule of interest Labeled DNA/binding molecule solution: labeled DNA solution containing DNA-binding molecule at twice the concentration desired in the photofootprinting experiment Additional reagents and equipment for photolysis (see Support Protocol 8) and mapping cleavage sites (see Support Protocol 9) 1. Mix equal volumes of [Rh(phi)2(bpy)]Cl3 and labeled DNA solutions for a total volume of 10 to 50 µL as described (see Support Protocol 8, steps 1 to 3). Set up dark and light controls of each DNA of interest. It is important to keep a 1:1 ratio between the rhodium complex and the DNA base pairs in the experiment to preserve the sequence neutrality of the cleavage reagent. Some selectivity is observed at low Rh/DNA ratios.
2. Set up separate reactions and controls using the labeled DNA/binding molecule solution. 3. Irradiate 5 to 10 min at 313 nm and treat as described (see Support Protocol 8, steps 4 to 8). 4. Perform electrophoresis and autoradiography as described (see Support Protocol 9). Sites of DNA binding by the molecule of interest will be obvious from the attenuation of rhodium complex cleavage.
5. Identify specific bases of protection by comparison with Maxam-Gilbert sequencing reactions as described in Support Protocol 9. For more complete characterization of a DNA-binding interaction, multiple footprinting reactions (with a set amount of DNA and rhodium complex) can be performed at different
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.21 Current Protocols in Nucleic Acid Chemistry
concentrations of DNA-binding molecule to titrate in the cleavage footprint. Weaker protection is observed at lower concentrations. SUPPORT PROTOCOL 6
SYNTHESIS OF BIS(9,10-PHENANTHRENEQUINONE DIIMINE)(2,2′-BIPYRIDYL)RHODIUM(III) TRICHLORIDE [Rh(phi)2(bpy)]3+ is synthesized in the two-step process shown in Figure 6.2.5. In this methodology, 9,10-diaminophenanthrene (phi) is used to generate a [Rh(phi)2Cl2]Cl complex (from Pyle et al., 1990). Before introduction of the third chelating ligand, the chloride ligands are removed with silver triflate. Preparation of [Rh(phi)2(bpy)]3+ is from Howells and McCown (1977) and Pyle et al. (1990). Although this reaction is described on a relatively large scale, it can be scaled down without difficulty. In addition to the procedure described here for the synthesis of [Rh(phi)2(bpy)]3+, a synthesis based on the condensation methodology described above for [Rh(phen)2(phi)]3+ (see Support Protocol 2) would also be an effective method to make this molecule. Although specific reaction conditions have not been determined to produce the bipyridyl complex, methods have been developed to make the very similar [Rh(phi) 2(phen)]3+. The conditions required to make this molecule are likely to be almost identical (Mürner et al., 1998). Additional Materials (also see Support Protocol 1) 9,10-Diaminophenanthrene Dimethylformamide Argon gas Absolute ethanol (anhydrous) Chloroform Silver trifluoromethanesulfonate (silver triflate) 2,2′-Bipyridine 50% (v/v) acetonitrile (reagent grade or better) in distilled water
1+
RhCl3•H2O H2N
H
1. reflux DMF/EtOH Hz•HCl
NH2
H N
N
Rh
N H
2. oxidation
N H
Cl Cl
3+
1. AgOTf DMF 70°C
H N
2. DMF 60°C
N H
N
Probing Nucleic Acid Structure
H
N N
Rh
H
N N
N
3+ Figure 6.2.5 Synthesis of [Rh(phi)2(bpy)] . AgOTf, silver triflate; DMF, dimethylformamide; Hz⋅HCl, hydrazine monohydrochloride.
6.2.22 Current Protocols in Nucleic Acid Chemistry
Sephadex QAE-25 anion-exchange resin 1 M HCl (APPENDIX 2A) Sephadex CM-C25 cation-exchange resin 1000-mL three-necked, round-bottom flasks Vacuum line with argon flush 100-, 250-,and 500-mL round-bottom flasks Septa to fit flasks Heat gun 60-mL gas-tight syringe with long needle 25-, 500-, and 1000-mL Erlenmeyer flasks 60-mL coarse- and fine-frit glass funnels Chromatography columns: 0.5 to 1 feet (15 to 30 cm) long, 1 to 1.5 in. (2.5 to 3.8 cm) diameter 1 to 2 feet (30 to 61 cm) long, 1 to 1.5 in. (2.5 to 3.8 cm) diameter Prepare [Rh(phi)2Cl2]Cl 1. Set up a reflux condenser with a temperature-controlled magnetic stir plate and an oil bath or heating mantle. 2. Combine 3.5 g of 9,10-diaminophenanthrene and 65 to 75 mg hydrazine monohydrochloride in a 1000-mL three-necked, round-bottom flask. 3. Add 250 mL dimethylformamide and immediately connect the flask to the reflux condenser, seal it with septa, and begin stirring the mixture to dissolve the reactants. 4. While stirring, deoxygenate the solution by pumping and backfilling with argon gas. Repeat deoxygenation at least three times. 5. In a 250-mL round-bottom flask, add 80 mL absolute ethanol to 1.6 g rhodium trichloride monohydrate. Dissolve the rhodium completely by swirling the flask and applying gentle heat with a heat gun. 6. Seal the flask with a septum and deoxygenate as described in step 4 by piercing the septum with a long syringe needle connected to the argon/vacuum line. 7. When both flasks are completely deoxygenated, transfer the rhodium solution to the three-neck reaction flask with a 60-mL gas-tight syringe. 8. Deoxygenate the reaction mixture again by repeated pumping down and backfilling with argon. 9. Heat the reaction at 80°C under argon for 16 to 24 hr. 10. Remove from heat and transfer to a 1000-mL Erlenmeyer flask. Open the mixture to air and stir for several days to allow oxidation of the coordinated amines to immine ligands. 11. Collect the product by filtration through a 60-mL coarse-frit glass funnel using a 250-mL filter flask and an aspirator or vacuum pump. 12. Wash with copious amounts (~100 mL each) of distilled water, ethanol, and chloroform (in that order). 13. Characterize the product by 1H NMR and mass spectrometry. Spectroscopic data are included below (see Compound Characterization). Spectroscopy shows that both a trans and a cis product are produced in this reaction. Both isomers, however, can go on to react with an additional chelating ligand to form the desired product.
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.23 Current Protocols in Nucleic Acid Chemistry
Prepare [Rh(phi)2(bpy)]3+ 14. Dissolve 1 g [Rh(phi)2Cl2]Cl in 40 mL dimethylformamide in a 100-mL round-bottom flask. 15. In a 25-mL Erlenmeyer flask, dissolve 1.25 g silver triflate in 8 mL dimethylformamide. 16. Add the silver triflate solution to the rhodium complex, deoxygenate the reaction, and heat to 70°C. Continue heating for 48 hr under argon. 17. Remove the dark silver chloride precipitate by vacuum filtration, first through a 60 mL coarse and then through a 60-mL fine frit, using a 250-mL filter flask and an aspirator or vacuum pump. 18. Rinse the precipitate with 50 mL dimethylformamide to bring the total volume of the filtrate to ∼100 mL. 19. Transfer the triflate intermediate solution to a 250-mL round-bottom flask and add 0.5 g (2 eq) of 2,2′-bipyridine chelating ligand. To ensure as complete a reaction as possible, an excess of ligand should be added.
20. Deoxygenate the reaction mixture several times, then stir at 60°C under argon for 24 hr. 21. Remove the solvent using a rotary evaporator with a vacuum pump, and dissolve the product in 10 to 20 mL of 50% acetonitrile for chromatography. The product is now ready for anion-exchange chromatography, which exchanges any of the triflate counter-ions in the reaction for chloride ions. This will improve the solubility of the complex in water and help in the subsequent cation-exchange purification column.
Purify by anion exchange 22. Swell a small amount (10 to 20 g) of Sephadex QAE-25 anion-exchange resin in 50 to 100 mL of 50% acetonitrile containing 0.1 M HCl in a 500-mL Erlenmeyer flask. 23. Pour the slurry into a 0.5- to 1-foot column and allow it to pack to a height of 5 to 7 inches (13 to 18 cm). 24. Wash the column with one to two column volumes of 50% acetonitrile. When the level of the solvent approaches the top of the resin, pour the product solution into the column and immediately begin collecting the effluent. Although it is desirable to keep the packing as undisturbed as possible, it is not necessary to worry too much about agitating the top of the column. It is necessary to begin collecting immediately because the reaction mixture will flow straight through.
25. Collect all the metal complex off the anion-exchange column by washing it with one to two column volumes of 50% acetonitrile. 26. Transfer the fraction to a 500-mL round-bottom flask and remove the solvent using a rotary evaporator.
Probing Nucleic Acid Structure
Purify by cation exchange 27. Swell 25 to 50 g Sephadex CM-C25 cation-exchange resin in 200 to 400 mL of 50% acetonitrile containing 0.1 M HCl. Pour a 1- to 2-foot column and allow the resin to settle. 28. Wash the column with one to two column volumes of 50% acetonitrile.
6.2.24 Current Protocols in Nucleic Acid Chemistry
The mixture of acetonitrile, water, and Sephadex CM resin is used instead of the SP resin used in previous protocols to prevent the complex from sticking irreversibly to the chromatography column.
29. Dissolve the dried product (step 25) in a minimum volume of 50% acetonitrile (just enough to dissolve all the solid) and load carefully to the top of the cation-exchange column, taking care not to disturb the resin surface. All the orange metal complex should adhere to the top 1⁄4 to 1⁄2 inch of column packing. If desired, a small layer of swelled resin may be added over the top of the product band using a long pipet. This protective layer prevents disruption of the product band when solvent is subsequently added.
30. Begin eluting the column with 50% acetonitrile. Increase the concentration of HCl first to 0.05 M and then gradually to 0.25 M to elute the charged metal complexes. Typically 100 to 200 mL of each molarity are used, but this depends of the results obtained. If not bands move at a given molarity, the entire volume need not be used. If bands move at a reasonable rate, the volume can be increased to 400 to 500 mL to elute a given product band. Any ligand or ligand breakdown products that remained in the reaction mixture will come off the column immediately. Any remaining rhodium phi starting materials should elute next. The [Rh(phi)2(bpy)]Cl3 will elute last as an intense orange band.
31. To isolate the product from the appropriate fraction, evaporate the solvent to dryness by rotary evaporation or lyophilization. Residual hydrochloric acid can be removed by repeatedly dissolving the material in methanol and redrying. If desired, the product can be more completely desalted by dissolving in water and using a Waters Sep-Pak reversed-phase cartridge as described earlier (see Support Protocol 1, steps 22 to 26).
32. Characterize the product by 1H NMR, UV/visible spectrometry, and mass spectrometry. Spectroscopic data are included below (see Compound Characterization).
SHAPE-SELECTIVE CLEAVAGE OF RNA Some of the complexes described above for the shape-selective recognition of DNA can also be applied to the study of RNA structural forms. Their utility as structural probes of RNA is considerable, and the sites targeted tend to be unique for current RNA chemical probes. [Rh(phen)2(phi)]3+ (see Basic Protocol 1) is particularly valuable as a probe of RNA structure. Because of the preference of RNA duplexes to adopt the A form, recognition by [Rh(phen)2(phi)]3+ becomes much more specific on RNA than on B-form DNA. Because of its width and depth, the A-form region is not bound by [Rh(phen)2(phi)]3+. Hence, on folded RNA, the complex can bind only to areas of tertiary interaction or to sites where the major groove has become substantially opened and accessible to intercalation. At such points (e.g., the D-T loop on tRNAPhe, triply bonded sites, and helix-loop junctions), the major groove of the nucleic acid is accessible. This selective identification of sites of tertiary interactions is a very useful tool in exploring the three-dimensional structure of an unknown segment of RNA. It is noteworthy that sites that are opened in the major groove are often specific targets for RNA-binding proteins. As a result, such structural information could find application not only in the study of the nucleic acids themselves but in the rapidly expanding study of RNA-protein interactions.
BASIC PROTOCOL 5
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.25 Current Protocols in Nucleic Acid Chemistry
[Rh(DIP)3]3+ (see Basic Protocol 2) recognizes and cuts RNA in areas of folded loops. This recognition is similar to the recognition of DNA cruciforms, which is assumed to be a mostly hydrophobic interaction with areas of the molecule that are not purely double or single stranded. Of greatest utility is the fact that [Rh(DIP)3]3+ recognizes GU mismatches in RNA within double-helical regions (Chow and Barton, 1992). Recognition of RNA by [Ru(TMP)3]2+ and [Ru(phen)3]2+ (see Basic Protocol 1) is much less informative than by the rhodium complexes. Although one might expect strong binding of [Ru(TMP)3]2+ to the A-form helices of the RNA, very little selective cleavage is observed. Instead, photolytic cleavage with [Ru(phen)3]2+ provides a practical and clean chemical G sequencing reaction. Application of [Ru(phen)3]2+ in this context as a sequencing reagent is therefore described below (see Basic Protocol 6). In all cases, use of these metal complexes to probe RNA structure is analogous to the experiments described in the previous sections for DNA. Although the types of information the complexes provide are different, the solutions, procedures, and equipment are the same. BASIC PROTOCOL 6
SINGLET OXYGEN-MEDIATED CLEAVAGE AT GUANINE RESIDUES IN DNA AND RNA The use of [Ru(phen)3]2+ to control for the shape-selective recognition properties of [Ru(TMP)3]2+ described in Basic Protocol 5 is not the only use of the complex as a DNA probe. As a result of its nonspecific binding behavior and general oxidative reactivity, [Ru(phen)3]2+ can also be used as a guanine-specific photosequencing reagent to complement the chemical Maxam-Gilbert methods. By simply applying the protocol described in Basic Protocol 1 to a DNA or RNA of interest, specific and clean determinations of guanine positions in DNA and RNA can be made.
SUPPORT PROTOCOL 7
Probing Nucleic Acid Structure
PREPARATION AND LABELING OF DNA AND RNA The nucleic acid samples used in studies with these probes have been prepared synthetically using standard solid-phase chemistries or excised from plasmids using restriction endonucleases, or, in the case or RNA, sometimes produced by enzymatic run-off transcription. All samples must be stringently purified either by reversed-phase high-performance liquid chromatography (RP-HPLC) or by PAGE and labeled with 32P on either the 5′ or 3′ end by standard enzymatic methods. Comprehensive protocols for these activities are included elsewhere in this book and in Current Protocols in Molecular Biology, and the reader is directed there for specific details (for nucleic acid synthesis, see Chapter 3 and APPENDIX 3C of this book; for restriction digestion of plasmid DNA, see CPMB UNIT 3.1; for run-off transcription of RNA, see CPMB UNIT 3.8; for purification of restriction fragments, see CPMB UNIT 2.7; for labeling of nucleic acids, see CPMB UNITS 3.5-3.15 and also UNIT 6.1 of this book). The labeled DNA solution should contain unlabeled, sonicated calf thymus DNA or oligonucleotide DNA (carrier DNA; see below) with enough radiolabeled DNA so that the specific activity of a single irradiation sample is between 20,000 and 200,000 cpm. The appropriate amounts of DNA required are discussed in each Basic Protocol and refer to the amount of carrier DNA, as labeled DNA (on the picomole scale) is assumed not to affect the total DNA concentration. Since photocleavage is performed by mixing DNA and metal complexes at a 1:1 (v/v) ratio, the solution should be made at twice the concentration desired in the photocleavage experiment. Appropriate buffers include ammonium acetate buffer, sodium cacodylate/NaCl buffer, Tris/acetate/NaCl buffer, Tris/NaCl buffer, Tris/NaCl/imidazole buffer (see recipes), and TBE electrophoresis
6.2.26 Current Protocols in Nucleic Acid Chemistry
buffer (APPENDIX 2A). Concentrated, unlabeled, DNA stock solutions in buffer can be stored refrigerated for several weeks. Solid DNA samples are stable, either refrigerated or frozen, for several months. DNA stocks including radioactivity should be used within one half-life, as longer storage times are associated with larger amounts of nonspecific (background) DNA damage. Sonicated calf thymus DNA can be obtained commercially and dissolved in an appropriate buffer (pH 7 to 9). The concentration of a dilution of this stock solution should be determined spectrophotometrically using a value of 13,200 M−1 cm−1 per base pair at 260 nm. The stock solution can then be diluted to a concentration appropriate for its designated use. For oligonucleotides of known sequence, extinction coefficients can be estimated using additive values for each base. One such set of values includes A, 8600; C, 6800; G, 9700; and T, 8400 M−1 cm−1, and is based on the absorbance characteristics of homopolymers. A more stringent treatment is to make calculations based on each base step in the DNA, as described by Wilson et al. (1997). Once an estimate of the extinction coefficient is made, the concentration of a stock solution can be determined spectrophotometrically and appropriate dilutions can be made. In the design of an experiment, it is important to consider what will be used as the unlabeled carrier nucleic acid. In most cases this unlabeled nucleic acid is required to increase the concentration of the polymer and ensure good binding of the probe molecule. It is always best that the carrier DNA be of identical sequence to the labeled DNA strand being observed in the experiment. Therefore, in the case of oligonucleotides, extra unlabeled material is simply added to the reaction stocks. Because it is not practical to produce large amounts of plasmid restriction fragments, however, it is not always possible to use DNA of identical sequence as carrier. In that case, another commercially available unlabeled polymer is typically used. The most common alternative is sonicated calf thymus DNA because it is relatively inexpensive. It is important to consider how the selection of a carrier DNA will affect an experiment, however, because the distribution of DNA sites in one sample of DNA is unlikely to be the same as that in a specific labeled restriction fragment. Similar considerations apply to carrier RNA. In the case of run-off transcripts (where only RNA of a single sequence is made), it may be possible to use identical-sequence RNA as carrier. More likely, however, commercially available tRNA will be used as carrier.
PHOTOLYSIS OF METAL COMPLEXES Although the exact photolysis procedures that will be successful under any given set of experimental conditions depend on the individual light source used, the following description includes many of the possible options used in the authors’ laboratory and is likely to be generally adaptable. It is recommended that a new light system be appropriately tested to ensure that it is performing adequately before attempting any experimental trials. The most straightforward method for performing such tests is the photolysis of commercially available supercoiled plasmid DNA in the presence and absence of the metal complex probes. The results of the photolysis are monitored by agarose gel electrophoresis as described (see Basic Protocol 2). A properly performing lamp system should produce a large fraction of nicked and linearized plasmid DNA in the presence of metal complex and little to no damage in the absence of the probe. If insufficient photocleavage or too much complex-independent damage is observed, the light source must be modified to correct the problems. These modifications include filtering out short-wavelength UV light (<300 nm), changing the distance between the samples and the light source, and increasing the amount of time the samples are irradiated.
SUPPORT PROTOCOL 8
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.27 Current Protocols in Nucleic Acid Chemistry
After irradiation, nucleic acids can be isolated from the solutions either by drying the samples in a Speedvac evaporator or by ethanol precipitation. It is important to consider the consequences of precipitating DNA when designing a photocleavage experiment and interpreting the results. When DNA is isolated in this manner, the longest polymers are preferentially precipitated. As a result, shorter photocleavage products may be left in the supernatant and discarded. To eliminate this possible complication, it is preferable to dry samples down rather than precipitate them if the electrophoresis of the reactions is not hindered by high salt concentrations or interference of the metal complex (see Critical Parameters and Troubleshooting). Details for precipitation are given in the steps below. Materials Metal complex solution (prepared at twice the concentration desired in the photocleavage experiment; see recipe) DNA or RNA stock solution containing radiolabeled and unlabeled carrier DNA or RNA (see Support Protocol 7; prepared in buffered solution at twice the desired concentration of all components in the photocleavage experiment) 9 mM base pairs calf thymus DNA solution in deionized water, buffered to pH 7 to 9 (optional; see Support Protocol 7) 7.5 M ammonium acetate solution in deionized water (optional) Absolute ethanol (200 proof, dehydrated; optional) Dry ice Loading buffer (e.g., formamide loading buffer or urea loading buffer; see recipes) 1.7- or 0.65-mL silanized microcentrifuge tubes Light source, such as: Hg-Xe arc lamp (e.g., Oriel) equipped with an infrared (IR) filter, monochromator, and ultraviolet (UV) cut-off filter (<300 nm) He-Cd laser (e.g., Linconix model 4200 NB; 442 nm, 22 mW) Transilluminating light box (e.g., Spectroline model TR302 from Spectronics) with a broad band of irradiation centered at 302 nm Speedvac evaporator (Savant) or lyophilizer Liquid scintillation counter 1. Combine equal volumes of metal complex and DNA or RNA stock solutions in a silanized microcentrifuge tube, using a total volume of between 5 and 50 µL. Agitate the solution well to ensure complete mixing and then centrifuge the solution to the bottom of the tube. Reactions typically contain between 10,000 and 200,000 cpm of end-labeled nucleic acid fragment. Total nucleic acid concentrations (including unlabeled carrier DNA or RNA) are typically between 5 and 100 mM base pairs, depending on the binding constant of the metal complex being used and the desired ratio between the probe and nucleic acid. Additional details for nucleic acid solutions are given in the specific protocols.
2. Set up light and dark control experiments for each nucleic acid and metal complex sample used to ensure the reliability of the experimental results. For dark controls, mix metal complex with nucleic acid as in step 1. For light controls, add the same amount of nucleic acid to the tubes, but add an equal volume of deionized water instead of metal complex.
Probing Nucleic Acid Structure
Dark controls are incubated in the absence of light (i.e., are not irradiated) to assess any nuclease or other contamination. In light controls, nucleic acid is irradiated at the experimental wavelength in the absence of metal complex to measure the level of metal-independent damage.
6.2.28 Current Protocols in Nucleic Acid Chemistry
3. Allow all samples and controls to equilibrate 5 to 15 min in the dark at ambient temperature before irradiation. To ensure consistency throughout an experiment, all samples and controls should be allowed to equilibrate for the same amount of time before irradiation. As a result, if irradiations are being performed serially on a single light source, solutions should be mixed one by one as the experiment progresses.
4. Irradiate samples by positioning the open tubes in the output of a light source for between 5 and 30 min. Place the sample as close as possible to the light source or to the beam focal point of a focused source to maximize exposure to the radiation. After each sample is irradiated, store it in the dark until the experiment is completed. Typical wavelengths are 313 and 365 nm. The selection of irradiation wavelength is discussed in the individual experimental protocols. The exact irradiation time is dependent upon the strength of a given light source, the efficiency of the probe, and the wavelength used for irradiation. When a given source or wavelength is first used for an experiment, an irradiation time series should be performed to select the optimum irradiation time. For reference, a given sample irradiated at the focal point of a 313-nm beam produced by a Hg-Xe arc lamp might require 10 min for sufficient cleavage, but require two- or three-fold longer on a transilluminator. The optimal experimental irradiation time consists of the minimal time required to observe a significant level of metal-dependent cleavage. By keeping irradiation as short as possible, metal-independent light damage is minimized. The lid of the microcentrifuge tube must be open with the light entering through the mouth of the tube. Plastic absorbs strongly at most wavelengths used for these experiments. This requirement also puts an upper constraint on the length of sample irradiation time, as evaporation will occur during the course of photolysis.
5. After irradiation of all samples is completed, dry samples in a Speedvac concentrator (and proceed directly to step 8) or precipitate nucleic acids by adding 1 to 5 µL of 9 mM calf thymus DNA, 1⁄6th to 1⁄4th vol of 7.5 M ammonium acetate, and 5 vol absolute ethanol. Mix well and incubate on dry ice for 30 min. For example, for a 25-mL sample, add 1 mL calf thymus DNA, 5 mL ammonium acetate, and 125 mL absolute ethanol. Make sure to mix precipitation samples well before chilling on dry ice. If the ethanol and the buffer are not sufficiently combined, the water layer will freeze and no precipitation will occur. If complete dissociation of the metal complex is desired before electrophoresis, heat the samples 2 min at 90°C after addition of the ethanol.
6. Microcentrifuge 12 min at maximum speed (14,000 rpm). 7. Carefully remove the supernatant, rinse the resulting pellets twice with cold 80% (v/v) ethanol, and dry under vacuum. 8. Count samples in a liquid scintillation counter and resuspend in loading buffer to a constant concentration of radioactivity before electrophoresis (see Support Protocol 9).
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.29 Current Protocols in Nucleic Acid Chemistry
SUPPORT PROTOCOL 9
MAPPING CLEAVAGE SITES ON THE NUCLEIC ACID The electrophoretic techniques used in identifying nucleic acid cleavage sites are identical to those used in standard DNA sequencing. As a result, the reader is directed to sections of Current Protocols in Molecular Biology for specific procedures (see CPMB UNITS 7.1 & 7.4 and APPENDIX 3B of this book). In the authors’ laboratory, dried or precipitated photocleavage samples are resuspended in denaturing loading buffer (e.g., formamide loading buffer or urea loading buffer; see recipes) and electrophoresed in TBE electrophoresis buffer (APPENDIX 2A) on 8% to 20% (w/v) denaturing polyacrylamide gels cast from commercially available stock solutions (National Diagnostics). The polyacrylamide gel is pre-run until it is warm to the touch. In addition, the samples are heated to 90°C for 5 min before loading to promote denaturation of the nucleic acid strands and dissociation of the metal complex probes. To identify the specific sites of nucleic acid cleavage, photocleavage samples are run alongside samples of the same polymers that have been chemically sequenced using standard MaxamGilbert sequencing reactions (Maxam and Gilbert, 1980; e.g., CPMB UNIT 7.5). Gels are read using standard techniques of autoradiography or phosphorimagery. Because both Maxam-Gilbert sequencing reactions and metal complex photocleavage result in the loss of a base at the cleavage site, chemical sequencing products comigrate with analogous photocleavage products. Enzymatic sequencing methods, which do not involve base loss, will result in a systematic one-base shift in the positioning of the cleavage bands. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Ammonium acetate buffer, 10× 19.3 g/L ammonium acetate (250 mM) Adjust to pH 9 with NH4OH and acetic acid Sterilize by filtering with a 0.22-µm filter Store up to several months at room temperature Formamide loading buffer 80% (v/v) deionized formamide 50 mM Tris base 50 mM boric acid 1 mM EDTA 0.025% (w/v) xylene cyanol 0.025% (w/v) bromphenol blue Adjust to pH 8.3 Store for up to several months at room temperature In the authors’ experience, this buffer performs better than the urea loading buffer for resuspending samples after drying or precipitation.
Probing Nucleic Acid Structure
Metal complex solution Prepare all metal complex solutions as concentrated stocks in deionized water from a sample of solid reagent immediately before use. Determine the concentration of the stock solution spectrophotometrically using the extinction coefficients provided in the Commentary (see Compound Characterization) and then make dilutions in deionized water to concentrations appropriate for the experiments. Store solutions in the dark for the duration of a single experiment and discarded immediately after use.
6.2.30 Current Protocols in Nucleic Acid Chemistry
Sodium cacodylate/NaCl buffer, 10× 15.9 g/L sodium cacodylate (100 mM) 23.4 g/L sodium chloride (400 mM) Adjust to pH 7 with NaOH and cacodylic acid Sterilize by filtering with a 0.22-µm filter Store up to several months at room temperature Tris/acetate buffer, 10× Prepare as for Tris/acetate/NaCl buffer (see recipe) but omit NaCl. Tris/acetate/NaCl buffer, 10× 60.6 g/L Tris base (500 mM) 16.4 g/L sodium acetate (200 mM) 10.5 g/L sodium chloride (180 mM) Adjust to pH 7 with NaOH and acetic acid Sterilize by filtering with a 0.22-µm filter Store up to several months at room temperature Tris/NaCl buffer, 10× 6.1 g/L Tris base (50 mM) 29.2 g/L sodium chloride (500 mM) Adjust to pH 7 with NaOH and HCl Sterilize by filtering with a 0.22-µm filter Store up to several months at room temperature Tris/NaCl/imidazole buffer, 10× 6.1 g/L Tris base (50 mM) 29.2 g/L sodium chloride (500 mM) 0.82 g/L imidazole (12 mM) Adjust to pH 7 with NaOH and HCl Sterilize by filtering with a 0.22-µm filter Store up to several months at room temperature Urea loading buffer 7 M urea 90 mM Tris base 90 mM boric acid 2.5 mM EDTA 0.025% (w/v) xylene cyanol 0.025% (w/v) bromphenol blue Adjust to pH 8.3 Store for up to several months at room temperature Refrigeration of this buffer is not recommended as the urea tends to crystallize out. In the authors’ experience, the formamide-based buffer performs better for resuspending samples after drying or precipitation.
COMMENTARY Background Information Recent results in NMR structure determination and X-ray crystallography of nucleic acids (Hartmann and Lavery, 1996) have shown that the structure of correctly matched, putatively canonical B-form DNA can be heterogeneous and polymorphic. When the additional helical
forms and the many possible DNA lesions (e.g., mismatches, bulges, abasic sites, DNA-drug adducts, and sites of base damage) are added, it is clear that what was once thought to be an uninteresting linear polymer can hide a wealth of structural variety. When the field of consideration is expanded to include RNA, whose
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.31 Current Protocols in Nucleic Acid Chemistry
Probing Nucleic Acid Structure
folding pathways and structures rival those of proteins in complexity, and the many heteroduplexes that now exist between DNA and other polymers, the structural determinations of nucleic acids becomes a very rich field indeed. Because of the time and resources that are involved in techniques for high-resolution structural determination, solution probes of nucleic acids have a very important role in the pursuit of a broad-based structural understanding of DNA and RNA. Unhindered by the concerns of size and system selection that sometimes make NMR difficult, and the need for crystals to determine a structure in the solid state, photochemical probes like those described here provide a rapid, relatively inexpensive method to probe the characteristics of DNA or RNA in solution. Furthermore, for larger macromolecules in particular, chemical probe studies can serve as a bridge to high-resolution structural information. The specific transition metal complex probes described in this unit therefore help fill a critical niche in the large body of chemical and photochemical probes for nucleic acid structure. Residing between probes that have been designed to identify specific sequences along a DNA or RNA strand or helix and the large body of chemical probes that cleave nucleic acids more nonspecifically at primarily solvent-accessible sites, it is the structure specificity of these complexes that makes them unique. The three-dimensional shapes of the complexes, which lead to their structural recognition properties, allow the identification of specific geometries without consideration of base sequence. One of the most well-defined rhodium complexes that has been used for a wide variety of purposes, [Rh(phen)2(phi)]3+, binds in the major groove of DNA by intercalation of the phi ligand between adjacent base pairs (Pyle et al., 1989). DNA strand cleavage is effected by direct hydrogen atom abstraction by the photoexcited phi ligand at the site of intercalation (Sitlani et al., 1992). The complex recognizes primarily two families of sites: 5′-pyrimidinepyrimidine-purine-purine-3′ and homopyrimidine-homopurine tracts. The two enantiomers of the complex demonstrate different recognition behaviors. The ∆ enantiomer, whose chirality is complementary to the right-handed DNA helix, favors the pyrimidine-pyrimidinepurine-purine tracts (Sitlani and Barton, 1994). Site selectivity of this complex is based on shape selection by the bulky aromatic phenanthroline ligands. Comparisons of crystallographic data on the sequences targeted by the
complex show that they are more open in the major groove, thus allowing both the stacking, intercalative interactions of the phi ligand and accommodation of the phen ancillary ligands. Enantioselectivity in cleavage correlates with the opening of sites and changes in the propeller twisting of the bases involved (Campisi et al., 1994). When [Rh(phen)2(phi)]3+ is applied to ribonucleic acids, the folding of complex RNA molecules can be assessed; in an application that crosses over the boundaries between the two types of nucleic acids, the same complex has been used to examine the structure of tDNAPhe (the DNA analog of tRNAPhe) to assess the similarities or differences between DNA copies and their RNA counterparts (Lim and Barton, 1993). The other molecule used as a geometrical probe of unknown DNA sequences, [Ru(TMP)3]2+, has very different behavior. It does not bind by intercalation but rather to the surface of the minor groove of DNA. The complex binds avidly to A-form helices and exhibits little binding to B- or Z-form synthetic oligonucleotides. There is some level of chiral discrimination in binding that favors the Λ-isomer of the complex (Mei and Barton, 1986). The basis of the binding selectivity of [Ru(TMP)3]2+, like [Rh(phen)2(phi)]3+, rests in shape selection, although in this case generated by a different type of interaction. The sterically bulky methyl groups on the TMP ligands not only prevent intercalation, which is observed for the unmethylated [Ru(phen)3]2+ parent complex, but also restrict binding to open and shallow areas of the minor groove. This explains the preferential binding to A-form DNA and RNA by [Ru(TMP)3]2+. In contrast to the direct photocleavage found with [Rh(phen)2(phi)]3+, where the cleavage chemistry involves reaction with the deoxyribose sugar and shows no sequence bias, irradiation of the ruthenium complexes sensitizes the formation of singlet oxygen, a diffusible species that reacts with nearby bases and preferentially with guanine residues (Mei and Barton, 1988). As a result, [Ru(TMP)3]2+ cleavage must be compared with [Ru(phen)3]2+, its sequence-neutral analog, to deduce areas of preferential TMP binding. The combination of [Rh(phen)2(phi)]3+ and [Ru(phen)3]2+ probes have been applied to mapping the conformation of DNA within an entire gene (Huber et al., 1991). The shape and structure specificity that is provided by simple steric exclusion has also been used for a variety of recognition tasks. The ability of [Rh(DIP)3]3+ to recognize cruciforms
6.2.32 Current Protocols in Nucleic Acid Chemistry
and other unusual tertiary structures is a relatively unique property that has been comprehensively explored in the authors’ laboratory. Experiments have shown that it is the cruciform structure, rather than any specific sequence, that is targeted by the molecule (Kirshenbaum et al., 1988). The molecule’s shape selectivity has also been applied to an examination of the DNA structures in the SV40 T antigen and adenovirus 2 E1A genes (Lee and Barton, 1993). This steric specificity mechanism has been applied to the design of the intercalating ligand [Rh(bpy)2(chrysi)]3+, which has been used to probe both the structures of DNA base mismatches (Jackson and Barton, 1997; Jackson et al., 1999) and abasic sites in DNA (B.A. Jackson and J.K. Barton, unpub. observ.). A comprehensive study of this molecule and its recognition behavior is currently in progress in the authors’ laboratory. When these probes are used to examine RNA structure, their unique properties lead to additional applications. From the studies performed on tRNAPhe, the authors’ laboratory has shown that the A-form configuration of RNA leads to much more specific recognition of ribonucleic acid structural features by this family of rhodium probes. The complex recognizes sites of tertiary structure, including bulges and triplex sites where the phi intercalating ligand can gain access to the major groove of the nucleic acid. It is becoming clear that RNA binding proteins also target sites that are opened in the major groove. As a result, use of [Rh(phen)2(phi)]3+ may find application in scanning unknown ribonucleic acids for possible protein-binding sites. The cleavage behavior of [Rh(phen)2(phi)]3+ has been used to examine structural domains in eukaryotic 5S rRNA (Chow et al., 1992) and to probe the conformations of TAR RNAs from human (Neenhold and Rana, 1995) and bovine (Lim and Barton, 1997) immunodeficiency viruses. Moving away from modes of structural recognition, the nonspecific [Rh(phi)2(bpy)]3+ complex is an important addition to the body of molecular tools for studying the interactions between DNA and other molecules. Because its DNA cleavage chemistry is photoinduced, it provides a unique opportunity to study systems where it is advantageous to combine all the components at one time and activate the footprinting reaction later. This compound has been used in the authors’ laboratory to footprint both DNA-binding proteins (like EcoRI) and small DNA binders (like distamycin; Uchida et al., 1989). Furthermore, [Rh(phi)2(bpy)]3+ is
unaffected by additives such as glycerol and Mg2+ at the levels that might be required to promote DNA binding by certain proteins. It can footprint both major and minor groove ligands, and, since no diffusible intermediate is involved in the cleavage, produces sharp, wellresolved footprint patterns. In spite of the breadth and number of tasks to which these reagents have been applied in the past, the many open questions about nucleic acid structure provide myriad opportunities where solution probes like those described here can be used. As recent crystal structures of DNA-protein complexes have made clear, the shape and geometry of a nucleic acid target site can play an instrumental role in specific DNAprotein interactions. As more is discovered about the unique structures of damaged or mispaired sites in DNA and the dramatic effects that DNA-binding proteins and small molecules can have on DNA geometry, the easily accessible information that can be provided by solution probes of structure becomes more and more valuable.
Compound Characterization Mapping DNA major and minor groove characteristics [Rh(phen)2(Cl2)]Cl 1H-NMR: d -dmso, 9.69 (d, 2H); 8.99 (d, 6 2H); 8.89 (d, 2H); 8.62 (t, 2H); 8.32 (t, 2H); 8.18, (t, 2H); 7.80 (d, 2H); 7.59 (t, 2H). [Rh(phen)2(NH3)2]Cl3 1H-NMR: d -acetone, 9.93 (d, 2H); 9.42 (d, 6 2H); 8.99 (d, 2H); 8.64 (m, 4H); 8.51 (d, 2H); 8.32 (d, 2H); 7.87 (dd, 2H); 5.18 (broad s, 6H). 13C-NMR: d -acetone, 154.1, 153.6, 147.6 6 q, 146.6 q, 142.9, 141.5, 133.2 q, 129.2, 129.1, 128.3, 127.7. UV/visible (acetonitrile/water 1:1, 1.9 × 10− −1 −1 −1 5 M): ε 302, 12,400 M cm , ε271, 51,500 M −1 −1 −1 cm , ε222, 51,800 M cm . [Rh(phen)2(phi)]Cl3 1H-NMR: d -dmso, 14.98 (s, N-H phi); 9.25 6 (d, H 4 phen); 9.12 d, H 2 phen); 9.03 (d, H 7 phen); 8.94 (d, H 5,6 phen); 8.54 (dd, H 4,5 phi); 8.47 (d, H 1,8 phi); 8.36 (dd, H 3 phen); 8.08 (d, H 9 phen); 7.91 (dd, H 8 phen); 7.84 (t, H 2,7 phi); 7.57 (t, H 3,6 phi). UV/visible: ε362, 19,400 M−1 cm−1 (pH isosbestic point); ε358, 19,400 M−1 cm−1; ε300, 34,000 M−1 cm−1; ε272, 116,200 M−1 cm−1.
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.33 Current Protocols in Nucleic Acid Chemistry
CD: (∆ enantiomer, acetonitrile, 1.47 × 10−5 M) 261 (99), 275 (-179), 346 (-24). [Ru(TMP)3]Cl2 1H-NMR: CDCl , 8.20 (s, H 5,6); 7.98 (s, 3 H, 2,9); 2.74 (s, CH3, 4,7); 2.37 (s, CH3, 3,8). UV/visible: ε210, 94,500 M−1 cm−1; ε233, 64,000 M−1 cm−1; ε269, 114,600 M−1 cm−1; ε299(shoulder), 26,400 M−1 cm−1; ε438, 24,500 M−1 cm−1. [Ru(phen)3]Cl2 1H-NMR: d -dmso, 8.8 (d, H 4,7); 8.7 (d, H 6 2,9); 8.4 (s H 5,6); 7.7 (dd, H 3,8). UV/visible: ε222, 87,600 M−1 cm−1; ε262, 107,000 M−1 cm−1; ε291(shoulder), 22,500 M−1 cm−1; ε447, 19,000 M−1 cm−1. Shape-selective cleavage of unusual structures in nucleic acids
Photofootprinting of DNA-binding molecules [Rh(phi)2Cl2]Cl 1H-NMR: d -dmso, trans product: 14.89 (s, 6 4H); 8.75 (d, 4H); 8.53 (d, 4H); 7.92 (t, 4H); 7.82 (t, 4H); cis product, 14.53 (s, 2H); 13.19 (s, 2H); 9.23 (d, 2H); 8.5 (t, 4H); 8.37 (d, 2H); 7.95 (d, 2H); 7.8 (d, 2H); 7.75 (t, 2H); 7.58 (t, 2H).
[Rh(DIP)3]Cl3 1H-NMR: CDCl , 9.01 (d, 6H); 8.30 (s, 6H); 3 8.07 (s, 6H); 7.71-7.69 (m, 10H); 7.61-7.53 (m, 20H). UV/visible: ε296, 116,000 M−1 cm−1; ε334 (shoulder), 49,200 M−1 cm−1; ε370, 19,400 M−1 cm−1.
[Rh(phi)2(bpy)]Cl3 1H-NMR: d -dmso, 14.13 (s, 2H, N-H); 6 13.63 (s, 2H, N-H); 8.92 (d, 2H); 8.66 (m, 6H); 8.45 (d, 6H); 7.84 (m, 6H); 7.61 (t, 4H). UV/visible (Tris-acetate, pH 7.0): ε250, 67,400 M−1 cm−1; ε262, 62,400 M−1 cm−1; ε270, 64,200 M−1 cm−1; ε292, 43,200 M−1 cm−1; ε312, 30,000 M−1 cm−1; ε378, 28,200 M−1 cm−1; ε350, 23,600 M−1 cm−1 (pH isosbestic point).
Recognition of destabilized structures in DNA
Critical Parameters and Troubleshooting
5,6-chrysenequinone 1H-NMR: CD Cl , 9.39 (d, 1H); 8.16 (m, 2 2 4H); 7.92 (d, 1H); 7.77 (t of d, 2H); 7.57 (t of d, 2H).
Syntheses of transition metal probes Other than the general safety concerns involved with working with flammable liquids and strong acids, most of the synthetic work described in this unit has few concerns. In those reactions that require vacuum lines and evacuation of glassware, care must be taken to select equipment that can withstand the required changes in pressure, and appropriate protective eyewear and a laboratory coat should be worn. All reactions, even those that do not specifically mention it, should be performed in a well-functioning fume hood whenever possible. In the authors’ hands, the reactions described in this work have performed to produce their intended products in reliable yields. In all cases, if a reaction is not performing as it should, the correct course of action is to assess the identity and purity of all starting materials and reagents and try the reaction again.
[Rh(bpy)2(Cl2)]Cl 1H-NMR: d -dmso, 9.69 (d, 2H); 9.0 (d, 6 2H); 8.89 (d, 2H); 8.63 (t, 2H); 8.33 (t, 2H); 8.17 (t, 2H); 7.80 (d, 2H); 7.59 (t, 2H). [Rh(bpy)2(NH3)2]Cl3 1H-NMR: d -acetone, 9.45 (d, 2H); 9.05 (d, 6 2H); 8.89 (d, 2H); 8.79 (td, 2H); 8.45 (td, 2H); 8.30 (td, 2H); 8.05 (d, 2H); 7.74 (td, 2H); 5.06, (broad s, 6H).
Probing Nucleic Acid Structure
135.7, 132.3, 132.2, 132.0, 131.8, 130.9, 130.8, 130.6, 130.2, 129.4, 127.7, 127.6, 127.3, 126.9, 126.8, 123.9, 122.9, 120.8, 118.6, 116.3, 114.1. UV/visible (water, 7.8 × 10−6 M): ε271, 63,800 M−1 cm−1 (pH isosbestic point); ε303, 57,000 M−1 cm−1; ε315, 52,200 M−1 cm−1; ε391, 10,600 M−1 cm−1.
[Rh(bpy)2(chrysi)]Cl3 1H-NMR: CD OD, 8.94 (t, 2H); 8.86 (t, 2H); 3 8.80 (d, 1H); 8.77 (d, 1H); 8.56 (split t, 2H); 8.44 (m, 5H); 8.40 (d, 1H); 8.15 (m, 1H); 8.03 (m, 1H); 7.95, (m, 3H); 7.86, (d, 1H); 7.81, (d, 1H); 7.64, (m, 5H). 13C-NMR: CD OD, 183.3, 177.3, 175.4, 3 157.4, 157.2, 157.2, 153.8, 153.2, 152.1, 144.6, 144.5, 143.9, 143.8, 139.7, 138.8, 138.4, 136.2,
DNA photocleavage experiments To achieve success with any of the protocols for assaying DNA structure, all reagents must be of the highest possible purity. The DNA or
6.2.34 Current Protocols in Nucleic Acid Chemistry
RNA fragments in use must be scrupulously purified. In the case of restriction fragments and RNA transcripts, the nucleic acids must be purified by polyacrylamide gel electrophoresis, and the smallest possible gel band containing the labeled fragment should be excised from the gel. After the labeled polymer is removed from the gel, it must be purified from the various components that were present in the electrophoresis buffers. This should be done by ethanol precipitation or by using one of the many commercially available reversed-phase purification cartridges. In the case of synthetic oligonucleotides, the synthesized materials must be purified by HPLC and, after end labeling, also purified by either denaturing or nondenaturing PAGE. In the authors’ laboratory, the last trityl protecting group is typically left at the end of DNA synthesis and used as a hydrophobic handle for HPLC purification. After removing the protecting group, a second HPLC run is performed to ensure purity. In both cases, only the top portion of the major product peak is collected. If an experiment is run with insufficiently pure nucleic acid, the most obvious symptom is an absence of clear, clean banding in all experimental lanes. Such behavior will also be observed in the Maxam-Gilbert sequencing reactions. The metal complexes used for photolysis reactions must also be of the highest possible purity. If they are contaminated by synthetic side products or other material, the concentrations of prepared solutions will be incorrect and other reactivity (unrelated to the desired rhodium or ruthenium probe) may be observed. It is customary in the authors’ laboratory to HPLC purify all metal complexes after ion-exchange chromatography or enantiomer resolution to ensure their purity. If desired, enantiomeric purity can be verified by CD spectroscopy or by use of a commercially available chiral HPLC column. Furthermore, even if metal solutions are made from analytically pure material, it is critical that the solutions used in photocleavage be made as close as possible to the time the experiments are performed. With extended storage in solution, especially with exposure to light, the various metal complex probes can degrade. The degradation products may interact with or cleave DNA, with different specificities than that of the intended probes. All buffer solutions should be made with the highest purity materials available, treated carefully to ensure continuing purity, and remade frequently. In the authors’ laboratory it is also customary to filter all solutions through 0.22-
µm sterilizing filter units before use. Nuclease contamination in buffers can occur through environmental factors and can result in observed cleavage that is independent of irradiation. This is of particular concern with work relating to RNA. Other contaminants in stock solutions can lead to light-dependent damage to the target DNA as well. All the concerns regarding reagent and solution purity underscore the importance of running control samples as part of each photocleavage experiment. These must include both dark controls (DNA incubated with a metal complex probe in the absence of light) and light controls (DNA irradiated in the absence of metal complex). Without these important comparisons, it is impossible to assess the value of any given experimental result. Furthermore, even with the best techniques and intentions, single samples sometimes become contaminated and give anomalous results. This underscores the importance of repeating trials when experimental or control samples display unexpected behavior. Depending on the light source that is available for metal complex irradiation and the specific experimental plan, some troubleshooting of each new system must be done to obtain optimal results with these DNA probes. In all cases, if insufficient cleavage is observed in a given experiment, appropriate responses are to increase the concentrations of the complexes and DNA to promote more complete binding, increase the irradiation times for the experimental samples, and, depending on the sensitivity of the autoradiography technique being employed, increase the amount of radioactivity loaded in each gel lane. On the other hand, if too much photocleavage is observed, one can no longer trust the data extracted from a DNA photocleavage experiment, because at high levels of cleavage there is no reasonable guarantee that single-hit conditions prevail (i.e., that each cleaved polymer has been cut only once by a single metal complex). In the authors’ laboratory, excessive cleavage is defined as an experiment where >10% of the full-length polymer has been cleaved. Under multiple-hit conditions, cleavage bands that correspond to shorter polymers will be enriched at the expense of longer cleavage products. In this case, the exact opposite course of action is required as when too little cleavage is observed (reduce metal complex concentration or decrease irradiation times). In addition to these general concerns for all photocleavage protocols, a few procedures
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.35 Current Protocols in Nucleic Acid Chemistry
Table 6.2.1 Photocleavage Experiment Troubleshooting Guide
Problem Gels do not show clean banding, including in Maxam-Gilbert sequencing lanes
Possible cause Contamination of desired labeled DNA by fragments or shorter oligonucleotide synthetic products produces labeled DNA sample of heterogeneous length
Unexpected cleavage bands observed in experimental lanes but not light or dark controls
Damage is dependent on the addition of the Purify metal complex again by ion exchange or HPLC metal solution and irradiation. Likely contamination of the metal solution by a synthetic by-product or breakdown product
Likely cause is a contaminant in one of the Unexpected cleavage bands observed in experimental samples buffer solutions or solvents used in making and light control but not dark control up the experimental samples that causes light-induced damage to the nucleic acid.
Solution Purify DNA more stringently before running the experiment to ensure homogeneity
Prepare fresh buffer solutions and stocks from new materials of known purity
This behavior has also been observed when Use silanized microcentrifuge tubes nonsilanized tubes are used for irradiation reactions. Presumably some component in the plastic leaches into solution in sufficient concentration to promote light-induced DNA damage Damage is dependent on addition of metal Prepare fresh metal solutions from Unexpected cleavage bands observed in experimental lanes and solution but not on irradiation. Likely cause new materials of known purity is contamination of the metal solution by a dark control but not light control nuclease of some type Unexpected cleavage bands observed in experimental lanes, dark control, and light control
Likely cause is contamination of DNA stock Prepare fresh DNA and metal (or buffer/water stocks) by a nuclease that solutions from new materials of cleaves DNA independent of irradiation known purity
Damage observed in dark control Likely due to exposure of dark control to that is similar to experimental lanes ambient lighting
Keep samples dark before and after irradiation
Smearing observed in experimental Most likely due to problems with lanes but not in sequencing or light renaturation of nucleic acid strands or control interaction between the metal complex and nucleic acids
Heat samples to 90°C (5 min) before loading gel and pre-run the gel until very hot to ensure strand denaturation. If necessary, reprecipitate samples to remove metal complex.
Bands of interest too high or low on Gel run for inappropriate time the gel for adequate clarity
Run gel longer or shorter to affect position of bands in gel. For best resolution of cleaved bases and different polymer ends, bands of interest should be in the lower third of the gel.
Band intensity varies dramatically from gel lane to gel lane, specifically in the uncleaved DNA band
Gel lanes were not loaded with the same amount of radioactivity
Ensure samples are resuspended well and at the same concentration of radioactivity
continued
Probing Nucleic Acid Structure
6.2.36 Current Protocols in Nucleic Acid Chemistry
Table 6.2.1 Continued
Problem Possible cause High concentrations of salt in samples As bands progress down the gel, they compress towards the center of loaded on the gel can affect the mobility of the DNA the lane or run anomalously compared to Maxam-Gilbert sequencing reactions
Solution Reduce the effect/amount of loaded salt by: (1) precipitating samples with ammonium acetate and drying thoroughly under vacuum, (2) increasing the amount of radioactivity in each irradiation sample so a smaller fraction of the sample is loaded on the gel, (3) running the gel at a very low wattage (25 watts) for 30-60 min at the beginning of a gel run to allow some salt to separate from DNA, or (4) reducing the volume of irradiation samples to reduce the amount of salt that is dried down into the DNA
Insufficient DNA cleavage observed Concentration may be too low for effective in experimental lanes binding of complexes to DNA
Increase concentration of both metal solution and DNA
Ratio of total DNA concentration to the Increase concentration of metal solution metal complex probe may be too high to cleave an observable fraction of the labeled DNA Light source may be too weak to promote enough DNA cleavage
Increase power of the light source or increase the irradiation time per sample Amount of radioactivity loaded per gel lane Increase amount of labeled DNA may be too low for sensitivity of the loaded per lane autoradiography technique
Too much cleavage, resulting in >10% depletion of the full-length DNA band
Too much metal complex in the reaction for Reduce concentration of metal a given reaction time, resulting in extensive complex in the solution or reduce cleavage. This will likely involve more than the irradiation time one cleavage event per DNA polymer making the data unreliable
have more specific considerations. In particular, experiments with [Rh(DIP)3]3+ must control carefully for the superhelicity dependence of cleavage and whether the cleavage is single or double stranded. In addition, it must be carefully noted that in the absence of an unusual structure and at long irradiation times [Rh(DIP)3]3+ does induce nonspecific oxidative damage at 5′-GG-3′ and 5′-GA-3′ sites. If this behavior is not adequately controlled, incorrect conclusions may be drawn from observed cleavage data. In a related manner, when using [Rh(phi)2(bpy)]3+ for DNA footprinting, the binding behavior of the metal complex must be considered to ensure that experimental results are meaningful. The concentration of the probe
must be kept high to ensure nonspecific binding. Irradiation times must be kept short enough that there are not problems with excessive cleavage at these high loadings. Finally, one must also consider that the binding constant of rhodium phi intercalators for DNA is in the range of 106 to 108 M−1, making it possible for them to displace weaker DNA-binding molecules from the polymer. A troubleshooting guide for photocleavage experiments is given in Table 6.2.1.
Anticipated Results Using the numerous reagents and techniques described in these protocols, questions about the structure of an unknown nucleic acid sequence or lesion can be approached. In all
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.37 Current Protocols in Nucleic Acid Chemistry
cases, rapid and relatively inexpensive techniques lead to identification of binding sites on the polymer of interest and direct information about the geometry and conformations of the molecule in solution. Because of the large variety of assays and probes included in this unit, it is more efficient to direct the reader to those original papers that include examples of experimental results than to reproduce examples here. Appropriate citations are included below (see Key References).
Time Considerations Once the apparatus has been collected and is in place, most of the synthetic reactions described in this protocol can be completed within a single day. There are a few exceptions to this, including the synthesis of complexes involving 9,10-diaminophenanthrene. These procedures can require between a few days and a week, although the amount of hands-on time during these periods is minimal. Complex purification columns and enantiomer separation require an additional 1 day to 2 weeks of chromatography time. However, these time investments must be viewed with the knowledge that a single synthetic cycle will produce enough material for several hundred DNA structure experiments. Once a given metal complex of interest is made and purified, the biological experiments proceed much more rapidly. DNA can be made and purified within 3 days, and labeled and isolated in another day. Irradiation reactions can be completed in 2 to 12 hr, depending on the number of samples involved. Electrophoresis and autoradiography on one set of samples requires an additional 1 to 2 days.
Acknowledgments The work described here was supported by the NIH (GM33309) and performed by many able graduate students who are referenced as indicated. B.A.J. would also like to acknowledge the NSF for a predoctoral fellowship.
Literature Cited
Probing Nucleic Acid Structure
Campisi, D., Morii, T., and Barton, J.K. 1994. Correlations of crystal structures of DNA oligonucleotides with enantioselective recognition by [Rh(phen)2(phi)]3+: Probes of DNA propeller twisting in solution. Biochemistry 33:41304139.
Cartwright, P.S., Gillard, R.D., and Sillanpåå, E.R.J. 1987. Optically active coordination compounds—XLVI. Resolution of tris-diimmine compounds of chromium(III) using fac(+)tris[L-cysteinesulphinato(2-)SN]cobaltate (III). Polyhedron 6:105-110. Chow, C.S. and Barton, J.K. 1992. Recognition of G-U mismatches by tris(4,7-diphenyl-1,10phenanthroline)rhodium(III). Biochemistry 31:5423-5429. Chow, C.S., Behlen, L.S., Uhlenbeck, O.C., and Barton, J.K. 1992. Recognition of tertiary structure in tRNAs by [Rh(phen)2(phi)]3+, a new reagent for RNA structure-function mapping. Biochemistry 31:972-982. Dollimore, L.S. and Gillard, R.D., 1973. Optically active co-ordination compounds. Part XXXII. Potassium (+) tris-[L-cysteinesulphinato(2-)SN]cobaltate(III): A versatile agent for resolution of 3+ species. J. Chem. Soc. Dalton Trans. (1973):934-940. Gidney, P.M., Gillard, R.D., and Heaton, B.T. 1972. 1,10-Phenanthroline and 2,2′-bipyridyl complexes of rhodium(III). J. Chem. Soc. Dalton Trans. (1972):2621-2628. Gillard, R.D., Osborn, J.A., and Wilkinson, G. 1965. Catalytic approaches to complex compounds of rhodium(III). J. Chem. Soc. Dalton Trans. (1965):1951-1965. Greabe, V.C. and Hönisberger, F. 1900. Ueber die Oxydationsproducte des Chrysens. Ann. Chem. 311:257-265. Hall, D.B., Holmlin, R.E., and Barton, J.K. 1996. Oxidative DNA damage through long range electron transfer. Nature 382:731-735. Hartmann, B. and Lavery, R. 1996. DNA structural forms. Q. Rev. Biophys. 29:309-368. Howells, R.D. and McCown, J.D. 1977. Trifluormethanesulfonic acid and derivatives. Chem. Rev. 77:69-92. Huber, P.W., Morii, T., Mei, H.-Y., and Barton, J.K. 1991. Structural polymorphism in the major groove of a 5S RNA gene complements the zinc finger domains of transcription factor IIIA. Proc. Natl. Acad. Sci. U.S.A. 88:10801-10805. Jackson, B.A. and Barton, J.K. 1997. Recognition of mismatches by a rhodium intercalator. J. Am. Chem. Soc. 199:12986-12987. Jackson, B.A., Alekseyev, V.A., and Barton, J.K. 1999. A versatile recognition agent: Specific cleavage of a plasmid DNA at a single base mispair. Biochemistry 38:4655-4662. Kirshenbaum, M.R., Tribolet, R., and Barton, J.K. 1988. [Rh(DIP)3]3+: A shape-selective metal complex which targets cruciforms. Nucl. Acids Res. 16:7943-7960. Lee, I. and Barton, J.K. 1993. A distinct intronDNA structure in simian virus 40 T-antigen and a d e n ov i r u s 2 E 1 A g e nes. Biochemistry 32:6121-6127.
6.2.38 Current Protocols in Nucleic Acid Chemistry
Lim, A.-C. and Barton, J.K. 1993. Chemical probing of tDNAPhe with transition metal complexes: A structural comparison of RNA and DNA. Biochemistry 32:11029-11034. Lim, A.-C. and Barton, J.K. 1997. Targeting the Tat-binding site of bovine immunodeficiency virus TAR RNA with a shape-selective rhodium complex. Bioorg. Med. Chem. 5:1131-1136. Lin, C-T., Böttcher, W., Chou, M., Creutz, C., and Sutin, N. 1976. Mechanism of the quenching of the emission of substituted polypyridineruthenium(II) complexes by iron(III), chromium(III), and europium(III) ions. J. Am. Chem. Soc. 98:6536-6544. Maxam, A. and Gilbert, W. 1980. Sequencing endlabeled DNA with base-specific chemical cleavages. Methods Enzymol. 65:499-560. Mei, H.Y. and Barton, J.K. 1986. Chiral probe for A-form helices of DNA and RNA: Tris(tetramethylphenanthroline)ruthenium(II). J. Am. Chem. Soc. 108:7414-7416. Mei, H.Y. and Barton, J.K. 1988. Tris(tetramethylphenanthroline)ruthenium(II): A chiral probe that cleaves A conformations. Proc. Natl. Acad. Sci. U.S.A. 85:1339-1343. Müller, B.C., Raphael, A.L., and Barton, J.K. 1987. Evidence for altered DNA conformations in the simian virus genome: Site-specific DNA cleavage by the chiral complex Λ-tris(4,7-diphenyl1,10-phenanthroline)cobalt(III). Proc. Natl. Acad. Sci. U.S.A. 84:1764-1768. Mürner, H., Jackson, B.A., and Barton, J.K. 1998. A versatile synthetic approach to rhodium(III) diimine metallointercalators: Condensation of o-quinones with coordinated cis-ammines. Inorg. Chem. 37:3007-3012. Neenhold, H.R. and Rana, T.M. 1995. Major groove opening at the HIV-1 Tat-binding site of TAR RNA evidenced by a rhodium probe. Biochemistry 34:6303-6309. Pyle, A.M., Long, E.C., and Barton, J.K. 1989. Shape-selective targeting of DNA by (phenanthrenequinone)rhodium(III) photocleaving agents. J. Am. Chem. Soc. 111:4520-4522. Pyle, A.M., Chiang, M.Y., and Barton, J.K. 1990. Synthesis and characterization of physical, electronic, and photochemical aspects of 9,10-phenanthrenequinone diimine complexes of ruthenium(II) and rhodium(III). Inorg. Chem. 29:4487-4495. Sitlani, A. and Barton, J.K. 1994. Sequence-specific recognition of DNA by phenathrenequinone diimine complexes of rhodium(III): Importance of steric and van der Waals interactions. Biochemistry 33:12100-12108. Sitlani, A., Long, E.C., Pyle, A.M., and Barton, J.K. 1992. DNA photocleavage by phenanthrenequinone diimine complexes of rhodium(III): Shape selective recognition and reaction. J. Am. Chem. Soc. 114:2303-2312.
Uchida, K., Pyle, A.M., Morii, T., and Barton, J.K. 1989. High resolution footprinting of EcoRI and distamycin with [Rh(phi)2(bpy)]3+, a new photofootprinting reagent. Nucl. Acids Res. 17:10259-10279. Wilson, W.D., Tanious, F.A., Fernandez-Saiz, M., and Rigl, C.T. 1997. Evaluation of drug-nucleic acid interactions by thermal melting curves. In Methods in Molecular Biology, Vol. 90: DrugDNA Interaction Protocols (K.R. Fox, ed.) pp. 219-240. Humana Press, Totowa, N.J. Yoshikawa, Y. and Yamasaki, K. 1979. Chromatographic resolution of metal complexes on Sephadex ion exchangers. Coord. Chem. Rev. 28:205-229.
Key References Campisi et al., 1994. See above. Examples of mapping the major and minor grooves of DNA with [Rh(phen)2(phi)]3+ and [Ru(TMP)3]2+. Chow et al., 1992. See above. Examples of mapping RNA structure using rhodium probes. Hartmann and Lavery, 1996. See above. This is an excellent review of recent work in DNA structural determinations of A-, B-, and Z-form DNA, mismatches, abasic sites, and bulges. As such, it provides an excellent overview of many of the targets these metal complex probes can be used to elucidate. Jackson and Barton, 1997. See above. Examples of using [Rh(bpy)2(chrysi)]3+ as a probe for DNA mismatches. Kirshenbaum et al., 1988. See above. Examples of site-selective cleavage of unusual structures in nucleic acids using [Rh(DIP)3]3+. Lim and Barton, 1997. See above. Example of [Ru(phen)3]2+ used as a guanine-specific sequencing reagent for nucleic acids. Uchida et al., 1989. See above. Examples of footprinting both major groove– and minor groove–binding molecules using rhodium probes.
Contributed by Brian A. Jackson and Jacqueline K. Barton California Institute of Technology Pasadena, California
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.2.39 Current Protocols in Nucleic Acid Chemistry
Probing RNA Structure by Lead Cleavage
UNIT 6.3
This unit describes the probing of RNA structure using Pb(II) ions. The Pb(II) hydroxide species has a relatively low pK a (∼7.7), which allows it to extract the proton from a 2′-hydroxyl group. In RNA, abstraction of a 2′ proton increases the nucleophilicity of the 2′ oxygen, facilitating attack on an adjacent phosphodiester bond. This transesterification reaction breaks the 5′,3′ phosphodiester backbone of the RNA to generate 2′,3′ cyclic phosphate and 5′-hydroxyl products. Since the efficiency of 2′ hydroxyl proton abstraction is related to steric and chemical constraints on particular 2′ OH groups embedded in the RNA structure, the Pb2+ cleavage rate can be used to examine the structure of individual nucleotides within an RNA molecule. This method is a particularly sensitive probe of the tertiary structure of an RNA, provided that appropriate Pb2+ binding sites are created in the tertiary structure. In order to detect the cleavage site and the cleavage rate by Pb2+, RNAs are labeled with 32 P at either the 5′ or 3′ end (see Support Protocol 1 and 2). The labeled RNA is then subjected to Pb2+ cleavage (see Basic Protocol). The reaction mixture is analyzed on denaturing polyacrylamide gels containing 8 M urea. Partial alkaline hydrolysis reaction products and nuclease T1 digests of the same labeled sample are run alongside the cleavage reaction products to accurately locate the cleavage site (see Support Protocol 4). For optimal results, RNA should be renatured after labeling to ensure conformational homogeneity (see Support Protocol 3 to determine proper renaturation conditions). Reverse transcription runoff (UNIT 6.1) can also be applied to detect cleavages in larger RNAs. CAUTION: All personnel should be trained in working with radioactivity. NOTE: Extremely careful precautions should be taken in doing RNA work. Wear disposable gloves at all times to prevent nuclease contamination from your hands. Use the highest-quality water available and autoclave water prior to all solution preparation. It is not usually necessary to treat the water with diethyl pyrocarbonate (DEPC) if all possible precautions are taken to avoid nuclease contamination. NOTE: Use silanized (low-adhesion) tubes for the cleavage reaction to minimize adsorption of RNA to the tube wall. This is especially critical if the reaction products are to be quantitated. PROBING THE RNA STRUCTURE BY Pb2+ CLEAVAGE If one is to obtain interpretable and reproducible results, there is much more to the Pb 2+ cleavage reaction than simply mixing all the components and taking aliquots over time. Since RNA can have different conformations depending on a variety of external factors, Pb2+ cleavage under one set of conditions may not accurately reflect the biologically relevant structures one seeks to analyze. Therefore, to ensure correct interpretation, optimization of reaction conditions should be performed (see Critical Parameters) prior to conducting the final experiment. This section describes the basic protocol for a standard Pb2+ cleavage reaction.
BASIC PROTOCOL
Materials 5′ or 3′ 32P-labeled RNA in water (see Support Protocols 1 and 2) 0.3 M buffer (see Critical Parameters) Urea loading buffer (APPENDIX 2A) Contributed by Tao Pan Current Protocols in Nucleic Acid Chemistry (2000) 6.3.1-6.3.9 Copyright © 2000 by John Wiley & Sons, Inc.
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.3.1
Pb(OAc)2 stock solution prepared at 10× desired final concentration in reaction (see recipe for 10 mM solution) Heating block 3-mm filter paper (Whatman) Phosphorimager with appropriate software and phosphor screens Additional reagents and equipment for RNA renaturation (see Support Protocol 3), partial alkaline hydrolysis and T1 nuclease digestion (see Support Protocol 4), and denaturing polyacrylamide gel electrophoresis (see APPENDIX 3B) Cleave RNA 1. If necessary, renature RNA sample (see Support Protocol 3 to determine proper renaturation conditions). Renaturation is advisable if the RNA or source cells have been subjected to any potentially denaturing conditions; see Support Protocol 3 for details. Renaturation, which refers to folding, is different for every RNA. To obtain proper tertiary structure in vitro, [RNA], [Mg2+], and temperature will need to be varied. The exact amount of RNA to use should be determined empirically, as should the other parameters, when experimental conditions are optimized; however, enough RNA to give a concentration of 0.1 to 1 mM in 50 mL is a good starting point. See Critical Parameters for further discussion.
2. Dilute RNA to 50 µL with water and enough 0.3 M buffer to provide a 20 mM final buffer concentration. Renaturation should leave the RNA appropriately diluted.
3. Take a 4.5-µL aliquot of the RNA/buffer mix and mix with an equal volume of denaturing-gel loading buffer. This is the time-zero RNA sample.
4. Add 5 µL of 10× Pb(OAc)2 solution to the remaining RNA to initiate cleavage. Pb(OAc)2 concentration should be optimized with other experimental conditions; see Critical Parameters for discussion.
5. Incubate at 20° to 40°C. See Critical Parameters for discussion of reaction temperature.
6. Remove samples at selected time points: e.g., 5-µL aliquots at 0.5, 1, 2, 4, 8, 15, 30, and 60 min. To stop the reaction, mix an equal volume of urea loading buffer with each aliquot as it is taken. 7. Perform partial alkaline hydrolysis and T1 nuclease digestion of the same RNA (see Support Protocol 4). 8. Analyze the Pb2+ reaction mixture on a high-resolution denaturing polyacrylamide gel containing 8 M urea. Include samples of partial alkaline hydrolysis and T1 nuclease digestion reactions on the same gel. The author uses a 15% gel for a 76-mer RNA. Probing RNA Structure by Lead Cleavage
9. Compare the cleavage product band with the partial alkaline hydrolysis and T1 nuclease reaction bands to determine the cleavage site.
6.3.2 Current Protocols in Nucleic Acid Chemistry
Analyze data 10. Transfer the sequencing gel to a piece of 3-mm Whatman filter paper, cover with Saran Wrap, and dry. 11. Expose the gel on a phosphorimager plate at ambient temperature. Exposing the gel in a freezer does not improve the result although exposure at 4°C can lead to sharper bands.
12. Scan the phosphor plate and quantitate the full-length RNA using volume count, cpm substrate (S), and any reaction products of interest, cpm product (P): %S = cpm(S)/[cpm(S) + cpm(P)] 13. Plot ln(%S) versus reaction time and fit with a linear equation. The slope of the curve is the observed cleavage rate (kobs). The cleavage reaction may be biphasic when all time points are plotted on the same graph. This observation is often due to the heterogeneity of RNA conformation (see Support Protocol 3). In this case, use only the early time points, until ∼20% to 50% of the RNA are cleaved.
14. Compare kobs values for reactions under different conditions for optimal interpretation (see Commentary). 5′ 32P-LABELING OF RNA INCLUDING DEPHOSPHORYLATION RNA can be 32P labeled at the 5′ end with [γ-32P]ATP and T4 polynucleotide kinase. Chemically synthesized oligoribonucleotides generally contain a free 5′ hydroxyl group, which can be labeled directly. RNAs made by in vitro transcription generally contain a 5′ triphosphate group that has to be removed with alkaline phosphatase prior to the 5′ labeling reaction.
SUPPORT PROTOCOL 1
Label enough RNA for both the cleavage reaction (Basic Protocol) and the preparation of T1 and partial alkaline hydrolysis standards for electrophoresis (Support Protocol 4). Materials RNA sample in water 1 M Tris⋅Cl, pH 8.0 (APPENDIX 2A) 1 U/µL calf-intestine alkaline phosphatase Soaking buffer: 50 mM potassium acetate ( APPENDIX 2A)/200 mM KCl, pH ∼7 1:1 (v/v) phenol/chloroform (APPENDIX 2A) Ethanol 10× T4 polynucleotide kinase buffer (see recipe) [γ-32P]ATP (use highest activity available) T4 polynucleotide kinase Urea loading buffer (see APPENDIX 2A) Polyacrylamide gel containing 8 M urea Sodium acetate, pH 5.2 (APPENDIX 2A) Micropipettor Speedvac evaporator (e.g., Savant) X-ray film RNase-free surgical blade Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (see APPENDIX 3B)
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.3.3 Current Protocols in Nucleic Acid Chemistry
NOTE: If phosphatase treatment is not needed, proceed directly to step 7. NOTE: All microcentrifugations are performed at full speed. Remove 5′ triphosphate group 1. To 10 to 50 pmol RNA dissolved in water, add 1.5 µL of 1 M Tris⋅Cl, pH 8.0, and water to 29 µL. 2. Add 1 µL of 1 U/µL calf-intestine alkaline phosphatase. Incubate 30 min at 37°C. Purify phosphatase-cleaved RNA 3. Add 20 µL soaking buffer; then extract alkaline phosphatase with 50 µL of 1:1 (v/v) phenol/CHCl3, mix, and let stand 1 min at room temperature. Microcentrifuge 1 min and collect the top layer into a fresh tube. 4. Add 140 µL ethanol and incubate 20 min at −20°C. 5. Microcentrifuge 20 min at 4°C. Align tubes in the microcentrifuge so that the hinges of the tubes are facing the center of the rotor. The pellet will adhere to the tube on the side opposite the hinge.
6. Carefully pipet out the liquid. Dry the RNA pellet in a Speedvac evaporator. The pellet may not be visible to the naked eye.
Label RNA at 5′ end with 32P 7. To the RNA add in following order (10 µL final volume): 1 µL 10× T4 polynucleotide kinase buffer H2O 1 to 1.5 mol equiv [γ-32P]ATP 3 to 5 U T4 polynucleotide kinase. 8. Incubate 30 min at 37°C. Purify 5′-labeled RNA 9. Add 10 µL denaturing-gel loading buffer and purify on a denaturing polyacrylamide gel containing 7 M urea. The author uses a 10% gel for a 76-nucleotide RNA; see APPENDIX 3B.
10. Locate the radioactive band by exposing the gel briefly on X-ray film. Use a fluorescent ruler for orientation. If the RNA did not become labeled with phosphatase in step 2.
32
P, repeat using a new batch of alkaline
11. Develop the film and make a template that will be used to locate the band containing your RNA. 12. Lay the template on the gel, using the rulers as a guide. Cut out the RNA band with an RNase-free surgical blade. Elute RNA in the soaking buffer for at least 1 hr at ambient temperature.
Probing RNA Structure by Lead Cleavage
13. Add sodium acetate, pH 5.2, to 0.25 M (final) and 2.5 vol ethanol to precipitate the RNA. Mix and incubate at least 5 min on crushed dry ice or 30 min at –20°C. Microcentrifuge to remove supernatant. Dry the pellet and redissolve in 10 µL water.
6.3.4 Current Protocols in Nucleic Acid Chemistry
3′ 32P LABELING OF RNA USING T4 RNA LIGASE AND [32P]pCp RNA can be labeled at the 3′ end with 5′ [32P]pCp and T4 RNA ligase. Steps 1 to 3 describe the synthesis of 5′ [32P]pCp. For additional detail on the reaction, see 3′ RNA labeling in UNIT 6.1. In general, the RNA should be labeled such that at least 20,000 cpm will be loaded per lane.
SUPPORT PROTOCOL 2
Be sure to label enough RNA to have extra for the preparation of the sizing standards for electrophoresis (Support Protocol 4) in addition to what will be needed for the Basic Protocol. This procedure is designed for labeling 20 to 100 pmol RNA. Materials 10× T4 polynucleotide kinase buffer (see recipe) [γ-32P]ATP 3′ cytosine monophosphate (3′ Cp) T4 polynucleotide kinase Dimethyl sulfoxide (DMSO) 100 µM nonradioactive ATP T4 RNA ligase RNA sample in water Soaking buffer: 50 mM potassium acetate (APPENDIX 2A)/200 mM KCl, pH ∼7 Urea loading buffer (APPENDIX 2A) Additional reagents and equipment for purification of ligated RNA (see Support Protocol 1) NOTE: If commercial 5′ [32P]pCp (e.g., New England Nuclear) is used, proceed directly to step 4. Depending on the concentration supplied, use a quantity sufficient to provide the same molar ratio as achieved with [γ-32P]ATP. 1. Mix in the following order (20 µl final volume): 2 µL 10× T4 polynucleotide kinase buffer H2O 1 mol equiv [γ-32P]ATP to 3′ Cp (∼1 to 2 mol equiv RNA to be ligated) 3 to 5 U T4 polynucleotide kinase. 2. Incubate 30 min at 37°C. 3. Heat at 75°C for 3 min to inactivate kinase, then immediately place on ice for at least 3 min. 4. Add the following (30 µL final volume including RNA added in step 5): 3 µL 10× T4 polynucleotide kinase buffer (1× final) 4.5 µL DMSO (15% final) 1.8 µL 100 µM nonradioactive ATP (6 µM final) 0.7 U/µL T4 RNA ligase 5. Add ≤3 µL of RNA sample in water and incubate 4 to 19 hr at 16°C. If the RNA sample is too dilute, concentrate the RNA first in a Speedvac.
6. Add 30 µL urea loading buffer. Purify the ligated RNA as described for 5′-labeled molecules (see Support Protocol 1, steps 9 to 12).
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.3.5 Current Protocols in Nucleic Acid Chemistry
SUPPORT PROTOCOL 3
OPTIMIZATION OF RNA RENATURATION It is generally not difficult to observe Pb2+ cleavage of an RNA; however, caution is necessary for proper interpretation. RNAs are capable of forming multiple conformations that may be kinetically trapped (Uhlenbeck, 1995). This is particularly true for RNAs that are purified from denaturing polyacrylamide gels. For those RNAs, a renaturation step is mandatory to ensure conformational homogeneity. For RNAs that are isolated from cell extracts and have not gone through any steps that are likely to cause denaturation, renaturation is probably not necessary. However, if the cell extract has been subjected to the presence of millimolar amounts of EDTA, high temperature (>60°C), or phenol extraction, the RNA may have been denatured and renaturation should be attempted. The following procedure should be used to determine optimal conditions for renaturing a particular RNA species. Materials RNA sample in water MES, MOPS, or HEPES buffer MgCl2 stock solution, prepared at 10× desired final concentration in reaction (APPENDIX 2A for 1 M stock solution) Heating block Additional materials for nondenaturing polyacrylamide gel electrophoresis (see e.g., CPMB UNIT 2.7) 1. Mix RNA with buffer and water to obtain a final RNA concentration of ∼0.1 to 1 µM. Prepare triplicate tubes. 2. Heat 2 min at 85° to 90°C using a heating block. This step should denature any alternate secondary-structure conformers in the absence of divalent ions.
3. Take the sample out of the heating block. Either leave tube at room temperature for 3 min and then microcentrifuge briefly, or turn the heat block off and let the sample slowly cool to room temperature. The best method for cooling the sample will depend on the RNA and has to be determined empirically. In general, it is best to start with the slow cooling.
4. Add 10× MgCl2 solution to the desired Mg2+ concentration. Incubate tubes at three different temperatures (recommended: ambient, 37°C, and 50°C) for 10 min. To prevent nonspecific Mg2+-induced hydrolysis, do not heat the RNA to above ∼60°C in the presence of Mg2+.
5. Add other components, if required (e.g., 0.1 M KCl, 1 mM spermine). Incubate again at 37°C for 5 min. 6. Analyze all samples on a nondenaturing polyacrylamide gel (see, e.g., CPMB UNIT 2.7). 7. Assess the gel to determine which conditions produced the best renaturation, and use these when renaturing samples of this RNA species.
Probing RNA Structure by Lead Cleavage
Properly renatured RNA should show the following general properties when run on a nondenaturing gel (although these may differ among individual RNAs): (1) a single band on the gel, suggesting conformational homogeneity, and (2) fast migration relative to RNA in the absence of Mg2+, suggesting compactness. Most, if not all, RNAs with tertiary structure require the presence of a divalent ion (e.g., Mg2+) to renature correctly.
6.3.6 Current Protocols in Nucleic Acid Chemistry
PARTIAL ALKALINE HYDROLYSIS AND NUCLEASE T1 DIGESTION Aliquots of the end-labeled RNA are used to generate markers to size and analyze the Pb cleavage products. A ladder of truncated RNAs is generated by alkaline hydrolysis, while the T1 nuclease cleaves the RNA at guanosines.
SUPPORT PROTOCOL 4
Materials 5′- or 3′-end-labeled RNA (≥200,000 cpm; see Support Protocols 1 and 2) 1 mg/mL E. coli tRNA mixture (Sigma), dissolved in water 5× alkaline hydrolysis (AH) buffer: 5 mM glycine/2 mM MgSO4, pH 9.5 1 U/µL ribonuclease T1, diluted in water from a 100 U/µl stock (store up to ∼1 month at −20°C) Urea loading buffer (APPENDIX 2A) 100°C water bath 1. To 0.2 to 1.0 µl 5′- or 3′-end-labeled RNA, add water to 3 µL. Prepare two duplicate tubes. The exact amount of RNA will depend on the level of radioactivity.
2. Add 1 µL E. coli tRNA to each tube. This ensures that all reactions are carried out at a similar ratio of RNA to nuclease T1.
3. To one tube (alkaline hydrolysis condition), add 1 µL of 5× AH buffer, then boil 1 min, in a 100°C water bath. 4. To the second tube (T1 digestion condition), add 1 µL nuclease T1 (1 U), then incubate 2 min at 65°C. 5. Quick-cool 3 min on ice. CAUTION: Do not keep the T1 reaction mixture for >30 min, as T1 continues to cut RNA, even on ice.
6. Microcentrifuge briefly. 7. Add 5 µL urea loading buffer to each tube. The samples are now ready for use as size standards for comparison with the Pb2+ cleavage products.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps; prepare all buffers with autoclaved water and then reautoclave. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Pb(OAc)2, 10 mM Dissolve solid Pb(OAc)2 in autoclaved water. Filter through 0.22-µm filters into sterile plastic tubes. Divide into aliquots and store at −70°C. Thaw and use one aliquot at a time; thawed aliquots can be stored at −20°C and used for ∼1 week. T4 polynucleotide kinase buffer, 10× 0.5 M Tris⋅Cl, pH 7.6 (APPENDIX 2A) 100 mM MgCl2 100 mM 2-mercaptoethanol Store up to ~1 year at –20°C
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.3.7 Current Protocols in Nucleic Acid Chemistry
COMMENTARY Background Information The basic mechanism whereby Pb2+ cleaves
Probing RNA Structure by Lead Cleavage
RNA is proton abstraction by the polyhydrated Pb(OH)+ species. Because of its low pKa (∼7.7), Pb(OH)+ is present in relatively high proportions at neutral pH. Compared to other divalent metal ions with higher pKa (e.g., Mg(H2O)5(OH)+ with a pKa of ∼11.4), Pb(OH)+ base induces cleavage of the RNA backbone much more frequently to result in faster “background rates,” defined as the cleavage rate of an RNA of any sequence independent of secondary or tertiary structure. For example, background rates have been measured at 50 µM Pb(OAc)2, 10 mM MgCl2, pH 7.0, 25°C to be ∼0.003 min−1 (half-life ∼4 hr; Pan and Uhlenbeck, 1992) and at 10 mM MgCl2, pH 7.5, 25°C to be ∼1 × 10−6 min−1 (half-life ∼11,000 hr; Hertel et al., 1994). Thus, the background rate for Pb2+ cleavage is within the time scale for an impatient experimentalist, and Pb2+ cleavage can be easily carried out in reasonable amount of time. It should be emphasized, however, that due to the peculiar properties of hydrated Pb(OH)+, the background rate does not necessarily increase linearly with total Pb2+ concentration (Kragten, 1978; Pan et al., 1994). The actual observed Pb2+ cleavage rate (kobs) is of course dependent on the RNA structure. The helical regions in RNA do not have the appropriate geometry for an in-line attack of the 2′ oxygen on its adjacent phosphate. Therefore, cleavage rates for nucleotide residues within an RNA helix are slower than the background rate. The completely unstructured regions in an RNA should be cleaved at the background rate. Hence, Pb2+ cleavage is a useful means for differentiating doublestranded from single-stranded regions in RNA. On the other hand, the single-stranded regions in RNA often have some kind of structure that can either enhance or inhibit the cleavage mechanism. The observed cleavage rates for nonhelical regions are often uneven. Nevertheless, without a specific Pb2+ binding site on the RNA (see below), cleavage of nonhelical regions rarely exceeds 10-fold the background rate. Pb2+ ion(s) bound at well-defined sites in an RNA can significantly increase the cleavage rate at specific sites. These so-called “specific cleavage” reactions can have cleavage rates >1000-fold above the background rate (Pan and Uhlenbeck, 1992). Cleavage of yeast tRNAPhe is the most extensively studied case of a specific
cleavage reaction by Pb2+. Three Pb2+ binding sites have been found in the crystal structure of this tRNA, only one of which, Pb(1), results in specific RNA cleavage (Brown et al., 1985). Pb(1) is directly coordinated to nucleotides U59 and C60 in the TψC loop and cleaves the phosphodiester between U17 and G18 in the D loop. Site-directed mutagenesis of yeast tRNAPhe shows that the cleavage rate is sensitive to subtle changes in the tertiary structure of tRNA (Behlen et al., 1990). The Pb2+ cleavage rates are well-suited to assessment of structural changes in the tRNA tertiary fold upon mutations or site-specific modifications.
Critical Parameters The three essential components of the cleavage reaction, RNA, Pb2+, and Mg2+, can act cooperatively to influence the cleavage rate. As a polyanion, RNA can bind divalent ions nonspecifically to decrease the concentration of free divalent ions in solution, so high concentrations of RNA should be avoided. On the other hand, nonspecific adsorption of RNA on the wall of a test tube can cause inaccurate measurement of cleavage rates, and this is exacerbated by low RNA concentration (the use of silanized tubes alleviates, but does not eliminate, this problem). Therefore, the RNA concentration in the reaction should be kept at ∼20 to 200 µM in phosphate (e.g., yeast tRNAPhe = 76 phosphates). Use of >5 mM Pb2+ is not recommended since higher concentrations generate insoluble polyhydroxides that can coaggregate RNA, and aggregated RNA may be cleaved differently than RNA that is free in solution. How much Pb2+ to use in the reaction will depend on the concentration of Mg2+, which can compete with Pb2+ binding to RNA. The Pb2+/Mg2+ ratio is especially important for specific Pb2+ cleavage reactions since Mg2+ may bind at the same site or at an overlapping site to exclude simultaneous binding of Pb2+. To maintain the RNA structure, Mg2+ concentration should be kept at 1 to 100 mM. A 1:50 [Pb2+]/[Mg2+] ratio may be a good starting point for experimental optimization. pH and temperature of the reaction also play a role in producing optimal results. Since the cleavage rate is often proportional to Pb(OH)+ concentration, the reaction rate can be log-linearly dependent with pH below the pKa of Pb(OH)+. At pH above ∼7.5, Pb2+ is more prone to form polyhydroxide aggregates. Therefore, Pb2+ cleavage should be carried out between
6.3.8 Current Protocols in Nucleic Acid Chemistry
pH 6.0 and 7.7 (neutral pH is a good compromise). For the 0.3 M buffer, use MES, MOPS, or HEPES buffers prepared at an appropriate pH in autoclaved water, divided into aliquots, and reautoclaved. Inorganic buffers such as phosphates should not be used, since they may form insoluble salts with Pb2+. Higher temperature accelerates the cleavage rate but may destabilize the RNA structure to result in unintended cleavage sites. On the other hand, kinetically trapped alternate conformers of RNA are more stable at low temperature. Therefore, the reaction temperature should be kept between 20° and 40°C. Interpretation of cleavage results should be based only on primary cleavages, defined as the first cleavage event within the full-length molecule. It is possible, or even likely, that cleaving the backbone at one site can result in a conformational change in RNA to allow it be cleaved at other sites. These secondary cleavage sites are characterized by an initial lag in the kinetic analysis of the cleavage product. Therefore, it is crucial to perform time courses to ensure that interpretation is not based on secondary cleavage reactions.
Anticipated Results The cleavage rates at individual sites can be used to deduce information on the structure of RNA and on structural changes resulting from mutations or other modifications. Cleavage of residues is generally faster in nonhelical regions than in helical regions, so cleavage rates can reflect the involvement of particular residues in the secondary structure. Cleavage at specific sites can be sensitive to structural changes. Based on Pb2+ cleavage alone, however, the precise molecular nature of such structural changes is difficult to understand; it requires further investigation by other methods. Finally, protein binding to an RNA can sterically hinder the access of the reaction site. Therefore, it should be possible to deduce some
kind of “footprint” of a protein binding site by Pb2+ cleavage. Protein binding may also change the conformation of the RNA in the complex to enhance cleavage at other sites. The enhanced cleavage rates at such sites may be an excellent indication of conformational changes of RNA upon protein binding.
Time Considerations All procedures in this unit are simple and can be completed in 2 days. Day 1 involves 32P labeling and gel purification (see Support Protocols 1 and 2). Day 2 involves Pb2+ cleavage and gel analysis (see Basic Protocol). An extra day is needed for optimizing renaturation conditions (see Support Protocol 3).
Literature Cited Behlen, L.S., Sampson, J.R., DiRenzo, A.B., and Uhlenbeck, C. 1990. Lead-catalyzed cleavage of yeast tRNAPhe mutant. Biochemistry 29:25152523. Brown, R., Dewan, J., and Klug, A. 1985. Crystallographic and biochemical investigation of the lead(II)-catalyzed hydrolysis of yeast phenylalanine tRNA. Biochemistry 24:4785-4801. Hertel, K.J., Herschlag, D., and Uhlenbeck, O.C. 1994. A kinetic and thermodynamic framework for the hammerhead ribozyme reaction. Biochemistry 33:3374-3385. Kragten, J. 1978. Atlas of Metal-Ligand Equilibria in Aqueous Solution. Halsted Press, Chichester, UK. Pan, T. and Uhlenbeck, O.C. 1992. A small metalloribozyme with a two-step mechanism. Nature 358:560-563. Pan, T., Dichtl, B., and Uhlenbeck, O.C. 1994. Properties of an in vitro selected Pb2+ cleavage motif. Biochemistry 33:9561-9565. Uhlenbeck, O.C. 1995. Keeping RNA happy. RNA 1:4-6.
Contributed by Tao Pan University of Chicago Chicago, Illinois
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.3.9 Current Protocols in Nucleic Acid Chemistry
Probing Nucleic Acid Structure with Nickeland Cobalt-Based Reagents
UNIT 6.4
Use and application of nickel and cobalt reagents are described for characterizing the solvent exposure of guanine residues within DNA and RNA. These reagents promote guanine oxidation in the presence of a peracid such as monopersulfate, and the extent of reaction indicates the steric and electronic environment surrounding the N7 and aromatic face of this residue. Together with the complementary reagents described elsewhere in this chapter, the secondary and tertiary solution structure of polynucleotides may be deduced. Very low concentrations of the metal reagents are sufficient to promote guanine oxidation under near physiological conditions (see Basic Protocol). Nucleic acid recognition and oxidation does not perturb target structure or result in direct strand scission. Therefore, secondary procedures are necessary to identify sites of reaction. For polynucleotides of <200 to 300 residues, guanine oxidation is most conveniently detected by its induction of strand fragmentation upon treatment with piperidine (DNA; see Support Protocol) or aniline acetate (RNA; see UNIT 6.1). For larger polynucleotides, guanine oxidation is detected by its characteristic termination of primer extension (UNIT 6.1). Although the nickel reagent demonstrates the greatest specificity, the cobalt reagent maintains its selectivity at high temperatures and salt concentrations that are not compatible with the nickel reagent. NICKEL- AND COBALT-DEPENDENT OXIDATION OF NUCLEIC ACIDS This protocol describes a method for selectively oxidizing guanine residues (RNA or DNA) that do not stack within a double-helical structure. After excess oxidant is quenched and removed, the modified guanines are detected by procedures common to nucleic acid sequencing. These include either direct strand scission by subsequent treatment with piperidine (DNA) or aniline acetate (RNA) or termination of primer extension. The resulting polynucleotides are then separated and identified by denaturing gel electrophoresis.
BASIC PROTOCOL
CAUTION: Perchlorate is often used as a counter ion to the nickel complexes. Perchlorate salts containing organic materials are potentially explosive and should be handled carefully and in small quantities. Materials 1 µg/µL carrier DNA or RNA 100 mM potassium phosphate buffer, pH 7 (APPENDIX 2A) 1 M NaCl in water RNA or DNA sample: 30,000 cpm end-labeled (UNIT 6.1 or UNIT 6.3) or 1 pmol unlabeled 60 µM [NiCR](PF6)2 solution (see recipe and Fig. 6.4.1) or 60 µM CoCl2 in water 0.6 mg/mL OXONE solution (see recipe) 20 mM HEPES/100 mM EDTA, pH 7, in water NaOAc/EDTA/Tris solution (see recipe) 1 mM EDTA, pH 8 (APPENDIX 2A) Microdialyzer Lyophilizer Contributed by Steven E. Rokita and Cynthia J. Burrows Current Protocols in Nucleic Acid Chemistry (2000) 6.4.1-6.4.7 Copyright © 2000 by John Wiley & Sons, Inc.
Chemical and Enzymatic Probes For Nucleic Acid Structure
6.4.1
Additional reagents and equipment for phenol/chloroform extraction and ethanol precipitation (APPENDIX 2A and, e.g., CPMB UNIT 2.1A), piperidine treatment (see Support Protocol), aniline acetate treatment (UNIT 6.1), PAGE (e.g., APPENDIX 3B or CPMB UNIT 7.6), partial alkaline hydrolysis and T1 nuclease digestion (UNIT 6.1), and primer extension (UNIT 6.1) Oxidize nucleic acid 1. Combine the following in a microcentrifuge tube: 2 µL 100 mM potassium phosphate buffer 2 µL 1 M NaCl 6 µL 1 µg/µL carrier RNA or DNA. Other buffers may also be used (see Critical Parameters).
2. Add 30,000 cpm end-labeled RNA or DNA, or 1 pmol unlabeled RNA or DNA (for primer extension), and sufficient deionized water to obtain a total volume of 18 µL. If the sample is <250 residues in length, it should be 5′ or 3′ end labeled with 32P, permitting detection of the modified sites after chemical cleavage and electrophoretic separation. Longer samples should not be end labeled; these will be analyzed via primer extension. CPMB provides additional labeling methods, see e.g., CPMB UNITS 3.5, 3.6 & 3.10.
3. Initiate reaction by adding the following (total 20 µL/tube), vortexing, and incubating for 30 min at room temperature: 1 µL 0.6 mg/mL OXONE solution 1 µL 60 µM [NiCR](PF6)2 or CoCl2 This amount of OXONE results in a final concentration of 0.1 mM potassium peroxymonosulfate (KHSO5) in the reaction. The extent of target modification can be controlled by adjusting a number of parameters (see Critical Parameters).
Stop reaction and purify sample 4a. For subsequent piperidine treatment: Stop reaction by adding 2 µL of 20 mM HEPES/100 mM EDTA and vortex. Dialyze oxidized samples in a microdialyzer against 1 mM EDTA, pH 8 (twice) and deionized water (once) to remove the inorganic salts. Lyophilize the dialysate. The resulting material may be analyzed immediately or stored at −20°C for >1 week.
4b. For subsequent primer extension: Stop reaction by adding 180 µL NaOAc/EDTA/ Tris solution and vortexing. Extract with phenol/chloroform (see APPENDIX 2A and CPMB UNIT 2.1A). Precipitate DNA by adding 3 vol ethanol, incubating 60 min at −20°C, and microcentrifuging 10 min at 4°C. Remove supernatant, rinse with ethanol, microcentrifuge, and remove supernatant. Dry the pellet. (Also see CPMB UNIT 2.1A.)
2+ R N N
Probing Nucleic Acid Structure with Nickel- and Cobalt-Based Reagents
N Ni N H NiCR
Figure 6.4.1
NH2
N
NH N
DNA
N
KHSO5
III N N Ni N N
O
guanosine oxidation
OOSO3
The mechanistic basis for specific oxidation of guanine.
6.4.2 Current Protocols in Nucleic Acid Chemistry
Dialysis can be omitted for large RNA and for large DNA that is easily precipitated; however, precipitation alone is often not sufficient to purify RNA/DNA for subsequent analysis. The resulting material may be analyzed immediately or stored at −20°C for >1 week.
Identify sites of oxidation 5a. Chemical cleavage: For polynucleotides of <250 residues, assess modification sites by their characteristic strand scission after treatment with piperidine (for DNA; see Support Protocol; see CPMB UNIT 7.5) or aniline acetate (for RNA; Peattie and Gilbert, 1980; see DMS modification and cleavage of RNA, steps 8 and 9, in UNIT 6.1). Determine the ultimate profile of strand fragmentation by denaturing polyacrylamide gel electrophoresis (PAGE; e.g., APPENDIX 3B or CPMB UNIT 7.6). For RNA samples, subject aliquots of labeled RNA to partial alkaline hydrolysis and T1 nuclease digestion to use as sizing standards on the gel (UNIT 6.1). 5b. Termination of primer extension: For polynucleotides >250 residues, assess modified sites by determining their ability to terminate extension of a radiolabeled primer. Use this procedure for both DNA and RNA (Woodson et al., 1993; see UNIT 6.1).
DNA STRAND SCISSION BY PIPERIDINE TREATMENT DNA samples ≤250 nucleotides in length that were modified at exposed guanines using the nickel and cobalt reagents described in Basic Protocol 1 can be analyzed by treatment with piperidine. Piperidine causes strand scission at the modified guanine residues. The resulting DNA fragments can then be analyzed by denaturing polyacrylamide gel electrophoresis.
SUPPORT PROTOCOL
Materials Modified DNA sample pellet (see Basic Protocol) 0.2 M piperidine, freshly prepared in sterile water 1.5-mL screw-cap microcentrifuge tubes 90°C water bath Speedvac evaporator 1. Resuspend modified DNA sample pellets in 20 µL of 0.2 M piperidine. 2. Transfer samples to 1.5-mL screw-cap microcentrifuge tubes, tightly cap the tubes, and incubate 30 min at 90°C. During this reaction, the tubes must be tightly sealed to prevent piperidine loss.
3. Microcentrifuge tubes briefly to collect any condensate from the sides of the tubes. Transfer the tubes to a Speedvac evaporator and evaporate the piperidine to dryness. 4. Resuspend dried samples in 30 µL sterile water, transfer to new tubes, and dry in a Speedvac evaporator. 5. Repeat step 4 using 30 µL sterile water. Failure to completely remove piperidine will result in smearing of bands on the sequencing gel. The samples are now ready for gel fractionation by PAGE (e.g., APPENDIX 3B or CPMB UNIT 7.6).
Chemical and Enzymatic Probes For Nucleic Acid Structure
6.4.3 Current Protocols in Nucleic Acid Chemistry
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
NaOAc/EDTA/Tris solution 0.3 M NaOAc 10 mM EDTA 10 mM Tris⋅Cl, pH 7.6 (APPENDIX 2A) 0.5% (w/v) SDS Store up to 1 month at 4°C [NiCR](PF6)2 solution, 60 mM This complex is prepared according to Karn and Busch (1969) except that NaPF6 is used in place of NaClO4 and HClO4 (see caution below). [NiCR](PF6)2 is also available from the authors’ laboratories upon written request. A concentrated aqueous solution of [NiCR](PF6)2 (0.3 mM; 0.36 mg/mL) is stable for several months when stored at 4°C and protected from ambient light. Aliquots may be diluted to a working stock solution of 60 µM as needed. [Ni(II)cyclam](ClO4)2 (see recipe) may be used as an alternative to [NiCR](PF6)2 . However, Ni(cyclam) is less efficient than [NiCR](PF6)2. CAUTION: Perchlorate is often used as a counter ion to the nickel complexes. Perchlorate salts containing organic materials are potentially explosive and should be handled carefully and in small quantities.
[Ni(II)cyclam](ClO4)2 Dissolve 0.110 g 1,4,8,11-tetraazacyclotetradecane (cyclam; 0.55 mmol) in 10 mL ethanol and heat to 60°C. In a separate container, dissolve 0.200 g (0.55 mmol) Ni(ClO4)2⋅6H2O in 5 mL ethanol, and add in a dropwise fashion to the warm cyclam solution. Stir the resulting orange suspension at 60°C for an additional 10 min and then cool to room temperature. Collect the yellow product by vacuum filtration and wash with ice-cold ethanol. Use in the same fashion as [NiCR](PF6)2. Ni(II)cyclam is easily prepared from commercially available materials (Martin et al., 1977) with a recovered yield of 0.243g (96%). CAUTION: Perchlorate is often used as a counter ion to the nickel complexes. Perchlorate salts containing organic materials are potentially explosive and should be handled carefully and in small quantities.
OXONE solution, 0.6 mg/mL Dissolve 6 mg 2KHSO5⋅KHSO4⋅K2SO4 (OXONE; Sigma, Aldrich) in 10 mL deionized water (final 2 mM KHSO5). Prepare fresh each day. The suggested concentration refers to KHSO5 only.
COMMENTARY Background Information
Probing Nucleic Acid Structure with Nickel- and Cobalt-Based Reagents
Chemical and enzymatic probes are essential for characterizing nucleic acid structure when nuclear magnetic resonance and crystallography are not appropriate or possible (Ehresmann et al., 1987; Knapp, 1989; Nielsen, 1990). The variable extent of modification induced by each reagent indicates the relative accessibility of its target site. Information on multiple sites may in turn be used to predict the three-dimensional structure of
a polynucleotide. Although a limited analysis may provide some conformational data, most reliable structures are derived from the results of a broad series of experiments using a variety of probes that examine each functional region of each residue. Such an effort is necessary because the accessibility of one region of a nucleotide will not necessarily predict the accessibility of another region. For example, exposure of the phosphoribose backbone and N7 of guanine in the Tetrahymena group I intron
6.4.4 Current Protocols in Nucleic Acid Chemistry
RNA were found to be independent (Chen et al., 1993). The use of multiple probes to characterize a particular site is also often recommended to compensate for the inherent limitations of each reagent (Zheng et al., 1998). The solvent accessibility of guanine N7 has typically been examined by its alkylation with dimethyl sulfate (Ehresmann et al., 1987; Nielsen, 1990; UNIT 6.1). Failure to react suggests that a polynucleotide tertiary structure blocks access to this site. In contrast, variation in secondary structure does not significantly affect this reaction. Diethylpyrocarbonate (DEPC; UNIT 6.1) is more sensitive to base stacking within helical polynucleotides, but it is also much more efficient at adenine N7 than guanine N7 modification (Conway and Wikens, 1989). The structural environment of guanine N7 in RNA may also be investigated with RNase T1 (UNIT 6.1), but this enzyme requires access to the N1, O6, and N7 positions of guanine for activity (Heinemann and Saenger, 1985) and may disrupt local structure during substrate turnover (Zheng et al., 1998). The nickel reagent appears to bind directly to guanine N7 and deliver its oxidizing equivalent specifically to this guanine (Fig. 6.4.1). The intermediate size of the nickel complex provides for greater selectivity than dimethyl sulfate, and provides access to more target sites than is possible for a macromolecule such as RNase T1. Nickel-dependent oxidation is selective for guanine residues that are not held within canonical A or B helices (Chen et al., 1992, 1993). Modification occurs at extrahelical and single-stranded guanines in addition to base-paired guanines at a helical terminus or junction (Chen et al., 1992, 1993). Since this probe does not affect the structure of its target, its relative ability to oxidize guanine has been a very strong indicator of both static and dynamic conformations of DNA and RNA (Chen et al., 1993; Shih et al., 1998). Cobalt provides a complementary probe for guanine structure by generating a diffusible rather than metal-bound oxidant. Simple inorganic salts of cobalt, in contrast to nickel, are sufficient to catalyze formation of the sulfate radical from KHSO5 (Muller et al., 1996). [CoII(H2O)5(OH)]+ + HSO5−→[CoIII(H2O)5O]+
+ SO•− 4 + H2O
This radical exhibits an inherent specificity for guanine relative to the other bases and the phosphoribose backbone. More importantly,
guanine residues with highly accessible aromatic faces are most rapidly oxidized. The profiles of cobalt- and nickel-induced modification are often similar (Muller et al., 1996; Zheng et al., 1998), but will differ significantly when the environments surrounding the N7 and aromatic face of guanine are not equivalent (e.g., tRNAPhe; Muller et al., 1996). Overall, the conformational selectivity of cobalt is less than that of nickel, as expected for the smaller size of the cobalt- versus nickel-dependent oxidants. Comparative investigations based on these two probes are also useful for determining the influence of electrostatic potential on modification. The anionic sulfate radical generated by cobalt and the cationic radical-nickel complex may respond to surface charge in the opposite manner.
Critical Parameters The oxidation protocol described in this unit may be used as a starting point or guide for characterizing a variety of polynucleotides under different conditions. Typically, nucleic acid structure should be examined under conditions optimized for the nucleic acid’s natural function rather than for chemical modification. Variations in reaction and quenching conditions are described in the cited literature and other studies based on these reagents. Buffers and temperature. Contamination of the nucleic acid may inhibit the oxidative process. Buffers such as phosphate, Tris, and cacodylate are compatible with the nickel- and cobalt-dependent reactions. However, the Good buffers (for example, MOPS, MES, HEPES), EDTA, and thiols quench the reaction. Borate also inhibits the guanine-specific reaction and seems instead to promote variable modification of all polynucleotides. In addition, the nickel reagent is inhibited by high concentrations of salt (>1 M NaCl or NaClO4) and appears to decompose in the presence of KHSO5 at temperatures above 35°C. Although the cobalt reagent is also inhibited by NaCl, it remains active in the presence of 4 M NaClO4 and at temperatures above 80°C. Both reagents tolerate the presence of MgCl2, a common counterion required to form native RNA structure. Finally, bromide salts should be avoided as KHSO5 will oxidize bromide, which in turn leads to modification of C residues (Ross and Burrows, 1996). Extent of reaction. Oxidation should be limited to approximately 10% of the initial target nucleic acid in order to minimize secondary reactions of the oxidized products. The extent
Chemical and Enzymatic Probes For Nucleic Acid Structure
6.4.5 Current Protocols in Nucleic Acid Chemistry
of target modification is best controlled by increasing or decreasing the concentration of KHSO5, although the concentration of nickel and cobalt reagents as well as incubation time and temperature may also be varied. A common cause for a lack of reaction is the use of a KHSO5 solution that is not prepared fresh daily or the use of a metal solution that has been stored for an excessive time (typically >3 months). Old reagents may also induce abnormal reactions of uridine residues in RNA. If primer extension is used to detect sites of oxidation, phenol/chloroform extraction of the polynucleotide is generally required. This procedure likely removes and quenches materials that might otherwise inhibit polymerase activity.
Anticipated Results
Probing Nucleic Acid Structure with Nickel- and Cobalt-Based Reagents
The nickel complexes most readily oxidize guanine residues with highly accessible N7 positions, and the extent of reaction is indicative of the relative accessibility and electrostatic environment surrounding this position. In Aand B-helical structures of DNA and RNA, guanine N7 resides on the surface of the major groove, but is protected from reaction with nickel. Guanine residues are similarly unreactive when stacked within a duplex structure and paired in a noncanonical manner (Schmidt et al., 1995; Zheng et al., 1998). In contrast, guanine residues that are base paired at a helical junction or terminus are subject to nickel-dependent oxidation. The N7 of guanine within a Z helix is also exposed and demonstrates predictably high reactivity (Burrows et al., 1995). Guanines exhibiting the greatest reactivity are commonly those in highly disordered loops or single-stranded regions, and all guanines within such regions are oxidized with an equally high efficiency. When multiple structures are formed by a target nucleic acid, modification reflects the relative accessibility of each guanine weighted by the fractional concentration of its alternative conformations (Shih et al., 1998). The cobalt reagent generates equivalent data on guanine, except the extent of reaction is more dependent on the exposure of its aromatic surface than its N7 position (Muller et al., 1996). The specificity for unstacked guanine residues is also lower for cobalt- than nickeldependent oxidation. In the absence of an exposed target, both reagents will begin to oxidize guanine residues within a duplex in accord with their ionization potential. Therefore, under these conditions, the 5′ guanine of a GG se-
quence will react preferentially (Sugiyama and Saito, 1996; Burrows and Muller, 1998). Only one anomalous result has yet been detected with NiCR, and this, too, may provide useful structural data. Uridine residues are subject to direct strand scission if they are on the 3′ side of an adjacent uridine that forms a wobble base pair with guanine (5′-UU paired to 3′-GA; Hickerson et al., 1998).
Time Considerations Probing the structure of nucleic acids with nickel and cobalt reagents can be accomplished very rapidly and performed in tandem with other typical modification reactions. Samples can be left overnight to dialyze if this method is used for purification. Product characterization by strand scission (using piperidine or aniline acetate) or primer extension, followed by gel electrophoresis, can be completed on the second day.
Literature Cited Burrows, C.J. and Muller, J.G. 1998. Oxidative nucleobase modifications leading to strand scission. Chem. Rev. 98:1109-1151. Burrows, C.J., Muller, J.G., Shih, H.-C., and Rokita, S.E. 1995. Recognition of B vs. Z-form DNA using nickel and cobalt complexes. In Supramolecular Stereochemistry (J.S. Siegel, ed.) pp. 57-62. Kluwer Academic Publishers, Dordrecht, the Netherlands. Chen, X., Burrows, C.J., and Rokita, S.E. 1992. Conformation specific detection of guanosine in DNA: Ends, mismatches, bulges and loops. J. Am. Chem. Soc. 114:322-325. Chen, X., Woodson, S.A., Burrows, C.J., and Rokita, S.E. 1993. A highly sensitive probe for guanine N7 in folded structures of RNA: Application to tRNAphe and Tetrahymena group I intron. Biochemistry 32:7610-7616. Conway, L. and Wickens, M. 1989. Modification interference analysis of reactions using RNA substrates. Methods Enzymol. 180:369-377. Ehresmann, C., Baudin, F., Mougel, M., Romby, P., Ebel, J.-P., and Ehresmann, B. 1987. Probing the structure of RNAs in solution. Nucl. Acids Res. 15:9109-9128. Heinemann, U. and Saenger, W. 1985. Mechanism of guanosine recognition and RNA hydrolysis by RNase T1. Pure Appl. Chem. 57:417-422. Hickerson, R.P., Watkins-Sims, C.D., Burrows, C.J., Atkins, J.F., Gesteland, R.F., and Felden, B. 1998. A nickel complex cleaves uridines in folded RNA structures: Application to E. coli tmRNA and related engineered molecules. J. Mol. Biol. 279:577-587.
6.4.6 Current Protocols in Nucleic Acid Chemistry
Karn, J.L. and Busch, D.H. 1969. Nickel(II) complexes of the new macrocyclic ligands meso- and rac-2,12-dimethyl-3,7,11,17-tetraazabicyclo[11.3.1]heptadeca-1(17),13,15-triene. Inorg. Chem. 8:1149-1153. Knapp, G. 1989. Enzymatic approaches to probing of RNA secondary and tertiary structure. Methods Enzymol. 180:192-212. Martin, L.Y., Sperati, C.R., and Busch, D.H. 1977. The spectrochemical properties of tetragonal complexes of high spin nickel(II) containing macrocyclic ligands. J. Am. Chem. Soc. 99:29682981. Muller, J.G., Zheng, P., Rokita, S.E., and Burrows, C.J. 1996. DNA modification promoted by [Co(H2O)6]Cl2: Probing temperature-dependent conformations. J. Am. Chem. Soc. 118:23202325. Nielsen, P.E. 1990. Chemical and photochemical probing of DNA complexes. J. Mol. Recognit. 3:1-25. Peattie, D.A. and Gilbert, W. 1980. Chemical probes for higher-order structure in RNA. Proc. Natl. Acad. Sci. U.S.A. 77:4679-4682. Ross, S.A., and Burrows, C.J. 1996. Cytosine-specific chemical probing of DNA using bromide and monoperoxysulfate. Nucl. Acids Res. 24:5062-5063. Schmidt, M., Zheng, P., and Delihas, N. 1995. Secondary structures of Escherichai coli antisense micF RNA, the 5′-end of the target ompF mRNA, and the RNA/RNA duplex. Biochemistry 34:3621-3631. Shih, H.-C, Tang, N., Burrows, C.J., and Rokita, S.E. 1998. Nickel-based probes of nucleic acid structure bind to guanine but do not perturb a dynamic equilibrium of extrahelical guanine residues J. Am. Chem. Soc. 120:3284-3288. Sugiyama, H. and Saito, I. 1996. Theoretical studies of GG-specific photocleavage of DNA via electron transfer: Significant lowering of ionization potential and 5′-localization of HOMO of GG bases in B-form DNA. J. Am. Chem. Soc. 118:7063-7068.
Woodson, S.A., Muller, J.G., Burrows, C.J., and Rokita, S.E. 1993. A primer extension assay for modification of guanine by Ni(II) complexes. Nucl. Acids Res. 21:5524-5525. Zheng, P., Burrows, C.J., and Rokita, S.E. 1998 Nickel- and cobalt-dependent reagents identify structural features of RNA that are not detected by dimethyl sulfate or RNase T1. Biochemistry 37:2207-2214.
Key References Burrows, C.J. and Rokita, S.E. 1994. Probing guanine structure in nucleic acid folding using nickel complexes. Acc. Chem. Res. 27:295-301. A comprehensive review on various nickel-based reagents. Burrows, C.J. and Rokita, S.E. 1995. Nickel complexes as probes of guanine sites in nucleic acid folding. In Metal Ions in Biological Systems (H. Sigel, ed.) pp. 537-560. Marcel Dekker, New York. The most recent review covering applications from various laboratories. Rokita, S.E., Zheng, P., Tang, N., Cheng, C.-C., Yeh, R.-H., Muller, J.G., and Burrows, C.J. 1995. Nickel complexes in modification of nucleic acids. In Genetic Response to Metals (B. Sarkar, ed.) pp. 201-216. Marcel Dekker, New York. A summary of initial mechanistic studies.
Contributed by Steven E. Rokita University of Maryland College Park, Maryland Cynthia J. Burrows University of Utah Salt Lake City, Utah
Chemical and Enzymatic Probes For Nucleic Acid Structure
6.4.7 Current Protocols in Nucleic Acid Chemistry
Probing RNA Structures with Hydroxyl Radicals
UNIT 6.5
Iron(II)-EDTA is the principal reagent whereby one can conveniently generate hydroxyl radicals to promote cleavage of RNA at nucleotide resolution. In this unit, the Basic Protocol describes the simplest cleavage conditions, whereby free radicals originate from solvated molecular oxygen, while the Alternate Protocol outlines a more elaborate procedure commonly used to generate free radicals from hydrogen peroxide added to solution. In both procedures, Fe(II)-EDTA is added to solution mixtures containing end-labeled RNA, either alone or equilibrated with another ligand (e.g., protein), to allow for oxidative strand scission of the RNA chain under neutral buffer conditions. A similar cleavage pattern is observed using either method. CAUTION: When working with radioactivity, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following the guidelines provided by the local radiation safety officer. NOTE: Experiments involving RNA require careful precautions to prevent contamination and RNA degradation (see APPENDIX 2A). NOTE: All reagents should be ≥99% purity. All solutions must be prepared fresh in sterile water immediately before use and should be discarded thereafter. STRAND SCISSION OF RNA USING O2-DERIVED FREE RADICALS For strand scission of RNA using free radicals derived from solvated oxygen, 5′- or 3′-end-labeled RNA is equilibrated under the desired buffer or ligand-binding conditions. The Fe(II)-EDTA mixture and a suitable reducing agent are added to the RNA mixture to initiate the strand scission reaction and, after a 1- to 2-hr incubation, the free-radical reaction is quenched. The RNA is then directly processed alongside sequencing standards on a denaturing polyacrylamide gel to resolve the cleavage products.
BASIC PROTOCOL
Materials End-labeled RNA with a specific activity ≥2 × 106 dpm/pmol (UNIT 6.1) Appropriate buffers and RNA-binding ligands (see Critical Parameters) 1,4-Dithiothreitol (DTT; store at −20°C) (NH4)2Fe(II)(SO4)2⋅6H2O powder (store at room temperature) 200 mM Na2EDTA, pH 8.0 (APPENDIX 2A), prepared in sterile water 100 mM thiourea in sterile water (store at 4°C) 2× urea loading buffer (APPENDIX 2A) T1 nuclease digest and alkaline hydrolysis of end-labeled RNA ( UNIT 6.1) Distilled, deionized water, autoclaved before use 0.5- and 1.5-mL polypropylene microcentrifuge tubes, sterile RNase-free micropipettors and tips Water bath or suitable incubator Radioanalytic detection instrument or autoradiography film Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (PAGE; see APPENDIX 3B) Chemical and Enzymatic Probes for Nucleic Acid Structure Contributed by Daniel W. Celander Current Protocols in Nucleic Acid Chemistry (2000) 6.5.1-6.5.6 Copyright © 2000 by John Wiley & Sons, Inc.
6.5.1
1. Combine 0.1 pmol end-labeled RNA, the buffer components (e.g., buffers or simple salts), any additional RNA-binding ligands (e.g., proteins), and water to a total volume of 8 µL in a 0.5-mL polypropylene tube. Prepare an identical tube as a control. Incubate the mixture at the desired temperature until equilibrium is achieved. The buffers, incubation temperature, and the time required to achieve equilibration of the folded RNA with any additional ligands must be determined independently by the investigator (see Critical Parameters).
2. During equilibration, dissolve 7.7 mg DTT in 1 mL sterile water (final 50 mM) in a 1.5-mL microcentrifuge tube. This solution should be made fresh immediately before use.
3. Add 39.4 mg (NH4)2Fe(II)(SO4)2⋅6H2O to a sterile microcentrifuge tube and store in the dark until use. 4. Once the RNA solution has achieved equilibrium, prepare the Fe(II)-EDTA solution for immediate addition to the RNA solution. Add 1 mL sterile water to the tube containing (NH4)2Fe(II)(SO4)2⋅6H2O (final 100 mM) and vortex well to completely dissolve. In a separate polypropylene microcentrifuge tube, combine 10 µL of this solution with 10 µL of 200 mM Na2EDTA solution and 80 µL sterile water, yielding 10 mM Fe(II) and 20 mM EDTA. Vortex well. 5. Immediately add 1 µL Fe(II)-EDTA solution and 1 µL of 50 mM DTT solution to the RNA mixture to yield a final volume of 10 µL containing 1 mM Fe(II), 2 mM EDTA, and 5 mM DTT. Add 2 µL sterile water to the control tube. The remaining DTT and Fe(II)-EDTA solutions should be discarded following their use.
6. Incubate the reaction mixture for 1 to 2 hr at the desired temperature. The indicated time period is sufficient to generate a partial digest pattern of end-labeled RNA under single-hit conditions (see Commentary). The cleavage reaction displays a modest temperature dependence; nevertheless, suitable cleavage profiles have been obtained for reactions performed at temperatures ranging from 0° to 50°C (Celander and Cech, 1991).
7. Add 1 µL of 100 mM thiourea to quench the free radical reaction. Also add thiourea to the control tube. 8. Add an equal volume of 2× urea loading buffer to each tube. If larger reaction volumes are used, the RNA should be precipitated from ethanol before the addition of urea loading buffer. At this point, the reaction samples can be stored up to several days at −20°C.
9. Fractionate the reaction mixture on a denaturing polyacrylamide gel (8% to 20%; APPENDIX 3B), using a T1 digest and an alkaline hydrolysis of the original end-labeled RNA as a sequencing standard for the free-radical cleavage reactions. 10. Dry the gel and expose it to film by autoradiography or to a phosphor plate for quantitation using a radioanalytic detection instruments.
Probing RNA Structures with Hydroxyl Radicals
6.5.2 Current Protocols in Nucleic Acid Chemistry
STRAND SCISSION OF RNA USING H2O2-DERIVED FREE RADICALS This procedure is similar in several details to that described in the Basic Protocol. However, since the free radicals originate from hydrogen peroxide rather than from solvated molecular oxygen, the reaction proceeds at a more rapid pace.
ALTERNATE PROTOCOL
Additional Materials (also see Basic Protocol) 0.6% (w/w) hydrogen peroxide (H2O2) in sterile water, freshly prepared from 30% (w/w) commercial solution Sodium ascorbate powder 1. Prepare and equilibrate the RNA mixture as described (see Basic Protocol, step 1) but use a total volume of 7 µL. 2. During equilibration, dissolve 1.98 mg sodium ascorbate in 1 mL sterile water (final 10 mM). This solution should be made fresh immediately before use. It may be easier to prepare 10 mL solution.
3. Add 39.4 mg (NH4)2Fe(II)(SO4)2⋅6H2O to a sterile microcentrifuge tube and store in the dark until use. 4. Once the RNA solution has achieved equilibrium, prepare the Fe(II)-EDTA solution (see Basic Protocol, step 4). 5. Immediately add the following to the reaction mixture (total 10 µL): 1 µL Fe(II)-EDTA solution 1 µL 0.6% (w/w) H2O2 1 µL 10 mM sodium ascorbate. Add 3 µL sterile water to the control tube. Final reagent concentrations are 1 mM Fe(II), 2 mM EDTA, 0.06% H2O2, and 1 mM sodium ascorbate. The remaining sodium ascorbate, H2O2, and Fe(II)-EDTA solutions should be discarded following their use.
6. Incubate the reaction mixture for 1 to 3 min at the desired temperature. 7. Quench, fractionate, and observe the reaction as described (see Basic Protocol, steps 7 to 10). COMMENTARY Background Information The RNA strand scission reaction described here is initiated by a free radical, which originates from the reduction of an oxygen species in solution. The oxidation of the Fe(II)-EDTA reagent provides the source of electrons needed for reduction of molecular oxygen or hydrogen peroxide to form the reactive free radicals. The inclusion of a reducing agent, such as dithiothreitol or ascorbate, allows for the recycling of Fe(III)-EDTA to Fe(II)-EDTA. The formation of free radicals from solvated mo-
lecular oxygen can proceed via a hydroperoxide intermediate. The direct employment of a suitable concentration of hydrogen peroxide bypasses the requirement for an initial electron transfer to oxygen to generate the superoxide precursor of the hydroperoxide intermediate, as well as the ensuing bimolecular reactions giving rise to the peroxide. Consequently, the free radical cleavage reaction proceeds more rapidly in the Alternate Protocol than in the Basic Protocol. Before the RNA is loaded onto a denaturing polyacrylamide gel, a free-radical
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.5.3 Current Protocols in Nucleic Acid Chemistry
scavenger, such as thiourea, is added to quench the highly reactive hydroxyl radicals that are generated during the reaction. These reactions are as follows:
Probing RNA Structures with Hydroxyl Radicals
An analysis of nucleic acid products generated by the Fe(II)-EDTA cleavage reagent is consistent with the mechanism of strand scission beginning with hydrogen atom abstraction occurring at the C4′ position of ribose (Hertzberg and Dervan, 1984). Following oxidative damage of the ribose ring, a β-elimination-like reaction decomposes the ribose ring and directly leads to strand scission of the RNA chain. The C4′ hydrogen atoms are equally accessible on single- and double-stranded forms of RNA; therefore, it is not surprising that free radical–induced cleavage of RNA occurs with little preference for primary sequence or secondary structure (Celander and Cech, 1990). The folding of RNA tertiary structure often results in ribose moieties having different extents of accessibility to solvent. In the case of tRNAPhe and the group I intron from Tetrahymena thermophila, different regions of the RNA were found to be protected from cleavage by the Fe(II)-EDTA reagent (Latham and Cech, 1989). A striking correlation was found to exist for both RNA molecules in which the ribose positions that were underrepresented in the Fe(II)-EDTA cleavage reactions were found to possess low solvent-accessible surface areas (Latham and Cech, 1989; Cate et al., 1996). Thus, the Fe(II)-EDTA reagent can be used to discriminate solvent inaccessible surfaces within RNA molecules. Such surfaces arise whenever RNA tertiary structure forms a compact structure or through the binding of a ligand (e.g., a protein or an antibiotic drug) to RNA. The Fe(II)-EDTA cleavage reagent has been used to characterize the folded conformation and folding pathways of several ribozymes, including group I introns (Latham and Cech, 1989; Celander and Cech, 1991;
Heuer et al., 1991), RNase P RNAs (Pan, 1995; Westhof et al., 1996), and the delta virus ribozyme (Rosenstein and Been, 1996). Likewise, RNA regions that are protected by bound ligands have been elucidated in many systems with the use of the Fe(II)-EDTA cleavage reagent (Darsillo and Huber, 1991; Westhof et al., 1996). The chief utility and power of the Fe(II)-EDTA cleavage reagent lies in its ability to generate oxidative damage at every ribose moiety along the RNA chain, resulting in a pattern of strand scission with single-nucleotide resolution. Other chemical and enzymatic cleavage reagents commonly used today display primary sequence or secondary structure preference in their cleavage profile, so every nucleotide position along the RNA cannot be simultaneously monitored in a single experiment. It must be emphasized that the Fe(II)EDTA cleavage reagent protocols described here should only be used to monitor the solvent-accessible surfaces of RNA when folded in solution or when bound by a ligand under conditions of thermodynamic equilibrium. This particular reagent is not well suited to monitor folding transitions that occur on a more rapid scale than free radical formation by Fe(II)-EDTA. A higher flux of free radical formation is required to monitor the kinetics of RNA conformational changes or protein binding that occur on the millisecond time scale. Radiolysis of water by high-energy X rays produces hydroxyl radicals and free electrons. Irradiation of solutions by the high-flux “white light” X-ray beam, such as the X-19C at the National Synchrotron Light Source, yields sufficient concentrations of hydroxyl radicals so that cleavage kinetics required for quantitative nuclease protection studies of RNA can be done with exposures as short as 50 msec (Sclavi et al., 1997, 1998). This and similar sources are now available for characterizing conformational changes in RNA on a more rapid time scale.
Critical Parameters The successful application of either protocol relies on three critical parameters: (1) the quality of the end labeled RNA, (2) the reduction or elimination of substances from the reaction cocktail that act as free radical scavengers, and (3) the freshness of the Fe(II)EDTA cleavage reagent and companion reducing agent.
6.5.4 Current Protocols in Nucleic Acid Chemistry
RNA quality Any chemical modification procedure that generates a degradation product as the principal species requires that the investigator uses RNA of known length. For end-labeled RNA molecules, it is imperative that full-length species are subject to Fe(II)-EDTA cleavage. The reason for this criterion is that end-labeled RNA transcripts that are broken fragments of fulllength molecules may adopt conformations that differ from the population of full-length molecules. Since the Fe(II)-EDTA cleavage reagent cannot discriminate among these different populations of RNA transcripts, two or more different cleavage profiles will be superimposed upon the entire end-labeled RNA population if the RNA is not homogenous. The net result is a pattern of cleavage that is difficult to interpret because it is difficult to relate the cumulative cleavage profile to each RNA population present in the reaction. RNA fragmentation occurs readily through the use of nucleasecontaminated enzymes or by adventitious cleavage of RNA by metal ions in solution. The full-length, end-labeled RNA transcripts can be readily purified from any smaller products using denaturing PAGE (e.g., CPMB UNIT 2.12). Freshly labeled RNA is also recommended for use in these protocols. The very high specific end-labeling of RNA with 32P (3000 to 6000 Ci/mmol) results in molecules that are prone to degradation by either direct or watermediated autoradiolysis over time. A subtle degradation profile begins to emerge following storage of end-labeled RNA molecules for a period of 10 to 14 days at −20°C. The degradation pattern would not generally interfere with many studies performed with radiolabeled RNA, yet the intensity of the profile is significant enough to obscure the interpretation of the subtle cleavage profiles generated by Fe(II)EDTA. Free-radical scavengers can protect the RNA from water-mediated autoradiolysis; however, their inclusion in RNA storage buffers is discouraged because such compounds will interfere with the strand scission reaction promoted by the Fe(II)-EDTA cleavage reagent (see below). Free radical scavengers Three broad classes of compounds that are routinely used in RNA preparations are known to inhibit strand scission by Fe(II)-EDTA. (1) Polyribose compounds, like bulk RNA and glycogen, are frequently used as carrier agents to coprecipitate end-labeled RNA species from ethanol. These compounds do not ordinarily
pose a significant liability for most biochemical reactions involving RNA; however, they act as direct competitive inhibitors to strand scission by Fe(II)-EDTA since they contain the same highly reactive ribose moieties found in the end-labeled RNA. RNA transcripts can be precipitated from ethanolic solutions that contain low concentrations of simple salts, like 0.25 M NaCl or 0.30 M NaOAc, as one alternate precipitation procedure. (2) Glycerol, a cryoprotectant found in most protein storage buffers, is another efficient scavenger of free radicals. Glycerol is such an effective quencher of the Fe(II)-EDTA reaction that it can be used in place of thiourea (Latham and Cech, 1989). Glycerol is usually included in storage buffers to promote long-term stability of RNA-binding proteins, yet proteins often remain stable under short-term conditions with either no glycerol or low concentrations of glycerol present. One should evaluate whether the optimal reaction conditions require the presence of glycerol for studies that involve RNA-binding proteins. In the worst-case scenario, one may be able to empirically determine a low final concentration of glycerol that can be used to maintain the RNA-binding protein’s biochemical activity and to allow for optimal strand scission by Fe(II)-EDTA generated free radicals. (3) Buffers that are enriched with primary alcohols, e.g., Tris, are effective free-radical scavengers since they more readily undergo hydrogen-atom abstraction than other buffering compounds. One should reduce the concentration of such buffers in solution or replace them with an alternative buffering component (e.g., MOPS) so that efficient RNA strand scission occurs (Celander and Cech, 1990). Fe(II)-EDTA cleavage reagent freshness The most important ingredient to success in the strand-scission reaction is the use of fresh Fe(II)-EDTA and reducing agents. A common mistake made by those who cannot generate a decent cleavage pattern with end-labeled RNA is that the Fe(II)-EDTA reagent was prepared well before use (an hour or longer) or that old solutions of DTT or ascorbate were stored frozen from use in cleavage reactions performed at an earlier date. An Fe(II)-EDTA solution will undergo rapid oxidation to Fe(III)-EDTA, resulting in a yellow-colored solution in <1 hr after preparation. A fresh Fe(II)-EDTA solution should have a colorless to faint-blue tint, and such solutions are best prepared immediately before use. The chosen reducing reagent should
Chemical and Enzymatic Probes for Nucleic Acid Structure
6.5.5 Current Protocols in Nucleic Acid Chemistry
also be prepared immediately before use and discarded thereafter.
Anticipated Results The interpretation of cleavage pattern on denaturing polyacrylamide gels is extremely straightforward. The cleavage pattern will be a fairly uniform banding pattern at each nucleotide position. The extent of cleavage varies little between reactions; most reactions generate 10% to 20% cleavage products. Since strand scission occurs with the loss of the ribose moiety for the affected nucleotide, the position at which the end-labeled RNA fragment migrates relative to sequencing standards is shifted by one nucleotide. For example, hydrogen atom abstraction at the ribose group of the seventeenth nucleotide in a 5′-[32P]-end-labeled RNA chain will generate an oxidative strandscission product that migrates like an end-labeled 16-mer. The usual controls should be included to ensure that the cleavage pattern obtained is correctly attributed to the Fe(II)EDTA reagent rather than to an adventitious cleavage event caused by another source.
Time Considerations This assay is extremely rapid to perform and requires little technical expertise or prior experience. The preparation of the end-labeled RNA is more time consuming than any aspect of the Fe(II)-EDTA cleavage protocols described herein. One can learn an enormous amount of structural information about a folded RNA molecule or its association with other ligands in the time it takes to conduct these reactions (<2 hr) and perform gel electrophoresis (3 to 5 hr).
Literature Cited Cate, J.H., Gooding, A.R., Podell, E., Zhou, K., Golden, B.L., Kundrot, C.E., Cech, T.R., and Doudna, J.A. 1996. Crystal structure of a group I ribozyme domain: principles of RNA packing. Science 273:1678-1685.
Celander, D.W. and Cech, T.R. 1990. Iron(II)ethylenediaminetetraacetic acid catalyzed cleavage of RNA and DNA oligonucleotides: Similar reactivity toward single- and double-stranded forms. Biochemistry 29:1355-1361. Celander, D.W. and Cech, T.R. 1991. Visualizing the higher order folding of a catalytic RNA molecule. Science 251:401-407. Darsillo, P. and Huber, P.W. 1991. The use of chemical nucleases to analyze RNA-protein interactions. The TFIIIA-5 S rRNA complex. J. Biol. Chem. 266:21075-21082. Hertzberg, R.P. and Dervan, P.B. 1984. Cleavage of DNA with methidiumpropyl-EDTA-iron(II): Reaction conditions and product analyses. Biochemistry 23:3934-3945. Heuer, T.S., Chandry, P.S., Belfort, M., Celander, D.W., and Cech, T.R. 1991. Folding of group I introns from bacteriophage T4 involves internalization of the catalytic core. Proc. Natl. Acad. Sci. U.S.A. 88:11105-11109. Latham, J.A. and Cech, T.R. 1989. Defining the inside and outside of a catalytic RNA molecule. Science 245:276-282. Pan, T. 1995. Higher order folding and domain analysis of the ribozyme from Bacillus subtilis ribonuclease P. Biochemistry 34:902-909. Rosenstein, S.P. and Been, M.D. 1996. Hepatitis delta virus ribozymes fold to generate a solventinaccessible core with essential nucleotides near the cleavage site phosphate. Biochemistry 35:11403-11413. Sclavi, B., Woodson, S., Sullivan, M., Chance, M.R., and Brenowitz, M. 1997. Time-resolved synchrotron X-ray "footprinting", a new approach to the study of nucleic acid structure and function: Application to protein-DNA interactions and RNA folding. J. Mol. Biol. 266:144-159. Sclavi, B., Sullivan, M., Chance, M.R., Brenowitz, M., and Woodson, S.A. 1998. RNA folding at millisecond intervals by synchrotron hydroxyl radical footprinting. Science 279:1940-1943. Westhof, E., Wesolowski, D., and Altman, S. 1996. Mapping in three dimensions of regions in a catalytic RNA protected from attack by an Fe(II)-EDTA reagent. J. Mol. Biol. 258:600-613.
Contributed by Daniel W. Celander Loyola University Chicago Chicago, Illinois
Probing RNA Structures with Hydroxyl Radicals
6.5.6 Current Protocols in Nucleic Acid Chemistry
Chemical Reagents for Investigating the Major Groove of DNA A wide array of chemical reagents have been developed over the past 25 years for characterizing DNA structure and its interaction with drugs and proteins. When first introduced, these reagents provided one of a very limited number of approaches for identifying nucleotide sequences that, for example, bound to proteins or formed unusual (non-B helical) structures. More recently, physical methods such as NMR (James, 1995; Addess and Feigon, 1996) and X-ray crystallography (Timsit and Moras, 1992; Sriram and Wang, 1996) have increasingly become viable alternatives since their application has become more routine, providing molecular details of unrivaled resolution. Despite these advances, chemical methods remain highly popular. Their lasting appeal is maintained in part by their low cost and ease of use. Few other techniques have proven as versatile for both defined and complex systems. Most chemical reagents used to characterize DNA either alkylate or oxidize a particular functional group within one or more of the four standard deoxyribonucleotides. A complementary set of reagents is therefore necessary if an analysis of all polynucleotide domains is desired. Although a single region of DNA such as its major groove may be examined successfully with a limited number of reagents, reliance on a single reagent is not recommended. Data from multiple reagents help to ensure that results truly reflect the state of DNA and are not a function of an unknown or unforeseen idiosyncracy of any one reagent. A number of excellent reviews describe the general specificity and utility of a broad range of reagents, and the most comprehensive of these cover the literature prior to the last decade (Nielsen, 1990; Tullius, 1991; Chow and Barton, 1992; Lilley, 1992). This field has continued to benefit from ongoing development of many additional reagents as illustrated throughout this chapter. The focus of this specific commentary centers on the most commonly used and readily available reagents that react in the major groove: dimethyl sulfate (DMS), diethylpyrocarbonate (DEPC), potassium permanganate (KMnO4), osmium tetroxide (OsO4), and bromine (Br2) formed in situ (Ross and Burrows, 1996) from bromide (Br–) and monoperoxysulfate (HSO5−). The characteristics of each reagent will be reviewed individually below, and then a sampling of their
UNIT 6.6
broad-ranging applications will be illustrated collectively in the final sections.
GENERAL CONSIDERATIONS FOR DATA ANALYSIS One of the most attractive features of experiments based on chemical modification is the ability to generate large quantities of data rapidly. However, accurate interpretation of the data is less assured. Assessment of the results requires some familiarity with the basic chemistry of modification, the origins of its specificity, and its potential for ambiguity. With these considerations, strategies may be chosen to distinguish between a limited set of alternative conformations or even explore unknown structures. Ambiguities may still arise when unanticipated variables dominate reaction or when initial modification of a target, or molecules bound to a target, promote one or more secondary reactions. Perhaps the greatest limitation generally affecting chemical modification is set by its ability to report on only the most reactive and not necessarily the most abundant species. These potential problems need not diminish the importance of this approach as long as caution is practiced during experimental design and data interpretation. For example, perturbations caused by secondary reactions can usually be avoided by simply adjusting conditions so that only a small fraction of molecules (<20%) are modified in each analysis. Of course no single experimental technique is ever likely to establish a significant conclusion without corroboration from alternative methods. Chemical modification provides a convenient complement to the more time-consuming procedures based on biological and physical methods.
EXPERIMENTAL PROCEDURES SUPPORTED BY CHEMICAL MODIFICATION Use of chemical probes typically follows one of three approaches identified as (1) modification protection or “footprinting,” (2) interference experiments, and (3) missing contact probing. The first essentially measures the extent of modification along a nucleotide sequence. Uniform reactivity indicates structural homogeneity within DNA, and, conversely, variable reactivity indicates structural heterogeneity at least to the extent of localized in-
Contributed by Steven E. Rokita Current Protocols in Nucleic Acid Chemistry (2001) 6.6.1-6.6.16 Copyright © 2001 by John Wiley & Sons, Inc.
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*
*
1. DMS 2. pyridine/heat
* * *
1. KMnO4 2. pyridine/heat
Figure 6.6.1 Protein-dependent suppression and activation of DNA modification by conformation-specific probes.
Chemical Reagants for Investigating the Major Groove of DNA
creases or decreases in steric or electrostatic repulsion. For example, assembly of a proteinDNA complex may be expected to block access to nucleotides in a certain region of a helix and prevent reaction at this site uniquely (Fig. 6.6.1). Likewise, partial unwinding of duplex DNA resulting from a helical junction or bound protein will increase access and concomitant reaction of a selected number of neighboring nucleotides. These sites of interest are identified in both cases by their deviation from the basal level of reactivity established over many nucleotides. Although certain reagents such as hydroxyl radicals induce polynucleotide strand scission directly, the reagents reviewed below modify the pyrimidine and purine bases without causing spontaneous strand scission. The modified bases are instead detected by their diagnostic ability to (1) cause strand scission after subsequent treatment with heat and piperidine (Maxam and Gillbert, 1980; see CPMB UNIT 7.5) or (2) terminate polymerasebased primer extension (see CPMB UNIT 7.4A; Htun and Johnston, 1992). Interference and missing contact experiments both rely on chemical reagents to generate statistical populations or libraries of DNA that collectively contain a particular modification at various sites along a nucleotide sequence to block or remove potential contacts between
DNA and proteins (Brunelle and Schleif, 1987; Wissmann and Hillen, 1991). For example, DMS methylation of guanine N7 (G N7) will not inhibit protein binding to the major groove except for the subpopulation of DNA that is modified within its recognition domain. The crucial sites required for binding are then identified from the product distribution of the protein-bound and free sequences (Fig. 6.6.2). Alternatively, this same methylation process can be used to induce depurination and delete certain interactions that stabilize protein-DNA assembly (Brunelle and Schleif, 1987). In this case, sequence recognition is identified by the subpopulation of apurinic DNA that exhibits reduced affinity for the protein of interest. Any abasic site has the additional potential to condense with neighboring lysine residues and form protein-DNA cross-links. This provides yet another tool for analyzing DNA-protein association (Mirzabekov et al., 1989; Bavykin and Pruss, 1997). For all of the methods described above, success depends on selection of the appropriate chemical reagents that modify DNA in a highly predictable, reproducible, and selective manner.
*
*
*
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chemical
…
modification
*
*
* …
+
* …
pyridine heat
Figure 6.6.2 An interference assay for characterizing protein-DNA interactions.
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INDIVIDUAL CHARACTERISTICS OF REAGENTS COMMONLY USED TO PROBE THE MAJOR GROOVE OF DNA Despite the ever-growing number of reagents known to react in the major groove of DNA, none have maintained more prominence than DMS, DEPC, and KMnO4. The popularity of these and most reagents is a function of their reliable specificity and ready availability. The following section provides a brief overview of these reagents, as well as OsO4 and Br−/HSO5−, and explores the or igins and limitations of their selectivity. Specific examples illustrating concurrent application of these reagents are left for the final sections of this unit.
Dimethyl Sulfate (DMS) The small and simple methylating agent DMS was among the first in a set of reagents frequently used to characterize DNA, and was central to many of the initial efforts in nucleotide sequencing, protection interference, and missing contact experiments. Application of DMS remains widespread due in part to this versatility. The predominant site of methylation in duplex DNA is the N7 position of guanine on the surface of the major groove (Singer and Grunberger, 1983). The resulting N7 methyl derivative is only metastable, and ultimately generates an abasic site that can be detected by its diagnostic fragmentation after treatment with piperidine and heat (Fig. 6.6.3). This chemistry is the basis for the now famous Maxam-Gilbert sequencing reaction for guanine (Maxam and Gilbert, 1980; also see CPMB UNIT 7.5). Methylation of G N7 does not inhibit polymerase chain extension, and thus piperidine-dependent strand scission is a necessary prerequisite for detecting this modification with the Klenow fragment of polymerase I (Saluz and Jost, 1989; Htun and Johnston, 1992). With the advent of thermostable polymerases like Taq polymerase, the heat of thermo-
cycling is sufficient to fragment the methylated DNA and may now supersede the need for piperidine treatment (Brewer et al., 1990). The same characteristics that made DMS appealing for sequencing have also made it attractive for many other applications. DMS modifies G N7 with equal proficiency in singleand double-stranded DNA and does not exhibit sequence-dependent activation or inhibition (Hartley, 1993). Consequently, a relatively homogeneous profile of methylation can be generated with DMS. This in turn facilitates the interpretation of all sequencing and footprinting experiments. The intrinsic reactivity of DMS is quite high, and, as with most reagents that act on DNA, precautions must be taken to avoid contact with it (see CPMB UNIT 7.5). The danger of DMS is further compounded by its neutral and lipophilic nature, two properties that are largely responsible for its desirable lack of sequence specificity. The small size of DMS relative to DNA also provides it with unencumbered access to G N7, and the phosphoribose backbone neither shields the reagent from nor attracts it to particular regions or conformations of DNA. Other sites in DNA are also methylated by DMS, although at far lower efficiencies. For example, the N3 position of adenosine (A N3), located in the minor groove of duplex DNA, is subject to modification but reacts at a rate ~4-fold slower than that of G N7 (Singer and Grunberger, 1983). Other potential sites of alkylation are blocked by the formation of duplex DNA. The profile of methylation of single-stranded DNA is consistent with the nucleophilicity and electrostatic potential of the nitrogen heteroatoms (Pullman and Pullman, 1981). G N7 is significantly more reactive than the next most reactive position, A N1, and these are followed by A N7 > A N3 > G N3 (Singer and Grunberger, 1983). Theoretical and experimental investigations additionally confirm that A N7 is not nearly as nucleophilic as G N7 (Pullman and
Figure 6.6.3 DMS-dependent methylation of G N7 and subsequent strand scission promoted by piperidine and heat.
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Pullman, 1981; Singer and Grunberger, 1983). DMS was even used to map the nucleophilic sites within the pyrimidine nucleobases, despite their modest reactivity at the nucleoside level and even weaker reactivity when assembled into duplex DNA (Singer and Grunberger, 1983). The preference of DMS for G N7 remains sufficient in most target sequences so that complications rarely arise from reaction at competing sites. Methylation is only effectively inhibited when the accessibility of G N7 is severely limited by drug, oligonucleotide, or protein binding in the major groove (Nielsen, 1990). This result in turn serves as the basis for interference studies that help to localize binding sites in DNA. Quadruplexes and other unusual structures of DNA that involve coordination of G N7 also exhibit protection from DMS (Sen and Gilbert, 1988; Williamson et al., 1989; Fig. 6.6.4). Two caveats that may effect data evaluation were noted in a review by Nielsen (1990), and are well worth repeating. First, DMS has the potential to disrupt DNA-ligand interaction by methylation of the ligand in competition with its methylation of DNA. Second, proteins have the potential to increase the local concentration of DMS since they may establish lipophilic binding pockets in the vicinty of DNA. The lack of charge on DMS also renders this reagent relatively insensitive to the electrostatic properties of DNA and the ionic strength of solution (Wurdeman and Gold, 1988). This is in direct contrast to the nature of other alkylating agents, such as nitrogen mustards, that modify G N7 through a cationic intermediate and consequently demonstrate electrostatic affinity for DNA (Hartley et al., 1990).
Diethylpyrocarbonate (DEPC) A N7 is the target most commonly associated with reaction of DEPC (Peattie, 1979; Nielsen, 1990). Initial carbethoxylation of N7
Chemical Reagants for Investigating the Major Groove of DNA
Figure 6.6.4
leads to cleavage of the imidazole ring and subsequent alkaline lability of the phosphoribose backbone (Fig. 6.6.5). Reaction profiles of DEPC are therefore typically examined after piperidine treatment of the modified DNA. Interest in DEPC as a probe for DNA originated from its earlier applications in determining RNA sequence and structure (Peattie, 1979; Peattie and Gilbert, 1980), which in turn evolved from its even earlier use as a histidinespecific reagent for protein modification (Lundblad, 1995). This ability to modify proteins should now remain a concern when using DEPC to map protein-DNA interactions. The conformational specificity of DEPC as characterized with RNA is very sensitive to base stacking and is severely inhibited in helical structures (Peattie and Gilbert, 1980; Ehrsemann et al., 1987; Weeks and Crothers, 1993). Single-stranded and nonhelical regions provide the most accessible targets of DEPC within RNA. Equivalent specificity was detected for DNA, although some reaction was additionally noted within duplex DNA (Herr et al., 1982; Furlong and Lilley, 1986). Adenine residues are not the exclusive target of DEPC. Guanine residues, particularly those with high solvent exposure at N7, may also react (Herr, 1985; Johnston and Rich, 1985; Runkel and Nordheim, 1986; Scholten and Nordheim, 1986). Early studies indicated that both pyrimidine and purine nucleosides were subject to carbethoxylation by DEPC (Leonard et al., 1971; Vincze et al., 1973), but only purines appear to maintain an observable reactivity in polynucleotides (Nielsen, 1990). Within helical DNA, DEPC generally favors modification of A over G when the accessibility of their N7 positions is similar (Herr, 1985). The origin of this selectivity has not yet been determined, and is certainly not based on nucleophilicity. Otherwise, guanine would have been most reactive as illustrated with DMS. Neither the general electrostatic nature of DNA
DMS reaction and protection for G-C and G-G base pairs and G-quartets.
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Figure 6.6.5 DEPC-dependent modification of A N7 and subsequent strand scission promoted by piperidine and heat.
nor the ionic conditions of reaction greatly affect the specificity of DEPC as expected for a neutral reactant (Klysik et al., 1990; NejedlO et al., 1998). The most predictable determinant for modification of A appears simply to be the steric accessibility of its N7 position.
Potassium Permanganate (KMnO4) and Osmium Tetroxide (OsO4) Both reagents selectively oxidize the C5, C6 double bond of thymine residues (Fig. 6.6.6) and support convenient methods for DNA sequencing and conformational analysis (Nielsen, 1990). The lack of charge on OsO4 minimizes its sensitivity to the electrostatic properties of DNA and may simplify interpretation of modification data relative to those based on KMnO4. However, OsO4 is somewhat volatile and quite hazardous (Pale?ek, 1992a), and is not applied as frequently as the safer alternative, KMnO4. Even though the ultimate oxidant MnO4− is charged, it remains cell-permeable for in vivo studies (Sasse-Dwight and Gralla, 1989). The anionic characteristic also suppresses reaction with DNA due to electrostatic
repulsion of DNA, despite the enduring focus in the general literature on the importance of sterics (Hänsler and Rokita, 1993). Oxidation of duplex DNA with KMnO4 and OsO4 is greatly inhibited relative to that of single-stranded DNA, and consequently these reagents are very useful for identifying regions of unusual and nonhelical structure. Perturbations of duplex DNA caused by association of proteins and drugs are also often sufficient for stimulating a local hyperreactivity (Fig. 6.6.2). Even minor distortions in DNA conformation resulting from single base mismatches can be detected by KMnO4 and OsO4 (Cotton et al., 1988; Roberts et al., 1997; Lambrinakos et al., 1999). The cis diol product of T oxidation is conveniently detected by either strand scission induced by piperidine and heat (Friedman and Brown, 1978; Rubin and Schmid, 1980) or termination of primer extension catalyzed by a variety of common DNA polymerases (Ide et al., 1985; Borowiec et al., 1987; Clark and Beardsley, 1987). The specificity and potential utility of MnO4− was noted very early in the quest for
Figure 6.6.6 KMnO4- and OsO4-dependent oxidation of thymine and subsequent strand scission promoted by piperidine and heat.
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Chemical Reagants for Investigating the Major Groove of DNA
chemical probes of nucleic acids. Access to the target C5, C6 of T is an understandable requirement for efficient reaction, and hence may explain the preference for single- versus doublestranded DNA (Hayatsu and Ukita, 1967). Residues within duplex DNA that became sensitive to MnO4− after supercoiling or binding to a protein were originally considered indicative of helix distortions and base unpairing (Borowiec et al., 1987; O’Halloran et al., 1989). However, the enhanced exposure of T anticipated from chemical modification is not always evident in the crystal structure of protein-DNA complexes (Bochkarev et al., 1998). In some cases, only subtle changes in conformation have appeared responsible for the high reactivity of thymine residues. These unusual sites of modification might reflect protein-dependent shielding of the anionic charge of DNA from the anionic reagent MnO4− (Hänsler and Rokita, 1993). Permanganate oxidation of duplex DNA can be accelerated 25-fold by merely increasing the ionic strength of the reaction solution from 0.1 to 4.0 M (Hänsler and Rokita, 1993). Under these conditions, the high concentration of cations effectively diminishes charge repulsion between the reactants. This dependence on ionic strength is not an inherent characteristic of MnO4− since the oxidation of neutral thymidine is enhanced only 1.3-fold in a comparable study (Hänsler and Rokita, 1993). Similarly, oxidation of DNA by OsO4 is unaffected by ionic strength (NejedlO et al., 1998). Permanganate then serves as an example of how reagents with a charge may respond to both steric and electrostatic properties of DNA. The specificity of MnO4− for T was first demonstrated using mononucleotides (Hayatsu and Ukita, 1967). Under conditions that consumed 95% of TMP, the purines dGMP and dAMP were nearly inert, and the pyrimidine dCMP was only marginally reactive. Even uridine 5′-phosphate that only lacks the 5-methyl of TMP oxidized at a rate ~10-fold slower than TMP. The resulting product of oxidation, the cis 5,6-diol of T, readily hydrolyzes to an abasic site (Howgate et al., 1968). These favorable characteristics led MnO4− to become a standard reagent for chemical sequencing of DNA (Rubin and Schmid, 1980; McCarthy, 1989; Williamson and Celander, 1990). Occasionally, background oxidation of duplex DNA has been observed particularly at guanine residues (McCarthy et al., 1990; McCarthy and Rich, 1991). The origin of this has not yet been identified, and controversy remains on whether or not 8-oxoguanine is a product of this minor
pathway (Akman et al., 1990; Nawamura et al., 1994). Perhaps the background reactions are a result of prior modification or contamination of the parent DNA. For example, 8-oxoguanine residues and their neighboring bases, as well as guanine side products encountered during oligonucleotide synthesis, are all targets of MnO4− oxidation (Yeung et al., 1988; Koizume et al., 1998). The reactivity of OsO4 is uniquely activated by the presence of tertiary amine ligands, and thus this reagent is often used in the presence of pyridine or 2,2′-bipyridine (Pale?ek, 1992a). Such ligands dramatically stabilize the osmium ester intermediate (Fig. 6.6.6). Unlike MnO4− and OsO4, which generate the diol derivative of thymine, the OsO4-pyridine complex proceeds only to the cyclic ester (Neidle and Stuart, 1976). This species hydrolyzes in a manner equivalent to the diol under standard alkaline conditions to yield strand fragmentation. Alternatively, a primer-extension assay can be used to identify the sites of modification. The conformational specificity of oxidation can be controlled in part by the choice of amine used to coordinate to the osmium (Pale?ek, 1992a). The ultimate extension of this strategy is illustrated by the sequence-directed oxidation of a single thymine using an osmium complex derived from a bipyridine-oligonucleotide conjugate (Ford et al., 1981; Nakatani et al., 2000).
Bromide (Br–) with Monoperoxysulfate (HSO5−) A very promising method of detecting cytosine residues in nonhelical DNA has been recently developed and relies on generation of bromine (Br2) in situ by oxidation of Br– in the presence of HSO5− (Ross and Burrows, 1996; Fig. 6.6.7). Extrahelical C residues exhibit a 10-fold increase in reactivity above a low background level in duplex DNA. In single-stranded structures, modification of C is minimally 4fold more efficient than that of T, G, or A. Preferential reaction of C is also observed after equivalent addition of Br2, although its selectivity is reduced compared to that of Br− / HSO5− (Ross and Burrows, 1996). Both conditions result in the intermediate formation of 5-bromo C. This is susceptible to further addition by Br2, which in turn induces DNA strand fragmentation after treatment with piperidine and heat (Ross and Burrows, 1996, 1997). As more investigators begin to explore the utility of Br–/HSO5− (Stevens and Glick, 1999; Kasparkova et al., 2000; Kostrhunova
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Figure 6.6.7 and heat.
Oxidation of C by Br−/HSO5− and subsequent strand scission promoted by piperidine
and Brabec, 2000), this technique will likely become the MnO4− or OsO4 equivalent for C. Two alternative reagents, hydroxylamine (Rubin and Schmid, 1980; Johnston and Rich, 1985; Johnston, 1992) and bisulfite (Hayatsu, 1976; Gough et al., 1986), have been known for many years to react with C in a conformation-selective manner, but their use has been very limited. This lack of popularity may be due in part to the very high concentration of reagents (2 M) that is required for sufficient modification (Johnston and Rich, 1985; Gough et al., 1986). Interest in bisulfite has recently revived since its selectivity for C over 5-methyl C has provided a convenient method for mapping methylation patterns of CpG islands in DNA (Rother et al., 1995; Kinoshita et al., 2000). Bisulfite reaction with C has also become central to a protocol for mapping chromatin in vivo in concert with methyltransferases (Kladde and Simpson, 1996).
APPLYING CHEMICAL REAGENTS TO PROBE THE MAJOR GROOVE OF DNA Sample investigations based on chemical approaches to characterizing DNA structure, and particularly its major groove, are described below. The goal of this section is to illustrate the range of information that is made available by these methods, rather than provide a comprehensive survey of all major advances that have occurred since previous review of this subject (Nielsen, 1990; Tullius, 1991; Lilley, 1992; Chow and Barton, 1992). Topics are organized into four sections, DNA conformation, drug-DNA association, protein-DNA association, and in vivo footprinting. Most examples involve the use of multiple reagents, since definitive conclusions are often difficult to establish with only a single reagent. Complementary techniques of footprinting (protection) and interference or missing contacts are also often performed concurrently to substantiate the re-
sults of individual analyses. Although chemical modification does not offer the highest atomic resolution, it represents a very expedient method for defining key structural features of DNA and identifying the functional groups responsible for recognition and binding to proteins and drugs.
Conformational Analysis of Duplex DNA Uniform double-helical DNA typically yields an equally uniform pattern of modification, and even minor perturbations of groove dimensions or base pairing have the potential to promote or inhibit reaction of conformationspecific probes. For example, biological methylation of A to form N6-methyl A appears to weaken A-T pairing and stacking in duplex DNA, as detected by the greater susceptibility of its N7 position to reaction with DEPC (Guo et al., 1995). Numerous investigators have also used DEPC to characterize the properties of A-tract DNA. The N7 position of A is more reactive in A-tracts than in canonical duplex DNA. This is consistent with the wider major groove and narrower minor groove associated with A-tract DNA (McCarthy et al., 1990, 1993; NejedlO et al., 1998). In contrast, the complementary T-tract does not exhibit high reactivity with either OsO4-bipyridine or MnO4− and suggests that the widened major groove still maintains considerable steric and electrostatic repulsion (McCarthy et al., 1993; NejedlO et al., 1998). Some deviations from this general reaction profile, and hence helical structure of DNA, have additionally been noted at certain A-tract junctions and in natural variants of A-tract DNA (McCarthy et al., 1993; Chang et al., 1994; NejedlO et al., 1998). The sensitivity of DEPC, OsO4-pyridine, and MnO4− to nucleotide stacking extends to parallel-stranded duplex DNA (Klysik et al., 1990), and its helix-coil transition may be monitored by exposure to OsO4-pyridine. An-
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tiparallel DNA forming a left-handed Z helix represents another highly unusual duplex structure that has been examined by chemical probes. DEPC readily reacts with the N7 positions of both A and G residues within Z-helical domains (Herr, 1985; Johnston and Rich, 1985; Runkel and Nordheim, 1986). This modification is relatively independent of base stacking, since the N7 position in Z DNA is oriented towards the outer edge of the helix and maintains much greater accessibility than it does in right-handed B DNA (Pullman and Pullman, 1981). B-Z and Z-Z helical junctions are readily modified by a variety of reagents including OsO4 and MnO4− in a manner consistent with a region of high disorder or conformational dynamics (NejedlO et al., 1985; Falazka et al., 1986; Jiang et al., 1991). All of the reagents introduced above except for DMS are quite useful in identifying extrahelical or weakly stacked bases within standard duplex DNA. DMS is insensitive to these perturbations because both single- and doublestranded DNA are methylated at G N7 with equal efficiency (Hartley, 1993). However, subtle changes in helical conformation caused by noncanonical structures such as base mismatches containing T may be detected by oxidation with MnO4− in the presence of tetralkylammonium salts (Gogos et al., 1990; Roberts et al., 1997; Lambrinakos et al., 1999). Addition of this type of salt appears to enhance selectivity for mismatches (Cotton, 1989). These conditions now serve as the basis for a protocol entitled “chemical cleavage of mismatch” for detecting point mutations (Gogos et al., 1990; Roberts et al., 1997; Lambrinakos et al., 1999). Interestingly, reaction is not limited to mispaired T. MnO4− additionally seems to oxidize some mismatched G and C residues, and both MnO4− and OsO4 oxidize normally paired T residues adjacent to mismatched bases (Cotton and Campbell, 1989; Lambrinakos et al., 1999).
Chemical Reagants for Investigating the Major Groove of DNA
Single-stranded regions of DNA formed by cruciform or hairpin structures are readily identified by conformational probes as well, and the presence of T, C, and A in these structures is typically identified by reaction with MnO4−, Br−/HSO5−, and DEPC, respectively (Scholten and Nordheim, 1986; Hänsler and Rokita, 1993; Stevens and Glick, 1999). These reagents have also been successfully applied to characterization of four-way helical junctions (Webb and Thomas, 1999), strand displacement and invasion by peptide nucleic acids (Egholm et al., 1995; Armitage et al., 1997), and equilibria of single-, double-, and triple-stranded species formed by naturally occurring triplet base repeats (Mäueler et al., 1998; Fig. 6.6.8). Once again, methylation of G N7 by DMS cannot discriminate between these forms of DNA. An alternative reagent for G N7, not discussed in this review but sensitive to secondary structure, involves nickel- or cobalt-dependent oxidation of G in the presence of HSO5− (UNIT 6.4). The first indication that G-rich telomeric sequences could form four-stranded structures was based on their unusual pattern of modification with DMS, MnO4−, and DEPC (Sen and Gilbert, 1988; Williamson et al., 1989; Venczel and Sen, 1993; Balagurumoorthy and Brahmachari, 1994). In this case, DMS was particularly useful, since the quadruplex assembled into a so-called G-quartet that is stabilized by Hoogsteen base pairs that involve G N7 (Sen and Gilbert, 1988; Williamson et al., 1989; Fig. 6.6.4). Thus, formation of the Gquartet inhibited methylation at this site and, conversely, methylation inhibited G-quartet formation. Nucleotides linking the G-rich regions were targets of DEPC and MnO4− reaction when looped between G-quartets, but were inert when stacked in the standard B duplex.
Drug-DNA Association The footprint of a low-molecular-weight drug on DNA often extends over only a few
Figure 6.6.8 Strand displacement increases exposure of target sequences to chemical probes. A peptide nucleic acid analog based on thymine and an anthraquinone intercalator (AQI) was used to recognize an A5 sequence (Armitage et al., 1997).
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base pairs, and consequently chemical modification data are often difficult to interpret during the initial stages of characterizing drug-DNA interactions. Instead, reagents are most useful when applied to well-defined oligo- or polynucleotide model systems that are designed to address specific questions on drug binding. The reagents highlighted in this unit are also not typically the first employed, since majorgroove recognition is rather rare for either synthetic or natural compounds. Two notable exceptions to this generalization are triplex-forming oligonucleotides that are currently under development as gene-targeted drugs (Miller, 1996; Giovannangeli and Hélène, 1997; Fox, 2000) and cisplatin derivatives that have already proven highly successful in treatment of certain cancers (Jamieson and Lippard, 1999; Wong and Giandomenico, 1999). Assembly of a third DNA strand into the major groove of duplex DNA protects A N7 and G N7 from DEPC and DMS reaction, respectively (Collier et al., 1991; Beal and Dervan, 1992). Similarly, platinum coordinates to DNA through G N7, and hence protects this site from DMS-dependent methylation (Sip et al., 1992; Kasparkova et al., 2000). Interstrand cross-linking at d(GC/CG) sites by platinum does not appear to cause significant unstacking of DNA, at least as indicated by the reaction profile of DEPC and OsO4 (Sip et al., 1992). Structural perturbations are limited to the two C residues directly adjacent to the platinum, as identified by reaction of hydroxylamine. Intrastrand crosslinking by cisplatin and a related distamycin conjugate induced only a localized hyperreactivity of MnO4−, DEPC, and Br−/HSO5−, and this was consistent with an asymmetric and local unwinding of DNA (Marrot and Leng, 1989; Kasparkova et al., 2000; Kostrhunova and Brabec, 2000; Fig. 6.6.9).
Figure 6.6.9 Local distortion of duplex DNA induced by a dinuclear platinum complex. Relative reaction is designated as high (h), medium (m), and low (l). Adapted from Kasparkova et al. (2000) with permission from The American Society for Biochemistry and Molecular Biology.
A variety of intercalating agents such as ethidium, 9-aminoacridine, and N,N-di(9-acridinyl)spermidine stimulate unwinding of helical DNA as determined in part by reaction with DEPC and MnO4− (Jeppesen and Nielsen, 1988). These same probes similarly helped to identify the sequence-dependent binding properties of the natural bisintercalator echinomycin (Bailly et al., 1994). Even drugs such as bleomycin that do not directly interact with the major groove may still cause sufficient structural distortion of DNA to affect its subsequent reactivity (Fox and Grigg, 1988). In particular, footprinting by DEPC and MnO4− suggests that bleomycin enhances exposure of the nucleotide directly 3′ to its binding site. As expected, DMS is generally insensitive to such binding, since no steric barrier is created around G N7.
Protein-DNA Association The great majority of protection and interference experiments designed to probe proteinDNA interactions within the major groove rely on DMS and MnO4−. All reagents have the potential to react with protein in a competitive manner, but the greatest number of complications likely arise from alternative reagents such as DEPC and OsO4 (Pale?ek, 1992b; Lundblad, 1995). Consequently, DEPC and OsO4 are applied with less frequency to study protein-DNA complexes (Dobi and Agoston, 1998; Mäueler et al., 1998). Investigations based on Br–/ HSO5− have only just begun, and therefore their potential to map protein-DNA interactions is not yet known. However, these conditions have the potential to oxidize a variety of amino acid side chains that might disrupt protein-DNA association. DMS and MnO4− alone can still provide considerable information on DNA-protein interactions, since their reaction specificities are quite complementary. Methylation of G N7 by DMS is unaffected by the helical conformation of DNA, but can be suppressed by loss of major groove accessibility due to protein binding. In contrast, oxidation of T by MnO4− is suppressed in helical DNA and activated by relaxation of its conformational and electrostatic constraints due to protein binding. Such effects were observed after lac repressor bound to its operator sequences (Borowiec et al., 1987). DMS protection was evident at two repressor binding sites, while MnO4− reaction was enhanced in an intervening AT-rich sequence. Similarly, an open transcription complex formed by RNA polymerase and MerR protein bound with Hg2+ conferred hypersensitivity to MnO4− at nine T
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residues along a 14-nucleotide sequence (O’Halloran et al., 1989). In contrast, the Hg2+MerR protein complex in the absence of the polymerase appeared to unwind duplex DNA without affecting MnO4− reaction, and the polymerase in the absence of the MerR complex induced MnO4− reaction at only five T residues (O’Halloran et al., 1989; Ansari et al., 1992). Each of the intermediate DNA-protein complexes were characterized by their unique patterns of DMS protection. Surprisingly, a small number of G residues also became hyperreactive to DMS. Since DMS is insensitive to DNA unwinding, reaction efficiency might have increased due to the accumulation of DMS in hydrophobic pockets of the complexes. Open transcription complexes formed at other DNA sequences also exhibit hyperreactivity with MnO4− (Jeppesen and Nielsen, 1989), and this characteristic has since served as the basis for determining which amino acids within a series of σ70 mutants help to stabilize the open complex (Fenton et al., 2000). In addition, quantitative analysis of the DNA fragments generated in related experiments yielded information on the population distribution of alternative protein-DNA conformers (Tsodikov et al., 1998). The hyperreactivity observed for these systems may originate in part from a protein-dependent flipping of adenosine bases out of the DNA helix to expose their complementary T residues (Fenton et al., 2000). Related reaction is evident when T is held in an extrahelical position by enzymes that utilize a base-flipping mechanism during their processing of DNA (Serva et al., 1998; Reddy and Rao, 2000; Fig. 6.6.10). DMS-dependent methylation generates an equally diagnostic pattern of hyper- and hyporeactive G residues within a complex of MutY and DNA, which also involves nucleotide flipping (Chepanoske et al., 1999). At least for MnO4−, the basis of protein-dependent hyperreactivity is not always so evident. Two protein-DNA complexes characterized by X-ray crystallography do not reveal the extensive unpairing of T at sites that were previously shown to have a high sensitivity to MnO4− (König et al., 1996; Bochkarev et al., 1998). A single T proximal to the pseudo-dyad axis of nucleosomes was similarly found to be hyperreactive, but not necessarily extrahelical (Fitzgerald and Anderson, 1999). Perhaps the lysine-rich histone tails were instead able to attenuate the electrostatic repulsion of the anionic reagent and DNA and allow for efficient oxidation of T (Hänsler and Rokita, 1993).
Figure 6.6.10 Base flipping can be detected by chemical modification. KMnO4 was used to detect an extrahelical T formed by duplex DNA in the presence and absence of DNA adenineN6 methyltransferase (M. TaqI), S-adenosyl methionine (SAM), S-adenosyl homocysteine (SAH), and sinefungin (Sin; adenosyl ornithine). Densitometry traces of lanes 2, 3, and 4 are shown to the right. Reprinted from Serva et al. (1998) with permission from Oxford University Press.
Just as a protein may influence chemical modification of DNA, so too may chemical modification of DNA affect its association with a protein (Fig. 6.6.2). Accordingly, an enhancer factor R from Epstein-Barr virus could have been characterized through its ability to protect a region of DNA from DMS-dependent methylation, but, instead, methylation of G N7 was used to determine which sites block binding of the enhancer (Gruffat et al., 1990). Oxidation of T to its glycol derivative (Fig. 6.6.6) has similarly been used for interference analysis of GCN4 binding to the major groove of DNA (Pu and Struhl, 1992). This type of interference experiment represents a convenient and complementary alternative to the protection studies described above. Together, these approaches can establish a compelling model for specific protein-DNA complexes and even help discover new DNAprotein systems. For example, an 80-bp region of DNA and two associated nuclear factors that are involved in transcription control of histone H10 were identified through a combination of techniques including chemical footprinting, UV cross-linking, DMS interference, and sitedirected mutagenesis (Breuer et al., 1993). When studies are based solely on interference
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assays, a series of reagents are often employed. In this manner, results based on DMS and DEPC were sufficient to detect distinct proteinDNA contacts with two consensus-sequence variants recognized by the M-lysozyme downstream enhancer (Nickel et al., 1995). The importance of using multiple reagents is further demonstrated by investigations on a series of hormone-responsive elements. DMS interference suggested that both the glucocorticoid and estrogen receptors form similar contacts to G N7, but MnO4− interference was able to determine that only the glucocorticoid receptor interacts with T (Truss et al., 1991; Fig. 6.6.11). Missing contact probing offers yet another complementary approach for examining protein-DNA complexes. In this case, chemical modification removes, rather than masks, functional groups of DNA that have the potential to interact with protein. A population of DNA lacking individual guanine bases can be generated by treatment with DMS and can be used to screen for individual G residues that are critical for stabilizing protein-DNA interactions. This was first applied to investigations
Figure 6.6.11 Protein contact sites can be mapped onto helical models of duplex DNA. Interference assays were used to differentiate DNA recognition by (A) the glucocorticoid receptor with a half-site of its responsive element and (B) the estrogen receptor with a half-site of its equivalent responsive element. DMS, KMnO4, and ethylnitrosourea were used to probe N7 of G, C5, C6 of T, and phosphate oxygens of the backbone, respectively. Reprinted from Truss et al. (1991) with permission from the American Society for Microbiology.
on the λ phage repressor protein (Brunelle and Schleif, 1987) and has more recently been combined with DMS and KMnO4 interference studies to ascertain the DNA binding properties of a nitrogen regulatory protein (Feng et al., 1993). Missing contact experiments are not limited to depurination of G by DMS. Depurination of G + A by formic acid, depyrimidation of T + C by hydrazine, and depyrimidation of T by KMnO4 have all been used to identify protein-DNA interactions (Brunelle and Schleif, 1987; McBoom and Sadowski, 1994). Any one of these procedures may be adequate to address a particular question on structure, but when used in concert with protection and interference analysis, a broad range of structural characteristics may be described in the absence of crystallography or NMR.
Applications of Chemical Modification In Vivo The vast majority of studies based on chemical modification are performed in vitro for obvious reasons including convenience and simplicity. However, questions regarding biological relevance can linger in the absence of complementary data for equivalent systems in vivo. One of the many procedures available to help forge the necessary connection to cellular conditions is none other than chemical modification. Such an approach might be initially dismissed out of concerns about cell permeability and target specificity, but numerous laboratories have successfully footprinted DNA in vivo. One of the first examples utilized methylation of G N7 by DMS to detect regulatory proteins binding to the lac operon (Nick and Gilbert, 1985). DNA modification depended on the ability of this neutral and low-molecularweight organic reagent to diffuse into bacterial cells. Subsequent mapping of the DNA reaction originally entailed chromatin isolation, restriction, separation, and Southern blotting. Primer extension soon replaced blotting (Borowiec and Gralla, 1986), and more recently ligationmediated PCR and use of AlkA protein, a glycosylase that hydrolyzes DNA at N3-methyl A and N7-methyl G, have been adopted to enhance the detection of chemical probing in vivo (Szabó et al., 2000). Despite the charge and high reactivity of MnO4−, this oxidant has similarly been used to examine complexes of lac promoter and RNA polymerase within Escherichia coli (SasseDwight and Gralla, 1989). The specificity of MnO4− for single-stranded DNA is especially
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Chemical Reagants for Investigating the Major Groove of DNA
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Nakatani, K., Hagihara, S., Sando, S., Miyazaki, H., Tanabe, K., and Saito, I. 2000. Site selective formation of thymine glycol-containing oligodeoxynucleotides by oxidation with osmium tetroxide and a bipyridine-tethered oligonucleotide. J. Am. Chem. Soc. 122:6309-6310. Nawamura, T., Negishi, K., and Hayatsu, H. 1994. 8-Hydroxyguanine is not produced by permanganate oxidation of DNA. Arch. Biochem. Biophys. 311:523-524. Neidle, S. and Stuart, D.I. 1976. The crystal and molecular structure of an osmium bispyridine adduct of thymine. Biochim. Biophys. Acta 418:226-231. NejedlO, K., Kwinkowski, M., Galazka, G., Klysik, J., and Pale?ek, E. 1985. Recognition of the structural distortions at the junctions between B and Z segments in negatively supercoiled DNA by osmium tetroxide. J. Biomol. Struct. Dyn. 3:467-478. NejedlO, K., Skorová, E., Diekmann, S., and Pale?ek, E. 1998. Analysis of a curved DNA constructed from alternating dAn:dTn-tracts in linear and supercoiled form by high resolution chemical probing. Biophys. Chem. 73:205-216. Nick, H. and Gilbert, W. 1985. Detection in vivo of protein-DNA interactions within the lac operon of Escherichia coli. Nature 313:795-798. Nickel, J., Short, M.L., Schmitz, A., Eggert, M., and Renkawitz, R. 1995. Methylation of the mouse M-lysozyme downstream enhancer inhibits heterotetrameric GABP binding. Nucl. Acids Res. 23:4785-4792. Nielsen, P.E. 1990. Chemical and photochemical probing of DNA complexes. J. Mol. Recogn. 3:1-25. O’Halloran, T.V., Frantz, B., Shin, M.K., Ralston, D.M., and Wright, J.G. 1989. The MerR heavy metal receptor mediates positive activation in a topologically novel transcription complex. Cell 56:119-129. Pale?ek, E. 1992a. Probing DNA structure with osmium tetroxide complexes in vitro. Methods Enzymol. 212:139-155. Pale?ek, E. 1992b. Probing of DNA structure in cells with osmium tetroxide-2,2′-bipyridine. Methods Enzymol. 212:305-318. Peattie, D.A. 1979. Direct chemical method for sequencing RNA. Proc. Natl. Acad. Sci. U.S.A. 76:1760-1764. Peattie, D.A. and Gilbert, W. 1980. Chemical probes for higher-order structure in RNA. Proc. Natl. Acad. Sci. U.S.A. 77:4679-4682.
Rahmouni, A.R. and Wells, R.D. 1989. Stabilization of Z-DNA in vivo by localized supercoiling. Science 246:358-363. Reddy, Y.V.R. and Rao, D.N. 2000. Binding of EcoP151 DNA methyltransferase to DNA reveals a large structural distortion within the recognition sequence. J. Mol. Biol. 298:597-610. Roberts, E., Deeble, V.J., Woods, C.G., and Taylor, G.R. 1997. KMnO4 and tetraethylammonium chloride are a safe and effective substitute for OsO4 in solid-phase fluorescent chemical cleavage of mismatch. Nucl. Acids Res. 25:33773378. Ross, S.A. and Burrows, C.J. 1996. Cytosine-specific chemical probing of DNA using bromide and monoperoxysulfate. Nucl. Acids Res. 24:5062-5063. Ross, S.A. and Burrows, C.J. 1997. Bromination of pyrimidines using bromide and monoperoxysulfate: A competition study between cytidine, uridine and thymidine. Tetrahedron Lett. 38:28052808. Rother, K.I., Silke, J., Georgiev, O., Schaffner, W., and Matsuo, K. 1995. Influence of DNA sequence and methylation status on bisulfite conversion of cytosine residues. Anal. Biochem. 231:263-265. Rubin, C.M. and Schmid, C.W. 1980. Pyrimidinespecific chemical reactions useful for DNA sequencing. Nucl. Acids Res. 8:4613-4619. Runkel, L. and Nordheim, A. 1986. Chemical footprinting of the interaction between left-handed Z-DNA and anti-Z-DNA antibodies by diethyl pyrocarbonate carbethoxylation. J. Mol. Biol. 189:487-501. Saluz, H. and Jost, J.-P. 1989. A simple high-resolution procedure to study DNA methylation and in vivo DNA-protein interactions on a single-copy gene level in higher eukaryotes. Proc. Natl. Acad. Sci. U.S.A. 86:2602-2606. Sasse-Dwight, S. and Gralla, J.D. 1989. KMnO4 as a probe for lac promoter DNA melting and mechanism in vivo. J. Biol. Chem. 264:80748081. Scholten, P.M. and Nordheim, A. 1986. Diethyl pyrocarbonate: A chemical probe for DNA cruciforms. Nucl. Acids Res. 14:3981-3993. Sen, D. and Gilbert, W. 1988. Formation of parallel four-stranded complexes by guanine-rich motifs in DNA and its implications for meiosis. Nature 334:364-366. Serva, S., Weinhold, E., Roberts, R.J., and Klimasauskas, S. 1998. Chemical display of thymine residues flipped out by DNA methyltransferease. Nucl. Acids Res. 26:3473-3479.
Chemical and Enzymatic Probes for Nucleic Acid Structure
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Singer, B. and Grunberger, D. 1983. Reactions of directly acting agents with nucleic acids. In Molecular Biology of Mutagens and Carcinogens (B. Singer and D. Grunberger, eds.), Chapter 4, pp. 45-96. Plenum Press, New York.
Vincze, A., Henderson, R.E.L., McDonald, J.J., and Leonard, N.J. 1973. Reaction of diethyl pyrocarbonate with nucleic acid components. Bases and nucleosides derived from guanine, cytosine and uracil. J. Am. Chem. Soc. 95:2677-2682.
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Stevens, S.Y. and Glick, G.D. 1999. Evidence for sequence-specific recognition of DNA by antisingle-stranded DNA autoantibodies. Biochemistry 38:560-568. Szabó, P., Pfeifer, G.P., Miao, F., O’Connor, T.R., and Mann, J.R. 2000. Improved in vivo dimethyl sulfate footprinting using AlkA protein: DNAprotein interactions at the mouse H19 gene promoter in primary embryo fibroblasts. Anal. Biochem. 283:112-116. Timsit, Y. and Moras, D. 1992. Crystallization of DNA. Methods Enzymol. 211:409-429. Truss, M., Chalepakis, G., Slater, E.P., Mader, S., and Beato, M. 1991. Functional interaction of hybrid response elements with wild-type and mutant steroid hormone receptors. Mol. Cell. Biol. 11:3247-3258. Tsodikov, O.V., Craig, M.L., Saecker, R.M., and Record, M.T. 1998. Quantitative analysis of multiple-hit footprinting studies to characterize DNA conformational changes in protein-DNA complexes. J. Mol. Biol. 283:757-769. Tullius, T.D. 1991. The use of chemical probes to analyze DNA and RNA structures. Curr. Opin. Struct. Biol. 1:428-434. Venczel, E.A. and Sen, D. 1993. Parallel and antiparallel G-DNA structures from a complex telomeric sequence. Biochemistry 32:62206228.
Williamson, J.R. and Celander, D.W. 1990. Rapid procedure for chemical sequencing of small oligonucleotides without ethanol precipatation. Nucl. Acids Res. 18:379. Williamson, J.R., Raghuraman, M.K., and Cech, T.R. 1989. Monovalent cation-induced structure of telomeric DNA: The G-quartet model. Cell 59:871-880. Wissmann, A. and Hillen, W. 1991. DNA contacts probed by modification protection and interference studies. Methods Enzymol. 208:365-379. Wong, E. and Giandomenico, C.M. 1999. Current status of platinum-based antitumor drugs. Chem. Rev. 99:2451-2466. Wurdeman, R.L. and Gold, B. 1988. The effect of DNA sequence, ionic strength, and cationic DNA affinity binder on the methylation of DNA by N-methyl-N-nitrosourea. Chem. Res. Toxicol. 1:146-147. Yeung, A.T., Dinehart, W.J., and Jones, B.K. 1988. Modification of guanine bases during oligonucleotide synthesis. Nucl. Acids Res. 16:45394554.
Contributed by Steven E. Rokita University of Maryland College Park, Maryland
Chemical Reagants for Investigating the Major Groove of DNA
6.6.16 Supplement 5
Current Protocols in Nucleic Acid Chemistry
Probing DNA Structure with Hydroxyl Radicals
UNIT 6.7
This unit describes how to use the hydroxyl radical (⋅OH) to gain information on the structure of a DNA molecule. Radiolabeled DNA is allowed to react with the hydroxyl radical, which causes strand breaks in the DNA. The broken DNA strands are separated by length via electrophoresis of the reaction products on a denaturing polyacrylamide gel. The cleavage pattern of the DNA is visualized by exposure of the gel to a phosphor imager plate. Quantitative analysis of the cleavage pattern results in an “image” of the shape of the surface of the DNA molecule. Expertise in the techniques of basic molecular biology is necessary to perform this procedure. CAUTION: This procedure should be performed only by personnel trained in the proper use of the 32P isotope, in Nuclear Regulatory Commission (NRC)–licensed sites. Standard precautions to prevent excessive exposure and radioactive contamination of personnel and equipment must be followed at all times. PREPARATION OF THE HYDROXYL RADICAL CLEAVAGE PATTERN OF A DNA MOLECULE This protocol describes how to perform hydroxyl radical cleavage of a DNA molecule and how to visualize the cleavage pattern by subjecting the reaction products to denaturing gel electrophoresis. A convenient source of the hydroxyl radical is the Fenton reaction of iron(II) EDTA with hydrogen peroxide (Fig. 6.7.1; Udenfriend et al., 1954). Sodium ascorbate is also present in the reaction, to reduce the iron(III) EDTA product of the Fenton reaction back to iron(II) EDTA. Thus, a catalytic cycle is established, which allows the use of very low concentrations of the iron reagent.
BASIC PROTOCOL
Materials 1 ng/µL radiolabeled DNA (5′-end-labeled with 32P on one strand; 3000 Ci/mmol; see, e.g., CPMB UNIT 3.5) dissolved in TE buffer, pH 8.0 (APPENDIX 2A) 1× Fe(II) EDTA (see recipe) 10 mM sodium ascorbate 0.3% (v/v) hydrogen peroxide (H2O2) 100 mM thiourea 3 M sodium acetate (APPENDIX 2A) 85% and 100% (v/v) ethanol Dry ice/acetone bath Formamide-containing dye mixture (e.g., loading buffer, UNIT 10.4) Denaturing poylacrylamide electrophoresis gel (∼30 × 40 × 0.04 cm; APPENDIX 3B) 1.5-mL microcentrifuge tubes, siliconized 25° and 90°C heating blocks SpeedVac evaporator
Figure 6.7.1 The Fenton reaction of iron(II) EDTA with hydrogen peroxide to generate the hydroxyl radical. Contributed by Thomas D. Tullius Current Protocols in Nucleic Acid Chemistry (2001) 6.7.1-6.7.8 Copyright © 2001 by John Wiley & Sons, Inc.
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Phosphor imager with storage phosphor plate that corresponds to the size of the gel Phosphor imaging plate scanner Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (PAGE; APPENDIX 3B), drying of high-percentage acryamide denaturing gels (see Support Protocol 1), and image analysis (optional; see Support Protocol 2) Perform hydroxyl radical cleavage reaction 1. Add 7 µL of 1 ng/µL radiolabeled DNA in 10 mM TE buffer, pH 8.0, to a 1.5-mL siliconized microcentrifuge tube. 2. Place separate drops of the following solutions on the wall of the microcentrifuge tube, above the DNA solution: 1 µL 1× Fe(II) EDTA 1 µL 10 mM sodium ascorbate 1 µL 0.3% H2O2. These solutions make up the cleavage reagent.
3. Close the tube, tap the tube to mix the three drops with the DNA solution, and briefly vortex the solution. 4. Incubate the reaction mixture for 2 min in a heating block at 25°C. The reaction time may need to be adjusted to prevent overdigestion of the sample (see Critical Parameters). If overdigestion is suspected, the intensity of the full-length band in the sample can be compared to a control (step 6) and the reaction time or concentration of cleavage reagents can be reduced as appropriate.
Stop reaction and isolate cleaved DNA 5. Add 7 µL of 100 mM thiourea to the mixture to stop the reaction. Briefly vortex the mixture. 6. Combine 7 µL unreacted radiolabeled DNA with 10 µL TE buffer in a second tube as a control. A control is used to ensure that the starting DNA is intact (i.e., that there is no prior cleavage) and to evaluate the extent of cleavage in the sample (see Critical Parameters).
7. Add 3 µL of 3 M sodium acetate and 100 µL of 100% ethanol to the sample and the control, and place in a dry ice/acetone bath for 1 hr. 8. Microcentrifuge 30 min at maximum speed, 4°C, to pellet the DNA. 9. Rinse the DNA pellet twice with 85% ethanol, centrifuging for 5 min after each rinse. 10. Dry the DNA pellet in a SpeedVac evaporator under vacuum. If the DNA molecule under study is a short oligonucleotide (fewer than ∼20 base pairs), ethanol precipitation often is inefficient at pelleting the DNA. Therefore, for short oligonucleotides, steps 7 to 10 are skipped and the DNA is isolated directly by lyophilization of the sample in a SpeedVac evaporator. The disadvantage of this procedure is that any salts present in the reaction mixture are dried down along with the DNA, and problems with denaturing gel electrophoresis may occur. Such problems include poorly shaped gel bands and marked narrowing of the lane near the bottom of the gel. If these problems occur, steps should be taken to minimize the size of the sample and the concentration of salts in the sample.
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Separate DNA cleavage products by denaturing gel electrophoresis 11. Dissolve the DNA pellet in 4 µL formamide-containing dye mixture. 12. To denature the DNA, heat the DNA sample at 90°C for 3 min, and then quickly cool it by plunging it into ice. 13. Load the DNA sample onto a denaturing polyacrylamide gel that has been prepared in an apparatus suitable for sequencing DNA (see APPENDIX 3B for details on denaturing PAGE). To study short DNA molecules (~20 base pairs), use a high-percentage gel (20% or 25% acrylamide). Longer DNA fragments require a lower-percentage gel (e.g., 8% or 10%). CAUTION: Acrylamide is a neurotoxin; always wear gloves, safety glasses, and a surgical mask when working with acrylamide powder.
14. Electrophorese the sample for a time appropriate to separate the cleavage products of interest (usually ∼2 hr). The time of electrophoresis depends on the percentage of poylacrylamide in the gel, the size of the DNA fragments to be separated, and the current (i.e., at constant voltage, the wattage) at which the gel is run. The current should be set so that the gel runs fairly “hot” (~65°C) to ensure that the DNA sample remains denatured during electrophoresis. Be aware, though, that too much heat will cause the glass plates to crack!
15. Dry the gel using a gel dryer (see Support Protocol 1). 16. Expose the gel to a phosphor imager plate. This can take several hours to a few days, depending on the level of radioactivity of the bands on the gel.
17. Scan the phosphor imager plate. If desired, analysis of the gel can be performed using image analysis software (see Support Protocol 2).
DRYING A HIGH-PERCENTAGE POLYACRYLAMIDE DENATURING GEL Drying a low-percentage polyacrylamide gel (8% to 10% polyacrylamide) is straightforward (see APPENDIX 3B). Drying a high-percentage gel (20% to 25% polyacrylamide) can be tricky, because such a gel does not stick well to the filter paper that is used to pull the gel off the glass plate before drying. High-percentage gels are used for high-resolution separation of shorter DNA molecules (Balasubramanian et al., 1998), and so are important for use in chemical probe experiments to determine DNA structure.The author’s laboratory has developed a procedure that works well for drying high-percentage gels (Shafer et al., 1989). The key to the procedure is to first transfer the gel to plastic wrap, which clings well and provides mechanical support to the gel.
SUPPORT PROTOCOL 1
Materials Gel (see Basic Protocol) Plastic wrap: 12- or 18-in. (∼30- or 45-cm) Saran Wrap or Reynolds film Whatman 3MM filter paper Gel dryer 1. After running the gel, carefully lift the larger glass plate off the gel. 2. Cover the gel with plastic wrap. Trim the edges of the plastic wrap so that the wrap extends 2 in. (5 cm) beyond the edges of the gel plate. 3. Using the plastic wrap to support the gel, turn the gel plate over so that the second glass plate is now on top.
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4. Carefully pull the gel away from the glass plate, starting at the top of the gel. The gel should remain on the plastic wrap.
5. Trim two pieces of Whatman 3MM filter paper to fit the gel, and place on the gel. 6. Carefully invert the filter paper/gel/plastic wrap sandwich so that the plastic wrap is on top. Remove air bubbles by brushing the plastic wrap with a piece of filter paper. 7. Dry the gel using a gel dryer. SUPPORT PROTOCOL 2
ANALYSIS OF THE CLEAVAGE PATTERN While the image of the gel produced in the Basic Protocol can provide much information, hydroxyl radical cleavage patterns often exhibit subtle features. Small differences in cleavage between one nucleotide and another can be difficult to discern by eye alone. Fortunately, the image of the gel that is obtained by the scanning device can be further analyzed using software programs for image analysis. This protocol describes the basic steps involved in quantitative analysis of the cleavage pattern using readily available software. The goal of this method is to deconvolute the closely spaced bands in a cleavage pattern, and integrate each band. The author’s laboratory has found that the Lorentzian function provides a very good analytical description of the bands that are produced by phosphor imager analysis of a gel (Shadle et al., 1997). Each band in the cleavage pattern is fitted by a Lorentzian function using nonlinear least-squares fitting methods. One parameter of the Lorentzian function is directly proportional to the area of the band. These band areas are plotted as a histogram to provide a quantitative cleavage pattern. One benefit of this method of analysis is that once the bands of a lane are deconvoluted and integrated, it is a simple matter to compare one cleavage pattern to another, for normalization, subtraction, and other purposes. A similar but more elaborate whole-band analysis method, using the software program GelExplorer, is described in Shadle et al. (1997). See Internet Resources for the URL of the GelExplorer manual. Materials Computerized image of gel obtained by scanning the storage phosphor plate (see Basic Protocol) Personal computer running software for image analysis: ImageQuant (Molecular Dynamics) Origin, with Peak Fitting Module (OriginLab) Microsoft Excel (optional) 1. Perform a linescan of the gel lane of interest, using ImageQuant. 2. Transfer the linescan to Origin. 3. Using the Peak Fitting Module of Origin, simultaneously fit a Lorentzian function to each peak in the cleavage pattern. 4. Plot the integrals of the bands, using Origin or Microsoft Excel.
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REAGENTS AND SOLUTIONS Use ultrapure (e.g., Milli-Q) water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Fe(II) EDTA, 1× 5× stock solution: Dissolve 0.098 g of ferrous ammonium sulfate hexahydrate [(NH4)2Fe(SO4)2⋅6H2O] and 0.186 g of Na2EDTA⋅2H2O in 50 mL of water. Vortex the solution for several minutes to dissolve as much solid as possible. Filter solution through a 0.45-ìm polypropylene filter—a very light greenish-yellow (or perhaps colorless) solution is obtained (concentration, 5 mM (NH4)2Fe(SO4)2/10 mM EDTA). Store in 100-µL aliquots in microcentrifuge tubes up to several months at –20°C. 1× working solution: Immediately before use, dilute 1 part 5× stock solution with 4 parts water (concentration, 1 mM (NH4)2Fe(SO4)2/2 mM EDTA). COMMENTARY Background Information The structure of any particular doublestranded DNA molecule differs from the canonical B-form structure, primarily due to stacking and other interactions between successive base pairs in the DNA double helix. The shape of the surface of a DNA molecule therefore is not perfectly regular, but instead reflects the underlying sequence-dependent structural variability of the DNA. Such structural variations in DNA, whether large or subtle, may contribute to the ability of a protein to recognize a particular DNA sequence, and so are important to characterize. DNA also can adopt structures radically different from the typical double helix, for example the four-stranded Holliday junction that is an intermediate in genetic recombination. Such unusual DNA structures can be difficult to study by standard methods. The hydroxyl radical has proven to be a highly useful probe of the shape of the surface of DNA. It works by causing strand breaks at each nucleotide in a DNA molecule. The main advantages of the hydroxyl radical as a chemical probe of DNA structure are that it reacts with every nucleotide in a DNA molecule that is exposed to the solvent, that it is a very small molecule, and that fine details of DNA structure can potentially be revealed. This highly reactive free radical causes a DNA strand to break by abstracting a hydrogen atom from a deoxyribose residue in the backbone of the DNA. The resulting strand breaks are detected by electrophoresis of the reaction products on a denaturing gel. Every nucleotide of a double-stranded DNA molecule is susceptible to attack by the hydroxyl radical. However, the nucleotides in a DNA molecule do not all react to precisely the same extent, because
the exposure of each deoxyribose residue to the solvent differs due to sequence-dependent structural variations in the DNA. Quantitation of the cleavage pattern provides an abstract “image” of the shape of the surface of the DNA molecule. Other chemical probes have been developed that recognize specific structural features of DNA, and are described in other units in Chapter 6. The hydroxyl radical has been used to study a wide variety of unusual DNA structures (Price and Tullius, 1992). Two examples are bent A-tract DNA (Burkhoff and Tullius, 1987, 1988; Price and Tullius, 1993) and the fourstranded Holliday junction recombination intermediate (Churchill et al., 1988; Kimball et al., 1990).
Critical Parameters Several parameters are critical for the success of the Basic Protocol. The DNA must be intact before the hydroxyl radical cleavage reaction is conducted. Because the hydroxyl radical cleaves DNA at every accessible nucleotide, any preexisting (background) cuts in the DNA strand interfere with analysis of the hydroxyl radical–induced cuts. To check the integrity of the starting DNA, include in the experiment a control sample of DNA that is not subjected to hydroxyl radical cleavage, and run this sample on the gel adjacent to the experimental samples. The control untreated sample should show no bands other than the band corresponding to the intact full-length DNA molecule. If cleavage is seen in this lane, the DNA should be further purified, or the DNA molecule should be prepared again. The reagents used to initiate hydroxyl radical cleavage must be active. Iron(II) EDTA is
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not indefinitely stable, because iron(II) is subject to oxidation to iron(III) and the eventual formation of a precipitate. Iron(II) EDTA stock solutions should be stored frozen, and thawed for use immediately before the experiment. Sodium ascorbate also is subject to oxidation, both in the solid state and in solution. Oxidized ascorbate will not reduce the iron(III) product of the reaction back to iron(II), and so the cleavage reagent is not active. A fresh bottle of sodium ascorbate is used to prepare a stock solution, which is divided into aliquots in microcentrifuge tubes and kept frozen until needed for an experiment. The DNA must not be overdigested by the hydroxyl radical. The goal is to have one or fewer cleavages in any individual DNA strand. This can be achieved operationally by limiting the reaction so that <30% of the DNA molecules in the sample are cleaved. Thus, 70% of the DNA in the sample will run as intact, fulllength DNA in the gel. If overdigestion is suspected, compare the intensity of the full-length band in a control, untreated DNA sample with the intensity of the full-length band in a sample treated with hydroxyl radical. This can be accomplished by a short exposure of the gel to the phosphor imager plate. The intensity of the treated sample’s full-length band should be no less than 70% of the control band’s intensity. If
Probing DNA Structure with Hydroxyl Radicals
the sample is overdigested, adjust the time of reaction or the concentration of the iron(II) EDTA in the reaction mixture. This unit describes the use of 5′-radiolabeled DNA in a hydroxyl radical cleavage experiment. DNA singly end-labeled at the 3′ end could in principle also be used in this protocol. However, very high-resolution gel electrophoresis shows that a major product of DNA cleavage by the hydroxyl radical is a DNA strand terminated by a 5′ aldehyde group (Fig. 6.7.2; Balasubramanian et al., 1998). A DNA strand with this type of terminus has an anomalous gel mobility, making detailed quantitation of the gel pattern difficult. Therefore 5′-radiolabeled DNA is best for studies of DNA structure using the hydroxyl radical.
Troubleshooting Common problems that may arise in the course of probing DNA structure with hydroxyl radicals, along with their possible causes and solutions, are listed in Table 6.7.1.
Anticipated Results The results of this protocol will be a ladder of bands on a denaturing polyacrylamide gel for a sample treated with the hydroxyl radical, and a single intense full-length band for the control untreated sample. The bands in the
Figure 6.7.2 Hydroxyl radical cleavage using 5′- and 3′-end-labeled DNA. The major product for each is a DNA strand terminated by phosphate at the site of strand breakage. With 3′-labeled DNA, the aldehyde-terminated strand also is produced in substantial amounts, leading to difficulties in quantitation.
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Current Protocols in Nucleic Acid Chemistry
Table 6.7.1 Troubleshooting Guide for Probing DNA Structure with Hydroxyl Radicals
Problem
Possible cause
Solution
No cleavage pattern is observed
Ascorbate reducing agent is inactive due to oxidation (see Critical Parameters)
Prepare a fresh ascorbate solution
DNA sample contains radical scavengers (glycerol is commonly used to stabilize proteins, and sometimes is used in other buffers, and is an excellent radical scavenger)
Remove radical scavengers from the buffer
DNA sample contains too much salt
Perform additional ethanol precipitations to remove salt, or use less salt in the reaction mixture Prepare fresh DNA. Do not store DNA for an extended time once it is radiolabeled, especially at high specific activity, as autoradiolysis can damage the DNA molecule.
Bands are poorly shaped, and become progressively more narrow toward the bottom of the gel Control (untreated) DNA shows cleavage
DNA sample has deteriorated
treated sample should be fairly uniform in intensity, since the hydroxyl radical cleaves each nucleotide in a B-form double helix to a nearly equal extent.
Time Considerations Radiolabeling the DNA, performing the hydroxyl radical cleavage reaction, and isolating the DNA can be accomplished in one working day or less. Preparing and running the denaturing gel to separate the cleavage products takes another day. The time of exposure of the gel to the phosphor plate varies depending on the radioactivity of the sample, but 1 to 3 days is typically sufficient. Quantitative analysis of a gel takes a day.
Literature Cited
Churchill, M.E.A., Tullius, T.D., Kallenbach, N.R., and Seeman, N.C. 1988. A Holliday recombination intermediate is twofold symmetric. Proc. Natl. Acad. Sci. U.S.A. 85:4653-4656. Kimball, A., Guo, Q., Lu, M., Cunningham, R.P., Kallenbach, N.R., Seeman, N.C., and Tullius, T.D. 1990. Construction and analysis of parallel and antiparallel Holliday junctions. J. Biol. Chem. 265:6544-6547. Price, M.A. and Tullius, T.D. 1992. Using the hydroxyl radical to probe DNA structure. Methods Enzymol. 212:194-219. Price, M.A. and Tullius, T.D. 1993. How the structure of an adenine tract depends on sequence context. A new model for the structure of TnAn DNA sequences. Biochemistry 32:127-136. Shadle, S.E., Allen, D.F., Guo, H., Pogozelski, W.K., Bashkin, J.S., and Tullius, T.D. 1997. Quantitative analysis of electrophoresis data: Novel curve fitting methodology and its application to the determination of a protein-DNA binding constant. Nucl. Acids Res. 25:850–861.
Balasubramanian, B., Pogozelski, W.K., and Tullius, T.D. 1998. DNA strand breaking by the hydroxyl radical is governed by the accessible surface areas of the hydrogen atoms of the DNA backbone. Proc. Natl. Acad. Sci. U.S.A. 95:97389743.
Shafer, G.E., Price, M.A., and Tullius, T.D. 1989. Use of the hydroxyl radical and gel electrophoresis to study DNA structure. Electrophoresis 10:397-404.
Burkhoff, A.M. and Tullius, T.D. 1987. The unusual conformation adopted by the adenine tracts in kinetoplast DNA. Cell 48:935-943.
Udenfriend, S., Clark, C.T., Axelrod, J., and Brodie, B.B. 1954. Ascorbic acid in aromatic hydroxylation. J. Biol. Chem. 208:731-739.
Burkhoff, A.M. and Tullius, T.D. 1988. Structural details of an adenine tract that does not cause DNA to bend. Nature 331:455-457.
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Key References Price and Tullius, 1992. See above. A comprehensive description of the use of the hydroxyl radical as a probe of DNA structure.
Internet Resources http://people.bu.edu/tullius/GelExplorer_Manual.pdf A detailed online manual that describes the use of the GelExplorer software for quantitative analysis of electrophoresis gel patterns.
Contributed by Thomas D. Tullius Boston University Boston, Massachussetts
Probing DNA Structure with Hydroxyl Radicals
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Probing RNA Structure and Metal-Binding Sites Using Terbium(III) Footprinting Metal ions play a crucial role in RNA tertiary structure folding by neutralizing and bridging the negatively charged phosphoribose backbone. A folded RNA is stabilized by specific and nonspecific interactions with metal ions. The current unit describes the use of the lanthanide metal ion terbium(III) as a probe of both RNA structure and high-affinity metal-binding sites. Thus, it complements UNIT 6.3, since both the lead (Pb2+) and terbium (Tb3+) aqueous hydroxo complexes cleave the RNA backbone in a similar fashion. Specifically, terbium(III) binds to the same sites on RNA as magnesium(II), but with two to four orders of magnitude higher affinity. Low (micromolar) concentrations of terbium(III) will therefore displace magnesium and result in hydrolytic backbone cleavage at high-affinity magnesium-binding sites. At high (millimolar) concentrations of terbium(III), cleavage occurs in a largely sequence-independent manner, preferentially cutting single-stranded or non-Watson-Crick base-paired regions. The aqueous terbium(III) complex has a near-neutral pKa (∼7.9), which enables it to hydrolyze the RNA backbone around physiological pH. This metal ion–promoted RNA backbone cleavage occurs via deprotonation of the 2′-hydroxyl group and nucleophilic attack of the resulting oxyanion on the adjacent 3′,5′-phosphodiester bond to form 2′,3′-cyclic phosphate and 5′-hydroxyl termini (Figure 6.8.1). Since cleavage is dependent on terbium(III) being able to access the 2′-hydroxyl group, the terbium(III) cleavage pattern can be used to probe the secondary and tertiary structure within an RNA molecule.
UNIT 6.8
BASIC PROTOCOL
In the protocol below, which is a standard procedure for performing terbium(III)-mediated footprinting of an RNA molecule, RNA that has been end-labeled with 32P at either the 5′ or 3′ end is prefolded in an appropriate buffer including divalent cations (generally magnesium). Varying concentrations of terbium(III) are added to the RNA mixture to initiate the strand-scission reaction, and the mixture is incubated 30 min to 2 hr. The reaction is quenched, the cleaved RNA is separated on a denaturing polyacrylamide gel, and the cleavage pattern is visualized by exposure of the gel to a phosphor imager screen. Partial alkaline hydrolysis and ribonuclease (RNase) T1 digestion reactions of the radiolabeled RNA are analyzed alongside the terbium(III)-mediated cleavage reaction to accurately locate the cleavage sites (see UNIT 6.1). The intensities of cleaved bands are quantified and a footprint of the overall RNA molecule is obtained. CAUTION: When working with radioactive materials, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following the guidelines provided by the local radiation safety officer. NOTE: As for experiments with RNA in general, care must be taken to avoid introducing ribonucleases (RNases) into the samples. This can be avoided by wearing gloves to avoid skin contact when handling samples and solutions; by using nuclease-free, sterilized pipet tips, sample tubes, and other disposable plasticware; by preparing all solutions from highest-purity (e.g., molecular-biology grade) components in double-deionized water (18 MΩ conductivity); and by filter-sterilizing (through a 0.22-µm filter) or autoclaving all solutions. Although less desirable, treatment with diethylpyrocarbonate (DEPC; APPENDIX 2A) may be used to block background nuclease activities. Chemical and Enzymatic Probes for Nucleic Acid Structure Contributed by Dinari A. Harris and Nils G. Walter Current Protocols in Nucleic Acid Chemistry (2003) 6.8.1-6.8.8 Copyright © 2003 by John Wiley & Sons, Inc.
6.8.1 Supplement 13
Materials RNA, gel-purified (UNIT 6.1), labeled at 5′ or 3′ end with 32P (conc. ≥50,000 cpm/µL; sp. act. ≥150,000 cpm/pmol) Appropriate buffers to fold RNA (usually Tris, MES, and/or HEPES buffer of desired pH; see Critical Parameters for discussion of optimization) 1 M MgCl2 (APPENDIX 2A) 100 mM TbCl3 (see recipe) 0.5 M EDTA, pH 8.0 (APPENDIX 2A) 3 M sodium acetate (APPENDIX 2A) 80% and 100% (v/v) ethanol Urea loading buffer (APPENDIX 2A) Heating block Water bath at optimized incubation temperature (25° to 45°C) Phosphor screens and phosphor imager with appropriate software (e.g., PhosphorImager Storm 840 with ImageQuant software; Molecular Dynamics) Additional reagents and equipment for partial alkaline hydrolysis and RNase T1 digestion of RNA (UNIT 6.1, Support Protocol 3), and denaturing polyacrylamide gel electrophoresis (APPENDIX 3B) Prepare reaction mixture and terbium(III) dilution series 1. Mix the desired amount of end-labeled RNA with appropriate buffer. Prepare a single pool containing 250,000 to 500,000 cpm (typically 0.5 to 2 pmol) RNA per aliquot, with a total volume that will be sufficient for duplicate 8-µL aliquots at each terbium(III) concentration (see steps 4 and 5). Denature 2 min in a 90°C heating block. A micromolar background of unlabeled RNA can be added to reduce nonspecific cleavage at low terbium(III) concentrations and to facilitate subsequent ethanol precipitation.
2. Slowly renature the RNA by incubating mixture at an optimized temperature between 25° and 45°C (see Critical Parameters) for ∼10 min. 3. Add MgCl2 from 1 M or other appropriately diluted stock solution to obtain desired Mg2+ concentration (see Critical Parameters) and continue to incubate at appropriate temperature for an additional 5 min. 4. Make a serial set of TbCl3 dilutions in water from the 0.1 M TbCl3 stock solution, ranging from micromolar to millimolar concentrations, calculated at 5× the final reaction concentration (see step 5). The TbCl3 dilutions in water should be made immediately prior to use. A serial set of dilutions is recommended to ensure consistency in cleavage band intensity between gel lanes. Use a fresh aliquot of 0.1 M TbCl3 stock solution each time. Final TbCl3 concentrations used in the cleavage reactions should be optimized together with other experimental conditions for the specific RNA and experimental goal (see Critical Parameters).
Run reaction 5. Initiate terbium(III)-mediated cleavage reactions by combining 8 µL RNA (step 3) with 2 µL of each TbCl3 dilution (step 4) to achieve the desired final Tb3+ concentrations (typically 0 to 5 mM). Perform each concentration in duplicate. Continue to incubate at the optimized temperature (25° to 45°C) for an optimized length of time (30 min to 2 hr; see Critical Parameters). Probing RNA Structure and Metal-Binding Sites Using Terbium
Incubation times between 30 min and 2 hr are sufficient to generate a partial digestion pattern of end-labeled RNA under single hit conditions. Avoid more extended incubation times to prevent secondary hits (see Critical Parameters).
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6. Terminate the cleavage reaction by adding 0.5 M EDTA, pH 8.0, to a final concentration of 50 mM (or at least a 2-fold excess over the total concentration of multivalent metal ions). Collect RNA and analyze cleavage patterns 7. Add 3 M sodium acetate to the reaction to final concentration of 0.3 M, then add 2 to 2.5 vol of 100% ethanol and incubate overnight −20°C to precipitate the RNA. Centrifuge 30 min at 12,000 × g, 4°C. Decant supernatant, wash pellet with 80% ethanol, decant supernatant, and dry RNA in a Speedvac evaporator. Redissolve samples in 10 to 20 µL of urea loading buffer. 8. Using separate samples of the same end-labeled RNA, perform partial alkaline hydrolysis and RNase T1 digestion reactions (see UNIT 6.1, Support Protocol 3). Add 10 µL urea loading buffer to an equal volume (10 µL) of the partial hydrolysis reaction and the T1 digestion, and load each directly onto the gel. 9. Place all samples in a heating block at 90°C for 5 min, then snap cool in ice water. Analyze the cleavage products on a high-resolution denaturing (8 M urea) polyacrylamide gel (APPENDIX 3B), using the partial alkaline hydrolysis and RNase T1 digestion reactions as sequencing ladders to identify the specific terbium(III) cleavage products at nucleotide resolution. The authors use a 15% wedged gel, run at a constant power of 80 W, for separating an 80-mer RNA. Identical samples can be loaded at different times on the same gel to resolve different regions of the RNA.
10. Expose the gel to a phosphor imager screen. This can take several hours to overnight, depending on the level of radioactivity of the bands on the gel.
11. Scan the phosphor screen and quantify the full-length RNA and cleavage-product bands using a volume count method. At every Tb3+ concentration, calculate a normalized extent of cleavage (Π) by substituting the peak intensities in the equation:
Π=
band intensity at nucleotide x Σ band intensity at nucleotide i i y [Tb3+ ] band intensity at nucleotide x Σ band intensity at nucleotide i i 0 mM [Tb3+ ]
where y is the terbium(III) concentration in a particular cleavage reaction, x is the analyzed nucleotide position of the RNA, and “0 mM [Tb3+]” signifies a control reaction containing no terbium (III), incubated in the same fashion as reactions containing terbium(III). A Π value of ≥2 indicates significant cleavage over background degradation. By dividing the ratio of a single band intensity over total RNA in the presence of terbium(III) by the ratio of a single band intensity over total RNA in the absence of terbium(III), one normalizes for the effect of nonspecific background degradation. Chemical and Enzymatic Probes for Nucleic Acid Structure
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REAGENTS AND SOLUTIONS Use double-deionized nuclease-free water (18 MΩ conductivity) in all recipes and protocol steps (also see General Considerations for Working with RNA in APPENDIX 2A). To eliminate traces of RNases on glassware, rinse in RNase-free water (APPENDIX 2A) and autoclave or bake 2 hr at 150°C. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Sodium cacodylate buffer, 5 mM (pH 5.5) Prepare 15.9 g/L sodium cacodylate (100 mM final). Adjust to pH 7 with NaOH or cacodylic acid. Dilute 100 mM stock solution to 5 mM, sterilize by filtering with a 0.22-µm filter, and store up to several months at room temperature. CAUTION: Be careful when handling the toxic cacodylate.
Terbium(III) chloride Dissolve 3.73 g solid TbCl3⋅6H2O (Aldrich; 99.9%) in 100 ml of 5 mM sodium cacodylate buffer (see recipe) to give a final concentration of 0.1 M. Filter through 0.22-µm filters into 1-mL aliquots and store up to 1 year at −20°C. The 0.1 M TbCl3 stock solution is prepared in a 5 mM sodium cacodylate buffer at pH 5.5 to prevent precipitation of terbium(III) hydroxide, which occurs at higher pH.
COMMENTARY Background Information
Probing RNA Structure and Metal-Binding Sites Using Terbium
Tertiary structure formation in RNA is highly influenced by metal ions, particularly divalents (Pyle, 2002). The lanthanide metal ion terbium(III) has proven to be a powerful probe of tertiary structure, as well as a tool for mapping metal-binding sites in RNA. Terbium(III) binds to sites on RNA in a manner similar to Mg2+, with an affinity two to four orders of magnitude higher (Walter et al., 2000). Terbium(III) has the ability to substitute for magnesium because hydrated Tb3+ ions have an ionic radius (0.92 Å) similar to that of hydrated Mg2+ (0.72 Å), and because terbium(III) and magnesium share the same preference for coordination to oxygen ligands (Saito and Suga, 2002). There are several classes of metal-binding sites found in RNA, ranging from diffuse to site-specific, and involving both inner- and outer-sphere coordination to the RNA. The most common ligands on the RNA are the nonbridging phosphate oxygens, the purine N7 positions, the base keto groups, and the ribose 2′-hydroxyls. Once bound to RNA, terbium(III) cleaves the phosphodiester backbone by abstracting the 2′-hydroxyl of a nearby nucleotide. This allows for nucleophilic attack of the resulting 2′-oxyanion on the juxtaposed phosphodiester bond, leading to strand cleavage (Ciesiolka et al., 1989; Matsumura and Komiyama, 1997; Figure 6.8.1). Tb3+ can effectively catalyze cleavage of an RNA backbone at physiological pH due to its lower pKa (∼7.9) compared to that of Mg2+ (Sigel et al., 2000).
By varying the concentration of Tb3+ used in the cleavage reaction, it is possible to obtain a map of metal-binding sites and a footprint of secondary and tertiary structure. Micromolar concentrations of Tb3+ bind to high-affinity metal binding sites within a folded RNA, leading to few specific cleavage sites. By contrast, millimolar concentrations of Tb3+ produce a footprinting pattern of solvent-accessible regions, mainly cleaving the RNA backbone in a sequence-independent manner. Under these conditions, the backbone is preferentially cut in single-stranded and non-Watson-Crick basepaired regions, possibly due to the better accessibility of ligands such as N7 positions of purines and the π-electron systems of all nucleobases (Walter et al., 2000). Therefore, it is critical to perform the cleavage reactions over a wide range of Tb3+ concentrations in order to acquire information on both high-affinity metal binding and secondary and tertiary structure folding. Terbium(III)-mediated footprinting and other properties of this lanthanide ion have provided for a versatile probe of RNA structure, function, and metal-ion binding. For example, micromolar concentrations of terbium(III) not only report a metal binding site in loop B of the hairpin ribozyme through backbone cleavage, but also permit the analysis of metal-binding affinity and kinetics through sensitized luminescence, and reversibly inhibit the ribozyme’s catalytic activity by competing for a crucial, yet nonselective, cation-binding site (Walter et al., 2000). Millimolar terbium(III) concentrations,
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O δ− O
base
O
5′
5′ OH
O
P
Tb(OH)(aq)2+
δ−
O −O
pro-Rp
pro-Sp
O
O CH2
5′
base
O
Tb(H2O)(aq)3+
P
O−
base
O
O
O
δ− P O
δ− O
O O
3′
base
OH
BH+
CH2
3′
O
base
OH
HO
CH2
3′
O
base
OH
Figure 6.8.1 Terbium(III)-mediated RNA backbone cleavage. The aqueous terbium(III) hydroxo complex Tb(OH)(aq)2+ deprotonates the 2′-hydroxyl group, allowing for nucleophilic attack of the resulting oxyanion on the adjacent phosphodiester bond. This leads to formation of a trigonal-bipyramidal transition state (center), which, upon protonation by a general acid (BH+), breaks down to form 2′,3′-cyclic phosphate and a 5′-hydroxyl terminus.
in contrast, footprint the hairpin ribozyme’s secondary and tertiary structure and reveal a solvent-protected core similar to that observed in hydroxyl radical footprinting (Walter et al., 2000; also see Figure 6.8.2). At low Tb3+ concentrations, cleavage of human tRNALys,3 is restricted to nucleotides that were previously identified from X-ray crystallography as specific metal-binding pockets (Hargittai and Musier-Forsyth, 2000). The use of higher Tb3+ concentrations resulted in an overall footprint of the L-shaped tRNA structure, showing increased cleavage in the loop regions (D and anticodon loop; Hargittai and Musier-Forsyth, 2000). HIV nucleocapsid protein could then be shown to result in the disruption of the tRNA’s metal-binding pockets and, at higher concentrations, to induce subtle structural changes in, for example, the tRNA acceptor-TψC stem minihelix (Hargittai et al., 2001). Other RNAs that have similarly been studied by terbium(III)-mediated footprinting include the hammerhead (Feig et al., 1999), aminoacyltransferase (Flynn-Charlebois et al., 2001; Vaidya and Suga, 2001), RNase P (Kaye et al., 2002), and group II intron ribozymes (Sigel et al., 2000).
Critical Parameters Tb3+ can be a very useful chemical probe of metal binding and tertiary structure folding in
RNA. However, there are several parameters that are important to consider in order to obtain a reliable and reproducible RNA footprint. Prefolding the RNA under the correct buffer conditions and magnesium concentrations will ensure conformational homogeneity. When optimizing these components, the pH of the cleavage reaction should be kept near physiological pH (7.0 to 7.5), allowing for the accumulation of the cleavage-active Tb(OH)(aq)2+ species (Walter et al., 2000). At a pH above 7.5, Tb3+ increasingly forms insoluble polynuclear hydroxo aggregates (Baes and Mesmer, 1976; Matsumura and Komiyama, 1997), which should be avoided. Precipitation of such insoluble polynuclear species lowers the concentration of the cleavage-active mononuclear species, Tb(OH)(aq)2+, and potentially co-precipitates the RNA. The temperature at which the cleavage reaction is performed must be experimentally determined. Higher temperatures result in faster cleavage rates but also increase the amount of background degradation. Therefore, typical reaction temperatures range from 25° to 45°C. Since Tb3+ has the ability to displace Mg2+ in high-affinity binding pockets while also footprinting solvent-accessible regions, identifying a metal-binding site must be done with care. An effective means of locating a high-affinity binding site is to first decrease the Tb3+
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A Lane no. S(dA −1) S(G+1A) Na+ Mg2+ Co(NH3)63+ spd3+ Tb3+ EDTA
B hinge H3
loop E motif H2
H4 H1
= strong hit = medium hit = weak hit = change upon tertiary structure folding = major hit at mM [Tb3+]
Probing RNA Structure and Metal-Binding Sites Using Terbium
Figure 6.8.2 Terbium(III)-mediated footprinting of the hairpin ribozyme (Walter et al., 2000). (A) Footprint of 5′-32P-labeled hairpin ribozyme (Rz) after incubation with terbium(III) for 2 hr at 25°C. Lanes 1 and 2, controls; lanes 3 and 4, sequencing ladders with RNase U2 (cutting after A) and RNase T1 (cutting after G); lane 5, partial alkali hydrolysis; lanes 6 to 19, order-of-addition experiments (numbers indicate order) with a 100-fold excess (500 nM) of noncleavable substrate analog S(dA–1) or S(G+1A), 100 mM Na+, 12 mM Mg2+, 12 mM Co(NH3)63+, and 12 mM spermidine (spd3+), 12 mM Tb3+, and 120 mM EDTA as chelator; lanes 20 to 27, incubation of 10 µM ribozyme-substrate complex, containing trace amounts of radiolabeled ribozyme, in the presence of 12 mM Mg2+ with varying concentrations of Tb3+, as indicated, and with noncleavable substrate analog, either S(dA–1) or S(G+1A). (B) Superposition of the observed backbone cleavage sites at 12 mM terbium(III) onto a schematic of the hairpin ribozyme. Strong, medium, and weak hits, together with changes caused by tertiary-structure formation, are indicated at the nucleotide 3′ to which cleavage occurs, as deduced from quantitation of lanes 8 (footprint of secondary structure; Tb3+ added before magnesium, thus preventing tertiary structure folding) and 10 (footprint of tertiary structure; magnesium added before Tb3+) in (A). The only major hit at micromolar concentrations of terbium(III) is 3′ to U37. Reproduced from Walter et al. (2000) with permission from Elsevier.
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concentration until a very narrow cleavage pattern is observed (typically at 10 to 100 µM Tb3+), and then to perform a competition experiment with increasing concentrations of Mg2+. This can be done as in the Basic Protocol, but the concentration of Mg2+ is increased against a constant Tb3+ concentration. The residual terbium(III)-mediated cleavage bands should diminish as the Mg2+ concentration is increased. The optimal Tb3+ concentration(s) to use for structure probing are also an important consideration and must be determined for each individual RNA. The trivalent terbium(III) has been shown to induce slight perturbations in the RNA structure (Hargittai and Musier-Forsyth, 2000), but careful titration will reveal the optimal terbium(III):RNA ratio needed for detecting secondary and tertiary structure features in a given RNA molecule.
Anticipated Results A typical result involving both terbium(III)mediated structure probing and metal-bindingsite detection is shown in Figure 6.8.2 for the hairpin ribozyme (Walter et al., 2000). The cleavage pattern on such a denaturing polyacrylamide gel normally shows many bands of varying intensities. These varying intensities (or extents of cleavage) at each nucleotide position relate to the structure of the RNA. The extent of total cleavage should generally be no more than 20% of the uncleaved or full-length band. This ensures that the RNA undergoes a single hit and minimizes the presence of products resulting from multiple cleavage events. The normalized extent of cleavage (Π) is best quantified according to a modified form of the equation used in nucleotide analog interference mapping (NAIM) analysis (Ryder and Strobel, 1999), as given above (see Basic Protocol, step 11). It is important to note that terbium(III) footprinting may not reveal all high-affinity metal binding sites due to the geometrical requirement of Tb3+ accessibility to the 2′-hydroxyl group on the ribose (Figure 6.8.1). The rate and extent of cleavage depend on distance between the terbium(III) and the 2′-hydroxyl of the nearby nucleotide, which is unfavorable in a standard A-type RNA helix. Therefore, strong metal sites that occur in RNA helical regions may go undetected by Tb3+ cleavage (Sigel et al., 2000).
Time Considerations The procedure outlined in the Basic Protocol provides an easy and sensitive means of probing a wide variety of RNA structures. All procedures in this unit can be completed in 2 to 3 days. The most time-consuming aspect involved is the end labeling and careful gel purification of the RNA (UNIT 6.1). The cleavage reactions themselves take no more than 2 hr and are followed by gel electrophoresis (3 to 5 hr) and phosphorimager analysis (6 to 24 hr).
Literature Cited Baes, C.F. and Mesmer, R.E. 1976. The Hydrolysis of Cations. John Wiley & Sons, New York. Ciesiolka, J., Marciniec, T., and Krzyzosiak, W. 1989. Probing the environment of lanthanide binding sites in yeast tRNA(Phe) by specific metal-ion-promoted cleavages. Eur. J. Biochem. 182:445-450. Feig, A.L., Panek, M., Horrocks, W.D. Jr., and Uhlenbeck, O.C. 1999. Probing the binding of Tb(III) and Eu(III) to the hammerhead ribozyme using luminescence spectroscopy. Chem. Biol. 6:801-810. Flynn-Charlebois, A., Lee, N., and Suga, H. 2001. A single metal ion plays structural and chemical roles in an aminoacyl-transferase ribozyme. Biochemistry 40:13623-13632. Hargittai, M.R. and Musier-Forsyth, K. 2000. Use of terbium as a probe of tRNA tertiary structure and folding. RNA 6:1672-1680. Hargittai, M.R., Mangla, A.T., Gorelick, R.J., and Musier-Forsyth, K. 2001. HIV-1 nucleocapsid p r o tein zinc fin ger structures ind uce tRNA(Lys,3) structural changes but are not critical for primer/template annealing. J. Mol. Biol. 312:985-997. Kaye, N.M., Zahler, N.H., Christian, E.L., and Harris, M.E. 2002. Conservation of helical structure contributes to functional metal ion interactions in the catalytic domain of ribonuclease P RNA. J. Mol. Biol. 324:429-442. Matsumura, K. and Komiyama, M. 1997. Enormously fast RNA hydrolysis by lanthanide(III) ions under physiological conditions: Eminent candidates for novel tools of biotechnology. J. Biochem. 122:387-394. Pyle, A.M. 2002. Metal ions in the structure and function of RNA. J. Biol. Inorg. Chem. 7:679690. Ryder, S.P. and Strobel, S.A. 1999. Nucleotide analog interference mapping. Methods 18:38-50. Saito, H. and Suga, H. 2002. Outersphere and innersphere coordinated metal ions in an aminoacyltRNA synthetase ribozyme. Nucl. Acids Res. 30:5151-5159. Sigel, R.K., Vaidya, A., and Pyle, A.M. 2000. Metal ion binding sites in a group II intron core. Nat. Struct. Biol. 7:1111-1116.
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Vaidya, A. and Suga, H. 2001. Diverse roles of metal ions in acyl-transferase ribozymes. Biochemistry 40:7200-7210. Walter, N.G., Yang, N., and Burke, J.M. 2000. Probing non-selective cation binding in the hairpin ribozyme with Tb(III). J. Mol. Biol. 298:539555.
Contributed by Dinari A. Harris and Nils G. Walter University of Michigan Ann Arbor, Michigan
Probing RNA Structure and Metal-Binding Sites Using Terbium
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Probing RNA Structure and Function by Nucleotide Analog Interference Mapping
UNIT 6.9
The chemistry of phosphorothioate interference has been expanded into a method to identify the specific chemical groups that are important for RNA activity (Eckstein, 1985; Gaur and Krupp, 1993; Strobel and Shetty, 1997). This approach, termed nucleotide analog interference mapping (NAIM), makes it possible to simultaneously, yet individually, test the contribution of a particular functional group at almost every position within the RNA molecule in an assay that is as simple as RNA sequencing. In a NAIM experiment the smallest mutable unit is not the base pair, but rather the individual functional groups that comprise the nucleotide. Because the deletion or modification of particular functional groups within an RNA can severely affect its activity, this approach makes it possible to determine the chemical basis of RNA structure and function (Ryder et al., 2000). Using phosphorothioate-tagged nucleotide analogs, all of the positions on the RNA can be simultaneously characterized. NAIM involves a four-step process: (1) the phosphorothioate-tagged nucleotide analog is randomly incorporated into the RNA by in vitro transcription; (2) the active members of the RNA population are selected from inactive transcripts; (3) the phosphorothioate linkage is cleaved by iodine treatment; and (4) the sites of analog substitution detrimental to activity are identified by gel electrophoresis and autoradiography. The approach is applicable to any RNA that can be transcribed in vitro and where active variants can be selected in some fashion. RNA functions include catalysis, protein or ligand binding, folding, and others.
STRATEGIC PLANNING Phosphorothioate-Tagged Nucleotide Analog Triphosphates (NTPαS) NAIM relies on the use of nucleotides in which one of the non-bridging α-phosphate oxygen atoms has been replaced with a sulfur (Eckstein, 1985). This modification does not drastically alter the RNA structure in the vast majority of cases, and any effects resulting from the sulfur are readily controlled. The NTPαS is randomly incorporated into the RNA molecule by in vitro transcription. The sulfur facilitates the direct detection of sites sensitive to analog incorporation because the phosphorothioate linkage can be selectively cleaved with iodine without affecting any of the other bonds within the RNA (Gish and Eckstein, 1988). The RNA fragments are separated by denaturing gel electrophoresis to identify the extent of analog incorporation and magnitude of interference at each position within the transcript. NAIM uses both parent and analog phosphorothioate-tagged nucleotides (Ryder et al., 2000). Any effects that arise from the phosphorothioate tag are controlled by comparison to the band intensity from the four parental 5 -O-(1-thio)nucleotide triphosphates (AαS, CαS, GαS, and UαS). The phosphorothioate substitution introduces an additional chiral center into the nucleotide. Synthesis of the phosphorothioate-tagged nucleotides results in a diastereomeric mixture of RP and SP isomers, of which only the SP diastereomer is recognized by RNA polymerase (Chamberlin et al., 1983; Griffiths et al., 1987). The transcription reaction proceeds with inversion of configuration at the α-phosphate, resulting in RNAs with RP phosphorothioate substitutions. It is possible to purify the SP diastereomer of the triphosphate by HPLC (Fischer et al., 1999), but the RP diastereomer is not recognized by the polymerase, nor does it inhibit the transcription reaction; therefore, its inclusion in the in vitro transcription reaction has no significant consequence. Contributed by Jesse C. Cochrane and Scott A. Strobel Current Protocols in Nucleic Acid Chemistry (2004) 6.9.1-6.9.21 C 2004 by John Wiley & Sons, Inc. Copyright
Chemical and Enzymatic Probes for Nucleic Aicd Structure
6.9.1 Supplement 17
Analog nucleotides contain modifications to the sugar or the base in addition to the phosphorothioate tag (Ryder et al., 2000). The range and variety of modifications are summarized in Table 6.9.1. Any analog that is amenable to transcriptional incorporation by RNA polymerase, even at low efficiency, can be used in this approach. To date, twelve adenosine analogs have been utilized in NAIM (Conrad et al., 1995; Hardt et al., 1996; Ortoleva-Donnelly et al., 1998; Soukup et al., 2002; Jones and Strobel, 2003; Schwans et al., 2003). Eight analogs modify the nucleotide base and four modify the ribose sugar. The base analogs include purine riboside (PurαS), N-methyladenosine (m6 AαS), 7-deazaadenosine (7dAαS), diaminopurine riboside (DAPαS), 2-aminopurine riboside (2APαS), formicin (FormαS), 3-deazaadenosine (c3 AαS), and 8-azaadenosine (n8 AαS). The ribose sugar analogs all modify the 2 -OH group and include 2 -deoxyadenosine (dAαS), 2 -deoxy-2 -fluoroadenosine (F AαS), 2 -O-methyladenosine (OMe AαS), and 2 deoxy-2 -thioadenosine (SH AαS). Each of these analogs provides specific information about the chemical basis of RNA activity at almost every incorporated position in the transcript. PurαS, 2APαS, and m6 AαS measure the effect of modifications to the N6 exocyclic amine of adenosine. The base analog 7dAαS replaces the N7 nitrogen with a C-H group. Interference with this analog indicates an important major groove contact to the ring nitrogen. Additionally, 7dAαS, FormαS, and n8 AαS can be used to probe important protonation events. DAPαS and 2APαS both add an additional amine to the C2 position of adenosine. dAαS interference identifies the 2 -OH groups important for RNA function, while F AαS delineates the role these 2 -OH groups play as either hydrogen bond donors or hydrogen bond acceptors. Seven guanosine analogs have been utilized in NAIM (Hardt et al., 1996; Strobel and Shetty, 1997; Basu et al., 1998; Kazantsev and Pace, 1998; Schwans et al., 2003). Four analogs modify the base functional groups of G, including inosine (IαS), N2 methylguanosine (m2 GαS), 7-deazaguanosine (7dGαS), and 6-thioguanosine (s6 GαS). Three analogs, 2 -deoxyguanosine (dGαS), 2 -O-methylguanosine (OMe GαS), and 2 deoxy-2 -thioguanosine (SH GαS), modify the 2 -OH of the ribose ring. IαS and m2 GαS both modify the N2 exocyclic amine of G. m2 GαS substitution is isoenergetic with G in the context of G·C, G·U, and G·A base pairs, and m2 G interference has only been observed at sites of tertiary hydrogen bonding (Rife et al., 1998). The base analog 7dGαS, like 7dAαS, replaces the N7 nitrogen with a C-H group. s6 GαS also modifies a major groove functional group. It introduces an oxygen-to-sulfur substitution at the O6 keto group of G. dGαS, like dAαS, replaces the 2 -OH with a proton. Thirteen pyrimidine analogs have been used, including seven cytidine and six uridine analogs (Ryder and Strobel, 1999b; Szewczak et al., 1999; Oyelere et al., 2002; Schwans et al., 2003). These include seven analogs that modify the 2 -OH group of the ribose: 2 -deoxyuridine (dUαS), 2 -deoxy-2 -fluorouridine (F UαS), 2 -Omethyluridine (OMe UαS), 2 -deoxy-2 -thiouridine (SH UαS), 2 -deoxycytidine (dCαS), 2 -O-methylcytidine (OMe CαS), and 2 -deoxy-2 -thiocytidine (SH CαS). This series of analogs can be used to address the same types of questions as outlined for the A analogs. Two additional uridine derivatives have been used in NAIM, 5-methyluridine (m5 UαS) and pseudouridine (αS). m5 UαS introduces steric bulk in the major groove and probes for possible hydrophobic contacts to the base. αS changes the 5 position of U to a nitrogen and is a C-linked nucleoside. Four cytidine analogs have been synthesized for use in NAIM experiments, 5-fluorocytidine (f5 CαS), 6-azacytidine (n6 CαS), pseudoisocytidine (iCαS), and zebularine (ZαS). All four of these analogs have altered pKa values. Nucleotide Analog Interference Mapping
Some of the phosphorothioate-tagged nucleotide analog triphosphates are available from a variety of commercial sources. TriLink Biotechnologies sells the parental and 2 -deoxyphosphorothioates, Amersham-Pharmacia sells two 2 -deoxy-phosphorothioates, and
6.9.2 Supplement 17
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Table 6.9.1 Phosphorothioate-Tagged Nucleotides for NAIM
[δTPαS] [NTP] (mM)b (mM)a
Polymerase (WT or Description Y639F)
Reference
0.05
1.0
WT
Christian and Yarus (1992)
0.05
1.0
WT
Replaces N7 with a carbon
Ortoleva-Donnelly et al. (1998)
m AαS (Sp )
0.1
1.0
WT
Ortoleva-Donnelly et al. (1998)
DAPαS (Sp )
0.025
1.0
WT
PurαS (Sp )
2.0
1.0
WT
2APαS (Sp )
0.5
1.0
WT
FormAαS
0.1
1.0
WT
c3 AαS
2.0
0.5
Y639F
Replaces a hydrogen on exocyclic amine with methyl Replaces H2 with exocylic amine Replaces exocyclic amine with a hydrogen Replaces exocyclic amine with hydrogen and H2 with exocylic amine Replaces N9 with carbon and C8 with nitrogen Replaces N3 with carbon
Ryder et al. (2001)
0.5
1.0
WT
Replaces C8 with nitrogen
Ryder et al. (2001)
Analog δTPαS AαS (Sp isomer only)c 7dAαS (Sp ) 6
8
n AαS
Strobel and Shetty (1997) Ortoleva-Donnelly et al. (1998) Ortoleva-Donnelly et al. (1998)
Ryder et al. (2001)
dAαS (Sp )
0.8
1.0
Y639F
Replaces 2 -OH with hydrogen Ortoleva-Donnelly et al. (1998)
OMe
2.0
0.2
Y639F
0.25
1.0
Y639F
Replaces hydrogen on 2 -OH with methyl Replaces 2 -OH with fluorine
F
AαS (Sp )
AαS (Sp )
SH
AαS
Ortoleva-Donnelly et al. (1998)
0.4
1.0
Y639F
0.05
1.0
WT
0.05
1.0
WT
Replaces N7 with a carbon
m GαS (Sp )
0.1
1.0
Y639F
IαS (Sp )
0.1
1.0
WT
s6 GαS (Sp , 4 mM Mn2+ ) dGαS (Sp )
0.25
1.0
WT
Replaces a hydrogen on Ortoleva-Donnelly et al. (1998) exocyclic amine with methyl Replaces exocyclic amine with Strobel and Shetty (1997) hydrogen Replaces O6 with sulfur Basu et al. (1998)
0.25
1.0
Y639F
Replaces 2 -OH with hydrogen Szewczak et al. (1998)
OMe
2.0
0.1
Y639F
0.5
1.0
Y639F
Replaces hydrogen on 2 -OH with methyl Replaces 2 -OH with 2 -SH
GαS (Sp isomer only)c 7dGαS (Sp ) 2
SH
GαS (Sp )
GαS
CαS (Sp isomer only)c f5 CαS
0.05
1.0
WT
1.0
1.0
WT
n6 CαS
1.0
0.5
WT
iCαS
0.1
1.0
WT
ZαS
1.0
0.5
Y639F
Replaces 2 -OH with 2 -SH
Ortoleva-Donnelly et al. (1998)
Schwans et al. (2003) Christian and Yarus (1992) Kazantsev and Pace (1998)
(unpub. observ.) Schwans et al. (2003) Christian and Yarus (1992)
Replaces C5 hydrogen with fluorine Replaces C6 with nitrogen
Oyelere and Strobel (2000) Oyelere and Strobel (2000)
Replaces N1 with carbon and Oyelere and Strobel (2000) C5 with nitrogen Replaces N1 with carbon and Szewczak et al. (1999) exocyclic amine with hydrogen continued
Chemical and Enzymatic Probes for Nucleic Aicd Structure
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Supplement 17
Table 6.9.1 Phosphorothioate-Tagged Nucleotides for NAIM, continued
Analog δTPαS
[δTPαS] [NTP] (mM)b (mM)a
Polymerase (WT or Description Y639F)
dCαS (Sp )
0.75
1.0
Y639F
Replaces 2 -OH with hydrogen Ryder and Strobel (1999b)
OMe
2.0
0.05
Y639F
0.4
1.0
Y639F
Replaces hydrogen on 2 -OH with methyl Replaces 2 -OH with 2 -SH
SH
CαS
CαS
Reference
Conrad et al. (1995) Schwans et al. (2003)
UαS (Sp isomer only)c m5 UαS
0.05
1.0
WT
1.0
1.0
WT
Replaces H5 with methyl
Ryder and Strobel (1999a)
αS
0.1
1.0
WT
Replaces C5 with amine
Ryder and Strobel (1999a)
Christian and Yarus (1992)
dUαS (Sp )
0.25
1.0
Y639F
Replaces 2 -OH with hydrogen Szewczak et al. (1998)
OMe
2.0
0.1
Y639F
0.25
1.0
Y639F
Replaces hydrogen on 2 -OH with methyl Replaces 2 -OH with fluorine
F
UαS
UαS
SH
UαS
0.4
1.0
Y639F
Replaces 2 -OH with 2 -SH
Conrad et al. (1995) Szewczak et al. (1998) Schwans et al. (2003)
a Final (1×) concentration of phosphorothioate-tagged analog in transcription reaction. b Final (1×) concentration of untagged, unmodified nucleotide in same transcription reaction. c Phosphorothioate-tagged parent nucleotide (NTPαS) used for control transcript.
Ambion sells the parental phosphorothioates. Currently, there are 18 phosphorothioatetagged nucleotide triphosphates available from Glen Research. Parental nucleotides are provided as the purified SP diastereomer, while nucleotide analogs are typically a diasteromeric mixture. Many more nucleotide analogs can be utilized in a NAIM experiment than are available commercially. In the vast majority of cases, it is relatively straightforward to synthesize the phosphorothioate-tagged triphosphate beginning with the unprotected nucleoside (Fischer et al., 1999; Ryder et al., 2000). A wide variety of nucleoside analogs are commercially available from sources such as Sigma, Berry & Associates, or RI Chemical. In the first step, the nucleoside is converted to the 5 -O-(1,1-dichloro-1thio)phosphorylnucleoside using PSCl3 . In the second step, the nucleotide is converted to the cyclotriphosphate intermediate by the stepwise addition of tributylammonium pyrophosphate, and the triphosphate is generated by hydrolysis with aqueous triethylammonium bicarbonate. A detailed synthesis with improved overall yields has been published by Fischer et al. (1999).
Nucleotide Analog Interference Mapping
RNA Transcripts for NAIM RNA molecules with randomly incorporated phosphorothioate nucleotides are readily prepared using run-off transcription from a DNA template, either a cleaved plasmid or a synthetic oligonucleotide. A Support Protocol presents a brief method for transcription and purification. Complete protocols for these activities are found elsewhere in this manual and in Current Protocols in Molecular Biology. (For digestion of plasmid DNA with restriction enzymes, see CPMB UNIT 3.1; for in vitro transcription of RNA, see CPMB UNIT 3.8.) For the most part, the analogs are incorporated into the RNA by the standard procedure for run-off transcription (see CPMB UNIT 3.8), except that a phosphorothioatetagged analog triphosphate is included in the reaction. Depending upon the analog, it may also be necessary to make slight adjustments to the standard transcription cocktail, such as reduction of the non-tagged nucleotide triphosphate concentration, inclusion of
6.9.4 Supplement 17
Current Protocols in Nucleic Acid Chemistry
Mn2+ in the buffer, or use of a mutant polymerase. These adjustments are summarized in Table 6.9.1 for each of 36 analogs. The conditions described in Table 6.9.1 should be viewed as a guideline. These values were derived from transcription of the Tetrahymena group I intron, but each RNA molecule has slightly different conditions under which it is efficiently transcribed. This necessitates transcriptional optimization of the RNA and optimization of the incorporation of individual analogs during the initial stages of the project. Phosphorothioate incorporation levels should be kept at ∼5% (one analog incorporated per 20 occurrences of that residue in the sequence). The nature of the modification affects the concentration at which the nucleotide must be included to achieve this level of incorporation. Incorporation levels <5% make it difficult to detect the cleavage signal, while over-incorporation can cause multiple cleavage events within a single RNA. This is manifested as a phosphorothioate cleavage ladder in which the bands at the bottom of the gel are substantially more intense than those closer to the top. The wildtype form of the T7 RNA polymerase is incapable of incorporating several analogs, most notably the 2 -deoxy nucleotides. Several of the transcriptionally resistant analogs can be incorporated using the Y639F mutant of the T7 RNA polymerase, a variant originally reported by R. Sousa and co-workers for its ability to synthesize DNA polymers (Sousa and Padilla, 1995; Padilla and Sousa, 1999). This mutant is able to incorporate several analogs with modifications in the minor groove (Soukup et al., 2002). It is sold commercially by Epicentre Technologies under the name T7 R&DNA Polymerase.
Radiolabeling RNA Transcripts and Substrates See UNIT 6.1 for both 3 -end-labeling using [32 P]pCp and T4 RNA ligase, and 5 -endlabeling using [γ-32 P]ATP and T4 polynucleotide kinase. The protocols outlined in UNIT 6.1 utilize calf intestinal alkaline phosphatase to dephosphorylate the 5 ends of the RNA transcripts prior to 5 -end-labeling. RNAs without a 5 -phosphate can also be successfully generated by including guanosine in the transcription reaction. A guanosine concentration is chosen that is two to three times that of the GTP concentration, up to the saturating guanosine concentration of 20 mM, to maximize the number of molecules with a free 5 -OH. Choice of Selection Method and Expanded Applications NAIM requires an assay to distinguish active from inactive variants within a substituted RNA population. In general, two types of selection method have been employed. The first is to physically separate active RNAs by gel mobility shift, column chromatography, filter binding, or denaturing polyacrylamide gel electrophoresis (Boudvillain and Pyle, 1998; Kazantsev and Pace, 1998; Basu and Strobel, 1999; Batey et al., 2000; Szewczak et al., 2002). This approach has been used in NAIM experiments to study protein-RNA interactions, RNA-RNA interactions, RNA folding, and ribozyme cleavage activity. The second approach is specific to analysis of ribozymes, wherein the inherent reactivity of the RNA is used to selectively radiolabel or change the size of active members in the population (Strobel and Shetty, 1997; Ryder et al., 2001). The Basic Protocols presented here outline the methodology used initially to define the important functional groups within an RNA molecule (Ryder and Strobel, 1999a). The first three protocols present methods for selecting RNA based on ligand binding activity (see Basic Protocol 1), ribozyme activity (see Basic Protocol 2), or RNA structure or folding (see Basic Protocol 3). Basic Protocol 4 describes how the sites of interference are identified after RNA selection is complete.
Chemical and Enzymatic Probes for Nucleic Aicd Structure
6.9.5 Current Protocols in Nucleic Acid Chemistry
Supplement 17
Four Alternate Protocols are also presented that expand upon the information gained from the basic NAIM experiment. These include: (1) identification of tertiary contacts within a folded RNA by nucleotide analog interference suppression (NAIS; see Alternate Protocol 1; Szewczak et al., 1998); (2) identification of residues with a functionally important pKa perturbation (see Alternate Protocol 2; Oyelere et al., 2002); (3) identification of functional groups likely to coordinate monovalent or divalent metal ions (see Alternate Protocol 3; Basu and Strobel, 2001); and (4) quantitation of the free energy contribution made by individual functional groups for ligand affinity (see Alternate Protocol 4; Cochrane et al., 2003). Many of the experiments suggested here can be optimized for the RNA of interest. The selectivity of the procedure can be adjusted using various salt concentrations, ligand concentrations, and temperatures as desired. Each case must be considered on an individual basis. In the experiments outlined below, a starting point for selectivity is provided, but this number is just a recommendation that provides a balance between being too permissive and providing a reasonable signal for NAIM analysis. BASIC PROTOCOL 1
SELECTION FOR LIGAND BINDING There are many ways in which selection for ligand binding can be accomplished. In this protocol, filter binding allows for the selective retention of molecules that are able to perform a particular binding activity. Column chromatography, in which the ligand has been immobilized on beads, can be used for the same purpose. Native gel electrophoresis (see Basic Protocol 3) is another way in which RNA molecules that are able to bind ligand can be separated from those that cannot. The selectivity of the ligand binding assay can be adapted to the peculiarities of any RNA-ligand system. Obviously, variations in the concentration of the RNA or the ligand will result in different levels of selectivity in the assay. A high RNA concentration and low ligand concentration will lead to the most selective conditions, while a high ligand concentration and low RNA concentration may mask all interferences. A ligand concentration approximately equal to the Kd for the complex and an RNA concentration ten-fold lower than the ligand results in ∼50% of the RNA molecules being bound in a noncompetitive selection, which is ideal for the first sets of interference experiments. Filter binding is a relatively straightforward method for separating molecules that are able to bind ligand from those that are not (Wong and Lohman, 1993). In this approach, the ligand must be able to bind efficiently to a nitrocellulose filter. The RNA is incubated with a ligand and then applied to a nitrocellulose membrane. The RNA is eluted from the filter, cleaved with iodine, and the fragments separated by gel electrophoresis. The specific activity and concentration of the RNA will dictate the volume of RNA-ligand mixture that will need to be applied to the membrane. The minimum activity required for visualization in a NAIM experiment is 5000 cpm in the final loaded sample.
Materials
Nucleotide Analog Interference Mapping
Wash buffer (see recipe) Transcribed ribozyme RNA containing phosphorothioate-tagged analogs (see Support Protocol and Strategic Planning) Stock of ligand 500 mM Tris/HEPES buffer, pH 7.5 (see recipe) 2 M KCl 1 M MgCl2 0.1 M dithiothreitol (DTT)
6.9.6 Supplement 17
Current Protocols in Nucleic Acid Chemistry
0.1% Igepal C-680 1 mg/mL tRNA Filter elution buffer (see recipe) Ethanol (EtOH), cold Nitrocellulose membrane (Fisher Scientific) Hybond-N+ nylon filter (Amersham Pharmacia Biotech) 90◦ C heating block 1.5-mL microcentrifuge tubes Whatman paper Glass plate Platform rocker Additional reagents and equipment for denaturing filter binding assays (see CPMB UNIT 12.8) and denaturing gel electrophoresis (see APPENDIX 3B) 1. Soak nitrocellulose and Hybond-N+ filters in wash buffer. The Hybond-N+ filter retains RNA not bound to ligand.
2. Heat the labeled RNA for 1 min at 90◦ C, then place on ice for at least 3 min. 3. In a 1.5-mL tube incubate:
0.1× Kd labeled RNA 1× Kd ligand 20 mM Tris/HEPES buffer, pH 7.5 200 mM KCl 10 mM MgCl2 1 mM DTT 0.01% Igepal C-680 0.1 mg/mL tRNA. Incubations should reach equilibrium. For many RNA-ligand complexes, 1 hr at room temperature is reasonable.
4. Assemble a filter binding apparatus as diagrammed in CPMB UNIT 12.8. Place a piece of Whatman paper directly on the bottom plate, then the Hybond-N+ membrane and then the nitrocellulose membrane. Place the top plate on the apparatus and clamp it. 5. Pipet 40 µL of the reaction mixture (from step 3) into the wells of the filter binding apparatus. For an experiment with ∼10 pM RNA, 10,000 cpm/pmol and a ligand concentration around the Kd , twelve wells are needed to have sufficient signal in the final stages of the experiment.
6. Wash each well with 100 µL wash buffer. 7. Remove the top plate of the filter binding apparatus. Allow the filters to air dry for 1 to 2 min. 8. Remove the nitrocellulose filter and place it on a glass plate. Cut out the individual dot blots from the filter. 9. Place the blots into a 1.5-mL microcentrifuge tube and add 700 µL filter elution buffer. Rock the tube for 1 hr to allow the RNA to elute from the filter. 10. Remove the elution buffer from the tube and divide into two equal fractions.
Chemical and Enzymatic Probes for Nucleic Aicd Structure
6.9.7 Current Protocols in Nucleic Acid Chemistry
Supplement 17
11. Ethanol precipitate the RNA by adding 3 vol cold EtOH and incubating at −80◦ C for 15 min. Centrifuge 15 min at 17,000 × g, 25◦ C. Save the pellet and allow the RNA to air dry for 5 to 10 min. 12. Determine the positions of interference (see Basic Protocol 4). BASIC PROTOCOL 2
ACTIVITY SELECTION FOR RIBOZYMES Active ribozymes can be selected from an RNA population based on a change in the size of the molecule or transfer of the radiolabeled substrate onto itself (Cech, 1990). A general set of conditions for performing a ribozyme catalysis reaction is presented here, but it should be recognized that some molecules are more reactive than others and may require other co-factors, higher or lower reaction temperatures, or longer incubation times to achieve between 20% and 40% reacted ribozyme. To use a ribozyme in a NAIM experiment, it must be self-catalytic either in ligation or cleavage (Jones et al., 2001). The main difference between these two activities is that the ribozyme must be labeled in the case of cleavage, while either the ribozyme or the substrate may be labeled in the case of the ligation. Ribozymes and substrates may be labeled at either the 5 or 3 end; however, during the course of the reaction, the label must either be retained by the ribozyme or be transferred to the ribozyme.
Materials Transcribed ribozyme RNA containing phosphorothioate-tagged analogs (see Support Protocol and Strategic Planning) 500 mM HEPES buffer, pH 7.0 10 mM MgCl2 10 mM Mn(OAc)2 Labeled substrate molecule (optional; see Strategic Planning) 2× formamide loading buffer (FLB; APPENDIX 2A) Gel elution buffer (see recipe) Ethanol (EtOH), cold 0.65-mL microcentrifuge tubes 37◦ and 50◦ C heating blocks Autoradiography film Platform rocker Additional reagents and equipment for denaturing gel electrophoresis (see APPENDIX 3B) 1. In a 0.65-mL microcentrifuge tube, mix:
5 µL 1.0 µM RNA transcript 1 µL 500 mM HEPES buffer, pH 7.0 3 µL 10 mM MgCl2 1 µL 10 mM Mn(OAc)2 . Incubate reaction 10 min at 50◦ C. Nucleotide Analog Interference Mapping
This is the RNA folding step. Alternative temperatures, incubation times, and salt concentrations should also be used as appropriate for the RNA of interest.
2. For ligating ribozymes, add 10 µL of preheated (37◦ C) 10 nM substrate dissolved in 50 mM HEPES buffer, pH 7.0, 3 mM MgCl2 , and 1 mM Mn(OAc)2 .
6.9.8 Supplement 17
Current Protocols in Nucleic Acid Chemistry
3. Incubate the reaction at 37◦ C for a time sufficient for 20% to 40% of the ribozyme to react. To determine the length of time required to achieve 20% to 40% reactivity, monitor the reaction by taking aliquots at time intervals over several minutes. Electrophorese the time points on a denaturing polyacrylamide gel and determine the fraction reacted over time.
4. Quench the reaction in 2 vol FLB. If the label was transferred from the substrate to the ribozyme, the sample can be used directly to determine the sites of interference. Otherwise, it is necessary to physically separate the active from inactive fraction based upon the size of the RNA using the radiolabel to visualize the active population.
5. For all experiments in which the initial label was on the ribozyme, separate the reaction products from the substrates by denaturing polyacrylamide gel electrophoresis (see APPENDIX 3B). 6. Expose the gel to autoradiography film and excise the RNA from the gel by physically cutting the products from the gel. Elute RNA in 700 µL gel elution buffer by rocking for 2 to 4 hr at room temperature. Shorter incubation times are used to minimize the extent of RNA degradation. Lower temperatures and inclusion of detergents (1% SDS) in the elution buffer can also be used toward this goal.
7. Ethanol precipitate the RNA by adding 3 vol cold EtOH and incubating 15 min at −80◦ C. Centrifuge 15 min at 17,000 × g, 25◦ C. Save the pellet and allow the RNA to air dry for 5 to 10 min. 8. Determine the extent of interference at all positions (see Basic Protocol 4).
SELECTION FOR STRUCTURE OR FOLDING Selecting for structured RNA molecules is a bit more difficult than selecting for RNA molecules that are able to perform a catalytic activity, as there is no length difference between active and inactive molecules. Native gel electrophoresis conditions that separate the RNA based upon shape must be identified (Basu and Strobel, 1999). Ideal separation conditions for a particular RNA system are often variable, but, in general, the gel solution should not contain denaturants such as urea, and should include polycations such as magnesium to promote folding of the RNA. The electrophoresis is typically performed at a lower voltage and the running buffer is cooled to promote retention of RNA structure within the gel.
BASIC PROTOCOL 3
Materials Transcribed ribozyme RNA containing phosphorothioate-tagged analogs (see Support Protocol and Strategic Planning) 500 mM Tris/HEPES buffer, pH 7.5 (see recipe) 10 mM MgCl2 50% glycerol Native gel mix (see recipe) Gel elution buffer (see recipe) Ethanol (EtOH), cold 0.65-mL microcentrifuge tubes 70◦ C heating block Autoradiography film Platform rocker Additional reagents and equipment for native gel electrophoresis (UNIT 11.4)
Chemical and Enzymatic Probes for Nucleic Aicd Structure
6.9.9 Current Protocols in Nucleic Acid Chemistry
Supplement 17
1. In a 0.65-mL microcentrifuge tube, mix:
5 µL 1 mM RNA transcript 2 µL 500mM Tris/HEPES buffer, pH 7.5 4 µL 10 mM MgCl2 4 µL 50% glycerol 5 µL ddH2 O. 2. Incubate the RNA 5 min at 70◦ C and cool slowly to room temperature. The folding conditions presented here are a suggestion, and should be optimized for the RNA molecule of interest.
3. Perform native gel electrophoresis as described in UNIT 11.4 using a gel mix optimized for the system of interest. 4. Expose the gel to autoradiography film and excise the RNA by physically cutting the products from the gel. Elute the RNA in 700 µL gel elution buffer by rocking for 2 to 4 hr at room temperature. Identification of the folded population can be made by comparing the substituted RNA reactions to those done with wild-type RNA.
5. Ethanol precipitate the RNA by adding 3 vol cold EtOH and incubating 15 min at −80◦ C. Centrifuge 15 min at 17,000 × g, 25◦ C. Save the pellet and allow the RNA to air dry for 5 to 10 min. 6. Determine the extent of interference at all positions (see Basic Protocol 4). SUPPORT PROTOCOL
TRANSCRIPTION AND PURIFICATION OF PHOSPHOROTHIOATE-TAGGED RNA This protocol describes run-off transcription for incorporating phosphorothioate-tagged nucleotides into the desired RNA. For each RNA incorporating a modified analog (δαS), a separate RNA incorporating the phosphorothioate-tagged parent nucleotide (NαS) will be needed as a NAIM control.
Materials 10× transcription buffer (see recipe) 1 M MgCl2 (store at 4◦ C) 20× NTPs: 20 mM each ATP, CTP, GTP, and UTP (see Table 6.9.1) 10× NTPαS: modified (δαS) or parent (NαS) phosphorothioate-tagged nucleotide (see Table 6.9.1) 1 µg/µL linearized template DNA, containing a T7 promoter upstream of the RNA gene to be transcribed 0.1 mg/mL inorganic pyrophosphatase (store at −20◦ C) 500 to 1000 U/µL T7 RNA polymerase (store at −20◦ C) 2× formamide loading buffer (FLB; APPENDIX 2A) Gel elution buffer (see recipe) 3 M sodium acetate (NaOAc), pH 5.2 (APPENDIX 2A) 100% ethanol (EtOH), prechilled at −20◦ C T10 E0.1 buffer, pH 7.5 (see recipe)
Nucleotide Analog Interference Mapping
1.5-mL microcentrifuge tubes 37◦ C heating block Platform rocker Additional reagents and equipment for denaturing gel electrophoresis (APPENDIX 3B)
6.9.10 Supplement 17
Current Protocols in Nucleic Acid Chemistry
1. In a 1.5-mL microcentrifuge tube, combine:
10 µL 10× transcription buffer 2 µL 1 M MgCl2 5 µL 20× NTPs 10 µL 10× NTPαS 2 µL 1 µg/µL linearized DNA template 1 µL 0.1 mg/mL inorganic pyrophosphatase 5 µL 500 to 1000 U/µL T7 RNA polymerase 65 µL ddH2 O. Incubate 1 to 2 hr at 37◦ C. The final 1× concentrations of tagged, modified nucleotides (δTPαS) are indicated in Table 6.9.1. For NAIM control transcripts, the 1× concentration of tagged parent nucleotide (AαS, CαS, GαS, or UαS) is 0.05 mM. For both modified and control transcripts, the 1× concentration of unmodified, untagged nucleotides (from the NTP mix) should be 1 mM in most cases, but see Table 6.9.1 for some exceptions. Any of the above concentrations can be altered to better suit the template being used in transcription, particularly the Mg2+ concentration. Larger volumes can be transcribed if more RNA is needed for the particular assay being run. The expected yield per transcription is ∼0.1 to 1 nmol/mL. If higher NTP concentrations are required for RNA transcription, proportional adjustments should be made to the analog and Mg2+ concentrations.
2. Add 100 µL of 2× formamide loading buffer to RNA transcripts and proceed with preparative denaturing gel electrophoresis (e.g., APPENDIX 3B). 3. Visualize the RNA using UV shadowing on a silica plate and physically remove the gel slice containing the RNA. Elute RNA in 700 µL gel elution buffer by rocking for 1 to 2 hr at room temperature. 4. Transfer the elution buffer to a clean 1.5-mL microcentrifuge tube and add 0.1 vol of 3 M NaOAc, pH 5.2, and 3 vol of 100% EtOH. Place 15 min at −80◦ C, then centrifuge 15 min at 17,000 × g, 25◦ C. 5. Carefully remove ethanol from tube, without disturbing the RNA pellet. Allow the pellet to dry for 5 to 10 min. 6. Resuspend the RNA pellet in 50 µL T10 E0.1 buffer, pH 7.5. 7. Store RNA at −20◦ C.
IDENTIFYING POSITIONS OF INTERFERENCE AND CONTROLLING FOR THE LEVEL OF INCORPORATION
BASIC PROTOCOL 4
In this protocol, the selected RNA pools are treated with iodine to cleave at the site of the phosphorothioate tag. The cleaved RNAs are then separated by denaturing gel electrophoresis, allowing identification of the sites of analog interference. The key to analysis of NAIM data is the inclusion of all relevant samples on a single gel. For every analog tested in a NAIM experiment, the relevant gel must also contain samples in which the parent nucleotide (i.e., unmodified NαS) has been tested in a parallel experiment. The gel will also have samples for both parent and analog nucleotides that have not been iodine treated. These serve as a control for nonspecific degradation of the RNA. Positions of strong degradation in this lane are uninformative in the NAIM assay, and cannot have any interference value determined.
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An additional element in data analysis is the inclusion of RNA transcripts that have not been subjected to any selection step, but are end-labeled, iodine-treated, and analyzed on a sequencing gel. These unselected RNA molecules control for the overall level of analog incorporation and the specific level of incorporation at a particular residue in the sequence. If the incorporation level is too high, the bands at the bottom of the gel will be much stronger than those at the top of the gel, as multiple iodine-induced cleavages occur in each RNA; this should be avoided. The level of incorporation of a particular phosphorothioate nucleotide is closely correlated to the relative concentrations of the phosphorothioate and untagged nucleotide in the transcription mixture. Therefore, if over-incorporation is a problem, the concentration of the analog should be decreased or the untagged nucleotide increased in the transcription mixture. It should be noted that the cleavage of the phosphorothioate bond by iodine has an end point of ∼15% reacted. Consequently, the majority of the RNA species will appear as full length on a sequencing gel unless there are too many substitutions in the RNA. The unselected RNA molecules also control for the level of incorporation at each position. There is sequencespecific variability for the extent of incorporation that is stronger for some analogs than for others. This includes sites of hyper- and non-incorporation, but information can still be gained from such positions because of the controls. These unselected controls do not have to be included on every gel, but must be performed every time an RNA molecule is transcribed and used in a NAIM reaction. Any interferences observed with the tagged parent nucleotide can be attributed to a phosphorothioate effect. In the strongest cases, interferences at these sites are uninformative for subsequent analog studies, and an effort should be made to reduce them as much as possible. The inclusion of manganese in the reaction buffer sometimes serves to minimize phosphorothioate effects. Quantitation of individual band intensities in the I2 cleavage ladder is used to identify sites of interference resulting from analog substitution. The data are also normalized for loading differences and variation in the extent of the reaction. The following protocol calculates the extent of analog interference at each position within the RNA.
Materials 2× formamide loading buffer (FLB; APPENDIX 2A) Selected RNA samples containing phosphorothioate-tagged analogs (see Basic Protocols 1, 2, and 3) Parallel unselected RNA samples (control) 50 mM I2 /EtOH: 50 mM I2 dissolved in ethanol Phosphorimager and screen (Molecular Dynamics) ImageQuant software Additional reagents and equipment for denaturing gel electrophoresis (APPENDIX 3B) 1. Add 2 vol FLB to the selected RNA containing phosphorothioate-tagged analogs. 2. Split the selected RNA into two fractions. To one, add 1/10 vol of 50 mM I2 /EtOH. 3. Heat the reaction 2 min at 90◦ C to denature the RNA. Nucleotide Analog Interference Mapping
4. Load all the reactions onto a denaturing polyacrylamide gel (see APPENDIX 3B). Run the gel for varying lengths of time, depending on the length of the RNA molecule and the section of the RNA that is being studied. 5. Dry the gel and expose to a phosphorimager screen.
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6. Quantitate the peak intensities at each position of incorporation for both the parental nucleotide (NαS) and the nucleotide analog (δαS) observed in the ligation reaction by phosphorimager analysis using area integration. Within the ImageQuant software, draw an equivalent line through every lane on the gel and graph the radioactive intensity versus distance. Carefully define the baseline and make sure it is consistently defined between each of the lanes. Find the peaks by adjusting the noise and sensitivity variables. Compute the area under each peak and use these values in Equation 6.9.1 for each of the different variables and sites of incorporation. 7. Calculate the extent of interference at each position by substituting the individual band intensities into Equation 6.9.1.
Equation 6.9.1
where at each site of incorporation, NαS experiment is the peak intensity for the parental selected RNA, δαS experiment is the peak intensity for the analog selected RNA, NαS unselected is the peak intensity for the parental unselected RNA, δαS unselected is the peak intensity for the analog unselected RNA, and NF is a normalization factor used to account for differences between loading and extent of reaction between lanes. Values are initially calculated without the normalization factor, but then are normalized against each other to calculate κ. NF is determined by calculating the average interference value for all positions that are within two standard deviations of the mean, and dividing the value at each position by this average. NF is typically between 0.8 and 1.2. Equation 6.9.1 provides a value defined as κ that indicates the extent of interference at each site of incorporation within the sequence. A κ value >2 defines a site of analog interference. A value <0.5 defines a site of enhancement, and a value of 1 defines a residue unaffected by the substitution. The noise in the κ measurement is such that values between 0.5 and 2 are positions of little or no interference, though this is a fairly conservative interpretation.
NUCLEOTIDE ANALOG INTERFERENCE SUPRESSION (NAIS) Some of the interferences that are observed in a traditional NAIM experiment arise from the loss of a hydrogen bond and the consequent destabilization of the structure of the RNA. NAIM identifies the functional groups involved in hydrogen bonding, but NAIS can map tertiary contacts by identifying which functional groups are paired to each other in the active state of the complex (Boudvillain and Pyle, 1998; Strobel et al., 1998; Szewczak et al., 1998; Strobel and Ortoleva-Donnelly, 1999; Szewczak et al., 1999; Soukup et al., 2002).
ALTERNATE PROTOCOL 1
NAIS can be used in conjunction with any of the previous types of selection. A mutation in the RNA or an alteration in a ligand can be used in combination with NAIM to identify the tertiary interactions within that complex. The approach utilizes a single mutation or functional group substitution at one important residue in the complex and phosphorothioate-tagged analogs throughout the RNA. NAIS has been used extensively to identify hydrogen-bonded functional groups within ribozymes. Chimeric RNA molecules are constructed from a synthetic oligonucleotide containing the site-specific deletion of a functional group of interest and a transcribed RNA. These chimeric RNAs are ligated as outlined in CPMB UNIT 3.15. The RNA is transcribed with a phosphorothioate-tagged nucleotide analog that deletes the other functional group predicted to be involved in the tertiary interaction. NAIS utilizes the prediction that there will not be a further energetic penalty incurred upon the deletion of the second
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member of a hydrogen-bonded pair and, as a result, the interference will be suppressed. This assay is dependent on the ability to reconstruct the selection assay such that there is activity despite the deletion of the functional group that initially gave rise to the interference. Positions of interference that arise from the hybrid molecules are identified and compared to those from RNAs that retain the specific functional group. Any interference that is suppressed upon the addition of the site-specific substitution is a strong candidate for involvement in a hydrogen bond with the modified functional group. Additional controls must be included to show that the suppression of the interference is specific to both the nature of the substitution and the site of suppression. Ideally, this should include the following demonstrations of specificity. (1) The suppression must be specific to the site-specifically modified functional group. Mutations or site-specific substitutions at other residues should not lead to the same suppression pattern. (2) The suppression must be specific to a single residue among additional sites of interference. (3) The suppression must be specific to a single or closely related set of analogs. For materials for selection, see Basic Protocol 1; for identification of sites of interference, see Basic Protocol 4; and for ligation of RNA molecules using T4 DNA ligase, see CPMB UNIT 3.15. 1. Use NAIM to identify key functional groups in the RNA. 2. Follow the protocol in CPMB UNIT 3.15 for ligation of RNA molecules using T4 DNA ligase. 3. Use the hybrid RNA in a selection reaction as outlined previously. Because of the inherently lower activity of the hybrid RNA caused by the site-specific substitution, the selection reaction may need to be modified. Many variables, such as temperature, salt, and pH, can be altered to achieve a higher level of activity. This assay can only be performed if the substituted RNA retains some residual level of activity.
4. Identify the sites of interference as described. Compare κ values obtained using wild type ribozymes to those obtained using the hybrid RNA molecules. Sites that have high κ values in the wild-type experiment and lower values in the hybrid experiment are excellent candidates for tertiary contacts with the modified or mutated residue. ALTERNATE PROTOCOL 2
Nucleotide Analog Interference Mapping
FUNCTIONAL GROUP IONIZATION DETERMINED BY NAIM Several of the analog nucleotides used in NAIM experiments contain base modifications that, in addition to altering a functional group in the molecule, have altered pKa values from the parent nucleotide (Oyelere and Strobel, 2000; Ryder et al., 2001; Oyelere et al., 2002; Jones and Strobel, 2003). The nucleotides used in this experiment are analogs of adenosine and cytosine. These analogs make it possible to screen for functionally important base ionization at every adenosine or cytosine residue in an RNA. The sites of ionization can play a role in either the structure of the RNA or, for those RNAs that are catalytic, in the chemistry of the molecule. The adenosine analogs used to probe protonation at the N1 position include: adenosine (pKa = 3.5), purine (pKa = 2.1), 8-azaadenosine (pKa = 2.2), formycin A (pKa = 4.4), and 7-deazaadeonsine (pKa = 5.2). The cytosine analogs used to identify protonation events at the N3 position include: cytidine (pKa = 4.2), 6-azacytidine (pKa = 2.6), 5-fluorocytidine (pKa = 2.3), and pseudoisocytidine (pKa = 9.4). These analogs are used in conjunction with each other to probe for the ionization of adenosine and cytosine nucleotides. On the basis of previous studies with these analogs, the following interference pattern is expected at individual adenosine residues whose ionization is functionally important.
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At elevated pH, interference is seen with PurαS and n8 AαS. The interference is at least partially rescued by lowering the pH of the buffer. Furthermore, enhanced activity may be seen with FormAαS or 7dAαS at sites of base ionization due to the increased basicity of these nucleosides, if base ionization is a limiting event in the reaction. At those positions where the N3 ionization of cytidine within the RNA is important for activity, both n6 CαS and f5 CαS have been shown to cause interference at high pH due to the increased acidity of both analogs (Oyelere et al., 2002). But as is the case for the adenosine analogs, the interference can be rescued as the pH is reduced. Conversely, iCαS will show enhanced activity at elevated pHs, but the effect will be diminished as the pH is reduced.
METAL BINDING FUNCTIONAL GROUPS IDENTIFIED BY NAIM The use of sulfur-substituted analogs enables NAIM to be used in metal ion rescue experiments (Basu et al., 1998; Basu and Strobel, 2001). Substitution with sulfur disturbs metal ion binding sites, as sulfur is unable to form stable complexes with magnesium or potassium (Heitner et al., 1972; Douglas et al., 1990). Interferences seen with parental nucleotides may arise from disruption of metal ion binding to the pro-RP oxygen of the RNA backbone. Identification of guanosine-O6 -carbonyl ligands for metal ions is done with the analog 6-thioguanosine phosphorothioate (s6 GαS). Inclusion of thiophilic metals such as Mn2+ , Cd2+ , Co2+ , Ni2+ , Zn2+ , and Tl+ in the selection reaction ameliorates the interference caused by the change from oxygen to sulfur.
ALTERNATE PROTOCOL 3
Materials RNA molecules transcribed in the presence of sulfur-substituted nucleotides 100 mM Mn(OAc)2 100 mM TlOAc Other mono- and divalent metals as required 2% 2-mercaptoethanol Additional reagents and equipment for selection (see Basic Protocol 1) and for identification of sites of interference (see Basic Protocol 4) 1. Perform the selection activity at various concentrations of thiophilic metal ion concentration, from 0.1 to 5 mM or higher, if needed. Any buffers or salts with chloride as a counter ion cannot be used for thallium rescue experiments, as thallium chloride is insoluble. CAUTION: Care should be taken when handling Tl+ as it is toxic.
2. Control for specificity of the metal ion rescue by using other mono- and divalent metal ions in the selection assay. To show that rescue is specific to the metal ion forming a stable complex with sulfur, and not merely overall structure stabilization by metal ions, other non-thiophilic metals such as K+ , NH4 + , Li+ , Na+ , Ca2+ , and Sr2+ at high concentrations (up to 100 mM) should be included in the reaction buffers.
3. Prepare the samples for loading on a denaturing polyacrylamide gel. For RNA samples containing s6 GαS, add 1/10 vol of 2% 2-mercaptoethanol directly before loading the samples on the gel in order to reduce streakiness in the gel. 4. Identify the sites of interference. Compare the κ values obtained for sites of interference at each metal ion concentration. Specific rescue of an interference by a single thiophilic metal suggests this functional group is a ligand for metal ion binding.
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ALTERNATE PROTOCOL 4
QUANTITATIVE NUCLEOTIDE ANALOG INTERFERENCE MAPPING (QNAIM) QNAIM is a series of NAIM experiments performed at several ligand concentrations (Cochrane et al., 2003), which allows characterization of the energetics of binding. The RNA concentration is held substantially below the dissociation constant of the complex (Kd ), and the ligand concentration is varied from one to several times the Kd concentration. Under these conditions, RNA-ligand binding is driven by the increase in the ligand concentration, which is already in vast excess. As a result, the RNAs do not compete with each other for binding; instead, the distribution of RNAs in the bound fraction results entirely from the intrinsic affinity of each RNA for the ligand. The population distribution in the bound fraction is revealed by the individual band intensities observed in the sequencing ladder that results from iodine treatment. By plotting the interference magnitude as a function of ligand concentration, it is possible to calculate the binding constant and free energy contribution ( G) of each functional group substitution at each position in the RNA. Because of slight differences in salt and ligand concentrations, the binding constant can vary between experiments and should be determined separately for each experiment. The binding constant for the bulk RNA population is determined using a dot blot method.
Materials 1× Kd RNA (labeled) 500 mM Tris/HEPES buffer, pH 7.5 (see recipe) 2 M KCl 1 M MgCl2 0.1 M DTT 0.1% Igepal C-680 1 mg/mL tRNA 200× Kd stock of ligand Ligand dilution buffer (see recipe) Wash buffer (see recipe) Whatman paper Cling film Phosphorimager and screen Additional reagents and equipment for dot blots (see CPMB UNIT 2.9B), for selection of active molecules based on ligand binding (see Basic Protocol 1), and for determining sites of interference (see Basic Protocol 4) Determine binding constant 1. Heat 120 µL of 1× Kd RNA to 90◦ C for 1 min; cool on ice for at least 3 min. 2. To the RNA, add:
Nucleotide Analog Interference Mapping
48 µL 500 mM Tris/HEPES, pH 7.5 120 µL 2 M KCl 12 µL 1 M MgCl2 15 µL 0.1 M DTT 120 µL 0.1% Igepal C-680 120 µL 1 mg/mL tRNA 645 µL ddH2 O.
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3. Make twelve 20-µL 1:2 serial dilutions from the 200× Kd stock of ligand into ligand dilution buffer. 4. Add 80 µL of the RNA mix from step 2 into each of the twelve serial dilutions of ligand. 5. Pipet 40 µL from each ligand concentration into a separate dot blot hole and wash with 100 µL wash buffer. 6. Dismantle the dot blot apparatus and allow the filters to air dry for 1 to 2 min. Place the filters on Whatman paper, cover with cling film, and expose to a phosphorimager screen. 7. Quantitate the data by comparing the density of counts in the bound fraction to the unbound fraction at every ligand concentration. Determine the binding constant (Kd ) by fitting the data to Equation 6.9.2:
fraction bound =
(M × [protein]) +L (K d bulk + [protein])
Equation 6.9.2
where Kd is the apparent equilibrium binding constant, M is the maximum fraction of RNA bound at the highest protein concentration (this varies from 0.7 to 0.9), and L is the minimum fraction of RNA bound at the lowest protein concentration (this is usually very close to 0).
Perform QNAIM 8. For each ligand concentration, follow the protocol outlined in Basic Protocol 1. 9. Determine the sites of interference by denaturing polyacrylamide gel electrophoresis as outlined in Basic Protocol 4. 10. Once the sites of interference have been found, determine 1/κ values for each site at every protein concentration. 11. Use these interference values to determine the relative binding constant for each RNA that has a substitution at a particular position:
(M × [protein]) 1 = +L κ (K interference + [protein]) d Equation 6.9.3
where Kd is the apparent equilibrium binding constant, M is the maximum level of rescue of the particular interference (or the upper baseline), and L is the maximum extent of interference for the same position (or the lower baseline). 12. Determine the change in binding free energy ( G) from Equation 6.9.4 using the measured bulk binding constant and the binding constant determined for the particular interference band:
∆∆G = RT × ln ( K d bulk/K d interference) Equation 6.9.4
where R is the gas constant and T is temperature.
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Filter elution buffer 300 mM sodium acetate (NaOAc), pH 5.2 (APPENDIX 2A) 1% SDS (APPENDIX 2A) T10 E0.1 buffer (see recipe) Store up to 3 months at 25◦ C Gel elution buffer 2% SDS (APPENDIX 2A) T10 E0.1 buffer (see recipe) Store up to 3 months at 25◦ C Ligand dilution buffer 20 mM Tris/HEPES buffer, pH 7.5 (see recipe) 200 mM KCl 10 mM MgCl2 0.1% Igepal C-680 Store up to 3 months at 4◦ C Native gel mix 6% acrylamide 1× TBE electrophoresis buffer (APPENDIX 2A) 3 mM MgCl2 Store up to 3 months at 25◦ C T10 E0.1 buffer, pH 7.5 10 mM Tris·Cl, pH 7.5 (APPENDIX 2A) 0.1 mM EDTA (APPENDIX 2A) Transcription buffer, 10× 40 mM Tris·Cl, pH 7.5 (APPENDIX 2A) 1 mM spermidine 5 mM dithiothreitol 0.01% Triton X-100 Store up to 3 months at −20◦ C Tris/HEPES buffer, 500 mM, pH 7.5 500 mM Tris 500 mM HEPES Adjust pH to 7.5 Store up to 3 months at 25◦ C
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Wash buffer 20 mM Tris/HEPES buffer, pH 7.5 (see recipe) 200 mM KCl 10 mM MgCl2 Store up to 3 months at 4◦ C
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COMMENTARY Background Information NAIM is a generalizable chemogenetic approach that identifies functional groups that are important for the activity of an RNA (Strobel and Shetty, 1997). This method uses a series of 5 -O-(1-thio) nucleoside analogs to probe the contribution specific functional groups make to RNA activity. NAIM utilizes a pool of randomly substituted RNAs generated by in vitro transcription, and the active RNAs are identified through a selection experiment. In this way, a particular RNA can be screened with multiple analogs in a time-efficient manner. NAIM is applicable to any RNA with a selectable function. It can be used to study RNA catalysis through cleavage or ligation (Strobel and Shetty, 1997; Boudvillain and Pyle, 1998; Kazantsev and Pace, 1998; Sood et al., 1998; Jones et al., 2001; Kaye et al., 2002), RNA interactions with protein and other ligands (Batey et al., 2001), as well as the steps of RNA folding (Basu and Strobel, 1999). It has even been used to look at RNP assembly in vivo (Szewczak et al., 2002). This approach should make it possible to identify the chemical groups important for a wide variety of RNA activities.
Critical Parameters NAIM requires an assay that distinguishes active members of a substituted RNA population from less active members. Two types of methods have been used to achieve this separation. The first is to physically separate active from inactive RNAs in the course of the assay. The second approach, specific to ribozymes, is to make use of inherent RNA activity to selectively radiolabel active members in the population. In general, the second type of assay is easier to perform, as one preparation of labeled substrate will allow interference mapping with many different analog-containing RNA transcripts. Experiments that require multiple gels are undesirable because they increase the time necessary for analysis. Additionally, longer experimental protocols could lead to increased levels of RNA degradation, which will interfere with quantitation of band intensity. It is important to design the RNA being studied by NAIM so that it is of sufficient length to contain several sites of analog substitution that do not show interference. These provide important internal controls in the interference experiments to account for differences in loading volume and extent of reaction between experiments.
These positions are used to calculate the normalization factor (NF) in Equation 6.9.1. NF becomes more accurate as the population of sites that do not show interference increases. These issues constrain the length of the RNA to no less than 40 to 50 nucleotides. Once an appropriate assay has been designed, it is necessary to optimize the reaction in order to obtain the desired level of selectivity. This can be done by varying several aspects of the experimental conditions. For example, it has been observed that high concentrations of divalent metal ions and lower reaction temperatures tend to mask sites of weaker interference in the group I intron. This can be attributed to overall improvement in the stability of the RNA fold. In all cases, a spectrum of reaction conditions should be explored with a subset of analogs prior to completion of a full screen. Furthermore, sites that show phosphorothioate interference are uninformative in a NAIM assay, so some effort should be made to find conditions that minimize these effects, such as the inclusion of Mn2+ in the reaction buffer.
Anticipated Results NAIM is a versatile method, used to quickly identify important functional groups within a structured RNA molecule. In conjunction with the other techniques outlined in this unit, a NAIM experiment can provide detailed structural and energetic information on any in vitro transcribed RNA molecule. A typical NAIM experiment will yield information about the contribution to catalysis and/or stability of an individual functional group at every possible position within an RNA molecule. The magnitude of the interference (κ) will provide additional information about the relative importance of that modification when compared to other sites within the same molecule. By using the full range of nucleotide analogs available, the key functional groups in the RNA molecule can be specifically elucidated. Furthermore, information about hydrogen bonding partners (NAIS), metal ion ligands, pKa -perturbed nucleotides, and the energetics of ligand binding (QNAIM) can be determined. NAIM data analysis requires the comparison of the experimental data to several reference data sets. By carefully controlling for the incorporation level at a particular site and the effect of the phosphorothioate tag, a wealth of detailed data can be obtained from a single NAIM experiment.
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Time Considerations The NAIM experiment itself can be completed in a single day, once the necessary materials are assembled. Due to the inherent instability of RNA molecules, it is preferable to complete the experiment as quickly as possible. However, RNA molecules in which phosphorothioate-tagged nucleotides have been incorporated are no more prone to degradation than normal RNA molecules, and may be slightly less so.
Acknowledgments We thank S.P. Ryder for critical comments on this manuscript. We also thank A.B. Kosek, A.A. Szewczak, A.K. Oyelere, and S. Basu for helpful discussion. This work was supported by NSF grant CHE-0100057 to S.A.S.
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Ryder, S.P., Oyelere, A.K., Padilla, J.L., Klostermeier, D., Millar, D.P., and Strobel, S.A. 2001. Investigation of adenosine base ionization in the hairpin ribozyme by nucleotide analog interference mapping. RNA 7:1454-1463.
Szewczak, A.A., Ortoleva-Donnelly, L., Zivarts, M.V., Oyelere, A.K., Kazantsev, A.V., and Strobel, S.A. 1999. An important base triple anchors the substrate helix recognition surface within the Tetrahymena ribozyme active site. Proc. Natl. Acad. Sci. U.S.A. 96:1118311188.
Schwans, J.P., Cortez, C.N., Olvera, J.M., and Piccirilli, J.A. 2003. 2 -Mercaptonucleotide interference reveals regions of close packing within folded RNA molecules. J. Am. Chem. Soc. 125:10012-10018.
Szewczak, L.B., DeGregorio, S.J., Strobel, S.A., and Steitz, J.A. 2002. Exclusive interaction of the 15.5 kD protein with the terminal box C/D motif of a methylation guide snoRNP. Chem. Biol. 9:1095-1107.
Sood, V.D., Beattie, T.L., and Collins, R.A. 1998. Identification of phosphate groups involved in metal binding and tertiary interactions in the core of the Neurospora VS ribozyme. J. Mol. Biol. 282:741-750.
Wong, I. and Lohman, T.M. 1993. A double-filter method for nitrocellulose-filter binding: Application to protein-nucleic acid interactions. Proc. Natl. Acad. Sci. U.S.A. 90:5428-5432.
Soukup, J.K., Minakawa, N., Matsuda, A., and Strobel, S.A. 2002. Identification of A-minor tertiary interactions within a bacterial group I intron active site by 3-deazaadenosine interference mapping. Biochemistry 41:10426-10438.
Contributed by Jesse C. Cochrane and Scott A. Strobel Yale University New Haven, Connecticut
Sousa, R. and Padilla, R. 1995. A mutant T7 RNA polymerase as a DNA polymerase. EMBO J. 14:4609-4621.
Chemical and Enzymatic Probes for Nucleic Aicd Structure
6.9.21 Current Protocols in Nucleic Acid Chemistry
Supplement 17
CHAPTER 7 Biophysical Analysis of Nucleic Acids INTRODUCTION he opening commentary (UNIT 7.1) by Tinoco defines the limitations of the current state of the art in the biophysical analysis of nucleic acids and sets the stage for the units that follow. In UNIT 7.2, James explains the steps to be followed in determining an NMR structure. He also discusses the care that is necessary to be sure that the structure determined is in fact the correct structure. The unit on optical methods (UNIT 7.3) by Plum is likely to be one of the most heavily used units in this volume and will be of particular importance to the novice. The section on determination of oligonucleotide extinction coefficients, in particular, is essential to anyone needing to know solution concentrations, and the section on analysis of equilibrium melting curves contains all of the relevant equations and a discussion of which are most appropriate to a given situation.
T
Calorimetric methods (UNIT 7.4) are becoming much more commonly used, and, while quite powerful, there are numerous sources of error. Pilch gives a thorough explanation of the care that is necessary to obtain meaningful results, as well as the relevant methods for analysis of the data produced. In UNIT 7.5, Cheatham, Brooks, and Kollman give an introduction to computer simulation of nucleic acids. This topic is continued in UNITS 7.8-7.10, which provide more in-depth coverage of energy and sampling, electrostatics and solvation, and setup and analysis. In UNIT 7.6, Doudna and Ferr´e-D’Amar´e include crystallization conditions for twenty RNA and RNA-protein complexes. The overall goal of the unit, however, is to provide general guidance, rather than to attempt to give a detailed protocol or set of protocols. The RNA theme continues in UNIT 7.7, which deals with NMR determination of RNA structure. Williamson focuses on the assignment of resonances and the calculation of structures. For preparation of the isotopically labeled RNA, references are given for published procedures. discusses analysis of DNA structure by circular dichroism (CD). For nucleic acids, CD is typically used to monitor electronic transitions in nucleobases. It is extremely sensitive to changes in environmental conditions such as pH, temperature, and ionic strength. This unit provides both procedural guidelines and representative spectra to illustrate the usefulness of the method.
UNIT 7.11
The antigene approach requires that a single-stranded oligonucleotide recognize doublestranded DNA to form a triplex structure. The two most common methods for evaluating, in vitro, the potency of modified oligonucleotides to associate with dsDNA are gel-shift electrophoresis and triplex melting. These methods, although conceptually simple, can be tricky to perform. They are described in detail in UNIT 7.12. Roger Jones and Piet Herdewijn
Biophysical Analysis of Nucleic Acids Contributed by Roger Jones and Piet Herdewijn Current Protocols in Nucleic Acid Chemistry (2007) 7.0.1 C 2007 by John Wiley & Sons, Inc. Copyright
7.0.1 Supplement 54
Biophysical Analysis of Nucleic Acids BIOPHYSICAL METHODS The determination of the structure of a nucleic acid, or of any molecule, is a necessary step in beginning to understand what the molecule can do. Before one can ask how reactive it is, what ligands it can bind, or what reactions it can catalyze, one must know the structures of its reactants, its products, and its intermediates. Structures are required before functions can be understood. This unit describes the main biophysical methods that have been used to study nucleic acids, and briefly mentions some of the powerful new methods that are just appearing. The first step in determining the structure of a nucleic acid is to establish its sequence—its primary structure. If the molecule was made by chemical or enzymatic synthesis, this may be a sufficient proof of sequence. For a natural DNA sample, the standard method is that of Sanger (e.g., see CPMB UNIT 7.4A), in which the DNA is replicated by DNA polymerase in the presence of one dideoxynucleoside triphosphate such as ddGTP. The ddGTP acts as a chain terminator, so that chain length determination by gel electrophoresis provides the sequence of G’s. The reaction is repeated with each ddNTP base to determine the complete sequence. For RNA the same method is used, except with reverse transcriptase as the DNA-synthesizing enzyme. Once a pure, single species of a nucleic acid with a known sequence has been obtained, its secondary structure (base pairing) is assessed. Measurement of its ultraviolet absorbance at 260 nm provides its approximate concentration. To further characterize the structure and to obtain a more accurate concentration, an absorbance melting curve is measured. Doublestranded nucleic acids have a lower absorbance per nucleotide than single-stranded nucleic acids. Stacked bases are said to be hypochromic (less absorbing) compared to unstacked bases. Thus, as a nucleic acid solution is heated, the absorbance increases as double strands melt and the bases in single strands unstack. A plot of absorbance versus temperature, or of the derivative of absorbance versus temperature, is called a melting curve. The shape of the curve—the number, widths, and positions of the maxima in the derivative curve—can reveal a great deal about the different conformations that are present in a nucleic acid, as well as the transitions that occur. Measurement of circular dichroism versus wavelength provides a more
detailed picture of the conformations present in the sample. The various double-stranded conformations of nucleic acids (A, B, and Z forms; see APPENDIX 1B) have characteristic signatures in circular dichroism. Absorbance and circular dichroism are thus used to measure concentrations and to qualitatively characterize the secondary structures of nucleic acids. They are also used to monitor covalent reactions or equilibrium binding with ligands. Reactions can also be followed by changes in the fluorescence of rare fluorescing bases in tRNA or, more commonly, of fluorescent ligands or reagents. All these spectroscopic measurements require on the order of 1 mL of a 0.1 mM nucleotide solution. The small amount of material needed and the ease of measurement make ultraviolet absorption and circular dichroism the methods of choice for qualitative characterization of nucleic acids. Atomic resolution structures—X, Y, Z coordinates for each atom—require either nuclear magnetic resonance (NMR) measurements in solution or X-ray diffraction measurements on single crystals. Both types of measurements take months of effort and at least an order of magnitude more material. X-ray diffraction depends on the ability to obtain single crystals that diffract well, which seems to depend largely on trial and error. Once a suitable crystal is found and the diffraction data analyzed, a complete structure of the nucleic acid is obtained. The structure includes coordinates of the molecule plus coordinates of tightly bound water molecules and ions. Very large molecules can be determined; for example, progress is being made on the structure of a ribosome. NMR studies in solution can provide useful information short of a three-dimensional structure. For example, the detailed base pairing in an RNA can be established from a one-dimensional imino spectrum using no more material than needed for a UV absorbance spectrum. However, for atomic coordinates, two-dimensional spectra and ten times the concentration are needed. Interpretation of NMR experiments is limited by resolution of the spectra. Twodimensional spectra and isotope labeling improve the resolution, but there is a limit to the size of nucleic acids that can be analyzed. The size is increasing as stronger magnets and newer techniques become available, but a
Contributed by Ignacio Tinoco, Jr. Current Protocols in Nucleic Acid Chemistry (2000) 7.1.1-7.1.8 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 7.1
Biophysical Analysis of Nucleic Acids
7.1.1
length of more than 100 nucleotides is very difficult at this time.
X-Ray Diffraction
Biophysical Analysis of Nucleic Acids
The field of structural biology can be considered to have started in 1953 when Watson and Crick (Watson and Crick, 1953) interpreted the fiber X-ray diffraction data of Wilkins (Wilkins et al., 1953) and Franklin (Franklin and Gosling, 1953) on DNA. The data were consistent with a double-helical structure with complementary paired bases on the inside of the helix and phosphates on the outside. The resolution was not high, but the structural information was sufficient to revolutionize the study of biology. X-ray diffraction of crystals now gives the most accurate information about the atomic structures of nucleic acids. Positions of atoms can be determined to 0.001 Å for small molecules and to 0.1 Å for macromolecules. Crystals of bases, nucleosides, nucleotides, dinucleotides, and oligonucleotides revealed the preferred torsion angles around the seven bonds that characterize the conformation of a nucleotide (see APPENDIX 1B & 1C). The nomenclature used to describe a nucleic acid is based on the structures determined by X-ray diffraction of crystals. B-form DNA has 2′-endo sugar puckers (pseudorotation phase angle = 162°), but a wide variation of pseudorotation angles is found. A-form RNA is more conformationally rigid, with 3′-endo sugar puckers (pseudorotation phase angle = 18°). The naturally occurring nucleotides have an anti conformation around the glycosidic bond in righthanded A-form and B-form nucleic acids, but left-handed Z-form nucleic acids have syn purines. Much of this work is summarized in Saenger (1984). Recently, the emphasis of X-ray diffraction studies on DNA-containing crystals has included unusual structures, such as G quartets (Kang et al., 1992), DNA-drug complexes (Chen et al., 1997), DNA-protein complexes (Passner and Steitz, 1997), and oligonucleotides with chemically damaged bases pertinent to DNA mutation and repair (Lipscomb et al., 1995). High-resolution X-ray structures of RNA molecules began with transfer RNAs in 1973 (Saenger, 1984). Several tRNA synthetasetRNA complexes, including tRNASer and its synthetase (Biou et al., 1994), have since been solved. Ribozymes are novel RNA enzymes whose mechanisms are being actively pursued. Structures of hairpin ribozymes (Pley et al., 1994; Scott et al., 1995, 1996) have provided
the conformation of the RNA and the location of crucial metal ions. The structure determined is that of the ground state, not the transition state, but the structure does allow plausible proposals about which torsion angles must change in order to form the transition state. The structure of the first RNA molecule large enough to have an inside and an outside has been published (160 nucleotides; Cate et al., 1996a). Here, inside indicates areas of the folded RNA that are not accessible to the solvent, as found in most proteins. The structure revealed several new types of RNA structural motifs (Tinoco and Kieft, 1997), including adenosine platforms and ribose zippers (Cate et al., 1996b). The importance of metal ion– binding sites in the folding of RNA molecules has become increasingly clear (Cate and Doudna, 1996; Cate et al., 1997; Correll et al., 1997). X-ray diffraction of crystals is the most powerful method to obtain accurate coordinates for molecular structures (Glusker et al., 1994). The conformation of the molecule is obtained, as are the locations of bound ions and bound water molecules, provided the resolution is high enough. Whether a nucleic acid molecule or complex will crystallize and whether the crystal will diffract to high resolution seems to be a matter of luck. One must try a wide range of crystallizing conditions, nucleic acid sequences, and chain lengths to obtain useful crystals. Once the crystal structure of a nucleic acid is known, one must ask how it is related to solution conformations or physiological conformations. DNA tends to crystallize as A-form double helices, since the low water activity used for crystallization solvents favors the A form. However, the physiological structure of DNA is close to the B form. The difference between crystal and solution structures for DNA is exemplified by a crystal that showed the X-ray diffraction of both A- and B-form DNA (Doucet et al., 1989). The crystalline oligodeoxynucleotide had an A-form structure, but the soluble oligodeoxynucleotide in the solvent-filled interstices of the crystal had the fiber diffraction pattern of oriented B-form DNA. RNA tetraloops are very stable, very common hairpin loops that have been characterized by NMR in solution. When a UUCG tetraloop and a GAAA tetraloop were each crystallized, each formed a double strand with an internal loop of eight nucleotides (Holbrook et al., 1991; Baeyens et al., 1996). In crystals, the double
7.1.2 Current Protocols in Nucleic Acid Chemistry
helix with an internal loop apparently packed better than an intramolecular hairpin loop. Crystal structures are static. There is local variation around the equilibrium positions of the atoms, which is quantitated by the temperature factor for each atom in the crystal. Some regions of the molecule may be so disordered that they cannot be determined in the crystal structure. Although the extent of variation and disorder can be interpreted in terms of dynamics of the molecule, the molecules are essentially frozen in the solid. However, the time dependence of a reaction, such as that catalyzed by an enzyme, can be measured in a crystal using the Laue X-ray diffraction method (Farber, 1997), which uses X rays with a wide range of wavelengths rather than the usual monochromatic X rays. For a chosen orientation of the crystal, the range of wavelengths produces a series of diffraction spots for the different crystal lattice spacing. The Laue diffraction method has the advantage that the full intensity of a synchrotron source can be incident on the crystal; no monochromator is needed. The synchrotron source provides short pulses of high-intensity radiation that can be used to take motion pictures of the crystal. One possible scenario is to start the reaction in the crystal with a short light pulse, then follow the changes in conformation with the synchrotron X-ray pulses.
Nuclear Magnetic Resonance Nucleic acid structures at atomic resolution and the dynamics of these structures can be measured in solution by NMR (UNIT 7.2; Wüthrich, 1986; Roberts, 1993; James, 1995; Wemmer, 2000). Two- and three-dimensional NMR experiments provide distances between protons (nuclear Overhauser effect spectroscopy, or NOESY) and torsion angles between protons separated by three bonds (correlated spectroscopy, or COSY). Distance measurements depend on the nuclear Overhauser effect (NOE), which is proportional to the inverse sixth power of the distance between the protons. The rapid decrease of the effect with distance means that protons separated by >5 Å do not have measurable NOEs. Clearly, NMR provides local structure (protons within 5 Å and three bonds). However, the local structure can also establish the global structure for compact molecules, such as many folded RNA molecules. It is less accurate for extended DNA molecules with slight bends.
NMR spectra in water provide an accurate secondary structure for a nucleic acid. The imino protons of guanine, thymine, and uracil resonate in a characteristic region between 9 and 15 ppm, but are only seen if they are exchanging slowly with water. Base-paired or otherwise protected imino protons exchange slowly; others exchange rapidly and are not measurable. The imino spectrum of a nucleic acid thus has only a few peaks and is straightforward to assign from the NOEs between adjacent base pairs. The assigned imino spectrum provides the sequence of the basepaired regions, which defines the secondary structure. To obtain a complete three-dimensional structure, nonexchangeable protons must be assigned. This is usually straightforward for molecules of <40 nucleotides (Varani and Tinoco, 1991; Allain and Varani, 1997). Proton spectra and natural abundance 13C spectra suffice to assign all base protons and the 1′, 2′, 2′′, and some 3′ protons. NOESY and COSY spectra of these protons can lead to atomic resolution structures. For molecules containing >20 nucleotides, isotope labeling is often required to allow the assignments necessary for structure determination. Uniform labeling with 13C (Batey et al., 1992; Nikonowicz et al., 1992) and specific 13C labeling (SantaLucia et al., 1995) allow three-dimensional and 13C-edited NMR experiments that can resolve overlapping two-dimensional proton spectra. Deuteration can simplify spectra by removing protons (Tolbert and Williamson, 1996); this can be done nonuniformly (Foldesi et al., 1996) to provide NMR windows for analyzing small regions of larger molecules. Current NMR studies have focused on unusual (non-B-form) DNA structures and on all types of RNA structures. Base-base mismatches in DNA double strands (Chou et al., 1997), DNA triple strands (Wang et al., 1992; Radhakrishnan and Patel, 1994), G quartets (Williamson, 1994), and DNA-antibiotic complexes (Wemmer and Dervan, 1997) are some of the systems studied. Base-base mismatches in RNA (Wu et al., 1995; Wu and Turner, 1996), RNA G quartets (Cheong and Moore, 1992), RNA-antibiotic complexes (Fourmy et al., 1996; Recht et al., 1996), RNA-peptide complexes (Battiste et al., 1994; Puglisi et al., 1995), RNA-protein complexes (Ramos et al., 1997; Varani, 1997), and RNA–metal ion complexes (Kieft and Tinoco, 1997) have all been studied. RNA molecules that have been selected from a random pool of sequences to
Biophysical Analysis of Nucleic Acids
7.1.3 Current Protocols in Nucleic Acid Chemistry
specifically bind ligands have very compact, rigid structures (Fan et al., 1996; Jiang et al., 1996; Dieckmann et al., 1997; Zimmerman et al., 1997). RNA-RNA interactions as seen in pseudoknots (Shen and Tinoco, 1995), kissing hairpins (Chang and Tinoco, 1997), and tetraloop receptors (Butcher et al., 1997) characterize the motifs that fold RNA into functional molecules. The dynamics of a nucleic acid molecule and of any part of the molecule can be assessed directly by NMR measurements. The width of each nuclear resonance peak (quantitated by T2, the spin-spin relaxation time) depends on the motion of the nucleus. The motion is a combination of the rotation of the molecule as a whole plus any motion of the part of the molecule containing the nucleus. Measurement of T2 plus T1 (the spin-lattice relaxation time) plus NOEs for each nucleus can provide a detailed picture of the nanosecond dynamics of the molecule (Lipari and Szabo, 1982). When there is exchange on a millisecond time scale between two different conformations of a molecule, the NMR spectrum can be used to quantitate the rate constants. Fast exchange produces a spectrum that is the average of the spectra in the two conformations, weighted by the amount of each conformation. Slow exchange produces a spectrum that is the weighted sum of the spectra of the two conformations. Intermediate exchange produces very broad resonances that are difficult to measure and interpret. In this context, fast means that the sum of the forward and reverse rate constants is large compared to the difference in resonance frequencies of a nucleus in the two conformations; slow means the opposite. For an NMR experiment at 600 MHz, a 0.1 ppm difference in resonance frequencies means a frequency difference of 60 sec–1. It is clear that, for rate constants in this range, some of the resonances can be in slow exchange and others in fast exchange. Thus, it is possible to learn which parts of a molecule are dynamic and undergoing changes in conformation. Reactions that occur in the range of minutes to hours can be monitored by a series of onedimensional NMR spectra as a function of time. Thus, NMR measurements can be used to study dynamics of molecules on time scales from nanoseconds to days.
Optical Spectroscopy Biophysical Analysis of Nucleic Acids
Absorption, circular dichroism, and fluorescence can be used to obtain structural information about molecules, but not at atomic resolu-
tion. Absorbance melting curves have long been used to study double-helix formation in nucleic acids (Gray et al., 1995; UNIT 7.3). Circular dichroism can characterize conformational changes on forming tertiary structure in RNA (Johnson and Gray, 1992). Fluorescence has been used to identify hybridization to arrays of sequences on solid supports (Pease et al., 1994). Fluorescence resonance energy transfer (FRET) has been used to measure distances between fluorophores in DNA (Murchie et al., 1989; Clegg et al., 1993) and RNA (Tuschl et al., 1994). Absorption and circular dichroism are very convenient for monitoring changes in conformation after a change, for example, in solvent, pH, salt concentration, or temperature. The kinetics of changes in structure and conformation of nucleic acids have been extensively studied by optical spectroscopy. Temperature-jump and stopped-flow kinetics (LeCuyer and Crothers, 1994; Maglott and Glick, 1997) provide data for reactions in the millisecond range.
THEORETICAL AND COMPUTATIONAL METHODS Theoretical analysis is indispensable in understanding the experimental methods used to obtain structure and dynamics. However, theoretical methods that can deduce structure directly from sequence are greatly hoped for and are being actively developed. Useful reviews of the computational methods in nucleic acid structure modeling can be found in Louise-May et al. (1996) and in UNIT 7.5. Many of the computational programs for calculating macromolecular structures are available on the internet. Secondary structure prediction programs for RNA (http://www. ibc.wustl.edu/∼zuker/rna/form1.cgi) a n d D N A (http://sun2.science.wayne.edu/ ∼jslsun2/servers/dna/form1.cgi) use experimentally determined thermodynamic parameters to find base-paired arrangements with the lowest and near-lowest free energies. M C - SYM (http://www.iro.umontreal.ca/ ∼major/mcsym.html) is a structural modeling program that uses published nucleic acid structures as a database to calculate three-dimensional structures from the sequence and experimental constraints. Standard double-helix geometry is used for the base-paired regions. A wide range of possible loops and bulges are modeled for the remaining regions based on published structures. Experimental data such as chemical reactivity of the nucleotides, cross-linking results, and incomplete NMR
7.1.4 Current Protocols in Nucleic Acid Chemistry
constraints are used to help the investigator choose between possible structures. A M B E R (http://www.amber.ucsf.edu/ amber/amber.html) and CHARMM (http:// yuri.harvard.edu/charmm/charmm.html) are programs that allow molecular dynamics simulations to be performed on nucleic acids and proteins. Coordinates and velocities are chosen at time zero for each atom in the macromolecule and solvent. After a time interval of ∼1 fsec, potential energy functions are used to calculate the force, and thus the acceleration, on each atom in its new position. After each time step, new positions and new forces are calculated. The molecular motion of the macromolecule in solution can be simulated over a time scale of picoseconds to one nanosecond. The potential functions used to calculate the forces are clearly the key to a useful simulation. At present the potentials cannot fold an RNA into a correct structure without further information. However, if potentials derived from NMR measurements on the molecule (proton-proton distances and torsion angles) are added, an NMRderived structure is obtained. The quality of the structure will depend on the number and precision of the NMR restraints. A goal to develop “Multiscale Modeling Tools for Structural Biology” is being supported by the NIH. The NIH Research Resource has a Web page (http://mmtsb.scripps.edu/) that has links to AMBER and CHARMM and also to Yammp, a molecular mechanics program for modeling structures such as ribosomes, viruses, and supercoiled DNA. Yammp uses a reduced representation, in which a sugar, base, or basepair is represented by a single “atom.” Helices and proteins can be modeled by cylinders or spheres. The ultimate goal is to be able to model any molecular biological structure at any scale of representation needed to answer the questions of interest.
SINGLE-MOLECULE METHODS Methods that measure the properties of single molecules can provide unique information. Measurements made in solution or in crystals are clearly the average properties of many molecules. With single-molecule detection, each different molecule can be studied in its own microenvironment. A review of the results from the early days of single-molecule research (1980s) has been published (Bustamante, 1991). In principle, any type of spectroscopy can be applied to a single molecule, but the sensitivity required to do NMR spectroscopy has yet to be attained. A single electron magnetic mo-
ment can be detected, but detection of a single proton magnetic moment requires about a 1000-fold increase in sensitivity (Rugar et al., 1994). Fluorescence and absorption spectroscopy have been measured on single molecules. Fluorescence energy transfer between two fluorophores depends on the distance (1/r6) and angle between the fluorophores. It is common to assume random angular orientation between the fluorophores, which leads to an uncertainty in the distance measurement. If only two fluorophores are observed, either within one molecule or in two molecules, the absolute orientation of each fluorophore can be measured from the polarization of the fluorescence. This measurement determines the angular orientation and produces a more accurate distance (Ha et al., 1996). Spectroscopy of individual molecules has revealed the existence of longlived “dark” excited states (excited states that do not emit light) in addition to the previously known excited singlet and triplet states (Dickson et al., 1996, 1997). Mechanical properties (such as elasticity) of single DNA molecules have been analyzed by measuring the force (piconewtons) necessary to stretch the molecule from a random coil to a stretched rod (Smith et al., 1992). Laser tweezers can be used to manipulate macromolecules in various ways; some recent applications have been illustrated on the muscle protein titin (Kellermayer et al., 1997; Rief et al., 1997). Scanning-probe microscopy includes atomic force microscopy (AFM) and scanning tunneling microscopy (STM; Miles, 1997). A very fine tip is scanned across the sample attached to a surface, and the force (AFM) or the current (STM) is measured. The AFM method, which is the most useful for biological molecules, provides the height of the molecule as a function of the position of the scanning probe. Measurements can be made with the sample immersed in solution, allowing biochemical reactions to be followed. The resolution of the method (a few angstroms) allows imaging of duplex DNA and of the transcription of DNA by RNA polymerase (Kasas et al., 1997; Rippe et al., 1997). By coating the scanning tip with different surfaces, chemical forces between the tip and the sample molecule can be measured. Hydrogen bonding in nucleic acid bases (Boland and Ratner, 1995) and protein-ligand interactions (Chilkoti et al., 1995) have been measured in this manner. Single-molecule methods are just being developed and applied. The ability to hold,
Biophysical Analysis of Nucleic Acids
7.1.5 Current Protocols in Nucleic Acid Chemistry
move, stretch, and observe individual nucleic acid and protein molecules allows, for the first time, direct measurement of the forces involved in biochemical reactions. The coupling of mechanical and chemical forces can be measured and controlled. Molecular motions can be directly observed, as demonstrated for the rotation of the ATP synthase enzyme (Noji et al., 1997).
LITERATURE CITED Allain, F.T. and Varani, G. 1997. How accurately and precisely can RNA structure be determined by NMR? J. Mol. Biol. 267:338-351. Baeyens, K.J., De Bondt, H.L., Pardi, A., and Holbrook, S.R. 1996. A curved RNA helix incorporating an internal loop with G⋅A and A⋅A nonWatson-Crick base pairing. Proc. Natl. Acad. Sci. U.S.A. 93:12851-12855. Batey, R.T., Inada, M., Kujawinski, E., Puglisi, J.D., and Williamson, J.R. 1992. Preparation of isotopically labeled ribonucleotides for multidimensional NMR spectroscopy of RNA. Nucl. Acids Res. 20:4515-4523. Battiste, J.L., Tan, R., Frankel, A.D., and Williamson, J.R. 1994. Binding of an HIV Rev peptide to Rev responsive element RNA induces formation of purine-purine base pairs. Biochemistry 33:2741-2747.
Chen, X., Ramakrishnan, B., and Sundaralingam, M. 1997. Crystal structures of the side-by-side binding of distamycin to AT-containing DNA octamers d(ICITACIC) and d(ICATATIC). J. Mol. Biol. 267:1157-1170. Cheong, C. and Moore, P.B. 1992. Solution structure of an unusually stable RNA tetraplex containing G- and U-quartet structures. Biochemistry 31:8406-8414. Chilkoti, A., Boland, T., Ratner, B.D., and Stayton, P.S. 1995. The relationship between ligand-binding thermodynamics and protein-ligand interaction forces measured by atomic force microscopy. Biophys. J. 69:2125-2130. Chou, S.H., Zhu, L., and Reid, B.R. 1997. Sheared purine⋅purine pairing in biology. J. Mol. Biol. 267:1055-1067. Clegg, R.M., Murchie, A.I., Zechel, A., and Lilley, D.M. 1993. Observing the helical geometry of double-stranded DNA in solution by fluorescence resonance energy transfer. Proc. Natl. Acad. Sci. U.S.A. 90:2994-2998. Correll, C.C., Freeborn, B., Moore, P.B., and Steitz, T.A. 1997. Metals, motifs, and recognition in the crystal structure of a 5S rRNA domain. Cell 91:705-712.
Biou, V., Yaremchuk, A., Tukalo, M., and Cusack, S. 1994. The 2.9 Å crystal structure of T. thermophilus Seryl-tRNA synthetase complexed with tRNASer. Science 263:1404-1410.
Dickson, R.M., Norris, D.J., Tzeng, Y.L., and Moerner, W.E. 1996. Three-dimensional imaging of single molecules solvated in pores of poly(acrylamide) gels. Science 274:966-969.
Boland, T. and Ratner, B.D. 1995. Direct measurement of hydrogen bonding in DNA nucleotide bases by atomic force microscopy. Proc. Natl. Acad. Sci. U.S.A. 92:5297-5301.
Dickson, R.M., Cubitt, A.B., Tsien, R.Y., and Moerner, W.E. 1997. On/off blinking and switching behaviour of single molecules of green fluorescent protein. Nature 388:355-358.
Bustamante, C. 1991. Direct observation and manipulation of single DNA molecules using fluorescence microscopy. Annu. Rev. Biophys. Biophys. Chem. 20:415-446.
Dieckmann, T., Butcher, S.E., Sassanfar, M., Szostak, J.W., and Feigon, J. 1997. Mutant ATPbinding RNA aptamers reveal the structural basis for ligand binding. J. Mol. Biol. 273:467-478.
Butcher, S.E., Dieckmann, T., and Feigon, J. 1997. Solution structure of a GAAA tetraloop receptor RNA. EMBO J. 16:7490-7499.
Doucet, J., Benoit, J.P., Cruse, W.B., Prange, T., and Kennard, O. 1989. Coexistence of A- and B-form DNA in a single crystal lattice. Nature 337:190192.
Cate, J.H. and Doudna, J.A. 1996. Metal binding sites in the major groove of a large ribozyme domain. Structure 4:1221-1230. Cate, J.H., Gooding, A.R., Podell, E., Zhou, K., Golden, B.L., Kundrot, C.E., Cech, T.R., and Doudna, J.A. 1996a. Crystal structure of a group I ribozyme domain: Principles of RNA packing. Science 273:1678-1685. Cate, J.H., Gooding, A.R., Podell, E., Zhou, K., Golden, B.L., Szewczak, A.A., Kundrot, C.E., Cech, T.R., and Doudna, J.A. 1996b. RNA tertiary structure mediation by adenosine platforms. Science 273:1696-1699. Biophysical Analysis of Nucleic Acids
Chang, K.-Y. and Tinoco, I. Jr. 1997. The structure of an RNA “kissing” hairpin complex of the HIV TAR hairpin loop and its complement. J. Mol. Biol. 269:52-66.
Cate, J.H., Hanna, R.L., and Doudna, J.A. 1997. A magnesium ion core at the heart of a ribozyme domain. Nature Struct. Biol. 4:553-558.
Fan, P., Suri, A.K., Fiala, R., Live, D., and Patel, D.J. 1996. Molecular recognition in the FMN-RNA aptamer complex. J. Mol. Biol. 258:480-500. Farber, G.K. 1997. Laue crystallography: Lights! Camera! Action! Curr. Biol. 7:R352-R354. Foldesi, A., Yamakage, S.I., Nilsson, F.P., Maltseva, T.V., and Chattopadhyaya, J. 1996. The use of non-uniform deuterium labelling [‘NMR-window’] to study the NMR structure of a 21mer RNA hairpin. Nucl. Acids Res. 24:1187-1194. Fourmy, D., Recht, M.I., Blanchard, S.C., and Puglisi, J.D. 1996. Structure of the A site of Escherichia coli 16S ribosomal RNA complexed with an aminoglycoside antibiotic. Science 274:1367-1371.
7.1.6 Current Protocols in Nucleic Acid Chemistry
Franklin, R.E. and Gosling, R.G. 1953. Molecular configuration in sodium thymonucleate. Nature 171:740-741.
Louise-May, S., Auffinger, P., and Westhof, E. 1996. Calculations of nucleic acid conformations. Curr. Opin. Struct. Biol. 6:289-298.
Glusker, J.P., Lewis, M., and Rossi, M. 1994. Crystal Structure Analysis for Chemists and Biologists. VCH Publishers, New York.
Maglott, E.J. and Glick, G.D. 1997. A new method to monitor the rate of conformational transitions in RNA. Nucl. Acids Res. 25:3297-3301.
Gray, D.M., Hung, S.-H., and Johnson, K.H. 1995. Absorption and circular dichroism spectroscopy of nucleic acid duplexes and triplexes. Methods Enzymol. 246:19-34.
Miles, M. 1997. Scanning probe microscopy. Probing the future. Science 277:1845-1847.
Ha, T., Enderle, T., Ogletree, D.F., Chemla, D.S., Selvin, P.R., and Weiss, S. 1996. Probing the interaction between two single molecules: Fluorescence resonance energy transfer between a single donor and a single acceptor. Proc. Natl. Acad. Sci. U.S.A. 93:6264-6268. Holbrook, S.R., Cheong, C., Tinoco, I. Jr., and Kim, S.-H. 1991. Crystal structure of an RNA double helix incorporating a track of non-Watson-Crick base pairs. Nature 353:579-581. Special issue: James, T.L. (ed.). 1995. Nuclear magnetic resonance and nucleic acids. Methods Enzymol. 261:1-644. Jiang, F., Kumar, R.A., Jones, R.A., and Patel, D.J. 1996. Structural basis of RNA folding and recognition in an AMP-RNA aptamer complex. Nature 382:183-186. Johnson, K.H. and Gray, D.M. 1992. Analysis of an RNA pseudoknot structure by CD spectroscopy. J. Biomol. Struct. Dyn. 9:733-745. Kang, C.H., Zhang, X., Ratliff, R., Moyzis, R., and Rich, A. 1992. Crystal structure of four-stranded oxytricha telomeric DNA. Nature 356:126-131. Kasas, S., Thomson, N.H., Smith, B.L., Hansma, H.G., Zhu, X., Guthold, M., Bustamante, C., Kool, E.T., Kashlev, M., and Hansma, P.K. 1997. Escherichia coli RNA polymerase activity observed using atomic force microscopy. Biochemistry 36:461-468. Kellermayer, M.S., Smith, S.B., Granzier, H.L., and Bustamante, C. 1997. Folding-unfolding transitions in single titin molecules characterized with laser tweezers. Science 276:1112-1116. Kieft, J.S. and Tinoco, I. Jr. 1997. Solution structure of a metal-binding site in the major groove of RNA complexed with cobalt (III) hexammine. Structure 5:713-721.
Murchie, A.I.H., Clegg, R.M., von Kitzing, E., Duckett, D.R., Diekmann, S., and Lilley, D.M.J. 1989. Fluorescence energy transfer shows that the fourway DNA junction is a right-handed cross of antiparallel molecules. Nature 341:763-766. Nikonowicz, E.P., Sirr, A., Legault, P., Jucker, F.M., Baer, L.M., and Pardi, A. 1992. Preparation of 13 C and 15N labelled RNAs for heteronuclear multi-dimensional NMR studies. Nucl. Acids Res. 20:4507-4513. Noji, H., Yasuda, R., Yoshida, M., and Kinosita, K. Jr. 1997. Direct observation of the rotation of F1-ATPase. Nature 386:299-302. Passner, J.M. and Steitz, T.A. 1997. The structure of a CAP-DNA complex having two cAMP molecules bound to each monomer. Proc. Natl. Acad. Sci. U.S.A. 94:2843-2847. Pease, A.C., Solas, D., Sullivan, E.J., Cronin, M.T., Holmes, C.P., and Fodor, S.P. 1994. Light-generated oligonucleotide arrays for rapid DNA sequence analysis. Proc. Natl. Acad. Sci. U.S.A. 91:5022-5026. Pley, H.W., Flaherty, K.M., and McKay, D.B. 1994. Three-dimensional structure of a hammerhead ribozyme. Nature 372:68-74. Puglisi, J.D., Chen, L., Blanchard, S., and Frankel, A.D. 1995. Solution structure of a bovine immunodeficiency virus Tat-TAR peptide-RNA complex. Science 270:1200-1203. Radhakrishnan, I. and Patel, D.J. 1994. Solution structure and hydration patterns of a pyrimidine⋅purine⋅pyrimidine DNA triplex containing a novel T⋅CG base-triple. J. Mol. Biol. 241:600619. Ramos, A., Gubser, C.C., and Varani, G. 1997. Recent solution structures of RNA and its complexes with drugs, peptides and proteins. Curr. Opin. Struct. Biol. 7:317-323.
LeCuyer, K.A. and Crothers, D.M. 1994. Kinetics of an RNA conformational switch. Proc. Natl. Acad. Sci. U.S.A. 91:3373-3377.
Recht, M.I., Fourmy, D., Blanchard, S.C., Dahlquist, K.D., and Puglisi, J.D. 1996. RNA sequence determinants for aminoglycoside binding to an A-site rRNA model oligonucleotide. J. Mol. Biol. 262:421-436.
Lipari, G. and Szabo, A. 1982. Model-free approach to the interpretation of nuclear magnetic resonance relaxation in macromolecules. J. Am. Chem. Soc. 104:4546-4559.
Rief, M., Gautel, M., Oesterhelt, F., Fernandez, J.M., and Gaub, H.E. 1997. Reversible unfolding of individual titin immunoglobulin domains by AFM. Science 276:1109-1112.
Lipscomb, L.A., Peek, M.E., Morningstar, M.L., Verghis, S.M., Miller, E.M., Rich, A., Essigmann, J.M., and Williams, L.D. 1995. X-ray structure of a DNA decamer containing 7,8-dihydro-8-oxoguanine. Proc. Natl. Acad. Sci. U.S.A. 92:719-723.
Rippe, K., Guthold, M., von Hippel, P.H., and Bustamante, C. 1997. Transcriptional activation via DNA-looping: Visualization of intermediates in the activation pathway of E. coli RNA polymerase x sigma 54 holoenzyme by scanning force microscopy. J. Mol. Biol. 270:125-138.
Biophysical Analysis of Nucleic Acids
7.1.7 Current Protocols in Nucleic Acid Chemistry
Roberts, G.C.K. 1993. NMR of Macromolecules. A Practical Approach. IRL Press, New York. Rugar, D., Zuger, O., Hoen, S., Yanonni, C.S., Vieth, H.-M., and Kendrick, R.D. 1994. Force detection of nuclear magnetic resonance. Science 264:1560-1563. Saenger, W. 1984. Principles of Nucleic Acid Structure. Springer-Verlag, New York. SantaLucia, J. Jr., Shen, L.X., Cai, Z., Lewis, H., and Tinoco, I. Jr. 1995. Synthesis and NMR of RNA with selective isotopic enrichment in the base moieties. Nucl. Acids Res. 23:4913-4921. Scott, W.G., Finch, J.T., and Klug, A. 1995. The crystal structure of an all-RNA hammerhead ribozyme: A proposed mechanism for the RNA catalytic cleavage. Cell 81:991-1002. Scott, W.G., Murray, J.B., Arnold, J.R.P., Stoddard, B.L., and Klug, A. 1996. Capturing the structure of a catalytic RNA intermediate: The hammerhead ribozyme. Science 274:2065-2069. Shen, L.X. and Tinoco, I. Jr. 1995. The structure of an RNA pseudoknot that causes efficient frameshifting in mouse mammary tumor virus. J. Mol. Biol. 247:963-978. Smith, S.B., Finzi, L., and Bustamante, C. 1992. Direct mechanical measurements of the elasticity of single DNA molecules by using magnetic beads. Science 258:1122-1126. Tinoco, I. Jr. and Kieft, J.S. 1997. The ion core in RNA folding. Nature Struct. Biol. 4:509-512. Tolbert, T.J. and Williamson, J.R. 1996. Preparation of specifically deuterated RNA for NMR studies using a combination of chemical and enzymatic synthesis. J. Am. Chem. Soc. 118:7929-7940. Tuschl, T., Gohlke, C., Jovin, T.M., Westhof, E., and Eckstein, F. 1994. A three-dimensional model for the hammerhead ribozyme based on fluorescence measurements. Science 266:785-789. Varani, G. 1997. RNA-protein intermolecular recognition. Acc. Chem. Res. 30:189-195.
Varani, G. and Tinoco, I. Jr. 1991. RNA structure and NMR spectroscopy. Q. Rev. Biophys. 24:479-532. Wang, E., Malek, S., and Feigon, J. 1992. Structure of a G⋅T⋅A triplet in an intramolecular DNA triplex. Biochemistry 31:4838-4846. Watson, J.D. and Crick, F.H.C. 1953. Molecular structure of nucleic acids. Nature 171:737-738. Wemmer, D. 2000. Structure and dynamics by NMR. In Nucleic Acids: Structures, Properties, and Functions (V.A. Bloomfield, D.M. Crothers, and I. Tinoco, Jr., eds.) pp. 111-163. University Science Books, Mill Valley, Calif. Wemmer, D. and Dervan, P. 1997. Targeting the minor groove of DNA. Curr. Opin. Struct. Biol. 7:355-361. Wilkins, M.H.F., Stokes, A.R., and Wilson, H.R. 1953. Molecular structure of deoxypentose nucleic acids. Nature 171:738-740. Williamson, J.R. 1994. G-quartet structures in telomeric DNA. Annu. Rev. Biophys. Biomol. Struct. 23:703-786. Wu, M. and Turner, D.H. 1996. Solution structure of (rGCGGACGC)2 by two-dimensional NMR and the iterative relaxation matrix approach. Biochemistry 35:9677-9689. Wu, M., McDowell, J.A., and Turner, D.H. 1995. A periodic table of symmetric tandem mismatches in RNA. Biochemistry 34:3204-3211. Wüthrich, K. 1986. NMR of Proteins and Nucleic Acids. John Wiley & Sons, New York. Zimmerman, G.R., Jenison, R.D., Wick, C.L., Simorre, J.-P., and Pardi, A. 1997. Interlocking structural motifs mediate discrimination by a theophylline-binding motif. Nature Struct. Biol. 4:644-648.
Contributed by Ignacio Tinoco, Jr. University of California Lawrence Berkeley National Laboratory Berkeley, California
Biophysical Analysis of Nucleic Acids
7.1.8 Current Protocols in Nucleic Acid Chemistry
NMR Determination of Oligonucleotide Structure NMR, in conjunction with appropriate computational searching algorithms, has become the method of choice for determining solution structures. Methodology is now available to determine an accurate, high-precision (i.e., high-resolution) structure of nearly any DNA or RNA double helix of up to 15 to 20 base pairs (bp) via NMR if sufficient care and effort are expended (Schmitz and James, 1995). Care is imperative, as it is also possible to obtain worthless structures; easy-to-use restrained molecular dynamics (rMD) programs currently available may yield a structure even if the user has provided them with poor experimental data or has used an inappropriate conformational searching protocol. It is also feasible to determine the low-resolution structure of even larger oligonucleotides (possibly up to the size of tRNA). Although one can model a chemically reasonable, even useful, structure from small amounts of data, this does not constitute structure determination. Before resorting to the relatively timeconsuming task of “determining a high-resolution structure,” however, one should consider the knowledge to be gained as a result. In the case of proteins or single-stranded nucleic acids that fold into a tertiary structure, some useful insights into function can be derived from low-resolution structures or even models. The fairly subtle structural variations in a DNA or RNA⋅DNA helix that are sequence dependent, and consequently guide protein, mutagen, or drug recognition, demand a detailed high-resolution structure to be very useful. The discussion that follows will emphasize the steps necessary for determination of a high-resolution structure, with some mention of the variations for determining structures of nucleic acids possessing significant tertiary structure. The discussion will also be very strongly biased in describing approaches that have been successful in our lab. One book has appeared that focuses solely on NMR of nucleic acids (James, 1995). The reader can find there details about many topics discussed below, much additional information about RNA and DNA structure determination, and references.
UNIT 7.2
OVERVIEW OF OLIGONUCLEOTIDE STRUCTURE DETERMINATION BY NMR Before embarking upon nucleic acid structure determination via NMR, it is advisable to assess the likelihood of success. The size limitation of the method has already been mentioned. Another question to consider is whether the nucleic acid possesses features suggesting that it may have a structure. As a first consideration, solubility in water implies that success is likely. A single-stranded DNA or RNA that does not have a sequence capable of forming at least four or five Watson-Crick base pairs in a helix is highly unlikely to possess a sufficiently stable structure in aqueous solution—although the i-motif for oligo-C sequences at low pH is an obvious exception to this rule (Leroy et al., 1993). Nucleic acids, fortunately, are highly soluble in water. They may, however, aggregate via nonspecific interactions, especially in the presence of multivalent counterions or ligands. Aggregation will result in broadening of NMR resonances, with consequent loss of signal intensity and ability to resolve individual signals. A flow chart describing the process by which oligonucleotide structure is determined is presented in Figure 7.2.1. As with proteins, assignment of resonances is often the bottleneck in structure determination for structures exhibiting tertiary structure, such as many RNA elements. Making proton resonance assignments in a nucleic acid duplex, however, is fairly quick and straightforward (van de Ven and Hilbers, 1988). The pattern of cross-peaks in a proton two-dimensional nuclear Overhauser effect (2D NOE) spectrum makes distinction between left- and right-handed helices obvious. In the case of a right-handed helix, through-space connectivities between protons in sequential residues are evident from the appearance of cross-peaks corresponding to the pertinent protons in the 2D NOE spectrum. Figure 7.2.2 demonstrates the major sequential assignment strategy. With data in hand, it is often possible to “walk through” up to 200 proton assignments in a day or so. If the sequence is bound by a ligand or contains loops, bulges, or other deviations from a simple righthanded helix, however, assignment of resonances may require many months of work and
Contributed by Thomas L. James Current Protocols in Nucleic Acid Chemistry (2000) 7.2.1-7.2.16 Copyright © 2000 by John Wiley & Sons, Inc.
Biophysical Analysis of Nucleic Acids
7.2.1
establish appropriate solution conditions: e.g., pH, salts, detergents (stable, not aggregated)
assign resonances obtain 2D NOE spectrum and acquire interproton distance restraints obtain experimental structure restraints with error bars
search conformational space to define all structures consistent with experimental data and holonomic constraints, using algorithm such as DG or rMD
obtain correlation spectroscopy NMR spectrum and calculate torsion-angle restraints
establish holonomic constraints: bond lengths and angles, atom connectivities, steric limitations
TIME-AVERAGED STRUCTURE
DYNAMIC STRUCTURE
consider dynamic information: e.g., NMR relaxation parameters, H/D exchange data, reliably inconsistent time-averaged restraints
Figure 7.2.1 Scheme for deriving oligonucleotide structure using experimental NMR data in conjunction with conformational searching procedures such as distance geometry (DG) and restrained molecular dynamics (rMD; sometimes called simulated annealing). 2D NOE, two-dimensional nuclear Overhauser effect.
NMR Determination of Oligonucleotide Structure
many different types of 2D NMR experiments, including at the least total correlation spectroscopy (TOCSY). If the structure is sufficiently large or complicated, it may be necessary to use 13C or 15N labeling of the sample and multidimensional (e.g., 3D) heteronuclear NMR. The structure of any molecule can be determined given a sufficient number of experimental structural restraints—e.g., internuclear distances and bond torsion angles, together with the holonomic constraints of bond lengths, bond angles, and steric limitations. A key to the determination of good-quality structures is to
use as many structural restraints as possible. For high-resolution structures, the restraints should also possess the best accuracy possible. A means to determine the bounds or error bars on these structural restraints is also quite valuable, as at least an estimate will be needed for structure refinement. These experimental structural restraints are used with algorithms, such as distance geometry (DG) and rMD, that search conformational space to define all structures consistent with the experimental restraints. If these resulting structures are closely related and there is confidence
7.2.2 Current Protocols in Nucleic Acid Chemistry
A
3′ end
B G1
ω1 (ppm)
A3 G2 H8 or 6
A3 G2 H1′ BASE
G1 5′ end
ω2 (ppm)
T4
aromatic resonances
2D NOE spectrum
H1′ resonances
Figure 7.2.2 Schematic representations of (A) the sequential connectivity between protons in the sugar ring (shown as H1′) and protons in the base (H6 for pyrimidines and H8 for purines) and (B) the “walk” between the congruous cross-peaks in the proton homonuclear two-dimensional nuclear Overhauser effect (2D NOE) spectrum. Note, for example, that a particular H1′ proton has a 2D NOE cross-peak at a frequency ω1 corresponding to the resonance frequency of an aromatic proton (H6 or H8) from its own residue as well as at the resonance frequency of the H6 or H8 from its 3′-neighboring residue. Likewise, each aromatic proton “sees” its own and its neighbor residue’s H1′. While there is some overlap of regions, the different proton types, e.g., H6, H1′, H2′, etc., have their own characteristic range of resonance frequencies. In a real spectrum, there are many additional cross-peaks; i.e., a cross-peak should be found between a given proton and any other proton within ∼5 Å. In addition to the H1′-aromatic proton walk, there is an analogous H2′- (and, for DNA, H2′′) -to-aromatic proton walk. Of course, intrasugar ring protons yield cross-peaks. There is also a walk between sequential imino protons. The ability to assign these hundreds of resonances devolves from the entire pattern of interrelated connectivities; all cross-peaks must be accounted for. The only serious difficulty in making resonance assignments is that as the number of residues increases and as the structure becomes more monotonous, there is more overlap of cross-peaks obscuring discernment of connectivities. The overlap problem has driven the use of isotopic labeling and use of 3D or 4D NMR in order to spread out the overlapping cross-peaks.
that pertinent parts of conformational space have not been neglected, one can conclude that the structure has been determined. It is preferable that the array of structures generated by the search algorithms be large enough to map out the conformational space that will accommodate all available experimental data. With the generation of a number of structures satisfying the experimental data, an assessment of the structures is in order. Typically, one compares the atomic coordinates of all of the satisfactory structures to obtain an assessment of the apparent precision or “resolution” of structure determination. In addition to visually inspecting structural features, it is often useful to calculate torsion angles and helical
parameters to compare with canonical or other nucleic acid structures.
BASIC NMR SPECTRAL PARAMETERS Before considering in detail the determination of oligonucleotide structures from NMR data, it is worthwhile to step back and briefly consider the nature of the NMR data that will be examined. A rigorous, mathematically oriented coverage of the principles can be found in texts by Abragam (1961) and Ernst et al. (1987), while a good introductory text is that by Derome (1987). With an eye towards protein structure determination, a good description of background and applications is given by Cavanagh et al. (1996).
Biophysical Analysis of Nucleic Acids
7.2.3 Current Protocols in Nucleic Acid Chemistry
The nuclear magnetic resonance phenomenon can be described succinctly as follows. If a sample is placed in a magnetic field and is subjected to radiofrequency (RF) energy at the appropriate frequency, nuclei in the sample can absorb the energy. The frequency of the radiation necessary for absorption of energy—i.e., the resonance frequency—depends on the type of nucleus (e.g., 1H and 31P have a different value for the gyromagnetic ratio γ) and also on the chemical environment of the nucleus. For example, the methyl, methylene, and hydroxyl protons of ethanol absorb at three different frequencies, and H2 protons of two different adenine residues in a DNA duplex absorb at different frequencies, since they are in different chemical environments by virtue of sequencedependent structural differences. After absorption of energy by the nuclei, the length of time and the manner in which the nuclei dissipate that energy can also be used to reveal information regarding a variety of dynamic processes. The magnetic field Blocal perceived by the nuclei in a molecule will be very slightly altered from the field Bo in which the sample is placed, as a result of currents in the molecule’s electrons. The frequency ν at which a nucleus absorbs RF energy depends linearly on the local magnetic field: ν = γBlocal / 2π Equation 7.2.1
NMR Determination of Oligonucleotide Structure
Equation 7.2.1 is often referred to as the Larmor equation. As the precise chemical environment will determine the local magnetic field, different nuclei of the same type will absorb energy at slightly different frequencies. Because the local magnetic field variations are small compared with that of the magnet in which the sample is placed, the frequency shifts will be relatively small. So if the absorption frequency ν is several hundred megahertz, differences in resonance frequencies for two different hydrogen nuclei will be on the order of several hertz. Although one cannot easily determine absolute RFs to an accuracy of ±1 Hz, it is possible to determine the relative positions of two signals in the NMR spectrum with even greater accuracy than this. Consequently, a reference signal is chosen, and the difference between the position of the signal of interest and that of the reference, termed the chemical shift, is assessed. The chemical shift is usually expressed in terms of parts per million (ppm), actually a dimensionless number, by
δ=
νref − νsample × 106 νref
Equation 7.2.2
where the difference between the resonance frequency of the reference and that of the sample nucleus (νref − νsample) measured in hertz (e.g., 90 Hz) divided by the spectrometer’s operating frequency (e.g., 600 MHz) gives the chemical shift (e.g., 0.15 ppm). Typical ranges in chemical shifts for nuclei in biochemically important samples are 15 ppm for 1H, 250 ppm for 13C, 400 ppm for 15N, and 35 ppm for 31P. A nucleus with a magnetic moment may interact with other nuclear spins resulting in mutual splitting of the NMR signal from each nucleus into multiplets. This is termed scalar coupling. For 1H, 13C, 15N, and 31P, the number of components into which a signal is split is n + 1, where n is the number of other nuclei interacting with the nucleus. For example, a nucleus (e.g., 13C or 1H) interacting with three methyl protons will give a signal split into a quartet. To a first approximation, the relative intensities of the multiplets are given by binomial coefficients: 1:1 for a doublet, 1:2:1 for a triplet, and 1:3:3:1 for a quartet. The difference between any two adjacent components of a multiplet is the same and yields the value of the spin-spin coupling constant J (in hertz). One important feature of spin-spin splitting is that it is independent of magnetic field strength. Therefore, increasing the magnetic field strength will increase the chemical shift difference between two peaks in hertz (not parts per million) but will not alter the coupling constant. To simplify a spectrum, especially with 13C and 15N NMR, it is common to employ decoupling. Strong irradiation of the protons at their resonance frequency will cause a collapse of the multiplet in the 13C or 15N resonance into a singlet. In oligonucleotide samples, the signal width may be broader than the splitting, so the latter may not always be apparent. However, measurement of J can provide valuable structural information (as discussed below; see section on Acquisition of Torsion-Angle Restraints). While it is possible to obtain chemical-shift and coupling-constant information from a onedimensional (1D) NMR spectrum for small organic molecules, it is usually necessary to employ 2D (or sometimes 3D and 4D) NMR for olignonucleotides. Just as a biochemist may employ two-dimensional gel electrophoresis to spread out peaks in two dimensions, it is possible to resolve the many additional NMR sig-
7.2.4 Current Protocols in Nucleic Acid Chemistry
nals coming from larger biopolymers by utilizing additional dimensions. While the plethora of different possible ways to achieve this can be complicated, basically, by using different combinations of pulses of RF energy, it is possible not only to spread the signals out over two or three dimensions, but also to establish relationships between the nuclei giving rise to the different signals. Nuclei that are scalar-coupled to one another can be identified by the appearance of cross-peaks in the 2D NMR spectrum. Nuclei that are in close proximity in the molecule can be identified via the nuclear Overhauser effect manifest in signal intensities.
ACQUISITION OF INTERPROTON DISTANCE RESTRAINTS All structural determinations to date have utilized interproton distance restraints from NOE data. Some have also utilized torsionangle restraints from the oligonucleotide backbone or sugar ring.
2D NOE Spectral Requirements The intensities of the cross-peaks in a 2D NOE spectrum are related to the distances between protons that are in close spatial proximity (5 to 6 Å) in a structure, and can be used to estimate those distances. One might anticipate that accurate cross-peak intensities would be desired in order to obtain accurate distances. Acquiring intensities accurately reflecting the interproton distance generally requires that (1) the pulse repetition time (TR) of the experiment be long compared with the longest proton T1 (spin-lattice relaxation time) value, i.e., such that relaxation be nearly complete; (2) contributions to the cross-peaks from multiple quantum coherences be minimal; (3) good flat baseplanes in the well-digitized spectrum be produced to enable good integration; and (4) peak overlap be minimal (or very reliable deconvolution software be available for spectral analysis). Note that these requirements are not very stringent. For example, if (to take an arbitrary figure) an error of 5% in the measured interproton distance r were acceptable, an error in cross-peak intensity of ∼35% could be tolerated. (This assumes only the first-order relationship that cross-peak intensity varies as r−6, as discussed below; see section on Extracting Interproton Distances from 2D NOE CrossPeak Intensities). The adenine H2 proton in a typical DNA duplex has a T1 value of ∼5 sec, and T1 values are even longer in RNA. Using a TR of 2 sec, as commonly seen in the literature, would a render cross-peak involving an adenine
H2 proton only 14% of its fully relaxed intensity. As other peaks will also have attenuated intensities, resulting distance errors will not be as large as this might imply, since relative peak intensities are used in making distance estimates (also discussed below). However, our lab has typically used TR values of 8 to 12 sec to minimize intensity distortions due to incomplete relaxation. More recently, it has also become possible to correct for intensity distortions caused by using a short TR (Liu et al., 1996). Due to the dynamic range problem, acquiring 2D NOE spectra for oligonucleotides in H2O solution, which is required for observation of imino and amino proton resonances, is more trouble than obtaining spectra for oligonucleotides in a D2O solution, which is sufficient for nonexchangeable proton data. Most structural studies of oligonucleotides have not utilized distances entailing exchangeable protons. That has generally been wise, since imino and amino protons of nucleic acids exchange with bulk water and the intensity of 2D NOE crosspeaks involving exchangeable protons can be strongly altered by that exchange. If such exchange is not taken into account, calculation of distances to exchangeable protons is prone to significant error (Landy and Rao, 1989). However, the additional structural information that can be obtained by using such distances is extremely valuable. For example, cross-strand distances define the spatial relationship between the two strands in duplex structures, but there are very few such distances entailing solely nonexchangeable protons. The bases in DNA and RNA have a paucity of protons, so structural data provided by those imino and amino protons is far more valuable than structural data emanating from the proton-rich sugars. We have shown that we can incorporate effects of exchange with bulk water in the case of exchangeable protons and hence extract these important distance restraints (Liu et al., 1993). There is a simple, useful method for measuring the exchange rate that we have found useful (Adams and Lerner, 1992); I strongly recommend making the effort, since the information gained is so valuable.
Extracting Interproton Distances from 2D NOE Cross-Peak Intensities The effect of cross-relaxation between two neighboring protons during the mixing time period τm of the 2D NOE experiment is to transfer magnetization between them (Macura and Ernst, 1980). The efficiency of this transfer
Biophysical Analysis of Nucleic Acids
7.2.5 Current Protocols in Nucleic Acid Chemistry
depends on (1) the length of τm, (2) the distance between the two protons, (3) surrounding protons, and (4) the rate of molecular motion, generally characterized by a correlation time τc (Keepers and James, 1984). With transfer of magnetization, the cross-peak intensities in the spectrum will be modified. Consequently, cross-peak intensities have structural information—i.e., distances—embedded in them. In interesting molecules, the two protons giving rise to a particular cross-peak are not the only protons in the molecule. Rather, they belong to an array of all protons in the molecule that, in principle, experience dipole-dipole interactions with all the others. So cross-relaxation between the two protons is part of a coupled relaxation network. There are different methods of analyzing 2D NOE spectra to obtain interproton distances. The commonly employed two-spin or isolated-spin-pair approximation (ISPA) can be used: 1
rij = rref(aref / aij)6 Equation 7.2.3
where rij is the interproton distance to be estimated and aij is the corresponding 2D NOE cross-peak intensity, while rref and aref are, respectively, a known interproton distance (e.g., cytosine H5-to-H6 distance = 2.46Å) and its cross-peak intensity. This equation results from truncation after the linear term of the Taylor-series expansion of the complete relaxation rate expression accounting for all proton dipole-dipole interactions: 1 a(τm ) = e−Rτm = 1 − Rτm + R2τ2m − ⋅ ⋅ ⋅ 2 Equation 7.2.4
NMR Determination of Oligonucleotide Structure
where a is the matrix of 2D NOE intensities and R is the matrix describing the complete dipole-dipole relaxation network (i.e., all interproton interactions). The off-diagonal element Rij (row i, column j) of the relaxation matrix R corresponds to the cross-relaxation rate between protons i and j due to the fluctuating (from Brownian motion) dipolar interaction between them, and hence depends on the internuclear distance as (rij)−6 and on the rate of reorientation of the ij vector relative to the magnetic field in which the nucleic acid sample has been placed. The truncation leading to Equation 7.2.3 is valid in the limit of short mixing time, which is equivalent to assuming each cross-peak intensity depends only on the cross-relaxation rate between the two pertaining protons, i.e.,
ISPA. ISPA can lead to sizable systematic, as well as random, distance errors due to multispin effects commonly called “spin diffusion” (Keepers and James, 1984; Borgias and James, 1989; Post et al., 1990). The general method of assessing the impact of spin diffusion has been to obtain 2D NOE buildup curves, i.e., crosspeak intensities as a function of mixing time. Spin diffusion is usually considered negligible if the buildup is initially linear. Constructing an NOE buildup curve requires that several NOE spectra be analyzed, and still cannot properly account for multispin effects (Keepers and James, 1984; Post et al., 1990; Thomas et al., 1991). For example, for mixing times generally accepted as being sufficiently short (i.e., 50 to 100 msec) and not including internal motions, ISPA can result in systematic errors of 45% to 80% in actual distances >3.5 Å, a range that is quite important in defining molecular structure. Limiting analysis to 2D NOE spectra with short mixing times also yields cross-peaks with lower signal-to-noise ratio, limiting the number of distances that can be extracted. Improving the accuracy of distances derived from NOE cross-peak intensities requires consideration of all structure-dependent relaxation pathways of the entire proton system, which can be done with complete relaxation matrix methods (Keepers and James, 1984; Borgias and James, 1989; Post et al., 1990). For example, theoretical NOE spectra including spin-diffusion effects can be calculated for a proposed structure. Conversely, this approach can be used to compute accurate distances from NOE intensities acquired at any mixing time. Rearrangement of Equation 7.2.4 leads to a(τm) −ln a(0) R= τm Equation 7.2.5
Complete relaxation matrix methods utilize linear algebra, so Equation 7.2.5 can be solved by matrix diagonalization techniques to yield the relaxation matrix R (Borgias and James, 1989, 1990). Some cross-peak intensities from NOE spectra of nucleic acids cannot be measured due to peak overlap and spectral signal-tonoise limitations, so it has been found expedient to utilize a hybrid matrix approach, where all experimentally unobserved NOEs are taken from a model structure that is presumed to be similar to the actual molecular structure (Boelens et al., 1989; Borgias and James, 1989, 1990; Post et al., 1990). The equation using the
7.2.6 Current Protocols in Nucleic Acid Chemistry
hybrid matrix composed of experimental and model intensities is solved iteratively. It has been found that the model structure utilized is not very significant (Borgias and James, 1989, 1990). Exact details of methods differ, but the elements of the relaxation matrix are varied until a consistent fit to the experimentally observed NOEs is obtained. The programs IRMA (Boelens et al., 1989) and MORASS (Post et al., 1990) integrate this process directly into the conformational search for the final structure via rMD, but the program MARDIGRAS (Borgias and James, 1989, 1990; Liu et al., 1995) simply varies cross-relaxation rates until the best solution is found. Interproton distances are readily obtained from the converged relaxation matrix elements. This latter approach offers the advantage that the conformational search methodology, force field, or penalty function does not influence the structural restraints, so the subsequent conformational search uses independent restraints that have not been biased by the search methodology itself. An additional advantage is that it is not necessary that all distances be satisfied by a single structure. A set of MARDIGRAS-derived distances can exhibit some mutually inconsistent restraints resulting from conformational flexibility and leading to dynamically averaged, and therefore possibly inconsistent, distances (Ulyanov et al., 1995). Consequently, examination of these distance inconsistencies may aid in recognition of and insights about conformational flexibility (Ulyanov et al., 1995; Tonelli and James, 1998). More information and access to MARDIGRAS can be found at the web page http://picasso. ucsf.edu/software.html. An important aspect of the MARDIGRAS algorithm is that all intensities are used to obtain all distances simultaneously—i.e., the entire cross-relaxation network is employed; this is in distinct contrast to the old paradigm of a one-to-one mapping of intensities and distances. This means that a cross-peak that one would ordinarily not observe at low mixing times can be reliably examined at longer mixing times; even though the cross-peak intensity itself may be dominated by so-called spin diffusion, the cross-relaxation rate and consequently distance may be reliably inferred from the overall pattern of intensities, since the existence of that pair of protons will influence the intensities of other cross-peaks (Borgias and James, 1990; Liu et al., 1995). Any method used for calculating interproton distances from NOE data requires an estimate
of the correlation time. A detailed assessment of methods for evaluating correlation times has been presented elsewhere (Lane, 1995). In general, an isotropic correlation time is assumed for the overall tumbling of a nucleic acid. This assumption is reasonable up to a length of ∼15 bp in a duplex, which might be considered as a cylinder. The assumption that one can treat a biopolymer as a sphere works quite well, as long as the molecule is not so distended that the ratio of the long molecular axis to the short molecular axis exceeds ∼2.5. Whenever possible, it is best to estimate the correlation time using 13C relaxation-time measurements, since proton relaxation times are influenced by the multispin effects noted above. Natural-abundance 13C relaxation-time measurements in nucleic acids can be time consuming, however. While subject to error, obtaining an estimate of τc from the ratio of T1 and T2 relaxation times for individual resolved protons is perhaps the easiest approach. A recent comparison of correlation times obtained using these two methods yielded comparable values (Tonelli et al., 1998). Although this rule of thumb has not been published anywhere, one can obtain a decent estimate of the correlation time for tumbling of a nucleic acid or protein in aqueous solution at approximately room temperature by taking the value of the molecular mass (daltons) in picoseconds and dividing by 2; for example, a structured 14,000-Da biopolymer has τc ≈ 7 nsec. Fortunately, τc accuracy is not paramount: an error by a factor of 2 translates to distance errors of no more than 12%, usually much less. Nevertheless, it is advisable to use both upper and lower bound estimates for τc in calculating distance restraints.
Estimation of Interproton Distance Bounds When searching conformational space to find structures consistent with a set of experimental data, it is necessary to have an estimate of the accuracy of the distance restraints. Error estimates on restraints are absolutely required for setting bounds in DG or flat-well size in rMD calculations. The refinement process should yield a final structure in the context of a conformational envelope reflecting the intrinsic limitations of the experimental data and the method. Estimates of the interproton distance error, reflected by the upper and lower bounds assigned, vary widely throughout the literature. Tighter distance bounds (smaller error bars) lead to a higher-resolution structure; however,
Biophysical Analysis of Nucleic Acids
7.2.7 Current Protocols in Nucleic Acid Chemistry
NMR Determination of Oligonucleotide Structure
distance bounds made tighter than warranted by experimental accuracy mislead to a highly precise (small atomic root-mean-square-deviation, RMSD) but incorrect structure (Thomas et al., 1991). So one needs bounds as tight as possible but not so tight that the real distance can lie outside them. What this means is that for some purposes it may be perfectly reasonable to utilize ISPA to estimate distances, but the spread between upper and lower bounds should be made quite large. More accurate distances obtainable via MARDIGRAS minimize the possibility of an estimated distance lying outside the bounds. MARDIGRAS can also aid our choice of bounds, set individually for each proton pair. In particular, it is recommended that bounds be set using the RANDMARDI option now incorporated into the program (Liu et al., 1995). This procedure uses an absolute noise error (e.g., conservatively using the size of the smallest NOE) and an additional relative error (e.g., 10% to 20%) in distance bounds determination. In addition, different motional models with varying correlation times are used in combination with different starting geometries. With this procedure, typical error bars amount to ∼10% for 3-Å distances but may be >30% for 5-Å distances, which derive from cross-peaks with lower signal to noise. The compensation for the broader bounds is that distances up to 7 Å (entailing methyls) can be reliably determined (Liu et al., 1995). Distances are determined from a single 2D NOE spectrum using MARDIGRAS rather than using buildup curves. It is still desirable to obtain spectra at a few different mixing times. Depending on the internuclear distance and proton environment, different mixing times will be optimum for different proton pairs. This enables distances determined independently from spectra acquired at different mixing times to be compared. Of course, the range of distance values measured from different spectra for any given proton pair can aid in the choice of bounds. MARDIGRAS calculates bounds for distances to protons undergoing motional or overlap averaging, i.e., methyl, methylene, and aromatic protons (Liu et al., 1992). With MARDIGRAS, distances are calculated from individual cross-relaxation rates in the converged rate matrix. Distances entailing protons averaged by motion or spectral overlap may be in serious error if the averaging is ignored. The cross-relaxation rates in these cases will depend on orientation as well as distance. MARDIGRAS does a second level of iteration, varying
the orientation and distances of all dipole-dipole interactions to find the best fit (Liu et al., 1992). Most importantly, however, MARDIGRAS lists the distances corresponding to the worst-case geometries, enabling upper and lower bounds to be set for distances involving protons averaged by either overlap or internal motions. As noted above, the effects of chemical exchange should also be taken into account in setting bounds for exchangeable protons (Liu et al., 1993).
ACQUISITION OF TORSION-ANGLE RESTRAINTS Bond torsion angles can be determined using Karplus correlations with vicinal coupling constants, which can be derived from various correlation spectroscopic techniques—e.g., E.COSY, PCOSY, and double-quantum-filtered COSY (2QF-COSY; Piantini et al., 1982; Marion and Bax, 1988). In principal, these should make it possible to determine deoxyribose-ring pucker conformations. A detailed description of the methodology (Schmitz and James, 1995) should be consulted by anyone planning on employing this technique. Broad lines prevent direct analysis of nearly all coupling constants in DNA oligomers greater than ∼8 bp in length. Consequently, most labs determine only those that can be readily measured, or determine sums of coupling constants—e.g., ΣH2′, the sum of all coupling constants involving the H2′ proton of a deoxyribose ring. We have found that limiting analysis to these can leave ambiguities, but fitting of experimental cross-peaks to cross-peaks simulated using the program SPHINX (Widmer and Wüthrich, 1987) enables extraction of vicinal coupling constants and, subsequently, torsion-angle restraints (Celda et al., 1989; Schmitz et al., 1990; Gonzàlez et al., 1994). This entails extensive peak-shape analysis, and typical errors range from ±0.3 Hz for the best-defined coupling constants, i.e., JH1′H2′′ and JH1′H2′, to ±1 Hz or more for JH3′H4′. The major difficulty is in establishing the correct linewidth to be employed. However, the choice of linewidth can usually be constrained such that limits (i.e., the bounds) on torsion angles describing sugar pucker can be established. Dipolar effects can influence the scalar-coupling-constant values (Harbison, 1993), with the effects becoming significant at larger correlation times (Zhu et al., 1995). Calculations (most unpublished) have shown that the dipolar effects are small for correlation times <5 nsec— i.e., not significant compared with experimen-
7.2.8 Current Protocols in Nucleic Acid Chemistry
tal errors. This means that for room-temperature studies of duplexes up to ∼14 bp in length, dipolar effects on the sugar three-bond coupling constants will not be detectable. For larger nucleic acids, the temperature can be raised to keep τc <5 nsec (Conte et al., 1996). For larger nucleic acids, it is also possible to obtain more limited, but useful, information: the sum of the vicinal coupling constants for a particular proton, e.g., ΣH1′, is not affected by dipolar relaxation due to compensatory effects for individual coupling constants, JH1′H2′′ and JH1′H2′ (Conte et al., 1996). The torsion angles for deoxyribose rings are determined using a parameterization of the relationship between torsion angles and coupling constants (Rinkel and Altona, 1987; Wijmenga et al., 1993; Gonzàlez et al., 1994; Schmitz and James, 1995). In studies of about a dozen DNA and RNA⋅DNA duplexes, examining all scalar coupling–based cross-peaks, it has been found that a single conformer will rarely account for all measured coupling constants within a sugar ring, but reasonable fits are obtained with a two-state model representing a rapid interconversion between S- (i.e., C2′-endo) and N-type (i.e., C3′-endo) sugar puckers (Altona and Sundaralingam, 1972). It should be evident in considering the shape of a Karplus curve (vicinal coupling constant versus dihedral angle) that conformational exchange will not average all coupling constants the same, so that if one has measured them all they cannot simultaneously fit a single conformation. This highlights the utility of using the more time-consuming SPHINX fitting of spectra to extract more coupling-constant data; with just a few coupling constants from each sugar ring, the data can be fit to a single averaged conformer, but one that will not necessarily correspond to any of the exchanging conformers (Gonzàlez et al., 1994). To employ the simplest model to account for the data, one can employ the two-state model. The pseudorotational phase angles of the major S- and minor N-conformers (PS and PN, respectively), as well as the relative populations and amplitudes of these conformers, are typically derived. For nonterminal nucleotides, we generally find that the S-conformer dominates, being populated 70% to 95% of the time with some exceptions.
STRUCTURE REFINEMENT The ability to determine solution structures by NMR is limited by the quantity, accuracy, and distribution of distance and torsion-angle restraints that can be extracted from the NMR
data. Obtaining enough restraints is a paramount requirement and will significantly counterbalance a lack of precisely determined restraints. As noted, interproton distances are essential for structure determination and a complete relaxation matrix approach will enable more numerous, more accurate, and more precise distances to be used. In principle, one should try to determine all structures that will satisfy the structural restraints taking into account experimental error—i.e., upper and lower bounds are specified, so that the accessible conformational space can be mapped out. Basically, a large number of structures are derived, all of which will fit the data. This is a source of irritation for scientists accustomed to the single structure typically reported in x-ray or modeling studies. (With arm-twisting, however, it is possible to come up with a single structure that will satisfy the biochemist or modeler.) In some sense, the family of structures gives a fuzzy picture of the structure. With more restraints, more accurate restraints, and tighter (but not unreasonable) bounds, the structural picture becomes sharper. Methods entailing systematic searches of conformational space have been advocated, but for biopolymers they are computationally too expensive at present. So an intelligently restricted search of conformational space is performed using a conformational search engine, which attempts to find a global minimum in fitting a structure with the requisite primary sequence connectivities, bond lengths, and bond angles and experimental restraint bounds; it is also common to include additional chemical force-field information as well. While it is possible simultaneously to refine restraints and structure iteratively, I advocate separating the restraint-determination step and the structuregeneration step. This should diminish any possibility of being trapped in a local energy minimum in the vicinity of early-iteration structures that are trying to satisfy inaccurate restraints. Also, individual assessment of the bounds for each structural restraint can be performed and then used in the structure-refinement procedure. And, as noted, one may also discern conformational exchange via inconsistent but accurate distance values. DG and rMD (also known as simulated annealing) calculations are commonly used for biopolymer structure generation and refinement. Although DG has been employed for nucleic acid structure determination, it typically produces structures that need subsequent refinement to account for molecular energetic
Biophysical Analysis of Nucleic Acids
7.2.9 Current Protocols in Nucleic Acid Chemistry
considerations, generally via rMD calculations, to obtain energetically feasible structures.
Restrained Molecular Dynamics Simulations The most commonly employed search technique is rMD (e.g., AMBER, GROMOS, or XPLOR). It should be understood that an rMD simulation is not the same as an MD simulation, which is generally concerned with following the molecular motions of a molecule over the femtosecond-to-nanosecond time scale. Rather, rMD uses the mathematics of MD to search conformational space: the kinetic energy in an MD simulation allows energy barriers of amplitude approximately kT (the Boltzmann constant multiplied by the absolute temperature) to be surmounted in a search for the global energetic minimum that maintains a balance between the “classical” energy terms and the experimental restraints. In rMD, the empirical force field is modified to incorporate a penalty term for not matching experimental NMR restraints: Vtotal = Vbond + Vangle + Vdihedral + Vvan der Waals + Vcoulomb + VH bond + VNOE + V J coupling Equation 7.2.6
NMR Determination of Oligonucleotide Structure
The first five terms monitor the classical potential energy of the molecule. There may or may not be an explicit term used to maintain hydrogen bonds; restraints may be experimentally justified, for example, by observation of particular imino proton signals in a spectrum. The final two terms serve as penalty functions, monitoring the NOE-derived distance restraints and scalar coupling–derived torsionangle restraints, respectively. Some studies have not incorporated torsion-angle restraints, but all have utilized distance restraints. Different functional forms have been employed for these penalty functions, but the consensus is a flat-well potential with quadratic boundaries beyond the experimental upper and lower distance bounds; for example, that is found in AMBER. The bounds determined via MARDIGRAS can thus determine the distance range for each individual proton pair over which no penalty will be exacted. More information regarding access to AMBER, X-PLOR, and GROMOS can be found at the web pages, respectively, http://www.amber.ucsf.edu/amber/amber.html, http://xplor.csb.yale.edu/xplor-info.html, and http://igc.ethz.ch/gromos.
While force constants to be used for the conventional energy terms are well established, that is not the case for the experimental restraint-energy terms. We have consequently examined the effects of changing the force constant for the restraint-energy terms in some detail for four different DNA duplexes. While the apparently optimum value varies somewhat for different duplexes, as long as the force constant is in the range 10 to 40 kcal mol−1 Å−2, for simulations near 300 K the exact value chosen is not too important. On the basis of our experience, I would recommend that a value of 20 kcal mol−1 Å−2 be used in the absence of an evaluation for any particular nucleic acid; this seems to work fine for RNA as well. The forceconstant value for torsion angle–restraint violations should be about four times larger than that for distance-restraint violations. Of course, in a simulated annealing protocol where the temperature is raised, the force constants should be commensurably increased. For rMD simulations, starting structures should be chosen in different regions of conformational space: e.g., for a DNA duplex, one would use A-DNA, B-DNA, and possibly some other model structure. In addition, different random initial trajectories should be used for each starting structure. The “final” structure reported for any particular rMD run is in fact not from the final step in the calculation. A trajectory (rMD simulation) might typically run for 30 psec in 1-fsec steps. An average set of atomic coordinates can be obtained from the coordinates of say 200 structures in the last 5 psec of the run when the search has stabilized. As this average set of coordinates may not represent a reasonable structure, that structure is subjected to restrained minimization—i.e., the energy is minimized with inclusion of the experimental restraint violation terms. For any method, successful refinement requires convergence: i.e., essentially the same final structure is obtained from different starting models and starting trajectories using a reasonable search protocol (for a detailed discussion see Schmitz and James, 1995). In reality, the structures resulting from different rMD runs will not be identical, but they should be similar—i.e., with small atomic RMSD between the individual structures little bigger than atomic displacements from librational motions (∼0.5 Å); generally <1.0 Å is considered adequate. For rMD, it is typically difficult to obtain convergence in reasonable computational times if the starting structure lies far from the final structure. For proteins, it has been found most
7.2.10 Current Protocols in Nucleic Acid Chemistry
efficient to use DG initially and then utilize rMD for subsequent refinement. For nucleic acid duplexes, using any right-handed helix, e.g., A or B form, is sufficient for rMD. Most DG programs do not work at all or do not work well for nucleic acids. For other RNA or DNA structures, we have found the program DYANA to work well and efficiently. DYANA performs torsion-angle dynamics, gaining its efficiency from using internal rather than Cartesian coordinates, which decreases the number of parameters by an order of magnitude. It is best to use rMD, e.g., with AMBER, for final refinement of each of the acceptable structures found by running DYANA. More information regarding access to DYANA can be found on the web page http://www.mol.biol.ethz.ch/dyana. The envelope of individual structures resulting from the multiple conformational search runs, which appears as a “fuzzy” structure, is perhaps a better representation of reality, but it is typically simpler to display a single final structure. That is obtained from the “final” structures resulting from the different trajectories being averaged and that average structure subjected to restrained minimization. There are methods other than those mentioned above for generating structures from NMR data, but there are few cases where different structure-refinement methods have been independently applied to the same data set and the resulting structures compared. A Monte Carlo search in torsion-angle space has been developed using generalized helical parameters, rather than Cartesian coordinates, to define DNA conformation for efficiency (Ulyanov et al., 1993). Using an idealized geometry for the aromatic rings eliminates the small distortions sometimes observed with structures emanating from rMD, which result from the force field permitting some distortion of bond angles and lengths in a compromise to fit the experimental restraints. Restrained Monte Carlo (rMC) calculations have been used on two DNA duplexes (Ulyanov et al., 1993; Tonelli et al., 1998). Convergence of final structures via rMC is easily achieved from A-DNA, B-DNA, or other right-handed DNA models (atomic RMSD <0.3 Å for all rMC simulations in one study) without use of torsion-angle restraints. For rMD, it is difficult (but sometimes possible) to achieve convergence to a global minimum starting from A-DNA without torsion-angle restraints. The structures resulting from rMC and rMD are in agreement—RMSD <0.5 Å for all structures generated—despite the different force fields used and despite the fact that the
rMD calculation used deoxyribose torsion-angle restraints. The structures generated with rMC and rMD were both in reasonable accord with experimental 2D NOE peak intensities and with experimental scalar-coupling data from 2QF-COSY spectra. Agreement with the scalar-coupling data is particularly satisfying, since those data were used in the rMD refinement but not the rMC refinement.
ASSESSMENT OF STRUCTURE QUALITY It is clear that the methodology exists to determine the structure of a small nucleic acid. Beyond adhering to the cautionary notes cited in the sections above, how can one be confident that the structure is correct? A detailed discussion of this question has been presented (James, 1994).
Number of Restraints We have examined the effect of the number of restraints available on the structure determined in one DNA duplex where we had on average 20 distance restraints and 5 torsion-angle restraints per residue (Weisz et al., 1994). While a structure is fairly well restrained by ∼10 distance restraints and 5 torsion angle restraints per nucleotide, we have found that it is better defined by ∼15 distance restraints per residue, along with the torsion angle restraints. More than ∼15 distance restraints provides redundant information, and the structure determined is little affected by additional restraints. It is assumed, of course, that the restraints are fairly evenly distributed across the molecule. If one has much less than ten restraints per residue, “structure determination” is the wrong term to apply; “modeling” would be more appropriate, as the chemical nature of the force field begins to dominate the resulting structure, with the experimental restraint data only providing some limits on that structure. As noted earlier, however, a model created with limited experimental restraints may still provide very valuable insight.
Atomic Root-Mean-Square-Deviation It is common to cite a value of the atomic RMSD among the ensemble of structures fitting the experimental data (as has been done earlier in this unit). One should be cautious, however, about interpreting the RMSD in terms of the quality of the structure derived or comparing RMSD values from one study to another. In fact, one should be careful about trying to push the value too low. It is possible to make the RMSD smaller by various means—e.g., by
Biophysical Analysis of Nucleic Acids
7.2.11 Current Protocols in Nucleic Acid Chemistry
initiation complex (Mujeeb et al., 1998), which is essential for packaging and replication of the HIV-1 virus. The two identical 23-nucleotide RNA strands each form hairpin structures with the loops of each hairpin containing the hexameric CGCGCG palindrome. The loops from two hairpins, as well the stem regions, can thus form a helix in creating a so-called kissing-loop structure. The 34 structures resulting from NMR structure determination are shown with the kissing-loop residues superimposed at top left in Figure 7.2.3 (RMSD of heavy atoms of bases 10 to 15 is 0.63 ± 0.17 Å for the 34 structures relative to a global average structure) and with the stem residues superimposed at top right (RMSD of heavy atoms of bases 1 to 6
the use of larger force constants for the restraint violation terms in rMD simulations, by inadequate sampling in DG calculations, and the selection of very tight restraint bounds. The structural inaccuracy engendered by restraint bounds made tighter than the experimental accuracy has already been mentioned. While it provides some insight, the atomic RMSD is definitely not a measure of accuracy and is only a modest descriptor of precision (Shriver and Edmondson, 1993). Another limitation of using the overall atomic RMSD to characterize a structure is demonstrated in Figure 7.2.3. Here NMR has been used to determine the structure of an RNA construct corresponding to the genomic dimer
1
3
5
9 7 A
GGCAAUGA -
CCGUUGC A
NMR Determination of Oligonucleotide Structure
C G C G C
G C G C G
16
A
19
21
23
CGUUGCC -
GUAACGG A A
Figure 7.2.3 Structure of an RNA construct of the HIV-1 genome dimer initiation complex. The lower scheme shows the complex of 46 nucleotides composed of two identical 23-nucleotide RNA strands that each form a stem-loop. The two loops interact via the palindromic CG hexamer, creating a helix from the interacting loops—i.e., a kissing-loop structure is formed. The 34 structures resulting from NMR structure determination are shown with the kissing loop residues superimposed at top left and with the stem residues superimposed at the top right. As the dimer initiation complex is symmetric, the other stem region can also be superimposed.
7.2.12 Current Protocols in Nucleic Acid Chemistry
and 18 to 23 is 0.87 ± 0.36 Å for the structures relative to a global average structure). As the dimer initiation complex is symmetric, the other stem region can also be superimposed with identical results. A casual glance at either the right or left ensemble of structures might lead one to conclude that a significant amount of the RNA was either unstructured or not determined very well. In fact, this appearance is due solely to the lower RMSD (1.63 ± 0.47 Å) in the region of the junction (G7, A8, A9, A16, and G17): some torsion angles in this junction region may possess some degree of flexibility or there may not be enough experimental restraints to define this region completely.
R-Factor Calculations Other figures of merit can be used to assess NMR structures, however. For example, one can compare the final structure, or even interim structures, with the experimental NMR data. In x-ray crystallography, an evaluation of the fit of a derived structure with the original electron densities via a residual index (R factor) is requisite for any published structure. But such a comparison of the derived structure with the original data is done only occasionally for NMR structures. However, one can quantitatively compare calculated 2D NOE spectral intensities for any proposed molecular structure—obtained, e.g., with the program CORMA that is embedded within the MARDIGRAS program package—with the experimental intensities. It would be possible to use a residual index analogous to the crystallographic R factors, but that gives equal weighting to all deviations between observed (ao)and calculated (ac) intensities such that stronger peaks (from very short distances, i.e., <2.5 Å) dominate. A sixth-root residual index Rx, however, permits longer interactions, e.g., ∼4 to 5 Å, to have a role in the calculated R factor. c (i)| ∑ |a1/o 6(i) − a1/6
Rx =
i o (i) ∑ a1/6
cule. Such analysis is capable of indicating regions of good (or bad) fit between any model structure and the actual solution structure. In principle, one might wish to generate a structure with the lowest R factor possible. However, prudence is advisable in any efforts to push an R factor to its lowest value. R factors are also of limited value when assessing the accuracy of a refined model. A small restraint set can be easily overfit to a low R value, although the structure is only poorly defined by the data. Other complications arise from the nonrandom nature of errors in experimental intensities and the limited knowledge of molecular motions. Although the exact nature of the molecular motions is not a strong determinant in the calculations of the 2D NOE intensities, motions can still influence the intensities (Keepers and James, 1984). Values for NMR R factors should not be compared with those for x-ray R factors, since the total range of the x-ray R factor (with a maximum value of 0.83) is much more limited than for an NMR R factor (with a maximum value of ∞). A free R factor Rfree has been proposed as an unbiased indicator for assessing x-ray crystal structures and NMR structures (Brünger and Nilges, 1993). In the case of NMR, Rfree measures the fit of a model structure’s NOE intensities to a randomly selected set of experimental NOE intensities that are not used in structure refinement— ∼10% of the total are in the test set; the other 90% are used in structure refinement. This avoids any model bias in calculating the R value. It is to be expected that Rfree will be larger than the usual R factor, since the test data were not used in structure generation. However, discounting noise in the data, if the test data are consistent with the structure determined, Rfree should not be too much larger. Determination of the exact amount that should be considered acceptable awaits more experience with real data. For one DNA decamer duplex, we found it to be 60% larger, and 50% to 100% larger has been reported for the few proteins examined so far.
i
Equation 7.2.7
Longer distance restraints are most important in structure determination, so it would be good to have their input into assessing structure quality. Unlike in x-ray crystallography, it is possible to define a subset of NOE cross-peaks and calculate Rx for some particular aspect of the structure, e.g., interresidue versus intraresidue or one selected region of the mole-
Other Assessment Criteria Consistency of the final structure with scalar coupling–based multidimensional NMR spectra may be assessed as well. For example, the RMSD between experimental and theoretical coupling constants can be calculated as (Jexp − Jtheor)2 Jrms = (1/N)√ Equation 7.2.8
Biophysical Analysis of Nucleic Acids
7.2.13 Current Protocols in Nucleic Acid Chemistry
where the summation is over N, constituting all or any subset, of the coupling constants (Ulyanov et al., 1993). Of course, this assumes that a well-parameterized Karplus relationship has been established between the dihedral angles and three-bond coupling constants (Wijmenga et al., 1993). A residual distance violation is the difference between a particular interproton distance in a structure and the closest of either the upper or lower bound. An evaluation of the structures resulting from any refinement technique should not exhibit substantial distance violations. Other criteria depend on the method of refinement. For structures resulting from rMD or rMC refinement, the restraints violation energies (VNOE and VJ coupling in Equation 7.2.6) should be low, and the total of the other terms should not be much greater than in the absence of any experimental restraints. Any structure generated should, of course, be consistent with all other available experimental data on the oligomer in solution. This includes chemical as well as physical data.
ANALYSIS OF NUCLEIC ACID STRUCTURE A detailed characterization of the resulting NMR structure should be performed, entailing determination of backbone torsion angles, sugar puckers, glycosidic torsion angles, and a series of helical (or base-orientation if not in a helix) parameters. The program CURVES 5.1 calculates structural parameters from the Cartesian coordinates of the atoms in a nucleic acid (Lavery and Sklenar, 1989, 1996). It calculates the structural disposition of individual bases—e.g., inclination, tip, x displacement, and y displacement—and uses that information to define a local helical axis. By building up from these local helical segments, the global helix axis Table 7.2.1
B-DNAb A-DNAb NMRc
is constructed. That global axis may have kinks or curvature that can be examined and measured. Further information about use of CURVES can be found on the web page http://plumber/ csb.yale.edu/userguides/datamanip/curves/doc. html. Dials and Windows is a program package convenient for computing and graphing the structural parameters output by rMD (Ravishanker et al., 1989); it works ideally in concert with CURVES. Some information can be found on the Dials and Windows web page at http://plumber.csb.yale.edu/userguides/ graphics/ dandw/dandw_descr. Dials and Windows is one part of the Molecular Dynamics Toolchest developed at Wesleyan University. An important reason for calculating the structural parameters is to assess the validity of structures generated by NMR. Unusual backbone torsion angles or sugar puckers could very well highlight inconsistencies among the restraints or even mistakes in the data that might have gone undetected by the assessment tools already mentioned. For example, helical parameters for duplex DNA solution structures should probably not deviate too much from canonical B-DNA or related crystal structures unless this is clearly justified by the original data. The same holds for double-helical regions of RNA, which should be very much like canonical A-form RNA. Table 7.2.1 lists some structural parameter values found for double-helical DNA; RNA should have the values of ADNA. Extreme values of structural parameters or unusual conformations should be examined very carefully. It is necessary to scrutinize the restraints and the rMD protocol to be certain that the source of the unusual structural feature lies in reliable experimental data.
Average Structural Parameters for DNA from Solution and Solid-State Dataa
Twistb (°)
Tilt (°)
Roll (°)
Shift (Å)
Slide (Å)
Rise (Å)
36.1 4.2 30.8 4.8 35.3 4.2
0.0 3.6 0.0 3.3 −0.1 2.5
−0.2 5.6 7.9 5.6 4.6 7.0
0.00 0.55 0.00 0.52 0.00 0.28
0.21 0.75 −1.57 0.38 −0.34 0.46
3.35 0.24 3.32 0.31 3.15 0.21
Propeller twist (°) −13.8 6.6 −9.8 5.6 −10.8 7.8
aThe first line of each entry gives the mean value, and the second line (in italics) gives the standard deviation in each
NMR Determination of Oligonucleotide Structure
individual set of data. Data are from a larger compilation that also lists sequence-dependence of the parameters (Ulyanov and James, 1995). The local helical parameters reported here conform to the Cambridge convention (Dickerson et al., 1989). bFrom high-resolution crystal structures. cFrom nine high-resolution NMR structures.
7.2.14 Current Protocols in Nucleic Acid Chemistry
ACKNOWLEDGMENTS I wish to express grateful appreciation to my co-workers who have carried out research that has led to insights presented in this chapter; they are authors on the references cited. I explicitly thank Dr. Anwer Mujeeb, who constructed the prototype of Figure 7.2.1. The research described in this article from our lab was supported by National Institutes of Health grants GM39247 and RR01081. Some of the computations were carried out at the Pittsburgh Supercomputing Center (supported by grant no. 1 P41 RR06009 from the NIH National Center for Research Resources).
LITERATURE CITED Abragam, A. 1961. Principles of Nuclear Magnetism. Oxford University Press, Oxford. Adams, B. and Lerner, L. 1992. A simple one-dimensional method for measuring proton exchange rates in water. J. Magn. Reson. 96:604-607. Altona, C. and Sundaralingam, M. 1972. Conformational analysis of the sugar ring in nucleosides and nucleotides. A new description using the concept of pseudorotation. J. Am. Chem. Soc. 94:8205-8212. Boelens, R., Koning, T.M.G., van der Marel, G.A., van Boom, J.H., and Kaptein, R. 1989. Iterative procedure for structure determination from proton-proton NOEs using a full relaxation matrix approach. Application to a DNA octamer. J. Magn. Reson. 82:290-308. Borgias, B.A. and James, T.L. 1989. Two-dimensional nuclear Overhauser effect: Complete relaxation matrix analysis. Methods Enzymol. 176:169-183.. Borgias, B.A. and James, T.L. 1990. MARDIGRAS—Procedure for matrix analysis of relaxation for discerning geometry of an aqueous structure. J. Magn. Reson. 87:475-487. Brünger, A. and Nilges, M. 1993. Computational challenges for macromolecular structure determination by X-ray crystallography and solution NMR-spectroscopy. Q. Rev. Biophys. 26:49-125. Cavanagh, J., Fairbrother, W.J., Palmer, A.G., III, and Skelton, N.J. 1996. Protein NMR Spectroscopy: Principles and Practice. Academic Press, San Diego. Celda, B., Widmer, H., Leupin, W., Chazin, W.J., Denny, W.A., and Wüfthrich, K. 1989. Conformational studies of d-(AAAAATTTTT)2 using constraints from nuclear Overhauser effects and from quantitative analysis of the cross-peak fine structures in two-dimensional 1H nuclear magnetic resonance spectra. Biochemistry 28:14621470. Conte, M.R., Bauer, C.J., and Lane, A.N. 1996. Determination of sugar conformations by NMR in larger DNA duplexes using both dipolar and scalar data: Application to d(CATGTGACGTCACATG)2. J. Biomol. NMR 7:190-206.
Derome, A. 1987. Modern NMR Techniques for Chemistry Research. Pergamon Press, Oxford. Dickerson, R.E., Bansal, M., Calladine, C.R., Diekmann, S., Hunter, W.N., Kennard, O., Lavery, R., Nelson, H.J.C., Olson, W.K., Saenger, W., Shakked, Z., Sklenar, H., Soumpasis, D.M., von Kitzing, E., Wang, A. H.-J., and Zhurkin, V.B. 1989. Definitions and nomenclature of nucleic acid structure parameters. EMBO J. 8:1-4. Ernst, R.R., Bodenhausen, G., and Wokaun, A. 1987. Principles of Nuclear Magnetic Resonance in One and Two Dimensions. Clarendon Press, Oxford. Gonzàlez, C., Stec, W., Kobylanska, A., Hogrefe, R., Reynolds, M., and James, T.L. 1994. Structural study of a DNA-RNA hybrid duplex with a chiral phosphorothioate moiety by NMR: Extraction of distance and torsion angle constraints and imino proton exchange rates. Biochemistry 33:11062-11072. Harbison, G.S. 1993. Interference between J-couplings and cross-relaxation in solution NMR spectroscopy: Consequences for macromolecular structure determination. J. Am. Chem. Soc. 115:3026-3027. James, T.L. 1994. Assessment of the quality of derived macromolecular structures. Methods Enzymol. 239:416-439. James, T.L. (ed.) 1995. Nuclear Magnetic Resonance and Nucleic Acids. Methods in Enzymology, Vol. 261. Academic Press, New York. Keepers, J.W. and James, T.L. 1984. A theoretical study of distance determinations from NMR. Two-dimensional nuclear Overhauser effect spectra. J. Magn. Reson. 57:404-426. Landy, S.B. and Rao, B.D.N. 1989. Dynamical NOE in multiple-spin systems undergoing chemical exchange. J. Magn. Reson. 81:371-377. Lane, A. N. 1995. Determination of fast dynamics of nucleic acids by NMR. Methods Enzymol. 261:413-35. Lavery, R. and Sklenar, H. 1989. Defining the structure of irregular nucleic acids: Conventions and principles. J. Biomol. Struct. Dyn. 6:655-667. Lavery, R. and Sklenar, H. 1996. CURVES 5.1. Helical Analysis of Irregular Nucleic Acids. Laboratoire de Biochimie Theoretique, Centre National de la Recherche Scientifique, Paris. Leroy, J.L., Gehring, K., Kettani, A., and Guéron, M. 1993. Acid multimers of oligodeoxycytidine strands: Stoichiometry, base-pair characterization, and proton exchange properties. Biochemistry 32:6019-6031. Liu, H., Thomas, P.D., and James, T.L. 1992. Averaging of cross-relaxation rates and distances for methyl, methylene and aromatic ring protons due to motion or overlap: Extraction of accurate distances iteratively via relaxation matrix analysis of 2D NOE spectra. J. Magn. Reson. 98:163-175.
Biophysical Analysis of Nucleic Acids
7.2.15 Current Protocols in Nucleic Acid Chemistry
Liu, H., Kumar, A., Weisz, K., Schmitz, U., Bishop, K.D., and James, T.L. 1993. Extracting accurate distances and bounds from 2D NOE exchangeable proton peaks. J. Am. Chem. Soc. 115:15901591. Liu, H., Spielmann, H.P., Ulyanov, N.B., Wemmer, D.E., and James, T.L. 1995. Interproton distance bounds from 2D-NOE intensities: Effect of experimental noise and peak integration errors. J. Biomol. NMR 6:390-402. Liu, H., Tonelli, M., and James, T.L. 1996. Correcting NOESY cross-peak intensities for partial relaxation effects enabling accurate distance determination. J. Magn. Reson. B 111:85-89. Macura, S. and Ernst, R.R. 1980. Elucidation of cross relaxation in liquids by 2D NMR spectroscopy. Mol. Phys. 41:95-117. Marion, D. and Bax, A. 1988. P.COSY, a sensitive alternative for double-quantum-filtered COSY. J. Magn. Reson. 80:528-533. Mujeeb, A., Clever, J.L., Billeci, T.M., James, T.L., and Parslow, T.G. 1998. Structure of the dimer initiation complex of the HIV-1 genomic RNA. Nature Struct. Biol. 5:432-436. Piantini, U., Sørensen, O.W., and Ernst, R.R. 1982. Multiple quantum filters for elucidating NMR coupling networks. J. Am. Chem. Soc. 104:68006801. Post, C.B., Meadows, R.P., and Gorenstein, D.G. 1990. On the evaluation of interproton distances for three-dimensional structure determination by NMR using a relaxation rate matrix analysis. J. Am. Chem. Soc. 112:6796-6803. Ravishanker, G., Swaminathan, S., Beveridge, D.L., Lavery, R., and Sklenar, H. 1989. Conformational and helicoidal analysis of 30 ps of molecular dynamics on the d(CGCGAATTCGCG) double helix: “Curves,” dials and windows. J. Biomol. Struct. Dyn. 6:669-699. Rinkel, L.J. and Altona, C. 1987. Conformational analysis of the deoxyribofuranose ring in DNA by means of sums of proton-proton coupling constants: A graphical analysis. J. Biomol. Struct. Dyn. 4:621-649. Schmitz, U. and James, T.L. 1995. How to generate accurate solution structures of double-helical nucleic acid fragments using nuclear magnetic resonance and restrained molecular dynamics. Methods Enzymol. 261:3-44. Schmitz, U., Zon, G., and James, T.L. 1990. Deoxyribose conformation. In [d(GTATATAC)]2: Evaluation of sugar pucker by simulation of double-quantum-filtered COSY cross-peaks. Biochemistry 29:2357-2368. Shriver, J. and Edmondson, S. 1993. Defining the precision with which a protein structure is determined by NMR. Application to motilin. Biochemistry 32:1610-1617.
Thomas, P.D., Basus, V.J., and James, T.L. 1991. Protein solution structure determination using distances from 2D NOE experiments: Effect of approximations on the accuracy of derived structures. Proc. Natl. Acad. Sci. U.S.A. 88:12371241. Tonelli, M. and James, T.L. 1998. Insights into the dynamic nature of DNA duplex structure via analysis of nuclear Overhauser effect intensities. Biochemistry 37:11478-11487. Tonelli, M., Ragg, E., Bianucci, A.M., Lesiak, K., and James, T.L. 1998. NMR structure of d(GCATATGATAG).d(CTATCATATGC): A consensus sequence for promoters recognized by σK RNA polymerase. Biochemistry 37:1174511761. Ulyanov, N.B. and James, T.L. 1995. Statistical analysis of DNA duplex structural features. In Nuclear Magnetic Resonance and Nucleic Acids, Vol. 261 (T.L. James, ed.) pp. 3-44. Academic Press, New York. Ulyanov, N.B., Schmitz, U., and James, T.L. 1993. Metropolis Monte Carlo calculations of DNA structure using internal coordinates and NMR distance restraints: An alternative method for generating high-resolution solution structure. J. Biomol. NMR 3:547-568. Ulyanov, N.B., Schmitz, U., Kumar, A., and James, T.L. 1995. Probability assessment of conformational ensembles: Sugar repuckering in a DNA duplex in solution. Biophys. J. 68:13-24. van de Ven, F.J.M. and Hilbers, C.W. 1988. Resonance assignments of non-exchangeable protons in B type DNA oligomers, an overview. Nucl. Acids Res. 16:5713-5726. Weisz, K., Shafer, R.H., Egan, W., and James, T.L. 1994. Solution structure of the octamer motif in immunoglobulin genes via restrained molecular dynamics calculations. Biochemistry 33:354366. Widmer, H. and Wüthrich, K. 1987. Simulated twodimensional NMR cross-peak fine structures for 1 H spin systems in polypeptides and polydeoxynucleotides. J. Magn. Reson. 74:316-336. Wijmenga, S.S., Mooren, M.M.W., and Hilbers, C.W. 1993. NMR of nucleic acids; from spectrum to structure. In NMR in Macromolecules (G.C. Roberts, ed.) pp. 217-288. IRL Press, Oxford. Zhu, L., Reid, B.R., and Drobny, G.P. 1995. Errors in measuring and interpreting values of coupling constants J from PE.COSY experiments. J. Magn. Reson. A 115:206-212.
Contributed by Thomas L. James University of California San Francisco, California
NMR Determination of Oligonucleotide Structure
7.2.16 Current Protocols in Nucleic Acid Chemistry
Optical Methods
UNIT 7.3
Nucleic acids participate in a variety of processes in which nucleic acid complexes are formed. The stabilization of the DNA duplex, its replication and recombination, its translation into RNA, and the folding of RNA structures all depend on specific interactions between nucleic acids. To understand these processes and to take advantage of the exquisite recognition capabilities of nucleic acids requires an understanding of the thermodynamics of nucleic acid complex formation. Methods for extraction of thermodynamic parameters from equilibrium optical melting curves are described in this unit. Additional procedures are presented for making important preliminary determinations of molar extinction coefficients and the number of oligonucleotides that combine to form a complex. Basic Protocol 1 presents a simple method for accurately determining the extinction coefficients of oligonucleotides and polynucleotides, while Basic Protocol 2 presents a method for simultaneously determining the extinction coefficient for a nucleic acid complex and the number of strands in the complex. Basic Protocol 3 describes a method for determining equilibrium melting curves of nucleic acids by monitoring the temperature dependence of UV absorbance. Methods for analysis of the equilibrium melting curves to extract values for ∆H° and ∆G° are described in Support Protocols 1 to 5. DETERMINATION OF OLIGONUCLEOTIDE MOLAR EXTINCTION COEFFICIENTS
BASIC PROTOCOL 1
The molar extinction coefficient (ε260) of an oligonucleotide is determined by means of a colorimetric phosphate assay (Snell and Snell, 1949; Griswald et al., 1951), whereby phosphate is released from the oligonucleotide enzymatically. A standard curve of color intensity (absorbance) produced versus phosphate concentration is used to quantify the released phosphate. Given that information, the extinction coefficient is calculated easily. Poly(rU) is used as a positive control to monitor completeness of the enzymatic digestion. To avoid contamination that would compromise the molar extinction coefficient determination, phosphate should be scrupulously avoided in every step of the preparation of the oligonucleotide. In addition, the oligonucleotide should be desalted subsequent to purification and prior to determination of the extinction coefficient. Materials 10 mM cacodylate buffer solution (see recipe) 1 mg/mL nuclease P1 solution (see recipe) 100 U/mL alkaline phosphatase solution (see recipe) Oligonucleotide to be analyzed 53 µM poly(rU) (Amersham Pharmacia Biotech) in 10 mM cacodylate buffer solution Standard phosphate solution (see recipe) ANS solution (see recipe) Molybdate solution (see recipe) 1-dram screw-capped glass vials 95°C water bath Single- or double-beam UV/visible spectrophotometer Matched quartz semimicro 10-mm spectrophotometer cuvettes Contributed by G. Eric Plum Current Protocols in Nucleic Acid Chemistry (2000) 7.3.1-7.3.17 Copyright © 2000 by John Wiley & Sons, Inc.
Biophysical Analysis of Nucleic Acids
7.3.1
NOTE: In all steps, samples should be prepared in 1-dram screw-capped glass vials. Plastic microcentrifuge tubes should not be used. 1. Prepare 1 mL oligonucleotide solution with an A260 of ∼0.4 to 0.5 in 10 mM cacodylate buffer solution and place in a quartz semimicro 10-mm cuvette. Fill another cuvette with 10 mM cacodylate buffer solution. 2a. For a single-beam instrument: Place the buffer-only cuvette in the instrument and zero the instrument at 260 nm. Replace cuvette with the oligonucleotide-containing cuvette and record absorbance at 260 nm. 2b. For a double-beam instrument: Place the buffer-only cuvette in the reference beam and the oligonucleotide-containing cuvette in the sample beam. Record the absorbance at 260 nm. 3. Repeat this procedure using a 1-mL sample of poly(rU) solution in place of the oligonucleotide. The selection of 260 nm is by convention, as it is at or near the absorbance maximum of most oligonucleotides. If the spectrophotometer has wavelength scanning capabilities, it is desirable to scan between 200 and 350 nm. If the oligonucleotide displays significant self-structure, as seen in a temperature-dependent absorbance, the absorbance measurement should be made at elevated temperature.
4. To each oligonucleotide and poly(rU) sample and to 1 mL of cacodylate buffer, add 3 µL of 1 mg/mL nuclease P1 solution and 10 µL of 100 U/mL alkaline phosphatase solution. Allow digestion to proceed overnight at room temperature. Nuclease P1 cleaves the phosphodiester backbone of the oligonucleotide, and alkaline phosphatase cleaves the phosphate group from the released nucleotides.
5. In separate tubes, prepare the following samples, then dilute each to 800 µL with cacodylate buffer solution: a. Phosphate standard curve: Prepare thirteen samples containing 0 through 120 µL standard phosphate solution at 10-µL intervals. b. Test samples: Prepare four parallel samples containing 50, 100, 150, and 200 µL enzyme-digested oligonucleotide solution. c. Positive control: Prepare four samples containing 50, 100, 150, and 200 µL enzyme-digested poly(rU) solution. d. Negative control: Prepare four samples containing 50, 100, 150, and 200 µL enzyme-containing negative control sample. The standard curve and controls (positive and negative) must be run with each assay as a quality control measure. If desired, additional oligonucleotides may be analyzed in parallel using the same standard curve and controls, provided that the same solutions are used and all samples are analyzed simultaneously.
6. Add 100 µL ANS solution and 100 µL molybdate solution to each sample. Heat for 10 min in a 95°C bath, then cool to 25°C. Upon heating, formation of a phosphomolybdate complex results in a blue tint to the solution that is proportional to the concentration of phosphate.
Optical Methods
7. Set the spectrophotometer to read absorbance at 820 nm. Fill a cuvette with cacodylate buffer solution. Measure and record A820 for each sample, using the cacodylate buffer solution for a reference as described in steps 2a and 2b. Thoroughly rinse and dry the cuvette and change pipet tips between samples.
7.3.2 Current Protocols in Nucleic Acid Chemistry
Figure 7.3.1
Standard curve for phosphate analysis.
8. Prepare a standard curve by plotting A820 versus phosphate concentration, [PO4], for the thirteen samples of standard phosphate solution. Confirm that A820 for the blank samples falls on the y intercept of a line through the data points. The amount of phosphate in each of the samples is simply the initial standard solution concentration multiplied by the volume of standard solution used: PO4 (mol) = [PO4]std (M) × Vstd (L). Because [PO4]std ≈ 100 mM and the volumes range from 10 to 120 mL, the amount of PO4 will be in the 1 to 12 nmol range. See Figure 7.3.1 for an example.
9. Determine the amount (in mol) of phosphate present in each oligonucleotide and poly(rU) sample by finding the measured A820 on the standard curve and projecting a line onto the abscissa. 10. Calculate the molar extinction coefficient of the oligonucleotide using the equation ε260 =
A260 b mol PO4 mol oligo V mol PO4
cm−1M−1
Equation 7.3.1
where b is the pathlength (1 cm), mol PO4 is from step 9, A260 is from step 2a or 2b, and the mole unit of the final value refers to the oligonucleotide. Obtain the final value of the denominator from the length of the oligonucleotide, defined by mol oligo = (no. bases − 1)−1 mol PO4 Equation 7.3.2
assuming the oligonucleotide is not end phosphorylated. 11. Confirm that the correct values for the extinction coefficient for poly(rU) positive controls are obtained. Because the polymer is of indeterminate length, a different mole unit must be used for the extinction coefficient, specifically, moles nucleotide/liter. The equation for ε260 reduces to the equation below because there is one phosphate per nucleotide. There are a number of slightly varying values for the poly(rU) extinction coefficient; a value of 9450 cm−1M−1 can be considered reliable (J. Völker, pers. comm.).
Biophysical Analysis of Nucleic Acids
7.3.3 Current Protocols in Nucleic Acid Chemistry
ε260 =
A260
mol PO4 b V
cm−1M−1
Equation 7.3.3
BASIC PROTOCOL 2
DETERMINATION OF MOLECULARITY AND EXTINCTION COEFFICIENTS OF OLIGONUCLEOTIDE COMPLEXES The same differences in UV absorbance between isolated oligonucleotides and the complexes they form that are exploited to produce melting profiles are used to determine the number of oligonucleotides in the complex. The method presented is based on careful preparation of mixing curves and measurement of absorbance as a function of the relative concentration of oligonucelotides. It is known as the method of continuous fractions (Felsenfeld et al., 1957). The resultant data are presented as a so-called Job plot. Multiple wavelengths are used because some wavelengths cannot detect formation of some complexes. Materials Two oligonucleotides, A and B, which are expected to form a complex Buffer Single- or double-beam UV/visible spectrophotometer Quartz semimicro spectrophotometer cuvette (1-cm pathlength) Additional reagents and equipment for determining molar extinction coefficients (see Basic Protocol 1) 1. Determine extinction coefficients for oligonucleotides A and B using Basic Protocol 1. 2. Prior to beginning the experiment, determine the minimum sample volume required for an accurate absorbance measurement in the particular spectrophotometer and cuvettes being used. The mounting of the cell holder in the spectrophotometer relative to the beam varies with the manufacturer and model. A volume of 600 mL is typically sufficient so that the meniscus does not impinge on the beam, but this should be confirmed. The measurement is easily done and need not be repeated, assuming the equipment used does not change. To determine the appropriate volume, prepare a solution with absorbance between 0.5 to 1.0 at 260 nm. It may be convenient to use a nucleic acid solution, although any solution with absorbance at the desired wavelength may be used. Begin with 500 mL of solution in the cell and measure the absorbance. Add 25-mL aliquots of the solution, each time noting the absorbance. The volume at which the absorbance becomes constant is the minimum usable volume. It is advisable to begin the experiment with a slightly larger volume.
3. Prepare 1.5 mL each of two oligonucleotide solutions, one with oligonucleotide A and another with oligonucleotide B, at identical concentrations. Use a concentration that gives an A260 of ~0.5 in a 1-cm-pathlength cuvette. A good choice of concentration is c = 0.5/εavg, where εavg is the average of the extinction coefficients for the two oligonucleotides. Because the solution conditions (salt concentration, pH, and temperature) determine the relative stabilities of complexes of various stoichiometries, the oligonucleotide solutions should be prepared in the buffer in which further experiments will be performed. Optical Methods
4. Fill the cuvette with buffer and record the absorbance at 220, 240, 260, 280, and 300 nm.
7.3.4 Current Protocols in Nucleic Acid Chemistry
5. Place 600 µL (or the experimentally determined minimum volume) of oligonucleotide A solution in the cuvette and place it in the spectrophotometer. Measure the absorbance at 220, 240, 260, 280, and 300 nm. 6. Add 100 mL oligonucleotide B solution to the cuvette. Mix thoroughly. Because semimicro 1-cm-pathlength cuvettes are used, below-cell magnetic stirring does not work well. The best alternative is an immersible mixing device, although care must be taken that the immersed stirrer does not scratch the cuvette and does not impinge on the light beam when absorbance measurements are made. Alternatively, mixing can be accomplished by repeated drawing and dispensing of the solution with a transfer pipet or by capping and sealing of the cell followed by inversion. In either case, great care must be taken to avoid a loss of solution. If standard 1-cm-pathlength cuvettes and a spectrophotometer equipped with a built-in magnetic stirrer in the cell holder are used, adequate stirring should be possible with a 7 × 2–mm Teflon-coated magnetic stir bar. The amount of each solution required increases by a factor of ∼3 when standard 1-cm cuvettes are used.
7. Measure the absorbance at 220, 240, 260, 280, and 300 nm. Repeat the absorbance measurements at 5-min intervals until constant values are observed. If significant self-structure or self-association is observed, as detected by temperaturedependent transitions in the absorbance of either oligonucleotide, it is necessary to include an annealing step. The cuvette is heated above the melting temperature of the complex and cooled slowly. If this step is included, the cells must be securely capped to minimize evaporation.
8. Repeat steps 6 and 7 five times using 100-µL aliquots of oligonucleotide solution B. 9. Clean and dry the cuvette. Repeat steps 4 through 8, beginning with 600 µL oligonucleotide solution B and adding 100-µL aliquots of oligonucleotide solution A. 10. Prepare a plot of A260 versus the mole fraction of oligonucleotide B (XB). Because oligonucleotide A and B solutions are at identical concentrations, XB = [VB/(VA + VB)], where VA and VB are the total volumes of oligonucleotide solutions A and B in the
Figure 7.3.2 Example of an oligonucleotide mixing curve. The filled circles represent additions of oligonucleotide B to oligonucleotide A. Open circles represent additions of oligonucleotide A to oligonucleotide B.
Biophysical Analysis of Nucleic Acids
7.3.5 Current Protocols in Nucleic Acid Chemistry
cuvette. An example is shown in Figure 7.3.2. The inflection point at XB = 0.5 in the example indicates the formation of an n:n complex. In the absence of contrary information, it is reasonable to assume that a 1:1 complex has formed. Because the concentration, c, of the complex is known from the concentrations of the oligonucleotide solutions, the extinction coefficient of the complex is easily determined from ε260 = (A260inf/bc), where A260inf is the absorbance at the inflection point and b is the pathlength. Failure of the two titration curves to meet indicates a probable error in one of the oligonucleotide extinction coefficients or in solution preparation. Excessive curvature may indicate failure to reach equilibrium or a very low association constant.
11. Repeat step 10 for data collected at remaining wavelengths to confirm that no other stoichiometries are observed. BASIC PROTOCOL 3
PREPARATION OF EQUILIBRIUM MELTING CURVES The absorbance of nucleic acids differs between the less structured isolated single strands observed at high temperature and the folded or complexed strands observed at low temperature. The thermal melting profiles obtained from a temperature-dependent absorbance experiment can be used to extract thermodynamic parameters. The concentration dependence of the melting temperature is particularly useful in determining the thermodynamic parameters (see Support Protocol 1) for complexes of molecularity >1; therefore, the experiment is described in terms of a concentrationdependent study. Materials Oligonucleotides in buffer solution (single folded chains or multimolecular complexes) Double-beam UV/visible spectrophotometer with temperature-controlled (stepping and/or scanning) cell holder Stoppered quartz spectrophotometer cuvettes (0.1-, 0.2-, 0.5-, and 1.0-cm pathlengths) with metal spacers (shorter pathlength cuvettes are useful, if available) Additional reagents and equipment for determining molar extinction coefficients (see Basic Protocol 1) and determining molecularity and extinction coefficients of complexes (see Basic Protocol 2) NOTE: Buffers with large heats of ionization, which includes most popular biochemical buffers, should be avoided, as the pH of solutions prepared with these buffers is temperature dependent. Tris solutions are particularly susceptible to this effect. Phosphate, cacodylate, and PIPES are acceptable choices. 1. Determine extinction coefficients for the individual oligonucleotides (see Basic Protocol 1). 2. If a multimolecular complex is to be examined, determine the molecularity of the complex and its extinction coefficient (see Basic Protocol 2). 3a. For a single melting experiment: Fill a stoppered, 1-cm-pathlength semimicro cuvette with 1300 µL of an oligonucleotide solution that gives an A260 of ~0.4 to 0.6. 3b. For a concentration-dependence study: Prepare eight to ten solutions of the oligonucleotide complex to cover as wide a range of concentrations as possible (≥200-fold). Calculate the concentration from c = (A260/ε260b).
Optical Methods
The oligonucleotide complex concentrations should be designed to span the A260 range of 0.1 in a 1.0-cm cuvette to 2.0 in a 0.1-cm cuvette. Depending on the quality of the spectrophotometer, this concentration range may be expanded.
7.3.6 Current Protocols in Nucleic Acid Chemistry
Table 7.3.1
ln(εc) −4.61 −4.08 −3.55 −3.02 −2.49 −1.96 −1.43 −0.90 −0.37 0.16 0.69
Selection of Concentration and Cuvette Pathlengtha
εc 0.01 0.017 0.029 0.049 0.083 0.14 0.24 0.41 0.69 1.2 2.0
b = 1 mm 0.01 0.017 0.029 0.049 0.083 0.14 0.24 0.41 0.69 1.2 2.0
A = εcbb b = 2 mm b = 5 mm 0.02 0.05 0.034 0.085 0.057 0.14 0.10 0.24 0.17 0.41 0.28 0.7 0.48 1.2 0.81 2 3.5 1.4 2.3 5.9 4.0 10.0
b = 10 mm 0.1 0.17 0.29 0.49 0.83 1.4 2.4 4.1 6.9 12.0 20.0
aTo use the table, select from each row a target absorbance (A) and a useable pathlength (b from colums 3, 4, 5, or 6). Use the known ε value to calculate c from the εc column. bBoldface indicates absorbance values that are usable with the pathlength b.
Because of the logarithmic dependence of the melting temperature on oligonucleotide complex concentration, it is desirable to distribute the concentrations evenly on a logarithmic scale. Selection of concentrations and cuvette pathlengths to produce solutions giving appropriate absorbance values is nontrivial. Table 7.3.1 provides a guide for solution preparation and pathlength selection. The desired 200-fold concentration range (from εc = 0.01 to 2.0) is covered with even steps on a logarithmic scale. Solutions should be prepared in sufficient volume to fill the selected cuvette. Table 7.3.2 provides the volume necessary to fill commonly available cuvettes.
4. Fill a cuvette (or cuvettes, if spectrophotometer is equipped with a multiple-cell holder) and securely seal with a stopper. Place cuvette in spectrophotometer cell holder. Use metal spacers to ensure proper alignment of the cuvette and good thermal contact with the thermostatically controlled cell holder. If stoppered cuvettes are not available, a small amount of silicon oil may be layered on top of the sample. It is good practice to heat the sample above Tm and slowly cool it to assure proper annealing of the complex. This annealing step need not be performed in the spectrophotometer.
5. Equilibrate the instrument at the starting temperature, usually 1° to 5°C. 6a. For scanning temperature: Increase the temperature at a rate of 0.5°C/min or less. Collect absorbance data at intervals such that four or more absorbance measurements are recorded for each sample for each degree increase in temperature. Collect data
Table 7.3.2 Approximate Filling Volumes for Common Cuvettes
Pathlength (cm) 1.0a 0.5a 0.2 0.1 aSemimicro cuvette.
Total volume (µL) 1300 600 600 300
Biophysical Analysis of Nucleic Acids
7.3.7 Current Protocols in Nucleic Acid Chemistry
until the temperature has increased well beyond the apparent melting temperature of the complex, usually to 95° to 100°C. Cool and remove the cuvette. 6b. For stepping temperature (preferred): Collect data at 0.25°C or smaller temperature intervals. Collect data until the temperature has increased well beyond the apparent melting temperature of the complex, usually to 95° to 100°C. Cool and remove the cuvette. 7. If conducting a concentration dependent study, repeat steps 4 to 6 until all samples have been heated in the spectrophotometer. 8. Analyze data (see Support Protocol 1 for concentration-dependent data or one of Support Protocols 2 through 4 and Support Protocol 5 for a single-melting study). ANALYSIS OF EQUILIBRIUM MELTING CURVES A number of methods for analyzing single or multiple equilibrium melting curves to extract thermodynamic parameters are described below. All of the calculations described are easily implemented in a spreadsheet. All of the thermodynamic parameters determined using Support Protocols 1 to 5 describe the association equilibrium. The preferred method of analysis for equilibrium melting curves is described in Support Protocol 1, which allows determination of the enthalpy change (∆H°), the entropy change (∆S°), and the free energy change (∆G°). If concentration-dependent curves are not available or if the process is monomolecular (hairpin or other folded single-chain structure) or pseudomonomolecular (polynucleotide), Support Protocols 2 through 4 may be used to extract ∆H° and Support Protocol 5 may be used to determine ∆G°. For pseudomonomolecular processes, which formally are bimolecular but show no concentration dependence, a value of 1 can be used for the molecularity in the following equations. In all of the equations that follow, the concentration CT refers to total oligonucleotide concentration. n
CT =
∑ [Ai] i=1
Equation 7.3.4
All methods of analysis described below depend on the experimental setup to ensure that [A1] = … [Ai] … = [An], thus [Ai] =
CT n
Equation 7.3.5
Due to statistical effects, the equations for the association constant differ for complexes comprised of non-self-complementary strands and for complexes comprised of self-complementary strands. When appropriate, two forms of the equations are presented. Although it is convenient to present data as a function of temperature in degrees Celsius, all calculations of thermodynamic parameters must be computed using temperature in Kelvin units.
Optical Methods
7.3.8 Current Protocols in Nucleic Acid Chemistry
Figure 7.3.3 Simulated UV-monitored oligonucleotide melting curve. The dashed lines represent the upper and lower baselines that are used to calculate the α(T) versus T curve.
CALCULATION OF ∆H° FROM CONCENTRATION-DEPENDENT MELTING CURVES
SUPPORT PROTOCOL 1
In this protocol, thermodynamic parameters are determined for short oligonucleotide complexes (i.e., duplexes with <12 bp) from a plot of 1/Tm versus lnCT as described by Marky and Breslauer (1987). 1. Define upper and lower absorbance baselines, AU(T) and AL(T), by fitting the linear portions of the absorbance versus temperature curve, A(T) versus T. Figure 7.3.3 shows an example of a melting curve and the fitted baselines.
2. Define a parameter α(T) to represent the relative fraction of the complex that remains in the initial state at temperature T. Plot α(T) versus T using the slopes (m) and intercepts (b) from the fitted baselines as follows α(T ) =
AU(T ) − A(T ) mUT + bU − A(T ) = AU(T ) − AL(T ) (mU − mL)T + bU − bL Equation 7.3.6
where AU(T) = mUT + bU and AL(T) = mLT + bL.
Figure 7.3.4 The α versus T curve. This curve shows the fraction of the initial complex as a function of temperature. Determination of Tm from the curve is shown.
Biophysical Analysis of Nucleic Acids
7.3.9 Current Protocols in Nucleic Acid Chemistry
3. Determine Tm, which is defined as the temperature at which α = 0.5. Figure 7.3.4 shows an a versus T curve and the determination of Tm. The melting temperature Tm is of limited value in comparing the stability of two duplexes or other complexes. Unfortunately, this is common practice. The Tm value depends on nucleic acid concentration for duplexes and higher molecularity complexes. Also, Tm comparisons assume implicitly that ∆Ho is identical for both processes. The solution conditions also influence Tm significantly. Because of the change in linear charge density associated with the transition from complex to single strands, the Tm depends on salt concentration. Dependence on pH is observed for C+GC-containing triple helices and some lesion-containing duplexes. All complexes are affected by extremes of pH (<5 or >9). Before any useful Tm comparison can be made, the nucleic acid concentrations and solution conditions must be identical.
4. Plot 1/Tm versus lnCT and determine ∆H° using the measured slope and the slope from the following equation [∆So − (n − 1)R ln2n] 1 (n − 1)R = lnCT + o Tm ∆H ∆Ho Equation 7.3.7
for non-self-complementary oligonucleotide complexes or [∆So − (n − 1)R ln2n + R lnn] 1 (n − 1)R = ln + C T Tm ∆Ho ∆Ho Equation 7.3.8
for self-complementary oligonucleotide complexes. In both cases, n is molecularity and R is the gas constant. Note that in both cases, 1/Tm versus lnCT is linear with a slope of (n − 1)R/∆H°. Thus, ∆H° is readily determined from the measured slope. Further note that when n = 1 the slope = 0. This is consistent with the observed concentration independence of monomolecular processes, such as hairpin melting.
5. Determine values of ∆S° from the intercept of the appropriate equation shown above. Determine values of ∆G° using ∆G° = ∆H° – T∆S°. Use caution in interpreting these values. The ∆S° and ∆G° values depend directly on the value of ∆H°. This coupling of the ∆S° and ∆G° values to ∆H° may result in apparent entropy-enthalpy compensation when none exists. For intermediate-length complexes, a concentration-dependent Tm is still observed. This dependence is reduced relative to that expected for complexes with the same ∆H° in a two-state equilibrium. Therefore, ∆H° is overestimated by the above equations. The length where this effect is observed depends on the molecularity, on the sequence, and possibly on the solution conditions. For DNA duplexes, the length is roughly between 12 and ~50 base pairs, where pseudomonomolecular behavior begins. SUPPORT PROTOCOL 2
CALCULATION OF ∆H° FROM =(T) VERSUS T PLOTS This method is from Marky and Breslauer (1987). 1. Calculate values for α(T) as described above (see Support Protocol 1, steps 1 and 2). 2. Plot α(T) as a function of 1/T as in Figure 7.3.5. 3. Examine the melting curves to get an indication of the enthalpy change associated with the transition.
Optical Methods
7.3.10 Current Protocols in Nucleic Acid Chemistry
Figure 7.3.5 Determination of ∆H° from a plot of α versus 1/T. The slope of the line shown is used to calculate ∆H°.
A steep slope indicates a large value of ∆H°, whereas a shallow one indicates a small value of ∆H°.
4. Fit a line to the linear portion of the melting curve centered at Tm using the linear least squares technique. Calculate the slope of this line. The linear region of the melting curve is typically found at α values between 0.3 and 0.7. Significant curvature in this region may indicate a complex dissociation process; that is, the two-state assumption is violated.
5. Calculate ∆H° using the equation ∂α(T ) ∂α(T ) = −(2n + 2)R ∆Ho = (2n + 2)RT 2m ∂ T ∂(1 / T) T=Tm T=Tm Equation 7.3.9
for non-self-complementary or self-complementary complexes. CALCULATION OF ∆H° BY DIRECT APPLICATION OF THE VAN’T HOFF EQUATION
SUPPORT PROTOCOL 3
This method is from Puglisi and Tinoco (1989). 1. Calculate values for α(T) as described (see Support Protocol 1, steps 1 and 2). 2. Use the value of α at any temperature T to calculate a value for the association constant, K(T), where the expressions for K(T) depend upon the number of molecules (strands), n, in the equilibrium and the sum of their concentrations, CT. For non-selfcomplementary complexes use the equation α(T )
K (T ) =
n−1
CT n
[1 − α(T )]n
Equation 7.3.10
and for non-self-complementary complexes use the equation K (T ) =
α(T )
nC n−1 T [1
− α(T )]n
Equation 7.3.11
Biophysical Analysis of Nucleic Acids
7.3.11 Current Protocols in Nucleic Acid Chemistry
3. Plot lnK(T) versus 1/T and use the slope of this line to determine ∆H° by direct application of the van’t Hoff equation. ∆Ho = −R
∂ lnK (T ) ∂(1 / T )
Equation 7.3.12
SUPPORT PROTOCOL 4
CALCULATION OF ∆H° FROM DIFFERENTIAL CURVES In this protocol, the shape of the curve defined by the derivative of α with respect to 1/T is used to make a robust determination of ∆H° (Marky and Breslauer, 1987). 1. Calculate values for α(T) as described (see Support Protocol 1, steps 1 and 2). 2. Construct the derivative curve, ∂α(T)/∂(1/T) versus T, in a spreadsheet using the method of Savitsky and Golay (1964). Because the Savitsky-Golay method requires evenly spaced values on the temperature axis, compute ∂α(T)/∂T and use the relation ∂α(T)/∂(1/T) = −T2[∂α(T)/∂T] to get the requisite derivative. 3. Determine Tmax as the temperature at which the α(T) versus T curve attains its maximum slope. Tmax is easily found as the maximum of the derivative curve. It is important to note that Tm = Tmax only for monomolecular transitions.
4. Define T1 and T2 as the temperatures at which ∂α(T)/∂(1/T) = 12 [∂α(T)/∂(1/T)]max. Figure 7.3.6 shows an example of the derivative curve with Tmax, T1, and T2 indicated.
5. Calculate the value of ∆H° for the derived temperatures using the following relations ∆Ho =
B 1 1 T − T 2 1
or ∆Ho =
B′ 1 1 − T max T2
Equation 7.3.13
where values for B and B′ depend on the molecularity, n, of the transition and are shown in Table 7.3.3. This method is not recommended for analysis of repeating-sequence polynucleotide melting. These transitions are extremely sharp, making accurate differential curves difficult to calculate. Use Support Protocols 2 or 3 for these molecules.
Table 7.3.3 Curves
Values for the Constants B and B′ Used in Calculating ∆H° Values from Derivative
Molecularity, n 1 2 3 4
B –7.00 –10.14 –12.88 −15.40
B′ −3.50 −4.38 −5.06 −5.63
Optical Methods
7.3.12 Current Protocols in Nucleic Acid Chemistry
Figure 7.3.6 Determination of ∆H° from a derivative curve. The value of Tmax, T1, and T2 are easily found and used to calculate the value of ∆H°.
CALCULATION OF ∆G° USING THE VAN’T HOFF EQUATION
SUPPORT PROTOCOL 5
This method is from Plum et al. (1999). 1. Calculate values for α(T) as described (see Support Protocol 1, steps 1 and 2). 2. Determine the value of ∆H° by one of the methods described in the other Support Protocols and use it to extrapolate the free energy change (∆G°) associated with the duplex disruption to a reference temperature, typically 37° or 25°C (310° or 298°K). Use the equation
∆Go =
∆Ho R
1 − 1 + (n − 1)ln CT 2n T Tm
Equation 7.3.14
for non-self-complementary oligonucleotide complexes and
∆Go =
∆Ho R
1 − 1 + (n − 1)ln nCT 2 T Tm
Equation 7.3.15
for self-complementary oligonucleotide complexes. There is an assumption implicit in these equations that the heat capacity change (∆Cp°) for the transition is zero and that ∆H° is temperature invariant. These equations allow one to calculate ∆G° at any temperature, although they are valid only for matched pairs of Tm and CT values. Complementary expressions in terms of Tmax also can be used (Plum et al., 1995a). These are particularly useful when either the upper or lower baseline is not well defined, thereby making Tm difficult to determine.
Biophysical Analysis of Nucleic Acids
7.3.13 Current Protocols in Nucleic Acid Chemistry
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Alkaline phosphatase solution, 100 U/mL Dilute 25 µL of the enzyme (1000 U/mL; Calbiochem) with 225 µL of 10 mM cacodylate buffer solution (see recipe). Store at –20°C for up to 6 to 12 months. 1-Amino-2-naphthol-4-sulfonic acid (ANS) solution Weigh 5.95 g sodium bisulfite and 0.2 g anhydrous sodium sulfite. Dissolve in 25 mL water. Add 0.1 g 1-amino-2-naphthol-4-sulfonic acid (ANS). Dilute to 100 mL final volume. Store for up to 1 to 2 weeks at room temperature, protected from air and light. Cacodylate buffer solution, 10 mM Dissolve 0.21 g NaC2H6AsO2⋅3H2O (sodium cacodylate trihydrate) in 800 mL water in a 1-liter beaker. Adjust the pH to 7.0 with 1 N HCl. Transfer to a 1-liter volumetric flask and add water to bring the final volume to 1 liter, giving a 10 mM cacodylate solution. Store at 4°C for up to 6 months. CAUTION: Be careful not to breath the dust or come into contact with solutions containing cacodylate as it is an arsenic-containing compound.
Molybdate solution Dissolve 2.5 g Na2MoO4⋅2H2O (sodium molybdate dihydrate) in 100 mL of 10 N sulfuric acid (28 mL of concentrated H2SO4 plus 72 mL water). Store at room temperature for up to 12 months. CAUTION: Add acid slowly to water. Significant heat is produced upon mixing. Place the vessel in a bucket to prevent spills in the event of breakage.
P1 nuclease solution, 1 mg/mL Dissolve 1 mg in 1 mL water in vial provided by supplier (Calbiochem). Store at –20°C for up to 6 to 12 months. Standard phosphate solution Dry KH2PO4 in an oven overnight or under vacuum over P2O5 overnight. Dissolve 13.6 mg KH2PO4 in 1 liter of water using volumetric glassware to produce a 100 µM PO4 solution. Store at room temperature for up to 1 to 2 weeks. Prepare this solution with utmost care, as the entire procedure is dependent on its accuracy. If necessary, make a higher concentration solution and dilute accordingly. Preparation of a solution of exactly 100 mM PO4 concentration is less important than knowing the concentration precisely.
COMMENTARY Background Information The well-known relation described by van’t Hoff between the temperature dependence of the equilibrium constant and the enthalpy change associated with the equilibrium is shown in the eponymous equation: ∆Ho = RT 2 Optical Methods
∂ lnK (T ) ∂ lnK (T ) = −R ∂T ∂(1 / T )
The applications of this equation described herein are predicated on several assumptions. (1) The system examined is at equilibrium. Most oligonucleotide duplexes equilibrate rapidly; however, longer duplexes and highermolecularity complexes (especially quadruplexes) may require long equilibration times. (2) The equilibrium described must be twostate; that is, no thermodynamically significant intermediate states may be present. In the case
Equation 7.3.16
7.3.14 Current Protocols in Nucleic Acid Chemistry
of nucleic acid melting transitions, only the fully formed complex (or structure if the process is monomolecular) or fully melted single strands may be present. (3) The enthalpy change, ∆H°, is assumed to be independent of temperature; that is, the heat capacity change (∆Cp°) is zero. A large ∆Cp° results in curved van’t Hoff plots, which typically are not observed for nucleic acid melting transitions. (4) The observable, here UV, absorbance must reflect linearly the global extent of the melting transition. The shape of the melting curve is related to the transition enthalpy change, ∆H°, for the thermodynamic cooperative unit. For short duplexes, the cooperative unit is comprised of the entire molecule; however, for polymeric duplexes, the cooperative unit may be only a small fraction of the entire duplex. In all cases, the mole unit of the van’t Hoff enthalpy refers to the cooperative unit. The only method for determining the size of the cooperative unit is by comparison of the van’t Hoff enthalpy to the model-independent enthalpy value determined by differential scanning calorimetry. For complexes of molecularity >1, the association constant (K) depends on oligonucleotide concentration. Because of the relationship between K and Tm, the melting temperature also depends on oligonucleotide concentration. This leads to the preferred method (when n > 1; see Support Protocol 1) for extraction of thermodynamic data from equilibrium melting curves. For monomolecular processes, one or more of the alternate methods (see Support Protocols 2 to 5), which are based on analysis of single melting curves, must be used. It is useful to consider the behavior of oligonucleotide complexes as the length of the constituent strands increases. As the oligonucleotides increase in length the transition curves become sharper. This increase in sharpness does not necessarily indicate an increase in cooperativity. Rather, it is a manifestation of the increase in ∆H° of the thermodynamic unit due to the increase in the number of base pairs. As long as the two-state assumption is valid, cooperativity is maximal, yet the curves become sharper as the length increases. Eventually, the length increases to a point where the two-state assumption fails and thermodynamically significant intermediate states participate in the equilibrium. The general appearance of the melting curves is unaffected and, for n ≥ 2, the Tm still depends strongly on concentration. Yet the assumptions underlying the methods of analysis described here are
invalid, and the resultant thermodynamic data are compromised. In this length regime, the only recourse is to study the system by direct calorimetric methods (see UNIT 7.4). The point at which one must abandon van’t Hoff methods for calorimetry is not well defined. One approach that, while not absolute, does help address this problem is a comparison of the results from several of the methods described here. If all of the assumptions underlying the van’t Hoff methodology are valid, ∆H° values derived from the various methods should be in good agreement. The various methods (particularly the concentration-dependent versus single-curve methods) vary in sensitivity to deviations from the van’t Hoff model. While small differences are expected, significant disparities among the data derived from the various methods provide a good indicator of equilibria that are not amenable to analysis by van’t Hoff methods. The point where disparities become significant is a matter of debate; however, disparities greater than 5% to 10% call into question whether the two-state van’t Hoff model has been applied appropriately. Typically, disparities among the methods begin to appear in duplexes >12 base pairs in length. As the duplex length increases further, the concentration dependence of Tm vanishes. This is directly related to the mechanism of duplex disruption. Duplex formation can be described, at least qualitatively, by two parameters: initiation and propagation. Initiation involves the encounter of the two separate strands and the formation of a few base pairs. Initiation is a bimolecular process and, therefore, depends on concentration. Short duplex formation is dominated by initiation. In fact, the two-state assumption implies that initiation is the only thermodynamically significant event. Propagation is the extension of base pairing beyond the initiation complex. The process does not involve a change in the number of strands in the complex and is therefore concentration independent. Polymer formation is dominated by propagation, resulting in the failure of polymer melting temperatures to change with concentration. Intermediate-length duplexes form with significant contributions from initiation and propagation. They exhibit concentrationdependent melting due to the contribution from initiation, but significant populations of partially paired intermediate states contribute to the melting profile. Rigorous treatment of complexes of intermediate length (that is, those displaying nontwo-state behavior) requires application of par-
Biophysical Analysis of Nucleic Acids
7.3.15 Current Protocols in Nucleic Acid Chemistry
tition function techniques in which all states are enumerated (Poland, 1974). A useful compromise can be achieved by combining the calorimetrically determined ∆H°cal and the concentration dependence of Tm (or Tmax) to obtain self-consistent ∆G° values (Plum et al., 1995a). The measured slope, m, of the 1/Tm versus lnCT plot is substituted into the following expression to find a value for the effective molecularity, neff. neff =
m∆H ocal +1 R
Equation 7.3.17
The thermal disruption of higher-order complexes is assumed to proceed by a mechanism similar to that of duplexes, although far fewer data address this issue for nucleic acid complexes composed of three or more strands. The length at which the van’t Hoff treatment fails will depend on the molecularity of the complex. For DNA triple helices, there is evidence that the van’t Hoff methods are not reliable, even for relatively short oligonucleotides (Plum et al., 1995b). Higher-order complexes have not yet received sufficient study to define even crudely the length range over which van’t Hoff methods may be applied with confidence. To ensure that reliable values for thermodynamic parameters are obtained for oligonucleotide duplexes >12 base pairs and for higherorder complexes, model-independent, calorimetric experiments are advisable in parallel to the optical characterizations described here.
Critical Parameters
Optical Methods
Accurate knowledge of nucleic acid concentration is necessary for determining values of the thermodynamic parameters that describe the stability (∆G°) of a nucleic acid structure and the temperature dependence (∆H°) of that stability. Knowledge of the molecularity of the nucleic acid complex also is necessary. Determination of both concentration and molecularity is dependent on accurate values of extinction coefficients. Accurate concentration measurements are necessary to prepare solutions so that oligonucleotides are present in the correct ratios for complex formation, and reliable molar extinction coefficients for oligonucleotides are necessary to ensure that solutions are prepared accurately. Estimation of the extinction coefficient based on the base sequence is an alternative to the phosphate analysis procedure described here (Puglisi and Tinoco, 1989). The
level of precision (±10%) is often not satisfactory for thermodynamic measurements, particularly if parallel calorimetric studies are contemplated. Assuming that the concentration of the standard phosphate solution is known accurately, the most common error in the extinction coefficient determination is incomplete enzymatic digestion of the oligonucleotide. In this case, the amount of phosphate is underestimated and thus the extinction coefficient is overestimated. The use of the poly(rU) positive control should alert one to this problem. Failure of the color to develop properly indicates that fresh solutions should be prepared. All of the methods for extracting thermodynamic data from equilibrium melting curves assume a model for the oligonucleotide complex. Therefore, it is vital that the molecularity (i.e., the number of oligonucleotides in the complex) be known. Mixing curves are used to confirm the molecularities of the complexes formed by oligonucleotides, to validate the single-strand molar extinction coefficient values, and to determine molar extinction coefficient values for the complexes. The critical parameters in the determination of the molecularity of the complex are the individual extinction coefficients. Failure of the curves to meet indicates errors in the extinction coefficients or errors in solution preparation. If the points on the absorbance axis where X = 0.0 or X = 1.0 are not colinear with the data where 0.0 < X < 1.0, an incorrect determination of the minimum necessary volume for the absorbance measurement is indicated. Time dependence of the measured absorbance may indicate the necessity of an annealing step. The most important parameter in the determination of thermodynamic parameters by the several analysis protocols presented here is the length of the nucleic acid. As described in detail in Background Information, the methods presented here are valid only for nucleic acid structures that melt via a two-state process. This requirement restricts application of the methods to very short (≤12 base pairs) and very long (demonstrating no concentration dependence of Tm) nucleic acid complexes. Monomolecular processes may or may not be two state. The only method for evaluating the validity of the twostate approximation is comparison of the model-dependent values calculated as described here to the model-independent calorimetric values.
7.3.16 Current Protocols in Nucleic Acid Chemistry
Anticipated Results The determination of the extinction coefficient described in Basic Protocol 1 is dependent on the calibration curve as shown in Figure 7.3.1. Mixing curves typically will look like Figure 7.3.2. When like numbers of strands A and B form a complex, the inflection point of the curve will be at 0.5. When different numbers of strands A and B form a complex, the inflection point will shift. For example, formation of an A2B complex is indicated by an inflection point at 0.33 and formation of AB2 complex by an inflection point at 0.67. The melting curve of most nucleic acids will look like Figure 7.3.3. This appearance is in itself not proof of two-state behavior. If multiple transitions are observed, as is frequently seen for triple helices, two-state behavior clearly cannot be assumed. It may be possible, however, to dissect multiphasic transitions as a series of two-state transitions. In the absence of calorimetric data, any assumption of two-state behavior must be considered tentative.
Time Considerations The determination of oligonucleotide extinction coefficients requires 4 to 5 hr divided over 2 days. The determination of complex molecularity and extinction coefficient by the method of continuous fractions requires ∼4 hr. The melting experiments require ∼2 hr to set up, while the running time can range from 3 to 12 hr (unattended spectrophotometer time) depending on the heating rate for each melting experiment. Temperature-programmable multiple-cell holders greatly accelerate data throughput.
Literature Cited Felsenfeld, G., Davies, D.R., and Rich, A. 1957. Formation of a three-stranded polynucleotide molecule. J. Am. Chem. Soc. 79:2023-2024.
Griswold, B.L., Humoller, F.L., and McIntyre, A.R. 1951. Inorganic phosphates and phosphate esters in tissue extracts. Anal. Chem. 23:192-194. Marky, L.A. and Breslauer, K.J. 1987. Calculating thermodynamic data for transitions of any molecularity from equilibrium melting curves. Biopolymers 26:1601-1620. Plum, G.E., Grollman, A.P., Johnson, F., and Breslauer, K.J. 1995a. Influence of the oxidatively damaged adduct 8-oxodeoxyguanosine on the conformation, energetics, and thermodynamic stability of a DNA duplex. Biochemistry 34:16148-16160. Plum, G.E., Pilch, D.S., Singleton, S.F., and Breslauer, K.J. 1995b. Nucleic acid hybridization: Triplex stability and energetics. Annu. Rev. Biophys. Biomol. Struct. Dyn. 24:319-350. Plum, G.E., Breslauer K.J., and Roberts, R.W. 1999. Thermodynamics and kinetics of nucleic acid association/dissociation and folding processes. In Comprehensive Natural Products Chemistry, Vol. 7 (E.T. Kool, ed.), pp. 15-53. Elsevier Science Ltd., Oxford. Poland, D. 1974. Recursion relation generation of probability profiles for specific-sequence macromolecules with long range interactions. Biopolymers 13:1859-1871. Puglisi, J.D. and Tinoco, I. Jr. 1989. Absorbance melting curves of RNA. Methods Enzymol. 180:304-325. Savitsky, A. and Golay, M.J.E. 1964. Smoothing and differentiation of data by simplified least squares procedures. Anal. Chem. 36:1627-39. Snell, F.D. and Snell, C.T. 1949. Phosphorous. In Colorimetric Methods of Analysis, 3rd ed., Vol. 2, pp. 630-681. Van Nostrand, New York.
Contributed by G. Eric Plum Rutgers University Piscataway, New Jersey
Biophysical Analysis of Nucleic Acids
7.3.17 Current Protocols in Nucleic Acid Chemistry
Calorimetry of Nucleic Acids
UNIT 7.4
A number of units in this chapter describe a collection of techniques for evaluating the structures of nucleic acids. These techniques have been employed to generate a substantial and rapidly expanding database of nucleic acid structures. Yet it has become apparent that thermodynamic as well as structural information is required to develop a clear understanding of the complex relationships between structure, energetics, and biological function. In recognition of this requirement, the number of studies designed to characterize the thermodynamics of a broad range of nucleic acid structures has increased dramatically in recent years. This unit describes the application of calorimetry to characterize the thermodynamics of nucleic acids, specifically, the two major calorimetric methodologies that are currently employed: differential scanning and isothermal titration calorimetry. Differential scanning calorimetry (DSC; Basic Protocol 1) is used to study thermally induced order-disorder transitions in nucleic acids. A DSC instrument measures, as a function of temperature (T), the excess heat capacity (C ex p ) of a nucleic acid solution relative to the same amount of buffer solution. A single DSC profile provides a wealth of both thermodynamic and extrathermodynamic information, much of which cannot be obtained by any other technique. Specifically, from a single curve of C ex p versus T, one can derive the following information (see Commentary): the transition enthalpy (∆H), entropy (∆S), free energy (∆G), and heat capacity (∆Cp); the state of the transition (two-state versus multistate); and the average size of the molecule that melts as a single thermodynamic entity (i.e., the size of the cooperative unit). Isothermal titration calorimetry (ITC; Basic Protocol 2) is used to study the hybridization of nucleic acid molecules at constant temperature. In a typical ITC experiment, small aliquots of a titrant nucleic acid solution are added to an analyte nucleic acid solution (with the analyte nucleic acid being complementary to the titrant nucleic acid) and the released heat is monitored. Judicious selection of nucleic acid concentrations and buffer conditions results in a titration curve that can be analyzed to yield the stoichiometry of the association reaction (n), the enthalpy of association (∆H), the equilibrium association constant (K), and thus the free energy of association (∆G). Once ∆H and ∆G are known, the entropy of association (∆S) can also be derived. Thus, as noted above for DSC, a single ITC experiment also yields a wealth of thermodynamic information about the association reaction. Repetition of the ITC experiment at a number of different temperatures yields the ∆Cp for the association reaction from the temperature dependence of ∆H. STRATEGIC PLANNING General Considerations The accuracy of the thermodynamic data derived from the calorimetric protocols described in this unit depends to a great extent on the purity of the nucleic acids being studied and the accuracy with which the concentrations of the experimental nucleic acid solutions have been determined. All nucleic acids should be devoid of protein contaminants, with oligonucleotides being purified by high-performance liquid chromatography (HPLC) and/or denaturing polyacrylamide gel electrophoresis (PAGE) prior to their use in any calorimetry experiments. The concentrations of all nucleic acids used in calorimetric studies should be determined spectrophotometrically using experimentally derived extinction coefficients. UNIT 7.3 describes an excellent method for determining the extinction coefficient of a nucleic acid by digesting the nucleic acid (either enzymatically or
Biophysical Analysis of Nucleic Acids
Contributed by Daniel S. Pilch
7.4.1
Current Protocols in Nucleic Acid Chemistry (2000) 7.4.1-7.4.9 Copyright © 2000 by John Wiley & Sons, Inc.
chemically) and subsequently performing colorimetric quantification of the phosphate concentration.
Differential Scanning Calorimetry The choice of buffer is important in performing DSC on a nucleic acid. One should not choose buffers whose pKa values exhibit large temperature dependencies (e.g., Tris⋅Cl; ∆pKa/°C = −0.028). Thus, the selection of a DSC buffer should depend not only on its buffering capacity at a desired experimental pH, but also on its pKa having a minimal temperature dependence. Examples of suitable DSC buffers include phosphoric acid (pKa2 at 25°C = 7.20; ∆pKa2/°C = −0.0028), citric acid (pKa3 at 25°C = 6.40; ∆pKa3/°C ≈ 0), and acetic acid (pKa at 25°C = 4.76; ∆pKa/°C = 0.002). The choice of salt (cation) concentration is another important consideration in designing a DSC experiment. The thermal stability of a polyanionic nucleic acid molecule depends on the concentration of cation in solution. In general, the higher the cation concentration, the greater the thermal stability of the nucleic acid. Thus, one can modulate the temperature range over which the melting transition of the nucleic acid occurs by varying the salt concentration. This ability can prove useful for ensuring sufficient pre- and post-transition baseline readings for accurate analysis of the data. Multivalent cations generally are more potent stabilizers of nucleic acid thermal stability than monovalent cations. Thus, conferring a desired thermal stability upon a nucleic acid molecule requires lower concentrations of salts containing multivalent cations—e.g., MgCl2 or Co(NH3)6Cl3—than salts containing monovalent cations—e.g., NaCl or KCl. Note that different cations of similar valence (e.g., Na + versus Li+, Mg2+ versus Ca2+, or Co3+ versus spermidine3+) can differ significantly in the extent to which they thermally stabilize a given nucleic acid structure. In spite of this variability in degree of cation-induced thermal enhancement, a good rule of thumb for oligonucleotides is to use cation concentrations in the following ranges: 50 mM to 1 M for monovalent cations, 1 to 15 mM for divalent cations, and 20 to 500 µM for cations with valences of ≥3. For longer DNA fragments, such as polynucleotides (particularly those having high GC contents), it may be necessary to use cation concentrations that fall below these ranges to ensure complete denaturation of the nucleic acid over the experimentally accessible temperature range, as well as to prevent cation-induced aggregation and/or precipitation of the nucleic acid.
Calorimetry of Nucleic Acids
A third important criterion in designing a DSC experiment is choosing an appropriate concentration of the nucleic acid to be studied. The dissociation heat (enthalpy) of a nucleic acid molecule depends on both its length and its base sequence. In general, the heat of dissociation decreases with decreasing fragment length. Thus, DSC experiments on short nucleic acid fragments require larger concentrations than those on longer nucleic acid fragments. The minimum concentration of a given nucleic acid fragment required for a DSC experiment will depend not only on the length and sequence of the fragment, but also on the sensitivity of the DSC instrument being employed. The Materials section of Basic Protocol 1 lists a range of nucleic acid concentrations that is suitable for most DSC instruments. Note that the thermal stabilities of short nucleic acid molecules (e.g., ≤20 base pairs, base triplets, etc.) with molecularities of two or more depend on their concentrations, with increasing concentrations resulting in increased thermal stabilities. Thus, in studies on such short multimolecular nucleic acids, not only can one modulate the temperature range over which the melting transition occurs by varying the salt concentration (as noted above), but also by varying the nucleic acid concentration.
7.4.2 Current Protocols in Nucleic Acid Chemistry
Isothermal Titration Calorimetry It is critical to the success of an ITC experiment that concentrations of the solution components (i.e., buffer, salt, chelating agent) in the two nucleic acid solutions be as close to identical as possible. Such solution components can have high heats of dilution and thereby can introduce substantial error to the measurement. Similarly, one should ensure that the pH values of the two nucleic acid solutions are as close to identical as possible, since buffers often have substantial heats of protonation. The best way to insure that all the components in the two nucleic acid solutions are identical is to prepare a single batch of buffer solution, lyophilize an appropriate amount of each of the two nucleic acid samples to dryness, and dissolve each of the dried nucleic acid samples in the appropriate volume of buffer. DIFFERENTIAL SCANNING CALORIMETRY OF NUCLEIC ACIDS This protocol describes the use of DSC to study thermally induced order-disorder transitions in nucleic acids. Steps 1 to 5 describe the acquisition of a buffer-versus-buffer DSC scan, while steps 6 to 9 describe the acquisition of the corresponding nucleic acid–versus-buffer scan. Both scans are required, since the first step in data analysis mandates the background correction of the nucleic acid-versus-buffer scan by subtracting from it the corresponding buffer-versus-buffer scan. Analysis of the resulting DSC data is described below (see Commentary).
BASIC PROTOCOL 1
Materials Appropriate buffer Buffer solution of purified nucleic acid (0.2 to 2.0 mM in nucleotide) Nitrogen (N2) gas Differential scanning calorimetry (DSC) instrument Vacuum source and side-arm flask (for degassing) 1. Rinse both the sample and reference chambers of the DSC instrument thoroughly with a copious volume (0.5 to 1.0 liter) of distilled water and dry with nitrogen gas. 2. Degas a sufficient volume (typically 3 to 4 mL) of buffer to fill both the sample and reference chambers. Maintain vacuum for ∼10 to 15 min. 3. Fill both the sample and reference chambers with degassed buffer, being careful not to introduce any air bubbles. 4. Select the desired temperature limits and scan rate. A typical scan rate for a DSC experiment on a nucleic acid is 1.0°C/min. However, if the kinetics of the order-disorder equilibrium being studied are slow, one may need to lower this temperature scan rate in order to ensure equilibrium conditions throughout the scan. Note that upon decreasing the scan rate, one must increase the nucleic acid concentration in order to maintain the same signal strength. A good rule of thumb is to double the nucleic acid concentration upon halving the scan rate.
5. Allow the instrument to equilibrate (typically 20 to 40 min) and start the scan. 6. Repeat step 1. 7. Prepare a sufficient volume (typically 1.5 to 2 mL) of buffer to fill the reference chamber and a sufficient volume (typically 1.5 to 2 mL) of nucleic acid solution in buffer to fill the sample chamber. Degas both solutions for ∼10 to 15 min.
Biophysical Analysis of Nucleic Acids
7.4.3 Current Protocols in Nucleic Acid Chemistry
8. Fill the sample chamber with the degassed nucleic acid solution and the reference chamber with the degassed buffer, taking care not to introduce any air bubbles. 9. Repeat steps 4 and 5. BASIC PROTOCOL 2
ISOTHERMAL TITRATION CALORIMETRY OF NUCLEIC ACIDS This protocol describes the use of ITC to study the hybridization of nucleic acid molecules at a constant temperature. Analysis of the resulting ITC data is described below (see Commentary). ITC cannot be used to study the thermodynamics of self-complementary nucleic acids. For evaluating the thermodynamics of self-complementary nucleic acids, DSC is the only calorimetric technique that can be employed. The choice of the concentration of the titrant nucleic acid (nucleic acid T) will depend on the expected stoichiometry of its interaction with the analyte nucleic acid (nucleic acid A). When this stoichiometry is 1:1 (as would be the case for two single strands hybridizing to form a duplex), a good rule of thumb is to use 25 times more titrant than analyte. This ratio will ensure that the lower and upper baselines of the resulting titration curve are well defined. Materials Appropriate buffer Buffer containing purified analyte nucleic acid (nucleic acid A; 0.1 to 1.0 mM in nucleotide) Buffer containing purified titrant nucleic acid (nucleic acid T), whose base sequence is complementary to that of nucleic acid A and whose concentration is ≥20 times that of nucleic acid A Isothermal titration calorimetry (ITC) instrument Vacuum source and side-arm flask (for degassing) 1. Set the ITC thermostat to the desired temperature. If the desired temperature is at or below ambient temperature, use a refrigerated circulating water bath to facilitate the temperature regulation of the sample cell, and set the bath to a temperature ≥5°C below the desired ITC thermostat temperature. When using an external water bath, let the bath and ITC instrument equilibrate for ∼24 hr prior to running an experiment.
2. Degas appropriate buffer or distilled water for ~10 to 15 min and use it to fill the reference cell, taking care not introduce any air bubbles. Unless more frequent changes are warranted by corresponding changes in buffer or solvent, the solution in the reference cell need only be changed once per month.
3. Rinse the sample cell of the ITC instrument thoroughly with a copious volume (0.5 to 1.0 liter) of distilled water. 4. Prepare a sufficient volume of buffer solution containing nucleic acid A to fill the sample cell and degas for 10 to 15 min. 5. Fill the sample cell with nucleic acid A solution, being careful not to introduce any air bubbles. 6. Allow the instrument to equilibrate (typically 20 to 40 min). Calorimetry of Nucleic Acids
7. While the instrument is equilibrating, prepare a sufficient volume of buffer solution containing nucleic acid T to fill the titrating syringe and degas for 10 to 15 min.
7.4.4 Current Protocols in Nucleic Acid Chemistry
8. Fill the titrating syringe with nucleic acid T solution, being careful not to introduce any air bubbles into the syringe. 9. Keeping the syringe as upright as possible, insert the syringe into the sample cell and firmly seat into place. Align the motor-driven piston with the syringe plunger, if not done automatically by computer control. 10. Start rotating the syringe at a speed of ∼400 rpm and allow the instrument to equilibrate for 20 to 40 min. 11. After the instrument has equilibrated (i.e., the baseline is essentially unchanging), set the run parameters—including the number of injections, the injection volume, and the time between injections—and begin the titration. To derive parameters such as K and n from a single ITC experiment, the resulting titration curve should be defined by ≥10 points (i.e., ≥10 injections).
12. Repeat steps 3 to 11, replacing the nucleic acid A solution in the sample cell with buffer alone. This control experiment will yield the sequential dilution heats associated with injection of nucleic acid T into buffer, which, in turn, must be subtracted from the corresponding experimental heats resulting from the titration of nucleic acid T into nucleic acid A.
13. Repeat steps 3 to 11, replacing the nucleic acid T solution in the injecting syringe with a solution of buffer alone. This control experiment will yield the sequential dilution heats, if any, associated with injection of buffer into nucleic acid A. These heats often are negligible; however, in cases where they are not, they must be subtracted from the corresponding experimental heats resulting from the titration of nucleic acid T into nucleic acid A.
COMMENTARY Background Information In recent years, it has become apparent that thermodynamic as well as structural information is essential for understanding the nature of the relationships between the structures, energetics, and biological functions of nucleic acids. Recognition of this need, coupled with the commercial availability of sensitive calorimetric instruments, has led to a profound increase in the number of studies exploring the thermodynamics of nucleic acids (Sturtevant, 1987; Breslauer et al., 1992; Plum and Breslauer, 1995). Differential scanning and isothermal titration calorimetry are the two major calorimetric techniques that are currently employed to characterize the thermodynamics of nucleic acids. The purpose of this unit is to provide specific protocols for applying these two calorimetric techniques to the study of nucleic acid hybridization. Consequently, no attempt is made to review the theory of calorimetry or to present a description of instrument features. For such information, refer to the instruction manuals for the calorimeters, previously published reviews
(Sturtevant, 1987; Freire et al., 1990; Breslauer et al., 1992; Plum and Breslauer, 1995), as well as the original research articles referenced in these manuals and reviews. Differential scanning calorimetry Differential scanning calorimetry (DSC) is used to study thermally induced order-disorder transitions in nucleic acids. A DSC instrument measures as a function of temperature (T) the excess heat capacity (C ex p ) of a nucleic acid solution relative to the same amount of buffer solution. As detailed below, a single DSC profile provides a wealth of both thermodynamic and extrathermodynamic information, much of which cannot be obtained by any other technique. Integration of the experimental curve of C ex p versus T (see example in Fig. 7.4.1) yields the transition enthalpy (∆H), since ∆H = ∫CpdT. Note that this calorimetrically determined transition enthalpy (∆Hcal) is model independent and therefore does not depend on the state of the transition (two-state, multistate). This characteristic distinguishes ∆Hcal from
Biophysical Analysis of Nucleic Acids
7.4.5 Current Protocols in Nucleic Acid Chemistry
model-dependent van’t Hoff transition enthalpies (∆HvH), which are derived from the temperature dependence of equilibrium properties, and are typically predicated on the assumption (model) that the transition proceeds in an allor-none (two-state) fashion, with no thermodynamic contributions from intermediate states. Citing disparities that often arise between ∆HvH and ∆Hcal, Sturtevant and co-workers (Naghibi et al., 1995) have stressed the importance of using model-independent calorimetric measurements rather than less-reliable, model-dependent van’t Hoff analyses to characterize the thermodynamics of biological macromolecules. The heat capacity change (∆Cp) for the transition can be derived from the difference between pre- and posttransition baselines in a DSC measurement (Edsall and Gutfreund, 1983). This difference is often negligible for nucleic acids (i.e., ∆Cp ≈ 0), as is the case for the DSC profile shown in Figure 7.4.1. In cases where the experimental curve of C ex p versus T yields an insufficient amount of pre- or posttransition baseline for accurate integration and/or ∆Cp analysis, the salt and/or nucleic acid concentration can be modified (see Strategic Planning) to shift the transition to a higher or lower temperature range, as needed. The experimental curve of C ex p versus T can be converted to C ex /T versus T by dividing the p data by T and then replotting the raw C ex p resulting values as a function of T (Marky and
Calorimetry of Nucleic Acids
Figure 7.4.1 DSC profile for the thermal denaturation of the d(CCTCTCCGGCTCTTC)⋅ d(GAAGAGCCGGAGAGG) duplex. This DSC measurement was conducted on a model 5100 Nano Differential Scanning Calorimeter (Calorimetry Sciences) using a temperature scanning rate of 60°C/hr. The DNA concentration was 50 µM in duplex and the solution conditions were 10 mM sodium cacodylate (pH 7.0), 100 mM NaCl, 10 mM MgCl2, and 0.1 mM EDTA.
Breslauer, 1987). Integration of this curve yields the transition entropy (∆S), since ∆S = ∫(Cp / T) dT. Thus, a single DSC curve can yield ∆H, ∆S, and ∆Cp. Once these data are known, the corresponding transition free energy (∆G) can be determined at any temperature (T) using the following general thermodynamic relationship (Edsall and Gutfreund, 1983; Breslauer et al., 1992): ∆G = ∆H − T∆S. Note that although ∆S and ∆G can be extracted from DSC data, these data are less reliable than the ∆H and ∆Cp values obtained directly, due to the coupling and propagation of errors (Krug et al., 1976). ∆HvH can be determined by analysis of the shape (either the full or half width at half height) of an experimental curve of C ex p versus T using either of the following two relationships (Gralla and Crothers, 1973; Breslauer, 1995): For full width at half height: ∆HvH =
B 1 1 − T1 T2
Equation 7.4.1
For upper half width at half height: ∆HvH =
B′ 1 Tmax
−
1 T2
Equation 7.4.2
where Tmax is the temperature at the maximum of the experimental curve of C ex p versus T, and T1 and T2 correspond to the lower and upper temperatures, respectively, at which C ex p is equal to one half of the maximum value. B and B′ are constants that depend on the molecularity of the melting process under investigation. For a bimolecular process, such as the thermally induced denaturation of a DNA duplex into two single strands (as depicted by the DSC curve shown in Fig. 7.4.1), B and B′ are equal to 10.14 and 4.38 cal/mol⋅K, respectively. A comparison of ∆HvH and ∆Hcal allows one to evaluate the state of the transition (Marky and Breslauer, 1987). Specifically, if ∆HvH = ∆Hcal, then the transition proceeds in a twostate, all-or-none fashion. Under such conditions, meaningful thermodynamic data can be obtained from van’t Hoff analyses of equilibrium data. However, if ∆HvH < ∆Hcal, then the transition involves intermediate states, thereby precluding the use of the two-state van’t Hoff model. If ∆HvH > ∆Hcal, then intermolecular cooperation (e.g., aggregation) is indicated. A comparison of ∆HvH and ∆Hcal also provides insight into the cooperative nature of the tran-
7.4.6 Current Protocols in Nucleic Acid Chemistry
sition. Specifically, the ratio ∆HvH/∆Hcal provides a measure of the fraction of the structure that melts as a single thermodynamic entity (i.e., the size of the cooperative unit). Isothermal titration calorimetry Isothermal titration calorimetry (ITC) is used to study the hybridization of nucleic acid molecules at a constant temperature (Wiseman et al., 1989; Freire et al., 1990; Breslauer et al., 1992; Wilson et al., 1994; Plum and Breslauer, 1995; Plum et al., 1995). In a typical ITC experiment, small aliquots of a titrant nucleic acid solution are injected into an analyte nucleic acid solution (with the analyte nucleic acid being complementary to the titrant nucleic acid) and the heat of hybridization is measured directly. Each injection results in a heat burst curve, as depicted in Figure 7.4.2A. The heat ∂(∆Q) 1 = ∆H × V + 2 ∂([Ttot])
(∆Q; expressed in kcal or kJ per mole of injected titrant) evolved from each injection can be determined by integration of the corresponding heat burst curves and subsequently plotted as a function of the molar ratio of the two interacting nucleic acids ([Ttot]/[Atot]; see Fig. 7.4.2B). The resulting titration curve can be analyzed by nonlinear least squares fitting to yield the stoichiometry of the association reaction (n), the enthalpy of association (∆H), and the equilibrium association constant (K) (Wiseman et al., 1989; Freire et al., 1990). For a reaction with a 1:1 stoichiometry (n = 1), as is the case when two complementary nucleic acid strands hybridize to form a duplex, a plot of ∆Q versus [Ttot]/[Atot] can be fit to the following equation to yield ∆H and K (Wiseman et al., 1989), where V is the volume of the sample cell. 1 1 + K[A ] tot [Ttot] 1− − 2 2[Atot]
2 [Ttot] 1 1 1− + 1− −2 K[Atot] [Atot] K[Atot]
√ 2
[Ttot] [Atot]
Equation 7.4.3
Once the value of K has been determined in this manner, the free energy (∆G) of association can be derived from the relationship ∆G = −RTlnK. The resulting value of ∆G then can be used in conjunction with ∆H to derive the entropy (∆S) of association using the following standard relationship: ∆S = (∆H − ∆G)/T. Thus, as noted above for DSC, a single ITC experiment also yields a wealth of thermodynamic information about the association reaction. The heat capacity change, ∆Cp, for the association reaction can be determined by repeating the ITC experiment at two different temperatures using the following thermodynamic relationship: ∆Cp =
∂∆H ∆HT2 − ∆HT1 = ∂T T2 − T1
Equation 7.4.4
where T1 and T2 are the two different temperatures at which the ITC experiments were conducted. Note that K can be accurately derived from an ITC titration curve (∆Q versus [Ttot]/[Atot]) only if K ≤ 108 M−1. When K > 108 M−1 (as is often observed when two complementary nucleic acid strands hybridize to form a duplex), the extreme sharpness of the ITC titration curve precludes accurately fitting for
K. In such cases, it is better to use DSC to evaluate the hybridization thermodynamics. Caution should be exercised when comparing the thermodynamic parameters derived from ITC data with those derived from DSC data, since the single-stranded states of the nucleic acids exist at lower thermal energy (kT) in ITC experiments than they do in DSC experiments. The Breslauer group (Vesnaver and Breslauer, 1991) has shown that, for DNA duplex formation, thermodynamic data derived from ITC and DSC experiments can be quite different due to differences in the low- and high-temperature states of the DNA single strands. Recent studies by Sarai and co-workers (Kamiya et al., 1996) have revealed that the conformational states of single-stranded DNA also play a role in the thermodynamics of DNA triplex formation.
Critical Parameters As noted above (see Strategic Planning), two of the most critical parameters for both the DSC and ITC protocols described in this unit are the purity of the nucleic acids being studied and the accuracy with which the concentrations of the experimental nucleic acid solutions have been determined. All nucleic acids should be devoid of protein contaminants, with oligonucleotides being purified by HPLC and/or PAGE
Biophysical Analysis of Nucleic Acids
7.4.7 Current Protocols in Nucleic Acid Chemistry
ence. Examples of suitable DSC buffers include phosphoric acid (pKa2 at 25°C = 7.20; ∆pKa2/°C = −0.0028), citric acid (pKa3 at 25°C = 6.40; ∆pKa3/°C ≈ 0), and acetic acid (pKa at 25°C = 4.76; ∆pKa/°C = 0.002). Also of importance in a DSC experiment is selection of a temperature scan rate that ensures equilibrium conditions throughout the scan. For ITC experiments, it is of critical importance that the concentrations of the solution components (i.e., buffer, salt, chelating agent) in both the titrant and analyte nucleic acid solutions be as close to identical as possible. Such solution components can have high heats of dilution and thereby can introduce substantial error to the measurement. Similarly, one also should ensure that the pH values of the two nucleic acid solutions are as close to identical as possible, since buffers often have substantial heats of protonation.
Anticipated Results Figure 7.4.2 (A) ITC profile for the hybridization of d(CGTGTCCAGC) and d(GCTGGACACG) at 20°C. This ITC measurement was conducted on a MicroCal model MCS Titration Calorimeter (MicroCal). Five-microliter aliquots of a d(CGTGTCCAGC) solution (233.5 µM in strand) were sequentially injected from a 100µL rotating syringe (400 rpm) into 1.31 mL of a d(GCTGGACACG) solution (8.4 µM in strand). The duration of each injection was 4.93 sec and the delay between injections was 200 sec. The solution conditions were 10 mM sodium cacodylate (pH 7.0), 10 mM KCl, 10 mM MgCl2, and 5 mM CaCl2. (B) Integrated areas of each heat burst curve in (A) plotted as a function of the molar ratio of d(CGTGTCCAGC) to d(GCTGGACACG). The solid line reflects the nonlinear least squares fit of the data to Equation 7.4.3, where K = 6.1 × 107 M–1 and ∆H = –52 kcal/mole.
Calorimetry of Nucleic Acids
prior to their use in any calorimetry experiments. The concentrations of all nucleic acids used in calorimetric studies should be determined spectrophotometrically using experimentally derived extinction coefficients. For DSC experiments, choosing the correct buffer is of critical importance. One should not choose buffers whose pKa values exhibit large temperature dependencies (e.g., Tris⋅Cl, ∆pKa/°C = −0.028). Thus, selecting a DSC buffer should depend not only on its buffering capacity at a desired experimental pH, but also on its pKa having minimal temperature depend-
DSC enables one to measure the heat capacity (Cp) of a nucleic acid in solution as a function of temperature. Integration of the resulting Cp-versus-T curve yields the enthalpy change associated with the thermal denaturation of the nucleic acid. Comparison of this enthalpy value with the corresponding van’t Hoff enthalpy value, which can be obtained by analyzing the shape of the Cp-versus-T curve, allows one to evaluate both the state (e.g., two-state versus multistate) and the cooperativity of the transition. Dividing the experimentally observed Cp by T and plotting the resultant data as a function of T produces a Cp/T-versus-T curve, the integration of which yields the transition entropy (∆S). Once ∆S and ∆H are known, the transition free energy (∆G) can be calculated. ITC allows one to study the hybridization of two non-self-complimentary nucleic acids at a constant temperature. Specifically, one can measure the heat evolved from sequential injections of a nucleic acid titrant solution into a solution containing the complimentary nucleic acid (the analyte nucleic acid) to the titrant. Using nonlinear least squares analysis, one can fit a plot of the sequential injection heats as a function of the molar ratio of the two nucleic acids to yield the stoichiometry (n), enthalpy (∆H), and equilibrium association constant (K) for the hybridization reaction. Once ∆H and K are known, the free energy (∆G) and entropy (∆S) of the hybridization reaction can be calculated. Running the ITC experiment at two different temperatures allows one to determine the
7.4.8 Current Protocols in Nucleic Acid Chemistry
heat capacity change (∆Cp) associated with the hybridization reaction.
Marky, L.A. and Breslauer, K.J. 1987. Calculating thermodynamic data for transitions of any molecularity from equilibrium melting curves. Biopolymers 26:1601-1620.
Time Considerations In general, both calorimetric protocols described in this unit are fairly rapid, often requiring no more than 1 day to complete.
Naghibi, H., Tamura, A., and Sturtevant, J.M. 1995. Significant discrepencies between van’t Hoff and calorimetric enthalpies. Proc. Natl. Acad. Sci. U.S.A. 92:5597-5599.
Literature Cited
Plum, G.E. and Breslauer, K.J. 1995. Calorimetry of proteins and nucleic acids. Curr. Opin. Struct. Biol. 5:682-690.
Breslauer, K.J. 1995. Extracting thermodynamic data from equilibrium melting curves for oligonucleotide order-disorder transitions. Methods Enzymol. 259:221-242. Breslauer, K.J., Freire, E., and Straume, M. 1992. Calorimetry: A tool for DNA and ligand-DNA studies. Methods Enzymol. 211:533-567. Edsall, J.T. and Gutfreund, H. 1983. Calorimetry, heat capacity, and phase transitions. In Biothermodynamics: The Study of Biochemical Processes at Equilibrium, pp. 210-227. John Wiley & Sons, New York. Freire, E., Mayorga, O.L., and Straume, M. 1990. Isothermal titration calorimetry. Anal. Chem. 62:950A-959A. Gralla, J. and Crothers, D.M. 1973. Free energy of imperfect nucleic acid helices III. Small internal loops resulting from mismatches. J. Mol. Biol. 78:301-319. Kamiya, M., Torigoe, H., Shindo, H., and Sarai, A. 1996. Temperature dependence and sequence specificity of DNA triplex formation: An analysis using isothermal titration calorimetry. J. Am. Chem. Soc. 118:4532-4538. Krug, R.R., Hunter, W.G., and Grieger, R.A. 1976. Enthalpy-entropy compensation. 1. Some fundamental statistical problems associated with the analysis of van’t Hoff and Arrhenius data. J. Phys. Chem. 80:2335-2341.
Plum, G.E., Pilch, D.S., Singleton, S.F., and Breslauer, K.J. 1995. Nucleic acid hybridization: Triplex stability and energetics. Annu. Rev. Biophys. Biomol. Struct. 24:319-350. Sturtevant, J.M. 1987. Biochemical applications of differential scanning calorimetry. Annu. Rev. Phys. Chem. 38:463-488. Vesnaver, G. and Breslauer, K.J. 1991. The contribution of DNA single-stranded order to the thermodynamics of duplex formation. Proc. Natl. Acad. Sci. U.S.A. 88:3569-3573. Wilson, W.D., Hopkins, H.P., Mizan, S., Hamilton, D.D., and Zon, G. 1994. Thermodynamics of DNA triplex formation in oligomers with and without cytosine bases: Influence of buffer species, pH, and sequence. J. Am. Chem. Soc. 116:3607-3608. Wiseman, T., Williston, S., Brandts, J.F., and Lin, L.-N. 1989. Rapid measurement of binding constants and heats of binding using a new titration calorimeter. Anal. Biochem. 179:131-137.
Contributed by Daniel S. Pilch University of Medicine and Dentistry of New Jersey Robert Wood Johnson Medical School Piscataway, New Jersey
Biophysical Analysis of Nucleic Acids
7.4.9 Current Protocols in Nucleic Acid Chemistry
Molecular Modeling of Nucleic Acid Structure Molecular modeling, loosely defined, relates to the use of models to investigate the three-dimensional structure, dynamics, and properties of a molecule or set of molecules. At the heart of this is specification of a molecular model, which provides a molecular structure at an appropriate level of granularity, usually in terms of three-dimensional atomic coordinates. Molecular modeling can be approached on many levels, ranging from energy minimization (finding the set of coordinates that minimizes the energy) with a complete ab initio quantum-mechanical treatment of the energetics, to sampling “reasonable” conformations with a simplified energy representation or potential, to the manipulation of physical models where no implicit energy representation is included. These methods serve not only as tools to aid in the interpretation of experimental data, but to directly complement such data by providing a relationship between the macroscropic behavior observed experimentally and the microscopic properties represented in the model or simulation. As discussed in previous units, various molecular modeling tools can serve as conformational search engines for sampling conformational space subject to the restraints inferred from nuclear magnetic resonance (NMR; see UNIT 7.2) and crystallography (see UNIT 7.1) experiments. This is a critical step in the refinement of three-dimensional atomic structure. Inclusion of some representation of the energy, such as through the use of a specially parameterized empirical force field, can aid in this endeavor by limiting sampling to more realistic (in terms of energy) conformations. As mentioned above, molecular mechanics methods can not only be used as a tool, but can directly complement experimental data. For instance, molecular dynamics simulations can be used to aid in the interpretation of NMR order parameters or to estimate anisotropic rotational diffusion. In addition, computer simulation techniques have the potential to give structural and dynamic insight into the atomic interactions occurring on a time scale (<µsec) typically not observable due to averaging in crystallography and NMR experiments. Ultimately, as methods are proven reliable, they can then be applied in cases where experimentation is limited, difficult, or unfeasible, such as study-
ing highly flexible systems, investigating proposed chemical modifications that have yet to be synthesized, or to represent extremes of pressure, temperature, and concentration. As will become apparent, the methods are steadily improving to the point that reliable predictions are emerging. A critical point that needs to be made at the outset is that these methods cannot be treated as a “black box” or hands-off procedure; there is no standard protocol that can be applied. Modeling is really more of an art. As each situation has differing requirements and needs, various choices need to be made as to what level of treatment to apply and what model to use. These choices rely on a critical understanding of the limitations in the methods. Therefore, the purpose of this discussion is to open up this black box a bit to allow some understanding of the options and choices a modeler makes, highlighting the tradeoffs that must be made in accuracy, system size, and time. The discussion here and in UNITS 7.8 to 7.10 is not meant to provide a complete review of nucleic acid modeling, nor to substitute for the more complete treatment discussed in the primary literature. Instead, these units are intended to provide a framework that describes molecular modeling of nucleic acids, points out common issues and limitations, and points the reader to other useful information sources. Implicit in this discussion is a realization that a molecular model is more than simply a representation of the covalent connectivity or static structure. The model may also include some representation of the energetics of the system and perhaps the dynamics over a particular time scale. Although it increases the utility, supplementing static structure with a representation of the energy and dynamics of molecular motion tremendously increases the cost of the modeling. For example, the simulations required to accurately represent the sequence-specific structure and molecular dynamics of a small, solvated nucleic acid duplex (<20 base pairs) on a nanosecond time scale would likely require weeks to months on available computer workstations, even with simple empirical energy representations. Of course, this added information may not always be necessary. For example, to investigate whether a proposed modification to a DNA base is steri-
Contributed by Thomas E. Cheatham, III, Bernard R. Brooks, and Peter A. Kollman Current Protocols in Nucleic Acid Chemistry (2000) 7.5.1-7.5.12 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 7.5
Biophysical Analysis of Nucleic Acids
7.5.1 Supplement 6
cally feasible may only require the crude manipulation of a physical model to see an effect. Therefore, it is critical to understand the applicability, reliability, and limitations of these methods. In other words, the choice of the model depends on the question being asked. The remainder of the discussion in this unit introduces the simplest levels of molecular modeling applied to nucleic acids. These include generation, evaluation, and characterization of the initial molecular model. At this simplest level, a nucleic acid model is limited to a static representation of the structure in the gas phase. Evaluation of this given model’s utility is therefore based on the chemical intuition of the modeler, where manipulations to the model are limited to rotation about single bonds. To move beyond this level, supplement units in this series will delve more deeply into the myriad of issues involved in the computer simulation of nucleic acids. These include describing the common energy representations for nucleic acids that may be applied (UNIT 7.8), and discussion of how to properly represent the electrostatic interactions and solvation effects (UNIT 7.9). Additionally, various methods to find more representative structures are introduced, with a focus on molecular dynamics simulation methodologies. Finally, a description of practical issues in nucleic acid simulations will be provided (UNIT 7.10), such as what force fields are appropriate to apply, how simulations of nucleic acid are set up with explicit solvent and counterions, and how crude relative free energy differences can be estimated from molecular dynamics simulations. In these discussions, the focus will be on the middle ground in terms of size, time scale, and accuracy—that is, the simulation of small nucleic acids (typically less than ~250 base pairs), with explicit representation of the environment (if feasible or necessary), empirical pairwise potential functions, and time scales ranging from the analysis of individual snapshots to nanosecond-length simulations. For those readers more interested in learning about the simulation of larger nucleic acid systems (∼1,000 to 15,000 base pairs), a variety of reviews can be consulted (Vologodski and Cozzarelli, 1994; Schlick, 1995; Olson, 1996).
Molecular Modeling of Nucleic Acid Structure
Figure 7.5.1 Schematic representation of molecular modeling analysis.
Prior to generating an initial molecular model, it is necessary to choose its representation or level of detail. For nucleic acids, the structural representation can be approached on many levels, ranging from the atomic level (including electrons) to coarser levels, such as those that model structure using a single point per base pair. The realism of the model directly depends on this choice of representation and further depends on what properties one is trying to represent. As shown in Table 7.5.1, modeling can be considered a tradeoff between the accuracy, the size and granularity of the system, and the time scale to be represented. If the model only concerns a single conformation or small set of conformations of a molecule of <100 atoms, a very accurate energy model and a description that includes all the atoms and electrons can be used (such as ab initio quantum mechanics with a fairly large basis set and even correlation). However, to investigate the supercoiling of a small DNA plasmid over a microsecond time scale, the system can no longer be represented at the atomic level, and a much simpler description of the energetics and a coarser representation of the structure must be imposed. However, this may be sufficient to represent the properties of interest. Between a full quantum mechanical treatment appropriate for small molecules and the coarse-grained single point per base pair model appropriate for large systems, molecular dynamics methods with an empirical potential may give reliable results as long as no “chemistry” is involved (such as bond forming, bond breaking, or electron transfer) and highly polarizable metal ions are treated at a very approximate level. These methods can give reliable insight into the sequence-specific structure and dynamics of a small nucleic acid duplex in solution.
MOLECULAR MODELING
The Static Structure Model
The practice of molecular modeling basically involves the generation of an initial molecular model, evaluation of the model’s utility, and perhaps manipulation of the molecular model (followed by further evaluation; see Figure 7.5.1).
At the simplest level, and where the representation of the model does not include any reality beyond the covalent connectivity, molecular modeling can be performed by creating and manipulating physical models. Physical
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Current Protocols in Nucleic Acid Chemistry
Table 7.5.1 Tradeoffs in Molecular Modeling
Effective potential
Time scale (decreasing) Microseconds
Molecular mechanics (implicit solvent) Molecular mechanics
Nanoseconds to microseconds Nanoseconds
Quantum mechanics
Individual snapshots
Accuracy (increasing)
models are available that can represent three levels of granularity. At the finest level, there are a variety of atomic and molecular orbital models that represent the atoms and electrons. These molecular orbital models are not really appropriate for larger and more complicated molecules (such as anything larger than perhaps benzene), and therefore their use is really limited to teaching. Much more useful for representing nucleic acid structure are models that represent the atoms and bonds and, therefore, the covalent connectivity of a molecule. There are a few common types of models in use that can be classified as either space-filling or bond-oriented. The most common space-filling models are of the Corey-Pauling-Koltun (CPK) variety, named after the researchers that developed them. These space-filling models represent the various atoms as cut-out spheres of a size proportional to the van der Waals radius, which are colored and shaped according to atom type and can be connected together (based upon the hybridization state and possible connectivity of the atom). The most common bond-filling models are polyhedral models. These provide a series of pieces that are in various polyhedral shapes with holes for pegs, which represent the bonds. The shape, color, and number of holes represent the various atom types (and hybridization state), and connecting pegs represent the bonds. Although these models are useful for teaching and for building models of small molecules, they are not appropriate for building macromolecular models, such as of a DNA duplex. To build a larger molecule, special-purpose and more durable physical models can be purchased. These provide larger building units (such as DNA bases) in addition to smaller atom/half-bond units, which can be connected together. The scale of these models is usually
System size (decreasing) Supercoiled DNA, plasmid <1000 base pairs <250 base pairs Nucleotide(s), few waters/ions
Granularity (finer grain) One point per base pair, elastic rod All atom, implicit solvent All atom, explicit solvent All atom plus electrons, implicit solvent
in the 1 cm to 1 inch per Å range. Some models that have been used successfully are the Maruzen models, such as the HGS Biochemistry Molecular Model (see Internet Resources). Coarser folded-chain models, such as protein models that represent a connection/bond for each α-carbon, are also in use. The physical bond-oriented models, although tedious to build and often very fragile, are very useful for gaining insight into atomic structure. In addition, the models can be manipulated (which can lead to problems with larger model structures, as they tend to deform). Although the models have rigid bonds and angles, they typically allow free rotation about single bonds. This can provide insight into the correlated conformational changes that occur upon change in a given coordinate. One example is the change in sugar pucker conformation from C2′-endo to C3′-endo, which lowers the rise between base pairs and shifts the conformation not only of the atoms in the ribose ring but also of the nucleic acid backbone. In fact, modeling B-DNA with physical models led to the formulation of Calladine’s rules, which suggest means to overcome strong steric hinderances between adjacent purines in opposite strands as the base pair propeller twist increases to improve stacking.
Computational Graphics and Energy Models A problem with physical models is that there is no reliable means to include a description of the energy. With these models, energy can only be represented rather crudely, such as by inhibiting free rotation because of the connectivity or by the addition of physical restraints to prevent rotation about double bonds. This allows a minimal interpretation of the intra-
Biophysical Analysis of Nucleic Acids
7.5.3 Current Protocols in Nucleic Acid Chemistry
Molecular Modeling of Nucleic Acid Structure
molecular or internal energetics of the system (related to the connectivity of the molecule). In addition to intramolecular interactions, a realistic depiction of the energy requires representation of the intermolecular interactions (e.g., van der Waals or steric repulsion and dispersion attraction interactions, hydrogen bonding, and electrostatic interactions). Although the solid-sphere models can represent steric repulsion, they cannot be used to accurately describe the total energy; however, a realistic treatment of the energetics can readily be calculated by computer. Coupled with molecular graphics (digital display of molecular models), computational energy models open the door for much more realistic and reliable molecular modeling. Prior to the advent of molecular graphics, physical models were routinely used as aids for crystallographic refinement. Molecular graphics programs are now abundant and allow very nice and realistic display of molecular structure. The generality of the programs removes some of the tedium and cost of building physical models. However, since the computer graphics display is two-dimensional, the ease of seeing the three-dimensional model is lost and needs to be recovered by coloring, shading, or rotating the model to project the third dimension. Alternatively, stereoview displays can be used, which allow threedimensional viewing with special glasses (either through shuttering, as with the Crystal Eyes display, or with coloring and shading). In addition to more general usage, adding a description of the conformational energy to the molecular model is easier on the computer. Including a picture of the energy along with the molecular graphics can provide greater insight and help aid in the evaluation of the model. Examples include coloring regions of a molecule based on favorable electrostatic potential or highlighting atoms that show significant steric overlap. The manipulations possible at the simplest level mirror those of physical models and include a variety of coordinate manipulations, such as rotating about bonds or chemically modifying the structure. However, rather than manipulating the model by hand as with physical models, hooks need to be provided in the molecular graphics software to allow selection and rotation of various parts of the molecule. Given a reliable initial model structure, molecular modeling with simple coordinate manipulations may be sufficient for many applications, such as suggesting that it is not feasible
to fit a particular drug into the minor groove of a double-helical nucleic acid without seriously distorting the duplex, or showing that a certain chemical modification to the phosphodiester backbone is incompatible with the model structure. Simple modeling and molecular graphics were used as a guide in the initial design of peptide nucleic acid (PNA), an isosteric and stable backbone modification to DNA proposed for use as an antisense therapeutic agent (Nielsen et al., 1991). Manipulation of molecular graphics or physical models, when coupled with an appropriate chemical/structural intuition, can give useful information. Examples include understanding steric effects, such as the interaction of drugs with the grooves or base pairs of nucleic acid duplexes or correlated changes in structure due to rotation about particular bonds. However, a major issue with this type of modeling is evaluation of the molecular models. Evaluation and interpretation of the meaning of the molecular model depends on the quality of the initial model, the reliability of the energy representation (if any), and the choice of coordinate manipulations to the model that might be made. Without a reliable guide into the conformational energetics and coordinate manipulations necessary to “improve” the model, evaluation of the model depends solely on the chemical intuition of the modeler. This intuition is necessary to rule out unfeasible or unrealistic models or to suggest manipulations to the model that may improve the property of interest. Because there is no easy way to judge the quality of these models within this simple modeling framework, the conclusions made are often tenuous in the absence of experimental verification. For example, the initial model may not have been at all representative of what is seen experimentally or structural manipulations may lead to a model structure that is energetically unreasonable. Although the situation, in principle, improves with more advanced treatments because the energy is included and unreasonable coordinate manipulations are avoided, there are still many limitations in the methods. This is compounded by the sheer complexity of rugged energy landscapes for biomolecular structures, which makes evaluation of the reliability of a model structure difficult. In this sense, it should not be immediately assumed that “better” results are seen with more advanced treatments only because more reliable methods are used. There is still an essential need to compare the model
7.5.4 Current Protocols in Nucleic Acid Chemistry
with experimental data and to critically evaluate the model. To aid the modeler with simple molecular modeling, perhaps the ultimate molecular modeling environment might involve viewing a molecular graphics depiction of the model as it updates in real time according to the underlying energy potential, while the model is manipulated according to the whims of the modeler. An example of this type of program is Sculpt (Surles et al., 1994), which allows real-time minimization of the structure as it is manipulated. Further enhancement to this environment could come from visual and aural feedback from the system, such as a bang sound and flash of red light, to discourage manipulations by the modeler that move atoms into sterically forbidden regions. More involved haptic feedback mechanisms are also possible, such as increasing the difficulty of performing a given manipulation in proportion to the energetic penalty. Ultimately, molecular modeling environments of this type will incorporate visual, aural, and tactile feedback mechanisms, coupled with stereoscopic three-dimensional display in a virtual reality “cave” (Cruz-Neira et al., 1992), to guide the modeler as the model is manipulated. Software to perform this type of real-time modeling has become available in recent years, although the complexity of the calculations limits the treatment, and therefore fairly approximate representations of the energetics must be employed. Nevertheless, this ultimate molecular modeling facility, with realistic energy representations and user feedback to steer the various molecular manipulations, unfortunately does not give a complete understanding of the molecular structure. The energy (enthalpy) alone is insufficient to describe the relative stability of various models, and care needs to be levied in judging the reliability of models based on differences in energy. In addition to describing the energy of the system, it is also necessary to include entropic effects. When entropic effects are included, free energy values may be obtained, providing the connection with reality and experimental measurement. With free energy, the modeler has a handle on the relative population of each state or can equivalently understand the various thermally accessible conformations of the molecule in its native environment. To add entropic effects, some means of sampling the space of accessible conformations (according to the relative probability of observing a given conformation or equivalently ac-
cording to the Boltzmann distribution) is needed. To do this, molecular dynamics (MD) or Monte Carlo (MC) simulation (discussed in more detail in UNIT 7.8) can be done with the given energy representation. This, however, tremendously increases the cost and complexity of modeling. Whether or not the sampled space of conformations is representative depends on the reliability of the energy description, the amount of conformational sampling, and the reliability of the initial model. However, it should be emphasized that more costly and detailed treatments do not always lead to “better” insight and are not always necessary to address the question at hand.
Generating the Initial Model The first step in any modeling endeavor is creation of the initial molecular model, where “model” refers to a particular set of three-dimensional coordinates that define the structure of interest. In this discussion, which concerns nucleic acid structure on an atomic level (as opposed to the more coarse-grained bead models appropriate for modeling larger nucleic acid structures), this model is the set of three-dimensional atomic coordinates. Generally, a model of the coordinates is built by hand or received from another source (such as a database of experimentally derived structures). As will become more apparent later in this overview, the quality of the modeling in large part relates to the quality of the initial model or the ability to find or sample the “correct” structure given the initial model. In this regard, studying an unknown RNA structure is likely to be unfeasible at present, since it is unrealistic to imagine correctly folding up the RNA structure in dynamics simulations (due to barriers to conformational transition that cannot be overcome during the time scale of the simulations, and to inaccuracies in the energetic representation). Although there has been tremendous progress in predicting RNA secondary structure, predicting the overall tertiary structure (i.e., threedimensional atomic coordinates) is still a major unsolved challenge. In spite of this, there have been a few attempts (for review see Brion and Westhof, 1997; Leclerc et al., 1997). Therefore, it is best to base the modeling on experimentally derived structures. Since DNA tends to adopt regular duplex structures, one can often use the canonical structures as an initial guess. The canonical models were derived from fiber diffraction studies of large DNA fibers and give an average idealized geometry and structure
Biophysical Analysis of Nucleic Acids
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Supplement 6
representative of DNA under specific conditions (such as A-DNA under low humidity and B-DNA under physiological conditions; Arnott and Hukins, 1972). Crystallography provides another source of high-resolution structures, such as the left-handed Z-DNA duplex (Wang et al., 1979). The common canonical forms of DNA (A-DNA, B-DNA and Z-DNA) are shown in Figure 7.5.2 as stereo views. A good resource (although somewhat out of date) for general information on the structure of DNA is Saenger’s excellent book (Saenger, 1984). High-resolution structures are also emerging from NMR spectroscopy (Ulyanov and James, 1995; also see UNIT 7.2). A more recent book surveying nucleic acid structure and interactions as well as NMR and crystallography studies is Bioorganic Chemistry: Nucleic Acids (Hecht, 1996). Many of the experimentally derived nucleic acid structures are freely available through either the Protein Data Bank (PDB; see Internet
Molecular Modeling of Nucleic Acid Structure
Resources; Abola et al., 1987) or the Nucleic Acid Database (NDB; see Internet Resources; Berman et al., 1992), both of which contain the coordinates for a variety of nucleic acid structures and protein-nucleic acid complexes derived from crystallography or NMR experiments. The NDB may be a more appropriate place to start, as (1) it has been specifically tailored to assemble and distribute structural information about nucleic acids, (2) it can be searched, and (3) it provides coordinates (in multiple formats) as well as information about the crystal parameters, packing, and experimental conditions. From both of these sources, coordinate files in the commonly used PDB format can be obtained. If an experimental structure is not available, it may still be possible to generate a reasonable model structure. A tool (or more accurately, a language for molecular manipulation) that can help develop such an initial model is Nucleic Acid Builder (NAB) developed by Tom Macke
Figure 7.5.2 Canonical structures of DNA shown as stereo views. Shown are canonical models of A-DNA and B-DNA of d[CCAACGTTGG]2 (Arnott and Hukins, 1972) and a 10-mer extended model of the Wang Z-DNA structure of d[CGCGCGCGCG]2 (Wang et al., 1979) as stereo views. Stereo views are common in the literature; these are wall-eyed stereo views as opposed to cross-eyed. Although some people can view these directly, most people resort to one of a variety of hand-held stereo viewers, such as those based on mirrors or better ones that use focusing lenses. The model of Z-DNA was built by overlaying the two 6-mers at the joining region to the root-meansquared (RMS) best fit overlapping CpG steps, and the A/B-DNA models were built using the NUCGEN module of AMBER 4.1 (Pearlman et al., 1995). The A-DNA and B-DNA models were all-atom RMS best fit to a common reference frame, and the view is into the major groove on top and the minor groove on the bottom.
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Current Protocols in Nucleic Acid Chemistry
and Dave Case (Macke and Case, 1998). The NAB molecular manipulation language allows a specification of rigid body translations, specification of restraints, distance geometry methods, and various other tools to aid in the generation of arbitrary structures. This has been used to generate model structures of synthetic Holliday junctions, protein-DNA complexes, RNA pseudoknots, supercoiled DNA, and other structures (Macke and Case, 1998). If the model shares properties with other known structures, such as common secondary structure elements or sequence, it may be possible to model by homology to the known structures or, alternatively, to build up the structure from a library of smaller pieces of known structure. This approach has been used to model RNA tertiary structure (Major et al., 1991) and the structure of DNA single strands (Erie et al., 1993). Recent surveys of crystal structures in the Cambridge Structure Database (which contains a variety of high-resolution structures of mononucleosides and mononucleotides; Allen et al., 1979) and the NDB (Berman et al., 1992) provide a set of parameters that can serve as the beginnings of a dictionary for standard nucleic acid geometry. These surveys investigate the geometry of the bases (Clowney et al., 1996) and the sugar and phosphate backbone (Gelbin et al., 1996; Schneider et al., 1997). Additionally, recent surveys have investigated the specific hydration of nucleic acids and interaction with metal ions (Schneider et al., 1993; Schneider and Kabelac, 1998). High-level theoretical techniques can also give useful information. Ab initio quantum-mechanical simulations with a reasonable basis set (6-31 G* or better) and some inclusion of correlation can accurately represent geometry and polarization effects, and therefore properly represent nucleic acid interaction with various ions, metals, or nucleic acid bases. Monte Carlo and molecular dynamics simulation can also be used to obtain specific insight into ion association and hydration.
Completing the Initial Model Often the experimentally determined structures obtained from the PDB or NBD lack explicit hydrogen atoms. Additionally, the nomenclature used is invariably different from that of the given modeling program, and the user has to impose various contortions to coerce the file into the expected naming and numbering conventions. Therefore it is fairly common to have to modify a PDB file to conform to the
particular program’s pedantic conventions and, additionally, to somehow add hydrogen atoms to the structure. Almost all of the modeling programs are equipped with some facility for adding missing atoms, particularly hydrogens. For more advanced treatments, solvent and counterions can also be added (discussed in UNIT 7.9). It is always a good idea to check the initial structure carefully to determine if the conformation and nomenclature is as expected and whether the hydrogens are added with the correct stereochemistry. It would be very disappointing to discover, after spending weeks running nanosecond-length molecular dynamics simulations of solvated DNA, that one of the H1′ atoms on a particular residue was inadvertently added with the wrong stereochemistry, leading to an α-glycosyl linkage rather than the expected β linkage. It is likewise critical to check the stereochemistry of the structure after manipulations to the molecular model are made. Under some conditions, such as when using distance geometry methods or when performing stringent minimization with large restraints, the structure can be distorted and the stereochemistry altered. Although not all modeling programs adhere to IUPAC naming conventions (JCBN, 1983; see APPENDIX 1C), these conventions are a good reference to check the naming, orientation, and placement of the various atoms. Additionally, there are a variety of tools for characterizing the nucleic acid structure, which are discussed in the next section. However, these methods do not necessarily check stereochemistry, depend on the use of correct hydrogen naming conventions, or enforce IUPAC naming conventions. Although the PDB format is a common and well-defined standard for three-dimensional atomic coordinates, not all programs understand the standard PDB format, and they instead rely on some subtle variant or expect another coordinate format entirely. To aid in converting between the large set of formats available for many of the various modeling tools, the program babel is very useful (see Internet Resources). Not only can this perform direct conversion among various coordinate file formats, it can assign connectivity, bond orders, and hybridization when this information is not present.
Characterizing Nucleic Acid Structure In order to characterize the quality of an initial molecular model or to later evaluate the conformational changes that occur as the model
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is manipulated (for example, during MD simulation), it is useful to characterize the overall three-dimensional structure. In proteins, one is typically only concerned with the φ and ψ backbone angles and perhaps some of the sidechain χ angles; the overall structure is characterized by the particular secondary structure elements and folding class. In contrast, with nucleic acids, there are many angles of interest. These range from the backbone angles α, β, γ, ε, and ζ , to the puckering conformation of the furanose ring, to the χ angle representing the orientation of the sugar to the base (Saenger, 1984; see APPENDIX 1B). To characterize the conformation of the sugar moeity (the furanose ring), the Altona and Sundaralingam concept of pseudorotation is generally used (Altona and Sundaralingam, 1972). This defines the sugar pucker amplitude (representing how far the ring is from planar) and the pseudorotation phase angle (representing the correlated values of the individual torsions making up the ring). Various values of the pseudorotation phase angle, more commonly referred to as the sugar pucker, represent different puckerings out of the plane (on the same side as the C5′ atom, endo, or to the opposite side, exo). Methods for calculating these values are straightforward and are typically included in most modeling packages. In addition to characterizing the overall backbone structure, sugar pucker, and χ angle
Molecular Modeling of Nucleic Acid Structure
Figure 7.5.3
of a single polynucleotide strand, it is also desirable to characterize the commonly occurring duplex structures that result from complementary base pairing between strands. Helicoidal analysis is typically applied to characterize global properties of the duplex (such as the helical repeat or overall helical twist), properties between adjacent base pairs (such as the rise), or properties of individual bases (such as the propeller twist). These properties represent the extent of rotation or translation of the bases or base pairs with respect to a common reference frame, typically the helical axis. The nomenclature and definitions were standardized at an EMBO workshop on DNA curvature and bending (Dickerson et al., 1989). See Figure 7.5.3 for a graphical description of these values. Despite the standard nomenclature and definitions, the precise details of the mathematics were not standardized. Therefore, among the variety of programs commonly used to analyze helicoidal structure, each differs in the details regarding the exact definition of the helical axis, reference frame, and pivot points. Commonly used programs include NEWHELIX by Richard Dickerson, Curves by Heinz Sklenar and Richard Lavery (Lavery and Sklenar, 1988), and programs by Marla Babcock and Wilma Olson (Babcock et al., 1994) among others. The most developed and consistent mathematical treatment of the helicoidal
Pictorial definition of the helicoidal parameters.
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Current Protocols in Nucleic Acid Chemistry
parameters is likely either that of Babcock and Olson or that of Elhassan and Calladine, which is fully reversible (Elhassan and Calladine, 1995). The former has symmetrical definitions on a uniform scale for the various rotations and defines pivot points or axes that minimize mathematically induced artifactual correlations between the various rotational and translational parameters. Despite the advantages of these programs, NEWHELIX and Curves are the most commonly used programs to calculate helicoidal parameters. Although these methods give qualitatively comparable results, care should be taken in quantitative comparison of helicoidal values calculated from different programs due to the sensitivity of the method to definition of the reference frame. This is discussed in more detail in recent work by Lu and Olson (Lu and Olson, 1999; Lu et al., 1999). A further distinction relates to global versus local helicoidal parameters; reference to a local helical axis typically relates to the axis between adjacent base pairs, whereas global helicoidal parameters are in reference to some best-fit global helical axis over the whole duplex. While the global parameters typically lead to more regular values (and less individual variation), the global axis may not be sufficiently determined for small duplexes (such as those with less than a full helical repeat) or distorted duplexes (such as an RNA duplex with a bulge), giving rise to misleading helicoidal parameters. The global axis may therefore not be appropriate. Moreover, given that the overall structure is determined by local interactions between adjacently stacked base pairs, local helicoidal parameters may be more representative. When comparing helicoidal values calculated during modeling to those in the literature, care should be taken to ensure that consistent reference frames (local versus global) and definitions of the values are applied. In addition to standard helicoidal analysis, groove structure is also commonly investigated, such as the relative width and depth of the minor or major groove (see, for example, Stofer and Lavery, 1994). Helicoidal analysis and calculation of the various backbone angles can also be applied to the individual coordinate snapshots (for like conformations) or a representative coordinateaveraged structure generated during modeling, such as from a molecular dynamics or Monte Carlo simulation. Although it is often the case that average backbone angles calculated as the average of individual values for each coordinate snapshot are close to the values determined from the average structure, this is not typically
true for helicoidal parameters, which are very sensitive to the conformation (Cheatham and Kollman, 1997). Modelers should keep in mind that the average structure obtained, such as that seen in crystallography or NMR experiments, hides the detailed dynamics. Moreover, coordinate-averaged conformations are not equivalent to torsion-averaged structures, which do not necessarily give average properties similar to that from the mean of the individual coordinate sets. Therefore, care should be taken in various coordinate comparisons. The common means to compare structures is through the use of best-fit root-mean-squared deviations (RMSd) between the coordinates or torsion angles. This indicator is very useful for determining the degree of similarity between two structures (when the RMSd values are small), but does less well at representing dissimilarity, since small differences in structure can lead to large root-mean-squared differences.
SUMMARY This unit has introduced molecular modeling of nucleic acids on the simplest level. The modeling process can be described in three stages: Generation. Create an initial model either by hand building it based on the molecular connectivity or by obtaining the coordinates from a depository of experimentally derived structures. In the absence of a complete experimental structure, base the structure on known (cannonical) structure and/or use tools (e.g., Nucleic Acid Builder) to complete the model. Evaluation. Is the structure valid? Judge this based on chemical/structural intuition and comparison with experimentally derived structures. The structure can be described in terms of the backbone angles, sugar pucker, glycosidic χ torsion, and helicoidal parameters. Additionally, it is important to check the stereochemistry and hydrogen placement. Manipulation. Coordinate manipulations can be made by simple rotation around chemical bonds. As possible, include some crude representation of the energy to avoid bad steric overlap and unrealistic rotations. Other units delve more deeply into methods for evaluating and manipulating the models and representations of nucleic acids that go beyond the single static gas-phase structure model. This includes a discussion of how to properly represent the long-range electrostatic interactions and how to include some representation of the effect of the environment (solvent and ionic strength effects; see UNIT 7.9). With a more
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realistic representation of the energy (UNIT 7.8), the energy can be used as a guide to suggest coordinate manipulations. Evaluation of the model depends on the reliability of the energy and how the system is represented, coupled with the chemical intuition of the modeler and comparison to experimental data.
Dickerson, R.E., Bansal, M., Calladine, C.R., Diekmann, S., Hunter, W., Kennard, O., von Kitzing, E., Lavery, R., Nelson, H.C.M., Olson, W.K., Saenger, W., Shakked, Z., Sklenar, H., Soumpasis, D.M., Tung, C.S., Wang, A.H., and Zhurkin, V.B. 1989. Definitions and nomenclature of nucleic acid structure components. Nuc. Acids Res. 17:1797-1803.
LITERATURE CITED
Elhassan, M.A. and Calladine, C.R. 1995. The assessment of the geometry of dinucleotide steps in double helical DNA; a new local calculation scheme. J. Mol. Biol. 251:648-664.
Abola, E.E., Bernstein, F.C., Bryant, S.H., Koetzle, T.F., and Weng, J. 1987. Protein Data Bank. In Crystallographic Databases—Information Content, Software Systems, Scientific Applications (F.H. Allen, G. Bergerhoff, and R. Sievers, eds.) pp. 107-132. Data commission of the international union of crystallography, Bonn/Cambridge/Chester. Allen, F.H., Bellard, S., Brice, M.D., Cartright, B.A., Doubleday, A., Higgs, H., Hummelink, T., Hummelink-Peters, B.G., Kennard, O., Motherwell, W.D.S., Rodgers, J.R., and Watson, D.G. 1979. The Cambridge Crystallographic Data Centre: Computer-based search, retrieval, analysis and display of information. Acta Crystallogr. B35:2331-2339. Altona, C. and Sundaralingam, M. 1972. Conformational analysis of the sugar ring in nucleosides and nucleotides. A new description using the concept of pseudorotation. J. Am. Chem. Soc. 94:8205-8212. Arnott, S. and Hukins, D.W. 1972. Optimised parameters for A-DNA and B-DNA. Biochem. Biophys. Res. Commun. 47:1504-1509. Babcock, M.S., Pednault, E.P., and Olson, W.K. 1994. Nucleic acid structure analysis. Mathematics for local Cartesian and helical structure parameters that are truly comparable between structures. J. Mol. Biol. 237:125-156. Berman, H.M., Olson, W.K., Beveridge, D.L., Westbrook, J., Gelbin, A., Demeny, T., Hsieh, S.H., Srinivasan, A.R., and Schneider, B. 1992. The nucleic acid database—A comprehensive relational database of 3-dimensional structures of nucleic acids. Biophys. J. 63:751-759. Brion, P. and Westhof, E. 1997. Hierarchy and dynamics of RNA folding. Annu. Rev. Biophys. Biomol. Struct. 26:113-137. Cheatham, T.E. III. and Kollman, P.A. 1997. Molecular dynamics simulations highlight the structural differences in DNA:DNA, RNA:RNA and DNA:RNA hybrid duplexes. J. Amer. Chem. Soc. 119:4805-4825. Clowney, L., Jain, S.C., Srinivasan, A.R., Westbrook, J., Olson, W.K., and Berman, H.M. 1996. Geometric parameters in nucleic acids: Nitrogenous bases. J. Amer. Chem. Soc. 118:509-518. Molecular Modeling of Nucleic Acid Structure
Cruz-Neira, C., Sandin, D.J., DeFranti, T.A., Kenyon, R.V., and Hart, J.C. 1992. The CAVE: Audio visual experience automatic virtual environment. Commun. ACM 35:65-72.
Erie, D.A., Breslauer, K.J., and Olson, W.K. 1993. A Monte Carlo method for generating structures of short single-stranded DNA sequences. Biopolymers 33:75-105. Gelbin, A., Schneider, B., Clowney, L., Hsieh, S.-H., Olson, W.K., and Berman, H.M. 1996. Geometric parameters in nucleic acids: Sugar and phosphate constituents. J. Amer. Chem. Soc. 118:519529. Hecht, S. 1996. Bioorganic Chemistry: Nucleic Acids (S. Hecht, ed.) pp. 512. Oxford University Press, New York. JCBN. 1983. IUPAC-IUB Joint Commission on Biochemical Nomenclature (JCBN). Abbreviations and symbols for the description of conformations of polynucleotide chains. Recommendations 1982. Eur. J. Biochem. 131:9-15. Lavery, R. and Sklenar, H. 1988. The definition of generalized helicoidal parameters and of axis curvature for irregular nucleic acids. J. Biomol. Struct. Dyn. 6:63-91. Leclerc, F., Srinivasan, J., and Cedergren, R. 1997. Predicting RNA structures: The model of the RNA element binding Rev meets the NMR structure. Folding Des. 2:141-147. Lu, X.-J. and Olson, W.K. 1999. Resolving the discrepancies among nucleic acid conformational analyses. J. Mol. Biol. 285:1563-1575. Lu, X.-J., Babcock, M.S., and Olson, W. K. 1999. Overview of nucleic acid analysis programs. J. Biomol. Struct. Dyn. 16:833-843. Macke, T. and Case, D.A. 1998. Modeling unusual nucleic acid structures. In Molecular Modeling of Nucleic Acids (N.B. Leontis and J. Santa Lucia, eds.) pp. 379-393. ACS, Washington, D.C. Major, F., Turcotte, M., Gautheret, D., LaPalme, G., Fillion, E., and Cedergren, R. 1991. The combination of symbolic and numerical computation for three-dimensional modeling of RNA. Science 253:1255-1260. Nielsen, P.E., Egholm, M., Berg, R.H., and Buchardt, O. 1991. Sequence-selective recognition of DNA by strand displacement with a thy mine-substituted polyamide. Science 254:1497-1500. Olson, W.K. 1996. Simulating DNA at low resolution. Curr. Opin. Struct. Biol. 6:242-256.
7.5.10 Supplement 6
Current Protocols in Nucleic Acid Chemistry
Pearlman, D.A., Case, D.A., Caldwell, J.W., Ross, W.S., Cheatham, T.E., Debolt, S., Ferguson, D., Seibel, G., and Kollman, P. 1995. AMBER, a package of computer programs for applying molecular mechanics, normal mode analysis, molecular dynamics and free energy calculations to simulate the structure and energetic properties of molecules. Comp. Phys. Comm. 91: 1-41. Saenger, W. 1984. Principles of Nucleic Acid Structure. Springer Advanced Texts in Chemistry (C.E. Cantor, ed.). Springer-Verlag, New York. Schlick, T. 1995. Modeling superhelical DNA: Recent analytical and dynamical approaches. Curr. Opin. Struct. Biol. 5:245-252. Schneider, B. and Kabelac, M. 1998. Stereochemistry of binding of metal cations and water to a phosphate group. J. Am. Chem. Soc. 120:161165.
http://honiglab.cpmc.columbia.edu/grasp The home page for the GRASP continuum electrostatics and molecular graphics display code developed by Anthony Nicholls. http://www.lobos.nih.gov/Charmm The CHARMM molecular mechanics/dynamics software home page at the National Institutes of Health. The root of this link discusses the LoBoS “lot’s of boxes on shelves” parallel computer developed at the NIH for use in molecular simulation. http://www.msi.com The home page for Molecular Simulations, which distributed X-Plor and the commercial version of CHARMM. http://www.intsim.com
Schneider, B., Cohen, D.M., Schleifer, L., Srinivasan, A.R., Olson, W.K., and Berman, H.M. 1993. A systematic method for studying the spatial distribution of water molecules around nucleic acid bases. Biophys. J. 65:2291-2303.
The home page for the company Interactive Simulations, which develops the Sculpt software. This program allows real-time molecular modeling with continuous energy minimization as the model is manipulated.
Schneider, B., Neidle, S., and Berman, H.M. 1997. Conformations of the sugar-phosphate backbone in helical DNA crystal structures. Biopolymers 42:113-124.
http://www.ks.uiuc.edu/Research/namd
Stofer, E. and Lavery, R. 1994. Measuring the geometry of DNA grooves. Biopolymers 34:337346. Surles, M.C., Richardson, J.S., Richardson, D.C., and Brooks, F.P. 1994. Scuplting proteins interactively—Continual energy minimization embedded in a graphical modeling system. Protein Sci. 3:198-210. Ulyanov, N.B. and James, T.L. 1995. Statistical analysis of DNA duplex structural features. Methods Enzymol. 261:90-120. Vologodski, A.V. and Cozzarelli, N.R. 1994. Conformational and thermodynamic properties of supercoiled DNA. Annu. Rev. Biophys. Biomol. Struct. 23:609-643.
The home page for the NAMD molecular mechanics/dynamics simulation package developed by Klaus Shulten’s group at the University of Illinois. http://dasher.wustl.edu/tinker The home page for the TINKER molecular mechanics/dynamics software. Includes an extensive list of WWW links to other MM/MD resources.
Model building and analysis tools, nucleic acid nomenclature http://www.scripps.edu/case The home page of Professor David Case at the Scripps Research Institute contains links to the NAB (Nucleic Acid Builder) software and manuals. http://www.eyesopen.com/babel.html
Wang, A.H., Quigley, G.J., Kolpak, F.J., Crawford, J.L., van Boom, J.H., van der Marel, G., and Rich, A. 1979. Molecular structure of a lefthanded double helical DNA fragment at atomic resolution. Nature 283:743-745.
The home page of the Molecular Structure Information Interchange Hub or the program babel developed in Professor Dan Dolata’s group by Pat Walters and Matt Stahl. This program is very useful for interconverting a variety of different molecular modeling program file formats.
INTERNET RESOURCES
http://www.chem.qmw.ac.uk/iupac
Simulation codes
A repository of many of the IUPAC naming conventions. This site has a very nice Web page describing in detail the notation and naming conventions that apply to nucleic acids.
http://www.amber.ucsf.edu/amber The home page for the AMBER suite of programs for molecular mechanics and dynamics. See also the subpage http://www.amber.ucsf.edu/amber/ polyA-polyT/ for a tutorial that describes in detail setting up, equilibrating, and running molecular dynamics simulations using AMBER on a small DNA duplex in solution. http://igc.ethz.ch/gromos The GROMOS molecular mechanics/dynamics software home page.
http://www.sphere.ad.jp/hgs The site for the company that makes the Maruzen physical molecular models (HGS). For protein and nucleic acids, of particular interest is the Maruzen Biochemistry Molecular Models.
Coordinate repositories and information resources http://www.rcsb.org/pdb
Biophysical Analysis of Nucleic Acids
7.5.11 Current Protocols in Nucleic Acid Chemistry
The Protein Data Bank server at the Research Collaboratory for Structural Bioinformatics (Rutgers, SDSC, NIST).
This page, sponsored by the Center for Molecular Modeling at the NIH, provides a nice introduction to macromolecular simulation.
http://ndbserver.rutgers.edu The Nucleic Acid Database server maintained by Helen Berman and others at Rutgers University. http://www.ccl.net/chemistry The computational chemistry list archives. This contains information about a number of modeling programs, conference listings, and job postings. http://cmm.info.nih.gov/intro_simulation/course_ for_html.html
Contributed by Thomas E. Cheatham, III and Bernard R. Brooks National Heart, Lung and Blood Institute, NIH Bethesda, Maryland Peter A. Kollman University of California San Francisco, California
Molecular Modeling of Nucleic Acid Structure
7.5.12 Current Protocols in Nucleic Acid Chemistry
Methods to Crystallize RNA Preparation of suitably large and well-ordered single crystals is usually the rate-limiting step in the determination of the three-dimensional structure of RNAs and their complexes with proteins by X-ray crystallography (reviewed by Holbrook and Kim, 1997). As illustrated by the examples of RNA and RNAprotein complexes in Table 7.6.1, successful crystallization conditions vary greatly for different molecules. A detailed protocol for the crystallization of even a limited set of RNAs cannot be written; therefore, we discuss a variety of experimental considerations relevant to obtaining RNA crystals for structure determination.
OVERVIEW Biological macromolecules, as well as small molecules or simple salts, are crystallized from their metastable supersaturated solutions. When the system relaxes, some of the macromolecules come out of solution, yielding a solid phase which is in equilibrium with a saturated solution. Depending on the properties of the solvated molecule, and the conditions under which supersaturation and relaxation are achieved, the solid phase can consist of amorphous precipitate, a “shower” of microscopic crystals, or ideally, large single crystals. Successful growth of single crystals of macromolecules typically requires fine-tuning a large number of interdependent variables, and is thus often a formidable optimization problem. The problem can be conquered by dividing it into a series of “unit operations” to be optimized in a recursive manner. First, a molecule or molecular complex must be designed or chosen to address the specific structural questions being posed and to have a good chance of being “crystallizable.” Second, the molecule must be purified so that it is covalently homogeneous. Third, conditions must be found under which the molecule is conformationally homogeneous or monodisperse, and has full biochemical activity. Fourth, with a covalently and conformationally homogeneous sample in hand, a series of solution conditions in which the macromolecule might form supersaturated solutions can be screened. Fifth, if crystals result from these screens, they can be characterized and crystallization conditions further optimized. If crystals are not forthcoming, further optimization of preceding steps might be
UNIT 7.6
carried out. In the sections that follow, each of these steps is discussed.
CONSTRUCT DESIGN The first consideration when crystallizing RNA is whether the molecule to be crystallized will be a short RNA (oligonucleotide) duplex, a non-duplex oligonucleotide, a larger RNA molecule, or an RNA-protein complex. Depending on the choice, the experimental strategies, as well as the likelihood of obtaining crystals of satisfactory quality with a given amount of effort, will vary. In general, oligonucleotide duplexes are the easiest to crystallize, and these crystals are most likely to be well ordered at the atomic level. Crystals of large RNAs are not necessarily difficult to obtain, but they are less likely to be well ordered. RNAprotein complexes fall somewhere in between in terms of their crystallizability and typical degree of order.
Oligonucleotide Duplexes The double helix is the basic unit of RNA structure. Because of the strength of basestacking interactions in aqueous solutions, crystals of short nucleic acid duplexes are typically dominated by pseudo-infinite stacks of helices that traverse the entire crystal. Side-byside helical packing involves weaker interactions of backbone, and occasionally base, functional groups (Dickerson et al., 1994; Berman et al., 1996; Holbrook and Kim, 1997). The strategy for the design of crystallizable oligonucleotides is based on these characteristics, and resembles the strategy for the crystallization of DNA-protein complexes (Aggarwal, 1990; Schultz et al., 1990). Duplexes are prepared that incorporate the RNA segment of biological or chemical interest and have additional base pairs on either or both ends to make their length consist of an integral number of helical turns, or some rational fraction thereof. The crystallographically allowed fractions are 1/2, 1/3, 1/4, and 1/6 (Burns and Glazer, 1990). In this way, formation of a pseudo-infinite helix with alignment of successive duplexes is promoted. The pitch of A- or B-form duplexes varies between 10 and 12 bp. The pitch of a crystallization candidate is unknown a priori; thus, the basic strategy for oligonucleotide crystallization is one of length variation.
Contributed by Adrian R. Ferré-D’Amaré and Jennifer A. Doudna Current Protocols in Nucleic Acid Chemistry (2000) 7.6.1-7.6.13 Copyright © 2000 by John Wiley & Sons, Inc.
Biophysical Analysis of Nucleic Acids
7.6.1 Supplement 1
Table 7.6.1
Illustrative Examples of RNA and RNA-Protein Complex Crystals
Space group
Compound “UUUG” RNA dodecamer (24 nt)
P41
RNA “dodecamer” with Shine-Delgarno sequence (24 nt) Loop E “dodecamer” (24 nt)
P1
HIV-1 TAR, RNA fragment (27 nt)
P1
Pseudoknot (28 nt)
P3221
Sarcin-ricin loop (29 nt)
P6122
Hammerhead ribozyme-2′OMe inhibited (41 nt)
P3121
C2
Hammerhead P3221 ribozyme-DNA inhibitor complex (47 nt) 5S rRNA fragment I P6122 (62 nt) Yeast tRNAPhe (76 nt) P21
Crystallization conditions
Tetrahymena group I intron fragment (247 nt)
dmin (Å)b
Reference
30% PEG 4000; 0.2 M NH4 acetate; 0.1 M citrate, pH 5.6; 8 mM spermine 30% MPDc; 400 mM MgCl2; 40 mM cacodylate, pH 6.5; 32°C
2.9
2.6
Baeyens et al. (1995)
2.0
2.8
Schindelin et al. (1995)
5%-25% MPD; 5-25 mM MgCl2; 50 mM cacodylate, pH 6.0; 20°C 20% PEG 4000; 2.5 mM MgCl2; 200 mM NH4Cl; 100 mM CaCl2; 50 mM cacodylate, pH 6.0; 19°C 18% sec-butanol; 5 mM MgCl2; 2 mM spermidine; 100 mM MOPS, pH 7.0; 25°C 3.0-3.2 M (NH4)2SO4; 20 mM MgCl2; 50 mM MOPS, pH 7.0; 1 mM spermine; 2 mM CoCl2; 20°C 23% PEG 6000; 10 mM Mg acetate; 100 mM NH4 acetate; 1 mM spermine; 30 mM cacodylate, pH 6.5; 20°C 1.9-2.2 M (NH4)2SO4; 0-100 mM MgCl2; 0-2 mM spermine; 10 mM cacodylate, pH 6.0; 4°C
2.8
1.5
Correll et al. (1997)
2.2
1.3
Ippolito and Steitz (1998)
2.0
1.6
Su et al. 1999
2.7
2.1
Correll et al. (1998)
3.2
3.1
Scott et al. (1995a)
4.8
2.6
Pley et al. (1993)
3.2
3.0
2.4
2.5
3.0
2.3
Kim (1992); Correll et al. (1997) Ladner et al. (1972); Robertus et al. (1974) Kim et al. (1971); Kim et al. (1974)
5.2
3.0
3.4
2.5
5.0
5.0
1.35 M MgSO4; 20 mM MES, pH 6.4; room temperature 10% dioxane; 5-15 mM MgCl2; 1-2 mM spermine; 4°C Yeast tRNAPhe (76 nt) P212121 10% iso-propanol or 10%-12% MPD; 10 mM MgCl2; 1 mM spermine; 4°-6°C Yeast initiator tRNA P6422 2 M (NH4)2SO4; 5 mM MgCl2; 2 mM spermine (76 nt) Tetrahymena group I intron P4-P6 domain (160 nt)
Vm (Å3/Da)a
P212121 17% MPD; 20 mM MgCl2; 10 mM NaCl; 20 mM cacodylate, pH 6.0; 0.2 mM spermine; 10 M Co (III) hexammine; 30°C P42212 19% MPD; 18 mM MgCl2; 50 mM KCl; 0.5 mM spermine; 50 mM cacodylate, pH 6.0; room temperature
Schevitz et al. (1979); Basavappa and Sigler, (1991) Doudna et al. (1993); Cate et al. (1996)
Golden et al. (1997)
(continued) Methods to Crystallize RNA
7.6.2 Supplement 1
Current Protocols in Nucleic Acid Chemistry
Table 7.6.1
Continued
Compound Tetrahymena group I intron fragment (247 nt) U1A-RNA complex (21 nt + 11 kDa protein) U2B′′U2A′-RNA complex (24 nt + 31 kDa protein) HDV ribozyme-U1A complex (72 nt + 11 kDa protein)
Yeast AspRS-tRNAAsp complex E. coli GlnRS-tRNAGln complex
Space group
Crystallization conditions
P42212 19% MPD; 18 mM MgCl2; 50 mM KCl; 0.5 mM spermine; 50 mM cacodylate, pH 6.0; room temperature P6522 1.8 M (NH4)2SO4; 40 mM Tris⋅Cl, pH 7.0; 5 mM spermine; 20°C P21212 1% PEG 600; 9 mM MgCl2; 50 mM NaCl 0.25 mM spermine; 0.25% octyl glucoside; 50 mM Tris⋅Cl, pH 7.3 14% PEG-MME 2000; 1 mM R32 MgCl2; 250 mM Li2SO4; 100 mM Tris⋅Cl, pH 7.0; 4.0 mM spermine; 0.2 mM Co (III) hexammine; 25°C P212121 62% sat.(NH4)2SO4; 5mM MgCl2; 40 mM Tris⋅Cl, pH 7.5; 4°C C2221 48% sat. (NH4)2SO4; 20 mM MgSO4; 80 mM PIPES, pH 7.4; 4 mM ATP; 17°C
Vm (Å3/Da)a
dmin (Å)b
Reference
5.0
5.0
Golden et al. (1997)
3.2
1.7
Oubridge et al. (1994)
2.7
2.4
Price et al. (1998)
3.2
2.2
Ferré-DAmaré et al. (1998a)
3.8
2.7
Ruff et al. (1988)
3.7
2.5
Rould et al. (1991)
aV
m is the Matthews number (see text). bd min is the best reported diffraction. cMPD is 2-methyl-2,4-pentanediol.
In a standard crystallization project, a series of duplexes are prepared that span the expected length of one (or several) helical turns. For instance, if the segment of interest is 8 bp, then oligonucleotides that range from 9 to 14 bp can be synthesized. The basic length screen should be augmented by varying the termini of the duplexes. Given the expectation that the duplexes will stack end-to-end, this variation is likely to have a strong influence on crystal growth and order. Overhanging ends, whose sequences might be complementary to those at the opposite end of the duplex, can be added to some of the duplexes, and the composition of the ends varied (if they are not part of the sequence whose structure is of interest). This can be achieved simultaneously with a length search by “mixing and matching” oligonucleotides of different lengths. The composition of the ends of oligonucleotides can have nontrivial effects because the stacking energies of different pairs of bases can vary by more than 10 kcal/mol (Saenger, 1984). Table 7.6.1 includes four examples of oligonucleotide duplex crystals. The crystals of
Baeyens et al. (1995), Schindelin et al. (1995), and Correll et al. (1997) are of duplex dodecamers which are almost exactly one helical turn in length. The HIV-1 trans-activation response region (TAR) fragment crystals of Ippolito and Steitz (1998) comprise a 15-nt strand with three bulged residues paired to a 12-nt strand. This effectively results in one complete turn of the atypical helix. The duplexes of Correll et al. (1997) and of Ippolito and Steitz (1998) incorporate 5′ overhangs. The other two are blunt-ended. When designing asymmetric (nonpalindromic) oligonucleotides for crystallization, particular attention should be paid to possible undesired duplexes that the RNAs might form. If significant self-complementarity is present, a homoduplex might unexpectedly be favored over the desired heteroduplex, and crystals of an irrelevant duplex may result. Limiting the lengths of duplexes to integral numbers of helix turns is favorable. The external surface of nucleic acid duplexes is dominated by the periodic negative charges of the phosphate backbone. In unlucky cases, du-
Biophysical Analysis of Nucleic Acids
7.6.3 Current Protocols in Nucleic Acid Chemistry
Supplement 1
plexes whose length is an irrational fraction of helical turns have produced beautiful, well diffracting crystals in which the helices stack head-to-tail in the crystal. On close inspection, some of these consist of infinite pseudo-continuous helices packed side-by-side, out of register with their sequence. If the structure can be solved, the electron density will consist of very well ordered sugar phosphate backbone (which produces the high-resolution diffraction) and poor base density which results from averaging the different bases which comprise the oligonucleotide. See Shah and Brunger (1999) for a detailed analysis of an example.
Complex Oligonucleotides
Methods to Crystallize RNA
Under this heading we include short (∼30 nt or less) RNAs whose biologically relevant structures are not simple duplexes, but have elements such as terminal loops or strand crossovers. These molecules usually comprise some sequences that are self-complementary, and a problem that often arises in crystallization is that the molecules disproportionate into duplexes, with the noncomplementary regions forming noncanonical base pairs. For instance, crystallization of isolated stem-loops incorporating “tetraloop” sequences has often been hampered by the hairpin-loop to duplex equilibrium favoring of the duplex at the high RNA concentrations required for crystallization. The equilibrium probably favors the duplex because, in crystals, stacking occurs at both ends of the molecule, while the stem-loops only stack at one end (e.g., Holbrook et al., 1991). This can be exploited to study the structure of the noncanonical base pairs formed between the “loop” residues of the two strands (e.g., the “UUUG” RNA crystals of Baeyens et al., 1995; Table 7.6.1). Despite this, some hairpin-loops have been crystallized. The 29-nt Sarcin-Ricin Loop crystals of Correll et al. (1998; Table 7.6.1) are stabilized by helix stacking at one end of the hairpin-loop, and by interactions between the loop nucleotides of adjoining molecules in the crystal. Similar interactions are observed in another case where a short stem-loop sequence was successfully crystallized (Perbandt et al., 1998). A 28-nt RNA which forms a classical pseudoknot (two helices stacked end-to-end, held by strand cross-overs in the major and minor grooves) has also been crystallized and its structure has been determined (Su et al., 1999; Table 7.6.1). These successful examples suggest that a reasonable strategy for obtaining crystals of
stem-loops might be to covalently constrain them from unpairing and duplexing. This might be achieved, for example, by introducing a disulfide bond linking the 5′ and 3′ termini (UNIT 5.4), or possibly by circularizing the RNA (Puttaraju and Been, 1992; UNIT 5.2). Stem-loops can also be stabilized by incorporating them into large RNA molecules.
Large RNAs Large “globular” RNAs are composed of double-stranded helices, packed together to form substantial solvent-inaccessible cores (Ferré-D’Amaré and Doudna, 1999). In a manner similar to that with proteins, it is possible to define structural and functional domains in large RNAs. Often, these are better crystallization targets than the parent, multidomain molecules. Proteolysis is a powerful technique for defining the boundaries of protein domains (Cohen, 1996). Some RNA domains have been defined with the help of nucleases (e.g., 5S RNA fragment I; Kim, 1992; Correll et al., 1997; Table 7.6.1). Today, however, multiple sequence and secondary-structure alignments based on natural or artificial phylogenies constitute an expeditious means of searching for domain boundaries. Because the double helices that dominate RNA structure often bring together segments that are distant in primary sequence, secondary structures, based on sequence covariation and/or biochemical probing are, in general, the best starting point for domain-boundary elucidation for large RNAs. The single-domain transfer RNAs were the first nucleic acids whose atomic structures were determined (Kim et al., 1974; Robertus et al., 1974). Other single-domain RNAs that have yielded crystal structures are the hammerhead ribozyme (Pley et al., 1994b; Scott et al., 1995b), Fragment I of the E. coli 5S rRNA (Correll et al., 1997), and a pseudoknot (Su et al., 1999; Table 7.6.1). An extensive body of biochemical information has guided the design of crystallization constructs based on the Group I self-splicing intron of Tetrahymena. This large catalytic RNA can be divided into several structural domains which can be prepared separately, and when mixed, these will assemble to produce a functional RNA (Doudna and Cech, 1995). One of these, the 160-nt P4-P6 domain produced crystals that diffracted X-rays anisotropically to 2.5 and 2.8 Å (Doudna et al., 1993; Cate et al., 1996; Table 7.6.1). P4-P6 is an autonomously folding domain that is the first to acquire its structure in the kinetic folding pathway
7.6.4 Supplement 1
Current Protocols in Nucleic Acid Chemistry
of the intron, and nucleates folding of the complete RNA. Work on larger multidomain constructs of this class of catalytic RNAs has been partially successful (Doudna et al., 1993; Golden et al., 1997, 1998). The design of crystallization constructs of the hammerhead ribozyme was also based on the biochemical knowledge of the system. Pley et al. (1993), and Scott et al. (1995a) varied the placement of the termini (and connectivity) of their molecules in the search for ribozymes that produced well-ordered crystals (Table 7.6.1). The crystallization of the hammerhead ribozymes also illustrates the use of different strategies for capturing one state of a catalytic molecule. While Pley et al. (1993) replaced the strand bearing the 2′- hydroxyl nucleophile with a DNA strand, Scott et al. (1995a) replaced the hydroxy group with a methoxy function. Both groups thus obtained structures of analogs of the ground-state of the catalytic RNA. As with proteins, binding of inhibitors and cofactors should be considered from the outset when crystallizing RNAs the interact with such molecules. In designing crystallizable constructs of an RNA domain, or a multidomain RNA, length and end variation as practiced with oligonucleotide duplexes (Anderson et al., 1996) can be combined with other strategies, such as the circular permutation employed for the hammerhead ribozymes. In both crystal forms of the hammerhead, the blunt double-stranded ends of symmetry-related molecules stack on each other, and tetraloops placed at the distal ends of duplex stems also make various crystal contacts (Pley et al., 1994a; Scott et al., 1995b). Crystallization of the same protein from several different species is a time-honored practice among crystallographers. Proteins from different sources often crystallize differently because the most phylogenetically variable residues in a folded macromolecule will lie on their surface. Surface residues are precisely those that will affect crystallization, as they will be responsible for crystal contacts and determine solubility. The same consideration applies to RNAs. Most of the variation in sequence between related RNAs occurs within duplex regions; some exposed variable residues will reside in helix terminal loops and can be varied to generate additional surface diversity (Golden et al., 1997). Surface variation can be taken one step further by incorporating crystallization modules into target RNAs (Ferré-D’Amaré et al., 1998b). These are moieties which are engi-
neered into the sequence of the target RNAs in solvent-exposed portions so that they are available to make intermolecular interactions that can lead to crystal formation. One such crystallization module involves a GAAA tetraloop placed at the end of a duplex harboring its 11-nt receptor sequence. Since the two elements are stacked coaxially, they cannot dock intramolecularly, but can interact to pack neighboring molecules together. The module should be placed in a solvent-exposed portion of the target RNA, in a location that does not affect the activity (and by inference, the structure) of the parent molecule. In employing this technique, the placement of the crystallization module and the number of “spacer” nucleotides can both be varied (similar to the length variation for oligonucleotides) to generate a series of related RNAs that can be subjected to crystallization trials. Example RNA constructs based on this approach are shown in Figure 7.6.1.
RNA-Protein Complexes Many biologically important RNAs carry out their functions in complexes with proteins. Crystals have been obtained both of complexes of unmodified full-length proteins and RNAs, and of engineered protein and RNA domains. Aminoacyl tRNA synthetase–tRNA complexes are examples of the former (e.g., Ruff et al., 1988; Rould et al., 1991; Table 7.6.1). In the two examples of an engineered complex in Table 7.6.1 (Oubridge et al., 1994; Price et al., 1998), both the proteins and the RNAs were optimized. Successful crystallization required careful definition of protein domain boundaries, mutational modification of the solvent-exposed surface of the protein, and a length search on the RNA moiety (Oubridge et al., 1995). RNA-binding proteins can be employed as part of a second type of crystallization module. Crystals of a hepatitis delta virus (HDV) ribozyme which diffract X-rays to 2.2 Å were obtained by engineering the catalytic RNA so that its solvent-exposed, functionally dispensable stem-loop P4 was replaced by a high-affinity binding site for the U1A protein RNA-binding domain (Ferré-D’Amaré et al., 1998a; Table 7.6.1). The presence of the bound protein greatly facilitated crystallization, presumably because the RNA-protein complex has a larger variety of surface functional groups available for making crystal contacts than the naked RNA. In searching for good cocrystals, the authors made a series of constructs differing in the length of the spacer helix between the crystallization module and the catalytic core of the
Biophysical Analysis of Nucleic Acids
7.6.5 Current Protocols in Nucleic Acid Chemistry
Supplement 1
Figure 7.6.1 Examples of constructs of a genomic hepatitis delta virus (HDV) ribozyme engineered for crystallization. The solvent-exposed P4 stem is dispensable for ribozyme function, and therefore unlikely to participate in the architecture of the catalytic core. Engineered crystallization constructs comprised a variety of substitute stem-loops incorporating a GAAA tetraloop, a tetraloop receptor (TR, in both orientations) and varying numbers of spacer base pairs (lower right). Additional variation was produced by progressive shortening of the 3′ terminus of the RNA (upper right). See text and Ferré-D’Amaré et al. (1998b) for details.
ribozyme, and also varied the termini of the ribozyme (Ferré-D’Amaré and Doudna, 2000). This strategy has been extended to other RNA targets.
Construct Design and the Phase Problem
Methods to Crystallize RNA
Once well-ordered crystals have been produced, the phase problem must be overcome in order to obtain the three-dimensional structure. Thought should be given to heavy-atom derivative preparation when designing constructs. If the structure is to be solved either by the multiple isomorphous replacement (MIR) or multiwavelength anomalous diffraction (MAD) method, heavy atoms must be introduced to the RNA or RNA-protein complex. Traditionally, heavy atom derivatives were obtained by “soaking and praying,” that is, by placing crystals in solutions of various heavy atom compounds at different concentrations, and then collecting X-ray data on the crystals to determine the effect of the soaks (Holbrook and Kim, 1985; Petsko, 1985). However, recombinant technology has made it possible to introduce heavy
atoms covalently into the target molecules in advance of crystallization. For proteins and RNA-protein complexes, biosynthetic substitution of methionine with selenomethionine often provides excellent scatterers for MAD phasing (Doublié, 1997; Smith and Thompson, 1998). If an RNA or a segment of RNA is prepared by solid-phase synthesis, it is possible to substitute uracil with 5-bromouracil or cytosine with 5-bromocytosine (for instance, Correll et al., 1998; Ippolito and Steitz, 1998). Bromine is also an effective scatterer for MAD phasing. Synthetic introduction of phosphorothioates or thiols (which can later be bound to mercury or other heavy metals) into RNA is another possible approach. For large RNAs that bind tightly to either a small nucleic acid or a small molecule, heavy atom substitution of the ligand provides another means of obtaining a derivative. If a large RNA can be split into fragments without affecting biochemical activity and crystallizability, a small fragment could be prepared by chemical synthesis and modified to incorporate heavy atoms.
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SAMPLE PREPARATION AND ANALYSIS The vast majority of RNAs that are subjected to crystallization are prepared in vitro either by stepwise solid-phase chemical synthesis or by runoff transcription, using bacteriophage RNA polymerases. Some RNAs have been isolated from cells, and when post-transcriptional modifications are functionally and structurally important, this might be the method of choice. Synthesis and purification of RNA are covered in APPENDIX 3C. Covalent homogeneity of starting materials is important for crystallization. In the case of synthetic RNA, substantial effort should be devoted to removing shorter impurities resulting from incomplete coupling. These can be removed either by preparative gel electrophoresis under denaturing conditions or by chromatographic methods. If phosphorothioates are being synthetically incorporated into the RNA, it might be desirable to resolve the diasteromers. Halogenated RNAs can be separated from molecules that have lost the halogen by anionexchange chromatography at high pH by taking advantage of the perturbed pKa of N3 of bromouracil. Runoff transcription with phage RNA polymerases results in molecules that are heterogeneous at their 3′, and sometimes their 5′ termini. In the case of shorter molecules, these impurities can be resolved by electrophoresis or chromatography. For larger RNAs (∼40 nt or longer), these methods cease to work on a preparative scale. For these, cis- or trans-acting ribozymes can be employed to homogenize the termini (Price et al., 1995; Ferré-D’Amaré and Doudna, 1996). Another possibility is to cleave the transcripts using RNase H and a guide oligonucleotide (Lapham and Crothers, 1996). If the crystallization target is a ribonucleoprotein complex, then attention should be paid to preparing a nuclease-free protein. Crystallization involves incubation of the proteinRNA complex for weeks to months and even a few contaminating nuclease molecules can hydrolyze large quantities of RNA in this time. The suitability of a protein preparation for cocrystallization with RNA should be evaluated by incubating the complex for a period of at least several days at room temperature. Once purified, samples should be analyzed (i.e., electrophoresis, chromatography, or mass spectrometry), not only for covalent homogeneity, but also for conformational homogeneity. Since most RNAs are purified by denaturing methods, they often need to be refolded (“an-
nealed”). This is usually accomplished by heating the RNA in the presence of divalent cations and buffer, then slowly cooling the solution to ambient temperature or lower. The success of a particular annealing protocol can be monitored by native gel electrophoresis, or physical methods such as dynamic light scattering (FerréD’Amaré and Doudna, 1997). In the case of larger RNAs with a well characterized biochemical activity, specific activity measurements provide a stringent indication of correct folding. For oligonucleotides, it is imp o r t a n t t o ascertain that the desired oligomerization state is the prevalent one, especially at high concentrations. Conformational homogeneity of the macromolecular sample is often an excellent predictor of crystallizability (D’Arcy, 1994; FerréD’Amaré and Burley, 1997). If an RNA or RNA-protein complex is not monodisperse, annealing conditions can be further explored, ligands or inhibitors added, or the construct reengineered to increase homogeneity. Molecules that are polydisperse may crystallize. However, the amount of effort that must be expended to get crystalline material usually far exceeds that required to find a different construct that is monodisperse.
CRYSTALLIZATION TRIALS Once pure, monodisperse, and fully biochemically active preparations are available in sufficient quantity, crystallization trials can begin. Depending on the solubility of the target molecule or complex, crystallization trials are carried out at concentrations of 1 to 20 mg/mL. There are a variety of experimental setups for screening crystallization conditions (reviewed in McPherson, 1990, 1999; Chayen et al., 1992; Ducruix and Geige, 1992; Weber, 1997). Of these, those favored by the authors are hanging-drop vapor diffusion and microbatch crystallization under oil for initial screens, because they require relatively small amounts of sample and can be set up quickly. The most efficient way to search for crystallization conditions is by means of sparse matrix screens or incomplete factorial screens. Several such screens have been formulated. In the authors’ laboratories, the screens of Doudna et al. (1993) and Scott et al. (1995a) are routinely employed for RNAs and the screen of Jancarik and Kim (1991), augmented with 1 mM spermine and varying concentrations of Mg2+, is used for protein-RNA complexes. Other published screens include those of Carter and Carter (1979), Cudney et al. (1994), and Berger et
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al. (1996). Because the goal of the crystallization setups is to achieve supersaturation, the concentration of the macromolecule is important. Generally, for initial screens, the authors consider it satisfactory when about one third of conditions produce precipitate within a few minutes of being mixed with the screen solution. If fewer conditions precipitate, the concentration of macromolecule should be raised, and vice versa. As can be seen from Table 7.6.1, RNAs and RNA-protein complex crystals have been obtained from three broad classes of solutions: organic solvent-water mixtures, salt–polyethylene glycol (PEG) mixtures, and concentrated salt solutions. In screening for crystallization conditions for RNA, the authors have found that the sparse matrices of Scott et al. (1995a) and Doudna et al. (1993) complement each other well; the former is rich in salt/PEG mixtures, while the latter samples many organic solvents. Because of the polyanionic nature of RNA, all of the crystallization conditions include some cations, most often Mg2+ and spermine. Spermine has been employed for the overwhelming majority of successful RNA and RNA-protein complex crystallizations; it should initially be included in all screens. Temperature has a profound effect on the crystallization behavior of macromolecules. At the very least, replicate screens should be set up at 4°C and room temperature (constant-temperature incubators at 20° to 25°C are preferable). Several RNAs crystallize better at temperatures higher than 30°C. Two examples in Table 7.6.1 are the Shine-Delgarno dodecamer of Schindelin et al. (1995) and the P4-P6 domain of Doudna et al. (1993). Replicate setups at 10° to 15°C and at 30°C are strongly recommended.
OPTIMIZATION OF CRYSTALLIZATION CONDITIONS
Methods to Crystallize RNA
In the authors’ experience, provided that construct design has been successful in producing monodisperse samples that have molecular surfaces favoring intermolecular interactions (such as the ends of oligonucleotide duplexes, or crystallization modules), at least some conditions in a crystallization screen should produce crystalline material within days to weeks. In extreme cases, crystals become visible upon minutes of setting up the screens. Most crystals (except those in cubic space groups) are optically birefringent in some directions. Therefore crystallization trials are examined through a stereomicroscope under polarizers crossed so that the field is dark and
birefringent objects show up brightly. If crystals have appeared, then the immediate goal is to characterize them to decide whether or not to thoroughly optimize the crystallization conditions. If the crystals are very small, some improvement of growth conditions might be needed before any characterization can be carried out. The first question to be asked of any crystals is whether they are comprised of the RNA or RNA-protein complex of interest. If crystalline material is found in a vapor diffusion setup, one should immediately inspect the reservoir solution; if the setup was not airtight and has dried out, crystals might also be found in the reservoir. Macromolecular crystals are much more fragile than crystals of simple salts because they are typically comprised of 50% or more solvent by volume, being, in this sense, better described as oriented gels rather than crystals in the minerological sense. Therefore, if a crystal is poked with a scalpel or a thin wire and shatters easily, it is likely to be macromolecular. When sufficiently large crystals (∼50 µm) become available, they should be washed free of mother liquor (which contains macromolecule in equilibrium with the crystal), dissolved, and analyzed by either electrophoretic, chromatographic, or mass spectrometric methods. Crystals can usually be washed with the screen solution modified to contain an additional 10% to 30% of the precipitant, be it the organic solvent, PEG, salt, or polyamine. Before washing the crystals, as much mother liquor as possible should be removed; this should be analyzed as well to determine if there has been any nucleolytic or proteolytic degradation. The crystals should be washed by adding 5 to 10 µL of wash solution and pipetting it away. Washing can be repeated 2 to 3 times. The wash solutions should be analyzed as well, to ensure that the crystals were not dissolving during the process. Finally, the crystals can be dissolved in deionized water or dilute EDTA. Analysis of the resulting solution will demonstrate the composition of the crystals. A rough estimate of how much RNA or (RNA and protein) to expect to find in the last solution can be gained from calculating the volume of the crystal or crystals being dissolved (based on their exterior dimensions), and then assuming a typical solvent content (∼60%) and a density of ∼1 g/mL. If much less macromolecule than expected is found, and the crystals were not dissolving during the wash process, then the macromolecule in the last solution could be residual contamination from the mother liquor.
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Crystallization consists of two distinct steps: nucleation and growth. In the first, thermodynamically unstable clusters of possibly as few as 100 molecules form. Once these nuclei grow beyond a certain point, growth becomes energetically favored (Stura and Wilson, 1990). Conditions under which nucleation and growth are favored are not necessarily the same, and can be experimentally separated. Seeding techniques are a powerful way to dissociate nucleation from growth, allowing conditions for both to be optimized separately. The techniques involved in protein crystal seeding have been reviewed elsewhere (Stura and Wilson, 1990) and apply to RNA and RNA-protein crystallization as well. Optimization of crystallization conditions involves (1) determining what variables are relevant to nucleation and growth, and (2) what ranges and combination of ranges are optimal. The screen conditions under which crystals were obtained are the starting point for determining these parameters. Initially, it is important to sample broadly, because sparse matrices provide only a limited sampling of parameter space. For instance, if crystals grew in high concentrations of ammonium sulfate, followup screens should explore the effectiveness of lithium or magnesium sulfate in producing crystals. If crystals appear in conditions with PEG of average molecular weight 4000 (PEG 4000), then the effect of PEGs as well as PEG derivatives (e.g., polyethylene glycol monomethyl ethers; PEG-MMEs) of different average molecular weights should be explored. If these follow-up screens demonstrate that crystal growth is very sensitive to divalent cation concentration, then a variety of these, such as magnesium, manganese, calcium, barium, strontium, and cadmium, as well as metal hexammines, which mimic solvated magnesium, should be investigated. If calcium ions appear to be important for crystal formation, then lanthanides, which have similar coordination properties, should be tested. Temperature and pH should be varied in small steps to evaluate their effects on crystal growth. Once the variables that are relevant to crystal formation have been determined, their optimal ranges and interactions must be investigated. This can be achieved by analytical techniques (i.e., response surface methods) such as those described by Carter (1997). However, the priority after discovering a new crystal form is not to find the optimal conditions for growth, but rather to obtain a few crystals which are large enough (∼100 µm on the small dimension) to be placed in an X-ray beam to determine their
degree of order. Because many RNA crystal forms do not diffract to high enough resolution to yield biochemical insights (see, for example, Ferré-D’Amaré et al., 1998b), too much effort should not be expended on a crystal form before it is known whether thorough optimization is warranted. Ultimately, the most important property of crystals is how well they diffract X-rays. In order for a structure to yield biochemical insight, it is usually necessary to have X-ray data extending to at least 3.5 Å. A well refined structure at this resolution will have an average precision of atomic coordinates of the order of 0.3 to 0.5 Å, so that only the approximate location of atoms can be inferred from it. At a resolution of 2.0 Å, the coordinate error will typically be of the order of 0.1 to 0.2 Å, so that the presence of hydrogen bonds, for example, can be ascertained. At a resolution of 1.5 Å, details of hydration and coordination become apparent (Richardson and Richardson, 1985; Swanson, 1988). The highest resolution to which a crystal form will ultimately diffract, when large (∼0.3 mm3) specimens are exposed to very bright and well collimated X-rays from a synchrotron radiation source under cryogenic conditions, is difficult to estimate from initial diffraction measurements. If after several rounds of optimization the flash-cooled (see below) crystals fail to diffract beyond ∼5 Å with a laboratory X-ray source with focusing mirrors, it is advisable to look for a different crystal form. The analysis of X-ray diffraction data falls beyond the scope of this unit (see, for instance, Blundell and Johnson, 1976; Drenth, 1994). However, three pieces of information that result from a preliminary analysis are listed for different crystal forms in Table 7.6.1. These are the space group, or the symmetry rules that the crystals obey (Wukovitz and Yeates, 1995), the Matthews number (Vm), and the best diffraction (dmin). The space group (technically, the point group), the unit cell dimensions, the molecular weight of the macromolecule, and the number of molecules in the asymmetric unit (Z), yield the volume occupied per dalton of macromolecule, or Vm (Matthews, 1985). If the density of solvent is assumed to be that of water, then it can be shown (Matthews, 1968) that the volume fractions of the crystal occupied by macromolecule and solvent are: _
Vmacromolecule = 1.66v /Vm
and Vsolvent = 1 − Vmacromolecule
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Methods to Crystallize RNA
_ where v is the partial specific volume of the macromolecule. This is ∼0.74 and 0.6 mL/g for proteins and RNA, respectively, so that for an RNA crystal, the solvent content is ∼1 − (1/Vm). Table 7.6.1 shows that Vm of RNA and RNAprotein complex crystals varies between 2.0 and 5.2. The best diffraction seen from a given crystal form correlates only loosely with solvent content (the regression coefficient for these 19 crystal forms is 0.62). Given the typical range of Vm, the number of molecules per asymmetric unit of a new crystal form can be guessed from the unit cell dimensions, the space group, and the molecular weight of the macromolecule. If the densities of the crystals and the crystallization solution are measured directly, then Z can be established rigorously (Matthews, 1985). After measuring the first data set from a new crystal form, especially an oligonucleotide duplex, intensity statistics should be calculated to ascertain that the crystals do not suffer from twinning. See Yeates (1997) and Shah and Brunger (1999) for details. Once a promising crystal form has been found from screens, a thorough optimization of growth conditions is warranted. In addition to a variety of possible additives (see above), seeding, analytical optimization methods, and modification of the crystallization setup should be attempted. For example, it is possible to change the crystallization kinetics of vapor diffusion setups by judicious use of oil (Chayen, 1997). Some crystal forms will grow better in a batch setup, or by microdialysis, rather than by vapor diffusion. The effects of varying the concentration of macromolecule and the kinetics of equilibration can be explored quickly in a diffusion setup by mixing different ratios of macromolecule stock and reservoir solutions when preparing the drops. If analysis of the RNA (and if present, protein) in the crystal reveals nicking of the macromolecules, and the crystals show enrichment for a nicked species relative to mother liquor, it may be worthwhile to characterize the position of the nick. The shortened or nicked species may be a better crystallization construct. Furthermore, if there is enrichment in the crystals of some modified form of the macromolecule, then recrystallization may result in better ordered crystals. Macromolecular crystals are damaged by exposure to X-radiation. This damage manifests itself as a decrease in intensity of the higher resolution reflections, and an increase in the mosaicity (Drenth, 1994) of the crystals.
Flash-cooling the crystals, so that the aqueous solutions surrounding the crystals and in the solvent channels of the crystals form an amorphous glass (Dubochet et al., 1988), permits data collection to be carried out at cryogenic temperatures (∼100 K) which minimizes radiation damage. Most mother liquors will not form a glass when flash cooled; therefore, crystals must be transferred to a “cryoprotectant” solution. Optimization of the composition of this solution and the transfer protocol are important in obtaining the highest possible resolution diffraction from a given crystal form. This has been reviewed elsewhere (Rodgers, 1997; Harp et al., 1998). The authors’ experience suggests that the cryoprotectants which have been successful for proteins and protein-DNA complexes will often work for RNAs and RNA-protein complexes.
CONCLUSION Successful crystallization is an iterative optimization process. In this unit we have emphasized construct design, because in our experience this is the single most important factor in obtaining well-ordered RNA and RNA-protein complex crystals.
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Carter C.W., Jr. and Carter, C.W. 1979. Protein crystallization using incomplete factorial experiments. J. Biol. Chem. 254:12219-12223. Cate, J.H., Gooding, A.R., Podell, E., Zhou, K., Golden, B.L., Kundrot, C.E., Cech, T.R., and Doudna, J.A. 1996. Crystal structure of a group I ribozyme domain: Principles of RNA packing. Science 273:1678-1685. Chayen, N.E. 1997. The role of oil in macromolecular crystallization. Structure 5:1269-1274. Chayen, N., Stewart, P.D.S., and Blow, D.M. 1992. Microbatch crystallization under oil—a new technique allowing many small-volume crystallization trials. J. Cryst. Growth122:176-80. Cohen, S.L. 1996. Domain elucidation by mass spectrometry. Structure 4:1013-1016. Correll, C.C., Freeborn, B., Moore, P.B., and Steitz, T.A. 1997. Metals, motifs, and recognition in the crystal structure of a 5S rRNA domain. Cell 91:705-712. Correll, C.C., Munishkin, A., Chan, Y.-L., Ren, Z., Wool, I.G., and Steitz, T.A. 1998. Crystal structure of the ribosomal RNA domain essential for binding elongation factors. Proc. Natl. Acad. Sci. U.S.A. 95:13436-13441. Cudney, B., Patel, S., Weisgraber, K., Newhouse, Y., and McPherson, A. 1994. Screening and optimization strategies for macromolecular crystal growth. Acta Crystallogr. Sect. D Biol. Crystallogr. 50:414-423. D’Arcy, A. 1994. Crystallizing proteins—a rational approach? Acta Crystallogr. Sect. D Biol. Crystallogr. 50:469-471. Dickerson, R.E., Goodsell, D.S., and Neidle, S. 1994. “... the tyranny of the lattice ...”. Proc. Natl. Acad. Sci. U.S.A. 91:3579-3583. Doublié, S. 1997. Preparation of selenomethionyl proteins for phase determination. Methods Enzymol. 276:523-530. Doudna, J.A. and Cech, T.R. 1995. Self-assembly of a group I intron active site from its component tertiary structural domains. RNA 1:36-45. Doudna, J., Grosshans, C., Gooding, A., and Kundrot, C.E. 1993. Crystallization of ribozymes and small RNA motifs by a sparse matrix approach. Proc. Natl. Acad. Sci. U.S.A. 90:78297833. Drenth, J. 1994. Principles of Protein X-ray Crystallography. Springer-Verlag, New York. Dubochet, J., Adrian, M., Chang, J.-J., Homo, J.-C., Lepault, J., McDowall, A.W., and Schultz, P. 1988. Cryo-electron microscopy of vitrefied specimens. Q. Rev. Biophys.21:129-228. Ducruix, A. and Geigé, R. (eds.) 1992. Crystallization of Nucleic Acids and Proteins: A Practical Approach. Oxford University Press, Oxford. Ferré-D’Amaré, A.R. and Burley, S.K. 1997. Dynamic light scattering in evaluating crystallizability of macromolecules. Methods Enzymol. 276:157-166.
Ferré-D’Amaré, A.R. and Doudna, J.A. 1996. Use of cis- and trans-ribozymes to remove 5′ and 3′ heterogeneities from milligrams of in vitro transcribed RNA. Nucl. Acids Res. 24:977-978. Ferré-D’Amaré, A.R. and Doudna, J.A. 1997. Establishing suitability of RNA preparations for crystallization. Determination of polydispersity. In Ribozyme Protocols (P.C. Turner, ed.) pp. 371-378. Humana Press, Totowa, N.J. Ferré-D’Amaré, A.R. and Doudna, J.A. 1999. RNA folds: Insights from recent crystal structures. Annu. Rev. Biophys. Biomol. Struct. 28:57-73. Ferré-D’Amaré, A.R. and Doudna, J.A. 2000. Crystallization and structure determination of a hepatitis delta virus ribozyme: use of the RNA-binding protein U1A as a crystallization module. J. Mol. Biol. 295:541-556. Ferré-D’Amaré, A.R., Zhou, K., and Doudna, J.A. 1998a. Crystal structure of a hepatitis delta virus ribozyme. Nature 395:567-574. Ferré-D’Amaré, A.R., Zhou, K., and Doudna, J.A. 1998b. A general module for RNA crystallization. J. Mol. Biol. 279:621-631. Golden, B.L., Podell, E.R., Gooding, A.R., and Cech, T.R. 1997. Crystals by design: A strategy for crystallization of a ribozyme derived from the Tetrahymena group I intron. J. Mol. Biol. 270:711-723. Golden, B.L., Gooding, A.R., Podell, E.R., and Cech, T.R. 1998. A preorganized active site in the crystal structure of the Tetrahymena ribozyme. Science 282:259-264. Harp, J.M., Timm, D.E., and Bunick, G.J. 1998. Macromolecular crystal annealing: Overcoming increased mosaicity associated with cryocrystallography. Acta Cryatallogr. Sect. D Biol. Crystallogr. 54:622-628. Holbrook, S.R. and Kim, S.-H. 1985. Crystallization and heavy-atom derivatives of polynucleotides. Methods Enzymol. 114:167-176. Holbrook, S.R. and Kim, S.-H. 1997. RNA crystallography. Biopolymers 44:3-21. Holbrook, S.R., Cheong, C., Tinoco, I.J., and Kim, S.-H. 1991. Crystal structure of an RNA double helix incorporating a track of non-Watson-Crick base pairs. Nature 353:579-581. Ippolito, J.A. and Steitz, T.A. 1998. A 1.3 Å resolution crystal structure of the HIV-1 trans-activation response region RNA stem reveals a metal ion-dependent bulge conformation. Proc. Natl. Acad. Sci U.S.A. 95:9819-9824. Jancarik, J. and Kim, S.-H. 1991. Sparse matrix sampling: A screening method for crystallization of proteins. J. Appl. Crystallogr.24:409-411. Kim, J.L. 1992. X-ray crystallographic studies of a ribonuclease resistant fragment of E. coli 5S RNA. Ph.D. Dissertation, Yale University, New Haven, Conn. Kim, S.-H., Quigley, G., Suddath, F.L., Rich, A. 1971. High-resolution X-ray diffraction patterns of crystalline transfer RNA that show helical regions. Proc. Natl. Acad. Sci. U.S.A. 68:841-845.
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Kim, S.-H., Suddath, F.L., Quigley, G.J., McPherson, A., Sussman, J.L., Wang, A.H.J., Seeman, N.C., and Rich, A. 1974. Three-dimensional tertiary structure of yeast phenylalanine transfer RNA. Science 185:435-440. Ladner, J.E., Finch, J.T., Klug, A., and Clark, B.F.C. 1972. High-resolution X-ray diffraction studies on a pure species of transfer RNA. J. Mol. Biol. 72:99-101. Lapham, J. and Crothers, D.M. 1996. RNase H cleavage for processing of in vitro transcribed RNA for NMR studies and RNA ligation. RNA 2:289-296. Matthews, B.W. 1968. Solvent content of protein crystals. J. Mol. Biol. 33:491-497. Matthews, B.W. 1985. Determination of protein molecular weight, hydration and packing from crystal densities. Methods Enzymol. 114:176-187. McPherson, A. 1990. Current approaches to macromolecular crystallization. Eur. J. Biochem. 189:123. McPherson, A. 1999. Crystallization of Biological Macromolecules. Cold Spring Harbor Press, Cold Spring Harbor, N.Y. Oubridge, C., Ito, N., Evans, P.R., Teo, C.-H., and Nagai, K. 1994. Crystal structure at 1.92 Å resolution of the RNA-binding domain of the U1A spliceosomal protein complexed with an RNA hairpin. Nature 372:432-438. Oubridge, C., Ito, N., Teo, C.-H., Fearnley, I., and Nagai, K. 1995. Crystallization of RNA-protein complexes II. The application of protein engineering for crystallization of the U1A proteinRNA complex. J. Mol. Biol. 249:409-423. Perbandt, M., Nolte, A., Lorenz, S., Bald, R., Betzel, C., and Erdmann, V.A. 1998. Crystal structure of domain E of Thermus flavus 5S RNA: A helical RNA structure including a hairpin loop. FEBS Lett. 429:211-215. Petsko, G.A. 1985. Preparation of isomorphous heavy atom derivatives. Methods Enzymol. 114:147-156. Pley, H.W., Lindes, D.S., DeLuca-Flaherty, C., and McKay, D.B. 1993. Crystals of a hammerhead ribozyme. J. Biol. Chem. 268:19656-19658. Pley, H.W., Flaherty, K.M., and McKay, D.B. 1994a. Model for an RNA tertiary interaction from the structure of an intermolecular complex between a GAAA tetraloop and an RNA helix. Nature 372:111-113. Pley, H.W., Flaherty, K.M., and McKay, D.B. 1994b. Three-dimensional structure of a hammerhead ribozyme. Nature 372:68-74. Price, S.R., Ito, N., Oubridge, C., Avis, J.M., and Nagai, K. 1995. Crystallization of RNA-protein complexes I. Methods for the large-scale preparation of RNA suitable for crystallographic studies. J. Mol. Biol. 249:398-408.
Methods to Crystallize RNA
Price, S.R., Evans, P.R., and Nagai, K. 1998. Crystal structure of the spliceosomal U2B′′U2A′ protein complex bound to a fragment of U2 small nuclear RNA. Nature 394:645-650.
Puttaraju, M. and Been, M.D. 1992. Group I permuted intron-exon (PIE) sequences self-splice to produce circular exons. Nucl. Acids Res. 20:5357-5364. Richardson, J.S. and Richardson, D.C. 1985. Interpretation of electron density maps. Methods Enzymol. 115:189-206. Robertus, J.D., Ladner, J.E., Finch, J.T., Rhodes, D., Brown, R.S., Clark, B.F.C., and Klug, A. 1974. Structure of yeast phenylalanine tRNA at 3Å resolution. Nature 250:546-551. Rodgers, D.W. 1997. Practical cryocrystallography. Methods Enzymol. 276:183-203. Rould, M.A., Perona, J.J., and Steitz, T.A. 1991. Structural basis of anticodon loop recognition by glutaminyl-tRNA synthetase. Nature 352:213-218. Ruff, M., Mitschler, A., Cavarelli, J., Giegé, R., Mikol, V., Thierry, J.C., Lorber, B., and Moras, D. 1988. A high resolution diffracting crystal form of the complex between yeast tRNAAsp and aspartyl-tRNA synthetase. J. Mol. Biol. 201:235236. Saenger, W. 1984. Principles of Nucleic Acid Structure. Springer-Verlag, New York. Schevitz, R.W., Podjarny, A.D., Krishnamachari, N., Hughes, J.J., Sigler, P.B., and Sussman, J.L. 1979. Crystal structure of a eukaryotic initiator tRNA. Nature 278:188-190. Schindelin, H., Zhang, M., Bald, R., Fürste, J.-P., Erdmann, V.A., and Heinemann, U. 1995. Crystal structure of an RNA dodecamer containing the Escherichia coli Shine-Delgarno sequence. J. Mol. Biol. 249:595-603. Schultz, S.C., Shields, G.C., and Steitz, T.A. 1990. Crystallization of Escherichia coli catabolite gene activator protein with its DNA binding site the use of modular DNA. J. Mol. Biol. 213:159166. Scott, W.G., Finch, J.T., Grenfell, R., Fogg, J., Smith, T., Gait, M.J., and Klug, A. 1995a. Rapid crystallization of chemically synthesized hammerhead RNA’s using a double screening procedure. J. Mol. Biol. 250:327-332. Scott, W.G., Finch, J.T., and Klug, A. 1995b. The crystal structure of an all-RNA hammerhead ribozyme: A proposed mechanism for RNA catalytic cleavage. Cell 81:991-1002. Shah, S.A. and Brunger, A.T. 1999. The 1.8 Å crystal structure of a statically disordered 17 base-pair RNA duplex: principles of RNA crystal packing and its effect on nucleic acid structure. J. Mol. Biol. 285:1577-88. Smith, J.L. and Thompson, A. 1998. Reactivity of selenomethionine--dents in the magic bullet? Structure 6:815-819. Stura, E.A. and Wilson, I.A. 1990. Analytical and production seeding techniques. Methods 1:38-49. Su, L., Chen, L., Egli, M., Berger, J.M., and Rich, A. 1999. Minor groove RNA triplex in the crystal structure of a viral pseudoknot involved in ribosomal frameshifting. Nature Struct. Biol. 6:285292.
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Swanson, S.M. 1988. Effective resolution of macromolecular X-ray diffraction data. Acta Crystallogr. A44:437-442. Weber, P.C. 1997. Overview of protein crystallization methods. Methods Enzymol. 276:13-22. Wukovitz, S.W. and Yeates, T.O. 1995. Why protein crystals favor some space-groups over others. Nature Struct. Biol. 2:1062-1067.
Contributed by Adrian R. Ferré-D’Amaré Fred Hutchinson Cancer Research Center Seattle, Washington Jennifer A. Doudna Yale University New Haven, Connecticut
Yeates, T.O. 1997. Detecting and overcoming crystal twinning. Methods Enzymol. 276:344-358.
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Recent Advances in RNA Structure Determination by NMR Recent progress in RNA structure determination by nuclear magnetic resonance (NMR) spectroscopy has led to the solution of the structure of a number of new RNA and RNAligand complexes. In addition, recent advances in isotopic labeling and the introduction of through-bond experiments to assign bases and phosphate backbones raise the hope of solving even larger and more complicated structures. The limiting step is still the gathering of a large number of nuclear Overhauser effect (NOE) and torsion restraints. Additional sources of information for the structure determination of larger RNA molecules have recently become available, and it is now possible to supplement NOE and J-coupling data with the measurement of dipolar couplings and cross-correlated relaxation rates in high-resolution NMR spectroscopy. The high quality of current structures of small- to medium-sized RNAs can be examined, and the prospects for solving larger RNA structures to equivalent levels of resolution can be assessed. The recent proliferation of complete RNA structure determinations using NMR spectroscopy is in part a result of the availability of rapid and simple methods for isotopic labeling of RNA molecules with either 13C or 15N, which permit heteronuclear experiments to be performed that resolve the severe spectral overlap inherent in the proton spectra of RNAs (Batey et al., 1992, 1995; Nikonowicz et al., 1992). The rapid development of pulse sequences tailored for RNA spin systems has facilitated many structure determinations, and it is now possible, in principle, to obtain complete sequential assignments of RNAs using only through-bond coherence transfer experiments. The process of solving a high-resolution RNA structure by NMR can be subdivided into three major steps as outlined in Figure 7.7.1. In the present discussion, the focus will be on steps II and III, addressing basic issues related to NMR data accumulation and structure calculation. After sequence-specific assignments of RNAs are obtained, the structure determination is based on collecting sufficient numbers of proton-proton distance restraints utilizing nuclear Overhauser effect correlation spectroscopy (NOESY) experiments. Potentially, these short-distance restraints between pairs of protons (<6Å) can be complemented with torsion
UNIT 7.7
angle information accessible through J-coupling constants. New experiments to measure orientational rather than distance dependent dipolar couplings, as well as cross-correlated relaxation rates, have been developed providing additional structural information (Reif et al., 1997; Tjandra and Bax, 1997a). In addition, NMR experiments have been introduced that allow the direct identification of donor and acceptor nitrogen atoms involved in hydrogen bonds (Dingley and Grzesick, 1998). These recently introduced parameters are especially important for structure determination of RNA due to the low proton density, and because a significant number of protons are potentially involved in exchange processes. There have been several recent reviews of RNA NMR methodology (Dieckmann and Feigon, 1994; Pardi, 1995; Wijmenga and van Buuren, 1998; Marino et al., 1999), including an exhaustive review by Varani et al. (1996). These reviews give an overview of the structure determination process including resonance assignment, isotopic labeling patterns, NOE identification, coupling constant measurement, and a discussion of recently solved structures. The determination of oligonucleotide structures by NMR is also discussed in UNIT 7.2.
ISOTOPIC LABELING OF RNA In order to perform a wide variety of heteronuclear NMR experiments, isotopic enrichment of RNA samples with NMR-active 13C and 15N nuclei is required. This greatly simplifies the resonance assignments and significantly extends the size limitation for structure determination of RNAs by NMR. Carbon and nitrogen cannot be readily studied with highresolution NMR techniques at natural abundance as the 12C isotope is NMR inactive and the 14N isotope possesses an electric quadrapole moment. Isotopic labeling techniques of RNAs will not be discussed in great detail, because optimized protocols for the preparation of labeled nucleotides have already been published (Batey et al., 1992, 1995; Nikonowicz et al., 1992); however, the standard method for preparation of isotopically labeled RNA is in vitro transcription from synthetic DNA oligonucleotide templates using T7 RNA polymerase and isotopically labeled nucleotide triphosphates (NTPs). Labeled NTPs can be
Contributed by M. Hennig, J.R. Williamson, A.S. Brodsky, and J.L. Battiste Current Protocols in Nucleic Acid Chemistry (2000) 7.7.1-7.7.30 Copyright © 2000 by John Wiley & Sons, Inc.
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Figure 7.7.1 Schematic protocol for RNA structure determination using heteronuclear NMR spectroscopy. Depending on the observed quality of the various NMR experiments, not all methods for the resonance assignment might be needed. Traditionally used restraint files consist of experimental distances and J-couplings only.
Recent Advances in RNA Structure Determination by NMR
isolated from bacteria grown on 13C- and/or 15N-enriched media (Batey et al., 1992, 1995; Nikonowicz et al., 1992). The advantage of enzymatic RNA synthesis is that uniform isotopic enrichment is relatively inexpensive and easy to accomplish; however, the disadvantage is the lack of selectivity in labeling. Chemical synthesis of RNA, on the other hand, has more potential for applications where selective labeling is desired. Methods for chemical synthesis of isotopically labeled RNA, however, are not as straightforward and are relatively expensive. Nevertheless, there has been recent progress in chemical synthesis methods, and they may become more utilized in the near future (Foldesi et al., 1992, 1996; Quant et al., 1994; SantaLucia et al., 1995; Tolbert and Williamson, 1996; Zhang et al., 1998).
ASSIGNMENT OF RNA NMR RESONANCES Before starting the detailed and time-consuming investigation of a chosen RNA by NMR, it is extremely important to optimize the sample conditions for data acquisition from the various required NMR experiments. It is critical to determine at the outset if the system is suitable for a high-resolution NMR structure elucidation. The imino proton region of the proton NMR spectrum of an unlabeled RNA sample in H2O provides a sensitive diagnostic for this purpose. A sample imino proton one-dimensional (1-D) spectrum for a correctly folded 30-mer RNA is shown in Figure 7.7.2. In this spectrum, one peak appears for each Watson-Crick base pair in the molecule. Since the imino protons exchange rapidly with the bulk H2O, the spectrum was recorded with a jump-return echo sequence that avoids presaturation, while providing the
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Current Protocols in Nucleic Acid Chemistry
Figure 7.7.2 Imino proton region of HIV-2 TAR RNA (inset) from a jump-return echo 1-D experiment recorded in 90% H2O/10% D2O at 298 K. Conditions used are 10 mM sodium phosphate buffer (pH 6.4), 50 mM sodium chloride, and 0.1 mM EDTA. The concentration is ∼3.0 mM RNA in 500 µL.
most efficient water suppression (Sklenar and Bax, 1987). The sample conditions to be surveyed are summarized in Figure 7.7.1. The goal is to obtain the narrowest linewidth and best chemical shift dispersion for the observable imino protons that report on secondary structure formation. A number of factors contribute to the imino proton linewidth, which are typically on the order of 10 to 30 Hz; most important is the rate of exchange with the solvent. The complex kinetics of base-pair opening and imino proton exchange in nucleic acids have been extensively reviewed in the literature and are beyond the scope of this unit (Gueron and Leroy, 1995). Assignment of RNA resonances has historically been achieved through identification of sequential base to ribose NOE patterns seen in helical regions of nucleic acid structure, in analogy to the procedure originally utilized for DNA studies in the 1980s (Wüthrich, 1986). With the advent of isotopic labeling for RNA, the basic NOE assignment approach was initially expanded to include multidimensional (3-D and 4-D) versions of the standard NOESY, correlation spectroscopy (COSY), and total
correlation spectroscopy (TOCSY) 2-D experiments, which simplified assignment and identification of NOEs (Nikonowicz and Pardi, 1992, 1993; Pardi and Nikonowicz, 1992). The NOE-based approach, however, relies on assumptions about structure and assignments and is somewhat susceptible to errors from structural bias. A methodology that achieves sequential assignment via unambiguous through-bond correlation experiments, as is the case for proteins, would be more ideal. The easiest extensions of homonuclear experiments, applicable with labeled samples, are one-bond heteronuclear single quantum coherence (HSQC)– or heteronuclear multiple quantum coherence (HMQC)–type experiments, where the proton resonances are separated in two-dimensional spectra according to the direct bound heteronuclei, e.g., 15N or 13C. These experiments utilize large 1H-15N or 1H-13C one-bond couplings rather than relatively small multiple-bond 1H homonuclear scalar couplings for magnetization transfer. The resulting two-dimensional HSQC or HMQC correlations between 1H and 13C or 15N spins are integral components of all multidimensional NMR experiments. The dra-
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Recent Advances in RNA Structure Determination by NMR
Figure 7.7.3 (Top) The aromatic region of a 13C-HSQC experiment of HIV-2 TAR RNA. Partial assignments are given in less crowded regions of the spectrum. The corresponding region from a 1-D 1H NMR spectrum recorded in 99.9% D2O at 298 K is shown on top. 1H,13C correlations arising from H2 and C2 resonances of adenosines are not shown. Correlations that arise from C6 carbon resonances in pyrimidines are split in the carbon dimension due to the homonuclear 1JCC coupling constant evolution to the C5 resonance. (Bottom) Ribose region of a 13C-HSQC experiment of HIV-2 TAR RNA. Nonspecific assignments for the different one-bond correlations are given. The corresponding region from a 1-D 1H NMR spectrum recorded in 99.9% D 2O at 298 K is shown on top. The aromatic H5 protons of pyrimidines that overlap with the H1′ proton region are not visible in the 1H,13C correlations, because the corresponding C5 resonates outside the depicted carbon spectral region. Further differences between the 1-D and 2-D experiments arise from incomplete suppression of the HDO and EDTA proton resonances.
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matic gain in resolution for RNA proton resonances bound to carbon is demonstrated in Figure 7.7.3, top panel, for the aromatic 1H,13C region and in Figure 7.7.3, bottom panel, for the ribose 1H,13C spectral region. There has been a proliferation of recent experiments that correlate RNA proton resonances through the heteronuclear spin systems of the base, the ribose ring, and phosphate backbone. Those experiments can be considered as extensions of basic two-dimensional HSQC or HMQC correlations. The new NMR experiments for resonance assignment can be conceptually organized into four classes (Fig. 7.7.4 and Table 7.7.1). Con-
ceptually, the names given to the through-bond correlation experiments are derived from the series of nuclei through which magnetization is transferred during the experiment. The nuclei that undergo chemical shift evolution during the experiment are given in capital letters, sugar and base resonances are specified using subscripted lower case letters, and intervening nuclei, through which magnetization is transferred but which are not directly observed, are enclosed in parentheses. For example, the Hs(CsNbCb)Hb experiment shown in Figure 7.7.4, panel C, correlates the H1′ sugar proton (Hs) via transfer of magnetization to the directly attached C1′ sugar carbon (Cs). Subsequently,
Figure 7.7.4 Schematic of coherence transfer for through-bond assignment experiments. (a) HCN correlation experiments for assignment of the base resonances. Closed and open arrows show HC-TOCSY-CNH and HNC-TOCSY-CH experiments, respectively. (b) HCP experiments for sequential ribose-ribose assignment. (c) HCN experiments for intranucleotide correlation of base and ribose resonances. Closed arrows show the sequential Hs-to-Hb correlation. Open arrows show one of the out-and-back experiments (HsCsNb). See Table 7.7.1 for a more complete listing of all types of experiments. Numbering conventions are included for each of the base and ribose positions.
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Table 7.7.1
Experiment
Through-Bond Correlation Experiments for Assignment of RNA Resonances
Nucleotide Coherence pathwaya
Reference
Base-base (carried out in H2O, 13C/15N-labeled sample) H(NC)-TOCSY-(C)H Guanosine I HT T I H1(t1)→N1→C6/C2→C8→H8(t2) H(C)-TOCSY-CH Adenosine I T T I H2/H8(t1)→C2/C8→C4/C6→C8/C2(t2)→H8/H2(t3) (H)N(C)-TOCSY-(C)H Adenosine HT HT T I H6→N6(t1)→C6→C8/C2→H8/H2(t2) H(NCCC)H Uridine I HT I I I H3(t1)→N3→C4→C5→C6→H6(t2) Cytosine HT HT I I I H4(t1)→N4→C4→C5→C6→H6(t2) I T I I HC-TOCSY-(CN)Hb Purine H8/H2(t1)→C8/C2(t2)→C6→N6/N1→H6/H1(t3) I T HT I H(C)-TOCSY-(CN)Hb Guanosine H8(t1)→C8→C6→N1→H1(t2) Uridine I HT I H6(t1)→C6→N3→H3(t2) Base-sugar (preferably carried out in D2O, 13C/15N-labeled sample) Hs(CsNbCb)Hbb All I I I I H1′(t1)→C1′→N9/N1→C8/C6→H8/H6(t2) All I I I I HsCsNbb H1′→C1′(t1)→N9/N1(t2)→C1′→H1′(t3) HbCbNbf All I I I I H8/H6→C8/C6(t1)→N9/N1(t2)→C8/C6→H8/H6(t3) All I I I I HsCs(NbCb)Hb H1′(t1)→C1′(t2)→N9/N1→C8/C6→H8/H6(t3) Purine I I I I HsCsNb H1′→C1′(t1)→N9(t2)→C1′→H1′(t3) HsCs(Nb)Cb Purine I I I I I I H1′→C1′→N9→C8(t1)→N9→C1′(t2)→H1′(t3) Purine I I I HbNbCb H8→N9(t1)→C8(t2)→H8(t3) Phosphate-backbone (preferably carried out in D2O, 13C-labeled sample) HCP All I I I I H4′→C4′→P(t1)→C4′(i+1)(t2)→H4′(i+1)′(t3) HCP-CCH-TOCSY All I I I T I H4′→C4′→P(t1)→C4(i+1)→C1′(i+1)(t2)→H1′(i+1)′(t3) P(CC)H-TOCSY All I T HT P(t1)→C4′(i,i+1)→C1′(i,i+1)→H1′(i,i+1)′(t2) P(H)H-TOCSY All HT T P(t1)→H3′/4′/5′(i,i+1)→H1′/2′/3′/4′/5′(i,i+1)′(t2) Sugar-sugar (preferably carried out in D2O, 13C-labeled sample) HCCH-COSY All I I I H1-5′→C1-5′(t1)→C(i±1)′(t2)→H(i±1)′(t3) HCCH-TOCSY All I T I H1-5′→C1-5′(t1)→C(i±n)′(t2)→H(i±n)′(t3) HCCH-fdTOCSY All I T I H1-5′→C1-5′(t1)→C(i+n)′(t2)→H(i+n)′(t3)
Simorre et al. (1996a) Legault et al. (1994); Marino et al. (1994a) Simorre et al. (1996b) Simorre et al. (1995) Simorre et al. (1995) Fiala et al. (1996) Sklenar et al. (1996) Sklenar et al. (1996)
Sklenar et al. (1993b) Heus et al. (1994); Marino et al. (1994b) Heus et al. (1994); Marino et al. (1994b) Farmer et al. (1993) Farmer et al. (1994) Farmer et al. (1994) Farmer et al. (1994)
Heus et al. (1994); Marino et al. (1994b) Marino et al. (1995) Wijmenga et al. (1995) Kellogg (1992)
Kay et al. (1990) Fesik et al. (1990) Hu et al. (1998); Schwalbe et al. (1995)
aThe letter above the arrow indicates the type of coherence transfer method used: I - INEPT, T-TOCSY, HT-Hetero-TOCSY. In parentheses are the (t , 1 t2, t3) dimensions for which chemical shift is detected. bExperiment titles have been slightly altered in order to be presented in a uniform fashion. Parentheses indicate a nucleus that coherence is transfered
through, but the chemical shift is not detected.
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magnetization is transferred via 1JNC couplings to the N9 (for purines) or N1 (for pyrimidines) nitrogens (Nb), and further to the directly attached C8 (for purines) or C6 (for pyrimidines) base carbons (Cb). Finally, magnetization is transferred to the base protons (Hb, H8/H6 for purines and pyrimidines, respectively) for detection (t1 chemical shift evolution period). Thus, the Hs(CsNbCb)Hb experiment is a twodimensional experiment correlating sugar to base protons. The CsNbCb pathway for magnetization transfer is required to connect the Hs and Hb, but is not directly recorded in the experiment. The HCN class correlates the protons of the bases (both exchangeable and non-exchangeable) to one another via JCC and JCN couplings (Sklenar et al., 1987; Simorre et al., 1995, 1996a,b; Fiala et al., 1996), while another HCN class correlates the base resonances (H8/H6) to the ribose H1′ proton through the intervening carbon (C8/C6/C1′) and nitrogen (N9/N1) resonances (Farmer et al., 1993, 1994; Sklenar et al., 1993a,b). An HCP class of experiments correlates the sugar resonances of sequential nucleotides through the phosphate backbone (Heus et al., 1994; Marino et al., 1994b, 1995; Varani et al., 1995; Wijmenga et al., 1995). Finally, the extended versions of Table 7.7.2
homonuclear COSY and TOCSY experiments utilizing large one-bond 1JHC and 1JCC coupling constants for the coherence transfer between ribose proton and carbon resonances comprise the HCCH class. Heteronuclear chemical shifts relevant for the experiments discussed below are summarized in Table 7.7.2.
Base HCN Correlation Experiments A set of HNCCH- and HCCNH-TOCSY experiments have been developed that correlate the exchangeable imino and amino proton resonances with the nonexchangeable base resonances for the complicated spin systems of all four nucleotides (Sklenar et al., 1987; Simorre et al., 1995, 1996a,b; Fiala et al., 1996; Wöhnert et al., 1999). In contrast to the poor chemical shift dispersion of the different ribose protons, the imino proton resonances show reasonably good dispersion mainly due to varying hydrogen-bonding networks and stacking effects of base pairing in RNAs as shown in Figure 7.7.5. Unfortunately, amino groups frequently undergo rotation around the carbon-nitrogen bonds, which can result in intermediate exchange for the attached NH2 proton resonances, resulting in a broad linewidth, as can be seen in Figure 7.7.6.
Heteronuclear Chemical Shifts in Nucleotides
Atom(s)
Chemical shift range
Carbon Purine C5 Purine C8 Purine C2,C4,C6 Pyrimidine C5 Pyrimidine C6 Pyrimidine C2,C4 Ribose carbons
∼120 ppm ∼140 ppm ∼150-160 ppm ∼100 ppm ∼145 ppm ∼155-170 ppm ∼70-90 ppm
Nitrogen Purine N9 Purine N7 Guanosine N1 Guanosine N2 Guanosine N3 Adenosine N1 Adenosine N3 Adenosine N6 Pyrimidine N1 Uridine N3 Cytosine N3 Cytosine N4
∼170 ppm ∼220-240 ppm ∼145-150 ppm ∼70-80 ppm ∼160-165 ppm ∼220 ppm ∼215 ppm ∼80-90 ppm ∼145-150 ppm ∼150-160 ppm ∼195-200 ppm ∼95 ppm
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Figure 7.7.5 Assignments of the imino region of the HIV-2 TAR argininamide complex from a jump-return HMQC experiment recorded in 90% H2O/10% D2O at 298 K. The terminal 5′G⋅C3′ G16 imino resonance as well as the U40 imino resonance from the base-pair that terminates the upper stem are missing due to unfavorable exchange properties with the solvent.
The HCN correlation experiments differ in the exact mechanism of magnetization transfer, but predominantly rely on one-bond heteronuclear coupling constants to transfer magnetization from protons to the heteronuclei of the ring, and homonuclear 13C-TOCSYs to transfer magnetization across the ring. There are two important issues which impinge upon the sensitivity of these experiments. One is the chemi-
Recent Advances in RNA Structure Determination by NMR
cal shift differences between the carbon resonances in the ring. The efficiency of homonuclear 13C-TOCSY transfers relies on the elimination of chemical shift differences as achieved by the spin-lock mixing schemes. The large bandwidth of carbon resonances associated with the different bases require radio frequency (rf) powers beyond current instrumental limitations for efficient carbon spin lock. Therefore,
Figure 7.7.6 Residue-type specific assignments of the amino region of the φ21 boxB RNA hairpin complexed to a 19–amino acid bacteriophage Nλ peptide (Cilley and Williamson, 1997) from an HSQC experiment recorded in 90% H2O/10% D2O at 298 K. In the case of cytidine amino residues involved in base-pairing interaction, the rotation around the C-N bond is sufficiently slow to observe two distinct proton resonance frequencies; however, the amino resonances of base-paired guanosines and adenosines are generally not observable due to intermediate exchange at 298 K. The only observable resonances are those of unpaired purine residues. Only one proton resonance is observable, indicative of fast exchange on the NMR time scale.
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magnetization is usually transferred via smaller two-bond carbon-carbon (2JCC) coupling constants (C8-C4-C6 for purines and C6-C4 for pyrimidines), which require long spin-lock mixing times for transfer and loss of sensitivity due to fast carbon transverse relaxation. Alternatively, sequential homonuclear insensitive nuclei enhanced by polarization (INEPT) transfers (C4-C5-C6) can be used rather than a 13C-TOCSY H(NCCC)H for pyrimidines (Simorre et al., 1995). For purines, a broad-band spin-locking scheme (Mohebbi and Shaka, 1991) can be used to increase the efficiency of one-bond carbon-carbon transfer and decrease mixing times (Simorre et al., 1996a,b). Adenines have a network of homonuclear 2JCC and 3J CC couplings, allowing the assignments of nonexchangeable H2 and H8 base proton resonances using HCCH-TOCSY experiments (Legault et al., 1994; Marino et al., 1994a). Magnetization between the H2 and H8 protons can be transferred through the intervening carbon using a broad-band spin-locking scheme (Mohebbi and Shaka, 1991). Another factor, which decreases the sensitivity, is rotational exchange broadening of exocyclic amino protons on adenine, guanine, or cytosine. More efficient 1H,15N-transfer techniques than the standard INEPT procedure have to be used when proton resonances are broadened by rotational exchange (Mueller et al., 1995). In addition, the nitrogen linewidths are narrower than the corresponding protons undergoing chemical exchange as shown in Figure 7.7.6, and detection of nitrogen chemical shift rather than proton is more efficient (Simorre et al., 1995, 1996b). The extra sensitivity gained by optimization is offset by increased experiment time for acquiring a separate data set for each nucleotide. Nevertheless, the added sensitivity probably will be essential for assigning the resonances of many larger RNAs.
Base-to-Ribose Correlation Experiments An important application of 13C/15N-labeling of RNA is through-bond intranucleotide correlation of base-to-ribose resonances, which historically has been achieved through NOE transfer. There are now many published experiments for achieving this through bond transfer (Farmer et al., 1993, 1994; Sklenar et al., 1993a,b), which are reviewed by Dieckmann and Feigon (1994) and Pardi (1995). A simple sequential INEPT transfer procedure with nonselective pulses can be used to provide sugarto-base correlations. Branching coherence
transfer pathways occurring due to 1JNC couplings of similar size and thus correlating N9/N1 to C8/C6 as well as C4/C2 resonances in purines and pyrimidines, however, reduce the sensitivity of the basic approach. In addition, small one-bond 1JNC couplings result in relatively inefficient magnetization transfer. Therefore, selective carbon pulses have been cleverly utilized to direct coherence predominantly along the desired pathways. Separate optimized sequences are often required because of sensitivity and overlap problems (Farmer et al., 1994). Acquiring several experiments detecting different heteronuclei (C8/C6, N9/N1, or C1′) may resolve overlap problems for larger RNA molecules. The experiments which will provide the best resolution may vary for different RNA molecules and can be best determined empirically from 2-D 1H-13C or 1H-15N HSQCs of the base and C1′/H1′ resonances.
Phosphate Backbone HP and HCP Correlation Experiments In order to link the resonances of each ribonucleotide sequentially without the use of NOEs, magnetization must be transferred through the phosphate backbone. For unlabeled RNAs, a number of relatively efficient 1H,31Pmultidimensional correlation schemes are available for sequential assignment of 31P and ribose 1H resonances. Magnetization can be transferred from excited 31P resonances to the 3J HP scalar coupled ribose protons for detection using either COSY (Sklenar et al., 1986) or heteronuclear TOCSY (Kellogg, 1992) transfer steps. The resulting two-dimensional H3′/H5′/H5′′, 31P-correlations can be concatenated with homonuclear 1H,1H NOESY or TOCSY experiments to transfer magnetization to potentially better-resolved resonances like H1′ or aromatic H8/H6 resonances (Kellogg and Schweitzer, 1993). A straightforward approach for 13C-labeled RNAs is HCP correlation via sequential INEPT transfers (i. e., 1H → 13C → 31P → 13C → 1H; Heus et al., 1994; Marino et al., 1994b) correlating nuclei of adjacent nucleotides i and i + 1. This approach has two limitations for complete assignment of RNA molecules over ∼20 nucleotides. One is severe overlap due to poor spectral dispersion of C4′i,i+1 and Pi resonances, particularly in A-form helical regions. Second, the better-resolved C3′i-Pi-C5′i+1 connectivity requires a separate HCCH-TOCSY experiment to connect the C3′ and C5′ resonances on the same nucleotide. Subsequent experiments, HCP-
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CCH-TOCSY (Marino et al., 1995) and P(CC)H-TOCSY (Wijmenga et al., 1995), resolve both problems by combining the HCP and HCCH-TOCSY experiments and thus resolving relevant correlations on the well-dispersed C1′/H1′ resonances. However, it is not clear whether these experiments will be generally useful for RNA molecules larger than ∼30 nucleotides (∼10 kDa), particularly with the severe line broadening of 31P resonances at higher molecular weights and higher magnetic fields.
Ribose-to-Ribose HCCH Correlation Experiments
Recent Advances in RNA Structure Determination by NMR
The information obtained from homonuclear 1H, 1H-COSY and -TOCSY experiments can be obtained more readily using HCCHCOSY and -TOCSY experiments on ribose rings uniformly labeled with 13C, which allows magnetization transfer and chemical shift evolution on the C1′ to C5′ carbons (Fesik et al., 1990; Kay et al., 1990; Pardi and Nikonowicz, 1992; Nikonowicz and Pardi, 1993; Pardi, 1995). The assignment of the severely overlapped ribose proton resonances is facilitated through the additional carbon dimension, which provides better shift dispersion. Furthermore, the magnetization transfer through the ribose proton spin systems was hampered due to the small 3JHH-vicinal coupling, present in most commonly populated A-form RNA, correlating the H1′ and H2′ resonances. HCCHtype experiments overcome this problem by making use of large and uniform one-bond 1JHC and 1JCC coupling constants for more efficient coherence transfer. In contrast to vicinal 3J couplings, one-bond couplings are, in general, much less biased due to different conformations present in various structural motifs, thus facilitating resonance assignments based on through-bond correlation experiments. The HCCH-type experiments transfer magnetization from the excited sugar protons to the directly attached carbons. Neighboring carbon resonances can be correlated through scalar 1J CC coupling constants utilizing either COSY, relayed-COSY, or TOCSY (Fesik et al., 1990) mixing schemes. Forward-direct TOCSY provides the best sensitivity for magnetization transfer through the whole ribose JCC coupling network (Schwalbe et al., 1995; Hu et al., 1998). Finally, all schemes transfer magnetization back to the protons for detection. Unlike triple-resonance experiments used for proteins, some of the triple-resonance experiments for RNA are not mandatory prereq-
uisites for assignments and structure determination. The HCN experiments for correlation of the base resonances are the most likely to succeed with medium-sized RNA molecules (30 to 40 nucleotides, ∼10 to 15 kDa) due to less spectral overlap and the relatively long relaxation times of the base resonances. Complete through-bond assignment will probably not be feasible for many medium-sized RNAs, since the HCP experiments are most susceptible to relaxation problems. A hybrid approach with HCN and NOESY experiments will likely be the optimal compromise. The HCN experiments can unambiguously determine the intranucleotide correlations within and between the base and ribose resonances, which will significantly reduce the ambiguity present in the complete NOESY-based assignment procedure. Even assuming no problems due to relaxation, it does not appear that there will be a simple and quick procedure for assignment of RNA molecules. Neglecting the problems with sensitivity or overlap, complete assignment with only through-bond methods still requires a large number of experiments, if all of the optimized sequences are performed (∼4 experiments for the bases, ∼3 experiments to correlate the base resonances to the ribose, and ∼2 to 3 experiments to correlate the ribose resonances). This results in a very rough estimate of ∼20 days measurement time (assuming an average of two days measurement time per experiment) for an RNA sample, with sample concentrations in the mM range and a molecular weight between 10 and 25 kDa, carried out on spectrometers with at least 500 MHz proton resonance frequency. Nevertheless, this is around the same number of experiments and days of measurement time that have been required for many RNA structures solved by the NOE-based method, since multiple samples with different specific labeling patterns (see below) were required to resolve ambiguities in the sequential NOEs (Battiste et al., 1995; Cai and Tinoco, 1996; Dieckmann et al., 1996; Mao and Williamson, 1999; Mao et al., 1999). Despite roughly the same number of experiments, analysis of the spectra for the through-bond experiments should hopefully be quicker and should lower the potential for misassignment inherent in the NOE method.
NOE MEASUREMENT The end goal of NMR analysis is usually a structure determination. Despite the method used for assignment of resonances, the main
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source of structural data is still obtained from NOEs, which provide distance restraints for pairs of hydrogen atoms in the RNA molecule. Only short proton-proton distances in the range <6 Å are accessible through NOESY experiments. The intensity of NOESY cross peaks is approximately proportional to the inverse of the averaged distance to the power of six, <1/rij6>, assuming an isolated pair of proton spins i and j. This assumption, however, is usually not valid, which complicates the quantitative treatment of NOESY cross-peak intensities. Potential problems with interpretation of obtained NOESY cross-peak intensities in terms of 1H1H distances in structure calculations arise mainly from a phenomenon called spin diffusion. Spin diffusion causes a breakdown of the isolated spin pair approximation, because other nearby protons provide competing indirect pathways for observing the direct NOE between the two protons. Spin diffusion effects play a role especially when longer NOESY mixing times (>100 msec) are used. This usually leads to damped NOESY cross-peak intensities that build up through the direct pathway, resulting in underestimated interproton distances. Additionally, multistep transfer pathways can occur, resulting in false NOE assignments. For example, the imino protons of guanines might show spin diffusion–mediated NOEs to the nonexchangeable aromatic H5 and H6 protons of cytidines in Watson-Crick base pairs through the cytidine amino protons. However, in an early stage of the assignment procedure based on NOESY correlations, spin diffu-
sion pathways can aid the identification of spin systems. Thus, for assignments, it is recommended to analyze NOESY spectra acquired with shorter (∼50 msec) and longer (∼150 msec) mixing times. The classification of NOE cross peaks for the structure calculation is discussed in more detail below. Sequential assignment of RNA proton resonances can be achieved for lower-molecular-weight systems (<10 kDa) by identification of NOE patterns seen in A-form helical regions. Homonuclear 1H-1H NOESY in H O and D O solution both 2 2 potentially contain valuable through-space correlations for the sequential assignment of adjacent nucleotides. Sequential walks using nonexchangeable ribose and aromatic protons can be achieved in canonical A-form helix through the combined analysis of intraresidual H1′-toH6/H8 (3.5 and 3.7 Å, respectively) and interresidual H1′-to-H6/H8 NOEs (4.3 Å). The relevant distances are illustrated in Figure 7.7.7. Additionally, the short interresidual distances between the H2′ ribose and aromatic H6/H8 proton resonances (1.7 Å) as shown in Figure 7.7.7 give rise to strong NOE cross-peaks. NOESY data recorded in H2O potentially reveal sequential connectivities between imino protons of residues i and i + 1 (5.2 Å). Additionally, interresidual imino proton to H6/H8 aromatic proton NOE cross-peaks (4.3 and 4.4 Å, respectively) might be observable. However, the NOE-based approach is limited and susceptible to errors from structural bias. Most of the expected A-form sequential through-space
Figure 7.7.7 Classical sequential assignment pathway for A-form helical RNA using intra- and interresidual correlations between H1′/H2′ (i) and H6/H8 (i,i+1) nonexchangeable proton resonances from NOESY experiments. NOESY cross-peaks due to interresidual H2′ (i) to H6/H8 (i+1) correlations are usually strong (dHH ∼ 1.7Å), while intra- and interresidual correlations connecting H1′ (i) and H6/H8 (i,i+1) protons give rise to weaker cross-peaks (d HH = 3.5Å or 4.3Å).
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connectivities are either missing or altered in noncanonical loop or bulge regions. Identification of NOEs can be aided by resolving the 1H-1H NOE connectivities that are essential for determining the structure into three and four dimensions through detection of the heteronuclear (13C/15N) chemical shifts of the proton-attached nuclei (Nikonowicz and Pardi, 1992, 1993). Even with the extra dimensions in 3-D NOESY-HSQC or 4-D HMQCNOESY-HSQC experiments, overlap can still be severe for medium-sized RNAs (∼30 to 40 nucleotides). An approach to aid in the identification of NOEs that has been very successful is selective isotopic labeling of RNA with either heteronuclei (13C/15N) or deuterium (2H). 13C/15N-labeled nucleotides can be separated and used individually in in vitro transcriptions with other unlabeled nucleotides (e.g., [13C]GTP plus unlabeled ATP, CTP, and UTP) to produce RNA molecules selectively labeled at only one nucleotide (Nikonowicz et al., 1992; Batey et al., 1995). There are two advantages to selective labeling for identification of NOEs. First, reduced overlap in 3-D NOESY experi-
ments, which permits unambiguous identification of more NOEs. Second, isotopic filtering experiments (Otting and Wüthrich, 1989a, 1990) can be performed to help resolve intranucleotide from internucleotide NOEs (Battiste et al., 1995). The introduction of residue-specific 13C-labeling in conjunction with isotopic filtering makes it possible to distinguish between different classes of NOEs, namely 13C-1H to 13C-1H, 12C-1H to 12C-1H, and 12C-1H to 13C1H, respectively. This is a particularly powerful approach that can also be utilized for sequential assignment of RNA where the through-bond methods described above are not successful. A similar approach has also been used with RNAs containing site-specific labels produced synthetically at the C8/C6 positions of the base (SantaLucia et al., 1995; Cai and Tinoco, 1996). One additional labeling methodology that has been developed is specific labeling of a short segment of nucleotides within a larger RNA sequence (Xu et al., 1996). Depending on the RNA being studied and the structural questions being addressed, each of these specific heteronuclear labeling methods will greatly assist in
A.
B.
Recent Advances in RNA Structure Determination by NMR
Figure 7.7.8 Effect of specific deuterium labeling on RNA NOESY spectra. (A) NOESY spectrum of unlabeled TAR RNA (30 nucleotides) in D2O. (B) Same region for d4-TAR RNA which has deuterium labels at the 3′, 4′, 5′, and 5′′ positions of the ribose ring. A dramatic reduction in spectral overlap is evident, and an unusual 2′-2′ NOE is observed that could not be identified in heteronuclear NOESY experiments of 13C-labeled TAR RNA.
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the assignment and structure determination of medium-sized RNA molecules. In general for medium to large RNAs, several labeled samples are required to obtain unambiguous assignments. Another way to remove or filter proton resonances is to specifically substitute them with deuterium. An advantage of deuteration, not present with specific 13C/15N-labeling, is the decreased relaxation times of protons in RNAs with reduced proton spin density. In addition, any 13C atoms attached to deuterium will have better relaxation properties with respect to the 13C-1H moieties, and 13C-13C transfers are more efficient (Dayie et al., 1998). Therefore, specific deuterium labeling should greatly aid in the study of larger RNA molecules. Two protocols have been published for specific deuterium labeling. One uses a combination chemical/enzymatic synthesis approach (Tolbert and Williamson, 1996), while the other uses all chemical synthesis (Foldesi et al., 1992, 1996; Glemarec et al., 1996). Figure 7.7.8 shows an example of the filtering affect on a NOESY spectrum of specific deuterium labeling of a 30-nucleotide RNA at the 3′, 4′, 5′, and 5′′ positions of the ribose ring (Tolbert and Williamson, 1996). In addition to the reduction in overlap, the relaxation times of the remaining protons in the proton-depleted environment were increased ∼2-fold. The disadvantage of deuterium labeling is the greatly reduced 1H-1H NOE-based information content essential for the structure determination. These disadvantages may in part be compensated because high degrees of deuteration allow longer NOESY mixing times to be used (>150 msec), since potential spin diffusion pathways are limited. This possibly extends the observable 1H-1H distance in NOESY experiments to ∼7 Å, thus adding rare long-range information for the structure determination, as has been shown for highly deuterated proteins. It is also possible, however, to make multiple samples with different deuteration patterns to circumvent this problem (Tolbert and Williamson, 1996). Alternatively, since many of the NOEs between the base and sugar protons that define the conformation of the helices are redundant, the reduced NOE data set from one specific deuteration pattern may still be useful for determining qualitative structural models of larger RNAs. A number of protons involved in relevant interactions in nucleic acids are often not observable due to exchange processes. An experimental approach to improving the sensitivity of
NOEs to amino protons in RNA has been developed (Krishnan and Rance, 1995; Mueller et al., 1995). As was noted above, similar concepts have also been utilized to improve the sensitivity of through-bond experiments for assignment of base amino protons (Simorre et al., 1995, 1996b). Easier assignment and identification of amino proton NOEs, which are often difficult to observe, has the potential to greatly increase the quality of structure determinations. This is particularly true for RNA-protein interactions, since amino protons line the grooves where proteins often make specific contacts. RNA structure tends to be very dynamic and flexible, and it is very common to observe NMR resonances that are broadened by conformational exchange.
HYDROGEN BONDS IN RNA NMR STRUCTURES Some of the most important interactions within RNA structures are hydrogen bonding interactions. These can now be directly determined unambiguously by NMR even in the case of the noncanonical base-pairing interactions that are the most biologically interesting and novel aspects of RNA structure. The existence of scalar couplings due to hydrogen bonds between imino proton donors and acceptor nitrogens in Watson-Crick base-pairs of RNA was recently shown (Dingley and Grzesiek, 1998). Hydrogen bonds have a partially covalent character that gives rise to scalar spin-spin couplings of the 2hJNN and 1hJHN type, which are an important additional NMR parameter for the structure determination of biomacromolecules in solution. In early stages of a structural study, these HNN-COSY-type experiments allow the rapid identification of basic secondary structural elements such as A-form Watson-Crick duplexes in RNA. The imino resonances of base-paired guanosines and uridines can be directly correlated with their acceptor nitrogens, N3 for cytosines and N1 for adenines, respectively, as shown in Figure 7.7.9. The corresponding 2hJNN coupling constant values across the hydrogen bonds are surprisingly large and on the order of 6 Hz (Dingley and Grzesiek, 1998; Pervushin et al., 1998a). Before the recent advance in measuring these scalar couplings, it was only possible to obtain indirect evidence for Watson-Crick base pairing in the A-form helix by the observation of upfield-shifted imino resonances, along with a network of cross-strand NOEs indicative of a typical base pair.
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Figure 7.7.9 Direct observation of scalar cross-hydrogen bond 2hJNN coupling constants in the HIV-2 TAR argininamide complex at 298 K in H2O using a quantitative J-correlation HNN–COSY experiment. Assignments for all observable imino resonances that are involved in Watson-Crick base pairs in HIV-2 TAR are given. Sequence and secondary structure of HIV-2 TAR is shown. The sequence is identical to HIV-1 TAR except for the deletion of the C24 bulge nucleotide.
Recent Advances in RNA Structure Determination by NMR
Unfortunately, some hydrogen bonding interactions are difficult to observe due to rapid imino proton exchange; however, a modified pulse scheme allows the observation of 2hJNN couplings in the absence of detectable imino protons (Hennig and Williamson, 2000). Instead of measuring the coupling by detection of the imino protons, the 2hJNN couplings can be observed via 2JHN correlations with nonexchangeable base protons. The experiment provides a sensitive measure of base-pairing interactions, even in D2O solution. This approach has led to the conformation of the existence of the U38⋅A27⋅U23 base triple in the HIV-2 transactivation response element (TAR)-argininamide complex as shown in Figure 7.7.10. Additionally, the chemical shifts of the unprotonated base nitrogens, which can be correlated to nonexchangeable aromatic protons via 2JHN couplings in a reasonably sensitive manner, might be useful to identify hydrogen-bonding interactions (Sklenar et al., 1994). Crystal structures of two different ribozymes have highlighted the important structural role of 2′-OHs (Pley et al., 1994a; Scott et al., 1995; Cate et al., 1996). Only a handful of 2′-OH protons have been observed by NMR, thus making any direct evidence for hydrogen bonding difficult (Allain and Varani, 1995). However, hydrogen-bonding interactions have been proposed based on a large number of NOEs in the region which localize possible
hydrogen bond acceptor and donor groups near the 2′-OH, as in the GNRA structures (where N is any nucleotide and R is a purine; Jucker et al., 1996). One additional consideration is suggested by the paromomyin-16S RNA structure, where some potential interactions appeared with heavy atom distances in the 4.0 to 4.5 Å range (Fourmy et al., 1996). This distance is generally considered too long for a hydrogen bond, but could possibly be due to a water-mediated hydrogen bond. These types of interactions are very difficult to determine even in protein NMR, where large hydrophobic cores help protect single water molecules from exchanging with the bulk solvent (Otting and Wüthrich, 1989b; Otting et al., 1991). The structural role of 2′-OHs and water molecules in RNA structure will be an important area of future investigations.
MEASUREMENT OF BACKBONE TORSIONS Many of the interesting RNA structures studied exhibit a wide variety of backbone conformations, suggesting that torsion restraints will be important to define RNA structure. RNA backbone conformations are characterized by six backbone torsion angles (α, β, γ, δ, ε, ζ) as illustrated in Figure 7.7.11. NMR analysis of the backbone conformation is complicated by the lack of useful 1H-1H NOE distance restraints available that define the back-
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A.
B.
Figure 7.7.10 Direct observation of scalar cross-hydrogen bond 2hJNN and intraresidue 2JNN coupling constants in the HIV-2 TAR argininamide complex at 298 K in D 2O. (A) Quantitative J-correlation 2JHN HNN-COSY spectrum. Assignments for all adenosine residues in HIV-2 TAR are given on top of the spectrum. The corresponding H2 and H8 proton resonance frequencies are highlighted by vertical wide and narrow dotted lines, respectively. Cross-peaks that are due to 2JNN couplings have opposite signs and are shown in boxes. A dashed horizontal line connects the N3 resonance frequency of U23 as obtained from (A) 2JHN HNN-COSY and (B) nJHN HMQC experiments. Arrows point to the A20-U42 Watson-Crick base-pair correlation, the A22-U40 correlation, and the A27-U23 correlation at the H8 proton resonance frequency of A27. Chemical shift regions for the nitrogen and proton resonances giving rise to the observable correlations are given next to the corresponding axis. Peaks marked with an asterisk are due to minor impurities from sample degradation. (B) Identification of cross-hydrogen bond coupling partners using intraresidual nJHN HMQC correlations. Assignments for all uridine residues in HIV-2 TAR are given. Connectivities for uridine residues are due to small intraresidue couplings (3JH5N1 ≈ 4.5 Hz, 3JH5N3 ≈ 2.5 Hz). The corresponding H5,N1 and H5,N3 cross-peaks are connected by vertical solid lines. Chemical shift regions for the nitrogen and proton resonances giving rise to the observable correlations are given next to the corresponding axis.
bone torsions. Alternatively, vicinal 3J coupling constants can provide useful structural information about torsion-angle conformations (Wijmenga and van Buuren, 1998; Marino et al., 1999). Vicinal 3J couplings, involving nuclei separated by three bonds, can be correlated to torsion angles using empirical Karplus relations (Karplus, 1959). With the introduction of isotopic labeling methods, a number of approaches have been developed that take advantage of both 1H-31P and 13C-31P couplings to measure these torsion angle restraints. Additionally, the ribose sugar pucker as well as the glycosidic torsion angle χ as shown in Figure 7.7.11 can be characterized utilizing vicinal 1H-1H and 13C-1H couplings.
Sugar Pucker and the δ Torsion The ribose sugar geometry is defined by five alternating torsion angles (ν0 through ν4, see Fig. 7.7.11). Usually, the ribose sugar adopts one of the energetically preferred C2′-endo (South) or the C3′-endo (North) conforma-
tions. A number of 1H,1H and 1H,13C scalar couplings are available to determine the sugar pucker qualitatively, with a combination of H1′-H2′ and H3′-H4′ coupling constants being the most useful for smaller RNAs. The 3JH1′H2′ vicinal coupling is >8 Hz for C2′-endo puckers and ∼1 Hz for the C3′-endo puckers, typically found in A-form helices. The opposite behavior is expected for the 3JH3′H4′ coupling constant with C2′-endo puckers associated with small and the C3′-endo puckers associated with relatively large coupling constant values. The easiest method to measure the H1′,H2′/H3′,H4′ coupling constant is through the use of 2-D 1H-1H COSY experiments (Varani and Tinoco, 1991). The absence of a H1′,H2′ COSY crosspeak is often qualitatively interpreted as the C3′-endo conformation; however, this approach may be misleading where broad lines lead to the cancellation of the antiphase crosspeaks. This pitfall, which becomes more and more important with increasing molecular weights associated with line-broadening ef-
Biophysical Analysis of Nucleic Acids
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Figure 7.7.11 The six backbone torsion angles are shown on a segment of RNA. Individual torsion angles are defined as follows: α (O3′i-1-P i- O5′i- C5′i), β (P i- O5′i- C5′i- C4′i), γ (O5′i- C5′i- C4′iC3′i), δ (C5′i- C4′i- C3′i- O3′i), ε (C4′i- C3′i- O3′i-P i+1), ε (C3′i- O3′i-P i+1-O5′i+1), ν0 (C4′-O4′-C1′-C2′), ν1 (O4′-C1′-C2′-C3′), ν2 (C1′-C2′-C3′-C4′), ν3 (C2′-C3′-C4′-O4′), ν3 (C3′-C4′-O4′-C1′), χ (pyrimidine) (O4′-C1′-N1-C2), χ (purine) (O4′-C1′-N9-C4).
fects, can be avoided with the use of the HCCHE.COSY (exclusive correlation spectroscopy) experiment (Schwalbe et al., 1994), where even small 1H-1H couplings can be measured. This experiment will also be adversely affected by broad lines, but because the measurement is not based on the absence of a peak, a more definitive measurement is possible. In the presence of severe spectral crowding, the HCCE.COSY-CCH TOCSY experiment, a variant which contains a TOCSY period to transfer magnetization to the well-resolved C1′, improves the spectral dispersion (Schwalbe et al., 1995; Glaser et al., 1996). An alternative method involves a collection of 1H-13C coupled HMQC-NOESY and HMQC-TOCSY spectra that are collected such that the sign of a number of 2JCH and 3JCH couplings can be determined and used to define the sugar pucker as shown in Table 7.7.3 (Hines et al., 1994). Often ribose groups are found with homonuclear H1′,H2′/H3′,H4′ coupling constants in the 3 to 6 Hz range, indicative of a conformation in between the C2′-endo and C3′-endo puckers. This mixed conformation is often left unrestrained, since it is averaging between the two major conformations (Aboul-ela et al., 1995; Brodsky and Williamson, 1997).
χ Torsion Recent Advances in RNA Structure Determination by NMR
Two heteronuclear vicinal 1H,13C couplings contain useful information about the glycosidic torsion angle χ. The 3JH1′C couplings involving the C4,C8 carbons in purines and the C2,C6 carbons in pyrimidines, respectively, all depend
on the χ torsion. The preferred orientation around χ in A-form helix is anti, which makes the base accessible for commonly found hydrogen-bonding interactions. The adopted conformation in RNA around χ strongly depends on the corresponding sugar pucker with the C3′endo puckers associated with the anti orientation. However, the different anti and syn conformations around the glycosidic torsion can usually be identified on the basis of intraresidual sugar-to-base, H1-3′-H8/H6, NOEs. The distance between H1′ and H8/H6 is shorter in the syn conformation, a corresponding strong NOE indicative for such a conformation.
ε and β Torsions
Considering the torsion angle β, the three low-energy staggered rotamers are characterized by a unique combination of vicinal 1H,31P and 13C,31P couplings. In a standard A-form helix, β is in the trans conformation (β = 180°) for which the 3JPH5′ and 3JPH5′′ are small and the 3JPC4′ and 3JPH4′ are large, as summarized in Table 7.7.3. Therefore, when one of the two 3JPH5 couplings is large, a gauche conformation (β = ± 60°) is present. This analysis of 3J PH5 couplings can also easily indicate the presence of conformational averaging as found in the HIV-2 TAR-argininamide complex where one of the nucleotides exhibited large couplings for both the 3JPH5′ and 3JPH5′′ (Brodsky and Williamson, 1997). A similar qualitative analysis exists for the ε torsion and associated staggered rotamers, which can be measured by monitoring the 3JC4′P, 3JC2′P, and 3JH3′P
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couplings. However, the 3JH3′P coupling does not change significantly for the different conformations and shows small values for both the trans (ε = 180°) and the gauche− (ε = −60°) conformers. Thus, in order to unambiguously define ε, the 3JC4′P and 3JC2′P need to be analyzed. The ε torsion is in the trans conformation in a standard A-form helix where the 3JC2′P coupling is small and the 3JC4′P coupling is large, as summarized in Table 7.7.3. The ε and β torsions can be determined by measuring a variety of 13C,31P and 1H,31P scalar couplings. Some of these torsions may be measured directly in 2-D 1H,31P heteronuclear COSY (HETCOR) experiments (Sklenar et al., 1986) and nonrefocused 1H,31P HSQCs if the phosphorus and proton resonances are sufficiently resolved; however, both the ribose proton and phosphorus resonances involved are generally overlapped for even moderate-sized RNAs. Accurate measurements for 13C,31P and 1H,31P couplings can be obtained from both phosphorus fitting of doublets from singlets (so called P-FIDS) (Schwalbe et al., 1993) or spin echo difference experiments (Legault et al., 1995; Hoogstraten and Pardi, 1998b; Szyperski et al., 1999). An extensive strategy has been introduced based on the intensity of cross-peaks in 3-D HCP and HPCH experiments (Varani et al., 1995). The intensity of cross-peaks in these spectra are proportional to the 3JPH5′, 3JPH5′′, and 3JPC4′ couplings. This approach suffers from the potential drawback that peak intensities may also be affected by other factors, such as dynamics. Recently, novel experiments for the measurement of 13C,31P and 1H,31P couplings using intensity modulations of cross-peaks due to coupling constant evolution in so-called quantitative J correlations have been introduced (Clore et al., 1998; Richter et al., 1998).
γ Torsion
Measurement of the γ torsion is difficult due to the need for stereospecific assignments of the H5′ and H5′′ proton resonances. The twobond C4′,H5′/H5′′ couplings can be used in conjunction with the vicinal H4′,H5′/H5′′ couplings to define γ (Hines et al., 1994; Schwalbe et al., 1994). Some couplings may be obtained from a collection of 1H,13C coupled versions of 3-D 13C-edited TOCSY and NOESY spectra (Hines et al., 1993, 1994). Unfortunately, both the C4′ and the H4′ in A-helical RNAs are extremely crowded, making correlations from these to the C5′/H5′ region, also showing poor dispersion, difficult. Therefore, experiments similar to the HCC-E.COSY-CCH TOCSY,
where the overlapped H4′-H5′ cross-peaks are correlated to the C1′/H1′, may be particularly useful if sufficient TOCSY transfer can be achieved (Schwalbe et al., 1994). In A-form helices, both the 3JH4′H5′ and 3JH4′H5′′ scalar couplings are small in the preferred staggered gauche+ (γ = 60°) conformation. Table 7.7.3 indicates that when γ changes to gauche− (γ = −60°) or trans (γ = 180°) conformations, either the 3JH4′H5′ or 3JH4′H5′′ coupling becomes larger; however, additional information is required to determine the stereospecific assignment. The sign of the 2JC4′H5′, 2JC4′H5′′, and 2JC5′H4′ couplings are very useful to help determine not only the stereospecific assignments but also the conformation as summarized in Table 7.7.3.
α and ζ torsions
Unfortunately, the α and ζ torsions are not accessible by J-coupling measurements because the involved 16O nuclei have no magnetic moment. Some groups have used 31P or 13C chemical shifts as a guide for loose constraints on these torsions; however, the correlation between 31P chemical shifts and the phosphodiester backbone conformation is not well understood in RNA. For example, in tRNA, a wide range of phosphorus chemical shifts are observed that are outside the ranges expected for variation of the α or ζ torsions, and other factors may be major contributors to phosphorous chemical shifts (Gueron and Shulman, 1975; Gorenstein and Luxon, 1979; Salemink et al., 1979). Furthermore, quantum calculations have shown that counterions also affect the chemical shift, further indicating that 31P chemical shifts may not be a reliable indicator of the conformation (Giessner-Prettre and Pullman, 1987). The understanding of different conformational contributions to the investigated C3′, C4′, and C5′ 13C chemical shifts remains incomplete (Xu et al., 1998); however, the use of heteronuclear chemical shifts in RNA structure calculations might have a greater impact in the future, especially in regions with low density of traditional restraints such as distances and torsion angles. Until a more complete chemical shift data base is created, restraints based on chemical shifts alone should probably be used with great caution.
The Value of Torsion Restraints in Structure Determination In protein NMR, torsion restraints help increase the precision of the structure, and it is expected that the precision of nucleic acid structures will also improve with the inclusion
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Table 7.7.3
Backbone J-Couplings in RNA
Angle
Rotamer
nJ-Coupling
J (Hz)
β
transa
H5′ - P(i) H5′′ - P(i) H4′ - P(i)b C4′ - P(i) H5′ - P(i) H5′′ - P(i) C4′ - P(i) H5′ - P(i) H5′′ - P(i) C4′ - P(i) H3′ - P(i+1) C2′ - P(i+1) C4′ - P(i+1) H3′ - P(i+1) H2′ - P(i+1)c C2′ - P(i+1) C4′ - P(i+1) H3′-P(i+1) H2′ - P(i+1)c C2′ - P(i+1) C4′ - P(i+1) H4′ - H5′ H4′ - H5′′ C4′ - H5′ C4 - H5′′ C5′ - H4′ H4′ - H5′ H4′ - H5′′ C4′ - H5′ C4 - H5′′ C5′ - H4′ H4′ - H5′ H4′ - H5′′ C4′ - H5′ C4 - H5′′ C5′ - H4′ H1′ - H2′ H2′ - H3′ H3′ - H4′ C1′ - H2′ C2′ - H3′ C3′ - H2′ C4′ - H3′ H1′ - H2′ H2′ - H3′ H3′ - H4′ C1′ - H2′ C2′ - H3′ C3′ - H2′ C4′ - H3′
1-4 1-3 ∼2 8-11 1-3 >15 <5 >15 1-3 <5 <5 <5 >10 <5 2-3 >10 <5 >15 2-3 <5 <5 1-3 3 − + + 3 9-11 + − − 9-11 5 − − − ∼1 ∼4 ∼9 − − + + ∼8 ∼4 ∼1 + + − −
gauche–
gauche+ ε
transa gauche−
gauche+
γd
gauche+ a
trans
gauche−
δd
C3′ endoa
C2′ endo
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aThe preferred rotamer populated in A-form helix. bThe 4J H4′P coupling can reach measurable values, ∼2 Hz, if β falls in the trans region and the corresponding torsion angle γ adopts a gauche+ conformation. cThe 4J H2′P coupling is ∼2 to 3 Hz, if ε adopts a gauche conformation. dThe characteristic sign patterns of involved 2J HC couplings contain particulary useful infor-
mation to restrict either the torsion γ or the sugar pucker.
Current Protocols in Nucleic Acid Chemistry
of torsion restraints. J-coupling restraints can be implemented in two different ways during the structure determination. They can be introduced qualitatively by restricting a torsion angle in a loose manner (±30°) to one of the three staggered rotamers along the phosphodiester backbone or defining the preferred ribose sugar pucker such as C2′-endo or C3′-endo. Alternatively, vicinal J-couplings can be quantitatively related to a certain torsion angle using parameterized Karplus relations (Karplus, 1959; Wijmenga and van Buuren, 1998; Marino et al., 1999). Preliminary studies indicate that only mild improvement is obtained with sufficiently large distance restraint data sets (Allain and Varani, 1995; Brodsky and Williamson, 1997). In fact, in the absence of any torsion restraints other than the sugar puckers, a structure with a root mean square deviation (rmsd) to the average of ∼1.2 Å can be defined with a large NOE restraint set for a 30-nucleotide RNA (Brodsky and Williamson, 1997). Inclusion of additional torsion restraints results in a modest improvement of the rmsd to ∼1.1 Å. Therefore, with a sufficient number of NOEs the structure may be reasonably well defined without torsion restraints that are increasingly difficult to measure for larger RNAs and RNA-protein complexes.
NOVEL PARAMETERS FOR LIQUID-STATE NMR STRUCTURE DETERMINATION In general, there is a practical difficulty in defining RNA structures precisely by NMR because NOE- and J-coupling-based structure calculation relies on either short-range distance (<6 Å) or local torsion angle information. Thus, RNAs often are elongated structures, that are better approximated as cylindrical rather than globular shapes. There is a lack of NOE information between distant ends of the molecule and, as a result, the relative orientations of helical segments at opposite ends of the molecule are poorly defined. There have been two recent advances in methodology that may help alleviate or overcome this shortcoming.
Relative Orientations of Bond Vectors Derived from Cross-Correlated Relaxation Cross-correlated relaxation rates have been recently introduced to high-resolution NMR as a tool for structure determination (Reif et al., 1997). For RNAs, these rates are measured using an experiment that belongs to the HCCH class, and result in the precise determination of
the ribose sugar pucker without the need of any empirical Karplus parameterization (Felli et al., 1999). For example, measurement of cross-correlated relaxation rates between neighboring 13C-1H dipoles within the ribose ring can be used to define the sugar pucker. The efficiency of cross-correlated relaxation in this case depends on 1⁄2(3cos2θ − 1), where θ is the projection angle between the two 13C-1H bond vectors. Thus, cross-correlated relaxation is most efficient for either a parallel (θ = 180°) or an orthogonal (θ = 90°) orientation of the two interactions and can also be zero in the case of θ ∼ 54.6°. The angle between 13C-1H bond vectors can be readily determined from HCCHtype cross-correlated relaxation measurements, and the resolution of this experiment can be further enhanced by combination with a CC-TOCSY transfer (Richter et al., 1999). The quantitative analysis of scalar J-couplings, especially in the case of homonuclear 3JHH couplings related to the ribose sugar pucker, becomes more and more difficult with increasing molecular weight. In contrast, the efficiency of cross-correlated relaxation pathways scales linearly with the overall correlation time of the molecule, which is related to its size. These new methods that exploit cross-correlated relaxation as a tool for structure determination should allow the characterization of conformations for larger RNA molecules, where J-coupling analysis is no longer feasible.
Residual Dipolar Couplings as a Probe for Long-Range Interactions In isotropic solutions, molecules are randomly oriented such that all orientations of a molecule with respect to the external magnetic field are equally probable. Since molecules undergo rapid random Brownian motion, dipolar couplings are not directly observable in NMR spectra because they are averaged to zero. Over the past few years, methods have been developed to create a slightly anisotropic environment for molecules tumbling in solution. This results in a small degree of alignment of the molecule, and the dipolar couplings no longer average to zero, while retaining the quality of high-resolution NMR spectra. The most promising systems for NMR studies of partially aligned systems redilute liquid crystalline bicelles (Tjandra and Bax, 1997a) or bacteriophage solutions (Hansen et al., 1998a). The reason for the preferred orientations of these macromolecules is interactions between the external magnetic field and orientation-dependent anisotropic magnetic susceptibility in
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the molecules. The huge rod-like phages, as well as the disk-like liquid crystalline bicelles, populate the orientational space in a nonuniform manner, and can thus induce alignment of dissolved RNAs. Bicelles used in high-resolution NMR studies are composed of mixtures of phospholipids such as dimyristoylphosphatidylcholine (DMPC) and lipids with detergentlike properties such as dihexanoylphosphatidylcholine (DHPC). They form an anisotropic liquid crystalline phase when prepared at 10 to 30 weight % lipid in aqueous solution over the 20° to 40°C range of temperatures (Prestegard, 1998). The filamentous phage Pf1 at concentrations between 10 and 60 mg/L also forms an aligned medium. The degree of alignment of the RNA molecule can be controlled by varying the concentration of the phage or bicelle solution. Higher phage or bicelle concentrations are associated with stronger alignments and produce larger residual dipolar couplings, while lower concentrations correspond to lower degrees of ordering, reflected in smaller dipolar couplings. There is a narrow useful range of alignments suitable for high-resolution NMR studies. Too much alignment gives larger dipolar couplings, but also results in line broadening to such an extent that high-resolution NMR is not possible. Dipolar couplings on the order of ±10 to 30 Hz can be introduced using phages or bicelles as cosolutes. In particular, dipolar couplings contain valuable structural information for the evaluation of the relative orientations of helices in RNA molecules. The size of dipolar couplings for an axially symmetric RNA molecule depends on the average value of an orientational function, 1⁄2(3cos2θ − 1), and the inverse cubic distance, 1/r3, between the coupled nuclei. Here, the angle θ characterizes the axial orientation of the internuclear vector that connects the coupled nuclei with respect to the principal axis system of the molecular alignment tensor. For a directly bonded pair of nuclei with known distance, such as 1H-13C or 1H-15N in labeled RNA, angular restraints can be extracted from dipolar coupling data and incorporated during the structure calculation (Tjandra et al., 1997). Such one-bond dipolar couplings can be measured in a straightforward and sensitive manner. The difference between scalar J coupling constant values measured in isotropic and anisotropic media gives the residual dipolar coupling. The simplest method for the measurement of these large one-bond couplings is the HSQC experiments recorded without proton decoupling. Several methods to measure 1JHC
or 1JHN more precisely have been published (Tjandra et al., 1996; Tjandra and Bax, 1997b). In addition, dipolar 1H,1H couplings are a potentially valuable source of long-range distance information (Hansen et al., 1998b). The transfer efficiency for dipolar couplings between protons falls off as a function of 1/r3 as opposed to the 1/r6 dependence of the NOE. Since those long-range interactions between dipolar coupled protons are obtainable in a straightforward manner, they can be measured using either COSY or TOCSY transfer schemes. Distances >5 Å could be easily measured for a 16-mer DNA duplex (Hansen et al., 1998b); however, since the interproton distance is not necessarily known in advance, the interpretation of the angular dependence for the interproton interaction is more complicated than in the case of the 1H-13C or 1H-15N vectors.
RNA STRUCTURES To generate a family of structures consistent with the NMR data, structures are traditionally refined against the distance and torsion restraints along with geometric and nonbonded terms using restrained molecular dynamics calculations (Brünger and Karplus, 1991; Nilges, 1996). A number of RNA structures have been determined utilizing many of the heteronuclear techniques discussed in this review. Many of the problems in RNA NMR studies revolve around the difficulty of gathering a large data restraint set due to chemical shift overlap along with dynamics issues. These issues also need to be considered during the molecular modeling process and for interpreting the resulting family of structures. Furthermore, the paucity of protons along the backbone leaves few distance restraints to define the many degrees of freedom of the RNA backbone. Neither distance nor J-coupling-based NMR methods are available for the direct determination of the backbone angles α and ζ. In general, traditionally applied restraints such as proton-proton distances or torsions accurately define local features of RNA structures, whereas global features like helical bending cannot be determined precisely using these restraint sets; however, parameters mentioned in the section above, such as residual dipolar couplings, hold the promise to provide global rather than local structural information for RNA structure determination. The data sets traditionally used as a restraint list in the structure determination include NOE and torsion restraints, and parameters such as cross-correlated relaxation rates and residual dipolar couplings have yet to be
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included in a published RNA structure determination.
Overall Quality of RNA NMR Structures In evaluating the quality of a family of RNA NMR structures, a number of statistics can be evaluated: rmsd, number of NOE and torsion restraints, residual distance and torsion violations, and the largest distance and torsion violations. The distance restraints are further dissected into the number of interresidue, intraresidue, and intermolecular NOEs. A listing of RNA NMR structures is shown in Table 7.7.4 with the rmsd’s shown in the context of the number of NOE and torsion restraints. The local rmsd is given because the overall global rmsd is usually in the 2.0 to 2.6 Å range, which might otherwise be indicative of poor convergence. This is because almost every RNA structure studied includes a region which is poorly defined, e.g., a disordered loop, terminal base pair, or a nucleotide without any internucleotide NOEs. This situation is comparable to protein NMR and X-ray studies, which often neglect the N- and C-terminal ends of proteins because of the lack of structural data from these regions. Therefore, the more useful rmsd to consider includes only the region of interest and is usually a more accurate description of the quality of the structure than the overall global rmsd. However, for RNA NMR studies, the region that is defined as the structured “core” can be a subjective decision, as the tendency is to find a combination of nucleotides which will give the lowest possible rmsd. One alternative is to define the well-ordered region by using an average standard deviation matrix which will identify the ordered and disordered regions (Kundrot, 1996). All the structures listed in Table 7.7.4 have low rmsd’s, indicating that a family of structures converged to a very similar structure, but the crucial question is whether the structures are accurate. This question was addressed in great detail using synthetic sets of restraints as derived from the ribozyme crystal structure (Allain and Varani, 1997). As in all structural biology techniques, correlations with other biochemical and structural data are used to help determine the accuracy of the structure. For RNA NMR studies, it is possible to make mutations of the RNA to determine whether any specific nucleotide alters the structure or complex, as has been done in a few cases (Puglisi et al., 1995; Ye et al., 1995; Dieckmann et al., 1996) to help support the proposed interactions.
In addition, the GNRA tetraloops (where N is any nucleotide and R is a purine) were first determined by NMR (Heus and Pardi, 1991) and have now been seen in both the Hammerhead crystal and P4/P5/P6 IVS crystal structures (Pley et al., 1994a,b; Cate et al., 1996). The GNRA tetraloops determined by NMR and X-ray crystallography are extremely similar and incorporate the same essential features. Furthermore, RNA NMR studies have been found to be very self-consistent, as all the general features and many of the specifics were very similar for the four structures that have been independently solved by two research groups.
Choosing a Molecular Modeling Protocol The goal of the molecular modeling calculations is to start from a sufficiently random initial model and then explore as much conformational space as possible. It is difficult to assess whether the protocols currently used are sampling all conformational space consistent with the data; therefore, this is an area of much study (Brünger and Karplus, 1991). Three major types of initial models are predominantly used as shown in Table 7.7.4: random coordinates, random torsions, and distance geometry. The major difference between these approaches is the computational expense required to generate a family of structures. Generally, random torsions offer high convergence rates in the 40% to 50% range (Aboul-ela et al., 1995; Allain and Varani, 1995; Allain et al., 1996; Jucker et al., 1996), while random coordinates require significant sorting of the atoms and generally generate lower convergence rates in the 10% to 20% range (Puglisi et al., 1995; Brodsky and Williamson, 1997). Distance geometry is an algorithm routinely used to sample conformational space in protein NMR studies with great success (de Vlieg and van Gunsteren, 1991). In RNA NMR studies, distance geometry has been found to be especially useful when many long-range, global fold type of NOEs are found as in peptide complexes and the highly folded ATP aptamer (Battiste et al., 1996; Dieckmann et al., 1996; Fan et al., 1996; Ye et al., 1996).
Determining Bounds for Distance Restraints A large difference in the number of NOEs per nucleotide is seen in the structures summarized in Table 7.7.4, in part because not all NOEs are always included in the distance re-
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straint data set for molecular modeling calculations. These apparent discrepancies occur because, although some intranucleotide NOEs are important in defining RNA structure, there are a number of intraribose and intrabase NOEs that do not contain any useful information, either because the range of sugar puckers do not change these distances significantly within the precision that the restraints can be accurately defined, or because the covalent structure of the nucleotide itself defines the restraints. The HIV-2 TAR argininamide NMR study did not use intrabase H5-H6 and H5-H4 NOEs (Brodsky and Williamson, 1997). Also, one of the BIV Tat-TAR and one of the ATP aptamer studies excluded many “conformationally unimportant” NOEs from the NOE restraint list, thus accounting for the small difference between the total number of NOEs and internucleotide NOEs shown in Table 7.7.3 (Puglisi et al., 1995; Dieckmann et al., 1996). Deciding what bounds to use for NOE restraints is a critical issue for RNA NMR structures. Modeling studies of protein structures using both synthetic and real data have demonstrated a correlation between the precision of a family of structures and the accuracy to a target structure (Liu et al., 1992). This correlation is critically dependent on the accuracy of the bounds used and the proper identification of NOEs; however, it is unclear whether this same correlation between rmsd and accuracy pertains to RNA structures which have fewer interresidue NOEs and lack the global fold, longrange NOEs often found in protein NMR studies. Unfortunately, the dynamic character of many RNA molecules on the NMR time scale prevents the accurate determination of tight bounds. For RNA NMR studies, NOE constraints are often determined semi-quantitatively and placed into four categories: strong, medium, weak, and very weak NOEs. Protein NMR studies have shown that with a sufficiently large restraint data set, a precise and accurate family of structures can be determined even when loose bounds are used (Liu et al., 1992). The RNA NMR studies listed in Table 7.7.4 have used different upper and lower bounds to define these categories. These differences lead to slightly different residual distance violation statistics, which may or may not reflect the quality of the structure. Each research group has used a slightly different methodology to determine the NOE bounds which also changes on a case-by-case basis depending on the quality of the spectra. The P1 helix, HIV TAR, and U1A studies set all the lower bounds
to 1.8 Å with upper bounds ranging from 3.0 Å for the most intense NOEs to 7.0 Å for the weakest NOEs found in H2O experiments (Aboul-ela et al., 1995; Allain and Varani, 1995; Allain et al., 1996; Brodsky and Williamson, 1997). One of the reasons for this very conservative approach is demonstrated by the extreme case of the HIV TAR studies where a weakbinding ligand caused a variety of conformational exchange issues. This precluded the use of accurate tight bounds such that independent groups used very conservative bounds (Aboulela et al., 1995; Brodsky and Williamson, 1997); however, this approach may be too conservative in some cases, as much of the information in the NOE data is not being used. An alternative approach is to use slightly tighter bounds where the lower bounds are set to 1.8, 2.5, and 3.5 Å, with upper bounds of 2.5, 3.5, and 5 Å for the strong, medium, and weak categories, respectively (Puglisi et al., 1995; Ye et al., 1995, 1996; Fan et al., 1996; Fourmy et al., 1996; Jucker et al., 1996). This strategy has been used in a number of cases with water NOESY spectra treated differently to account for possible spin diffusion effects of close amino protons. Both these approaches lead to a family of converged structures as shown in Table 7.7.4; however, when looser bounds were used, a significant number of restraints were used and thus presumably overcame the larger conformational space allowed by each bound as illustrated by the P1 helix, U1A, and HIV TAR studies (Aboul-ela et al., 1995; Allain and Varani, 1995; Allain et al., 1996; Brodsky and Williamson, 1997). The danger of using tight bounds is that errors can be made, some of which may lead to the wrong structure or can limit the rate of convergence, making the structure calculations difficult. Some techniques have been developed to help limit the amount of spin diffusion and therefore increase the accuracy of tight bounds by limiting the effects of spin diffusion, thus increasing the precision and presumably the accuracy of the resulting structures (Hoogstraten and Pardi, 1998a).
PROSPECTS FOR LARGE RNA MOLECULES Most NMR studies have focused on small hairpin elements (∼20 to 40 nucleotides) that contain unusual base-pairing elements and/or are the recognition sites for proteins or small molecules. Expanding the size of RNA molecules that can be studied by NMR to the 70 to 80 nucleotide range would greatly enhance the ability to study biologically significant RNAs.
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The two significant technical problems that must be overcome in order to study large macromolecules are the chemical shift overlap in spectra from the increased number of resonances and the increased transverse relaxation times that significantly reduce the sensitivity of multidimensional heteronuclear experiments. The first problem can be most easily overcome by the selective segmental and/or type-specific 13C/15N-labeling strategies outlined earlier, though strategies that allow more flexibility in the labeling patterns will probably be necessary. The relaxation problem is more fundamental and is difficult to overcome, but promising strategies have recently been developed.
Utilizing TROSY in RNA Assignments and Structure Determinations
Recent Advances in RNA Structure Determination by NMR
A new class of NMR experiments based on transverse relaxation-optimized spectroscopy (TROSY) promises a several-fold increase of the molecular size of RNA structures accessible (Pervushin et al., 1997; Wüthrich, 1998). Different relaxation mechanisms usually contribute to the rapid decay of magnetization during NMR experiments. The TROSY approach uses interference effects between different relaxation mechanisms in a constructive manner, resulting in partial cancellation of transverse relaxation effects, thus dramatically increasing the sensitivity of NMR. The efficiency of this mutual cancellation is a function of the spin pairs studied, the molecular weight, and the external magnetic field available. For example, for a 1H-15N spin pair in a molecule with a molecular weight of 150 kDa, nearly complete cancellation of magnetization decay can be achieved with a 900 MHz (proton resonance frequency) spectrometer, which will soon become available. Basically all NMR experiments that excite and detect imino proton resonances in RNA potentially benefit from the TROSY approach, which was originally proposed for 1H-15N amide moieties in proteins. Furthermore, recent experiments show significant gains in sensitivity for 1H-13C moieties in aromatic spin systems (Brutscher et al., 1998; Pervushin et al., 1998b; Meissner and Sorensen, 1999a). These applications have an even greater impact on the experiments commonly used for the investigation of RNA by NMR, because HCCH-type experiments for the assignment of the ribose resonances or HSQCtype experiments for either ribose or base 1H13C spin pairs potentially benefit, opening the field for NMR studies on larger RNA systems in the future. Finally, most recent studies indi-
cate sensitivity gains for the collection of NOE data, essential for a structure determination in NOESY experiments (Meissner and Sorensen, 1999b, 2000; Pervushin et al., 1999).
Sensitivity Enhancement Through Multiple Quantum Line Narrowing Two approaches can be applied to obtain one-bond 1H-13C correlations, HSQC or HMQC experiments. Recent studies show that the multiple quantum (MQ) experiments are more sensitive for correlating carbon and proton nuclei in RNA (Marino et al., 1997). The reason for this is similar to considerations given in the previous section. The main sources of relaxation for CH and CH2 moieties are not effective during multiple quantum evolution times; thus, MQ lifetimes are increased with respect to SQ (single quantum) coherences, present in HSQC experiments. The enhancement was demonstrated to be about a factor of 3 for a 36-mer RNA hairpin (Marino et al., 1997). This encouraging result has implications for through-bond assignment experiments, mentioned in earlier sections. Optimized HCN pulse schemes for the through-bond correlation of ribose and base resonances utilizing MQ instead of SQ evolution periods have been proposed and show significant sensitivity gains, essential for successful investigations of larger RNA systems (Fiala et al., 1998; Sklenar et al., 1998). In addition, specific deuteration of ribose or base resonances is likely to be beneficial. Full sensitivity can be retained at a particular position while still mitigating relaxation effects. New labeling patterns that incorporate both 13C and 2H will probably be needed, so that multidimensional experiments can be performed to resolve overlap problems. In addition, 13C labels in the ribose ring, along with deuteration of all the protons except at the H1′ position, has the potential to improve the feasibility of HCP sequential assignment experiments for larger RNA through increased efficiency of 13CTOCSY transfer (Dayie et al., 1998). To achieve a level of precision equivalent to that which has been commonly found in protein NMR of medium-sized (<20-kD) proteins, relatively more data is expected to be required and used in the structure determination process (Liu et al., 1992). Given the difficulties in acquiring any data for larger systems, this presents a particularly difficult situation. In RNA NMR, where good precision for even mediumsized (<40 nucleotide) RNAs is a difficult and tedious task, the challenge may be even greater.
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However, the new methods described above, especially dipolar couplings, hold the promise of great impact on the structure determination of even larger RNA systems. Perhaps dependence on other biochemical and biophysical techniques like transient electric birefringence and fluorescence resonance energy transfer measurements to gain additional structural restraints to aid in the structure determination process will become useful and necessary to solve large RNA structures.
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Fiala, R., Jiang, F., and Patel, D.J. 1996. Direct correlation of exchangeable and nonexchageable protons on purine bases in 13C,15N-labeled RNA using a HCCNH-TOCSY experiment. J. Am. Chem. Soc. 118:689-690.
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Foldesi, A., Nilsson, F.P.R., Glemarec, C., Gioeli, C., and Chattopadhyaya, J. 1992. Synthesis of 1′# , 2′, 3′, 4′#, 5′, 5′′-2H6-beta-D-ribonucleosides and 1′#, 2′, 3′, 4′#, 5′, 5′′-2H7-beta-D2′-deoxyribonucleosides for selective suppression of proton resonances in partially deuterated oligo-DNA, oligo-RNA and in 2,4A core (1HNMR window). Tetrahedron 48:9033-9072. Foldesi, A., Yamakage, S.-I., Nilsson, F.P.R., Maltseva, T.V., and Chattopadhyaya, J. 1996. The use of non-uniform deuterium labelling [“NMRwindow”] to study the NMR structure of a 21mer RNA hairpin. Nucl. Acids Res. 24:1187-1194. Fourmy, D., Recht, M.I., Blanchard, S.C., and Puglisi, J.D. 1996. Structure of the A site of Escherichia coli 16S ribosomal RNA complexed with an aminoglycoside antibiotic. Science 274:1367-1371. Giessner-Prettre, C. and Pullman, B. 1987. Quantum mechanical calculations of NMR chemical shifts in nucleic acids. Q. Rev. Biophys. 20:113172. Glaser, S.J., Schwalbe, H., Marino, J.P., and Griesinger, C. 1996. Directed TOCSY, a method for selection of directed correlations by optimal combinations of isotropic and longitudinal mixing. J. Magn. Reson. B112:160-180. Glemarec, C., Kukel, J., Foldesi, A., Maltseva, T., Sandstrom, A., Kirsebom, L.A., and Chattopadhyaya, J. 1996. The NMR structure of 31mer RNA domain of Escherichia coli RNase P RNA using its non-uniformly deuterium labelled counterpart [the “NMR-window” concept]. Nucl. Acids Res. 24:2022-2035. Gorenstein, D.G. and Luxon, B.A. 1979. High-resolution phosphorus nuclear magnetic resonance spectra of yeast phenylalanine transfer ribonucleic acid. melting curves and relaxation effects. Biochemistry 18:3796-3804. Gueron, M. and Leroy, J.L. 1995. Studies of base pair kinetics by NMR measurement of proton exchange. Methods Enzymol. 261:383-413. Gueron, M. and Shulman, R.G. 1975. 31P magnetic resonance of tRNA. Proc. Natl. Acad. Sci. U.S.A. 72:3482-3485. Hansen, M.R., Mueller, L., and Pardi, A. 1998a. Tunable alignment of macromolecules by filamentous phage yields dipolar coupling interactions. Nature Struct. Biol. 5:1065-1074. Recent Advances in RNA Structure Determination by NMR
Hansen, M.R., Rance, M., and Pardi, A. 1998b. Observation of long-range 1H-1H distances in solution by dipolar coupling interactions. J. Am. Chem. Soc. 120:11210-11211.
Heus, H.A., Wijmenga, S.S., van de Ven, F.J.M., and Hilbers, C.W. 1994. Sequential backbone assignment in 13C-labeled RNA via through-bond coherence transfer using three-dimensional triple resonance spectroscopy(1H, 13C, 31P) and twodimensional hetero TOCSY. J. Am. Chem. Soc. 116:4983-4984. Hines, J.V., Varani, G., Landry, S.M., and Tinoco, J.I. 1993. The stereospecific assignment of H5′ and H5′′ in RNA using the sign of two-bond carbon-proton scalar coupling. J. Am. Chem. Soc. 115:11002-11003. Hines, J.V., Landry, S.M., Varani, G., and Tinoco, J.I. 1994. Carbon-proton scalar couplings in RNA: 3D heteronuclear and 2D isotope-edited NMR of a 13C-labeled extra-stable hairpin. J. Am. Chem. Soc. 116:5823-5831. Hoogstraten, C.G. and Pardi, A. 1998a. Improved distance analysis in RNA using network-editing techniques for overcoming errors due to spin diffusion. J. Biomol. NMR 11:85-95. Hoogstraten, C.G. and Pardi, A. 1998b. Measurement of carbon-phosphorus J coupling constants in RNA using spin-echo difference constanttime HCCH-COSY. J. Magn. Reson. 133:236240. Hu, W., Kakalis, L.T., Jiang, L., Jiang, F., Ye, X., and Majumdar, A. 1998. 3D HCCH-COSY-TOCSY experiment for the assignment of ribose and amino acid side chains in 13C labeled RNA and protein. J. Biomol. NMR 12:559-564. Jiang, F., Kumar, R.A., Jones, R.A., and Patel, D.J. 1996. Structural basis of RNA folding and recognition in an AMP-RNA aptamer complex. Nature 382:183-186. Jucker, F.M. and Pardi, A. 1995. Solution structure of the CUUG hairpin loop: A novel RNA tetraloop motif. Biochemistry 34:14416-27. Jucker, F.M., Heus, H.A., Yip, P.F., Moors, E.H.M., and Pardi, A. 1996. A network of heterogeneous hydrogen bonds in GNRA tetraloops. J. Mol. Biol. 264:968-980. Karplus, M. 1959. Contact electron-spin coupling of nuclear magnetic moments. J. Chem. Phys. 30:11-15. Kay, L.E., Ikura, M., and Bax, A. 1990. Proton-proton correlation via carbon-carbon couplings: A three-dimensional NMR approach for the assignment of aliphatic resonances in proteins labeled with carbon-13. J. Am. Chem. Soc. 112:888-889. Kellogg, G.W. 1992. Proton-detected heteroTOCSY experiments with application to nucleic acids. J. Magn. Reson. 98:176-182.
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Kellogg, G.W. and Schweitzer, B.I. 1993. Two- and three-dimensional 31P-driven NMR procedures for complete assignment of backbone resonances in oligodeoxyribonucleotides. J. Biomol. NMR 3:577-95. Krishnan, V.V. and Rance, M. 1995. Influence of chemical exchange among homonuclear spins in heteronuclear coherence-transfer experiments in liquids. J. Magn. Reson. A A116:97-106. Kundrot, C. 1996. Rapid identification of ordered and disordered domains in NMR structures. J. Am. Chem. Soc. 118:8725-8726. Legault, P., Farmer, B.T., II, Mueller, L., and Pardi, A. 1994. Through-bond correlation of adenine protons in a 13C-labeled ribozyme. J. Am. Chem. Soc. 116:2203-2204. Legault, P., Jucker, F.M., and Pardi, A. 1995. Improved measurement of 13C, 31P J coupling constants in isotopically labeled RNA. FEBS Lett. 362:156-160. Liu, Y., Zhao, D., Altman, R., and Jardetzky, O. 1992. A systematic comparison of three structure determination methods from NMR data: Dependence upon quality and quantity of data. J. Biomol. NMR 2:373-388. Mao, H. and Williamson, J.R. 1999. Assignment of the L30-mRNA complex using selective isotopic labeling and RNA mutants. Nucl. Acids Res. 27:4059-4070. Mao, H., White, S.A., and Williamson, J.R. 1999. A novel loop-loop recognition motif in the yeast ribosomal protein L30 autoregulatory RNA complex [see comments]. Nature Struct. Biol. 6:1139-1147. Marino, J.P., Prestegard, J.H., and Crothers, D.M. 1994a. Correlation of adenine H2/H8 resonances in uniformly 13C labeled RNAs by 2D HCCHTOCSY: A new tool for 1H assignment. J. Am. Chem. Soc. 116:2205-2206. Marino, J.P., Schwalbe, H., Anklin, C., Bermel, W., Crothers, D.M., and Griesinger, C. 1994b. A three-dimensional triple-resonance 1H, 13C, 31P experiment: Sequential through-bond correlation of ribose protons and intervening phosphorous along the RNA oligonucleotide backbone. J. Am. Chem. Soc. 116:6472-6473. Marino, J.P., Schwalbe, H., Anklin, C., Bermel, W., Crothers, D.M., and Griesinger, C. 1995. Sequential correlation of anomeric ribose protons and intervening phosphorus in RNA oligonucleotides by a 1H,13C,31P triple resonance experiment: HCP-CCH-TOCSY. J. Biomol. NMR 5:87-92. Marino, J.P., Diener, J.L., Moore, P.B., and Griesinger, C. 1997. Multiple-quantum coherence dramatically enhance the sensitivity of CH and CH2 correlations in uniformly 13C-labeled RNA. J. Am. Chem. Soc. 119:7361-7366. Marino, J.P., Schwalbe, H., and Griesinger, C. 1999. J-coupling restraints in RNA structure determination. Acc. Chem. Res. 32:614-623.
Meissner, A. and Sorensen, O.W. 1999a. Optimization of three-dimensional TROSY-type HCCH NMR correlation of aromatic 1H-13C groups in proteins. J. Magn. Reson. 139:447-450. Meissner, A. and Sorensen, O.W. 1999b. Suppression of diagonal peaks in TROSY-type 1H NMR NOESY spectra of 15N-labeled proteins. J. Magn. Reson. 140:499-503. Meissner, A. and Sorensen, O.W. 2000. Three-dimensional protein NMR TROSY-type (15)N-resolved (1)H(N)-(1)H(N) NOESY spectra with diagonal peak suppression. J. Magn. Reson. 142:195-198. Mohebbi, A. and Shaka, A.J. 1991. Improvements in carbon-13 broadband homonuclear cross-polarization for 2D and 3D NMR. Chem. Phys. Lett. 178:374-378. Mueller, L., Legault, P., and Pardi, A. 1995. Improved RNA structure determination by detection of NOE contacts to exchange-broadened amino protons. J. Am. Chem. Soc. 117:1104311048. Nikonowicz, E.P. and Pardi, A. 1992. Three-dimensional heteronuclear NMR studies of RNA. Nature 355:184-186. Nikonowicz, E.P. and Pardi, A. 1993. An efficient procedure for assignment of the proton, carbon and nitrogen resonances in 13C/15N labeled nucleic acids. J. Mol. Biol. 232:1141-1156. Nikonowicz, E.P., Sirr, A., Legault, P., Jucker, F.M., Baer, L.M., and Pardi, A. 1992. Preparation of 13 C and 15N labelled RNAs for heteronuclear multi-dimensional NMR studies. Nucl. Acids Res. 20:4507-4513. Nilges, M. 1996. Structure calculation from NMR data. Curr. Opin. Struct. Biol. 6:617-623. Otting, G. and Wüthrich, K. 1989a. Extended heteronuclear editing of 2D 1H NMR spectra of isotope-labeled proteins, using the X(ω1, ω2) double half filter. J. Magn. Reson. 85:586-594. Otting, G. and Wüthrich, K. 1989b. Studies of protein hydration in aqueous solution by direct NMR observation of individual protein-bound water molecules. J. Am. Chem. Soc. 111:18711875. Otting, G. and Wüthrich, K. 1990. Heteronuclear filters in two-dimensional [1H,1H]-NMR spectroscopy: Combined use with isotope labelling for studies of macromolecular conformation and intermolecular interactions. Q. Rev. Biophys. 23:39-96. Otting, G., Liepinsh, E., and Wüthrich, K. 1991. Protein hydration in aqueous solution. Science 254:974-980. Pardi, A. 1995. Multidimensional heteronuclear NMR experiments for structure determination of isotopically labeled RNA. Methods Enzymol. 261:350-380. Pardi, A. and Nikonowicz, E.P. 1992. Simple procedure for resonance assignment of the sugar protons in 13C-labeled RNAs. J. Am. Chem. Soc. 114:9202-9203.
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Pervushin, K., Riek, R., Wider, G., and Wüthrich, K. 1997. Attenuated T2 relaxation by mutual cancellation of dipole-dipole coupling and chemical shift anisotropy indicates an avenue to NMR structures of very large biological macromolecules in solution. Proc. Natl. Acad. Sci. U.S.A. 94:12366-71. Pervushin, K., Ono, A., Fernandez, C., Szyperski, T., Kainosho, M., and Wüthrich, K. 1998a. NMR scalar couplings across Watson-Crick base pair hydrogen bonds in DNA observed by transverse relaxation-optimized spectroscopy. Proc. Natl. Acad. Sci. U.S.A. 95:14147-14151. Pervushin, K., Riek, R., Wider, G., and Wüthrich, K. 1998b. Transverse relaxation-optimized spectroscopy (TROSY) for NMR studies of aromatic spin systems in 13C-labeled proteins. J. Am. Chem. Soc. 120:6394-6400. Pervushin, K., Wider, G., Riek, R., and Wüthrich, K. 1999. The 3D NOESY-[1H,15N,1H]-ZQTROSY NMR experiment with diagonal peak suppression. Proc. Natl. Acad. Sci. U.S.A. 96:9607-9612. Pley, H., Flaherty, K.M., and McKay, D.B. 1994a. Three-dimensional structure of a hammerhead ribozyme. Nature 372:68-74. Pley, H.W., Flaherty, K.M., and McKay, D.B. 1994b. Model of an RNA tertiary interaction from the structure of an intermolecular complex between a GAAA tetraloop and an RNA helix. Nature 372:111-113. Prestegard, J.H. 1998. New techniques in structural NMR—Anisotropic interactions. Nature Struct. Biol. 5:517-522. Puglisi, J.D., Chen, L., Blanchard, S., and Frankel, A.D. 1995. Solution structure of a bovine immunodeficiency virus tat TAR RNA-peptide complex. Science 270:1200-1203. Quant, S., Wechselberger, R.W., Wolter, M.A., Wörner, K.-H., Schell, P., Engels, J.W., Griesinger, C., and Schwalbe, H. 1994. Chemical synthesis of 13C-labelled monomers for the solidphase and template controlled enzymatic synthesis of DNA and RNA oligomers. Tetrahedron Lett. 35:6649-6652. Reif, B., Hennig, M., and Griesinger, C. 1997. Direct measurement of angles between bond vectors in high-resolution NMR. Science 276:12301233. Richter, C., Reif, B., Wörner, K.-H., Quant, S., Marino, J.P., Engels, J.W., Griesinger, C., and Schwalbe, H. 1998. A new experiment for the measurement of nJ(C,P) coupling constants including 3J(C4′i,Pi) and 3J(C4′i,Pi+1) in oligonucleotides. J. Biomol. NMR 12:223-230.
Recent Advances in RNA Structure Determination by NMR
Richter, C., Griesinger, C., Felli, I.C., Cole, P.T., Varani, G., and Schwalbe, H. 1999. Determination of sugar conformation in large RNA oligonucleotides from analysis of dipole-dipole cross-correlated relaxation by solution NMR spectroscopy. J. Biomol. NMR 15:241-250.
Salemink, P.J.M., Swarthof, T., and Hilbers, C.W. 1979. Studies of yeast phenylalanine-accepting transfer ribonucleic acid backbone structure in solution by phosphorous-31 nuclear magnetic resonance spectroscopy. Biochemistry 18:34773485. SantaLucia, J., Shen, L.X., Cai, Z., Lewis, H., and Tinoco, I. 1995. Synthesis and NMR of RNA with selective isotopic enrichment in the bases. Nucl. Acids Res. 23:4913-4921. Schwalbe, H., Samstag, W., Engels, J.W., Bermel, W., and Griesinger, C. 1993. Determination of 3J(C,P) and 3J(H,P) coupling constants in nucleotide oligomers with FIDS-HSQC. J. Biomol. NMR 3:479-486. Schwalbe, H., Marino, J.P., King, G.C., Wechselberger, R., Bermel, W., and Griesinger, C. 1994. Determination of a complete set of coupling constants in 13C-labeled oligonucleotides. J. Biomol. NMR 4:631-644. Schwalbe, H., Marino, J.P., Glaser, S.J., and Griesinger, C. 1995. Measurement of H,H-coupling constants associated with v1, v2, and v3 in uniformly 13C labeled RNA by HCC-TOCSYCCH-E. COSY. J. Am. Chem. Soc. 117:72517252. Scott, W.G., Finch, J.T., and Klug, A. 1995. The crystal structure of an all-RNA hammerhead ribozyme: A proposed mechanism for RNA catalytic cleavage. Cell 81:991-1002. Simorre, J.-P., Zimmermann, G.R., Pardi, A., Farmer, B.T., II, and Mueller, L. 1995. Triple resonance HNCCCH experiments for correlating exchangeable and nonexchangeable cytidine and uridine base protons in RNA. J. Biomol. NMR 6:427-432. Simorre, J.P., Zimmermann, G.R., Mueller, L., and Pardi, A. 1996a. Correlation of the guanosine exchangeable and nonexchangeable base protons in 13C-/15N-labeled RNA with an HNCTOCSY-CH experiment. J. Biomol. NMR 7:153156. Simorre, J.-P., Zimmermann, G., Mueller, L., and Pardi, A. 1996b. Triple-resonance experiments for assignment of adenine base resonances in 13 15 C/ N-labeled RNA. J. Am. Chem. Soc. 118:5316-5317. Sklenar, V. and Bax, A. 1987. Spin echo water suppression for the generation of pure phase two-dimensional NMR spectra. J. Magn. Reson. 74:469-479. Sklenar, V., Miyashiro, H., Zon, G., Miles, H.T., and Bax, A. 1986. Assignment of the 31P and 1H resonances in oligonucleotides by two-dimensional NMR spectroscopy. FEBS Lett. 208:9498. Sklenar, V., Brooks, B.R., Zon, G., and Bax, A. 1987. Absorption mode two-dimensional NOE spectroscopy of exchangeable protons in oligonucleotides. FEBS Lett. 216:249-252.
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Sklenar, V., Peterson, R.D., Rejante, M.R., and Feigon, J. 1993a. Two- and three-dimensional HCN experiments for correlating base and sugar resonances in 15N,13C-labeled RNA oligonucleotides. J. Biomol. NMR 3:721-7. Sklenar, V., Peterson, R.D., Rejante, M.R., Wang, E., and Feigon, J. 1993b. Two-dimensional tripleresonance HCNCH experiment for direct correlation of ribose H1′ and base H8, H6 protons in 13 15 C, N-labeled RNA oligonucleotides. J. Am. Chem. Soc. 115:12181-12182. Sklenar, V., Peterson, R.D., Rejante, M.R., and Feigon, J. 1994. Correlation of nucleotide base and sugar protons in a 15N-labeled HIV-1 RNA oligonucleotide by 1H-15N HSQC experiments. J. Biomol. NMR 4:117-22. Sklenar, V., Dieckmann, T., Butcher, S.E., and Feigon, J. 1996. Through-bond correlation of imino and aromatic resonances in 13C, 15N-labeled RNA via heteronuclear TOCSY. J. Biomol. NMR 7:83-87. Sklenar, V., Dieckmann, T., Butcher, S.E., and Feigon, J. 1998. Optimization of triple-resonance HCN experiments for application to larger RNA oligonucleotides. J. Magn. Reson. 130:119-124.
Tolbert, T.J. and Williamson, J.R. 1996. Preparation of specifically deuterated RNA for NMR studies using a combination of chemical and enzymatic synthesis. J. Am. Chem. Soc. 118:7929-7940. Varani, G. and Tinoco, J.I. 1991. RNA structure and NMR spectroscopy. Q. Rev. Biophys. 24:479532. Varani, G., Aboul-ela, F., Allain, F., and Gubser, C.C. 1995. Novel three-dimensional 1H-13C-31P triple resonance experiments for sequential backbone correlations in nucleic acids. J. Biomol. NMR 5:315-320. Varani, G., Aboul-ela, F., and Allain, F. H.-T. 1996. NMR investigation of RNA structure. Prog. Nucl. Magn. Reson. Spectrosc. 29:51-127. Wijmenga, S.S. and van Buuren, B.N.M. 1998. The use of NMR methods for conformational studies of nucleic acids. Prog. Nucl. Magn. Reson. Spectrosc. 32:287-387. Wijmenga, S.S., Heus, H.A., Leeuw, H.A.E., Hoppe, H., van der Graaf, M., and Hilbers, C.W. 1995. Sequential backbone assignment of uniformly 13C-labeled RNAs by a two-dimensional P(CC)H-TOCSY triple resonance NMR experiment. J. Biomol. NMR 5:82-86.
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Xu, J., Lapham, J., and Crothers, D.M. 1996. Determining RNA solution structure by segmental isotopic labeling and NMR: Application to Caenorhabditis elegans spliced leader RNA 1. Proc. Natl. Acad. Sci. U.S.A. 93:44-48. Xu, X.-P., Chiu, W.-L.A.K., and Au-Yeung, S.C.F. 1998. Chemical shift and structure relationship in nucleic acids: Correlation of backbone torsion angles γ and α with 13C chemical shifts. J. Am. Chem. Soc. 120:4230-4231. Ye, X., Kumar, R.A., and Patel, D.J. 1995. Molecular recognition in the bovine immunodeficiency virus tat peptide TAR RNA complex. Chem. Biol. 2:827-840. Ye, X., Gorin, A., Ellington, A.D., and Patel, D.J. 1996. Deep penetration of an a-helix into a widened RNA major groove in the HIV-1 rev peptide-RNA aptamer complex. Nature Struct. Biol. 3:1026-1033. Zhang, X., Gaffney, B.L., and Jones, R.A. 1998. 15N NMR of RNA fragments containing specifically labeled tandem GA pairs. J. Am. Chem. Soc. 120:6625-6626.
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Contributed by M. Hennig and J.R. Williamson The Scripps Research Institute La Jolla, California A.S. Brodsky Dana-Farber Cancer Institute Boston, Massachusetts J.L. Battiste Harvard Medical School Boston, Massachusetts We thank all the scientists with whom we have interacted and have enjoyed fruitful discussions in helping us learn and think about RNA NMR. This work was supported by a grant from the NIH (GM-47467). M. Hennig acknowledges support of the Human Frontier Science Program.
Recent Advances in RNA Structure Determination by NMR
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Molecular Modeling of Nucleic Acid Structure: Energy and Sampling
UNIT 7.8
Building on the introduction to molecular modeling of nucleic acid structure presented in UNIT 7.5, this unit discusses computational and theoretical methods aimed at giving greater insight into the structure and energy of nucleic acids. This includes describing means for representing the energy in model nucleic acid structures and methods for sampling relevant conformational states. THE ENERGY REPRESENTATION To aid a modeler in judging the reliability of a given model structure, it is useful to include some representation of the energy. The presumption is that structures with lower energies—such as those with less steric overlap, less distortion, and more favorable interactions—will be more representative. As discussed (UNIT 7.5), adding a representation of the energy to a physical model is difficult; however, some implementation of the energy for a given molecular model can be easily programmed on a computer. In common use, the level of theory applied spans the range from highly accurate ab initio quantum mechanical (QM) to simpler, empirical molecular mechanical energy treatments. As the accuracy of the energy increases, so does the computational cost. The increase in computational cost is tremendous and limits the level of theory that may be applied. For calculating the energy of a single model structure, very accurate methods can be applied; however, if one is interested in dynamics or investigating the energy of many configurations of a given model, less accurate energy representations are generally necessary. Quantum Mechanical (QM) Treatments To provide an accurate and complete theoretical description of the energy (as a function of the atomic and electronic configuration or structure) of a molecular system, ab initio QM treatments can be applied. Standard codes for performing QM calculations include Gaussian, Jaguar, Q-Chem, and GAMESS (see Internet Resources). Performing calculations with these programs is fairly straightforward, even for those without a broad theoretical background in the methods. For reviews of QM methods, see the books by Szabo and Ostlund (1989) or Levine (1991). A serious drawback of these methods is that they are extremely computationally demanding, which typically limits accurate QM calculations to small model systems (<100 atoms). Recent improvements in the methods, coupled with the availability of greater computational power, have led to more involved and highly accurate QM calculations. Accuracy is improved with the use of larger and higher level basis sets and with the inclusion of electron correlation. More recently there has also been an increase in the use of density functional methods, which allow the investigation of larger systems (and implicitly include some electron correlation); however, even with these improvements, tractable model systems are limited to individual base pairs or stacked nucleotides. The effect of the solvent, if included at all, can only be included implicitly (via a mean field or continuum treatment) or through the inclusion of a very small number of explicit waters around the molecule in the QM calculation. Prior to using these methods to investigate nucleic acid models, it would be wise to consult the detailed literature for similar examples. Some recent examples of QM treatments of nucleic acids include the investigation of base-pairing energetics, base stacking, the interaction of metal ions with base pairs, and even base solvation in simulations of individual bases surrounded by a small amount of explicit water (for review see Hobza and Sponer, 1999). Very few QM calculations have been applied directly in macromolecuContributed by Thomas E. Cheatham, III, Bernard Brooks, and Peter A. Kollman Current Protocols in Nucleic Acid Chemistry (2001) 7.8.1-7.8.20 Copyright © 2001 by John Wiley & Sons, Inc.
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lar modeling applications. An exception to this is the critical role of QM methods in the development, evaluation, and critique of empirical potential functions (as are introduced later in this section); this probably represents the largest use of QM methods in nucleic acid modeling. As mentioned in UNIT 7.5, although the QM treatments have the potential to provide a very high level of accuracy, this level of accuracy is not always required, and faster less-accurate techniques may be appropriate; however, the commonly applied approximations come at a cost. Without a QM treatment, it is generally not possible to accurately represent processes that involve chemical changes (i.e., chemical reaction, bond breaking, or bond forming), excited states (i.e., electronic transitions), or electron transfer. In practice this is not a major limitation, since with nucleic acids the investigator is most often simply interested in the structure, dynamics, and relative importance of a given model. The middle ground between pure ab initio QM techniques and the faster empirical potentials (discussed in the next section; see Molecular Mechanics: Empirical Potential Energy Functions) are semi-empirical treatments. These apply a quantum mechanical formalism where significant approximations are applied in the calculation to decrease the computational cost, while accuracy lost through the approximations is offset by the addition of empirical parameters. The semi-empirical methods allow investigation of ~2500 atoms for geometry optimization or ~100 atoms for dynamics. Their use in biomolecular simulations has been limited in part due to the difficulty in properly representing hydrogen-bonded systems; however, the future holds promise with the development of semi-empirical parameterizations for biomolecular systems, better methods for treating large systems (Thiel, 1997), and semi-empirical codes designed to run on massively parallel computers (Dixon and Merz, 1997). Standard semi-empirical codes include MOPOC, MNDO4, AMSOL, Argus, and ZINDO (see Internet Resources) among others, and utilize various parameterizations including AM1, PM3, and MNDO. Reviews on semi-empirical methods discuss the codes and parameterizations in greater detail (Stewart, 1990; Zerner, 1991). Despite the higher levels of accuracy possible with the quantum mechanical and semi-empirical treatments, the most commonly applied treatment of energy uses simplified molecular mechanical (MM) potential energy functions. Given an appropriate parameterization, these MM potential functions are able to accurately reproduce nucleic acid structure at the atomic level in a very computationally efficient manner. This allows rapid evaluation of not only energy, but forces (or the first derivative of the energies with respect to the positions), thereby allowing analysis of many configurations in a reasonable time frame.
Molecular Modeling of Nucleic Acid Structure: Energy and Sampling
Before leaving this discussion of ab initio and semi-empirical methods, it is important to mention that there has been significant progress in the development of hybrid methods. These treat part of the system quantum mechanically (the part undergoing chemical change) and the remainder with a significantly faster molecular mechanical (empirical) potential. The hybrid QM/MM treatments allow representation of larger systems (such as enzymes) with explicit representation of the environment, while still treating the region of chemical interest (such as the active site) quantum mechanically. Drawbacks of these methods are that they are extremely computationally demanding (depending in large part on the size of the QM region) and that there are a number of open research questions, such as how best to merge the QM and MM regions, how best to decide what part of the system should be represented quantum mechanically, and what level of treatment to apply in the QM region (i.e., ab initio versus specifically parameterized semi-empirical) or MM regions (Field et al., 1990; Stanton et al., 1995; Gao, 1996; Cummins and Gready, 1997; Chatfield et al., 1998). Although most of the QM/MM applications have been limited to enzyme systems or small molecules in solution, a recent application involved model
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reactions for ribozymes (Lahiri and Nilsson, 1997). Code for performing QM/MM is in many of the standard molecular dynamics programs, including AMBER (Pearlman et al., 1995), CHARMM (Brooks et al., 1983), and GROMOS (van Gunsteren and Berendsen, 1987). Molecular Mechanics: Empirical Potential Energy Functions The most commonly applied methods for describing energy use an empirically derived or molecular mechanics (MM) potential function. This involves the application of a simplified potential function that has been parameterized to properly model the structures of interest. The specific parameterization, or force field, needs to represent not only the intramolecular interactions (based on the covalent structure of the molecule) but the intermolecular interactions between all the atoms and molecules. Most of the commonly applied empirical derived force fields describe the intermolecular interactions in a similar manner. The complete and true atomic energy representation, U(r1, r2,...., rN), involves the interactions of all N atoms (at positions r1 to rN) in the system. Following Allen and Tildesley (1987), this energy representation can be decomposed into a sum of pairs, triples, quadruples, and higher interactions between atom centers: N
N N
N N
U (r1,r2, . . . , rN) = ∑U (ri) + ∑ ∑ U (ri,rj) + ∑ ∑ i=1
i=1 j>i
N
∑ U (ri,rj,rk) + . . .
i=1 j>i k>j>i
Equation 7.8.1
Approximations are then applied to simplify the representation. Given that the individual interactions for the terms from the quadrupolar interaction and higher-order interactions are usually very small, these are often neglected. The first-order term, U(ri), involves interactions with external fields and, therefore, is also generally not included. This leaves the dominant terms, specifically the pair interactions (a function of the distance between the atoms; rij) and the three-body (atom-dipolar) interactions. The three-body interactions are less often included explicitly in biomolecular simulation at present due to the increased cost of calculation and the nonadditivity of the energy; however, these interactions are not completely neglected since their effect is implicitly included in the molecular mechanical force field during the explicit parameterization of the pairwise interactions. To summarize, the most commonly applied empirical force fields use an additive pairwise potential (with the nonadditivity and three-body terms omitted). Intramolecular Interactions The intramolecular interactions describe the covalent structure of the molecule. This includes the bonds, angles, dihedrals, and overall connectivity and flexibility of the model. A reasonable MM representation can be obtained either with full atomic or internal coordinate representations. Atomic representations imply that each atom center is allowed to move independently and explicit bond, angle, and dihedral terms are added to represent the connectivity. Two common representations are applied: all-atom force fields treat each atom as independent, whereas united-atom force fields fold multiple atomic centers into a single particle, such as treating a methylene group (–CH2–) as a single center. Internal coordinate representations, on the other hand, attempt to represent the inherent motions and flexibility of the system via rotations about naturally rigid groups of atoms. These can be useful since there are effectively fewer degrees of freedom, with flexibility only included where necessary to represent the natural motion of the molecule. For example, with nucleic acids the bases can often be treated as rigid units, as can most of the backbone (except for ε and ζ), with rotations and translations set up to represent deviations from a common helical axis system, such as rotations involving inclination,
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tip and twist, and translations (e.g., x- and y-displacement from the helical axis and rise between base pairs). To include the flexibility of the sugar pucker, the bonds at the C4′-O4′ atoms can be broken, leading to two torsions and three angles to describe the ring. This internal coordinate representation is used in the program JUMNA (junction minimization of nucleic acids; Lavery et al., 1995). Alternative treatments may use different rotations (e.g., twist, roll, and tilt; Gorin et al., 1990), more complicated treatments (Nesterova et al., 1997), or could (as is done in protein simulations) consider the bonds and angles as fixed while allowing free rotation around all the torsions. Ideally, a good internal coordinate representation will attempt to minimize the number of degrees of freedom while still retaining good structure and dynamics. Reduction of the number of degrees of freedom is desirable since this has clear benefits when trying to sample the possible conformations (as will become more apparent later). Although the internal coordinate treatments lead to fewer degrees of freedom, using this kind of representation in molecular dynamics simulations is difficult because accurate treatment requires inverting the moment of inertia tensor (or mass matrix) or the application of computationally demanding holonomic constraints (that maintain the fixed structure within rigid groups and properly equalize or propagate the forces). In all-atom or united-atom treatments, because each atom center is free to move, the mass matrix is diagonal and thus, trivially invertable. Even though there are approximations that effectively treat the moment of inertia tensor when it is not diagonal with order N rather than N3, accuracy is lost. Based on these efficiency issues, internal coordinate treatments are generally only used with minimization or Monte Carlo simulation (as described more fully in the next section). A further potential difficulty with the internal coordinate representation is that it requires specification of the rigid units; if the unit is rigid, it cannot distort structurally, in contrast to what might be the expected behavior under certain conditions. Thus, care has to be taken not to rigidify a part of the molecule that may not be rigid in practice. Moreover, rigid rotation about a given bond effectively leads to higher rotational barriers since there is no coupling to other modes (i.e., the bonds or angles cannot open up to facilitate rotation). For example, the gauche, gauche rotational barrier for butane is roughly twice as large when the bonds and angles are held rigid. Of course, this artifact is not a problem in practice since the force fields are parameterized to compensate; this just points out that it is not possible to directly mix intramolecular force fields designed for use with internal coordinates with those designed for use in all-atom treatments. The alternative to using an internal coordinate treatment is to not treat any of the internal coordinates as rigid, so that each atom is free to move. In this case, to represent the covalent structure and intramolecular energetics, explicit energetic representations for the bonds, angles, and dihedrals need to be added. In the simplest form, this is typically done with harmonic potentials to maintain bond lengths and angles, and Fourier terms for the torsion angles to represent rotation about bonds (Figure 7.8.1). Either harmonic or Fourier terms are also commonly added to maintain planarity or prevent rotation about double bonds. Higherorder terms can also be added as necessary for more detailed representation. A common form for the intramolecular part of the mechanical potential function is as follows: Uintramolecular = ∑ kb(r − req)2 + ∑ kθ(θ − θeq)2 + ∑ bonds
angles
∑
dihedrals η
Vη 2
[1 + cos(ηφ − γ)] + . . .
Equation 7.8.2 Molecular Modeling of Nucleic Acid Structure: Energy and Sampling
The parameterization, or force field, refers to the specific equilibrium geometry values for the various terms (such the equilibrium bond length, req) and the force constants representing the energy (or vibrational frequency, kb) of distortion away from the equi-
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Figure 7.8.1 Schematic of the interactions in a pairwise additive molecular mechanics force field.
librium geometry. The parameterization is typically performed on molecular fragments with the implicit assumption that the force field parameters are transferable to other fragments. In other words, a carbon-carbon bond in propane is the same as a carbon-carbon bond in pentane. Bond lengths and bond angles are determined via experiment (crystallography or other spectroscopic techniques) with force constants for the vibration inferred from microwave, infrared (IR), and other spectroscopic data. In the parameterization, the least well-determined parameters (based on experimental information) are the dihedral terms, since these terms effectively include not only the equilibrium torsion value but 1-4 atom interactions and other implicit interactions explicitly omitted. Given this, the dihedral part of the force field is usually the last part to be parameterized (based on QM and empirical data) and modification of these parameters can be used to fix up deficiencies from the other intramolecular and intermolecular interactions. More complex molecular mechanical representations are also possible, such as those including cubic and quartic terms for bond stretching, cubic angle terms for anharmonic bending, bond-torsion, angle-torsion, bend-bend, and other terms. Although these higherorder terms can aid in parameterization efforts to better represent vibrational spectra, structure, and heats of formation in a diverse set of molecules (including strained molecules), these are not typically included in biomolecular force fields. This is because biomolecular force fields are primarily parameterized to represent structure and secondarily relative conformational energetics in as simple and transferable a means as possible. An adequate representation is obtained without the added complexity, as will become apparent. For more information about the more strongly parameterized class 2 and class
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3 force fields that include the more complicated intramolecular energy representation, see discussions of the following force fields: MMFF94 (Halgren, 1996), MM3 and MM4 (Allinger et al., 1989, 1996), and QMFF (Maple et al., 1994). Intermolecular Interactions The standard form for the intermolecular pair energy involves a Lennard-Jones potential to represent the electron cloud repulsion (rij−12), dispersion attraction (rij−6) interactions, and a Coulombic term (with atom point charges qi and qj, and dielectric constant ε) representing the electrostatic interactions between all the atom pairs (where rij is the distance between atoms i and j). Aij Uintermolecular = 12 − rij
Bij r6ij
+
qiqj εrij
Equation 7.8.3
Note that the dielectric constant (ε) is shown in a simplified form that implicitly includes the 4πε0 leading factor, where ε0 is the permittivity of free space. Also note that self-interactions (1-1), interactions between bonded atoms (1-2), and interactions between the atoms involved in angles (1-3) are most often omitted, since their interaction has already been included in the intramolecular part of the force field. Interactions between terminal atoms involved in a dihedral angle (1-4 interactions) are sometimes scaled. Polarization effects are also often included implicitly in the parameterization, although explicit polarization (as is discussed later) can be included at additional cost. The specific parameters (Aij, Bij, qi, qj) in large part determine the reliability of the intermolecular potential. There are many philosophies regarding how to “best” derive these parameters, ranging from total reliance on high-level QM treatments of fragments to parameterization based entirely on empirical data, or some combination of each. A recent trend has been to derive the Lennard-Jones parameters for particular atom types from simulations of neat liquids (Jorgensen et al., 1996). Like with the bond, angle, and dihedral parameters, there is an implicit assumption that (in general) the Lennard-Jones parameters are transferable. It should be noted that the parameters Aij and Bij for the Lennard-Jones part of the equation above represent mixed van der Waal parameters for atoms i and j. Since the literature is sometimes confusing with presentation of the mixed parameters as Aij and Bij (with or without pre-exponentiation) or in terms of r* (the minimum of the potential well in Å) or σ (the zero of the potential in Å), and since this is further complicated by application of different combining rules, it is worth a brief digression. The two forms of the van der Waals energy in terms of r* and σ are as follows: ∗ 12 12 6 r∗ 6 r σ σ ELJ = 4ε − = ε − 2 r rij rij rij ij Equation 7.8.4
This implies that: 2 r∗ = σ√ 6
Equation 7.8.5 *
Molecular Modeling of Nucleic Acid Structure: Energy and Sampling
Given an r value (representing the van der Waal radius) for two atoms i and j that are not the same, it is necessary to define a mixed van der Waal radius. There are two methods in common usage, arithmetic or Lorentz-Bertholet combining rules where:
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r∗ij
r∗i + r∗j = 2
Equation 7.8.6
(as applied in AMBER and CHARMM) and geometric mean combining rules where:
r∗ij = √ r∗i r∗j Equation 7.8.7
(as applied in GROMOS). In both cases, a geometric mean is used for the well depth. These differences again point to the critical need to be careful when trying to adapt force fields applied in one program for use in another or, alternatively, in mixing parameters from different force fields. Other forms for the intermolecular potential can also be employed to represent the intermolecular interactions, such as replacing the repulsive (12) potential by an exponential, as in MM2 (Allinger, 1977), or replacing the Lennard-Jones potential by a buffered 7-14 potential for the van der Waal interactions and shifting the electrostatics to prevent infinite attractive electrostatics from dominating the finite van der Waal interactions at short range, as employed in MMFF94 (Halgren, 1996). Although these more complicated functional forms arguably work much better for the varied small molecules of interest to pharmaceutical companies, these force fields have not seen significant use in biomolecular simulation applied to nucleic acids. For the electrostatic interactions, most MM methods assume fixed atomic point charges. Polarization is only included implicitly via the construction of point charge values that lead to effectively larger dipoles. The point charges are one part of the MM model that is not very transferable since a given atomic charge depends critically on its environment; therefore, charges are typically calculated for individual fragments (such as each individual nucleic acid or amino acid) rather than for specific atom types. With the lack of explicit polarization, a major limitation of the additive pairwise force fields is in the treatment of transition metals or multivalent ions (which may not be treated properly with standard additive empirical force fields); in this case, accurate treatment may require the inclusion of explicit polarization effects or even some QM treatment. Although polarization (induced dipole) effects have often been neglected to date, these effects can be included in the MM potential energy function. This can be done by including inducible dipoles on each center (µi) reacting to the electrostatic field (Ei0) on center i arising from all other fixed charges and representing the polarization energy (Upol) as follows: Upol = −
1 2
∑ µiEi
0
i
Equation 7.8.8
This is evaluated self-consistently, solving for the inducible dipole based on the polarizability of atom i (αi) and the total electrostatic field (Ei) at the polarizable center, where µi = αi × Ei, noting that the total electrostatic field: Ei = E0i + ∑ Tijµj i≠j
Equation 7.8.9
and Tij is the dipole tensor:
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1 Tij = 3 rij
3Hij
Hij r2ij
− 1
Equation 7.8.10
Additional terms can be added to represent three-body exchange repulsions (Caldwell et al., 1990). Alternative treatments of polarization include the fluctuating charge model that treats the charges as dynamic variables in the simulation (Rick et al., 1994). In the published literature, the inclusion of explicit polarization effects in molecular mechanics treatments has generally only included simulations of small polarizable molecules or ions in polarizable water. Polarization effects were added to better represent molecule association or free energies of solvation. A fully polarizable treatment has not been applied to large biomolecules in solution, in part because adding explicit polarization tremendously increases the cost of the calculations, but also because a consistent polarizable force field for nucleic acids or proteins has not been developed. This will likely change in the near future due to the availability of faster and better methods to include explicit polarization, which in turn will lead to the development of more reliable nucleic acid force fields that are parameterized for use with explicit polarization. The Total Energy Added together, the two energy terms (Uintra + Uinter) represent the total potential energy of the system. An important, but often overlooked, point is that it is often meaningless— despite the use of common units for energy such as kcal/mol—to compare absolute molecular mechanical energies between different molecules (or force fields) due to the lack of a common zero point energy. Moreover, many of the commonly employed force fields were not parameterized to very accurately estimate heats of formation, so even with a complete specification of a reaction linking two molecules together, it is likely that the relative energy between two different molecules will not be accurate. Despite this warning, it is possible to compare the relative energies of different conformations of the same molecule (under the same conditions, e.g., same force field, same number of explicit waters), remembering that this energy difference is not a free energy but only a relative energy or enthalpy. Although low energy structures are likely more representative, it is important to remember that this is not always the case (at normal temperatures) due to possibly large differences in entropy. For the simulation of nucleic acid systems, the most reasonable representation of the structure comes from the force fields specifically parameterized to represent nucleic acids. Current all-atom force fields for nucleic acids that perform reasonably well include the Cornell et al. (1995) and recent variants (Cheatham et al., 1999; Wang et al., 2000), BMS (Langley, 1998), and CHARMM all-27 nucleic acid (Foloppe and MacKerell, 2000) force fields. In addition to all-atom force fields, the JUMNA internal coordinate force field and others previously mentioned also perform reasonably well. BEYOND ENERGY EVALUATION
Molecular Modeling of Nucleic Acid Structure: Energy and Sampling
Evaluating the energy of a given model does not suggest anything about the relative stability or appropriateness of that model; however, differences in the relative energy between two conformations of the same model structure can suggest which structure is more enthalpically reasonable. Coupled with some representation of the relative entropy, this can give insight into the relative importance of each conformation. In general, however, it is not easy to estimate the entropy for a given conformation. Although there are some approximate methods for estimating relative conformational free energies (Kollman et al., 2000), these require knowing a priori what the representative structures are.
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Ideally, the investigator would like to know the conformation that represents the lowest energy (enthalpy) and ultimately the lowest free energy structures, since these structures are more representative of the true conformation of the molecule. A simple way in principle to find low-energy structures is to find the set of coordinates that minimizes the potential energy. In practice, this is limited due to the complexity of the potential energy hypersurface and the large number of degrees of freedom. Without exhaustive sampling of all possible conformations of a molecule, in general, it is impossible to determine if a given low-energy structure represents the true “global” minimum structure or whether it is simply a “local” minimum of the potential energy (where minimum energy structure means a structure that is at the bottom of one of many possible wells in the energy representation). Without knowing what the global minimum is, it is impossible to determine if a given model structure is at all representative of what might be expected at room temperature. Even with a knowledge of the global minimum energy structure, a molecule may have a number of other low-energy structures nearby that may be populated at room temperature. To find representative conformations, it may be desirable to apply methods that sample according to the expected probability of observing a given conformation (at a given temperature). Examples of methods that do this are molecular dynamics (MD) or Monte Carlo (MC) methods. This is sampling according to the Boltzmann distribution and, in the limit of infinite sampling, this gives a complete representation of all the possible conformations and their relative probabilities of occurrence, pi, where εi is the total energy of the ith state, T is the temperature, and kB is the Boltzmann constant. pi =
e−εi / kBT −εi / kBT
∫e
Equation 7.8.11
The term in the denominator is the partition function and specifies an integral over all phase space or the complete set of possible coordinates and momenta. Of course, in practice, sampling is limited and the sheer complexity of the accessible conformational space may prevent finding all the low-energy conformations and, therefore, full determination of the partition function. The complexity of the potential energy hypersurface, and the exponential explosion of the number of possible conformations as the number of degrees of freedom increases, limit exhaustive search of this space to systems that possess only a few degrees of freedom. In 1990, none of a variety of systematic and random conformational search methods in both internal (torsion) and all-atom coordinate frames was independently able to find all the relevant low-energy conformers (within 3 kcal/mol) of the cyclically constrained cycloheptadecane molecule, which formally has 147 degrees of freedom (all atom; Saunders et al., 1990). Although computer power has increased tremendously in this time, order-of-magnitude computational speed advances do not significantly improve the effective conformational sampling due to the exponential explosion. A simple way to estimate the effective complexity is to assume that the number of possible minima or low-energy conformations relates to the simplistic set of three low-energy rotations about a given single bond (i.e., the trans, gauche+, and gauche− states) or 3n − 1, where n is the number of rotatable bonds. Although this is typically less than the total number of degrees of freedom, it is still large! With fewer degrees of freedom, there are less minima and sampling is easier. Moreover, minimization algorithms are less likely to become stuck in less representative local minima; however, the complexity quickly grows; even with the internal coordinate treatments for nucleic acids described, there are still ~20 degrees of freedom per nucleotide (representing hundreds of possible conformations
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for a given nucleotide assuming three low-energy states for each of approximately five rotatable bonds). This places exhaustive sampling out of reach for any system with more than a few nucleotides. This difficulty in finding the global minimum (or set of coordinates that leads to the lowest energy) is often termed the multiple minima problem. In the context of the effective amount of sampling attainable during MD simulation, this problem is often termed the conformational sampling problem and relates to the improbability of overcoming large energy barriers. The multiple minima or conformational sampling problem is why it is desirable to choose reasonable initial structures (i.e., experimentally derived structures or valid model structures) when modeling. With reasonable structures, likely MD or MC (see below) simulation will sample reasonably well near the initial structure; however, large conformational changes, such as B-DNA to Z-DNA transitions or RNA folding will not likely be seen in a reasonable time. Of course, these limits in sampling imply that even with unreasonable structures, such as the imaginary and perhaps metastable B-RNA structure, MD or MC will likely sample reasonably well near the initial model structure. This is indeed the case, as shown in state-of-the-art all-atom solvated MD simulations where B-RNA is stable for >10 nsec and B-RNA to A-RNA transitions are not observed unless artificial means are applied to force the conformational transition (Cheatham and Kollman, 1997). In the next sections, minimization, Monte Carlo, and molecular dynamics methods will be discussed in more detail; more detailed treatments can be found elsewhere (Valleau and Whittington, 1977; Allen and Tildesley, 1987; McCammon and Harvey, 1987; Brooks et al., 1988; van Gunsteren and Berendsen, 1990; Leach, 1997.) Minimization Minimizing the potential energy corresponds to instantaneously “freezing” the system or dropping to the bottom of the nearest potential energy well (as shown schematically in Figure 7.8.2A). Minimization is a standard optimization problem that can be approached using tools of various complexity ranging from simple and inefficient zero order methods, such as grid search, which only require evaluation of the energy for a particular conformation, to more efficient but complicated nth-order methods, which use information about all the derivatives of the energy function up to the nth order. Thus, to perform minimization with an nth-order method requires analytic derivatives of the potential energy function up through the (n−1)th derivative (since the nth can be approximated by finite difference methods). In practice, a variety of first- and second-order methods are typically applied since these provide a reasonable balance between functionality (i.e., finding a minimum) and efficiency. Finding the set of coordinates that minimizes the potential energy does not guarantee that the conformation found represents the lowest energy structure due to the presence of multiple minima. As discussed, the molecule can get trapped into a local minimum, and this local minimum may not be representative of the “true” conformation.
Molecular Modeling of Nucleic Acid Structure: Energy and Sampling
In spite of the local minimum problem, minimization is still a useful tool for modeling, since after making a particular modification to a given model, say the replacement of the phosphodiester backbone of DNA by a poly-amide (PNA) backbone, the conformation can be minimized. This will remove gross steric overlap and relax strained bonds and angles. While this will not say much about the relative stability of this backbone modification, it can suggest whether the backbone replacement is at least sterically feasible for the given nucleic acid conformation. Coupled with a good chemical intuition, this may provide sufficient information to the modeler to suggest whether this backbone modification is potentially useful or not. For example, consider related backbone modifications to PNA that include one more or one less methyl group along the backbone. Simple minimization may suggest that the shorter backbone will strain the nucleic acid, leading to a lower rise between base pairs, and the longer backbone will increase the
7.8.10 Supplement 4
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A
B
C
Figure 7.8.2 Schematic representations of the sampling of various methods. These plots represent the energy of the system along an arbitrary reaction coordinate. The wells represent energy minima in the phase space. The state of the system is depicted by the location of the ball. (A) Minimization. The system moves to the bottom of the nearest well and barriers are not overcome. (B) Monte Carlo. Each configuration of the system is represented by a number and barrier crossing relates to the move set and total number of moves. (C) Molecular dynamics. The state of the system evolves due to force according to Newton’s equations of motion. In short simulations, large barriers will not be surmounted.
separation between base pairs, which might suggest that these backbone modifications are less reasonable. While some insight can be gained, it is important to remember that entropic effects are not included (such as the likely greater configurational entropy loss on binding for the larger PNA) and that the conformation sampled by the minimization is only a local minimum. In addition to being a useful tool for modeling, minimization is necessary prior to running molecular dynamics simulations to relieve any large steric overlap or remove strained bonds or angles that might otherwise lead to initially large forces. During the dynamics, the large forces lead to large displacements, which in turn may lead to more overlaps and more large displacements; this cascade of events can lead to local hot spots or failure of the integrator during MD. There are two common first-order minimization methods in common usage: the steepest descent (Wiberg, 1965) and conjugate gradient (Fletcher and Reeves, 1964) methods. These use the first derivative to give information about the slope of the potential (but not the curvature). In steepest descent, movement is made parallel to the net force. Given reasonable step sizes, this method will not cross barriers and will readily traverse down
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to the bottom of the nearest potential energy well. Although this method is not very efficient, it is very stable and is therefore often used to initially minimize the structure when there are large energies. Often this is used as the first step in modeling, particularly when there are potentially large van der Waals overlaps to initially relax the most drastic energetic penalties. The conjugate gradient method improves upon the steepest descent by using the gradients from previous steps to further guide the minimization. This method is more efficient than steepest descent and is appropriate to apply after initial minimization (e.g., to remove the largest steric overlaps). Near the bottom of the harmonic well, or after some initial minimization, use of second order methods (which assume an approximately quadratic relationship to the energy) such as Newton-Raphson (TNPACK; Schlick and Overton, 1987) can be used to speed up convergence at the expense of greater memory usage and the need for second derivatives (either by finite difference or analytical). These methods are typically more expensive since they require inverting the second derivative matrix. The expense can be significantly reduced by limiting the space sampled to regions where there is significant movement in the energy, thus limiting the size of the second derivative matrix (which is calculated by finite difference; Brooks et al., 1983). The key point is that minimization is a very useful tool, but that care should be taken when analyzing the validity of a particular “low-energy” structure. Because of the limits in sampling and the great likelihood of minimizing to a local minimum (which may or may not be representative of the favored low-energy structures), the validity needs to be judged based on the chemical intuition of the modeler and known reliability of the initial model. To better sample space, Monte Carlo or molecular dynamics methods can be applied; although these methods avoid the problem of getting trapped in local minima, limits in sampling only allow partial sampling of conformations “near” the initial structure. Despite the limited sampling, minimization is an extremely useful tool to characterize model structures. Note that care should be taken when applying restraints along with minimization, such as when using NMR-derived distance restraints, to balance the restraint force constants with those of the force field. If the restraint force constants are too large relative to the force field, unrealistic distortion of the structure may result. As a final practical comment about minimization, it is important to initially use small step sizes during the minimization of high-energy structures. This prevents the minimizer from jumping out of the current well to a potentially higher energy surface, which with repeated large jumps can lead to instability. Moreover, if the step size is too large, the minimizer may effectively get “trapped” in a cycle, jumping back and forth across the bottom of the well and preventing the minimizer from converging. Monte Carlo
Molecular Modeling of Nucleic Acid Structure: Energy and Sampling
The Metropolis Monte Carlo method is essentially an algorithm for generating a random walk in conformational space such that the conformations obtained are distributed according to the probabilities expected for the equilibrium Boltzmann distribution (Metropolis et al., 1953; Valleau and Whittington, 1977). The algorithm is very simple and requires a single evaluation of the potential energy (and no force calculation) for each step (Figure 7.8.2B). At each step, a random movement is made. In an all-atom representation, this may be the movement of a single particle; with an internal coordinate representation, it is a movement along the normal coordinates (such as a rotation of one of the rigid units). The potential energy difference (∆E) between the new conformation and the old is calculated. If the energy of the new conformation is lower, the move is accepted. If not, the move is accepted if e−∆E / k T is less than a random number drawn from the interval [0,1]; otherwise the old conformation is retained. The length of the simulation B
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relates to the number of attempted moves or configurations. The length of the simulation does not relate to the time scale and only represents the number of configurations sampled. Use of this procedure leads to a representative set of structures. With sufficient sampling, this will converge to the equilibrium distribution and in principle allow reasonable estimation of any ensemble average, assuming the forces between particles are velocity independent (Metropolis et al., 1953). This assumption allows one to separate out the momenta or kinetic energy from the total energy such that only the potential energy is evaluated in the above expression; in practice, the kinetic component is always ignored. Since uphill moves are only accepted randomly (and not according to any deterministic process), there is no implicit time evolution and the progression of the sampling gives no information about the dynamics. The success of this procedure (with finite sampling) largely relates to the “move set”, or set of possible moves. When an all-atom treatment is applied with single particle moves, the stiff internal degrees of freedom can lead to large energy changes and therefore a small step size needs to be used. With small step sizes, many more configurations may need to be evaluated to generate an appropriate ensemble, decreasing the efficiency. Moreover, for systems with disparate frequencies in different degrees of freedom—i.e., both stiff internal degrees of freedom (bonds, angles) and softer modes (correlated movements, dihedral rotation)—such as nucleic acids or single particle or atom moves, this leads to poor sampling. Even for liquid simulation, small atom-based moves lead to low acceptance ratios and poor sampling. In liquid simulation this can be overcome by using moves along normal coordinates, such as bond rotations, coupled with rotations and translations of the entire molecule. In general, it is desirable to avoid move sets that overly reject moves, and a roughly 40% acceptance rate represents a reasonable balance (Jorgensen and Tirado-Rives, 1996). The difficulty in choosing a proper move set led to the early impression that MD simulations were up to ten times more efficient than MC at generating conformations of the small protein BPTI (Northrup and McCammon, 1980); however, this simulation used an extremely inefficient move set based on movement of atomic centers. Much better behavior is seen with a move set based on internal coordinate rotations about bonds (with rigid bond length and angles; Noguti and Go, 1985). For liquid simulation, MC is likely the most efficient method for generating reasonable ensembles; an excellent MC program for liquid simulation is BOSS developed by the Jorgensen group (Jorgensen, 1995). Use of the MC procedure for the simulation of neat liquids led to the development of very reliable van der Waals parameters for the OPLS force field (Jorgensen et al., 1996). A direct comparison of MC and MD simulation of liquid hexane suggests that MC is roughly three times more efficient than MD when an appropriate move set for the MC is applied (Jorgensen and Tirado-Rives, 1996). Although MC is more efficient for liquid simulations, it is not clear this generalization follows to the simulation of biomolecules, particularly in solution. For example, with internal coordinate moves with flexible long-chain molecules (such as nucleic acids), a small rotation about a central dihedral angle can lead to a large displacement of the end of the chain. In explicit water, this might lead to extreme van der Waals overlap and likely very high rejection rates (for these types of moves). Although the use of correlated moves (such as crankshaft rotations about two bonds) can counter this effect (Dodd et al., 1993; Deem and Bader, 1996), there has been little published use of MC methods in all-atom simulation of nucleic acids in explicit solvent. For simulations without explicit solvent, MC simulation is widely used for nucleic acid simulation particularly with internal coordinate representations. Examples include its use in structure predictions as previously mentioned (Erie et al., 1993), investigating DNA bending in polyadenine tracts (Zhurkin et al., 1991), and investigating counterion distribution about DNA (Young et al., 1997).
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Molecular Dynamics Molecular dynamics simulation refers to integrating numerically the classical equations of motion (Newton’s equations) for all the atoms in the system (Figure 7.8.2C). A simulation is started by assigning random momenta (velocities, vi) for each of the N particles (of mass mi) from a Boltzmann distribution about a given temperature, T. N
∑ miv2i =
1 2
i
3NkBT 2
Equation 7.8.12
Then the dynamics are propagated by integrating Newton’s equations of motion, which for the pairwise potential, Ui, and its first derivative: N
∇i ∑ Uij j
Equation 7.8.13
is represented by the following: mi
d2ri dt2
N
= −∇i ∑ Uij j
Equation 7.8.14
The integration is typically performed through the use of one of a variety of first-order integration algorithms, such as leap-frog or velocity Verlet (Allen and Tildesley, 1987). This requires the calculation of the forces at each step, so a typical MD step involves calculation of the energy, forces, and velocities, and integration to obtain the coordinates for the next step. An important assumption is the ergodic hypothesis that, in the limit of complete sampling, the time average (that obtained by molecular dynamics) is equivalent to the ensemble average. Given sufficient sampling, it appears that the ergodic hypothesis is valid in practice since both MD and MC lead to equivalent ensembles for liquid hexane even though they use rather different sampling mechanisms (Jorgensen and Tirado-Rives, 1996). Also note that, unlike MC, the MD sampling is based on a dynamic propagation of the molecular mechanical forces; therefore, it is important that the forces are analytical or exact derivatives of the energy (i.e., the forces and energies should match). This is not always true in practice since the force calculation in some cases may be too expensive (particularly for nonpairwise forces), requiring some approximations. This means that the sampling is done on a surface different than the molecular mechanical energy surface.
Molecular Modeling of Nucleic Acid Structure: Energy and Sampling
Issues in MD simulation include the need for stable, reversible, and ideally symplectic (which implies loosely that the algorithm conserves energy and momentum) integrators. The simple first-order Verlet, velocity Verlet, and leap-frog integration algorithms satisfy these conditions in proper usage, in contrast to the more complex higher-order integrators such as the Gear predictor. Stability of the integration directly relates to the integration time step that in turn relates to the expected frequency of motion. A simple rule of thumb for Verlet integrators is that the integration catastrophe, or the time step where the integrator blows up, is the period of the highest frequency motion divided by π. For an all-atom simulation, the highest-frequency motions involve bond stretching of bonds to hydrogen. In the absence of rigid bond lengths (or constrained bonds), time steps are limited to the <1 fsec range. With SHAKE (Ryckaert et al., 1977) applied to constrain the lengths of bonds involving hydrogen, time steps in the 2 fsec range are routinely applied. Larger time steps are possible by limiting the high-frequency motion, such as through the
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use of rigid units, which then necessitates inverting the inertia tensor as with internal coordinate treatments or the need for imposition of iteratively solved holonomic constraints. Effectively larger time steps are also possible through the application of multiple time step methods, which treat the slowly varying forces (which ideally represent the more computationally demanding part of the energy and force evaluation) with a longer time step, such as the long-range pairwise forces (Tuckerman et al., 1992; Biesiadecki and Skeel, 1993). With increased masses on hydrogen atoms to limit high-frequency motion, time steps as large as 5 fsec have been applied to systems with explicit water. The symplectic nature of the integrator relates to preserving the Hamiltonian (or loosely the energy representation) of the system during integration. Not only should this be a property of the integrator, but molecular dynamics simulation in general should conserve energy. This is an excellent test of the methods. Note that in common usage, MD programs do not always necessarily conserve energy. This is often due to SHAKE tolerances that are not stringent enough, integration time steps that are too large, the use of the weak-coupling algorithm for constant pressure, and neglect of pair interactions. In order to speed the calculation, the effective number of pair interactions is often reduced to only those within a given range and a list of in-range pair interactions is maintained for each atom. For speed this “pair list” is not updated every step. Unless a buffer is maintained to not omit (include) pair interactions moving into (out of) range which is conservatively or heuristically updated, energy conservation may not be maintained. How the pair interactions are limited to finite range can also have important consequences, as can temperature or pressure coupling for constant T,P ensembles. In order to deterministically integrate Newton’s equations of motion, it is important that systematic force errors be avoided (such as can occur with lack of energy conservation and temperature scaling) since these can lead to artifactual behavior, such as violation of equipartition (Harvey et al., 1998; Chiu et al., 2000). For example, with energy drains and the application of temperature adjustment by uniform scaling of the velocities of all the atoms, energy accumulates in low-frequency modes. This can lead to a growth of the center of mass translation. Given this, and since random velocity assignment likely leads to nonzero center of mass translational motion, it is advisable to remove this motion at the beginning of an MD run. Random force errors, such as those resulting from the slow accumulation of errors due to finite numerical precision, do not seem to lead to artifacts since they are equally likely to add as subtract from the total energy. To this end, differences obtained between sequential and parallel MD runs due to differences in the order of operations do not lead to significant differences and simply manifest the inherently chaotic nature of the integration (Braxenthaler et al., 1997). Variants of standard MD include Langevin and Brownian dynamics. These MD methods are used to implicitly include the diffusive effects of solvent. In practice, this involves adding to the standard forces stochastic forces from a heat bath (via random fluctuation of forces) and dissipative forces to balance them. For in vacuo simulations, this represents thermal collisions with other molecules and allows coupling of energy among the different internal degrees of freedom. This can be very useful since, in the absence of Langevin forces, during deterministic dynamics, memory of the initial conditions may persist in the form of various correlated motions, and some low-frequency correlated motions may not be able to couple back into other modes of motion (since there are no collisions with other molecules). This can lead to poor sampling. In the limit of very high friction, Langevin dynamics become Brownian dynamics. Brownian dynamics are purely diffusive and effectively add a random coordinate displacement. This method is typically used to model the diffusional motion of molecules (such as the encounter of a substrate into an enzyme binding site). More detailed discussion of MD and its variants can be found in the previously cited books and a very useful published discussion of stochastic dynamics methods (Pastor, 1994).
Biophysical Analysis of Nucleic Acids
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With molecular dynamics, each particle has a finite kinetic energy; a direct implication is that the rate of barrier crossing will be proportional to the temperature and the length of the simulation. This implies that the probability of crossing large energy barriers during MD is rather small (as is represented schematically in Figure 7.8.2C). A back-of-the-envelope estimation of the rate of barrier crossing can be obtained from transition state theory with some basic approximations. ‡
k ≅ κν × e−Gact / RT Equation 7.8.15
The rate k is related to the transmission coefficient κ, which represents the ratio of successful transitions over the barrier, an effective rate at the top the barrier or equivalently a factor loosely representing collisions with the barrier, ν, times a Boltzmann-weighted “free energy of activation,” which represents the height of the barrier. For simplicity, it can be assumed that at the top of the barrier, the particle always crosses the barrier rather than reflecting back, leading to κ = 1 (in contrast to expected values of 0.4 to 1.0 in solution). By classical equipartition (E ≅ kBT) and from E = hν this barrier rate is: ν=
kBT h
Equation 7.8.16
where T is the temperature, h is Plank’s constant, and kB is the Boltzmann constant. At room temperature, RT is ∼0.6 kcal/mol and ν is ∼6.2 psec−1. Remembering that this is only an approximation, this suggests that barriers of ∼1 kcal/mol can be surmounted in picoseconds (∼1.2 psec−1) and ∼5 kcal/mol in nanoseconds (∼1.5 nsec−1), but that barriers >10 kcal/mol may take microseconds or longer. This relates to the conformational sampling problem and is a significant limitation of MD. In MC simulations, this problem can be overcome by increasing the size of the moves (at the expense of lower acceptance ratios) or by designing clever move sets. In a similar manner, with MD simulation various methods can be applied to effectively lower barriers to conformational transition, such as adding biasing potentials to lower specific torsion barriers, increasing masses, increasing temperature, smoothing the overall potential surface, or applying mean field approximations, such as with the locally enhanced sampling (LES) methodology. The LES methodology shows much promise in biomolecular simulation, such as by reducing the barriers to conformational transition and thereby allowing transition from an “incorrect” to correct RNA hairpin loop conformation (Simmerling et al., 1998). For review of enhanced sampling methods, see Straub (1996).
Molecular Modeling of Nucleic Acid Structure: Energy and Sampling
The attainable MD simulation time scale relates to the complexity of the potential (and therefore the time required to evaluate the energy and forces) and the integration time step. With longer time steps, longer simulations are possible. Through the deterministic procedure and specification of an integration time step, there is a direct relationship between the number of MD steps and the effective time scale (time step × the number of steps = total time). The current state of the art for MD simulation of nucleic acids in explicit solvent involves simulations of <50 nsec. Although in principle this time scale is “accurately” represented by the dynamics or integration of the classical equations of motion, whether a 1 nsec simulation of DNA actually accurately represents 1 nsec of real motion depends on the empirical potential employed. For equilibrium or ensemble properties (such as the energy, free energy, heat capacity, or density) this exact time scale is not important, assuming independence of the forces on the velocity of a given particle. This is implicitly the case with the standard molecular mechanics potentials. For ensemble properties, what matters is the effective amount of sampling. In principle, one can obtain equivalent ensemble averages even if the masses on all the hydrogens are increased. This
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will allow a larger time step and therefore a longer sampling. Although this does not affect the ensemble properties, this will drastically affect time-averaged properties, such as water diffusion or the rate of specific conformational transitions. The point is that these dynamic properties are very sensitive to the potential (and atomic masses). Since the force field is primarily parameterized to represent structure, it is not clear how accurate the dynamic properties are. Clearly in the absence of viscous damping forces or explicit solvent, the rate of dynamics may be enhanced relative to simulations in explicit solvent. Similarly, under high pressure conditions, such as in an isolated solvent droplet, the dynamics may be reduced. In explicit solvent, MD simulations of proteins and nucleic acids display thermal parameters that are in good accord with experiment and in general properly represent fast (picosecond) time scale motions. Whether these methods can accurately represent longer time scale motions is unclear since realistic MD simulation is currently limited to the nanosecond time scale. Evidence from the simulations is that some properties are well represented; however, there is a clear overestimation of harmonic motion in solvated proteins at low temperature (i.e., suggesting motion that is too slow; Steinbach and Brooks, 1994). On the other hand, one of the most commonly used water models, TIP3P (Jorgensen et al., 1983) diffuses at twice the experimental value. The point of this discussion of time scale and dynamics is not to criticize the methods but to remind potential modelers of the various issues and sensitivity not only to the force field but the representation. Although in general the results are within range, it is important to understand how the methods have been validated by comparison to experiment for a given property before making firm claims about exact details of the time scale for conformational transition. For example, estimating the free energy of binding for a particular water to DNA based on the lifetime of its bound state is likely to be inaccurate and very dependent on the specific water model. Additionally, in order to make claims about the time scale for a given process based on MD, the time scale of the simulation should be significantly longer than the relaxation time of the process of interest. Likely the simulation should be at least an order, if not two orders, of magnitude longer than the relaxation time and, when attempting to make statistically valid claims about the rate of a given transition or specific correlation, many events need to be observed. These and related issues regarding the validation of simulation results are presented in a recent review (van Gunsteren and Mark, 1998). SUMMARY In this unit, the basic principles of the common energy representations for nucleic acid models have been presented, and methods for exploring these energy surfaces have been discussed in some detail. To move beyond simple energy evaluation of a single model structure, it is fairly clear that a large number of energy (and possibly force) evaluations will be necessary. This becomes computationally demanding, and it is desirable to limit the computational cost as much as possible without sacrificing accuracy. For systems of reasonable size, there is a need to use empirical potentials as discussed; however, even with the simple pairwise potentials, the number of pair interactions quickly grows as the number of atoms is increased. Thus, for larger systems the authors limit the effective number of intermolecular interactions by limiting the range of the interaction (e.g., by limiting interactions to distances less than some cutoff or utilizing hierarchical approaches that coalesce groups of distant atoms into an approximate single effective particle). Additional complications relate to the representation of nucleic acids since water and associated counterion-ions (salt) are an integral part of nucleic acid structure (see UNIT 7.9).
Biophysical Analysis of Nucleic Acids
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LITERATURE CITED Allen, M.P. and Tildesley, D.J. 1987. Computer Simulation of Liquids. Oxford University Press, Oxford. Allinger, N.L. 1977. Conformational analysis 130. MM2. A hydrocarbon force field utilizing V1 and V2 torsional terms. J. Am. Chem. Soc. 99:8127-8134. Allinger, N.L., Yuh, Y.H., and Lii, J.H. 1989. Molecular mechanics. The MM3 force field for hydrocarbons.1. J. Am. Chem. Soc. 111:8551-8566. Allinger, N.L., Chen, K., and Lii, J.H. 1996. An improved force field (MM4) for saturated hydrocarbons. J. Comp. Chem. 17:642-668. Biesiadecki, J.J. and Skeel, R.D. 1993. Dangers of multiple time step methods. J. Comp. Phys. 109:318-328. Braxenthaler, M., Unger, R., Auerbach, J., Given, A., and Moult, J. 1997. Chaos in protein dynamics. Proteins 29:417-425. Brooks, B.R., Bruccoleri, R.E., Olafson, B.D., States, D., Swaminathan, J.S., and Karplus, M. 1983. CHARMM: A program for macromolecular energy, minimization, and dynamics calculations. J. Comp. Chem. 4:187-217. Brooks, C.L., III, Karplus, M., and Pettitt, B.M. 1988. Proteins. A Theoretical Perspective of Dynamics, Structure, and Thermodynamics. John Wiley & Sons, New York. Caldwell, J.W., Dang, L.X., and Kollman, P.A. 1990. Implementation of nonadditive intermolecular potentials by use of molecular dynamics: Development of a water-water potential and water-ion cluster interactions. J. Am. Chem. Soc. 112:91449147. Chatfield, D.C., Eurenius, K.P., and Brooks, B.R. 1998. HIV-1 protease cleavage mechanism: A theoretical investigation based on classical MD simulation and reaction path calculations using a hybrid QM/MM potential. Theochem. J. Mol. Struct. 423:79-92. Cheatham, T.E. III. and Kollman, P.A. 1997. Molecular dynamics simulations highlight the structural differences in DNA:DNA, RNA:RNA and DNA:RNA hybrid duplexes. J. Am. Chem. Soc. 119:4805-4825. Cheatham, T.E. III, Cieplak, P., and Kollman, P.A. 1999. A modified version of the Cornell et al. force field with improved sugar pucker phases and helical repeat. J. Biomol. Struct. Dyn. 16:845-862. Chiu, S.W., Clark, M., Subramaniam, S., and Jakobsson, E. 2000. Collective motion artifacts arising in long-duration molecular dynamics simulations. J. Comp. Chem. 21:121-131.
Molecular Modeling of Nucleic Acid Structure: Energy and Sampling
Cornell, W.D., Cieplak, P., Bayly, C.I., Gould, I.R., Merz, K.M., Ferguson, D.M., Spellmeyer, D.C., Fox, T., Caldwell, J.W., and Kollman, P.A. 1995. A second generation force field for the simulation of proteins, nucleic acids, and organic molecules. J. Am. Chem. Soc. 117:5179-5197.
Cummins, P.L. and Gready, J.E. 1997. Coupled semiempirical molecular orbital and molecular mechanics model (QM/MM) for organic molecules in aqueous solution. J. Comp. Chem. 18:1496-1512. Deem, M.W. and Bader, J.S.. 1996. A configurational bias Monte-Carlo method for linear and cyclic peptides. Mol. Phys. 87:1245-1260. Dixon, S.L. and Merz, K.M.J. 1997. Fast, accurate semiempirical molecular orbital calculations for macromolecules. J. Chem. Phys. 107:879-893. Dodd, L.R., Boone, T.D., and Theodorou, D.N. 1993. A concerted rotation algorithm for atomistic Monte-Carlo simulation of polymer melts and glasses. Mol. Phys. 78:961-996. Erie, D.A., Breslauer, K.J., and Olson, W.K. 1993. A Monte Carlo method for generating structures of short single-stranded DNA sequences. Biopolymers 33:75-105. Field, M., Bash, P., and Karplus, M. 1990. A combined quantum mechanical and molecular mechanical potential for molecular dynamics simulation. J. Comp. Chem. 11:700-733. Fletcher, R. and Reeves, C.M. 1964. Function minimization by conjugate gradients. Computer J. 7:149-153. Foloppe, N. and MacKerell, A.D.J. 2000. All-atom empirical force field for nucleic acids. 1) Parameter optimization based on small molecule and condensed phase macromolecular target data. J. Comp. Chem. 21:86-104. Gao, J.L. 1996. Hybrid quantum and molecular mechanical simulations–An alternative avenue to solvent effects in organic chemistry. Acc. Chem. Res. 29:298-305. Gorin, A.A., Ulyanov, N.B., and Zhurkin, V.B. 1990. S-N transition of the sugar ring in B-form DNA. Molekulyarnaya Biologiya 24:1300-1313. Halgren, T.A. 1996. Merck molecular force field 1. Basis, form, scope, parameterization, and performance of MMFF94. J. Comp. Chem. 17:490519. Harvey, S.C., Tan, R.K.Z, and Cheatham, T.E. III. 1998. The flying ice cube: Velocity rescaling in molecular dynamics simulations leads to violation of equipartition. J. Comp. Chem. 19:726740. Hobza, P. and Sponer, J. 1999. Structure, energetics, and dynamics of the nucleic acid base pairs: Nonempirical ab initio calculations. Chem. Rev. 11:3247-3276. Jorgensen, W. 1995. BOSS, Version 3.6. Yale University, New Haven. Jorgensen, W.L. and Tirado-Rives, J. 1996. Monte Carlo vs Molecular dynamics for conformational sampling. J. Phys. Chem. 100:14508-14513. Jorgensen, W.L., Chandrasekhar, J., Madura, J.D., Impey, R.W., and Klein, M.L. 1983. Comparison of simple potential functions for simulating liquid water. J. Chem. Phys. 79:926-935.
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Jorgensen, W.L., Maxwell, D.S., and Tirado-Rives, J. 1996. Development and testing of the OPLS all-atom force field on conformational energetics and properties of organic liquids. J. Am. Chem. Soc. 118:11225-11236. Kollman, P.A., Massova, I., Reyes, C., Kuhn, B., Huo, S., Chong, L., Lee, M., Lee, T., Duan, Y., Wang, W., Donni, D., Cieplak, P., Srinivasan, J., Case, D.A., and Cheatham, T.E. III 2000. Calculating structures and free energies of complex molecules: Combining molecular mechanics and continuum models. Acc. Chem. Res. 33:889-897. Lahiri, A. and Nilsson, L. 1997. Properties of dianionic oxyphosphorane intermediates from hybrid QM/MM simulations: Implications for ribozyme reactions. J. Mol. Struct. 419:51-55.
Pearlman, D.A., Case, D.A., Caldwell, J.W., Ross, W.S., Cheatham, T.E., Debolt, S., Ferguson, D., Seibel, G., and Kollman, P. 1995. AMBER, a package of computer programs for applying molecular mechanics, normal mode analysis, molecular dynamics and free energy calculations to simulate the structure and energetic properties of molecules. Comp. Phys. Comm. 91:1-41. Rick, S.W., Stuart, S.J., and Berne, B.J. 1994. Dynamical fluctuating charge force fields–application to liquid water. J. Chem. Phys. 101:61416156. Ryckaert, J.P., Ciccotti, G., and Berendsen, H.J.C. 1977. Numerical integration of the cartesian equations of motion of a system with constraints: Molecular dynamics of n-alkanes. J. Comp. Phys. 23:327-341.
Langley, D.R. 1998. Molecular dynamics simulations of environment and sequence dependent DNA conformation: The development of the BMS nucleic acid force field and comparison with experimental results. J. Biomol. Struct. Dyn. 16:487-509.
Saunders, M., Houk, K.N., Wu, Y.-D., Still, W.C., Lipton, M., Chong, G., and Guida, W.C. 1990. Conformations of cycloheptadecane. A comparison of methods for conformational searching. J. Am. Chem. Soc. 112:1419-1427.
Lavery, R., Zakrzewska, K., and Sklenar, H. 1995. JUMNA (junction minimisation of nucleic acids). Comp. Phys. Comm. 91:135-158.
Schlick, T. and Overton, M. 1987. A powerful truncated Newton method for potential energy minimization. J. Comp. Chem. 8:1025-1039.
Leach, A.R. 1997. Molecular Modeling: Principles and Applications. Addison-Wesley, Reading, Mass.
Simmerling, C., Miller, J.L., and Kollman, P.A. 1998. Combined locally enhanced sampling and particle mesh Ewald as a strategy to locate the experimental structure of a non-helical nucleic acid. J. Am. Chem. Soc. 120:7149-7155.
Levine, I.N. 1991. Quantum Chemistry. Prentice Hall, Englewood Cliffs, N.J. Maple, J.R., Hwang, M.J., Stockfisch, T.P., Dinur, U., Waldman, M., Ewig, C.S., and Hagler, A.T. 1994. Derivation of class II force fields. 1. Methodology and quantum force field for the alkyl functional group and alkane molecules. J. Comp. Chem. 15:162-182. McCammon, J.A. and Harvey, S.C. 1987. Dynamics of Proteins and Nucleic Acids. Cambridge University Press, Cambridge. Metropolis, N., Rosenbluth, A.W., Rosenbluth, M.N., Teller, A.H., and Teller, E. 1953. Equation of state calculations by fast computing machines. J. Chem. Phys. 21:1087-1092. Nesterova, E.N., Federov, O.U., Poltev, V.I., and Chuprina, V.P. 1997. The study of possible A and B conformations of alternating DNA using a new program for conformational analysis of duplexes (CONAN). J. Biomol. Struct. Dyn. 14:459-474. Noguti, T. and Go, N. 1985. Efficient Monte Carlo method for simulation of fluctuating conformations of native proteins. Biopolymers 24:527546. Northrup, S.H. and McCammon, J.A. 1980. Simulation methods for protein structure fluctuations. Biopolymers 19:1001-1016. Pastor, R.W. 1994. Techniques and applications of Langevin dynamics simulations. In The Molecular Dynamics of Liquid Crystals (G. Luckhurst and C. Veracini, eds.) pp. 85-138. Kluwer Academic Publishers, Amsterdam, The Netherlands.
Stanton, R.V., Little, L.R., and Merz, K.M. 1995. An examination of a Hartree-Fock molecular mechanical coupled potential. J. Phys. Chem. 99:17344-17348. Steinbach, P.J. and Brooks, B.R. 1994. Protein simulation below the glass-transition temperature. Dependence on cooling protocol. Chem. Phys. Lett. 226:447-452. Stewart, J.J.P. 1990. Semiempirical molecular orbital methods. In Reviews in Computational Chemistry (K.B. Lipkowitz and D.B. Boyd, eds.) pp. 45-81. VCH, New York. Straub, J.E. 1996. Optimization techniques with applications to proteins. In New Developments in Theoretical Studies of Proteins (R. Elber, ed.) pp. 137-196. World Scientific, Singapore. Szabo, A. and Ostlund, N.S. 1989. Modern Quantum Chemistry. McGraw-Hill, New York. Thiel, W. 1997. Computational methods for large molecules. J. Mol. Struct. 398:1-6. Tuckerman, M., Berne, B.J., and Martyna, G.J. 1992. Reversible multiple time scale molecular dynamics. J. Chem. Phys. 97:1990-2001. Valleau, J.P. and Whittington, S.G. 1977. A Guide to Monte Carlo for Statistical Mechanics: 1. Highways. In Statistical Mechanics A. A Modern Theoretical Chemistry (B.J. Berne, ed.) pp. 137168. Plenum Press, New York. van Gunsteren, W.F. and Berendsen, H.J.C. 1987. Groningen molecular simulation (GROMOS) library manual. BIOMOS, Nijenborgh, Groningen, The Netherlands.
Biophysical Analysis of Nucleic Acids
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van Gunsteren, W.F. and Berendsen, H.J.C. 1990. Computer simulation of molecular dynamics: Methodology, applications, and perspectives in chemistry. Angew. Chem., Int. Ed. Engl. 29:9921023. van Gunsteren, W.F. and Mark, A.E. 1998. Validation of molecular dynamics simulation. J. Chem. Phys. 108:6109-6116. Wang, J., Cieplak, P., and Kollman, P.A. 2000. How well does a restrained electrostatic potential (RESP) model perform in calculating conformational energies of organic and biological molecules? J. Comp. Chem. 21:1049-1074. Wiberg, K.B. 1965. A scheme for strain energy minimization. Application to the cycloalkanes. J. Am. Chem. Soc. 87:1070-1078. Young, M.A., Jayaram, B., and Beveridge, D.L. 1997. Intrusion of counterions into the spine of hydration in the minor groove of B-DNA: Fractional occupancy of electronegative pockets. J. Am. Chem. Soc. 119:59-69. Zerner, M.C. 1991. Semiempirical molecular orbital methods. In Reviews in Computational Chemistry (K.B. Lipkowitz and D.B. Boyd, eds.) pp. 313-365. VCH, New York. Zhurkin, V.B., Ulyanov, N.B., Gorin, A.A., and Jernigan, R.L. 1991. Static and statistical bending of DNA evaluated by Monte Carlo calculations. Proc. Natl. Acad. Sci. U.S.A. 88:70467050.
Internet Resources http://www.ccl.net or http://www.netsci.org/Resources/ Software/Modeling Lists of available software. http://www.msg.ameslab.gov/GAMESS/GAMESS. html GAMESS website.
http://www.dl.ac.uk/CCP/CCP1/gamess.html GAMESS-UK website. http://www.gaussian.com Gaussian program website. http://www.schrodinger.com Jaguar website. http://www.wavefunction.com, MNDO4 is available in many different codes from this website and others. http://www.ccl.net MOPAC 6.0 and 7.0 are in the public domain and are available from this (the Computational Chemistry mailing list site) and other sites. It is also available from the Quantum Chemistry Program Exchange, Department of Chemistry at the University of Indiana. http://www.q-chem.com Q-chem website. http://www.msi.com ZINDO website.
Contributed by Thomas E. Cheatham, III University of Utah Salt Lake City, Utah Bernard R. Brooks National Heart Lung & Blood Institute, NIH Bethesda, Maryland Peter A. Kollman University of California San Francisco, California
Molecular Modeling of Nucleic Acid Structure: Energy and Sampling
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Current Protocols in Nucleic Acid Chemistry
Molecular Modeling of Nucleic Acid Structure: Electrostatics and Solvation
UNIT 7.9
UNITS 7.5 & 7.8 introduced the modeling of nucleic acid structure at the molecular level. This
included a discussion of how to generate an initial model, how to evaluate the utility or reliability of a given model, and ultimately how to manipulate this model to better understand the structure, dynamics, and interactions. Subject to an appropriate representation of the energy, such as a specifically parameterized empirical force field, the techniques of minimization and Monte Carlo simulation, as well as molecular dynamics (MD) methods, were introduced as means to sample conformational space for a better understanding of the relevance of a given model. From this discussion, the major limitations with modeling, in general, were highlighted. These are the difficult issues in sampling conformational space effectively—the multiple minima or conformational sampling problems—and accurately representing the underlying energy of interaction. In order to provide a realistic model of the underlying energetics for nucleic acids in their native environments, it is crucial to include some representation of solvation (by water) and also to properly treat the electrostatic interactions. These are discussed in detail in this unit. ELECTROSTATICS AND SOLVATION Accurately modeling the structure and dynamics of nucleic acids with standard ab initio or empirical potentials presents special difficulties due to the highly charged phosphate backbone and the observation that nucleic acids are essentially always hydrated. Even under extremely dehydrating conditions, DNA still has very tightly associated water. To apply an accurate model, some representation of this structural and solvating water is likely necessary. Water has a structural role as both a donor and acceptor of hydrogen bonds, and can not only be specifically associated with the nucleic acid backbone but can specifically and nonspecifically associate into both the major and minor grooves. The importance of the structural water was readily observed in the first crystal structures that showed a clear spine of hydration in the minor groove (Drew and Dickerson, 1981). In addition to the structural role, water has a number of special properties. In addition to hydrodynamic properties, random thermal excitations, and viscous damping forces (which are likely to be important considerations when representing transport properties or investigating the dynamics), there are strong polarization effects. The relatively high permittivity, or dielectric constant, of water (~80) strongly screens the electrostatic interactions. On a microscopic level, this screening results from reorientation of permanent dipoles and electronic polarizability (or the creation of induced dipoles in the presence of an electric field). In addition to the specific structural and polarizing effects, another important interaction of the solvent derives from nonspecific entropic effects. Nonpolar molecules, those without hydrogen bonding capability or charges to interact with the water, attempt to minimize their exposure to water and tend to associate; this hydrophobic effect is a large driving force in protein folding (Dill, 1990; Spolar and Record, 1994) and appears to be a larger driving force in the association of ligands into the minor groove of DNA than electrostatic effects (Misra et al., 1994). In addition to solvent, the high charge density from the polyionic backbone and profound salt dependence on the structure of nucleic acids are likely to necessitate some representation of ionic screening beyond simple screening by solvent. This can be included through addition of explicit counterions. However, even with the inclusion of explicit solvent and counterions in nucleic acid simulation and the use of a reasonable nucleic acid force field, if the long-range forces are not properly represented, nucleic acid Contributed by Thomas E. Cheatham, III, Bernard R. Brooks, and Peter A. Kollman Current Protocols in Nucleic Acid Chemistry (2001) 7.9.1-7.9.21 Copyright © 2001 by John Wiley & Sons, Inc.
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7.9.1 Supplement 5
structure will be unstable during molecular dynamics simulation (Cheatham et al., 1995). This is due to the presence of high charge and large electrostatic interactions that decay only slowly with distance due to the long-range character of the Coulombic potential. Standard approximations to limit the effective range (for more tractable computation) often lead to instability (as will be discussed in greater detail later). All of these effects—i.e., improper representation of the solvent and ionic effects and improper treatment of the long-range electrostatic interactions—unduly influence the structure and dynamics of nucleic acids during simulation. However, as discussed in previous units, the level of simulation one applies represents a tradeoff in speed and accuracy, and in some cases it may be unfeasible to fully represent the effect of solvent, ions, and long-range electrostatics. Until fairly recently, computer power was not sufficient to allow fully detailed treatments with explicit water and complete representation of the long-range electrostatics. Therefore, the first MD of DNA was performed in vacuo (without any representation of screening by solvent or ions; Levitt, 1983). In order to prevent the duplex from distorting, it was necessary to remove the charge on the phosphates. Unfortunately, this approximation removes one of the essential features of polymeric nucleic acids. There are, however, some simple approximations that can be applied and that reasonably represent solvent screening; these are discussed in the next section. This will be followed by discussion of more accurate implicit solvent representations, followed finally by the more computationally demanding (but even more accurate) treatments that include explicit solvent. IN VACUO REPRESENTATIONS
Molecular Modeling: Electrostatics and Solvation
Simulations without any explicit solvent, or in vacuo simulations, are typically very rapid (with an empirical potential) since the omission of explicit solvent reduces the requisite number of atoms. Therefore, all pairwise interactions can typically be calculated, and cutoffs that limit the number of pairs included in the intermolecular interactions are not necessary. However, unless the nucleic acid environment is actually in the gas phase (where the dielectric constant, ε, is equal to 1), simulation tends to overemphasize the charge interactions compared to what is expected in solution. The significant phosphate repulsion will therefore tend to destabilize the structure. Although this can in part be remedied by reducing the charges on the ionic groups (as in the early MD), this is unrealistic. Alternatively, it is possible to increase the effective dielectric (ε) of the system up to the permittivity of water (ε ≈ 80) this, however, leads to too much screening at short distances, leading to destabilization of short-range charge interactions, such as hydrogen bonds. Ideally we want a method that allows the full charges at short range (ε = 1) and bulk solvent screening at longer distances (i.e., ε ≈ 80 at 20 Å).The simplest way to do this is through the application of “effective” dielectric constants or modified dielectric functions (Davis and McCammon, 1990). The most common and simplest form is the distance-dependent dielectric constant, where (ε = krij with k in the range from 1 to 4. A better form which does not screen as drastically at short range uses a more complex sigmoidal dielectric function (Hingerty et al., 1985; Ramstein and Lavery, 1988; Daggett et al., 1991) that tapers the short-range screening more slowly. Alternatively, a dielectric function that increases exponentially with distance (consistent with Debye-Hückel theory) has been applied in the simulation of nucleic acids (von Kitzing and Diekmann, 1987; Sarai et al., 1988). A drawback of the distance-based effective dielectric functions is the uniformity of the screening regardless of the proximity to solvent or environment. This is a poor approximation for a macromolecule that tends to have a lower effective dielectric in the interior of the molecule compared to bulk. Moreover, these functions tend to cause the molecule to compact during the dynamics and suppress motion (Harvey, 1989; Steinbach and Brooks, 1994).
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Despite these issues, these treatments have been routinely applied in modeling nucleic acids. In MD simulation, these approaches lead to good representation of DNA duplexes, and when computers were less powerful, these methods were widely used (Beveridge et al., 1993). Less reasonable behavior is observed with higher-order nucleic acid structures, such as tRNA. When coupled with specifically parameterized force fields and internal coordinate treatments, sigmoidal dielectric functions provide a fast means to investigate nucleic acid structure. With simplified treatments such as this, counterion damping of the charge interactions can be accounted for effectively by reducing the charge on the phosphates or including explicit counterions. If explicit counterions are used, the standard ion parameters appropriate for solvent are not applicable; instead, larger ion radii are needed to effectively represent the first solvation shell of ion hydration (Singh et al., 1985). Recent applications of simplified internal coordinate treatments with effective dielectric constants include investigation of extreme stretching of DNA (Lebrun and Lavery, 1996; Lebrun et al., 1997) and DNA A-tract bending (Zhurkin et al., 1991). These simplified methods are powerful enough to characterize and evaluate various nucleic acid force fields in common usage (Flatters et al., 1997). In spite of these successes, it is important to note that such a simple form for the dielectric screening is unlikely to accurately represent the dielectric response of the surrounding medium, which is dependent on the position of all charges rather than a uniform scaling based on simple pairwise distances. Investigations of DNA suggest that no one form of an effective dielectric can reasonably represent all types of pair interactions (Friedman and Honig, 1992). In minimal nucleic acid models, despite the limitations, these treatments are very useful for rapid characterization of the structure. Additionally, although the DNA is not perfectly represented, this level of representation is often sufficient for use in the refinement of structure based on restraints from NMR data. IMPLICIT SOLVENT MODELS Although the effective dielectric functions can partially represent solvent screening, there is no representation of any reorientational polarization. In addition, the screening is unrealistically uniform and there is no representation of hydrophobic or hydrodynamic effects. To correct some of these deficiencies, various implicit solvent models may be applied. These have the benefit that the calculation of the influence of solvent is very rapid and moreover is typically represented in terms of a solvation free energy. This is possible due to the uniformity of the dielectric continuum which represents the solvent ensemble properties directly, rather than as a sampling of many distinct solvent configurations. Treating all the water as “bulk” through the use of a dielectric continuum may not be advisable in all conditions, since it is clear that some waters are structurally important and that an explicit representation is therefore necessary. However, some explicit structural water can be included along with an implicit “bulk” water representation outside the explicit system.
Surface Area Approaches From the observation that the free energy of solvation for a saturated hydrocarbon is linearly related to the solvent-accessible surface area, one can create a set of empirical parameters to effectively represent the hydrophobic effect. For protein simulation, Eisenberg and McLachlan developed a series of effective atomic solvation parameters (σi) based on the water/octanol transfer free energy for a variety of amino acid analogs (Eisenberg and McLachlan, 1986). Using these parameters, which implicitly include the effective polarization in the parameterization, the free energy of interaction of a given residue with water (∆Gresidue) is related to the change in solvent-accessible surface area of that residue (in the folded state) or ∆Ai:
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∆Gresidue = ∑ σi∆Ai atoms,i
Scheraga and co-workers also developed this method instead using vapor/water transfer free energies (Ooi et al., 1987; Kang et al., 1988; Williams et al., 1992). Given the derivative of the solvent-accessible surface area (or exposed van der Waals surface) necessary for calculation of the forces, this term can be included in MD simulation (Wesson and Eisenberg, 1992; Schiffer et al., 1993). In practice, depending on the speed of calculating the solvent-accessible surface area (and derivatives), adding in these terms costs anywhere from ~1 to 4 times standard MD in vacuo. The relative costs relate to the accuracy of the derivatives; exact treatments are rather expensive, but more approximate treatments (which have errors in the 1% to 5% range) can be calculated much more rapidly (LeGrand and Merz, 1993; Sridharan et al., 1995; Fraczkiewicz and Braun, 1998). Although the calculation is fairly rapid compared to explicit solvent, inclusion of these derivatives into MD simulation has not been used extensively in biomolecular simulation to date. This is because the results are very sensitive to the parameters used—a good parameter set has not emerged from the studies—and solvent screening or polarization needs to be included (either with a rudimentary distance-dependent dielectric or more complex treatments). Moreover, although it may be argued that the hydrophobic driving force for protein folding might be reasonably represented by such a term, it is not clear if this approach will prove useful in nucleic acid molecular dynamics. With nucleic acids, the stability and conformational preferences are largely due to self-association and base stacking, hydrogen bonding with the base pairs and water, and electrostatic interactions related to the phosphates, associated ions, and solvent. Since base stacking is clearly a major driving force in nucleic acid stabilization, a solvent-accessible surface area term might allow global characterization of large-scale transitions, such as a single- to double-stranded DNA transition. However, subtle conformational differences, such as between A DNA and B DNA, which differ only very slightly in solvent-accessible surface area (Alden and Kim, 1979), will probably not be accurately estimated.
Adding in Effective Polarization Rather than fitting the solvation free energy (∆Gsol) to a single property, such as the solvent-accessible surface area, one can in principle partition ∆Gsol into a more natural and easily developed solvent-solvent and solute-solvent polarization term (∆Gpol), a solute cavity or first solvation layer term related to the formation of a cavity in the solvent (∆Gcavity), and a term related to immersing the uncharged solute into this cavity (∆Gvdw). The latter two terms, which loosely represent the hydrophobic effect, are typically represented by a solvent-accessible surface term parameterized to represent uncharged system. However, before discussing the polarization term in more detail, note that there is no way to uniquely partition the solvation free energy into these subparts, since each partition is not a state function; despite this, given appropriate parameterization, the solvation free energy can be reasonably estimated. The simplest way to represent the effective solvent polarization is by solving the Poisson equation. The simplest case is for an isolated point charge (q) within a spherical cavity of radius r and dielectric εint immersed in a dielectric continuum with dielectric constant ε. This is the Born equation (Born, 1920).
Molecular Modeling: Electrostatics and Solvation
∆GBorn =
q2 1 1 − 2r εint ε
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The value of this is a large quantity; for an isolated point charge (q = 1, εint = 1) with a radius of 5 Å immersed in a dielectric like water (ε ≈ 80) this represents approximately –66 kcal/mol. A similar expression for a neutral dipolar system (with total dipole moment µ assuming εint = 1) was developed by Onsager (1936). ∆GOnsager = −
2(ε − 1) µ 1 µ× 3 2 2ε + 1 r
In the above form, and a number of extensions (Kirkwood, 1939; Friedman, 1975), this is also referred to as a reaction field. A more general form applicable to real molecules (i.e., not isolated point charges or dipoles) is the Generalized Born (GB) formalism. For a series of overlapping atoms with charges qi immersed in a dielectric continuum with 1 dielectric constant ε, where αij are the Born radii [with αij = (αiαj) ⁄2] an approximate form is: ∆GGB =
qiqj 1 1 1 − ∑ ∑ 2 2 2 2 2 −r /(2αij) 1 / 2 ε ) i j (rij + αije ij
For superimposed charges, this gives the Born term, a Born plus Coulomb dielectric polarization for two spheres at longer distances, and, approximately, the Onsager reaction field at short range. When included with the surface area models discussed previously, this provides a very rapid and fairly accurate (depending on the parameterization) method to estimate solvation free energy for general molecules. In the context of molecular mechanics, this is the GBSA model (Still et al., 1990; Bashford and Case, 2000). This formalism has also been applied with reasonable success in the context of molecular orbital (semiempirical) theory (see Cramer and Truhlar, 1995, and references therein). This latter approach also defines a reasonable method for estimating the Born radii that is generalizable to molecular mechanics, uses the exposed van der Waals surface (instead of the solvent-accessible surface area), and calculates solvation free energies within ~0.55 kcal/mol over a test set of 255 molecules (Hawkins et al., 1997). This is available in the program AMSOL (Hawkins et al., 1996). A drawback of standard implementations of GBSA is that often the derivatives are simplified for efficiency reasons by assuming that the effective Born radii, which depend on all the pairwise distances, are constant over a given interval during the simulation; this reduces to sampling based on an effective dielectric with forces inconsistent with the GBSA potential. For a more complete and general treatment of the polarization by solvent, still in the context of a continuum representation (i.e., uniform bulk dielectric outside the “solute” or explicit system) or macroscopic solvent representation, the Poisson equation needs to be solved. If the effects of salt are to be included, the Poisson-Boltzmann equation can be applied. Except for some very simple cases (such as the sphere with independent ionizable groups, as discussed above, or cylinders), analytical solutions are not possible. Instead, finite difference or other numerical methods are applied. For a complete description of Poisson-Boltzmann (PB) methods, see any of a variety of reviews (Harvey, 1989; Sharp and Honig, 1990; Gilson, 1995). There are a variety of programs for solving the PB equation in common usage, such as Delphi (Gilson et al., 1987), UHBD (Madura et al., 1995), and MEAD (Bashford and Karplus, 1990), among others. Technical issues include the sensitivity to the approximating grid and radii for specifying the molecular surface and how to represent the hydrophobic effect (typically through the addition of a surface area term). Although calculation of the free energy of solvation is rapid compared to free energy perturbation in explicit solvent, solution of the PB equation is still moderately time-consuming. Therefore, although the methods have been incorporated into MD (Gilson and Honig, 1991; Sharp, 1991; Zauhar, 1991; Gilson et al., 1993), the
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complexity of the calculation (and need for first derivatives) has limited calculation to short time scales, and very few reports of its use in MD of macromolecules have been reported. Issues with the representation are that the model contains no microscopic description of water (although some limited explicit water can be included) and the results are strongly dependent not only on the bulk dielectric but on the internal dielectric within the solute. For proteins and nucleic acids, controversy surrounds what, precisely, the numerical value of the effective dielectric is (or whether it is even constant; Harvey, 1989; Warshel and Aqvist, 1991). Despite this, the PB model has proven very useful in a variety of applications; a nonexhaustive list includes investigating the salt dependence of DNA ligand interaction (Misra et al., 1994), electrostatic contributions to the B-to-Z DNA transition (Misra and Honig, 1996), base stacking (Friedman and Honig, 1995), and analysis of the stability of adenine bulge DNA conformations (Zacharias and Sklenar, 1997). Alternative methods for representing the solvent, intermediate between a continuum model and fully explicit water, are the Langevin dipole and protein-dipole Langevin dipole methods developed by Warshel and co-workers (Warshel and Levitt, 1976; Warshel and Russell, 1984; Warshel and Aqvist, 1991; Papazyan and Warshel, 1997). The idea here is that the most important effect of the solvent is the polarization rather than the detailed structural properties. Therefore, it is possible to replace the water by fixed-point polarizabilities (reorienting dipoles responding to the local electrostatic field according to the Langevin equation) on a cubic grid which is ultimately surrounded by a continuum. Since the orientation of the Langevin dipoles—based on the mean field of the local electrostatic field—depends on the local electrostatic field, these equations are typically iterated until self-consistency; however, a noniterative technique has also been applied (Lee et al., 1993). The method has been used for investigation of solvation free energies and electrostatic components in enzyme catalysis, and appears to be generally useful; however, the general simulation community has not extensively adopted its use.
Molecular Modeling: Electrostatics and Solvation
In addition to the methods discussed above, other techniques for representing the interaction with the continuum are possible, such as dielectric shielding (Luo et al., 1997). An important consideration with implicit solvent models is the lack of a microscopic understanding of the role of solvent, particularly any structural or dynamic role. Since the structural role is likely to be very important in modeling endeavors, this suggests that some explicit representation of the solvent is desirable. Therefore, although the implicit solvent models can give insight into the energetics and relative stability, the lack of microscopic detail limits its utility and in fact can be misleading. For example, simulations of a DNA triplex with implicit solvent models (either with a distance-dependent dielectric constant or a much more detailed Poisson-Boltzmann treatment) predict that an antiparallel third strand with reverse Hoogsteen base pairs would be more stable than a parallel third strand with Hoogsteen base pairs in d[CG-G]7 triplexes, in contrast to simulation in explicit solvent and experiment (Cheng and Pettitt, 1992, 1995). Of course, the corollary to this is that results of simulations in explicit water are not necessarily better! In some cases, short simulations of DNA with distance-dependent dielectric models agree better with experiment than simulations in explicit solvent (Fritsch et al., 1993). This points out the critical role of the nucleic acid force field, solvent parameters, stability of the methods, and the myriad of details that will become apparent in the next section. A final point is that various hybrid models can also be employed that treat an interior core (solute plus some water) explicitly with an implicit representation outside this core, for example, surrounding a nucleic acid in a blob of water by a reaction field or more detailed PB treatment.
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SIMULATIONS IN EXPLICIT SOLVENT There are a variety of considerations that become apparent when including an explicit solvent representation, and they boil down not only to how to represent the water and how much to add, but what type of boundary conditions to apply and how to make the calculations tractable. To make the computations tractable, we want to limit the effective number of interactions both by reducing the number of waters in the simulation and by applying cutoffs to limit the pair interactions to shorter range; however, as mentioned earlier, we want to keep in mind that misrepresenting the long-range interactions can lead to artifactual behavior. When considering how much solvent to use, we want not only enough water to solvate the solute (or nucleic acid of interest), but enough (either implicit or explicit) water surrounding this to ideally represent “bulk” water away from the solute. To allow investigation of the ensemble properties in this explicit system, we want to apply as simple and efficient a model as possible while still reasonably representing the structural, dynamic, and bulk properties of water. To represent the ensemble properties now requires detailed sampling over the various possible solvent configurations. This implies that explicit solvent should only be added for simulations employing Monte Carlo or molecular dynamics methods (UNIT 7.8), because minimization with explicit solvent will trap the structure into a less representative ice structure, rather than properly represent the ensemble properties of the solvent. In these simulations, various boundary conditions can be applied and are classified as either nonperiodic boundary conditions—e.g., surrounding the explicit system by a vacuum interface or a continuum—or as periodic boundary conditions—which effectively eliminate this interface. The most commonly used explicit water models are the rigid three-point water models such as TIP3P (Jorgensen et al., 1983) and SPC/E (Berendsen et al., 1987). These are the simplest models that can be applied that still retain the structural, energetic, and dynamic properties of water; for example, these models adequately reproduce the density, interaction energy, and first peak of the radial distribution function for water. The rigidity of the model requires the addition of constraints during molecular dynamics simulation; this is typically performed with the SHAKE procedure (Ryckaert et al., 1977). Although these are not necessarily the best water models, they are the most widely used. Drawbacks of the three-point models are an underestimation of compressibility, absence of much structure beyond the first peak of the radial distribution function, and less tetrahedrality than expected. The latter could have implications in simulation, since, although the bulk properties are acceptable, the reduced tetrahedrality could have structural implications. It should also be noticed that the two commonly used models were parameterized with different observables in mind. Although the TIP3P model accurately reproduces the interaction energies, it diffuses roughly twice what is expected. SPC/E, on the other hand, diffuses at the expected rate at the expense of slightly higher interaction energies. Neither of these models are polarizable, and therefore they are effectively prepolarized (through a larger fixed dipole moment); this is compatible with the pairwise force fields which do not include explicit polarization but implicitly represent the polarization with enhanced charges. Although slightly better water properties are obtained with the rigid four-point TIP4P (Jorgensen et al., 1983) model (which adds an extra charge on the OH bisector 0.15 Å from the oxygen), this model has not seen extensive usage in biomolecular simulation. In addition to the rigid water models, there have been reports of a variety of flexible water models; however, again these models have not seen extensive usage. As a final point, it should be noted that the commonly employed water models were parameterized for use with rather small cutoffs (in the 7 to 10 Å range). Yet, with the prescribed or even longer cutoffs, and the application of reliable cutoff methods such as atom-based force shifting in the 12 to 14 Å range, the neglect of the long-range electrostatic interactions can lead to artifacts in the transport properties and unexpected structure in
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the radial distribution function near the cutoff (Feller et al., 1996). This is particularly notable in simulations of polyionic systems. As will be discussed, much better behavior is instead observed in simulations that do not neglect the long-range electrostatic interactions. To make the calculations tractable, in practice the solute (or nucleic acid of interest) is only surrounded by roughly 5 to 20 Å of water in each direction. While this may appear to be a minimal representation, especially given the high charge typical on nucleic acids, good behavior is observed. Molecular dynamics simulations with as little as 5 Å of water surrounding the DNA in each direction give similar average structures and properties in nanosecond-length simulations compared to simulations with more water (Cheatham and Kollman, 1998; Norberto de Souza and Ornstein, 1997). Moreover, although it might be expected that the nucleic acid has far-reaching influence on the neutralizing counterion distribution, Manning theory suggests that the necessary neutralizing ions are contained within ~7 Å of the surface of duplex DNA, with “bulk” counterion distribution outside this range (Manning, 1978; Jayaram and Beveridge, 1996). Although the observation from these simulations and arguments about the nearness of the neutralizing counterion distribution can in part justify systems with as little as ~5 Å of water surrounding the nucleic acid, ~10 Å is a more common and tolerable value, despite the additional cost, that allows near “bulk” water away from the solute. Slightly more solvent may be necessary for nonperiodic systems, due to the surface/interface effects. Although in principle even more water may be desirable to better represent bulk solvent away from the nucleic acid and to allow effectively less concentrated solutions, the computational requirements typically limit the system size to less water. Even with a limited amount of solvent surrounding the nucleic acid model, it is necessary to further limit the number of pairwise interactions to make the calculations tractable. This is done through the application of a cutoff that is typically applied in the 9 to 15 Å range. Although this is likely an appropriate range for van der Waals interactions, it is typically insufficient for electrostatics, particularly with highly charged systems. Therefore, methods need to be applied to minimize artifacts from the truncation of the long-range electrostatic interactions (as will be discussed later).
Molecular Modeling: Electrostatics and Solvation
Nonperiodic Boundary Conditions The earliest molecular dynamics simulations of nucleic acids in explicit solvent used nonperiodic boundary conditions and surrounded a small piece of a DNA duplex, d[CGCGA], by a small blob of 806 TIP3P waters and 8 Na+ counterions with a vacuum interface outside of the explicit water (Seibel et al., 1985). The results from these simulations were similar to earlier in vacuo simulations, but provided the added insight that comes from a detailed investigation of hydration. Although “good” behavior was observed, this 100-psec-length simulation was too short to definitively verify the methods or to sample much of the accessible conformational space. It was also too short for the instabilities due to the cutoff of the electrostatic interactions to become apparent. Despite the apparent stability, however, even in this short run the water blob had distorted to such a degree that the terminal residues on one side of the duplex were nearly exposed to the vacuum. This can be prevented in principle by restraining the solute to remain at the center of the blob with spherical boundary potentials added at the surface to prevent solvent evaporation. The major issue with this type of representation is that the small size of the solvent blob, coupled with the reorientation of the surface waters in response to the vacuum interface, leads to a large surface tension. This surface tension, compounded by the standard boundary potentials added to prevent waters from drifting away, leads to an effectively large pressure at the center of the blob which can inhibit fluctuations, particularly at the center of the blob and at the surface where the waters are strongly
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ordered due to the vacuum interface (Fox and Kollman, 1996; Steinbach and Brooks, 1994). A crude estimate of the effective pressure increase is R∆P ≈ 15,000 Å-atm, where R is the radius of the solvent droplet. Because of the large surface tension and large pressures, this boundary representation is not recommended unless the surface effects are modulated, either through the use of a very large solvent blob (which is unfeasible) or by applying some means to break up the surface ordering and/or restore the effective polarization. An effective way to do this is to add stochastic forces to waters near the surface. This is shown schematically in Figure 7.9.1. This can be done via the Langevin equation (Berkowitz and McCammon, 1982) and leads to efficient simulations of proteins where only a core part of the system is solvated (Brooks et al., 1985, 1988). For proteins, this may be an adequate treatment since most of the strong ionic interactions are limited to the surface of the protein, and the water at the surface of the protein is not likely to play as large a structural and stabilizing role, nor the electrostatics as large a destabilizing role, as with nucleic acids. Nevertheless, this method has been applied in a limited fashion in the simulation of nucleic acids, e.g., to investigate the glass transition temperature of DNA (Norberg and Nilsson, 1996) and the hydration of guanyl-5′-3′-uridine (Norberg and Nilsson, 1995) in an ~20 Å sphere of water. Although applying random forces can partially eliminate the surface tension, the strong influence of the vacuum interface prevents its complete elimination. The surface ordering is fairly significant, and attempts to design restraining functions that break up this surface order have led to very complicated size, water model, and system-specific expressions that only work moderately well (Essex and Jorgensen, 1995). In addition to removing the ordered water at the surface, it is also important to include polarization by the bulk solvent outside the explicit system. This is not well reproduced by the random fluctuations of the surface water in stochastic boundary treatments. To include polarization terms in addition to radial constraints, Warshel and co-workers developed the SCAAS model (King and Warshel, 1989) to modulate the properties of the surface waters at a fixed radius of solvent. This model allows good simulation of the static (such as the radial distribution function) and dynamic (such as diffusion) properties for the water in spheres as small as 8 to 10 Å. The SCAAS model apparently also allows fairly size-independent charging free energies of ions in solution (King and Warshel, 1989; Aqvist, 1990); however, the fixed radius disallows characteristic density and volume fluctuations, and reportedly the total dipole of the system is unstable during the simulation. Also, in each of these cases, the solvent boundary is effectively fixed and parameterized for a particular radius. A related method has also been reported that allows the solvent boundary to move during the simulation (thereby not inhibiting solute volume
Figure 7.9.1 Representations of the system with nonperiodic boundary simulations. The picture on the left shows schematically what happens with stochastic boundary conditions, compared to a dielectric continuum, represented on the right.
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changes) and models the implicit solvent outside the explicit system with a reaction field treatment (Beglov and Roux, 1994). Given a particular implementation, these spherical “solvent boundary potentials” are parameterized to represent bulk water outside the explicit system and to minimize the surface water ordering effects. This parameterization is fairly system- and size-specific, and in the general case the model may not be appropriate unless the boundary potential is refined. A final concern is that none of the methods completely remove the effect of the interface. In spite of this, the methods have proved useful in applications ranging from ionic charging free energies and ion parameter development (Aqvist, 1990) to understanding peptide folding in solution (Mohanty et al., 1997), including the stochastic boundary applications to nucleic acid systems mentioned previously. Despite the apparent success of these methods in reproducing solvent properties and charging free energies, these methods have seen limited use in large-scale biomolecular simulation. Part of the reason is that, for biomolecular simulation, fairly large solvent spheres are necessary to completely surround the solvent. Since periodic boundary simulations can be run at a similar cost (and include shapes that are nearly spherical), periodic boundary conditions may be more appropriate. Otherwise, for small solutes and/or partially solvated macromolecules (i.e., where only a part of the system is treated in detail with explicit solvent), the solvent blobs with an appropriate boundary potential can be applied with good success. Given the expense of the simulations and issues related to the interface between the explicit and implicit system, periodic boundary conditions may be the more appropriate choice. This is particularly true for large systems, since most of the accurate solvent blob potentials discussed above are not amenable to a standard cutoff treatment because typically all pairwise interactions are represented. This makes the calculations rather expensive. However, to counter the computational expense, various groups have recently started to apply fast multipole methods, which allow more tractable calculation of all pairwise interactions. The fast multipole method recursively groups distant atoms into multipoles based on hierarchical trees (Greengard, 1988, 1994; Greengard and Rokhlin, 1989). This tremendously increases the efficiency for large systems. For even greater efficiency (at the expense of accuracy) the cell-multipole method can also be applied (Ding et al., 1992). Ewald methods can also be adapted for simulation of finite nonperiodic systems and may provide an alternative rapid treatment for representing all pairwise interactions (Hockney and Eastwood, 1981; Pollock and Glosli, 1996).
Molecular Modeling: Electrostatics and Solvation
Periodic Boundary Conditions An artificial construction that removes the problem of the vacuum or continuum interface at the edge of the explicit system is the imposition of toroidal or periodic boundary conditions (PBC; Allen and Tildesley, 1987). This, in principle, eliminates the vacuum interface by imposing a lattice structure on the molecular system, so that, effectively, a molecule leaving one side of the periodic box enters the opposite side (see Fig. 7.9.2). With this construction, the volume and pressure are rigorously defined. Scaling of the box size (and also, typically, the relative positions of the atoms/molecules) can be used to effectively couple the pressure to define an isobaric ensemble. Similarly, scaling the velocities of the atoms can be used to control temperature, and, together, coupling the temperature and pressure allows simulation of the constant temperature and pressure ensemble (as opposed to the constant volume and energy ensemble discussed so far). Temperature and pressure scaling have typically been performed using the weak-coupling method (Berendsen et al., 1984). More recently, various simulation codes have started to incorporate more accurate coupling methods that properly simulate the isothermal/isobaric ensemble, such as Nose-Hoover thermostated chains (Nose, 1984; Hoover, 1985) or the Langevin piston method (Feller et al., 1995).
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Figure 7.9.2 Periodic boundary conditions.
A potential issue with PBC is the imposition of true periodicity. In principle, a given atom should feel the influence of all its periodic images (see Fig. 7.9.3); for the long-range interactions, such as the electrostatics, this can be a strong influence (Valleau and Whittington, 1977). For example, we can consider a freely rotating dipole in a periodic box. Due to the influence of the periodic images, the dipole may not freely reorient. Similarly, imagine two charged particles separated by half the box length in a periodic box. Depending on the charge of the particles, they should either be attracted or repulsed. However, in a periodic lattice, due to the influence of the image particles, no net force will be felt by the particles. Additionally, due to the spatial periodicity, there may be correlation in the fluctuations or time-averaged properties; for example, a ripple or wave will not continue outward from the center unimpeded as in a nonperiodic system, but will chaotically interfere with the same ripple or wave from the image unit cells. These effects could, in principle, lead to drastic artifacts that may be more severe than the vacuum or continuum interface discussed in the previous section. In practice, with simulations including solvent with a sufficiently high permittivity (such as water) in a reasonably sized box and under equilibrium conditions, the artifacts appear minor in simulations that are truly periodic (Bader and Chandler, 1992; Smith and Pettitt, 1996; Smith et al., 1997). Also, in practice it is not necessary to treat the system as truly periodic. This is possible through the application of a cutoff that limits the pair interactions to those within the unit cell. In this way, no atom directly feels the influence of its neighbor; however, this comes at the cost of neglecting the long-range interactions.
A
B
Figure 7.9.3 Potential artifacts from imposition of true periodicity. (A) Freely rotating dipole versus a dipole confined to a periodic lattice. (B) Free charges versus charges in a periodic lattice.
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Figure 7.9.4
The various cutoff schemes.
There are two standard cutoff-truncation schemes in common usage (see Fig. 7.9.4). The standard cutoff scheme limits pair interactions about a given atom to only those atoms within a given radius. Alternatively, we can apply minimum image truncation, which includes all interactions within the unit cell centered on the given atom. However, this is not recommended because it tends to over-represent the interactions in the corners of the unit cell and leads to damped, anisotropic reorientational motion (Roberts and Schnitker, 1995). Note also that most molecular-mechanics programs, although using a radial cutoff, are limited to minimum image conditions, limiting the effective cutoff in periodic systems to half the shortest box length. The neglect of long-range interactions leads to a number of artifacts. The largest relates to the nature of the truncation. Standard atom- or charge group–based truncation (where the latter is performed to avoid splitting up charge groups such that if one atom from the group is within the cutoff, all the atoms within that group are assumed to be within the cutoff) simply neglects all the interactions outside some finite range, typically within ~8 to 20 Å. At this range the electrostatic interactions are still rather large, particularly with polyionic systems such as nucleic acids. The truncation leads to force discontinuities as atoms/groups enter or leave the cutoff, and these in turn can lead to large instabilities during the dynamics. For example, even with cutoffs in the 9 to 20 Å range and state-of-the-art force fields, group-based truncation leads to complete disruption of a explicitly solvated DNA duplex within ~500 psec (Cheatham et al., 1995). To avoid these problems, various techniques have been applied to smooth the discontinuity in the energy or forces, either by switching the potential (e.g., by adding a spline to bring the potential from the truncated value to zero over a short range, typically 2 to 4 Å) or shifting the entire potential to zero. However, given that the dynamics are largely dictated by the forces, these methods do not necessarily lead to better behavior. In the case of atom-based switching functions over a short range, fluctuations are completely inhibited. Much better behavior is obtained by shifting the electrostatic forces on an atom basis (and switching the van der Waals energies over a range of ~2 Å); this allows stable nanosecond-length MD of nucleic acids in solution (MacKerell et al., 1995; MacKerell, 1997; MacKerell and Banavali, 2000), although water transport properties may be misrepresented (Feller et al., 1996). For a more thorough discussion of cutoffs, switching, and shifting, see the published reports (Steinbach and Brooks, 1994; Levitt et al., 1995). In addition to smoothing the interactions to the cutoff, it might be expected that better behavior will be seen with larger cutoffs. To this end, dual-cutoff or multiple-time-step methods can be applied, which treat the short-range interactions in the typical fashion but also keep track of interactions out to a larger cutoff and update these longer-range interactions less frequently (Tuckerman et al., 1992; Biesiadecki and Skeel, 1993). However, the artifactual behavior of the cutoff is not monotonically related to the size of the cutoff (i.e., longer is not necessarily better) and is not completely eliminated by smoothing the force artifacts. Molecular Modeling: Electrostatics and Solvation
Despite the presence of significant artifacts from neglect of the long-range electrostatic interactions, cutoff approximations have been widely applied in the simulation of biomolecules such as nucleic acids. Part of the reason for this is the large size of typical
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biomolecular systems, which necessitates the use of cutoffs to limit the number of pair interactions to make the calculations more tractable. Until more recently, the time scales of the simulations (~100 psec) were not long enough to expose the deficiencies. Despite the neglect of long-range interactions, reasonable simulation of proteins was routinely observed in the earlier simulations and still continues to date (even with rather poor cutoff methods). This was not true of nucleic acid simulations, which were routinely plagued by instabilities and necessitated tricks to keep the nucleic acid stable, such as WatsonCrick hydrogen bonding restraints, reduced phosphate charges (McConnell et al., 1994), and restrained counterions (Tapia and Velazquez, 1997). Since it has more recently become possible to perform truly periodic simulations at a cost that is comparable to reasonable cutoff simulations, it is worthwhile to discuss the artifactual behavior that is seen when the long-range interactions are ignored, and compare this to the artifactual behavior that might be expected in truly periodic simulations (which fully treat the long-range electrostatic interactions). The artifacts due to the neglect or absence of the long-range electrostatic interactions are well known and include long-range orientational correlation, such as a strong anticorrelation of dipolar fluctuations (Neumann, 1983), and decreased rotational and translational motion (van Gunsteren et al., 1978). Additionally, ion pairing between chloride ions has been observed (Dang and Pettitt, 1987), as has an attractive potential of mean force between two Fe2+ ions in solution resulting from the cutoff (despite the use of a spline smoothed potential at the cutoff; Bader and Chandler, 1992). Despite the imposition of true periodicity (which might be expected to give force artifacts due to interaction with the periodic images), Ewald methods show the expected behavior (Bader and Chandler, 1992) and no Cl– ion pairing (Hummer et al., 1993). In addition to the force artifacts, cutoff methods (with or without switched potentials) massively distort the electrostatic potential around charged groups in comparison with what is expected, in contrast to Ewald methods (Smith and Pettitt, 1991). Perhaps the most dramatic example of the problems with cutoffs, which further demonstrate that the effect of truncation is not monotonically related to the length of the cutoff, is shown in simulations of an α-helical peptide by Schreiber and Steinhauser (1992a). In these simulations, the helix was unstable at an unreasonably short cutoff (6 Å), as might be expected. The helix was stable at an intermediate and commonly employed cutoff (10 Å), similar to what was seen in Ewald simulations. However, the helix was again unstable at a longer cutoff (14 Å). What is particularly interesting in this case is the fortuitous agreement with Ewald simulations when an intermediate cutoff value was used. This is not restricted to solvated protein simulation, as fortuitous agreement has also been seen in simulation of ions in water where the orientational correlation functions and transport properties were in good agreement with results from Ewald simulation (Roberts and Schnitker, 1995). This fortuitous cancellation of errors masks the deficiencies, and has further rationalized the use of cutoff methods in protein simulation. However, for polyionic systems, this fortuitous agreement is not typically seen. For example, in salt solution the correlations between like charged ions lead to higher than expected radial populations near the cutoff, compared to a corresponding depletion of oppositely charged ions (Auffinger and Beveridge, 1995). Also, as mentioned previously, except when the forces are smoothed at the cutoff, DNA duplexes will tend to distort (Cheatham et al., 1995). However, even with smooth forces at the cutoff, such as with the application of an atom-based force shifting method, long-range order appears in the radial distribution function, and lower translational diffusion and greater viscosity of water than expected are seen; this behavior is not seen in Ewald simulations (Feller et al., 1996). The presence of all these artifacts argues for the routine use of truly periodic methods or, alternatively, the inclusion of an implicit representation of the long-range electrostatics outside the cutoff. While the former may seem drastic, the lack of artifacts in simulations to date and the relative speed have made
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truly periodic methods the new “standard.” In spite of this, there has been some use of reaction field methods that partially offset some of the effects of the cutoff (even within the context of periodic boundary conditions). For free energy simulations, a simple Born correction may be appropriate. More generality comes from application of a reaction field within periodic boundary simulations to represent the interactions outside the cutoff (Schreiber and Steinhauser, 1992b; van Gunsteren et al., 1978; Tironi et al., 1995, 1997) despite the potential for force/energy instabilities from molecules entering or leaving the cutoff and difficulties representing net-charged systems. These methods have seen limited usage in biomolecular simulation.
Ewald and True Periodicity To move beyond minimum image conventions in periodic systems, it is necessary to extend the long-range interactions to a sum over the unit cells in the lattice. For truly periodic methods, this involves adding more and more unit cells into the summation until the potential converges. The electrostatic interactions are a special case, since the sum is conditionally convergent. In other words, the limiting value depends on the order of summation (for example, summing by adding more and more 2-D slabs, versus adding more and more spherical layers around the unit cell, leads to a different result). A method to evaluate the infinite sum is via the Ewald method (Ewald, 1921; Allen and Tildesley, 1987). This converts the infinite sum into a sum of a self energy plus two absolutely converging series—the screened real space or direct space interactions, and a series in reciprocal space—with an additional term (which represents the conditionally convergent part of the series) that depends on the surface and the shape of the sum at the limits. This later term, which results when the summation is represented by adding roughly spherical layers, depends on the dielectric boundary in the limit and the dipole of the unit cell (DeLeeuw et al., 1980). Under tin-foil boundary conditions, an infinite dielectric is assumed at the boundary (also called conducting boundary conditions) and this surface term vanishes. This is likely to be the appropriate boundary condition for liquids (since the liquid surface structure or polarization should disappear in the limit), whereas a dielectric boundary with dielectric constant ε = 1 may be more appropriate for crystals. Tin-foil boundary conditions are also appropriate in net-charged systems (due to the ill-defined dipole; Bogusz et al., 1998) and are the most common dielectric boundary conditions (since this surface term is zero). In practice, application of these boundary conditions does not appear to lead to significant artifacts in the simulation of nucleic acids. However, recent simulations demonstrate that these boundary conditions may overstabilize the correlation between dipoles at long distances; this suggests using dielectric boundary conditions that more closely match the system, depending on what properties are to be represented in the simulation (Boresch and Steinhauser, 1997). The bulk of the Ewald computation relates to the calculation of the direct and reciprocal space sums. The direct space sum represents the standard Coulombic potential screened to short range through the complementary error function, or erfc(). Since this has the same form as the standard Coulombic potential (with charges qi and qj), it is often calculated in the same manner through the application of pairlists that include all the pair interactions within a given spherical cutoff. Edirect = Molecular Modeling: Electrostatics and Solvation
1 2
∑ ∑ ∑′ i
j |n |≠0
qiqjerfc(κ|rij + n |) |rij + n |
In the above equation, the inner sum is over the unit cells, n = naA + nbB + ncC (for integers na,, nb, and nc where A, B, and C are the unit cell lengths) and the prime (′) in the sum over
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lattice vectors means omit the i = j interactions when n = 0 and the outer sums are over all atoms. The reciprocal space sum, so named since the summation is performed in reciprocal space, provides the remainder of the Ewald potential (correcting for the screening of the direct space sum). 1 Ereciprocal = 2πV
∑ m≠0
2 2 2 e−π m / κ
m2
|S(m)2| where S(m) = ∑qie2πim⋅ri i=1
In the above expression, V denotes the volume and m the reciprocal vectors . The self energy is: Eself = −
κ π √
∑q2i i
The parameter κ determines the width of the screening potential (typically a gaussian) that partitions the work between the reciprocal and direct space interactions. For a smaller κ value, more of the work is in the direct space interactions; this implies that a longer cutoff is necessary to accurately represent this term. With larger κ values, more of the work is partitioned into the reciprocal space. This requires more reciprocal vectors for the reciprocal sum (and shorter cutoffs in the direct space) for equivalent accuracy. More information on the relative speed and accuracy can be found in the published reports (Petersen, 1995; Pollock and Glosli, 1996; Toukmaji and Board, 1996; Darden et al., 1997). The additional work due to the explicit sum over the reciprocal vectors increases 3 the computational cost of Ewald sums (optimally order N ⁄2 where N is the number of particles). Until fairly recently, due to the increased computational cost, simulation with Ewald potentials were limited to fairly small systems. A significant advance in the past few years has been the emergence of fast Ewald methods. These significantly speed up the calculation of the Ewald sum by utilizing fast Fourier transforms (FFT) to speed the calculation of the reciprocal space interactions. Various methods have emerged, such as the particle mesh Ewald (Darden et al., 1993; Essmann et al., 1995), the particle-particle particle mesh Ewald (Luty et al., 1995; Pollock and Glosli, 1996), and the fast Fourier Poisson (York and Yang, 1994) methods, all of which are generalizations of the formulation by Hockney and Eastwood (1981). The speedup is made possible by effectively interpolating the irregularly spaced reciprocal space charges onto a regular grid, which then allows an FFT to perform the necessary convolutions. Achieving comparable accuracy to standard Ewald methods requires interpolating grids of sufficient density and reasonable interpolation of the charges; when this is done, equivalent accuracy and energy conservation are maintained. The use of the FFT changes the scaling of the Ewald method down to Nlog(N), which significantly increases the relative performance. In fact, since shorter cutoffs can be used for the direct space interactions (~9 Å) while still completely representing the long-range electrostatic interactions, these methods are typically faster than comparable simulations with atombased force shifts at the cutoff in the 12 to 14 Å range. For simulations up to ~50,000 atoms, these fast Ewald methods are the most efficient way to calculate the long-range interactions. For larger systems, periodic fast multipole methods can be applied (Schmidt and Lee, 1997), although these are slightly more complicated than standard fast multipole methods. With these methods, there is considerable controversy as to the relative accuracy and relative speed, so the 50,000-atom break-even point should not be taken as an absolute. Some of these issues are discussed in more detail in the literature (Petersen, 1995; Solvason et al., 1995; Challacombe et al., 1997). To date, almost all of the realistic simulations of biomolecular systems in the nanosecond time range that have applied a truly periodic method have involved the fast Ewald methods, and in particular the particle
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mesh Ewald method. This later code has been extensively used due to its availability in common MD codes (AMBER, CHARMM), its facility for constant pressure, and the smooth nature of the errors. As mentioned previously, application of the truly periodic methods comes at a cost: that of imposing true periodicity where all atoms in the unit cell now freely interact with their periodic images. Fortunately, for simulations in water the imposition of true periodicity does not seem to lead to significant artifacts. Dipolar rotation is not strongly inhibited; the difference between free versus hindered motion of a model dipole in water is less than kT (Smith and Pettitt, 1996) and the rotation of a small zwitterionic peptide is not inhibited (Smith et al., 1997). Also, as mentioned before, no like ion pairing is seen in water (Bader and Chandler, 1992; Hummer et al., 1993). Moreover, recent simulations show that the conformational potential of mean force of a blocked trialanine in a 26 Å box is similar in a truly periodic box to that calculated in the absence of true periodicity. However, care should be taken to understand the nature of potential artifacts, particularly for simulations that only include a limited amount of water. A recent study of a helical peptide suggested that when there is insufficient solvation, the peptide is artificially stabilized into an α-helical conformation (Weber et al., 2000). Since nucleic acids are typically well hydrated and in a dielectric medium with a sufficiently high permittivity, artifacts from the true periodicity are likely to be minor. Care should be taken with the application of these truly periodic methods in solvents with a lower dielectric, such as ethanol. However, in spite of possible artifacts from true periodicity, reasonable representation of DNA in ethanol/water mixtures has been reported in MD simulation (Cheatham et al., 1997; Sprous et al., 1998). Care should also be taken with net-charged, truly periodic systems (where, in principle, the Coulombic energy should diverge!) and particularly when the charges change during the simulation, such as in free energy perturbation simulations; this is discussed in a number of publications investigating free energy of ionic hydration (Figueirido et al., 1997; Hummer et al., 1997; Bogusz et al., 1998). For a more thorough discussion of the success of nucleic acid simulation using these methods, see the authors’ detailed review (Cheatham and Kollman, 2000). LITERATURE CITED Alden, C.J. and Kim, S.-H. 1979. Solvent-accessible surfaces of nucleic acids. J. Mol. Biol. 132:411-434. Allen, M.P. and Tildesley, D.J. 1987. Computer Simulation of Liquids. Oxford University Press, Oxford. Aqvist, J. 1990. Ion-water interaction potentials derived from free energy perturbation simulations. J. Phys. Chem. 94:8021-8024. Auffinger, P. and Beveridge, B.L. 1995. A simple test for evaluating the truncation effects in simulations of systems involving charged groups. Chem. Phys. Lett. 234:413-415. Bader, J.S. and Chandler, D. 1992. Computer simulation study of the mean forces between ferrous and ferric ions in water. J. Phys. Chem. 96:6423-6427. Bashford, D. and Case, D.A. 2000. Generalized Born models of molecular solvation effects. Annu. Rev. Phys. Chem. 51:129-152. Bashford, D. and Karplus, M. 1990. pKa’s of ionizable groups in proteins: Atomic detail from a continuum electrostatic model. Biochem. 29:10219-10225. Beglov, D. and Roux, B. 1994. Finite representation of in infinite bulk system: Solvent boundary potential for computer simulations. J. Chem. Phys. 100:9050-9063. Berendsen, H.J.C., Postma, J.P.M., van Gunsteren, W.F., DiNola, A., and Haak, J.R. 1984. Molecular dynamics with coupling to an external bath. J. Comp. Phys. 81:3684-3690. Berendsen, H.J.C., Grigera, J.R., and Straatsma, T.P. 1987. The missing term in effective pair potentials. J. Phys. Chem. 91:6269-6274. Molecular Modeling: Electrostatics and Solvation
Berkowitz, M.L. and McCammon, J.A. 1982. Molecular dynamics with stochastic boundary conditions. Chem. Phys. Lett. 90:215-217.
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Beveridge, D.L., Swaminathan, S., Ravishanker, G., Withka, J.M., Srinivasan, J., Prevost, C., Louise-May, S., Langley, D.R., DiCapua, F.M., and Bolton, P.H. 1993. Molecular dynamics simulations on the hydration, structure and motions of DNA oligomers. In Water and Biological Molecules (E. Westhof, ed.) pp. 165-225. Macmillan Press, New York. Biesiadecki, J.J. and Skeel, R.D. 1993. Dangers of multiple time step methods. J. Comp. Phys. 109:318-328. Bogusz, S., Cheatham, T.E. III, and Brooks, B.R. 1998. Removal of pressure and free energy artifacts in charged periodic systems via net charge corrections to the Ewald potential. J. Chem. Phys. 108:7070-7084. Boresch, S. and Steinhauser, O. 1997. Presumed versus real artifacts of the Ewald summation technique: The importance of dielectric boundary conditions. Ber. Bunsenges. Phys. Chem. 101:1019-1029. Born, M. 1920. Volumen der Hydratationswärme der Ione. Z. Phys. Chem. 1:45-48. Brooks, C.L., Brunger, A., and Karplus, M. 1985. Active site dynamics in protein molecules: A stochastic boundary-molecular dynamics approach. Biopolymers 24:843-865. Brooks, C.L. III, Karplus, M., and Pettitt, B.M. 1988. Proteins. A theoretical perspective of dynamics, structure, and thermodynamics. In Advances in Chemical Physics, Vol. 71 (I. Prigogine, and S.A. Rice, eds.). John Wiley & Sons, New York. Challacombe, M., White, C., and Head-Gordon, M. 1997. Periodic boundary conditions and the fast multipole method. J. Chem. Phys. 107:10131-10140. Cheatham, T.E. III and Kollman, P.A. 1998. Molecular dynamics simulation of nucleic acids in solution: How sensitive are the results to small perturbations in the force field and environment. In Structure, Motion, Interactions and Expression of Biological Macromolecules (M. Sarma and R. Sarma, eds.) pp. 99-116. Adenine Press, Schenectady, New York. Cheatham, T.E. III and Kollman, P.A. 2000. Molecular dynamics simulation of nucleic acids. Annu. Rev. Phys. Chem. 51:435-471. Cheatham, T.E. III, Miller, J.L., Fox, T., Darden, T.A., and Kollman, P.A. 1995. Molecular dynamics simulations on solvated biomolecular systems: The particle mesh Ewald method leads to stable trajectories of DNA, RNA and proteins. J. Am. Chem. Soc. 117:4193-4194. Cheatham, T.E. III, Crowley, M.F., Fox, T., and Kollman, P.A. 1997. A molecular level picture of the stabilization of A-DNA in mixed ethanol-water solutions. Proc. Natl. Acad. Sci. U.S.A. 94:9626-9630. Cheng, Y.-K. and Pettitt, B.M. 1992. Hoogsteen versus reverse-Hoogsteen base pairing in DNA triplex helices. J. Am. Chem. Soc. 114:4465-4474. Cheng, Y.-K. and Pettitt, B.M. 1995. Solvent effects on model d(CG-G)7 and d(TA-T)7 DNA triplex helices. Biopolymers 35:457-473. Cramer, C.J. and Truhlar, D.G. 1995. Continuum solvation models: Classical and quantum mechanical implementations. In Reviews in Computational Chemistry, Vol. 6 (K.D. Lipkowitz and D.B. Boyd, eds.) pp. 1-72. VCH, New York. Daggett, V., Kollman, P.A., and Kuntz, I.D. 1991. Molecular dynamics simulations of small peptides: Dependence on dielectric model and pH. Biopolymers 31:285-304. Dang, L.X. and Pettitt, B.M. 1987. Chloride ion pairs in water. J. Am. Chem. Soc. 109:5531-5532. Darden, T.A., York, D.M., and Pedersen, L.G. 1993. Particle mesh Ewald: An N log(N) method for Ewald sums in large systems. J. Chem. Phys. 98:10089-10092. Darden, T.A., Pedersen, L.G., Toukmaji, A.Y., Crowley, M.F., and Cheatham, T.E. III. 1997. Eighth SIAM conference on parallel processing for scientific computing, Minneapolis, Minn. Society for Industrial and Applied Mathematics, Philadelphia. Davis, M.E. and McCammon, J.A. 1990. Electrostatics in biomolecular structure and dynamics. Chem. Rev. 90:509-521. DeLeeuw, S.W., Perram, J.M., and Smith, E.R. 1980. Simulation of electrostatic systems in periodic boundary conditions. I. Lattice sums and dielectric constants. Proc. R. Soc. Lond. A373:27-56. Dill, K.A. 1990. Dominant forces in protein folding. Biochemistry 29:7133-7155. Ding, H.Q., Karasawa, N., and Goddard, W.A. 1992. Atomic level simulations on a million particles: The cell multipole method for Coulomb and London nonbond interactions. J. Chem. Phys. 97:4309-4315. Drew, H.R. and Dickerson, R.E. 1981. Structure of a B-DNA dodecamer. III. Geometry of hydration. J. Mol. Biol. 151:535-556. Eisenberg, D. and McLachlan, A.D. 1986. Solvation energy in protein folding and binding. Nature 319:199203. Essex, J.W. and Jorgensen, W.L. 1995. An empirical boundary potential for water droplet simulations. J. Comp. Chem. 16, 951-972. Essmann, U., Perera, L., Berkowitz, M.L., Darden, T., Lee, H., and Pedersen, L.G. 1995. A smooth particle mesh Ewald method. J. Chem. Phys. 103:8577-8593.
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Ewald, P. 1921. Investigations of crystals by means of Roentgen rays. Ann. Phys. (Leipzig) 64:253-264. Feller, S.E., Zhang, Y., Pastor, W., and Brooks, B.R. 1995. Constant pressure molecular dynamics simulation: The Langevin piston method. J. Chem. Phys. 103:4613-4621. Feller, S.E., Pastor, R.W., Rojnuckarin, A., Bogusz, S., and Brooks, B.R. 1996. Effect of electrostatic force truncation on interfacial and transport properties of water. J. Phys. Chem. 100:17011-17020. Figueirido, F., Del Buono, G.S., and Levy, R.M. 1997. On finite-size corrections to the free energy of ionic hydration. J. Phys. Chem. 101:5622-5623. Flatters, D., Zakrzewska, K., and Lavery, R. 1997. Internal coordinate modeling of DNA: Force field comparisons. J. Comp. Chem. 18:1043-1055. Fox, T. and Kollman, P.A. 1996. The application of different solvation and electrostatic models in molecular dynamics simulations of ubiquitin: How well is the X-ray structure “maintained.” Proteins 25:315-334. Fraczkiewicz, R. and Braun, W. 1998. Exact and efficient analytical calculation of the accessible surface areas and their gradients for macromolecules. J. Comp. Chem. 19:319-333. Friedman, H. 1975. Image approximation to the reaction field. Mol. Phys. 29:1533-1543. Friedman, R.A. and Honig, B. 1992. The electrostatic contribution to DNA base-stacking interactions. Biopolymers 32:145-159. Friedman, R.A. and Honig, B. 1995. A free energy analysis of nucleic acid base stacking in aqueous solution. Biophys. J. 69:1528-1535. Fritsch, V., Ravishanker, G., Beveridge, D.L., and Westhof, E. 1993. Molecular dynamics simulations of poly(dA)-poly(dT): Comparisons between implicit and explicit solvent representations. Biopolymers 33:1537-1552. Gilson, M.K. 1995. Theory of electrostatic interactions in macromolecules. Curr. Opin. Struct. Biol. 5:216-223. Gilson, M.K. and Honig, B. 1991. The inclusion of electrostatic hydration energies in molecular mechanics calculations. J. Comp. Aided Mol. Des. 5:5-20. Gilson, M.K., Sharp, K.A., and Honig, B.H. 1987. Calculating the electrostatic potential of molecules in solution: Method and error assessment. J. Comp. Chem. 9:327-335. Gilson, M.K., Davis, M.E., Luty, B.A., and McCammon, J.A. 1993. Computation of electrostatic forces on solvated molecules using the Poisson-Boltzmann equation. J. Phys. Chem. 97:3591-3600. Greengard, L. 1988. The Rapid Evaluation of Potential Fields in Particle Systems. MIT Press, Cambridge, Mass. Greengard, L. 1994. Fast algorithms for classical physics. Science 265:909-914. Greengard, L. and Rokhlin, V. 1989. On the evaluation of electrostatic interactions in molecular modeling. Chem. Scrip. 29A:139-144. Harvey, S.C. 1989. Treatment of electrostatic effects in macromolecular modeling. Proteins 5:78-92. Hawkins, G.D., Lynch, G.C., Giesen, D.J., Rossi, I., Storer, J.W., Liotard, D.A., Cramer, C.J., and Thular, D.G. 1996. AMSOL (QCPE Bull. 16, 11), Minnesota, Minn. Hawkins, G.D., Cramer, C.J., and Truhlar, D.G. 1997. Parameterized model for aqueous free energies of solvation using geometry-dependent atomic surface tensions with implicit electrostatics. J. Phys. Chem. B. 101:7147-7157. Hingerty, B.E., Ritchie, R.H., Ferrell, T.L., and Turner, J.E. 1985. Dielectric effects in biopolymers: The theory of ionic saturation revisited. Biopolymers 24:427-439. Hockney, R.W. and Eastwood, J.W. 1981. Computer simulation using particles. McGraw-Hill, New York. Hoover, W.G. 1985. Canonical dynamics: Equilibrium phase distributions. Phys. Rev. A. 31:1695-1697. Hummer, G., Soumpasis, D.M., and Neumann, M. 1993. Computer simulations do not support Cl-Cl pairing in aqueous NaCl solution. Mol. Phys. 81:1155-1163. Hummer, G., Pratt, L.R., and Garcia, A. 1997. Ion sizes and finite-size corrections for ionic-solvation free energies. J. Chem. Phys. 107:9275-9277. Jayaram, B. and Beveridge, D.L. 1996. Modeling DNA in aqueous solutions: Theoretical and computer simulation studies on the ion atmosphere of DNA. Annu. Rev. Biophys. Biomol. Struct. 25:367-394. Jorgensen, W.L., Chandrasekhar, J., Madura, J.D., Impey, R.W., and Klein, M.L. 1983. Comparison of simple potential functions for simulating liquid water. J. Chem. Phys. 79:926-935. Molecular Modeling: Electrostatics and Solvation
Kang, Y.K., Nemethy, G., and Scheraga, H.A. 1988. Free energies of hydration of solute molecules. 4. Revised treatment of the hydration shell model. J. Chem. Phys. 79:926-935. King, G. and Warshel, A. 1989. A surface constrained all-atom solvent model for effective simulations of polar solutions. J. Chem. Phys. 91:3647-3661.
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Kirkwood, J.G. 1939. The dielectric polarization of liquids. J. Chem. Phys. 7:911-919. Lebrun, A. and Lavery, R. 1996. Modelling extreme stretching of DNA. Nucl. Acids Res. 24:2260-2267. Lebrun, A., Shakked, Z., and Lavery, R. 1997. Local DNA stretching mimics the distortion caused by TATA box-binding protein. Proc. Natl. Acad. Sci. U.S.A. 94:2993-2998. Lee, F.S., Chu, Z.T., and Warshel, A. 1993. Microscopic and semimicroscopic calculations of electrostatic energies in proteins by the Polaris and Enzymix programs. J. Comp. Chem. 14:161-185. LeGrand, S.M. and Merz, K.M. 1993. Rapid approximation to molecular surface area via the use of boolean logic and look-up tables. J. Comp. Chem. 14:349-352. Levitt, M. 1983. Computer simulation of DNA double-helix dynamics. Cold Spring Harbor. Symp. Quant. Biol. 47:251-262. Levitt, M., Hirshberg, M., Sharon, R., and Daggett, V. 1995. Potential energy function and parameters for simulations of the molecular dynamics of proteins and nucleic acids in solution. Comp. Phys. Comm. 91:215-231. Luo, R., Moult, J., and Gilson, M.K. 1997. Dielectric screening treatment of electrostatic solvation. J. Phys. Chem. B. 101:11226-11236. Luty, B.A., Tironi, I.G., and van Gunsteren, W.F. 1995. Lattice-sum methods for calculating electrostatic interactions in molecular simulations. J. Chem. Phys. 103:3014-3021. MacKerell, A.D., Jr. 1997. Influence of magnesium ions on duplex DNA structural, dynamic, and solvation properties. J. Phys. Chem. B101:646-650. MacKerell, A.D. Jr. and Banavali, N. 2000. All-atom empirical force field for nucleic acids. 2. Application to molecular dynamics simulations of DNA and RNA in solution. J. Comp. Chem. 21:105-120. MacKerell, A.D. Jr., Wiorkiewicz-Kuczera, J., and Karplus, M. 1995. An all-atom empirical energy function for the simulation of nucleic acids. J. Am. Chem. Soc. 117:11946-11975. Madura, J.D., Briggs, J.M., Wade, R.C., Davis, M.E., and McCammon, J.A. 1995. Electrostatics and diffusion of molecules in solution—Simulations with the University of Houston brownian dynamics program. Comp. Phys. Comm. 91:57-95. Manning, G.S. 1978. The molecular theory of polyelectrolyte solutions with applications to the electrostatic properties of polynucleotides. Quart. Rev. Biophys. 2:159-246. McConnell, K.J., Nirmala, R., Young, M.A., Ravishanker, G., and Beveridge, D.L. 1994. A nanosecond molecular dynamics trajectory for a B DNA double helix - Evidence for substates. J. Am. Chem. Soc. 116:4461-4462. Misra, V.K. and Honig, B. 1996. The electrostatic contribution to the B to Z transition of DNA. Biochem. 35:1115-1124. Misra, V.K., Sharp, K.A., Friedman, R.A., and Honig, B. 1994. Salt effects on ligand-DNA binding. Minor groove binding antibiotics. J. Mol. Biol. 238:245-263. Mohanty, D., Elber, R., Thirumalai, D., Beglov, D., and Roux, B. 1997. Kinetics of protein folding: Computer simulations of SYPFDV and peptide variants in water. J. Mol. Biol. 272:423-442. Neumann, M. 1983. Dipole moment fluctuation formulas in computer simulations of polar systems. Mol. Phys. 50:841-858. Norberg, J. and Nilsson, L. 1995. NMR relaxation times, dynamics, and hydration of a nucleic acid fragment from molecular dynamics simulations. J. Phys. Chem. 99:14876-14884. Norberg, J. and Nilsson, L. 1996. Glass transition in DNA from molecular dynamics simulations. Proc. Natl. Acad. Sci. U.S.A. 93:10173-10176. Norberto de Souza, O. and Ornstein, R.L. 1997. Effect of periodic box size on aqueous molecular dynamics simulation of a DNA dodecamer with particle-mesh Ewald method. Biophys. J. 72:2395-2397. Nose, S. 1984. A molecular dynamics method for simulations in the canonical ensemble. Mol. Phys. 52:255-268. Onsager, L. 1936. Electric moments of molecules in liquids. J. Am. Chem. Soc. 58:1486-1493. Ooi, I., Oobatake, M., Nemethy, G., and Scheraga, H.A. 1987. Accessible surface areas as a measure of the thermodynamic parameters of hydration of peptides. Proc. Natl. Acad. Sci. U.S.A. 84:3086-3090. Papazyan, A. and Warshel, A. 1997. Continuum and dipole-lattice models of solvation. J. Phys. Chem. B 101:11254-11264. Petersen, H.G. 1995. Accuracy and efficiency of the particle mesh Ewald method. J. Chem. Phys. 103:36683679. Pollock, E.L. and Glosli, J. 1996. Comments on P3M, PMM, and the Ewald method for large periodic Coulombic systems. Comp. Phys. Comm. 95:93-110.
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Ramstein, J. and Lavery, R. 1988. Energetic coupling between DNA bending and base pair opening. Proc. Natl. Acad. Sci. U.S.A. 85:7231-7235. Roberts, J.E. and Schnitker, J. 1995. Boundary conditions in simulations of aqueous ionic solutions: A systematic study. J. Phys. Chem. 99:1322-1331. Ryckaert, J.P., Ciccotti, G., and Berendsen, H.J.C. 1977. Numerical integration of the cartesian equations of motion of a system with constraints: Molecular dynamics of n-alkanes. J. Comp. Phys. 23:327-341. Sarai, A., Mazur, J., Nussinov, R., and Jernigan, R.L. 1988. Origin of DNA helical structure and its sequence dependence. Biochemistry 27:8498-8502. Schiffer, C.A., Caldwell, J.W., Kollman, P.A., and Stroud, R.M. 1993. Protein structure prediction with a combined solvation free energy-molecular mechanics force field. Mol. Sim. 10:121-149. Schmidt, K.E. and Lee, M.A. 1997. Multipole Ewald sums for the fast multipole method. J. Stat. Phys. 89:411-424. Schreiber, H. and Steinhauser, O. 1992a. Cutoff size does strongly influence molecular dynamics results on solvated polypeptides. Biochemistry 31:5856-5860. Schreiber, H. and Steinhauser, O. 1992b. Taming cutoff induced artifacts in molecular dynamics studies of polypeptides. The reaction field method. J. Mol. Biol. 228:909-923. Seibel, G.L., Singh, U.C., and Kollman, P.A. 1985. A molecular dynamics simulation of double-helical B-DNA including counterions and water. Proc. Nat. Acad. Sci. U.S.A. 82:6537-6540. Sharp, K.A. 1991. Incorporating solvent and ion screening into molecular dynamics using the finite-difference Poisson-Boltzmann approach. J. Comp. Chem. 12:454-468. Sharp, K.A. and Honig, B. 1990. Electrostatic interactions in macromolecules: Theory and applications. Annu. Rev. Biophys. Biophys. Chem. 19:301-332. Singh, U.C., Weiner, S.C., and Kollman, P.A. 1985. Molecular dynamics simulations of d(C-G-C-G-A)-d(TC-G-C-G) with and without “hydrated” counterions. Proc. Natl. Acad. Sci. U.S.A. 82:755-759. Smith, P.E. and Pettitt, B.M. 1991. Peptides in ionic solution—A comparison of the Ewald and switching function techniques. J. Chem. Phys. 95:8430-8441. Smith, P.E. and Pettitt, B.M. 1996. Ewald artifacts in liquid state molecular dynamics simulations. J. Chem. Phys. 105:4289-4293. Smith, P.E., Blatt, H.D., and Pettitt, B.M. 1997. On the presence of rotational Ewald artifacts in the equilibrium and dynamical properties of a zwitterionic tetrapeptide in aqueous solution. J. Phys. Chem. 101B:38863890. Solvason, D., Kolafa, J., Petersen, H.G., and Perram, J.W. 1995. A rigorous comparison of the Ewald method and the fast multipole method in two dimensions. Comp. Phys. Comm. 87:307-318. Spolar, R.S. and Record, M.T. 1994. Coupling of local folding to site-specific binding of proteins to DNA. Science 263:777-784. Sprous, D., Young, M.A., and Beveridge, D.L. 1998. Molecular dynamics studies of the conformational preferences of a DNA double helix in water and an ethanol/water mixture: Theoretical considerations of the A-B transition. J. Phys. Chem. B. 102:4658-4667. Sridharan, S., Nicholls, A., and Sharp, K.A. 1995. A rapid method for calculating derivatives of solvent accessible surface areas of molecules. J. Comp. Chem. 16:1038-1044. Steinbach, P.J. and Brooks, B.R. 1994. New spherical-cutoff methods for long-range forces in macromolecular simulation. J. Comp. Chem. 15:667-683. Still, W.C., Tempczyk, A., Hawley, R.C., and Hendrickson, T. 1990. Semi analytical treatment of solvation for molecular mechanics and dynamics. J. Am. Chem. Soc. 112:6127-6128. Tapia, O. and Velazquez, I. 1997. Molecular dynamics simulations of DNA with protein’s consistent GROMOS force field and the role of counterions’ symmetry. J. Am. Chem. Soc. 119:5934-5938. Tironi, I.G., Sperb, R., Smith, P.E., and van Gunsteren, W.F. 1995. A generalized reaction field method for molecular dynamics simulations. J. Chem. Phys. 102:5451-5459. Tironi, I.G., Luty, B.A., and van Gunsteren, W.F. 1997. Space-time correlated reaction field: A stochastic dynamical approach to the dielectric continuum. J. Chem. Phys. 106:6068-6075. Toukmaji, A.Y. and Board, J.A.J. 1996. Ewald summation techniques in perspective: A survey. Comp. Phys. Comm. 95:73-92. Tuckerman, M., Berne, B.J., and Martyna, G.J. 1992. Reversible multiple time scale molecular dynamics. J. Chem. Phys. 97:1990-2001. Molecular Modeling: Electrostatics and Solvation
Valleau, J.P. and Whittington, S.G. 1977. A guide to Monte Carlo for statistical mechanics: 1. Highways. In Statistical Mechanics A: A Modern Theoretical Chemistry, Vol. 5-6 (B.J. Berne, ed.), pp. 137-167. Plenum Press, New York.
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van Gunsteren, W.F., Berendsen, H.J.C., and Rullman, J.A.C. 1978. Inclusion of a reaction field in molecular dynamics: Application to liquid water. Faraday Disc. 66:58-70. von Kitzing, E. and Diekmann, S. 1987. Molecular mechanics calculations of dA12-dT12 and the curved molecule d(GCTCGAAAAA)4-d(TTTTTCGAGC)4. Eur. Biophys. J. 15:13-26. Warshel, A. and Aqvist, J. 1991. Electrostatic energy and macromolecular function. Annu. Rev. Biophys. Biophys. Chem. 20:267-298. Warshel, A. and Levitt, M. 1976. Theoretical studies of enzyme reactions: Dielectric, electrostatic and steric stabilization of the carbonium ion in the reaction of lysozyme. J. Mol. Biol. 103:227-249. Warshel, A. and Russell, S.T. 1984. Calculation of electrostatic interactions in biological systems and in solution. Quart. Rev. Biophys. 17:283-422. Weber, W., Hunenberger, P.H., and McCammon, J.A. 2000. Molecular dynamics simulations of a polyalanine octapeptide under Ewald boundary conditions: Influence of artificial periodicity on peptide conformation. J. Phys. Chem. B. 104:3668-3675. Wesson, L. and Eisenberg, D. 1992. Atomic solvation parameters applied to molecular dynamics of proteins in solution. Prot. Sci. 1:227-235. Williams, R.L., Vila, J., Perrot, G., and Scheraga, H.A. 1992. Empirical solvation models in the context of conformational energy searches: Application to bovine pancreatic trypsin inhibitor. Proteins 14:110-119. York, D. and Yang, W. 1994. The fast Fourier Poisson method for calculating Ewald sums. J. Chem. Phys. 101:3298-3300. Zacharias, M. and Sklenar, H. 1997. Analysis of the stability of looped-out and stacked-in conformations of an adenine bulge in DNA using a continuum model for solvent and ions. Biophys. J. 73:2990-3003. Zauhar, R.J. 1991. The incorporation of hydration forces determined by continuum electrostatics into molecular mechanics simulations. J. Comp. Chem. 12:575-583. Zhurkin, V.B., Ulyanov, N.B., Gorin, A.A., and Jernigan, R.L. 1991. Static and statistical bending of DNA evaluated by Monte Carlo calculations. Proc. Natl. Acad. Sci. U.S.A. 88:7046-7050.
Contributed by Thomas E. Cheatham, III University of Utah Salt Lake City, Utah Bernard R. Brooks National Heart, Lung, and Blood Institute Bethesda, Maryland Peter A. Kollman University of California San Francisco, California
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Molecular Modeling of Nucleic Acid Structure: Setup and Analysis
UNIT 7.10
ISSUES IN THE SIMULATION OF NUCLEIC ACIDS From the information presented in previous units (UNITS 7.5, 7.8 & 7.9), one should have a reasonable understanding of the various trade-offs that are necessary to model nucleic acid structures. For instance, when choosing an energy representation and a means to sample relevant conformations, there is a trade-off between detail, sampling, and computational cost. Although the discussions thus far have presented basic means for modeling nucleic acids, some important details have not been sufficiently addressed. Reasonable questions that remain are: (1) what empirical molecular mechanical force field is appropriate; (2) if one wants to run an accurate simulation of a nucleic acid in an explicit solvent, how does one set up and equilibrate the system; and (3) how does one analyze molecular dynamics trajectories? This unit provides the answers to some of these questions and outlines a protocol for accurate simulation of nucleic acids. Which Molecular Mechanics Force Field Is Appropriate? This is a difficult question. The answer is complicated, often contentious, and in part depends on what representation (i.e., implicit versus explicit solvent, internal coordinate versus all-atom) is applied. With an internal coordinate representation, where bonds and angles are fixed, the most widely used molecular mechanical force field for nucleic acids is the FLEX force field within JUMNA (Lavery et al., 1995). Reasonable success has also been observed with internal coordinate force fields developed by Zhurkin et al. (1980, 1991). With internal coordinate molecular mechanical force fields, solvent is rarely included explicitly and is instead modeled via a simple implicit solvent model. Effective dielectric treatments such as the distance-dependent dielectric (ε = 4rij) or sigmoidal dielectric functions are commonly applied (as discussed in more detail in UNIT 7.9). More recently, increases in computer power have led to the incorporation of more accurate and implicit solvent models, such as finite difference Poisson-Boltzmann methods (Zacharias and Sklenar, 1997). Although it is not directly possible to mix and match the internal coordinate force fields with those designed for all-atom simulation, the FLEX force field has been shown to agree well with the all-atom nucleic acid force field described by Cornell et al. (1995) and Flatters et al. (1997). For more information on the differences between internal coordinate and all-atom representations, see UNIT 7.8. For simulations where each atom is free to move (known as all-atom simulation), force fields have steadily and continually improved in recent years. For simulations including explicit solvent and a proper treatment of the electrostatic interactions, the most widely applied force fields for nucleic acids have been the Cornell et al. (1995) force field, the MacKerell force fields in CHARMM (Mackerell et al., 1995), and the force field in GROMOS (van Gunsteren and Berendsen, 1987). The latter force field performs poorly in nanosecond-length simulation (Tapia and Velazquez, 1997), presumably due to improper treatment of the long-range electrostatic interactions. The tendency to fail when the long-range electrostatic interactions are not properly included or omitted appears to be a general property of most of the available empirical force fields (Cheatham et al., 1995). As discussed in UNIT 7.9, to treat the electrostatic interactions correctly, it is recommended that a smooth cut-off method be applied (such as an atom-based forceshifted cut-off) or that the electrostatic interactions be fully included via an Ewald treatment. Biophysical Analysis of Nucleic Acids Contributed by Thomas E. Cheatham, III, Bernard R. Brooks, and Peter A. Kollman Current Protocols in Nucleic Acid Chemistry (2001) 7.10.1-7.10.17 Copyright © 2001 by John Wiley & Sons, Inc.
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Each of the currently available force fields for nucleic acids has various strengths and weaknesses. Direct comparison of all is difficult since, to date, there has not been a published systematic study of each, with consistent simulation protocols, to benchmark their relative performances. Despite this caveat, based on the authors’ analysis of the literature and the authors’ experience with many of these force fields, some insight can be found. The earlier MacKerell force field (Mackerell et al., 1995), also referred to as the all22 force field in CHARMM, accurately represents canonical A-form DNA structures. This force field does not properly stabilize canonical B-form DNA structures since slow transitions to A-DNA are seen in solutions with low salt conditions (Norberg and Nilsson, 1996a; Feig and Pettitt, 1997; MacKerell, 1997, 1998). This has been remedied in a more recent parameterization (all27; Foloppe and MacKerell, 2000; MacKerell and Banavali, 2000) that does a better job on B-DNA structures, albeit with less sequencespecific minor groove narrowing than expected. More consistent behavior with nucleic acids is observed with the Cornell et al. force field in conjunction with a particle mesh Ewald treatment of the electrostatic interactions. With this force field, spontaneous A-DNA to B-DNA transitions are seen with a variety of sequences as expected (Cheatham and Kollman, 1996, 1998), and B-DNA to A-DNA transitions are observed with phosphoramidate-modified backbones (Cieplak et al., 1997), consistent with those seen in experiments. This observation is very exciting since it suggests that conformational sampling is not overly inhibited under truly periodic boundary conditions for DNA in explicit solvents. The force field also predicts A-RNA to be stable, although B-RNA to A-RNA transitions do not occur spontaneously (Cheatham and Kollman, 1997b). This brings up the issue of poor sampling of RNA in nanosecond-length simulation since B-RNA is stable for >10 nsec unless concerted changes in the sugar pucker are forced. This is likely due to the larger barriers to conformational transition due to 2′-O interactions with the backbone and larger barriers to sugar repuckering. With this force field (and likely others), modelers should be aware that sampling is more limited in molecular dynamics simulations of RNA. The Cornell et al. force field also reproduces sequencespecific structures, such as the expected TpG step bends in the major groove or the narrowing of the minor groove in polyadenine (A-tract) regions, and shows good agreement with crystal data (Young et al., 1997b). Expected structural differences between DNA-DNA, DNA-RNA, and RNA-RNA duplexes are well modeled, as are modified nucleic acids such as phosphoramidates (Cieplak et al., 1997) and photo-damaged DNA (Miaskiewicz et al., 1996; Spector et al., 1997). More recently, the Cornell et al. force field has also demonstrated the ability to model (with the same force field and simulation protocol) changes in nucleic acid structure that result from changes in the solvent environment. This includes the stabilization of A-DNA in a water and ethanol solution (Cheatham et al., 1997) and spontaneous B-DNA to A-DNA transitions in the presence of hexaammine cobalt(III) (Cheatham and Kollman, 1997a). Limitations of this force field include lower-than-expected sugar pucker, χ angles, and helical twist. This has been improved in a more recent parameterization (called parm98 or parm99) of the dihedral terms (Cheatham et al., 1999). Another force field worthy of notice is the BMS (Bristol Meyer Squibb) force field for nucleic acids (Langley, 1998), which was explicitly parameterized in order to properly simulate A-DNA/B-DNA equilibria under various conditions including water/ethanol and high salt.
Molecular Modeling of Nucleic Acid Structure: Setup and Analysis
Note that with all of these force fields the terminal phosphates are generally not included and, moreover, there is a distinction between internal, free, and 3′- or 5′-terminal residues. With each force field or simulation program, there are procedures to handle this distinction either through the use of different names for the residues (as in AMBER) or using various procedures designed to patch the terminal residues to shift charge and delete the terminal phosphate group (as in CHARMM).
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Given the constantly changing landscape, the limited application of some of these force fields, and variations in the applied methods, it is difficult to evaluate which force field is “best”. Experience plays a part in the selection of a force field and, therefore, careful evaluation of the published reports is important. A final point is that not all force fields are compatible with a given simulation code. The Cornell et al. force field is released with AMBER, the MacKerell force fields with CHARMM, and other force fields with other codes such as GROMOS or Insight/Discover. Since CHARMM and AMBER use a similar molecular mechanics potential and equivalent Lennard-Jones combining rules, these force fields can be interconverted; this is not easily possible for GROMOS, which uses geometric-mean combining rules and has a different form for the bond- and angle-stretching terms. Besides choosing which force field to use, an additional question is what to do if parameters are missing. This is a difficult question to answer in general terms, however, as most force fields have a specific protocol that should be followed to develop new parameters. For example, with the Cornell et al. force field, new intramolecular parameters are chosen by analogy to be consistent with existing parameters, general van der Waals parameters are obtained from simulations of neat liquids (e.g., OPLS; Jorgensen et al., 1996), and restrained electrostatic potential (RESP) fit charges from ab initio calculations are obtained for each new nucleotide, residue, or substructure. For more information about adding missing parameters, one should read all of the relevant force field literature and search for guides or repositories of existing parameters on the Internet. It is also possible to pose questions to the various e-mail reflectors for each program to ask if other investigators have already developed parameters for the system of interest. Additionally, it may be appropriate to contact the corresponding author of the force field papers for more information on the specific protocol for developing new parameters. Balance is also an important requirement for a given force field; this means that in addition to accurately modeling the intra-DNA interaction, it is important to have a balanced representation of the DNA with solvent. The current force fields (Cornell et al., MacKerell, and Langley) appear reasonably balanced. In addition to balance of the DNA with the explicit solvent, these force fields, in general, allow simulation of protein systems. Reasonable representation of protein–nucleic acid structures has been seen with all three of the major force field derivatives (Cornell et al., MacKerell, and BMS; Wang et al., 2001). In addition to being balanced, the force fields do not appear to be overly sensitive to small changes in the force field in nanosecond-length simulation (Cheatham and Kollman, 1998). For example, with addition or removal of water, use of modified water that diffuses two times as fast as TIP3P, various low-salt environments, or other small changes, there does not appear to be a major systematic alteration in the observed structure. However, there are clear issues with the imposition of periodicity, as discussed in UNIT 7.9. Setting Up a Nucleic Acid System With Explicit Water And Counter-Ions For Molecular Dynamics or Monte Carlo Simulation A molecular dynamics (MD) or Monte Carlo (MC) simulation (see UNIT 7.8) is generally broken up into two sequential phases. The initial part of the simulation is the equilibration phase, and this is followed by the production or sampling phase. The equilibration phase includes the beginning parts of the simulation, often including both minimization and dynamics, performed in order to obtain a “stable” simulation, as measured by a varied series of parameters such as the root-mean-squared atomic deviation from the starting structure, the temperature, or the total molecular mechanical energy. Once the simulation is stable, the production phase is performed. The production phase is the part of the dynamics or sampling that is extensively analyzed. The omission of the equilibration
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phase from this detailed analysis is necessary to avoid bias. The precise definition of what the production and equilibration phases entail are somewhat ambiguous. For example, when one parameter, such as root-mean-squared deviation (RMSd) of the solute to its starting structure or the temperature, has stabilized around an equilibrium value, this does not imply that all variables have equilibrated. Whether a given observable property has fully equilibrated depends on how long that observable takes to relax from its initial value to its equilibrium value. Many observable properties, such as the distribution of ions or sampling of thermally accessible conformational substates, take considerable amounts of time to equilibrate, times often longer than those that current state-of-the-art simulations can achieve. Therefore, when discussing equilibration, it should be referenced to a particular observable property (e.g., temperature, pressure, volume, potential energy). Equilibration is necessary in molecular dynamics simulations to relax structural distortions and remove large forces that may bias the dynamics. This means that equilibration is necessary to thermalize the system to put a comparable amount of kinetic energy into each degree of freedom. When this is not done, large forces may result at the distortions that, in turn, lead to large collisions on the local scale and create local “hot spots”. These hot spots may move the structure in unrealistic ways. Therefore, the goal of the equilibration procedure is to relax the system as much as possible to avoid biasing it away from the starting geometry. This is generally done through a series of minimization and molecular dynamics simulations where the temperature or kinetic energy is gradually increased. Setting up an initial in vacuo model, with or without inclusion of an implicit solvent model, is straightforward. Models can be generated in a variety of ways, usually based on known experimental structure. Issues related to model building are discussed in greater detail in UNIT 7.5. In molecular dynamics simulation of a model in vacuo, limited equilibration is necessary. As long as the structure is not significantly distorted, small distortions in the structure can be relaxed by short minimizations (∼100 to 1000 steps). Generally, a simple first-order method, such as steepest descent, is applied first to remove the largest forces, followed by a faster directed minimization method, such as conjugate gradient minimization. Minimization is performed until the change in energy or gradient between each minimization step is small (~0.1 to 0.0001 kcal/mol). Careful thermalization of the system, via a series of short molecular dynamics simulations where the temperature is gradually raised, is usually not necessary for in vacuo simulations since the system equilibrates rapidly (due to significantly fewer degrees of freedom than corresponding simulations with explicit solvent). If explicit ions are included in the in vacuo simulation, more careful equilibration may be necessary to relax the ion atmosphere.
Molecular Modeling of Nucleic Acid Structure: Setup and Analysis
Adding explicit solvent significantly increases the complexity and computational cost, and also necessitates a more stringent equilibration protocol (see Basic Protocol). To perform a simulation with explicit solvent using an appropriate nucleic acid model starting structure (in vacuo), an initial configuration of the solvated model is necessary. Moreover, it is typically desirable to include at least enough explicit salt to neutralize the system and any excess salt as desired. Adding solvent is typically performed by first completely surrounding the desired system by a set of pre-equilibrated solvent “boxes” representing the coordinates of a unit cell of “bulk” water, and then deleting those waters that overlap the model or extend beyond the boundary of the system under consideration. The only problem with this approach is knowing how to place solvent inside interior cavities or at interfaces, such as a protein-DNA interface. Unfortunately, there is currently no clear consensus on how best to do this in the absence of experimental structural data. However, in the authors’ experience, the water structure relaxes rapidly and is able to diffuse efficiently even into tight interfaces where proteins bind DNA within short (100-psec)
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simulations. In the absence of a pre-equilibrated solvent box, it is possible to simply add a crystalline representation of the solvent equally spaced at approximately the correct density; in this case, longer equilibration of the solvent may be necessary to fully relax the system and remove the crystalline bias. As discussed in UNIT 7.9, enough water should ideally be added to not only fully hydrate the model but to represent bulk water some distance away from the model. However, adding water tremendously increases the cost of the simulation. Typically, when periodic boundary simulations are applied, ∼8 to 12 Å of water surrounding the model in each direction is appropriate, although more adventurous souls could use more or less water. Given the issues and potential for artifacts with nonperiodic systems, particularly for highly charged systems such as nucleic acids, nonperiodic boundary conditions are not recommended for nucleic acid simulations except under specific conditions. These include the desire to represent a minimally hydrated nucleic acid in vacuo or a very large nucleic acid where only a core part of the structure (surrounded by a blob of explicit water with stochastic boundary conditions) is of interest, as applied in various protein simulations (Brooks et al., 1985; Steinbach and Brooks, 1993) and has been applied in some simulations of nucleic acids (Norberg and Nilsson, 1996b; Mazur, 1998). Nonperiodic boundary conditions may also be appropriate for very large and irregularly shaped models where the regular shapes of periodic boundary conditions may require too much water or could, in principle, inhibit motion. However, in these cases, implicit solvent models are more likely appropriate. In the opinion of the authors, it is wise to avoid methods that include explicit solvent but also dampen the electrostatic interactions through the application of a distance dielectric function, since these methods tend to significantly dampen the conformational fluctuations. For standard nucleic acid models representing a folded structure or 10 to 25 bp of a linear duplex, periodic boundary conditions are more appropriate. In principle, there is no reason to limit the shape of the periodic unit cell to a cubic shape, as any uniform space-filling shape is appropriate. Possible unit cell types are shown in Table 7.10.1. For long linear duplexes, the most appropriate unit cell might be the orthorhombic or hexagonal unit cells with one of the dimensions longer than the other two. This allows reasonable solvation of the duplex without large uninteresting regions that contain only solvent. However, an important consideration is that the rotational correlation time of small duplexes in solution is in the nanosecond time range. Therefore, during the dynamics in this type of orthorhombic box, the model may rotate to span the short edge of the box where the model can then interact directly with its periodic image (assuming a truly periodic method). This can also happen with constant pressure simulations where each box length is free to change. As shown in Figure 7.10.1, this can, in principle, lead to distortion of the model structure (as observed in long simulations of B-RNA in solution; Cheatham, unpub. observ.). This rotation can, in principle, be removed (at the expense of adding an uncorrected net torque to the system, which can lead to artifacts) or be inhibited by restraining the top and bottom of the duplex with weak restraints (which may inhibit bending). An additional issue is that interactions with periodic images in these long narrow boxes could, in principle, inhibit bending. In practice, inhibition of bending does not seem to be a major issue. Young and Beveridge (1998) reproduced expected sequence- and salt-specific bending in various phased A-tract models based on simulations of 24-mers in long rectangular boxes. To avoid the possible bias from inhibited rotation, more voluminous cubic boxes can be applied. This, however, leads to much more water in the corners than is necessary. Therefore, more modelers have shifted towards using more “spherical” unit cells, such as the 14-sided truncated octahedron (Allen and Tildesley, 1987) and 12-sided rhombic dodecahedron (see Figure 7.10.2). These unit cells limit the volume while maintaining distance between periodic images. Of course, adding the
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Table 7.10.1 Standard Unit Cells Appropriate for Periodic Boundary Conditionsa
Restrictions on unit cell parameters, Volume Cubic, a = b = c, α = β = γ = 90.0°,V = a3 Tetragonal, a = b, α = β = γ = 90.0°, V = ca2 Orthorhomic, α = β = γ = 90.0°, V = abc Monoclinic, α = γ = 90°, V =abc × sin(β) Triclinic, No restrictions, see legend Hexagonal, a = b Rhombohedral (trigonal), a = b = c, α = β = γ < 120.0°, V = a3 × [1 – cos(α)] × [1 + 2cos(α)]1/2 Octahedral (truncated octahedral), a = b = c, α = β = γ = 109.47122063449, V = (4(3)1/2/9)a3 Rhombic dodecahedral, a = b = c, α = γ = 60°, β = 90.0°, V =(1/2)1/2a3 a Restrictions on the unit cells lengths (a, b, c) and angles (α, β, γ) are presented along with the volumes for a variety of simulation cells. The volume of a triclinic cell is V = abc × [1 – cos(α)2 – cos(β)2 – cos(γ)2 + 2cos(α)cos(β)cos(γ)]1/2.
solvent is a little more tricky in nonorthorhombic unit cells since, when overlaying a larger solvent box, it is not a trivial procedure to remove waters outside the cell by simply checking if the water has coordinates larger than the box in a given dimension. In practice, a simple solution is to keep a set of the original coordinates with solvent, perform periodic imaging with the new unit cell type, and then, by comparison with the saved coordinates, delete any waters that have moved. Although, in principle, adding salt is as easy as adding solvent, it is slightly more complicated in practice. In an ideal case, random waters might be replaced by explicit salt ions up to the desired concentration and then molecular dynamics or Monte Carlo simulation can be performed to equilibrate the salt. However, unlike water, which is at a relatively high concentration and equilibrates rather rapidly, specific association of ions to the nucleic acid and relaxation of the ion distribution, in principle, may take a significant amount of time. Therefore, this approach may be impractical. This is particularly true for
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Figure 7.10.1 Orthorhombic unit cells and duplex rotation or unit cell size changes.
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Figure 7.10.2 A model B-DNA/Z-DNA junction in a solvated rhombic dodecahedron unit cell after ∼2 nsec of molecular dynamics with particle mesh Ewald (PME) in CHARMM.
the closely associated net-neutralizing counter-ions and multivalent ions that tend to have long water-exchange lifetimes and less rapid diffusion. Given the less rapid equilibration of multivalent ions, it is wise to make sure that each of these ions is sufficiently hydrated. This is wise because any direct binding of a multivalent ion may not exchange with water during nanosecond-length simulations. In the absence of any explicit information suggesting direct binding of an ion to the nucleic acid, placing ions that are directly bound (without bridging water) should be avoided, otherwise, the structure may distort under the influence of the bound ion. Placing ions within a hydration shell is reasonable since, in many cases, the interaction of a cation with a nucleic acid involves bridging water, such as with magnesium (Buckin et al., 1994) or barium (Sternglanz et al., 1976). In spite of this, specific interaction of ions with nucleic acids are seen. For example, ions have been observed in the minor groove of A-tract B-DNA (Hud and Feigon, 1997; Shui et al., 1998), interacting with the bases in the major groove of A-form structures (Robinson and Wang, 1996), or involved in phosphate interactions. Direct interaction is also seen with divalent ions in RNA that are known to stabilize the tertiary structure. Before solvating the system, it is advisable to place the net-neutralizing counter-ions. If available, structural information regarding the placement of ions can be used as an initial guide. However, more often than not, this information is not available. The net-neutralizing counter-ions are added to balance the charge on the phosphates. Sodium ions are typically the ion of choice because of their fairly rapid diffusion, small size and atomic number, and relatively rapid exchange times. Precise placement is not a major issue. Reasonable molecular mechanical potentials exist for treating these ions, and a variety of parameterizations are in common use (Straatsma and Berendsen, 1988; Aqvist, 1990; Smith and Dang, 1994). In the absence of structural information, a simple procedure commonly used is to place the ions some distance (∼5 Å) from the phosphate along the bisector of the phosphate oxygens. This works reasonably well for B-DNA structures since the phosphates are regularly spaced and the vector from the phosphate to the bisector points away from the surface of duplex DNA. For A-form structures or folded structures, the phosphates may be closer together and the vector from the phosphate to the bisector may point to interior regions or other phosphates, which can lead to overlapping ions. These initial guesses can be easily relaxed for more favorable positions that avoid the overlap by using a quick in vacuo minimization with the nucleic acid held fixed. However, during the in vacuo minimization, it is important to avoid the direct approach of the ions to the nucleic acid, which will happen because water has not been included at this stage and the ion parameters for use in explicit water have a small van der Waals radius and large charge. This can be accomplished by alterring the ion parameters to represent the ion as being effectively hydrated (Singh et al., 1985). This can be done by increasing the
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van der Waals radius (rmin ≈ 5 Å) and decreasing the well depth (ε ≈ 0.1 kcal/mol). For simulation with sodium ions (Na+), given their reasonable diffusion, this procedure leads to reasonable interaction with the nucleic acid. With folded structures, care should be taken to make sure an ion is not buried where it should not be. A more elaborate and perhaps better means to place the ions might be to use the electrostatic potential as a guide, such as that calculated from Poisson-Boltzmann treatments. For example, the program GRASP (http://www.honiglab.cpmc.columbia.edu/ grasp) can display the electrostatic potential and ions can be placed in favorable positions. Alternatively, the individual grid elements can be investigated and ions placed based on grid elements with low energies. A simple method along these lines is to place the ions, based just on the electrostatic potential, on a grid representing the Coulombic potential of the nucleic acid interacting with an ion charge (qion) at each grid point (r) atoms
Egrid = ∑ j
qj qion
ε rj − r
where Egrid is the interaction energy of the ion with the grid element, qj is the charge on each atom, rj is the position of each atom, and ε is the dielectric constant. This is typically done assuming uniform screening (ε = 1), although an effective dielectric could also be used. For placing ions around a specific group, such as a phosphate, a small grid is typically created. To avoid van der Waals overlap, the van der Waals energies can also be calculated for each grid element. The ion is then placed at the grid element with the lowest energy. To place all the ions, a larger and coarser grid can be built over the whole model (deleting grid elements within the solvent-accessible surface area or including van der Waals energy), and then the ions can be iteratively placed at low-energy grid points based on the energy, reevaluating the energy at each grid element after the placement of each ion. Alternatively, for placing positive ions in a previously solvated simulation, the DNA to water oxygen interaction energy can be used as a guide, with low interaction energies representing favorable positions in which the modeler can swap ions. The standard MD codes include procedures that allow placement of ions based on the phosphate bisector or other empirical rules, and some of these programs also have facilities for placing ions based on an energy grid. After addition of these ions, the system can be explicitly solvated. After this is done, it is a good idea to check the model structure to make sure no buried water appears where it is not appropriate and that the ions are explicitly and completely hydrated (unless directly bridging).
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After placing the net-neutralizing ions, any excess salt can be placed after solvating the system by replacing random waters some distance away from the nucleic acid and other ions. The key question is how much excess salt should be added. Adding excess cation also means that balancing anions should be added. Since the system is being modeled at an atomic level with unit cell lengths of ∼25 to 100 Å, very small changes in the unit cell size or the number of counter-ions have a large influence on the effective molarity. For a given ion, the molarity as reported in simulation literature is based on either the total amount of a specific ion present in the simulation (not just excess salt) or the total number of excess ions per the total volume (converted to moles/liter). Since the system is not a bulk macroscopic system, this concentration may be much higher than expected under periodic boundary conditions. One can also refer to the ionic strength of the system; this includes all the ionized groups (including phosphates). Given the small size of the unit cells and relatively large number of phosphates to added salt, these simulations are most often performed at high ionic strength. Despite the sensitivity of molarity to unit cell size, when looking at monovalent ions, there is very little salt dependence on dynamics or structure over the range of no salt (including no net-neutralizing salt) to ∼1 M salt in
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1-nsec-length simulations with the Cornell et al. force field (Cheatham and Kollman, 1998). It is not until high salt concentrations (>3 to 4 M) are reached that transitions in DNA duplex structure are seen; these transitions have been observed in simulations with the BMS nucleic acid force field (Langley, 1998). Divalent and multivalent ions, on the other hand, have much more direct influence on structure. For example, only four Co(NH3)63+ ions are necessary to observe B-DNA to A-DNA transitions with the Cornell et al. force field (Cheatham and Kollman, 1997a). Magnesium may also effect bending (Young and Beveridge, 1998). Equilibrating Simulations With Explicit Solvent After generating initial ion and solvent positions, it is necessary to equilibrate the system. This relaxes the system to the expected density and allows the water and ions to react to the presence of the nucleic acid. Minimization to remove unrealistic energies is an essential first step to this process; however, it is not sufficient. Given the multiple minima problem (UNIT 7.8) and, moreover, the fact that minimized water or “ice” is not what is really desired, it is necessary to sample possible configurations via molecular dynamics or Monte Carlo simulation. Since the solvent was most likely placed suboptimally, there may be holes or gaps in the solvation. Furthermore, the “pre-equilibrated” water will not have reacted to the presence of the nucleic acid. Thus, the system will most likely not be at the correct density. To remedy this, constant pressure equilibration under periodic boundary conditions is likely necessary. The initial solvent (and ion) equilibration is the most important part of any equilibration protocol prior to production MD. Given appropriate simulation methodologies, if the solvent and ionic atmosphere is well equilibrated, the simulation will likely be stable. In this case, the precise and intricate details of the remainder of the equilibration protocol are likely to be unimportant. This has been shown in molecular dynamics simulations of a DNA duplex where, after equilibration, there was little observable effect of varied ion placement when comparing three different mechanisms for placing sodium counter-ions (Young et al., 1997a). It should be noted that “equilibration” in this context refers to the generation of a more reasonable solvent structure and initial configuration that (1) does not contain local hot spots with unreasonable forces, (2) is at the correct pressure and density, and (3) has a reasonably stable potential energy. This equilibration does not refer to complete equilibration of the nucleic acid model, a process that may take significantly longer. A standard procedure (see Basic Protocol) is to first perform minimization to remove any large energies (which will lead to initially large forces), and then perform ∼25 to 100 psec of dynamics of constant pressure, with the nucleic acid held fixed or restrained to the initial model structure to relax the water and ion environment. The progress of the equilibration procedure is typically monitored by plotting the potential energy, density, and pressure. Equilibration is thought to be complete when these (and other) values have stabilized (see Figure 7.10.3). After this phase, minimization is performed on the entire system with the restraints on the initial nucleic acid model structure gradually reduced. Then, dynamics are performed on the entire system, slowly raising the temperature. Various modelers use different protocols (for examples, see Cheatham and Kollman, 1997b; MacKerell, 1997; Norberto de Souza and Ornstein, 1997; Young et al., 1997b), although, in practice, all these protocols seem to work consistently well. For more complicated systems, such as those involving high concentrations of salt or mixed solvent (such as ethanol and water), longer equilibration protocols are necessary. Note that this type of equilibration protocol tends to support water and ion conformations that stabilize the initial model structure. This can inhibit conformational transitions to other structures, such as B-DNA to A-DNA transitions in high salt, since the initial configurations are optimized to the “fixed” initial structure (Langley, 1998). A final note is that constant
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pressure simulation methodologies are plagued with pitfalls. If the forces restraining or fixing the initial conformation of the nucleic acid are not properly included with the calculated pressure, the pressure may be overestimated, leading to box expansion upon pressure scaling in longer equilibration simulations. For a more detailed discussion of this and other issues, see Cheatham and Brooks (1998). BASIC PROTOCOL
EQUILIBRATION This protocol describes the constraint/restraint of solute, relaxation of restraints, and equilibration (without restraints). Minimization is performed for ∼50 to 10,000 steps, although less minimization (∼200 to 500 steps) may be acceptable. Initial equilibration in molecular dynamics simulation takes ∼10 to 100 psec. It is important to avoid large force constants when applying harmonic restraints in molecular dynamics, since these may lead to high frequencies and require shorter time steps for proper equilibration. Force constants in the range of 5.0 to 15.0 kcal/mol/Å2 are reasonable. This protocol is intended to serve only as a guide. The primary literature, program manuals, and available resources on the Internet should be consulted for more information. For AMBER, see http://www.amber.ucsf.edu/amber/tutorial. Constrain or restrain solute (optional for in-vacuo simulation) Minimization: 1a. If necessary, turn off SHAKE constraints (that fix bond lengths involving hydrogen). The need for this step depends on the minimizer, force field, and SHAKE algorithm used.
2a. Perform steepest descent minimization until energy change at each step is less than ∼1.0 kcal/mol. 3a. Continue with conjugate gradient until energy change is less than ∼0.1 kcal/mol. 4a. If initial dynamics “blow up” (e.g., through SHAKE failures or sudden large increases in energy), perform additional minimization. Molecular dynamics: 1b. Apply an integration time step of 1 to 2 fsec for most available molecular mechanics force fields. 2b. Apply SHAKE on hydrogen atoms if using a rigid three-point water model. 3b. Apply constant pressure if using periodic boundaries in explicit solvent (UNIT 7.9). 4b. Maintain desired kinetic energy or slowly raise (ramp up) the kinetic energy to desired values. Perform molecular dynamics simulation for ~10 to 100 psec. Longer equilibration times are likely necessary with high salt conditions, multivalent ions, mixed solvents, or slowly diffusing solvents.
Relax restraints (optional) 5. Perform minimization only (∼50 to 5000 steps) with restraint force constants gradually moved to zero, or perform cycles of minimization (∼50 to 5000 steps) and dynamics (∼1 to 50 psec) with restraint force constants gradually moved to zero (i.e., 15 to 10 to 5 to 0.0 kcal/mol/Å2). Molecular Modeling of Nucleic Acid Structure: Setup and Analysis
Peform equilibration without restraints 6. Perform minimization or molecular dynamics as described above.
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a. Make certain that the overall rotational and translational kinetic energy is removed after initial velocity assignment and at periodic intervals as necessary (Harvey et al., 1998). b. If molecular dynamics blow up due to SHAKE failure or large energy change, try more minimization. If dynamics continue to fail, try decreasing the integration time step. If it continues to fail, look for a strong overlap of atoms and/or improper treatment of the electrostatic interactions. c. Monitor potential energy and the root-mean-squared deviation (RMSd) from starting structure, temperature, pressure, density, and volume in order to judge progress of equilibration. Begin production dynamics when these and other interesting observables appear to stabilize. At this point it is possible to change from an NPT (constant pressure, constant temperature) ensemble to an NVE (constant volume, constant energy) ensemble.
ANALYZING THE RESULTS After equilibration, production simulations are run for as long as computationally feasible or necessary. Current state-of-the-art simulations of small solvated biomolecules (representing on the order of ∼10,000 to 25,000 atoms) are performed for on the order of 1 to 25 nsec. At regular intervals, the configuration of the system (including the values of various energy terms and the atomic coordinates) should be saved and recorded to file(s). There are a variety of means to analyze the results. Analysis is performed not only for the purpose of extracting useful information about the structure but also to check the simulations for any aberrant behavior. In general, in order to obtain meaningful statistics when monitoring a particular observable, the simulation should be run on a time scale that is at least an order of magnitude longer than the correlation time of that particular observable. Some properties relax very quickly, such as various equilibrium properties of
Figure 7.10.3 Graphs monitoring the equilibration of a simulation of a DNA (polyA-polyT 10-mer) in explicit water. The first part of the simulation (25 psec) was performed with a cutoff. The second part (25 to 50 psec) was performed applying the particle mesh Ewald method. The DNA was held fixed and only the ions and water were allowed to move.
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water, i.e., density and average potential energy, which converge in short simulations (10 to 100 psec). Other properties, such as diffusion of salt, structural relaxation, or folding, may occur on a very long time scale. As discussed earlier, not all properties of a given system may fully equilibrate within the time scale of the simulation. Figure 7.10.3 shows a representative graph of various properties during a molecular dynamics simulation. From the graphs shown, it is clear that the simulation properties monitored fully converged during these short simulations. Common properties to monitor include the RMSd from the starting and average structures (created by performing a straight coordinate average of RMSd fit configurations over a stable portion of the trajectory) and helicoidal parameters (UNIT 7.5), among others. In addition to investigating the time series, various correlation functions are also appropriate to extract information that can be compared more directly to experimental results. Although the production dynamics typically take the most time to perform, the major effort of any modeling project is often spent in analyzing the results. In large part, the type of analysis methods applied depends on what one is trying to learn from the simulation and what experimental data is available for comparison. There is not a specific protocol that can be summarized here. For more information and to get a better handle on various analysis tools, the primary literature should be consulted. A good source of information on modeling and analysis is provided by Leach (2001). Additional information is available with each of the simulation programs and on their respective Web sites. An excellent discussion of the major issues in simulation and analysis is provided by Mark and van Gunsteren (van Gunsteren, 1992; van Gunsteren and Mark, 1998). Although it is beyond the scope of this unit to discuss all the means for analyzing molecular dynamic trajectories in detail, an important tool worthy of discussion is the means to judge the importance of sampled conformations from a molecular dynamics or Monte Carlo simulation. INEXPENSIVE METHODS TO ESTIMATE CRUDE RELATIVE FREE ENERGY DIFFERENCES Given two different conformational states of the same molecule (using the same force field) sampled in Monte Carlo or molecular dynamics simulation, an estimate of the relative free energy can be obtained either by characterizing the set of configurations that represents each sampled state or by characterizing the minimum energy conformation that best resembles each sampled state (Kollman et al., 2000). This characterization involves estimating the relative free energy. In this context, the free energy is the sum of the enthalpy and a temperature-weighted entropy term. Determining the energy or enthalpy for a given state is relatively straightforward; it comes directly from the molecular mechanics energy function, either as an average over the configurations or as the minimum energy of a representative conformation for each state. As discussed in UNIT 7.8, it is not directly possible to compare molecular mechanical energies among different molecules (due to different zero-point energies) or with different force fields (due to possible different scales and different zero-point energies). Therefore, in this unit, the reference to relative energy and free energy differences are for the same molecule. For different molecules, other techniques may be more appropriate, such as free energy perturbation (discussed briefly at the end of this section). Typically, the solvent is not included explicitly (as discussed in more detail below) but can be represented implicitly. Molecular Modeling of Nucleic Acid Structure: Setup and Analysis
In contrast to enthalpy, the entropy is less straightforward to estimate because it is an ensemble property. Although it can be calculated directly (at considerable cost), it is most often approximated and calculated independently for the solute and solvent. There are two basic methods for approximating the entropy; both comprise translational, rotational,
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and vibrational components. The translational and rotational components are calculated for a rigid rotor approximation or by some other means. The vibrational component can be estimated via two methods. The first involves the use of a “representative” minimum energy conformation. For this conformation, the normal modes of vibration are calculated using a harmonic approximation. These normal mode frequencies can then be used to estimate the vibrational components of the entropy based on the local fluctuations in the neighborhood of the minimum energy conformation. This can give crude estimates, assuming that (1) the conformation is truly at the energy minimum, (2) the single minimum-energy conformation represents the state of interest, and (3) the anharmonic effects are small. However, the entropy typically involves more than local fluctuations within a given potential energy well for macromolecules, such as the entropy from larger scale conformational rearrangements. If the state of interest is characterized by a number of substates, which is likely the case (Poncin et al., 1992), the approximation of a single representative state may break down. Therefore, a set of representative states may be more reliable. However, counting the number of effective states and estimation of the energy based on a complete enumeration of the partition function necessitates reasonable sampling that may, in practice, not be feasible. The alternative procedure to calculate the vibrational entropy uses a quasiharmonic approach with vibrational frequencies estimated from the fluctuations observed during molecular dynamics. This allows estimation based only on the relatively important fluctuations in the representative set of states (Karplus and Kushick, 1981). Given the sampling difficulties, entropic effects are difficult to estimate and lead to the calculation of “crude” relative free energies. Note that these approaches are only valid (in practice) for estimation of the entropy of the solute. In spite of the difficulties in estimating entropy, various groups have started to use data from a series of molecular dynamics simulations to estimate relative free energy differences. As a sample case study, consider the simulation of a canonical A-form RNA model and a canonical B-form RNA model, both of which are stable in multi-nanosecond-length state-of-the-art simulation including explicit solvent (Cheatham and Kollman, 1997b). Since both simulations are stable and no spontaneous conformational transition is seen, it is impossible to determine based on the MD results which conformation, B-RNA (a molecule that has never been experimentally observed) or A-RNA, is preferred. This is an important question since it would be valuable to better characterize the force field and to understand whether B-RNA is overstabilized by the Cornell et al. force field or is actually less favored than A-RNA. If A-RNA is indeed the energetically favored molecule, this implies that B-RNA to A-RNA transitions are not observed on a nanosecond time scale due to large conformational sampling barriers and insufficient sampling. In general, the ability to rank the various “models” can be used to judge the utility or importance of a given model. Since the same sequence and force field is used, it is possible to directly compare the molecular mechanical energies (although, note that the solventsolvent energies may have to be normalized if the two different simulations contain differing numbers of waters). The easiest way to estimate the relative free energy is to break up the total into contributions from the solvent (typically done implicitly) and the solute from the MD or MC simulation. Given a series of representative configurations from the dynamics, it is possible to determine the average intrasolute energy (or enthalpy). As mentioned, the entropy of the solute is a little more problematic to estimate, although the procedures specified above can be used. In general, a clear consensus on how best to estimate the entropic component has not emerged. One might consider making the assumption that the differences in entropy are largely represented in the solvation terms and not due to differences in the configurational entropy of the solute; however, this is likely invalid in most cases. With the A/B-RNA case study, in principle and as a first approximation, an estimation of the entropy can be obtained using the vibrational partition
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function and a harmonic approximation to the normal modes to estimate the frequencies in representative minimum-energy conformations. A problem with this approach is that in vacuo minimization of nucleic acids (without the water) leads to distortion of the structure away from the structure represented in solution; therefore, the calculated entropy may not accurately represent the entropy as estimated from the various snapshots in the respective trajectories. It will, however, reasonably estimate the entropy of the gas phase model structure. For the model average structures of [CCAACGTTGG]2 A-RNA and B-RNA (averaged over nanosecond portions of the respective trajectories taken at 1-psec intervals), minimization moved the models 1.9 Å and 2.4 Å, respectively, from the average structure. Estimation of the entropy difference using the normal modes favors A-RNA by ∼3.0 kcal/mol (at 300 K); although this is not a large difference, it is significant (Cheatham, 1997). The free energy of solvation can be estimated more directly. Under the assumption of linear response, a simple approximation to the solvation free energy in explicitly solvated simulations (also assuming that the bulk of this energy is represented by “close” waters, so that normalization for the total number of waters is not necessary) equates this free energy with half the solute-solvent interaction energy, 1⁄2 Esolute-solvent. A better estimate might be obtained by stripping the explicit water from each configuration and then performing a quick calculation on this conformation with an implicit solvent treatment. This will give an estimate of the solvation free energy. Recall that the implicit water models are typically parameterized to reproduce the free energy of solvation directly with a polarization component from Poisson-Boltzmann or a generalized Born treatment of the electrostatics and nonpolar contributions from a surface area term. This type of treatment has been applied to investigate a small turn-forming peptide based on long solvated MD trajectories (Bashford et al., 1997), and has recently been applied by various groups not only on the A/B-RNA case study but also the A/B-DNA equilibrium under various conditions, as well as a variety of other applications (Kollman et al., 2000). These techniques are very useful tools for post-processing MD or MC trajectories to give further insight. For more accurate configurational free energy differences, it is necessary to explicitly sample the configuration of accessible conformations connecting the end points, or states of interest, in a single simulation or series of simulations. This is typically done by adding biasing potentials (umbrella sampling) to force sampling along a particular path (Valleau and Whittington, 1977). Typically, multiple simulations are applied with different biasing potentials along a given path or reaction coordinate, and the results from the various simulations are accumulated and unbiased through a procedure such as the weighted histogram method (Kumar et al., 1992, 1995; Roux, 1995). This procedure has been used to characterize protein folding (Boczko and Brooks, 1995), but has been used infrequently for nucleic acid simulation. The use of these biasing potentials requires some understanding of the reaction path between the two states of interest, and therefore is not straightforward and is very computationally demanding. Calculating relative free energies of different molecules upon small chemical changes, free energy perturbation techniques, or thermodynamic integration techniques can be applied; for detailed reviews see Beveridge and DiCapua (1989) and Kollman (1993). Using data from an MD simulation can also give some qualitative insight (Radmer and Kollman, 1997). Molecular Modeling of Nucleic Acid Structure: Setup and Analysis
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SUMMARY The methods and tools for accurate simulation of small nucleic acids in solution have advanced considerably in recent years. A summary of the highlights are presented in recent reviews (Cheatham and Kollman, 2000; Kollman et al., 2000). In general, when simulating a polyionic system such as nucleic acids, it is necessary to not only provide a proper representation of the long-range electrostatic interactions through atom-based force-shifted cutoffs or an Ewald treatment, but also include some representation of the surrounding environment (i.e., water and salt). Tremendous strides have been made in recent years, including accurate representation of A-tract bending, specific ion association, and sequence-specific structure and dynamics. The current generation of force fields still retains some systematic errors and clearly more computer power is necessary to begin to tackle larger-scale problems and longer simulation times. However, the future holds tremendous promise. LITERATURE CITED Allen, M.P. and Tildesley, D.J. 1987. Computer Simulation of Liquids. Oxford University Press, Oxford. Aqvist, J. 1990. Ion-water interaction potentials derived from free energy perturbation simulations. J. Phys. Chem. 94:8021-8024. Bashford, D., Case, D.A., Choi, C., and Gippert, G.P. 1997. A computational study of the role of solvation effects in reverse turn formation in the tetrapeptides APGD and APGN. J. Amer. Chem. Soc. 119:4964-4971. Beveridge, D.L. and DiCapua, F.M. 1989. Free energy via molecular simulation. Annu. Rev. Biophys. Biophys. Chem. 18:431-492. Boczko, E.M. and Brooks, C.L. III. 1995. First-principles calculation of the folding free energy of a three-helix bundle protein. Science 269:393-396. Brooks, C.L. III, Brunger, A., and Karplus, M. 1985. Active site dynamics in protein molecules: A stochastic boundary-molecular dynamics approach. Biopolymers 24:843-865. Buckin, V.A., Kankiya, B.I., Rentzeperis, D., and Marky, L.A. 1994. Mg2+ recognizes the sequence of DNA through its hydration shell. J. Amer. Chem. Soc. 116:9423-9429. Cheatham, T.E. III. 1997. Ph.D. dissertation. Realistic simulation of nucleic acids in solution. University of California at San Francisco. Cheatham, T.E. III and Brooks, B.R. 1998. Recent advances in molecular dynamics simulation towards the reliable representation of biomolecules in solution. Theor. Chem. Acc. 99:279-288. Cheatham, T.E. III and Kollman, P.A. 1996. Observation of the A-DNA to B-DNA transition during unrestrained molecular dynamics in aqueous solution. J. Mol. Biol. 259:434-444. Cheatham, T.E. III and Kollman, P.A. 1997a. Insight into the stabilization of A-DNA by specific ion association: Spontaneous B-DNA to A-DNA transitions observed in molecular dynamics simulations of d[ACCCGCGGGT]2 in the presence of hexaammine cobalt(III). Structure 5:1297-1311. Cheatham, T.E. III and Kollman, P.A. 1997b. Molecular dynamics simulations highlight the structural differences in DNA:DNA, RNA:RNA and DNA:RNA hybrid duplexes. J. Amer. Chem. Soc. 119:4805-4825. Cheatham, T.E. III and Kollman, P.A. 1998. Molecular dynamics simulation of nucleic acids in solution: How sensitive are the results to small perturbations in the force field and environment. In Structure, Motion, Interactions and Expression of Biological Macromolecules (M. Sarma and R. Sarma, eds.) pp. 99-116. Adenine Press, Schenectady, N.Y. Cheatham, T.E. III and Kollman, P.A. 2000. Molecular dynamics simulation of nucleic acids. Annu. Rev. Phys. Chem. 51:435-471. Cheatham, T.E. III, Miller, J.L., Fox, T., Darden, T.A., and Kollman, P.A. 1995. Molecular dynamics simulations on solvated biomolecular systems–The particle mesh Ewald method leads to stable trajectories of DNA, RNA and proteins. J. Amer. Chem. Soc. 117:4193-4194. Cheatham, T.E. III, Crowley, M.F., Fox, T., and Kollman, P.A. 1997. A molecular level picture of the stabilization of A-DNA in mixed ethanol-water solutions. Proc. Natl. Acad. Sci. U.S.A. 94:9626-9630. Cheatham, T.E. III, Cieplak, P., and Kollman, P.A. 1999. A modified version of the Cornell et al. force field with improved sugar pucker phases and helical repeat. J. Biomol. Struct. Dyn. 16:845-862. Cieplak, P., Cheatham, T.E. III, and Kollman, P.A. 1997. Molecular dynamics simulations find that 3′ phosphoramidate modified DNA duplexes undergo a B to A transition and normal DNA duplexes an A to B transition. J. Amer. Chem. Soc. 119:6722-6730.
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Cornell, W.D., Cieplak, P., Bayly, C.I., Gould, I.R., Merz, K.M., Ferguson, D.M., Spellmeyer, D.C., Fox, T., Caldwell, J.W., and Kollman, P.A. 1995. A second generation force field for the simulation of proteins, nucleic acids, and organic molecules. J. Amer. Chem. Soc. 117:5179-5197. Feig, M. and Pettitt, B.M. 1997. Experiment vs force fields: DNA conformation from molecular dynamics simulations. J. Phys. Chem. 101:7361-7363. Flatters, D., Zakrzewska, K., and Lavery, R. 1997. Internal coordinate modeling of DNA: Force field comparisons. J. Comp. Chem. 18:1043-1055. Foloppe, N. and MacKerell, A.D. Jr. 2000. All-atom empirical force field for nucleic acids. (1) Parameter optimization based on small molecule and condensed phase macromolecular target data. J. Comp. Chem. 21:86-104. Harvey, S.C., Tan, R.K.-Z., and Cheatham, T.E. III. 1998. The flying ice cube: Velocity rescaling in molecular dynamics simulations leads to violation of equipartition. J. Comp. Chem. 19:726-740. Hud, N.V. and Feigon, J. 1997. Localization of divalent metal ions in the minor groove of DNA A-tracts. J. Amer. Chem. Soc. 119:5756-5757. Jorgensen, W.L., Maxwell, D.S., and Tirado-Rives, J. 1996. Development and testing of the OPLS all-atom force field on conformational energetics and properties of organic liquids. J. Amer. Chem. Soc. 118:11225-11236. Karplus, M. and Kushick, J.N. 1981. Method for estimating the configurational entropy of macromolecules. Macromolecules 14:325-332. Kollman, P.A. 1993. Free energy calculations–Applications to chemical and biochemical phenomena. Chem. Rev. 93:2395-2417. Kollman, P.A., Massova, I., Reyes, C., Kuhn, B., Huo, S., Chong, L., Lee, M., Lee, T., Duan, Y., Wang, W., Donini, O., Cieplak, P., Srinivasan, J., Case, D.A., and Cheatham, T.E. III. 2000. Calculating structures and free energies of complex molecules: Combining molecular mechanics and continuum models. Acc. Chem. Res. 33:889-897. Kumar, S., Bouzida, D., Swendsen, R.H., Kollman, P.A., and Rosenberg, J.M. 1992. The weighted histogram analysis method for free-energy calculations on biomolecules. 1. The method. J . Comp. Chem. 13:1011-1021. Kumar, S., Rosenberg, J.M., Bouzida, D., Swendsen, R.H., and Kollman, P.A. 1995. Multidimensional free-energy calculations using the weighted histogram analysis method. J. Comp. Chem. 16:1339-1350. Langley, D.R. 1998. Molecular dynamics simulations of environment and sequence dependent DNA conformation: The development of the BMS nucleic acid force field and comparison with experimental results. J. Biomol. Struct. Dyn. 16:487-509. Lavery, R., Zakrzewska, K., and Sklenar, H. 1995. JUMNA (junction minimisation of nucleic acids). Comp. Phys. Comm. 91:135-158. Leach, A.R. 2001. Molecular Modeling: Principles and Applications, 2nd ed. Pearson Education Limited, Essex, England. MacKerell, A.D. Jr. 1997. Influence of magnesium ions on duplex DNA structural, dynamic, and solvation properties. J. Phys. Chem. B101:646-650. MacKerell, A.D. Jr. 1998. Observations on the A versus B equilibrium in molecular dynamics simulations of duplex DNA and RNA. In Molecular Modeling of Nucleic Acids (N.B. Leontis and J. Santa Lucia, eds.) pp. 304-311. American Chemical Society, Washington, D.C. MacKerell, A.D. Jr. and Banavali, N. 2000. All-atom empirical force field for nucleic acids. (2) Application to molecular dynamics simulations of DNA and RNA in solution. J. Comp. Chem. 21:105-120. Mackerell, A.D. Jr., Wiorkiewicz-Kuczera, J., and Karplus, M. 1995. An all-atom empirical energy function for the simulation of nucleic acids. J. Amer. Chem. Soc. 117:11946-11975. Mazur, A.K. 1998. Accurate DNA dynamics without accurate long-range interactions. J. Amer. Chem. Soc. 120:10928-10937. Miaskiewicz, K., Miller, J., Cooney, M., and Osman, R. 1996. Computational simulations of DNA distortions by a cis,syn-cyclobutane thymine dimer lesion. J. Amer. Chem. Soc. 118:9156-9163. Norberg, J. and Nilsson, L. 1996a. Constant pressure molecular dynamics simulations of the dodecamers: d(GCGCGCGCGCGC)2 and r(GCGCGCGCGCGC)2. J. Chem. Phys. 104:6052-6057. Norberg, J. and Nilsson, L. 1996b. Glass transition in DNA from molecular dynamics simulations. Proc. Natl. Acad. Sci. U.S.A. 93:10173-10176.
Molecular Modeling of Nucleic Acid Structure: Setup and Analysis
Norberto de Souza, O. and Ornstein, R.L. 1997. Effect of warmup protocol and sampling time on convergence of molecular dynamics simulations of a DNA dodecamer using AMBER 4.1 and particle mesh Ewald method. J. Biomol. Struct. Dyn. 14:607-611. Poncin, M., Hartmann, B., and Lavery, R. 1992. Conformational sub-states in B-DNA. J. Mol. Biol. 226:775-794.
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Radmer, R. and Kollman, P.A. 1997. Free energy calculation methods: A theoretical and empirical comparison of numerical errors and a new method for qualitative estimates of free energy changes. J. Comp. Chem. 18:902-919. Robinson, H. and Wang, A.H.-J. 1996. Neomycin, spermine and hexaamminecobalt(III) share common structural motifs in converting B- to A-DNA. Nucl. Acids Res. 24:676-682. Roux, B. 1995. The calculation of the potential of mean force using computer simulations. Comp. Phys. Comm. 91:275-282. Shui, X., McFail-Isom, L., Hu, G.G., and Williams, L.D. 1998. The B-DNA dodecamer at high resolution reveals a spine of water on sodium. Biochemistry 37:8341-8355. Singh, U.C., Weiner, S.C., and Kollman, P.A. 1985. Molecular dynamics simulations of d(C-G-C-G-A)-d(TC-G-C-G) with and without “hydrated” counterions. Proc. Natl. Acad. Sci. U.S.A. 82:755-759. Smith, D.E. and Dang, L.X. 1994. Computer simulations of NaCl association in polarizable water. J. Chem. Phys. 100:3757-3766. Spector, T.I., Cheatham, T.E. III, and Kollman, P.A. 1997. Unrestrained molecular dynamics of photodamage DNA in aqueous solution. J. Amer. Chem. Soc. 119:7095-7104. Steinbach, P.J. and Brooks, B.R. 1993. Protein hydration elucidated by molecular dynamics simulation. Proc. Natl. Acad. Sci. U.S.A. 90:9135-9139. Sternglanz, H., Subramanian, E., Lacey, J.C.J., and Bugg, C.E. 1976. Interactions of hydrated metal ions with nucleotides: The crystal structure of barium adenosine 5′-monophosphate heptahydrate. Biochemistry 15:4797-4802. Straatsma, T.P. and Berendsen, H.J.C. 1988. Free energy of ionic hydration: Analysis of a thermodynamic integration technique to evaluate free energy differences by molecular dynamics simulations. J. Chem. Phys. 89:5876-5886. Tapia, O. and Velazquez, I. 1997. Molecular dynamics simulations of DNA with protein’s consistent GROMOS force field and the role of counterions’ symmetry. J. Amer. Chem. Soc. 119:5934-5938. Valleau, J.P. and Whittington, S.G. 1977. A guide to Monte Carlo for statistical mechanics. 1. Highways. In Statistical Mechanics A: A Modern Theoretical Chemistry, Vol. 5-6 (B.J. Berne, ed.). Plenum Press, New York. van Gunsteren, W.F. 1992. On the interpretation of biochemical data by molecular dynamics computer simulation. Eur. J. Biochem. 204:947-961. van Gunsteren, W.F. and Berendsen, H.J.C. 1987. Groningen molecular simulation (GROMOS) library manual. BIOMOS, Nijenborgh, Groningen, The Netherlands. van Gunsteren, W.F. and Mark, A.E. 1998. Validation of molecular dynamics simulation. J. Chem. Phys. 108:6109-6116. Wang, W., Donini, O., Reyes, C.M., and Kollman, P.A. 2001. Biomolecular simulations: Recent developments in force fields, simulations of enzyme catalysis, protein-ligand, protein-protein, and protein-nucleic acid noncovalent interactions. Annu. Rev. Biophys. Biomol. Struct. 30:211-243. Young, M.A. and Beveridge, D.L. 1998. Molecular dynamics simulations of an oligonucleotide duplex with adenine-tracts phased by a full helix turn. J. Mol. Biol. 281:675-687. Young, M.A., Jayaram, B., and Beveridge, D.L. 1997a. Intrusion of counterions into the spine of hydration in the minor groove of B-DNA: Fractional occupancy of electronegative pockets. J. Amer. Chem. Soc. 119:59-69. Young, M.A., Ravishanker, G., and Beveridge, D.L. 1997b. A 5-nanosecond molecular dynamics trajectory for B-DNA: Analysis of structure, motions and solvation. Biophys. J. 73:2313-2336. Zacharias, M. and Sklenar, H. 1997. Analysis of the stability of looped-out and stacked-in conformations of an adenine bulge in DNA using a continuum model for solvent and ions. Biophys. J. 73:2990-3003. Zhurkin, V.B., Poltev, V.I. and Florent’ev, V.L. 1980. Atom-atom potential functions for conformational calculations of nucleic acids. Mol. Biol. 14:1116-1130. Zhurkin, V.B., Ulyanov, N.B., Gorin, A.A., and Jernigan, R.L. 1991. Static and statistical bending of DNA evaluated by Monte Carlo calculations. Proc. Natl. Acad. Sci. U.S.A. 88:7046-7050.
Contributed by Thomas E. Cheatham, III University of Utah Salt Lake City, Utah Bernard R. Brooks National Institutes of Health Bethesda, Maryland Peter A. Kollman University of California San Francisco, California Current Protocols in Nucleic Acid Chemistry
Biophysical Analysis of Nucleic Acids
7.10.17 Supplement 6
Characterization of DNA Structures by Circular Dichroism Chiral substances are distinguishable by their interactions with polarized light at wavelengths near and distant from absorption energies (Lowry, 1964). Circular dichroism (CD) is the difference in absorbance by a substance of right- and left-handed circularly polarized light (Crabbe, 1965; Drake, 2001). CD is a powerful tool for the study of the secondary structures and conformations adopted by nucleic acids and proteins (Tinoco and Cantor, 1970; Ivanov et al., 1973; Greenfield, 1996; Venyaminov and Yang, 2000). In the case of nucleic acids, the CD of electronic transitions of the bases is typically monitored. CD measures the asymmetry in nucleic acid systems resulting from an induced CD in the symmetric bases. CD arises from the asymmetric backbone sugars and by the helical structures often adopted by nucleic acids. CD spectroscopy is extremely sensitive to nucleic acid conformation, especially within the 180 to 320 nm wavelength range (Ivanov et al., 1973; Steely et al., 1986; Sun et al., 1999; Porumb et al., 2002). The CD of nucleic acids can in principle be calculated using quantum mechanics, but such calculations are not presently reliable enough to be used in the quantitative interpretation of experimental spectra. As a result, CD of nucleic acids is commonly used in an empirical manner to provide a signature for a given secondary structure. CD is particularly powerful for monitoring structural changes resulting from changes in environmental conditions such as temperature, ionic strength, and pH. A number of reviews of CD studies of nucleic acids have appeared (Johnson, 1985, 2000a,b; Gray et al., 1995; Maurizot, 2000), including a detailed compilation of CD spectra of nucleic acids by Johnson (1990). A number of commercially available instruments are available for the measurement of CD spectra. Since the details of instrumental operation differ for each machine, this unit provides a descriptive guide, rather than a strict protocol. It describes considerations that the experimentalist will face when attempting to examine nucleic acid structures by CD spectropolarimetry. Specific details are also given for cell filling and cleaning, and for instrumental calibration. Finally, several examples of CD spectra are given to provide useful reference points for a variety of nucleic acid structures, ranging
UNIT 7.11
from single-stranded to triplex and quadruplex forms.
GUIDELINES FOR CARRYING OUT A CD EXPERIMENT Initial Considerations The measurement of CD spectra of a variety of nucleic acids is straightforward and is operationally similar to recording an absorbance spectrum. However, since CD spectra are actually the result of taking the difference of two large numbers, subtle variations in nucleic acid structure as induced by solution pH, temperature, ionic composition, and sample degradation can lead to significant variations in the resulting spectra, especially in the 180 to 260 nm wavelength range. Great care must go into the preparation of samples, including the elimination of contaminants by proper cleaning of sample cells. Instrumental calibration is another important consideration when collecting CD spectra of nucleic acids.
CD Instrumentation A number of reliable circular dichroism spectrometers are available. Manufacturers include Jasco (http://www.jascoinc.com), OnLine Instruments (http://www.olisweb.com), Aviv Instruments (http://www.avivinst.com), and Applied Photophysics (http://www.apltd. co.uk). Like all precision spectrometers, CD instruments should be well maintained if they are to function properly, and their calibration should be checked regularly (see Calibration with (+)-10-Camphorsulfonic Acid). Investigators should realize that CD spectrometers measure small differences in absorbance (∆A) between two large and similar numbers, typically with ∆A ≈ 0.001. During operation, CD spectropolarimeters should be continuously flushed with dry nitrogen gas, both to preserve the optics and to purge oxygen and ozone from the system in order to decrease background absorbance and permit measurements down to wavelengths <200 nm. Dry nitrogen (N2) gas is prepared by passing ultrapure N2 through two gas-tight canisters containing anhydrous calcium sulfate (available as Drierite from Sigma).
Contributed by G. Reid Bishop and Jonathan B. Chaires Current Protocols in Nucleic Acid Chemistry (2002) 7.11.1-7.11.8 Copyright © 2002 by John Wiley & Sons, Inc.
Biophysical Analysis of Nucleic Acids
7.11.1 Supplement 11
Measuring Circular Dichroism
Characterization of DNA Structures by Circular Dichroism
Excellent reviews of the basic theory and practice of measuring CD spectra can be found in Drake (2001) and Woody (1995). Molar circular dichroism is defined as the difference in extinction coefficients (∆ε) between left- and right-handed circularly polarized light, ∆ε = εL − εR. If the difference in absorbance between left- and right-handed circularly polarized light is ∆A = AL − AR, then ∆ε = ∆A/cl, where c is the molar concentration and l is the pathlength. While ∆ε is the preferred way to express circular dichroism, authors frequently report data as molar ellipticity (Θ). Ellipticity is related to ∆A by the equation ∆A = Θ/32.98. CD spectra are typically reported as ellipticity in degrees as a function of wavelength. Molar ellipticity and molar circular dichroism are related by the expression [Θ] = 3298∆ε (Woody, 1995). Since CD instruments utilize photomultiplier tube (PMT) detectors, they measure the intensity of transmitted light. It is important that the total absorbance (i.e., cell, sample, and solvent combined) be kept in the analytical range between 0.2 to 1.2 AU. An absorbance of 0.87 AU gives the optimal signal-to-noise ratio in a CD experiment. Investigators should keep in mind that a nucleic acid sample with an absorbance of 1.0 at 260 nm in a 10-mm-pathlength cell will have much higher absorbance at wavelengths shorter than 220 nm. In order to accurately record nucleic acid spectra at wavelengths below 220 nm, cells with shorter pathlengths (or more dilute solutions) must generally be used. Most CD instruments are single-beam spectrometers. Hence, sample spectra should be corrected by subtracting a baseline spectrum of the cell plus solvent obtained under identical acquisition conditions. It is essential to record such a baseline in each working session. Instrumental settings should be carefully chosen and maintained for each working session. There are several key parameters that must be controlled. (1) The spectral range of measurement should be chosen to include ∼20 nm before the onset of differential absorbance by the sample. This will establish, after baseline correction, a flat region where the CD signal is zero. For most nucleic acid samples, a spectral range of 350 to 200 nm is a good choice. It is highly desirable to go to wavelengths below 200 nm, if possible, since nucleic acid structures show distinctive features in that region (Johnson, 2000a). Remaining instrumental settings are adjusted to optimize the signal-tonoise ratio. (2) The spectrometer slit width
(spectral band width) may be adjusted to allow more light to pass through the sample, reducing noise, but potentially decreasing spectral resolution. Since solution-phase ultraviolet features are not particularly sharp, a spectral bandwidth of ≥1 nm can be used for CD of nucleic acid samples. Even larger spectral bandwidths may be used safely if required because of weak CD signals or low sample concentrations. (3) The scan speed and instrumental time constant should be selected with care. The choice of one of these parameters dictates the allowable range of the other. Most instrument manuals provide a table of acceptable scan speeds for a given time constant. The time constant is the signal averaging period and governs the amount of electrical damping of the signal. Longer time constants lower the noise. A longer time constant should be balanced by a slower scan rate, otherwise distortions of the CD spectrum may result. In order to obtain reliable data below 200 nm, longer time constants and very slow scan speeds are usually required. This is compounded by the observation that absorption features at shorter wavelengths tend to occur over a narrower bandwidth and are thus prone to distortion. (4) Most CD data acquisition software now permit signal averaging in which multiple spectra are recorded and then averaged. Signal averaging reduces noise at the cost of the time to record the multiple scans. Since noise reduction is proportional to the square root of the number of scans, four scans are required to reduce the noise by a factor of two compared to a single scan. (5) Finally, since most modern CD instruments collect digital data through their computerized acquisition system, it is necessary to select a value for the data resolution, that is, how frequently data are obtained and recorded in the spectrum. A data resolution of 0.2 nm per point is usually adequate for nucleic acid samples. The primary concerns in the measurement of nucleic acid CD spectra are optimizing the signal-to-noise ratio and accurately defining spectral maxima and minima. Large slit widths, long time constants, slow scan speeds, and sample absorbance near 0.8 AU generally yield quality spectra. Time and sample constraints may necessitate modifications to be decided by each investigator.
Cell Selection, Cleaning, and Filling A variety of spectrophotometric sample cells may be used to collect CD spectra of nucleic acids. The only unavoidable feature of the cells is that they must be fabricated from
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optical grade fused quartz. Quartz-Suprasil (typically designated by the abbreviation QS) has an optimal transmission range between 200 to 2500 nm and is adequate for the study of nucleic acids. Although QS cells are only rated down to 200 nm, they can be used down to 185 nm if increased noise in the experimental results can be tolerated. Cells fabricated with cemented joints should be avoided since the joints can weaken during rigorous cleaning. A water-jacketed cylindrical cell with a 10mm pathlength is the most versatile cell for CD studies of nucleic acids. If it is desirable to collect CD data at wavelengths <200 nm, it will probably be necessary to use a cell with a smaller pathlength (i.e., 0.1 to 1.0 mm). Nucleic acids exhibit higher absorbances (and molar ellipticities) in this region compared to the 200 to 300 nm range. Although cylindrical cells are typically used for CD work, there is really no reason why a standard QS-grade rectangular cuvette with a 10-mm pathlength cannot also be used at longer wavelengths (i.e., 220 to 350 nm). The only real limitations in cell selection are the design of the sample compartment of the spectrometer and the quality of the quartz from which the cell is made. Sample cells described above can be purchased from Hellma Cells (http://www.hellmausa.com). The 10-mm-pathlength cylindrical cell requires ∼750 to 1000 µL sample to fill it, whereas the 0.1-mm pathlength cell only requires ∼75 to 100 µL. Regardless of the pathlength chosen, the sample concentration should be adjusted so that the sample exhibits a UV absorbance in the analytical range (i.e., 0.2 to 1.2 AU). If lower concentrations are used, then signal averaging may be necessary for optimal results. Other issues related to cell selection may be found in the review by Tinoco and Cantor (1970). As with any optical spectroscopic technique, recording an accurate CD spectrum requires clean sample cells (Tinoco and Cantor, 1970) . This is especially important in the determination of the circular dichroism of nucleic acids and oligonucleotides, which routinely require observation over a broad UV range (e.g., 200 to 350 nm). Even slight amounts of dust and debris can scatter light and cause a decrease in the signal-to-noise ratio. Obstructions to the light path (such as smears, fingerprints, and exterior stains resulting from improper sample and cell handling and poor cleaning techniques) can lead to spectral artifacts, especially at wavelengths ≤300 nm. For a rigorous initial cleaning, prepare ∼10 to 50 mL of a 1:1 (v/v)
mixture of concentrated nitric and sulfuric acids in a fume hood. In the hood, wash the entire sample cell carefully by completely submerging it in the acid solution for variable times depending upon the degree of cleaning desired (protective eyewear and gloves should be worn). Excellent results are generally obtained by cleaning 1 to 5 min. In general, it is best to completely remove the acid cleaning solution from the vessel containing the cleaned cell with a serological pipet and to rinse several times with high-quality nanopure water. Detergent solutions, 10% by weight, can be used to clean cells that are already moderately clean. Soak the sample cells in detergent solutions for several hours. After removal of the detergents, the cells must be rinsed extensively with nanopure water to remove detergent residues. High pH detergents can etch the cells and render them useless. After thorough rinsing, excess water and residual organic materials can be removed from the cell by several rinses with polar solvents such as methanol, 95% ethanol, or neat acetone, followed by evaporation under a stream of dinitrogen or high-quality air. Solvents must be spectrophotometric grade to prevent the deposit of residues. After cleaning, it is critical to avoid touching the windows of the sample cells. It best to remove drops of solvent from the windows with a lint-free tissue such as a Kimwipe. If acetone is used to remove cleaning solvents, it must be completely removed by evaporation because of its strong UV absorption, which will cause interference in the acquisition of accurate and reproducible spectra. High-grade sample cells should not routinely be heated above 100oC to remove solvents as high temperatures can result in deformation of the quartz and loss of structural integrity of the cell. Cleaning solutions containing chromic acid should be avoided since they can leave behind chromium ions that may interact with nucleic acids and alter their solution structure and CD spectrum. To avoid contamination and dilution of the sample, it is best to start with a completely dry cell when changing samples. If it is undesirable to wait the time necessary to dry the sample cells after cleaning, it is acceptable to rinse the cell a few times with small aliquots of the sample under study if the sample is not too precious. When performing concentration-dependence studies, it is always best to work from the least to the most concentrated samples to avoid extensive and time-consuming cleaning peri-
Biophysical Analysis of Nucleic Acids
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ods between sample loading and spectral acquisitions. Residue from a concentrated sample remaining in the cell can lead to misleading measurements. The most efficient way to fill and empty the sample cell is with a disposable fine-tip polyethylene transfer pipet such as that manufactured by Corning-Samco and available from VWR International. Such pipets allow the sample to be completely removed from the cell and will not scratch the interior window surfaces of the sample cell. Biological samples will not adhere to polyethylene pipets, as may happen with unsilanized glass Pasteur pipets.
Calibration With (+)-10-Camphorsulfonic Acid
Characterization of DNA Structures by Circular Dichroism
Any time the CD spectrophotometer has undergone any maintenance such as lamp replacement, cleaning, movement or replacement of any of the mirrors in the optical bench, or movement of the instrument itself, it is necessary to check the instrument calibration with a standard sample. Loss of calibration of the instrument is typically due to misalignment or degradation of the light source, mirrors, PMT, and/or monochromator. A variety of standards can be selected to check the instrument calibration over the wavelength range between 190 to 800 nm. However, since this unit is limited to the CD of nucleic acids, which have no observable absorbance or CD above 320 nm, only calibration of the UV range will be considered. In all calibration determinations, set the instrument measurement parameters to a standard set of conditions as indicated by the manufacturer. The standard conditions recommended by Jasco for the J500 series of CD spectropolarimeters are: sensitivity = 20 m° cm−1 (millidegrees per centimeter), chart speed = 5 cm min−1, wavelength expansion = 10 nm cm−1, time constant = 0.5 sec, and scan rate = 10 nm min−1. The monochromator should be calibrated using the methods outlined in the Jasco J500 instrument manual. For this purpose, several standards must be employed to cover both the ultraviolet and visible wavelength ranges. Each of the following solutions must be prepared using the highest-quality solutes, nanopure water, Class A volumetric glassware, and precision balances. Prior to weighing, reagents should be oven-dried for 24 hr at ∼120°C to remove water. If possible, reagents should be dried under a vacuum. A 0.15 mg/mL solution of D-(−)-pantoyl lactone in water should have a CD maximum at 220 nm, a 0.6 mg/mL solu-
tion of (+)-10-camphorsulfonic acid (a.k.a. D10-camphorsulfonic acid or CSA) in pure water must have a CD minimum at 190.5 and a maximum at 290.5 nm, and a 0.5 mg/mL solution of androsterone dissolved in dioxane should have a CD maximum at 304 nm. Deviations in the observed wavelength from these values indicate that the monochromator should be adjusted. (Adjustment should be accomplished by appropriate technical personnel or by consulting the instrument manufacturer). The alignment and quality of the lamp, mirrors, and PMT between 190 to 320 nm can be checked using a sample of 0.1 mg/mL aqueous CSA and a 10-mm-pathlength cell (Gray et al., 1995). The concentration of CSA can be determined spectrophotometrically using its extinction coefficient (i.e., molar absorptivity) at 285 nm (ε285 = 34.5 M−1 cm−1; Venyaminov and Yang, 2000). The ∆ε at 290.5 nm should be 2.36 M−1 cm−1 and at 190.5 nm should be between −4.9 and −4.7 M−1 cm−1. Alternatively, the molar ellipticity (Θ) at 290.5 and 190.5 should be 7.8 and −15.6 deg cm2 dmol−1, respectively (−15.6/7.8 = −2.0). The ratio of the intensity of the CD bands at 192.5/290.5 nm should be between −2.0 and −2.1. (Deviations of ∼3% from measurements taken in the far UV range of 190 to 240 nm may be within the experimental error of the instrument). If the ratio drops as low as −1.9, then this is an indication that the xenon lamp is beginning to fail. If any of the spectral criteria listed above are not met during calibration, then the basic remedy requires realignment or possible replacement of the xenon lamp or mirrors. Each spectrometer is different and the details of handling these problems should be determined by consultation with the manufacturer. Since spectral quality also depends upon the scan rate, it is always best to adjust the scan rate before realigning the mirrors or replacing a lamp. Scan rates that are set too high relative to a given time constant can lead to spectral deformation (see Measuring Circular Dichroism).
SAMPLE CD SPECTRA Figures 7.11.1 to 7.11.3 show representative spectra for a variety of nucleic acid structures. These spectra were acquired using a 15-yearold Jasco J500A CD spectrometer, an instrument no longer supported by its manufacturer. These spectra show that even an older instrument with aged optics can produce quality spectra over the 220 to 320 nm wavelength range. All of the spectra shown were obtained using a spectral band width of 2 nm, a time
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Current Protocols in Nucleic Acid Chemistry
constant of 32 sec, and a scan rate of 2 nm min−1. Each spectrum required 50 min to acquire. A single scan was recorded with a time constant of 32 sec with no digital signal averaging and no numerical smoothing. Figure 7.11.1 shows the CD spectrum of E. coli DNA, and illustrates the features of standard right-handed duplex DNA. There is a CD maximum near 280 nm, a CD minimum near 250 nm, and a cross-over point (from positive to negative intensity) at 260 nm. Figure 7.11.1 shows the typical spectrum for standard B-form DNA. The exact positions and intensities of the peaks are sensitive to base composition and sequence, but this spectrum provides a reference signature for standard duplex DNA. Figure 7.11.2 shows CD spectra and signatures for a wide variety of nucleic acid structures, ranging from single strands to triplexes. For the spectra shown in Figure 7.11.2, synthetic polynucleotides containing only adenine and thymine (or uracil) were used. Figure 7.11.2A shows spectra for single-stranded poly(dA) and poly(dT), along with the dramatically different spectrum that results when these are co mb ined to form the duplex poly(dA):poly(dT). Compare the spectrum for poly(dA):poly(dT) (dotted line) with that of
standard duplex DNA (Fig. 7.11.1). The difference in the spectra reflects the fact that poly(dA):poly(dT) adopts a nonstandard righthanded conformation with a helical pitch of 10.0 bp/helix turn instead of the normal value of 10.4 bp/helix turn. Figure 7.11.2B compares the CD spectrum of duplex DNA with an RNA:DNA hybrid and a duplex RNA form. These are from the polynucleotides poly(dA):poly(dT), poly(rA):poly(dT), and poly(rA):poly(rU), and were recorded under the identical conditions presented above. The spectrum of the RNA is characteristic of the A-form helix (Steely et al., 1986), with a maximum near 270 nm and only a shallow negative peak. The spectrum of the RNA:DNA hybrid structure is intermediate between that of the pure DNA and RNA spectra, retaining the negative peak near 250 nm found in DNA, but showing distinctive differences from DNA in the region of 260 to 290 nm. Finally, Figure 7.12.2C compares duplex poly(dA):poly(dT) with triplex poly(dA):[poly(dT)]2. The triplex shows somewhat subtle but distinctive differences from the duplex structure. This system provides an opportunity to observe the distinctive changes in CD spectra over a range of polynucleotide structures. In those cases, the
1.5
Molar ellipticity × 10−3
1.0
0.5
0.0
−0.5
−1.0
−1.5
−2.0 220
240
260
280
300
320
Wavelength (nm)
Figure 7.11.1 Circular dichroism spectrum of E. coli DNA in 1 mM Na2EDTA, 185 mM NaCl, and 6 mM Na2HPO4/2 mM NaH2PO4, pH 7.0.
Biophysical Analysis of Nucleic Acids
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structures including duplex, triplex, quadruplex, and i-motif polynucleotide structures. A more comprehensive compilation of nucleic acid CD spectra may be found in articles by Johnson (1990, 2000a,b), which are key and essential references.
polynucleotide base sequences are identical and the spectral differences are due entirely to secondary structural differences. Figure 7.11.2 provides a useful set of reference spectra. Figure 7.11.3 shows spectra for two quadruplex structures. The oligonucleotide 5′AGGG(TTAGGG)3 spontaneously folds into an antiparallel quadruplex structure under these conditions (Sun et al., 1999). A distinctive CD spectrum, with a positive maximum near 295 nm and a negative minimum near 260 nm, results. The oligonucleotide 5′T2G20T2 spontaneously self-associates into an intermolecular four-stranded parallel structure. The parallelstranded quadruplex shows a distinctive CD spectrum with a positive maximum near 270 nm and only a shallow minimum (Porumb et al., 2002). Figures 7.12.1 to 7.12.3 clearly show the utility of CD as a tool for the empirical characterization of nucleic acid secondary structure. Table 7.11.1 provides a rough guide for the CD spectral characteristics of several secondary
LITERATURE CITED Crabbe, P. 1965. Optical Rotatory Dispersion and Circular Dichroism in Organic Chemistry. Holden-Day, San Francisco, Calif. Drake, A.F. 2001. Circular dichroism. In ProteinLigand Interactions: Structure and Spectroscopy (S.E. Harding and B.Z. Chowdhry, eds.) pp.123167. Oxford University Press, Oxford, U.K. Gray, D.M., Hung, S.W., and Johnson, K.H. 1995. Absorption and circular dichroism spectroscopy of nucleic acid duplexes and triplexes. Methods Enzymol. 246:19-71. Greenfield, N.J. 1996. Methods to estimate the conformation of proteins and polypeptides from circular dichroism data. Anal. Biochem. 235:1-10.
A 2 0 −2 −4 −6
Molar ellipticity × 10−3
B 8 6 4 2 0 −2 −4 −6
C 2 0 −2 −4 −6 −8 220
240
260
280
300
320
Wavelength (nm)
Characterization of DNA Structures by Circular Dichroism
Figure 7.11.2 Circular dichroism spectra of nucleic acid structures in 1mM Na2EDTA, 185 mM NaCl, and 6 mM Na2HPO4/2mM NaH2PO4, pH 7.0. (A) Poly(dT) (dashed line), poly(dA) (solid line), and poly(dA):poly(dT) duplex (dotted line). (B) Poly(dA):poly(dT) DNA duplex (solid line), poly(rA):poly(dT) RNA:DNA hybrid (dotted line), and poly(rA):poly(rU) RNA duplex (dashed line). (C) Poly(dA):poly(dT) DNA duplex (solid line) and poly(dA):[poly(dT)]2 DNA triplex (dotted line).
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Table 7.11.1 Peak and Cross-over Bands for Representative Duplex, Triplex, Quadruplex, and i-Motif Nucleic Acid Structuresa
λmin (–) (nm) λcrossover (nm)
λmax (+) (nm)
Calf thymus DNAb (A form) Calf thymus DNAc (B form) [Poly(dGdC)]2d (A form) [Poly(dGdC)]2b (B form) [Poly(dGdC)]2e (Z form) Triplex
210 244 210, 300 205, 252 195, 290
256 258 244, 288 268 217, 278
266 216, 274 273 276 267
[Poly(dA)]2:poly(dT)f Poly(dA):[poly(dT)]2f Quadruplex
248 246
234, 276 253
222, 286 257, 278
Oligo(dG12) (parallel)f Oligo(dG4dT4dG4) (antiparallel)f i-motif
239 264
247 251, 278
257 241, 296
[poly(dC)]4f
265
274
290
Nucleic acid Duplex
a All spectral features are approximations and will vary with solution conditions. b 10.0 mM sodium phosphate buffer, pH 7.0. c 0.7 mM sodium phosphate buffer, pH 7.0, with 80% (v/v) 2,2,2-trifluoroethanol. d 0.67 mM sodium phosphate buffer, pH 7.0, with 80% (v/v) 2,2,2-trifluoroethanol. e 10.0 mM sodium phosphate buffer, pH 7.0, with 2.0 mM sodium perchlorate. f 1.0 mM Na EDTA, 185.0 mM NaCl, and 6.0 mM disodium (dibasic) phosphate/2.0 mM monosodium (monobasic) 2
phosphate, pH 7.0.
12 10 8 6
Molar ellipticity × 10−3
4 2 0 −2 −4 −6 −8 −10 220
240
260
280
300
320
Wavelength (nm)
Figure 7.11.3 Circular dichroism spectra of quadruplex structures. The intramolecular folded antiparallel structure formed by the oligonucleotide 5′AGGG(TTAGGG)3 is shown by the solid line. The four-strand, intermolecular parallel quadruplex structure [5′T2G20T2]4 is shown by the dotted line. The solution conditions are the same as in Figure 7.11.2.
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Ivanov, V.I., Minchenkova, E., Schyolkina, A.K., and Poletayev, A.I. 1973. Different conformations of double-stranded nucleic acid in solution as revealed by circular dichroism. Biopolymers 12:89-110. Johnson, W.C. Jr. 1985. Circular dichroism and its empirical application to biopolymers. Meth. Biochem. Anal. 31:61-163. Johnson, W.C. Jr. 1990. Spectroscopic and kinetic data: Physical data I. Section 3.1: Electronic circular dichroism (CD) spectroscopy of nucleic acids. In Landolt-Bornstein Numerical Data and Functional Relationships in Science and Technology, Vol. VII/1/c (W. Saenger, ed.) pp. 1-24. Springer-Verlag, Heidelberg, Germany. Johnson, W.C. Jr. 2000a. CD of nucleic acids. In Circular Dichroism: Principles and Applications, 2nd ed. (N. Berova, K. Nakanishi, and R.W. Woody, eds.) pp. 703-718. Wiley-VCH, New York. Johnson, W.C. Jr. 2000b. Determination of the conformation of nucleic acids by electronic CD. In Circular Dichroism and The Conformational Analysis of Biomolecules (G.D. Fasman, ed.) pp. 433-468. Plenum Press, New York. Lowry, T.M. 1964. Optical Rotatory Power. Dover Publications, New York. Maurizot, J.C. 2000. Circular dichroism of nucleic acids: Nonclassical conformations and modified oligonucleotides. In Circular Dichroism: Principles and Applications, 2nd ed. (N. Berova, K. Nakanishi, and R.W. Woody, eds.) pp. 719-739. Wiley-VCH, New York. Porumb, H., Monnot, M., and Fermandjian S. 2002. Circular dichroism signatures of features simultaneously present in structured guanine-rich oligonucleotides: A combined spectroscopic and electrophoretic approach. Electrophoresis 23:1013-1020.
Steely, H.T. Jr., Gray, D.M., and Ratliff, R.L. 1986. CD of homopolymer DNA-RNA hybrid duplexes and triplexes containing A-T or A-U base pairs. Nucl. Acids Res. 14:10071-10090. Sun, X.G., Cao, E.H., He, Y.J., and Qin, J.F. 1999. Spectroscopic comparison of different DNA structures formed by oligonucleotides. J. Biomolec. Struct. Dynam. 16:863-872. Tinoco, I. Jr., and Cantor, C. 1970. Application of optical rotatory dispersion and circular dichroism to the study of biopolymers. Meth. Biochem. Anal. 18:81-203. Venyaminov, S.Y. and Yang, J.T. 2000. Determination of protein secondary structure. In Circular Dichroism and the Conformational Analysis of Biomolecules (G.D. Fasman, ed.) pp. 69-107. Plenum Press, New York. Woody, R. 1995. Circular Dichroism. Methods Enzymol. 246:19-71.
KEY REFERENCES Johnson, 1985. See above. A detailed practical guide for measuring CD spectra.
Johnson, 1990. See above. A valuable compilation of nucleic acid CD spectra. Rodger, A. and Norden, B. 1997. Circular Dichroism and Linear Dichroism. Oxford University Press, Oxford, U.K. A brief lucid introduction to the theory of circular dichroism, with many examples of nucleic acid spectra.
Contributed by G. Reid Bishop Mississippi College Clinton, Mississippi Jonathan B. Chaires University of Mississippi Medical Center Jackson, Mississippi
Characterization of DNA Structures by Circular Dichroism
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Biophysical Analysis of Triple-Helix Formation
UNIT 7.12
Formation of a DNA triple helix between a triplex-forming oligonucleotide (TFO) and double-stranded DNA (dsDNA) is a very important process that can be potentially exploited for artificial regulation of gene expression on the level of genomic DNA. Analysis of DNA triple-helix formation in vitro is the first step for any study regarding practical applications of triple helices. This unit contains procedures for two methods for accomplishing such analysis: nondenaturing polyacrylamide gel-shift electrophoresis and DNA thermal denaturation. The first method (Basic Protocol 1) is based on the slower migration of the high-molecular-weight triple-helical complex compared to the target duplex. The second method (Basic Protocol 2) is based on the hypochromic effect upon the association of the third strand to its target DNA. These methods are specific for triple-helix analysis and are not described in previous units of this book. For more details on the treatment of equilibrium melting curves, see UNIT 7.3. NOTE: All reagents used in the following protocols should be of the highest purity grade accessible. They can be purchased from Sigma-Aldrich, Acros Organics, or VWR-Merck, if not otherwise indicated. Oligonucleotides, including fluorescently labeled ones, can be purchased from Eurogentec (http://www/eurogentec.com).
STRATEGIC PLANNING Gel-Shift Assay Choice of oligonucleotide sequences The test system consists of two components: (1) a complementary DNA duplex that contains the target oligopurine-oligopyrimidine tract and (2) the triplex-forming oligonucleotide (TFO). The target consists of two separated complementary strands in equimolar concentrations. From a practical point of view, it is better to use a hairpin structure in which the two strands are linked by at least four thymidine or cytidine moieties. Such a construction minimizes concentration errors and ensures that both strands are in equimolar concentrations. The structure of the TFO depends on the specific task of the assay. It can contain only purines, only pyrimidines, or a mixture of the two. Binding can occur in the parallel or antiparallel direction relative to the oligopurine strand of the target, depending on the base content. Figure 7.12.1 provides an example of two standard assay systems: (1) the polypurine tract of the HIV proviral DNA (Giovannangeli et al., 1997) consisting of a 16-bp polypurine tract in the form of a 62-nt covalent hairpin duplex; and (2) a model DNA sequence for topoisomerase-assisted DNA cleavage (Arimondo et al., 2002) in the form of two separated duplex strands. In both cases, the third strand is a 16-mer pyrimidine oligonucleotide. Labeling the target duplex The simplest method for duplex labeling using commercial oligonucleotides employs [γ-32 P]ATP and T4 polynucleotide kinase (see CPMB UNIT 3.10). For this type of experiment, a final specific activity of ∼0.1 to 1 Ci/mol is suitable for the radioactive duplex. When two separated strands are used, the unlabeled strand should be added in a slight excess in order to avoid the presence of a labeled band corresponding to single-stranded DNA, which may also interact with the third strand. Better labeling is usually obtained with an oligopyrimidine sequence, as compared to an oligopurine sequence. When a hairpin duplex is used, it must be denatured just before labeling, because the polynucleotide
Contributed by Alexandre S. Boutorine and Christophe Escud´e Current Protocols in Nucleic Acid Chemistry (2007) 7.12.1-7.12.16 C 2007 by John Wiley & Sons, Inc. Copyright
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Figure 7.12.1 Sequences for (A) a fragment of the HIV polypurine tract (PPT) in the form of a hairpin covalent duplex, and (B) a model oligopurine-oligopyrimidine duplex for topoisomeraseassisted DNA cleavage studies in the form of two separate strands. The third oligopyrimidine strand is shown above each duplex. The PPT is indicated by italics. In order to stabilize the triplex, T can be replaced by 5-propinyl-U and C by 5-methyl-C in the triplex-forming oligonucleotide.
kinase has much better affinity for single-stranded nucleic acids. Alternatively, enzymatic phosphorylation may be facilitated by introduction of a 5 protruding end. Fluorescent detection can be used instead of radioactive labeling, as the presence of a fluorescent label at the oligonucleotide terminus does not affect triplex formation. Several companies sell fluorescently labeled oligonucleotides. Fluorescent labeling offers several advantages, among them safety with respect to the experimenter and a higher stability of the label. Terminally phosphorylated oligonucleotides can also be labeled in the laboratory with any fluorescent label from the repertoire of Molecular Probes (Invitrogen). Methods for the terminal labeling of oligonucleotides have been described in Grimm et al. (2000) and Boutorine and Sun (2005), as well as in UNITS 4.2, 4.3, 4.5, 4.6, 4.9 & 4.10. Both radioactive and fluorescent labels can be analyzed using the Amersham Typhoon 9410 Variable Mode Imager, supplied with ImageQuant software (Molecular Dynamics).
Conditions for triple-helix formation In the majority of cases using unmodified oligonucleotides, triple-helix formation proceeds at low temperature in buffers containing monovalent and bivalent metal ions (Na+ and Mg2+ ). However, since the main application of triple-helix studies is for gene expression regulation under physiological conditions, tests at physiological temperature and neutral pH with modified TFOs are needed. The choice of buffer depends on the pH required for triplex formation. Commonly used buffering compounds for gel electrophoresis include 2-(N-morpholino)ethanesulfonic acid (MES) for pH ∼6, 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) for pH ∼7, and tris(hydroxymethyl)aminomethane (Tris) for pH ∼8. In order to enhance triplex stability and mimic physiological conditions, monovalent cations (e.g., 50 to 100 mM NaCl) and divalent cations (e.g., 5 mM MgCl2 ) are usually added to the buffer.
Biophysical Analysis of Triple-Helix Formation
Triple-helix formation is sometimes associated with very slow kinetics, so a rather long incubation time can be necessary. Triplex formation can take several hours (Roug´ee et al., 1992; Grimm et al., 2002). Incubation of the labeled duplex with the TFO is usually carried out overnight. The temperature depends on the type of triplex and varies between 4◦ and 37◦ C. When nothing is known about the stability or affinity of the triplex, it is better to fix the labeled duplex concentration (10 to 60 nM) and apply a series of concentrations of TFO in
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the nanomolar to micromolar range at low temperature and eventually at slightly acid pH. The characteristic value of the TFO concentration when half of the target duplex is bound in a triplex can be determined. An apparent dissociation constant can be calculated from these data; however, it will not be a real thermodynamic dissociation constant because the gel electrophoretic separation of the complex is not under equilibrium conditions.
Triplex Melting Choice of oligonucleotide sequences and concentrations This test system consists of the same two components: the complementary DNA duplex and the TFO. The principles for choosing oligonucleotide sequences are the same as for the gel-shift assay, except that no oligonucleotide labeling is needed. It is possible to use either two separate strands or a hairpin oligonucleotide for the duplex. From a practical point of view, the use of two separate strands is preferable, as the synthesis is easier and it will be possible to follow both triplex and duplex melting. The length of the target sequence is usually chosen between 20 and 45 bp. The target sequence should ideally be put in its natural context, as it has been shown that sequences adjacent to a triple helix can influence its stability. If the triple helix is very stable, a longer target will be preferred in order to clearly differentiate between the melting of the triplex and the dissociation of the duplex. The covalent linkage of both strands in the hairpin results in an important increase of the melting temperature of the duplex, and complete melting of the hairpin duplex can be difficult to achieve at temperatures below 90◦ C. Three principles should be considered for oligonucleotide concentrations: (1) the concentrations of oligonucleotides must be close to equimolar; (2) the concentrations must be high enough to provide quantitative complex formation, where practically all the components are involved in a triple helix (in this sense, knowledge of the dissociation constant of the triplex is useful); and (3) the total optical density of the final solution must be in the range of 0.3 to 1.0 OD units/mL/cm. This means that, for an oligonucleotide of 15 to 30 nt, the molar concentration of each component should be 0.5 to 1.5 µM, depending on the molar extinction of the individual oligonucleotides.
Choice of buffer A variety of buffers can be used for melting experiments. Note, however, that the range of experimental temperatures is very high (generally between 5◦ and 95◦ C), so the pH can change drastically during the experiment. The best buffer for thermal dissociation experiments is cacodylate buffer. Its maximal buffering capacity is at a pH close to 6, and the change in pH is only 0.16 over the temperature intervals used (from 6.13 at 5◦ C to 6.29 at 95◦ C). Choice of instrument and thermal denaturation program For thermal denaturation experiments with UV absorption detection, a multiwavelength UV spectrophotometer with a thermostatted cell holder and the ability to follow the exact temperature in the cells is needed. In addition, a mechanical device that automatically changes samples in the spectrophotometer chamber is necessary if several samples are studied simultaneously. Several companies produce such instruments. The authors use the UVIKON XL instrument (Secomam, a Nova Analytic Company; http://www.secomam.com) with thermostatted mobile cell holder for twelve cells, linked to a computer with LifePower TM software (DuSoTec GmbH; http://www.dusotec.de) for programming and managing the thermal denaturation program. The other necessary component is a programmable thermostat. A large variety of programmable thermostats is available (e.g., ThermoElectron, Lauda, Haake, Heto, Neslab). As already mentioned, formation of triple helices is a slow process, so, to keep an equilibrium state at any temperature, slow heating and cooling are necessary. Usually the
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7.12.3
temperature gradient is 0.1◦ to 0.5◦ C/min, so the complete cycle of cooling/heating takes 6 to 15 hr. The most suitable cells are quartz Suprasil Hellma QS114B cells with a 10-mm light path and a total volume of 1400 µL, with black side walls and Teflon hermetic caps. The cells must be hermetically closed during the experiment to avoid evaporation. In addition, a “hot start” of the experiment is preferable. This means that, to avoid complications connected with secondary structures of the tested samples, it is best to start the cycle from the highest temperature, when all the components are denatured. BASIC PROTOCOL 1
NONDENATURING GEL-SHIFT STUDIES OF TRIPLE-HELIX FORMATION The gel-shift separation is based on the different migration rates of duplex and triplex DNA in a nondenaturing polyacrylamide gel in an electric field. A triplex has a higher molecular weight than the target duplex alone, so it migrates more slowly in the gel. In the present protocol, preparation of the sample and nondenaturing gel electrophoresis are described in detail. The optimal gel concentration depends on oligonucleotide length (between 12% and 18% for a 40- to 60-mer duplex). The gel is typically loaded with samples of increasing concentrations of TFO, with control labeled duplex (without TFO) in the first and last wells. In order to prevent eventual triplex dissociation, electrophoresis is performed at 4◦ to 10◦ C and at a power setting of 5 to 6 W to avoid heating of the gel. The gel is directly scanned on a fluorescence scanner or dried and exposed with X-ray film or a phosphor screen. The image can be then analyzed using scanning image analysis software such as ImageQuant. In the majority of cases, it is necessary to determine the affinity of the TFO for the target and to calculate an affinity constant between the TFO and its target (see Strategic Planning). In this case, triplex formation is calculated as a function of TFO concentration at a fixed duplex concentration. The procedure that follows provides a specific example, in which seven samples are prepared with a constant concentration of labeled duplex (60 nM) and increasing concentrations of TFO (50 nM, 100 nM, 250 nM, 500 nM, 750 nm, 1 µM, and 1.5 µM). Each sample contains 15 µL of the initial labeled duplex solution and is brought to a final volume of 20 µL with TFO and water. This example provides a guideline that can be used to recalculate the proposed experimental scheme for any oligonucleotide concentration and any number of samples.
Materials 50 to 100 µM fluorescently or radioactively labeled duplex (target DNA) in H2 O (two separate strands or a single hairpin oligonucleotide) 50 to 100 µM triplex-forming oligonucleotide (TFO) in H2 O 10× MES buffer (see recipe) or HEPES buffer (see recipe) 30% (w/v) sucrose with 0.025% (w/v) bromphenol blue and 0.025% (w/v) xylene cyanol FF 40% acrylamide/N,N -methylene-bis-acrylamide (19:1) N,N,N N -Tetramethylethylenediamine (TEMED) 10% (w/v) ammonium persulfate
Biophysical Analysis of Triple-Helix Formation
0.5- to 1.5-mL polypropylene microcentrifuge tubes (e.g., Eppendorf) 90◦ C water bath or dry heating block 50- to 100-mL side-arm flask and rubber or silicone stopper Sintered-glass funnel filter, no. 3 or 4 Vertical electrophoresis system with glass plates (e.g., 16.5 × 14–cm plates with 0.3- to 0.6-mm spacers and 12- to 16-well comb with 0.5- to 0.8-cm-long teeth)
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High-voltage electrophoresis power supply with power regulation Whatmann 3MM chromatographic paper Gel dryer Fluorescence gel scanner or radioactive phosphor imager (e.g., Phosphor Imager or Typhoon instrument from GE Healthcare) Storage phosphor screens and cassettes for exposure to radioactive gels () Scanning image analysis software (e.g., ImageQuant v. 5, Molecular Dynamics) Additional reagents and equipment for gel electrophoresis (APPENDIX 3B) CAUTION: Radioactive labeling of oligonucleotides must be done in specially equipped laboratories that have permission to work with isotopes such as 32 P or 33 P. CAUTION: Acrylamide is toxic and must be handled using gloves and appropriate eye protection.
Prepare test samples To test triplex formation 1a. Dilute labeled duplex solution to 1.2 µM and TFO solution to 5 µM with water. 2a. Add the following to each of two microcentrifuge tubes:
2 µL 10× MES (or HEPES) buffer 4 µL 30% sucrose with bromphenol blue and xylene cyanol 1 µL 1.2 µM labeled duplex (final 60 nM) 1 to 4 µL 5 µM TFO (final 0.25 to 1 µM) H2 O to 20 µL. 3a. Mix well by vortexing and proceed to step 4. The TFO concentration should be 0.25 to 1 µM, depending on the expected stability of the triplex. The duplex concentration can range from 10 to 100 nM.
To test dependence of triplex formation on TFO concentration 1b. Dilute labeled duplex solution to 1.2 µM and TFO solution to 10 µM with water. Take 5 µL of the 10 µM TFO and dilute 1:10 with water to obtain a 1.0 µM concentration. 2b. Prepare the initial labeled duplex solution in a 1.5-mL microcentrifuge tube by mixing the following components:
20 µL 10× MES (or HEPES) buffer 40 µL 30% sucrose with bromphenol blue and xylene cyanol 10 µL 1.2 µM labeled duplex solution 80 µL H2 O (final 150 µL). Mix well by vortexing. Nine samples will be applied (seven experimental and two control). Taking into account possible errors in the pipetting, it is prudent to prepare a total volume corresponding to n+1 samples, in this case 150 µL. Concentrations of buffer, sucrose, and duplex must be calculated for a final volume of 200 µL (20 µL per sample), taking into consideration the subsequent addition of TFO and/or water.
3b. Distribute 15 µL of the mixture into each of nine microcentrifuge tubes and add TFO solutions and water according to Table 7.12.1. Proceed to step 4. Biophysical Analysis of Nucleic Acids
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Table 7.12.1 Sample Preparation for Testing Triple-Helix Formation as Function of TFO Concentration
Tube
Final TFO concentration (µM)
Volume 1.0 µM TFO (µL)
Volume 10 µM TFO (µL)
Water (µL)
1
0
—
—
5
2
0.05
1
—
4
3
0.10
2
—
3
4
0.25
5
—
—
5
0.50
—
1
4
6
0.75
—
1.5
3.5
7
1.0
—
2
3
8
1.5
—
3
2
9
0
—
—
5
Denature and renature samples to facilitate triplex formation 4. Place the samples in a dry heating block or water bath and incubate 3 min at 90◦ C. 5. Turn off the heating device and allow the samples to return slowly to room temperature as the device cools. 6. When the temperature in the tubes reaches room temperature, transfer the samples to a refrigerator and incubate at least 3 hr at 4◦ C, preferably overnight.
Prepare 12% nondenaturing polyacrylamide gel 7. Combine the following and mix on a magnetic stirrer: 6 mL 40% acrylamide/N,N -methylene-bis-acrylamide solution (19:1) 2 mL 10× MES buffer (or HEPES buffer, depending on pH of complex formation) 12 mL H2 O. Preparation of the gel and electrophoresis are similar to methods described in APPENDIX 3B, but without the denaturing agent and with other modifications indicated below.
8. Filter the mixture through a sintered glass filter using a side-arm flask and vacuum aspirator. 9. Close the flask with a rubber or silicone stopper and degas the solution 3 to 5 min under vacuum. 10. Add 12 µL TEMED and 120 µL of 10% ammonium persulfate solution, mix well, and immediately pour the gel solution between the glass electrophoresis plates with spacers. Insert the comb to form the wells for sample loading. 11. Remove the comb and rinse the wells with 1× MES or HEPES buffer. Install the gel in the electrophoresis apparatus in a cold room, fill the electrophoresis chambers with 1× MES or HEPES buffer, and turn the current on. Adjust the power to 10 W and leave the gel to pre-electrophorese for ∼1 hr. Biophysical Analysis of Triple-Helix Formation
Load samples and run gel 12. Turn off the power supply, remove the buffer from the chambers, and replace it with fresh buffer. Wash the wells with 1× MES or HEPES buffer.
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13. Load the 7- to 10-µL samples into the wells. Start electrophoresis, adjusting the power to 6 W maximum. Approximately one-half of the sample should be reserved in case the gel needs to be run again. The MES and HEPES buffers do not have enough buffering capacity to support long electrophoresis, as Tris-borate buffer does. Thus, the pH in the chambers changes quickly during electrophoresis. It is therefore necessary either to replace the buffer every hour or to remove the buffer from both cells into one beaker, mix the solution well, and return it to the chambers.
14. Stop the electrophoresis when the bromphenol blue reaches ∼2/3 of the gel length (∼3 to 6 hr). 15. Remove the gel from the glass plates.
Scan results For fluorescently labeled samples 16a. Place the gel on the scanning plate of a fluorescence scanner and scan the gel according to manufacturer’s instructions. 17a. Proceed to step 18.
For radioactively labeled samples 16b. Transfer the gel to Whatmann 3MM paper and cover it with plastic wrap. Place the wrapped gel on the plate of a vacuum gel dryer heated to 80◦ C. Cover with the plastic cover and dry 15 min under vacuum. 17b. Place the gel in the exposure cassette with the storage phosphor screen and leave overnight. Scan the screen on the Phosphor Imager or Typhoon scanning instrument according to manufacturer’s instructions. Proceed to step 18.
Analyze results 18. Analyze the fluorescence or phosphor image using any scanning software (e.g., ImageQuant). A typical image obtained from a gel-shift experiment is shown in Figure 7.12.2. The resulting data represent the percentage of the signal in the upper band (corresponding to triplex) relative to total fluorescence or radioactivity in the lane. From
Figure 7.12.2 Gel-shift analysis of triple-helix formation between oligonucleotide 5 C*U*C*U*C*U*C*U*C*U*U*U*U*U*U*U*p-3 and DNA duplex 5 -GATAGAGAGAGAGAAAAAAA GAGAAGATC-3 / 5 -32 P-GATCTTCTCTTTTTTTCTCTCTCTCTATC-3 . U* is 5-propinyluridine and C* is 5-methylcytosine. Analysis performed on a 12% nondenaturing polyacrylamide gel in 1× HEPES buffer (see Reagents and Solutions). Duplex concentration, 60 nM; third-strand concentrations, 0, 0.01, 0.05, 0.1, 0.5, 1, 5, and 0 µM (lanes 1 to 8, respectively). Preliminary incubation of the triplex in the buffer was 2 hr at 37◦ C. The Typhoon 9410 variable-mode imager was used as a scanner. The lower band corresponds to the duplex and the higher band to the triplex.
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Figure 7.12.3 Determination of the apparent dissociation constant of a triple helix using the data from a gel-shift experiment. See explanation in text.
the corresponding graph, a value of the TFO concentration when half of the duplex is involved in a triplex can be determined.
19. Calculate the apparent dissociation constant. This calculation is based on the equation for the dissociation constant (Kd ):
where D is duplex concentration, S is TFO concentration, and T is triplex concentration. If D0 and S0 are the initial duplex and TFO concentrations, and x is the part of the duplex involved in a triplex, the equation is transformed to:
Conversion of this equation gives:
Thus, logKd can be determined as the y intersection of a graph built on the following coordinates:
This task can be facilitated by creation of a calculation list in a graphing software program such as Microsoft Excel or KaleidaGraph, where the best parameters for fitting of experimental data to the linear function (fourth equation) can be determined. An example of the graph is shown in Figure 7.12.3. BASIC PROTOCOL 2
Biophysical Analysis of Triple-Helix Formation
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THERMAL DENATURATION STUDIES OF TRIPLE-HELIX FORMATION This method (also called triplex melting) permits determination of the thermodynamic characteristics of a triple helix (Roug´ee et al., 1992). In routine work, the most interesting parameter that can be determined directly from a melting curve is the temperature of the triplex-to-duplex transition. The melting temperature (Tm ) is the temperature where half of the triplex has dissociated into duplex plus third strand. Triplex melting can be followed by UV spectrophotometry. The extinction coefficient of the triple helix is lower than that of the dissociated complex, so UV absorbance is increased upon heating (dissociation) of the triplex. This increase is cooperative and has a sharp transition that permits detection of the half-dissociation temperature with quite high precision. Current Protocols in Nucleic Acid Chemistry
Materials 10× cacodylate buffer (see recipe) 130 µM target DNA in H2 O (oligonucleotide duplex in the form of two complementary strands in equimolar concentrations) 130 µM triplex-forming oligonucleotide (TFO) in H2 O Mineral oil 0.5- to 1.5-mL polypropylene microcentrifuge tubes (e.g., Eppendorf) UV-Vis spectrophotometer with thermostatted cells (e.g., UVIKON XL), linked to a computer with data-acquisition software Programmed thermostat (e.g., ThermoElectron, Lauda, Haake, Heto, Neslab) Vacuum desiccator Quartz spectrophotometer cells with black side walls and hermetic Teflon caps (e.g., Hellma QS114B quartz cells) Graphing software: e.g., Microsoft Excel or KalaidaGraph (Synergy Software; http://www.synergy.com/) CAUTION: Cacodylic acid and its salts are highly toxic. Observe all precautions for handling and disposal of this material, including the use of gloves and appropriate eye protection.
Prepare test samples 1. Prepare the following mixture in a microcentrifuge tube for each sample: 60 µL 10× cacodylate buffer 6 µL 130 µM duplex solution (or 6 µL of each strand solution if separated) 6 µL 130 µM TFO solution H2 O to 600 µL. Mix by vortexing. The minimum volume that can be measured in a QS114B cell is 600 µL. These concentrations may be adapted for very short or very long sequences. It is a good idea to choose concentrations that will provide an optical density between 0.3 and 1.0. The hypochromic effect due to triplex formation is not as strong as for duplex formation. Moreover, sequences for triplex formation are usually rather short (12 to 20 bp). Therefore, the melting will often be more clearly observed at higher concentrations. The UVIKON XL spectrophotometer has room for twelve cuvettes, but three of them cannot contain samples: one is used for the temperature sensor and two are used as a reference for the light beam. Therefore, nine samples can be recorded at the same time. It is advised that the melting curve of the duplex alone be recorded, as well as that of the third strand alone. Transitions observed in these experiments indicate the formation of alternative structures that may compete with triple-helix formation, and that will in any case complicate the analysis of the triplex melting profiles.
2. Check the spectra of the samples. If the maximal absorption at 260 nm is not in the range of 0.3 to 1.0, adjust the DNA concentration by adding more DNA or by diluting with 1× cacodylate buffer.
Program thermostat 3. Program the thermostat as described in the operator’s manual and according to the following guiding principles: a. The starting point must be the highest temperature of the denaturation cycle (e.g., 95◦ C).
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b. The lowest point of the cycle must be at least 10◦ C lower than the presumed triplex melting point (if known), for example, 15◦ C. c. A broad interval of temperatures, from 5◦ to 90◦ or 95◦ C, is used to observe both triplex and duplex melting. To observe just triplex melting, a narrower interval and lower temperatures can be used. d. The rate of temperature variation must take into account the slow kinetics of triple-helix formation. The whole length of the cycle can reach more than 8 hr. For example, if the total temperature interval during the experiment is 72◦ C (from 20◦ to 92◦ C) and the rate of change is 0.1◦ C/min, the cycle will take 24 hr.
Program spectrophotometer detection unit 4. Program the detection unit for data acquisition according to the instructions for the spectrophotometer and programming/acquisition software. Select the wavelength, the time of acquisition, and the time intervals between data points using the following guiding principles. a. Even in simple experiments when the absorbance changes are followed at a single maximum absorption wavelength (260 nm for standard oligonucleotides), a control background measurement must be taken at a wavelength where nucleic acids do not absorb (typically 340 or 580 nm). Subtraction of background absorption from the experimental data helps to avoid errors due to the temperature-dependent changes of optical properties of solvent or other components often present in the sample. In addition, cytosine protonation can be followed at 295 nm. At this wavelength, triplex formation is associated with an increase in absorbance. b. The temperature inside the spectrophotometer cells must be thoroughly controlled by installing a thermosensor into a separate control cell. The data from the thermosensor are registered together with each measurement of optical density in the acquisition file. c. The frequency of data acquisition should be set to every 150 to 600 sec at each wavelength. The frequency usually depends on the temperature variation rate. It can be convenient to set one acquisition per 1◦ C. d. The complete cycle can be repeated. A good coincidence of two consecutive renaturation-denaturation curves, and the absence of hysteresis, demonstrate that the thermal denaturation has been done under conditions of thermodynamic equilibrium. Nevertheless, one must be careful that heating did not result in evaporation of the sample. e. If experimental temperatures are lower than room temperature, air can be blown through the cell chamber to help prevent condensation of water on the cell walls.
Run and save baseline 5. Fill the control and the experimental cells with 1× cacodylate buffer. Run the baseline in the range of wavelengths that will be used for data acquisition, then remove buffer from all the cells except the control cells. Degas samples and start experiment 6. Open the sample tubes, place in a vacuum desiccator, and degas the solutions with the help of a vacuum aspirator for 2 to 3 min. To avoid loss of sample due to boiling up in vacuo, leave tubes closed but perforate the caps with a needle.
Biophysical Analysis of Triple-Helix Formation
Alternatively, to remove dissolved oxygen without vacuum degassing, when heating the cells to highest temperature (step 8), some bubbles will likely form, and these can be removed by tapping the cell with one’s fingers.
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7. Transfer the samples into the quartz spectrophotometer cells. Add five to seven drops of mineral oil above each sample solution to minimize solvent evaporation. Close the cells hermetically with the Teflon caps. 8. Install the cells in the cell holder of the spectrophotometer. Place the control cells into the corresponding control compartments. Close the chamber. Start the thermostat and begin blowing air through the cell chamber, then wait until the temperature in the cells reaches the highest point in the cycle. Remove any bubbles that form. 9. Start the spectrophotometer and the thermostat program at the same time.
Stop experiment and register data 10. Export the numeric data into a Microsoft Excel or KaleidaGraph calculation list according to the software export method. In LifePower TM software, simply use the menu option “Export to Excel.” After the end of the experiment, treatment of the thermal denaturation data is necessary for exact determination of the triplex melting point. The acquisition software permits the numeric data tables to be exported into calculation lists compatible with graphic software such as Microsoft Excel or KaleidaGraph.
Figure 7.12.4 Thermal denaturation experiment for a triple helix formed from a duplex of two complementary strands, 5 -CCACTTTTTAAAAGAAAAGGGGGGACTGG-3 and 3 GGTGAAAAATTTTCTTTTCCCCCCTGACC-5 , and the third strand, 5 -TTTTCTTTTCCCCCCT3 . (A) Typical thermal denaturation curve. (B) First derivative of the thermal denaturation curve calculated using Microsoft Excel. Both curves show the melting points of the triple helix (1) and duplex (2). The experiment was done in a standard cacodylate buffer (see text).
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11. Open the exported file with Microsoft Excel or KaleidaGraph software. Use the graphical functions to prepare a melting curve as demonstrated in Figure 7.12.4A. 12. Using the derivative functions of the software (Menu Insertion → Function → Slope in Excel or Menu Macros → Derivative in KaleidaGraph), build the derivative dA/dT◦ , where A is optical density and T◦ is temperature. To obtain a smooth derivative curve (Fig. 7.12.4B), use an average slope of eight to ten points around any point.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Cacodylate buffer, 10× Dissolve 21.4 g sodium cacodylate trihydrate [NaO2 As(CH3 )2 ·3H2 O] in 250 mL of 2 M NaCl. Add 12.5 mL of 2 M MgCl2 . Add water to 350 to 400 mL and titrate to pH 6.0 with 5 M HCl. Adjust buffer volume to 500 mL with water. Store 10× buffer several months at 4◦ C; store 1× working buffer up to 1 month at 4◦ C. Final concentrations: 200 mM cacodylate, 1 M NaCl, and 50 mM MgCl2 .
HEPES buffer, 10× Dissolve 59.6 g 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) in 125 mL of 2 M NaCl. Add water to 350 to 400 mL and titrate with 5 M NaOH to pH 7.5. Add 12.5 mL of 2 M MgCl2 . Adjust pH to 7.5, if necessary. Adjust total buffer volume to 500 mL with water. Store 10× buffer several months at 4◦ C; store 1× working buffer up to 1 month at 4◦ C. Final concentrations: 500 mM HEPES, 500 mM NaCl, and 50 mM MgCl2 .
MES buffer, 10× Dissolve 48.8 g 2-(N-morpholino)ethanesulfonic acid (MES) in 125 mL of 2 M NaCl. Add water to 350 to 400 mL and titrate with 5 M NaOH to pH 6.0. Add 12.5 mL of 2 M MgCl2 . Adjust pH to 6.0, if necessary. Adjust buffer volume to 500 mL with water. Store 10× buffer several months at 4◦ C; store 1× working buffer up to 1 month at 4◦ C. Final concentrations: 500 mM MES, 500 mM NaCl, and 50 mM MgCl2 .
COMMENTARY Background Information
Biophysical Analysis of Triple-Helix Formation
Triple-helix formation was first described by Felsenfeld and Rich (1957). However, the idea of applying triple–helix formation for regulation of gene expression on the level of dsDNA appeared when two independent groups (those of C. H´el`ene and P. Dervan) demonstrated triple-helix formation by synthetic oligonucleotides and chemical modification of dsDNA using modified triple-helixforming oligonucleotides (TFO; Le Doan et al., 1987; Moser and Dervan, 1987). During the last 20 years, different types of triple helices have been described (for reviews, see Praseuth et al., 1999; Guntaka et al., 2003; Rogers et al., 2005), and the biological effects of TFOs in living cells have been
demonstrated (Faria et al., 2000; Brunet et al., 2006). In a DNA triple helix, the third strand of the triplex is bound within the major groove of an oligopurine-oligopyrimidine tract of the target duplex by Hoogsteen or reversed Hoogsteen hydrogen bonds. The orientation of the third strand and the nature of the base triplets are different for these two types of triple helices (Sun et al., 1996). Ionic and pH conditions that favor triplex formation are also different. In particular, the formation of Hoogsteen-type triple helices sometimes requires protonation of cytosines, which is facilitated by a slightly acid pH. The requirement for an oligopurine tract and the poor stability of certain triple helices under physiological conditions have
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been regarded as the primary limitations for practical applications of triple helices. Significant improvements that overcome these restrictions have been achieved by introducing various chemical modifications into the TFOs, either at the DNA backbone (Gryaznov and Banait, 2002; Jepsen et al., 2004) or at the nucleobases (Morvan et al., 1995; Giovannangeli et al., 1996; Lacroix and Mergny, 2000; Sollogoub et al., 2002). Numerous efforts to design universal artificial bases that can recognize any DNA sequence at physiological pH have been realized (Rusling et al., 2005). For a general review, see Fox (2000). Melting curves are usually recorded for Hoogsteen-type triple helices. Although rare, the presence of detectable hypochromism upon triplex formation has been reported for reverse Hoogsteen triple helices. This method is widely used for estimation of triple-helix stability, especially for the study of chemical modifications of the third strand and triplexstabilizing agents (Escud´e et al., 1993, 1995, 1996). The gel-shift method is a general method for detecting different nucleic acid complexes that are stable under nondenaturing gel electrophoresis conditions, such as nucleic acid– nucleic acid, nucleic acid–protein, or nucleic acid–ligand complexes. In this sense, it is universal and can be applied for any type of triple helix. These two methods provide complementary information on the parameters of triple helix formation. Nevertheless, the gelshift method does not provide the results under thermodynamic equilibrium, as the thermal denaturation method does.
Critical Parameters and Troubleshooting Gel-shift assay The gel-shift method of triplex formation analysis is not very complicated and is accessible for any molecular biology laboratory familiar with gel electrophoresis techniques, radioactive or fluorescent labeling of nucleic acids, and nucleic acid analysis. It requires no additional reagents and equipment beyond those needed for DNA sequencing and fluorescent or radioactive probe analysis. Some critical points in the procedure must be noted. The best target duplex is a covalent hairpin. This eliminates problems that can arise from pipetting errors, as both strands of the duplex are always in equimolar concentrations. However, the yield of phosphate incorporation
during the radiolabeling process can be low due to the perfect double-stranded structure of the 5 terminus. Duplex denaturation before labeling can be helpful. Heat the oligonucleotide for 3 min at 90◦ C and chill quickly in an ice bath. A protruding 5 terminus of labeled oligonucleotide (2 to 3 nucleotides) may also facilitate successful labeling. It is easier to label single-stranded oligonucleotides, the oligopyrimidine strand being preferable for labeling. After labeling, the duplex strands must be mixed and reannealed in exactly equimolar concentrations (or even with an excess of unlabeled strand) to avoid the appearance of radioactive bands of unannealed labeled strand. Another problem is the formation of structures that compete with triple-helix formation. This is particularly crucial with G-rich oligonucleotides (Olivas and Maher, 1995) and alternating GA sequences (Noonberg et al., 1995), and is more important when the third strand contains these sequences or when separate oligopurine and oligopyrimidine strands are used as a target duplex. In this case, preliminary hybridization of the target duplex prior to the experiment will be helpful. Although the zwitterionic character of MES and HEPES buffers should prevent buffer migration, some pH changes in the electrophoresis chambers can be noticed during electrophoresis. It is very important to check pH every hour and change electrophoresis buffer, if necessary. A change in the color of the bromphenol blue dye (to yellow) or a distortion of the migration front indicate a significant pH variation in which the buffer in the lower chamber is becoming very acidic. Sometimes, especially in the case of fluorescent detection, the presence of bromphenol blue and xylene cyanol interfere with signal detection. In this case, it is better to avoid these colorants in the experimental samples and include them only in empty wells located on either side of the experimental wells. The temperature regime during electrophoresis is very important. It can be monitored with special adhesive temperaturesensitive strips on liquid crystals that change color with temperature. It is advised that the first experiments be done at low temperature in a cold room (4◦ to 6◦ C). Since electrophoresis heats gels, high-voltage electrophoresis should not be used. The best method for controlling temperature is to adjust the power and not the voltage or the current. Usually, a power below 6 W does not cause strong heating of the gel. If the triplex dissociation temperature has
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been determined (see Basic Protocol 2), it is easier to find the optimal temperature for gel electrophoresis. However, due to nonequilibrium conditions in the gel, it is advised that the gel temperature be kept much lower than the determined triplex melting point. Prior experience with routine techniques for manipulating nucleic acids, especially electrophoresis, is required. General laboratory safety is also of primary concern when hazardous materials (acrylamide, organic solvents, radioactivity) are involved.
Biophysical Analysis of Triple-Helix Formation
Melting curves Aside from the requirement for specific equipment and software, the melting temperature method is not complicated and only a few technical problems should be noted. The first is the appearance of bubbles in heated solutions. To prevent this problem, a degassing step or manual removal of bubbles after sample heating is necessary. Condensation of liquid on the cell walls can be prevented by blowing compressed air into the cell chamber. The minimal volume of sample solution analyzed in the Hellma QS114B quartz cells is 600 µL. The aqueous solution must be covered by 3 to 4 mm of mineral oil and the cell must be hermetically closed with a Teflon cap; otherwise, the evaporation of the water will dramatically affect the results. Do not leave too much air in the cell when closing the cap tightly, as air that expands upon heating will push the cap out of the cell. The criteria of reproducibility of the results and equilibrium conditions of the experiment are met when there is perfect coincidence of the curves recorded at both cooling and heating steps. The presence of a hysteresis indicates the slow kinetics of association or dissociation of the complex and nonequilibrium processes. This hysteresis can be used for studies of kinetic parameters (Roug´ee, 1992). To study thermodynamic parameters at equilibrium, the rate of temperature variation should be decreased. As already noted, a “hot start” of the experiment is preferable because it favors hybridization of nucleic acids and helps to dissociate possible competing structures. However, formation by separate strands of alternative structures competing with triple-helix formation can seriously complicate the results. Such structures must be controlled by control melting of separated components of the triplex. Prior experience with routine nucleic acid manipulation techniques is required. General
laboratory safety is also of primary concern when hazardous materials are involved.
Anticipated Results Gel-shift assays provide information about the formation and stability of triple-helix complexes. Apparent dissociation constants can also be calculated. Upon variation of electrophoresis temperature, buffer pH, and component concentrations, these experiments will indicate the conditions and limits of triplex formation for any particular oligonucleotide (natural or modified) and its DNA targets. The kinetics of triplex formation may also be studied by gel-shift assay, as described in Novopashina et al. (2005). Melting curves are used to determine the triplex dissociation point. Thermodynamic and kinetic parameters of triple helices can also be extracted from melting curves; however, this is not the aim of these experimental protocols. Interested readers are referred to original articles (Roug´ee et al., 1992; Mergny and Lacroix, 2003) and to UNIT 7.3 for analysis of equilibrium melting curves by optical methods.
Time Considerations All procedures for the gel-shift assay can be accomplished in 1 to 2 days. Typically, samples are prepared for analysis in the evening and are left at the selected incubation temperature overnight in order to ensure triplex formation. The next morning, the gel is prepared, pre-electrophoresed for 1 hr, and then electrophoresed 3 to 6 hr. Scanning of a fluorescent gel is very quick. For radioactive gels, drying (with all the preliminary procedures) can take up to 1 hr, and exposure with the storage phosphor screen takes from 20 min to overnight, depending on the specific radioactivity of the labeled DNA. All procedures for melting curves can be accomplished in less than 1 day, although this can increase to 2 to 3 days if several cycles of renaturation-denaturation are performed. It is convenient to prepare the samples for analysis in the evening and leave the melting experiments to run overnight.
Literature Cited Arimondo, P.B., Boutorine, A.S., Baldeyrou, B., Bailly, C., Kuwahara, M., Hecht, S.M., Sun, J.S., Garestier, T., and H´el`ene, C. 2002. Design and optimization of camptothecin conjugates of triple helix–forming oligonucleotides for sequence-specific DNA cleavage by topoisomerase. J. Biol. Chem. 277:3132-3140.
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Boutorine, A.S. and Sun, J.S. 2005. Postsynthetic functionalization of triple helix–forming oligonucleotides. In Oligonucleotide Synthesis: Methods and Applications (P. Herdewijn, ed.) pp. 251-260. Humana Press, Totowa, N. J. Brunet, E., Corgnali, M., Cannata, F., Perrouault, L., and Giovannangeli, C. 2006. Targeting chromosomal sites with locked nucleic acid-modified triplex-forming oligonucleotides: Study of efficiency dependence on DNA nuclear environment. Nucl. Acids Res. 34:4546-4553. Escud´e, C., Franc¸ois, J.C., Sun, J.S., Ott, G., Sprinzl, M., Garestier, T., and H´el`ene, C. 1993. Stability of triple helices containing RNA and DNA strands: Experimental and molecular modeling studies. Nucl. Acids Res. 21:5547-5553. Escud´e, C., Nguyen, C.H., Mergny, J.L., Sun, J.S., Bisagni, E., Garestier, T., and H´el`ene, C. 1995. Selective stabilization of DNA triple helices by benzopyridoindole derivatives. J. Am. Chem. Soc. 117:10212-10219. Escud´e, C., Giovannangeli, C., Sun, J.S., Lloyd, D.H., Chen, J.K., Gryaznov, S.M., Garetier, T., and H´el`ene, C. 1996. Stable triple-helices formed by N3 →P5 oligophosphoramidates inhibit in vitro transcription elongation. Proc. Natl. Acad. Sci. U.S.A. 93:4365-4369. Faria, M., Wood, C.D., Perrouault, L., Nelson, J.S., Winter, A., White, M.R.H., H´el`ene, C., and Giovannangeli, C. 2000. Targeted inhibition of transcription elongation in cells mediated by triplexforming oligonucleotides. Proc. Natl. Acad. Sci. U.S.A. 97:3862-3867. Felsenfeld, G. and Rich, A. 1957. Studies on the formation of two- and three-stranded polyribonucleotides. Biochim. Biophys. Acta 26:457-468. Fox, K.R. 2000. Targeting DNA with triplexes. Curr. Med. Chem. 7:17-37. Giovannangeli, C., Perrouault, L., Escud´e, C., Thuong, N.T., and H´el`ene, C. 1996. Specific inhibition of in vitro transcription elongation by triplex-forming oligonucleotide-intercalator conjugates targeted to HIV proviral DNA. Biochemistry 35:10539-10548. Giovannangeli, C., Diviacco, S., Labrousse, V., Gryaznov, S., Charneau, P., and H´el`ene, C. 1997. Accessibility of nuclear DNA to triplexforming oligonucleotides: The integrated HIV-1 provirus as a target. Proc. Natl. Acad. Sci. U.S.A. 94:79-84. Grimm, G.N., Boutorine, A.S., and H´el`ene, C. 2000. Rapid routes of synthesis of oligonucleotide conjugates from non-protected oligonucleotides and ligands possessing different nucleophilic or electrophilic functional groups. Nucleosides Nucleotides Nucleic Acids 19:19431965. Grimm, G.N., Boutorine, A.S., Lincoln, P., Nord´en, B., and H´el`ene, C. 2002. Formation of DNA triple helices by an oligonucleotide conjugated to a fluorescent ruthenium complex. Chembiochem. 3:324-331. Gryaznov, S.M. and Banait, N.S. 2002. DNA and RNA analogues: Oligonucleotide phosphorami-
dates with bridging nitrogen. Expert Opin. Ther. Patents 12:543-559. Guntaka, R.V., Varma, B.R., and Weber, K.T. 2003. Triplex-forming oligonucleotides as modulators of gene expression. Int. J. Biochem. Cell Biol. 35:22-31. Jepsen, J.S., Sorensen, M.D., and Wengel, J. 2004. Locked nucleic acid: A potent nucleic acid analog in therapeutics and biotechnology. Oligonucleotides 14:130-146. Lacroix, L. and Mergny, J.L. 2000. Chemical modification of pyrimidine TFOs: Effect on imotif and triple helix formation. Arch. Biochem. Biophys. 381:153-163. Le Doan, T., Perrouault, L., Praseuth, D., Habhoub, N., D´ecout, J.L., Thuong, N.T., Lhomme, J., and H´el`ene, C. 1987. Sequence-specific recognition, photocrosslinking and cleavage of the DNA double helix by an oligo-[α]-thymidilate covalently linked to an azidoproflavine derivative. Nucl. Acids Res. 15:7749-7760. Mergny, J.L. and Lacroix, L. 2003. Analysis of thermal melting curves. Oligonucleotides 13:515537. Morvan, F., Chaix, C., Zeissler, A., Rayner, B., and Imbach, J.L. 1995. Triple helix forming αoligonucleotides containing 5-methylcytosine and/or 5-bromouracil. Nucleosides Nucleotides 14:975-977. Moser, H.E. and Dervan, P.B. 1987. Sequencespecific cleavage of double helical DNA by triple helix formation. Science 238:645-650. Noonberg, S.B., Francois, J.C., Garestier, T., and Helene, C. 1995. Effect of competing selfstructure on triplex formation with purine-rich oligodeoxynucleotides containing GA repeats. Nucl. Acids Res. 23:1956-1963. Novopashina, D.S., Sinyakov, A.N., Ryabinin, V.A., Venyaminova, A.G., Halby, L., Sun, J.S., and Boutorine, A.S. 2005. Sequence-specific conjugates of oligo(2 O-methylribonucleotides) and hairpin oligocarboxamide minor-groove binders: Design, synthesis, and binding studies with doublestranded DNA. Chem. Biodivers. 2:936-952. Olivas, W.M. and Maher, L.J. III. 1995. Competitive triplex/quadruplex equilibria involving guaninerich oligonucleotides. Biochemistry 34:278284. Praseuth, D., Guieysse, A.L., and H´el`ene, C. 1999. Triple helix formation and the antigene strategy for sequence-specific control of gene expression. Biochim. Biophys. Acta 1489:181206. Rogers, F.A., Lloyd, J.A., and Glazer, P.M. 2005. Triplex-forming oligonucleotides as potential tools for modulation of gene expression. Curr. Med. Chem. 5:319-326. Roug´ee, M., Faucon, B., Mergny, J.L., Barcelo, F., Giovannangeli, C., Garestier, T., and H´el`ene, C. 1992. Kinetics and thermodynamics of triplehelix formation: Effects of ionic strength and mismatches. Biochemistry 31:9269-9278.
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Rusling, D.A., Powers, V.E., Ranasinghe, R.T., Wang, Y., Osborne, S.D., Brown, T., and Fox, K.R. 2005. Four base recognition by triplexforming oligonucleotides at physiological pH. Nucl. Acids Res. 33:3025-3032. Sollogoub, M., Darby, R.A.J., Cuenoud, B., Brown, T., and Fox, K.R. 2002. Stable DNA triple helix formation using oligonucleotides containing 2 -aminoethoxy,5-propargylamino-U. Biochemistry 41:7224-7231. Sun, J.S., Garestier, T., and H´el`ene, C. 1996. Oligonucleotide directed triple helix formation. Curr. Opin. Struct. Biol. 6:327-333.
Contributed by Alexandre S. Boutorine and Christophe Escud´e Mus´eum National d’Histoire Naturelle Paris, France
Biophysical Analysis of Triple-Helix Formation
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CHAPTER 8 Nucleic Acid Binding Molecules INTRODUCTION he association of nucleic acids with other intracellular molecules, including proteins and small molecules, is vitally important to nucleic acid function. Understanding the biological consequences of these interactions is significantly aided by knowing the mode, kinetics, and thermodynamics of these binding interactions. Moreover, it is extremely important to understand xenobiotic interactions with nucleic acids, including drugs and carcinogens.
T
This chapter has been initiated with units discussing two important aspects of small molecule–DNA binding: determination of the mode of interaction (UNIT 8.1) and determination of the binding constant (UNIT 8.2). The technique most widely used to determine binding constants is spectrophotometric titration. UNIT 8.2 describes the use of UV/vis spectroscopy to quantify ligand binding. Details are given for extracting the binding constant from the titration data using a Scatchand plot and nonlinear regression analysis. This method works successfully for those molecules whose UV/vis spectrum or fluorescence emission spectrum changes on binding to nucleic acids. Typically this includes molecules that bind by intercalation. Knowledge of the binding constant is necessary for determination of the binding mode as outlined in UNIT 8.1. This unit provides three complementary techniques that are indicative of an intercalative binding mode. This includes DNA lengthening by viscometry, assessment of DNA unwinding by plasmid viscometry, and changes in spectrophotometric properties of the ligand on binding. If the results of all three techniques are positive, then one can assume that the mode of binding is intercalation. Future units will deal with other modes of nucleic acid binding. In rational drug design, initial success depends in part on the availability of simple, rapid assays for activity that can be used to screen candidate drugs. UNIT 8.3 describes such a screen that was developed for assessing the selective binding of small molecules to particular nucleic acid structures (e.g., single-stranded, duplex, triplex molecules). This competition dialysis assay provides easily interpretable data in much less time than is required to obtain melting curves or binding constants. Molecules that bind to the minor groove of dsDNA can be conjugated to oligodeoxyribonucleotides. UNIT 8.4 describes various types of minor groove binders (MBs), their conjugation to oligodeoxyribonucleotides (ODNs), and binding modes of MB-ODN conjugates to either ssDNA or dsDNA. Characterization and applications of hybrids involving MB-ODN conjugates are also described. The feasibility of screening a library of different dsDNA sequences to establish binding specificity of a dsDNA binding molecule or to screen libraries of molecules for their ability to bind specific sequences would be significantly enhanced by the availability of high-throughput assays. Tse and Boger provide protocols for such an assay in UNIT 8.5. The authors describe a fluorescent intercalator displacement (FID) assay in 96-well-plate format that is both simple and fast. The assay utilizes hairpin oligonucleotides from Nucleic Acid Binding Molecules Contributed by Donald E. Bergstrom Current Protocols in Nucleic Acid Chemistry (2005) 8.0.1-8.0.2 C 2005 by John Wiley & Sons, Inc. Copyright
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which a bound intercalator (ethidium bromide or thiazole orange) can be displaced by an analyte. The decrease in fluorescence is directly related to the binding constant of the analyte. The binding constant for a particular dsDNA sequence and analyte pair can then be rapidly determined by an FID titration assay, which is also described. Donald E. Bergstrom
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Determination of Binding Mode: Intercalation
UNIT 8.1
This unit describes several techniques that can be used to establish the role of intercalation in the binding of small molecules to DNA. In very general terms, a molecule can be presumed to bind to DNA by intercalation between base pairs if it causes lengthening and unwinding of the DNA helix and undergoes changes in its spectral properties, such as DNA-induced hypochromism and quenching of its UV absorbance. DNA lengthening and unwinding can be determined from the change in viscosity of a solution of linear or plasmid DNA, respectively, upon addition of ligand (see Basic Protocols 1 and 2 and Alternate Protocols 1 and 2). Changes in the optical properties upon intercalation of the ligand can be monitored as described in Basic Protocol 3. A Support Protocol detailing the preparation of appropriately sized and purified DNA needed for Basic Protocols 1 and 3 and Alternate Protocol 1 is also included. If one of the above-mentioned criteria is not satisfied, then classical intercalation cannot be assumed. Ultimately, biochemical evidence for intercalation should be corroborated by nuclear magnetic resonance (NMR) or crystallographic studies. Of critical importance to these binding mode experiments is knowledge of the thermodynamics of binding of the molecule to DNA. The determination of binding constants is presented in UNIT 8.2.
DETERMINATION OF DNA LENGTHENING BY VISCOMETRY The mechanical properties of DNA require that the helix be lengthened to accommodate the positioning of a molecule between DNA base pairs. The simplest method to measure this lengthening is viscometry, as the viscosity of a DNA solution increases with the length of a DNA molecule. Using a capillary viscometer suspended in a constant-temperature water bath, the rate of flow of a solution of short DNA fragments is measured in the absence and presence of a candidate intercalator to determine the relative change in viscosity of the solution.
BASIC PROTOCOL 1
Materials 10% HNO3 10 mM buffer (Tris⋅Cl, HEPES, or other buffer) containing 1 mM EDTA, pH 7 (for TE buffer see APPENDIX 2A) 0.8 to 2.5 mM (base pairs) sonicated S1 nuclease-treated calf thymus DNA of ∼200-bp average length, in above buffer (see Support Protocol) Ligand of interest dissolved in above buffer Semi-micro capillary viscometer (Cannon-Ubehold or Cannon-Manning semi-micro type 75, see Fig. 8.1.1) 0.4-µm membrane filtration apparatus (e.g., Millipore Ultrafree MC or Centricon filters) Constant-temperature water bath, 25° ± 0.1°C Stopwatch (± 0.01 sec) or Wescan fiber-optic detection unit and timer (Wescan Instruments) Micropipettor with extension or extended microsyringe assembly (Hamilton or Stoelting) Contributed by Peter C. Dedon Current Protocols in Nucleic Acid Chemistry (2000) 8.1.1-8.1.13 Copyright © 2000 by John Wiley & Sons, Inc.
Nucleic Acid Binding Molecules
8.1.1
Figure 8.1.1 Cannon-Manning semi-micro viscometer. As described in Basic Protocol 1, solution is placed in tube A and suction is applied to tube B to draw solution above mark C. Flow time is determined from the time it takes for the meniscus to pass from mark C to mark D.
NOTE: All solutions should be passed through a 0.4-µm membrane filter to remove particulate matter that could clog the viscometer capillary and thus produce aberrant flow times. Determine buffer flow time 1. Clean the viscometer overnight with 10% HNO3, rinse thoroughly with deionized water, and dry prior to use. Place 1 mL of 10 mM buffer in a semi-micro capillary viscometer (tube A in Fig. 8.1.1) and equilibrate to 25°C for 5 min in a constant-temperature water bath. To conserve reagents, volumes as low as 750 mL can be used in certain viscometers, such as the Cannon 75C15. Be sure to use the same volume in all determinations. The viscometer should remain immersed in the water bath at all times.
2. Apply gentle suction to the other end of the viscometer (tube B in Fig. 8.1.1) with a pipet bulb and draw the buffer into the arm with the capillary until the meniscus lies above the upper marking (etched line) on the viscometer (mark C in Fig. 8.1.1). 3. Release the suction and start the stopwatch when the meniscus reaches the upper marking (mark C). Stop the stopwatch when the meniscus reaches the lower marking (mark D). Alternatively, the fiber-optic detection unit can be used to automatically determine flow times.
4. Perform several repetitions of the flow-time determination until the flow times fall within ±0.5 sec. Calculate the average flow time for at least three determinations. 5. Remove the buffer completely from the viscometer using a micropipettor with an extension or a long microsyringe to withdraw the remaining droplets. Determination of Binding Mode: Intercalation
Each solution must be completely removed from the viscometer prior to the addition of the next solution to prevent dilution of the DNA or ligand concentration.
8.1.2 Current Protocols in Nucleic Acid Chemistry
Optimize flow time for DNA solution 6. Prepare 1 mL of sonicated, S1 nuclease-treated calf thymus DNA at 1 mM base pairs in buffer. 7. Dispense the solution into the viscometer and equilibrate to 25°C in the water bath for 5 min. 8. Determine flow times as described in steps 2 to 4. Optimize the flow time by repeating this step with varying concentrations of DNA. The flow time should vary between 70 sec and 150 sec depending on the particular viscometer used. The acceptable range of DNA concentrations is 0.5 to 2.5 mM base pairs.
9. Prepare a 5- to 10-mL filtered stock of the final DNA solution. This will avoid small changes in viscosity due to pipetting errors.
Determine flow time without ligand 10. Place 1 mL of the appropriate DNA solution into the viscometer and equilibrate to 25°C. 11. Perform several repetitions of the flow-time determination (steps 2 to 4) until the flow times fall within ±0.5 sec. Calculate the average flow time for at least three determinations. Determine flow time in the presence of ligand 12. Add a small, filtered aliquot of the ligand dissolved in buffer (1 to 2 µL) to the DNA solution in the viscometer and mix the solution by repeated gentle pipetting or by gently bubbling nitrogen through the solution. Equilibrate to 25°C for 5 min. It is best to start with a ligand concentration that results in 3 to 10 bound ligand molecules per 100 base pairs of DNA. This concentration can be calculated from the binding constant determined as described in UNIT 8.2. Alternatively, a broad range of ligand concentrations can be examined; however, this is time consuming. Particulate matter should be removed from the ligand solution by filtration.
13. Determine the average flow time as in steps 2 to 4. 14. Repeat steps 12 and 13 until the total added volume of ligand is no more than 15 µL. The small dilution of DNA by added ligand will be taken into account in the calculation of relative viscosities described shortly.
15. If additional ligand concentrations are required, rinse the viscometer with deionized water, dry, and start with a more concentrated ligand stock and a fresh 1-mL volume of DNA solution. Repeat steps 10 and 11, and use this flow time in subsequent calculations of relative viscosity changes with the new ligand concentrations. Calculate relative viscosities and analyze data 16. Calculate the specific viscosity of the DNA solution without added ligand (η0) using the flow times of buffer without (t0) and with (t) DNA and the equation η0 = (t − t0)/t0. 17. Calculate the specific viscosity of the DNA solution with added ligand (η) using the flow times of buffer alone (t0) and buffer with DNA plus ligand (tl) and the equation η = (tl − t0)/t0. Correct the flow times of samples diluted by serial addition of ligand by multiplying the value by the degree of dilution. 18. Plot (η/η0)1/3, which is the cubed root of the relative viscosity, as a function of the moles of ligand bound per base pair of DNA (r).
Nucleic Acid Binding Molecules
8.1.3 Current Protocols in Nucleic Acid Chemistry
Figure 8.1.2 Viscometry data for ethidium bromide binding to sonicated calf thymus DNA. According to the theory of Cohen and Eisenberg (1969), a plot of the cubed root of the specific relative viscosity versus r should yield a slope of 1 for a classical intercalator. The slope of 1.07 determined for ethidium bromide is in reasonable agreement with this value.
The latter value is calculated from the binding constant determined in UNIT 8.2. See Commentary for a discussion of the interpretation of this plot in terms of intercalation. An example of such a plot for the binding of ethidium bromide is shown in Figure 8.1.2. ALTERNATE PROTOCOL 1
DETERMINATION OF DNA LENGTHENING BY VISCOMETRY FOR LIGANDS SOLUBLE IN ORGANIC SOLVENTS DNA-binding ligands are often water insoluble, which necessitates addition of small amounts of organic solvent to DNA solutions. However, organic solvents significantly affect the viscosity of the DNA solution. To avoid solvent anomalies, viscosity studies, in this case, must be performed at a constant concentration of organic solvent and each ligand concentration must be assessed independent of other concentrations; serial addition of aliquots of the ligand solution will affect the total solvent concentration and thus the viscosity. Additionally, some organic solvents have a tendency to produce air bubbles in aqueous solutions, which will produce anomalous flow times. This problem is solved by degassing the solutions prior to viscometric studies; volatile solvents should be added after the buffer is degassed. Additional Materials (also see Basic Protocol 1) Ligand of interest dissolved in an organic solvent that is miscible with water Organic solvent Microcentrifuge tubes Small vacuum desiccator with water aspirator Micro stir bars to fit in microcentrifuge tubes Magnetic stirring plate
Determination of Binding Mode: Intercalation
Degas solutions 1. In microcentrifuge tubes, prepare 1-mL solutions of buffer, DNA, and DNA with each concentration of ligand to be tested, making sure that each solution has the same final concentration of organic solvent.
8.1.4 Current Protocols in Nucleic Acid Chemistry
If a volatile solvent must be used, degas the aqueous solutions first (steps 1 to 3) and then add solvent (with or without ligand) after degassing. The solvent concentration should be the same as that used in the determination of the DNA binding constant for the ligand (UNIT 8.2). This permits calculation of the proportion of DNA-bound ligand during the viscosity measurements. In addition, the DNA and organic solvent concentrations in the samples must be carefully controlled to ensure meaningful results. This can be accomplished by preparing one large DNA stock solution and dividing it into aliquots for each ligand concentration, followed by addition of ligand in solvent. Positive-displacement pipettors can be used to ensure that precise quantities of organic solvent are delivered to the solutions.
2. Place a micro stir bar in each tube and place the tubes, with lids open, upright in a small beaker with a piece of tissue to support the tubes. 3. Place the beaker in a vacuum desiccator atop a magnetic stirring plate and, while stirring, apply a vacuum for 5 to 10 min. A water aspirator will supply adequate vacuum to degas the solutions. The solutions must be stirred to prevent bumping of the contents during evacuation.
Determine flow times and analyze the viscosity data 4. Optimize flow times for the DNA solution without ligand as described (see Basic Protocol 1, steps 6 to 9). 5. Determine flow times of buffer, DNA alone, and DNA with ligand as described (see Basic Protocol 1, steps 1 to 5 and 10 to 15), but replace each solution with the next higher ligand concentration instead of adding additional ligand to increase the concentration. There is likely to be greater scatter in the viscosity data with this protocol due to the necessity of using separate DNA solutions for each ligand concentration. This problem is controlled by performing multiple determinations for each ligand concentration.
6. Calculate relative viscosities as described (see Basic Protocol 1, steps 16 to 18). ASSESSMENT OF DNA UNWINDING BY PLASMID VISCOMETRY The second hallmark of intercalation is unwinding of the DNA helix to accommodate the intercalating ligand. Among the several techniques available to assess helical unwinding, plasmid viscometry is perhaps the easiest. The technique is based on changes in DNA topology, and thus the viscosity of the DNA solution, when an intercalator binds to closed circular plasmid DNA. Knowledge of the DNA binding constant (UNIT 8.2) under the conditions of the plasmid viscometry allows direct calculation of the angle of unwinding for each bound intercalator. The unwinding angle can also be calculated by comparison to ethidium bromide without knowledge of the binding constant. Both analyses are described below. Additionally, the same binding constant can be applied to all of the studies in this unit if identical buffer, solvent, and temperature conditions are employed. Materials DNA: Phage DNA M13mp, PM2, or ∅X174; plasmids COL E1 or pBR325; or any similarly sized plasmid (5 to 7 kbp); each prepared by standard alkaline lysis (e.g., CPMB UNIT 1.7) or similar procedures, and suspended at 1 mg/mL in TE (APPENDIX 2A) or other buffer 10 mM buffer (Tris⋅Cl, HEPES, or other buffer) containing 1 mM EDTA, pH 7 (for TE buffer see APPENDIX 2A) Ligand of interest dissolved in above buffer (~1 mM) 1 to 2 mM ethidium bromide solution (see recipe)
BASIC PROTOCOL 2
Nucleic Acid Binding Molecules
8.1.5 Current Protocols in Nucleic Acid Chemistry
Semi-micro capillary viscometer (Cannon-Ubehold or Cannon-Manning semi-micro type 75, see Fig. 8.1.1) 0.4-µm membrane filtration apparatus (e.g., Millipore Ultrafree MC or Centricon filters) Constant-temperature water bath, 25° ± 0.1°C Stopwatch (± 0.01 sec) or Wescan fiber-optic detection unit and timer (Wescan Instruments) Micropipettor with extension or extended microsyringe assembly (Hamilton or Stoelting) CAUTION: Ethidium bromide is a mutagen and an environmental hazard. It should be handled carefully with gloves and disposed of properly. Methods of disposal may vary between institutions. Consult with the institution’s environmental safety office for the preferred means of storage and disposal of ethidium bromide. Determine flow times 1. Determine the average flow time for a 1-mL solution of DNA at a concentration of 0.1 to 0.6 mM nucleotides (30 to 200 µg/mL) as described (see Basic Protocol 1, steps 6 to 11). The choice of DNA concentrations depends on the dissociation constant of the ligand. For dissociation constants in the micromolar range, a DNA concentration 0.2 to 0.6 mM (nucleotides) ensures virtually complete binding of the ligand, thus obviating the need to determine the quantity of free and bound ligand.
2. Add the ligand to the DNA solution in the viscometer in 5- to 10-µL increments and mix the solution by repeated pipetting or by bubbling nitrogen through the solution. The incremental addition of ligand to the DNA solution will produce minor alterations in the viscosity of the solution compared to the changes in DNA topology caused by ligand binding. However, the dilution must be accounted for in the calculation of the unwinding angle, which are based on actual ligand and DNA concentrations.
3. Following temperature equilibration, determine the average flow time (see Basic Protocol 1, steps 2 to 4) of the solution for each addition of ligand. Additions of ligand should continue well past the point at which the viscosity reaches a maximum; this indicates complete relaxation of the supercoiled plasmid. Further addition of the intercalating ligand should result in positive supercoiling and a reduction in viscosity. If there is a bell-shaped dependence of viscosity (flow time) on ligand concentration, then the ligand may be assumed to cause unwinding of the DNA helix. If no change in viscosity is observed after ten aliquots of ligand are added, then the ligand either binds with low affinity or with a small unwinding angle (i.e., DNA unwinding is minimal); or, less likely, binds with very high affinity or with a very large unwinding angle (i.e., the first aliquot of ligand produced maximal positive supercoiling). To overcome this problem, increase (or decrease) the ligand concentration 5-fold and repeat steps 1-3 until a bell-shaped curve is obtained.
Determine the unwinding angle
Determination of Binding Mode: Intercalation
With prior knowledge of the binding constant: 4a. Repeat the flow-time determinations for a serial addition of ethidium bromide to a fresh 1-mL DNA solution and determine the concentration of ethidium bromide that produces maximal relaxation of the plasmid DNA (maximal flow time).
8.1.6 Current Protocols in Nucleic Acid Chemistry
Ethidium bromide serves as a standard for helical unwinding, with each bound molecule unwinding the helix by ∼26°. As a starting point, ethidium bromide should produce full relaxation and positive supercoiling over a concentration range of 10 to 40 µM with a 6to 7-kbp negatively supercoiled plasmid at ∼0.3 mM base pairs concentration.
5a. Calculate the molar binding ratio (ν; moles of bound ligand per mole of nucleotides) under conditions of maximal relaxation for both ethidium bromide and ligand. Calculate the concentration of bound ligand using the binding constant and equation for the neighbor exclusion model discussed in UNIT 8.2. 6a. Calculate the unwinding angle (relative to ethidium bromide) from the equation ∅L = ∅Et(νEt/νL), where ∅L is the unwinding angle for the ligand, ∅Et is the known 26° unwinding angle for ethidium bromide, and νEt and νL are the molar binding ratios for ethidium bromide and ligand, respectively, at concentrations producing maximal plasmid relaxation. Without prior knowledge of the binding constant: 4b. Determine the concentration of ethidium bromide and ligand that produce maximal relaxation (maximal flow times) for four to five different DNA concentrations covering a four- to five-fold range (e.g., 0.5 to 2.5 mM nucleotides). 5b. Plot the total concentration of ligand (CT) versus the total DNA concentration (NT) according to the equation CT = νNT + CF, where the slope (ν) is the molar binding ratio (moles of ligand bound per mole of nucleotides) and the y intercept (CF) is the concentration of free ligand. 6b. Calculate the unwinding angle (relative to ethidium bromide) from the equation: ∅L = ∅Et(νEt/νL), where ∅L is the unwinding angle for the ligand, ∅Et is the known 26° unwinding angle for ethidium bromide, and νEt and νL are the molar binding ratios determined from the slopes of the plots in step 5b. ASSESSMENT OF DNA UNWINDING BY PLASMID VISCOMETRY FOR LIGANDS SOLUBLE IN ORGANIC SOLVENTS
ALTERNATE PROTOCOL 2
The problems associated with studying ligands in the presence of organic solvents are identical to those discussed in Alternate Protocol 1 for DNA lengthening determined by viscometry. To avoid changes in organic solvent concentration associated with incremental addition of a ligand dissolved in the solvent, the viscosity studies must be performed at a constant concentration of organic solvent and each ligand concentration must be assessed independent of other concentrations. The materials and procedures for plasmid viscometric studies with ligands in organic solvent are exactly as described earlier. The flow time for each ligand concentration must be determined in a fresh DNA solution. Large quantities of DNA (∼10 mg) are required for determination of the unwinding angle if the binding constant is not known because of the need to perform flow-time determinations for five to ten ligand and ethidium bromide concentrations at four to five different DNA concentrations. ASSESSMENT OF CHANGES IN THE OPTICAL PROPERTIES OF LIGANDS UPON BINDING TO DNA The third criterion for intercalative DNA binding is a change in the optical properties of the DNA-bound chromophore. The UV/visible absorbance of classical DNA intercalators is typically reduced by 40% to 60% when the chromophore inserts between DNA base pairs. Furthermore, there is often a shift in the absorbance maximum to longer wavelengths (i.e., a red shift) as the chromophore interacts with adjacent DNA bases. The
BASIC PROTOCOL 3
Nucleic Acid Binding Molecules
8.1.7 Current Protocols in Nucleic Acid Chemistry
following protocol assumes that the molar extinction (i.e., extinction coefficient) for the ligand can be determined and that the absorbance maximum (λmax) of the ligand is significantly different from that of DNA (>300 nm). Materials Sonicated, S1 nuclease–treated calf thymus DNA (see Support Protocol) 10 mM buffer (Tris⋅Cl, HEPES, or other buffer) containing 1 mM EDTA, pH 7 (for TE buffer see APPENDIX 2A) Ligand of interest dissolved in above buffer Digital or double-beam UV/visible spectrophotometer capable of recording absorbance spectra Determine extinction coefficient of ligand in the absence of DNA 1. Prepare five 1-mL solutions of ligand at 1, 3, 10, 30, and 100 µM in the desired buffer. Since the molar extinction varies as a function of factors such as ionic strength and pH, these studies should be performed in the same buffer used in the viscometric studies and in the binding constant studies described in UNIT 8.2. The wide range of ligand concentrations ensures that several concentrations will produce an absorbance value between 0.01 and 1 absorbance unit (AU).
2. Blank a spectrophotometer against the buffer either digitally or by placing the buffer in one of the compartments of a double-beam instrument. 3. Place a cuvette containing the first ligand solution into the spectrophotometer and record the spectrum from 200 to 800 nm. 4. Record spectra for each of the ligand concentrations. 5. For spectra containing absorbance peaks in the range of 0.01 to 1 AU, note the wavelength and exact absorbance for a major peak or shoulder that lies above the absorbance of DNA (>300 nm). 6. Calculate the extinction coefficient (ε) at this wavelength from the Beer-Lambert law: A = εlc, where A is the absorbance value, l is the cuvette path length in cm, and c is the molar ligand concentration. The extinction coefficient is valid only under the conditions used in its determination (e.g., buffer type, pH, ionic strength, temperature, and so on). The coefficient should be calculated from triplicate determinations of at least three different concentrations.
Determine extinction coefficient of ligand in the presence of DNA 7. Prepare four solutions each containing (1) a ligand concentration that will produce an absorbance of ∼0.2 to 0.3 and (2) a DNA concentration (nucleotides) that is 3-, 10-, 30-, and 100-fold greater than the ligand concentration. Also prepare the same DNA solutions without ligand. It is important to use S1 nuclease–treated DNA to avoid possible problems with spectral changes upon binding to single-stranded DNA.
8. Blank the spectrophotometer against one of the DNA solutions without ligand. Determination of Binding Mode: Intercalation
9. Record the absorbance spectrum of the ligand-containing solution with the same DNA concentration. 10. Repeat steps 8 and 9 for the other DNA concentrations.
8.1.8 Current Protocols in Nucleic Acid Chemistry
11. At the wavelength used for determining the extinction coefficient of unbound ligand in step 6, plot the absorbance for each ligand-containing solution versus the DNA concentration (in nucleotides) and determine the minimum absorbance value. As the DNA concentration increases, the ligand absorbance should decrease to a minimum value indicative of complete ligand binding. There should be at least two DNA concentrations that produce the minimum absorbance value. If not, repeat steps 8 and 9 with a higher DNA concentration.
12. Use the minimum absorbance value to calculate the extinction coefficient for the DNA-bound ligand, as in step 6. For a ligand with an association constant greater than ~104 M–1, a 100-fold excess of DNA should result in the minimum absorbance indicative of complete ligand binding
Determine change in absorbance with DNA 13. Calculate the ratio of extinction coefficients in the absence and presence of DNA. A classical intercalator will typically experience a 40% to 60% reduction in absorbance upon binding to DNA. A shift in absorbance maximum of the ligand upon binding to DNA can be calculated by obtaining the absorbance spectrum of the ligand without DNA and in the presence of enough DNA to cause complete binding of the ligand.
PREPARATION OF GENOMIC DNA FRAGMENTS FOR VISCOMETRY To avoid possible complications from ligand-induced changes in DNA flexibility, the viscometry studies in Basic Protocol 1 and Alternate Protocol 1 should be performed with DNA fragments roughly 200 to 250 bp long. DNA of this length can be prepared by sonication of calf thymus DNA and size fractionation by gel-filtration chromatography. This protocol is also used to prepare DNA needed for the optical studies in Basic Protocol 3.
SUPPORT PROTOCOL
Materials Calf thymus DNA (sodium salt) Phosphate/EDTA buffer: 50 mM sodium phosphate with 1 mM EDTA, pH 7 Nitrogen gas Gel loading buffer (APPENDIX 2A) 1% (w/v) agarose gel in TBE buffer TBE buffer (APPENDIX 2A) DNA size markers, 100 to 1000 bp (e.g., HaeIII-digested ∅X174 DNA) 10 mM sodium acetate, pH 5/100 mM NaCl/30 µM ZnCl2 S1 nuclease Buffered phenol or 1:1 (v/v) phenol/chloroform (APPENDIX 2A) 3 M sodium acetate, pH 7 100%, 95%, and 70% (v/v) ethanol TE buffer, pH 7.5 (APPENDIX 2A) 10 mg/mL DNase-free RNase A (see recipe) 20 mg/mL proteinase K in water (store in single-use aliquots at −20°C) 24:1 (v/v) chloroform/isoamyl alcohol 1 µg/mL ethidium bromide solution (see recipe) Sonicator (e.g., Branson model 450) 3000- to 10,000-MWCO dialysis tubing 3 × 60–cm column of Sepharose 4B or Sephacryl C-500-HR
Nucleic Acid Binding Molecules
8.1.9 Current Protocols in Nucleic Acid Chemistry
UV transilluminator Additional reagents and equipment for agarose gel electrophoresis (e.g., CPMB UNIT 2.5A) and phenol/chloroform extraction (APPENDIX 2A) Sonicate DNA to produce short fragments 1. Dissolve 100 mg calf thymus DNA in 50 mL phosphate/EDTA buffer by stirring overnight at 4°C. 2. Place the beaker containing the DNA solution on ice and gently bubble nitrogen through the solution for 5 min to remove oxygen. The presence of nitrogen and removal of oxygen reduces the size of DNA fragments generated in subsequent sonication.
For lengthening/unwinding studies: 3. Sonicate the DNA for a total of 30 min in 5-min intervals at full power using the 0.5-in. end of a disrupter horn (for a Branson model 450 or similar sonicator) with 5-min cooling periods between sonication periods. Gently bubble nitrogen through the solution during the cooling periods to remove oxygen. For smaller volume (<50 mL), a microprobe sonicator tip can be used. However, to avoid damage to the probe, the energy setting must be reduced according to the manufacturer’s specifications. For the spectral studies of Basic Protocol 3, sonicate as in step 3 but reduce the time from 30 min to 10 to 15 min and proceed with step 6.
4. Assess the size of DNA fragments by removing 1 µg of DNA from the sonication mixture, mixing with gel loading buffer, and resolving the DNA on a 1% agarose gel in TBE buffer until the bromophenol blue dye has migrated approximately one-half the length of the gel. Include a lane of DNA size markers covering the range of 100 to 1000 bp. This ensures that an adequate quantity of 200- to 250-bp DNA fragments is present in the mixture of sheared DNA fragments.
5. If the fragment sizes are too large, sonicate the mixture for another 10 to 15 min and repeat the gel analysis. Remove single-stranded DNA: 6. Dialyze the DNA three times against 4 L (each time) of 10 mM sodium acetate, pH 5/100 mM NaCl/30 µM ZnCl2 at 4°C. Use 3000- to 10,000-MWCO dialysis tubing and perform at least one of the dialysis steps overnight. 7. Digest the single-stranded DNA with S1 nuclease (20 IU) 1 hr at 37 °C. Extract the DNA several times with phenol and chloroform (see APPENDIX 2A). 8. Add 3 M sodium acetate to 0.3 M and 2 vol of 100% ethanol to precipitate. Centrifuge 15 min at 16,000 × g. Wash the DNA twice with 70% ethanol and once with 95% ethanol. Determination of Binding Mode: Intercalation
9. Dry the DNA either by vacuum centrifugation or by overnight evaporation on a bench top. Redissolve the dried DNA pellet in 50 mL TE buffer and proceed with step 7. The dialysis can be performed after the sonication step.
8.1.10 Current Protocols in Nucleic Acid Chemistry
Purify DNA 10. Remove the DNA from the dialysis tubing and add sodium acetate to 300 mM followed by DNase-free RNase to 0.1 mg/mL. Incubate 1 hr at 37°C. 11. Add proteinase K to 0.1 mg/mL and incubate 2 hr at 37°C. 12. Extract the solution four times with an equal volume of buffered phenol or 1:1 (v/v) phenol/chloroform to remove proteins ( APPENDIX 2A), and twice with 24:1 (v/v) chloroform/isoamyl alcohol to remove residual phenol. 13. Precipitate the DNA by adding 2 vol of 100% ethanol and centrifuging 15 min at 16,000 × g, 4°C. 14. Wash the DNA twice with 70% ethanol and once with 95% ethanol. Dry the DNA either by vacuum centrifugation or by overnight evaporation on a bench top. Size fractionate DNA 15. Redissolve the DNA in phosphate/EDTA buffer at 4 mg/mL and apply half the solution to a 3 × 60–cm Sepharose 4B or Sephacryl S-500-HR gel-filtration column to resolve the DNA fragments by size. The other half of the DNA solution can be stored at −20°C for later chromatographic purification. The most convenient means to monitor the elution of DNA from the column is to use a complete liquid chromatographic system with a peristaltic pump, UV detector (254 nm), chart recorder, and fraction collector.
16. Mix a 5- to 10-µL aliquot of each fraction with gel loading buffer and resolve the DNA on a 1% agarose gel as in step 4a. The aliquot should contain 50 to 300 ng of DNA based on absorbance at 260 nm, assuming that 1 A260 = 50 µg/mL DNA.
17. Stain the gel with 1 µg/mL ethidium bromide solution and visualize the DNA fragments by UV illumination. CAUTION: Ethidium bromide is a mutagen and environmental hazard. It should be handled carefully with gloves and disposed of properly. Methods of disposal may vary between institutions. Consult with the institution’s environmental safety office for the preferred means of storage and disposal.
18. Identify the fractions containing DNA with an average size of ∼200 bp, pool these fractions, and dialyze the solution three times against 4 L of TE buffer at 4°C with at least one overnight dialysis step. 19. Add 0.1 vol of 3 M sodium acetate and precipitate the DNA with 2 vol of 100% ethanol by centrifuging 30 min at 16,000 × g, 4°C. 20. Wash and dry the pellet as in step 14. 21. Dissolve the DNA in the desired buffer (see Basic Protocol 1 and Basic Protocol 3) at 3 mg/mL and store at −20°C. The DNA can be stored several months at –20°C. If the DNA is to be kept for long periods, it can be stored in screw-cap tubes at –80°C for at least a year.
Nucleic Acid Binding Molecules
8.1.11 Current Protocols in Nucleic Acid Chemistry
REAGENTS AND SOLUTIONS Use deionized water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
DNase-free RNase A, 10 mg/mL Dissolve DNase-free RNase A in TE buffer (APPENDIX 2A) at 1 mg/mL and boil 10 to 30 min. Store in aliquots at −20°C to prevent microbial growth. For additional details on this enzyme, see CPMB UNIT 3.13. Ethidium bromide solution, 1 to 10 mg/mL Prepare a 1 to 10 mg/mL stock solution of ethidium bromide in water. Determine the exact concentration of ethidium bromide in this crude stock by measuring its absorbance at 480 nm and using the molar extinction coefficient (ε480) = 5600 M−1cm−1. Store for up to 6 months at 4°C protected from light. For assessment of DNA unwinding (see Basic Protocol 2), dilute stock to 1 to 2 mM in the desired buffer. For staining agarose gels (see Support Protocol), dilute to 1 µg/mL in TBE buffer (APPENDIX 2A). COMMENTARY Background Information The three sets of experiments presented in this unit are designed to define the ability of a DNA-binding ligand to intercalate between the base pairs of DNA. If a ligand increases the viscosity of a solution of linear DNA fragments (Lerman, 1961; Waring, 1970), produces a bellshaped viscosity profile with plasmid DNA (Révet et al., 1971), and undergoes DNA-induced quenching of its absorbance, then it can be presumed to bind to DNA by intercalation. Alone, the individual observations are insufficient to establish intercalation since there are exceptions to each. For example, cisplatin, a DNA cross-linker that binds to the N7 position of guanine, and irehdiamine, a steroidal groove-binder, both unwind supercoiled plasmid DNA (Saucier 1977; Cohen et al., 1979), but neither causes an increase in the viscosity of a DNA solution. The reader is referred to the work of Satyanarayana et al. (1993) for a rigorous comparison of the viscometric and spectral behaviors of several groove binders and intercalators. This biophysical evidence for intercalation is best corroborated by NMR or X-ray crystallographic studies.
Critical Parameters
Determination of Binding Mode: Intercalation
Several parameters must be controlled to avoid erroneous results. First, the viscometer should be cleaned after each use by overnight soaking in 10% to 50% nitric acid, followed by thorough rinsing with deionized water and drying. Particulate matter present in all of the solutions should be removed by filtration through a ≥0.4-µm filter.
Second, the temperature, ionic composition, pH, and concentration of organic solvents must be precisely controlled in viscosity studies. Small changes in organic solvent concentration, for example, can significantly alter flow times in the viscometer. These same conditions must be employed for the determination of binding constants discussed in UNIT 8.2, since the thermodynamics of ligand binding vary according to the properties of the solution. Third, it is important to check for changes in flow time of the buffer and control DNA solutions regularly throughout the viscometric studies. This ensures that there are no anomalies caused by residual solvents or components sticking to the walls of the viscometer capillary. Finally, the length of the linear DNA fragments must be short enough to avoid problems arising from ligand-induced changes in the flexibility of the DNA. The use of ∼200-bp DNA fragments, roughly the persistence length of DNA, prevents problems with flexibility. Significantly shorter fragments (<50 bp) will produce unacceptably short flow times.
Troubleshooting The most likely cause of highly variable flow times is a dirty or clogged viscometer. Regular cleaning of the viscometer and filtration of the solutions will prevent this problem. The presence of small air bubbles in the viscometer capillary can be avoided by degassing the solutions, especially those containing nonvolatile organic solvents. Care must be taken to avoid precipitation of the DNA fragments that can be caused by extremely high concentrations
8.1.12 Current Protocols in Nucleic Acid Chemistry
of some ligands as indicated by visible precipitates or a sudden decrease in flow time. Furthermore, high concentrations of water-insoluble ligands can result in aggregation and precipitation of the ligand, which may also create problems. One way to check for viscosity changes due to ligand insolubility is to perform flow-time measurements with high concentrations of the ligand in buffer alone. Although flow times will depend on the type of viscometer used, excessively long flow times (>180 sec) can be avoided by reducing the DNA concentration.
Anticipated Results The results of the plasmid viscosity experiments are self explanatory: if the viscosity of the DNA solution increases and then decreases in a bell-shaped curve as a function of ligand concentration, then the ligand can be presumed to unwind the DNA helix. For the DNA lengthening studies, the theory of Cohen and Eisenberg (1969) predicts that a plot of (η/η0)1/3 versus r (moles of ligand bound per mole of DNA base pairs) will give a slope ≈ 1 for a “classical” intercalator (for example, see Fig. 8.1.2). This result is based on the predicted 3.4-Å increase in the length of the DNA helix when an intercalator is positioned between DNA base pairs. Most classical intercalators produce a slope lying between 0.5 and 1 in such a plot (Wilson and Jones, 1982), while “classical” groove binders such as Hoechst 33258 produce a slope of zero (Satyanarayana et al., 1993). With regard to changes in ligand absorbance upon binding to DNA, the results can be highly variable. Classical intercalators, as well as many groove-binding ligands, will show a reduced absorbance of light when bound to DNA. However, a shift to longer wavelengths does not always accompany intercalation, as illustrated by the binding of the intercalating enediyne esperamicin A1 to DNA (Yu et al., 1994).
organic solvents, and the experience of the researcher. At least 1 week of work should be anticipated to complete each protocol. Although the preparation of DNA fragments and plasmid DNA will require additional time, sufficiently large quantities of these reagents can be prepared at one time for use in many different studies.
Literature Cited Cohen, G. and Eisenberg, H. 1969. Viscosity and sedimentation study of sonicated DNA-proflavine complexes. Biopolymers 8:45-55. Cohen, G.L., Bauer, W.R., Barton, J.K., and Lippard, S.J. 1979. Binding of cis- and trans-dichlorodiammineplatinum(II) to DNA: Evidence for unwinding and shortening of the double helix. Science 203:1014-1016. Lerman, L.S. 1961. Structural considerations in the interaction of DNA and acridines. J. Mol. Biol. 3:18-30. Révet, B.M., Schmir, M., and Vinograd, J. 1971. Direct determination of the superhelix density of closed circular DNA by viscometric titration. Nature 229:10-13. Satyanarayana, S., Dabrowiak, J.C., and Chaires, J.B. 1993. Tris(phenanthroline)ruthenium(II) enantiomer interactions with DNA: Mode and specificity of binding. Biochemistry 32:25732584. Saucier, J.-M. 1977. Physicochemical studies on the interaction of irehdiamine A with bihelical DNA. Biochemistry 16:5879-5889. Waring, M. 1970. Variation in the supercoils in closed circular DNA by binding of antibiotics and drugs: Evidence for molecular models involving intercalation. J. Mol. Biol. 54:247-279. Wilson, W.D. and Jones, R.L. 1982. Intercalation in biological systems. In Intercalation Chemistry (M.S. Whittingham and A.J. Jacobson, eds.) pp. 445-501. Academic Press, New York. Yu, L., Golik, J., Harrison, R., and Dedon, P. 1994. The deoxyfucose-anthranilate of esperamicin A1 confers intercalative DNA binding and causes a switch in the chemistry of bistranded DNA lesions. J. Am. Chem. Soc. 116:9733-9738.
Time Considerations The time to complete the viscosity studies is highly variable and depends on the number of ligand concentrations studied, the need for
Contributed by Peter C. Dedon Massachusetts Institute of Technology Cambridge, Massachusetts
Nucleic Acid Binding Molecules
8.1.13 Current Protocols in Nucleic Acid Chemistry
Determination of Binding Thermodynamics
UNIT 8.2
This unit serves as a starting point for exploring the thermodynamic properties of small molecule–DNA interactions, in theory and practice. The treatment of thermodynamics here will be limited to determination of simple, apparent association/dissociation constants, a necessary limitation due to the complexity of DNA as a “receptor” and the variety of different mechanisms by which small molecules recognize DNA sequence, structure, and dynamics. The spectrum of drug-DNA interactions ranges from sequence specific to sequence nonselective. For example, the intercalator ethidium bromide is relatively sequence nonselective in its binding to DNA compared to the enediyne calicheamicin, which binds to the 3′ ends of purine tracts. For both molecules, the DNA-binding affinity can be determined on a global scale for a mixture of DNA sequences with a spectrum of different binding sites. This type of thermodynamic study is useful for defining general modes of binding such as intercalation, as described in UNIT 8.1, or as a first step in structure-function studies with ligand congeners. However, the model used to calculate binding constants must take into account the fact that there is a spectrum of binding sites with differing affinities for the ligand, and that binding of a ligand molecule to one site may influence subsequent binding of other ligand molecules (i.e., cooperative interactions). The basic approach described in this unit consists of data gathering and curve fitting. The first step in all cases is to determine, under a defined set of conditions, both the concentration of DNA-bound ligand and the concentration of ligand “free” in solution. This information can be obtained by any technique that measures a change in some property of the ligand or DNA that occurs upon binding—including UV/visible (UV/vis) and fluorescence spectroscopy, circular dichroism, and DNA cleavage analysis—or by a technique that allows separation of bound and unbound ligand, such as equilibrium dialysis, centrifugation, or solvent partitioning. Of necessity, this unit will focus on a single method, spectroscopic titration, to quantify bound and unbound ligand. A binding constant is then extracted from the data using any of several mathematical models for DNA-ligand thermodynamics, including the neighbor-exclusion models of McGhee and von Hippel. SPECTROSCOPIC TITRATION OF FIXED DNA CONCENTRATIONS WITH VARYING LIGAND CONCENTRATIONS
BASIC PROTOCOL
There are several applications in which ligand binding thermodynamics must be determined in solutions containing many copies of a long (>50 bp) “random” DNA sequences, such as in the intercalation studies of UNIT 8.1 with plasmid DNA or sonicated calf thymus DNA. In these cases, the calculation of apparent binding constants is performed with mathematical models that take into account the possibility of cooperative interactions and the presence of many sites with differing affinities. The binding of a ligand to DNA can be assessed by UV/vis or fluorescence spectroscopy if the ligand possesses optical properties that change upon binding to DNA. Furthermore, the optical properties must differ from those of DNA, with significant absorbance at wavelengths >300 nm. Table 8.2.1 contains examples of the optical properties of two intercalating ligands, ethidium bromide and daunomycin, that can be exploited to determine DNA binding thermodynamics. In this protocol, varying ligand concentrations are added to a fixed concentration of DNA and the absorbance changes are recorded to determine the degree of ligand binding. Nucleic Acid Binding Molecules Contributed by Peter C. Dedon Current Protocols in Nucleic Acid Chemistry (2000) 8.2.1-8.2.8 Copyright © 2000 by John Wiley & Sons, Inc.
8.2.1
Table 8.2.1
Optical Properties of Ethidium Bromide and Daunomycin
Property
Ethidium bromidea
Daunomycinb
λmax free λmax bound ε480 free ε480 bound Isosbestic pointsc Relative fluorescenced
479 nm 517 nm 5600-5800 M−1cm−1 ∼2500 M−1cm−1 390 and 510 nm free = 1; bound ∼15
480 nm 505 nm 11500 M−1cm−1 7000 M−1cm−1 540 nm (ε = 5100 M−1cm−1) free = 1; bound = 0.05
aData from Waring, 1965; Hinton and Bode, 1975; Chaires et al., 1982. bData from Chaires et al., 1982; conditions: 20°C, 6 mM Na HPO , 2 mM NaH PO , 1 mM Na EDTA, 185 mM NaCl, 2 4 2 4 2
pH 7; calf thymus DNA. cIsosbestic point: free and bound ligand share the same absorbance at this wavelength. dFor daunomycin, λ = 555 nm and λ = 480 nm; for ethidium bromide, λ = 605 nm and λ = 525 nm. Values apply em ex em ex
to unbound ligand.
Materials Assay buffer: 10 mM Tris⋅Cl, HEPES, or other buffer, pH 7, containing 1 mM EDTA Calf thymus DNA, sonicated (see recipe) Ligand of interest dissolved in assay buffer UV/vis spectrophotometer with temperature-controlled cuvette holder 1-mL cuvette Software capable of linear regression analysis (e.g., Microsoft Excel) Additional reagents and equipment for determination of extinction coefficients of free and DNA-bound ligands (UNIT 8.1, Basic Protocol 3) Determine extinction coefficients of bound and unbound ligand 1. Using a UV/vis spectrophotometer with temperature-controlled cuvette holder and a 1-mL cuvette, determine the extinction coefficient of the ligand free in solution and fully bound to DNA as described in UNIT 8.1, Basic Protocol 3. 2. Determine if an isosbestic point exists by examining the absorbance spectra obtained in step 1 for a wavelength at which the absorbance is the same in the presence and absence of DNA. Determine the extinction coefficient at the isosbestic wavelength for a least three different DNA and ligand concentrations (e.g., vary each by a factor of three). This value allows calculation of total ligand concentration in the presence of any concentration of DNA.
Perform spectral titrations 3. Blank the spectrophotometer against 1 mL of a 1 mM DNA solution (molarity calculated as base pairs). 4. Add 1 µL of a 10 mM solution of the ligand and mix thoroughly.
Determination of Binding Thermodynamics
For ligands with extinction coefficients >20,000 M−1cm−1, a 10 mM ligand concentration should provide an absorbance of >0.2, which is adequate for the titration. For ligands with extinction coefficients <20,000 M−1cm−1, start with higher ligand concentrations (e.g., 50 mM). This will ensure an adequate absorbance value for the ligand. Mixing is accomplished by repeated pipetting or, ideally, on a continuous basis with a stirring bar in spectrophotometers equipped with a magnetic stirrer.
8.2.2 Current Protocols in Nucleic Acid Chemistry
5. Record the absorbance at the λmax of the ligand or perform a wavelength scan that encompasses the λmax. If there is an isosbestic point associated with ligand binding to DNA, then record the absorbance at this wavelength as well.
6. Repeat steps 4 and 5 for each of three additional 1-µL aliquots of the 10 mM ligand solution. 7. Repeat steps 3 to 6 for DNA concentrations decreasing in half-log intervals (i.e., 300, 100, 30, . . . µM DNA base pairs) to a final DNA concentration of 1 nM. The broad range of DNA concentrations ensures a rigorous and complete binding isotherm.
Analyze data 8. Adjust all absorbance values to account for dilution of the sample that occurred upon addition of each aliquot of ligand. 9. For each aliquot of ligand added to the DNA solution, calculate the total ligand concentration in solution (Ct), either directly from knowledge of the concentration of the ligand stock solution or, to avoid pipetting errors, from the absorbance at the isosbestic wavelength using Beer’s law (i.e., A = εisoslCt, where A is the absorbance, εisos is the extinction coefficient at the isosbestic wavelength, l is the cuvette pathlength in centimeters, and Ct is the total concentration of the ligand). 10. For each ligand concentration, calculate the concentration of bound ligand (Cb) according to the following equation: Cb =
εflCt − A εfl − εbl
Equation 8.2.1
where A is the absorbance of the solution, εf is the extinction coefficient of the free (unbound) ligand, εb is the extinction coefficient of the fully bound ligand, Ct is the total ligand concentration, and l is the pathlength of the cuvette in centimeters. This equation is derived from Beer’s law in which the total absorbance is the sum of contributions from the free and bound ligand: A = εflCf + εblCb. The equation is solved for Cb by substitution from the equation Ct = Cf + Cb.
11. Calculate Cf according to the following equation: Cf = Ct − Cb
Equation 8.2.2
12. Calculate r, the number of moles of ligand bound per DNA base pair, according to the following equation: r=
Cb CDNA
Equation 8.2.3
where CDNA is the concentration of DNA as base pairs. 13. Plot values of r/Cf versus r for each ligand concentration at each DNA concentration (i.e., a Scatchard plot).
Nucleic Acid Binding Molecules
8.2.3 Current Protocols in Nucleic Acid Chemistry
Figure 8.2.1 Example of a DNA binding isotherm with data fit to the McGhee and von Hippel equation. Esperamicin A1 is an enediyne antibiotic that intercalates in DNA; esperamicin C is an analog of A1 missing the intercalating moiety. Adapted with permission from Yu et al. (1994). Copyright 1994, American Chemical Society.
If there is no positive cooperativity in the binding of ligand to DNA, the plot should appear as a downward-sloping curve as demonstrated in Figure 8.2.1. Positive cooperativity will be apparent as an initial upward slope in the plot followed by a downward-sloping curve.
14. Using any of several software programs that allow nonlinear regression analysis (e.g., Microsoft Excel), calculate the apparent binding (association) constant from the r/Cf and r values according to the neighbor exclusion model of McGhee and von Hippel: 1 − nr r = Kobs(1 − nr) Cf 1 − (n − 1)r
Equation 8.2.4
where Kobs is the intrinsic (apparent) binding constant (units of M−1) for the ligand and the DNA species studied, and n is the neighbor exclusion parameter that represents the size of the ligand binding site in base pairs. The software should calculate both Kobs and n for the fitted data. One can vary the values of either n or Kobs and observe the effect of the changes on the standard deviation of the fit. For further information, the reader is referred to the work by Johnson and Faunt (1992), which provides a general review of least-squares fitting methods.
15. Finally, fit the data to an extended version of the above equation that takes into account cooperative interactions between ligands: n−1
(2ω − 1)(1 − nr) + r − R r = Kobs(1 − nr) Cf s(ω − 1)(1 − nr)
2
1 − (n + 1)r + R 2(1 − nr)
Equation 8.2.5
where ω is the cooperativity parameter (ω >1 indicates positive cooperativity and ω <1 indicates negative cooperativity).
Determination of Binding Thermodynamics
The use of Equation 8.2.5 is justified only if the standard deviation of the fitted line is smaller than that derived from Equation 8.2.4 or if there is prior knowledge of cooperativity of the ligand/DNA interaction. If there is little difference between binding constants obtained by either equation, use the values obtained from Equation 8.2.4.
8.2.4 Current Protocols in Nucleic Acid Chemistry
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Calf thymus DNA, sonicated Prepare sonicated calf thymus DNA in assay buffer (10 mM Tris⋅Cl, HEPES, or other buffer, pH 7, containing 1 mM EDTA) as described in UNIT 8.1, Support Protocol; use the preparation recommended for Basic Protocol 3 in that unit—i.e., 10 to 15 min sonication and no size fractionation. COMMENTARY Background Information The experiments in this unit represent just one of many approaches to quantifying the DNA binding affinity of a ligand. While the unit focuses on the use of UV/vis spectroscopy, virtually any technique can be used as long as some property of either the DNA or the ligand changes upon interaction of the two, or the unbound ligand can be physically separated from bound ligand. Examples of applicable techniques include: equilibrium dialysis (Chaires et al., 1982); fluorescence titration (Chaires et al., 1982); centrifugation (Minton, 1990; Yu et al., 1994); solvent partitioning (Waring et al., 1975); and DNA cleavage (Krishnamurthy et al., 1995). The calculation of Cf, Cb, and r from data obtained with these methods is straightforward. For example, for fluorescent ligands whose fluorescence decreases upon binding to DNA (e.g., daunomycin), Cf can be calculated as follows: F − Fmin Cf = Ct F0 − Fmin
Equation 8.2.6
where F is the fluorescence of the ligand in the presence of some quantity of DNA, Fmin is the fluorescence of the ligand in the presence of a large excess of DNA (i.e., fully bound ligand), and F0 is the fluorescence of the ligand in the absence of DNA. For a fluorescent ligand whose fluorescence increases upon binding to DNA (e.g., ethidium bromide), Cb can be calculated as follows: F − F0 Cb = Ct Fmax − F0
Equation 8.2.7
where Fmax is the fluorescence of the ligand in the presence of a large excess of DNA (i.e., fully bound ligand). In either case, Cb or Cf is calculated by difference.
Ideally, the binding data should be acquired by more than one method, since the physicochemical properties of the ligand may affect the data obtained with different techniques. This is illustrated in studies performed by Chaires et al. (1982), in which the binding of daunomycin to DNA was assessed by solvent partitioning and spectral titration techniques. Once the concentrations of bound and free ligand have been determined, the binding affinity can be quantified by analysis of the ligandDNA isotherm. In most cases, a Scatchard plot of the data (i.e., a plot of r/Cf versus r) will not produce a straight line as one would expect for ligand binding to a simple receptor with one or a few discrete binding sites. Instead, there is usually a pronounced downward curvature to the plot, as shown in Figure 8.2.1. The curvature arises from the binding of one drug molecule that excludes the binding of other ligand molecules nearby along the DNA polymer, a phenomenon referred to as neighbor exclusion. Extraction of a binding constant in the face of the neighbor-exclusion phenomenon can be achieved by several different numerical analysis methods, the most convenient of which are the equations derived by McGhee and von Hippel. Equation 8.2.4 is applied to the binding of a ligand to a DNA lattice of identical, noninteracting binding sites. The value n, the exclusion parameter, describes the size of the binding site in base pairs and it is best determined with data approaching saturation of the DNA binding sites with ligand (Fig. 8.2.2). However, Equation 8.2.4 does not address cooperative interactions between ligands. To account for positive and negative cooperativity in ligand binding, McGhee and von Hippel (1974) introduced an additional parameter, ω, into Equation 8.2.4 to yield Equation 8.2.5. If ω >1, binding of one ligand molecule will promote the binding of another ligand nearby (positive cooperativity), while ω <1 implies that binding of one ligand molecule will inhibit
Nucleic Acid Binding Molecules
8.2.5 Current Protocols in Nucleic Acid Chemistry
Figure 8.2.2 Plot of log(Cf) versus r for the binding of esperamicin A1 to DNA to demonstrate completeness of the binding isotherm. The curve adopts a roughly sigmoidal shape and becomes nearly asymptotic at low and high r values, which reflects complete binding and saturation, respectively. Data adapted from Yu et al. (1994).
Determination of Binding Thermodynamics
the binding of another molecule nearby (negative cooperativity). The case of negative cooperativity is extremely difficult to distinguish from the neighbor-exclusion phenomenon, and the reader is referred to the work of Correia and Chaires (1994) for approaches to resolving the contributions of these two features of ligand binding. Furthermore, the use of the more complicated Equation 8.2.5 can sometimes produce a poorer fit of the data than Equation 8.2.4. The best approach is to fit the data to both equations and compare the standard deviation of the plots. One can also apply the F test to the resulting standard deviations to determine the goodness of fit (Chaires, 1992). At this point, it is important to discuss the allosteric binding model of Dattagupta et al. (1980). This model is more rigorous for small molecules than the McGhee and von Hippel model, but it requires greater sophistication in statistical-mechanical approaches to fitting data. The McGhee and von Hippel model assumes that the conformation of DNA is not affected by ligand binding and that positive cooperativity arises instead from ligand-ligand interactions, such as that occurring between protein ligands. The allosteric model of Dattagupta et al. (1980), however, takes into account changes in DNA conformation caused by ligand binding, which is usually what is occurring. Positive cooperativity in this model arises when ligand binding changes DNA conformation in such a way as to promote binding of other ligands. The only drawback to the model is that the data cannot be fit
using simple nonlinear regression programs and instead requires a statistical-mechanical model to extract the binding constant. Interested experimentalists are encouraged to explore the work of Dattagupta et al. (1980). Further extraction of thermodynamic information from the binding isotherms, such as enthalpy and entropy, is beyond the scope of this unit. However, one can begin to parse the contributions made by electrostatic and nonelectrostatic interactions by measuring the binding constants in the presence of varying concentrations of NaCl. For each salt concentration, the apparent binding constant, Kobs, can be used to calculate the observed Gibbs free energy (∆Gobs°) from the relation: ∆G°obs = −RTlnKobs Equation 8.2.8
where R is the gas constant and T is the temperature in degrees Kelvin. The ∆Gobs° can be viewed as the sum of two contributions: ∆G°obs = ∆Gnonel + ∆Gel Equation 8.2.9
where ∆Gnonel and ∆Gel represent the nonelectrostatic and electrostatic contributions to free energy. For several different NaCl concentrations, it has been shown by Record et al. (1978) that a plot of Kobs versus salt concentration yields a slope of Zϕ according to the equation: δlnKobs δln(Na+)
= −Zϕ
Equation 8.2.10
where Z is the charge of the ligand, and thus represents the number of counterions released during drug binding, and ϕ is the fraction of Na+ associated with each DNA phosphate. The electrostatic contribution to the free energy of ligand binding can then be calculated from: ∆Gel = ZϕRTln(Na+)
Equation 8.2.11
In this way, one can begin to determine the individual contributions of electrostatic and nonelectrostatic (e.g., hydrophobic) contributions to the DNA binding energetics of a ligand.
Critical Parameters There are several factors critical to the determination of binding constants of small molecules by the technique of spectroscopic titration
8.2.6 Current Protocols in Nucleic Acid Chemistry
described here, and more generally for all methods used to assess binding thermodynamics. The most obvious is that the UV/vis absorbance properties of the ligand must change when the ligand binds to DNA. A 10% to 15% difference between the absorbance of bound and unbound ligand can be considered a minimum for the rigorous determination of a binding constant. The ligand should have a λmax > 300 nm to avoid interference from DNA. More generally, the titration of ligand and DNA must be performed over a broad range of DNA concentrations to ensure a complete isotherm. The completeness of the binding isotherm can be assessed by plotting log(Cf) versus r for the entire set of data. Ideally, the data will fall on a sigmoidal curve as illustrated in Figure 8.2.2 for the enediyne antibiotic esperamicin A1. At high r values, the curve becomes asymptotic due to the saturation of DNA binding sites. Though not illustrated well in Figure 8.2.2, the plot becomes asymptotic at low r values where most of the ligand is bound to DNA and the Cf becomes vanishingly small. Another critical factor is the purity of the DNA. The presence of contaminating salts, metals, buffers, proteins, and other molecules can affect the DNA binding energetics of the ligand or block binding sites. Ideally, the DNA should be thoroughly extracted with phenol/chloroform and desalted by dialysis into the appropriate buffer; alternatively, the extracted DNA can be purified by gel-filtration chromatography as described in the Support Protocol in UNIT 8.1.
Troubleshooting Several problems may be encountered during the collection of binding data. First, there may be significant scatter in the data at the extremes of the binding isotherm due to systematic errors in determination of the extinction coefficients and the sensitivity of the spectrophotometer. It is best to use data in which the concentration of bound ligand as a fraction of total ligand concentration is between 0.2 and 0.8. Equipment errors may also pose problems. Sequential additions of ligand should be limited to only a few aliquots to avoid changes in absorbance due to drift in the spectrophotometer electronics. Pipetting errors will also introduce significant scatter in the binding data. For ligands soluble only in organic solvents, the vapor pressure created during pipetting of aliquots of the ligand in a volatile solvent can
affect the actual volume of ligand transferred into the spectrophotometer cuvette. The use of nonvolatile solvents (e.g., DMSO) or positivedisplacement pipets helps to prevent this problem. It is also important to use pipets that are both accurate and precise in the volume range used in the studies (1 to 10 µL).
Anticipated Results An example of the anticipated results of these experiments is shown in the Scatchard plot in Figure 8.2.1 for the enediyne antibiotics esperamicins A1 and C. The range of r values obtained will vary as a function of the binding affinity of the ligand.
Time Considerations A complete binding isotherm will require at least 2 weeks to obtain. The bulk of the time will be spent preparing reagents such as the DNA substrate, which must be quite pure to prevent artifacts from contaminating salts and proteins. Collection of the binding data by spectroscopic titration will require ∼1 week, with the largest fraction of the time spent determining the optimal range of concentrations of ligand and DNA.
Literature Cited Chaires, J.B. 1992. Application of equilibrium binding methods to elucidate the sequence specificity of antibiotic binding to DNA. In Advances in DNA Sequence Specific Agents, Vol. 1 (L. Hurley, ed.) pp. 3-23. JAI Press, New York. Chaires, J.B., Dattagupta, N., and Crothers, D.M. 1982. Studies on interaction of anthracycline antibiotics and deoxyribonucleic acid: Equilibrium binding studies on interaction of daunomycin with deoxyribonucleic acid. Biochemistry 21:3933-3940. Correia, J.J. and Chaires, J.B. 1994. Analysis of drug-DNA binding isotherms: A Monte Carlo approach. Methods Enzymol. 240:593-614. Dattagupta, N., Hogan, M., and Crothers, D.M. 1980. Interaction of netropsin and distamycin with deoxyribonucleic acid: Electric dichroism study. Biochemistry 19:5998-6005. Hinton, D.M. and Bode, V.C. 1975. Ethidium bromide affinity of circular λ-deoxyribonucleic acid determined fluorimetrically: The effect of NaCl concentration on supercoiling. J. Biol. Chem. 250:1061-1070. Johnson, M.L. and Faunt, L.M. 1992. Parameter estimation by least-squares methods. Methods Enzymol. 210:1-37. Krishnamurthy, G., Brenowitz, M.D., and Ellestad, G.A. 1995. Salt-dependence of calicheamicinDNA site-specific interactions. Biochemistry 34:1001-1010.
Nucleic Acid Binding Molecules
8.2.7 Current Protocols in Nucleic Acid Chemistry
McGhee, J.D. and von Hippel, P.H. 1974. Theoretical aspects of DNA-protein interactions: Co-operative and non-co-operative binding of large ligands to a one-dimensional homogeneous lattice. J. Mol. Biol. 86:469-489.
Waring, M.J., Wakelin, L.P.G., and Lee, J.S. 1975. A solvent-partition method for measuring the binding of drugs to DNA: Application to the quinoxaline antibiotics echinomycin and triostin A. Biochim. Biophys. Acta 407:200-212.
Minton, A.P. 1990. Quantitative characterization of reversible molecular associations via analytical centrifugation. Anal. Biochem. 190:1-6.
Yu, L., Golik, J., Harrison, R., and Dedon, P. 1994. The deoxyfucose-anthranilate of esperamicin A1 confers intercalative DNA binding and causes a switch in the chemistry of bistranded DNA lesions. J. Am. Chem. Soc. 116: 9733-9738.
Record, M.T., Jr., Anderson, C.F., and Lohman, T.M. 1978. Thermodynamic analysis of ion effects on the binding and conformational equilibria of proteins and nucleic acids: The role of ion association or release, screening, and ion effects on water activity. Q. Rev. Biophys. 11:103-178. Waring, M.J. 1965. Complex formation between ethidium bromide and nucleic acids. J. Mol. Biol. 13:269-282.
Contributed by Peter C. Dedon Massachusetts Institute of Technology Cambridge, Massachusetts
Determination of Binding Thermodynamics
8.2.8 Current Protocols in Nucleic Acid Chemistry
A Competition Dialysis Assay for the Study of Structure-Selective Ligand Binding to Nucleic Acids
UNIT 8.3
DNA is polymorphic and exists in a variety of secondary and tertiary structures that are dictated by both sequence and environmental conditions. Unique DNA structures represent potential targets for small molecules, and this provides a promising new avenue for drug development, as small molecules that selectively bind to a particular structure could interfere with whatever biological function requires that structure. Recently, for example, multistranded triplex and tetraplex (quadruplex) nucleic acid structures have attracted considerable attention as targets for small molecules. Attempts to rationally design small molecules that bind selectively to a particular structure are hampered by the lack of a rapid and convenient assay for structural selectivity. Typically, thermal denaturation studies monitored by UV absorbance are used (UNIT 7.3), but such an assay is time consuming and cannot easily be used to screen a large number of different structures simultaneously. In addition, melting curves in the presence of bound ligand are often complex and multiphasic, making the analysis and interpretation of data difficult. Alternatively, binding constants for the interaction of ligands with the target structure can be measured directly (UNIT 8.2) and compared to standard duplex DNA, but such measurements are even more time consuming and tedious than obtaining optical melting curves. In order to circumvent these difficulties, the authors recently devised a novel competition dialysis assay for rapidly screening ligand binding to a variety of nucleic acid structures. Determination of structure-selective ligand binding using competition dialysis is described here (see Basic Protocol). The competition dialysis assay is simple, straightforward, and rapid once stock solutions of the nucleic acid structures of interest have been prepared (see Support Protocol). DETERMINATION OF STRUCTURE-SELECTIVE LIGAND BINDING BY COMPETITION DIALYSIS In the competition dialysis assay, the interaction of a small molecule of interest with an array of nucleic acid structures is studied. The exact structures to be included in the array are dictated by the interests of the investigator. The competition dialysis assay is based on the fundamental thermodynamic principle of equilibrium dialysis. The approach is easily summarized as follows. A nucleic acid solution is placed inside a dialysis unit containing a semipermeable membrane whose pore size allows small molecules to pass while large macromolecules are retained. The dialysis unit is then suspended in a solution containing small ligand molecules. At equilibrium, the chemical potential of the free ligand must be equal inside and outside the dialysis unit, and any excess ligand on the macromolecule side of the membrane can be attributed to binding. In the competition dialysis experiment, several nucleic acid samples of differing structure, at identical concentrations, are dialyzed against a common ligand solution. The volume of the ligand solution must be much larger than the total volume of nucleic acid samples within the dialysis units in order to keep the free ligand concentration relatively constant. At equilibrium, more ligand will accumulate into the sample(s) with the structures that have the highest relative binding affinity. The simple experimental setup is shown in Figure 8.3.1.
BASIC PROTOCOL
Nucleic Acid Binding Molecules Contributed by Jonathan B. Chaires Current Protocols in Nucleic Acid Chemistry (2002) 8.3.1-8.3.8 Copyright © 2002 by John Wiley & Sons, Inc.
8.3.1 Supplement 11
20 mm
7 mm
Figure 8.3.1 Experimental setup for the competition dialysis experiment. Slide-a-Lyzer MINI dialysis units, held in their floatation device and containing nucleic acid samples, are in contact with a common ligand solution. The diagram on the right shows the dimensions of the dialysis units.
The choice of dialysis membrane to be used in the assay is dictated primarily by the size of the nucleic acid samples. Dialysis units with regenerate cellulose membranes with a molecular weight cutoff (MWCO) of 1,000 to 10,000 Da are available. A MWCO of 3500 Da was selected for routine use, since such membranes would retain the smallest oligonucleotides included in the assay, yet would allow passage of the small molecules studied. Materials Dialysate solution: 1 µM test ligand in BPES (see recipe for BPES) Nucleic acid samples (see Support Protocol) 10% (w/v) SDS Slide-a-Lyzer MINI dialysis units with appropriate molecular weight cutoff (e.g., 3500 Da; Pierce), and floatation device Submicro spectrophotometer cell (Starna Cells) with 1-cm path length and 160-µL geometric volume Graphics software (e.g., Microcal Origin) 1. For each competition dialysis assay, place 400 mL dialysate solution (containing 1 µM test ligand in BPES) into a 1-L beaker. Add a stir bar of appropriate size. The exact volume of dialysate solution required may be varied, and depends on the number of nucleic acid samples to be studied and the size of the float employed to hold the dialysis units. The choice of 1 ìM concentration is somewhat arbitrary and may also be varied. This concentration is convenient for absorbance detection of most DNA-binding ligands.
2. Pipet 200 µL of each nucleic acid sample into a separate Slide-A-Lyzer MINI dialysis unit with the appropriate MWCO membrane (e.g., 3500 Da). The exact array of nucleic acid structures used will vary according to the interests of the investigator. The nucleic acids listed in Table 8.3.1 are suggestions, and should be seen as a point of departure.
A Competition Dialysis Assay for Structure-Selective Ligand Binding to Nucleic Acids
3. Place all dialysis units in a MINI dialysis floatation device and then place the whole floatation unit in the beaker containing the dialysate solution. Cover the beaker with Parafilm, wrap it with foil, and allow the system to equilibrate with continuous stirring for 24 hr at room temperature (20° to 22°C). In early tests of the method, dialysis periods of 24, 48, and 72 hr were compared. All these times yielded identical results, indicating that dialysis equilibrium was attained by 24 hr.
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Table 8.3.1
Nucleic Acid Samples Used in Competition Dialysis Experimentsa
Conformation
Nucleic acid
λ (nm)
ε (M-1cm-1) Monomeric unit
Single-strand purine Single-strand pyrimidine Duplex DNA
poly(dA) poly(dT)
257 264
8,600 8,520
Nucleotide Nucleotide
C. perfringens (31% GC) calf thymus (42% GC) M. lysodeikticus (72% GC) poly(dA):poly(dT) [poly(dAdT)]2 [poly(dGdC)]2 poly(rA):poly(dT) poly(rA):poly(rU) poly(dA):[poly(dT)]2 (5′T2G20T2)4 5′AG3TTAG3TTAG3TTAG3 [poly(dC)]4
260 260 260 260 262 254 260 260 260 260 260 274
12,476 12,824 13,846 12,000 13,200 16,800 12,460 14,280 17,200 39,267 73,000 29,600
Base pair Base pair Base pair Base pair Base pair Base pair Base pair Base pair Triplet Quartet Quartet Quartet
DNA-RNA hybrid Duplex RNA Triplex DNA Quadruplex DNA 1 Quadruplex DNA 2 i-motif
aKey: λ, wavelength; ε, molar extinction coefficient at wavelength λ, expressed in terms of the monomeric unit specified.
4. At the end of the equilibration period, carefully collect 180 µL of each DNA sample and transfer to microcentrifuge tubes. Add 20 µL of 10% (w/v) SDS to make the final concentration 1%. SDS is widely used to dissociate bound ligands from DNA. It is used here to avoid any complexities resulting from differences in absorbance of free and bound ligand. Ligand dissociation by SDS is usually rapid (milliseconds to seconds) and should be complete within 5 min.
5. Spectrophotometrically measure the absorbance of each sample at a wavelength that is appropriate for the ligand, but that is greater than 320 nm. Use a submicro spectrophotometer cell with 1-cm path length and 160-µL geometric volume to measure the small volume used in this version of the assay. The appropriate wavelength to use for each ligand is determined from information in the literature, if available. If no information is available, preliminary UV-Vis spectroscopic experiments must be done to determine an appropriate wavelength. In the worst case, an extinction coefficient at an appropriate wavelength must be determined by using careful analytical techniques and dry weight measurements. These procedures are beyond the scope of this unit. Preliminary spectrophotometric studies should be done to verify that the ligand absorbs light above 320 nm. Below that wavelength, the nucleic acid samples (which absorb strongly) will preclude an accurate measurement of ligand concentration by absorbance. If absorbance measurements cannot be used, fluorescence or any other appropriate analytical method can be used to determine the total ligand concentration as dictated by the properties of the particular ligand under investigation.
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DODC
Ethidium i-motif quadruplex 2 quadruplex 1 triplex DNA poly(rA):poly(rU) poly(rA):poly(dT) [poly(dGdC)]2 [poly(dAdT)]2 poly(dA):poly(dT)
M. lyso. DNA CT DNA
C. perf. DNA poly(dT) poly(dA) 0
2
4
6
C bound (µM)
8
10
0
1
2
3
4
Cbound (µM)
Figure 8.3.2 Sample competition dialysis data for ethidium bromide and 3,3′-diethyloxadicarbocyanine (DODC). Data are presented as a bar graph in which the amount of ligand bound to each nucleic acid structure is plotted. See Table 8.3.1 for further definition of nucleic acids analyzed.
6. Correct each absorbance reading by multiplying by 1.11 (i.e., 200 µL/180 µL) to account for the slight dilution resulting from the addition of the SDS solution. Calculate the total ligand concentration using Beer’s law: Ct = A/εl, where A is the absorbance, ε is the extinction coefficient of the ligand, and l is the path length of the cuvette. If the extinction coefficient of the ligand of interest is not known, it must be determined as described in UNIT 8.1 (see assessment of changes in the optical properties of ligands).
7. Determine the free ligand concentration (Cf) spectrophotometrically using an aliquot of the dialysate solution, using the same procedures described in steps 4 to 6 above. Usually the free ligand concentration should not vary appreciably from the initial 1 ìM.
8. Determine the amount of ligand bound to each nucleic acid structure (Cb) by calculating the difference: Cb = Ct − Cf. 9. Plot the data as a bar graph using any convenient graphics software. The author uses Origin Version 5.1 for plotting and analysis, but any package will do. Figure 8.3.2 shows representative data for two different ligands. SUPPORT PROTOCOL
A Competition Dialysis Assay for Structure-Selective Ligand Binding to Nucleic Acids
PREPARATION OF NUCLEIC ACID SOLUTIONS The most time-consuming step in the competition dialysis method is the preparation of nucleic acid stock solutions for use in the assay. The exact nucleic acids to be used depend on the interests of the investigator. The only requirements are that the nucleic acid structure of interest is stable under the ionic conditions used and that it is large enough to be retained by the particular dialysis membrane used in the assay. Table 8.3.1 shows one set of samples and provides a point of departure that can be changed and supplemented as needed. Natural duplex DNA samples require the most preparation. Commercial polynucleotides and oligonucleotides may generally be used without further purification, and are simply dissolved in buffer. The quality of all samples must be checked by UV
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spectroscopy (UNIT 7.3), circular dichroism spectroscopy (UNIT 7.12), and optical melting studies (UNIT 7.3). Materials Natural DNA samples from Clostridium perfringens, calf thymus, and Micrococcus lysodeikticus (Sigma) BPES (see recipe) Synthetic polydeoxyribonucteotides (Amersham Pharmacia Biotech): poly(dA), poly(dT), poly(dC), poly(dA):poly(dT), poly(dAdT), and poly(dGdC) Synthetic polyribonucleotides (Sigma): poly(rA) and poly(rA):poly(rU) Custom synthetic deoxyoligonucleotides 5′T2G20T2 and 5′AG3(T2AG3)3 (Research Genetics or Oligos Etc.) Additional reagents and equipment for preparing genomic DNA fragments for viscometry (UNIT 8.1), and for UV absorbance (UNIT 7.3) and circular dichroism (UNIT 7.12) spectroscopy 1. Prepare natural DNA samples from Clostridium perfringens, calf thymus, and Micrococcus lysodeikticus using the protocol for preparation of genomic DNA fragments for viscometry described in UNIT 8.1. At the final step, dissolve samples in BPES at a concentration of ∼1 mg/mL. 2. Individually dissolve the following synthetic DNA and RNA polynucleotides in BPES at a concentration of ∼1 mg/mL: poly(dA) poly(dT) poly(dC) [poly(dAdT)]2 [poly(dGdC)]2 poly(dA):poly(dT) poly(rA) poly(rA):poly(rU). As stated in the introduction, synthetic DNA and RNA polynucleotides are used without further purification.
3. Prepare the poly(rA):poly(dT) DNA:RNA hybrid and poly(dA):[poly(dT)]2 DNA triplex by mixing poly(rA) or poly(dA):poly(dT) with poly(dT) in a 1:1 molar ratio, heating to 90°C, and slowly cooling to room temperature. For additional details on preparing these solutions at a 1:1 molar ratio, see determination of molecularity and extinction coefficients of oligonucleotide complexes in UNIT 7.3.
4. Prepare (5′T2G20T2)4 (parallel quadruplex DNA), 5′AG3(T2AG3)3 (antiparallel quadruplex DNA), and [poly(dC)]4 (i-motif DNA) by dissolving the oligonucleotide samples in BPES, heating the samples to 90°C for 2 min, and slowly cooling to room temperature. 5. Determine the concentrations of all of the above sample solutions by UV spectrophotometry using the wavelengths and extinction coefficients provided in Table 8.3.1. If the extinction coefficient of a nucleic acid sample of interest to the investigator is not known, it must be determined using the protocol described in UNIT 7.3.
6. Characterize the quality of all samples by recording their UV absorbance spectrum (see UNIT 7.3), circular dichroism spectrum (see UNIT 7.12), and melting curve (see UNIT 7.3).
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Reference data for comparison is available for free online from the American Chemical Society as supporting information for published articles. See http://www.pubs.acs.org/ cgi-bin/suppinfo.pl?ja9934955.
7. Prepare working stock solutions of each nucleic acid by appropriate dilution with BPES to yield a final concentration of 75 µM in terms of the monomeric unit. Store up to 6 months at 4°C. Monomeric unit means nucleotides (nt) for single-stranded forms, base pairs (bp) for duplex forms, base triplets for triplex forms, and base tetrads for quadruplex forms (Table 8.3.1). These working stock solutions may be stored at 4°C. It is imperative that all nucleic acid samples be at identical concentrations, but the concentration chosen can vary. The key requirement is to have a large excess of potential nucleic acid binding sites relative to the free ligand concentration. Early tests in the author’s laboratory used nucleic acid concentrations of 75, 100, 150, and 200 ìM. All gave suitable results. The choice of 75 ìM represents a compromise that ensures a sufficient excess of sites but economizes on the cost of nucleic acid samples.
REAGENTS AND SOLUTIONS Use distilled, deionized water or other ultrapure water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
BPES (buffered phosphate/EDTA/saline) 6 mM Na2HPO4 2 mM NaH2PO4 1 mM Na2EDTA 185 mM NaCl, pH 7.0 Store up to 6 months at 4°C COMMENTARY Background Information
A Competition Dialysis Assay for Structure-Selective Ligand Binding to Nucleic Acids
The competition dialysis method described here evolved from a simpler method first described by Müller and Crothers (1975). They described a three-chamber dialysis apparatus that proved useful for quantitative studies of the base selectivity of ligand-DNA interactions. In the original method, two different natural DNA samples with differing base composition were dialyzed against a common ligand-containing dialysate solution. If the test compound preferentially binds to GC over AT base pairs, more ligand would accumulate in the sample with the higher GC content. The method predated chemical and enzymatic footprinting methods for the identification of ligand binding sites on DNA, and allowed rather detailed quantitative inferences about the nature of preferred binding sites to be made (Müller and Crothers, 1975; Chaires, 1992). The assay was extended to studies of structure-selective ligand binding by comparing binding to standard duplex DNA with binding to a DNA:RNA hybrid (Becker and Dervan, 1979), to left-handed Z DNA (Chaires, 1985; Satyanarayana et al., 1992),
and to triplex DNA (Haq et al., 1996). The current, expanded version of the competition dialysis assay was introduced in 1999 (Ren and Chaires, 1999). The assay has since been applied to investigate a wide variety of compounds with several types of nucleic acid structures, both in the author’s laboratory (Ren et al., 2000, 2001; Ren and Chaires, 2000, 2001) and in other laboratories (Perry and Jenkins, 1999; Alberti et al., 2001; Lisgarten et al., 2002). There are several advantages of the competition dialysis assay for the study of structureselective ligand binding. First, the method allows a variety of nucleic acid structures to be studied simultaneously, allowing for a more complete comparison than is typically available from melting or binding studies. Second, the method is thermodynamically sound, and is based on the fundamental principle of equilibrium dialysis. Finally, the method is comparatively rapid and, while it is not yet a true high-throughput screen, unambiguous results may be obtained within 24 hr once the necessary stock solutions are prepared. Few disadvantages are evident so far. Perhaps the greatest
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potential disadvantage is that a particular nucleic acid structure of interest may not be stable under certain ionic conditions, such as the BPES used here. The assay is flexible, however, and any type of buffer can be used, although the stability of the array of nucleic acid structures would need to be reevaluated for any new buffers chosen for use.
Critical Parameters In order for the competition dialysis assay to be quantitative, it is crucial that the concentration of potential binding sites in the nucleic acid samples be identical. Since these are in equilibrium with the same solution containing the free ligand, differences in nucleic acid concentrations would result in differences in the total amount of ligand bound, obscuring the underlying binding specificity. A comment is needed here about the choice of concentration units. Concentration units are defined in terms of the monomeric unit of the poly- or oligonucleotide used to produce a given structure. This means nucleotides for single-stranded forms, base pairs for duplex forms, triplets for triplex forms, and tetrads for quadruplex forms. These monomeric units represent the minimal potential binding site. An advantage of that choice for the concentration scale is that it negates differences in length among the various samples. A critical consideration, however, is that because of actual differences in lengths among the various poly- and oligonucleotides used, the concentration of ends differs dramatically among the samples. This would present a problem only in the case of a ligand that binds only to an end site and avoids internal sites. In such an unusual case, the assay would be flawed, and the bias would manifest itself as greater apparent binding to the samples with the shortest lengths.
Troubleshooting In the author’s experience with the assay over a four-year period, few problems have been encountered. On a few occasions, some mini-dialysis units were found to leak, causing the nucleic acid sample to leak from the unit. Such cases can be diagnosed by a loss of the UV absorbance from the retentate solution (solution inside the dialysis unit). A second problem encountered on occasion resulted from ligand binding to surfaces, including the dialysis membrane, the stir bar, and the interior surface of the beaker. Minor adsorption can be tolerated, since it represents only another competing equilibrium. Any ligand bound to the
dialysis unit itself will not be measured, since the retentate solution is removed from the unit prior to treatment with SDS and determination of the ligand bound to the nucleic acid. Since only the free ligand concentration dictates the amount bound to the various nucleic acid structures, the assay remains valid as long as there is appreciable free ligand in solution. In such cases, the free ligand concentration would decrease relative to the 1 µM starting concentration. Comparison of binding to structures within that particular assay would remain valid, but comparison to a separate assay with a different free ligand concentration would not. In the worst case, all of the ligand would be adsorbed to the beaker, stir bar, and dialysis membrane surfaces (encountered only once out of over 150 compounds studied). In such a case, the method is useless. Finally, on occasion some retentate will accumulate on the cap of the dialysis unit. These few drops should be carefully pipetted off and mixed with the retentate. Low-speed centrifugation of the dialysis unit to remove liquid adhering to the cap is not recommended, since retentate solution might be lost through the membrane, or the membrane might break.
Anticipated Results Figure 8.3.2 shows representative results for two different nucleic acid binding ligands: ethidium and 3,3′-diethyloxadicarbocyanine (DODC). Ethidium, the classic DNA intercalator, is relatively nonspecific, while DODC interacts only with multistranded structures. Interpretation and analysis of competition dialysis is n or mally straightforward and unambiguous by design, and simple inspection of the results in graphical format reveals binding preferences. The data of Figure 8.3.2 provide a wealth of information about the comparative nucleic acid binding of ethidium and DODC. First, neither ethidium nor DODC bind to single-stranded DNA—poly(dA) and poly(dT). Ethidium binds to all duplex forms present in the assay, while DODC does not bind to any. Among the duplex forms present, ethidium binds with a slight preference to the DNA:RNA hybrid—poly(rA):poly(dT)—and to duplex RNA—poly(rA):poly(rU). The weak binding of ethidium to the poly(dA):poly(dT) duplex arises from the well-known unusual structure of that polynucleotide (Herrera and Chaires, 1989). Ethidium binds to triplex DNA as well as it does to natural duplex DNA, but binds only slightly to quadruplex forms. DODC binds preferentially to the DNA triplex
Nucleic Acid Binding Molecules
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poly(dA):[poly(dT)]2, and also interacts to a lesser extent with parallel and antiparallel quadruplex structures. Neither ethidium nor DODC binds to the four-stranded i-motif. The data of Figure 8.3.2 show the power of the method and demonstrate how clear indications of structural selectivity can be obtained rapidly and efficiently in a single experiment. More elaborate discussions of the interpretation of results are presented in the original published papers (Ren and Chaires, 1999, 2000, 2001).
Time Considerations The most time-consuming part of the assay is the preparation and characterization of the nucleic acid stock solutions. Preparation of these will take several days, but no more than one week. Once stock solutions are prepared and available, the assay proceeds rapidly. Setting up a single assay consisting of 12 to 20 nucleic acid samples takes only 30 to 60 min, at which point the dialysis may proceed unattended for the required 24-hr period. Removing the samples, adding SDS, and measuring absorbance should take only 60 to 90 min. It is, of course, possible and even desirable to run several competition dialysis assays with different ligands in parallel. This expands the time needed to set up and analyze the results, but leads to more efficient gathering of primary data.
Literature Cited Alberti, P., Ren, J., Teulade-Fichou, M.P., Guittat, L., Riou, J.F., Chaires, J., Helene, C., Vigneron, J.P., Lehn, J.M., and Mergny, J.L. 2001. Interaction of an acridine dimer with DNA quadruplex structures. J. Biomol. Struct. Dyn. 19:505-513. Becker, M.M. and Dervan, P.B. 1979. Molecular recognition of nucleic acids by small molecules. Binding affinity and structural specificity of bis(methdium)spermine. J. Am. Chem. Soc. 101:3664-3666. Chaires, J.B. 1985. Long-range allosteric effects on the B to Z equilibrium by daunomycin. Biochemistry 24:7479-7486. Chaires, J.B. 1992. Application of equilibrium binding methods to elucidate the sequence specificity of antibiotic binding to DNA. In Advances in DNA Sequence Specific Agents (L. H. Hurley, ed.) pp. 3-23. JAI Press, Greenwich, Conn.
Haq, I., Ladbury, J.E., Chowdhry, B.Z., and Jenkins, T.C. 1996. Molecular anchoring of duplex and triplex DNA by disubstituted anthracene-9,10diones: Calorimetric, UV melting and competition dialysis studies. J. Am. Chem. Soc. 118:10693-10701. Herrera, J.E. and Chaires, J.B. 1989. A premelting conformational transition in poly(dA)-poly(dT) coupled to daunomycin binding. Biochemistry 28:1993-2000. Lisgarten, J.N., Coll, M., Portugal, J., Wright, C.W., and Aymami, J. 2002. The antimalarial and cytotoxic drug cryptolepine intercalates into DNA at cytosine-cytosine sites. Nat. Struct. Biol. 9:5760. Müller, W. and Crothers, D.M. 1975. Interactions of heteroaromatic compounds with nucleic acids. 1. The influence of heteroatoms and polarizability on the base specificity of intercalating ligands. Eur. J. Biochem. 54:267-277. Perry, P.J. and Jenkins, T.C. 1999. Recent advances in the development of telomerase inhibitors for the treatment of cancer. Exp. Opin. Invest. Drugs. 8:1981-2008. Ren, J. and Chaires, J.B. 1999. Sequence and structural selectivity of nucleic acid binding ligands. Biochemistry 38:16067-16075. Ren, J. and Chaires, J.B. 2000. Preferential binding of 3,3′-diethyloxadicarbocyanine to triplex DNA. J. Am. Chem. Soc. 122:424-425. Ren, J. and Chaires, J.B. 2001. Rapid screening of structurally selective ligand binding to nucleic acids. Methods Enzymol. 340:99-108. Ren, J., Bailly, C., and Chaires, J.B. 2000. NB-506, an indolocarbazole topoisomerase I inhibitor, binds preferentially to triplex DNA. FEBS Lett. 470:355-359. Ren, J., Qu, X., Dattagupta, N., and Chaires, J.B. 2001. Molecular recognition of a RNA:DNA hybrid structure. J. Am. Chem. Soc. 123:67426743. Satyanarayana, S., Dabrowiak, J.C., and Chaires, J.B. 1992. Neither delta- nor lambda-tris(phenanthroline)ruthenium(II) binds to DNA by classical intercalation. Biochemistry . 31:9319-9324.
Jonathan B. Chaires University of Mississippi Medical Center Jackson, Mississippi
A Competition Dialysis Assay for Structure-Selective Ligand Binding to Nucleic Acids
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Chemistry of Minor Groove Binder–Oligonucleotide Conjugates The interaction of minor groove binder molecules (MBs) with double-stranded DNA (dsDNA) is a rich area of chemistry. The synthetic chemistry, biochemistry, molecular biology, and medicinal chemistry of MBs have been developed, and much is known about their DNA binding mechanisms. The antibiotic activity of naturally occurring MBs has prompted the development and examination of many synthetic analogs. Generally, MBs are planar, polyaromatic molecules that can form crescent-shaped conformations in aqueous solution. Figure 8.4.1 shows examples of several types of MB structures that are described later in this unit (see Types of MB-ODNs). The strongest-binding MBs are isohelical with the minor groove formed in B-form DNA duplexes, and are stabilized by hydrophobic interactions, van der Waals forces, or hydrogen bonds to nucleotide bases in the floor of the groove. MBs can have high affinity for dsDNA with binding constants that are orders of magnitude larger than other DNA binding agents such as intercalating agents (for more on intercalation, see UNIT 8.1). Interactions of MBs with dsDNA have been reviewed (Zimmer and Wahnert, 1986), and the biological activity of anticancer MB compounds has been described in detail (Cory, 1995). Recently, various types of MBs have been attached to synthetic oligodeoxynucleotides (ODNs). Interactions of these MBODN conjugates with DNA have been studied and are reviewed here. Applications of fluorogenic MB-ODNs in quantitative PCR assays are also presented. MB small molecules have DNA sequence preferences, with A/T-rich dsDNA sequences being especially good targets for binding (Zimmer and Wahnert, 1986). Recently, MBs have been developed that have improved sequence recognition through a dimer-binding motif in which two linked MBs occupy the same binding site (Kielkopf et al., 1998). Sequence-selective interactions between alkylating MB natural products (analogs of the antibiotic CC1065) and dsDNA have also been reviewed (Boger et al., 1997). These highly toxic molecules react rapidly with N3 of adenine residues in dsDNA when bound to certain sequences. Other natural products and synthetic MB analogs are even more potent. The high binding
UNIT 8.4
strength of the MBs to dsDNA (Ka ∼1 × 107 to 1 × 109) can make them effective “DNA delivery agents” that target specific molecular functions such as DNA-cleaving agents (Boger and Zhou, 1993) or alkylating agents (Inga et al., 1999) to the minor groove of DNA. The conjugation chemistry of distamycin-type MBs has been reviewed (Bailly and Chaires, 1998). The authors, as well as others, have studied the hybridization performance of MB-ODN conjugates. Short pieces of synthetic DNA with attached MBs have enhanced DNA affinity, and have improved the hybridization properties of sequence-specific DNA probes. Short MBODNs hybridize with single-stranded DNA (ssDNA) strands to give more stable DNA duplexes than unmodified ODNs with similar lengths. Mismatch discrimination of short MBODNs is enhanced in comparison to longer unmodified ODNs. The stronger binding of MB-ODNs allows for more stringent hybridization conditions to be used in DNA probe– based assays. MB-ODNs are especially useful in quantitative “real-time” PCR assays since they bind efficiently during the high-temperature primer extension cycle. The synthesis and biophysical chemistry of MB-ODN conjugates are reviewed here. Four published structural classes of MB-ODNs and their various dsDNA binding modes are discussed. The well-characterized DPI3-type MBODNs and their interactions with ssDNA target strands are described in detail. First, general methods of synthesis of DPI3-ODNs are presented. Next, hybridization properties, mismatch discrimination, and predictability of binding of these conjugates are described. Finally, synthesis and applications of fluorogenic DPI3-ODNs in quantitative PCR assays are discussed.
DNA BINDING MODES AND STRUCTURE OF MB-ODN CONJUGATES Binding Modes of MB-ODNs MB-ODN conjugates have three distinctive molecular components that can be altered to give sequence-specific DNA probes: the MB, ODN, and linker (Fig. 8.4.2). By varying the MB component, dsDNA binding affinity can be tuned. Altering the length and type of repeat-
Contributed by Igor Kutyavin, Sergey Lokhov, Eugene Lukhtanov, and Michael W. Reed Current Protocols in Nucleic Acid Chemistry (2003) 8.4.1-8.4.21 Copyright © 2003 by John Wiley & Sons, Inc.
Nucleic Acid Binding Molecules
8.4.1 Supplement 13
netropsin
CC-1065
Hoechst 33258
Chemistry of Minor Groove BinderOligonucleotide Conjugates
distamycin
imidazole-lexitropsin
CDPI3 methyl ester
ImPyPy-γ-PyPyPy-Dp
Figure 8.4.1 Examples of minor groove binders (MBs). The planar, aromatic MB molecular structures are drawn in the presumed crescent shape conformation that binds to DNA duplexes. The inner (concave) edge of each MB faces the minor groove in B-form DNA duplexes. Netropsin, distamycin, and imidazole-lexitropsin are of the distamycin type; CC-1065 is an alkylating CPI type; CDPI3 methyl ester is of the DPI type; Hoechst 33258 represents the Hoechst type; and ImPyPyγ-PyPyPy-Dp is of the dimer-forming type. See text for a description of each type of MB.
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R CH3
N H N
O
distamycin-type MB-ODN
N CH3 H O ODN O
O
O P
N
H
N
N
H N CH3
O
H
H N
N
CH3
O
N
O
N
CH3
NH2 N
O
DPI-type MB-ODN N
O ODN O
P
O
O
O
N H
O
H
N N
N
H
N H
O
H
CH3 N
O ODN-O P O-ODN
Hoechst-type MB-ODN N
H N S
H
H
N
H N
N O
N
O N
CH3 H N
CPI-DPI-type MB-ODN N
O O
O N H
N
N O
H
O O
P
O-ODN O
Figure 8.4.2 Published examples of different types of MB-ODN structures. Structures of the MB, the ODN, and the linker all affect hybridization performance. MB-ODN conjugates were prepared by treating an ODN containing a linking group with a suitable reactive group on the MB. The different linker structures are described in the text. R, dabcyl chromophore.
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5′ 3′ ssDNA 5′ 3′ ssDNA 5′ 3′ ssDNA 5′ 3′ 5′
dsDNA
5′ 3′ 5′
dsDNA
5′ 3′ 5′
dsDNA
mode 1: terminal MB duplex, 5′-folding
mode 2A: internal MB duplex, 5′-folding
mode 2B: internal MB duplex, 3′-folding
mode 3A: terminal MB triplex, 5′-folding
mode 3B: terminal MB triplex, 3′-folding
mode 4: terminal MB triplex, intercalating linker
5′ 3′ 5′
dsDNA
mode 5: terminal MB triplex, linked MB dimers
Figure 8.4.3 DNA binding modes of MB-ODNs. The drawings illustrate “unwound” DNA helices with 10-mer MB-ODNs. Mode 1 has the most predictable properties since the MB can only fold into a single dsDNA binding site after hybridization. The outer (convex) edge of the MB is shown as a black oval covering ∼5 bp in the minor groove. Longer linkers are required for the triplex binding modes since the ODN is bound in the major groove.
Chemistry of Minor Groove BinderOligonucleotide Conjugates
ing subunits in the MB allows the balance between DNA affinity and sequence specificity to be optimized. The ODN component of MBODNs provides sequence-specific interactions with dsDNA via natural Watson-Crick hydrogen bonding. Since most MBs bind more tightly to A/T-rich sites, the base content of the ODN can affect MB binding after hybridization. The position of MB attachment and the length and type of linker combine to confine the MB to various local regions of dsDNA. MB-ODNs can recognize dsDNA in a variety of binding modes as illustrated in Figure 8.4.3 and discussed below. In addition to simple DNA duplex formation with ssDNA targets, MB-ODNs have been used to improve binding of triplex-forming oligonucleotides (TFOs). The different binding modes of MB-ODNs have different linker requirements, and each
linker must be properly designed for optimum performance. Understanding the effects of the MB on DNA hybridization allows DNA probe performance to be predicted as described below. Terminal MBs and ssDNA targets (binding mode 1) In binding mode 1, MB-ODNs form DNA duplexes and the MB linked to the terminus folds into the newly formed minor groove. In Figure 8.4.3, a 3′-linked MB is illustrated, but the binding mechanisms are analogous for 5′linked MBs. The linker needs to be long enough to allow the MB to orient itself in the groove, but not so long that it can find other DNA binding sites. This is an important issue since predictable duplex stability requires the MB binding site to be known. Binding mode 1
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allows predictable hybridization properties for DNA probe applications, and is described in detail for the terminally modified DPI3-type MB-ODNs (see Synthesis and Hybridization of DPI3-Type MB-ODNs and Fig. 8.4.2).
alkylation efficiency with triplex targeting. Unfortunately, this CPI-DPI conjugate gave high nonspecific binding and alkylation of the DNA. Non-TFO-directed binding is a general problem with the triplex-directed MBs.
Internal MBs and ssDNA targets (binding mode 2) This binding mode was explored with the Hoechst-type MBs and involves internal linkers to the ODN (Fig. 8.4.3; Wiederholt et al., 1996, 1997). This mode is complicated by the ability of the MB to find one of two possible binding sites—towards either the 5′ terminus (mode 2A) or the 3′ terminus (mode 2B) of the hybridized MB-ODN. This made binding to ssDNA targets unpredictable since the “twostate” model (equilibrium of ssDNA and dsDNA) no longer operates. In addition, the internal linkage via a phosphoramidate bond introduced stereoisomers that had specific directional effects for the various isomers. Despite this complexity, certain internally modified MB-ODNs could form strong duplexes with appropriate ssDNA targets. Fluorescent signal correlated well with dsDNA stability, but the unpredictable nature of these probes and the complexity of the synthesis made MB-ODNs that operate via mode 2 less attractive.
Other MB binding modes Other MB binding modes have also been explored with TFOs. In order to prevent nonspecific binding of alkylating CPI-type MBs, a novel “threading intercalator” linker was developed (Dempcy et al., 1999). In this motif, the CPI was delivered to the minor groove by threading through the helix, presumably after triplex binding (mode 4; Fig. 8.4.3). Sequencespecific DNA alkylation of up to 88% was observed on synthetic targets. The strongest intercalating linker showed some nonspecific alkylation. Another MB binding motif has been described for TFOs conjugated with dimer-forming MBs synthesized from mixed pyrrole and imidazole heterocycles (Kielkopf et al., 1998). After triplex formation, the sequence-specific MB dimers targeted mixed-base stretches adjacent to the TFO sequence (mode 5; Fig. 8.4.3). In one example (Szewczyk et al., 1996a), two TFOs were required where one of the MB dimers was at the 3′ end of one TFO and the other was at the 5′ end of the adjacent TFO. The MBs could interact only after both TFOs hybridized. In theory, this could improve sequence specificity of triplex-forming MBODNs. The mode has severe sequence limitations, and requires design and synthesis of MB dimers for targeted delivery to each dsDNA sequence of interest. In another example (Szewczyk et al., 1996b), a linked MB dimer was directly conjugated to a TFO and allowed each sequence-specific element to interact separately. A matching set of MB dimers must be prepared for optimum binding to each specific DNA sequence, and suitable chemistry has been described (Baird and Dervan, 1996). The threading intercalators and dimer-type MBs are not described further in this unit.
Terminal MB linkers and dsDNA targets (binding mode 3) Triplex-forming oligonucleotides (TFOs) recognize and bind to homopurine sequences in the major groove of dsDNA (Thuong and Helene, 1993). As a result, the terminal MB must be attached to a linker that is long enough to reach around the sugar-phosphate backbone to the minor groove of the dsDNA target. The triplex-stabilizing binding modes are illustrated as mode 3 in Figure 8.4.3. MBs conjugated to TFOs have been shown to improve binding affinity to dsDNA (Robles et al., 1996; Rajur et al., 1997; Robles and McLaughlin, 1997). These MB-ODN conjugates used the Hoechst-type MBs. The various long linkers studied allowed the MB in triplexes to either fold back towards itself (mode 3A) or extend out toward the neighboring duplex region (mode 3B). Alkylating CPI-type analogs of MBs have also been conjugated to TFOs (Lukhtanov et al., 1997), and their interaction with dsDNA has been explored. As much as 98% targeted alkylation of dsDNA was observed for optimized linkers. Addition of a DPI subunit (see Fig. 8.4.2) was required for this high DNA
Types of MB-ODNs Numerous types of MBs are available for conjugation to synthetic ODNs, and the specific molecular interactions responsible for dsDNA binding vary. MB conjugate synthesis requires a reactive linking group on the MB. This requires synthetic organic chemistry to develop the “conjugatable” MBs. The MB linking group then reacts with another linking group on the oligonucleotide. These linking
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groups are readily introduced during DNA synthesis. Structural designs of several published MB-ODN conjugates are shown in Figure 8.4.2. The most notable structural similarity between the MBs is their ability to form stable crescent-shaped conformations in aqueous solution that can fit snugly in the minor groove of B-form DNA. However, each type of MBODN has unique physical properties and chemistry. Investigators chose the MBs shown in Figures 8.4.1 and 8.4.2 because the synthetic chemistry had been developed and the binding mechanisms had been studied. The distamycintype and Hoechst-type MBs have oriented heteroatom-bonded protons on the inside of the crescent-shaped structures that allow hydrogen bonds to form with functional groups on the floor of the minor groove. The DPI3-type MBs are more hydrophobic and bind to the minor groove mostly because of differences in the energies of MB-DNA and MB-water interactions (that is, reduction of MB-water “interface” and surface tension; Mikheikin et al., 2001). Synthesis of the MB-ODN conjugates can be complicated, but has been improved significantly as discussed below. Structural analogs of MB-ODNs with various types of MBs, attachment points (internal versus terminal), and linker structures have been explored. The effect of MB and linker lengths on biophysical properties of these classes of MB-ODN conjugates is described below. The distamycin and DPI3 conjugates are noteworthy since the repeating subunits can be prepared from common heterocyclic precursors, and assembled using welldeveloped peptide chemistry. This simplified the synthesis of these complex biomolecules, and allowed MB analogs of various lengths (DNA affinity) to be studied. The synthesis and hybridization properties of the DPI3 conjugates are described in detail in later sections.
Chemistry of Minor Groove BinderOligonucleotide Conjugates
Distamycin-type MBs The first MB-ODN conjugates described in the literature were the distamycin type (Sinyakov et al., 1995). Distamycin and netropsin are naturally occurring antibiotics that contain repeating subunits of N-methylpyrrolecarboxamide (MPC). Distamycin has three repeating MPC subunits whereas netropsin has two MPC subunits (Fig. 8.4.1). The top structure in Figure 8.4.2 is an example of a distamycin-type MBODN with five MPC subunits. Both distamycin and netropsin also contain a positively charged amidine group that aids water solubility and provides additional DNA affinity. The synthetic
chemistry for preparing and linking the MPC subunits had been developed earlier, and the stepwise peptide coupling chemistry allowed ready access to MPC peptides of various lengths. A reactive alkylamine linker arm at the N-terminus of the peptide was introduced for conjugation to ODNs. Similar conjugation chemistry was later used to directly attach netropsin and distamycin to ODNs (Levina et al., 1996). UV melting measurements of DNA duplexes formed with T8-MB-ODNs and complementary ssDNA strands (binding mode 1) showed dramatic increases in melting temperature (Tm). The MB-ODN conjugates were well behaved in standard physiological buffer conditions. Melting curves showed good cooperativity, typical of unmodified DNA strand melting. The results of these studies are shown in Figure 8.4.4A. The first derivatives of the melting curves showed a stepwise increase in Tm as the number of MPC subunits increased from two to five. MB-ODN conjugates with five repeating MPC subunits showed an increase in Tm of 44°C in comparison to an unmodified DNA duplex. In comparison, conjugation of distamycin-type MBs to a G/C-rich 8-mer ODN gave no significant increase in duplex stability as shown in Figure 8.4.4B. Conjugation of the hydrophobic distamycin-type MBs to ODNs improved the water solubility of the MB significantly and allowed hybridization to occur in standard aqueous buffers. This is a significant advantage, since many MBs are sparingly soluble in water and conjugation to ODNs allows their exploration as DNA-targeting agents. For example, the stabilization of unconjugated MPC molecules could not be studied due to insolubility. In addition, the local concentration of hydrophobic MBs at the attachment point to the ODN is much higher than can be achieved by adding free MB to DNA duplexes in solution. Although hydrophobic MBs initially complicated the conjugation to hydrophilic ODNs, a broader range of potential MB structures could ultimately be explored. Although no direct evidence was presented that showed minor groove binding of the distamycin-type MB-ODN conjugates, the A/T specificity of the stabilization and the stepwise increase in Tm with MB length was compelling. Also consistent with an MB mechanism was the relative lack of affinity to RNA of the distamycin-type MB-ODNs. Presumably, the mechanism of binding first involves duplex formation of the MB-ODN with its comple-
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A/T-rich sequences are efficiently alkylated, and much is known about the sequence-specific binding (and bonding) mechanisms involved (Boger and Johnson, 1995). The structure of CC-1065 has a repeating 1,2-dihydro-3H-pyrrolo[3,2-e]indole-carboxylate (DPI) subunit that provides dsDNA affinity. The alkylating cyclopropapyrroloindole (CPI) subunit of CC1065 can react with A3 of adenine in B-form DNA, and provides the structural basis of the CPI-type MB-ODNs (see CPI-type MBs below). DPI dimers, trimers, and tetramers were prepared as high-affinity, noncovalent B-DNA minor groove–binding agents (Boger et al., 1987). Unlike the distamycin-type MBs, the angular tricyclic structure of the DPI subunit pre-organizes these MBs to form crescentshaped conformations that are isosteric with the
mentary strand. The conjugated MB can then fold back into the newly created minor groove where it is stabilized by hydrophobic interactions. For the distamycin-type MBs, additional stability is provided by hydrogen bonds from the amide hydrogens to N3 of adenine or O2 of thymine. Lack of stabilization of G/C-rich sequences is presumably due to the steric hindrance and relative hydrophilicity provided by the N2 of guanine. Lack of binding to RNA is presumably due to the wide, shallow minor groove in the A-form duplexes of DNA/RNA hybrids. This is typical of distamycin-type MBs. DPI-type MBs The natural product CC-1065 is an alkylating MB with potent antibiotic activity. Certain
A 1 3
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3
B
1 0.2
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5 4 0.1
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Figure 8.4.4 UV melting curves of DNA duplexes formed from distamycin-type MB-ODNs and their complements. First derivative plots of helix-coil transitions at 260 nm for 3′-MB-ODN duplexes are shown. Distamycin-type MBs of various lengths were prepared with zero to five MPC subunits and conjugated to dT8 or G/C-rich 8-mer ODNs. (A) TTTTTTTT-MPCn (n = 0 to 5). (B) CATCCGCTMPCn (n=0,1,3,4,5). The A/T-rich duplex shows a stepwise increase in duplex melting termperature (Tm) with increasing MB length, whereas the G/C-rich duplex shows no increase in stability. Reprinted from Sinyakov et al. (1995) with permission from the American Chemical Society.
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minor groove of B-form DNA. The well-developed synthetic chemistry of the DPI-type MBs allowed exploration of their ODN conjugates soon after the distamycin-type MB-ODNs (Lukhtanov et al., 1995). Conjugation methods were developed (see below) that simplified the synthesis of DPI conjugates in comparison to the distamycin type, and this allowed a variety of analogs to be examined. Most of these early DPI analogs retained the N-terminal carbamoyl group and were abbreviated as CDPI. It was later shown (Lukhtanov et al., 1996a) that the carbamoyl group was not necessary, and other linker groups could be added at this position. The synthesis and properties of the DPI-type MB-ODNs are described in detail below. The major advantage of the DPI-type MBs is the lack of hydrogen bonding groups in the inner surface of the crescent-shaped conformation. DNA duplex stabilization by DPI-type MBs is primarily due to close atomic contacts (van der Waals forces). The importance of hydrophobic interactions and van der Waals forces (Chang and Cheng, 1996) in the DPI-type MBs has been discussed (Boger et al., 1990). Unlike the distamycin-type MBs, the DPI-type MBs can significantly stabilize G/C-rich duplexes. In addition, the lack of hydrogen bonding in the minor groove makes hybridization properties of DPI-type MBs more predictable.
Chemistry of Minor Groove BinderOligonucleotide Conjugates
Hoechst-type MBs The Hoechst-type MBs are based on a benzimidazole dye structure with interesting biophysical properties (Hoechst 33258; Fig. 8.4.1). This compound is known to prefer A/Trich binding sites in dsDNA, but analogs have been prepared that show some G/C tolerance (Singh et al., 1992). The compounds are moderately fluorescent in aqueous solutions, but quantum yield increases some 20-fold in the presence of dsDNA. The DNA binding and fluorogenic properties of the Hoechst-type MBs made them interesting candidates for targeted delivery by ODNs. Enhanced stability was first explored by preparing TFOs with tethered Hoechst-type MBs (Robles et al., 1996; Robles and McLaughlin, 1997). As described above for binding mode 3, a long linker between the MB and ODN is a critical component for the TFO binding mode to be successful. An 18-atom hexa(ethylene glycol) linker was used, and the resulting three-stranded complex showed increased stability (Tm values of the triplexes increased by as much as 18°C). In addition, these three-stranded complexes
showed increased fluorescence, consistent with minor groove binding. Duplex-stabilizing effects with internally linked Hoechst-type MB-ODNs were also explored (Wiederholt et al., 1996, 1997). As described above for binding mode 2, this approach allows several potential types of complexes to be formed. The linker structure of the internal modification is shown in Figure 8.4.2. The ODN conjugates were formed by first introducing a protected thiol linker at an internal phosphate through a phosphoramidate bond. This introduced two stereoisomers at this position that influence the direction in which the MB is oriented (towards the major or minor groove). The MB-ODN was prepared by reacting the thiol with a bromoacetyl derivative of the Hoechst-type MB. Despite difficulties described for isolation, enough conjugate was isolated to study the hybridization properties. These internal MB-ODNs showed increased duplex stability, with dramatic differences between the stereoisomers (increased Tm values of 6°C versus 18°C). Increased fluorescence upon hybridization was consistent with these increases in MB binding efficiency. Finally, duplex-stabilizing effects with terminally linked Hoechst-type MB-ODNs were explored (binding mode 1). The same 18-atom hexa(ethylene glycol) linker that was developed for triplex targeting was used for the duplex system (Rajur et al., 1997). Introduction of the tethered MB to the 5′ terminus of the ODN was accomplished by forming a 5′-phosphoramidite on the protected ODN after DNA synthesis. This was treated with the alcohol form of the linker-MB, and the conjugate was deprotected with ammonia as usual. This resulted in better yields of Hoechst-type MBODNs, and allowed various duplex systems to be studied. As expected, increases in duplex stability were observed. The Tm increased as much as 14°C for A/T-rich 15-mer sequences. There was little increase in binding to G/C-rich targets. The differences in Tm corresponded to differences in fluorescence (higher Tm gave greater fluorescence). It was noted that the long linker allowed the MB to reach A/T-rich binding sites far removed from the terminus. A shorter tri(ethylene glycol) linker allowed the tethered MB to reach binding sites four bases removed from the end of the duplex. As described above, this can improve binding affinity for some sequences, but the stabilization is unpredictable.
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F method A solution-phase synthesis
H
F
F
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O
ODN-NH 2 N
N
F H
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DMTrO
ODN
O
O method B solid-phase synthesis
DPI3
NH2 DMSO, triethylamine
N H
N
DNA synthesis
O O
linker- CPG
H O
Figure 8.4.5 Synthesis methods for DPI3-type MB-ODNs. Method A involves acylation of aminemodified ODNs using DPI3-activated esters. Method B uses DPI3-modified glass beads that allow rapid synthesis of MB-ODNs with only slight modification of standard automated DNA synthesis methods. DPI3-ODNs are easily purified by reversed-phase HPLC methods. Both 5′- and 3′-labeled conjugates can be synthesized from the same CPG reagent by using the appropriate 5′ or 3′ phosphoramidite nucleoside bases. CPG, controlled-pore glass; DMSO, dimethyl sulfoxide; DMTr, 4,4′-dimethoxytrityl.
CPI-type MBs As described for binding mode 3, MBs containing the alkylating subunit of CC-1065 have been used to site-specifically alkylate certain DNA sequences. It is amazing that nature evolved molecular machinery that could create such efficient dsDNA-targeting molecules. The left-hand subunit of the CPI-DPI-type MBODN in Figure 8.4.2 contains a hybridizationtriggered cyclopropyl functional group that can alkylate N3 of adenine in dsDNA. A single “conjugatable” CPI subunit was developed, ODN conjugates were prepared, and ssDNA targets were alkylated with the simple CPI subunit (Lukhtanov et al., 1996b). In a later report, a detailed analysis of the kinetics of alkylation of ssDNA and targets showed the sequence specificity of various CPI and CPIDPI analogs (Lukhtanov et al., 1997). Triplex binding and alkylation with CPI-type MBODNs required the additional DPI subunit (see Fig. 8.4.2) for efficient alkylation (Lukhtanov et al., 1997). The CPI-type MBs are more aptly described as minor groove “bonding” agents, and are not discussed further in this unit.
SYNTHESIS AND HYBRIDIZATION OF DPI3-TYPE MB-ODNs Conjugation Chemistry As described earlier, conjugation of the MPC peptides to short ODNs presented technical hurdles (Sinyakov et al., 1995). MPC derivatives containing an aminohexanoyl linker were prepared with one to five MPC subunits. Conjugation was accomplished by activating a 3′ phosphate on A8, T8, or mixedsequence 8-mer ODNs in organic solvents and utilized the organic-soluble cetyltrimethylammonium (CTAB) salts of the ODNs. Since the simple MPC analogs lacked the positively charged amidine group present in distamycin, organic solvent (dimethylformamide) was required to dissolve the hydrophobic MPC derivatives. Preparation and handling of the CTAB salts was not described in detail, but isolated yields of 30% to 50% were reported. Despite the complexity of synthesis, distamycin-type MB-ODN conjugates were isolated as pure organic compounds. Purification of the conjugates was aided by a dabcyl chromophore (R group in Fig. 8.4.2) that was present on the C-terminal MPC subunit. Preparation of DPI-activated esters (Lukhtanov et al., 1995) allowed easy access to the convenient conjugation methods shown in Figure 8.4.5. The simple formation of amide
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5′-phosphate
Figure 8.4.6 Solution structure of the minor groove binding site in a DPI3-ODN/DNA duplex was determined by NMR. The rendering above shows the 5′-DPI3, the linker structure, and the terminal T-A base pair as solid bonds in a space-filling DNA duplex. The DPI3 covers 5 to 6 bp in the minor groove. A 5′-DPI3-ODN with sequence 5′-TGATTATCTG was made by method A (Fig. 8.4.5) and hybridized to a complementary DNA strand. The DPI3-ODN/DNA duplex had the same B-form conformation as an unmodified 10-mer DNA duplex (Kumar et al., 1998). The atomic coordinates for this structure were deposited in the Protein Data Bank as PDB ID: 1AUL.
Chemistry of Minor Groove BinderOligonucleotide Conjugates
bonds in organic solvents was easily adapted to synthesis of ODN conjugates since an aminohexyl linker arm can be conveniently added to either end of the strand during DNA synthesis. The linked DPI subunits have strong UV absorbance at ∼340 nm, which simplifies analysis of the conjugation chemistry and aids in characterization of the DPI-type MB-ODNs. The DPI-type MBs are more hydrophobic than the distamycin-type MBs, and reversed-phase HPLC is a convenient method for analysis and purification of DPI-type MB-ODNs. Conjugation method A used solution-phase conjugation chemistry (in organic solvents). The described method (Lukhtanov et al., 1995) still required that CTAB salts of the ODN be prepared, but formation of amide bonds by reacting the DPI3-activated ester with aminohexyl-modified ODNs was simpler than the distamycin-type MB conjugation chemistry. The solution-phase conjugation chemistry was improved significantly when it was found that triethylammonium (TEA) salts of ODNs are soluble in anhydrous dimethyl sulfoxide (DMSO; Milesi et al., 1999). The purified TEA salts of ODNs are easily isolated by reversedphase HPLC, dried in vacuo, and treated directly with activated esters. The method was described in detail for preparation of DPI3-
ODNs. This acylation method couples HPLCpurified aminohexyl-ODNs with DPI3 TFP (tetrafluorophenyl) or PFP (pentafluorophenyl) ester to give the HPLC-purified DPI3ODN as a single, well-resolved peak. Conjugation method B describes DNA synthesis directly from DPI3-modified controlledpore glass (CPG) supports. This method is the most versatile if many different DPI3-ODN sequences are required, since each synthesis is performed in a separate column and can be easily automated. A flexible aminodiol linker allows simultaneous attachment of the MB to a dimethoxytrityl (DMTr)–protected hydroxyl group that forms the starting point for DNA synthesis. If standard phosphoramidite chemistry is used, 3′-DPI3 conjugates can be prepared. If 5′-DPI3 conjugates are desired, reverse (3′-DMTr) phosphoramidites can be used. The DPI3 is able to survive optimized DNA synthesis conditions. For example, iodination of the DPI subunits during the oxidation cycle is a possible side reaction for the DPI3-ODNs, but can be prevented by using a lower concentration of iodine. It was noted that truncated failure sequences also contain the hydrophobic DPI3, but these side products (and those corresponding to damaged DPI) are easily separated by reversed-phase HPLC purification. Various
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linker structures were used in construction of other DPI3-CPGs. Another method described direct peptide coupling of the DPI subunits on CPG to prepare the DPI3-CPG (Lukhtanov et al., 1996a). Although the method simplified assembly of the DPI-type MBs, excess DPI reagent was required at each peptide coupling step, and quality was more difficult to control. A related conjugation method involves treatment of the CPG-bound ODN after removal of a 5-monomethoxytrityl (5-MMTr) protecting group from an aminohexyl linker arm (Lukhtanov et al., 1995). This method has the advantage that the heterocyclic bases in the immobilized ODN strands are still protected, and allows the use of organic solvents to introduce the DPI-type acylating agents. The disadvantage of this method is that the DPI3 must be able to survive the harsh ammonia deprotection conditions. Fortunately, the amide bonds in the DPI3-ODNs survive these conditions with little hydrolysis. In summary, all of the DPI conjugation methods are useful, and selection of a synthetic strategy depends on the scale and desired level of purity. For example, method A proved to be the most efficient for larger-scale synthesis of a 5′-DPI3 10-mer ODN for NMR studies (Kumar et al., 1998). In this synthesis method, the valuable DPI3 TFP ester was used in only threefold excess, and the DPI3 was not subjected to harsh deprotection or DNA synthesis conditions. The MMTr-on oligonucleotide was purified by HPLC, deprotected, and repurified by HPLC. A yield of 4 mg of the DPI3-ODN conjugate was isolated as the sodium salt after precipitation from sodium perchlorate in acetone (42% yield starting from the amine-modified ODN).
Structure of a DPI3-ODN/DNA Duplex A 5′-linked DPI3-ODN conjugate with sequence 5′-TGATTATCTG was hybridized to a complementary ODN strand and the solution structure was determined by NMR as shown in Figure 8.4.6 (Kumar et al., 1998). Under suitable aqueous conditions, the complementary DNA strands first assemble by natural WatsonCrick hydrogen bonding to the preferred Bform double helix. The 5′-linked hydrophobic DPI3 explores conformational space (i.e., it swings around) and positions itself in the newly formed (hydrophobic) minor groove. Analysis by NMR showed that the DPI3-ODN/DNA duplex has the same B-form structure as an unmodified 10-mer duplex of the same sequence. The crescent-shaped DPI3 is a “perfect
fit” for B-form DNA, and stabilizes the duplex by forming close contacts with both DNA strands in the minor groove. The electron density of the hydrophobic DPI3 is well-contained within the sugar-phosphate “walls” of the minor groove. The NMR structure of the DPI3ODN duplex in solution shows the minor groove–binding region covers 5 to 6 bp of the DNA duplex. As described below, thermodynamic analysis shows that DNA duplex stabilization provided by the conjugated DPI3 molecule is due mainly to an increase in the entropy (∆S) of the hydrated duplex system (see Thermodynamics of Hybridization). The minor groove of B-form DNA contains different types of oriented interand intra-strand water bridges (water of hydration; Savage, 1993). Crystallography of highly hydrated B-form DNA duplexes shows a wellordered “spine of hydration” in the minor groove of A/T-rich sites (Dickerson et al., 1982). Although interaction of DPI3 in the minor groove implies a certain degree of unfavorable loss of entropy, the total hydrated system increases in entropy. This entropy compensation is a result of two different entropic factors. First, an increase in entropy accompanies expulsion of the well-ordered water of hydration in the minor groove. A single crescent-shaped DPI3 molecule presumably displaces ten to thirteen organized water molecules in the spine (Dickerson et al., 1982). Second, as mentioned above, the organized water interface around the unbound hydrophobic DPI3 is significantly reduced. After binding, the planar surface area of DPI3 is shielded in the minor groove, and a simplified water layer can form on top of the molecule in the binding region. The NMR structure shows the hydrophobic DPI3 molecule deeply buried within the A/T-rich minor groove. For G/C-rich binding sites, the N2 group of guanine protrudes into the minor groove and prevents the deep binding of DPI3. In that case, the geometry of the water layer over the MB site is more complex, and entropic stabilization of the system is decreased.
HYBRIDIZATION PROPERTIES OF DPI3-ODNs DNA Duplex Stabilization The dsDNA binding properties of the DPItype MB-ODNs have many similarities to those of distamycin-type MB-ODNs. The UV melting curves of DPI-ODNs in Figure 8.4.7 show the same increase in DNA binding affinity with increasing numbers of DPI subunits shown in
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0.08
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T m(°C) Figure 8.4.7 Differential UV melting curves for complexes formed with DPI-type MB-ODNs. dT 8-mers with the following 3′-DPIn conjugate groups were melted with poly(dA) in 140 mM KCl, 10 mM MgCl2, and 20 mM HEPES⋅HCl (pH 7.2). (1) no DPI, (2) DPI1, (3) DPI2, (4) DPI3. The concentration of each ODN was 2 × 10–6 M and the A/T ratio was 1:1. Curve 5 shows the melting of the DPI2 conjugate alone in the same buffer. Anomalies in the melting curves may stem from self-association and/or uncharacterized minor transitions. Reprinted from Lukhtanov et al. (1995) with permission from the American Chemical Society.
Table 8.4.1 Melting Temperatures (°C) of MB-ODN Duplexes Formed by Poly(dA) and Poly(rA) with Octathymidylate Strands Terminally Linked to DPI1-3 MBsa
Poly(dA) Octathymidylate derivative (dTp)8-O(CH2)6NH2 (dTp)8-DPI1 (dTp)8-DPI2 (dTp)8-DPI3 NH2(CH2)6-(pdT)8 DPI1-(pdT)8 DPI2-(pdT)8 DPI3-(pdT)8
Tm 25 34 50 68 (65) 24 31 49 68
Poly(rA)
∆Tmb
9 25 43 (40) 7 25 44
Tm 13 18 —c 32 (31) 12 14 —c 35
∆Tmb
5 19 (18) 2 23
aThe MB conjugates were prepared by method A (Fig. 8.4.5). For comparison, data
Chemistry of Minor Groove BinderOligonucleotide Conjugates
obtained with (dTp)8-DPI3 conjugates synthesized by method B are presented in parentheses. bDifference between ODNs with and without conjugated MBs. cNo melting transition was observed.
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60
DPI3-ODN ODN
50
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8 A/T
Figure 8.4.8 Effect of A/T content on melting temperature (Tm) of 3′-modified DPI3-ODNs versus unmodified 8-mer ODNs is shown. Unmodified ODNs (light bars) show a linear decrease in Tm as A/T content increases, while DPI3-ODNs (dark bars) show only a slight drop in Tm (Tm leveling effect). Sequences (5′ to 3′) of the 8-mers are as follows: GCCGCCGC, GTCGCCGC, GTCGCTGC, GTCACTGT, GTTACTGT, GTTACTAT, GTTATTAT, ATTATTAT. The concentration of each ODN was 1 × 10–6 M, and the buffer used was 0.5× SSPE.
Figure 8.4.4A. It is noteworthy that DPI3 and MPC5 have similar Tm values when hybridized to poly(dA) targets, and that the crescentshaped conformation of each MB structure fits in a 5- to 6-bp length of dsDNA. The DPI 8-mers described in Figure 8.4.7 were synthesized by postsynthetic conjugation of a 3′hexylamine-modified ODN with the TFP ester (Fig. 8.4.5, method A). The DPI3 8-mer gave the most stable complex (Tm = 68°C). Other hybridization properties of DPIODNs are apparent from the list of Tm data in Table 8.4.1. The conjugate formed with the linker structure from method B (synthesis from DPI3-CPG) showed a similar Tm (65°C), indicating that the DNA synthesis and ammonia deprotection conditions did not compromise the structural integrity of the DPI3. This was an important finding, since it validated that synthesis of 3′-DPI3-ODNs could be efficiently prepared from CPG supports. As described above, this is a major advantage of the DPI3type MBs, since it allows a variety of ODN sequences to be easily accessed by automated
DNA synthesis. It was noted that 5′-DPI3ODNs could be prepared from DPI3-CPG by using 5′-phosphoramidites instead of the standard 3′-phosphoramidites in DNA synthesis (Lukhtanov et al., 1995).
Backbone Effects on Hybridization Another advantage of the DPI3-ODNs is improved binding to RNA targets. Both 3′- and 5′-DPI3 conjugates increased stability with poly(rA) target strands, but the stabilization was not as great as with DNA targets (Table 8.4.1). Surprisingly, DPI2-ODN conjugates formed no detectable hybrids with poly(rA). It is not clear why this phenomenon was observed, but the DPI2 8-mer ODNs appeared to self-associate as shown in melting curve 5 in Figure 8.4.7. The stabilization of RNA by certain DPI-type MBs has been reported. For example, unconjugated CC-1065 was shown to bind with RNA targets (Kim et al., 1995a) and could improve antisense activity of ODNs (Kim et al., 1995b). DPI3-ODN conjugates were studied with other nucleic acid backbones
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such as phosphorothioates and 2′-O-methyl RNA (Kutyavin et al., 1997). In virtually all systems studied, DPI3-ODNs stabilized duplexes when conjugated at either the 3′ or 5′ terminus. However, the duplexes with DNA targets were much more stable. This is consistent with the preferential binding of low-molecular-weight DPI analogs with B-form DNA duplexes.
Base Composition Effects on Hybridization (Tm Leveling) Most of the early studies on MB-ODNs used dT 8-mer ODNs with an MB at either the 3′ or 5′ terminus. Since these early studies were used to optimize the MB conjugation chemistry, it was easier to focus on a single strong binding sequence for development of the synthetic chemistry. However, a G/C-rich 3′-MB conjugate with the sequence CATCCGCT-MB was prepared for both the distamycin-type and the DPI3-type MB (Kutyavin et al., 1997). Melting curve analysis of 8-mer duplexes showed no
increase in stability with the distamycin-type MB-ODNs, whereas the DPI3-ODN gave a 16°C increase in Tm. This stabilization of G/Crich duplexes is a major advantage of the DPI3ODNs. With the availability of a standardized DPI3CPG support chemistry, various DPI3-ODN sequences could be readily prepared via automated DNA synthesis. A more rigorous study of DNA binding affinity versus base sequence was undertaken. DPI3 molecules have an affinity for A/T-rich sequences, and this same property is observed in the DPI3-ODNs. Since the linker between DPI3 and the ODN is short, the end of the DPI3 does not stretch past six base pairs. If the six terminal bases at the linker end of the ODN are A/T rich, then this duplex will likely make a good DPI3 binding site. Figure 8.4.8 shows the huge increase in affinity that can be attained with A/T-rich DPI3-ODNs. The all-A/T DPI3-ODN had a Tm of 48°C, whereas the corresponding unmodified ODN had a Tm of 4°C. The stepwise increase in ∆Tm as the
7 DPI3-ODN ODN 6
∆∆G °50 (kcal/mol)
5
4
3
2
1
0 T/G A/C A/C G/T T/G A/C G/T
A/C C/A A/C T/G A/C
A/C
DPI3 site
Chemistry of Minor Groove BinderOligonucleotide Conjugates
Figure 8.4.9 Relative free energy difference between match and mismatch. Detection of mismatches by a 3′-modified 13-mer DPI3-ODN (light bars) and an unmodified 13-mer ODN (dark bars) for various DNA targets. UV melting experiments were used to calculate a free energy difference (∆∆G°50) for each mismatch type and location. Mismatch discrimination for each duplex is shown in relationship to the DPI3 binding site. The sequence of the 13-mer is TAAGTAGACATAA.
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Table 8.4.2 Accuracy of Predicted Increases in DNA Duplex Stability for Various Fluorogenic 3′-DPI3-ODNs
Sequencea (5′ to 3′ direction)
Calculated Tm Tm (°C)b Tm (°C)b ODN DPI3-ODN (°C)c DPI3-ODN
Error (°C)
CTGTAAGTAGATATAAC GGCAAGATATATAG GTGACGCAGATTCC GTAAGTAGACATAAC CAGGGAGCTTTGGA CACTCGTGAAGCTG GTAAGTAGGCATAAC CCGGATGTAGGATC GATTACCTGGATTT CGGCTACAGCTGG GTTCATGGGTGTAAT
51.8 50.2 61.3 52.1 59.9 60.9 55.7 57.5 50.6 61.1 57.5
0.8 –0.8 –1.8 –1.3 –2.9d –1.7 –0.9 0.7 0.1 1.4 1.9
65.9 66.4 77.0 64.6 74.4 74.0 66.9 69.3 62.3 71.6 66.9
66.7 65.6 75.2 63.3 71.5 72.3 66.0 70.0 62.2 73.0 68.8
aRepresentative sequences from a list of 50 test ODNs are shown. All DPI -ODNs were modified with 5′ 3
fluorescein and a 3′ quencher as shown in Figure 8.4.10. DPΙ3-ODN sequences contained a G analog (PPG). bMelting temperatures (T ) of ODNs and DPI -ODNs were determined from A m 3 260 melting curves using PCR buffer and a DNA duplex concentration of 1 µM. cThermodynamic parameters (∆H and ∆S) were determined for each duplex sequence using nearest-neighbor base pair models. Entropy (∆S) for each DPI3 binding site was determined and used to calculate the predicted Tm. dLargest error obtained for all sequences tested.
DPI3 binding site becomes more A/T-rich is especially striking and resulted in a “Tm leveling” effect for this series of 8-mer sequences.
Mismatch Discrimination with DPI3-ODNs Short DPI3-ODNs have another advantage for detection of mismatches in DNA targets. As shown in Figure 8.4.9, if there is a mismatch in the DNA duplex, especially in the DPI3 binding site, the DPI3-ODN/DNA duplex is greatly destabilized. This can be used to advantage for detecting certain “difficult” mismatches. For example, T/G mismatches are known to be difficult to detect with natural DNA probes. Figure 8.4.9 compares probe specificity versus mismatch position for 13-mers with and without DPI3 at the 3′ terminus. The higher values for ∆∆G of formation for the DPI3-ODN 13mer indicate higher specificity. The T/G mismatch under the DPI3 site is almost three times easier to detect (i.e., has an almost three-foldhigher ∆∆G) in comparison to the unmodified ODN. Note that the comparison of 13-mer DPI3ODNs in Figure 8.4.9 does not account for the fact that much longer (and less discriminating) unmodified ODNs would be required to reach the Tm of the DPI3-ODNs. The Tm of the perfect match 13-mer DPI3-ODN is 60°C, whereas the
Tm of the unmodified ODN is 42°C. In order to get maximum mismatch detection with the unmodified ODN, the assay would need to be run at a temperature 18°C lower than that used for the DPI3-ODN. As pointed out below (see Thermodynamics of Hybridization), hybridization assays such as PCR require the higher temperatures where DPI3-ODNs perform best.
Thermodynamics of Hybridization The effects of the sequence and structure of DPI3-ODNs on hybridization performance was rigorously examined by UV melting curve analysis. As described above, DPI3-ODNs hybridize to A/T-rich ssDNA targets more strongly than to G/C-rich targets, thus leveling the Tm values of DPI3-ODN/DNA duplexes. A/T-rich targets require long ODN sequences since A-T base pairs bind more weakly. The shorter DPI3-ODNs gave better mismatch discrimination than longer unmodified ODNs as described below. The objective of the authors’ research was to develop methods for predicting hybridization properties of DPI3-ODNs and to improve their performance in hybridizationbased assays. Thermodynamic parameters were determined that allow accurate prediction of melting temperatures of DNA duplexes formed with MB probes. Fluorogenic 3′-DPI3ODNs (see next section) were used for these
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F
DNA target
Taq digestion F 3′
MGB TaqMan mechanism
DNA target hybridization
5′
MB
Q
MB
Q
Q
F
F
Q MB
MGB Eclipse mechanism
MB
Figure 8.4.10 Typical assay formats for hybridization detection using fluorogenic DNA probes. F is a fluorescent reporter and Q is a nonfluorescent quencher. Quench release by nuclease digestion (TaqMan mechanism; Roche and Applied Biosystems) or hybridization (Eclipse mechanism; Epoch Biosystems) gives a fluorescent signal. The stylized MB-ODN structure shows a coiled ODN conformation with MB, Q, and F in close contact. The two mechanisms are described in detail in Afonina et al. (2002).
Chemistry of Minor Groove BinderOligonucleotide Conjugates
experiments since there are commercial applications for PCR probes with these specific structures. DPI3-ODNs hybridize with A/T-rich targets to give B-form DNA duplexes. As described above, the hydrophobic DPI3 molecule stabilizes DPI3-ODN/DNA duplexes due to the increased entropy of the hydrated duplex system. Base pairing is not disturbed in the minor groove binding region (5 to 6 bp) and conformation of the sugar-phosphate backbone is not distorted. Analysis of UV melting curves and Van’t Hoff plots showed that the decreased free energy (more negative ∆G) of the DPI3ODN/DNA duplex is mainly due to an increase in entropy (less negative ∆S). These entropic p ar am eters were measured for DPI3ODN/DNA duplexes with varying A/T- and G/C-rich sequences. Specific sequences of ODNs were used to develop parameters for all possible DPI3 binding sites. Since the B-form conformation and natural Watson-Crick bonding are undisturbed for DPI3-ODNs, thermodynamic parameters for DNA duplex formation could be used in the unbound region. The derived thermodynamic parameters for DPI3-ODN/DNA duplex and natural DNA duplex binding were used to develop computer software that can accurately predict Tm values for fluorogenic DPI3-ODNs. Additional ther-
modynamic parameters were derived for a nonaggregating pyrazolo[3,4-d]pyrimidine guanine analog (PPG) in DNA duplexes since it was found that PPG enhanced hybridization performance of G-rich ODNs and DPI3-ODNs (Kutyavin et al., 2002). Table 8.4.2 shows experimental and calculated Tm values for selected sequences. The accuracy of the prediction program is generally within 2°C. Although the thermodynamic parameters were first derived for 3′-DPI3-ODN/DNA duplexes, parameters have since been derived for fluorogenic 5′-DPI3-ODN conjugates. Details of these experiments will be described elsewhere. Tm prediction algorithms for fluorogenic 3′− DPI3-ODNs are incorporated in TaqMan MGB probe design software (Applied Biosystems). Algorithms for fluorogenic 5′-DPI3-ODNs are incorporated in MGB Eclipse probe design software (Epoch Biosciences).
APPLICATIONS OF DPI3-ODNs IN PCR ASSAYS In contrast to longer unmodified ODNs, DPI3-ODNs allow accurate mismatch detection at high temperature. At low temperatures, DNA targets can ball up or aggregate. These intramolecular structures can prevent accurate genetic analysis of many sequences. For instance, PCR is the most widely used DNA
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12-mer probe (3′-DPI3)
Relative fluorescence units
27-mer probe
Tm (match) = 65°C Tm (C/T mismatch) = 61°C 10
10
match
8 6
Tm (match) = 66°C Tm (C/T mismatch) = 46°C
mismatch
6
4
4
2
2
0
0
match
8
10 20 30 40 PCR cycle number
50
0
mismatch 0
10 20 30 40 PCR cycle number
50
Figure 8.4.11 Effect of 3′-DPI3 on single-base mismatch discrimination in the TaqMan assay. Fluorogenic 27-mer and 12-mer (3′-DPI3) probes with similar Tm values were prepared (Kutyavin et al., 2000). The sequence of the 12-mer probe was FAM-GGCAATTTAAAG*-DPI3 and the sequence of the 27-mer probe was FAM-GGATCCAAAGAATTGGGCAATTTAAAG*-DPI3, where FAM is fluorescein amidite, the underline indicates the site of the C/T mismatch, and G* indicates the attachment point of a fluorescent quencher (TAMRA). Melting studies with mismatched complements showed improved discrimination with the shorter 3′-DPI3 probe. Fluorogenic PCR was performed with an extension temperature of 60°C. Each diagram shows a real-time PCR fluorescent curve with either match (upper curve) or mismatch (lower curve) plasmids as templates.
amplification method. PCR requires short DNA primers to bind to DNA targets at high annealing and extension temperatures (ideally ∼65°C) to prevent false priming. Although PCR works well, very long primers are sometimes required to bind at these high temperatures. Even at these high temperatures, some undesired complexes can remain in the target or probes. If the DNA target is inaccessible to primers, then PCR can be inefficient. PCR applications of DPI3-ODN probes and primers were explored soon after the DPI3-type MB-ODNs were developed. It was found that 5′-DPI3 primers could be extended by Taq polymerase, and primers as short as 8-mers were useful (Afonina et al., 1997). Another application took advantage of the inability of Taq polymerase to efficiently process through a 5′-DPI3-ODN probe (Afonina et al., 1996). When Taq polymerase encountered a blocking 5′-MB-ODN “clamp”, PCR was arrested. This sequence-specific clamping of PCR with DPI3ODNs only occurred when Taq polymerase encountered the DPI3-ODN probe from the 5′ end, and only when primer extension temperature was low enough to allow efficient binding of the DPI3-ODN.
Real-Time PCR with Fluorogenic DPI3-ODN Probes DNA probes or primers with fluorescent labels have a wide range of applications in molecular biology laboratories. Fluorescence has rapidly replaced radioactivity as a label in hybridization assays. Fluorescence is easily measured, presents few safety hazards, and can be measured in a multiplexed detection scheme, allowing in-process controls or higher assay throughput. Recently, molecular biologists and nucleic acid chemists have developed versatile hybridization assays that use fluorogenic probes. These assays are especially useful for researchers studying clinical samples since the closed-tube format reduces sample handling and possible contamination. Fluorogenic probes such as hairpin probes (Tyagi et al., 1998), 5′-nuclease probes (Livak et al., 1995), or Invader probes (Lyamichev et al., 1999) can be used in thermal-cycling fluorometers for quantitation of DNA samples. Fluorogenic probes can also be used economically with simple fluorescence plate readers for end-point analysis (de Kok et al., 2002). In the real-time PCR assay, fluorescent signal is measured at each PCR cycle and the curve generated from these data can be analyzed to determine the starting copy number of DNA. This technique
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has become standard in modern molecular biology (Walker, 2002). Fluorogenic DNA probes function in one of two molecular mechanisms as shown in Figure 8.4.10. In either format, sensitivity relies on efficient quenching of the intact fluorogenic probes. A molecular fragment with desired fluorescent properties (fluor, F) is linked to the DNA probe of interest, and a quenching molecule (Q) is attached to the other terminus. Quenching of the excited state of the fluor involves fluorescence resonance energy transfer (FRET). FRET has historically been used to measure distances between suitable fluor and quencher pairs (see UNIT 11.10), but has become widely used with fluorogenic probes for DNA detection assays (Didenko, 2001). This quenching mechanism requires the fluor and quencher to pass within a specific distance (the Forster radius; see UNIT 11.10) during the lifetime of the excited state of the fluor. Suitable fluors and quenchers are selected based on overlap of the emission spectrum of the fluor and the absorbance spectrum of the quencher. Recently, the quenching mechanisms of FRET probes have been explored and nonfluorescent quenchers have been described (Johansson et al., 2002).
Improved Sequence Specificity of Fluorogenic DPI3-ODN Probes in a Model PCR System
Chemistry of Minor Groove BinderOligonucleotide Conjugates
To demonstrate the utility of fluorogenic DPI3-ODNs in real-time PCR, probes with and without DPI3 were compared in the 5′-nuclease (TaqMan) assay (Kutyavin et al., 2000). In these early experiments, TAMRA was used as a quencher molecule and attached between the ODN and the terminal DPI3 via a 3′-terminal PPG linker (see UNIT 1.8). Probes were designed to bind to the same strand in the center of an 81-bp PCR product. The model PCR system used two plasmid templates that varied by only a single base pair. Each fully matched probe was designed to have a Tm value close to the optimal temperature of the Taq polymerase extension step (65° to 72°C). Comparison of a 12-mer DPI3-ODN with an unmodified 27-mer ODN showed similar Tm values for the matched duplexes, but a dramatic difference in mismatch discrimination (see Figure 8.4.11). The 12-mer DPI3-ODN showed a 20°C drop in Tm for the mismatch target, whereas the 27-mer ODN showed only a 4°C drop in Tm. The short DPI3ODN probe had much better mismatch discrimination than the longer unmodified ODN.
The fluorogenic probes used in Figure 8.4.11 were compared for their ability to function in the 5′-nuclease assay. PCR was conducted with a two-step cycle, with annealing and extension at the same temperature. In separate experiments (Kutyavin et al., 2000), primer extension temperature was varied between 55°C and 70°C, and the DPI3-ODN showed excellent discrimination over the entire temperature range. Ideally, the Tm of the probe with perfectly matched template should be slightly higher than the extension temperature, and the Tm of a probe with the single nucleotide mismatched template should be lower. Because of the large ∆Tm for the DPI3-ODN probes, these criteria were easily met. The real-time PCR fluorescence curves for the DPI3-ODN 12-mer probe showed excellent mismatch discrimination. In contrast, discrimination by the ODN 27-mer probe was difficult to achieve. Some discrimination was seen at 67°C, but the increase in fluorescence was low. At 65°C the 27-mer showed poor discrimination, and at 70°C there was no signal. Another remarkable feature of the DPI3ODN probes was the low background fluorescence (at cycle 1). As seen in Figure 8.4.11, the background of the intact DPI3-ODN probe was several times lower than that of the 27-mer ODN probe. Quenching of 5′-fluorescein emission by FRET requires close proximity of the reporter and quencher dyes during the excited state lifetime of the reporter fluor. The random coil structure of the probes in solution allows FRET despite the long distance between the dyes (Livak et al., 1995). The lower background observed for the DPI3-ODN probe is presumably due to the short probe length, but there are other structural factors that contribute to more efficient quenching (Johansson et al., 2002). As a result, the dynamic range of fluorogenic DPI3ODN probes in real-time PCR curves is much greater than with conventional unmodified ODN probes. The short length and efficient quenching of fluorogenic DPI3-ODN probes also make them ideal as “hybridization-triggered” probes (Fig. 8.4.10, Eclipse mechanism). Recently, application of this technology to real-time PCR has been reported (Afonina et al., 2002). Since the 5′-DPI3-ODN Eclipse probes are not digested during PCR (Afonina et al., 1996), the amplified DNA strands can be analyzed in the same wells by fluorogenic melting curve analysis. This allows PCR to be run in standard thermal cyclers, and more accurate data analysis to be
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obtained on various commercial thermal-cycling fluorometers.
SUMMARY Studies of MB-ODNs have shown that they can have advantages in comparison to unmodified ODNs. The DPI3-type MB-ODNs are preferred since the conjugation chemistry has been developed to allow rapid synthesis of various sequences. The shorter DPI3-ODNs have advantages for many hybridization assays, especially for A/T-rich DNA targets and detection of single nucleotide polymorphisms (SNPs). The biophysical properties of DPI3-ODNs have been elucidated, and this allows the Tm values for specific DPI3-ODN sequences to be predicted. Fluorogenic DPI3-ODN probes have been developed that show improved performance in real-time PCR assays, and this application is being exploited in modern molecular biology laboratories.
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Boger, D.L., Coleman, R.S., and Invergo, B.J. 1987. Studies on the total synthesis of CC-1065: Preparation of a synthetic, simplified 3-carbamoyl1,2-dihydro-3H-pyrrolo[3,2-e]indole dimer/trimer/tetramer (CDPI dimer/trimer/tetramer) and development of methodology for PDE-I dimer methyl ester formation. J. Org. Chem. 52:1521-1530. Boger, D.L., Invergo, B.J., Coleman, R.S., Zarrinmayeh, H., Kitos, P.A., Thompson, S.C., Leong, T., and McLaughlin, L.W. 1990. A demonstration of the intrinsic importance of stabilizing hydrophobic binding and non-covalent van der waals contacts dominant in the non-covalent CC1065/B-DNA binding. Chem. Biol. Interact. 73:29-52. Boger, D.L., Boyce, C.W., Garbaccio, R.M., and Goldberg, J.A. 1997. CC-1065 and the duocarmycins: Synthetic studies. Chem. Rev. 97:787828. Chang, D.K. and Cheng, S.F. 1996. On the importance of van der Waals interaction in the groove binding of DNA with ligands: Restrained molecular dynamics study. Int. J. Biol. Macromol. 19:279-285. Cory, M. 1995. DNA minor-groove binding compounds as antitumo r ag ents. Cancer Chemotherapeutic Agents: ACS Professions Reference Book Chapter 8:311-344. de Kok, J.B., Wiegerinck, E.T., Giesendorf, B.A., and Swinkels, D.W. 2002. Rapid genotyping of single nucleotide polymorphisms using novel minor groove binding DNA oligonucleotides (MGB probes). Hum. Mutat. 19:554-559. Dempcy, R.O., Kutyavin, I.V., Mills, A.G., Lukhtanov, E.A., and Meyer, R.B. 1999. Linkers designed to intercalate the double helix greatly facilitate DNA alkylation by triplex-forming oligonucleotides carrying a cyclopropapyrroloindole reactive moiety. Nucl. Acids Res. 27:29312937. Dickerson, R.E., Drew, H.R., Conner, B.N., Wing, R.M., Fratini, A.V., and Kopka, M.L. 1982. The anatomy of A-, B-, and Z-DNA. Science 216:475-485. Didenko, V.V. 2001. DNA probes using fluorescence resonance energy transfer (FRET): Designs and applications. Biotechniques 31:11061121. Inga, A., Chen, F.X., Monti, P., Aprile, A., Campomenosi, P., Menichini, P., Ottaggio, L., Viaggi, S., Abbondandolo, A., Gold, B., and Fronza, G. 1999. N-(2-Chloroethyl)-N-nitrosourea tethered to lexitropsin induces minor groove lesions at the p53 cDNA that are more cytotoxic than mutagenic. Cancer Res. 59:689-695. Johansson, M.K., Fidder, H., Dick, D., and Cook, R.M. 2002. Intramolecular dimers: A new strategy to fluorescence quenching in dual-labeled oligonucleotide probes. J. Am. Chem. Soc. 124:6950-6956. Nucleic Acid Binding Molecules
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Kielkopf, C.L., White, S., Szewczyk, J.W., Turner, J.M., Baird, E.E., Dervan, P.B., and Rees, D.C. 1998. A structural basis for recognition of A-T and T-A base pairs in the minor groove of BDNA. Science 282:111-115. Kim, D.-Y., Shih, D.S., Cho, D.Y., and Swenson, D.H. 1995a. Helix-stabilizing compounds CC1065 and U-71,184 bind to RNA-DNA and DNA-DNA duplexes containing modified internucleotide linkages and stabilize duplexes against thermal melting. Antisense Res. Dev. 5:49-57. Kim, D.-Y., Swenson, D.H., Cho, D.Y., Taylor, H.W., and Shih, D.S. 1995b. Helix-stabilizing agent, CC-1065, enhances suppression of translation by an antisense oligodeoxynucleotide. Antisense Res. Dev. 5:149-154. Kumar, S., Reed, M.W., Gamper, H.B., Gorn, V.V., Lukhtanov, E.A., Foti, M., West, J., Meyer, R.B., and Schweitzer, B.I. 1998. Solution structure of a highly stable DNA duplex conjugated to a minor groove binder. Nucl. Acids Res. 26:831838. Kutyavin, I.V., Lukhtanov, E.A., Gamper, H.B., and Meyer, R.B. 1997. Oligonucleotides with conjugated dihydropyrroloindole tripeptides: Base composition and backbone effects on hybridization. Nucl. Acids Res. 25:3718-3723. Kutyavin, I.V., Afonina, I.A., Mills, A., Gorn, V.V., Lukhtanov, E.A., Belousov, E.S., Singer, M.J., Walburger, D.K., Lokhov, S.G., Gall, A.A., Dempcy, R., Reed, M.W., Meyer, R.B., and Hedgpeth, J. 2000. 3′-Minor groove binderDNA probes increase sequence specificity at PCR extension temperatures. Nucl. Acids Res. 28:655-661. Kutyavin, I.V., Lokhov, S.G., Afonina, I.A., Dempcy, R., Gall, A.A., Gorn, V.V., Lukhtanov, E., Metcalf, M., Mills, A., Reed, M.W., Sanders, S., Shishkina, I., and Vermeulen, N.M. 2002. Reduced aggregation and improved specificity of G-rich oligodeoxyribonucleotides containing pyrazolo[3,4-d]pyrimidine guanine bases. Nucl. Acids Res. 30:4952-4959. Levina, A.S., Metelev, V.G., Cohen, A.S., and Zamecnik, P.C. 1996. Conjugates of minor groove DNA binders with oligonucleotides: Synthesis and properties. Antisense Nucleic Acid Drug Dev. 6:75-85. Livak, K.J., Flood, S.J., Marmaro, J., Giusti, W., and Deetz, K. 1995. Oligonucleotides with fluorescent dyes at opposite ends provide a quenched probe system useful for detecting PCR product and nucleic acid hybridization. PCR Methods Appl. 5:357-362.
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Lukhtanov, E.A., Podyminogin, M.A., Kutyavin, I.V., Meyer, R.B., and Gamper, H.B. 1996b. Rapid and efficient hybridization-triggered crosslinking within a DNA duplex by an oligodeoxyribonucleotide bearing a conjugated cyclopropapyrroloindole. Nucl. Acids Res. 24:683687. Lukhtanov, E.A., Kutyavin, I.V., Gorn, V.V., Reed, M.W., Adams, A.D., Lucas, D.D., and Meyer, R.B. Jr. 1997. Sequence and structure dependence of the hybridization-triggered reaction of oligonucleotides bearing conjugated cyclopropapyrroloindole. J. Am. Chem. So c. 119:6213-6225. Lyamichev, V., Mast, A.L., Hall, J.G., Prudent, J.R., Kaiser, M.W., Takova, T., Kwiatkowski, R.W., Sander, T.J., de Arruda, M., Arco, D.A., Neri, B.P., and Brow, M.A. 1999. Polymorphism identification and quantitative detection of genomic DNA by invasive cleavage of oligonucleotide probes. Nature Biotechnol. 17:292-296. Mikheikin, A.L., Zhuze, A.L., and Zasedatelev, A.S. 2001. Molecular modelling of ligand-DNA minor groove binding: Role of ligand-water interactions. J. Biomol. Struct. Dyn. 19:175-178. Milesi, D., Kutyavin, I., Lukhtanov, E.A., Gorn, V.V., and Reed, M.W. 2000. Synthesis of oligonucleotide conjugates in anhydrous dimethyl sulfoxide. Methods Enzymol. 313:164-173. Rajur, S.B., Robles, J., Wiederholt, K., Kuimelis, R.G., and McLaughlin, L.W. 1997. Hoechst 33258 tethered by a hexa(ethylene glycol) linker to the 5′-termini of oligodeoxynucleotide 15mers: Duplex stabilization and fluorescence properties. J. Org. Chem. 62:523-529. Robles, J. and McLaughlin, L.W. 1997. DNA triplex stabilization using a tethered minor groove binding Hoechst 33258 analogue. J. Am. Chem. Soc. 119:6014-6021. Robles, J., Rajur, S.B., and McLaughlin, L.W. 1996. A parallel-stranded DNA triplex tethering a Hoechst 363258 analogue results in complex stabilization by simultaneous major groove and minor groove binding. J. Am. Chem. Soc. 118:5820-5821. Savage, H.F.J. 1993. Water structure. In Water and Biological Macromolecules (E. Westhof, ed.) pp. 3-39. CRC Press, Boca Raton, Fla. Singh, M.P., Joseph, T., Kumar, S., Bathini, Y., and Lown, J.W. 1992. Synthesis and sequence-specific DNA binding of a topoisomerase inhibitory analog of Hoechst 33258 designed for altered base and sequence recognition. Chem. Res. Toxicol. 5:597-607.
Lukhtanov, E.A., Kutyavin, I.V., Gamper, H.B., and Meyer, R.B. Jr. 1995. Oligodeoxyribonucleotides with conjugated dihydropyrroloindole oligopeptides: Preparation and hybridization properties. Bioconjugate Chem. 6:418-426.
Sinyakov, A.G., Lokhov, S.G., Kutyavin, I.V., Gamper, H.B., and Meyer, R.B. 1995. Exceptional and selective stabilization of A-T rich DNADNA duplexes by N-methypyrrole carboxamide peptides conjugated to oligodeoxynucleotides. J. Am. Chem. Soc. 117:4995-4996.
Lukhtanov, E.A., Kutyavin, I.V., and Meyer, R.B. 1996a. Direct, solid phase assembly of dihydropyrroloindole peptides with conjugated oligonucleotides. Bioconjugate Chem. 7:564-567.
Szewczyk, J.W., Baird, E.E., and Dervan, P.B. 1996a. Cooperative triple-helix formation via a minor groove dimerization domain. J. Am. Chem. Soc. 118:6778-6779.
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Szewczyk, J.W., Baird, E.E., and Dervan, P.B. 1996b. Sequence-specific recognition of DNA by a major and minor groove binding ligand. Angew. Chem. Int. Ed. Engl. 35:1487-1489. Thuong, N. and Helene, C. 1993. Sequence-specific recognition and modification of double-helical DNA by olgionucleotides. Angew. Chem. 32:666-690. Tyagi, S., Bratu, D.P., and Kramer, F.R. 1998. Multicolor molecular beacons for allele discrimination. Nature Biotechnol. 16:49-53.
Wiederholt, K., Rajur, S.B., and McLaughlin, L.W. 1997. Oligonucleotides tethering Hoechst 33258 derivatives: Effect of the conjugation site on duplex stabilization and fluorescence properties. Bioconjugate Chem. 8:119-126. Zimmer, C. and Wahnert, U. 1986. Nonintercalating DNA-binding ligands: Specificity of the interaction and their use as tools in biophysical, biochemical and biological investigations of the genetic material. Prog. Biophys. Mol. Biol. 47:31-112.
Walker, N. 2002. A technique whose time has come. Science 296:557-559. Wiederholt, K., Rajur, S.B., Giuliano, J. Jr., O’Donnell, M.J., and McLaughlin, L.W. 1996. DNAtethered Hoechst groove binding agents: Duplex stabilization and fluorescence characteristics. J. Am. Chem. Soc. 118: 7055-7062.
Contributed by Igor Kutyavin, Sergey Lokhov, Eugene Lukhtanov, and Michael W. Reed Epoch Biosciences Bothell, Washington
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A Fluorescent Intercalator Displacement Assay for Establishing DNA Binding Selectivity and Affinity
UNIT 8.5
The fluorescent intercalator displacement (FID) assay is a rapid, high-resolution, and technically nondemanding technique for establishing DNA binding selectivity and affinity. The assay utilizes the displacement of DNA-bound ethidium bromide (or other fluorescent intercalator). Hairpin oligodeoxyribonucleotides are treated with the intercalator, yielding a fluorescence increase upon binding. Addition of a DNA-binding compound results in a decrease in fluorescence due to the displacement of the bound intercalator, where the percent fluorescence decrease is directly related to the extent of binding. In a 96-well format (see Basic Protocol 1), the FID assay provides for the high-throughput evaluation of a single compound against a library of DNA sequences (to establish sequence selectivity) or for the high-throughput selection of high-affinity binders for a defined sequence from a library of compounds. Alternatively, FID titrations (see Basic Protocol 2) provide detailed information on single compounds and their binding to individual sequences, including quantitative binding constants, stoichiometry of binding, and binding site size. This latter application as well as selection screening against a single sequence is amenable to examination of any sequence length, notably is not limited to small molecule assessments, and has been used with a variety of ligands, including proteins and triplex-forming oligonucleotides (for more detailed information on its application, see Tse and Boger, 2004). Unlike complementary techniques, the FID assay is nondestructive, providing the opportunity for hairpin immobilization onto reusable supports (chips, beads, or glass slides), and thus removing the barrier to comprehensive and repeated screening of longer sequences than are presently exemplified.
STRATEGIC PLANNING Hairpin Oligodeoxyribonucleotides Hairpin oligodeoxyribonucleotides (Broude, 2002; Fig. 8.5.1) have proved especially useful in the FID assay. Embedded in the hairpin are two complementary 5 -to-3 sequences connected by a loop, avoiding the requirement for two separate strands and the associated additional quantitation and mixing. The number of hairpins required to create a library of sequences is half the number of sequence permutations, if the position of the variable region and its orientation within the hairpin are not considered. For example, only 512 hairpins are required for a library of all possible 5-bp sequences, which number 1024 sequences in total (Fig. 8.5.1). Moreover, the hairpins were established to provide stable duplexes at working temperatures (i.e., 25◦ C) in UV thermal melting studies independent of the duplex sequence. Intercalator Following the characterization of the intrinsic fluorescence increase that accompanies DNA intercalation (LePecq and Paoletti, 1967), the displacement of DNA-bound ethidium bromide has been used in a variety of ways to qualitatively establish DNA binding. Ethidium is nonselective, equilibrates rapidly, and has a low binding affinity (∼105 M−1 ) that allows assessment of compounds with low binding affinity or assessment of lowaffinity sites. These aspects are not easily addressed by footprinting or electrophoretic mobility shift assay (EMSA). For tight binding sequences, the displacement of ethidium proceeds in a virtually noncompetitive manner, permitting quantitative assessments of binding. Contributed by Winston C. Tse and Dale L. Boger Current Protocols in Nucleic Acid Chemistry (2005) 8.5.1-8.5.11 C 2005 by John Wiley & Sons, Inc. Copyright
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Figure 8.5.1 Structures of hairpin oligodeoxyribonucleotides, ethidium bromide, and thiazole orange. Reprinted from Tse and Boger (2004) with permission from the American Chemical Society.
Thiazole orange (Lee et al., 1986; Nygren et al., 1998) may be used as an effective alternative intercalator that addresses three issues: (1) its excitation and emission maxima are distinct from those of ethidium, (2) its fluorescent enhancement upon intercalation exceeds that of ethidium, and (3) its binding displays less sequence dependence, albeit with higher affinity. For a more complete discussion of thiazole orange in the FID assay, refer to Boger and Tse (2001).
Ethidium Bromide Concentration The concentration of ethidium bromide in solution will be contingent on the DNA length and concentration; there should be one ethidium bromide per every two base pairs. For the example of the 5-bp variable region library used in Basic Protocol 1, there are 8 bp in the stem region (including 3 capping base pairs), which equates to four ethidium bromide molecules for every hairpin DNA oligodeoxyribonucleotide. Thus, for a 1.5 µM oligodeoxyribonucleotide solution, a final concentration of 6 µM ethidium bromide is desired. Typically, 70 µL will be added to each well, requiring an 8.57 µM stock solution to achieve a final concentration of 6 µM. The solution should be protected from light at all times as a precaution against fading/quenching. BASIC PROTOCOL 1
Fluorescent Intercalator Displacement Assay For DNA Binding
96-WELL SINGLE-POINT ASSAY In the 96-well format, the assay is rapid, comprehensive, and technically nondemanding (Fig. 8.5.2; for a review, see Tse and Boger, 2004). It may take two forms. (1) For a single compound, the technique permits establishment of a rank-order binding profile for all possible 5-bp sites, comprehensively defining the sequence selectivity in a single experiment (Boger et al., 2001; Tse et al., 2003). (2) For a defined sequence, it permits the highthroughput identification of binding agents from a library of compounds (Boger et al., 2000b). In the former assay, a single compound or select compounds are assayed against a library of oligodeoxyribonucleotides that represent all possible sequence permutations. In the latter, a single oligodeoxyribonucleotide or select oligodeoxyribonucleotides are employed to select the highest-affinity members from a library of compounds.
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Figure 8.5.2 General procedure for the 96-well FID assay. Reprinted from Tse and Boger (2004) with permission from the American Chemical Society.
The following protocol is an example for assaying a single compound at two concentrations against a library of hairpin oligodeoxyribonucleotides containing all possible 512 variations of a 5-bp variable region (Fig. 8.5.1) for determination of sequence selectivity by measuring the displacement of the fluorescent intercalator ethidium bromide. This may be easily extended to libraries of larger variable regions or for assaying a library of compounds against a single oligodeoxyribonucleotide.
Materials 8.57 µM ethidium bromide in Tris buffer Tris buffer: 0.1 M Tris·Cl, pH 8.0 (APPENDIX 2A), with 0.1 M NaCl 15 µM solutions of each DNA hairpin oligodeoxyribonucleotide in H2 O (100 µL each) Agent blank solution: Tris buffer containing 10% dimethylsulfoxide (DMSO) 7.5 µM and 10.0 µM compound of interest in Tris buffer containing 10% DMSO Nucleic Acid Binding Molecules
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96-well assay plates: tissue culture (TC)-treated, black, flat-bottom, polystyrene microtiter plates (e.g., Costar 3916) Top-reading fluorescence plate reader (e.g., Molecular Devices Spectra Max Gemini) NOTE: To benefit measurement accuracy, ensure that the same percentage of DMSO is in the agent blank solution and each agent solution (typically 10% DMSO in stock solutions).
Perform assay 1. Prepare all solutions the day prior to running the assay. 2. Add 70 µL of 8.57 µM ethidium bromide solution (final assay concentration 6 µM) to every well of the 96-well TC-treated black assay plates. Once the ethidium bromide solution has been added, protect plates from light. The number of plates will vary with the experimental design. This assay design will require 43 plates, with one column of wells for each of 512 sequences plus three columns for DNA blanks. For 1.5 µM final DNA hairpin concentration, a 6 µM ethidium bromide concentration is desired to achieve 1 molar equivalent of ethidium bromide per 2 DNA bp.
3. Add 10 µL H2 O to 24 wells (3 columns) as 0% controls. The 0% control wells contain neither DNA nor the compound of interest.
4. Add 10 µL of 15 µM hairpin DNA stock solutions (final assay concentration 1.5 µM) to the appropriate wells. For convenience, load each individual sequence into all eight wells of a single column (512 sequences will thus require 512 columns). For each of the 512 sequences, there will be duplicate 100% control wells (with DNA but no compound) and triplicates of two concentrations of the compound of interest. Note that a 512-sequence library of a 5-bp variable region represents all possible sequences if the compounds examined are not interacting with any other part of the hairpin structure. See Strategic Planning for further elaboration of this point. DNA stocks are typically pre-made and stored frozen in 96-well plates. To ensure that the hairpins are annealed properly after storage, plates are thawed, briefly heated, and cooled slowly back to room temperature. Plates are then spun down in a centrifuge equipped with microtiter plate carriers (e.g., Beckman Coulter Allegra 6R centrifuge) to remove condensation from the lids.
5. Add 20 µL of agent blank solution to two wells containing DNA for each sequence to be analyzed (100% controls). For convenience, add to rows A and B for each sequence. Also add agent blank solution to each of the 0% control wells. 6. Add 20 µL of 7.5 µM and 10.0 µM compound solutions (final assay concentration 1.5 µM and 2.0 µM, respectively) to triplicate wells for each sequence (e.g., rows C through E and F through H, respectively). Typical compound concentrations are 1.5 and 2.0 µM versus 1.5 µM DNA. These concentrations are chosen to achieve a dynamic range in which the assay shows measurable differences for very strong DNA binders without losing observable differences for weak DNA binders.
7. Allow to equilibrate for 30 min on an orbital shaker at room temperature. Fluorescent Intercalator Displacement Assay For DNA Binding
8. Read the plates using a top-reading fluorescence plate reader. For ethidium bromide, typical wavelengths are 545 nm for excitation and 595 nm for emission, with a cut-off of 590 nm. On the Gemini Spectra Max, also adjust settings for six measurement points/well sensitivity and 5 sec automix before the plate is read.
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Analyze data 9. Average the fluorescence of the 24 0% wells (ethidium bromide alone) and subtract it from the fluorescence of each substrate well and each 100% control well. This accounts for the background fluorescence and normalizes all subsequent data.
10. For each sequence, calculate the %F by dividing the median normalized compound fluorescence measurement (at each concentration) by its corresponding averaged normalized 100% measurement. The 100% readings (ethidium, DNA, no compound) are used to account for differences in ethidium affinity to DNA of differing sequences. The %F is calculated for each sequence relative to its own average 100% fluorescence value. Since the assay is a high-throughput experiment, errors are expected to occur. Aside from the concentration of the hairpin oligodeoxyribonucleotides, the next greatest source of error is perceived to be in pipetting.The key to obtaining an accurate rank-order binding profile is in minimizing the affect of aberrant data on the observed sequence selectivity. Several methods have been examined, with three (triplicate) data points preferred over two (duplicate). The method currently recommended is the use of the median value of triplicate substrate agent readings and the average value for the 100% wells.
11. Order the sequences by %F into a rank binding order. Then, determine the ensemble of sequences, if any, that the compound has affinity towards. Use one of the following two methods: a. identify an ensemble of sequences and calculate the average rank of the constituent members; or b. identify an ensemble of sequences and calculate the average score of the constituent members. A score is assigned to be reflective of its %F relative to the %F of the best sequence (i.e., the one with the lowest %F). If a sequence had a %F of 55% and the best sequence had a %F of 40%, then its score would be (100 − 55)/(100 − 40) = 0.75. By definition, the best sequence carries a score of 1. The calculated score is a better representation of the binding profile.
FID TIRATION ASSAY Quantitative displacement of ethidium bromide from a hairpin oligodeoxyribonucleotide provides a well-defined titration curve that is useful for establishing binding constants and stoichiometry of binding (Boger et al., 2001; Boger and Tse, 2001; Tse and Boger, 2004). In some cases, FID titrations may provide additional information to distinguish modes of binding and binding site size (Woods et al., 2002a,b). FID titrations have proved useful not only for small molecule assessments but for use with a variety of ligands, including proteins (Ham et al., 2003) and triplex-forming oligonucleotides (Yeung et al., 2003).
BASIC PROTOCOL 2
Materials Ethidium bromide in Tris buffer (see Basic Protocol 1) Hairpin oligodeoxyribonucleotide of interest Compound of interest at 0.05 M in Tris buffer containing 10% DMSO 3-mL quartz cuvette Fluorospectrophotometer (e.g., JY Horiba FluoroMax-3 with Hamilton autotitration apparatus) Nucleic Acid Binding Molecules
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Perform titration 1. Load a 3-mL quartz cuvette with 2.5 mL ethidium bromide in Tris buffer. Place in the spectrophotometer and begin stirring. The concentration of the ethidium bromide will be contingent on the structure and concentration of the hairpin oligodeoxyribonucleotide employed, but should yield 1 molar equivalent of ethidium per 2 DNA stem bp (see Strategic Planning).
2. Measure background fluorescence for normalized 0% reading. 3. Add hairpin oligodeoxyribonucleotide of interest to a final concentration of 1 µM. 4. Measure fluorescence for normalized 100% reading. 5. Titrate 3-µL aliquots of the compound of interest (0.05 mM in DMSO) and measure the resultant fluorescence decrease (Fx ) after a 5-min equilibration time. Add compound by manual pipetting or via syringe pump. Continue additions until the system reaches saturation and the fluorescence remains constant with subsequent compound additions.
Determine stoichiometry of binding 6. Plot the change in fluorescence (Fx ) versus equivalents of compound (ex ). This provides a titration curve from which the stoichiometry of binding may be experimentally derived mathematically.
7. Simultaneously solve the equations representing the pre- and post-saturation regions of the titration curve (Fig. 8.5.3A). Determine the point at which the two lines intersect. The molar equivalents of the compound (ex ) at this point is the experimental stoichiometry of binding. The change in fluorescence (F) at the point of stoichiometry is defined as Fsat .
Determine binding constant by Scatchard analysis 8. Calculate the concentration of free agent for the observed change in fluorescence at every titrant point (Fx ). a. Divide the change in fluorescence at each titrant point (Fx ) by the change in fluorescence at the point where DNA is saturated with ligand (Fsat ). b. For every titrant point, calculate X by dividing the molar equivalents of compound added (ex ) by the molar equivalents at saturation (stoichiometry of binding) calculated in step 7. c. For every titrant point, calculate the concentration of free agent:
where [free agent] = concentration of free agent, [DNA]T = total concentration of DNA in hairpin duplexes, and X = molar equivalents of compound versus hairpin DNA at a given titrant point. Fluorescent Intercalator Displacement Assay For DNA Binding
For every titrant point, the fractions of free agent versus DNA-bound agent can also be calculated as:
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Figure 8.5.3 (A) Titration of netropsin versus a hairpin oligodeoxyribonucleotide containing the 5-bp variable region 5 -AATTT at 1.5 µM utilizing ethidium bromide as the fluorescent intercalator. (B) Scatchard plot for determining the binding constant (Ka ). Reprinted from Tse and Boger (2004) with permission from the American Chemical Society.
9. Generate a Scatchard plot by plotting Fx /[free agent] versus Fx (Fig. 8.5.3B). Calculate the slope of the region immediately preceding complete saturation of the system, which provides the negative of the association binding constant (−Ka ).
COMMENTARY Background Information The regulation of gene expression is based on the sequence-selective recognition of nucleic acids by repressor, activator, and enhancer proteins. A full understanding of the proteins involved, the delineation of the
sequences to which they bind, and the discovery of the genes that they regulate holds significant promise in therapeutic medicine (Choo et al., 1994; Neidle and Thurston, 1994; Browne et al., 1996; Matteucci and Wagner, 1996; Neidle, 1997; Thurston, 1999). Thus,
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Fluorescent Intercalator Displacement Assay For DNA Binding
extensive efforts continue to be directed at understanding the transcriptional process, and are being increasingly directed at the discovery of small molecules that selectively bind DNA and activate (block a repressor) or inhibit (block an activator) gene expression (Knudsen and Nielsen, 1996; Trauger et al., 1996; Gottesfeld et al., 1997; Chiang et al., 1998; Werstuck and Green, 1998). Complicating such studies is the recognition that it is not a single sequence that is ideally targeted, but rather an ensemble of related sites that compose the consensus binding sequence of a nuclear receptor or transcription factor. Consequently, the understanding of known compounds or the design of new sequence-specific DNA-binding agents has been slow due to the complexity associated with understanding small molecule–DNA and protein–DNA interactions and the technically demanding techniques involved in the determination of their binding affinity and selectivity for any given sequence, much less an ensemble of related sequences. Of the techniques commonly used to establish the DNA-binding properties of small molecules and proteins, most are technically challenging, require the knowledge of specialized biochemical procedures, and are time and labor intensive. The most widely used methods are footprinting (Dervan, 1986) and affinity cleavage (Taylor et al., 1984). Because of the power of the technique, a number of such methods have been introduced, including footprinting with DNase I (Galas and Schmitz, 1978), exonuclease III (Royer-Pokora et al., 1981), MPE-Fe(II) (Van Dyke et al., 1982), 1,10phenanthroline-Cu(I) (Kuwabara and Sigman, 1987), and EDTA-Fe(II) (Tullius et al., 1987). Complementary approaches for disrupting binding (interference footprinting; Hayashibara and Verdine, 1992) by specific base or phosphate modifications have been introduced to probe base contacts and their locations. Less general techniques that capitalize on a compound’s intrinsic properties— such as DNA cleavage (e.g., bleomycin, endiynes), alkylation/thermal cleavage (Boger and Johnson, 1996; e.g., CC-1065, duocarmycins), alkylation (site-specific inhibition of in vitro transcription), or cross-linking (Rajski and Williams, 1998; Tao et al., 2000; e.g., mitomycin)—have been powerfully but restrictively applied to selected classes of molecules. Recently, the introduction of techniques for expanding the sequence coverage of footprinting (Hardenbol et al., 1997) and the use of electrophoretic mobility shift assays (EMSAs or gel retardation assays; Chaltin
et al., 2003) and even DNase I footprinting (Guelev et al., 2000; Hamy et al., 2000) for the iterative deconvolution of mixture libraries have been disclosed, expanding the scope on which they can be applied. Inherent in these methods is the characterization of the highest affinity sites within a limited-size and customdesigned segment of DNA. Similarly, the DNA-binding properties of proteins (Larson and Verdine, 1996) are typically assessed by selection screening (Blackwell et al., 1990; Blackwell and Weintraub, 1990; Thiesen and Bach, 1990), footprinting (Galas and Schmitz, 1978), or EMSAs (Chodosh et al., 1986). The former provides exhaustive sequence coverage for deducing the preferred binding site(s), but it selects only the highest-affinity sites and does not provide quantitative binding information. Footprinting and, to a lesser extent, EMSAs have been used to define, or at least refine, the binding selectivity of a protein, but their most frequent uses have been to provide qualitative distinctions and quantitative comparisons among candidate binding sites or those constructed to assess single base pair substitutions. Throughout such studies, footprinting has emerged as the most general technique for the study of either small molecule–DNA or protein–DNA binding. The authors have introduced and reviewed a complementary technique, a fluorescent intercalator displacement (FID) assay (Boger et al., 2000a; Boger et al., 2001; for a full review, Tse and Boger, 2004) useful for establishing DNA binding affinity, comprehensive sequence selectivity, and binding stoichiometry. The assay is non-destructive, technically nondemanding, and amenable to high-throughput screening. Unlike other techniques, the former feature would permit DNA (hairpin) immobilization onto reusable supports for repetitive use and expansion of the sequence space beyond that presently exemplified (all 5-bp sites). For a single compound, the technique permits establishment of a rank-order binding profile for all possible 5-bp sites, comprehensively defining the sequence selectivity in a single experiment. For a defined sequence, it permits the high-throughput identification of binding agents from a library of compounds or quantitative titrations for establishment of binding constants. The 96-well high-throughput assay and the cuvette-based titrations detailed here may be used in a complementary fashion, one intended for a high-throughput screen capable of handling large libraries of compounds or providing comprehensive sequence preference data, and the other for detailed elucidation
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of DNA-ligand interactions and establishing quantitative binding constants. The assay is not limited to small molecule assessments and has been used with a variety of ligands including proteins (Ham et al., 2003) and triplex-forming oligonucleotides (Yeung et al., 2003).
Critical Parameters Oligodeoxyribonucleotide quality. The most critical variable to the success of the assay, and most likely to be responsible for avoidable errors, is the quality of the hairpin DNAs. In addition to the obvious concern of their constitution and purity, concentration is critical and may be determined by measuring the UV absorption (260 nm) of the denatured, single-stranded DNA at 80◦ to 95◦ C. Since the hairpins exist in a construct representing a combination of double- (stem) and singlestranded (loop) DNA at 25◦ C, calculations based on the UV absorption at 25◦ C utilizing the standard coefficients for single-stranded oligodeoxyribonucleotides underestimate the concentration by as much as 25%. It has also been found that the UV absorbance (A260 ) at 25◦ C may be measured and converted to an accurate concentration by adjusting the millimolar extinction coefficients for ssDNA. Excluding neighboring effects, a 5-bp variable region library contains only six combinations of base compositions. These correspond to 5 bp A/T, 4 bp A/T and 1 bp G/C, 3 bp A/T and 2 bp G/C, 2 bp A/T and 3 bp G/C, 1 bp A/T and 4 bp G/C, and 5 bp G/C. Utilizing millimolar extinction coefficients published by Invitrogen, correction factors relating A260 at 90◦ C to A260 at 25◦ C were determined to be 1.18, 1.13, 1.12, 1.12, 1.10, and 1.06 for the combinations listed above, respectively. Blind testing found that errors resulting from these measurements were consistently <2% and always <4%. 96-well assay plates. The use of tissue culture–treated (TC-treated), black, opaque, flat-bottom polystyrene plates (e.g., Costar 3916) is recommended for the assay. It was found that TC-treated plates produced more consistent readings than non-treated plates. This is believed to be a consequence of the hydrophobic interactions between the polystyrene and the planar aromatic compounds of interest, as well as the interaction of the DMSO/buffer solution with the polystyrene. DMSO/buffer meniscus. An additional issue related to the type of 96-well plate used is the percentage of DMSO used in the assay wells. The menisci of all wells should be uni-
form for optimum consistency of the readings by a top-reading fluorometer. While readings are affected by differing types of microtiter plates, the most drastic effect results from variations in the percentage of DMSO in buffer. For best consistency of fluorescence measurements, the same percentage of DMSO should be present in each well. Alternative excitation/emission wavelengths. In the case of overlapping fluorescence spectra between that of the reporter ethidium bromide and the compound of interest, the intercalator reporter may be switched to thiazole orange. Typical settings are excitation at 509 nm and emission at 527 nm with a cut-off of 504 nm. Alternatively, several other excitation/emission pairs have been shown to be useful with ethidium bromide (e.g., excitation at 510 nm, emission at 590 nm).
Anticipated Results Scatchard analysis. The most reliable means of determining K and the stoichiometry of binding has proved to be Scatchard analysis (Scatchard, 1949). The method described in the above protocol, analogous to that introduced by Bruice with Hoechst 33258 (Browne et al., 1993; Satz and Bruice, 2000), is easily extended to analyzing higher-order 2:1 and 3:1 complexes. For 1:1 binding, binding constants are established by Scatchard analysis of the equilibrium portion of the titration curve generating a plot of F/[free agent] versus F, yielding a linear section where the slope is the negative of the association binding constant Ka . A critical element to obtaining accurate association constants from Scatchard analysis is the collection of enough data in the area of the curve to ensure linearity and accuracy of the resultant Scatchard plot. Bruice has emphasized that such reciprocal plotting techniques are restrictive and, in more complex systems (e.g., 2:1 or 3:1 binding), can lead to errors in interpreting the data. In such cases, iterative curve fitting of the experimental points provides reliable results and can be used to dissect multiple equilibria. For the analysis of 1:1 binding as illustrated for netropsin in Figure 8.5.3, this is unnecessary. Both the Scatchard and the curve-fitting analysis provide nearly identical values for K. Given the relative ease of the use of this technique and the reliability with which visual inspection of the plotted data can confirm the assumption of 1:1 binding, the use of Scatchard analysis of the titration binding curves for determining K is recommended.
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Binding constants can be assessed indirectly by measuring the displacement of ethidium bromide or directly by measuring the fluorescent increase of the DNA binding compound itself (i.e., in the absence of a competitive binding agent). For the compounds examined to date (e.g., DAPI and Hoechst 33258), the direct and indirect methods provided comparable results that do not appear to be affected by the sequence-dependent affinity of ethidium bromide, or the stoichiometry of binding displacement (Boger et al., 2000).
Time Considerations The time needed to complete the 96-well plate assay (see Basic Protocol 1) is variable and depends on the number of compounds and compound concentrations tested. Typically, 1 to 2 days are set aside for material and reagent preparation and 1 full day is needed to execute the protocol. It should be noted that the preparation of the hairpin oligodeoxyribonucleotides will require significant additional time, but they may be prepared in sufficient quantities for use in many different studies. The time to complete a single titration run (see Basic Protocol 2) is highly dependent on the number of titrant points desired. The time anticipated to complete a titration protocol may range from 1 to 10 hr.
Literature Cited Blackwell, T.K. and Weintraub, H. 1990. Differences and similarities in DNA-binding preferences of MyoD and E2A protein complexes revealed by binding site selection. Science 250:1104-1110. Blackwell, T.K., Kretzner, L., Blackwood, E.M., Eisenman, R.N., and Weintraub, H. 1990. Sequence-specific DNA binding by the c-Myc protein. Science 250:1149-1151.
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binding agents and development of a rapid, highthroughput screen for determining relative DNA binding affinity or DNA binding sequence selectivity. J. Am. Chem. Soc. 122:6382-6394. Boger, D.L., Fink, B.E., Brunette, S.R., Tse, W.C., and Hedrick, M.P. 2001. A simple, highresolution method for establishing DNA binding affinity and sequence selectivity. J. Am. Chem. Soc. 123:5878-5891. Broude, N.E. 2002. Stem-loop oligonucleotides: A robust tool for molecular biology and biotechnology. Trends Biotechnol. 20:249-256. Browne, M.J. and Thurlbey, P.L. 1996. Genomes, Molecular Biology and Drug Discovery. Academic Press, London. Browne, K.A., He, G.X., and Bruice, T.C. 1993. Microgonotropens and their interactions with DNA. 2. Quantitative evaluation of equilibrium constants for 1:1 and 2:1 binding of dien-microgonotropen-a, -b, and -c as well as distamycin and Hoechst-33258 to d(GGCGCAAATTTGGCGG)/d(CCGCCAAA TTTGCGCC). J. Am. Chem. Soc. 115:70727079. Chaltin, P., Borgions, F., Van Aerschot, A., and Herdewijn, P. 2003. Comparison of library screening techniques used in the development of dsDNA ligands. Bioorg. Med. Chem. Lett. 13:4750. Chiang, S.Y., Azizkhan, J.C., and Beerman, T.A. 1998. A comparison of DNA-binding drugs as inhibitors of E2F1- and Sp1-DNA complexes and associated gene expression. Biochemistry 37:3109-3115. Chodosh, L.A., Carthew, R.W., and Sharp, P.A. 1986. A single polypeptide possesses the binding and transcription activities of the adenovirus major late transcription factor. Mol. Cell Biol. 6:4723-4733. Choo, Y., Sanchez-Garcia, I., and Klug, A. 1994. In vivo repression by a site-specific DNA-binding protein designed against an oncogenic sequence. Nature 372:642-645. Dervan, P.B. 1986. Design of sequence-specific DNA-binding molecules. Science 232:464-471.
Boger, D.L. and Johnson, D.S. 1996. CC-1065 and the duocarmycins: Understanding their biological function through mechanistic studies. Angew. Chem. Int. Ed. Engl. 35:1438-1474.
Galas, D.J. and Schmitz, A. 1978. DNase footprinting: A simple method for the detection of protein-DNA binding specificity. Nucl. Acids Res. 5:3157-3170.
Boger, D.L. and Tse, W.C. 2001. Thiazole orange as the fluorescent intercalator in a high resolution FID assay for determining DNA binding affinity and sequence selectivity of small molecules. Bioorg. Med. Chem. 9:2511-2518.
Gottesfeld, J.M., Neely, L., Trauger, J.W., Baird, E.E., and Dervan, P.B. 1997. Regulation of gene expression by small molecules. Nature 387:202205.
Boger, D.L., Dechantsreiter, M.A., Ishii, T., Fink, B.E., and Hedrick, M.P. 2000a. Assessment of solution-phase positional scanning libraries based on distamycin A for the discovery of new DNA binding agents. Bioorg. Med. Chem. 8:2049-2057. Boger, D.L., Fink, B.E., and Hedrick, M.P. 2000b. Total synthesis of distamycin A and 2640 analogs: A solution-phase combinatorial approach to the discovery of new, bioactive DNA
Guelev, V.M., Harting, M.T., Lokey, R.S., and Iverson, B.L. 2000. Altered sequence specificity identified from a library of DNA-binding small molecules. Chem. Biol. 7:1-8. Ham, Y.W., Tse, W.C., and Boger, D.L. 2003. Highresolution assessment of protein DNA binding affinity and selectivity utilizing a fluorescent intercalator displacement (FID) assay. Biorg. Med. Chem. Lett. 13:3805-3807. Hamy, F., Albrecht, G., Florsheimer, A., and Bailly, C. 2000. An ARE-selective DNA minor groove
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binder from a combinatorial approach. Biochem. Biophys. Res. Commun. 270:393-399. Hardenbol, P., Wang, J.C., and Van Dyke, M.W. 1997. Identification of preferred distamycinDNA binding sites by the combinatorial method REPSA. Bioconjugate Chem. 8:617-620. Hayashibara, K.C. and Verdine, G.L. 1992. Template-directed interference footprinting of cytosine contacts in a protein-DNA complex: Potent interference by 5-aza-2 -deoxycytidine. Biochemistry 31:11265-11273. Knudsen, H. and Nielsen, P.E. 1996. Antisense properties of duplex- and triplex-forming PNAs. Nucl. Acids Res. 24:494-500. Kuwabara, M.D. and Sigman, D.S. 1987. Footprinting DNA-protein complexes in situ following gel retardation assays using 1,10-phenanthrolinecopper ion: Escherichia coli RNA polymeraselac promoter complexes. Biochemistry 26:72347238. Larson, C.J. and Verdine, G.L. 1996. Bioorganic Chemistry: Nucleic Acids (S.M. Hecht, ed.) pp. 324–346. Oxford University Press, Oxford. Lee, L.G., Chen, C.H., and Chiu, L.A. 1986. Thiazole orange: A new dye for reticulocyte analysis. Cytometry 7:508-517. LePecq, J.B. and Paoletti, C. 1967. A fluorescent complex between ethidium bromide and nucleic acids. Physical-chemical characterization. J. Mol. Biol. 27:87-106. Matteucci, M.D. and Wagner, R.W. 1996. In pursuit of antisense. Nature 384:20-22. Neidle, S. 1997. Recent developments in triple-helix regulation of gene expression. Anticancer Drug Des. 12:433-442. Neidle, S. and Thurston, D.E. 1994. New Targets for Cancer Chemotherapy (D.J. Kerr and P. Workman, eds.). CRC Press, Boca Raton, Fla. Nygren, J., Svanvik, N., and Kubista, M. 1998. The interactions between the fluorescent dye thiazole orange and DNA. Biopolymers 46:39-51. Rajski, S.R. and Williams, R.M. 1998. DNA crosslinking agents as antitumor drugs. Chem. Rev. 98:2723-2796. Royer-Pokora, B., Gordon, L.K., and Haseltine, W.A. 1981. Use of exonuclease III to determine the site of stable lesions in defined sequences of DNA: The cyclobutane pyrimidine dimer and cis and trans dichlorodiammine platinum II examples. Nucl. Acids Res. 9:4595-4609. Satz, A.L. and Bruice, T.C. 2000. Synthesis of fluorescent microgonotropens (FMGTs) and their interactions with dsDNA. Bioorg. Med. Chem. 8:1871-1880.
Taylor, J.S., Schultz, P.G., and Dervan, P.B. 1984. DNA affinity cleaving. Sequence specific cleavage of DNA by distamycinEDTA-iron(II) and EDTA-distamycin-iron(II). Tetrahedron 40:457-465. Thiesen, H.J. and Bach, C. 1990. Target detection assay (TDA): A versatile procedure to determine DNA binding sites as demonstrated on SP1 protein. Nucl. Acids Res. 18:3203-3209. Thurston, D.E. 1999. Nucleic acid targeting: Therapeutic strategies for the 21st century. Br. J. Cancer 80(Suppl 1):65–85. Trauger, J.W., Baird, E.E., and Dervan, P.B. 1996. Recognition of DNA by designed ligands at subnanomolar concentrations. Nature 382:559-561. Tse, W.C. and Boger, D.L. 2004. A fluorescent intercalator displacement assay for establishing DNA binding selectivity and affinty. Acc. Chem. Res. 37:61-69. Tse, W.C., Ishii, T., and Boger, D.L. 2003. Comprehensive high-resolution analysis of hairpin polyamides utilizing a fluorescent intercalator displacement (FID) assay. Biorg. Med. Chem. 11:4479-4486. Tullius, T.D., Dombroski, B.A., Churchill, M.E., and Kam, L. 1987. Hydroxyl radical footprinting: A high-resolution method for mapping protein-DNA contacts. Methods Enzymol. 155:537-558. Van Dyke, M.W., Hertzberg, R.P., and Dervan, P.B. 1982. Map of distamycin, netropsin, and actinomycin binding sites on heterogeneous DNA: DNA cleavage-inhibition patterns with methidiumpropyl-EDTA-Fe(II). Proc. Natl. Acad. Sci. U.S.A. 79:5470-5474. Werstuck, G. and Green, M.R. 1998. Controlling gene expression in living cells through small molecule-RNA interactions. Science 282:296298. Woods, C.R., Ishii, T., Wu, B., Bair, K.W., and Boger, D.L. 2002a. Hairpin versus extended DNA binding of a substituted β-alanine linked polyamide. J. Am. Chem. Soc. 124:2148-2152. Woods, C.R., Ishii, T., and Boger, D.L. 2002b. Synthesis and DNA binding properties of iminodiacetic acid-linked polyamides: Characterization of cooperative extended 2:1 side-by-side parallel binding. J. Am. Chem. Soc. 124:10676-10682. Yeung, B.K.S., Tse, W.C., and Boger, D.L. 2003. Determination of binding affinities of triplex forming oligonucleotides using a fluorescent intercalator displacement (FID) assay. Biorg. Med. Chem. Lett. 13:3801-3804.
Scatchard, G. 1949. The attractions of proteins for small molecules and ions. Ann. N.Y. Acad. Sci. 51:660-672.
Contributed by Winston C. Tse Gilead Sciences Foster City, California
Tao, Z.F., Saito, I., and Sugiyama, H. 2000. Highly cooperative DNA dialkylation by the homodimer of imidazole-pyrrole diamide-CPI conjugate with vinyl linker. J. Am. Chem. Soc. 122:1602-1608.
Dale L. Boger The Scripps Research Institute La Jolla, California
Nucleic Acid Binding Molecules
8.5.11 Current Protocols in Nucleic Acid Chemistry
Supplement 20
CHAPTER 9 Combinatorial Methods in Nucleic Acid Chemistry INTRODUCTION
I
n vitro selection methodology entails the generation of randomized populations of nucleic acids (often termed a library) followed by the screening of these nucleic acids for molecules possessing specific properties. For example, it is possible select nucleic acids that bind other molecules (aptamers), catalyze chemical reactions (ribozymes), or possess novel physicochemical properties (e.g., enhanced thermal stability). The nucleic acids identified through in vitro selection have a wide range of utilities, particularly in the areas of biotechnology and medicine. For example, aptamers have been employed as biosensors for the detection of analytes. These nucleic acid sensors have similar specificity profiles to traditional sensors, which are often monoclonal antibodies. However, the nucleic acid sensors offer a unique advantage over antibody sensors: in vitro selection experiments can be conducted in days-to-weeks rather than the 3 to 8 months needed to generate a monoclonal antibody.
This chapter is designed to provide the reader with a comprehensive understanding of nucleic acid in vitro selection experiments. The units presented in the core volume provide the reader with state-of-the-art protocols for DNA/RNA in vitro selection experiments to generate either aptamers or molecules with functional properties. UNIT 9.1 sets the stage for this chapter by providing a comprehensive theoretical treatment of in vitro selection. Building on the first unit, UNIT 9.2 provides the reader with guidance on how to design the initial random nucleic acid libraries so that adequate sampling of both sequence and conformational space is insured. Library design is a critical parameter in all in vitro selection experiments. The material presented in UNIT 9.2 is applicable to all subsequent units in this chapter. UNITS 9.3 & 9.4 described the actual process of in vitro selection, in this case to generate aptamers and ribozymes, respectively. UNIT 9.5 presents specialized procedures for in vitro selection of RNA aptamers to a small molecule target. UNIT 9.6 provides specialized procedures for using an RNA pool that incorporates modified nucleotides. In addition to procedures for generating the modified RNA pool and performing the selection, this unit presents control experiments that are necessary to verify that the modifications will not adversely affect the procedure. Gary D. Glick
Combinatorial Methods in Nucleic Acid Chemistry Contributed by Gary D. Glick Current Protocols in Nucleic Acid Chemistry (2002) 9.0.1 Copyright © 2002 by John Wiley & Sons, Inc.
9.0.1 Supplement 8
Theoretical Principles of In Vitro Selection Using Combinatorial Nucleic Acid Libraries Over the past decade, a new paradigm for drug discovery (Gold, 1995) and biological research (Gold et al., 1995) has been developed from technologies that integrate combinatorial chemistry with rounds of selection and amplification, a technique that is called in vitro selection. Systematic Evolution of Ligands by EXponential enrichment, or SELEX (Ellington and Szostak, 1990; Tuerk and Gold, 1990) is a flexible and extremely successful form of this technology that uses combinatorial libraries of oligonucleotides containing regions of randomized sequence as potential ligands. Oligonucleotide libraries (containing randomized regions) provide, after selection, compounds that bind tightly to the intended target. The process of in vitro selection was called SELEX by Tuerk and Gold (1990), while the selected compounds were called aptamers by Ellington and Szostak (1990). SELEX and in vitro selection (from oligonucleotide libraries) are identical. The selected and amplified bonding site (SAAB) technology (Blackwell and Weintraub, 1990) is a specialized form of SELEX directed toward finding naturally occurring sequences that bind proteins in vivo; however, the number of unique sequences used for SAAB analysis is usually much smaller than that used in most SELEX experiments, since the size of the binding area is usually well defined and thus the number of mutagenized nucleotides is small. SELEX and other adaptive molecular evolution techniques, such as phage display (Cwirla et al., 1990; Scott and Smith, 1990; Kay, 1994; Winter et al., 1994), gain much of their power from their ability to isolate individual molecules from vast molecular pools without resorting to cumbersome deconvolution or tagging methods commonly used in combinatorial chemistry schemes. Rather, these methods utilize iterative rounds consisting of ligand selection from combinatorial libraries followed by amplification of these selected ligands to form new libraries enriched for the particular function of interest, e.g., affinity binding or catalytic function. Such techniques enable quite rapid searches of enormous libraries (typically greater than 1015 potential ligands in the case of SELEX). SELEX has been used to discover high-affinity ligands to a wide variety of different molecular targets, including nucleic acid binding proteins, non–nucleic acid
binding proteins, peptides, and small organic molecules (reviewed in Klug and Famulok, 1994; Gold, 1995; Gold et al., 1995). This unit presents a theoretical overview of in vitro affinity selection using SELEX technology. A schematic representation of the SELEX process is shown in Figure 9.1.1 and may be used to describe SELEX performed with libraries of RNA, RNA derivatives, or DNA. For the purposes of developing a mathematical model, the SELEX process for affinity binding may be summarized in four steps: (1) generation of a library of potential ligands, (2) binding of the library to the target molecule, (3) partitioning of the bound ligands from the unbound ligands, and (4) amplification of the partitioned ligands to generate a new, enriched library, leading again to step 1. Repeated application of steps 2 to 4 results in an enriched pool composed of the sequences of interest. For selection of single-stranded DNA (UNIT 9.2), the two strands of the PCR-amplified pool of dsDNA must be denatured, and one of the strands isolated before binding with the target. For selection of RNA and RNA derivatives (UNIT 9.3), the PCRamplified pool of dsDNA must be transcribed to form a pool of RNA before binding with the target. The partitioned RNA must then be reverse-transcribed into DNA before PCR amplification. For the present analysis, these enzymatic transformations—reverse transcription (RT), PCR, and transcription—are all assumed to be perfect, meaning that they do not affect the relative concentrations of the ligands. However, see Mathieu-Daudé et al., (1996) regarding imperfect amplification due to concentration differences, and Sun et al. (1996) for mathematical modeling of the amplification process taking stochastic effects into account. SELEX is a very forgiving technology. High-affinity ligands to nearly any desired target may be found even when the selection conditions (protein and RNA concentrations, for example) are far from optimal. However, great savings in time and material, or perhaps even success with difficult targets, may be achieved by working at the optimal conditions. Determining what these conditions are demands a deeper understanding of the mechanisms of SELEX. We present such a theoretical model here. We first describe the characteristics of a ligand library, comprised of oligonu-
Contributed by Barry Vant-Hull, Larry Gold, and Dominic A. Zichi Current Protocols in Nucleic Acid Chemistry (2000) 9.1.1-9.1.16 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 9.1
Combinatorial Methods in Nucleic Acid Chemistry
9.1.1
Figure 9.1.1 Schematic representation of the SELEX in vitro selection methodology. The initial random pool is derived from synthesized DNA oligonucleotides that are used directly for DNA SELEX or converted to double-stranded templates for transcription for RNA SELEX. Once the initial pool is created, the steps for a round of affinity binding SELEX are presented in the squares; (1) pool generation, (2) incubation with target, (3) partitioning, and (4) enzymatic amplification.
cleotide sequences, that are relevant to the model. We then describe the equilibrium selection model central to SELEX, incorporating these library characteristics. A demonstration of the model applied to experimental data is then presented. Analytical expressions for the optimal nucleic acid and protein concentrations are derived, these being two parameters easily varied during SELEX experiments. However, the formulas for optimal concentrations unfortunately depend on parameters that cannot easily be determined experimentally. We therefore introduce a new parameter, the signal-to-noise ratio, which allows the determination of nearoptimal conditions based only on parameters that are easily determined experimentally.
NUCLEIC ACID LIBRARIES
Theoretical Principles of In Vitro Selection
In vitro selection is performed with nucleic acid libraries containing vast numbers of unique molecules, typically ∼1015 sequences. Such large libraries are desirable in order to saturate the sequence space of longer randomized regions—a useful goal, as SELEX is often directed toward non–nucleic acid–binding proteins that are unlikely to have sites with high affinity and specificity to short sequence regions. Even with known nucleic acid–binding proteins, longer randomized regions may pro-
vide a larger contact surface, often making it possible to find sequences that bind with higher affinity than the wild-type binding sequences. The only practical limitation on library size is imposed by the volumes of material manipulated experimentally; 1015 random sequences are easily synthesized and readily processed. Each sequence in the library is composed of a random region of variable length sandwiched between two regions of fixed sequence used for primer binding sites during enzymatic processing. The length of the random region varies considerably among selection experiments. For affinity binding, most studies use between 20 and 60 nucleotides (Gold et al., 1995), while researchers performing catalytic selections typically use much larger random regions, the largest comprising >200 nucleotides (Hager et al., 1996; Breaker, 1997). The motivation for the difference in sequence length is that binding interactions with proteins and small molecules may require smaller molecular arrangements than those needed to carry out enzymatic activity. It is commonly believed that typical catalytic oligonucleotides have multiple secondary structural domains that may be required for activity, but this hypothesis still awaits rigorous proof.
9.1.2 Current Protocols in Nucleic Acid Chemistry
A basic tenet of in vitro selection experiments is that the selected function of oligonucleotide molecules is conferred through their three-dimensional structures. These structures, usually supported by stacking interactions between adjacent base pairs, are a consequence of the individual sequences. The identification of conserved primary structural units (residues) and secondary structural units (e.g., helices and loops) from those sequences sharing a selected function allows one to define a motif required for the function. Once a motif is defined, it is easy to compute its frequency of occurrence in the initial pool. For example, SELEX-isolated sequences that bind with high affinity to the E. coli rho factor, displayed in Figure 9.1.2a, define the hairpin motif shown in Figure 9.1.2b. There are 44 combinations of base pairs forming the central stem (N-N′) and 2 bases (C/U) tolerated at position 15 out of 20 contiguous nucleotides defining the motif. Since the motif can start at 11 different positions within the 30-nucleotide random region used in this experiment, the sequence for such a motif occurs in the initial pool with a frequency of (44 × 2 × 11)/420 = 5 × 10–9. As discussed below, such an estimate is always an upper limit on the frequency of actual high-affinity ligands. In addi-
tion, the actual frequency of high-affinity ligands is likely to be different than that calculated from the consensus motif, since this motif is usually defined from a sampling of relatively few sequences and is thus underdetermined. We show below that the frequency of selected motifs within the initial pool plays a central role in the progress of SELEX experiments. For a continuous motif of length m, increasing the random region by n bases results in an n-fold increase for representation of that motif in the random sequence library. However, calculating the frequency of occurrence of a particular motif within the original sequence pool is certainly an upper limit on the number of active molecules with that motif, since this estimate does not take into account the likelihood that a particular sequence will fold into the motif of interest. As the length of the random region increases, the number of possible secondary and three-dimensional structures formed increases, decreasing the likelihood that the motif of interest is thermodynamically accessible in any particular sequence in which it occurs (Sabeti et al., 1997). It is difficult to estimate the loss of activity due to alternative folds, but, clearly, as the length of a sequence
Figure 9.1.2 Consensus sequences and motif obtained from E. coli rho SELEX (Schneider et al., 1993). The sequences (a) are aligned to reveal the consensus motif (b), a hairpin loop. Primer binding sites are denoted by lower case letters in (a) and N-N′ denote any Watson-Crick or G-U base pair in (b).
Combinatorial Methods in Nucleic Acid Chemistry
9.1.3 Current Protocols in Nucleic Acid Chemistry
Theoretical Principles of In Vitro Selection
increases, these losses become more significant. Complete coverage of all sequences for a given-size random region can only be achieved with relatively small random regions. Since the number of unique molecules in the starting pool is practically limited to 1015 molecules and there are 4N sequences for a random region containing N positions, the set of all possible sequences (commonly called the sequence space) for libraries with N > 25 is not fully represented at the outset of the experiment (Ciesiolka et al., 1996). Further, most in vitro selection experiments do not rely on mutation to alter the makeup of sequences after selection; therefore the molecules possessing the desired functional activity must be present in the starting pool. As the size of the random region grows, coverage of the full sequence space diminishes exponentially, compounding the problem of culling rare sequences from a limited initial pool. To help overcome this problem of limited sequence space coverage for libraries with large random regions, optimization of “lead” sequences isolated with in vitro selection is commonly performed with further rounds of selection, starting with a biased pool. Typically, the best sequence resulting from a selection is isolated from the enriched pool. A second pool of molecules is then constructed from this lead sequence by biasing the nucleotide content at each position to contain, for example, 70% original sequence and 10% the remaining three bases. This is a common strategy employed in catalytic RNA selection schemes; a rare sequence isolated from pools with >100 random bases is usually not optimal but is a good starting point for further selection experiments. Such strategies have resulted in increased activities more than several-fold over that of the starting sequence (Hager et al., 1996; Breaker, 1997). Of course, such a tactic never guarantees that the best sequence within the overall sequence space has been isolated. The primary focus of the theoretical development to follow will be on in vitro affinity selection experiments—so-called SELEX. Mutation and recombination events are not deliberately included in most SELEX methodologies, although lack of perfect enzymatic fidelity during transcription, reverse transcription, and PCR amplification certainly introduces a small number of mutation events. The effects of these mutations will be ignored in the present analysis, and are expected to be small in any event. SELEX experiments are usually performed to
isolate those sequences present in the initial pool that have the highest binding affinities to the target of interest. Restricting our discussion to these in vitro selection schemes allows for an enormous reduction in complexity of representation; the vast library of sequences can be formally mapped onto affinity distributions with no loss in generality. Coupling this reduction with an equilibrium model for binding, that is easily achieved experimentally, we can accurately model in vitro selection experiments and, therefore, identify those features of the process most critical to experimental success.
AFFINITY PROBABILITY DISTRIBUTIONS It is now commonly accepted that nucleic acid sequences can fold into complex threedimensional shapes bolstered by their secondary structures. It is the three-dimensional display of functional groups on nucleic acid oligomers that is responsible for their differential binding affinities to wide-ranging target molecules. This is true for small-molecule targets, where the oligonucleotides typically fold to engulf the target, as well as protein targets, where extensive surfaces of the macromolecules are in direct contact. Regardless of the particular target, the three-dimensional structures adopted by the sequences, in both their free and bound states, can in principle be mapped to free energies of binding for the target of interest. This is our basis for mapping linear sequences onto a probability distribution of binding affinities. For an equilibrium model of in vitro selection, all those sequences with the same binding affinities will partition in precisely the same way between target-bound and free in solution. In other words, averages computed over distinct sequences are mathematically identical to averages over the binding affinity distribution constructed from the vast (1015) sequence library. This formally exact reduction in complexity is key to our theoretical development presented below. In the following discussion, we will denote an average over an affinity distribution p(Ka) by <. . .> , i.e., = ∫dKa p(Ka) f(Ka), where f(Ka) is some function of affinity and ∫dKa p(Ka) has been normalized to one. In order to construct an equilibrium model for in vitro selection, it is necessary to explicitly define the affinity distribution of the library for the target of interest. Clearly, the details of such distributions will change depending on the target of interest. We show below that certain features of this distribution are key for assessing the progress of selection experiments. This
9.1.4 Current Protocols in Nucleic Acid Chemistry
probability distribution is most conveniently cast in terms of association constants, p(Ka), but can also be cast in terms of binding free energies, since ∆G = −kBT ln(Ka), where kB is the Boltzmann constant and T is the absolute temperature. An experimental determination of p(Ka) is quite difficult and there exist very little data on which to base estimates. For a model of double-stranded DNA-protein interactions based upon independent base-pair contributions to affinity, the correlation between nucleic acid information content and protein binding affinity leads to a binding free energies that are normally distributed (Berg and von Hippel, 1987; Stormo and Yoshioka, 1991). Consequently, the probability distribution of Ka values for this model are log-normal. Such a profile clusters the majority of sequences around <∆G> (), while those sequences with most favorable ∆G of binding [ln(Ka)] occur relatively rarely. The frequency of the highestaffinity molecules, the so-called “winners,” in the pool depends on the width of the distribution and the difference in affinity between the majority of the sequences and the rare winners. Whether the true affinity distributions are lognormal or skewed in some fashion, we expect the general profiles to be consistent with these features. Although direct experimental determination of p(Ka) is not practical, some useful data can be extracted from binding curves of the sequence pool. A pool binding curve obtained with a constant, small nucleic acid concentration and varying protein concentrations yields some limited data for the distribution of affinities. Specifically, the affinity measured using a standard binding curve analysis for bimolecular association yields a value for Kbulk = e; that is, the measured affinity reflects the ln(Ka) averaged over the distribution. The initial asymptotic behavior of the pool binding curve, corresponding to low total protein concentration, is best described by , the average of Ka over the distribution, and this may be quite different from the Kbulk. Similarly, the behavior of the binding curve where protein is in excess follows , whose inverse may differ considerably from and Kbulk. Figure 9.1.3 illustrates this for two distributions of p(Ka). We show in the next section that progress of SELEX during initial rounds is relatively insensitive to the shape of the distribution p(Ka), but depends on the following three key aspects of p(Ka): (1) the of the pool, (2) the association constant Kw of the highest-affinity sequences in the pool (winners), and (3)
the frequency fw of these winners. We show below that for a round of SELEX and Kw are the critical features of p(Ka) that determine the increase in the population of the winners, whereas the frequency of winners in the initial pool sets the scale for the number of rounds required for completion. In general, Kw is a measure of the amount of winning molecules bound to protein, while is a measure of the amount of total molecules bound. Therefore a ratio of these affinities reflects the possible enrichment during a round of SELEX. In general, we believe that the average affinity can be from two to twenty times higher than Kbulk for affinity distributions p(Ka) of initial pools. For the purpose of testing the mathematical model of SELEX (described below) against completed experiments, we must define the key features of p(Ka). There is currently no good way of experimentally determining directly without knowing the affinity distribution, for which there is little data. We can, however, rely on assumptions to fill this gap and make reasonable choices for parameters describing p(Ka). Kbulk may be easily measured experimentally and used to establish a lower limit for . Since the binding curve of a pool of nucleic acids usually closely approximates that of a single defined ligand (see, e.g., Schneider et al., 1993), we assume that the vast majority of ligands cluster around a “bulk” affinity. The affinities of the winning ligands, Kw, however, are usually several orders of magnitude greater than this bulk affinity. Kw is easily obtained at the completion of SELEX by cloning and sequencing the enriched pool and measuring the affinity of individual clones for the target. As previously described, further sequence analysis of the enriched pool often leads to the identity of a winning motif, which then allows for an estimation of the winner’s frequency in the initial pool. These frequencies are typically on the order of 10−9 to 10−13 (Gold et al., 1995) and provide some information about the affinity distribution. Assuming that binding affinities are distributed log-normally, and having determined Kbulk, Kw, and fw, a value for the width of the distribution, and therefore p(Ka), can be determined. Though there are many possible distributions, for the purposes of this discussion, we will consider the lognormal distribution. The log-normal affinity distribution of ligands for the target protein has the probability density function:
Combinatorial Methods in Nucleic Acid Chemistry
9.1.5 Current Protocols in Nucleic Acid Chemistry
Figure 9.1.3 Affinity distributions and their resulting binding curves. A log-normal distribution of binding affinities is presented in (a) and a Poisson distribution in is presented in (c). Calculated binding curves for these distributions are displayed in (b) and (d) and denoted by open circles. The one-component binding curves corresponding to affinities e, , and computed from the distributions are displayed as solid lines. Note that the binding curve derived from best fits the overall binding curve generated from the distributions, while the low and high protein asymptotes are best fit by and , respectively.
p(lnKa) =
2 2 1 e−(lnKa − ) /(2σ ) √ 2πσ
Equation 9.1.1
Theoretical Principles of In Vitro Selection
where lnKbulk = is related to the bulk affinity of the initial pool, σ is the standard deviation of the log-normal distribution, and p(lnKa) dlnKa is the fraction of ligands with affinities having logarithms equal to lnKa ± (1/2)dlnKa. In Equation 9.1.1), Kbulk = e is the affinity corresponding to the peak (and the midpoint) of the log-normal distribution. Since can be determined experimentally, σ in Equation 9.1.1 is determined to match fw at Kw.
A log-normal distribution for p(Ka) allows an analytical calculation of the difference between and Kbulk, given by the following expression: 2
〈Ka〉 = Kbulke(σ / 2) Equation 9.1.2
For instance, if a random pool of RNA has a measured bulk affinity of 106 M−1, and has a winning motif with an affinity of 109 M−1 in the pool at a frequency of 10−10 (integrating the affinity distribution from ln Kw to infinity), then the average pool affinity will be approximately three times greater than the measured bulk affinity.
9.1.6 Current Protocols in Nucleic Acid Chemistry
AN EQUILIBRIUM MODEL FOR SELEX For most SELEX applications, we have found the equilibrium model of Irvine et al. (1991) to be a good description of the in vitro selection process. This model assumes that there are no interactions among the different nucleic acid molecules in the pool and that no multiprotein aggregates form. Further, we replace the impractical summation over distinct sequences (on the order of 1015) with an integral over the affinity probability distribution p(Ka) for a particular protein, P. It is important to note that once p(Ka) has been defined, there are no adjustable parameters in the model; the remainder of the variables are determined by the experimental conditions or are evaluated explicitly. The total concentration of sequences with an affinity Ka is given by: Lt(Ka) = p(Ka)Lt Equation 9.1.3
where Lt is the total concentration of nucleic acid ligands. Concentrations of the ligand:protein complexes for a particular affinity Ka are given by: [P : L(Ka)] = Ka[P][L(Ka)] Equation 9.1.4
By applying mass conservation: Pt = [P] + ∫[P: L(K a)]dK a
(a)
Lt(Ka) = [L(Ka)] + [P: L(Ka)]
(b)
Equation 9.1.5
along with the equilibrium condition expressed in Equation 9.1.4, it is easily shown that the free protein concentration is given by: [P] =
Pt Pt = KaLt KaLt(Ka) 〉 1+∫ dKa 1 + 〈 1 Ka[P] + (n) 1 + Ka[P] Equation 9.1.6
where Pt is the total concentration of protein, both free and bound in complexes. Note that the integrated term in Equation 9.1.5a denotes the total amount of complex formed for all affinity species obtained by summing over the affinity distribution; a similar interpretation holds for the integrated term in Equation 9.1.6. Application of Equation 9.1.3 allows the integral in Equation 9.1.6 to be replaced with angle brackets (<...>). The only unknown in Equation 9.1.6 is [P] and may be conveniently solved for
iteratively to self-consistency by choosing an initial free protein concentration of zero. Once [P] is determined, concentrations of all other species are easily determined with Equation 9.1.4 and Equation 9.1.5. In the ideal case, all ligands that are complexed with protein would be carried into the next round of SELEX and all noncomplexed ligands would be lost. The optimal strategy for such an ideal partitioning would then be trivial: smaller concentrations of protein would always lead to better selection of the highest-affinity ligands over all other ligands. Unfortunately, no partitioning method is ideal. Only a fraction of the protein:ligand complexes are recovered, and some portion of the noncomplexed ligands is partitioned along with the complexed ones. We call the fraction of correctly partitioning complexes the partitioning efficiency, eff. In the case of using nitrocellulose filters as a partitioning method, for example, the efficiency for different proteins may vary from closely approaching unity to as low as 0.1, where only 10% of the input protein (and complexes) is captured. For a given protein, the partitioning efficiency can be measured. We usually assume that this efficiency is the same for all protein:nucleic acid complexes for a particular protein. We define the background partitioning, bg, as that fraction of noncomplexed ligands which are recovered by the partitioning method. For nitrocellulose filters, the background is easily determined and is found to be typically on the order of 1% to 0.01%. We usually assume that this background value is the same for all ligands (however, see Concluding Remarks regarding ligands to nitrocellulose filters). We will show below that these deviations from ideality make the question of finding optimal conditions for SELEX an interesting one. Application of the above considerations leads directly to an equation for determining the frequency, or probability p(n+1)(Ka), at which ligands L(Ka) with an affinity Ka will exist in the sequence pool at round n+1 following a cycle of SELEX with a pool composition of p(n)(Ka) (Irvine et al., 1991): p(n+1)(Ka) =
eff Ka[P][L(Ka)] + bg[L(Ka)]
∫(n){eff Ka[P][L(Ka)] + bg[L(Ka)]}dKa Equation 9.1.7
where ∫(n) ... dKa indicates a summation over the affinity distribution at round n, and thus represents the total amount of nucleic acid partitioned at round n. Note that Equation 9.1.7 is
Combinatorial Methods in Nucleic Acid Chemistry
9.1.7 Current Protocols in Nucleic Acid Chemistry
Figure 9.1.4 Comparison of enrichment obtained with experiment and theory. Enrichment, as measured by pool affinities determined during a SELEX experiment against E. coli rho factor (open circles) is compared to that calculated with the equilibrium model (filled diamonds). All parameters for this simulation, with no adjustable parameters, were obtained from Schneider et al. (1993). A log-normal initial distribution, parameterized as described in the text, was used in the simulation. The initial winner frequency is the parameter with the highest uncertainty in determination, and thus this parameter was varied as described below. Closed diamonds represent the simulation performed at an initial winner frequency of 10−10. Bars above and below represent simulations using initial winner frequencies of 10−9 and 10−11, respectively. The error bars on the experimental data are 95% confidence intervals found by fitting binding curves to the data of Schneider et al. (1993).
Theoretical Principles of In Vitro Selection
cast in terms of [P], determined from Equation 9.1.6, and [L(Ka)], determined from Equations 9.1.4 to 9.1.6, both evaluated with the pool at round n, and affinity distribution p(n)(Ka). The repeated application of this equation, with chosen concentrations of total protein and nucleic acid for each round, allows the initial affinity distribution to be propagated through multiple rounds of SELEX. The exponential enrichment is a consequence of equilibrium binding, i.e., Ka = e–∆G/kBT, and is reflected in the changes in the affinity distributions from one round p(n)(Ka) to the next p(n+1)(Ka). Equation 9.1.7 assumes that all partitioned sequences will be enzymatically processed with the same efficiency and with complete fidelity. In order to incorporate the effects of mutations during the RT, PCR, and transcription steps, a model for computing affinity as a function of sequence is required. As previously discussed, affinity selection using SELEX is designed to isolate those high-affinity sequences that exist in the initial pools, so treatment of mutations is not included here. To accurately model in vitro evolution, however, where mutations are a key characteristic of the technique, such a mapping is essential (Kauffman and Macready, 1995; Schuster, 1995). In our mathematical description here, however, we are concerned with selection and not evolution.
This completes our description of the mathematical model for SELEX. To assess the utility of this model, we present a comparison of simulated results to SELEX experimental results to find high-affinity oligonucleotides binding to E. coli Rho factor (Fig. 9.1.2), a transcription terminator (Schneider et al., 1993). For the simulated data, a log-normal distribution for p(Ka) is assumed and parameterized with experimental data as described above. The remaining parameters, bg, eff, and the protein and RNA concentrations were taken from the SELEX experimental work. Progress of the enrichment during rounds of SELEX is monitored by pool affinity measured with binding curves. The simulated results compare quite well with the experiment (Fig. 9.1.4); the essential features of in vitro selection are quantitatively captured in the equilibrium model developed by (Irvine et al., 1991) and presented here in terms of affinity distribution functions. We now address the issue of how a particular distribution affects the progress of SELEX experiments. We have constructed three distinct affinity distributions that have identical values for , Kw, and fw, and yet different overall distributions p(Ka). We compare a two-point distribution consisting of the affinities and Kw, a log-normal distribution discussed in detail above (Fig. 9.1.3a), and a Poisson distribution p(lnKa) = α2xe−αx, where x = lnKa−
9.1.8 Current Protocols in Nucleic Acid Chemistry
Figure 9.1.5 Comparison of three different initial affinity distributions versus progress of enrichment. The enrichment is measured here by the frequency of the winning ligands in the pool at each round for a two-point distribution (marked with x’s) a log-normal distribution (marked with open circles) and a Poisson distribution (marked with triangles) of binding affinities.
(Fig. 9.1.3c). The log-normal distribution spans the largest range of Ka values, followed by the Poisson distribution, and finally the two-point distribution. These initial distributions are used to initiate 12 rounds of SELEX performed in the absence of background and with identical conditions throughout. The results are displayed in Figure 9.1.5 for three different values of fw. Sensibly, the in vitro selection is completed most rapidly for the case where the winning frequency is highest in the starting pool. In all cases of fw, the enrichment for the first round, where the , Kw, and fw are well matched, is essentially the same for all three distributions, illustrating the importance that these features of the distribution have for enrichment. As the SELEX progresses, however, the distributions begin to differ in and fw, and some differences in enrichment progress begin to emerge. The two-point distribution moves towards full enrichment (100% Kw) fastest, followed by the Poisson, and finally the log-normal distribution. This is due to increased competition for binding reflected in the width of the distributions. Fewer species competing for limited binding helps drive the two-point distribution to completion, while the normal distribution takes many more rounds, on average, to completely saturate the high-affinity binders. A simulation of SELEX requires that the affinity profile for the initial random pool of nucleic acids be defined. Although little is known about fw and Kw at the outset of an
experiment, there is, fortunately, a great deal of useful theory to help guide the design of in vitro selection experiments. In the next section, we present theoretical guides for SELEX that have been derived from the equilibrium model of SELEX presented here, even without a detailed knowledge of the affinity distribution. This set of analytical, theoretical results is, at least to first appearances, independent of the shape of the affinity profile.
OPTIMAL CONDITIONS FOR IN VITRO SELECTION A variety of useful predictions may be derived from the SELEX theory as presented thus far without having to resort to computer simulations. The more useful of these predictions have to do with the conditions for optimal enrichment of the highest-affinity ligands. These conditions depend on the background, bg, the partitioning efficiency, eff, the affinity of the winning ligands, Kw, and the average pool affinity, . It is important to remember that the pool affinity used below is not necessarily the same as the affinity that is measured experimentally (see Fig. 9.1.3). This is especially true of the earliest rounds of SELEX. There are two dimensionless terms that occur often in SELEX theory. We define these as: ε=
eff bg
Equation 9.1.8
Combinatorial Methods in Nucleic Acid Chemistry
9.1.9 Current Protocols in Nucleic Acid Chemistry
and k=
Kw Kpool
Equation 9.1.9
As defined, both the partitioning effectiveness ε and the affinity ratio k should be much greater than unity in most cases. The pool affinity is defined by the total complexed [P:L(Ka)] and un-complexed [L(Ka)] molecules summed over p(Ka): Kpool = ∫ [P:L(Ka)] dKa/{[P] ∫ [L(Ka)]dKa}.
In other words, Kpool is the affinity for the pool as a whole that would result from a measurement of the total amount of ligand bound for the given protein and ligand concentration and distribution of affinities embodied in p(Ka). It is important to note, however, that Kpool is not a true equilibrium constant since its value changes with total protein and ligand concentration. As the total amount of free protein goes to zero (excess nucleic acid), Kpool approaches . Approximating Kpool by is actually quite good for most SELEX conditions since these conditions usually correspond to excess ligand concentrations. The enrichment is defined as the factor by which the frequency of the “winner” in the affinity distribution changes between rounds of SELEX. Clearly, choosing experimental conditions to optimize enrichment leads to the greatest progress during a round of SELEX. Most of the discussion that follows focuses on guidelines to experimental conditions that maximize enrichment and therefore lead to most efficient in vitro selection schemes. It can be shown (Irvine et al., 1991) that the overall maximum enrichment is given by: 2
εk 1 = √ E opt = √ ε + k √
Equation 9.1.10
This optimum may be achieved at any given total nucleic acid concentration by adjusting the total protein concentration, whereas the converse is not true. There are total protein concentrations whose corresponding optimal nucleic acid concentrations do not achieve the maximal enrichment possible; this behavior is discussed in detail below.
Optimal Concentrations Theoretical Principles of In Vitro Selection
The effect of protein concentration on enrichment at several different background levels is shown in Figure 9.1.6a. At high protein con-
centrations (above ) nearly all ligands are bound to an equal extent, and no selection takes place. At low concentrations, the amount of nucleic acid partitioned in the background overwhelms the complexed nucleic acid molecules, and again there is no selection. Clearly, in the absence of background, the optimal strategy is simply to use the lowest reasonable concentration of protein. However, even in the case of no background, it is important to note that the maximum enrichment is fixed at k, or the ratio of Kw to Kpool. As selection proceeds, this ratio decreases since Kpool approaches Kw and enrichment slows. The effect of protein concentration on enrichment at several different values of k is shown in Figure 9.1.6b. The bold curve in this plot (and in all plots in these two sets) is for conditions identical to the bold curve in Figure 9.1.6a. The above argument for the effect of protein concentration on enrichment holds here as well. What is startling is the similarity between the effects of k and the effects of ε on enrichment. It is clear from Equation 9.1.10 that the effects of k and ε are interchangeable for optimal enrichment. In fact, it can be shown that, in the general expression for enrichment as a function of total protein and nucleic acid concentration, the dimensionless quantities k and ε are mathematically interchangeable as well. At a given total ligand concentration, the total protein concentration that leads to optimal enrichment is closely approximated by: 1 )/√ εk Popt = (Lt + K −pool
Equation 9.1.11
where Popt is the optimal total protein concentration, and Lt is the total concentration of ligand (Irvine et al., 1991). As Kw increases compared to Kpool (increasing k) the optimal protein concentration can be reduced, increasing the so-called stringency of selection. This reduction in protein concentration favors the high-affinity molecules in their competition for binding over lower-affinity species, thereby increasing enrichment. In general, the protein concentration that maximizes enrichment is calculated with respect to either Lt or K−1 pool. For typical nucleic acid concentrations ∼10−6 M and high-affinity pools (>106 M−1), the total ligand concentration Lt dominates the choice of protein concentration, whereas for low-affinity pools (<106 M–1) the affinity of the pool K−1 pool sets the concentration range for total protein. For targets with little measurable binding to nucleic acid pools, high protein concentrations should be
9.1.10 Current Protocols in Nucleic Acid Chemistry
Figure 9.1.6 Enrichment as a function of key parameters for SELEX. The effects of protein concentration on enrichment are displayed for various values of the partitioning effectiveness ε (a) and the affinity ratio k (b) with fixed nucleic acid concentration. Similar plots for enrichment as a function of nucleic acid concentration are displayed in (c) and (d) and as a function of signal-to-noise ratio in (e) and (f). The optimal enrichments for each curve calculated with the approximations discussed in the text are denoted by the open diamonds. The bold solid line in each plot represents identical selection conditions (Pt = 10−8 M, Lt = 10−6 M, Kpool = 106 M−1, Kw = 109 M−1, bg = 0.1%, eff = 1), aside from the independently varied parameter for each plot.
employed; in extreme cases, protein can even be in excess with respect to nucleic acid! The open symbols in Figures 9.1.6a and 9.1.6b represent the optimal enrichment and protein concentrations, calculated from Equation 9.1.10 and Equation 9.1.11, respectively.
Even though Equation 9.1.11 is approximate, Figure 9.1.6 illustrates that it is clearly quite good. It is important to note that at high protein concentrations, the enrichment is reduced to one (i.e., no enrichment); all nucleic acid is bound by the protein whose concentration is
Combinatorial Methods in Nucleic Acid Chemistry
9.1.11 Current Protocols in Nucleic Acid Chemistry
in excess of even the highest dissociation constants (lowest affinity binders) and no enrichment occurs. At low protein concentration, the background overwhelms the correctly partitioned species, and again no enrichmentoccurs. In the absence of background, as noted above, the enrichment plateau at E = k is observed for low protein concentration. The effect of nucleic acid concentration on enrichment at several different values of ε and k are shown in Figure 9.1.6c and d. As in Figures 9.1.6a and b, the effects of k and ε on enrichment are identical. For conditions in which nucleic acid is in excess, enrichment is dominated by the competition of high-affinity binders to the limited protein molecules; the same behavior is observed here for fixed protein concentration as for fixed nucleic acid previously discussed, although excess ligand appears to the right of center in Figures 9.1.6c and d, whereas it is to the left of center in Figures 9.1.6a and b. As seen in the fixed nucleic acid case, for zero background the limiting enrichment approaches k, as nucleic acid concentration far exceeds that of protein. As total nucleic acid concentration decreases and protein is in excess, enrichment is qualitatively different than that observed above for fixed nucleic acid concentration. Here, enrichment is seen to plateau at protein excess, the plateau value depending on the fixed protein concentration. The enrichment plateau is governed by Equation 9.1.12: (1 + εkp)(1 + p) E(Lt → 0) = (1 + εp)(1 + kp) Equation 9.1.12
where p = PtKpool. At concentrations where Equation 9.1.12 is valid, protein is in excess over nucleic acid and there is no competition for binding among the ligands. Selection is driven purely by differences in affinity. It is important to note that, even under these conditions, enrichment occurs; these may be conditions necessary for early rounds of SELEX against targets with low overall pool affinity to nucleic acids and in situations of high background. The optimal total nucleic acid concentration, Lopt, at a given total protein concentration is closely approximated by: εk − K−1 Lopt = Pt√ pool Theoretical Principles of In Vitro Selection
Equation 9.1.13
Equation 9.1.13 has an interesting symmetry with respect to Equation 9.1.11 for optimal
protein concentration. For a given Pt, only one concentration Lt exists which maximizes enrichment as given by Equation 9.1.10. Similarly, that concentration Lt determines the same corresponding concentration Pt in order to maximize enrichment. It is important to note that the expression for optimal enrichment contains no dependence on either Pt or Lt, and yet there exist pairs of Popt and Lopt that achieve this global enrichment maximum. In fact, for any given Lt, there always exists a Pt that allows for global enrichment. The converse, however, is not true. When k or ε are small, Lopt becomes negative for certain Pt values, indicating that there is no local optimal, as seen in Figures 9.1.6c and d, for k or ε = 10; maximal global enrichment can never be achieved for those values of Pt.
The Signal-to-Noise Ratio A quantity that is routinely (and easily) measured at the bench is the signal-to-noise ratio, S, which is defined as the amount of oligonucleotide recovered during partitioning in the presence of protein, divided by the amount recovered in the absence of protein, which is due to background. The effect of signal-to-noise on enrichment at several different values of ε and k is shown in Figure 9.1.6, panels e and f. The signal-to-noise ratio that gives optimal enrichment is closely approximated by: Sopt = 1 +
√ εk
Equation 9.1.14
which has the pleasing characteristic of being independent of either protein or ligand concentration. Indeed, S promises to be the reduced variable of choice for guiding SELEX experiments, as the enrichment E may be expressed as a function of S independent of protein or ligand concentration: E=
εk(S − 1) + (ε − S) Sk(S − 1) + S(ε − S)
Equation 9.1.15
It is obvious from inspection of Equation 9.1.15 that the enrichment is sensibly reduced to 1, either when S = 1 (representing no partitioned complex), or when S = ε (when all nucleic acid is partitioned). Equation 9.1.14 may be derived by setting the derivative of E with respect to S in Equation 9.1.15 equal to zero and solving for S, and assuming that ε, k >> 1. As suggested by Equation 9.1.14 and
9.1.12 Current Protocols in Nucleic Acid Chemistry
Figure 9.1.7 Comparison of optimal enrichment to signal-to-noise guidelines. The enrichments obtained using conditions that give signal-to-noise specified by Equation 9.1.16 are compared to the optimal enrichments as a function of the partitioning effectiveness ε for various values of the affinity ratio k in (a) while those obtained using conditions that fix S at two are displayed in (b). Note that enrichment is usually better than 50% of the optimal for these two approximations.
Equation 9.1.15, and by Figures 9.1.6, panels e and f, the effects of k and ε on enrichment as a function of signal-to-noise are not identical, as contrasted with protein and nucleic acid concentrations discussed above. Decreasing the background by a certain factor effectively shifts the enrichment curve up by this factor for large values of S. Decreasing k (decreasing the difference between Kpool and Kw) causes the enrichment curve to asymptotically approach a straight line (on a log/log plot) with intercepts on both the enrichment and signal-to-noise axes equal to ε. This suggests a simple geometric argument for choosing a near optimal signalto-noise. A signal-to-noise ratio may be picked which is one-tenth of the way from the y axis (signal-to-noise = 1) to the x intercept (signal-
to-noise = ε). To this, 1 is added to avoid the sudden drop-off in enrichment near the y axis. Formally, this is expressed as: ε S ∗ = 1 + 10√ Equation 9.1.16
which has a form somewhat similar to Equation 9.1.14. A key feature of Equation 9.1.16 is that it is independent of k; no information about the winning affinity is required to find the optimal signal-to-noise. This information, however, is implicitly accounted for in the determination of S. A plot of the enrichment achieved using Equation 9.1.16, divided by the maximum enrichment as a function of ε and k, is shown in Figure 9.1.7a. Even in the worst-case scenario, an enrichment that is at least one third
Combinatorial Methods in Nucleic Acid Chemistry
9.1.13 Current Protocols in Nucleic Acid Chemistry
of the maximum possible enrichment is achieved by picking conditions based only on easily measured quantities (signal-to-noise and background). In later rounds of SELEX, as k decreases (i.e., as the affinity of the pool approaches that of the winner), this strategy promises an enrichment at least 80% of the maximum. Yet the signal-to-noise ratio allows an even simpler strategy. Setting the signal-to-noise ratio to 2 guarantees an enrichment that is at least 50% of the maximum value (Fig. 9.1.7b). Together, these plots suggest the following strategy. Initially, conditions are used that yield S close to 2 in the early rounds, and, as movement is seen in the bulk affinity of the pool, S is increased to values determined by Equation 9.1.16. This strategy can be realized by evaluating S for different ratios of Pt and Lt at each round of SELEX and carrying forward those conditions that closely match the desired value of S. The relative insensitivity of enrichment to the signal-to-noise ratio at S >2 is this parameter’s most exciting property. There is no effect of protein or nucleic acid concentration on these plots. Although one or both of these concentrations would have to be varied in order to obtain the desired signal-to-noise, it does not matter how this is done (a higher signal-to-noise could be obtained by lowering nucleic acid concentration or raising protein concentration, or a combination of both), and it is not necessary to know what these concentrations actually are.
Theoretical Principles of In Vitro Selection
The range of signal-to-noise ratios that give near-optimal enrichment is also relatively insensitive to either background or the ratio of the winner affinity to the pool affinity. In nearly all cases, enrichment close to the optimal may be achieved by using signal-to-noise ratios between 2 and 4. Amazingly enough, the signalto-noise ratio provides a way to optimize SELEX even while thrashing around in the dark, so to speak. That is, based on the signal-to-noise ratio, an experimenter may select conditions that lead to enrichment of the highest-affinity ligand that is within a factor of two of the best enrichment achievable, and this is possible without the benefit of any affinity data whatsoever! To illustrate this point, we present a comparison of three simulated SELEX experiments. In each simulation, we begin with the same initial distribution chosen to be a log-normal affinity profile. For each profile, the SELEX experiments are performed as follows. The first SELEX simulation follows a set of conditions that optimize pool enrichment at each round, while the second two simulations utilize the signal-to-noise prescriptions outlined above. For the first signal-to-noise simulation, conditions are selected to yield S of 2, while the second signal-to-noise simulation employs Equation 9.1.16 to select appropriate conditions at each round. Results from these simulations are displayed in Figure 9.1.8. The signal-to-noise procedure performs remarkably well, tracking the optimal-condition SELEX
Figure 9.1.8 Progress of SELEX by different guidelines. The progress for rounds of SELEX performed under optimal conditions (filled diamonds) is compared to two signal-to-noise S guidelines, choosing conditions with fixed S of 2 (open squares) and choosing conditions that yield S given by Equation 9.1.16 (open diamonds). Both S guidelines yield progress remarkably close to optimal.
9.1.14 Current Protocols in Nucleic Acid Chemistry
throughout the simulation. Even though we expect the signal-to-noise optimal to average only 75% maximal enrichment, the results in Figure 9.1.8 demonstrate significantly better performance. This is presumably due to the fact that enrichment that is less than optimal in one round leads to a potentially greater enrichment maximum in the next round compared to the corresponding round of fully optimized SELEX. Even though the near-optimal falls behind, it is able to keep up with the optimal conditions. We are currently working to further validate these simulation results with experimental verification for a number of protein targets. As noted above, it is easy to experimentally evaluate the signal-to-noise for a particular set of conditions for a round of SELEX. Since we have shown that signal-to-noise can be used to adjust conditions to achieve near-maximal global enrichment, guiding automated SELEX is an obvious application of signal-to-noise theory; indeed, this was the motivation for exploring enrichment as a function of signalto-noise. An automated system might easily be programmed to evaluate the signal-to-noise ratio and adjust protein or nucleic acid concentrations (or both) appropriately to achieve nearoptimal enrichment. As all the other manipulations in SELEX follow standard protocols, the addition of a protocol for choosing selection conditions should allow for efficient automation of the entire SELEX process with performance close to the theoretical optimal.
CONCLUDING REMARKS We have presented a powerful mathematical model for equilibrium in vitro selection experiments that allows for the identification of conditions leading to optimal enrichment for each round of the iterative SELEX procedure. We have cast the astronomical problem of summing over distinct sequences into a tractable integration over affinity distributions. Although a detailed knowledge of such distributions is currently unavailable, we have demonstrated the utility of our approach by both comparing simulations to experiment and by deriving an optimization guide that is independent of the details of the affinity distributions. We have demonstrated that optimization of SELEX conditions may be easily achieved by monitoring experimental signal-to-noise, a readily measured quantity. This approach promises to be quite powerful for choosing near-optimal conditions for equilibrium in vitro selection experiments.
Recent focus on the automation of SELEX has led to the development of microtiter plate formats for affinity partitioning. In addition to being a convenient format for automation, plate-SELEX offers the possibility of adding a kinetic-selection step to the process, less encumbered by rebinding events as compared to column formats. Ligands could therefore be subjected to selection pressures for kinetic characteristics, such as long off-rates, in addition to the usual high-affinity selection pressures. This is easily achieved through washing after high-affinity ligands have been captured by immobilized targets on the plates. The effects of such kinetic pressures can easily be included in the mathematical model for SELEX; dramatic enrichment for such kinetic characteristics is theoretically possible (Levitan, 1998). There is some experimental evidence to support this theory and more experiments are currently underway. At this point, it is worth emphasizing that the above theory is predicated on the assumption that the background consists of nonspecific (not affinity-related) partitioning of the input oligonucleotides. Although this is a reasonable assumption, there are at least two other possible sources of background. Background could include outside contamination or artifactual, nonamplifiable signal (such as unbound radioactive label). In these cases, the actual background would be lower than what is measured, and the above strategies would have to be adjusted accordingly. Background could also, however, consist of sequences selected for their binding affinity to targets other than the target of interest. For example, sequences that bind to nitrocellulose, or sequences that bind to contaminants in the target preparation, may be partitioned along with specifically bound sequences. In this case, the behavior of the SELEX experiment is better described by a generalization of the SELEX theory to encompass multiple targets (Vant-Hull et al., 1998). Such a theory suggests that high-affinity ligands will evolve independently to each of the various targets in proportion to the concentrations of the respective targets. The SELEX experiment is still likely to be successful, however, as long as: (1) the concentration of the undesired target is not so great that ligands to the desired target cannot be found, (2) the partitioning of the undesired ligands does not preclude partitioning of the desired ligands (as may be the case when binders to the partitioning matrix are being evolved), or (3) the partitioning efficiency for the desired target is not low com-
Combinatorial Methods in Nucleic Acid Chemistry
9.1.15 Current Protocols in Nucleic Acid Chemistry
pared to the undesired targets. In this final case, ligands to the desired target may only be selected after a great many rounds (however, low partitioning efficiency has little effect in the case of a single target). In any case, the best overall strategy for in vitro selection is to make background as low as possible.
ACKNOWLEDGMENTS We would like to thank our many colleagues at NeXstar Pharmaceuticals and the University of Colorado for the intellectual stimulus leading to this work. In particular, we wish to thank Doug Irvine for the initial development of the theory described here and for useful comments on this work, and Ed Brody for his careful reading of the manuscript. Finally, we acknowledge Matt Wecker-Wintersquash for timely supplies of Girl Scout cookies during the writing of these acknowledgments.
LITERATURE CITED Berg, O.G. and von Hippel, P.H. 1987. Selection of DNA binding sites by regulatory proteins: Statistical-mechanical theory and application to operators and promoters. J. Mol. Biol. 193:723750. Blackwell, T.K. and Weintraub, H. 1990. Differences and similarities in DNA-binding preferences of MyoD and E2A protein complexes revealed by binding site selection. Science 250:1104-1110. Breaker, R.R. 1997. In vitro selection of catalytic polynucleotides. Chem. Rev. 97:371-390. Ciesiolka, J., Illangasekare, M., Majerfeld, I., Nickles, T., Welch, M., Yarus, M., and Zinnen, S. 1996. Affinity selection-amplification from randomized ribooligonucleotide pools. Methods Enzymol. 267:315-335. Cwirla, S.E., Peters, E.A., Barrett, R.W., and Dower, W.J. 1990. Peptides on phage: A vast library of peptides for identifying ligands. Proc. Natl. Acad. Sci. U.S.A. 87:6378-6382. Ellington, A.D. and Szostak, J.W. 1990. In vitro selection of RNA molecules that bind specific ligands. Nature 346:818-822. Gold, L. 1995. Oligonucleotides as research, diagnostic, and therapeutic agents. J. Biol. Chem. 270:13581-13584. Gold, L., Polisky, B., Uhlenbeck, O., and Yarus, M. 1995. Diversity of oligonucleotide functions. Annu. Rev. Biochem. 64:763-797. Hager, A.J., Pollard, J.D., Jr., and Szostak, J.W. 1996. Ribozymes: Aiming at RNA replication and protein synthesis. Chem. Biol. 3:717-725.
Irvine, D., Tuerk, C., and Gold, L. 1991. SELEXION: Systematic evolution of ligands by exponential enrichment with integrated optimization by nonlinear analysis. J. Mol. Biol. 222:739-761. Kauffman, S.A. and Macready, W.G. 1995. Search strategies for applied molecular evolution. J. Theor. Biol. 173:427-440. Kay, B.K. 1994. Biologically displayed random peptides as reagents in mapping protein-protein interactions. Persp. Drug Discovery Design 2:251-268. Klug, S.J. and Famulok, M. 1994. All you wanted to know about SELEX. Mol. Biol. Rep. 20:97-107. Levitan, B. 1998. Stochastic modeling and optimization of phage display. J. Mol. Biol. 277:893916. Mathieu-Daudé, F., Welsh, J., Vogt, T., and McClelland, M. 1996. DNA rehybridization during PCR: the ‘C0t effect’ and its consequences. Nucl. Acids Res. 24:2080-2086. Sabeti, P.C., Unrau, P.J., and Bartel, D.P. 1997. Accessing rare activities from random RNA sequences: The importance of the length of molecules in the starting pool. Chem. Biol. 4:767-774. Schneider, D., Gold, L., and Platt, T. 1993. Selective enrichment of RNA species for tight binding to Escherichia coli rho factor. FASEB J. 7:201-207. Schuster, P. 1995. How to search for RNA structures. Theoretical concepts in evolutionary biotechnology. J. Biotechnol. 41:239-257. Scott, J.K. and Smith, G.P. 1990. Searching for peptide ligands with an epitope library. Science 249:386-390. Stormo, G.D. and Yoshioka, M. 1991. Specificity of the mnt protein determined by binding to randomized operators. Proc. Natl. Acad. Sci. U.S.A. 88:5699-5703. Sun, F., Galas, D., and Waterman, M.S. 1996. A mathematical analysis of in vitro molecular selection-amplification. J. Mol. Biol. 258:650-660. Tuerk, C. and Gold, L. 1990. Systematic evolution of ligands by exponential enrichment: RNA ligands to bacteriophage T4 DNA polymerase. Science 249:505-510. Vant-Hull, B., Payano-Baez, A.R., Davis, R.H., and Gold, L. 1998. The mathematics of SELEX against complex targets. J. Mol. Biol.278:579597. Winter, G., Griffiths, A.D., Hawkins, R.E., and Hoogenboom, H.R. 1994. Making antibodies by phage display technology. Annu. Rev. Immunol. 12:433-455.
Contributed by Barry Vant-Hull, Larry Gold, and Dominic A. Zichi NeXstar Pharmaceuticals Boulder, Colorado
Theoretical Principles of In Vitro Selection
9.1.16 Current Protocols in Nucleic Acid Chemistry
Design, Synthesis, and Amplification of DNA Pools for In Vitro Selection
UNIT 9.2
This unit describes the design, synthesis, and amplification of a random sequence DNA pool. Functional nucleic acid–binding or catalytic species can be selected from these random sequence pools. In designing the DNA pool, careful consideration should be given both to the degree of randomization and the length of the random sequence region (see Strategic Planning). Following pool design, chemical synthesis on a commercial DNA synthesizer will yield a single-stranded DNA pool. The newly synthesized oligonucleotide pool can then be purified (see Basic Protocol 1). Prior to amplification, the initial complexity of the pool should be determined (see Support Protocol 1), the skewing of the pool should be determined (see Support Protocol 2), and amplification reaction conditions should be optimized (Support Protocol 3). If the nascent synthetic oligonucleotide is judged to be suitable for large-scale amplification, it can be enzymatically converted into a double-stranded DNA library (see Basic Protocol 2). Multiple copies of a singlestranded DNA pool can be derived from each double-stranded DNA library, or the library
Figure 9.2.1
Flow chart outlining pool design, synthesis, and large-scale amplification.
Contributed by Jack Pollard, Sabine D. Bell, and Andrew D. Ellington Current Protocols in Nucleic Acid Chemistry (2000) 9.2.1-9.2.23 Copyright © 2000 by John Wiley & Sons, Inc.
Combinatorial Methods in Nucleic Acid Chemistry
9.2.1
can be transcribed to yield a RNA pool or a modified RNA pool (see UNIT 9.3). Figure 9.2.1 outlines the procedure. STRATEGIC PLANNING Designing the Initial DNA Pool The nucleic acid pools used for in vitro selection experiments typically contain a randomized central core flanked by constant sequences that are required for enzymatic manipulations, such as PCR amplification, in vitro transcription, or restriction digestion (see also Fig. 9.2.2). Since a pool is relatively expensive to synthesize, both in terms of time and cost, some effort should be devoted to pool design. There are many subtle parameters to consider that can greatly influence the outcome of a selection experiment, including the degree of randomization, pool length, and pool modularity (see Table 9.2.1 for references to selection experiments that have previously been successfully executed with different types and sizes of pools). Type of selection and degree of randomization Most researchers who carry out in vitro selection experiments wish to either better define or optimize a known binding site (binding-site selection), or to identify a novel binding site (aptamer selection). Each of these tasks in turn requires the synthesis of different types of pools. The sequences and structures that contribute to known binding sites are frequently best defined by selections that start from partially randomized pools. One example of binding-site definition that started from a partially randomized pool was a selection that defined critical residues of the Rev-responsive element (RRE) of HIV-1 Rev (Bartel et al., 1991). This experiment is also described in more detail below. Biased pools can also be used for the optimization of a previously isolated motif. For example, aptamers that could bind to the Rex protein of HTLV-1 were selected from a partially randomized pool based on the wild-type Rex-binding element (XBE) but in the end bound Rex 9-fold better than the XBE (Baskerville et al., 1995). In contrast, completely random sequence pools explore a much wider swath of sequence space and are more useful for the isolation of novel binding species (aptamers) or catalytic species (Breaker, 1997; Jaeger, 1997). There are many examples of the selection of novel binding sites from completely random sequence pools (reviewed in Gold et al., 1995; Osborne and Ellington, 1997). Even when a natural binding site is known in advance, a completely different binding site may be selected from a random sequence pool; for
DNA Pools for In Vitro Selection
Figure 9.2.2 Two examples of pools used in in vitro selection. Primers are shown above and below the sequence of the pool. The T7 promoter is delineated in bold. Restriction sites are underlined, with their enzymes listed.
9.2.2 Current Protocols in Nucleic Acid Chemistry
Table 9.2.1
Selection Experiments with Different Types and Sizes of Pools
Target DNA/RNA Length of random region Reference Bacteriophage T4 DNA RNA 8 Tuerk and Gold (1990) polymerase HIV-1 Rev RNA 66, doped (65% wild type, Bartel et al. (1991) 30% non-wild type, 5% deleted) Ribozyme RNA 120 Bartel and Szostak (1993) HIV-1 Rev RNA 30 Tuerk and MacDougal-Waugh (1993) HIV-1 Rev RNA 4 and 6, segmental; 6-9 Giver et al. (1993) and 6-9, segmental RNA 120 Conrad et al. (1994) PKCβ HTLV-1 Rex RNA 43, doped (70% wild type, Baskerville et al. (1995) 30% non-wild type)
example, Tuerk and MacDougal-Waugh (1993) isolated unique binders to Rev that bound better than the wild-type RBE sequence in vitro. Completely random sequence pools can also be used to extract aptamers that bind to proteins not normally thought to bind to nucleic acids; an example of this is the selection of an RNA aptamer that bound and inhibited the β isoform of protein kinase C (Conrad et al., 1994). Completely random sequence pools can also be used for the selection of novel nucleic acid catalysts. For example, starting from a pool with a 220-position random region, Bartel and Szostak (1993) isolated a novel ribozyme capable of RNA ligation. Generally, selections for catalysis require pools with a random region greater than 90 residues, while binding selections use pools with a random region of less than 70 residues. Intermediate between partially random and completely random sequence pools are segmentally random sequence pools. In a segmentally random pool, short tracts of sequence are completely randomized. Segmental randomization thus allows all possible sequences within a short region or set of residues to be examined. Thus, if a natural binding site is known, but a portion of that binding site is suspected to be particularly important for function, then a segmentally random pool can be used to identify all possible, functional sequences within the wild-type sequence context. For example, Tuerk and Gold (1990) selected aptamers that bound T4 DNA polymerase from a pool that contained 8 random sequence positions flanked by wild-type residues. Similarly, many binding sites are known to be presented within a particular structural context, such as a stem-loop or stem-bulge structure. In these cases, a portion of the structure can be completely randomized, and all possible functional stem-loops or stem-bulges can be identified. For example, the Rev-binding element was known to form a stem-internal loop-stem structure. Giver et al. (1993) segmentally randomized only the internal loop portion of the structure and selected Rev-binding species. Many of the anti-Rev aptamers had sequences that were significantly different than the wild-type, yet were still presented in the context of a stem-internal loop-stem structure. Partially random (doped) pool design (binding site selection) The most important issue in the synthesis of a doped pool is the level of randomization (the probability of sequence substitution/position). As a general rule, the substitution frequency of a doped pool should roughly correspond to the number of positions thought to be required for function. For example, if 10 residues within a nucleic acid binding site
Combinatorial Methods in Nucleic Acid Chemistry
9.2.3 Current Protocols in Nucleic Acid Chemistry
are thought to be functional, then the rate of substitution might be set to yield single mutants at least half the time. If the substitution frequency is set too low, there may be too few varying residues or combinations of residues to yield information about functional sequences or structures. In contrast, if the substitution frequency is set too high, the sequence space nearest the wild-type motif will only be sparsely sampled, and many of the highly mutated molecules may be nonfunctional because their sequences will have diverged too far from the wild-type. For example, an in vitro genetic analysis has been used to uncover the critical structural interactions between the HIV-1 Rev protein and its primary RNA binding site, the Rev-binding element (Bartel et al., 1991). The RBE had previously been mapped by deletion analysis to a short segment of HIV-1. Bartel and his co-workers assumed that the minimal RBE was smaller even than the region identified by deletion analysis, and thus decided to heavily dope a portion of a 66-nucleotide sequence at a frequency of 35% substitution/position. The initial RRE library contained ∼1013 molecules that had an average of 23 substitutions/template (0.35 probability substitution/position × 66 positions = ∼23 substitutions); less than 1 in 1012 molecules were completely wild-type. Following selection, a 20-nucleotide core-binding site within the 66-nucleotide pool was readily defined by sequence conservations and covarying residues. A lower substitution rate might not have precisely defined the relatively small binding site, while an even higher substitution rate might have created a mutational load that would have limited the selection of functional molecules or even have allowed the selection of novel, non-wildtype anti-Rev aptamers (Giver et al., 1993; Tuerk and MacDougal-Waugh, 1993). Conversely, if the binding site were larger than originally hypothesized, the relatively high rate of substitution might have meant that few functional molecules could have survived the selection unscathed. The number and type of sequence substitutions, as opposed to the probable target size for mutation, can also be used to plan the synthesis of a doped sequence pool, as described by the following equations. Typically, a 1-µmol synthesis of a 100-residue template yields a pool of ∼1015 amplifiable molecules. Regardless of the degree of partial randomization or the precise doping strategy employed, the number of different mutational combinations is given by: 3n{L!/[n!(L − n)!]} where n is the number of sequence substitutions/template in a template of length L. For example, in the case of the 66-nucleotide RRE pool discussed earlier, there were ∼2.17 × 109 possible 5-residue substitutions and ∼1.25 × 1016 possible 10-residue substitutions. To calculate what fraction of a given set of substitutions are actually contained in a doped pool, the binomial probability distribution can be used: P(n,L,f) = {L!/[n!(L − n)!]}(fn )(1 − f)(L−n) where P is the fraction of the template population when f is the probability of substitution/position. If primarily single-base substitutions are desired, then f should be maximized for n = 1; if multiple mutations (e.g., double or triple substitutions) are desired, then f should be correspondingly higher. If the doping strategy is optimized for n substitutions, then this number of substitutions will occur most frequently, “n − 1” and “n + 1” substitutions will occur less frequently but in roughly equal numbers, and so forth. Higher levels of sequence substitution skew the mutant frequency distribution, allowing the sampling of some regions of sequence space at the exclusion of others (Fig. 9.2.3). DNA Pools for In Vitro Selection
9.2.4 Current Protocols in Nucleic Acid Chemistry
Figure 9.2.3 or 35%.
Comparison of substitution distributions for a 66-nucleotide pool doped to either 18%
Therefore, in the RRE example already cited, a pool of 1 × 1013 molecules doped at a frequency of 35% would contain few 5-residue substitutions [1 × 1013 × P(5,66,0.35) = ∼1.82 × 106 5-residue substitutions out of ∼2.17 × 109 possible 5-residue substitutions]. In contrast, if the pool were doped at a frequency of 18%, all 5-residue substitutions would almost certainly be included [1 × 1013 × P(5,66,0.18) = ∼9.3 ×1010 5-residue substitutions]. Note that in a pool of only 1 × 1013 total molecules, neither doping scheme would yield all possible 10-residue substitutions. Completely random pool design (aptamer selection) Completely random sequence pools are used to initiate selection experiments when no functional nucleic acid sequence or structural motif is known in advance. There is really only one parameter to consider when designing a completely random pool: the length of the random region. While we will consider this parameter in detail below, we must first dismiss a frequent bogey of selection neophytes, the issue of complexity and representation. Random sequence space is a vast landscape of possibilities of which only a vanishingly small fraction can be sampled by either nature or man. Assuming a 4-monomer repertoire from which pools can be constructed, there are ∼1.6 × 1060 unique individual sequences in a sequence space bounded by a 100-residue template (4100 = ∼1.6 × 1060), a quantity of nucleic acid greater than an Avogadro’s number of Earth masses. While this grotesquely large value is clearly beyond the realm of experimental possibility, modern methods of chemical nucleic acid synthesis do allow the sampling of nearly as much sequence information as may be contained in the Earth’s biosphere. As a back-of-the-envelope calculation, consider that there are on the order of ∼1× 109 species in the biosphere, each with ∼1 × 105 genes. If each of these genes in turn is composed of ∼1 × 103 residues, then there are ∼1 × 1017 residues worth of information in a biosphere. In contrast, a typical 1-µmol synthesis of a 100-residue random sequence pool would contain 1 × 1015
Combinatorial Methods in Nucleic Acid Chemistry
9.2.5 Current Protocols in Nucleic Acid Chemistry
molecules × ∼1 × 102 residues/molecule = ∼1 × 1017 unique residues or roughly 1 biosphere’s worth of information. Obviously, the connection and ordering of sequence information in organisms is important as well. Typically, a random sequence pool contains ∼1 × 1015 molecules, and thus can potentially sample on the order of all possible 25-mers (415 = ∼1.1 × 1015). In fact, since different 25-mers can be found in different “reading frames,” a slightly larger sequence space will likely be sampled. Because of this physical restriction, it is sometimes thought that random sequence pools should be no more than 25 residues in length—any longer, and only a fractional sampling would be possible, and many potential sequences would be lost. While this is true, it should be realized that longer pools do not lose any of the numerical complexity of smaller pools (except in those instances where long syntheses are extremely inefficient) and in fact gain access to some fraction of longer sequence and structural motifs as well. For example, tRNA molecules are roughly 76 nucleotides in length. It might prove more difficult to select tRNA mimics from a random sequence population containing 30 randomized residues than from a pool spanning 80 randomized residues. However, any short functional tRNA mimics present in the shorter population should also be present in equal or greater number in the longer population. In most instances, the relative completeness of the pool is not a consideration in the success of a selection. Indeed, it has been shown that functional nucleic acids are not extremely rare (for recent reviews see Gold et al., 1995; Fitzwater and Polisky, 1996) and can be isolated both from “complete” pools that span 20 random sequence positions and from very “incomplete” pools that span 90 random sequence positions. Having dismissed considerations of complexity and representation, the one guiding principle that emerges from this analysis is that longer pools are more generally useful for selection experiments than shorter pools. However, this principle must be applied with appropriate caveats. First, aptamers derived from shorter pools are easier to analyze. Sequence and structural motifs embedded within a 30-nucleotide random sequence region are much more readily apparent than sequence and structural motifs embedded within a 90-nucleotide random sequence region, especially if the motifs are not colinear. Second, longer pools are more difficult to synthesize than shorter pools. Finally, longer pools are more likely to yield amplification or other selection artifacts than shorter pools. For example, pools that contain random regions greater than 90 nucleotides in length can form self-aggregates that precipitate from solution upon prolonged incubation, and thus require immobilization on a solid support prior to selection (Bartel and Szostak, 1993; Lorsch and Szostak, 1994). Because of these considerations, pools used for the in vitro selection of aptamers typically contain from 20 to 80 random sequence positions.
DNA Pools for In Vitro Selection
Longer pools are not only desirable but are likely required in selections for complex functions, such as catalysis. Pools used for the selection of ribozymes typically contain from 50 to 220 random sequence positions (for recent reviews see Gold et al., 1995; Fitzwater and Polisky, 1996). The optimal length of the random region is an active area of research (Sabeti et al., 1997) where many of the fundamental parameters remain to be defined. Practically, though, longer pools must be synthesized as oligonucleotides of 150 residues or fewer in length because of the constraints of DNA synthetic chemistry. For this reason, pools longer than 150 bases are typically generated in a modular fashion by ligating together individual, synthetic oligonucleotides (Bartel and Szostak, 1993). Segments of shorter DNAs can be stitched together by the inclusion of unique restriction sites (Bartel and Szostak, 1993). Asymmetric restriction sites, such as AvaI (C|YCGRG), BanI (G|GYRCC), and StyI (C|CWWGG), are very useful for this task since they minimize intra-pool dimerization via self-ligation. Also, these enzymes are cost-effective for digesting large amounts of DNA. Alternatively, an overlapping region can be included at
9.2.6 Current Protocols in Nucleic Acid Chemistry
the 3′ end of each synthetic oligonucleotide and mutually primed synthesis (e.g., CPMB of a longer template can be carried out. After assembling pool modules, the complexity (yield) of the new, aggregate pool will need to be freshly assessed. The upper bound of the complexity of an assembled pool (e.g., 1011 100-mer modules × 1011 100-mer modules) will likely be much larger than its actual complexity (e.g., 100 micrograms of ligated 200-mer, 9.12 × 1014 molecules).
UNIT 8.2)
Segmentally random pool design (binding site and aptamer selection) In general, the rules governing the design of segmentally random pools are idiosyncratic, depending on experimental purpose. If the desire is to better define a known binding site, then relatively short sequence tracts (i.e., from four to ten residues) should be completely randomized. The randomization of longer sequence tracts may lead to the selection of novel binding sites rather than variants of a known binding site. The residues can either be colinear (as is the case for many DNA binding sites) or dispersed (as is the case for many RNA binding sites). If the desire is to identify a binding site within the context of a known structural element, then from four to twenty residues can be completely randomized. In this instance, the fewer the number of residues that are randomized, the more likely it will be that the selected sequences will resemble a wild-type binding site or retain an engineered structure. The greater the number of residues that are randomized, the more likely it will be that a novel aptamer sequence or structure will be discovered. Primer design Generally, the constant sequences at the 5′ and 3′ ends of a pool function as primer-binding sites and can be almost any sequence or length. Primers of 20 nucleotides in length are convenient because their melting temperatures are convenient for the PCR and they can easily be synthesized in high yields. In designing constant sequences and complementary primers, obvious artifacts associated with the PCR, such as secondary-structure formation or self-association that could lead to the production of primer dimers, should be avoided. Computer programs such as the Genetic Computer Group’s PRIME or the Whitehead’s PRIMER3 assist in designing constant regions. Other primer design programs include Amplify (Bill Engels, Dept. of Genetics, University of Wisconsin, Madison) and Oligo (National Biosciences). As a rule of thumb, one should try to avoid using the same triplet sequence more than once in either constant region. Beyond these basal considerations, there are two schools of thought regarding the sequence of the priming site itself. On the one hand, designing primers to possess a 3′ clamp of 5′-WSS-3′ (IUB codes: W = A or T, S = C or G), such as ACC, ensures good extension by polymerases. On the other hand, the inclusion of A/T-rich regions at the 3′ termini of primers reduces the frequency of mispriming and allows virtually "infinite" multiplication of DNA amplicons (Crameri and Stemmer, 1993). The inclusion of restriction sites within primer regions can facilitate cloning of selected nucleic acids, although palindromes adjacent to the 3′ ends can also facilitate the genesis of primer-dimers. Finally, primers for partially randomized pools should be designed so that they do not conflict with the folding or accessibility of a known DNA or RNA binding site. It is suggested that the secondary structure of the wild-type binding site with any appended primer-binding sites be determined using an algorithm such as Mulfold (Jaeger et al., 1989). If the native or wild-type structure of the binding site is not among the most common folds, then the primers should be redesigned. If an RNA pool is to be constructed, runoff RNA transcripts for in vitro selection are frequently made with T7 RNA polymerase. There are several known promoters for T7
Combinatorial Methods in Nucleic Acid Chemistry
9.2.7 Current Protocols in Nucleic Acid Chemistry
RNA polymerase (Milligan et al., 1987), but the following minimal sequence gives good yields: −17 −1 5′-TAA-TAC-GAC-TCA-CTA-TA-3′ Addition of a G and C residue at the −18 and −19 positions of the minimal promoter helps to close the DNA duplex and stabilize the 5′ end of the promoter region, thereby increasing transcriptional yields. Transcription initiation is optimal when there are stretches of purines in the +1 and +2 positions, with GG being the best initiator (Milligan et al., 1987). Transcriptional yields also increase if uridine does not appear in the transcript before position 6. Typical pool designs incorporating all the elements described are shown in Figure 9.2.2. Chemically Synthesizing the Pool While pools of genomic DNA sequences have been used for selection (Singer et al., 1997), partially or completely random sequence pools must be chemically synthesized. Modern DNA synthesizers utilize phosphoramidite chemistry (UNIT 3.3) or H-phosphonate chemistry (UNIT 3.4) and can routinely produce usable amounts of DNA up to 150 nucleotides in length. Longer oligonucleotides can also be synthesized, but side reactions such as branching and depurination accumulate throughout the synthesis and the amount of final, usable product recovered can be vanishingly small. Since stepwise coupling efficiencies for a long oligonucleotide are on average ≥98%, the typical yield of a 100-base synthesis that starts with a 1-µmol column is 13.5%, or 13.5 nmol, or 1 × 1016 different molecules, of which ∼10% to 30% can be enzymatically elongated or amplified. Several strategies can be used to enhance the synthetic yield of oligonucleotides that are longer than 100 bases (see APPENDIX 3C). Further, if a pool longer than ∼150 nucleotides is desired, smaller pools can be modularly synthesized and coupled by ligation or mutually-primed synthesis (see discussion of completely random pool design, above). During synthesis it is wise to prevent the cross-contamination of primers with their corresponding pool. It has recently been discovered (A. Friedman, pers. comm.) that when pools and primers are synthesized on identical ports of a DNA synthesizer, there is some mixing of the molecules. The contamination is sufficient to yield a positive signal following extensive (30 to 50 PCR cycles) amplification of a no-template negative control. The unprogrammed interleaving of pools and primers can lead to extreme skewing of amplified materials, such that only a few species from the original pool may comprise a significant fraction of a subsequent amplification reaction. Therefore, pools and their cognate primers should be synthesized on different synthesizer ports and/or the machine should be extensively flushed with acetonitrile between syntheses.
DNA Pools for In Vitro Selection
Most synthesizers can be programmed for in-line, degenerate mixing of bases. While this method is useful when only a few positions must be randomized, because of the extremely fast reaction of the activated phosphoramidite with the newly deprotected 5′ hydroxyl, random sequences will be skewed towards the phosphoramidite that first enters the column. Therefore, for longer pools or pools that should contain a statistically random distribution of nucleotides, it is better to manually mix the phosphoramidites off-line and use this mixture for the synthesis of degenerate sequence positions. A more stochastic distribution can be obtained by including larger amounts of A and C phosphoramidites in the mix to compensate for the faster coupling times of G and T phosphoramidites (Zon et al., 1985). Suggested ratios include a 3:3:2:2.4 molar ratio of A:C:G:T phosphoramidites (D.P Bartel, pers. comm.), and a 1.5:1.25:1.15:1 molar ratio of A:C:G:T (see User’s Manual for PE Biosystems Models 392 and 394 DNA/RNA Synthesis).
9.2.8 Current Protocols in Nucleic Acid Chemistry
Table 9.2.2 Representative Calculations Based on the Masses and Efficiencies for Couplings that Utilize the Canonical Tetrazole Activation Chemisty and Phosphoramidites Bearing Standard Protecting Groups
Phosphoramidite 5′-CE-dA 5′-CE-dC 5′-CE-dG 5′-CE-dT
Molecular mass (g/mol)
Mass correction
858 834 840 745
0.87 0.89 0.89 1.00
Coupling efficiency correction 0.67 0.67 1.00 0.83
Overall correction 0.58 0.60 0.89 0.83
Doped pools are perhaps the most difficult to synthesize (Hermes et al., 1989; Bartel et al., 1991). Doping can be accomplished by using phosphoramidite mixtures that have been adjusted to ensure the proper level of partial randomization of a given nucleotide. For example, if a doped pool is to be synthesized in which non-wild-type residues are included at a rate of 10%/position, then for the adenosine bottle a molar ratio of 33.43:1.50:1.00:1.21 of A:C:G:T phosphoramidites should be used. These ratios were derived by first adjusting for the relative molecular mass and coupling differentials of the individual phosphoramidites and then mixing the phosphoramidite solutions on a percent volume basis to yield the desired extent of doping. This process is described in greater detail below. To normalize the coupling of different phosphoramidites, relative correction factors that take into account different coupling efficiencies and molecular masses must be calculated. Multiplying together these correction factors gives an overall correction factor to provide equal molar coupling of each phosphoramidite. Table 9.2.2 displays representative calculations based on the masses and efficiencies for couplings that utilize the canonical tetrazole activation chemistry (UNIT 3.3) and phosphoramidites bearing standard protecting groups [cyanoethyl for the phosphates along either isobutyryl (N-2 of guanine) or benzoyl (N-6 of adenine and N-4 of cytidine) groups; see UNIT 2.1]. Other chemistries and protections may require the substitution of other correction factors. Most modern synthesizers require that ∼1 g of phosphoramidite be dissolved in ∼20 mL of acetonitrile to be used in the coupling reaction. Applying this constraint along with the combined mass-coupling (overall) correction factor gives the volumes shown in Table 9.2.3 to dissolve 1 g of each phosphoramidite. Therefore, if equal volumes of each of these solutions are mixed, equal molar coupling should occur since the molar concentrations have been adjusted to account for both the mass and coupling differentials. As in the example above, if a doped pool is to be synthesized in which non-wild-type residues are included at a rate of 10%/position, then the amidites should be mixed as in Table 9.2.4.
Table 9.2.3 Volumes of Acetonitrile Needed to Dissolve 1 g of Phosphoramidite
Phosphoramidite 5′-CE-dA 5′-CE-dC 5′-CE-dG 5′-CE-dT
Dissolved in X mL of acetonitrile 11.6 12.0 17.8 16.6
Combinatorial Methods in Nucleic Acid Chemistry
9.2.9 Current Protocols in Nucleic Acid Chemistry
Table 9.2.4 Amidite Mixtures for Synthesis of Doped Pool in Which Non-Wild-Type Residues are Included at a Rate of 10% /Position
Phosphoramidite A C G T A C G T
Total Mutagenesis Volume (%) (mL) 10 10 10 10 10 10 10 10 20 10 20 10 20 10 20 10
Volume each amidite to mix (mL) A
C
G
T
9.00 0.33 0.33 0.33 8.00 0.67 0.67 0.67
0.33 9.00 0.33 0.33 0.67 8.00 0.67 0.67
0.33 0.33 9.00 0.33 0.67 0.67 8.00 0.67
0.33 0.33 0.33 9.00 0.67 0.67 0.67 8.00
In addition to varying nucleotide composition, it is also possible to vary the length of random sequence that is synthesized. Deletions can be stochastically incorporated during a synthesis by replacing the capping step with an acetonitrile wash (Bartel et al., 1991). It is more difficult to stochastically incorporate insertions, but the lengths of segmental random sequences in a pool can be mixed. For example, in Giver et al. (1993), four columns were used to generate a pool with two random regions of 6 to 9 positions separated by a constant domain. The first column was synthesized with 6 random positions, the second with 7 random positions, etc. Following the addition of the intervening constant sequence, the synthesis was stopped, the four columns were opened, and the resins from the four columns were mixed. The mixed resins were then equally redivided into four new columns and the synthesis was resumed. The first column incorporated 6 positions, the second column 7 positions, etc. Thus, the first column contained oligonucleotides in which the first random segment was 6, 7, 8, or 9 residues long and a second random segment that was uniformly 6 residues long. The second column contained oligonucleotides in which the first random segment was 6, 7, 8, or 9 residues long and a second random segment was uniformly 7 residues long, and so forth. Following the completion of all four syntheses, the reactions were combined to generate the final random sequence pool. BASIC PROTOCOL 1
PURIFICATION OF A RANDOM SEQUENCE POOL A newly synthesized oligonucleotide pool should be purified on a denaturing polyacrylamide gel (see e.g., CPMB UNIT 2.12) prior to amplification. Oligonucleotides can also be purified using an HPLC or commercially available spin columns, but HPLC purification is not recommended for ssDNA pools, due to concerns about cross-contamination. Since oligonucleotides of equivalent length but different sequence migrate at slightly varying rates (see User’s Guide for PE Biosystems Expedite Nucleic Acid Synthesis System), a pool should appear as a broader band than a homogeneous sequence. In fact, because of the presence of capped failure sequences and depurinated, cleaved fragments, it is likely that the oligonucleotide product will appear even more heterogeneous. As a general note, since sequences exist as single copies prior to amplification, individual species can be easily lost. Therefore, it is important to wash and elute the various filters, tubes, and tips described below one or more times. The eluates can then be pooled for a final precipitation and eventual amplification. Contamination of primers or other solutions with a synthesized or isolated pool should be avoided by using aerosol barrier tips. Similarly, gel plates used during purification
DNA Pools for In Vitro Selection
9.2.10 Current Protocols in Nucleic Acid Chemistry
should be washed thoroughly to ensure that they are free of contamination with other pools or primers. Materials DNA pool Ammonium hydroxide n-butanol TE buffer, pH 8.0 (APPENDIX 2A) Urea loading buffer, 2× (APPENDIX 2A) 5 M NaCl Ethanol Fluorescent TLC plate (VWR), wrapped in plastic wrap UV lamp Razor blades Small-bore syringes 13-mL centrifuge tubes capable of withstanding temperature extremes (Sarstedt) 90°C water bath Rotary shaker Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (e.g., APPENDIX 3B or CPMB UNIT 2.12) 1. After synthesis, deprotection, and cleavage from the solid support, lyophilize the oligonucleotide solution (in concentrated ammonium hydroxide) to dryness or precipitate with a 10-fold volume of n-butanol. The n-butanol precipitation can occur quite quickly at room temperature for longer oligonucleotides. Shorter (<20 base) oligonucleotides may require longer or colder incubations. To ensure more efficient recoveries of oligonucleotides it is safest to precipitate for ≥1 hr at −70°C.
2. Pour a denaturing polyacrylamide gel (e.g., APPENDIX 3B or CPMB UNIT 2.12). To allow for good separation of near-full-length from non-full-length products, the acrylamide concentration should be chosen so that the full-length oligonucleotide will migrate approximately one-half to three-fourths of the way into the gel by the time the loading dye reaches the bottom.
3. Resuspend the lyophilized or precipitated pellet in ∼100 to 200 µL of water or buffer (e.g., TE buffer, pH 8.0) and add an equal volume of 2× loading dye. Heat denature samples at 75°C for 5 min prior to loading. Load ∼20% of a 1-µmole synthesis per 2 cm × 2 cm × 1.6 mm well and perform electrophoretic separation. 4. Place gel on a fluorescent TLC plate that has been wrapped in plastic wrap and excise the oligonucleotide product from the gel with the aid of a UV lamp, using razor blades. The desired oligonucleotide product is generally the darkest, shadowed band on the gel (excluding UV-absorbing material that runs at the dye front). If stepwise synthetic efficiency has been low, the product will appear as a smear instead of as a clear band. Since many of the N-1, N-2, etc. products can be converted into full-length products by the polymerase chain reaction, a fairly wide band of near full-length products can be cut from the gel. The excision should be carried out relatively quickly, since unnecessarily long UV exposure can damage the oligonucleotide product.
Combinatorial Methods in Nucleic Acid Chemistry
9.2.11 Current Protocols in Nucleic Acid Chemistry
The full-length oligonucleotide product should be the slowest-migrating band. However, if deprotection has been incomplete, lighter bands that migrate considerably above the major fully deprotected band may be observed. Unpolymerized acrylamide absorbs strongly at 211 nm and may cause shadowing at the edges and wells of the gel. This can obscure the resolution or recovery of bands in the outer lanes.
5. Elute the oligonucleotide from the gel slices as follows. a. To aid in the diffusion of the oligonucleotide from the acrylamide matrix, chop gel slabs into fine particles by forcing the gel through a small-bore syringe. b. Place the crushed gel slabs in a 13-ml centrifuge tube capable of withstanding temperature extremes. c. Add 3 mL of TE buffer, pH 8.0, per 0.5 mL of gel slab (typically corresponding to one to two wells), and place the sample at −80°C for 30 min or until it is frozen solid. d. Quickly thaw the tube in a hot water bath and then let it soak at 90°C for 5 min. Elute the DNA overnight at room temperature on a rotary shaker. This freeze-rapid thaw approach (Chen and Ruffner, 1996) allows ice crystals to break apart the acrylamide matrix, increasing yield and decreasing elution time. Typically, 80% of a 20-mer oligonucleotide can be recovered after 3 hr of rotary shaking, making this technique comparable to electroelution (see, e.g., CPMB UNIT 2.7). Because elution is a diffusion-controlled process, higher elution volumes or serial elutions from the same gel slice can increase the amount of DNA recovered. Longer oligonucleotides diffuse from the gel more slowly than shorter sequences. Samples of especially long synthetic DNAs and RNAs that are particularly resistant to elution with aqueous buffers may be eluted more easily in 6 vol of formamide (>5 hr at room temperature), followed by a brief elution with an aqueous buffer (∼1 hr). Isoamyl alcohol extraction (e.g., CPMB UNIT 2.12) can be used to bring the extracts to a convenient volume for subsequent precipitation.
6. Precipitate the eluted oligonucleotide pool by adjusting the salt concentration to 0.3 M using a 5 M NaCl stock solution, then adding 3 vol of ethanol. Keep at –20°C for 3 hr, then microcentrifuge at maximum speed 4°C. Lyophilize to dryness. Resuspend the synthetic pool in TE buffer, pH 8.0 (to protect against nuclease contamination or drastic pH changes). If the volume of the eluted oligonucleotide is too large to conveniently precipitate, concentrate the sample by extracting against an equal volume of n-butanol. Remove the upper butanol layer and repeat until the aqueous volume is convenient for precipitation. About 1/5 of the aqueous layer is extracted into the organic butanol layer for every volume of butanol used. If too much butanol is used, thereby completely extracting the aqueous layer into the butanol, add more water and repeat the concentration. SUPPORT PROTOCOL 1
DNA Pools for In Vitro Selection
DETERMINING THE POOL COMPLEXITY The number of different molecules present in a population can affect the outcome of a selection experiment (see Troubleshooting). If the pool complexity is too low for a given application, the pool will have to be resynthesized. Pool complexity is, in turn, a function of yield and of the number of molecules in the pool that can be fully extended by a polymerase. The overall yield of the synthesis can be calculated by determining the UV absorption of the pool. However, deletions, incompletely deprotected residues, or backbone lesions that arise during chemical synthesis decrease by 10% to 40% the fraction of molecules in a synthetic pool that can be fully extended by polymerases. For example, the rate of insertions (presumably due to DMT
9.2.12 Current Protocols in Nucleic Acid Chemistry
cleavage via tetrazole) has been measured to be as high as 0.4% per position, and the rate of deletions (presumably due to incomplete capping) has been found to be as high as 0.5% per position (A. Keefe and D. Wilson, pers. comm.). The number of usable DNA molecules that are actually present in a nascent pool can be calculated by determining the fraction of the pool that can be extended by Taq polymerase. Materials Purified ssDNA pool and labeled primers 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2A) 10 mM MgCl2 5 mM DTT [γ-32P]ATP (>3000 Ci/mmol) T4 polynucleotide kinase 1:1 phenol/chloroform (APPENDIX 2A) Chloroform 4.0 M ammonium acetate Taq DNA polymerase TE buffer, pH 8.0 (APPENDIX 2A) PCR amplification buffer (APPENDIX 2A) 2× formamide loading buffer (APPENDIX 2A) 15 × 17–cm denaturing polyacrylamide gel (APPENDIX 3B) Thermal cycler Phosphor imager plate and phosphor imager Additional reagents and equipment for quantitation of DNA (e.g., CPMB APPENDIX 3D), end-labeling of DNA (e.g., CPMB UNIT 3.10), phenol/chloroform and chloroform extraction of DNA ( APPENDIX 2A), PCR amplification (e.g., CPMB Chapter 15), and denaturing polyacrylamide gel electrophoresis (APPENDIX 3B) 1. Quantitate DNA by UV absorption assuming that A260 of 1.0 indicates ∼37 µg/ml of single stranded DNA. Also see, e.g., CPMB APPENDIX 3D.
2. Label the 5′ end of the 3′ PCR primer with [γ-32P]ATP by preparing the following reaction mixture. For 30-ml reaction (volume of reaction and concentration of DNA and [g32 P]ATP will vary depending on application): 50 mM Tris⋅Cl, pH 7.5 10 mM MgCl2 5 mM DTT 1 to 50 pmol dephosphorylated DNA, 5′ ends 50 pmol (150 µCi) [γ-32P]ATP 50 µg/ml BSA 20 U T4 polynucleotide kinase Incubate 60 min at 37°C, then stop reaction by adding 1 µl of 0.5 M EDTA. Phenol/chloroform and chloroform extract the labeled oligonucleotide (see recipe for phenol/chloroform/isoamyl alcohol in APPENDIX 2A), and precipitate by adding an equal volume of 4.0 M ammonium acetate and 2 vol ethanol. Microcentrifuge to collect the pellet, remove the supernatant, and redissolve the labeled DNA pellet in 10 µL of TE buffer, pH 8.0.
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This procedure ensures that most of the unincorporated label remains in the supernatant.
3. Incubate ∼50 pmol of labeled primer with a 2- to 5-fold molar excess of pool in a 50-µL extension reaction, under the same conditions that will be used in the final amplification, in a thermal cycler as follows (see, e.g., CPMB UNIT 15.1 for PCR). a. Denature and anneal the primer and template DNA in PCR amplification buffer (usually 94°C for the denaturation step and ∼50°C for the annealing step). b. Add Taq or other DNA polymerase (scaled to the anticipated enzyme concentration to be used in the large-scale amplification), then ramp the temperature to 72°C for 20 min. It may be useful to take time points to determine whether the reaction has gone to completion.
c. Finally, terminate the reaction by the addition of 2× formamide loading buffer. 4. Heat the extension reaction to 90°C for 3 min and load the reaction on a 15 × 17–cm denaturing polyacrylamide gel with appropriate radiolabeled size markers. Electrophorese until the dye is at or near the bottom of the gel, but do not let the radiolabeled primers run off. It is also useful to load a separate well with an aliquot of the primer alone. Choose an acrylamide percentage that allows efficient separation of small primers from larger extended products.
5. Dry and expose the gel to a phosphor imager plate. Using a phosphor imager, quantify the control primer band and the extended product band. There may be a smear leading up to the extended band. One should use one’s best judgement in determining how much near-full-length material will be included in the quantitation. Calculate the percent extension by dividing counts of labeled, extended product by counts of labeled primer. Percent extension for a gel-purified ssDNA pool can range from 10% to 30%. The complexity of the pool is then the yield (determined in step 1) multiplied by the extension efficiency (percent extension determined above). If the complexity of the pool is insufficient for planned experiments, then the pool must be resynthesized. SUPPORT PROTOCOL 2
DETERMINING THE POOL BIAS
SUPPORT PROTOCOL 3
SMALL-SCALE PCR OPTIMIZATION OF POOL AMPLIFICATION
Following extension, the reaction should be repeated using a cold primer and the nonradioactive double-stranded DNA pool should be amplified in a PCR reaction, cloned (e.g., using a TA cloning kit from Invitrogen), and individual members sequenced to determine the degree of partial or completely randomness. The cloning step could also be carried out following PCR optimization (see Support Protocol 3). From 20 to 30 clones should be sequenced to determine the base composition of the starting pool. The random region should be composed of roughly 25% of each base. A pool with the random region skewed toward one or more bases (>30%) should be resynthesized.
To enhance yield and further avoid bias, the amplification conditions for a pool should be optimized prior to carrying out a large-scale amplification. Moreover, since amplifying a pool is costly in terms of both time and money, any optimization of the PCR should first take place on a small scale. The more involved large-scale amplification can then be carried out with confidence.
DNA Pools for In Vitro Selection
9.2.14 Current Protocols in Nucleic Acid Chemistry
Materials dNTPs (APPENDIX 2A) Taq DNA polymerase (e.g., Boehringer Mannheim) PCR amplification buffer containing 1.5 mM Mg2+ (APPENDIX 2A) dsDNA mass markers (e.g., Life Technologies) 4% Nu Sieve agarose gell (FMC Bioproducts) Thermal cycler Densitometer Additional reagents and equipment for agarose gel electrophoresis (e.g., CPMB UNIT 2.5) 1. Carry out a 0.1 mL PCR reaction using 2 nM of synthetic pool oligonucleotide as template, 2 µM primers, and PCR buffer with 1.5 mM magnesium. Use the manufacturer’s suggested quantity of Taq (e.g., 2.5 U of Boehringer Mannheim Taq) in a reaction containing 200 µM dNTPs. A suggested temperature regime is: 10 to 15 cycles: 2 min 1 min 3 min
95°C (denaturation) 55°C (annealing) 72°C (extension).
After 10 to 15 cycles of amplification, check the length and purity of the amplified DNA on a 4% Nu Sieve agarose gel in 1× TBE buffer (e.g., CPMB UNIT 2.5). Annealing temperature may need to be adjusted to as low as 45°C depending on primer composition (e.g., for a small or AU-rich primer). A 0.1 mL reaction typically yields ∼1 ìg, but the amount can vary from 0.1 to 10 ìg. A fuzzy band may indicate that too many cycles of PCR have been carried out. In this case, set up the reaction again and perform fewer cycles.
2. Dilute the double-stranded PCR DNA product 1:128, and repeat the PCR reaction, removing a 5- to 10-µL aliquot during the last 10 sec of the cycle-7 extension step. Serially dilute the amplified product 1:2, 1:4, ... 1:128. Electrophorese all of the samples on a large agarose gel. Note that it is quite difficult to accurately pipet solutions at 72°C. It may therefore be desirable to pipet an amount slightly larger than that intended for use in the serial dilution.
3. Calculate the average PCR efficiency by identifying to what extent the cycle-7 PCR reaction is the result of progressive doublings of the original synthetic DNA. Determine which dilution lanes lack detectable DNA. The largest dilution that lacks detectable DNA is also the dilution that is a minimum estimate of the number of doublings. For example, if the 1/64 dilution is the largest dilution without detectable DNA, this implies that 6 “doublings” of the synthetic DNA yielded at least 64-fold more DNA. This is expressed as follows:
(average efficiency)no. of theoretical doublings (i.e., PCR cycles) = fold increase in DNA Thus, if 7 cycles of PCR were performed, then the average number of doublings per cycle is ∼1.81 [from (∼1.81)7 = 64].
4. Modulate PCR conditions to enhance PCR efficiency. If the pool’s average number of doublings per cycle is <1.8, then the PCR conditions chosen may skew the representation of the pool. In that case PCR conditions should be modulated to enhance PCR efficiency. The following parameters or variables are most amenable to modification. It is best to begin the optimization with a single set of reaction conditions, modify individual parameters relative to this one reference reaction, and then combine all advantageous alterations into a single reaction. In addition, one may wish to consult CPMB UNIT 15.1.
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Supplement 8
Theoretically PCR can proceed until the primers or dNTPs are depleted. Therefore, primer and dNTP concentrations should be well above those used for the amplification of small amounts of DNA. Primer concentrations from 1 ìM to as high as 5 ìM have been used (although concentrations >5 ìM are generally not helpful). It may be useful to scan both above and below 2.5 ìM in 0.5-ìM increments. Magnesium concentration affects both primer annealing and the fidelity of Taq (which decreases with increasing magnesium concentration). Starting at the magnesium supplied in the PCR buffer (usually 1.5 mM), scan in 1-mM increments toward 5 mM as a maximal concentration. DNA denaturation at temperatures above 95°C is usually impractical since this greatly reduces Taq’s half-life. While other thermostable polymerases can be more resistant to higher temperatures, they usually have a lower extension efficiency and are more expensive than Taq. Annealing temperatures are dependent upon both primer sequence and length. The primer annealing temperatures should already be known from the primer design p rocess, o r may be calculated via an algorithm that can be found at http://paris.chem.yale.edu/extinct.html. This algorithm takes into account nucleotide composition, stacking energies (according to Turner’s rules), and empirical data. An annealing temperature ∼5°C less than the calculated annealing temperature is a good place to begin optimization. The amplification is more efficient at a lower annealing temperature, but mispriming and secondary structural problems are more pronounced. Higher temperatures improve the specificity, but decrease the overall yield of the reaction. To determine the optimum annealing temperature for a given primer and magnesium concentration, one should scan in both directions around the annealing temperature in 5°C increments. Finally, extension temperatures are modulated by the properties of Taq, which will extend (although inefficiently) at temperatures as low as 65°C. When extending at temperatures above Taq’s optimum temperature (70° to 75°C) somewhat more polymerase may be required; scanning of the enzyme quantity should be done in 2.5-U increments. However, too much Taq may be harmful to structured single-stranded nucleic acids (Lyamichev et al., 1993).
5. Confirm the results of the extension reaction described in Support Protocol 1 by the optimization method as follows. After optimizing pool PCR conditions for >1.8 average number of doublings per cycle, determine the pool complexity by performing another 0.1-ml PCR reaction with 2 nM of the original, synthetic pool oligonucleotide under the now optimized reaction conditions. After 7 or more cycles of PCR, perform agarose gel electrophoresis on serial dilutions of the PCR reaction adjacent to serial dilutions of dsDNA mass markers. Calculate the amount of amplified DNA using either a densitometer or by estimating which dilutions are most similar. Calculate the approximate pool complexity as follows: g of PCR DNA afterN cycles of PCR = g of starting extendable ssDNA g avg no. of doublings per cycle (see step 4) g of starting extendable ssDNA 330 g / mole × (no. of bases in fullM length product) = mol starting extendable ssDNA mol starting extendable ssDNA × (6.02 × 1023) = molecules of star ting extendablessDNA molecules of starting extendable ssDNA = fraction of extendable ssDNA starting molecules fraction of extendable ssDNA × no. of synthetic pool molecules = pool complexity DNA Pools for In Vitro Selection
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PCR efficiency should be optimized to balance the average number of doublings per cycle against the total reaction volume. A pool of 1 × 1015 molecules (∼1.7 × 109 mol) at a starting template concentration of 2 nM will require 0.85 L for amplification. Therefore, it is greatly desirable to amplify the pool at the highest template concentration that still gives a reasonable number of doublings per cycle. The amplification should generate at least 8 copies of pool DNA if the pool complexity is to be archived and preserved (see Basic Protocol 2).
LARGE-SCALE PCR AMPLIFICATION OF POOL DNA Very long and complex pools often require PCR amplification on a multiple-milliliter scale. Large-scale PCR differs from conventional PCR in that it is typically conducted in water baths using 15 mL, 17 × 120–mm, screw-capped (Sarstedt) thermostable tubes to accommodate the larger volumes. Amplification reactions of up to 2.5 L have been carried out in this way. Medium-scale amplifications can sometimes be carried out in thermal cyclers that can accommodate multiple samples (e.g., 96-well PCR plates).
BASIC PROTOCOL 2
Materials Purified ssDNA pool and primers EDTA 1:1 phenol/chloroform (APPENDIX 2A) Chloroform 4 M ammonium acetate Ethanol TE buffer, pH 8.0, containing 50 mM of a salt such as potassium chloride Thermal cycler or three water baths (one must be a circulating water bath) 96-well PCR plate or 13-mL thermostable tubes (Sarstedt) Thermometer Styrofoam racks Spectrophotometer or fluorometer Additional reagents and equipment for PCR amplification ( CPMB UNIT 15.1; see Support Protocol 3 for determination of conditions on a small scale) and phenol/chloroform and chloroform extraction of DNA (APPENDIX 2A) Plan the reaction Since large-scale reactions are quite expensive in terms of nucleotides and enzyme, preparedness and planning for the large-scale amplification cannot be overemphasized. Primers <20 bases in length usually do not need to be gel purified and can instead be purified by precipitation. 1. After identifying the optimal PCR conditions on a small scale (see Support Protocol 3), prepare reagents for the large-scale reaction. Set aside time for the large-scale amplification, which will probably consume an entire day. The size of the large-scale reaction will be determined in part by the amount of DNA pool to be amplified and by the number of copies of the library that are desired. For example, assume that 100 (extendable) mg of a pool are to be amplified 16-fold. Since the typical amount of DNA recovered from a 100-mL PCR reaction is 1 mg, then each 100-mL reaction should have 1 mg/16 = 60 ng of DNA. 100 mg total/60 ng/100 mL = 1667 × 100 mL, or a 167-mL reaction.
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Choose how the amplification will be carried out If the volume of the large-scale amplification reaction is to be ≤100 mL 2a. Use a commercially available thermal cycler repetitively. Set the reaction mixture up in advance, and pipet 100-µL aliquots into individual wells of a 96-well PCR plate. 3a. Carry out several small amplification reactions in advance to ensure that the optimized conditions determined in Support Protocol 3 work with the PCR plate format, and that amplification is uniform across the PCR plate. 4a. Perform thermal cycling on the entire reaction using eleven PCR plates. For larger volumes Reactions will be divided into aliquots in 13-mL thermostable (Sarstedt) tubes and amplified in a series of water baths. Construct floating racks by cutting off the bottom of the tubes’ Styrofoam packing material. Reinforce these racks by wrapping their edges with heavy tape. Place the racks iteratively in three circulating or static water baths held at the denaturation, annealing, and elongation temperatures previously determined (see Support Protocol 3). 2b. Determine how long it will take for the reaction mixture in a tube to come to thermal equilibrium by constructing a temperature probe, placing a thermometer through the top of a Sarstedt tube filled with 10 mL of water. Place the probe in a rack with other, similar tubes. Typical equilibration times range from 2 to 8 min, depending on the temperature differential. Annealing, and extension times of 5, 6, and 7 min are typical. It should be noted that these ramping temperature profiles are very slow relative to a commercial PCR machine and can yield more amplification artifacts.
3b. To ensure that the reaction conditions actually work as planned, fill the rack with tubes of water, a single amplification reaction, and the temperature probe. Denature the sample for 30 min, and then add Taq after the first annealing step. Take aliquots at each cycle to monitor the progress of the reaction. 4b. When reaction conditions have been confirmed, proceed with the remaining amplification reactions. Allow the final extension step to proceed for at least 20 min to ensure that all templates are completely double-stranded. Do not be alarmed if the solution becomes cloudy; the detergent in the buffer causes the turbidity. Amplification efficiencies of 3 to 4 doublings in 5 cycles can typically be achieved using this method.
5. Following the amplification, pool the reactions from the individual wells or tubes. Chelate the magnesium in the buffer by adding 1.1 molar equivalents of ETDA, pH 8.0. The reactions can be left at 4°C overnight.
6. Add an equal volume of 2-butanol and extract to concentrate the reaction to a manageable volume (usually 10- to 20-fold). Mix the layers by vortexing and then separate by centrifuging 5 min at 1200 × g at room temperature, then discard the upper, butanol layer. Repeat as necessary. About one-fifth of the aqueous layer is extracted into the organic butanol layer for each volume of butanol used. DNA Pools for In Vitro Selection
9.2.18 Current Protocols in Nucleic Acid Chemistry
7. After concentrating the DNA, carry out a phenol/chloroform extraction, followed by two successive chloroform extractions (see recipe for phenol/chloroform/isoamyl alcohol in APPENDIX 2A). At this point, it should be possible to easily precipitate the DNA. Be sure to temporarily save all of the organic layers in case of a mishap. Falcon tubes (50 mL) work well for these extractions, as they are conveniently sized and have a small surface area. Alternatively, a Teflon extraction funnel may be useful since nucleic acids will not stick to its surface.
8. Precipitate the DNA by adding an equal volume of 4 M ammonium acetate (final concentration, 2 M) and 2 vol ethanol in 13-mL Sarstedt tubes if possible. If larger tubes are required, prepare a set of Beckman 250-mL high-speed centrifugation jars. Wash the jars with 15 mL of 3% hydrogen peroxide for 30 min and then rinse three times with 100 mL of distilled water to remove any residual DNases that may remain from previous use (typically bacterial cell pelleting).
9. Resuspend the amplified DNA in TE buffer, pH 8.0, containing 50 mM of a salt such as potassium chloride. It is unwise to resuspend a double-stranded DNA pool in water, since the random segments may denature, reassort, and become transcriptionally incompetent. If it is suspected that the pool has become denatured (for example, if a large single-stranded DNA component is seen on a nondenaturing agarose gel), simply repeat one to two cycles of PCR.
10. Quantitate the PCR DNA. This can be done by carrying out gel electrophoresis in parallel with a DNA ladder of known concentration. The concentration can also be determined spectrophotometrically or by monitoring the change in absorbance of an intercalated fluorescent dye, Hoechst 33258 (Sigma) on a fluorometer (e.g., DyNA Quant 200, Amersham Pharmacia Biotech). These latter methods are much more quantitative (although the fluorometer method may not be accurate for sequences <100 nucleotides in length). However, these methods may not distinguish precipitated double-stranded DNA from residual, precipitated nucleotides or single-stranded primers. Determine the overall PCR efficiency and the final number of DNA molecules. The amount of DNA obtained from large-scale amplification is often referred to in terms of the number of copies of the original synthetic pool’s complexity. For example, if the starting pool had a complexity of 1 × 1015 molecules and 8 × 1015 total DNA molecules were recovered, then, on average, 8 copies of the original starting pool were obtained from the amplification. It should be noted that skewing that may arise during amplification, and sampling errors that occur during the use of the amplified pool, may cause this estimation to be grossly inaccurate; nevertheless, it is empirically useful.
11. Following large-scale amplification, store at least 4 copies of the pool at −80°C. Because of the aforementioned sampling errors, archiving at least 4 copies worth of the pool DNA ensures the preservation of most of the pool’s complexity. The amount of preserved pool complexity can be calculated using the following equation: % of the pool complexity in a given sample = 100 × {1−[( x − y)/x]x} where x is total number of pool copies, and y is the number of pool copies archived. Therefore, in the example given above, if 4 of the 8 copies of the pool generated through amplification are archived, then ∼99.6% of the original starting pool’s complexity is preserved. Similarly, at least 4 copies of the pool should be used whenever manipulations such as ligation, transcription, or biotinylation, are carried out, so that the original complexity is also manifest in the manipulated or synthesized copies.
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Supplement 4
COMMENTARY Critical Parameters Synthesis Depending on the size of the pool to be synthesized, the operation of the DNA synthesizer may first need to be optimized. Short pools (<80 total nucleotides in length) can be synthesized using standard protocols (see, e.g., PerSeptive Biosystems, 1997). In order to synthesize longer pools (>80 total nucleotides in length), all reagents should be fresh and special care should be taken to exclude water from the synthesis (see APPENDIX 3C). To ensure equimolar base incorporation in the random region of longer pools, the phosphoramidites must be mixed in a skewed ratio (see Strategic Planning). Coupling efficiency should be monitored throughout the synthesis by following the trityl output (see APPENDIX 3C). Amplification Optimization of PCR conditions according to established protocols is vital to the success of the large-scale amplification. Cycle temperatures and times, as well as the concentrations of polymerase, primers, and dNTPs (see, e.g., CPMB UNIT 15.1) should be addressed prior to the large-scale workup. Most importantly, since extremely large quantities of relatively expensive reagents (e.g., Taq polymerase) may be required, care should be taken to make sure that all reagents and procedures are in readiness. Different priming sequences often require distinct PCR buffers for optimal extension efficiency; the best buffer for a given pool and primer combination can be easily and systematically identified through the use of a PCR optimization kit (e.g., the PCR Optimizer Kit from Invitrogen).
Troubleshooting
DNA Pools for In Vitro Selection
The most common problem with the synthesis of a random sequence pool is the overall synthetic yield. However, researchers should carefully decide how many sequences are really necessary for their selection experiments. In selection experiments from a pool with a relatively limited potential diversity (i.e., a segmentally random pool with only 1 × 1011 possible sequences or less), even a low synthetic yield should be sufficient. However, in vitro selection from a pool with a very high potential diversity (i.e., a completely random pool with 1 × 1015 possible sequences or more) should use at least 1 × 1014 different sequences initially in order
to adequately sample the potential sequence space. Pools that contain fewer than 1 × 1013 possible sequences should not be used. The most likely sources of low yields and coupling efficiencies are old (i.e., water-contaminated) synthesis reagents. Thus, instead of attempting to amplify an incomplete pool, the pool should be resynthesized with fresh reagents; the old and new pools can then be combined, if desired. If fresh synthesis reagents do not significantly raise yields, then more serious problems, such as line or valve blockage, may be the cause, and the instrument service representative should be contacted. The second most common problem is that the base composition of a partially or completely random region is skewed. Unfortunately, skewing cannot be detected until after completion of a large-scale amplification. Fortunately, unless the degree of skewing is extreme, it should not seriously affect the outcome of a selection. Moreover, if the degree of skewing is known in advance of a selection, it can be taken into account when analyzing the results of the selection. For example, Baskerville et al. (1995) selected functional Rexbinding elements from a partially randomized pool. Despite the fact that the initial pool did not contain equimolar representation of nonwild-type bases at partially randomized positions, these authors were able to determine the relative importance of individual residues by comparing the degree of conservation or variance before and after selection. If a researcher decides that extant skewing of base ratios is unacceptable, this can only be fixed by adjustment of the randomized phosphoramidite mixture and resynthesis of the pool. The third most common problem is that the pool fails to efficiently elongate. With the proviso that the efficiency of extension may be as low as 10% of the available pool, it should not be much lower (i.e., 1% of the available pool). If extension or PCR efficiency is dauntingly low, the PCR conditions should be reexamined and optimized as described, including buffer and enzyme concentrations, temperatures, and extension times. Switching to a different thermostable polymerase, or to a combination of polymerases, will sometimes improve primer extension. If all possible PCR optimization conditions have been addressed, poor extension efficiency could reflect a problem with the synthetic DNA. For example, the pool may not have been completely deprotected or
9.2.20 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Figure 9.2.4 Typical extension reaction. The pool used (N30P) is shown below the figure of the gel. Lane 1 is a size standard, lanes 2 and 3 show control reactions, and lanes 4, 5, and 6 follow the extension efficiency after different incubation times.
a primer binding site may have become largely depurinated during the course of a long synthesis. Although incomplete deprotection is rarely a problem, small aliquots of the pool can be further treated with ammonia, and extension and amplification can again be assessed. If additional deprotection instead yields oligonucleotide degradation, then it is likely that apurinic sites have accumulated, and the pool will have to be resynthesized.
Anticipated Results It is apparent from the discussion earlier in this unit that there is no one correct way to design and amplify a random sequence pool. However, by following the protocols described above, results similar to the following should be observed. If the integrity of the nascent, synthetic pool is good, then the primer extension efficiency (described in Support Protocol 1) should be relatively high. Figure 9.2.4 shows a typical extension reaction for a pool synthesized in the authors’ laboratory (N30P, a segmentally random pool). Lane 1 is an RNA size standard; lanes 2 and 3 show a control reaction with “no template” and “no enzyme,” respectively. In these lanes, only the radiolabeled 3′ primer (24 nucleotides in length) is visible. Lanes 4, 5, and 6 show the primer extension reaction at various incubation times. Molecules that were incapable of full extension make up the smear leading to the full-length product. By determining the number of counts in the full-length product relative to the radiolabeled primer, the extension efficiency for the N30P pool was calculated to be ∼8%. Moreover, it appeared as
though the extension reaction had gone to completion within 2 min. Assuming that the nascent pool is intact and can serve as a template for the primer extension reaction, then it should be possible to amplify the pool via the polymerase chain reaction. Figure 9.2.5 shows the results of an amplification "cycle course" for a different pool (N71, with a 71-nucleotide random sequence core). An 8-mL PCR reaction was placed in a 15-mL Falcon tube and cycled through a series of three water baths. The samples in the figure were drawn at 0, 2, 4, 6, 8, and 10 cycles. This initial PCR reaction was only a trial, and for the final, large-scale amplification of the entire pool, a 150-mL PCR reaction was distributed to 18 Falcon tubes and 7 PCR cycles were carried out. Following amplification, a portion of the N71 pool was cloned into a TA cloning vector (Invitrogen) and ten clones were sequenced. The proportions of different nucleotides in the final pool reflected almost perfect equimolar coupling efficiencies: A, 25.22%; C, 25.37%; G, 25.82%, and T, 23.58%.
Time Considerations The amount of time required for the protocols described in this section should not be underestimated. Pool design will require at least one day, depending on the degree of background research. It is strongly recommended that pool design be discussed with one or more colleagues prior to synthesis. The synthesis of oligonucleotides <150 bases in length can be easily accomplished in one day, allowing 1 hr to ensure proper instrument setup. Pool purification and optimization of PCR conditions
Combinatorial Methods in Nucleic Acid Chemistry
9.2.21 Current Protocols in Nucleic Acid Chemistry
Figure 9.2.5 Cycle course. The gel follows amplification of the N71 pool after 0, 2, 4, 6, 8, and 10 cycles. The pool used in the cycle course is depicted below the figure of the gel.
should take 1 to 2 additional weeks. Finally, the actual large-scale amplification and subsequent isolation of the dsDNA pool will require the researcher’s undivided attention for ∼2 days.
Literature Cited Bartel, D.P. and Szostak, J.W. 1993. Isolation of new ribozymes from a large pool of random sequences. Science 261:1411-1418. Bartel, D.P., Zapp, M.L., Green, M.R., and Szostak, J.W. 1991. HIV-1 Rev regulation involves recognition of non-Watson-Crick base pairs in viral RNA. Cell 67:529-536. Baskerville, S., Zapp, M., and Ellington, A.D. 1995. High-resolution mapping of the human T-cell leukemia virus type 1 rex-binding element by in vitro selection. J. Virol. 69:7559-7569. Breaker, R.R. 1997. In vitro selection of catalytic polynucleotides. Chem. Rev. 97:371-390. Chen, Z. and Ruffner, D.E. 1996. Modified crushand-soak method for recovering oligodeoxynucleotides from polyacrylamide gel. BioTechniques 21:820-822. Conrad, R., Keranen, L.M., Ellington, A.D., and Newton, A.C. 1994. Isozyme-specific inhibition of protein kinase C by RNA aptamers. J. Biol. Chem. 269:32051-32054. Crameri, A. and Stemmer, W.P.C. 1993. 1020-fold aptamer library amplification without gel purification. Nucl. Acids Res. 21:4410. Fitzwater, T. and Polisky, B. 1996. A SELEX primer. Methods Enzymol. 267:275-301. Giver, L., Bartel, D., Zapp, M., Pawul, A., Green, M., and Ellington, A.D. 1993. Selective optimization of the Rev-binding element of HIV-1. Nucl. Acids Res. 21:5509-5516. Gold, L., Polisky, B., Uhlenbeck, O., and Yarus, M. 1995. Diversity of oligonucleotide functions. Annu. Rev. Biochem. 64:763-797.
Hermes, J.D, Parekh, S.M., Blacklow, S.C., Koster, H., and Knowles, J.R. 1989. A reliable method for random mutagenesis: The generation of mutant libraries using spiked oligodeoxyribonucleotide primers. Gene 84:143-151. Jaeger, J.A., Turner, D.H., and Zuker, M. 1989. Predicting optimal and suboptimal secondary structure for RNA. Methods Enzymol. 183:281306. Jaeger, L. 1997 The new world of ribozymes. Curr. Opin. Struct. Biol. 7:324-335. Lorsch, J.R. and Szostak, J.W. 1994. In vitro evolution of new ribozymes with polynucleotide kinase activity. Nature 371:31-36. Lyamichev, V., Brow, M.A., and Dahlberg, J.E. 1993. Structure-specific endonucleolytic cleavage of nucleic acids by eubacterial DNA polymerases. Science 260:778-783. Milligan J.F., Groebe D.R., Witherell G.W., and Uhlenbeck O.C. 1987. Oligoribonucleotide synthesis using T7 RNA polymerase and synthetic DNA templates. Nucl. Acids Res. 15:8783-8798. Osborne, S.E. and Ellington, A.D. 1997. Nucleic acid selection and the challenge of combinatorial chemistry. Chem. Rev. 97:349-370. PerSeptive Biosystems. 1998. Expedite Nucleic Acid Synthesis System: User’s Guide. PerSeptive Biosystems, Framingham, Mass. Sabeti, P.C., Unrau, P.J., and Bartel, D.P. 1997. Accessing rare activities from random RNA sequences: The importance of the length of molecules in the starting pool. Chem. Biol. 4:767-774. Singer, B.S., Shtatland, T., Brown, D., and Gold, L. 1997. Libraries for genomic SELEX. Nucl. Acids Res. 25:781-786. Tuerk, C. and Gold, L. 1990. Systematic evolution of ligands by exponential enrichment: RNA ligands to baceriophage T4 DNA polymerase. Science 249:505-510
DNA Pools for In Vitro Selection
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Tuerk, C. and MacDougal-Waugh, S. 1993. In vitro evolution of functional nucleic acids: High affinity RNA ligands of HIV-1 proteins. Gene 137:33-39. Zon, G., Gallo, K.A., Samson, C.J., Shao, K., Summers, M.F., and Byrd, R.A. 1985. Analytical studies of "mixed sequence" oligodeoxyribonucleotides synthesized by competitive coupling of either methyl- or β-cyanoethyl-N,N-diisopropylamino phosphoramidite reagents, including 2′deoxyinosine. Nucl. Acids Res. 13:8181-8196.
Contributed by Jack Pollard Harvard University Cambridge, Massachusetts Sabine D. Bell and Andrew D. Ellington University of Texas Austin, Texas
Combinatorial Methods in Nucleic Acid Chemistry
9.2.23 Current Protocols in Nucleic Acid Chemistry
In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
UNIT 9.3
While there are multiple possible configurations for in vitro selection experiments, this unit will describe one of the most common: selection of aptamers that bind to a protein target from a single-stranded RNA pool. Aptamers generated from these types of selection experiments can potentially function as protein inhibitors, and may find applications as therapeutic or diagnostic reagents. In short, a double-stranded DNA pool (see UNIT 9.2) will be transcribed to generate a single-stranded RNA pool (Basic Protocol 1). The initial concentration of protein target to be used is determined by labeling an aliquot of the pool (see Support Protocol 1) and performing the binding assay as described in Support Protocol 2. Following purification, the pool is mixed with the protein target. Binding species are separated from nonbinding species by filtration (see Basic Protocol 2). RNA:protein complexes are then eluted from the filter and binding species are amplified by a combination of reverse transcription, the polymerase chain reaction (PCR), and in vitro transcription (see Basic Protocol 3). The progress of the selection will be monitored by assaying the affinity of the radiolabeled RNA pool for the protein target after several rounds of selection (see Support Protocol 3). These steps are then repeated until a
Figure 9.3.1
Steps involved in in vitro selection of RNA aptamers.
Contributed by Sulay D. Jhaveri and Andew D. Ellington Current Protocols in Nucleic Acid Chemistry (2000) 9.3.1-9.3.25 Copyright © 2000 by John Wiley & Sons, Inc.
Combinatorial Methods in Nucleic Acid Chemistry
9.3.1
significant increase in binding is observed or until the diversity of the pool has been completely plumbed. The procedure is summarized in Figure 9.3.1. BASIC PROTOCOL 1
TRANSCRIPTION AND ISOLATION OF RNA POOLS The following protocol describes the preparation of the RNA pool to be used for selection. Starting from the dsDNA pool, the RNA is transcribed and purified by denaturing polyacrylamide gel electrophoresis. Recovery of the RNA from the gel is followed by ethanol precipitation of the RNA. Additional instructions can be found in CPMB UNIT 3.8. The directions provided here are specific for the isolation of nucleic acid pools. As is the case for the original amplification of DNA pools (UNIT 9.2), many of the procedures described here can potentially lead to the cross-contamination of different RNA selection experiments or different generations of the same selection experiment. To avoid crosscontamination, it is wise to always use barrier tips and to use disposable plastic Pasteur pipets rather than automatic micropipettors for large-volume transfers. Materials Double-stranded DNA pool (UNIT 9.2) Transcription mix (see recipe) RNase-free DNase (e.g., RQ1 DNase; Promega) 10% polyacrylamide denaturing gel (see recipe and APPENDIX 3B) 2× denaturing dye (see recipe) TBE buffer (APPENDIX 2A) 5 M NaCl 90% and 100% ethanol TE buffer, pH 8.0 (APPENDIX 2A) 37° to 42°C and 65° to 75°C water baths Fluorescent TLC plate (VWR) wrapped in plastic wrap Spectrophotometer Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (APPENDIX 3B) NOTE: All solutions and buffers should be freshly treated with DEPC (APPENDIX 2A). Use sterile, disposable plasticware where possible. See APPENDIX 2A for guidelines on standard methods to protect against contaminating RNases. Perform initial round of transcription Use the double-stranded DNA pool generated in transcription with T7 RNA polymerase.
UNIT 9.2
as a template for in vitro
1. Add ∼1 µg of double-stranded DNA template generated as in UNIT 9.2 to transcription mix for a 20 µL total reaction volume. Incubate reaction overnight at 37° to 42°C. Depending on the length and initial complexity of the pool, 1 mg of double-stranded DNA will represent ∼1013 different sequences. The transcription reaction will yield ∼10 to 50 mg of RNA, and thus from 20 to 100 copies of each sequence originally present. If more RNA is desired for initial or subsequent rounds of selection, a proportionately larger transcription reaction should be attempted. In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
The success of the transcription reaction can sometimes be monitored by observing the formation of a precipitate over time. This precipitate is likely a complex between magnesium and the pyrophosphate released from each polymerized ribotide. However, this rule is not absolute: many successful transcription reactions have no precipitate; some unsuccessful reactions have a precipitate.
9.3.2 Current Protocols in Nucleic Acid Chemistry
In general, though, the authors have found that much higher yields of RNA can be obtained from commercial transcription kits (e.g., Ampliscribe from Epicentre Technologies or Megashortscipt from Ambion) than from home-made in vitro transcription reactions. The use of such kits reduces the amount of time necessary to generate an adequate amount of transcript (1 to 4 hr rather than overnight). Depending on the amount of RNA that is desired for each round of selection, such kits may also (surprisingly) represent a more cost-efficient alternative. In some instances it will be desirable to radiolabel the RNA. For example, it is relatively easy to determine whether and how much RNA binds to a filter in the presence or absence of a protein target by radiolabeling the initial pool (Support Protocol 1). An [a-32P]nucleoside triphosphate—e.g., 0.5 ml [a-32P]UTP (NEN Life Science Products) in a 20-mL total volume—can be included in the reaction mixture in addition to all the other reagents. Varying the proportion of “hot” to “cold” nucleoside triphosphates can control the specific activity of the RNA pool. Since the overall yield of the transcription reaction will generally be important, the specific activity of the nucleoside triphosphate mixture should be varied by increasing the amount of radioactive nucleotide added, rather than by decreasing the amount of unlabeled nucleotide present. Again, commercial transcription kits can be obtained that are geared towards the incorporation of labeled nucleoside triphosphates (RiboScribe, Epicentre).
2. In order to remove DNA from the transcription reaction, after the transcription incubation has been completed, add 5 to 10 U of RNase-free DNase and incubate 25 min at 37°C. Because individual members of the double-stranded DNA library can potentially bind nonspecifically to either the target or to the selection matrix and subsequently be amplified, the DNA template should be removed from the transcription reaction according to this step, prior to proceeding with the selection. It is essential that RNase-free DNase, such as RQ1 DNase (Promega) be used, otherwise contaminating ribonucleases may destroy the newly transcribed RNA pool. An alternative would be to add RNase inhibitors to impure DNases, but such inhibitors themselves frequently contain endogenous ribonucleases that can be released during the incubation.
Purify the RNA pool The RNA pool should generally be purified by denaturing gel electrophoresis 3. Prepare a 0.75-mm thick, denaturing 10% acrylamide gel (see Reagents and Solutions and, e.g., APPENDIX 3B). A 10% acrylamide concentration is convenient for the purification of RNA molecules from 45 to 70 nucleotides in length. However, the concentration of acrylamide used to separate the full-length transcript from incomplete transcripts is ultimately contingent upon the size of the RNA and should be chosen so that the RNA will migrate approximately half-way through the gel when the loading dye has reached the bottom (see APPENDIX 3B). If the RNA sample contains a significant amount of nascent structure (for example, a doped sequence population that is based on a tightly folded secondary structure) it may not fully denature. Thus, it may be advisable to warm the gel to ∼55°C by first pre-running the gel at a higher voltage (300 to 400 V). The temperature of the gel can be monitored using adherent thermometers (VWR). In some cases, very large amounts of RNA may need to be purified (for example, the initial transcription of an extremely complex DNA library may yield upwards of a milligram or more of an RNA library). In these instances, it may be desirable to purify the RNA library by either gel-filtration or ion-exchange chromatography (e.g., Qiagen RNA kit). However, the purification of the initial or subsequent pools should never be neglected, as foreshortened amplicons can arise and overtake selected populations.
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9.3.3 Current Protocols in Nucleic Acid Chemistry
4. Fully denature the RNA pool by adding an equal volume of 2× denaturing dye, and heat the RNA-dye mix 3 min at 65° to 75°C. Although each species in the pool has a different sequence and shape, they should migrate similarly when fully elongated. Using a higher temperature or longer denaturing time risks hydrolysis of the RNA into smaller fragments by the high concentration of Mg2+ present in the transcription buffer.
5. Thoroughly rinse each well with TBE buffer using a plastic Pasteur pipet prior to loading (to remove urea, which will otherwise leach into the wells and form a barrier between the loaded sample and the gel). Load samples directly on the gel at 10 to 20 µg per 1-cm-wide lane (i.e., load a single transcription reaction in 2 to 3 lanes of the gel). Run electrophoresis for 1 to 2 hr at 150 to 250 V, until the bromophenol blue dye front reaches the bottom of the gel. If the wells are not cleaned prior to loading, the resolution of the separation can be compromised, especially if large amounts of RNA are being isolated.
6. Visualize the RNA bands by UV shadowing on a fluorescent TLC plate covered with plastic wrap, then excise the bands. Be sure to cut with a sharp razor blade and cut only the shadowed regions that contain the bulk of the RNA. There may be extra bands in the lane that correspond to incomplete transcripts or undigested DNA. The use of a size standard in a neighboring lane is recommended. Note, however, that the size standard should not itself be amplifiable, as cross-contamination of a single sequence with the RNA pool would drastically skew the distribution of sequences in the purified pool. Similarly, the razor blade used for excision should not have come into contact with other potentially amplifiable sequences, and should either be fresh or should have been cleaned extensively. Finally, if multiple selections are being carried out in parallel they should not be purified on the same gel.
7. Immerse the gel slices in RNase-free water at ∼400 µL water/cm2 of gel (typically, slices from 2 lanes) and incubate at 37°C overnight with agitation to elute the RNA pool. For a quicker elution step, incubate the slices at 65° to 75°C for 1 hr. However, the amount of RNA recovered will be lower, and there is a greater risk of degradation. The gel can be macerated to increase the speed or efficiency of recovery, but in this case small fragments of acrylamide may remain in the eluant. The eluate can be filtered through an 0.45-µm nitrocellulose membrane to remove acrylamide fragments.
Collect and quantitate the RNA 8. Use a plastic Pasteur pipet to separate the RNA-containing eluate from the gel slice. Add NaCl (from 5 M stock) to a final concentration of 0.3 M and ethanol precipitate the RNA by adding 2 vol ethanol. Mix and incubate at −20°C for 30 min or −70°C for 10 min. Microcentrifuge 20 to 40 min at maximum speed, 4°C, to recover the precipitate. Smaller RNA molecules (20 to nucleotides in length) can be more efficiently precipitated with 2.5 vol of ethanol. The authors frequently include 1 mL of a 1 mg/mL glycogen solution to increase the yield of nucleic acid precipitate and to better visualize the pellet. If the selection target binds to or interacts with glycogen, then this step should be omitted. Transfer RNA can also be used as a carrier, but will obfuscate the quantification of the pool RNA (see below). In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
9. Wash the RNA pellet with cold 90% ethanol and dry the pellet. The pellet can be air dried, dried under a nitrogen or argon stream, or dried in a SpeedVac evaporator. The first method is least likely to result in cross-contamination of nucleic acid
9.3.4 Current Protocols in Nucleic Acid Chemistry
species; the last method is least likely to lead to degradation. In any event, keep the tube covered with Parafilm to avoid inadvertent nuclease contamination (poke holes in the Parafilm with a sterile needle to allow evaporation to proceed). If the RNA pool is particularly short (≤50 nucleotides) use cold 95% ethanol for the wash step.
10. Resuspend the RNA pellet in 25 µL TE buffer, pH 8.0. To avoid disturbing the composition of the selection buffer, the pellet can also be resuspended in RNase-free water (APPENDIX 2A). However, the small amount of EDTA present in TE buffer will limit ribonuclease degradation of the pool, since ribonucleases frequently require a divalent metal. In some instances, though (e.g., small-volume PCR reactions), the presence of EDTA may have to be compensated for by adding more magnesium to the reaction.
11. Estimate the quantity of the RNA photometrically by measuring the absorbance at 260 nm. Use an extinction coefficient of 0.025 mL cm−1mg−1 (see, e.g., CPMB APPENDIX 3D).In practical terms, measure the A260 of a 1:500 dilution of the sample (2 mL dissolved in 1 mL RNase-free water) and multiply the absorbance by 20 to obtain the number of mg/mL in the original sample. Do not attempt to calculate concentrations using absorbance readings less than ∼0.03. The A260/A280 ratio should be somewhere between 1.8 and 2.2. Ratios outside of this range make the purity of the original RNA sample suspect (with residual acrylamide being the most likely contaminant), and the sample should be reprecipitated prior to use.
RADIOLABELING THE RNA FOR USE IN AN INITIAL AFFINITY ASSAY Radioactive RNA can be generated either by incorporation of an [α-32P]nucleoside triphosphate during transcription or by transfer of the terminal phosphate of γ-32P ATP to the 5′ terminus of a dephosphorylated RNA molecule. The authors tend to prefer the latter method, despite the additional labor involved in preparation, because the specific activity of the sample is higher, less RNA is required for assays, and dissociation constants are correspondingly easier to compute.
SUPPORT PROTOCOL 1
Materials RNA pool (see Basic Protocol 1) 10× alkaline phosphatase buffer (Boehringer Mannheim) Calf alkaline phosphatase (Boehringer Mannheim) 1:1 phenol/chloroform (APPENDIX 2A) Chloroform 5 M NaCl 90% and 100% ethanol 10× PNK buffer (New England Biolabs) T4 polynucleotide kinase (PNK; New England Biolabs) 167 mCi/ml [γ-32P]ATP (7000 Ci/mmol; ICN) 4 M ammonium acetate 42° and 75°C water baths NOTE: All solutions and buffers should be freshly treated with DEPC (APPENDIX 2A). Use sterile, disposable plasticware where possible. See APPENDIX 2A for guidelines on standard methods to protect against contaminating RNases. Combinatorial Methods in Nucleic Acid Chemistry
9.3.5 Current Protocols in Nucleic Acid Chemistry
Dephosphorylate the 5′ triphosphate termini of the isolated RNA pool 1. Mix the following components: 1 µg RNA in <3.5 µL volume 0.5 µL 10× alkaline phosphatase buffer 1 µL (1 U) calf alkaline phosphatase x µL RNase-free water for a total reaction volume of 5 µL. The RNA sample may need to be reprecipitated to obtain an adequately concentrated sample. If so, the precipitate can be resuspended directly in the reaction buffer or mixture. Calf alkaline phosphatase is preferred over bacterial alkaline phosphatase because the activity can be heat-killed (see step 4) prior to the addition of the radiolabel.
2. Incubate at 42°C for 20 min to 2 hr. 3. Add 95 µL RNase-free water. 4. Heat denature the calf alkaline phosphatase 10 min at 75°C. 5. Perform a phenol/chloroform extraction (see Basic Protocol 2, step 10). If the sample will be gel-isolated, this step can be omitted. If the radiolabeled sample will merely be precipitated prior to use, this step should be included.
6. Ethanol precipitate the RNA in the presence of 0.3 M NaCl and wash the pellet with 90% ethanol (see Basic Protocol 1, steps 8 and 9). Avoid precipitating RNA in the presence of ammonium acetate since ammonium ions inhibit the T4 polynucleotide kinase used in the next step.
7. Resuspend the dried pellet in a minimal volume (3 to 10 µl) of RNase-free water. Perform kinase reaction 8. Set up the kinase reaction as follows: 0.5 to 3 µL dephosphorylated RNA pool (from step 7) 0.5 µL 10× PNK buffer 1 µL (10 U) T4 polynucleotide kinase (PNK) 0.5 µL (83 µCi) [γ-32P]ATP (7000 Ci/mmol, ICN) x µL RNase-free H2O for a total volume of 5 µL. Only a very small amount of RNA will be used in the binding assay (∼50 pM in a 100 mL reaction). Unless multiple experiments are contemplated, the specific activity of the sample can be kept quite high by using a very small amount of RNA in the kinase reaction.
9. Incubate for 1 hr at 37°C. 10. Add 95 µL RNase-free water. If the sample will be gel-isolated, this step can be omitted.
11. Perform a phenol/chloroform extraction (see Basic Protocol 2, step 10). In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
If the sample will be gel-isolated, this step can be omitted.
12. Ethanol precipitate the RNA in the presence of 2.0 M ammonium acetate (i.e., by adding an equal volume of 4.0 M ammonium acetate).
9.3.6 Current Protocols in Nucleic Acid Chemistry
The use of ammonium acetate inhibits the precipitation of nucleotides and small transcripts. However, if the RNA pool is short, the precipitation may also be inefficient. If the sample will be gel-isolated, then this step can be omitted.
13. Optional: In order to fully separate the radiolabeled RNA pool from unincorporated nucleotides, partially degraded transcripts, and enzymes, isolate the transcript as described in Basic Protocol 1, steps 3 to 9). If this is done, the phenol/chloroform extractions and the final precipitation of the RNA (steps 10 to 12 of this protocol) can be omitted. The chief disadvantages of gel isolation are the time required for sample preparation and the relatively low efficiency of recovery of the radiolabeled RNA pool. However, since only a small amount of RNA pool is required for the binding assay such low yields can frequently be tolerated. The authors frequently gel isolate radiolabeled RNA pools to ensure the integrity of RNA samples prior to carrying out binding assays.
BINDING ASSAY WITH THE END-LABELED RNA POOL TO DETERMINE THE OPTIMAL PROTEIN CONCENTRATION FOR SELECTION
SUPPORT PROTOCOL 2
To determine the initial concentration of a protein target to be used in a selection experiment, it is necessary to measure the affinity of the unselected pool for the protein target (for theoretical considerations, see UNIT 9.1). The aggregate dissociation constant of the pool:protein complex can be calculated by determining the fraction of radioactively labeled RNA that can be bound at various protein concentrations. The radiolabeled RNA is incubated in the binding buffer and protein solutions are added. The binding reaction is filtered through a vacuum manifold containing nitrocellulose and nylon membranes and the fraction of RNA bound to the target is calculated to obtain a value for the dissociation constant. The nitrocellulose membrane will capture RNA:protein complexes, while the nylon membrane will capture all free RNA that flows through the nitrocellulose membrane. Materials Radiolabeled RNA pool (Support Protocol 1) Binding buffer (see Critical Parameters) Target protein 65° to 75°C water bath Milliblot apparatus (Schleicher & Schuell) Nylon transfer membrane (Hybond N+, Amersham Pharmacia Biotech) 0.45-µm nitrocellulose transfer and immobilization membrane (Midwest Scientific) Glass plate Phosphorimager (Molecular Dynamics) and screen or X-ray film and densitometer Graphing software (e.g., Kaleidograph from Synergy Software) NOTE: All solutions and buffers should be freshly treated with DEPC (APPENDIX 2A). Use sterile, disposable plasticware where possible. See APPENDIX 2A for guidelines on standard methods to protect against contaminating RNases. Set up binding reactions 1. Collect the RNA precipitate by centrifugation and resuspend the radiolabeled RNA in a minimal volume (i.e., 5 to 10 µL) of RNase-free water. Dilute the RNA sample with binding buffer to a final concentration of 100 pM.
Combinatorial Methods in Nucleic Acid Chemistry
9.3.7 Current Protocols in Nucleic Acid Chemistry
The concentration can be very roughly estimated by assuming full recovery of the RNA sample. Differences between estimated and actual concentrations are less important because the RNA sample will be limiting relative to the amount of protein sample present in the binding reaction. The binding assay will yield 11 data points in triplicate (see below). Since each data point will be generated from a 50-mL binding reaction, 2 mL of the RNA solution should be adequate. If the specific activity of the RNA is not high enough, a higher concentration of RNA may be used, but that will complicate the assumption that RNA is limiting and hence the calculation of the Kd.
2. To ensure that each species in the RNA pool folds into the most accessible or most stable conformation, heat the RNA pool in 25-µL binding buffer to 65° to 75°C for 3 min and then allow the sample to cool to room temperature over ∼10 min. 3. Add 25 µL of the protein target in binding buffer to the thermally equilibrated RNA from step 2. Use ten different protein concentrations ranging from 1 µM to 50 pM. Also include one data point with no protein to measure the filter-binding ability of the pool itself. The original protein solution should be sufficiently concentrated for all of the dilutions. To ensure consistency between samples, serial dilutions of the 1 mM sample can be made. The authors suggest the following concentrations: 1 mM, 333 nM, 111 nM, 37 nM, 12 nM, 4.1 nM, 1.4 nM, 460 pM, 152 pM, 51 pM, (i.e., 1 mM, and subsequent 1/3 dilutions) and a “no protein” control. For statistically significant results perform the binding assay in triplicate.
4. Incubate the binding reaction at room temperature for 15 min to 1 hr (see Critical Parameters). Perform filter binding 5. Assemble the Milliblot apparatus (Fig. 9.3.2). Lay the nylon transfer membrane on top of the perforations in the middle section. Moisten the nylon membrane and lay the nitrocellulose membrane on top of the nylon membrane, taking care to avoid the formation of bubbles between the two membranes. Cover and tighten the brackets. Prior to filtering the binding reactions, wash the wells that will be used with binding buffer and check for leaks. When the manifold is used in conjunction with an aspirator, turn the
In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
Figure 9.3.2
Assembly of the Milliblot apparatus used for binding assays.
9.3.8 Current Protocols in Nucleic Acid Chemistry
water faucet to a level that causes liquid to pass slowly through the membranes (i.e., 100 mL every 3 sec). Since there are so many binding reactions, it is more convenient to use a manifold apparatus that can accommodate multiple filtrations (up to 96 slots) than to assemble 33 individual filter holders.
6. Filter the binding reactions and wash three times with binding buffer. When pipetting onto the manifold, dispense the liquid slowly and evenly. Try to keep the membrane constantly hydrated during each wash step. Keep the pipet tip close to the membrane to avoid bubble formation, but not so close as to risk damaging the membrane.
7. Disassemble the manifold apparatus and transfer the membranes to a clean paper towel. Air dry for ∼5 min. Handle membranes with a clean pair of forceps or tweezers. The forceps or tweezers can be quickly cleaned with ethanol and RNase-free water prior to contacting the filters.
8. Transfer the membranes to a glass plate, cover with plastic wrap, and expose to a phosphor screen (e.g., Phosphorimager) or X-ray film for 4 to 12 hr. If the samples have a very high specific activity, the exposure time can be reduced to 5 to 60 min.
9. Measure the radioactivity using the Phosphorimager or a densitometer if X-ray film was used to develop the image, and calculate the binding percentages as follows: Fraction bound = cpm on nitrocellulose/(cpm on nitrocellulose + cpm on nylon) If X-ray film was used to develop the image, then a digitizer (densitometer) should yield similar results to those obtained with a Phosphorimager.
10. Plot the fraction bound as a function of the concentration of unbound protein. Fit the points to a curve using graphing software (e.g., Kaleidograph) and obtain a value for the aggregate parent dissociation constant. Within the Kaleidograph program, fit the curve using the equation y = m1m0/(m0 + m2), where y = the fraction of RNA bound, m0 = concentration of unbound protein, m1 = the extrapolated activity of the RNA at an infinite protein concentration (maximal value of fraction bound), and m2 = the apparent dissociation constant. The apparent Kd is equal to the concentration of unbound protein at half the maximal value of fraction bound.
ISOLATING A FUNCTIONALLY ENRICHED POOL In the following protocol, the RNA pool is partitioned to isolate those species that bind to the target protein and not to the filter. RNAs that are coimmobilized with the target are eluted off the filter under denaturing conditions and subsequently isolated and amplified. Materials RNA pool (see Basic Protocol 1) Binding buffer (see Critical Parameters) Elution buffer (see recipe) 1:1 phenol/chloroform (see APPENDIX 2A), ice-cold Chloroform Isopropanol 65° to 75°C and 100°C water baths Filter holders (Nuclepore)
BASIC PROTOCOL 2
Combinatorial Methods in Nucleic Acid Chemistry
9.3.9 Current Protocols in Nucleic Acid Chemistry
13-mm, 0.45-µm HAWP nitrocellulose disk filters (Millipore) 5-ml syringe Vacuum manifold NOTE: All solutions and buffers should be freshly treated with DEPC (APPENDIX 2A). Use sterile, disposable plasticware where possible. See APPENDIX 2A for guidelines on standard methods to protect against contaminating RNases. Partition the pool 1. Use ∼5 µg of the RNA pool (∼1013 to 1014 sequences) for selection. Using significantly lower quantities of RNA may affect the diversity of the population in the initial rounds of selection. Using significantly higher quantities may lead to precipitation of the nucleic acid pool. Irvine et al. (1991) have devised a formula to determine the optimum protein and RNA concentration in order to minimize the number of rounds of selection, based on the Kd of the starting pool, the desired Kd,, and the fraction of free RNA molecules that partitions as nonspecific background versus the fraction of RNA molecules that forms specific RNA:protein complexes. Refer to UNIT 9.1 for additional details. Empirically, the concentrations of many available protein targets will be in the nanomolar range, and a 1- to 10-fold excess of the RNA pool should suffice for early rounds of selection. If only a small amount of RNA pool is initially recovered from the gel, be sure to save at least some sample for the “no protein” control (see below).
2. To ensure that each species in the RNA pool folds into the most accessible or most stable conformation, heat the RNA pool in 50 to 100 µL binding buffer (see Critical Parameters for discussion on choosing a binding buffer) to 65° to 75°C for 3 min and then allow the sample to cool to room temperature over ∼10 min. Since ionic strength, monovalent and divalent cation concentrations, pH, temperature, and buffer concentrations can all influence interactions with the target, it is usually wise to keep all of these parameters constant during the early rounds of selection when productive binding species are accumulating. Hence, the binding buffer, equilibration time, and preparation of the RNA for selection should be kept uniform until a significant interaction between pool and target is observed (see Critical Parameters for discussion of stringency of selection). Higher temperatures can be used for thermal equilibration, but the presence of divalent metal ions in the selection buffer can lead to RNA degradation.
3. Prior to the addition of the protein target, perform a negative selection to remove any filter-binding species that may be in the population. Moisten a filter disk with buffer and lock it into a filter holder (Fig. 9.3.3).
In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
Negative selection to remove filter-binding species is an extremely important step in the selection procedure. Filter-binding species are typically more numerous in a naive RNA population than are aptamers. If filter-binding species are not efficiently sieved from the population, they will quickly accumulate to the point where it may be difficult (and likely impossible) to select protein-binding species. If the potential for accumulating filter-binding species is large (i.e., the target has a low initial affinity for a pool, or selections with DNA pools), then repeat the preselection filtration to remove any filter-binding species that may persist or carry out a post-selection filtration (see optional steps 14 to 17, below). If filter-binding species do accumulate during a selection experiment, it is usually wisest to repeat the selection starting with a different pool that can be amplified with different primers. In addition to filter-binding species, replication parasites (see Critical Parameters for discussion on parasites) can accumulate in and overrun a selected population. A separate regime is required to avoid these selection predators.
9.3.10 Current Protocols in Nucleic Acid Chemistry
Figure 9.3.3
Components and assembly of filter holder used during selection.
4. Load the binding buffer onto the filter. Place the pipet tip just above the filter to avoid the formation of any bubbles. Lock a 5-ml syringe to the top of the filter holder and apply gentle pressure to force the liquid out of the filter holder and into a collecting tube. Prior to filtering the RNA, it is important to wash the nitrocellulose filter disk with binding buffer and check for leaks in the assembled filter holder. The syringe should form a tight seal with the filter holder. The pressure applied should be just enough to force the liquid through without rupturing the membrane. The formation of foam at the bottom of the filter holder or the presence of a hissing sound when pressure is applied indicates that the pressure is too high, and the integrity of the seal or the membrane may have been breached. Test for leaks every time the filter holder is assembled in order to avoid substantial loss of sample.
5. Load the RNA solution onto the filter. Place the pipet tip just above the filter to avoid the formation of any bubbles. Lock a 5-ml syringe to the top of the filter holder and apply gentle pressure to force the liquid out of the filter holder and into a collecting tube. Since there will still be some amount of liquid retained by the filter and filter holder, it is necessary to wash the filter with an equal amount of binding buffer to maximize the collection of non-filter-binding species. Discard the filter. Table 9.3.1
Round 1 2 3 4 5 6
Progress of N30 Selection Against bFGFa
Input (RNA) nM 800 800 800 800 800 800
Input (BFGF) (RNA):(bFGF) nM 760 1.05 760 1.05 76 10.5 76 10.5 13 61.5 13 61.5
% bound to protein 2.1 — — 6.0 — 17.0
% bound to filter 2.3 — — 4.0 — 0.4
aPools were assayed in a 50-µL reaction at a concentration of 75 nM in the presence and absence of equimolar protein.
Combinatorial Methods in Nucleic Acid Chemistry
9.3.11 Current Protocols in Nucleic Acid Chemistry
6. Add the protein target and any competitors, specific and/or non-specific, to the filtrate. Allow the binding reaction to equilibrate. In selection experiments that targeted the cytokine bFGF, the authors used an equimolar protein-to-RNA ratio for the first two rounds of selection and decreased it 10-fold after two rounds and 60-fold after another two, yielding a functionally-enriched pool after 6 rounds of selection and amplification (Table 9.3.1). The final volume of the binding reaction should be from 100 to 200 mL. In addition, to ensure that the selected RNAs are actually binding to the target and not to the filter, a parallel binding reaction in the absence of protein can be carried out intermittently. The authors strongly suggest that “no protein” controls be scrutinized after 0, 5, 8, and 11 rounds of selection. The choice of selection conditions is probably the second most important determinant (following the choice of target) for whether a selection experiment will succeed or fail. While general guidelines for modulating the stringency of selection can be recommended (see Critical Parameters for comments on the stringency of selection), every target and every selection are different and no precise guidelines for success can be provided. In general, the stringency of selection should be lower in the early rounds of selection and higher in the later rounds. This will give binding species an opportunity to establish themselves in the population relative to filter-binding species. It should be noted that there is some danger of cross-contaminating the selected pool with the “no protein” control. Basically, executing the “no protein” control is identical to selecting for protein-independent (filter) binding species, hence DNA arising from the “no protein” control should be handled with care.
7. Attach the filter holder to a vacuum manifold (which is used here to maintain a constant negative pressure during filtration, so that each round of selection is similar and reproducible). Apply a negative pressure of 5 in. of Hg to the filter holder. Pipet the binding reaction directly onto the filter with the tip just above the filter, avoiding the formation of bubbles, which may lead to an uneven application of the sample to the filter and impede the flow of liquid through the filter. Wash the filter with 3 vol of binding buffer. Varying the strength of the vacuum, uneven application of the sample, and formation of bubbles during wash steps may result in inefficient sieving of binding from nonbinding species, and hence may reduce the efficiency of an individual round of selection. However, the selection as a whole is fairly robust with respect to changes in these parameters. In other words, even if steps are not performed perfectly, the selection can be carried forward.
Elute RNA off the filter 8. Remove the filter containing RNA:protein complexes from the filter holder using sterile forceps and place it in a 1.5-mL microcentrifuge tube. Transfer the filter quickly, in order to avoid ribonuclease contamination from the surrounding environment. 9. Add 200 µL of elution buffer and heat for 5 min at 100°C to elute RNA molecules from the protein and filter. Remove the eluate to a tube and repeat with fresh elution buffer. Two shorter, smaller-volume elutions will more efficiently recover intact RNA than one long, large-volume elution. In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
10. To remove residual peptide fragments or proteins that may have coeluted with the RNA, add an equal volume (i.e., 400 µl) of cold, 1:1 phenol/chloroform. Vortex, then microcentrifuge 1 min at maximum speed to separate the liquid phases (the RNA should be in the top, aqueous phase). Transfer the aqueous phase to a new 1.5-mL microcentrifuge tube.
9.3.12 Current Protocols in Nucleic Acid Chemistry
Avoid transferring phenol/chloroform with the aqueous layer, as it can interfere with subsequent enzyme reactions. Nevertheless, the aqueous phase will sometimes appear milky, especially at low temperatures, due to the presence of dissolved phenol-chloroform.
11. Extract the eluate with a similar volume of chloroform to remove any residual phenol. Avoid transferring chloroform with the aqueous layer, as it can interfere with subsequent enzyme reactions.
12. Dilute the eluate with an equal volume (∼400 µL) of RNase-free water and add 800 µL of isopropanol, then chill 20 min at –20°C to precipitate. A carrier such as glycogen can be added to aid precipitation. The elution buffer contains a high concentration of urea. Dilution with 400 mL water and precipitation with isopropanol is necessary to avoid the formation of salt precipitates, which appear as oily, unstable droplets in the bottom of the microcentrifuge tube following centrifugation. If such “salt pellets” appear, additional water should be added to the sample, the mixture should be homogenized, and the precipitation repeated.
13. Microcentrifuge 30 min at maximum speed, remove the supernatant, and resuspend the RNA sample in 12 ml sterile RNase-free water. Perform an additional negative selection (optional steps) An extremely effective method for ridding the population of filter-binding species is to carry out an additional negative selection following the selection for binding species, but prior to amplification. However, at early stages of the selection, an additional post-selection filtration step may reduce the complexity of the selected population. Therefore, it is recommended that post-selection filtration only be carried out following the second round of selection. Post-selection filtration can also be used to successfully remove filter-binding species that have begun to accumulate and overrun a selected population. However, once filter-binding species have established themselves, even a combination of pre- and post-selection filtrations may not allow specific binding species to regain a selective advantage. If a simple regime of pre- and post-filtration negative selections does not succeed in drastically reducing or eliminating established filter-binding species, the selection should be repeated with a different RNA pool that can be amplified with different primers, as recommended above. 14. Resuspend the selected RNA pellet in 50 µL binding buffer. 15. Assemble the filter holder with a fresh filter disk as described above. 16. Filter the sample and wash as described above. 17. Discard the filter disk and ethanol precipitate the RNA filtrate. A carrier (glycogen) can be added to improve the efficiency of precipitation. If the binding buffer contains a high (>0.5 M) salt concentration, dilute the filtrate with an equal volume of RNase-free water and precipitate with isopropanol instead.
AMPLIFYING SELECTED BINDING SPECIES In the following steps, RNA species that survived the positive and negative selection steps are reverse transcribed to generate a cDNA library, which is subsequently amplified by PCR. The double-stranded DNA resulting from these steps comprises the pool from which the next round of selection will begin. While the authors have found that reverse transcription and PCR steps can be combined (steps 1a to 3a) for some of our selections, this is not universally true. To obtain the highest yield of RNA and DNA products, it is frequently desirable to carry out separate reverse transcription and PCR reactions (steps 1b to 3b).
BASIC PROTOCOL 3
Combinatorial Methods in Nucleic Acid Chemistry
9.3.13 Current Protocols in Nucleic Acid Chemistry
Materials Selected RNA pool (Basic Protocol 2) TE buffer, pH 8.0, or RNase-free water RT-PCR mix (see recipe) 10× RT buffer (see recipe) 20 µM 3′-end primer 20 µM 5′-end primer PCR mix (see recipe) 4 mM dNTP mix (APPENDIX 2A) AMV reverse transcriptase (USB) 6× nondenaturing dye: 0.6% bromphenol blue in TBE buffer 4% NuSieve agarose gel (FMC Bioproducts; also see e.g., CPMB UNIT 2.6) 10 µg/mL ethidium bromide solution (APPENDIX 2A) TBE buffer (APPENDIX 2A) 4 M ammonium acetate 100% ethanol Thermal cycler (e.g., MJ Research) Additional reagents and equipment for the polymerase chain reaction (CPMB Chapter 15) and agarose gel electrophoresis (e.g., CPMB UNIT 2.6) NOTE: All solutions and buffers should be freshly treated with DEPC (APPENDIX 2A). Use sterile, disposable plasticware where possible. See APPENDIX 2A for guidelines on standard methods to protect against contaminating RNases. Amplify selected binding species To amplify selected binding species via combined RT-PCR reactions 1a. Resuspend the RNA in 12 µL TE buffer or RNase-free water and add 4 µL of this RNA suspension to 96 µL RT-PCR mix. Since only 1/3 of the total sample recovered is used for amplification this will obviously restrict the proportion of successful species that are carried into the next round of selection. This is only a potential problem in the early rounds of selection. For example, if the diversity of the RNA pool was such that each species was represented only a few times on average, then a population bottleneck is unavoidable. For this reason, it is always desirable to start with an RNA pool in which each species is represented numerous times. However, the amount of sample that is committed to amplification should probably not exceed one-half to two-thirds of the sample. If the reverse transcription or any subsequent steps are unsuccessful, then the archived RNA serves as an inviolate reservoir for proceeding forward in the selection experiment. Otherwise, one will have to return to material from an earlier round.
2a. Run the following controls in parallel with the amplification of selected RNA species in order to detect nonspecifically bound RNA species and replication parasites (see Critical Parameters for discussion of parasites). a. No template control: To ensure that none of the stock solutions have been contaminated with exogenous RNA or DNA amplicons, set up a RT-PCR reaction without adding any template.
In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
b. No RT control: To ensure that amplified products are in fact derived from selected RNA species and not from endogenous or cross-contaminating DNA molecules, set up a RT-PCR reaction without the reverse transcriptase. 3a. Run the RT-PCR reaction on the thermal cycler as follows: a. 10 min at 65°C
9.3.14 Current Protocols in Nucleic Acid Chemistry
b. c. d. e.
10 min at 50°C 45 sec at 94°C 60 sec at 50°C 90 sec at 72°C.
Repeat steps c to e six more times, then follow with: f. 150 sec at 72°C. Step a allows for primers to anneal to the RNA, while step b allows the RT to generate cDNA. Steps c through e comprise the PCR cycle, and the final elongation step (f) at 72°C completes the extension of any incomplete DNA templates. The number of cycles that should initially be carried out is considered below. It should be noted that the listed conditions have been optimized for the pool used in the selections described, the N30 pool. Different pools and primers may require very different amplification conditions. (see CPMB UNIT 15.4 for comments on primer selection and, for the experimental parameters that govern reverse transcription and PCR). If using a PCR machine without a heated bonnet, cover the amplification reaction with mineral oil (e.g., Mallinckrodt). In order to avoid the accumulation of replication parasites, it may be desirable to use one of a number of “hot-start” methods for the PCR reaction (see CPMB Chapter 15). The technically simplest of these is to add reverse transcriptase following heating to 65°C in step a, and to add Taq or another thermostable polymerase following heating to 94°C in step c.
To amplify selected binding species via separate RT and PCR reactions 1b. Resuspend the RNA in 12 µL TE buffer or RNase-free water, and set up the RT reaction as follows: 4 µL RNA, diluted as described above 2 µL 10× RT buffer 10 µL 20 µM 3′-end primer 4 µL 4 mM dNTP mix 0.3 µL (10 U) AMV reverse transcriptase. Also set up “no template” control without the RNA template and “no RT” control without the reverse transcriptase. 2b. Incubate reactions 30 min at 42°C. 3b. Add 10 µL of each RT reactions to an individual tube containing 100 µL PCR mix. Conduct the PCR reaction as follows: a. 45 sec at 94°C b. 60 sec at 50°C c. 90 sec at 72°C. Repeat steps a and b six more times, then follow with: d. 150 sec at 72°C. See, e.g., CPMB UNIT 15.1 for additional information on PCR amplification.
Check for the presence of amplified, double-stranded DNA 4. Add 1.5 µL of 6× non-denaturing dye to 5 to 10 µL of the PCR reaction. Load the sample onto a 4% NuSieve agarose gel which has been presoaked in 10 µg/mL
Combinatorial Methods in Nucleic Acid Chemistry
9.3.15 Current Protocols in Nucleic Acid Chemistry
ethidium bromide solution for 10 min (e.g., CPMB UNIT 2.6). Run the gel in TBE at 125 V for 15 min. Look for products with a hand-held UV lamp or UV light box. An estimate of the minimal number of cycles needed to visualize a product band on the agarose gel can be roughly calculated. Consider that, of the 5 mg of RNA added to the selection, ∼3% likely binds to the filter and is lost during the negative selection step. Approximately 1% of the population may bind to the target. When the selected RNA is precipitated, one-third of the sample is used for RT-PCR. Therefore: (5.0 mg)(0.97) (0.01)/3 =0 .016 mg RNA. Assuming that every thermal cycle doubles the amount of DNA, a minimum of seven thermal cycles would be necessary to obtain 1 to 2 mg of DNA. This would imply that 0.1 to 0.2 mg could be loaded and readily visualized on the ethidium bromide–stained agarose gel. Thus, from 7 to 8 thermal cycles should initially be carried out and the products analyzed by gel electrophoresis. The authors frequently find this rough estimate to be true.
5. If no product bands are apparent, then carry out an additional 4 to 5 thermal cycles and again analyze the products by gel electrophoresis. If only faint product bands are apparent, then one may want to accumulate additional template via an additional 2 to 4 thermal cycles. The accumulation of double-stranded DNA is closely monitored in order to avoid “overPCR” of the sample and the concomitant accumulation of high-molecular-weight species. DNA that has been over-amplified will look blurry and disperse following analysis by gel electrophoresis. These large DNA molecules are often the result of the 3′ end of a single-stranded DNA folding back and internally priming its own extension, resulting in a long stem-loop that can be amplified by a single PCR primer (also known as single-primer artifacts). Overamplified DNA templates can also yield RNA molecules of the incorrect size following transcription. Adding 2 mL of the RT-PCR reaction to 100 mL of a fresh PCR mix and carrying out 2 to 3 additional thermal cycles can clean up DNA that has been over-amplified. If one primer is more abundant or efficient than the other is, a smaller, single-stranded DNA band or bands may also be present. The hiatus between carrying out the amplification reaction and running the agarose gel allows the sample to cool to room temperature, and can potentially result in mispriming and the accumulation of replication parasites. However, this is unavoidable and is not as serious for samples that have been partially amplified as it is for samples that are just beginning the amplification procedure. To avoid this potential problem, it is sometimes desirable to take one-fourth to one-third of the selected RNA and carry out a “ranging” RT-PCR reaction to establish the optimal number of cycles for amplification. Another one-fourth to one-third of the selected RNA can then be continuously amplified to this optimum level.
In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
The various controls (“no protein,” “no template,” “no RT”) should be amplified in parallel with the actual sample. If specifically bound RNA is templating the accumulating amplicons, then the “No RT” sample should lag the RT-PCR reaction by at least three cycles. It is devoutly hoped that no bands will be observed in the “no template” control, but if they do arise, they should lag the RT-PCR reaction by at least five cycles. If bands do arise, a distinction should be made between full-length PCR products (indicating contaminating replicons) and smaller products (likely primer amplification artifacts). If product bands in the control lanes are as prominent as product bands in the experimental lanes, then it is necessary to check or remake reagents and go back and repeat the previous round of selection. There is one exception to this rule: in the initial rounds, it is common to see a band in the “no protein” control lane because the proportion of the population that binds to the filter is typically greater than the proportion that binds specifically to the target. However, subsequent rounds of selection should result in the diminution or disappearance of the “no protein” band.
9.3.16 Current Protocols in Nucleic Acid Chemistry
Although we will consider methods for closely monitoring the progress of the selection experiment, observing the number of thermal cycles needed to visualize a double-stranded DNA band can loosely monitor the progress of the selection. The number of thermal cycles should be roughly proportional to the amount of RNA pool that originally binds to the protein. Therefore, if the RNA eluted from the “no protein” control requires more thermal cycles for full amplification than does the RNA selected in the presence of protein, it can be tentatively assumed that the selected RNA is binding to the protein. Occasionally, in the early rounds of selection, this may not be true, since a very small fraction of the pool will bind to the protein relative to the small fraction of the pool that adheres to the filter. Counting PCR cycles is, however, only a very rough (and frequently inconsistent) measure of success. In fact, it is common for the number of thermal cycles required to fully amplify selected nucleic acids to vary greatly between rounds. Direct binding assays of the RNA pool (Support Protocol 3) are a much more accurate and useful gauge of the progress of a selection experiment.
6. When a product band does appear, precipitate the PCR reaction by adding an equal volume of 4 M ammonium acetate, and, to the resulting mixture, an equal volume of ethanol (i.e., 2× the original PCR reaction volume). If a large amount of sample has been used for gel analysis (for example, if only ∼50 mL of the original RT-PCR reaction remains), then one may wish to return to the selected RNA reservoir and amplify a new DNA template using the already determined “optimal” number of thermal cycles.
Use amplified DNA template for the next round of selection 7. Centrifuge the sample and resuspend in 10 to 20 µL TE buffer. Proceed with the next round of selection starting with step 1 of Basic Protocol 1. A 100-mL RT-PCR reaction yields ∼1 to 2 mg DNA, so approximately half of the resuspended DNA sample should be used for the next transcription reaction. The remaining DNA can serve as a long-term, archival sample.
ASSAYING THE ACCUMULATION OF BINDING SPECIES In order to verify that the RNA pool has been or is being winnowed to those few sequences that bind the protein target with high affinity and specificity, the selected RNA pool should periodically be assayed for its ability to bind the target protein. The authors recommend an initial binding assay after five rounds of selection and amplification, then again every three additional rounds (the same recommendation that was made with regard to checking for filter-binding species; the two tests can be carried out in parallel). While the initial binding assay is carried out at a series of protein concentrations to gauge the amount of protein that should be used in the selection, the progress of the selection can be most simply monitored by radiolabeling the RNA and determining how much binds to a single, convenient concentration of the protein target.
SUPPORT PROTOCOL 3
Materials Pool of dsDNA after n rounds of selection Binding buffer Target protein 13-mm, 0.45-µm HAWP nitrocellulose disk filters (Millipore) Filter holders (Nuclepore) Vacuum manifold Glass plate Plastic wrap Phosphoimager and screen or X-ray film
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9.3.17 Current Protocols in Nucleic Acid Chemistry
Additional reagents and equipment for purifying a radiolabeled DNA pool (see Basic Protocol 1) and performing the filter binding assay (see Support Protocol 2) 1. Generate radiolabeled RNA pool via a “hot transcription” with α-labeled nucleoside triphosphates and purify as described in the final annotation to step 1 of Basic Protocol 1. 2. Thermally equilibrate 1 µg of the radiolabeled RNA pool after a round of selection in binding buffer as described in Support Protocol 2, step 2. 3. Add an equimolar amount of protein to the RNA pool. Incubate the binding reaction under conditions similar to those used for selection (see Support Protocol 2, steps 3 and 4). If the amount of protein sample is limited or limiting, less protein can be used in the binding reaction. However, one should be cognizant of the fact that less than 100% binding is possible. Alternatively, less protein and less RNA sample can be used, although the diminution of both components will mean that one is assaying binding under conditions more stringent than those actually used for selection. While the volume of the binding reaction could also be diminished to conserve protein, it is difficult to uniformly apply volumes less than 30 mL to the filter.
4. Prior to filtration, take a small aliquot of the binding reaction (i.e., 5 µL out of a 100 µL binding reaction) to determine the total amount of radioactive RNA in the binding reaction. Pipet the sample onto a nitrocellulose filter disk and set the disk aside on a glass plate. 5. Filter the binding reaction and wash 3 times with 200 µL binding buffer (see Support Protocol 2, steps 5 and 6). 6. Place the filters on the glass plate next to the initial aliquots of the binding reaction. 7. Cover the membranes with plastic wrap and expose to a phosphor screen (e.g., Phosphor imager or X-ray film for 4 to 12 hr). 8. Count the radioactivity using the Phosphorimager or a densitometer and calculate the fraction bound as follows: Fraction bound = (cpm of filtered solution)/[(cpm of aliquot from step 4) × (vol. of filtered solution/vol. of aliquot from step 4)]. A good result at this point would be 0.15 to 0.20 fraction bound above background (see Table 9.3.1. round 6). If binding to filter alone is too high, then filter binders are being selected and more negative selection is needed.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Denaturing dye, 2× TBE buffer (APPENDIX 2A) containing: 0.1% (w/v) bromphenol blue 7 M urea Store up to 6 months at –20°C In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
Denaturing polyacrylamide gel, 10% TBE buffer (APPENDIX 2A) containing: 10% (w/v) acrylamide
9.3.18 Current Protocols in Nucleic Acid Chemistry
0.5% (w/v) bisacrylamide 7 M urea See APPENDIX 3B for full details on pouring and running the gel.
Elution buffer 7 M urea 100 mM sodium citrate 3 mM EDTA Store up to 3 months at –20°C Prepare with RNase-free water.
PCR mix 10 mM Tris⋅Cl, pH 8.4 (APPENDIX 2A) 50 mM KCl 1.5 mM MgCl2 0.2 mM each dNTP 5% (w/v) acetamide 0.05% (v/v) Nonidet P-40 (NP-40) 0.5 µM each primer 0.2 U Taq DNA polymerase (Promega) RT buffer, 10× 500 mM Tris⋅Cl, pH 8.0 (APPENDIX 2A) 400 mM KCl 60 mM MgCl2 Store up to 6 months at –20°C RT-PCR mix 10 mM Tris⋅Cl, pH 8.4 (APPENDIX 2A) 50 mM KCl 1.5 mM MgCl2 0.2 mM each dNTP 5% (w/v) acetamide 0.05% (v/v) Nonidet P-40 (NP-40) 0.5 µM each primer 0.2 U Taq polymerase (Promega) 10 U AMV reverse transcriptase (USB) Transcription mix 40 mM Tris⋅Cl, pH 7.9 (APPENDIX 2A) 26 mM MgCl2 0.01% (v/v) Triton X-100 2.5 mM spermidine trihydrochloride 5 mM dithiothreitol 2.5 mM each ribonucleotide triphosphate 20 U RNasin (Promega) 100 U T7 RNA polymerase (New England Biolabs) Prepare fresh COMMENTARY Background Information Sol Speigelman and coworkers developed a working system for the in vitro replication and evolution of small RNA molecules over 25
years ago (Mills et al., 1967; Levisohn and Spiegelman, 1969; Kramer et al., 1974). The development of more advanced (although conceptually identical) methods for in vitro evolu-
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9.3.19 Current Protocols in Nucleic Acid Chemistry
tion as described in this chapter was potentiated by advances in the chemical synthesis of oligonucleotides and the amplification of nucleic acids, such as PCR, in vitro transcription, and 3SR (Guatelli et al., 1990). The adaptation of these methods to in vitro evolution of RNA molecules was partially due to a recognition that early evolutionary events, such as the genesis of ribozymes, could be recapitulated in a test tube, and partially due to a recognition that the ability to tailor RNA binding species and catalysts might have numerous biotechnological applications. Following the publication of key papers outlining and proving selection technologies (Ellington and Szostak, 1990; Tuerk and Gold, 1990), a much wider array of selection experiments has been attempted. To date, RNA molecules that can bind targets as small as zinc and as large as viruses and organs have been selected. RNA molecules that interact with both nucleic acid–binding proteins and non-nucleic acid binding proteins can be selected with almost equal facility from random sequence populations. These results have been thoroughly reviewed in numerous recent publications (Gold et al., 1995; Uphoff et al., 1996; Famulok and Jenne, 1998).
Critical Parameters
In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
Choosing protein targets As briefly described above, a wide variety of proteins have proven to be successful targets for selection experiments, including enzymes, transcription factors, cytokines, antibodies, and viral capsids (Gold et al., 1995; Uphoff et al., 1996; Famulok and Jenna, 1998). There is no common functional theme uniting these targets, nor can many generalities be drawn regarding their biochemistry or structure. However, it is safe to say that “good” selection targets tend to fall into two classes. First, proteins that normally bind nucleic acids will also be able to extract aptamers from a random sequence pool. The notion of a nucleic acid–binding protein can to some extent be expanded to include proteins that bind nucleotides. For example, kinases and dehydrogenases bind nucleotide cofactors and have proven to be good selection targets. Second, proteins that for whatever reason contain basic patches in their primary sequences or on their surfaces also frequently yield high-affinity aptamers. For example, many cytokines and other signal-transduction proteins bind heparin or other sulfated oligosaccharides, and can also be used to select
aptamers from random sequence populations. The anti-cytokine aptamers frequently bind to the same sites as heparin (Jellinek et al., 1993). Similarly, proteins that bind phosphate or phosphomonoester or phosphodiester bonds frequently have positively charged active sites and can be used to elicit aptamers. For example, anti-phosphatase aptamers have been selected from random sequence pools (Bell et al., 1998). This is not to say that proteins that do not fall into these categories will of necessity be poor selection targets, merely that they are not sure selection targets. For example, antibodies have frequently proven to be excellent selection targets irrespective of whether or not they bind negatively charged antigens (Keene, 1996). This likely implies that proteins that have large pockets or clefts on their surface are good selection targets. This hypothesis is further bolstered by another line of reasoning. Aptamers selected to bind proteins frequently inhibit protein function. That is, anti-antibody aptamers block interactions with antigens, anti-enzyme aptamers inhibit enzymatic activities, and so forth. This so-called ’homing principle’ may be due to the fact that aptamers have to not only form a surface that is chemically complementary to a target, but also must fold into a structure that properly presents the chemically complementary surface. The most informationally parsimonious way to achieve both functions is to fit into a pocket on a target, rather than to form a “grasping” structure that can enfold a surface protrusion of a target. Thus, the most common (and most highly represented) aptamers may be those that fit into surface crevices. In contrast, antibodies have a preformed structure for the presentation of chemically complementary surfaces, and thus can more easily grasp protruding epitopes and less easily fit into surface crevices. Overall, though, researchers should be guided not so much by these considerations, but by the results of initial binding assays with their particular protein target. If the target binds to the filter (not a given, since small, acidic proteins such as the Rop protein from E. coli will frequently pass through the filter) and shows some affinity for a random sequence pool, then it is highly probable that there will be some sequences or structures within the pool with greatly enhanced affinities for the target. Choosing a binding buffer The binding buffer should promote specific binding of nucleic acids to a protein target. The first consideration in choosing a buffer is to
9.3.20 Current Protocols in Nucleic Acid Chemistry
identify conditions under which the protein is active, or at least stable. In addition, if the selected nucleic acid species are to eventually be used in a particular environment, the selection buffer should reflect this environment. For example, if the selected nucleic acids are to be expressed in a cell, then the selection buffer should be at physiological pH and contain physiological ion concentrations. Second, there are a variety of parameters that can be used to make the RNA pool more or less “sticky.” These parameters are discussed in much greater detail in the following section on the stringency of selection. A typical binding reaction is built from one of the commonly used buffers, such as Tris⋅Cl, phosphate, or HEPES, which can hold the pH near 6 to 8, together with 50 to 200 mM NaCl or KCl and 1 to 10 mM MgCl2. However, these are merely suggestions, and aptamers have in fact been selected under a variety of buffer conditions. For example, in the selection that targeted bFGF, phosphate-buffered saline was used even though it lacked divalent cations. Similarly, ribozyme selections have been carried out in which a variety of divalent metal ions are mixed, and nascent ribozyme species “decide” which combination of metals most enhance their activities (Lehman and Joyce, 1993). An equivalent strategy could be used for the selection of aptamers. Selection matrices Due to the tremendous ratio of matrix surface area to protein surface area, matrix-binding aptamers can quickly and easily eclipse target-binding aptamers. Proteins are likely captured on nitrocellulose or modified cellulose filters via hydrophobic interactions. Nucleic acids are, by and large, too hydrophilic or charged to be similarly captured. This distinction is the basis for most filter-binding assays. However, the nucleobases of nucleic acids obviously contain large hydrophobic surface areas, and it is easy to select nucleic acids that can present nucleobases and be captured by the filter. Selected filter-binding sequences frequently contain purine (especially guanosine) tracts presented as single-stranded loops or bulges. Interestingly, hydrophobic-binding sequences selected on one hydrophobic matrix are frequently cross-reactive with other hydrophobic matrices: i.e., microtiter plate–binding species can bind tubes and filters, filter-binding species can bind tubes and microtiter plates, and so forth.
In order to avoid filter-binding sequences, the authors have filtered RNA samples multiple times in the absence of protein, and in some cases filtered samples following selection but prior to the RT-PCR step. Matrix-binding sequences can also be avoided by altering the matrices used for selection. For example, techniques such as gel mobility shifts, immunoprecipitation, and affinity chromatography have all been successfully used to sieve pools and select target-binding aptamers (Conrad et al., 1996). If filter-binding species predominate in a population even after appropriate precautions are taken, these alternative selection techniques can be used either to rid the selected population of the filter-binding species or, better yet, to restart the selection. For example, if the immunoprecipitation of RNA:protein complexes has been worked out in advance, then immunoprecipitation can be interspersed with rounds of filter-binding. Even though the selection of filter-binding sequences can be a problem, filter binding is still generally recommended as the technique of choice for most selections. Gel mobility shift experiments tend to be much more sensitive to parameters such as sample preparation, ionic strength, pH, and electrophoresis conditions than are filter-binding experiments. Moreover, just as filter-binding species can be inadvertently selected during filtration selection, RNA species with altered electrophoretic mobilities (e.g., dimers) can be selected during gel-mobility shift selections. Immunoprecipitation experiments require an additional protein reagent and in consequence anti-antibody rather than anti-target aptamers are frequently selected. Affinity chromatography or similar techniques generally require that very large amounts of target proteins be committed to the preparation of affinity matrices. If affinity elution is to be used, then even larger amounts of target proteins will be required. Moreover, aptamers that bind to agarose matrices can be selected almost as easily as aptamers that bind to nitrocellulose or modified cellulose filters (although the two, thankfully, do not cross-bind to one another’s matrices). Finally, microtiter plate panning selections encourage the accumulation of the same sorts of matrix-binding aptamers that are elicited by filter-binding selections. Stringency of selection Overall, most selection experiments are generally competitions between specifically and nonspecifically binding nucleic acid species. The authors tend to initially choose con-
Combinatorial Methods in Nucleic Acid Chemistry
9.3.21 Current Protocols in Nucleic Acid Chemistry
In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
servative binding conditions in hopes of promoting the early establishment of binding species in the population. While this may mean that low-affinity species are isolated from the pool along with high-affinity species, the lowaffinity species can eventually be removed by increasing the stringency of selection. In essence, time (the number of cycles required to purify high-affinity species) can be traded for the assurance that filter-binding species will not accumulate and predominate. A variety of parameters can be modulated in order to increase or decrease the stringency of a selection experiment. These parameters should initially be chosen based on the results of Support Protocol 2, which assays the affinity of the pool for the target and should be made progressively more stringent based on the results of Support Protocol 3. The amount of protein target. The more protein there is to bind, the easier it is to capture nucleic acid binding species. Using low amounts of protein increases competition among binding species. However, the amount of protein target available to researchers is usually limited, and thus it is easier to use a set amount of protein (usually from 0.1 to 10.0 µm per binding reaction) and to vary the RNA:protein ratio. RNA:protein ratio. By increasing the ratio of pool to target, more binding species will compete for a smaller number of targets. Typically, after a few initial rounds with an equimolar pool-to-target ratio, the ratio is increased to between 10:1 and 100:1. This increase can be effected either by increasing the amount of RNA or by decreasing the amount of protein. Because of the underlying competition between specifically binding species and nonspecifically binding species, increasing the amount of RNA is preferable to decreasing the amount of protein. For a more detailed treatment of this subject, see Irvine et al. (1991). However, the general conclusions of these mathematical models are similar to the empirical advice given here. Competitors. High concentrations of nonspecific, non-amplifiable competitors such as tRNA or bulk cellular RNA will compete with low-affinity binding species that adhere to basic patches on the surface of a protein. Typically, a 100-fold excess of tRNA is used. Similarly, specific competitors can be used to block the access of low-affinity binding species to a preferred site. Wild-type nucleic acid ligands can be used to block the binding sites of nucleic acid binding proteins. For example, during the
selection of anti-Rev aptamers, Giver et al. (1993) included a 10-fold excess of the wildtype Rev-binding element. The anti-Rev aptamers that were obtained could bind with high affinity to the RNA-binding domain of Rev and could effectively compete with the wild-type Rev-binding element. Other ligands or substrates can also be used to block the binding or catalytic sites of non-nucleic acid binding proteins. For example, during the selection of antibFGF aptamers, Jellinek et al. (1993) included heparin, a natural ligand for bFGF. The antibFGF aptamers that were obtained could bind with high affinity to the heparin binding site and could effectively compete with heparin. Cation concentration. Monovalent cations (such as Na+) and divalent cations such as Mg2+ stabilize the structure of RNA molecules and contribute to both specific and nonspecific binding. Decreasing monovalent and/or divalent cation concentrations therefore can therefore increase the stringency of the selection. However, it is unclear, in advance, whether specific or nonspecific binding species will be more favored by such a change. Moreover, since binding species that require a monovalent and/or divalent cation to fold into shapes that are chemically complementary to a target may be favored in the early rounds of selection, potentially high-affinity binding species may be lost by changing the binding buffer late in the selection experiment. It is better to attempt to change the buffer dependency of aptamers by partial randomization and re-selection following the initial selection experiment, rather than to attempt to change the buffer dependency during the selection. Conversely, higher concentrations of monovalent cations (generally sodium or potassium) increase the structural integrity of folded nucleic acids by neutralizing the close approach of nucleic acid strands. However, higher monovalent ion concentrations also suppress electrostatic interactions with targets. Thus, paradoxically, both “low” and “high” monovalent ion concentrations can be used to increase the stringency of a selection experiments. Higher concentrations of divalent cations such as magnesium help to maintain the structural integrity of RNA molecules and potentially facilitate the formation of salt bridges between acidic residues and the phosphate backbone. Equilibration time. Longer equilibration times give stronger binding species a greater chance to bind to the target, since weaker binding species more quickly dissociate from the target. In general, though, species with nano-
9.3.22 Current Protocols in Nucleic Acid Chemistry
molar dissociation constants or lower can be readily selected by allowing the reaction to equilibrate for 5 min or more. The authors usually allow up to 30 min for the binding reaction in order to allow for slow folding or refolding steps in the presence of the target. However, longer equilibration times may not be possible for proteins that are inherently unstable or that themselves undergo slow, bufferor temperature-induced conformational changes. Dilution of binding buffer. Similarly, diluting the binding reaction by 10- to 20-fold just prior to filtration will favor the selection of RNA:protein complexes with low dissociation constants over RNA:protein complexes with higher dissociation constants. Baskerville et al. (1995) have successfully used this technique to select high affinity anti-Rex aptamers. Amount and composition of wash. Increasing the number of times a filter is washed and the volume of the buffer used for the washes should preferentially increase the retention of high-affinity binding species relative to low-affinity and nonspecific binding species. It is generally recommended that the same buffer be used for selection and for wash steps, in order to avoid changing the conditions under which aptamers are selected. However, the stringency of the selection can potentially be manipulated by changing the buffer used for the wash steps. For example, if monovalent cation concentrations are limited in the binding buffer due to requirements for the stability or activity of a protein target, a separate wash buffer that contains a higher salt concentration can be used to challenge captured RNA:protein complexes. Parasites Replication parasites differ from matrixbinding aptamers, but can interfere with the selection of target-binding aptamers in the same way. Reverse transcriptase, Taq polymerase, and T7 RNA polymerase all have some preference for which sequences they will copy or reproduce. These preferences are generally not obvious when constant sequence nucleic acids are being synthesized. However, in selection experiments many cycles of amplification are carried out, and differences in the rates of synthesis are also proportionately amplified, leading to the selection of sequences that have no function other than to replicate optimally. For example, during the polymerase chain reaction if a primer designed to bind to a constant sequence region instead recognizes a partially complementary sequence within a random se-
quence region, it can set down and generate a smaller amplicon. The smaller amplicon will generally be amplified more quickly than the larger amplicon, and thus can potentially outcompete full-length species selected for binding function. Depending on the relative advantage of the replication parasite relative to an aptamer, even if the replication parasite is partially removed from the population during each selection step, enough molecules may remain to overrun the amplification reaction and displace the functionally selected aptamer. This is especially true if the amplification parasite also happens to be a filter-binding species. It is for this reason that the authors of this unit strongly recommend that DNA templates and/or RNA molecules be size-selected in each round. The nascent reproductive differences between nucleic acid species can be grossly amplified by amplification methods that allow continuous reproduction of the nucleic acids, such as isothermal amplification or 3SR (Guatelli et al., 1990). For example, Breaker and Joyce (1994) generated an extremely robust replication parasite, RNA Z, during a selection designed to generate catalytic variants of a group II intron. Similarly, the authors have generated replication parasites of isothermal amplification reactions from completely random sequence pools (K. Marshall, pers. comm.). Interestingly, these isothermal amplification parasites were actually larger than the initial RNA species and represented recombination events between individual members of the pool. Airborne copies of these replication parasites can readily “seed” isothermal amplification reactions and overrun pool molecules that are initially present in even million-fold excess. In this respect, the replication parasites of isothermal amplification reactions resemble the midi-variants or “monsters” of Qβ replicase amplification reactions, and are equally hard to vanquish, once established. It is for this reason that the authors strongly recommend the sometimes tedious but inherently faithful regime of reverse transcription, PCR, and in vitro transcription for the amplification of RNA pools. However, successful selections have been carried out that have relied upon isothermal amplification (see, for example, Breaker et al., 1994; Wright and Joyce, 1997; Wlotzka and McCaskill, 1997), and this admonition can most confidently challenged if the starting pool is a partially randomized binding site or ribozyme. The reason is that isothermal amplification parasites are more likely to be found in or derived from a “deep random” pool than in
Combinatorial Methods in Nucleic Acid Chemistry
9.3.23 Current Protocols in Nucleic Acid Chemistry
a pool that centers on a given functional sequence.
Anticipated Results Table 9.3.1 shows the progression of a selection carried out in the authors’ lab against bFGF with an RNA pool with a 30 nucleotide randomized region. In order to evaluate the success of a selection experiment, it was necessary to compare the affinity of the selected pool versus the affinity of the unselected pool for the protein target (Support Protocol 3). When assaying the pool after a round of selection, it was necessary to validate the fraction of the pool that bound to the protein by including a no protein control. If the accumulation of matrix-binding species had been evident, more stringent negative selections could have potentially been used to control or reduce their numbers.
Time Considerations
In Vitro Selection of RNA Aptamers to a Protein Target by Filter Immobilization
The time required to go from one pool of selected DNA templates to the next is ∼24 to 72 hr, depending on the researcher and the demands of the particular selection experiment. Minimally, a transcription reaction takes ∼4 hr, and the ensuing DNAse, heat denaturation and gel purification steps can take another 2 to 3 hr. Elution for 8 to 10 hr yields an adequate amount of RNA to be used it the subsequent binding reaction. After precipitation and quantification of the RNA (1 hr), the preselection filtration, incubation with target, and selection steps can be performed in 2 hr. Elution of protein-RNA complexes, subsequent extractions, and another precipitation step take another 2 hr. The amount of time needed to see a DNA product varies according to the number of PCR cycles needed to amplify the pool to a certain amount, and that number is inversely related to the abundance of target-binding species that survived the selection. Nevertheless, the RT-PCR steps, followed by precipitation of the DNA templates that can be added to the transcription mix, should consume ∼3 to 4 hr. The amount of time it takes to carry out the entire selection is contingent upon the number of rounds needed to accumulate target-binding species. That number, in turn, varies depending upon the initial affinity of the unselected pool for the target and on the stringency with which each round of the selection is carried out. When additional steps such as radiolabeling and assaying unselected and selected pools are taken into account, an entire selection experiment can take up to 2 to 3 weeks. It is for this reason that
the authors have recently developed automated methods for selection experiments (Cox et al., 1998) that can speed the entire process by an order of magnitude.
Literature Cited Baskerville, S., Zapp, M., and Ellington, A.D. 1995. High resolution mapping of the human T-cell, leukemia virus type 1 rex-binding element by in vitro selection. J. Virol. 69:7559-7569. Bell, S.D., Denu, J., Dixon, J.E., and Ellington, A.D. 1998. RNA molecules that bind to and inhibit the active site of a tyrosine phosphatase. J. Biol. Chem. 273:14309-14314. Breaker, R. and Joyce, G.F. 1994. Emergence of a replicating species from an in vitro RNA evolution reaction. Proc. Natl. Acad. Sci. U.S.A. 91:6093-6097. Breaker, R., Banerji, A., and Joyce, G.F. 1994. Continuous in vitro evolution of bacteriophage RNA polymerase promoters. Biochemistry 33:1198011986. Conrad, R.C., Giver, L., Tian, Y., and Ellington, A.D. 1996. In vitro selection of nucleic acid aptamers that bind proteins. Methods Enzymol. 267:336367. Cox, J.C., Rudolph, P., and Ellington, A.D. 1998. Automated DNA selection. Biotechnol. Prog. 14:845-850. Ellington, A.D. and Szostak, J.W. 1990. In vitro selection of RNA molecules that bind specific ligands. Nature 346:818-822. Famulok, M and Jenne, A. 1998. Oligonucleotide libraries--variatio delectat. Curr. Opin. Chem. Biol. 2:320-327. Giver, L., Bartel, D., Zapp, M., Green, M., and Ellington, A.D. 1993. Selective optimization of the Rev-binding element of HIV-1. Nucl. Acids Res. 23:5509-5516. Gold, L., Polisky, B., Uhlenbeck, O., and Yarus, M. 1995. Diversity of oligonucleotide functions. Annu. Rev. Biochem. 64:763-797. Guatelli, J., Whitfield, K., Kwoh, D., Barringer, K.J., Richman, D., and Gingeras, T.R. 1990. Isothermal, in vitro amplification of nucleic acids by a multienzyme reaction modeled after retroviral replication. Proc. Natl. Acad. Sci. U.S.A. 87:1874-1878. Irvine, D., Tuerk, C., and Gold, L. 1991. SELEXION: Systematic evolution of ligands by exponential enrichment with integrated optimization by non-linear analysis. J. Mol. Biol. 222:739761. Jellinek, D., Lynott, C., Riata, D., and Janjic, N. 1993. High affinity RNA ligands to basic fibroblast growth factor inhibit receptor binding. Proc. Natl. Acad. Sci. U.S.A. 90:11227-11231. Keene, J.D. 1996. RNA surfaces as mimetics of proteins. Chem. Biol. 3:505-513.
9.3.24 Current Protocols in Nucleic Acid Chemistry
Kramer, F.R., Mills, D.R., Cole, P.E., Nishihara, T., and Speigelman, S. 1974. Evolution of in vitro sequence and phenotype of a mutant RNA resistant to ethidium bromide. J. Mol. Biol. 89:719736. Lehman, N. and Joyce, G.F. 1993. Evolution in vitro of an RNA enzyme with altered metal dependence. Nature 361:182-185. Levisohn R. and Spiegelman, S. 1969. Further extracellular Darwinian experiments with replicating RNA molecules: Diverse variants isolated under different selective conditions. Proc. Natl. Acad. Sci. U.S.A. 63:805-811. Mills, D.R., Peterson, R.L., and Speigelman, S. 1967. An extracellular Darwinian experiment with a self-duplicating nucleic acid molecule. Proc. Natl. Acad. Sci. U.S.A. 58:217-224. Tuerk, C. and Gold, L. 1990. Systematic evolution of ligands by exponential enrichment: RNA ligands to bacteriophage T4 DNA polymerase. Science 249:505-510. Uphoff, K., Bell, S., and Ellington, A.D. 1996. In vitro selection of aptamers: The dearth of pure reason. Curr. Opin. Struct. Biol. 6:281-288.
Wlotzka, B. and McCaskill, J.S. 1997. A molecular preator and its prey: Coupled isothermal amplification of nucleic acids. Chem. Biol. 4:25-33. Wright, C. and Joyce, G.F. 1997. Continuous in vitro evolution of catalytic function. Science 276:614617.
Key References Conrad et al., 1996. See above. Conrad, R.C., Brück, F.M., Bell, S., and Ellington, A.D. 1998. In vitro selection of nucleic acid ligands. In Nucleic Acid-Protein Interactions: A Practical Approach (W.J. Christopher, ed.). Oxford University Press, New York, pp. 285-315. The above two papers also provide protocols for the selection of aptamers via filter immobilization as well as by other means.
Contributed by Sulay D. Jhaveri and Andrew D. Ellington University of Texas Austin, Texas
Combinatorial Methods in Nucleic Acid Chemistry
9.3.25 Current Protocols in Nucleic Acid Chemistry
Selection for Catalytic Function with Nucleic Acids
UNIT 9.4
Existing protocols for in vitro selection of catalytic polynucleotides are largely derived from the judicious assembly of experimental techniques routinely used in molecular biology. Each new protocol requires the use of the specific techniques that will facilitate the isolation of ribozymes or deoxyribozymes with the desired catalytic activity—that is, the protocol must be designed so as to harness this catalytic activity to help propel the selection process itself. This unit outlines several representative protocols for those who wish to repeat successful selection experiments and for those who seek a conceptual basis to aid in the design and implementation of new selective-amplification protocols for nucleic acids. STRATEGIC PLANNING Amplifying RNA and DNA Molecules Several methods for amplifying RNA and DNA molecules are commonly used in molecular biology and, with little or no modification, can be made to play a central role in the selective amplification of catalytic nucleic acids. RNA molecules can be synthesized by chemical means (Wincott et al., 1995) or prepared in large quantities by enzymatic synthesis from DNA templates using T7 RNA polymerase (Milligan and Uhlenbeck, 1989). The conversion of RNA to DNA can be achieved using reverse transcriptase, which is a component of RT-PCR protocols for RNA amplification (Rashtchian, 1994). The details of these methods are described elsewhere (see UNITS 9.2 & 9.3) and therefore are not covered in this unit. Creating Variants of Existing Ribozymes Basic Protocols 1, 2, and 4 detail the in vitro selection of existing ribozymes. Basic Protocols 1 and 2 describe the selection of group I and RNase P RNA-cleaving ribozyme variants, respectively. Basic Protocol 4 describes the propagation of RNA ligase ribozyme variants by continuous evolution. The natural “self-cleaving” ribozymes are quite small (Symons, 1992), and as a result libraries of variant ribozymes can be prepared by in vitro transcription from synthetic DNA templates that carry partially or fully randomized sequence domains (see UNIT 9.3). However, given the sizes of group I, group II, and RNase P ribozymes, to introduce mutants into larger catalysts it is necessary to use alternative methods. Fortunately, a number of excellent protocols exist for constructing starting pools of large ribozymes that allow the incorporation of the most desirable mutational patterns. The protocol most commonly used involves performing the polymerase chain reaction (PCR) under mutagenic conditions (Cadwell and Joyce, 1992; Vartanian et al., 1996). These and related DNA amplification protocols are typically used to introduce mutations over the entire length of larger ribozymes. Alternatively, mutations can be introduced in a more focused manner through the targeted integration of mutagenized oligonucleotides into ribozyme-encoding templates (Joyce and Inoue, 1989). Different combinations of fixed, mutagenized, and random-sequence domains can be joined using conventional DNA ligation methods. Alternatively, the DNA shuffling methods (Stemmer, 1994; Zhao et al., 1998) developed for the directed evolution of proteins should also be useful for the preparation of RNA populations by recombination of related ribozymes.
Contributed by Ronald R. Breaker Current Protocols in Nucleic Acid Chemistry (2000) 9.4.1-9.4.17 Copyright © 2000 by John Wiley & Sons, Inc.
Combinatorial Methods in Nucleic Acid Chemistry
9.4.1
Creating New Ribozymes from Random-Sequence Pools One of the more exciting applications of in vitro selection is the isolation of new classes of catalytic RNAs and DNAs. Usually, new catalysts are isolated from random-sequence pools, or from pools that are biased in favor of a preexisting functional domain. Basic Protocol 3 details the methods used to isolate ribozymes with polynucleotide kinase activity. In its original manifestation (Lorsch and Szostak, 1994), this selection began with an RNA pool that, in addition to random-sequence domains, carried a mutagenized domain that binds ATP. This creates a biased pool wherein greater numbers of molecules are expected to bind the substrate for the reaction. This strategy could be used for other selection efforts, although a starting pool made exclusively of random-sequence molecules may be adequate for many selection goals. BASIC PROTOCOL 1
IN VITRO SELECTION OF GROUP I RIBOZYME VARIANTS A prolific method for the directed evolution of the group I ribozyme was outlined by Joyce and Inouye (1989) and has been used both to probe the structure and function of group I ribozymes and to demonstrate various features of the in vitro selection process. This method exploits the fact that certain constructs of the group I ribozyme catalyze the reversal of the second step of RNA splicing, thereby generating a 3′-tagged ribozyme (Fig. 9.4.1). These self-modifying RNAs are selectively amplified using self-sustained sequence replication (3SR; Guatelli et al., 1990; Fahy et al., 1991) and the resulting RNAs are used to initiate the next round of selection. Although the wild-type ribozyme cleaves DNA substrates poorly (Robertson and Joyce, 1990; Herschlag and Cech, 1990), the isolation of ribozyme variants with improved DNA cleavage activity was one of the earliest achievements for catalytic RNA selections. The details of this in vitro selection protocol are given below. The following protocol has been adapted from that reported by Beaudry and Joyce (1992). For different desired outcomes, the ribozyme pool may be uniquely mutagenized and subjected to a variation of the protocol described here. It is important to note that the nucleotide sequence of the substrate RNA or DNA used with the ribozyme reaction must be made complementary to the internal guide sequence (IGS) of the ribozyme (Cech, 1990). NOTE: Buffer solutions, reagents, and oligonucleotides used in all protocols described in this unit should be stored frozen at −20°C and thawed only when needed. Protein enzymes should be stored according to the vendor’s instructions.
Selection for Catalytic Function with Nucleic Acids
Materials 2× selection buffer: 60 mM EPPS [4-(2-hydroxethyl)-1-piperazinepropanesulfonic acid; pH 7.5 at 23°C)/20 mM MgCl2 Substrate oligonucleotide solution (200 pmol/µL) 100% ethanol, −20°C TE buffer, pH 7.5 at 23°C (APPENDIX 2A) 10× 3SR buffer: 500 mM Tris⋅Cl (APPENDIX 2A; pH 7.5 at 23°C)/100 mM MgCl2/50 mM dithiothreitol (DTT) 10× 3SR NTP/dNTP mix: 20 mM each of the four ribonucleoside-5′-triphosphates and 2 mM each of the four deoxyribonucleoside-5′-triphosphates (see APPENDIX 2A for details of NTP and dNTP preparation and storage) DNA primer 1: complementary to the 3′ tail of the modified ribozymes (20 pmol/µL) DNA primer 2: homologous to the 5′ end of each ribozyme (20 pmol/µL) 50 U/µL T7 RNAP (New England Biolabs)
9.4.2 Current Protocols in Nucleic Acid Chemistry
Figure 9.4.1 An in vitro selection scheme for the directed evolution of group I ribozymes. A pool of group I ribozyme variants (each carrying a 3′-terminal guanidyl residue) are incubated with a substrate oligonucleotide. Those ribozymes that catalyze the cleavage of the substrate via a phosphoester transfer reaction acquire a fragment of the substrate at their 3′ terminus. This oligonucleotide tail acts as a primer-binding site that tags the functional ribozyme variants for amplification by self-sustained sequence replication (3SR) using two primer DNAs (1 and 2). PCR is subsequently performed using a different set of DNA primers (2 and 3) that regenerate the desired 3′ terminus upon transcription of the resulting double-stranded DNA (dsDNA) template. Although the 3SR process is a significant source of mutations, additional sequence variation can be created through the use of mutagenic PCR (Support Protocol 1). T7 RNAP and RT represent T7 RNA polymerase and reverse transcriptase, respectively. Inset: sequence of substrate, including the tag sequence that is transferred to the 3′ terminus of the ribozyme upon cleavage.
10 U/µL avian myoblastosis virus reverse transcriptase (AMV RT; Amersham) 0.3 N NaOH 3 M sodium acetate, pH 5.2 (APPENDIX 2A) 10× PCR buffer: 100 mM Tris⋅Cl (pH 8.3 at 23°C)/15 mM MgCl2/500 mM KCl/0.1% (w/v) gelatin 10× PCR dNTP mix: 2 mM each of the four deoxyribonucleoside-5′-triphosphates (see APPENDIX 2A for details of dNTP preparation and storage) DNA primer 3: complementary to the original 3′ terminus of the unmodified ribozymes (20 pmol/µL) 5 U/µL Taq DNA polymerase (Boehringer Mannheim) diluted 20:1 in distilled H2O to 0.25 U/µL final concentration Additional reagents and equipment for agarose gel electrophoresis (e.g., CPMB UNIT 2.5A) and DNA sequencing (APPENDIX 3B)
Perform the ribozyme reaction 1. Prepare mutagenized RNA pool for group I ribozyme selection, at 10 pmol/µL.
Combinatorial Methods in Nucleic Acid Chemistry
9.4.3 Current Protocols in Nucleic Acid Chemistry
Due to its large size, starting pools for group I ribozyme selection cannot be prepared easily by chemical synthesis. However, there are several methods, such as mutagenic PCR or site-directed mutagenesis, that can be used for this purpose (see discussion above). Beaudry and Joyce (1992) made use of four mutagenic oligonucleotides that were used to form a pool of DNA templates that encode group I ribozyme variants. The mutations brought about by these oligonucleotides are located within the catalytic core of the ribozyme and were introduced at these locations to screen more efficiently for variant ribozymes with altered catalytic function. This protocol for the introduction of a specific level of sequence degeneracy over defined regions in DNA templates is described in detail elsewhere (Joyce and Inoue, 1989; Beaudry and Joyce, 1992). The resulting pool of DNA templates is transcribed in vitro using T7 RNAP (UNIT 9.3; Milligan and Uhlenbeck, 1989), and purified by polyacrylamide gel electrophoresis (APPENDIX 3B; Sambrook et al., 1989). Alternatively, mutagenic PCR (Support Protocol 1, Cadwell and Joyce, 1992) or hypermutagenic PCR (Vartanian et al., 1996) methods can be used to introduce mutations throughout the entire molecule.
2. Combine 100 µL of 2× selection buffer, 70 µL distilled water, and 20 µL of 10 pmol/µL ribozyme solution. Allow mixture to equilibrate several minutes at 37°C. 3. Initiate the ribozyme reaction by the addition of 10 µL of 200 pmol/µL substrate oligonucleotide. Incubate 1 hr at 37°C. 4. Precipitate the RNA by the addition of 500 µL of 100% ethanol, −20°C. Pellet the precipitate by centrifugation for 15 min at 15,000 × g, 4°C. Resuspend the pelleted RNA in 20 µL TE buffer. During the first few rounds of selection it may be best to incubate the ribozyme reaction for extended times to allow suboptimal ribozymes greater opportunity to react. Because of the inherent mutagenic nature of the polymerase enzymes used for RNA and DNA amplification, or through the use of mutagenic PCR, the suboptimal ribozymes may acquire mutations that improve catalytic performance. In later rounds, reducing the reaction time will favor those ribozymes with faster catalytic rates. The time required to complete each round may vary as a result of alterations in the incubation times or of the intermittent use of accessory methods such as mutagenic PCR or additional purification steps. In practice, the completion of a single round of selection that includes the following amplification components requires 2 to 3 days.
Selectively amplify by 3SR 5. Combine 20 µL resuspended pool RNA with the following reagents: 35 µL distilled H2O 10 µL 10× 3SR buffer 10 µL 10× 3SR NTP/dNTP mix 5 µL 20 pmol/µL DNA primer 1 5 µL 20 pmol/µL DNA primer 2 10 µL 50 U/µL T7 RNAP 5 µL 10 U/µL AMV RT. Incubate mixture 1 hr at 37°C.
Selection for Catalytic Function with Nucleic Acids
6. Terminate reaction by the addition of 10 µL of 3 M sodium acetate, pH 5.2, and then precipitate the RNA/DNA mixture by the addition of 275 µL of 100% ethanol, −20°C. Pellet the resulting nucleic acid precipitate by centrifugation as described in step 4. 7. To destroy the remaining RNA, resuspend the resulting pellet in 200 µL of 0.3 N NaOH and heat to 90°C for 10 min.
9.4.4 Current Protocols in Nucleic Acid Chemistry
8. Precipitate the DNA by adding 20 µL of 3 M sodium acetate, pH 5.2, and 550 µL of 100% ethanol, −20°C, then pellet by centrifugation as described in step 4. Resuspend the pelleted DNA in 20 µL TE buffer. Perform nested PCR amplification of cDNA 9. Combine 20 µL cDNA with the following reagents: 48 µL distilled H2O 10 µL 10× PCR buffer 10 µL 10× PCR dNTP mix 1 µL DNA primer 2 1 µL DNA primer 3 10 µL 0.25 U/µL Taq DNA polymerase. In order to restore the original untagged terminus of the RNA pool, cDNA products of the 3SR reaction are amplified by PCR using a new DNA primer combination (primers 2 and 3). Primer 3 is complementary to the 3′-terminal bases of the ribozyme immediately preceding the tag sequence that was acquired during the preceding ribozyme reaction. This nested PCR reaction deletes the tag sequence and restores the guanosine residue required by the ribozyme for catalytic activity in the next selection cycle.
10. Perform up to 30 thermal cycles, each consisting of repetitive denaturing, annealing, and extension incubations (e.g., 30 sec at 92°C, 30 sec at 45°C, 30 sec at 72°C). Optimal temperatures and incubation times must be established for different template and primer combinations.
11. Analyze the PCR products by agarose gel electrophoresis (e.g., Sambrook et al., 1989, or CPMB UNIT 2.5A) to confirm the presence of template DNA. 12. The resulting double-stranded DNA is now ready to be transcribed (Milligan and Uhlenbeck, 1989; see also UNIT 9.3) to create the RNA pool for the next round of selection, thereby completing the selective-amplification cycle. Alternatively, the DNA can be cloned and sequenced (APPENDIX 3B) in order to examine individual RNAs that populate the selected pool. MUTAGENIC PCR Nucleic acid amplification processes such as RNA polymerization, reverse transcription, and the polymerase chain reaction unavoidably mutagenic, and therefore facilitate the accumulation of genetic diversity as in vitro selection progresses. However, in most cases it is beneficial to increase the frequency of mutations that are being introduced into the population in order to more rapidly and thoroughly explore sequence variations. The following support protocol provides a substantial increase in the error rate of DNA synthesis during the polymerase chain reaction. This protocol provides a mutation rate of approximately 0.7% per position with no strong bias for base substitutions. However, variation of the reaction conditions, particularly the concentrations of manganese and the relative concentrations of the dNTPs can significantly affect the mutagenic character of the reaction. The following protocol for mutagenic PCR is adapted from Cadwell and Joyce, 1992.
SUPPORT PROTOCOL 1
Materials DNA Primer 2: homologous to the 5′ end of each ribozyme (20 pmol/µL) DNA Primer 3: complementary to the original 3′ terminus of the unmodified ribozymes (20 pmol/µL) DNA template: 5 pmole/µL
Combinatorial Methods in Nucleic Acid Chemistry
9.4.5 Current Protocols in Nucleic Acid Chemistry
10× mutagenic PCR buffer: 100 mM Tris•Cl (pH 8.3 at 23°C)/70 mM MgCl2/500 mM KCl/0.1% (w/v) gelatin 10× mutagenic PCR dNTP mix: 2 mM each of dGTP and dATP/10 mM each of dCTP and dTTP 5 mM MnCl2 5 U/µL Taq DNA polymerase (Boehringer Mannheim) diluted 20:1 in dH2O to 0.25 U/µL final concentration 1. Combine 4 µL DNA template with the following reagents: 54 µL distilled H2O 10 µL 10× mutagenic PCR buffer 10 µL 10× mutagenic PCR dNTP mix 1 µL DNA Primer 2 1 µL DNA Primer 3 10 µL 5 mM MnCl2 10 µL 0.25 U/µL Taq DNA polymerase. To prevent precipitation of reagents, the MnCl2 should be added just prior to the addition of Taq DNA polymerase and initiation of the reaction.
2. Perform up to 30 thermal cycles as described in step 10 of Basic Protocol 1. Analyze the products and transcribe RNA as described in steps 11 and 12 of Basic Protocol 1. BASIC PROTOCOL 2
Selection for Catalytic Function with Nucleic Acids
IN VITRO SELECTION OF RNase P RIBOZYME VARIANTS Several groups have reported the use of RNase P for the in vitro selection of new or improved substrates (Yuan and Altman, 1994; Liu and Altman, 1994; Pan, 1995) or for the selection of variant ribozymes themselves (Frank et al., 1996; Frank and Pace, 1997). In the latter examples, a self-cleaving RNA system consisting of a ribozyme/substrate construct was generated to facilitate the separation of active from inactive ribozyme variants. Since RNase P catalyzes the hydrolysis of RNA substrates, in vitro selection schemes for this ribozyme cannot take advantage of a self-ligation reaction. In contrast, mutagenized populations of this construct, termed TP292∆, are immobilized on a thiophilic column via a phosphorothioate nucleotide attached to each RNA molecule (Fig. 9.4.2). Separation of active TP292∆ variants is achieved upon RNA self-cleavage, which liberates ribozymes from the matrix—a process that has been termed “catalytic elution” (Breaker, 1997a). As a result of the specific methods used for this selection, specific changes must be made to the conventional protocols for RNA preparation and manipulation; these alterations are highlighted in the protocol given here. This procedure, adapted from Frank et al. (1996), can be broadly applied for the study of RNase P structure and function using in vitro selection. In vitro transcription must be carried out under specialized conditions in order to include the 5′-thiophosphate moiety and to minimize the fraction of precursor RNAs that undergo self cleavage prior to the selection phase of the process. The efficiency of RNA transcription by T7 RNAP is substantially improved for templates that encode for the presence of a 5′-terminal guanosine nucleotide in the RNA transcript. As a result, different chemical groups can be incorporated into RNA during the transcription process if modified guanosine nucleotides are included in the reaction. This fact is exploited here to create pool RNAs that begin with a 5′-phosphorothioate residue. The in vitro transcription protocol is an adaptation of that described by Milligan and Uhlenbeck (1989) for the enzymatic synthesis of RNA.
9.4.6 Current Protocols in Nucleic Acid Chemistry
Figure 9.4.2 A selective-amplification strategy for the isolation of RNase P ribozyme variants. A pool of double-stranded DNA templates that encode for TP292∆ variants is transcribed in vitro using T7 RNA polymerase (T7 RNAP) in the presence of 5′-guanosine monophosphorothioate (GMPS). The resulting RNA pool is immobilized on a thiophilic beaded-agarose matrix. Competent variants of the ribozyme-substrate fusion are recovered from the matrix under permissive reaction conditions (added Mg2+) and amplified by RT-PCR. The cycle is repeated until RNAs with the desired activity dominate the amplified population. (−) and (+) represent the T7 RNAP promoter sequences that reside on the nontemplate and template strands, respectively. Shaded box represents the agarose matrix. Arrowhead identifies the site of ribozyme cleavage.
Materials 10× transcription buffer (see recipe) 10× transcription NTP mix: 10 mM each of the four ribonucleoside-5′-triphosphates 50% (v/v) aqueous glycerol Double-stranded template DNA (at 10 pmol/µL) 100 mM guanosine-5′-phosphorothioate (GMPS; Amersham Life Sciences–NucleixPlus) 10 mCi/mL [α-32P]GTP (3000 Ci/mmol) 50 U/µL T7 RNAP (New England Biolabs) 0.5 M EDTA, pH 8.0 (APPENDIX 2A) 100% ethanol, −20°C 2× urea loading buffer (APPENDIX 2A) Gel elution buffer (see recipe) TE buffer, pH 7.5 (APPENDIX 2A) Sulfolink gel (aqueous slurry of beaded agarose; Pierce) Loading buffer: 40% methanol/20 mM sodium phosphate (pH 8.9)/0.1% SDS (APPENDIX 2A)/5 mM EDTA Nonpool RNAs: e.g., Bacillus subtilis rRNA Wash buffer: 3 M NaCl/50 mM Tris⋅Cl (pH 8; APPENDIX 2A)/5 mM EDTA Reaction buffer −Mg2+ (see recipe) 1 M MgCl2 Glycogen Sterile razor blade Liquid scintillation counter
Combinatorial Methods in Nucleic Acid Chemistry
9.4.7 Current Protocols in Nucleic Acid Chemistry
Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (PAGE; e.g., APPENDIX 3B or CPMB UNIT 7.6) and RT-PCR (UNIT 9.3) Perform in vitro transcription 1. Combine in a 1.5-mL microcentrifuge tube (100 µl total volume): 30.5 µL distilled H2O 10 µL 10× transcription buffer 10 µL 10× transcription NTP mix 15 µL 50% aqueous glycerol 2 µL 10 pmol/µL template DNA 7.5 µL 100 mM GMPS 5 µL 10 mCi/mL [α-32P]GTP 20 µL 50 U/µL T7 RNAP. Incubate mixture 12 hr at 12°C. 2. Terminate reaction by the addition of 3 µL of 0.5 M EDTA, pH 8.0. 3. Recover the resulting RNA products by precipitation with 250 µL of 100% ethanol at −20°C, centrifuge as described in step 4 of Basic Protocol 1, and resuspend the pelleted RNA in 40 µL of 2× urea loading buffer. Preparation of the RNA pool differs in two main ways from the method described previously (Milligan and Uhlenbeck, 1989). First, an excess (7.5:1 ratio) of GMPS to GTP is used to favor the incorporation of the thiophosphate-containing nucleotide at the start of most transcripts. Second, in vitro transcription is conducted under reduced temperature in order to minimize cleavage (and loss) of functional ribozymes during preparation. The RNase P ribozyme, like most catalytic nucleic acids, requires Mg2+ to display catalytic function. As a result, it is important to terminate the transcription reaction with EDTA to preclude continuing self cleavage of the RNA products.
Gel purify RNA transcription products 4. Separate the transcription products on a 5% denaturing polyacrylamide gel (e.g., APPENDIX 3B or CPMB UNIT 7.6). 5. Visualize the RNA precursor band by UV shadowing or autoradiography, and excise using a sterile razor blade. 6. Recover the isolated RNAs by crush/soaking the gel overnight at 4°C in gel elution buffer. 7. Precipitate recovered RNAs by the addition of 2.5 vol of 100% ethanol, −20°C, and pellet by centrifugation as described in step 4 of Basic Protocol 1. 8. Resuspend pelleted RNA in TE buffer, quantitate by liquid scintillation counting, and store at −20°C. Perform ribozyme selection and amplification 9. Place 150 µL of Sulfolink gel slurry in a 1.5-mL microcentrifuge tube, add 300 µL loading buffer, and swirl to wash the beads. Pellet beads by brief centrifugation. Repeat for total of three washes. Selection for Catalytic Function with Nucleic Acids
10. Cover the beads with 300 µL loading buffer containing 16 µg Bacillus subtilis rRNA (or other nonpool RNA) and let stand 45 min at 25°C. This prebinding serves to occupy nonspecific RNA binding sites on the agarose matrix.
9.4.8 Current Protocols in Nucleic Acid Chemistry
11. To refold RNAs, preincubate 100 pmol 5′-thiophosphate-terminated RNA molecules (from step 8) in 300 µL loading buffer for 3 min at 65°C, then for 1 min at 50°C. 12. Combine washed gel beads and preincubated RNA solution, and incubate 1 to 2 hr at room temperature. 13. Remove unbound RNAs from the matrix with four successive washes each using 300 µL wash buffer as described in step 9, followed by four additional washes each using 300 µL reaction buffer −Mg2+. 14. Resuspend the final washed RNA/bead slurry in 225 µL reaction buffer −Mg2+, giving a total volume of ∼370 µl. Allow the mixture to equilibrate at 50°C for 1 min. Initiate reactions by the addition of 9 µL of 1 M MgCl2 (giving a final concentration of ∼25 mM cofactor). 15. After the desired incubation time, terminate reaction by the addition of 30 µL EDTA. 16. Add 5 µg glycogen, then precipitate with ethanol as described in step 4 of Basic Protocol 1 to recover the ribozymes. The glycogen aids in precipitating even small amounts of RNA.
17. Resuspend the resulting pellet in 25 µL TE buffer. 18. Use the RNAs as a substrate for RT-PCR using appropriate DNA oligomers (to convert the RNAs into cDNA and amplify them; see UNIT 9.3 for RT-PCR procedure). The entire protocol, beginning with in vitro transcription, is repeated in an iterative fashion until satisfactory catalytic activity is observed with the RNA pool. Molecules present in the final pool are cloned and sequenced to facilitate further analysis of the resulting ribozyme variants.
IN VITRO SELECTION OF NEW KINASE RIBOZYMES FROM RANDOM SEQUENCE
BASIC PROTOCOL 3
Due to the enormous numbers of oligonucleotides that can be screened simultaneously by in vitro selection, it is practical to isolate entirely new and complex ribozymes from random-sequence pools. This was first demonstrated by Bartel and Szostak (1993), who isolated a series of “ligase” ribozymes from a large pool of random-sequence RNAs. Lorsch and Szostak (1994) expanded on this approach by creating a biased pool that carried both random-sequence and preexisting functional RNA domains in their successful search for “kinase” ribozymes (Fig. 9.4.3). The selection protocol for the latter study is described below. Materials 2× kinase selection buffer (see recipe) 100 mM adenosine 5′-O-(3-thiotriphosphate) (ATP-γS; Sigma) 100% ethanol, −20°C Binding buffer: 1 mM EDTA/25 mM HEPES (pH 7.4 at 23°C) Thiopyridine-activated thiopropyl Sepharose 6B (Amersham Pharmacia Biotech) Wash buffer: 1 M NaCl/5 mM EDTA/25 mM HEPES (pH 7.4 at 23°C) Urea solution: 3 M urea/5 mM EDTA 0.1 M 2-mercaptoethanol 3 M sodium acetate, pH 5.2 (APPENDIX 2A) Chromatographic column
Combinatorial Methods in Nucleic Acid Chemistry
9.4.9 Current Protocols in Nucleic Acid Chemistry
Selection for Catalytic Function with Nucleic Acids
Figure 9.4.3 Scheme for the isolation of self-phosphorylating ribozymes from a biased RNA pool that includes random-sequence domains. The pool is composed of three regions of random nucleotide sequence that are intermixed with a mutagenized (d = 0.15; Breaker and Joyce, 1994a) ATP-binding domain. Pool RNAs must be dephosphorylated each round to remove the 5′-triphosphate moiety that results from preparative in vitro transcription. RNAs that acquire the thiophosphate moiety of ATP-γS in the selection reaction (inset) are selectively captured by a thiophilic resin. RNAs bound by the resulting disulfide bond are specifically eluted with 2-mercaptoethanol and are amplified by RT-PCR. The variable d represents the nucleotide degeneracy at each of the specified positions in the polynucleotide chain (e.g. d = 0.15 reflects a 0.85 probability of finding the wild-type nucleotide and a 0.15 probability of finding one of the three remaining nucleotides at a given position).
9.4.10 Current Protocols in Nucleic Acid Chemistry
1. Prepare mutagenized RNA pool for kinase ribozyme selection, at 20 pmol/µL. In the original study reported by Lorsch and Szostak (1994), a mutagenized ATP-binding domain was integrated with random-sequence domains to create the RNA pool. Briefly, two synthetic DNAs that correspond to the two halves of the final RNA pool (Fig. 9.4.3) were made double stranded by PCR, treated with the restriction enzyme BanI, and joined using T4 DNA ligase. The ligated pool of DNA templates was transcribed in vitro using T7 RNAP, and the resulting pool of RNA transcripts was purified by polyacrylamide gel electrophoresis. At this stage, each RNA molecule carried a 5′-terminal triphosphate moiety that would block autophosphorylation at this position. Therefore, the RNA pool was treated exhaustively with alkaline phosphatase prior to the selection reaction in order to generate RNA precursors each carrying a hydroxyl group at their 5′ terminus. Alternatively, a pool composed exclusively of random-sequence RNAs can be prepared (see UNIT 9.2).
2. Combine 500 µL of 2× selection buffer, 440 µL distilled water, and 50 µL of 20 pmol/µL pool RNA solution. Allow mixture to equilibrate at 23°C for several minutes. 3. Initiate the ribozyme reaction by the addition of 10 µL ATP-γS and incubate 20 hr at 23°C. 4. Precipitate the RNA by the addition of 2.5 mL of 100% ethanol, −20°C, followed by centrifugation as described in step 4 of Basic Protocol 1. Resuspend the pelleted RNA in 500 µL binding buffer. 5. Combine the RNA solution with an equal volume of a slurry of the activated thiopropyl Sepharose and incubate 30 min at 23°C. 6. Prepare a column (e.g. 0.8 × 4 cm poly-prep., Bio-Rad) with the RNA/Sepharose mixture. Remove unbound RNAs from the matrix by washing with 20 bed volumes each of wash buffer, distilled H2O, and urea solution. 7. Elute disulfide-linked RNAs with 1 column volume 0.1 M 2-mercaptoethanol. 8. Add 3 M sodium acetate, pH 5.2, to the recovered RNA solution to give a final concentration of 0.3 M. Precipitate the RNA by the addition of 2.5 vol of 100% ethanol, −20°C, followed by centrifugation as described in step 4 of Basic Protocol 1. As stated in Basic Protocol 1, during the first few rounds of selection it may be best to incubate the ribozyme reaction for extended times to allow suboptimal ribozymes greater opportunity to react. In later rounds, reducing the reaction time will favor those ribozymes with faster catalytic rates. Other reaction conditions, such as concentrations of ATP-γS or divalent metal ions, can be adjusted to favor ribozymes that have greater affinities for these agents.
CONTINUOUS EVOLUTION OF LIGASE RIBOZYMES A new method for the continuous evolution of ribozymes has proven to be a powerful means to isolate ligase ribozyme variants with modified catalytic activity. Continuous evolution is a dynamic process whereby ribozymes that best serve as catalysts and as templates for cDNA synthesis and RNA transcription dominate the population of amplifying RNAs. Therefore, any variations in the catalysis and amplification cycle can be made so long as the protein enzymes that facilitate RNA and DNA synthesis are not rendered inactive. Outlined below is a protocol used by Wright and Joyce (1997) to propagate variants of RNA ligase ribozymes.
BASIC PROTOCOL 4
Combinatorial Methods in Nucleic Acid Chemistry
9.4.11 Current Protocols in Nucleic Acid Chemistry
Materials 10× reaction buffer (see recipe) 10× 3SR NTP/dNTP mix: 20 mM each of the four ribonucleoside-5′-triphosphates and 2 mM each of the four deoxyribonucleoside-5′-triphosphates (see APPENDIX 2A for NTP and dNTP preparation details) DNA primer 1: complementary to the 3′ tail of the pool RNAs (20 pmol/µL) Substrate oligonucleotide: DNA/RNA chimera that encodes the T7 promoter sequence (20 pmol/µL) 40 U/µL T7 RNAP (New England Biolabs) 200 U/µL Moloney murine leukemia virus reverse transcriptase (MMLV RT; Amersham) 1. Prepare mutagenized RNA pool for continuous evolution. Initial pools for the first round of continuous evolution can be prepared by in vitro transcription from synthetic DNAs (mutagenized during chemical synthesis) or from DNA templates that resulted from RT-PCR amplification of the selected ribozymes (see UNIT 9.3). A primary concern when initiating the continuous evolution experiment is the catalytic speed of the starting ribozyme. The prototype ribozyme used by Wright and Joyce (1997) to generate the starting RNA pool for their RNA ligase selection experiment was an inefficient catalyst that could not compete successfully with selfish RNAs. As a result, the authors subjected a pool of 1014 ribozyme variants to several conventional rounds of in vitro selection followed by an intermediate form of continuous evolution to enrich the RNA population with highly active ribozymes. This more efficient population of ribozymes subsequently was used to initiate the continuous evolution process. Similarly, individual ribozymes that meet this activity threshold must be present in the starting RNA population in order to initiate the continuous evolution process.
2. Combine in a 1.5-mL microcentrifuge tube (20 µL total volume): 5.5 µL distilled H2O 1 µL pool RNA 2 µL 10× reaction buffer 2 µL 10× 3SR NTP/dNTP mix 2.5 µL 20 pmol/µL DNA primer 1 5 µL 20 pmol/µL substrate oligonucleotide 1 µL 40 U/µL T7 RNAP 1 µL 200 U/µL MMLV RT. Incubate 1 hr at 37°C. 3. Dilute an aliquot of the reaction mixture 50-fold with distilled H2O. Either proceed with the next serial transfer or store the diluted sample at −20°C for future use.
Selection for Catalytic Function with Nucleic Acids
Iterations involving serial dilution of the pool followed by repetition of the reaction can be carried out indefinitely. This continuous evolution format favors catalytic RNAs that perform the ligation reaction most rapidly. As the selective-amplification cycles proceed and the ribozymes become more efficient, stages of the cycle other than the catalytic step may become limiting to the overall process. Propagation of ribozyme variants that overcome whatever is the rate-limiting (catalytic or amplification) step will dominate the population of amplifying RNAs. This feature of continuous evolution could be exploited to examine the mechanisms and kinetics of nucleic acid amplification in addition to catalysis.
9.4.12 Current Protocols in Nucleic Acid Chemistry
REAGENTS AND SOLUTIONS Use distilled, deionized water or other ultrapure water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Gel elution buffer 0.3 M sodium acetate 10 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) 1 mM EDTA (APPENDIX 2A) 0.5% (w/w) sodium dodecyl sulfate (SDS; APPENDIX 2A) Autoclave and store indefinitely at room temperature Reaction buffer for continuous evolution, 10× 500 mM EPPS [4-(2-hydroxethyl)-1-piperazinepropanesulfonic acid], pH 8.5 at 23°C 500 mM KCl 250 mM MgCl2 40 mM dithiothreitol (DTT) 20 mM spermidine Store indefinitely at –20°C Reaction buffer −Mg2+ (for ribozyme selection) 3 M sodium acetate 44.5 mM Tris⋅Cl (APPENDIX 2A) 16 mM piperazine-N,N’-bis(2-hydroxypropanesulfonic acid) (PIPES) 0.05% SDS (APPENDIX 2A) Adjust pH to 8 with NaOH Store indefinitely at –20°C Selection buffer, 2× 50 mM HEPES (N-2-hydroxyethylpiperazine-N’-2-ethanesulfonic acid), pH 7.4 at 23°C 100 mM MgCl2 10 mM MnCl2 800 mM KCl Store indefinitely at –20°C Transcription buffer for in vitro selection, 10× 400 mM Tris⋅Cl (APPENDIX 2A), pH 8.1 at 37°C 60 mM MgCl2 50 mM DTT 10 mM spermidine 0.1 % (v/v) Triton X-100 50 µg/mL bovine serum albumin (BSA) Store indefinitely at –20°C COMMENTARY Background Information Catalytic nucleic acids and in vitro selection An essential characteristic of any class of polymers that is capable of supporting diverse catalytic activity is the ability to form a nearly endless array of higher-ordered folding patterns. This structural sophistication allows the polymer to form precise active sites for specific
binding of substrates and for catalysis of chemical transformations. Beginning with the determination of the three-dimensional structure of tRNAPhe (Kim et al., 1973; Robertus et al., 1974), it has become increasingly clear over the last two decades that RNA has enormous potential for forming unique secondary and tertiary structures (Gold et al., 1995; Osborne and Ellington, 1997). Evidence that this structural
Combinatorial Methods in Nucleic Acid Chemistry
9.4.13 Current Protocols in Nucleic Acid Chemistry
Selection for Catalytic Function with Nucleic Acids
complexity could be exploited to form catalytic RNAs was first obtained upon the discovery of natural ribozymes that self splice (Kruger et al., 1982) and that function as RNA endonucleases (Guerrier-Takada et al., 1983). The collection of natural and artificial ribozymes has expanded rapidly in recent years to include a wide range of catalytic activities (Breaker, 1997a; Jaeger, 1997), providing strong support for the view that the structure-forming potential of RNA is more than adequate to allow the formation of a great diversity of ribozymes that carry precisely formed active sites. The many examples of catalytic RNAs also provide compelling evidence for the notion that an “RNA world” may have preceded the protein-dominated metabolic processes that are in operation today (White, 1976; Gilbert, 1986; Cech, 1993; Benner et al., 1989). Although always a topic of debate, this theory has inspired many who study nucleic acids to aggressively seek new ribozymes that catalyze reactions believed to be central to the origin and evolutionary progression of life. In addition, researchers continue to probe the limits of biocatalysts both under cellular conditions and when the biochemical rules and reaction conditions are no longer set by living organisms. Even DNA can be forced to function as an enzyme outside the confines of living cells (Breaker, 1997b), suggesting that the perceived biological limitations for catalytic nucleic acids may be overcome simply by using the latest strategies for the design of novel nucleic acids. Almost without exception, the search for new ribozymes and deoxyribozymes makes use of in vitro selection methods, which have proven to be the most effective approach for producing new or altered RNA catalysts (Breaker, 1997a). In vitro selection relies on the probability that only one or perhaps a few individuals amongst trillions of random-sequence molecules can promote the chemical reaction of interest. This process makes use of selection and amplification procedures that are performed repetitively to recover these rare catalysts from the pool of inactive molecules. Critical to the success of an in vitro selection experiment is the judicious design of the selection and amplification protocol. As with natural selection, these “test-tube evolution” experiments are unforgiving and can give rise to spurious products unless great care and forethought are given to the design of the experimental methodology (see UNIT 9.3 for further discussion on “molecular parasites”). Unfortunately, methods for the selection of catalytic
RNAs and DNAs must be revised for each new catalytic activity sought, each time forcing the careful reevaluation of the protocols to be employed. Given the expected functional diversity of RNA and DNA, a wide range of chemical and molecular biological techniques will be needed to successfully and efficiently expand the catalytic repertoire of nucleic acids. The protocols described herein are representative of the many existing methods that have been used successfully to isolate ribozymes with new or improved catalytic function. Future efforts in this field no doubt will expand the catalytic repertoire of RNA and DNA, probe the limits of function with existing catalysts, and create superior RNA and DNA catalysts that have utility beyond basic science. Whether the experimenter’s specific goal is to identify new classes of catalytic nucleic acids, or to focus the selection pressure precisely on a individual kinetic parameter of an existing enzyme, new or modified selection strategies and methods most likely will need to be devised. Early selection experiments with group I ribozymes One of the first ribozymes to be subjected to in vitro selection is the group I self-splicing ribozyme. This ribozyme forms an intron of the 26S ribosomal RNA precursor from the ciliated protozoan Tetrahymena thermophila, which is known to self process (Kruger et al., 1982). The Tetrahymena ribozyme has been used as a starting point for a number of pioneering in vitro selection experiments that tested the methods, possible strategies, and the limitations of different selective-amplification processes. Although this ribozyme normally promotes two successive RNA phosphoester transfer reactions that produce spliced exons, reorganized forms of the enzyme have been made to catalyze several related reactions including RNA cleavage, oligonucleotide polymerization/depolymerization, and dephosphorylation reactions (Cech, 1990). More recently, in vitro selection has been used to isolate variants of group I ribozymes that have new functions such as DNA cleavage (Beaudry and Joyce, 1992; Tsang and Joyce, 1996) and altered metal-ion dependence (Lehman and Joyce, 1993), or that have new or altered structural domains (e.g., Green et al., 1990; Williams et al., 1994; Costa and Michel, 1997). Similar investigations are now being conducted with other large ribozymes.
9.4.14 Current Protocols in Nucleic Acid Chemistry
Continuous evolution of ribozymes Most in vitro selection experiments proceed in a punctuated fashion, meaning that the experimenter must actively participate in the selection process through manual intervention or automated processing. For example, the selection and amplification steps of most simulated evolutionary processes each require time-consuming manipulations such as gel purification, column chromatography, gel elution, nucleic acid precipitation, or separate enzyme reactions for the amplification of selected products. Without this intervention the selection process would cease and only a portion of a given round of selection would be completed. Depending on the complexity of the selection process, this demand for manipulation adds days or weeks to the time that the catalytic molecules actually are performing as catalysts, and these manipulations can become annoyingly repetitive. For one class of reactions at least, this process has been simplified by employing a selective-amplification method that almost completely eliminates the need for experimenter intervention. “Continuous evolution” of catalytic RNA function has been demonstrated (Wright and Joyce, 1997) using a variant of the ligase ribozymes isolated by Bartel and Szostak (1993). This scheme (Fig. 9.4.4) was designed to couple the catalysis and amplification processes in order to create a self-sustaining system whereby the fastest ribozymes become available for selective amplification sooner than their molecular competitors (Breaker and Joyce, 1994b). The resulting amplification process creates molecular progeny molecules that again need to promote the ligation reaction in order to replicate. Infinite cycles of coupled selection and amplification can proceed in a continuous fashion using a serial transfer procedure, whereby a portion of the preceding reaction is used to initiate a fresh reaction mixture. Coupled catalysis and amplification processes, however, are prone to the generation of selfish nucleic acids that replicate without the need for catalysis (Breaker and Joyce, 1994; Breaker et al., 1994). Therefore great care must be taken to prevent these molecules from overtaking the RNA population.
Critical Parameters There are an almost overwhelming number of issues that must be considered when initiating an in vitro selection project. Unfortunately, how many of these critical parameters should be addressed will change depending on the goals of the project. Of foremost importance is
Figure 9.4.4 Scheme for the continuous in vitro evolution of ligase ribozymes. A mutagenized population of self-ligating ribozymes is incubated in the presence of reverse transcriptase (RT), RNA polymerase (RNAP), and oligonucleotide substrate. Functional ribozymes in this mixture undergo a sequence of ribozyme- and protein enzyme–mediated reactions in order to replicate. RNAs that catalyze the formation of a new 3′-5′ linkage to a substrate oligonucleotide (at the expense of the 5′-terminal triphosphate moiety) acquire a promoter sequence that is specific for the constituent RNA pol ymerase. Upon reverse transcription, the chimeric RNA-DNA hybrids formed from active ribozymes now carry a double-stranded promoter element. RNA polymerase initiates trans cr iption for m this double-stranded sequence, producing new RNA enzymes each with a regenerated 5′triphosphate terminus. These new RNAs, some of which carry new mutations as a result of the inherent infidelity of the polymerizing enzymes, can participate in the next round of selective amplification.
the catalytic activity sought by the researcher. For example, one may want to have some confidence that molecules within the pool can catalyze the target reaction before committing time and resources to an in vitro selection effort. In many cases, in vitro selection is limited to isolating catalysts with a rate constant of at least 10−5 min−1—which equates to a half life for the precursor of ∼5 days for self-modifying enzymes. To effectively isolate enzymes with poorer rate constants, even longer incubation times for the selection would be necessary to allow a significant number of enzymes to react. In addition, most catalysts isolated from random-sequence pools catalyze the target reaction with rates that are <1-million-fold above the equivalent uncatalyzed reaction, although in many instances additional rounds of mutation and selection do result in greater rate en-
Combinatorial Methods in Nucleic Acid Chemistry
9.4.15 Current Protocols in Nucleic Acid Chemistry
hancements. Therefore, targeting a reaction that has a rate constant for the uncatalyzed reaction of 10−11 or lower is expected to be very challenging. In general, maximizing both the pool diversity and the size of the mutagenized domains will increase the probability that the catalysts of interest will be represented. However, these parameters must be balanced with practical considerations such as the ease and expense of pool construction and the technical challenges posed by the amplification and manipulation of larger oligonucleotide constructs. The introduction of mutations into preexisting ribozymes is also of great importance. Too small a number of mutations will not allow sampling of ribozyme variants that have the catalytic parameters of interest, while too large a number of mutations may create variant that are so distant from the original ribozyme that entirely new catalytic motifs with the same catalytic function will be isolated. Extreme care must be exercised during all stages of the selection protocol, as most technical problems can result in a reduction of the sequence diversity comprising the pool. This is particularly important during the early stages of any selection project. In many cases, a catalytic “signal” from the population that could be used to confirm satisfactory progress may not appear until several rounds of selection have been completed. Since many in vitro selection protocols are assembled using routine molecular biology techniques, precautions used to ensure success with these techniques should be exercised faithfully throughout the entire selection process.
Anticipated Results
Selection for Catalytic Function with Nucleic Acids
If successful, a typical in vitro selection protocol yields a population of functional molecules within three to ten rounds of selective amplification. The exact performance of each selection experiment will depend upon a number of factors, such as the frequency of active molecules in the original pool and the stringency of the selection process. However, it must be noted that ∼50% of all reported ribozyme-selection experiments to date have yielded molecules that survive the selection protocol by some mechanism other than that originally intended by the investigators. These unexpected “answers” range from simple “molecular parasites” that do not perform chemical reactions, to catalysts that promote reactions that are distinct from the desired reaction. If these undesirable molecules emerge during in
vitro selection, each must be assessed and dealt with on an individual basis given the innumerable possible mechanisms by which they may survive.
Time Considerations The time required to successfully complete an in vitro selection experiment will vary widely depending on the complexity of the selection protocol and on the challenge of the reaction that is presented to the pool of molecules. Most of the standard molecular biology techniques typically used when assembling selection protocols can be completed within several hours. However, more complicated in vitro selection protocols may be comprised of a dozen or more techniques for each round that together may require several days to complete. For most in vitro selection efforts, the researcher should be prepared to devote several months to the selection phase of the study.
Literature Cited Bartel, D.P. and Szostak, J.W. 1993. Isolation of new ribozymes from a large pool of random sequences. Science 261:1411-1418. Beaudry, A.A. and Joyce, G.F. 1992. Directed evolution of an RNA enzyme. Science 257:635-641. Benner, S.A., Ellington, A.D., and Tauer, A. 1989. Modern metabolism as a palimpsest of the RNA world. Proc. Natl. Acad. Sci. U.S.A. 86:70547058. Breaker, R.R. and Joyce, G.F. 1994a. Inventing and improving ribozyme function: rational design versus iterative selection methods. Trends Biotechnol. 12:268-275. Breaker, R.R and Joyce, G.F. 1994b. Emergence of a replicating species from an in vitro RNA evolution reaction. Proc. Natl. Acad. Sci. U.S.A. 91:6093-6097. Breaker, R.R., Banerji, A., and Joyce, G.F. 1996. Continuous in vitro evolution of bacteriophage RNA polymerase promoters. Biochemistry 33:11980-11986. Breaker, R.R. 1997a. In vitro selection of catalytic polynucleotides. Chem. Rev. 97:371-390. Breaker, R.R. 1997b. DNA aptamers and DNA enzymes. Curr. Opin. Chem. Biol. 1:26-31. Cadwell, R.C. and Joyce, G.F. 1992. Randomization of genes by PCR mutagenesis. PCR Methods Appl. 2:28-33. Cech, T.R. 1990. Self-splicing of group I introns. Annu. Rev. Biochem. 59:543-568. Cech, T.R. 1993. The efficiency and versatility of catalytic RNA: Implications for an RNA world. Gene 135:33-36.
9.4.16 Current Protocols in Nucleic Acid Chemistry
Costa, M. and Michel, F. 1997. Rules for RNA recognition of GNRA tetraloops deduced by in vitro selection: Comparison with in vivo evolution. EMBO J. 16:3289-3302. Fahy, E., Kwoh, D.Y., and Gingeras, T.R. 1991. Self-sustained sequence replication (3SR): An isothermal transcription-based amplification system alternative to PCR. PCR Methods Appl. 1:25-33. Frank, D.N. and Pace, N.R. 1997. In vitro selection for altered divalent metal specificity in the RNase P RNA. Proc. Natl. Acad. Sci. U.S.A. 94:14355-14360. Frank, D.N., Ellington, A.E., and Pace, N.R. 1996. In vitro selection of RNase P RNA reveals optimized catalytic activity in a highly conserved structural domain. RNA 2:1179-1188. Gilbert, W. 1986. The RNA world. Nature 319:618. Gold, L., Polisky, B., Uhlenbeck, O., and Yarus, M. 1995. Diversity of oligonucleotide functions. Annu. Rev. Biochem. 64:736-797. Green, R., Ellington, A.D., and Szostak, J.W. 1990. In vitro genetic analysis of the Tetrahymena selfsplicing intron. Nature 347:406-408. Guatelli, J.C., Whitfield, K.M., Kwoh, D.Y., Barringer, K.J., Richman, D.D., and Gingeras, T. R. 1990. Isothermal, in vitro amplification of nucleic acids by a multienzyme reaction modeled after retroviral replication. Proc. Natl. Acad. Sci. U.S.A. 87:1874-1878.
Milligan, J.F. and Uhlenbeck, O.C. 1989. Synthesis of small RNAs using T7 RNA polymerase. Methods Enzymol. 180:51-62. Osborne, S.E. and Ellington, A.D. 1997. Nucleic acid selection and the challenge of combinatorial chemistry. Chem. Rev. 97:349-370. Pan, T. 1995. Novel RNA substrates for the ribozyme from Bacillus subtilis ribonuclease P identified by in vitro selection. Biochemistry 34:8458-8464. Rashtchian, A. 1994. Amplification of RNA. PCR Meth. Appl. S83-S91. Robertson, D.L. and Joyce, G.F. 1990. Selection in vitro of an RNA enzyme that specifically cleaves single-stranded DNA. Nature 344:467-468. Robertus, J.D., Ladner, J.E., Finch, J.T., Rhodes, D., Brown, R.S., Clark, B.F., and Klug, A. 1974. Structure of yeast phenylalanine tRNA at 3 Å resolution. Nature 250:546-551. Sambrook, J., Fritsch, E.F. and Maniatis, T. (eds.) 1989. Molecular Cloning, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Stemmer, W.P.C 1994. Rapid evolution of a protein in vitro by DNA shuffling. Nature 370:389-391. Symons, R.H. 1992. Small catalytic RNAs. Annu. Rev. Biochem. 61:641-671. Tsang, J. and Joyce, G.F. 1996. Specialization of the DNA-cleaving activity of a group I ribozyme through in vitro evolution. J. Mol. Biol. 262:31-42.
Guerrier-Takada, C., Gardiner, K., Marsh, T., Pace, N., and Altman, S. 1983. The RNA moiety of ribonuclease P is the catalytic subunit of the enzyme. Cell 35:849-857.
Vartanian, J.-P., Henry, M., and Wain-Hobson, S. 1996. Hypermutagenic PCR involving all four transitions and a sizeable proportion of transversions. Nucleic Acids Res. 14:2627-2631.
Herschlag, D. and Cech, T.R. 1990. DNA-cleavage catalysed by the ribozyme from Tetrahymena. Nature 344:405-409.
White, H.B. III 1976. Coenzymes as fossils of an earlier metabolic state. J. Mol. Evol. 7:101-104.
Jaeger, L. 1997. The new world of ribozymes. Curr. Opin. Struct. Biol. 7:324-335. Joyce, G.F. and Inouye, T. 1989. A novel technique for the rapid preparation of mutant RNAs. Nucleic Acids Res. 17:711-722 Kim, S.H., Quigley, G.J., Suddath, F.L., McPherson, A., Sneden, D., Kim, J.J., Weinzierl, J., and Rich, A. 1973. Three-dimensional structure of yeast phenylalanine transfer RNA: Folding of the polynucleotide chain. Science 179:285-288. Kruger, K., Grabowski, P.J., Zaug, A.J., Sands, J., Gottschling, D.E., and Cech, T.R. 1982. Selfsplicing RNA: Autoexcision and autocyclization of the ribosomal RNA intervening sequence of Tetrahymena. Cell 31:147-157. Lehman, N. and Joyce, G.F. 1993. Evolution in vitro of an RNA enzyme with altered metal ion dependence. Nature 361:182-185. Liu, F. and Altman, S. 1994. Differential evolution of substrates for an RNA enzyme in the presence and absence of its protein cofactor. Cell 77:1093-1100. Lorsch, J.R. and Szostak, J.W. 1994. In vitro evolution of new ribozymes with polynucleotide kinase activity. Nature 371:31-36.
Williams, K.P., Imahori, H., Fujimoto, D.N., and Inoue, T. 1994. Selection of novel forms of a functional domain within the Tetrahymena ribozyme. Nucleic Acids Res. 22:2003-2009. Wincott, F., DiRenzo, A., Shaffer, C., Grimm, S., Tracz, D., Workman, C., Sweedler, D., Gonzalez, C., Scaringe, S., and Usman, N. 1995. Synthesis, deprotection, analysis and purification of RNA and ribozymes. Nucleic Acids Res. 23:26772684. Wright, M.C. and Joyce, G.F. 1997. Continuous in vitro evolution of catalytic function. Science 267:614-617. Yuan, Y. and Altman, S. 1994. Selection of guide sequences that direct efficient cleavage of mRNA by human ribonuclease P. Science 263:1269-1273. Zhao, H., Giver, L., Shao, Z., Affholter, J.A., and Arnold, F. H. 1998. Molecular evolution by staggered extension process (StEP) in vitro recombination. Nature Biotechnol. 16:258-261.
Contributed by Ronald R. Breaker Yale University New Haven, Connecticut
Combinatorial Methods in Nucleic Acid Chemistry
9.4.17 Current Protocols in Nucleic Acid Chemistry
In Vitro Selection of RNA Aptamers to a Small Molecule Target
UNIT 9.5
This unit describes the selection of aptamers that bind to a small molecule target from a single-stranded RNA pool. Aptamers generated by this type of selection experiment can potentially function as receptors or biosensors for small molecules, and may be useful in applications ranging from medical diagnostics to environmental monitoring. As described in UNIT 9.3, aptamers are artificially evolved through an iterative process of selection and amplification. A pool of RNA is transcribed from a synthetic pool of double-stranded DNA (UNITS 9.2 & 9.3). The RNA pool is partitioned to isolate species that can bind to a target on a solid support matrix. Two modes of selection are presented in this unit: (1) selection by column filtration (see Basic Protocol), and (2) selection by batch absorption (see Alternate Protocol). Bound species are eluted off and amplified for the next round of selection using a combination of reverse transcription, PCR, and in vitro transcription (UNIT 9.3). The progress of the selection is monitored by assaying for the accumulation of binding species (see Support Protocol). Examples of selections that have targeted small molecules are shown in Table 9.5.1. STRATEGIC PLANNING Affinity Matrices for Column Selection If an affinity matrix with an immobilized target is not commercially available, an appropriate activated resin must be functionalized with the target molecule or derivative. The presentation and loading of a target molecule on an affinity matrix can greatly influence the course of a selection experiment. There are many different coupling chemistries available for attachment, and each is highly dependent on the functional groups that are available on both the target and the resin. A few examples of matrices that have been successfully used for the selection of aptamers against small molecule targets are provided in Table 9.5.1. Instructions for conjugating targets to resins are found in the references given in Table 9.5.1, and are typically detailed in instructions from the resin manufacturers. See Critical Parameters for further discussion of derivatization. It is especially important to make sure that the resin itself does not bind the RNA pool. For example, unless it was appropriately blocked, an amine-coated resin would electrostatically interact with the negatively charged backbone of RNA, causing a major nonspecific binding problem. It is also useful to determine what fraction of the RNA pool initially binds to both the underivatized matrix and to the prepared affinity matrix (see Support Protocol). If too much (>10%) of the RNA pool binds to the underivatized matrix, a different conjugation chemistry or binding buffer should be chosen. If too much (>10%) binds to the targetbound affinity matrix, the stringency of the selection conditions should be increased (see Critical Parameters). Functionalized Beads for Batch Selection If magnetic beads with the desired target are not available, they must be functionalized with the target or derivative. Magnetic beads bearing amine, carboxylate epoxy, and tosyl-activated surfaces are available (Dynal) and can be derivatized with the appropriate target molecules. The manufacturer’s instructions provide information on the concentra-
Contributed by Sulay Jhaveri and Andrew Ellington Current Protocols in Nucleic Acid Chemistry (2002) 9.5.1-9.5.14 Copyright © 2002 by John Wiley & Sons, Inc.
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Resins and Attachment Chemistry for Small Molecule Aptamer Targetsa
Table 9.5.1
Selection reference
Commercially available resin
Small molecule target
Resin
Attachment chemistry/preselection
Aminoglycosides
Agarose
Immobilization of targets via coupling of primary or secondary amines to epoxy activated agarose; Preselection performed on glycine-agarose
L-Arginine
Thiopropyl Sepharose 6B
L-Arginine-L-cysteine
dipeptiede linked Connell et al. through the sulfhydryl group to the matrix; (1993) Preselection on L-cysteine-thiopropyl Sepharose
ATP
Agarose
Cyanogen bromide activation, C8 linkage, 9-atom spacer; Preselection on agarose, and GTP agarose
Cibaron blue
4% cross-linked agarose
Epoxy activation, 21-atom spacer; Ellington and Preselection on oligo (dT)12-18 on cellulose Szostak (1990)
Sigma; Amersham Pharmacia Biotech
L-Citrulline
Sepharose 6B
Coupling to N-hydroxysuccinimide-activated agarose (CH-4B), 11-atom spacer; Preselection on glycine derivatized epoxy-agarose
Famulok (1994)
Amersham Pharmacia Biotech
Guanosine
Affi-Gel 102 agarose
GDP-βS linked through sulfur to resin (Bio-Rad) derivatized with bromoacetyl N-hydroxysuccinimide; Preselection on acylated agarose column
Connell and Yarus (1994)
N-Methylmesoporphyrin (NMM)
Acrylic
NMM reaction with oxirane-acrylic beads; Li et al. (1996) Preselection on 2-mercaptoethanol-blocked (DNA) oxirane beads
Neomycin
Sepharose
Neomycin sulfate coupled to epoxy-activated Sepharose; Preselection on glycine-agarose column
Nicotinamide
Agarose
Cyanogen bromide activation, attachment Lauhon and Szostak (1995) to ribose hydroxyl, 11-atom spacer; Preselection on ATP-agarose, and elution with quinoxaline-2-carboxylic acid prior to affinity elution
Sigma
Reactive blue
4% cross-linked agarose
Triazine coupling; Ellington and Preselection on oligo (dT)12-18 on cellulose Szostak (1990)
Sigma; Amersham Pharmacia Biotech
Riboflavin
Agarose
Lauhon and N3-carboxymethyl riboflavin attached at Szostak (1995) the carboxyl group to adipic dihydrazide-modified agarose using EDC; Preselection on adipic dihydrazide agarose, elution with quinoxaline-2-carboxylic acid prior to affinity elution
Lato et al. (1995) Pierce
Jhaveri et al. (2000) (Fluorescent RNA)
Wallis et al. (1995)
Sigma Sigma
Amersham Pharmacia Biotech
aInstructions for binding targets to resins are found in the references given, and are clearly detailed in instructions from resin manufacturers.
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tion of active functional groups on the magnetic beads. For example, 200 µL of aminecoated Dynabeads corresponds to 1 µmol of active functional groups. Details on attachment of targets to beads can be obtained from the manufacturer (also see Critical Parameters for a discussion of derivatization). Alternatively, beads and targets can be coupled via hetero-bifunctional cross-linkers (e.g., Pierce). It is also useful to determine what fraction of the RNA pool initially binds to both the underivatized beads and the beads that contain immobilized target (see above). ISOLATING A FUNCTIONALLY ENRICHED POOL BY COLUMN SELECTION
BASIC PROTOCOL
In the following protocol, an RNA pool is passed through two different columns in series: a preselection column that removes nonspecifically bound material, and an affinity column derivatized with a target molecule that collects RNAs that bind specifically to the target. RNA species captured on the column are washed off with elution buffer containing a molar excess of target (i.e., affinity elution). This method can be done in batch mode, especially if only a small quantity of resin is available. However, one must be very careful when separating the RNA pool bound on the solid phase from the remaining unbound RNA. For this reason, the authors prefer the use of magnetic beads for batch selection (see Alternate Protocol). NOTE: All solutions and buffers should be prepared fresh and should be autoclaved or treated with diethylpyrocarbonate (DEPC). Sterile disposable plasticware should be used wherever possible. See APPENDIX 2A for guidelines on how to protect against contaminating RNases. Materials Resin (Table 9.5.1) Affinity resin with immobilized target molecule (see Strategic Planning and Table 9.5.1) Binding buffer (see Critical Parameters) RNA pool (UNIT 9.3) Affinity elution buffer (see Critical Parameters) 5 M NaCl 100% and 90% (v/v) ethanol, room temperature and ice cold, respectively 2-mL, 0.8 × 4–cm columns with cap and lid (e.g., Poly-Prep or Econo-Prep columns, Bio-Rad) 65° to 75°C water bath 13-mL collection tubes (e.g., Sarstedt) Prepare columns and RNA 1. Prepare two 0.8 × 4–cm columns: a preselection column that contains resin, and a selection column that contains a target molecule conjugated to the resin. Pack each column with resin to obtain a 1-mL bed volume. Use disposable sterile columns, caps, and lids. Do not reuse. Column size can range from 100 ìL to 4 mL. The concentration of target can range from <1 to 20 mM. See Critical Parameters for further discussion of the size of the column, the concentration of target on the column, and the stringency of selection. Combinatorial Methods in Nucleic Acid Chemistry
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2. Rinse each column five times with 5 mL binding buffer. Prior to the application of the RNA pool to the column, the resin should be equilibrated in binding buffer in order to remove any undesired compounds that may initially be present in the resin (e.g., preservatives such as sodium azide, often found in commercially available resins; or salts, organics, or leaving groups that remain from coupling the target to the resin). Take care not to dry out the column while washing the resin. Dry columns develop cracks and channels that can divert solutions away from the immobilized target molecules.
3. Before loading the RNA pool, make sure that there are no bubbles in the matrix. These, too, can divert solution and impede flow. To ensure a uniform matrix, the resin in the column can be briefly vortexed in the presence of binding buffer or can be gently pipetted up and down.
4. Use ∼10 µg of the RNA pool (∼1014 sequences) for selection. Refer to UNITS 9.1 & 9.3 for discussions of the amount of RNA to be used. Using significantly lower amounts of RNA may result in insufficient representation of individual sequences, while significantly larger quantities of RNA may lead to viscosity problems or precipitation of the RNA pool.
5. To ensure that each species in the RNA pool folds into the most accessible or most stable conformation, heat the RNA pool in 100 µL binding buffer 3 min at 65° to 75°C, and then allow the sample to cool to room temperature over ∼10 min. Keep the binding buffer temperature and composition (i.e., the ionic strength, monovalent and divalent cation concentrations, pH) constant during the early rounds of selection when productive binding species are accumulating, as these parameters can influence interactions with the target. Higher temperatures for thermal equilibrations are cautioned against, due to the degradative effects of divalent cations on RNA.
Select RNA 6. Cap the tip of the washed preselection column and load 100 µL of solution containing the RNA pool onto the column. Be careful to load the RNA onto the column gently, disturbing the interface as little as possible. Some columns come with plastic frits that can be placed on top of the resin to reduce surface perturbation.
7. Cover the column with the lid and incubate the RNA pool with the underivatized or appropriately blocked matrix for 10 min at room temperature. Negative selection to remove matrix-binding species is an extremely important step in any selection procedure. Initially, the target will have a low affinity for the aggregate pool, and the surface area of the target will be much less than the surface area of the matrix, so the probability of accruing matrix-binding species is relatively high. If matrix-binding species are not effectively sieved from the population, they will quickly accumulate to a point where it may be extremely difficult to select aptamers. During later rounds of selection, using an affinity column containing a structurally similar target as the negative selection will enhance the specificity of the aptamer for its cognate target. For example, in the course of a selection against ATP immobilized on agarose, GTP-agarose beads could be used in place of agarose beads as a negative selection in later rounds. In Vitro Selection of RNA Aptamers to a Small Molecule Target
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8. Collect the eluate from the column in a 13-mL collection tube. Rinse the column three times with 0.5 mL binding buffer and collect the eluate in the same tube. Rinsing the preselection column with larger volumes of binding buffer might elute RNAs that have an intrinsic affinity for the matrix.
9. Cap the tip of the prewashed selection column and load ∼1.6 mL of the eluate from the preselection column, being careful not to disturb the surface of the resin. 10. Cover the top of the column and incubate the RNA pool with the affinity matrix for up to 1 hr at room temperature. See Critical Parameters for a discussion of incubation times.
11. Remove the cap, allow the eluate to flow out of the column, and discard the eluate. At this point, RNA molecules that do not bind to the target have been removed from the pool. Further washing of the column will result in the removal of weakly bound species.
12. Wash the column at least two times with at least two column volumes (each) of binding buffer. The volume of wash buffer directly affects the stringency of the selection (see Critical Parameters for a discussion of stringency). It is recommended that the volume of washes be low during the early rounds, so binding species have a chance to accumulate. The volume can then be increased as the selection progresses. During later rounds of selection, a larger volume of washes will remove RNAs that have faster koff and/or lower Kd values from the column. The cumulative effect of more washing will be to retain only those RNA molecules that can form tight complexes with their targets. As before, be careful to avoid disturbing the surface of the resin. This is more important during wash steps than during loading steps, since any resin kicked into solution will initially be removed from the immobile phase, allowing matrix-binding species to accumulate near the top of the column.
Elute and precipitate enriched RNA 13. Apply 1 mL affinity elution buffer to the column and incubate 10 min, room temperature. Collect the eluate in a 13-mL collection tube. See Critical Parameters for a discussion of the composition of the elution buffer. The elution buffer should contain a significant molar excess of the target in the mobile phase to ensure that released RNA molecules remain in the mobile phase. It may also be possible to release RNA molecules from the affinity column by denaturation.
14. Add at least two column volumes of elution buffer and allow to equilibrate 5 min. Collect wash and add to the eluted fraction from step 13. Repeat wash at least once more. If radiolabeled RNA is used, the flow-through, wash, and elution fractions can be calculated (see Support Protocol). This will allow the progress of the selection to be easily monitored.
15. Add 5 M NaCl to the eluate at a final concentration of 0.3 M and then add 2 vol of 100% ethanol to precipitate the RNA. Vortex briefly and incubate 30 min at −20°C or 5 min at −80°C. Centrifuge at 10,000 × g, 4°C, to recover the precipitate. It is a common practice to add 1 to 2 ìg of glycogen to increase the yield of the nucleic acid precipitate and to better visualize the pellet. If the selection target binds to or interacts with glycogen, it should be omitted from the precipitation. Combinatorial Methods in Nucleic Acid Chemistry
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16. Rinse the RNA pellet with 1 mL ice-cold 90% ethanol and remove any excess liquid from the tube, taking care not to damage the pellet with the pipet tip. Dry the pellet. The pellet can be air dried, dried under a nitrogen or argon stream, or dried in a SpeedVac evaporator. Cover the tube with Parafilm and poke holes with a sterile needle to allow evaporation to proceed while avoiding inadvertent nuclease contamination.
17. Resuspend the pellet in 30 µL RNase-free sterile water and proceed with amplification of the pool via reverse transcription and PCR (UNIT 9.3). It is usually a good idea to save some of the pool (a third to a half of the resuspended eluate) for archival purposes. If subsequent amplification or selection reactions fail for any reason, the procedure can be started from the archival pool. ALTERNATE PROTOCOL
ISOLATING A FUNCTIONALLY ENRICHED POOL BY BATCH SELECTION The isolation of aptamers against small molecule targets via batch selection is very similar, in principle, to column selection. However, instead of using chromatography, RNA samples are simply mixed with resins and the pool is partitioned between solid and liquid phases (as opposed to immobile and mobile phases). As before, the RNA pool is incubated with a preselection resin followed by a target-bound resin. Bound RNA is competed off the resin with elution buffer. Although affinity resins can be readily adapted to batch selections, magnetic beads may be used as a convenient solid support, as described here. The use of magnetic beads also makes this protocol amenable to automated selection. Additional Materials (also see Basic Protocol) Magnetic beads (Dynal), with and without immobilized small molecule target (Dynal; see Strategic Planning) Magnet or magnetic concentrator (Dynal) Prepare magnetic beads 1. Add 200 µL of each magnetic bead solution to 1.5-mL microcentrifuge tubes. Set up one tube for preselection beads that contain no immobilized target and one tube for selection beads that contain immobilized target. The amount of resin used can to some extent be regarded as a parameter that can modulate the stringency of selection. During later rounds, the amount of beads (and thus binding sites) used for selection can be reduced, thereby increasing the competition between RNA species. This can also be achieved by lowering the concentration of target conjugated to the beads. For a fuller discussion of stringency, see Critical Parameters.
2. Add 1 mL binding buffer to each tube and place the tube next to a magnet, or in a magnetic concentrator, for 1 min to allow beads to concentrate on the side of the tube. Gently remove supernatant with a transfer pipet. Repeat for a total of five washes. After the final wash, resuspend the beads in a final volume of 100 µL binding buffer. Do not let the beads dry out. Also, note that the beads are easily disturbed by turbulance and use this opportunity to practice removing supernatant without disturbing the beads.
Select RNA 3. Prepare the RNA pool (see Basic Protocol, steps 4 and 5) and add to the batch of preselection beads. Gently pipet up and down several times to disperse the RNA among the beads. This procedure should be carried out away from the magnet. In Vitro Selection of RNA Aptamers to a Small Molecule Target
4. Incubate the RNA with the preselection beads for 10 min at room temperature.
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5. Concentrate the magnetic beads to one side of the tube (step 2) and transfer the supernatant to the tube containing beads with immobilized target. Disperse the beads by pipetting up and down. 6. Incubate the RNA pool with the target-bound beads for 1 hr at room temperature. Place the tube on a benchtop rotator or shaker to prevent the beads from settling out. See Critical Parameters for a discussion of incubation times.
7. Magnetically concentrate the beads and discard the supernatant. At this step, RNA molecules that do not bind to the target have been removed from solution.
8. Wash the beads two times with 200 µL binding buffer. Weakly bound RNA molecules are removed at this step. As suggested in the Basic Protocol, keep the number or volume of washes low during the earlier rounds of selection, and increase as the selection progresses.
Elute and precipitate RNA 9. Add 200 µL affinity elution buffer to the beads and incubate 10 min at room temperature. Concentrate the beads and collect the eluate. 10. Repeat the elution and pool the two fractions of RNA competed off the beads. 11. Precipitate and amplify the eluted RNA as described (see Basic Protocol, steps 15 to 17). ASSAYING THE ACCUMULATION OF BINDING SPECIES In order to determine if the RNA pool is becoming selectively enriched with sequences that bind the small molecule target, the pool should be assayed periodically. A radiolabeled RNA pool is applied to the resin, and the fraction of the RNA pool that is retained on the resin is measured. Additionally, the fraction of the RNA pool that can be eluted from the resin with soluble target is measured. Assaying the RNA pool for its ability to bind to both the preselection resin and the selection resin (containing immobilized target) should be performed in parallel to evaluate whether the selected RNA pool specifically binds to the target.
SUPPORT PROTOCOL
While this procedure applies primarily to the Alternate Protocol, it can also be carried out with the Basic Protocol by substituting column fractions for aliquots washed from the beads. Although this protocol is a simple way to determine the extent to which the pool binds to an immobilized target, binding affinity constants for individual aptamers can also be measured using techniques such as equilibrium gel filtration, microdialysis, or changes in anisotropy of fluorescent ligands. Additional Materials (also see Basic Protocol and Alternate Protocol) Radiolabeled RNA pool (UNIT 9.3) 0.45-µm, 13-mm HAWP nitrocellulose filter disks (Millipore) Glass plate Phosphorimager screen or X-ray film Phosphorimager or densitometer 1. Thermally equilibrate 1 µg radiolabeled RNA pool in 100 µL binding buffer (see Basic Protocol, step 5). Combinatorial Methods in Nucleic Acid Chemistry
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2. Remove 5 µL to determine the total amount of radiolabeled RNA present in the binding reaction (A). Pipet this sample onto a 0.45-µm nitrocellulose filter disk and set the disk aside on a glass plate. 3. Add the remaining radiolabeled RNA to a 100-ìL solution of affinity resin (see Alternate Protocol, steps 1 and 2). Pipet up and down several times to disperse the beads. 4. Incubate the RNA pool with the beads for 1 hr at room temperature. Place the tube on a benchtop rotator or shaker to prevent the beads from settling out. 5. Magnetically concentrate the beads and remove a 5-µL aliquot from the supernatant to measure the amount of RNA that did not bind to the beads (B). Pipet this sample onto a filter disk and discard the remaining supernatant. 6. Wash the beads twice with 200 µL binding buffer. Combine the washes and remove a 5-µL aliquot to measure the fraction of the pool that has been washed off (C). Pipet onto a filter disk and discard the remaining washes. The number or volume of washes can be varied to roughly determine the affinity of the RNA for the immobilized target. The number and volume of washes should reflect those used at the conclusion of the selection.
7. Add 200 µL affinity elution buffer to the beads and incubate for 10 min at room temperature. Transfer the supernatant containing RNA species that have been eluted from the beads into a separate tube. 8. Repeat the elution and combine with the first eluent. Take a 5-µL aliquot to measure the amount of RNA that has been eluted off (D). Pipet onto a filter disk. 9. Resuspend the beads in 200 µL binding buffer, make sure that the beads are evenly dispersed, and take a 5-µL aliquot containing beads and any remaining radioactivity (E). Pipet onto a nitrocellulose filter disk. The aliquot of beads must be taken before the beads are allowed to settle. It may prove difficult to manipulate the beads onto the solid support. This procedure is only semi-quantitative, and thus the loss of some beads (e.g., on the inside of the pipet tip) is acceptable.
10. With all of the nitrocellulose disks on the glass plate, cover the plate with plastic wrap and expose the plate (plastic side up) to a Phosphorimager screen or X-ray film for 4 to 12 hr. The exposure time will vary greatly depending on the specific activity and decay of the radioactivity incorporated into the RNA pool.
11. Count the radioactivity on the filters using a Phosphorimager or densitometer and calculate the fraction of the RNA pool bound to the resin, the fraction washed off, the fraction eluted off, and the fraction retained on the beads following elution: Fraction bound = (19A − 39B)/19A Fraction eluted = 80D/19A Fraction washed = 80C/19A Fraction retained = 40E/19A The values 19, 39, 40, and 80 are dilution factors used with the different 5-ìL aliquots. The eluted, washed, and retained fractions should add up to equal the total bound, and the bound and unbound (39B/19A) should equal 1. In Vitro Selection of RNA Aptamers to a Small Molecule Target
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The initial fraction of the RNA pool that binds to the underivatized beads or to the beads that contain immobilized target should be <10%. If the fraction of the RNA pool that can be eluted off with the target approaches 10% to 15% above the initial fraction (e.g., 2.5% of the pool initially bound, 14% bound after five rounds of selection), the pool should be cloned and sequenced to isolate individual aptamers.
COMMENTARY Background Information Although the principle of in vitro selection was expounded more than 30 years ago (Mills et al., 1967; Levisohn and Speigleman, 1969; Kramer et al., 1974), the development of modern molecular biology techniques such as PCR and the chemical synthesis of DNA greatly potentiated the development of workable methods. In 1990, Ellington and Szostak first implemented the selection technology to enrich a pool of RNA molecules to bind to small organic dyes (Ellington and Szostak, 1990). Since then, in vitro selection methods have been used to identify aptamers against a wide variety of small molecules, including small inorganic ions such as metal ions (Kawakami et al., 2000); metabolites such as ATP (Sassanfar and Szostak, 1993; Huizenga and Szostak, 1995) and theophylline (Jenison et al., 1994); and large organic compounds such as porphyrins (Li et al., 1996; Okazawa et al., 2000) and short peptides (Xu and Ellington, 1996). In fact, researchers have selected aptamers against a wide variety of small molecules of biological importance (reviewed in Ellington, 1994; Gold et al., 1995; Famulok and Jenne, 1998).
Critical Parameters Choice of target There is seemingly no common functional theme uniting the molecules that have so far proven to be good targets for in vitro selection. However, upon closer inspection, some commonalities can be identified (see also Ellington, 1994). Small molecules that are already known to interact with RNA should obviously make good selection targets. In this respect, aminoglycoside antibiotics function by interacting with ribosomal RNA and have yielded aptamers (Lato et al., 1995; Wallis et al., 1995; Berens et al., 2001). Small molecules that contain chemical functionalities that are complementary to those of RNA should also be good selection targets. For example, selections that target charged aromatic compounds, heterocyclic rings, or compounds resembling or containing nucleotides have frequently been successful. Finally, larger targets seem to elicit
tighter-binding aptamers than smaller ones, possibly due to a larger number of contact sites between the ligand and the RNA pocket. Structural analyses of aptamers have revealed that the selected RNAs generally use short helical segments to form simple pockets around their cognate ligands (Patel and Suri, 2000). This may explain why aromatic and heterocyclic rings are readily bound by nucleic acids, since flat shapes can easily fit into helical grooves (as opposed to more bulky compounds, e.g., cubane). Indeed, the tightest-binding aptamers that have been discovered thus far recognize nucleotides or heterocyclic aromatic compounds resembling nucleotides. Although these considerations may assist in the choice of a small molecule target, it is likely that a researcher will already have a target in mind, irrespective of whether or not it meets these criteria. Therefore, it is worthwhile to assess the chances for success by performing a quick binding assay with the original random sequence pool and an appropriate affinity matrix or derivatized bead (see Support Protocol). If the random sequence pool shows even a slight affinity for the target (>1% to 2% retention), there is a good chance that there will be at least some species in that pool with high affinities for the target (Kd <10 µM). Making an affinity column Choosing a matrix. There are three considerations when choosing a matrix: the chemical nature of the matrix itself, the spacer arm, and the functional group. Various chemical permutations of agarose beads have been consistently used as affinity matrices. Agarose is a particularly good choice for affinity chromatography because of its stability under various pH, ionic, and temperature conditions, and because the hydroxyl group on the sugars can be easily derivatized for the chemical conjugation of target molecules (Axen et al., 1967). Acrylic (Li et al., 1996) and cross-linked polystyrene beads (from Dynal) have also been used as selection matrices. Derivatization chemistries. Although agarose can be derivatized with ligands, there are a number of chemically modified matrices
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In Vitro Selection of RNA Aptamers to a Small Molecule Target
(e.g., Celite, acrylic, polyacrylamide, Affi-Gel 10) that are sold specifically for the preparation of affinity columns. These matrices typically present either a spacer arm with a pendant functional group (e.g., a sulfhydryl or primary amine) or a preactivated functional group (e.g., tosyl-activated thiols, N-hydroxysuccinimide (NHS)–activated carboxylates, as well as oxirane, vinylsulfone, thiopropyl, and periodate groups). Depending on the chemistries available on the target of choice, it may be possible to merely mix the target with matrix in order to generate an affinity matrix. For example, a target that contains a primary amine can be directly mixed with a matrix containing NHSactivated carboxylate to give an amide linkage between the target and the matrix. These matrices should be used according to their manufacturer’s instructions. Concentration of target. Following derivatization, the concentration of the target on the matrix can range from submillimolar up to 20 mM. The actual amount is dependent on the concentration of functional groups that were originally available and the yield of the coupling reaction. Ideally, the concentration of the target on the resin should be a controllable feature. In practice, it is sometimes very difficult to control or even determine the concentration of a derivatized target. Depending on the target molecule, it may be possible to assay the amount of target remaining in solution following derivatization, and to thereby calculate the approximate concentration on the column. For example, if the target molecule has a strong absorbance, then the optical density of the solution containing the target can be determined before and after derivatization. It should be noted, though, that many leaving groups have strong absorbances in their own right, and thus can disguise the level of derivatization that has occurred. Alternatively, if there is a convenient biochemical assay for the target molecule, the concentration pre- and postderivatization can again be determined. If no signal from or assay for a target molecule is available, or if the signal or assay is somehow disguised, then a control reaction can be set up in parallel with the target-conjugation reaction. For example, amine derivatives of fluorescein can be readily observed, and could be conjugated in a separate but parallel reaction to an amine-bearing target that had no easily observable signal. While the extrapolation of concentration via this method is only approximate, it is nonetheless a useful control to ensure that the conjugation chemistry is working.
Blocking reactive groups. Functional or activated groups that were not originally conjugated to the target molecule should be subsequently blocked with an inert functional group. For example, Li et al. (1996) blocked oxirane-acrylic beads with 2-mercaptoethanol, while the authors’ laboratory typically blocks underivatized NHS-carboxylates with glycine or ethanolamine. The concentration of the blocking ligand should be in gross excess relative to the concentration of the functional or activated groups on the matrix to ensure that all remaining reactive sites are efficiently blocked. Column size. Column volumes can range from as little as 100 µL up to 4 mL. With significantly smaller volumes, problems with reproducibility and flow may arise due to the resin sticking to the walls of the column, tubes, or pipet tips. If only a small amount of affinity resin is available for experiments, volumes <100 µL should be used in a batch selection format. Conversely, a significantly larger column (i.e., >10 mL) will make the manipulation of washing and elution volumes unwieldy. Irrespective of the column size, the amount of target on the column should generally be in excess of the RNA pool in order to ensure the complete capture of binding species (however, see discussion below on controlling the stingency of selection). Negative selection Members of the RNA pool can potentially bind to the selection matrix, to linker between the matrix and the target molecule, to the target molecule itself, or to some combination of these. To encourage the recognition of the target molecule, it is important that the selection matrix and the linker do not nonspecifically bind to RNA. In particular, affinity columns that utilize CNBr coupling to agarose typically leave behind a positively charged linkage that can result in high levels of nonspecific binding. For this reason, commercially available affinity matrices (e.g., from Sigma) should be carefully scrutinized prior to use. The level of nonspecific binding for a given matrix can be emperically determined using the methods described in the Support Protocol. Irrespective of the initial level of nonspecific binding to the matrix, preselection can be used to minimize or decrease the enrichment of matrix-binding RNA species. The negative selection matrix should contain exactly the same matrix, spacer arm, and conjugation chemistry as the affinity matrix (see Making an affinity column, above), except that an “inert” ligand
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should be derivatized to the matrix. In general, the inert ligand can be the same compound used to block the affinity matrix. However, when attempting to select for extremely specific aptamers, the ligand on the negative selection matrix can be related to the target ligand. For example, a selection for highly specific antiadenosine aptamers might utilize an affinity matrix containing guanosine as the negative selection matrix. However, it should be noted that increasing the stringency of the negative selection will increase the stringency of the selection overall (see Controlling the stingency of selection, below). Thus, decreasing the total number of adenosine-binding species in the pool may paradoxically lead to the accumulation of matrix-binding molecules (e.g., agarose-binding species). If, during the course of selection, the RNA pool is overrun by matrix-binding species (as determined using the Support Protocol), it may still be possible to recover productive targetbinding species. This is generally not possible when filtration is used as a partition method (see UNIT 9.3). The stingency of the negative selection can be increased by passing the pool over multiple preselection columns and/or reducing the number or volume of washes of the preselection column. Alternatively, the relative contributions of matrix-binding and targetbinding species to the population as a whole can be altered by increasing the concentration of target on the column (see Making an affinity column, above). If matrix-binding species prove to be intractable, it may be possible to alternate between affinity chromatography and other selection modalities, such as filtration (assuming the target molecule binds to a filter; UNIT 9.3) or gel-shift assays. Matrix-binding species that have an affinity for agarose are unlikely to have an affinity for either modified cellulose or acrylamide, and vice versa. Choosing binding conditions The binding buffer should promote tight and specific binding of the RNA to the small molecule target. Two simple considerations are that the buffer should not cleave the target from the matrix and should not degrade the RNA. In this respect, it is important to once again note that all buffers should be treated to deactivate contaminating RNases. Monovalent cations (such as Na+ or K+) and divalent cations (such as Ca2+ or Mg2+) are common ingredients in selection buffers, because they stabilize the structure of RNA molecules and can contribute to both specific and nonspecific binding to target mole-
cules. Both monovalent and divalent cations act to stabilize the structural integrity of folded aptamers by neutralizing the close approach of nucleic acid strands, and can participate in salt bridges with target molecules. More specific divalent cation–binding sites can evolve, and these may promote the formation of specific aptamer structures or interactions with target molecules. A typical binding buffer contains 5 to 20 mM of compounds that can hold pH near neutrality, such as Tris, HEPES, or phosphate. Monovalent salts such as NaCl, potassium acetate, or LiCl have been used at concentrations of 50 to 200 mM. Divalent salts such as MgCl2 or CaCl2 are frequently (although not always) included at concentrations of 1 to 10 mM. Biopolymers such as bovine serum albumin (BSA) or, better, tRNA can be included in order to saturate nonspecific binding sites on the matrices and to increase the stringency of the selection (also see Controlling the stingency of selection, below). While there have been no real systematic studies of the effects of buffer conditions on aptamer-target interactions, it is likely that selected nucleic acids will bind near optimally in the presence of their binding buffer. If there is a need to change the buffer conditions following selection (for example, to better approximate physiological conditions), it is best to partially randomize the aptamer and reselect for binding in the presence of the new buffer. Incubation time is also a variable during binding conditions. Species with high affinity can be readily selected by allowing the reaction to equilibrate for only 5 min. However, to account for slow folding or refolding in the presence of the target molecule, incubation times as long as 1 hr may be used. Choosing elution conditions There are a number of ways that aptamers can be eluted from the affinity matrix. For example, since RNA structure is dependent on monovalent and divalent cations, changing the ionic composition of the buffer can destabilize the aptamer-target interactions. Increasing the temperature of the elution buffer can cause RNA molecules to unfold and hence be released from the column. However, the preferred method is ligand-specific elution of RNA (affinity elution), as this ensures that the selected RNAs bind to the free ligand and its immobilized affinity derivative at the same site (Connell and Yarus, 1994). In general, aptamers can be specifically eluted off with soluble target at concentrations ranging from 1 to 50 mM. The
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concentration of the target in solution should generally be higher than the concentration of the target on the affinity column. Just as preselection using a related target can potentially increase the specificities of selected RNA molecules, it is possible that an initial elution with a related target will increase the specificities of RNA molecules that are eventually eluted with the cognate target (Lauhon and Szostak, 1995). Finally, in some instances it may be possible to simultaneously detach both target and nucleic acid from a column. If the spacer arm connecting the target to the column contains a labile moiety such as a disulfide bond, the disruption of this moiety (e.g., with 2-mercaptoethanol) will concomitantly detach any bound aptamers. The advantage of this method is that one is assured of recovering even the tightest-binding aptamers. Time is also an important consideration for the elution of specifically bound species. RNA molecules trapped on an affinity column will be able to interact with a mobile phase containing soluble target according to their rates of dissociation, or koff values. The tighest-binding species will have very long residence times, and thus might be missed with a short wash step. Therefore, it is frequently useful to run in the elution buffer, cap the bottom of the column, and let the mixture equilibrate prior to collecting the eluate. Alternatively, bound species can be repartitioned with target in the solution phase by heat denaturation, as described below. Different elution methods can be combined to yield aptamers with specific properties. As an example, Geiger et al. (1996) used a combination of heat denaturation and specific elution to identify aptamers that were highly specific for arginine. After washing an arginine affinity column with buffer, the authors included an elution step with a noncognate amino acid, citrulline. The column-bound RNA was heat denatured and renatured in the presence of citrulline. The column was then developed with arginine, and another heat denaturation/renaturation step was employed to repartition RNA species between immobile and mobile phases. The authors’ rationale was that the tightest binders, i.e., the molecules with slow dissociation rates, would separate from the immobilized arginine when heat denatured and renature in the presence of arginine, in solution.
In Vitro Selection of RNA Aptamers to a Small Molecule Target
Controlling the stringency of selection Based on the parameters described above, general rules for how to increase (or decrease) the stringency of a selection experiment as a
function of the round of selection can be advanced. Normally, selection stringency will be gradually increased throughout the course of a selection experiment in order to identify the tightest-binding aptamers. The authors typically monitor selections every three to five rounds using the methods described here (see Support Protocol; also see Anticipated Results) and increase stringency if it seems that a significant number (>15% to 20% of binding species) have accumulated. Occasionally, selection stringency will need to be decreased, for example, to increase the representation of target-binding species relative to matrix-binding species. Affinity matrix. Selection stringency can be increased by decreasing the concentration of the small molecule on the matrix in the later rounds of selection. Competition for a smaller number of binding sites should favor tighter binding aptamers. Since it is practically difficult to control the concentration of the target on the column, it is sometimes useful to dilute a premade batch of affinity resin with negative selection resin in order to reduce the aggregate concentration of target on a column. Binding buffer. Increasing the ionic strength of the binding buffer may decrease nonspecific electrostatic interactions with the column matrix. However, the affinity of an aptamer for a charged or polar target may also decrease at higher ionic strengths. Since it is unclear, in advance, whether the quest for enhanced specificity will be detrimental to aptamer affinity, selections should be initiated with a generic buffer recipe. Nonetheless, as aptamers begin to accumulate in a selection (as determined by a binding assay, see Support Protocol), the authors routinely increase the ionic strength of binding (and wash) buffers to further parse the selected population and identify only the tightest-binding species. The use of nonspecific molecular competitors and noncognate molecular competitors to increase specificity is mentioned above (see Choosing binding conditions and Choosing elution conditions, respectively). Both of these methods also apply to increasing the stringency of a selection: the more nonspecific or noncognate molecular competitor present in the binding (or wash) buffers, the more one is likely to remove weakly bound but specific aptamers from the column. As a final consideration, specific competitors can also be used. For example, AMP might be included in a binding buffer to enhance the affinities of selected anti-ATP aptamers.
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Wash steps. As indicated above, many of the considerations that apply to the composition of binding buffers to increase stringency also apply to wash buffers. In addition, though, increasing the number of washes over the course of a selection prior to affinity elution will increase the stringency of the selection. During the early rounds, washing with as little as two column volumes will be enough to clear the column of weakly binding species while retaining the tighter ones. An excellent example of this can be found in Wallis et al. (1995). The enrichment of neomycin-specific RNAs is brought about by progressively increasing the number of washes from 5 column volumes in the first round, to 30, and then to 50. Affinity elution. As might be imagined, many of the considerations that apply to the composition of binding buffers to increase stringency also apply to elution buffers. For example, aptamers can be eluted at increasing ionic strengths over the course of a selection experiment. For affinity elution, the concentration of affinity eluant can initially be quite high, guaranteeing partition to the solution phase, and can then be decreased over the course of a selection experiment to identify tighter-binding species. The rationale for this approach is that species that have differential affinity for free ligand (for example, those species whose binding may be partially inhibited by the presence of a spacer arm) may more successfully compete for increasingly limited amounts of free ligand. However, affinity elution with low concentrations of soluble target should probably be performed only after it has been well established that the pool prolifically binds to its target. Overall, this suggestion will likely work best if long incubation times or transient heating are used to potentiate partitioning of bound species between immobile and mobile phases.
Anticipated Results In order to monitor the progress of a selection, the pool should be periodically assayed for the accumulation of binding species (see Support Protocol). The authors suggest that this be done after the first five rounds and every three rounds thereafter. It is recommended that the accumulation of specific target-binding species be validated by running assays in parallel with negative selection columns. If no aptamers have accumulated after approximately twelve rounds, the selection should be started again with much less stringent conditions.
Aptamers against “good” targets will typically have dissociation constants on the order of 100 nM to 10 µM. Aptamers against “hard” targets (i.e., isoleucine; Majerfeld and Yarus, 1998) may have dissociation constants that are only slightly lower than might be expected for purely nonspecific hydrophobic interactions. It is entirely possible that aptamers will not be found at all for some target classes (e.g., fatty acids).
Time Considerations The time required for one round of selection (i.e., to go from one pool of selected DNA templates to the next) can range from 24 to 72 hr depending on the researcher and the demands of the particular selection experiment. While a significant portion of that time is required for amplification steps, the actual affinity selection, either on column or in batch, can take ∼2 to 4 hr. After the quantitation of the RNA (15 to 30 min), heat denaturation/renaturation steps can take another 15 min. This is ample time to prepare and wash the next column or batch of resin. Negative selection will take on the order of 30 min. The actual selection—including incubation times and wash and elution steps—can take anywhere from 30 min to 2 hr. This time can increase as the selection progresses, especially if a greater number of washes are being performed on a slow-flowing column. Overall, the amount of time it takes to carry out an entire selection experiment is contingent upon the number of rounds needed to accumulate target-binding species. The number of rounds depends in turn on the initial affinity of the pool for the target and the stringency regime used for selection. Typically, a selection experiment requiring twelve rounds plus radiolabeling and assaying the pool can take up to 25 or 30 days. This time can be significantly decreased by using automated methods (see UNIT 9.3).
Literature Cited Axen, R., Porath, J., and Ernback, S. 1967. Chemical coupling of peptides and proteins to polysaccharides by means of cyanogen halides. Nature 214:1302-1304. Berens, C., Thain, A., and Schroeder, R. 2001. A tetracycline-binding RNA aptamer. Bioorg. Med. Chem. 9:2549-2556. Connell, G.J. and Yarus, M. 1994. RNAs with dual specificity and dual RNAs with similar specificity. Science 264:1137-1141.
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Connell, G.J., Illangesekare, M., and Yarus, M. 1993. Three small ribooligonucleotides with specific arginine sites. Biochemistry 32:54975502.
Lauhon, C.T. and Szostak, J.W. 1995. RNA aptamers that bind flavin and nicotinamide redox cofactors. J. Am. Chem. Soc. 117:1246-1257.
Ellington, A.D. 1994. Empirical explorations of sequence space—host-guest chemistry in the RNA world. Berichte Der Bunsen-Gesellschaft-Physical Chemistry Chemical Physics 98:1115-1121.
Levisohn, R. and Speigleman, S. 1969. Further extracellular Darwinian experiments with replicating RNA molecules: Diverse variants isolated under different selective conditions. Proc. Natl. Acad. Sci. U.S.A. 63:805-811.
Ellington, A.D. and Szostak, J.W. 1990. In vitro selection of RNA molecules that bind specific ligands. Nature 346:818-822.
Li, Y.F., Geyer, C.R., and Sen, D. 1996. Recognition of anionic porphyrins by DNA aptamers. Biochemistry 35:6911-6922.
Famulok, M. 1994. Molecular recognition of amino acids by RNA aptamers: An L-citrulline binding RNA motif and its evolution into an L-arginine binder. J. Am. Chem. Soc. 116:1698-1706.
Majerfeld, I. and Yarus, M. 1998. Isoleucine:RNA sites with associated coding sequences. RNA 4:471-478.
Famulok, M. and Jenne, A. 1998. Oligonucleotide libraries—variatio delectat. Curr. Opin. Chem. Biol. 2:320-327. Geiger, A., Burgstaller, P., von der Eltz, H., Roeder, A., and Famulok, M. 1996. RNA aptamers that bind L-arginine with sub-micromolar dissociation constants and high enantioselectivity. Nucl. Acids Res. 24:1029-1036. Gold, L., Polisky, B., Uhlenbeck, O., and Yarus, M. 1995. Diversity of oligonucleotide functions. Annu. Rev. Biochem. 64:763-797. Huizenga, D.E. and Szostak, J.W. 1995. A DNA aptamer that binds adenosine and ATP. Biochemistry 34:656-665.
Mills, D.R., Peterson, R.L., and Speigelman, S. 1967. An extracellular Darwinian experiment with a self-duplicating nucleic acid molecule. Proc. Natl. Acad. Sci. U.S.A. 58:217-224. Okazawa, A., Maeda, H., Fukusaki, E., Katakura, Y., and Kobayashi, A. 2000. In vitro selection of hematoporphyrin binding DNA aptamers. Bioorg. Med. Chem. Lett. 10:2653-2656. Patel, D.J. and Suri, A.K. 2000. Structure, recognition, and discrimination in RNA aptamer complexes with cofactors, amino acids, drugs, and aminoglycoside antibiotics. J. Biotechnol. 74:39-60. Sassanfar, M. and Szostak, J.W. 1993. An RNA motif that binds ATP. Nature 364:550-553.
Jenison, R.D., Gill, S.C., Pardi, A., and Polisky, B. 1994. High resolution molecular discrimination by RNA. Science 263:1425-1429.
Wallis, M.G., Vonahsen, U., Schroeder, R., and Famulok, M. 1995. A novel RNA motif for neomycin recognition. Chem. Biol. 2:543-552.
Jhaveri, S.D., Rajendaran, M., and Ellington, A.D. 2000. In vitro selection of signaling aptamers. Nat. Biotechnol. 18:1293-1297.
Xu, W. and Ellington, A.D. 1996. Anti-peptide aptamers recognize amino acid sequence and bind a protein epitope. Proc. Natl. Acad. Sci. U.S.A. 93:7475-7480.
Kawakami, J., Imanaka, H., Yokota, Y., and Sugimoto, N. 2000. In vitro selection of aptamers that act with Zn2+. J. Inorg. Biochem. 82:197-206. Kramer, F.R., Mills, D.R., Cole, P.E., Nishihara, T., and Speigelman, S. 1974. Evolution of in vitro sequence and phenotype of a mutant RNA resistant to ethidium bromide. J. Mol. Biol. 89:719736. Lato, S.M., Boles, A.R., and Ellington, A.D. 1995. In vitro selection of RNA lectins—using combinatorial chemistry to interpret ribozyme evolution. Chem. Biol. 2:291-303.
Contributed by Sulay Jhaveri Naval Research Laboratory Washington, D.C. Andrew Ellington University of Texas at Austin Austin, Texas
In Vitro Selection of RNA Aptamers to a Small Molecule Target
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In Vitro Selection Using Modified or Unnatural Nucleotides
UNIT 9.6
In vitro selection is the process by which a pool of nucleic acids is enriched via iterative selection and amplification for those species that are capable of performing a particular task. Nucleic acids have been selected that bind to particular targets (aptamers), catalyze reactions (ribozymes or deoxyribozymes), or act as molecular switches (aptazymes). Instructions for carrying out in vitro selection experiments have been detailed elsewhere in this Chapter (e.g., UNITS 9.3 & 9.4). This unit augments these other units by describing how modified or unnatural nucleotides can potentially be incorporated into a selection. It is strongly recommended that the researcher be conversant with a “normal” in vitro selection experiment prior to attempting selections with modified nucleotides. A normal in vitro selection experiment is already fraught with problems and pitfalls, and the addition of modified nucleotides adds an extra level of difficulty. For simplicity, this unit focuses on the transcription and amplification of RNA pools; however, similar procedures can be used for DNA pools. CAUTION: When working with radioactivity, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following the guidelines provided by the local radiation safety officer. NOTE: Experiments involving RNA require careful precautions to prevent contamination and degradation by RNases (see APPENDIX 2A). The water used to make all solutions and buffers should be treated with diethylpyrocarbonate (DEPC; APPENDIX 2A). Use sterile, disposable plasticware where possible. STRATEGIC PLANNING When to Use Modified Nucleotides for In Vitro Selection As discussed in other units (UNITS 9.3 & 9.4), the desired outcome of a selection experiment should be determined in advance and the selection strategy should be designed accordingly. The decision to include modified nucleotides in a selection experiment and the selection of which ones to use should be based on how the nucleotides might benefit the selection. There are various advantages to using modified nucleotides, the chief one being to increase the stability of the selected nucleic acid to nuclease degradation. For example, the incorporation of pyrimidines modified at the 2′ position with amino or fluoro functional groups has been shown to drastically increase the stabilities of transcribed RNA molecules. Selections from nuclease-resistant pools have yielded aptamers against human neutrophil elastase (Lin et al., 1994), vascular endothelial growth factor (Green et al., 1995; Ruckman et al., 1998), basic fibroblast growth factor (Jellinek et al., 1995), human keratinocyte growth factor (Pagratis et al., 1997), and interferon γ (Kubik et al., 1997), as well as a sequence-specific phosphodiesterase (Beaudry et al., 2000). Aptamers selected from such pools have been shown to be stable in sera or urine for as much as 2 days (Pieken et al., 1991; Lin et al., 1994; Green et al., 1995). In addition, modified nucleotides potentially expand the chemical functionality of nucleic acids. Modified nucleotides have been included in selections for catalytic nucleic acids, and the resultant catalysts have been shown to be highly dependent upon the modifications for activity (Tarasow et al., 1997; Wiegand et al., 1997; Beaudry et al., 2000; Santoro et Contributed by Scott M. Knudsen, Michael P. Robertson, and Andrew D. Ellington Current Protocols in Nucleic Acid Chemistry (2001) 9.6.1-9.6.21 Copyright © 2001 by John Wiley & Sons, Inc.
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Table 9.6.1 Successful In Vitro Selections Using Modified Nucleotides
Modified nucleotide
Class
Target/functiona
Citation
5-(1-Pentynyl)-deoxyuracil N-6-Aminohexyl adenosine
Deoxy aptamer Ribozyme
Thrombin Ligation to its 5′ end
Latham et al. (1994) Teramoto et al. (2000)
5-Imidizole-uracil 5-Pyridylmethylcarboxamid-uracil
Ribozyme Ribozyme
Amide synthase Diels-Alderase
Wiegand et al. (1997) Tarasow et al. (1997)
C5-Imidizole-uracil analog (unnamed) Deoxyribozyme 8-[2-(4-Imidazolyl)ethylamino]deoxyadenosine Deoxyribozyme and 5-(3-aminoallyl)-deoxyuracil
Sequence directed RNase Santoro et al. (2000) RNase (internal rC with Perrin et al. (2001) no Mg2+)
5-(3′-aminopropynyl)-deoxyuracil aS NTPs (phosphorothioate linked RNA)
Deoxy aptamer Aptamer
ATP bFGF
Battersby et al. (1999) Jhaveri et al. (1998)
aS dNTPs (phosphorothioate linked DNA) 2′-Fluoro pyrimidines
Deoxy aptamer Aptamer
NF-IL6 VEGF(165)
King et al. (1998) Ruckman et al. (1998)
2′-Amino pyrimidines 2′-Amino pyrimidines
Aptamer Ribozyme
VPF/VEGF Trans cleavage of RNA
Green et al. (1995) Beaudry et al. (2000)
2′-Amino pyrimidines
Aptamer
Human neutrophil elastase
Lin et al. (1994)
2′-Amino pyrimidines 2′-Fluoro pyrimidines
Aptamer Aptamer
bFGF hKGF
Jellinek et al. (1995) Pagratis et al. (1997)
2′-Amino pyrimidines 2′-Amino pyrimidines
Aptamer Aptamer
hKGF IFN-γ
Pagratis et al. (1997) Kubik et al. (1997)
2′-Fluoro pyrimidines
Aptamer
IFN-γ
Kubik et al. (1997)
aAbbreviations: bFGF, basic fibroblast growth factor; hKGF, human keratinocyte growth factor; IFN-γ, interferon γ; NF-IL6, nuclear factor for human
interleukin 6; VEGF, vascular endothelial growth factor; VPF, vascular permeability factor.
al., 2000). Interestingly, no study has yet been published in which a selection with a modified nucleotide was done side-by-side with one without the modified nucleotide to compare the relative chemical advantage imparted by the modification. In this respect, it is interesting to note that while Tarasow et al. (1997) suggested that the inclusion of the modified nucleotide was the only reason they were able to select a Diels-Alder synthetase, Seelig et al. (1999) were later able to readily select such enzymes from an unmodified pool. To the extent that a modified nucleotide will be included to hopefully enhance catalytic functionality, the choice of modification should complement the desired function. For example, an imidizole ring (with a pKa near neutrality; Battersby et al., 1999) may be beneficial in a pool to be screened for catalysis of an acid/base reaction, while the incorporation of a thiolated residue (Jhaveri et al., 1998) could allow nucleic acids to participate in disulfide bond formation or rearrangement, reactions normally denied to them.
In Vitro Selection Using Modified or Unnatural Nucleotides
Overall, the most important considerations in deciding whether to use a modified nucleotide in a selection experiment are purely technical ones, i.e., can the modification be incorporated, and will it inhibit amplification of the nucleic acid pool? A number of modified nucleotides have been incorporated with varying degrees of success into selection experiments (Table 9.6.1). However, in the authors’ experience, at least in some instances (e.g., the incorporation of 5-Br-dUTP with Taq DNA polymerase), modified nucleotides decrease the replicability of sequences into which they are incorporated, and therefore there is an inherent and unintended selection in favor of those members of a population that have fewer modified nucleotides. While most of the modifications mentioned thus far retain an unmodified Watson-Crick pairing face, this need not always
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be the case. For example, Piccirilli et al. (1990) have shown that diaminopyrimidine and xanthosine can form a stable base pair. Lutz et al. (1998), using xanthosine in the template, showed that diamino pyrimidine triphosphate was incorporated by the 9°N-7 thermostable DNA polymerase (Southworth et al., 1996) in a sequence-specific fashion. However, the fidelity of this reaction was low, with only 45% incorporation of the correct nucleotide analog. This may mean that it would be difficult to carry out multiple rounds of selection with such modified nucleotides. While it is difficult to predict in advance which combinations of polymerases and modified nucleotides will work well together (also see Critical Parameters), there is at least one strategic consideration that can be taken into account in the design of selection experiments that include modified nucleotides. While the length and composition of a random sequence pool can vary according to the type of selection that is carried out (also see UNIT 9.2), some attention should be paid to the choice of constant sequence regions. RNA polymerases tend to have difficulty incorporating modified nucleotides prior to positions 8 to 10 following the transcription start site (Milligan and Uhlenbeck, 1989; Kujau et al., 1997), most likely because the attempted incorporation of modified nucleotides leads to increased abortive initiation. Thus, it is wise to design a pool that lacks or limits the incorporation of the modified base within this region. Inasmuch as the first 18 or more nucleotides of a transcribed pool typically remain constant to allow for secondstrand DNA synthesis, this limitation should not affect the overall diversity of the pool. Preparation and Use of Modified Nucleotides This unit presents procedures in two main sections: determination of the suitability of modified nucleotides for in vitro selection, followed by the use of modified nucleotides in in vitro selection. Since the main consideration is whether a given nucleotide analog can serve as a substrate and template for various enzymes and thus be selected, it is critical to determine this prior to beginning the selection. In the first set of procedures, transcription reactions are performed to show that full-length transcripts can be obtained from the modified nucleotides (see Basic Protocol 1). The prevalence of the modified nucleotide in the transcript pool is then assessed (see Basic Protocol 2). Since selected transcripts must be amplified, it is also important to verify that the modified RNAs are suitable templates for reverse transcriptase (see Basic Protocol 3). Finally, a mock selection should be performed and the products cloned and sequenced to determine the fidelity of replication (see Basic Protocol 4 for a discussion). Once it has been determined that a given modified nucleotide(s) is suitable for use in a selection, the starting pool can be constructed and selection can be performed. In the second set of protocols, double-stranded DNA that codes for the modified pool (as generated in UNIT 9.2) is first transcribed and the products are gel purified (see Basic Protocol 5). A general discussion of special considerations in designing an in vitro selection procedure for modified nucleotides is then presented (see Basic Protocol 6), followed by a procedure for amplification of the selection products (see Basic Protocol 7).
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DETERMINING WHETHER A MODIFIED NUCLEOTIDE IS SUITABLE FOR IN VITRO SELECTION BASIC PROTOCOL 1
Evaluation and Optimization of Transcription with a Modified Nucleotide In the case of RNA selections, the ability of a modified nucleotide to be incorporated into a transcript should be evaluated. This is best done by performing a transcription reaction in which the modified nucleotide completely substitutes for its natural counterpart (e.g., 2′-fluoro-cytidine in place of cytidine). A control reaction is performed in parallel on the same template with all natural nucleotides. The comparative success of these reactions can be determined by including α-32P-labeled nucleotides in the reaction mix, separating transcripts by denaturing polyacrylamide gel electrophoresis, and visualizing the fulllength transcripts with a phosphorimager or via autoradiography. Materials Double-stranded template DNA (UNIT 9.2) and randomly mutagenized DNA template (UNIT 9.4) 3000 Ci/mmol α-32P-labeled ATP (e.g., NEN Life Sciences) Transcription mix (see recipe) 2× denaturing stop dye (see recipe) 10% (w/v) denaturing polyacrylamide gel (see recipe, APPENDIX 3B), 0.75 mm thick 37° to 42° and 70°C water baths or heating blocks, or a programmable thermal cycler Gel blotting paper (Bio-Rad) Plastic wrap Gel dryer with vacuum (e.g., Bio-Rad) Phosphorimager screen Phosphorimager and image analysis software (e.g., ImageQuant) Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (PAGE; APPENDIX 3B) Perform transcription with modified and unmodified nucleotides 1. Combine the following in two separate tubes: ∼0.5 µg double-stranded DNA template (first tube) and randomly mutagenized DNA template (second tube) 0.5 µL 3000 Ci/mmol α-32P-labeled ATP transcription mix for a total reaction volume of 10 µL. Begin to incubate at 37° to 42°C. It is advisable to amplify a small portion of the pool that will actually be used in the selection for these test transcriptions. As this is a test reaction, do not commit the entire pool. Using high-yield transcription kits (e.g., Ambion or Epicentre Technologies) instead of a homemade mix can result in increased yields with shorter reaction times, and is advised wherever possible.
2. At several time points (e.g., 2, 4, 8, and 12 hr), vortex the tubes, briefly centrifuge to bring down the liquid, and transfer 1 µL to separate tubes containing 5 µL of 2× denaturing stop dye. In Vitro Selection Using Modified or Unnatural Nucleotides
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Vortexing and centrifuging the reaction ensures that each sample will contain a consistent portion of the reaction. The accumulation of condensation and the precipitation of pyrophosphate during transcription can sometimes lead to variations in reaction volumes or concentrations. If a commercial transcription kit is utilized, the time points can be much shorter (e.g., 0.25, 0.5, 1, and 2 hr).
3. Denature the time point samples by heating 3 min to 70°C and separate transcripts from aborted transcripts and unincorporated nucleotides on a 0.75-mm-thick, 10% denaturing polyacrylamide gel (APPENDIX 3B). The concentration of acrylamide can be varied to efficiently separate different length products, depending on their sizes (APPENDIX 3B). Prerunning the gel will heat it up and help to denature molecules that contain significant secondary structure. Nonetheless, pools can sometimes appear “fuzzy” on a gel due to the presence of molecules that differ slightly in length or that have different secondary structures.
Analyze transcription reactions 4. Remove one glass plate and place a sheet of gel blotting paper against the gel. Peel the gel off of the other glass plate (it will stick to the paper) and cover it with plastic wrap. Dry the gel under heat and vacuum using a commercial gel dryer. Drying the gel can be omitted, and a wet gel can be directly used for exposure. If this is done, leave the gel against one glass plate and carefully wrap with plastic wrap to avoid contaminating the phosphorimager screen and the exposure cassette with radiation.
5. Expose the dried gel to a phosphorimager screen for 1 hr. The exposure time may need to be increased or decreased, depending on the specific activity of the labeled nucleotide and the amount of transcript that is produced. Gels can also be exposed to X-ray film, although this makes quantitation somewhat more cumbersome. A standard laboratory densitometer can be used for quantitation.
6. Develop the phosphorimager screen and quantitate the relative amounts of RNA transcripts produced using image analysis software such as ImageQuant. 7. Compare the intensities of product bands for the transcription reaction that includes the modified nucleotide to those for transcription with the normal nucleotide to determine the relative efficiency of incorporation for the modified nucleotide under the conditions tested. 8. Plot the amount of product that accumulates versus time to determine the optimal time for further transcriptions. Optimize transcription reaction 9. Repeat steps 1 to 6 using different concentrations of the modified nucleotide. Take samples only at the optimal time determined in step 8. Since it is likely that the modified nucleotide will not work as well as the normal nucleotide because it binds to the polymerase more poorly, higher modified nucleotide concentrations may be necessary to enhance transcription. Therefore, care should be taken that the amount of modified nucleotide does not unnecessarily restrict the magnesium concentration in solution. If the total nucleotide concentration approaches that of magnesium, or the concentration of modified nucleotide exceeds the normal concentration used for unmodified (i.e., 2.5 mM), then additional magnesium should be added to the reaction. The initial sequence of the transcription product can also influence the incorporation of a modified nucleotide (see Strategic Planning for examples and references).
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10. Plot the amount of product (band intensity) versus modified nucleoside triphosphate concentration. This step is included both to maximize the yield of RNA produced with the new substrate and to efficiently use the modified nucleotide, which could potentially be expensive or difficult to produce. To conserve modified nucleotide, the concentration of modified nucleotide should be chosen at either a maximum in the above plot or at the lowest concentration at which near-maximal levels of RNA are produced. BASIC PROTOCOL 2
Confirmation of the Presence of Modified Nucleotides The appearance of a full-length product in the transcription reaction (see Basic Protocol 1) is encouraging, but does not indicate the level at which a modified nucleotide has been incorporated into a pool. For example, full-length RNAs could be members of the original population that incorporated the modified nucleotide at only a few positions, could have been synthesized via contaminating natural nucleotides, could have been synthesized via the misincorporation of a natural nucleotide (e.g., via G:U mismatches), or could be the result of some combination of these possibilities. In order to determine with certainty the level of modified nucleotide incorporation, transcription products should be isolated, fully digested, and separated by HPLC to identify the component nucleosides. This protocol assumes that one has already gel-purified and quantitated full-length transcription products, isolated as below. The RNA will be digested to mononucleotides using nuclease P1, and then treated with alkaline phosphatase to generate nucleosides. Separation of individual mononucleosides will be carried out by reversed-phase HPLC. Digestions will be compared to standards containing both modified and unmodified nucleosides. Sample HPLC data are given in Figure 9.6.1.
A
A 260 (au)
0.06
A = 22.0% C = 24.8% G = 29.0% U = 24.2%
CU 0.04
G
A = 22.5% C = 25.5% G = 31.4% U = <0.5% im-U = 22.5%
A
A = 23.0% C = 26.1% G = 30.2% U = <0.5% pm-U = 20.7%
0.06 C 0.04 0.02
0
0
im-U G
D 0.08
hm−U
0.06
A 260 (au)
A
0.08
0.02
B C G
0.04
A = 22.4% C = 25.1% A G = 29.4% U = <0.5% hm-U = 23.1%
0.08 0.06
0.02
0
0 5
10
15 20 25 Time (min)
30
35
C
0.04
0.02
0
In Vitro Selection Using Modified or Unnatural Nucleotides
C
A 260 (au)
0.08
A 260 (au)
A
G pm-U
0
5
10
15 20 25 Time (min)
30
35
Figure 9.6.1 HPLC elution profile of pool RNA that has been treated with nuclease P1 and alkaline phosphatase as described in Basic Protocol 2. (A) Unmodified pool, (B) pool containing 5-hydroxymethyluridine, (C) pool containing 5-imidizolemethyluridine, (D) pool containing 5-phenolmethyluridine (Robertson, 2001).
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Materials Transcribed RNA pools, modified and unmodified (see Basic Protocol 1) Nuclease P1 digestion mix (see recipe) Alkaline phosphatase reaction mix (see recipe) HPLC mobile phase solution: 5% (v/v) methanol in 0.1 M sodium phosphate, pH 6.0 (APPENDIX 2A) 37° and 50°C water baths High-performance liquid chromatograph (HPLC) with reversed-phase C18 column (5-µm, 250 × 4.5-mm; Alltech Associates) and UV detector 1. Combine 200 pmol of each RNA pool (modified and unmodified) in separate reactions with nuclease P1 digestion mix for a total reaction volume of 5 µL and incubate 1.5 hr at 37°C. This step will digest the nucleic acid into its component nucleotides. Nuclease P1 will digest both RNA and DNA. It is possible that nuclease P1 will not digest after a specific modified nucleotide. If incomplete digestion is seen compared to an unmodified sample digestion, this in itself is convincing evidence of modified nucleotide incorporation. The unmodified transcription reaction should be run separately through the same column using the same conditions to provide a series of standard retention times for comparison. This control can also be used to obtain the extinction coefficient of the modified nucleoside, if it is not otherwise known.
2. Combine each entire nuclease P1 digestion reaction with alkaline phosphatase reaction mix in a final volume of 25 µL. Incubate 1.5 hr at 37°C, followed by 1 hr at 50°C. 3. Inject 2.5 µL of one reaction onto a reversed-phase C18 HPLC column and separate using a mobile phase flow rate of 1 mL/min and detection at 260 nm. Repeat with the other reaction. If the pool initially had an equimolar mix of nucleotides, then the digested RNA should contain ∼25% of the modified nucleoside and no peak corresponding to the replaced nucleoside. Of course, the actual composition of pools can vary greatly, depending on the synthesis of the pool (e.g., see UNIT 9.2). Therefore, the composition of the original pool should first be determined either by sequencing individual clones (e.g., see CPMB, Chapter 7) or by first digesting and separating a transcription product made solely from natural nucleotides. Nucleotides are detected at 260 nm as this is close to the absorbance maxima for their aromatic bases. Nucleotides with heavily modified bases that disrupt their aromaticity may absorb at different maxima; however, A260 should still be used because the RNA will likely be mostly unmodified.
Evaluation of Modified RNA as a Template for Reverse Transcriptase During each round of in vitro selection, the selected RNAs must be amplified. Therefore, it is necessary to determine if an RNA transcript containing modified bases will serve as a suitable template for reverse transcription, the first step of the amplification process. A control reaction can be performed on the same template with all natural nucleotides. The relative success of the reverse transcription reactions can again be visualized by incorporation of α-32P-labeled nucleotides, gel electrophoresis, and analysis on a phosphorimager.
BASIC PROTOCOL 3
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Materials Transcribed RNA pools, modified and unmodified (prepared as in Basic Protocol 1 but without radiolabeled ATP) 3000 Ci/mmol α-32P-labeled dATP (e.g., NEN Life Sciences) RT reaction mix (see recipe) AMV reverse transcriptase (e.g., USB) 2× denaturing stop dye (see recipe) 10% (w/v) denaturing polyacrylamide gel (see recipe; APPENDIX 3B), 0.75 mm thick 42° and 70°C water baths or heating blocks, or a programmable thermal cycler Gel blotting paper (Bio-Rad) Plastic wrap Gel dryer with heat and vacuum (e.g., Bio-Rad) Phosphorimager screen Phosphorimager Image analysis software (e.g., ImageQuant) Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (PAGE; APPENDIX 3B) Optimize RT reaction 1. Combine the following in two separate tubes: 10 pmol transcribed RNA pool (modified in one tube, unmodified in the other) 0.5 µL α-32P-labeled dATP RT reaction mix for a total of 10 µL. Incubate 2 min at 42°C. Do not use radiolabeled RNA as the input. It may be difficult to differentiate product from input if both are labeled.
2. Add 10 U AMV reverse transcriptase per 10 µL RT reaction mix and incubate 50 min at 42°C. The success of the reverse transcriptase reaction will depend primarily on the reverse transcriptase that is used. While reverse transcriptases in general tend to be somewhat forgiving with respect to template chemistry (e.g., they can use both RNA and DNA as templates), different reverse transcriptases may have distinct abilities to utilize diverse modified bases. If one reverse transcriptase does not prove efficient in copying a particular RNA template, then another one should be used, or they should all be compared at the outset.
3. Remove 2 µL and add it to 5 µL of 2× denaturing stop dye. 4. Denature by heating 3 min to 70°C and run on a 0.75-mm-thick 10% denaturing polyacrylamide gel (APPENDIX 3B). A sample of radiolabeled RNA, as produced in the test transcriptions, can be used as a convenient size standard. RNA runs slightly slower on a gel than DNA of the same size. The concentration of acrylamide can be varied to efficiently separate different products (APPENDIX 3B).
Analyze RT reaction 5. Remove one glass plate and place a sheet of gel blotting paper against the gel. Peel the gel off of the other glass plate (it will stick to the paper) and cover with plastic wrap. Dry the gel in a gel dryer under heat and vacuum. In Vitro Selection Using Modified or Unnatural Nucleotides
Drying the gel can be omitted, and a wet gel can be directly exposed to the phosphorimager plate. If this is done, carefully wrap the gel with plastic wrap to avoid contaminating the screen and exposure cassette with radiation.
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6. Expose the dried gel to a phosphorimager screen for 1 hr. The exposure time may need to be increased or decreased, depending on the specific activity of the labeled nucleotide and the amount of transcript produced. Alternatively, gels can be exposed to X-ray film, and the film developed to visualize bands.
7. Develop the phosphorimager screen and view the output with image analysis software such as ImageQuant. Full-length cDNA should have been produced from both modified and unmodified RNA templates. It is possible that smaller, discrete bands will be observed. These may be due to the failure of reverse transcriptase to read past secondary structures in the RNA. It is unlikely that discrete bands will be observed with a pool that contains many different RNA molecules, unless the secondary structures are in the constant regions. However, as a selection progresses, species that give rise to discrete stops may become fixed in a population. In this instance, the reverse transcription should be carried out at a slightly higher temperature. In extreme cases, a thermophilic DNA polymerase, Tth polymerase, has been shown to have significant reverse transcriptase activity, and can potentially be used to copy recalcitrant RNA molecules. Of course, it should be realized that even though strong stops cannot be seen early in selection, certain RNA species may drop out of a population because they cannot be copied by reverse transcriptase.
Confirmation of the Fidelity of Replication If Basic Protocols 1, 2, and 3 give successful results, then it is likely that the random sequence pool can make it through the preparative steps leading to the selection. However, it is possible that the incorporation of modified nucleotides may alter the dynamics of selection experiments in one of two ways: first, modified nucleotides may in and of themselves be extremely mutagenic; second, it may be difficult to either make full-length transcripts that contain modified nucleotides or to fully copy transcripts containing modified nucleotides, in which case there will be an unseen selection against transcripts that contain modified nucleotides over the course of several cycles. In order to guard against these possibilities, it may be worthwhile to carry out at least one round of mock selection and to clone and sequence the products.
BASIC PROTOCOL 4
Cloning and sequencing PCR products is beyond the scope of this protocol, but can be found, for example, in CPMB Chapter 7. However, the protocols in the next section can be used to simulate a round of selection (omitting the selective steps). Two different types of selections can be carried out, depending on whether the mutagenic or replicative potential of modified nucleotides is being tested. To test the mutagenic potential, a single sequence cloned from the pool should be subjected to a round of selection. Following the selection, ∼30 clones should be sequenced and the number of mutations counted. To test the replicative potential, the pool should be subjected to a round of selection. Before and after the selection, ∼30 clones should be sequenced and the base composition of the pool should be analyzed. If the base composition following selection is highly skewed relative to before selection, then selection against a given residue may prove to be a problem. It should be recalled that even small skewing can prove to be significant over many rounds of selection. For example, if a modified cytidine is present at 25% of the random sequence positions in a starting pool, but is present at only 22.5% of the positions following one round of selection, after 10 rounds of selection its frequency may fall to 0.25[(0.225/0.25)10] = 0.087, or <9% of the random sequence positions in a pool.
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UTILIZING MODIFIED NUCLEOTIDES IN IN VITRO SELECTION BASIC PROTOCOL 5
Preparation of the RNA Pool The pool will be transcribed using the conditions determined during the trial experiments. The RNA will be gel purified, eluted from the gel, and ethanol precipitated. Alternatively, very large amounts of RNA can be purified by size-exclusion chromatography (e.g., Qiagen RNA kit). Following quantitation, it will be ready to use in the first round of selection. Transcription reactions in later rounds will be performed in much the same way, except that smaller amounts of DNA template may be used in later rounds as the diversity of the pool becomes constricted. Materials Double-stranded, modified template DNA (UNIT 9.2) Transcription mix (see recipe) RNase-free DNase (e.g., RQ1 DNase, Promega) 2× denaturing stop dye (see recipe) 10% (w/v) denaturing polyacrylamide gel (see recipe, APPENDIX 3B), 1.5 mm thick 0.3 M NaCl (APPENDIX 2A) 70% and 100% (v/v) ethanol TE buffer, pH 7.5 to 8.0 (APPENDIX 2A) 37° to 42° and 70°C water baths or heating blocks, or a programmable thermal cycler Fluorescent thin-layer chromatography (TLC) plate (e.g., Aldrich), wrapped in plastic wrap 0.45-µm syringe filter with disposable syringe Spectrophotometer Additional reagent and equipment for denaturing polyacrylamide gel electrophoresis (APPENDIX 3B) Prepare RNA 1. Add ∼1 µg double-stranded, modified template DNA to 20 µL of transcription mix. Incubate at 37° to 42°C for the predetermined optimal time (see Basic Protocol 1). If the pool contains >1 ìg of template DNA, scale up the transcription reaction accordingly. The complexity of the pool for the initial round of selection can be determined as described in UNIT 9.3. At least 1013 different sequences should be used for the first round of selection. By increasing the complexity of the population to ≥1015 sequences, it may be possible to isolate rare binding or catalytic species (e.g., see Bartel and Szostak, 1993). In later rounds of selection, as the population is winnowed, there will be many copies of each selected species in the pool, and smaller amounts of DNA template (∼1 ìg/round) and RNA (∼10 ìg/round) can be utilized. Although it is not necessary to produce radiolabeled RNA at this stage, it may be useful to have a labeled sample of the original pool RNA. This can be accomplished by adding α-32P-labeled ATP to the transcription reaction mix.
2. Remove template DNA by adding 5 to 10 U RNase-free DNase and incubating 15 min at 37°C. Stop the reaction by adding an equal volume of 2× denaturing stop dye. The DNase treatment is necessary to ensure that selected RNA molecules, rather than DNA carryover from a previous amplification reaction, is preferentially amplified.
In Vitro Selection Using Modified or Unnatural Nucleotides
In some cases, amplification artifacts can overrun selection experiments (e.g., see Marshall and Ellington, 1999). In these instances, it may prove worthwhile to include dUTP in PCR amplification reactions, and to eliminate unwanted templates by the addition of uracil N-glycosylase.
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3. Denature the samples 3 min at 70°C, and load the entire reaction on a 1.5-mm-thick, 10% denaturing polyacrylamide gel (APPENDIX 3B). No more than 150 ìg of RNA should be loaded per 1-cm-wide lane. For a thinner gel, this amount should be proportionally decreased. This value corresponds to ∼50 ìL of a very efficient transcription reaction. Since later rounds need not contain as much template (see step 1), a 0.75-mm-thick gel may suffice to purify transcripts.
4. Visualize the product band by UV shadowing against a fluorescent TLC plate covered in plastic wrap. Excise the product band with a clean razor blade. Use fresh plastic wrap and razor blades every time to avoid contaminating the pool. It is important to only cut out the product band, as bands that migrate lower on the gel are likely to be aborted transcripts or template DNA that has not been completely digested. As some modified nucleotides may be especially sensitive to UV light, it is best to expose the pool for as short a time as necessary to define the bands. It may prove useful to lay an additional sheet of plastic wrap over the gel and to draw a line around the product band before excision.
Elute RNA 5. Submerge the gel slice in ∼400 µL of 0.3 M NaCl per cm2 of gel. Elute RNA overnight at 37°C with agitation. Increased recovery of RNA can be achieved by cutting the gel slice into smaller pieces, or by pulverizing it with a clean glass stir rod. Elution at higher temperatures can speed recovery time, but may lead to increased fragmentation of the pool RNA.
6. Pipet the supernatant off of the gel slice and filter it through a 0.45-µm filter attached to a syringe to remove small pieces of acrylamide. Add 2.5 vol of 100% ethanol and incubate 30 min at −20°C or 10 min at −80°C. Filtering out small pieces of acrylamide is especially important if the gel slice was pulverized. These pieces will precipitate with the RNA and will cause errors in determining the final concentration of RNA, as the presence of acrylamide can interfere with the determination of the sample’s UV absorbance.
7. Place the precipitation reaction in a 1.5-mL microcentrifuge tube and microcentrifuge 20 to 30 min at maximum speed, 4°C. For larger-scale precipitations, such as purification of the original pool, 10- to 12-mL Sarstedt tubes can be used, and the reaction centrifuged 40 min at 13,000 × g, 4°C.
8. Carefully decant the supernatant without disturbing the pellet. Wash the pellet with 70% ethanol and dry the pellet. If the pellet is dislodged during the wash step, microcentrifuge an additional 5 min in the 70% ethanol to reattach it to the side of the tube. The pellet can be air dried or dried under vacuum. When air drying, cover the tube with Parafilm and poke holes in the Parafilm with a clean needle in order to guard against contamination while allowing evaporation to proceed.
9. Resuspend the RNA pellet in 25 µL TE buffer, pH 7.5 to 8.0. If RNA is to be used in a reaction whose volume is small relative to the volume of RNA added, it may be necessary to supplement the reaction with magnesium to avoid complete chelation by EDTA. Alternatively, RNA can be resuspended in sterile water; however, this renders the sample more susceptible to RNase degradation.
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10. Determine the amount of RNA recovered by determining the absorbance of the solution at 260 nm. The concentration of an RNA solution = (A260 × dilution factor)/(10,625 × n) M for a 1-cm cell, where n is the length of the RNA. A convenient dilution factor is 500, corresponding to 2 ìL of RNA solution diluted in 1 mL water. The extinction coefficient 10,625 M−1cm−1 is the average of the extinction coefficients for the four nucleobases, C = 7100, U = 8400, G = 12,000, and A = 15,000 M cm−1. However, it should be noted that modification could drastically affect absorbance at 260 nm. For example, if an aromatic group is appended to a base, then the extinction coefficient of the pool would be higher than expected. In such a case, it would be wise to use the extinction coefficient for the modified base to create a new average. The RNA is now ready for a round of selection (see Basic Protocol 6). Repeated rounds of selection are performed by generating a new (selected) DNA template (see Basic Protocol 7) and repeating this protocol to generate the new (selected) RNA pool. BASIC PROTOCOL 6
Isolation of a Functionally Enriched Pool by In Vitro Selection The procedure used to isolate active species from a pool is highly dependent on what function is being selected, i.e., binding versus catalysis, binding proteins versus binding small organics. Ideally, selections should be designed in which only those individuals that are capable of performing the desired function are isolated from the remainder of the pool. This is best done by a series of negative and positive selections (also see UNITS 9.2 & 9.3). A positive selection is performed with the intention of isolating active sequences. For example, if binding a small organic molecule is the goal, the pool may be passed over a column that presents the desired target, and binding species eluted with the target in solution (i.e., affinity chromatography). Most methods of isolating functional sequences can also isolate nonfunctional sequences for other reasons, such as binding to the matrix rather than to the target. For this reason, a negative selection generally contains all elements of the positive selection with the exception of the specific target or substrate. Negative selection steps, in general, reduce the number of sequences that artifactually survive the positive selection steps. As rounds progress, both negative and positive selections are made more stringent by adjusting reaction concentrations or times. More detailed discussions of the design of selection experiments to yield binding or catalytic species can be found in UNITS 9.3 & 9.4, respectively. Conceptually, the design of a selection using modified nucleotides is not different from any other selection. However, special consideration should be given to the prospect that a modified nucleotide can influence the relative number of molecules that will survive a negative selection step. For example, a more hydrophobic nucleotide might be bound more readily to a modified cellulose filter even in the absence of a protein target. Moreover, as previously discussed, it is possible that discrimination against modified nucleotides at any point in a replicative cycle (e.g., failure to be incorporated by RNA polymerse, failure to be utilized as a template by reverse transcriptase, failure to be utilized as a substrate or a template by Taq polymerase) could lead to artifactual selection of those sequences that replicate best with few or no modified nucleotides, rather than to the actual selection of binding or catalytic species containing modified nucleotides.
In Vitro Selection Using Modified or Unnatural Nucleotides
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Current Protocols in Nucleic Acid Chemistry
Amplification of a Functionally Enriched Pool Once an RNA pool undergoes a selection function, the selected pool, presumably enriched in functional molecules, must be amplified. In the case of an RNA selection, this is best done by a combination of reverse transcription, PCR, and in vitro transcription. An unintended benefit of this convoluted process is that it makes it more difficult for molecular parasites to accumulate, as there are few molecular species that can be preferentially amplified by reverse transcriptase, Taq polymerase, and T7 RNA polymerase (Marshall and Ellington, 1999).
BASIC PROTOCOL 7
Materials Selected RNA pool (see Basic Protocol 6) RT reaction mix (see recipe) AMV reverse transcriptase (e.g., USB) PCR reaction mix (see recipe) Taq DNA polymerase (e.g., Promega) 6× nondenaturing dye solution (see recipe) 4% (w/v) agarose gel: e.g., NuSieve in tris/borate/EDTA (TBE) electrophoresis buffer (available precast or see, e.g., CPMB UNIT 2.6) Size standard 5 M NaCl (APPENDIX 2A) 100% and 70% (v/v) ethanol TE buffer, pH 7.5 to 8.0 (APPENDIX 2A) 10 mg/ml ethidium bromide solution (APPENDIX 2A) 42°C water bath or heating block Programmable thermal cycler (e.g., MJ Research) Additional reagents and equipment for agarose gel electrophoresis (e.g., CPMB UNIT 2.5) Perform reverse transcription 1. Add a sample of a selected RNA pool to RT reaction mix to a total volume of 10 µL. Incubate 2 min at 42°C. 2. Add 10 U AMV reverse transcriptase per 10 µL reaction mix. Incubate 50 min at 42°C. In some instances, RNA samples may be attached to beads, such as streptavidin-coated agarose beads. If so, the RNA can be directly amplified from the beads, but it is helpful to first wash the support with 1× RT buffer. The RT reaction may also need to be scaled up to ensure sufficient volume to completely cover the support.
Perform amplification 3. Seed 100 µL PCR reaction mix with 2 µL of the RT reaction mix (or 1⁄5 the RT reaction volume). Hot start the reaction with 2.5 U Taq DNA polymerase. 8 to 10 cycles:
30 sec 94°C (denaturation) 30 sec 55°C (annealing) 60 sec 72°C (extension).
The annealing temperature is dependent on primer composition and may need to be lowered. Adding polymerase at high temperatures (or “hot starting”) helps to prevent mispriming events that can lead to the accumulation of artifacts. The easiest way to hot start a reaction is to wait until the initial 94°C step to add Taq DNA polymerase. If the RT reaction was performed on beads, these should also be added to the PCR reaction. It is best to limit the bead volume to no more than 20 ìL in a 100-mL PCR reaction.
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4. Withdraw 5 µL of the reaction and mix with 1 µL of 6× nondenaturing dye solution. 5. Run on a 4% agarose gel, preferably next to a size standard. If only a faint band of the proper size is visible, run the reaction for one to two more cycles. If no band is visible, continue the amplification reaction for four more cycles and check again. Continue to amplify and monitor until a band is visible on the gel. Following the reaction in this manner helps to avoid overamplification, which can lead to accumulation of larger-than-normal artifactual products. This somewhat laborious method of monitoring reactions can be greatly simplified if a laboratory has access to a real-time PCR machine and has designed appropriate primer pairs in advance. Although the selected RNA pool can be amplified by a combined reverse transcription PCR reaction, the authors have found that separate reactions yield more consistent results. However, if effort is expended on optimizing a combined RT-PCR reaction, then the time required for a round of selection can be proportionately shortened.
6. Run four identical polymerase chain reactions with the remainder of the RT reaction. Use the optimal number of thermal cycles determined in step 5. Purify template DNA 7. Combine the five amplification reactions. Add 30 µL of 5 M NaCl to a final concentration of 0.3 M. Add 2.5 volumes of 100% ethanol. Incubate 30 min at −20°C or 10 min at −80°C. If a support such as agarose beads was included in the reaction, filter these out with a Millipore filter column prior to precipitation.
8. Place the precipitation reaction in a 1.5-mL microcentrifuge tube and microcentrifuge 20 to 30 min at maximum speed, 4°C. 9. Carefully decant the supernatant without disturbing the pellet. Wash the pellet with 70% ethanol and dry the pellet. If the pellet is dislodged during the wash step, microcentrifuge an additional 5 min in 70% ethanol to reattach it to the side of the tube. The pellet can be air dried or dried under vacuum. Cover the tube with Parafilm and poke holes in the Parafilm with a clean needle in order to guard against contamination while allowing evaporation to proceed.
10. Resuspend the pellet in 25 µL TE buffer, pH 7.5 to 8.0. 11. Dilute 1 µL template DNA sample in 4 µL water and add 1 µL of 6× nondenaturing dye solution. 12. Run the sample on an agaraose gel next to a ladder or size standard of known concentration and stain with 10 µg/mL ethidium bromide solution. Visualize the bands on a UV light table. Estimate the concentration of the original template DNA stock. This DNA will be used to transcribe RNA for the next round of selection (see Basic Protocol 5). Laboratories that have access to a digital camera with quantitation software can more accurately estimate the amount of DNA in a given sample.
In Vitro Selection Using Modified or Unnatural Nucleotides
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REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Alkaline phosphatase reaction mix 50 mM Tris⋅Cl, pH 8.5 (APPENDIX 2A) 0.1 mM EDTA 2 U alkaline phosphatase per 25 µL reaction Store buffer without enzyme indefinitely at −20°C. Add enzyme fresh. Denaturing polyacrylamide gel, 10% (w/v) 1× TBE electrophoresis buffer (APPENDIX 2A) containing: 10% (w/v) acrylamide 0.5% (w/v) bisacrylamide 7 M urea Prepare fresh See APPENDIX 3B for full details on pouring and running gels.
Denaturing stop dye, 2× Deionized formamide containing: 0.1% (w/v) bromphenol blue 50 mM EDTA Store indefinitely at −20°C. Daily-use aliquot may be stored up to 1 month at room temperature. Nondenaturing dye, 6× 60% (v/v) glycerol 0.1% (w/v) bromophenol blue Store indefinitely at −20°C. Daily-use aliquot may be stored up to 1 month at room temperature. Nuclease P1 digestion mix 15 mM sodium acetate, pH 5.2 (APPENDIX 2A) 0.1 mM zinc sulfate 1 U P1 nuclease per 5 µL reaction Store buffer indefinitely at −20°C. Add enzyme fresh. PCR reaction mix 10 mM Tris⋅Cl, pH 8.4 (APPENDIX 2A) 50 mM KCl (APPENDIX 2A) 1.5 mM MgCl2 (APPENDIX 2A) 0.2 mM each dNTP 0.5 µM each primer 2.5 U/100 µL reaction Taq DNA polymerase The Tris⋅Cl, KCl, and MgCl2 can be combined at ten times the indicated concentrations to make a 10× PCR buffer and stored indefinitely at −20°C. The remainder of the reagents should be added fresh.
RT reaction mix 100 mM Tris⋅Cl, pH 8.0 (APPENDIX 2A) 50 mM KCl (APPENDIX 2A) 10 mM MgCl2 (APPENDIX 2A) 10 mM dithiothreitol (DTT) 0.4 mM each dNTP 5 µM reverse (3′) primer continued
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The Tris⋅Cl, KCl, and MgCl2 can be combined at ten times the indicated concentrations to make a 10× RT buffer and stored indefinitely at −20°C. The remainder of the reagents should be added fresh.
Transcription mix 40 mM Tris⋅Cl, pH 7.9 (APPENDIX 2A) 26 mM MgCl2 (APPENDIX 2A) 0.01% (v/v) Triton X-100 2.5 mM spermidine trihydrochloride 5 mM dithiothreitol (DTT) 2.5 mM each NTP 20 U/20 µL reaction RNasin (Promega) 100 U/20 µL reaction T7 RNA polymerase The efficiency of transcription will be one of the variables that will require the greatest optimization. It is again suggested that a commercial transcription kit may give the greatest yields and best results, especially with modified nucleotides.
COMMENTARY Background Information
In Vitro Selection Using Modified or Unnatural Nucleotides
Nucleic acid chemistry is constrained by the relatively limited chemistry of the five canonical nucleotides. For example, nucleic acids, especially RNA, tend to be much more chemically labile than proteins. In the case of RNA, stability is impaired by the presence of a 2′-hydroxyl group that can serve as a general base poised to attack the phosphodiester bond. To offset this problem, Eckstein and co-workers developed 2′-modified nucleotides that were resistant to chemical and enzymatic hydrolysis. Hammerhead ribozymes modified with 2′amino or 2′-fluoro groups were shown to retain activity, and had a 1000-fold increase in stability in rabbit serum (Pieken et al., 1991). While the riboside triphosphates could be incorporated into RNA, the kcat/KM values for incorporation of 2′-modified ribonucleotides were substantially higher than for unmodified nucleotides, with a preference order of 2′-OH > 2′-NH2 > 2′-F > 2′-H (Brieba and Sousa, 2000). In order to improve incorporation, some groups have begun to engineer or evolve polymerases that can incorporate modified nucleotides. For example, a single mutation in T7 RNA polymerase (Y639F) reduces discrimination at the 2′ position, and allows more efficient incorporation of deoxyribonucleotides into RNA (Kostyuk et al., 1995). It was subsequently shown that this mutation also allows the incorporation of NTPs with fluoro, amino, and thio groups at their 2′ positions. The Y639F mutant of T7 RNA polymerase has been shown to have up to 24-fold greater specificity for incorporation of 2′-modified NTPs, with a preference order of 2′-OH > 2′-F > 2′-H > 2′-NH2 (Brieba
and Sousa, 2000). Several 2′-modified ribonucleoside triphosphates are now commercially available (Jena BioScience, Trilink); Y639F T7 polymerase is available from Epicentre Technologies. The replacement of the ribose 2′-OH group with other chemical moieties interferes with the primary mechanism for nuclease cleavage of RNA, attack of the 2′-hydroxyl on the bridging phosphate. However, substitutions on the backbone, such as replacing the phosphate with a phosphorothioate, have also been shown to increase oligonucleotide stability in the presence of nucleases (Zon and Geiser, 1991). An additional benefit is that phosphorothioate nucleotides have been shown to be incorporated into an elongating transcript by T7 polymerase with little or no increase in KM (Griffiths et al., 1987). While DNA is not as vulnerable to hydrolysis as RNA, it is nonetheless susceptible to cleavage by a variety of deoxyribonucleases and phosphodiesterases. The stability of DNA can also be increased by the incorporation of phosphorothioate nucleotides, and these can be readily incorporated by Taq DNA polymerase (Nakamaye et al., 1988). These modified ribonucleoside and deoxyribonucleoside triphosphates are also commercially available (NEN Life Sciences, Sigma, Life Technologies). Nucleic acid selection experiments have generated a wide variety of binding species (aptamers) and catalysts (ribozymes and deoxyribozymes). However, nucleic acids are by and large not as functional as proteins: aptamers bind their ligands less well than antibodies, for the most part, while ribozymes are slower than
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protein enzymes, for the most part. While it is possible that these limitations are a function of the relative newness of aptamer and catalyst selections relative to a well-established field like protein engineering, it is also possible that functional nucleic acids are inherently limited by the diversity of the genetic alphabet. The binding interactions and chemical reactions performed by a nucleic acid biopolymer may be constrained by the functional groups they contain. The ability to expand the functional groups available to a DNA or RNA polymer through the incorporation of modified nucleotides could potentially open up realms of chemistry and binding interactions that were previously inaccessible. Researchers are currently trying to assess this hypothesis and demonstrate the utility of modified bases in functional nucleic acids. For example, the first RNA capable of catalyzing the formation of carbon-carbon bonds utilized 5-pyridylmethylcarboxamid-UTP in place of UTP (Tarasow et al., 1997). When the most active isolate from this selection was transcribed with unmodified UTP, it was inactive. Similar results were obtained for a ribozyme that catalyzed amide bond formation (Wiegand et al., 1997) whose activity was dependent on the incorporation of 5-imidizole-UTP. Additionally, two sequence-specific RNase deoxyribozymes have been selected that were dependent on the incorporation of a 5-imidizole-TTP (Santoro et al., 2000), and both 8-2-(4-imidazolyl)ethylamino-2′-dATP and 5-(3-aminoallyl)-2′-dUTP (Perrin et al., 2001). However, none of the sequences, motifs, or activities found in these selection experiments was directly compared with ribozymes that contained canonical nucleotides and that were sieved from the same pool using the same selection conditions. Therefore, at present time, it is unclear whether modified nucleotides truly improve RNA catalysis. Surprisingly, though, there are at least a few counterexamples that would suggest that modified nucleotides do not greatly contribute to binding or catalysis relative to natural nucleotides. Santoro and colleagues (Santoro and Joyce, 1997; Santoro et al., 2000) selected deoxyribozymes with RNA hydrolysis activity from different aliquots of the same, unamplified random sequence pool. The selection performed with natural nucleotides produced a much faster catalyst. Ultimately, it is unknown whether this indicates the superiority of natural nucleotides for this pool and this function, or whether the fraction of the original pool used
for the selection of the natural catalyst just contained a “jackpot” sequence. An additional example of a catalytic activity selected for with both modified and unmodified pools is the Diels-Alder catalyzing ribozymes (Tarasow et al., 1997; Seelig and Jaschke, 1999, respectively). The modified selection yielded a catalyst with a kcat/KM of ∼4 M−1sec−1, while the unmodified selection yielded a catalyst with a kcat/KM of 167 M−1sec−1. However, these selections were performed by different research groups with different pools, and thus are not directly comparable. For a final example, see Critical Parameters, discussion of multiple approaches.
Critical Parameters Optimizing preparative reactions A good portion of this protocol has been devoted to testing reactions and optimizing conditions. Although this process will likely take several days, it is time wisely spent. The first round of selection is by far the most important, as this is the round when selection conditions will query the greatest number of possible answers. Concomitantly, creating the largest possible number of unique sequences (1013 to 1015) for the first round of selection is a critical task. However, because creating a large library requires a large amount of effort and large volumes of relatively expensive reagents, initial optimization and practice should always be performed on a small scale. Similarly, it is expected that only a small number of the input molecules will survive any given round; therefore, it is essential that these successful sequences be efficiently carried to the next round. Inefficient reverse transcription or amplification reactions may lead to a selection for molecules that can be efficiently replicated (amplification artifacts) rather than to molecules that are highly functional. Archiving reactions In vitro selection experiments are particularly grueling because the ultimate outcome, the successful isolation of binding species or catalysts, may not be known for days, weeks, or even months. This is especially true when working with modified nucleotides, because of the strong possibility that one or more amplification reactions or selection steps will not work at some point during the course of the selection. Therefore, it is desirable to keep an archived copy of each round. After the initial round, there should be multiple copies of each winning
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sequence present in the selected population. Thus, it is not necessary to use all of the RNA, cDNA, or double-stranded DNA template in a given selection or amplification step. Rather, some portion of the pool should be saved at any convenient point (e.g., as a double-stranded PCR product). In the event an unsuccessful round of selection should occur, it will not be necessary to repeat the entire experiment. Rather, selection can continue from the last successful round, with a minimal loss of time and effort.
In Vitro Selection Using Modified or Unnatural Nucleotides
Understanding modified nucleotide chemistry The incorporation of additional functional groups into the context of an RNA backbone is expected to increase the diversity of its available interactions, both with itself and with its desired substrates. Thiol groups, for example, can participate directly in catalysis as nucleophiles. Additionally, disulfide bonds could be formed intramolecularly between thiols. This may add to the structural diversity of nucleic acids, normally limited to hydrogen bonding, salt bridges with metals, and stacking interactions. Charged groups, such as a lysine-like side chain, could potentially add to the structural repertoire of nucleic acids by allowing the formation of electrostatic interactions and salt bridges. The inclusion of chemical moieties with a pKa closer to neutrality, such as an imidizole group, is also expected to benefit nucleic acid chemistry. Natural nucleotide bases contain no functional groups with unperturbed pKa values between 3.5 and 9.2, inherently limiting the proton “push-pull” chemistry found in so many protein enzymes. The introduction of these and other functional groups can also potentially increase the abilities of nucleic acids to bind metals. While attempts to associate nucleotide functionality with binding or catalytic properties are at best still guesswork, it is nonetheless true that failing to appreciate the chemical properties of modified nucleotides can potentially adversely affect how a selection experiment proceeds. For example, the inclusion of thiolated nucleotides can potentially lead to the unwanted formation of disulfide-bonded products if care is not taken to include a reducing agent in enzyme and/or selection reactions. For example, deoxyribozyme ligases that form an unnatural internucleotide linkage between a 5′-iodinated pool and an oligonucleotide substrate with a 3′ phosphorothioate have been selected from random sequence pools (Levy and Ellington, 2001a,b).
Unless reducing agents are kept in the selection reaction, the oligonucleotide substrates will dimerize, reducing their effective concentration. Similarly, the inclusion of modified nucleotides that have altered pKas could very easily change the pH of a concentrated stock solution, and care should be taken to make sure that all such stocks are at the desired pH and appropriately buffered. The accidental alteration of pH in enzyme or selection reactions can of course lead to decreases in product yield. Finally, the inclusion of modified nucleotides that have new metal binding or chelating properties may alter the available metal concentrations in an enzyme or selection reaction. In particular, the chelation of magnesium can lead to large changes in the efficiency of product formation by many different polymerases. Multiple approaches As has been apparent throughout, the in vitro selection of functional nucleic acids that contain modified nucleotides is still in its infancy, and thus there are few hard and fast rules regarding what will and will not work. Because of this, researchers need to be somewhat versatile in their approach to selection experiments involving modified nucleotides. If a given approach does not work, this does not mean that the selection experiment inherently has no chance of working, but instead indicates that it may be necessary to alter one or more parameters. In particular, to improve the incorporation of modified nucleotides into a pool there are three variables that should be adjusted as needed. First, the buffer conditions for incorporation can be adjusted. Padilla and Sousa (1999) have systematically investigated several buffer conditions that aid the incorporation of nucleotides modified specifically at the 2′ position. These authors find better incorporation upon supplementing a transcription reaction with 0.5 mM MnCl2, 1 U/µL pyrophosphatase, and either 8 mM spermidine for plasmid templates or 8 mM spermine for short DNA templates. Second, different polymerases clearly have different potentialities for the incorporation of modified nucleotides. For example, the Benner group has tested several thermostable DNA polymerases for their ability to incorporate a variety of modified nucleotides (Lutz et al., 1998, 1999). Finally, the sequence of the pool itself can have a surprisingly large effect on selection experiments. The authors originally selected a relatively fast ribozyme ligase from an RNA pool that contained 90 random
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sequence positions (Robertson and Ellington, 1999; Robertson et al., 2001). A variant of this pool was generated in which only four positions in the constant region were altered, substituting a GAAA tetraloop in place of a UUCG tetraloop. Selection experiments for ribozyme ligases were initiated from amplified aliquots of this pool; one aliquot was used for the selection of ribozymes containing canonical nucleotides, other aliquots of identical complexity were used for the selection of ribozymes containing one of three different modified nucleotides (Robertson, 2001). After six rounds of selection and amplification, all of the pools had collapsed to contain a relatively few winning sequences. However, the rates of all of these winning sequences were remarkably slow (500-fold slower) compared to the ribozyme ligase originally selected from the slightly different pool. There was no discernable difference between the ribozymes that contained canonical or modified nucleotides; they were all very slow. The authors hypothesized that the pools had somehow become artificially narrowed during the course of the selection, and therefore repeated the selection experiments with new aliquots of the original, amplified pool. Again, only very slow ribozymes were obtained.
be incorporated between rounds in a selection to more efficiently explore sequence space. However, it should be noted that in the latter case, one does not want the mutation rate to be so high that a few of each winning sequence from one round do not survive unchanged into the next round. One benefit of using modified nucleotides rather than standard mutagenic PCR is that it is easy to control the amount of mutation introduced into the sample simply by adjusting the relative rate of modified to unmodified nucleotide in the amplification process. As such, a higher rate of mutation can be achieved than with mutagenic PCR, which would be particularly useful for diversifying an initial pool. Zaccolo et al. (1996) describe such a system using both a purine and pyrimidine analog in a PCR reaction, and have quantitated the frequencies of each base transition. It should be mentioned that when using mutagenic modified nucleotides, one would not want them to be included in the active pool, as their decreased fidelity would make it less likely that they would be in the same position during the next round. For example, if a DNA selection were being performed, a second amplification of the pool would be required using only natural nucleotides.
Modified nucleotides and mutations Although a selection starts with a large number of sequences, this number is usually a small fraction of the total number of sequences possible for the length of the random region. Additionally, with each round, the number of sequences within the pool is diminished. As such, it may be useful to explore the sequence space around the selected winners in order to discover functional variants. To some extent, this occurs in any selection due to the inherent mutations that arise in the amplification process; however, this background mutation rate is small, and one may wish to increase the frequency of mutations. For these purposes, the mutagenic potential of a nucleotide analog that serves as a nonspecific template can be used to increase the diversity of a pool. Similar to mutagenic PCR (UNIT 9.4), these techniques can be used at the outset of a selection to mutate an existing ribozyme or aptamer for either optimization or reselection for altered specificity. For example, Kore et al. (2000) have used modified nucleotides to create a degenerate pool based on the hammerhead ribozyme, from which they selected a variant that cleaves at an alternate sequence. Additionally, a mutagenic step can
Anticipated Results As with any selection experiments (UNITS 9.3 is virtually impossible to anticipate the outcome of any given experiment. This is especially true when considering the incorporation of modified nucleotides, since relatively few selection experiments have so far been carried out with modified residues. However, if the protocols for the production of RNA and DNA pools that have been outlined are followed, it can be anticipated that it should be possible to generate and purify upwards of at least 1014 different nucleic acid sequences (∼10 µg) that contain a particular modified nucleotide. For successful selection experiments, the population should be sieved by a factor of 100 to 1000 each round. That is, <1% of the total population will be recovered following a round of selection. Thus, most successful selections will show significant functional improvement or be completed within 5 to 10 rounds. However, in some selections it may take very successful functional sequences a longer period of time to overcome a great sea of mediocre functional sequences. For example, it took 18 rounds of selection to isolate cGMP-dependent aptazymes from a random sequence population
& 9.4), it
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(Koizumi et al., 1999). Following selection, the population should contain a relatively small number (1 to 10) of sequence classes. A sequence class is a set of related sequences that contain a common motif. To date, the sequences of aptamers or ribozymes selected from pools containing modified nucleotides do not correspond to the sequences of aptamers or ribozymes selected from pools that contain canonical nucleotides. Thus, any selection that includes modified nucleotides is likely to produce new sequence motifs.
Time Considerations The preparative portions of each round of selection can be expected to take several days. A transcription could take several hours to overnight, followed by ∼2-hr gel purification, and an overnight elution from the excised gel pieces. The amount of time spent on the selective steps will vary greatly with each individual selection. However, in initial rounds, the reaction time alone is typically several hours to an entire day. The selective segregation of winning sequences from losing sequences will vary from a few to several hours, depending on the method used. Reverse transcription followed by PCR, isolation, and quantitation of new template DNA will take a few more hours, depending mainly on the number of cycles required for amplification. The largest variable, the number of rounds of selection required, will vary greatly depending on the activity sought, from only a few to ≥12. Cloning and assaying of individual sequences will likely also take several days. Thus, even if no problems are encountered during a selection, it is likely to take ≥1 month to complete.
Literature Cited Bartel, D.P. and Szostak, J.W. 1993. Isolation of new ribozymes from a large pool of random sequences. Science 261:1411-1418. Battersby, T.R., Ang, D.N., Burgstaller, P., Jurczyk, S.C., Bowser, M.T., Buchanan, D.D., Kennedy, R.T., and Benner, S.A. 1999. Quantitative analysis of receptors for adenosine nucleotides obtained via in vitro selection from a library incorporating a cationic nucleotide analog. J. Am. Chem. Soc. 121:9781-9789. Beaudry, A., DeFoe, J., Zinnen, S., Burgin, A., and Beigelman, L. 2000. In vitro selection of a novel nuclease-resistant RNA phosphodiesterase. Chem. Biol. 7:323-334. In Vitro Selection Using Modified or Unnatural Nucleotides
Brieba, L.G. and Sousa, R. 2000. Roles of histidine 784 and tyrosine 639 in ribose discrimination by T7 RNA polymerase. Biochemistry 39:919-923.
Green, L.S., Jellinek, D., Bell, C., Beebe, L.A., Feistner, B.D., Gill, S.C., Jucker, F.M., and Janjic, N. 1995. Nuclease-resistant nucleic acid ligands to vascular permeability factor/vascular endothelial growth factor. Chem. Biol. 2:683695. Griffiths, A.D., Potter, B.V., and Eperon, I.C. 1987. Stereospecificity of nucleases towards phosphorothioate-substituted RNA: Stereochemistry of transcription by T7 RNA polymerase. Nucl. Acids Res. 15:4145-4162. Jellinek, D., Green, L.S., Bell, C., Lynott, C.K., Gill, N., Vargeese, C., Kirschenheuter, G., McGee, D.P., Abesinghe, P., Pieken, W.A., et al. 1995. Potent 2′-amino-2′-deoxypyrimidine RNA inhibitors of basic fibroblast growth factor. Biochemistry 34:11363-11372. Jhaveri, S., Olwin, B., and Ellington, A.D. 1998. In vitro selection of phosphorothiolated aptamers. Bioorg. Med. Chem. Lett. 8:2285-2290. King, D.J., Ventura, D.A., Brasier, A.R., and Gorenstein, D.G. 1998. Novel combinatorial selection of phosphorothioate oligonucleotide aptamers. Biochemistry 37:16489-16493. Koizumi, M., Soukup, G.A., Kerr, J.N., and Breaker, R.R. 1999. Allosteric selection of ribozymes that respond to the second messengers cGMP and cAMP. Nat. Struct. Biol. 6:1062-1071. Kore, A.R., Vaish, N.K., Morris, J.A., and Eckstein, F. 2000. In vitro evolution of the hammerhead ribozyme to a purine-specific ribozyme using mutagenic PCR with two nucleotide analogues. J. Mol. Biol. 301:1113-1121. Kostyuk, D.A., Dragan, S.M., Lyakhov, D.L., Rechinsky, V.O., Tunitskaya, V.L., Chernov, B.K., and Kochetkov, S.N. 1995. Mutants of T7 RNA polymerase that are able to synthesize both RNA and DNA. FEBS Lett. 369:165-168. Kubik, M.F., Bell, C., Fitzwater, T., Watson, S.R., and Tasset, D.M. 1997. Isolation and characterization of 2′-fluoro-, 2′-amino-, and 2′-fluoro/amino-modified RNA ligands to human IFNgamma that inhibit receptor binding. J. Immunol. 159:259-267. Kujau, M.J., Siebert, A., and Wolfl, S. 1997. Design of leader sequences that improve the efficiency of the enzymatic synthesis of 2′-amino-pyrimidine RNA for in vitro selection. J. Biochem. Biophys. Methods 35:141-151. Latham, J.A., Johnson, R., and Toole, J.J. 1994. The application of a modified nucleotide in aptamer selection: Novel thrombin aptamers containing 5-(1-pentynyl)-2′-deoxyuridine. Nucl. Acids Res. 22:2817-2822. Levy, M. and Ellington, A.D. 2001a. In vitro selection of a deoxyribozyme that can utilize multiple substrates. J. Mol. Evol. In press. Levy, M. and Ellington., A.D. 2001b. Section of deoxyribozyme ligases that catalyze the formation of an unnatural internucleotide linkage. Bioorg. Med. Chem. 9:2581-2587.
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Lin, Y., Qiu, Q., Gill, S.C., and Jayasena, S.D. 1994. Modified RNA sequence pools for in vitro selection. Nucl. Acids Res. 22:5229-5234. Lutz, M.J., Horlacher, J., and Benner, S.A. 1998. Recognition of a non-standard base pair by thermostable DNA polymerases. Bioorg. Med. Chem. Lett. 8:1149-1152.
Robertson, M.P., Hesselberth, J.R., and Ellington, A.D. 2001. Optimization and optimality of a short ribozyme ligase that joins non-WatsonCrick base pairings. RNA 7:513-523.
Lutz, S., Burgstaller, P., and Benner, S.A. 1999. An in vitro screening technique for DNA polymerases that can incorporate modified nucleotides. Pseudo-thymidine as a substrate for thermostable polymerases. Nucl. Acids Res. 27:2792-2798.
Ruckman, J., Green, L.S., Beeson, J., Waugh, S., Gillette, W.L., Henninger, D.D., ClaessonWelsh, L., and Ja njic, N. 1998. 2′Fluoropyrimidine RNA-based aptamers to the 165-amino acid form of vascular endothelial growth factor (VEGF165). Inhibition of receptor binding and VEGF-induced vascular permeability through interactions requiring the exon 7-encoded domain. J. Biol. Chem. 273:20556-20567.
Marshall, K.A. and Ellington, A.D. 1999. Molecular parasites that evolve longer genomes. J. Mol. Evol. 49:656-663.
Santoro, S.W. and Joyce, G.F. 1997. A general purpose RNA-cleaving DNA enzyme. Proc. Natl. Acad. Sci. U.S.A. 94:4262-4266.
Milligan, J.F. and Uhlenbeck, O.C. 1989. Synthesis of small RNAs using T7 RNA polymerase. Methods Enzymol. 180:51-62.
Santoro, S.W., Joyce, G.F., Sakthivel, K., Gramatikova, S., and Barbas, C.F. III. 2000. RNA cleavage by a DNA enzyme with extended chemical functionality. J. Am. Chem. Soc. 122:2433-2439.
Nakamaye, K.L., Gish, G., Eckstein, F., and Vosberg, H.P. 1988. Direct sequencing of polymerase chain reaction amplified DNA fragments through the incorporation of deoxynucleoside alpha-thiotriphosphates. Nuc l. Acids Res. 16:9947-9959. Padilla, R. and Sousa, R. 1999. Efficient synthesis of nucleic acids heavily modified with non-canonical ribose 2′-groups using a mutantT7 RNA polymerase (RNAP). Nucl. Acids Res. 27:15611563. Pagratis, N.C., Bell, C., Chang, Y.F., Jennings, S., Fitzwater, T., Jellinek, D., and Dang, C. 1997. Potent 2′-amino- and 2′-fluoro-2′-deoxyribonucleotide RNA inhibitors of keratinocyte growth factor. Nat. Biotechnol. 15:68-73. Perrin, D.M., Garestier, T., and Helene, C. 2001. Bridging the gap between proteins and nucleic acids: A metal-independent RNAseA mimic with two protein-like functionalities. J. Am. Chem. Soc. 123:1556-1563. Piccirilli, J.A., Krauch, T., Moroney, S.E., and Benner, S.A. 1990. Enzymatic incorporation of a new base pair into DNA and RNA extends the genetic alphabet. Nature 343:33-37. Pieken, W.A., Olsen, D.B., Benseler, F., Aurup, H., and Eckstein, F. 1991. Kinetic characterization of ribonuclease-resistant 2′-modified hammerhead ribozymes. Science 253:314-317. Robertson, M.P. 2001. Ph.D. dissertation. Engineered regulation of an RNA ligase ribozyme. University of Texas, Austin. Robertson, M.P. and Ellington, A.D. 1999. In vitro selection of an allosteric ribozyme that transduces analytes to amplicons. Nat. Biotechnol. 17:62-66.
Seelig, B. and Jaschke, A. 1999. A small catalytic RNA motif with Diels-Alderase activity. Chem. Biol. 6:167-176. Southworth, M.W., Kong, H., Kucera, R.B., Ware, J., Jannasch, H.W., and Perler, F.B. 1996. Cloning of thermostable DNA polymerases from hyperthermophilic marine Archaea with emphasis on Thermococcus sp. 9°N-7 and mutations affecting 3′-5′ exonuclease activity. Proc. Natl. Acad. Sci. U.S.A. 93:5281-5285. Tarasow, T.M., Tarasow, S.L., and Eaton, B.E. 1997. RNA-catalysed carbon-carbon bond formation. Nature 389:54-57. Teramoto, N., Imanishi, Y., and Ito, Y. 2000. In vitro selection of a ligase ribozyme carrying alkylamino groups in the side chains. Bioconjug. Chem. 11:744-748. Wiegand, T.W., Janssen, R.C., and Eaton, B.E. 1997. Selection of RNA amide synthases. Chem. Biol. 4:675-683. Zaccolo, M., Williams, D.M., Brown, D.M., and Gherardi, E. 1996. An approach to random mutagenesis of DNA using mixtures of triphosphate derivatives of nucleoside analogues. J. Mol. Biol. 255:589-603. Zon, G. and Geiser, T.G. 1991. Phosphorothioate oligonucleotides: Chemistry, purification, analysis, scale-up and future directions. Anticancer Drug Des. 6:539-568.
Contributed by Scott M. Knudsen, Michael P. Robertson, and Andrew D. Ellington University of Texas Austin, Texas
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CHAPTER 10 Purification and Analysis of Synthetic Nucleic Acids and Components INTRODUCTION urification and analysis are obviously essential to studies involving synthetic nucleic acids, but choice of the most appropriate technique, which ultimately depends on the final use for the oligonucleotide, is not always obvious. An overview unit (UNIT 10.3) discusses alternative techniques and provides information necessary for the reader to determine the most appropriate technique for a particular application of interest. For example, oligonucleotides to be used for sequencing or as PCR primers do not need to be as rigorously pure as those to be used for cloning or X-ray crystallography studies. The vast majority of oligonucleotides are purified by three techniques: polyacrylamide gel electrophoresis (PAGE; UNIT 10.4), high-performance liquid chromatography (HPLC; UNIT 10.5), or commercially available cartridges (UNIT 10.7). The overview unit contains basic information about synthesis, including evaluation of coupling yields, and options for deprotection, workup, purification, and quantification.
P
The most common technique for purification of oligonucleotides to be used as primers in PCR and sequencing is polyacrylamide gel electrophoresis. UNIT 10.4 provides detailed procedures for sample preparation, running gels, and visualization and isolation of oligonucleotides. UNIT 10.5 on HPLC analysis and purification of synthetic nucleic acids includes basic information and procedures using both reversed-phase and anion-exchange HPLC. Cartridge techniques for oligonucleotide purification and desalting have become very popular because of their speed and efficiency. UNIT 10.7 provides the reader the background necessary to effectively use this technique.
Another method for separation of oligonucleotides is capillary electrophoresis (CE). Because of the small capillary diameters and high separating voltages used, CE is fast and offers high sensitivity and resolution (e.g., single-nucleotide resolution for nanogram quantities of fragments up to several hundred base pairs). UNIT 10.9 describes theory, instrumentation, and methods for analysis of oligonucleotides by CE. Mass spectrometry has become a pervasive and indispensable technique in nucleic acid chemistry. Indeed, mass spectrometry is now particularly useful in the characterization of nucleoside phosphoramidites, which are the most widely used synthons in contemporary solid-phase oligonucleotide synthesis. The molecular weight determination of these compounds has been problematic at both low and high resolution, presumably because of their sensitivity to mild acids. This limitation has been overcome with the recent development of a novel matrix system that has enabled the rapid and reliable molecular weight determination of nucleosidic and non-nucleosidic phosphoramidites by liquid secondary ion mass spectrometry or fast-atom bombardment mass spectrometry. This valuable procedure is featured in UNIT 10.11. The composition of synthetic oligonucleotides should always be confirmed. Recently, mass spectrometry has developed as the most efficient tool for rapid analysis of short
Purification and Analysis of Synthetic Nucleic Acids and Components
Contributed by Donald E. Bergstrom
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Current Protocols in Nucleic Acid Chemistry (2006) 10.0.1-10.0.2 C 2006 by John Wiley & Sons, Inc. Copyright
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oligonucleotides (<40 nt). A molecular ion mass determination along with either PAGE or HPLC data is generally considered sufficient for the confirmation of oligonucleotide integrity and purity. MALDI-TOF mass analysis has become especially commonplace and is now routinely used by commercial oligonucleotide synthesis vendors. Two different mass spectrometry techniques, matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF; UNIT 10.1) and electrospray ionization (ESI) mass spectrometry (UNIT 10.2) are particularly useful. The differences between the techniques, as well as the advantages and disadvantages of each, are compared. Both units contained detailed instruction for sample preparation and useful troubleshooting tips. Both techniques can be used to sequence oligonucleotides, and the fragment pattern in ESI mass spectrometry may be helpful in confirming the identity of oligonucleotide modifications. This kind of a more detailed analysis may be important for characterization of modified oligonucleotides for which conventional gel-based sequencing assays are not informative. An innovative method for sequencing synthetic oligonucleotides consists of extracting coupling failure sequences from crude syntheses of oligonucleotides functionalized with a 5 -O-(4,4 -dimethoxytrityl) group. Assessment of mass differences between these shorter-length sequences by MALDI-TOF mass spectrometry allows the identification of specific nucleobases or structural modifications. The details of this creative approach to oligonucleotide sequence determination are presented in UNIT 10.10. Base composition analysis, described in UNIT 10.6, is an important tool for establishing the presence and integrity of modified bases in synthetic oligonucleotides. Snake venom phosphodiesterase, a 3 to 5 exonuclease, is capable of degrading oligonucleotides containing modified bases. In conjunction with monomer standards, HPLC nucleoside analysis of enzyme-degraded oligonucleotides is a powerful tool for confirming the presence and quantifying modified nucleosides. The oxidation of DNA is mediated by many different xenobiotics, including DNAdamaging drugs, substances present in the environment, and radiation. Determination of the pathway of oxidation is an important step in understanding the mechanism of DNA degradation by these substances. In UNIT 10.8, Awada and Dedon provide experimental methods for determining the site and, hence, mechanism of oxidation of DNA. These methods allow one to distinguish pathways that involve 1 , 3 , 4 , and 5 hydrogen atom abstraction. Donald E. Bergstrom
Introduction
10.0.2 Supplement 26
Current Protocols in Nucleic Acid Chemistry
Analysis of Oligonucleotides by Matrix-Assisted Laser Desorption/Ionization Time-of-Flight Mass Spectrometry
UNIT 10.1
Matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS; Karas et al., 1987; Karas and Hillenkamp, 1988) is one of the most useful techniques available for determining biomolecule mass. MALDI-MS offers high mass accuracy, good sensitivity, simplicity, and speed, all of which make it suitable for the analysis of oligonucleotides. Conceptually, sample analysis using MALDI-MS is relatively straightforward: the analyte of interest is mixed with an appropriate matrix and spotted on a sample plate, and the plate is placed in the instrument for analysis. Normally, singly charged ions of oligonucleotides are observed, which makes MALDI-MS spectra very easy to interpret and simplifies the analysis of mixtures (Fig. 10.1.1). Nowadays commercial MALDI instruments, usually interfaced with time-of-flight (TOF) mass analyzers, are easy to operate and are routinely used for many types of oligonucleotide analysis. A complementary approach to oligonucleotide analysis is electrospray ionization mass spectrometry (ESIMS); this approach is presented, along with a comparison of the two techniques, in UNIT 10.2. At present MALDI-MS is utilized routinely for the analysis of 120-mer and smaller oligonucleotides. Polymerase chain reaction (PCR) and restriction enzyme digestion products larger than 120-mers have not yet been routinely mass analyzed, although 500-mer and larger oligodeoxynucleotides have been ionized and observed (Tang et al., 1994a; Liu et al., 1995a; Berkenkamp et al., 1998). Poor ionization efficiencies and fragmentation during the desorption/ionization process have been blamed for the lack of success with higher-molecular-weight oligonucleotides (Nordhoff et al., 1993; Schneider and Chait, 1993; Zhu et al., 1995). However, MALDI-MS has become a useful method to rapidly identify and characterize oligonucleotides up to 120-mers, and may soon replace the combination of Maxam-Gilbert chemical degradation and subsequent polyacrylamide gel electrophoresis (PAGE) for sequence characterization of these samples. Only high femtomole to low picomole per microliter amounts of oligonucleotides are needed for an analysis. Because MALDI-MS can be used to analyze mixtures, synthetic oligonucleotides (usually <30-mers) can be characterized by their failure sequences (Keough et al., 1993; Limbach, 1996). Exonuclease digestion has also proven to be a versatile alternative approach for determining the sequences of moderate-length oligonucleotides (Pieles et al., 1993; Limbach, 1996; Smirnov et al., 1996). A still unproven, but promising, strategy for determining oligonucleotide sequences is the analysis of prompt and metastable fragment ions (Nordhoff et al., 1995; Juhasz, 1996). Because of MALDI-MS’s potentially greater accuracy and speed of analysis compared to electrophoresis-based methods, considerable effort has been expended on DNA sequencing reaction (Sanger reaction) product readout (Murray, 1996). One strategy to overcome the ionization and fragmentation limitations is to replace normal nucleotides with nucleotide analogs (7-deaza- and 2′-fluoropurines) that are more stable during MALDI-MS analysis and are tolerated by the polymerase(s) used during chain extension and termination reactions (Kirpekar et al., 1995; Schneider and Chait, 1995; Tang et al., 1997b). Another potential application of MALDI-MS is the analysis of PCR products as a means to characterize disease-specific genetic mutations (Bai et al., 1994b; Ch’ang et al., 1995; Hurst et al., 1996; Wada and Yamamoto, 1997). Affinity probes based on hybridization between complementary strands of oligonucleotides to capture specific oligonucleotides fragments (e.g., DNA sequencing fragments, PCR products) for poten-
Purification and Analysis of Synthetic Nucleic Acids and Components
Contributed by Chau-Wen Chou and Patrick A. Limbach
10.1.1
Current Protocols in Nucleic Acid Chemistry (2000) 10.1.1-10.1.25 Copyright © 2000 by John Wiley & Sons, Inc.
Figure 10.1.1 MALDI mass spectrum of oligonucleotide mixture (a) prior to desalting and (b) after desalting with ammonium-activated cation-exchange resin beads.
tial automated MALDI-MS analysis are being developed (Jurinke et al., 1997; Little et al., 1997; Ross and Belgrader, 1997; Ross et al., 1997). In this unit the practical aspects of oligonucleotide analysis by MALDI-TOF MS are discussed, with protocols provided for certain specific procedures. Sample preparation— including purification of the oligonucleotide, preparation of the matrix and co-matrix, and preparation of the sample for analysis—is discussed in detail. Protocols for oligonucleotide purification using ammonium-activated cation-exchange resin beads (see Basic Protocol 1), spin columns (see Basic Protocol 2), C18 reversed-phase cartridges (see Alternate Protocol 1), and molecular-weight-cutoff filters (see Alternate Protocol 2) are included. Also covered are steps for preparing matrix and co-matrix solutions (see Basic Protocol 3) and the matrix/analyte samples for analysis (see Basic Protocol 4). In addition, an introduction to the instrumentation and its calibration is provided, along with discussions of the technique’s application to molecular-weight measurement, oligonucleotide sequencing—including a method for exonuclease-based sequencing of oligonucleotides using MALDI-TOF MS (see Basic Protocol 5)—and analysis of PCR reactions.
Analysis of Oligonucleotides
10.1.2 Current Protocols in Nucleic Acid Chemistry
OLIGONUCLEOTIDE PURIFICATION AND DESALTING The most important criterion for successful MALDI-MS analysis of oligonucleotides is sample preparation. Oligonucleotide sample contaminants must be reduced or removed prior to analysis. The primary challenge to accurate molecular-weight measurements of oligonucleotides and nucleic acids is the presence of salt adducts, which must be reduced or completely removed. In solution, the phosphodiester backbone is completely ionized at pH >1, and the solvent acts as a dielectric shield to reduce the repulsive Coulombic charging. During the ionization process, this Coulombic protection is lost, which eventually results in the adduction of any cations that may be present in the sample solution. These adducts shift the ion signal to higher m/z values and interfere with accurate determination of molecular weights (Fig. 10.1.1). Sources of Contamination Cations can arise from any source with which the oligonucleotide sample comes into contact, such as solvents, buffers, the mass spectrometer sample probe, solution storage containers, and pipet tips. To reduce cation concentration, ultrapure water—water that has been doubly distilled and deionized (e.g., “Nanopure” water, purified using a Barnstead/Thermolyne Nanopure system)—should always be used. Organic solvents to be used in preparing matrix solutions may need to be desalted by distillation or using activated ion-exchange resin beads, such as Bio-Rad AG501-X8(d) resin beads (Simmons and Limbach, 1998). Often the sample holder can be a source of contaminants. The sample probe and plates must be cleaned carefully before use so that cross-contamination with the previous sample will not interfere with the newly deposited oligonucleotides. Sonicating and washing in organic solvents, such as methanol and acetone, effectively removes most matrices and analytes. Further, stainless steel sample plates can be cleaned by polishing followed by acid oxidation in a dilute inorganic solution such as 0.1 N nitric acid. The sample plates should be rinsed with ultrapure water to eliminate residual cations. Nonmetallic sample substrates, such as nitrocellulose and Nafion films, can be used as alternatives; these have been demonstrated to reduce salt adducts and enhance the ion signal of oligonucleotides, especially for those of >100-mer size (Bai et al., 1994a,b, 1995; Liu et al., 1995a). Many buffers simply interfere with ion formation without forming intense adduct peaks. Investigation of the tolerance limits for several common buffers (Shaler et al., 1996) determined that oligonucleotides could be analyzed in the presence of up to 10 mM salt or 500 mM buffer in the sample solution, and that positive ions were less affected than negative ions by impurities. Purification Methods Table 10.1.1 summarizes the numerous oligonucleotide purification schemes available. The most common approaches are presented as protocols within this unit. In all cases, after purification, the sample may be analyzed immediately or stored (at –20°C) for future use. Optimal MALDI-MS results will be obtained if the sample is analyzed immediately after purification. A popular and useful method is to treat both the sample and matrix solutions with cation-exchange resin beads, such as Bio-Rad AG50W-X8 resin beads (see Basic Protocol 1; Nordhoff et al., 1992). Although these beads are usually obtained from the manufacturer in the free acid form, the ammonium-activated form is preferred for MALDIMS; a procedure for activating the beads is described below (see Support Protocol 1). A simple method to remove minor cations (at concentrations <10 mM) is to add an ammonium salt, such as ammonium citrate, tartrate, or fluoride, to the matrix solution
Purification and Analysis of Synthetic Nucleic Acids and Components
10.1.3 Current Protocols in Nucleic Acid Chemistry
(Currie and Yates, 1993; Pieles et al., 1993; Zhu et al., 1996b, Cheng, 1996). Except with 2′,4′,6′-trihydroxyacetophenone (THAP), the concentration of ammonium salt should be ≤10 mM (see section on Matrix/Analyte Preparation); otherwise, the matrix will not crystallize well, limiting the ability to find “hot spots” that yield abundant ion signal. A method that has been useful for desalting oligonucleotides in ESI-MS (see UNIT 10.2) has been adapted for MALDI-MS (Simmons and Limbach, 1997, 1998). In this approach organic bases, such as triethylamine or piperidine, are used as co-matrices—i.e., are combined with the matrix; these organic bases have high cation affinities and serve to reduce or eliminate cation adduction at levels exceeding those obtainable with cation-exchange resin beads for extremely salty samples. The organic bases should be treated with ion-exchange beads, such as Bio-Rad AG501-X8(d) resin beads, before use to eliminate impurities. A number of vendors (Millipore, Amersham Pharmacia Biotech, Boehringer Mannheim, Amicon) offer prepackaged purification systems. These purification devices are typically 0.5- to 2.5-mL microcentrifuge tubes containing coated particles or filters that retain the oligonucleotide and allow contaminants to be washed off the sample. C18 reversed-phase cartridges (see Alternate Protocol 1) utilize the strong binding affinity of oligonucleotides for the stationary-phase packing material in the presence of 2 M triethylamine as the means of trapping the sample for purification. The oligonucleotides are eluted with 20% aqueous acetonitrile. Sephadex G-25 spin-column purification (see Basic Protocol 2) is a very useful, quick method (∼5 min) to eliminate most salts and buffers if the oligonucleotide is larger than a 10-mer; generally one pass through the column yields satisfactory results. Spin columns must be equilibrated in ultrapure water before desalting. Molecular-weight-cutoff filters or membranes (see Alternate Protocol 2) retain oligonucleotides above a certain molecular weight and allow contaminants to be washed through. The membrane or filter is then inverted and the oligonucleotide is eluted and lyophilized. For samples that contain a high concentration of inorganic impurities (such as PCR products), reversed-phase high-performance liquid chromatography (HPLC), anion-exchange chromatography, microdialysis, or ethanol precipitation (see UNIT 10.2) followed Table 10.1.1 Common Approaches for Desalting and Purifying Oligonucleotides and Matrixes Prior to MALDI-TOF MS Analysis
Means of purification (and selected manufacturers) Oligonucleotides Cation-exchange resin beads Ammonium salt co-matrices Organic base co-matrices C18 reversed-phase cartridges (Waters, Perkin Elmer, Glen Research) Sephadex G-25 spin columns (Amersham Pharmacia Biotech, Boehringer Mannheim) Molecular-weight cutoff filters (Amicon, Millipore, Gelman Sciences) Microdialysis HPLC Analysis of Oligonucleotides
Matrices Recrystallization Cation-exchange resin beads
Reference(s) Nordhoff et al. (1992); Basic Protocol 1 Pieles et al. (1993); Basic Protocol 3 Simmons and Limbach (1997); Basic Protocol 3 Alternate Protocol 1 Roskey et al. (1996); Basic Protocol 2 Shaler et al. (1995); Alternate Protocol 2
Nordhoff et al. (1992); Basic Protocol 3
10.1.4 Current Protocols in Nucleic Acid Chemistry
by lyophilization are reasonable purification techniques. Several appropriate HPLC buffers for oligonucleotide purification are presented in Table 10.1.2. Biotin-linked oligonucleotides can be purified using affinity chromatography (Jurinke et al., 1997); this approach is often suitable (and preferred) for enzymatic digestion products. BASIC PROTOCOL 1
Cation-Exchange Resin Purification of Oligonucleotides Ammonium-activated resin beads are used to reduce salt contamination of oligonucleotide mixtures prior to MALDI-MS analysis. This method can also be used to desalt matrix solutions. Materials Oligonucleotide sample solution, 100 to 400 µM in ultrapure water Ammonium-activated cation-exchange resin beads (see Support Protocol 1), either dry or slurry CAUTION: RNA samples are easily degraded by adventitious nucleases. If RNA is to be purified or analyzed, all laboratory equipment must be sterilized prior to use. Use of gloves is recommended during preparation of nucleic acid samples and solutions to avoid contamination of glassware and other equipment. Autoclaved distilled water and the highest available grade of salts must be used during solution preparation. 1. Place oligonucleotide sample in a test tube and add 10 to 20 mg of resin per 20 µL sample solution. 2. Let mixture stand 5 to 20 min. Decant supernatant. 3. The sample is now ready for analysis (see Basic Protocol 4). Overnight incubation of an oligonucleotide sample with resin beads should be avoided, as this leads to degradation of oligonucleotide spectra (Fitzgerald et al., 1993a). Best results are obtained if the sample is analyzed immediately, but it may instead be stored at –20°C for later analysis. Table 10.1.2 Common HPLC Buffers Suitable for Oligonucleotide Purification
Buffer Reversed phase A = 25 mM triethylammonium bicarbonate (TEAB), pH 6 B = 40% acetonitrile (AcCN) Anion exchange A = 25 mM TEAB/20% (v/v) AcCN, pH 6.4
B = 1 M TEAB/20% (v/v) AcCN, pH 7.6
Preparation Add 7 mL triethylamine to container and fill with water to 2 L; bubble CO2 gas (from sublimation of dry ice) through solution for ~1.5 hr until pH reaches 6 Add 80 mL AcCN to container and fill with water to 2 L Add 7 mL triethylamine and 400 mL AcCN to container and fill with water to 2 L; bubble CO2 gas through solution for ~2.5 hr until pH reaches 6.4 Add 280 mL triethylamine and 400 mL AcCN to container and fill with water to 2 L; bubble CO2 gas through solution for at least 2.5 hr until pH reaches 7.6
Purification and Analysis of Synthetic Nucleic Acids and Components
10.1.5 Current Protocols in Nucleic Acid Chemistry
SUPPORT PROTOCOL 1
Ammonium Activation of Cation-Exchange Resin Beads This protocol describes the procedure for converting free-acid cation-exchange resin beads to the preferred ammonium form for use in Basic Protocol 1. Materials Ammonium acetate, HPLC grade Ultrapure (e.g., Nanopure) water Cation-exchange resin beads in free-acid (H-) state (e.g., Bio-Rad AG50W-X8 beads) 1. Add 30 g ammonium acetate to 250 mL ultrapure water. Stir for 1 hr. 2. Transfer the supernatant solution into a small container. 3. Add resin beads to solution, ensuring that the solution completely covers the beads and that the mixture can be stirred easily. Stir on a stir plate overnight. 4. Filter excess solution from resin beads and air-dry the beads if desired. Transfer the beads to a closed container and store at room temperature for future use. The resin beads can be used for up to 12 months after activation.
BASIC PROTOCOL 2
Spin-Column Purification of Oligonucleotides Oligonucleotides >10-mers can be purified prior to MALDI-MS analysis using a spin column. Alternative methods using C18 reversed-phase cartridges and molecular weight cutoff filters are presented below (see Alternate Protocols 1 and 2). Sephadex G-25 spin columns are efficient (95% salt free, 85% average recovery) and fast (<5 min). Materials Microspin column (e.g., Sephadex G-25) Ultrapure (e.g., Nanopure) water Oligonucleotide sample solution, 500 to 1000 µM in ultrapure water CAUTION: RNA samples are easily degraded by adventitious nucleases. If RNA is to be purified or analyzed, all laboratory equipment must be sterilized prior to use. Use of gloves is recommended during preparation of nucleic acid samples and solutions to avoid contamination of glassware and other equipment. Autoclaved distilled water and the highest available grade of salts must be used during solution preparation. 1. Resuspend the resin in the microspin column by vortexing gently. 2. Loosen the cap of the column and open bottom closure. 3. Place the column in a 1.5-mL microcentrifuge tube for support. 4. Prespin the column for 1 min at recommended g force. If the centrifuge is marked in rpm (revolutions per minute), using the following equation to convert the force (× g) to rpm: 2
1.12 × r × r pm 1000
RCF (in g) = Analysis of Oligonucleotides
where r = radius in millimeters measured from center of spindle to bottom of rotor bucket.
10.1.6 Current Protocols in Nucleic Acid Chemistry
5. Discard the solution to waste in the 1.5-mL tube support. Resuspend the resin in 200 µL ultrapure water to rinse off residual impurities and spin 1 min at the same speed. 6. Place the column in a new 1.5-mL tube and slowly apply the sample to the center of the angled surface of the compacted resin bed, being careful not to disturb the resin bed. Do not allow any of the sample to flow around the sides of the bed. 7. Spin the column for 2 min at the recommended g force. The purified sample will collect at the bottom of the support tube.
8. Lyophilize the purified sample. After lyophilization, the sample is ready for analysis (see Basic Protocol 4). Best results are obtained if the sample is analyzed immediately, but it may instead be stored at –20°C for later analysis.
C18 Reversed-Phase Cartridge Purification of Oligonucleotides
ALTERNATE PROTOCOL 1
This protocol describes the use of oligonucleotide purification cartridges, commonly used to purify machine-synthesized trityl-on oligonucleotides, for desalting oligonucleotides. Materials Oligonucleotide purification cartridge Acetonitrile (AcCN), HPLC grade 2.0 M triethylammonium acetate (TEAA), HPLC grade 10 nmol oligonucleotide sample, dry Ultrapure (e.g., Nanopure) water CAUTION: RNA samples are easily degraded by adventitious nucleases. If RNA is to be purified or analyzed, all laboratory equipment must be sterilized prior to use. Use of gloves is recommended during preparation of nucleic acid samples and solutions to avoid contamination of glassware and other equipment. Autoclaved distilled water and the highest available grade of salts must be used during solution preparation. 1. Slowly pass first 5 mL acetonitrile, then 5 mL of 2 M TEAA, through the purification cartridge to a waste container. 2. Dissolve the oligonucleotide sample in 1 to 3 mL ultrapure water, 0.1 M TEAA, or other aqueous solution. The loading solution should not contain any organic solvents or ammonium hydroxide.
3. Pass this solution through the oligonucleotide purification cartridge at a rate of ∼1 drop per second exiting the cartridge, and collect the eluate. 4. Pass the eluate through the cartridge a second time. 5. Pass 10 mL ultrapure water through the cartridge to waste. 6. Elute and collect the desalted oligonucleotide by slowly passing, drop by drop, 1 mL of 50% acetonitrile in ultrapure water through the cartridge. 7. Lyophilize the desalted sample. After lyophilization, the sample is ready for analysis (see Basic Protocol 4). Best results are obtained if the sample is analyzed immediately, but it may instead be stored at –20°C for later analysis.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.1.7 Current Protocols in Nucleic Acid Chemistry
ALTERNATE PROTOCOL 2
Molecular-Weight-Cutoff-Filter Purification of Oligonucleotides This protocol describes the procedure for purifying oligonucleotides larger than 25-mers prior to MALDI-MS analysis. It is essential to concentrate the sample to a small volume prior to desalting. Materials Molecular-weight-cutoff-filter column Oligonucleotide sample solution, 100 to 400 µM in ultrapure water Ultrapure (e.g., Nanopure) water CAUTION: RNA samples are easily degraded by adventitious nucleases. If RNA is to be purified or analyzed, all laboratory equipment must be sterilized prior to use. Use of gloves is recommended during preparation of nucleic acid samples and solutions to avoid contamination of glassware and other equipment. Autoclaved distilled water and the highest available grade of salts must be used during solution preparation. 1. Select a filter column with nucleotide cutoff equal to or smaller than the size of the nucleic acid to be desalted (consult the manufacturer’s instructions). 2. Insert the sample reservoir into a microcentrifuge tube or into the tube supplied by manufacturer. 3. To concentrate the sample, pipet up to 500 µL of sample solution into the reservoir. Spin under the conditions (time and g force) recommended by the manufacturer, being careful not to exceed the recommended force. 4. To exchange salt, dilute the concentrated sample to 500 µL with ultrapure water. Spin under the conditions (time and g force) recommended by the manufacturer. To reduce salt concentration further, repeat this step. 5. Remove reservoir from vial and invert into a new vial (save and store filtrate at –20°C until sample has been analyzed). 6. Spin for 2 min at 500 to 1000 × g to recover oligonucleotide sample in the vial. 7. Remove reservoir. 8. Lyophilize sample in vial. After lyphilization, the sample is ready for analysis (see Basic Protocol 4) Best results are obtained if the sample is analyzed immediately, but it may instead be stored at –20°C for later analysis.
MATRIX/ANALYTE PREPARATION Matrix and Co-Matrix Selection The essence of MALDI-MS is the matrix—typically a small acidic organic compound that has a relatively high molar absorptivity (ελ) at the wavelength of the laser. Common MALDI-MS matrices have molar absorptivities on the order of 103 to 105 M−1 cm−1. A large number of potential matrices have been investigated over the years, with a relatively small number becoming the standards for MALDI-MS of oligonucleotides (Fig. 10.1.2). Analysis of Oligonucleotides
3-Hydroxypicolinic acid (3-HPA) has been the most useful oligonucleotide matrix, as it consistently results in the least fragmentation (Wu et al., 1993). Picolinic acid (PA) and
10.1.8 Current Protocols in Nucleic Acid Chemistry
OH N
3-hydroxypicolinic acid
O
N
COOH
picolinic acid
CH3
OH
OH
O
OH
COOH N nicotinic acid
CH3
HN S
2',4',6'-trihydroxyacetophenone
COOH
N H
N
6-azathiothymine
COOH NH2 anthranilic acid (2-aminobenzoic acid)
Figure 10.1.2 Chemical structures of the matrices most commonly used for analyzing oligonucleotides with MALDI-TOF MS.
3-aminopicolinic (3-APA) acid have also been investigated (Tang et al., 1994b), but are less practical choices, because PA works best at 266 nm and 3-APA is still not commercially available (Juhasz et al., 1996). However, a mixture of PA and 3-HPA is often used for larger oligonucleotides (Liu et al., 1995a). Other commonly used matrices include 2′,4′,6′-trihydroxyacetophenone (THAP; Pieles et al., 1993; Zhu et al., 1996a), a mixture of anthranilic acid and nicotinic acid (AA/NA; Nordhoff et al., 1992), and 6-aza-2thiothymine (ATT; Lecchi et al., 1995). These are useful for smaller oligonucleotides, up to 25 bases in length (Nordhoff et al., 1996). The popular peptide and protein matrices, such 2,5-dihydroxybenzoic acid (DHBA; Strupat et al., 1991) and sinapinic acid (SA; Beavis and Chait, 1989), should not be used as they yield intense prompt and metastable fragmentation that limits the molecular ion abundance (Parr et al., 1992; Schneider and Chait, 1993).
Matrix and Co-Matrix Preparation
BASIC PROTOCOL 3
Fresh matrix solutions should be used within a week of preparation, before they start to degrade, and it is best to prepare them daily before use. Because most matrices have to be dissolved in suitable solvents prior to mixing with the analyte solution, the solvent(s) should be chosen based on the solubilities of the matrices. In all cases, a large molar excess of matrix to analyte (103 to 105:1) is used when preparing oligonucleotides for analysis. This protocol describes the procedure for preparing MALDI-MS matrix solutions. Note: Optimal results are obtained when the highest-purity matrix and co-matrix are used. Materials One of the following matrices: 3-Hydroxypicolinic acid (3-HPA; Aldrich or Fluka) 3-Hydroxypicolinic acid and picolinic acid (Aldrich or Fluka; 3-HPA/PA)
Purification and Analysis of Synthetic Nucleic Acids and Components
10.1.9 Current Protocols in Nucleic Acid Chemistry
2′,4′,6′-Trihydroxyacetophenone (THAP; Aldrich or Fluka) 6-Aza-2-thiothymine (ATT; Aldrich) Co-matrix: dibasic ammonium citrate or imidazole (Aldrich or Fluka) Organic solvent, HPLC grade: 100% methanol (for THAP) or 50% aqueous acetonitrile (for all other matrices) Ultrapure (e.g., Nanopure) water Ammonium-activated cation-exchange resin beads (see Support Protocol 1) CAUTION: RNA samples are easily degraded by adventitious nucleases. If RNA is to be purified or analyzed, all laboratory equipment must be sterilized prior to use. Use of gloves is recommended during preparation of nucleic acid samples and solutions to avoid contamination of glassware and other equipment. Autoclaved distilled water and the highest available grade of salts must be used during solution preparation. 1. Weigh one of the following amounts of matrix and place in a 1.5-mL tube: 0.03 g 3-HPA; 0.03 g 3-HPA and 0.0075 g PA; 0.05 g THAP; or 0.04 g ATT. 2. Add 500 µL of appropriate organic solvent to matrix (Table 10.1.3). Vortex for 30 sec. Except for THAP (which can be used directly), allow solutions to settle and remove supernatant for use with sample. Matrix solution should be used quickly after preparation. Storage for >3 days is not recommended.
3. Add ~50 mg of ammonium-activated cation-exchange resin beads to matrix solution. Stir gently and allow to stand 30 min. Remove supernatant for use with sample. Resin beads are included in matrix to reduce the formation of salt adducts. Using resin beads is strongly recommended when highest-purity matrix (purity >98%) is not available.
4. Optionally, prepare co-matrix solution by adding either 0.23 g dibasic ammonium citrate or 0.20 g imidazole to 10 mL water and vortexing until dissolved. Ammonium citrate solution should be made fresh prior to use; storage for >3 days is not recommended. Imidazole solution can be stored with Bio-Rad AG501-XG(d) resin beads for up to 3 months. Co-matrices are included in samples to reduce salt-adduct formation. Ammonium citrate is recommended for samples that are not expected to contain nonvolatile (e.g., sodium or potassium) salts, while imidazole is recommended for samples known to contain nonvolatile salts. Use of co-matrices is recommended even if the sample has been previously Table 10.1.3 Guidelines for Preparation of UV Matrix Solutions
Matrix 3-Hydroxypiconilic acid (3-HPA) 3-Hydroxypiconilic acid/piconilic acid (PA) 2′,4′,6′-Trihydroxyacetophenone (THAP) Analysis of Oligonucleotides
6-Aza-2-thiothymine (ATT)
Solution Saturated (60 g/L) 3-HPA in 50:50 (v/v) acetonitrile/water or in 25:75 (v/v) methanol/water Saturated 3-HPA and 15 g/L PA in 50:50 (v/v) acetonitrile/water 100 g/L THAP in 50:50 (v/v) methanol (or ethanol)/water or in 100% methanol 80 g/L ATT in 50:50 (v/v) acetonitrile/water
10.1.10 Current Protocols in Nucleic Acid Chemistry
purified, as it tends to improve the quality of the mass spectral results, possibly through a cooling mechanism during the desorption process (Simmons and Limbach, 1998).
Matrix/Analyte Sample Preparation
BASIC PROTOCOL 4
Typically oligonucleotides up to 20-mers in size can be analyzed at concentrations down to the low femtomoles per microliter. To analyze larger oligonucleotides, more concentrated solutions—as much as 5 nmol/µl—are required. It is not critical that the sample, matrix, and co-matrix (if used) be mixed prior to spotting on the sample plate, although this is often done. A “two-layer” sample deposition method, similar to the sample preparation method for peptides suggested by Vorm et al. (1994), is widely used for oligonucleotides. In this protocol, the matrix solution is first deposited on the sample plate and allowed to dry. The oligonucleotide solution, with or without additional matrix solution, is then added on top of the dried matrix. The sample may be air-dried or drying may be assisted by blowing a slow stream of gas (house air or nitrogen) at room temperature over the sample. Backbone-modified oligonucleotides in which the phosphodiester backbone is neutralized (e.g., methylphosphonates) or nonexistent (e.g., peptide nucleic acids) should be prepared and analyzed with MALDI-MS matrices used for peptides or protein analysis (Keough et al., 1993, 1996; Butler et al., 1996; Ross et al., 1997). If the backbone-modified oligonucleotide is not neutralized (e.g., phosphorothioates; Schuette et al., 1995; Polo et al., 1997), the oligonucleotides should be prepared as described for unmodified oligonucleotides. This protocol details the procedure for preparing MALDI-MS sample solutions. If 2′,4′6′-trihydroxyacetopherone (THAP) is used as the matrix, follow steps 2b and 4b in place of steps 2a and 4a. Materials Ultrapure (e.g., Nanopure) water Matrix solution (see Basic Protocol 3) Co-matrix solution (optional; see Basic Protocol 3) Oligonucleotide sample, dry, lyophilized or as a 100- to 400-µM solution in ultrapure water CAUTION: RNA samples are easily degraded by adventitious nucleases. If RNA is to be purified or analyzed, all laboratory equipment must be sterilized prior to use. Use of gloves is recommended during preparation of nucleic acid samples and solutions to avoid contamination of glassware and other equipment. Autoclaved distilled water and the highest available grade of salts must be used during solution preparation. 1. If sample is in solid form, dissolve oligonucleotide sample in ultrapure water to a concentration of 100 to 400 µM. Lower concentrations are appropriate for small (<25-mer) oligonucleotides and higher concentrations are necessary for large (>50-mer) oligonucleotides.
2a. To a 0.5-mL tube, add 4 µL matrix solution, 1 µL oligonucleotide solution, and (optionally) 0.5 µL co-matrix solution. 2b. Alternative procedure for THAP: To a 0.5 mL tube, add 4 µL matrix solution and 1 µL co-matrix solution.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.1.11 Current Protocols in Nucleic Acid Chemistry
3. Mix by withdrawing and expelling the solution with the pipet 10 times. 4a. Spot 2 µL of solution on the sample plate and dry. To allow homogeneous crystallization, do not disturb the spotted samples after they start to crystallize. 4b. Alternative procedure for THAP: Spot 2 µL of matrix/co-matrix solution on the sample plate. Immediately spot 1 µL of the sample solution on top of the matrix solution on the sample plate before the matrix begins to crystallize. To allow homogeneous crystallization, do not disturb the spotted samples after they start to crystallize. IMPORTANT NOTE: Do not load the sample plate into the mass spectrometer before the plate is dry. Good signal can be obtain anywhere around the edges of the crystallized spot.
INSTRUMENTATION The Role of the Matrix It is now agreed that the matrix plays several roles in promoting the formation of intact molecular ions from the sample being analyzed: analyte isolation, absorption of laser radiation, analyte desorption, and analyte ionization. The ability of the matrix to serve as a solvent for the analyte is critical to the success of MALDI-MS. The matrix has molar absorptivities much higher than the analyte, so the laser energy is preferentially deposited in (absorbed by) the matrix. This process reduces subsequent fragmentation of the analyte, improving the production of intact molecular ions. While the exact matrix characteristics that facilitate analyte desorption are still the subject of investigation, it is presumed that the laser energy absorbed by the matrix allows desorption of the first few layers of the sample. Current research suggests heats of sublimation are an important consideration in analyte desorption (Vertes et al., 1993; Bencsura et al., 1997). One of the more contentious issues in MALDI-MS is the role of the matrix in the ionization process. The scenario most commonly reported involves photoionization of the matrix molecule, which can then react, by one of a number of possible pathways, to yield the analyte ion (Ehring et al., 1992). A number of investigations aimed at better understanding the exact mechanism of analyte ion formation have been performed, and presently there is no consensus whether any one particular ion-formation pathway dominates the process (Dreisewerd et al., 1995; Gimon-Kinsel et al., 1997). Most likely, analyte ion formation is influenced largely by the chemical properties of the analyte and matrix used, and more than one pathway may lead to the ultimate formation of the analyte ions detected in the mass spectrum. Laser Source Pulsed-nitrogen lasers (λ = 337 nm) are most commonly used in MALDI-MS and are the standard lasers available on commercial instruments. IR lasers were used in some early studies of MALDI-MS, and have recently received additional attention because they give results strikingly similar to those of UV-MALDI lasers (Niu et al., 1998). In some cases, IR lasers are more successful than UV lasers at characterizing higher-molecular-weight samples (Berkenkamp et al., 1997, 1998).
Analysis of Oligonucleotides
Operationally, laser fluences on the order of 106 to 107 W/cm2 with spot sizes around 10 to 100 µm are common in MALDI-MS. The best results are achieved by working at or just above the threshold irradiance necessary to generate analyte signal. (Threshold laser-power density is the minimum laser-power density necessary to obtain reproducible signal.) To achieve the appropriate laser irradiance, various means of attenuating the laser output are typically employed; these are common accessories on commercial instruments.
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Figure 10.1.3
Schematic diagram of a linear MALDI-TOF mass spectrometer.
Time-of-Flight Mass Analyzer The most common instrument configuration is a MALDI ion source coupled to a time-of-flight (TOF) mass analyzer (Fig. 10.1.3). Separation of ions of different m/z values in TOF mass spectrometry is accomplished by accelerating the ions through a short 20- to 30-kV electric field, allowing them to drift through a field-free region, and measuring the total flight time from ion formation to impact on an electron-multiplier detector. Lower-mass ions have shorter flight times than higher-mass ions and thus, by calibrating the ions’ flight times through the instrument using standards of known mass, a mass spectrum can be obtained. An ion’s flight time is proportional to the square root of the mass-to-charge ratio of the ion and is proportional to the length of its flight path, as shown in the following equation: m L t=√ 2zeVacc
where t is the ion’s flight time in seconds, m is its mass in kilograms, z is the number of charges on the ion, e is the fundamental charge (1.6 × 10−19 C), Vacc is the accelerating voltage in the ion source in volts, and L is the total flight path in meters. The mass spectral resolution (R) between two ions, m and m + ∆m, is proportional to the flight time of the ion divided by twice the time interval of ion arrival at the detector: R=
m t = ∆m 2∆t
As is evident from this equation, longer flight paths typically result in higher mass-spectral resolution than do shorter flight paths. To avoid detector saturation by the matrix ions present in large excess in the sample, most commercial instruments are equipped with a low-mass ion gate. Low-mass matrix ions are deflected by an electrode that has a low potential of the same polarity as the ions and that is located in front of the detector, or the detector is turned on immediately after these low-mass ions reach it. The most important instrumental factor limiting MALDI mass analysis of oligonucleotides is the instability of gas-phase oligonucleotide ions, which results in a continuous
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O
A, G, C, or T
O
O
O N
A, G, C, or T O
N
O
O H O P OH O
O
NH2
X*
O
O
A, G, C, or T
H
O P OH O
Y
5' H2C
O
O
A, G, C, or T O O
H
H
H
O
H
O
O
3'
-Gua
O P O O
A, G, C, or T O
O NH
N
O
O HO P OH O
O H O P OH O
O
H O H O P OH
A, G, C, or T O
A, G, C, or T O
H O H HO P OH O X
O HO P O O
H H A, G, C, or T O
O
H Y*
Figure 10.1.4 (a) Nordhoff (Nordhoff et al., 1995) and (b) McLuckey (McLuckey et al., 1992) nomenclature for the dissociation products of oligonucleotides. The McLuckey nomenclature is generally followed in the literature because of its applicability to ESI-MS as well as MALDI-MS. (c) Prevalent oligonucleotide fragmentation mechanism in UV-MALDI-MS (Nordhoff et al., 1995). Base protonation initiates base loss at purine sites as a result of cleavage at the N-glycosidic bond, which leads to subsequent cleavage of the phosphodiester backbone at the 3′-CO bond. Fragment ions here are denoted using the Nordhoff nomenclature.
fragmentation process beginning immediately as the ions are formed and continuing until they are detected (Fig. 10.1.4). There are essentially four differing time scales for desorption/ionization-induced dissociations in MALDI time-of-flight mass spectrometry (MALDI-TOF MS): prompt, fast, fast metastable, and metastable. Analysis of Oligonucleotides
1. Prompt dissociations occur on a time scale equal to or less than that of the desorption event.
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2. Fast dissociations occur after the desorption event, but before or at the beginning of the acceleration event. 3. Fast metastable decays occur on the time scale of the acceleration event. 4. Metastable decays occur after the acceleration event, during the field-free flight time of the ion. In addition, the initial kinetic energy distribution and the presence of salt adducts can reduce the quality of mass-spectral results. To date, instrumentation improvements are able to reduce the deleterious effects of kinetic energy spread and ion formation. As mentioned above, reduction of the fragmentation processes can be achieved through proper matrix selection or through modification of the oligonucleotide structure. Adduct peaks can only be eliminated by appropriate purification techniques prior to mass analysis. The sensitivity, resolution, and mass accuracy depend on the particular TOF configuration used. MALDI was first coupled with basic linear TOF (L-TOF) instruments (Fig. 10.1.3), in which ions are continuously accelerated for less than a few hundred nanoseconds. Less fragmentation is observed than in other instrument configurations: fragmentation that occurs after ions leave the acceleration region and enter the field-free region will not be detected, because the L-TOF instrument cannot differentiate the parent and fragment ions. A significant disadvantage of continuous-extraction L-TOF is that it cannot reduce peak broadening occurring from the initial velocity distribution of ions. Typically (e.g., see Wu et al., 1994) this results in poor resolution and mass accuracy, with oligonucleotides larger than 50-mers yielding resolutions on the order of 20 to 60 FWHM (full-width half-maximum). Time-lag focusing, commonly referred to as delayed extraction (DE), was coupled to TOF as a means of reducing the initial velocity spread of ions while they are still in the accelerating region of the mass spectrometer (Colby et al., 1994; Brown and Lennon, 1995; Vestal et al., 1995). In the DE mode of operation, a second electrostatic gate is included in the ion-source region. During the laser pulse, this electrode is held at a potential sufficient to prevent the acceleration of the laser-desorbed ions into the field-free region. After a suitable delay period, which is mass dependent (Vestal et al., 1995), the ions are accelerated into the field-free region and analyzed as usual. DE L-TOF instruments achieve very reasonable resolution (∼1000 FWHM) and lower mass errors (∼0.01% to 0.1%) for oligonucleotides up to 50-mers (Juhasz et al., 1996). A common instrumental approach for increasing the flight path, which has the added advantage of addressing a higher-order ion-focusing problem, is the addition of an electrostatic mirror at the end of the flight tube that reverses the ion’s direction and refocuses it toward the detector. This “reflectron” TOF (re-TOF) mass analyzer increases the path-length of the ion, yielding longer flight times and hence higher mass-spectral resolution. The electrostatic ion mirror compensates for the initial kinetic energy spread of the ions, thereby improving the focusing of ions of a single m/z more effectively at the detector (Fig. 10.1.5). In addition, delayed-extraction can be combined with a reflectron time-of-flight mass analyzer to yield a “high-performance” MALDI mass spectrometer with resolution approaching 15,000 FWHM and part-permillion mass errors. The use of a reflectron can resolve metastable dissociations, but unless certain parameters are adjusted (Spengler, 1997), these ions appear as uninformative broad peaks.
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Figure 10.1.5 Schematic diagram of a reflectron MALDI-TOF mass spectrometer. Higher-energy ions penetrate the ion mirror further than lower-energy ions, allowing them to be focused to a common point at the detector.
INSTRUMENT CALIBRATION
Analysis of Oligonucleotides
The mass axis in TOF-MS is normally calibrated via a two-point calibration procedure. Calibration can be performed externally or internally; the latter provides higher mass accuracies. As is typical in TOF-MS, the two calibrant masses should bracket the mass(es) of interest. For small oligonucleotides (<10-mers), matrix peaks can serve as the lowermass calibration point. Doubly charged molecular ions often appear at abundances significant enough to serve as a check for the mass calibration. For larger oligonucleotides, the two calibration masses must be from ions other than those from the matrix. Peptides and proteins are not suitable as calibrants for oligonucleotides. The concentration of the calibrants should be the same as the concentration of the analytes, especially when internal calibration is used. Because the initial ion velocities depend on the laser power density (Spengler and Bokelmann, 1993) and the matrix (Juhasz et al., 1997; see also references therein), internal calibration is necessary in L-TOF-MS, because this configuration cannot correct for the initial velocity distribution of the ions. On the other hand, DE-TOF and re-TOF instruments can mitigate those factors and external calibration
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yields satisfactory results; internal calibration can still be used, however, and will provide better accuracy. Caution should be exercised when assigning accurate molecular weights of the calibrants. In L-TOF, the average molecular weight should be used, because this configuration will not resolve the 13C isotope peaks. The centroid of a peak (the mass value at the center of the peak) is preferred to the peak top when assigning molecular weights to ions whose isotopes have not been resolved. The centroid at 50% of the peak height may be used to avoid including trace cation-adduct peaks in the molecular weight assignment. Peak tops may be used to assign molecular weights of calibrants for instrument configurations that resolve the 13C isotope peaks. Calibrants producing abundant cation adducts should be avoided to minimize errors in molecular-weight assignments. MOLECULAR WEIGHT DETERMINATION USING MALDI-MS MALDI-MS analysis of oligonucleotides is typically performed in the negative-ion mode, due to its higher sensitivity, higher resolution, and lower fragmentation compared to positive-ion-mode analysis (Stemmler et al., 1995; Tang et al., 1997a). The best massspectral data is obtained at near-threshold laser-power densities. Threshold laserpower density varies by matrix (Wu et al., 1993; Gusev et al., 1995) and is usually higher for oligonucleotide analysis than for peptide analysis (Wu et al., 1993). Laser power higher than threshold may produce higher oligonucleotide-ion abundance. However, as mentioned previously, the fragmentation and peak resolution are dependent on the laser-power density, and operating at laser powers significantly above threshold may degrade the quality of the mass spectrum. One should always be aware that signal abundance will vary from spot to spot within the same sample, due to the heterogeneous matrix/analyte crystals. The large shot-to-shot variability typically precludes the use of MALDI-MS in quantitative analysis, except under special experimental conditions (Bruenner et al., 1996). OLIGONUCLEOTIDE AND NUCLEIC ACID SEQUENCING USING MALDI-MS MALDI-MS is becoming the method of choice for rapid and accurate analysis of smaller synthetic oligonucleotides after the products are purified (Keough et al., 1993; Wang and Biemann, 1994), and is suitable as a quality control method in oligonucleotide synthesis (Ball and Packman, 1997). Sequence information from synthetic oligonucleotides can be obtained using several different approaches, of which the two most successful are failure-sequence analysis and exonuclease digestion—both dependent on the formation of a “mass ladder” (Limbach, 1996; Nordhoff et al., 1996) and hence well-suited to MALDI-MS. Sequence information arising from the formation of a mass ladder is obtained by determining the mass difference between successive peaks in the mass spectrum (Table 10.1.4 and Fig. 10.1.6). As is the case for all sequence-determination Table 10.1.4 Characteristic Mass Losses for Naturally Occurring Deoxynucleotides (dX) and Ribonucleotides (rX)a
Nucleotide dA dG dC dT
Mass loss 313.27 329.27 289.25 304.26
Nucleotide rA rG rC rU
Mass loss 329.27 345.27 305.25 306.26
aSeen during failure sequence analysis or exonuclease digestion. All mass values are atomic weight based.
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Figure 10.1.6 MALDI mass spectrum of 3′-to-5′ exonuclease digestion of d(GCCCAAGCTG). After 25 min of digestion, nearly all of the oligonucleotide sequence can be determined.
methods that rely on the mass measurements of successive n-mers, oligodeoxynucleotides are easier to characterize than oligoribonucleotides due to the relatively large differences in mass among the four oligodeoxynucleotide residues. Because of the small mass difference between U- and C-containing nucleotides (only 1 u), ribonucleotide analysis requires a higher mass accuracy to correctly distinguish Us from Cs. Furthermore, all mass-ladder methods have a distinct advantage for sequence determination because it is the difference in two mass measurements that results in the desired information—the identity of the nucleotide residue. Detection of failure sequences from the original synthesis step is one of the simplest methods for characterizing the chain length and sequence of synthetic oligonucleotides (Keough et al., 1993; Butler et al., 1996; Juhasz et al., 1996). This simple and straightforward method takes advantage of the fact that automated solid-phase synthesis of oligonucleotides, especially those containing modified internucleotide linkages such as methylphosphonates or phosphorothioates, is not 100% efficient. The mass spectrum will therefore contain a series of peaks that correspond to the final product and to each failure sequence, each of which differs in mass by the appropriate nucleotide residue value. The sequence of the oligonucleotide is determined in the 5′-to-3′ direction from the mass ladder of the synthesis failure products. The use of exonucleases to generate mass ladders of oligonucleotides that are suitable for analysis by mass spectrometry is now a standard method for sequencing small to moderate-length oligonucleotides—up to 30-mers for L-TOF without DE and 50-mers for DE-TOF (Pieles et al., 1993; Limbach, 1996). Unlike the failure-sequence analysis method described above, this approach is suitable for the sequence analysis of naturally occurring oligonucleotides. Two exonucleases are commonly used. Phosphodiesterase I, or snake venom phosphodiesterase (SVP), is a 3′-to-5′ exonuclease that is inhibited by a 3′-terminal phosphate and will generate 3′-to-5′ sequence information, while phosphodiesterase II, or calf spleen phosphodiesterase (CSP), is a 5′-to-3′ exonuclease that is inhibited by a 5′-terminal phosphate and will generate 5′ to 3′ sequence information. Analysis of Oligonucleotides
SVP digestion of an oligonucleotide is generally more rapid than CSP digestion and is not inhibited by the presence of base- or sugar-modified nucleosides. CSP is
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inhibited by 2′-O-methyl-modified nucleosides (Pieles et al., 1993). Such inhibition may be advantageous when base- and sugar-methyl-modifications must be distinguished in the overall sequence. Digestion with SVP would locate methyl-modified nucleosides, and digestion with CSP would distinguish between a base or sugar modification. Typically, complete sequence coverage of moderate-length (15 to 50 nt) oligonucleotides can be achieved within a 20- to 40-min reaction period, depending on the enzyme and digestion conditions used (Bentzley et al., 1996; Tolson and Nicholson, 1998).
Oligonucleotide or Nucleic Acid Sequencing Using Sequential Exonuclease Digestion and MALDI-TOF MS
BASIC PROTOCOL 5
This protocol describes the procedure for exonuclease sequencing of oligonucleotides or nucleic acids using MALDI-TOF MS. SVP, which cleaves oligonucleotides in the 3′-to-5′ direction, should be used to generate 3′ sequence information while CSP, which cleaves in the opposite direction, should be used if 5′ sequence information is required. During the first few minutes of analysis, information can be gathered on the first five or so nucleotides in the sequence. After longer reaction times, sequence information can be obtained for nucleotides further along the chain (Limbach, 1996; Smirnov et al., 1996). The ultimate time for analysis is dependent on the size of the oligonucleotide and the rate of digestion, and is determined empirically. The buffer solutions used here are compatible with the matrices, and therefore matrix desalting is typically not required. Sample purification should be performed as needed just prior to sequencing. Materials 5-µg oligonucleotide sample, lyophilized Ultrapure (e.g., Nanopure) water 50 g/L dibasic ammonium citrate, pH ∼6.4 (unadjusted) 10% (v/v) aqueous ammonium hydroxide 1 × 10−4 U/µL phosphodiesterase I (from snake venom; SVP; Sigma) or 1 × 10−3 unit/µL phosphodiesterase II (from calf spleen; CSP; Sigma or Worthington Biochemical) in ultrapure water Matrix (see Basic Protocol 3) Dry ice CAUTION: RNA samples are easily degraded by adventitious nucleases. If RNA is to be purified or analyzed, all laboratory equipment must be sterilized prior to use. Use of gloves is recommended during preparation of nucleic acid samples and solutions to avoid contamination of glassware and other equipment. Autoclaved distilled water and the highest available grade of salts must be used during solution preparation. 1. Dissolve oligonucleotide sample in ultrapure water to a concentration of 100 to 400 µM (200 µM is optimum). 2. If performing 3′-to-5′ sequencing using SVP: Adjust ammonium citrate solution to pH 9.4 with 10% aqueous ammonium hydroxide. 3. Mix 4 to 5 µL phosphodiesterase and 1 µL ammonium citrate solution with 5 µL of oligonucleotide solution in a microcentrifuge tube. Close the tube and vortex the reaction mixture.
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4. Place reaction mixture in 37°C water bath and allow temperature to equilibrate. 5. Every 10 min, remove a 1- to 2-µL aliquot of the reaction mixture. 6a. Quickly mix the aliquot with appropriate volume of matrix solution and place in dry ice to quench reaction. Either immediately spot samples on the sample plate or retain them to be tested all together after all digestion aliquots are obtained. 6b. Alternatively, analyze the reaction mixture aliquot immediately by MALDI-TOF MS. Aliquots must be either quenched or analyzed immediately to limit further enzymatic digestion.
Future Developments: Full-Scale Oligonucleotide Sequencing As mentioned earlier, there are essentially four differing time scales for desorption/ionization-induced dissociations in MALDI-TOF MS: prompt, fast, fast metastable, and metastable. In theory, dissociations occurring during any one of these time scales will generate fragment ions that could be used to determine the sequence of the oligonucleotide. In practice, unless certain instrumental parameters are manipulated, most of these fragments result in a broadening of the molecular ion peak with a concomitant loss of resolution and sensitivity. Because of these drawbacks and because this sequencing method provides little control over the extent of fragmentation, there are few reports of the use of desorption/ionizationinduced fragmentation to determine oligonucleotide sequences. As mentioned earlier, instrument configurations will to a large extent determine the type of fragment ions that can be detected in the mass spectrum. Currently, the use of desorption/ionization-induced fragment ions for sequence determination of oligonucleotides is limited. The mechanisms of prompt fragment-ion production have been studied in some detail (Fig. 10.1.4; Nordhoff et al., 1995; Zhu et al., 1995), and fragmentation patterns appear to be similar to those found in ESI–tandem mass spectrometry (ESI-MS/MS) studies of oligonucleotides (see UNIT 10.2), suggesting that routine use of this method for sequence determination may be possible in the near future. As mentioned previously, analogs of nucleotides are more stable than natural oligonucleotides in MALDI-TOF MS. 7-Deazapurines have a carbon instead of a nitrogen at position 7 in the nucleobase—thought to be the site at which the protonation leading to oligonucleotide fragmentation occurs in the gas phase (Nordhoff et al., 1993; Schneider and Chait, 1993). The modification on 2′-fluoro-nucleotides makes the N-glycosidic bond stronger, preventing nucleobase loss (Zhu et al., 1995). Although DE-TOF MS has been used for the complete sequence assignment of the desorption/ionization-induced fragments of an 11-mer oligodeoxynucleotide (Juhasz et al., 1996), the fragmentation mechanisms are not understood and no other oligonucleotides have been characterized using this approach.
Analysis of Oligonucleotides
There has been considerable interest in the use of MALDI-TOF MS to replace current gel-based separation and radioactive or fluorescent detection schemes for enzymatically synthesized oligonucleotides. By overlapping four sets of Sanger sequencing products that are terminated at guanosine, adenosine, cytidine, and thymidine, respectively, the sequence of the oligonucleotide of interest can be determined. In the first report of this approach (Fitzgerald et al., 1993b), a mock sequencing experiment was performed in which machine-synthesized oligodeoxynucleotides, corresponding to the expected nested set of oligodeoxynucleotides in a Sanger approach, were analyzed using MALDI-TOF MS. The sequence of a potential DNA fragment could be determined up to 18 bases past the primer (a 35-mer overall) before the signal ceased to be observed.
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Because of the buffers and other components in these reaction mixtures, however, the use of MALDI-MS to analyze actual Sanger dideoxy-termination sequencing reactions has proven to be much more difficult. Shaler et al. (1995) characterized the products from a dideoxy reaction catalyzed by a modified T7 DNA polymerase system. Working with picomole quantities of material, they were able to read the sequence out to the 19th base past the 12-mer primer. Using a 21-mer as the primer, the complete sequence could be determined for a 45-base synthetic template. Roskey and coworkers have analyzed the primer extension products from the Sanger reaction of a 50-base template with a 13-base primer; the use of a higher-resolution DE-MALDI-TOF mass spectrometer for the analyses afforded sequence information out to the 32nd base past the primer (Roskey et al., 1996). Mouradian and coworkers have performed mass spectrometric analysis of the primer-extension products that are generated from the M13 bacteriophage DNA template that is commonly used in actual Sanger sequencing (Mouradian et al., 1996; Roskey et al., 1996) and obtained sequence information out to the 18th base past the primer. Clearly more work is needed in this area before mass spectrometry in combination with the Sanger method becomes a viable alternative to traditional gel- or capillary-based separation and detection schemes.
ANALYSIS OF POLYMERASE CHAIN REACTION PRODUCTS USING MALDI-MS A promising application of MALDI-MS is in the identification of DNA fragments amplified using the polymerase chain reaction (PCR). PCR allows selected regions of DNA extracted from a variety of sample sources to be amplified to a detectable level. PCR primers may be designed to produce amplified DNA products of anywhere from ten to hundreds of nucleotide residues. One impediment to high-accuracy analysis of PCRamplified fragments is the presence of buffers, primers, and nucleoside triphosphates, which can interfere with the mass-spectral analysis. A number of investigators have analyzed PCR-amplified fragments by MALDI-MS (Limbach, 1996). The combination of PCR and mass spectrometry shows the potential to be used as a rapid and accurate method for DNA diagnostics. Applications have included the analysis of human and bacterial DNA fragments. MALDI mass spectrometric analysis of PCR-amplified fragments has been used to distinguish a three-base deletion site in the cystic fibrosis transmembrane conductance regulator, which is characteristic of patients with cystic fibrosis (Ch’ang et al., 1995), and to distinguish genetic polymorphisms in human DNA (Liu et al., 1995b). Legionella pneumophila, the causative agents of Legionnaire’s disease, has been identified using MALDI-MS (Hurst et al., 1996).
SUMMARY The speed and sensitivity of mass spectrometric analyses make this an attractive approach to analyzing oligonucleotides and nucleic acids. With the advent of “soft” ionization techniques such as MALDI-MS and ESI-MS, the ability to analyze these molecules in their intact form has brought mass spectrometry into the forefront as a viable means for characterizing them. MALDI-MS is well-adapted for rapid characterization of oligonucleotides because of its relatively high tolerance of sample impurities, ease of spectral interpretation (resulting from the fact that predominantly singly charged ions are formed), and ability to handle complex mixtures of oligonucleotides. The rapid pace of technological development and the expected developments in understanding and reducing the causes of oligonucleotide fragmentation, should result in significant experimental advances in this area of analysis in the near future.
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LITERATURE CITED Bai, J., Liu, Y.-H., Cain, T.C., and Lubman, D.M. 1994a. Matrix-assisted laser desorption/ionization using an active perfluorosulfonated ionomer film substrate. Anal. Chem. 66:3423–3430. Bai, J., Liu, Y.H., Lubman, D.M., and Siemieniak, D. 1994b. Matrix-assisted laser desorption/ionization mass spectrometry of restriction enzyme–digested plasmid DNA using an active Nafion substrate. Rapid Commun. Mass Spectrom. 8:687-691. Bai, J., Liu, Y.-H., Liang, X., Zhu, Y., and Lubman, D.M. 1995. Procedures for detection of DNA by matrix-assisted laser desorption/ionization mass spectrometry using a modified Nafion film substrate. Rapid Commun. Mass Spectrom. 9:1172-1176. Ball, R.W. and Packman, L.C. 1997. Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry as a rapid quality control method in oligonucleotide synthesis. Anal. Biochem. 248:185-194. Beavis, R.C. and Chait, B.T. 1989. Cinnamic acid derivatives as matrices for ultraviolet laser desorption mass spectrometry of peptides. Rapid Commun. Mass Spectrom. 3:432-436. Bencsura, A., Navale, V., Sadeghi, M., and Vertes, A. 1997. Matrix-guest energy transfer in matrix-assisted laser desorption. Rapid Commun. Mass Spectrom. 11:679-682. Bentzley, C.M., Johnston, M.V., Larsen, B.S., and Gutteridge, S. 1996. Oligonucleotide sequence and composition determined by matrix-assisted laser desorption/ionization. Anal. Chem. 68:2141-2146. Berkenkamp, S., Menzel, C., Karas, M., and Hillenkamp, F. 1997. Performance of infrared matrix-assisted laser desorption/ionization mass spectrometry with lasers emitting in the 3-µm wavelength range. Rapid Commun. Mass Spectrom. 11:1399-1406. Berkenkamp, S., Kirpekar, F., and Hillenkamp, F. 1998. Infrared MALDI mass spectrometry of large nucleic acids. Science 281:260-262. Brown, R.S. and Lennon, J.J. 1995. Mass resolution improvement by incorporation of pulsed ion extraction in a matrix-assisted laser desorption ionization time-of-flight mass spectrometer. Anal. Chem. 67:19982003. Bruenner, B.A., Yip, T.T., and Hutchens, T.W. 1996. Quantitative analysis of oligonucleotides by matrix-assisted laser desorption/ionization mass spectrometry. Rapid Commun. Mass Spectrom. 10:1797-1801. Butler, J.M., Jiang-Baucom, P., Huang, M., Belgrader, P., and Girard, J. 1996. peptide nucleic acid characterization by MALDI-TOF mass spectrometry. Anal. Chem. 68:3283-3287. Ch’ang, L.Y., Tang, K., Schell, M., Ringelberg, C., Matteson, K.J., Allman, S.L., and Chen, C.H. 1995. Detection of ∆F508 mutation of the cystic fibrosis gene by matrix-assisted laser desorption/ionization mass spectrometry. Rapid Commun. Mass Spectrom. 9:772-774. Cheng, S.-W. and Chan, T.-W.D. 1996. Use of ammonium halides as co-matrices for matrix assisted laser desorption ionization studies of oligonucleotides. Rapid Commun. Mass. Spectrom. 10:907-910. Colby, S.M., King, T.B., and Reilly, J.P. 1994. Improving the resolution of matrix-assisted laser desorption/ionization time-of-flight mass spectrometry by exploiting the correlation between ion position and velocity. Rapid Commun. Mass Spectrom. 8:865-868. Currie, G.J. and Yates, J.R., III 1993. Analysis of oligodeoxynucleotides by negative-ion matrix-assisted laser desorption mass spectrometry. J. Am. Soc. Mass Spectrom. 4:955-963. Dreisewerd, K., Schürenberg, M., Karas, M., and Hillenkamp, F. 1995. Influence of the laser intensity and spot size on the desorption of molecules and ions in matrix-assisted laser desorption/ionization with a uniform beam profile. Int. J. Mass Spectrom. Ion Processes 141:127-148. Ehring, H., Karas, M., and Hillenkamp, F. 1992. Role of photoionization and photochemistry in ionization processes of organic molecules and relevance for matrix-assisted laser desorption ionization mass spectrometry. Org. Mass Spectrom. 27:472-480. Fitzgerald, M.C., Parr, G.R., and Smith, L.M. 1993a. Basic matrices for the matrix-assisted laser desorption/ionization mass spectrometry of proteins and oligonucleotides. Anal. Chem. 65:3204-3211. Fitzgerald, M.C., Zhu, L., and Smith, L.M. 1993b. The analysis of mock DNA sequencing reactions using matrix-assisted laser desorption/ionization mass spectrometry. Rapid Commun. Mass Spectrom. 7:895897. Gimon-Kinsel, M., Preston-Schaffter, L.M., Kinsel, G.R., and Russell, D.H. 1997. Effects of matrix structure/acidity on ion formation in matrix-assisted laser desorption ionization mass spectrometry. J. Am. Chem. Soc. 119:2534-2540. Gusev, A.L., Wilkinson, W.R., Proctor, A., and Hercules, D.M. 1995. Improvement of signal reproducibility and matrix/comatrix effects in MALDI analysis. Anal. Chem. 57:1034-1044. Analysis of Oligonucleotides
Hurst, G.B., Doktycz, M.J., Vass, A.A., and Buchanan, M.V. 1996. Detection of bacterial DNA polymerase chain reaction products by matrix-assisted laser desorption/ionization mass spectrometry. Rapid Commun. Mass Spectrom. 10:377-382.
10.1.22 Current Protocols in Nucleic Acid Chemistry
Juhasz, P., Roskey, M.T., Smirnov, I.P., Haff, L.A., Vestal, M.L., and Martin, S.A. 1996. Applications of delayed extraction matrix-assisted laser desorption ionization time-of-flight mass spectrometry to oligonucleotide analysis. Anal. Chem. 68:941-946. Juhasz, P., Vestal, M.L., and Martin, S.A. 1997. On the initial velocity of ions generated by matrix-assisted laser desorption ionization and its effect on the calibration of delayed extraction time-of-flight mass spectra. J. Am. Soc. Mass Spectrom. 8:209-217. Jurinke, C., van den Boom, D., Collazo, V., Luchow, A., Jacob, A., and Köster, H. 1997. Recovery of nucleic acids from immobilized biotin-streptavidin complexes using ammonium hydroxide and applications in MALDI-TOF mass spectrometry. Anal. Chem. 69:904-910. Karas, M., Bachmann, D., Bahr, U., and Hillenkamp, F. 1987. Matrix-assisted ultraviolet laser desorption of non-volatile compounds. Int. J. Mass Spectrom. Ion Processes 78:53-68. Karas, M. and Hillenkamp, F. 1988. Laser desorption ionization of proteins with molecular masses exceeding 10,000 daltons. Anal. Chem. 60:2299-2301. Keough, T., Baker, T.R., Dobson, R.L.M., Lacey, M.P., Riley, T.A., Hasselfield, J.A., and Hesselberth, P.E. 1993. Antisense DNA oligonucleotides II: The use of matrix-assisted laser desorption/ionization mass spectrometry for the sequence verification of methylphosphonate oligodeoxyribonucleotides. Rapid Commun. Mass Spectrom. 7:195-200. Keough, T., Shaffer, J.D., Lacey, M.P., Riley, T.A., Marvin, W.B., Scurria, M.A., Hasselfield, J.A., and Hesselberth, E.P. 1996. Detailed characterization of antisense DNA oligonucleotides. Anal. Chem. 68:3405-3412. Kirpekar, R., Nordoff, E., Kristiansen, K., Roepstorff, P., Hahner, S., and Hillenkamp, F. 1995. 7-Deaza purine bases offer a higher ion stability in the analysis of DNA by matrix-assisted laser desorption/ionization mass spectrometry. Rapid Commun. Mass Spectrom. 9:525-531. Lecchi, P., Le, H.M.T., and Pannell, L.K. 1995. 6-Aza-2-thiothymine: A matrix for MALDI spectra of oligonucleotides. Nucleic Acids Res. 23:1276-1277. Limbach, P.A. 1996. Indirect mass spectrometric methods for characterizing and sequencing oligonucleotides. Mass Spectrom. Rev. 15:297-336. Little, D.P., Cornish, T.J., O’Donnell, M.J., Braun, A., Cotter, R.J., and Köster, H. 1997. MALDI on a chip: Analysis of arrays of low-femtomole to subfemtomole quantities of synthetic oligonucleotides and DNA diagnostic products dispensed by piezoelectric pipet. Anal. Chem. 69:4540-4546. Liu, Y.-H., Bai, J., Liang, X., Lubman, D.M., and Venta, P.J. 1995a. Use of a nitrocellulose film substrate in matrix-assisted laser desorption/ionization mass spectrometry for DNA mapping and screening. Anal. Chem. 67:3482-3490. Liu, Y.-H., Bai, J., Zhu, Y., Liang, X., Siemieniak, D., Venta, P.J., and Lubman, D.M. 1995b. Rapid screening of genetic polymorphisms using buccal cell DNA with detection by matrix-assisted laser desorption/ionization mass spectrometry. Rapid Commun. Mass Spectrom. 9:735-743. McLuckey, S.A., Van Berkel, G.J., and Glish, G.L. 1992. Tandem mass spectrometry of small, multiply charged oligonucleotides. J. Am. Soc. Mass Spectrom. 3:60-70. Mouradian, S., Rank, D.R., and Smith, L.M. 1996. Analyzing sequencing reactions from bacteriophage M13 by matrix-assisted laser desorption/ionization mass spectrometry. Rapid Commun. Mass Spectrom. 10:1475-1478. Murray, K.M. 1996. DNA sequencing by mass spectrometry. J. Mass Spectrom. 31:1203-1215. Niu, S., Zhang, W., and Chait, B.T. 1998. Direct comparison of infrared and ultraviolet wavelength matrix-assisted laser desorption/ionization mass spectrometry of proteins. J. Am. Soc. Mass Spectrom 9:1-7. Nordhoff, E., Ingendoh, A., Cramer, R., Overberg, A., Stahl, B., Karas, M., Hillenkamp, F., and Crain, P.F. 1992. Matrix-assisted laser desorption/ionization mass spectrometry of nucleic acids with wavelengths in the ultraviolet and infrared. Rapid Commun. Mass Spectrom. 6:771-776. Nordhoff, E., Cramer, R., Karas, M., Hillenkamp, F., Kirpekar, F., Kristiansen, K. and Roepstorff, P. 1993. Ion stability of nucleic acids in infrared matrix-assisted laser desorption/ionization mass spectrometry. Nucleic Acids Res. 21:3347-3357. Nordhoff, E., Karas, M., Cramer, R., Hahner, S., Hillenkamp, F., Lezius, A., Kirpekar, F., Kristiansen, K., Muth, J., Meier, C., and Engels, J.W. 1995. Direct mass spectrometric sequencing of low pmol amounts of oligodeoxynucleotides up to 21 bases by matrix-assisted laser desorption/ionization mass spectrometry. J. Mass Spectrom. 30:99-112. Nordhoff, E., Kirpekar, F., and Roepstorff, P. 1996. Mass spectrometry of nucleic acids. Mass Spectrom. Rev. 15:67-138. Parr, G.R., Fitzgerald, M.C., and Smith, L.M. 1992. Matrix-assisted laser desorption/ionization mass spectrometry of synthetic oligodeoxyribonucleotides. Rapid Commun. Mass Spectrom. 6:369-372.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.1.23 Current Protocols in Nucleic Acid Chemistry
Pieles, U., Zürcher, W., Schär, M., and Moser, H.E. 1993. Matrix-assisted laser desorption ionization time-of-flight mass spectrometry: A powerful tool for the mass and sequence analysis of natural and modified oligonucleotides. Nucleic Acids Res. 21:3191-3196. Polo, L.M., McCarley, T.D., and Limbach, P.A. 1997. Chemical sequencing of phosphorothioate oligonucleotides using matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Anal. Chem. 69:1107-1112. Roskey, M.T., Juhasz, P., Smirnov, I.P., Takach, E.J., Martin, S.A., and Haff, L.A. 1996. DNA Sequencing by delayed extraction-matrix-assisted laser desorption/ionization time of flight mass spectrometry. Proc. Natl. Acad. Sci. U.S.A. 93:4724-4729. Ross, P.L. and Belgrader, P. 1997. Analysis of short tandem repeat polymorphisms in human DNA by matrix-assisted laser desortpion/ionization mass spectrometry. Anal. Chem. 69:3966-3972. Ross, P.L., Lee, K., and Belgrader, P. 1997. Discrimination of single-nucleotide polymorphisms in human DNA using peptide nucleic acid probes detected by MALDI-TOF mass spectrometry. Anal. Chem. 69:4197-4202. Schneider, K. and Chait, B.T. 1993. Matrix-assisted laser desorption mass spectrometry of homopolymer oligodeoxyribonucleotides. Influence of base composition on the mass spectrometric response. Org. Mass Spectrom. 28:1353-1361. Schneider, K. and Chait, B. 1995. Increased stability of nucleic acids containing 7-deaza-guanosine and 7-deaza-adenosine may enable rapid DNA sequencing by matrix-assisted laser desorption mass spectrometry. Nucleic Acids Res. 23:1570-1575. Schuette, J.M., Pieles, U., Maleknia, S.D., Srivatsa, G.S., Cole, D.L., Moser, H.E., and Afeyan, N.B. 1995. Sequence analysis of phosphorothioate oligonucleotides via matrix-assisted laser desorption ionization time-of-flight mass spectrometry. J. Pharmaceut. Biomed. Anal. 13:1195-1203. Shaler, T.A., Tan, Y., Wickham, J.N., Wu, K.J., and Becker, C.H. 1995. Analysis of enzymatic DNA sequencing reactions by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Rapid Commun. Mass Spectrom. 9:942-947. Shaler, T.A., Wickham, J.N., Sannes, K.A., Wu, K.J., and Becker, C.H. 1996. Effect of impurities on the matrix-assisted laser desorption mass spectra of single-stranded oligodeoxynucleotides. Anal. Chem. 68:576-579. Simmons, T.A. and Limbach, P.A. 1997. The use of a co-matrix for improved MALDI-TOFMS analysis of oligonucleotides. Rapid Commun. Mass Spectrom. 11:567-572. Simmons, T.A. and Limbach, P.A. 1998. Influence of co-matrix proton affinity on oligonucleotide ion stability in matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. J. Am. Soc. Mass Spectrom. 9:668-675. Smirnov, I.P., Roskey, M.T., Juhasz, P., Takach, E.J., Martin, S.A., and Haff, L.A. 1996. Sequencing oligonucleotides by exonuclease digestion and delayed extraction matrix-assisted laser desorption ionization time-of-flight mass spectrometry. Anal. Biochem. 238:19-25. Spengler, B. 1997. Post-source decay analysis in matrix-assisted laser desorption/ionization mass spectrometry of biomolecules. J. Mass Spectrom. 32:1019-1036. Spengler, B. and Bokelmann, V. 1993. Angular and time resolved intensity distributions of laser-desorbed matrix ions. Nuclear Instruments and Methods in Physics Res. B. 82:379-385. Stemmler, E.A., Buchanan, M.V., Hurst, G.B., and Hettich, R.L. 1995. Analysis of modified oligonucleotides by matrix-assisted laser desorption/ionization fourier transform mass spectrometry. Anal. Chem. 67:29242930. Strupat, K., Karas, M., and Hillenkamp, F. 1991. 2,5-Dihydroxybenzoic acid: A new matrix for laser desorption-ionization mass spectrometry. Int. J. Mass Spectrom. Ion Process. 111:89-102. Tang, K., Taranenko, N.I., Allman, S.L., Chang, L.Y., and Chen, C.H. 1994a. Detection of 500-nucleotide DNA by laser desorption mass spectrometry. Rapid Commun. Mass Spectrom. 8:727-730. Tang, K., Taranenko, N.I., Allman, S.L., Chen, C.H., Ch’ang, L.Y., and Jacobson, K.B. 1994b. Picolinic acid as a matrix for laser mass spectrometry of nucleic acids and proteins. Rapid Commun. Mass Spectrom. 8:673-677. Tang, W., Nelson, C.M., Zhu, L., and Smith, L.M. 1997a. Positive ion formation in the ultraviolet matrix-assisted laser desorption/ionization analysis of oligonucleotides by using 2,5-dihydroxybenzoic acid. J. Am. Soc. Mass Spectrom. 8:218-224. Tang, W., Zhu, L., and Smith, L.M. 1997b. Controlling DNA fragmentation in MALDI-MS by chemical modification. Anal. Chem. 69:302-312. Analysis of Oligonucleotides
Tolson, D.A. and Nicholson, N.H. 1998. Sequencing RNA by a combination of exonuclease digestion and uridine specific chemical cleavage using MALDI-TOF. Nucleic Acids Res. 26:446-451.
10.1.24 Current Protocols in Nucleic Acid Chemistry
Vertes, A., Irinyi, G., and Gijbels, R. 1993. Hydrodynamic model of matrix-assisted laser desorption mass spectrometry. Anal. Chem. 65:2389-2393. Vestal, M.L., Juhasz, P., and Martin, S.A. 1995. Delayed extraction matrix-assisted laser desorption time-offlight mass spectrometry. Rapid Commun. Mass Spectrom. 9:1044-1050. Vorm, O., Roepstorff, P., and Mann, M. 1994. Improved resolution and very high sensitivity in MALDI-TOF of matrix surfaces made by fast evaporation. Anal. Chem. 66:3281-3287. Wada, Y. and Yamamoto, M. 1997. Detection of single-nucleotide mutations including substitutions and deletions by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Rapid Commun. Mass Spectrom. 11:1657-1660. Wang, B.H. and Biemann, K. 1994. Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry of chemically modified oligonucleotides. Anal. Chem. 66:1918-1924. Wu, K.J., Steding, A., and Becker, C.H. 1993. Matrix-assisted laser desorption time-of-flight mass spectrometry of oligonucleotides using 3-hydroxypicolinic acid as an ultraviolet-sensitive matrix. Rapid Commun. Mass Spectrom. 7:142-146. Wu, K.J., Shaler, T.A., and Becker, C.H. 1994. Time-of-flight mass spectrometry of underivatized singlestranded DNA oligomers by matrix-assisted laser desorption. Anal. Chem. 66:1637-1645. Zhu, L., Parr, G.R., Fitzgerald, M.C., Nelson, C.M., and Smith, L.M. 1995. Oligodeoxynucleotide fragmentation in MALDI/TOF mass spectrometry using 355-nm radiation. J. Am. Chem. Soc. 117:6048-6056. Zhu, Y.F., Chung, C.N., Taranenko, N.I., Allman, S.L., Martin, S.A., Haff, L., and Chen, C.H. 1996a. The study of 2,3,4-trihydroxyacetophenone and 2,4,6-trihydroxyacetophenone as matrices for DNA detection in matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Rapid Commun. Mass Spectrom. 10:383-388. Zhu, Y.F., Taranenko, N.I., Allman, S.L., Martin, S.A., Haff, L., and Chen, C.H. 1996b. The effect of ammonium salt and matrix in the detection of DNA by MALDI-TOF mass spectroscopy. Rapid Commun. Mass Spectrom. 10:1591-1596.
Contributed by Chau-Wen Chou and Patrick A. Limbach Louisiana State University Baton Rouge, Louisiana
Purification and Analysis of Synthetic Nucleic Acids and Components
10.1.25 Current Protocols in Nucleic Acid Chemistry
Analysis of Oligonucleotides by Electrospray Ionization Mass Spectrometry
UNIT 10.2
Because of the high molecular weights and thermal lability of biomolecules such as nucleic acids and proteins, the analysis of these molecules by mass spectrometry is a rather difficult process. Two major problems in such analyses are the need for a “soft” ionization method capable of generating intact molecular ions and the limited upper mass range of most mass analyzers. In the mid 1980s, Fenn and co-workers demonstrated that electrospray ionization (ESI) could be used to analyze molecules with molecular weights (Mr) larger than the mass-to-charge ratio (m/z) limit of the mass analyzer (Fenn et al., 1989). Later work on oligonucleotides opened the door to accurate, high-resolution analysis of these compounds by ESI-MS (Covey et al., 1988). ESI allows for the analysis of high-molecular-weight compounds through the generation of multiply charged ions in the gas phase. Because the basis of the mass spectrometric measurement is the m/z value of the molecule, the presence of multiple charges on the molecule will result in a decrease in the m/z values and allow characterization using mass analyzers with limited m/z ranges. A typical example of the electrospray mass spectra one obtains for oligonucleotides is shown in Figure 10.2.1. The sample contains three unique
Figure 10.2.1 Representative electrospray mass spectrum of a mixture of oligonucleotides. X = 17-mer, Y = 16-mer, and Z = 15-mer. The characteristic feature of an electrospray mass spectrum is the presence of multiply charged ions of each analyte.
Purification and Analysis of Synthetic Nucleic Acids and Components
Contributed by Lenore M. Polo and Patrick A. Limbach
10.2.1
Current Protocols in Nucleic Acid Chemistry (2000) 10.2.1-10.2.20 Copyright © 2000 by John Wiley & Sons, Inc.
Figure 10.2.2
Diagram of a typical electrospray source.
oligonucleotides—a 17-mer, a 16-mer, and a 15-mer—designated X, Y, and Z, respectively. As is illustrated in this spectrum, each of the oligonucleotide species are detected as multiply charged negative ions. Thus, even though the molecular weights of each oligonucleotide are greater than the m/z range of the mass analyzer (here 2000), the sample can still be identified due to the presence of the multiply charged peaks. A discussion on identification of oligonucleotide-ion charge state and molecular weight will be presented later in this unit. A necessary requirement for ESI is that the analyte molecules be charged in solution. The negatively charged phosphate backbone of oligonucleotides allows for negative-ion-mode analysis of ESI-generated ions. A diagram of a typical electrospray source is shown in Figure 10.2.2. Transfer of ions from solution phase to the gas phase is accomplished by generating an electric field between a spraying needle that is held at a high negative potential and a counter-electrode held at ground or a positive potential some distance from the needle. The solution being sprayed exits the needle as a conical distribution of droplets (“Taylor cone”), each containing excess negative charge. A heated drying gas, such as nitrogen, is typically used to assist evaporation of the solvent sheath from the ion. The desolvated, multiply charged ion is then introduced into the mass spectrometer for analysis (Gaskell, 1997). A number of applications of ESI to oligonucleotide and nucleic acid analysis have been reported since the introduction of this technique. Among the applications that will be discussed here are molecular mass measurement, sequence identification, and analysis of noncovalent complexes. In addition, the ESI source can be readily coupled with a number of separation techniques such as liquid chromatography (Apffel et al., 1997a,b; Glover et al., 1995), capillary electrophoresis (Barry et al., 1996), and capillary electrochromatography (Ding and Vouros, 1997) for the analysis of mixtures of oligonucleotides. ESI-MS VERSUS MALDI-MS FOR OLIGONUCLEOTIDE ANALYSIS
Analysis of Oligonucleotides by Electrospray Ionization Mass Spectrometry
Matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS) for oligonucleotide analysis is discussed in UNIT 10.1. Here, a brief comparison of the two approaches for oligonucleotide analysis is presented; a summary is provided in Table 10.2.1. The advantages of ESI-MS include higher mass accuracies, ease of coupling to on-line separation methods, and the ability to characterize noncovalent interactions; it is also the preferred method for gas-phase MS sequencing (tandem MS). MALDI-MS is more tolerant of sample contaminants, can handle complex mixture analysis, and is the preferred method for sequencing by an exonuclease digestion protocol. In general, both methods are capable of providing molecular weight and sequence information from oligonucleotides, and the choice of ionization method depends on the available instrumentation, type of analyses desired, and user preference.
10.2.2 Current Protocols in Nucleic Acid Chemistry
Table 10.2.1 Summary of the Characteristics of MALDI-MS Versus ESI-MS
Parameter or characteristic Mass errors Upper mass limit Feasibility of: Mixture analysis Easy coupling to separation methods Exonuclease sequencing Sequencing via gas-phase dissociation (MS/MS) Characterization of noncovalent interactions
MALDI-MS 1.0–0.05% ~36,000
ESI-MS 0.1–0.005% ~40,000
Yes No Preferred Possible
Difficult Yes Possible Preferred
No
Yes
CONVENTIONAL VERSUS NANOELECTROSPRAY CONFIGURATION ESI is inherently a solution-based ionization technique. The sample solution flows through the charged ESI needle, resulting in the formation of the Taylor cone mentioned above. Currently, there are two different flow-rate/needle configurations utilized in ESI-MS: conventional electrospray, and micro- or nanoelectrospray. Conventional electrospray is performed using stainless steel needles (with 0.15 to 0.41-mm-i.d. orifices) and the sample solution is delivered at a flow rate of 1 to 10 µL/min. Many of the original ESI-MS oligonucleotide analysis results were obtained using this configuration. Recently it has been shown that spraying capillaries with a 1- to 2-µm-i.d. spraying orifice have several advantages over the traditional ESI needle, particularly in biomolecular analyses (Wilm and Mann, 1996). This “nanoelectrospray” source is essentially a capillary that has been pulled to a fine tip. The small spraying orifice requires low flow rates (nL/min), which aids in sample conservation. In addition, the overall efficiency of desolvation, ionization, and transfer is increased. The small size of the droplets generated means that one analyte molecule is present per droplet. Because the size of the droplets is significantly reduced, desolvation is easily achieved without the need of a drying gas. The overall charge-to-volume ratio of the droplets is also higher than in conventional ESI, which improves ionization. Transfer of ions into the mass analyzer is improved because the spraying tip can be placed close to the orifice of the analyzer (1 to 2 mm away). Another important advantage of the nanoelectrospray source is that it has a higher tolerance to salt adduction than conventional ESI. In addition, it can operate at high pH. As described in the section on Sample Preparation, below, both of these characteristics can be useful in nucleic acid analysis. MASS ANALYZER CONFIGURATION ESI can be coupled with a large variety of mass analyzers. Table 10.2.2 lists the analyzers most often coupled to the ESI source. Commercial versions of these instruments are available for each of the different mass analyzers listed. Quadrupole mass analyzers are the most popular configuration due to their ease of operation and low cost. However, they suffer from poor resolution as well as a limited m/z range (∼3000). Triple-quadrupole instruments can be used to perform tandem MS experiments, and therefore are suitable when gas-phase sequencing experiments are of interest. Two common mass analyzers that are useful for several types of analysis of oligonucleotides are the quadrupole ion-trap and the Fourier-transform ion-cyclotron resonance mass spectrometer (FTICR-MS; see Table 10.2.3 for definitions of the instrument configurations discussed in this unit). These instruments have an added advantage over
Purification and Analysis of Synthetic Nucleic Acids and Components
10.2.3 Current Protocols in Nucleic Acid Chemistry
Table 10.2.2 Mass Analyzers Typically Coupled to ESI Sourcesa
Analyzerb
Resolution
Sensitivity
Mass accuracy
TOF Quadrupole Triple-quadrupole Double-focusing sector Ion-trap FTICR
Good Fair Fair High
High Good High Fair
High Fair Fair High
Fair Very high
Good Good
Fair Very high
Upper m/z limit ~40,000 3,000 3,000 10,000 3,000 10,000
Tandem MS capability No No Yes Yes Yes Yes
aThe relative performance aspects of each analyzer are also included. bSee Table 10.2.3 for definitions of instrument configurations.
the other mass analyzers listed in Table 10.2.2 that they operate on the principle of ion trapping. With trapped-ion mass analyzers, the analyte ions are confined to a particular region of space and their subsequent analysis is performed as a function of time. In contrast, analysis of ions with quadrupole, triple-quadrupole, sector, and time-of-flight mass analyzers is performed by separating the analysis regions in space. Thus, trapped-ion instruments are ideal for performing multiple stages of tandem mass spectrometry because no additional hardware is needed (McLuckey and Habibi-Goudarzi, 1993; Little et al., 1994a; Habibi-Goudarzi and McLuckey, 1995; Little and McLafferty, 1995). Another advantage is that these instruments permit alternative dissociation schemes—be-
Table 10.2.3 Glossary of Mass Spectrometry Terms Used in This Unit
Collision-induced dissociation (CID) Double-focusing mass spectrometer Fourier-transform ion-cyclotron resonance mass spectrometer (FTICR-MS)
Ion-trap mass spectrometer Quadrupole mass spectrometer
Tandem mass spectrometer (MS/MS) Time-of-flight mass spectrometer (TOF-MS)
Triple-quadrupole mass spectrometer Analysis of Oligonucleotides by Electrospray Ionization Mass Spectrometry
Process in which an ion is dissociated as a result of interaction with a target neutral species Mass spectrometer that uses direction and velocity to focus ions of the same mass and charge Mass spectrometer in which ions are confined by an electric and magnetic field and detected on the basis of their cyclotron frequency at a particular excitation energy in the high magnetic field Mass spectrometer in which ions can be confined for extended periods of time by the use of a quadrupolar electric field Mass spectrometer in which ions with a desired m / z are focused towards the detector by the use of a static and a high-frequency electric field Mass spectrometer in which ions of selected m / z pass through two or more stages of analysis Mass spectrometer which operates on the principle that ions of different m / z with the same initial energy require different times to travel through a field-free region Mass spectrometer composed of three sets of quadrupole rods, the first and last set operated in a manner similar to the quadrupole mass spectrometer and the second (middle) set used as a collision cell to allow for MS/MS experiments
10.2.4 Current Protocols in Nucleic Acid Chemistry
sides the common collision-induced dissociation (CID) method—to be employed (Little et al., 1994b, 1996). Quadrupole ion-trap MS typically gives poorer resolution and mass accuracy than FTICR-MS. FTICR-MS offers the highest resolution and mass accuracy, but the instrumentation is not only more expensive, but also more difficult to operate. Two additional mass analyzers can be used with ESI. A promising combination is the use of a time-of-flight (TOF) mass analyzer with an ESI source. At the time of writing, only first-generation ESI-TOF mass spectrometers are available and the ultimate effectiveness of the combination is not yet known. However, their high sensitivity, extended upper m/z range, adequate resolution and mass accuracy, and high duty cycle suggests that ESI-TOF mass analyzers may soon replace quadrupole mass analyzers as the instruments of choice for molecular weight determinations using ESI-MS. Double-focusing sector instruments are suitable for use with electrospray sources, have high sensitivities and mass resolution, and have tandem MS capabilities. However, their complexity and the small number of vendor offerings has limited their use for oligonucleotide analysis by ESI-MS. SAMPLE PREPARATION The success of oligonucleotide analyses using ESI depends largely on sample preparation. When dealing with nucleic acids, the two key factors in sample preparation are reduction of cation adducts (oligonucleotide purification and desalting) and selection of a proper solvent system. Purification Methods The greatest difficulty in achieving accurate mass measurement of oligonucleotides and nucleic acids is the formation of cation adducts. The negative charge on the nucleic acid phosphate backbone results in a large degree of Coulombic strain. In solution, solvent molecules help reduce these Coulombic interactions. In the gas phase, where solvent molecules are absent, relief of the strain is achieved by neutralization or cation adduction. Figure 10.2.3 is a representative electrospray mass spectrum of an oligonucleotide with a large number of cation adducts. The presence of nonvolatile cation adducts such as sodium and potassium results in spectral peaks broadened by the adduction. In addition, the ion current is shared over a larger number of peaks, thereby decreasing sensitivity. As illustrated in Figure 10.2.3b, a number of cation-adduct combinations may be present for a particular charge state, and a complex mass spectrum results. A number of purification techniques have been applied to nucleic acid samples (Stults and Marsters, 1991; Emmett and Caprioli, 1994; Little et al., 1994a; Potier et al., 1994; Greig and Griffey, 1995; Limbach et al., 1995; Liu et al., 1996, 1997; Muddiman et al., 1996a). Although these methods differ in protocol, the objective and end result are the same—to replace the nonvolatile (e.g., sodium and potassium) adducts with volatile ones. Most commonly, ammonium salts are exchanged for the nonvolatile adducts. Purification techniques that reduce or eliminate the presence of cation adducts can be classified into two groups: off-line and on-line methods (Table 10.2.4). The former include ammonium acetate precipitation , dialysis, and HPLC purification. The latter include on-line microdialysis, on-line micro-HPLC, and solvent additives. By far the most common purification method is ammonium acetate precipitation (see Basic Protocol 1; Stults and Marsters, 1991; Limbach et al., 1995). In this procedure, the oligonucleotide is mixed with ammonium acetate and the oligonucleotide in its ammonium salt form is precipitated from ethanol. This procedure can be repeated a number of times, and it is advisable to do so, as multiple precipitations increase the overall amount of salt exchange.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.2.5 Current Protocols in Nucleic Acid Chemistry
Figure 10.2.3 Representative electrospray mass spectrum used to illustrate the detrimental effects of cation adduction on mass spectral quality.
Analysis of Oligonucleotides by Electrospray Ionization Mass Spectrometry
Another routine approach to reducing salt adduct formation is to sequester the adducts using chelating agents and/or organic bases (see Basic Protocol 2; Potier et al., 1994; Greig and Griffey, 1995; Limbach et al., 1995; Muddiman et al., 1996a). The strategy here is to add chelating agents such as trans-1,2-diaminocyclohexane-N,N,N′N′-tetraacetic acid (CDTA; Limbach et al., 1995) or organic bases (Greig and Griffey, 1995; Muddiman et al., 1996a) such as triethylamine (TEA) that have a higher cation affinity than the nucleic acid. The cations will bind preferentially to these additives, leaving the nucleic acid in its free acid form. It is important to note that none of these purification techniques alone can remove all cation adducts, and using a combination of techniques—for example, ammonium acetate precipitation followed by CDTA and TEA addition—will improve results (Greig and Griffey, 1995; Limbach et al., 1995).
10.2.6 Current Protocols in Nucleic Acid Chemistry
Table 10.2.4 Techniques for Purifying Nucleic Acid Samples Prior to ESI-MS Analysis
Method Off line Ammonium acetate precipitation Reversed-phase HPLC Microdialysis On line Addition of co-matrix and/or chelating agent Microdialysis Microscale liquid chromatography
Reference Stults and Marsters (1991); Limbach et al. (1995) Little et al. (1994a) Liu et at. (1997) Potier et al. (1994); Greig and Griffey (1995); Limbach et al. (1995) Liu et al. (1996) Emmett and Caprioli (1994)
Removal of Nonvolatile Cation Adducts from Oligonucleotide or Nucleic Acid Samples by Ammonium Acetate Precipitation
BASIC PROTOCOL 1
The amount of salt adduction in nucleic acid samples can be reduced by exchanging the nonvolatile cations with the volatile ammonium cation (see section on Purification Methods, above). The effectiveness of the cation exchange increases with repetition of the precipitation procedure (Stults and Marsters, 1991; Limbach et al., 1995). This protocol should only be used for oligonucleotides larger than 25 bases—smaller oligonucleotides are difficult to precipitate and large sample losses will occur. Materials Oligonucleotide or nucleic acid sample, dry 10 M ammonium acetate Absolute and 70% ethanol, ice cold CAUTION: RNA samples are easily degraded by adventitious nucleases. If RNA is to be purified or analyzed, all laboratory equipment must be sterilized prior to use. Use of gloves is recommended during preparation of nucleic acid samples and solutions to avoid contamination of glassware and other equipment. Autoclaved distilled water and the highest available grade of salts must be used during solution preparation. 1. Dissolve the nucleic acid in 40 to 70 µL ultrapure water to generate a 250 to 400 µM solution. 2. Add 1⁄3 volume of 10 M ammonium acetate. 3. Add 2.5 vol ice-cold absolute ethanol to precipitate the nucleic acid. Store at least 3 hr at −20°C. 4. Centrifuge the suspension 15 min at 12,500 rpm, 25°C. 5. Decant the supernatant, working carefully to avoid disturbing the pellet. 6. Wash the pellet with 70% aqueous ethanol. Store the suspension at least 2 hr at −20°C. 7. Centrifuge the suspension 15 min at 12,500 rpm. 8. Decant the supernatant and lyophilize the pellet. 9. Dissolve the pellet in ultrapure water to a final concentration of 300 to 1000 µM.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.2.7 Current Protocols in Nucleic Acid Chemistry
BASIC PROTOCOL 2
Removal of Nonvolatile Cation Adducts from Oligonucleotide or Nucleic Acid Samples with Chelating Agents and/or Organic Bases and ESI-MS Sample Preparation This protocol describes the steps for preparing oligonucleotide and nucleic acid solutions for analysis by ESI-MS. The addition of chelating agents or organic bases serves to sequester cations, thereby reducing the extent to which adducts are observed in the mass spectra (see section on Purification Methods, above). Each of these agents will reduce the amount of adduct formation, but the results are usually better when both materials are used. This protocol is the recommended choice for analyzing small oligonucleotides (≤25 bases). For larger oligonucleotides, this procedure should be used in conjunction with ammonium acetate precipitation (see Basic Protocol 1). Materials Oligonucleotide or nucleic acid sample trans-1,2-Diaminocyclohexane-N,N,N′,N′-tetraacetic acid (CDTA) Ethylenediaminetetraacetic acid (EDTA) 0.1% aqueous triethylamine Appropriate solvent: 30% to 70% aqueous acetonitrile, isopropanol, or methanol (see section on Solvent Selection, below, for discussion of how to choose an appropriate solvent.) CAUTION: RNA samples are easily degraded by adventitious nucleases. If RNA is to be purified or analyzed, all laboratory equipment must be sterilized prior to use. Use of gloves is recommended during preparation of nucleic acid samples and solutions to avoid contamination of glassware and other equipment. Autoclaved distilled water and the highest available grade of salts must be used during solution preparation. 1. For oligonucleotides >25 bases, carry out ammonium acetate precipitation (see Basic Protocol 1). 2. Prepare a 1 to 10 µM solution of oligonucleotide in the appropriate organic solvent. 3. Add 5 µL of 0.1% triethylamine to each 100 µL of oligonucleotide solution. 4. If analyzing intact nucleic acids, add CDTA or EDTA to the sample so as to obtain a final CDTA or EDTA molar concentration three times that of the nucleic acid sample. 5. This solution is now ready for immediate analysis by ESI-MS. Solvent Selection
Analysis of Oligonucleotides by Electrospray Ionization Mass Spectrometry
A second aspect of sample preparation is the selection of an appropriate solvent for electrospray. Two important criteria are (1) that the solvent readily evaporate to facilitate the transfer of the ion from the liquid to the gas phase, and (2) that the solvent allow the generation of a large number of ions. It has been shown that as the organic composition of the solvent increases, the signal intensity increases as well (Bleicher and Bayer, 1994). In that particular study, the best ion signals were seen with acetonitrile, although other solvents such as isopropanol or methanol have been used. Typically, the aqueous oligonucleotide solution is mixed in a 1:1 ratio with the organic solvent. This allows the oligonucleotide to remain solubilized while assisting the ESI process by improving evaporation. The second consideration in solvent selection is that the solvent must allow the ready formation of ions. The main factor in this case is solution pH: as pH increases, more negative ions are produced and the signal intensity rises (Bleicher and Bayer, 1994). This
10.2.8 Current Protocols in Nucleic Acid Chemistry
increase in pH can be achieved by adding an organic base such as TEA to the nucleic acid solution, which also reduces cation adduction, as previously discussed. Finally, the pH can be manipulated to change the charge-state distribution of the peaks observed in the spectra (Stults and Marsters, 1991; Cheng et al., 1995; Griffey et al., 1997). As the pH increases, the charge-state distribution shifts to higher charge states or lower m/z. Thus, when using a mass analyzer with a limited m/z range, it is preferable to work at high solution pH. The typical procedure for preparing samples for ESI-MS analysis is as follows. After purification of the oligonucleotide or nucleic acid sample by one of the off-line techniques mentioned above, the oligonucleotide sample is dissolved in ultrapure (e.g., Nanopure) water to a stock concentration of 50 to 100 µM. An aliquot of the stock solution is then diluted into the organic solvent of interest to a final concentration of 1 to 10 µM. Chelating agents or organic bases such as TEA can be added to this solution prior to analysis. The deciding criteria for final concentration and organic content of solvent depend on the particular type of information desired from the mass spectrometric step. When the molecular weight or sequence of single-stranded oligonucleotides or intact nucleic acids are to be determined, the percentage organic solvent should be between 50% and 90%; smaller oligonucleotides are more soluble at the higher percentages of organic solvent. The best concentration will vary by instrument. When noncovalent interactions of double-stranded oligonucleotides or intact nucleic acids are to be investigated, the organic content should remain as low as possible while still generating a stable electrospray (typically 10% to 35%). Analyte concentrations in these cases are typically higher (∼25 µM). MOLECULAR WEIGHT DETERMINATION USING ESI-MS Molecular weight determination of oligonucleotides and intact nucleic acids is one of the more common applications of ESI, as well as one of the simplest. This approach can be used to verify the expected base composition of an oligonucleotide by comparing the measured mass to the mass corresponding to the predicted composition. The analysis of small oligonucleotides (<10,000 Da) has become routine. In fact, if constraints can be placed on the number of A, C, G, or U/T residues allowed in the molecule, then the base composition of an oligonucleotide up to the 14-mer level can be determined unambiguously by accurate mass measurement alone (Pomerantz et al., 1993). For larger oligonucleotides and nucleic acids, however, extremely high mass accuracies are required to confirm base composition assignments. The most impressive molecular weight measurements have been obtained using FTICRMS (Chen et al., 1995; Cheng et al., 1996a). With this high-resolution mass analyzer, DNA molecules in the 110-MDa range have been detected with a 10% uncertainty in the molecular weight value (Chen et al., 1995). Other relatively large oligonucleotides and nucleic acids in the molecular weight region between 15,000 and 40,000 Da have also been analyzed. Among the applications of ESI-MS to these molecules are the analysis of intact tRNA and rRNA (Limbach et al., 1995) and PCR products (Naito et al., 1995; Muddiman et al., 1996b, 1997). In these cases, the errors in mass measurement are typically ∼0.01% or less. Because ESI produces multiply charged ions, the resulting spectra may be complicated by the presence of several ions of the same mass but different charge and their respective cation adducts. Determination of the charge state of a particular peak may be accomplished using mathematical equations. For a negatively charged peak, p1, one may write Equation 10.2.1:
Purification and Analysis of Synthetic Nucleic Acids and Components
10.2.9 Current Protocols in Nucleic Acid Chemistry
p1z1 = Mr − Maz1 = Mr − 1.0079 z1 Equation 10.2.1
where p1 is the m/z value of the peak, z1 is the charge of the peak, Mr is the molecular weight of the sample, and Ma is the molecular weight of the charge-carrying species. Typically, the charge-carrying species is a proton; therefore, Ma = 1.0079. Similarly, for a second peak, p2, that is j peaks away from p1, one may write Equation 10.2.2: p2(z1 − j) = Mr − Ma(z1 − j) = Mr − 1.0079(z1 − j) Equation 10.2.2
Solving Equations 10.2.1 and 10.2.2, Equation 10.2.3 is obtained for z1: z1 = −j(p2 + 1.0079) / | p1 − p2 | Equation 10.2.3
Once the charge state is known, Equation 10.2.1 can be used to determine the molecular weight of the sample being analyzed. The data in Figure 10.2.1 will serve to illustrate how charge-state and molecular weight determinations are made. To determine the charge state, for example, of the peak at 1348.9, the following values can be substituted into Equation 10.2.3: p1 = 1348.9, p2 = 1798.5, and j = 1. By substituting these values into Equation 10.2.3, a value of z1 of −4 is obtained for the charge state. Using Equation 10.2.1, Mr is found to be 5399.6. To obtain a more accurate value for Mr, it should be calculated from the values measured for a number of adjacent peaks and the calculated values then averaged, as shown in Figure 10.2.1. The value one obtains from solving Equation 10.2.3 should be an integer; if it is not, a different pair of m/z values should be chosen. For example, if one chooses p1 = 1348.9, p2 = 1697.7, and j = 1, a value of z1 = −4.87 is obtained for the charge state. Because this value is not an integer, one can conclude that the m/z values of 1348.9 and 1697.7 do not compose an oligonucleotide of the same mass. In some cases, multiple peaks are not present in the electrospray series or are difficult to discern (such as in tandem MS experiments), and some other means of determining the charge state of the ion is necessary. On a high-resolution instrument such as a sector or FTICR mass spectrometer, the ability to resolve isotope peaks is helpful in determining the charge state of a particular peak. If the spacing between a peak A and its consecutive isotope peak A+1 is 1 (e.g., for 13C), then this peak will correspond to the singly charged species. If, for example, the difference between the A and A+1 peaks detected in the mass spectrum is 0.20 u, then the A peak corresponds to the species carrying a −5 charge (i.e., 1⁄ 5 = 0.20 u). In general, if the difference between the A and A+1 peaks is 1/x u, the A peak will correspond to the −x charge state. Low-resolution instruments typically cannot resolve 13C isotopes for multiply charged peaks where z > 3. Often, the charge state of the particular ion can be determined by the mass shift of a cation adduct. For example, a single sodium adduct will appear at an m/z value 22 u higher than the unadducted m/z value for singly charged ions, 22⁄2 = 11 u higher for doubly charged ions, 22⁄3 = 7.3 u higher for triply charged ions, and so forth (see Fig. 10.2.3b).
Analysis of Oligonucleotides by Electrospray Ionization Mass Spectrometry
OLIGONUCLEOTIDE AND NUCLEIC ACID SEQUENCING USING ESI-MS A powerful application of ESI-MS in the analysis of oligonucleotides and nucleic acids is sequence determination. The two most common techniques for sequencing are tandem MS and indirect sequencing using enzyme digestions (Limbach, 1996). In tandem MS,
10.2.10 Current Protocols in Nucleic Acid Chemistry
A w3
x3
z3
y3
B1
O
O
O
P
c1
O O
O
a2
d1
b2
c2
d2
a3
1,2 elimination B2
d3
O
O O P O O
O
B2
O O
O O
OH
c3
b3
O
O
O H O P O O O
O
OH
Ade
O
3'
P
OH
OH
b1
z1
y1
B4
O
P
HO
x1
w1
B3
O
a1
z2
y2
B2
5'
B
x2
w2
O
O
a - B fragment
H O
O
HO P O O
O
O
B2
1,2 elimination
HO P O O
O
B2
O
w - type fragment
Figure 10.2.4 (a) Nomenclature for the dissociation products of oligonucleotides. (b) Prevalent oligonucleotide fragmentation mechanism, especially with ion-trap and FTICR-MS instruments (McLuckey and Habibi-Goudarzi, 1993). The mechanism involves two elimination reactions, the first resulting in loss of the base and the second resulting in cleavage at the sugar to yield (a-B)-type and w-type fragments.
molecules are allowed to dissociate in the gas phase, and analysis of the resulting fragmentation pattern yields insight into the structure and, in the case of oligonucleotides, sequence of the molecule. Figure 10.2.4a gives the nomenclature for assigning dissociation products of oligonucleotides. As can be seen in the figure, a large number of fragmentation pathways are possible. This large number of pathways results in highly complicated spectra, and as oligonucleotide length increases, so does the spectral complexity. Figure 10.2.4b shows the most common fragmentation pathway oligonucleotides are found to follow during tandem MS experiments. Generally, the resulting mass spectrum will be composed primarily of a-B fragments and the w-type fragments, although the other dissociation products may also be present (McLuckey and HabibiGoudarzi, 1993). Tandem MS analysis of oligonucleotides using ESI have been carried out using triple-quadrupole (Barry et al., 1995, 1996; Wolter and Engels, 1995; Boschenok
Purification and Analysis of Synthetic Nucleic Acids and Components
10.2.11 Current Protocols in Nucleic Acid Chemistry
and Sheil, 1996; Ni et al., 1996a), ion-trap (McLuckey et al., 1992, 1995; McLuckey and Habibi-Goudarzi, 1993; Habibi-Goudarzi and McLuckey, 1995), and FTICR mass analyzers (Little and McLafferty, 1995; Little et al., 1995, 1996). Although the techniques and the information generated by each of these mass analyzers are similar, there do exist substantial differences, primarily in the preferred order of base loss. For instance, in ion-trap and FTICR-MS studies, adenine is usually lost, and this leads to cleavage of the 3′ C-O bond of the sugar, generating w-type and (a-B)-type fragments (Fig. 10.2.4; McLuckey et al., 1992). In contrast, triple-quadrupole studies show no preference for loss of any one base over another (Barry et al., 1995; Boschenok and Sheil, 1996; Ni et al., 1996a). Recently, McCloskey and co-workers (Ni et al., 1996b) developed an algorithm for the sequence identification of oligonucleotides using a triple-quadrupole mass spectrometer equipped with an ESI source. The algorithm is based on the analysis of relative ion abundances and charge-state distributions of ions generated using low-energy collisionally induced dissociation (CID). In their preliminary study, the sequence of an “unknown” 15-mer was determined. In a subsequent study, the algorithm was employed to determine the sequence of oligonucleotides in combinatorial libraries composed of mixtures of 8-mers or 12-mers containing two and three unknown nucleotides in the sequence, respectively (Pomerantz et al., 1997). The greatest advantage presented by this technique for this application is the ability to sequence isomeric oligonucleotides. When analyzing mixtures using tandem MS, components of a mixture are analyzed by selecting (one at a time) each molecular ion in the mass spectrum. Once isolated, it is allowed to undergo CID, and the fragment-ion spectrum is analyzed to determine the sequence. If, however, the molecular ion selected in the primary mass spectrum represents two or more components of equal mass, the CID spectrum becomes more difficult to interpret. By employing the sequencing algorithm, the isomeric oligonucleotides can be distinguished and sequenced. The algorithm is not without its limitations, however. In the case of isomeric mixtures, as the number of components represented by the molecular ion in the primary mass spectrum increases, it becomes nearly impossible to determine the individual oligonucleotides unambiguously without a previous fractionation step. The task becomes even more difficult if the number of components in the mixture is unknown. In addition, as one of the factors used for sequence determination is relative ion abundance, the analysis of minor components becomes difficult in a mixture where one or more components are in a molar excess. Table 10.2.5 Characteristic Mass Losses During Exonuclease Digestion for Naturally Occurring Deoxynucleotides (dX) and Ribonucleotides (rX)a
Nucleotide
Analysis of Oligonucleotides by Electrospray Ionization Mass Spectrometry
dA dG dC dT rA rG rC ru
∆ mass (singly charged) 313.27 329.27 289.25 304.26 329.27 345.27 305.25 306.26
∆ mass (doubly charged) 156.64 164.64 144.63 152.13 164.64 172.64 152.62 153.13
∆ mass (triply charged) 104.42 109.76 96.417 101.42 109.76 115.09 101.75 102.09
aThe particular values are listed for several different charge states, and will be determined by the charge states of the two oligonucleotides used to find ∆ mass. All values are atomic weight based.
10.2.12 Current Protocols in Nucleic Acid Chemistry
Apart from these specific drawbacks in analyzing mixtures, it is uncertain whether the algorithm will be applicable when other mass analyzers or high-energy CID are employed. Nonetheless, its development represents a large step forward in the simplification of tandem MS analyses of oligonucleotides. Although it is questionable whether this technique will be applicable to larger molecules, it can clearly be used to analyze oligonucleotides up to the 15-mer range. In addition, the sequence location as well as the nature (i.e., sugar or base) of modifications can be determined. The second approach for sequencing using ESI-MS involves the partial enzymatic digestion of oligonucleotides or nucleic acids followed by mass-spectral analysis of the digestion products (see Basic Protocol 3). Enzymatic digestion may consist of either exonuclease or endonuclease digestion. Sequence identification using exonuclease digestion is a relatively straightforward, simple method, particularly for smaller oligonucleotides (Limbach et al., 1994; Glover et al., 1995). Exonuclease digestions generate consecutive backbone cleavages which, when analyzed using mass spectrometry, create a “mass ladder.” The sequence of the molecule is then determined from the mass difference between adjacent peaks in the spectrum. Table 10.2.5 lists the expected mass difference for loss of each of the deoxy- and ribonucleotides. 5′ and 3′ exonucleases can be used to generate 5′ and 3′ sequence information, respectively. Modified nucleosides can be easily identified with this method from the anomalous mass shifts caused by the modifications.
Oligonucleotide or Nucleic Acid Sequencing Using Sequential Exonuclease Digestion and ESI-MS
BASIC PROTOCOL 3
This protocol describes the procedure for on-line sequencing of oligonucleotides or nucleic acids using ESI-MS. To generate 3′ sequence information, phosphodiesterase I should be used, as it cleaves oligonucleotides in the 3′-to-5′ direction. Similarly, if 5′ sequence information is required, phosphodiesterase II should be used. During the first few minutes of analysis, information can be gathered on the first five or so nucleotides in the sequence. After longer reaction times, sequence information can be obtained for nucleotides further along the chain (Limbach et al., 1994). This protocol is most effective for oligonucleotides <30-mers. Materials Oligonucleotide or nucleic acid sample, dry Phosphodiesterase I (from snake venom, SVP; Sigma) or phosphodiesterase II (from calf spleen, CSP; Sigma or Worthington) dissolved to 0.1 U/µL in water 5.0 M ammonium acetate (APPENDIX 2A), pH 9.8 to 10.4 for phosphodiesterase I or 6 to 7 for phosphodiesterase II Organic solvent (see section on Solvent Selection, above) Variable-flow-rate syringe pump 1. Mix ∼1.2 nmol dry oligonucleotide, 10 µL (1.0 U) phosphodiesterase, and 0.5 M ammonium acetate at the appropriate pH to adjust the pH for optimum enzyme digestion. 2. Dilute the solution in an appropriate percentage of organic solvent to a concentration of 10 pmol/µL. 3. Draw the digestion mixture into a syringe and infuse the mixture into the ESI-MS at a flow rate of ∼2 µL/min using a syringe pump.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.2.13 Current Protocols in Nucleic Acid Chemistry
Digestion will occur in the syringe as the data is acquired. Data can be acquired continuously or every 2 min until digestion is complete.
4. The syringe may be cooled (to 0°C to 10°C) or heated (to 30°C to 45°C) in a water bath in order to decrease or increase digestion times, respectively. Although direct analysis of the digestion mixtures can be advantageous because the sequence information is obtained from a difference in mass rather than an absolute value, it is sometimes preferable to separate the digestion products before MS analysis (Glover et al., 1995). For example, one may wish to separate digestion products when dealing with oligonucleotides ≥30-mers, since the large number of peaks generated can decrease sensitivity.
Analysis of RNA Modifications Using Endonuclease Digestion and ESI-MS For larger nucleic acids, the generation of smaller fragments using endonucleases facilitates sequence assignments. One of the more significant advances in this area has been achieved by McCloskey and co-workers. This group has developed a protocol based on site-specific endonuclease digestions to determine modifications in transfer RNA (tRNA) and ribosomal RNA (rRNA; Crain, 1990; Bruenger et al., 1993; Kowalak et al., 1993, 1994, 1995, 1996). The steps involved in this sequencing method are outlined in Figure 10.2.5, and the enzymatic digestions used in this method are outlined in the protocols below. The purified RNA sample is divided in two. One of the samples is digested with nuclease P1 and alkaline phosphatase to generate its component nucleosides (see Basic Protocol 4). Liquid chromatography mass spectrometry (LC-MS) is then used to characterize the nucleosides present in the mixture, as well as to determine if any modifications are present (evidenced by a difference in mass from the “naturally” occurring nucleosides;
Analysis of Oligonucleotides by Electrospray Ionization Mass Spectrometry
Figure 10.2.5
Endonuclease-based RNA sequencing protocol.
10.2.14 Current Protocols in Nucleic Acid Chemistry
step 1a). The other RNA sample is subjected to RNase T1 digestion (see Basic Protocol 5) to generate oligonucleotide fragments terminating in a 3′-Gp (step 1b). The resulting oligonucleotides are then separated based on their chain length using DEAE-HPLC (step 2b). Each of the collected fractions (which typically contain more than one oligonucleotide) is then separated into two batches. One set is subjected to nuclease P1 and alkaline phosphatase digestion as in step 1a and the nucleosides and modified nucleosides present in each fraction are determined (step 3a). Each oligonucleotide fraction in the second set is analyzed using ESI-MS (step 3b). The accurate mass measurement obtained is then compared to the predicted masses from the known gene sequence from which the nucleotide sequence, as well as sites of modifications, can be determined (step 4b). Because the number of RNase T1 fragments that are generated can be quite large, an alternative procedure is to isolate regions of interest selectively by hybridizing complementary oligodeoxynucleotides to the region and using mung bean nuclease to digest the unhybridized sections of RNA (Kowalak et al., 1995). The purified oligoribonucleotides remaining may then be subjected to the protocol detailed in Figure 10.2.5.
Total Nucleoside Digestion of RNA
BASIC PROTOCOL 4
The following procedure can be used to digest RNA to its component nucleosides. This is a necessary step in the determination of post-transcriptional modifications using the method developed by McCloskey and co-workers. The amounts of enzyme specified are those required to completely digest ∼20 µg (0.5 A260 units) of RNA (Crain, 1990). Materials RNA sample, dry Nuclease P1 (Sigma), dissolved to 2 U/µL in 0.05 M ammonium acetate, pH 5.3 Snake venom phosphodiesterase I (SVP) or bovine intestinal phosphodiesterase (both from Sigma; the former is preferred), dissolved to 0.001 U/µL in water Bacterial alkaline phosphatase (Sigma), suspended in 2.5 M ammonium sulfate 1 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) 0.1 M ammonium acetate, pH 5.3 1 M ammonium bicarbonate, pH 7.8 37° to 100°C water bath CAUTION: RNA samples are easily degraded by adventitious nucleases. If RNA is to be purified or analyzed, all laboratory equipment must be sterilized prior to use. Use of gloves is recommended during preparation of nucleic acid samples and solutions to avoid contamination of glassware and other equipment. Autoclaved distilled water and the highest available grade of salts must be used during solution preparation. 1. Dissolve the RNA sample in 1 mM Tris⋅Cl, pH 7.4, to a concentration of 1 to 3 µg/µL. 2. Heat the solution 3 min at 100°C to denature the nucleic acid, and immediately chill on ice. 3. Add 1⁄10 vol of 0.1 M ammonium acetate, pH 5.3. 4. Add 2 U nuclease P1 and incubate the solution 2 hr at 45°C. If thiolated nucleosides are present, include 1⁄10 vol of 1 mM dithiothreitol in the buffer to reduce sulfur loss.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.2.15 Current Protocols in Nucleic Acid Chemistry
5. Add 1⁄10 vol of 1 M ammonium bicarbonate, pH 7.8, and 0.002 U phosphodiesterase I, and incubate 2 hr at 37°C. The pH of ammonium bicarbonate will increase with time, so the pH of this buffer should be checked, and adjusted if necessary by addition of acetic acid, prior to use.
6. Add 0.5 U alkaline phosphatase and incubate 1 hr at 37°C. Bacterial alkaline phosphatase from Sigma is provided as a suspension in 2.5 M ammonium sulfate. The ammonium sulfate does not typically interfere with subsequent LC/MS analysis. If dry alkaline phosphatase is used, dissolve the enzyme to 0.5 U/mL in 2.5 M ammonium sulfate.
7. The solution is ready for immediate analysis by LC/MS. For details of LC-MS analysis of RNA, consult Pomerantz and McCloskey (1990). The digest can be stored at –20°C before use; in this case it must first be neutralized with acetic acid. BASIC PROTOCOL 5
RNase T1 Digestion of RNA Total nucleoside digestion of RNA will yield information on the kinds of post-transcriptional modifications present in RNA, but analysis of RNase T1 products is required to identify the sequence location of these modifications. The following is the procedure for generating RNase T1 fragments of RNA (Kowalak et al., 1993). Materials RNA sample, dry RNase T1 (Sigma or Ambion), suspended in 3.2 M ammonium sulfate, pH ~6 1 mM EDTA/50 mM Tris⋅Cl, pH 7.5 (see APPENDIX 2A for preparation of EDTA and Tris⋅Cl stock solutions) 37°C water bath CAUTION: RNA samples are easily degraded by adventitious nucleases. If RNA is to be purified or analyzed, all laboratory equipment must be sterilized prior to use. Use of gloves is recommended during preparation of nucleic acid samples and solutions to avoid contamination of glassware and other equipment. Autoclaved distilled water and the highest available grade of salts must be used during solution preparation. 1. Dissolve 8 nmol RNA in 1 mM EDTA/50 mM Tris⋅Cl, pH 7.5, to a concentration of 0.2 nmol/µL. 2. Heat the solution 3 min at 100°C to denature the nucleic acid, and immediately chill on ice. 3. Add 2000 U RNase T1 and incubate 30 min at 37°C. RNase T1 from Sigma is supplied as a suspension in 3.2 M ammonium sulfate. The ammonium sulfate does not typically interfere with subsequent HPLC analysis. If dry RNase T1 is to be used, dissolve the enzyme to 1000 U/mL in 3.2 M ammonium sulfate.
4. The solution is ready for immediate analysis by HPLC. Analysis of Oligonucleotides by Electrospray Ionization Mass Spectrometry
For details of HPLC analysis of RNA, consult Kowalak et al., 1993. The digest can be stored at –20°C prior to HPLC, although long-term storage (>6 months) is not recommended as it may lead to RNA degradation.
10.2.16 Current Protocols in Nucleic Acid Chemistry
ANALYSIS OF NONCOVALENT COMPLEXES USING ESI-MS Among the advantages of ESI in nucleic acid analysis is that this is what is termed a “soft” ionization technique in the sense that the ionization process itself does not cause molecular fragmentation. There are several classes of noncovalent complexes that can be studied using ESI-MS (Przybylski and Glocker, 1996; Loo, 1997). Among these are specific complex stoichiometry, competition of complex components, specificity of solution conditions for complex formation, and gas-phase stability of complex ions. Each of these properties can be studied by varying certain ESI parameters. For example, studies on complex stoichiometry can be achieved by changing solution concentrations, ESI temperature, and ESI voltage. Competitions between components in a complex can be studied by inducing modifications in the components and monitoring the changes effected on the ESI spectra. Solution requirements are determined by changing such variables as buffer pH. Lastly, the gas-phase stability of the complexes may be determined using CID experiments (Przybylski and Glocker, 1996). In addition, ESI-MS has several advantages over the other techniques traditionally used to study noncovalent complexes (e.g., NMR, X-ray crystallography), including the ability to accurately measure molecular mass and the smaller amounts of sample required. The earliest studies of nucleic acid complexes were carried out on oligonucleotide duplexes. Typically, the observation of oligonucleotide duplexes is achieved with samples dissolved in 10 M ammonium acetate. The duplex is usually observed at low charge states (high m/z), requiring the use of an analyzer with an extended mass range. Besides small oligonucleotide duplexes, double-stranded PCR products have also been observed (Wunschel et al., 1996). The complexes studied consisted of 100 to 105 bp. However, accurate mass measurement of such large complexes is difficult to obtain even on a high-resolution instrument because of the large amount of cation adduction present in these samples. Quadruplex structures of DNA have also been observed using ESI-MS (Goodlett et al., 1993). However, formation of these complexes requires the presence of high concentrations of monatomic cations (e.g., Na+) which, as previously discussed, degrade spectral quality. In addition to base-paired nucleic acid hybridizations, ESI-MS has also been used to analyze the binding of proteins to DNA and RNA. Among the studies that have been conducted on protein-nucleic acid complexes are determination of gas-phase stability (Gale and Smith, 1995), recognition studies (Sannes-Lowery et al., 1997), measurement of dissociation constants (Greig et al., 1995), determination of binding stoichiometry (Cheng et al., 1996b) and specificity (Gao et al., 1995), and competitive binding studies (Cheng et al., 1996c). ESI-MS for the study of noncovalent complexes is still a developing technique. Apart from the fundamental studies of noncovalent complexes, this methodology may have more far-reaching applications. Among the proposed applications (Przybylski and Glocker, 1996) are sequence determination by hybridization, analysis of antisense oligonucleotide complexes, and analysis of intercalation complexes. SUMMARY The speed and sensitivity of mass spectrometric analyses make this technique an attractive approach to analyzing oligonucleotides and nucleic acids. With the advent of “soft” ionization techniques such as MALDI and ESI, the analysis of these molecules in their intact form has brought mass spectrometry into the forefront as a viable method for characterization of nucleic acids. Although MALDI has a greater tolerance for salt adducts than ESI, the latter technique has the advantage that the generation of multiply charged ions allows the analysis of these molecules using more routine analyzers such as
Purification and Analysis of Synthetic Nucleic Acids and Components
10.2.17 Current Protocols in Nucleic Acid Chemistry
quadrupoles that may have a limited m/z range. In addition, ESI-MS allows the analysis of intact noncovalent complexes. This characteristic permits the study of these complexes in the gas-phase with minimal sample amounts, and provides complementary information to the classic solution phase methods. LITERATURE CITED Apffel, A., Chakel, J.A., Fischer, S., Lichtenwalter, K., and Hancock, W.S. 1997a. Analysis of oligonucleotides by HPLC–electrospray ionization mass spectrometry. Anal. Chem. 69:1320-1325. Apffel, A., Chakel, J.A., Fischer, S., Lichtenwalter, K., and Hancock, W.S. 1997b. New procedure for the use of high-performance liquid chromatography–electrospray ionization mass spectrometry for the analysis of nucleotides and oligonucleotides. J. Chromatogr. A 777:3-21. Barry, J.P., Vouros, P., Schepdael, A.V., and Law, S.-J. 1995. Mass and sequence verification of modified oligonucleotides using electrospray tandem mass spectrometry. J. Mass Spectrom. 30:993-1006. Barry, J.P., Muth, J., Law, S.-J., Karger, B.L., and Vouros, P. 1996. Analysis of modified oligonucleotides by capillary electrophoresis in a polyvinylpyrrolidone matrix coupled with electrospray mass spectrometry. J. Chromatogr. A 732:159-166. Bleicher, K. and Bayer, E. 1994. Various factors influencing the signal intensity of oligonucleotides in electrospray mass spectrometry. Biol. Mass Spectrom. 23:320-322. Boschenok, J. and Sheil, M.M. 1996. Electrospray tandem mass spectrometry of nucleotides. Rapid Commun. Mass Spectrom. 10:144149. Bruenger, E., Kowalak, J.A., Kuchino, Y., McCloskey, J.A., Mizushima, H., Stetter, K.O., and Crain, P.F. 1993. 5S rRNA modification in the hyperthermophilic archaea Sulfolobus solfataricus and Pyrodictium occultum. FASEB J. 7:196-200. Chen, R., Cheng, X., Mitchell, D.W., Hofstadler, S.A., Wu, Q., Rockwood, A.L., Sherman, M.G., and Smith, R.D. 1995. Trapping, detection, and mass determination of coliphage T4 DNA ions of 10 8 Da by electrospray ionization Fourier transform ion cyclotron resonance mass spectrometry. Anal. Chem. 34:1159-1163. Cheng, X., Gale, D.C., Udseth, H.R., and Smith, R.D. 1995. Charge state reduction of oligonucleotide negative ions from electrospray ionization. Anal. Chem. 67:586-593. Cheng, X., Camp, D.C., II, Wu, Q., Bakhtiar, R., Springer, D.L., Morris, B.J., Bruce, J.E., Anderson, G.A., Edmonds, C.G., and Smith, R.D. 1996a. Molecular weight determination of plasmid DNA using electrospray ionization mass spectrometry. Nucl. Acids Res. 24:2183-2189. Cheng, X., Harms, A.C., Goudreau, P.N., Terwilliger, T.C., and Smith, R.D. 1996b. Direct measurement of oligonucleotide binding stoichiometry of gene V protein by mass spectrometry. Proc. Natl. Acad. Sci. U.S.A. 93:7022-7027. Cheng, X., Morin, P.E., Harms, A.C., Bruce, J.E., Ben-David, Y., and Smith, R.D. 1996c. Mass spectrometric characterization of sequence-specific complexes of DNA and transcription factor PU.1 DNA binding domain. Anal. Biochem. 239:35-40. Covey, T.R., Bonner, R.F., Shushan, B.I., and Henion, J. 1988. The determination of protein, oligonucleotide, and peptide molecular weights by ion-spray mass spectrometry. Rapid Commun. Mass Spectrom. 2:249-256. Crain, P.F. 1990. Preparation and enzymatic hydrolysis of DNA and RNA for mass spectrometry. Methods Enzymol. 193:782-790. Ding, J. and Vouros, P. 1997. Capillary electrochromatography and capillary electrochromatography—mass spectrometry for the analysis of DNA adduct mixtures. Anal. Chem. 69:379-384. Emmett, M.R. and Caprioli, R.M. 1994. Micro-electrospray mass spectrometry: Ultra-high sensitivity analysis of peptides and proteins. J. Am. Soc. Mass Spectrom. 5:605-613. Fenn, J.B., Mann, M., Meng, C.K., Wong, S.F., and Whitehouse, C.M. 1989. Electrospray ionization for mass spectrometry of large biomolecules. Science 246:64-71. Gale, D.C. and Smith, R.D. 1995. Characterization of noncovalent complexes formed between minor groove binding molecules and duplex DNA by electrospray ionization–mass spectrometry. J. Am. Soc. Mass Spectrom. 6:1154-1164. Analysis of Oligonucleotides by Electrospray Ionization Mass Spectrometry
Gao, Q., Cheng, X., Smith, R.D., Yang, C.F., and Goldberg, I.H. 1996. Binding specificity of post-activated neocarzinostatin chromophore drug-bulged DNA complex studied using electrospray ionization mass spectrometry. J. Mass Spectrom. 31:31-36. Gaskell, S.J. 1997. Electrospray: Principles and practice. J. Mass Spectrom. 32:677-688.
10.2.18 Current Protocols in Nucleic Acid Chemistry
Glover, R.P., Sweetman, G.M.A., Farmer, P.B., and Roberts, G.C.K. 1995. Sequencing of oligonucleotides using high performance liquid chromatography and electrospray mass spectrometry. Rapid Commun. Mass Spectrom. 9:897-901. Goodlett, D.R., Camp, D.G., II, Hardin, C.C., Corregan, M., and Smith, R.D. 1993. Direct observation of a DNA quadruplex by electrospray ionization mass spectrometry. Biol. Mass Spectrom. 22:181-183. Greig, M. and Griffey, R.H. 1995. Utility of organic bases for improved electrospray mass spectrometry of oligonucleotides. Rapid Commun. Mass Spectrom. 9:97-102. Greig, M.J., Gaus, H., Cummins, L.L., Sasmor, H., and Griffey, R.H. 1995. Measurement of macromolecular binding using electrospray mass spectrometry. Determination of dissociation constants for oligonucleotide–serum albumin. J. Am. Chem. Soc. 117:831-832. Griffey, R.H., Sasmor, H., and Greig, M.J. 1997. Oligonucleotide charge states in negative ionization electrospray–mass spectrometry are a function of solution ammonium ion concentration. J. Am. Soc. Mass Spectrom. 8:155-160. Habibi-Goudarzi, S. and McLuckey, S.A. 1995. Ion trap collisional activation of the deprotonated deoxymononucleoside and deoxydinucleoside monophosphates. J. Am. Soc. Mass Spectrom. 6:102-113. Kowalak, J.A., Pomerantz, S.C., Crain, P.F., and McCloskey, J.A. 1993. A novel method for the determination of posttranscriptional modification in RNA by mass spectrometry. Nucl. Acids Res. 21:4577-4585. Kowalak, J.A., Dalluge, J.J., McCloskey, J.A., and Stetter, K.O. 1994. Role of posttranscriptional modification in stabilization of transfer RNA from hyperthermophiles. Biochemistry 33:7869-7876. Kowalak, J.A., Bruenger, E.B., and McCloskey, J.A. 1995. Posttranscriptional modification of the central loop of domain V in E. coli 23S ribosomal RNA. J. Biol. Chem. 270:17758-17764. Kowalak, J.A., Bruenger, E., Hashizume, T., Peltier, J.M., Ofengand, J., and McCloskey, J.A. 1996. Structural characterization of U*-1915 in domain IV from Escherichia coli 23S ribosomal RNA as 3-methylpseudouridine. Nucl. Acids Res. 24:688-693. Limbach, P.A. 1996. Indirect mass spectrometric methods for characterizing and sequencing oligonucleotides. Mass Spectrom. Rev. 15:297-336. Limbach, P.A., Crain, P.F., and McCloskey, J.A. 1994. Enzymatic sequencing of oligonucleotides with electrospray mass spectrometry. Nucl. Acids Res. Symp. Ser. 31:127-128. Limbach, P.A., Crain, P.F., and McCloskey, J.A. 1995. Molecular mass measurement of intact ribonucleic acids via electrospray ionization quadrupole mass spectrometry. J. Am. Soc. Mass Spectrom. 6:27-39. Little, D.P. and McLafferty, F.W. 1995. Sequencing 50-mer DNAs using electrospray tandem mass spectrometry and complementary fragmentation methods. J. Am. Chem. Soc. 117:6783-6784. Little, D.P., Chorush, R.A., Spier, J.P., Senko, M.W., Kelleher, N.L., and McLafferty, F.W. 1994a. Rapid sequencing of oligonucleotides by high-resolution mass spectrometry. J. Am. Chem. Soc. 116:4893-4897. Little, D.P., Speir, J.P., Senko, M.W., O’Conner, P.B., and McLafferty, F.W. 1994b. Infrared multiphoton dissociation of large multiply charged ions for biomolecule sequencing. Anal. Chem. 66:2809-2815. Little, D.P., Thannhauser, T.W., and McLafferty, F.W. 1995. Verification of 50- to 100-mer DNA and RNA sequences with high-resolution mass spectrometry. Proc. Natl. Acad. Sci. U.S.A. 92:2318-2322. Little, D.P., Aaserud, D.J., Valaskovic, G.A., and McLafferty, F.W. 1996. Sequence information from 42-108-mer DNAs (complete for a 50-mer) by tandem mass spectrometry. J. Am. Chem. Soc. 118:93529359. Liu, C., Muddiman, D.C., Tang, K., and Smith, R.D. 1997. Improving the microdialysis procedure for electrospray ionization mass spectrometry of biological samples. J. Mass Spectrom. 32:425-431. Liu, C., Wu, Q., Harms, A.C., and Smith, R.D. 1996. On-line microdialysis sample cleanup for electrospray ionization mass spectrometry of nucleic acid samples. Anal. Chem. 68:3295-3299. Loo, J.A. 1997. Studying non-covalent protein complexes by electrospray ionization mass spectrometry. Mass Spectrom. Rev. 16:1-23. McLuckey, S.A. and Habibi-Goudarzi, S. 1993. Decompositions of multiply charged oligonucleotide anions. J. Am. Chem. Soc. 115:12085-12095. McLuckey, S.A., Van Berkel, G.J., and Glish, G.L. 1992. Tandem mass spectrometry of small, multiply charged oligonucleotides. J. Am. Soc. Mass Spectrom. 3:60-70. McLuckey, S.A., Viadyanathan, G., and Habibi-Goudarzi, S. 1995. Charged vs. neutral nucleobase loss from multiply charged oligonucleotide anions. J. Mass Spectrom. 30:1222-1229. Muddiman, D.C., Cheng, X., Udseth, H.R., and Smith, R.D. 1996a. Charge state reduction with improved signal intensity of oligonucleotides in electrospray ionization mass spectrometry. J. Am. Soc. Mass Spectrom. 7:697-706.
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10.2.19 Current Protocols in Nucleic Acid Chemistry
Muddiman, D.C., Wunschel, D.S., Liu, C., Pasa-Tolic, L., Fox, K.F., Fox, A., Anderson, G.A., and Smith, R.D. 1996b. Characterization of PCR products from Bacilli using electrospray ionization FTICR mass spectrometry. Anal. Chem. 68:3705-3712. Muddiman, D.C., Anderson, G.A., Hofstadler, S.A., and Smith, R.D. 1997. Length and base composition of PCR-amplified nucleic acids using mass measurements from electrospray ionization mass spectrometry. Anal. Chem. 69:1543-1549. Naito, Y., Ishikawa, K., Koga, Y., Tsuneyoshi, T., Terunuma, H., and Arakawa, R. 1995. Molecular mass measurement of polymerase chain reaction products amplified from human blood DNA by electrospray ionization mass spectrometry. Rapid Commun. Mass Spectrom. 9:1484-1486. Ni, J., Pomerantz, S.C., and McCloskey, J.A. 1996a. Rapid sequencing of modified oligonucleotides using tandem mass spectrometry with electrospray ionization. Nucl. Acids Symp. Ser. 35:113-114. Ni, J., Pomerantz, S.C., Rozenski, J., Zhang, Y., and McCloskey, J.A. 1996b. Interpretation of oligonucleotide mass spectra for determination of sequence using electrospray ionization and tandem mass spectrometry. Anal. Chem. 68:1989-1999. Pomerantz, S.C. and McCloskey, J.A. 1990. Analysis of RNA hydrolysates by liquid chromatography–mass spectrometry. Methods Enzymol. 193:796-824. Pomerantz, S.C., Kowalak, J.A., and McCloskey, J.A. 1993. Determination of oligonucleotide composition from mass spectrometrically measured molecular weight. J. Am. Soc. Mass Spectrom. 4:204-209. Pomerantz, S.C., McCloskey, J.A., Tarasow, T.M., and Eaton, B.E. 1997. Deconvolution of combinatorial oligonucleotide libraries by electrospray ionization tandem mass spectrometry. J. Am. Chem. Soc. 119:3861-3867. Potier, N., Dorsselaer, A.V., Cordier, Y., Roch, O., and Bischoff, R. 1994. Negative electrospray ionization mass spectrometry of synthetic and chemically modified oligonucleotides. Nucl. Acids Res. 22:3895-3903. Przybylski, M. and Glocker, M.O. 1996. Electrospray mass spectrometry of biomacromolecular complexes with noncovalent interactions—new analytical perspectives for supramolecular chemistry and molecular recognition processes. Angew. Chem. Intl. Ed. Engl. 35:806-826. Sannes-Lowery, K.A., Mack, D.P., Hu, P., Mei, H.-Y., and Loo, J.A. 1997. Positive ion electrospray ionization mass spectrometry of oligonucleotides. J. Am. Soc. Mass Spectrom. 8:90-95. Stults, J.T. and Marsters, J.C. 1991. Improved electrospray ionization of synthetic oligodeoxynucleotides. Rapid Commun. Mass Spectrom. 5:359-363. Wilm, M. and Mann, M. 1996. Analytical properties of the nanoelectrospray ion source. Anal. Chem. 68:1-8. Wolter, M.A. and Engels, J.W. 1995. Nanoelectrospray mass spectrometry/mass spectrometry for the analysis of modified oligoribonucleotides. Eur. Mass Spectrom. 1:583-590. Wunschel, D.S., Fox, K.F., Fox, A., Bruce, J.E., Muddiman, D.C., and Smith, R.D. 1996. Analysis of double-stranded polymerase chain reaction products from the Bacillus cereus group by electrospray ionization Fourier transform ion cyclotron resonance mass spectrometry. Rapid Commun. Mass Spectrom. 10:29-35.
Contributed by Lenore M. Polo and Patrick A. Limbach Louisiana State University Baton Rouge, Louisiana
Analysis of Oligonucleotides by Electrospray Ionization Mass Spectrometry
10.2.20 Current Protocols in Nucleic Acid Chemistry
Overview of Purification and Analysis of Synthetic Nucleic Acids Chapter 10 presents information on evaluating, analyzing, and purifying nucleic acids, a broad class of biologically essential molecules, ranging from nucleobases, nucleosides, nucleotides, and oligonucleotides to native, highmolecular-weight, duplex DNA and RNA. Additionally, these methods will be useful for nucleic acid analogs, which possess many biological, diagnostic, and therapeutic effects. Emphasis is placed on the 5 to 10 million oligonucleotides synthesized each year currently for a large array of molecular biology applications. Steady improvements in the efficiency of synthesis chemistry and in the automated DNA synthesizers have made production of oligonucleotides routine. To produce an adequate level of quality and purity, rapid and convenient analytical methods are necessary for the dozens of oligonucleotides produced each day by a DNA synthesis laboratory. Basic principles are discussed to guide selection of the appropriate protocols to ensure functionality of oligonucleotides in the myriad of molecular biology applications. Synthetic oligonucleotides should be evaluated for concentration and purity before use, and purified if necessary. At the current levels of synthesizer efficiency and reagent quality, the crude oligonucleotide may be in an appropriate form and sufficient purity level to function as a sequencing or PCR primer—two relatively low-stringency applications. However, careful purification, analysis, quantitation, and other preparations may be necessary for oligonucleotides to perform in more stringent experiments such as mutagenesis, gene construction, and those requiring the cloning and expression of synthetic oligonucleotide sequences. Post-synthesis protocols for preparing ready-to-use oligonucleotides are provided in this chapter. Practical nucleic acid separations are based on four physical principles: velocity in an electric field (electrophoresis), sorption, size exclusion, and ultrafiltration. Useful methods of nucleic acid separation should identify samples accurately on the basis of molecular size (length) and possess near-single-nucleotide resolution. The analytical and purification methods detailed in this chapter are (1) polyacrylamide gel electrophoresis (PAGE; UNIT 10.4); (2) high performance liquid chroma-
UNIT 10.3
tography (HPLC; UNITS 10.5 & 10.6); (3) cartridge purification (UNIT 10.7); (4) mass spectroscopy (UNITS 10.1 & 10.2). Other useful methods, such as gel capillary electrophoresis, will be covered in future supplements.
SELECTING ANALYSIS/PURIFICATION METHODS Analysis and purification methods differ in complexity, expense, and time requirements. It is advisable to become familiar with all available options. Some factors to consider when selecting strategies for analysis and purification are (1) quantity needed for a particular experiment(s); (2) purity level required for the experiment; (3) time constraints; (4) equipment required; (5) sample composition (charge, hydrophobicity, solubility, sequence, length, salt form, labels, and modifications). Highly efficient syntheses yielding pure oligonucleotides lessen the demands on purification and facilitates analysis. Using the standard synthesis routine of automated, solid-phase, phosphoramidite chemistry (Beaucage and Iyer, 1992; Caruthers and Beaucage, 1983; Caruthers and Matteucci, 1984; UNIT 3.3; APPENDIX 3C), the oligonucleotide can be prepared with a final average yield per base addition of ∼98%. At this level of efficiency, a 20-nt oligonucleotide will result in ∼70% of the theoretical yield. Depending on the detection method and definition of purity, the crude sample will also be ∼70% pure by mass. The remaining ~30% is a heterogeneous population of failure sequences, shorter oligonucleotides that failed to couple during synthesis and were effectively capped to prevent further extension. Size (length) of the oligonucleotide is usually the discriminating factor which allows efficient separations. The sequence or base content of an oligonucleotide does not generally affect synthesis performance or the choices in analysis and purification methods. During synthesis, the four bases, as A,G,C,T phosphoramidite monomers, react at comparable rates and little compensation for their differences is necessary. Similarly, during analysis and purification, the chemical differences imparted by the bases are not usually significant. Exceptions are evident primarily under nondenaturing conditions, such as reversed-phase
Purification and Analysis of Synthetic Nucleic Acids and Components
Contributed by Alex Andrus and Robert G. Kuimelis
10.3.1
Current Protocols in Nucleic Acid Chemistry (2000) 10.3.1-10.3.6 Copyright © 2000 by John Wiley & Sons, Inc.
Supplement 1
HPLC. Occasionally, some sequences may show anomalous behavior, probably caused by hydrogen-bonding or secondary structure. For example, sequences containing four or more contiguous G bases are often problematic. Relative to other analytes, the narrow range of chromatographic and electrophoretic differences amongst oligonucleotides make this class of molecules relatively straightforward to analyze and purify.
POST-SYNTHESIS PROCEDURES FOR OLIGONUCLEOTIDES Post-synthesis processing, completed before analysis and purification, includes dimethoxytrityl (DMTr) assay, cleavage/deprotection, UV yield quantitation, desalting, and any other manipulations required to prepare the oligonucleotide for the experiment. Some of these steps may be omitted after success has been demonstrated in a particular application. For example, in some repetitive, well controlled sequencing or PCR experiments, the primers are simply dried from the ammonium hydroxide solution, after cleavage and deprotection, and dissolved in buffer, assuming an approximate concentration that contains the presumed amount of crude oligonucleotide. Caution is required when using oligonucleotides without analysis or purification. At a minimum, the DMTr fractions from synthesis and UV quantitation should be checked visually. Solutions of crude oligonucleotides should be assumed to contain no more than 70% of the correct sequence when making experimental calculations.
Synthesis Yield
Overview of Purification and Analysis of Synthetic Nucleic Acids
The flexibility of DNA/RNA synthesizers allows the choice of different synthesis scales. The smallest scale, ∼40 nmol, provides more than enough oligonucleotide for common applications such as sequencing and PCR primers. At the opposite end of the scale spectrum, the 10-µmol scale is used primarily for physical studies, antisense experiments, or for commercially produced probes and primers. At the 40-nmol scale, synthesis of a 20-nt crude oligonucleotide typically yields 5 to 10 OD units (165 to 330 µg), enough for hundreds of PCR and sequencing experiments (Table 10.3.1). An optical density (OD) unit is the absorbance of a 1-mL solution, typically in water, measured at 260 nm (A260), in a 1-cm path-length cuvette. The actual amount of pure oligonucleotide that can be attained after purification is dependent on the synthesis efficiency and can vary for
numerous reasons. In general, the small synthesis scales are more efficient and generate purer oligonucleotide products. The approximate yields (Table 10.3.1) are applicable for phosphorothioate or normal, phosphodiester sequences. The conversion [33 µg oligonucleotide per OD unit (A260nm)] between absorption (OD units) and mass depends on an averaged extinction coefficient (ε) per base of 10,000. Table 10.3.1 gives the approximate crude product yield at each scale to be expected when all instrument and reagent parameters are optimized.
DMTr Assay Determining the stepwise yield of coupling reactions by the DMTr (dimethoxytrityl, or trityl) assay during synthesis is a useful yet indirect indication of final product quality (Ellington, 1995; APPENDIX 3C). The DMTr group is the hydroxyl protecting group at the 5′ terminus of the oligonucleotide (UNIT 2.3). The DMTr assay is useful for immediate feedback on the performance of an automated DNA synthesizer as the DMTr cation is liberated at each detritylation step in the synthesis cycle and can be quantitated by absorbance measurement on a spectrophotometer. Most DNA synthesizers can deliver the DMTr cation–containing detritylation effluent from each cycle to a fraction collector, advanced by a signal from the synthesizer. The ISCO Cygnet fraction collector is convenient for one-column instruments, and the ISCO Retriever II, with a four-column adaptor, is suitable for multi-column instruments. Both can accommodate various test tube sizes. The DMTr fraction is diluted to 10.0 mL (5.0 mL for 40-nmol scale synthesis) using 0.1 M p-toluenesulfonic acid (Aldrich) in acetonitrile. The solution is mixed well and the absorbance read in a 1.0-cm path length cuvette at 490 nm, near the absorbance maxima for the bright orange dimethoxytrityl cation. The total yield is calculated by converting the final DMTr absorbance to a percent of the initial DMTr absorbance. The average stepwise yield is calcu-
Table 10.3.1 Approximate Yield of a Crude 20mer Oligonucleotide
Scale 40 nmol 0.2 µmol 1 µmol 10 µmol
Average overall yield >75% 50%-75% 50% 40%
OD unita
Amount
5-10 20-30 100 800
165-330 µg 660-1000 µg 3.3 mg 26 mg
a33 µg single-stranded DNA = 1 OD unit (A
260nm).
10.3.2 Supplement 1
Current Protocols in Nucleic Acid Chemistry
lated by raising the total yield to the power of the inverse number of trityl fractions (Andrus, 1992b). Overall yield = lowest / highest (OY = An/A2) Stepwise yield = Overall yield1/couplings(SY = OY1/n)
Some synthesizers feature an automated conductivity measurement of the DMTr cation, which calculates real-time average stepwise yields (Andrus, 1992a; Kaufman et al., 1993) while other instrument designs measure the absorbance of the detritylation effluent. An unusually low trityl absorption value corresponding to the detritylation of the first nucleoside on the column is sometimes noted. Typically, this first base absorption value is slightly less than the second due to spontaneous detritylation of the support-bound nucleoside during storage of the column. The DMTr cation cleaved in this manner is lost in the initial washes of the synthesis cycle and is not collected in the DMTr cation fraction. While this has no effect on the synthesis, it can affect the trityl assay. Therefore, the first trityl value is not used in calculations. A coupling failure with efficient capping would be detected in the trityl assay as a large drop in absorbance at one specific fraction, with subsequent fractions showing equally low absorbance. The oligonucleotides that failed to couple in the low yield coupling reaction were completely capped, thus eliminating their ability to react in subsequent coupling reactions. A failure of this type can be confirmed by the appearance of a major peak or band in HPLC, gel capillary electrophoresis (Andrus, 1992c, 1994), or PAGE analysis. A coupling failure that occurs with the first coupling, noted by a large decrease in absorbance of the second trityl fraction, could be indicative of inadequate reagent delivery (e.g., base, tetrazole, or trichloroacetic acid), most likely stemming from a failure to purge the phosphoramidite and tetrazole lines prior to synthesis. Unfortunately, the DMTr cation assay will not detect several conditions that may be detrimental to the final product quality. In some situations, the DMTr assay may give positive results while the product oligonucleotide is of poor quality, or there may be no product at all. For example, if oxidation is inefficient due to impaired flow, an empty bottle, or degraded reagent, then the internucleotide phosphite triester will be cleaved by the acidic detritylating reagent which is the next step in the synthesis cycle. The DMTr cation is liberated normally, giving a false indication of synthesis efficiency.
Stepwise yields from phosphoramidite synthesis on commercial synthesizers should be in the 98 ± 0.5% range. Yields below 97% may indicate synthesizer or reagent problems or some other unoptimized condition. Calculation and documentation of DMTr-assay yield data is very useful in the early diagnosis of instrument-related problems. Low DMTr assay average stepwise yields will inform the user of inferior product quality more quickly than analysis of the oligonucleotide. However, the DMTr assay should only be regarded as indirect evidence of purity and yield. Direct analysis of the oligonucleotide mixture is the only way to positively confirm product quality.
Cleavage/Deprotection After the repetitive addition of phosphoramidite monomers to complete the extension of the growing oligonucleotide, the ester linkage to the solid support is cleaved, usually with concentrated ammonium hydroxide (Ellington, 1995; APPENDIX 3C). Cleavage is automated on most synthesizers. Typically, ∼90% of the succinate ester linkages (UNIT 3.2) are cleaved and the protected oligonucleotide is liberated into solution within 30 min. To ensure complete cleavage, 1 hr at room temperature is required. Cleavage on a polystyrene support is slightly slower than on controlledpore-glass (CPG; McCollum and Andrus, 1991; UNITS 3.1 & 3.2). On the other hand, CPG dissolves slightly in ammonium hydroxide during the typical 1-hr cleavage, leaving a white residue of glass with the oligonucleotide upon concentration. During the cleavage period, removal of the other oligonucleotide protecting groups begins. Complete deprotection of an oligonucleotide continues in the same concentrated ammonium hydroxide cleavage solution and entails removing the protecting groups from the phosphate groups and the exocyclic amino groups on the bases. No extra addition of ammonium hydroxide is required. Deprotection requires ∼8 hr at 55°C when the standard set of nucleobase protection is used (Abz, Gibu, Cbz, T). The time period may be halved with each 10°C increase in temperature. For example, deprotection may alternatively be carried out for 4 hr at 65°C. Incomplete deprotection of synthetic oligonucleotides will impede analysis and purification, as well as adversely affect biological activity. Alternative reagents and conditions for cleavage and deprotection are sometimes employed (Bellenson and Smith, 1992; Reynolds and Buck, 1992; Boal et al., 1996). Since the
Purification and Analysis of Synthetic Nucleic Acids and Components
10.3.3 Current Protocols in Nucleic Acid Chemistry
Supplement 1
isobutyryl group on G bases is by far the slowest protecting group on the oligonucleotide to be removed, substitution of it with dimethylformamide (DMF) and use of the proven benzoyl protecting group on A and C has the benefits of fast deprotection, high yield, and high purity (Theisen et al., 1993). It is important that only fresh ammonium hydroxide be used. Concentrated ammonium hydroxide should be purchased in small bottles (500 mL or less) and stored at 4°C. Each bottle should be dated when opened and kept refrigerated. Bottles that have been open for more than one month or that have warmed to room temperature should not be used for deprotection. Laboratory personnel should be warned that keeping concentrated ammonium hydroxide sealed at high temperatures causes very high pressure of a hazardous reagent. Eye protection, gloves, and shields should be used when handling samples.
Quantitation
Overview of Purification and Analysis of Synthetic Nucleic Acids
Quantitation of oligonucleotides by UV absorbance is the most common and practical method to determine synthesis yield. The theoretical yield of any particular oligonucleotide can be calculated based on length and synthesis scale. The measurement of absorbance of light of a sample of DNA can be accurately used to convert to mass and thus molar amounts. The method is nondestructive and the sample is easily recovered. According to Beer’s Law, A = εCl (where A is the absorbance; ε is the molar extinction coefficient; C is the concentration in mol/L; and l is the path length in cm; typically, a 1-cm path-length cuvette is used). The conditions are defined at a certain wavelength and temperature and with a certain medium, all of which influence ε. The aromatic bases of DNA and RNA strongly absorb light with maxima near 260 nm. An average molar extinction coefficient of ε = 10,000 for each of the 4 bases is a useful approximation in a neutral pH range from 7.0 to 7.4. Using this, and other approximations, the absorbance can be translated to mass and concentration of oligonucleotides. One OD unit represents ∼33 µg of singlestranded oligodeoxynucleotide (DNA). For example, 1 mg of an oligonucleotide is ∼30 OD units. Conversely, for concentration purposes, 1 µmol of oligonucleotide will absorb 10 times the number of bases, in OD units. For example, 0.2 µmol of an 18-mer would be ∼36 OD units. There is a strong correlation between the measured yield (crude OD units), yield as a percentage of the theoretical yield (scale of synthesis), and oligonucleotide purity (qual-
Figure 10.3.1 Absorbance spectra of four natural deoxynucleosides, dA, dC, dG, and T in aqueous solution at pH of 7.0.
ity). For example, an 18-nt oligonucleotide made at the 0.2 µmol scale, which gives 20 crude OD units is probably significantly more pure than one which yields 10 crude OD units. A low overall yield may indicate poor coupling reactions during synthesis, incomplete cleavage from the support, or inaccurate calculation from the absorbance reading. The absorbance spectra of each of the four nucleosides are shown in Figure 10.3.1. A typical spectrum of a crude 18-nt oligonucleotide is shown in Figure 10.3.2. The average absorbance maximum of the four nucleosides is ∼260 nm. Oligonucleotides that are very rich in either purines or pyrimidines could have absorbance maxima above or below 260 nm, depending upon the base composition.
Figure 10.3.2 Absorbance spectra of an 18nt oligonucleotide 5′ TCA CAG TCT GAT CTC GAT 3′ in 0.1 M TEAA, pH 7.0.
10.3.4 Supplement 1
Current Protocols in Nucleic Acid Chemistry
Desalting The crude mixture, dissolved in ammonium hydroxide from the final deprotection, contains a variety of failure oligonucleotides along with some extraneous ammonium salts from the removed protecting groups. The crude mixture includes (1) the desired full-length oligonucleotide product; (2) shorter, failure-sequence oligonucleotides resulting from incomplete coupling and subsequent capping; (3) a collection of byproduct oligonucleotides resulting from other low-level side reactions, which were cleaved during ammoniolysis; (4) small molecule byproducts of synthesis including: benzamide, isobutyramide, ammonium acetate, ammonium trichloroacetate, pyridine, acetonitrile, and other solvents/reagents; and (5) trace amounts of other impurities. Although desalting the crude oligonucleotide to remove these contaminants may be unnecessary for some applications, one may wish to consider desalting the crude sample before analysis and purification. At the very least, this provides for more accurate quantitation and subsequent concentration calculations. It also provides the opportunity to exchange the ammonium counterions for others, such as sodium, which may be better enzymatic substrates or confer different solubility properties. There are several methods of desalting oligonucleotides, including precipitation, size-exclusion gel media (UNIT 10.7), and affinity cartridges (UNIT 10.7). The preferred method depends on the quantity of oligonucleotide to be desalted, the materials, and the time available. Precipitation of oligonucleotides in an aqueous alcohol medium is a quick, efficient, and inexpensive desalting method, with the capacity to desalt large quantities. Only one precipitation is needed to remove virtually all soluble salts, byproducts, and short failure-sequence oligonucleotides. A convenient and efficient protocol for precipitation of oligonucleotides is as follows: 1. Dissolve the oligonucleotide in 30 µL water (20 µL at large scale) and 5 µL of 3 M sodium acetate per OD unit of oligonucleotide. 2. Add 100 µL of ethanol per OD unit of oligonucleotide and mix by vortexing. For very short oligonucleotides (<15mers), isopropanol may be substituted for ethanol to ensure complete precipitation. 3. Store at 4°C or –20°C for ∼30 min, then centrifuge at high speed, i.e., ∼10,000 × g, for 5 min. 4. Remove the supernatant with a pipet or micropipet, or decant, being careful not to dis-
turb the pellet. Small quantities (<100 µg) may not be visible. 5. Add another 100 µL of ethanol, mix briefly, and centrifuge for 1 to 5 min. 6. Remove the supernatant and discard, being careful not to disturb the pellet. The oligonucleotide pellet can be dried by vacuum centrifugation, or air dried. 7. Resuspend the detritylated, desalted oligonucleotide in aqueous medium and quantitate by A260nm.
MIXED-BASE OLIGONUCLEOTIDES Oligonucleotides that have degenerate, or mixed-base sites, produce a more complex crude product mixture. The heterogeneity of mixed-base-containing oligonucleotides can present ambiguities and problems in analysis and purification. Equivalent incorporation of each base at a mixed-base site where more than one base has been delivered is usually desired. The four phosphoramidites, being different molecules, couple at slightly different rates. The order of reactivity, T>C>G>A, is largely compensated for by the order of the phosphoramidite positions on the valve block of some synthesizers, where A is positioned closest to the column and T the furthest. Virtually equal incorporation of the four bases in the oligonucleotide can result by concomitant delivery of the four equimolar and equivolume monomers. More precise and controlled incorporation of bases will result by premixing the desired phosphoramidites in a single bottle. Since mixed-base oligonucleotides are of the same length and contain the same number of phosphate anions, methods that separate on the basis of charge (electrophoresis and anion-exchange HPLC) will be more predictable than those that separate by hydrophobicity difference (reversed-phase HPLC).
MODIFIED OLIGONUCLEOTIDES Although standard oligonucleotides make up the vast majority of chemically synthesized nucleic acids, modified oligonucleotides (containing one or many modifications) are becoming increasingly important (see Chapter 4). Modifications can include fluorescent moieties and other types of labels attached to the 3′ or 5′ terminus, or attached at internal positions. Other common modifications include amino or thiol linkers, which are used for subsequent attachment schemes. Nucleobase, carbohydrate, and phosphodiester modifications are commonly used to alter the physical or enzyme
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substrate properties of an oligonucleotide. Many of these modifications also alter the chromatographic or electrophoretic behavior of the oligonucleotide and therefore influence the ultimate choice of a purification or analytical scheme. For example, fluorescent labels (and their associated linkers) usually impart substantial hydrophobicity to an oligonucleotide, and accordingly, affinity or reversed-phase separations would be employed. Certain special applications of oligonucleotides, especially modified oligonucleotides, may require very rigorous purification. In these situations it can be helpful to utilize an orthogonal purification scheme, whereby multiple purifications are performed in series on the same sample, but different modes of separation are employed at each step. For example, one might first use anion-exchange HPLC (charge-based separation) followed by reversed-phase HPLC (hydrophobicity-based separation). Performing the reversed-phase HPLC after ion exchange provides the added benefit of desalting the oligonucleotide. Each separation technique removes different types of impurities. Although time-consuming, orthogonal purification can be very effective when necessary.
LITERATURE CITED Andrus, A. 1992a. AutoAnalysis: Trityl monitoring of DNA synthesizer operation by conductivity. Research News, Applied Biosystems, Foster City, Calif. Andrus, A. 1992b. Evaluating and Isolating Synthetic Oligodeoxynucleotides. Applied Biosystems, Foster City, California. Andrus, A. 1992c. Oligodeoxynucleotide analysis by gel capillary electrophoresis. In Methods: A Companion to Methods in Enzymology, Vol. 4 (J. Wiktorowicz, ed.) pp. 213-226. Academic Press, San Diego. Andrus, A. 1994. Gel-capillary electrophoresis analysis of oligonucleotides. In Protocols for Oligonucleotide Conjugates (S. Agrawal, ed.) pp. 277-300. Humana Press, Totowa, N.J.
Bellenson, J. and Smith, A.J. 1992. Increasing DNA synthesizer throughput via off-instrument cleavage and deprotection. BioTechniques 12:219220. Boal, J.H., Wilk, A., Harindranath, N., Max, E., Kempe, T., and Beaucage, S.L. 1996. Cleavage of oligodeoxyribonucleotides from controlledpore glass supports and their rapid deprotection by gaseous amines. Nucleic. Acids Res. 24:311517. Caruthers, M.H. and Beaucage, S.L. 1983. Phosphoramidite compounds and processes. United States Patent 4,415,732. Caruthers, M.H. and Matteucci, M.D. 1984. Process for preparing polynucleotides. United States Patent 4,458,066 Ellington, A. 1995. Synthesis and purification of oligonucleotides. In Current Protocols in Molecular Biology, Vol. 1, (F. Ausubel et al, eds.) Unit 2.11. John Wiley & Sons, New York. Kaufman, J., Le, M., Ross, G., Hing, P., Budiansky, M., Yu, E., Campbell, E., Yoshimura, V., Fitzpatrick, V., Nadimi, K., and Andrus, A. 1993. Trityl monitoring of automated DNA synthesizer operation by conductivity: A new method of realtime analysis. BioTechniques 14:834-839. McCollum, C. and Andrus, A. 1991. An optimized polystyrene support for rapid, efficient oligonucleotide synthesis. Tetrahedron Lett. 32:40694072. Reynolds, T.R. and Buck, G.A. 1992. Rapid deprotection of synthetic oligonucleotides. BioTechniques 12:518-521. Theisen, P., McCollum, C., and Andrus, A. 1993. N-6-Dialkylformamidine-2′-deoxyadenosine phosphoramidites in oligodeoxynucleotide synthesis. Rapid deprotection of oligodeoxynucleotides. Nucleosides Nucleotides 12:10331046.
Contributed by Alex Andrus PE Applied Biosystems Foster City, California Robert G. Kuimelis Phylos, Inc. Lexington, Massachusetts
Beaucage, S.L. and Iyer, R.P. 1992. Advances in the synthesis of oligonucleotides by the phosphoramidite approach. Tetrahedron 48:22232311.
Overview of Purification and Analysis of Synthetic Nucleic Acids
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Polyacrylamide Gel Electrophoresis (PAGE) of Synthetic Nucleic Acids
UNIT 10.4
This unit describes methods and protocols for denaturing polyacrylamide gel electrophoresis (PAGE) for the analysis and purification of synthetic oligonucleotides. The analytical information is nonquantitative and recorded usually by a photograph (UV shadow or staining), autoradiography by radiolabeling ( CPMB UNITS 3.10 & 6.4), or digital methods, e.g., densitometry scanning. The materials and equipment for PAGE are relatively inexpensive and available in most molecular biology laboratories. The most convenient and familiar format, the vertical slab gel apparatus, is commercially available in different dimensions and configurations. The cross-linked polyacrylamide gel matrix is held between two glass plates and exposed on the top and bottom to chambers containing buffered electrolyte solution through which an electric field of controlled voltage or current is applied. The major operations involved in conducting PAGE are (1) sample preparation, (2) electrophoresis, (3) visualization of the gel, and (4) isolation of the product (purification). The protocols for analysis (see Basic Protocol 1) differ from those used for purification (see Basic Protocol 2); therefore, they are discussed separately. One method of staining gels for purposes of visualization is described in Support Protocol 1; radiolabeling of oligonucleotides (prior to separation by PAGE) for visualization by autoradiography is described in Support Protocol 2. Oligonucleotides made by the automated phosphoramidite method (UNIT 3.3 & APPENDIX 3C) should be quantitated prior to PAGE by UV absorbance (A260) so as not to overload or underload the gel. Initial analysis by MALDI-TOF (UNIT 10.1) or electrospray mass spectrometry analysis (UNIT 10.2) may be advised in some laboratories to ensure the presence of the full-length oligonucleotide. CAUTION: Radioactive materials require special training and handling; all supernatants must be considered radioactive waste and disposed of appropriately. [γ-32P]ATP should be handled behind a β shield. CAUTION: Acrylamide and bisacrylamide monomers are hazardous. Solutions of acrylamide deteriorate quickly, especially when exposed to light or left at room temperature. ANALYSIS OF SYNTHETIC NUCLEIC ACIDS BY PAGE The steps in PAGE analysis of oligonucleotides include (1) preparation of the gel and setup of the gel apparatus, (2) electrophoretic separation, and (3) detection of the oligonucleotides on the gel. There are two alternatives for detection--staining with methylene blue or SYBR Green, and radiolabeling analysis. For radiolabeling analysis, the oligonucleotides are first labeled at the 5′ end using [γ-32P]ATP and T4 polynucleotide kinase. Following electrophoresis, the radiolabeled oligonucleotides are detected by exposing the gel to X-ray film (autoradiography) or by use of a PhosphorImager (Molecular Dynamics; not described here). Materials 38% (w/v) acrylamide/2% (w/v) bisacrylamide (see recipe) 1× formamide/dye mix (see recipe for 10×; dilute in TBE buffer) Loading buffer: 9:1 formamide/1× TBE buffer
Contributed by Alex Andrus and Robert G. Kuimelis Current Protocols in Nucleic Acid Chemistry (2000) 10.4.1-10.4.10 Copyright © 2000 by John Wiley & Sons, Inc.
BASIC PROTOCOL 1
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Vertical electrophoresis system, such as Hoefer Model SE 400 or SE 600 in the 16 × 18–cm format with power supply (Pharmacia Biotech) Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (APPENDIX 3B) Set up gel 1. Assemble the gel plates, spacers, and combs as described in APPENDIX 3B, or following manufacturer’s instructions. 2. Determine the optimum acrylamide concentration for the range of oligonucleotide lengths as shown in Table 10.4.1. Prepare the desired acrylamide gel solution as described in APPENDIX 3B in a side-arm Erlenmeyer flask, adding everything but the TEMED. 3. Stopper the flask and apply a weak vacuum (using a water aspirator) through the side arm for ∼5 min to degas and remove oxygen. 4. Vent the system and add 50 µL TEMED while mixing. After 30 sec, pour the gel(s) as described in APPENDIX 3B. A 10- to 25-mL pipet with a rubber bulb is a convenient dispensing device.
5. Insert the comb. Dislodge any trapped air bubbles, especially in the wells, by tapping gently on the glass plates. Perform electrophoresis 6. After gel has polymerized (usually for ≥1 hr), remove bottom spacer of gel sandwich. Remove extraneous polyacrylamide from around combs. Clean spilled urea and acrylamide solution from outer plate surfaces with water. If gels are to be stored for several days before use, remove the comb upon polymerization, fill the wells with water, and place a piece of Parafilm or UV-transparent plastic wrap over the top of the gel sandwich to prevent drying. For further detail regarding gel preparation and handling see APPENDIX 3B.
7. Place the gel on the electrophoresis apparatus. Add enough 1× TBE buffer to the upper and lower chambers, to submerge the gel and wells by ∼3 cm. 8. Flush the wells with 1× TBE buffer using a Pasteur pipet to remove debris and urea before the sample is applied. 9. Set the power supply to a constant power of 20 to 40 W (for a 16 × 18–cm format). For other formats, set the power to draw 20 to 40 V/cm.The ideal wattage of the gel should generate enough heat so that the gel plates are warm, but not too hot to touch (~50°C).
10. Prerun with a blank sample of 5 to 10 µL of 1× formamide/dye mix in any one or more of the lanes for ∼30 min to ensure the lanes are running straight and to predict the time required for appropriate sample migration.
Table 10.4.1 Polyacrylamide Gel Electrophoresis (PAGE) of Synthetic Nucleic Acids
Selecting the Acrylamide Concentration
Oligonucleotide size (nt) <25 25-50 50-90 >90
Polyacrylamide concentration (%) 20 15 12 8
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Prerunning also increases the temperature of the gel, which helps to denature secondary structures, elutes impurities from the gel, and serves as visible confirmation that the gel is functioning properly.
11. Dissolve each sample oligonucleotide (0.5 to 2 OD units) in 10 µL of loading buffer by vortexing. Microcentrifuge the dissolved sample briefly to collect it at the bottom of the microcentrifuge tube. These conditions are appropriate where visualization is to be done by UV shadowing or staining analysis (using a 0.8-mm thick gel). For detection by autoradiography, load 2 to 20 pmol of crude radiolabeled oligonucleotide (5 to 10 µL of labeling reaction; Support Protocol 2) on a 0.4-mm thick, 40-cm long gel.
12. Flush the sample wells again with 1× TBE buffer and load the samples under the surface of the buffer and just above the surface of the well. A small, 10- to 25-mL syringe with a 25-G needle, or a flat-tipped micropipettor can be used to load the sample.
13. Start electrophoresis immediately using the conditions described in step 9. Electrophoresis should be started immediately after loading, to avoid band broadening by diffusion
14. Run until the position of the marker dye indicates that the DNA has been sufficiently resolved. For radiolabeled samples, dry the gel and perform autoradiography as described in APPENDIX 3B. To visualize unlabeled oligonucleotides by methylene blue staining, see Support Protocol 1. For detection by UV shadowing, see Basic Protocol 2, steps 5 to 11. Oligonucleotides longer than ∼80 nt often do not form well-shaped, detectable product bands, and should be analyzed by radiolabeling/PAGE or gel capillary electrophoresis. A photograph can be taken under UV light for a permanent record.
VISUALIZATION USING METHYLENE BLUE STAINING Direct visualization in the gel matrix can be done with a DNA-specific stain such as methylene blue or SYBR Green (Molecular Probes, 1996). Staining can be particularly useful if the gel is to be preserved by drying, but is not recommended for preparative purposes when the analyzed sample is to be used in subsequent applications. As an analytical tool, staining has limitations because some sequences in low concentrations may not be visible. Nevertheless, staining is more sensitive than UV shadowing and is considerably easier than radiolabeling. The stained gel can be photographed under ambient light, as opposed to short-wavelength UV light.
SUPPORT PROTOCOL 1
Additional Materials (also see Basic Protocol 1) 0.02% (w/v) methylene blue (Aldrich) in H2O 1. Electrophorese ∼1 OD unit of oligonucleotide per lane until the product has migrated about two-thirds down the gel (use bromphenol blue and xylene cyanol FF as marker dyes). 2. Disassemble the gel sandwich and carefully place the gel in a shallow pan containing enough 0.02% methylene blue to cover the gel. 3. Let stand for 15 to 30 min, then decant the stain and gently rinse the gel in water. If the background staining is too high, some destaining can be effected by soaking the gel in water.
4. Photograph the gel under ambient light against a white background.
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SUPPORT PROTOCOL 2
RADIOLABELING OF OLIGONUCLEOTIDES USING T4 POLYNUCLEOTIDE KINASE It is not necessary to prepurify or desalt sample oligonucleotides for radiolabeling analysis, as they may be used after drying down directly from the ammonia deprotection solution. The amount of each sample to be labeled can vary according to the activity of the [32P]ATP and the desired autoradiography film development time. The following protocol provides for loading samples in duplicate and will result in an approximate film development time of ∼1 to 2 hr. Label the following quantities of oligonucleotides according to length for 5′ 32P-radiolabeling analysis: <40 nt: 5 pmol; 40 to 70 nt: 10 pmol; >70 nt: 20 to 50 pmol. Convert absorbance to molar quantities by the approximation of 1 µmol oligonucleotide = 10 OD units/nt. For example, 5 pmol of a 20-mer is 0.001 OD unit. Typically 2 to 20 pmol of crude radiolabeled oligonucleotide are loaded onto a 0.4-mm thick, 40-cm long gel for optimum resolution. It will be necessary to make a serial dilution of the sample in water to avoid pipetting submicroliter aliquots. Materials Radiolabeling master mix (see recipe) 10× formamide/dye mix (see recipe) Additional reagents and equipment for preparing and running a polyacrylamide gel (Basic Protocol 1) 1. Dry quantity of oligonucleotide to be labeled in a microcentrifuge tube. 2. Vortex the master mix and microcentifuge briefly to collect any drops from the sides of the tube. 3. Pipet 6 µL of the master mix into each sample tube. 4. Vortex, then microcentrifuge briefly. Incubate at 37°C for 45 to 60 min. 5. Chill on ice for ~2 min, then microcentrifuge briefly. Add 14 µL of 10× formamide/dye and mix. Vortex, then microcentrifuge briefly. 6. Load 5 to 10 µL of each sample into the flushed wells of a prerun gel (see Basic Protocol 1). The dye-containing samples are stable, may be stored at 4°C/-20°C for weeks, and are ready for loading on the gel. With a half-life of 14 days, 32P-labeled oligonucleotides should be used within a few weeks.
BASIC PROTOCOL 2
Polyacrylamide Gel Electrophoresis (PAGE) of Synthetic Nucleic Acids
PURIFICATION OF SYNTHETIC NUCLEIC ACIDS USING PAGE AND UV SHADOWING The key steps in the PAGE purification process are (1) electrophoretic separation, (2) UV-shadow visualization, (3) excision of the product band from the gel, and (4) recovery of the pure product from the gel matrix (Efcavitch, 1990; Andrus, 1992; Ellington and Pollard, 1998). Crude samples of long oligonucleotides (>50 nt) contain substantially more truncated sequences due to the increased number of coupling cycles. A crude long
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oligonucleotide is in a lower-purity state than a shorter one, therefore, with a long oligonucleotide, more sample must be loaded on a preparative gel to visualize and isolate a useful amount. There is a limit to how much oligonucleotide can be loaded on a gel before resolution is lost and distortion prevails. Desalting prior to purification is recommended, e.g., by precipitation, for oligonucleotides >50 nt. Materials (also see Basic Protocol 1 and APPENDIX 3B) 20 × 20–cm fluorescent TLC plate, wrapped with UV-transparent plastic wrap Hand-held, short-wavelength UV light (240 to 300 nm) 1. Prepare a denaturing polyacrylamide gel as described in Basic Protocol 1 and APPENDIX 3B. The highest percentage gel for a given length is recommended to obtain maximum resolution (Table 10.4.1). Typically, 1.5- to 3.0-mm thick preparative gels are used. Longer oligonucleotides require longer migration distances, i.e., longer gels, to obtain sufficient resolution. The 16-cm height gel for oligonucleotides up to 50 nt and 40-cm gel height for oligonucleotides >50 nt are useful. The width of the comb teeth should be 2 to 3 cm and the maximum sample loading is ∼10 to 20 OD units per lane for the aforementioned dimensions.
2. Quantitate and prepare the DNA samples for purification as for analysis (see Basic Protocol 1). Dissolve the dried oligonucleotide sample in loading buffer to a concentration of ∼1 to 2 OD units/µL. Prepurification desalting is recommended to maximize resolution, e.g., precipitation, for >50 nt. Overloading a gel can lead to considerable distortion. Samples that do not readily dissolve in the loading buffer should be heated to ∼60°C and mixed. It is recommended that tracking dyes not be loaded in the same lane as the sample to avoid possible contamination and masking of the sample during UV shadowing.
3. Load samples and tracking dyes on the gel. Load tracking dyes in the outermost wells of the gel as a gauge to monitor the migration of the oligonucleotide. The crude long oligonucleotide is in a lower-purity state than a shorter one; it is necessary to load a larger sample of a long oligonucleotide on a preparative gel so that the product can be visualized and isolated in a useful amount.
4. Run the gel as described in Basic Protocol 1, steps 9 to 13. The gel conditions and concentrations are essentially the same for purification as they are for analysis, although the thicker gels require a higher power setting. The product oligonucleotide to be isolated should migrate as far down the gel as possible to achieve optimum product separation. This is unlike analysis, where more of the faster-running failure sequences need to be observed. Because the purpose is to separate the oligonucleotide from the crude mixture, it is acceptable to run the failure sequences off the bottom of the gel. Ideally the product band should be allowed to migrate two-thirds or more of the length of the gel. As a guideline, refer to the migration of dyes and oligonucleotides versus acrylamide concentration shown in Figure 10.4.1.
5. Remove the gel from the glass plates and place on top of a fluorescent TLC plate covered with UV-transparent plastic wrap. 6. Visualize the oligonucleotide under a short-wavelength UV light by holding the lamp directly overhead. The UV-absorbing oligonucleotide masks the fluorescent emission of the plate and appears as a shadow. Minimize the exposure of the oligonucleotide to UV light, which can cause degradation.
7. Slice the gel on the perimeter of the product band with a clean razor blade.
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Figure 10.4.1
Migration of dyes and oligonucleotides versus acrylamide concentration.
If the oligonucleotide does not have mixed-base sites, the cuts should be slightly to the interior of the product band to eliminate N-1 contamination. To avoid missing some of the possible sequences in a degenerate product mixture, the cuts should be be at or slightly outside of the diffuse product band.
8. Transfer the excised gel fragment(s) with tweezers to a small vessel (e.g., 15-mL conical, disposable tube) and crush the fragments with a spatula or rod. If more than one sample is run on the same gel, use caution when handling, to prevent cross-contamination.
9. Add just enough extraction solution (water or an aqueous, neutral buffer, such as 50 mM triethylammonium acetate) to cover the gel fragment. Vortex briefly and let stand for 8 to 24 hr at room temperature. Alternatively, the oligonucleotide can be isolated from the gel fragment by electroelution (UNIT 5.4). 10. Withdraw the oligonucleotide-containing solution and desalt by precipitation, sizeexclusion filtration, or affinity cartridge protocols (UNIT 10.6). 11. Quantitate by UV absorbance. Typical recovery yields vary somewhat, but are usually ∼50%.
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REAGENTS AND SOLUTIONS Use distilled, deionized water or other ultrapure water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Acrylamide/bisacrylamide, 38% (w/v)/2% (w/v) Dissolve 380 g acrylamide and 20 g bisacrylamide in deionized water to a volume of 1 L. Filter through a 0.5 µM membrane and store at 4°C, protected from light. Formamide/dye mix, 10× Prepare in deionized formamide: 0.1% (w/v) bromphenol blue 0.1 % (w/v) xylene cyanol FF Store up to 3 months at –20°C Kinase buffer, 10× 0.25 M Tris⋅Cl, pH 7.6 (APPENDIX 2A) 0.1 M MgCl2 0.1 M DTT Store at –20°C; discard after 6 months Radiolabeling master mix For each sample, prepare a master mix consisting of the following: 4 µL of 100 µM cold ATP in water (400 pmol) 1 µL of kinase buffer (see recipe) 0.2 µL 10 mM spermidine 1 µL (10 U) T4 polynucleotide kinase (New England Biolabs) 1.5 µL 5 mCi/ml [γ-32P]ATP [(3000 Ci; 111 TBq]/mmol; NEN Life Science Products)] COMMENTARY Background Information Charged molecules, such as oligonucleotides, migrate through the gel matrix under the influence of the applied electric field. The exact equation for electrophoretic velocity is complex, but the overwhelming factors are charge and mass. The charge-to-mass ratio for oligonucleotides remains virtually constant, independently of length. Molecular shape, net hydrophobicity, size, interactions with the gel matrix, and other parameters allow separation due to the net difference in velocity of sieving through the gel matrix. The elution order of an inverse relationship between length and velocity is established at ∼10 nt in length. For very short oligonucleotides, from 2 to ∼10 nt, the charge-to-mass ratio changes rapidly and the elution order is reversed. For example, a dimer migrates very slowly and a trimer slightly faster under denaturing gel conditions (7 M urea). Synthetic oligonucleotides of common lengths between 15 and 50 nt generally separate into regular, predictable band patterns in a lane on the polyacrylamide slab gel. The length, and, to a lesser degree, the sequence of an oligonucleotide affect the mobil-
ity of DNA through the gel. The order of migration contributed by the bases is C>A>T>G, with C being the fastest (Maniatis et al., 1975; Efcavitch, 1990). Oligonucleotides of the same length but with different sequence and base content will not migrate at exactly the same rate. Therefore, comparison of an "oligonucleotide size marker" with the sample oligonucleotide to verify the correct length gives only an approximate comparison. Denaturing conditions should be maintained with single-stranded DNA (oligonucleotides) to minimize the formation of duplexes and other secondary structures. Secondary structures can significantly alter the expected mobility of an oligonucleotide. Both the high concentration of urea (7 M) in the gel and formamide dissolving the sample serve as denaturants. Colored tracking dyes, such as bromphenol blue and xylene cyanol FF, can be loaded in the outside lanes to help determine the migration of the oligonucleotides in the inside lanes, which are not visible during electrophoresis. The relative migration rates of bromphenol blue and xylene cyanol FF to oligonucleotides changes as a function of acry-
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lamide concentration. Figure 10.4.1 shows the location of the dyes in a certain percentage of acrylamide relative to an oligonucleotide. The dyes determine the location of the oligonucleotides during electrophoresis and indicate the point at which the run should be terminated. For example, in a 15% gel, a 43-mer will run at the same rate as xylene cyanol FF. UV shadowing, the visualization technique most commonly used for purification, can also be used for analysis of crude mixtures. UV shadowing is less labor-intensive than radiolabeling and often provides sufficient resolution for routine analysis. Although a relatively higher oligonucleotide concentration is required for visualization by UV-shadowing (0.5 to 2 OD units), this is only a fraction of a typical synthesis. Also, it is possible to utilize UV shadowing to analyze a preparative/purification gel run. A photograph can be taken under UV light for a permanent record. Oligonucleotides longer than ∼80 nt often do not form well-shaped, detectable product bands, and should be analyzed by radiolabeling PAGE or gel capillary electrophoresis.
Critical Parameters and Troubleshooting
Polyacrylamide Gel Electrophoresis (PAGE) of Synthetic Nucleic Acids
Sometimes the oligonucleotide sequence can have a marked affect on the resolution of the gel. Usually the problems are due to secondary structures of palindromic, self-complementary sequences and sequences with contiguous G nucleosides. Such oligonucleotides can show multiple or diffuse bands, which are nonreproducible. PAGE analysis of mixed-base oligonucleotides, where certain base positions contain a mix of 2 or more bases, also can be complex. The product is a collection of sequences of equal length, and may resolve into multiple bands or appear as a broad band. When purifying mixed-base oligonucleotides, a wide excision should be made to ensure recovery of all represented sequences. Before the oligonucleotide undergoes electrophoresis, it should be quantitated by absorbance, desalted if necessary, and dried. The sample for PAGE is dissolved in ∼10 µL per 1.5-cm well for analytical-size gels (0.8 mm) of a mixture of formamide/1× TBE at a ratio of 9:1. This medium denatures, dissolves oligonucleotides rapidly, and is denser than 1× TBE, the electrophoresis buffer, so the sample falls through the buffer and settles on the bottom of the well and remains there until the power is turned on. Other denaturing loading media serve the same purpose. If an oligonucleotide is thought to form secondary structures, heating
to 90°C for several minutes prior to loading may help to give normal elution patterns and a single product band. Nucleic acid staining with dyes such as methylene blue or SYBR Green obviates the hazards and rapid decay of radioisotope labeling. Although ethidium bromide staining of DNA fragments is a well established technique for visualizing double-stranded DNA fragments, it is not useful for visualizing short, singlestranded DNA fragments such as oligonucleotides. The UV irradiation required to visualize ethidium bromide damages DNA and compromises the usefulness of this stain. Besides its toxicity hazards, the intercalation of ethidium bromide into short, single-stranded DNA is highly sequence-dependent and is sometimes undetectable in sequences less than 25 nt long. T4 polynucleotide kinase catalyzes the phosphorylation of an oligonucleotide 5′hydroxyl by ATP (Wallace and Miyada, 1987). Detection of radiolabeled oligonucleotides is far more sensitive than the UV shadowing or staining methods. Desalting of the sample oligonucleotide is not necessary for UV shadow analysis but may be necessary prior to 3′- or 5′-end enzymatic radiolabeling. After radiolabeling, desalting or purification is not necessary prior to electrophoresis. The excess [32P]ATP and other non-oligonucleotidic radioactive species will expose the film, but in most cases will not obscure the region of the labeled oligonucleotides of interest. If necessary, the radiolabeled oligonucleotides can be separated from the other components of the kinase reaction by gel-filtration, size-exclusion chromatography on Sephadex media (UNIT 10.7). The alternative radioisotope [33P]ATP is more stable (25 day half-life) and less hazardous (less penetrating); it is also available as [γ-33P]ATP and other nucleotides for labeling (NEN Life Science Products). Oligonucleotides with the DMTr group still on the 5′ end will not phosphorylate with kinase. Also, oligonucleotides with residual base-protecting groups remaining will appear as diffuse, slower-migrating species compared to deprotected samples, because the baseprotecting groups retard migration and add mass. Additionally, the base-protecting groups also hamper the ability of T4 kinase to end-label the product molecules. Excessive salt in the oligonucleotide sample may affect migration and may interfere with enzymatic labeling reactions. The presence of certain organic contaminants, such as acetic acid, can grossly distort electrophoretic mobility. Fresh kinase re-
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action buffer should be used, because this buffer contains DTT, which can degrade on storage. For analyzing crude compounds, there must be an excess of ATP in the reaction for competitive labeling of each oligonucleotide species in a sample. With a limiting amount of ATP (excess of oligonucleotide), the kinase enzyme will preferentially phosphorylate oligonucleotides of ∼10 to 20 nt before it will phosphorylate longer sequences. This effect will skew the analysis and result in false representation of the sample composition. The usual amount of [γ-32P]ATP is at a low concentration and is not sufficient by itself for efficient phosphorylation. The molar excess of ATP is provided by the addition of unlabeled (cold) ATP. Also, oligonucleotides can be 3′-radiolabeled by chain extension with radiolabeled nucleotides and terminal transferase (Sambrook et al., 1989). Other sample-related problems are due to inaccurate quantification of the sample, inactive enzyme, and inaccurate ATP concentrations. Oligonucleotide analogs, such as phosphorothioates, are not good substrates for kinase and will not radiolabel efficiently by phosphate transfer. Autoradiography is used to visualize the electrophoresis of radiolabeled oligonucleotides. The radioactive oligonucleotide molecules emit β particles which expose an X-ray film placed on top of the gel in the dark. To speed up the exposure process, an intensifying screen may be used. The film is developed in the dark with photographic developer and fixer and dried. Film exposure time is critical. An extended exposure will overrepresent failure sequences, since the product band will be saturated and will no longer respond to β emissions. An exposure that is too short will not sufficiently reveal failure sequences. Figure 10.4.2 is an autoradiogram of a collection of crude 32P-labeled oligonucleotides, 70 to 140 nt. The amount of full-length oligonucleotide relative to shorter, failure sequences can be seen to decrease with increase in length. Careful quantitation of samples is necessary to make comparisons, since the dynamic range of the film is low. Once a band fully exposes the film, it does not further darken while the lighter bands containing less radioisotope continue to expose and darken the film. A useful alternative to autoradiography is phosphor imaging. In this technique, a reusable phosphor storage screen is exposed to the gel instead of an X-ray film. The storage screen is then laser-scanned and the emitted light is detected, digitized, and quantitated. Phosphor imaging is faster, more
Figure 10.4.2 Autoradiogram of 32P-labeled ATP kinase gel, crude, 70- to 140-nt oligonucleotides.
sensitive, and provides a much broader dynamic range than autoradiography.
Anticipated Results Following the above protocols, it can be expected that for average-length (20 to 40 nt) and reasonably pure crude oligonucleotides, single-base separation will be observed by PAGE. For example, a 30-nt oligonucleotide will be completely resolved from the mixture of N-1 29-nt failure sequences and visible as distinct bands. Radiolabeling will give better analytical results and resolution, especially for longer oligonucleotides. While PAGE has less capacity than HPLC for purification purposes, up to 1 mg (30 OD units) of highly pure oligonucleotide can be isolated from a single gel.
Time Considerations The protocols here can be completed in 1 to 2 days. Typical electrophoresis runs require 1 to 6 hr. The post-electrophoresis purification protocol usually requires an overnight treatment of soaking the gel pieces followed by 1 to 2 hr for product isolation.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.4.9 Current Protocols in Nucleic Acid Chemistry
Supplement 1
Literature Cited Andrus, A. 1992. Evaluating and Isolating Synthetic Oligodeoxynucleotides. PE Biosystems Division of Perkin-Elmer, Foster City, Calif. Available upon request. Efcavitch, J.W. 1990. The electrophoresis of synthetic oligonucleotides. In Gel Electrophoresis of Nucleic Acids—A Practical Approach (D. Rickwood and B.D. Hames, eds.) pp. 125-149. Oxford University Press, Oxford. Ellington, A and Pollard, J.D., Jr. 1998. Purification of oligonucleotides using denaturing polyacrylamide gel electrophoresis. In Current Protocols in Molecular Biology, vol. 1 (F. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 2.12.12.12.7. John Wiley & Sons, New York. Maniatis, T., Jeffrey, A., and van deSande, H. 1975. Chain length determination of small double- and single-stranded DNA molecules by polyacrylamide gel electrophoresis. Biochemistry 14:3787-3794.
Molecular Probes, 1996. Handbook of Fluorescent Probes and Research Chemicals, 6th ed., pp. 164. Sambrook, J., Fritsch, E.F., and Maniatis, T. 1989. Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. Wallace, R.B. and Miyada, C.G. 1987. Oligonucleotide probes for the screening of recombinant DNA libraries. In Methods in Enzymology, Vol. 152, Guide to Molecular Cloning Techniques (S.L. Berger and A.R. Kimmel, eds.) pp. 432442. Academic Press, San Diego.
Contributed by Alex Andrus PE Applied Biosystems Foster City, California Robert G. Kuimelis Phylos, Inc. Lexington, Massachusetts
Polyacrylamide Gel Electrophoresis (PAGE) of Synthetic Nucleic Acids
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Current Protocols in Nucleic Acid Chemistry
Analysis and Purification of Synthetic Nucleic Acids Using HPLC
UNIT 10.5
This unit describes a collection of methods and protocols for high-performance liquid chromatography (HPLC) of synthetic nucleic acids (Pingoud et al., 1988; Newton, 1990; Zon, 1990; Warren and Vella, 1993; Oefner and Bonn, 1994; Andrus and Bloch, 1998). For 30 years, HPLC has been a powerful and popular method for the analysis and purification of small molecules, and, more recently, for biopolymers. A considerable industry exists to supply equipment, columns, and technical advice for the practice of HPLC. This unit will emphasize protocols for the analysis and purification of synthetic oligonucleotides. Reversed-phase (see Basic Protocol) and anion-exchange (see Alternate Protocol) are the reigning methods for HPLC of oligonucleotides. Typically anion-exchange gives better resolution and a more predictable elution pattern than reversed-phase. Reversed-phase has advantages in higher capacity purification and the use of volatile buffer systems for easier product recovery. A protocol for detritylating and precipitating oligonucleotides after HPLC purification is also presented (see Support Protocol). REVERSED-PHASE CHROMATOGRAPHY This protocol describes analysis and purification of oligonucleotides under reversedphase conditions. Oligonucleotides undergo hydrophobic interactions with the alkylsilylbonded groups on the reversed-phase adsorbent, resulting in elution patterns that correlate with their overall hydrophobicity. The popular reversed-phase method described in this protocol employs an acetonitrile gradient elution in dilute triethylammonium acetate (TEAA) at neutral pH on a porous hydrocarbon (C4 to C18) silica or polystyrene-divinylbenzene (PS-DVB) adsorbent. The hydrophobicity of triethylammonium, the oligonucleotide counterion, is a significant hydrophobic interactant in the reversed-phase method. Oligonucleotides can be chromatographed tritylated (with the 5′ DMTr group attached; DMTr ON) or detritylated (DMTr OFF).
BASIC PROTOCOL
Reversed-phase analysis and purification of tritylated and detritylated oligonucleotides are effective on a column such as Aquapore RP-300 with a moderately hydrophobic, octylsilyl (C8), bonded phase and a large 300 Å average pore size, or a more hydrophobic octadecylsilyl (C18), column, such as the Spheri-5 RP-18, 5 µm spherical silica (both available from PE Applied Biosystems). A standard-size column or cartridge (4.6 × 220 mm) is suitable for analysis of ∼0.3 to 1.0 OD units of crude mixtures. A maximum of ∼10 OD units can be applied to this size column for purifications. Smaller bore columns— 2.1-mm, or even microbore, 1.0-mm—offer some advantages in speed and reduced mobile-phase consumption. A larger column (7 × 250 mm or 10 × 250 mm) gives better separation with larger loadings. To purify the entire product from a 10-µmole scale synthesis requires many preparative injections. Materials Oligonucleotide sample Triethylamine 0.1 M triethylammonium acetate, pH 7 (TEAA; PE Applied Biosystems) for dissolving sample Mobile phase solution A: 0.1 M TEAA, pH 7.0 Mobile phase solution B: acetonitrile
Contributed by Alex Andrus and Robert G. Kuimelis Current Protocols in Nucleic Acid Chemistry (2000) 10.5.1-10.5.13 Copyright © 2000 by John Wiley & Sons, Inc.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.5.1 Supplement 1
HPLC instrumentation, capable of both analysis and purification up to about several milligrams oligonucleotide per injection with the following features and specifications: Injector: autosampler (preferred) or manual syringe Pumping system: ternary (preferred) or binary, 0.1-5 mL/min Detector: UV/fluorescence (preferred) or UV/VIS variable between 190 to 600 nm Data: integrating data system (preferred) or chart recorder Gradient system: displays and stores for redisplay and reformatting (preferred) or programmable Column: 4.6 × 220-mm Aquapore RP-300 (Applied Biosystems) or Spheri-5 RP-18 (Applied Biosystems) Fraction collector 1. Dry the quantity of oligonucleotide needed for analysis (0.05 to 0.1 OD unit) or purification (>0.1 OD unit) in a vessel (e.g., microcentrifuge tube) under vacuum. For tritylated oligonucleotides, add a drop of triethylamine periodically during the evaporation process as a preservative. To prevent inadvertent detritylation of tritylated oligonucleotides, avoid heat and acid during the drying step. Evaporation of ammonium hydroxide should be done at room temperature when the 5′ trityl is being preserved on the oligonucleotide.
2. Dissolve the sample immediately in sufficient 0.1 M triethylammonium acetate (TEAA) pH 7 for injection. Typical injection volumes for analysis are 10 to 50 mL, or 10 ml to 2 ml for purification limited by the sample loop size of the injector. Sample loops as large as 2 mL are available for purification injections.
3. Store tritylated oligonucleotides, if necessary, in the TEAA buffer, adding ∼10% (v/v) triethylamine as a preservative for tritylated oligonucleotides. It is difficult or impossible to keep the trityl intact on oligonucleotides upon prolonged storage under any conditions. The trityl group is also unstable if the tritylated oligonucleotide is stored on the solid support, prior to cleavage.
4. Program the gradient system to start with 100% mobile phase solution A (0.1 M TEAA), increasing the percentage of mobile phase solution B (acetonitrile) with time (Table 10.5.1). Ensure that sufficient mobile phase has been installed to keep intakes covered during run. The mobile phase most often used for tritylated and detritylated oligonucleotides is an increasing gradient of acetonitrile in 0.1 M TEAA, pH 7.0 (Table 10.5.1). Most crude
Table 10.5.1 Gradient and Mobile Phase for Reversed-Phase HPLCa
Elapsed time (min) 0 24 34 Analysis and Purification of Synthetic Nucleic Acids Using HPLC
% mobile phase B at elapsed time 8 20 40
aGradient conditions are based on a flow rate of 1 mL/min using a Spheri-5 RP-18, 5 µm diameter spherical silica, 4.6 × 200 mm, or Aquapore RP-300 C-8, 7 µm diameter, 300 Å pore, 4.6 × 220 mm, (all available from PE Applied Biosystems) column at ambient temperature with a 60 min injection cycle.
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reaction mixtures in the range of 15 to 40 nt can be analyzed and purified using the linear gradient system in Table 10.5.1. This simple gradient is useful for tritylated and detritylated oligonucleotide analysis and purifications.
5. Equilibrate the HPLC system with the starting mobile phase composition until a flat baseline is achieved at the desired detection wavelength. 6. Inject the sample. 7. Collect the desired fractions either with an automated fraction collector, or by observing the chromatogram in real-time and manually collecting the eluate. Tritylated oligonucleotides will elute at ∼25 to 35 min and detritylated oligonucleotides at ∼12 to 15 min under the recommended conditions (Table 10.5.1).
ANION-EXCHANGE HPLC This protocol describes analysis and purification of oligonucleotides under anion-exchange conditions. The anion-exchange method consists of a salt-gradient elution on a tertiary or quaternary ammonium-derivatized porous silica or polymeric adsorbent. Denaturing cosolvents such as formamide may be added to minimize hydrogen bonding.
ALTERNATE PROTOCOL
Additional Materials (also see Basic Protocol) Mobile phase solutions A and B for anion exchange (see recipe) Aquapore AX-300 (PE Applied Biosystems) or DNAPac Pa-100 (Dionex) column 1. Dry the quantity of oligonucleotide needed for analysis (0.05 to 0.1 OD units) or purification (≤1 O.D. units) in a vessel (e.g., microcentrifuge tube) under vacuum. Anion-exchange samples are typically analyzed and purified detritylated. Crude oligonucleotides in the ammonium hydroxide solution do not have to be dried when using polymeric adsorbent anion-exchange columns.
2. Dissolve the sample in sufficient mobile phase A (10 mM NaClO4) for injection. Typical injection volume for analysis is 10 to 50 mL, or 10 ml to 2 ml for purification, limited by the sample loop size of the injector. Sample loops as large as 2 mL are available for purification injections.
3. Program the gradient system to start with 100% mobile phase A (10 mM NaClO4), increasing the percentage of mobile phase B (300 mM NaClO 4) with time (Table 10.5.2). Ensure that sufficient mobile phase has been installed to keep intakes covered during run. 4. Equilibrate the HPLC system with the starting mobile phase composition until a flat baseline is achieved at the desired detection wavelength. 5. Inject the sample. 6. Collect the desired fractions either with an automated fraction collector, or by observing the chromatogram in real-time and manually collecting the eluate. Oligonucleotides will elute at ∼15 to 25 min under the recommended conditions (Table 10.5.2).
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Supplement 1
Table 10.5.2
Gradient for Anion-Exchange HPLCa
Elapsed time (min) 0 20 25 26
% mobile phase B at elapsed timeb 0 70 70 0
aGradient is based on a 1 mL/min flow rate at 25°C using an
Aquapore AX-300 (PE Applied Biosystems) or DNAPac PA-100 (Dionex) column with a 35 min injection cycle. bMobile phase solution A is 10 mM NaClO and mobile phase 4 solution B is 300 mM NaClO4. See recipe in Reagents and Solutions.
SUPPORT PROTOCOL
POST-HPLC DETRITYLATION AND PRODUCT ISOLATION After collection of the purified tritylated oligonucleotide, the trityl group must be removed. The evaporated sample, which could appear as an oil, is detritylated with acetic acid at room temperature. The acidic treatment does not cause depurination because nucleotides are much more stable to acid after deprotection. Direct precipitation removes acetic acid, the DMTr cation, and byproducts, and ultimately yields the purified oligonucleotide in sodium salt form, dried as a pellet. A final analytical separation will give confirmation of purity. Post-HPLC isolation is simple for collected fractions of detritylated oligonucleotides by reversed-phase purification because detritylation is not required (see step 2 of this protocol). The combined purified fractions are dried to remove water and the volatile salts (TEAA), and can be used directly in most applications. The purified product can also be ethanol precipitated with concomitant counterion exchange, e.g., sodium for triethylammonium, by the following protocol whereby each OD unit of oligonucleotide is dried and dissolved in 20 µL water and carried forward from step 3 (Andrus, 1992). Materials Purified oligonucleotide 80% acetic acid 3 M sodium acetate Absolute ethanol or 2-propanol 1. Dry the combined, purified oligonucleotide fractions (see Basic Protocol and Alternate Protocol) under vacuum. Incorporate all fractions in a small vessel, such as a 1.5-mL microcentrifuge tube. If the evaporated sample leaves a white film on the bottom on the vial, probably some residual TEAA remains in the sample. Add a small amount (∼100 µL) of distilled, deionized water and evaporate to remove these salts (several evaporations may be necessary). 2. Dissolve the tritylated oligonucleotide in 20 µL 80% acetic acid per OD unit, with vortexing, at room temperature. Let stand 20 min. Omit this step for detritylated samples. Phosphorothioate oligonucleotides may require brief heating at 65°C.
Analysis and Purification of Synthetic Nucleic Acids Using HPLC
3. Add 5 µL 3 M sodium acetate and 60 µL absolute ethanol per OD unit oligonucleotide and mix by vortexing. For very short oligonucleotides (<15-mers), substitute 2propanol for ethanol to ensure complete precipitation.
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4. Chill the precipitation at ≤4°C for ∼30 min. Microcentrifuge 5 min at ∼10,000 × g, at ambient temperature or colder. 5. Remove the supernatant with a pipet, micropipet, or by decanting, being careful not to disturb the pellet. Small quantities (<100 µg) may not be visible.
6. Add another 60 µL absolute ethanol per OD unit to the pellet, mix briefly, and microcentrifuge 1 to 5 min, ~10,000 × g, at ambient temperature or colder. 7. Discard the supernatant, being careful not to disturb the pellet. Dry the purified oligonucleotide pellet by vacuum centrifugation or by leaving open on the benchtop. 8. Resuspend the detritylated, desalted oligonucleotide in aqueous medium and quantitate by A260. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Mobile phase solutions A and B for anion exchange Mobile phase solution A (10 mM NaClO4, pH 8.5): Dissolve 1.4 g NaClO4.H2O (10 mM final; Fluka) in 1 L distilled or deionized water. Adjust pH to 8.5 with 0.1 N NaOH. Mobile phase solution B (300 mM NaClO4, pH 8.5): Dissolve 42.1 g NaClO4.H2O (300 mM final) in 1 L distilled or deionized water. Adjust pH to 8.5 with 0.1 N NaOH. COMMENTARY Background Information HPLC embodies the combination of analysis and purification. The analytical chromatogram gives information about (1) purity quantitation, (2) identity by comparison with known samples, and (3) synthesis efficiency. HPLC is a scaleable method, with the largest capacity of any method for oligonucleotide purification. Other aspects of HPLC are a high degree of automation, high resolution, high sensitivity, and ease of pure product recovery. Parameters can be tailored for each separation, as there are many commercial adsorbents (columns and cartridges) for evaluation and purification of nucleic acids. HPLC technology is evolving in the direction of smaller-diameter, nonporous, spherical adsorbent particles packed in smaller columns, with benefits in separation speed, resolution, throughput, and automation. Because of the resolution limitations of both reversed-phase and anion-exchange HPLC for longer oligonucleotides, HPLC is a useful system for the reliable analysis and purification of oligonucleotides up to ∼50 nt in length. Longer oligonucleotides are best analyzed by electro-
phoretic methods such as slab gel (PAGE, UNIT 10.4) or gel capillary electrophoresis. At the current level of synthesizer efficiency and reagent quality, the crude oligonucleotide may be sufficiently pure to function in some experiments. However, careful purification, analysis quantitation, and other preparations are necessary for longer oligonucleotides and for experiments where oligonucleotide purity is critical, especially those requiring cloning and expression. Furthermore, failure to document oligonucleotide purity (e.g., by HPLC) may cloud the interpretation of applications with ambiguous results, as well as the quality control aspects of a laboratory. Automated, solid-phase, phosphoramidite chemistry (Alul, 1993) gives oligonucleotides with a final average yield per base addition of ∼98%, lessening slightly at the larger scales (see Table 10.3.1). At this level of efficiency, synthesis of a 20 nt oligonucleotide will result in ∼70% of the theoretical yield. Depending on the detection method and definition of purity, the sample will also be ∼70% pure. The major side-products are shorter oligonucleotides that failed to couple during synthesis and were ef-
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10.5.5 Current Protocols in Nucleic Acid Chemistry
Supplement 4
fectively capped to prevent further extension. These failure products typically elute earlier in HPLC than the full length product. Initial analysis by MALDI-TOF (UNIT 10.1) or electrospray mass analysis (UNIT 10.2) may be advised in some labs to ensure the presence of the full-length oligonucleotide. Other side reactions during synthesis lead to small amounts of later-eluting, more hydrophobic, or highly charged impurities. The yield after HPLC purification is ∼50% of the crude yield (see Table 10.3.1). Reversed-phase (RP) columns separate by hydrophobic differences. The solid particles within the reversed-phase column are usually silica bonded with hydrocarbon chains via silanol linkages. The polar, aqueous mobile phase and the hydrophobic adsorbent are considered to be in the “reversed” phase, compared to traditional, normal-phase chromatography which uses nonpolar organic solvents and underivatized, polar silica gel. Anion-exchange columns (discussed below) separate oligonucleotides based on charge differences. The particles are typically inorganic silica, derivatized with charged groups such as alkyl ammonium (typically weak or strong anion exchangers). Polymeric particles are also commercially available and useful for reversed-phase and anion-exchange column adsorbents.
Analysis and Purification of Synthetic Nucleic Acids Using HPLC
Reversed-phase Interactions between the oligonucleotide sample (analyte) and the stationary adsorbent packed in the column determine the rate and order of elution. The composition of the liquid mobile phase is typically altered throughout the run (gradient), as opposed to being held constant (isocratic). The mobile phase is aqueous, with dissolved organic solvents or ionic salts, that compete with the sample for interactive adsorbent sites, or otherwise influence elution of the sample. Components of the mobile phase may also act as denaturants in minimizing hydrogen bonding. Stable intramolecular or intermolecular conformations, known as secondary structures, may exist for oligonucleotides with certain sequences. The large, hydrophobic dimethoxytrityl (DMTr) group may be left on or removed from the 5′ end of the oligonucleotide, according to the option selected by the DNA synthesizer user. Highly efficient synthesis (high coupling yield, minimum side reactions, complete capping and detritylation) gives crude oligonucleotides with only the desired, full-length product bearing a DMTr group. Theoretically,
the only oligonucleotide bearing the terminal 5′ DMTr group is the full-length product. All the other sequences that failed to couple in the crude reaction mixture will have been acetylated at the 5′ position during capping. Upon deprotection in ammonia, the failure sequences will have 5′-hydroxyl groups and be separable from the product by a large difference in hydrophobicity due to the presence of the terminal 5′ DMTr group on the full-length product. With the proper gradient and mobile phase conditions, reversed-phase HPLC will easily separate the two types of oligonucleotides. The tritylated product, being more hydrophobic, will be retained longer on the column than the shorter, nontritylated failure sequences and other components of the crude reaction mixture. The retention time differences between tritylated and nontritylated oligonucleotides in the crude oligonucleotide mixture are quite large, especially by the reversed-phase method. Since reversed-phase columns have much greater capacity for tritylated oligonucleotides, this is the preferred mode for purification. However, the trityl group is labile and it can be problematic to preserve it quantitatively on the oligonucleotide during sample preparation or storage. For this reason, and for greater resolution, HPLC of detritylated oligonucleotides is preferred for analysis. Polymeric adsorbents, such as highly cross-linked, rigid polystyrene beads (Ikuta et al., 1984; Germann et al., 1987; Huber et al., 1993), are useful alternatives to hydrocarbon-bonded silica in reversed-phase columns and can use the same gradients and mobile phases (Table 10.5.1). The polymeric columns (Polymer Laboratories, Hamilton, Sarasep, and Rapp Polymere) have the advantages of stability toward high temperature, pH extremes, and aggressive solvents. They typically exhibit longer lifetimes than their silica counterparts and show very high capacity for tritylated oligonucleotides. High pH of the mobile phase and heated columns, act as denaturants in the reversed-phase process. These denaturing conditions eliminate some of the secondary structure problems mentioned above. Anion-exchange Anion-exchange HPLC is another efficient technique for analysis and purification (Drager and Regnier, 1985; Maisano et al., 1989; Bergot and Egan, 1992; Huber et al., 1996). Separation is largely based on charge differences. Each oligonucleotide in the crude mixture has a different net charge based on the number of phosphate groups in the molecule (base length) and
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on the respective charges on the bases (Krstulovic, 1987). Therefore, base composition can affect the ion-exchange separation process. Resolution is excellent in the analysis and purification of small quantities of detritylated oligonucleotides. Single-base resolution can usually be achieved for oligonucleotides up to 25 nt in length and sometimes longer, depending on the sequence. The advantages of anion-exchange are ease of sample preparation, without the need to preserve the trityl group, and recovery of the purified compound in a ready-to-use form. The detritylated oligonucleotide sample can be redissolved in deionized water and stored in a refrigerator or freezer indefinitely. Anion-exchange polymeric adsorbents allow for high-pH mobile phase analysis, which neutralizes nucleobases and denatures secondary structures. Separation of the crude mixture is accomplished by slowly increasing the ionic strength (concentration) of the mobile phase (Table 10.5.2). The longer, more highly charged, oligonucleotides will thus elute later than shorter ones, with the product peak eluting last since it is the most highly charged. Phosphate groups are occasionally masked by secondary structures and will not exhibit the effect of a full charge, causing deviation from the predictable size-dependent elution pattern. Oligonucleotides have a hydrophilic, polyanionic backbone decorated with hydrophobic and hydrogen-bonding nucleobases, capable of complex interactions with the adsorbent. Stable intra or intermolecular conformations, secondary structures from Watson/Crick base-pairing or from Hoogsteen base-pairing in G-rich sequences can result in complex chromatograms and variable retention times.(Kang et al., 1992; Wang, 1993). The secondary-structure effects can be minimized by adding a strong denaturant, such as formamide, to the mobile phase, or by heating the column up to ∼70°C. Alternatively, in the case of anion exchange HPLC, alkaline mobile phases (pH 12) can be employed. Resolution decreases as larger oligonucleotides are eluted, because the relative difference in charge between longer oligonucleotides is less than in the shorter size ranges. Also, there is inevitable peak broadening as a function of increasing oligonucleotide length. The retention times of oligonucleotides on ionexchange HPLC, using identical mobile phase conditions, will usually be quite reproducible and predictable. Therefore, sequence length can be confirmed with reliability. Crude oligonucleotides may be injected (0.02 to 0.2 OD units per run) directly onto polymeric columns
from ammonium hydroxide solution (Ausserer and Biros, 1995). Labeled oligonucleotides, RNA, and analogs Covalent attachment of molecules, such as fluorescent dyes, biotin, proteins, and others, can be made at virtually any site on the oligonucleotide (Goodchild, 1990; Alul, 1993; Andrus, 1995). These include the 5′ and 3′ ends, through the bases, and through an internucleotide phosphate modification. HPLC of internucleotide analog DNA, such as phosphorothioate, methylphosphonate, phosphorodithioate, and phosphoramidate, has been investigated (Zon 1993; Andrus and Bloch, 1998). Analysis and purification by HPLC are feasible for most of these derivatized oligonucleotides, with only slight modifications to the methods. Many of these analogs are very hydrophobic and require higher organic modifier concentration in the mobile phase for reversedphase separations. Some analog DNAs have chiral phosphorus atoms, leading to complex diastereomeric mixtures. The retention time and elution pattern of conjugated oligonucleotides will be affected by the hydrophobicity of the attached species. For example, biotin, especially with a long linking chain, imparts significant additional hydrophobicity and increases the retention time. Synthetic oligoribonucleotides (RNA) are well-behaved HPLC analytes (Sproat et al., 1995). Extra care is warranted for analysis and purification since the synthesis efficiency is lower, reagent costs are higher, and most of the biological applications for RNA are more purity-critical than for DNA. The standard 2′-Otert-butyldimethylsilyl protecting group of oligoribonucleotides is usually removed with tetrabutylammonium fluoride in the final deprotection operation and requires careful separation of the copious tetrabutylammonium salts from the RNA. New reagents and post-synthesis methods have been developed to address this problem (Applied Biosystems, 1995). RNA is more likely to exhibit secondary-structure effects than DNA, especially those sequences designed with intramolecular hydrogen-bonding, e.g., ribozymes (Noller, 1984; Symons, 1992). Sometimes heating the column in an oven gives a better indication of purity due to denaturation of secondary structures and the resulting simplification of the chromatogram. RNA is more hydrophilic than DNA, which decreases the loading capacity on HPLC columns.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.5.7 Current Protocols in Nucleic Acid Chemistry
Supplement 1
Critical Parameters and Troubleshooting
Analysis and Purification of Synthetic Nucleic Acids Using HPLC
HPLC is meant to be an automated, largely unattended operation. However, the hardware is relatively complex and expensive, compared to PAGE (polyacrylamide gel electrophoresis, see UNIT 10.4). A minimum of down time and consistent, optimal results can be attained by observing the following proven HPLC maintenance operations: 1. Sample preparation: Filter or centrifuge to remove insoluble particulate matter. 2. Pump maintenance: Clear all salts and buffers from the pumps at the end of a session. Mobile phases should be maintained under a positive pressure of ∼5 psi of inert gas, e.g., helium, to prevent bubble formation and evaporation. 3. Data storage: Store chromatograms in personal computer–based systems. 4. Mobile phases: Filter and degas. 5. Guard columns: Mount in line between the injector and the column. A guard column can extend the column lifetime by trapping insoluble impurities and chemically damaging species in the sample. Manufacturers of HPLC equipment often provide useful literature and references for learning to use and maintain the system, as well as for specific applications. In configuring an HPLC system, it is imperative that extra column dead volumes be minimized, especially for analytical-scale and microbore HPLC. Most systems are configured primarily for analysis; therefore dead volumes and mixing points are often minimized only up to the flow cell of the detector. For preparative work, dead volumes and mixing points must be eliminated between the flow cell and point of collection. Nucleic acids—DNA and RNA—absorb light strongly at ∼260 nm due to their chromophoric, conjugated pyrimidine and purine ring systems. Detecting and quantitating the absorbance of the ultraviolet light can be done precisely. A finite period of time is required for a compound to pass from the detector to the collection port. This time interval can be determined by injecting any UV-absorbing compound, collecting the fractions at specific time points, and measuring the absorbance of the fractions individually. The time corresponding to the fraction containing the strongest UV absorption minus the time of the peak maximum on the chart recorder will be the time lag before samples should be collected. It is important that this time interval be kept under 10 to 20 sec for most accurate preparative work. It is also very help-
ful to have a digital readout of the absorbance on a real-time basis. This can be extremely important in the purification of complex mixtures where the points of collection play a key role in obtaining pure material. The HPLC purification process is different from the analysis process in the following ways. Typically a larger column and higher flow rates are employed for isolating milligram amounts of oligonucleotides. The resolution requirements for purification are not as strict as in analysis. Because the amount of product needed for most applications is quite small, only a portion of the peak need be collected. Since most molecular biology applications require only picomole quantities of pure oligonucleotide, it is better to be more conservative in the collection process and get less quantity of the pure product than to try for a larger amount of less pure product. Hence, good collection techniques and post-purification analysis can routinely produce pure product even when the analytical chromatogram shows a complex mixture.
Anticipated Results For conventional UV absorbance detection at or near 260 nm, ng to µg quantities of nucleic acid in 0.1 to 1 mL peak volumes give signals between the state-of-the-art detection limit (10−5 AU) and the 1 to 2 AUFS (absorbance units full scale) upper bound of detector linear dynamic range. For milligram-scale separations, detection may be set off the absorbance maximum for DNA at 280 to 300 nm to keep the plot on scale. Analytical-scale, 2.1 to 4.6 mm i.d. (non-microbore) HPLC columns show good resolution and recovery of ng to mg quantities of nucleic acid. Larger-scale separations for industrial manufacture of ribozyme or antisense therapeutics will require significant scale-up, a topic beyond the scope of this unit. A typical tritylated oligonucleotide chromatogram is shown in Figure 10.5.1. The initial peak at 5 min retention time is benzamide, the by-product from deprotection of the deoxycytidine and deoxyadenosine bases. Isobutyramide, from the deoxyguanosine base, does not have a chromophore and will not appear in the chromatogram, with the usual UV absorbance detection at ∼260 nm. The next group of peaks between 14 and 18 min are the detritylated (trityl OFF), failure sequences. Any oligonucleotide that has accidentally lost its trityl group will also appear in this region. The major peak at 30 min is the tritylated (trityl ON) product. The number of peaks in the region
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Current Protocols in Nucleic Acid Chemistry
Figure 10.5.1 Chromatogram of reversedphase analysis of tritylated 18-nt oligodeoxyribonucleotide, 5′-DMTr-TCACAGTCTGATCTCGAT-3′, using an Aquapore RP-300 column.
where trityl-bearing compounds elute indicates heterogeneity in tritylated species due to the imperfections of DNA synthesis. Longer tritylated oligonucleotides have less overall hydrophobicity. A typical chromatogram of a detritylated oligonucleotide is shown in Figure 10.5.2. The major peak at 14.5 min is the 20-nt product. This peak is surrounded by minor peaks which constitute the various failure sequences and are eluted earlier than shorter tritylated oligonucleotides due to increased charge and less net hydrophobicity. It is hard to identify peaks corresponding to by-products of DNA synthesis from relative retention time because of the
Figure 10.5.2 Chromatogram of reversedphase analysis of detritylated 20-nt oligodeoxyribonucleotide, 5′-CGAGTACTCCAA AACTAATC3′, using an Aquapore RP-300 column.
complex chromatographic effect of base composition. Oligonucleotides of shorter length than the product are usually eluted earlier than the product. Benzamide appears here at 5.9 min. Detection at 0.01 to 0.1 AUFS is typical for routine examination of the chromatogram (Fig. 10.5.1 and 10.5.2). Figure 10.5.3 shows a crude 29-nt oligonucleotide synthesized with low efficiency, 96.7% final average step-wise yield, and analyzed by anion-exchange HPLC (see Alternate Protocol) with the gradient of Table 10.5.2. The failure sequences are well resolved and separated from
Figure 10.5.3 Chromatogram of anion-exchange analysis of 29-nt oligodeoxyribonucleotide, 5′-CCATGAAGCTTTGACCATGAAAATGGAGA-3′, using a DNAPac PA-100 column.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.5.9 Current Protocols in Nucleic Acid Chemistry
Supplement 1
the product. Resolution by this system is excellent out to at least 50 bases. Sample carryover from one chromatogram to the next may occur due to buildup of sample and other hydrophobic impurities, but treatment with 50% acetonitrile will help clean the column. Oligonucleotides purified by ion-exchange HPLC must be desalted since the amount of nonvolatile salts collected with the oligonucleotide product will be substantial. Desalting can be achieved by the procedures in UNIT 10.7. Figure 10.5.4 shows an 18-nt oligonucleotide conjugated with biotin through the 5′ end using the cartridge and gradient of Table 10.5.1 (see Basic Protocol). The biotinylated product elutes at 15.2 min. A small amount of 5′-OH oligonucleotide is present as the smaller preceding peak at 12.9 min. Benzamide is present at 5.9 min. Fluorescent dyes, most commonly attached through the 5′ terminus, also impart additional hydrophobicity to the oligonucleotide (Oefner et al., 1994). The dye often is charged, either positive or negative. Like biotinylated oligonucleotides, fluorescent dye-labeled oligonucleotides elute later than their 5′ OH counter-
Analysis and Purification of Synthetic Nucleic Acids Using HPLC
parts. Fluorescence detection is a powerful aid to analysis. Figure 10.5.5 shows a crude 5′ fluorescent dye 20-nt, labeled with 6-carboxyhexachlorofluorescein oligodeoxyribonucleotide phosphoramidite-HEX (Theisen et al., 1992), eluting at 26 min using the Aquapore RP-300 column and the gradient of Table 10.5.1 (see Basic Protocol). A small amount of unlabeled 20 nt is visible at 21 min, well separated from the product. Purification of oligonucleotides Reversed-phase HPLC of tritylated oligonucleotides is the method with the largest capacity for general oligonucleotide purification and is also the most easily scaled-up (Huang and Krugh, 1990). A large cartridge or column (PRP-1, 10 × 250 mm, 10 µm diameter; Hamilton) at a flow rate of 4.5 mL/min can purify the entire product, ∼20-30 mg, from a 10 µmolscale synthesis, albeit with considerable loss of resolution relative to analytical-scale HPLC. The collection process during purification is important, since impurities can elute both before and after the product peak. Any unwanted tritylated peaks will usually elute later in the
Figure 10.5.4 Chromatogram of reversed-phase analysis of oligodeoxyribonucleotide 5′-biotinTCACAGTCTGATCTCGAT-3′, using an Aquapore RP-300 C-8 column.
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Current Protocols in Nucleic Acid Chemistry
Figure 10.5.5 Figure 10.5.5 Chromatogram of reversed-phase analysis of 5′-HEXACATCTCCCCTACCGCTATA-3′, using an Aquapore RP-300 C-8 column.
chromatogram, appearing as separate peaks or as shoulders of the main peak. The collection should start at 50% of the peak maximum on the upside of the peak to 50% of the peak maximum on the downside. The preparative chromatogram in Figure 10.5.6 was performed at 290 nm, far off the absorbance maxima of
∼260 nm, to allow for better visualization of the resolution under such overloaded conditions (Table 10.5.1). In the chromatogram, benzamide elutes at 6 min, followed by an off-scale broad absorbance by trityl-off failure sequences at 11 to 16 min. The tritylated 22 nt product elutes as an off-scale broad peak from
Figure 10.5.6 Chromatogram of reversed-phase preparative purification of 100 OD units crude tritylated 22-nt oligodeoxyribonucleotide, 5′-DMTr-GAATCACAGTCTGATCTCGATT-3′, using an Aquapore RP-300, 10 × 250–mm, 20 µm particle diameter (ABI), a flow rate of 4.5 mL/min, and detection at 290nm.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.5.11 Current Protocols in Nucleic Acid Chemistry
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26 to 31 min. The peak from 31 to 33 min is a set of tritylated failure sequences, more hydrophobic than the product since they are shorter.
Time Considerations Most HPLC runs can be conducted in less than 1 hr, including time to reequilibrate the mobile phase and conduct the attendant maintenance operations. With an autosampler, HPLC analysis can be conducted virtually around the clock. “Smart” fraction collectors can automatically collect the purified product.
Literature Cited Alul, R. 1993. Chemical synthesis of DNA and DNA analogs. In DNA Probes, 2nd ed. (G. Keller and M. Manak, eds.) pp. 69-136. Stockton Press, New York. Andrus, A. 1992. Evaluating and Isolating Synthetic Oligodeoxynucleotides. Applied Biosystems, Foster City, Calif. Available upon request. Andrus, A. 1995. Chemical methods for 5′ non-isotopic labelling of PCR probes and primers. In PCR II: A Practical Approach (M. McPherson, B. Hames, and G. Taylor, eds.) pp. 39-54. Oxford University Press, Oxford. Andrus, A. and Bloch, W. 1998. HPLC of oligonucleotides and polynucleotides. In HPLC of Macromolecules: A Practical Approach (R.W.A. Oliver, ed.) pp. 141-70, Oxford University Press, Oxford. Applied Biosystems. 1995. Improved 1-µmol RNA synthesis, analysis and purification. User Bulletin No. 91. Ausserer, W. and Biros, M. 1995. High-resolution analysis and purification of synthetic oligonucleotides with strong anion-exchange HPLC. Biotechniques 19:136-39. Bergot, B.J. and Egan, W. 1992. Separation of synthetic phosphorothioate oligodeoxynucleotides from their oxygenated (phosphodiester) defect species by strong-anion-exchange high-performance liquid chromatography. J. Chromatogr. 599:35-42. Drager, R.R. and Regnier, F.E. 1985. High-performance anion-exchange chromatography of oligonucleotides. Anal. Biochem. 45:47-56. Germann, M.W., Pon, R.T., and van de Sande. H. 1987. A general method for the purification of synthetic oligodeoxyribonucleotides containing strong secondary structure by reversed-phase high-performance liquid chromatography on PRP-1 resin. Anal. Biochem. 165:399-405. Goodchild, J. 1990. Conjugates of oligonucleotides and modified oligonucleotides: A review of their synthesis and properties. Bioconjugate Chem. 1:165-187. Analysis and Purification of Synthetic Nucleic Acids Using HPLC
Huang, G. and Krugh, T.R. 1990. Large-scale purification of synthetic oligonucleotides and carcinogen-modified oligodeoxynucleotides on a reverse-phase (PRP-1) column. Anal. Biochem. 190:21-25. Huber, C.G., Oefner, P.J., and Bonn, G.K. 1993. High-resolution liquid chromatography of oligonucleotides on nonporous alkylated styrenedivinylbenzene copolymers. Anal. Biochem. 212:351-358. Huber, C., Stimpf, E., Oefner, P., and Bonn, G. 1996. A comparison of micropellicular anion-exchange and reversed-phase stationary phases for HPLC analysis of oligonucleotides. LC/GC 14:114-127. Ikuta, S., Chattopadhyaya, R., and Dickerson, R.E. 1984. Reverse-phase polystyrene column for purification and analysis of DNA oligomers. Anal. Chem. 56:2253-2256. Kang, C., Zhang, X., Ratliff, R., Moyzis, R., and Rich, A. 1992. Crystal structure of four-stranded Oxytricha telomeric DNA. Nature 356:126. Krstulovic, A. 1987. Nucleic acids and related compounds. In Handbook of Chromatography, Vol. 1, Parts A and B (G. Zweig and J. Sherma, ed.) pp. 161-169; 35-64. CRC Press, Boca Raton, Fla. Maisano, F., Parente, D., Velati Bellini, A., Carrera, P., Zamai, M., and Grandi, G. 1989. A rapid and efficient method for the purification of synthetic oligonucleotides by high performance anion-exchange chromatography in volatile buffer. Biochromatography 4:279-281. Newton, P. 1990. Complex biological matrices: Column capacity and separation strategy. LC/GC 8:116-122. Noller, H.F. 1984. Structure of ribosomal RNA. Annu. Rev. Biochem. 53:119-162. Oefner, P. and Bonn, G. 1994. High-resolution liquid chromatography of nucleic acids. Am. Lab. June 28C-28J. Oefner, P.J., Huber, C.G., Umlauft, F., Berti, G-N., Stimpfl, E., and Bonn, G.K. 1994. High-resolution liquid chromatography of fluorescent dyelabeled nucleic acids. Anal. Biochem. 223:39-46. Pingoud, A., Fliess, A., and Pingoud, V. 1988. HPLC of oligonucleotides. In HPLC of Macromolecules (R.W.A. Oliver, ed.) pp. 183. IRL Press, New York. Sproat, B., Colonna, F., Mullah, B., Tsou, D., Andrus, A., Hampel, A., and Vinayak, R. 1995. An efficient method for the isolation and purification of oligoribonucleotides. Nucleosides Nucleotides 14:255-273. Symons, R.H. 1992. Small catalytic RNAs. Ann Rev. Biochem. 61:641-671. Theisen, P., McCollum, C,. and Andrus, A. 1992. Fluorescent dye phosphoramidite labelling of oligonucleotides. Nucl. Acid Symp. Ser. No. 27, pp. 99-100.
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Current Protocols in Nucleic Acid Chemistry
Warren, W.J. and Vella, G. 1993. Analysis and purification of synthetic oligonucleotides by highperformance liquid chromatography. In Protocols for Oligonucleotide Conjugates (S. Agrawal, ed.) pp. 233-64. Humana Press, Totowa, N.J.
Zon, G. 1993. Oligonucleoside phosphorothioates In Protocols for Oligonucleotides and Analogs (S. Agrawal, ed.) pp. 165-189. Humana Press, Totowa, N.J.
Wang, K.Y., McCurdy, S., Shea, R.G., Swaminathan, S., and Bocton, P.H. 1993. A DNA aptamer which binds to and inhibits thrombin exhibits a new structural motif for DNA. Biochemistry 32:1899-1904.
Contributed by Alex Andrus PE Applied Biosystems Foster City, California
Zon, G. 1990. Purification of synthetic oligodeoxyribonucleotides. In High Performance Liquid Chromatography in Biotechnology (W.S. Hancock, ed.) pp. 301. John Wiley & Sons, New York.
Robert G. Kuimelis Phylos, Inc. Lexington, Massachusetts
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Supplement 1
Base Composition Analysis of Nucleosides Using HPLC
UNIT 10.6
The protocol described in this unit, involving nuclease degradation of an oligonucleotide, followed by dephosphorylation and HPLC analysis of the monomers on a reversed-phase C-18 column, can detect and quantitate a wide variety of nucleobase modifications in oligonucleotides (Eadie et al., 1987; Li and Swann, 1989; Singhal et al., 1989; Andrus, 1992). The integrated areas of the nucleoside chromatogram give precise quantitation of the nucleoside composition of the oligonucleotide when the relative extinction coefficient cofactors (Table 10.6.1) are applied to the sum of the areas of the four bases. The protocol is also useful for analyzing oligonucleotides containing conjugated moieties, such as fluorophores or linkers, and carbohydrate modifications. Inclusion of nucleotide analogs in an oligonucleotide with special phosphoramidite monomers can be verified by the enzymatic digestion and base-composition analysis. Oligoribonucleotides (RNA) also undergo enzymatic digestion and can be similarly analyzed by HPLC. The elution order of nucleosides in this method is C,U,G,A (as opposed to dC, dG, T, dA). Since ribonucleosides are more hydrophilic than their corresponding deoxynucleosides, the gradients are slightly different (Table 10.6.2), but excellent results are still obtained. SAMPLE PREPARATION, DIGESTION, HPLC Oligonucleotides, DNA or RNA, can be analyzed in a crude or purified state but more accurate quantitative results can be obtained from purified oligonucleotides. The samples are digested using a master mix that contains snake venom phosphodiesterase (SVP) and bacterial alkaline phosphatase (BAP). Snake venom phosphodiesterase is an exonuclease which cleaves 3′-5′ internucleotide phosphate bonds from the 3′ terminus, yielding nucleotide 5′-monophosphates. Bacterial alkaline phosphatase effects the hydrolysis of 5′ phosphates from the nucleotide 5′-monophosphates resulting from SVP cleavage, yielding nucleoside monomers. The enzymes are active under the digest conditions with crude oligonucleotides dried directly from the ammonium hydroxide cleavage/deprotection reagent. Salts remaining from other sources, such as anion-exchange HPLC, may result in incomplete hydrolysis to nucleosides and give ambiguous results. An oligonucleotide in the range of 0.2 to 1.0 OD units is sufficient for one or more injections of the digested sample.
BASIC PROTOCOL
Materials 0.2 to 1.0 OD units (A260) oligonucleotide sample Master mix (see recipe) 3 M sodium acetate 95% ethanol Acetonitrile/triethylammonium acetate buffer/water gradient (Table 10.6.2 and UNIT 10.5)
Table 10.6.1
Molar Extinction Coefficients of Nucleosidesa
Nucleoside dA dG T dC
Extinction coefficient 15,400 11,700 8800 7300
aMeasured at pH 7.0, 25°C, 260 nm.
Contributed by Alex Andrus and Robert G. Kuimelis Current Protocols in Nucleic Acid Chemistry (2000) 10.6.1-10.6.6 Copyright © 2000 by John Wiley & Sons, Inc.
Cofactor 2.11 1.60 1.21 1.00
Purification and Analysis of Synthetic Nucleic Acids and Components
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Table 10.6.2 Gradients and Mobile Phase for DNA and RNA Digest HPLCa
Elapsed time (min) 0 5 (DNA) 15 (RNA) 35 65 70 72
%B at elapsed time 0 0 0 10 100 100 0
aColumn, Applied Biosystems Spheri-5 RP-18 # 0711-0017 (cartridge); flow rate, 0.5 mL/min; mobile phase A, 3% acetonitrile in 0.1 M triethylammonium acetate (92:5:3 deionized water: 2 M TEAA:acetonitrile); mobile phase B, 90% acetonitrile (9:1 acetonitrile:water); sample injection: 10 to 50 µL; injection cycle: 85 min.
HPLC system (UNIT 10.5) with C-18 reversed-phase column Additional reagents and equipment for reversed-phase HPLC (UNIT 10.5) Digest the samples 1. Evaporate 0.2 to 1.0 OD unit of oligonucleotide sample to dryness under vacuum in an appropriate vessel, e.g., a 1.5-mL microcentrifuge tube. 2. Vortex the master mix. Pipet 28 µL of master mix into each sample at room temperature, leaving the remainder in the tube to use as the digest blank. 3. Vortex each sample and centrifuge briefly to collect the liquid at the bottom of the tube. 4. Incubate the samples at 37°C for 8 to 24 hr. Prepare the digested samples for HPLC analysis 5. Add 4 µL of 3 M sodium acetate and 100 µL of ethanol to each sample. 6. Vortex and chill samples on dry ice for at least 10 min. 7. Centrifuge in a benchtop centrifuge at maximum speed, 5 min at room temperature. Carefully remove each supernatant with a pipet and transfer to a new, labeled tube. Discard the original tubes containing the pellets. 8. Add 300 µL ethanol to each sample. Vortex and chill samples on dry ice for at least 10 min. 9. Centrifuge in a benchtop centrifuge at maximum speed, 5 min at room temperature. Carefully remove each supernatant with a pipet, being careful not to disturb the pellet, and transfer to a new, labeled tube. If necessary leave some of the solution rather than remove any of the pellet.
10. Evaporate the samples to complete dryness under vacuum. Samples containing traces of ethanol will elute too early.
11. Dissolve the dried samples in 60 µL water and vortex each sample for at least 30 sec. If using an autosampler, transfer the samples into appropriate vials. Base Composition Analysis of Nucleosides Using HPLC
Perform HPLC of digested samples 12. Perform reversed-phase HPLC using a C-18 column (UNIT 10.5) on the digest samples using ∼0.2 OD unit per injection (this quantity is sufficient for optimum resolution
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Current Protocols in Nucleic Acid Chemistry
and sensitivity) and eluting with an acetonitrile-triethylammonium acetate buffer– water gradient as specified in Table 10.6.2. Space sample injections 85 min apart (Table 10.6.2). Set detector to 260 nm. Digestion of larger amounts of sample gives the opportunity for multiple injections.
13. Collect data for 60 min. Run a digest blank to establish an absorbance baseline profile. Even after the ethanol precipitation, the digest cocktail contains some absorbing artifacts, which are displayed as small peaks in the HPLC.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Bacterial alkaline phosphatase (BAP) The enzyme (Pharmacia Biotech) comes as a stabilized suspension in ammonium sulfate. To prepare the stock solution for the digest protocol, centrifuge the tube to pellet the salt and remove the supernatant, containing BAP. Transfer the supernatant to a tube and dilute with water to a final activity of 10 µL/unit. Store at ≤−20°C. Avoid prolonged exposure to room temperature conditions. Master mix The volumes below are given for each sample. Multiply each volume by the number of samples to be analyzed. Add one to the number to use as an HPLC digest blank control. Pipet this total volume into a tube. For example, if analyzing 10 samples, add 11 times each volume into the mixture. 20 µL freshly deionized or distilled water 0.4 µL 1.0 M MgCl2 2 µL 0.5 M Tris⋅Cl, pH 7.5 (APPENDIX 2A) 4 µL bacterial alkaline phosphatase (BAP; see recipe) 2.4 µL snake venom phosphodiesterase (SVP; see recipe) Total = 28.8 µL Prepare mix just before digestion and keep on ice Snake venom phosphodiesterase (SVP) The enzyme (Pharmacia Biotech) comes as a lyophilized powder. Dissolve the sample to 1 mg/mL. Store at ≤−20°C. Avoid prolonged exposure to room temperature conditions. COMMENTARY Background Information Although the oligonucleotide bears protecting groups during synthesis, the very reactive reagents, such as activated phosphoramidites, acetic anhydride, and iodine, can modify various sites. Modifications of the nucleobases can have a profound effect on biological and hybridization activity. Base composition analysis by enzymatic digestion and HPLC analysis make it possible to check the integrity of the nucleobases. Alternatively, one can check the integrity of oligonucleotides by MALDI mass spectrometry (UNIT 10.1).
Critical Parameters and Troubleshooting Snake venom phosphodiesterase is a very indiscriminate and robust 3′-5′ exonuclease, but occasionally base or other modifications are encountered that cause difficulty for enzyme activity. In these cases, as evidenced by incomplete digestion, alternative nucleases (e.g., S1 nuclease) are added to the mix or used separately. Some SVP samples contain varying amounts of adenosine deaminase activity. This results in a low-level conversion of adenosine or deoxyadenosine to inosine or deoxyinosine. Especially where RNA is concerned, it would
Purification and Analysis of Synthetic Nucleic Acids and Components
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Figure 10.6.1 Chromatogram after enzymatic digest of an incompletely deprotected 29-nt oligonucleotide. Obtained using Spheri 5 RP-18 column (Applied Biosystems).
be prudent to take special precautions to avoid nuclease contamination of laboratory equipment. For certain nucleoside analogs or conjugates, it may be necessary to set the detector to a wavelength other than 260 nm in order to observe the peak.
Base Composition Analysis of Nucleosides Using HPLC
Anticipated Results Figure 10.6.1 shows the chromatogram of a digest of a 29-nt oligonucleotide with incomplete deprotection of the bases. The protected deoxynucleosides, dGibu, dAbz, and dCbz elute later than the fully deprotected bases. Note that the G bases are the slowest to deprotect. Figure 10.6.2 shows the HPLC of a fully deprotected
Figure 10.6.2 Chromatogram after enzymatic digest of a completely deprotected 61-nt oligonucleotide, A13G14C14T20. Obtained using Spheri 5 RP-18 column (Applied Biosystems).
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Table 10.6.3 Calculation of Base Composition of a 61-nt Oligonucleotide (A 13G14C14T20) by Digestion and Quantitative Nucleoside HPLC Analysis
Nucleoside dA dG dC T Total
Integrated area Corrected areaa 234,732 197,491 120,783 211,529
111,247 123,432 120,783 174,812 530,274
Corrected % area 21.0 23.3 22.8 33.0 100
Found
Actual
12.8 14.2 13.9 20.1 61
13 14 14 20 61
aThe corrected area was determined by dividing the integrated area by the appropriate cofactor (Table 10.6.1).
61-nt oligonucleotide with the expected deoxynucleosides and the protecting group byproduct, benzamide. The actual base composition of an oligonucleotide can be calculated from the integrated areas of the nucleosides. Using the relative extinction coefficient cofactors (Table 10.6.1), one can divide the integrated area by the cofactor and then divide this number by the sum of the four corrected areas to find the corrected area percent (Table 10.6.3). Multiply the corrected area percent by the length of the oligonucleotide to find the base composition of each nucleoside in the oligonucleotide. In the 61-nt oligonucleotide example shown (Figure 10.6.2), the measured base composition (found) agrees closely with that expected (actual). This calculation is able to show that a particular phosphoramidite is not coupling efficiently or if a certain nucleoside is undergoing chemical modification. Also, this calculation can confirm the desired incorporation level when synthesizing oligonucleotides with mixed-base sites. This quantitative test is dependent on precise controls, so a standard mixture of known-concentration nucleoside standards should be run with each set of digestion samples to calibrate the system. The assigned cofactors are equal to the ratio of the extinction coefficient for each nucleoside to the extinction coefficient of dC. These ratios may vary with the concentration, temperature, pH, and the buffer/solvent composition. The relative extinction coefficient cofactors will be slightly different in each HPLC system, mobile phase, and gradient. Accurate quantitation of base composition is dependent on calibration with known standards. Comparison of the peak areas for a nucleoside mixture of known concentration allows one to determine the precise cofactor needed for one’s particular HPLC system and conditions. In certain contexts, the snake venom phosphodiesterase enzyme will release dinucleotides instead of mononucleotides. This can
happen, for example, in the case of 3′- or 5′conjugated oligonucleotides or when the enzyme encounters certain internucleotide modifications. This should be kept in mind when one is comparing retention times with authentic standards or quantitatively evaluating the base composition. Also, by omitting the phosphatase enzyme from the digestion mixture, it is possible to directly analyze the 5′-monophosphates (nucleotides) that make up the oligonucleotide. This approach also allows one to identify the terminal 5′ base, since it will be the only nucleoside in the mixture. Apart from sequencing the 5′-terminal base, there is no real advantage to analyzing the nucleotides, since they are very hydrophilic and are not reversed-phase HPLC columns. Anion-exchange HPLC is the dominant technique for nucleotide analysis (Krstulovic, 1987).
Time Considerations The protocol requires 85 min between each run. An HPLC system with an autosampler and data system can thus run unattended.
Literature Cited Andrus, A. 1992. Evaluating and Isolating Synthetic Oligodeoxynucleotides, Appendix 1. Applied Biosystems, Foster City, Calif. Available upon request. Eadie, J.S., McBride, L.J., Efcavitch, J.W., Hoff, L.B., and Cathcart, R. 1987. High-performance liquid chromatographic analysis of oligodeoxyribonucleotide base composition. Anal. Biochem. 165:442-447. Krstulovic, A. 1987. Nucleic acids and related compounds. In Handbook of Chromatography, Vol. 1, Part A and B (G. Zweig and J. Sherma, eds.) pp. 161-69 (A), 35-64 (B). CRC Press, Boca Raton, Fla. Li, B. and Swann, P.F. 1989. Synthesis and characterization of oligodeoxynucleotides containing O6-methyl-,O6-ethyl-, and O6-isopropylguanine. Biochemistry 28:5779-5786.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.6.5 Current Protocols in Nucleic Acid Chemistry
Supplement 1
Singhal, R.P., Landes, P., Singhal, N.P., Brown, L.W., Anevski, P.J., and Toce, J.A. 1989. Highperformance liquid chromatography for trace analysis of DNA and kinetics of DNA modification. Biochromatography 4:78-88.
Contributed by Alex Andrus PE Applied Biosystems Foster City, California Robert G. Kuimelis Phylos, Inc. Lexington, Massachusetts
Base Composition Analysis of Nucleosides Using HPLC
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Current Protocols in Nucleic Acid Chemistry
Cartridge Methods for Oligonucleotide Purification
UNIT 10.7
This unit provides protocols for DMTr (trityl)-selective (see Basic Protocol 1) and affinity desalting (see Basic Protocol 2) purification of oligonucleotides. The protocols are applicable for many of the convenient, disposable products for rapid oligonucleotide purification, clean-up by selective adsorption, and elution on solid-phase media. Many of the products are prepackaged, one-time use cartridges or columns filled with affinity or size-exclusion media. DMTr-SELECTIVE CARTRIDGE PURIFICATION This protocol describes purification of tritylated oligonucleotides with the Oligonucleotide Purification Cartridge (OPC, PE Applied Biosystems). The OPC can desalt, detritylate, and purify tritylated oligonucleotides directly from the ammonium hydroxide solution after cleavage/deprotection (McBride et al., 1988; Applied Biosystems, 1991; Blocker et al., 1991).
BASIC PROTOCOL 1
The purity level achieved by OPC is appropriate for such common applications as sequencing primers, PCR primers, and hybridization probes. Upon loading, the tritylated oligonucleotide is retained by the hydrophobic polystyrene medium, while the salts, non-tritylated oligonucleotides, and other impurities are not retained, and are washed away. The trityl group of the OPC-bound oligonucleotide is removed with a brief acid treatment, and the full-length, detritylated oligonucleotide is eluted with ∼1 mL of a 20% acetonitrile solution. Materials 20% (v/v) and 100% acetonitrile 2 M triethylammonium acetate (TEAA) Tritylated oligonucleotide in ammonia deprotection solution 3% (v/v) trifluoroacetic acid (TFA) 1.5 M ammonium hydroxide Oligonucleotide Purification Cartridges (OPC; PE Biosystems) Vacuum manifold (Analytichem or National Scientific; optional) with an in-line trap for waste and a water aspirator pump or small vacuum pump 5-mL disposable plastic syringes with exposed luer tips Prepare the cartridge 1. Mount the OPC with a clamp and insert an empty 5-mL syringe barrel on the top of the cartridge. Multiple cartridges can optionally be used with a vacuum manifold system.
2. Fill the syringe barrel with 5 mL of 100% acetonitrile. Insert and depress the plunger to pass the reagent through the OPC (into an appropriate waste container) over an elapsed time of ∼15 to 30 sec. All reagents should be eluted from the OPC cartridge dropwise.
3. Repeat step 2 with 5 mL of 2 M TEAA. Prepare the sample 4. Dilute the oligonucleotide ammonia solution (up to 20 OD units) with an equal volume of water (e.g., normally ~1 mL).
Purification and Analysis of Synthetic Nucleic Acids and Components
Contributed by Alex Andrus and Robert G. Kuimelis
10.7.1
Current Protocols in Nucleic Acid Chemistry (2000) 10.7.1-10.7.5 Copyright © 2000 by John Wiley & Sons, Inc.
Supplement 1
Apply the sample to the cartridge 5. Pass the diluted solution through the OPC at a rate of ∼1 drop/sec. Collect the eluate and pass it through a second time. 6. Wash the OPC with 5 mL of 1.5 M ammonium hydroxide, then with 10 mL deionized water, allowing the effluents to pass to waste. 7. Fill the syringe barrel with 5 mL of 3% trifluoroacetic acid (TFA) and pass ~2 mL through the OPC to waste. Let stand for 5 min before passing the remainder through to waste. 8. Pass 10 mL of deionized water through the OPC to waste. 9. For oligonucleotides >40 nt, pass 5 mL of 1.5 M ammonium hydroxide through the column, to waste, then pass 5 mL of water through the OPC to waste. If oligonucleotide is ≤40 nt, skip to step 10. Collect the purified oligonucleotide 10. Elute the column, with 1 mL of 20% acetonitrile dropwise, and collect the purified oligonucleotide in a 1.5-mL microcentrifuge tube. Quantify by OD measurement. BASIC PROTOCOL 2
AFFINITY CARTRIDGE DESALTING OF DETRITYLATED OLIGONUCLEOTIDES This protocol details the use of an affinity cartridge to desalt and concentrate detritylated oligonucleotides, with the capacity to yield up to 50 OD units of desalted oligonucleotide per cartridge. No purification is effected, beyond removal of species that fail to bind to the cartridge medium, which typically include salts, solvents, reagents, and very short failure oligonucleotides. The OPC is one example of several commercially available products that can render oligonucleotides free of salts, trace amounts of solvents, and other small molecules by immobilizing, washing, and eluting on an affinity medium. This desalting protocol is useful for (1) trityl-off oligonucleotides; (2) fluorescent dye–labeled, biotin-labeled, and other labeled oligonucleotides; (3) RNA oligonucleotides, after desilylation/deprotection; (4) polyacrylamide gel purification extracts; and (5) anion-exchange HPLC purification fractions. An alternative to this protocol is ethanol precipitation (UNIT 10.3). Materials 0.1 M triethylammonium acetate (TEAA) 50% (v/v) acetonitrile Additional reagents and equipment for cartridge purification (see Basic Protocol 1) 1. Mount cartridge and wash with acetonitrile and TEAA (see Basic Protocol 1, steps 1 to 3). 2. Concentrate up to 50 OD units of the oligonucleotide to dryness under vacuum. For maximum binding capacity, all organic solvents and ammonium hydroxide must be removed.
3. Dissolve the completely dried oligonucleotide in 2 mL of 0.1 M TEAA. Cartridge Methods for Oligonucleotide Purification
4. Pass the oligonucleotide solution through the OPC at a rate of ∼1 drop/sec. Collect the eluate and pass it through a second time. 5. Pass 15 mL of 0.1 M TEAA through the OPC to waste.
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Current Protocols in Nucleic Acid Chemistry
6. Elute the column with 1 mL of 50% acetonitrile dropwise, and collect the desalted oligonucleotide in a 1.5-mL microcentrifuge tube. Quantify by OD measurement. COMMENTARY Background Information DMTr-selective cartridge purification DMTr-selective cartridge purification separates oligonucleotides by the same hydrophobic interaction principle as reversed-phase HPLC (UNIT 10.5). Instead of a constant gradient elution with an organic modifier (i.e., acetonitrile) as in HPLC, the cartridge method is a batch-loading, washing, and elution process. The costs and complexity of the cartridge methods, in general, are far less than HPLC or PAGE (UNIT 10.4). However, the cartridge methods do not provide concomitant analytical information and are limited in capacity and efficiency of purification. Other cartridges, with similar protocols, are commercially available for purification and desalting of oligonucleotides. Check with the manufacturers for their optimum protocols. The DMTr group at the 5′ terminus is very hydrophobic and allows for specific binding of the tritylated oligonucleotide with the stationary, hydrophobic polystyrene adsorbent of OPC. Therefore, the product can be separated from other components of the crude synthesis mixture which do not bind. The crude tritylated oligonucleotide is conveniently loaded from the concentrated ammonium hydroxide cleavage/deprotection solution onto the OPC. Other cartridges packed with reversed-phase silica adsorbent, are not compatible with the low- and high-pH reagents used in Basic Protocol 1. Affinity desalting The affinity desalting protocol (Basic Protocol 2) takes advantage of the hydrophobic interactions of even untritylated oligonucleotides with the cartridge adsorbent. In the absence of the denaturing solution, ammonium hydroxide, which limits binding, the desalting protocol can bind a relatively large quantity of oligonucleotide and remove large quantities of impurities. Desalting removes impurities, such as deprotection byproducts, which may hinder some applications. Also, removal of UV-absorbing small molecules results in more accurate UV quantitation. Basic Protocol 2 is particularly useful to remove the large quantities of salts from anion-exchange HPLC purification fractions (UNIT 10.5) and the buffer and gel debris from PAGE purified oligonucleotides (UNIT 10.4).
Many labeled oligonucleotides can be partially separated from unlabeled oligonucleotides by affinity cartridges (Basic Protocol 2). Fluorescent dyes, biotin, and many other labels impart added hydrophobicity to confer reasonable binding capacity. When attempting to separate labeled oligonucleotides from unlabled oligonucleotides, it may be necessary to replace step 5 with 5 mL of 3% to 12% acetonitrile in 0.1 M TEAA. HPLC can be used to help determine the best conditions for a particular label. Many cartridge purifications can be conducted in parallel with a simple, commercially available vacuum manifold, e.g., Vac Elut SPS 24, or Vac Elut 10-place manifold (Analytichem) with a 12 × 75-mm test tube rack. The 12-position manifold (National Scientific) is also suitable. In addition to vastly increased throughput, manifolds require fewer manual manipulations. The cartridges are mounted to the suction element of the manifold and the syringe barrels are merely filled with reagents, remaining connected to the cartridge at all times. Many commercial units give excellent results, with no cross-contamination of samples (Applied Biosystems, 1991). Size-exclusion filtration Oligonucleotides can be separated from smaller molecules by passage through a medium, such as Sephadex, which allows separation based on molecular weight (this protocol is not provided here). Large molecules follow a shorter mean path length, due to exclusion from the particle pores, than smaller molecules. Unlike hydrophobic media in affinity methods, size-exclusion media have little or no hydrophobic or ion-exchange interactions with oligonucleotides. Size-exclusion filtration, also called gel-filtration or gel-permeation chromatography, is rapid and convenient but has limited separating power across a narrow mass range. The Sephadex medium is available as dry powder to be preswollen in aqueous solution, or prepacked in columns. The MicroSpin and NAP columns (Pharmacia Biotech) have quick protocols and require minimal labor. Sample volumes of 100 to 150 µL are optimally processed in a MicroSpin S-200 HR column in a microcentrifuge, whereas sample volumes between 0.2 and 2.5 mL require the appropri-
Purification and Analysis of Synthetic Nucleic Acids and Components
10.7.3 Current Protocols in Nucleic Acid Chemistry
Supplement 1
ately scaled NAP column under gravity flow. The medium is equilibrated in the solution of choice, such as water. The oligonucleotide should be loaded in a minimum volume, such as 200 µL water, and all eluate should be collected. The oligonucleotide will elute in the void volume, while smaller species will be slightly retained. A related technique utilizes disposable, membrane-based, ultrafiltration devices, e.g., Microcon (Amicon), that operate in a microcentrifuge. In this method, the “large” oligonucleotides (and other, higher-molecular-weight nucleic acids) are retained above the membrane, whereas the “small” impurities flow through and are discarded. In effect, the oligonucleotide is continuously concentrated in a diminishing volume, which is an added benefit. Disposable devices are available that can accommodate a variety of different volumes and mass ranges.
Critical Parameters and Troubleshooting
Cartridge Methods for Oligonucleotide Purification
The efficiency of DMTr-selective cartridge purification (Basic Protocol 1) depends largely on the efficiency of the synthesis process. When the detritylation, capping, and oxidation steps during synthesis are efficient, only the correctsequence oligonucleotide should bear the DMTr group, and a DMTr-selective purification protocol can furnish a very pure product. Moderately low coupling efficiency (95% to 97% average yield per cycle) will not limit purification as long as these other steps in the synthesis process are quantitative. Other side reactions, such as depurination and extraneous growth off the synthesis support, lead to trityl heterogeneity and limit purification. Shorter tritylated oligonucleotides have higher affinity for the OPC than longer tritylated oligonucleotides because they are more hydrophobic and tend to be more pure than long oligonucleotides. For example, a 20-nt oligonucleotide may yield 5 OD units, whereas a 100-nt oligonucleotide may yield only 1 OD unit of purified product. Normally, when oligonucleotides are purified (Basic Protocol 1) soon after synthesis, e.g., within a few days, the trityl group is stable in the ammonium hydroxide solution. However, upon prolonged storage, even in a freezer, the trityl group will cleave regardless of the solution it is stored in. Therefore, DMTr-selective purifications (see Basic Protocol 1; also see UNIT 10.5 for reversed-phase HPLC) should be conducted within a few days after synthesis. Triethylammonium is the re-
sulting counterion of oligonucleotides purified by Basic Protocols 1 and 2. If it is necessary to exchange the triethylammonium counterion with sodium, the oligonucleotide can be precipitated in ethanol and sodium acetate (UNIT 10.5) after cartridge purification. In Basic Protocols 1 and 2, the initial acetonitrile wash wets and swells the adsorbent. The triethylammonium acetate wash removes the acetonitrile and provides triethylammonium counter-ions to help bind the oligonucleotide. The presence of ammonia in the loading solution of the oligonucleotide provides the desired denaturing medium but limits the binding capacity of tritylated oligonucleotides. If ammonia is partially removed (by leaving for several minutes in a Speedvac evaporator or letting the sample remain uncapped for 15 min), the binding capacity of DMTr oligonucleotides will increase to yield ∼10 OD units of purified, detritylated oligonucleotide. A dilute ammonia wash follows loading to insure that non-DMTr oligonucleotides have been removed. The water wash removes residual ammonia so it will not neutralize the TFA (trifluoroacetic acid) in the following step. Detritylation with 5 mL of 3% TFA requires 5 min. The detritylation can sometimes be observed as a slight orange or pink blush to the support. Even though it no longer has a trityl group, at neutral and low pH, the purified oligonucleotide is retained while washing with water to remove residual acidic TFA. For oligonucleotides >40 nt, the second dilute ammonia wash serves to remove shorter oligonucleotides while allowing longer oligonucleotide to be retained. A water wash follows to remove residual ammonium hydroxide. Elution of the purified oligonucleotide occurs with 20% acetonitrile. The cleaved dimethoxytrityl alcohol remains on the OPC and does not contaminate the purified oligonucleotide. Reuse of any purification matrix can lead to problems with carryover and sample contamination. Avoiding these problems is especially important in synthetic DNA applications because of the potential for errant priming and hybridization. Analytical HPLC studies demonstrate that ∼5% of an oligonucleotide product is carried over as a contaminant into a second, different oligonucleotide sequence upon reuse of a purification cartridge. Also, the DMTr that is released upon TFA treatment is very tightly bound, resulting in diminished capacity and selectivity for DMTr oligonucleotides. The saf-
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est, most reliable solution is a disposable onetime use format. For longer-term storage (i.e., >6 months), oligonucleotides that have been purified by OPC (see Basic Protocol 1) should be stored as dry solids at –15°C.
Anticipated Results In Basic Protocol 1, 10 to 20 OD units of a crude, tritylated oligonucleotide (i.e., an entire 40 nmol scale and most, or all of a 0.2 µmol scale synthesis) can be loaded, to yield 1 to 10 OD units of purified product. The yield depends on initial purity, length, and concentration of ammonium hydroxide in the loading solution. Yield is measured in OD units by UV absorbance at 260 nm, converted to mass by using the approximation of 33 µg per OD unit. Oligonucleotides purified by Basic Protocol 1 are typically >90% pure by integrated peak area HPLC analysis, which compares well with HPLC (UNIT 10.5) and PAGE (UNIT 10.4) methods. In Basic Protocol 2, the yield can be as high as 50 OD units of desalted product. Synthetic oligoribonucleotides (RNA) can also be desalted and purified by DMTr-selective, affinity cartridges (Mullah and Andrus, 1996). Purification of phosphorothioate oligonucleotide analogs by the cartridge method is also very efficient. Since phosphorothioates are more hydrophobic, use 1 mL of 35% acetonitrile to elute them (Basic Protocol 1, step 10).
Time Considerations
Basic Protocol 1 requires ∼15 to 20 min. Basic Protocol 2 requires ∼10 to 15 min. Use of a vacuum manifold will speed up purification of even a single oligonucleotide and results in large time savings when purifying a larger number. For example, 10 oligonucleotides can be purified by Basic Protocol 1 on a vacuum manifold in <30 min.
Literature Cited Applied Biosystems. 1991. New Applications for the Oligonucleotide Purification Cartridge. User Bulletin No. 59. Blocker, H., Frank, R., Heisterberg-Moutsis, G., Kurth, G., and Meyerhans, A. 1991. Improved Process for the Purification of Synthetic Oligonucleotides. U.S. Patent No. 4,997,927. McBride, L., McCollum, C., Davidson, S., Efcavitch, J.W., Andrus, A., and Lombardi, S. 1988. A new, reliable cartridge for the rapid purification of synthetic DNA. Biotechniques 6:362-367. Mullah, B. and Andrus, A. 1996. Purification of 5′-O-trityl-On oligoribonucleotides. Investigation of phosphate migration during purification and detritylation. Nucleosides Nucleotides 15:419-430.
Contributed by Alex Andrus PE Applied Biosystems Foster City, California Robert G. Kuimelis Phylos, Inc. Lexington, Massachusetts
Purification and Analysis of Synthetic Nucleic Acids and Components
10.7.5 Current Protocols in Nucleic Acid Chemistry
Supplement 1
Analysis of Oxidized DNA Fragments by Gel Electrophoresis
UNIT 10.8
This unit describes electrophoretic techniques that can be used to characterize the chemistry of deoxyribose oxidation in DNA. The techniques exploit the unique electrophoretic mobility of DNA fragments possessing the various products of deoxyribose oxidation on their termini. Oxidation of each position in deoxyribose produces a spectrum of different products, some of which remain covalently attached to the DNA strand (Fig. 10.8.1). These fragments either accelerate (e.g., 3′-phosphoglycolate) or retard (e.g., 5′-nucleoside aldehyde) the mobility of the DNA fragment in a denaturing polyacrylamide gel. By altering the mobility characteristics with several simple chemical derivatizations, deoxyribose oxidation at the 3′-, 4′-, and 5′-positions can be defined and quantified with great accuracy. This technique has been used to characterize the chemistry of deoxyribose oxidation produced by radiation (Henner et al., 1983), rhodium complexes (Sitlani et al., 1992), bleomycin (Steighner and Povirk, 1990), neocarzinostatin (Dedon and Goldberg, 1990; Kappen et al., 1991), calicheamicin (Hangeland et al., 1992; Dedon et al., 1993), esperamicin (Christner et al., 1992; Yu et al., 1994), C-1027 (Xu et al., 1994), and Fe-EDTA (Balasubramanian et al., 1998), among other oxidizing agents. DETERMINATION OF 4′-OXIDATION OF DEOXYRIBOSE Oxidation of the 4′ position of deoxyribose by a variety of oxidizing agents and DNAdamaging drugs results in the formation of either of two sets of products, as shown in Figure 10.8.1. One set consists of a phosphoglycolate residue attached to the 3′ end of the broken DNA strand. This negatively charged two-carbon fragment increases the mobility
B
3'-Phosphoglycolate
4'-Oxidation
P CH2 B
P
B
P
O
O
.
P
.O
H
.
R
B
O Base propenal
O
P P B
5'-Oxidation .R H H
O H P O O
B
P O
+
B
H
O P
B
O
B
P P =
O O P O– O–
O P Nucleoside-5'-aldehyde
R.
P
O
O
H
P
+ B
.R
P Deoxyribonolactone abasic site
3'-Oxidation
O
P
4'-Keto-1'-aldehyde abasic site
O
1,4-dioxo-2-butene
H
O
P
B
P
O
+
1'-Oxidation
3'-Formylphosphate
+
O
O
O
P
P
–
BASIC PROTOCOL 1
3'-Phosphoglycoaldehyde
B
O P
+ CH2 H
O
O–
Base propenoic acid
Figure 10.8.1 Products formed by oxidation of deoxyribose in DNA. This compilation is based on studies performed with the DNA-cleaving antibiotic bleomycin and members of the enediyne antibiotic family (Dedon and Goldberg, 1992), as well as DNA-cleaving rhodium complexes (Sitlani et al., 1992).
Purification and Analysis of Synthetic Nucleic Acids and Components
Contributed by Mohamad Awada and Peter C. Dedon
10.8.1
Current Protocols in Nucleic Acid Chemistry (2001) 10.8.1-10.8.11 Copyright © 2001 by John Wiley & Sons, Inc.
Supplement 4
of the DNA fragment in a sequencing gel. The other product consists of a 4′-keto-1′-aldehyde abasic site (Fig. 10.8.1). By treatment with hydrazine, this product can be converted to a phosphopyridazine residue attached to the 3′ end of the resulting break, which retards the migration of the fragment on a sequencing gel. The presence of one or both of these oxidation products is consistent with oxidation of the 4′ position of deoxyribose. With the anticancer drug bleomycin, the ratio of the two products serves as an index of oxygen tension, since one pathway is oxygen-dependent while the other is not (Stubbe and Kozarich, 1987). The ratio may also serve as an index of the reducing environment in the vicinity of the lesion, given the decrease in the ratio of glycolate to abasic site as thiol concentrations increase (Dedon and Goldberg, 1992). Materials 5′-32P-end-labeled DNA or duplex oligonucleotide (e.g., see CPMB UNIT 3.10) 1 mg/mL sonicated calf thymus DNA (UNIT 8.1) 50 mM Tris⋅Cl (APPENDIX 2A), HEPES, or other buffer (not phosphate), containing 1 mM EDTA, pH 7 DNA-damaging agent 3 M sodium acetate, pH 7 (APPENDIX 2A) 100% (v/v) ethanol, –20°C 70% and 95% (v/v) ethanol 1 M putrescine dihydrochloride (Prepare immediately before use) 1 M aqueous hydrazine (Prepare immediately before use) Formamide sequencing gel loading buffer without NaOH (e.g., see CPMB UNIT 7.4A) Maxam-Gilbert sequencing markers prepared from the end-labeled DNA (e.g., see CPMB UNIT 7.5) 20% to 25% (w/v) polyacrylamide gel, 0.4 mm thick, containing urea and TBE electrophoresis buffer Tris/borate/EDTA (TBE) electrophoresis buffer (see, e.g., CPMB UNIT 7.6) Sequencing gel fixing solution (10% methanol, 10% acetic acid) Sephadex G-25 spin column (e.g., see CPMB UNIT 3.4) Sequencing gel apparatus (30 × 40 cm) with 0.4 mm spacers and 2000 V power supply Boiling water bath Whatman 3MM paper, larger than the size of the gel Sequencing gel drier, solvent trap, and vacuum pump (e.g., see CPMB UNIT 7.6) X-ray film or phosphor imager CAUTION: Hydrazine is extremely toxic. It should be handled with gloves, in a fume hood, and disposed of properly. Methods of disposal may vary between different institutions. Consult with the institution’s environmental safety office for the preferred means of storage and disposal of hydrazine. Perform DNA damage reaction 1. Prepare 48 µL of a solution containing ∼50,000 cpm of 32P-labeled DNA, 31.2 µg/mL calf thymus DNA (add from 1 mg/mL stock), and 50 mM buffer. 2. Add 2 µL of a solution containing the DNA-damaging agent and allow the reaction to proceed at the desired temperature. Analysis of Oxidized DNA Fragments by Gel Electrophoresis
The final concentration of the DNA-damaging agent must be determined empirically, and should provide an adequate band intensity for sequencing gel analysis. However, care should be taken to avoid loss of more than 50% of the parent DNA substrate. For example,
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Dedon et al. (1993) used calicheamicin (final concentration, 0.2 mM) with 32P-labeled DNA to induce damage adequate for visualization on sequencing gels.
3. At the desired end-point, stop the reaction as necessary and purify the DNA either by ethanol precipitation or by passage of the reaction mixture over a Sephadex G-25 spin column (to remove salts, metals, etc.) followed by ethanol precipitation. Perform ethanol precipitation as follows: a. Add 1/10 vol 3 M sodium acetate and then 2 to 3 vol 100% ethanol, –20°C. b. Centrifuge ≥30 min at 16,000 × g, 4°C. c. Carefully decant the supernatant or remove it with a pipet. Rinse the pellet with 70% ethanol (to remove salts), and then rinse again with 95% ethanol. The need to terminate the DNA-damaging reaction depends entirely on the damaging species and its concentration in the reaction mixture (i.e., for agents present in vast excess of the DNA substrate). For example, enediyene antibiotics are labile species that produce only one DNA-damage event per molecule (Dedon et al., 1993). Reactions with enediyenes are thus self-terminating. However, reactions with redox cycling metals such as Fe(EDTA) may require termination with a metal-chelating agent, such as thiourea in this case (Balasubramanian et al., 1998), to avoid excessive DNA damage.
Prepare DNA sample for chemical derivatization reactions 4. Dry the sample under vacuum and resuspend the DNA in 30 µL deionized water. 5. Split the sample into three 10-µL portions and place one portion on ice as the control. 6. To another 10-µL portion, add 1/10 vol of 1 M putrescine dihydrochloride, and incubate the sample for 1 hr at 37°C to cleave abasic sites. 7. To the final 10-µL portion, add 1/10 vol of 1 M hydrazine solution and incubate the sample 1 hr at room temperature to convert the 4′-keto-1′-aldehyde abasic site to a phosphopyridazine residue. The phosphoglycolate, if present will form in all three samples.
8. Precipitate all three DNA samples with ethanol as described in step 3. 9. Dry the DNA under vacuum and dissolve in 4 µL formamide sequencing gel loading buffer. It is important to use a loading buffer that is not alkaline (e.g., that contains no NaOH). Alkaline pH will cause hydrolysis of abasic sites in DNA and lead to erroneous conclusions regarding the quantity of phosphate-ended fragments and abasic sites in the damaged DNA.
Load and run sequencing gel 10. Prerun a 20% to 25% polyacrylamide gel (0.4 mm thick) at 1500 to 2000 V for ∼1 hr to heat the gel and glass plates to 40° to 50°C. Use TBE electrophoresis buffer as the running buffer. 11. Place a 2-µl aliquot of each of the DNA samples and Maxam-Gilbert sequencing markers in a capped centrifuge tube, place tubes in a boiling water bath for 1 min, and plunge the tubes into ice water. 12. Load the 2-µL DNA samples onto the gel, bracketing the samples with 2-µL aliquots of the Maxam-Gilbert sequencing markers containing roughly equal quantities of radioactivity as the DNA samples. 13. Using the bromphenol blue and xylene cyanol dyes as DNA size indices, run the sequencing gel at the maximal voltage that does not cause the temperature of the glass
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Supplement 4
plates to exceed 60°C, and until the DNA fragment size of interest has migrated to within 10 cm of the bottom of the gel. It is important to run the gel until the DNA fragment size of interest has migrated as far as possible without running off the gel. An optimal run will allow the glycolate-ended fragments to be well resolved from the phosphate-ended fragments. On a 20% sequencing gel, the bromphenol blue migrates as if it were an 8-nucleotide DNA fragment while the xylene cyanol migrates as if it were 28 nucleotides long.
14. Dismantle the electrophoresis apparatus, allow the glass plates to cool to room temperature, and then pry the plates apart. Pry the plates apart carefully to avoid ripping the gel. Scrupulous cleaning of the glass plates is essential, with regular overnight soaking in 1 M NaOH. It is often helpful to coat one of the plates with a silanizing reagent such as Sigmacote (Sigma).
15. Place the gel and its attached plate into a tray containing a sheet of Whatman 3MM paper and enough fixing solution to cover the gel, and gently agitate until the gel detaches from the plate and floats freely in the solution. If the gel is to be dried, fixation is essential, since 20% sequencing gels will crack unless soaked in the fixation solution to remove the urea. Alternatively, the wet gel on its plate can be wrapped in plastic wrap and exposed to X-ray film or a phosphor imager plate. However, the thickness of the wet gel will reduce the resolution of the sequencing gel by increasing the fuzziness of the bands.
16. Carefully slide the glass plate out from underneath the gel, allowing the gel to settle onto the Whatman paper, and remove the fixation solution using vacuum suction. While removing the fixation solution from the tray, use gloved hands to keep the gel positioned over the Whatman paper.
17. Carefully lift the paper out of the tray and dry it in a gel drier set to ramp slowly up to 70°C. Do not break the vacuum until the gel is completely dry, or the gel will crack extensively. The gel is dry if the entire surface is hot to the touch and the gel no longer has any perceptible thickness; cool spots represent evaporating water.
18. Remove the dried gel from the drier and wrap it in plastic wrap for exposure to X-ray film or a phosphor imager plate. The length of exposure will vary depending on the quantity of radioactivity. An overnight exposure of X-ray film should be adequate for a gel containing lanes with 10,000 to 20,000 cpm of radioactivity; shorter times are needed for a phosphor imager plate. As illustrated in Figure 10.8.2, phosphoglycolate-ended DNA fragments in the control lane will appear as a band migrating ∼1/4 nucleotide faster than the phosphate-ended fragment co-migrating with the Maxam-Gilbert sequencing standard. Treatment with hydrazine converts the 4′-keto-1′-aldehyde abasic site to a strand break with a phosphopyridazine residue attached to the 3′ end. The phosphopyridazine-ended fragment migrates 1 to 2 nucleotides more slowly than the phosphate-ended fragment (Fig. 10.8.2). Putrescine cleaves all types of abasic sites (simple and oxidized; Lindahl and Andersson, 1972; Dedon et al., 1992) to phosphate-ended fragments that co-migrate with the phosphate-ended fragments of the Maxam-Gilbert sequencing reactions. BASIC PROTOCOL 2 Analysis of Oxidized DNA Fragments by Gel Electrophoresis
DETERMINATION OF 5′-OXIDATION OF DEOXYRIBOSE As with 4′-oxidation, oxidation of the 5′ position of deoxyribose results in the formation of either of two sets of products, as shown in Figure 10.8.1. One set consists of a formylphosphate residue attached to the 3′ end of the broken DNA strand. The
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Current Protocols in Nucleic Acid Chemistry
Figure 10.8.2 Gel electrophoretic analysis of the products formed by oxidation of the 4′ position of deoxyribose in DNA. All studies were performed with a 5′-32P-labeled DNA molecule. (A) 20% sequencing gel analysis of products formed by esperamicin C cleavage at the A of CAT; the fragment is 21 nucleotides long. Lane C: Control DNA showing a phosphate-ended fragment (1) and a 3′-phosphoglycolate-ended fragment (2). Lane H: Treatment with hydrazine converts a 4′-keto-1′aldehyde abasic site to a 3′-phosphopyridazine-ended fragment (3). Lane P: Treatment with putrescine converts the abasic site to a 3′-phosphate-ended fragment (1). (B) 20% sequencing gel analysis of products formed by reaction of bleomycin (Bleo) and neocarzinostatin (Neo) at the T of GTT; the fragment is 10 nucleotides long. Bleomycin produced only a 3′-phosphoglycolate-ended fragment (2) while neocarzinostatin produced both phosphate- (1) and phosphoglycolate-ended (2) fragments. (C) 25% sequencing gel analysis of products formed by calicheamicin cleavage at the C of AGGATC; the resulting fragment is 34 nucleotides long. The sample was treated with hydrazine, so the gel shows fragments containing phosphate (1), phosphoglycolate (2), and phosphopyridazine (3) residues. Lanes marked AG and CT represent Maxam-Gilbert chemical sequencing standards, all with 3′-phosphate groups.
formyl group is highly reactive and undergoes rapid hydrolysis or transfer to nucleophiles in solution (e.g., Tris; Chin et al., 1987), and so will not be apparent in a sequencing gel. The other product is a strand break containing a nucleoside aldehyde residue attached to the 5′ end (Fig. 10.8.1). This residue causes the fragment to migrate 1 to 4 nucleotides more slowly than a phosphate-ended fragment. The nucleoside aldehyde can be reduced with sodium borohydride to a nucleoside, which migrates slightly more rapidly than the nucleoside aldehyde, though more slowly than a phosphate-ended fragment. The nucleoside aldehyde is also alkali labile, so that treatment of the damaged DNA with 0.5 M piperidine or 100 mM sodium hydroxide will result in removal of the nucleoside aldehyde and formation of a phosphate-ended fragment. CAUTION: Piperidine is a toxic chemical. It should be handled with gloves, in a fume hood, and disposed of properly. Methods of disposal may vary between different institutions. Consult with the institution’s environmental safety office for the preferred means of storage and disposal of piperidine stock solutions. Materials 3′-32P-end-labeled DNA or duplex oligonucleotide (e.g., see CPMB UNIT 3.6) 1 M HEPES buffer, pH 7 1 M sodium borohydride (prepare immediately before use) 0.5 M piperidine (freshly prepared) 8% (w/v) polyacrylamide gel, 0.4 mm thick, containing urea and TBE electrophoresis buffer 90°C water bath
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10.8.5 Current Protocols in Nucleic Acid Chemistry
Supplement 4
Additional reagents and equipment for determination of 4′-oxidation of DNA (see Basic Protocol 1) Perform DNA damage and chemical derivatization 1. Proceed with the DNA damage reaction as described above (see Basic Protocol 1, steps 1 to 5). 2. Precipitate the DNA in one of the 10-µL portions as described (see Basic Protocol 1, step 3); this is the control sample. Reduce DNA in one sample with sodium borohydride 3. To one of the 10-µL portions, add 35 µL water and 5 µL of 1 M HEPES buffer with mixing. 4. Add 2.5 µL of freshly prepared 1 M sodium borohydride and incubate 15 min on ice. 5. Repeat the addition of another 2.5-µL aliquot of freshly prepared 1 M sodium borohydride and incubate the sample for 15 min on ice. 6. Precipitate the DNA by adding 4 µL of 3 M sodium acetate and 150 µL of 100% ethanol, and then process the DNA as described (see Basic Protocol 1, steps 3b and 3c). Dry sample under vacuum. The sodium borohydride contributes a 100 mM concentration of sodium, so add only enough 3 M sodium acetate to bring the sodium concentration to 300 mM for the ethanol precipitation. Alternatively, the DNA can be desalted by passage over a Sephadex G-25 spin column and purified by ethanol precipitation.
Treat DNA in one sample with piperidine 7. Evaporate the third 10-µL portion of the damaged DNA to dryness under vacuum, resuspend the DNA in 50 µL of 0.5 M piperidine, and heat the sample at 90°C for 15 min. 8. Evaporate the piperidine solution from sample under vacuum and resuspend the DNA in 10 µL water. Repeat the evaporation and addition of water twice more, ending with a dried sample after the third evaporation. Load and run sequencing gel 9. Resuspend the DNA from steps 2, 6, and 8 each in 4 µL of formamide sequencing gel loading buffer without NaOH. 10. Load and run an 8% sequencing gel as described (see Basic Protocol 1; steps 10 to 18). The large shifts in DNA fragment mobility associated with the nucleoside aldehyde residue and its reduced nucleoside form usually do not require high resolution (e.g., 20%) sequencing gels for analysis. As illustrated in the control lane of Figure 10.8.3, a DNA fragment possessing a nucleoside-5′-aldehyde at the 5′ end will migrate 1 to 4 nucleotides more slowly than the corresponding phosphate-ended fragment (i.e., a fragment lacking the nucleoside aldehyde). Reduction of the nucleoside-5′-aldehyde with sodium borohydride results in the formation of a nucleoside-ended DNA fragment that migrates slightly faster than the nucleoside-aldehyde-ended fragment. Piperidine treatment results in removal of the nucleoside-aldehyde moiety and leaves a phosphate-ended fragment. Analysis of Oxidized DNA Fragments by Gel Electrophoresis
10.8.6 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Figure 10.8.3 Gel electrophoretic analysis of the products formed by oxidation of the 5′ position of deoxyribose in DNA. This is an 8% polyacrylamide sequencing gel containing the products formed by neocarzinostatin cleavage at the T of ACT. Lane C: Control DNA showing a phosphate-ended fragment (1) and a fragment possessing a nucleoside-5′-aldehyde aldehyde (2). Lane Pip: Treatment with piperidine cleaves the nucleoside-5′-aldehyde to produce a phosphate-ended fragment. Lane NaB: Reduction of the nucleoside-5′-aldehyde with sodium borohydride produces a nucleoside-ended fragment (3) that migrates slightly ahead of the nucleoside aldehyde.
DETERMINATION OF 3′-OXIDATION OF DEOXYRIBOSE As shown in Figure 10.8.1, oxidation of the 3′ position of deoxyribose results in the formation of a phosphoglycoaldehyde residue attached to the 3′ end of the resulting DNA fragment. The glycoaldehyde can be reduced with sodium borohydride to glycol, which causes the DNA fragment to migrate slightly faster than the glycoaldehyde-ended fragment, but more slowly than the corresponding phosphate-ended fragment. Oxidation of the glycoaldehyde with sodium chlorite results in the formation of a glycolate residue that causes the fragment to migrate slightly faster than the corresponding 3′-phosphateended fragment as discussed in Basic Protocol 1.
BASIC PROTOCOL 3
CAUTION: Piperidine is a toxic chemical. It should be handled with gloves, in a fume hood, and disposed of properly. Methods of disposal may vary between different institutions. Consult with the institution’s environmental safety office for the preferred means of storage and disposal of piperidine stock solutions. Materials 0.1 M sodium chlorite in 20 mM potassium phosphate buffer, pH 4 (see APPENDIX 2A for buffer) 0.1 M sodium sulfite (Na2SO3) Additional reagents and equipment as for 4′- and 5′-oxidation of DNA (see Basic Protocols 1 and 2) Prepare damaged DNA for chemical derivatization 1. Proceed with the DNA damage reaction as described above (see Basic Protocol 1, steps 1 to 3). 2. Dry the sample under vacuum and resuspend the DNA in 40 µL of deionized water. Split the solution into four 10-µL aliquots. 3. Precipitate the DNA in one of the 10-µL portions (see Basic Protocol 1, step 3); this is the control sample. Oxidize DNA in one sample with sodium chlorite 4. To one of the 10-µL portions, add 1 µL of 0.1 M sodium chlorite solution and incubate at room temperature for 6 hr. 5. Quench the reaction by adding 1 µL of 0.1 M sodium sulfite.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.8.7 Current Protocols in Nucleic Acid Chemistry
Supplement 4
6. Remove salts by adding 30 µL of water and passing the sample over a Sephadex G-25 spin column. 7. Precipitate the DNA from the column filtrate as described (see Basic Protocol 1, step 3). Reduce DNA in one sample with sodium borohydride 8. Treat one of the 10-µL DNA aliquots with sodium borohydride as described above (see Basic Protocol 2, steps 3 to 6). Convert phosphoglycoaldehyde residues to phosphate-ended fragments 9. Treat the fourth 10-µL portion with piperidine as described (see Basic Protocol 2, steps 7 and 8). Load and run sequencing gel 10. Resuspend the DNA in samples from steps 3, 7, 8, and 9 in 4 µL of formamide sequencing gel loading buffer without NaOH. 11. Load and run a 20% sequencing gel as described (see Basic Protocol 1, steps 10 to 18). As illustrated in the control lane of Figure 10.8.4, a 16-nucleotide DNA fragment possessing a 3′-phosphoglycoaldehyde residue will migrate ∼1 nucleotide more slowly than the corresponding phosphate-ended fragment (i.e., a fragment lacking the phosphoglycoaldehyde). Reduction of the phosphoglycoaldehyde with sodium borohydride results in the formation a 3′-phosphoglycol-ended DNA fragment that migrates slightly faster than the 3′-phosphoglycoaldehyde-ended fragment but more slowly than the corresponding 3′phosphate-ended fragment. Oxidation of the glycoaldehyde to glycolic acid with sodium chlorite results in the formation of a fragment that migrates faster than the corresponding 3′-phosphate-ended fragment. This behavior was described for products of oxidation of the 4′ position of deoxyribose in Basic Protocol 1. Piperidine treatment results in removal of the glycoaldehyde moiety and leaves a phosphate-ended fragment.
Analysis of Oxidized DNA Fragments by Gel Electrophoresis
Figure 10.8.4 Gel electrophoretic analysis of the products formed by oxidation of the 3′ position of deoxyribose in DNA. This is a 20% polyacrylamide sequencing gel containing a synthetic 16-mer oligonucleotide possessing a 3′-phosphoglycoaldehyde residue and the associated products of chemical derivatization. Lane GA: 3′-phosphate-ended Maxam-Gilbert chemical sequencing standards for G and A; the band marked 1 is the 3′-phosphate-ended 16-mer fragment. Lane C: Control DNA fragment with a 3′-phosphoglycoaldehyde residue. Lane Red: Treatment of the glycoaldehydeended fragment with sodium borohydride produces a 3′-phosphoglycol-ended fragment (2). Lane Ox: Oxidation of the glycoaldehyde-ended fragment with sodium chlorite produces a 3′-phosphoglycolate-ended fragment (3). The other band present in this lane represents an unidentified intermediate in chlorite oxidation of 3′-phosphoglycoaldehyde. The product can be avoided by longer incubation with chlorite.
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COMMENTARY Background Information The experiments presented in this unit are intended as a first step in defining the chemistry of deoxyribose oxidation produced by hydrogen atom abstraction at the 3′-, 4′-, and 5′-positions. The technique exploits the unique shifts in polyacrylamide gel migration of DNA fragments with damage residues attached to the 3′ and 5′ ends. The change in mobility is judged against the phosphate-ended fragments generated by Maxam-Gilbert chemical sequencing reactions. Gel mobility shift assays have been used successfully to define the deoxyribose oxidation chemistry produced by agents such as the enediyne antibiotics neocarzinostatin (Dedon and Goldberg, 1990; Kappen et al., 1991), esperamicin (Christner et al., 1992; Yu et al., 1994), calicheamicin (Hangeland et al., 1992; Dedon et al., 1993), and C-1027 (Xu et al., 1994), as well as the antitumor agent bleomycin (Steighner and Povirk, 1990), rhodium complexes (Sitlani et al., 1992), ionizing radiation (Henner et al., 1983), and Fe-EDTA (Balasubramanian et al., 1998). Gel electrophoretic analysis of deoxyribose oxidation products finds utility in several aspects of DNA damage research. The first is primary estimation of the DNA damage chemistry. The position of deoxyribose oxidation by a DNA-cleaving agent with limited sequence selectivity (i.e., nonrandom cleavage) can be estimated using the mobility shift assay with chemical derivatization. To avoid the ambiguities associated with a single gel shift observation, each protocol involves several chemical derivatization steps to fully characterize the products associated with oxidation at each position in deoxyribose. Even with the inclusion of additional chemical derivatization steps, however, the gel shift assay cannot replace the rigor of complete chemical characterization of the deoxyribose degradation products. Furthermore, the assay does not provide definitive evidence for the deoxyribonolactone abasic site produced by 1′-hydrogen atom abstraction from deoxyribose (Yu et al., 1994), though this abasic site is resistant to hydrazine derivatization and sensitive to putrescine cleavage (Yu et al., 1994). Once the chemical identity of the damage has been established, a second application involves quantitative and qualitative comparison of deoxyribose oxidation chemistry at various damage sites. For example, the enediyne antibiotic neocarzinostatin cleaves DNA to pro-
duce lesions caused by both 4′- and 5′-oxidation at the same site (Kappen et al., 1991). The shuttling between the 4′ and 5′ positions can be directed by site-specific deuterium labeling, with quantitation of the isotope effect defined by gel shift analysis of the various products of 4′- and 5′-oxidation chemistry (Kappen et al., 1991). A third application involves an assessment of the partitioning of chemical reactions occurring at a single site in deoxyribose. For example, the chemistry of a bleomycin-induced 4′hydrogen atom undergoes abstraction partitioning along two pathways to produce either a phosphoglycolate-ended fragment or a 4′keto-1′-aldehyde abasic site (Fig. 10.8.1). This partitioning varies as a function of the oxygen concentration since the glycolate is oxygen dependent while the abasic site does not require oxygen for its formation (McGall et al., 1987). Another example involves neocarzinostatin and calicheamicin. These enediynes produce double-strand lesions at several sites in DNA with 4′-deoxyribose oxidation on one strand producing both glycolate and the abasic site. However, in the single-strand lesions produced at the same site, the sole product is glycolate (Dedon et al., 1993). This suggests that the partitioning of enediyne-mediated 4′-chemistry involves interaction between lesions on each strand. Combined with the sensitivity of phosphor imager analysis, the gel shift assay allows sensitive and quantitative determination of deoxyribose oxidation chemistry at multiple sites and within a single site.
Critical Parameters There are several factors critical to the successful application of the gel shift technique. An important factor is the length of the sequencing gel used to resolve glycolate- from phosphate-ended fragments. A 40-cm sequencing gel will allow resolution of these species for DNA fragments up to 30 to 40 bp, with migration of the fragments of interest at least to the bottom quarter of the gel. Better resolution of 30- to 40-bp glycolate- and phosphateended fragments can be achieved on a 60-cm gel, though this may require running times of 24 to 48 hr. Interpretation of the bands representing cleavage products also requires a reasonable separation of cleavage sites. Optimal results are obtained with an agent that cleaves DNA at sites separated from one another by more than 3 to
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4 nucleotides. In the “random” cleavage patterns produced by ionizing radiation and FeEDTA, the proximity of glycolate-ended DNA fragments to their phosphate-ended counterparts permits ready identification of these fragments. However, nucleoside aldehyde residues and phosphopyridazine derivatives migrate several nucleotides distant to their phosphateended counterparts and there is more significant variation in their mobility as a function of length relative to glycolate-ended fragments. This makes their identification difficult due to the complexity of the band pattern of closely spaced fragments. The stability of several of the deoxyribose oxidation products presents a problem for quantitative studies. The abasic sites and the nucleoside aldehyde residue are susceptible to degradation at alkaline pH and elevated temperature. To retain these lesions, samples should be processed quickly, with the DNA samples kept on ice and manipulations minimized to avoid degradation of the labile structures. It is important to add HEPES buffer to avoid the alkaline pH generated during the sodium borohydride reduction. Finally, it is important to use freshly prepared DNA substrates for these studies. Radiolytic degradation of the DNA will lead to a high background of strand breaks possessing a variety of fragment end-chemistries. Minimization of 32P-mediated degradation can be achieved by (1) preparing the DNA within 24 hr of use; (2) storing the DNA in Tris, HEPES, or other amine-containing buffer; and (3) diluting the DNA as much as possible for storage.
Troubleshooting Several problems may be encountered during the experiments. Inadequate resolution of DNA fragments can be solved by decreasing the length of the DNA substrate under study or by increasing the length of the gel as discussed above. Excessive background due to cleavage produced by 32P can be handled by proper storage conditions (see Critical Parameters). If salts are not removed adequately by Sephadex chromatography and ethanol precipitation, the sequencing gel will show progressively narrower lanes with fuzzy, poorly resolved bands.
Anticipated Results Analysis of Oxidized DNA Fragments by Gel Electrophoresis
The experiments described in this unit will produce results similar to those shown in Figure 10.8.2, Figure 10.8.3, and Figure 10.8.4. The reader can also review the cited references for other examples of the application of gel mobil-
ity analysis to the solution of DNA oxidation chemistry.
Time Considerations Preparation of DNA substrates will require one day while performance of the damage reactions and processing of the DNA samples will require an additional day. The length of time required to run the sequencing gels will depend on the length of the DNA fragment. Oligonucleotides of 10 to 30 bp require several hours to adequately resolve glycolate-ended fragments from phosphate-ended fragments on a 40-cm gel; longer DNA fragments necessitate runs of as much as 24 to 48 hr on a 60-cm sequencing gel. Processing of the sequencing gels may require several hours to allow for fixation and drying, with exposure of X-ray film or phosphor imager plates requiring several hours to overnight.
Literature Cited Balasubramanian, B., Pogozelski, W.K., and Tullius, T.D. 1998. DNA strand breaking by the hydroxyl radical is governed by the accessible surface areas of the hydrogen atoms of the DNA backbone. Proc. Natl. Acad. Sci. U.S.A. 95:9738-9743. Chin, D.-H., Kappen, L.S., and Goldberg, I.H. 1987. 3′-Formyl phosphate-ended DNA: High energy intermediate in antibiotic-induced DNA sugar damage. Proc. Natl. Acad. Sci. U.S.A. 84:70707074. Christner, D.F., Frank, B.L., Kozarich, J.W., Stubbe, J., Golik, J., Doyle, T.W., Rosenberg, I.E., and Krishnan, B. 1992. Unmasking the chemistry of DNA cleavage by the esperamicins: Modulation of 4′-hydrogen abstraction and bistranded damage by the fucose-anthranilate moiety. J. Am. Chem. Soc. 114:8763-8767. Dedon, P.C. and Goldberg, I.H. 1990. Sequencespecific double-strand breakage of DNA by neocarzinostatin within a staggered cleavage site. J. Biol. Chem. 265:14713-14716. Dedon, P.C. and Goldberg, I.H. 1992. Influence of thiol structure on neocarzinostatin activation and expression of DNA damage. Biochemistry 31:1909-1917. Dedon, P.C., Jiang, Z.-W., and Goldberg, I.H. 1992. Neocarzinostatin-mediated DNA damage in a model AGT.ACT site: Mechanistic studies of thiol-sensitive partitioning of C4′ DNA damage products. Biochemistry 31:1917-1927. Dedon, P.C., Salzberg, A.A., and Xu, J. 1993. Exclusive production of bistranded DNA damage by calicheamicin. Biochemistry 32:3617-3622. Hangeland, J.J., De Voss, J.J., Heath, J.A., Townsend, C.A., Ding, W.-D., Ashcroft, J.S., and Ellestad, G.A. 1992. Specific abstraction of the 5′(S)- and 4′-deoxyribosyl hydrogen atoms from DNA by calicheamicin γ1I. J. Am. Chem. Soc. 114:9200-9202.
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Henner, W.D., Rodriguez, L.O., Hecht, S.M., and Haseltine, W.A. 1983. Gamma ray induced deoxyribonucleic acid strand breaks: 3′ glycolate termini. J. Biol. Chem. 258:711-713. Kappen, L.S., Goldberg, I.H., Frank, B.L., Worth, L.J., Christner, D.F., Kozarich, J.W., and Stubbe, J. 1991. Neocarzinostatin-induced hydrogen atom abstraction from C-4′ and C-5′ of the T residue at a d(GT) step in oligonucleotides: Shuttling between deoxyribose attack sites based on isotope selection effects. Biochemistry 30:20342042. Lindahl, T. and Andersson, A. 1972. Rate of chain breakage at apurinic sites in double-stranded deoxyribonucleic acid. Biochemistry 11:36183623. McGall, G., Rabow, L., Stubbe, J., and Kozarich, J.W. 1987. Incorporation of 18O into glycolic acid obtained from the bleomycin-mediated degradation of DNA: Evidence for 4′-radical trapping by 18O2. J. Am. Chem. Soc. 109:2836-2837. Sitlani, A., Long, E.C., Pyle, A.M., and Barton, J.K. 1992. DNA photocleavage by phenanthrenequinone diimine complexes with rhodium (III): Shape-selective recognition and reaction. J. Am. Chem. Soc. 114:2303-2312.
Steighner, R.J. and Povirk, L.F. 1990. Bleomycininduced DNA lesions at mutational hot spots: Implications for the mechanism of double-strand cleavage. Proc. Natl. Acad. Sci. U.S.A. 87:83508354. Stubbe, J. and Kozarich, J.W. 1987. Mechanisms of bleomycin-induced DNA degradation. Chem. Rev. 87:1107-1136. Xu, Y.-J., Zhen, Y.-S., and Goldberg, I.H. 1994. C1027 chromophore, a potent new enediyne antitumor antibiotic, induces sequence-specific double-strand DNA cleavage. Biochemistry 33:5947-5954. Yu, L., Golik, J., Harrison, R., and Dedon, P. 1994. The deoxyfucose-anthranilate of esperamicin A1 confers intercalative DNA binding and causes a switch in the chemistry of bistranded DNA lesions. J. Am. Chem. Soc. 116:9733-9738.
Contributed by Mohamad Awada and Peter C. Dedon Massachusetts Institute of Technology Cambridge, Massachusetts
Purification and Analysis of Synthetic Nucleic Acids and Components
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Capillary Electrophoresis of DNA
UNIT 10.9
DNA fragments are traditionally separated and analyzed by slab gel electrophoresis. The gel matrices are usually either polyacrylamide (UNIT 10.4) or agarose, and separations are achieved in the presence (for ssDNA) or the absence (for dsDNA) of dissociating agents such as urea or formamide. The slab gel systems have the advantage of analyzing multiple samples in the same separation at low cost, but normally take several hours to complete. The DNA is typically visualized with stains, UV shadowing, intercalating dyes such as ethidium bromide, and on occasion by radioactivity. Capillary electrophoresis (CE), an alternative to conventional slab gel electrophoresis, has developed over the past few years into a very powerful tool for the separation of DNA fragments. CE offers a number of advantages over slab gel separations in terms of speed, resolution, sensitivity, and data handling. This is partly because the CE separation occurs inside a small-diameter (50- to 100-µm) quartz capillary in the presence of high (kilovolt-level) separating voltages. Separation times are generally only a few minutes. The DNA is detected either by UV absorption or by fluorescent labeling, both of which eliminate the need to use mutagenic substances (e.g., ethidium bromide) or dispose of radioactive waste. The quantity of DNA required for the separation is in the nanogram range. Single-base resolution can be readily obtained on fragments up to several hundred base pairs in size. In the presence of appropriate standards, fragments can be accurately sized, based on relative electrophoretic mobility. The separation of DNA fragments by CE occurs within the walls of a fused-silica capillary. Since the negatively charged nature of this surface has a dramatic impact on the resolution achieved during the separations, the vast majority of CE separations are done in “coated” capillaries whose surface has been modified to be chemically inert to the DNA. The capillaries are filled with a sieving matrix, and the DNA fragments are separated on the basis of size, analogously to standard slab gel separations. The matrix is either a chemically cross-linked gel, such as polyacrylamide, or a flowable polymer, such as modified cellulose or non-cross-linked polyacrylamide. Single-stranded DNA (ssDNA) fragments as small as 5 bases are readily separated with single-base resolution. The analysis of synthetic oligonucleotides in a flowable matrix is described in this unit (see Basic Protocol 1) as an example of this type of application. Fragments of double-stranded DNA (dsDNA) as large as 20 kb are also separated, although not with single-base-pair resolution. The only difference between these separations is the separation matrix. CE has found increasing use in a number of analytical applications where DNA separations are required. These include assessment of the purity of synthetic oligonucleotides and their modifications, analysis of PCR products, sequencing of fluorescent DNA, analysis of restriction maps, accurate sizing of restriction fragments for genetic analysis, forensic analysis of biological samples, genotyping, and analysis of conformational polymorphisms. Additional applications continue to be developed. An area of growing interest is the ability to analyze low levels of PCR products in biological fluids, as presented below (see Basic Protocol 2). Rapid progress is also being made in the development of multicapillary automated DNA sequencing instruments using laser fluorescence detection. CE is an analytical technique rarely used in preparative mode. This is largely because only small quantities of DNA can be loaded onto a capillary. Amplifying DNA by PCR after separation can circumvent this problem. In general, however, preparative separation of DNA fragments is best achieved by slab gel electrophoresis (UNIT 10.4) or high-performance liquid chromatography (HPLC) methods (UNIT 10.5). Contributed by Alan Smith and Robert J. Nelson Current Protocols in Nucleic Acid Chemistry (2003) 10.9.1-10.9.16 Copyright © 2003 by John Wiley & Sons, Inc.
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INSTRUMENTATION CE separation in its simplest form can be achieved by passing a high voltage between two buffer reservoirs that are joined by a fused silica capillary filled with liquid or gel. This results in an electric field that drives the molecules of interest from one end of the capillary to the other. The capillaries are generally 20 to 80 cm long and 50 to 100 µm in internal diameter, with total volumes in the 1- to 2-µl range. For comparison, the volume of a slab gel lane is ∼1000 µl. The capillaries are thin walled, which allows for dissipation of the Joule heating resulting from the high voltages (10 to 30 kV) that are necessary for high-performance electrophoretic separations. This minimizes convective effects that could result in band broadening during electrophoresis. The fused-silica capillary is coated on the outside with a polyimide layer that eliminates oxidation of the fused-silica glass and confers excellent tensile strength to the otherwise fragile capillary. The polyimide sheathing is carefully burned from a small portion of the capillary to expose a section of the silica. This clear section of the capillary is inserted into the light path of a UV or fluorescence detector and becomes the on-column flow cell. As the DNA molecules migrate through the capillary as a result of the electric field, they pass through the detector light path and are measured by UV or fluorescence detection. In effect, the separation column itself becomes a very-low-volume flow cell. IMPORTANT NOTE: Removal of the polyimide coating makes the capillaries susceptible to breakage. Capillaries that are not provided in cartridges by the manufacturer should be handled with care to avoid breakage. The combination of high field strength and large surface-area-to-volume ratio of the capillaries results in rapid and very efficient separations of both ssDNA and dsDNA. Sample loading can be accomplished from as little as 1 µl, with starting sample concentrations of ∼1 µg/ml for UV detection and ∼1 pg/ml or less for laser-induced fluorescence detection. Clearly, with respect to sensitivity, speed, and versatility, CE offers significant advantages over gel electrophoresis for the separation of nucleic acids.
to data aquisition
detector
temperature control region
(–)
high voltage
capillary column
(+)
anode
cathode buffer and sample
Capillary Electrophoresis of DNA
buffer
Figure 10.9.1 Schematic of a CE instrument configured for DNA separations.
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Table 10.9.1
Capillary Electrophoresis Systems
Manufacturer Model numbera
Autosampler
Detection methodb
Column temperature
Agilent
CE System
Diode array UV-vis
15° to 60°C
Amersham
MegaBACE 500 16, 32 or 48-capillary DNA sequencer
48-position, 10° to 40°C 96-well plate, no cooling
4-color scanning LIF detection
27° to 44°C
MegaBACE 1000 96capillary DNA sequencer
Single 96-well plate, no cooling
4-color scanning LIF detection
27° to 44°C
MegaBACE 4000 384-capillary DNA sequencer Prism 310
Single 384-well plate, no cooling 482- or 96-sample position, cooled 96-well
4-color scanning LIF detection 4-color LIF with CCD
27° to 44°C
Prism 3100 16-capillary DNA sequencer
96-well plate with sample CCD cooling
26° to 65°C
Prism 3730 96-capillary DNA sequencer P/ACE MDQ DNA System
96-well plates with autoloader Up to 96-well plate
CCD
18° to 70°C
UV and diode array detection
15° to 60°C
CEQ 8000 8-capillary DNA sequencer
96-well plate with cooling 4-color CCD and heating to 90°C
Applied Biosystems
Beckman Coulter
30° to 60°C
30° to 60°C
aAll units listed have a single capillary unless otherwise noted. bLIF, laser-induced fluorescence detector, used with either a PMT (photomultiplier tube) or a CCD (charge-coupled device) camera for visualization; UV, ultraviolet detector.
As stated previously, in its simplest form capillary electrophoretic separation can be achieved by passing a current between cathodic and anodic buffer reservoirs via a liquid-filled glass capillary. In practice, the basic CE instrument also requires a suitable sample injection module, a detector, adequate temperature control, and isolation for the user from the high voltages used for the separations. A schematic of the basic instrument is shown in Figure 10.9.1. There have been a number of changes recently in the features of commercially available instruments; these instruments and their capabilities are summarized in Table 10.9.1. SEPARATION THEORY CE is part of the family of electrophoretic techniques that separate species based upon their size and ionic properties. An ion (i) placed in an electric field will move in the direction parallel to the field towards the oppositely charged electrode with a velocity (vi) defined as follows: vi = µiE = µiV/L where µi is the mobility of the ion, E is the electric field in volts per centimeter, V is the voltage across the column, and L is the total column length. The electrophoretic mobility of a given ion is equal to: mi = qi/6πηai where qi is the charge on the ion, η is the viscosity of the buffer or gel matrix, and ai is the radius of the ion.
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Since DNA has a constant size-to-charge ratio, a sieving matrix must be added to the capillary in order to discriminate based only on size, rather than charge and size. In CE there are two types of gels employed in DNA separations: cross-linked gels (static gels) and non-cross-linked gels (flowable polymers or polymer networks). Cross-linked gels are fixed gels that are polymerized inside the capillary, usually covalently bound to the capillary surface, and are not removed from the capillary between runs. Flowable polymers are viscous hydrophilic polymer solutions that can be pumped into the capillary. The same flowable polymer matrix can be used repeatedly when small molecules such as synthetic oligonucleotides are being analyzed. The time between injections is sufficient for the preceding sample to clear the detector. Alternatively, the polymer can be used once, discarded, and replaced with fresh matrix prior to the next injection. This approach is preferred where larger DNA molecules are present in the samples—e.g., for fragment analysis and DNA sequencing analysis. Usually, a coated capillary is utilized to eliminate the charge effects that are contributed by the native silica surface. With cellulose-derived polymers or some specially modified acrylamides, however, uncoated capillaries may be used, because of the strong interaction of the polymer with the inner surface of the bare fused-silica capillary, in essence forming its own coating. With either a cross-linked or non-cross-linked gel in the capillary, the matrix offers a frictional resistance to the movement of the DNA through the gel medium that is proportional to the size of the species. The frictional resistance can vary with the molecular weight, concentration, and chemical composition of the flowable gel polymer or the pore size in the cross-linked gel, and must be optimized for the particular size of the DNA to be separated. A detailed description of the theory of DNA motility in entangled polymer solutions can be found in Grossman (1991). STRATEGIC PLANNING The most common approach to the separation of both ssDNA and dsDNA by CE uses a coated capillary and an uncharged sieving matrix. This is very similar to slab gel electrophoresis, but in a silica capillary. The separation matrix, as mentioned, can take the form of a cross-linked polyacrylamide gel or flowable polymer such as hydroxypropyl methyl cellulose (HPMC), hydroxyethylcellulose (HEC), polyethylene oxide (PEO), or non-cross-linked linear polyacrylamide. The cross-linked gel is polymerized directly inside the capillary and can be reused for 30 to 100 separations before losing resolution. The capillary is then discarded, since the polyacrylamide gel cannot be regenerated. The flowable polymer has the advantage that it can be expelled from the capillary by pressure at the end of each electrophoretic separation; fresh matrix is then reloaded into the capillary prior to the next separation. These capillaries have lifetimes of several hundred injections. The eventual loss of the surface coating is the major reason for replacement; another common reason is mechanical breakage. The selection of the appropriate matrix can significantly affect the quality of the separation. Cross-linked polyacrylamide is best used for the separation of synthetic oligonucleotides—both native and modified versions. However, flowable polymers can also be used for oligonucleotide analysis and for the separations of automated sequencing ladders. Where dsDNA fragment analysis is required, only flowable polymers are routinely used. The general rule for matrix selection is that the larger the DNA fragment, the weaker the sieving capabilities of the matrix.
Capillary Electrophoresis of DNA
Separation buffers frequently are variants of Tris/borate/EDTA (TBE) mixtures and are buffered at alkaline pH. Urea is often included in the buffer, as a denaturant, when analyzing ssDNA (e.g., synthetic oligonucleotides). Samples are loaded onto the capillary by electrokinetic, or pressure, injection. Separation times range from 10 to 45 min, at
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voltages between 1 and 10 kV. DNA fragments are detected in the UV spectrum at 260 nm, either in the presence or absence of ethidium bromide. Sensitivity can be increased by at least two orders of magnitude through the use of fluorescence detection. Since DNA possesses no native fluorescence, intercalating dyes such as cyanine derivatives (Zhu et al., 1994) or rhodamine derivatives must be added to the electrophoresis buffer, or covalently attached to the DNA prior to the electrophoretic separation. In addition to increasing the sensitivity of detection, these intercalating dyes can improve resolution and sharpening of the bands by physically disrupting the DNA structure. The selection of specific dyes is dictated by their excitation and emission spectra and the compatibility with the detection systems of individual instruments. Specific examples of intercalating dyes are: thiazole orange (Aldrich), YO-PRO-1, YOYO-1, and Sybr Green (Molecular Probes). The added sensitivity is particularly useful when analyzing PCR products that have been amplified from biological fluids. Targets of interest are frequently present in small amounts, and the presence of salts and proteins make their direct analysis by CE impractical. However, after completion of the PCR reaction, the sample can be diluted with water and an aliquot analyzed using fluorescent detection. The mobility of a given DNA fragment may not be constant over a series of injections. This variability can have a variety of causes: aging of the polymer (polyacrylamide), loss of capillary coating, or depletion of the conductivity of the running buffers. The absolute mobility of DNA in a given sample will be dependent upon the salt content (and hence the conductance) of that sample. The presence of high salt will significantly reduce the electrophoretic mobility of the DNA. One solution is to dilute the sample in water and load for longer times; alternatively, the sample can be desalted (UNIT 10.7) prior to injection. Where accurate sizing is important, it is essential to incorporate sizing standards into the sample prior to electrophoresis. The CE analysis of synthetic oligonucleotides requires the selection of a matrix that optimizes resolution of low-molecular-weight oligonucleotides. The separation of fluorescently labeled fragments from an automated sequencing ladder represents a specialized CE application and requires the selection of a matrix with a greater resolution range. These ladders range in size from 20 to more than 1000 bases and can be separated with single-base resolution to a high degree of accuracy. Currently, three automated CE instruments are commercially available as DNA sequencers (see Table 10.9.1). A detailed discussion of the general principles associated with dideoxy sequencing can be found in Ausubel et al. (2003). SEPARATION OF OLIGONUCLEOTIDES In this protocol synthetic oligonucleotides are analyzed for purity by CE using a replaceable, flowable polymer as the separation matrix. The running buffer and separation matrix contain 7 M urea to keep the DNA in its single-stranded configuration. The sample is loaded onto the capillary at the cathode by electrochemical injection. After loading, the sample vial is replaced by the cathode buffer reservoir, and the electrophoresis is continued. The matrix does not need to be replaced between each separation, but should be replaced at the beginning of each series of separations, e.g., at the beginning of each day. Each time fresh matrix is loaded, the capillary must be equilibrated before samples are run. No further equilibration is required between samples. The electrophoretic separation should provide single-base resolution for DNAs of at least 100 bases. A poly(A)40-60 size ladder (see Fig. 10.9.2) should be analyzed initially in order to confirm that resolution is optimal. NOTE: The filled capillary can be stored on the instrument overnight, but if >1 day elapses between runs, the capillary should be stored at 4°C with both ends capped.
BASIC PROTOCOL 1
Purification and Analysis
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NOTE: The following protocol demonstrates the use of a P/ACE 5510 CE instrument from Beckman Coulter. However, other instruments (Table 10.9.1) are capable of comparable separations when operated in accordance with manufacturers’ instructions. NOTE: The selection of a matrix is often instrument-dependent. It is recommended that a kit be used initially for a reference separation. Linear polyacrylamide is frequently used for oligonucleotide separations. Materials ssDNA 100-R separation kit (Beckman Coulter) including: 60-cm, 100-µm-i.d. coated capillary ssDNA 100-R separation gel solution Running buffer: Tris-borate electrophoresis buffer (reconstitute and store up to 30 days at 4°C) Poly(A)40-60 sizing standard (dissolve at 100 µg/ml [3 OD260 units/ml] in water and store indefinitely at −20°C) Dried ssDNA oligonucleotide sample CE instrument (e.g., Beckman Coulter P/ACE 5510 or equivalent; see Table 10.9.1) 1. Reverse standard polarity of the CE instrument electrodes (see manufacturer’s instructions). 2. Rinse capillary on-instrument with deionized water for 5 min. 3. Fill the capillary with ssDNA 100-R gel solution using a 20-min pressure rinse from the matrix vial (based on a 20-psi rinse pressure). This solution can be stored on-instrument for 5 days, but should then be discarded.
0.006
A 260
0.004
0.002
0.000
–0.001 20
24
28
32
36
40
Time (min)
Capillary Electrophoresis of DNA
Figure 10.9.2 CE separation of a standard poly(A)40-60 mixture of synthetic oligonucleotides.
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4. Equilibrate the capillary in running buffer by running a voltage ramp from 0 to 8.1 kV over 20 min and holding at 8.1 kV for 10 min. 5. Replace the inlet reservoir with a container of water and inject for 1 sec at 7.5 kV. 6. Position the sizing standard vial at the inlet and inject for 10 sec at 7.5 kV. 7. Replace with running buffer reservoir and carry out the electrophoresis at 8.1 kV for 40 min at 30°C. 8. At completion of run, confirm that the separation of the standards is satisfactory by comparison with the example provided in the kit. 9. Prepare samples to be analyzed by dissolving in water to ∼10 µg/ml. 10. Load samples onto autosampler and inject for 10 sec at 7.5 kV. 11. Carry out the electrophoresis at 8.1 kV for 40 min at 30°C. 12. Repeat steps 10 and 11 until resolution begins to deteriorate (at least 15 runs); then replace separation gel by returning to step 2. QUANTITATIVE PCR ANALYSIS Quantitative PCR can be used in conjunction with CE separation to amplify and quantitate any DNA target sequence—by the use of either an intercalating dye (coinjected with the samples) or a covalently modified, fluorescently labeled oligonucleotide primer. The size of the expected product is determined by coinjection of sizing standards. Quantitation is achieved by the coamplification of a second target sequence of known concentration, or by the addition of a known quantity of DNA to each sample.
BASIC PROTOCOL 2
Another application of this method is the direct measurement of viral load by reverse transcription (RT)-PCR of the viral RNA. This is achieved by the procedure known as competitive PCR analysis (Piatak, 1993). A known amount of a standard DNA template is included in the reaction mixture to compete for amplification with the target DNA. The sequence of the competing DNA is designed such that the PCR product is similar, but not identical, in size to the target DNA. The small quantities of DNA that are produced by this process require coinjection of an intercalating dye (e.g., YO-PRO-1 or Sybr Green I) as well as fluorescence detection. Since dsDNA is being analyzed in these applications, it is not necessary to include denaturant in the electrophoresis buffer. Materials LIFluor dsDNA 1000 kit (Beckman Coulter) containing: Gel buffer mixture (containing separating gel and Tris/borate/EDTA buffer) EnhanCE intercalating dye 65-cm, 100-µm-i.d. coated capillary Standard sizing ladder: HaeIII restriction digest of φX-174 DNA (dissolve at 10 µg/ml in deionized water and store at −20°C) PCR reaction mixes containing amplicon CE instrument with fluorescent detection (e.g., Beckman Coulter P/ACE 5510 or equivalent) 1. Prepare gel buffer mixture according to manufacturer’s instructions and add 0.4 µg/ml intercalating dye. Store up to 30 days at 4°C.
2. Reverse standard polarity of CE instrument electrodes (see manufacturer’s instructions). Purification and Analysis
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3. If the capillary is new, rinse with gel buffer for 10 min at high (20 psi) pressure. This step is not necessary between runs.
4. Place standard sizing ladder in inlet position and load for 10 sec at low (0.5 psi) pressure. See Figure 10.9.3 for a chromatogram of this standard. The buffer ions become depleted over a series of injections. Consequently the inlet gel reservoir should be replaced after 30 injections.
5. Perform electrophoresis at 9.4 kV for 30 min at 25°C. 6. At the completion of the separation, replace the gel matrix using a 3-min high-pressure wash. 7. Assess the resolution and the linearity of area quantitation of the electrophoretic separation, and compare with the expected profile. 8. Dilute the PCR reaction mixes 10-fold with water and analyze by repeating steps 4 to 6, as with the sizing ladder. Some applications may require the coinjection of the sizing ladder with the sample. In these instances the sizing ladder is loaded for 10 sec, followed by the sample for 10 sec.
64 1363
48
Fluorescence
1078
32
872
503
16 281
271
310 194 234
72
118
0 10
Capillary Electrophoresis of DNA
12
14
16 Time (min)
18
20
Figure 10.9.3 A sizing ladder from a HaeIII digest of φX-174 DNA. Fragment size in bp is indicated.
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COMMENTARY Background Information The application of capillary electrophoresis to the separation of DNA fragments is a relatively recent development. The inherent advantages of this technology—high resolution, excellent sensitivity, and rapid separation times— provide significant improvements over the conventional slab gel electrophoresis technology. This has led to rapid acceptance of CE as an essential tool for the analysis of DNA fragments of all sizes. When applied to the analysis of synthetic oligonucleotides (see Basic Protocol 1), the technique’s ability to obtain single-base resolution has proven extremely useful in diagnosing synthesis problems and in obtaining overall purity estimates for the final products (Cohen et al., 1988). The chemistry of DNA synthesis involves sequential addition of bases to the growing oligonucleotide chain in a prescribed order (APPENDIX 3C). After each addition, the growing chain is “capped” in order to terminate the portion of the oligonucleotide that has not completed the coupling reaction. This yields a truncation series. If present in sufficient quantity, these species will compete with the full-
length oligonucleotide in some applications. Since they are shorter than the full-length oligonucleotide they can readily be resolved and quantitated by CE. After synthesis has been completed, the bases have to be deprotected. If deprotection is incomplete it can interfere with the base-pairing properties of the oligonucleotide. These species, which appear larger than the fulllength oligonucleotide, can also be readily resolved by CE. Figure 10.9.4 illustrates a crude oligonucleotide containing coupling failures (shorter migration times) and incompletely deprotected species (longer migration times). The resolution capacity of CE using a matrix optimized for oligonucleotides can extend to the analysis of oligonucleotides in excess of 100 bases. An example of the separation of a standard oligonucleotide mixture ranging from 40 to 60 bases is presented in Figure 10.9.2. Single-base resolution is readily observed. An increasing number of oligonucleotide applications require modification of the basic oligonucleotide probe—e.g., biotinylation, fluorescent dye modification, phosphorylation, b ase mo dification, addition of phos-
A 260
0.100
0.050
0.000
–0.020 0
10
20
30
40
Time (min)
Figure 10.9.4 CE separation of a synthetic 20-mer oligonucleotide. The coupling failure products (shorter migration times) and incomplete deprotection products (longer migration times) are clearly visible.
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phorothioate (antisense) backbones, DNARNA hybridization, and 5′-amino- or 5′-thiolmodification—prior to conjugation to other molecular species. In all instances, the extent of modification to produce these unique oligonucleotides can be readily assessed by CE due to its high resolution. Recent developments in the area of quantitative gene expression measurements (Freeman et al., 1999) and single nucleotide polymorphism (SNP) detection have utilized the principle of fluorescence energy transfer via oligonucleotide probes. Assays based on quantitative RT-PCR have been developed—e.g., TaqMan (Applied Biosystems), Invader (Third Wave), and Black Hole Quenchers (BioSearch Technologies)—that require either paired single-labeled or dual-labeled oligonucleotides. These reagents are readily analyzed for purity by CE using Basic Protocol 1. CE is particularly useful in this instance since MALDI mass spectrometry (UNIT 10.1), a commonly used alternative, fragments the fluorescent label, resulting in multiple molecular species. Small interfering RNAs (siRNA) can be used for the down-regulation of individual genes (Elbashir et al., 2001). These molecules all share the same features: 2 thymidine deoxynucleotides at the 3′ end followed by 19 ribonucleotides, terminating in a 5′-hydroxyl. These siRNA probes are readily characterized for purity by CE using Basic Protocol 1. The development of matrices to extend this single-base resolution to ssDNA that ranges in size from 20 to 1000 bases has allowed for development of important applications in the field of automated fluorescence-based DNA sequencing. The most commonly performed
sequencing chemistry is the “dye terminator” chemistry, in which the sequence-terminating dideoxy nucleotide also contains the fluorescent reporter group. Consequently, the sequence ladder is labeled in the 3′−hydroxyl position. This means that sequencing reactions can be performed using a primer of any sequence. The alternate “dye primer” chemistry employs a primer that is labeled at the 5′-hydroxyl and is restricted to a small number of commonly used vector sequences—e.g., M13 forward and reverse, T3, T7, and SP6 sequences. An example of this type of application is shown in Figure 10.9.5. Significant effort is being directed towards the development of matrix formulations that will further extend the length of the current sequence reads. Commercial CE sequencing instruments all have shorter run times than slab gel–based automated sequencers and retain comparable sensitivity and accuracy. Each of these applications requires the analysis of ssDNA and is performed under dissociating conditions in the presence of a flowable matrix. The sizing of larger fragments that have been amplified from genomic DNA has proven to be a very effective method for studying genetic variability in populations. These fragments can be analyzed by capillary gel electrophoresis either under dissociating conditions (short fragments, high size accuracy) or as dsDNA (large fragments, lower resolution). Genomic DNA from eukaryotes contains a large number of tandem-repeating sequences that vary in size from 2 to several dozen bp in size. This polymorphism can be used to advantage when studying human identity or individual heredity. The smaller repeats, commonly referred to as
CCACAGAATCAGGGGATAACGCAGGAAAGAACATGTGAGCAAAA 430 440 450 460
Capillary Electrophoresis of DNA
Figure 10.9.5 A portion of the chromatographic output from a PE Biosystems model 373A DNA sequencer using dye terminator sequencing chemistry. Although shown in black and white here, the direct output is color-coded to more clearly illustrate the peaks corresponding to each base in the sequence.
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microsatellite or short tandem repeats (STR), are best analyzed as ssDNA (Butler et al., 1994). To determine identity or heredity it is necessary to accurately determine the number of copies of (2- to 5-bp-long) identical sequence repeats in the selected fragment. Fragment lengths are usually <300 bp and require single-nucleotide resolution. Larger repeats are commonly referred to as variable number of tandem repeats (VNTR) and produce larger-sized fragments that are analyzed as dsDNA. The sizes of the repeats are 10 to 12 nucleotides and up, and fragment sizes can exceed 4000 bp. The required resolution for this type of separation is on the order of 4%. In contrast to the accurate sizing of DNA, fragment pattern matching can also be a very useful analytical tool. The loss or gain of a restriction site between individuals is sufficiently common that restriction fragment length polymorphism (RFLP) analysis can demonstrate individual identity to a high degree of certainty (Ulfelder et al., 1992). These patterns are readily analyzed by CE. RFLP has found application in the fields of forensic medicine and pedigree testing. Where a polymorphic mutation produces a detectable phenotype, it can also be used as a diagnostic for inherited diseases. It requires the separation of dsDNA in the presence of an intercalating dye for visualization. The matrix is generally a flowable polymer, and fragment lengths are from hundreds to thousands of bp in size. Mutation detection can also be performed using ssDNA. A denatured DNA fragment can adopt a sequence-specific conformation upon refolding, which will affect its electrophoretic mobility. This property is taken advantage of in single-stranded conformational polymorphism (SSCP) separations (Ren et al., 1997). The DNA region of interest is PCR-amplified from genomic or cDNA to give sufficient copies of a small (<300-bp) fragment. The strands are separated and allowed to refold. The wild-type and mutant fragments adopt different conformations and are resolved with single-base-pair resolution by CE using a nondissociating medium and a flowable polymer matrix. CE can accurately quantitate and size DNA fragments (Rossomando et al., 1991; Fasco et al., 1995). This can be of considerable value when used to quantitate levels of viruses and pathogenic bacteria. The difficulty arises when quantitation is attempted in the presence of high levels of other DNA, RNA, and proteins. Fluorescence detection coupled with PCR—which
can amplify very low levels of DNA in a highly specific manner—can be used to surmount this problem. This amplification can be combined with the inherent sensitivity of CE through the incorporation of fluorescently labeled primers into the amplified DNA. In this fashion samples such as blood can be analyzed at high dilution, thereby reducing the levels of interfering substances to manageable levels. RNA can be amplified by RT-PCR to give the dsDNA fragment that is subsequently quantitated. This is ideal for measuring low levels of viral messenger RNA, i.e., viral load. The fragments are <1000 bp and separations are usually performed under nondenaturing conditions. The use of appropriate quantitative calibration standards is essential. One such standard, a sizing ladder, is shown in Figure 10.9.3, and an example of this technique is presented in Basic Protocol 2. A majority of CE separations are performed at the standard alkaline pH in the presence of borate, which is an effective buffer in this pH range. The buffer may also contain 6 to 8 M urea, a denaturant that keeps the DNA in its simple single-stranded conformation when required. The urea is omitted from the buffer for analyses where secondary structure plays an important role in the separation, e.g., single-nucleotide polymorphisms or conformational polymorphisms. The type of matrix that is selected for the actual separation can dramatically influence the quality of the separation that is achieved for a given application. Instrument manufacturers frequently supply kits that have been optimized for a particular application (see Table 10.9.2). These can be very helpful as a first, and maybe the only step required for optimizing individual applications. In general, cross-linked polyacrylamide that is polymerized inside, and covalently attached to, the capillary is best suited for smaller-fragment separations. However, since the column is reused multiple times, many things can reduce resolution—for example, capillary plugging, bubble formation, or drying of the capillary end—and require that the capillary be discarded. In the absence of such external parameters, the lifetime of the capillary is ultimately dependent on the breakdown of the polyacrylamide (or other hydrophilic polymer) matrix. Flowable polymers have the advantage of wide fragment-separation ranges. These polymers can have a variety of chemical origins (see Table 10.9.2) and have the advantage of being replaced after each separation. This is achieved by applying pressure to the inlet end of the
Purification and Analysis
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Table 10.9.2
Supplies and Kits for CE Systems
Manufacturer
Part number
Supplies and kits
Agilent
192-1311
Amersham
25-6001-0
Applied Biosystems
402838
µPAGE-10 (10% T, 0% C) 100-µm i.d. µPAGE-5 (5% T, 5% C) 75-µm i.d. µPAGE-3 (3% T, 3% C) 75-µm i.d. MegaBACE SNuPe genotyping application kit POP-4
4316355
POP-4
4313087
POP-5
402844
POP-6
4316357
POP-6
477480 477407
eCAP ssDNA 100-R kit, including gel, caps, and standard eCAP dsDNA 1000 kit
477486
eCAP dsDNA 20,000 kit
477409
EnhanCE Dye
192-5211 192-3211
Beckman Coulter
Capillary Electrophoresis of DNA
capillary. However, some of these polymers are quite viscous and require considerable pressure within the instrument (up to 1600 psi) to load the capillary. Linear polyacrylamide, which contains no crosslinker, is a flowable polymer that can produce a very effective sieving matrix. The sieving properties are dependent upon both the polymer concentration and the average chain length (molecular weight). In general, lower polymer concentrations of higher average molecular weight are preferred for the separation of high molecular weight DNA, the reverse being true for lower molecular weight DNA. The selection of an optimal combination of these conditions must be balanced by viscosities that are compatible with the pumping capabilities of the CE instrumentation. It is essential to avoid purchasing incompatible
Application information
Single-base primer extension for SNP analysis Gel for microsatellite, SNP, differential display, AFLP, and other genotyping applications using capillary #402839 and buffer #402824 on the 310 platform Gel for microsatellite, SNP, differential display, AFLP, and other genotyping applications using capillary array #4315930 and buffer #402824 on the 3100 platform Gel for high-throughput DNA sequencing on the 3730 platform Sequencing applications on the 310 platform, including template suppression reagent (TSR) for 67-cm capillary #402840 and buffer #402824 Sequencing applications on the 3100 platform with 50-cm capillary #4315930 and buffer #402824 Oligonucleotides, RNA, and antisense DNA from 10 to 100 bases Analysis of dsDNA fragments from 72 to 1000 bp Analysis of dsDNA fragments from 1,000 to 20,000 bp Intercalating dye for LIF applications
components by matching the instrument capabilities to the matrix that has been selected for the given application. With bare fused silica, either cellulose-based gels (e.g., hydroxyethylcellulose; Aldrich) or acrylamide-based gels (PE Biosystems) can be used. Replacing coated or gel-filled capillaries can be quite expensive; as premature failure of the capillary can normally be attributed to excessive voltage during separation or to inappropriate storage, it is worthwhile to take precautions for avoiding these problems. All manufacturers of CE instrumentation offer UV absorbance detection systems that will work for most general applications. Beckman Coulter and Bio-Rad offer laser-based fluorescence detectors for highsensitivity CE applications. PE Biosystems offers a single-capillary laser fluorescence instru-
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ment for genetic analysis and DNA sequencing, while Beckman Coulter offers similar instrumentation in an eight-capillary format.
Critical Parameters Oligonucleotide purity The most common size of synthetic oligonucleotide probes is in the range of 20 to 30 bases. However, some applications require synthetic probes that are 100 bases or more in length. Single-base resolution over this range is essential in order to obtain an accurate assessment of purity. The salt content of the oligonucleotide should be kept to a minimum (<50 mM) in order to obtain optimal resolution. The oligonucleotide should be dissolved in water, or serially diluted in water from the stock TBE solution, prior to electrophoresis. Concentrations should be in the 1 µg/20 ml range for UV absorbance detection. The presence of alkaline-pH buffers and urea in the separation matrix and electrophoresis buffer are essential for single-stranded separations. It is important to run a standard— poly(A)40-60—at the beginning of each set of analyses in order to confirm that the electrophoretic resolution is optimal (see Fig. 10.9.2). DNA sequencing In addition to the items above, this application requires single-base resolution over the range of 20 to 1000 bases. Automated, DNA sequencing instruments rigidly control the separation conditions to minimize temperature and power fluctuations and to eliminate variability in the matrix loading protocols. Consequently, most of the critical parameters are associated with template and primer quality and quantity issues. High-quality, salt-free plasmid, cosmid, or PCR-derived DNA is essential. The optimal molar ratio of template to primer is 1:1 and an imbalance of either component outside a ratio of 4:1 will give unusable sequences. Template concentrations should be in the 1 µg/20 µl range. The presence of proteins, salts, detergents, etc. in the template can inhibit the DNA polymerase and kill the reaction. It is essential to remove the excess fluorescent primers or dideoxynucleotides before separating the sequencing ladder on the CE. Further details on the essential parameters of the sequencing reaction itself can be found in Ausubel et al. (2003). Fragment sizing Selection of an appropriate separation matrix is probably the most important issue when
analyzing microsatellite repeats. Resolution of the smaller fragments at the one to two nucleotide level is required. The DNA should be single-stranded and separated with denaturants. The use of internal size standards is essential since the absolute mobilities can change from run to run and from sample to sample. When analyzing the VNTR fragments, which have larger repeating units, base pair resolution in the 3% to 6% range is required. These fragments are analyzed as double-stranded DNA without denaturants. Since the fragments are longer, lower matrix concentrations are preferred. Again, internal standards are essential. The fragments are either obtained directly from genomic DNA, or PCR-amplified DNA. Considerable care has to be taken to remove particulate matter and salts from these samples prior to CE separation. The internal diameter of most capillaries is only 50 to 100 µm, and they are sensitive to plugging. Sometimes plugged capillaries can be salvaged by highpressure back-flushing, but replacement is usually necessary. Salt concentrations should be <50 mM to minimize sample loading problems. RFLP mutation screening The same criteria for VNTR analysis are applied to mutation screening—the major exception being that the application requires pattern matching between control and mutant DNA, rather than absolute sizing. The doublestranded fragments must assume their sequence-dependent conformations. The addition of 10% glycerol to the flowable polymer significantly improves the discrimination of the electrophoretic separation. Quantitation (RT-PCR) The readout from this application is the absolute level of viral RNA in a biological sample. The PCR amplification is specific, but the target is present in a mixture of other nucleic acids, proteins and salts. The use of a fluorescent label or the addition of an intercalating dye, such as YO-PRO-1 or Sybr Green I, as well as the ability to dilute out the interfering substances, are essential features of this application. The fragments are separated as doublestranded PCR products and are quantitated by peak height on the electropherogram. The presence of salts in the starting material and in the PCR reaction can cause variable sample loading if electrokinetic sample injection is used. Sample loading by pressure injection bypasses these problems.
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Absolute quantitation of the PCR product has to take into account both the length of the DNA and the degree of incorporation of the intercalating dye. This is achieved by coinjecting a standard of known concentration with the sample. If this standard contains a ladder of fragments of known length then the quantity of the amplicon can be accurately determined. This procedure carries the added advantage of confirming the correct fragment size. The difficulties encountered with this type of quantitation are not associated with the CE quantitation but with the PCR amplification step. It is essential that a dilution series is performed on the PCR mix to ensure that the reaction is linear with respect to cycle number. In the case of competitive PCR, it is essential that the competing template concentration approximates that of the target template. Alternatively, expression levels can be quantitated relative to a “housekeeping” gene. This is a gene, such as β-actin, that is constitutively expressed in the cells of interest. In this procedure the β-actin gene is coamplified with the gene of interest prior to quantitation by CE. As indicated previously, this field has recently undergone major expansion. The introduction of automated PCR-based instrumentation and the availability of high-quality fluorescently labeled oligonucleotides has produced a number of robust RT-PCR-based kits. They have superseded the intercalating dye approach for some applications.
Troubleshooting
Capillary Electrophoresis of DNA
Mechanical and electrical problems that might be encountered during instrument use are addressed in the troubleshooting sections of the manufacturers’ manuals. One difficulty is the loss of electrical continuity during a run. This loss of current can be caused if a small bubble forms in the column during injection or if the power generated by the run is so great that the solution boils or outgasses. Purging the column after a failed run removes any bubbles in the column. The vial containing the sample must have sufficient liquid to cover the end of the column during injection to prevent introduction of air into the column. On the autosampler, the vials can dry out during a long run, so sample temperature control or the use of the correct cap on the vial is required to slow evaporation. Reducing the buffer concentration or the run voltage can eliminate bubble formation during a run. Degassing the buffer is also useful if the outgassing problems continue.
On occasion capillaries can become plugged. This is normally due to the capillary matrix being allowed to dry out. When not in use, the capillary can remain installed on the CE instrument, but both ends must remain submerged in buffer. If it is removed from the instrument the capillary should be stored refrigerated, with both ends capped to prevent evaporation. The capillary can also become plugged from insoluble material that is present in the sample. Sometimes it is possible to apply pressure to the anode outlet and blow out the plug. More likely, however, the capillary will need to be replaced. Loss of resolution on a separation of standards can have several causes: e.g., the buffer, the separation matrix, or the capillary. A process of elimination should follow the course of first trying fresh buffer, then new matrix, and finally changing out the capillary. Where separation of standards is normal, but resolution or signal strength is poor, the cause is likely excess salts and/or buffers in the samples. Examining the behavior of the electrical current during the run will probably assist in troubleshooting. Most CE instruments simultaneously monitor both detector output and current flow throughout the electrophoresis run. A drop in sensitivity or signal strength can be observed when multiple injections are made from a single standard vial (especially if the standard is dissolved in deionized water). This phenomena can be due to salt contamination of the sample by residual matrix buffer on the outside of the capillary (Guttman and Schwartz, 1995). Keep in mind that any source of additional ions to the sample (e.g., salt, buffer, or other ion species) will affect an electrokinetic injection. The ions of the sample will be effectively diluted by the ions from the contaminant. Sample preparation to remove variable salt contamination will improve the consistency of injections (Ruiz-Martinez et al., 1998).
Anticipated Results The separation of oligonucleotides should show single-base resolution from 5 bases to >100 bases. The synthetic efficiency of the assembly process is in excess of 99% (Pon et al., 1996), such that the yield of full-length product for a 100-mer should still exceed 30% of the total DNA present. For a 25-mer the yield of full-length product should exceed 80%. The addition of various functional moieties to the oligonucleotide will normally be seen as an increase in size (or decrease in mobility). The
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addition of a group as small as a phosphate group at the 5′-hydroxyl will make the oligonucleotide appear longer by a single base. The size increase, however, is insufficient proof of successful modification. This must be verified by obtaining a molecular weight by mass spectrometry (e.g., MALDI-TOF; UNIT 10.1). The DNA-sequencing application (as stated previously) can have single-base resolution up to 1000 bases or more, though performance today on commercial multicapillary instruments is ∼600 bases or more in 2 hr. The smallest sequencing fragments will be detected as sharp, tight bands usually ∼1 to 3 sec in width. The larger (slower-moving) fragments will show considerable band broadening due to diffusion as well as reduced separation efficiency of the matrix. The microsatellite and VNTR analyses will show well-resolved peaks for the various multimers that are reflective of the number of repeats in the sample. These should be well resolved from the sizing control that has been added to, or coinjected with, the sample of interest. The variability of these repeats at a given locus should give sufficient information to clearly identify relatedness between individuals. The expression levels of various isotypes should be sufficiently well discriminated to use this technique as a potential diagnostic test for disease states. Quantitative PCR data should be sufficiently accurate to detect a 3- to 5-fold change in viral load level. The linearity of response of the standard and target DNAs should extend to the low to sub-nanogram level.
Time Considerations These will vary dependent upon the type of separations that are required. Preparation of the stock solutions can take up to 1 hr. Filling the capillaries with fresh gel can take 1 hr. The analysis of synthetic oligonucleotides is normally completed within 30 min and sample preparation time is minimal. The DNA sequencing application probably requires the longest separation times. A 2-hr cycle time from injection to injection is normally sufficient to obtain 500 to 600 bases of high-quality sequence. Regeneration and loading times are <15 min. Preparing the stock solutions from the kit takes 24 hr. Filling the capillary with gel takes 30 min. Analysis of microsatellite fragments can take as little as 15 min, whereas the analysis of VNTR fragments takes up to 30 min. Regeneration times are <10 min. The analysis
component of quantitative PCR is similar. However, this application requires multiple runs in order to obtain linearity curves for the PCR products and the calibration standards.
Literature Cited Ausubel, F.M., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (eds.) 2003. Current Protocols in Molecular Biology. John Wiley & Sons, Hoboken, N.J. Butler, J., McCord, B., Jung, J., and Allen, R. 1994. Rap id analysis of short tandem repeat HUMTH01 by capillary electrophoresis. BioTechniques 10:1062-1068. Cohen, A., Najarian, D., Paulus, A., Guttman, A., Smith, J., and Karger, B. 1988. Rapid separation and purification of oligonucleotides by high-performance, capillary, gel electrophoresis. Proc. Natl. Acad. Sci. U.S.A. 85:9660-9663. Elbashir, E.M., Harborth, J., Lendeckel, W., Yalkcin, A., Weber, K., and Tuschl, T. 2001. Duplexes of 21-nucleotide RNAs mediated RNA interference in cultured mammalian cells. Nature 411:494498. Fasco, M., Treanor, C., Spivack, S., Figge, H., and Kaminsky, L. 1995. Quantitative RNA-polymerase chain reaction-DNA analysis by capillary electrophoresis and laser induced fluorescence. Anal. Biochem. 224:140-147. Freeman, W.M., Walker, S.J., and Vrana, K.E. 1999. Quantitative RT-PCR: Pitfalls and potential. BioTechniques 26:112-125. Grossman, P.D. 1991. Effect of molecular orientation and entangled polymer additives on the electrophoresis of biopolymers in free solution. Ph.D. Thesis, University of California, Berkeley. Guttman, A. and Schwartz, H. 1995. Artifacts related to sample introduction in capillary gel electrophoresis affecting separation performance and quantitation. Anal. Chem. 67:2279-2283. Jarcho, J. 1994. Restriction fragment length polymorphism analysis. In Current Protocols in Human Genetics (N.C. Dracopoli, J.L. Haines, B.R. Korf, D.T Moir, C.C. Morton, C.E. Seidman, J.G. Seidman, and D.R. Smith, eds.) pp. 2.7.12.7.15. John Wiley & Sons, New York. Piatak, M., Luk, K.-C., Williams, B., and Lifson, J.D. 1993. Quantitative competitive PCR for accurate quantitation of HIV DNA and RNA species. BioTechniques 14:70-81. Pon, R., Buck, G., Hager, K., Naeve, C., Niece, R., Robertson, M., and Smith, A. 1996. Multifacility survey of oligonucleotide synthesis and an examination of the performance of unpurified primers in automated DNA sequencing. BioTechniques 21:680-685. Ren, J., Ulvik, A., Ueland, P.U., and Refsum, H. 1997. Analysis of single-strand conformation polymorphism by capillary electrophoresis with laser-induced fluorescence detection using short-chain polyacrylamide as sieving medium. Anal. Biochem. 245:79-84.
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Rossomando, E., White, L., and Ulfelder, K. 1991. Capillary electrophoresis: Separation and quantitation of reverse transcriptase polymerase chain reaction products from polio virus. J. Chromatogr. B. 656:159-168. Ruiz-Martinez, M., Salas-Solano, O., Carilho, E., Kotler, L., and Karger, B. 1998. A sample purification method for ruffed and high-performance DNA sequencing by capillary electrophoresis using replaceable polymer solutions. A. Development of the cleanup protocol. Anal. Chem. 70:1516-1527. Sozer, A.C., Kelly, C.M., Demers, D.B. 1998. Molecular analysis of paternity. In Current Protocols in Human Genetics (N.C. Dracopoli, J.L. Haines, B.R. Korf, D.T Moir, C.C. Morton, C.E. Seidman, J.G. Seidman, and D.R. Smith, eds.) pp. 14.4.1-14.4.26. John Wiley & Sons, New York. Ulfelder, K., Schwartz, H., Hall, J., and Sunzeri, F. 1992. Restriction fragment length polymorphism analysis of ERBB2 oncogene by capillary electrophoresis. Anal. Biochem. 200:260-267.
Zhu, H., Clark, S., Benson, S., Rye, H., and Glazer, A. 1994. High sensitivity capillary electrophoresis of double stranded DNA fragments using monomeric and dimeric fluorescent intercalating dyes. Anal. Chem. 66:1941-1949.
Key Reference Ulfelder, K.J. and McCord, B. 1997. Chapter 11. Separation of DNA by capillary electrophoresis. In Handbook of Capillary Electrophoresis, 2nd ed. (J.P. Landers, ed.) pp. 347-378. CRC Press, Boca Raton, Fla. An excellent reference on all aspects of capillary electrophoresis separations. Chapter 11, on DNA, goes into much greater depth than is possible here on the theory of separation, selection of buffers, and selection of gel matrices.
Contributed by Alan Smith Stanford University Stanford, California Robert J. Nelson Dakota Scientific Sioux Falls, South Dakota
Capillary Electrophoresis of DNA
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Sequencing Oligonucleotides by Enrichment of Coupling Failures Using Matrix-Assisted Laser Desorption/Ionization Time-of-Flight Mass Spectrometry
UNIT 10.10
This unit describes a procedure for sequencing production-scale synthetic oligonucleotides. Coupling failure sequences in a crude synthesis are isolated from the fulllength 5 -O-DMTr-oligonucleotide using a DMTr-selective C18 purification cartridge (see UNIT 10.7). The extraction is subsequently analyzed by matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrometry to determine the mass difference between failure ions for identification of a particular base or structural modification. The methods presented here are based on MALDI-TOF-MS procedures in UNIT 10.1, which should be consulted for additional details.
EXTRACTION, DESALTING, AND CONCENTRATION OF FAILURE SEQUENCES FROM A CRUDE OLIGONUCLEOTIDE
BASIC PROTOCOL 1
This protocol describes the extraction of coupling failure sequences from the full-length 5 -O-DMTr-oligonucleotide of a crude synthesis. A crude synthesis is first prepared for loading onto a C18 reversed-phase purification cartridge. Coupling failures are then eluted from the full-length oligonucleotide, which binds more strongly to the cartridge due to the hydrophobicity of its attached DMTr protecting group. The eluate is then desalted and concentrated, resulting in a series of failure products readily detectable by MALDI-TOF analysis. Refer also to UNIT 10.7 regarding DMTr-selective cartridge purification and affinity cartridge desalting of detritylated oligonucleotides.
Materials Crude 5 -O-(4,4 -dimethoxytrityl)-protected (DMTr) oligonucleotide (1 to 2 mg from an 80- to 750-µmol scale synthesis), ammoniacal solution 0.1 and 2.0 M triethylammonium acetate (TEAA), pH 7.0, HPLC grade Acetonitrile (MeCN), HPLC grade Ultrapure (e.g., Milli-Q) water 17% or 18% (v/v) MeCN in 0.1 M TEAA, pH 7.0 (see recipe) Vacuum centrifugation system (Speedvac) Temperature-controlled heating block 5-mL disposable plastic syringes with exposed Luer tips C18 reversed-phase oligonucleotide purification cartridges (e.g., Sep-Pak Plus, Waters Corporation) Prepare crude oligonucleotide 1. Evaporate the ammoniacal solution of a crude 5 -O-DMTr-oligonucleotide synthesis to dryness under vacuum using a Speedvac overnight. 2. Add 2 mL of 0.1 M TEAA to the dried oligonucleotide. 3. Heat sample at 95◦ C for 10 min, then vortex into solution. The sample may contain residual material that does not go into solution. Do not overheat, as this can result in removal of the DMTr group from the full-length oligonucleotide.
Contributed by David Alazard and James Russell Current Protocols in Nucleic Acid Chemistry (2005) 10.10.1-10.10.7 C 2005 by John Wiley and Sons, Inc. Copyright
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10.10.1 Supplement 23
Extract failure sequences 4. Using a 5-mL disposable plastic syringe, pass 5 mL MeCN and then 5 mL of 2 M TEAA through a C18 reversed-phase oligonucleotide purification cartridge into a waste container. All reagents should be eluted from the cartridge in a dropwise fashion. Multiple cartridges can optionally be used with a vacuum manifold system (Analytichem or National Scientific; optional) with an in-line trap for waste and a water aspirator pump or small vacuum pump.
5. Slowly pass the dissolved oligonucleotide solution at approximately 1 drop/second through the cartridge to the waste. 6. Pass 10 mL of ultrapure water through the cartridge to the waste. 7. Elute and collect failure sequences with 2 mL of 17% or 18% (v/v) MeCN in 0.1 M TEAA, pH 7.0. Use 17% MeCN in 0.1 M TEAA for shorter sequences (≤35 bases) and 18% MeCN in 0.1 M TEAA for longer sequences (>35 bases).
8. Dilute the eluate with 5 mL of 0.1 M TEAA.
Desalt and concentrate failure sequences 9. Pass 5 mL MeCN and then 5 mL of 2 M TEAA through a new purification cartridge into a waste container. 10. Slowly pass the diluted eluate (containing failure sequences) at approximately 1 drop/second through the new cartridge to the waste. 11. Pass 10 mL ultrapure water through the cartridge to the waste. 12. Elute and collect desalted failure sequences with 1 mL MeCN. 13. Concentrate extracted failure sequences under vacuum using a Speedvac to ∼20 to 50 µL. Do not allow the sample to dry completely. BASIC PROTOCOL 2
PREPARATION OF OLIGONUCLEOTIDE FAILURE SEQUENCES FOR MALDI-TOF ANALYSIS This protocol describes the preparation of extracted failure sequences for MALDI-TOF analysis. Diluted oligonucleotide coupling failure sequences are mixed with ammonium citrate buffer and 3-hydroxypiconilic acid for spotting on a sample plate well. Refer also to UNIT 10.1 for alternative procedures for matrix/co-matrix/analyte preparation.
Materials Concentrated oligonucleotide failure sequences (see Basic Protocol 1) Ultrapure (e.g., Milli-Q) water 50 mg/mL ammonium citrate buffer, pH 9.4 (from Sequazyme Oligonucleotide Sequencing Kit, PerSeptive Biosystems, or see recipe) 50 mg/mL 3-hydroxypiconilic acid (3-HPA) matrix (see recipe) Ammonium-activated cation-exchange resin beads (see UNIT 10.1) Parafilm Sequencing Oligonucleotides Using MALDITOF-MS
1. Dilute concentrated oligonucleotide failure sequences 1:10 with ultrapure water. 2. On a small piece of Parafilm, combine 2 µL of diluted failure sequences, 1 µL of 50 mg/mL ammonium citrate buffer (pH 9.4), and 7 µL of 3-HPA matrix.
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3. Add approximately 0.1 mg of ammonium-activated cation-exchange resin beads (a small spatula tip full) to cover the bottom of the solution droplet. Do not overload the beads, which can completely absorb the droplet.
4. Mix droplet and beads by withdrawing and expelling the solution 10 times with a pipettor. 5. Allow beads to settle for 30 sec and then spot 2 µL of the supernatant on a sample plate well. 6. Allow spot to crystallize at room temperature for approximately 10 min. Do not load the sample plate until the spot has crystallized to dryness.
MALDI-TOF ANALYSIS Experimental Conditions Conditions for MALDI-TOF analysis are based on operation of the linear Voyager-DE mass spectrometer (PerSeptive Biosystems) equipped with a 337-nm laser. Instrument parameters depend in part on sequence length.
BASIC PROTOCOL 3
For shorter oligonucleotides (≤35 bases):
Positive ion mode 25,000 acceleration voltage 92.5% grid voltage 0.15% guide wire voltage 250 nsec delay extraction time. For longer oligonucleotides (>35 bases):
Positive ion mode 25,000 acceleration voltage 91.5% grid voltage 0.15% guide wire voltage 400 nsec delay extraction time. Spectra are obtained using laser intensities ranging from 2300 to 2600 µJ, or just above the threshold of ion detection. Typically, 50 to 100 scans are collected. Smoothing is performed using 19 points and a polynomial order of 2. Instrument parameters may need optimization according to the particular mass spectrometer configuration Table 10.10.1 Characteristic Molecular Masses for Deoxyribonucleotides, 2 -O-Methylribonucleotides, and a Non-Nucleosidic Linkera
Nucleotide
Symbol
Mass (Da)
A
313.2
C
289.2
G
329.2
T
304.2
a
343.3
c
319.2
g
359.2
2 -O-Methyluridine-5 -phosphate
u
320.2
Non-nucleosidic linker
X
266.1
2 -Deoxyadenosine-5 -phosphate
2 -Deoxycytidine-5 -phosphate
2 -Deoxyguanosine-5 -phosphate 2 -Deoxythymidine-5 -phosphate
2 -O-Methyladenosine-5 -phosphate
2 -O-Methylcytidine-5 -phosphate
2 -O-Methylguanosine-5 -phosphate
a Adapted from Alazard et al. (2002) with permission from Elsevier.
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(e.g., continuous extraction or a reflected pathlength). Manufacturer specifications and recommendations from UNIT 10.1 (e.g., negative ion mode) may also influence the sensitivity and resolution.
Interpretation of Mass Spectrometry Data After obtaining the mass of individual oligonucleotide coupling failures, the mass difference between successive spectral ions is calculated. The identification of a particular
Figure 10.10.1 Positive-ion MALDI-TOF mass spectrum of extracted failure sequences from the crude synthesis of a 16-mer (5 gga aXG TCA GGC AAC AT 3 ). Uppercase letters denote deoxyribonucleotides, lowercase letters denote 2 -O-methylribonucleotides, and X denotes the non-nucleosidic linker. See Table 10.10.2 for molecular mass differences obtained for base assignment. Table 10.10.2 Molecular Mass Differences of Enriched Failure Sequences from a 16-mer Composed of Deoxyribonucleotides, 2 -O-Methylribonucleotides, and a Non-Nucleosidic Linker
Observed mass (M + H)+
Molecular mass difference (Da)
Base assignmenta
1166.8
314.4
A
1481.1
314.3
A
1771.4
290.3
C
2101.5
330.1
G
2431.4
329.9
G
2745.3
313.9
A
3035.3
290.0
C
3340.5
305.2
T
3669.3
328.8
G
3936.6
267.3
X
4281.9
341.9
a
4623.8
345.1
a
4981.0
357.2
g
5341.0
360.0
g
852.4
Sequencing Oligonucleotides Using MALDITOF-MS
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a Uppercase letters denote deoxyribonucleotides, lowercase letters denote 2 -O-
methylribonucleotides, and X denotes the non-nucleosidic linker. Current Protocols in Nucleic Acid Chemistry
nucleotide or structural modification is determined based on its characteristic mass (Table 10.10.1). Sequencing information is read from the 5 →3 direction, as demonstrated for a sample 16-mer with a mixed sugar backbone and non-nucleosidic linker (Table 10.10.2 and Fig. 10.10.1).
REAGENTS AND SOLUTIONS Use ultrapure (e.g., Milli-Q-purified) water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Acetonitrile (MeCN) in 0.1 M triethylammonium acetate (TEAA), pH 7.0, 17% or 18% (v/v) Measure 1.7 or 1.8 mL of MeCN in a 15-mL centrifuge tube. Add 0.1 M TEAA, pH 7.0 (see recipe) to a volume of 10 mL and mix. Prepare fresh prior to use and keep at room temperature. Storage for greater than 3 days is not recommended.
Ammonium citrate, pH 9.4, 50 mg/mL Dissolve 5.0 g dibasic ammonium citrate in 100 mL ultrapure water. Adjust pH to 9.4 with ammonium hydroxide. Store up to 1 month at 4◦ C. 3-Hydroxypiconilic acid (3-HPA), 50 mg/mL Weigh 0.05 g of 3-HPA in a 1.5-mL microcentrifuge tube. Add 1 mL ultrapure water, then vortex into solution. Store up to 1 month at 4◦ C. Triethylammonium acetate (TEAA), pH 7.0, 0.1 M Measure 5.0 mL of 2.0 M TEAA, pH 7.0 (commercially available), in a 100-mL storage bottle. Add ultrapure water to a volume of 100 mL and mix. Prepare fresh prior to use and keep at room temperature. Storage for greater than 3 days is not recommended.
COMMENTARY Background Information Since the early 1990s, mass spectrometry has emerged as a successful alternative for the sequence determination of oligonucleotides. Various strategies—such as collision-induced dissociation (McLuckey et al., 1992; Little et al., 1996; Ni et al., 1996), Sanger termination reactions (K¨oster et al., 1996; Roskey et al., 1996; Taranenko et al., 1998), chemical cleavage (Polo et al., 1997; Isola et al., 1999), and enzymatic digestion (Limbach et al., 1994; Wu and Aboleneen, 2000) using electrospray ionization (UNIT 10.2) and MALDI-TOF (UNIT 10.1)—are currently being employed. However, each technique has its own limitation dependent on the interpretation of data, the oligonucleotide length, and structural modifications. The generation of oligonucleotide fragments using exonucleases is the more commonly used approach for MALDI-TOF
sequencing (Pieles et al., 1993; Schuette et al., 1995; Smirnov et al., 1996). With this procedure, “mass ladders” (UNIT 10.1) are created by the partial, sequential hydrolysis of nucleotides using a 5 →3 calf spleen phosphodiesterase (CSP) or a 3 →5 snake venom phosphodiesterase (SVP). The mass difference between successive fragment ions is calculated to identify a particular base. This technique is practical and easy to interpret. However, backbone, sugar, and terminal modifications are resilient to phosphodiesterase digestion. Also CSP and SVP cleave only single-stranded oligonucleotides; regions with a high degree of intra- or intermolecular structure will also resist hydrolysis. Another straightforward approach to MALDI-TOF sequencing utilizes crude synthetic oligonucleotides (Juhasz et al., 1996; Keough et al., 1996; Alazard et al., 2002). Similar to exonuclease digestion, the mass
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difference between spectral ions is calculated, but the coupling failures generated during synthesis are used for the calculation. However, with a highly efficient synthesis, the amount of failure species can fall below the detection limit of MALDI-TOF analysis, especially with the predominant, full-length product still present. Therefore, enrichment of the failure sequences using DMTr-selective C18 purification cartridges is quite beneficial, since it enhances the signal and resolution of failure ions, allowing for an accurate confirmation of an oligonucleotide sequence. This technique is useful for routine quality control testing or as a process verification. It is helpful for sequencing modified oligonucleotides, since it depends on the coupling efficiency of the synthetic process and not structural alterations affecting enzymatic digestion. Modified and non-nucleosidic amidites will, if anything, have a reduced coupling efficiency, thus increasing their failure signal.
Critical Parameters and Troubleshooting
Sequencing Oligonucleotides Using MALDITOF-MS
After the crude ammoniacal solution of an oligonucleotide synthesis is evaporated overnight, 0.1 M TEAA is added and the sample is heated slightly to help dissolve the dried material. Overheating must be avoided since it will cause detritylation, which in turn will cause a large amount of the full-length material to elute with the coupling failures during extraction. The ensuing MALDI-TOF analysis will result in an excess of the full-length signal, suppressing the ions of the desired sequencing failures. Some insoluble material may remain after vortexing. It should be allowed to settle, and only the dissolved solution should be loaded on the cartridge to prevent clogging. The extraction procedure utilizes the increased hydrophobicity of the full-length 5 -ODMTr-protected oligonucleotide. Failure sequences are eluted using 17% or 18% MeCN in 0.1 M TEAA, pH 7.0, depending on their length. MeCN percentages greater than 18% will co-elute the DMTr-on product with the failures. Highly hydrophobic modifications may require a stepwise increase in the MeCN percentage to elute failure species. When concentrating extracted failure sequences under vacuum centrifugation, it is important to prevent the solution from drying completely. A residual concentrate of approximately 20 to 50 µL will avoid difficulty dissolving a dried sample back into solution. A 1:10 dilution of the concentrate with water will improve crystallization with the matrix,
enhancing ionization of the coupling failure sequences during MALDI-TOF analysis. Sequence information is obtained from the 5 end to the beginning of the 3 end. As mass signals approach the smaller 3 sequences (<1000 Da), matrix and doubly or triply charged ions interfere with their detection. Complete sequencing at the 3 end can be obtained using SVP hydrolysis if applicable (UNIT 10.1). Also, adduct peaks can complicate the reading of sequencing failures; desalting with a C18 cartridge and cation-exchange beads will help reduce the formation of these additional ions.
Anticipated Results Following the above protocols, strong mass spectral signals and resolution of failure sequence ions by MALDI-TOF mass spectrometry are obtained. Sequencing information for oligonucleotides (≤60 bases) from the 5 end up to three to four nucleotides at the 3 end is expected. Backbone, sugar, and terminal (e.g., phosphorothioate, 2 -O-methylribose, fluorescent labeling) modifications can also be identified using this procedure.
Time Considerations The evaporation of an ammoniacal solution of a crude oligonucleotide synthesis is best done overnight prior to the extraction procedure and MALDI-TOF analysis, since vacuum centrifugation can take up to 6 to 8 hr depending on the sample volume. The extraction and desalting of the oligonucleotide failure sequences typically requires 1 hr. The concentration of these purified coupling failure sequences to approximately 20 to 50 µL requires ∼2 hr. The sample preparation for MALDI-TOF analysis and data interpretation generally takes 30 to 60 min.
Acknowledgment The authors acknowledge Dr. Huynh Vu of Gen-Probe, Inc. for the synthesis of the crude 16-mer oligonucleotide.
Literature Cited Alazard, D., Filipowsky, M., Raeside, J., Clarke, M., Majlessi, M., Russell, J., and Weisburg, W. 2002. Sequencing of production-scale synthetic oligonucleotides by enriching for coupling failures using matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Anal. Biochem. 301:57-64. Isola, N.R., Allman, S.L., Golovlov, V.V., and Chen, C.H. 1999. Chemical cleavage sequencing of DNA using matrix-assisted laser
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desorption/ionization time-of-flight mass spectrometry. Anal. Chem. 71:2266-2269. Juhasz, P., Roskey, M.T., Smirnov, I.P., Haff, L.A., Vestal, M.L., and Martin, S.A. 1996. Applications of delayed extraction matrix-assisted laser desorption ionization time-of-flight mass spectrometry to oligonucleotide analysis. Anal. Chem. 68:941-946. Keough, T., Shaffer, J.D., Lacey, M.P., Riley, T.A., Marvin, W.B., Scurria, M.A., Hasselfield, J.A., and Hesselberth, E.P. 1996. Detailed characterization of antisense DNA oligonucleotides. Anal. Chem. 68:3405-3412. K¨oster, H., Tang, K., Fu, D.J., Braun, A., van den Boom, D., Smith, C.L., Cotter, R.J., and Cantor, C.R. 1996. A strategy for rapid and efficient DNA sequencing by mass spectrometry. Nat. Biotechnol. 14:1123-1128. Limbach, P.A., McCloskey, J.A., and Crain, P.F. 1994. Enzymatic sequencing of oligonucleotides with electrospray mass spectrometry. Nucleic Acids Symp. Ser. 31:127-128. Little, D.P., Aaserud, D.J., Valaskovic, G.A., and McLafferty, F.W. 1996. Sequence information from 42-108-mer DNAs (complete for a 50-mer) by tandem mass spectrometry. J. Am. Chem. Soc. 118:9352-9359. McLuckey, S.A., Van Berkel, G.J., and Glish, G.L. 1992. Tandem mass spectrometry of small, multiply charged oligonucleotides. J. Am. Soc. Mass Spectrom. 3:60-70. Ni, J., Pomerantz, C., Rozenski, J., Zhang, Y., and McCloskey, J.A. 1996. Interpretation of oligonucleotide mass spectra for determination of sequence using electrospray ionization and tandem mass spectrometry. Anal. Chem. 68:1989-1999. Pieles, U., Z¨urcher, W., Sch¨ar, M., and Moser, H.E. 1993. Matrix-assisted laser desorption ionization time-of-flight mass spectrometry: A powerful tool for the mass and sequence analysis of natural and modified oligonucleotides. Nucl. Acids Res. 21:3191-3196.
Polo, L.M., McCarley, T.D., and Limbach, P.A. 1997. Chemical sequencing of phosphorothioate oligonucleotides using matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Anal. Chem. 69:1107-1112. Roskey, M.T., Juhasz, P., Smirnov, I.P., Takach, E.J., Martin, S.A., and Haff, L.A. 1996. DNA sequencing by delayed extraction-matrix-assisted laser desorption/ionization time of flight mass spectrometry. Proc. Natl. Acad. Sci. U.S.A. 93:4724-4729. Schuette, J.M., Pieles, U., Maleknia, S.D., Srivatsa, G.S., Cole, D.L., Moser, H.E., and Afeyan, N.B. 1995. Sequence analysis of phosphorothioate oligonucleotides via matrix-assisted laser desorption ionization time-of-flight mass spectrometry. J. Pharm. Biomed. Anal. 13:1195-1203. Smirnov, I.P., Roskey, M.T., Juhasz, P., Takach, E.J., Martin, S.A., and Haff, L.A. 1996. Sequencing oligonucleotides by exonuclease digestion and delayed extraction matrix-assisted laser desorption ionization time-of-flight mass spectrometry. Anal. Biochem. 238:19-25. Taranenko, N.I., Allman, S.L., Golovlev, V.V., Taranenko, N.V., Isola, N.R., and Chen, C.H. 1998. Sequencing DNA using mass spectrometry for ladder detection. Nucl. Acids Res. 26:2488-2490. Wu, H. and Aboleneen, H. 2000. Sequencing oligonucleotides with blocked termini using exonuclease digestion and electrospray mass spectrometry. Anal. Biochem. 287:126-135.
Contributed by David Alazard and James Russell Gen-Probe, Inc. San Diego, California
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Mass Determination of Phosphoramidites
UNIT 10.11
Mass spectrometry (MS) is now used as a very common and indispensable tool in the field of nucleic acid chemistry. Nucleoside phosphoramidites are the most widely used building blocks in contemporary solid-phase synthesis of oligonucleotides (UNITS 3.3 & 3.5; Caruthers et al., 1987; Reese, 2002). Although MS has been effective and is extensively used for the characterization of nucleoside phosphoramidites, this approach has been problematic owing to the acid-labile properties of phosphoramidites and the lack of general methods. In view of the importance of nucleoside phosphoramidites and their analogs, this unit provides a convenient, rapid, and reliable method for measuring the molecular weights (MW) of various nucleoside and non-nucleoside phosphoramidites. The key feature of the method is the use of a triethanolamine (TEOA)/NaCl matrix system. A combination of TEOA and NaCl is, to our knowledge, the first effective matrix for analysis of phosphoramidites using soft ionization techniques. TEOA/NaCl can be employed for liquid secondary ion (LSI) or fast-atom bombardment (FAB) ionization (Ashcroft, 1997) with currently available double-focusing mass spectrometers. All accurate MW measurements of nucleoside and non-nucleoside phosphoramidites described in this unit are obtained using either LSIMS or FABMS (Basic Protocol). The accurate molecular weights of various phosphoramidites appeared as adduct ions [M+Na]+ with an average mass error smaller than 0.2 ppm. The Alternate Protocol provides a modified sample/matrix preparation for enhanced molecular-related ions (MRIs).
ACCURATE MASS MEASUREMENT OF PHOSPHORAMIDITES USING TRIETHANOLAMINE/NaCl
BASIC PROTOCOL
Phosphoramidite analysis is routinely performed on a commercial LSIMS or FABMS. There are no special requirements for the standard LSIMS or FABMS procedures or system configurations. Low- or high-resolution LSIMS is performed on a double-focusing magnetic sector mass spectrometer equipped with a cesium ion source. The sample is introduced into the ion source using a direct inlet system. A Cs+ ion primary beam with an energy of 15 keV is used, and the secondary sample ions are accelerated to 6 kV. Low-resolution LSIMS mass spectra are acquired at 8-sec intervals from m/z 0 to 1500 in the positive ion mode, and about ten scans are averaged. The low mass resolution of LSIMS is 2000 to 3000 (10% valley definition). Low- or high-resolution FABMS experiments are carried out on a double-focusing magnetic sector mass spectrometer equipped with a Xe gun. The sample is introduced into the ion source using a direct inlet system, and a fast-atom Xe beam is generated from Xe+ ions that are accelerated to 5 keV. The sample-ion accelerating voltage is 10 kV, and the emission current of the FAB gun is 1 mA. The low mass resolution of FABMS is 2000 to 3000 (10% valley definition). All of the high-resolution MS data are obtained by accelerating voltage scans. Polyethylene glycol (PEG) 600, PEG 1000, and their Na+ adduct ions are used as references to produce accurate masses. Two reference ions, one lower and one higher in mass than the measured sample ion, are selected as calibrators. The two reference peaks and the sample peak are measured by continuous voltage scans even when the accelerating voltage is turned off. As a consequence of the ability to switch reference and sample, it is not necessary for the sample and reference to be ionized at the same time. The postacquisition data processing program automatically groups scans of each of the three
Purification and Analysis of Synthetic Nucleic Acids and Components
Contributed by Shinya Harusawa, Mihoyo Fujitake, Takushi Kurihara, Zheng-yun Zhao, and David M.J. Lilley
10.11.1
Current Protocols in Nucleic Acid Chemistry (2006) 10.11.1-10.11.16 C 2006 by John Wiley & Sons, Inc. Copyright
Supplement 26
ions being analyzed (low reference, high reference, and sample). The high mass resolution of LSIMS is 3000 (10% valley definition), while that of FABMS is 8000 (10% valley definition). The phosphoramidites listed in Tables 10.11.3 and 10.11.4 (see Commentary) were analyzed as described in this protocol. IMPORTANT NOTE: The mass spectrometer must be maintained to its best operational conditions at all times to obtain optimal performance of the two ionization methods. In most cases, sufficient MRIs are obtained using this protocol, although decreases in ion intensities are observed somewhat compared to those of the Alternate Protocol (see Table 10.11.5 entry 7). In this protocol approximate amounts (0.02 to 0.05 µmol) of the phosphoramidites, saline (0.2 to 0.3 µL), and chloroform (0.5 to 1 µL) are used. The ratio of sample, saline, and TEOA corresponds approximately to entry 6 in Table 10.11.5 (see Commentary for discussion).
Materials Phosphoramidite to be analyzed (see Critical Parameters and Troubleshooting) Chloroform (>99% [GC]; Aldrich or TCI) Triethanolamine (TEAO; FABMS and LSIMS grade; TCI) Saline (0.9% NaCl in ultrapure water) 2-µL disposable capillary pipets (e.g., Minicaps; Hirshmann Laborger¨ate) Small glass tube NOTE: Phosphoramidites S.1a-d were synthesized in the laboratory (Araki et al., 2004, 2005). Phosphoramidites in Figures 10.11.4-10.11.6 are available from Cruachem (S.2a, S.2b, S.2d, S.2f, and S.3c), Proligo (S.2c, S.2e, S.2g, S.3d, S.3f, and S.3h), Glen Research (S.3a, S.3b, S.3e, S.3g, S.5a-5d, S.6a-d, S.7-16, and S.18), Dharmacon (S.4a-d), and Amersham Biosciences (S.17). Synthesis of S.1e and S.1f is unpublished. Matrices composed of glycerol (G), m-nitrobenzyl alcohol (NBA), diethanolamine (DEOA), 3:1 dithiothreitol/dithioerythritol (magic bullet, MB), triethanolamine (TEOA), or 1:1 dithiothreitol/α-thioglycerol (DTT/TG) can be purchased from TCI (Tokyo Kasei Kogyo). Chloroform and TEOA may be purchased from various suppliers. 1. Draw up ∼0.5 µL TEOA using a 2-µL disposable capillary pipet and apply it to the instrument’s sample target. 2. Place ∼0.02 to 0.1 mg phosphoramidite sample in a small glass tube and add ∼1 µL chloroform using another 2-µL disposable capillary pipet. Immediately draw up half of this solution (using the same pipet) and apply it to the TEOA on the target. This operation should be carried out quickly to avoid evaporation of chloroform.
3. To the prepared target, add 0.2 to 0.3 µL saline using a 2-µL disposable capillary pipet. Homogenize the solution on the target by drawing it in and expelling it out of the capillary pipet. KCl may be substituted for NaCl (see Background Information and Table 10.11.6).
4. Perform low-resolution LSIMS or FABMS analysis. 5. Prepare a new sample as described in steps 1 to 3 and analyze by high-resolution LSIMS or FABMS to obtain an accurate MW for the sample. The composition of the sample is determined by accurate mass measurement within 3 ppm of the theoretical mass of the sample. Mass Determination of Phosphoramidites
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OPTIMIZED MASS MEASUREMENT OF PHOSPHORAMIDITES USING TRIETHANOLAMINE/NaCl
ALTERNATE PROTOCOL
The best results may be obtained when phosphoramidite samples and matrix are prepared as described in this protocol under optimized conditions (see Commentary). When the Basic Protocol gives a low relative intensity (RI) for a sample, this protocol is recommended to enhance the RI (e.g., Table 10.11.7, entries 3, 6, 7).
Additional Materials (also see Basic Protocol) Microbalance (0.001 mg accuracy; e.g., Mettler ME30) 1-mL measuring safety pipet (e.g., Pyrex, Iwaki glass) 0.2- to 2-µL, 0.5- to 10-µL, and 5- to 50-µL micropipets (e.g., Finnpipette; Thermo Labsystems) 1. Draw up 0.5 µL TEOA using a 2-µL disposable capillary pipet and apply it to the instrument’s sample target. 2. Accurately weigh 10 µmol of a phosphoramidite sample to the nearest microgram using a microbalance. Place the sample in a small glass tube and add 0.5 mL chloroform using a 1-mL measuring safety pipet and micropipets. Immediately dispense 0.5 µL of this solution to the prepared TEOA on the target using a fresh 2-µL disposable capillary pipet. Given the difficulty of accurately weighing microgram samples, it is best to precisely weigh ∼10 µg of the sample using a microbalance and then adjust its concentration to 0.01µmol/0.5 µL by adding chloroform. This operation should be carried out quickly to avoid evaporation of chloroform.
3. To the prepared target, add 0.4 µL saline using a 2-µL disposable capillary pipet. Homogenize the solution on the target by drawing it in and expelling it out of the capillary pipet. KCl may be substituted for NaCl (see Background Information and Table 10.11.6).
4. Perform low-resolution LSIMS or FABMS analysis. 5. Prepare a new sample as in steps 1 to 3 and analyze by high-resolution LSIMS or FABMS to obtain an accurate MW for the sample. The composition of the sample is determined by accurate mass measurement within 3 ppm of the theoretical mass of the sample.
COMMENTARY Background Information DNA and RNA oligonucleotides are usually synthesized through phosphoramidite chemistry by sequential addition of nucleoside building blocks to a growing oligonucleotide chain that is covalently attached to a solid polymeric support (UNITS 3.1-3.3 and APPENDIX 3C; Caruthers et al., 1987; Reese, 2002). Nucleoside 3 -O-(2-cyanoethyl-N,Ndiisopropyl)phosphoramidites are the starting materials in automated solid-phase oligonucleotide synthesis (K¨oster et al., 1984; Usman et al., 1987; Wincott et al., 1995). The tert-butyldimethylsilyl (TBDMS) group is the most widely used protecting group for the 2 hydroxy function of RNA phosphoramidite
monomers. The N,N-diisopropyl phosphoramidite group can be activated by mild acidic treatment and reacts readily with nucleophiles (e.g., the free 5 -hydroxy group of the growing support-bound nucleotide) to generate a phosphite triester linkage that, upon oxidation, is rapidly converted to a relatively stable phosphotriester linkage. Upon completion of the synthesis, the 5 -O-dimethoxytrityl (DMTr) group is removed under acidic conditions, whereas the nucleobase and 2-cyanoethyl phosphate protecting groups are cleaved under basic conditions. These properties render the phosphoramidite building blocks inherently sensitive toward acids as well as basic nucleophiles.
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Figure 10.11.1 Incorporation of an imidazole moiety into RNA oligonucleotides using phosphoramidite S.1a. DMTr, 4,4 -dimethoxytrityl; POM, pivaloyloxymethyl; TBDMS, tert-butyldimethylsilyl. Reprinted from Fujitake et al. (2005) with permission from Elsevier.
Mass Determination of Phosphoramidites
To investigate the function of imidazole as a pseudo-nucleobase in ribozyme catalysis, an imidazole moiety was recently incorporated into RNA oligonucleotides (Fig. 10.11.1; Zhao et al., 2005; Wilson et al., 2006). A novel C4linked imidazole ribonucleoside phosphoramidite S.1a (Harusawa et al., 1996; Araki et al., 2004, 2005), functionalized with a basesensitive pivaloyloxymethyl (POM) group and a 2 -O-TBDMS group, was developed as a key ribonucleoside variant for probing general acid and base catalysis in ribozymes. However, MS characterization of S.1a has been difficult, because of its fragility as it decomposed even on a standard silica gel TLC plate (e.g., Merck 60 F254 ). Since crystallization of phosphoramidites is not generally possible and MS has been effective, the latter is used extensively for the characterization of phosphoramidites. Routine MS analyses require only nanogram amounts of material relative to the several milligrams needed for elemental analysis. The application of nanoelectrospray MS to the analysis of six standard 2 -deoxyribonucleoside phosphoramidites has been reported (Kele et al., 1999). Although FAB and LSI (Ashcroft, 1997) have been effective for MS analysis of phosphoramidites (Toren et al., 1986; Kele et al., 1999), it is not always possible to detect the molecular-related ions (MRIs) of various phosphoramidites by these methods. This limitation prompted the authors to develop the methods described in this unit.
Identification of a suitable matrix Initial MS analysis of phosphoramidite S.1a using conventional electron ionization (EI) and chemical ionization (CI) methods did not produce the MRI of S.1a (Table 10.11.1, entries 1 to 3), only that of the DMTr group as the base peak. Investigations were then directed toward LSIMS, which has been exploited for its mild and soft ionization of various types of labile materials. The matrix employed for LSIMS analysis plays a significant role as a supporting base (a relatively nonvolatile solvent holding the sample in position on the sample target) to achieve effective and stable ionization (Takayama, 1994; Ashcroft, 1997). Over the years, a number of matrices (Fig. 10.11.2) have been investigated for LSIMS. Glycerol (G) and m-nitrobenzyl alcohol (NBA) are popular choices, but they failed to give any useful MRIs for S.1a (Table 10.11.1, entries 4 to 5). Matrices composed of diethanolamine (DEOA), a 3:1 mixture of dithiothreitol/dithioerythritol (magic bullet, MB), a 1:1 mixture of dithiothreitol and αthioglycerol (DTT/TG), and triethanolamine (TEOA) also failed to provide MRIs for S.1a (entries 6 to 9). Cationization of molecules is an important analytical tool because of its simplicity and because of the abundant information that can be obtained from it (Teesch and Adams, 1992; Madhusudanan, 1995). Cationization is achieved easily by LSIMS and FABMS because in situ derivatization occurs when
10.11.4 Supplement 26
Current Protocols in Nucleic Acid Chemistry
a
Table 10.11.1 MS Conditions for Analysis of Phosphoramidite S.1a
Entry
Ionization Method
Matrixb
MRIc
RId (%)
1
EI (70 eV)
—
ND
2
EI (20 eV)
—
ND
3
CI (i-C4 H10 )
—
ND
4
LSIMS
G
—
ND
5
NBA
—
ND
6
DEOA
—
ND
7
MB
—
ND
8
DTT/TG
—
ND
9
TEOA
—
ND
10
G + NaCl
[M+Na]+
0.2
11
NBA + NaCl
[M+Na]+
0.6
12
DEOA + NaCl
[M+Na]+
0.7
13
MB + NaCl
[M+Na]+
0.4
14
DTT/TG + NaCl
[M+Na]+
0.3
15
TEOA + NaCl
[M+Na]+
20.1
G
—
ND
G + NaCl
—
16
FAB
17
TEOA + NaCl
18 19 20
[M+Na]
ESI MALDI-TOF
[M+Na]
Error f (ppm)
953.4625
0.4
953.5199
60.6
ND +
— THAP
Observed masse (m/z)
2.6 ND
+
—
g
a Reprinted from Fujitake et al. (2005) with permission from Elsevier. b DEOA, diethanolamine; DTT/TG, 1:1 dithiothreitol/thioglycerol; G, glycerol; MB (magic bullet), 3:1 dithiothreitol/dithioerythritol; NBA,
m-nitrobenzyl alcohol; TEOA, triethanolamine; THAP, 2 ,4 ,6 -trihydroxyacetophenone. c MRI, molecular-related ion. d RI, intensity relative to base peak ion (100%); ND, not detected (<0.1%). e Measured by HRMS. Theoretical mass for S.1a (C H N O PSi): (m/z) = 953.4621 [M+Na]+ . 50 71 4 9 f Error (ppm) = 106 × (observed mass - theoretical mass)/theoretical mass. g Not determined due to numerous peaks (mol. wt. <500) generated from THAP.
a sample is simply dissolved in a matrix. Thus, when NaCl was added to G, NBA, DEOA, MB, or DTT/TG as a 0.9% saline solution, the desired MRI peaks could be detected as [M+Na]+ with low relative intensities (RI < 0.7%; Table 10.11.1, entries 10 to 14). A combination of TEOA and NaCl produced a marked increase in the intensity of the MRI peak to 20.1% (entry 15). The MRI of S.1a by high-resolution MS (HRMS) appeared at m/z 953.4625 in the externally calibrated spectrum (polyethylene glycol), and the error on the observed mass determination was only 0.4 ppm from the expected mass value. The combination of TEOA and NaCl is the first effective matrix for analysis of phosphoramidites using soft ionization techniques. In addition, this matrix has the advantage of being useful for FABMS (Table 10.11.1, Current Protocols in Nucleic Acid Chemistry
entry 18). However, LSIMS analysis of S.1a in the negative mode using various matrices (G, NBA, TEOA, G+NaCl, NBA+NaCl, and TEOA+NaCl) did not lead to the MRI. Use of electrospray ionization (ESI) for S.1a did not show any useful peaks either (entry 19). Matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) MS was also investigated for analysis of S.1a. After examination of various matrices, the use of 2 ,4 ,6 -trihydroxyacetophenone (THAP) led to a [M+Na]+ for S.1a at m/z 953.5199 with an observed mass determination error of 60.6 ppm from the theoretical mass value (entry 20). The error using MALDI-TOF is 152 times larger than that obtained using the TEOA/NaCl matrix on LSIMS (0.4 ppm). Since determination of molecular composition requires mass errors
Purification and Analysis of Synthetic Nucleic Acids and Components
10.11.5 Supplement 26
Figure 10.11.2
Chemical structures of matrices used for LSIMS and FABMS.
Figure 10.11.3 Positive ionization LSIMS spectrum of S.1a (mol. wt. 930) introduced in a TEOA/NaCl matrix. Reprinted from Fujitake et al. (2005) with permission from Elsevier.
Mass Determination of Phosphoramidites
10.11.6 Supplement 26
Current Protocols in Nucleic Acid Chemistry
of less than ±3 ppm, MALDI-TOF-MS cannot give the molecular composition of S.1a. Because of the relatively large mass determination errors obtained from MALDI-TOFMS analyses, the method has limitations in the determination of molecular composition (Korshun et al., 2002; Ashraf et al., 2003; Mckeen et al., 2003; Zhu and Schmidt, 2003; Kirk and Bohn, 2004; Lu et al., 2004; Zatsepin et al., 2004). The low-resolution LSIMS spectrum of S.1a using the TEOA/NaCl matrix (Fig. 10.11.3) characterized its structure; the fragment ions at m/z 839 and 651 correspond to the Na+ -adducted ions generated upon removal of POM or DMTr from S.1a, together with a base peak due to the DMTr+ ion at m/z 303. Application of TEOA/NaCl matrix to C- and N-nucleoside phosphoramidites Phosphoramidites of uridine (S.2a) and thymidine (S.3a; Fig. 10.11.4), representing RNA and DNA monomers, respectively, have been analyzed similarly under various MS conditions (Table 10.11.2). EI-MS and CI-MS are ineffective for determining the mass of S.2a and 3a (Table 10.11.2, entries 1 to 3). Use of DEOA, MB, and TEOA matrices on LSIMS shows only 0.1% to 0.2% abundance of MRIs for S.2a in the absence of NaCl, and nothing for the DNA phosphoramidite monomer S.3a under the same conditions (entries 6 to 8). Conversely, the TEOA/NaCl matrix on LSIMS (entry 13) provides excellent MRI peaks for S.2a (19.1%) and S.3a (35.9%), with small errors on the observed mass determinations (0.1 and −0.3 ppm, respectively). Efficiency of the TEOA/NaCl matrix system in FABMS (entry 16) is evidenced by MRI peaks for S.2a or S.3a, although these have lower intensities (2.5% and 4.2%, respectively).
From the above results, the scope and limitation of the TEOA/NaCl matrix for LSIMS analysis of nucleoside phosphoramidites have been examined. Table 10.11.3 shows the observed and theoretical masses of various phosphoramidites whose chemical structures are shown in Figure 10.11.5. Use of LSIMS to analyze the unstable C4-linked imidazole Cnucleoside phosphoramidites S.1b-f (Araki et al., 2004, 2005) successfully provides the MRIs with low-ppm-level mass accuracy (error: <1.0 ppm; Table 10.11.3, entries 1 to 5). Commercially available ribonucleoside phosphoramidites S.2b-g, which are employed as building blocks in typical solidphase RNA synthesis programs, blended with the TEOA/NaCl matrix to give the expected LSIMS MRIs (entries 6 to 11). LSIMS MRIs of standard deoxyribonucleoside phosphoramidites S.3b-h, which are used for routine synthesis of DNA oligonucleotides, were also obtained under similar conditions (entries 12 to 18). Newly improved RNA synthons, 5 -OBzH-2 -O-ACE-protected phosphoramidites S.4a-d (Scaringe et al., 1998; Scaringe, 2000), were subjected to LSIMS analysis conditions to give the corresponding MRIs (entries 19 to 22), as were the 5 -O-DMTr-2 O-triisopropylsilyloxymethyl (TOM) ribonucleoside phosphoramidites S.5a-d (Pitsch et al., 2001; UNIT 3.8; entries 23 to 26). Further, the matrix was successfully applied to mass determination of the 5 -O-DMTr-2 O-Me RNA phosphoramidite monomers S.6ad (Beigelman et al., 2000; Roy and Tang, 2000; entries 27 to 30), which were designed to produce synthetic oligoribonucleotides exhibiting nuclease resistance properties. The method was extended to mass determination of phosphoramidites bound to a
Figure 10.11.4 Structures of phosphoramidites S.2a and S.3a. DMTr, 4,4 -dimethoxytrityl; TBDMS, tert-butyldimethylsilyl. Reprinted from Fujitake et al. (2005) with permission from Elsevier.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.11.7 Current Protocols in Nucleic Acid Chemistry
Supplement 26
a
Table 10.11.2 MS Conditions for Analysis of N-Nucleoside Phosphoramidites S.2a and S.3a
2ac MRIb
RIb (%)
EI (70 eV)
—
ND
ND
2
EI (20 eV)
—
ND
ND
3
CI (i-C4 H10 )
—
ND
ND
4
LSIMS
G
—
ND
ND
NBA
—
Entry
Ionization method
1
Matrixb
3ad
5 6
DEOA
7
MB
ND
ND
[M+H]
0.2
ND
[M+H]
+
0.1
ND
+
0.1
ND
TEOA
[M+H]
9
G + NaCl
—
ND
[M+Na]
0.1
0.2
[M+Na]
+
3.1
3.5
[M+Na]
+
2.5
ND
TEOA + NaCl
[M+Na]
+
19.1
G
—
G + NaCl
—
DEOA + NaCl MB + NaCl
12 13 14
ND +
NBA + NaCl
11
FAB
15
TEOA + NaCl
16
RIb (%)
+
8 10
Errorb (ppm)
[M+Na]
+
0.1
35.9
ND
ND
ND
ND
2.5
4.2
Errorb (ppm)
−0.3
a Reprinted from Fujitake et al. (2005) with permission from Elsevier. b See Table 10.11.1. c Theoretical mass for S.2a (C H N O PSi): (m/z) = 883.3840 [M+Na]+ ; observed: 883.3841. 45 61 4 9 d Theoretical mass for S.3a (C H N O P): (m/z) = 767.3182 [M+Na]+ ; observed: 767.3180. 40
49
4
8
variety of functional groups (Fig. 10.11.6). As shown in Table 10.11.4, the accurate masses of these functionalized phosphoramidites were measured with low average mass errors (<0.3 ppm) when the TEOA/NaCl matrix was employed for LSIMS analyses. In some cases, high-intensity [M+Na]+ peaks were observed (entries 4, 5, 10, and 11). These results demonstrate the potential value of this method.
Mass Determination of Phosphoramidites
Optimal molar ratios of phosphoramidites, NaCl, and TEOA During the course of the study, it became clear that the MRIs of phosphoramidites are influenced by the concentration of NaCl in the matrix. Thus, the relationship between MRI intensity and the molar ratio of NaCl to phosphoramidite sample was examined while holding the amount of TEOA constant at 0.5 µL. As shown in Figure 10.11.7, the MRI intensities of S.3a increased with increasing molar ratios of NaCl/S.3a until the RI reached a maximum of 30.4% at a ratio of 6 (see Table 10.11.5, entry 7). Phosphoramidites S.2a and S.4b also produced the highest RIs (25.8% and 7.7%, respectively) under these conditions, indicat-
ing that a NaCl/phosphoramidite molar ratio of 6 is optimal. A combination of 0.01 µmol phosphoramidite, 0.06 µmol NaCl, and 0.5 µL TEOA is therefore recommended as a sample/matrix composition for LSIMS analysis of phosphoramidites. Effect of metal ion in the matrix for LSIMS measurement of phosphoramidites Use of a matrix in the presence of a metal ion has been known to be effective in the formation of an abundant metal ion adduct via LSIMS and FABMS (Teesch and Adams, 1992; Madhusudanan, 1995). Thus, alkali and alkaline earth metal cationization of phosphoramidites has been studied using metal ions such as Li+ , Na+ , K+ , Rb+ , Cs+ , Mg2+ , and Ca2+ under the most suitable matrix condition (Table 10.11.5, entry 7). Addition of LiCl to a solution of phosphoramidite S.2a in TEOA yielded only a 2.5% RI for the [M+Li]+ ion (Table 10.11.6, S.2a, entry 1). Addition of NaCl or KCl to the solution increased the RIs to 25.8% and 12.9%, respectively (entries 2 and 3). Conversely, the presence of RbCl, CsCl, MgCl2 , or CaCl2 led to low RIs for
10.11.8 Supplement 26
Current Protocols in Nucleic Acid Chemistry
Table 10.11.3 LSIMS Measurements Using TEOA/NaCl Matrix: Accurate Molecular Weights of Synthetic and a Commercial Nucleoside Phosphoramidites
Theoretical massc
Observed massc
RId (%)
Errord (ppm)
C47 H61 N4 O9 P
879.4070
879.4069
2.2
−0.1
S.1c
C44 H57 N4 O8 P
823.3809
823.3801
3.5
−1.0
3
S.1d
C50 H71 N4 O9 PSi
953.4621
953.4618
14.6
−0.3
4
S.1e
C52 H75 N4 O9 PSi
981.4934
981.4935
1.6
0.1
5
S.1f
C52 H75 N4 O9 PSi
981.4934
981.4932
0.4
−0.2
6
S.2b (Ac-C)
C47 H64 N5 O9 PSi
924.4105
924.4106
3.6
0.1
7
S.2c (tac-C)
C57 H76 N5 O10 PSi
1072.4993
1072.4991
3.0
−0.2
8
S.2d (Bz-A)
C53 H66 N7 O8 PSi
1010.4373
1010.4375
5.5
0.2
9
S.2e (tac-A)
C58 H76 N7 O9 PSi
1096.5105
1096.5109
4.8
0.4
10
S.2f (iBu-G)
C50 H68 N7 O9 PSi
992.4479
992.4476
7.7
−0.3
11
S.2g (tac-G)
C58 H76 N7 O10 PSi
1112.5054
1112.5056
6.1
0.2
12
S.3b (Ac-dC)
C41 H50 N5 O8 P
794.3291
794.3290
1.3
−0.1
13
S.3c (Bz-dC)
C46 H52 N5 O8 P
856.3448
856.3448
2.1
0.0
14
S.3d (tac-dC)
C51 H62 N5 O9 P
942.4179
942.4182
2.8
0.3
15
S.3e (Bz-dA)
C47 H52 N7 O7 P
880.3561
880.3560
15.3
−0.1
16
S.3f (tac-dA)
C52 H62 N7 O8 P
966.4292
966.4294
15.5
0.2
17
S.3g (iBu-dG)
C44 H54 N7 O8 P
862.3666
862.3669
8.6
0.3
18
S.3h (tac-dG)
C52 H62 N7 O9 P
982.4241
982.4242
9.3
0.1
19
S.4a (U)
C44 H70 N3 O16 PSi3
1034.3695
1034.3699
3.5
0.4
20
S.4b (Ac-C)
C46 H73 N4 O16 PSi3
1075.3961
1075.3963
2.4
0.2
21
S.4c (iBu-A)
C49 H77 N6 O15 PSi3
1127.4386
1127.4387
6.9
0.1
22
S.4d (iBu-G)
C49 H77 N6 O16 PSi3
1143.4335
1143.4340
9.3
0.4
23
S.5a (U)
C49 H69 N4 O10 PSi
955.4414
955.4416
11.0
0.2
24
S.5b (Ac-C)
C51 H72 N5 O10 PSi
996.4680
996.4680
1.3
0.0
25
S.5c (Ac-A)
C52 H72 N7 O9 PSi
1020.4792
1020.4791
15.4
−0.1
26
S.5d (Ac-G)
C52 H72 N7 O10 PSi
1036.4741
1036.4742
7.6
0.1
27
S.6a (U)
C40 H49 N4 O9 P
783.3132
783.3133
20.7
0.1
28
S.6b (Bz-C)
C47 H54 N5 O9 P
886.3554
886.3551
10.9
−0.3
29
S.6c (Bz-A)
C48 H54 N7 O8 P
910.3666
910.3670
12.1
0.4
30
S.6d (DMAM-G)
C44 H55 N8 O8 P
877.3775
877.3777
17.4
0.2
Entry
Phosphoramiditeb
Formula
1
S.1b
2
a Reprinted from Fujitake et al. (2005) with permission from Elsevier. b Abbreviations: Ac, acetyl; Bz, benzoyl; DMAM, N,N-dimethylaminomethylene; iBu, isobutyryl; tac, 4-(tert-butyl)phenoxyacetyl. c [M+Na]+ ; MS measurements carried out as described in the Basic Protocol. d See Table 10.11.1.
Purification and Analysis of Synthetic Nucleic Acids and Components
10.11.9 Current Protocols in Nucleic Acid Chemistry
Supplement 26
Figure 10.11.5 Synthetic and commercial nucleoside phosphoramidites analyzed by LSIMS using the TEOA/NaCl matrix system. Bz, benzoyl; DMTr, 4,4 -dimethoxytrityl; i-Bu, isobutyryl; i-Pr, isopropyl; POM, pivaloyloxymethyl; tac, 4-(tertbutyl)phenoxyacetyl; TBDMS, tert-butyldimethylsilyl; TMS, trimethylsilyl; TOM, [(triisopropylsilyl)oxy]methyl. Reprinted from Fujitake et al. (2005) with permission from Elsevier.
10.11.10 Supplement 26
Current Protocols in Nucleic Acid Chemistry
Figure 10.11.6 Nucleoside and non-nucleoside phosphoramidites analyzed by LSIMS using the TEOA/NaCl matrix system: 3-nitropyrrole 2 -deoxynucleoside S.7 as a universal nucleoside, two halogenated-nucleosides S.8 and S.9, 5 -phosphoramidite S.10, 5 -amino modified S.11, photocleavable biotin phosphoramidite S.12 (Olejnik et al; 1996), phosphoramidite synthon S.13 for oligonucleotide dendrimers, disulfide thiol modifier S.14, amino-modified C6 dC S.15, fluorescein phosphoramidite S.16, Cy3 phosphoramidite S.17, and abasic phosphoramidite S.18 (Shishkina and Johnson, 2000). Reprinted from Fujitake et al. (2005) with permission from Elsevier.
10.11.11 Current Protocols in Nucleic Acid Chemistry
Supplement 26
Table 10.11.4 LSIMS Measurements Using TEOA/NaCl Matrix: Accurate Molecular Weights of Nucleoside and Nona nucleoside Phosphoramidites
Theoretical massb
Observed massb
C39 H47 N4 O8 P
753.3026
753.3024
8.5
−0.3
S.8
C39 H46 BrN4 O8 P
831.2131
831.2129
4.5
−0.2
3
S.9
C41 H49 FN5 O8 P
812.3197
812.3200
8.3
0.4
4
S.10
C40 H49 N4 O8 P
767.3183
767.3180
63.3
−0.4
5
S.11
C14 H25 N3 O3 PF3
394.1482
394.1482
42.0
0.0
6
S.12
C55 H72 N7 O9 PS
1060.4744
1060.4747
2.6
0.3
7
S.13
C64 H79 N4 O10 P
1117.5427
1117.5425
6.0
−0.2
8
S.14
C42 H61 N2 O5 S2 P
791.3654
791.3655
1.0
0.1
9
S.15
C53 H68 F3 N8 O9 P
1071.4692
1071.4688
5.3
−0.4
10
S.16
C46 H58 N3 O10 P
866.3754
866.3750
100.0
−0.5
917.5131
917.5130
76.1
−0.1
1033.5713
1033.5712
7.5
−0.1
Entry
Phosphoramidite
1
S.7
2
Formula
−
11
S.17
C58 H70 N4 O4 P(Cl )
12
S.18
C54 H91 N2 O8 PSi3
RIc (%)
Errorc (ppm)
a Reprinted from Fujitake et al. (2005) with permission from Elsevier. b [M+Na]+ except for compound S.17, which was detected as [M–Cl− ]+ ; MS measurements carried out as described in the Basic Protocol. c See Table 10.11.1.
Figure 10.11.7 Relationship between RIs of MRIs and molar ratios of NaCl/sample. TEOA constant at 0.5 µL. Reprinted from Fujitake et al. (2005) with permission from Elsevier.
S.2a, ranging from 1.1% to 8.0% (entries 4 to 7). Similar trends for phosphoramidites S.3a and S.4b were observed under identical conditions. These results show that NaCl and KCl are superior to other alkali and alkaline earth metals for enhancement of the RIs of MRIs. KCl may be used as a substitute for NaCl in the matrix for phosphoramidite mass analysis.
Mass Determination of Phosphoramidites
FABMS measurements of phosphoramidites under optimal conditions LSIMS and FABMS have been used most extensively to form metal ion adducts to confirm relative molecular masses (Ashcroft, 1997). Thus, the usefulness of the present method for FABMS was further confirmed us-
ing the optimal conditions described in the Alternate Protocol. The ten phosphoramidites listed in Table 10.11.7 gave MRIs with small errors (<1.2 ppm). Phosphoramidites S.4b, S.12, and S.14 produced RIs of only 2.4%, 2.6%, and 1.0%, respectively, when analyzed by LSIMS as described in the Basic Protocol (Table 10.11.3, entry 20, and Table 10.11.4, entries 6 and 8), but their FABMS measurements under the Alternate Protocol markedly enhanced their RIs to 44.4%, 10.2%, and 10.7%, respectively (Table 10.11.7, entries 3, 6, and 7). These results show that LSIMS and FABMS are complementary approaches to phosphoramidite analysis using the present matrix system.
10.11.12 Supplement 26
Current Protocols in Nucleic Acid Chemistry
Table 10.11.5 Suitable Molar Ratios for NaCl and Phosphoramidite Samplea,b
1
2
4
5
6
7
0.1
0.1
0.1
0.05
0.02
0.01
0.01
0.01
NaCl (µmol)
0
0.02
0.03
0.03
0.03
0.03
0.06
0.06
NaCl/sample
0
0.15
0.3
0.6
1.5
3
6
S.2a
0
2.9
5.5
7.6
12.9
24.7
25.8
21.2
S.3a
0
6.6
9.6
13.7
17.1
25.4
30.4
27.1
S.4b
0
1.1
2.4
3.8
2.4
4.7
7.7
6.3
Entry Phosphoramidite (µmol)
Relative intensity of [M+Na]+
3
8
12
a TEOA fixed at 0.5 µL. b Reprinted from Fujitake et al. (2005) with permission from Elsevier.
Table 10.11.6 Effect of Metal Ion in TEOA Matrix for MRI of Phosphoramiditesa
S.2ab
S.3ab
S.4bb
Entry
MRI
RIc (%) (m/z mass)d
RIc (%) (m/z mass)d
RIc (%) (m/z mass)d
1
LiCl
2.5 (867)
6.9 (751)
2.4 (1059)
2
NaCl
25.8 (883)
30.4 (767)
7.7 (1075)
3
KCl
12.9 (899)
35.8 (783)
3.1 (1091)
4
RbCl
2.7 (945)
10.3 (829)
2.8 (1137)
5
CsCl
6 7
8.0 (993)
4.6 (877)
2.7 (1185)
e
1.1 (1032)
2.6 (916)
0.1 (1224)
e
3.3 (1048)
4.6 (932)
0.1 (1240)
MgCl2 CaCl2
a MS measurements carried out as described in the Alternate Protocol using 0.01 µmol phosphoramidite, 0.06 µmol
metal (I or II) chloride, and 0.5 µL TEOA. b S.2a (C H N O PSi): mol. wt. 860; S.3a (C H N O P): mol. wt. 744; S.4b (C H N O PSi ): mol. wt. 1052. 40 49 4 8 46 73 4 16 3 45 61 4 9 c See Table 10.11.1. d Unless otherwise noted, mass shown as [M+Metal]+ . e MRIs observed as [M+TEOA+MCl ]+ . 2
Critical Parameters and Troubleshooting The use of the combined TEOA/NaCl matrix is key, and is a more important factor than the ratio of the phosphoramidite, saline, and TEOA. The purity of the synthetic phosphoramidites is also an important factor. Standard synthetic N-nucleoside phosphoramidites, which contain acid-labile moieties, can generally be purified by column chromatography on silica gel using several kinds of solvents containing low concentrations of triethylamine for elution. In most cases, the phosphoramidites can be analyzed immediately after purification or can be stored at −20◦ C for future use. Optimal MS results will be obtained if the sample is analyzed immediately after purification. All phosphoramidites in the present unit, except the newly synthesized S.1a-f, are
commercially available building blocks used in typical automated synthesizer programs. The imidazole C-nucleoside phosphoramidites S.1a-d and the carbon-elongated imidazole C-nucleosides S.1e-f (Fig. 10.11.1 and Fig. 10.11.5; Araki et al., 2004, 2005) were synthesized for probing general acid and base catalysis in ribozymes (Zhao et al., 2005; Wilson et al., 2006; see Background Information). These compounds are more labile and fragile than ordinary N-nucleoside phosphoramidites, decomposing even on standard TLC plates (e.g., Merck 60 F254 ). Therefore, purification of S.1a-f employs a special basic (N-H) silica gel, Chromatorex NH-DM 1020 (Fuji Silisia Chemical). These phosphoramidites should be handled very carefully during purification (Araki et al., 2005). Purified imidazole nucleoside S.1a has been stored for several months at −20◦ C to evaluate its
Purification and Analysis of Synthetic Nucleic Acids and Components
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Table 10.11.7 RIs and Errors on FABMS Measurements of Phosphoramidites Under Optimal Conditionsa
Entry
Phosphoramidite
RI (%)b
1
S.2a
29.2
2
S.3a
75.4
3
S.4b
44.4 (2.4)
4
S.5c
34.2
5
S.6d
71.4
6
S.12
Error (%)b 0.6 1.2
c
−0.1 −0.5 1.0
10.2 (2.6)
c
0.8
c
0.6
7
S.14
10.7 (1.0)
8
S.15
10.7
−0.1
9
S.16
100.0
0.5
10
S.17
100.0
0.1
a Measurements carried out according to the Alternate Protocol. b See Table 10.11.1. c Numbers in parentheses from LSIMS measurements using the Basic Protocol. See Table 10.11.3, entry 20 (S.4b) and Table 10.11.4, entries 6 (S.12) and 8 (S.14).
long-term stability; 31 P and 1 H NMR data did not show any substantial decomposition under these conditions. Nonetheless, the authors recommend that imidazole phosphoramidites S.1a-f be analyzed immediately after purification to obtain the best results. The synthesis of S.1a-f should be performed only by personnel trained and experienced in organic synthesis. Standard precautions to prevent excessive exposure to toxic chemicals and solvents should be followed. All reactions should first be performed on a small scale.
Anticipated Results
Mass Determination of Phosphoramidites
The TEOA-NaCl matrix on LSIMS or FABMS equipped with a double-focusing mass spectrometer was quite useful for molecular weight measurements of phosphoramidites. The present method rapidly and easily measures the accurate molecular weights of various phosphoramidites as adduct ions [M+Na]+ with average mass error smaller than 0.2 ppm. All of the 45 phosphoramidites investigated using the Basic Protocol gave MRIs using the present method (Tables 10.11.3 and 10.11.4). The Alternate Protocol describes the MS measurement of phosphoramidites under optimized conditions for phosphoramidite, NaCl, and TEOA (Table 10.11.5, entry 7). Indeed, phosphoramidites showing low RI signals by LSIMS measurements under the conditions described in the Basic Protocol revealed markedly enhanced RI signals by FABMS
(Table 10.11.7, entries 3, 6, and 7) under the optimized conditions outlined in the Alternate Protocol. Hence, the conditions described in the Alternate Protocol are effective in enhancing RI signals that might otherwise be low under the conditions described in the Basic Protocol.
Time Considerations MS measurement of each of the phosphoramidites described in the Basic Protocol may be carried out within 30 min. The procedures in the Alternate Protocol can take 3 or 4 hr per sample for precise adjustment of the molar ratio of sample, NaCl, and TEOA.
Literature Cited Araki, L., Harusawa, S., Yamaguchi, M., Yonezawa, S., Taniguchi, N., Lilley, D.M.J., Zhao, Z., and Kurihara, T. 2004. Synthesis of C4-linked imidazole ribonucleoside phosphoramidites with pivaloyoxymethyl (POM) group. Tetrahedron Lett. 45:2657-2661. Araki, L., Harusawa, S., Yamaguchi, M., Yonezawa, S., Taniguchi, N., Lilley, D.M.J., Zhao, Z., and Kurihara, T. 2005. Synthesis of novel C4-linked imidazole ribonucleoside phosphoramidites for probing general acid and base catalysis in ribozyme. Tetrahedron 61:11976-11985. Ashcroft, A.E. 1997. Ionization Methods in Organic Mass Spectrometry. The Royal Society of Chemistry, Cambridge, U.K. Ashraf, M.A., Notta, J.K., and Snaith, J.S. 2003. Solid phase synthesis of peptide dimers and trimers linked through an N-terminal lysine residue. Tetrahedron Lett. 44:9115-9119. Beigelman, L., Haeberli, P., Sweedler, D., and Karpeisky, A. 2000. Improved synthetic
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approaches toward 2 -O-methyl-adenosine and guanosine and their N-acyl derivatives. Tetrahedron 56:1047-1056. Caruthers, M.H., Barone, A.D., Beaucage, S.L., Dodds, D.R., Fisher, E.F., McBride, L.J., Matteucci, M., Stabinsky, Z., and Tang, J.-Y. 1987. Chemical synthesis of deoxyoligonucleotides by the phosphoramidite method. Methods Enzymol. 154:287-313.
Pitsch, S., Weiss, P.A., Jenny, L., Stutz, A., and Wu, X. 2001. Reliable chemical synthesis of oligoribonucleosides (RNA) with 2 -O[(triisopropylsilyl)oxy]methyl (2 -O-tom)protected phosphoramidites. Helv. Chim. Acta. 84:3773-3795. Reese, C.B. 2002. The chemical synthesis of oligoand poly-nucleotides: A personal commentary. Tetrahedron 58:8893-8920.
Fujitake, M., Harusawa, S., Araki, L., Yamaguchi, M., Lilley, D.M.J., Zhao, Z.-Y., and Kurihara, T. 2005. Accurate molecular weight measurements of nucleoside phosphoramidites: a suitable matrix of mass spectrometry. Tetrahedron 61:4689-4699.
Roy, S.K. and Tang, J.-Y. 2000. Efficient large scale synthesis of 2 -O-alkyl pyrimidine ribonucleosides. Org. Process Res. Dev. 4:170-171.
Harusawa, S., Murai, Y., Moriyama, H., Imazu, T., Ohishi, H., Yoneda, R., and Kurihara, T. 1996. Efficient and β-stereoselective synthesis of 4(5)-(β-D-ribofuranosyl)- and 4(5)-(2deoxyribofuranosyl)imidazoles. J. Org. Chem. 61:4405-4411.
Scaringe, S.A., Wincott, F.E., and Caruthers, M.H. 1998. Novel RNA synthesis method using 5 -Osilyl-2 -O-orthoester protecting groups. J. Am. Chem. Soc. 120:11820-11821.
Kele, Z., Kupih´ar, Z., Kov´acs, L., Jan´aky, T., and Szab´o, P.T. 1999. Electrospray mass spectrometry of phosphoramidites, a group of acid-labile compounds. J. Mass Spectrom. 34:1317-1321. Kirk, J.S. and Bohn, P.W. 2004. Surface adsorption and transfer of organomercaptans to colloidal gold and direct identification by matrix assisted laser desorption/ionization mass spectrometry. J. Am. Chem. Soc. 126:59205926. Korshun, V.A., Stetsenko, D.A., and Gait, M.J. 2002. Novel uridin-2 -yl carbamates: Synthesis, incorporation into oligodeoxyribonucleotides, and remarkable fluorescence properties of 2 -pyren-1-ylmethylcarbamate. J. Chem. Soc. 1:1092-1104. K¨oster, H., Biernat, J., McManus, J., Wolter, A., Stumpe, A., Narang, C.K., and Sinha, N.D. 1984. Polymer support oligonucleotide synthesis–XV. Synthesis of oligodeoxynucleotides on controlled pore glass (CPG) using phosphate and a new phosphite triester approach. Tetrahedron 40:103-112. Lu, X., He, X., Feng, L., Shi, Z., and Gu, Z. 2004. Synthesis of pyrrolidine ring-fused metallofullerene derivatives. Tetrahedron 60:37133716. Madhusudanan, K.P. 1995. Alkali metal cationization and its effect on the collision-induced decomposition of taxol. J. Mass Spectrom. 30:703707. McKeen, C.M., Brown, L.J., Nicol, J.T.G., Mellor, J.M., and Brown, T. 2003. Synthesis of fluorophore and quencher monomers for use in scorpion primers and nucleic acid structural probes. Org. Biomol. Chem. 1:2267-2275. Olejnik, J., Krzyma´nska-Olejnik, E., and Rothschild, K.J. 1996. Photocleavable biotin phosphoramidite for 5 -end-labeling, affinity purification and phosphorylation of synthetic oligonucleotides. Nucl. Acids Res. 24:361366.
Scaringe, S.A. 2000. Advanced 5 -silyl-2 orthoester approach to RNA oligonucleotide synthesis. Methods Enzymol. 317:3-19.
Shishkina, I.G. and Johnson, F. 2000. A new method for the postsynthetic generation of abasic sites in oligomeric DNA. Chem. Res. Toxicol. 13:907912. Takayama, M. 1994. Fast atom bombardment mass spectrometry: matrix effects on ion formation and fragmentation. J. Mass Spectrom. Soc. Jpn. 42:249-275. Teesch, L.M. and Adams, J. 1992. Metal ions as special reagents in analytical mass spectrometry. 1992. Org. Mass Spectrom. 27:931-943. Toren, P.C., Betsch, D.F., Weith, H.L., and Coull, J.M. 1986. Determination of impurities in nucleoside 3 -phosphoramidites by fast atom bombardment mass spectrometry. Anal. Biochem. 152:291-294. Usman, N., Ogilvie, K.K., Jiang, M.-Y., and Cedergren, R.J. 1987. Automated chemical synthesis of long oligoribonucleotides using 2 -Osilylated ribonucleoside 3 -O-phosphoramidites on a controlled-pore glass support: Synthesis of a 43-nucleotide sequence similar to the 3 half molecule of an Escherichia coli formylmethionine tRNA. J. Am. Chem. Soc. 109:78457854. Wincott, F., DiRenzo, A., Shaffer, C., Grimm, S., Tracz, D., Workman, C., Sweedler, D., Gonzalez, C., Scaringe, S., and Usman, N. 1995. Synthesis, deprotection, analysis and purification of RNA and ribozymes. Nucl. Acids Res. 23:2677-2684. Wilson, T., Ouellet, J., Zhao, Z, Harusawa, S., Araki, L., Kurihara, T., and Lilley, M.J. 2006. Nucleobase catalysis in the hairpin ribozyme. RNA 12:980-987. Zatsepin, T.S., Ivanova, Y.M., and Oretskaya, T.S. 2004. Synthesis of (2 S)- and (2 R)-2 -deoxy2 -[(2-methoxyethoxy)amino]pyridine nucleosides and oligonucleotides. Chem. Biodiver. 1:1537-1545. Zhao, Z., Mcleod, A., Harusawa, S., Araki, L., Yamaguchi, M., Kurihara, T., and Lilley, D.M.J. 2005. Nucleobase participation in ribozyme catalysis. J. Am. Chem. Soc. 127:5026-5027.
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Zhu, X. and Schmidt, R.R. 2003. Selective conversion of N-trichloroethoxycarbonyl (Troc) groups into N-acetyl groups in the presence of N-tertbutoxycarbonyl (Boc) protecting groups. Synthesis 2003:1262.
Contributed by Shinya Harusawa, Mihoyo Fujitake, and Takushi Kurihara Osaka University of Pharmaceutical Sciences Osaka, Japan Zheng-yun Zhao and David M.J. Lilley University of Dundee Dundee, Scotland
Mass Determination of Phosphoramidites
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Current Protocols in Nucleic Acid Chemistry
CHAPTER 11 RNA Folding Pathways INTRODUCTION lucidating the mechanisms by which macromolecules such as RNA fold into their three-dimensional biologically active conformations is among the most challenging research endeavors at the chemistry-biology interface. Because of the connection between structure and biological function, defining the factors that govern RNA folding should lead to a better understanding of how ribonucleic acids perform their functions in vivo. In practical terms, this understanding will also facilitate efforts to predict the threedimensional structure of complex RNAs from primary sequence and to design RNA molecules that adopt stable structures possessing specific binding and catalytic properties. Such RNA molecules hold considerable promise as biochemical tools and diagnostic reagents, and as starting points for the development of therapeutics for human disease.
E
A complete understanding of folding requires knowledge of the structures and energetics of each conformational state of a given molecule along the pathway from the unfolded structure to the final native functional state. However, much of the research on RNA folding conducted over the last 25 years has focused on elucidating the structures and stability of folded RNA molecules. Recent advances in X-ray crystallography, coupled with improved methods for both the synthesis and analysis of RNA, have facilitated significant advances in efforts to elucidate RNA folding pathways. The goal of this chapter is to present the reader with a current understanding of the problems in the field of RNA folding and the most useful and cutting-edge approaches to solving these problems. is an introductory commentary that addresses important considerations such as the questions that one wishes to ask about RNA structure, the difference between RNA and proteins in structure analysis, and the difference between folding and unfolding. UNIT 11.2 focuses on the prediction of RNA secondary structure using RNAstructure and mfold. UNIT 11.3 uses thermal analysis to study the tertiary structure of an RNA as it unfolds. In UNIT 11.4, different conformers of an RNA are separated based on structure-dependent electrophoretic properties and their activity is assayed in the gel, providing a simple means to correlate structure and function. UNIT 11.5 describes the use of circular dichroism and urea to look at RNA structure transition, which can be used to accurately and rapidly determine thermodynamic parameters in a wide variety of conditions. UNIT 11.6 presents an elegant use of X-rays for time-resolved hydroxyl radical footprinting of RNA. UNIT 11.7 uses magnesium chelation to study real-time tertiary unfolding of RNA. UNIT 11.8 & 11.9 discuss the use of fluorescence and chemical modification for studying the kinetics of RNA folding. UNIT 11.10 complements UNIT 11.8 by presenting the use of fluorescence resonance energy transfer to study the kinetics as well as the structural basis of a conformational change. Finally, UNIT 11.11 presents methods for synthesizing large pyrene-labeled RNAs that can be used in fluorescence experiments. UNIT 11.1
Gary D. Glick
RNA Folding Pathways Contributed by Gary D. Glick Current Protocols in Nucleic Acid Chemistry (2004) 11.0.1 C 2004 by John Wiley & Sons, Inc. Copyright
11.0.1 Supplement 19
RNA Folding Pathways OVERVIEW OF THE QUESTIONS The questions to be answered in the study of RNA folding are simple. What is the final structure? How does the molecule get there? How long does it take? If only the answers were equally simple. In cells, RNA is constantly being synthesized and folded while it undergoes continuous conformational switching and degradation. A vast time domain is covered, from picoseconds and nanoseconds for motions in individual nucleotides, to microseconds for forming small hairpin helices, to milliseconds for forming relatively compact tertiary structures such as in tRNA, to minutes for folding more complex molecules such as introns. The process of undoing incorrect folds, or renaturation, can take hours, days, or much longer. The final folded state is usually well defined, but the starting state and transient intermediates are likely to be very polymorphic. The number of possible folding pathways is huge for large molecules, and a correspondingly complex time spectrum can be expected. Different regions of the molecule may have quite distinct time constants for reaching their final form, so it is essential that the experiments be capable of resolution in both time and space. As complex as this problem may seem, there is some consolation in the fact that it is probably simpler than the protein folding problem.
HOW RNA AND PROTEINS DIFFER IN THE FOLDING PROBLEM Secondary structure is a stronger organizing principle in RNA than in proteins. For example, imagine that an αβ-dimer-forming protein is mixed with all the proteins in the cell, and the solution is subjected to thermal or chemical denaturation, causing separation of the subunits. Then it is slowly annealed back to native conditions. There is only a small probability that native αβ dimer will emerge from this messy process. Yet, if the same experiment were done with a double-helical RNA in the presence of all the RNA in the cell, the chances of recovering the duplex with native pairing are quite good, as verified by the widespread use of hybridization to identify DNA and RNA sequences in complex nucleic acid mixtures. (In such hybridizations, of course, an excess of strand α is added in order to drive the kinetics of formation of the αβ duplex.)
UNIT 11.1 Double-helical nucleic acids are stabilized primarily by the free energy of stacking the bases together. However, this interaction is relatively nonspecific, since there are only modest variations in the enthalpy of base pairing, depending on the nature of the dinucleotide step (Breslauer et al., 1986; Xia et al., 1998; UNIT 11.2). Specificity arises from the highly polar base-pairing interactions. The result of these strong, directional forces is that the secondary structure elements of RNA, i.e., local hairpin helices, are thermodynamically stable in the absence of other interactions. This is in contrast to proteins, in which α helices usually have only marginal stability by themselves. RNAs are capable of forming stable hairpin helices rapidly, followed by slower formation of the noncanonical interactions in the tertiary structure (Brion and Westhof, 1997; Tinoco and Bustamante, 1999). In these simple cases, the folding pathway through the energy “landscape” is well defined, although even apparently simple cases can involve some rearrangement of secondary structure upon tertiary structure formation (Wu and Tinoco, 1998). In more extreme cases, the high stability of improper double helices can cause RNAs to take long detours through metastable states in deep local energy minima that are characterized as “denatured” by their distinctive mobility on electrophoresis gels. The comparatively large per-residue enthalpy of double helix formation means that elevated temperature is frequently needed to overcome the activation barrier for melting the incorrect structure to produce the native form.
HOW WE KNOW THE FINAL STRUCTURE In only a few cases does crystallography provide knowledge of the structure of RNA molecules large enough to encode most of the complexity of the folding problem. The classic example is tRNA, augmented by more recent work on ribozymes such as the P4-P6 domain of the Tetrahymena ribozyme (Cate et al., 1996). For molecules of this size, NMR spectroscopy is not yet a useful tool, nor is any other spectroscopic method able to provide a general solution of the structural problem. The primary source of definitive information about the secondary structure and some tertiary structural elements in naturally occur-
Contributed by Donald M. Crothers Current Protocols in Nucleic Acid Chemistry (2000) 11.1.1-11.1.5 Copyright © 2000 by John Wiley & Sons, Inc.
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11.1.1 Supplement 2
ring RNAs is phylogenetic analysis of the conservation of pairing interactions. The basic idea is that base pairs in a structure have been conserved over evolution, even when particular sequences were not. Thus, if mutation of A to G at one site is strongly correlated with mutation of T to C at another, it is likely that the two positions are base paired, probably by WatsonCrick pairing. This logic enabled, for example, the work that defined the secondary structure of 23S ribosomal RNA (Noller et al., 1981), the tertiary structure of RNase P (Harris and Pace, 1995), and the secondary structures of Y RNAs (Farris et al., 1999) and telomerase RNA (Chen et al., 2000). An alternative approach is to compare the predicted thermodynamic stability of different possible secondary structures, and select the most stable of these. The empirical parameters and software necessary for this analysis have recently been updated by Turner and Zucker and their collaborators (UNIT 11.2; Xia et al., 1998; Matthews et al., 1999). This method has the disadvantage of not accounting for tertiary structure that can influence the relative free energy of structures that differ in secondary structure. The current methods predict ∼73% of the known base pairs (Matthews et al., 1999). However, knowledge of relative secondary structure free energies can be vitally important when assessing the possible importance of alternative transient structures in the folding pathway. Finally, there is the set of low-resolution structural methods that come under the heading of footprinting. These take advantage of the variation in chemical and enzymatic reactivity depending on the structural state of individual nucleotides. Of particular interest for the folding problem are those that can be carried out rapidly enough to follow the folding reaction in real time (see Resolution in Space and Time).
FOLDING VERSUS UNFOLDING
RNA Folding Pathways
The earliest studies of the RNA folding problem involved observation of thermal melting curves, e.g., for tRNAs (Riesner et al., 1969, 1970, 1973; Cole and Crothers, 1972; Cole et al., 1972). It might be asked how one can learn about folding by studying unfolding, but it should be recognized that relaxation kinetic measurements yield both forward and reverse reaction rate constants, so that information about the folding steps is inherent in the kinetic data. Direct measurement of folding rates for the tertiary structure in yeast tRNAPhe by
stopped-flow mixing with Mg2+ to induce folding verifies the convergence of the two approaches (Maglott and Glick, 1997). A perhaps more serious reservation has to do with the series of equilibrium states examined in relaxation kinetics. Small temperature jumps clearly do not correspond to the intracellular process, nor to the way folding is triggered in nature. But neither is a sudden jump in Mg2+ concentration to induce folding a physiologically relevant mechanism. The pathway of RNA folding in the cell is a largely unexplored experimental problem. Understanding of the thermodynamics of RNA secondary structure is now sufficiently advanced that individual components of multiphasic melting curves for small model RNAs can generally be assigned to melting of specific helices (UNIT 11.3; Draper and Gluick, 1995). For somewhat more complicated molecules, the disagreement between predicted and observed melting curves can confidently be assigned to tertiary structural interactions, although often of unknown nature (Draper and Gluick, 1995; UNIT 11.3).
RESOLUTION IN SPACE AND TIME The general objective of studies of RNA folding pathways is to chart the state of individual nucleotides, helices, or tertiary structural elements as a function of time after a perturbation alters the conformational state. The early work on the dynamics of tRNA tertiary structure, referred to above, exploited the instability of the tertiary structure in absence of Mg2+, thus making the tertiary structure unfolding transition the first component of the thermal melting curve (UNIT 11.3). At 100 to 200 mM monovalent counterion concentration, the time constants for tRNA tertiary structure formation are typically in the range of 10 to 100 msec. Faster signals at higher temperature correspond to melting of hairpin helices, which can be identified by a combination of NMR and relaxation kinetics (Crothers et al., 1974; Hilbers et al., 1976). Imino proton exchange of hairpin helices results in line broadening of the corresponding resonances when the helix lifetime is less than ∼5 msec (Crothers et al., 1974). The measured temperature dependence of the helix dissociation rate from relaxation kinetics can be extrapolated to lower temperatures, to match a 5 msec lifetime to a temperature zone for broadening of a group of resonances corresponding to a specific helix. In this way, the structural origin of the relaxation signals can
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be established (Crothers et al., 1974; Hilbers et al., 1976). Another important clue for determining the structural origin of relaxation signals is derived from the fortuitous presence of 4-thiouridine (S4U) bases in natural tRNAs. For example, the S4U residue at position 8 in tRNAfMet, between the acceptor stem and DHU helices, shows a large hyperchromism in the tertiary melting transition (Cole and Crothers, 1972; Cole et al., 1972). Modern synthetic methods, which permit substitution of modified bases at any position, should allow examination of the dynamics of specific sites in any given RNA using absorbance or fluorescence detection. Two important methods for resolving RNA folding in space and time have been applied to the Group I ribozyme from Tetrahymena thermophila. This RNA has played a pivotal role in the field of RNA catalysis, starting with the report by Cech and co-workers (Kruger et al., 1982) that Tetrahymena pre-rRNA is capable of excising its own intron. The ribozyme can be split into two domains, one of which, P4-P6, can fold independently, while the other, P3-P9, requires P4-P6 for formation of its native tertiary structure (Doherety and Doudna, 1997). Both the intrinsic interest of the system and the potential for in-depth analysis of the kinetic measurements has been greatly increased by solution of the crystal structure of the P4-P6 subunit of the ribozyme (Cate et al., 1996). Given the size and complex folding pattern of this RNA, it was necessary to develop new methods in order to analyze the kinetic properties at nucleotide resolution. Sclavi et al. (1998) described the application to RNA folding of a rapid nucleic acid footprinting method that uses a synchrotron source to generate a burst of hydroxyl radicals by radiolysis of water. The X-ray source is combined with a quenched-flow device that allows folding to be triggered by mixing with buffer containing Mg2+ ions, followed by a variable time delay before application of the X-ray pulse. Local kinetic parameters can be determined for any nucleotide whose hydroxyl radical reactivity changes upon Mg2+ addition on a time scale substantially slower than the fastest resolvable time, i.e.,∼10 msec. Another important technical innovation, employed by Williamson and co-workers, is the use of deoxyoligonucleotide hybridization probes and RNase H cleavage to monitor accessibility of regions of the RNA sequence that are blocked by tertiary structure formation (Zarrinkar and Williamson, 1994; Treiber et al.,
1998). In this method the reaction is initiated by Mg2+ addition, followed after a variable time delay by addition of the deoxyoligonucleotide probe and RNase H. The kinetics of processes that occur on a time scale of ∼1 min or longer can be resolved, allowing examination of tertiary structure formation. A general conclusion of these and other folding studies on the Tetrahymena ribozyme is that formation of the final conformation is slowed by the necessity for disrupting a competing structure.
CONFORMATIONAL SWITCHING It is often proposed that RNA switches between two or more stable conformations in the course of its biological function. Study of such processes probably has more biological significance than in vitro examination of the rate of folding an intact transcript, which starts from very different boundary conditions than found in the cell, where folding events accompany RNA synthesis. In addition, cellular proteins are likely to affect the pathway and rate of folding reactions (Weeks and Cech, 1995). Only a few systems of this kind have been intensively studied. For example, the kinetics of a conformational switch that involves two competing secondary structures in a spliced leader RNA was studied by LeCuyer and Crothers (1994). Using stopped-flow and T-jump methods, the authors evaluated the rate constants for the switch, which has a time constant on the order of 100 msec. The estimated activation energy was ∼25 kcal/mol, considerably less than would be expected if the mechanism involved dissociation of the 13- to 14-bp of structure 1 before structure 2 begins to form. To explain the results, a branch migration mechanism was proposed, in which the alternate helix is nucleated and grows while the competing structure shrinks. An example of an even slower conformational switch, whose detailed structural basis is not obvious but probably involves tertiary interactions, comes from the work of Gluick et al. (1997). They observed two electrophoretically distinguishable conformations of a 112-nt fragment containing the first ribosome initiation site of E. coli α mRNA. The corresponding relaxation time for the conformational switch is ∼2000 sec (∼30 min), or ∼2 × 104 fold slower than the spliced leader RNA switch. The RNA conformational switch most intensively studied to date is the reorganization of the P1, or substrate (because it contains the splice site), helix, and its docking as part of the tertiary structure of the Tetrahymena ribozyme.
RNA Folding Pathways
11.1.3 Current Protocols in Nucleic Acid Chemistry
Supplement 2
The early stopped-flow experiments of Turner and associates (Bevilacqua et al., 1992) used fluorescently labeled substrate oligonucleotides to form the P1 helix by hybridization with the IGS in trans. Two reaction steps could be resolved kinetically, which were interpreted as formation of the P1 helix followed by the slower docking reaction. Woodson and colleagues adopted the strategy of providing the substrate in cis, forming pre-rRNA, and examined the ability of alternate substrate secondary structures to affect the rate of folding (Emerick and Woodson, 1994; Pan and Woodson, 1999). A fraction of the RNA usually renatures rapidly to the form active in splicing, but a variable portion of primary transcripts renatured at physiological temperatures is found in inactive states. The active form of pre-rRNA has the highest electrophoretic mobility, corresponding to a compact structure. The inactive form is found to have an alternate structure, called alt P3 instead of P3, in the catalytic core of the ribozyme. The presence of this non-native structure is coupled with formation of an alternate hairpin helix, P(-1), in the 5′ exon (Pan and Woodson, 1999). The switch to the P1-P3 form from the P(-1)-alt P3 structure limits the rate of folding of the P3-P9 domain. The P(-1) structure is modeled as a 12-bp dumbbell helix, whereas P(1) is a 9-bp hairpin (Pan and Woodson, 1999). The activation energy for the folding reaction, ∼12 kcal/mol, is much smaller than would be expected if the mechanism involved full melting of incorrect structure before initiating formation of the native form. In this case, as in the spliced leader RNA switch (LeCuyer and Crothers, 1994), it is likely that some form of branch migration pathway is involved, allowing coexistence of portions of each structure in the transition state. An impressive degree of mechanistic detail at the molecular level has been achieved for Tetrahymena pre-rRNA and its corresponding ribozyme. However, it is also striking that a very significant effort by several laboratories was necessary to reach this level of understanding. The methods that have been developed will aid in attacking other RNA folding/switching problems, but present indications are that these systems are idiosyncratic, making each problem a new one and posing a major challenge to formulation of simplifying generalizations.
LITERATURE CITED Bevilacqua, P.C., Kierzek, R., Johnson, K.A., and Turner, D.H. 1992. Dynamics of ribozyme binding of substrate revealed by fluorescence-detected stopped-flow methods. Science 258:13551358. Breslauer, K.J., Frank R., Blocker, H., and Marky, L.A. 1986. Predicting DNA duplex stability from the base sequence. Proc. Natl. Acad. Sci. U.S.A. 83:3746-3750. Brion, P. and Westhof, E. 1997. Hierarchy and dynamics of RNA folding. Annu. Rev. Biophys. Biomol. Struct. 26:113-137. Cate, J.H., Gooding, A.R., Podell, E., Zhou, K., Golden, B.L., Kundrot, C.E., Cech, T.R., and Doudna, J.A. 1996. Crystal structure of a group I ribozyme domain: Principles of RNA packing. Science 273:1678-1685. Chen, J.L., Blasco, M.A., and Greider, C.W. 2000. Secondary structure of vertebrate telomerase RNA. Cell 100:503-514. Cole, P.E. and Crothers, D.M. 1972. Conformational changes of transfer ribonucleic acid. Relaxation kinetics of the early melting transition of methionine transfer ribonucleic acid (Escherichia coli). Biochemistry 11:4368-4374. Cole, P.E., Yang, S.K., and Crothers, D.M. 1972. Conformational changes of transfer ribonucleic acid. Equilibrium phase diagrams. Biochemistry 11:4358-4368. Crothers, D.M., Cole, P.E., Hilbers, C.W., and Shulman, R.G. 1974. The molecular mechanism of thermal unfolding of transfer RNA. J. Mol. Biol. 87:63-88. Doherty, E.A. and Doudna, J.A. 1997. The P4-P6 domain directs higher order folding of the Tetrahymena ribozyme core. Biochemistry 36:31593169. Draper, D.E. and Gluick, T.C. 1995. Melting studies of RNA unfolding and RNA-ligand interactions. Methods Enzymol. 259:281-305. Emerick, V.L. and Woodson, S.A. 1994. Fingerprinting the folding of a group I precursor RNA. Proc. Natl. Acad. Sci. U.S.A. 91:9675-9679. Farris, A.D., Koelsch, G., Pruijn, G.J., van Venrooij, W.J., and Harley, J.B. 1999. Conserved features of Y RNAs revealed by automated phylogenetic secondary structure analysis. Nucl. Acids Res. 27:1070-1078. Gluick, T.C., Gerstner, R.B., and Draper, D.E. 1997. Effects of Mg2+, K+, and H+ on an equilibrium between alternative conformations of an RNA pseudoknot. J. Mol. Biol. 270:451-463. Harris, M.E. and Pace, N.R. 1995-1996. Analysis of the tertiary structure of bacterial RNase P RNA. Mol. Biol. Rep. I22:115-123. Hilbers, C.W., Robilard, G.T., Shulman, R.G., Blake, R.D., Webb, P.K., Fresco, R., and Riesner, D. 1976. Thermal unfolding of yeast glycine transfer RNA. Biochemistry 15:1874-1882.
RNA Folding Pathways
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Current Protocols in Nucleic Acid Chemistry
Kruger, K., Grabowski, P.J., Zaug, A.J., Sands J., Gottschling, D.E., and Cech, T.R. 1982. Selfsplicing RNA: Autoexcision and autocyclization of the ribosomal RNA intervening sequence of Tetrahymena. Cell 32:147-157. LeCuyer, K.A. and Crothers, D.M. 1994. Kinetics of an RNA conformational switch. Proc. Natl. Acad. Sci. U.S.A. 91:3373-3377. Maglott, E.J. and Glick, G.D. 1997. A new method to monitor the rate of conformational transitions in RNA. Nucl. Acids Res. 25:3297-3301. Matthews, D.H., Sabina, J., Zucker, M., and Turner, D.H. 1999. Expanded sequence dependence of thermodynamic parameters improves prediction of RNA secondary structure. J. Mol. Biol. 288:911-940. Noller, H.F., Kop, J., Wheaton, V., Brosius, J., Gutell, R.R., Kopylov, A.M., Dohme, F., Herr, W., Stahl, D.A., Gupta, R., and Waese, C.R. 1981. Secondary structure model for 23S ribosomal RNA. Nucl. Acids Res. 9:6167-6189. Pan, J. and Woodson, S.A. 1999.The effect of longrange loop-loop interactions on folding of the Tetrahymena self-splicing RNA. J. Mol. Biol. 294:955-965. Riesner, D., Romer, R., and Maass, G. 1969. Thermodynamic properties of the three conformational transitions of alanine specific transfer RNA from yeast. Biochem. Biophys. Res. Commun. 35:369-376. Riesner, D., Romer, R., and Maass, G. 1970. Kinetic study of the three conformational transitions of alanine specific transfer RNA from yeast. Eur. J. Biochem. 15:85-91.
Riesner, D., Maass, G., Thiebe, R., Philippsen, P., and Zachau, H.G. 1973. The conformational transitions in yeast tRNAPhe as studied with tRNAPhe fragments. Eur. J. Biochem. 36:76-88. Sclavi, B., Sullivan, M., Chance, M.R., Brenowitz, M., and Woodson, S.A. 1998. RNA folding at millisecond intervals by synchrotron hydroxyl radical footprinting. Science 279:1940-1943. Tinoco, I., Jr. and Bustamante, C. 1999. How RNA folds. J. Mol. Biol. 293:271-281. Treiber, D.K., Rook, M.S., Zarrinkar, P.P., and Williamson, J.R. 1998. Kinetic intermediates trapped by native interactions in RNA folding. Science 279:1943-1946. Weeks, K.M. and Cech, T.R. 1995. Protein facilitation of group I intron splicing by assembly of the catalytic core and the 5′ splice site domain. Cell 82:221-230. Wu, M. and Tinoco, I., Jr. 1998. RNA folding causes secondary structure rearrangement. Proc. Natl. Acad. Sci. U.S.A. 95:11555-11560. Xia, T., SantaLucia, J., Jr., Burkard., M.E., Kierzek, R,. Schroeder, S.J., Jiao, X., Cox, C., and Turner, D.H. 1998. Thermodynamic parameters for an expanded nearest-neighbor model for formation of RNA duplexes with Watson-Crick base pairs. Biochemistry 37:14719-14735. Zarrinkar, P.P. and Williamson, J.R. 1994. Kinetic intermediates in RNA folding. Science 265:918924.
Contributed by Donald M. Crothers Yale University New Haven, Connecticut
RNA Folding Pathways
11.1.5 Current Protocols in Nucleic Acid Chemistry
Supplement 2
RNA Secondary Structure Prediction
UNIT 11.2
This unit details the steps for predicting the secondary structure of an RNA sequence and for predicting the equilibrium binding affinity of a complementary RNA or DNA oligonucleotide to an RNA target (see Basic Protocol 2). Two protocols are given for secondary structure prediction—one for the Windows computer program RNAstructure (see Basic Protocol 1) and another for the mfold Webserver (see Basic Protocol 3). The mfold server is an internet adaptation of the mfold package for Unix computers. This package is available for use on Unix platforms and is described elsewhere (Zuker et al., 1999; http://www.rpi.edu/∼zukerm/). RNAstructure is for personal computers (PCs) with Microsoft Windows and also includes OligoWalk for predicting oligonucleotide binding affinities (see Basic Protocol 2). The mfold server is accessed via the internet using a standard browser (e.g., Microsoft Internet Explorer, Apple Safari, or Mozilla Firefox). Both programs have similar structure prediction accuracy. The minimum free energy structure and a set of suboptimal structures with low free energies are predicted by both programs. The mfold program can predict secondary structures for circular RNAs, but RNAstructure cannot. RNAstructure includes the ability to constrain secondary structure prediction using chemical modification data, but mfold does not.
PREDICTING SECONDARY STRUCTURE AND PREDICTING BASE-PAIR PROBABILITIES
BASIC PROTOCOL 1
Secondary structure prediction by free energy minimization is the core functionality of RNAstructure. The secondary structure prediction algorithm predicts the lowest free energy structure, which is the single best prediction of the secondary structure. It also predicts low free energy structures, called suboptimal structures, which suggest possible alternative structures (Zuker, 1989). Low free energy structures can be color-annotated according to base-pair probabilities determined by a partition function calculation, and these probabilities imply confidence in the prediction of pairs (Mathews, 2004). This protocol describes the basic use of the program on a Microsoft Windows platforom. Many other options are available, and these are described in detail in the online help manual, which can be accessed from the program by choosing the Help Topics item from the Help menu. The protocol requires some basic familiarity with Windows. RNAstructure is known to run on Apple Macintosh computers with Virtual PC and on the Linux operating system using WINE. However, RNAstructure has not been extensively tested under those operating systems. Instructions for installing RNAstructure on Linux and Mac OS X are beyond the scope of this unit, but the use of the program is identical.
Materials Hardware: A personal computer running Microsoft Windows is required. RNAstructure is compatible with Windows XP, Windows 2000, and Windows ME. RNAstructure will be kept up-to-date when Windows Vista is released. Table 11.2.1 shows the computation times and memory requirements for secondary structure prediction and partition function calculation as a function of sequence length. Table 11.2.1 should be consulted to determine the memory (RAM) requirement for the sequence of interest. Software: The RNAstructure package must be downloaded at http://rna.urmc. rochester.edu. User registration is requested so that e-mail can be sent when significant upgrades are available or if significant bugs are found. The list of registered users is not shared with others. RNA Folding Pathways Contributed by David H. Mathews, Douglas H. Turner, and Michael Zuker Current Protocols in Nucleic Acid Chemistry (2007) 11.2.1-11.2.17 C 2007 by John Wiley & Sons, Inc. Copyright
11.2.1 Supplement 28
Table 11.2.1 Time and Memory Requirements for Secondary Structure Prediction and Base-Pair Probability Prediction Using RNAstructurea
Time for structure prediction
RR1664 tRNA
77
<1 sec
15
<1 sec
28
Bacillus stearothermophilus SRP RNA
268
3 sec
15
5 sec
34
Tetrahymena thermophila group I intron
433
11 sec
17
16 sec
40
Saccharomyces cerevisiae A5 group II intron
631
36 sec
18
49 sec
51
E. coli 16S rRNA
1542
8 min
34
12 min
148
E. coli 23S rRNA
2904
55 min
152
105 min
440
Sequence
Memory for Time for base-pair structure probability prediction (Mb) calculation
Memory for base-pair probability calculation (Mb)
Length (nucleotides)
a Calculations were performed on a computer with a 3.4 GHz Pentium 4 processor and 1 Gb of RAM, running Microsoft Windows XP.
Calculation times are less with a faster processor or with more memory and slower with a slower processor or less memory. The calculation time scales according to O(N3 ), where N is the length of the sequence. Therefore, doubling the length of the sequence requires roughly eight times the calculation time. Memory use scales according to O(N2 ), so doubling the length of the sequence required roughly four times the RAM. In practice, the time and memory scaling deviate from those expected because there is a baseline time and memory cost associated with each calculation.
Install RNAstructure 1. Download RNAstructure as RNAstructure.zip. Windows XP can open the .zip file directly. Windows 2000 and Windows ME require the use of another program to open the zip file, such as WinZip (http://www.winzip.com) or WinRAR (http://www.rarlab.com). Open RNAstructure.zip and doubleclick the file setup.exe. This will launch the installation routine. During the installation, the user can choose the installation destination. Most users will want to accept the default location. The installation will add RNAstructure to the start menu in the folder RNAstructure X.Y, where X.Y is the current version of the software. Enter a sequence 2. Start RNAstructure by choosing it from the Windows start menu. Open the sequence editor by choosing New Sequence at the top of the File menu. Figure 11.2.1 shows the layout of the sequence editor. The name or a short description of the sequence can be entered in the field labeled Title. This information will be displayed with predicted structures. The field labeled Comment is available for entering longer comments that are stored with the sequence, but will not be shown with the predicted structure. The sequence is entered in the field labeled Sequence.
3. Sequences can be entered manually or pasted from another program. To paste a sequence, copy it to the clipboard, click with the mouse in the Sequence field, then choose Paste from the Edit menu option or press Ctrl-V. Save the sequence by choosing Save Sequence from the File menu.
RNA Secondary Structure Prediction
The sequence should consist of A (adenine), C (cytidine), G (guanine), U (uridine), T (treated as uridine in RNA folding), and X (a nucleotide that neither base-pairs nor stacks). Note that nucleotides entered as lowercase characters are forced single-stranded in the structure prediction, so most nucleotides should be uppercase. Spaces and carriage returns are ignored during structure prediction.
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Current Protocols in Nucleic Acid Chemistry
Figure 11.2.1 Screen shot of the RNAstructure sequence editor. The D. melanogaster 5S rRNA sequence is entered and formatted using the Format Sequence button.
The sequence editor has several features. Clicking the Format Sequence button on the editor window will automatically format the sequence into lines of 50 nucleotides with a space after every fifth nucleotide. The sequence is recited aloud over speakers (if available) when the Read Sequence button is clicked. This provides a convenient method for checking the accuracy of a manually entered sequence. Clicking the Fold as RNA button saves the sequence and opens the secondary structure prediction window. Clicking Fold as DNA saves the sequences and opens a structure prediction window that predicts secondary structure of a DNA sequence.
Predict the secondary structure 4. After saving the input sequence (above), choose Fold RNA Single Strand from the File menu. The RNA secondary structure prediction window will open, as shown in Figure 11.2.2. 5. Choose the name of the sequence file by clicking the Sequence File button. This will open a standard Windows Open File dialog box for selecting the file. After the file has been selected, the name of the file will appear next to the Sequence File button, and default values will have been entered in all other fields. The output of the calculation (a predicted secondary structure and a set of low free energy structures) will be stored in a CT (connection table) file. The default CT file is the name of the sequence file chosen, but with a .ct file extension. The default name can be changed by clicking the CT File button. Three parameters control the generation of suboptimal structures. The first parameter is maximum percent energy difference of suboptimal structures as compared to the lowest free energy structure. Larger percent energy differences allow the prediction of more suboptimal structures, whereas a maximum percent of zero allows only prediction of the lowest free energy structure. The maximum percent energy difference can be changed from the default by manually typing in the text box adjacent to Max % Energy Difference.
RNA Folding Pathways
11.2.3 Current Protocols in Nucleic Acid Chemistry
Supplement 28
Figure 11.2.2
Screen shot of the RNA secondary structure prediction window.
The second parameter is the maximum number of structures. This places an absolute limit on the number of suboptimal structures and can be changed manually by typing in the text box adjacent to Max Number of Structures. The third parameter is the window size, which specifies how different each suboptimal structure must be, as compared to all other predicted structures. Each structure must have at least window number of pairs separated by at least window nucleotides from all pairs in all other structures. A window size of zero does not place any restriction. Larger window sizes result in structures with greater difference, but also result in fewer predicted structures. The default window parameter can be manually changed by typing in the text box adjacent to Window Size. The check box next to Generate Save File is checked by default. This will generate a Save file that can be used to show energy dot plots or to predict a different set of suboptimal structures using different suboptimal structure parameters. The online help contains entries labeled Dot Plot and Refolding a Saved Sequence that explain these functions. The name of the Save file is the same as the CT file, except that the file has a .sav extension instead of .ct.
6. RNAstructure can predict secondary structures with user-specified constraints. These are entered by choosing the appropriate menu item under the Force menu.
RNA Secondary Structure Prediction
a. Base Pair is used to specify required base pairs in the structure. b. Chemical Modification is used to specify nucleotides that are accessible to chemical modification, i.e., single-stranded, at the end of a helix, or in or adjacent to a GU pair. c. Double Stranded is used to specify nucleotides that must be base-paired, without specifying to which nucleotides they are paired. d. FMN Cleavage is used to indicate Us that are in GU base pairs. e. Single Stranded is used to indicate unpaired nucleotides. f. Prohibit Basepairs is used to indicate specific base pairs that are not allowed.
11.2.4 Supplement 28
Current Protocols in Nucleic Acid Chemistry
Figure 11.2.3 The lowest free energy secondary structure predicted for the D. melanogaster 5S rRNA sequence as drawn by RNAstructure.
Each of these options opens a dialog box for entering the specified constraints. In the dialog box, “OK” can be clicked to keep the dialog box open to enter more constraints, “Cancel” can be clicked in order not to record the constraints, or “OK and Close” can be clicked when all constraints are entered into the dialog box. The entered constraints are displayed on the screen if Current is chosen from the Force menu. Reset removes all entered constraints. Save Constraints can be used to save all constraints to a file (with a .con extension). Constraints can be read from a file by choosing Restore Constraints.
7. The default temperature of the prediction, 37◦ C, can be changed by choosing the temperature menu item. This opens a dialog box in which the desired temperature of structure prediction can be entered in Kelvin. The stability of RNA secondary structure is best determined at 37◦ C and is poorly determined at temperatures far from 37◦ C (Lu et al., 2006). For temperatures <20◦ C or >40◦ C, a more accurate secondary structure can be predicted by using the default folding temperature of 37◦ C.
8. Click the button labeled Start to begin the secondary structure prediction calculation. A window will open with a progress indicator. When the calculation ends, a dialog box opens with the options Draw Structures and Cancel. Click Draw Structures to display the predicted secondary structures. Figure 11.2.3 shows the secondary structure predicted for the D. melanogaster 5S rRNA sequence (with no user-specified constraints). The functionality of the RNAstructure drawing program is detailed in the later steps of this protocol.
Predict base-pair probabilities with a partition function calculation 9. Open the partition function window by choosing Partition Function RNA from the File menu. Choose the sequence name by clicking the Sequence Name button. For the example of the D. melanogaster 5S rRNA, choose the sequence file that contains that sequence. After the calculation, the base-pair probability data are saved to disk
RNA Folding Pathways
11.2.5 Current Protocols in Nucleic Acid Chemistry
Supplement 28
in a Save (.pfs) file. A default Save file name automatically appears in the field next to the Save File button after a sequence has been chosen. The default name can be changed by clicking the Save File button. 10. Enter constraints and change the temperature to that used for secondary structure prediction. The partition function calculation can accommodate the same structure restraints as secondary structure prediction (step 6). The partition function can also be performed at temperatures other than the default 37◦ C in the same way as structure prediction (step 7). To get the most information from base pair probabilities, these settings should be set the same as for structure prediction.
11. Start the calculation by clicking the button labeled Start. A window will open to show the progress of the calculation. When it is completed, a probability dot plot will be displayed that indicates the probability of all valid canonical base pairs (AU, GC, and GU). First, reduce the probability range of base pairs by choosing Plot Range under the Draw menu option. This will open a dialog box in which a min and max range are entered. Dots are registered by –log10 (Base Pair Probability). Set the maximum to 2, so that all base pairs with pairing probability >0.01 (1%) will be displayed. Click “OK.” The plot will now resemble the plot in Figure 11.2.4.
RNA Secondary Structure Prediction
Figure 11.2.4 The probability dot plot for base pairs in the D. melanogaster 5S rRNA sequence. Each dot in this plot is a single base pair and is associated with a base-pairing probability. For pairs between nucleotides i and j, with i < j, i is down the right-hand side of the plot and j is across the top. In RNAstructure, dots are colored to indicate a probability interval, with the most probable pairs in red and the least probable in blue. The current base pairs displayed have pairing probability of ≥1%. A color key is drawn at the bottom of the window (not shown in this view). After clicking on a dot, the pair and –log10 (base-pair probability) are indicated at the bottom of the RNAstructure window. The current display shows that the base pair between G75 and C102 was clicked and has a –log10 (base-pair probability) of 5.33232 × 10−2 (88.4%). For a color version of this figure, see http://www.currentprotocols.com.
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Current Protocols in Nucleic Acid Chemistry
The probability dot plot window provides several features for analyzing the dot plot information. By clicking on a dot, the message window at the bottom of RNAstructure provides the identity of the base pair and –log10 of the base-pair probability. The size of the plot in the window can be changed by choosing Zoom under the Draw menu. Alternatively, pressing Ctrl-left arrow zooms out and pressing Ctrl-right arrow zooms in. The base-pair probabilities can be written to a tab-delimited text file for analysis with other programs by choosing Output Text File under the Output menu option. The text file contains the –log10 of base-pair probability for all base pairs and not just the pairs that are currently displayed on the screen. Secondary structures composed of only probable base pairs can be output by choosing Output Probable Structure under the Output menu option. The probability dot plot window is created from the data stored in the Save (.pfs) file. To draw the probability dot plot again at a later time, choose Dot Plot Partition Function from the File menu. This will open a Windows file dialog box from which the Save file can be chosen.
Color annotate the predicted secondary structures according to base-pair probabilities 12. Return to the structure drawing window that contains the predicted secondary structures for the D. melanogaster 5S rRNA sequence. If the window has been closed, the secondary structure can be redrawn using the data stored in the CT file. A new drawing window can be opened by choosing Draw under the File menu. This will open a Windows file dialog box from which the CT file can be chosen. 13. To color annotate a predicted secondary structure for which a partition function calculation has also been performed, choose Add Color Annotation from the Draw menu with the drawing window open. This will open a Windows file dialog box from which the Save (.pfs) file can be selected. The secondary structure will then have color annotation with the most probable base pairs in red and the least probable pairs in violet. To show a key that indicates the association between color and probability range, choose Show Color Annotation Key under the Draw menu. Figure 11.2.5 shows the predicted lowest free energy structure for the D. melanogaster 5S rRNA with color annotation. The drawing window has several functions to facilitate the analysis of secondary structures. When suboptimal structures are included in the CT file, the displayed structure can be changed by choosing Structure Number under the Draw menu. This will open a window that indicates the current structure number and allows that number to be changed. The lowest free energy structure is structure 1, and folding free energy increases with the structure number. Alternatively, pressing Ctrl-up arrow increases the number of the currently displayed structure and pressing Ctrl-down arrow lowers the number of the currently displayed structure. The number of the currently displayed structure is indicated in the upper left-hand corner of the window. As indicated in Figures 11.2.3 and 11.2.5, there are a total of three low-energy secondary structures predicted for the D. melanogaster 5S rRNA using the default parameters. The size of the structures on the screen can be changed by choosing Zoom from the Draw menu. Alternatively, pressing Ctrl-left arrow zooms out and pressing Ctrl-right arrow zooms in.
14. To produce publication-quality drawings of secondary structures, base pairs predicted by RNAstructure can be exported to the drawing program XRNA, which is available for free download from the Santa Cruz RNA Center at http://rna.ucsc.edu/rnacenter/xrna/xrna.html. XRNA is Java-based and will run on most computers. To export helix locations in a file that is readable by XRNA, choose Export Structure to Text File from the Draw menu. RNA Folding Pathways
11.2.7 Current Protocols in Nucleic Acid Chemistry
Supplement 28
Figure 11.2.5 The lowest free energy secondary structure predicted for the D. melanogaster 5S rRNA sequence with color annotation. The color annotation key is shown in the lower right-hand corner. The most probable base pairs are red and least probable are violet. The predicted pairs with the highest base pair probabilities are more likely to be correctly predicted pairs (Mathews, 2004). For a color version of this figure, see http://www.currentprotocols.com.
BASIC PROTOCOL 2
PREDICTING BINDING AFFINITIES OF OLIGONUCLEOTIDES COMPLEMENTARY TO AN RNA TARGET WITH OLIGOWALK RNAstructure includes the OligoWalk program for predicting the binding affinity of complementary oligonucleotides to an RNA target. For an RNA sequence of N nucleotides, OligoWalk predicts an overall free energy of binding of all (N – L + 1) oligonucleotides of length L that are complementary to the target. Hence, the binding region considered is walked down the length of the sequence. The overall free energy of binding, G◦37 overall includes the effects of self structure in the target and self structure in the oligonucleotides. This protocol uses RNAstructure on a Microsoft Windows platform. RNAstructure is also known to run on Apple Macintosh computers with Virtual PC and on the Linux operating system using WINE. However, RNAstructure has not been extensively tested under those operating systems. Instructions for installing RNAstructure on Linux and Mac OS X are beyond the scope of this unit, but the use of the program is identical.
Materials Hardware: A personal computer running Microsoft Windows is required. RNAstructure is compatible with Windows XP, Windows 2000, and Windows ME. RNAstructure will be kept up to date when Windows Vista is released. Table 11.2.2 indicates typical calculation times for two different length target sequences. RNA Secondary Structure Prediction
Software: The RNAstructure package must be downloaded at http://rna.urmc. rochester.edu. User registration is requested so that e-mail can be sent when significant upgrades are available or if significant bugs are found. The list of registered users is not shared with others.
11.2.8 Supplement 28
Current Protocols in Nucleic Acid Chemistry
Table 11.2.2 Time and Memory Requirements for OligoWalk Calculationsa
Target length
Oligonucleotide length
Saccharomyces cerevisiae A5 group II intron
631
Saccharomyces cerevisiae A5 group II intron
Target
Mode
Time
Memory (Mb)
10
Break Local Structure; Do Not Include Suboptimal Structures
2 sec
21
631
20
Break Local Structure; Do Not Include Suboptimal Structures
5 sec
21
Saccharomyces cerevisiae A5 group II intron
631
20
Break Local Structure; Include Suboptimal Structures
9 sec
21
Saccharomyces cerevisiae A5 group II intron
631
20
Do Not Consider Target Structure
4 sec
21
Saccharomyces cerevisiae A5 group II intron
631
20
Refold Whole RNA; Include Suboptimal Structures
6 hr, 17 min
26
E. coli 16S rRNA
1542
20
Break Local Structure; Include Suboptimal Structures
39 sec
25
a For calculations that break local structure or do not consider target structure, the calculation scales according to O(NL3 ) where N is the length of the target and L is the length of the oligonucleotide. Doubling the length of the target only doubles the length of the calculation. A doubling in the length of the oligonucleotide requires eight times as much calculation time. For calculations in which the whole target RNA is refolded for each oligonucleotide, the calculation scales roughly according to O(N4 ). A doubling in the length of the target requires 16 times as much computation time, but lengthening the oligonucleotide results in little change in the calculation time.
Install RNAstructure 1. Install RNAstructure and predict the secondary structure of the target RNA (see Basic Protocol 1). Start the OligoWalk calculation 2. Open the OligoWalk input window (shown in Fig. 11.2.6) by choosing OligoWalk from the File menu. 3. Click on the button labeled CT File and choose the target secondary structure (stored in CT format; see Basic Protocol 1) using the Windows file dialog box. A default name is then chosen for output of the thermodynamic parameters. This name is the same as the CT file, but with the .ct extension changed to .rep. The default name can be changed by clicking on the button labeled Report File. The report file stores the calculated parameters as tab-delimited text that can be opened in most spreadsheet programs, such as OpenOffice or Microsoft Excel.
4. Choose a Mode for the calculation by clicking the button (see Fig. 11.2.6) adjacent to one of the three options: Break Local Structure, Refold Whole RNA for Each Sequence, or Do Not Consider Target Structure. Do Not Consider Target Structure is the fastest mode, but is not recommended because the target RNA secondary structure is neglected. Refold Whole RNA for Each Sequence predicts a new lowest free energy structure after oligonucleotide binding by predicting a structure with the nucleotides that are bound by the oligonucleotide forced to be unpaired. This mode is the slowest, but best approximates equilibrium. Break Local Structure is much faster than Refold Whole RNA for Each Sequence, and it calculates the secondary structure formation free energy change of the original structure with any pairs
RNA Folding Pathways
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Supplement 28
Figure 11.2.6
The OligoWalk input window.
that involve the oligonucleotide-bound nucleotides broken. For most applications, Break Local Structure is a good compromise between accuracy and calculation time.
5. Below the Mode options (see Fig. 11.2.6), note the check box labeled Include Target Suboptimal Structures in Free Energy Calculation. When checked, the cost (in free energy change) of opening base pairs in the target structure is calculated for each suboptimal secondary structure. The contribution made by each suboptimal structure to the total cost of opening target self structure is weighted according to the folding free energy of each structure. In general, it is recommended that this box be checked, so that alternative possible secondary structures are considered.
Input information about the oligonucleotides 6. Enter the oligonucleotide length in the text box to the right of Oligo Length (see Fig. 11.2.6). Oligonucleotides can be either RNA or DNA; the choice between these is indicated by the radio button in the Oligomer Chemistry box. Also choose an oligonucleotide concentration. The concentration units can be changed between mM, µM, nM, and pM by clicking the down arrow to the right of the displayed units. For the example shown in Figure 11.2.6, DNA nucleotides of length 18 are considered at a concentration of 1 µM.
RNA Secondary Structure Prediction
7. Next, the region for oligonucleotide binding can be reduced by adjusting the Start and Stop locations (see Fig. 11.2.6, bottom of dialog box). Start refers to the target RNA nucleotide bound to the 3 end of the first oligonucleotide, and Stop refers to the nucleotide bound to the 5 end of the last oligonucleotide in the walk. By default, these limits are set to the 5 and 3 ends of the target sequence. To adjust the limits, use the up and down arrows next to the value fields. Limiting the area of interest on the target RNA strand reduces the calculation time.
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Figure 11.2.7 The OligoWalk output display. For a color version of this figure, see http://www. currentprotocols.com.
8. The temperature of the calculation can be changed from the default of 37◦ C by choosing the Temperature menu item. This opens a dialog that takes the temperature in Kelvin. 9. Finally, start the calculation by clicking the Start OligoWalk button. A window will open to show the progress of the calculation. When the calculation is complete, RNAstructure will open the OligoWalk output window as shown in Figure 11.2.7.
Navigating the OligoWalk output window 10. The OligoWalk output window provides an interactive method for displaying the calculated thermodynamic parameters. The target sequence is drawn left to right across the window in a 5 to 3 direction. Red nucleotides are predicted to be basepaired in the lowest free energy structure. Black nucleotides are predicted to be single stranded. The currently displayed oligonucleotide is above the target sequence in a 3 to 5 direction. The position along the target of the currently displayed oligonucleotide is indicated in the upper left-hand corner of the display. Oligonucleotides are numbered according to the 5 -most binding nucleotide in the target; therefore, the oligonucleotides are numbered from 1. Below the current oligonucleotide is the backbone chemistry (RNA or DNA) and concentration of the oligonucleotide. 11. The thermodynamic parameters at the top of the screen are free energy changes at 37◦ C in kcal/mol. “Overall G◦37 ” is the total free energy change of binding for a structured target and structured oligonucleotide. “Duplex G◦37 ” is the free energy change of duplex formation between the oligonucleotide and target, without the cost of opening self-structure. “Break Targ. G◦37 ” is the free energy cost of opening target secondary structure. “Oligo Self G◦37 ” and “Oligo-Oligo G◦37 ” are the free energy costs of opening unimolecular and bimolecular self-structure in the
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oligonucleotide, respectively. The Tm is the melting temperature of duplex formation in ◦ C, not accounting for self-structure in target or oligonucleotide. The graph, by default, shows the “Overall G◦37 ” and “Duplex G◦37 ” profile along the target sequence. The graph color is identical to the text color above, except that the color for “Overall G◦37 ” for the current oligonucleotide is in red. The free energy term that is graphed can be changed by selecting Free Energy under the Graph menu option. Each of the free energy terms can be graphed, and a check on the menu shows the current selection.
12. The currently displayed oligonucleotide can be changed in several ways. The left and right arrow keys move the displayed oligonucleotide 5 and 3 , respectively. This can also be done by clicking the buttons labeled < or >. The currently displayed nucleotide is skipped ten nucleotides by clicking the buttons labeled or . By clicking the Go button, a navigation window is reached. In the navigation window, a specific oligonucleotide for display can be indicated, or a button labeled Most Stable can be clicked to display the oligonucleotide with the tightest binding. 13. For oligonucleotides with self structure, the self structure can be drawn on the screen by double-clicking the oligonucleotide sequence. For oligonucleotides with both bimolecular and unimolecular structure, a window opens to allow user selection of the structure type to display. BASIC PROTOCOL 3
PREDICTING A SECONDARY STRUCTURE WITH THE mfold SERVER The mfold server takes advantage of the Web to make RNA secondary structure prediction available to a large audience. It requires no special setup and is much more user-friendly than the unadorned Unix version of mfold. This protocol outlines the steps involved in predicting a secondary structure on the mfold server using a standard browser, e.g., Microsoft Internet Explorer, Mozilla Firefox, or Apple Safari. This protocol assumes basic familiarity with the Web.
Access the server and enter the required information The mfold server is accessed at http://www.bioinfo.rpi.edu/applications/mfold/rna/form1. cgi. This page gathers all the necessary information required to predict an RNA secondary structure. There are many fields for entering information; each is labeled, and many of these labels link to explanations of the information required in that field. This protocol describes the information that is required, starting at the top of the page and working down. Be sure to follow the link that reads “Notice” at the top of the page. This provides information to mfold users on accessing results, how long results are saved, and security on the server. Other links at the top of the page lead to an RNA page with helpful links to other RNA-related sites, the Zuker Lab Homepage (http://www.bioinfo.rpi.edu/∼zukerm/), or e-mail to Michael Zuker. A name for the sequence must be provided in the first field. An unnamed sequence will be assigned a name according to the date and time of submission. The next field takes the sequence, which can be pasted from other locations or typed directly. As the caption explains, blanks and nonalphabetic characters are ignored. N can be used to represent a nucleotide that will neither pair nor stack, and four Ns can be used to join regions of sequence. T is interpreted as U. Currently, a maximum of 6000 nt is allowed. RNA Secondary Structure Prediction
The next field is for optional folding constraints. These constraints might be determined experimentally with enzymatic cleavage (see UNIT 6.1). Five constraints are possible: (1) force nucleotides to be double stranded, (2) force specific base pairs, (3) force
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nucleotides to be single stranded, (4) prohibit specific base pairs, and (5) prohibit a nucleotide between positions i and j from pairing to nucleotides between positions k and l. Each of these constraints is described in detail on the page that is linked by “constraint information.” Next, choose whether the RNA strand is linear or circular; the default is linear. The temperature of structure prediction is limited to 37◦ C because that is the temperature at which RNA structure stability is best understood. Secondary structure prediction is possible at other temperatures using a set of thermodynamic parameters from prior studies (Walter et al., 1994) and a link to mfold 2.3 is available at the top of the page. The stability of RNA secondary structure is best determined at 37◦ C and is poorly determined at temperatures far from 37◦ C. For temperatures <20◦ C or >40◦ C, a more accurate secondary structure can be predicted by using the default folding temperature of 37◦ C. Next, enter the percent suboptimality, upper bound, and window size for suboptimal structure generation. Increasing the percent suboptimality and the upper bound will increase the number of suboptimal structures that are output. Increasing the window size will increase the structural difference in the suboptimal structures and can reduce the number of suboptimal structures. A window size of zero places no requirement on how much suboptimal structures must differ from each other and will result in the largest number of possible suboptimal structures. Choose whether the prediction should be an immediate job (predicted while one waits) or a batch job (an e-mail message is sent when the folding has been completed). Currently, a limit of 800 nt is allowed for immediate jobs. If a batch job is chosen, enter an e-mail address to receive a message when the prediction is complete. The remaining fields affect the drawing of the predicted structures. These settings are used for generating a compressed file that contains drawings of all the predicted structures and are not used for online viewing of structures. An image width of low, medium, large, Xlarge, or huge can be chosen. Larger widths produce structure images of higher resolution. The structure format can be selected as automatic, bases, or outline. A structure drawn in outline format does not specify the identity of each nucleotide in the structure, whereas the bases format shows each nucleotide identity explicitly. Automatic mode will choose either outline or bases depending on the length of the sequence. A base numbering frequency other than default can be chosen. This indicates the interval for labeling the nucleotides in the structures that are output. A structure annotation method can be selected (Zuker and Jacobson, 1998). These annotations can be used as an indication of confidence in a predicted base pair (Zuker and Jacobson, 1995). Finally, start the prediction by following the link labeled “Fold RNA” at the bottom of the page. The program will provide notification of the status of the job, either in the browser for immediate jobs or by e-mail for batch jobs. The other link at the bottom of the page, labeled “Reset form,” will return all values to default and erase the sequence. RNA secondary structure prediction with mfold is fast. A 433-nt Group I intron is folded in <1 min, and the 1542-nt E. coli 16S rRNA is folded in <10 min of CPU time (Table 11.2.1). The actual time for predicting a structure varies depending on how busy the server is.
View the results When the secondary structure prediction is complete, the output page will be available. For immediate jobs, the mfold server moves directly to this page. For batch jobs, the output is available by following the link labeled “View previous foldings” at the top of
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the mfold server homepage. All previous jobs within 2 days will be listed. The results must be viewed at the same computer used to submit the job. The top of the output page lists the sequence folded and the suboptimal structures generated. The rest of the output page is composed of links to various methods for displaying the predicted structures. The first method is the energy dot plot, which plots all base pairs that are contained in its structures within a prescribed free energy increment from the lowest free energy. The approximate lowest free energy possible for a structure that contains a pair is indicated by the color used to plot that pair. The dot plot provides an indication of how well the secondary structure is determined (Zuker and Jacobson, 1995). Three formats are available. Text is a machine-readable text file that can be used for further analysis. The PostScript format can be viewed only with software designed to display PostScript, can be printed on PostScript-capable printers, and can be modified with vector graphics programs. The png jpg format is native to Web browsers, and following the “png jpg” link will display the dot plot on the screen in an interactive mode. This mode allows the user to resize the plot, change the characteristics of the plot, and click on dots to obtain the exact free energy indicated. For this and other interactive plots, pop-up windows should be unblocked. All the predicted structures are available to download in a single compressed file. Choose a compression method and a file format and press “create” to download all structures. The compression is either “gzipped tar” or “zip.” Windows XP can open a zip file directly. On previous versions of Windows, these files can be decompressed with a decompression program such as WinZip, available as shareware at http://www.winzip.com. On a Unix machine, gzipped tar files are decompressed with the gunzip and then tar commands. The structures can be formatted as PostScript, pdf, png, jpg, ct file, Vienna, RNAML, RnaViz ct, Mac ct, GCG, or XRNA ss. A description of these formats is found at the “file formats” link. When in doubt, use the jpg format, because a Web browser can read and display this format. Figure 11.2.8 shows a pdf rendering of the D. melanogaster 5S rRNA secondary structure prediction on the mfold server. Next, structural information in the form of ss-count is available. This indicates the propensity of a nucleotide to be single stranded, and counts the number of times a nucleotide appears single stranded in the set of computed suboptimal structures. This information is available as a text file at the “(ss-count file)” link, or as a plot in PostScript, pdf, png, or jpg that is calculated by pressing the “View plot” button. The online plot can be averaged over several nucleotides or magnified by changing the default values under “View ss-count information.” The individual suboptimal structures are available in several formats, under “View Individual Structures.” The formats are the same as those available for downloading the whole set of structures. When in doubt, choose the jpg format, because it will be displayed in the Web browser. Finally, the last method for displaying the predicted structure information is the dot plot folding comparison. This creates a plot of i versus j, where each dot indicates a base pair in a suboptimal structure. The plot indicates regions of overlap of the suboptimal structures. It is available as either PostScript, png, or jpg. The png and jpg formats are interactive.
RNA Secondary Structure Prediction
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Figure 11.2.8 The secondary structure of D. melanogaster 5S rRNA as predicted by the mfold server and displayed as pdf. For a color version of this figure, see http://www.currentprotocols.com.
COMMENTARY Background Information RNAstructure and mfold predict RNA secondary structures on the basis of free energy minimization. The lowest free energy structure is the structure that is predicted most likely to occur at equilibrium. The secondary structure formation free energy change is predicted using a set of empirical nearestneighbor parameters, determined from optical melting experiments on model systems (Xia et al., 1998; Mathews et al., 1999b, 2004). The partition function in RNAstructure is likewise built from free energy changes for structure formation and implicitly considers all possible secondary structures when calculating base-pair probabilities (Mathews, 2004). RNAstructure and mfold use dynamic programming algorithms that guarantee that the lowest free energy structure will be found. Current Protocols in Nucleic Acid Chemistry
Essentially, the structure prediction problem is divided into smaller problems, and recursion builds the complete secondary structure. Two recent reviews are available that explain dynamic programming in detail (Eddy, 2004; Mathews and Zuker, 2004). The partition function is also calculated with a dynamic programming algorithm. The algorithms used, however, cannot predict pseudoknotted (nonnested) base pairs. Other dynamic programming algorithms exist that can predict pseudoknotted pairs (Rivas and Eddy, 1999; Dirks and Pierce, 2003), but these calculations take considerably longer. On average, only 1.4% of base pairs are pseudoknotted in a database of diverse RNA structures (Mathews et al., 1999b), but this percentage can be much higher for some classes of RNA structures, such as RNase P (Brown, 1999) and tmRNA (Williams and Bartel, 1996).
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Another computer program, the Vienna RNA package (Hofacker, 2003), also predicts lowest free energy secondary structures by dynamic programming. RNAstructure, mfold, and the Vienna package differ slightly in the implementation of the nearest-neighbor parameters for multibranch loops and exterior loops (loops that contain the ends of the sequence). RNAstructure explicitly considers both coaxial stacking of adjacent helices and helices separated by a single mismatch. The Vienna package considers adjacent helix coaxial stacking in free energy minimization, but does not include coaxial stacking in the partition function prediction of base-pair probabilities. In contrast, mfold does not consider coaxial stacking in the dynamic programming algorithm, but a second step, efn2, recalculates the free energy change of folding for each structure, including coaxial stacking of adjacent helices and helices separated by a single mismatch. Note, however, that the default behavior of the mfold server is to not rearrange the predicted structures according to the revised efn2-calculated free energy changes. Because of these differences in the energy models, the programs are not guaranteed to predict the same lowest free energy structure. A recent benchmark, however, showed that the programs have similar average accuracy when mfold is run with efn2 rearrangement (Dowell and Eddy, 2004).
Anticipated Results
RNA Secondary Structure Prediction
Free energy minimization, on average, predicts 73% of known base pairs in the lowest free energy structure for a diverse set of sequences <700 nt and with known secondary structure (Mathews et al., 1999b, 2004). However, using a different set of sequences with known structures, including longer sequences, RNAstructure only predicted 56% of known base pairs (Dowell and Eddy, 2004). Therefore, secondary structure prediction should be viewed as a method for developing structure hypotheses. Suboptimal structures are thus alternative hypotheses for the secondary structure. Constraints on the possible structures can be specified, and it has been shown that the use of constraints based on experimental data improves the accuracy of secondary structure prediction for sequences that would have poorly predicted structures without constraints (Mathews et al., 1999b, 2004). RNAstructure and mfold can utilize constraints based on enzymatic cleavage (revealing paired or unpaired nucleotides; Knapp, 1989). In ad-
dition, RNAstructure can use constraints derived from FMN cleavage (revealing U’s in GU pairs; Burgstaller and Famulok, 1997) or chemical modification (revealing nucleotides that are unpaired, at the ends of helices, or in or adjacent to GU pairs; Ehresmann et al., 1987). UNIT 6.1 discusses the use of enzymes and chemical reagents to probe RNA structures. The base-pair probabilities can be used to determine confidence in a predicted base pair (Mathews, 2004). On average, 66% of predicted base pairs in the lowest free energy structure are in the known structure for a diverse set of sequences. However, when only base pairs with predicted pairing probability ≥0.90 are considered, 83% of predicted pairs are in the known structure. For a probability threshold of 0.99, this accuracy increases to 91%. On average, nearly one quarter of predicted base pairs in the lowest free energy structure have pairing probability of at least 0.99. OligoWalk provides an estimate of binding affinity of structured oligonucleotides to a structured RNA target (Mathews et al., 1999a). For an oligonucleotide to bind tightly, not only should the duplex free energy be low (more negative), but the magnitude of the cost of opening the target structure should also be minimized. It has been shown that the duplex formation free energy and oligonucleotide self-structure terms correlate with antisense oligonucleotide efficacy (Matveeva et al., 2003). Recent reports have shown that self-structure of RNA targets is an important siRNA design criterion (Bohula et al., 2003; Far and Sczakiel, 2003; Petch et al., 2003; Heale et al., 2005).
Literature Cited Bohula, E.A., Salisbury, A.J., Sohail, M., Playford, M.P., Riedemann, J., Southern, E.M., and Macaulay, V.M. 2003. The efficacy of small interfering RNAs targeted to the type 1 insulin-like growth factor receptor (IGF1R) is influenced by secondary structure in the IGF1R transcript. J. Biol. Chem. 278:15991-15997. Brown, J.W. 1999. The ribonuclease P database. Nucleic Acids Res. 27:314. Burgstaller, P. and Famulok, M. 1997. Flavindependent photocleavage of RNA at G.U base pairs. J. Am. Chem. Soc. 119:1137-1138. Dirks, R. and Pierce, N. 2003. A partition function algorithm for nucleic acid secondary structure including pseudoknots. J. Comput. Chem. 24:1664-1677. Dowell, R.D. and Eddy, S.R. 2004. Evaluation of several lightweight stochastic context-free grammars for RNA secondary structure prediction. BMC Bioinformatics 5:71.
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Eddy, S.R. 2004. How do RNA folding algorithms work? Nat. Biotechnol. 22:1457-1458. Ehresmann, C., Baudin, F., Mougel, M., Romby, P., Ebel, J., and Ehresmann, B. 1987. Probing the structure of RNAs in solution. Nucleic Acids Res. 15:9109-9128. Far, R.K. and Sczakiel, G. 2003. The activity of siRNA in mammalian cells is related to structural target accessibility: A comparison with antisense oligonucleotides. Nucleic Acids Res. 31:4417-4424.
Walter, A.E., Turner, D.H., Kim, J., Lyttle, M.H., M¨uller, P., Mathews, D.H., and Zuker, M. 1994. Coaxial stacking of helixes enhances binding of oligoribonucleotides and improves predictions of RNA folding. Proc. Natl. Acad. Sci. U.S.A. 91:9218-9222. Williams, K.P. and Bartel, D.P. 1996. Phylogenetic analysis of tmRNA secondary structure. RNA 2:1306-1310.
Heale, B.S., Soifer, H.S., Bowers, C., and Rossi, J.J. 2005. siRNA target site secondary structure predictions using local stable substructures. Nucleic Acids Res. 33:e30.
Xia, T., SantaLucia, J. Jr., Burkard, M.E., Kierzek, R., Schroeder, S.J., Jiao, X., Cox, C., and Turner, D.H. 1998. Thermodynamic parameters for an expanded nearest-neighbor model for formation of RNA duplexes with Watson-Crick pairs. Biochemistry 37:14719-14735.
Hofacker, I.L. 2003. Vienna RNA secondary structure server. Nucleic Acids Res. 31:3429-3431.
Zuker, M. 1989. On finding all suboptimal foldings of an RNA molecule. Science 244:48-52.
Knapp, G. 1989. Enzymatic approaches to probing RNA secondary and tertiary structure. Methods Enzymol. 180:192-212.
Zuker, M. and Jacobson, A.B. 1995. “Welldetermined” regions in RNA secondary structure predictions. Applications to small and large subunit rRNA. Nucleic Acids Res. 23:27912798.
Lu, Z.J., Turner, D.H., and Mathews, D.H. 2006. A set of nearest neighbor parameters for predicting the enthalpy change of RNA secondary structure formation. Nucleic Acids Res. 34:4912-4924. Mathews, D.H. 2004. Using an RNA secondary structure partition function to determine confidence in base pairs predicted by free energy minimization. RNA 10:1178-1190. Mathews, D.H. and Zuker, M. 2004. Predictive methods using RNA sequences. In Bioinformatics: A Practical Guide to the Analysis of Genes and Proteins, 3rd ed. (A. Baxevenis and F. Oullette, eds.) pp. 143-170. John Wiley & Sons, Hoboken, N.J. Mathews, D.H., Burkard, M.E., Freier, S.M., Wyatt, J.R., and Turner, D.H. 1999a. Predicting oligonucleotide affinity to nucleic acid targets. RNA 5:1458-1469. Mathews, D.H., Sabina, J., Zuker, M., and Turner, D.H. 1999b. Expanded sequence dependence of thermodynamic parameters provides improved prediction of RNA secondary structure. J. Mol. Biol. 288:911-940.
Zuker, M. and Jacobson, A.B. 1998. Using reliability information to annotate RNA secondary structures. RNA 4:669-679. Zuker, M., Mathews, D.H., and Turner, D.H. 1999. Algorithms and thermodynamics for RNA secondary structure prediction: A practical guide. In RNA Biochemistry and Biotechnology (J. Barciszewski and B.F.C. Clark, eds.) pp. 1143. Kluwer Academic Publishers, Boston.
Key References Mathews et al., 1999b, 2004. See above. Derives the thermodynamic parameters used by the secondary structure prediction algorithm and tabulates the accuracy of the algorithm with a large database of structures from sequence comparisons. Zuker, 1989. See above. Explains the method for predicting suboptimal structures using a dynamic programming algorithm.
Internet Resources
Mathews, D.H., Disney, M.D., Childs, J.L., Schroeder, S.J., Zuker, M., and Turner, D.H. 2004. Incorporating chemical modification constraints into a dynamic programming algorithm for prediction of RNA secondary structure. Proc. Natl. Acad. Sci. U.S.A. 101:7287-7292.
http://www.bioinfo.rpi.edu/applications/mfold/ This site contains links to the mfold server and to the download site for mfold.
Matveeva, O.V., Mathews, D.H., Tsodikov, A.D., Shabalina, S.A., Gesteland, R.F., Atkins, J.F., and Freier, S.M. 2003. Thermodynamic criteria for high hit rate antisense oligonucleotide design. Nucleic Acids Res. 31:4989-4994.
http://rna.ucsc.edu/rnacenter/xrna/xrna.html The XRNA homepage at the Santa Cruz RNA Center is the source of XRNA, which can be used for creating publication-quality RNA secondary structure diagrams.
Petch, A.K., Sohail, M., Hughes, M.D., Benter, I., Darling, J., Southern, E.M., and Akhtar, S. 2003. Messenger RNA expression profiling of genes involved in epidermal growth factor receptor signalling in human cancer cells treated with scanning array-designed antisense oligonucleotides. Biochem. Pharmacol. 66:819-830.
Contributed by David H. Mathews and Douglas H. Turner University of Rochester Rochester, New York
Rivas, E. and Eddy, S.R. 1999. A dynamic programming algorithm for RNA structure prediction including pseudoknots. J. Mol. Biol. 285:20532068.
Michael Zuker Rensselaer Polytechnic Institute Troy, New York
Current Protocols in Nucleic Acid Chemistry
http://rna.urmc.rochester.edu The Mathews Lab homepage is the source for downloading RNAstructure.
RNA Folding Pathways
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Thermal Methods for the Analysis of RNA Folding Pathways RNAs form a remarkable variety of structures based on standard Watson-Crick helices, noncanonical pairings (e.g., as found in some internal and hairpin loops; Heus and Pardi, 1991; Correll et al., 1997), and tertiary interactions that join a loop or bulge to another part of the RNA. An objective of current RNA physical studies is to identify noncanonical and tertiary folding motifs and evaluate the factors responsible for their stability. Thermal melting analysis was used in the 1970s to establish the existence of a distinct set of tertiary interactions in transfer RNAs (Cole et al., 1972; Römer and Hach, 1975; Stein and Crothers, 1976) and has become a useful tool for examining the folding of unusual RNA structures up to ∼100 nucleotides in length. Suppose one has identified an RNA or RNA fragment that adopts a specific, functional conformation, e.g., the minimal sequence containing a ribozyme activity or a protein recognition site. The first task in determining the RNA structure is to devise a model of the secondary structure. A good approximation of canonical base pairing in an RNA is fairly easy to obtain from a combination of comparative sequence analysis (Gutell et al., 1992), computer prediction programs based on nearest-neighbor thermodynamic parameters (Zuker, 1989; Serra and Turner, 1995; UNIT 11.2), and “structure mapping” experiments (Ehresmann et al., 1987). Compensatory base changes may be able to establish the existence of specific helix segments required for function (Tang and Draper, 1989). At this point, it is worthwhile asking whether the deduced secondary structure accounts for all the intramolecular interactions of the RNA, or whether noncanonical and tertiary interactions might make the structure much more stable than predicted, or link parts of the structure in unexpected ways. One may also be interested in determining, as an aid to further structural or functional analysis, the pH, salt, and temperature ranges under which the RNA adopts a stably folded structure. Relatively simple melting experiments can answer these questions, and more extensive melting analysis can frequently provide a complete unfolding pathway for an RNA. This unit describes procedures for applying melting analysis to an RNA. The discussion assumes that the secondary structure has al-
UNIT 11.3
ready been established by a combination of prediction and experiment (see Chapter 6). After comments on sample preparation and instrumentation, a framework for describing the thermal denaturation of an RNA that unfolds in several steps is derived. An analysis of transfer RNA unfolding then illustrates the application of this framework to UV and calorimetry data sets and also shows how error analysis can point out uncertainties in the derived thermodynamic parameters. Lastly, comments are offered on experiments that can resolve ambiguities in the thermodynamic analysis and help identify unfolding transitions with specific structures in the unfolding pathway.
EXPERIMENTAL CONSIDERATIONS RNA Samples Transcription of DNA templates by T7 RNA polymerase (UNIT 9.3) is the easiest route to milligram quantities of RNAs longer than 15 to 20 nucleotides; methods have been reviewed elsewhere (Draper et al., 1988; Gurevich, 1996). Chemical synthesis is preferred for shorter RNAs, and may be necessary if modified nucleotides are required (Scaringe et al., 1990; Goodwin et al., 1996; APPENDIX 3C; also see Chapter 3). Aggregation of RNAs is a common problem, particularly in the presence of Mg2+, and samples should always be checked for the presence of multimers by nondenaturing gel electrophoresis (Chory and Pollard, 2000). A buffer of 100 mM potassium acetate and 30 mM Tris acetate (pH 7.3), with or without 5 mM magnesium acetate, is convenient (Gluick et al., 1997). Weak multimerization that is not detected by gel electrophoresis will appear as a concentration dependence of the melting temperatures. UV melting curves are typically obtained at RNA concentrations on the order of 5 to 10 µg/mL, while calorimetry requires about two orders of magnitude higher concentrations. Thus, comparison of melting temperatures in the two experiments should reveal problems with weak intermolecular interactions. Buffers for melting experiments should have small ionization enthalpies to avoid large pH changes with temperature. Tris should be avoided on this account; phosphate or cacody-
Contributed by David E. Draper, Yury V. Bukhman, and Thomas C. Gluick Current Protocols in Nucleic Acid Chemistry (2000) 11.3.1-11.3.13 Copyright © 2000 by John Wiley & Sons, Inc.
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late salts are preferred. Some of the Good buffers (Good and Izawa, 1972; APPENDIX 2A) are suitable for UV absorbance.
UV Absorbance Commercial UV spectrophotometers are available with accessories for collecting UV absorbance as a function of temperature, or a programmable water bath and temperature probe may be added to instruments with suitably thermostatted cells. Refer to UNIT 7.3 for experimental details of UV absorbance melting curves. Peltier devices may seem more convenient than circulating water baths for temperature control, but usually have a limited temperature range (e.g., 10° to 80°C) and a relatively short lifetime when repeatedly cycled over the full range. Instruments with a five- or six-cell carriage that can be automatically cycled for measurement will speed up data collection by a corresponding factor. The cells must be sufficiently surrounded by the heating block that large temperature gradients do not develop. Lastly, a temperature probe should be placed inside a dummy cuvette, and not in the heating block itself. The temperature difference between the block and the cuvette interior should not exceed ∼5°C at the highest temperatures. It is important to test for reversibility of the melting curve by running both heating and cooling experiments on the same sample. Folding kinetics at low temperatures can be slow (for example, see Gluick et al., 1997), and Mg2+ ion–induced hydrolysis of RNA at high temperatures is also a common problem. The latter can be minimized by using the lowest Mg2+ ion concentrations possible, and by working in a neutral to acid pH range (i.e., 6.0 to 7.0).
A
Thermal Methods for the Analysis of RNA Folding Pathways
Renaturation of an RNA preparation before every experiment is necessary to avoid kinetic artifacts. Rather than devise renaturation protocols for each new RNA and salt concentration, the authors have used the following protocol for data collection. RNA samples are diluted into the desired buffers in stoppered cuvettes and then incubated for 15 min at 70°C in the spectrophotometer. The temperature is then ramped at 0.8° to 1°C/min to ∼10°C, and then up to ∼90°C, while temperature and absorbance at 260 nm are read every 0.4°C. In some cases, it may be necessary to slow the temperature ramp at temperatures less than ∼20°C to achieve equilibrium. Example data are shown in Figure 11.3.1, panel A. The first unfolding transition in this RNA is a set of tertiary interactions whose stability is of interest. The coincidence of the heating and cooling curves demonstrate reversibility, and the two curves provide duplicate sets of data in the temperature range of most interest.
Scanning Calorimetry Prior to calorimetry experiments (see UNIT 7.4 for experimental details), it is advisable to thoroughly wash the cells to remove any trace of ribonuclease. A series of extensive rinses with RNase ZAP (Ambion), hot soapy water (using a commercial dishwashing soap such as Dove), and distilled water is adequate for this purpose. The amount of RNA needed for an experiment depends on the number of transitions and their enthalpies; typically ∼1.0 mg/ml is used with the most sensitive calorimeters now available from MicroCal and CSC, though 0.2 mg/ml may be adequate for RNAs with single, sharp transitions. After thorough dialysis of the sam-
B
Figure 11.3.1 UV absorbance melting curve. (A) Cooling curve (starting at ∼70°C, filled symbols) and heating curve (open symbols) for the same RNA sample. (B) First derivative calculated for the data set shown in panel A, using linear regression and a window size of 4°C.
11.3.2 Supplement 2
Current Protocols in Nucleic Acid Chemistry
ple against the desired buffer, two or three baseline scans using the dialysis buffer should be run. These ought to be reproducible. If not, check for degradation of buffer components. The sample should be renatured before being run; 65°C for 15 min, followed by slow cooling, is adequate for most RNAs, but higher temperatures or longer times may be needed. Renaturation conditions should be surveyed by UV experiments prior to calorimetry. A temperature ramp of 1°C/min is sufficiently slow for most RNAs; again, experience with UV melting experiments will indicate what ramp speed is appropriate. Calculations of transition enthalpies depend on accurate measurements of RNA concentration in the calorimetry sample. It is therefore advisable to determine the extinction coefficient of the RNA under the buffer conditions used for the experiment. To do this, prepare an RNA sample and a buffer blank in stoppered cuvettes. After measuring the absorbance of the sample, add concentrated NaOH to a final concentration of 1 M and incubate at 37°C overnight. Check for evaporation by weighing the cuvettes before and after incubation. The extinction of the RNA can then be calculated from the absorbance of the hydrolyzed sample and extinction coefficients of the mononucleotides in alkali (ε260 × 103 values are: AMP, 15.0; GMP, 11.2; CMP, 7.6; and UMP, 7.4).
constant, the expression for K can be written as: ∆H 1 1 K = exp Tm − T R Equation 11.3.2
A melting curve is simply a plot of the fraction of molecules that are unfolded as a function of temperature, or: FU =
Sequential Unfolding Model This section presents a framework for describing RNA unfolding as a function of temperature. To a good approximation, most short RNA hairpins unfold in a two-state manner, i.e., no partially unfolded intermediates are found in significant concentrations at equilibrium. The equilibrium between RNA in its fully folded, native structure (N) and fully unfolded RNA (U) is then simply: K
NU Equation 11.3.1
Because the enthalpy change (∆H) for unfolding is positive, an increase in temperature drives the reaction to the unfolded state. The Tm of the reaction is the temperature at which half of the molecules are unfolded, [N] = [U] and K = 1. From this definition and the van’t Hoff relation for the temperature dependence of K, dln(K)/d(1/T) = ∆H/R, where R is the ideal gas
=
K
1+K
Equation 11.3.3
Thus, there are two variables, ∆H and Tm, that describe the melting curve for a simple hairpin, and from which the equilibrium constant K can be calculated at any temperature. The ∆H found from application of Equations 11.3.2 and 11.3.3 to a melting curve is called a two-state or van’t Hoff enthalpy. Note that ∆H is assumed to be independent of temperature. RNAs larger than hairpins unfold in several steps; for example, there may be several helical segments separated by internal loops or junctions loops, and, of particular interest, additional tertiary interactions that “cross-link” different loops and/or helical segments. In the latter case, it may be obligatory that a set of tertiary interactions unfold before helical segments: K1
THEORETICAL BACKGROUND
[U] [U] + [N]
K2
K3
Kn+1
T S1 S2 S3 . . . Sn U Equation 11.3.4
In this sequential unfolding scheme, T is the completely folded RNA, with intact tertiary interactions, and the S states are RNAs with successively fewer helical segments. Studies of tRNA unfolding in the 1970s showed that its unfolding pathway was best analyzed by a sequential model, with a total of four steps— i.e., tertiary structure plus one of the cloverleaf stems in the first step, followed by the remaining three cloverleaf stems in order of their stabilities (Stein and Crothers, 1976). The alternative to a sequential unfolding pathway is one in which each set of interactions unfolds independently, for example, a series of hairpins linked by single-stranded RNA and having no interactions with each other. It can be shown that use of a sequential pathway to analyze an independently unfolding RNA will, in most cases, yield an excellent approximation of the unfolding thermodynamics. Since most RNAs will probably unfold by a mixed pathway, in which the first, tertiary unfolding step
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is obligatory, but subsequent melting of secondary structures may be largely independent, it is reasonable to use a sequential pathway for an initial analysis in most cases. This point has been considered in some detail by Draper and Gluick (1995), who found that the errors expected in using a sequential pathway to model independent unfolding are on the order of only 5%. Equation 11.3.3, which gives the fraction of unfolded RNA as a function of temperature for a single unfolding step, now must be modified to account for multiple transitions. It is convenient to use a partition function to do this. If the fully folded RNA in Equation 11.3.4, state T, is assigned as the reference state with a statistical weight of 1, the partition function Q for this sequential pathway is: Q = 1 + K1 + K1K2 + K1K2K3 . . . Equation 11.3.5
From the definition of equilibrium constants, each successive term represents the ratio of the concentrations of states S1, S2...Sn to that of state T. Q is therefore a sum of the relative probabilities of finding all of the different possible conformations of the RNA. The fraction of RNAs in which any particular helices are unfolded can be easily calculated from Q. For example, the fraction (F) of RNAs with only the tertiary structure and the first set of secondary interactions unfolded would be: F2 =
K1K2 Q
Equation 11.3.6
Thermal Methods for the Analysis of RNA Folding Pathways
Each K in Equations 11.3.5 and 11.3.6 represents a single two-state transition, and has a temperature dependence with an associated ∆H and Tm (as in Equation 11.3.2). Readers unfamiliar with the use of partition functions to describe conformational equilibria may wish to consult Wyman and Gill (1990) for a detailed discussion. Equations 11.3.2, 11.3.4, and 11.3.5 are all that are needed to describe the unfolding of many complex RNAs. The two simplifying assumptions that have been made are (i) the RNA unfolds in a sequential manner, and (ii) each transition is two-state. These assumptions were adequate for analysis of tRNA unfolding (Crothers et al., 1974; Stein and Crothers, 1976), and are a good starting place for the analysis of any RNA thought to have a well-defined structure. Specific application of these equations to UV hyperchromicity and cal-
orimetry experiments are developed in the next two sections.
UV Melting Profiles In a plot of absorbance versus temperature, it is usually difficult to pick out individual melting transitions or to see changes in transition Tm and sharpness between different plots. For this reason, the first step in analyzing UV melting data is to take the first derivative of the absorbance with respect to temperature; individual transitions then appear as peaks that are more easily distinguished. Two methods are available to do this. The simplest is to define a temperature “window,” W, usually between 4° and 6°C. For each absorbance data point, a linear least-squares line is fit to all the points within ±W/2°C. The slopes of the lines are plotted as the derivative; an example curve is shown in Figure 11.3.1, panel B. The potential disadvantage of this method is that very sharp melting transitions may be flattened and the associated ∆H will be underestimated. A method that avoids this problem is the SavitskyGolay convolution (Press et al., 1992). If the data points are evenly spaced in temperature (or have been made evenly spaced by interpolation), then a polynomial fit to the points in a temperature window can be found by multiplying the points by a matrix of appropriate weights. Using a second-order polynomial, derivatives of very sharp transitions are accurately calculated. A disadvantage of this method is that broad transitions are smoothed much less effectively than by linear regression. The authors’ experience has been that RNA melting transitions are rarely sharp enough to warrant fitting a second-order polynomial. The authors usually normalize the absorbance and derivative curves, so that melts done with different RNA concentrations can be directly compared. This is simply done by dividing all the absorbance readings by the absorbance at some arbitrarily chosen temperature. A low temperature at which the RNA is completely folded is preferable, as the absorbance readings at very high temperatures can be variable due to RNA hydrolysis. All the points within ±W/2°C of the chosen normalization temperature are averaged. In order to analyze an absorbance melting curve, Equation 11.3.6 of the preceding section (which gives the fraction of molecules with a specific structure unfolded) must be modified to relate fraction unfolding to hyperchromicity. First, each unfolding transition must be assigned a hyperchromicity (i.e., the ratio of un-
11.3.4 Supplement 2
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folded- to folded-state absorbance), and the contribution of each transition to the overall hyperchromicity summed. The complete equation is: ∆At =
∆A1K1 (∆A1 + ∆A2)K1K2 + Q Q +
(∆A1 + ∆A2 + ∆A3)K1K2K3 +.... Q Equation 11.3.7
where ∆At is the total fractional hyperchromicity (the same as the normalized absorbance) and ∆A1, ∆A2...∆An are the fractional hyperchromicities associated with each specific transition K1, K2...Kn. Q is the partition function for sequential unfolding, Equation 11.3.5. Each term in the numerator represents the hyperchromicity associated with a successive unfolded state in Equation 11.3.4. Upper- and lower-temperature baselines must be added to Equation 11.3.7, as the absorbances of completely folded or completely unfolded RNA are usually weakly temperaturedependent. After all base pairs are broken at high temperature, the absorbance continues to increase with temperature as bases unstack further. This increase is approximately linear over normally accessible temperature ranges. There is usually a similar increase in absorbance at low temperatures as base-pair stacking within helices decreases with temperature. (RNAs with stable tertiary structures tend to have temperature-independent absorbances at low temperatures, perhaps because the additional structure reduces motions within helices.) In the case of a single unfolding transition, the baselines are taken into account by the following equation (Albergo et al., 1981; Petersheim and Turner, 1983): 1 K A = BF + BU 1 + K 1 + K BF = AF,0 + mF(T ); BU = AU,0 + mU(T ) Equation 11.3.8
BF and BU are the low- and high-temperature baselines, respectively. AF,0 and AU,0 are the absorbances of folded and unfolded RNA extrapolated to a reference temperature (e.g., 0°C) and it is assumed that the baselines are linear, with slopes mF and mU. With multiple unfolding transitions, it is not as clear how to devise appropriate baselines for intermediate states. Fortunately, it is only necessary to fit the highand low-temperature baselines, because calculated values for enthalpies are quite insensitive
to the choice of baselines for folding intermediates, as long as the baseline values are in between the high and low temperature values. The authors have found the following formula convenient in carrying out least-squares fits; it uses the enthalpy of a transition to calculate a weighted average of the high- and low-transition baselines (∆Htot is the total enthalpy of unfolding for the RNA): B = BF + BU − BF ∆H1K1 + (∆H1 + ∆H2)K1K2 . . . Q ∆Htot Equation 11.3.9
The sum of Equations 11.3.7 and 11.3.9, (∆At + B), can be fit to a melting profile using standard nonlinear regression methods; the authors’ programs use the Levenburg-Marquardt algorithm (Press et al., 1992). Only the three variables associated with each transition (∆A, ∆H, and Tm) are fitted, and it is left to the user to provide high- and low-temperature baselines, either from linear least-squares analysis of the absorbance data or from experience. It should be noted that, when fitting curves to normalized first-derivative melting profiles, only the two slopes, mF(T) and mU(T), are needed, and not AF,0 or AU,0. In RNAs with several unfolding transitions, the breadth and spacing of the transitions are frequently such that some of the peaks are not individually resolved. In that case, it may not be obvious how many transitions should be fit to the melting profile because, unless the Tm of each transition is fixed by a separate maximum in the melting profile, there are usually too many degrees of freedom (three variables per transition) to obtain a unique fit. This problem can sometimes be circumvented by collecting data at two different wavelengths. RNA structures frequently differ in their hyperchromicities at different wavelengths; G-C rich helices, for instance, have much larger hyperchromicities at 280 nm than do A-U rich helices (Puglisi and Tinoco, 1989). One then has to fit two melting curves simultaneously, using the same set of ∆H and Tm values for each curve but allowing for different hyperchromicities. While this approach may still result in ambiguities, the benefits of a two-wavelength analysis can be substantial and the experiment should always be tried. An example of two-wavelength analysis applied to transfer RNA melting is shown in a following section, and more extensive methods for ascertaining
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and improving the reliability of a fit are also discussed below.
Calorimetry Calorimetry experiments detect enthalpy directly. The equations used to describe RNA unfolding are similar to those discussed above for absorbance, with ∆H replacing hyperchromicity. Thus, Equation 11.3.7 for sequential unfolding becomes: ∆H1K1 (∆H1 + ∆H2)K1K2 + Q Q
∆H(T ) = +
(∆H1 + ∆H2 + ∆H3)K1K2K3 + ... Q Equation 11.3.10
where ∆H(T) is the total enthalpy change in going from fully folded RNA to the collection of partially unfolded molecules at temperature T. The actual measurement made is the heat capacity of the RNA, ∆CP(T) = [∂∆H(T)/∂T]P, so it is the first derivative of Equation 11.3.10 that is fit to the data. The elimination of hyperchromicities as unknown parameters in Equation 11.3.10 simplifies its use for fitting data. Unfortunately, this advantage is partially offset by new parameters that must be introduced to describe the calorimetry baseline. To see this, first consider a single, two-state unfolding reaction. The completely general equation for the heat capacity curve is: 1 K K ∆H Cp = CF + CU 1 + K + 2 2 1 + K RT (1 + K) Equation 11.3.11
The last term is the temperature derivative of Equation 11.3.10 for a single transition. CF and CU are the intrinsic heat capacities of the native and unfolded states, and generally have small temperature dependencies: dCF CF = CF,0 + (T − T0) dT dCU CU = CU,0 + (T − T0) dT Equation 11.3.12
Thermal Methods for the Analysis of RNA Folding Pathways
where CF,0 and CU,0 are the heat capacities at reference temperature T0. Equations 11.3.11 and 11.3.12 are similar to Equation 11.3.8 for an absorbance melting curve, with heat capacity replacing absorbance in the description of the baseline. In both sets of equations, six variables need to be specified (for a heat capacity curve, the variables are ∆H, Tm, CF,0, CU,0,
and the temperature dependencies of the last two variables). It is possible to extract all of these parameters from a heat-capacity curve that extends over a large enough temperature range. Problems arise when the overall change in heat capacity between native and unfolded RNA, ∆CP = CU,0 − CF,0, is significant and there are several overlapping transitions. In principle, each folding intermediate has an associated heat capacity, CI,0, and a temperature dependence of the heat capacity, dCI/dT. The latter value is usually small, and can be assigned an arbitrary value in between those of the native and unfolded states without introducing significant error. The difficulty comes in determination of CI,0, as illustrated in Figure 11.3.2 for a heat-capacity curve with overlapping transitions. Very different baselines and total enthalpies of unfolding are calculated, depending on whether CI,0 is close to CF,0 (giving a larger total ∆H) or CU,0 (smaller total ∆H). This is a similar problem to that encountered with absorbance melting curves, i.e., three variables associated with each unfolding transition (in this case, ∆H, Tm, and CI,0) is, in many cases, too many degrees of freedom to fit a unique curve to a data set. A small ∆Cp is expected for RNA unfolding, due to the temperature dependence of base stacking; the fully unfolded, single-stranded state of an RNA will have less base stacking (and less enthalpy) at higher temperatures. Thus, RNA unfolding reactions will yield progressively larger ∆H values as the temperature of the measurement is raised; this temperature dependence of ∆H gives a positive ∆CP to any RNA unfolding. The effect is small, on the order of 0.1 kcal/mol per base pair in short helices (Freier et al., 1983; Petersheim and Turner, 1983). However, the authors find the heat capacity baseline problem can be a serious problem for larger RNAs with multiple unfolding transitions, for two reasons: (i) when melting takes place over a wide temperature range, small adjustments in the Cp values associated with each transition can have a significant effect on the total enthalpy, even if the overall ∆CP is relatively small; (ii) in some situations, the overall ∆CP of unfolding an RNA tertiary structure is quite substantial, i.e., much larger than ∆CP for unfolding a protein of similar molecular weight (Laing and Draper, 1994). Though the curve in Figure 11.3.2 might seem an exaggerated case, it is in fact based on actual data and is a realistic illustration of problems that may be encountered. Why some RNAs have
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Current Protocols in Nucleic Acid Chemistry
Figure 11.3.2 The baseline problem in fitting heat-capacity data. The heat-capacity curve was generated using Equation 11.3.10 and three transitions with total enthalpy 150 kcal/mol and overall ∆Cp 5 cal/mol⋅K. The two different baselines were obtained from fits of the heat capacity curve with three transitions (∆H = 135 kcal/mol) or four transitions (thinner line; ∆H = 173 kcal/mol); there was no significant difference in the quality of the fits to the data.
dramatically larger ∆CP compared to others is not presently understood.
EXAMPLE ANALYSIS: tRNA UNFOLDING Analysis of melting data requires considerable effort to ensure that a useful thermodynamic description of the unfolding has been obtained. As an example of problems that may be encountered, this section describes the analysis of transfer RNA melting data and calculations that test the reliability of a proposed fit. The next section describes experiments that can further test a proposed melting pathway.
Initial Fitting of The UV Melting Profile Figure 11.3.3, panel A, shows the UV melting profile of yeast tRNAPhe at 260 and 280 nm. The data clearly distinguish three transitions with different ratios of hyperchromicities at the two wavelengths, and a least-squares fitting program readily finds three sets of transition parameters that give an excellent fit to the data. Note that the low- and high-temperature baselines were fixed manually, and not by the fitting program; the transition parameters are relatively insensitive to the exact baseline values chosen. The first test of a fit is whether the total unfolding enthalpy is consistent with the RNA
secondary structure. From compiled nearestneighbor base-stacking enthalpies (Serra and Turner, 1995), the four tRNAPhe cloverleaf helices alone are predicted to have a total unfolding enthalpy of 190 kcal/mol. “Dangling bases” at the ends of helices and coaxial stacking of helices could add as much as 43 kcal/mol (Serra and Turner, 1995), and further stacking of bases within the anticodon loop and tertiary structure could also add a significant amount of enthalpy. The total enthalpy change associated with the three transitions in Figure 11.3.3, panel A, ∼140 kcal/mol, is thus much too small to be consistent with unfolding of the tRNA. When four transitions are used, the fitting program converges on the set of transitions shown in Figure 11.3.3, panel B. The total enthalpy is now 232 kcal/mol, more in line with the expected value, and the fit has improved significantly (2.7-fold smaller value for χ2). Further tests are now needed to see if this fit is unique and if all variables are well determined. Determining if an optimum fit has been found With a large number of variables, it is quite possible for a least-squares fitting routine to converge on a local minimum, rather than the actual optimum set of parameters. To circumvent this problem, the fitting program can be run a large number of times (>100) using randomized initial values of the parameters. Starting with the transition parameters of the fit in Figure 11.3.3, panel B, a random number generator was used to increase or decrease ∆H and ∆A values by factors of up to 4, and varied Tm values by ±5°C. Forty-seven of the 100 trials converged; one had a larger value for χ2, and the rest had χ2 values not significantly different from that of the fit shown in Figure 11.3.3, panel B. From several such sets of trials, using different ranges of the transition parameters, it is clear that a better fit (smaller χ2 value) has not been missed; however, some of the parameters varied substantially. ∆H3, for instance, has a range of 14 to 167 kcal/mol. This suggests that the two melting profiles do not have sufficient information to constrain all parameters for four melting transitions. Determining if the data support unique values for all of the parameters The problem of whether all the transition parameters are well constrained by a set of melting profiles can be approached by “bootstrap” analysis (Press et al., 1992). This is a Monte Carlo method, in which synthetic data
RNA Folding Pathways
11.3.7 Current Protocols in Nucleic Acid Chemistry
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A
B
C
ure 11.3.3B, and the linear correlation coefficients of each fit parameter versus all other parameters were obtained. These values are summarized in Table 11.3.1. All coefficients with absolute values greater than 0.8 are in boldface. The last two lines of the table give the average and standard deviation (1σ) for each parameter. There are a large number of strong correlations. Particularly worrisome are ∆H3, A2603, and A2803, which strongly correlate with a number of parameters and also have unusually large standard deviations. A plot of ∆H3 versus A2603 is in Figure 11.3.2C; it is clear from the wide range of values and strong correlation (R = 0.95) that the enthalpy and hyperchromicity of transition 3 are not uniquely determined. The authors conclude that the UV melting data sets have enough information to constrain three melting transitions, but not enough information to determine four transitions. Additional data are needed to interpret the unfolding thermodynamics.
Calorimetry Analysis
Figure 11.3.3 UV analysis of yeast tRNAPhe melting (buffer contained 100 mM KCl and 5 mM potassium cacodylate). (A) Fit obtained assuming three sequential transitions (Equation 11.3.7). Data were obtained at 260 nm (open symbols) and 280 nm (closed symbols). (B) Fit of four sequential transitions to the same data set as in panel A. (C) Correlation plot, ∆H3 versus A2603, for the four transition fit (Table 11.3.1).
Thermal Methods for the Analysis of RNA Folding Pathways
sets are generated by randomly drawing the same number of points from the original data set as contained in that set. Some points will be duplicated, and other points will be missing; different optimum fits will be found for each synthetic set, as the data points will be differently weighted. Statistics compiled on a large number (200 to 1000) of such trials give the confidence intervals for any one variable. A useful aspect of this analysis is the ability to ask whether different variables are correlated. If changes in one fit parameter are paralleled by changes in another parameter, then the data contain insufficient information to specify unique values of both parameters. In this analysis, 500 synthetic data sets were generated from the two melting profiles in Fig-
Calorimetry provides additional information that can be used to interpret the UV melting profile. Figure 11.3.4 shows a heat capacity curve for tRNAPhe under the same buffer conditions as used for the UV melting experiment in Figure 11.3.3. As discussed above, the uncertainty in analyzing these data is interpolation of a baseline. For this curve, the authors used the following procedure: 1. Low- and high-temperature baselines were determined by linear least-squares fitting of data over 10°C ranges (10° to 20°C and 90° to 100°C). 2. An initial set of transition parameters (including Cp values that determine the baseline) was obtained by a fitting routine that apportions the overall ∆Cp between the transitions according to their enthalpies (χ2 = 6.05; total ∆H = 239.6 kcal/mol). A minimum of four transitions was required to fit the data reasonably. 3. Fitting was continued after relaxing the constraint in step 2 (above) and substituting the constraint that ∆Cp from one transition to the next must be positive; a slightly better fit was obtained (χ2 = 5.72; total ∆H = 241.9 kcal/mol). 4. The fit parameters were randomly varied (as described above for UV melting profiles) to see if significantly different baselines could be drawn, subject to the constraint that ∆Cp values must be positive. Of 107 converged fits, none was a significantly improved fit (minimum χ2 = 5.71). Among these fits, the maximum and
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Current Protocols in Nucleic Acid Chemistry
0.0826
0.0027
1.0000
1.0000
1.0000
1.0000
1.0000
0.3
Α2804
0.981
-0.975
0.813
0.7423
0.989
1.0000
41.5
Α2604
-0.981
-0.827
0.922
0.703
0.152
0.132
1.0000
0.0032
0.0777 1.3
71.8 0.1
53.9 0.004
0.103 0.005
0.133
7.14
75.2
0.3
62.3
0.0066
0.0357
0.0088
0.0409
-0.940
1.42
46.96
0.440
-0.590
-0.253
0.5
72.2
0.908
0.499
-0.896
-0.342
0.0046
0.0576
-0.917
-0.914
-0.448
0.898
0.351
0.283
0.0061
0.0936
-0.873
-0.921
-0.900
-0.493
0.879
0.336
0.259
Σ∆
Αϖε
Τm4
∆H4
Α2803
Α2603
Τm3
∆H3
Α2802
Α2602
Τm2
∆H2
Α2801
Α2601
Τm1
-0.950
∆H1
-0.224
-0.611
-0.270
0.463
0.178
0.158
0.184
0.179
–0.116
1.000
-0.632
-0.182
0.469
0.170
0.143
0.167
0.166
–0.102
0.488
-0.540
-0.460
-0.179
-0.150
-0.175
-0.176
0.108
0.491
-0.579
-0.335
-0.047
-0.090
-0.116
-0.096
0.081
1.0000
0.526
-0.701
0.0716
0.0003
-0.037
-0.045
-0.006
0.432
-0.712
0.104
0.0264
-0.005
-0.015
-0.034
0.972
0.398
-0.642
-0.387
-0.373
-0.354
0.288
1.0000
0.642
0.111
0.114
0.141
0.152
-0.074
0.556
-0.710
-0.683
-0.633
-0.624
0.612
0.510
-0.721
-0.705
-0.690
0.665
-0.757
0.162
0.190
-0.148
1.0000
0.696
-0.558
-0.268
-0.926
1.0000
∆A2604 ∆A2804
0.115
Tm4
0.727
∆H4
1.0000
∆A2803
0.715
∆A2603
0.930
Tm3
1.0000
∆H3
0.894
∆A2802
0.906
∆A2602
1.000
Tm2
-0.912
∆H2
-0.860
∆A2801
∆A2601
Tm1
UV melting data from Figure 11.3.3 were used in 500 cycles of bootstrap analysis, with four sequential transitions; both 260 and 280 nm wavelength data sets were used simultaneously. The table shows the results of statistical analysis of the 492 cycles that converged on a fit; average and standard deviation for each of the variables are shown in the bottom two lines of the table, and linear correlation coefficients for each pair of variables are in the body of the table. Correlation coefficients with absolute values greater than 0.8 are in bold face type. ∆H units are kcalmol, and Tm values are in ˚C.
aThe
1.3
42.1
1.000
∆H1
Table 11.3.1 Correlation Coefficients from Bootstrap Analysisa
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Supplement 2
minimum total ∆H differed by only 9 kcal/mol (236.3 to 245.2); these curves have the most extreme possible positions of the baseline (Fig. 11.3.4). This source of error is probably comparable to the reproducibility of the heat capacity curve. 5. Bootstrap analysis showed no significant correlations between parameters, and small confidence intervals. The relatively small ∆Cp for tRNA melting is well within the range expected for unfolding of the four cloverleaf helical segments, and is small enough that uncertainties in the Cp values contribute less than ±2% to the overall error in measuring unfolding ∆H. Note that the assumption that ∆Cp > 0 for any one transition is essential; otherwise, there is not enough information in the curve to determine the parameters of all four transitions. Since the source of ∆Cp in this example is the temperature-dependent stacking of bases in the unfolded state, it is reasonable to assume ∆Cp > 0. In another RNA examined by the authors, ∆Cp is on the order of 5 kcal/mol-K. The assumption of positive ∆Cp values is then made with less confidence, since the origin of the large ∆Cp is unknown.
Agreement of UV Melting and Calorimetry Data Sets A simple test for consistency between UV and heat capacity data sets is to refit the UV data using fixed enthalpy and Tm values from the calorimetry analysis. When the calorimetric parameters obtained in Figure 11.3.4 were applied to the UV data in Figure 11.3.3, the hyperchromicities adjusted to give an excellent
Thermal Methods for the Analysis of RNA Folding Pathways
fit. A more general method is simply to fit both UV and heat capacity data sets simultaneously. Bootstrap error analysis can then be run on the combined data sets. With the tRNAPhe data, the parameters obtained are very close to those found by analysis of the calorimetry data alone, and the only significant differences from the UV analysis are the enthalpies of transitions 2 and 3.
Computer Programs Global Melt Fit, a program for simultaneous curve fitting of UV absorbance and calorimetry data sets, and for applying bootstrap analysis, is available on the Web (http://www.jhu.edu/ ∼chem/draper/) or from the authors.
EXPERIMENTAL TESTS OF AN UNFOLDING PATHWAY An analysis of UV and calorimetry data will invariably yield a set of two-state unfolding transitions whose total enthalpy is at least as large as the predicted nearest-neighbor basestacking enthalpy. The next question is the physical significance of the transitions, i.e., can individual transitions be identified with specific structures? Though it might seem that each transition should correspond to the unfolding of a single helical segment or set of tertiary interactions, there is no reason why this has to be true. Secondary structure units may melt in different orders by several pathways, so that the unfolding pathway is branched instead of strictly sequential. A single “state” in the sequential unfolding analysis may then represent a mixture of partially unfolded states that have about the same stability (Laing and Draper,
Figure 11.3.4 Calorimetry analysis of tRNAPhe under the same buffer conditions as in Figure 11.3.3. The solid curves represent the total Cp, baseline, and individual transitions for the fit with the smallest total enthalpy: ∆H1 = 48.7; Tm1 = 40.5; Cp1 = 3.95; ∆H2 = 82.9; Tm2 = 53.2; Cp2 = 4.70; ∆H3 = 56.7; Tm3 = 59.1; Cp3 = 5.37; ∆H4 = 49.0; Tm4 = 70.0; Cp4 = 5.39 (∆H in kcal/mol, Tm in °C, and Cp in kcal/mol-K). The dashed line is the baseline for a fit with the largest overall enthalpy, and differs principally in its smaller value for Cp3 (4.83).
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1994; Draper and Gluick, 1995). An additional problem is whether the two-state assumption is warranted for a transition. This section suggests additional experiments that can test a hypothesized unfolding mechanism.
Substitution of Base Pairs Substitution of a helix A-U base pair with a G-C pair, or vice versa, destabilizes or stabilizes the helix by a predictable amount (Serra and Turner, 1995). The resulting change in helix Tm is usually several degrees, which can be easily detected in a melting experiment. If the unfolding pathway is simple (i.e., branches are not significantly populated) then the ∆H and Tm of only a single transition will change significantly in the mutant. Doing this kind of experiment for each helix in the secondary structure is a rigorous test of the unfolding pathway. Similar substitution experiments can be done for sets of tertiary interactions, if specific bondings are known or suspected. Too little is known about tertiary structure to predict the outcome of such an experiment quantitatively, but it can be useful as a way to confirm the existence of an interaction within a tertiary structure and to probe the range of substitutions that can be accommodated by a structure. Such experiments have been done for a base pair linking D and variable loops within tRNA tertiary structure (Hou et al., 1995), and for a putative base triple interaction in a ribosomal RNA fragment (Conn et al., 1998). Any time a variant RNA is made, the new sequence should be run through a program that predicts secondary structure, such as Mulfold (Zuker, 1989; see also http://www. rpi.edu/∼zukerm/). It is not uncommon that a one- or two-base sequence change favors an alternative secondary structure with very different unfolding properties.
Melting of Subfragments Larger RNAs can usually be divided into several smaller fragments that should retain the same secondary structure as the larger fragment under study (Liang and Draper, 1994; Gluick and Draper, 1997). The melting behavior of such small fragments, with only one or two transitions, is usually easy to analyze and should correspond to transition(s) in the melting of the larger RNA. This is a particularly useful way to find out if the two-state approximation is valid for a specific helix. Note that coaxial stacking and simple entropic effects may alter the stability of a helix when it is taken
out of context of a larger, extended hairpin (Draper and Gluick, 1995).
Deviations from Two-State Behavior The model developed above for RNA unfolding assumes that the RNA behaves as a collection of two-state unfolding events. This assumption is difficult to test rigorously. The usual criterion for two-state behavior is that ∆H calculated from van’t Hoff analysis of the unfolding (as described above, Equation 11.3.3) agrees with the calorimetric ∆H measured from the area under the heat-capacity curve. Only in the case of a small RNA with a single transition, or an RNA with very well-resolved transitions, is it possible to carry out this quantitative test. But frequently an RNA can be divided into several segments which can be individually tested (see above). Deviations from two-state behavior can be anticipated from the sequence of a helix. Using nearest-neighbor base-stacking parameters and loop entropies (Serra and Turner, 1995), a separate equilibrium constant can be calculated for removal of each successive base pair from either end of a helical segment (the so-called “zipper” model of duplex unwinding). The terms for the fully folded state and all the partially unfolded states are then included in a partition function (Equation 11.3.5), and the melting curve is predicted. If the two-state approximation is valid, the calculated curve will not deviate much from a curve predicted using only a single equilibrium constant for folded and unfolded forms. Long helices (more than 6 to 8 base pairs), or helices containing runs of A-U or G-U pairs, tend to “fray” significantly. Fraying may be substantially reduced when the helix is adjacent to loops or helices in a larger structure. An example in which deviations from twostate behavior were identified is an RNA pseudoknot that contains two coaxially stacked helices (Gluick and Draper, 1997). This RNA melted in two well-resolved transitions. However, the heat capacity curve could only be fit with three two-state transitions; the two peaks of the melting curve were associated with large enthalpy transitions, and a third transition of much lower enthalpy was spread between the two main transitions. Several clues suggested that the low enthalpy transition might be an artifact of non-two-state behavior: 1. The total enthalpy calculated from twostate analysis of UV melting curves increased with salt concentration, even though the enthalpy of helix unfolding is usually independent of salt (Williams et al., 1985). Higher
RNA Folding Pathways
11.3.11 Current Protocols in Nucleic Acid Chemistry
Supplement 2
salt concentrations can reduce the proportion of molecules with “frayed” helices, promoting two-state melting. 2. An RNA fragment representing hairpin 1 of the pseudoknot melted in a transition with lower Tm at 260 than at 280 nm. This suggested that some A-U pairs (with larger hyperchromicities at 260 nm) melted before G-C pairs. 3. The two-state enthalpies of both component hairpins increased with salt concentration. The UV and heat-capacity melting profiles of the pseudoknot could be fit simultaneously with two transitions if the ratio of van’t Hoff and calorimetric ∆H values, ∆HvH/∆Hcal, were included to allow the transitions to be broader than expected based on the total enthalpy of unfolding. This ratio is a measure of the deviation from two-state behavior, and approaches 1 for a perfectly two-state transition. For the two transitions of the pseudoknot, the ratios varied from 0.68 to nearly 1, depending on the salt concentration (Gluick and Draper, 1997).
LITERATURE CITED Albergo, D.D., Marky, L.A., Breslauer, K.J. and Turner, D.H. 1981. Thermodynamics of (dGdC)3 double-helix formation in water and deuterium oxide. Biochemistry 20:1409-1413. Chory, J. and Pollard, J.D. Jr. 2000. Separation of small DNA fragments by conventional gel electrophoresis. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 2.7.1.-2.7.8. John Wiley & Sons, New York. Cole, P.E., Yang, S.K., and Crothers, D.M. 1972. Conformational changes of transfer ribonucleic acid: Equilibrium phase diagrams. Biochemistry 11:4358-4368. Conn, G.L., Gutell, R.R., and Draper, D.E. 1998. A functional ribosomal RNA tertiary structure involves a base triple interaction. Biochemistry 37:11980-11988. Correll, C.C., Freeborn, B., Moore, P.B. and Steitz, T.A. 1997. Metals, motifs, and recognition in the crystal structure of a 5S rRNA domain. Cell 91:705-711. Crothers, D.M., Cole, P.E., Hilbers, C.W. and Shulman, R.G. 1974. The molecular mechanism of thermal unfolding of Escherichia coli formylmethionine transfer RNA. J. Mol. Biol. 87:63-88. Draper, D.E. and Gluick, T.C. 1995. Melting studies of RNA unfolding and RNA-ligand interactions. Methods Enzymol. 250:281-305.
Thermal Methods for the Analysis of RNA Folding Pathways
Draper, D.E., White, S.A., and Kean, J.M. 1988. Preparation of specific ribosomal RNA fragments. Methods Enzymol. 164:221-237.
Ehresmann, C., Baudin, F., Mougel, M., Romby, P., Ebel, J.-P., and Ehresmann, B. 1987. Probing the structure of RNAs in solution. Nucl. Acids Res. 15:9109-9128. Freier, S.M., Burger, B.J., Alkema, D., Neilson, T. and Turner, D.H. 1983. Effects of 3′ dangling end stacking on the stability of GGCC and CCGG double helices. Biochemistry 22:6198-6202. Gluick, T.C. and Draper, D.E. 1997. Folding of an mRNA pseudoknot required for stop codon readthrough: Effects of mono- and divalent ions on stability. Biochemistry 36:16173-16186. Gluick, T.C., Gerstner, R.G. and Draper, D.E. 1997. Effects of Mg2+, K+, and H+ on an equilibrium between alternative conformations of an RNA pseudoknot. J. Mol. Biol. 270:451-463. Good, N.E. and Izawa, S. 1972. Hydrogen ion buffers. Methods Enzymol. 24:53-68. Goodwin, J.T., Osborne, S.E., Scholle, E.J., and Glick, G.D. 1996. Design, synthesis, and analysis of yeast tRNAPhe analogs possessing intraand inter-helical disulfide cross-links. J. Am. Chem. Soc. 118:5207-5215. Gurevich, V.V. 1996. Use of bacteriophage RNA polymerase in RNA synthesis. Methods Enzymol. 275:382-397. Gutell, R.R., Power, A., Hertz, G.Z., Putz, E.J., and Stormo, G.D. 1992. Identifying constraints on the higher-order structure of RNA: Continued development and application of comparative sequence analysis methods. Nucl. Acids Res. 20:5785-5795. Heus, H.A. and Pardi, R. 1991. Structural features that give rise to the unusual stability of RNA hairpins containing GNRA loops. Science 252:191-194. Hou, Y.M., Sterner, T. and Jansen, M. 1995. Permutation of a pair of tertiary nucleotides in a transfer RNA. Biochemistry 34:2978-2984. Laing, L.G. and Draper, D.E. 1994. Thermodynamics of RNA folding in a highly conserved ribosomal RNA domain. J. Mol. Biol. 237:560-576. Petersheim, M. and Turner, D.H. 1983. Base-stacking and base-pairing contributions to helix stability: Thermodynamics of double-helix formation with CCGG, CCGGp,CCGGAp, CCGGUp, and ACCGGUp. Biochemistry 22:265-263. Press, W.H., Teukolsky, S.A., Vetterling, W.T., and Flannery, B.P. 1992. Numerical Recipes in C. Cambridge University Press, Cambridge. Puglisi, J.D. and Tinoco, I., Jr. 1989. Absorbance melting curves of RNA. Methods Enzymol. 180:304. Römer, R. and Hach, R. 1975. tRNA conformation and magnesium binding: A study of yeast phenylalanine-specific tRNA by a fluorescent indicator and differential melting curves. Eur. J. Biochem. 55:271-284.
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Current Protocols in Nucleic Acid Chemistry
Scaringe, S.A., Francklyn, C. and Usman, N. 1990. Chemical synthesis of biologically active oligoribonucleotides using beta-cyanoethyl protected ribonucleoside phosphoramidites. Nucl. Acids Res. 18:5433-441.
Williams, A.P., Longfellow, C.E., Freier, S.M., Kierzek, R. and Turner, D.H. 1985. Laser temperature-jump, spectroscopic, and thermodynamic study of salt effects on duplex formation by dGCATGC. Biochemistry 28:4283-4291.
Serra, M.J. and Turner, D.H. 1995. Predicting thermodynamic properties of RNA. Methods Enzymol. 259:242-261.
Wyman, J. and Gill, S. 1990. Binding and Linkage. Functional Chemistry of Biological Macromolecules. University Science Books, Mill Valley, Calif.
Stein, A. and Crothers, D.M. 1976. Conformational changes of transfer RNA: The role of magnesium(II). Biochemistry 15:160-167.
Zuker, M. 1989. On finding all suboptimal foldings of an RNA molecule. Science 244:48-52.
Tang, C.K. and Draper, D.E. 1989. An unusual mRNA pseudoknot structure is recognized by a protein translational repressor. Cell 57:531-536.
Contributed by David E. Draper, Yury V. Bukhman, and Thomas C. Gluick Johns Hopkins University Baltimore, Maryland
RNA Folding Pathways
11.3.13 Current Protocols in Nucleic Acid Chemistry
Supplement 2
Probing RNA Folding Pathways by RNA Fingerprinting
UNIT 11.4
Many RNA sequences form more than one stable secondary or tertiary structure under physiological conditions. As these conformations frequently exchange with one another in a period of minutes or seconds, solution-based methods are often inadequate to resolve differences in their structure and activity. This unit describes methods for separating RNA conformers by native polyacrylamide gel electrophoresis. The activity of each electrophoretic species is assayed while the RNA is still immobilized in the gel matrix. The protocols given here were initially developed to discriminate between active and inactive conformation of the self-splicing RNA from Tetrahymena thermophila (Emerick and Woodson, 1994), but can be applied to a wide variety of catalytic RNAs and RNA-protein complexes. Native polyacrylamide gel electrophoresis is used to separate uniformly 32P-labeled RNA on the basis of shape or hydrodynamic radius (see Basic Protocol 1). Once optimal electrophoretic separation has been achieved, the catalytic activity of each conformer is determined in situ by soaking substrates into the gel matrix (see Basic Protocol 2). Alternatively, the RNA can be eluted from the gel and analyzed directly (see Alternate Protocol). The secondary structure of the RNA is probed by addition of base-modification reagents to the RNA in the gel (see Basic Protocol 3). CAUTION: This procedure should be performed only by personnel trained in the use of radioactive materials and in NRC-licensed sites. Standard precautions to minimize exposure and prevent radioactive contamination of personnel and equipment should be followed at all times. 32P is a high-energy β emitter and requires shielding of personnel. NOTE: Care must be taken to avoid introducing ribonucleases into the samples. All solutions should be prepared with deionized water (18 MΩ) that is free of pyrogens and organic contaminants. Although less desirable, water treated with diethylpyrocarbonate (DEPC; APPENDIX 2A) may be used if a supply of RNase-free water is not available. DEPC must be allowed to decompose completely before use (e.g., by autoclaving), as it reacts with nucleic acids and Tris base. Solutions should be sterilized by filtration (0.2 µm) or autoclaved. Gloves must be worn while handling samples and preparing solutions, and pipet tips and sample tubes must be free of nucleases. This can usually be achieved by purchasing good quality machine-packaged disposable plasticware, avoiding contact with bare skin, and storing disposables in clean, dust-free containers. NATIVE GEL ELECTROPHORESIS OF 32P-LABELED RNA This protocol was developed to separate different structural forms of a 657-nt self-splicing RNA (Emerick and Woodson, 1994). The protocol can be adapted to shorter transcripts by increasing the percentage of the polyacrylamide gel. Once the best conditions for separating the conformers of interest have been determined, this protocol is used to prepare the RNA for further analysis as described in Basic Protocols 2 and 3.
BASIC PROTOCOL 1
Materials 70% ethanol 40% (w/v) 29:1 acrylamide/bisacrylamide, 4°C 10× THEM buffer (see recipe) 10% (w/v) ammonium persulfate, 4°C TEMED, 4°C
RNA Folding Pathways
Contributed by Sarah A. Woodson
11.4.1
Current Protocols in Nucleic Acid Chemistry (2000) 11.4.1-11.4.17 Copyright © 2000 by John Wiley & Sons, Inc.
Supplement 2
5× splicing buffer (see recipe) 32 P-labeled RNA, desalted (see Support Protocol 1) 30 mM MgCl2 5× glycerol loading buffer (see recipe) 20 × 20–cm glass plates with 0.5-mm spacers and comb (1 or 2 sets) Vacuum source and 50-mL side-arm flask (for degassing) 20 × 20–cm vertical gel electrophoresis apparatus with recirculating cooling reservoir (e.g., Owl Scientific Penguin 10DS or equivalent) Refrigerated recirculating bath to connect to gel apparatus Spatula or razor blade 3MM filter paper (Whatman) Vacuum gel drying apparatus X-ray film for autoradiography or phosphorescent imager CAUTION: Acrylamide and N,N′-methylenebisacrylamide are neurotoxins and suspected carcinogens. Preparation of solutions with these compounds should be performed in a well-ventilated fume hood, and extreme precautions taken to minimize contact with solids or solutions. Cast native 6% polyacrylamide gel 1. Thoroughly clean two glass plates and remove any streaks with 70% ethanol. 2. Assemble glass plates, placing 0.5-mm spacers between glass on each side. Clamp the sides of the sandwich with aluminum binder clamps and seal bottom with tape. The bottom of plates may be left open. The gel solution will be retained by capillary action.
3. Mix the following in a graduated cylinder (25 mL total volume per gel): 3.75 mL 40% 29:1 acrylamide/bisacrylamide 2.5 mL 10× THEM buffer 250 µL 10% ammonium persulfate Deionized water up to 25 mL. 4. Transfer to 50-mL side-arm flask and degas mixture by swirling 1 to 3 min under a gentle vacuum (water aspirator). 5. Add 16 µL TEMED and swirl to initiate polymerization. Immediately pour solution into the top of the casting frame while holding glass plates at a very slight incline with respect to the bench. It is important to pour the solution evenly to avoid introducing bubbles.
6. Insert comb into top of frame to form desired number of sample wells; allow gel to polymerize in a horizontal position (i.e., ~20 min). Pre-run gel 7. Remove comb, flushing wells with water or running buffer. Remove any tape from bottom of gel. Place gel on electrophoresis apparatus as instructed by manufacturer.
Probing RNA Folding Pathways by RNA Fingerprinting
8. Prepare 1× THEM buffer by diluting 10× stock with deionized water. Add sufficient 1× THEM buffer to upper and lower chambers to cover top and bottom of gel. 9. Pre-run at 15 W per gel for 15 to 30 min. Adjust temperature of circulating bath to maintain surface of plates at 10°C.
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Prechilling the bath decreases the time required for gel to reach desired temperature. Bath temperatures of –5° to 0°C may be required to cool gel sufficiently; add 50% ethylene glycol to bath to prevent freezing. CAUTION: Personnel should be shielded from β radiation during steps 10 to 14.
Prepare and load RNA samples 10. Add the following to a 0.5-mL microcentrifuge tube (8 µL total volume): 2 µL 5× splicing buffer 1 µL 32P-labeled RNA (100,000 cpm) 5 µL deionized H2O. 11. Place 2 µL of 30 mM MgCl2 on lid of microcentrifuge tube and close firmly. The drop of buffer will remain on the lid by surface tension.
12. To renature RNA, place closed tube in a heating block at 95°C for 1 min. Transfer to microcentrifuge and immediately spin for 1 min. Place on ice. For unrenatured RNA samples, omit the incubation at 95°C. The drop of MgCl2 will mix with the warm RNA solution at the bottom of the tube during centrifugation. This procedure minimizes metal ion–catalyzed hydrolysis of the RNA during the high-temperature incubation. Place the heating block next to the microcentrifuge to minimize the time required to transfer the samples.
13. Add 2.5 µL of 5× glycerol loading buffer and mix well. Place samples on ice. 14. Load 2 µL of each sample into wells of native gel and run at 15 W per gel for 5 to 6 hr at 10°C. For the best resolution, the sample should be no more than 2 to 3 mm deep after loading. The electrophoresis time will vary with the size of the RNA and the percentage of polyacrylamide. Gels are typically run until xylene cyanol FF is at the bottom of the gel or beyond.
Detect 32P-labeled bands of RNA 15. Unclamp gel from apparatus and remove spacers. Gently pry the glass plates apart with a thin spatula or razor blade and remove one glass plate. Do this step as soon as the run is complete.
16. Transfer gel from the second glass plate to dry Whatman 3MM filter paper cut a little larger than the gel. Cover completely with plastic wrap, and dry 15 to 30 min under vacuum using a heated gel-drying apparatus. Do not do this step if planning to do two-dimensional gel electrophoresis (Basic Protocol 2).
17. Expose dried gel to X-ray film or a phosphorescent screen overnight to obtain an image of the radioactive bands. ASSAY OF CATALYTIC ACTIVITY BY TWO-DIMENSIONAL GEL ELECTROPHORESIS Following separation of RNA conformers by native gel electrophoresis (see Basic Protocol 1), the catalytic activity of each species is determined by addition of substrate while the RNA is immobilized in the gel matrix. The products are analyzed by two-dimensional electrophoresis under denaturing conditions (Branch et al., 1989). The spliced products are detected as shorter RNAs on the denaturing gel.
BASIC PROTOCOL 2
RNA Folding Pathways
11.4.3 Current Protocols in Nucleic Acid Chemistry
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Materials 0.1 mM GTP in splicing buffer (see recipe) 2× urea loading buffer (see recipe) 40% (w/v) 29:1 acrylamide/bisacrylamide, 4°C 10× TBE buffer (APPENDIX 2A) Urea 10% (w/v) ammonium persulfate TEMED Bed of ice (size of gel) X-ray film Razor blades Glass plates (e.g., 20 × 20 cm or larger) Additional reagents and equipment for native gel electrophoresis (see Basic Protocol 1) Excise samples from native polyacrylamide gel (first dimension) 1. Separate RNA conformers by native gel electrophoresis (see Basic Protocol 1, steps 1 to 15). In step 10, use ≥500,000 cpm per sample and reduce the volumes by half (i.e., total 5 µL). Load 3 to 5 µL. Skipping wells or leaving a large space between lanes will make it easier to excise the bands from gel without cross-contamination in step 4.
2. After removing top glass plate, cover gel with plastic wrap. Place gel (glass side down) on a bed of ice. The gel should be maintained on ice when possible during the following steps to retard diffusion of the RNA in the gel. Steps 3 to 5 should be performed as quickly as possible.
3. In a darkroom or light-tight box, place a sheet of X-ray film on gel for 2 to 15 min. Mark gel and film for realignment. Develop film after the desired time. 4. Align gel (glass side down) over the autoradiogram. Using the autoradiogram to locate the RNA, excise each lane with a new razor blade, being certain to include the entire region of the lane that contains the bands of interest. Invert an acrylic radiation shield over gel to block 32P radiation and protect samples from airborne ribonucleases. The gel pieces should be ∼0.6 × 4 cm.
5. Transfer gel pieces to a glass plate (e.g., 20 × 20 cm or larger), laying them horizontally along the bottom edge as shown in right panel of Figure 11.4.1. Forceps should be RNase-free or flamed with ethanol just before use.
Perform self-splicing in situ 6. Pipet 5 µL of 0.1 mM GTP in splicing buffer as evenly as possible over the surface of each gel piece. Allow to stand 2 min at room temperature. 7. Add 10 µL of 2× urea loading buffer. Urea quenches the self-splicing reaction. Probing RNA Folding Pathways by RNA Fingerprinting
Assemble and run denaturing polyacrylamide gel (second dimension) 8. Complete assembly of gel frame for the second-dimension electrophoresis by placing spacers on each side and a second glass plate on top of samples from step 7. Clamp sides of frame and seal bottom with tape.
11.4.4 Supplement 2
Current Protocols in Nucleic Acid Chemistry
I
N
1 I N 2
6% native gel
1 8% denaturing gel
Figure 11.4.1 Two-dimensional gel electrophoresis of self-splicing RNA. After native gel electrophoresis (first dimension), the lane is excised and placed on the bottom of the second casting frame. The active form of the RNA (N) is spliced when GTP substrate is added to the gel slice. The products are resolved in a denaturing polyacrylamide gel (second dimension).
It may be necessary to use spacers that are slightly thicker than those used for the first-dimension gel to avoid squashing the gel pieces.
9. Prepare solution for an 8% denaturing sequencing gel (25 mL total volume for 0.5 mm × 20 × 20 cm). 5 mL 40% 29:1 acrylamide/bisacrylamide 2.5 mL 10× TBE buffer 12 g urea Deionized water up to 25 mL. Dissolve urea at 50 to 65°C. Transfer to a sidearm flask and degas. 10. Add 0.5 mL of 10% ammonium persulfate and 20 µL TEMED to begin polymerization. 11. Immediately pour solution into gel frame from top, allowing solution to run slowly down the side and then across the bottom, covering gel slices without introducing air bubbles. It is not necessary to insert a comb in the top of gel.
12. After gel has polymerized, clamp it on gel box with first-dimension samples at the bottom. Add 1× TBE buffer to upper and lower reservoirs. 13. Connect the positive electrode to top of gel and the negative electrode to the bottom. The gel is run in reverse so that the RNA migrates from bottom to top.
14. Run at 25 to 30 W (40° to 50°C) until the xylene cyanol has run off the top of gel. 15. Disassemble, dry, and expose gel to X-ray film (see Basic Protocol 1, steps 15 to 17).
RNA Folding Pathways
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ALTERNATE PROTOCOL
ELUTION OF SPLICED PRODUCTS FROM NATIVE POLYACRYLAMIDE GELS As an alternative to two-dimensional electrophoresis, the molecular weight of the RNAs in a native gel band may be determined by eluting the RNA from the gel matrix. The activity of the conformer can be determined by adding substrate to the gel slice before eluting the RNA. After elution, the RNA is concentrated by ethanol precipitation and analyzed on a denaturing polyacrylamide gel. Additional Materials (also see Basic Protocols 1 and 2) TEN buffer (see recipe) 10 mg/mL carrier tRNA (Sigma) 70% and 100% ethanol, –20°C TE buffer, pH 7.5 (APPENDIX 2A) Rocker or orbital mixer 4% to 6% denaturing polyacrylamide gel (see Basic Protocol 2) Recover RNA from native polyacrylamide gel 1. Separate RNA conformers by native gel electrophoresis (see Basic Protocol 1, steps 1 to 14), using 200,000 cpm [32P]RNA per sample. 2. Excise bands of RNA from native gel (see Basic Protocol 2, steps 1 to 4). Instead of excising the entire lane, individually remove each band to be analyzed and place in separate 1.5- or 2-mL microcentrifuge tubes. 3. Place 1 to 2 µL of 0.1 mM GTP in splicing buffer on gel slice and incubate for 2 min, room temperature. Add 2 to 4 µL of 2× urea loading buffer. It is useful to prepare control samples, in which this step is omitted, to determine whether any splicing occurred before native gel electrophoresis.
4. Freeze excised gel slices on dry ice 1 to 5 min. Thaw at room temperature or in a 25°C water bath. A freeze/thaw cycle improves recovery of RNA.
5. Add a sufficient volume of TEN buffer to cover gel piece (e.g., 0.3 to 0.6 mL). Place tightly sealed tubes on a rocker or orbital shaker and soak overnight at 4°C. In some cases, 30 to 60 min at 65°C improves recovery of RNA.
6. Microcentrifuge 1 min at 10,000 × g, room temperature. Carefully transfer supernatant to a clean tube. Avoid transfering pieces of polyacrylamide. Solution may be filtered using a small spin-filtration device (0.45 mm) that is nuclease free.
7. Add a small amount of TEN buffer (e.g., 50 to 100 µL) to gel slices, mix thoroughly, and remove as in step 6. Pool supernatants. Elution of radiolabeled RNA from gel is easily checked using a standard survey meter.
Probing RNA Folding Pathways by RNA Fingerprinting
Concentrate RNA by ethanol precipitation 8. Precipitate RNA by adding 1 µL of 10 mg/mL carrier tRNA and three volumes ice-cold 100% ethanol to supernatants. Chill at –20°C 1 to 2 hr or overnight, or at –70°C for 10 min. 9. Recover RNA by microcentrifuging 20 to 30 min at 10,000 × g, 4°C. Discard supernatant with a pipet, being careful to avoid pellet.
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Current Protocols in Nucleic Acid Chemistry
10. Wash pellet with 70% ethanol and dry under vacuum. Analyze RNA by denaturing polyacrylamide gel electrophoresis 11. Dissolve dried pellets in 2 µL water and add 2 µL of 2× urea loading buffer. Adjust volumes as required so that the concentration of mately equal.
32
P in each sample is approxi-
12. Prepare one or more reference samples by diluting the original RNA sample (or molecular weight markers) in TE buffer, pH 7.5, so that the concentration of radioactivity matches that of the samples in step 11. Add an equal concentration of 2× urea loading buffer. 13. Load 4 µL of each sample into the wells of a 20 × 20–cm 4% to 6% denaturing polyacrylamide gel (see Basic Protocol 2, steps 9 to 10) and run at 30 W, 50°C, until xylene cyanol FF tracking dye reaches the bottom. Samples may be heated 1 min at 80°C before loading if desired.
CHEMICAL MODIFICATION OF RNA IN A NATIVE POLYACRYLAMIDE GEL
BASIC PROTOCOL 3
After native gel electrophoresis, RNA is modified in situ by dimethyl sulfate (DMS) or kethoxal. DMS reacts with unpaired adenosines (N1) and cytosines (N3). Kethoxal reacts with the N1 and N2 of unpaired guanosines. The position of the modified bases is determined by primer extension with reverse transcriptase. Additional Materials (also see Basic Protocols 1 and 2 and Alternate Protocol) Unlabeled RNA (see Support Protocol 2; see also UNIT 6.1) Dimethyl sulfate (DMS, Aldrich; obtain fresh and store at –20°C) 50% and 100% ethanol Stop buffer A (see recipe) 70 mg/mL kethoxal (ICN Biochemicals or Research Organics) in 20% ethanol (store at –20°C) Stop buffer B (see recipe) Buffered phenol (APPENDIX 2A) 24:1 (v/v) chloroform/isoamyl alcohol Additional reagents and equipment for primer extension (see Support Protocol 3) and for sequencing gels (UNIT 6.1 and APPENDIX 3B) CAUTION: Dimethyl sulfate (DMS) is a carcinogen and should be handled in a fume hood with gloves that are impermeable to nonpolar solvents.
Separate RNA conformers by native gel electrophoresis 1. Prepare a native gel (see Basic Protocol 1, steps 1 to 9). 2. Prepare 2 samples of 32P-labeled RNA (500,000 cpm) in 10 µL splicing buffer (see Basic Protocol 1, steps 10 to 12). 3. Prepare 3 samples containing 2 pmol each unlabeled RNA in the same manner. 4. Load samples into adjacent wells of a native polyacrylamide gel (see Basic Protocol 1, steps 13 to 15). When electrophoresis is complete, disassemble gel and obtain an autoradiogram (see Basic Protocol 2, steps 2 and 3).
RNA Folding Pathways
11.4.7 Current Protocols in Nucleic Acid Chemistry
Supplement 2
5. Using the position of labeled RNA in the autoradiogram as a guide, excise the regions of adjacent lanes containing unlabeled RNA that contain the bands of interest (see Basic Protocol 2, step 2). This step and all subsequent steps are carried out on ice.
6. Transfer each gel slice (containing a single band) to a separate 1.5- or 2-mL microcentrifuge tube. There should be three duplicate sets of gel slices containing unlabeled RNA.
Modify RNA with DMS 7. Dilute DMS 1:40 in 50% ethanol. Add 20 µL diluted DMS to each slice originating from one lane of native gel. Distribute as evenly as possible over the surface. Incubate 0.5 to 2 min on ice. The concentration of DMS should be adjusted so that an even ladder of primer extension products is obtained in steps 16 to 19.
8. Quench reaction with 300 µL stop buffer A. Modify RNA with kethoxal 9. Apply 20 µL of 70 mg/mL kethoxal to each gel piece from a second lane of the native gel. Incubate 3 to 8 min on ice. The incubation time should be adjusted as above.
10. Quench reaction with 300 µL stop buffer B. Elute RNA from gel 11. Add 300 µL stop buffer B to each gel piece from the third lane of the native gel. These will serve as unmodified controls.
12. Soak gel pieces several hours or overnight at 4°C. Recover RNA (see Alternate Protocol, steps 5 to 6). 13. Extract supernatants once with an equal volume of buffered phenol. Microcentrifuge 1 min at 10,000 × g, 4°C, to separate layers and carefully transfer all of upper aqueous phase to a clean microcentrifuge tube. 14. Extract aqueous phases with 24:1 chloroform/isoamyl alcohol and transfer upper aqueous phase to a new set of microcentrifuge tubes. 15. Precipitate RNA with 3 volumes of 100% ethanol and dry pellets (see Alternate Protocol, steps 8 to 10). RNA may be stored at −20°C as an ethanol suspension or as a dried pellet for up to a week.
Detect modified bases by primer extension 16. Dissolve RNA pellets in 2 to 10 µL water, so that their concentrations are roughly equal (neglecting any carrier RNA that may have been added). The relative amount of RNA present in each band of native gel can be estimated by quantifying distribution of radioactive samples in step 4. Probing RNA Folding Pathways by RNA Fingerprinting
17. Perform primer extension using AMV-RT (see Support Protocol 3). RNA that was not subjected to native gel electrophoresis can be used as a template for dideoxynucleotide sequencing reactions.
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Current Protocols in Nucleic Acid Chemistry
If sufficient RNA is recovered from the native gel, each sample can be divided into aliquots and used for several primer extension reactions.
18. Separate primer extension products on a 6% or 8% polyacrylamide sequencing gel (APPENDIX 3B). 19. Detect primer extension products by autoradiography or by using a phosphorescent imaging screen. Compare primer extension pattern of modified RNAs to unmodified controls to determine which bands are specific for reactions with DMS or kethoxal. Determine relative intensity of specific primer extension bands for each RNA conformer by comparing products of RNA samples that were recovered from different regions of the native gel. Differences in the extent of modification may be quantified using a phosphorimager or densitometer.
IN VITRO SYNTHESIS OF 32P-LABELED PRE-RNA WITH T7 RNA POLYMERASE
SUPPORT PROTOCOL 1
Uniformly radiolabeled pre-RNA is prepared by run-off transcription of plasmid DNA or oligonucleotide templates (Milligan and Uhlenbeck, 1980) with purified phage T7 RNA polymerase (Davanloo et al., 1984). The template DNA must be cleaved downstream of the desired RNA sequence with a restriction enzyme. Transcription is carried out for short times (i.e., 20 to 30 min) at 30°C to minimize self-splicing (Emerick and Woodson, 1993). Materials 1 µg/µL plasmid DNA containing a T7 promoter, digested with appropriate restriction enzyme(s) (e.g. Feinbaum, 2000; Tabor, 2000) 10× T7 RNAP buffer (see recipe) 20 mM spermidine⋅HCl (store at –20°C) 10× NTPs “low A” (see recipe) 10 to 20 µCi [α-32P]ATP (store at –20°C) 500 to 1000 U/µL T7 RNA polymerase (store at –20°C) TE buffer, pH 7.5 (APPENDIX 2A) or another buffer 30°C water bath Gel-filtration spin column (e.g., TE-100, Clontech), 4°C Liquid scintillation counter Scintillation fluid and vials NOTE: Buffers and reagents should be kept on ice after thawing. Steps 1 to 4 are carried out at 4°C except where noted.
Transcribe RNA 1. Add the following to a 1.5-mL microcentrifuge tube (40 µL total volume): 1 µL 1 µg/µL digested plasmid DNA 4 µL 10× T7 RNAP buffer 4 µL 20 mM spermidine⋅HCl 4 µL 10× NTPs “low A” 1 to 2 µL 10 µCi/µL [α-32P]ATP 25 to 24 µL H2O. For RNAs that are not self-splicing, the additional spermidine should be omitted.
RNA Folding Pathways
11.4.9 Current Protocols in Nucleic Acid Chemistry
Supplement 4
2. Mix gently by tapping with finger. Spin briefly in a microcentrifuge to bring contents to the bottom of tube, if necessary. 3. Add 1 µl of 500 to 1000 U/µL T7 RNA polymerase. Mix well. 4. Incubate at 30°C for 20 to 30 min. Short reaction times reduce accumulation of spliced products. For RNAs that do not self-cleave, better yields will be obtained by incubating 1 hr at 37°C.
Remove unincorporated nucleotide triphosphates 5. While the transcription reaction is proceeding, prepare a spin column at room temperature for each sample. Pre-equilibrate column with TE buffer, pH 7.5 (or another buffer). After the column has drained, spin 3 min at 3000 × g, room temperature, to remove all excess buffer. Avoid trapping air bubbles in column matrix. Drain column in clean microcentrifuge tube and avoid touching tip.
6. When transcription reaction is complete, immediately load entire sample onto top of spin column. 7. Spin 5 min at 3000 × g, room temperature, using a clean RNase-free tube to collect sample. 8. Discard column as solid radioactive waste. 9. Determine the yield of RNA by counting 1 or 2 µL eluate in a liquid scintillation counter. Place RNA on ice, or store at –20°C. One should obtain approximately 105 cpm/mL. The RNA may also be purified by denaturing gel electrophoresis. SUPPORT PROTOCOL 2
LARGER SCALE IN VITRO SYNTHESIS OF UNLABELED PRE-RNA WITH T7 RNA POLYMERASE Larger quantities (i.e., 0.5 to 2 nmol) of unlabeled RNA may prepared by increasing the scale of in vitro transcription reactions. Abortive initiation products and unincorporated nucleotide triphosphates are removed by gel-filtration chromatography. For some applications, self-splicing products must be removed by preparative polyacrylamide gel electrophoresis. Materials 1 µg/µL digested plasmid DNA (10 µg total) 10× T7 RNAP buffer (see recipe) 1 M spermidine⋅HCl 50 or 100 mM each of ATP, CTP, GTP, and UTP 500 to 1000 U/µL T7 RNA polymerase 0.5 M EDTA (APPENDIX 2A) 3 M sodium acetate, pH 5.0 (APPENDIX 2A) 100% ethanol G-50 gel-filtration chromatography resin (see recipe) TEN buffer (see recipe)
Probing RNA Folding Pathways by RNA Fingerprinting
15-mL sterile polypropylene culture tubes Empty 10-mL disposable column with frit (e.g., Pharmacia Biotech PD-10 or BioRad Econo column)
11.4.10 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Transcribe RNA 1. Add the following to a 15-mL sterile culture tube (2 mL total volume): 10 µL 1 µg/µL digested plasmid DNA 200 µL 10× T7 RNAP buffer 1 µL 1 M spermidine⋅HCl 20 µL each of 100 mM ATP, CTP, GTP, and UTP (1 mM each NTP final) 1.70 mL H2O. For RNAs that are not self-splicing, spermidine should be omitted.
2. Mix thoroughly by vortexing. Centrifuge briefly in a benchtop centrifuge to bring contents to bottom of tube. 3. Add 10 µl T7 RNA polymerase. Mix well (do not vortex). 4. Incubate 1 hr at 30°C. Non-self-cleaving RNAs may be transcribed over 2 hr at 37°C.
5. Add the following and mix well, 5 µL 0.5 M EDTA 0.2 mL 3 M sodium acetate, pH 5.0 6 mL 100% ethanol. Chill 2 hr or overnight at –20°C. Remove unincorporated nucleotide triphosphates 6. Recover RNA by centrifuging 30 min at 12,700 × g (9000 rpm in a Beckman JS13.1 rotor), 4°C. Drain pellet and dry completely under vacuum. 7. Resuspend pellet in 100 to 300 µL TEN buffer. If pellet does not dissolve easily, carefully disrupt solid material using the end of a disposable, nuclease-free pipet.
8. To prepare gel-filtration column, clamp empty 10-mL column body to ring stand or buret holder. Place a beaker underneath to catch effluent. 9. Pipet slurry of equilibrated G-50 resin into column, allowing buffer to drain freely as resin settles. Continue to add slurry until the bed volume is 10 mL. The column should be free of discontinuities or air bubbles. Pre-packed columns (5- to 10-mL) may be substituted if commercially available.
10. Wash column with an additional 2 to 3 mL TEN buffer. Allow buffer to drain until just level with the top of the column. Stop the flow by placing tight-fitting cover over the bottom tip of the column. The flow rate should be ~0.5 mL/min. The column must not be allowed to run dry.
11. Apply sample to a pre-equilibrated gel-filtration column and elute with additional TEN buffer. After the void volume (∼3 mL), collect eluate in ten 0.5-mL fractions. 12. Determine absorbance of fractions at 260 nm (dilute 100-fold), and pool the peak fractions. The RNA should elute in the first 2 to 4 fractions after the void volume.
13. Recover the nucleic acid from pooled fractions by ethanol precipitation and determine final yield from absorbance at 260 nm.
RNA Folding Pathways
11.4.11 Current Protocols in Nucleic Acid Chemistry
Supplement 2
SUPPORT PROTOCOL 3
DETECTION OF MODIFIED BASES BY PRIMER EXTENSION The positions of modified bases are determined by extension of a 32P-labeled primer with reverse transcriptase using the modified RNA from Basic Protocol 3 as a template. Positions of the modified bases are determined by running the primer extension reactions on a sequencing gel and comparing to sequencing lanes prepared using ddNTPs for chain termination. Differences in the secondary and tertiary structure of the RNA conformers probed in Basic Protocol 3 can be determined by comparing the intensities of the termination products. Refer to UNIT 6.1 for an alternate protocol and more comprehensive discussion of primer extension methods. Additional Materials (also see Basic Protocols 1 and 3) 5× annealing buffer (see recipe) 1 pmol/µL 5′ 32P-labeled sequencing primer (UNIT 6.1) Diluted RNA for primer extension (see Basic Protocol 3) Unmodified RNA (see Support Protocol 2) 5× ddATP, ddCTP, ddGTP, and ddTTP solutions, prepared separately at 0.4 mM in 1× annealing buffer 5× RT buffer (see recipe) Avian myoblastosis virus reverse transcriptase (AMV-RT; Life Sciences or Seikageiku America) 5× 4dNTP mix: 2 mM each of dATP, dCTP, dGTP, and dTTP combined in 1× annealing buffer 2× formamide loading buffer (see recipe) 48° and 65°C water baths or heating blocks Additional reagents and equipment for preparing and running 6% or 8% polyacrylamide sequencing gel (APPENDIX 3B) NOTE: The following steps are carried out on ice except where indicated. Thawed reagents should kept on ice throughout. Anneal complementary primer to RNA 1. Prepare primer cocktail (total 20 µL) by mixing the following: 4 µL 5× annealing buffer 8 µL 1 pmol/µL 5′ 32P-labeled sequencing primer 8 µL deionized water. Volumes given are for 20 reactions and can be adjusted as needed.
2. For each extension reaction, add 1 µL primer cocktail (0.4 pmol primer) to 2 µL diluted RNA (see Basic Protocol 3, step 16) in a 0.5-mL microcentrifuge tube. It is often helpful to carry out control reactions using 0.1 to 0.5 pmol RNA that has been modified with DMS or kethoxal in solution (UNIT 6.1).
3. Heat 3 min in a water bath or heating block at 65°C and place immediately on ice. Microcentrifuge briefly at maximum speed before opening tubes. Keep samples on ice. Better results are obtained with some primer/template combinations by heating 2 min at 80°C. Probing RNA Folding Pathways by RNA Fingerprinting
Prepare sequencing reactions 4. Mix the following in a 0.5-mL tube:
11.4.12 Supplement 2
Current Protocols in Nucleic Acid Chemistry
2 pmol unmodified RNA 2 µL 5× annealing buffer 2 µL 1 pmol/µL 5′ 32P-labeled primer deionized water to 10 µL. 5. Anneal primer as in step 3. 6. Divide the annealing reaction from step 5 into five 2-µL aliquots. To each aliquot add 1 µL of one the following: 1× annealing buffer, 5× ddATP, 5× ddCTP, 5× ddGTP, and 5× ddTTP. The four reactions containing dideoxynucleotide triphosphates serve as sequencing lanes; the reaction containing only buffer serves as a blank. The blank should have no visible bands on the sequencing gel, unless the RNA is partially degraded or has strong secondary structure that causes the reverse transcriptase to terminate.
Extend primer with reverse transcriptase 7. Prepare a RT cocktail by mixing the following: 6 µL 5× RT buffer 23 µL deionized water 1 µL AMV-RT (25 to 50 U). Add the enzyme last. Resuspend enzyme thoroughly in the buffer, but do not vortex and avoid foaming. 8. Add 30 µL of 5× 4dNTP mix (total 60 µL) and mix gently. 9. Begin primer extension by adding 2 µL RT cocktail to each 3 µL annealing reaction (steps 3 and 6). Immediately place each tube in a water bath at 48°C. Incubate 15 min. 10. Stop reactions by adding 5 µL of 2× formamide loading buffer. Samples are now ready to be analyzed on a sequencing gel (see Basic Protocol 3, steps 18 and 19). Samples may be stored at –20°C for one week, if desired.
REAGENTS AND SOLUTIONS Use deionized or distilled nuclease-free water in all recipes and protocol steps. To eliminate traces of RNase on glassware, rinse in RNase-free water and bake 2 hr or overnight at 150°C. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Annealing buffer, 5× 200 µL 1 M Tris⋅Cl, pH 8.3 (APPENDIX 2A; 250 mM final) 60 µL 4 M NaCl (300 mM final) 40 µL 1 M dithiothreitol (DTT; 50 mM final) 500 µL deionized water (800 µL total volume) Store up to 6 months at −20°C Formamide loading buffer, 2× 940 µL deionized formamide 40 µL 0.5 M EDTA, pH 8.3 (APPENDIX 2A; 20 mM final) 20 µL 2% (w/v) xylene cyanol 20 µL 2% (w/v) bromphenol blue Store up to one year at −20°C
RNA Folding Pathways
11.4.13 Current Protocols in Nucleic Acid Chemistry
Supplement 2
G-50 gel-filtration chromatography resin Gently disperse 5.5 g of G-50 M (medium) chromatography resin (Amersham Pharmacia Biotech) in 100 mL TEN buffer (see recipe). Allow to swell 1 to 4 hr, room temperature. Swirl mixture, allow to settle, and decant fines (see manufacturer’s instructions). Repeat if necessary. Adjust the volume of TEN buffer so that the total volume is 100 mL or twice that of the settled resin. Autoclave and store indefinitely at room temperature. Glycerol loading buffer, 5× 500 µL glycerol 20 µL 2% (w/v) xylene cyanol Bring up to 1 mL with H2O Store up to 1 year at –20ºC or 1 month at room temperature GTP in splicing buffer, 0.1 mM Dilute a 100 mM stock solution of GTP to 0.1 mM with 1× splicing buffer (see recipe) containing 6 mM MgCl2. Store in aliquots at –20° or –80°C. NTPs “low A,” 10× Obtain concentrated stocks (e.g., 50 or 100 mM) of each nucleoside triphosphate (ATP, CTP, GTP, and UTP). These may be prepared by dissolving 100 mg solid in 2 mL water at 4°C. Adjust the pH to 7.0 using small aliquots (i.e., 10 to 50 µL) of concentrated NaOH. The pH may be monitored with a microelectrode or by spotting 5 µL onto calibrated pH indicator strips. Determine the concentration from the UV absorbance of a 1:1000 dilution, and adjust to the desired final concentration with additional water. Store in aliquots at –80°C. To prepare working solution, mix 1.25 µL 100 mM ATP, 6.25 µL 100 mM CTP, 6.25 µL 100 mM GTP, 6.25 µL 100 mM UTP and bring final volume to 0.5 µL with water. Store at –20°C. Final concentrations are 0.25 mM ATP and 1.25 mM each CTP, GTP, and UTP.
RT (reverse transcriptase) buffer, 5× 100 µL 1 M Tris⋅Cl, pH 8.3 (APPENDIX 2A; 250 mM final) 30 µL 4 M NaCl (300 mM final) 20 µL 1 M dithiothreitol (DTT; 50 mM final) 60 µL 1 M magnesium acetate (150 mM final) 190 µL deionized water (400 µL total volume) Store up to 6 months at −20°C Splicing buffer, 5× 250 µL 1 M NaHEPES, pH 7.5 (final concentration 250 mM) 500 µL 1 M (NH4)2SO4 (final concentration 500 mM) 10 µL 0.5 M sodium EDTA, pH 8.3 (APPENDIX 2A; final concentration 5 mM) 240 µL distilled H2O (total volume 1 mL) Store up to 1 year at –20°C
Probing RNA Folding Pathways by RNA Fingerprinting
Stop buffer A 14 µL 2-mercaptoethanol (final concentration 0.2 M) 100 µL 3 M sodium acetate (APPENDIX 2A; final concentration 0.3 M) 40 µL 0.5 M sodium EDTA, pH 8.3 (APPENDIX 2A; final concentration 20 mM) 846 µL distilled H2O Mix well and store up to 3 months at –20°C
11.4.14 Supplement 2
Current Protocols in Nucleic Acid Chemistry
Stop buffer B 100 µL 3 M sodium acetate (APPENDIX 2A; final concentration 0.3 M) 40 µL 0.5 M sodium EDTA, pH 8.3 (APPENDIX 2A; final concentration 20 mM) 860 µL distilled H2O. Mix well and store up to 1 year at –20°C T7 RNAP (RNA polymerase) buffer, 10× 400 µL 1 M Tris⋅Cl, pH 7.5 (APPENDIX 2A; final concentration 400 mM) 150 µL 1 M MgCl2 (APPENDIX 2A; final concentration 150 mM) 20 µL 1 M spermidine⋅HCl (final concentration 20 mM) 50 µL 1 M dithiothreitol (DTT, APPENDIX 2A; final concentration 50 mM) 380 µL distilled H2O (total volume 1 mL) Mix well and store up to 3 months at –20°C TEN (Tris/EDTA/NaCl) buffer 10 mM Tris⋅Cl, pH 7.5 (APPENDIX 2A) 1 mM disodium EDTA (APPENDIX 2A) 250 mM sodium chloride Autoclave and store indefinitely at room temperature or up to 2 months after opening THEM (Tris/HEPES/EDTA/MgCl2) buffer, 10×, pH 7.4 157.08 g HEPES (final concentration 660 mM) 41.14 g Tris base (final concentration 340 mM) 0.36 g disodium EDTA (final concentration 1 mM) 20.32 g MgCl2 (final concentration 100 mM) 1 L H2O (final volume) Autoclave and store up to 1 year at room temperature Urea loading buffer, 2× 1.1 g urea (final concentration 10 M) 4 µL 2% (w/v) xylene cyanol 980 µL distilled H2O Dissolve urea at 50°C Store in 0.5-mL aliquots up to 1 year at –20°C COMMENTARY Background Information Native polyacrylamide gel electrophoresis is now well established as a method for characterizing stable nucleic acid–protein complexes (Fried and Crothers, 1981). Complexes that normally dissociate within several minutes were found to be stable during many hours of electrophoresis, apparently due to a “caging” effect of the polyacrylamide matrix (Fried and Crothers, 1981). More recently, this approach has been used to quantify the binding of small RNA substrates to the Tetrahymena ribozyme (Pyle et al., 1990). It has also been used to monitor conformational changes in large and small RNAs (e.g., LeCuyer and Crothers, 1993; Emerick and Woodson, 1994). The mobility of native RNA is correlated with its hydrodynamic radius (Orr et al., 1998), with compact structures migrating more rapidly
than extended forms. As a result, this method is most useful when analyzing RNA sequences that fold into defined tertiary structures. Although these protocols are optimized for analysis of a 657-nt RNA, electrophoretic conditions can be readily varied depending on the RNA to be studied. The author has successfully used gels containing 4% to 12% polyacrylamide and 3 to 10 mM MgCl2. Nonionic substrates such as guanosine may be incorporated into the gel matrix. RNA conformers or complexes that exchange on the 1- to 10-min time scale at room temperature can usually be resolved if the appropriate conditions are chosen. This method is not suitable for trapping shortlived intermediates, as 15 to 30 sec are required for samples to enter the polyacrylamide matrix (Pan and Woodson, 1998).
RNA Folding Pathways
11.4.15 Current Protocols in Nucleic Acid Chemistry
Supplement 2
Another advantage of native gel electrophoresis is that it is easily combined with a variety of methods for probing RNA structure, including chemical modification and photo-crosslinking (Branch et al., 1989). Chemical reagents are commonly used to map regions of the RNA that are accessible to the chemical probe (Ehresmann et al., 1987). The modified positions are either detected by treatment of RNA with aniline acetate, which results in strand scission, or by extension of a complementary DNA primer with reverse transcriptase (Inoue and Cech, 1985; Moazed et al., 1986). A number of organic and inorganic compounds and enzymes that modify nucleic acids have been used to footprint protein-DNA complexes within polyacrylamide gels (Law et al., 1987; Straney and Crothers, 1987), and these methods were extended to the analysis of RNA structure (Emerick and Woodson, 1994). The advantage of modifying RNA in the gel matrix is that the rate of conformational exchange remains low. The disadvantage is that reagents must diffuse through the gel, making it difficult to control reaction time. In addition, some reagents, such as diethylpyrocarbonate, are strongly inhibited by the gel components. An alternative approach that avoids some of these pitfalls is to modify the RNA before electrophoresis, as in modification interference (Pan and Woodson, 1998).
Critical Parameters An important parameter that affects separation of different conformational forms of RNA by electrophoresis is whether or not the confor-
mational change induces a significant change in the overall shape of the molecule (i.e., by altering tertiary interactions). Beyond this, it is important to ensure that the folded RNA is stable in the electrophoresis and sample buffers, and that the internal temperature of the gel remains below 12°C. For the Tetrahymena preRNA, the bands representing active and inactive conformers become blurred at 15°C, and are no longer separated at 20°C. Two-dimensional electrophoresis and in situ chemical modification require extensive handling of RNA. Even very slight levels of ribonuclease contamination can result in undesirable background at the end of the experiment. To obtain interpretable results, the concentrations of chemical modifying reagents and times of exposure must be adjusted empirically for each RNA sequence, so that each molecule is modified only once on average.
Anticipated Results Typical results from native gel electrophoresis of the Tetrahymena pre-RNA are illustrated in Figure 11.4.2. Addition of GTP and RNA substrates to the gel showed that only the more rapidly migrating species was competent to self-splice (Emerick and Woodson, 1994). Consistent with this result, nucleotides in the catalytic core of the intron were more susceptible to modification with DMS and kethoxal in the inactive conformer than in the active form. In general, a qualitative test of ribozyme activity is relatively straightforward, if a reasonable degree of separation in the native gel is obtained. In situ chemical modification of the
wt mut I N Probing RNA Folding Pathways by RNA Fingerprinting
Figure 11.4.2 Native 6% polyacrylamide gel of 32P-labeled Tetrahymena pre-RNA (657-nt). The RNA was annealed at 95°C before loading. Abbreviations: I, inactive pre-RNA; N, native (active) RNA; wt, wild type; mut, mutation (170C) that destablizes the active RNA structure.
11.4.16 Supplement 2
Current Protocols in Nucleic Acid Chemistry
Time Considerations
Law, R., Kuwabara, M.D., Briskin, M., Fasel, N., Hermanson, G., Sigman, D.S., and Wall, R. 1987. Protein-binding site at the immunoglobulin mu membrane polyadenylation signal: Possible role in transcription termination. Proc. Natl. Acad. Sci. U.S.A. 84:9160-9164.
All of the protocols described here can be carried out in 1 to 3 days. Two-dimensional gel electrophoresis requires a continuous 10- to 12-hr block of time.
LeCuyer, K.A. and Crothers, D.M. 1993. The Leptomonas collosoma spliced leader RNA can switch between two alternate structural forms. Biochemistry 32:5301-5311.
Literature Cited
Milligan, J.F. and Uhlenbeck, O.C. 1980. Synthesis of small RNAs using T7 RNA polymerase. Methods Enzymol. 180:51-62.
RNA is more challenging, and one can expect that several trials will be required before the optimal conditions are found.
Branch, A.P., Benedfeld, B.J., Paul, C.P., and Robertson, H.P. 1989. Analysis of ultraviolet-induced RNA-RNA cross-links: A means for probing RNA structure-function relationships. Methods Enzymol. 180:418-442.
Moazed, D., Stern, S., and Noller, H.F. 1986. Rapid chemical probing of conformation in 16 S ribosomal RNA and 30S ribosomal subunits using primer extension. J. Mol. Biol. 187:399-416.
Davanloo, P., Rosenberg, A.H., Dunn, J.J., and Studier, F.W. 1984. Cloning and expression of the gene for bacteriophage T7 RNA polymerase. Proc. Natl. Acad. Sci. U.S.A. 81:2035-2039
Orr, J.W., Hagerman, P.J., and Williamson, J.R. 1998. Protein and Mg2+-induced conformational changes in the S15 binding site of 16S ribosomal RNA. J. Mol. Biol. 275:453-464.
Ehresmann, C., Baudin, F., Mougel, M., Romby, P., Ebel, J.P., and Ehresmann, B. 1987. Probing the structure of RNAs in solution. Nucl. Acids Res. 15:9109-9128.
Pan, J. and Woodson, S.A. 1998. Folding intermediates of a self-splicing RNA: Mispairing of the catalytic core. J. Mol. Biol. 280:597-609.
Emerick, V.L. and Woodson, S.A. 1993. Self-splicing of the Tetrahymena pre-rRNA is decreased by misfolding during transcription. Biochemistry 32:14062-14067. Emerick, V.L. and Woodson, S.A. 1994. Fingerprinting the folding of a group I precursor RNA. Proc. Natl. Acad. Sci. U.S.A. 91:9675-9679. Feinbaum, R. 2000. Introduction to plasmid biology. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 1.5.1-1.5.17. John Wiley & Sons, New York. Fried, M. and Crothers, D.M. 1981. Equilibria and kinetics of lac repressor-operator interactions by polyacrylamide gel electrophoresis. Nucl. Acids Res. 9:6505-6525. Inoue, T. and Cech, T.R. 1985. Secondary structure of the circular form of the Tetrahymena rRNA intervening sequence: A technique for RNA structure analysis using chemical probes and reverse transcriptase. Proc. Natl. Acad. Sci. U.S.A. 76:1670-1764.
Pyle, A.M., McSwiggen, J.A., and Cech, T.R. 1990. Direct measurement of oligonucleotide substrate binding to wild-type and mutant ribozymes from Tetrahymena. Proc. Natl. Acad. Sci. U.S.A. 87:8187-8191. Straney, D.C. and Crothers, D.M. 1987. Comparison of the open complexes formed by RNA polymerase at the Escherichia coli lac UV5 promoter. J. Mol. Biol. 193:279-292. Tabor, S. 2000. DNA-dependent RNA polymerases. In Current Protocols in Molecular Biology (F.M. Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith, and K. Struhl, eds.) pp. 3.8.1-3.8.4. John Wiley & Sons, New York.
Contributed by Sarah A. Woodson Johns Hopkins University Baltimore, Maryland
RNA Folding Pathways
11.4.17 Current Protocols in Nucleic Acid Chemistry
Supplement 2
Characterization of Tertiary Folding of RNA by Circular Dichroism and Urea
UNIT 11.5
Since X-ray crystallography of tRNA was first performed in the 1970s, the study of RNA folding has extended into the characterization of tertiary organization. Although circular dichroism (CD) spectroscopy cannot yet provide structural detail about tertiary organization, it can be used to monitor RNA tertiary folding transitions, which may not be observable by absorbance spectroscopy (Pan and Sosnick, 1997). With the use of computer-controlled titrators, data can be acquired rapidly and accurate thermodynamic parameters can be obtained over a wide variety of conditions. Hence, CD provides a nice complement to site-resolved methods such as complementary oligonucleotide hybridization (Zarrinkar and Williamson, 1994), hydroxyl radical footprinting (Sclavi et al., 1997), or chemical modification (Banerjee and Turner, 1995). This unit provides a basic outline for using CD and urea to measure and characterize tertiary RNA folding transitions to extract thermodynamic parameters. MEASUREMENT OF A CIRCULAR DICHROISM (CD) SPECTRUM The CD spectrum provides information about the amount of secondary and tertiary structure in the RNA. The changes in the spectrum can be used to identify folding transitions and determine their thermodynamic parameters. Some of these transitions, in particular tertiary transitions, may be unobservable by more standard absorbance measurements.
BASIC PROTOCOL 1
Materials Unfolded RNA sample UV-VIS spectrophotometer 1-cm-path-length quartz cuvette 90°C water bath Circular dichroism (CD) spectrometer capable of far-UV measurements (e.g., Jasco or AVIV Associates) Magnetic stir bar to fit cuvette Additional reagents and equipment for RNA renaturation (UNIT 6.3) Prepare sample 1. Prepare an unfolded RNA sample with absorbance at 260 nm (A260) of ∼0.3 to 0.8 AU for optimal signal-to-noise ratio. This corresponds to ∼10 to 20 µg/mL when using a 1-cm-path-length cuvette (1.5 mL). Increase concentration accordingly for shorter path lengths. The same cuvette can be used for the UV-VIS and CD spectrophotometers. Decreasing the RNA concentration will extend measurements further into the UV, but this also decreases the signal. Increasing the concentration will increase the signal, but this decreases the amount of light passing through the sample, and hence lowers the signal quality. Keep buffer and chloride concentrations to a minimum as these reagents absorb in the far-UV regions.
2. Perform a renaturing step (UNIT 6.3) involving heating to 90°C for 3 min, to remove residual structure formed during purification steps. RNA Folding Pathways Contributed by Tobin R. Sosnick Current Protocols in Nucleic Acid Chemistry (2001) 11.5.1-11.5.10 Copyright © 2001 by John Wiley & Sons, Inc.
11.5.1 Supplement 4
Measure CD and absorbance spectra 3. Set up the CD spectrometer using acquisition parameters approximately as follows: a. Set resolution or bandwidth at 2 nm. All features in the RNA spectrum are broad and resolution need not be higher than this. Decreasing the resolution increases the amount of light passing through the sample and improves signal quality.
b. Set time constant or response time (time period over which the signal is averaged for each data point) according to the manufacturer’s recommendation. Generally, this period should be small enough that the CD signal from wavelengths differing by an amount greater than the resolution (e.g., 2 nm) is not averaged in. Shorter time constants will result in decreased averaging and lower signal quality. For example, a scan rate of 20 nm/min is well matched to a response time of 4 sec at 2 nm resolution.
4. Scan from 200 to 320 nm. This is the spectral range which contains the RNA structural information. The range from 250 to 320 nm, however, generally contains enough information to characterize the relevant folding transitions. The wavelength scan speed (e.g. 20 nm/min) should be slow, to obtain good signal quality. If the quality is low, multiple scans can be averaged together. For a 20 µg/mL RNA sample with an A260 of ∼0.5 in a 1-cm-path-length cuvette, a 100-nm scan at 2 nm resolution with reasonable signal quality can be taken in ∼5 min using an instrumental time constant of 4 sec.
5. Record CD and absorbance simultaneously. Most commercially available CD spectrometers simultaneously record CD and the amplitude of the applied photomultiplier voltage (high tension). The latter can be converted to absorbance, enabling one to measure transitions by both CD and absorbance without any additional measurements. See the manufacturer’s operational manual for the procedure for converting from high tension to absorbance.
6. Put CD spectrum on an absolute scale normalized to the number of nucleotides, in order that proper comparisons to other spectrum can be made, using the formula for the molar circular dichroic absorption (∆ε in cm2/mmol): ∆ε =
θ 32,980 × C × L × N
Equation 11.5.1
where θ is the measured (raw) CD amplitude in mdeg, C is the sample concentration in mol/L, L is the cell path length in cm, and N is the number of nucleotides of the RNA. BASIC PROTOCOL 2
Characterization of Tertiary Folding of RNA by Circular Dichroism and Urea
MEASUREMENT AND ANALYSIS OF Mg2+-INDUCED FOLDING TRANSITIONS The tertiary folding of many RNAs is intimately coupled to the binding of divalent cations, typically Mg2+ (Fang, et al., 1999). After the 90°C renaturing step conducted in the absence of cations, the initial state of the RNA may contain some residual secondary structures. Upon addition of metal, further secondary structure as well tertiary structure will form. Near-UV circular dichroism can be used to follow the Mg2+ dependence of the folding transitions. The protocol is performed using two titrations. In the first, a coarse Mg2+ titration is used to determine which and how many wavelengths are appropriate for monitoring each folding transition. A second, finely spaced Mg2+ titration is then performed. Thermodynamic parameters of each transition are obtained from an analysis of the changes in signal as a function of Mg2+ concentration. The data from the second
11.5.2 Supplement 4
Current Protocols in Nucleic Acid Chemistry
titration are fit to a binding equation to obtain two parameters, the dissociation constant, Kd, and Hill coefficient, n. The Mg2+ transition midpoint will be written as KD (units of molarity) in this unit and is related to the dissociation constant by (KD)n = Kd (Fig. 11.5.1). Materials Unfolded RNA sample 10 mM and 1 M magnesium stock solutions (ultrapure, autoclaved) Plastic capillary tubing Calibrated gas-tight glass syringe Magnetic stir bar to fit cuvette Additional reagents and equipment for determining CD spectrum (see Basic Protocol 1) 1. Perform an initial coarse Mg2+-dependence series starting from the unfolded, Mg2+free RNA followed by a series of spectra at increasing Mg2+ concentrations. Add small volumes (e.g., 2 to 20 µL) of a concentrated Mg2+ solution to a single sample and correct for the resulting dilution. To avoid excessive dilution, add stock solution of 10 mM MgCl2 until the RNA sample is near 100 mM Mg2+, at which point the 1 M MgCl2 solution should be used to increase Mg2+ concentration. Monitor absorbance and CD (see Basic Protocol 1) at each concentration point. Generally, an unfolded to intermediate transition occurs at micromolar Mg2+ concentrations and is readily monitored by changes in absorbance (A260) and CD at 260 nm (∆ε260). These signals are primarily sensitive to the formation of helical structure. Subsequent tertiary transitions generally occur in the millimolar Mg2+ range. These may only be measurable by changes in CD.
2. Identify number of structural transitions and choose wavelengths to monitor each transition based upon its particular spectral changes. An increase in the CD signal and a decrease in the absorbance near 260 nm are primarily due to base stacking accompanying helix formation. Changes in the CD signal in the region from 275 to 290 nm are useful to monitor tertiary folding transitions. The magnitude of the CD changes depends upon wavelength and can be in the range of 0.1 to 3 cm2/mmol. The magnitude of the absorbance changes can be 0.01—0.3 × 106 M–1 cm–1. Often the changes are <10% of the initial CD or absorbance signal. Hence, high-quality data and accurate dilution corrections are a requisite for proper analysis of the folding transitions.
Figure 11.5.1 Sample Mg2+-induced folding transition fit using Equation 11.5.4 and n1 = 1.0, n2 = 4, KD1 = 0.4 mM, and KD2 = 6.1 mM Mg2+.
RNA Folding Pathways
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3. Record CD and absorbance (see Basic Protocol 1) at wavelengths determined with the coarse Mg2+ titration. Add the least volume possible of concentrated Mg2+ solution so that correction for dilution of the RNA will be minimal. Use multiple Mg2+ stock solutions (e.g., 10 mM and 1 M Mg2+) to optimize coverage of each transition. The sensitive titration may require that over fifty separate measurements be conducted, each following the addition of small quantities (<10 µL) of the concentrated Mg2+ stock solutions. Rapid additions without opening the sample compartment can be made through a small-bore HPLC-style capillary tubing connected to calibrated glass syringe. To further increase accuracy, the glass syringe can be attached to manual push-button repeating dispenser (available from Hamilton) that can reliable dispense 1/50 of the syringe volume per increment. To further expedite the titration process, an electronically controlled titrator (also from Hamilton) can be programmed and interfaced to the spectrophotometer.
4. Correct the CD signal for dilution of RNA due to added MgCl2. Fit the Mg2+-induced transition between states A and B to a binding curve according to the following equation: θ(Mg2+) = θA +
[Mg2+]n [Mg2+]n + (KD)n
(θB − θA)
Equation 11.5.2
where θA and θΒ are the signals of states A and B, respectively. For two well-separated transitions Α→B and B→C, fit the signal as the sum of two single transitions: θ(Mg2+) = θA +
[Mg2+]n1 [Mg2+]n1 + (KD1)n1
(θB − θA) +
[Mg2+]n2 [Mg2+]n2 + (KD2)n2
(θC − θB)
Equation 11.5.3
where n1 and KD1, and n2, and KD2 are the Hill coefficients and midpoints of the first and second transitions, respectively, and θA, θB, and θC are the CD signals of states A, B, and C (Fig. 11.5.1). For two overlapping transitions where all three species are significantly populated, fit the signal to reflect this linked equilibrium according to: θ(Mg2+) =
θA + θB([Mg2+]/KD1)n1 + θC([Mg2+]/KD1)n1([Mg2+]/KD2)n2 1 + ([Mg2+]/KD1)n1 + ([Mg2+]/KD1)n1([Mg2+]/KD2)n2 Equation 11.5.4
Equation 11.5.4 reduces to Equation 11.5.3 when KD1 << KD2. BASIC PROTOCOL 3
MEASUREMENT AND ANALYSIS OF A UREA TITRATION The nonionic denaturant urea can be used to denature both secondary and tertiary RNA structures and a urea titration can determine the free energy and size of a folding transition (Shelton et al., 1999). This protocol involves measuring the CD signal at the appropriate wavelength for each sample at varying urea concentrations over an extended range, e.g., 0 to 7 M. The required concentration range will depend upon the size of the RNA and must be wide enough that accurate pre- and post-transition baselines can be observed. Other solvent conditions such as Mg2+ concentration and pH must remain constant.
Characterization of Tertiary Folding of RNA by Circular Dichroism and Urea
11.5.4 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Materials Unfolded RNA sample Urea (ultrapure; filter through 0.2-µm filter) Additional reagents and equipment for determining CD spectrum (see Basic Protocol 1) Prepare urea/RNA stock solutions 1. Prepare two urea solutions, one at low (e.g., 0.7 M) and one at high (e.g., 7 M) concentration. 2. Make a stock of 10× concentrated unfolded RNA in either the low- or high-molarity urea solution. 3. Add one part of the 10× concentrated sample to nine parts of each of the low- and high-molarity urea solutions. Final RNA concentration should be such that A260 is equal to 0.3 to 0.8 absorbance units. This ensures that each urea solution contains the same RNA concentration, thereby ensuring that mixtures of these two solutions are at the same RNA concentration and that the CD signal will not need to be corrected for varying RNA concentration.
Measure CD spectra on a series of urea concentrations 4a. Method a: Prepare individual samples at varying urea concentrations by mixing the two stock solutions at varying ratios (see Table 11.5.1). Measure each of these samples separately (see Basic Protocol 1). To make up 21 separate 1.5-mL samples, at least 15 mL of each stock solution are required.
Table 11.5.1
Sample number 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21
Preparation of Individual Samples for Urea Titration
Vol. 0.7 M urea solution (mL) 1.5 1.425 1.35 1.275 1.2 1.125 1.05 0.975 0.9 0.825 0.75 0.675 0.6 0.525 0.45 0.375 0.3 0.225 0.15 0.075 0
Vol. 7.0 M urea solution (mL) 0 0.075 0.15 0.225 0.3 0.375 0.45 0.525 0.6 0.675 0.75 0.825 0.9 0.975 1.05 1.125 1.2 1.275 1.35 1.425 1.5
Final urea conc. (M) 0.7 1.015 1.33 1.645 1.96 2.275 2.59 2.905 3.22 3.535 3.85 4.165 4.48 4.795 5.11 5.425 5.74 6.055 6.37 6.685 7
RNA Folding Pathways
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Supplement 4
4b. Method b: Prepare single sample initially containing the dilute urea solution and measure spectrum. Add small aliquots (e.g., 50 µL) of the concentrated urea solution (see Table 11.5.2) and measure spectrum after each addition (see Basic Protocol 1). This method produces more accurate data and uses ∼5- to 10-fold less sample than method a. The size of each aliquot determines the urea concentration spacing of the data. Additions through a small-bore HPLC-style capillary tubing connected to calibrated glass syringe can improve both speed and accuracy of this method.
4c. Method c: Perform as in step 4b, but remove a small aliquot prior to the addition of an equivalent volume aliquot (e.g., remove 50 µL, then add 50 µL) so that the total sample volume remains constant. This variation reduces the total amount of sample required while increasing the range of the urea titration for a given number of additions (see Table 11.5.3). A two-syringe programmable titrator capable of both removal and additions can greatly facilitate the implementation of this technique.
Analyze data 5. To determine the free energy between states A and B in the absence of urea, ∆GoH2O, and its dependence on denaturant according to ∆Go (urea) = ∆GoH2O, fit the data to: θ(urea) =
o θA + θBe(−∆GH2O−m [urea])/RT o
1 + e(−∆GH2O−m [urea])/RT Equation 11.5.5
where R is the gas constant (1.987 cal/K/mol), T is the absolute temperature (K), and θA and θB are the baseline values of states A and B, respectively (Fig. 11.5.2). Table 11.5.2
Add-Aliquot Variation for Preparation of Samples for Urea Titrationa
Addition number
Characterization of Tertiary Folding of RNA by Circular Dichroism and Urea
0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20
Net vol. 7.0 M urea solution added (mL) 0 0.05 0.1 0.15 0.2 0.25 0.3 0.35 0.4 0.45 0.5 0.55 0.6 0.65 0.7 0.75 0.8 0.85 0.9 0.95 1
Final urea conc.b (M) 0.7 0.90 1.09 1.27 1.44 1.6 1.75 1.89 2.03 2.16 2.28 2.39 2.5 2.60 2.70 2.8 2.89 2.98 3.06 3.14 3.22
a
Starting condition is 1.5 mL of 0.7 M urea. Calculated according to C = [Volinit × (Cinit + Voladd) × Cadd)/(Volinit + Voladd), where C is the urea concentration, Volinit and Voladd are the initial and added volumes, and Cinit and Cadd are the urea concentrations of the initial and added solutions, respectively. b
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Current Protocols in Nucleic Acid Chemistry
Table 11.5.3 Remove/Add-Aliquot Variation for Preparation of Samples for Urea Titrationa
Final urea conc.b (M) 0.70 0.93 1.14 1.35 1.55 1.75 1.94 2.12 2.29 2.46 2.62 2.78 2.93 3.07 3.21 3.35 3.48 3.61 3.73 3.84 3.96
Addition no. 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20
a Starting condition is 1.5 ml of 0.7 M urea, solution added is 7 M urea, with removal and addition in 50-µL aliquots. b Calculated according to the recursive relation Ci = [(Volinit – Voladd) × Ci-1 +Voladd × Cadd]/Volinit, where C is the urea concentration, Volinit and Voladd are the initial and added volumes, and Ci−1 and Cadd are the urea concentrations of the previous and added solutions, respectively.
a. The baseline values may be a constant value, or may be approximated as a line, e.g., θA(urea) = mA[urea]+ θHA O. For two transitions from A→B and B→C with stabilities ∆GoA→B (urea) = ∆GoA→B (0)+moA→B[urea ] and ∆GoB→C(urea) = ∆GoB→C (0)+moB→C[urea], the signal can be fit according to: 2
Figure 11.5.2 Sample urea denaturation profile fit using Equation 11.5.5 with ∆Go= 9.3 kcal/mol and mo = 2.5 kcal/mol/M.
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o
θ(urea) =
o
θA + e−∆GA−>B(urea)/RT(θB + θCe−∆GB→C(urea)/RT) o
o
1 + e−∆GA→B(urea)/RT(1 + e−∆GB→C(urea)/RT) Equation 11.5.6
where θA, θB, and θC are the ellipticities of states A, B, and C (e.g., θA(urea) = mA[urea] + θHA O). 2
b. For broad transitions, it can be difficult to simultaneously fit the m value and a sloping baseline with confidence. An alternative method to determine the m value is to use multiple Mg2+ titrations to measure the KD at different, fixed concentrations of urea. The change in free energy due to the addition of urea at any given Mg2+ concentration can be written as the difference of the stability before and after addition of urea: n O H2O 2 K KH D D o o o = −nRT ln ∆∆G (urea) = ∆GH O − ∆G (urea) = −RT ln 2 KD(urea) KD(urea)
Equation 11.5.7
A plot of ∆∆G (urea) versus urea concentration can be fit to the linear relation shown in Equation 11.5.11. o
COMMENTARY Background Information
Characterization of Tertiary Folding of RNA by Circular Dichroism and Urea
The characterization of the thermodynamic parameters of a tertiary RNA is fundamental to understanding its folding behavior. A common method for measuring the stability of RNAs is thermal denaturation. From thermal melting curves or differential scanning calorimetry, the enthalpy, free energy, and entropy can be determined, making these powerful tools in the study of RNA thermodynamics. However, thermal denaturation of Mg2+-dependent RNAs may not be reversible due to Mg2+-induced degradation at high temperatures. Further, the thermal melting of an RNA tertiary structure can be difficult to separate from the melting of the secondary structure. Consequently, other methods to measure RNA secondary and tertiary stability are desirable. This unit presents spectroscopic methods for obtaining the Mg2+ midpoint and the effective number of Mg2+ ions bound in the transition. These properties can be used to obtain the tertiary stability of the RNA. The use of urea titrations provides an alternative to obtain these quantities, as well as a measure of the amount of surface buried in the transition, the urea m value. Much of the analysis required for the proper extraction of the apparent thermodynamic parameters requires the use of an analysis program capable of nonlinear least-squares fitting. Many excellent plotting and analysis packages
are commercially available (e.g., Origin by Microcal). The basis of Equation 11.5.2, Equation 11.5.3, and Equation 11.5.4 is the following, which assumes that Mg2+ ions are required and that they specifically bind in the folding of a tertiary RNA. For a single transition between states A and B where n Mg2+ ions are bound cooperatively: Kd
A + nMg2+ ← B Equation 11.5.8
The dissociation constant, Kd, and the Mg2+ midpoint, KD, can be written as: Kd = (KD)n =
[Mg2+]n [A] [B]
Equation 11.5.9
The Mg2+-dependent equilibrium free energy, ∆Go, between states A and B is the logarithm of the ratio of their populations and can be written: ∆Go = −RT ln Keq = −RT ln
[B]
[A]
n
[Mg2+] [Mg2+] = −RT ln = −nRT ln K KD D Equation 11.5.10
where Keq is the equilibrium constant between states A and B.
11.5.8 Supplement 4
Current Protocols in Nucleic Acid Chemistry
It should be realized that the Hill coefficient identifies the minimal number of Mg2+ ions involved in the transition. However, a Hill coefficient of unity can be attributed to multiple, independent site binding of approximately the same Kd, so that the total number of Mg2+ ions bound is unknown. This number can be determined by other methods including equilibrium dialysis. Urea denatures RNAs by preferentially stabilizing the unfolded state; however, the precise mechanism is unknown. Urea may promote unfolding either by forming hydrogen bonds with newly exposed carbonyls and nitrogens on the bases, increasing the structure of water and making the exposure of hydrophobic regions (e.g., the aromatic rings) less favorable, or by weakly binding to the RNA, with the number of urea binding sites increasing upon unfolding (Makhatadze and Privalov, 1992). Regardless of which modes are operational, empirically the stability of secondary and tertiary RNA structures can be well approximated with a linear dependence upon urea concentration at a fixed Mg2+ concentration: ∆Go(urea) = ∆GoH2O − mo[urea] = −RT ln Keq(urea) = −RT ln
[B] [A]
θ(urea) − θA = −RT ln θB − θ(urea) Equation 11.5.11
For proteins, the slope mo is proportional to the amount of urea-sensitive surface buried upon folding and is directly related to the size of the protein (Myers et al., 1995). Ongoing studies in the author’s laboratory suggest that this is true for RNAs as well (Shelton et al., 1999).
Critical Parameters and Troubleshooting RNA is very sensitive to degradation by ribonucleases. Caution should be used to avoid contamination, including the use of latex gloves and autoclaved reagents. Sample cuvettes and other equipment such as HPLC tubing should be designated for RNA usage only, to whatever extent feasible. Measure buffer blanks in the same cuvette orientation for proper background subtractions. With viscous urea solutions, extreme care must be taken to ensure adequate mixing of the different-density solutions. Ensure thorough mixing with a magnetic stirrer. A simple method to test for proper mixing is to confirm
that the trace for dilutions going from low to high urea concentration matches that for dilutions from high to low concentration. This procedure also confirms that the folding is reversible. Thermodynamic measurements require that the system be reversible and in complete equilibrium during the measurement. Tertiary RNA folding transitions can take from tens of minutes to hours, and may be very sensitive to experimental conditions (e.g., temperature, Mg2+, and urea concentration). Slow transitions often have a high activation enthalpy and can be greatly accelerated with temperature.
Anticipated Results These experiments require tens of micrograms of RNA per titration. From the measurement of a Mg2+ titration, the number of equilibrium folding transitions and their Mg2+ midpoints and Hill coefficients can be determined. From this information, the stability of an RNA can be determined at any Mg2+ concentration. Similarly, from a urea melting measurement starting from a folded RNA, the stability and the amount of urea-sensitive surface area buried (mo value) in the transition can be determined at a given temperature and Mg2+ concentration.
Time Considerations These measurements can be performed in a day or less. The use of an automated titrator will greatly accelerate the rate of data collection and multiple titrations can be conducted in a day.
Literature Cited Banerjee, A.R. and Turner, D.H. 1995. The time dependence of chemical modification reveals slow steps in the folding of a group I ribozyme. Biochemistry 34:6504-6512. Fang, X., Pan, T., and Sosnick, T.R. 1999. A thermodynamic framework and cooperativity in the tertiary folding of a Mg2+-dependent ribozyme. Biochemistry 38:16840-16846. Makhatadze, G.I. and Privalov, P.L. 1992. Protein interactions with urea and guanidinium chloride. A calorimetric study. J. Mol. Biol 226:491-505. Myers, J.K., Pace, C.N., and Scholtz, J.M. 1995. Denaturant m values and heat capacity changes: Relation to changes in accessible surface areas of protein unfolding [published erratum appears in Protein. Sci. 1996 May;5(5):981]. Protein Sci. 4:2138-2148. Pan, T. and Sosnick, T.R. 1997. Intermediates and kinetic traps in the folding of a large ribozyme revealed by circular dichroism and UV absorbance spectroscopies and catalytic activity. Nat. Struct. Biol. 4:931-938.
RNA Folding Pathways
11.5.9 Current Protocols in Nucleic Acid Chemistry
Supplement 4
Sclavi, B., Woodson, S., Sullivan, M., Chance, M.R., and Brenowitz, M. 1997. Time-resolved synchrotron X-ray “footprinting,” a new approach to the study of nucleic acid structure and function: Application to protein-DNA interactions and RNA folding. J. Mol. Biol. 266:144-159. Shelton, V., Sosnick, T.R, and Pan, T. 1999. Applicability of urea in the thermodynamic analysis of secondary and tertiary RNA folding. Biochemistry 38:16831-16839.
Zarrinkar, P.P. and Williamson, J.R. 1994. Kinetic intermediates in RNA folding. Science 265:918924.
Contributed by Tobin R. Sosnick University of Chicago Chicago, Illinois
Characterization of Tertiary Folding of RNA by Circular Dichroism and Urea
11.5.10 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays
UNIT 11.6
This unit describes the use of a synchrotron X-ray beam to carry out hydroxyl radical footprinting of RNA. The cleavage reactions typically require 10 to 50 msec of exposure to the X-ray beam. Hence, this method is suitable for probing kinetic folding intermediates or other transient conformational states that appear on the 50-msec to 100-sec timescales. The protocols given here were developed to probe the folding pathway of the Tetrahymena ribozyme (Sclavi et al., 1997, 1998a), but are applicable to a variety of catalytic RNAs or RNA-protein complexes. A facility for X-ray footprinting is operated by the Center for Synchrotron Biosciences at beamline X28C of the National Synchrotron Light Source (NSLS) at Brookhaven National Laboratory. Equilibrium experiments are conducted by exposing the RNA to the X-ray beam using a sample holder and electronic shutter (see Basic Protocol 1). Kinetic experiments are carried out using a commercial rapid mixing apparatus modified to withstand high-flux X-radiation (see Basic Protocol 2). RNA folding reactions are initiated by rapidly mixing the RNA with buffer containing MgCl2. After a defined interval, the RNA is pushed into a flow cell in the X-ray beam and irradiated for a short time. Cleavage products are collected and analyzed by polyacrylamide gel electrophoresis (see Basic Protocol 3). CAUTION: Prospective users must inform themselves of the training and safety requirements at individual synchrotron facilities. The following procedures should only be performed by personnel who have been trained in the operation of X-ray beamlines and the use of radioactive materials, and received authorization by the operating facility. Standard precautions to minimize exposure to β rays and prevent radioactive contamination of personnel and equipment should be followed at all times. NOTE: Care must be taken to avoid introducing ribonucleases into the samples. This can usually be achieved by purchasing good-quality machine-packaged disposable plasticware, using gloves while handling samples and preparing solutions, and storing disposables in clean, dust-free containers. All solutions should be prepared with deionized water (18 MΩ) that is free of pyrogens and organics (e.g., tissue culture grade). Although less desirable, water treated with diethylpyrocarbonate (DEPC; APPENDIX 2A) may be used if a source of nuclease-free water is not available. Solutions should be sterilized by filtration (0.2-µm filter) or autoclaved. NOTE: Nucleic acid or protein samples within the experimental hutch (except the sample in use) should be placed in a lead-lined container before activating the X-ray beam, to guard against degradation due to scattered radiation. STRATEGIC PLANNING These experiments require access to a synchrotron X-ray beamline that has been suitably modified for footprinting studies. Requirements for the beamline configuration are discussed briefly at the end of this unit and elsewhere (Sclavi et al., 1998b; Ralston et al., 2000b). Protocols in this unit describe experiments conducted at beamline X28C at the NSLS, using equipment available to beamline users. Information regarding facilities maintained by the Center for Synchrotron Biosciences at the NSLS can be obtained at http://www.aecom.yu.edu/home/csb/. RNA Folding Pathways Contributed by Sarah A. Woodson, Michael L. Deras, and Michael Brenowitz Current Protocols in Nucleic Acid Chemistry (2001) 11.6.1-11.6.24 Copyright © 2001 by John Wiley & Sons, Inc.
11.6.1 Supplement 6
Synchrotron experiments must be planned 2 to 6 months in advance to allow users sufficient time to complete safety and equipment training before beginning work. Advance planning is particularly important if radioactive materials are to be used. It is essential to ensure that the required equipment is available and, if necessary, to allow for construction or installation of additional equipment. Before undertaking experiments at the synchrotron, it is important to determine whether the RNA or RNA-protein complex of interest can be studied effectively by hydroxyl radical footprinting. This is best determined by conducting preliminary experiments with Fe(II)-EDTA, which also generates hydroxyl radicals (UNIT 6.5). The cleavage patterns induced by Fe(II)-EDTA and X-rays are indistinguishable, although Fe(II)-EDTA reactions proceed more slowly. The experimental conditions (i.e., temperature, Mg2+ concentration, and protein concentration) should be adjusted to maximize the extent of protection. In addition, one should determine whether the transition is complete within 60 sec. Conformational changes that take longer than 60 sec are more easily probed by chemical methods (see Commentary). It is recommended that one begins with X-ray footprinting experiments under equilibrium conditions (see Basic Protocol 1). These experiments can be used to confirm that the optimum conditions have been selected, and establish the equilibrium parameters of the transition. The required X-ray exposure time for the sample must be determined from a dose-response curve and consideration of the current in the synchrotron ring at the time of the experiment (see Support Protocol 2). This information is indispensable in planning time-resolved experiments (see Basic Protocol 2). BASIC PROTOCOL 1
EQUILIBRIUM X-RAY FOOTPRINTING OF RNA This protocol is useful for performing titrations with Mg2+, Na+, urea, or protein. A temperature-controlled aluminum block is used to hold individual samples in the X-ray beam. An electronic shutter controls the exposure time. Materials 32 P end-labeled RNA in 6-µCi aliquot (see Support Protocol 1) CE buffer, pH 7.5 (see recipe) or other appropriate buffer (see Critical Parameters) 1 M MgCl2 (APPENDIX 2A; optional) Precipitation cocktail (see recipe) 100% ethanol Aluminum sample holder with electronic shutter and support stand (see Commentary, Figure 11.6.1) Support table Controller cable Detector for the automatic vertical alignment device Refrigerated recirculating bath with attachment tubes 1.5-mL and 0.5-mL microcentrifuge tubes with captive screw caps and O-ring seals (Rainin) Linagraph paper (Kodak) Masking tape Temperature-controlled heating block or water bath Lead sample box
Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays
CAUTION: Cacodylic acid is an arsenic compound and is toxic.
11.6.2 Supplement 6
Current Protocols in Nucleic Acid Chemistry
Set up shutter and sample holder 1. Install the shutter and sample holder assembly so that the sample tube is aligned with the X-ray beam (Figure 11.6.1). At beamline X28C, bolt the stand assembly to the support table, and connect the controller cable to the shutter. Attach the detector for the automatic vertical alignment device to the sample holder and plug in its output cable. Check that the shutter and stand are aligned with each other and horizontally aligned with the end of the beampipe. 2. Attach the tubes from a refrigerated recirculating water bath to the aluminum sample holder. Turn on the water bath and adjust the temperature as desired. Water should enter the bottom port on the sample block and exit the top port.
3. Extend the retractable flight tube from the end of the beampipe to the face of the shutter assembly. The flight tube reduces scattered X-ray radiation (see Commentary).
4. Engage the beamline interlock system, exit the hutch, and enable the X-ray beam. Experimental hutches at the NSLS are equipped with an interlock system to prevent accidental exposure of personnel. Procedures for enabling the beam are established by the synchrotron facility administration.
5. Perform the semi-automatic alignment protocol according to instructions on the Center for Synchrotron Biosciences website (http://www.aecom.yu.edu/home/csb/). Shut off the beam, and place the shutter in a closed position. 6. Confirm alignment of the beam manually by covering the sample tube hole with linagraph paper and masking tape. Activate the beam, and expose the paper for 1 sec. The burn mark on the paper should be centered over the hole, and should cover the area that is occupied by the sample.
Prepare RNA samples 7. Thaw one 20-µL aliquot of 32P end-labeled RNA (6 µCi) at room temperature. Microcentrifuge 15 sec at maximum speed. RNA samples should be prepared and shipped to the beamline a day in advance (see Support Protocol 1). Store at −20°C or on dry ice in a lead container when not in use.
8. Add 30 µL of the appropriate buffer (typically CE buffer) to each aliquot, bringing the total volume per aliquot to 50 µL. Vortex the samples and microcentrifuge 15 sec at maximum speed.
Figure 11.6.1 Stand with aluminum sample holder and automatic shutter for equilibrium footprinting experiments. Adapted from Ralston et al. (2000b) with permission from Harcourt.
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9. Label twenty 0.5-mL microcentrifuge tubes. Add 2.5 µL RNA from step 8 to each tube. 10. Add the desired buffer or reagents (2.5 µL) to each tube. For example, to measure the Mg2+-dependence of RNA folding, prepare a series of solutions containing 2× MgCl2 in CE buffer. Add 2.5 mL of 2× MgCl2 to eighteen samples. Add 2.5 mL of CE (no Mg2+) to two samples to serve as unfolded controls. The total volume of each sample should not exceed 5 mL, to ensure that the entire sample remains within the cross-section of the beam.
11. Anneal the RNA by incubating the samples at the desired temperature until the folding reaction has reached equilibrium, or by heating 1 to 2 min at 85° to 95°C and cooling to the desired temperature. As an example, the Tetrahymena ribozyme should be incubated 20 min at 42°C or 2 to 4 hr at 30°C. Prolonged incubation of RNA at temperatures >50°C should be avoided to minimize hydrolysis.
Expose RNA to X-ray beam 12. Set aside one of the samples without MgCl2. This sample will not be placed in the X-ray beam and will serve as an unexposed control.
13. Place one tube in the sample block with the large end facing the beam. 14. Interlock and exit the hutch. Turn on the X-ray beam, and then activate the shutter to expose the sample. Turn off the X-ray beam once the exposure is complete. Each sample should be exposed for the same time. Exposure times are typically 10 to 50 msec at X28C, depending on the beam current. The optimal time should be determined from a dose-response experiment (see Support Protocol 2).
15. Re-enter the hutch and remove the sample from the holder. Add 15 µL distilled water, 5 µL precipitation cocktail, and 75 µL of 100% ethanol. Store the sample in a lead box. 16. Repeat steps 13 to 15 until all the samples have been irradiated. Clean up 17. Store samples at −20°C or on dry ice in a lead container. 18. Turn off and disconnect the water bath and the shutter controller. 19. Dismount the sample holder and shutter, and store the entire assembly in a secure place. 20. At the end of the experiments, complete all safety checks and disable the beamline. BASIC PROTOCOL 2
Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays
TIME-RESOLVED X-RAY FOOTPRINTING OF RNA This protocol describes a standard method for time-resolved X-ray-dependent hydroxyl radical cleavage of RNA. A Kin-Tek rapid mixing device is used to initiate the reaction of interest (such as Mg2+-induced folding). After a programmed delay, the RNA is pushed into a flow cell and exposed to the X-ray beam. A progress curve is produced by varying the delay time. Samples are processed and analyzed as described in Basic Protocol 3.
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Materials CE buffer, pH 7.5 (see recipe) or other appropriate buffer (see Critical Parameters) 2 to 8 µCi 32P end-labeled RNA, divided between two 20-µL aliquots (see Support Protocol 1) 100% ethanol Precipitation cocktail (see recipe) CE20 buffer (see recipe) Additional buffers or salts as desired (see Critical Parameters) 0.6 µCi RNA for prefolded controls (see Support Protocol 1) Rapid-quench apparatus, modified for X-ray footprinting experiments (e.g., RQF-3, Kin-Tek; see Commentary, Figure 11.6.2, and Support Protocol 3) 1-mL and 5-mL Luer-lok disposable syringes Temperature-controlled heating block or water bath 1.5-mL microcentrifuge tubes with captive screw caps and O-ring seals (Rainin) 13-G needles 15-mL sterile disposable culture tubes Lead-lined box Refrigerated recirculating water bath NOTE: It is important to become familiar with the operation of the rapid-quench apparatus (Figure 11.6.2) and the valve settings illustrated in Figure 11.6.3 (LOAD syringes, LOAD sample, FIRE, and FLUSH). For RNA folding experiments, drive syringe B is loaded with buffer (e.g., CE or CE20 buffer), and the RNA sample is placed in the bottom right sample loop. Drive syringe A and the bottom left sample loop are loaded with CE or CE20 buffer. The third Quench syringe (C) is not used in the standard footprinting protocol. The plunger for the C syringe should remain fully depressed. CAUTION: Care must be taken to protect personnel from β radiation and prevent contamination of work area. Gloves, equipment, and work surfaces should be frequently monitored using a hand-held survey meter. Safety procedures pertaining to use of the beamline, such as the interlock system, must be observed at all times. Caution should be used in handling the waste container, sample syringe port, and exit line, which may be contaminated with radioactive material.
Figure 11.6.2 Modified rapid-quench apparatus for time-resolved footprinting. Adapted from Sclavi et al. (1998b) with permission from Harcourt.
RNA Folding Pathways
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Figure 11.6.3 Valve settings for Kin-Tek rapid chemical quench. Adapted with permission from manufacturer’s manual.
Adjust drive platform 1. Set up the rapid-quench apparatus as described in Support Protocol 3. Open a software routine that allows one to adjust the position of the drive platform (option 2 in the control software). 2. Use the remote actuate (small red) button on the stepper-motor to raise the platform. The platform should just contact the plungers of drive syringes A and B when they are completely filled. One may calibrate and mark the desired starting position for the drive platform in advance.
Load drive syringes 3. If using a Kin-Tek apparatus, turn the top row of valves to the LOAD syringe position, and inject 1× CE buffer using a 5-mL disposable syringe. Work the solution back and forth to remove bubbles. Fill the A and B drive syringes in the same manner. 4. Turn the top valves so they are aligned vertically (FIRE). Remove the 5-mL disposable syringes. Anneal RNA 5. Thaw one 20-µL aliquot of 32P end-labeled RNA (1 to 4 µCi) at room temperature. Mix gently. 6. If desired, anneal the RNA by incubating 1 min at 95°C in a heating block. Microcentrifuge 15 sec at maximum speed. 7. Add 120 µL of 1× CE buffer to the sample (140 µL final volume). Mix well (vortex) and microcentrifuge briefly at maximum speed. This yields enough sample for a “priming shot” and nine experimental trials. Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays
Load sample syringes 8. Using a 1-mL disposable syringe, pull the plunger back and draw in 0.3 mL air. Then, draw up the entire diluted RNA sample prepared in step 7.
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The final drop of RNA solution can be transferred to the syringe tip using a pipet. The air in the syringe will be used to push the entire sample into the sample loop of the rapid-quench apparatus.
9. Turn the two bottom sample loop valves to the LOAD sample position. Attach the RNA sample syringe to the right sample port, and fill the tubing between the port and the valve. Leave the 1-mL syringe in place, as it will be used for subsequent experiments. The top valves should be in the FIRE position. Be sure that the vacuum line is disconnected from the exit tube.
10. Using a 1-mL syringe, fill the left sample loop with CE buffer. 11. Turn the bottom valves to the FIRE position. All valves should be in the FIRE position, except for drive syringe valve C, which remains in the LOAD position.
12. Remove the cap from a spare 1.5-mL screw-cap microcentrifuge tube and punch a hole through the cap using a 13-G needle. Thread the exit line of the quench-flow through the cap (it should fit snugly). Screw an empty microcentrifuge tube onto the exit line adapter, and place it in a tube rack on the aluminum shelf under the rapid-quench apparatus. This tube will be used to collect waste. An initial shot primes the drive lines and should be performed every time the drive syringes or sample loops are filled. It also avoids a timing error that occurs when the BASIC control program is started. (Only the first shot is affected by this error.) CAUTION: The sample syringe containing RNA should be covered with either leaded Plexiglas or separate Plexiglas and lead shields to protect the sample from X-rays and the investigator from b radiation.
Load sample loops and fire a priming shot 13. Turn the upper and lower valves to LOAD sample. Advance exactly 10 µL from each sample syringe into the left and right sample loops, respectively. The valve and sample loop should be precalibrated and marked to indicate the proper position of the fluid meniscus. The amount of solution is determined by the volume of the sample loops (see Commentary).
14. Turn all valves to the FIRE position. The valve to drive syringe C should remain in the LOAD syringes position, unless a quench solution is used.
15. Open the software routine for the time-resolved footprinting experiment. Set speed and distance of the stepper motor to achieve the desired folding delay and exposure times. For the dummy shot, enter a delay time of 0.01 sec. Each push of the drive platform is defined by the speed (rpm) of the stepper motor and a distance (the number of turns of the motor). Refer to Table 11.6.1 for a standard “four push, one pause” routine for synchrotron hydroxyl radical footprinting using a Kin-Tek apparatus. Refer to Table 11.6.2 for setting exposure times.
16. Once the parameters are set, type “G” at the controller keyboard to fire the drive motor. Before firing, be sure that a microcentrifuge tube is attached to the exit line. RNA Folding Pathways
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Table 11.6.1
Sample Parameters for Time-Resolved X-ray Footprinting at X28C
Speed (rpm) 180
Distance 400
Eventa Mix
0
0
Fold
Variable
600
Expose
180
500
Exit
50
500
Eject
a
Sample position Advanced through mixing T up to exposure cell Variable pause to allow additional folding time Advanced through exposure cell Speed determines exposure time Advanced up to the end of the exit line Ejected into collection tube
The time required for each step is given in commentary (see Background Information, discussion of instruction).
Flush mixing and exit loops 17. Turn the lower valves to FLUSH, and allow any sample remaining in the exit line to drain into the waste or sample collection tube. Alternatively, turn the valve under syringe C 180 degrees, and expel the sample from the exit line by blowing air through port C with an empty 1-mL syringe. Close the valve (return to LOAD syringes). The lines of the rapid mixing apparatus are flushed and rinsed after each shot.
18. Gently unscrew the microcentrifuge tube from the exit line. 19. Attach the vacuum to the exit line using the Luer adapter. If the valves are not set to the FLUSH position before attaching the vacuum, the contents of the sample syringes will be aspirated into the waste.
20. With values in the FLUSH position, dip the wash inlet (tubing with a T joint protruding from the bottom of the rapid-quench apparatus) into a 15-mL tube of deionized water for 5 sec, and then into a 15-mL tube of 100% ethanol for 5 sec. Allow the vacuum to pull air through the system for 30 sec to dry. The vacuum will draw the solutions through the mixing and exit loops.
21. While holding the exit line, carefully detach the vacuum. Hang the end of the vacuum line (with Luer adapter) in an empty, disposable 15-mL tube to avoid contaminating the work area. Screw the waste tube onto the exit line. CAUTION: The exit line and the end of the vacuum line should be handled with care to keep them free of ribonuclease, and to avoid transferring radioactive material to gloves and work surfaces.
Table 11.6.2
Beam current, 2.8 GeV (mA)
Speed (rpm)
Exposure (msec)
300-200
240 160 120 96 30 60 48 40
10 15 20 25 20 40 50 60
200-150 Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays
Setting Exposure Times at X28C
150-100 —
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In a typical experiment (20 reactions), an unexposed control and a reaction without Mg2+ are acquired first (reactions 1 and 2). The left side drive syringe and the bottom left sample loop are then refilled with CE20, and 16 folding time points (reactions 3 to 18) are acquired by varying the folding delay time. Finally, two controls with prefolded RNA are performed (reactions 19 and 20).
22. Dispense 20 µL precipitation cocktail into each of 20 screw-cap microcentrifuge tubes. Label tubes 1 through 20 (or as desired). Recap tubes and hold at room temperature until needed. These tubes will be used to collect samples. It is important to avoid introducing nucleases or other contaminants.
Perform a “no exposure” control 23. Screw tube no. 1 onto the exit line adapter. Set the valves to LOAD sample, and inject 10 µL RNA and 10 µL CE buffer into the right and left sample loops as in steps 13 and 14. 24. Set the folding delay time to 0.01 sec, and repeat steps 15 and 16. 25. Expel any sample remaining in the exit line into the collection tube as in step 17. 26. Remove the tube from the exit line, and flush lines with deionized water and 100% ethanol as described in steps 18 through 21. 27. Add 600 µL of 100 % ethanol to the expelled sample, recap, and store at 4° to −20°C in a lead-lined box. It is important that the screw caps form a leak-proof seal to prevent loss of sample during shipment.
Perform a control with unfolded RNA 28. Attach tube no. 2 to the exit line. Reload the sample loops as in steps 13 and 14, except this time the interlock system should be enabled as the user exits the hutch. 29. Enter a delay of 0.01 sec in the controller software. 30. Turn on the X-ray beam when the interlock safety alarm has turned off. 31. Once the beam is on, fire the rapid-quench apparatus (type “G”). Turn the beam off as soon as the software displays a message indicating that the experiment has finished. The time that the beam is on should be kept to a minimum to limit X-ray damage to samples and equipment.
32. Collect the sample and flush the mixing apparatus as in steps 17 to 21. Precipitate the RNA as in step 27. Exchange buffer in drive syringe A 33. Close the top valves to drive syringes A, B, and C (LOAD syringes). Close the bottom left valve (180° from LOAD sample) and open bottom right valve (as in LOAD sample). Carefully remove the syringe containing RNA, and set aside for future use. To probe Mg2+-induced folding, the CE buffer in the left side of the mixing apparatus (drive syringe A) must be exchanged for CE20 buffer.
34. Attach the vacuum to the exit port and open the bottom left valve (LOAD sample). To empty drive syringes, turn the bottom and lower valves to FIRE. Slowly depress the plunger of syringe A and B until the syringe is empty. Open the lines connecting drive syringes to air (opposite of FIRE).
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35. Disconnect the vacuum, and return all valves to LOAD syringes. 36. Using the control software, move the drive platform up as in steps 1 and 2. 37. Fill drive syringe A with CE20 buffer as described in steps 3 and 4. Fill drive syringe B with additional CE buffer. 38. Load the bottom left sample loop with CE20 buffer (using a fresh 1-mL syringe) as in steps 9 through 11. Replace the syringe containing RNA on the right sample port. 39. Perform a priming shot and flush the mixing and exit lines, as described in steps 13 to 21. Expose samples with variable folding delays 40. Using collection tubes no. 3 to 18, acquire time-resolved data by repeating steps 28 to 32 for each sample. Vary the length of the folding delay by entering the appropriate value into the software before firing each shot. If the folding delay is longer than the time required for the interlock safety shutter to open (3 sec), begin the folding routine, then turn on the X-ray beam when 3 sec of the delay remains. A standard laboratory timer can be used to measure folding delays of ≥10 sec.
41. When the first aliquot of RNA is expended (typically after the ninth sample), prepare the second aliquot as in steps 5 to 7. Remove the right sample syringe, and replace it with a fresh 1-mL syringe of RNA (steps 8 to 9). Fire a priming shot, then repeat step 40 for the remaining reactions, no.10 through 18. 42. When the experimental reactions are complete, open the right sample syringe to the vacuum (it should point to the right), and aspirate any remaining RNA. Flush the lines as in steps 17 to 21. Remove the vacuum from the exit line, and the 1-mL syringe from the right sample port. Alternatively, the sample syringe can be removed and remaining RNA recovered before the system is flushed with water and 100% ethanol.
Acquire controls with prefolded RNA 43. Prepare a 0.6-µCi (50-µL) aliquot of RNA by annealing in MgCl2 under conditions in which the RNA is known to fold completely. For the Tetrahymena ribozyme, this is done by placing 5.5 mL of 100 mM MgCl2 on the inside lid of the microcentrifuge tube, incubating 1 min at 95°C, then immediately microcentrifuging 1 min.
44. Load the annealed RNA into the right sample loop, and fire a priming shot (steps 8 to 9, and steps 13 to 21). 45. Using collection tubes no. 19 and 20, perform two shots with a 0.01-sec folding delay, as in steps 28 to 32. 46. Store samples in a lead container at −20°C or on dry ice. Samples should be stored at −20° or −70°C until they are ready to be analyzed by polyacrylamide gel electrophoresis (see Basic Protocol 3). They should be analyzed as soon as possible after exposure, ideally within 1 week. Samples may be shipped to the investigator’s home laboratory for analysis. Users should consult with the synchrotron facility staff regarding procedures for packaging and shipping radioactive materials. Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays
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47. When the experiment is complete, thoroughly flush and disassemble the rapid-quench apparatus (see Support Protocol 3). At the end of the day, complete all safety checks, and disable the beamline. SAMPLE WORK-UP AND DATA ANALYSIS Radiolabeled products of X-ray-induced hydroxyl radical cleavage are analyzed directly by electrophoresis on a polyacrylamide sequencing gel. The intensity of the bands is quantified using a storage phosphorescence imager (Molecular Dynamics or equivalent). Equilibrium constants or kinetic rate constants are obtained by fitting changes in the relative extent of cleavage (or protection) to appropriate models.
BASIC PROTOCOL 3
Materials Irradiated 32P end-labeled RNA (see Basic Protocols 1 and 2) Unirradiated 32P end-labeled RNA (see Support Protocol 1) 2× formamide loading buffer (APPENDIX 2A) or loading buffer containing urea RNase T1 cocktail (see recipe) 0.5 U/mL RNase T1 (USB) CE buffer, pH 7.5 (see recipe) or TE buffer (APPENDIX 2A; see Critical Parameters) Prefolded control RNA (see Basic Protocol 2) Imaging system with large exposure cassettes (Phosphorimager, Molecular Dynamics; or equivalent) Densitometer Image analysis software for personal computer (PC) or Macintosh computer (ImageQuant, Molecular Dynamics; NIH Image; or equivalent) Spreadsheet software (Microsoft Excel or equivalent) Graphical fitting software (KaleidaGraph, SigmaPlot, or equivalent) Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (APPENDIX 3B) NOTE: Samples should be stored at −20° or −70°C until they are ready to be analyzed. This should be done as soon as possible after X-ray exposure, ideally within 1 week. Prepare RNA for gel electrophoresis 1. Microcentrifuge irradiated and unirradiated 32P end-labeled RNA samples 30 min at maximum speed, 4°C. Samples should be kept on ice or at 4°C at all times.
2. Carefully remove the supernatant using a pipet or aspirator, avoiding contact with the RNA pellet. Dispose of the supernatant as radioactive waste. If using a standard pipet, the solution in the pipet tip can be checked with a survey meter to make sure that the RNA pellet is not lost. If using an aspirator, it should be connected to a trap that can contain radioactive liquid.
3. Recap the tubes and microcentrifuge for an additional 5 sec at 4°C. Use a pipet tip to remove any remaining supernatant. 4. Dry the samples in a Speedvac evaporator at room temperature. 5. Redissolve each sample in 6 µL of 1× loading buffer with urea or formamide. Vortex briefly, and microcentrifuge 15 sec at maximum speed, room temperature. Hold samples at room temperature or 4°C.
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Run sequencing gel 6. Cast a 33 × 42–cm sequencing gel using an appropriate concentration of polyacrylamide (APPENDIX 3B). Use a comb that forms separate, flat-bottomed wells ∼0.8-cm wide. The authors have obtained good results with 6% to 10% (19:1 mono/bis) polyacrylamide gels.
7. Mount the polymerized gel on an electrophoresis apparatus. Preheat gel for 10 to 20 min at 55 W. 8. Prepare a sequencing ladder by digestion with RNase T1. Combine the following: 7 µL RNase T1 cocktail 3 µL 32P 5′-end-labeled RNA (∼100,000 cpm) 1 µL 0.5 U/mL RNase T1. Incubate 12 min at 50°C. Place on ice. The concentration of RNase T1 should be adjusted to produce an even ladder of fragments. Samples should be loaded as soon as possible (within 10 min) to prevent overdigestion. Ribonuclease reactions can be held up to 60 min on dry ice.
9. If desired, prepare an undigested control by combining 3 µL of 5′-end-labeled RNA with 2 µL of 1× CE or TE buffer and 5 µL of 2× loading buffer. 10. Immediately before loading, heat samples 1 min at 85° to 90°C and place on ice. 11. Load samples, with the RNase T1 digest next to lanes containing prefolded controls (see Basic Protocol 2, tube no. 19 and 20). Run the gel at 55 W for the desired length of time. Several electrophoresis runs may be necessary to resolve all of the cleavage products if the RNA is >150 nt.
12. Disassemble the sequencing apparatus, transfer the gel to Whatman 3MM filter paper, and dry under vacuum. Place the gel in a large Phosphorimager cassette. Depending on the radioactivity of the samples, exposure may require 1 to 5 days.
Data analysis 13. Scan the gel on a densitometer and display the results using image analysis software. 14. Identify residues that are protected from cleavage in the prefolded RNA control lanes by comparison with the pattern of RNase T1 digestion. A similar pattern should be visible in experimental lanes (e.g., with increasing MgCl2 concentration or longer folding times).
15. Define a rectangular area within the lane that includes a group of protected residues that behave similarly to each other over the course of the experiment. Use the copy feature of the image analysis software to place identical boxes in each lane of the gel. Great care must be taken to define the contours of the protected region accurately, as inclusion of adjacent bands in the area to be quantified will considerably reduce the quality of the analysis. A tilted rectangle or rhomboid can be used to account for “smiling” effects at the edge of the sequencing gel.
16. Repeat step 15 for all residues that are protected and that are resolved by the gel. Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays
17. In the same manner, box one or more bands of constant intensity.
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Figure 11.6.4 Organization of data from image analysis program using Excel.
These bands, which usually correspond to nucleotides that are not protected from cleavage, will serve as a reference.
18. Integrate the volume of all of the rectangles (objects). Export the results of the integration to a spreadsheet program. The average background intensity may be subtracted from each pixel if desired (see Commentary).
19. Use the drag-and-drop editing features of the spreadsheet program to arrange the volume (cpm) of each band or collection of bands into a table, as illustrated in Figure 11.6.4. It is usually easiest to arrange the data so that each column represents a set of residues (across the lanes of the gel), and each row corresponds to an experimental variable, such as folding time or MgCl2 concentration (represented by one lane). Enter the time or concentration in the first column of the spreadsheet. Label the columns with the sequence of the protected residues or reference bands.
20. Calculate the “protection ratio” for each column of data, according to P=1−
cpm protected cpm reference
where cpm protected is the volume integral of the protected region of interest and cpm reference is the volume integral of an appropriate reference band. 21. Export the data as tab-delimited text to a scientific graphing program, such as KaleidaGraph or SigmaPlot. Plot the protection ratio versus time (or concentration), and fit the data to the desired rate expression or binding isotherm (see Commentary for further discussion). It is useful to normalize the extent of protection by defining the fractional Y as Y=
P − Pmin Pmax − Pmin
where Pmin and Pmax are the values obtained for unfolded and fully folded RNA. These should correspond to the upper and lower baselines, respectively, of the transition.
RNA Folding Pathways
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SUPPORT PROTOCOL 1
PREPARATION OF RADIOLABELED RNA Unlabeled RNA is prepared by in vitro transcription of plasmid DNA with T7 RNA polymerase (UNIT 11.4), according to standard methods (Milligan et al., 1987; Milligan and Uhlenbeck, 1989). The RNA must be treated with phosphatase before labeling at the 5′ end with polynucleotide kinase. After gel purification, the labeled RNA is dispensed into aliquots (usually 10 µCi, 20 µL each) to guard against total loss of sample at the beamline. Samples are shipped in leak-proof screw-cap tubes. Materials 10 pmol/µL RNA, treated with calf intestinal phosphatase (UNIT 6.3) T4 polynucleotide kinase and 10× kinase buffer 6000 Ci/mmol (10 µCi/µL) γ-32P]ATP TE buffer, pH 7.5 (optional; APPENDIX 2A) CE buffer, pH 7.5 (see recipe) Microcentrifuge tubes with captive screw caps and O-ring seals (Rainin) Additional reagents and equipment for phosphorylation reaction (UNIT 6.3), for preparing and running a preparative polyacrylamide sequencing gel (APPENDIX 3B), for phenol/chloroform extraction and ethanol precipitation (APPENDIX 2A and, e.g., CPMB UNIT 2.1A) NOTE: Radioactive materials must be labeled and shipped in compliance with federal and state regulations. Consult with the radiation safety officer of your home institution and the receiving institution before planning to ship radioactive materials. Prepare 32P end-labeled RNA 1. Phosphorylate 40 pmol phosphatase-treated RNA using T4 polynucleotide kinase and 50 µCi [γ-32P]ATP (UNIT 6.3). Unlabeled or 3′-end-labeled RNA may also be used.
2. Purify the labeled RNA by denaturing polyacrylamide gel electrophoresis (APPENDIX 3B) on a 20 × 20–cm gel. 3. Elute RNA, extract with phenol/chloroform, and precipitate with ethanol (APPENDIX 2A and, e.g., CPMB UNIT 2.1A). Dissolve the final pellet in TE or CE buffer and store up to 3 months at −20°C. 4. Measure the activity of the purified RNA by counting 1 or 2 µL of the final solution in a liquid scintillation counter (1 µCi = 2.2 × 106 cpm). Typically, 1 mL contains 106 cpm. The solution should minimally contain 250,000 cpm/mL or 0.1 mCi/mL.
Dispense aliquots of RNA for shipping 5. Determine the total amount of RNA that will be required for the footprinting experiments. Each equilibrium or time-dependent footprinting experiment requires 4 mCi, or 200,000 to 400,000 cpm, per sequencing gel lane. Extra solution may be left in the home laboratory to serve as a control for background hydrolysis of the RNA during shipping and handling at the beamline. Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays
6. Dilute the required amount of RNA in 1× CE buffer at a final concentration of 3 µCi/20 µL, or 6 µCi/20 µL for equilibrium experiments. Mix thoroughly, and pipet 20-µL aliquots into leak-proof screw-cap microcentrifuge tubes.
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A range of 2 to 4 mCi per aliquot, or 4 to 8 mCi per aliquot for equilibrium experiments, may be used. Higher concentrations of 32P provide better signal-to-noise ratios.
7. Prepare additional aliquots for prefolded control reactions by diluting 0.6 µCi of labeled RNA (from stock solution) into a final volume of 50 µL CE buffer. Place the solution in a leak-proof screw-cap microcentrifuge tube. It may be necessary to make serial dilutions of the stock solution in order to accurately transfer 0.6 mCi. This is sufficient for two control reactions.
8. Prepare additional 0.6-µCi aliquots as needed (at least one aliquot per time-resolved experiment). Package RNA samples for shipment to the beamline 9. Store samples at −20° or −70°C until ready for shipment to the beamline. 10. Package samples for shipment on dry ice, according to the specific instructions provided by beamline personnel. In general, a Radiation Shipping Notice must be included inside the package. The outside should be labeled with (1) a “limited quantity notification” (49 CFR 173.421), if applicable, (2) a dry ice hazard diamond, and (3) the overnight shipper’s label, with shipping and return addresses. One may be required to obtain an authorization number from the recipient. Detailed instructions for shipping radioactive materials to beamline X28C at the NSLS should be obtained from the beamline supervisor. As these protocols are subject to change, users should verify that they are using the correct protocol before each shipment.
DETERMINE OPTIMAL EXPOSURE TIME The optimal time that samples should be exposed to the X-ray beam is determined by measuring the fraction of RNA that is cleaved after variable exposure times. The resulting dose-response curve is used to determine how much irradiation is required to cleave 10% to 30% of the RNA strands. This should be done in the buffer used for RNA folding experiments to control for the effect of solutes.
SUPPORT PROTOCOL 2
Additional Materials (also see Basic Protocols 1 and 3) 1 to 2 µCi 5′-32P-labeled RNA in 10 µL CE buffer Prepare and expose samples 1. Dilute RNA with 40 µL CE buffer (final 50 µL). Mix thoroughly and briefly microcentrifuge. CE20 buffer can also be used, if desired.
2. Divide the sample into five 10-µL aliquots. Place aliquots in screw-cap microcentrifuge tubes with O-ring seals. 3. Set up the sample holder and shutter as described (see Basic Protocol 1, steps 1 to 7). 4. Expose four aliquots to the X-ray beam for varying times (see Basic Protocol 1, steps 14 and 15). Reserve one aliquot as an unexposed control. At X28C, exposure times typically range from 10 to 100 msec.
5. Store irradiated and control aliquots on dry ice if they will not be analyzed right away (store up to 3 days). RNA Folding Pathways
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Determine extent of RNA cleavage 6. Cast and preheat a small (20-cm) 5% to 8% polyacrylamide sequencing gel. 7. Thaw aliquots of RNA (if frozen) and add 8 µL of 2× formamide loading buffer to each. Heat samples briefly (30 to 60 sec) at 85°C. 8. Load on the gel and run the gel so that the full-length RNA migrates at least one third of the way down. The object is to separate the full-length (uncleaved) RNA from the products.
9. Disassemble the gel and dry under vacuum. Expose the gel to a Phosphorimager screen long enough so that the full-length RNA appears as a dark band, but does not saturate the Phosphorimager screen. 10. Use the volume integration feature of the image analysis software to quantify the amount of uncut RNA remaining after cleavage. Prepare a plot of the fraction of the natural logarithm of uncut RNA versus exposure time. This plot should yield a straight line. The dose depends on the beam current (or flux) as well as exposure time. The exposure time must be adjusted to account for variations in the beam current during each 24-hr period. SUPPORT PROTOCOL 3
SET UP RAPID-QUENCH MIXING APPARATUS A modified Kin-Tek RQF-3 rapid-quench apparatus is used for X-ray footprinting at X28C (see Commentary). The mixing apparatus must be installed at the beginning of the experiment. Users should consult the manufacturer’s literature for more detailed information on the maintenance and use of the rapid-quench device. This protocol should be adapted to suit equipment available at individual facilities. Additional Material (also see Basic Protocol 2) Detergent (e.g., Absolve, NEN) Plastic-backed absorbent bench paper Diaphragm vacuum pump (details) Vacuum/vent filter, 0.2 µm (Millipore Millex 50 mm, or equivalent) Side-arm flask with one-hole stopper and Teflon tube Thick-walled soft tubing (e.g., Tygon, Nalgene) to fit Teflon tube Adapter (male M6 to male Luer) to connect soft vacuum tubing with 1⁄16-in.-o.d. polypropylene tubing (exit line of rapid-quench) 5-mL syringes Install rapid-quench apparatus 1. Mount the Kin-Tek rapid-quench apparatus (in polycarbonate water jacket) on the stepper motor frame (Figure 11.6.2). 2. Use the provided bolts to attach a horizontal aluminum plate on the Unistrut frame, near the bottom of the mixing apparatus. Mount a second vertical aluminum plate on the rear of the rapid-quench box so that it rests against the first plate. The horizontal plate will support sample tube racks. The vertical plate shields the sample exit lines from the X-ray beam.
Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays
3. Cover all exposed surfaces of the work area with plastic-backed absorbent bench paper to prevent contamination of hard surfaces in case a radioactive sample is spilled.
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4. Outside the hutch, turn on the controller and begin running the control software. If using a personal computer to interface with the controller, open the Kin-Tek software (or equivalent) in DOS and download the desired BASIC control program. Calibration settings and mixing parameters for individual mixers can be stored between experiments and imported into the Kin-Tek control software. The control code modified for X-ray footprinting at X28C is available from the Center for Synchrotron Biosciences.
Connect water bath and vacuum lines 5. Optional: Remove nucleases by soaking sample ports and internal tubing for 1 hr or overnight in detergent. Before use, remove all traces of detergent by thorough rinsing with RNase-free water followed by 100% ethanol. 6. Inside the hutch, connect the recirculating water bath to the jacket surrounding the rapid-quench apparatus. Set the temperature as desired. 7. Connect the vacuum pump to the side arm of a 0.5-L side-arm flask, through a 0.2-µm filter. CAUTION: This flask will collect the waste between kinetic shots, and therefore will be contaminated with radioactive material and should be labeled accordingly. Be extremely careful when handling liquid waste containers, and be sure that secondary trap and vacuum lines are secured. Always wear gloves, and monitor them frequently using a survey meter.
8. Attach thick-walled soft tubing to a Teflon tube in the top of the side-arm flask. Insert a male M6 to male Luer adapter into the opposite end of the tubing. The vacuum tubing should be stored in a clean, covered box to minimize risk of RNase contamination. The adapter should fit snugly over the 1⁄16-in.-o.d. tubing that is used as the exit line of the rapid-quench apparatus.
9. Turn on the vacuum pump. Attach the vacuum to the exit line, and aspirate any solution from the sample loops and drive syringes. Align mixing apparatus with X-ray beam 10. Secure the automatic alignment device to the registration pins on the back of the mixing box, and attach the output cable. 11. Enable the X-ray beam and follow the instructions for the automatic alignment procedure described on the Center for Biosciences website (http://www.aecom. yu.edu/home/csb/). If the beam is not aligned correctly with the exposure port, the extent of RNA cleavage may be drastically reduced. Although not necessary, it is prudent to check the alignment of the mixer at the end of an experimental session. This test will help troubleshoot the experiment in the event of poor results.
12. Remove the alignment device from the rapid-quench apparatus and extend the flight tube to the back of the rapid quench. Failure to do so will prevent the samples from being exposed to X-rays during the experiment.
13. Optional: Manually check the alignment by taping Kodak Linagraph paper over the exposure cell port on the rear of the rapid-quench apparatus. Expose the paper to the X-ray beam for 1 sec, and verify that the burn mark on the paper is centered in a depression caused by the outline of the exposure port. The apparatus is now ready for use as described in Basic Protocol 2. RNA Folding Pathways
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Clean up 14. At the end of the experiment, discard the screw cap on the exit line as radioactive waste. 15. Attach the vacuum to the exit line, and aspirate any remaining solutions from the sample loops and drive syringes. Leave the vacuum on. 16. With a fresh 5-mL syringe of deionized water, rinse out all the syringes and sample loops. Use the vacuum to aspirate each line after rinse. 17. Repeat step 15 with a 5-mL syringe of 100% ethanol. 18. Remove the vacuum line and turn off the pump and water bath. Remove the mixing apparatus from the support stand and store in a covered plastic box. 19. Exit the control software and shut off the controller. SUPPORT PROTOCOL 4
CALIBRATE RAPID-QUENCH MIXING APPARATUS Each mixing apparatus should be calibrated when first installed, or when tubing is changed. The parameters for the drive platform distances can be stored in the controller software. The performance of the rapid-quench mixing apparatus is evaluated visually using a water-soluble dye such as bromphenol blue. Additional Materials (also see Basic Protocol 2 and Support Protocol 3) 0.25% (w/v) bromphenol blue in water Dental mirror Small flashlight 1. Set up a rapid mixing apparatus (see Support Protocol 3). Load drive syringes with water or CE buffer, as described (see Basic Protocol 2, steps 1 to 4). 2. Load 0.25% bromphenol blue solution into the right sample loop (instead of RNA), as described in Basic Protocol 2, steps 8 to 11. Fill the left sample loop with water or CE buffer. 3. Set up a mock footprinting experiment with a long folding delay (>10 sec) and a very long exposure time (10 sec). Install a microcentrifuge tube at the exit tube to collect the ejected sample. 4. Exit the hutch without enabling the beam. Type “G” to go. 5. Quickly re-enter the hutch. Inspect the flow of colored solution in the tubing using a dental mirror and flashlight. If the dye and water are not mixing correctly, empirically adjust the distance values in the Kin-Tek control program. After the first push, the dye should advance up to the exposure cell, but should not enter it. The dye should mix uniformly with water. Once the “exposure” push starts, the dye should smoothly pass through the exposure cell and into the microcentrifuge tube.
REAGENTS AND SOLUTIONS Use RNase-free deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays
CE (sodium cacodylate/EDTA) buffer, 1× (pH 7.5) 1.0 mL 1 M sodium cacodylate 20 µL 0.5 M EDTA, pH 8.3 (APPENDIX 2A) Deionized water to a final volume of 100 mL
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Filter sterilize through a 0.2-µm filter Store up to 6 months at room temperature CE20 buffer, 1× Prepare as for 1× CE buffer (see recipe), but include 2 mL of 1 M MgCl2 (APPENDIX 2A; final 20 mM). Precipitation cocktail, 10× 8 mL 4 M NaCl (APPENDIX 2A) 0.5 mL 0.5 M EDTA, pH 8.3 (APPENDIX 2A) 100 µL 10 mg/mL carrier tRNA 150 µL 20 mg/mL glycogen Bring to 10 mL with deionized water Store in 1-mL aliquots up to 1 year at −20°C RNase T1 cocktail 100 µL 10 M urea (see recipe) 3 µL 1 M sodium citrate, pH 3.5 1.5 µL 0.1 M EDTA, pH 8.3 (APPENDIX 2A) 2 µL 2% (w/v) bromphenol blue and xylene cyanol 3 µL 10 mg/mL carrier tRNA Store in 0.2-mL aliquots up to 1 year at −20°C Urea, 10 M Mix 0.22 g urea in 200 µL water. Warm at 50°C and vortex to dissolve. Store up to 1 year at –20°C. COMMENTARY Background Information X-ray-dependent hydroxyl radical footprinting Hydroxyl radicals have been widely used to probe the conformation of nucleic acids and nucleic acid–protein complexes (Tullius and Dombroski, 1985; Latham and Cech, 1989; Dixon et al., 1991; Strahs and Brenowitz, 1994). Reaction of hydroxyl radical with ribose results in oxidation of the sugar and elimination of phosphate groups, leading to strand cleavage. Cleavage is relatively insensitive to base sequence and secondary structure of the RNA (Celander and Cech, 1990). However, the susceptibility of individual nucleotides to cleavage in the presence of a hydroxyl radical correlates well with the solvent accessibility of the ribose C4′ (Latham and Cech, 1989; Cate et al., 1996; Balasubramanian et al., 1998). Nucleotides that are inaccessible to bulk solvent due to RNA tertiary structure or interactions with a protein are protected from cleavage. The resulting footprint provides information about the conformation of the nucleic acid, and can be quantified
to determine the fraction of structured molecules in the population. Chemical methods for generating hydroxyl radicals, such as the Fenton-Haber reaction (Dixon et al., 1991) or disproportionation of peroxynitrous acid (King et al., 1993), typically require several seconds or longer to cleave 20% to 30% of the RNA (Chaulk and MacMillan, 2000; Hampel and Burke, 2001). This unit describes the use of a synchrotron X-ray beam to generate hydroxyl radicals in aqueous solution. The advantage of this method is that cleavage reactions can typically be completed in 10 to 50 msec. Therefore, “X-ray footprinting” can be used to probe kinetic folding intermediates or other transient conformations. The protocols given here were initially developed to probe the folding pathway of the Tetrahymena ribozyme (Sclavi et al., 1997, 1998a), but can be applied to a variety of catalytic RNAs or RNA-protein complexes. Recent improvements to the method are described in Ralston et al. (2000b) and Dhavan et al. (2001). To carry out cleavage reactions on nucleic acids in the millisecond timescales, the beam must deliver a sufficient flux of photons to the sample so that the steady-state concentration of
RNA Folding Pathways
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hydroxyl radicals is 0.5 to 1.2 µM (Ralston et al., 2000b). Hydroxyl radicals are produced after excitation of water molecules by absorbed photons, according to the following equation (Klassen, 1987). hν
H2O
H2O → H2O+ + e−dry → H3O+ + • OH + e−aq
The steady-state concentration of hydroxyl radicals and the dose required to cleave the sample depend on many factors (Ralston et al., 2000b), including the energy absorbed by the sample. A bending magnet beamline at the NSLS operating at 2.8 GeV delivers 5 × 1014 photons/sec to a 10-µL sample, with an absorption maximum near 7.5 keV (Ralston et al., 2000b). Absorption of 10 keV produces 287 hydroxyl radicals (Klassen, 1987). The steadystate concentration of hydroxyl radical can be estimated from the photon flux, and is ∼10−6 M for NSLS footprinting beamlines (Sclavi et al., 1998b). The effective concentration of hydroxyl radicals will be reduced by free radical scavengers in the sample (see Critical Parameters).
Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays
Instrumentation The design of equipment for X-ray footprinting experiments has been described previously (Sclavi et al., 1998b; Ralston et al., 2000b; Dhavan et al., 2001). Equilibrium experiments are carried out using an aluminum sample holder and stand, which places a standard microcentrifuge tube horizontally in the path of the beam (Figure 11.6.1). The sample is vertically aligned with the beam using a photodiode detector placed behind the sample block and a motorized stage. The alignment apparatus operates with a resolution of 50 µm. This level of precision is necessary to obtain reproducible extents of RNA cleavage at X28C. The exposure time is regulated by an electronic shutter placed between the sample block and the end of the beampipe. The electronic shutter is controlled by a microprocessor placed outside of the experimental hutch. The path of the beam between the end of the beampipe and the shutter is covered by a retractable flight tube. This reduces the amount of scattered radiation in the experimental hutch. The aluminum sample block is connected to a water bath to maintain constant temperature. This is sufficient to dissipate the small amounts of heat generated over short (<100 msec) irradiation times. The beam causes an ∼0.5°C/sec rise in the temperature of the sample (Sclavi et al., 1998b). A modified Kin-Tek stopped-flow rapidquench apparatus has been installed by the Center
for Synchrotron Biosciences staff at beamline X28C for rapid mixing experiments (Figure 11.6.2). The mixing valve present in a standard rapid-quench apparatus is replaced with a flow cell mounted on a steel plate at the back of the box. This enables the flow cell to be placed close to the end of the beampipe (Dhavan et al., 2001). The flow cell itself is constructed of Vespel to minimize damage from X-rays. Sample and buffer is driven through the lines by syringes at the top of the apparatus. After samples (10 to 20 µL) flow through a mixing T, they are aged in a 60-µL loop before being pushed through the flow cell. If desired, a quench solution can be delivered via a third syringe. However, a chemical quench is not normally necessary, because the hydroxyl radical concentration decreases rapidly (<1 msec) as soon as the beam is turned off (Sclavi et al., 1998b). The Kin-Tek mixers at X28C are configured for either 10 µL or 20 µL sample volumes. Larger volumes yield improved precision, but increase the mixing dead time. If the volume of the sample is larger than the capacity of the X-ray exposure cell, the exposure times will increase. The Kin-Tek RQF-3 uses a stepper motor and platform to advance the pistons of the drive syringes. The distance traveled by the sample is determined by the number of turns of the stepper motor. The time required to advance the sample over this distance depends on the motor speed. Typical parameters for RNA footprinting at X28C are given in Table 11.6.1. The exposure time is regulated by controlling the speed at which the sample flows past the exposure port. The exposure time is given by txray (msec) =
(0.04 rev) 60 sec × 1000 × 1 min (s rpm)
Hence, a 10-msec exposure would require a motor speed, s, of 0.024 rpm.
Critical Parameters X-ray-dependent hydroxyl radical footprinting has been used successfully to probe Mg2+-dependent folding of ribozymes, urea-induced denaturation of RNA, and the formation of protein-DNA complexes (Sclavi et al., 1998a; Deras et al., 2000; Ralston et al., 2000a; Dhavan et al., 2001). Although synchrotron X-ray footprinting provides sequence-specific information about the tertiary conformation of RNAs that cannot be presently obtained by other methods, several variables will determine the likelihood of success. The critical parame-
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ters discussed below include conformational stability of the RNA, optimum exposure times, factors influencing signal-to-noise ratios of product bands, and sample handling. Conformational stability X-ray-dependent footprinting will be successful only if the hydroxyl radical protection pattern produced by the folded RNA or RNAprotein complex is well defined. It is important that the RNA tertiary structure or protein complex of interest is stable under the final conditions of the experiment. If the RNA tertiary structure is unstable, or if the RNA folds into multiple conformations, the footprint will be faint and difficult to interpret. Nonspecific RNA-protein complexes obscure native interactions and make interpretation of the data difficult. It is important to optimize the conditions using Fe(II)-EDTA-dependent footprinting before undertaking experiments at the synchrotron beamline. Parameters that should be evaluated include buffer, pH, Mg2+ concentration, temperature, protein concentration, and protocols for annealing the RNA. Hydroxyl radicals react with nucleic acid bases as well as the ribose moiety (Dizdaroglu and Bergtold, 1986; Breen and Murphy, 1995). In general, these modifications do not lead to strand scission and are not detected by the protocols described here. The most common base modifications, 8-oxo-G and 5-hydroxy-C, are not expected to drastically destabilize RNA secondary structure (Wallace, 1998). Nonetheless, it is important to be aware that specific base modifications could induce unfolding of the RNA, and this could alter the observed cleavage pattern. Similarly, rapid oxidation of amino acid side chains may destabilize RNAprotein complexes and cause them to dissociate during the X-ray exposure. Exposure time To interpret footprinting experiments quantitatively, it is important that the extent of cleavage be adjusted so that each molecule is cleaved no more than once, on average. This is usually achieved by limiting the extent of cleavage to 10% to 30% of the starting material (Brenowitz et al., 1986). The exposure time required to cleave 10% to 30% of the RNA depends on the energy and beam current at the time of the experiment (Sclavi et al., 1998b). It also depends on the length of the RNA (longer RNAs require shorter exposure times). Ideally, the exposure time should be determined experimentally for each substrate by acquiring a dose-
response curve (see Support Protocol 2). The deadtime of the experiment will be determined largely by the exposure time required to generate the cleavage pattern. The extent of cleavage will also be affected by the presence of free radical scavengers in the sample, such as Tris and glycerol. Inorganic buffers such as phosphate or cacodylate give excellent results and should be used when possible. Concentrations <30 mM of Tris can be tolerated, as long as the exposure time is lengthened to compensate for the reduced rate of cleavage. Carboxylic acids (such as EDTA) and urea do not interfere with hydroxyl radical cleavage (Ralston et al., 2000a). Signal-to-noise ratio The ability to quantify the extent of protection from a sequencing gel will depend in large part on the intensity of the bands relative to the background activity in the lane. Several factors influence the signal-to-noise ratio. First, it is essential that a sufficient amount of 32P be used, so that individual bands can be detected using a phosphorescent screen or other imaging device. The authors found that, for the Tetrahymena ribozyme (388 nt), quantitation of the protection pattern improved as the amount of sample increased from 200,000 to 400,000 cpm per lane. By contrast, experiments using <150,000 cpm per lane failed to yield interpretable results. Since the cpm per band depends on the number of products, less radiolabeled material is required to analyze shorter RNAs. It is important that the cross-section of the beam is large enough to cover the entire sample or flow cell. Otherwise, only part of the sample will be irradiated. Similarly, air bubbles can exclude sample from the beam, and the dissolved oxygen will alter the distribution of cleavage products. The resolution of sequencing gels is improved by using 19:1 mono/bis acrylamide, and by precipitating samples with ethanol to remove excess salt. Background RNA cleavage Another critical parameter for enhancing the signal-to-noise ratio is the extent of background cleavage. Samples should be protected from scattered X-radiation when the beam is on. Recent improvements at X28C (Dhavan et al., 2001) have reduced scattered radiation. The flight tube was extended so that it reaches the face of the stopped-flow apparatus, and a lead and Plexiglas housing was constructed to cover the sample-loading syringe.
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Figure 11.6.5 Results of X-ray footprinting experiments showing Mg2+-dependent folding of the P4-P6 domain of the Tetrahymena ribozyme. Reprinted from Deras et al. (2000) with permission from the American Chemical Society.
Trace contamination by ribonucleases presents another source of background cleavage, and should be minimized to the extent possible. Even 2% cleavage by nucleases can lower the quality of the data. Besides incautious preparation of buffers and samples, common trouble spots are mishandling of the exit tube and sample-loading syringe on the Kin-Tek stopped-flow apparatus, prolonged exposure of tips and samples to airborne contaminants, and contaminated O-rings and lids in screw-cap microcentrifuge tubes. Additional control reactions can be added to the protocol to troubleshoot sources of background cleavage. If the facility is to be used for several different applications, such as protein and nucleic acid footprinting, it is helpful to maintain separate mixing apparatuses for each type of experiment. This reduces the chance of RNase contamination in RNA folding experiments. Nucleases can be removed by soaking the sample ports and internal tubing for 1 hr or overnight in a detergent such as Absolve (NEN). The detergent must be removed by thorough rinsing with RNase-free water and ethanol.
Anticipated Results
Time-Resolved Hydroxyl Radical Footprinting of RNA with X-Rays
Quantitative analysis of footprinting experiments has been discussed in detail elsewhere (Brenowitz et al., 1986). Nucleotides that become protected from hydroxyl radical cleavage because of tertiary structure or protein binding should be visible as clear regions within a ladder of bands on the gel. Comparison with a sequence ladder should permit assignment of the protected regions to particular sequences within the RNA. Interpretation of the data is improved considerably if a three-dimensional structure or model is available.
The relative extent of protection is determined by comparing the intensity of cleavage products in unfolded and fully folded controls. A detailed presentation of the integration method described in Basic Protocol 3 can be found in CPMB UNIT 12.4. The equilibrium folding of RNA as a function of Mg2+ concentration can be described by the Hill equation (Celander and Cech, 1991; Sclavi et al., 1997). The fractional saturation of a protected region, Y , is fit to Y=
(Kapp / [Mg2+])nH P − Pmin = Pmax − Pmin 1 + (Kapp / [Mg2+])nH
where Kapp is an apparent dissociation constant corresponding to the midpoint of the transition, nH is the Hill coefficient, and Pmin and Pmax are the upper and lower baselines of the transition (see Basic Protocol 3, steps 19 and 20). An example of this type of data is given in Figure 11.6.5. Time-dependent experiments should be fit to an appropriate rate expression, as illustrated in Figure 11.6.5. For single first-order transitions, this yields Y=
P − Pmin = 1 − e−kobsT Pmax − Pmin
The first equality holds only if the upper plateau of the kinetic transition defined by Pmax represents the full extent of folding, as determined from equilibrium experiments. Variations in sample loading can be controlled by normalizing the radioactivity in the bands of interest to another band in the gel whose intensity remains constant over the experiment. In general, the authors find that extreme variations in sample recovery increase the scatter of the data, even after normalization. Transitions that occur on the 0.1- to 10-sec timescales are easily monitored by the current state of equipment at beamline X28C. Slower
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transitions (1 to 10 min) can be more difficult to detect, because the long aging times in the sample loop lead to poor sample recovery. Processes occurring on timescales of <50 msec cannot be easily probed by X-ray footprinting using the current experimental protocol. It is hoped that future improvements to the beamline will shorten the deadtime of the experiment to several milliseconds.
Time Considerations Experiments should be planned 1 to 2 months in advance. Preparation of samples for shipment to the beamline requires 3 days, and samples must be shipped at least 24 hr in advance. Careful planning and execution are essential, as experiments at the beamline are often scheduled over a 1- to 3-day period. Roughly 1 week should be allowed for gel electrophoresis and for analysis of the data.
Literature Cited Balasubramanian, B., Pogozelski, W.K., and Tullius, T.D. 1998. DNA strand breaking by the hydroxyl radical is governed by the accessible surface areas of the hydrogen atoms of the DNA backbone. Proc. Natl. Acad. Sci. U.S.A. 95:9738-9743. Breen, A.P. and Murphy, J.A. 1995. Reactions of oxyl radicals with DNA. Free Radic. Biol. Med. 18:1033-1077. Brenowitz, M., Senear, D.F., Shea, M.A., and Ackers, G.K. 1986. Quantitative DNase footprint titration: A method for studying protein-DNA interactions. Methods Enzymol. 130:132-181. Cate, J.H., Gooding, A.R., Podell, E., Zhou, K., Golden, B.L., Kundrot, C.E., Cech, T.R., and Doudna, J.A. 1996. Crystal structure of a group I ribozyme domain: Principles of RNA packing. Science 273:1678-1685. Celander, D.W. and Cech, T.R. 1990. Iron(II)ethylenediaminetetraacetic acid catalyzed cleavage of RNA and DNA oligonucleotides: Similar reactivity toward single- and double-stranded forms. Biochemistry 29:1355-1361. Celander, D.W. and Cech, T.R. 1991. Visualizing the higher order folding of a catalytic RNA molecule. Science 251:401-407. Chaulk, S.G. and MacMillan, A.M. 2000. Kinetic footprinting of an RNA-folding pathway using peroxynitrous acid. Angew. Chem. Int. Ed. Engl. 39:521-523. Deras, M.L., Brenowitz, M., Ralston, C.Y., Chance, M.R., and Woodson, S.A. 2000. Folding mechanism of the Tetrahymena ribozyme P4-P6 domain. Biochemistry 39:10975-10985.
Dhavan, G.M., Chance, M.R., and Brenowitz, M. 2001. Kinetics analysis of DNA-protein interactions by time resolved synchrotron X-ray footprinting. In Practical Applications. In press. Dixon, W.J., Hayes, J.J., Levin, J.R., Weidner, M.F., Dombroski, B.A., and Tullius, T.D. 1991. Hydroxyl radical footprinting. Methods Enzymol. 208:380-413. Dizdaroglu, M. and Bergtold, D.S. 1986. Characterization of free radical-induced base damage in DNA at biologically relevant levels. Anal. Biochem. 156:182-188. Hampel, K.J. and Burke, J.M. 2001. Time-resolved hydroxyl-radical footprinting of RNA using Fe(II)-EDTA. Methods 23:233-239. King, P.A., Jamison, E., Strahs, D., Anderson, V.E., and Brenowitz, M. 1993. ‘Footprinting’ proteins on DNA with peroxynitrous acid. Nucl. Acids Res. 21:2473-2478. Klassen, N.V. 1987. Primary products in radiation chemistry. In Radiation Chemistry: Principles & Applications (I. Farhatazis and M.A. Rodgers, eds.) pp. 29-61. VCH Publishers, New York. Latham, J.A. and Cech, T.R. 1989. Defining the inside and outside of a catalytic RNA molecule. Science 245:276-282. Milligan, J.F. and Uhlenbeck, O.C. 1989. Synthesis of small RNAs using T7 RNA polymerase. Methods Enzymol. 180:51-62. Milligan, J.F., Groebe, D.R., Witherell, G.W., and Uhlenbeck, O.C. 1987. Oligoribonucleotide synthesis using T7 RNA polymerase and synthetic DNA templates. Nucl. Acids Res. 15:8783-8798. Ralston, C.Y., He, Q., Brenowitz, M., and Chance, M.R. 2000a. Stability and cooperativity of individual tertiary contacts in RNA revealed through chemical denaturation. Nat. Struct. Biol. 7:371-374. Ralston, C.Y., Sclavi, B., Sullivan, M., Deras, M.L., Woodson, S.A., Chance, M.R., and Brenowitz, M. 2000b. Time-resolved synchrotron X-ray footprinting and its application to RNA folding. Methods Enzymol. 317:353-368. Sclavi, B., Woodson, S., Sullivan, M., Chance, M.R., and Brenowitz, M. 1997. Time-resolved synchrotron X-ray “footprinting”, a new approach to the study of nucleic acid structure and function: Application to protein-DNA interactions and RNA folding. J. Mol. Biol. 266:144-159. Sclavi, B., Sullivan, M., Chance, M.R., Brenowitz, M., and Woodson, S.A. 1998a. RNA folding at millisecond intervals by synchrotron hydroxyl radical footprinting. Science 279:1940-1943. Sclavi, B., Woodson, S., Sullivan, M., Chance, M., and Brenowitz, M. 1998b. Following the folding of RNA with time-resolved synchrotron X-ray footprinting. Methods Enzymol. 295:379-402.
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Strahs, D. and Brenowitz, M. 1994. DNA conformational changes associated with the cooperative binding of cI-repressor of bacteriophage lambda to OR. J. Mol. Biol. 244:494-510. Tullius, T.D. and Dombroski, B.A. 1985. Iron(II) EDTA used to measure the helical twist along any DNA molecule. Science 230:679-681. Wallace, S.S. 1998. Enzymatic processing of radiation-induced free radical damage in DNA. Radiat. Res. 150:S60-79.
Contributed by Sarah A. Woodson and Michael L. Deras Johns Hopkins University Baltimore, Maryland Michael Brenowitz Albert Einstein College of Medicine of Yeshiva University Bronx, New York
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Rapid Magnesium Chelation as a Method to Study Real-Time Tertiary Unfolding of RNA
UNIT 11.7
This unit describes a method to measure the unfolding of RNA tertiary structure on a millisecond timescale. A stopped-flow instrument is designed to mix two or more solutions (generally up to four) on a millisecond time scale. After mixing, the resulting solution is moved by a compressed gas into a flow cell for observation (e.g., by UV, CD, or fluorescence spectrometry). Using this methodology, rate constants for unfolding of tertiary to secondary structure can be obtained over a range of temperatures and these values can be used to construct Arrhenius or Eyring plots (Maglott et al., 1999), from which parameters such as activation energy (Ea), the Arrhenius pre-exponential factor (A; Cole et al., 1972; Cole and Crothers, 1972), the enthalpy of activation (∆H†), and the entropy of activation (∆S†) can be obtained (Maglott and Glick, 1997). These data provide information about the energy of the transition state and the energy barriers between secondary and tertiary structure. This information is necessary to be able to predict RNA tertiary structure from secondary structure, since there can be significant kinetic barriers to tertiary folding even if there is only a small energy difference between the secondary and tertiary structures. This procedure requires prior expertise in the use of a stopped-flow spectrophotometer. General information on this subject may be found, e.g., in Fersht (1985). NOTE: All procedures should be conducted using sterile technique. NOTE: When handling RNA, suitable precautions should be taken to avoid RNase contamination (see APPENDIX 2A). MEASUREMENT OF UNFOLDING RATES OF RNA TERTIARY STRUCTURE
BASIC PROTOCOL
A stopped-flow spectrophotometer is used to measure the rate of unfolding of an RNA tertiary structure that is stabilized by the presence of Mg2+ ions. Unfolding of tertiary structure elements that occurs in the first few milliseconds, as well as at longer time intervals, after removal of the Mg2+ by a chelator (EDTA) can be observed using this methodology. Stopped-flow instruments are available from a number of vendors, listed below. The most important features are low deadline (≤2 msec), microsample volume (e.g., 50 µL), and the ability for multiple observation techniques (commonly, UV, CD, and fluorescence). Materials EDTA buffer (see recipe) at EDTA concentration needed to chelate all Mg2+ in RNA sample Mg2+ buffer (see recipe) Folded RNA sample (see Support Protocol 1) Syringe filters, 0.22-µm 10-mL and 3-mL luer-tip syringes (e.g., Fisher) Parafilm Stopped-flow spectrophotometer (e.g., Applied Photophysics SX18.MV or equivalent instruments from Olis Instruments or Hi-Tech Scientific) Side-arm Erlenmeyer flask RNA Folding Pathways Contributed by Emily J. Maglott and Gary D. Glick Current Protocols in Nucleic Acid Chemistry (2001) 11.7.1-11.7.11 Copyright © 2001 by John Wiley & Sons, Inc.
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Additional reagents and equipment for folding RNA (see Support Protocol 1) and analyzing kinetic traces to determine unfolding rates and activation parameters (see Support Protocol 3) Prepare solutions 1. On the basis of pilot experiments, choose a concentration of EDTA that is sufficient to chelate all Mg2+ from the RNA sample during the deadtime of the stopped-flow spectrophotometer. It is critical that all of the Mg2+ be chelated in the deadtime of the instrument (i.e., before the observation begins). This will always require that EDTA (or whatever chelating agent is used) be present in excess relative to the Mg2+. The way this can be judged is to measure the unfolding rates with increasing amount of EDTA. When the rate does not change, there is enough EDTA. As a check of this, one should compare the total change in absorbance (or whatever physical parameter is being measured) and compare that change to value measured at equilibrium. For absorbance, one would measure (in a normal spectrophotometer) the absorbance of an RNA solution in the presence and absence of Mg2+. The difference should be within error of the value measured as described above. IMPORTANT NOTE: The [EDTA] needed to fulfill this requirement may vary with the [Mg2+] and the deadtime of the instrument. For an instrument with a deadtime of 1.4 msec, a 6-fold excess of EDTA to Mg2+ was found to give reproducible results with a high signal-to-noise ratio. Lower EDTA concentrations gave similar kinetic results (rates and activation parameters) with lower signal-to-noise ratios due to the higher pH differential needed to maintain a constant final pH (Maglott, 1998; Maglott et al., 1998). Experiments should be conducted with a variety of [EDTA]:[Mg2+] ratios to help verify that all Mg2+ is being removed during the deadtime of the instrument and that tertiary unfolding is monitored (Maglott and Glick, 1997; Maglott et al., 1998).
2. Filter both the Mg2+ buffer and the EDTA buffer using 0.22-µM syringe filters. 3. Degas the EDTA buffer by loading ∼6 mL of the EDTA buffer into a 10-mL luer-tip syringe, holding a piece of Parafilm over the end of the syringe with a finger, and (holding the syringe tip up) pulling back on the plunger slowly, allowing all air bubbles to move to the top of the syringe. Remove the Parafilm and gently expel all of the air from the syringe. Repeat the procedure 2 to 3 times until the air has been removed. Set up instrument 4. Load the degassed EDTA buffer into one of the two drive syringes of the stopped-flow spectrophotometer, being careful not to introduce air bubbles into the drive syringe. In principle, it does not matter into which of the two drive syringes the EDTA solution is introduced; however, if one of the drive syringes has a larger prime volume than the other on the particular stopped-flow instrument being used, use this drive syringe for the EDTA solution. Then the drive syringe with the smaller prime volume will be used for the RNA solution; thus, less RNA will be required to conduct the experiments.
5. Load Mg2+ buffer into the second drive syringe. 6. Set the wavelength of the stopped-flow spectrophotometer to 268 nm.
Rapid Magnesium Chelation as a Method to Study Real-Time Teriary Unfolding of RNA
The usual RNA absorbance maximum of 260 nm cannot be used for these experiments because the EDTA buffer absorbs at this wavelength and creates a significant background absorbance. At 268 nm the absorbance due to the buffer substantially decreases, while the absorbance of the RNA remains sufficient. While this protocol describes the use of hyperchromicity changes to monitor the unfolding of RNA tertiary structure, either fluorescence or circular dichroism could also be used if tertiary unfolding is accompanied by a suitable spectral change.
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7. Acquire 10 to 15 kinetic traces of EDTA buffer versus Mg2+ buffer to fill the observation cell with a blank solution. This procedure primes the flow line to the cell with the EDTA solution. The number of acquisitions may need to be adjusted depending on the length and volume of the flow lines.
8. Equilibrate the sample cell and the drive syringes of the stopped-flow spectrophotometer at the temperature where the unfolding rate will be measured. The temperature range over which the unfolding rates will be measured is determined from the stability of both tertiary and secondary structure. In this temperature range, tertiary structure should be adopted in the presence of Mg2+ while in the absence of Mg2+ the secondary structure is stable. Furthermore, the rate of tertiary unfolding must be slow enough at these temperatures to be measured by the particular instrument being used. For yeast tRNAPhe this range was 28° to 35°C in a 32 mM Na+ buffer (Maglott and Glick, 1997) while for unmodified yeast tRNAPhe the range was 20° to 32°C in a 100 mM Na+ buffer (Maglott et al., 1998).
9. Zero the absorbance reading of the blank solution. Be sure that the lamp has been warmed up and its output has stabilized. Follow the manufacturer’s instructions.
Perform unfolding experiment 10. Remove the Mg2+ buffer from the second drive syringe and load the folded RNA sample (see Support Protocol 1) into the drive syringe. Degas after folding but before loading into syringe. The RNA solution can be degassed as in step 3, but using a 3-mL luer-tip syringe.
11. Equilibrate both the RNA solution and the EDTA solution (in their respective drive syringes) at the desired temperature for 15 min. 12. Acquire three kinetic traces to prime the line containing the RNA solution. The number of acquisitions needed to prime the solution in the observation cell may vary with the instrument. See the manufacturer’s instructions. To collect the RNA sample as it is used, have the output line in a small side-arm Erlenmeyer flask. Recovered RNA samples can be checked for degradation by denaturing PAGE, repurified if necessary, and reused by first dialyzing to remove the EDTA and Mg2+ and then dialyzing into the desired Mg2+-containing buffer.
13. At the highest resolution possible, acquire as many kinetic traces as sample quantity allows. Typically it is desirable to have seven good kinetic traces at each set of conditions. The number of traces obtained from a 1-mL sample will depend on the volume of sample used in each acquisition. To maximize the use of the RNA samples, the lowest possible volume with a reasonable deadtime should be used. For example, with an Applied Photophysics SX18.MV, only 50 mL of RNA is needed for a single trace, and the deadtime of this instrument is 1.4 msec.
14. Analyze the kinetic traces (see Support Protocol 3). FOLDING AND EQUILIBRATION OF RNA SAMPLES It is important that the RNA being used in the kinetic experiments be folded into the native tertiary structure. This protocol describes an optimized procedure for folding of both yeast tRNAPhe and unmodified yeast tRNAPhe. For other RNAs, optimized folding protocols can be developed by changing either the temperature (bath temperature and/or equilibration
SUPPORT PROTOCOL 1
RNA Folding Pathways
11.7.3 Current Protocols in Nucleic Acid Chemistry
Supplement 6
temperature) or the time spent at each temperature to ensure that the RNA has adopted the native tertiary structure. Materials Dialyzed RNA sample (see Support Protocol 2) Mg2+ buffer (see recipe) 75°C water bath Spectrophotometer 1. Equilibrate a water bath at ∼75°C. 2. Place a sample of RNA that has been dialyzed into the desired Mg2+ buffer (see Support Protocol 2) in a microcentrifuge tube. Dilute to 1 mL total volume with additional Mg2+ buffer. 3. Measure the absorbance of the sample at 268 nm in a regular laboratory spectrophotometer. The absorbance of the sample should be high enough that the absolute absorbance change observed upon unfolding will be >0.002 OD268. For example, 0.6 OD260/mL of yeast tRNAPhe was used to study tertiary unfolding in a 32 mM Na+ buffer (Maglott and Glick, 1997) and the observed absorbance changes were from 4 to 20 mOD268 in a temperature range of 28° to 34°C. More RNA can be used, but the background absorbance will increase as well.
4. Place the closed microcentrifuge tube into the 75°C water bath for 5 min. 5. Remove the sample from the water bath and equilibrate it at room temperature for 30 min. The folded RNA cannot be stored and must be used immediately. SUPPORT PROTOCOL 2
FLOW DIALYSIS OF RNA SAMPLES This protocol describes a procedure for equilibrium flow dialysis of the RNA sample into the Mg2+ buffer used in the kinetic experiment. Use of this procedure ensures that the total concentration of Mg2+ in the RNA sample is accurately known. The total [Mg2+] of an RNA sample that has simply been diluted into the Mg2+ buffer may be influenced by the presence of Mg2+ ions or chelators that are present due to previous handling of the RNA sample (e.g., PAGE purification or ethanol precipitation). Materials Mg2+ buffer (see recipe) Argon source Dry RNA sample of interest
Rapid Magnesium Chelation as a Method to Study Real-Time Teriary Unfolding of RNA
Bottle-top filtration apparatus containing a 0.22-µm filter (e.g., Corning) 10-well microdialyzer (Spectrum) Peristaltic pump Cellulose ester membrane for use with 10-well microdialyzer (Spectrum), MWCO 5000 1. Filter 2 L of Mg2+ buffer under vacuum using a bottle-top filtration device containing a 0.22-µm filter.
11.7.4 Supplement 6
Current Protocols in Nucleic Acid Chemistry
The Mg2+ buffer used in these experiments was designed for unmodified yeast tRNAPhe to satisfy the requirement that the secondary structure of the tRNA form in the absence of Mg2+ and the tertiary structure form in the presence of Mg2+. For other RNAs, the [Mg2+] may need to be altered, and the composition of the buffer may also need to be changed to meet the structural requirements of the experiment.
2. Bubble argon gas through the buffer for at least 30 min. 3. Assemble a 10-well microdialyzer with a peristaltic pump according to manufacturer’s instructions using a 5000 MWCO cellulose ester membrane. This procedure has been optimized for tRNA samples with molecular weights of ∼25 kDa. For larger or smaller RNAs the MWCO of the membrane may need to be changed.
4. Fill the chamber of the microdialyzer with Mg2+ buffer, being careful not to allow formation of air bubbles under the wells. Holding the dialyzer at a slant with the buffer inlet at the bottom and the buffer outlet at the top allows the air within the chamber to be forced out of the outlet without allowing air bubbles to become trapped under the wells.
5. Dissolve a dry RNA sample in 100 to 200 µL of Mg2+ buffer and transfer the solution to a well of the microdialyzer. The amount of RNA that can be used in this procedure depends on the amount of RNA needed to conduct the experiments. Both small and large quantities of RNA can be prepared using this procedure; however, if the RNA sample being used already contains a high concentration of salt, the volume of the RNA solution may significantly expand during dialysis and the final concentration of the RNA may be too low to be useful. It is recommended that the samples be desalted by either ethanol precipitation or flow dialysis against TE buffer, pH 7.0 (see APPENDIX 2A for recipe), followed by ethanol precipitation, to prevent this difficulty.
6. Rinse the tube with another 100 µL of Mg2+ buffer and add this solution to the RNA sample in the dialyzer. 7. Dialyze the RNA against the Mg2+ buffer at a flow rate of 5 mL/min (controlled by a peristaltic pump). A slower flow rate may also be used and may be useful for RNA samples containing higher concentrations of residual salts, to prevent significant volume expansion of the solution as it reaches equilibrium in the new buffer system.
8. When ∼50 mL of the Mg2+ buffer remains in the reservoir, change the flow rate to 0 mL/min. Reserve this buffer for dilution of the sample to the appropriate concentration during the kinetic experiments. 9. Using a micropipettor, remove the RNA solution from the microdialyzer and place into a microcentrifuge tube. Rinse the well of the dialyzer at least two times with Mg2+ buffer and add to the microcentrifuge tube. 10. Use sample immediately or store at 4°C for up to ∼1 week. RNA samples kept in solution for >1 week may begin to degrade. It is not advisable to freeze the sample in solution, as repeated freeze-thawing of an RNA sample may also cause degradation.
RNA Folding Pathways
11.7.5 Current Protocols in Nucleic Acid Chemistry
Supplement 6
SUPPORT PROTOCOL 3
DETERMINATION OF TERTIARY UNFOLDING RATES AND ACTIVATION PARAMETERS Materials Graphical analysis software (e.g., Kaleidagraph from Synergy Software) Statistical analysis software (e.g., SAS from SAS Institute) 1. Overlay the individual traces, discarding any that contain artifacts or are particularly different from the majority. Do not include the kinetic traces obtained while priming the RNA solution.
2. Average these overlaid traces into a single trace. 3. Attempt to fit this average trace (excluding any datapoints collected during the deadtime of the experiment) to a single exponential curve: absorbance = a × exp(−rate × t), where a is the amplitude of the kinetic trace. All stopped-flow instruments come with fitting software; see manufacturer’s instructions to carry out these steps. If the process is single-exponential, then the residual to the fit should not show any distinct patterns and should fluctuate between 10−4 and 10−3. The best fit is the one that has the lowest residual value.
4. If there is a pattern to the residual (i.e., sinusoidal) or if the residual is large (>10−3), then repeat the fitting procedure to increasing numbers of exponential components until a good fit is obtained. For example, for a two-exponential fit, absorbance = a1 × exp(−rate × t) + a2 × exp(−rate × t), where an is the amplitude of one phase of the kinetic trace. If the tertiary unfolding kinetics are not single-exponential, then analysis of activation parameters is more complicated and a mechanism for the unfolding must be established.
5. Obtain the standard deviation of the rate of unfolding at each set temperature from the unfolding rates of each of the individual traces used to obtain the average trace. The rate (k) is the average rate for each temperature obtained from the individual fitted rate data. During the course of doing individual fits, one may notice that some traces are not of high quality (e.g., with high signal-to-noise ratios or air bubble artifacts). It may be desirable to create an average trace that excludes one or more traces if this occurs. However, if most of the traces contain artifacts, the source of the artifacts should be determined and corrected, and the experiment should be repeated.
6. Construct an Eyring plot, from which transition-state parameters can be obtained, using the rates obtained at several different temperatures. To obtain ∆H†, ∆S†, and ∆G†, plot ln[k/(kBT/h)] versus 1/T (where T is expressed in K, degrees Kelvin) and where kB = Boltzmann’s constant and h = Planck’s constant. Use the slope of this line, ∆H†/R, and the intercept, ∆S†/R (where R is the gas constant) to calculate ∆G†, using the relation ∆G† = ∆H† − T∆S†.
Rapid Magnesium Chelation as a Method to Study Real-Time Teriary Unfolding of RNA
The error on ∆G† can be obtained by calculating the variance on both the slope and the intercept as well as the covariance between the slope and the intercept and using the following equation: var(∆G†) = R2[var(slope)] + (0.298R)2[var(intercept)] + 2R(0.298R)(covar) where var(slope) is the variance on the value of the slope, var(intercept) is the variance on the value of the intercept, and covar is the covariance between the slope and the intercept (Walpole and Myers, 1978).
11.7.6 Supplement 6
Current Protocols in Nucleic Acid Chemistry
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. Use >99% pure reagents containing <0.0005% divalent metal ion impurities. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Disodium EDTA stock solution, 150 mM Dissolve 55.8 g of disodium EDTA in 1 L of water. Stable for several months at room temperature. EDTA buffer 1.25 mL of 200 mM sodium cacodylate stock (see recipe; 10 mM final) 0.15 mL of 5 M NaCl (APPENDIX 2A; 30 mM final) 5 mL of 150 mM disodium EDTA stock (see recipe; 30 mM final) Dilute to 25 mL with deionized water Adjust to pH 6.8 with HCl or NaOH Prepare fresh Alternate EDTA buffers can be made by adjusting the [NaCl] so that the total [Na+] remains at 100 mM. The pH of the EDTA solution should be adjusted so that upon 1:1 mixing with the Mg2+ buffer the final pH remains at the pH of the original Mg2+ buffer.
Mg2+ buffer 100 mL of 200 mM sodium cacodylate stock (see recipe; 10 mM final) 36 mL of 5 M NaCl (APPENDIX 2A; 90 mM final) 50 mL of 200 mM MgCl2 stock (see recipe; 5 mM final) Dilute to 2 L with deionized water Adjust to pH 6.8 with HCl or NaOH Prepare fresh 5 mM is the minimum Mg2+ concentration needed to fold the particular RNA described here. The Mg2+ concentration needed to fold a sample will vary with the RNA and must be optimized (see UNIT 6.3, Support Protocol 3).
MgCl2 stock solution, 200 mM Dissolve 10.2 g of magnesium chloride hexahydrate in 250 mL water. Stable for several months at room temperature. Sodium cacodylate stock solution, 200 mM Dissolve 32.0 g of sodium cacodylate in 1 L of water. Stable for several months at room temperature. COMMENTARY Background Information Since the tertiary structures of RNAs are necessary for biological activity, understanding the folded structures of RNAs and the interactions that stabilize them is clearly important. However, knowledge of the final folded structure cannot reveal the processes by which these molecules fold, beginning from a primary nucleotide sequence. Elucidating the energetics involved in folding of an RNA from secondary structure to tertiary structure should enable a better understanding of why one structure forms in preference to another, and may provide insight into conformational changes that occur during biological processes such as protein
synthesis, which may involve local unfolding of tertiary structure elements (LeCuyer and Crothers, 1993, 1994). Furthermore, knowledge of the energetics of tertiary structure formation will also enable prediction of tertiary structure from secondary structure. Algorithms are available to predict secondary structures from primary sequence information (Zuker, 1989), and expansion of these algorithms to include prediction of tertiary structure will allow for the construction of RNAs with defined structures and, possibly, defined functions. However, predictions of tertiary structure formation from secondary structure must also include the energy barriers to folding. To fully
RNA Folding Pathways
11.7.7 Current Protocols in Nucleic Acid Chemistry
Supplement 6
Rapid Magnesium Chelation as a Method to Study Real-Time Teriary Unfolding of RNA
understand RNA folding, it is necessary to characterize the structure of the folded RNA, any intermediates on the folding pathway, and the rate-determining transition state. Relatively little is known about the rates of tertiary folding and unfolding of RNA, particularly on timescales faster than seconds. Recent approaches to investigate tertiary folding have used both the intrinsic properties of native structure to report on folding transitions, such as self-splicing for ribozymes (Jaeger et al., 1993; Emerick and Woodson, 1994; Emerick et al., 1996; Pan and Sosnick, 1997), as well as indirect techniques like oligonucleotide hybridization (Zarrinkar and Williamson, 1994; Zarrinkar et al., 1996), UV cross-linking (Downs and Cech, 1996), and chemical modification (Emerick and Woodson, 1994; Banerjee and Turner, 1995; Chance et al., 1997; Sclavi et al., 1997). However, these methods cannot monitor transitions that occur faster than about several events per minute. Several stopped-flow mixing techniques have been developed that can monitor folding transitions on the millisecond timescale. For example, conformational changes associated with substrate binding to ribozymes have been monitored using substrates tagged with a fluorescent adduct (Bevilacqua et al., 1992, 1994; Qin and Pyle, 1997). An especially interesting new approach couples rapid folding, initiated by addition of Mg2+, with hydroxyl radical footprinting (Sclavi et al., 1998; UNIT 11.6). This technique assesses regions of an RNA that become inaccessible to hydroxyl radicals as folding proceeds over time. However, generation of the hydroxyl radicals must occur faster than folding, which is achieved by generating the radicals using a synchrotron X-ray source. A more general method follows folding, again initiated by addition of Mg2+ to an RNA void of divalent metals, using UV (UNIT 11.3) and CD (UNIT 11.5) spectroscopies as well as catalytic activity (Pan and Sosnick, 1997). Since tertiary folding of a ribozyme is required for catalytic activity, rates of catalysis are readily correlated to rates of folding. On the other hand, both UV and CD spectroscopy are sensitive to basestacking and base-pairing, interactions that occur in both secondary and tertiary structure formation. However, verification that the spectroscopic signal being monitored arises solely from tertiary structure formation requires determination of the structure of the RNA in both the presence and absence of Mg2+ under the specific conditions (i.e., buffer and temperature) of the experiment.
Temperature-jump (T-jump) relaxation is one of the most general methods to measure unfolding transitions in RNA that occur on a timescale faster than seconds. The T-jump technique involves rapid heating of an RNA solution, and unfolding of the RNA is monitored spectroscopically as the solution establishes an equilibrium at the higher temperature. Much of our understanding of RNA folding comes from the classic work of Crothers (Cole et al., 1972; Yang and Crothers, 1972; Crothers et al., 1974; Crothers and Cole, 1978), Maass (Riesner et al., 1970, 1973; Urbanke et al., 1973, 1975; Coutts et al., 1975), and Biltonen (Levy et al., 1972), who used T-jump to study the folding and unfolding kinetics of transfer RNAs (tRNA). While T-jump is an extremely powerful technique, it does have some limitations. T-jump equipment is expensive and not widely available, current commercial instrumentation uses relatively large sample volumes (∼1 mL), and photobleaching can lead to sample degradation. Furthermore, some conformational changes are not thermally accessible, and these transitions cannot be monitored using T-jump. For example, tertiary folding of the P4-P6 domain of the Tetrahymena ribozyme is in a Mg2+dependent equilibrium (Szewczak and Cech, 1997) and, since unfolding of secondary and tertiary structure tends to be coupled in the presence of Mg2+, the tertiary-secondary unfolding transition is not thermally accessible. The Basic Protocol describes a simple, noninvasive method to determine the rate and activation parameters associated with unfolding of native RNA tertiary structure. This method exploits the fact that formation of RNA tertiary structure often requires Mg2+ or other divalent metals (Uhlenbeck, 1987; Celander and Cech, 1991; Kazakov and Altman, 1991; Christian and Yarus, 1993; Pyle, 1993; Butcher and Burke, 1994; Laing et al., 1994; Lu and Draper, 1994; Draper et al., 1995; Bassi et al., 1996; Narlikar and Herschlag, 1996), and it entails mixing an RNA solution containing Mg2+ with excess EDTA in a stopped-flow instrument under conditions of temperature and ionic strength where tertiary structure unfolds after chelation of Mg2+. Low-millisecond resolution can be achieved, and the unfolding transition can be observed by standard absorbance measurements. The results presented show that the stopped-flow technique is a robust and versatile method that in principle can be used to examine a range of systems.
11.7.8 Supplement 6
Current Protocols in Nucleic Acid Chemistry
Critical Parameters
Anticipated Results
The most important consideration in this experiment is that, in the same buffer system, the tertiary structure of the RNA form in the presence of Mg2+ and the secondary structure of the RNA form in the absence of Mg2+. Secondly, the unfolding of tertiary structure to the secondary structure must be accompanied by a spectroscopic change (e.g., absorbance, fluorescence, or circular dichroism). Finally, the chelation of the Mg2+ by the EDTA must occur completely and irreversibly (the rate of disassociation of Mg2+ from EDTA is 0.55 sec−1) within the deadtime of the spectrophotometer so that the rate of the transition observed will reflect only unfolding. Control experiments should be conducted to verify that the signal change being observed is due to tertiary unfolding. These may include (1) replacing the EDTA solution with a sodium acetate solution of equivalent anionic charge concentration, and (2) decreasing the temperature of the experiment below where tertiary unfolding will occur upon removal of Mg2+. Furthermore, rapid mixing of the Mg2+ buffer with the EDTA buffer should not give rise to any absorbance change at 268 nm.
The unfolding of the tertiary structure of unmodified yeast tRNAPhe was characterized by single exponential kinetics with rate constants ranging from 200 sec–1 to 900 sec–1 over a temperature range of 20° to 32°C (Maglott et al., 1998). There was a 3% change in absorbance observed upon unfolding of the tRNA. In this temperature range, the Arrhenius and Eyring plots constructed from these data were linear, and activation parameters were obtained with a ∆G† of tertiary unfolding of 14 kcal/mol. The rate, absorbance change, and activation parameters obtained for a specific RNA will probably vary from the results that the authors have obtained using unmodified yeast tRNAPhe.
Troubleshooting Table 11.7.1 describes several problems that can arise in the course of tertiary unfolding experiments, along with their possible causes and solutions.
Table 11.7.1
Time Considerations The kinetic experiments described here can be conducted in a few hours. Preparation of the sample and equilibration take ~45 min. Acquisition of the individual kinetics traces takes only a few minutes. Analysis of the kinetic traces and any subsequent data handling can be done at any time after the data is collected and can take several hours. Preparation of the RNA sample in the desired buffer using continuous flow dialysis usually takes 6 hr, although the time needed to effect complete buffer transfer will vary depending on the concentration and identity of the buffers being exchanged.
Troubleshooting Guide for Tertiary Unfolding Kinetics
Problem No hyperchromic change observed
Artifacts observed in kinetic traces
Possible cause RNA was not folded
Solution Fold the RNA
[Mg2+] is not sufficiently high to Increase [Mg2+] in buffer fold the RNA Tertiary structure does not unfold at the temperature used to conduct the experiment Increase the temperature at which the experiments are conducted Dissolved oxygen in the buffers Degas all buffer solutions Particulate in the buffers Residue in the sample cell
Filter all buffer solutions with 0.2-µm syringe filters Clean the sample cell following manufacturer’s directions
RNA Folding Pathways
11.7.9 Current Protocols in Nucleic Acid Chemistry
Supplement 6
Literature Cited Banerjee, A.R. and Turner, D.H. 1995. The time dependence of chemical modification reveals slow steps in the folding of a group I ribozyme. Biochemistry 34:6504-6512. Bassi, G.S., Murchie, A.I.H., and Lilley, D.M.J. 1996. The ion-induced folding of the hammerhead ribozyme: Core sequence changes that perturb folding into the active conformation. RNA 2:756-768. Bevilacqua, P.C., Kierzek, R., Johnson, K.A., and Turner, D.H. 1992. Dynamics of ribozyme binding of substrate revealed by fluorescence-detected stopped-flow methods. Science 258:13551358. Bevilacqua, P.C., Li, Y., and Turner, D.H. 1994. Fluorescence-detected stopped flow with a pyrene labeled substrate reveals that guanosine facilitates docking of the 5′ cleavage site into a high free energy binding mode in the Tetrahymena ribozyme. Biochemistry 33:11340-11348. Butcher, S.E. and Burke, J.M. 1994. A photo-crosslinkable tertiary structure motif found in functionally distinct RNA molecules is essential for catalytic function of the hairpin ribozyme. Biochemistry 33:992-999. Celander, D.W. and Cech, T.R. 1991. Visualizing the higher order folding of a catalytic RNA molecule. Science 251:401-407. Chance, M.R., Sclavi, B., Woodson, S.A., and Brenowitz, M. 1997. Examining the conformational dynamics of macromolecules with timeresolved synchrotron X-ray footprinting. Structure 5:865-869. Christian, E.L. and Yarus, M. 1993. Metal coordination sites that contribute to structure and catalysis in the group I intron from Tetrahymena. Biochemistry 32:4475-4480. Cole, P.E. and Crothers, D.M. 1972. Conformational changes of transfer ribonucleic acid: Relaxation kinetics of the early melting transition of methionine transfer ribonucleic acid (Escherichia coli). Biochemistry 11:4368-4374. Cole, P.E., Yang, S.K., and Crothers, D.M. 1972. Conformational changes of transfer ribonucleic acid: Equilibrium phase diagrams. Biochemistry 11:4358-4368. Coutts, S.M., Riesner, D., Römer, R., Rabl, C.R., and Maass, G. 1975. Kinetics of conformational changes in tRNAPhe (yeast) as studied by the fluorescence of the Y-base and of formycin substituted for the 3′-terminal adenosine. Biophys. Chem. 3:275-289. Crothers, D.M. and Cole, P.E. 1978. Conformational changes of tRNA. In TransferRNA (S. Altman, ed.) pp. 196-247. MIT Press, Cambridge, Mass. Rapid Magnesium Chelation as a Method to Study Real-Time Teriary Unfolding of RNA
Downs, W.D. and Cech, T.R. 1996. Kinetic pathway for folding of the Tetrahymena ribozyme revealed by three UV-inducible crosslinks. RNA 2:718-732.
Draper, D.E., Xing, Y., and Laing, L.G. 1995. Thermodynamics of RNA unfolding: Stabilization of a ribosomal RNA tertiary structure by thiostrepton and ammonium ion. J. Mol. Biol. 249:231238. Emerick, V.L. and Woodson, S.A. 1994. Fingerprinting the folding of group I precursor RNA. Proc. Natl. Acad. Sci. U.S.A.91:9675-9679. Emerick, V.L., Pan, J., and Woodson, S.A. 1996. Analysis of rate-determining conformational changes during self-splicing of the Tetrahymena intron. Biochemistry 35:13469-13477. Fersht, A. 1985. Enzyme Structure and Mechanism, 2nd Ed., p. 122. W.H. Freeman, New York. Jaeger, L., Westhof, E., and Michel, F. 1993. Monitoring of the cooperative unfolding of the sunY group I intron of bacteriophage T4. J. Mol. Biol. 234:331-346. Kazakov, S. and Altman, S. 1991. Site-specific cleavage by metal ion cofactors and inhibitors of M1 RNA, the catalytic subunit of RNase P from Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 88:9193-9197. Laing, L.G., Gluick, T.C., and Draper, D.E. 1994. Stabilization of RNA structure by Mg ions: Specific and non-specific effects. J. Mol. Biol. 237:577-587. LeCuyer, K.A. and Crothers, D.M. 1993. The Leptomonas collosoma spliced leader RNA can switch between two alternate structural forms. Biochemistry 32:5301-5311. LeCuyer, K.A. and Crothers, D.M. 1994. Kinetics of an RNA conformational switch. Proc. Natl. Acad. Sci. U.S.A.91:3373-3377. Levy, J., Rialdi, G., and Biltonen, R. 1972. Thermodynamic studies of transfer ribonucleic acids. II. Characterization of the thermal unfolding of yeast phenylalanine-specific transfer ribonucleic acid. Biochemistry 11:4138-4144. Lu, M. and Draper, D.E. 1994. Bases defining an ammonium and magnesium ion-dependent tertiary structure within the large subunit ribosomal RNA. J. Mol. Biol. 244:572-585. Maglott, E.J. 1998. Structural and kinetic studies of tertiary folding of an unmodified transfer RNA. Ph.D. Thesis, University of Michigan. Maglott, E.J. and Glick, G.D. 1997. A new method to monitor the rate of conformational transitions in RNA. Nucl. Acids Res. 25:3297-3301. Maglott, E.J., Deo, S.S., Pryzkorska, A., and Glick, G.D. 1998. Conformational transitions of an unmodified tRNA: Implications for RNA folding. Biochemistry46:16349-16359. Maglott, E.J., Goodwin, J.T., and Glick, G.D. 1999. Probing an RNA tertiary structure unfolding transition state. J. Am. Chem. Soc. 121:74617492. Narlikar, G.J. and Herschlag, D. 1996. Isolation of a local tertiary folding transition in the context of a globally folded RNA. Nat. Struct. Biol. 3:701-709.
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Current Protocols in Nucleic Acid Chemistry
Pan, T. and Sosnick, T.R. 1997. Intermediates and kinetic traps in the folding of a large ribozyme revealed by circular dichroism and UV absorbance spectroscopies and catalytic activity. Nat. Struct. Biol. 4:931-938. Pyle, A.M. 1993. Ribozymes: A distinct class of metalloenzymes. Science 261:709-714. Qin, P.Z. and Pyle, A.M. 1997. Stopped-flow fluorescence spectroscopy of a group II intron ribozyme reveals that domain 1 is an independent folding unit with a requirement for specific Mg2+ ions in the tertiary structure. Biochemistry 36:4718-4730. Riesner, D., Römer, R., and Maass, G. 1970. Kinetic study of the three conformational transitions of alanine specific transfer RNA from yeast. Eur. J. Biochem. 15:85-91. Riesner, D., Maass, G. Thiebe, R., Philippsen, P., and Zachau, H.G. 1973. The conformational transitions in yeast tRNAPhe as studied with tRNAPhe fragments. Eur. J. Biochem. 36:76-88. Sclavi, B., Woodson, S., Sullivan, M., Chance, M.R., and Brenowitz, M. 1997. Time-resolved synchrotron X-ray “footprinting,” a new approach to the study of nucleic acid structure and function: Application to protein-DNA interactions and RNA folding. J. Mol. Biol. 266:144-159. Sclavi, B., Sullivan, M., Chance, M.R., Brenowitz, M., and Woodson, S. 1998. RNA folding at millisecond intervals by synchrotron hydroxyl radical footprinting. Science 279:1940-1943. Szewczak, A.A. and Cech, T.R. 1997. An RNA internal loop acts as a hinge to facilitate ribozyme folding and catalysis. RNA 3:838-849.
Uhlenbeck, O.C. 1987. A small catalytic oligoribonucleotide. Nature 328:596-601. Urbanke, C., Römer, R., and Maass, G. 1973. The binding of ethidium bromide to different conformations of tRNA: Unfolding of tertiary structure. Eur. J. Biochem. 33:511-516. Urbanke, C., Römer, R., and Maass, G. 1975. Tertiary structure of tRNAPhe (yeast): Kinetics and electrostatic repulsion. Eur. J. Biochem. 55:439444. Walpole, R.E. and Myers, R.H. 1978. Probability and Statistics for Engineers and Scientists, 2nd ed. Macmillan, New York. Yang, S.K. and Crothers, D.M. 1972. Conformational changes of transfer ribonucleic acid: Comparison of the early melting transition of two tyrosine-specific transfer ribonucleic acids. Biochemistry 11:4375-4381. Zarrinkar, P.P. and Williamson, J.R. 1994. Kinetic intermediates in RNA folding. Science 265:918924. Zarrinkar, P.P., Wang, J., and Williamson, J.R. 1996. Slow folding kinetics of RNase P RNA. RNA 2:564-573. Zuker, M. 1989. On finding all suboptimal foldings of an RNA molecule. Science 244:48-52.
Contributed by Emily J. Maglott and Gary D. Glick University of Michigan Ann Arbor, Michigan
RNA Folding Pathways
11.7.11 Current Protocols in Nucleic Acid Chemistry
Supplement 6
Use of Fluorescence Spectroscopy to Elucidate RNA Folding Pathways In this unit, some of the practical aspects of performing fluorescence-detected experiments on RNA are described. Fluorescence spectroscopy has proven to be a powerful tool for studying RNA folding, (for examples, see Bevilacqua et al., 1992; Clegg et al., 1992; Turner et al., 1996; Walter and Burke, 1997; Walter et al., 1998, 1999; Silverman and Cech, 1999). Either the ribozyme or a small oligonucleotide substrate may be labeled with a fluorescent probe. Fluorescence spectroscopy affords detection of rapid and slow kinetic processes, with real time acquisition of data. The first step for performing fluorescencedetected RNA folding experiments involves preparation of modified RNA. A discussion of the advantages and disadvantages of various pendant probes and nucleotide analogs is presented. Then, methods for observing changes in fluorescence are discussed. Some considerations on instrument setup and samples to test are provided. Control experiments that should be performed to assess whether the modification has perturbed the system are also discussed. A variety of kinetic, thermodynamic, and structure-probing experiments are suggested. Lastly, an overview of experiments for obtaining kinetic parameters are presented. General experimental methods and designs for accurately obtaining microscopic rate constants are discussed.
PREPARATION OF FLUORESCENTLY LABELED RNA The four natural bases in RNA are not sufficiently fluorescent for most applications; therefore, a fluorophore must be covalently attached to the RNA for study. In the case of two-piece systems involving a large ribozyme and a short substrate, the fluorophore may be engineered into the ribozyme or the substrate. The particular site of attachment is dictated by the scientific question of interest. The chosen fluorophore should have a strong excitation peak that does not overlap with the four bases. In general, excitation anywhere above 310 nm is preferable. Fluorophores used in RNA folding experiments may be separated into two classifications: pendant probes and nucleotide analogs.
UNIT 11.8
Pendant Probes Pendant probes such as fluorescein and pyrene can provide large changes in fluorescence emission upon folding (up to 25-fold has been observed in the case of pyrene). They may be readily attached to the ends of large or small RNAs prepared enzymatically or by chemical synthesis. For example, the 5′ end of RNA may be modified with a thiophosphate by suitable priming of the T7 RNA polymerase transcription of RNA. A wide variety of commercially available thiol-reactive probes may then be coupled to the RNA; Molecular Probes (http://www.probes.com) is an ex cellent source. Alternatively, oligonucleotides can be synthesized with a 5′ amino group and conjugated to an amine-reactive derivative of a fluorescent dye (Kierzek et al., 1993). The 3′ end of the RNA may be modified by reaction with periodate to ring open the cis-diol to a dialdehyde. A variety of amine-containing fluorophores can then be coupled to the 3′ end of the RNA (Reines and Cantor, 1974). Coupling reactions should include a control in which an unmodified RNA is used, and lack of reactivity with the probe should be confirmed. Specific conditions for modifications can be found in the literature from Molecular Probes. Pendant probes can also be inserted internally in the RNA. A short oligomer can be chemically synthesized with a 2′ amino or thiophosphate (available from Dharmacon; http://www.dharmacon.com) and then coupled to a pendant probe. Modified oligomers can be inserted into a large RNA by ligation. Alternatively, coupling of the probe to the modified RNA could be performed after ligation. Coupling before ligation offers the advantage of easier purification of a modified RNA that is short, although it has the potential disadvantage of interfering with the ligation reaction if it is near the site of ligation. Several factors can help guide the choice of a suitable fluorophore. Experiments with modified RNA have shown that pyrene emission is strongly quenched by pyrimidines (Kierzek et al., 1993; Manoharan et al., 1995; Netzel et al., 1995), while fluorescein emission is strongly quenched by purines (Walter and Burke, 1997). Thus, if a stretch of pyrimidines changes from single to double stranded upon folding, then attachment of a nearby pyrene may be ideal. In
Contributed by Philip C. Bevilacqua and Douglas H. Turner Current Protocols in Nucleic Acid Chemistry (2002) 11.8.1-11.8.6 Copyright © 2002 by John Wiley & Sons, Inc.
RNA Folding Pathways
11.8.1 Supplement 9
general, however, there is no way to know for certain if a fluorophore will have favorable properties. A wide variety of reactive probes are commercially available; thus, once an RNA with a reactive group has been prepared, a series of fluorophores should be tested with it. An important consideration is the length of the linker arm used to attach the fluorophore to the RNA. Short linkers provide the least opportunity for interacting at distant sites, intercalating into a helix, and perturbing the system. Moreover, changes in fluorescence for probes attached by a short linker arm are more likely to reflect changes in local environment, such as protection from solvent. Longer linker arms could be advantageous in cases where the 5′ and 3′ ends are far removed from the site of folding. Potential for intercalation of the probe into a duplex involving the strand to which it is attached can be judged from molecular modeling of the duplex, including molecular dynamics simulations and energy minimization using commercially available computer programs. Disadvantages of pendant probes include the potential for perturbing the folding process by interacting with neighboring helices in an undesirable fashion, and the inability to directly interpret changes in fluorescence in structural detail.
cial sources for synthetic RNA are now available (Dharmacon).
Purification of Labeled Oligoribonucleotides Labeled oligoribonucleotides should be purified by HPLC on a C8 column and stored at −20°C in an EDTA-containing buffer such as TE, pH 7.5 (APPENDIX 2A). Periodically, the RNA should be monitored by HPLC (UNIT 10.5), TLC (APPENDIX 3D), or polyacrylamide gel electrophoresis (UNIT 10.4) to check for degradation, and the fluorescence spectrum should be checked for any alteration. As when working with any RNA sample, latex gloves should be worn and reagents should be RNase-free. Often, large RNAs are refolded or renatured before use, in case they have been trapped in a non-native conformation during purification and storage. For example, one renaturation involves heating the RNA for 10 min at 50°C in the presence of Mg2+, and another involves heating the RNA for 3 min at 95°C in the absence of Mg2+ followed by slow cooling on the bench. Renaturations should be carried out the day of the experiment, and only the RNA to be used that day should be renatured. To prevent sticking of hydrophobic probes, microcentrifuge tubes and pipet tips should be silanized before use (see APPENDIX 2A).
Nucleotide Analogs
Use of Fluorescence Spectroscopy to Elucidate RNA Folding Pathways
Nucleotide analogs can be incorporated into oligomers by chemical synthesis. Modified oligomers can then be used directly in experiments, or covalently attached to a large RNA with DNA ligase. Useful analogs include epsilon adenosine (εA) and 2-amino purine. For example, positioning of εA on the 5′ or 3′ end of a pyrimidine substrate of a ribozyme led to ∼2-fold changes in fluorescence upon binding (Sugimoto et al., 1989; Profenno et al., 1997). The analog 2-amino purine has an ∼2-fold decrease in fluorescence upon base pairing (Menger et al., 1996). Nucleotide analogs offer the advantages that they minimize the potential for perturbing the system, and provide localized information about changes in RNA folding (Jean and Hall, 2001). Several disadvantages exist for nucleotide analogs. In general, the fluorescence sensitivity is lower than for pendant probes, and the change in fluorescence is roughly 10-fold lower (Cantor and Schimmel, 1980). This can lead to poor signal-to-noise ratios and a need for large amounts of sample. Other drawbacks include the extra steps needed for chemical synthesis of modified RNA, although reliable commer-
MEASURING CHANGES IN FLUORESCENCE The first experiment should test whether there is a change in fluorescence upon folding. A fluorometer with emission and excitation monochromators is used, and wavelengths are chosen that give maxima in excitation and emission. Opening the slits to the monochromators can improve the signal-to-noise ratio for the following experiment. A thermostatted sample holder that requires a small volume (several hundred microliters) is ideal. A series of substrates should be tested to look for those that have a large change in fluorescence upon folding. RNA folding or unfolding is initiated and the time dependence of any fluorescence change is measured. Folding or unfolding may be initiated by the addition of substrate or by the addition or removal of divalent ions or a chemical denaturant such as urea. With practice, the dead time for mixing can be as little as 5 to 10 sec. Once the system no longer shows a change in fluorescence, an emission spectrum is taken and the change in fluorescence intensity and any wavelength shift are determined. Absence of a change in fluorescence shows that
11.8.2 Supplement 9
Current Protocols in Nucleic Acid Chemistry
the probe is not good for measuring equilibrium events, but it remains possible that an intermediate state with a different fluorescence emission exists; this can be tested by stopped-flow mixing. After a probe with a change in fluorescence has been identified, the kinetics of the system should be examined. Processes that occur over nanoseconds to hours can be monitored by fluorescence (Table 11.8.1). Processes with half-lives, t1⁄2, as fast as 1 nsec can be studied by temperature-jump methods; processes with t1⁄2 from 1 msec to 1 min can be studied by stopped-flow mixing methods; and processes with t1⁄2 as fast as 1 min can be studied by manual mixing. The temperature-jump method has the advantage that the same sample can be used for many runs in order to increase the signal-to-noise ratio.
Instrumentation: Mixing Methods Experiments performed by the authors employed a KinSim stopped-flow mixing apparatus. Fluorescence emission should be collected through either a bandpass or a cut-off filter to maximize the signal that reaches the photomultiplier tube (PMT). Alternatively, stoppedflow apparatuses exist that acquire a rapid emission spectrum. This method has the potential to reveal intermediate states with altered spectra, but has the disadvantage that the signal is weaker and must be averaged over more shots, thus requiring more sample. Sample requirements are modest, however, and a few hundred microliters can allow three or more shots to be averaged. The entire apparatus is thermostatted, allowing the temperature dependence of the kinetics to be determined. Attention should be given to the possibility of photobleaching the sample. A mock sample can be exposed to the light beam to look for any changes in fluorescence. Photobleaching can often be reduced by using a less powerful lamp, by narrowing the monochromator slits, or by closing the slits between time points. If it cannot be fully eliminated, photobleaching should be corrected by subtracting a suitable blank. Table 11.8.1
Testing if the System is Perturbed Most researchers are interested in observing the folding properties of unperturbed systems. Thus, attention must be paid to the possibility of perturbing the system of study. An ideal test for perturbation is an activity assay such as bond cleavage by a ribozyme. Modified and unmodified substrate can be radiolabeled and kcat/Km, the specificity constant, can be determined under various conditions (where kcat is the observed first-order rate constant under conditions of substrate saturation and Km is the Michaelis constant). The binding of modified and unmodified substrates to complementary oligomers can be compared to test perturbation of secondary structure formation. Methods of testing include UV melting to look at thermodynamic parameters and stopped-flow mixing to look at kinetic parameters. Folding of the ribozyme can be tested by structure mapping experiments such as chemical modification and enzymatic cleavage. Overall changes in folding may be revealed by circular dichroism (CD). In particular, CD should be measured between 220 and 300 nm (where the RNA absorbs) and over the wavelength region where the fluorophore absorbs. Ellipticity in the region of fluorophore absorbance implies that the fluorophore is in a chiral environment and may be directly interacting with the RNA. Large changes in cleavage or binding rate constants, thermodynamic parameters, or RNA structure suggest that the probe is not ideal for the particular system. The choice of probe should be reevaluated. Varying the site of attachment, linker arm, and type of fluorophore should be considered.
Obtaining Kinetic Parameters Once a nonperturbing fluorophore that is sensitive to the environment has been prepared, the kinetics of folding may be monitored. A number of articles describing the details of such experiments have been published (see Johnson, 1992). This section presents a brief introduction to the types of experiments that can be
Methods for Studying Folding Kinetics by Fluorescence Detection
Timescale (t1/2)
Method
Reference
1 nsec ≥1 nsec 1 msec to 1 min >1 min
Lifetime Temperature-jump Stopped-flow mixing Manual mixing
Walter et al. (2001) Eigen and DeMaeyer (1963); Turner (1986) Johnson (1992) RNA Folding Pathways
11.8.3 Current Protocols in Nucleic Acid Chemistry
Supplement 9
Use of Fluorescence Spectroscopy to Elucidate RNA Folding Pathways
performed and the information on rate constants that can be obtained. One of the first experiments to perform is to initiate the RNA folding process and observe the time dependence of fluorescence change. For two-piece systems, such as ribozyme-substrate systems, the kinetics are greatly simplified if pseudo-first-order binding conditions are maintained. This requires having one reagent in large excess. In general, the largest change in signal will be observed if the fluorescent molecule is the limiting reagent. If, however, the fluorophore undergoes a very large change in fluorescence, as in the case of pyrene-labeled oligomers, this condition is not absolutely necessary. Two-piece systems offer the additional advantage that the rate of binding can be adjusted to some extent by varying concentrations. If, for example, the rate is too fast to be observed, concentrations can be lowered to slow the rate of binding. The signal, however, will diminish if ribozyme concentration drops below the equilibrium dissociation constant (Kd) for the interaction. Special attention should be paid to experiments in which the ribozyme is maintained in excess over the substrate. Under these conditions, the multimerization state of the large RNA should be monitored by native gel electrophoresis. For example, under certain conditions, the Tetrahymena ribozyme can dimerize, and a large pseudoknot has also been seen to dimerize (P.C. Bevilacqua and D.H. Turner, unpub. observ.; P. Babitzke, P.C. Bevilacqua, and J.E. Schaak, unpub. observ.). Such dimerization processes may lead to further changes in fluorescence. Typically, dimerization can be avoided by renaturing the RNA at higher temperatures prior to the experiment. The effect of various temperatures and salts on the oligomerization state of the RNA can be monitored by native gel electrophoresis of a radiolabeled sample. Once changes in fluorescence as a function of time are obtained, the data are fit to a firstor higher-order exponential equation to obtain observed rate constants. Because data are generated in real time and acquired on a computer, it is practical to fit the data immediately after mixing. Observed rate constants are then measured at various concentrations of the excess reagent. Plots of observed rate constants versus concentration are fit to curves dictated by a particular kinetic mechanism to obtain microscopic rate constants. By monitoring the concentration dependence of the observed rate constants, further experiments can be rationally
designed and carried out in rapid fashion. The data may also be globally fit by numerical integration to obtain the rate constants using the programs KINSIM and FITSIM (Barshop et al., 1983; Zimmerle and Frieden, 1989; Frieden, 1993). Rate constants obtained by graphical and global fitting methods should be in reasonable agreement. The interested reader is referred to an excellent review containing numerous schemes and their rate equations (Johnson, 1992). Through these approaches, it is possible to accumulate data consistent with a binding mechanism involving numerous uniand bimolecular steps. In many cases, dissociation rate constants are too small to be obtained by such experiments and require use of the pulse-chase experiment. In the pulse-chase experiment, a fluorescent substrate is in equilibrium with a ribozyme in one mixing syringe (the pulse), and excess amounts of unlabeled substrate are present in the other syringe (the chase). Upon mixing, dissociation of the labeled substrate from the ribozyme is effectively irreversible since the binding site for the labeled molecule is now occupied by an unlabeled molecule. Data are fit to a single exponential equation to obtain a rate constant for dissociation that can be either a microscopic rate constant or a mathematical combination of several rate constants. Data should be obtained at several concentrations of chase to ensure that the observed rate constant is for substrate dissociation and is not dependent on chase concentration. In cases where the y intercept is close to zero for mixing experiments, pulse-chase experiments can allow determination of all the individual rate constants. In cases where the y intercept is well determined by mixing experiments, pulse-chase experiments confirm the value of the dissociation rate constant and allow its direct determination in a single experiment. For example, in the case of pyrene substrate binding to the Tetrahymena ribozyme, the rate constant was the rate of undocking of the P1 duplex—a tertiary RNA unfolding event (Bevilacqua et al., 1992). By varying parameters such as temperature, salt, and pH during the pulse-chase experiment, insight into the thermodynamic and structural details of the unfolding event may be obtained. An interesting variation of the pulse-chase experiment is the so-called reversed pulsechase experiment, in which an unlabeled substrate is forced to dissociate from the ribozyme by addition of excess labeled substrate. Because the labeled substrate is in excess, it must have a large change in fluorescence to be ob-
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Current Protocols in Nucleic Acid Chemistry
servable. The rate of dissociation of unlabeled substrate is then directly visualized by the increase in fluorescence upon binding of labeled substrate. Rates may be compared for the pulsechase and reversed pulse-chase experiments to look for any effects of fluorophore upon binding (Bevilacqua et al., 1993). Moreover, such experiments offer the advantage that dissociation behavior of a variety of chemically modified unlabeled substrates can be measured with a single fluorescent substrate.
CONCLUSION Fluorescence spectroscopy is a powerful method for studying the folding of large RNAs and offers a number of unique advantages. A variety of RNA folding processes may be monitored by fluorescence detection, including secondary and tertiary structure formation. For example, fluorescence detection has been used to look at binding of oligomer substrate to a ribozyme, and the mechanism was determined to involve base pairing of the substrate followed by docking of the helix into the core of the ribozyme (Bevilacqua et al., 1992). In addition, rates for bond cleavage have been determined by fluorescence detection. Data acquisition is in real time and processes that occur over nanoseconds to hours may be monitored. Fluorescence spectroscopy is a sensitive method; depending on the quantum yield of fluorophore, concentrations down to the nanomolar regime can be used. True equilibrium constants can be measured by fluorescence methods—an advantage over nonequilibrium methods such as gel mobility shift assays. Determination of equilibrium constants is an essential part of determining RNA folding mechanisms, and accurate thermodynamic parameters have allowed insight into the details of RNA tertiary folding. Limitations of fluorescence spectroscopy include the low level of detail of structural information that may be inferred. Other methods described in this series, such as hydroxyl radical cleavage and chemical modification, provide considerably more insight into detailed structural changes in the RNA. Perhaps the most difficult and time-consuming part of fluorescence experiments on RNA is identifying a fluorescent probe with favorable properties. An ideal probe should have a large change in fluorescence and minimal perturbation of RNA properties. Once a suitable fluorescent RNA is prepared, however, data may be obtained quickly for rapid and slow kinetic processes. The wide variety and increas-
ing number of commercially available fluorophores makes it feasible to rapidly prepare a large series of end-labeled RNAs for examination.
LITERATURE CITED Barshop, B.A., Wrenn, R.F., and Frieden, C. 1983. Analysis of numerical methods for computer simulation of kinetic processes: Development of KINSIM: A flexible, portable system. Anal. Biochem. 130:134-145. Bevilacqua, P.C., Kierzek, R., Johnson, K.A., and Turner, D.H. 1992. Dynamics of ribozyme binding of substrate revealed by fluorescence-detected stopped-flow methods. Science 258:13551358. Bevilacqua, P.C., Johnson, K.A., and Turner, D.H. 1993. Cooperative and anticooperative binding to a ribozyme. Proc. Natl. Acad. Sci. U.S.A. 90:8357-8361. Cantor, C.R. and Schimmel, P.R. 1980. Biophysical Chemistry Part II: Techniques for the Study of Biological Structure and Function. pp. 433-465. W.H. Freeman and Company, New York. Clegg, R.M., Murchie, A.I., Zechel, A., Carlberg, C., Diekmann, S., and Lilley, D.M. 1992. Fluorescence resonance energy transfer analysis of the structure of the four-way DNA junction. Biochemistry 31:4846-4856. Eigen, M. and DeMaeyer, L. 1963. Relaxation methods. In Techniques of Organic Chemistry, Vol. VIII, Part 2 (S.L. Friess, E.S. Lewis, and A. Weissberger, eds.), pp. 895-1054. John Wiley & Sons, New York. Frieden, C. 1993. Numerical integration of rate equations by computer. Trends Biochem. Sci. 18:58-60. Jean, J.M. and Hall, K.B. 2001. 2-Aminopurine fluorescence quenching and lifetimes: Role of base stacking. Proc. Natl. Acad. Sci. U.S.A. 98:37-41. Johnson, K.A. 1992. Transient state kinetic analysis of enzyme reaction pathways. In The Enzymes, Vol. 20 (D. Sigman, ed.), pp. 1-61. Academic Press, New York. Kierzek, R., Li, Y., Turner, D.H., and Bevilacqua, P.C. 1993. 5′-Amino pyrene provides a sensitive, non-perturbing fluorescent probe of RNA secondary and tertiary structure formation. J. Am. Chem. Soc. 115:4985-4992. Manoharan, M., Tivel, K.L., Zhao, M., Nafisi, K., and Netzel, T.L. 1995. Base-sequence dependence of emission lifetimes for DNA oligomers and duplexes covalently labeled with pyrene. Relative electron-transfer quenching efficiencies of A-nucleoside, G-nucleoside, C-nucleoside, and T-nucleoside toward pyrene. J. Phys. Chem. 99:17461-17472. Menger, M., Tuschl, T., Eckstein, F., and Pörschke, D. 1996. Mg(2+)-dependent conformational changes in the hammerhead ribozyme. Biochemistry 35:14710-14716.
RNA Folding Pathways
11.8.5 Current Protocols in Nucleic Acid Chemistry
Supplement 9
Netzel, T.L., Zhao, M., Nafisi, K., Headrick, J., Sigman, M.S., and Eatson, B.E. 1995. Photophysics of 2′-deoxyuridine (dU) nucleosides covalently substituted with either 1-pyrenyl or 1pyrenoyl-observation of pyrene-to-nucleoside charge-transfer emission in 5-(1-pyrenyl)-dU. J. Am. Chem. Soc. 117:9119-9128. Profenno, L.A., Kierzek, R., Testa, S.M., and Turner, D.H. 1997. Guanosine binds to the Tetrahymena ribozyme in more than one step, and its 2′-OH and the nonbridging pro-Sp phosphoryl oxygen at the cleavage site are required for productive docking. Biochemistry 36:12477-12485. Reines, S.A. and Cantor, C.R. 1974. New fluorescent hydrazide reagents for the oxidized 3′-terminus of RNA. Nucl. Acids Res. 1:767-786. Silverman, S.K. and Cech, T.R. 1999. RNA tertiary folding monitored by fluorescence of covalently attached pyrene. Biochemistry 38:14224-14237. Sugimoto, N., Sasaki, M., Kierzek, R., and Turner, D.H. 1989. Binding of a fluorescent oligonucleotide to a circularized intervening sequence f or Tetrahymena thermophila. Chem. Lett. (1989):2223-2226. Turner, D.H. 1986. Temperature-jump methods. In Investigations of Rates and Mechanisms of Reactions, Vol. 6, Part 2 (C. F. Bernasconi, ed.), pp. 141-189. John Wiley & Sons, New York. Turner, D.H., Li, Y., Fountain, M., Profenno, L., and Bevilacqua, P.C. 1996. Dynamics of a group I ribozyme detected by spectroscopic methods. In Nucleic Acids and Molecular Biology (F. Eckstein and D.M.J. Lilley, eds.), pp. 19-32. Springer-Verlag, Berlin. Walter, N.G. and Burke, J.M. 1997. Real-time monitoring of hairpin ribozyme kinetics through basespecific quenching of fluorescein-labeled substrates. RNA 3:392-404. Walter, N.G., Hampel, K.J., Brown, K.M., and Burke, J.M. 1998. Tertiary structure formation in the hairpin ribozyme monitored by fluorescence resonance energy transfer. EMBO J. 17:23782391.
Walter, N.G., Burke, J.M., and Millar, D.P. 1999. Stability of hairpin ribozyme tertiary structure is governed by the interdomain junction. Nat. Struct. Biol. 6:544-549. Walter, N.G., Chan, P.A., Hampel, K.J., Millar, D.P., and Burke, J.M. 2001. A base change in the catalytic core of the hairpin ribozyme perturbs function but not domain docking. Biochemistry 40:2580-2587. Zimmerle, C.T. and Frieden, C. 1989. Analysis of progress curves by simulations generated by numerical integration. Biochem. J. 258:381-387.
INTERNET RESOURCES http://chemgenes.com ChemGenes site with catalog of precursors for synthesis of nucleic acids with fluorescent bases. http://www.dharmacon.com Dharmacon Research site with catalog offering synthesis of labeled nucleic acids. http://www.glenres.com Glen Research site with catalog of precursors for synthesis of labeled nucleic acids. http://www.probes.com Molecular Probes site with catalog and technical information for fluorophores. http://www.trilinkbiotech.com TriLink BioTechnologies site with catalog of fluorescent nucleoside triphosphates.
Contributed by Philip C. Bevilacqua The Pennsylvania State University University Park, Pennsylvania Douglas H. Turner University of Rochester Rochester, New York
Use of Fluorescence Spectroscopy to Elucidate RNA Folding Pathways
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Current Protocols in Nucleic Acid Chemistry
Use of Chemical Modification To Elucidate RNA Folding Pathways Chemical modification reagents that react at the hydrogen-bonding groups of nucleotides, su ch as β-ethoxy-α-ketobutyraldehyde (kethoxal), dimethyl sulfate (DMS), and 1-cyclohexyl-3-(2-morpholinoethyl)carbodiimide metho-p-toluenesulfonate (CMCT), react at nucleotides that are either single stranded or are at the end of helices (see UNIT 6.1; Inuoe and Cech, 1985; Moazed et al., 1986; Ehresmann et al., 1987). However, not all nucleotides that are not in Watson-Crick pairs are modified, because these reagents act based on the nucleotide’s accessibility to solvent, which is sensitive to both secondary and tertiary structure (Banerjee et al., 1993; Mathews et al., 1997). Under pseudo-first-order reaction conditions (i.e., reagent concentration is much greater than RNA concentration), a linear relationship exists between rate of nucleotide modification and the fraction of accessible nucleotides; therefore, chemical modification can be used to measure the kinetics of RNA folding (Banerjee and Turner, 1995). This unit discusses the use of chemical modification reagents to measure RNA folding kinetics.
UNIT 11.9
CHOOSING A TRANSITION The first step in understanding the folding pathways of an RNA molecule is to identify the transitions of interest. A good starting point is to follow UV absorbance to identify transitions. RNA structure influences the extinction coefficient for the base transitions at 260 and 280 nm (Tinoco, 1959). For example, formation of secondary structure decreases the extinction coefficient. This phenomenon is often referred to as hypochromicity. Circular dichorism (CD) is an optical measurement that is even more sensitive to structural changes (Pan and Sosnick, 1997; UNIT 11.5). CD spectroscopy measures the difference in extinction coefficient for left and right circularly polarized light. The extinction coefficient and CD spectrum can be measured for the RNA as a function of the denaturant that will be used to induce the transition. For example, an optical melting experiment is used if structural transitions are to be induced by changes in the temperature of the sample. Alternatively, a titration curve of extinction coefficient and/or CD as a function of Mg2+ concentration could be determined if an adjustment in Mg2+ concentration will be used to affect the transition of interest. The structural transitions are revealed by changes of extinc-
Normalized absorbance or derivative
1.0 0.9 0.8 0.7 0.6 0.5 0.4 0.3 0.2 0.1 0 0
10
20
30
40
50
60
70
80
90
Temperature (°C)
Figure 11.9.1 Optical melting profile of the T. thermophila group I ribozyme. The solid line is the normalized absorbance as a function of temperature and the dotted line is the normalized derivative of absorbance (dA/dT). Structural transitions are centered around the derivative maxima.
RNA Folding Pathways
Contributed by David H. Mathews and Douglas H. Turner
11.9.1
Current Protocols in Nucleic Acid Chemistry (2002) 11.9.1-11.9.4 Copyright © 2002 by John Wiley & Sons, Inc.
Supplement 9
tion coefficient and/or CD. Structured RNAs typically have at least two transitions in optical melting curves (Crothers et al., 1974; Hilbers et al., 1976; Banerjee et al., 1993; Jaeger et al., 1993; Mathews et al., 1997). For example, Figure 11.9.1 shows the melting curve and its first derivative for the Tetrahymena thermophila group I ribozyme (Banerjee et al., 1993). The derivative reveals at least two transitions, one centered at ∼52°C and the other at 64°C.
CHARACTERIZING THE TRANSITION Once the transitions are identified, the structure should be mapped at equilibrium with chemical reagents at conditions both above and below the transitions (see UNIT 6.1). Consider the map of nucleotide accessibility to kethoxal in
A C A G A CA G C U A G C 200 G C G G U 120 P5a C A UU U G P5 C G C G U A C G C G G G U AA U A U A A A AU A A A 180 G C C G C G P5c G U A U A G AC 260 C G A AA G G G C C P4 C G CU A G UU C C G A U A 140 G U G A U U UA P6 G 160 G C A C G A U C A G C P5b U A A C U G A G U G 220 G U C G U A A G C G P6a AA C G U U A A A G U U A C U A U A U P6b C G A A G C 240 A U UCU
Use of Chemical Modification to Elucidate RNA Folding Pathways
the T. thermophila group I ribozyme shown in Figure 11.9.2 (based on data from Banerjee et al., 1993). The structural transition centered at 52°C involves solvent exposure of many nucleotides, and is consistent with disruption of tertiary structure but retention of most secondary structure. For example, nucleotides G73, G264, and G382 are strongly modified at 65°C, but not extensively modified at 50°C. Before determining folding kinetics, it is also advisable to test the effects of proposed perturbations on activity. For example, if folding will be initiated by lowering the temperature, it is useful to know if this perturbation results in fully functional RNA.
U G CA A A G C U A P9.1a G C A U A G G G A 340 U G A G C 360 P9.1G C C G G C A U P9 380 U G P9.2 C GG UGGG A G A A C U A A U UU G U A U G C G A A A UUCCUC U U G A U U A GU U A U A U G AA U 320 U 400 G G A G U U C 3′ C A G G C U G A U
CA G A C P7 U A
A U
5′ G A G G G
A UU G G C A U A A U 80 A C C G C G A U A U P2.1 A U U A U AA U A 60 C G U G G C A A A A A A A U G C U G U A P2 A U U G C G A U G C 40 G C A C A C UG
A A A U A C G U U G G C G A 100C C G P3 A U A G C U 300 A G C G 280 U G C G U A P8 U A C G U A U U A UG
50 °C 65 °C weak medium strong
Figure 11.9.2 The map of kethoxal-accessible nucleotides at 50° (triangles) and 65°C (circles). The modification intensity is quantified as weak, medium, and strong.
11.9.2 Supplement 9
Current Protocols in Nucleic Acid Chemistry
DETERMINING FOLDING KINETICS To determine the kinetics of changing accessibility, equilibrate the RNA in a state of interest and then apply a structural perturbation along with modification reagent. As the structure reequilibrates, remove aliquots of modified RNA at a series of time points and quench the modification reaction. The time points are chosen to cover the time scale of folding under consideration. The extent of reaction as a function of time is later quantified by reverse transcription. When interpreting the kinetics, the reactions of interest are: k
k
F R → F ← U M
where each nucleotide is folded (F), unfolded (U), or modified (M). Only unfolded nucleotides are accessible to modification. Two rate constants apply: kF, the rate of folding, and kR, the rate of reaction with modification reagent. This scheme assumes that the folding of the nucleotide is first order and that the concentration of the modification reagent is much larger than the concentration of nucleotides, so that modification is pseudo first order. The fraction (ft) of modified nucleotides at time t after exposure to the reagent will be:
ft = [
kR kR + kF
][1 − e
− kF t
+ k R te
− kF t
] + f0
where f0 is the apparent background modification that is a result of spontaneous strand cleavage and inefficiencies of reverse transcription (Banerjee and Turner, 1995). For nucleotides that do not become unreactive with time, kF = 0 and this equation reduces to:
ft = kR t + f0 The concentration of reagent should be chosen such that the number of modifications per RNA molecule is less than one, so that kRt << 1. Then, for nucleotides that become protected with time, the fraction modified will be:
ft = [
kR kR + kF
][1 − e
−kF t
] + f0
The fraction of modified nucleotide as a function of time can be fit to these equations to determine the rate constants. For example, Banerjee and Turner (1995) studied the tertiary folding of the T. thermophila group I ribozyme from equilibrium at 60°C to equilibrium at 15°C. The RNA was preequilibrated in 1 mM Mg2+ buffer at 60°C and then transferred to a larger volume of 10 mM Mg2+ buffer containing kethoxal at 15°C. Modification was quantified at 5 sec and 10, 20, 40, 60, 120, 180, 240, 360, and 480 min to determine folding rate constants from 0.048 to 0.008 min−1. The 5 sec time point was used to determine f0. Modification was visualized by reverse transcription of the RNA with labeled primer and the extent of modification was quantified with a phosphorimager. On the basis of the tertiary folding time scales, the guanines shown in Figure 11.9.2 were placed into four classes: (1) guanines already protected or that fold with half-lives of less than 10 min, (2) guanines that fold with time scales of tens of minutes, (3) guanines that continue to fold after an hour, and (4) guanines that never fold (Banerjee and Turner, 1995). Guanines that fold immediately include G44, G73, G341, G358, G360, G368, G382, and G384. These are clustered in loops at the 5′ and 3′ ends of the molecule, suggesting that the fast folding may involve interactions between the ends of the molecule. Guanines that do not fold demonstrate a linear increase in the fraction modified as a function of time (third equation). Such guanines are typically single stranded or at the ends of helices.
CONCLUSION Chemical modification is sensitive to solvent accessibility and can therefore be used to determine folding pathways. Measurements at equilibrium can reveal conditions inducing transitions. Measurements after a suitable perturbation can be used to determine the rates of these transitions at nucleotide resolution. Coupled with other methods, such as stopped-flow mixing (Bevilacqua et al., 1992), hydroxyl radical cleavage (Sclavi et al., 1998; Chaulk and MacMillan, 2000; Kent et al., 2000; UNIT 11.6), and oligonucleotide hybridization (Zarrinkar and Williamson, 1994), chemical modification can help provide a picture of the steps involved in folding a full-length RNA. It is quite possible, however, that the folding pathway for a full-length RNA will differ from that of the same RNA during transcription.
RNA Folding Pathways
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LITERATURE CITED Banerjee, A.R. and Turner, D.H. 1995. The time dependence of chemical modification reveals slow steps in the folding of a group I ribozyme. Biochemistry 34:6504-6512. Banerjee, A.R., Jaeger, J.A., and Turner, D.H. 1993. Thermal unfolding of a group I ribozyme: The low temperature transition is primarily disruption of tertiary structure. Biochemistry 32:153163. Bevilacqua, P.C., Kierzek, R., Johnson, K.A., and Turner, D.H. 1992. Dynamics of ribozyme binding of substrate revealed by fluorescence-detected stopped-flow methods. Science 258:13551358. Chaulk, S.G. and MacMillan, A.M. 2000. Characterization of the Tetrahymena ribozyme folding pathway using the kinetic footprinting reagent peroxynitrous acid. Biochemistry 39:2-8. Crothers, D.M., Cole, P.E., Hilbers, C.W., and Schulman, R.G. 1974. The molecular mechanism of thermal unfolding of Escherichia coli formylmethionine transfer RNA. J. Mol. Biol. 87:63-88. Ehresmann, C., Baudin, F., Mougel, M., Romby, P., Ebel, J., and Ehresmann, B. 1987. Probing the structure of RNAs in solution. Nucl. Acids Res. 15:9109-9128. Hilbers, C.W., Robillard, G.T., Shulman, R.G., Blake, R.D., Webb, P.K., Fresco, R., and Riesner, D. 1976. Thermal unfolding of yeast glycine transfer RNA. Biochemistry 15:1874-1882. Inuoe, T. and Cech, T.R. 1985. Secondary structure of the circular form of the Tetrahymena rRNA intervening sequence: A technique for RNA structure analysis using chemical probes and reverse transcriptase. Proc. Natl. Acad. Sci. U.S.A. 82:648-652. Jaeger, L., Westhof, E., and Michel, F. 1993. Monitoring of cooperative unfolding of the sunY group I intron of bacteriophage T4. J. Mol. Biol. 234:331-346.
Kent, O., Chaulk, S.G., and MacMillan, A.M. 2000. Kinetic analysis of the M1 RNA folding pathway. J. Mol. Biol. 304:699-705. Mathews, D.H., Banerjee, A.R., Luan, D.D., Eickbush, T.H., and Turner, D.H. 1997. Secondary structure model of the RNA recognized by the reverse transcriptase from the R2 retrotransposable element. RNA 3:1-16. Moazed, D., Stern, S., and Noller, H.F. 1986. Rapid chemical probing of conformation in 16S ribosomal RNA and 30S ribosomal subunits using primer extension. J. Mol. Biol. 187:399-416. Pan, T. and Sosnick, T.R. 1997. Intermediates and kinetic traps in the folding of a large ribozyme revealed by circular dichroism and UV absorbance spectroscopies and catalytic activity. Nat. Struct. Biol. 4:931-938. Sclavi, B., Sullivan, M., Chance, M.R., Brenowitz, M., and Woodson, S.A. 1998. RNA folding at millisecond intervals by synchrotron hydroxyl radical footprinting. Science 279:1940-1943. Tinoco, I. Jr. 1959. Hypochromism in polynucleotides. J. Am. Chem. Soc. 82:4785-4790. Zarrinkar, P.P. and Williamson, J.R. 1994. Kinetic intermediates in RNA folding. Science 265:918924.
KEY REFERENCES Banerjee and Turner, 1995. See above. Presents the kinetics of folding of the T. thermophila group I ribozyme as determined by chemical modification with kethoxal. Ehresmann et al., 1987. See above. Reviews the activity and use of many popular chemical modification reagents.
Contributed by David H. Mathews and Douglas H. Turner University of Rochester Rochester, New York
Use of Chemical Modification to Elucidate RNA Folding Pathways
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Probing RNA Structural Dynamics and Function by Fluorescence Resonance Energy Transfer (FRET)
UNIT 11.10
RNA is a ubiquitous biopolymer with complex functions in the maintenance, transfer, processing, and regulation of genetic information. Its secondary structure provides a comparably stable scaffold onto which long-range tertiary interactions are built. RNA folding pathways describe the possible trajectories by which an RNA molecule may acquire a functional tertiary structure. Often, biological function of RNA is mediated by cyclic switching between two or more (meta-)stable arrangements of tertiary structure. Fluorophore labeling of RNA offers a unique view into these folding and conformational switching events, since a fluorescence signal is sensitive to its molecular environment and can be continuously monitored in real time to produce kinetic rate information. Many of these general aspects are discussed in UNIT 11.8. The current chapter complements UNIT 11.8 by focusing on the practical implications of using fluorescence resonance energy transfer (FRET) to probe RNA structural dynamics and function. FRET is a particularly powerful fluorescence technique since, in addition to kinetic data, it provides insights into the structural basis of a conformational rearrangement. The current unit provides protocols that describe how to postsynthetically label RNA for FRET (see Basic Protocol 1) and how to acquire and analyze FRET data (see Basic Protocol 2). Support Protocols describe methods for deprotecting synthetic RNA (see Support Protocols 1 and 2) and for purifying RNA by gel electrophoresis and HPLC (see Support Protocols 3 and 4, respectively). Considerations for selecting appropriate RNA, fluorophores, and labeling strategies are discussed in detail in the Commentary (see Critical Parameters). NOTE: As for any experiments with RNA, care must be taken to avoid introducing ribonucleases (RNases) into the samples. Since most techniques described here involve commercially available synthetic RNA, the major source of RNases is postsynthetic contamination. This can be avoided by wearing gloves to avoid skin contact when handling samples and solutions; by using nuclease-free sterilized pipet tips, sample tubes, and other disposable plasticware (stored in clean, autoclaved, dust-free containers); by preparing all solutions from the highest purity (e.g., molecular-biology grade) components in doubly deionized water (18 MΩ conductivity); and by sterile filtering (0.22 µm) or autoclaving all solutions. When these suggestions are followed, inactivation of RNases using DEPC is not necessary and not particularly recommended because contaminating decay products may lead to fungal growth. POSTSYNTHETIC LABELING OF AMINO- OR THIOL-MODIFIED RNA To perform FRET experiments, the RNA sample must be labeled with both a donor and an acceptor fluorophore. This can be accomplished by a variety of strategies that are discussed in detail in the Commentary (see Critical Parameters). In short, fluorophores can be added either during oligoribonucleotide synthesis or by postsynthetic conjugation to a modified oligoribonucleotide. This protocol describes a method for postsynthetic labeling of a modified RNA. RNA modified with primary or secondary amino and thiol functionalities can be conveniently labeled under mild conditions with succinimidyl ester and maleimide derivatives of a fluorophore, respectively, available from companies such as Molecular Probes, Contributed by Nils G. Walter Current Protocols in Nucleic Acid Chemistry (2002) 11.10.1-11.10.23 Copyright © 2002 by John Wiley & Sons, Inc.
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Sigma-Aldrich, or Amersham Pharmacia Biotech. Since RNA purified by reversed-phase HPLC contains trace amounts of triethylamine that hydrolyze succinimidyl esters and, to a lesser extent, maleimides in a catalytic fashion, the labeling protocol consists of two steps: (1) chloroform extraction and ethanol precipitation to remove triethylamine and (2) coupling of the fluorophore derivative to the amino- or thiol-modified RNA. Excess fluorophore is then removed by C8-reversed phase HPLC purification (see Support Protocol 4). This method can be used for single or double labeling of an RNA. Only one fluorophore need be added, for instance, if the other fluorophore was added during synthesis or if the donor and acceptor are conjugated to opposite strands of the RNA (see Critical Parameters). If the RNA already contains a fluorophore, it should be protected from light at all times to prevent photobleaching. If both donor and acceptor are to be added postsynthetically to the same strand, the RNA must contain both an amino and a thiol modification. In this case, the thiol functionality should be reacted first, since maleimides provide for higher selectivity than succinimidyl esters. Materials Gel- and HPLC-purified RNA sample with amino or thiol functionality (see Support Protocol 4) Chloroform, buffered (see recipe) 3 M sodium acetate, pH 5.2 (APPENDIX 2A) 100% and 80% (v/v) ethanol Succinimidyl ester (for amino-modified RNA) or maleimide derivative (for thiol-modified RNA) of fluorophore of choice (e.g., Molecular Probes, Sigma-Aldrich, Amersham Pharmacia Biotech; for selection of fluorophores, see Critical Parameters) Dimethylsulfoxide (DMSO), anhydrous (e.g., Fisher) 100 mM sodium tetraborate, pH 8.5 (for amino-modified RNA, see recipe) or 100 mM HEPES-KOH, pH 7.0 (for thiol-modified RNA, see recipe) 100 mM ATP or GTP (APPENDIX 2A; recipe for dNTPs) Speedvac evaporator (e.g., Savant) Aluminum foil Tube shaker (e.g., Fisher) Chloroform extract and precipitate RNA 1. Bring the RNA sample to be labeled up to 100 µL with water. This volume is large enough to handle easily and small enough to ensure a good precipitation yield. Up to 100 ìg RNA can be labeled in one reaction using this protocol. It is wise to keep 50% of the original material for a second labeling in case of unexpected loss.
2. Extract RNA solution with 1 vol buffered chloroform, microcentrifuge briefly at maximum speed to separate phases, and transfer the top (aqueous) phase to a fresh tube. 3. Add 1⁄10 vol (10 µL) 3 M sodium acetate, pH 5.2, and 2.5 vol (250 µL) 100% ethanol, vortex, and precipitate at −70°C for 1 hr.
Probing RNA Structural Dynamics and Function by FRET
4. Microcentrifuge 30 min at maximum speed, 4°C, to collect RNA. Decant supernatant, wash with 80% ethanol, decant supernatant, and dry RNA in a Speedvac evaporator. Cover the evaporator with aluminum foil if the RNA already contains a fluorophore.
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Label RNA 5. Dissolve 200 µg fluorophore succinimidyl ester or maleimide in 14 µL DMSO. If both an amino- and a thiol-modification will be used to attach two fluorophores, perform the reaction at the thiol functionality first. In many cases it is easier to dissolve a whole vial of the fluorophore derivative at once than to weigh out a small amount of fluorophore. In this case, dispense fluorophore solution in 14-ìL aliquots and store up to 6 months at –20°C wrapped in aluminum foil. For long-term storage, dissolve fluorophore derivative in methanol or acetonitrile, dispense in 200-ìg aliquots per tube, dry in a Speedvac evaporator, and store up to 2 years at –20°C wrapped in aluminum foil.
6a. For amino-modified RNA: Dissolve RNA pellet from step 4 in 11 µL water, add 75 µL of 100 mM sodium tetraborate, pH 8.5, and transfer to the tube containing the fluorophore succinimidyl ester stock solution from step 5. 6b. For thiol-modified RNA: Dissolve RNA pellet from step 4 in 11 µL water, add 75 µL of 100 mM HEPES-KOH, pH 7.0, and transfer to the tube containing the fluorophore maleimide stock solution from step 5. 7. Vortex tube, wrap in aluminum foil, and tumble on a tube shaker overnight (16 to 20 hr) at room temperature. The exact incubation time and temperature may have to be optimized for a given reaction. For example, sterically hindered secondary amines (such as a 2′-amino modification on a ribose) tend to give higher labeling yields when incubated overnight at 4°C.
8. Add 1⁄10 volume (10 µL) 3 M sodium acetate, pH 5.2, 1⁄40 volume (2.5 µL) 100 mM ATP or GTP as carrier, and 2.5 volumes (250 µL) ethanol, vortex, and precipitate at –70°C for 1 hr. 9. Microcentrifuge 30 min at maximum speed, 4°C, to collect RNA. Decant supernatant, wash twice with 80% ethanol, decant supernatant, and dry RNA in a Speedvac evaporator covered with aluminum foil. 10. Resuspend dried RNA in 90 µL water and store in the dark at –20°C (stable at least 2 years). Before use for FRET (see Basic Protocol 2), remove excess fluorophore by C8 reversed-phase HPLC (see Support Protocol 4). 11. Optional: For attachment of a second fluorophore at the amino functionality, repeat steps 5 to 10. MILD DEPROTECTION OF STANDARD RNA OLIGONUCLEOTIDES WITH NH4OH/ETHANOL AND TRIETHYLAMINE TRIHYDROFLUORIDE This protocol describes a mild deprotection scheme for RNA oligonucleotides synthesized using standard β-cyanoethyl phosphoramidites. Synthesis is as described in UNIT 3.5 and APPENDIX 3C; phosphoramidites can be purchased from companies such as Glen Research, ChemGenes, Amersham Pharmacia Biotech, BD Biosciences (Clontech), CPG, Cruachem, Dalton Chemical Laboratories, or PE Applied Biosystems. This scheme is compatible with fluorophores (particularly fluorescein and the cyanine dyes) or linker modifications incorporated during synthesis. It uses a 3:1 mixture of concentrated ammonium hydroxide and ethanol to cleave the RNA from the solid support, perform a β-elimination of the cyanoethyl phosphodiester-protecting group, and remove the exocyclic N-acyl base-protecting groups. A second deprotection with triethylamine trihydrofluoride removes the tert-butyldimethylsilyl group on the 2′-hydroxyl functionality (also see UNIT 3.6). Many commercially available RNAs have already been cleaved from the support and deprotected at the base and phosphodiester moiety. If this is the case, the desilylation reaction is all that needs to be performed. If a fluorophore was already
SUPPORT PROTOCOL 1
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attached during synthesis, the RNA should be kept protected from light as much as possible to avoid photobleaching. Materials RNA oligonucleotide attached to solid synthesis support (1-µmol scale), made using standard β-cyanoethyl phosphoramidites (see above for suggested suppliers and UNIT 3.5 for synthesis protocols) 29% (v/v) ammonium hydroxide (e.g., Fisher) 100% and 80% (v/v) ethanol (e.g., Fisher) Triethylamine trihydrofluoride (Aldrich or Acros) N,N-Dimethylformamide (e.g., Fisher; optional) 1-Butanol (e.g., Fisher) 1.7-mL screw-top tube (e.g., Eppendorf Safe-Twist) Parafilm (e.g., Fisher) Aluminum foil Heating block (e.g., Fisher) 14-mL Falcon centrifuge tube (e.g., Fisher) Speedvac evaporator (e.g., Savant) Tube shaker (e.g., Fisher) Cleave from support and remove cyanoethyl and N-acyl protecting groups 1. Transfer the dried solid support beads with the attached RNA oligonucleotide from the synthesis cartridge to a 1.7-mL Safe-Twist screw-top tube. Most commercial suppliers of RNA provide material that already has undergone steps 1 through 6. In this case, proceed directly to desilylation in step 7.
2. Add 750 µL of 29% ammonium hydroxide and 250 µL 100% ethanol to the tube, screw the cap on tightly, wrap the top with Parafilm, and place in a heating block for 4 hr at 65°C. If a fluorophore is already attached to the RNA, cover with aluminum foil to protect from light. For RNA containing cyanine dyes, an even milder incubation (20 hr at 25oC) is preferable. Due to its high vapor pressure, the concentrated ammonium hydroxide solution is easiest to pipet when stored in the freezer at –20°C until use. This precaution also minimizes a gradual decrease over time in ammonia concentration due to degassing.
3. Remove the tube from the heating block, place it on ice, and wait 10 min for it to cool down. This procedure avoids loss of contents when opening the tube.
4. Microcentrifuge the tube 30 sec at maximum speed to compact the solid support beads, then pipet the supernatant into a 14-mL Falcon tube. 5. Add 1 mL water to the beads, vortex thoroughly, microcentrifuge for 30 sec, and add the supernatant to the Falcon tube. Repeat this wash a second time and again add the supernatant to the Falcon tube. 6. Evaporate the combined supernatants in a Speedvac evaporator. Cover the evaporator with aluminum foil if the RNA contains a fluorophore. To avoid subsequent solubility problems, be careful not to overdry the RNA. Probing RNA Structural Dynamics and Function by FRET
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Remove tert-butyldimethylsilyl protecting group 7. Add 800 µL triethylamine trihydrofluoride to the Falcon tube, wrap the top with Parafilm, and tumble on a tube shaker overnight (16 to 20 hr) at room temperature. Wrap in aluminum foil if the RNA contains a fluorophore. To increase the solubility of long RNA (>30 nt), 25% dimethylformamide may be added to the reaction.
8. Quench the desilylation reaction by adding 160 µL water to the Falcon tube. 9. Add 8 mL of 1-butanol and chill the solution at –20°C for 45 min. 10. Microcentrifuge 5 min at 3000 rpm, 4°C, and gently decant the 1-butanol from the precipitated RNA. 11. Wash RNA pellet with 80% ethanol, decant, and repeat wash. Evaporate remaining liquid from the Falcon tube in a Speedvac evaporator. Cover the evaporator with aluminum foil if the RNA contains a fluorophore. Completely removing the 1-butanol by ethanol washes and evaporation is important to improve the separation during subsequent gel purification (see Support Protocol 3). If necessary, the dried RNA can be stored at −20°C until gel purification (stable at least 1 month). RNA recovery at this stage should be at least 50% of the total synthesis scale.
MILD DEPROTECTION OF 2′-ACE-PROTECTED RNA OLIGONUCLEOTIDES WITH ACETIC ACID
SUPPORT PROTOCOL 2
This protocol describes a specific deprotection scheme required for RNA oligonucleotides purchased from Dharmacon Research. Dharmacon uses a very different 5′-silyl-2′-orthoester protection chemistry, and typically supplies RNA after the support cleavage and base deprotection step. Mildly acidic conditions are then used to remove the 2′-orthoester protecting groups. Dharmacon provides a limited but increasing number of modifications (including 5′ fluorescein and cyanine dyes) that are all compatible with the deprotection protocol outlined below. Again, if a fluorophore was already attached during the synthesis, the RNA should be kept protected from light as much as possible to avoid photobleaching. Additional Materials (see Support Protocol 1) RNA oligonucleotide made using 5′-silyl-2′-orthoester protection chemistry, already cleaved from solid support and base deprotected (Dharmacon Research) 100 mM acetic acid/TEMED, pH 3.8 (see recipe) 1. Briefly microcentrifuge the tube containing the RNA oligonucleotide to ensure that the RNA pellet is at the bottom. 2. Add 400 µL of 100 mM acetic acid/TEMED, pH 3.8, per 0.1 µmol synthesis material. Dissolve the pellet by pipetting up and down and vortexing. Centrifuge the solution to the bottom of the tube. 3. Wrap tube top with Parafilm and place the tube in a heating block for 30 min at 60°C. If a fluorophore is already attached to the RNA, cover tube with aluminum foil to protect from light. 4. Remove the tube from the heating block, place it on ice, and wait 10 min for it to cool down. This procedure avoids loss of contents when opening the tube. RNA Folding Pathways
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5. Evaporate buffer in a Speedvac evaporator. Cover the evaporator with aluminum foil if the RNA contains a fluorophore. If necessary, the dried RNA can be stored at −20°C until gel purification (stable at least 1 month). RNA recovery at this stage should be at least 50% of the total synthesis scale. SUPPORT PROTOCOL 3
GEL PURIFICATION OF RNA OLIGONUCLEOTIDES Polyacrylamide gel electrophoresis (PAGE) is the most effective way of purifying a full-length synthetic RNA oligonucleotide from shorter synthesis byproducts. Since PAGE in general is described in UNIT 10.4 and APPENDIX 3B, the following procedure focuses on a streamlined standard purification of a synthetic RNA oligonucleotide (up to 80 nt in length). The steps in preparative PAGE are: (1) preparation of the gel and gel apparatus, (2) electrophoretic separation, and (3) detection of the RNA by UV shadowing and elution from the gel. CAUTION: Acrylamide is a known neurotoxin and should be handled with care. Wear safety goggles and gloves when handling solutions or a solidified gel. Work in a fume hood when weighing powdered acrylamide. To avoid the health risk from working with powdered acrylamide, it is advisable to purchase a premade acrylamide solution. Materials Urea (e.g., Fisher) 38% (w/v) acrylamide/2% (w/v) bisacrylamide (e.g., Fisher; UNIT 10.4) 10× TBE electrophoresis buffer (APPENDIX 2A) 50% (w/v) APS (see recipe) N,N,N′,N′-Tetramethylethylenediamine (TEMED; Fisher) Deprotected RNA sample (see Support Protocol 1 or 2) 2× formamide loading buffer (see recipe) Elution buffer (see recipe) Chloroform, buffered (see recipe) 100 mM ATP or GTP (APPENDIX 2A; recipe for dNTPs) 100% and 80% (v/v) ethanol Vertical slab gel electrophoresis apparatus (e.g., 20 × 16–cm system from CBS Scientific), including glass plates, 1-mm spacers, fitting seal, 1-mm one- or two-well comb, clamps, and aluminum plate Power supply (e.g., Fisher) 60-mL syringe with bent 18-G needle (e.g., Fisher) Heating compartment (e.g., Fisher) filled with copper shot (e.g., Fisher), or other temperature-controlled heating block, set at 95oC Large-volume gel-loading pipet tips (e.g., Fisher) Aluminum foil Plastic wrap (e.g., Saran wrap) 20 × 20–cm TLC plate with fluorescent indicator (e.g., Fisher) 312- or 254-nm hand-held UV lamp (e.g., Fisher) Empty Poly-Prep chromatography column (Bio-Rad) Tube shaker (e.g., Fisher) 14-mL Falcon centrifuge tube (e.g., Fisher) Speedvac evaporator (e.g., Savant)
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Prepare gel 1. Assemble the gel plates, spacers, and seal following the manufacturer’s instructions (see also APPENDIX 3B). 2. Combine in a beaker in the following order: 24 g urea 25 mL 38% acrylamide/2% bisacrylamide 5 mL 10× TBE electrophoresis buffer 5 mL water. Stir, heat in a microwave oven for 20 to 30 sec (not longer), and continue stirring until urea is dissolved. This recipe produces a 20% acrylamide/8 M urea solution that should be prepared fresh for each gel. It can be scaled up for a larger gel or for several normal-sized gels. Degassing may be performed, if desired. If hexachlorofluorescein has been incorporated during RNA synthesis, urea should be left out of the gel mixture and 10 mL water added instead, as urea leads to loss of the chlorine substituents on the fluorophore.
3. Make sure that acrylamide solution is stirring at room temperature. In quick succession add 35 µL of 50% APS and 35 µL TEMED to the solution and continue stirring for 10 sec. 4. Immediately pour gel solution between glass plates either from the beaker or using a syringe as described in APPENDIX 3B. With notched glass plate on top, hold plate sandwich at a 45° angle from the benchtop and slowly pour acrylamide solution between the plates down one side. Adjust angle of plates such that gel solution flows slowly and continuously without forming bubbles. 5. When solution reaches top of notched plate, lower gel sandwich to lie flat on an empty disposable pipet tip rack. Insert thin side of a 1-mm one- or two-well comb into the solution and slowly push in until it fits snugly with the notch of the upper glass plate. Be careful to avoid introducing bubbles. Let sit for 1 hr to polymerize. For a typical 1-ìmol standard RNA synthesis (see Support Protocol 1) or a 0.2-ìmol synthesis from Dharmacon (see Support Protocol 2), one well of a two-well comb is sufficient for good separation. If more material is expected, use a one-well comb instead. Before continuing, make sure that the remaining acrylamide solution in the beaker, tilted for easier subsequent removal of polyacrylamide, has indeed polymerized.
Set up electrophoresis apparatus 6. Once acrylamide is polymerized, remove seal or bottom spacer of gel sandwich. Thoroughly clean outside of glass plates under running tap water to remove any polyacrylamide and urea residue. Remove comb gently without tearing top of gel. 7. Place gel sandwich in electrophoresis apparatus and clamp plates to upper reservoir. Clamp aluminum plate for heat distribution on front plate. 8. Tilt gel apparatus and fill bottom reservoir with 1× TBE buffer. Adjust tilt of gel apparatus so that no buffer is spilled while air is displaced from bottom of gel. Fill lower reservoir so that gel plates are submerged 2 to 3 cm. Remove any air bubbles at bottom of gel by squirting buffer between plates from one side using a 60-mL syringe with a bent 18-G needle. 9. Fill upper reservoir of gel apparatus with 1× TBE buffer to ∼1 cm from rim. 10. Prerun gel ∼15 to 30 min at 25 W constant power. In general, gels should be electrophoresed at ∼40 V/cm.
RNA Folding Pathways
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Load and run gel 11. Turn off power. Rinse well(s) with 1× TBE buffer just prior to loading gel to remove urea that has leached into the well. Use the same 60-mL syringe and bent 18-G needle used to remove bubbles from bottom of gel. 12. Dissolve the dried, deprotected RNA sample in 200 µL water and add an equal amount of 2× formamide loading buffer. Heat 1 min at 95°C, place in an ice water bath, and let chill. 13. Load whole sample in a single well using a gel-loading tip. Use a fresh tip for each sample to avoid cross-contamination. If two samples are run on a two-well gel, mark them by placing labeled sticky tape on the outside glass plate. Fold an edge of the tape over so that it can be removed easily.
14. Fill upper reservoir of gel apparatus with 1× TBE buffer to ∼1 cm from rim. If the RNA contains a fluorophore, cover gel apparatus with aluminum foil to protect the fluorophore from light. CAUTION: Obviously, the aluminum foil must not contact the electrophoresis buffer; modern buffer reservoirs have covers to prevent that.
15. Run gel at 25 W constant power until bromphenol blue has reached an appropriate distance (typically ∼1 to 3 cm) from the bottom of the gel (∼90 to 120 min). Bromphenol blue co-migrates with ∼8-mer RNA.
Visualize RNA by UV shadowing 16. Turn off power. Disassemble the gel apparatus and carefully open the gel sandwich from an unnotched corner using a spatula. Place gel on one end of a piece of plastic wrap and wrap the gel completely. If labeled sticky tape is used to mark the sample(s), transfer it onto the plastic wrap.
17. Wrap a 20 × 20–cm TLC plate with fluorescent indicator in plastic wrap. Place the wrapped gel on top of the TLC plate and visualize the RNA oligonucleotide under a 312-nm UV lamp held directly over the TLC plate. The UV-absorbing RNA blocks excitation and thus fluorescence emission of the indicator and appears as a shadow. If sensitivity is not high enough, switch to a 254-nm UV lamp; minimize exposure to short-wavelength UV light to avoid photo-induced cross-linking. When RNA contains a fluorophore, its absorbance (color) and fluorescence emission will indicate the location of labeled RNA.
18. Mark desired RNA band on the plastic wrap with a permanent marker. Remove from under the UV lamp. Normally, the slowest migrating band will be the most prevalent main product. Sometimes, however, there are branched oligonucleotides present that migrate even more slowly. Depending on the coupling efficiency of the modifications introduced, there may be exceptionally strong bands that migrate faster than the main product.
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Elute RNA from gel 19. Slice the gel on the perimeter of the product band with a clean razor blade. Cut into ∼2 × 2–mm pieces. 20. Transfer the excised gel fragments into an empty Poly-Prep chromatography column, add 4 mL elution buffer, and close tightly. If a fluorophore is already attached to the RNA, wrap column in aluminum foil. If labeled sticky tape was used to mark the sample(s), transfer it onto the column.
21. Tumble on a tube shaker overnight (16 to 20 hr) at 4°C. 22. Invert the Poly-Prep column, break off the bottom seal, uncap, and transfer the elution buffer into a 14-mL Falcon tube by gravity flow. If yield is critical, a second gel elution typically yields an additional ∼20% of the first elution.
23. Extract SDS from the elution buffer by thoroughly vortexing with an equal volume of buffered chloroform and centrifuge 10 min at 13,000 × g, 4°C, to separate the phases. Transfer the top (aqueous) phase to a fresh Falcon tube. If a pronounced interphase is carried over, a second chloroform extraction may be used to minimize it.
24. Add 100 mM ATP or GTP to a final concentration of 1 mM, then add 2 to 2.5 vol of 100% ethanol, and precipitate RNA overnight at –20°C or 2 hr at −70°C. Using ATP or GTP as a carrier increases the precipitation yield.
25. Centrifuge 30 min at 13,000 × g, 4oC, to collect RNA precipitate. Decant supernatant, wash with 80% ethanol, decant supernatant, and dry RNA in a Speedvac evaporator covered with aluminum foil. Dried RNA can be stored in a freezer at −20°C until HPLC purification (stable at least 1 month). Typical yields at this point are 20 to 100 nmol RNA from a 1-ìmol scale synthesis. Save all supernatants until you know that RNA has been recovered.
26. Resuspend dried RNA in 90 µL water for subsequent C8 reversed-phase HPLC (see Support Protocol 4). C8 REVERSED-PHASE PURIFICATION OF RNA OLIGONUCLEOTIDES C8 reserved-phase HPLC is an efficient way to separate the desired RNA from not fully deprotected material and small organic molecule contaminants from previous steps. Reversed-phase HPLC of synthetic nucleic acids is described in general in UNIT 10.5. This protocol therefore focuses on the specific parameters optimized and streamlined for modified RNA.
SUPPORT PROTOCOL 4
Materials Gel-purified RNA sample (see Support Protocol 3) 100 mM TEAA buffer, pH 7 (see recipe) Acetonitrile (see recipe) Centrifugal filtration unit (0.45-µm; Amicon) HPLC system (UNIT 10.5) with 4.6 × 250–mm Microsorb 100 C8 analytical column (5-µm particle size; Varian) and optional guard column Speedvac evaporator (e.g., Savant) Aluminum foil Spectrophotometer (220 to 800 nm) Additional reagents and equipment for reversed-phase HPLC (UNIT 10.5)
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Table 11.10.1 RNAa
Common Mobile Phase Gradients for Reversed-Phase HPLC Purification of
Sample
Elapsed time (min)
% Mobile phase B at elapsed time
RNA without fluorophores
0 24 34 0 50 0 20 60 70
0 20 40 0 60 0 20 40 60
RNA with fluorophores RNA with Cy5 and Cy3
aGradient conditions are based on a flow rate of 1 mL/min using a 4.6 × 250–mm Microsorb 100 C8 column (5-µm particle
size; Varian) at ambient temperature, and should be similar on other C8 reversed-phase columns.
1. For an analytical run, dilute 5 µL gel-purified RNA in 50 µL of 100 mM TEAA buffer, pH 7. For a preparative run, use 50% of the gel-purified RNA sample directly. Remove any particles with a centrifugal filtration unit. It is always wise to first perform an analytical run to determine the elution volume of a newly synthesized RNA so that the fraction collector can be properly programmed for the preparative run. Keeping 50% of the RNA sample for a second preparative run is advised in case of unexpected loss.
2. Start and equilibrate the HPLC system with 100% mobile phase A (100 mM TEAA buffer) according to manufacturer’s instructions. Program the gradient system (see step 4) and fraction collector (see step 5). A guard column may be used to prolong the life of the separation column.
3. Inject the RNA sample, making sure that the sample loop volume is sufficiently large (typically 100 µL). 4. Increase the percentage of mobile phase B (acetonitrile) with time according to one of the gradients listed in Table 11.10.1, depending on the particular sample. Eluted RNA is detected by UV absorbance at 254 nm. Using the gradients listed in Table 11.10.1 on a Microsorb C8 column leads to typical elution volumes between 12 and 18 min at a flow rate of 1 mL/min. If ATP or GTP was used as carrier for the ethanol precipitation, an additional peak is expected at ∼10 min. An attached fluorophore considerably retards elution of an RNA (by several min) compared to the unlabeled control. If RNA is purified after a fluorophore labeling reaction, additional peaks at high elution times (typically >18 min) will originate from excess free fluorophore.
5. Collect peak fractions and dry in a Speedvac evaporator. Cover the evaporator with aluminum foil if the RNA contains a fluorophore. At a flow rate of 1 mL/min, three to four fractions should be collected per min.
6. Dissolve each fraction in a suitable volume of water (20 to 200 µL) and combine all fractions that originate from the same HPLC elution peak. Probing RNA Structural Dynamics and Function by FRET
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7. Obtain a UV absorption spectrum of a 1:100 (v/v) diluted sample from 220 to 800 nm and calculate concentration from the peak absorption at 260 nm (1 A260 unit = 0.037 mg/mL RNA). Typical yields are 10% to 50% of the starting material. Alternatively, the molar extinction coefficient at 260 nm (ε260) of an RNA of given sequence can be estimated as the sum of extinction coefficients of the composing nucleotides, where ε260 (U) = 9,900 L mol−1 cm−1, ε260 (A) = 15,200 L mol−1 cm−1, ε260 (C) = 7,050 L mol−1 cm−1, and ε260 (G) = 12,010 L mol−1 cm−1. To calculate the concentration of a fluorophore-containing RNA, the additional absorbance of the fluorophore(s) at 260 nm should be subtracted. For fluorescein, A260/A492 = 0.3; for hexachlorofluorescein, A260/A535 = 0.3; for tetramethylrhodamine, A260/A554 = 0.49.
8. Store purified RNA (stable at least 2 years) at –20°C. If a fluorophore is already attached to the RNA, wrap tube in aluminum foil to protect from light. DATA ACQUISITION AND ANALYSIS FOR STEADY-STATE FRET Successful application of fluorescence methods in general hinges on paying close attention to experimental detail. There are numerous artifacts that can distort results and distract from obtaining meaningful results. Erroneous signals may arise from background fluorescence of solvents, stray light through the fluorometer, Raleigh and Raman scatter, higher-order light diffraction by the monochromator, and others. For more information on the origin of these artifacts, see Lakowicz (1999). In order to avoid these potential pitfalls, it is important to prepare samples and buffer solutions carefully, implement proper measurement and sample controls, and analyze data thoughtfully and on a case-by-case basis for each new RNA system. The following protocol describes generalized guidelines for: (1) preparation of samples and buffers, (2) getting started (the first steady-state fluorescence experiments), and (3) steady-state FRET data analysis.
BASIC PROTOCOL 2
To observe a conformational change by steady-state FRET, the system needs to be perturbed in a way that results in a distance change between the fluorophore pair. Steps 4 through 10 describe how this can be accomplished. Steady-state FRET refers to a continuous excitation of the donor as well as continuous recording of the donor and acceptor fluorescence signals. Such measurements can be done with widely available equipment to yield kinetic rate information, and should be the first experiments to be performed on a new RNA system. For analysis strategies in more sophisticated applications such as nanosecond time-resolved FRET for measurements of fluorophore distance distributions, please see Klostermeier and Millar (2001b). Materials Fluorophore-labeled RNA sample of choice (see Basic Protocol 1) Buffer of choice Argon gas (optional) Contrad 70 detergent (e.g., Fisher) Water or oil pump connected to side-arm Erlenmeyer flask (optional) Luer-tip syringe (e.g., Fisher) Centrifugal filtration unit (0.45-µm; Amicon) Quartz microcuvette (fill volume 120 to 150 µL; e.g., Starna) Research-type spectrofluorometer (e.g., Thermo Spectronic AMINCO-Bowman Series 2 or equivalent instrument from Jobin Yvon, Hitachi, or others), ideally with temperature control and stopped-flow equipment for fast kinetics Large-volume gel-loading tips (e.g., Fisher) RNA Folding Pathways
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NOTE: If a circulating water bath is used with the spectrofluorometer, the temperature difference between the bath and the cuvette content should be calibrated. Prepare samples and buffers 1. Use only highly purified RNA(s) (see Support Protocols 3 and 4) with fluorophores of choice attached (see Basic Protocol 1 and Commentary). 2. Choose the buffer of interest for the particular experiment. Prepare buffers using only high-quality nonfluorescent components. Most buffer components will be compatible with fluorescence experiments. Exceptions include (1) high concentrations of transition metal ions, iodide, and some aromatic compounds that quench fluorophores; (2) components that themselves are fluorescent; or (3) components that make the solution very viscous or turbid. High viscosity complicates mixing and lowers fluorophore mobility, thus increasing fluorescence anisotropy. High turbidity leads to strong light scattering.
3. Carefully remove oxygen from all buffer stock solutions used to make up the final sample by one or several of the following methods: a. Loosen container cap and heat solution in a microwave oven until close to boiling. Tighten cap, shake, loosen cap to release pressure, and repeat until no more bubbles emanate from the solution. b. Degas solution under partial vacuum by applying a water or oil pump to a side-arm Erlenmeyer flask that contains the solution in a bottle or tube with loose cap. c. For smaller volumes (e.g., up to 30 mL), load a luer-tip syringe with the solution, place Parafilm over the tip, and pull the plunger, allowing released air bubbles to move to the syringe tip. Remove the Parafilm and release all of the air. Repeat two to three times until no more air bubbles form. d. Bubble clean argon gas from a gas cylinder through the solution for 30 min. Get started: perform the first steady-state fluorescence experiments 4. Anneal the fluorophore-labeled RNA in 145 µL buffer of choice by heating to 70°C for 2 min and cooling to room temperature over 5 min. Fluorophore concentrations of 20 to 50 nM typically give a good signal-to-noise ratio. If separate RNA strands are annealed, unlabeled strands should be kept at an excess sufficient to saturate the labeled one. If donor and acceptor are coupled to two different segments of an RNA, the complex should be purified from any excess of the individual strands (see Commentary). Most standard fluorophores tolerate denaturation temperatures of 70°C. To anneal highly base-paired RNAs, higher temperatures may be necessary.
5. Centrifuge filter to remove all particles that may scatter light. 6. Clean a quartz microcuvette with 5% (v/v) Contrad 70 or other detergent and rinse thoroughly with clean, autoclaved RNase-free water. Transfer RNA solution into the cuvette. Equilibrate at temperature of choice for 15 min.
Probing RNA Structural Dynamics and Function by FRET
7. Measure emission and excitation spectra of the sample. For the emission spectrum, excite the donor at its peak absorbance and scan emission from 10 to 150 or 200 nm above this wavelength. For the excitation spectrum, detect at the acceptor peak emission and scan the excitation from 150 or 200 to 10 nm below this wavelength. Make sure that emission and excitation peaks for the specific fluorophore pair appear at the expected wavelengths. If the signal is too weak, adjust excitation and emission slit widths and signal amplification; if this is insufficient, raise the RNA concentration.
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If a strong signal is observed where it is not expected, make sure that it is not due to stray light (from light leaks in the fluorometer), Raleigh scattering (from a turbid solution), Raman scattering (will also be observed in a fluorophore-free control), or a second-order light diffraction in the monochromator (will have double the excitation wavelength). If a photomultiplier tube is used for detection, make sure it is not overloaded and damaged. If effects from the dependence of the monochromator transmission efficiency on sample polarization are to be avoided, e.g., for accurate donor-acceptor distance measurements, magic angle polarizer conditions need to be chosen at the expense of emission signal (Lakowicz, 1999).
8. Excite the donor at its excitation peak wavelength, and record its fluorescence trace at the emission peak wavelength over a sufficient time course (initially this may be 10 to 30 min at 1 datum per second). Ideally, record the acceptor signal simultaneously. Make sure that both signals are stable over time. If the signals are not stable, make sure that the solution is not visibly changing (i.e., forming a precipitate, evaporating), or ensure that a dust particle is not drifting through the light path. If one or both signals slowly decrease, fluorophore photobleaching may be the cause. This can be tested by decreasing the excitation slit width, i.e., lowering the excitation intensity. If the signal decrease becomes slower or less pronounced, photobleaching is likely the cause. In many cases photobleaching can be reduced by using a lower excitation intensity, by more completely removing oxygen (see above), or by adding a radical quencher and singlet oxygen scavenger such as 25 mM dithiothreitol.
9. Start a new time course, wait for 100 sec to ensure that the fluorescence signal is indeed stable, then close the emission shutter and access the sample to add 5 µL of an additive that perturbs the RNA system. To achieve fast manual mixing, use a small-volume pipet with a sequencing gel-loading tip to add the additive to the bottom of the cuvette and quickly pipet up and down twice with a large-volume (200-µL) pipet with gel-loading tip. Be sure not to place air bubbles in the cuvette. Close the fluorometer and open the emission shutter to continue recording. Possible additives include an additional RNA strand that binds to the fluorophore-labeled RNA (e.g., a substrate binding to a catalytic RNA), Mg2+ to assist RNA folding, or EDTA to chelate Mg2+ already present. The concentration of the additive should be in saturating excess so that simplified (i.e., pseudo-first order) reaction kinetics can be expected. If the FRET change is too fast for manual mixing, employ stopped-flow fluorescence equipment. This consumes considerably larger quantities of material, due to increased requirements both in concentration and volume.
10. Record the complete time course for both donor and acceptor emission and repeat experiment to acquire multiple data sets for calculating a standard deviation. Since all RNA molecules are synchronized with respect to the time of disturbance, any change in fluorophore distance and FRET in response to mixing will be observed as an ensemble-averaged relaxation to the new equilibrium.
Analyze steady-state FRET data 11. A meaningful steady-state FRET change is characterized by donor and acceptor signal changes in opposite directions. For a simple way to analyze these changes, calculate the ratio of acceptor to donor fluorescence over time as a relative measure for FRET efficiency. Discard any traces that contain artifacts or are particularly different from the majority. To extract rate constants, follow the procedure outlined in UNIT 11.7 (protocol for determination of tertiary folding rates and activation parameters). 12. To understand the origin of the observed FRET change(s), design control experiments in which specific experimental parameters are systematically altered, such as concentration or nature of additive, temperature, sequence of the RNA, and so on. See Commentary for explicit examples that can be found in the literature.
RNA Folding Pathways
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REAGENTS AND SOLUTIONS Use deionized, distilled, RNase-free water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Acetic acid/TEMED, 100 mM, pH 3.8 57.8 µL glacial acetic acid 5 mL water Adjust to pH 3.8 with N,N,N′,N′-tetramethylethylenediamine (TEMED) Bring to 10 mL with water Store up to 1 year at –20°C Acetonitrile Sterile filter and degas HPLC-grade acetonitrile through a 0.2-µM, organic-solventresistant bottletop filter (e.g., Millipore Millicup HV). Store up to 1 year at room temperature. Ammonium persulfate (APS), 50% (w/v) 5 g ammonium persulfate Bring up to 10 mL with water Store up to 1 year at –20°C Chloroform, buffered Mix 96 mL chloroform with 4 mL isoamyl alcohol. Overlay with 1⁄5 vol of 1× TE buffer (APPENDIX 2A) and mix again. Store up to 1 year at room temperature. For chloroform extraction, use organic (lower) phase. Elution buffer 6.67 mL 7.5 M ammonium acetate (final 500 mM) 1 mL 10% (w/v) SDS (APPENDIX 2A; final 0.1%) 20 µL 0.5 M EDTA, pH 8.0 (APPENDIX 2A; final 0.1 mM) Bring to 100 mL with water Sterilize by filtration Store up to 6 months at room temperature Formamide loading buffer, 2× 9 mL deionized formamide 1 mL 1× TBE electrophoresis buffer (APPENDIX 2A) 100 µL 2.5% (w/v) bromphenol blue Store up to 1 year at –20°C HEPES-KOH, 100 mM, pH 7.0 238.3 mg HEPES 8 mL water Adjust to 7.0 pH with 1 M KOH Bring to 10 mL with water Store up to 1 year at –20°C Sodium tetraborate, 100 mM, pH 8.5 381 mg sodium tetraborate decahydrate 8 mL water Adjust to 8.5 pH with half-concentrated HCl Bring to 10 mL with water Store up to 1 year at –20°C Probing RNA Structural Dynamics and Function by FRET
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Triethylammonium acetate (TEAA) buffer, 100 mM, pH 7 1.8 L water 12.6 mL glacial acetic acid 28 mL triethylamine Adjust to pH 7.0; when approaching pH 7.0, proceed slowly with addition of acetic acid or triethylamine so as not to overshoot Bring to 2 L with water Sterile filter and degas through a 0.22-µm bottletop filter Store up to 1 year at room temperature COMMENTARY Background Information The structure, dynamics, and function of several catalytic RNAs have been studied recently by FRET, including the hammerhead (Tuschl et al., 1994; Perkins et al., 1996; Bassi et al., 1997, 1999; Singh et al., 1999), hairpin (Murchie et al., 1998; Walter et al., 1998b, 1999, 2000, 2001; Pinard et al., 1999; Klostermeier and Millar, 2000, 2001a; Zhuang et al., 2002), hepatitis delta virus (Pereira et al., 2002), Varkud Satellite (Lafontaine et al., 2001a,b, 2002), and Tetrahymena ribozymes (Zhuang et al., 2000), as well as tRNA (Chan et al., 1999), a three-helix junction from E. coli 16S rRNA (Ha et al., 1999), and the RNA four-helix junction of U1 snRNA (Walter et al., 1998a). FRET techniques are applicable to any number of RNAs or RNA-protein complexes, and literature examples of their applications are increasing rapidly. FRET is the nonradiative transfer through space of the excited-state energy of an excited donor fluorophore to an acceptor fluorophore. When the two fluorophores are covalently tethered to defined sites on a biopolymer or a macromolecular complex, FRET can be used as a molecular ruler to estimate the distance between them. The donor normally emits at a shorter wavelength (higher energy) than the acceptor, which makes it easy to optically distinguish and quantify their relative emissions. The energy transfer efficiency (ET) strongly depends on the distance R between the two interacting fluorophores:
ET =
R06 R6 + R06
Equation 11.10.1 where R0 is the Förster distance at which 50% of the donor energy is transferred. R0 accounts for all factors besides distance that influence the rate of energy transfer, including the overlap
of the emission spectrum of the donor with the absorption spectrum of the acceptor, the donor quantum yield, and the relative orientation of the donor and acceptor transition dipole moments:
R06 = 8.79 × 10
−28
× Φ D × κ 2 × n −4 × J ( λ )
(in Å 6 ) Equation 11.10.2
where ΦD is the donor fluorescence quantum yield in the absence of acceptor, κ2 is the orientation factor of the interacting transition dipole moments, n is the refractive index of the medium, and J(λ) is the spectral overlap integral of donor emission and acceptor absorption. For most donor-acceptor pairs R0 has a value of 15 to 90 Å. FRET can be used to estimate the distance between the two fluorophores when that distance is in the range of ∼0.5 to 2 times R0. This feature makes FRET an ideal tool to complement other techniques that are applied to biological macromolecules to measure global architectures and their changes, such as NMR spectroscopy and electron microscopy, which are most sensitive at smaller and larger distances, respectively. The accessible range of rate constants that can be measured by FRET is limited only by the available fluorometer equipment and its dead time. Manual mixing has a typical dead time of 5 sec, limiting the measurable rate constants to less than ∼5 min–1. Rapid-mixing (stopped-flow) techniques reach dead times of 2 msec and below. Continuous-flow mixing techniques reach microsecond dead times. Relaxation techniques such as laser-induced temperature jumps can reach nanosecond dead times. All of these techniques can be coupled with FRET detection of structural changes.
RNA Folding Pathways
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Critical Parameters Strategic planning is critical for a successful application of FRET to RNA, since each system is quite idiosynchratic, making generalizations difficult. The effort invested in planning is often more than compensated by the unique insights gained from a successful experiment. Figure 11.10.1 summarizes a practical strategy to adapt a new RNA system to the use of FRET techniques.
Selection of RNA sequence First, one has to decide on the RNA sequence to study. Often, it might be useful to start with a wild-type or previously characterized sequence, but it is advisable to computationally check whether this sequence is predicted to fold homogeneously, e.g., by using Michael Zuker’s online RNA folding software mfold version 3.1 (http://www.bioinfo.rpi.edu/applications/ mfold/old/rna/form1.cgi) or Douglas Turner’s PC version RNAstructure (http://rna.chem. rochester.edu; see UNIT 11.2; Mathews et al.,
design RNA of interest with two fluorophores attached
no
Absence of sterical clash of fluorophores with known structural elements? yes synthesize deprotect, purify, label RNA
Does RNA ± fluorophores structurally and functionally behave the same?
no
yes
no
no
Are the alternative structures likely part of the folding pathway? yes characterize isomers structurally, kinetically, thermodynamically
Probing RNA Structural Dynamics and Function by FRET
Does the RNA fold homogeneously by steady-state FRET, gel shift, etc.?
yes
Do co-factors induce structural changes? no introduce mutations expected to induce structural changes
Figure 11.10.1 Iterative procedure to adapt a new RNA system to the use of FRET for studying structural dynamics. Adapted from Walter (2001) with permission from Academic Press.
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1999). In case of the hairpin ribozyme, for example, self-complementarity of the wildtype substrate was discovered and could be eliminated by careful redesign of the substrate and substrate-binding strands, under retention of all conserved base positions (Butcher et al., 1995; Walter and Burke, 1997). This modification led to improved catalytic (Esteban et al., 1997) and structural behavior (Hampel et al., 1998), which, in turn, allowed for a detailed characterization by FRET (Walter et al., 1998b, 1999; Zhuang et al., 2002). A simple computational check of all RNA strands for undesired secondary structures becomes particularly important when dividing a large RNA into several smaller strands that are synthesized to site-specifically introduce fluorophores. Fluorescent labeling strategies: Labeling during or after synthesis Unlike proteins, most RNAs do not contain intrinsic fluorophores (Walter and Burke, 2000). The most efficient way to site-specifically introduce the two fluorophores necessary for FRET is by synthesizing the fluorophorecontaining RNA strand(s). The fluorophore can be added during or after synthesis; in the latter case, a functional group is introduced during synthesis that allows for subsequent coupling to a reactive fluorophore derivative. Detailed descriptions of the many possible synthetic strategies are given elsewhere (Qin and Pyle, 1999; Walter and Burke, 2000). In general, solid-phase RNA synthesis in the 3′→5′ direction based on phosphoramidite chemistry (UNIT 3.5 and APPENDIX 3B) can be modified to introduce: 1. On the 5′ end: Fluorophore derivatives that are resistant to RNA deprotection chemistry (e.g., fluorescein or cyanine phosphoramidites). 2. On the 3′ end or internally: Fluorophores that are resistant to both the coupling and deprotection chemistries (e.g., fluorescein), using column supports and nucleotide phosphoramidites, respectively, modified with linkers carrying the fluorophore. 3. On the 5′ or 3′ ends or internally: Aliphatic amino or thiol linkers that can be postsynthetically coupled under mild conditions with derivatives (e.g., amino-reactive succinimidyl esters or thiol-reactive maleimides) of chemically sensitive fluorophores (e.g., rhodamines). New variations of these themes are continuously being developed, for example, the use of a site-specific phosphorothioate modification
for internal labeling (Konarska, 1999). Assembling the RNA on an automated DNA/RNA synthesizer (e.g., PE Applied Biosystems; http://www.appliedbiosystems.com) is quite straightforward using β-cyanoethyl phosphoramidites supplied by companies such as Glen Research (http://www.glenres.com), C hemGenes (http://www.chemgenes.com), Amersham Pharmacia Biotech (http://www. apbiotech.com), BD Biosciences (http://www. clontech.com), CPG (http://www.cpg-biotech. com), Cruachem (http://www.cruachem.com), Dalton Chemical Laboratories (http://www. dalton.com), or PE Applied Biosystems. More expensive, yet very convenient, is the option to buy from commercial suppliers of synthetic R NA such as Dharmacon Research (http://www.dharmacon.com), Xeragon (http:// www.xeragon.com), Midland Certified Reagent Co. (http://www.mcrc.com), IBA GmbH (http://www.iba-go.com), the Keck Foundation’s Biotechnology Resource Laboratory at the Yale University School of Medicine (http://www.info.med.yale.edu/wmkeck/oligos), Oligos Etc. (http:// www.oligosetc.com), CPG, or Cruachem. Some of these suppliers, such as Dharmacon and Xeragon, use alternative 2′OH protection chemistries to increase synthesis yield and maximum length of the synthesized RNA. Many reactive fluorophore derivatives for postsynthetic labeling of RNA are available from Molecular Probes (http://www.probes. com) or Sigma-Aldrich (http://www.sigmaaldrich.com). Succinimidyl esters of the cyanine dyes may be currently obtained only from Amersham Pharmacia Biotech. Fluorophore labeling strategies: Coupling sites and fluorophores At present, the length of a synthetic RNA is restricted to ∼50 to 80 nt due to limited coupling efficiency per nucleotide cycle. One way to overcome this limitation is to change the RNA strand connectivity; in the case of the hairpin ribozyme, this led to the use of a two-partite ribozyme with fluorophores on the 5′ and 3′ ends of the 32-nt 5′ segment. The ribozyme assembles by hybridization with a 21-nt 3′ segment and a 14-nt substrate (Walter et al., 1998b). Other approaches have used donor and acceptor on two different hybridized segments of an RNA; in this case, either the complex must be purified from any excess of individual strands (Bassi et al., 1997; Murchie et al., 1998; Lafontaine et al., 2001a) or the analysis must discriminate against an excess of (typically) the acceptor-labeled strand (Klostermeier and Mil-
RNA Folding Pathways
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Probing RNA Structural Dynamics and Function by FRET
lar, 2000, 2001a). In an elegant single-molecule FRET study, the donor fluorophore was coupled to an external substrate, while the acceptor was introduced through a DNA oligonucleotide complementary to a 3′ extension of the Tetrahymena ribozyme (Zhuang et al., 2000). The DNA oligonucleotide was also surface bound, which allowed any excess donor-labeled substrate to be washed away. Alternatively, a long contiguous RNA strand bearing an internal FRET fluorophore pair can be constructed by ligating, e.g., a synthetic fluorophore-labeled strand with one or two RNAs derived from natural sources or by in vitro transcription using T4 DNA ligase and a DNA splint (for reviews see Moore, 1999; Qin and Pyle, 1999). Additional enzymatic and chemical ligation methods are currently being developed. The choice of the two labeling sites, fluorophores, tether lengths, and adjacent RNA sequences depends on the unique features of the system to be studied. In general, the following considerations should be taken into account: 1. Available information on folding pathways and three-dimensional structures should be used to choose labeling sites expected to minimally interfere with biological function. 2. If structural transitions are to be observed, the attachment sites of the two fluorophores should be chosen to maximize the expected distance changes. 3. Each donor-acceptor pair is characterized by a specific Förster distance, R0 (Wu and Brand, 1994; Lakowicz, 1999). According to Equation 11.10.1, changes in FRET efficiency are at a maximum for distance changes that approximate R0. Hence, choosing a fluorophore pair whose R0 is close to the measured distance (e.g., within approximately two fold) will increase the sensitivity toward distance changes. 4. For certain requirements, specific donoracceptor pairs may be better suited than others. The following criteria need to be weighed against each other to choose fluorophores for a given problem. If quantity is an issue, high labeling yields can be obtained with fluorophores incorporated during synthesis. A large separation of the donor and acceptor emission peaks simplifies their optical distinction, if required by the available detection filters. High absorption, fluorescence quantum yield, and photostability lead to enhanced sensitivity, as required, e.g., for single-molecule FRET applications. Some fluorophore pairs that have been successfully used are listed below (see discussion of fluorophore pairs for FRET).
5. The labeling chemistry has to allow for site-specific incorporation of the two fluorophores. There are four options to accomplish such specificity. (1) Both fluorophores can be incorporated during the synthesis of a single RNA strand. (2) One fluorophore can be incorporated during and one after synthesis of a single RNA strand. (3) Both fluorophores can be incorporated after synthesis of a single RNA strand that carries two modifications allowing for distinct and specific (orthogonal) labeling chemistries, such as one thiol and one amino functionality. (4) The fluorophores can be incorporated into separate RNA strands that are assembled by hybridization or ligation. The choice of approach depends in part on the fluorophores used (see discussion of fluorophore pairs for FRET below). 6. A sufficient length of the fluorophore tether is necessary to assure high conformational dynamics. Such fast isotropic motion of the fluorophores is important for absolute distance measurements between donor and acceptor from Equation 11.10.1, since the Förster distance, R0, depends on the orientation factor κ2 of the interacting transition dipole moments (see Equation 11.10.2). The following values are normally well defined for a given system: ΦD, the donor fluorescence quantum yield in the absence of acceptor; n, the refractive index of the medium; and J(λ), the spectral overlap integral of donor emission and acceptor absorption. In principle, κ2 can range from 0 to 4 and is defined for a fixed relative fluorophore orientation; however, this is difficult to achieve in a dynamic solution-based system. The only other case in which κ2 adopts a well-defined value (of 2⁄3) is if the transition dipole moments of the two interacting fluorophores have an isotropic (random) relative orientation, i.e., show low fluorescence anisotropy after excitation with polarized light due to high conformational dynamics (Lakowicz, 1999; Klostermeier and Millar, 2001b; Parkhurst et al., 2001). In this case, R0 for a given FRET pair can be determined from Equation 11.10.2 so that Equation 11.10.1 yields the fluorophore distance. 7. Often, the emission of an excited fluorophore is quenched by a proximal nucleobase through an outer-sphere electron transfer between the two species. According to Marcus theory (Marcus, 1964), the rate of quenching is governed by the frequency of diffusional encounters of the excited fluorophore with the quenching base, as well as the activation barrier of their redox chemistry (Seidel et al., 1996).
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Therefore, the RNA sequence immediately adjacent to the fluorophore (and, to some extent, the fluorophore tether length; Dapprich et al., 1997) has an influence on the extent of quenching. For example, both fluorescein (Walter and Burke, 1997) and tetramethylrhodamine (Widengren et al., 1997), which constitute a popular FRET pair, are quenched by guanine. Such nucleobase-specific quenching has been utilized to observe RNA secondary structure formation (Walter and Burke, 1997; Walter et al., 1998b), yet it is best avoided to allow for absolute distance measurements by time-resolved FRET (Walter et al., 1999; Walter and Burke, 2000). Fluorophore pairs for FRET Typical examples for fluorophore pairs used on RNA are: 1. Fluorescein (exmax = 490 nm; emmax = 520 nm; quantum yield, qy = 71%) and hexachlorofluorescein (exmax = 538 nm; emmax = 551 nm). Both can be incorporated during or after synthesis. Pinard et al. (1999); Walter et al. (1998b, 2000, 2001). 2. Fluorescein and tetramethylrhodamine (exmax = 554 nm; emmax = 573 nm; qy = 28%). Any of a number of other rhodamines can also be used, but this is probably the most widely used and photophysically best understood FRET pair. Tetramethylrhodamine should be incorporated after synthesis. Clegg (1992); Tuschl et al. (1994); Perkins et al. (1996); Chan et al. (1999); Singh et al. (1999); Walter et al. (1999); Klostermeier and Millar (2000). 3. Fluorescein and cyanine 3 (Cy3; exmax = 554 nm; emmax = 573 nm; qy = 14%). Cy3 can be incorporated on the 5′ end during synthesis or anywhere after synthesis. Bassi et al. (1997, 1999); Murchie et al. (1998); Ha et al. (1999); Lafontaine et al. (2001a,b, 2002). 4. Tetramethylrhodamine and Cy5 (exmax = 652 nm; emmax = 672 nm; qy = 18%). Cy5 can be incorporated on the 5′ end during synthesis or anywhere after synthesis. The photophysics of Cy5 seem to involve fast photobleaching and dark states that are not yet well understood. Deniz et al. (1999). 5. Cy3 and Cy5 (Klostermeier and Millar, 2001a). This FRET pair is particularly useful for single-molecule experiments (Zhuang et al., 2000, 2002). Preparation of RNA and initial experiments Once the RNA sequence, strand connectivity, and modifications are chosen, the RNA is synthesized, deprotected, and purified. Post-
synthetically attached fluorophores are coupled at this stage. After a final HPLC purification step, the fluorophore-labeled RNA should be compared to unlabeled RNA to ensure that the fluorophore modification does not interfere with biological function (Fig. 11.10.1). If there are specific activity assays available, such as those for catalytic RNAs, they should be performed and compared in the presence and absence of the fluorophores. In principle, any biophysical or biochemical technique that generates a signature of proper folding and function may be used. As a broadly applicable method, unimpaired native folding may be tested on a nondenaturing polyacrylamide gel as described in UNIT 11.4, and may be detected either by autoradiography of radiolabeled RNA and/or by gel-based FRET analysis of fluorophore-labeled RNA (Ramirez-Carrozzi and Kerppola, 2001; Pereira et al., 2002). Such initial experiments may well yield the first evidence for RNA structural changes. For example, native gels may reveal alternate tertiary structures as part of an RNA folding pathway, if their interconversion rate is slow (Emerick and Woodson, 1994; Juneau and Cech, 1999; Pan et al., 2000; Pinard et al., 2001). Such structures may then be further characterized by FRET. Alternatively, RNA structural changes may be induced by the addition of co-factors (Emerick and Woodson, 1994) or substrates (Walter et al., 1998b), or by the introduction of mutations known to interfere with activity (Bassi et al., 1996; Pinard et al., 2001). By following the strategic plan in Figure 11.10.1 and combining complementary biochemical and biophysical techniques with FRET, one is likely to capture the most relevant and functionally important RNA structural changes.
Anticipated Results A typical result for a steady-state FRET assay is exemplified in Figure 11.10.2 for the hairpin ribozyme-substrate complex. A minimal reaction pathway of this ribozyme that cleaves an external substrate is composed of three reversible steps: substrate binding, cleavage, and product dissociation (Hegg and Fedor, 1995; Esteban et al., 1997; Walter et al., 1997). Previous linker insertion studies of the junction between the two independently folding domains A and B of the hairpin ribozyme-substrate complex (Feldstein and Bruening, 1993; Komatsu et al., 1994) suggested that a conformational change, docking of the two domains, occurred after binding and before the chemical step of the reaction. To observe this structural
RNA Folding Pathways
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Rz
Fluorescence (arbitrary units)
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+S(dA−1)
acceptor
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5
k dock k undock
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A D
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D
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extended
3 donor 0
200
400 Time (sec)
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Figure 11.10.2 Fluorescence signals over time as a result of tertiary structure folding of the hairpin ribozyme-substrate complex. The doubly labeled ribozyme displays a strong signal for the acceptor (A) and a weaker one for the donor (D) fluorophore. Upon manual addition of a ten-fold excess of noncleavable substrate analog S(dA–1), significant quenching of the acceptor fluorescence is observed due to rapid ribozyme-substrate complex formation. Subsequently, the acceptor signal increases, while the donor signal decreases at the same rate, reflecting the reversible transition from a flexible extended (low-FRET) to a docked (high-FRET) and catalytically active conformation (inset; short arrow, catalytic site). Adapted from Walter (2001) with permission from Academic Press.
Probing RNA Structural Dynamics and Function by FRET
transition by FRET, a two-strand version of the hairpin ribozyme was utilized and labeled with a 5′-hexachlorofluorescein acceptor and a 3′fluorescein donor. A ten-fold molar excess of the 3′ ribozyme segment was annealed with the 5′ segment by heating to 70°C for 2 min, followed by cooling to room temperature over 5 min, to saturate the fluorescently labeled strand. Steady-state fluorescence spectra and intensities were recorded as described in Basic Protocol 2 on an AMINCO-Bowman Series 2 spectrofluorometer from Thermo Spectronic in a cuvette with 3-mm excitation and emission path lengths (120-µl minimal fill volume). This instrument allows for the parallel detection of both fluorophores during a time course by continually shifting the emission monochromator back and forth. A noncleavable substrate analog with a deoxy modification at the cleavage site,
S(dA–1), and the assembled ribozyme were preincubated separately for 15 min in standard buffer (50 mM Tris⋅Cl, pH 7.5, 12 mM MgCl2, 25 mM DTT), at 25oC. Fluorescence data acquisition was started, and hairpin ribozymesubstrate complex was formed by manually mixing 145 µl ribozyme solution in the fluorometer cuvette with 5 µl substrate stock solution (supplying a saturating ten-fold substrate excess). Fluorescence emission values (one datum per second) for both donor (F515 nm) and acceptor (F560 nm) were recorded using the fluorometer software package. Upon substrate addition, a strong acceptor quench and slight donor dequench were observed (Fig. 11.10.2). At the same time, an increasing fluorescence anisotropy of the acceptor revealed decreasing fluorophore mobility upon substrate binding (Walter et al., 1998b). Under the conditions used, substrate binding was fast and
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occurred within the manual mixing time (Walter and Burke, 1997). Because the fluorescence decrease was observed only with cognate substrate, the authors concluded that the rapid acceptor quench is mostly due to quenching of hexachlorofluorescein in the ribozyme-substrate complex, presumably by a base-specific electron transfer mechanism involving the 3′terminal uracils of the substrate (Walter and Burke, 1997). Subsequently, the donor fluorescence decreased over several minutes, while the acceptor fluorescence increased at the same rate. This observation strongly suggested that the underlying molecular process involves increasing FRET between the two fluorophores, as expected for their approach upon domain docking in the ribozyme-substrate complex (inset of Fig. 11.10.2). From the temporal change of the ratio Q = F560/F515, the rate constant of the transition between the extended and docked conformations was extracted and its dependence on RNA and buffer modifications was studied (Walter et al., 1998b). It should be noted that for a reversible docking step, the observed docking rate constant kdock,obs of 0.64 ± 0.04 min−1 under standard conditions is a linear combination of the elementary docking and undocking rate constants: kdock,obs = kdock + kundock × kdock,obs can only be dissected further by an independent measurement of either the docking equilibrium constant (possible by time-resolved FRET; Klostermeier and Millar, 2001b) or the undocking rate constant (possible in single-molecule experiments; Zhuang et al., 2002). These studies on the hairpin ribozyme illustrate some of the principles that can be utilized to probe RNA structural dynamics and function by FRET. Each unique RNA system will require specific modification of these procedures to provide novel and often unanticipated results.
Time Considerations Typically, 2 to 3 weeks are needed from the design of an RNA construct to beginning the first steady-state FRET experiments with the deprotected, purified, and fully labeled material. Most of this initial work will require only a few hours of effort each day. Steady-state FRET experiments, once they are set up, will be more time consuming and will be best accomplished in 4- to 6-hr time blocks.
Literature Cited Bassi, G.S., Murchie, A.I., and Lilley, D.M. 1996. The ion-induced folding of the hammerhead ribozyme: Core sequence changes that perturb folding into the active conformation. RNA 2:756768. Bassi, G.S., Murchie, A.I., Walter, F., Clegg, R.M., and Lilley, D.M. 1997. Ion-induced folding of the hammerhead ribozyme: A fluorescence resonance energy transfer study. EMBO J. 16:74817489. Bassi, G.S., Mollegaard, N.E., Murchie, A.I., and Lilley, D.M. 1999. RNA folding and misfolding of the hammerhead ribozyme. Biochemistry 38:3345-3354. Butcher, S.E., Heckman, J.E., and Burke, J.M. 1995. Reconstitution of hairpin ribozyme activity following separation of functional domains. J Biol. Chem. 270:29648-29651. Chan, B., Weidemaier, K., Yip, W.T., Barbara, P.F., and Musier-Forsyth, K. 1999. Intra-tRNA distance measurements for nucleocapsid proteindependent tRNA unwinding during priming of HIV reverse transcription. Proc. Natl. Acad. Sci. U.S.A. 96:459-464. Clegg, R.M. 1992. Fluorescence resonance energy transfer and nucleic acids. Methods Enzymol. 211:353-388. Dapprich, J., Walter, N.G., Salingue, F., and Staerk, H. 1997. Base-dependent pyrene fluorescence used for in-solution detection of nucleic acids. J. Fluor. 7:87S-89S. Deniz, A.A., Dahan, M., Grunwell, J.R., Ha, T., Faulhaber, A.E., Chemla, D.S., Weiss, S., and Schultz, P.G. 1999. Single-pair fluorescence resonance energy transfer on freely diffusing molecules: Observation of Forster distance dependence and subpopulations. Proc. Natl. Acad. Sci. U.S.A. 96:3670-3675. Emerick, V.L. and Woodson, S.A. 1994. Fingerprinting the folding of a group I precursor RNA. Proc. Natl. Acad. Sci. U.S.A. 91:9675-9679. Esteban, J.A., Banerjee, A.R., and Burke, J.M. 1997. Kinetic mechanism of the hairpin ribozyme. Identification and characterization of two nonexchangeable conformations. J. Biol. Chem. 272:13629-13639. Feldstein, P.A. and Bruening, G. 1993. Catalytically active geometry in the reversible circularization of ‘mini-monomer’ RNAs derived from the complementary strand of tobacco ringspot virus satellite RNA. Nucl. Acids Res. 21:1991-1998. Ha, T., Zhuang, X., Kim, H.D., Orr, J.W., Williamson, J.R., and Chu, S. 1999. Ligand-induced conformational changes observed in single RNA molecules. Proc. Natl. Acad. Sci. U.S.A. 96:9077-9082. Hampel, K.J., Walter, N.G., and Burke, J.M. 1998. The solvent-protected core of the hairpin rib ozy me-sub strate complex. Biochemistry 37:14672-14682. RNA Folding Pathways
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Hegg, L.A. and Fedor, M.J. 1995. Kinetics and thermodynamics of intermolecular catalysis by hairpin ribozymes. Biochemistry 34:1581315828. Juneau, K. and Cech, T.R. 1999. In vitro selection of RNAs with increased tertiary structure stability. RNA 5:1119-1129. Klostermeier, D. and Millar, D.P. 2000. Helical junctions as determinants for RNA folding: Origin of tertiary structure stability of the hairpin ribozyme. Biochemistry 39:12970-12978.
Pereira, M.J., Harris, D.A., Rueda, D., and Walter, N.G. 2002. Reaction pathway of the trans-acting hepatitis delta virus ribozyme: A conformational change accompanies catalysis. Biochemistry 41:730-740.
Klostermeier, D. and Millar, D.P. 2001a. Tertiary structure stability of the hairpin ribozyme in its natural and minimal forms: Different energetic contributions from a ribose zipper motif. Biochemistry 40:11211-11218.
Perkins, T.A., Wolf, D.E., and Goodchild, J. 1996. Fluorescence resonance energy transfer analysis of ribozyme kinetics reveals the mode of action of a facilitator oligonucleotide. Biochemistry 35:16370-16377.
Klostermeier, D. and Millar, D.P. 2001b. Timeresolved fluorescence resonance energy transfer: A versatile tool for the analysis of nucleic acids. Biopolymers 61:159-179.
Pinard, R., Lambert, D., Walter, N.G., Heckman, J.E., Major, F., and Burke, J.M. 1999. Structural basis for the guanosine requirement of the hairpin ribozyme. Biochemistry 38:16035-16039.
Komatsu, Y., Koizumi, M., Nakamura, H., and Ohtsuka, E. 1994. Loop-size variation to probe a bent structure of a hairpin ribozyme. J. Am. Chem. Soc. 116:3692-3696.
Pinard, R., Hampel, K.J., Heckman, J.E., Lambert, D., Chan, P.A., Major, F., and Burke, J.M. 2001. Functional involvement of G8 in the hairpin ribozyme cleavage mechanism. EMBO J. 20:6434-6442.
Konarska, M.M. 1999. Site-specific derivatization of RNA with photocrosslinkable groups. Methods 18:22-28. Lafontaine, D.A, Norman, D.G., and Lilley, D.M. 2001a. Structure, folding and activity of the VS ribozyme: Importance of the 2-3-6 helical junction. EMBO J. 20:1415-1424. Lafontaine, D.A., Wilson, T.J., Norman, D.G., and Lilley, D.M. 2001b. The A730 loop is an important component of the active site of the VS ribozyme. J. Mol. Biol. 312:663-674. Lafontaine, D.A., Norman, D.G., and Lilley, D.M. 2002. The global structure of the VS ribozyme. EMBO J. 21:2461-2471. Lakowicz, J.R. 1999. Principles of Fluorescence Spectroscopy, 2nd ed. Kluwer Academic/Plenum Publishers, New York. Marcus, R.A. 1964. Chemical and electrochemical electron-transfer theory. Annu. Rev. Phys. Chem. 15:155-196. Mathews, D.H., Sabina, J., Zuker, M., and Turner, D.H. 1999. Expanded sequence dependence of thermodynamic parameters improves prediction of RNA secondary structure. J. Mol. Biol. 288:911-940. Moore, M.J. 1999. Joining RNA molecules with T4 DNA ligase. Methods Mol. Biol. 118:11-19. Murchie, A.I., Thomson, J.B., Walter, F., and Lilley, D.M. 1998. Folding of the hairpin ribozyme in its natural conformation achieves close physical proximity of the loops. Mol. Cell 1:873-881.
Probing RNA Structural Dynamics and Function by FRET
Parkhurst, L.J., Parkhurst, K.M., Powell, R., Wu, J., and Williams, S. 2001. Time-resolved fluorescence resonance energy transfer studies of DNA bending in double-stranded oligonucleotides and in DNA-protein complexes. Biopolymers 61:180-200.
Pan, J., Deras, M.L., and Woodson, S.A. 2000. Fast folding of a ribozyme by stabilizing core interactions: Evidence for multiple folding pathways in RNA. J. Mol. Biol. 296:133-144.
Qin, P.Z. and Pyle, A.M. 1999. Site-specific labeling of RNA with fluorophores and other structural probes. Methods 18:60-70. Ramirez-Carrozzi, V.R. and Kerppola, T.K. 2001. Gel-based fluorescence resonance energy transfer (gelFRET) analysis of nucleoprotein complex architecture. Methods 25:31-43. Seidel, C.A.M., Schulz, A., and Sauer, M.H.M. 1996. Nucleobase-specific quenching of fluorescent dyes. 1. Nucleobase one-electron redox potentials and their correlation with static and dynamic quenching efficiencies. J. Phys. Chem. 100:5541-5553. Singh, K.K., Parwaresch, R., and Krupp, G. 1999. Rapid kinetic characterization of hammerhead ribozymes by real-time monitoring of fluorescence resonance energy transfer (FRET). RNA 5:1348-1356. Tuschl, T., Gohlke, C., Jovin, T.M., Westhof, E., and Eckstein, F. 1994. A three-dimensional model for the hammerhead ribozyme based on fluorescence measurements. Science 266:785-789. Walter, N.G. 2001. Structural dynamics of catalytic RNA highlighted by fluorescence resonance energy transfer. Methods 25:19-30. Walter, N.G. and Burke, J.M. 1997. Real-time monitoring of hairpin ribozyme kinetics through basespecific quenching of fluorescein-labeled substrates. RNA 3:392-404. Walter, N.G. and Burke, J.M. 2000. Fluorescence assays to study structure, dynamics, and function of RNA and RNA-ligand complexes. Methods Enzymol. 317:409-440. Walter, N.G., Albinson, E., and Burke, JM. 1997. Probing structure formation in the hairpin ribozyme using fluorescent substrate analogs. Nucl. Acids Symp. Ser. 36:175-177.
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Walter, F., Murchie, A.I., Duckett, D.R., and Lilley, D.M. 1998a. Global structure of four-way RNA junctions studied using fluorescence resonance energy transfer. RNA 4:719-728.
Widengren, J., Dapprich, J., and Rigler, R. 1997. Fast interactions between Rh6G and dGTP in water studied by fluorescence correlation spectroscopy. Chem. Phys. 216:417-426.
Walter, N.G., Hampel, K.J., Brown, K.M., and Burke, J.M. 1998b. Tertiary structure formation in the hairpin ribozyme monitored by fluorescence resonance energy transfer. EMBO J. 17:2378-2391.
Wu, P. and Brand, L. 1994. Resonance energy transfer: Methods and applications. Anal. Biochem. 218:1-13.
Walter, N.G., Burke, J.M., and Millar, D.P. 1999. Stability of hairpin ribozyme tertiary structure is governed by the interdomain junction. Nat. Struct. Biol. 6:544-549. Walter, N.G., Yang, N., and Burke, J.M. 2000. Probing non-selective cation binding in the hairpin ribozyme with Tb(III). J. Mol. Biol. 298:539555. Walter, N.G., Chan, P.A., Hampel, K.J., Millar, D.P., and Burke, J.M. 2001. A base change in the catalytic core of the hairpin ribozyme perturbs function but not domain docking. Biochemistry 40:2580-2587.
Zhuang, X., Bartley, L.E., Babcock, H.P., Russell, R., Ha, T., Herschlag, D., and Chu, S. 2000. A single-molecule study of RNA catalysis and folding. Science 288:2048-2051. Zhuang, X., Kim, H., Pereira, M.J., Babcock, H.P., Walter, N.G., and Chu, S. 2002. Correlating structural dynamics and function in single ribozyme molecules. Science 296:1473-1476.
Contributed by Nils G. Walter University of Michigan Ann Arbor, Michigan
RNA Folding Pathways
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Site-Specific Fluorescent Labeling of Large RNAs with Pyrene
UNIT 11.11
RNA molecules that adopt specific folded conformations participate in many biochemical processes. A fundamental description of RNA folding pathways will allow for a more complete understanding of the roles that RNA plays in biochemistry. UNIT 11.8 provides an overview of fluorescence studies of RNA folding, and UNIT 11.10 specifically describes the application of fluorescence resonance energy transfer (FRET) methods. This unit focuses on the preparation of large RNAs—those generally >80 nucleotides (nt) in length—with covalently attached pyrene as a fluorescence probe. This approach uses just one chromophore per RNA molecule and is not based on FRET. Typically, the emission intensity of the pyrene chromophore is monitored as a function of solution conditions (e.g., Mg2+ concentration), and these emission intensity measurements are used to infer the populations of various RNA conformational states (Silverman and Cech, 1999b). Although this approach does not provide high-resolution information such as that obtained from synchrotron hydroxyl radical footprinting (UNIT 11.6), fluorescence emission intensity data can provide a simple yet meaningful overview of an RNA folding pathway. Furthermore, such data are generally easy to obtain with common and relatively inexpensive spectrometers, as long as the fluorescently labeled RNAs can be prepared. The specific task of preparing pyrene-labeled RNA is the focus of this contribution. This unit describes protocols for synthesizing large RNAs that are derivatized with pyrene attached via a short tether at a specific internal nucleotide position. Basic Protocol 1 describes the procedure for pyrene derivatization of the 2 -amino group of a short synthetic RNA oligonucleotide (<40 nt in length). This is required as the first step in preparation of large pyrene-labeled RNAs. The second step is to join the pyrene-derivatized oligonucleotide with a second, unlabeled RNA fragment to form the desired full-length RNA. Basic Protocol 2 describes a ligation procedure for this purpose. Basic Protocol 3 describes an annealing procedure to be used when the pyrene-labeled oligonucleotide does not need to be covalently joined to the remainder of the large RNA for it to adopt proper tertiary structure. Each of these protocols has been developed using the pyr3 chromophore shown in Figure 11.11.1 attached at the 2 -position of specific RNA nucleotides (Silverman and Cech, 1999b). This and other pyrene derivatives are useful in terms of fluorescence response when placed at various nucleotides within the P4-P6 domain of the Tetrahymena group I intron RNA (M.K. Smalley and S.K. Silverman, unpub. observ.). The P4-P6 RNA (Cate et al., 1996) has often been used as a test system for studies of RNA folding by fluorescence (Silverman and Cech, 1999b, 2001; Silverman et al., 2000; Young and Silverman, 2002), among other experiments. Pyrene labeling has not yet been studied systematically in large RNAs other than P4-P6. The protocols described here will assist such experiments. NOTE: For all protocols in this unit, samples should be mixed in the order indicated. After each addition of reagents, the sample tube should be mixed gently (unless otherwise indicated) and microcentrifuged briefly to collect all of the sample at the bottom of the tube. See the note at the beginning of UNIT 11.10 regarding general precautions necessary for working with RNA. Any pyrene-containing samples should be kept away from light as much as possible to avoid photodegradation of the chromophore.
RNA Folding Pathways Contributed by Mary K. Smalley and Scott K. Silverman Current Protocols in Nucleic Acid Chemistry (2004) 11.11.1-11.11.12 C 2004 by John Wiley & Sons, Inc. Copyright
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Figure 11.11.1 Reaction of the sulfotetrafluorophenyl (STP) ester pyr3-STP with a 2 -amino group, leading to pyr3derivatized RNA, which is shown as a pyrene-labeled uridine nucleotide.
STRATEGIC PLANNING Preparing Pyrene-Labeled RNA Oligonucleotides: Direct Solid-Phase Synthesis Versus Postsynthetic Labeling The P4-P6 RNA is 160 nt in length, which is too large to be prepared directly by solidphase synthesis. Therefore, to incorporate a pyrene chromophore at a specific internal position, at least two RNA fragments must be assembled. As shown in Figure 11.11.2A, pyrene is initially incorporated into a smaller portion of the RNA either by solid-phase synthesis using a phosphoramidite monomer (Silverman et al., 2000) or by derivatizing a short RNA oligonucleotide with a suitable reagent such as the sulfotetrafluorophenyl (STP) ester pyr3-STP (Fig. 11.11.1). The latter derivatization procedure is described in Basic Protocol 1. At present, the only pyrene-labeled nucleotide that is available via a commercial solidphase RNA synthesis service is pyr3-labeled uridine (Fig. 11.11.1), which is offered by Dharmacon (http://www.dharmacon.com; 2 -P-U nucleotide). Dharmacon also provides RNA oligonucleotides containing unlabeled 2 -amino-2 -deoxyuridine or 2 -amino-2 deoxycytidine; currently, the adenosine and guanosine derivatives are not commercially available. Either of these amino-derivatized nucleotides may be postsynthetically derivatized with pyr3-STP according to Figure 11.11.1 to form pyrene-labeled RNA. It is anticipated that as the pyrene fluorescence method is more widely adopted (e.g., Blount and Tor, 2003), the direct solid-phase synthesis of pyrene-labeled RNA oligonucleotides will become more readily available through commercial sources. In addition, ChemGenes (http://www.chemgenes.com) offers the pyrimidine 2 -amino-2 -deoxynucleotide phosphoramidite monomers for solid-phase RNA synthesis by individual investigators.
Site-Specific Fluorescent Labeling of Large RNAs with Pyrene
Assembling a Large Pyrene-Labeled RNA: Ligation Versus Annealing Once a pyrene-labeled RNA oligonucleotide is in hand, it can be used in the assembly of a large RNA. For P4-P6, two assembly approaches have been reported: ligation (Silverman and Cech, 1999b) and annealing (Golden et al., 1996). In the ligation approach, the short pyrene-labeled RNA oligo is covalently joined using T4 DNA ligase to a second RNA fragment that encompasses the remaining nucleotides of P4-P6 (Fig. 11.11.2B; Basic Protocol 2). Alternatively, in the annealing approach, the ligation reaction is omitted because the short oligonucleotide bearing the pyrene label can hybridize noncovalently with the remainder of the RNA, as shown schematically in Figure 11.11.2C. After the derivatization, the annealing step of Basic Protocol 3 is performed, using a modest excess of the unlabeled RNA to ensure that all of the labeled RNA becomes part of a full-length RNA complex.
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Figure 11.11.2 Synthetic approaches for large pyrene-labeled RNAs. (A) Preparing a short pyrene-labeled RNA oligonucleotide either by solid-phase synthesis or by derivatizing a 2 -aminoRNA with pyr3-STP (Basic Protocol 1). (B) Covalently ligating the short pyrene-labeled RNA to form a full-length labeled RNA (Basic Protocol 2). (C) Noncovalently annealing the short pyrenelabeled RNA to the remainder of the large RNA to form full-length RNA without ligation (Basic Protocol 3).
The ligation procedure of Basic Protocol 2 is required when the RNA backbone must be intact. In contrast, if a nick in the backbone is tolerated—that is, if the RNA fragment bearing the pyrene chromophore does not have to be covalently joined to the remainder of the RNA for proper structure—then the procedure of Basic Protocol 2 is unnecessary, and Basic Protocol 3 is performed instead. The choice of assembly strategies will be determined by the specific requirements of the large RNA being studied, as discussed in the Commentary (see Critical Parameters).
DERIVATIZATION OF A 2 -AMINO RNA OLIGONUCLEOTIDE WITH PYRENE Derivatizing the RNA oligonucleotide with pyrene requires that the RNA have an appropriately reactive 2 -amino group and that a suitably activated pyrene reagent be available. This protocol describes the use of pyr3-STP to derivatize a 2 -amino group, forming a pyr3-labeled RNA oligonucleotide, as shown in Figure 11.11.1. If the pyrene-labeled RNA is obtained directly from solid-phase synthesis, then most of Basic Protocol 1 is not necessary, and the preparation can be continued with Basic Protocol 2 or Basic Protocol 3
BASIC PROTOCOL 1
RNA Folding Pathways
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as appropriate. In all cases, however, the oligonucleotide should be purified by denaturing polyacrylamide gel electrophoresis as described below.
Materials RNA oligonucleotide with a 2 -amino group 500 mM sodium phosphate buffer, pH 8.0 (APPENDIX 2A) 10 mM EDTA, pH 8.0 (APPENDIX 2A) 50 mM pyrene 4-sulfotetrafluorophenyl ester (pyr3-STP; Molecular Probes or Gee et al., 1999) in N,N-dimethylformamide (DMF) 3 M NaCl (prepared directly or diluted from 5 M NaCl; see APPENDIX 2A) Absolute ethanol (e.g., Fisher) 80% formamide gel loading buffer with dye (see recipe) TEN buffer (see recipe) 1.7-mL RNase-free microcentrifuge tubes (e.g., Eppendorf or Fisher) Speedvac evaporator (Savant) Glass rod 50-mL plastic tube (e.g., Fisher) Platform rocker (Clay-Adams Nutator or equivalent) 0.45-µm syringe filter (e.g., Fisher) 40-mL polypropylene tube (e.g., Oak Ridge tube from Fisher) Tabletop centrifuge (e.g., IEC HN-SII), chilled to 4◦ C Refrigerated centrifuge (e.g., Beckman J2-HS with Beckman JS-13.1 rotor) Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (PAGE; APPENDIX 3B) Derivatize oligonucleotide 1. Dissolve the 2 -amino-RNA in water to make a 500 µM solution. This RNA may be obtained either from a commercial supplier or by solid-phase synthesis in an individual laboratory (see Strategic Planning).
2. Combine the following in a 1.7-mL microcentrifuge tube (total 125 µL):
20 µL 500 µM RNA from step 1 (10 nmol) 50 µL 500 mM sodium phosphate buffer, pH 8.0 5 µL 10 mM EDTA, pH 8.0 50 µL H2 O. If the RNA stock solution is at a concentration other than 500 µM, adjust the amount of water accordingly. 3. Add 125 µL of 50 mM pyr3-STP in DMF, and incubate the resulting clear solution at room temperature for 24 hr. The final concentrations during incubation are 40 µM RNA, 100 mM sodium phosphate, 0.2 mM EDTA, 25 mM pyr3-STP, and 50% (v/v) DMF.
4. Remove 125 µL of the 250-µL sample to a second 1.7-mL microcentrifuge tube. To each tube add 175 µL water, to a total volume of 300 µL. 5. To each tube add 30 µL of 3 M NaCl and 900 µL absolute ethanol, and vortex the sample for 10 sec. Freeze the samples on dry ice for at least 20 min or in a −80◦ C freezer for at least 1 hr. Site-Specific Fluorescent Labeling of Large RNAs with Pyrene
6. Microcentrifuge the samples at least 30 min at maximum speed (∼16,000 × g), 4◦ C, to precipitate the crude pyrene-labeled RNA.
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7. Remove the supernatants with a micropipettor and dry the pelleted samples in a Speedvac evaporator for 5 min. Redissolve each pellet in 50 µL water and combine the two samples.
Purify derivatized RNA 8. To the 100-µL sample, add 250 µl of 80% formamide gel loading buffer with dye. Purify the sample by denaturing PAGE (APPENDIX 3B) on a 20% gel that is 3 mm thick on 26 × 20–cm glass plates. Use one lane of a three-lane gel for a single derivatization sample. Set the power supply at 30 W and electrophorese the sample for 4 to 5 hr. 9. Excise the band that corresponds to the derivatized RNA (see gel purification method in UNIT 11.10). The desired band migrates slightly more slowly than the band corresponding to underivatized RNA (see Commentary).
10. Manually crush the excised gel band with a glass rod in a 50-mL plastic tube. 11. To extract the RNA, add 6 to 7 mL TEN buffer and place the sample tube on a platform rocker at 4◦ C for 6 to 12 hr. Centrifuge the sample at least 10 min at 200 × g (1000 rpm in IEC HN-SII), 4◦ C, and filter through a 0.45-µm syringe filter into a 40-mL polypropylene tube. 12. Extract again as in step 11 and combine the two extracts for a total eluted volume of 10 mL. The extraction will require a total of ∼12 to 13 mL of TEN buffer, of which 2 to 3 ml remains absorbed into the polyacrylamide pellet and is unrecoverable.
13. Add 27 mL cold absolute ethanol to the combined TEN elutions and place the sample at −20◦ C for at least 6 hr. 14. Precipitate the RNA by centrifuging 30 min at ∼10,000 × g (8000 rpm in Beckman JS-13.1 rotor), 4◦ C. 15. Wash the pellet by adding 1 ml cold 75% ethanol, centrifuging 10 min as in step 14, and removing the ethanol by pipetting. Dry under vacuum for 10 min. Dissolve sample in 300 µL water and quantitate by UV absorbance (A260 ). A typical isolated yield of derivatized RNA is 3 to 5 nmol starting from 10 nmol at step 1.
ASSEMBLY OF THE LARGE RNA BY LIGATION When the full-length RNA must be assembled using covalent ligation (Fig. 11.11.2B), this protocol is used. Ligation is performed using T4 DNA ligase and a complementary DNA splint that extends for at least ten nucleotides on either side of the ligation junction (Moore and Sharp, 1992; Moore and Query, 1998; Silverman and Cech, 1999a). Because yields from such splint ligation reactions can vary widely, optimization is necessary (see Commentary).
BASIC PROTOCOL 2
Materials PAGE-purified pyrene-derivatized RNA oligonucleotide (see Basic Protocol 1) DNA splint oligonucleotide (e.g., Integrated DNA Technologies or other commercial supplier) Polynucleotide that constitutes the remaining portion of the large RNA (e.g., transcribed from a plasmid DNA template, as described in Silverman and Cech, 1999a) 10× annealing buffer (see recipe) 10× ligation buffer (see recipe)
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T4 DNA ligase (e.g., USB; see Commentary) 80% formamide gel loading buffer with dye (see recipe) 1.7-mL RNase-free microcentrifuge tubes (e.g., Eppendorf or Fisher) Dry heating block Additional reagents and equipment for denaturing polyacrylamide gel electrophoresis (PAGE; APPENDIX 3B) 1. Place 2.0 nmol of pyrene-derivatized RNA oligonucleotide into a 1.7-mL microcentrifuge tube. 2. Add 3.0 nmol (1.5 eq) of the DNA splint and 3.2 nmol (1.6 eq) of the polynucleotide that constitutes the remaining portion of the large RNA. Evaporate the sample to dryness on a Speedvac evaporator and redissolve in 90 µl water. Alternatively, if the sample volume after mixing all components is less than 90 µL, omit the evaporation step and add sufficient water to raise the sample volume to 90 µL. Lower yields of RNA ligation are observed if an excess of the DNA splint and RNA polynucleotide is not used (see Commentary).
3. Add 10 µl of 10× annealing buffer. Anneal the sample by heating in a dry heating block to 95◦ C for 3 min and cooling on ice for 5 min. 4. Equilibrate the sample by heating in a water bath at 37◦ C for 2 min. 5. Add 12.5 µl of 10× ligation buffer followed by 12.5 µL of T4 DNA ligase. Incubate the sample in a water bath at 37◦ C for 4 hr. 6. Add 200 µl of 80% formamide gel loading buffer with dye. Purify the sample by denaturing PAGE (APPENDIX 3B) on a 12% gel that is 1.5 mm thick on 26 × 20–cm glass plates. Use one lane of a three-lane gel for a single ligation sample. Set the power at 30 W and electrophorese the sample for 7 to 8 hr. The separation between unligated and ligated RNA must be optimized depending on the exact sizes of the RNAs (see Commentary).
7. Excise the band that corresponds to the ligated RNA (see gel purification method in UNIT 11.10) and extract and precipitate the RNA (see Basic Protocol 1, steps 9 to 15). A typical isolated yield of ligated RNA is 0.5 to 0.8 nmol starting from 2.0 nmol at step 1.
8. Proceed to the fluorescence experiments using the RNA sample. BASIC PROTOCOL 3
ASSEMBLY OF THE LARGE RNA BY ANNEALING When the full-length RNA can be assembled by noncovalently hybridizing the pyrenelabeled fragment to the remainder of the RNA (Fig. 11.11.2C), this protocol is used. As for most annealing procedures, this protocol should not be followed until fluorescence experiments are ready to be performed on the annealed sample (see UNIT 11.8). Typically, an excess of the unlabeled RNA relative to pyrene-labeled RNA is used. This ensures that all of the pyrene label is found within a full-length RNA assembly.
Materials
Site-Specific Fluorescent Labeling of Large RNAs with Pyrene
PAGE-purified pyrene-derivatized RNA oligonucleotide (see Basic Protocol 1) Polynucleotide that constitutes the remaining portion of the large RNA (e.g., transcribed from a plasmid DNA template, as described in Silverman and Cech, 1999a) 10× TB buffer (equivalent to 10× TBE but omitting EDTA; see APPENDIX 2A) 1.7-mL RNase-free microcentrifuge tubes (e.g., Eppendorf or Fisher) Dry heating block
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1. Place 0.2 nmol of the pyrene-derivatized RNA oligonucleotide from Basic Protocol 1 into a 1.7-mL microcentrifuge tube. 2. Add 0.8 nmol (4 eq) of the polynucleotide that constitutes the remaining portion of the large RNA. 3. Add sufficient water to obtain a total volume of 603 µL. 4. Add 67 µL of 10× TB buffer, providing a total sample volume of 670 µL in 1× TB buffer. 5. Anneal the sample by heating to 60◦ C for 10 min in a dry heating block and cooling at room temperature on the benchtop for 10 min. This volume of sample is sufficient to provide at least 650 µL for transfer to a standard fluorescence cuvette. The heating temperature during annealing is kept down to 60◦C to avoid nonspecific RNA degradation. EDTA is omitted from the sample during this process, because fluorescence experiments often involve addition of small amounts of Mg2+ as part of a detailed titration series (Silverman and Cech, 1999b). The buffer used may be varied (see Commentary).
6. Proceed to the fluorescence experiments using the RNA sample.
REAGENTS AND SOLUTIONS Use deionized RNase-free water in all recipes and protocol steps. Do not use DEPC-treated water (see UNIT 11.10 for precautions to avoid RNase contamination). For common stock solutions, see APPENDIX 2A. For suppliers, see SUPPLIERS APPENDIX.
10× annealing buffer 50 µL 1 M Tris·Cl, pH 7.5 (APPENDIX 2A; 50 mM final) 2 µl 0.5 M EDTA, pH 8.0 (APPENDIX 2A; 1 mM final) Dilute to 1 mL with H2 O Store up to 1 year at room temperature Formamide gel loading buffer with dye (80% formamide) Place 25 mg each of xylene cyanol and bromphenol blue into an autoclaved 125-mL glass bottle. Add 80 mL formamide (e.g., Fisher), 10 mL 10× TBE buffer, and 10 mL 0.5 M EDTA (see APPENDIX 2A for the latter two reagents). Swirl the components to mix completely. Do not autoclave, which will decompose the formamide. Store up to 1 year at 4◦ C. Ligation buffer, 10× 500 µL 1 M Tris·Cl, pH 8.0 (APPENDIX 2A; 500 mM final) 100 µL 1 M MgCl2 (100 mM final) 300 µL 250 mM dithiothreitol (DTT; 75 mM final) 100 µL 100 mM ATP (10 mM final) Store up to 1 year at −20◦ C (avoid repeated freeze-thaw cycles, which may decompose the ATP) TEN (Tris/EDTA/NaCl) buffer 4 mL 1 M Tris·Cl, pH 8.0 (APPENDIX 2A; 10 mM final) 800 µL 0.5 M EDTA, pH 8.0 (APPENDIX 2A; 1 mM final) 40 mL 3 M NaCl (or 24 mL 5 M NaCl; APPENDIX 2A; 300 mM final) Dilute to 400 mL with H2 O Sterilize by autoclaving Store up to 1 year at room temperature
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COMMENTARY Background Information
Site-Specific Fluorescent Labeling of Large RNAs with Pyrene
Fluorescence methods are widely used throughout biochemistry to monitor conformational changes (Whitaker, 2000; Mollova, 2002). In particular, fluorescence approaches have been employed in many cases with RNA. The 2-aminopurine nucleobase is often used (Millar, 1996), but the fluorescence changes upon RNA folding are frequently small; the responses are largest when the nucleobase changes its stacking during the RNA folding event. Several reports have demonstrated that pyrene is a useful probe of RNA tertiary structure. In principle, a pyrene chromophore may be incorporated at either the 5 -terminus (Bevilacqua et al., 1992; Kierzek et al., 1993) or 3 -terminus (see Chapter 4 for terminal modifications of oligonucleotides). Alternatively, pyrene may be incorporated at an internal nucleotide position (Silverman and Cech, 1999b, 2001; Silverman et al., 2000; Young and Silverman, 2002; Blount and Tor, 2003). Although site-specific labeling at a terminus is generally possible for any RNA sequence, each RNA necessarily has many more internal positions than termini. These internal positions offer greater opportunity for monitoring different phases of RNA folding depending on the location of the chromophore. Circular permutation (Pan and Uhlenbeck, 1993; Pan, 2000) can convert an internal RNA position into a terminus, thereby overcoming this limitation. However, some sites of interest for derivatization will require maintaining the local backbone connectivity for proper RNA structure, and thus these positions are unsuitable for the circular permutation approach. This necessitates an assembly strategy in which the chromophore is incorporated internally into the large RNA. The assembly strategies shown in Figure 11.11.2 are in principle applicable to any large RNA. However, several practical issues must be addressed. First, solid-phase RNA synthesis has a limit (∼80 nt) that is much smaller than many large RNAs, which may have up to several hundred nucleotides. Therefore, the major challenge in preparing large site-specifically modified RNAs is the assembly of multiple RNA fragments, one or more of which have nonstandard nucleotides. This assembly requires either the ligation or annealing strategy of Figure 11.11.2B or C, depending on whether or not the RNA tolerates a break in the backbone at the chosen location (see Critical Parameters). Second, if the site of chro-
mophore incorporation is not near one of the termini, and therefore the short pyrene-labeled RNA oligonucleotide lies near the middle of the large RNA, multiple ligation reactions may be necessary to prepare the full-length modified RNA. Although numerous ligations can be performed sequentially, the final yields necessarily suffer (Silverman and Cech, 1999a). Fluorescence experiments require sufficiently large amounts of pyrene-labeled RNA (typically nanomoles) that multiple ligations are to be avoided whenever possible. The fluorescence experiments reported to date that use RNA ligation require just one ligation reaction to provide properly folded RNA. Once a reasonable synthesis strategy has been designed and the pyrene-labeled RNA has been prepared, either equilibrium titration or stopped-flow experiments may be performed (Bevilacqua et al., 1992; Kierzek et al., 1993; Silverman and Cech, 1999b, 2001; Silverman et al., 2000; Young and Silverman, 2002). It is also possible to monitor other fluorescence characteristics of the pyrene, such as its fluorescence lifetime and anisotropy (Preuss et al., 1997; Walter et al., 1998). The protocols described in this unit should facilitate applying these methods to an increasing array of interesting large RNA structures.
Critical Parameters RNA derivatization with pyrene The derivatization method described in Basic Protocol 1 has been optimized for the specific reaction shown in Figure 11.11.1. The reactions used a 15-mer 2 -amino-modified oligonucleotide corresponding to the first 15 nt of P4-P6, in which the 2 -amino group was on U107, the sixth nucleotide from the 5 -end. In the initial report (Silverman and Cech, 1999b), an N-hydroxysuccinimide (NHS) ester was used as the activated pyrene reagent; subsequently, the more water-soluble 4-sulfotetrafluorophenyl (STP) ester (Gee et al., 1999) was found to provide faster labeling with higher yield (M.K. Smalley and S.K. Silverman, unpub. observ.). For individual small-scale reactions, purchasing the pyr3STP reagent is likely the most expedient option. For larger-scale reactions or for multiple derivatizations, preparing the reagent according to the published procedure (Gee et al., 1999) would be worthwhile. The authors have also found that a 24mer 2 -amino-modified oligonucleotide corresponding to the last 24 nt of P4-P6
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(Golden et al., 1996) may be derivatized successfully under the same conditions when the 2 -amino-nucleotide is located at various sites within the 24-mer. For unrelated 2 -aminoRNAs that would be required for other large RNAs, the derivatization efficiency should be checked by analytical-scale assays before committing a large amount of material to the procedure. The variables to adjust include the buffer identity and pH (sodium phosphate, pH 7 to 8, or bicine, pH 8 to 9), reaction temperature (16◦ to 60◦ C), and incubation time (1 to 24 hr). To facilitate the quantification of these assays, a small portion of the oligonucleotide should be 5 -32 P-radiolabeled (e.g., UNITS 6.1 & 10.4), which permits the assay to be monitored using analytical-scale PAGE and a PhosphorImager (Silverman and Cech, 1999b). An important concern in the derivatization protocol is the extent of nonspecific labeling of various nucleophilic sites on the oligonucleotide other than the 2 -amino group. The extent of nonspecific labeling may be assessed by performing the derivatization reaction on an unmodified oligonucleotide that has the same RNA sequence but lacks the 2 -amino group. For the unmodified 15-mer oligonucleotide corresponding to the first 15 nt of P4-P6, nonspecific labeling occurs to the extent of 5% to 8% under the conditions of Basic Protocol 1. This is compared with specific labeling to the extent of ∼75% for an RNA oligonucleotide of the same sequence with a single 2 -amino group. To avoid decomposition, small aliquots of pyr3-STP stock solution (<100 µL) should be stored in a −80◦ C freezer and kept on ice when not in immediate use. If the derivatization reaction is unsuccessful, then the purity of the pyr3-STP should be checked by thin-layer chromatography (APPENDIX 3D), and the sample should be discarded or repurified by silica gel column chromatography (APPENDIX 3E) as necessary. RNA ligation The yields of T4 DNA ligase–mediated splint ligations vary widely depending on the exact sequence and structural context of the ligation site. Therefore, for any new RNA ligation, the efficiency must be optimized on a small scale using 32 P-radiolabeled samples as required. The ratio of the two RNA fragments and the DNA splint are generally chosen such that one RNA fragment is the limiting reagent and the other two nucleic acids are in modest excess (a 1:1:1 ratio of the three components
usually gives poor results). The amounts and ratios reported in Basic Protocol 2 were chosen based on empirical data, with the consideration that the pyrene-derivatized RNA oligonucleotide is typically the most precious of the three fragments and is therefore the limiting reagent. Despite significant optimization attempts, it is sometimes the case that a particular splint ligation cannot be achieved in high yield (e.g., Sontheimer et al., 1999; Strobel and OrtolevaDonnelly, 1999). If so, the most expedient strategy may be to redesign the experiment to avoid that particular ligation site, because moving the ligation site by even just one nucleotide can have an unpredictable and often substantial effect on the ligation efficiency. Ongoing efforts with deoxyribozymes (Flynn-Charlebois et al., 2003) may alleviate some of the difficulties associated with T4 DNA ligase–mediated splint ligations. Several commercial suppliers provide T4 DNA ligase. However, the enzyme from some sources may be ineffective owing to insufficient degree of purification or insufficient concentration. Because the availability of T4 DNA ligase from various manufacturers changes with time, this must be tested empirically. The authors generally use His6 -tagged T4 DNA ligase prepared in-house from a plasmid clone provided by Scott Strobel (Yale University). The amount of T4 DNA ligase required for a specific scale of ligation reaction is optimized for each batch of enzyme. After the ligation reaction has been performed, the sample is purified by denaturing PAGE (APPENDIX 3B). The percentage of acrylamide and the running time must be optimized for each ligation reaction, depending on the sizes of the unligated and ligated RNA. Generally, because the pyrene-derivatized fragment is a relatively short oligonucleotide (<40 nt) and the remainder of the RNA is often very large (>100 nt), the physical separation on the gel between unligated and ligated RNA may be small. For ligation of the pyrene-derivatized 15-mer oligonucleotide to the 145-mer corresponding to the remainder of P4-P6 (Silverman and Cech, 1999a,b), the observed separation requires a higher-than-usual percentage of acrylamide (12%) for that length of RNA, and consequently a longer running time (∼7 to 8 hr for a 1.5-mm-thick gel). For a lower percentage of acrylamide (e.g., 6%), which would be more typical for such a large RNA, the separation between unligated and ligated RNA is insufficient on a preparative scale. For RNAs other than P4-P6, the details of the electrophoresis
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must be determined empirically to identify adequate separation conditions. After preparing a sample by ligation according to Basic Protocol 2 and before fluorescence spectroscopy, one can consider whether a renaturation (annealing) step is necessary (see UNIT 11.8). At this point, the annealing procedure of Basic Protocol 3 may be performed if desired. For P4-P6, the authors have performed fluorescence experiments both with and without a renaturation step. In equilibrium fluorescence experiments the authors have observed indistinguishable results, and typically do not renature the sample for this purpose. However, for stopped-flow fluorescence experiments, modest differences in kinetics have been observed, so it would be reasonable to renature the sample before performing the experiments. In general, the need for renaturation after ligation should be checked experimentally, and the renaturation may be omitted if it is shown to be unnecessary.
Site-Specific Fluorescent Labeling of Large RNAs with Pyrene
RNA annealing The Tris-borate buffer reported for annealing in Basic Protocol 3 (1× TB) is the same as that typically used for nondenaturing gel electrophoresis of structured RNAs (Silverman and Cech, 1999a). This buffer has also proven to be suitable for pyrene fluorescence spectroscopy, although various buffer identities and strengths are acceptable (Silverman and Cech, 1999b). For other large RNAs, the effects of buffer composition on fluorescence spectroscopy should be tested explicitly, as bufferdependent differences in folding rates have been observed (Silverman et al., 2000). In the annealing procedure, the heating temperature should be high enough to denature any undesired RNA secondary structures that might be present, but it should not be so high that nonspecific RNA degradation is promoted. The annealing buffer does not contain EDTA to chelate any stray divalent metal ions that may be present, because EDTA may interfere with Mg2+ titrations during subsequent fluorescence experiments. Therefore, the commonly used annealing temperature of 90◦ to 95◦ C (in the presence of EDTA; e.g., Basic Protocol 2, step 3) is unwise in this context because any trace amount of a divalent ion such as Ca2+ could lead to degradation during the heating step. The temperature of 60◦ C reported in Basic Protocol 3 is a compromise between the need to denature and the need to avoid degradation. For other large RNAs, the heating temperature required for reproducible results must be determined empirically.
Choice of site to derivatize the RNA with pyrene The choice of a specific nucleotide to be labeled with pyrene according to Figure 11.11.1 may be guided by available X-ray crystallography or NMR spectroscopy data. For example, the X-ray crystal structure of P4-P6 (Cate et al., 1996) allowed sites for 2 -derivatization to be chosen in a rational manner (Silverman and Cech, 1999b; M.K. Smalley and S.K. Silverman, unpub. observ.). These results have led to the general prediction that many sites within other large RNAs should be good candidates for modification, even without guidance from high-resolution structural information. If the large RNA of interest has no highresolution structure available, then a reasonable strategy is to choose multiple sites and prepare several pyrene-labeled versions of the RNA in parallel. An appropriate structural or functional assay (e.g., nondenaturing gel electrophoresis or catalytic activity) should then be performed to determine if the pyrene derivatizations have disrupted the RNA structure significantly. Only those pyrene-labeled RNAs that function well in the appropriate assay should be pursued further using fluorescence experiments. Choice of site(s) to divide the RNA for assembly Each of the two strategies of Figure 11.11.2 requires choosing at least one site at which to divide the RNA. If the derivatization site is very close to the ligation junction (Fig. 11.11.2B), then T4 DNA ligase–mediated joining of the two RNA fragments may not work well. This problem is observed when a pyr3 modification is included merely three nucleotides from the ligation site within P4-P6 (M.K. Smalley and S.K. Silverman, unpub. observ.). Because the success of splint ligation is difficult to predict as described above, it is best to avoid designing a ligation site very close to the derivatization site. Regardless of the location of the ligation site, the ligation efficiency should be checked on the analytical scale using 32 P-radiolabeled RNAs before committing large amounts of material to preparative reactions. Alternatively, if the annealing strategy of Figure 11.11.2C can be used instead of ligation, then this obviates the issue of derivatization interfering with ligation.
Anticipated Results Once tractable derivatization and ligation or annealing conditions are established, synthesis of pyrene-labeled RNA is usually
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uneventful. Starting from 10 nmol of unlabeled 2 -amino oligonucleotide and assuming average yields of derivatization and ligation, ∼1 to 2 nmol of pyrene-labeled RNA is expected. At a concentration of 300 nM, which is typical for equilibrium fluorescence studies, this is sufficient for three to six experiments. Stoppedflow experiments require more material, and the preparative reactions may need to be scaled up several-fold. Typical results from both equilibrium and stopped-flow fluorescence experiments using pyrene-labeled P4-P6 are found in the published papers of the authors (Silverman and Cech, 1999b, 2001; Silverman et al., 2000; Young and Silverman, 2002).
Time Considerations
Kierzek, R., Li, Y., Turner, D.H., and Bevilacqua, P.C. 1993. 5 -Amino pyrene provides a sensitive, nonperturbing fluorescent probe of RNA secondary and tertiary structure formation. J. Am. Chem. Soc. 115:4985-4992. Millar, D.P. 1996. Fluorescence studies of DNA and RNA structure and dynamics. Curr. Opin. Struct. Biol. 6:322-326. Mollova, E.T. 2002. Single-molecule fluorescence of nucleic acids. Curr. Opin. Chem. Biol. 6:823828. Moore, M.J. and Query, C.C. 1998. Use of sitespecifically modified RNAs constructed by RNA ligation. In RNA-Protein Interactions: A Practical Approach (C.W.J. Smith, ed.) pp. 75-108. Oxford University Press, Oxford. Moore, M.J. and Sharp, P.A. 1992. Site-specific modification of pre-mRNA: The 2 -hydroxyl groups at the splice site. Science 256:992-997.
Once the appropriate 2 -amino RNA oligonucleotide(s) and pyr3-STP are in hand, testing the derivatization reaction typically takes several days. Testing the ligation reactions should require a similar length of time, although several testing rounds may be needed to find suitable sites. Finally, assaying the derivatized and ligated RNA in structural or functional assays may take several additional weeks. Once appropriate derivatization and (if necessary) ligation sites have been identified, then equilibrium or stopped-flow fluorescence experiments may be initiated. These are anticipated to take several weeks at minimum.
Pan, T. 2000. Probing RNA structure and function by circular permutation. Methods Enzymol. 317:313-330.
Literature Cited
Silverman, S.K. and Cech, T.R. 2001. An early transition state for folding of the P4-P6 RNA domain. RNA 7:161-166.
Bevilacqua, P.C., Kierzek, R., Johnson, K.A., and Turner, D.H. 1992. Dynamics of ribozyme binding of substrate revealed by fluorescence-detected stopped-flow methods. Science 258:1355-1358. Blount, K.F. and Tor, Y. 2003. Using pyrene-labeled HIV-1 TAR to measure RNA-small molecule binding. Nucl. Acids Res. 31:5490-5500. Cate, J.H., Gooding, A.R., Podell, E., Zhou, K., Golden, B.L., Kundrot, C.E., Cech, T.R., and Doudna, J.A. 1996. Crystal structure of a group I ribozyme domain: Principles of RNA packing. Science 273:1678-1685. Flynn-Charlebois, A., Wang, Y., Prior, T.K., Rashid, I., Hoadley, K.A., Coppins, R.L., Wolf, A.C., and Silverman, S.K. 2003. Deoxyribozymes with 2 -5 RNA ligase activity. J. Am. Chem. Soc. 125:2444-2454. Gee, K.R., Archer, E.A., and Kang, H.C. 1999. 4Sulfotetrafluorophenyl (STP) esters: New watersoluble amine-reactive reagents for labeling biomolecules. Tetrahedron Lett. 40:1471-1474. Golden, B.L., Gooding, A.R., Podell, E.R., and Cech, T.R. 1996. X-ray crystallography of large RNAs: Heavy-atom derivatives by RNA engineering. RNA 2:1295-1305.
Pan, T. and Uhlenbeck, O.C. 1993. Circularly permuted DNA, RNA and proteins; A review. Gene 125:111-114. Preuss, R., Dapprich, J., and Walter, N.G. 1997. Probing RNA-protein interactions using pyrenelabeled oligodeoxynucleotides: Qβ replicase efficiently binds small RNAs by recognizing pyrimidine residues. J. Mol. Biol. 273:600-613. Silverman, S.K. and Cech, T.R. 1999a. Energetics and cooperativity of tertiary hydrogen bonds in RNA structure. Biochemistry 38:8691-8702. Silverman, S.K. and Cech, T.R. 1999b. RNA tertiary folding monitored by fluorescence of covalently attached pyrene. Biochemistry 38:14224-14237.
Silverman, S.K., Deras, M.L., Woodson, S.A., Scaringe, S.A., and Cech, T.R. 2000. Multiple folding pathways for the P4-P6 RNA domain. Biochemistry 39:12465-12475. Sontheimer, E.J., Gordon, P.M., and Piccirilli, J.A. 1999. Metal ion catalysis during group II intron self-splicing: Parallels with the spliceosome. Genes & Dev. 13:1729-1741. Strobel, S.A. and Ortoleva-Donnelly, L. 1999. A hydrogen-bonding triad stabilizes the chemical transition state of a group I ribozyme. Chem. Biol. 6:153-165. Walter, N.G., Hampel, K.J., Brown, K.M., and Burke, J.M. 1998. Tertiary structure formation in the hairpin ribozyme monitored by fluorescence resonance energy transfer. EMBO J. 17:23782391. Whitaker, M. 2000. Fluorescent tags of protein function in living cells. BioEssays 22:180-187. Young, B.T. and Silverman, S.K. 2002. The GAAA tetraloop-receptor interaction contributes differentially to folding thermodynamics and kinetics for the P4-P6 RNA domain. Biochemistry 41:12271-12276.
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Key Reference Silverman and Cech, 1999b. See above. Presents the derivatization and ligation approaches to synthesizing pyrene-labeled P4-P6 RNA.
Contributed by Mary K. Smalley and Scott K. Silverman University of Illinois at Urbana-Champaign Urbana, Illinois
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CHAPTER 12 Nucleic Acid-Based Microarrays and Nanostructures INTRODUCTION ucleic acid chemistry at the material science interface has rapidly gained importance as chemists develop new strategies for nucleic acid array-based detection systems. The fabrication of DNA microarrays requires attachment of both biologically and chemically derived nucleic acids to a variety of surfaces. Surface-attached nucleic acids can be modified in ways that will allow optical or electrical detection of hybridization, extension, or other nucleic acid–based transformations. The chemistries of nucleic acid surface attachment and modification are also related to efforts to develop nucleic acids as material for fabrication of nanostructures. Their highly specific self-assembling properties, coupled with their potential for synthetic remodeling, provide the opportunity to use nucleic acid–related substances as scaffolds as well as functioning components of mechanical, electronic, or photonic devices. This chapter will provide a means to highlight advances in the field and provide detailed protocols for some of the potentially more useful applications.
N
To initiate the chapter, one of the pioneers of the field, Nadrian Seeman, provides a perspective of the advances in his laboratory. In UNIT 12.1, “Key Experimental Approaches in DNA Nanotechnology,” Dr. Seeman describes the construction of supramolecular structures based on DNA self-assembly. Important issues concerning synthesis strategy, junction design, and branching motifs are described. The characterization of complex DNA supramolecular structures is discussed in detail. This information provides important guidelines for future experimenters pursuing the construction of self-assembling supramolecular structures based on nucleic acids. In UNIT 12.2, “Preparation of Gold Nanoparticle-DNA Conjugates,” Andrew Taton provides experimental protocols for the preparation of gold nanaparticles containing a single tethered oligonucleotide, as well as gold nanoparticles coated with a dense layer of oligonucleotides. The unit includes protocols that provide specific guidelines for preparation of nanoparticles of different sizes, as well as their characterization and properties. Recent research on gold nanoparticle–linked oligonucleotides suggests that they may be of significant use as components of hybridization-based diagnostic probes. “Synthesis of 5 -Phosphoramidites Containing a Photolabile 3 -O-Protecting Group,” is the first of a series of units on building oligonucleotides on surfaces. In UNIT 12.3, Markus Beier and J¨ org Hoheisel outline procedures for the construction of deoxyribonucleoside 5 -phosphoramidites containing a photolabile protecting group on the 3 -hydroxyl group. These protected monomers allow one to build oligonucleotides in the 5 -to-3 direction, which yields surface-bound oligonucleotides with a 3 -terminus that is accessible for enzymatic extension. UNIT 12.3,
In the protocol that follows, UNIT 12.4, these authors describe procedures for derivatization of glass and polypropylene surfaces. The unique feature of the surface modification strategy described in this unit is the dendrimeric layer formed through a series of coupling reactions between 4-nitrophenyl chloroformate or acryloyl chloride and
Contributed by Donald E. Bergstrom Current Protocols in Nucleic Acid Chemistry (2005) 12.0.1-12.0.2 C 2005 by John Wiley & Sons, Inc. Copyright
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polyamines. Dendrimeric layers terminating in an amino group may be used directly for in situ DNA synthesis or, after transformation to an acrylamide or 4-nitrophenylformate moiety, as a substrate for attachment of amino-labeled oligonucleotides. Finally, in UNIT 12.5, Beier and Hoheisel show how to construct arrayed 5 -linked oligonucleotides using the 3 -O-NPPOC-protected 5 -phosphoramidites described in UNIT 12.3. The authors describe the construction of a flow cell–type apparatus for in situ synthesis on a glass microscope slide. Reagents, including phosphoramidites, are delivered to the apparatus by linking it directly to a conventional DNA synthesizer. A transparent cover allows site-specific light-mediated deprotection. Methods for linking oligonucleotides to glass surfaces for making arrays continue to improve. In UNIT 12.6, Melynk and coworkers outline procedures for making semicarbazide glass slides and alpha-oxo aldehyde conjugated oligonucleotides. The alpha-oxo aldehyde reacts rapidly with the semicarbazide to yield a highly stable alpha-oxo semicarbazone linkage. This results in arrays with superior sensitivity. Nucleic acid microarray technology has advanced considerably in the past decade. Now this technology can be directly translated to the fabrication of protein microarrays by making use of complementary nucleic acids for address-specific localization of proteins on a surface. In UNIT 12.7, Wacker and Niemeyer describe methodology for DNA-directed immobilization (DDI) of proteins from DNA-conjugated streptavidin and biotinylated antibodies. The biotin-labeled antibody is allowed to bind to the DNA-conjugated streptavidin, which in turn can be linked to specific sites on a surface bearing complementary DNA sequences. Preparation of the biomolecular conjugates as well as their attachment to an array is described in the unit. Donald E. Bergstrom
Introduction
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Key Experimental Approaches in DNA Nanotechnology Nanotechnology is concerned with controlling the structure of matter on the nanometer scale. There are many different chemistries that can lead to this endpoint. One that has proved to be very effective is DNA nanotechnology. The chemical features of DNA that make it such a successful molecule when functioning as the genetic material of living organisms can be exploited conveniently to produce a tractable system for nanotechnology. Of course, doublehelical DNA is a linear molecule, and it would be hard to produce interesting or useful structural species from it. However, it is possible to design synthetic sequences that hybridize to produce unusual motifs based on DNA branching. DNA nanotechnology is an endeavor that combines these unusual DNA motifs with cohesive (sticky) ends to yield nanoscale structures (Seeman, 1982). These structures include stick polyhedra (Chen and Seeman, 1991a; Zhang and Seeman, 1994), knots (Mueller et al., 1991; Du et al., 1995a), links (Mao et al., 1997), molecular devices (Mao et al., 1999a; Yurke et al., 2000), and both crystalline (Winfree et al., 1998; Liu et al., 1999; Mao et al., 1999b; LaBean et al., 2000; Sha et al., 2000a) and aperiodic (Mao et al., 2000) arrays in one and two dimensions. DNA is clearly a nanometer-scale object. Its diameter is ∼20 Å, and its helical repeat consists of 10 to 10.5 nucleotide pairs spaced 3.4 Å apart. It should be clear that this scale is roughly an order of magnitude larger than the scale that is exploited typically by synthetic organic chemists. What purposes could be served by construction on this scale? The key targets that have been described so far utilize DNA as architecturally robust and convenient scaffolding for other species; the expectation is that this system will be used in the crystallization of biological molecules that cannot otherwise be crystallized (Seeman, 1982) and in the construction of extremely dense nanoelectronic devices (Robinson and Seeman, 1987). Molecular devices are being developed from DNA with the goal of nanorobotics, and aperiodic assembly is likely to contribute to DNA-based computation. Figure 12.1.1 is a schematic illustrating a branched DNA molecule, similar to a Holliday (1964) junction in genetic recombination. The methods currently used to select sequences to
produce stable branched molecules are described below. The molecule is shown with four sticky ends, and the right half of the diagram illustrates four of these branched molecules cohering to produce a quadrilateral. The quadrilateral contains sticky ends on its periphery, so the quadrilateral could serve as the nucleus of an infinite two-dimensional lattice. Most readers of this series are familiar with the fact that in appropriate conditions complementary sticky ends will cohere to bring two DNA molecules together. Less well appreciated is the fact that sticky ends cohere to produce B-DNA when they pair (Qiu et al., 1997). Thus, not only does DNA generate motifs that can serve to produce connected networks (Wells, 1977) and objects with predictable topology, but sticky ends lead to locally predictable geometry when they cohere. The extent of predictability available from DNA is contrasted usefully with that available from, for example, antibodies. Although the structures of these proteins have been well established, the relative orientations of an antibody and its antigen cannot be predicted reliably and must be determined from experiments such as X-ray crystallography. In addition to the remarkable predictability of the geometry of intermolecular interactions available from DNA, these interactions have numerous other advantages as a nanotechnological system. Prominent amongst these is the ease with which DNA may be synthesized using extremely robust chemistry (Caruthers, 1985); in addition to standard DNA, many unusual bases and derivatized bases are readily available for special purposes. It is easy to purify targets and to troubleshoot experiments using the DNA-modifying enzymes that are also commercially available, including exonucleases that digest all but cyclic molecules, restriction enzymes that cleave at specific sequences, ligases that can cement sticky-ended cohesion to covalency, and topoisomerases that can solve linkage issues. The existence of these enzymes makes it easy to troubleshoot experiments. Other backbones, such as RNA or peptide nucleic acids (PNAs; Neilsen, 1995; UNIT 4.11), may ultimately be superior for specific applications, but the absence of similar enzymes for these systems militates at present for concentrating on DNA. In addition, DNA is locally a stiff molecule: its persistence length
Contributed by Nadrian C. Seeman Current Protocols in Nucleic Acid Chemistry (2002) 12.1.1-12.1.14 Copyright © 2002 by John Wiley & Sons, Inc.
UNIT 12.1
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is ∼500 Å under standard conditions (Hagerman, 1988), and the typical lengths used in DNA nanotechnology are two to three turns between branch points (∼70 to 100 Å). A further feature of DNA is that it has an externally readable code, even when its strands are wrapped tightly into double helices (Seeman et al., 1976).
els B and C are identical. However, if one performs the operation twice, five isomers are formed, including features that distinguish the fusion of parallel or antiparallel strands and the separation of the cross-over points (Figure 12.1.3; see legend for abbreviations describing cross-over events). Although the single-crossover molecules shown in Figures 12.1.1 and 12.1.2 are structurally flexible in the angles around their branchpoints (Petrillo et al., 1988), the double-cross-over (DX) molecules (Fu and Seeman, 1993) shown in Figure 12.1.3 have been shown to be quite stiff (Li et al., 1996). The DAE+J species in Figure 12.1.3, which contains an extra helical domain, is also stiff and can be incorporated into periodic arrays (Winfree et al., 1998) to produce patterns that are visible by atomic force microscopy (AFM) if its extra helix axis is directed normal to the plane containing the other helices. In addition to molecules containing doublecross-overs, DNA motifs have been developed with three co-planar helices, known as triplecross-over (TX) molecules (LaBean et al., 2000), and even a six-helix bundle has been reported (Mathieu et al., 2001). The antiparallel molecules with short separations between their cross-over points are much more robust than parallel molecules (Fu and Seeman, 1993); however, a parallel species in which every possible cross-over is present, known as paranemic cross-over (PX) DNA (Seeman, 2001), is well behaved. Paranemic DNA refers to the interaction of DNA duplex molecules with single- or
THE GENERATION OF NEW DNA MOTIFS Branching of DNA Molecules It was noted above that the DNA molecule is inherently linear because its helix axis is unbranched. However, it is easy to produce DNA molecules branched at the level of their secondary structures by performing reciprocal cross-overs between two double helices. Figure 12.1.2A illustrates the nature of the reciprocal cross-over operation that leads to DNA branching (Seeman, 2001). The effects of the reciprocal cross-over event between two duplex molecules are shown in Figures 12.1.2B and 12.1.2C. The antiparallel orientation of the DNA strands leads to two possible ways to perform the operation: between strands of the same polarity, leading to a parallel molecule (panel B), and between strands of opposite polarity, leading to an antiparallel molecule (panel C). Parallel and antiparallel can be understood as terms that describe the phasing of the connections between helices. In the case of a single exchange event, the molecules in pan-
Y′ X
Y′ X
X′
X′ Y
X
X′ Y
Key Experimental Approaches in DNA Nanotechnology
Y′
Y
Figure 12.1.1 Formation of a two-dimensional lattice from a four-arm junction with sticky ends. A sticky end and its complement are indicated by X and X′, respectively. The same relationship exists between Y and Y′. Four of the monomeric junctions on the left are complexed in parallel orientation to yield the structure on the right. DNA ligase can close the gaps left in the complex. Note that the complex has maintained open valences, so that it could be extended by the addition of more monomers.
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A
reciprocal exchange resolve
B +
reciprocal exchange resolve
C +
reciprocal exchange resolve
Figure 12.1.2 Reciprocal exchange between DNA strands. (A) Reciprocal exchange between two juxtaposed helical half turns. A black and a gray hairpin are shown. The helix axis is horizontal in this view and the dyad axis is vertical. Arrowheads on strands indicate the 3′ ends. A negative node is formed in the rightward reaction, where the strands have retained their initial shading. (B,C) Reciprocal exchange generates a four-arm junction from two double helices. Panel B shows exchange between strands of the same polarity, and panel C shows exchange between strands of opposite polarity. Symmetry elements are indicated by arrows in B.
double-stranded molecules without strand exchange.
Sequence Selection The concept of complementary DNA base pairing—A with T and G with C—is a familiar one. It is also well known that a strand with a given sequence will pair with a strand that is complementary to it to form a double helix. How is a branch point formed? The procedure is predicated on the assumption that DNA molecules will form Watson-Crick base pairs in double-helical structures in preference to all other types of base pairing. However, the concept of complementarity can be broadened to include more than a single strand (Seeman, 2000). If the way that a strand can maximize its Watson-Crick base pairing is to form a branch point, it seems that it will do so (Kallenbach et al., 1983; Wang et al., 1991). Sequences are designed according to the approach illustrated in Figure 12.1.4. Each
strand of the structure is broken up into a series of overlapping elements. In Figure 12.1.4, each of the 16-mer strands of the branched junction shown is divided into thirteen elements of four nucleotides (black boxes). Each element is required to be unique. In addition, no element that is designated to go around a corner (e.g., the boxed CTGA) is permitted to have its complement present. A further restriction is that twofold symmetry cannot flank the branch point of a four-arm junction, so that branch migration cannot occur (Seeman, 1982). With these constraints in place, competition with the target octamer structures shown in each of the arms of the junction in Figure 12.1.4 comes only from trimers or smaller, such as the ATG sequences drawn in gray boxes. This approach is based on the cooperativity of DNA hybridization, and it has a thermodynamic basis (Seeman and Kallenbach, 1983). However, the application of thermodynamic criteria has been largely abandoned in recent years (Seeman,
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DPE
DPOW
DAE
DAO
DPON
DAE+J
Figure 12.1.3 The structural isomers of DNA double-cross-over (DX) molecules. DPE, DPOW, and DPON are the three parallel DX molecules. The DAE and DAO molecules are the antiparallel isomers. Symmetry in DAO is between the thick and thin lines. DAE+J is a molecule in which the cyclic strand of DAE is extended to form a three-arm bulged junction. Arrowheads indicate 3′ ends. Symmetry elements are shown by arrows, and line thickness is related by symmetry in each drawing. In abbreviations, A is antiparallel, P is parallel, E and O refer to even and odd numbers of double-helical half turns between cross-overs, W and N indicate wide or narrow groove spacings for the odd half turn, and J is junction.
1990), because the systems used have been sufficiently robust to sustain the numerological approach described above. A program that applies this approach, SEQUIN, is available from the author.
CONSTRUCTION METHODS Synthesis and Purification
Key Experimental Approaches in DNA Nanotechnology
Solid-support methods for the synthesis of DNA containing designated sequences (Caruthers, 1985) are the central enabling methodology for the pursuit of DNA nanotechnology. This field is still at the stage where the key principles are being elucidated. Consequently, it is both economical and convenient to perform syntheses on relatively small scales (e.g., 200 nM). On a well-tuned synthesizer, one can achieve apparent trityl-based step yields of ∼99.2%, leading to an ∼45% yield of
crude target product in the synthesis of a 100mer. Despite numerous attempts at alternative approaches, purification by denaturing gel remains the most reliable (and tedious) method of separating target molecules from failure products. It is worth pointing out that DNA of this length is frequently damaged during the synthesis, so reproducing (and amplifying) it with PCR may be advisable in cases where high chemical purity is an issue (Wang et al., 1998). Future issues involving synthesis will likely entail scaling up to larger quantities for specific applications.
Hybridization In the author’s experience, the design algorithm is very reliable in producing unusual DNA motifs. However, it is predicated on estimates of equilibrium structures, and it is key that kinetic traps be avoided. Consequently, all
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matic phosphorylation produces completely phosphorylated material. If this is critical to an experiment, the most highly phosphorylated strands can be produced by restricting a molecule containing the sequence of interest (Podtelezhnikov et al., 2000). Ligation of DNA is central to many aspects of DNA nanotechnology. The author has found that enzymatic ligation is superior to chemical ligation (Du et al., 1995b), but it is still not very effective (Chen and Seeman, 1991a). Virtually stoichiometric quantities of ligase are often needed to produce relatively limited yields of product. The origin of this problem is not well understood and may arise from the presence of unphosphorylated ends or branch points titrating out the ligase. Minimizing the number of ligations of branched molecules in a preparative protocol is strongly advised. The topological state of a ligation product is often a key feature of a target molecule. If the molecule contains unpaired segments, it can be useful to employ topological protection (Seeman, 1992; Fu et al., 1994a). This is a technique whereby the single-stranded regions
samples should be heated to 90°C for 5 min and then cooled slowly. Relatively quick protocols entail a number of stages in heating blocks, such as 20 min at 65°, 45°, 37°C, room temperature, and perhaps a lower temperature. To form simpler arrays, the 90°C solution can be put in a Styrofoam box and cooled to room temperature over ∼40 hr. For more complex arrays, a thermocycling protocol is often used with preformed tiles. The presence of Mg2+ or other multivalent cations in solution appears to be required for the stability of small branched molecules on gels (Seeman et al., 1985).
Phosphorylation and Ligation Molecules that are to be ligated must contain phosphates on their 5′ ends. Phosphates may be added chemically as a final step in the synthesis or enzymatically using polynucleotide kinase. The author has found that phosphorylation of unpurified DNA with 32P-labeled phosphate may occasionally result in the labeling of a failure sequence (i.e., one whose secondary structure is more accessible to the enzyme than the target strand). Neither chemical nor enzy-
I
1
C G C A A T C C
G C G T T A G G
4
G C A C G A G T
T G A T A C C G
C G T G C T C A
A C T A T G G C
II
IV
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III Figure 12.1.4 The design of a stable four-arm branched junction. The junction shown is composed of four strands of DNA, as indicated by Arabic numerals. The 3′ end of each strand is indicated by a half arrow. Each strand is paired with two other strands to form double-helical arms, labeled with Roman numerals. There is no homologous two-fold sequence symmetry flanking the central branch point, thereby stabilizing its position. Tetrameric elements are boxed in black; trimeric elements are boxed in gray. The tetramers are all unique, and there is no complement to any tetramer flanking the junction. Competition with the target octamers can only occur from trimers, such as the ATG sequences.
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are paired temporarily with complements during ligation, thus eliminating any braiding they may undergo. This procedure has been successful in the synthesis of specific catenanes, but has not been as useful in the synthesis of knots (Du and Seeman, 1994), because the singlestranded segments consist of oligo(dT); the addition of oligo(dA) leads to the formation of DNA triplexes, whose rotation defeats the purpose of topological protection.
Solid-Support Methodology Control over the products of DNA ligation is critical for the construction of complex motifs. Our initial complex object, a DNA cube, was synthesized entirely in solution (Chen and Seeman, 1991a). This approach afforded only limited control over the edges that were designed to ligate at any particular step: they could be phosphorylated or not. After the first partial products were obtained, even this level of control was lacking, because there was no control over which 5′ ends were phosphorylated during the second step. A solution to the lack of control was the development of a solid-support methodology, allowing successive double-helical edges of the target molecule to be ligated individually (Zhang and Seeman, 1992). The start-
restrict 1 2
1
ing unit was added to the solid support, with all of its sticky ends protected as DNA hairpins. These could be deprotected individually by restriction enzymes and then ligated to another molecule whose only unclosed helix contained the complementary sticky end. Thus, every target intermediate is topologically closed, permitting intermediate purification by exonuclease treatment. One major advantage of the solid-support approach is that intermolecular reactions can be conducted at high concentrations, but crosslinking between growing objects during intramolecular reactions can be minimized. It is worth noting that sticky ends should not be self-complementary, as this unacceptably decreases control. Figure 12.1.5 illustrates the synthesis of a quadrilateral using this overall approach, which has been utilized to construct a truncated octahedron (Zhang and Seeman, 1994).
CHARACTERIZATION IN DNA NANOTECHNOLOGY Characterization of Motifs DNA nanotechnology has produced a large variety of complex motifs derived by merging
ligate
restrict 1
2
2
ligate 2
1
1
2
1
2
restrict 1 1
Key Experimental Approaches in DNA Nanotechnology
1
ligate
restrict 1,2
ligate
1 1
Figure 12.1.5 Protocol for the solid-support synthesis of a quadrilateral. Beginning with the support containing a closed strand, alternate cycles of restriction and ligation are performed, always at the position indicated as site 1. Selection of the target product (e.g., triangle, quadrilateral, pentalateral) is determined by the point at which one chooses to restrict at site 2, exposing a sticky end complementary to that exposed by restriction at site 1. Note that the final closure converts a simple cyclic molecule to a catenane.
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the notions of reciprocal exchange between strands (Seeman, 2001) and the generalization of complementarity (Seeman, 2000). If one produces a set of strands designed to form such a motif, one must demonstrate that the target motif actually has been obtained. These species are quite large and are not readily tractable to techniques that produce unambiguous structural results, such as X-ray crystallography or NMR. Consequently, a battery of analytical procedures has been developed or adapted to establish the features of these unusual DNA motifs. Partial-product gels The very first thing to do with a new motif is to demonstrate that it forms cleanly. A routine way to do this is to prepare a nondenaturing gel containing the stoichiometric complex and its subcomplexes; for example, for a four-arm junction, such a gel would have lanes containing each strand, plus each of the six strand pairs, the four three-strand complexes, and the fourstrand target complex (Kallenbach et al., 1983). It is important to work out the stoichiometry of the strands first in a pairwise fashion. Titrating each strand against one of its partial complements on gels is straightforward: the end point is the ratio that shows no excess of either strand. It is not uncommon for individual strands to migrate as multimers, and this occurrence should not be regarded as a problem. The SEQUIN algorithm is designed for the final complex, not the partial products. The successful assembly of a motif is characterized by a single band of roughly the expected molecular weight. Migration under nondenaturing conditions is a function of many factors, including surface area, molecular weight, and shape. Nevertheless, target motifs tend to migrate roughly in the vicinity of linear duplex markers of similar molecular weight. Ferguson analysis (see discussion of shape analysis, below) should always be performed to ensure that the results are not affected by the choice of gel percentage. There are three ways that a motif can fail this analysis. (1) It can be unstable, producing bands that migrate more rapidly than the target band. (2) It can produce bands that correspond to multimers of the target complex. Multimers can arise from a system that is stressed by torsional or electrostatic features of the molecular design; for example, parallel DX molecules generate eight-strand dimers, twelve-strand trimers, and sixteen-strand tetramers at high concentrations (Fu and Seeman, 1993). Mul-
timers can be avoided at high concentrations if the target complex is converted to a topologically closed form (Fu et al., 1994b). Sticky ends can be an artificial source of multimer bands (Li et al., 1996). In this case a new motif with blunt ends should be assayed, or the gels used for analysis should be run at temperatures that preclude intermotif cohesion. (3) A third form of failure is a smear, resulting from the formation of an open complex (i.e., strand 1:2:3:4:1:2:3...) rather than a closed complex (i.e., strand 1:2:3:4). This behavior was seen with the four-strand antijunction complex at high concentrations (Du et al., 1992). In addition to knowing the constituents of a new motif, it is also important to establish the stoichiometry of the complex. This is easily done for an n-strand complex by titrating an (n−1)-strand stoichiometric complex with the missing strand. For example, in a four-strand motif, a 1:1:1 three-strand complex can be titrated with the fourth strand (Kallenbach et al., 1983). The titration process is monitored readily on a nondenaturing gel. Assuming 1:1:1:1 stoichiometry, a 1:1:1:0.5 mixture will show bands for both the partial complex and the target motif. A 1:1:1:1 mixture should show a band containing only the target motif, and a 1:1:1:2 mixture should show bands corresponding both to the target motif and to the excess single strand. Shape analysis The qualitative shape of a DNA motif can be compared usefully and conveniently with standards by means of a Ferguson plot (Rodbard and Chrambach, 1971). This is a plot of log(mobility) versus acrylamide concentration, and its slope is proportional to the friction constant of the molecule. Ferguson analysis has been used to characterize the stacked character of four-arm junctions in comparison to three-, five-, and six-arm junctions (Wang et al., 1991) and to demonstrate that the addition of helices to linear duplex and DX molecules results in similar changes in the friction constant (LaBean et al., 2000). Likewise, the qualitative structure of PX DNA has been compared usefully to that of DX molecules to establish their similar shapes. Both denaturing and nondenaturing Ferguson analyses have been applied to a DNA cube-like molecule and its subcatenanes (Chen and Seeman, 1991b). Cooper and Hagerman (1987) have developed a method for the analysis of DNA branched junctions. By adding long reporter double helices pairwise to the arms of four-arm
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transient electric birefringence (Cooper and Hagerman, 1989) and fluorescence resonance energy transfer (FRET) to establish details of the branched-junction structure and dynamics (Murchie et al., 1989; Eis and Millar, 1993).
junctions, they established the basic structural features of DNA branched junctions. Their results suggest that gel mobility is a monotonically increasing function of the angle between the two extended arms. This approach was modified by Lilley and colleagues (Duckett et al., 1988) for the conventional Holliday junction, and was utilized effectively as well for Bowtie junctions containing 3′,3′ and 5′,5′ linkages in the cross-over strands of the branched species (Sha et al., 1999). The same pairwise approach has been applied to junctions using
*1 (5′)
*1 (3′) 20
10
Thermal analysis The relative stabilities of new motifs are often usefully analyzed by melting them while monitoring a feature such as OD260 (Kallenbach et al., 1983) or circular dichroism (Seeman et al., 1985). Melting a complex DNA motif
20
*2 (5′)
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2 10 G T T C A G C C T TA G T
50 60 70 C C A C A G T C A C G G AT G G A C T C G ATA G C C A A
C A A G T C G G A AT C A 70 60 20 TT T C T G G A C T C C TT A G A C C T G A G G
G G T G T C A G T G C C TA C C T G A G C TAT C G G T T 50 10 1 30 40 10 T G G C AT C T C AT T C G C A G G A C A G G TA G T T A C C G TA G A G TA A G C G T C C T G T C C AT C TT
10
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4
Key Experimental Approaches in DNA Nanotechnology
20 35 G TA G A G C A G G A A C G C A A A G C G G T TA G G T C
TTCGG
C AT C T C G T C C T T G C G T T T C G C C A AT C C A G 50 35
AAGCC 20
50 ACGCTCGT TGCGAGCA
3
Figure 12.1.6 The hydroxyl radical autofootprinting pattern of a DNA triple-cross-over (TX) molecule. The top portion of the figure contains densitometer scans of autoradiograms for each strand of the TX molecule. The data for each strand are shown twice, once for its 5′ end and once for its 3′ end, as indicated above the appropriate panel. Susceptibility to hydroxyl radical attack is compared for each strand when incorporated into the TX molecule (TX) and when paired with its traditional Watson-Crick complement (DS). Nucleotide numbers are indicated above every tenth nucleotide. The two nucleotides flanking expected cross-over positions are indicated by two Js. Note the correlation between the Js and protection in all cases. Additional protection is seen at further locations (arrows), indicating occlusion a turn away from the cross-over points on the cross-over strands, and about four nucleotides 3′ to the cross-overs on the helical strands, as noted previously. The data are summarized on a molecular drawing below the scans. Sites of protection are indicated by triangles pointing towards the protected nucleotide; the extent of protection is indicated qualitatively by the sizes of the triangles. Asterisks indicate labeled strands.
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often reveals features that are more characteristic of the compositions of individual helical domains than of the complex itself. It is often useful to pay particular attention to premelting transitions, because they may be more diagnostic of features of the complex that differentiate it from linear duplex DNA or other topologies of the same composition. Another technique of great value is analysis of the motif on a perpendicular denaturing gradient gel (Fischer and Lerman, 1979). This method enables one to see parts of the complex fall apart, rather than monitoring the details of the intradomain destacking transition. Calorimetry has also been applied usefully to DNA motifs (Marky et al., 1987). Hydroxyl radical autofootprinting The highest-resolution technique that is used conveniently to characterize the structures of unusual DNA motifs is hydroxyl radical autofootprinting. Hydroxyl radical autofootprinting (Churchill et al., 1988) is a variation on the technique of hydroxyl radical footprinting, which is used to establish the binding sites of proteins on DNA. Branched junctions (Churchill et al., 1988; Wang et al., 1991), tethered junctions (Kimball et al., 1990), antijunctions and mesojunctions (Du et al., 1992), and DX (Fu and Seeman, 1993) and TX (LaBean et al., 2000) molecules have all been analyzed by this
method. The analysis is performed by labeling a component strand of the complex and exposing it to hydroxyl radicals. The key feature noted at cross-over sites in these analyses is decreased susceptibility to attack when comparing the pattern of the strand as part of the complex relative to the pattern of the strand derived from linear duplex DNA. Decreased susceptibility is interpreted to suggest that access of the hydroxyl radical may be limited by steric factors at the sites where it is detected (Balasubramanian et al., 1998). Likewise, similarity to the duplex pattern at points of potential flexure is assumed to indicate that the strand has adopted a conventional helical structure in the complex, whether or not it is required by the secondary structure. In studies of junctions, DX molecules, and mesojunctions, protection has been seen particularly at the cross-over sites, but also at non-cross-over sites where strands from two adjacent parallel or antiparallel domains appear to occlude each other’s surfaces, thereby preventing access by hydroxyl radicals (Kimball et al., 1990; Fu and Seeman, 1993; LaBean et al., 2000). Thus, cross-over sites can be located reliably by hydroxyl radical autofootprinting analysis, but it is not always possible to distinguish them unambiguously from juxtapositions of backbone strands. The technique is particularly powerful as a method to establish whether
ligated DAE
ligated DAE+J
ligated DAO
Figure 12.1.7 Ligation products of antiparallel double-cross-over molecules DAE, DAO, and DAE+J. One domain has been capped by hairpins. Ligation of the DAE molecule leads to a reporter strand, which is drawn more darkly. Ligation of the DAE+J molecule also leads to a reporter strand, similar to the one in DAE. However, ligation of the DAO molecule produces a polycatenated structure.
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cross-overs occur where they are expected to occur. Those who wish to construct new motifs are advised strongly to ascertain by hydroxyl radical autofootprinting that they have formed the molecular topologies that they have designed. An example of the hydroxyl radical protection pattern of a DNA TX molecule is shown in Figure 12.1.6.
Characterization of Ligation Targets Denaturing gels Ligated species are extremely difficult to analyze. The simplest ligated species are linear multimers of a given motif. Junctions (Ma et al., 1986; Petrillo et al., 1988), DX molecules (Li et al., 1996; Winfree et al., 1998), and DNA triangles (Yang et al., 1998) are all examples of unusual DNA motifs that have been oligomerized in one dimension. The trick to analyzing these systems is for them to contain a reporter strand whose fate reports the fate of the complex. Examples of reporter strands are shown in Figure 12.1.7, where the ligation products of DAE, DAE+J, and DAO molecules are shown. The thick strand in the DAE and DAE+J ligations is a reporter strand, but there is no reporter strand in the DAO ligation. It is important to extract the reporter strands from other strands in the ligation product to which they might be catenated; this is usually done by restriction. The reporter strand can be sized in comparison with linear and cyclic markers (e.g., Li et al.,
−
Key Experimental Approaches in DNA Nanotechnology
1996). In addition to providing an estimate of the products, ligation experiments can be used to estimate the stiffness of a given motif, either qualitatively (Ma et al., 1986; Petrillo et al., 1988; Li et al., 1996) or quantitatively (Podtelezhnikov et al., 2000). More complex target ligation products are likely to be catenanes or perhaps knots. This is a consequence of the plectonemic (interwound) nature of the DNA strands; indeed, DNA is almost the ideal synthon for topological construction. Catenanes are usually separable on denaturing gels of one (Chen and Seeman, 1991a,b) or two (Wang, et al., 1998) dimensions. Catenanes of greater linking number migrate more rapidly on denaturing gels than similar catenanes of lower linking number. This seems to be true of knots as well, although it is key to ensure that a fortuitous acrylamide percentage has not been selected for the analysis. A reliable way to ensure independence from gel artifacts is to perform a denaturing sedimentation analysis in comparison with markers. If material appears to behave anomalously on denaturing gels, it is useful to check that the sample is completely denatured. The FischerLerman 100% denaturing conditions (7 M urea and 40% formamide; Fischer and Lerman, 1979) often denature samples that misbehave on conventional denaturing gels.
=
Figure 12.1.8 Restriction analysis of a DNA cube. The cube is drawn with strands of three different thicknesses, with the darkest in front and the lightest at the rear. The linear triple catenane shown at the center was the starting material for the last step of the construction. It corresponds to the left, front, and right sides of the cube. Its removal from the cube by restricting at the left-front and right-front edges leaves the top-back-bottom linear triple catenane as a product. Each edge of the cube contains a unique restriction site.
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Restriction analysis Complex constructions, such as DNA polyhedra (Chen and Seeman, 1991a; Zhang and Seeman, 1994) or Borromean rings (Mao et al., 1997), must be designed carefully from the start, so that proof of construction can be obtained. As noted above, crystallography is not readily available to characterize these species, and molecular weights of 150 to 800 kDa are beyond the range of current NMR capabilities. At present, the best characterizations of these systems are topological rather than structural. The analysis entails the judicious insertion of restriction sites in the molecules. These sites are utilized to break the products down to smaller catenanes that can be synthesized independently and used for comparison. Thus, the DNA cube, a hexacatenane (Chen and Seeman, 1991a), was built from a linear triple catenane corresponding to the ultimate left-front-right faces of the target. Figure 12.1.8 shows that cleavage of the left-front and right-front edges of the cube leads to the top-back-bottom linear triple catenane. Likewise, the cleavage of any of the strands of the Borromean rings leads to two single strands (Mao et al., 1997).
Characterization of Devices DNA nanomechanical devices hold great p ro mise in th e d evelo ping field of nanorobotics. Devices may consist of ligated (Mao et al., 1999a) or annealed (Yurke et al., 2000; Yan et al., 2002) components. The requirements for characterization of the system are no less stringent for a device than for a static target. In addition, one must demonstrate that the device responds to the triggers for which it has been designed. Mechanical motion is the hallmark of a device, and this intramolecular motion leads to a change in some distance within the molecule that can be monitored by FRET. FRET has been used successfully to demonstrate the action of devices predicated on the B-Z transition (Mao et al., 1999a) and on the binding and removal of a specific strand (Yurke et al., 2000). A sequence-specific device based on hybridization topology has also been demonstrated recently by AFM. An edge-sharing half hexagon attached roughly perpendicular to the device provides a 17-nm lever arm whose repositioning is readily visible at the 7to 10-nm resolution of the AFM (Yan et al., 2002).
Characterization of DNA Arrays Periodic arrays The use of reporter strands to establish the extent that a system has ligated in one dimension was described above. However, such an approach is of less utility in two dimensions (or three), because it will not reveal the nature or extent of faults very well. Direct observation of the array is necessary. The technique used to date is AFM. Periodic arrays of DX molecules (Winfree et al., 1998), TX molecules (LaBean et al., 2000), and conventional (Mao et al., 1999b) and Bowtie (Sha et al., 2000a) DNA parallelograms have all been characterized by this method. As with all microscopy, the investigator can usually find the structure that is sought; consequently, any observations must be challenged by making a molecular-level change that leads to a predictable change in the AFM pattern. For example, DAE molecules can be interspersed with DAE+J molecules (Figure 12.1.3) whose extra helical domain is oriented out of the plane of the array. For DAE molecules of dimensions 4 × 16 nm, alternating DAE molecules with DAE+J molecules leads to a striped feature every 32 nm; if there is only a single DAE+J molecule in every four molecules, then the stripes will be separated by 64 nm. Likewise, differently shaped cavities can be designed for arrays made up of DNA parallelograms. To ensure reliable results, changes in molecular design must lead to predictable changes in AFM patterns. Studies of periodic DNA arrays using other microscopic techniques such as transmission electron microscopy, scanning transmission electron microscopy, or near-field scanning optical microscopy are ongoing, but no data have yet emerged in the literature. Similarly, attempts are going forward to produce three-dimensional crystalline materials, but no successes have been reported. X-ray diffraction clearly will be required to characterize threedimensional periodic DNA constructions. Aperiodic arrays The founding of experimental DNA-based computation by Adleman (1994) has led to an interest in other types of DNA self-assembly. Winfree (1996) pointed out that branched DNA motifs could be used to do logical computation. The idea is that sticky ends can be assigned a logical value, and that their assembly can then be used to perform the computation. This suggestion has been implemented successfully in one dimension, where a reporter strand ap-
DNA Nanotechnology
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proach was used to extract the answer to the computation (Mao et al., 2000). Algorithmic assembly can also be used to program patterns in two or three dimensions, but patterns of this complexity have not yet been obtained. Characterization in two dimensions may be amenable to AFM analysis, but new methods, beyond AFM and diffraction, will be needed in three dimensions to confirm the presence of designed aperiodic patterns.
DNA NANOTECHNOLOGY AS A TOOL IN BIOCHEMISTRY The applications of DNA nanotechnology are certainly not limited to technological goals like solution of the macromolecular crystallization problem or the facilitation of nanoelectronics or nanorobotics. DNA nanotechnology has already served as a valuable tool in the characterization of components of biological systems. DX molecules have been used to establish the thermodynamics of Holliday junction cross-over isomers (Zhang and Seeman, 1994) and branch migratory isomers (Sun et al., 1998). They have also been used to establish Holliday junction cross-over topology (Fu et al., 1994a), the cutting sites of resolvases (Sha et al., 2000c), and the spontaneity of both Holliday cross-over isomerization (Li et al., 1997) and antiparallel branch migration (Sha et al., 2000b). Two-dimensional parallelogram arrays have been used to measure the interdomain angles in Holliday (Mao et al., 1999b) and Bowtie (Sha et al., 2000a) junctions; V-shaped parallelogram arrays were used to confirm the parallel orientation of Bowtie junction domains (Sha et al., 2000a). Similarly, an RNA knot was used to demonstrate that E. coli DNA topoisomerase III is an RNA topoisomerase, although topoisomerase I is not (Wang et al., 1996, 1998). Other applications certainly exist and are not just limited to the analysis of Holliday junctions; they are limited only by the imagination.
CONCLUSIONS
Key Experimental Approaches in DNA Nanotechnology
This unit is intended to give a sense of the nature, goals, and experimental techniques that characterize DNA nanotechnology. DNA nanotechnology that utilizes the detailed geometry that results from the predictable molecular structures formed by sticky-ended cohesion has been emphasized. Other approaches to nanotechnology using DNA have been performed utilizing only the complementarity aspects of DNA, without the concomitant structural features inherent in sticky-ended interac-
tions (e.g., Niemeyer et al., 1994; Alivisatos et al., 1996; Mirkin et al., 1996; Shi and Bergstrom, 1997). However, these investigators have brought an important new component to DNA nanotechnology: the combination of DNA and heteromolecules, such as proteins and nanocrystals. It is likely that as more investigators enter this field, heteromolecules will be added to geometrically precise DNA constructs in a way that will incorporate the functionality that this system requires if it is to have a major impact.
LITERATURE CITED Adleman, L. 1994. Molecular computation of solutions to combinatorial problems. Science 266:1021-1024. Alivisatos, A.P., Johnsson, K.P., Peng, X., Wilson, T.E., Loweth, C.J., Bruchez, M.P., and Schultz, P.G. 1996. Organization of ‘nanocrystal molecules’ using DNA. Nature 382:609-611. Balasubramanian, B., Pogozelski, W.K., and Tullius, T.D. 1998. DNA strand breaking by the hydroxyl radical is governed by the accessible surface areas of the hydrogen atoms of the DNA backbone. Proc. Natl. Acad. Sci. U.S.A. 95:97389743. Caruthers, M.H. 1985. Gene synthesis machines: DNA chemistry and its uses. Science 230:281285. Chen, J. and Seeman, N.C. 1991a. The synthesis from DNA of a molecule with the connectivity of a cube. Nature 350:631-633. Chen, J. and Seeman, N.C. 1991b. The electrophoretic properties of a DNA cube and its substructure catenanes. Electrophoresis 12:607611. Churchill, M.E.A., Tullius, T.D., Kallenbach, N.R., and Seeman, N.C. 1988. A Holliday recombination intermediate is twofold symmetric. Proc. Natl. Acad. Sci. U.S.A. 85:4653-4656. Cooper, J.P. and Hagerman, P.J. 1987. Gel electrophoretic analysis of the geometry of a DNA four-way junction. J. Mol. Biol. 198:711-719. Cooper, J.P. and Hagerman, P.J. 1989. Geometry of a branched DNA structure in solution. Proc. Natl. Acad. Sci. U.S.A. 86:7336-7340. Du, S.M. and Seeman, N.C. 1994. The construction of a trefoil knot from a DNA branched junction motif. Biopolymers 34:31-37. Du, S.M., Zhang, S., and Seeman, N.C. 1992. DNA junctions, antijunctions and mesojunctions. Biochemistry 31:10955-10963. Du, S.M., Stollar, B.D., and Seeman, N.C. 1995a. A synthetic DNA molecule in three knotted topologies. J. Am. Chem. Soc. 117:1194-1200. Du, S.M., Wang, H., Tse-Dinh, Y.-C., and Seeman, N.C. 1995b. Topological transformations of synthetic DNA knots. Biochemistry 34:673-682.
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Duckett, D.R., Murchie, A.I., Diekmann, S., von Kitzing, E., Kemper, B., and Lilley, D.M.J. 1988. The structure of the Holliday junction, and its resolution. Cell 55:79-89.
Mao, C., Sun, W., and Seeman, N.C. 1999b. Designed two-dimensional DNA Holliday junction arrays visualized by atomic force microscopy. J. Am. Chem. Soc. 121:5437-5443.
Eis, P.S. and Millar, D.P. 1993. Conformational distributions of a four-way DNA junction revealed by time-resolved fluorescence resonance energy transfer. Biochemistry 32:13852-13860.
Mao, C., LaBean, T., Reif, J.H., and Seeman, N.C. 2000. Logical computation using algorithmic self-assembly of DNA triple crossover molecules. Nature 407:493-496.
Fischer, S.G. and Lerman, L.S. 1979. Length-independent separation of DNA restriction fragments in two-dimensional gel electrophoresis. Cell 16:191-200.
Marky, L.A., Kallenbach, N.R., McDonough, K.A., Seeman, N.C., and Breslauer, K.J. 1987. The melting behavior of a nucleic acid junction: A calorimetric and spectroscopic study. Biopolymers 26:1621-1634.
Fu, T.-J. and Seeman, N.C. 1993. DNA double crossover structures. Biochemistry 32:32113220. Fu, T.-J., Tse-Dinh, Y.-C., and Seeman, N.C. 1994a. Holliday junction crossover topology. J. Mol. Biol. 236:91-105. Fu, T.-J., Kemper, B., and Seeman, N.C. 1994b. Endonuclease VII cleavage of DNA double crossover molecules. Biochemistry 33:38963905. Hagerman, P.J. 1988. Flexibility of DNA. Annu. Rev. Biophys. Biophys. Chem. 17:265-286. Holliday, R. 1964. A mechanism for gene conversion in fungi. Genet. Res. 5:282-304. Kallenbach, N.R., Ma, R.-I., and Seeman, N.C. 1983. An immobile nucleic acid junction constructed from oligonucleotides. Nature 305:829831. Kimball, A., Guo, Q., Lu, M., Kallenbach, N.R., Cunningham, R.P., Seeman, N.C., and Tullius, T.D. 1990. Conformational isomers of Holliday junctions. J. Biol. Chem. 265:6544-6547.
Mathieu, F., Mao, C., and Seeman, N.C. 2001. A DNA nanotube based on a six-helix bundle motif. J. Biomol. Struct. Dyn. 18:907-908. Mirkin, C.A., Letsinger, R.L., Mucic, R.C., and Storhoff, J.J. 1996. A DNA-based method for rationally assembling nanoparticles into macroscopic materials. Nature 382:607-609. Mueller, J.E., Du, S.M., and Seeman, N.C. 1991. The design and synthesis of a knot from singlestranded DNA. J. Am. Chem. Soc. 113:63066308. Murchie, A.I., Clegg, R.M., von Kitzing, E., Duckett, D.R., Diekmann, S., and Lilley, D.M.J. 1989. Fluorescence energy transfer shows that the four-way DNA junction is a right-handed cross of antiparallel molecules. Nature 341:763-766. Neilsen, P.E. 1995. DNA analogs with non-phosphodiester backbones. Annu. Rev. Biophys. Biophys. Chem. 14:167-183. Niemeyer, C.M., Sano, T., Smith, C.L., and Cantor, C.R. 1994. Oligonucleotide-directed self-assembly of proteins. Nucl. Acids Res. 22: 5530-5539.
LaBean, T., Yan, H., Kopatsch, J., Liu, F., Winfree, E., Reif, J.H., and Seeman, N.C. 2000. The construction, analysis, ligation and self-assembly of DNA triple crossover complexes. J. Am. Chem. Soc. 122:1848-1860.
Petrillo, M.L., Newton, C.J., Cunningham, R.P., Ma, R.-I., Kallenbach, N.R., and Seeman, N.C. 1988. The ligation and flexibility of four-arm DNA junctions. Biopolymers 27:1337-1352.
Li, X., Yang, X., Qi, J., and Seeman, N.C. 1996. Antiparallel DNA double crossover molecules as components for nanoconstruction. J. Am. Chem. Soc. 118:6131-6140.
Podtelezhnikov, A., Mao, C., Seeman, N.C., and Vologodskii, A.V. 2000. Multimerization-cyclization of DNA fragments as a method of conformational analysis. J. Biophys. 79:2692-2704.
Li, X., Wang, H., and Seeman, N.C. 1997. Direct evidence for Holliday junction crossover isomerization. Biochemistry 36:4240-4247.
Qiu, H., Dewan, J.C., and Seeman, N.C. 1997. A DNA decamer with a sticky end: The crystal structure of d-CGACGATCGT. J. Mol. Biol. 267:881-898.
Liu, F., Sha, R., and Seeman, N.C. 1999. Modifying the surface features of two-dimensional DNA crystals. J. Am. Chem. Soc. 121:917-922. Ma, R.-I., Kallenbach, N.R., Sheardy, R.D., Petrillo, M.L., and Seeman, N.C. 1986. Three-arm nucleic acid junctions are flexible. Nucl. Acids Res. 14:9745-9753.
Robinson, B.H. and Seeman, N.C. 1987. The design of a biochip: A self-assembling molecular-scale memory device. Protein Eng. 1:295-300. Rodbard, D. and Chrambach, A. 1971. Estimation of molecular radius, free mobility, and valence using polyacrylamide gel electrophoresis. Anal. Biochem. 40:95-134.
Mao, C., Sun, W., and Seeman, N.C. 1997. Assembly of Borromean rings from DNA. Nature 386:137-138.
Seeman, N.C. 1982. Nucleic acid junctions and lattices. J. Theor. Biol. 99:237-247.
Mao, C., Sun, W., Shen, Z., and Seeman, N.C. 1999a. A DNA nanomechanical device based on the B-Z transition. Nature 397:144-146.
Seeman, N.C. 1990. De novo design of sequences for nucleic acid structure engineering. J. Biomol. Struct. Dyn.8:573-581. Seeman, N.C. 1992. The design of single-stranded nucleic acid knots. Mol. Eng. 2:297-307.
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Seeman, N.C. 2000. In the nick of space: Generalized nucleic acid complementarity and the development of DNA nanotechnology. Synlett (2000):1536-1548. Seeman, N.C. 2001. DNA nicks and nodes and nanotechnology. NanoLetters 1:22-26. Seeman, N.C. and Kallenbach, N.R. 1983. Design of immobile nucleic acid junctions. J. Biophys. 44:201-209. Seeman, N.C., Rosenberg, J.M., and Rich, A. 1976. Sequence specific recognition of double helical nucleic acids by proteins. Proc. Natl. Acad. Sci. U.S.A. 73:804-808. Seeman, N.C., Maestre, M.F., Ma, R.-I., and Kallenbach, N.R. 1985. Physical characterization of a nucleic acid junction. In Progress in Clinical and Biological Research, Vol. 172A: The Molecular Basis of Cancer (R. Rein, ed.) pp. 99-108. Alan R. Liss, New York. Sha, R., Liu, F., Bruist, M.F., and Seeman, N.C. 1999. Parallel helical domains in DNA branched junctions containing 5′,5′ and 3′,3′ linkages. Biochemistry 38:2832-2841. Sha, R., Liu, F., Millar, D.P., and Seeman, N.C. 2000a. Atomic force microscopy of parallel DNA branched junction arrays. Chem. Biol. 7:743-751. Sha, R., Liu, F., and Seeman, N.C. 2000b. Direct evidence for spontaneous branch migration in antiparallel DNA Holliday junctions. Biochemistry 39:11514-11522. Sha, R., Iwasaki, H., Liu, F., Shinagawa, H., and Seeman, N.C. 2000c. Cleavage of symmetric immobile DNA junctions by Ruv C. Biochemistry 39:11982-11988. Shi, J. and Bergstrom, D.E. 1997. Assembly of novel DNA cycles with rigid tetrahedral linkers. Angew Chem. Int. Ed. Engl. 36:111-113. Sun, W., Mao, C., Liu, F., and Seeman, N.C. 1998. Sequence dependence of branch migratory minima. J. Mol. Biol. 282:59-70. Wang, Y.L., Mueller, J.E., Kemper, B., and Seeman, N.C. 1991. Assembly and characterization of five-arm and six-arm DNA branched junctions. Biochemistry 30:5667-5674.
Wang, H., Di Gate, R.J., and Seeman, N.C. 1998. The construction of an RNA knot and its role in demonstrating that E. coli DNA topoisomerase III is an RNA topoisomerase. In Structure, Motion, Interaction and Expression of Biological Macromolecules (R.H. Sarma and M.H. Sarma, eds.) pp. 103-116. Adenine Press, Schenectady, New York. Wells, A.F. 1977. Three-dimensional nets and polyhedra. John Wiley & Sons, New York. Winfree, E. 1996. On the computational power of DNA annealing and ligation. In DNA Based Computing (R.J. Lipton and E.B. Baum, eds.) pp. 199-219. American Math Society, Providence, R.I. Winfree, E., Liu, F., Wenzler, L.A., and Seeman, N.C. 1998. Design and self-assembly of two-dimensional DNA crystals. Nature 394:539-544. Yan, H., Zhang, X., Shen, Z., and Seeman, N.C. 2002. A robust DNA mechanical device controlled by hybridization topology. Nature 415:6265. Yang, X., Wenzler, L.A., Qi, J., Li, X., and Seeman, N.C. 1998. Ligation of DNA triangles containing double crossover molecules. J. Am. Chem. Soc. 120:9779-9786. Yurke, B., Turberfield, A.J., Mills, A.P. Jr., Simmel, F.C., and Neumann, J.L. 2000. A DNA-fueled molecular machine made of DNA. Nature 406:605-608. Zhang, S. and Seeman, N.C. 1994. Symmetric Holliday junction crossover isomers. J. Mol. Biol. 238:658-668. Zhang, Y. and Seeman, N.C. 1992. A solid-support methodology for the construction of geometrical objects from DNA. J. Am. Chem. Soc. 114:26562663. Zhang, Y. and Seeman, N.C. 1994. The construction of a DNA truncated octahedron. J. Am. Chem. Soc. 116:1661-1669.
Contributed by Nadrian C. Seeman New York University New York, New York
Wang, H., Di Gate, R.J., and Seeman, N.C. 1996. An RNA topoisomerase. Proc. Natl. Acad. Sci. U.S.A. 93:9477-9482.
Key Experimental Approaches in DNA Nanotechnology
Dr. Seeman would like to thank the students, postdocs, and colleagues whose work is cited above; their efforts have created DNA nanotechnology. This work has been supported by grants GM-29554 from the National Institute of General Medical Sciences, N00014-98-1-0093 from the Office of Naval Research, NSFCCR-97-25021 from DARPA/National Science Foundation, and F30602-98-C0148 from the Information Directorate of the Rome, N.Y. Air Force Research Laboratory.
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Preparation of Gold Nanoparticle–DNA Conjugates
UNIT 12.2
This unit outlines the preparation of covalent conjugates between short synthetic oligonucleotides (10 to 100 nucleotides in length) and gold nanoparticles (5 to 50 nm in diameter). These conjugates are formed readily between aqueous gold colloid solutions and synthetic oligonucleotides bearing free thiol or disulfide groups at their ends. The oligonucleotidefunctionalized nanoparticles can then be isolated from starting materials and side products by centrifugation or gel electrophoresis. The two protocols presented here correspond to two distinct types of gold nanoparticle–oligonucleotide conjugates: nanoparticles functionalized with just one or a few oligonucleotide strands (Basic Protocol 1) and nanoparticles functionalized with a dense layer of many oligonucleotide strands (Basic Protocol 2). The physical and chemical properties of these two types of conjugates are different, and the relative stability and utility of the nanoparticles in different environments are discussed below (see Strategic Planning). In addition, the Support Protocol describes a simple synthesis of the aqueous gold colloid used as a starting material in the synthesis of DNA-nanoparticle conjugates. NOTE: Use ultrapure water (e.g., Nanopure; R > 18 MΩ) in all solutions and protocol steps. STRATEGIC PLANNING Properties of Gold Nanoparticle–DNA Conjugates Despite the thematic similarity between the two types of oligonucleotide-nanoparticle conjugates described here, their physical properties are fairly different. Gold nanoparticles with a single or a few attached oligonucleotides, originally described by Alivisatos and Schultz (Alivisatos et al., 1996), have a discrete and characterizeable number of DNA molecules attached to each particle (Zanchet et al., 2001). The remaining surface of these particles is passivated with a monolayer of anionic phosphine molecules, which protect the particles from aggregating with each other and precipitating from solution. Conjugates synthesized by the method described in Basic Protocol 1 are not stable under extended exposure to high temperatures (e.g., >60°C) or in buffers with high ionic strength (e.g., 1 M Na+). In addition, fairly long oligonucleotides (e.g., >50 nucleotides) must be used in order for the different particle-DNA conjugates to be electrophoretically separated from unreacted oligonucleotide and from each other. The synthesis and electrophoretic purification of these conjugates described in Basic Protocol 1 are nearly identical to those reported by Alivisatos and Schultz (Loweth et al., 1999). Gold nanoparticles functionalized with a layer of many attached oligonucleotides, on the other hand, are further stabilized against flocculation and precipitation at high temperature and ionic strength (Storhoff et al., 1998). Using the method described in Basic Protocol 2, reported originally by Mirkin and Letsinger (Mirkin et al., 1996), gold nanoparticles with diameters ranging from a few to tens of nanometers can be conjugated with thiol-terminated oligonucleotides containing 10 to 100 base pairs. The attached oligonucleotides still hybridize selectively to complementary DNA sequences, and the conjugates are stable under high salt concentrations (e.g., ≤2 M Na+) and high temperatures (e.g., for hours at 80°C).
DNA Nanotechnology Contributed by T. Andrew Taton Current Protocols in Nucleic Acid Chemistry (2002) 12.2.1-12.2.12 Copyright © 2002 by John Wiley & Sons, Inc.
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Disulfide- and Thiol-Containing Oligonucleotides The starting oligonucleotide, bearing a disulfide or thiol group at the 3′ or 5′ terminus, can be purchased (e.g., Integrated DNA Technologies, Sigma-Genosys) or synthesized in the laboratory. If the oligonucleotide is purchased, it is extremely important that the product be purified from other thiols, such as dithiothreitol (DTT) or dithioerythritol (DTE), which are sometimes added as stabilizers. This can be achieved via size-exclusion chromatography, HPLC, or preparative gel electrophoresis (e.g., CPMB UNIT 2.5A). If the oligonucleotide is synthesized on a DNA synthesizer (e.g., APPENDIX 3C) using the phosphoramidite method (UNIT 3.3), a disulfide functionality may be generated in a number of ways. 3′-Disulfide-containing oligonucleotides can be synthesized by beginning with a 3′-thiol modifier controlled-pore glass (CPG) or by beginning with a universal support CPG and adding a disulfide modifier phosphoramidite as the first monomer in the sequence. 5′-Disulfide-containing oligonucleotides can be synthesized by ending the synthesis with a disulfide modifier phosphoramidite. In all cases, the 4,4′-dimethoxytrityl protecting group should be removed from the 5′-hydroxy terminus under acidic conditions (80% acetic acid for 30 min; UNIT 10.5) before conjugation of the DNA to the particles. Oligonucleotide synthesis reagents are available from a number of suppliers (e.g., Glen Research, TriLink BioTechnologies, ChemGenes). Oligonucleotides with a free 5′-thiol can also be generated by ending the synthesis with a 5′-tritylthiol modifier phosphoramidite, and these oligonucleotides can be conjugated to particles (Storhoff et al., 1998). However, this protocol requires that the 4,4′-dimethoxytrityl protecting group be removed from the thiol before conjugation. This deprotection procedure is described in Storhoff et al. (1998) and can be found at the Glen Research Web site (http://www.glenresearch.com/ProductFiles/Technical/ tech_questions.html#44). In addition, because free thiol groups are not stable to prolonged storage, the strands should be used immediately after deprotection. BASIC PROTOCOL 1
Preparation of Gold Nanoparticle-DNA Conjugates
PREPARATION OF GOLD NANOPARTICLE–DNA CONJUGATES CONTAINING ONE TO SEVERAL DNA STRANDS PER PARTICLE This protocol describes the synthesis and isolation of gold nanoparticles with one to several synthetic oligodeoxyribonucleotides attached. The synthetic oligonucleotide must be modified to contain a thiol or disulfide group to attach the strand to the gold surface of the particle. A strong, covalent Au-S bond is formed spontaneously between the nanoparticles and DNA by simply mixing the two components. In practice, this procedure generates particles with a statistically distributed number of oligonucleotides attached to each particle; particles with a single attached oligonucleotide can be separated from unmodified and multiply modified particles by preparative horizontal gel electrophoresis. The protocol works best for nanoparticles with diameters between 5 and 20 nm. The resulting DNA-particle conjugates can then be characterized by UV/visible spectroscopy. Materials Oligonucleotide: ∼l mM synthetic 5′- or 3′-disulfide-containing or thiol-containing oligonucleotide (see Strategic Planning), dissolved in water Aqueous gold nanoparticle solution (British Biocell, Ted Pella; or see Support Protocol) Phosphine: 4,4′-(phenylphosphinidene)bis(benzenesulfonic acid), dipotassium salt hydrate (Aldrich), solid and 0.5 M aqueous solution NaCl, solid and 1 M aqueous solution Methanol 5× TBE electrophoresis buffer (APPENDIX 2A) 30% (v/v) glycerol
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UV/vis spectrophotometer Quartz cuvette Razor blade, sterilized Glass-fiber filter paper, 1.2-µm retention (e.g., GF/C; Whatman) Dialysis membrane, MWCO 10,000 (e.g., SpectraPor Biotech RC; Spectrum Laboratories) Centrifugal filter device, 0.45-µm pore size (e.g., Ultrafree-MC; Amicon) Additional reagents and equipment for agarose gel electrophoresis (e.g., CPMB UNIT 2.5A) Quantitate oligonucleotide solution 1. Prepare a 100-fold dilution of an ∼1 mM oligonucleotide solution in water. Synthesized DNA, after deprotection and lyophilization, should be powdery and white and should readily dissolve in water. If necessary, mild heating (∼60°C) and/or brief sonication (1 min) can be used to completely dissolve the DNA. Insoluble material may cause some turbidity but is not a cause for concern. Steps 1 to 3 can be skipped if the concentration of the oligonucleotide solution is already known (e.g., if the modified oligonucleotide was purchased).
2. Use a properly calibrated UV/vis spectrophotometer and quartz cuvette to measure the absorbance at 260 nm (A260) of the diluted oligonucleotide. 3. Calculate the DNA concentration (co in M) in the stock solution using a rearrangement of Beer’s Law: co = (A260 × 100)/(εo × b)
where 100 is the dilution factor, b is the path length of the cuvette (typically 1 cm), and εo is the extinction coefficient of the oligonucleotide at 260 nm. Extinction coefficient calculators for oligonucleotide sequences are available online (see Internet Resources). Alternatively, extinction coefficients can be calculated using nearestneighbor approximations (Breslauer et al., 1986; Sugimoto et al., 1996). See also UNIT 7.3.
Quantitate nanoparticle solution 4. Using the spectrophotometer, measure the A520 of an aliquot of an aqueous gold nanoparticle solution. If the measured absorbance is >1, dilute the aliquot by increments of ten with water until the absorbance is <1. If the nanoparticles are purchased, the supplier may also include a measured concentration of the particle solution, and steps 4 and 5 can be skipped.
5. Estimate the concentration of the nanoparticle solution (cn in M), again using Beer’s Law: cn = ( A520 × d ) /(εn × b)
where εn is the molar extinction coefficient of the nanoparticle at 520 nm and d is the dilution factor used in step 4. Estimated values for εn are sufficient for this protocol and can be obtained from Table 12.2.1. The calculated nanoparticle concentration from this step may range from micromolar to picomolar.
Complex nanoparticles with phosphine 6. To 10 mL gold nanoparticle solution, add 2 mg phosphine (final 0.5 M). Rotate this solution 10 hr at room temperature on an orbital shaker at low speed.
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Table 12.2.1
Estimated Extinction Coefficients for Gold Nanoparticles at 520 nma
Particle diameter (nm)
Extinction coefficient (ε520, M-1cm-1)
2 5 10 20 50
8 × 105 1 × 107 1 × 108 1.2 × 109 3 × 1010
aFrom Yguerabide and Yguerabide (1998); British Biocell International Technical Data Sheet.
The synthesis can be readily scaled up to larger volumes of nanoparticle solution. For best results, divide the total volume into several 1-mL microcentrifuge tubes.
7. Add solid NaCl until the solution turns from deep burgundy to a lighter purple color (usually at ∼2 to 3 M NaCl). 8. Centrifuge the particle solution 30 min at 500 × g, room temperature, to pellet the particles. 9. Discard supernatant and resuspend particles in 1 mL of 0.5 mM phosphine.
cathode
10. Add 0.5 mL methanol to precipitate the particles and centrifuge again 30 min at 500 × g, room temperature.
5 4 3 2
anode
1
Preparation of Gold Nanoparticle-DNA Conjugates
0
Figure 12.2.1 Schematic results for horizontal gel electrophoresis of oligonucleotide-nanoparticle conjugates. The first, third, and fifth wells were loaded with unmodified gold nanoparticle standards, and the second and fourth wells with the nanoparticle/conjugate mixture (see Basic Protocol 1, step 19). The numbers to the right of the gel indicate the number of attached oligonucleotides. The unmodified particles migrate at the zero mark. Each band of successively decreasing mobility corresponds to a nanoparticle with an additional oligonucleotide covalently attached.
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11. Discard supernatant and resuspend particles in 1 mL of 0.5 mM phosphine. 12. Add 110 µL of 5× TBE electrophoresis buffer to this solution (0.5× TBE final). 13. Repeat steps 4 and 5 to determine the concentration of this nanoparticle solution. 14. Calculate the number of moles of nanoparticles in the solution by multiplying this concentration by the solution volume (1.1 mL). Conjugate nanoparticles and oligonucleotide 15. Dilute an aliquot of disulfide-functionalized oligonucleotide solution containing 0.9 mol eq DNA (relative to moles of nanoparticle in step 14) in enough 0.5× TBE to make the solution 50 µM in oligonucleotide. 16. Combine the oligonucleotide and nanoparticle solutions and mix well. 17. Add 0.05 vol of 1 M NaCl to bring the solution to 50 mM NaCl. Place the solution on an orbital shaker at low speed (<1 Hz) and incubate 16 hr at room temperature. Purify conjugates 18. Add 0.2 vol of 30% glycerol (5% final) to the conjugate mixture. 19. Load aliquots of this solution into alternating wells of a horizontal agarose gel in 0.5× TBE. Also load a gel standard, prepared by diluting 20 µL unmodified particles (step 12) with 4 µL of 30% glycerol. A precast horizontal agarose gel (e.g., Ready Agarose Mini-Gel, Bio-Rad) can be used.
20. Electrophorese 90 min at 15 V/cm, or until the red nanoparticle bands approach the bottom of the gel. The resulting gel should look something like the drawing in Figure 12.2.1. Successful resolution of the oligonucleotide-nanoparticle conjugates into discrete bands in the gel is somewhat sequence dependent. See Critical Parameters and Troubleshooting for more details.
21. Disconnect the power to the gel apparatus and lift out the gel tray. With a sterilized razor blade, cut slits in the gel just beneath the bands of oligonucleotide conjugate to be isolated. The bands that migrate just slower than the unmodified particle standards contain particles with only one attached oligonucleotide (Fig. 12.2.1). Nanoparticles with more than one oligonucleotide attached can be specifically isolated by cutting beneath another (higher) band of the agarose gel and electroeluting appropriately.
22. Insert a piece of glass-fiber filter paper and a strip of dialysis membrane into the slit, with the filter paper facing the band. The pink color of the band is easily visualized as it transfers from the gel to the filter paper.
23. Return the gel tray to the electrophoresis apparatus and electrophorese at 15 V/cm until the band has electroeluted into the paper. 24. Remove the filter paper and dialysis membrane together from the gel and place them, filter paper face down, into a 0.45-µm centrifugal filter device. 25. Add 250 µL of 0.5× TBE to the device and centrifuge 3 min at 14,000 × g, room temperature (according to manufacturer’s instructions), to elute the particles into the device receiver. DNA Nanotechnology
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Quantitate and store conjugates 26. Repeat steps 4 and 5 to determine the concentration of particle conjugates. The extinction coefficient of the nanoparticles at 520 nm is not significantly affected by the attached oligonucleotide.
27. Store the conjugates at 4°C in the dark. Conjugates are stable for at least 1 month. Do not freeze nanoparticle conjugate solutions made by this method. BASIC PROTOCOL 2
PREPARATION OF GOLD NANOPARTICLE–DNA CONJUGATES CONTAINING MANY DNA STRANDS PER PARTICLE This protocol describes the synthesis and isolation of gold nanoparticles with a layer of many synthetic oligodeoxyribonucleotides attached. The number of oligonucleotides attached can vary widely; for a detailed discussion of this topic, see Background Information. As in Basic Protocol 1, each oligonucleotide is bound to the particle surface by a strong, covalent Au-S bond. However, the dense oligonucleotide layer formed in these conjugates obviates the need for a protective shell of anionic phosphine ligands. Also, because of the large number of oligonucleotides attached to each particle, it is not possible to separate nanoparticles bearing slightly different numbers of oligonucleotides from each other. As a result, the DNA-particle conjugates can be isolated from free, unreacted thiol-oligonucleotides by centrifugation and characterized by UV/visible spectroscopy. Materials Oligonucleotide: ∼1 mM synthetic 5′- or 3′-disulfide-containing or thiol-containing oligonucleotide (see Strategic Planning), dissolved in water Aqueous gold nanoparticle solution (British Biocell or see Support Protocol) 1 M NaCl 0.1 M sodium phosphate buffer, pH 7 (APPENDIX 2A) 0.1 M NaCl/10 mM sodium phosphate buffer, pH 7 0.3 M NaCl/0.01% (w/v) sodium azide/10 mM sodium phosphate buffer, pH 7 Additional reagents and equipment for quantitating oligonucleotide, nanoparticle, and conjugate solutions (see Basic Protocol 1) CAUTION: Sodium azide is poisonous and explosive in solid form; wear gloves and handle with care. Quantitate oligonucleotide and nanoparticle solutions 1. Quantitate oligonucleotide and aqueous gold nanoparticle solutions (see Basic Protocol 1, steps 1 to 5). Oligonucleotide sequences with a high fraction of guanosine bases form less-stable nanoparticle conjugates than do other sequences. Such conjugates may not survive exposure to the high salt concentrations or centrifugation used to isolate the particles and may frequently precipitate and fuse during preparation. If the concentration of the oligonucleotide and nanoparticle solutions are known (i.e., they were purchased), this step can be skipped.
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Calculate amount of oligonucleotide to attach 2. Calculate the required number of moles of oligonucleotide needed to conjugate to the particles by multiplying the total nanoparticle surface area available for reaction by the expected surface density of nanoparticle-bound oligonucleotide as follows: mol conjugated oligonucleotide = An × cn × D × V = 4 πr 2 × cn × 35 pmol/cm 2 × V
where An is the surface area of the nanoparticle, cn is the concentration of the nanoparticle solution (step 1, in nanoparticles per liter), Do is the oligonucleotide density on each particle (∼35 pmol oligonucleotide/cm2), V is the volume of nanoparticle solution (in L), and r is the radius of the nanoparticles. The value used for Do is based on Demers et al. (2000); see Background Information for more details. As an example of how this equation might be used, the amount of disulfideterminated oligonucleotide needed to functionalize 5 mL of a 17-nM solution of 13-nm-diameter gold nanoparticles would be:
conjugated oligonucleotide −
= 4 π(6.5 nm)2 × 17 nM × 6.02 × 10 23 mol 1 × 35 pmol / cm 2 × 0.005 L = 9.5 nmol Conjugate nanoparticles and oligonucleotide 3. To the solution of gold nanoparticles, add an amount of oligonucleotide solution containing 1.5 times the moles of oligonucleotide calculated in step 2 (i.e., a 0.5-molar excess). The volume of nanoparticle solution is not critical; volumes from microliters to hundreds of milliliters can be used.
4. Rotate the solution 16 hr at room temperature on an orbital shaker at low speed (<1 Hz). This step works best if conducted in a stoppered, glass container that has been covered to prevent unnecessary exposure to light.
5. Add 0.125 vol each of 1 M NaCl and 0.1 M sodium phosphate buffer (0.1 M NaCl and 10 mM phosphate buffer final). Rotate at low speed for 24 hr at room temperature. For larger particles (i.e., >20 nm diameter), the synthesis may be more successful if the transition from pure water to salt solution is made gradually, with multiple small additions of NaCl and buffer solutions.
Table 12.2.2
Centrifugation Conditions for Nanoparticle-Oligonucleotide Conjugates
Particle diameter (nm)
Relative centrifugal force (× g)
Duration (min)
5 10 20 50
64,000 20,000 10,000 4,000
60 30 20 15
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Isolate conjugates 6. Use a benchtop or high-speed centrifuge to centrifuge the suspension to form a red oil of nanoparticles beneath a clear solution of excess oligonucleotide. Choose (or interpolate) the speed and duration of centrifugation from Table 12.2.2, based on the diameter of the nanoparticles. For total solution volumes >1 mL, it is best to divide the volume into several microcentrifuge tubes before centrifuging for optimal separation of the oil from the supernatant. Failure to form stable nanoparticle-DNA conjugates, for any reason, will result in the formation of a solid pellet rather than a red oil.
7. Carefully remove the clear supernatant and resuspend the oil in the same volume of 0.1 M NaCl/10 mM sodium phosphate buffer. 8. Repeat centrifugation, removal of supernatant, and resuspension twice more, but resuspend the last time in 0.3 M NaCl/0.01% sodium azide/10 mM sodium phosphate buffer. CAUTION: Sodium azide is poisonous and explosive in solid form; wear gloves and handle with care.
Quantitate and store conjugates 9. Determine the concentration of the product conjugate solution (see Basic Protocol 1, steps 4 and 5). 10. Store conjugate solution at 4°C in the dark. Conjugates are stable for at least 1 month. Although conjugate solution prepared in this manner can be frozen without effect, do not store the conjugates below 0°C. SUPPORT PROTOCOL
SYNTHESIS OF AQUEOUS CITRATE-PROTECTED GOLD COLLOID This protocol describes a simple synthesis of aqueous gold nanoparticles surrounded by shells of coordinated citrate anions. These particles can be used as starting material in Basic Protocols 1 and 2. A variety of simple protocols for synthesizing gold nanoparticles of various sizes has been reported recently (Brown et al., 2000; Jana et al., 2001). The particles in this protocol are synthesized by reducing hydrogen tetrachloroaurate (HAuCl4) with a citrate salt (Frens, 1973; Grabar et al., 1995). The protocol typically yields particles with diameters of ∼15 nm and narrow size distributions; however, both particle diameter and size dispersity vary from preparation to preparation. In general, both particle diameter and monodispersity are influenced by the type of reducing agent and gold precursor salt used, the concentrations of these reagents, the reaction temperature and duration, and the postreaction workup. The resulting particle diameters and dispersities can be quantified by transmission electron microscopy and statistical analysis of collections of particles. However, because of the reliability of this protocol, such characterization is not strictly necessary for conjugate synthesis. Materials Aqua regia: 3:1 (v/v) concentrated HCl/concentrated HNO3 1 mM HAuCl4 (Aldrich) 38.8 mM sodium citrate (Aldrich)
Preparation of Gold Nanoparticle-DNA Conjugates
1-L round-bottom flask Reflux condenser Heating mantle 0.45-µm nylon filter
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CAUTION: Aqua regia is noxious and extremely caustic. Handle with extreme care in a well-ventilated fume hood. 1. Wash a 1-L round-bottom flask, a reflux condenser, and a large stir-bar with first aqua regia and then thoroughly with water. 2. Assemble the glassware on a heating mantle and a magnetic stirrer, and put the stir-bar in the flask. 3. Charge the flask with 500 mL of 1 mM HAuCl4 and bring the solution to reflux (100°C) with vigorous stirring. 4. Add 50 mL of 38.8 mM sodium citrate all at once. Continue to reflux the solution for 20 min. The sodium citrate solution should be added to the mixture as rapidly as possible. If the addition is done correctly, the solution should turn from yellow to purple and then to deep red.
5. Cool the solution to room temperature and filter through a 0.45-µm nylon filter. 6. Store the solution at room temperature in a glass container in the dark. Gold nanoparticles synthesized and stored in this way are stable almost indefinitely. In fact, gold nanoparticles synthesized by Michael Faraday in 1857 are still on display in the British Museum.
A
B a
Au
a′b′
Au
a
b
Au
b
Au
(complementary template)
Au
Au
a′b′
Au
Au
Au
Au
Figure 12.2.2 Assembly of gold nanoparticle conjugates, functionalized with oligonucleotide sequences a and b, onto complementary oligonucleotide templates a′b′. (A) Nanoparticle conjugates bearing only one oligonucleotide strand assemble selectively into dimers in the presence of the template. More complex structures can be generated from templates with additional recognition segments. (B) Nanoparticle conjugates bearing many oligonucleotide strands assemble into polymeric macrostructures in the presence of the complementary template. The optical changes associated with this polymeric assembly make this system particularly effective as a colorimetric DNA hybridization sensor.
DNA Nanotechnology
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COMMENTARY Background Information As suggested above, the structure, stability, and potential uses of the two types of DNAnanoparticle conjugates described here are very different. Particles with a single attached oligonucleotide, synthesized by the method outlined in Basic Protocol 1, are extremely promising for building tailored nanostructures containing just a few predetermined nanoparticle components. For example, Alivisatos and coworkers have selectively synthesized clusters of two and three nanoparticles in high yields by hybridizing the attached strands to a complementary template (Loweth et al., 1999), as shown in Figure 12.2.2A. However, the instability of these conjugates to the conditions frequently encountered in DNA hybridization protocols limits their application in DNA sequence analysis and in situ hybridization. In addition, gel electrophoresis will not provide adequate separation of conjugates made with short oligonucleotide sequences (i.e., <50 nucleotides) and even some long ones. On the other hand, conjugates with a dense layer of attached oligonucleotides, as described in Basic Protocol 2, have been used in homogeneous and heterogeneous schemes for DNA sequence analysis that require flexible control over hybridization conditions. The number of oligonucleotide strands bound to each particle depends on the surface area of the nanoparticles and the length and sequence of the DNA (Demers et al., 2000). For example, conjugates between 16-nm-diameter particles and thiolmodified 12-mer oligonucleotides were found to bear ∼160 oligonucleotides per particle, but conjugates between the same particles and a 32-mer oligonucleotide bore 70 oligonucleotides per particle (Demers et al., 2000). It is important to note that not all of these oligonucleotides are available for hybridization to complementary DNA strands, due to steric compression and electrostatic repulsion at the
S(CH2)3O
Au + HO(CH2)6SS(CH2)3O
Au DNA
Preparation of Gold Nanoparticle-DNA Conjugates
nanoparticle surface (Levicky et al., 1998). For example, of the 160 12-mer oligonucleotides attached to each 16-nm particle described above, only 6 oligonucleotides (4%) were found to hybridize to complementary oligonucleotides under standard hybridization conditions (Demers et al., 2000). This fraction can be increased by inserting a noncomplementary “tether” sequence between the thiol group and the intended hybridization sequence; adding a 20-adenine-long tether to the 12-mer increased the hybridization efficiency from 4% to 44%. Regardless of nanoparticle size, oligonucleotide length or sequence, the highest surface density of nanoparticle-bound oligonucleotides that has been reported is ∼35 pmol/cm2 (or ∼0.2 oligonucleotides/nm2). This highest value is used as Do in step 2 of Basic Protocol 2 as an estimate of how much DNA will be needed to completely functionalize every particle in solution. If the oligonucleotide being used is particularly dear, the precise density of functionalization can be measured (Demers et al., 2000) and a more accurate value substituted for Do in subsequent preparations. Because each particle synthesized by this method bears multiple strands and thus multiple opportunities for hybridization, template strands bearing sequences complementary to different particles will typically assemble those particles into macroscopic assemblies, as shown in Figure 12.2.2B. The dramitic red-toblue color change that accompanies this particle assembly in solution has been used to identify specific DNA target sequences in solution (Elghanian et al., 1997). In addition, using these conjugates as DNA labels has been found to increase the selectivity of DNA sequence analysis by oligonucleotide arrays (Taton et al., 2000, 2001). Basic Protocols 1 and 2 describe the reaction of a disulfide-modified oligonucleotide with the nanoparticle surface. Oligonucleotide
S(CH2)6OH
Figure 12.2.3 Scheme for conjugation of gold nanoparticles with disulfide-modified oligonucleotides. An organic mercaptoalcohol is also incorporated into the conjugate.
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strands with other types of sulfur modifications, such as alkylthiol and phosphorothioate groups, would also be expected to form stable nanoparticle conjugates. Disulfide-modified starting materials have the advantage that they are stable to long-term storage and that they spontaneously react with the gold nanoparticle surface. (Alkylthiol-modified strands, on the other hand, typically bear trityl protecting groups, which must be removed before reacting with Au.) The reaction of a disulfide-terminated strand actually adds both the oligonucleotide and a mercaptoalcohol molecule to the particle (Figure 12.2.3), but this additional group does not appear to affect the stability or physical properties of the conjugates.
Critical Parameters and Troubleshooting The success of both protocols is somewhat dependent on the sequence used. As noted above, Basic Protocol 1 provides isolatable conjugates only for sequences longer than 50 nucleotides. In addition, secondary structure caused by self-complementarity in the oligonucleotide sequence may result in broadened bands in the preparative agarose gel (and thus poorer separation). Oligonucleotides shorter than seven nucleotides will generally fail to provide stable conjugates by Basic Protocol 2. In addition, guanosine-rich sequences also fail to form stable conjugates by this method. Conjugates synthesized by Basic Protocol 2 are stable to storage over very long periods of time, but bacterial growth and degradation of hybridization activity have been observed in conjugate solutions that did not contain sodium azide as a preservative. Conjugates made from particles >10 nm may settle from solution with time, but can easily be resuspended by swirling the conjugate solution.
Anticipated Results Basic Protocols 1 and 2 both yield red solutions of nanoparticle conjugates in which the hybridization properties of the attached oligonucleotide are fully active. This can be readily confirmed by adding a fluorescently labeled, complementary oligonucleotide to the conjugate solution; gold nanoparticles are exceptional quenchers of fluorescence (Gersten and Nitzan, 1981), and hybridization of the two components leads to a significant decrease in the solution fluorescence. The conjugates can also be analyzed by transmission electron microscopy or by dynamic light scattering to confirm particle sizes and size distributions.
Time Considerations If the disulfide-modified oligonucleotide and nanoparticle solution are prepared or purchased in advance, then conjugates made by either protocol can easily be prepared and isolated within 3 days.
Literature Cited Alivisatos, A.P., Johnsson, K.P., Peng, X., Wilson, T.E., Loweth, C.J., Bruchez, M.P. Jr., and Schultz, P.G. 1996. Organization of ‘nanocrystal molecules’ using DNA. Nature 382:609-611. Breslauer, K.J., Frank, R., Bloecker, H., and Marky, L.A. 1986. Predicting DNA duplex stability from the base sequence. Proc. Natl. Acad. Sci. U.S.A. 83:3746-3750. Brown, K.R., Walter, D.G., and Natan, M.J. 2000. Seeding of colloidal Au nanoparticle solutions. 2. Improved control of particle size and shape. Chem. Mater. 12:306-313. Demers, L.M., Mirkin, C.A., Mucic, R.C., Reynolds, R.A. III, Letsinger, R.L., Elghanian, R., and Viswanadham, G. 2000. A fluorescencebased method for determining the surface coverage and hybridization efficiency of thiol-capped oligonucleotides bound to gold thin films and nanoparticles. Anal. Chem. 72:5535-5541. Elghanian, R., Storhoff, J.J., Mucic, R.C., Letsinger, R.L., and Mirkin, C.A. 1997. Selective colorimetric detection of polynucleotides based on the distance-dependent optical properties of gold nanoparticles. Science 277:1078-1081. Frens, G. 1973. Controlled nucleation for the regulation of the particle size in monodisperse gold suspensions. Nature Phys. Sci. 241:20-22. Gersten, J. and Nitzan, A. 1981. Spectroscopic properties of molecules interacting with small dielectric particles. J. Chem. Phys. 75:1139-1152. Grabar, K.C., Freeman, R.G., Hommer, M.B., and Natan, M.J. 1995. Preparation and characterization of Au colloid monolayers. Anal. Chem. 67:735-743. Jana, N.R., Gearheart, L., and Murphy, C.J. 2001. Seeding growth for size control of 5-40 nm diameter gold nanoparticles. Langmuir 17:67826786. Levicky, R., Herne, T.M., Tarlov, M.J., and Satija, S.K. 1998. Using self-assembly to control the structure of DNA monolayers on gold: A neutron reflectivity study. J. Am. Chem. Soc. 120:97879792. Loweth, C.J., Caldwell, W.B., Peng, X., Alivisatos, A.P., and Schultz, P.G. 1999. DNA-based assembly of gold nanocrystals. Angew. Chem. Int. Ed. Engl. 38:1808-1812. Mirkin, C.A., Letsinger, R.L., Mucic, R.C., and Storhoff, J.J. 1996. A DNA-based method for rationally assembling nanoparticles into macroscopic materials. Nature 382:607-609. DNA Nanotechnology
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Storhoff, J.J., Elghanian, R., Mucic, R.C., Mirkin, C.A., and Letsinger, R.L. 1998. One-pot colorimetric differentiation of polynucleotides with single base imperfections using gold nanoparticle probes. J. Am. Chem. Soc. 120:1959-1964.
Yguerabide, J. and Yguerabide, E.E. 1998. Lightscattering submicroscopic particles as highly fluorescent analogs and their use as tracer labels in clinical and biological applications. Anal. Biochem. 262:137-156.
Sugimoto, N., Nakano, S., Yoneyama, M., and Honda, K. 1996. Improved thermodynamic parameters and helix initiation factor to predict stability of DNA duplexes. Nucl. Acids Res. 24:4501-4505.
Zanchet, D., Micheel, C.M., Parak, W.J., Gerion, D., and Alivisatos, A.P. 2001. Electrophoretic isolation of discrete Au nanocrystal/DNA conjugates. Nano Lett. 1:32-35.
Taton, T.A., Mirkin, C.A., and Letsinger, R.L. 2000. Scanometric DNA detection with nanoparticle probes. Science 289:1757-1760.
Internet Resources
Taton, T.A., Lu, G., and Mirkin, C.A. 2001. Twocolor labeling of oligonucleotide arrays via sizeselective scattering of nanoparticle probes. J. Am. Chem. Soc. 123:5164-5165.
www.basic.nwu.edu/biotools/oligocalc.html Provides an extinction coefficient calculator for oligonucleotides.
Contributed by T. Andrew Taton University of Minnesota Minneapolis, Minnesota
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Synthesis of 5-O-Phosphoramidites with a Photolabile 3-O-Protecting Group
UNIT 12.3
In recent years, DNA microarrays have become a diagnostic assay system of everincreasing importance to a wide range of biotechnical and biomedical applications. Among the various technologies used for their production, the light-controlled in situ synthesis of oligonucleotide arrays has proved to be especially versatile. This methodology has permitted oligonucleotide synthesis in only the 3 →5 direction, however, thus linking the 3 terminus to the solid support (Beier and Hoheisel, 1999). For several enzymatic reactions, and especially for polymerase extension, the availability of the 3 -hydroxyl is a prerequisite. Thus, reversing the direction of synthesis is advantageous or even essential to various chip-based applications, such as highly parallel DNA sequencing or the creation of microarrays containing double-stranded DNA probes. This unit describes the chemical synthesis of phosphoramidite building blocks that carry a photolabile protecting group at the 3 position. These inversely oriented synthons expose a 2-(2-nitrophenyl)propoxycarbonyl (NPPOC) group as the photolabile protecting group of choice. Among other applications, the building blocks can be employed for in situ synthesis of DNA microarrays in which each oligonucleotide is immobilized at the 5 terminus, leaving the 3 terminus available as a polymerase substrate. This unit describes the synthesis of 3 -O-NPPOC-protected nucleosides (see Basic Protocol 1) and 3 -ONPPOC-5 -O-phosphoramidites (see Basic Protocol 2), as well as the synthesis of the required acylating reagent (see Support Protocol). NOTE: All reactions should be performed using dry reagent-grade solvents and should be carried out under an inert atmosphere unless otherwise specified. NOTE: All reactions described involve photolabile compounds. All reactions should be performed and all photolabile derivatives should be stored in the dark (e.g., by covering reaction flasks with aluminum foil).
INTRODUCTION OF THE PHOTOLABILE 2-(2-NITROPHENYL)PROPOXYCARBONYL GROUP INTO 5 -O-DIMETHOXYTRITYLATED NUCLEOSIDES
BASIC PROTOCOL 1
The reaction scheme for the introduction of the photolabile 2-(2nitrophenyl)propoxycarbonyl (NPPOC) group into the 3 -O position is depicted in Figure 12.3.1. Chemical synthesis starts from commercially available 5 -Odimethoyxtrityl (DMTr) nucleosides (S.1a-S.1d). The photolabile NPPOC protecting group is attached to the 3 -O position by a mild acylation reagent that is generated in situ from 2-(2-nitrophenyl)propoxycarbonyl chloride and N-methylimidazole (see Support Protocol). Subsequently, the 5 -O-DMTr moiety is removed by acid treatment without the need to isolate the 5 -O-DMTr-3 -O-NPPOC-protected intermediate nucleoside compounds. The 3 -O-NPPOC-protected derivatives (S.2a-S.2d) are purified by flash chromatography.
Materials Nitrogen source 2-(2-Nitrophenyl)propoxycarbonyl-N-methylimidazolium chloride solution (S.6; see Support Protocol), prepare fresh Dichloromethane, anhydrous Contributed by Markus Beier and J¨org D. Hoheisel Current Protocols in Nucleic Acid Chemistry (2004) 12.3.1-12.3.10 C 2004 by John Wiley & Sons, Inc. Copyright
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Figure 12.3.1 Synthesis of 2-(2-nitrophenyl)propoxycarbonyl (NPPOC)-protected nucleosides (S.2a-S.2d) and phosphoramidites (S.3a-S.3d). S.6 is the acylating reagent shown in Figure 12.3.2. Average yields are given in parentheses. DMTr, 4,4 -dimethoxytrityl.
Synthesis of 5 -OPhosphoramidites with a Photolabile 3 -O-Protection
N-[(4-tert-Butylphenoxy)acetyl]-5 -O-(4,4 -dimethoxytrityl) deoxyribonucleosides: N6 -tac-5 -O-DMTr-2 -deoxyadenosine (S.1a; e.g., Proligo, ChemGenes; see Background Information) N4 -tac-5 -O-DMTr-2 -deoxycytidine (S.1b; e.g., Proligo, ChemGenes; see Background Information) N2 -tac-5 -O-DMTr-2 -deoxyguanosine (S.1c; e.g., Proligo, ChemGenes; see Background Information) 5 -O-DMTr-thymidine (S.1d; e.g., Proligo) Toluene Ethyl acetate 5% (v/v) aqueous HCl Sodium sulfate (Na2 SO4 ) 10% (v/v) trichloroacetic acid in dichloromethane Methanol Saturated aqueous sodium bicarbonate (NaHCO3 ) solution Silica gel (30 to 60 µm; e.g., Baker) for flash chromatography 100-mL three-neck round-bottom flask with drying tube Dropping funnel Balloons 100-mL two-neck flask Thin-layer chromatography (TLC) silica gel 60 plates (Merck) 254-nm UV lamp
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500-mL separatory funnels Rotary evaporator equipped with a water aspirator Additional reagents and equipment for TLC (APPENDIX 3D), flash chromatography (APPENDIX 3E), nuclear magnetic resonance (NMR), and mass spectrometry (MS) Prepare 3 -O-NPPOC-protected deoxyribonucleosides 1. Equip a 100-mL three-neck round-bottom flask with a drying tube, a dropping funnel, and a balloon filled with nitrogen. Under a positive flow of nitrogen, transfer the entire S.6 solution to this flask. 2. Chill solution in an ice bath to 0◦ C. 3. In a 100-mL two-neck flask, also under a positive flow of nitrogen, dissolve one of the following compounds in 40 mL dry dichloromethane:
2.73 g (3.67 mmol) S.1a 2.65 g (3.67 mmol) S.1b 2.79 g (3.67 mmol) S.1c 2.00 g (3.67 mmol) S.1d. 4. Connect the flask to the dropping funnel of the chilled S.6 and add the deoxyribonucleoside dropwise over 10 min, while stirring. Stir overnight at 0◦ C. 5. Check the reaction progress by TLC (APPENDIX 3D) on a silica gel 60 plate. Co-spot the starting material for comparison. For solvents and Rf values, see step 13 (analytical data). Visualize by exposing the plate to a 254-nm UV lamp. If the reaction is not complete, more reagent should be added, and the reaction should be allowed to continue. The amount to add must be determined empirically, although with experience this shouldn’t be necessary.
Work up product 6. Wash the reaction mixture with 100 mL of 5% aqueous HCl and separate the phases using a 500-mL separatory funnel. 7. Dry the organic phase with ∼2 g Na2 SO4 . 8. Add 70 mL of 10% (v/v) trichloroacetic acid in dichloromethane to the organic phase and stir for 2 min. 9. Check the reaction progress by TLC (step 5). If the reaction is not complete, more reagent should be added, and the reaction should be allowed to continue.
10. Wash the red reaction mixture with 100 mL saturated aqueous NaHCO3 solution and separate the phases using a 500-mL separatory funnel. 11. Dry the organic phase with ∼2 g Na2 SO4 and evaporate the organic phase to dryness on a rotary evaporator equipped with a water aspirator.
Purify nucleosides 12. Dissolve the residue in a minimal amount of dichloromethane and purify by flash chromatography (APPENDIX 3E). Prepare a 3 × 30–cm column using 90 g silica gel, and elute (50-mL fractions) with the following solvents (∼2 liters total): Nucleic Acid–Based Microarrays and Nanostructures
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Adenosine derivative: 1:1 (v/v) toluene/ethyl acetate with 0% to 4% (v/v) methanol Cytidine derivative: 1:1 (v/v) toluene/ethyl acetate with 0% to 4% (v/v) methanol Guanosine derivative: 3:1 (v/v) ethyl acetate/methanol Thymidine derivative: 5:4 (v/v) toluene/ethyl acetate with 0% to 10% (v/v) methanol. 13. Identify fractions containing product by TLC and check purity by NMR and MS. N6 -[(4-tert-Butylphenoxy)acetyl]-3 -O-[2-(2-nitrophenyl)propoxycarbonyl]-2 deoxyadenosine (S.2a): Yield, 86%. TLC (toluene/ethyl acetate, 1:1 [v/v]): Rf = 0.17. 1 H NMR (DMSO, δ): 10.78 (br, NH), 8.66 (m, H-C(2), H-C(8)), 7.83 (m, 1H o to NO2 ), 7.70 (m, 1H m to NO2 , 1H p to NO2 ), 7.49 (m, 1H m to NO2 ), 7.30 (m, 2H o to tert-butyl), 6.89 (m, 2H m to tert-butyl), 6.43 (m, H-C(1 )), 5.28 (m, H-C(3 )), 5.14 (m, HO-C(5 )), 4.98 (s, CH2 O), 4.34 (m, OCH2 CH), 4.12 (m, H-C(4 )), 3.60 (m, 2 H-C(5 ), CHCH3 ), 3.02 (m, H-C(2 )), 2.57 (m, H-C(2 )), 1.31 (d, CHCH3 ), 1.24 (s, C(CH3 )3 ). HRMS (FAB, M + H+ ): calcd. for C32 H36 N6 O9 , 649.2621; found, 649.2644. ESI-MS: 649 (M + H+ ), 671 (M + Na+ ), 1297 (2M + H+ ), 1319 (2M + Na+ ). N4 -[(4-tert-Butylphenoxy)acetyl]-3 -O-[2-(2-nitrophenyl)propoxycarbonyl]-2 deoxycytidine (S.2b): Yield, 73%. TLC (toluene/ethyl acetate, 1:4 [v/v]): Rf = 0.50. 1 H NMR (DMSO, δ): 10.87 (br, NH), 8.29 (d, H-C(6)), 7.82 (m, 1H o to NO2 ), 7.69 (m, 1H m to NO2 , 1H p to NO2 ), 7.48 (m, 1H m to NO2 ), 7.29 (m, 2H m to tert-butyl), 7.13 (d, H-C(5)), 6.84 (m, 2H o to tert-butyl), 6.08 (m, H-C(1 )), 5.10 (m, H-C(3 ), HO-C(5 )), 4.77 (s, CH2 O), 4.33 (m, OCH2 CH), 4.10 (m, H-C(4 )), 3.62 (m, 2 H-C(5 )), 3.53 (m, CHCH3 ), 2.48 (m, H-C(2 )), 2.21 (m, H-C(2 )), 1.29 (d, CHCH3 ), 1.24 (s, C(CH3 )3 ). HRMS (FAB, M + H+ ): calcd. for C31 H36 N4 O10 , 625.2509; found, 625.2495. ESI-MS: 625 (M + H+ ), 647 (M + Na+ ), 1249 (2M + H+ ), 1271 (2M + Na+ ). N2 -[(4-tert-Butylphenoxy)acetyl]-3 -O-[2-(2-nitrophenyl)propoxycarbonyl]-2 deoxyguanosine (S.2c): Yield, 81%. TLC (ethyl acetate/methanol, 3:1 [v/v]): Rf = 0.18. 1 H NMR (DMSO, δ): 11.74 (br, 2 NH), 8.23 (2s, H-C(8)), 7.83 (m, 1H o to NO2 ), 7.70 (m, 1H m to NO2 , 1H p to NO2 ), 7.49 (m, 1H m to NO2 ), 7.30 (m, 2H o to tert-butyl), 6.89 (m, 2H m to tert-butyl), 6.18 (m, H-C(1 )), 5.13 (m, H-C(3 )), 5.06 (m, HO-C(5 )), 4.81 (2s, CH2 O), 4.34 (m, OCH2 CH), 4.04 (m, H-C(4 )), 3.55 (m, 2 H-C(5 ), CHCH3 ), 2.83 (m, H-C(2 )), 2.49 (m, H-C(2 )), 1.29 (d, CHCH3 ), 1.25 (s, C(CH3 )3 ). HRMS (FAB, M + H+ ): calcd. for C32 H36 N6 O10 , 665.2570; found, 665.2582. ESI-MS: 665 (M + H+ ), 687 (M + Na+ ), 1329 (2M + H+ ), 1351 (2M + Na+ ). 3 -O-[2-(2-Nitrophenyl)propoxycarbonyl]thymidine (S.2d): Yield, 93%. TLC (toluene/ ethyl acetate, 1:2 [v/v]): Rf = 0.21. 1 H NMR (DMSO, δ): 11.26 (br, NH), 7.82 (m, 1H o to NO2 ), 7.69 (m, H-C(6), 1H m to NO2 , 1H p to NO2 ), 7.49 (m, 1H m to NO2 ), 6.11 (m, H-C(1 )), 5.09 (m, H-C(3 ), HO-C(5 )), 4.33 (m, CHCH2 O), 3.96 (m, H-C(4 )), 3.59 (2m, 2 H-C(5 )), 3.52 (m, CHCH2 O), 2.24 (m, 2 H-C(2 )), 1.77 (2s, CH3 ), 1.29 (d, CHCH3 ). HRMS (FAB, M + H+ ): calcd. for C20 H23 N3 O9 , 450.1512; found, 450.1524. ESI-MS: 450 (M + H+ ), 472 (M + Na+ ), 899 (2M + H+ ), 921 (2M + Na+ ). BASIC PROTOCOL 2
PREPARATION OF 3 -O-[2-(2-NITROPHENYL)PROPOXYCARBONYL]PROTECTED 5 -O-PHOSPHORAMIDITES The reaction scheme for synthesis of the 3 -O-NPPOC-protected phosphoramidite building blocks (S.3a-S.3d) is depicted in Figure 12.3.1. For conversion of the 3 -O-NPPOC nucleosides into the corresponding phosphoramidites, a bifunctional phosphitylating reagent is utilized in combination with pyridine hydrochloride. Alternatively, other activators (e.g., tetrazole) may be employed.
Synthesis of 5 -OPhosphoramidites with a Photolabile 3 -O-Protection
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Materials Nitrogen source 3 -O-NPPOC-protected nucleosides (S.2a-S.2d; see Basic Protocol 1) Acetonitrile, anhydrous 2-Cyanoethyl-N,N,N,N-tetraisopropylphosphorodiamidite 0.5 M pyridine hydrochloride in anhydrous acetonitrile, dried over 4 Å molecular sieves Toluene Ethyl acetate Dichloromethane, anhydrous Saturated aqueous sodium bicarbonate (NaHCO3 ) solution Sodium sulfate (Na2 SO4 ) Silica gel for flash chromatography (30 to 60 µm; Baker) 100-mL three-neck round-bottom flask with drying tube Dropping funnel Balloons 100-mL two-neck flask Thin-layer chromatography (TLC) silica gel 60 plates (Merck) 254-nm UV lamp 500-mL separatory funnel Rotary evaporator equipped with a water aspirator Additional reagents and equipment for TLC (APPENDIX 3D), flash chromatography (APPENDIX 3E), nuclear magnetic resonance (NMR), and mass spectrometry (MS) 1. Connect a 100-mL three-neck round-bottom flask equipped with a drying tube to a dropping funnel and a balloon filled with nitrogen. Under a positive flow of nitrogen, dissolve one of the following 3 -O-NPPOC-protected nucleosides in 15 mL dry acetonitrile:
1.90 g (2.93 mmol) S.2a 1.51 g (2.41 mmol) S.2b 1.0 g (1.5 mmol) S.2c 1.53 g (3.4 mmol) S.2d. 2. Chill solution in an ice bath to 0◦ C. 3. In a 100-mL two-neck flask, also under a positive flow of nitrogen, mix the following amounts of 2-cyanoethyl-N,N,N,N-tetraisopropylphosphorodiamidite (A) and a dry solution of 0.5 M pyridine hydrochloride in acetonitrile (B):
for S.2a derivative: 1.2 mL A plus 2.9 mL B for S.2b derivative: 0.9 mL A plus 2.6 mL B for S.2c derivative: 0.9 mL A plus 1.75 mL B for S.2d derivative: 1.2 mL A plus 3.4 mL B. 4. Connect the flask, under a positive stream of nitrogen, to the dropping funnel of the three-neck round-bottom flask. Add the phosphitylating solution within 10 min dropwise to the chilled nucleoside reaction solution. Stir 1 hr at 0◦ C. 5. Check the reaction progress by TLC (APPENDIX 3D) on a silica gel 60 plate. Visualize by exposing the plate to a 254-nm UV lamp. For solvents and Rf values, see step 9 (analytical data).
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If the reaction is not complete, more reagent should be added, and the reaction should be allowed to continue. The amount to add must be determined empirically, although with experience this shouldn’t be necessary.
6. Dilute reaction mixture with 100 mL dichloromethane. Wash with 100 mL saturated aqueous NaHCO3 solution and separate the phases using a 500-mL separatory funnel. 7. Dry the organic phase with ∼2 g Na2 SO4 and evaporate the organic phase to dryness on a rotary evaporator equipped with a water aspirator. 8. Dissolve the residue in a minimal amount of dichloromethane and purify by flash chromatography (APPENDIX 3E). Prepare a 2 × 20–cm column using 25 g silica gel, and elute with (20- to 25-mL fractions) 0% to 30% (v/v) ethyl acetate/toluene (∼500mL total). 9. Identify fractions containing product by TLC and check purity by NMR and MS. N6 -[(4-tert-Butylphenoxy)acetyl]-3 -O-[2-(2-nitrophenyl)propoxycarbonyl]-2 deoxyadenosine-5 -O-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite] (S.3a): Yield, 65%. TLC (toluene/ethyl acetate, 1:1 [v/v]) Rf = 0.53. 1 H NMR (DMSO, δ ): 11.70 (br, NH), 8.64 (m, H-C(2), H-C(8)), 7.83 (m, 1H o to NO2 ), 7.71 (m, 1H m to NO2 , 1H p to NO2 ), 7.48 (m, 1H m to NO2 ), 7.29 (m, 2H o to tert-butyl), 6.88 (m, 2H m to tert-butyl), 6.45 (m, H-C(1 )), 5.33 (m, H-C(3 )), 4.98 (s, OCH2 ), 4.36 (m, OCH2 CH2 ), 4.24 (m, H-C(4 )), 4.78 (m, 2 H-C(5 ), OCH2 CH2 CN), 3.53 (m, 3 CHCH3 ), 3.12 (m, H-C(2 )), 2.74 (m, CH2 CH2 CN), 2.60 (m, H-C(2 )), 1.30 (d, CH3 ), 1.11 (m, 7 CH3 ). 31 P NMR (DMSO, δ ): 149.24, 149.20, 149.16. HRMS (FAB, M + H+ ): calcd. for C41 H53 N8 O10 P, 849.3700; found, 849.3723. ESI-MS: 849 (M + H+ ), 871 (M + Na+ ), 1697 (2M + H+ ), 1719 (2M + Na+ ). N4 -[(4-tert-Butylphenoxy)acetyl]-3 -O-[2-(2-nitrophenyl)propoxycarbonyl]-2 deoxycytidine-5 -O-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite] (S.3b): Yield, 81%. TLC (toluene/ethyl acetate, 1:1[v/v]) Rf = 0.43. 1 H NMR (DMSO, δ ): 10.90 (br, NH), 8.14 (m, H-C(6)), 7.82 (m, 1H o to NO2 ), 7.69 (m, 1H m to NO2 , 1H p to NO2 ), 7.48 (m, 1H m to NO2 ), 7.29 (m, 2H o to tert-butyl), 7.14 (m, H-C(5)), 6.83 (m, 2H m to tert-butyl), 6.07 (m, H-C(1 )), 5.10 (m, H-C(3 )), 4.77 (s, OCH2 ), 4.30 (m, OCH2 CH2 , H-C(4 )), 3.75 (m, 2 H-C(5 ),OCH2 CH2 CN), 3.55 (m, 3 CHCH3 ), 2.73 (m, CH2 CH2 CN), 2.55 (m, H-C(2 )), 2.25 (m, H-C(2 )), 1.29 (m, CH3 ), 1.15 (m, 7 CH3 ). 31 P NMR (DMSO, δ): 149.35. HRMS (FAB, M + H+ ): calcd. for C40 H53 N6 O11 P, 825.3587; found, 825.3568. ESI-MS: 825 (M + H+ ), 847 (M + Na+ ), 1649 (2M + H+ ), 1671 (2M + Na+ ). N2 -[(4-tert-Butylphenoxy)acetyl]-3 -O-[2-(2-nitrophenyl)propoxycarbonyl]-2 deoxyguanosine-5 -O-[(2-cyanoethyl)-N,N-diisopropylphosphoramidite] (S.3c): Yield, 91%. TLC (toluene/ethyl acetate/methanol, 5:4:1 [v/v/v]) Rf = 0.39. 1 H NMR (DMSO, δ): 11.45 (br, 2 NH), 8.15 (m, H-C(8)), 7.83 (m, 1H o to NO2 ), 7.70 (m, 1H m to NO2 , 1H p to NO2 ), 7.49 (m, 1H m to NO2 ), 7.29 (m, 2H o to tert-butyl), 6.88 (m, 2H m to tert-butyl), 6.19 (m, H-C(1 )), 5.23 (m, H-C(3 )), 4.79 (m, OCH2 ), 4.36 (m, OCH2 CH2 ), 4.20 (m, H-C(4 )), 3.73 (m, 2 H-C(5 ), OCH2 CH2 CN), 3.55 (m, 3 CHCH3 ), 2.87 (m, CH2 CH2 CN), 2.76 (m, H-C(2 )), 2.57 (m, H-C(2 )), 1.30 (d, CH3 ), 1.15 (m, 7 CH3 ). 31 P NMR (DMSO, δ): 149.46, 149.41. HRMS (FAB, M + H+ ): calcd. for C41 H53 N8 O11 P, 865.3649; found, 865.3660. ESI-MS: 865 (M + H+ ), 887 (M + Na+ ), 1751 (2M + Na+ ).
Synthesis of 5 -OPhosphoramidites with a Photolabile 3 -O-Protection
3 -O-[2-(2-Nitrophenyl)propoxycarbonyl]thymidine-5 -O-[(2-cyanoethyl)-N,Ndiisopropyl phosphoramidite] (S.3d): Yield, 88%. TLC (toluene/ethyl acetate, 1:1 [v/v]) Rf = 0.37. 1 H NMR (DMSO, δ): 11.26 (br, NH), 7.82 (m, 1H o to NO2 ), 7.69 (m, 1H m to NO2 , 1H p to NO2 ), 7.55-7.47 (m, H-C(5), 1H m to NO2 ), 6.08 (m, H-C(1 )), 5.09 (m, H-C(3 )), 4.35-4.27 (m, OCH2 CH2 ), 4.12 (m, H-C(4 )), 3.83-3.70 (m, 2 H-C(5 ), OCH2 CH2 CN), 3.59-3.49 (m, 3 CHCH3 ), 2.75 (m, CH2 CH2 CN), 2.29 (m, 2 H-C(2 )), 1.78 (m, CH3 ), 1.28 (d, CH3 ), 1.24-1.09 (m, 7 CH3 ). 31 P NMR (DMSO, δ): 149.36, 149.33, 149.29. HRMS (FAB, M + H+ ): calcd. for C29 H40 N5 O10 P, 650.2590; found, 650.2576. ESI-MS: 649 (M + H+ ), 672 (M + Na+ ), 1321 (2M + Na+ ).
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PREPARATION OF 2-(2-NITROPHENYL)PROPOXYCARBONYL-NMETHYLIMIDAZOLIUM CHLORIDE
SUPPORT PROTOCOL
The preparation of the acylating reagent used in Basic Protocol 1 is illustrated in Figure 12.3.2. CAUTION: Trichloromethyl chloroformate (diphosgene) is a hazardous reagent; upon reaction, it decomposes into phosgene gas, which is extremely toxic. Therefore, all preparations should be performed in a well-ventilated fume hood, and extreme precautions should be taken to avoid inhalation of diphosgene or phosgene fumes. NOTE: S.4 and subsequent compounds are light sensitive and thus require special handling to prevent photolysis.
Materials Nitrogen source Tetrahydrofuran (THF), dry Trichloromethyl chloroformate (diphosgene) (e.g., Fluka, Sigma) 2-(2-Nitrophenyl)propanol (S.4; see Uhlmann and Pfleiderer, 1981) N-Methylmorpholine Toluene Ethyl acetate Liquid nitrogen Methanolic KOH solution: 5% (w/v) KOH in MeOH, prepare fresh N-Methylimidazole Dichloromethane, dry Molecular sieves, 4 Å 100- and 250-mL three-neck round-bottom flasks Drying tubes and septa for 100- and 250-mL flasks Dropping funnel for 250-mL flasks Balloons 20-mL syringes and 18- to 22-G needles Thin-layer chromatography (TLC) silica gel 60 plates (Merck) Glass frit 250- and 500-mL two-neck Schlenk-type flasks with glass stoppers (neoLab)
Figure 12.3.2 Synthesis of the acylating reagent 2-(2-nitrophenyl)propoxycarbonyl-Nmethylimidazolium chloride. THF, tetrahydrofuran.
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Vacuum tubing Gas inlets Vacuum pump with appropriate tubing 2-mL gas-tight syringe with disposable needles Additional reagents and equipment for TLC (APPENDIX 3D) Prepare S.5 1. Connect a 250-mL three-neck round-bottom flask with a drying tube, a dropping funnel, and a septum. Flush the apparatus with nitrogen. 2. Under a positive flow of nitrogen from a balloon filled with nitrogen, add 25 mL dry THF and cool to 0◦ C in an ice bath. 3. Using a 20-mL syringe and an 18- to 22-G needle, add 14.2 mL (118 mmol) diphosgene through the septum. 4. In a separate flask, under a positive flow of nitrogen, dissolve 20.8 g (114 mmol) S.4 and 12.5 mL (114 mmol) N-methylmorpholine in 25 mL dry THF. 5. Connect this flask, under a positive flow of nitrogen, to the dropping funnel and add solution dropwise over 30 min to chilled diphosgene solution. Stir 1 hr at 0◦ C. A white solid precipitate is formed.
6. Check the reaction progress by TLC (APPENDIX 3D) on silica gel 60 plates using 1:1 (v/v) toluene/ethyl acetate (S.4 Rf = 0.2; S.5 Rf = 0.7) If the reaction is not complete, allow to continue for an additional 30 min and check again. If it is still not complete, add more diphosgene. The amount to add must be determined empirically, although with experience this shouldn’t be necessary.
7. Remove ice bath and let the reaction slowly warm to ambient temperature while stirring. Continue stirring 1 hr at ambient temperature. 8. Attach a glass frit to a 250-mL two-neck Schlenk-type flask and add a magnetic stir-bar. Connect this flask with vacuum tubing to a 500-mL two-neck Schlenk flask (with stir bar) connected to a gas inlet. Using a vacuum pump attached by appropriate tubing, apply a slight vacuum to the second Schlenk flask and cool this flask in liquid nitrogen. 9. Filter the reaction mixture into the first Schlenk flask by extending the slight vacuum to the first Schlenk flask. 10. Wash precipitate on the glass frit with dry THF. 11. Remove glass frit and replace with a glass stopper. 12. Apply an initial modest vacuum to the whole apparatus to remove liquid from the filtrate, while stirring, until the vigorous bubbling ceases. CAUTION: If the initial vacuum is too high, most of the phosgene gas will not be condensed within the second Schlenk flask, but will be sucked into the vacuum pump.
13. Increase the vacuum and remove all liquid from the filtrate.
Synthesis of 5 -OPhosphoramidites with a Photolabile 3 -O-Protection
14. Finally, apply high vacuum overnight to remove the last remaining solvent residues. The product, 2-(2-nitrophenyl)propyl carbonochloridate (S.5) is a brown oil (23.3 g; 84%) that can be used without further purification.
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15. To destroy the remaining phosgene gas, which was condensed in the second flask by cooling with liquid nitrogen, slowly add methanolic KOH solution to the condensate while stirring vigorously.
Convert to S.6 16. Connect a 100-mL three-neck round-bottom flask to a gas inlet, a drying tube, and a septum. Flush the apparatus with a positive flow of nitrogen. 17. Under a continuous flow of nitrogen, dissolve in the flask 1.24 mL (15.5 mmol) N-methylimidazole in 40 mL dry dichloromethane over 1 to 2 g of 4 Å molecular sieves. Chill to 0◦ C in an ice bath. 18. Load a 2-mL gas-tight syringe with 1.07 mL (4.4 mmol) S.5 and add dropwise within 10 min through the septum to the chilled reaction solution. Stir an additional 30 min at 0◦ C. This reaction solution, 2-(2-nitrophenyl)propoxycarbonyl-N-methylimidazolium chloride (S.6), is obtained in quantitative yield and is used directly to introduce the photolabile protecting group to the 3 -O position as described (see Basic Protocol 1). It must be used immediately.
COMMENTARY Background Information
Photolabile 3 -O-[2-(2-nitrophenyl)propoxycarbonyl]-protected 5 -O-phosphoramidites (S.3a-S.3d) are synthesized mainly for an alternative mode of light-directed production of oligonucleotide arrays (Beier et al., 2001). Because of their characteristics, light-controlled in situ DNA synthesis— performed by photolithography (Pease et al., 1994) or micro-mirror control (Singh-Gasson et al., 1999; Baum et al., 2003)—occurs in a 5 →3 direction, conforming to the orientation of enzymatic synthesis. The resulting oligonucleotides are attached to the surface via their 5 termini, leaving the 3 -hydroxyls available as substrates for enzymatic reactions, such as primer extension upon hybridization of a DNA template. The production of such oligonucleotide chips adds new procedural avenues to the growing number of applications of DNA microarrays. The starting monomers are commercially available 5 -O-DMTr-protected nucleosides. The derivatives described here utilize a labile 4-(tert-butylphenoxy)acetyl (tac) protecting group for protection of the exocyclic amino groups (Fig. 12.3.3; Sinha et al., 1993). This
Figure 12.3.3
results in a shorter and milder deprotection procedure when the 3 -O-NPPOC-protected 5 -O-phosphoramidites are employed for in situ synthesis of DNA microarrays (Beier and Hoheisel, 2000). N-tac-5 -O-DMTr-protected nucleosides are available from Proligo and ChemGenes (under the name tBPAC instead of tac). The authors have also had excellent success using the phenoxyacetyl (pac) protecting group to synthesize standard NPPOCprotected 3 -O-phosphoramidites for synthesis in the standard direction. It is reasonable to conclude that working with pac protection would give good results with the reverse 5 -Ophosphoramidites, although this has not been tested. The N-pac-5 -O-DMTr-protected nucleosides are available from ChemGenes. The synthesis of the photolabile NPPOC protecting group (see Support Protocol) starts out from 2-(2-nitrophenyl)propanol (S.4). The 2-(2-nitrophenyl)propyl carbonochloridate (S.5) is preferentially synthesized employing the less-dangerous liquid diphosgene reagent (Giegrich et al., 1998) instead of phosgene gas (Hasan et al., 1997). The mild acylating reagent 2-(2-nitrophenyl)propoxycarbonylN-methylimidazolium chloride (S.6) is
Structure of the 4-(tert-butylphenoxy)acetyl group used for N protection.
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generated from S.5 and N-methylimidazole. The introduction of the NPPOC protecting group into the 3 -O position is best performed by employing S.6. When the carbonochloridate (S.5) is employed directly, the reaction produces several side products and the yields drop significantly. The addition of molecular sieves to the reaction increases the yields further. Having installed the NPPOC group, the DMTr group is cleaved by a short acid treatment. The straightforward synthesis of the phosphoramidites (S.3a–d) utilizes a mild bifunctional phosphitylating reagent combined with an acidic activator. Instead of employing tetrazole, the authors prefer to use pyridine hydrochloride as the activator of choice because it is easier to handle and gives slightly higher yields (Beier and Pfleiderer, 1999). In the analysis, the phosphoramidites showed additional peaks in their nuclear magnetic resonance (NMR) spectra. These were due to the additional chiral center within the NPPOC moiety.
Critical Parameters Keeping moisture from entering the reactions is the most critical factor that can significantly affect yield and quality. S.6 (see Support Protocol) must be prepared immediately before use.
Anticipated Results Average yields of the various products are shown in Figure 12.3.1. All products are seen as single spots by TLC, with no detectable sideproducts (peaks) in 1 H NMR or mass spectra. If the products are not sufficiently pure, use flash chromatography (APPENDIX 3E) to purify them further.
Time Considerations Each of the intermediates and products described can be prepared and isolated within 1 or 2 days.
Literature Cited
Synthesis of 5 -OPhosphoramidites with a Photolabile 3 -O-Protection
Baum, M., Bielau, S., Rittner, N., Schmid, K., Eggelbusch, K., Dahms, M., Schlauersbach, A., Tahedl, H., Beier, M., G¨uimil, R., Scheffler, M., Hermann, C., Funk, J.M., Wixmerten, A., Rebscher, H., H¨onig, M., Andreae, C., B¨uchner, D., Moschel, E., Glathe, A., J¨ager, E., Thom, M., Greil, A., Bestvater, F., Obermeier, F., Burgmaier, J., Thome, K., Weichert, S., Hein, S.,
Binnewies, T., Foitzik, V., M¨uller, M., St¨ahler, C.F., and St¨ahler, P.F. 2003. Validation of a novel, fully integrated and flexible microarray benchtop facility for gene expression profiling. Nucl. Acids Res. 31:e151. Beier, M. and Hoheisel, J.D. 1999. Versatile derivatisation of solid support media for covalent bonding on DNA-microchips. Nucl. Acids Res. 27:1970-1977. Beier, M. and Hoheisel, J.D. 2000. Production by quantitative photolithographic synthesis of individually quality checked DNA microarrays. Nucl. Acids Res. 28:e11. Beier, M. and Pfleiderer, W. 1999. Pyridinium salts—an effective class of catalysts for oligonucleotide synthesis. Helv. Chim. Acta 82:879887. Beier, M., Stephan, A., and Hoheisel, J.D. 2001. Synthesis of photolabile 5 -O-phosphoramidites for the production of microarrays of inversely oriented oligonucleotides. Helv. Chim. Acta 84:2089-2095. Giegrich, H., Eisele-B¨uhler, S., Hermann, C., Kvasyuk, E., Charubala, R., and Pfleiderer, W. 1998. New photolabile protecting groups in nucleoside and nucleotide chemistry—synthesis, cleavage mechanisms and applications. Nucleosides Nucleotides 17:1987-1996. Hasan, A., Stengele, K.P., Giegrich, H., Cornwell, P., Isham, K.I., Sachleben, R., Pfleiderer, W., and Foote, R.S. 1997. Photolabile protecting groups for nucleosides: Synthesis and photodeprotection rates. Tetrahedron 53:4247-4264. Pease, A.C., Solas, D., Sullivan, E.J., Cronin, M.T., Holmes, C.P., and Fodor, S.P.A. 1994. Lightgenerated oligonucleotide arrays for rapid DNA sequence analysis. Proc. Natl. Acad. Sci. U.S.A. 91:5022-5026. Singh-Gasson, S., Green, R.D., Yue, Y.J., Nelson, C., Blattner, F., Sussman, M.R., and Cerrina, F. 1999. Maskless fabrication of light-directed oligonucleotide microarrays using a digital micromirror array. Nat. Biotechnol. 17:974-978. Sinha, N.D., Davis, P., Usman, N., Perez, J., Hodge, R., Kremsky, J., and Casale, R. 1993. Labile exocyclic amine protection of nucleosides in DNA, RNA, and oligonucleotide analog synthesis facilitating N-deacylation, minimizing depurination and chain degradation. Biochimie 75:1323. Uhlmann, E., and Pfleiderer, W. 1981. Helv. Chim. Acta 64:1688-1704.
Contributed by Markus Beier and J¨org D. Hoheisel Deutsches Krebsforschungszentrum Heidelberg, Germany
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Derivatization of Glass and Polypropylene Surfaces
UNIT 12.4
This unit describes the derivatization of solid support media with a versatile linker system. The procedure permits the production of various linker types whose characteristics can be tailored to the requirements of the eventual application. The protocols described here have been used extensively on both nonporous (glass, polypropylene foil) and porous (nylon or polypropylene membrane) substrates. If required, a dendrimeric structure can be formed, thereby increasing the loading capacity of the surface in a stepwise fashion, which is especially useful for glass surfaces. A prerequisite for derivatization is the presence of anchoring groups (hydroxyl or amino groups) on the support surface. These anchoring groups may be intrinsic to the medium (e.g., amino functions on nylon) or may be introduced by chemical modification. Standard modifications include silanization reactions on glass (see Support Protocol 1) or plasma-amination on polypropylene. The latter process is a standard surface modification technique in the plastics industry. The authors obtained plasma-aminated material from AIMS Scientific Products, which processed polypropylene foils provided by the authors. Generally, the linker synthesis described below (see Basic Protocol) consists of a two-step process (Fig. 12.4.1) that may be repeated in an iterative way. In a first step, the anchoring groups on the surface are activated by an acylation reagent (e.g., acryloyl chloride, 4nitrophenyl chloroformate). Subsequently, in a second step, these activated intermediates are reacted with an amine component. Preferably, a polyamine is employed, since this increases the surface loading. By careful selection of the polyamine component, the surface properties (e.g., linker length, hydrophobicity, charge) may be customized and adapted to specific requirements. The performance of the derivatization process can be
Figure 12.4.1 Linker synthesis on an aminated support. A two-step process of acylation (activation; top) and reaction with various amines (bottom) is used to derivatize the solid support with an appropriate linker. The process is depicted using activation with (a) 4-nitrophenyl chloroformate and (b) acryloyl chloride.
Contributed by Markus Beier and J¨org D. Hoheisel Current Protocols in Nucleic Acid Chemistry (2004) 12.4.1-12.4.8 C 2004 by John Wiley & Sons, Inc. Copyright
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monitored by analyzing the density of amino functions on the surface via bromphenol blue staining (see Support Protocol 2). For the immobilization of existing nucleic acids, an additional activation step is required to generate a covalent linkage between biomolecule and linker. For that purpose, bifunctional cross-linking reagents are used (see Support Protocol 3). In situ synthesis of DNA microarrays does not require such activation but can proceed directly, since the terminal groups of the linker serve as starting points for oligonucleotide synthesis. NOTE: Use deionized, distilled water in all recipes and protocol steps. Use dry reagentgrade solvents for all reactions. BASIC PROTOCOL
DERIVATIZATION OF SOLID SUPPORT The process described below permits the modification of glass and polypropylene surfaces forming a linker of dendrimeric structure. The derivatization is used either for attaching prefabricated DNA-oligonucleotides, PCR products, and peptide nucleic acid (PNA) oligomers, or for the in situ synthesis of DNA microarrays (UNIT 12.3).
Materials Acetone Nitrogen Diisopropylethylamine (DIPEA) Acetonitrile, anhydrous 4-Nitrophenyl chloroformate or acryloyl chloride Dichloroethane, dry Amine compound—e.g., tetraethylenepentamine; 1,4-bis-(3-aminopropoxy)butane; 4-aminomethyl-1,8,-octadiamine; 4,7,10-trioxa-1,13-tridecandiamine; N,N-dimethyl-1,6-hexadiamine; 2-(2-aminoethoxy)ethanol; jeffamine; 3-amino-1,2-propandiol Dimethylformamide (DMF), dry and amine free Methanol, reagent grade 18 × 8–cm polypropylene vessels with tight-fitting lids Solid support medium (e.g., glass, polypropylene foil, nylon membrane, polypropylene membrane) with anchoring groups in place on surface (e.g., see Support Protocol 1 for silanizing glass) Orbital shaker Activate solid support 1. Clean an 18 × 8–cm polypropylene vessel carefully with acetone and dry thoroughly. Any polypropylene vessel can be used if it is of sufficient size to act as a reaction container for the derivatization of the support medium in bulk. It must also have a lid that fits tightly to prevent moisture from entering the reaction chamber. An 18 × 8–cm container can be used to process up to six microscope slides in parallel or one ∼17 × 7.5–cm sheet of polypropylene foil, and will have a wash volume of ∼30 to 40 ml.
2. Flush the container with nitrogen and fill with a solution of 171 µl (1 mmol) DIPEA in 30 mL anhydrous acetonitrile. 3. Add one of the following: Derivatization of Glass and Polypropylene Surfaces
192 mg (1 mmol) 4-nitrophenyl chloroformate 81 µl (1 mmol) acryloyl chloride. See Background Information for a discussion of these activating agents.
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4. Add the solid support medium (with anchoring groups) and close the lid carefully to seal the reaction chamber. 5. Incubate 2 hr at ambient temperature on an orbital shaker. 6. Remove the reaction solution, wash the supports thoroughly (two times) with 30 to 40 ml dichloroethane, and then dry under a flush of nitrogen. It is not advisable to store the activated supports. It is best to proceed directly to the reaction with the amine component.
React with amine 7. Clean a second polypropylene container carefully with acetone and dry thoroughly. 8. Flush the container with nitrogen and fill it with a solution of 1 mmol of the desired amine component in 30 mL anhydrous amine-free DMF. In principle, any amine reagent can be employed in this reaction (see Commentary and materials section for suggestions).
9. Add the activated solid support medium to the amine reaction solution and close the lid carefully to seal the reaction chamber. 10. Incubate at ambient temperature on an orbital shaker for 12 hr if 4-nitrophenyl chloroformate was used for activation or 24 hr if acryloyl chloride was used.
NH2
O step 1
i -Pr2NEt CH2Cl2
step 2
DMF
step 3
i -Pr2NEt CH2Cl2
step 4
DMF
Cl
C
H N
H2 N
N H
H N
NH2
O Cl
C
H2 N
O
O
NH2
dendrimeric linker system Figure 12.4.2 By an iterative process of activation and amination, linker structures of different complexity and diverse inherent characteristics can be synthesized.
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The nucleophilic substitution of the 4-nitrophenyl moiety by an amine produces a colored group, which allows monitoring of the reaction by visual inspection.
11. Remove the reaction solution and wash the support media thoroughly with DMF, then methanol, and finally acetone. Dry under a flush of nitrogen. 12. If desired, perform another round of activation and amine reaction in order to increase surface loading/dendrimeric character of the support or to alter the surface properties. For the derivatization of microarray glass surfaces used for the immobilization of nucleic acids or in situ synthesis of DNA oligonucleotides, the best results are obtained with two reaction cycles consisting of acryloyl chloride (activation), tetraethylenepentamine (amine derivatization), acryloyl chloride (activation), and 1,4-bis-(aminopropoxy)butane (amine derivatization). This is illustrated in Figure 12.4.2.
13. Store derivatized support in a dry place up to several weeks at 4◦ C SUPPORT PROTOCOL 1
SILANIZATION OF GLASS SLIDES Prior to the actual silanization, the glass is etched, thereby improving the overall quality of the process. Both processes are described below.
Materials 10% (w/v) aqueous NaOH 1% (v/v) aqueous HCl Methanol, reagent grade 3% (w/v) aminopropyltrimethoxysilane in 95% (v/v) methanol Nitrogen Glass microscope slides (e.g., Menzel-Gl¨aser, Germany) Several glass vessels with lids (e.g., hematology staining jar) Orbital shaker Polypropylene vessel with lid Bath sonicator 110◦ C oven Etch glass surface 1. Immerse underivatized glass microscope slides in a glass vessel filled with a 10% (w/v) NaOH solution and close the lid. Etching can be performed in the type of glass staining jar frequently used in hematology for labeling experiments. These allow eight to ten slides to be placed standing within the vessel.
2. Shake gently overnight on an orbital shaker. 3. Remove slides from the NaOH bath using tweezers. 4. Wash by immersing the slides in a glass vessel filled with water and shaking briefly (2 to 3 min) on an orbital shaker. 5. Wash in another glass vessel, filled with 1% (v/v) HCl, by shaking 2 to 3 min on an orbital shaker. 6. Wash 2 to 3 min in water.
Derivatization of Glass and Polypropylene Surfaces
7. Immerse the slides in a glass vessel filled with methanol and shake 2 to 3 min on an orbital shaker. Proceed directly to the silanization reaction.
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Silanize etched surface 8. Fill a polypropylene container with 3% (w/v) aminopropyltrimethoxysilane in 95% (v/v) methanol. 9. Immerse etched glass slides in the container so that they just cover the bottom in a single layer. Close the lid. A single layer is used to prevent adherence between the slides.
10. Transfer the container to a bath sonicator. Sonicate 15 min at ambient temperature. CAUTION: The reaction mixture becomes hot during sonication. Open lid carefully.
11. Remove slides with tweezers. 12. Wash by immersing the slides in a glass vessel filled with methanol and shaking 2 to 3 min on an orbital shaker. Use very gentle agitation to prevent the slides from moving and adhering to one another. 13. Wash 2 to 3 min in water on an orbital shaker. 14. Dry slides under a stream of nitrogen gas. 15. Transfer slides to a preheated 110◦ C oven for 15 min. 16. Remove slides from oven and store up to several weeks at a convenient temperature.
QUALITY CONTROL OF DERIVATION REACTIONS To control the efficiency of the derivatization procedure, a bromphenol blue staining reaction is employed. The quality is assessed by measuring the amount of surface-bound amino functions and hence the surface loading. This assay is preferably carried out on a control strip made of the appropriate support material and added to the derivatization reactions. After each amination step, a part of the control strip is removed and analyzed.
SUPPORT PROTOCOL 2
For glass supports, only qualitative assessments can be made because of the low loading capacity of glass. Because the blue color achieved with glass can barely be visualized by eye, a polypropylene control strip is used in the same reactions with the glass slides. If a strong loading (blue color) is achieved on the polypropylene control strip, it can be assumed that the derivatization reaction in general was successful and that the slides are usable. Quantitative assessments can be made with polypropylene foils and membranes, and with nylon membranes, all of which have sufficiently high loading to obtain reliable readings by spectrophotometry. For quantitative purposes, the control strip must be of the identical support medium used for derivatization.
Additional Materials (also see Basic Protocol) Control strip: hydroxylated or aminated solid support (see Basic Protocol 1) Amine-free DMF containing 0.05% bromphenol blue Ethanol, reagent grade 20% piperidine in DMF Spectrophotometer, 605 nm 1. Add a control strip to the reactions described in the Basic Protocol. For quantification, use the identical support material that was derivatized for use. 2. At each step in the derivatization, remove an
∼1-cm2
portion of the control strip.
For quantification and comparison purposes, always take identical portions.
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3. Transfer to a new vessel and wash four times with 10 to 15 mL amine-free DMF. 4. Transfer to 2 mL amine-free DMF containing 0.05% (w/v) bromphenol blue and incubate 5 to 10 min with gentle agitation. In the presence of amino functions on the support surface, the support will turn blue and the staining solution will turn yellow.
5. Transfer to a new vessel and wash three times with ethanol to remove any residual stain. 6. Transfer to 20% piperidine in amine-free DMF to destain. Agitate gently until the blue color of the control strip is completely removed. 7. Collect the destain solution and analyze in a spectrophotometer at 605 nm (ε605 = 95,000 M−1 cm−1 ). If necessary, dilute solution with 20% piperidine in DMF. 8. Compare the blue color/absorption of different rounds of derivatization to monitor surface loading. For polypropylene, the actual number of amino functions per area can be calculated. For glass, only relative measurements across reactions are possible. SUPPORT PROTOCOL 3
IMMOBILIZATION OF 5 -AMINO-LABELED NUCLEIC ACID COMPOUNDS While in situ DNA synthesis can proceed directly on the linker resulting from two subsequent reaction cycles with acryloyl chloride, tetraethylenepentamine, acryloyl chloride, and 1,4-bis-(aminopropoxy)butane (see Basic Protocol), an additional activation step is required for the covalent attachment of prefabricated 5 -amino-labeled nucleic acid compounds (oligonucleotides, PCR products, peptide nucleic acids).
Additional Materials (also see Basic Protocol) Phenylene diisothiocyanate (PDITC) or dimethylsuberimidate dihydrochloride (DMS) 10% (v/v) anhydrous pyridine in amine-free DMF (for PDITC) Saturated aqueous sodium bicarbonate (NaHCO3 ; for DMS) 5 -Amino-labeled nucleic acid Diisopropylethylamine (DIPEA) 1 mM Tris·Cl, pH 7.5 (see APPENDIX 2A; optional) 6-Amino-1-hexanol Amino-functionalized glass slides or polypropylene sheets (see Basic Protocol) 37◦ C humid chamber Activate linker For activation with PDITC 1a. Fill an 18 × 8–cm polypropylene container with 192 mg (1 mmol) PDITC in 40 mL of 10% (v/v) anhydrous pyridine in DMF. 2a. Immerse amino-functionalized glass slides or polypropylene sheets in activation solution and incubate for 2 hr with gentle agitation on an orbital shaker. 3a. Wash twice with 30 to 40 mL amine-free DMF, shaking gently for 2 to 3 min each. Derivatization of Glass and Polypropylene Surfaces
4a. Wash twice as above with dichloroethane. 5a. Dry slides under a stream of nitrogen gas.
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For activation with DMS 1b. Fill an 18 × 8–cm polypropylene container with 273 mg (1 mmol) DMS in 40 mL saturated NaHCO3 . 2b. Immerse the amino-functionalized glass slides or polypropylene sheets in activation solution and incubate for 1 hr with gentle agitation on an orbital shaker. 3b. Wash twice with 30 to 40 mL water, shaking gently for 2 to 3 min each. 4b. Wash twice as above with acetone. 5b. Dry slides under a stream of nitrogen gas.
Spot nucleic acids onto activated surfaces 6. Take up a 5 -amino-labeled nucleic acid in 1% (w/v) DIPEA in water or 1 mM Tris·Cl, pH 7.5. 7. Administer droplets of 0.1 to 50 nl onto the activated support. 8. Incubate in a humid chamber at 37◦ C overnight. 9. Wash twice with 30 to 40 mL water, shaking gently for 2 to 3 min each. 10. Wash twice as above with methanol. 11. Deactivate by incubating 2 hr in a freshly made solution of 50 mM 6-amino-1-hexanol and 150 mM DIPEA in DMF. 12. Wash the DNA arrays successively (two times each) with DMF, acetone, and water. 13. Store arrays dry for 4 to 8 weeks at 4◦ C.
COMMENTARY Background Information This unit describes the synthesis of a flexible linker system on glass and polypropylene support media (Beier and Hoheisel, 1999). The reaction scheme consists of a two-step procedure: an initial activation followed by a reaction with an amine, preferentially a polyamine. In the first step, a surface-bound anchoring group is reacted with 4-nitrophenyl chloroformate to form an activated ester, or with acryloyl chloride to form an α,β-unsaturated carbonyl derivative (Fig. 12.4.1). Both products represent activated species that are reactive towards an amine derivative. The reaction of the nitrophenyl ester with an amine proceeds quickly, whereas the α,β-unsaturated carbonyl derivative needs a prolonged 24-hr reaction time. However, because the latter leads to better final hybridization results, activation with acryloyl chloride is preferred. After having completed a first cycle of derivatization (activation plus amine coupling), the very same procedure may be repeated with the same or a different amine reagent. By simple variation of the amino components, different linker types can be synthesized. The linker properties can be mod-
ulated in many ways, from merely increasing the distance between the support surface and the biomolecule to be attached, to multiplying the loading capacity. If a polyamine such as tetraethylenepentamine is utilized, for example, the loading capacity of the support should be increased approximately five-fold. Since polyamines contain both primary and secondary amino functions, one unique product is not generated, but rather a mixture of compounds. Furthermore, the number of amino functions incorporated during linker synthesis controls the number of positive charges at neutral pH. Employing the amines found in the materials section of the Basic Protocol, a large number of different linker types have been synthesized by this reaction scheme (Beier and Hoheisel, 1999). The support characteristics can be tailored to the requirements of the intended applications. Many linker systems were found to be well suited for binding nucleic acids; however, one system in particular proved to be excellent under all conditions and was used for attaching prefabricated DNA oligonucleotides, PCR products, and peptide nucleic acid (PNA) oligomers, as well as for in situ synthesis of
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DNA microarrays. It is generated preferably on aminated (glass) surfaces by sequential reactions with acryloyl chloride, tetraethylenepentamine, acryloyl chloride, and 1,4-bis(aminopropoxy)butane (Fig. 12.4.2). For in situ synthesis of DNA microarrays (Pease et al., 1997; Beier and Hoheisel, 2000), no further derivatization step is required. The terminal amino group of the linker serves directly as the starting point for oligonucleotide synthesis. For immobilization of prefabricated nucleic acids, an additional activation is needed to generate a covalent linkage between biomolecule and support. For that purpose, bifunctional cross-linking reagents like phenylene diisothiocyanate or dimethylsuberimidate are utilized (Beier and Hoheisel, 1999). Activating the solid-phase surface prevents any crossreaction between the nucleic acids prior to immobilization. Prior to spotting onto the support, a base is added to the nucleic acid solution, since the cross-linking agents have their optimal reactivity in a basic milieu. Because of its non-nucleophilic character, diisopropylethylamine is employed, since it does not compete for the reactive sites on the support. However, immobilization also works at neutral pH (i.e., in water). Since small droplets evaporate quickly after application, the reaction is run to completion by incubating the DNA arrays in a humid chamber at 37◦ C after administration of the spots. Alternatively, betaine may be used to reduce evaporation (Diehl et al., 2001).
Critical Parameters Overall, the procedure is very robust. The most critical parameter in actual routine processing is preventing moisture from entering the reaction chamber during the two-step derivatization. Also, there should be no lengthy time interval between activation and amination.
a critical factor for glass supports in particular. With the protocols for coupling nucleic acids as detailed above, an increase by ten-fold has been observed.
Time Considerations The overall preparation time depends on the number of iterative activation and amination steps performed and is mainly determined by the incubations times. About 1 day should be considered for each cycle of activation and amination. However, even if several iterative cycles are performed, throughput is no major limitation, since surface modification can be performed in bulk.
Literature Cited Beier, M. and Hoheisel, J.D. 1999. Versatile derivatisation of solid support media for covalent bonding on DNA-microchips. Nucl. Acids Res. 27:1970-1977. Beier, M. and Hoheisel, J.D. 2000. Production of quantitative photolithographic synthesis of individually quality checked DNA microarrays. Nucl. Acids Res. 28:e11. Diehl, F., Grahlmann, S., Beier, M., and Hoheisel, J.D. 2001. Manufacturing DNA-microarrays of high spot homogeneity and reduced background signal. Nucl. Acids Res. 29:e38. Pease, A.C., Solas, D., Sullivan, E.J., Cronin, M.T., Holmes, C.P., and Fodor, S.P.A. 1994. Lightgenerated oligonucleotide arrays for rapid DNA sequence analysis. Proc. Natl. Acad. Sci. U.S.A. 91:5022-5026.
Contributed by Markus Beier Febit AG Mannheim, Germany J¨org D. Hoheisel Deutsches Krebsforschungszentrum Heidelberg, Germany
Anticipated Results Building a dendrimeric structure increases the loading capacity significantly, which can be
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DNA Microarray Preparation by Light-Controlled In Situ Synthesis
UNIT 12.5
Although array technology has been used in genomic research for only a few years, it has already become a standard tool in molecular biology laboratories, and it is expected to reshape the way in which future genomic research is conducted. Because a lot of sequence information is already available, and because even more will be available from public databases within the next few years, oligonucleotide arrays that require only sequence data for setup will become especially important. With such microarrays, produced by in situ synthesis using straightforward phosphoramidite chemistry, it is possible to address many applications, including resequencing, typing of single-nucleotide polymorphisms (SNPs), transcriptional profiling, and the investigation of variations in gene splicing. Spatial control during synthesis can be achieved either by light-directed activation or by using a robotic inkjet printer. This unit presents the light-directed activation process for producing oligonucleotide microarrays.
STRATEGIC PLANNING With regard to materials and equipment, there are four requirements for performing lightdirected in situ synthesis of DNA microarrays: (1) a solid support (preferably a glass substrate); (2) phosphoramidite building blocks carrying photolabile protecting groups; (3) a DNA synthesizer; and (4) an irradiation apparatus. Furthermore, for the subsequent use and analysis of the in situ–synthesized DNA microarrays, a hybridization chamber and a fluorescence scanner (preferably a laser- or CCD-based system) are needed.
Solid Support In principle, any flat solid support that is suitable for oligonucleotide synthesis via phosphoramidite chemistry may be used. Although plastic surfaces have been used successfully, nearly all of the more widely used supports are made of glass. Ideal anchoring groups for starting the oligonucleotide synthesis process include hydroxyl and amino groups, with the former being preferred. This preference arises because phosphoramidite building blocks form more stable phosphodiester linkages to hydroxyl linkers compared with amino linkers. If desired, a linker may be placed between the anchoring group and the starting monomer in the oligonucleotide synthesis. It has been shown that long linkers lead to increased yields from in situ syntheses on solid supports. Aside from solid supports consisting of dendrimeric linker systems (Beier and Hoheisel, 1999; see UNIT 12.4), quite a few other solid support media that are suited to in situ synthesis can be found in the literature (e.g., Southern et al., 1994; McGall et al., 1997).
Phosphoramidite Building Blocks Carrying Photolabile Protecting Groups Several types of photolabile protecting groups have been introduced for the light-controlled synthesis of DNA microarrays. The more prominent ones include [(2-nitroveratryl)oxy]carbonyl (NVOC; Fodor et al., 1991; Pease et al., 1994), [(αmethyl-2-nitropiperonyl)oxy]carbonyl (MeNPOC; Pirrung and Bradley, 1995; McGall et al., 1997), dimethoxybenzoincarbonate (DMBOC; Pirrung et al., 1998), [2-(2nitrophenyl)propoxy]carbonyl (NPPOC; Hasan et al., 1997; Giegrich et al., 1998), and [2-(2-nitrophenyl)ethyl]sulfonyl (NPES; B¨uhler et al., 2004). Nearly all of these photolabile protecting groups are typically employed at the 5 position, thereby enabling
Contributed by Markus Beier and J¨org D. Hoheisel Current Protocols in Nucleic Acid Chemistry (2005) 12.5.1-12.5.10 C 2005 by John Wiley & Sons, Inc. Copyright
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Figure 12.5.1 Monomeric phosphoramidite building blocks. The photolabile [2-(2nitrophenyl)propoxy]carbonyl (NPPOC) protecting group can be placed at the 5 or 3 position. B, nucleobase.
the preparation of DNA microarrays that expose a free 5 end while leaving the 3 terminus attached to the solid support. More recently, DNA microarrays that expose a free 3 hydroxyl group have gathered special interest, opening the way for a new generation of applications that make use of enzymatic on-chip reactions (Beier et al., 2001). This unit focuses on experimental conditions that have been tested for the in situ synthesis of oligonucleotide microarrays using NPPOC-protected phosphoramidites (Fig. 12.5.1; Beier and Hoheisel, 2000; see UNIT 12.3). Conditions (e.g., buffer composition, irradiation time, power of the light source, irradiation wavelength) for cleavage of a different protecting group would have to be optimized specifically for that protecting group. The authors have also performed this protocol using MeNPOC (Affymetrix) as a protecting group. While NPPOC is best removed using an irradiation buffer containing a non-nucleophilic base such as acetonitrile, MeNPOC groups have to be removed under “dry” conditions. NPPOC also leads to a significantly higher yield than does MeNPOC.
DNA Synthesizer Any conventional DNA synthesizer can be employed for light-directed synthesis of DNA microarrays. To perform an in situ synthesis on a flat surface, a flow cell–like holder for the solid support is needed (for an example, see Fig. 12.5.2). The inlet and outlet of this holder are attached to the DNA synthesizer in the same way a normal synthesis column is attached. To customize the DNA synthesizer for in situ synthesis on a solid support, the instrument’s synthesis protocols have to be adapted to the flow cell holder. For high yields to be achieved, it has to be guaranteed that a sufficient volume of reagent solution is transported to the solid support and subsequently removed. Special care must be taken to ensure that no gas bubbles are generated on the solid support within the flow cell. In principle, all steps of the synthesis protocols presented here are the same as for standard DNA synthesis on a conventional synthesis column, except for the detritylation step. In light-controlled synthesis, detritylation is replaced by an irradiation step. In this step, the flow cell is flushed with an irradiation buffer, and defined regions of the solid support (where chain elongation is desired) are activated by irradiation.
DNA Microarray Preparation by Light-Controlled In Situ Synthesis
Irradiation Apparatus The irradiation device is responsible for illuminating defined regions of the solid support, thus removing the photolabile protecting groups present in those regions. At these deprotected positions, oligonucleotide chain elongation takes place during the next round of
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Figure 12.5.2 Flow cell apparatus for light-controlled in situ synthesis of DNA microarrays. The main components of the apparatus are the reactor and the mask. A rubber seal (b) is fitted around a glass chip (i.e., derivatized slide; c), which is then placed on top of a Teflon reactor (d) that sits on an aluminum base (e). The slide is positioned with its derivatized side facing down, such that it is illuminated through its “back” side. There will be a gap between the front (derivatized) side of the slide and the bottom of the reactor; its depth (∼0.5 mm) is defined by the thickness of the Teflon sealing (outer edge) of the reactor. The inner dimensions of the reactor should be smaller than the corresponding dimensions of the glass slide (so that the edges of the slide rest securely on the raised outer edges of the reactor), but larger than the area over which the oligonucleotide spots will be distributed. The inlet/outlet ports of the flow cell are drilled into the reactor itself at the front and back, and lead to the sealed space between the slide and the reactor. The actual ports at the outside of the reactor are made to fit the tubing of the DNA synthesizer that is used to control the flow of reagents. To complete the assembly of the apparatus, the stainless steel top plate (a), a single solid piece containing 16 to 64 holes, is placed on top of the seal and glass slide, and components a through e are fastened together. The mask will touch the back of the slide. For a full-color version of this figure, go to http://www.currentprotocols.com/colorfigures.
the synthesis cycle. The most important parameter is the spatial control of the irradiation pattern. Additional critical factors include intensity, wavelength, and duration of light exposure. Since most photolabile protecting groups absorb within the UV spectrum, the light source of choice is a mercury lamp. To set the irradiation wavelength for NPPOC removal, a 365-nm interference filter is typically employed. Several techniques can be used to control the accuracy of the light pattern during the irradiation step. Among them, photolithography, pioneered by Affymetrix (Fodor et al., 1991; Fodor et al., 1993), represents the most common technology for generating oligonucleotide microarrays. Spatial resolution is obtained by implementing a new photolithographic mask for each synthesis step; for a typical DNA microarray with 25-mer oligonucleotides, 75 to 85 photolithographic masks are commonly required (as many as 100 masks may be required). Another more flexible option is the use of a so-called spatial light modulator, such as a digital micromirror device (DMD). Using a DMD, the irradiation pattern can be controlled electronically via a computer interface, which is advantageous when a larger set of possible microarray designs is desired (Gao et al., 1998; Singh-Gasson et al., 1999). Both photolithography and DMD-aided irradiation represent high-end technologies, and only the latter can be performed using a commercially available benchtop instrument (Baum et al., 2003).
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Figure 12.5.3 Schematic drawing of an assembly (consisting of a DNA synthesizer and an irradiation apparatus) for performing light-directed synthesis of oligonucleotide microarrays.
BASIC PROTOCOL 1
ASSEMBLY OF AN APPARATUS FOR LIGHT-DIRECTED SYNTHESIS OF OLIGONUCLEOTIDE MICROARRAYS The low-tech apparatus described here (Fig. 12.5.3) represents a simple combination of the lithographic and DMD-based approaches and allows the production of oligonucleotide microarrays of low complexity (see below). Instead of replacing the photolithographic mask after every synthesis step, a single mask is directly mounted onto the flow cell and used throughout (Fig. 12.5.2). For the synthesis of a 16-feature microarray, for example, this unique mask would have 16 holes. The light pattern projected onto the microarray surface during each synthesis cycle is obtained by closing certain holes mechanically; this can be achieved by applying removable black tape to the desired holes to prevent irradiation of the surface underneath. As a result, the photolabile protecting groups at these positions (whether attached to the support or to the growing oligonucleotide) are not removed during the next synthesis step. The irradiation apparatus consists of a mercury lamp within an air-cooled housing mounted on an optical bench. To select the right wavelength for removal of the NPPOC groups, a 365-nm interference filter is placed between the mercury lamp and the flow cell, which is also mounted on the optical bench. The term low complexity refers to the low number of different sequences that can be synthesized on a single microarray (i.e., 16 to 64 distinct oligonucleotide probes). In principle, long oligonucleotides can be produced using this apparatus; however, because the method represents a low-tech approach, the overall yield corresponds directly to the precision of the instrumentation. This approach is not recommended for the synthesis of oligonucleotide probes longer than 25 bp. Although rather simple and not automated, this method permits the low-throughput production of oligonucleotide microarrays of lower complexity without the need for expensive equipment.
Materials
DNA Microarray Preparation by Light-Controlled In Situ Synthesis
50 mM iodine in 7:1:2 (v/v/v) acetonitrile/pyridine/water Flow cell apparatus (custom-made; Fig. 12.5.2), including the following: Teflon reactor Aluminum base plate Rectangular rubber seal
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Nonderivatized microscope slide (for optimization of DNA synthesizer program) Derivatized microscope slide (for synthesis of microarray; see UNIT 12.4) Stainless steel top plate with inner mask (16 to 64 holes) DNA synthesizer (e.g., Eppendorf D200) 100-watt mercury lamp (e.g., Leica) 365-nm interference filter (e.g., Owis) Optical shutter (e.g., Owis) Optical bench (e.g., Owis) and assorted optical hardware Assemble the chemical apparatus 1. Assemble a flow cell holder containing a derivatized microscope slide as follows (also see Fig. 12.5.2). a. Place the Teflon reactor on top of the aluminum base plate. b. Fit a rectangular rubber seal around a nonderivatized glass microscope slide. Place the slide on top of the Teflon reactor. c. Place the stainless steel top plate/mask on top of the slide. d. Secure the components of the flow cell apparatus to each other by tightening with thumbscrews, taking care not to break the glass slide. 2. Using standard fittings provided with the DNA synthesizer, attach the synthesizer tubing to the inlet and outlet ports on the Teflon reactor. The DNA synthesizer tubing that is normally connected to the bottom of a synthesis column should be connected to the reactor’s inlet port such that the inlet port is on the bottom when the flow cell apparatus is mounted on an optical bench, and the synthesizer tubing that is normally connected to the top of a synthesis column should be connected to the reactor’s outlet port.
Modify DNA synthesizer protocols 3. Modify a standard, preprogrammed DNA synthesizer protocol by replacing the detritylation step with a waiting step of 300 sec, corresponding to the duration of irradiation. Ensure that this modified program also includes cleavage of the final photolabile protecting group. During the irradiation step, irradiation buffer is pumped into the flow cell until it is completely filled, without any bubbles. After the 300-sec waiting step, argon is pumped into the flow cell until the irradiation buffer is completely removed. This is followed by a washing step.
4. Fill all synthesizer reagent bottles with 50 mM iodine in 7:1:2 (v/v/v) acetonitrile/pyridine/water. The use of this colored solution allows the passage of liquid through the system to be monitored visually.
5. Run the modified DNA synthesizer program. Using the movement of the colored iodine solution as a guide, adjust the time and volume of each step (coupling, washing, capping, oxidation, and irradiation) to the volume of the flow cell.
Assemble the optical setup 6. Mount the mercury lamp, interference filter, and optical shutter onto the optical bench as shown in Figure 12.5.3. The distance between the lamp and the flow cell will depend on the type of lamp used. Generally, the lamp should be as close to the flow cell as possible. It is critical that the flow cell be illuminated completely.
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7. To automate the illumination process, connect the optical shutter to the fraction collector port on the DNA synthesizer. The synthesizer will trigger the opening of the shutter just before the flow cell is flushed with irradiation buffer, and will hold it open for the programmed irradiation period. The type of connector needed depends on the DNA synthesizer used; the authors use the DNA synthesizer’s trityl cable for this purpose.
8. Replace the nonderivatized slide in the flow cell apparatus with the derivatized slide on which the microarray is to be generated (installing the slide with its derivatized surface facing down), and then mount the flow cell onto the optical bench. For this purpose, the authors use a slider that is mounted to the optical table and is fitted with a screw that couples to a hole drilled in the aluminum base of the flow cell.
9. Align the mercury lamp, filter, shutter, and flow cell such that the illumination of the mask on the front of the flow cell is optimized. BASIC PROTOCOL 2
LIGHT-DIRECTED IN SITU SYNTHESIS OF OLIGONUCLEOTIDE MICROARRAYS In this protocol, the sequence programmed into the DNA synthesizer should resemble the sum of all of the different oligonucleotide probe sequences that are to be synthesized on the microarray. The simplest way to design the synthesis program is to perform a separate coupling step for each position of each sequence, uncovering the appropriate holes in order. As an example, for a simple “array” containing the two 4-bp sequences 5 -GTCG-3 (sequence 1) and 5 -GATG-3 (sequence 2), the program GGCTTAGG can be used, where monomers are added starting at the 3 end with G for sequence 1, G for sequence 2, C for sequence 1, T for sequence 2, and so on. With careful planning, however, the program can be optimized to reduce the number of coupling cycles by coupling the same monomer on more than one sequence at a time. Thus, in the same example, the program used could be GCTAG, where both sequences start with a 3 -G, followed by C for sequence 1, T for both sequences, A for sequence 2, and finally G for both sequences. In this manner, three coupling cycles have been saved. In either approach, to avoid mistakes, it is advisable to make a careful plan for which holes are to be opened or closed in each step.
Materials
DNA Microarray Preparation by Light-Controlled In Situ Synthesis
Standard DNA synthesizer reagents (Proligo): Activator (e.g., pyridine hydrochloride, dicyanoimidazole, tetrazole) Anhydrous acetonitrile (for wash steps) Oxidizing reagent Capping reagents Irradiation buffer: 50 mM diisopropylethylamine (DIPEA) in acetonitrile NPPOC-protected phosphoramidite solutions in acetonitrile (dA, dC, dG, and T; concentration, 0.1 M for each; see UNIT 12.3) 25% (v/v) ammonia in water Nitrogen stream Assembly for light-directed oligonucleotide microarray synthesis (see Basic Protocol 1) Removable black tape (e.g., standard laboratory tape): one precut piece of sufficient size to cover the entire stainless steel mask, plus precut pieces to cover each individual hole in the mask Polypropylene jar with lid Orbital shaker Additional reagents and equipment for automated DNA synthesis (APPENDIX 3C)
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NOTE: Immediately before being used in this protocol, phosphoramidites (prepared as in UNIT 12.3) should be dried and dissolved in acetonitrile. Care should be taken to prevent phosphoramidite solutions from absorbing humidity from the atmosphere.
Perform light-directed synthesis 1. Fill each reagent bottle of the DNA synthesizer with the appropriate standard DNA synthesizer reagent, irradiation buffer (to replace detritylation solution), or NPPOCprotected phosphoramidite solution (see Fig. 12.5.3). 2. Program the desired nucleotide sequence into the DNA synthesizer (see above), using the program modified for irradiation (see Basic Protocol 1). 3. Cover all holes in the flow cell mask using a single piece of removable black tape. 4. Turn on the mercury lamp. The lamp remains on throughout the entire synthesis, with the shutter being triggered to open just before the flow cell is flushed with irradiation buffer. Thus, each irradiation step starts with the opening of the shutter and ends 300 sec later with the closing of the shutter.
5. Start the DNA synthesizer program. 6. After the first coupling step (but before the first irradiation step begins), remove the single piece of black tape covering the flow cell mask. Use a separate, precut piece of black tape to cover each microarray spot that is to be protected from irradiation during the first synthesis cycle, and leave all other holes uncovered. At the conclusion of the first irradiation step, carefully cover each uncovered hole in the mask with a separate, precut piece of removable black tape. 7. In each subsequent synthesis cycle, remove the tape from each microarray spot at which chain elongation is desired during that cycle. Continue until all desired sequences have been completed. The accurate removal and attachment of tape can be complicated with a 16-hole setup, and even more challenging when a 64-hole setup is used. Careful experimental planning is necessary, and it is particularly advisable to make notes in advance regarding each irradiation pattern to be used.
Perform final deprotection 8. After the synthesis program has been completed, remove the microscope slide from the flow cell holder. 9. Fill a polypropylene jar with 25% (v/v) ammonia. Immerse the microscope slide in the ammonia solution, cover with a lid, and agitate 2 hr using an orbital shaker. This removes both the base-protecting groups and the cyanoethyl groups.
10. Remove the microscope slide from the deprotection solution, wash with water, and dry under a stream of nitrogen.
HYBRIDIZATION TO OLIGONUCLEOTIDE MICROARRAYS This protocol describes the hybridization of target oligonucleotides to oligonucleotide microarrays produced by light-directed in situ synthesis. Hybridization is performed using a simple coverslip procedure, which is easy to perform and cost-effective.
BASIC PROTOCOL 3
Materials Target oligonucleotide sample SSARC hybridization buffer (see recipe) Microscope slide containing oligonucleotide microarray (see Basic Protocol 2) Nitrogen stream
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Coverslip Polypropylene vessel with lid Fluorescence scanner, preferably laser or charge-coupled device (CCD), and image analysis software 1. Prepare the hybridization solution by dissolving the target oligonucleotide sample in SSARC hybridization buffer at room temperature. A final oligonucleotide concentration ranging from 2 to 100 nM should be appropriate, depending on the microarray used and the specific assay being performed.
2. Apply 50 µL hybridization solution onto the microarray on the microscope slide and then cover with a coverslip, taking care to avoid bubble formation underneath the coverslip. 3. Place the microscope slide face-up on a damp tissue in a polypropylene vessel. Close the lid and allow hybridization to occur at the required temperature for the desired length of time (usually overnight). The damp tissue ensures a humid environment for hybridization.
4. Using tweezers, remove the coverslip from the slide. 5. Wash the slide carefully by briefly immersing in cold SSARC hybridization buffer, and then dry under a stream of nitrogen. 6. Insert the slide into a fluorescence scanner and analyze in accordance with the manufacturer’s instructions. 7. To reuse the slide with a new target oligonucleotide, denature duplexes by incubating the slide for 30 sec at 95◦ C in 2.5 mM Na2 HPO4 containing 0.1% (w/v) SDS. Repeat with fresh stripping solution if needed. Wash with water to remove salt, and dry the slide before storing. The reuse of slides after storage for >3 months is not recommended. The original target oligonucleotides must be completely removed before the slide is rehybridized. One or two incubations is usually sufficient; repeated or extended incubations have only been required for a strong positive charge on the solid support. Complete target removal should be verified by fluorescence scanning.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
SSARC hybridization buffer 600 mM NaCl 60 mM sodium citrate 7.2% (v/v) sodium N-laurylsarcosine (Sigma) Adjust pH to 7.0 using 1.0 M citric acid or 1.0 M NaOH Store 6 months at room temperature COMMENTARY Background Information DNA Microarray Preparation by Light-Controlled In Situ Synthesis
There are two techiques for producing microarrays: (1) spotting/deposition of prefabricated DNA sequences (10 to 70 bp) or PCR products, and (2) in situ synthesis of oligonucleotides (0 to 70 bp). Whereas the spotting
approach is used primarily when sequence information is not available, the in situ approach is superior due to its flexibility. Microarrays generated by in situ synthesis are usually fabricated from the 3 end to the 5 end, yielding microarrays that are valuable
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tools for straightforward hybridization-based assays. Building an oligonucleotide chain from the 5 end to the 3 end leaves a free 3 -OH group, which has an additional use in microarrays: with a free 3 -OH, enzyme-based assays (e.g., using primer extension) become possible. Synthesis in this “reverse” direction can be achieved using the appropriate NPPOCprotected monomers described here and in UNIT 12.3. The low-tech approach presented here is not useful for applications such as gene expression profiling, where hundreds or thousands of genes—each represented by up to 20 oligonucleotides—must be investigated. However, the simplicity and low cost associated with the preparation of these low-complexity microarrays make this approach ideal for applications in which the analysis of a small number of oligonucleotides can result in meaningful findings (e.g., SNP typing, virus typing, basic microarray technology studies).
Critical Parameters
scher, H., H¨onig, M., Andreae, C., B¨uchner, D., Moschel, E., Glathe, A., J¨ager, E., Thom, M., Greil, A., Bestvater, F., Obermeier, F., Burgmaier, J., Thome, K., Weichert, S., Hein, S., Binnewies, T., Foitzik, V., M¨uller, M., St¨ahler, C.F., and St¨ahler, P.F. 2003. Validation of a novel, fully integrated and flexible microarray benchtop facility for gene expression profiling. Nucl. Acids Res. 31:e151. Beier, M. and Hoheisel, J.D. 1999. Versatile derivatisation of solid support media for covalent bonding on DNA-microchips. Nucl. Acids Res. 27:1970-1977. Beier, M. and Hoheisel, J.D. 2000. Production by quantitative photolithographic synthesis of individually quality checked DNA microarrays. Nucl. Acids Res. 28:e11. Beier, M., Stephan, A., and Hoheisel, J.D. 2001. Synthesis of photolabile 5 -O-phosphoramidites for the production of microarrays of inversely oriented oligonucleotides. Helv. Chim. Acta 84:2089-2095. B¨uhler, S., Lagoja, I., Giegrich, H., Stengele, K.P., and Pfleiderer, W. 2004. New types of very efficient photolabile protecting groups based upon the [2-(2-nitrophenyl)propoxy]carbonyl (NPPOC) moiety. Helv. Chim. Acta 87:620-659.
The critical parameters for this method have been discussed elsewhere (see Strategic Planning and UNIT 12.3 and UNIT 12.4).
Fodor, S.P., Read, J.L., Pirrung, M.C., Stryer, A., Liu, A., and Solas, D. 1991. Light-directed, spatially addressable parallel chemical synthesis. Science 251:767-773.
Anticipated Results
Fodor, S.P., Rava, R.P., Huang, X.C., Pease, A.C., Holmes, C.P., and Adams, C.L. 1993. Multiplexed biochemical assays with biological chips. Nature 364:555-556.
The product yield is determined by the coupling efficiency as well as the efficiency of removal of the photolabile protecting groups. Whereas the observed coupling efficiency is comparable to that for standard DNA synthesis methods (98% to 99%), the trigger for high yields—and, therefore, the primary factor that determines the rate of failure sequences—is the type of photochemistry used. It has been shown, for example, that NPPOC protecting groups are removed more efficiently than are MeNPOC protecing groups.
Time Considerations For a 25-mer array, the maximum number of synthesis cycles is 4 × 25 = 100 (i.e., the number of possible nucleotide choices at a given position times the number of nucleotides in the synthesized oligomers). Typically, 40 to 70 cycles are needed to build a 25-mer array, depending on the choice of sequences. Assuming the use of 10-min irradiation steps and 5-min synthesis cycles, a complete microarray synthesis can be expected to take 10 to 17 hours.
Literature Cited Baum, M., Bielau, S., Rittner, N., Schmid, K., Eggelbusch, K., Dahms, M., Schlauersbach, A., Tahedl, H., Beier, M., G¨uimil, R., Scheffler, M., Hermann, C., Funk, J.M., Wixmerten, A., Reb-
Gao, X., Yu, P., LeProust, E., Sinigo, L., Pellios, J.P., and Zang, H. 1998. Oligonucleotide synthesis using solution photogenerated acids. J. Am. Chem. Soc. 120:12698-12699. Giegrich, H., Eisele-B¨uhler, S., Hermann, C., Kvasyuk, E., Charubala, R., and Pfleiderer, W. 1998. New photolabile protecting groups in nucleoside and nucleotide chemistry—synthesis, cleavage mechanisms and applications. Nucleosides Nucleotides 17:1987-1996. Hasan, A., Stengele, K.P., Giegrich, H., Cornwell, P., Isham, K.I., Sachleben, R., Pfleiderer, W., and Foote, R.S. 1997. Photolabile protecting groups for nucleosides: Synthesis and photodeprotection rates. Tetrahedron 53:4247-4264. McGall, G.H., Barone, A.D., Diggelman, M., Fodor, S.P., Gentalen, E., and Ngo, N. 1997. The efficiency of light-directed synthesis of DNA arrays on glass substrates. J. Org. Chem. 119:5081-5090. Pease, A.C., Solas, D., Sullivan, E.J., Cronin, M.T., Holmes, C.P., and Fodor, S.P. 1994. Lightgenerated oligonucleotide arrays for rapid DNA sequence analysis. Proc. Natl. Acad. Sci. U.S.A. 91:5022-5026. Pirrung, M.C. and Bradley, J.C. 1995. Comparison of methods for photochemical phosphoramiditebased DNA synthesis. J. Org. Chem. 60:62706276.
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Pirrung, M.C., Fallon, L., and McGall, G. 1998. Proofing of photolithographic DNA synthesis with 3 ,5 -dimethoxybenzoinoxycarbonylprotected deoxynucleoside phosphoramidites. J. Org. Chem. 63:241-246. Singh-Gasson, S., Green, R.D., Yue, Y.J., Nelson, C., Blattner, F., Sussman, M.R., and Cerrina, F. 1999. Maskless fabrication of light-directed oligonucleotide microarrays using a digital micromirror array. Nat. Biotechnol. 17:974-978. Southern, E.M., Case-Green, S.C., Elder, J.K., Johnson, M., Mir, K., Wang, L., and Williams, J.C. 1994. Arrays of complementary oligonucleotides for analysing the hybridisation behaviour of nucleic acids. Nucl. Acids Res. 22:1368-1373.
Contributed by Markus Beier and J¨org D. Hoheisel Deutsches Krebsforschungszentrum Heidelberg, Germany
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Preparation of α -Oxo Semicarbazone Oligonucleotide Microarrays
UNIT 12.6
This unit presents the fabrication of α-oxo semicarbazone oligodeoxynucleotide (ODN) microarrays by site-specific ligation of α-oxo aldehyde ODNs to semicarbazide glass slides. The microarray methodology requires the preparation of α-oxo-aldehydefunctionalized ODNs and of semicarbazide glass slides. The α-oxo semicarbazone linkage is resistant to hydrolysis. Ligation and immobilization of glyoxylyl ODNs occurs spontaneously during the printing step. The microarrays can be used directly in detection experiments and do not require capping of the remaining semicarbazide groups after the printing step. Compared to aldehyde/amino ODN chemistry on glass slides, α-oxo semicarbazone microarrays allow a gain in sensitivity of ∼18-fold in model hybridization experiments using fluorescence detection (Olivier et al., 2003). The preparation of α-oxo aldehyde ODNs begins with a standard solid-phase oligonucleotide synthesis on a CPG support (Fig. 12.6.1; see Basic Protocol 1). The ODN chain is modified at the 5 end by a commercially available monomethoxytrityl (MMTr)–protected amino phosphoramidite. The deprotection is followed by acylation with a tartaric acid derivative. The tartaramide oligodeoxynucleotide (ODN-tar) is the precursor of the αoxo aldehyde ODN (ODN-COCHO). The ODN-tar is cleaved and deprotected from the CPG support by aminolysis. Finally, a periodic oxidation in aqueous solution allows the oxidative cleavage of the vicinal diol of the tartaramide and the formation of the glyoxylyl group. The preparation of semicarbazide-functionalized glass slides (Fig. 12.6.2; see Basic Protocol 2) involves a multistep procedure, which starts with the careful cleaning and activation of commercial glass microscope slides. The glass surface is modified by an amino silane, whose amine group is transformed in situ into semicarbazide. Semicarbazide glass slides are subjected to quality control prior to use (contact angle measurements, control of reactivity using labeled dipeptides). The preparation of α-oxo semicarbazone microarrays is described in Basic Protocol 3 (Fig. 12.6.3). Ligation of the ODNs to the surface is spontaneous; however, incubation of the microarrays at 30◦ C in a humid chamber improves the yield of immobilization.
SYNTHESIS AND CHARACTERIZATION OF GLYOXYLYL OLIGODEOXYNUCLEOTIDES
BASIC PROTOCOL 1
ODNs are assembled using standard phosphoramidite protocols on a controlled-pore glass support (Gait, 1984; UNITS 3.3, 4.2 & 4.10). The ODN is modified at the 5 end by a PEG-like spacer obtained by coupling two times with 18-(O-dimethoxytrityl)hexaethyleneglycol1-[(2-cyanoethyl)-(N,N-diisopropyl)]phosphoramidite and then one time with 6-(4monomethoxytritylamino)hexyl-(2-cyanoethyl)-(N,N-diisopropyl)phosphoramidite. The glyoxylyl ODN (ODN-COCHO) is obtained in four steps starting from the supportbound and MMTr-protected 5 -amino ODN (Fig. 12.6.1). The MMTr group is selectively removed using trichloroacetic acid in dichloromethane. Acylation with an excess of (+)-diacetyl-L-tartaric anhydride in the presence of 2,6-lutidine leads to support-bound ODN-tar. Aminolysis of the CPG-bound ODN with ammonium hydroxide removes all the ODN protecting groups and the O-acetyl groups on the tartaramide, and yields the deprotected ODN-tar in solution. The oligonucleotides are concentrated in vacuo, purified Contributed by Oleg Melnyk, Christophe Olivier, Nathalie Ollivier, Yves Lemoine, David Hot, Ludovic Huot, and Catherine Gouyette Current Protocols in Nucleic Acid Chemistry (2004) 12.6.1-12.6.19 C 2004 by John Wiley & Sons, Inc. Copyright
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Figure 12.6.1 Synthesis of tartaramide and glyoxylyl oligodeoxyribonucleotides. Reprinted from Olivier et al. (2003) with permission from the American Chemical Society.
by RP-HPLC, and lyophilized. In the final step, the tartaramide group is converted into an α-oxo aldehyde group by treatment with aqueous sodium periodate.
Materials
Preparation of α-Oxo Semicarbazone Oligonucleotide Microarrays
18-(O-Dimethoxytrityl)hexaethyleneglycol-1-[(2-cyanoethyl)-(N,Ndiisopropyl)]phosphoramidite (Glen Research) 6-(4-Monomethoxytritylamino)hexyl-(2-cyanoethyl)-(N,Ndiisopropyl)phosphoramidite (Glen Research) 3% (v/v) trichloroacetic acid in dichloromethane (TCA/DCM) Acetonitrile Argon Compressed air 2,6-Lutidine Anhydrous tetrahydrofuran (THF) (+)-Diacetyl-L-tartaric anhydride (Aldrich) 32% concentrated ammonia solution HPLC buffer A: 99:1 (v/v) 10 mM tetraethylamine acetate (TEAA), pH 6.5, in acetonitrile HPLC buffer B: 95:5 (v/v) acetonitrile/deionized water 3-Hydroxypicolinic acid (3-HPA) matrix 100 mM sodium phosphate buffer, pH 6.6 1 mM sodium periodate in deionized water Tartaric acid
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Tri-n-butylphosphine D-Mannitol Synthesis column with DMTr-protected controlled-pore glass (CPG) support (1-µmol scale) 1-mL glass syringes 1.5-mL screw-cap vials 55◦ C oven UV spectrophotometer and cuvettes 5-mL flask Rotary evaporator 250 × 4.6–mm and 300 × 12.5–mm C18 RP-HPLC columns Lyophilizer 1.5-mL microcentrifuge tube Additional reagents and equipment for automated DNA synthesis (UNIT 3.3, APPENDIX 3C), RP-HPLC (UNIT 10.5), MALDI-TOF-MS (see Support Protocol 1 and UNIT 10.1), and quantitation of DNA (UNIT 10.3) Detrilylate support-bound 5 -amino ODNs 1. Using standard phosphoramidite methods for automated DNA synthesis, prepare the desired protected ODN on a 1-µmol scale on a DMTr-protected CPG support. For a spacer, include cycles to modify the 5 end two times with 18(O-dimethoxytrityl)hexaethyleneglycol-1-[(2-cyanoethyl)-(N,N-diisopropyl)]phosphoramidite and then one time with 6-(4-monomethoxytritylamino)hexyl-(2cyanoethyl)-(N,N-diisopropyl)phosphoramidite. For standard methods, refer to Gait (1984), UNIT 3.3, and APPENDIX 3C. The authors have thus far only performed these reactions starting with a 1-µmol scale.
2. Remove the MMTr protecting group on the DNA synthesizer using a continuous flow of 3% TCA/DCM for 2 min. 3. Wash the column with a continuous flow of acetonitrile. This eliminates the yellow color.
4. Dry with argon and compressed air. 5. Repeat steps 2 to 4 two additional times. This deprotection can be performed manually. The acylation step must be performed immediately after the removal of the MMTr protecting group in order to avoid side reactions, in particular, acetylation of the 5 -amino group.
Introduce tartaramide moiety (acylation) 6. Remove CPG column from the synthesizer and connect a 1-mL glass syringe to the column (syringe A). 7. Mix 20 µL of 2,6-lutidine with 400 µL THF (solution 1). 8. Dissolve 15 mg (+)-diacetyl-L-tartaric anhydride in 300 µL THF (solution 2). 9. Take up 210 µL (86 equiv) of solution 1 with a new 1-mL glass syringe (syringe B) and connect to the other end of the column. Push solution 1 into the column and then remove syringe B. 10. Take up 200 µL (46 equiv) of solution 2 with a new 1-mL glass syringe (syringe C) and connect to the free end of the column. Push solution 2 into the column, and then push and pull the pistons of the syringes for 5 min (at a rate of ∼1 cycle/sec) to create an artificial flow in the synthesis column.
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11. Remove the syringes and place the column into the synthesizer. 12. Wash five to six times with THF, drying with argon after each THF wash. After the last THF/argon cycle, dry again with compressed air. This washing step can be performed manually.
13. Repeat steps 6 to 12, but increase the reaction time to 10 min in step 10.
Deprotect and cleave oligonucleotide (aminolysis) 14. Load 1 mL of 32% concentrated ammonia in a 1-mL glass syringe and connect to one end of the CPG column. Connect another 1-mL syringe to the other extremity. 15. Inject one-fourth of the ammonia solution (250 µL) by pushing the piston manually. 16. Wait 15 min (at room temperature) and then inject another 250 µL ammonia. This will simultaneously transfer the old solution (containing cleavage products) to the opposite syringe.
17. Repeat two additional times. 18. Transfer the cleavage solution from the syringe to a 1.5-mL screw-cap vial and place in a 55◦ C oven overnight.
Quantify ODN-tar by UV spectroscopy 19. Place the vial at 4◦ C to avoid evaporation and process immediately. 20. Take 10 µL of the ODN/ammonia solution (e.g., for an 18-mer), dilute with 2000 µL deionized water, and read the absorbance at 260 nm (A260 ) in a spectrophotometer against a blank of deionized water.
Analyze crude ODN-tar 21. Transfer the ammonia solution to a 5-mL flask and evaporate to dryness with a rotary evaporator. Add 1 mL deionized water. 22. Analyze by RP-HPLC (UNIT 10.5) on a 250 × 4.6–mm C18 column, with detection at 260 nm, using a linear gradient of 0% to 100% HPLC buffer B over 70 min at a flow rate of 1 mL/min. 23. Analyze by MALDI-TOF-MS (UNIT 10.1) on a 3-HPA matrix (see Support Protocol 1 for preparation of oligonucleotides for MALDI-TOF analysis).
Purify ODN-tar 24. Purify by RP-HPLC on a preparative 300 × 12.5–mm C18 column, with detection at 260 nm, using a linear gradient of 0% to 100% buffer B over 215 min at a flow rate of 3 mL/min. 25. Pool the fractions containing the pure ODN-tar as determined by MS and RP-HPLC analyses. Freeze at −20◦ C, lyophilize, and store at −20◦ C. The product can be stored at least 1 week at −20◦ C without detectable decomposition.
Quantify and analyze purified ODN-tar 26. Add 1 mL deionized water to lyophilized sample of purified ODN-tar.
Preparation of α-Oxo Semicarbazone Oligonucleotide Microarrays
27. Dilute 5 µL ODN-tar solution (e.g., for a 18-mer) with 400 µL deionized water, read the absorbance at 260 nm against a blank of deionized water, and determine the amount of ODN (UNIT 10.3). 28. Analyze by RP-HPLC and MALDI-TOF-MS as described in steps 22 and 23.
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Oxidize ODN-tar 29. In a 1.5-mL microcentrifuge tube, pipet an adequate volume of aqueous ODN-tar solution to give the desired quantity of oligonucleotide to be oxidized (deduced from UV quantitation). Freeze and lyophilize this solution. 30. Add an adequate volume of 100 mM sodium phosphate buffer, pH 6.6, to give a 0.1 to 1 mM final concentration of ODN. 31. Add 1.7 equiv sodium periodate from a 1 mM solution in deionized water. Stir 1 hr at room temperature on a magnetic stirrer. The added volume should be negligible to minimize the dilution.
32. Quench excess sodium periodate with 3.4 equiv tartaric acid. The purification of the resulting oligonucleotides should be done immediately after the oxidation step in order to avoid side reactions.
Purify and analyze ODN-COCHO 33. Purify by RP-HPLC (step 24). Record the percentage of buffer B during the elution of the desired oligonucleotide. 34. Pool the pure fractions of the desired ODN-COCHO. This ODN-COCHO solution is stable for at least 1 month at −80◦ C.
35. Prepare a blank of the same buffer composition as noted for the elution in step 33. Quantify the product in the pooled fractions by absorbance at 260 nm against this blank. 36. Add 0.005% (v/v) tri-n-butylphosphine and 6.7 µg D-mannitol/µg ODN to the pooled fractions. Freeze at −20◦ C, lyophilize, and store at −20◦ C. The ODN-COCHO must be used rapidly (<1 week) after preparation. Adding tri-n-butylphosphine and D-mannitol prevents degradation of the α-oxo aldehyde group during lyophilization.
37. Add 1 mL deionized water and quantify at 260 nm against a blank of deionized water. 38. Analyze by RP-HPLC and MALDI-TOF-MS (as in steps 22 and 23).
PREPARATION OF ODNs FOR MALDI-TOF-MS The analysis of ODNs by MALDI-TOF mass spectrometry is very sensitive to the presence of salts. It is thus important to exchange the solution of ODNs on a cation-exchange resin before analysis.
SUPPORT PROTOCOL 1
Materials Dowex 1×8 H+ (100 to 200 mesh) cation-exchange resin 1 M ammonium acetate Acetone n-Hexane 3-Hydroxypicolinic acid (3-HPA) ODN sample(s) to be analyzed (see Basic Protocol 1) 0.1 M diammonium citrate 250-mL Erlenmeyer flask 500-mL filtration flask 70-mm-diameter Buchner funnel Vacuum pump
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Parafilm 0.5-mL microcentrifuge tubes MALDI-TOF mass spectrometer (PE Biosystems) Convert H+ Dowex resin to NH4 + Dowex resin 1. In a 250-mL Erlenmeyer flask, mix 1 vol Dowex resin (e.g., 5 g) with 2 vol of 1 M ammonium acetate. Add a stir bar and mix overnight at room temperature. 2. Filter with a 500-mL filtration flask, 70-mm-diameter Buchner funnel, and vacuum pump. 3. Wash the resin two times successively with 20 mL each of deionized water, acetone, and n-hexane. 4. Dry by suction and store in a glass bottle. Under these conditions, NH4 + Dowex resin is stable several months.
Prepare sample for MALDI-TOF analysis 5. On a piece of parafilm, deposit several NH4 + Dowex resin beads. 6. In a 0.5-mL microcentrifuge tube, mix 7 µL of 3-HPA, 1 µL of ODN solution (5 to 10 OD/mL), and 1 µL of 0.1 M diammonium citrate. Pipet this mixture onto the resin beads and mix 3 to 4 min. 7. Wash the MALDI-TOF sample plate with 0.1 M diammonium citrate. 8. Take up the mixture of ODN without the beads with a pipet and deposit it on the sample plate. Let dry and analyze. BASIC PROTOCOL 2
PREPARATION AND CHARACTERIZATION OF SEMICARBAZIDE-FUNCTIONALIZED GLASS SLIDES The preparation of semicarbazide glass slides involves silanization of glass microscope slides with 3-aminopropyltrimethoxysilane to form an amine layer, conversion of amino groups into isocyanate groups, reaction of isocyanate with Fmoc-protected hydrazine, and, finally, removal of the Fmoc group in the presence of a base (Fig. 12.6.2). The amino groups are generated at the surface of the glass slide by employing a typical procedure used for the derivatization of silica supports by aminoalkoxysilanes (Balladur et al., 1997). The amine layer is converted into an isocyanate layer by reaction with triphosgene in the presence of diisopropylethylamine. Nucleophilic addition to the isocyanate layer by Fmoc-hydrazine (Zhang et al., 1991) yields the Fmoc-semicarbazide layer, which is deprotected with a nitrogen base. The chemical reactivity of the surface is checked using two labeled peptides, one modified by the COCHO group and the other modified by an amide group that does not react with semicarbazide. The surface tension of the modified glass substrate is checked using contact angle measurements (Kwok and Neumann, 1999). Drops of reference liquids (water, formamide, diiodomethane) are deposited on the substrate, and the angles between the base of the drops and the tangents at the solid-liquid-air interface are measured.
Preparation of α-Oxo Semicarbazone Oligonucleotide Microarrays
CAUTION: All syntheses must be performed in a well-ventilated laboratory fume hood, and gloves should be worn throughout the procedure.
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Figure 12.6.2 Preparation of semicarbazide glass slides. Reprinted from Olivier et al. (2003) with permission from the American Chemical Society.
Materials 25% hydrazine hydrate in water Fluorenylmethylchloroformate (Fmoc-Cl) Acetonitrile Absolute ethanol Phosphorus pentoxide (P2 O5 ) 35% (v/v) hydrogen peroxide (H2 O2 ) in water Sulfuric acid (H2 SO4 ) Methanol 3-Aminopropyltrimethoxysilane Triphosgene 1,2-Dichloroethane Diisopropylethylamine (DIPEA) N,N-Dimethylformamide (DMF) Piperidine 1,8-Diazabicyclo[5.4.0]undec-7-en (DBU) Formamide Diiodomethane Peptide 1: Rho-Lys-Arg-NH-(CH2 )3 -NH-COCHO (see Support Protocol 2) Peptide 2: Rho-Lys-Arg-CONH2 (see Support Protocol 3) 100 mM sodium acetate buffer, pH 5.5 5% aqueous K2 HPO4 0.1 M tris(hydroxymethyl)aminomethane acetate (Tris acetate), pH 5.5, containing 0.1% (v/v) Tween 20 2-L flask with a 500-mL dropping funnel Rotary evaporator equipped with a vacuum pump Water condenser Oil bath 1-L filter flask with a filter adapter and fritted glass
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Glass microscope slides (e.g., 75 × 25–mm, frosted, ESCO Precleaned Micro Slide) Glass staining dishes for 20 slides, with covers, slide racks, and slide rack handle (e.g., Wheaton) 750-mL Teflon PFA dish (15 × 10 × 5–cm, e.g., Bioblock) 1-L Teflon PFA beaker with handle (e.g., Nalgene Labware) Teflon tweezers 500- and 1000-mL Erlenmeyer flasks Sonicator, 40 kHz (e.g., Branson) 110◦ C drying oven Desiccator Vacuum pump Dust-proof container for slides Goniometer Microarray scanner with Cy3 channel Fluorescence analysis software Synthesize Fmoc-NH-NH2 1. In a laboratory fume hood, pour 60 mL of 25% hydrazine hydrate into a 2-L flask. 2. Solubilize 6.00 g Fmoc-Cl in 1500 mL acetonitrile and add dropwise using a 500-mL dropping funnel, with stirring, to the solution of hydrazine hydrate. 3. Allow to react for 30 min and then concentrate with a rotary evaporator equipped with a vacuum pump. 4. Connect a water condenser to the 2-L flask, add 250 mL absolute ethanol, and heat until ebullition with an oil bath. After solubilization in boiling ethanol, stop heating and allow Fmoc-hydrazine to crystallize slowly overnight. 5. Filter using a 1-L filter flask with a filter adapter and fritted glass. Wash the white solid with 100 mL absolute ethanol. Dry the solid in vacuo in a desiccator overnight in the presence of P2 O5 . Store the solid for months at room temperature.
Synthesize amine-functionalized glass slides 6. Arrange 39 microscope slides alternately straight across and diagonally in the glass slide rack. Place the slide rack into a 750-mL Teflon dish containing a magnetic stir bar. 7. In a 1-L Teflon beaker, with agitation, add successively 225 mL of 35% H2 O2 and 225 mL H2 SO4 . After 10 sec of agitation, add this solution to the dish containing the slides and agitate for 1 hr, allowing the solution to cool to room temperature. CAUTION: The addition of H2 SO4 to H2 O2 is highly exothermic. The mixture (known as piranha solution) is highly corrosive and must be handled with great care. After activation, the solution must be eliminated in a waste container dedicated to this aqueous acidic solution. The mixture of piranha solution with organic solvents must be strictly avoided.
8. Remove the slide rack with Teflon tweezers and dip successively, under agitation, in glass staining dishes containing:
500 mL deionized water, three times for 3 min each 500 mL methanol for 3 min. Preparation of α-Oxo Semicarbazone Oligonucleotide Microarrays
9. In a 500-mL Erlenmeyer flask mix 15 mL of 3-aminopropyltrimethoxysilane, 460.5 mL pure methanol, and 24.5 mL deionized water. Add this solution to a glass staining dish.
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10. Use the slide rack handle to remove the slide rack from the methanol solution and dip it in the silane solution. Sonicate for 30 min. 11. Remove the slide rack from the silane solution and dip successively, under agitation, in glass staining dishes containing:
500 mL methanol for 3 min 500 mL deionized water, two times for 3 min each 500 mL methanol for 3 min. 12. Remove the slide rack from the methanol solution, air dry for 2 min, and place in a 110◦ C drying oven for 15 min. 13. Store the slides one night under vacuum in a desiccator.
Synthesize semicarbazide-functionalized glass slides 14. Place the slide rack into a glass staining dish containing a stir bar. 15. In a 1000-mL Erlenmeyer flask, add successively, under agitation, 12.5 g triphosgene (weighed under an efficient laboratory fume hood), 391.3 mL of 1,2-dichloroethane, and 58.7 mL DIPEA. After 10 sec of agitation, add this solution to the dish containing the slides and sonicate for 2 hr. CAUTION: This mixture is highly toxic.
16. In a 1000-mL Erlenmeyer flask, add successively 2.52 g Fmoc-NH-NH2 , 450 mL DMF, and 4.5 mL absolute ethanol. After 10 sec of agitation, add this solution to a glass staining dish and immerse the slides in this bath. Sonicate for 2 hr. 17. Remove the slide rack from the Fmoc-NH-NH2 solution and dip successively, under agitation, in glass staining dishes containing:
500 mL DMF, two times for 3 min each 450 mL DMF with 1 mL piperidine and 10 mL DBU for 30 min 450 mL DMF for 3 min 500 mL deionized water, two times for 3 min each 500 mL methanol for 3 min. 18. Dry the slides in vacuo in a desiccator for 1 hr (they can be left overnight). Store at room temperature in a dust-proof container until use.
Measure contact angle on semicarbazide glass slides 19. Determine the contact angle of three reference liquids (water, diiodomethane, and formamide) using one semicarbazide glass slide for each liquid. With a goniometer, deposit ten 1-µL drops of the liquid on the slide, wait 10 sec, and measure the contact angle for each drop at the solid-liquid-air interface at 20◦ C. 20. Verify that the mean contact angle for each liquid matches the following reference contact angles: water, 36.3 ± 0.5; diiodomethane, 30.9 ± 0.2; and formamide, 15.6 ± 0.9. Reference angles are from Duburcq et al. (2004).
Perform chemical characterization of semicarbazide glass slides 21. Prepare separate solutions of peptides 1 and 2, each at a concentration of 10−4 M in 100 mM sodium acetate buffer, pH 5.5. Prewarm to 37◦ C for 30 min. 22. Immerse one semicarbazide glass slide in each solution and stir 1 hr at 37◦ C. Wash the slides individually with deionized water.
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23. Immerse the slides individually in 5% aqueous K2 HPO4 and sonicate 2 hr. Wash the slides individually two times with deionized water. 24. Immerse the slides individually in 0.1 M Tris acetate, pH 5.5, containing 0.1% Tween 20. Sonicate 30 min. 25. Wash the slides individually, with agitation, two times for 3 min each in water and then one time for 3 min in methanol. Dry the slides in vacuo for 30 min. 26. Scan the slides with a microarray scanner using the Cy3 channel. Use identical scanner settings, laser power, and photomultiplier level (below fluorescence saturation) for the two slides. Quantify the mean fluorescence for each slide. 27. Confirm that the ratio of mean fluorescence for peptide 1 to mean fluorescence for peptide 2 is >5. If it is not, discard the slides. SUPPORT PROTOCOL 2
SYNTHESIS OF PEPTIDE 1: RHO-LYS-ARG-NH-(CH2 )3 -NH-COCHO Synthesis of glyoxylyl-peptide 1 is performed using a manual peptide synthesis reactor and the Fmoc/tert-butyl strategy (Fields et al., 1990) on a PEG-PS type resin (e.g., Novasyn TG resin available from Novabiochem). Prior to peptide elongation, the resin is modified by an isopropylidene tartrate–based linker (Melnyk et al., 2001). The addition of each successive amino acid in the peptide chain is controlled by a TNBS test for the presence of free amines. After the peptide elongation, the peptidyl-resin is deprotected using TFA in the presence of scavengers. A periodic oxidation liberates the peptide in solution, which is immediately purified by RP-HPLC and lyophilized. NOTE: The synthesis of modified peptides requires prior chemical expertise. Reagents and solvents should be handled with care under a laboratory fume hood with gloves.
Materials
Preparation of α-Oxo Semicarbazone Oligonucleotide Microarrays
Novasyn TG resin (Novabiochem) SurfaSil siliconizing fluid (Interchim) Dichloromethane (DCM) N,N-Dimethylformamide (DMF) Fmoc-Val-OH 2-(1H-Benzotriazole-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HBTU) N-Hydroxybenzotriazole (HOBT) Diisopropylethylamine (DIPEA) 2,4,6-Trinitrobenzenesulfonic acid (TNBS) 20% piperidine in DMF Dimethyl-2,3-O-isopropylidene-L-tartrate (Aldrich) 1,8-Diazabicyclo[5.4.0]undec-7-ene (DBU, 97%, Aldrich) 4-Dimethylaminopyridine (DMAP) Benzotriazole-1-yl-oxy-tris-pyrrolidinophosphonium hexafluorophosphate (PyBOP) 1,3-Diaminopropane Fmoc-Arg(Pbf)-OH Fmoc-Lys(Boc)-OH 5(6)-Carboxytetramethylrhodamine (Rho) Trifluoroacetic acid (TFA) Thioanisole Phenol 1,2-Ethanedithiol
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Triisopropylsilane (TIS) 33% acetic acid (AcOH) NaIO4 Ethanolamine Acetonitrile Eluent A: water containing 0.05% (v/v) TFA Eluent B: water/25% (v/v) acetonitrile containing 0.05% TFA 60-mL glass manual solid-phase peptide synthesis reactor, closed with an inert screw cap at the top and a sealed-in fritted glass at the bottom, connected by a lower stopcock (customized by Vasse Industries) Vacuum flask and Teflon vacuum pump Automatic shaker Analytical and preparative HPLC systems with 300 × 12.5–mm and 250 × 4.6–mm C18 columns Lyophilizer Vacuum pump Additional reagents and equipment for TNBS test (Chan and White, 1999), RP-HPLC (UNIT 10.5), and mass spectrometry Synthesize isopropylidene tartrate–based linker on the resin 1. Introduce 0.1 mmol Novasyn TG resin in a 60-mL glass manual solid-phase synthesis reactor that has been silanized with SurfaSil siliconizing fluid prior to use. Solvents and reagents are introduced by the top and removed after each step through the fritted glass at the bottom using a vacuum flask connected to a Teflon vaccum pump.
2. Wash the resin two times for 2 min each with DCM and repeat with DMF. The solvents are eliminated by filtration using the fritted glass that is located on the bottom of the manual reactor. Throughout the procedure, perform all washes with a 20-mL volume unless otherwise indicated.
3. Dissolve 135.7 mg (4 equiv) Fmoc-Val-OH, 151.7 mg (4 equiv) HBTU, 54 mg (4 equiv) HOBT, and 0.21 mL (12 equiv) DIPEA in 2.5 mL DMF. Add to the resin and agitate on a shaker for 45 min. 4. Wash the resin two times for 2 min each with DMF and repeat with DCM. 5. Perform a TNBS test (Chan and White, 1999) for the presence of free amines. If the test is positive (presence of orange beads), wash the resin two times for 2 min each with DMF and repeat steps 3 to 5. 6. Wash the resin two times for 2 min each with DMF. 7. Add 5 mL of 20% piperidine in DMF to the resin and incubate 5 min. Repeat once for 15 min. 8. Wash the resin four times for 2 min each with DMF. 9. In a separate vessel, mix 0.76 mL (40 equiv) of dimethyl-2,3-O-isopropylidene-Ltartrate, 7.2 µL (4 equiv) water, and 60 µL (4 equiv) DBU. Stir 60 min and then add to the resin. 10. Immediately add 4.88 mg (0.4 equiv) DMAP as a solid, followed by 0.208 g (4 equiv) PyBOP dissolved in 2 mL of DMF. Agitate on a shaker for 60 min. 11. Wash the resin four times for 2 min each with DMF and two times for 2 min with DCM.
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12. Perform a TNBS test. If the test is positive, repeat steps 9 to 12. 13. Wash the resin two times for 2 min each with DMF. 14. Mix 649 µL (77 equiv) 1,3-diaminopropane and 351 µL DMF, add to the resin, and agitate on a shaker for 20 min. 15. Wash the resin two times for 2 min each with DMF.
Perform peptide elongation 16. Dissolve 648.78 mg (10 equiv) Fmoc-Arg(Pbf)-OH, 379.3 mg (10 equiv) HBTU, 135.12 mg (10 equiv) HOBT, and 0.523 mL (30 equiv) DIPEA in 2.5 mL DMF. Add to the resin and agitate on a shaker for 45 min. 17. Wash the resin two times for 2 min each with DMF and repeat with DCM. 18. Perform a TNBS test. If the test is positive, repeat steps 16 to 18. 19. Wash the resin two times for 2 min each with DMF. 20. Add 5 mL of 20% piperidine in DMF to the resin and incubate 5 min. Repeat once for 15 min. 21. Wash the resin four times for 2 min each with DMF. 22. Dissolve 187 mg (4 equiv) Fmoc-Lys(Boc)-OH, 151 mg (4 equiv) HBTU, 54 mg (4 equiv) HOBT, and 0.209 mL (12 equiv) DIPEA in 2.5 mL DMF. Add to the resin and agitate on a shaker for 45 min. 23. Wash the resin two times for 2 min each with DMF and repeat with DCM. 24. Perform a TNBS test. If the test is positive, repeat steps 21 to 24. 25. Wash the resin two times for 2 min each with DMF. 26. Add 5 mL of 20% piperidine in DMF to the resin and incubate 5 min. Repeat once for 15 min. 27. Wash the resin four times for 2 min each with DMF. 28. Dissolve 47.3 mg (1.1 equiv) 5(6)-carboxytetramethylrhodamine, 41.7 mg (1.1 equiv) HBTU, 14.9 mg (1.1 equiv) HOBT, and 57.5 µL (3.3 equiv) DIPEA in 2.5 mL DMF. Add to the resin and agitate on a shaker for 45 min. 29. Wash the resin two times for 2 min each with DMF and repeat with DCM. Dry the resin in vacuo.
Deprotect peptide 30. Prepare the following solution: 10 mL TFA 306.7 µL thioanisole 306 mg phenol 306.7 µL water 153.7 µL 1,2-ethanedithiol 61.3 µL TIS. Add to the resin and agitate on a shaker for 2 hr. Preparation of α-Oxo Semicarbazone Oligonucleotide Microarrays
31. Wash the resin four times for 2 min each with DCM. Remove the solvent and dry the resin in vacuo.
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Cleave peptide from resin 32. Wash the resin for 5 min with 10 mL of 33% AcOH. 33. Add 5 mL of 33% aqueous AcOH and 0.128 g (6 equiv) NaIO4 solubilized in 2 mL of 16% aqueous AcOH. Shake for 2 min. 34. Add 144.6 µL (24 equiv) ethanolamine and collect the cleavage solution. 35. Wash the resin two times with 3 mL water and collect the washes.
Purify peptide and lyophilize 36. Pool all solutions and add 30 mL deionized water. 37. Purify by RP-HPLC (UNIT 10.5) on a 300 × 12.5–mm C18 preparative column, with detection at 230 nm, using a linear gradient of 0% to 17% eluent B over 5 min, 17% to 24% eluent B over 7 min, and 24% to 44% eluent B over 40 min at a flow rate of 3 mL/min. Peptide 1 is composed of two isomers (5- and 6-carboxytetramethylrhodamine derivatives). Depending on the purification conditions, they may appear as one or two peaks on the chromatogram.
38. Identify the fractions containing pure peptide 1 (both isomers) by MALDI-TOF mass spectroscopy and analytical RP-HPLC. Purify by RP-HPLC (isomer 1 = 13.3 min and isomer 2 = 14.8 min) on a 250 × 4.6–mm C18 nucleosil column, with detection at 215 nm, using a linear gradient 0% to 100% eluent B over 30 min at a flow rate of 1 mL/min at 50◦ C. Mass analysis: [M+H]+ calculated monoisotopic 827.4, found 827.4.
39. Combine the pure fractions, lyophilize, and store at −20◦ C.
SYNTHESIS OF PEPTIDE 2: Rho-Lys-Arg-NH2 Synthesis of peptide 2 is performed on a manual peptide synthesis reactor using the Fmoc/tert-butyl strategy (Fields et al., 1990) and a Rink amide type resin (e.g., Novasyn TGR resin available from Novabiochem). The couplings are controlled by a TNBS test as for peptide 1 (see Support Protocol 2). After peptide elongation, the peptide is deprotected and cleaved from the resin using TFA in the presence of scavengers. The peptide is precipitated in diethyl ether/n-heptane, purified by RP-HPLC, and lyophilized prior to use.
SUPPORT PROTOCOL 3
NOTE: The synthesis of modified peptides requires prior chemical expertise. Reagents and solvents should be handled with care under a laboratory fume hood with gloves.
Additional Materials (also see Support Protocol 2) Novasyn TGR resin (Novabiochem) 1:1 (v/v) diethyl ether/n-heptane Perform peptide elongation 1. Introduce 0.1 mmol Novasyn TGR resin to a manual solid-phase synthesis reactor. 2. Wash the resin two times for 5 min each with DMF. Throughout the procedure, perform washes with a 20-mL volume unless otherwise indicated.
3. Dissolve 648.78 mg (10 equiv) Fmoc-Arg(Pbf)-OH, 379.3 mg (10 equiv) HBTU, 135.12 mg (10 equiv) HOBT, and 0.523 mL (30 equiv) DIPEA in 2.5 mL DMF. Add to the resin and agitate on a shaker for 45 min.
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4. Wash the resin two times for 2 min each with DMF and repeat with DCM. 5. Perform a TNBS test. If the test is positive, wash the resin two times for 2 min each with DMF and repeat steps 3 to 5. 6. Wash the resin two times for 2 min each with DMF. 7. Add 5 mL of 20% piperidine in DMF to the resin and incubate 5 min. Repeat once for 15 min. 8. Wash the resin four times for 2 min each with DMF. 9. Dissolve 187 mg (4 equiv) Fmoc-Lys(Boc)-OH, 151 mg (4 equiv) HBTU, 54 mg (4 equiv) HOBT, and 0.209 mL (12 equiv) DIPEA in 2.5 mL DMF. Add to the resin and agitate on a shaker for 45 min. 10. Wash the resin two times for 2 min each with DMF and repeat with DCM. 11. Perform a TNBS test. If the test is positive, repeat steps 8 to 11. 12. Wash the resin two times for 2 min each with DMF. 13. Add 5 mL of 20% piperidine in DMF to the resin and incubate 5 min. Repeat once for 15 min. 14. Wash the resin four times for 2 min each with DMF. 15. Dissolve 47.3 mg (1.1 equiv) 5(6)-carboxytetramethylrhodamine, 41.7 mg (1.1 equiv) HBTU, 14.9 mg (1.1 equiv) HOBT, and 57.5 µL (3.3 equiv) DIPEA in 2.5 mL DMF. Add to the resin and agitate on a shaker for 45 min. 16. Wash the resin two times for 2 min each with DMF and repeat with DCM. Dry the resin in vacuo.
Deprotect peptide and cleave from the resin 17. Prepare the following solution: 10 mL TFA 306.7 µL thioanisole 306 mg phenol 306.7 µL water 153.7 µL 1,2-ethanedithiol 61.3 µL TIS. Add to the resin and agitate on a shaker for 2 hr. Collect the cleavage solution. 18. Wash the resin with 5 mL TFA, collect the TFA solution, and combine with the cleavage solution.
Purify peptide and lyophilize 19. Add the peptide solution dropwise to a stirring solution of 120 mL of 1:1 (v/v) diethyl ether/n-heptane. Isolate the precipitate by centrifuging 15 min at 1600 × g, 4◦ C. 20. Purify the precipitate by RP-HPLC on a C18 preparative column using a water/acetonitrile gradient containing 0.05% (v/v) TFA.
Preparation of α-Oxo Semicarbazone Oligonucleotide Microarrays
Peptide 2 is composed of two isomers (5- and 6-carboxytetramethylrhodamine derivatives). Depending on the purification conditions, they may appear as one or two peaks on the chromatogram.
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21. Identify the fractions containing pure peptide 2 (both isomers) by MALDI-TOF mass spectroscopy and analytical RP-HPLC. Purify by RP-HPLC (isomer 1 = 13.3 min and isomer 2 = 15.0 min) on a 250 × 4.6–mm C18 nucleosil column, with detection at 215 nm, using a linear gradient 0% to 100% eluent B over 30 min at a flow rate of 1 mL/min at 50◦ C. Mass analysis: [M+H]+ calculated monoisotopic 714.4, found 714.4.
22. Combine the pure fractions, lyophilize, and store at −20◦ C.
PREPARATION OF α-OXO SEMICARBAZONE ODN MICROARRAYS This protocol describes the preparation of α-oxo semicarbazone microarrays by printing ODN-COCHO onto semicarbazide glass slides (Fig. 12.6.3). The ODN-COCHO are dissolved in 3× SSC at pH 5.5. The printing can be done using either a pin-and-ring or a piezoelectric robot. The ligation of the ODNs to the glass surface is spontaneous, but better yields of immobilization are obtained by incubating the microarrays in a humid chamber at 30◦ C. Glass slides are washed with aqueous SDS to remove the unbound ODNs and are dried before use.
BASIC PROTOCOL 3
Materials ODN-COCHO (glyoxylyl ODN; see Basic Protocol 1) 20× SSC, pH 7.0 (APPENDIX 2A) 1 N HCl Saturated NaCl 0.2% (w/v) SDS Speedvac evaporator (Savant) 96-well microtiter plates, V-shaped, low profile (ABGene) Semicarbazide glass slides (see Basic Protocol 2)
Figure 12.6.3 α-Oxo semicarbazone immobilization chemistry. Reprinted from Olivier et al. (2003) with permission from the American Chemical Society.
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Arrayer (e.g., Affymetrix 427 Arrayer) Metallic slide rack Hermetic plastic box Temperature-controlled incubator 1. Dry the ODN-COCHO in vacuo using a Speedvac evaporator. 2. Prepare 3× SSC from a 20× SSC stock solution and adjust to pH 5.5 with 1 N HCl. 3. Dissolve the ODN-COCHO in 3× SSC to a final concentration of 50 µM and transfer to a 96-well microtiter plate before spotting. 4. Place semicarbazide glass slides and 96-well plates containing ODN-COCHO in the arrayer and perform the glass printing according to manufacturer’s instructions. 5. Place the printed slides vertically in a metallic slide rack inside a hermetic plastic box containing saturated NaCl. Incubate box 14 hr at 30◦ C. Avoid putting the glass slides in contact with water.
6. To remove unbound ODNs, wash slides 5 min with 0.2% SDS at 42◦ C, then 5 min with fresh 0.2% SDS at room temperature, and finally 1 min with distilled water at room temperature. 7. Dry slides vertically by centrifuging 5 min at 200 × g. Store spotted glass slides in vacuo until use. The slides can be used directly in hybridization reactions for gene expression experiments (Schena et al., 1995; DeRisi et al., 1996). SUPPORT PROTOCOL 4
QUANTITATION OF ODN-COCHO IMMOBILIZED ONTO SEMICARBAZIDE GLASS SLIDES To assess the immobilization efficiency, 3 -radiolabeled ODN-COCHO is prepared using [α32 P]ddATP and terminal deoxynucleotide transferase. The labeled ODN is deposited onto semicarbazide glass slides with a micropipet. The bound ODN is detected and quantified with a phosphorimaging device.
Additional Materials (also see Basic Protocol 3)
Preparation of α-Oxo Semicarbazone Oligonucleotide Microarrays
[α32 P]ddATP (Amersham Biosciences) 5× terminal transferase reaction buffer: 1 M potassium cacodylate, 125 mM Tris·Cl, 1.25 mg/mL bovine serum albumin, pH 6.6 at 25◦ C Terminal transferase Qiaquick nucleotide removal kit (Qiagen) which includes: PN solution Qiaquick nucleotide removal column PE buffer DNase- and RNase-free distilled water 1.5- and 2-mL microcentrifuge tubes 37◦ C water bath Benchtop centrifuge Scintillation counter Vacuum desiccator Plastic film (e.g., Saran Wrap) Phosphorimager (e.g., Cyclone Storage Phosphor System, Packard BioScience)
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Radiolabel ODN-COCHO with [α32 P] 1. In a 1.5-mL reaction tube, mix 200 pmol ODN-COCHO and 33 pmol [α32 P]ddATP (100 µCi) into 1× terminal transferase reaction buffer (total volume 18 µL). 2. Add 50 U terminal transferase to obtain a final concentration of 2.5 U/µL and incubate 2.5 hr in a 37◦ C water bath.
Purify radiolabeled ODN-COCHO 3. Add 10 vol PN solution to the terminal transferase reaction mixture. Set aside a 2-µL sample in a separate tube to measure incorporation. 4. Apply the remaining sample to a Qiaquick nucleotide removal column to bind ODNCOCHO. Place the column in a 2-mL microcentrifuge tube and centrifuge 1 min at 8300 × g in a benchtop centrifuge at room temperature. 5. Place the column in a clean 2-mL collection tube, add 500 µL PE buffer onto the column, and centrifuge 1 min at 8300 × g, room temperature. 6. Repeat step 5. 7. Discard the wash buffer from the collection tube and return the column back to the same collection tube. Centrifuge 1 min at 18,000 × g, room temperature, to completely remove any trace of wash buffer. 8. Elute ODN from the column with 100 µL distilled water adjusted to pH 7.0. Centrifuge 1 min at 18,000 × g. 9. Determine specific activity (cpm/µL) of labeled ODN-COCHO by measuring the radioactivity of a 2-µL sample in a scintillation counter. Also measure the 2-µL sample from step 3 to calculate incorporation. Remember to account for the dilution factor when determining incorporation.
Evaluate immobilization of ODN-COCHO on semicarbazide slides 10. Prepare four reaction tubes containing different quantities of unlabeled ODNCOCHO (e.g., 2, 1, 0.2, and 0.02 nmol) in 20 µL of DNase- and RNase-free distilled water. 11. Add ∼1 × 106 cpm of labeled ODN-COCHO to each tube of unlabeled ODNCOCHO and dry the solution in a vacuum desiccator. 12. Dissolve the dry pellet in 20 µL of 3× SSC, pH 5.5, and calculate the specific radioactivity (cpm/fmol) as in step 9. 13. Deposit four 1-µL beads of each concentration of ODN-COCHO on a semicarbazide slide with a micropipet and let dry. 14. Place the slides vertically in a metallic rack inside a hermetic plastic box containing saturated NaCl. Incubate box 14 hr in a 30◦ C incubator. Avoid putting the glass slides in contact with water.
15. To remove unbound ODNs, wash slides with 0.2% SDS for 5 min at 42◦ C, then with fresh 0.2% SDS for 5 min at room temperature, and finally with distilled water for 1 min at room temperature. 16. Dry slides vertically by centrifuging 5 min at 200 × g, room temperature. 17. Prepare several dilutions of [α32 P]ddATP, deposit 1 µL of the different dilutions on the slide, and let dry. These spots constitute the reference for quantitation.
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18. Place the slides in plastic film and expose them to a phosphor screen inside a cassette. 19. Scan the screen in the phosphorimager and analyze the image with the associated software to obtain spot intensities and surrounding background evaluation. 20. Compare measured intensity values of labeled ODN spots to the ones obtained with the calibration scale spotted on the same slide. Use the calculated specific activities (step 12) to evaluate quantity (in fmol) of ODN immobilized on the glass. Trace a diagram of cpm as a function of the [α 32 P]ddATP dilutions that have been deposited in step 17. Calculate the regression line equation and use it to transform cpm measurements in quantity of [α 32 P]ddATP. Go back to the quantity of ODN using the specific radioactivity calculated in step 12. Graphical representations of fixed ODN (in fmol) as a function of the starting ODN concentration can then be obtained. Curves generally show that optimal fixation of ODN-COCHO on semicarbazide-functionalized glass slides is obtained for a small concentration (∼10 µM).
COMMENTARY Background Information
Preparation of α-Oxo Semicarbazone Oligonucleotide Microarrays
One of the most widely used strategies for microarray fabrication is to attach characterized and purified synthetic probes to a surface. In this case, the performance of the array relies mainly on—besides the selection of the probes and their mode of immobilization—the chemical and physicochemical properties of the array. Glass slides are now widely used for the preparation of DNA arrays. Glass is an inexpensive support medium that has a low intrinsic fluorescence and a relatively homogeneous chemical surface. Its properties have been studied in detail and diverse silanization chemistries are available for derivatization (e.g., UNIT 12.4). The ideal glass surface for microarray fabrication must respond to several specifications. The glass coating must (1) be homogeneous, (2) be stable upon storage, (3) be chemically inert towards biomolecules such as nucleic acids or proteins, but allow the covalent and dense immobilization of functionalized probes, so as to reach high signal-tonoise ratios, and (4) minimize the adsorption of biomolecules, so that a blocking procedure can be avoided following the printing step. On the other hand, the modified probes must (1) be stable upon storage and (2) be highly reactive towards the surface. Finally, the overall procedure for microarray fabrication must be robust to guarantee a high level of reproducibility. The method described by Podyminogin et al. (2001) is based on the printing of benzaldehyde ODNs onto semicarbazide glass slides. The glass slides are prepared differently and no quality control is described. The benzaldehyde semicarbazone is labile at 65◦ C in HEPES buffer, pH 7.5.
The methodology described here meets most of the above requirements and allows the preparation of high-quality microarrays. The method is particularly useful when high signal-to-noise ratios are needed. The quality controls allow rejection of low-quality batches. The yield of immobilization is high, and the linkage is stable at 65◦ C in HEPES buffer, pH 7.5. ODN-COCHO probes were used in printing experiments with different printing buffers. The observations led to the use of a salinetype buffer such as saline sodium citrate (SSC). When using arrayers equipped with a pinand-ring spotting system (e.g., Affymetrix 427 Arrayer), solvent such as dimethylsulfoxide or formamide led to nonuniform spot shape and/or intensity. Consequently, cosolvents must be avoided during the printing step. Most of the immobilization occurs in the first 3 hr after printing, as shown by the experiments using radiolabeled ODNs. However, incubating the slides for 14 hr in a watersaturated atmosphere at 30◦ C (as described in Basic Protocol 3) is recommended to achieve optimal immobilization yields.
Critical Parameters The quality of semicarbazide glass slides is highly dependent on the efficiency of the first cleaning step using piranha solution. One or two slides can be used after this treatment for contact angle measurements. The contact angle of water on cleaned glass must be below 15◦ . The use of ethanol as cosolvent during the reaction with Fmoc-NH-NH2 is crucial. Semicarbazide glass slides are resistant to air pollution as shown by contact angle
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Current Protocols in Nucleic Acid Chemistry
measurements during aging experiments. Nonetheless, storing the slides in a nitrogen atmosphere is recommended. The use of acetone or volatile carbonyl compounds that are able to react with semicarbazide must be avoided in the laboratory. The use of detergents for floor cleaning should be avoided since some detergents contain large quantities of aldehyde (formaldehyde), which can deactivate the semicarbazide surfaces.
Anticipated Results The quality control based on the incubation of the semicarbazide surface with peptides 1 and 2 provides a picture of the degree and homogeneity of the slide functionalization. Several batches should be prepared for the determination of reference values. Typically, the ratio of the mean fluorescence of peptide 1 to the mean fluorescence of peptide 2 is >5 and can be as high as 10. The fluorescence intensity is also an important parameter. Typically, using the Affymetrix 418 array scanner at L30/PMT45 sensitivity, the mean fluorescence obtained with glyoxylyl peptide 1 is ∼40,000. The coefficient of variation (CV = standard deviation/mean × 100) is usually 10%. The quality of the starting microscope glass slides is probably important, but a systematic study has not been performed. The slides are chemically stable at least 3 months at room temperature. Glyoxylyl peptides 1 and 2 are stable for months in the lyophilized form or in solution. Storage at −20◦ C is recommended. Microarray preparation is very simple and no capping step is required before the hybridization experiments. A gain in sensitivity of ∼18 fold was observed compared to standard aldehyde glass slide/amino ODN immobilization chemistry in model hybridization experiments using 18-mers. This gain is the result of both a better yield of immobilization (6 fold) and a reduced background (3 fold).
Chan and P.D. White, eds.) pp. 61-62. Oxford University Press, Oxford. DeRisi, J., Penland, L., Brown, P.O., Bittner, M.L., Meltzer, P.S., Ray, M., Chen, Y., Su, Y.A., and Trent, J.M. 1996. Use of a cDNA microarray to analyse gene expression patterns in human cancer. Nat. Genet. 14:457-460. Duburcq, X., Olivier, C., Desmet, R., Halasa, M., Carion, O., Grandidier, B., Heim, T., Stievenard, D., Auriault, C., and Melnyk, O. 2004. Polypeptide semicarbazide glass slide microarrays: Characterization and comparison with amine slides in serodetection studies. Bioconjug. Chem. 15:317-325. Fields, G.B. and Noble, R.L. 1990. Solid phase peptide synthesis utilizing 9-fluorenylmethoxycarbonyl amino acids. Int. J. Pept. Protein Res. 35:161-214. Gait, M.J. 1984. An introduction to modern methods of DNA synthesis. In Oligonucleotide Synthesis: A Practical Approach (M.J. Gait, ed.) pp. 1-22. IRL Press, Oxford. Kwok, D.Y. and Neumann, A.W. 1999. Contact angle measurement and contact angle interpretation. Adv. Colloid Interface Sci. 81:167-249. Melnyk, O., Fruchart, J.S., Grandjean, C., and GrasMasse, H. 2001. Tartric acid-based linker for the solid-phase synthesis of C-terminal peptide αoxo aldehydes. J. Org. Chem. 66:4153-4160. Olivier, C., Hot, D., Huot, L., Ollivier, N., El-Madhi, O., Gouyette, C., Huynh-Dinh, T., Gras-Masse, H., Lemoine, Y., and Melnyk, O. 2003. α-Oxo semicarbazone peptide or oligodeoxynucleotide microarrays. Bioconjug. Chem. 14:430-439. Podyminogin, M.A., Lukhtanov, E.A., and Reed, M. 2001. Attachment of benzaldehyde-modified oligodeoxynucleotide probes to semicarbazidecoated glass. Nucl. Acids Res. 29:5090-5098. Schena, M., Shalon, D., Davis, R.W., and Brown, P.O. 1995. Quantitative monitoring of gene expression patterns with a complementary DNA microarray. Science 270:467-470. Zhang, R.E., Cao, Y.L., and Hearn, M.W. 1991. Synthesis and application of Fmoc-hydrazine for the quantitative determination of saccharides by reversed-phase high-performance liquid chromatography in the low and subpicomole range. Anal. Biochem. 195:160-167.
Time Considerations Fmoc-NH-NH2 must be prepared 2 days before glass slide preparation to allow the reagent to crystallize and then dry efficiently. The preparation of the semicarbazide glass slides takes 2 days. Using two sonicators (four slide racks), 160 slides can be prepared at one time.
Contributed by Oleg Melnyk, Christophe Olivier, and Nathalie Ollivier Universit´e de Lille 2 Lille, France
Literature Cited
Yves Lemoine, David Hot, and Ludovic Huot Institut Pasteur de Lille Lille, France
Balladur, V., Theretz, A., and Mandrand, B. 1997. Determination of the main forces driving DNA oligonucleotide adsorption onto aminated silica wafers. J. Colloid Interface Sci. 194:408-418. Chan, W.C. and White, P.D. 1999. Fmoc Solid Phase Peptide Synthesis: A Practical Approach (W.C.
Catherine Gouyette Institut Pasteur Paris, France
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Synthesis of Covalent OligonucleotideStreptavidin Conjugates and Their Application in DNA-Directed Immobilization (DDI) of Proteins
UNIT 12.7
The development of immunoassays performed on a small scale and in parallel, based on antigen and antibody microarray technology, is currently of tremendous interest for a broad range of applications in biomedical diagnostics, where several parameters of an individual sample must be determined simultaneously from a limited amount of material (Kusnezow and Hoheisel, 2002; Schweitzer and Kingsmore, 2002; Templin et al., 2002). While microarray-based analyses of nucleic acids have made progress towards routine application (Pirrung, 2002), the production and use of protein chip devices are hampered by the intrinsic instability of many proteins. Although protein microarrays have been prepared for high-throughput antibody screening (Lueking et al., 1999), the analysis of antibody-antigen interactions (de Wildt et al., 2000; Angenendt et al., 2002, 2003), and the identification of protein targets of small molecules (MacBeath and Schreiber, 2000), the stepwise robotic immobilization of multiple proteins on chemically activated surfaces is often obstructed by the instability of most proteins, which usually reveal a significant tendency for denaturation and, thus, loss of functionality. To circumvent these obstacles, the authors have developed the method of DNA-directed immobilization (DDI) of proteins (Niemeyer et al., 1994) using covalent conjugates synthesized from
Figure 12.7.1 Schematic representation of the modular preparation of functional conjugates employed in the DDI-method. (A) Self-assembly of DNA-STV conjugate S.1 with biotinylated antibody S.2 to give the DNA-antibody conjugate S.3. (B) Site-specific immobilization of the conjugates S.3A-D on their complementary oligonucleotides on a solid support. Contributed by Ron Wacker and Christof M. Niemeyer Current Protocols in Nucleic Acid Chemistry (2005) 12.7.1-12.7.15 C 2005 by John Wiley & Sons, Inc. Copyright
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12.7.1 Supplement 22
single-stranded DNA and streptavidin (STV) as molecular connectors for tagging biotinylated proteins with single-stranded DNA oligomers (Niemeyer et al., 1994, 1999). As shown in Figure 12.7.1, the preparation of functional DNA-antibody conjugates (S.3) is accomplished through coupling of the DNA-STV conjugate (S.1) with a biotinylated antibody (S.2). The DNA-antibody conjugate can then be hybridized to arrays containing oligonucleotides with complementary sequences. DDI is a chemically mild process for site-selective adsorption of delicate proteins to a solid support, using DNA-functionalized substrates as an immobilization matrix. Because the lateral surface structuring can be carried out at the level of stable nucleic acid oligomers, the DNA-functionalized substrate can be produced and stored almost indefinitely. Stored substrates can be functionalized with proteins of interest via DDI immediately prior to use in a microscaled fluorescence immunoassay (µFIA; Wacker and Niemeyer, 2004; Wacker et al., 2004). Since both the production of the functional protein conjugates and their immobilization on a solid support are based entirely on self-assembly, the setup of DDI-µFIA is readily configured from the modular reagents used, i.e., covalent STV-DNA conjugates, biotinylated proteins, and a microarray containing complementary DNA capture oligomers. The method presented here describes the immobilization of biotinylated antibodies, but can theoretically be applied to any protein (or substance) that can be biotinylated. BASIC PROTOCOL 1
SYNTHESIS OF SEMISYNTHETIC DNA-STREPTAVIDIN CONJUGATES BY COVALENT COUPLING OF THIOL-MODIFIED OLIGONUCLEOTIDES Covalent conjugates are synthesized from streptavidin (STV) and thiol-modified oligonucleotides. The covalent attachment of an oligonucleotide moiety provides the STV with a specific binding domain for complementary nucleic acids in addition to its four native binding sites for biotin. This bispecificity of the DNA-STV conjugate S.1 allows it to serve as a universal, efficient, and highly selective connector in the oligonucleotidedirected assembly of proteins (Niemeyer et al., 1994, 1999; Wacker and Niemeyer, 2004; Wacker et al., 2004) and other molecular and colloidal components, e.g., gold nanoparticles (Niemeyer, 2002). The covalent attachment of the oligonucleotide to STV is carried out with the aid of the heterobispecific crosslinker sulfosuccinimidyl-4-(Nmaleimidomethyl)cyclohexane-1-carboxylate (sSMCC, Fig. 12.7.2). The addition of thiols to maleimides yields stable thioether conjugates. This crosslinking method is chosen because 5 -thiol-modified oligonucleotides can be prepared on a standard DNA synthesizer. The ε-amino groups of the Lys sidechains of STV are first derivatized with the sSMCC crosslinker to provide maleimide functional groups, which are subsequently coupled with the thiolated oligonucleotide. The crosslinked products are pre-purified by ultrafiltration, and then fractionated by anion-exchange chromatography using an automated FPLC system. The latter step allows for quantitative separation of the initial DNASTV conjugates, which contain different numbers of attached DNA strands, to obtain the mono-adduct S.1, which contains only one oligonucleotide per STV (Fig. 12.7.3). Subsequently, the migrating properties of the conjugates are characterized by native PAGE. The conjugation of one DNA strand to STV enhances its mobility due to the addition of negative charges, whereas the higher adducts reveal reduced electrophoretic mobility, likely due to the increased molecular size of the conjugate (Fig. 12.7.4).
Materials
OligonucleotideStreptavidin Conjugates for Immobilization of Proteins
1 M dithiothreitol (DTT) Thiolated oligonucleotides (Table 12.7.1, e.g., Thermo Electron) TE buffer, pH 7.5 (APPENDIX 2A) PBSE buffer (see recipe) Sulfosuccinimidyl-4-(N-maleimidomethyl)cyclohexane-1-carboxylate (sSMCC; Pierce)
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Figure 12.7.2
Covalent crosslinking of STV and 5 -thiolated oligonucleotides.
Dimethylformamide (DMF) 100 µM streptavidin, recombinant (STV; Roche) in PBS1 (see recipe) 1 M 2-mercaptoethanol Tris·Cl (APPENDIX 2A): 20 mM at pH 6.3, 1.5 M at pH 8.8, and 1 M at pH 6.8 TBSE (see recipe) 37.5%:1% (w/v) acrylamide/bisacrylamide solution 10% (w/v) ammonium persulfate Tetramethylethylenediamine (TEMED) 1-Butanol (pure) Running buffer (see recipe) Loading buffer (see recipe) 123-bp DNA ladder (Invitrogen) SybrGold (Molecular Probes) 37◦ C heating block or water bath Fast protein liquid chromatography (FPLC) system (Amersham Biosciences) with Superdex peptide column, fraction collector, and detector Gel filtration columns (e.g., NAP5 and NAP10, Amersham Biosciences) Molecular cut-off ultrafiltration unit (e.g., Centricon 30, Millipore) Anion-exchange column (e.g., MonoQ HR5/5, Amersham Biosciences) Photometer (e.g., BioPhotometer, Eppendorf) Electrophoresis device (e.g., Bio-Rad) Container for gel staining Transilluminator SybrGold camera filter (Biozym) AlphaImager 2200 gel documentation device (Biozym) Activate and purify oligonucleotide 1. Add 60 µL of 1 M DTT solution to 100 µL (100 µM) of the thiol-modified oligonucleotide (see Table 12.7.1) in TE buffer. 2. Mix briefly and incubate for 2 hr at 37◦ C.
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Figure 12.7.3
Chromatograph of conjugate purification by anion-exchange chromatography.
Figure 12.7.4 Gel pictures of DNA-STV conjugate. M: 123-bp DNA ladder; lane 1: 1:1 DNA:STV; lane 2: 2:1 DNA:STV (also containing 3:1); lane 3: 3:1 and 4:1 DNA:STV.
3. Purify the activated oligonucleotide by gel filtration chromatrography using a Superdex peptide column connected to an FPLC system. Inject 160 µL of activated oligonucleotide and elute the sample with PBSE buffer using a flow rate of 0.7 mL/min. Detect the absorbance at 260 and 280 nm and collect peaks in 0.55-mL fractions. OligonucleotideStreptavidin Conjugates for Immobilization of Proteins
4. Pool the collected fractions of the main peak at 260 nm (elution volume ∼10 to 12 mL), which contain the activated oligonucleotide. Use the oligonucleotide as soon as possible in the cross-linking reaction since dimer formation through disulfide bridges might occur.
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Table 12.7.1 Sequences of Thiolated Oligonucleotide Library for the Generation of DNA-Protein Conjugates and Complementary Amino-Modified Capture Oligomersa
Name
Sequence
Modification
Thiolated oligonucleotide library tA
TCC TGT GTG AAA TTG TTA TCC GCT
5 Thiolink (C6)
tB
ACC TCA AGT GAT CTA CCT ACC TCA G
5 Thiolink (C6)
tC
CTC ACA TCC AAC AAT ACA GGT CAC AT
5 Thiolink (C6)
tD
TGA GCG TTC GTG GGA TAG T
5 Thiolink (C6)
Complementary amino-modified capture oligomers cA
AGC GGA TAA CAA TTT CAC ACA GGA
5 Aminolink (C6)
cB
CTG AGG TAG GTA GAT CAC TTG AGG T
5 Aminolink (C6)
cC
ATG TGA CCT GTA TTG TTG GAT GTG AG
5 Aminolink (C6)
cD
ACT ATC CCA CGA ACG CTC A
5 Aminolink (C6)
a Sequences are shown in 5 - 3 direction.
Activate and purify STV 5. Dissolve 2 mg sSMCC in 60 µL DMF. If the sSMCC does not dissolve readily, shake at 35◦ C.
6. Add the sSMCC solution to 200 µL of 100 µM STV in PBS. Incubate 1 hr at room temperature in the dark. 7. Remove the top and bottom caps of two disposable gel filtration columns (NAP5 and NAP10) and pour off the conserving liquid. Support each column over a suitable receptacle and equilibrate the columns three times by completely filling with PBSE and allowing to flow through by gravity. 8. Apply the 260 µL activated STV on top of the filterplate of the NAP5 column and allow the liquid to completely enter the gel bed. 9. Adjust the sample volume to 500 µL by applying 240 µL PBSE. 10. Elute the activated STV with 1 mL PBSE and collect 1 mL of filtrate. 11. Apply the collected filtrate sample onto the NAP10 column. Elute the activated STV with 1.5 mL PBSE. Use the activated STV as soon as possible in the crosslinking reaction since the reactivity of the maleimide will reduce over time.
Crosslink STV and oligonucleotide 12. Mix the purified, activated STV with the oligonucleotide fractions and incubate the solution for 1.5 hr at room temperature in the dark. 13. Transfer the mixture into a Centricon 30 ultrafiltration unit and reduce the volume to ∼600 µL by alternating steps of centrifuging 4 min at 2900 × g (4000 rpm) and shaking 5 min on an orbital shaker (total two to six cycles). 14. Quench the reaction by adding 1 µL of 1 M 2-mercaptoethanol and further reduce the volume to ∼200 µL. 15. Perform buffer exchange by adding 1 mL of 20 mM Tris·Cl, pH 6.3, and repeat filtration until the sample volume is ∼200 µL.
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Table 12.7.2 Buffer Gradient Used for FPLC Purification of DNA-STV Conjugate
Volume (mL)
[NaCl] (M)
Flow rate (mL/min)
0–4 min
0.3
0.2
4–8 min
0.3 + 0.015/min
0.2
8–40 min
0.45 + 0.01/min
0.2
Purify STV-oligonucleotide conjugate 16. Purify S.1 by anion-exchange chromatography using an appropriate column (MonoQ HR5/5) and a suitable purification program (see Table 12.7.2). Collect 0.55-mL fractions. 17. Record the absorbance at 260 and 280 nm on a photometer. 18. Pool the fractions from the major peak corresponding to the mono-adduct (S.1; Fig. 12.7.3), and then exchange the buffer and concentrate the conjugate by a twofold ultrafiltration (step 13) using 500 µL each of TBSE. Store the fractions at 4◦ C until further use (up to 6 months).
Quantitate conjugate yield 19. Quantitate the concentrated conjugate fraction by measuring the absorbance at 260 and 280 nm. 20. Determine the ratios α and β of the absorbance at 260 and 280 nm for DNA and STV, respectively. Assume these ratios are constant values. α = A260(DNA) /A280(DNA) β = A260(STV) /A280(STV) 21. Using the conjugate A260 and A280 values measured in step 19, calculate the corrected absorbance at 280 nm for STV: A280(STV) = [(A280 × α) − A260 ]/(α − β) The derivation of the above equation is based on the assumption that the measured values arise from the absorbance values of the STV and the oligonucleotide in the sample: (1) A280 = A280(STV) + A280(DNA) (2) A260 = A260(STV) + A260(DNA) Inserting the equations for α and β into equations (1) and (2), respectively, gives: (3) A280 = A280(STV) + (A260(DNA) × 1/α) (4) A260 = A260(DNA) + (A280(STV) × β) Inserting (4) into (3) gives: (5) A280 = A280(STV) + [A260 − (A280(STV) × β)]/α This equation is then rearranged to solve for A280(STV) .
22. Calculate the concentration using the Lambert-Beer law: c = A/(ε × l) where the extinction coefficient (ε) of recombinant tetrameric STV at 280 nm is 142,400 M−1 cm−1 . OligonucleotideStreptavidin Conjugates for Immobilization of Proteins
The molar conjugate concentration is equivalent to the molar STV concentration. The concentration of ssDNA can be calculated from the approximation at 260 nm: 1 OD of ssDNA = ∼33 µg/mL. Use a DNA concentration of ∼1 µM for these measurements.
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Table 12.7.3 Preparation of Separating and Stacking Gels
8.5% separating gel
5% stacking gel
Distilled H2 O
3.7 mL
1600 µL
1.5 M Tris·Cl, pH 8.8
2 mL
—
1 M Tris·Cl, pH 6.8
—
285 µL
37.5%:1% (w/v) acrylamide/bisacrylamide
2266 µL
374 µL
10% ammonium persulfate
40 µL
22.5 µL
Tetramethylethylenediamine
4 µL
2.25 µL
Total
∼8 mL
∼2.3 mL
Add immediately before use:
Prepare gel for nondenaturing PAGE 23. Prepare an 8.5% separating gel in a glass beaker using the reagents specified in Table 12.7.3. Mix the components in the order shown. Mix gently but thoroughly, avoiding air bubbles. CAUTION: Wear gloves when working with acrylamide.
24. Assemble the gel electrophoresis pouring device according to the manufacturer’s instructions. Pour the separating gel solution into the gap between the glass plates. 25. Carefully pipet 100 µL of 1-butanol on top of the gel and allow the gel to polymerize for at least 1 hr. The gel can be stored wrapped in a plastic film for several weeks at 4◦ C.
26. Wash the top of the gel several times with ddH2 O, then fix the gel in the electrophoresis device according to the manufacturer’s instructions. Insert a comb between the plates. 27. Prepare the 5% stacking gel solution in a disposable plastic tube using the reagents specified in Table 12.7.3. Mix the components in the order shown. Mix gently but thoroughly, avoiding air bubbles. 28. Pipet the solution into the gap between the glass plates with a Pasteur pipet and allow the stacking gel to polymerize for 30 min. 29. Remove the comb. Flush the wells by adding running buffer and pipetting up and down to remove any unpolymerized acrylamide.
Electrophorese conjugate 30. Dilute the peak fractions (step 18) to a final STV concentration of 1.33 µM in a total volume of 17 µL with TE buffer and incubate 20 min at room temperature. Add 3 µL of loading buffer and mix thoroughly. 31. Fill the electrophoresis device with running buffer according to the manufacturer’s instructions. Load 15 µL of the sample and a 123-bp DNA ladder as a marker (final concentration 100 µg/mL). 32. Run the gel at a constant 150 V for ∼90 min, until the bromphenol blue dye front has moved to ∼1.5 cm from the bottom of the gel.
Stain and analyze gel 33. Make a 1× SybrGold staining solution by diluting the SybrGold stock solution 10,000-fold in TE buffer to a final volume of 25 mL. Transfer to a suitable container for staining the gel, and protect from light by covering the container with aluminum foil.
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34. Carefully disassemble the electrophoresis device, remove the stacking gel, and transfer the separating gel to the 1× SybrGold staining solution. Gently agitate the gel ∼15 min at room temperature. All bands of the gel containing DNA are stained with the SybrGold intercalating dye.
35. Image the stained acrylamide gel using a transilluminator at 300 nm and a SybrGold camera filter (Fig. 12.7.4). Optimal photographic conditions need to be determined experimentally with the gel documentation device. BASIC PROTOCOL 2
SEMISYNTHETIC DNA-STV CONJUGATES AS MOLECULAR LINKERS IN DNA-DIRECTED IMMOBILIZATION OF BIOTINYLATED ANTIBODIES ON DNA MICROARRAYS The covalent DNA-STV conjugates S.1A-D prepared in Basic Protocol 1 are employed as molecular connectors in the DNA-directed immobilization (DDI) of biotinylated proteins (Fig. 12.7.5). As an alternative to synthesizing the covalent DNA-STV conjugates, these compounds are commercially available from Chimera Biotec as ready-to-use reagents. DDI enables the production of microstructured protein arrays (Wacker and Niemeyer, 2004; Wacker et al., 2004). As shown in Figure 12.7.1A, the preparation of functional DNA-antibody conjugates S.3 is accomplished through coupling of the DNA-STV conjugate S.1 with a biotinylated antibody S.2 (for examples, see Table 12.7.4). The covalent modification of the proteins (antibodies) with biotinyl groups is achieved using sulfo-(Nhydroxysuccinimid)biotin (sulfoNHS-biotin). This standard modification reagent readily reacts with terminal amino groups and the amino groups of Lys sidechains of a protein. Subsequent to mixing and incubating S.1 and S.2, the conjugate product S.3 can be hybridized to arrays containing oligonucleotides with complementary sequences (Table 12.7.1) without further purification (Fig. 12.7.1B). Excess antibody, not connected to the STV conjugate, is simply removed by washing the array after hybridization. The homogeneous immobilization of S.3 is demonstrated by the binding of a species-specific antibody-Cy5 conjugate that binds to the immobilized antibodies. Alternatively, the protein microarray generated by DDI might be used in various immunological assay systems, such as a sandwich ELISA (Wacker and Niemeyer, 2004; Wacker et al., 2004).
Materials
OligonucleotideStreptavidin Conjugates for Immobilization of Proteins
PBS2 (see recipe) Antibodies (Table 12.7.4, Sigma-Aldrich) Sulfo-(N-hydroxysuccinimid)biotin (sulfoNHS-biotin; Pierce) Dimethylformamide (DMF) Blocking solution (Chimera Biotec) DNA-STV conjugates S.1 (Table 12.7.4; see Basic Protocol 1 or Chimera Biotec) TE buffer, pH 7.5 (APPENDIX 2A) Conjugate dilution buffer (Chimera Biotec) TETBS (see recipe) Rabbit anti–goat IgG conjugated with Cy5 (e.g., Chimera Biotec) Gel filtration columns (NAP10, Amersham Biosciences) Molecular cut-off ultrafiltration unit (Centricon 30, Millipore) Photometer (e.g., BioPhotometer, Eppendorf) Microarray slides (e.g., HP -slides; Chimera Biotec) functionalized with covalently coupled oligonucleotides complementary to DNA-STV conjugates (see bottom of Table 12.7.1) Slide boxes (e.g., Chimera Biotec) Microplate centrifuge (e.g., 5804R with rotor A-2-DWP, Eppendorf)
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Figure 12.7.5
Schematic drawing of the DDI method.
Table 12.7.4 Components of DNA-Antibody Conjugates (S.3)
Conjugate
Oligonucleotide moietya
Biotinylated antibody
S.3A
tA (S.1A)
Goat anti–mouse IgG (S.2A)
S.3B
tB (S.1B)
Goat anti–human IgG (S.2B)
S.3C
tC (S.1C)
Goat anti–rat IgG (S.2C)
S.3D
tD (S.1D)
Goat anti–guinea pig IgG (S.2D)
a For sequences of thiolated oligonucleotides tA to tD, see Table 12.7.1.
Adhesive hybridization chambers, 25 µL (e.g., Chimera Biotec) Microscope slide holder (for centrifuge; e.g., Chimera Biotec) Microarray laser scanning system (e.g., Axon 4000B, Axon) Biotinylate antibody 1. Remove the top and bottom caps of a disposable gel filtration column (NAP10) and pour off the conserving liquid. Support the column over a suitable receptacle and equilibrate the column by gravity flowthrough of three complete fillings with PBS2. 2. Apply 0.5 mg of an antibody (see Table 12.7.4) on top of the filterplate of the NAP10 column and allow the liquid to completely enter the gel bed. 3. Adjust the sample volume to 1 mL with PBS2. 4. Elute the antibody with 1.5 mL of PBS2 and collect the 1.5 mL of filtrate. 5. Transfer the mixture into a Centricon 30 ultrafiltration unit and reduce the volume to ∼200 µL by centrifuging 5 min at 2900 × g (4000 rpm) with a subsequent 5-min shaking step at room temperature. Collect the filtrate. 6. Dissolve 0.1 mg of NHS-biotin in 1 mL DMF (final 180 µM). Always prepare the NHS-biotin solution immediately before use.
7. Add 0.5 µL of 180 µM NHS-biotin solution to the antibody. Incubate for at least 30 min at room temperature with orbital shaking.
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Purify and quantitate biotinylated antibody 8. Transfer the mixture into a Centricon 30 ultrafiltration unit and reduce the volume to ∼50 µL by centrifuging 5 min at 2900 × g (4000 rpm). 9. Add 1 mL of PBS2 to the ultrafiltration unit and reduce the volume to ∼200 µL by centrifuging another 8 min. 10. Perform gel filtration (see steps 1 to 4). 11. Transfer the 1.5 mL filtrate into another Centricon 30 ultrafiltration unit and reduce the volume to ∼200 µL by centrifuging 5 min at 2900 × g (4000 rpm) with a subsequent 5-min shaking step at room temperature. Collect the filtrate. This protocol is suitable for most proteins, assuming that the protein is stable in phosphate buffer.
12. Determine the absorbance of biotinylated antibody at 280 nm. Calculate the concentration using the Lambert-Beer law: c = A/(ε × l) where the extinction coefficient (ε) of antibodies at 280 nm is 203,000 M−1 cm−1 . The final concentration of S.2 should be ∼10 µM.
Prepare DNA microarrays 13. Remove microarray slides from the package and, if stored in the freezer, equilibrate to room temperature. Microarrays with custom-immobilized oligonucleotides can be purchased from various manufacturers. In this protocol, HP -slides (Chimera Biotec) functionalized with covalently coupled oligonucleotides (see bottom of Table 12.7.1) are used. The slides are stored at −20◦ C until use.
14. Place slides in a slide box, add ∼22 mL blocking solution, and incubate 30 min with gentle agitation. For best results, the prehybridization of the microarrays should be performed with blocking solution in a large volume to ensure equal blocking of the whole slide surface. The use of a slide box such as the one from Chimera Biotec is recommended for blocking and washing steps. Up to 5 arrays can be processed in parallel.
15. Remove the microarrays from the blocking solution, place in a new slide box, and wash with ∼22 mL ddH2 O for 1 min. 16. Dry the microarrays by centrifuging in a microplate centrifuge using a microscope slide holder for 1 min at 400 × g (1500 rpm). Drying slides with compressed nitrogen or air is also possible but may lead to higher fluorescent background and may adversely effect signal and background uniformity.
17. Fix adhesive hybridization chambers on top of the slides to avoid losing solution later during hybridization. Store the microarrays at 4◦ C until use (up to 6 weeks).
Prepare capture conjugate 18. Prepare the capture conjugates S.3A-D by mixing 0.01 mM stock solution of DNA-STV conjugates S.1A-D (Table 12.7.4) with equimolar amounts (0.01 mM stock solution) of one of the biotinylated antibodies S.2A-D (Table 12.7.4), each in a total volume of 10 µL TE buffer, as shown in Figure 12.7.1A. Incubate 15 min at room temperature. OligonucleotideStreptavidin Conjugates for Immobilization of Proteins
12.7.10 Supplement 22
19. Dilute each conjugate S.3A-D to 500 nM with conjugate dilution buffer and incubate for an additional 10 min at room temperature. 20. Adjust S.3A-D to a final concentration of 25 nM each in conjugate dilution buffer. Proceed directly to the next step. Current Protocols in Nucleic Acid Chemistry
Figure 12.7.6 Fluorescent images of the microarrays obtained from DDI. For the color version of this figure go to http://www.currentprotocols.com.
Perform DDI and detection 21. Add 25 µL of the diluted capture conjugates per array and hybridize for 120 min at room temperature. 22. Wash two times in a slide box with ∼22 mL TETBS for 4 min under gentle agitation. Never rinse arrays with ddH2 O after hybridization.
23. Add 25 µL per array of a 25 nM dilution of rabbit anti–goat IgG Cy5 conjugate in conjugate dilution buffer and incubate 60 min at room temperature. 24. Wash two times in a slide box with ∼22 mL TETBS for 4 min under gentle agitation. 25. Dry microarrays by centrifuging in a microplate centrifuge using a microscope slide holder for 1 min at 400 × g (1500 rpm). 26. Measure the fluorescence intensity of the signals (Fig. 12.7.6) using a microarray laser scanning system (e.g., Axon, by a pmt of 500 and 100% laser power). It is best to scan the arrays immediately after processing. If this is not possible, store at 4◦ C protected from light.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Loading buffer 50 mM Tris·Cl, pH 7 10% glycerol 100 mM DTT 0.1% bromphenol blue Store up to 1 year at 4◦ C PBS1 5.552 g KH2 PO4 8.947 g K2 HPO4 400 ml ddH2 O pH 7.5 Store up to 1 year at room temperature Current Protocols in Nucleic Acid Chemistry
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PBS2 16.7 mM KH2 PO4 83.3 mM K2 HPO4 150 mM NaCl pH 7.3 Store up to 1 year at room temperature PBSE 6.4 mM KH2 PO4 63.6 mM K2 HPO4 150 mM NaCl 5 mM EDTA, pH 7.5 Adjust to pH 6.8 with HCl Store up to 1 year at room temperature Running buffer 25 mM Tris·Cl, pH 7 192 mM glycine Store up to 6 months at room temperature TBSE 20 mM Tris·Cl, pH 7 150 mM NaCl 1 mM EDTA, pH 7.5 pH 7.3 Store up to 1 year at room temperature TETBS 20 mM Tris·Cl, pH 7 150 mM NaCl 5 mM EDTA, pH 7.5 0.05% Tween-20 Adjust to pH 7.5 with NaOH Store up to 6 months at room temperature COMMENTARY Background Information
OligonucleotideStreptavidin Conjugates for Immobilization of Proteins
Covalent DNA-STV conjugates In this unit, covalent conjugates of DNA and streptavidin (STV) are synthesized from 5 -thiolated oligonucleotides and recombinant streptavidin by chemical crosslinking, and the chief products—hybrids containing a single oligonucleotide moiety per STV—are purified by chromatography. The use of these basic components as universal adaptors for tagging antibodies with a DNA oligomer has several advantages. STV (mol. wt. ∼56 kDa) is a homotetrameric protein from Streptomyces avidinii that demonstrates a remarkable and unique interaction with its low-molecular-weight ligand D-biotin. The molecular structure of STV
(Fig. 12.7.7) is highly conserved and is homologous to the homotetrameric protein avidin (mol. wt. ∼60 kDa). The biomolecular recognition of the water-soluble molecule biotin (vitamin H, Fig. 12.7.7) by streptavidin is characterized by the extraordinary affinity constant of ∼1014 dm3 mol−1 , indicating the strongest ligand-receptor interaction currently known (Weber et al., 1989). Another great advantage of STV is its extreme chemical and thermal stability. Streptavidin is resistant to many proteases including proteinase K under physiological conditions, and can be heated repeatedly at temperatures needed for PCR cycling with no apparent damage. It survives extreme pH conditions and can still bind biotin, for example, even in the presence of 7 M urea.
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Figure 12.7.7
Molecular structure of STV (Weber et al., 1989) and D-biotin.
Short DNA oligonucleotides are also powerful tools in biomedical diagnostics because of their specificity during stringent hybridization, which allows any unique DNA sequence (16 to 20 nt) in a target with the complexity of a mammalian genome (∼3 × 109 bp) to be detected specifically and, in principle, isolated. The power of DNA as a molecular tool is enhanced by the ability to synthesize virtually any DNA sequence by automated methods, and to amplify any DNA sequence from microscopic to macroscopic quantities by PCR. Another very attractive feature of DNA is the immense rigidity of short double helices (15 to 60 bp), which behave effectively like a rigid rod spacer between two tethered binding sites on both ends. Immobilization efficiency and site selectivity Various studies (Niemeyer et al., 1994, 1999; Wacker et al., 2004) on the covalent DNA-STV conjugates show the high immobilization efficiency of DDI, which is only slightly affected by the size of the biotinylated protein attached. For example, comparison to covalent immobilization methods indicated that the reversible DDI method is much more efficient than conventional irreversible immobilization techniques (Niemeyer et al., 1999; Wacker et al., 2004). This was attributed to the reversibility of DNA hybridization, enabling denser packing during formation of the protein layer on the substrate. In addition, the lean structure of the rigid double-helical DNA spacer between the surface and protein may also contribute to a larger effective surface
area. Also, a higher biological activity was demonstrated, possibly as a result of the larger distance between the surface and immobilized protein, enabling a more homogeneous type of reaction during enzymatic substrate transformation. The detailed comparison of various immobilization techniques for generating antibody microarrays, taking particular consideration of the DNA-directed immobilization of antibodyDNA conjugates as a method to generate versatile and robust protein arrays, was recently reported by Wacker et al. (2004). The DDI method was compared to direct spotting of antibody as well as immobilization using biotinstreptavidin interaction. Although all three differently prepared antibody arrays allowed for detection of low antigen quantities (as low as 150 pg/mL), the microarrays prepared by DDI and direct spotting displayed the best fluorescence intensities and allowed sensitive detection of antigens even at low concentrations of the capture reagent. The DDI-based microarray also revealed the best spot homogeneity and the lowest intra- and inter-assay standard deviations. For optimization of DDI, the generation of non-crossreacting DNA sequences is a point of particular interest. The functionality of the sequences in DDI-based assays essentially depends on large and uniform hybridization efficiencies combined with low non-specific crossreactivity between the individual sequences, as these parameters define the site-selective immobilization and the efficiency of DDI (Feldkamp et al., 2004).
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Previous studies have shown that shortening of the oligonucleotide lengths results in decreased binding (Niemeyer et al., 1998). Moreover, the solid-phase hybridization efficiency decreases with increasing stability of sequence-specific secondary structures, i.e., the formation of homodimers and intramolecular hairpin loops, which preferentially occur in GC-rich oligomers. To address this problem, the in vitro evaluation of in silico–designed oligonucleotide sequences has recently been used by Feldkamp et al. (2004). This was the first study to demonstrate the comparison of sequence motif properties theoretically predicted by their performance in real life, and thus represents one crucial step towards the systematic development of software tools for DNA sequence design for applications in supramolecular self-assembly such as DDI-based microarray technologies (Winssinger et al., 2001; Niemeyer, 2002), DNA computing (Parker, 2003), and DNA nanoconstruction (Seeman, 2003). Reversibility DDI allows for loading distinct amounts of biomolecules and for complete regeneration of DNA-coated surfaces (Niemeyer et al., 1998). Hence, an advantage of the DDI technique is its usefulness in recovering and reconfiguring expensive BIAcore sensor chips or glass substrates used in ellipsometric measurements.
OligonucleotideStreptavidin Conjugates for Immobilization of Proteins
Applications A DDI micro-fluorescence-immunoassay (µFIA) was used by Wacker and Niemeyer (2004) in a model system for the simultaneous detection and quantification of four different protein antigens, the tumor marker human carcinoembryonic antigen (CEA), recombinant mistletoe lectin rViscumin (rVis), ceruloplasmin (CEP), and complement-1inactivator (C1A) in human blood serum samples. With the DDI-µFIA, it was possible to detect low antigen quantities (as low as 400 pg/mL) comparable to the sensitivity of conventional enzyme-amplified microplate immunoassays. Moreover, the DDI-µFIA also revealed an extraordinary robustness and reproducibility, which can also be improved by using an internal standard, employing proteins CEP or C1A, which intrinsically occur in human blood serum. In particular, this normalization allows compensation for inter-assay deviations resulting from the use of various batches of microarray slides. Furthermore, the study
demonstrated that the DDI-µFIA can be carried out in a single step, taking advantage of the specificity of DDI of immuno-complexes formed in solution, thereby significantly reducing the time and costs of analyses.
Critical Parameters and Troubleshooting The synthesis of covalent conjugates of DNA and STV (see Basic Protocol 1) is relatively short, straightforward, and efficient. However, some handling expertise is essential. First, it is strongly recommended to precisely quantify the purchased oligonucleotides and to analyze their degree of modification. In addition, problems may occasionally arise from loss of intermediates or products during the various gel filtration steps. These are normally related to clogged membranes and may be avoided by extensive repeated shaking of the filtration tubes. In addition, during the use of the FPLC system, problems might arise from the quality of the column material, leading to an insufficient separation of product peaks. It is strongly recommended to use original columns by Amersham Biosciences and to carefully design the purification method, i.e., flow rate and buffer gradient. The DDI of proteins (see Basic Protocol 2) is an unproblematic method based on self-assembly. Nevertheless, two points need consideration. On the one hand, it is important to use microarrays with a very high density of reactive groups on the surface (e.g., HP -slides) to optimize the immobilization efficiency of the DDI method. On the other hand, the design of sequences for the simultaneous immobilization of more than four proteins is difficult because of the nonstringent conditions that must be used, but may lead to an enhanced level of cross-reactivity of the oligonucleotides. To overcome the aforementioned problems of the conjugate synthesis and the development of DDI-based protocols, purchasing the DNASTV conjugates and other components for DDI applications from Chimera Biotec may be a consideration.
Anticipated Results Since the synthesis of the covalent DNASTV conjugates is well established, one may expect good to moderate isolated yields of the conjugates. The application of the DNA-STV conjugates in DDI should lead to highly active protein microarrays that can be used for a wide variety of immunoassays in both fundamental research and clinical diagnostic projects.
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Time Considerations The synthesis of the covalent conjugates of DNA and STV can be accomplished in 1 to 2 days. The time required for DDI of proteins on DNA microarrays is 3 to 4 hr, assuming that microarrays with complementary oligonucleotides to the DNA-STV conjugates are available.
Literature Cited Angenendt, P., Glokler, J., Murphy, D., Lehrach, H., and Cahill, D.J. 2002. Toward optimized antibody microarrays: A comparison of current microarray support materials. Anal. Biochem. 309:253-260.
Niemeyer, C.M., Boldt, L., Ceyhan, B., and Blohm, D. 1999. DNA-directed immobilization: Efficient, reversible, and site-selective surface binding of proteins by means of covalent DNAstreptavidin conjugates. Anal. Biochem. 268:5463. Parker, J. 2003. Computing with DNA. EMBO Rep. 4:7-10. Pirrung, M.C. 2002. How to make a DNA chip. Angew. Chem. Int. Ed. Engl. 41:1276-1289; Angew. Chem. 114: 326-1341. Schweitzer, B. and Kingsmore, S.F. 2002. Measuring proteins on microarrays. Curr. Opin. Biotechnol. 13:14-19. Seeman, N.C. 2003. DNA in a material world. Nature 421:427-431.
Angenendt, P., Glokler, J., Sobek, J., Lehrach, H., and Cahill, D.J. 2003. Next generation of protein microarray support materials: Evaluation for protein and antibody microarray applications. J. Chromatogr. 1009:97-104.
Templin, M.F., Stoll, D., Schrenk, M., Traub, P.C., Vohringer, C.F., and Joos, T.O. 2002. Protein microarray technology. Trends Biotechnol. 20:160166.
de Wildt, R.M., Mundy, C.R., Gorick, B.D., and Tomlinson, I.M. 2000. Antibody arrays for highthroughput screening of antibody-antigen interactions. Nat. Biotechnol. 18:989-994.
Wacker, R. and Niemeyer, C.M. 2004. DDI-µFIA— A readily configurable microarray-fluorescence immunoassay based on DNA-directed immobilization of proteins. Chem. Bio. Chem. 5:453459.
Feldkamp, U., Wacker, R., Banzhaf, W., and Niemeyer, C.M. 2004. Microarray-based in vitro evaluation of DNA oligomer libraries designed in silico. Chem. Phys. Chem. 5:367-372. Kusnezow, W. and Hoheisel, J.D. 2002. Antibody microarrays: Promises and problems. BioTechniques Suppl:14-23. Lueking, A., Horn, M., Eickhoff, H., Bussow, K., Lehrach, H., and Walter, G. 1999. Protein microarrays for gene expression and antibody screening. Anal. Biochem. 270:103-111. MacBeath, G. and Schreiber, S.L. 2000. Printing proteins as microarrays for high-throughput function determination. Science 289:1760-1763. Niemeyer, C.M. 2002. The developments of semisynthetic DNA-protein conjugates. Trends Biotechnol. 20:395-401. Niemeyer, C.M., Sano, T., Smith, C.L., and Cantor, C.R. 1994. Oligonucleotide-directed self-assembly of proteins: Semisynthetic DNA– streptavidin hybrid molecules as connectors for the generation of macroscopic arrays and the construction of supramolecular bioconjugates. Nucl. Acids Res. 22:5530-5539.
Wacker, R., Schroeder, H., and Niemeyer, C.M. 2004. Performance of antibody-microarrays fabricated by either DNA-directed immobilization, direct spotting or streptavidin-biotin attachment: A comparative study. Anal. Biochem. 330:281287. Weber, P.C., Ohlendorf, D.H., Wendoloski, J.J., and Salemme, F.R. 1989. Structural origins of highaffinity biotin binding to streptavidin. Science 243:85-88. Winssinger, N., Harris, J.L., Backes, B.J., and Schultz, P.G. 2001. From split-pool libraries to spatially addressable microarrays and its application to functional proteomic profiling. Angew. Chem. Int. Ed. Engl. 40:3152-3155.
Contributed by Ron Wacker Chimera Biotec GmbH Dortmund, Germany Christof M. Niemeyer Universit¨at Dortmund, Fachbereich Chemie Dortmund, Germany
Niemeyer, C.M., B¨urger, W., and Hoedemakers, R.M. 1998. Hybridization characteristics of biomolecular adaptors, covalent DNA– streptavidin conjugates. Bioconjugate Chem. 9:168-175.
Nucleic Acid-Based Microarrays and Nanostructures
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CHAPTER 13 Nucleoside Phosphorylation and Related Modifications atural nucleic acids (DNA, RNA) contain phosphodiester groups, dinucleoside pyrophosphates (as intermediates in nucleic acid ligation), and dinucleoside triphosphates (mRNA caps). Nucleoside triphosphates are substrates for the biochemical synthesis of nucleic acids, and nucleoside analog triphosphates are frequently used as substrates for the synthesis of modified nucleic acids and as components in tools for molecular techniques in nucleic acid biology. Derivatives and analogs of natural nucleosides are frequently studied as their mono-, di-, and triphosphate derivatives in studies of biochemical processing. In many cases these studies stem from interest in these analogs as therapeutics, in other cases as tools for probing and dissecting the function of phosphorylated nucleoside intermediates in biochemical pathways including nucleoside metabolism, co-factor function, and signal transduction.
N
There are many effective chemical reagents and strategies for transforming nucleosides to nucleoside monophosphates (nucleotides). However, the transformation of nucleotides or nucleotide derivatives to their di- and triphosphates is often a challenge. The nature of the nucleobase as well as the presence of certain kinds of substituents can strongly influence the outcome of both chemical and enzymatic phosphorylation. In addition, there is interest in analogs derived from modified phosphates, including phosphorothioates and phosphoramidates. There is no one optimal way to construct all of the different types of phosphorylated nucleosides. The goal of this chapter will be to provide a cross-section of the best methods for different analogs and situations. The first unit, UNIT 13.1, provides a complete overview of nucleoside phosphate and polyphospate synthesis. The unit critically discusses the issues and problems associated with these syntheses. Classical chemical methods for synthesis of nucleoside monophosphates are discussed, and strategies for synthesis of diphosphates, triphosphates, and cyclic phosphates are presented. A substantial portion of the unit is devoted to nucleotide analogs of interest to nucleic acid chemists, including thio-phosphorous derivatives, phosphonates, and imidophosphates. A section on 32 P-radiolabeled derivatives is included as well. Finally, there are sections on phosphosulfate and enzymatic nucleotide synthesis. The most common rapid method for synthesis of nucleoside triphosphates is the Yoshikawa procedure that is often done as a single continuous operation. One of the most effective versions of this process is provided in UNIT 1.5. However, analogs containing sensitive modified bases cannot always be phosphorylated by the Yoshikawa procedure. In UNIT 13.2, Wu et al. outline effective procedures for the synthesis of riboand deoxyribonucleoside 5 -triphosphates of azole carboxamide-containing nucleoside analogs using a combination of chemical and enzymatic methods. The synthesis of oligonucleotides containing phosphorus-nitrogen linkages within the backbone provides an interesting challenge for nucleic acid chemists. UNIT 4.7 describes the step-wise chemical synthesis of oligonucleotide N3 -P5 phosphoramidates via the phosphoramidites of N-trityl-protected 3 -deoxy-3 -amino nucleoside analogs. In contrast, UNIT 13.3 outlines the synthesis of oligonucleotide N5 -P3 phosphoramidates by way of Nucleoside Phosphorylation and Related Modifications Contributed by Donald E. Bergstrom Current Protocols in Nucleic Acid Chemistry (2006) 13.0.1-13.0.2 C 2006 by John Wiley & Sons, Inc. Copyright
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DNA polymerase-mediated synthesis using triphosphate derivatives of 2 ,5 -dideoxy-5 aminoribonucleosides. The syntheses of the individual 5 -amino nucleoside derivatives are described, as well as a simple procedure for their direct conversion to triphosphate derivatives via reaction with trimetaphosphate. These nitrogen-containing analogs of nucleoside triphosphates are demonstrated to be effective substrates for Klenow polymerase (exo− ). Dinucleotide pyrophosphates containing a variety of heterocyclic and monosaccharide modifications are of considerable interest for biochemical studies on enzymes and pathways that utilize the natural dinucleoside pyrophosphates NAD and FAD. UNIT 13.4 provides experimental details for the classical phosphorimidazolide route to these compounds. Two procedures are included for the preparation of phosphorimidazolides, which differ based on the structure of the monosaccharide, and their coupling to monophosphates to yield dinucleoside pyrophospates is described. The synthesis of methylenebis(phosphonate) analogs of dinucleotide pyrophosphates requires a different strategy. As described in UNIT 13.5, a methylenebis(phosphonate) moiety is introduced by allowing the 2 ,3 -isopropylidine derivative of the first nucleoside to react with methylenbis(phosphonic acid dichloride) to yield, after aqueous hydrolysis, a mononucleotide 5 -methylenbis(phosphonate). This is then coupled to a second nucleoside using diisopropylcarbodiimde to yield the nonsymmetrical dinucleotide pyrophosphate. The method is general and can be extended to alcohols other than nucleosides. Phosphoramidite-based reagents for solid-phase monophosphorylation of oligonucleotides such as (2-cyanoethoxy)-2-(2 -O-4,4 -dimethoxytrityloxyethylsulfonyl)ethoxyN,N-diisopropylaminophosphine are commercially available. In UNIT 13.6, synthesis and phosphorylation procedures are described for an alternative reagent, bis[S-(4,4 dimethoxytrityl)-2-mercaptoethyl]-N,N-diisopropylphosphoramidite. The advantage of this latter reagent is that it allows one to produce both phosphates and thiophosphates at room temperature, a property that is important for the preparation of oligonucleotides containing other temperature-sensitive functional groups. Donald E. Bergstrom
Introduction
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Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates Phosphorylated nucleosides play a dominant role in biochemistry. Primary metabolism, DNA replication and repair, RNA synthesis, protein synthesis, signal transduction, polysaccharide biosynthesis, and enzyme regulation are just a handful of processes involving these molecules. Literally thousands of enzymes use these compounds as substrates and/or regulators. The need to obtain such compounds in both labeled and unlabeled forms, as well as a burgeoning need for analogs, has driven the development of a myriad of chemical and enzymatic synthetic approaches. As chemical entities, few molecules possess the wide array of densely packed functionality present in phosphorylated nucleosides. This poses a formidable challenge to the synthetic chemist, one that has not yet been fully overcome. This overview will address some common methods (synthetic and enzymatic) used to construct phosphorylated nucleosides. Particular emphasis is placed upon the advantages, limitations, and generality of the methods. Additionally, the synthesis of phosphorylated analogs such as phosphorylsulfates, phosphoramidates, phosphonates, thiophosphates, and imidophosphates will be addressed. This overview is not intended as a comprehensive review, but rather as a guide to general principles associated with the synthesis of phosphorylated nucleosides.
ISSUES ASSOCIATED WITH NUCLEOSIDE PHOSPHORYLATON In nature, the conversion of nucleosides to mono-, di-, or triphosphates is accomplished by specific enzyme-catalyzed reactions. Invariably, a nucleoside hydroxyl group functions as a nucleophile in coupling with an electrophilic phosphoryl donor (ATP, phosphoenol pyruvate, acetyl phosphate, and others). Binding constraints in the active site of the enzyme guarantee high regio- and chemoselectivity in the ensuing reaction. During chemical syntheses of nucleoside phosphates, the nucleoside can serve as either an electrophile or a nucleophile. Despite this flexibility, chemical syntheses are fundamentally more problematic than enzymatic ones. Phosphorylation procedures using electrophilic phosphorus reagents generally do not show high regioselectivity. Thus, it is often necessary to use protecting groups for the sugar, adding yet another level of complex-
ity to the synthesis. Side reactions can occur, such as depurination of the nucleoside, phosphorylation of the nucleobase, as well as chemical alteration of nucleobase analogs. Due to their intrinsic reactivity, the synthesis of phosphoanhydride bonds is also synthetically challenging. Phosphate anhydrides are phosphorylating reagents that are readily degraded under acidic conditions. Finally, purification of synthetic nucleotides can be problematic. Ionic reagents, starting materials, and mixtures of regioisomers (2′-, 3′-, 5′-phosphates) can be particularly difficult to separate from the desired product. In spite of the many potential difficulties associated with nucleoside phosphorylation and polyphosphorylation, a certain amount of success has been achieved in these areas. Given the wealth of phosphorylating reagents available, simple phosphorylation of nucleosides at any of the hydroxyl groups is possible, although selective phosphorylation at any particular hydroxyl group may require specific protection strategies. With sufficient attention to the nature of the nucleobase and protecting groups, the correct choice of phosphorylating agent should lead to a successful and high-yield synthesis. The synthesis of nucleoside diphosphates (NDPs) can also be succesfully performed in high yield. The excellent strategy introduced some time ago by Poulter for nucleophilic displacement of tosylates with tetran-butylammonium salts of pyrophosphate continues to be a very useful and successful strategy for simple preparation of diphosphates. While this is a multi-step procedure (particularly if protected nucleosides are used), the overall high yields, relative simplicity, and high reliability of this method make it one of the most useful preparative strategies available for the preparation of NDPs. In contrast, the preparation of nucleoside triphosphates (NTPs) and polyphosphate diesters remains somewhat less satisfactorily addressed. There is considerable hope in this area, however. Properly activated nucleoside monophosphates and their derivatives appear to be good acceptors for pyrophosphate and phosphate esters. The major problems here appear to be the highly variable yields reported for specific methods. This may simply be a consequence of the idiosyncratic nature of nucleoside
Contributed by David C. Johnson II and Theodore S. Widlanski Current Protocols in Nucleic Acid Chemistry (2003) 13.1.1-13.1.31 Copyright © 2003 by John Wiley & Sons, Inc.
UNIT 13.1
Nucleoside Phosphorylation and Related Modifications
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vinyl amine nucleophiles
ureido nucleophiles
O E
NH N
O
E
NH
H
O
O
O
N
NH
E
O
N
aryl amine nucleophiles E N
N
N
O
E O
ureido nucleophiles
H
NH2
NH2
N
N
E
N
E O
N
N
[Ox] O
N
O O
Figure 13.1.1 Some side reactions of nucleobases.
chemistry (in that different nucleosides are chemically very different), or may be related to the inherent difficulty in the reactions themselves. Many laboratories that use these methods are not ideally set up for organic synthesis or have little experience in doing this kind of chemistry. Developing robust chemistry for polyphosphate synthesis that can be routinely performed in a minimum of time, with a minimum number of steps, remains a daunting task for nucleoside chemists.
CHEMICAL REACTIVITY OF NUCLEOSIDES Nucleosides as Nucleophiles
Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
Both components of nucleosides, the sugar and nucleobase, possess nucleophilic functionality. The hydroxyl groups present in the sugar moiety of nucleosides are more acidic and less nucleophilic than corresponding primary or secondary alcohols in non-nucleosides. The sugar component of a nucleoside ionizes in the pH range of 12 to 13 for ribo- and deoxyribonucleosides (Albert, 1973). The reduced nucleophilicity of sugar hydroxyl groups may permit nucleophilic moieties in the nucleobase to compete for electrophiles. Accordingly, understanding the chemical properties of nucleobases is essential for implementing successful syntheses of nucleoside phosphates. The traditional nucleobases, as well as common derivatives, share chemical functionality of similar reactivity (Fig. 13.1.1). Aryl and vinyl amines are common nucleophiles present
in the nucleobases. Many bases also contain nucleophiles such as lactam, ureido, and guanidinium moieties. These moieties have different ionization characteristics. For this reason, Hayakawa and co-workers (Uchiyama et al., 1993) have grouped the common nucleobases into two different categories based upon their acidity. In group I nucleosides, the sugar hydroxyls are the most acidic functionality present. In group II nucleosides, the nucleobase has functionality more acidic than the sugar hydroxyls. Adenosine and cytidine belong to group I, whereas guanosine, thymidine, and uridine belong to group II. Unfortunately, these groupings do not directly correlate with the nucleophilicity of the nucleobase functionality. Nucleobases have different nucleophilicity towards electrophiles, even under identical reaction conditions. Thus, many phosphorylation procedures rely upon protecting groups (UNIT 2.1) to facilitate regioselective phosphorylation.
Nucleosides as Electrophiles Nucleosides can be phosphorylated by a displacement reaction between phosphate and an electrophilic carbon of a nucleoside. To render a carbon electrophilic, the hydroxyl group must be converted into a leaving group of some kind (e.g., a halogen or sulfonate ester). There are two common approaches to convert the 5′-hydroxyl into an electrophile: (1) conversion to a relatively stable leaving group, or (2) in-situ activation of the 5′-hydroxyl.
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O O HO P O P O O
O
B
O O O P O P OH O O 1
TsO
O
B
O O O O P O P O P OH O O O 2
O OH
OH
CH3CN
OH
OH
O O O HO P O P O P O O O O
CH3CN
3 B = Ade 4 B = Cyt 5 B = Uri 6 B = Gua
B
O OH
OH
7 B = Ade
Figure 13.1.2 Nucleosides as electrophiles. Displacement of tosylate (Ts) by various phosphate species.
Nucleotides via stable intermediates Halogenation is one common method of activating a 5′-hydroxyl group. For example, synthesis of 5′-iodides is readily achieved with methyltriphenoxyphosphonium iodide (MTPI; Verheyden and Moffatt, 1970) as well as a wide variety of other reagents. However, tosylation of the 5′-hydroxyl with toluene sulfonylchloride has proven to be one of the best activation methods. Synthesis of nucleoside 5′-tosylates of adenosine, guanosine, uridine, cytidine, and thymidine has been described (Davisson et al., 1987). Poulter introduced the synthesis of 5′-nucleoside mono-, di-, and triphosphates by displacement of the tosylate leaving group (Davisson et al., 1987; Wu et al., 2003; also see UNIT 13.2). Displacement of tosylate by bis(tetra-nbutylammonium) phosphate or tris(tetra-nbutylammonium) pyrophosphate (S.1) gives the corresponding nucleoside 5′-phosphate or 5′-diphosphate (S.3-S.6; Fig. 13.1.2), respectively. Similarly, displacement of tosylate by tetrakis(tetra-n-butylammonium) triphosphate (S.2) gives the nucleoside 5′-triphosphate. However, this latter reaction has only been useful for the synthesis of ATP (S.7; Burgess and Cook, 2000). To successfully execute a displacement reaction, care must be exercised in preparation and handling of the hygroscopic tetra-n-butylammonium phosphate salts. One nice feature of displacement reactions is that either protected or unprotected nucleosides can be phosphorylated. Ketal-protected nucleosides combine with tris(tetra-n-butylammonium) pyrophosphate much more rapidly than their unprotected counterparts. Reactions with pro-
tected nucleosides are complete in just 2 hours, whereas 2 to 4 days are often necessary with unprotected nucleosides. It is possible that the vicinal diols of ribonucleosides participate in H-bonding with the pyrophosphate reagent, thus slowing the reaction. Additionally, the yields for the displacement reaction are higher for protected nucleosides (Dixit and Poulter, 1984), suggesting that in appropriate cases it may be worthwhile to use protecting groups in spite of the added synthetic steps. Nucleotides via in-situ activation: The Mitsunobu reaction The Mitsunobu reaction (Fig. 13.1.3) exemplifies in-situ electrophilic activation of an alcohol functional group (Kimura et al., 1979). Activation is achieved by conversion of alcohol S.8 to the corresponding alkoxyphosphonium salt (S.9). In-situ displacement of the alkoxyphosphonium salt by dibenzylphosphate gives nucleotide triester S.10 and triphenylphosphine oxide (S.11). This reaction is driven by the formation of triphenylphosphine oxide and a strong phosphate ester bond. Deprotection via hydrogenolysis of the benzyl protecting groups affords the nucleoside monophosphate. Protected pyrimidine nucleosides can be phosphorylated in dioxane or tetrahydrofuran (THF), usually at an elevated temperature (60°C). However, phosphorylation of unprotected pyrimidine nucleosides is usually conducted at room temperature in polar aprotic solvents such as N,N-dimethylformide (DMF) or hexamethylphosphorous triamide (HMPT), in yields exceeding 75% (Kimura et al., 1979). Syntheses involving purine nucleosides in polar aprotic solvents are complicated by intra-
Nucleoside Phosphorylation and Related Modifications
13.1.3 Current Protocols in Nucleic Acid Chemistry
Supplement 15
O
O EtO HO
O
B
OH (H)
OH 8
N N H PPh3
OEt
DMF (B = U, C, T) or Pyr (B = A, G)
Ph Ph P O Ph
B
O
O BnO P O
O (BnO)2 P O
O
B
OBn OH
OH (H)
OH
9
10
OH (H)
O + Ph P Ph Ph 11
Figure 13.1.3 Electrophilic activation of nucleoside alcohols. The Mitsunobu phosphorylation. Bn, benzyl; DMF, N,N-dimethylformamide.
NH2 N
N O (BnO)2 P O
N
N O OH
OH 12
Figure 13.1.4 The N3,5′-cyclonucleoside salt of adenosine, the product of the Mitsunobu reaction in polar aprotic solvent. Bn, benzyl.
molecular cyclization, forming N3,5′-cyclonucleoside salts (Fig. 13.1.4). Formation of the purine cyclonucleosides can be avoided by conducting the phosphorylation reaction in anhydrous pyridine (Saady et al., 1995a). Phosphorylation of nucleosides with modified purine nucleobases (e.g., N6-chloro- and N6-azidoadenine) are successful in pyridine. However, anhydrous pyridine is not a suitable solvent for phosphorylation of unprotected pyrimidine nucleosides, as low yields are observed (Kimura et al., 1979). Finally, the Mitsunobu reaction is unsuitable for nucleoside 3′-phosphorylation due to competing intramolecular cyclization, resulting in the formation of anhydronucleosides.
CHEMICAL REACTIVITY OF PHOSPHORUS Phosphorus as a Nucleophile Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
Phosphorous acids and ethers are ambident nucleophiles (Fig. 13.1.5), reactive on phosphorus or oxygen. Accordingly, the phosphorus coordination number, degree of esterification,
and reaction conditions dictate the site of alkylation. These criteria are especially important for three-coordinate compounds of phosphorus, which will be considered first. Phosphites are three-coordinate (σ3) phosphorus species containing three total bonds (λ3) and are exceptionally useful for nucleoside chemistry. Phosphites exist as either a configurationally stable structure or a pair or tautomeric structures, depending upon the extent of esterification (Fig. 13.1.5). Trialkyl phosphites are configurationally stable structures characterized by a nucleophilic phosphorus and are incapable of tautomerizing to an H-phosphonate form. These species are easily oxidized by electrophilic oxidants and are subject to air oxidation as well. In contrast, both dialkyl phosphites (S.15) and monoalkyl phosphites (S.16) exist primarily in the fourcoordinate H-phosphonate (σ4,λ5) tautomer. The H-phosphonate tautomer is resistant to oxidation. Therefore, oxidation of four-coordinate H-phosphonates proceeds by way of the three-coordinate phosphite tautomer. In con-
13.1.4 Supplement 15
Current Protocols in Nucleic Acid Chemistry
ambident nucleophiles O RO P OR′′ OR′
O RO P R′′ OR′
[Ox] RO P OR′′ OR′
13 R = R′= alkyl; R′′ = H
O Pyr, AcCl
15 R = R′ = alkyl; R′′= H 16 R = alkyl; R′ = R′′ = H
14 R = R′= alkyl; R′′ = H
OR O P OR′ 17 R = R′= alkyl
Figure 13.1.5 The principle reactive species of phosphites.
phosphoramidite ( σ3, λ3 )
phosphoramidate ( σ4, λ5) O RO P N OR
RO P N RO 18
R = alkyl
N
19
Figure 13.1.6 Examples of electrophilic amino-phosphorus compounds important in nucleoside chemistry.
trast to oxidation, O-alkylation occurs primarily through the H-phosphonate tautomer. Acylation of H-phosphonates is affected by the strength of the base used. In the presence of weak bases such as pyridine, mono- and dialkyl phosphites are typically O-acylated forming O-acylphosphite ester intermediates (S.17). Transesterification of O-acylphosphite esters readily occurs in alcoholic solvents. However, in the presence of a strong base, competitive P-acylation of the phosphonates can occur, forming acylphosphonates (UNIT 3.4). Acylphosphonates readily degrade without undergoing transesterification. Another phosphorus nucleophile is the fourcoordinate, five-bond (σ4,λ5) phosphoric acid. Unlike H-phosphonates, phosphoric acid is exclusively O-alkylated. However, O-alkylation of phosphoric acid has found only modest application in the synthesis of nucleoside phosphates, due to the limited solubility of phosphoric acid in organic solvents. Biphasic solvent systems and phase-transfer catalysts have been used to facilitate phosphorylation. Alkylammonium salts of phosphoric acid are quite soluble in organic solvents and are often used in
nucleotide synthesis (see Nucleosides via stable intermediates).
Phosphorus as an Electrophile There are two coordination states common to electrophilic phosphorus, the phosphite (three-coordinate) and phosphate (four-coordinate) states. Two important amine-containing phosphorus compounds that can function as electrophiles are phosphoramidites (σ3,λ3) and phosphoramidates (σ4,λ5) (Fig. 13.1.6). Phosphoramidites are three-coordinate species with sigma bonds to two alkoxy groups and an amino functionality. Phosphoramidates are four-coordinate species containing two alkoxy groups and an amino group bound to a phosphoryl group. The amino moiety of these electrophilic phosphorus species has a P-N bond that is considerably weaker than the P-O bonds (70 versus 86 kcal/mol; Corbridge, 1995). Further, the amino moieties retain their basicity due to weak pπ-dπ bonding interactions. The amino moieties can become activated under acidic conditions to become good leaving groups. However, the basicity of the amino group can be affected by delocalization of the nitrogen
Nucleoside Phosphorylation and Related Modifications
13.1.5 Current Protocols in Nucleic Acid Chemistry
Supplement 15
N HN N N
RO
RO
20a
P N
N N N N
P N H
RO
N N N N
RO
N P N N N RO RO
20b path a
21
18
22 Nuc OH
Nuc OH
path b RO P O Nuc RO 23
Figure 13.1.7 Mechanistic rationale of N,N-diisopropyl dialkyl phosphoramidite coupling with a nucleoside.
O O RO P O P OR OR OR
R′OH N
O RO P N OR
O RO P OR′ OR
R′OH
26
25
24
Figure 13.1.8 Tetrakis(alkyl) pyrophosphates break down in pyridine to active phosphorylating agents.
O RO P OH OR
+
O Nuc P O
O
O
O O RO P O P O
B Nuc
Pα
OR
O
OH 28 pKa ≅ 2
29
O
B Nuc
Pβ
O RO P Nuc OR
O HO P O
O
B
O OH
OH 27
+
30
31 pKa ≅ 7
Figure 13.1.9 Selectivity in nucleophilic attack upon tris(substituted) nucleoside pyrophosphates. Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
13.1.6 Supplement 15
Current Protocols in Nucleic Acid Chemistry
lone pair into aryl substitution on the amine, engendering stability under acidic conditions. Accordingly, aliphatic amines are essential. Diisopropylamine is widely used owing to its ease and purity in preparation, as well as the balance between its chemical stability and reactivity (McBride and Caruthers, 1983). In phosphoramidate chemistry, the amino moiety is often morpholine or imidazole. Phosphoramidates can directly combine with nucleophiles, but do so very sluggishly. In the presence of catalysts such as acidic azoles or divalent metals, phosphoramidates react much more readily with nucleophiles. Azoles such as imidazole, tetrazole, 1-methylimidazole, 4,5-dicyanoimidazole, and other nitrosubstituted variations are useful as acidic catalysts. Unlike phosphoramidates (σ4,λ5), phosphoramidites (σ3,λ3) require activation by an acidic azole (e.g., 1H-tetrazole S.20a, pKa = 4.76) to form two electrophilic intermediates: protonated phosphoramidite S.21 and tetrazolide S.22 (Fig. 13.1.7). Protonated phosphoramidite S.21 can react with tetrazole anion S.20b to form a tetrazolide intermediate (path a) or can directly capture a nucleoside (path b). Interestingly, breakdown of phosphoramidite S.18 is zero order in alcohol and second order in tetrazole (Nurminen et al., 1998). Other common electrophilic phosphorus motifs are phosphorus halides and phosphate anhydrides. Unlike phosphorus amines, phosphorous halides and phosphate anhydrides do not require activation for rapid reaction with nucleophiles. The general structure of a tetrakis(alkyl) pyrophosphate S.24 is shown in Figure 13.1.8. Because the reaction of an alcohol with tetrakis(alkyl) pyrophosphates is sluggish, a base is generally used to reduce the time of reaction. Pyridine cleaves tetrakis(alkyl) pyrophosphates at either phosphoryl center, forming dialkyl phosphate and the phosphorylpyridinium species S.25, an active phosphorylating agent. Unlike tetrakis(alkyl) pyrophosphate, in tris(substituted) nucleoside pyrophosphates (S.27; Fig. 13.1.9) either phosphoryl center (Pα or Pβ) can be attacked by a nucleophile. The selectivity is based upon which phosphoryl center is the better leaving group. A phosphoryl center becomes a better leaving group when it is part of the more acidic phosphate (e.g., S.28 versus S.31). Increased electron withdrawing characteristics of the alkyl substituents will make a phosphate more acidic and, hence, a better leaving group. Therefore, the phosphoryl center of the weaker acid (i.e., S.29) is attacked
by nucleophiles in tris(substituted) nucleoside pyrophosphates (Fig. 13.1.9). In principle, phosphates such as phosphoric acid and its cyclic-ester trimer (trimetaphosphate) contain electrophilic phosphoryl centers and should undergo transesterification reactions with alcohols. In practice, this reaction finds limited application. However, the reaction of trimetaphosphate with unprotected ribonucleosides is noteworthy. Monophosphorylation of either the 2′- or 3′-hydroxyl group of unprotected ribonucleosides (A, C, G, U) can be achieved in aqueous base. The reaction requires elevated temperatures if sodium trimetaphosphate is used (Tsuhako et al., 1984), but can be conducted at room temperature with tris(tetramethylammonium) trimetaphosphate (Saffhill, 1970). Regardless of which trimetaphosphate is used, prolonged reaction times are required (up to 4 days). Aside from the prolonged reaction times, the reaction suffers from a lack of regioselectivity. Mixtures of 2′-monophosphate and 3′-monophosphate nucleosides are obtained. Interestingly, deoxyribonucleosides are not phosphorylated by trimetaphosphate.
REAGENTS USED IN SYNTHETIC NUCLEOTIDE SYNTHESIS Phosphorus Oxychloride Phosphorylation of nucleosides with phosphorus oxychloride (POCl3) initially yields a nucleoside phosphorodichloridate. Phosphorodichloridates such as S.32 (Fig. 13.1.10) can be hydrolyzed, providing the nucleoside monophosphate (not shown). The reaction is conducted in a trialkylphosphate solvent with an excess of POCl3 at a low temperature (−5°C). However, phosphorylation under these conditions leads to mixtures of 5′- and 2′(3′)-nucleoside phosphates. Protection of the vicinal diols in ribonucleosides alleviates regioselectivity problems. Thus, phosphorylation of ketal-protected nucleosides proceeds in high yield (Yoshikawa et al., 1967). Unprotected nucleosides can be regioselectively 5′-phosphorylated by modifying the reaction conditions described above. The addition of water to the phosphorylating reagent results in selective 5′-phosphorylation of unprotected nucleosides in moderate to high yield (Yoshikawa et al., 1967). Despite the acidity of the reaction medium, depurination has not been reported to be a significant side-reaction. Phosphorus oxychloride is also used to synthesize nucleoside triphosphates. However, construction of nucleoside diphosphates has
Nucleoside Phosphorylation and Related Modifications
13.1.7 Current Protocols in Nucleic Acid Chemistry
Supplement 15
O O HO P O P O O O HO P O O
O HO P O O
B
O OH 33
OH
Cl
N
Cl
O P O Cl
B
O OH
O O HO P O P O O O
O O O HO P O P O P O O O O
OH
OH
32
OH
Cl Cl
H
B
34
Cl
35
O
O O P O O O P O O P O O
N H
B
O OH
35
OH
36 H2O
NTP
Figure 13.1.10 Synthesis of nucleoside triphosphates from phosphorodichloridates via trimetaphosphate intermediates.
Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
not been reported with this reagent. To understand why, it is beneficial to understand the mechanistic rationale behind the synthesis of nucleoside triphosphates using POCl3. As shown in Figure 13.1.10, nucleoside triphosphates are synthesized by adding a tributylamine/DMF solution of either inorganic ortho-phosphate (Mishra and Broom, 1991) or pyrophosphate (Ludwig, 1987) to the nucleoside phosphorodichloridate S.32. ortho-Phosphate or pyrophosphate displaces chloride from nucleoside phosphorodichloridate S.32, forming intermediate S.33 or S.34, respectively. Excess POCl3 present in the reaction medium combines with DMF to form the Vilsmeier reagent (S.35), which promotes the dehydration of intermediates S.33 and S.34, resulting in the formation of nucleoside 5′-trimetaphosphate (S.36; Glonek et al., 1974). Hydrolysis of the trimetaphosphate intermediate under buffered conditions gives the NTP as a stable product. Phosphorylation of nucleosides containing non-natural nucleobases is possible with the POCl3/trialkylphosphate reagent combination (Nairne et al., 2002). However, modifications of the nucleobase that are sensitive to acid (e.g., alkynyl substituents) are not always compatible with the POCl3 reagent. Addition of the proton sponge 1,8-bis(dimethylamino)naphthalene has been reported to mitigate side-reactions
promoted by the acidity of the reaction medium. Unexpectedly, the addition of the proton sponge reduced the reaction time for phosphorylation from 12 hours to 2 hours (Kovacs and Otvos, 1988). Finally, the POCl3 methodology is not particularly suitable for selectively introducing radiolabels at the Pβ or Pγ phosphoryl of nucleoside triphosphates. This is because hydrolysis of trimetaphosphate S.36 is nonselective. Hydrolysis can occur at either the Pβ or Pβ′ position, leading to a mixture of products.
Phosphorochloridites and Phosphorochloridates In general, both phosphorochloridite [(RO)2PCl] and p ho sphorochloridate [(RO)2POCl] reagents are O-protected. There are many protecting groups that have been developed. In the case of phosphorochloridite reagents, the protecting groups are removed after oxidation of the phosphite triester. Therefore, protecting groups need to be stable to oxidants such as peroxides, organic peracids, and molecular halogens. Protecting groups have been developed that are labile to acid, base, or alternate conditions (e.g., reductive fission or photolysis). The functionality inherent in the nucleoside will determine which protecting group is most suitable for any given application. Nucleosides with modifications in the nucleobase require the most judicious
13.1.8 Supplement 15
Current Protocols in Nucleic Acid Chemistry
PGO
O
B
(EtO)2PCl
37
PGO
B
O
I2/H2O
38 OH
OPG EtO
PGO
O
B
39 O P
OPG O EtO P O OEt
OPG OEt
I2
PGO
O
B
40 O OPG O P I
1)
O
2) Br2 3) base 4) deprotection
OEt
HO
O
B
41
O OH HO P O O Br
Figure 13.1.11 Phosphitylation as a method for NMP synthesis. PG, protecting group.
choice of protecting group, as the common nucleobases (A, C, T, G, U) are fairly compatible with many deprotection conditions. Phosphorochloridite reagents can be used to synthesize nucleoside monophosphates. The general strategy is outlined in Figure 13.1.11. Phosphitylation of protected nucleoside S.37 with diethyl phosphorochloridite produces the phosphite triester S.38. Oxidation of the phosphite triester is usually effected by peroxide, organic peracid, or aqueous iodine. Following oxidation, the trialkyl phosphate S.39 is then deprotected to provide the nucleoside monophosphate (not shown). However, phosphite intermediate S.38 can be oxidized with iodine in situ to give halophosphate S.40. Various nucleophiles can be trapped by S.40, and appropriate deprotection then affords the nucleoside diester S.41. This dual phosphitylation/phosphorylation strategy is useful for preparation of nucleoside diesters that are inaccessible by standard phosphoramidite methodology, such as nucleoside-enol diester S.41 (Stowell and Widlanski, 1995). The synthesis of nucleoside monophosphates via phosphorochloridites is conceptually straightforward, but can be experimentally challenging. First, phosphorochloridites are air-sensitive reagents that are usually prepared in low yield and purified by vacuum distillation. Second, phosphite triesters are labile towards acid. This can be problematic if the phosphite
triester is purified by silica-gel chromatography. Including triethylamine (1% to 2%) in the eluent, however, usually prevents significant decomposition of the phosphite triester during purification. Last, phosphite triesters can airoxidize, which is essentially an irreversible chemical modification. Phosphorochloridites can be used to synthesize nucleoside triphosphates as well. In Figure 13.1.12, phosphitylation of a protected nucleoside (S.42) with salicyl phosphorochloridite (S.43) is carried out in pyridine/dioxane or pyridine/DMF. The phosphite triesters produced are diastereomers and are not isolated; rather, transesterification of the salicyl moiety is effected by adding a solution of bis(tri-nbutylammonium) pyrophosphate/tributylamine in DMF. Complete transesterification leads to the formation of the cyclic phosphite species S.45. Oxidation of phosphite S.45 with aqueous iodine produces nucleoside 5′-trimetaphosphate. Concomitant hydrolysis of the nucleoside 5′-trimetaphosphate yields protected nucleoside triphosphates as the product. The ester protecting groups used to protect the nucleoside are removed by aminolysis. By this process, TTP was synthesized from the corresponding protected nucleoside in 72% yield (Ludwig and Eckstein, 1989). Phosphorochloridite reagents are known to phosphitylate nucleobases at ambient temperatures but not at low temperatures (−78°C; Imai
Nucleoside Phosphorylation and Related Modifications
13.1.9 Current Protocols in Nucleic Acid Chemistry
Supplement 15
O O HO
O
T
O 43
P
O Cl
O P O O
T
OAc
OAc 42
O
O O HO P O P OH O O
O O P O O P O O P O O
O
T
1) I2 /H 2O 2) NH3
TTP
OAc
45
44
Figure 13.1.12 Salicyl phosphorochloridite. A useful reagent for NTP synthesis.
Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
and Torrence, 1981). Given that temperature is a factor in nucleobase phosphitylation, the experimental conditions under which salicyl phosphorochloridite is used may promote nucleobase phosphitylation. The extent to which salicyl phosphorochloridite may phosphitylate nucleobases has not been thoroughly investigated. Experimental data in the original paper do not conclusively rule out nucleobase phosphitylation as a source of reduced yield. As an aside, N-protected nucleobases are recommended during construction of nucleoside H-phosphonates using salicyl phosphorochloridite (UNIT 2.6). Nucleobase phosphitylation can be of particular concern when constructing nucleotides that have nucleobase modifications. Typical yields for phosphorylation of nucleosides with modified nucleobases are <45%, with yields in the 15% to 30% range being common (Jurczyk et al., 1999; Trevisiol et al., 2000). Nucleobase phosphitylation may also be a concern when using salicyl phosphorochloridite with oligodeoxyribonucleotides. Recent application of the salicyl phosphorochloridite methodology on solid support provided a low yield (15% to 30%) of the desired 5′-triphosphate-capped oligonucleotide (Lebedev et al., 2001). Finally, due to the intermediacy of the nucleoside 5′trimetaphosphate, this methodology is unlikely to be used for selective radiolabeling of nucleoside triphosphates at the Pβ or Pγ positions. Phosphorochloridates share many similarities with phosphorochloridites. For example, many of the same protecting groups are used and similar side reactions are observed. However, phosphorochloridates provide direct access to nucleoside dialkyl-phosphates without the need for oxidation. Reactions with phos-
phorochloridates can be sluggish, requiring prolonged reaction times. The reaction time can be reduced through use of a nucleophilic catalyst such as 1-methylimidazole or iodide ion (Stromberg and Stawinski, 1987). These catalysts function by forming a more electrophilic phosphorylating agent. However, increasing the electrophilicity of the phosphorochloridate also results in increased nucleobase phosphorylation. Protecting groups that are strongly electron withdrawing also increase the electrophilicity of the phosphorochloridate. Unlike their three-coordinate counterparts, four-coordinate phosphorochloridates can be used to synthesize nucleoside 3′- or 5′-monophosphates from N-unprotected nucleosides. Chemo- and regioselective phosphorylation can be affected through “functional group activation” (Uchiyama et al., 1993). Treatment of an appropriate hydroxyl-protected, N-unprotected nucleoside with tert-butylmagnesium chloride results in the formation of a nucleoside alkoxide (Fig. 13.1.13). Group I nucleosides (adenosine and cytidine) require only one equivalent of strong base, whereas group II nucleosides (guanosine, thymidine, uridine) require two equivalents of base. Upon treatment of the alkoxide with phosphorochloridate, the nucleoside-dialkyl-phosphate is obtained in high yield. Competition experiments demonstrate the preferential deprotonation of the hydroxyl functionality over nucleobase functionality for group I nucleosides. This deprotonation is expected because the sugar hydroxyl is more acidic than the nucleobase functionality. Chemoselectivity for group II nucleosides is not explicable based upon acidity. The 3′magnesium alkoxide bis-protected thymidine S.54 (Fig. 13.1.14) is preferentially phospho-
13.1.10 Supplement 15
Current Protocols in Nucleic Acid Chemistry
Group I: adenosine and cytidine HO
A
O
t BuMgCl (1 eq)
ClMgO
O (EtO)2 P Cl
A
O
O (EtO)2 P O
A
O
THF O
O
O
O
O
47
46
O
48
Group II: thymidine, uridine and guanosine TBSO
49
O
T
TBSO
t BuMgCl (1 eq) THF
OH
50
O
O (EtO)2 P Cl
T
MgCl
51
O
O
53 O (EtO)2 P
OMgCl
TBSO
TBSO
T
(EtO)2
O P Cl
TBSO
52
OH
T
O
O
O T P (OEt)2
OH
Figure 13.1.13 Grignard-promoted chemoselective phosphorylations based upon acidity (group I) or reversibility (group II). TBS, tert-butyldimethylsilyl.
N TBSO
N
O
O MgCl
O CH3
N
N
O
N
TBSO
O
O
(EtO)2
TBSO
N
CH3 O
O
(0.1 eq)
+ OMgCl
O
O P Cl
OTBS
1 eq
50 eq
54
55
(75%)
O EtO P O OEt
56
Figure 13.1.14 Competition experiment establishing alkoxide as the more reactive phosphorochloridate acceptor. TBS, tert-butyldimethylsilyl.
rylated by a limiting amount of diethyl phosphorochloridate in the presence of a 50-fold excess of N5-magnesium iminoxide bis-protected thymidine S.55. This result corresponds to the more basic magnesium alkoxide being phosphorylated in preference to the weaker magnesium iminoxide base. Interestingly, treatment of 5′-protected thymidine S.49 with one equivalent of base results in phosphorylation of the iminoxide functionality (Fig. 13.1.13). However, formation of phosphoramidate S.52 is reversible. Intermolecular intercon-
version provides exclusively the O-phosphorylated product S.53 after 96 hr.
Phosphoramidites and Phosphoramidates Phosphoramidites (σ3,λ3) can be used for the synthesis of NMPs, although their most popular application is found in the synthesis of oligodeoxyribonucleotides (UNIT 3.3). Unlike phosphoramidates (σ4,λ5), the construction of a nucleotide from phosphoramidites requires a minimum of two synthetic steps. The first step
Nucleoside Phosphorylation and Related Modifications
13.1.11 Current Protocols in Nucleic Acid Chemistry
Supplement 15
Classic phosphorimidazolidate approach–suitable for dNTP synthesis O O HO P O
O
T
O N P O
N
CDI
O
O
O P O P OH T
O
O
O TTP
O 57
58
OH
OH
Reverse approach–suitable for NTP synthesis
O
O
HO P O O 60
G
O
CDI R= H
O
RO P O O
O
G
O 61 H3CO P N O ZnCl2
N O O O H3CO P O P O P O O O O
O
G
O 59
O
OH
OH
62
P O OH
R=
O
OH
OH
Figure 13.1.15 Phosphorimidazolidates as a phosphorylation strategy. CDI, carbonyldiimidazole.
O O P OH O O HO P O
O
B
O OH 63
(PhO)2
O P Cl
O O PhO P O P O OPh O
O
OH
OH
NDP
B
OH
64
O O O P O P OH O O
NTP
Figure 13.1.16 The anion exchange strategy for synthesis of NDPs and NTPs.
Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
is the coupling of a phosphoramidite to a nucleoside, forming a nucleoside-dialkylphosphite, followed by oxidation to form the nucleoside-dialkyl-phosphate. A nucleophilic functionality in the nucleobase may be oxidized under such conditions. The N1 and N3 positions of adenosine and cytidine, respectively, are examples of nucleobase moieties prone to oxidation. Fortunately, appreciable oxidation
of the nucleobase is not observed when mild oxidants are used (e.g., I2/H2O or tBuOOH). Phosphoramidate NMPs are finding recent application as NMP prodrugs (Zemlicka, 2002), as well as precursors for constructing NDPs and NTPs. A classic method for the synthesis of dNTPs via nucleoside phosphoramidates involves the activation of an NMP with carbonyldiimidazole (CDI) to give the nucleoside phosphorimidazolidate S.58
13.1.12 Supplement 15
Current Protocols in Nucleic Acid Chemistry
HO
O O O HO P O P O P S O O O
O
A
O O O HO P O P O P S O O O
I
B
O
66
U
O OTBS HO P O 67 S B = Uri; R = OH
B = Ade; R = H OH
O
OH
R
65
HO
O O
HO P O S
U
OTBS
O OH
U
OH
68
Figure 13.1.17 Synthesis of nucleoside phosphorothiol esters by the displacement strategy. TBS, tert-butyldimethylsilyl.
(Fig. 13.1.15). Addition of acidic pyrophosphate activates the phosphorimidazolidate toward nucleophilic displacement by pyrophosphate, giving the dNTP (Hoard and Ott, 1965). This method is rarely used for the synthesis of ribonucleoside triphosphates. Ribonucleosides with unprotected vicinal diols (S.59) readily form cyclic carbonates (S.60) when treated with CDI. Additionally, nucleoside phosphorimidazolidates can react sluggishly with phosphate or pyrophosphate, requiring prolonged reaction times or the use of catalysts such as 1-methylimidazole or divalent cations (MnII, CdII). Reversal of the general strategy, such that the NDP is the nucleophile and the phosphorimidazolidate (e.g., S.61) is the electrophile, results in a high yield of the desired NTP without the need for long reaction times. This strategy has been used with success for the synthesis of Pγ-methyl-capped guanosine 5′triphosphates (e.g., S.62) in the presence of ZnCl2 (Kadokura et al., 1997).
Di- and Triphosphates from Monophosphates: Use of Phosphorus Anhydrides Michelson (1964) described a general method termed “anion exchange” for the synthesis of NDP, NTP, and P1-nucleoside-5′-P2sugars. The method is compatible with unprotected nucleotides and gives good yields. In short, an alkylammonium NMP is phosphorylated with diphenyl phosphorochloridate (Fig. 13.1.16). The resultant nucleoside mixed phos-
phoanhydride (S.64) is exposed to the alkylammonium salt of phosphate or pyrophosphate in pyridine solution to give the NDP or NTP, respectively. The reaction is quite selective for the displacement of diphenylphosphate from the mixed phosphoanhydride S.64. The presence of the anionic oxygen on the Pα of S.64 renders the nucleoside phosphate a weaker acid than diphenylphosphate. The product pyrophosphate is resistant to cleavage because the product possesses relatively poor leaving groups.
NUCLEOTIDE ANALOGS Analogs may be derived by modification of the sugar, base, or phosphorus component of a nucleotide. Phosphate analogs have great utility for studying the mechanism and stereospecificity of enzyme-catalyzed reactions and for investigating biochemical pathways. Sugar and nucleobase analogs are also used widely as therapeutic agents and mechanistic probes. A method for synthesis of the triphosphate of modified nucleosides by a variation of the Yoshikawa procedure is presented in UNIT 1.5. The following section will focus on methodologies and issues associated with the synthesis of analogs of the phosphorus component of nucleotides.
Thio-Phosphorus Derivatives Synthesis of thio-phosphorus derivatives of nucleotides are particularly problematic. The sulfur atom can occupy either a bridging (phos-
Nucleoside Phosphorylation and Related Modifications
13.1.13 Current Protocols in Nucleic Acid Chemistry
Supplement 15
Cl N P MMTrO
O SH
B
OCE
MMTrO
O
69 N
S P
B
HO
O
B
S 71 HO P O OH
70 OCE
Figure 13.1.18 A phosphoramidite approach toward the synthesis of phosphorothiol esters. CE, cyanoethyl; MMTr, 4-monomethoxytrityl.
ing phosphoramidite methodology and displacement reactions at electrophilic 5′-carbons. However, the majority of bridging phosphorothiol esters are synthesized in oligonucleotides. Nevertheless, the methods should be applicable to the synthesis of nucleosides containing a phosphorothiol ester linkage. Both nucleoside 5′-tosylates and 5′-iodides have been displaced with phosphorothioates. The reactions are usually conducted in DMF and require prolonged reaction times (24 to 48 hr). Both purine and pyrimidine bases have been used with or without base protection. Syntheses of 5′-thiotriphosphates of uridine and adenosine have been described. Unlike the synthesis of ATP, synthesis of 2′,5′-dideoxyadenosine-5′-thiotriphosphate (S.66; Fig. 13.1.17) by displacement of the 5′-iodide of nucleoside S.65 was accomplished in poor yield (21%). A similar yield was reported for 5′-deoxyuridine-5′-thiotriphosphate (19%; Patel and Eckstein, 1997). Construction of dinucleotides containing a 5′-thioester linkage (e.g., S.68) have also been carried out by displacement reactions. Yields are similar for displacement of 5′-tosylates or 5′-iodides by de-
phorothiol ester) or a non-bridging (phosphorothioate) position in the phosphate component of the nucleotide. To appreciate the difficulties associated with thio-phosphorus nucleotide synthesis, a brief discussion of the properties of thio-phosphorus acids is warranted. Thio-phosphorus acids have similar acidity to the corresponding phosphate. For example, diethyl phosphate and O,O-diethyl phosphorothioate have pKa’s of 1.37 and 1.49, respectively (Quin, 2000). There are several competing factors that affect the acidity of a phosphorothioate. Although the sulfur atom is larger and should stabilize the conjugate base better than oxygen, competing effects such as solvation, substituent effects, and bond strengths also affect the acidity of thio-phosphorus acids. Synthesis of nucleoside phosphorothiol esters Bridging phosphorothiol esters are attractive as phosphate surrogates because they are achiral, thus reducing molecular complexity and easing purification. Several methods have been used to access these compounds, includ-
O S S O
Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
O
72
Figure 13.1.19 3H-1,2-Benzodithiol-3-one 1,1-dioxide. A soluble alternative to elemental sulfur.
13.1.14 Supplement 15
Current Protocols in Nucleic Acid Chemistry
O H P O
TMSO O
B
P O
TMSCl
OH
O
TMSO
B
S8
X HO P O
B
O
SH 75 X = O
OH
OH
OH
73
74
1) I2, H2O 2) deprotection 75 X = O
HO
O
S O P OH
B
H
+
S O P O
O P Cl
(EtO)2
1) S8 2) deprotection 79 X = S
or
O
B
H OPG
OPG 76
77
78
Figure 13.1.20 Synthesis of phosphorothioate nucleosides from H-phosphonate precursors. PG, protecting group; TMS, trimethylsilyl.
HS O O O P OR HO P O P O OH
O +
OH
O
O
HO P O P O OH OH
80 P(R)
SH P
OR
NDP kinase or hexokinase
O
HS
HO P O P O OH OH
81 P(S)
R=
O
O P
OR
80 P(R)
O OH
A
OH
Figure 13.1.21 Selective enzymatic degradation of contaminating diastereomers is a common method to separate mixtures of P(R) and P(S) triphosphorothioates.
o xy rib o- and r ibo nu cleoside 3′-phosphorothioates, respectively (Chladek and Nagyvary, 1972; Thomson et al., 1996). The phosphoramidite approach appears to be a more viable entry into 3′- or 5′-thionucleotides. However, most applications using phosphoramidites are relevant to the construction of oligonucleotides containing bridging phosphorothiol esters and not 5′-thionucleotides. One approach to obtain bridging phosphorothiol esters is outlined in Figure 13.1.18 (Cosstick and Vyle, 1990). Modification of this approach should be suitable for high-yield syntheses of 3′- or 5′-thionu-
cleotides, although this avenue has not been fully explored. Synthesis of nucleoside phosphorothioates Nucleoside phosphorothioates are commonly accessed either by oxidation of trialkyl phosphites with sulfur, or by condensation with protected thiophosphoryl agents. Hydrolysis of the alkyl protecting groups produces the nucleotide phosphorothioate. Nucleoside trialkylphosphites are commonly accessed via phosphoramidite methodology (see Phosphoramidites and Phosphoramidates). Oxidation of the trialkyl
Nucleoside Phosphorylation and Related Modifications
13.1.15 Current Protocols in Nucleic Acid Chemistry
Supplement 15
HO
O OH 82
Cl
B
S P Cl Cl
Cl
S P O
B
Cl
(RO)3 P O
OH (H)
O OH
O O P O S O P O O P O O
O O HO P O P O O O DMF
OH (H)
B
O
OH (H)
OH
84
83
H 2O
O
O HO P O S
B
O OH
O (PhO)2 P Cl
(PhO)2
O O P O P O
B
O
O
O
HO P O P O O O
S
OH
OH
86
O
O
HO P O P O P O O S O
OH
OH (H)
B
O
OH (H)
85
87
Figure 13.1.22 Synthesis of 5′-(1-thio)triphosphates via anion exchange or through thiophosphorochloridates.
O O S CEO P O P O P O S
O
T
O 2
O O 1) CEO P O P OH S O 89 2)
OAc 90
O O P O O
O
T
1)
2)
S8
OAc
O O P O S O P O O P O
O O HO P O P OH O O S8
O
S
O
O
OAc
1) Li2S 2) NH3
T
O O S HO P O P O P O O
S
O
O
T
S OH
OH 91
T
92
88
HO
O O O HO P O P O P O
O
93
Figure 13.1.23 Salicyl phosphorochloridite provides versatility. Chiral and achiral thio-phosphorus analogs can be synthesized from a common precursor. CE, cyanoethyl. Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
13.1.16 Supplement 15
Current Protocols in Nucleic Acid Chemistry
O HO P O O O (PhO)2 P
1)
18O 18O
P O
O
A
A
O O
18O
O
95 OMe
S OH
OH
HS
O
O HO P O O
2) chromatography
94
A
O
P
1) NaIO4 OH
OH A
O
SH O P O P O HO O
18O
2) H+ 3) HO
A
O OH
OH
97 P(S)
96 P(S) O
O OMe
Figure 13.1.24 Synthesis of terminal chiral phosphates of known stereochemical configuration.
phosphite can be carried out with elemental sulfur. However, oxidation with elemental sulfur is sluggish due to poor solubility of the sulfur oxidant. Alternative sulfur oxidants such as 3H-1,2-benzodithiol-3-one 1,1,-dioxide (S.72; Fig. 13.1.19) have been introduced to eliminate the solubility problems associated with elemental sulfur (Iyer et al., 1990). Unlike elemental sulfur, S.72 is soluble in acetonitrile, oxidizes phosphites readily, and does not modify nucleobases on prolonged exposure (Regan et al., 1992). However, the full scope and utility of oxidant S.72 has yet to be assessed. It is not known, for example, whether unprotected nucleosides are compatible with this reagent. Another method to access nucleoside phosphorothioates is by oxidation of H-phosphonates (Fig. 13.1.20). Thus, phosphorous acid can be condensed with an unprotected nucleoside in the presence of N,N′-ditolylcarbodiimide to give predominately the nucleoside 5′-H-phosphonate S.73. Silylation of the nucleoside 5′H-phosphonate with trimethylsilyl chloride (TMSCl) forms the intermediate silylphosphite S.74. Oxidation with elemental sulfur in pyridine then gives the nucleoside 5′-phosphorothioate S.75 (Chen and Benkovic, 1983). Although no protecting groups are necessary, this method is limited by the long reaction times needed for the condensation (∼3 days) and the low yields of the nucleoside 5′-H-phosphonates produced (30% to 64%). Additionally, some nucleoside di-H-phosphonates are produced as byproducts.
An alternate procedure to access 5′-phosphorothioates makes use of 9-fluorenemethyl H-phosphonothioate S.77. This reagent condenses readily (<3 min) with protected nucleosides in the presence of diethyl phosphorochloridate to give nucleoside H-phosphonothioate diester S.78 (Jankowska et al., 1997). The salient feature of this method is that phosphorothioates can be accessed via aqueous iodine oxidation of H-phosphonothioate S.78, or phosphorodithioates (S.79) can be obtained by sulfur oxidation of S.78. The method is suitable for many deoxyribonucleosides and gives good to high yields of the phosphorothioates and phosphorodithioates (Jankowska et al., 1998). Although nucleoside 5′-phosphorothioates are more complex to synthesize than nucleoside 5′-phosphates, the synthesis of nucleoside 5′di- and 5′-triphosphorothioates is even more complicated. Esterification of a nucleoside 5′phosphorothioate results in the formation of a new chiral center. The majority of chemical methods available to access nucleoside 5′-diand 5′-triphosphorothioates are nonstereoselective. Therefore, a mixture of P(S) and P(R) diastereomers are produced during the synthesis. Separation of the diastereomers by conventional chromatography is non-trivial. One popular strategy for obtaining a pure diastereomer is to enzymatically degrade the contaminating isomer (Fig. 13.1.21). For example, both nucleoside diphosphate kinase and hexokinase selectively hydrolyze the P(S) isomer of
Nucleoside Phosphorylation and Related Modifications
13.1.17 Current Protocols in Nucleic Acid Chemistry
Supplement 15
O RO P OR
O P OH OR
98 R = Bu4N 100 R = Bn
O RO P OR
O P
O P OH
OR
OR
99 R = Bu4N 101 R = Bn
Figure 13.1.25 Structural variations of phosphonates make them suitable for use with either displacement strategies or Mitsunobu reaction. Bn, benzyl; Bu, butyl.
Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
ATPαS leaving the pure P(R) isomer of ATPαS untouched (Eckstein, 1979). Despite being nonstereoselective, many chemical methods used to synthesize nucleoside triphosphates are applicable to the construction of nucleoside 5′-(1-thio)triphosphates. Unprotected nucleosides can be mixed with thiophosphoryl trichloride in a trialkylphosphate solvent to form the corresponding thiophosphorodichloridate (S.83; Fig. 13.1.22). Introduction of pyrophosphate to the reaction mixture results in formation of the nucleoside thio-trimetaphosphate S.84. Hydrolysis of thio-trimetaphosphate S.84 produces the nucleoside 5′-(1-thio)triphosphate S.85. This method has been applied to the synthesis of several nucleosides, including dA, dG, T, and U. Although it is an expedient route to nucleoside 5′-(1-thio)triphosphates, yields are typically low (Arabshahi and Frey, 1994). Michelson’s technique of anion exchange has also been applied to the synthesis of nucleoside 5′-(1-thio)triphosphates (Fig. 13.1.22). Thus, activation of AMPS and GMPS (S.86) with diphenyl phosphorochloridate followed by displacement with pyrophosphate gives the ATPαS and GTPαS in reasonable yield (Chen et al., 1983). The salicyl phosphorochloridite methodology has been applied to the synthesis of nucleoside 5′-(1,3-bisthio)triphosphates and nucleoside 5′-(1,1-dithio)triphosphates (Fig. 13.1.23; Ludwig and Eckstein, 1991). Transesterification of nucleoside phosphite triester S.88 with excess of P1-O-(cyanoethyl)-P1-thiopyrophosphate (S.89) followed by oxidation with elemental sulfur gives the branched pentaphosphate S.90. Selective hydrolysis along with concomitant deprotection of the cyanoethyl protecting group occurs under basic conditions to give thymidine 5′-(1,3-bisthio)triphosphate S.91. Selectivity for the hydrolysis follows the principles outlined earlier (see Phosphorus as an Electrophile).
For construction of nucleoside 5′-(1,1dithio)triphosphates, transesterification of nucleoside phosphite triester S.88 with pyrophosphate followed by oxidation with elemental sulfur leads to the formation of thiotrimetaphosphate S.92. Selective cleavage of thiotrimetaphosphate S.92 at Pα occurs in DMF with lithium sulfide. Selectivity is not observed when pyridine/dioxane is used as the solvent. Aminolysis of the protecting groups then provides thymidine 5′-(1,1-dithio)triphosphate S.93. Although the method produces S.93 in very low yield (15%), S.93 may serve as a good achiral phosphorothioate derivative. Frey has introduced a very elegant method for the synthesis of ATP with chiral α, β, or γ terminal phosphates of known configuration. The phosphates are rendered chiral by containing sulfur and heavy oxygen (18O). Thus, adenosine 5′-(1-thio,1-18O)diphosphorothioate, adenosine 5′-(2-thio,2-18O)diphosphorothioate, and adenosine 5′-(3-thio,3-18O)triphosphorothioate have been synthesized (Richard et al., 1978). The basic strategy is to selectively thiophosphorylate adenosine with thiophosphoryl chloride followed by hydrolysis with H218O, yielding S.94 (Fig. 13.1.24). Michelson’s anion-exchange reaction is used to displace diphenylphosphate with 2′,3′methoxy-methylidene AMP (S.95), providing diadenosine S.96. Separation of the diastereomers at this point ensures configurational purity of the phosphorus stereocenter in the product. Sodium periodate oxidation of the unprotected vicinal diols of S.96 cascades into complete degradation of the adenosine moiety, providing (2-thio)diphosphorothioate S.97. Similar methodology is used to obtain adenosine 5′-(1-thio,1-18O)diphosphorothioate and adenosine 5′-(3-thio,3-18O)triphosphorothioate.
13.1.18 Supplement 15
Current Protocols in Nucleic Acid Chemistry
Michaelis-Arbuzov chemistry
(BnO)2POMe
O BnO P
Cl
O O P OMe BnO P OBn OBn
103
OBn
O
KCN
104
102
O
P OH BnO P OBn OBn 105
Selenophosphonate anion chemistry
Se MeO P
Li
Cl2PNMe2
N P
Se MeO P
OBn 106
OBn
Se P OMe
1) BnOH, tetrazole 2) mCPBA
O MeO P
OBn
O P
O P OMe
OBn OBn OBn
107
108
Figure 13.1.26 Strategies to access phosphonates. mCPBA, meta-chloroperoxybenzoic acid.
Phosphonate Derivatives Replacing a bridging oxygen with a methylene functionality in pyrophosphate produces a pyrophosphonate (Fig. 13.1.25). The bond dissociation energy for a C-P bond is 65 kcal/mol, which is considerably less than the P-O bond dissociation energy (86 kcal/mol; Corbridge, 1995). Despite this, pyrophosphonates have considerably more thermal stability than phosphoanhydrides. In addition, pyrophosphonates are more resistant to hydrolysis, a consequence arising from the reduced electrophilicity of the phosphoryl group when bonded to the methylene unit. However, pyrophosphonate is neither a geometric nor an electronic isostere of pyrophosphate. The oxygenoxygen nonbonded distance in a pyrophosp honate is ∼16 % greater than in a pyrophosphate. Pyrophosphonate is not capable of accepting a hydrogen bond to the bridging atom like pyrophosphate can. Despite these differences, nucleoside pyrophosphonates are used as enzymatically stable, nonhydrolyzable surrogates for nucleoside pyrophosphates (Engel, 1977). Nucleoside di- and triphosphonates have been prepared either with a single bridging methylene unit replacing the bridging α-β or β-γ oxygens, or with total oxygen replacement in these bridging positions. Direct displacement reactions conducted on nucleoside 5′-tosylates with tris(tetrabutylammonium) pyrophosphonate (S.98) give the nucleoside 5′-py-
rophosphonates in good yield (Dixit et al., 1984). Similarly, displacement of 5′-tosylates with bis-methylene triphosphonates (S.99) give the nucleoside analogs with α-β and β-γ bridging methylenes (Stock, 1979). Significantly better yields are obtained for the displacement reaction when performed in acetonitrile at room temperature, as opposed to heating in DMF, although acetonitrile requires longer reaction times. The nice feature about accessing nucleoside phosphonates in this fashion is that no protecting groups are needed on either the nucleoside or the phosphonate. The Mitsunobu reaction has also been used in the construction of nucleoside phosphonates. Unlike the displacement methodology, protected nucleosides as well as protected phosphonates are usually used under Mitsunobu conditions. The reaction has been perfected such that both purine and pyrimidine nucleosides can be used without significant competing cyclization to the anhydro-nucleoside. Phosphorylation of protected nucleosides via the Mitsunobu reaction gives higher yields with tris(benzyl) pyrophosphonate (S.100) than with tetrakis(benzyl) bis-methylene triphosphonate (S.101). The yields for the phosphorylation step with tris(benzyl) pyrophosphonate are comparable to the displacement method on comparable nucleosides. Slightly better yields for the phosphorylation step with tetrakis(benzyl) bis-methylene triphosphonate are observed for the Mitsunobu reaction than with the
Nucleoside Phosphorylation and Related Modifications
13.1.19 Current Protocols in Nucleic Acid Chemistry
Supplement 15
O HO P O
A
O
(PhO)2
O P Cl
(PhO)2
O OH
OH
O
O O O H HO P N P O P O O O O
A
O O H O P N P OH O O
OH
OH
111 APPNP
O O H HO P N P O
creatine kinase phosphoenol pyruvate
A
O
O
O O O H HO P O P N P O O
OH
OH
O
B
O
O
OH
OH
113
112
A
O
OH
110
O OH
O O H O P N P OH O O
A
O
OH
109
TsO
O O P O P O O
OH
114 APNPP
creatine kinase O Cl HO
O OH
A
OH
115
Cl
P N P Cl Cl Cl 116
Cl
O Cl P N P O Cl
O
A
Cl
(MeO)3 P O
TEA-bicarb buffer
O O H HO P N P O O
OH
OH
117
O
A
O OH
OH
118
Figure 13.1.27 The common strategies used to access imidophosphate derivatives of adenosine. TEA, triethylamine.
Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
displacement method. Deprotection of the nucleoside benzyl-protected phosphonates is routinely performed with trimethylsilyl bromide or catalytic hydrogenolysis. Either method seems to work equally well. Protected pyrophosphonates can be constructed by the Michaelis-Arbuzov reaction (Saady et al., 1995b) or by phosphorylation of methane selenophosphonate anions (Eymery et al., 1999). The Michaelis-Arbuzov reaction allows ready access to tris(benzyl) pyrophosphonate (S.105; Fig. 13.1.26). However, experimental reproducibility of the MichaelisArbuzov reaction can be tricky. Recently, selenophosphonate anion chemistry has been developed that allows efficient access to bismethylene triphosphonates (Klein et al., 2002). Methaneselenophosphonate anion S.106 couples with dichlorophosphoroamidite producing selenophosphoramidite S.107. Owing to the reduced basicity of the selenophosphoryl moiety, competing deprotonation of the selenophosphoramidite intermediates is not observed. Tetrazole-promoted esterification of phosphoramidite S.107 is followed by oxidation to give bis-methylene triphosphonate S.108. The
diacid form of triphosphonate S.108 is revealed upon selective demethylation with potassium cyanide.
Imidophosphate Derivatives The chemistry of the imidodiphosphate and diimidotriphosphate functional groups closely mirrors that of the phosphoramidate functional group (see Phosphoramidites and Phosphoramidates). Thus, imidodiphosphates and diimidotriphosphates are stable in alkaline media, but hydrolyze in acidic media (Tomasz et al., 1988). Despite its acid lability, the imidophosphate functional group has been used in nucleotides as a phosphate analog. When either the α/β or β/γ oxygen of ATP is replaced with nitrogen, the result is adenosine imidotriphosphate, abbreviated APNPP and APPNP, respectively. If both the α/β and β/γ oxygens are replaced by nitrogen, the result is adenosine diimidotriphosphate, APNPNP. Adenosine imidotriphosphate mimics ATP for many applications. This mimicry may arise because the physical properties of the imidodiphosphate and pyrophosphate functional groups are quite similar (Larsen et al., 1969).
13.1.20 Supplement 15
Current Protocols in Nucleic Acid Chemistry
Although imidodiphosphate is not a geometric isostere in the truest sense, the bond angle and bond lengths are very close to those of pyrophosphate. For example, the P-N-P and P-O-P bond angles are 127.2° and 128.6°, respectively. The bond lengths are also very similar: 1.68 Å (P-N) and 1.63 Å (P-O). Unlike a phosphate ester linkage, an imidophosphate can, in principle, accept or donate a H-bond. However, the shortened P-N bond length suggests partial P=N characteristic of the imidophosphate, which mitigates the H-bond donating and accepting abilities. Various nucleosides with the imidophosphate structural motif are inhibitors of HIV-RT1 (Li et al., 1996) and ATP/GTP phosphatases (Batra et al., 1987), but are substrates for E. coli alkaline phosphatase (Yount et al., 1971). Nucleoside imidophosphates are widely used in their triphosphate form. As a result, synthetic methods have focused principally on the triphosphate form as the target. There are several approaches to synthesize nucleoside imidophosphates, and no single approach is superior. All methods give similar, yet variable, yields for attachment of the imidophosphate functionality to the nucleoside. Synthesis of nucleoside-β,γ-imidotriphosphates can be accomplished by Michelson’s anion-displacement methodology (Yount et al., 1971). Activation of AMP with diphenyl phosphorochloridate followed by displacement of diphenyl phosphate with imidodiphosphate gives APPNP (S.111; Fig. 13.1.27). This method has been used successfully to incorporate heavy nuclides of oxygen and nitrogen into the phosphate component by the labeling of imidodiphosphate (Reynolds et al., 1983). The preparation of nucleoside-α,β-imidotriphosphates is generally carried out in a two-step procedure. The nucleoside-α,β-imidodiphosphate is chemically synthesized and then enzymatically phosphorylated. Displacement of the tosylate leaving group in adenosine 5′-tosylate S.112 by tris(tetrabutylammonium) imidodiphosphate gives access to adenosineα,β-imidodiphosphate S.113 (Ma et al., 1988). Enzymatic phosphorylation of S.113 by creatine kinase using phosphoenol pyruvate as the phosphoryl donor provides access to APNPP (S.114). An alternate strategy uses imidophosphorylation of unprotected nucleosides with trichloro-[(dichlorophosphoryl)imido]phos phorane (S.116; Tomasz et al., 1988). Both ribo- and deoxyribonucleosides have been imidophosphorylated using phosphorane S.116. Phosphorane S.116 is also a good chlorinating
reagent, however, and yields nucleoside 5′chlorides as byproducts. Many purine and py rimidine nucleoside-α,β-imidodiphosphates obtained through imidophosphorylation have been converted to their triphosphate counterparts using creatine kinase (Li et al., 1996). Access to nucleoside-α,β-β,γ-diimidotriphosphates is very challenging; more so than imidotriphosphates. In fact, no satisfactory method is yet available. Adenosine-α,β-β,γ-diimidotriphosphate has been accessed by reaction between adenosine 5′-tosylate and the tetrabutylammonium salt of diimidotriphosphate in acetonitrile (Ma et al., 1990). The desired product was obtained in poor yield (7%) along with another diimidotriphosphate, AP(NP)2 (6%). Several imidophosphate derivatives of adenosine and guanosine are commercially available. However, use of commercial preparations requires HPLC purification prior to use to remove the contaminating phosphoramidate APPN (Penningroth et al., 1980; Batra et al., 1987). APPN does not reappear after prolonged storage (8 months) of repurified APPNP, suggesting the phosphoramidate contaminant is a byproduct of preparation. 32
P-Radiolabeled Derivatives
This section will focus on radiolabeled compounds containing 32P modifications to the phosphate component of nucleotides. There are several important provisos to consider when choosing a method to synthesize radiolabeled derivatives of nucleotides. First is the location of the radiolabel. Second is the availability of reagents in labeled form. Third is the time required for synthesis to be complete. Because the half-life of 32P is 14 days, prolonged syntheses reduce the specific activity of the labeled nucleotide. Taken together, these provisos place severe restrictions on the methodologies that are available to synthesize radiolabeled nucleotides. Nucleotides with radiolabeled phosphates are commercially available for common nucleoside mono- and triphosphates (C, G, T, dA, and others). The nucleotide triphosphates are widely available with 32Pα and 32Pγ labels. However, there is a conspicuous lack of 32Pβlabeled triphosphates. Therefore, synthetic methods are necessary to obtain 32Pβ-labeled nucleotides. Synthetic methods are also necessary to obtain radiolabeled mono-, di-, and triphosphates of nucleosides with sugar and/or nucleobase modifications. There are many methods available for the synthesis of nucleo-
Nucleoside Phosphorylation and Related Modifications
13.1.21 Current Protocols in Nucleic Acid Chemistry
Supplement 15
HO
119
O OH
1) Cl
A
OH
O O P O P Cl Cl Cl
2) H2O, pH= 7
O HO P O OH 120
A
O O
DCC ∆
O
O HO P O OH 121
A
O O
O
P O
OH HO P O OH
TEA-SO3
O O HO S O P O O OH 123
O
A
O OH HO P O OH
ribonuclease-T2
O O HO S O P O O OH 122
bovine spleen phosphodiesterase II
A
O
O O HO S O P O O
O
O
124
P O
OH
O
A
OH OH O O P OH OH
Figure 13.1.28 Combination methodology for the synthesis of PAPS. DCC, dicyclohexylcarbodiimide; TEA-SO3, triethylamine-N-sulfonic acid.
O N N H
N
125 Figure 13.1.29 4-Morpholine-N,N′-dicyclohexyl carboxamidine. A useful base for increasing solubility of NMPs.
Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
tides with 31P that cannot be used with 32P, because 32P is not available in as many coordination states as 31P. The most readily available form of 32P is an aqueous solution of orthophosphoric acid. Consequently, many of the phosphorylation reactions with 32P are conducted in dimethyl sulfoxide (DMSO), dimethylformamide (DMF), or water. Nucleoside phosphate mixed anhydrides and nucleoside phosphoramidates are electrophilic phosphorus species that are suitable for use with 32P-ortho-phosphoric acid. Treatment of NDPs with ethylchloroformate produces a nucleoside phosphate mixed anhydride. Reac-
tion of the mixed anhydride with 32P-orthophosphoric acid produces the 32P-labeled nucleoside triphosphate. This method is compatible with various NDPs (A, C, G, U) for the construction of 32Pγ-labeled NTPs. The reaction is complete in a short period of time, but requires extensive handling of the 32P reagent (Janecka et al., 1980). This method does not allow for the construction of 32Pβ-labeled NDPs, as the NMP mixed anhydride does not react to an appreciable extent with phosphoric acid (Michelson, 1964). The phosphoramidate approach relies upon ready access to the nucleoside phosphormor-
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pholidate. Condensation of AMP or ADP with morpholine in the presence of dicyclohexylcarbodiimide (DCC) gives the corresponding nucleoside phosphormorpholidates. 32Pβ- or 32Pγlabeled ATP can be obtained from AMP- and ADP-morpholidate, respectively. Reaction of AMP-morpholidate with 32Pβ-ρ-nitrobenzyl diphosphate gives 32Pβ-labeled ATP in low yield. Reaction between 32P-ortho-phosphoric acid and ADP-morpholidate gives 32Pγ-labeled ATP in good yield. The nice feature of this method is that commercially available unprotected nucleosides can be used. A drawback of the method is the vast excess of reagents used throughout the process. Both DCC and 32P-ortho-phosphoric acid are used in excess. This is disadvantageous because the urea byproduct of DCC can be difficult to separate from nucleotides, and excess 32P reagent contributes to additional radioactive waste. Additionally, the reaction requires prolonged reaction times, which results in reduced activity of the 32P-labeled nucleotides (Werhli et al., 1965). Introduction of the 32P label at the Pα of NDP or NTP is possible by condensation of a 2′,3′isopropylidene ribonucleoside with 32P-orthophosphoric acid in the presence of trichloroacetonitrile and triethylamine. Following the removal of the protecting group, the 32P-labeled monophosphate can be phosphorylated with (2-cyanoethyl)phosphoryl- or (2-cyanoethyl)pyrophosphoryl imidazolidate, providing 32Pα-NDPs and 32Pα-NTPs, respectively (Symons, 1970). Extension of this methodology with unprotected nucleosides is possible, although the first condensation step produces a mixture of 2′(3′)- and 5′-nucleoside monophosphates. This mixture can be selectively phosphorylated with enzymes to yield the desired 32P -NTPs or 32P -dNTPs (Symons, 1974). α α In addition to the purely chemical and mixed chemical/enzymatic syntheses of 32Pα-NTPs, a
complete enzyme-mediated process has been developed. The enzymatic process has the advantages that the reaction is performed in the container that the 32P is shipped in and the intermediates do not need to be purified. A consequence, though, is that the final purification can be difficult (Walseth and Johnson, 1979; Walseth et al., 1991). As with any enzymatic synthesis of nucleotides, if the phosphoryl donor is ATP, contamination of the product NTPs with ATP is possible. This is particularly problematic with 32Plabeled ATP, as the specific activity can be reduced significantly (Furuichi and Shatkin, 1977). This can be addressed by use of alternate phosphoryl donors such as phosphoenol pyruvate or acetyl phosphate. Enzyme systems have been developed for the synthesis of 32Pβ-NTPs from phosphorylation of NMPs (Furuichi et al., 1977) and by polynucleotide digestion with polynucleotide phosphorylase (Kaufmann et al., 1980).
3′-PHOSPHOADENOSINE-5′-PHOSPHOSULFATE (PAPS) 3′-Phosphoadenosine-5′-phosphosulfate (PAPS) is the sulfate source used by sulfotransferases for the sulfation of steroids, drugs, proteins, and saccharides (Klaassen et al., 1997). PAPS has been synthesized for direct use as a cofactor and to study biological processes. Methodologies developed for the synthesis of PAPS reflect the application. Methods combining both chemical synthesis and enzymes have been detailed. Synthetic methods typically have fallen short of providing PAPS in pure form, as both the 2′- and 3′-phosphate regioisomers are produced. Thus, synthesis of PAPS by pure synthetic methods will not be addressed.
Ph Ph 1) HO
126
O OH
B
OH
O OMe P O OMe OMe 127
2) H2O
HO
128
B
O O
O P
O
OH
Figure 13.1.30 Selective transesterification of unprotected nucleosides by oxyphosphorane leads to regioselective formation of 2′,3′-cyclic phosphates.
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Enzymatic Methods Pure enzymatic synthesis of PAPS is particularly suitable when PAPS is a cofactor used in sulfation reactions. In these instances a large quantity of PAPS is not necessary. Rather, what is needed is a means to reconvert PAP back to the coenzyme PAPS. Systems have been developed that directly convert PAP to PAPS (Burkart et al., 1999) or that accomplish this conversion through several enzymatic steps (i.e., PAP → AP → APS → PAPS; Lin et al., 1995). The one real disadvantage of this methodology is its level of sophistication. Although very elegant, multienzymatic regeneration of PAPS requires construction of a plasmid, as well as overexpression and purification of several enzymes (Burkart et al., 1999). These procedures are routine in molecular biology laboratories, but may be intimidating to the synthetic chemist.
Combination Methods Figure 13.1.28 summarizes methods to obtain PAPS by combination methodology. The key to several combination methods is the availability of 2′,3′-cyclic phosphoadenosine 5′phosphosulfate (S.122). Several approaches are available to access this intermediate. Adenosine (S.119) can be phosphorylated with pyrophosphoryl chloride followed by hydrolysis to give 2′,3′-cyclic phosphoadenosine 5′phosphate (S.120; Horwitz et al., 1977). Alternately, cyclic phosphate S.120 can be obtained by cyclizing a commercial mixture of 2′,5′- and 3′,5′-bis(phospho)adenosine (S.121) with DCC at elevated temperature (Sekura, 1981). Sulfation of cyclic phosphate S.120 is most commonly effected with triethylamine-N-sulfonic acid (Horwitz et al., 1980; Sekura, 1981). 5′-Phosphosulfate nucleosides are quite prone to hydrolysis and decomposition. Therefore, yields for the sulfation step tend to vary (46% to 75%). Cyclic phosphate S.122 can be cleaved by either bovine spleen phosphodiesterase II or ribonuclease-T2 to provide 2′- or 3′-phosphoadenoside-5′-phosphosulfate, respectively (Horwitz et al., 1980; Sekura, 1981). The enzymatic cleavage of cyclic phosphates is tolerant to minor structural modifications of the nucleobase (Horwitz et al., 1981).
Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
CYCLIC NUCLEOSIDE PHOSPHATES AND PHOSPHOROTHIOATES Two important types of nucleoside cyclic phosphates are 2′,3′- and 3′,5′-cyclic phosphates. There are two common approaches to
obtain these cyclic phosphates: cyclization by dehydration or cyclization by nucleophilic attack on electrophilic phosphorus. The latter is often promoted by adding base to the reaction medium. Cyclization by dehydration is commonly affected with DCC, although chloroformate reagents and trifluoroacetic anhydride have also been used. Preparation of cAMP, cCMP, cGMP, and cUMP have been carried out by heating the corresponding NMP-4-morpholine-N,N′-dicyclohexylcarboxamidinium salts in pyridine/DCC (Smith et al., 1961). Unlike tri-n-butylamine or tri-n-octylamine, the base 4-morpholine-N,N′-dicyclohexylcarboxamidine S.125 (Fig. 13.1.29) imparts better solubility upon nucleoside monophosphates at high temperatures. Despite the better solubility of the carboxamidinium salts, guanosine and cytidine need N-benzoyl protecting groups for full dissolution. The yields for this reaction are generally good. DCC-promoted dehydration can also be used for the synthesis of cyclic-2′,3′-phosphates. Cyclization of a commercial mixture of 2′,5′- and 3′,5′-bis(phospho)adenosine with DCC at elevated temperature provides cyclic2′,3′-phosphoadenosine-5′-phosphate (S.120; Fig. 13.1.28; Sekura, 1981). Similarly, reaction of unprotected nucleosides (A, C, G, U) with pyrophosphoryl chloride followed by buffered hydrolysis (pH = 7) results in the formation of cyclic-2′,3′-phosphonucleoside-5′-phosphates (Fig. 13.1.28; Simoncsits and Tomasz, 1975). This reaction is successful for adenosine, cytidine, and guanosine. However, cyclic-2′,3′phosphouridine-5′-phosphate is obtained in low yield via this procedure. These reactions have the inherent drawback that the bis-phosphate is produced instead of pure cyclic-2′,3′phosphate. This limitation has been overcome by the use of the five-coordinate oxyphosphorane S.127 (σ5,λ5) instead of pyrophosphoryl chloride. Treatment of an unprotected nucleoside with oxyphosphorane S.127 in pyridine followed by hydrolysis provides the nucleoside cyclic-2′,3′-phosphate (S.128; Fig. 13.1.30). In this manner, both adenosine and cytidine cyclic-2′,3′-phosphates have been regioselectively synthesized in high yield (Chen et al., 1997). Entrée into 3′,5′-cyclic phosphates is possible through base-promoted cyclization of phosphorodichloridates. Treatment of an unprotected nucleoside with POCl3 produces a phosphorodichloridate that is hydrolyzed with an aqueous solution of KOH in acetonitrile at 0°C
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S P O
Cl
O
B
Cl
O S P O O
KOH
S O B
O P O O
t -BuOK O B
(P(R) major) OH
O2 N
(P(S) major)
OH
129
HO
HO
130 P(R)
131 P(S)
S O P O
B
O
2
OH
OH
132
1) K, CS2 2) NaOH 3) NH3, MeOH
O O P O O
O B
1) Ph3P-CCl4 2) PhNH2 3) chromatography
Ph
Ph
O N P O H O
O B
NH
O P O O
HO
HO
HO
133
134
135
O B
Figure 13.1.31 Synthesis of cyclic phosphorothioates.
to produce the cyclic phosphate in low yield (22% to 49%). This procedure varies slightly from the Yoshikawa procedure (Fig. 13.1.10) in that water is omitted from the reaction milieu during formation of the phosphorodichloridate. Omission of water during the initial phosphorylation step furnishes mixtures of nucleoside2′(3′)-5′-diphosphates (Yoshikawa et al., 1967). Therefore, the reduced yields obtained for this cyclization reaction may be due to lack of regio-control in the initial phosphorylation step. As secondary messengers, nucleoside cyclic-3′,5′-phosphates are important in many biological pathways. Investigation of the pathways could benefit from the ready availability of nucleoside cyclic phosphate analogs that have increased enzymatic and hydrolytic stability. Phosphorothioate derivatives of cyclic3′,5′-phosphates such as cAMPS are, in fact, hydrolyzed more slowly than the parent phosphate (Eckstein, 1979). Additionally, the stereochemical course of enzyme-catalyzed reactions has been determined by using nucleoside cyclic-2′,3′-phosphorothioates (Usher et al., 1970).
The synthesis of cAMPS and other cyclic phosphorothioates is difficult because the phosphorothioate functional group is part of a sixmembered ring. The phosphorus center is therefore stereogenic. Both the P(R) and P(S) stereoisomers are configurationally stable. As shown in Figure 13.1.31, cyclic-3′,5′-phosphorothioates can be accessed by base-promoted cyclization of phosphorothiodichloridates (S.129; Genieser et al., 1988) or nucleoside 5′-bis(ρ-nitrophenyl)phosphorothioates (S.132; Eckstein and Kutzke, 1986). Cyclic phosphorothioates synthesized by these methods are mixtures of phosphorus stereoisomers. Interestingly, ester displacement reactions give predominately the P(S) stereoisomer, whereas chloride displacement results in primarily the P(R) stereoisomer (Fig. 13.1.31). Resolution of the individual stereoisomers from these reactions is possible, but the process is tedious and the yields are generally poor. An alternate method to access the individual stereoisomers is through the stereospecific Horner-Wadsworth-Emmons reaction of nucleoside phosphoranilidates (S.134 or S.135) with potassium reduced carbon disulfide (Fig. 13.1.31; Stec, 1983). This approach has the
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HO
O
B
nucleoside monophosphate kinase
nucleoside kinase NMP
OH
OH
(base-specific)
nucleoside diphosphate kinase NDP
NTP
(base-specific)
Figure 13.1.32 Enzyme-catalyzed phosphorylation of nucleosides proceeds in a stepwise fashion.
advantage that commercially available nucleoside cyclic phosphates can be used. The reaction sequence is outlined in Figure 13.1.31 and has been applied to adenosine, cytidine, guanosine, and uridine. Additionally, the separation of the nucleoside phosphoranilidate stereoisomers is easier than separation of the nucleoside cyclic phosphorothioates. However, the chemical synthesis of the nucleoside phosphoranilidates proceeds in low yield.
ENZYMATIC NUCLEOTIDE SYNTHESIS
Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
In principle, enzymes can be used to synthesize NMPs, NDPs, or NTPs. However, cell-free enzyme-catalyzed reactions are most routinely used for the synthesis of NTPs. Enzymes catalyze the phosphorylation of nucleosides to NTP in a stepwise fashion by the successive action of nucleoside kinase, nucleoside monophosphate kinase, and nucleoside diphosphate kinase (Fig. 13.1.32). The first two enzymes in this pathway are nucleoside specific, whereas, nucleoside diphosphate kinase is not nucleotide specific. Nucleoside kinase, nucleoside monophosphate kinase, and nucleoside diphosphate kinase primarily use ATP as the phosphate donor. Therefore, ATP has to be supplied or regenerated, depending upon scale, during the nucleotide syntheses (Chenault et al., 1988). Due to the use of ATP as phosphate donor, pure nucleotide product can be difficult to obtain. Pure NTP product (where N = C, T, U, G) can be especially difficult to separate from ATP. Purity of product NTPs can be increased by several modifications of the general enzymatic scheme outlined in Figure 13.1.32. Use of commercially available NMPs and regeneration of catalytic amounts of ATP are two approaches to reduce contamination by ATP/ADP. Another way is to use alternate enzymes/phosphate donors to effect the phosphorylation. For example, by choosing conditions that favor the backwards reactions of pyruvate or acetate kinase, phosphoenol pyruvate and acetyl phosphate
can be used as phosphate donors. Phosphoenol pyruvate is more stable in solution than acetyl phosphate and is used extensively for the phosphorylation of NDPs. Hydrolytically stable phosphate donors are especially important when the nucleoside is phosphorylated at slow rates (such as the case with modified nucleosides). Nucleosides containing modified nucleobases cannot always be phosphorylated using synthetic methods due to incompatibility with the reaction conditions. Enzymatic phosphorylation offers an alternative to synthetic methods because the reaction conditions are generally mild and phosphorylations are regio- and chemoselective. However, there has been limited success for enzyme-mediated phosphorylation of modified nucleosides. Unnatural purine- and pyrimidine-like nucleobases are phosphorylated with reduced efficiency (kcat/Km) by D. melanogaster deoxynucleoside kinase (by a factor of 102 to 104; Wu et al., 2002). Phosphorylation of sugar-modified nucleosides can also be problematic. The kinases that phosphorylate AZT to AZT-TP do so with reduced efficiency (Van Rompay et al., 2000). For example, NDP kinase phosphorylates AZT-DP with reduced efficiency (a factor of 103 to 104; Schneider et al., 2001). This is interesting because NDP kinase has little specificity for either the nucleobase or the ribose or deoxyribose of natural NDPs. However, critical H-bonds are provided by the 3′-hydroxyl group to stabilize a conformation of the nucleotide that facilitates phosphorylation. Thus, a minor modification in the architecture of the nucleoside (3′-OH to 3′-N3) results in the destabilization of the active conformation of the nucleotide within the kinase. In contrast to the problems associated with enzymatic phosphorylation of AZT-DP, NDP kinase has been used successfully to phosphorylate nucleoside diphosphates containing nonnatural nucleobases (Wu et al., 2003). Diphosphates of azole carboxamide deoxyribonucleo sides (o r azo le car boxamide
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ribonucleosides) are phosphorylated by NDP kinase using ATP as the phosphate donor (see UNIT 13.2). Good to high yields of the desired azole carboxamide nucleoside triphosphates are obtained. Precursor diphosphates are obtained by way of Poulter’s tosylate displacement methodology. This combination synthetic/enzymatic method is notable for several reasons. First, the method is of interest for its potential as a general means to phosphorylate nucleoside diphosphates containing non-natural nucleobases. Second, the method addresses contamination of the desired nucleoside triphosphate product with ATP/ADP. Use of an ATP regeneration system involving phosphoenol pyruvate and pyruvate kinase minimizes the amount of ATP in the reaction milieu. Nevertheless, contaminating ATP/ADP is conveniently removed by retention on a boronate affinity gel column. The boronate affinity gel forms a complex with the vicinal diols of ribonucleotides; thus, azole carboxamide deoxyribonucleotide products conveniently pass through the gel. There are two issues unique to enzyme-catalyzed phosphorylations that are not encountered during chemical syntheses: enzyme stability and product inhibition. The source of the enzyme can greatly affect the required reaction conditions. For example, acetate kinase from E. coli contains a thiol group that can be oxidized. To prevent inactivation due to oxidation, reactions are often conducted in the presence of a reducing agent (DTT) or under an inert atmosphere. Interestingly, acetate kinase from B. stearothermophilus cannot be autooxidized because it does not contain a thiol group (Kim and Whitesides, 1987). Aside from enzyme inactivation, inhibition of the enzymes can also cause a decrease in the reaction rate. The phosphorylation reactions catalyzed by NMP and NDP kinases are reversible. The consequence of this reversibility is that a large buildup of NDP can inhibit NMP kinase. Thus, the rate at which NMP kinase produces NDP product slows dramatically. Product inhibition is another complicating factor when designing enzyme-catalyzed phosphorylations.
STATE OF THE ART In a 1958 Nobel Prize speech, Sir Alexander Todd made the following observation, “we are thus still seeking an ideal method for unsymmetrical pyrophosphate synthesis” (Todd, 1958). Despite the passage of almost 50 years, the challenge of an “ideal method” for nucleo-
tide synthesis has yet to be satisfactorily met. Nominally, an ideal method permits the regioand chemoselective phosphorylation of all nucleosides and nucleoside analogs (base or sugar modifications alike). Additionally, an ideal method should facilitate the synthesis of any phosphate-chain analog (thio-, borano-, imido-, and others) from common intermediates or an appropriate reagent system. This overview has presented some common methods and reagents used to construct nucleotides. Despite the elegant design of the many methods available, it is clear that no single method is “ideal” under the criteria outlined above. This is due in part to the variation in chemical properties of the nucleobases. Chemoselectivity may be improved by transiently masking the nucleophilic moieties of the nucleobases. Alternately, finding conditions that selectively degrade any phosphorylated (phosphitylated) nucleobase moieties shows potential for a general solution. Progress has been made on chemoselective phosphitylation of N-unprotected nucleosides using phosphoramidite reagents. Both transient protection via protonation of basic nucleobases (Sekine et al., 2003) and selective cleavage of phosphitylated nucleobases (Gryaznov and Letsinger, 1991) have been successfully employed in deoxyoligonucleotide syntheses. Modification of these methods may ultimately be useful for the synthesis of nucleoside mono-, di-, and triphosphates. However, a general solution for the regio- and chemoselective phosphorylation of nucleosides and analogs without invoking the use of protecting groups is a very difficult problem that awaits a general solution.
LITERATURE CITED Albert, A. 1973. Ionization constants of pyrimidines and purines. In Synthetic Procedures in Nucleic Acid Chemistry (W.W. Zorbach and R.S. Tipson, eds.) pp. 1-46. Wiley-Interscience, New York. Arabshahi, A. and Frey, P.A. 1994. A simplified procedure for synthesizing nucleoside 1thiotriphosphate: dATPαS, dGTPαS, UTPαS, and dTTPαS. Biochem. Biophys. Res. Commun. 204:150-155. Batra, J.K., Lin, C.M., and Hamel, E. 1987. Nucleotide interconversions in microtubule protein preparations, a significant complication for accurate measurement of GTP hydrolysis in the presence of adenosine 5′-(β,γ-imidotriphosphate). Biochemistry 26:5925-5931. Burgess, K. and Cook, D. 2000. Syntheses of nucleoside triphosphates. Chem. Rev. 100:20472059.
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Burkart, M.D., Izumi, M., and Wong, C.-H. 1999. Enzymatic regeneration of 3′-phosphoadenosine-5′-phosphosulfate using aryl sulfotransferase for the preparative enzymatic synthesis of sulfated carbohydrates. Angew. Chem. Int. Ed. Engl. 38:2747-2750. Chen, J.-T. and Benkovic, S.J. 1983. Synthesis and separation of diastereomers of deoxynucleoside 5′-O-(1-thio)triphosphates. Nucl. Acids Res. 11:3737-3751. Chen, X., Zhang, N.-J., Li, Y.-M., Jiang, Y., Zhang, X., and Zhao, Y.-F. 1997. Direct phosphorylation of nucleosides by oxyphosphorane. Tetrahedron Lett. 38:1615-1618. Chenault, H.K., Simon, E.S., and Whitesides, G.M. 1988. Cofactor regeneration for enzyme-catalysed synthesis. In Biotechnology and Genetic Engineering Reviews (G.E. Russell, ed.) pp. 221-270. Intercept, Wimborne, Dorset, England. Chladek, S. and Nagyvary, J. 1972. Nucleophilic reactions of some nucleoside phosphorothioates. J. Am. Chem. Soc. 94:2079-2085. Corbridge, D.E.C. 1995. Phosphorous. An Outline of its Chemistry, Biochemistry and Uses. Elsevier, Amsterdam. Cosstick, R. and Vyle, J.S. 1990. Synthesis and properties of dithymidine phosphate analogues containing 3′-thiothymidine. Nucl. Acids Res. 18:829-835. Davisson, V.J., Davis, D.R, Dixit, V.M., and Poulter, C.D. 1987. Synthesis of nucleotide 5′-diphosphates from 5′-O-tosyl nucleosides. J. Org. Chem. 52:1794-1801. Dixit, V.M. and Poulter, C.D. 1984. Convenient syntheses of adenosine 5′-diphosphate, adenosine 5′-methylenediphosphonate, and adenosine 5′-triphosphate. Tetrahedron Lett. 25:40554058. Eckstein, F. 1979. Phosphorothioate analogues of nucleotides. Acc. Chem. Res. 12:204-210. Eckstein, F. and Kutzke, U. 1986. Synthesis of nucleoside 3′,5′-cyclic phosphorothioates. Tetrahedron Lett. 27:1657-1660. Engel, R. 1977. Phosphonates as analogues of natural phosphates. Chem. Rev. 77:349-367. Eymery, F., Iorga, B., and Savignac, P. 1999. Synthesis of phosphonates by nucleophilic substitution at phosphorus: The SN P(V) reaction. Tetrahedron 55:13109-13150. Furuichi, Y. and Shatkin, A.J. 1977. A simple method for the preparation of [β-32P]purine nucleoside triphosphates. Nucl. Acids Res. 4:3341-3355. Genieser, H.-G., Dostmann, W., Bottin, U., Butt, E., and Jastorff, B. 1988. Synthesis of nucleoside3′,5′-cyclic phosphorothioates by cyclothiophosphorylation of unprotected nucleosides. Tetrahedron Lett. 29:2803-2804. Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
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Gryaznov, S.M. and Letsinger, R.L. 1991. Synthesis of oligonucleotides via monomers with unprotected bases. J. Am. Chem. Soc. 113:5876-5877. Hoard, D.E. and Ott, D.G. 1965. Conversion of mono- and oligodeoxyribonucleotides to 5′triphosphates. J. Am. Chem. Soc. 87:1785-1788. Horwitz, J.P., Neenan, J.P., Misra, R.S., Rozhin, J., Huo, A., and Philips, K.D. 1977. Studies on bovine adrenal estrogen sulfotransferase III. Facile synthesis of 3′-phospho- and 2′-phosphoadenosine 5′-phosphosulfate. Biochim. Biophys. Acta 480:376-381. Horwitz, J.P., Misra, R.S., Rozhin, J., Helmer, S., Bhuta, A., and Brooks, S.C. 1980. Studies on bovine adrenal estrogen sulfotransferase. V. Synthesis and assay of analogs of 3′-phosphoadenosine 5′-phosphosulfate as cosubstrates for estrogen sulfurylation. Biochim. Biophys. Acta 613:85-94. Horwitz, J.P., Neenan, J.P., Misra, R.S., Rozhin, J., Huo, A., and Philips, K.D. May 1981.U.S. patent 4,266,048. Synthesis of Analogs of 3′-Phosp ho aden osine 5′-Phosphosulfate (PAPS). USPTO Patent Full-Text and Image Database. USA, The United States of America as represented by the Department of Health. Imai, J. and Torrence, P.F. 1981. Bis(2,2,2-trichloroethyl) phosphorochloridite as a reagent for the phosphorylation of oligonucleotides: Preparation of 5′-phosphorylated 2′,5′-oligoadenylates. J. Org. Chem. 46:4015-4021. Iyer, R., Phillips, L.R., Egan, W., Regan, J.B., and Beaucage, S.L. 1990. The automated synthesis of sulfur-containing oligodeoxyribonucleotides using 3H-1,2-benzodithiol-3-one 1,1-dioxide as a sulfur-transfer reagent. J. Org. Chem. 55:46934699. Janecka, A., Panusz, H., Pankowski, J., and Koziolkiewicz, W. 1980. Chemical synthesis of nucleoside-γ-[32P]triphosphates of high specific activity. Prep. Biochem. 10:27-35. Jankowska, J., Cieslak, J., and Kraszewski, A. 1997. 9-Fluorenemethyl H-phosphonothioate, a versatile reagent for the preparation of H-phosphonothioate, phosphorothioate, and phosphorodithioate monoesters. Tetrahedron Lett. 38:2007-2010. Jankowska, J., Sobkowska, A., Cieslak, J., Sobkowski, M., Krazewski, A., Stawinski, J., and Shugar, D. 1998. Nucleoside H-phosphonates. 18. Synthesis of unprotected nucleoside 5′-Hphosphonates and nucleoside 5′-H-phosphonothioates and their conversion into the 5′phosphorothioate and 5′-phosphorodithioate monoesters. J. Org. Chem. 63:8150-8156. Jurczyk, S.C., Kodra, J.T., Park, J.-H., Benner, S.A., and Battersby, T.R. 1999. Synthesis of 2′-deoxyisoguanosine 5′-triphosphate and 2′-deoxy-5methylisocytidine 5′-triphosphate. Helv. Chim. Acta 82:1005-1015.
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Kadokura, M., Wada, T., Urashima, C., and Sekine, M. 1997. Efficient synthesis of γ-methyl-capped guanosine 5′-triphosphate as a 5′-terminal unique structure of U6 RNA via a new triphosphate bond formation involving activation of methyl phosphorimidazolidate using ZnCl2 as a catalyst in DMF under anhydrous conditions. Tetrahedron Lett. 38:8359-8362. Kaufmann, G., Choder, M., and Groner, Y. 1980. Synthesis of carrier-free β-32P-nucleosidetriphosphate in almost quantitative yields. Anal. Biochem. 109:198-202. Kim, M.-J. and Whitesides, G.M. 1987. Enzymecatalyzed synthesis of nucleoside triphosphates from nucleoside monophosphates. Appl. Biochem. Biotechnol. 16:95-108. Kimura, J., Fujisawa, Y., Yoshizawa, T., Fukuda, K., and Mitsunobu, O. 1979. Studies on nucleosides and nucleotides. VII. Preparation of pyrimidine nucleoside 5′-phosphates and N3,5′-purine cyclonucleosides by selective activation of the 5′hydroxyl group. Bull. Chem. Soc. Jpn. 52:11911196. Klaassen, C.D. and Boles, J.W. 1997. The importance of 3′-phosphoadenosine 5′-phosphosulfate (PAPS) in the regulation of sulfation. FASEB J. 11:404-418. Klein, E., Mons, S., Valleix, A., Mioskowski, C., and Lebeau, L. 2002. Synthesis of enzymatically and chemically non-hydrolyzable analogues of dinucleoside triphosphate Ap3A and Gp3G. J. Org. Chem. 67:146-153. Kovacs, T. and Otvos, L. 1988. Simple synthesis of 5-vinyl- and 5-ethynyl-2′-deoxyuridine-5′triphosphates. Tetrahedron Lett. 29:4525-4528. Larsen, M., Willett, R., and Yount, R.G. 1969. Imidodiphosphate and pyrophosphate: Possible biological significance of similar structures. Science 166:1510-1511. Lebedev, A.V., Koukhareva, I.I., Beck, T., and Vaghefi, M.M. 2001. Preparation of oligodeoxynucleotide 5′-triphosphates using solid support approach. Nucleosides Nucleotides Nucleic Acids 20:1403-1409. Li, R., Muscate, A., and Kenyon, G.L. 1996. Synthesis, characterization, and inhibitory activities of nucleoside α,β-imido triphosphate analogues on human immunodeficiency virus-1 reverse transcriptase. Bioorg. Chem. 24:251-261. Lin, C.-H., Shen, G.-J., Garcia-Junceda, E., and Wong, C.-H. 1995. Enzymatic synthesis and regeneration of 3′-phosphoadenosine 5′-phosphosulfate (PAPS) for regioselective sulfation of oligosaccharides. J. Am. Chem. Soc. 117:80318032. Ludwig, J. 1987. A simple one flask synthesis of nucleoside 5′-triphosphates from unprotected nucleosides via nucleoside 5′-cyclotriphosphates. In Biophosphates and Their Analogues—Synthesis, Structure, Metabolism and Activity (K.S. Bruzik and W.J. Stec, eds.) pp. 201-204. Elsevier Science Publishers B V, Amsterdam.
Ludwig, J. and Eckstein, F. 1989. Rapid and efficient synthesis of nucleoside 5′-O-(1-thiotriphosphates), 5′-triphosphates and 2′,3′-cyclophosphorothioates using 2-chloro-4H-1,3,2-benzodioxaphosphorin-4-one. J. Org. Chem. 54:631635. Ludwig, J. and Eckstein, F. 1991. Synthesis of nucleoside 5′-O-(1,3-dithiotriphosphates) and 5′O-(1,1-dithiotriphosphates). J. Org. Chem. 56:1777-1783. Ma, Q.-F., Babbitt, P.C., and Kenyon, G.L. 1988. Adenosine 5′-[α,β-imido]triphosphate, a substrate for T7 RNA polymerase and rabbit muscle creatine kinase. J. Am. Chem. Soc. 110:40604061. Ma, Q.-F., Kenyon, G.L., and Markham, G.D. 1990. Specificity of S-adenosylmethionine synthetase for ATP analogues mono- and disubstituted in bridging positions of the polyphosphate chain. Biochemistry 29:1412-1416. McBride, L.J. and Caruthers, M.H. 1983. An investigation of several deoxynucleoside phosphoramidites. Tetrahedron Lett. 24:245-248. Michelson, A.M. 1964. Synthesis of nucleotide anhydrides by anion exchange. Biochim. Biophys. Acta 91:1-13. Mishra, N.C. and Broom, A.D. 1991. A novel synthesis of nucleoside 5′-triphosphates. J. Chem. Soc., Chem. Comm. 18:1276-1277. Nairne, R.J.D., Pickering, L., and Smith, C.L. 2002. Synthesis of pyrrole carboxamide nucleotide triphosphates—putative labeled nucleotide analogues. Tetrahedron Lett. 43:2289-2291. Nurminen, E.J., Mattinen, J.K., and Lonnberg, H. 1998. Kinetics and mechanism of tetrazole-catalyzed phosphoramidite alcoholysis. J. Chem. Soc., Perkin Trans. 2:1621-1628. Patel, B.K. and Eckstein, F. 1997. 5′-Deoxy-5′thioribonucleoside-5′-triphosphates. Tetrahedron Lett. 38:1021-1024. Penningroth, S.M., Olehnik, K., and Cheung, A. 1980. ATP formation from adenyl-5′-yl imidodiphosphate, a nonhydrolyzable ATP analog. J. Biol. Chem. 255:9545-9548. Quin, L.D. 2000. Organophosphorus Chemistry. Wiley-Interscience, New York. Regan, J.B., Phillips, L.R., and Beaucage, S.L. 1992. Large-scale preparation of the sulfurtransfer reagent 3H-1,2-benzodithiol-3-one 1,1dioxide. Org. Prep. Proc. Int. 24:488-492. Reynolds, M.A., Gerlt, J.A., Demou, P.C., Oppenheimer, N.J., and Kenyon, G.L. 1983. 15N and 17 O NMR studies of the proton binding sites in imidodiphosphate, tetraethyl imidodiphosphate, and adenylyl imidodiphosphate. J. Am. Chem. Soc. 105:6475-6481. Richard, J.P., Ho, H.-T., and Frey, P.A. 1978. Synthe sis of nuc leos ide [18O]pyrophosphorothioates with chiral [18O]phosphorothioate groups of known configuration. Stereochemical orientations of enzymatic phosphorylations of chiral [18O]phosphorothioates. J. Am. Chem. Soc. 100:7756-7757.
Nucleoside Phosphorylation and Related Modifications
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Saady, M., Lebeau, L., and Mioskowski, C. 1995a. Synthesis of adenosine-5′-phosphates and 5′-alkylphosphonates via the Mitsunobu reaction. Tetrahedron Lett. 36:2239-2242. Saady, M., Lebeau, L., and Mioskowski, C. 1995b. Synthesis of di- and triphosphate ester analogs via a modified Michaelis-Arbuzov reaction. Tetrahedron Lett. 36:5183-5186. Saffhill, R. 1970. Selective phosphorylation of the cis-2′,3′-diol of unprotected ribonucleosides with trimetaphosphate in aqueous solution. J. Org. Chem. 35:2881-2883. Schneider, B., Babolat, M., Xu, Y.W., Janin, J., Vernon, M., and Deville-Bonne, D. 2001. Mechanism of phosphoryl transfer by nucleoside diphosphate kinase. Eur. J. Biochem. 268:19641971. Sekine, M., Ohkubo, A., and Seio, K. 2003. Protonblock strategy for the synthesis of oligodeoxynucleotides without base protection, capping reaction and P-N bond cleavage reaction. J. Org. Chem. 68:5478-5492.
Tomasz, J., Vaghefi, M.M., Ratsep, P.C., Willis, R.C., and Robins, R.K. 1988. Nucleoside imidodiphosphate synthesis and biological activities. Nucl. Acids Res. 16:8645-8664. Trevisiol, E., Defrancq, E., Lhomme, J., Laayoun, A., and Cros, P. 2000. Synthesis of nucleoside triphosphates that contain an aminooxy function for “Post-Amplification Labeling”. Eur. J. Org. Chem. 1:211-217. Tsuhako, M., Fujimoto, M., Ohashi, S., Nariai, H., and Motooka, I. 1984. Phosphorylation of nucleosides with sodium cyclo-triphosphate. Bull. Chem. Soc. Jpn. 57:3274-3280. Uchiyama, M., Aso, Y., Noyori, R., and Hayakawa, Y. 1993. O-selective phosphorylation of nucleosides without N-protection. J. Org. Chem. 58:373-379. Usher, D.A., Richardson, D.I. Jr., and Eckstein, F. 1970. Absolute stereochemistry of the second step of ribonuclease action. Nature 228:663-665.
Sekura, R.D. 1981. [53] Adenosine 3′-phosphate 5′-phosphosulfate. In Detoxification and Drug Metabolism: Conjugation and Related Systems (W.B. Jakoby, ed.) pp. 413-415. Academic Press, New York.
Van Rompay, A.R., Johansson, M., and Karlsson, A. 2000. Phosphorylation of nucleosides and nucleoside analogs by mammalian nucleoside monophosphate kinases. Pharmacol. Ther. 87:189198.
Simoncsits, A. and Tomasz, J. 1975. Simple onestep synthesis of ribonucleoside 2′,3′-cyclic phosphate 5′-phosphates. Biochim. Biophys. Acta 395:74-79.
Verheyden, J.P.H. and Moffatt, J.G. 1970. Halo sugar nucleosides. I. Iodination of the primary hydroxyl groups of nucleosides with methyltriphenoxyphosphonium iodide. J. Org. Chem. 35:2319-2326.
Smith, M., Drummond, G.I., and Khorana, H.G. 1961. Cyclic phosphates. IV. Ribonucleoside3′,5′ cyclic phosphates. A general method of synthesis and some properties. J. Am. Chem. Soc. 83:698-706. Stec, W.J. 1983. Wadsworth-Emmons reaction revisited. Acc. Chem. Res. 16:411-417. Stock, J.A. 1979. Synthesis of phosphonate analogues of thymidine di- and triphosphate from 5′-O-toluenesulfonylthymidine. J. Org. Chem. 44:3997-4000. Stowell, J.K. and Widlanski, T.S. 1995. Mechanismbased inactiviation of ribonuclease A. J. Org. Chem. 60:6930-6936. Stromberg, R. and Stawinski, J. 1987. Iodide and iodine catalysed phosphorylation of nucleosides by phosphorodiester derivatives. Nucleosides & Nucleotides 6:815-820. Symons, R.H. 1970. Practical methods for the routine chemical synthesis of 32P-labelled nucleoside di- and triphosphates. Biochim. Biophys. Acta 209:296-305. 32
Symons, R.H. 1974. Synthesis of [α- P]ribo- and deoxyribonucleoside 5′-triphosphates. Methods Enzymol. 29:102-115.
Overview of the Synthesis of Nucleoside Phosphates and Polyphosphates
Todd, A. 1958. Synthesis in the study of nucleotides. Science 127:787-792.
Thomson, J.B., Patel, B.K., Jimenez, V., Eckart, K., and Eckstein, F. 1996. Synthesis and properties of diuridine phosphate analogues containing thio and amino modifications. J. Org. Chem. 61:6273-6281.
Walseth, T.F. and Johnson, R.A. 1979. The enzymatic preparation of [α-32P]nucleoside triphosphates, cyclic [32P]AMP, and cyclic [32P]GMP. Biochim. Biophys. Acta 526:11-31. Walseth, T.F., Yuen, P.S.T., and Moos, M.C. Jr. 1991. Preparation of α-32P-labeled nucleoside triphosphates, nicotinamide adenine dinucleotide, and cyclic nucleotides for use in determining adenylyl and guanylyl cyclases and cyclic nucleotide phosphodiesterase. Methods Enzymol. 195:29-44. Werhli, W.E., Verheydden, D.L.M., and Moffatt, J.G. 1965. Dismutation reactions of nucleoside polyphosphates. II. Specific chemical syntheses of α-, β-, and γ-32P-nucleoside 5′-triphosphates. J. Am. Chem. Soc. 87:2265-2277. Wu, Y., Fa, M., Tae, E.L., Schultz, P.G., and Romesberg, F.E. 2002. Enzymatic phosphorylation of unnatural nucleosides. J. Am. Chem. Soc. 124:14626-14630. Wu, W., Bergstrom, D.E., and Davisson, V.J. 2003. A combination chemical and enzymatic approach for the preparation of azole carboxamide nucleoside triphosphate. J. Org. Chem. 68:38603865. Yoshikawa, M., Kato, T., and Takenishi, T. 1967. A novel method for phosphorylation of nucleos id es to 5′-nucleotides. Tetrahedron Lett. 50:5065-5068.
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Yount, R.G., Babcock, D., Ballantyne, W., and Ojala, D. 1971. Adenylyl imidodiphosphate, an adenosine triphosphate analog containing a P-NP linkage. Biochemistry 10:2484-2489. Zemlicka, J. 2002. Lipophilic phosphoramidates as antiviral pronucleotides. Biochim. Biophys. Acta 1587:276-286.
Contributed by David C. Johnson II and Theodore S. Widlanski Indiana University Bloomington, Indiana
Nucleoside Phosphorylation and Related Modifications
13.1.31 Current Protocols in Nucleic Acid Chemistry
Supplement 15
Chemoenzymatic Preparation of Nucleoside Triphosphates
UNIT 13.2
This unit presents protocols for the preparation of azole carboxamide deoxyribo- and ribonucleoside triphosphates by enzyme-catalyzed phosphorylation of the cognate diphosphates. The latter are synthesized from nucleoside 5′-O-tosylates by displacement with tris(tetra-n-butylammonium) pyrophosphate. Purification procedures using boronate affinity gel to yield highly purified nucleoside triphosphates are also presented. A general procedure for the preparation of the azole carboxamide deoxyribonucleoside triphosphates is described first (see Basic Protocol 1). This method involves selective 5′-tosylation of the azole carboxamide deoxyribonucleoside, followed by displacement of the 5′-tosylate with tris(tetra-n-butylammonium) pyrophosphate to yield the corresponding nucleoside diphosphate. Enzymatic phosphorylation utilizes ATP as the phosphate donor and nucleoside diphosphate kinase as the catalyst, coupled with phosphoenolpyruvate (PEP) and pyruvate kinase as an ATP regeneration system. Next, a general procedure is presented for the synthesis of azole carboxamide ribonucleoside triphosphates (see Basic Protocol 2). This method includes protection of 2′,3′hydroxyl groups of the ribonucleoside, 5′-tosylation, displacement of the 5′-tosylate with tris(tetra-n-butylammonium) pyrophosphate, and deprotection of the 2′,3′-hydroxyls to yield ribonucleoside diphosphate. Enzyme-catalyzed phosphorylation uses PEP as the phosphate donor and pyruvate kinase as the catalyst. SYNTHESIS OF AZOLE CARBOXAMIDE DEOXYRIBONUCLEOSIDE TRIPHOSPHATES
BASIC PROTOCOL 1
This protocol outlines a general procedure for the synthesis and purification of azole carboxamide deoxyribonucleoside triphosphates (Fig. 13.2.1; Wu et al., 2003). Specific instructions are given for the synthesis of TzA4 triphosphate (S.10); however, synthesis of the other azole carboxamide deoxyribonucleoside triphosphates in Figure 13.2.1 (S.11 and S.12) can be accomplished using the same procedure (see Critical Parameters and Troubleshooting). Materials Azole carboxamide deoxyribonucleoside (Fig. 13.2.1): S.1 (N = TzA4; Makabe et al., 1977), S.2 (N = TzA3; Witkowski et al., 1975), or S.3 (N = Tz2A4; Makabe et al., 1977) Pyridine, anhydrous Argon p-Toluenesulfonyl chloride (TsCl) Ethyl acetate, ACS reagent grade 5% (w/v) phosphomolybdic acid solution (see recipe) Methanol, ACS reagent grade 200- to 400-mesh silica gel 60 Hexanes, ACS reagent grade Phosphorus pentoxide (P2O5) Dowex AG 50W-X8 cation-exchange resin (100 to 200 mesh, Bio-Rad) 1 M HCl Disodium dihydrogen pyrophosphate 1 M and concentrated (28%) ammonium hydroxide (NH4OH) 40% (w/v) tetra-n-butylammonium hydroxide solution (Aldrich) Contributed by Weidong Wu, Donald E. Bergstrom, and V. Jo Davisson Current Protocols in Nucleic Acid Chemistry (2004) 13.2.1-13.2.19 Copyright © 2004 by John Wiley & Sons, Inc.
Nucleoside Phosphorylation and Related Modifications
13.2.1 Supplement 16
O
O N
HO
TsO
a
O
N O
O P
b
O
O OH 1 N = TzA4 2 N = TzA3 3 N = Tz2A4 CONH2
N TzA 4 N =
N
CONH2
N TzA 3 N =
N N
O
OH
OH
4 N = TzA4 5 N = TzA3 6 N = Tz2A4
7 N = TzA4 8 N = TzA3 9 N = Tz2A 4 ATP
pyruvate
nucleoside diphosphate kinase
PEP O
ADP
O−
O
O
HO P O P CONH2
N
O
O
pyruvate kinase
N
P
O−
O
P
N
O
O
O− OH
Tz2A 4 N =
N
N N
10 N = TzA4 11 N = TzA3 12 N = Tz2A 4
Figure 13.2.1 General procedure for the synthesis of azole carboxamide deoxyribonucleoside triphosphates (see Basic Protocol 1). Reagents: (a) TsCl, pyridine, room temperature (steps 5 and 6); (b) (NBu4)3HP2O7, CH3CN, room temperature (steps 27 and 28). Abbreviations: TzA3, (1H)1,2,4-triazole-3-carboxamide; TzA4, (1H)-1,2,3-triazole-4-carboxamide; Tz2A4, (2H)-1,2,3-triazole4-carboxamide.
Chemoenzymatic Preparation of Nucleoside Triphosphates
Acetonitrile, anhydrous Deuterated acetonitrile (Aldrich) CF11 fibrous cellulose powder (Whatman) 50 mM, 100 mM, and 500 mM ammonium bicarbonate (NH4HCO3) solutions (no pH adjustment) 1% (w/v) sulfosalicylic solution (see recipe) 0.2% (w/v) ferric chloride solution (see recipe) Triethanolamine hydrochloride (Sigma) Magnesium chloride hexahydrate (MgCl⋅6H2O) Potassium chloride (KCl) Adenosine triphosphate (ATP, disodium salt, Sigma) Sodium phosphoenolpyruvate monohydrate (PEP; Sigma) 1 M sodium hydroxide (NaOH) Nucleoside diphosphate kinase (see recipe) Pyruvate kinase (see recipe) Mobile phase A: 0.1 M potassium phosphate buffer (pH 6.0; APPENDIX 2A) containing 4 mM tetrabutylammonium dihydrogenphosphate (TBAP, Aldrich; added from a 1.0 M stock) Mobile phase B: 70:30 (v/v) mobile phase A/methanol, pH 7.2 Adenosine diphosphate (ADP, sodium salt, Sigma)
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Affi-Gel 601 boronate affinity gel (Bio-Rad) 1 M ammonium bicarbonate (NH4HCO3), pH 8.5 (see recipe) Carbon dioxide source (e.g., dry ice) Q Sepharose FF anion-exchange resin (Bio-Rad) 1 M potassium chloride (KCl) 10- and 20-mL oven-dried one-neck round-bottom flasks with rubber septa Rotary evaporator equipped with a water aspirator and a temperature-controlled water bath (45°C) Capillary tubes TLC plates: Silica gel 60 F254 polyester-backed TLC plates (Aldrich) Cellulose TLC plates (EM Science) Heat gun 254-nm UV lamp 2.0 × 25–cm, 2.5 × 5–cm, 2.5 × 10–cm, 2.5 × 20–cm, and 2.5 × 25–cm chromatography columns Vacuum desiccator 250-mL and 1-L beakers 250-mL round-bottom flasks Lyophilizer TLC sprayer (Analtech) 15-mL polystyrene round-bottom tube HPLC system (e.g., Beckman System Gold) including: 128 solvent module 166 detector set at 230 nm 25-cm × 4.6-mm × 5-µm-i.d. Supelcosil LC-18-T column 25-mm-diameter, 0.2-µm nylon syringe filter (Fisher) Peristaltic pump Medium-pressure liquid chromatography (MPLC) system (e.g., ISCO LC system) with following equipment: Model 2360 gradient programmer Model 2350 HPLC pump V4 variable wavelength absorbance detector Additional reagents and equipment for thin-layer chromatography (TLC, APPENDIX 1 13 31 3D), column chromatography (APPENDIX 3E), and H NMR, C NMR, P NMR, and ESI-MS NOTE: All the nucleosides used in the reaction are dried under reduced pressure (0.1 Torr) in the presence of phosphorus pentoxide at 50°C in an Abderhalden drying apparatus (Ace Glass) overnight prior to reaction. Prepare 5′-tosylate of TzA4 deoxyribonucleoside (S.4) 1. Add 100 mg (0.44 mmol) TzA4 azole carboxamide deoxyribonucleoside (S.1) to an oven-dried 20-mL one-neck round-bottom flask. 2. Add 5 mL anhydrous pyridine to the flask and evaporate to dryness with a 45°C rotary evaporator equipped with a water aspirator. Repeat this procedure three times. 3. Put a magnetic stir bar into the reaction flask, insert a rubber septum, and place the flask on top of a magnetic stir plate. 4. Evacuate the reaction flask using a vacuum line, then flush with argon. Repeat this procedure three times, then attach the flask to an argon line.
Nucleoside Phosphorylation and Related Modifications
13.2.3 Current Protocols in Nucleic Acid Chemistry
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5. Quickly remove the septum, add 100 mg (0.53 mmol) TsCl into the reaction flask, and immediately reinsert the septum. 6. Transfer 4 mL anhydrous pyridine to the flask under argon and stir the reaction at room temperature. 7. Monitor the progress of the reaction by analytical TLC (APPENDIX 3D) as follows: a. Withdraw a small sample using a capillary tube and spot on a silica gel 60 F254 polyester-backed TLC plate. b. To remove pyridine in the sample, heat the plate for several minutes using a heat gun until the diffuse dark spot visualized under a 254-nm UV lamp disappears. Be careful not to burn the TLC plate (the polymer will begin to shrink). It is very important to remove pyridine as completely as possible to allow visualization of the product.
c. Develop the TLC plate in ethyl acetate. d. Visualize by dipping the plate into 5% phosphomolybdic acid solution and heating with a heat gun. The starting material and product appear as blue or dark blue spots on a green background (Rf = 0.30 for S.1; Rf = 0.50 for S.4).
Workup and purify S.4 8. When TLC analysis indicates the reaction is complete (∼4 days), remove the magnetic stir bar from the flask and remove the solvent under reduced pressure with a rotary evaporator. 9. Add 10 mL methanol to the flask and evaporate to dryness. Repeat this procedure three times to completely remove pyridine. 10. Dissolve the obtained residue in 10 mL methanol and add 1.5 g of 200- to 400-mesh silica gel 60 to the solution. Evaporate the mixture to dryness. The crude product is absorbed on silica gel.
11. Pack a 2.0 × 25–cm chromatography column with 25 g silica gel in 1:1 (v/v) hexanes/ethyl acetate for flash column chromatography (APPENDIX 3E). 12. Add the silica gel containing the crude product to the top of the packed column. 13. Elute with a stepwise gradient of 1:1 hexanes/ethyl acetate to 100% ethyl acetate. Collect 10-mL fractions and analyze by TLC as described in step 7. 14. Combine the product-containing fractions and evaporate to dryness with a rotary evaporator. 15. Dry the product overnight over phosphorus pentoxide in a vacuum desiccator at 0.1 Torr. 16. Confirm the desired product by 1H NMR, 13C NMR, and ESI-MS.
Chemoenzymatic Preparation of Nucleoside Triphosphates
Flash chromatography gives 105 mg (62%) S.4 as a white solid. 1H NMR (DMSO-d6, 300 MHz): 8.59 (1H, s), 7.88 (1H, s), 7.68 (2H, d), 7.52 (1H, s), 7.39 (2H, d), 6.37 (1H, t, J = 6.0 Hz), 5.58 (1H, d, D2O exchangeable), 4.38 (1H, m), 4.20 (1H, m), 4.03 (2H, m), 2.64 (1H, m), 2.40 (4H, m). 13C NMR (DMSO-d6, 75 MHz): 161.3, 145.0, 143.1, 130.1, 127.5, 125.5, 88.0, 84.3, 70.1, 69.9, 21.1. MS (ESI): 383 [M+H]+. Anal. calcd. for C15H18O6N4S: C 47.11, H 4.74, N 14.65; found: C 47.00, H 4.64, N 14.52. Characterization data for S.5 and S.6 can be found in Wu et al. (2003).
13.2.4 Supplement 16
Current Protocols in Nucleic Acid Chemistry
Prepare tris(tetra-n-butylammonium) hydrogen pyrophosphate 17. Pack Dowex AG 50W-X8 cation-exchange resin in water into a 2.5 × 10–cm column and elute the column in sequence with 150 mL of 1 M HCl and 150 mL water. The cation-exchange resin is transformed to the hydrogen form.
18. Dissolve 3.33 g (15.0 mmol) disodium dihydrogen pyrophosphate in 13.5 mL water and 1.5 mL concentrated (28%) aqueous ammonium hydroxide. 19. Pass the solution through the cation-exchange column. 20. Elute the column with 100 mL deionized water and collect the eluent in a 250-mL beaker. 21. Titrate the eluent to pH 7.3 with 40% (w/w) aqueous tetra-n-butylammonium hydroxide (∼50 mL). The resulting solution is ∼150 mL in total volume.
22. Transfer the solution to a 250-mL round-bottom flask and lyophilize for 24 hr to dry. After lyophilization, 13.1 g (97%) of a hygroscopic white solid is obtained. The product can be stored up to one year in a desiccator at −20°C.
Prepare nucleoside diphosphate S.7 23. Add 214 mg (0.56 mmol) nucleoside 5′-tosylate (S.4) to an oven-dried 10-mL one-neck round-bottom flask. 24. Add 5 mL anhydrous acetonitrile to the flask and evaporate to dryness with a rotary evaporator. Repeat this procedure three times. 25. Put a magnetic stir bar into the reaction flask, insert a rubber septum, and place the flask on top of a magnetic stir plate. 26. Evacuate the reaction flask via a vacuum line, then flush with argon. Repeat this procedure three times and attach the flask to an argon line. 27. Quickly remove the septum, add 1.0 g (1.12 mmol) tris(tetra-n-butylammonium) hydrogen pyrophosphate (step 22) into the reaction flask, and immediately reinsert the septum. 28. Transfer 1.5 mL anhydrous acetonitrile to the flask under argon and stir the reaction at room temperature. 29. Monitor the progress of the reaction by 1H NMR as follows: a. Withdraw 50 µL reaction mixture and dilute with 0.5 mL deuterated acetonitrile. b. Transfer the sample to a dried NMR tube and acquire a 1H NMR spectrum. c. Monitor the progress of the reaction by the change in the 1H NMR spectrum between 7 and 9 ppm, where the tosylate moiety appears as an AA′XX′ spin system. The four-line pattern for the tosylate moves upfield by ∼0.1 to 0.2 ppm upon conversion to the tosylate anion. TLC is not suitable for monitoring the progress of the reaction because the tetra-n-butylammonium salt does not give a well-defined spot. Nucleoside Phosphorylation and Related Modifications
13.2.5 Current Protocols in Nucleic Acid Chemistry
Supplement 16
Workup and purify S.7 30. When the 1H NMR indicates that the reaction is complete (∼24 hr), add 5 mL deionized water to the reaction mixture. 31. Pack Dowex AG 50W-X8 cation-exchange resin in water into a 2.5 × 10–cm column. Transform the cation-exchange resin to the ammonium form by washing the column with 150 mL (3 column volumes) of the following solutions in sequence: 1 M HCl H2O 1 M NH4OH H2O. 32. Load the sample solution from step 30 onto the column. 33. Elute with 100 mL (2 column volumes) water and collect in a 250-mL round-bottom flask. Lyophilize to dryness. The 1H NMR spectrum of the crude product shows only a trace of the signals from the tetra-n-butylammonium cation.
34. Pack a Whatman CF11 fibrous cellulose column as follows: a. Mix 500 mL (dry volume) CF11 fibrous cellulose power with 350 mL water in a 1-L beaker by vigorous stirring with a glass rod. b. Slurry-pack the cellulose into a 2.5 × 25–cm chromatography column. c. Wash the column with 300 mL water, 300 mL acetonitrile, and 300 mL of 1:1 (v/v) acetonitrile/water in sequence. d. Equilibrate the column with 300 mL of 7:3:2 (v/v/v) acetonitrile/100 mM ammonium bicarbonate/concentrated (28%) ammonium hydroxide. 35. Extract the crude product (from step 33) with 5 mL of the same acetonitrile/ammonium bicarbonate/ammonium hydroxide buffer. Pellet the precipitate. The product is soluble in the buffer and the white precipitate formed is excess inorganic pyrophosphate.
36. Load the soluble material onto the cellulose column and elute with the same buffer by flash chromatography, collecting 10-mL fractions. 37. Analyze every second fraction on a cellulose TLC plate as follows: a. Spot the sample on the cellulose TLC plate and develop in 7:3:2 acetonitrile/100 mM ammonium bicarbonate/concentrated ammonium hydroxide. b. Spray the plate with 1% sulfosalicylic acid solution until thoroughly wetted but not dripping, and allow to air dry 5 min. c. Lightly spray 0.2% ferric chloride solution onto the plate and visualize the spots. When visualized by sulfosalicylic acid/ferric chloride spray, phosphate-containing compounds appear as white spots on a pink background. A second light spray with ferric chloride may be necessary to make the spots pronounced. For S.7, the Rf value is 0.2 and is similar to the other nucleoside diphosphates.
38. Combine the product-containing fractions and remove acetonitrile by rotary evaporation with the bath temperature below 30°C. 39. Lyophilize the resulting aqueous solution to dryness. Chemoenzymatic Preparation of Nucleoside Triphosphates
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40. Confirm the product by 1H NMR, 13C NMR, 31P NMR, and ESI-MS. Flash cellulose chromatography gives 200 mg (81%) S.7 as a white fluffy powder. 1H NMR (D2O, 300 MHz): 8.68 (1H, s), 6.43 (1H, t, J = 6.0 Hz), 4.14 (1H, m), 3.98 (2H, m), 2.68 (1H, m), 2.52 (1H, m). 13C NMR (D2O, 75 MHz): 164.5, 141.9, 125.9, 89.5, 86.7, 70.6, 64.9, 39.9. 31P NMR (D2O, 121 MHz): –9.85 (d, Jp,p = 21.8 Hz), –13.38 (d, Jp,p = 21.8 Hz). HRMS (ESI): calcd. for C8H15N4O10P2 389.0263 [M+H]+; found 389.0247. Characterization data for S.8 and S.9 can be found in Wu et al. (2003).
Perform enzyme phosphorylation 41. Dissolve the following in 8 mL water in a 15-mL polystyrene round-bottom tube: 154 mg (0.83 mmol) triethanolamine hydrochloride 34.5 mg (0.17 mmol) MgCl⋅6H2O 50 mg (0.67 mmol) KCl 180 mg (0.4 mmol) S.7 220 mg (0.4 mmol) ATP 152 mg (0.8 mmol) PEP. 42. Adjust to pH 7.6 with 1 M NaOH, then add water to make the final volume 10 mL. 43. Add 50 µL (100 U) nucleoside diphosphate kinase and 80 µL (200 U) pyruvate kinase. Incubate the reaction mixture at 37°C. 44. Monitor the progress of the reaction by analytical HPLC. Take 2-µL aliquots, dilute with mobile phase A, and run on a Supelcosil LC-18-T column at a flow rate of 1.5 mL/min using the following gradient conditions: 0% to 100% mobile phase B over 10 min 100% mobile phase B for 8 min 100% to 0% mobile phase B over 2 min. The retention times of TzA4DP (S.7) and TzA4TP (S.10) should be ∼7.7 and ∼10.4 min, respectively. The retention time of PEP should be ∼3.8 min. Special care should be taken to prevent the column from being damaged by the ion-pairing reagent TBAP. The column should first be washed with an adequate volume (≥10 column volumes) of buffer A without TBAP, then with pure water (≥10 column volumes) before equilibrating the column with organic solvent (i.e., MeOH) for storage at room temperature.
45. After S.7 is completely converted to triphosphate S.10 (∼4 hr), add 180 mg (0.4 mmol) ADP to the mixture and continue to incubate at 37°C. 46. Monitor the consumption of PEP by HPLC as described in step 44. 47. Once the excess PEP is completely consumed, pass the mixture through a 25-mmdiameter 0.2-µm nylon syringe filter and lyophilize the filtrate to dryness. Purify S.10 on a boronate affinity gel column 48. Mix 5 g Affi-Gel 601 boronate affinity gel with water and slurry-pack into a 2.5 × 5–cm column. 49. Equilibrate the column with 100 mL of 1 M ammonium bicarbonate, pH 8.5. 50. Dissolve the residual solid from step 47 in 10 mL of 1 M ammonium bicarbonate, pH 8.5, and load the sample solution onto the boronate affinity gel column using a peristaltic pump set at 1 mL/min. Alternatively, sample loading and column elution can be done by gravity.
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51. Elute the column with the same buffer using a peristaltic pump set at 1 mL/min, with UV detection at 230 nm. 52. Collect and pool the fractions containing product in the first two column volumes. Bubble carbon dioxide into the solution until the pH reaches 7.2. Carbon dioxide may be conveniently generated from dry ice in a filtering flask with a stopper and a side hose outlet.
53. Lyophilize the solution to dryness. Dissolve in 50 mL water, adjust to pH 7.2 with carbon dioxide, and repeat lyophilization. Two or three lyophilization cycles may be required to completely remove excess ammonium bicarbonate.
Purify S.10 on a Q Sepharose FF anion-exchange column 54. Pack a 2.5 × 20–cm Q Sepharose FF anion-exchange column and elute with the following using a peristaltic pump at 5 mL/min: 300 mL 1 M KCl 300 mL H2O 300 mL 50 mM ammonium bicarbonate solution. 55. Connect the column to an MPLC system equipped with a programmable gradient pump system and a UV detector. 56. Dissolve the solid from step 53 in 10 mL of 50 mM ammonium bicarbonate solution and load the sample solution onto the column. 57. Elute the column with a linear gradient from 50 mM ammonium bicarbonate to 500 mM ammonium bicarbonate over 2 hr at a flow rate of 5 mL/min, with UV detection at 230 nm. 58. Analyze the appropriate fractions by HPLC as described in step 44. 59. Combine fractions containing triphosphate S.10 and lyophilize to dryness. 60. Confirm the product by 1H NMR, 13C NMR, 31P NMR, and ESI-MS. The final product, TzA4TP S.10 (ammonium salt), is obtained as a white fluffy powder with a yield of 120 mg or 58%. 1H NMR (D2O, 300 MHz): 8.64 (1H, s), 6.42 (1H, t, J = 6.0 Hz), 4.14 (1H, m), 4.01 (2H, m), 2.68 (1H), 2.50 (1H, m). 13C NMR (D2O, 75 MHz): 164.5, 141.9, 125.9, 89.5, 86.6, 70.4, 65.2, 39.9. 31P NMR (D2O, 121 MHz): –6.26 (d, Jp,p = 19.4 Hz), –10.97 (d, Jp,p = 19.4 Hz), –21.88 (t, Jp,p = 19.4 Hz). HRMS (ESI): calcd. for C8H14N4O13P3 466.9770 [M–H]–; found 466.9766. Characterization data for S.11 and S.12 can be found in Wu et al. (2003). BASIC PROTOCOL 2
SYNTHESIS OF AZOLE CARBOXAMIDE RIBONUCLEOSIDE TRIPHOSPHATES This protocol outlines a general procedure for the synthesis and purification of the azole carboxamide ribonucleoside triphosphate (Fig. 13.2.2; Wu et al., 2003). Specific protocols are given for the synthesis of rTz2A4 nucleoside triphosphate (S.19). Synthesis of the other azole carboxamide ribonucleoside triphosphate in Figure 13.2.2 (S.20) can be accomplished using the same procedure (see Critical Parameters and Troubleshooting).
Chemoenzymatic Preparation of Nucleoside Triphosphates
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O
O N
HO
TsO
O
OH
N O
a, b
O P c, d
13 N = Tz 2A4 14 N = TzA 4
O
O
H
OCH3
O
OH
OH
15 N = Tz 2A4 16 N = TzA 4
17 N = Tz 2A4 18 N = TzA 4 pyruvate
CONH2 N
N
O
P O
O
OH
Tz2A4 N =
O
pyruvate kinase
N N
PEP CONH2
N TzA4 N =
O
HO P O P
N N
O−
O
O
O−
O
P
N
O
O
O− OH
OH
19 N = Tz 2A4 20 N = TzA 4
Figure 13.2.2 General procedure for the synthesis of azole carboxamide ribonucleoside triphosphates (see Basic Protocol 2). Reagents: (a) (CH3O)3CH, TsOH, THF, room temperature (steps 1 and 2); (b) TsCl, DMAP, CH2Cl2, room temperature (steps 6 and 7); (c) (NBu4)3HP2O7, CH3CN, room temperature (steps 25 and 26); (d) TFA/H2O, pH 2.0, then NH4OH/H2O, pH 8.5 (steps 28 and 29). Abbreviations: TzA4, (1H)-1,2,3-triazole-4-carboxamide; Tz2A4, (2H)-1,2,3-triazole-4-carboxamide.
Materials Azole carboxamide ribonucleoside (Fig. 13.2.2): S.13 (N = rTz2A4; Lehmkuhl et al., 1972) or S.14 (N = rTzA4; Lehmkuhl et al., 1972) p-Toluenesulfonic acid monohydrate (TsOH; Aldrich) Tetrahydrofuran (THF), anhydrous Trimethyl orthoformate [(CH3O)3CH; Aldrich], anhydrous Methylene chloride, anhydrous (distill from phosphorus pentoxide and store over 4-Å molecular sieves) Methanol, ACS reagent grade 5% phosphomolybdic acid solution (see recipe) Pyridine, anhydrous p-Toluenesulfonyl chloride (TsCl) 4-N,N-Dimethylaminopyridine (DMAP) Ethyl acetate, ACS reagent grade 200- to 400-mesh silica gel 60 Hexane, ACS reagent grade Phosphorus pentoxide (P2O5) Acetonitrile, anhydrous
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Argon Tris(tetra-n-butylammonium) hydrogen pyrophosphate (see Basic Protocol 1) Deuterated acetonitrile (Aldrich) Dowex AG 50W-X8 cation-exchange resin (100 to 200 mesh, Bio-Rad) 1 M HCl 1 M and concentrated (28%) ammonium hydroxide (NH4OH) Trifluoroacetic acid (TFA) Mobile phase A: 0.1 M potassium phosphate buffer, pH 6.0 (APPENDIX 2A) containing 4 mM tetrabutylammonium dihydrogenphosphate (TBAP, Aldrich; added from a 1.0 M stock) Mobile phase B: 70:30 (v/v) mobile phase A/methanol, pH 7.2 CF11 fibrous cellulose (Whatman) 50, 100, and 500 mM ammonium bicarbonate (NH4HCO3) solutions (no pH adjustment) 1% (w/v) sulfosalicylic solution (see recipe) 0.2% (w/v) ferric chloride solution (see recipe) Triethanolamine Magnesium chloride hexahydrate (MgCl⋅6H2O) Potassium chloride (KCl) Sodium phosphoenolpyruvate monohydrate (PEP; Sigma) 1 M sodium hydroxide (NaOH) Pyruvate kinase (see recipe) Ammonium bicarbonate Ammonium hydroxide, concentrated Affi-Gel 601 boronate affinity gel (Bio-Rad) 1 M ammonium bicarbonate (NH4HCO3), pH 8.5 (see recipe) Carbon dioxide source (e.g., dry ice) Q Sepharose FF anion-exchange resin (Bio-Rad) 1 M potassium chloride (KCl)
Chemoenzymatic Preparation of Nucleoside Triphosphates
20-mL one-neck round-bottom flasks, oven dried TLC plates: Silica gel 60 F254 polyester-backed TLC plates (Aldrich) Cellulose TLC plates (EM Science) Heat gun 254-nm UV light lamp Rotary evaporator with adjustable temperature and water aspirator 2 × 25–cm, 2.5 × 5–cm, 2.5 × 10–cm, 2.5 × 20-cm, and 2.5 × 25–cm chromatography columns Vacuum desiccator 250-mL round-bottom flask Lyophilizer HPLC system (e.g., Beckman System Gold) including: 128 Solvent Module 166 Detector set at 230 nm 25-cm × 4.6-mm × 5-µm-i.d. Supelcosil LC-18-T column 1-L beaker TLC sprayer (Analtech) 25-mm-diameter, 0.2-µm nylon syringe filter (Fisher) Peristaltic pump Medium-pressure liquid chromatography (MPLC) system (e.g., ISCO LC system) with following equipment: Model 2360 gradient programmer
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Model 2350 HPLC pump V4 variable wavelength absorbance detector Additional reagents and equipment for thin-layer chromatography (TLC, APPENDIX 1 13 31 3D), column chromatography (APPENDIX 3E), and H NMR, C NMR, P NMR, and ESI-MS Protect 2′,3′-hydroxyl groups of rTzA4 (S.13) and prepare 5′-tosylate (S.15) 1. Add 250 mg (1.0 mmol) rTz2A4 azole carboxamide ribonucleoside (S.13) and 210 mg (1.1 mmol) TsOH to a 20-mL one-neck round-bottom flask equipped with a magnetic stir bar. 2. Add 5 mL anhydrous THF and 0.55 mL (5.0 mmol) trimethyl orthoformate to the flask and stir the reaction mixture at room temperature. The suspension becomes clear after 20 min.
3. Monitor the progress of the reaction by analytical TLC (APPENDIX 3D) as follows: a. Withdraw a small sample using a capillary tube and spot on a silica gel 60 F254 polyester-backed TLC plate. b. Develop the TLC plate in 10:1 (v/v) methylene chloride/methanol. c. Visualize by dipping the plate into 5% phosphomolybdic acid solution and heating with a heat gun. The starting material and product appear as blue or dark blue spots on a green background.
4. When TLC indicates the reaction is complete (∼2 hr), add 0.5 mL pyridine to the solution. 5. Remove the solvent using a rotary evaporator equipped with a water aspirator. 6. Dissolve the residue in 5 mL anhydrous methylene chloride. 7. Add 230 mg (1.2 mmol) TsCl and 160 mg (1.3 mmol) DMAP and stir the reaction mixture at room temperature. 8. Monitor the progress of the reaction by TLC on a silica gel plate in 10:1 (v/v) ethyl acetate/methanol as described in step 3. 9. When TLC indicates the reaction is complete (∼24 hr), remove the magnetic stir bar from the flask. 10. Pack 30 g of 200- to 400-mesh silica gel 60 into a 2 × 25–cm chromatography column in 1:1 (v/v) hexanes/ethyl acetate. 11. Load the reaction mixture on top of the silica gel column and elute with 10:9:1 (v/v/v) hexane/ethyl acetate/methanol. 12. Collect 10-mL fractions and check by silica gel TLC as described in step 3. 13. Combine fractions containing product and evaporate to dryness. 14. Dry the product in a desiccator over phosphorus pentoxide in vacuo. Flash chromatography yields 310 mg (70%) of S.15 as white solid. 1H NMR for diastereomer (DMSO-d6, 300 MHz): 8.09, 8.10 (1H, s), 7.89 (1H, s), 7.61 (1H, s), 7.58 (2H, d), 7.35 (2H, d), 6.33 (1H, d, J = 6.0 Hz), 6.03, 6.14 (1H, s), 5.20 (2H, m), 4.49 (1H, m), 4.06 (2H, m), 3.21, 3.29 (3H, s), 2.39 (3H, s). MS (ESI): 463 [M+Na]+. Anal. calcd. for C17H20O8N4S: C 46.36, H 4.58, N 12.72; found: C 46.74, H 4.55, N 12.61. Characterization data for S.16 can be found in Wu et al. (2003).
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Prepare 2′,3′-protected diphosphate 15. Add 350 mg (0.8 mmol) S.15 to an oven-dried 20-mL one-neck round-bottom flask. 16. Coevaporate S.15 with 5 mL anhydrous acetonitrile three times. 17. Evacuate and flush the flask with argon. 18. Add 1.4 g (1.6 mmol) tris(tetra-n-butylammonium) hydrogen pyrophosphate to the flask. 19. Transfer 0.5 mL anhydrous acetonitrile into the reaction flask and stir the reaction mixture at room temperature under argon. 20. Monitor the progress of the reaction by 1H NMR as follows: a. Withdraw 50 µL reaction mixture and dilute with 0.5 mL deuterated acetonitrile. b. Transfer the sample to a dried NMR tube and acquire a 1H NMR spectrum. c. Monitor the progress of the reaction by the change in the 1H NMR spectrum between 7 and 9 ppm, where the tosylate moiety appears as an AA′XX′ spin system. The four-line pattern for the tosylate moves upfield by ∼0.1 to 0.2 ppm upon conversion to the tosylate anion. TLC is not suitable for monitoring the progress of the reactions because the tetra-n-butylammonium salt does not give a well-defined spot.
21. When 1H NMR indicates the reaction is complete (∼2 days), dilute the reaction mixture with 5 mL water. 22. Pack Dowex AG 50W-X8 cation-exchange resin in water into a 2.5 × 10–cm column. Transform the cation-exchange resin to the ammonium form by washing the column with 150 mL (3 column volumes) of the following solutions in sequence: 1 M HCl H2O 1 M NH4OH H2O. 23. Load the solution from step 21 onto the column and elute with 100 mL (2 column volumes) of deionized water. Collect the eluent in a 250-mL round-bottom flask. 24. Lyophilize the solution to dryness. Deprotect 2′,3′-hydroxyl groups 25. Dissolve the resulting solid in 20 mL deionized water and adjust the pH to 2.0 with TFA. 26. Stir the solution 2 hr at room temperature. 27. Monitor the progress of the reaction by analytical HPLC. Take 2-µL aliquots, dilute with mobile phase A, and run on a Supelcosil LC-18-T column at a flow rate of 1.5 mL/min using the following gradient conditions: 0% to 100% mobile phase B over 10 min 100% mobile phase B for 8 min 100% to 0% mobile phase B over 2 min. Chemoenzymatic Preparation of Nucleoside Triphosphates
The retention times of the 2′,3′-protected diphosphate should be 12.9 and 13.4 min (mixture of diastereomers); the retention times of the acid deprotection intermediate should be 10.7 and 11.1 min (mixture of 2′- and 3′-formate). For care of the column, see Basic Protocol 1, step 44.
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28. When HPLC indicates that hydrolysis is complete (∼3 hr), adjust the pH of the solution to 8.5 with 1 M NH4OH. 29. Stir the solution at room temperature and monitor the progress of the reaction by HPLC. The retention time of the fully deprotected diphosphate S.17 should be 8.6 min.
30. When HPLC indicates that deprotection is complete (∼2 hr), lyophilize the reaction mixture to dryness. Purify S.17 31. Pack a Whatman CF11 fibrous cellulose column as follows: a. Mix 500 mL (dry volume) CF11 fibrous cellulose power with 350 mL water in a 1-L beaker by vigorous stirring with a glass rod. b. Slurry-pack the cellulose into a 2.5 × 25–cm chromatography column. c. Wash the column with 300 mL water, 300 mL acetonitrile, and 300 mL of 1:1 (v/v) acetonitrile/water in sequence. d. Equilibrate the column with 300 mL of 7:3:2 (v/v/v) acetonitrile/100 mM ammonium bicarbonate/concentrated (28%) ammonium hydroxide. 32. Extract the solid obtained from step 30 with 5 mL of the same acetonitrile/ammonium bicarbonate/ammonium hydroxide buffer. Pellet the precipitate. The product is soluble in the buffer and the white precipitate formed is excess inorganic pyrophosphate.
33. Load the sample solution onto the column and elute with the same buffer by flash chromatography, collecting 10-mL fractions. 34. Analyze every second fraction on a cellulose TLC plate as follows: a. Spot the sample on the cellulose TLC plate and develop in 7:3:2 (v/v/v) acetonitrile/100 mM ammonium bicarbonate/concentrated ammonium hydroxide. b. Spray the plate with 1% sulfosalicylic acid solution until thoroughly wetted but not dripping, and allow to air dry 5 min. c. Lightly spray 0.2% ferric chloride solution onto the plate and visualize the spots. When visualized by sulfosalicylic acid/ferric chloride spray, phosphate-containing compounds appear as white spots on a pink background. A second light spray with ferric chloride may be necessary to make the spots pronounced (Rf = 0.30 for S.17).
35. Combine the fractions containing product and remove acetonitrile using a rotary evaporator with the bath temperature set below 30°C. 36. Lyophilize the resulting aqueous solution to dryness. 37. Confirm the product by 1H NMR, 13C NMR, 31P NMR, and ESI-MS. Flash cellulose chromatography yields 254 mg (70%) S.17 as a fluffy white powder. 1H NMR (D2O, 300 MHz): 8.01 (1H, s), 5.98 (1H, d, J = 3.6 Hz), 4.49 (1H, m), 4.18 (1H, m), 3.93 (2H, m). 13C NMR (D2O, 75 MHz): 164.3, 143.2, 136.5, 95.6, 84.4, 74.6, 70.7, 65.4. 31 P NMR (D2O, 121 MHz): –7.78 (d, Jp,p = 20.6 Hz), –10.79 (d, Jp,p= 20.6 Hz). HRMS (ESI): calcd. for C8H15N4O11P2 405.0213 [M+H]+; found 405.0208. Characterization data for S.18 can be found in Wu et al. (2003). Nucleoside Phosphorylation and Related Modifications
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Perform enzyme phosphorylation 38. Dissolve the following in 8 mL deionized water: 154 mg (0.83 mmol) triethanolamine 34.5 mg (0.17 mmol) MgCl⋅6H2O 50 mg (0.67mmol) KCl 180 mg (0.4 mmol) S.17 152 mg (0.8 mmol) PEP. 39. Adjust the pH to 7.6 with 1 M NaOH, then add deionized water to adjust the final volume to 10 mL. 40. Add 16 µL (40 U) pyruvate kinase and incubate the reaction mixture at 37°C. 41. Monitor the progress of the reaction by analytical HPLC as described in step 27. Retention times are ∼ 9.1 and ∼10.6 min for rTzA4DP and rTzA4TP, respectively.
42. When HPLC indicates complete conversion of S.17 to triphosphate S.19 (∼4 hr), add 790 mg ammonium bicarbonate to the solution. 43. Adjust the pH to 8.5 with concentrated (28%) ammonium hydroxide and filter the mixture through a 25-mm-diameter, 0.2-µm nylon syringe filter. Purify S.19 on a boronate affinity gel column 44. Mix 5 g Affi-Gel 601 boronate affinity gel with water and slurry-pack into a 2.5 × 5–cm column. 45. Equilibrate with 100 mL of 1 M ammonium bicarbonate , pH 8.5. 46. Load the sample solution from step 43 onto the column using a peristaltic pump at 1 mL/min. Alternatively, sample loading and column elution can be done by gravity.
47. Elute the column with 90 mL of 1 M ammonium bicarbonate using a peristaltic pump at 1 mL/min with UV detention at 230 nm. Excess PEP and enzyme reaction buffer are eluted out in this step.
48. Elute the column with deionized water at 2 mL/min, collecting 5-mL fractions with UV detection at 230 nm. S.19 is eluted with water.
49. Combine all the fractions containing S.19 and bubble carbon dioxide into the solution until the pH reaches 7.2. Carbon dioxide may be conveniently generated from dry ice in a filtering flask with a stopper and a side hose outlet.
50. Lyophilize the solution to dryness. Resuspend in 50 mL water and adjust to pH 7.2 with carbon dioxide. Repeat lyophilization. Two or three lyophilization cycles may be required to completely remove excess ammonium bicarbonate.
Chemoenzymatic Preparation of Nucleoside Triphosphates
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Purify S.19 on a Q Sepharose FF anion-exchange column 51. Pack a 2.5 × 20–cm Q Sepharose FF anion-exchange column and elute with the following using a peristaltic pump at 5 mL/min: 300 mL 1 M KCl 300 mL H2O 300 mL 50 mM ammonium bicarbonate solution. 52. Connect the column to an MPLC system equipped with a programmable gradient pump system and a UV detector. 53. Dissolve the solid from step 50 in 10 mL of 50 mM ammonium bicarbonate solution and load the sample solution onto the column. 54. Elute the column with a linear gradient from 50 mM ammonium bicarbonate to 500 mM ammonium bicarbonate over 2 hr at a flow rate of 5 mL/min with UV detection at 230 nm. 55. Analyze the appropriate fractions by HPLC as described in step 27. 56. Combine fractions containing S.19 and lyophilize to dryness. 57. Confirm the product by 1H NMR, 13C NMR, 31P NMR, and ESI-MS. Purification yields 78 mg (72%) S.19 (ammonium salt) as a white fluffy solid. 1H NMR (D2O, 300 MHz): 8.05 (1H, s), 6.02 (1H, d, J = 3.9 Hz), 4.24 (1H, m), 4.03 (2H, m). 13C NMR (D2O, 75 MHz): 163.9, 142.8, 136.1, 95.1, 84.1, 74.1, 70.4, 65.3. 31P NMR (D2O, 121 MHz): –9.11 (d, Jp,p = 19.4 Hz), –11.18 (d, Jp,p = 19.4 Hz), –22.56 (t, Jp,p = 19.4 Hz). HRMS (ESI): calcd. for C8H16N4O14P3 484.9876 [M+H]+; found 484.9869. Characterization data for S.20 can be found in Wu et al. (2003).
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX. Ammonium bicarbonate solution, 1 M, pH 8.5 Dissolve 39.5 g ammonium bicarbonate (NH4HCO3) in 400 mL water and adjust to pH 8.5 with concentrated (28%) ammonium hydroxide (NH4OH). Dilute to 500 mL with water. Make fresh every time. Ferric chloride solution, 0.2% (w/v) Dissolve 0.2 g ferric chloride in 100 mL of 4:1 (v/v) ethanol/water. Use within 6 hr. Nucleoside diphosphate kinase Purchase nucleoside diphosphate kinase (EC 2.7.4.6; from baker’s yeast; Sigma) as a lyophilized powder. Reconstitute with water to 2 U/µL. Store indefinitely at −20°C. Phosphomolybdic acid solution, 5% (w/v) Dissolve 25 g of phosphomolybdic acid in 500 mL of 95% (v/v) ethanol. Store indefinitely in an amber reagent bottle at room temperature. Pyruvate kinase Purchase pyruvate kinase (EC 2.7.1.40; Type VII from rabbit muscle; Sigma) at 2.5 U/µL in 50% glycerol containing 0.01 M phosphate, pH 7.0. Store indefinitely at 4°C.
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Sulfosalicylic solution, 1% (w/v) Dissolve 1.0 g sulfosalicylic acid in 100 mL of 3:2 (v/v) ethanol/water. Use within 6 hr. COMMENTARY Background Information Synthetic modified nucleobases as alternative substrates for DNA and RNA polymerases Synthetic modified nucleobases designed to pair in unusual ways with the natural nucleic acid base have many potential applications in nucleic acid biochemistry, ranging from biochemical tools for probing nucleic acid structures or protein–nucleic acid interactions to tools for re-engineering DNA and ultimately proteins. One class of materials of particular interest is the azole carboxamide nucleobases. Designed as universal bases, they have the interesting property of displaying multiple conformations and can mimic different sets of natural bases in the context of DNA replication. The variation in electronic distribution in these compounds offers a range of hydrogen-bonding features (Hoops et al., 1997; Zhang et al., 1998). The azole carboxamide deoxyribonucleoside triphosphates (see Fig. 13.2.1) are used as alternate substrates for DNA polymerase to probe enzyme fidelity during DNA replication; similarly, the azole carboxamide ribonucleoside triphosphates (see Fig. 13.2.2) can be used as alternate substrates for RNA polymerase to probe structure and function of RNA polymerase.
Chemoenzymatic Preparation of Nucleoside Triphosphates
Synthesis of nucleoside triphosphates The general utility of the “one-pot threestep” procedures for the synthesis of purine or pyrimidine nucleoside triphosphates has been established (Ludwig, 1981; Mishra and Broom, 1991; Burgess and Cook, 2000). Other multistep methods that rely on activated nucleoside monophosphates have also had long-standing application for these more standard nucleotides (Moffatt and Khorama, 1961; Hoard and Ott, 1965; Simoncsits and Tomasz, 1975; Tomasz et al., 1978). These methods involve the use of the Yoshikawa procedure or a variation of the Yoshikawa procedure which employs conditions for selective 5′-phosphorylation with phosphorus oxychloride as the primary donor (Yoshikawa et al., 1967). This reactive electrophilic phosphorus reagent presents limitations with many heterocycles, such as azole carbox-
amides, since attack at the carboxamide leads to side reactions. Alternative strategies that use other electrophilic P3 or P5 reagents are equally limiting because many heterocycles are susceptible to addition reactions with these reagents (van Boom et al., 1975; Ludwig and Eckstein, 1989). Enzyme-catalyzed synthesis of nucleoside triphosphates has proven to be a general method for a variety of purine and pyrimidine nucleoside analogs (Wong et al., 1983; Simon et al., 1988). Nucleoside diphosphate kinase (NDPK) catalyzes the transfer of the γ-phosphate from a nucleoside triphosphate to a nucleoside diphosphate. The enzyme is remarkably nonspecific with regard to the nucleotide substrate; it uses di- and triphosphate nucleotides with either deoxyribose or ribose and any of the natural purine and pyrimidine bases (Ratliff et al., 1964; Mourad and Parks, 1966). A number of the unnatural purine and pyrimidine nucleotide analogs can also serve as substrates (Miller et al., 1992; Kamiya and Kasai, 1999). The authors’ experiments with NDPK show that the azole carboxamide nucleoside diphosphates are substrates for NDPK. The reaction equilibrium offers a general method for the preparation of nucleoside triphosphates from diphosphates. In the case of the azole carboxamide deoxyribonucleoside diphosphates, incorporation of an ATP regeneration system using phosphoenolpyruvate (PEP) and pyruvate kinase allows for efficient conversion to the triphosphates (Hirschbein et al., 1982). In the case of azole carboxamide ribonucleoside diphosphates, preliminary studies show that NPDK and dATP coupled with dATP regeneration by pyruvate kinase and PEP could efficiently convert azole carboxamide nucleoside diphosphates to triphosphates. However, pyruvate kinase was later found to directly catalyze the phosphorylation of the azole carboxamide ribonucleoside diphosphates, allowing a simplified system to complete the transformation. Most biochemical applications of nucleoside triphosphates require a high degree of purity, especially with respect to other contaminating nucleotides. An efficient purification procedure was optimized for the desired nucleoside triphosphates following enzymatic
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phosphorylation. In the case of the deoxyribonucleoside triphosphate synthesis, separation from ATP is necessary, which is not readily achieved by conventional ion-exchange chromatography. Boronate affinity gel has been used to separate ribonucleoside 5′-phosphates from deoxyribonucleotides due to the formation of a complex between the borate group and cis-diol of the ribonucleoside (Schott, 1972; Schott et al., 1973). Passing the enzyme phosphorylation mixture through the boronate affinity gel at high pH gives efficient separation of the resulting azole carboxamide deoxyribonucleoside triphosphate from ATP. Purification of the azole carboxamide ribonucleoside triphosphate using boronate affinity gel is also more straightforward. The methods described in this unit for chemoenzymatic preparation of nucleoside triphosphates extend the use of nucleoside diphosphate synthesis (Davisson et al., 1987) to a general route for nucleoside triphosphate synthesis. It is likely that this strategy will be applicable as a general protocol for preparation of triphosphates of a wide variety of base- and sugar-modified nucleoside analogs.
Critical Parameters and Troubleshooting Since the nucleoside diphosphates and triphosphates prepared in these procedures are both acid and base labile, the pH of the product-containing solution must stay between 2.0 and 9.0, and all products must be stored at −20°C. Preparation of nucleoside diphosphates. High concentrations of tosylate and pyrophosphate are necessary for the displacement reaction to proceed at reasonable rates. Cellulose TLC is critical for detecting diphosphate-containing fractions after flash cellulose chromatography. Using EM Science TLC plates and following the procedure given for staining the plates will give the best results. Enzyme phosphorylation. When constructing the azole carboxamide deoxyribonucleoside triphosphates, it is very important to completely convert excess PEP to pyruvate by adding ADP after phosphorylation of the diphosphate is complete (see Basic Protocol 1). Excess PEP interferes with the final purification of triphosphate on a Q Sepharose FF anion-exchange column. For ribonucleosides, the excess PEP is easily separated from the triphosphate by boronate affinity gel chromatography. The PEP passes through the column using 1 M
ammonium bicarbonate, and the triphosphate is then eluted using water. Purification of triphosphates. It is necessary to use a slow flow rate (1 mL/min) for the boronate affinity gel column since faster flow rates may not allow complete binding of the ribonucleotides to the boronate affinity gel. Synthesis of alternate nucleoside triphosphates. Synthesis of the other azole carboxamide deoxyribonucleoside triphosphates shown in Fig. 13.2.1 (i.e., S.11 and S.12) can be accomplished using the exact conditions and quantities presented in the steps of Basic Protocol 1. Simple adjustments to the reaction times for the tosylation and diphosphate displacement are needed, and different chromatography solvents are used to purify the S.5 and S.6 tosylates; these adjustments can be found in Wu et al. (2003). For synthesis of the other azole carboxamide ribonucleoside triphosphate shown in Fig. 13.2.2 (S.20), there is one major modification with regard to the reaction conditions for synthesis of 2′,3′-protected 5′tosylate S.16 from nucleoside S.14; this modification is also detailed in Wu et al. (2003). Preparation of the nucleoside diphosphate S.18 and enzyme phosphorylation to S.20 can be accomplished using the exact conditions and quantities presented in the steps of Basic Protocol 2.
Anticipated Results The protocols described in this unit allow preparation of the azole carboxamide deoxyribo- and ribonucleoside triphosphates in good yields and high purity. In the first method (see Basic Protocol 1), selective tosylation of the azole carboxamide deoxyribonucleosides should give 5′-O-tosyl nucleosides in yields between 60% and 70%. Preparation of nucleoside diphosphates from 5′-O-tosyl nucleosides should give yields between 70% and 90%. Enzyme phosphorylation and subsequent purification should give deoxyribonucleoside triphosphates in yields between 58% and 89%. In the second method (see Basic Protocol 2), protection of the 2′,3′-hydroxyl groups of the azole carboxamide ribonucleosides and subsequent tosylation should give 5′-O-tosyl nucleosides in 52% to 70% yields. Diphosphate displacement and deprotection of 2′,3′-hydroxyl groups should give ribonucleoside diphosphates in yields between 66% and 70%. Enzyme phosphorylation and subsequent purification should give ribonucleoside triphosphates in ∼70% yield.
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13.2.17 Current Protocols in Nucleic Acid Chemistry
Supplement 16
Time Considerations In the first method (see Basic Protocol 1), preparation of 5′-O-tosyl nucleosides may take 2 to 4 days (4 days for S.4, or 1 to 2 days for the alternate tosylates S.5 and S.6). Preparation of nucleoside diphosphates should take 3 to 4 days for the displacement reaction, cation exchange, and cellulose chromatography. Enzyme phosphorylation and purification of resulting triphosphates should take ∼7 days to complete. In the second method (see Basic Protocol 2), preparation of 5′-O-tosyl nucleosides may take 3 to 4 days for protection of 2′,3′-hydroxyl groups and tosylation. Preparation of nucleoside diphosphates requires 5 days for the displacement reaction, cation exchange, deprotection of 2′,3′-hydroxyl groups, and cellulose chromatography. Enzyme phosphorylation and purification of resulting triphosphates needs ∼6 days to complete.
Literature Cited Burgess, K. and Cook, D. 2000. Syntheses of nucleoside triphosphates. Chem. Rev. 100:20472060. Davisson, V.J., Davis, D.R., Dixit, V.M., and Poulter, C.D. 1987. Synthesis of nucleotide 5′-diphosphates from 5′-O-tosyl nucleosides. J. Org. Chem. 52:1794-1801. Hirschbein, B.L., Mazenod, F.P., and Whitesides, G.M. 1982. Synthesis of phosphoenolypyruvate and its use in ATP cofactor regeneration. J. Org. Chem. 47:3765-3766. Hoard, D.E. and Ott, D.G. 1965. Conversion of mono- and oligodeoxyribonucleotides to 5′triphosphates. J. Am. Chem. Soc. 87:1785-1788. Hoops, G.C., Zhang, P., Johnson, W.T., Paul, N., Bergstrom, D.E., and Davisson, V.J. 1997. Template directed incorporation of nucleotide mixtures using azole-nucleobase analogs. Nucl. Acids Res. 25:4866-4871. Kamiya, H. and Kasai, H. 1999. Preparation of 8-hydroxy-dGTP and 2-hydroxy-dATP by a phosphate transfer reaction by nucleosidediphosphate kinase. Nucleos. Nucleot. Nucl. Acids 18:307-310. Lehmkuhl, F.A., Witkowski, J.T., and Robins, R.K. 1972. Synthesis of 1,2,3-triazole nucleosides via the acid-catalyzed fusion procedure. J. Heterocyclic Chem. 9:1195-1201.
Chemoenzymatic Preparation of Nucleoside Triphosphates
Makabe, O., Suzuki, H., and Umezawa, S. 1977. Syntheisi of D-arabinofuranosyl and 2′-deoxy-Dribofuranosyl 1,2,3-triazolecarboxamides. Bull. Chem. Soc. Jpn. 50:2689-2693. Miller, W.H., Daluge, S.M., Garvey, E.P., Hopkins, S., Reardon, J.E., Boyd, F.L., and Miller, R.L. 1992. Phosphorylation of carbovir enantiomers by cellular enzymes determines the stereoselectivity of antiviral activity. J. Biol. Chem. 267:21220-21224. Mishra, N.C. and Broom, A.D. 1991. A novel synthesis of nucleoside triphosphates. J. Chem. Soc. Chem. Comm. 1276-1277. Moffatt, J.G. and Khorama, H.G. 1961. Nucleoside polyphosphates. X1. The synthesis and some reactions of nucleoside-5′ phosphoromorpholidates and related compounds. Improved methods for the preparation of nucleoside-5′ polyphosphates. J. Am. Chem. Soc. 83:649-658. Mourad, N. and Parks, R.E. Jr. 1966. Erythrocytic nucleoside diphosphokinase. II. Isolation and kinetics. J. Biol. Chem. 241:271-278. Ratliff, R.C., Weaver, R.H., Lardy, H.A., and Kuby, S.A. 1964. Nucleoside triphosphate-nucleoside diphosphate transphosphorylase (nucleoside diphosphokinase). I. Isolation of the crystalline enzyme from brewer’s yeast. J. Biol. Chem. 239:301-309. Schott, H. 1972. New dihydroxyboryl-substituted polymers for column-chromatographic separation of ribonucleoside-deoxyribonucleoside mixtures. Angew. Chem. Int. Ed. Engl. 11:824825. Schott, H., Rudloff, E., Schmidt, P., Roychoudhury, R., and Kossel, H. 1973. A dihydroxyboryl-substituted methacrylic polymer for the column chromatographic separation of mononucleotides, oligonucleotides, and transfer ribonucleic acid. Biochemistry 12:932-938. Simon, E.S., Bednarski, M.D., and Whitesides, G.M. 1988. Generation of cytidine 5′-triphosphate using adenylate kinase. Tetrahedron Lett. 29:1123-1126. Simoncsits, A. and Tomasz, J. 1975. Nucleoside 5′-phosphordiamidates, synthesis and some properties. Nucl. Acids Res. 2:1223-1233. Tomasz, J., Simoncsits, A., Kajtar, M., Krug, R.M., and Shatkin, A.J. 1978. Chemical synthesis of 5′-pyrophosphate and triphosphate derivatives of 3′-5′ ApA, ApG, GpA and GpG. CD study of the effect of 5′-phosphate groups on the conformation of 3′-5′ GpG. Nucl. Acids Res. 5:2945-2957.
Ludwig, J. 1981. A new route to nucleoside 5′triphosphates. Acta Biochim. Biophys. Acad. Sci. Hung. 16:131-133.
van Boom, J.H., Crea, R., Luyten, W.C., and Vink, A.B. 1975. 2,2,2-Tribromoethyl phosphoromorpholinochloridate: A convenient reagent for the synthesis of ribonucleoside mono-, di- and triphosphates. Tetrahedron Lett. 16:2779-2782.
Ludwig, J. and Eckstein, F. 1989. Rapid and efficient synthesis of nucleoside 5′-O-(1-thiotriphosphates), 5′-triphosphates and 2′,3′-cyclophosphorothioates using 2-chloro-4H-1,3,2-benzodioxaphosphorin-4-one. J. Org. Chem. 54:631635.
Witkowski, J.T., Fuertes, M., Cook, P.D., and Robins, R.K. 1975. Nucleosides of 1,2,4-triazole-3carboxamide. Synthesis of certain pentofuranosyl, deoxypentofuranosyl, and pentopyranosyl 1,2,4-triazoles. J. Carbohydr. Nucleos. Nucleot. 2:1-36.
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Current Protocols in Nucleic Acid Chemistry
Wong, C.H., Haynie, S.L., and Whitesides, G.M. 1983. Preparation of a mixture of nucleoside triphosphates from yeast RNA: Use in enzymic synthesis requiring nucleoside triphosphate regeneration and conversion to nucleoside diphosphate sugars. J. Am. Chem. Soc. 105:115-117.
Key References
Wu, W., Bergstrom, D.E., and Davisson, V.J. 2003. A combination chemical and enzymatic approach for the preparation of azole carboxamide nucleoside triphosphate. J. Org. Chem. 68:38603865.
This paper describes synthesis of nucleoside diphosphates from 5′-O-tosyl nucleosides.
Yoshikawa, M., Kato, T., and Takenishi, T. 1967. A novel method for phosphorylation of nucleos ides to 5′-nucleotides. Tetrahedron Lett. 50:5065-5068. Zhang, P., Johnson, W.T., Klewer, D., Paul, N., Hoops, G., Davisson, V.J., and Bergstrom, D.E. 1998. Exploratory studies on azole carboxamides as nucleobase analogs: Thermal denaturation studies on oligodeoxyribonucleotide duplexes containing pyrrole-3-carboxamide. Nucl. Acids Res. 26:2208-2215.
Burgess and Cook, 2000. See above. This paper gives a review on syntheses of nucleoside triphosphates. Davisson et al., 1987. See above.
Wu et al., 2003. See above. This paper describes synthesis of the azole nucleoside triphosphate and compound characterization.
Contributed by Weidong Wu, Donald E. Bergstrom, and V. Jo Davisson Purdue University West Lafayette, Indiana
Nucleoside Phosphorylation and Related Modifications
13.2.19 Current Protocols in Nucleic Acid Chemistry
Supplement 16
Synthesis and Polymerase Incorporation of 5-Amino-2,5-Dideoxy-5-N-Triphosphate Nucleotides
UNIT 13.3
This unit describes procedures for synthesizing 5 -amino-2 ,5 -dideoxy analogs of four common 2 -deoxynucleosides (A, C, G, and T) and corresponding nucleotides, and the efficient incorporation of the latter into DNA (Wolfe et al., 2002). Since the 5 -amino functional group is much more reactive than the native 5 -hydroxyl group, these analogs can be exploited for various applications where differential chemical reactivity between natural and modified nucleotides is desirable. Unlike modifications on diverse heterocyclic structures, the 5 -modification on the sugar moiety is common to all nucleotides. Consequently, the preparation of these analogs is more likely to be generalizable among all four bases, and the products are more likely to exhibit similar traits in chemical reactivity and substrate specificity. Through the intermediacy of 5 -azido nucleosides (N3 -dNs), 5 -amino nucleosides (NH2 dNs) are conveniently prepared through robust chemical reactions such as tosylation, azide exchange, and the Staudinger reaction. Efficient conversion of NH2 -dNs to corresponding NH2 -dNTPs is achieved using an elegant one-step reaction with trisodium trimetaphosphate (TMP), which was first reported in the 1970s for the preparation of NH2 -dTTP (Letsinger et al., 1976a and b). By adding tris(hydroxymethyl)aminomethane (Tris) to neutralize protons generated during the reaction, the yield and stability of NH2 dNTPs have been significantly improved. Basic Protocols 1 and 2 describe detailed procedures for displacing the 5 -hydroxyl group of pyrimidines and purines, respectively, with a 5 -azido group. Tosylation at the 5 -hydroxyl followed by azide exchange is applied for purines (Fig. 13.3.1; see Basic Protocol 1), whereas a one-step reaction using carbon tetrabromide, triphenylphosphine, and lithium azide is utilized for pyrimidines (Fig. 13.3.2; see Basic Protocol 2). Because it is necessary to use protected dA, dC, and dG for these reactions, an additional hydrolysis step is required to unmask the exocyclic amino groups. The reduction of N3 -dNs to generate NH2 -dNs is achieved by the Staudinger reaction (Mungal et al., 1975), and is described in Basic Protocol 3 (Fig. 13.3.3). Efficient conversion of these nucleosides to corresponding NH2 -dNTPs is accomplished through a one-step reaction with TMP (Fig. 13.3.3) in the presence of Tris base (see Basic Protocol 4). Basic Protocol 5 outlines procedures for template-directed polymerase incorporation of NH2 -dNTPs. Each NH2 -dNTP participates in a DNA replication reaction in place of the corresponding dNTP, either completely or partially, through its exclusive or supplementary use. Mild acid treatment of the resulting DNA generates polynucleotide fragments that arise from specific cleavage at each modified nucleotide, providing a sequence ladder for each base when analyzed by polyacrylamide gel electrophoresis. Preparation methods for all four different NH2 -dNTPs are described in the protocols. If only one analog is needed for a desired application, NH2 -dTTP should be the first choice because NH2 -dT is commercially available; hence, Basic Protocols 1 and 2 can be skipped. Among the remaining analogs, pyrimidine NH2 -dC is somewhat more convenient to prepare than the purine analogs NH2 -dA and NH2 -dG. Nucleoside Phosphorylation and Related Modifications Contributed by Jia Liu Wolfe and Tomohiko Kawate Current Protocols in Nucleic Acid Chemistry (2004) 13.3.1-13.3.17 C 2004 by John Wiley & Sons, Inc. Copyright
13.3.1 Supplement 18
BASIC PROTOCOL 1
PREPARATION OF 5 -AZIDO-2 ,5 -DIDEOXYPURINES This protocol provides streamlined procedures to prepare N3 -dA and N3 -dG (Fig. 13.3.1). The synthetic route involves three commonly used chemical transformations and has been previously applied for synthesizing derivatives of N3 -dA and N3 -dG (Mag and Engels, 1989). Starting materials N4 -benzoyl-dA and N2 -isobutyryl-dG are tosylated at the 5 -hydroxyl group, then reacted with lithium azide to afford 5 -azido derivatives. Treatment with ammonia removes the protecting groups on the exocyclic amines of N3 dA and N3 -dG. N3 -dG is purified from the isobutyramide side product using solvent partitioning alone; however, purification of N3 -dA from benzoylamide requires column chromatography. CAUTION: All chemical reactions should be carried out in a fume hood to avoid exposure to toxic vapors. NOTE: In order to achieve satisfactory results, anhydrous reagents should be used and experiments should be performed under an inert atmosphere.
Materials
5 -Amino-2 ,5 Dideoxy-5 -NTriphosphate Nucleotides
Dry nitrogen (N2 ) or argon (Ar) N6 -Benzoyl-2 -deoxyadenosine (N6 -Bz-dA; S.1) Pyridine, anhydrous p-Toluenesulfonyl chloride (TsCl) N2 -Isobutyryl-2 -deoxyguanosine (N2 -i-Bu-dG; S.5) Ethyl acetate (EtOAc) Saturated aqueous sodium bicarbonate (NaHCO3 ) Saturated aqueous sodium chloride (brine) Sodium sulfate (Na2 SO4 ), anhydrous Dichloromethane (CH2 Cl2 ) Silica gel 60, 230 to 400 mesh Methanol (MeOH) Lithium azide (LiN3 ), concentrated to dryness from a 20% solution in dH2 O Dimethyl sulfoxide (DMSO), anhydrous Ammonium hydroxide (NH4 OH), concentrated aqueous solution Chloroform (CHCl3 ) Dry ice 250-, 100-, 50-, and 25-mL round-bottom flasks, oven dried Inert atmosphere/vacuum manifold with dry ice/2-propanol trap Balloons Tubing adaptor 10-mL syringes 18-G needles Rotary evaporator with a built-in dry ice/2-propanol trap, attached to a vacuum pump 250-mL separatory funnels 250-mL Erlenmeyer flasks Fritted funnels 250-mL glass chromatography columns Test tubes Thin-layer chromatography (TLC) plates, 0.25-mm silica gel 60F-254 on glass plates UV light source 60◦ and 50◦ C oil baths Vacuum pump
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Figure 13.3.1
Preparation of 5 -azido-dA and 5 -azido-dG.
10-mL screw-capped micro-vials (e.g., Accuform vials, Kimble) Lyophilizer Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) 5 -Tosylate nucleoside For adenosine derivative 1a. Under a N2 or Ar atmosphere, add 994 mg (2.8 mmol) of N6 -Bz-dA (S.1) and 20 mL anhydrous pyridine to an oven-dried 100-mL round-bottom flask containing a 3/4 -in. magnetic stir bar. Place the flask in an ice water bath on top of a magnetic stir plate. A N2 or Ar atmosphere can be provided through a tubing adaptor that connects the flask to a balloon filled with N2 or Ar, or an inert atmosphere/vacuum manifold attached to a regulated N2 or Ar tank.
2a. In a separate flask dissolve 802 mg (4.2 mmol, 1.5 eq.) TsCl in 10 mL anhydrous pyridine under N2 or Ar.
For guanosine derivative 1b. Under N2 or Ar atmosphere, add 777 mg (2.30 mmol) of N2 -i-Bu-dG (S.5) and 20 mL anhydrous pyridine in an oven-dried 100-mL round-bottom flask containing a 3/4 -in. magnetic stir bar. Place the flask in an ice water bath on top of a magnetic stir plate. 2b. In a separate flask dissolve 574 mg (3.01 mmol) TsCl in 5 mL anhydrous pyridine under N2 or Ar. 3. Transfer the TsCl solution to a 10-mL syringe through an 18-G needle and slowly add it to the cooled, stirring nucleoside in pyridine under N2 or Ar. Allow reaction to proceed 1 hr on ice.
Work up product 4. Remove the ice-water bath and transfer the flask to a 4◦ C refrigerator or cold room; continue stirring overnight under N2 or Ar provided through a balloon. 5. Place the reaction flask back into an ice bath, add 3 mL of dH2 O to the mixture, and continue to stir on ice for 1 hr.
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6. Concentrate the solution to near dryness on a rotary evaporator. 7. To the residue, add 150 mL of EtOAc and 30 mL of dH2 O. Stir the mixture using a magnetic stir plate to partition the products between organic and aqueous phases. 8. Transfer the biphasic mixture to a 250-mL separatory funnel to separate the organic and aqueous layers. 9. Collect the EtOAc (top) layer and wash it sequentially in the separatory funnel as follows: two times with 30 mL NaHCO3 , once with 30 mL dH2 O, and two times with 30 mL brine. 10. Collect the resulting EtOAc solution in a 250-mL Erlenmeyer flask containing ∼15 g of anhydrous Na2 SO4 and allow it to dry for 30 min. 11. Remove the drying agent by filtration using a fritted funnel. 12. Concentrate the EtOAc solution on a rotary evaporator to afford a white solid residue. 13. Dissolve the residue in ∼2 mL of CH2 Cl2 and load it onto a 100-mL chromatography column filled with 40 g of silica gel (APPENDIX 3E). Elute the column using CH2 Cl2 followed by 5:95 (v/v) MeOH/CH2 Cl2 and collect fractions in test tubes. 14. Analyze all fractions by TLC (APPENDIX 3D) on silica gel plates. Develop TLC plates with 10:90 (v/v) MeOH/CH2 Cl2 and analyze them under UV illumination. 15. Combine fractions that contain the desired product (Rf ≈ 0.5 for dA; Rf ≈ 0.44 for dG) and concentrate to dryness using a rotary evaporator to afford the product as a light yellow or off-white foam. 5 -TsO-N6 -Bz-dA (S.2): yield 1092 mg (72%). 1 H NMR (DMSO-d6 ): δ 2.32 (3H, s, Me), 2.37 (1H, ddd, J = 4.4, 6.7, 13.5 Hz, 2 a), 2.85 (1H, td, J = 6.5, 13.6 Hz, 2 b), 4.01 (1H, m, 4 ), 4.22 (1H, dd, J = 6.8, 10.7 Hz, 5 a), 4.31 (1H, dd, J = 3.6, 10.7 Hz, 5 b), 4.47 (1H, m, 3 ), 5.57 (1H, d, J = 6.6 Hz, 3 OH), 6.42 (1H, t, J = 6.6 Hz, 1 ), 7.30 (2H, d, J = 8.1 Hz, Ts), 7.55 (2H, m, Ph), 7.63 (3H, m, Ts & Ph), 8.05 (2H, d, J = 7.4 Hz, Ph), 8.54 (1H, s, 2), 8.66 (1H, s, 8), 11.24 (1H, s, NH). 5 -TsO-N2 -i-Bu-dG (S.6): yield 822 mg (73%). 1 H NMR (DMSO-d6 ): δ 1.11 (3H, d, J = 6.8 Hz, Me), d 1.12 (3H, d, J = 6.8 Hz, Me), 2.29 (1H, m, CHMe2 ), 2.35 (3H, s, Me(Ts)), 2.63 (1H, td, J = 6.5, 13.3 Hz, 2 a), 2.74 (1H, m, 2 b), 3.96 (1H, m, 4 ), 4.16 (1H, dd, J = 6.8, 10.8 Hz, 5 a), 4.24 (1H, dd, J = 3.3, 10.8 Hz, 5 b), 4.37 (1H, m, 3 ), 5.50 (1H, d, J = 6.6 Hz, 3 OH), 6.17 (1H, t, J = 6.6 Hz, 1 ), 7.32 (2H, d, J = 8.1 Hz, Ts), 7.66 (2H, d, J = 8.2 Hz, Ts), 8.06 (1H, s, 8), 11.56 (1H, s, NH), 12.04 (1H, d, J = 5.0 Hz, NH).
Convert to 5 -azido derivative 16a. For dA: Transfer 1063 mg (2.1 mmol) 5 -TsO-N6 -Bz-dA to a 50-mL round-bottom flask containing a magnetic stir bar, 514 mg (10.5 mmol, 5 eq.) LiN3 , and 11 mL DMSO. 16b. For dG: Transfer 811 mg (1.65 mmol) 5 -TsO-N2 -i-Bu-dG to a 25-mL round-bottom flask containing a magnetic stir bar, 411 mg (8.39 mmol, 5 eq.) LiN3 , and 8 mL DMSO. 17. Place the flask in a 60◦ C oil bath that is heated on top of a magnetic stir plate, and stir the reaction mixture for 5 hr while the oil bath temperature is maintained at ∼60◦ C.
-Amino-2 ,5
5 Dideoxy-5 -NTriphosphate Nucleotides
18. Concentrate the reaction mixture to ∼5 mL using a vacuum pump while heating the flask in a 50◦ C oil bath. 19. Dissolve the residual material in 100 mL of EtOAc. Wash the solution with 20 mL NaHCO3 followed by 20 mL brine using a 250-mL separatory funnel.
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Current Protocols in Nucleic Acid Chemistry
20. Transfer the EtOAc solution to a 250-mL Erlenmeyer flask containing ∼10 g of anhydrous Na2 SO4 and allow it to dry for 30 min. 21. Collect the organic solution by filtration through a fritted funnel. 22. Concentrate the EtOAc solution on a rotary evaporator to afford a pale yellow foam product. 23. Dissolve the crude product in ∼2 mL of CH2 Cl2 and apply it to a 100-mL chromatography column filled with 40 g of silica gel. Elute the column with CH2 Cl2 followed by 5:95 (v/v) MeOH/CH2 Cl2 . 24. Analyze all fractions by TLC on silica gel, developing the TLC plates with 10:90 (v/v) MeOH/CH2 Cl2 . 25. Collect the fractions containing the desired product (Rf ≈ 0.4 for dA; Rf ≈ 0.37 for dG) and concentrate to dryness. 5 -N3 -N6 -Bz-dA (S.3): yield 764 mg (92%). 1 H NMR (DMSO-d6 ): δ 2.40 (1H, ddd, J = 4.1, 6.6, 13.4 Hz, 2 a), 2.97 (1H, td, J = 6.6, 13.3 Hz, 2 b), 3.53 (1H, dd, J = 3.9, 13.1 Hz, 5 a), 3.65 (1H, dd, J = 7.0, 13.0 Hz, 5 b), 4.01 (1H, m, 4 ), 4.48 (1H, m, 3 ), 5.54 (1H, d, J = 3.7 Hz, 3 OH), 6.51 (1H, t, J = 6.7 Hz, 1 ), 7.54 (2H, m, Ph), 7.64 (1H, t, J = 7.3 Hz, Ph), 8.03 (2H, d, J = 7.5 Hz, Ph), 8.69 (1H, s, 2), 8.76 (1H, s, 8), 11.20 (1H, s, NH). 5 -N3 -N2 -i-Bu-dG (S.7): yield 419 mg (70%). 1 H NMR (DMSO-d6 ): δ 1.11 (6H, d, J = 6.8 Hz, Me), 2.31 (1H, ddd, J =3.7, 6.1, 13.3 Hz, 2 a), 2.75 (2H, m, CHMe2 & 2 b), 3.50 (1H, dd, J = 4.2, 13.1 Hz, 5 a), 3.58 (1H, dd, J = 6.8, 13.1 Hz, 5 b), 3.94 (1H, m, 4 ), 4.34 (1H, m, 3 ), 5.48 (1H, d, J = 3.7 Hz, 3 OH), 6.24 (1H, t, J = 6.8 Hz, 1 ), 8.24 (1H, s, 8), 11.64 (1H, s, NH), 12.07 (1H, d, J = 4.7 Hz, NH).
Deprotect exocyclic amine 26a. For dA: Transfer 599 mg (1.6 mmol) of 5 -N3 -N6 -Bz-dA to a 10-mL screw-capped micro-vial containing a magnetic stir bar and add 4 mL of MeOH to dissolve the solid. 26b. For dG: Transfer 409 mg (1.13 mmol) of 5 -N3 -N6 -i-Bu-dG to a 10-mL screwcapped micro-vial containing a magnetic stir bar and add 4 mL of MeOH to dissolve the solid. 27. Transfer 2 mL (half) of the resulting solution to a second 10-mL screw-capped micro-vial containing a magnetic stir bar. 28. Add 3 mL of concentrated NH4 OH to each micro-vial, place the vials in a ∼60◦ C oil bath on top of a magnetic stir plate, and stir overnight (13 to 15 hr) at ∼60◦ C. 29. Cool the vials on ice. Transfer the reaction mixtures to a round-bottom flask and concentrate to dryness using a rotary evaporator to afford a light yellow or off-white solid residue. 30. Dissolve the crude product in 60 mL of dH2 O, and extract the resulting aqueous solution five times with 5 mL CHCl3 using a separatory funnel. Collect the aqueous (top) layer and save the CHCl3 extracts. 31. Back extract the combined CHCl3 solution two times with 5 mL dH2 O using the separatory funnel. Collect the aqueous solutions.
For N3 -dA 32a. Combine all the aqueous solutions in a 250-mL round-bottom flask and freeze on dry ice. Concentrate to dryness using a lyophilizer.
Nucleoside Phosphorylation and Related Modifications
13.3.5 Current Protocols in Nucleic Acid Chemistry
Supplement 18
33a. Dissolve the crude N3 -dA product in CH2 Cl2 and purify on a chromatography column containing 40 g of silica gel. Elute the column with 2:98 to 10:90 (v/v) MeOH/CH2 Cl2 . 34a. Analyze all fractions by TLC, developing the silica gel plates with 10:90 (v/v) MeOH/CH2 Cl2 . 35a. Collect the fractions containing the desired product (Rf ≈ 0.2) and concentrate it to dryness to afford the product as light yellow foam. 5 -Azido-2 ,5 -dideoxyadenosine (N3 -dA; S.4): yield 350 mg (82%). 1 H NMR (DMSOd6 ): δ 2.30 (1H, ddd, J = 3.7, 6.4, 13.3 Hz, 2 a), 2.92 (1H, td, J = 6.7, 13.4 Hz, 2 b), 3.48 (1H, ddd, J = 3.9, 12.9 Hz, 5 a), 3.65 (1H, ddd, J = 7.3, 12.9 Hz, 5 b), 3.96 (1H, m, 4 ), 4.41 (1H, m, 3 ), 5.48 (1H, d, J = 4.0 Hz, 3 OH), 6.37 (1H, t, J = 6.9 Hz, 1 ), 7.29 (2H, s, NH2 ), 8.14 (1H, s, 2), 8.33 (1H, s, 8).
For N3 -dG 32b. Combine all the aqueous solutions in a 250-mL round-bottom flask and freeze on dry ice. Concentrate to dryness using a lyophilizer to afford the product as a white solid. Use this material to prepare NH2 -dG without further purification. 5 -Azido-2 ,5 -dideoxyguanosine (N3 -dG; S.8): yield 340 mg (100%). 1 H NMR (DMSOd6 ): δ 2.22 (1H, ddd, J = 3.2, 6.1, 13.2 Hz, 2 a), 2.70 (1H, td, J = 6.3, 13.5 Hz, 2 b), 3.46 (1H, dd, J = 4.2, 13.0 Hz, 5 a), 3.61 (1H, dd, J = 7.2, 13.0 Hz, 5 b), 3.91 (1H, m, 4 ), 4.29 (1H, m, 3 ), 5.44 (1H, br, 3 OH), 6.14 (1H, t, J = 6.9 Hz, 1 ), 6.53 (2H, brs, NH2 ), 7.90 (1H, s, 8), 9.48 (1H, brs, NH). BASIC PROTOCOL 2
PREPARATION OF 5 -AZIDO-2 ,5 -DIDEOXYPYRIMIDINES N3 -dC is prepared based on a one-step azidation procedure (Fig. 13.3.2) that utilizes a mixture of triphenylphosphine, carbon tetrabromide, and lithium azide (Yamamoto et al., 1980). This same procedure is applicable for the preparation of N3 -dT, although the latter is commercially available. An extra aqueous ammonia deprotection step is used in the synthesis of N3 -dC to remove the exocyclic amino protection that is required during the azidation procedure. CAUTION: All chemical reactions should be carried out in a fume hood to avoid toxic vapors. NOTE: In order to achieve satisfactory results, anhydrous reagents should be used and experiments should be performed under an inert atmosphere.
Materials N4 -Benzoyl-2 -deoxycytidine (N4 -Bz-dC; S.9)
5 -Amino-2 ,5 Dideoxy-5 -NTriphosphate Nucleotides
Figure 13.3.2
Preparation of 5 -azido-dC.
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Dimethylformamide (DMF), anhydrous Lithium azide (LiN3 ), concentrated to dryness from a 20% solution in dH2 O Triphenylphosphine (Ph3 P) Dry nitrogen (N2 ) or argon (Ar) Carbon tetrabromide (CBr4 ) Silica gel 60, 230 to 400 mesh Chloroform (CHCl3 ) Methanol (MeOH) Dichloromethane (CH2 Cl2 ) Pyridine, anhydrous Ammonium hydroxide (NH4 OH), concentrated aqueous solution Ethyl acetate (EtOAc) Diethyl ether (Et2 O) Dry ice 25-, 100-, and 250-mL round-bottom flasks, oven dried Rotary evaporator with dry ice/2-propanol trap, attached to a vacuum pump Vacuum pump Inert atmosphere/vacuum manifold with dry ice/2-propanol trap Tubing adaptor Balloons 250-mL glass chromatography column Thin-layer chromatography (TLC) plates, 0.25-mm silica gel 60F-254 on glass UV light source 10-mL screw-capped micro-vials (e.g., Accuform vials, Kimble) 60◦ C oil bath 100-mL separatory funnel Lyophilizer Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) Convert to 5 -azido derivative 1. Mix 603 mg (1.82 mmol) N4 -Bz-dC (S.9) and 5 mL of anhydrous DMF in a 25-mL round-bottom flask and concentrate the solution to dryness using a rotary evaporator. 2. Add a 1/2 -in. magnetic stir bar to the flask and attach the flask to a vacuum pump through a vacuum manifold to remove residual solvent. 3. Add 272 mg LiN3 (5.56 mmol, 3 eq.), 621 mg Ph3 P (2.37 mmol, 1.3 eq.), and 10 mL anhydrous DMF to the flask under a N2 or Ar atmosphere. Stir the mixture on a magnetic stir plate for a few minutes to afford a slightly cloudy solution (LiN3 does not dissolve completely). A N2 or Ar atmosphere can be provided through a tubing adaptor that connects the flask to a balloon filled with N2 or Ar, or an inert atmosphere/vacuum manifold attached to a regulated N2 or Ar tank.
4. Add 789 mg (2.34 mmol, 1.3 eq.) CBr4 to the solution and stir the mixture overnight (19 hr) at room temperature. 5. Remove DMF using a rotary evaporator to afford an orange residue. 6. Purify the residue using a 250-mL chromatography column containing 40 g of silica (APPENDIX 3E). Elute the column with CHCl3 followed by 4:100 (v/v) MeOH/CHCl3 . 7. Analyze all fractions by TLC (APPENDIX 3D) on silica gel plates. Develop TLC plates with 10:90 (v/v) MeOH/CH2 Cl2 .
Nucleoside Phosphorylation and Related Modifications
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8. Collect the fractions containing the major product (Rf ≈ 0.4) and concentrate to dryness to afford the product as a white foam. 5 -N3 -N4 -Bz-dC (S.10): yield 477 mg (74%). 1 H NMR (DMSO-d6 ): δ 2.19 (1H, td, J = 6.4, 13.2 Hz, 2 a), 2.29 (1H, ddd, J = 4.5, 6.2, 13.2 Hz, 2 b), 3.60 (1H, dd, J = 4.0, 13.2 Hz, 5 a), 3.67 (1H, dd, J = 6.4, 13.2 Hz, 5 b), 3.94 (1H, m, 4 ), 4.17 (1H, m, 3 ), 5.48 (1H, d, J = 4.3 Hz, 3 OH), 6.19 (1H, t, J = 6.4 Hz, 1 ), 7.39 (1H, brs, 5), 7.50 (2H, m, Ph), 7.61 (1H, t, J = 7.3 Hz, Ph), 7.99 (2H, d, J = 7.5 Hz, Ph), 8.18 (1H, d, J = 7.2 Hz, 6), 11.29 (1H, s, NH).
Deprotect exocyclic amine 9. Transfer 454 mg (1.27 mmol) 5 -N3 -N4 -Bz-dC to a 10-mL screw-capped micro-vial containing a magnetic stir bar. Add 5 mL pyridine and 5 mL concentrated NH4 OH. 10. Stir the resulting mixture to dissolve the solids, and then transfer 5 mL of the solution to another 10-mL screw-capped micro-vial containing a magnetic stir bar. 11. Place both vials in a ∼60◦ C oil bath heated on top of a magnetic stir plate and stir at ∼60◦ C for 8.5 hr. 12. Cool the vials on ice, transfer the reaction mixture to a 100-mL round-bottom flask, and concentrate it to dryness using a rotary evaporator. 13. To the residue, add 30 mL dH2 O and 3 mL of EtOAc, then partition the resulting mixture in a 100-mL separatory funnel. 14. Collect the aqueous solution and extract it three times with 3 mL Et2 O followed by two times with 5 mL of 1:2 (v/v) EtOAc/Et2 O. 15. Freeze the aqueous solution on dry ice and concentrate it using a lyophilizer to afford the product as a light yellow solid. 5 -Azido-2 ,5 -dideoxycytosine (N3 -dC; S.11): yield 316 mg (98%). 1 H NMR (DMSOd6 ): δ 2.07 (2H, m, 2 ), 3.51 (1H, dd, J = 4.4, 13.1 Hz, 5 a), 3.56 (1H, dd, J = 6.2, 13.1 Hz, 5 b), 3.83 (1H, m, 4 ), 4.12 (1H, m, 3 ), 5.38 (1H, d, J = 4.3 Hz, 3 OH), 5.73 (1H, d, J = 7.4 Hz, 5), 6.21 (1H, t, J = 6.8 Hz, 1 ), 7.16 (1H, brs, NH), 7.20 (1H, brs, NH), 7.60 (1H, d, J = 7.4 Hz, 6). BASIC PROTOCOL 3
PREPARATION OF 5 -AMINO-2 ,5 -DIDEOXYNUCLEOSIDES This protocol describes a common procedure useful for the preparation of all four 5 -amino-2 ,5 -dideoxynucleosides (NH2 -dNs) from corresponding 5 -azido derivatives (Fig. 13.3.3). Treatment with triphenylphosphine (Ph3 P) followed by hydrolysis with aqueous ammonia quantitatively converts 5 -azido nucleosides to 5 -amino nucleosides (Wolfe et al., 2002). The protocol uses the preparation of NH2 -dG as an example, but is also applicable for the synthesis of NH2 -dA and NH2 -dC. Specific experimental conditions for each of these analogs are listed in Table 13.3.1. NH2 -dT is commercially available and is not described here (interested readers are referred to Tetzlaff et al., 1998).
Materials
5 -Amino-2 ,5 Dideoxy-5 -NTriphosphate Nucleotides
N3 -dA, N3 -dG, and N3 -dC (see Basic Protocols 1 and 2) Pyridine, anhydrous Triphenylphosphine (Ph3 P) Dry nitrogen (N2 ) or argon (Ar) Ammonium hydroxide (NH4 OH), concentrated aqueous solution Ethyl acetate (EtOAc) 25-mL round-bottom flask Rotary evaporator with dry ice/2-propanol trap, attached to a vacuum pump
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Figure 13.3.3
Preparation of 5 -amino-dNs, 5 -amino-dNTPs, and an oligonucleotide containing a single P-N linkage.
Table 13.3.1 Synthesis Conditions Used for Each NH2 -dNa
N3 -dN (step 1)
Ph3 P/Pyr (step 3)
NH2 -dA
0.12 mmol
0.38 mmol/1 mL/7 hr 0.2 mL/15 hr RT/1 hr 55◦ C
5 mL
NH2 -dC
0.31 mmol
0.93 mmol/2 mL/7 hr 0.2 mL/15 hr RT
5 mL
NH2 -dG
0.14 mmol
NH4 OH (steps 4 to 5)
H2 O (step 6)
◦
0.43 mmol/1 mL/5 hr 0.3 mL/17 hr RT/1 hr 55 C
6 mL
a Pyr, pyridine; RT, room temperature.
Inert atmosphere/vacuum manifold with dry ice/2-propanol trap Tubing adaptor Balloon 55◦ C oil bath Fritted funnel Lyophilizer 1. Place 40 mg (0.14 mmol) N3 -dG (S.8) in a 25-mL round-bottom flask (also see Table 13.3.1). 2. Add 2 mL anhydrous pyridine and concentrate it to dryness on a rotary evaporator. Repeat. 3. Add 3 molar equiv. (111 mg, 0.43 mmol) Ph3 P, 1 mL anhydrous pyridine, and a 1/ -in. magnetic stir bar to the flask. Under a N or Ar atmosphere, stir the reaction 2 2 mixture using a magnetic stir plate 5 hr at room temperature. A N2 or Ar atmosphere can be provided through a tubing adaptor that connects the flask to a balloon filled with N2 or Ar, or an inert atmosphere/vacuum manifold attached to a regulated N2 or Ar tank.
4. Add 0.3 mL concentrated NH4 OH and stir overnight (17 hr) at room temperature. 5. Place the flask in a 55◦ C oil bath on top of a magnetic stir plate and continue to stir for 1 hr. 6. Cool the flask to room temperature and add 6 mL dH2 O. Remove the resulting precipitate using a fritted funnel and rinse with 1 mL dH2 O. 7. Collect the filtrate (aqueous solution) and extract it with 10 mL EtOAc. 8. Concentrate the resulting aqueous solution to dryness using a lyophilizer to afford a crude product which can be used in Basic Protocol 4 without further purification.
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5 -Amino-2 ,5 -dideoxyguanosine (NH2 -dG; S.12): yield 50 mg (100%) as pale yellow solid. 1 H NMR (DMSO-d6 ): δ 2.19 (1H, m, 2 a), 2.57 (1H, td, J = 6.6, 13.4 Hz, 2 b), 2.82 (2H, brs, 5 ), 3.79 (1H, brs, 4 ), 4.37 (1H, brs, 3 ), 5.76 (3H, brs, NH2 , & 3 OH), 6.11 (1H, td, J = 6.9 Hz, 1 ), 6.90 (2H, brs, NH2 ), 7.88 (1H, s, 8). 5 -Amino-2 ,5 -dideoxyadenosine (NH2 -dA; S.12): yield 63 mg (>100%) as white solid. H NMR (DMSO-d6 ): δ 2.23 (1H, ddd, J = 3.3, 6.1, 13.1 Hz, 2 a), 2.80 (3H, m, 2 b & 5 ), 3.82 (1H, m, 4 ), 4.42 (1H, m, 3 ), 5.70 (3H, br, NH2 & 3 OH), 6.32 (1H, t, J = 6.6 Hz, 1 ), 7.29 (2H, brs, NH2 ), 8.14 (1H, s, 2), 8.35 (1H, s, 8).
1
5 -Amino-2 ,5 -dideoxycytosine (NH2 -dC; S.12): yield 104 mg (>100%) as light brown foam. 1 H NMR (DMSO-d6 ): δ 1.95 (1H, td, J = 6.6, 13.4 Hz, 2 a), 2.09 (1H, ddd, J = 3.9, 5.7, 13.2 Hz, 2 b), 2.75 (2H, brs, 5 ), 3.69 (1H, m, 4 ), 4.15 (1H, m, 3 ), 5.14 (3H, br, NH2 & OH), 5.73 (1H, d, J = 7.3 Hz, 5), 6.14 (1H, t, J = 6.7 Hz, 1 ), 7.31 (2H, br, NH2 ), 7.72 (1H, d, J = 7.3 Hz, 6). Yields >100% are likely due to the incomplete removal of water.
BASIC PROTOCOL 4
PREPARATION AND HPLC ANALYSIS OF 5 -AMINO-2 ,5 -DIDEOXY-5 -N-TRIPHOSPHATE NUCLEOTIDES This protocol describes a common procedure for synthesizing NH2 -dNTPs from corresponding NH2 -dNs. As shown in Figure 13.3.3, the procedure utilizes a facile reaction between primary amines and trisodium trimetaphosphate (TMP). This straightforward approach was previously applied for the preparation of NH2 -dTTP with some success. However, the conversion yield was moderate and the product was unstable, which was likely caused by the inherently low stability of NH2 -dTTP (Letsinger et al., 1976b). By adding a commonly used, sterically hindered primary amine (Tris) to the reaction mixtures, all four NH2 -dNTPs have been prepared in dramatically improved yields (Wolfe et al., 2002). This protocol uses the preparation of NH2 -dTTP as an example, and the reaction conditions and yields for other analogs are summarized in Table 13.3.2.
Materials NH2 -dA, NH2 -dC, NH2 -dG (see Basic Protocol 3), and NH2 -dT (Sigma) Trisodium trimetaphosphate (TMP) 0.5 M aqueous Tris base HPLC buffers A and B (see recipes) 1.5-mL microcentrifuge tubes Vortex mixer C18 column (Waters Nova-pak; 3.9 × 150 mm, 4 µm) Additional reagents and equipment for HPLC (UNIT 10.5)
Table 13.3.2 Reagents and Conditions Used for the Synthesis of Each NH2 -dNTP
5 -Amino-2 ,5 Dideoxy-5 -NTriphosphate Nucleotides
Nucleosidea (step 1)
TMP (step 1)
0.5 M Tris base (step 2)
Conversion Reaction time yield by HPLC (step 3) (step 5)
NH2 -dATP
47 µmol
234 µmol
468 µL
7 days
85.6%
NH2 -dCTP
86 µmol
429 µmol
859 µL
5 days
78.9%
NH2 -dGTP
86 µmol
430 µmol
860 µL
5 days
83.2%
NH2 -dTTP
112 µmol
560 µmol
1120 µL
5 days
91.5%
a For NH -dA, NH -dC, and NH -dG, the starting material quantities are based on the assumption that N -dN was 2 2 2 3
quantitatively converted to NH2 -dN using Basic Protocol 3.
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1. Place 27.0 mg (112 µmol) of NH2 -dT and 171.4 mg (560 µmol, 5 molar equiv.) TMP in a 1.5-mL microcentrifuge tube (also see Table 13.3.2). 2. Add 1120 µL of 0.5 M aqueous Tris base (560 µmol, 5 molar equiv.) to the tube and dissolve the solids using a vortex mixer. 3. Allow the resulting solution to stand 5 days at room temperature to generate NH2 dTTP. The 100 mM NH2 -dNTP/500 mM Tris solutions (based on 100% conversion) are used directly for experiments described in Basic Protocol 5. The following steps for HPLC analysis are optional. HPLC analysis is not necessary for application described in this unit and is recommended only when the knowledge of a specific conversion yield is deemed useful. The product is stable at room temperature and can be stored for weeks without losing activity. However, to avoid water loss and slow oxidation of Tris base, the mixture should be stored at –20o C for a longer term (up to 1 year). 31
P NMR spectra were recorded on a Bruker DPX-400 NMP spectrometer using 80% H3 PO4 as an external standard. A doublet around –1 ppm was observed for all four triphosphate samples, corresponding to a phosphoramidate (Pα-N) resonance. In all spectra the most prominent peak was at –21 ppm, corresponding to excess TMP. Due to excess TMP and its breakdown products, Pβ and Pγ were not assigned. 31 P NMR (D2 O): NH2 -dATP, –0.99 ppm (d, J = 20.8 Hz); NH2 -dCTP, –0.94 ppm (d, J = 19.5 Hz); NH2 -dGTP, –1.00 ppm (d, J = 20.8 Hz); NH2 -dTTP, –0.97 ppm (d, J = 20.1 Hz).
4. Monitor the reaction mixture using an HPLC system equipped with a C18 column (UNIT 10.5). Elute the column with a mobile phase of HPLC buffers A and B at a flow rate of 1.00 mL/min. Vary the buffer gradient from 98% A to 100% B to achieve separation of NH2 -dT from NH2 -dTTP: 0 to 8 min 8 to 9 min 9 to 10 min 10 to 11 min 11 to 15 min
98% A to 70% A 70% A to 0% A 0% A 0% A to 98% A 98%A.
5. Compare peak areas corresponding to NH2 -dT (retention time ≈ 4.0 min) and NH2 dTTP (retention time ≈ 10.7 min) in the HPLC chromatograms to provide the yield of conversion (91.5%). Retention times for NH2 -dA and NH2 -dATP: ≈5.5 min and ≈14 min, respectively; for NH2 -dC and NH2 -dCTP: ≈2.0 min and ≈5.0 min, respectively; for NH2 -dG and NH2 dGTP ≈3.0 min and ≈10.5 min, respectively.
POLYMERASE INCORPORATION OF NH2 -dNTPS This protocol describes procedures for incorporating each NH2 -dNTP into DNA through a DNA polymerase–catalyzed template-directed primer extension reaction. The procedure is similar to routine primer extension reactions, with the exceptions that NH2 -dNTP is used to replace or supplement its naturally occurring counterpart to afford complete or statistical analog substitution in the DNA product, respectively, and that an alkaline pH 9.5 buffer is used to improve the NH2 -dNTP stability. Among the many commercially available polymerases tested, Klenow polymerase (exo– ) provides the most efficient NH2 dNTP incorporation. In this protocol, NH2 -dNTPs are used in higher concentrations than those of native dNTPs to compensate for their reduced incorporation efficiency compared to dNTPs. Mild acid treatment of the substituted DNA product generates fragments that correspond to site-specific cleavage at each modified nucleotide, which provides direct evidence of successful NH2 -dNTP incorporation as well as the sequence of the original DNA template (Wolfe et al., 2002).
BASIC PROTOCOL 5
Nucleoside Phosphorylation and Related Modifications
13.3.11 Current Protocols in Nucleic Acid Chemistry
Supplement 18
Materials 20 µM oligonucleotide primer 1.67 µM [γ-32 P]ATP (6000 Ci/mmol; Perkin-Elmer) 10 U/µL T4 polynucleotide kinase and 10× buffer (New England Biolabs) TE buffer, pH 8.0 (APPENDIX 2A) Single-stranded DNA templates: ∼100 or 500 nucleotides in length (for complete or partial NH2 -dNTP substitution, respectively) 200 mM MgCl2 500 mM sodium acetate (NaOAc) 10× polymerase extension buffers A and B (see recipes) 100 mM NH2 -dNTP solution(s) in 0.5 M Tris (see Basic Protocol 4) 2 -Deoxynucleoside triphosphates (dATP, dCTP, dGTP, dTTP) 5 U/µL Klenow fragment of DNA polymerase (exo– ; New England Biolabs) 1% and 10% (v/v) acetic acid Formamide loading buffer (Life Technologies) Denaturing polyacrylamide gel mix and buffer solutions (National Diagnostics) 1× TBE buffer (APPENDIX 2A) 10 mM EDTA, pH 8 (APPENDIX 2A) 0.5-mL microcentrifuge tubes Heat block at 37◦ and 90◦ C Sephadex G-50 columns, preswollen in dH2 O Speedvac evaporator Additional reagents and equipment for polyacrylamide gel electrophoresis (PAGE; UNIT 10.4) Label and anneal oligonucleotide primer 1. Label a primer using T4 polynucleotide kinase and [γ-32 P]ATP. In a 0.5-mL microcentrifuge tube mix the following: 1 µL 20 µM oligonucleotide primer 1 µL 1.67 µM [γ-32 P]ATP (6000 Ci/mmol) 1 µL 10× T4 polynucleotide kinase buffer 6 µL dH2 O 1 µL 10 U/µL T4 polynucleotide kinase. 2. Incubate the mixture for 45 to 60 min at 37◦ C. 3. Add 10 µL TE buffer and purify the solution using a Sephadex G-50 column to afford an ∼1 µM primer solution (assuming 100% recovery).
Perform primer extension with complete NH2 -dNTP substitution 4a. In a 0.5-mL microcentrifuge tube mix the following: 2 µL ∼1 µM labeled primer 10 µL ∼0.24 µM ssDNA template (∼100 nt in length) 2 µL 200 mM MgCl2 2 µL 500 mM of NaOAc 4 µL dH2 O.
5 -Amino-2 ,5 Dideoxy-5 -NTriphosphate Nucleotides
5a. Heat the solution in a heat block 2 min at 90◦ C, then allow the heat block to cool down to room temperature to afford a solution containing ∼0.1 M DNA duplex in 20 mM MgCl2 /50 mM NaOAc.
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6a. In a 0.5-mL microcentrifuge tube mix the following:
1 µL annealed DNA duplex solution 1 µL 10× polymerase extension buffer A 1 µL 100 mM DTT 0.4 µL 100 mM NH2 -dATP/0.5 M Tris (or other NH2 -dNTP) 1 µL 1 mM each dCTP/dGTP/dTTP (or any 3 dNTPs that complement the NH2 -dNTP) 4.6 µL dH2 O 1 µL 5 U/µL Klenow (exo– ) DNA polymerase. The resulting mixture contains 45 mM Tris·Cl, pH 9.5, 10 mM DTT, 22 mM MgCl2 , 5 mM NaOAc, 4 mM NH2 -dNTP, 0.1 mM each of the other three dNTPs, and 5 U Klenow (exo– ) polymerase.
7a. Incubate 1 hr at 37◦ C. 8a. Add 30 µL TE buffer and purify using Sephadex G-50 columns to afford ∼50 µL of modified DNA solution. Store up to 2 weeks at −20◦ C. Prepare G-50 columns using G-50 resin preswollen in dH2 O. G-50 resin that has been pre-equilibrated in Tris buffers may alter the amount of acetic acid needed in step 9a.
Perform site-specific cleavage at NH2 -dN for complete substitution 9a. In a 0.5-mL microcentrifuge tube mix 8 µL of each modified DNA solution with 2 µL of 1% acetic acid and incubate 30 min at 37◦ to 40◦ C. 10a. Add 100 µL dH2 O to each sample, and concentrate the resulting solution to dryness on a Speedvac evaporator. 11a. Dissolve each sample in dH2 O. Adjust volume depending on the radioactivity of the samples. 12a. Transfer 2 µL of each solution to another microcentrifuge tube and add 2 µL formamide loading buffer. 13a. Analyze the samples by electrophoresing on a 12% denaturing polyacrylamide gel using standard procedures (UNIT 10.4) and 1× TBE buffer.
Perform primer extension with partial (statistical) NH2 -dNTP substitution 4b. Anneal labeled primer to a ssDNA template as in steps 4a and 5a, but use a template that is ∼500 nt in length. 5b. In a 0.5-mL microcentrifuge tube mix the following:
1 µL annealed DNA duplex solution 1 µL 10× polymerase extension buffer B 0.4 µL 100 mM NH2 -dATP/0.5 M Tris (or other NH2 -dNTP) 4 µL 0.1 mM dATP (or 2.6 µL dCTP, 1.6 µL dGTP, 4.6 µL dTTP) 2 µL 2 mM each dCTP/dGTP/dTTP (or any 3 dNTPs that complement the NH2 -dNTP) dH2 O to 9 µL 1 µL 5 U/µL Klenow (exo– ) DNA polymerase. The resulting mixture contains 50 mM Tris·Cl, pH 9.5, 5 mM DTT, 22 mM MgCl2 , 5 mM NaOAc, 4 mM NH2 -dNTP, 0.016 to 0.046 mM of the corresponding dNTP (0.04 mM dATP, 0.026 mM dCTP, 0.016 mM dGTP, 0.046 mM TTP), 0.4 mM each of the other three dNTPs, and 5 U of Klenow (exo– ) polymerase.
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13.3.13 Current Protocols in Nucleic Acid Chemistry
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6b. Incubate 1 hr at 37◦ C. 7b. Add 1 µL of 1 mM (each) dATP/dCTP/dGTP/dTTP and incubate for an additional 15 min at 37◦ C. 8b. Add 20 µL TE buffer and store the resulting solutions up to 2 weeks at −20◦ C.
Perform site-specific cleavage at NH2 -dN for partial substitution 9b. In a 0.5-mL microcentrifuge tube mix the following: 2 µL modified DNA solution 1 µL 10 mM EDTA, pH 8 6 µL dH2 O 1 µL 10% acetic acid. 10b. Incubate 10 min at 37◦ C. 11b. Proceed with analyis as in steps 10a to 13a.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
HPLC buffer A Add 10 mL of 1 M Tris·Cl, pH 9 (APPENDIX 2A), and 50 mL of 2 M TEAA buffer (see recipe) to ∼900 mL dH2 O in a 1-L graduated cylinder. Add trace amounts of triethylamine and/or acetic acid to adjust the pH to 9 (measure using pH paper). Bring volume to 1 L with dH2 O (final 10 mM Tris/100 mM TEAA). Store up to several weeks at room temperature. HPLC buffer B Mix 750 mL HPLC buffer A (see recipe) and 250 mL MeOH. Store up to several weeks at room temperature. Polymerase extension buffer A, 10× 0.25 M Tris·Cl, pH 9.5 0.2 M MgCl2 Store up to several months at −20◦ C Polymerase extension buffer B, 10× 0.3 M Tris·Cl, pH 9.5 0.2 M MgCl2 50 mM dithiothreitol (DTT) Store up to several months at −20◦ C TEAA (triethylammonium acetate) buffer, 2 M Chill triethylamine and acetic acid on ice. In a fume hood, mix 278.8 mL of triethylamine and 114.4 mL of acetic acid in a 1-L graduated cylinder. Add dH2 O to 1 L. Store up to several months at room temperature. 5 -Amino-2 ,5 Dideoxy-5 -NTriphosphate Nucleotides
13.3.14 Supplement 18
Current Protocols in Nucleic Acid Chemistry
COMMENTARY Background Information Due to their notably increased chemical reactivity and limited structural deviation from naturally occurring nucleosides, 5 amino-2 ,5 -dideoxy nucleosides (NH2 -dNs) and nucleotides (NH2 -dNTPs) have been exploited for various applications. For example, derivatives of NH2 -dT have been used for model studies on DNA replication (Luo et al., 1998), template-directed chemical amplification (Zhan and Lynn, 1997), mechanistic studies on polymerases and reverse transcriptases (Lutz et al., 1997), and the construction of combinatorial peptide-DNA hybrids (Bergmann and Bannwarth, 1995) and their libraries (Koppitz et al., 1998; Tetzlaff et al., 1998). In addition, NH2 -dTTP and NH2 dCTP have been used to incorporate acid-labile phosphoramidate (P-N) linkers into DNA for high-throughput detection of single nucleotide polymorphisms using matrix-assisted fragmentation (Shchepinov et al., 2001), while NH2 -dTTP has been applied in a dinucleotide DNA cleavage method developed for discovery of single nucleotide polymorphisms (Wolfe et al., 2003). Most of the aforementioned applications have been demonstrated utilizing NH2 -dT or its triphosphate NH2 -dTTP, perhaps because NH2 -dT is the only commercially available analog among the commonly used nucleosides and nucleotides. The ready availability of all four NH2 -dNs and NH2 -dNTPs will help expand the use of this class of compounds. The synthetic strategy described here takes advantage of common chemical features of all NH2 dNTPs and utilizes synthetic methods including tosylation (also described in UNIT 13.2), azide exchange, and the Staudinger reaction. Because this approach does not require special equipment such as a Parr hydrogenation apparatus, these compounds can be conveniently made in many minimally equipped chemistry laboratories. Besides polymerase incorporation of NH2 dNTPs, P-N linkers can also be introduced into DNA through solid-phase oligonucleotide synthesis using 5 -NH2 -modified phosphoramidate building blocks (Mag and Engels, 1989; Shchepinov et al., 2001). This approach provides an opportunity to place P-N linkers at any specific position within an oligonucleotide, which is of importance for applications where predetermined localization of P-N bonds is desirable. However, solid-phase synthesis is not compatible with applications
where limited DNA sequence information is available, as is the case for DNA sequence or polymorphism discovery (Wolfe et al., 2002, 2003). In addition, due to the lower efficiency of chemical coupling relative to enzymatic elongation, oligonucleotides generated by solid-phase synthesis have a very short length limit (up to ∼150 bases) in comparison to replication reactions catalyzed by polymerases (up to several kilobases). For example, the authors have generated a 7.2-kb long NH2 dT-containing polymerization product using Klenow (exo− ) polymerase and the M13 plasmid as template DNA (Wolfe et al., 2002). Furthermore, solid-phase synthesis requires access to an automated DNA synthesizer, as well as 5 -NH2 -modified phosphoramidate building blocks that are not available commercially.
Critical Parameters and Troubleshooting For chemical reactions described in Basic Protocols 1 through 3, it is critical to avoid moisture in all reaction mixtures. In the authors’ experience, sealed bottles of anhydrous solvents obtained directly from vendors are sufficiently dry for these reactions. Starting material (nucleosides) should be routinely coevaporated with anhydrous solvents followed by vacuum drying before use. In Basic Protocol 4, using a freshly prepared 0.5 M Tris base solution will ensure sufficient buffering capacity. The NH2 -dNTP products should be stored and used as prepared; isolation or dilution is not recommended because they may reduce the amount of 5 -N-triphosphate nucleotides (Letsinger et al., 1976b). Although the concentration of the NH2 -dNTP from individual attempted reactions may vary slightly, the overall reproducibility of this reaction is very high. HPLC analysis can provide information on the actual amount of the desired product, but is not required. In Basic Protocol 5, the authors routinely use Klenow (exo– ) polymerase from New England Biolabs. Polymerases from other vendors may also be applicable, but reaction conditions may need to be optimized since other polymerase products may have different enzymatic activities. Because the purity and concentration of each nucleotide may affect its polymerase incorporation efficiency, slight modifications of the dNTP/NH2 -dNTP ratios may be necessary for statistical incorporation
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experiments to achieve even incorporation of NH2 -dNTPs. The efficiency of acid cleavage reactions may be very sensitive to the amount of Tris in each DNA sample and the actual concentration of acetic acid used. Minor adjustments in acid concentration and/or reaction time may be necessary to accommodate these variations.
Since HPLC analysis of NH2 -dNTPs requires instrumentation and is time consuming, users may choose to skip this procedure. In the authors’ experience, the preparation of NH2 dNTPs from NH2 -dNs has been very reproducible.
Literature Cited Anticipated Results For Basic Protocols 1 through 3, the yields of chemical reactions are generally very high, although the recovered yield of purified products may be variable and considerably lower, depending on the scale or the method of purification. Basic Protocol 4 is highly reproducible and the reaction mixtures can be directly used for downstream enzymatic reactions. Polymerase incorporation of one of the NH2 -dNTPs as complete replacement of its naturally occurring counterpart is very reproducible, as long as the Klenow (exo− ) polymerase is fresh and relatively short DNA templates are used. Acid cleavage of these products may need to be optimized to achieve even cleavage bands. Statistical incorporation of NH2 -dNTPs and subsequent acid cleavage provide a sequencing ladder from long DNA templates. Using harsher cleavage conditions than described (i.e., higher concentration of acetic acid, longer reaction time, or higher temperature) will enrich shorter DNA fragments. Insufficient cleavage will enrich longer DNA fragments and the full-length extension product.
Time Considerations
5 -Amino-2 ,5 Dideoxy-5 -NTriphosphate Nucleotides
The approximate time scale for the sequence of reactions and procedures described in this unit is 1 to 2 days for tosylation; 2 to 3 days for azidation and deprotection; 1 day for Ph3 P reduction and hydrolysis; 5 to 7 days for triphosphate formation (mostly hands-off time); 2 days for annealing of primer to template, followed by polymerase incorporation of NH2 dNTPs and subsequent chemical cleavage reactions; and 1 to 2 days for PAGE analysis of the products. A major uncertainty in time consumption is related to column chromatography. The optimum solvent composition may depend on column diameter, quality of silica gel, purity of eluting solvents, and rate of elution. If removal of impurity is not completely successful in the first attempt, another column chromatography purification may be necessary.
Bergmann, F. and Bannwarth, W. 1995. Solidphase synthesis of directly linked peptideoligodeoxynucleotide hybrids using standard synthesis protocols. Tetrahedron Lett. 36:18391842. Koppitz, M., Nielsen, P.E., and Orgel, L.E. 1998. Formation of oligonucleotide-PNA-chimeras by template-directed ligation. J. Am. Chem. Soc. 120:4563-4569. Letsinger, R.L., Hapke, B., Petersen, G.R., and Dumas, L.B. 1976a. Enzymatic synthesis of duplex circular phiX174 DNA containing phosphoramidate bonds in the (−) strand. Nucl. Acids Res. 3:1053-1063. Letsinger, R.L., Wilkes, J.S., and Dumas, L.B. 1976b. Incorporation of 5 -amino-5 deoxythymidine-5 -phosphate in polynucleotides by use of DNA polymerase I and a phiX174 DNA template. Biochemistry 15:2810-2816. Luo, P.Z., Leitzel, J.C., Zhan, Z.Y.J., and Lynn, D.G. 1998. Analysis of the structure and stability of a backbone-modified oligonucleotide: Implications for avoiding product inhibition in catalytic template-directed synthesis. J. Am. Chem. Soc. 120:3019-3031. Lutz, M.J., Benner, S.A., Hein, S., Breipohl, G., and Uhlmann, E. 1997. Recognition of uncharged polyamide-linked nucleic acid analogs by DNA polymerases and reverse transcriptases. J. Am. Chem. Soc. 119:3177-3178. Mag, M. and Engels, J.W. 1989. Synthesis and selective cleavage of oligodeoxyribonucleotides containing non-chiral internucleotide phosphoramidate linkages. Nucl. Acids Res. 17:59735988. Mungal, W.S., Greene, G.L., Heavner, G.A., and Letsinger, R.L. 1975. Use of the azido group in the synthesis of 5 -terminal aminodeoxythymidine oligonucleotides. J. Org. Chem. 40:16591662. Shchepinov, M.S., Denissenko, M.F., Smylie, K.J., Worl, R.J., Leppin, A.L., Cantor, C.R., and Rodi, C.P. 2001. Matrix-induced fragmentation of P3 -N5 phosphoramidate-containing DNA: High-throughput MALDI-TOF analysis of genomic sequence polymorphisms. Nucl. Acids Res. 29:3864-3872. Tetzlaff, C.N., Schwope, I., Bleczinski, C.F., Steinberg, J.A., and Richert, C. 1998. A convenient synthesis of 5 -amino-5 -deoxythymidine and preparation of peptide-DNA hybrids. Tetrahedron Lett. 39:4215-4218. Wolfe, J.L., Kawate, T., Belenky, A., and Stanton, V. Jr. 2002. Synthesis and polymerase
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incorporation of 5 -amino-2 ,5 -dideoxy-5 -Ntriphosphate nucleotides. Nucl. Acids Res. 30:3739-3747. Wolfe, J.L., Wang, B., Kawate, T., and Stanton, V.P. Jr. 2003. Sequence-specific dinucleotide cleavage promoted by synergistic interactions between neighboring modified nucleotides in DNA. J. Am. Chem. Soc. 125:10500-10501. Yamamoto, I., Sekine, M., and Hata, T. 1980. Onestep synthesis of 5 -azido-nucleosides. J. Chem. Soc. Perkin I 306-310. Zhan, Z.Y.J. and Lynn, D.G. 1997. Chemical amplification through template-directed synthesis. J. Am. Chem. Soc. 119:12420-12421.
Contributed by Jia Liu Wolfe and Tomohiko Kawate Massachusetts General Hospital Cambridge, Massachusetts
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Nucleoside-5 -Phosphoimidazolides: Reagents for Facile Synthesis of Dinucleoside Pyrophosphates
UNIT 13.4
Natural nucleoside pyrophosphates, such as NAD, NADH, FAD, FADH2 , NADP, and NADPH (Fig. 13.4.1), play a critical role as enzyme co-factors in biochemical redox reactions. Analogs of these nucleoside pyrophosphates that do not act in redox processes serve as valuable tools for investigating the mechanisms of these enzymes. For example, tiazofurin adenine dinucleotide (TAD), an NAD analog that does not participate in hydride transfer, is a potent inhibitor of inosine monophosphate dehydrogenase (IMPDH). TAD and similar dinucleoside pyrophosphates such as benzamide adenine dinucleotide (BAD) have been used to generate X-ray crystallographic data of IMPDH complexes and have been useful tools in elucidation of the mechanism of action of IMPDH (Li et al., 1994a,b; Pankiewicz et al., 2002a,b, 2004; Hedstrom et al., 2003). This unit contains a basic procedure for synthesis of dinucleoside pyrophosphates from a nucleoside 5 phosphoimidazolide and a nucleoside 5 -monophosphate. The preparation of nucleoside 5 -phosphoimidazolides from nucleoside 5 -phosphates is described in the first method (see Basic Protocol 1). When free nucleoside 5 monophophates are not available, nucleoside 5 -monophosphates protected at the 2 and 3 positions can be used (Alternate Protocol). The Alternate Protocol can also be used to prepare nucleoside 5 -phosphoimidazolides that do not have a cis-1,2-diol moiety, such as 2 -deoxyriboside-5 -monophosphoimidazolides. The use of phosphoimidazolides in the synthesis of dinucleoside pyrophosphates is described in Basic Protocol 2. The resulting pyrophosphates are stable to conditions used in removal of standard protecting groups such as 2 ,3 -isopropylidene (acetonide) and 2 ,3 -acyl protecting groups. NOTE: Although NAD and related compounds are dinucleotides because of the presence of a phosphate group (or rather pyrophosphate), dinucleoside pyrophosphate is used here as a general term for two nucleosides connected by a pyrophosphate. CAUTION: Carry out all operations involving organic solvents and reagents in a wellventilated fume hood. Wear appropriate protective clothing and glasses.
PREPARATION OF NUCLEOSIDE 5 -PHOSPHOIMIDAZOLIDES FROM NUCLEOSIDE 5 -MONOPHOSPHATES
BASIC PROTOCOL 1
Nucleoside 5 -phosphoimidazolides have been used previously to prepare dinucleoside pyrophosphates (Cramer and Neunhoeffer, 1962; Scheit, 1980; Gebeyehu et al., 1985; Zatorski et al., 1993, 1995, 1996). The protocol described here is based on the method of Cramer and Neunhoeffer. Many nucleoside 5 -monophosphates are commercially available from suppliers such as Sigma-Aldrich and Berry & Associates. Those that are not commercially available can be synthesized by the Yoshikawa reaction (Yoshikawa et al., 1967). RP-HPLC purification can be performed on any HPLC instrument.
Materials Nucleoside 5 -monophosphate (commercially available or by Yoshikawa reaction; Yoshikawa et al., 1967) Anhydrous N,N-dimethylformamide (DMF; Aldrich)
Nucleoside Phosphorylation and Related Modifications
Contributed by Liqiang Chen, Dominik Rejman, Laurent Bonnac, Krzysztof W. Pankiewicz, and Steven E. Patterson
13.4.1
Current Protocols in Nucleic Acid Chemistry (2005) 13.4.1-13.4.10 C 2005 by John Wiley & Sons, Inc. Copyright
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Figure 13.4.1
Natural and synthetic dinucleoside pyrophosphates.
Anhydrous N,N-dimethylformamide-d7 (DMFd7 ; Aldrich) 1,1 -Carbonyldiimidazole (CDI), 95% pure 0.1 M triethylammonium bicarbonate (TEAB), pH 7.3 (see recipe), or 0.1 M triethylamine (TEA), pH 9.0 0.04 M TEAB, pH 7.3 (see recipe) Acetonitrile (CH3 CN), HPLC grade Water (deionized; resistivity > 18 Mcm) NMR spectrometer and 5-mm NMR tubes HPLC system with: C18 HPLC column (Varian microsorb; 250 × 41.4–mm; 8-µm particle size, ◦ 100 A pore size) UV/Vis detector (260 nm) Rotary evaporator equipped with a vacuum pump Lyophilizer Additional reagents and equipment for NMR and RP-HPLC 1. Prepare a 0.1 M solution of the nucleoside 5 -monophosphate in anhydrous DMF or DMFd7 . Synthesis of Dinucleoside Pyrophosphates
To monitor the reaction by NMR, the reaction mixture should contain sufficient DMFd7 to obtain a deuterium lock in the NMR spectrometer (10% DMFd7 in DMF is sufficient).
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Figure 13.4.2
Preparation of nucleoside 5 -phosphoimidazoles.
2. Add 4 equiv CDI and stir at room temperature. 3. Perform the reaction in an NMR tube and monitor by proton-decoupled 31 P NMR as illustrated by the reaction of adenosine 5 -monophosphate with CDI (Figs. 13.4.2 and 13.4.3). If the total reaction volume is between 0.7 and 1.0 mL, perform the entire reaction in a 5-mm NMR tube. For larger reaction volumes, to monitor the reaction by NMR, place a 0.7-mL aliquot in a 5-mm NMR tube. a. The proton-decoupled 31 P NMR spectrum of the parent monophosphate has a single resonance at δ +1.75. b. About 2 min after addition of CDI at room temperature, the 31 P NMR spectrum (Fig. 13.4.3A) will have a singlet upfield from the monophosphate due to formation of intermediate S.1 (δ −5.71). c. After approximately 20 min (Fig. 13.4.3B), two additional singlets appear in the spectrum, corresponding to the carbonate S.2 (δ −6.10) and AMP-imid S.3 (δ −7.56). d. After 2.5 to 3 hr (Fig. 13.4.3D), the spectrum will be simplified to a single resonance at δ −7.76 (S.4), signifying completion of the reaction.
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Figure 13.4.3 31 P NMR spectra of the reaction of AMP with CDI. Reprinted from Zatorski et al. (1995) with permission from the American Chemical Society.
4. Hydrolyze the resulting nucleoside-2 ,3 -carbonate-5 -imidazolide (S.4) to the desired nucleoside 5 -imidazolide (S.3) by adding 1 mL of 0.1 M TEAB, pH 7.3, or 1 mL of 0.1 M TEA, pH 9.0 per mL of reaction mixture, and stirring for 2 hr at room temperature. TEAB is more readily removed by rotary evaporation than triethylammonium hydrochloride and other commonly used ion-pairing agents that require ion exchange. Many C18 columns are also unstable at the higher pH of triethylammonium hydroxide.
5. Purify the resulting mixture by RP-HPLC using the following conditions:
Solvent A: 0.04 M TEAB, pH 7.3 Solvent B: 70% acetonitrile/water Gradient: isocratic 15% B Flow rate: 40 mL/min UV detection: 260 nm ◦ C18 column: Varian microsorb, 250 × 41.4 mm, 8-µm particle size, 100 A pore size. Synthesis of Dinucleoside Pyrophosphates
Adenosine 5 -phosphoimidazolide: TR = 6.7 min.
6. Combine fractions containing the desired imidazolide and concentrate on a rotary evaporator.
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7. Add 2 vol water to the residue and remove triethylamine by co-evaporation. Dissolve in another 3 vol water and remove water on a rotary evaporator. Finally, dissolve in 2 vol water and lyophilize to give the imidazolide as a solid triethylammonium salt. After lyophilization, the resulting solid nucleoside 5 -phosphoimidazolide salts can be stored for 1 week at −20◦ C. These imidazolides slowly decompose to their parent nucleoside 5 -monophosphates and imidazole in the presence of water. At ambient temperature, 5 mM solutions of imidazolides in water at neutral pH have a half-life of ∼4 days.
8. Characterize by 1 H NMR and 31 P NMR. Adenosine 5 -phosphoimidazolide (S.3): yield 82%.31 P NMR (DMSO-d6 ): singlet at δ −7.56. 1 H NMR (DMSO-d6 ): δ 1.00 (t, 9H), 2.70 (q, 6H), 3.60-3.76 (m, 2H), 3.89-3.91 (m, 1H), 3.99-4.06 (m, 1H), 4.57 (pseudo t, 1H), 5.88 (d, J = 6.1 Hz, 1H), 6.85 (s, 1H), 7.10 (s, 1H) 7.65 (s, 1H), 7.25 (s, 2H), 8.12 (s, 1H), 8.40 (s, 1H).
PREPARATION OF NUCLEOSIDE 5 -PHOSPHOIMIDAZOLIDES FROM 2 ,3 -PROTECTED NUCLEOSIDE-5 -MONOPHOSPHATES
ALTERNATE PROTOCOL
While the above procedure provides a general method for unprotected nucleoside 5 monophosphates, those that do not have free 2 - and 3 -hydroxyls will not give the 2 ,3 -carbonates (S.2 and S.4). This alternate procedure is used in such cases. It is compatible with deoxynucleoside-5 -monophosphates, 2 ,3 -isopropylidenenucleoside5 -monophosphates, acetyl-protected monophosphates, and silyl-protected monophosphates. 1. Prepare a 0.1 M solution of the nucleoside 5 -monophosphate in anhydrous DMF or DMFd7 . To monitor the reaction by NMR, the reaction mixture should contain sufficient DMFd7 to obtain a deuterium lock in the NMR spectrometer (10% DMFd7 in DMF is sufficient).
2. Add 1.25 equiv CDI and stir at room temperature. 3. Monitor the reaction by 31 P NMR (see Basic Protocol 1, step 3). Using the reaction of 2 ,3 -isopropylidenetiazofurin with CDI as an example, the spectrum (not shown) reveals that: a. The resonance of the parent phosphate (δ 1.63) disappears within 5 min and a new resonance emerges at δ −6.53. This intermediate corresponds to intermediate S.1 in Figure 13.4.2. b. The strength of the resonance at δ −6.53 diminishes over time with simultaneous increase in strength of a new signal at δ −7.79. This corresponds to the desired 5 -imidazolide, with 2 - and 3 -hydroxyl protection still intact. c. After 45 min, only the resonance at δ −7.79 is detected in the spectrum, signifying completion of the reaction. 4. Purify the resulting mixture by RP-HPLC using the following conditions:
Solvent A: 0.04 M TEAB, pH 7.3 Solvent B: 70% acetonitrile/water. Gradient: isocratic 27% B Flow rate: 40 mL/min UV detection: 260 nm ◦ C18 column: Varian microsorb, 250 × 41.4 mm, 8-µm particle size, 100 A pore size. 2 ,3 -Isopropylidenetiazofurin-5 -phosphoimidazolide: TR = 7.8 min.
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5. Combine fractions containing the desired imidazolide and concentrate on a rotary evaporator. 6. Add 2 vol water to the residue and remove triethylamine by co-evaporation. Dissolve in another 3 vol water and remove water on a rotary evaporator. Finally, dissolve in 2 vol water and lyophilize to give the imidazolide as a solid triethylammonium salt. 7. Characterize by 1 H NMR and 31 P NMR. 2 ,3 -Isopropylidenetiazofurin-5 -phosphoimidazolide: yield 79%. 31 P NMR (D2 O): singlet at δ −7.80. 1 H NMR (D2 O): δ 1.07 (t, 9H), 1.42 (s, 3H), 1.61 (s, 3H), 2.68 (q, 6H), 3.92-3.97 (m, 2H), 4.42-4.57 (m, 1H), 4.79-4.81 (m, 1H), 5.18 (dd, J = 3 .4 Hz, 6.0 Hz, 1H), 5.33 (d, 1H), 7.03 (s, 1H), 7.11 (s, 1H), 7.75 (s, 1H) 8.17 (s, 1H). BASIC PROTOCOL 2
PREPARATION OF DINUCLEOSIDE PYROPHOSPHATES The protocol described here is a modification of the method of Cramer and Neunhoeffer (1962). The modifications in this procedure include the use of triethylammonium salts of phosphates and imidazolides in place of trioctylammonium salts, the omission of pyridine from the reaction mixture, and the use of preparative RP-HPLC to purify and isolate the products. The reaction time in this procedure is ∼3 hr, compared to ∼4 days in the Cramer and Neunhoffer method. Many nucleoside 5 -monophosphates are commercially available from suppliers such as Sigma-Aldrich and Berry & Associates. Those that are not commercially available can be synthesized by the Yoshikawa reaction (Yoshikawa et al., 1967). Because nucleoside 5 -phosphoimidazolides slowly decompose to the parent phosphate and imidazole in the presence of water (see Basic Protocol 1, step 7), it is best that the imidazolides be used soon after preparation. If necessary, imidazolides may be stored at −20◦ C after lyophilization, but care should be used to exclude moisture from the imidazolides. Deprotection is performed by standard methods after formation of the pyrophosphate. The phosphoimidazolide is not stable to deprotection methods used to remove the isopropylidene group or to remove acyl protecting groups. To ensure good yields, anhydrous DMF should be used in reactions. It can be purchased from a commercial supplier (e.g., Aldrich) and used without further purification. RP-HPLC purification can be performed on any HPLC instrument.
Materials Nucleoside 5 -phosphoimidazolide (see Basic Protocol 1 or Alternate Protocol) Anhydrous N,N-dimethylformamide (DMF; Aldrich) Anhydrous N,N-dimethylformamide-d7 (DMFd7 ; Aldrich) Nucleoside 5 -monophosphate (commercially available or by Yoshikawa reaction; Yoshikawa et al., 1967) 0.04 M triethylammonium bicarbonate (TEAB), pH 7.3 (see recipe) Acetonitrile (CH3 CN), HPLC grade Water (deionized; resistivity > 18 Mcm) Dowex 50WX8-200 H+ and Na+ forms
Synthesis of Dinucleoside Pyrophosphates
NMR spectrometer and 5-mm NMR tubes Rotary evaporator Lyophilizer HPLC system with: C18 HPLC column (Varian microsorb, 250 × 41.4 mm, 8-µm particle size, ◦ 100 A pore size) UV/Vis detector (260 nm) Additional reagents and equipment for NMR, RP-HPLC, and ion-exchange chromatography
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1. Prepare a 0.1 M solution of the nucleoside 5 -phosphoimidazolide in anhydrous DMF or DMFd7 . 2. Add 1.2 equiv of a nucleoside 5 -monophosphate and stir at room temperature. 3. Monitor the reaction by 31 P NMR (see Basic Protocol 1, step 3). Disappearance of the resonance due to the imidazolide and appearance of two signals at ∼10 ppm indicates the reaction is complete. In some cases the resonances of the two phosphorous atoms are not resolved, which gives a single resonance at ∼10 ppm. Typical reaction times are from 1 to 3 days.
4. Remove the solvent at reduced pressure either on a rotary evaporator or by lyophilization. 5. Dissolve the resulting residue in a minimum volume 0.4 M TEAB and purify by HPLC as illustrated for the purification of P1 -(2,6-diaminopurine-2 ,3 isopropyliden-5 -yl)-P2 -(tiazofurin-2 ,3 -isopropyliden-5 -yl) pyrophosphate:
Solvent A: 0.04 M TEAB, pH 7.3 Solvent B: 70% (v/v) acetonitrile/water. Gradient: isocratic 25% B Flow rate: 40 mL/min UV detection: 260 nm ◦ C18 column: Varian microsorb, 250 × 41.4 mm, 8-µm particle size, 100 A pore size. For pyrophosphates containing more lipophilic substituents, the amount of solvent B in the mobile phase may need to be increased. Conversely, for pyrophosphates containing substituents that are less lipophilic, the amount of solvent A in the mobile phase may need to be increased. In addition, the detector wavelength may need to be altered depending on the absorbance spectrum of the product.
6. Combine the fractions containing the desired pyrophosphate and concentrate on a rotary evaporator. 7. Add 2 vol water to the residue and remove triethylamine by co-evaporation. Dissolve in another 3 vol water and remove water on a rotary evaporator. Finally, dissolve in 2 vol water and lyophilize to give the pyrophosphate as a solid triethylammonium salt. 8. Characterize by 1 H NMR and 31 P MNR. P1 -(2 ,3 -Isopropylidenetiazofurin-5 -yl)-P2 -(2 ,3 -isopropylidene-2-aminoadenin-5 -yl) pyrophosphate: yield 67% relative to the phosphoimidazolide. 31 P NMR (D2 O): δ –10.54. 1 H NMR (D2 O, 300 MHz): δ 8.03 (s, 1H), 8.01 (s, 1H), 6.02 (d, J = 3.6 Hz, 1H), 5.30 (dd, J = 6.3, 3.6 Hz, 1H), 5.19 (dd, J = 6.0, 2.4 Hz, 1H), 5.14-5.08 (m, 1H), 4.90-4.86 (m, 2H), 4.58-4.50 (m, 1H), 4.38 (pseudo brs, 1H), 4.13 (pseudo brs, 2H), 4.00 (pseudo brs, 2H), 2.71 (q, J = 7.2 Hz, 12H), 1.63 (s, 3H), 1.59 (s, 3H), 1.41 (s, 3H), 1.35 (s, 3H), 1.06 (t, J = 7.2 Hz, 18H).
9. Remove the protecting groups by standard methods and perform ion-exchange chromatography with Dowex 50WX8-200 (Na+ form) to give the desired pyrophosphates as the sodium salts, as illustrated by removal of the acetonide protecting groups of P1 -(2,6-diaminopurine-2 ,3 -isopropylidin-5 -yl)-P2 -(tiazofurin-2 ,3 isopropylidin-5 -yl) pyrophosphate: a. Dissolve 0.1 mmol pyrophosphate in 2 mL water. b. Add 500 mg Dowex 50WX8-200 resin (H+ form). c. Stir or shake gently for 17 hr.
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d. Apply the mixture to a column containing 20 equiv (based on the pyrophosphate) of Dowex 50WX8-200 resin (Na+ form). Elute with water and monitor fractions at 260 nm. e. Combine the fractions containing the pyrophosphate (those with UV absorbance at 260 mn) and lyophilize to give the pyrophosphate as the solid sodium salt. 10. Characterize by 1 H NMR and 31 P NMR. P1 -(Tiazofurin-5 -yl)-P2 -(2-aminoadenin-5 -yl) pyrophosphate: yield 64%. 31 P NMR (D2 O) δ –10.27. 1 H NMR (D2 O, 300 MHz): δ 8.05 (s, 1H), 7.96 (s, 1H), 5.86 (d, J = 5.7 Hz, 1H), 5.03 (d, J = 4.5 Hz, 1H), 4.65 (pseudo t, J = 5.6 Hz, 1H), 4.44 (dd, J = 4.8, 3.9 Hz, 1H), 4.31 (pseudo brs, 1H), 4.26-4.02 (m, 7H).
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Triethylamonium bicarbonate (TEAB), pH 7-7.5 Bubble CO2 gas through a mixture of triethylamine (1 mol) in water (800 mL) until the pH is between 7 and 7.5. Add water until the total volume is 1 L. Store in a refrigerator at 4◦ C for up to 3 months. Appropriate dilution of this 1 M solution gives the 0.04 M solution used in HPLC separation.
COMMENTARY Background Information
Synthesis of Dinucleoside Pyrophosphates
Phosphoimidazolides were first used in synthesis of nucleoside pyrophosphates by Cramer and Neunhoeffer, who reacted adenosine 5 -phosphoimidazolide with riboflavin monophosphate to give FAD (Cramer and Neunhoeffer, 1962). The studies and methods described here were developed to study the mechanism of enzymes that use NAD or its reduced form, NADH, as co-factors. These dinucleoside pyrophosphates are common in nature (Dolphin et al., 1987) and serve as substrates in numerous biochemical processes such as ribosylation of proteins (monoand poly-ADP-ribosylation), cell signaling, synthesis of cylicADP-ribose (Lee, 2002), DNA repair, and recombination. They also act as acetyl group acceptors in histone deacetylation catalyzed by the Sir2 family of NAD-dependent deacetylases (Denu, 2003). In addition, it is well established that a great number of enzymes in cells use nicotinamide coenzymes in oxidation-reduction processes. Several hundred NAD-dependent cellular dehydrogenases (including isoforms) are known, and all of them bind these co-factors. Since the co-factor-binding domain is conserved among many of these proteins, one might expect that selective inhibition of a particular dehydrogenase by a co-factor analog would be difficult if not impossible to achieve. This would mean
that nonselective inhibition of NAD enzymes would occur, leading to severe toxicity. Contrary to this expectation, it has been shown that some co-factor-type inhibitors, such as pyrophosphates, can inhibit a specific enzyme, inosine monophosphate dehydrogenase (IMPDH) with very high (>1,000-fold) selectivity (Pankiewicz et al., 2002a,b). This discovery has led to development of a new class of inhibitors, the novel mycophenolic adenine dinucleotide (MAD) analogs (Fig. 13.4.4). These are metabolically stable bisphosphonate analogs of pyrophosphates that bind at the co-factor-binding domain of human IMPDH and show good to excellent selectivity in inhibition of the human enzyme. Recent reviews discuss the clinical potential of such inhibitors (Saunders and Raybuck, 2000; Pankiewicz and Goldstein, 2003; Pankiewicz et al., 2004) and of selective inhibition of other dehydrogenases by dinucleoside pyrophosphates (Pankiewicz et al., 2002a,b). While the natural dinucleoside pyrophosphates all contain an adenine base, dinucleoside pyrophosphates containing other natural bases (G, C, U) in place of adenine can be prepared. However, the most important synthetic analogs contain an adenine or a substituted adenine moiety, which seems to be important for activity.
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Figure 13.4.4
Mycophenolic adenine dinucleotide (MAD).
Compound Characterization NMR spectra were recorded on a 300-MHz Varian Mercury NMR spectrometer in DMFd7 . Chemical shifts (δ) are given in ppm and referenced to tetramethylsilane as an internal standard and 85% phosphoric acid (H3 PO4 ) as an external standard. Coupling constants (J) are given in Hertz and refer to apparent multiplicities.
Critical Parameters and Troubleshooting While the synthesis of dinucleoside pyrophosphates is relatively short and straightforward, it requires careful attention to details of basic organic synthesis procedures. Efficient preparation of these compounds requires prior experience with routine organic chemistry laboratory techniques such as solvent evaporation, extraction, TLC, and HPLC. Characterization of the products demands knowledge of 1 H and 31 P NMR spectroscopy, and UV and electrospray ionization (ESI) or fast-atom-bombardment (FAB) mass spectrometry. General laboratory safety is also of primary concern when hazardous materials are involved. Strict adherence to the outlined methods is therefore highly recommended. The reaction of phosphoimidazolides with nucleoside monophosphates to give dinucleoside pyrophosphates requires dry conditions. Low yields are often due to contamination of reaction mixtures with atmospheric moisture. If the phosphoimidazole intermediates are not used immediately after preparation, care should be used to exclude moisture from the samples and the samples should be stored at or below −20◦ C. Anhydrous DMF should be used in the reaction.
Anticipated Results These procedures are suitable for preparation of milligram to ∼200 milligram amounts
of the final pyrophosphates. These final compounds are stable for months when stored as solids; no decomposition of the sodium salt of BAD was detected when stored at −20◦ C for 12 months. Aqueous solutions of final pyrophosphates at room temperature were also found to be quite stable. No decomposition was detected after four days by HPLC (solvent A: 0.04 M TEAB, pH 7.3; solvent B: 70% acetonitrile/water; isocratic 10% B, flow rate 0.5 mL/min, UV detection at 260 nm; Varian microsorb C18 column: 250 × 4.6 mm, 5-µm ◦ particle size, 100 A pore size). The reaction of a nucleoside monophosphate with imidazole and trichloroacetonitrile provides an alternative method for preparation of nucleoside phosphoimidazoles (Cramer and Neunhoeffer, 1962). However, in our hands, the use of 1,1 -carbonyldiimidazole provides a more rapid and more efficient route to phosphoimidazoles.
Time Considerations The synthesis of the dinucleoside pyrophosphates starting from nucleoside 5 monophosphates can be accomplished in 2 to 3 weeks depending on the nature of the side chain and whether protecting groups are required. Additional time will be required to remove protecting groups in the final products.
Literature Cited Cramer, F. and Neunhoeffer, H. 1962. Reaktionen von adenosine-5 -phosphorsaure-imidazolideine neue synthese von adenosindiphosphat und flavin-adenin-dinucleotid. Chem. Ber. 95:1664-1669. Denu, J.M. 2003. Linking chromatin function with metabolic networks: Sir2 family of NAD(+)dependent deacetylases. Trends Biochem. Sci. 28:41-48. Dolphin, D., Poulson, R., and Avramovic, O. 1987. Pyridine Nucleotide Coenzymes: Chemical, Biochemical, and Medical Aspects. John Wiley & Sons, New York.
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Gebeyehu, G., Marquez, V.E., Van Cott, A., Cooney, D.A., Kelley, J.A., Jayaram, H.N., Ahluwalia, G.S., Dion, R.L., Wilson, Y.A., and Johns, D.G. 1985. Ribavirin, tiazofurin, and selenazofurin: Mononucleotides and nicotinamide adenine dinucleotide analogues. Synthesis, structure, and interactions with IMP dehydrogenase. J. Med. Chem. 28:99-105. Hedstrom, L., Gan, L., Schlippe, Y.G., Riera, T., and Seyedsayamdost, M. 2003. IMP dehydrogenase: The dynamics of drug selectivity. Nucl. Acids Res. 3:97-98. Lee, H.C. 2002. Cyclic ADP-Ribose and NAADP: Structure, Metabolism and Functions. Plenum, New York. Li, H., Hallows, W.A., Punzi, J.S., Marquez, V.E., Carrell, H.L., Pankiewicz, K.W., Watanabe, K.A., and Goldstein, B.M. 1994a. Crystallographic studies of two alcohol dehydrogenasebound analogs of thiazole-4-carboxamide adenine dinucleotide (TAD), the active anabolite of the antitumor agent tiazofurin. Biochemistry 33:23-32. Li, H., Hallows, W.H., Punzi, J.S., Pankiewicz, K.W., Watanabe, K.A., and Goldstein, B.M. 1994b. Crystallographic studies of isosteric NAD analogues bound to alcohol dehydrogenase: Specificity and substrate binding in two ternary complexes. Biochemistry 33:1173411744. Pankiewicz, K. and Goldstein, B.M. 2003. Inosine Monophosphate Dehydrogenase: A Major Therapeutic Target. ACS Symposium Series No. 839. Oxford University Press, Oxford, U.K. Pankiewicz, K.W., Watanabe, K.A., LesiakWatanabe, K., Goldstein, B.M., and Jayaram, H.N. 2002a. The chemistry of nicotinamide adenine dinucleotide (NAD) analogues containing C-nucleosides related to nicotinamide riboside. Curr. Med. Chem. 9:733-741. Pankiewicz, K.W., Lesiak-Watanabe, K.B., Watanabe, K.A., Patterson, S.E., Jayaram, H.N., Yalowitz, J.A., Miller, M.D., Seidman, M., Majumdar, A., Prehna, G., and Goldstein, B.M. 2002b. Novel mycophenolic adenine bis(phosphonate) analogues as potential differentiation agents against human leukemia. J. Med. Chem. 45:703-712.
Pankiewicz, K.W., Patterson, S.E., Black, P.L., Jayaram, H.N., Risal, D., Goldstein, B.M., Stuyver, L.J., and Shinazi, R.F. 2004. Cofactor mimics as selective inhibitors of NADdependent inosine monophosphate dehydrogenase (IMPDH)—the major therapeutic target. Curr. Med. Chem. 11:887-900. Saunders, J.O. and Raybuck, S.A. 2000. Annual Reports in Medicinal Chemistry. Academic Press, San Diego. Scheit, K.H. 1980. Nucleotide Analogs. Synthesis and Biological Function. John Wiley & Sons, New York. Yoshikawa, M., Kato, T., and Takenishi, T. 1967. A novel method for phosphorylation of nucleosides to 5 -mononucleotides. Tetrahedron Lett. 8:5065-5068. Zatorski, A., Lipka, P., Mollova, N., Schram, K.H., Goldstein, B.M., Watanabe, K.A., and Pankiewicz, K.W. 1993. Synthesis of thiazole4-carboxamide-adenine difluoro-methylenediphosphonates substituted with fluorine at C-2 of the adenosine. Carbohydr. Res. 249:95-108. Zatorski, A., Goldstein, B.M., Colby, T.D., Jones, J.P., and Pankiewicz, K.W. 1995. Potent inhibitors of human inosine monophosphate dehydrogenase type II. Fluorine-substituted analogues of thiazole-4-carboxamide adenine dinucleotide. J. Med. Chem. 38:1098-1105. Zatorski, A., Watanabe, K.A., Carr, S.F., Goldstein, B.M., and Pankiewicz, K.W. 1996. Chemical synthesis of benzamide adenine dinucleotide: Inhibition of inosine monophosphate dehydrogenase (types I and II). J. Med. Chem. 39:24222426.
Contributed by Liqiang Chen, Dominik Rejman, Laurent Bonnac, Krzysztof W. Pankiewicz, and Steven E. Patterson Center for Drug Design The University of Minnesota Minneapolis, Minnesota
Synthesis of Dinucleoside Pyrophosphates
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Synthesis of Methylenebis(phosphonate) Analogs of Dinucleotide Pyrophosphates
UNIT 13.5
Natural nucleoside pyrophosphates such as NAD, NADH, FAD, FADH2 , NADP, and NADPH play a critical role as enzyme cofactors in biochemical redox reactions (Fig. 13.5.1). Analogs of these nucleoside pyrophosphates that do not act in redox processes serve as valuable tools for investigating the mechanisms of these enzymes (see UNIT 13.4). In this unit, analogs with a methylenebis(phosphonate) linkage are synthesized. The preparation of 2 ,3 -O-ispropylidenenucleoside 5 -methylenebis(phosphonate)s from 2 ,3 -O-isopropylidene nucleosides by reaction with the commercially available methylenebis(phosphonic acid dichloride) in triethyl phosphate is based on the procedure of Patterson et al. (2004a) and is described in the first method (Basic Protocol 1). Alternate Protocol 1 can be used for reactions starting from nucleosides having protecting groups other than 2 ,3 -O-isopropylidene or even from unprotected nucleosides. Protected or unprotected carbohydrates or non-nucleoside alcohol derivatives are also suitable for reaction with methylenebis(phosphonic acid dichloride) using this protocol. The alternative methods for the synthesis of P1 -monosubstituted methylenebis(phosphonate)s, including nucleoside 5 -methylenebis(phosphonate)s, were originally described by Poulter and colleagues (Davisson et al., 1987) and by Lesiak et al. (1998a,b). The use of these methylenebis(phosphonate) monoesters in synthesis of the methylenebis(phosphonate) P1 ,P2 -diesters is described in Basic Protocol 2 and is based on procedures described by Lesiak et al. (1997) and Pankiewicz et al. (1997a,b). The methylenebis(phosphonate) monoesters and the corresponding methylenebis(phosphonate) diesters are stable to conditions used for removal of standard protecting groups such as 2 ,3 -isopropylidene (acetonide), 2 ,3 -acyl protecting groups, tetrahydropyranyl groups, and others. Preparation of diesters from non-nucleoside methylene (biophosphates) is described in Alternate Protocol 2. CAUTION: Carry out all operations involving organic solvents and reagents in a wellventilated chemical fume hood. Wear appropriate protective clothing and glasses.
PREPARATION OF 2 ,3 -O-ISOPROPYLIDENENUCLEOSIDE 5 -METHYLENEBIS(PHOSPHONATE)S FROM 2 ,3 -O-ISOPROPYLIDENENUCLEOSIDES
BASIC PROTOCOL 1
Nucleoside 5 -methylenebis(phosphonate)s have been prepared previously (Davisson et al., 1987; Lesiak et al., 1998a,b). The protocol described here is based on the methods described by Lesiak et al. (1997), Pankiewicz et al. (2002a), and Patterson et al. (2004a); a substantially similar report was subsequently published by Kalek et al. (2005). The reaction is illustrated in Figure 13.5.2. Many 2 ,3 -O-isopropylidene nucleosides are commercially available from suppliers such as Sigma-Aldrich and Berry & Associates. Those that are not commercially available can be synthesized by the well-established isopropylidenation reaction (Greene and Wuts, 1999). They can be purified by crystallization or silica gel column chromatography.
Materials 2 ,3 -O-Isopropylidene nucleoside (e.g., Sigma-Aldrich or Berry & Associates, http://www.berryassoc.com) Triethyl phosphate (EtO)3 PO, anhydrous Methylenebis(phosphonic acid dichloride) (Sigma-Aldrich) Contributed by Krzysztof W. Pankiewicz, Guangyao Gao, and Steven E. Patterson Current Protocols in Nucleic Acid Chemistry (2006) 13.5.1-13.5.13 C 2006 by John Wiley & Sons, Inc. Copyright
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13.5.1 Supplement 24
Figure 13.5.1 Natural nucleoside pyrophosphates and tiazole-3-carboxamide and benzamide adenine dinucleotides (TAD and BAD).
Solvent A: 0.04 M triethylammonium bicarbonate (TEAB), pH 7.0 to 7.5 (see recipe for 1 M) Solvent B: 70% (v/v) acetonitrile in H2 O 1 M TEAB, pH 7.0 to 7.5 (see recipe) Ethyl acetate (EtOAc)
Synthesis of Methylenebis (phosphonate) Analogs of Dinucleotide Pyrophosphates
HPLC system with: C18 RP-HPLC analytical column (Varian Microsorb-MV 100-5, 250 × ◦ 4.6–mm; 5-µm particle size, 100 A pore size) C18 RP-HPLC preparative column (Varian Microsorb, 250 × 41.4–mm; 8-µm ◦ particle size, 100 A pore size) UV-Vis detector Separatory funnel Rotary evaporator Lyophilizer
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Figure 13.5.2
Reaction of 2 ,3 -O-isopropylideneadenosine with methylenebis(phosphonic acid dichloride).
1. Prepare a 0.04 M solution of the 2 ,3 -isopropylidene nucleoside in anhydrous (EtO)3 PO and cool to 0◦ C. 2. Prepare a 0.12 M solution of methylenebis(phosphonic acid dichloride) in anhydrous (EtO)3 PO, cool to 0◦ C, and then add to the nucleoside solution in a 1:1 (v/v) ratio. Stir for 4 hr. 3. Monitor the reaction by analytical RP-HPLC using the following conditions:
Solvent A: 0.04 M TEAB, pH 7.0 to 7.5 Solvent B: 70% (v/v) acetonitrile/H2 O ◦ Column: Microsorb-MV 100-5, 250 × 4.6–mm; 5-µm particle size, 100 A pore size Gradient: isocratic, 15% B Flow rate: 0.5 mL/min UV detection at 260 nm. 4. Quench by adding 10- to 20-mL aliquots of 1 M TEAB, pH 7.0 to 7.5, with continuous stirring until the effervescence has ceased and the mixture has been neutralized. 5. Extract the mixture twice, each time with 1/3 vol of EtOAc. Evaporate the aqueous layer to dryness on a rotary evaporator equipped with a water aspirator. 6. Purify the resulting product by preparative RP-HPLC using the above conditions and a Varian Microsorb C18 preparative column. 7. Concentrate the fractions containing the desired bis(phosphonate) on a rotary evaporator. 8. Remove excess triethylamine by coevaporating three times, each time with 100 to 200 mL water, then lyophilize to give the desired 2 ,3 -O-isopropylidenenucleoside 5 -methylenebis(phosphonate) as the solid triethylammonium salt. After lyophilization, the resulting solid nucleoside 5 -methylenebis(phosphonate) salts can be stored for months at room temperature.
9. Characterize by 1 H and 31 P NMR, and mass spectrometry. 2 ,3 -O-Isopropylideneadenosine 5 -methylenebis(phosphonate): yield, 85% to 99%. 1 H NMR (D2 O): δ 1.18-1.21 (t, 18H, Et3 N), 1.42 (s, 3H, iPr), 1.64 (s, 3H, iPr), 2.04-2.10 (t, 2H, H5 ), 3.10-3.11 (two q, 12H, Et3 N), 4.06 (m, P-CH2 -P), 4.57 (s, H4 ), 5.18 (m, 1H, H3 ), 5.33 (s, bro, 1H, H2 ), 6.12 (s, bro, 1H, H1 ), 8.06, 8,37 (two 2H singlets, H2, H8). 31 P NMR (D2 O): δ 15.5 (d, J = 19.2 Hz, P-CH2 -P), 19.46 (d, J = 19.2 Hz, P-CH2 -P). MS (ESI): m/z 464 (M – H)− . 2 ,3 -O-Isopropylideneadenosine 5 -methylenebis(phosphonate) has been described previously (Pankiewicz et al., 1997a,b).
Nucleoside Phosphorylation and Related Modifications
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ALTERNATE PROTOCOL 1
PREPARATION OF METHYLENEBIS(PHOSPHONATE) MONOESTERS FROM OTHER STARTING COMPOUNDS While the above procedure is general for reaction of 2 ,3 -O-protected nucleosides with methylenebis(phosphonic acid dichloride), 2 - or 3 -deoxynucleosides should be protected differently, and a variety of groups are suitable for protecting OH groups other than the 5 -OH. Among others, acetyl, benzoyl, benzyl, tetrahydropyranyl (Lesiak et al., 1997), and hindered silyl ethers such as the tert-butyldimethylsilyl group can be used. Nucleosides protected with these groups can be used to make methylenebis(phosphonate) monoesters using this protocol. Unprotected nucleosides can also be converted to their corresponding 5 -methylenebis(phosphonate)s by reaction with methylenebis(phosphonic acid dichloride) in triethyl or trimethyl phosphate. This reaction is not very selective, however, and the formation of the corresponding methylenebis(phosphonate)s at other than the 5 -hydroxyl groups is observed (Patterson et al., 2004b). This requires careful monitoring of the reaction progress by HPLC. Sometimes it is difficult to separate the desired 5 -bis(phosphonate), and yields are usually lower than those reported for suitably protected nucleosides. Numerous carbohydrate derivatives and non-nucleoside alcohols can also react with methylenebis(phosphonic acid dichloride) in triethyl or trimethyl phosphate to give the corresponding methylenebis(phosphonate) monoesters, yielding, for example, C2-mycophenolic alcohol methylenebis(phosphonate) (Fig. 13.5.3; Pankiewicz et al., 2002a). Many protected nucleosides are commercially available from suppliers such as SigmaAldrich and Berry & Associates. Those that are not commercially available can be synthesized by methods in the literature.
Additional Materials (also see Basic Protocol 1) Alcohol derivative (e.g., 7-O-benzyl-C2-mycophenolic alcohol; Pankiewicz et al., 2002a), carbohydrate, 2 - or 3 -protected nucleoside, or unprotected nucleoside Methanol (MeOH) 10% palladium catalyst on activated carbon (Sigma-Aldrich) 1. Prepare a 0.04 M solution of the alcohol derivative (e.g., 7-O-benzyl-C2mycophenolic alcohol) in anhydrous (EtO)3 PO and cool to 0◦ C. 2. Prepare a 0.12 M solution of methylenebis(phosphonic acid dichloride) in anhydrous (EtO)3 PO, cool to 0◦ C, and then add to the solution of starting derivative in a 1:1 (v/v) ratio. Stir for 4 hr. 3. Monitor the reaction by analytical RP-HPLC using the following conditions:
Solvent A: 0.04 M TEAB, pH 7.5 Solvent B: 70% (v/v) acetonitrile/H2 O ◦ Column: Microsorb-MV 100-5, 250 × 4.6–mm; 5-µm particle size, 100 A pore size Gradient: isocratic, 15% B Flow rate: 40 mL/min UV detection at 260 nm. Synthesis of Methylenebis (phosphonate) Analogs of Dinucleotide Pyrophosphates
4. Quench by adding 10- to 20-mL aliquots of 1 M TEAB, pH 7.0 to 7.5, with continuous stirring until the effervescence has ceased and the mixture has been neutralized. 5. Extract the mixture twice, each time with 1/3 vol of EtOAc. Evaporate the aqueous layer to dryness on a rotary evaporator equipped with a water aspirator.
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Figure 13.5.3
Reaction of non-nucleoside alcohol with methylenebis(phosphonic acid dichloride).
6. Purify the resulting product by preparative RP-HPLC using the above conditions and a Varian Microsorb C18 preparative column. 7. Concentrate the fractions containing the desired alcohol bis(phosphonate) on a rotary evaporator. 8. Remove excess triethylamine by coevaporating three times, each time with 100 to 200 mL water, then lyophilize to give the desired methylenebis(phosphonate) as the solid triethylammonium salt. 9. Characterize by 1 H and 31 P NMR, and mass spectrometry. [7-Benzyloxy-6-(2-hydroxyethyl)-5-methoxy-4-methylphthalan-1-one-2-yl]methylenebis (phosphonic) acid: yield, 87%. 1 H NMR (D2 O): δ 1.85 (t, 2H, P-CH2 P, J = 19.6 Hz), 2.07 (s, 3H, CH3 ), 2.76 (t, 2H, CH2 (1), J1-2 = 6.8 Hz), 3.69 (s, 3H, OCH3 ), 3.79 (q, 2H, CH2 (2), JH,H = JH,P = 6.8 Hz), 5.06 and 5.20 (two 2H, CH2 -lactone, CH2 Ph), 7.29-7.37 (m, 5H, Ph). 31 P NMR (D2 O): δ15.84 (d, 1P, J = 7.7 Hz), 17.71 (d, 1P). MS (FAB): m/z 485 (M – H)− . After lyophilization, the resulting solid nucleoside 5 -methylenebis(phosphonate) salts can be stored for months at room temperature.
10. Remove the 7-O-benzyl protecting group from the derivative obtained in step 8 (100 mg) in MeOH (20 mL) over a palladium catalyst on activated carbon (5 to 10 mg) under atmospheric pressure and with stirring. Remove the catalyst by filtration, then remove the MeOH in vacuo. 11. Characterize by 1 H and 31 P NMR, and mass spectrometry. [7-Hydroxy-6-(2-hydroxyethyl)-5-methoxy-4-methylphthalan-1-one-2-yl]methylenebis (phosphonic) acid: yield, 99%. 1 H NMR (D2 O): δ 2.07 (t, 2H, P-CH2 -P, J = 19.6 Hz), 2.07 (s, 3H, CH3 ), 2.76 (t, 2H, CH2 (1), J1-2 = 7.0 Hz), 3.69 (s, 3H, OCH3 ), 3.79 (q, 2H, CH2 (2 ), JH,H = JH,P = 7.0 Hz), 5.20 (s, 2H, CH2 -lactone, CH2 Ph). 31 P NMR (D2 O): δ 15.84 (d, 1P, J = 7.7 Hz), 17.71 (d, 1P). MS (FAB): m/z 395 (M – H)− (10%); the sodium adduct: m/z 417 (5%).
PREPARATION OF METHYLENEBIS(PHOSPHONATE) ANALOGS OF DINUCLEOSIDE PYROPHOSPHATES The protocol described here, developed by Pankiewicz et al. (1997a,b), is based on 31 P-NMR-controlled coupling of nucleoside 5 -methylenebis(phosphonate)s with nucleosides, alcohols, and carbohydrates in the presence of a dehydrating agent such as dicyclohexylcarbodiimide (DCC) or preferably diisopropylcarbodiimide (DIC). The first step of this reaction, illustrated in Figure 13.5.4, is formation of the tetraphosphonate anhydride (S.1) from adenosine methylenebis(phosphonate). S.1 is further dehydrated to the bicyclic trisanhydride (BTA; S.2). The BTA contains four chiral phosphorus atoms and therefore shows multiple signal resonances in its 31 P NMR spectrum (Fig. 13.5.5). Addition of another nucleoside (ROH) to the solution of BTA results in nucleophilic attack at the sterically unhindered P2 and P3 atoms only, to give the P1 ,P2 ,P3 ,P4 -tetrasubstituted tetraphosphonate anhydride (S.3). Upon hydrolysis, this affords the desired Current Protocols in Nucleic Acid Chemistry
BASIC PROTOCOL 2
Nucleoside Phosphorylation and Related Modifications
13.5.5 Supplement 24
Figure 13.5.4 Reaction of nucleoside bis(phosphonate) monoester with DIC and nucleosides, carbohydrates, and nonnucleoside alcohols.
methyelenbis(phosphonate) analog of dinucleoside pyrophosphate (S.4). As an example, this protocol describes the synthesis of the methylenebis(phosphonate) analog of benzamide adenine dinucleotide (BAD) starting from 2 ,3 -O-isopropylideneadenosine 5 -methylenebis(phosphonate) (see Basic Protocol 1) and 3-(2 ,3 -O-isopropylidene-βD-ribofuranos-5-yl)benzamide.
Synthesis of Methylenebis (phosphonate) Analogs of Dinucleotide Pyrophosphates
Many nucleoside 5 -methylenebis(phosphonate)s can be synthesized by the reaction of 2 ,3 -O-isopropylidene-protected nucleosides with methylenebis(phosphonic acid dichloride) as described in Basic Protocol 1 or by the alternative methods of Davisson et al. (1987) or Lesiak et al. (1998a,b). These compounds can also be prepared from nucleosides containing a free 5 -OH and protected with a variety of protecting groups other than isopropylidene. In addition, unprotected nucleoside 5 -methylenebis(phosphonate)s can be synthesized first and then protected with groups such as orthoesters (Marquez et al., 1986) and possibly others for further DCC or DIC coupling. The monosubstituted 5 -methylenebis(phosphonate)s are usually used in the form of their triethylammonium salts obtained after HPLC purification with TEAB buffer. These compounds are stable in water for at least days or weeks, depending on their protecting groups— for example, the isopropylidene group will be cleaved if 2 ,3 -O-ispropylideneadenosine
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Figure 13.5.5
31
P NMR multi-signal resonances of BTA.
5 -methylenebis(phosphonate) is stored as an aqueous solution of the free acid. Strictly anhydrous conditions are required to ensure good yields. Anhydrous pyridine and completely dried nucleoside derivatives should be used in reactions. RP-HPLC purification can be performed on any HPLC instrument.
Materials 2 ,3 -O-Isopropylidenenucleoside 5 -methylenebis(phosphonate): e.g., 2 ,3 -Oisopropylideneadenosine 5 -methylenebis(phosphonate) (Basic Protocol 1) Anhydrous pyridine, HPLC grade Anhydrous pyridine-d5 Diisopropylcarbodiimide (DIC) Protected nucleoside containing free 5 -hydroxyl group: e.g., 3-(2 ,3 -O-isopropylidene -β-D-ribofuranos-5-yl)benzamide Solvent A: 0.1 M triethylammonium bicarbonate (TEAB), pH 7.0 to 7.5 (see recipe) Solvent B: 70% (v/v) acetonitrile in H2 O Dowex 50WX8-200, H+ form Dowex 50WX8-200, Na+ form HPLC system with: C18 HPLC analytical column (Varian Microsorb-MV 100-5, 250 × 4.6–mm; ◦ 5-µm particle size, 100 A pore size) C18 RP-HPLC preparative column (Varian Microsorb, 250 × 41.4–mm; 8-µm ◦ particle size, 100 A pore size) UV-Vis detector Filter paper or Buchner funnel Rotary evaporator Lyophilizer Glass chromatography column
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1. Prepare a 0.1 M solution of the nucleoside 5 -methylenebis(phosphonate) in anhydrous pyridine/10% pyridine-d5 containing 3.3 eq of DIC and stir (2 to 5 hr) until 31 P NMR of the reaction mixture shows multi-signal resonances (see Fig. 13.5.5), indicating that the formation of BTA is complete. The reaction mixture should contain sufficient pyridine-d5 to obtain a deuterium lock in the NMR spectrometer. A 10% solution of pyridine-d5 in pyridine is sufficient.
2. Add 1.2 eq of a protected nucleoside containing a free 5 -hydroxyl group. Heat at 60◦ to 65◦ C until two multiplets at δ 8-9 and δ 18-21 are observed (could range from 2 hr to 2 days). 3. Quench by adding 20 to 30 mL water per 100 mg, and stir at room temperature for 1 to 3 hr. If necessary, heat at 60◦ to 80◦ C until hydrolysis of S.3 is complete. Monitor the progress of hydrolysis and formation of the desired methylenebis(phosphonate) diester by 31 P NMR or by analytical RP-HPLC using the following conditions:
Solvent A: 0.1 M TEAB, pH 7.0 to 7.5 Solvent B: 70% (v/v) acetonitrile ◦ Column: Microsorb-MV 100-5, 250 × 4.6–mm; 5-µm particle size, 100 A pore size Gradient: linear Flow rate: 40 mL/min UV detection at 260 nm. By NMR, a further simplification of the signals to double-doublet-like should be observed.
4. Filter the mixture using a Buchner funnel or filter paper, and then concentrate in vacuo using a rotary evaporator and water aspirator. 5. Dissolve the resulting residue in a minimum volume 0.04 M TEAB and purify by HPLC using the following conditions:
Solvent A: 0.1 M TEAB, pH 7.0 to 7.5 Solvent B: 70% (v/v) acetonitrile/H2 O Column: Varian Microsorb C18 RP-HPLC preparative column (250 × ◦ 41.4–mm; 8-µm particle size, 100 A pore size) Gradient: solvent A followed by a linear gradient of 70% solvent B Flow rate: 40 mL/min UV detection at 260 nm. The abovementioned columns and conditions pertain specifically to the purification of P1 -[3-(2,3-O-isopropylidene-β-D-ribofuranos-5-yl)benzamide] P2 -(2 ,3 -O-isopropylideneadenosine) 5 -methylenebisphosphonate (Pankiewicz et al., 1997a). A replacement of benzamide riboside by substituents with different lipophilicity will require appropriate modification of the mobile phase. More lipophilic compounds will require more CH3 CN and less lipophilic compounds less CH3 CN.
6. Concentrate the fractions containing the desired bis(phosphonate) (i.e., those with absorbance at 260 nm) on a rotary evaporator. 7. Remove excess triethylamine by coevaporating three times, each time with 20 to 50 mL water, then lyophilize to give the desired methylenebis(phosphonate) diester as the solid triethylammonium salt. Synthesis of Methylenebis (phosphonate) Analogs of Dinucleotide Pyrophosphates
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8. Characterize by 1 H and 31 P NMR. P1 -[3-(2,3-O-Isopropylidene-β-D-ribofuranos-5-yl)benzamide] P2 -(2 ,3 -O-isopropylideneadenosine) 5 -methyelenebisphosphonate (S.4) as the ditriethylammoniun salt: yield, 79%. 1 H NMR (D2 O): δ 1.27-1.31 (t, 18H, Et3 N), 1.40 (s, 3H, iPr), 1.46 (s, 3H, iPr), 1.64 (s, 3H, iPr), 1.68 (s, 3H, iPr), 2.05-2.20 (m, 2H, PCH2 P), 3.21 (q, 12H, Et3 N), 4.09-4.12 [m, 4H, H5 , H5 (B), H5 (A)], 4.26-4.28 [m, 1H, H4 (B)], 4.59-4.64 [m, 2H, H4 (A), H2 (B)], 4.80 [1H, H1 (B)], 4.88 [dd, 1H, H3 (B), J1 ,2 = 5.7 Hz, J2 ,3 = 6.5 Hz], 5.20 [m, 1H, H3 (A)], 5.30 [dd, 1H, H2 (A), J1 ,2 = 3.0 Hz, J2 ,3 = 6.5 Hz], 6.12 [d, 1H, H1 (A)], 7.39 [pseudo t, 1H, H5(B)], 7.46 [d, 1H, H4(B), J4,5 = 7.8 Hz], 7.64 [d, 1H, H6(B), J5,6 = 7.8 Hz], 7.67 (s, 1H, H2(B)], 8.15, 8.40 [two 1H singlets, H2(A), H8(A)].
9. Remove the protecting groups by standard methods (Green and Wuts, 1999). For removal of the benzyl protecting group, use treatment with 10% palladium catalyst on activated carbon in MeOH under a hydrogen atmosphere (20 psi) in a Parr hydrogenation apparatus for 1 hr. Remove the catalyst by filtration through a Buchner funnel and evaporate the MeOH in vacuo. Dissolve the residue in a mixture consisting of 3 mL MeOH, 3 mL water, and 1 mL CF3 COOH per 100 mg residue. Stir 1 hr, evaporate to dryness on rotary evaporator, coevaporate three times, each time with 20 mL toluene, then dissolve in water and lyophilize. 10. Perform ion-exchange chromatography with Dowex 50 (sodium form) to give the desired methylenebis(phosphonate) as the sodium salt. a. b. c. d.
Dissolve 0.1 mmol of the methylenebis(phosphonate) in 2 mL water. Add 500 mg Dowex 50WX8-200 (H+ form). Stir or shake the mixture gently for 17 hr at ∼18◦ C. Apply the mixture to a glass chromatography column containing Dowex 50WX8200 [20 eq based on the methylenebis(phosphonate); Na+ form] and elute with water. e. Combine fractions containing the methylenebis(phosphonate), i.e., those with UV absorbance at 260 nm, and lyophilize to give the pyrophosphate as the solid sodium salt. The above instructions pertain specifically to the removal of the acetonide protecting groups to give P1 -(3-β-D-ribofuranos-5-ylbenzamide) P2 -adenosine 5 -methylenebis (phosphonate).
11. Characterize by 1 H and 31 P NMR, and mass spectrometry. Methylenebis(phosphonate) analog of BAD: yield, 95%. 1 H NMR (D2 O): δ 2.27 (pseudo t, 2H, PCH2 P), 4.05 [dd, 1H, H2 (B), J1 ,2 = 7.0 Hz, J2 ,3 = 5.1 Hz], 4.17-4.20 [m, 4H, H5 , H5 (A), H5 (B)], 4.25-4.27 [m, 2H, H3 (B), H4 (B)], 4.37 [pseudo t, 1H, H3 (A)], 4.66 [pseudo t, H2 (A)], 4.80 [d, 1H, H1 (B)], 6.06 [d, 1H, H1 (A), J1 ,2 = 4.9 Hz], 7.41 [pseudo t, 1H, H5(B)], 7.56 [d, 1H, H4(B), J4,5 = 7.8 Hz], 7.63 [d, 1H, H6(B), J5,6 = 7.8 Hz], 7.69 [s, 1H, H2(B)], 8.32, 8.57 [two 1H singlets, H2(A), H8(A)]. 31 P NMR (D2 O): 20.94 (s). MS (ES): m/z 659 (M – H)− .
PREPARATION OF P1 ,P2 -NON-SYMMETRICALLY SUBSTITUTED METHYLENEBIS(PHOSPHONATE) DIESTERS
ALTERNATE PROTOCOL 2
The reaction using a non-nucleoside methylenebis(phosphonate) proceeds essentially as in Basic Protocol 2, and is presented here for the preparation of the methylenebis (phosphonate) analog of mycophenolic adenine dinucleotide (MAD) from the alcohol 5 -methylenebis(phosphonate) (see Alternate Protocol 1) and 2 ,3 -O-isopropylideneadenosine. This compound could also be prepared from 2 ,3 -O-isopropyladenosine 5 -methylenebis(phosphonate) and C2-mycophenolic alcohol as in Basic Protocol 2 and shown in Figure 13.5.4.
Nucleoside Phosphorylation and Related Modifications
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Figure 13.5.6 Reaction of non-nucleoside methylenebis(phosphonate) with DIC and 2 ,3 -O-isopropylideneadenosine. An intermediate BTA (not shown) with similar multi-signal resonances as in Fig. 13.5.5 was formed first and then reacted with the adenosine derivative.
Proceed as in Basic Protocol 2 with the variations described below at the corresponding steps.
Additional Materials (also see Basic Protocol 2) Alcohol 5 -methylenebis(phosphonate) (Alternate Protocol 1) Protected nucleoside containing free 5 -hydroxyl group: e.g., 2 ,3 -O-isopropylideneadenosine Trifluoroacetic acid (CF3 COOH) Parr hydrogenation apparatus (Parr Instrument) and hydrogen source 1a. Use an alcohol 5 -methylenebis(phosphonate) in place of the 2 ,3 -O-isopropylidenenucleoside 5 -methylenebis(phosphonate). 8a. Characterize the product of the reaction (7-O -benzyl-C2-MAD; Fig. 13.5.6) by 1 H and 31 P NMR. P1 - (2 ,3 -O - Isopropylideneadenosin- 5 - yl) - P2 - [7- benzyloxy- 6- (2-hydroxyethyl) -5methoxy-4-methylphthalan-1-one-2-yl]methylenebis(phosphonate): yield, 70% to 80%. 1 H NMR (D2 O): δ 1.18 (t, 18H, Et3 N), 1.45 and 1.60 (two s, 3H each, isopropylidene), 2.05 (s, 3H, CH3 ), 2.51 (dt, 2H, P-CH2 -P, J = 21.2 Hz, J = 8.8 Hz), 3.10 (q, 12H, Et3 N), 3.40-3.48 (m, 2H, H5 , H5 ), 3.60 [t, 2H, CH2 (1), J = 6.4 Hz], 3.62 (s, 3H, OCH3 ), 4.10-4.12 [m, 3H, CH2 (2), H4 ], 4.54 (m, 1H, H3 ), 4.67 (m, 1H, H2 ), 4.95 (s, 2H, CH2 -lactone), 5.14, (s, 2H, CH2 -benzyl), 6.01 (d, 1H, H1 , J1 ,2 = 2.4 Hz), 7.94 (t, 2H, phenyl, J = 7.6 Hz), 7.96, 8.27 (two 1H singlets, H2, H8-adenine), 8.42 (t, 1H, phenyl, J = 7.6 Hz), 8.69-8.71 (m. 2H, phenyl). 31 P NMR (D2 O): δ 16.06-17.79 (AB system, J = 16.6 Hz).
11a.Characterize the product by 1 H and 31 P NMR, and mass spectrometry. Methylenebis(phosphonate) analog of MAD: yield, 90%. 1 H NMR (D2 O): δ 1.69 (s, 3H, CH3 ), 2.07 (t, 2H, P-CH2 -P, J = 20.0 Hz), 2.61 [t, 2H, CH2 (1), J = 6.8 Hz], 3.52 (s, 3H, OCH3 ), 3.73 [q, 2H, CH2 (2), JH.H = 6.4 Hz, JH.P = 6.4 Hz], 4.02 (m, 1H, H5 , J5 ,5 = 12.4 Hz), 4.10 (m, 1H, H5 ), 4.17 (m, 1H, H4 ), 4.34 (t, 1H, H3 , J2 ,3 = 4.8 Hz, J3 ,4 = 5.8 Hz), 4.47 (t, 1H, H2 , J1 ,2 = 4.4 Hz), 4.74 (s, 2H, CH2 -lactone), 5.77 (d, 1H, H1 ), 7.85, 8.21 (two 1H singlets, H2, H8-adenine). 31 P NMR (D2 O): δ 17.20-17.50 (two overlapping doublets). MS (FAB): m/z 644 (M – H)− (55%).
Synthesis of Methylenebis (phosphonate) Analogs of Dinucleotide Pyrophosphates
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REAGENTS AND SOLUTIONS Use deionized water (resistivity >18 M-cm) in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Triethylammonium bicarbonate (TEAB), 1 M, pH 7 to 7.5 Prepare a mixture of 1 mol triethylamine in 800 mL water. Bubble CO2 through this mixture until the pH is between 7 and 7.5. Add water until the total volume is 1 L. Store up to 3 months at 4◦ C. Appropriate dilution of this 1 M solution gives the 0.04 and 0.1 M solutions used in HPLC separation.
COMMENTARY Background Information
NAD+ , its reduced form NADH, and their 2 -phosphorylated analogs (NADP, NADPH) serve as coenzymes in oxidation-reduction processes. These compounds also serve as substrates in numerous chemical reactions such as ribosylation of proteins (monoand poly-ADP-ribosylation), cell signaling (cADP-ribose; Shuto and Matsuda, 2004), and DNA repair and recombination, and also act as the acetyl group acceptors in histone deacetylation catalyzed by the Sir2 family of NADdependent deacetylases (Sauve and Schramm, 2004). In their natural pyrophosphate form, these coenzymes and their analogs are vulnerable to cleavage by cellular phosphodiesterases and cannot penetrate the cell membrane. In contrast, methylenebis(phosphonate) diesters are metabolically stable analogs that can penetrate cells. Thus, methylenebis(phosphonate) analogs are useful tools for studies of biological effects of such compounds in vitro and in vivo, and have demonstrated potential value as chemotherapeutics. Tiazofurin, benzamide riboside (Jayaram et al., 2002; Pankiewicz et al., 2002b), mycophenolic acid (Bentley, 2000), and their corresponding adenosinemethylenebis(phosphonate) esters selectively inhibit inosine monophosphate dehydrogenase (IMPDH) and show therapeutic potential in treatment of cancer (Patterson et al., 2004a) and as immunosuppressants (Pankiewicz et al., 1999). Tiazofurin (TR), used clinically for treatment of patients in blast crisis of chronic myelogenous leukemia (CML; Grifantini, 2000), and the new anticancer drug candidate benzamide riboside (BR; Gharehbaghi et al., 2002) are nucleoside prodrugs that are converted in cells into tiazofurin adenine dinucleotide (TAD) and benzamide adenine dinucleotide (BAD), respectively, which are potent inhibitors of inosine monophosphate dehydro-
genase (IMPDH; see UNIT 13.4 for additional information). However, the activated forms, TAD and BAD, are metabolically unstable, and are easily cleaved by phosphodiesterases into the corresponding mononucleotides, which are then cleaved by phosphatases to the inactive parent nucleosides TR and BR. Resistance to TR and BR develops due to decreased ability of resistant cells to synthesize TAD and BAD, and also due to the markedly higher activity of phosphodiesterases (including a specialized TADase) that cleave the pyrophosphate bond (Pankiewicz et al., 2002a). Replacement of the pyrophosphate oxygen by a methylene group, such as in methylenebis(phosphonate)s of TAD and BAD, results in derivatives with a similar potency against IMPDH (Lesiak et al., 1997; Pankiewicz et al., 1997b). Since these compounds are not cleaved by phosphodiesterases, they show potent activity against tiazofurin- or benzamide-resistant CML cells. The most potent inhibitor of IMPDH is mycophenolic acid (MPA), which binds at the nicotinamide mononucleotide (NMN) subdomain of the NAD binding pocket, leaving the adenosine mononucleotide subdomain empty (Sintchak et al., 1996). However, MPA is not active against cancer, because of extensive glucuronidation (Franklin et al., 1996). Attaching an MPA moiety to adenosine 5 -methylenebis(phosphonate) yields a whole cofactor mimic, mycophenolic adenine dinucleotide (MAD), in the bis(phosphonate) form (Lesiak et al., 1998b). MAD is a potent inhibitor of IMPDH, is resistant to glucuronidation, and shows potent anticancer activity against K562 cells (a cancer cell line derived from a CML patient; Lesiak et al., 1998b) as well as in SCID mice xenotransplanted with K562 leukemia cells (Patterson et al., 2004a).
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Compound Characterization NMR spectra were recorded on a 300-MHz Varian Mercury NMR spectrometer in D2 O or DMSO-d6 . Chemical shifts (δ) are given in ppm and referenced to tetramethylsilane as an internal standard and 85% phosphoric acid (H3 PO4 ) as an external standard. Coupling constants (J) are given in Hertz (Hz) and refer to apparent multiplicities.
Critical Parameters and Troubleshooting While the synthesis of methylenebis(phosphonate) diesters is relatively short and straightforward, it requires careful attention to details of basic organic synthesis procedures. Efficient preparation of the various compounds requires prior experience with routine organic chemistry laboratory techniques such as solvent evaporation, extraction, TLC, and HPLC. The reaction of methylenebis(phosphonate) monoesters with DIC and alcohols to give nonsymmetrical methylenebis(phosphonate) diesters requires dry conditions. Low yields are often due to contamination of reaction mixtures with atmospheric moisture. Characterization of the products demands knowledge of 1 H and 31 P NMR spectroscopy, UV spectrophotometry, and electrospray or FAB mass spectrometry. General laboratory safety is also of primary concern when hazardous materials are involved. Strict adherence to the outlined methods is therefore highly recommended.
Anticipated Results These procedures are suitable for preparation of milligram up to ∼200 mg quantities of the final products. Scale-up is limited by the HPLC purification of the intermediates and final products.
Time Considerations The synthesis of the methylenebis(phosphonate) diesters starting from nucleosides, carbohydrates, or alcohols can be accomplished in 2 to 3 weeks depending on the nature of the nucleoside or other alcohol and whether protecting groups are required. Additional time will be required to remove protecting groups in the final products.
Literature Cited Synthesis of Methylenebis (phosphonate) Analogs of Dinucleotide Pyrophosphates
Bentley, R. 2000. Mycophenolic acid: A one hundred year odyssey from antibiotic to immunosuppressant. Chem. Rev. 100:3801-3826.
Davisson, V.J., Davis, D.R., Dixit, V.M., and Poulter, C.D. 1987. Synthesis of nucleotide 5 -diphosphates from 5 -O-tosyl nucleosides. J. Org. Chem. 52:1794-1801. Franklin, T.J., Jacobs, V.N., Jones, G., Ple, P., and Bruneau, P. 1996. Glucuronidation associated with intrinsic resistance to mycophenolic acid in human colorectal carcinoma cells. Cancer Res. 56:984-987. Gharehbaghi, K., Grunberger, W., and Jayaram, H.M. 2002. Studies on the mechanism of action of benzamide riboside: A novel inhibitor of IMP dehydrogenase. Curr. Med. Chem. 9:743-748. Greene, T.W. and Wuts, P.G.M. 1999. Protective Groups in Organic Synthesis, 3rd ed., pp. 207213. John Wiley & Sons, New York. Grifantini, M. 2000. Tiazofurine ICN pharmaceuticals. Curr. Opin. Investig. Drugs 1:257-262. Jayaram, H.N., Yalowitz, J.A., Arguello, F., and Greene, J.F. Jr. 2002. Toxicity and efficacy of benzamide riboside in cancer chemotherapy models. Curr. Med. Chem. 9:787-792. Kalek, M., Stepinski, J.J., Stolarski, R., and Darzynkiewicz, E. 2005. A direct method for the synthesis of nucleoside 5 methylenebis(phosphonate)s from nucleosides. Tetrahedron Lett. 46:2417-2421. Lesiak, K., Watanabe, K.A., Majumdar, A., Seidman, M., Vanderveen, K., Goldstein, B.M., and Pankiewicz, K.W. 1997. Synthesis of nonhydrolyzable analogues of thiazole-4-carboxamide and benzamide adenine dinucleotide containing fluorine atom at the C2 of adenine nucleoside: Induction of K562 differentiation and inosine monophosphate dehydrogenase inhibitory activity. J. Med. Chem. 40:2533-2538. Lesiak, K., Watanabe, K., George, J. and Pankiewicz, K.W. 1998a. 2-(4-Nitrophenyl)ethyl methylene-bis(phosphonate): A versatile reagent for the synthesis of nucleoside 5 methylene-bis(phosphonate)s. J. Org. Chem. 63:1906-1909. Lesiak, K., Watanabe, K.A., Majumdar, A., Powell, J., Seidman, M., Vanderveen, K., Goldstein, B.M., and Pankiewicz, K.W. 1998b. Synthesis of a methylenebis(phosphonate) analogue of mycophenolic adenine dinucleotide: A glucuronidation-resistant MAD analogue of NAD. J. Med. Chem. 41:618-622. Marquez, V.E., Tseng, C.K., Gebeyehu, G., Cooney, D.A., Ahluwalia, G.S., Kelley, J.A., Dalal, M., Fuller, R.W., Wilson, Y.A., and Johns, D.G. 1986. Thiazole-4-carboxamide adenine dinucleotide (TAD): Analogues stable to phosphodiesterase hydrolysis. J. Med. Chem. 29:1726-1731. Pankiewicz, K.W., Lesiak, K., and Watanabe, K.A. 1997a. Efficient synthesis of methylenebis (phosphonate) analogues of P1 ,P2 -disubstituted pyrophosphates of biological interest: A novel plausible mechanism. J. Am. Chem. Soc. 119:3691-3695.
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Pankiewicz, K.W., Lesiak, K., Zatorski, A., Goldstein, B.M., Carr, S.F., Sochacki, M., Majumdar, A., Seidman, M., and Watanabe, K.A. 1997b. The practical synthesis of a methylenebisphosphonate analogue of benzamide adenine dinucleotide: Inhibition of human inosine monophosphate dehydrogenase (type I and II). J. Med. Chem. 40:1287-1291. Pankiewicz, K.W., Lesiak-Watanabe, K., Watanabe, K.A., and Malinowski, K. 1999. Novel mycophenolic adenine bis(phosphonate)s as potential immunosuppressants. Curr. Med. Chem. 6:629-634. Pankiewicz, K.W., Lesiak-Watanabe, K.B., Watanabe, K.A., Patterson, S.E., Jayaram, H.N., Yalowitz, J.A., Miller, M.D., Seidman, M., Majumdar, A., Prehna, G., and Goldstein, B.M. 2002a. Novel mycophenolic adenine bis(phosphonate) analogues as potential differentiation agents against human leukemia. J. Med. Chem. 45:703-712. Pankiewicz, K.W., Watanabe, K.A., LesiakWatanabe, K., Goldstein, B.M., and Jayaram, H.N. 2002b. The chemistry of nicotinamide adenine dinucleotide (NAD) analogues containing C-nucleosides related to nicotinamide riboside. Curr. Med. Chem. 9:733-741. Patterson, S.E., Mason, C.J., and Pankiewicz, K.W. 2004a. The reaction of alcohols and nucleosides with methylenebis(phosphonic acid dichloride): Facile synthesis of methylenebis(phosphonic acids) monoesters. In Frontiers in Nucleosides and Nucleic Acids (R.F. Schinazi and D.C. Liotta, eds.) pp. 221-226. IHL Press, Arlington, Mass.
Patterson, S.E., Black, P.L., Clark, J.L., Risal, D., Goldstein, B.M., Jayaraman, H.N., Schinazi, R.F., and Pankiewicz, K.W. 2004b. The mechanism of action and antileukemic activity of bis(phosphonate) analogue of mycophenolic adenine dinucleotide (C2-MAD): An alternative for Tiazofurin? In Frontiers in Nucleosides and Nucleic Acids (R.F. Schinazi and D.C. Liotta, eds.) pp. 447-456. IHL Press, Arlington, Mass. Sauve, A.A. and Schramm, V.L. 2004. SIR2: The biochemical mechanism of NAD(+)-dependent protein deacetylation and ADP-ribosyl enzyme intermediates. Curr. Med. Chem. 11:807-826. Shuto, S. and Matsuda, A. 2004. Chemistry of cyclic ADP-ribose and its analogs. Curr. Med. Chem. 11:827-845. Sintchak, M.D., Fleming, M.A., Futer, O., Raybuck, S.A., Chambers, S.P., Caron, P.R., Murcko, M.A., and Wilson, K.P. 1996. Structure and mechanism of inosine monophosphate dehydrogenase in complex with the immunosuppressant mycophenolic acid. Cell 85:921-930.
Contributed by Krzysztof W. Pankiewicz, Guangyao Gao, and Steven E. Patterson The University of Minnesota Center for Drug Design Minneapolis, Minnesota
Nucleoside Phosphorylation and Related Modifications
13.5.13 Current Protocols in Nucleic Acid Chemistry
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Chemical Phosphorylation of Deoxyribonucleosides and Thermolytic DNA Oligonucleotides
UNIT 13.6
This unit provides a detailed preparation of bis[S-(4,4 -dimethoxytrityl)-2-mercaptoethyl]-N,N-diisopropylphosphoramidite (see Basic Protocol 1) and describes the use of this reagent in the solid-phase synthesis of nucleoside 5 -/3 -phosphate/-thiophosphate monoesters (see Basic Protocol 2), oligonucleotide 5 -phosphate/-thiophosphate monoesters (see Alternate Protocol 1), and a model dinucleoside phosphorothioate prodrug functionalized with a heat-sensitive thiophosphate protecting group (see Alternate Protocol 2). This dinucleotide model demonstrates that the phosphorylating reagent enables the preparation of thermosensitive oligonucleotide prodrugs under conditions that will not induce premature thermolytic cleavage of thiophosphate protecting groups to a significant extent. Given that commercial phosphorylating reagents usually require elevated temperature conditions (∼55◦ C) to produce phosphate/thiophosphate monoesters within an acceptable period of time, these conditions are incompatible with the preparation of oligonucleotides functionalized with thermosensitive phosphotriester groups. In this regard, bis[S-(4,4 -dimethoxytrityl)-2-mercaptoethyl]-N,N-diisopropylphosphoramidite is unique in that it generates phosphate/thiophosphate monoester derivatives efficiently and rapidly at 25◦ C. Thus, this unit describes an attractive and general method for the solid-phase phosphorylation of nucleosides and oligonucleotides under mild temperature conditions.
PREPARATION OF BIS[S-(4,4 -DIMETHOXYTRITYL)-2MERCAPTOETHYL-N,N-DIISOPROPYLPHOSPHORAMIDITE
BASIC PROTOCOL 1
Bis[S-(4,4 -dimethoxytrityl)-2-mercaptoethyl]-N,N-diisopropylphosphoramidite (S.4; see Fig. 13.6.1) has been designed specifically to generate heat-sensitive phosphate/ thiophosphate monoesters toward the development of a new class of thermolytic oligonucleotide prodrugs. This protocol outlines a general method for the preparation of bis[S-(4,4 -dimethoxytrityl)-2-mercaptoethyl]-N,N-diisopropylphosphoramidite in three steps from commercial methyl thioglycolate (S.1; see Fig. 13.6.1) and diisopropylphosphoramidous dichloride (Aus´ın et al., 2005).
Materials 4,4 -Dimethoxytrityl chloride (Aldrich) Anhydrous pyridine (Aldrich) Dry argon gas cylinder (Matheson) Methyl thioglycolate (Aldrich) Anhydrous diethyl ether (Fisher) Anhydrous sodium sulfate (Baker) Toluene (Fisher) Anhydrous tetrahydrofuran (Aldrich) Lithium aluminum hydride (LiAlH4 ; Aldrich) Ethyl acetate (EM Science) Celite (Aldrich) Hexane (Fisher) Dichloromethane (CH2 Cl2 , EM Science) Triethylamine (Et3 N, Aldrich)
Contributed by Cristina Aus´ın, Andrzej Grajkowski, Jacek Cie´slak, and Serge L. Beaucage Current Protocols in Nucleic Acid Chemistry (2006) 13.6.1-13.6.21 C 2006 by John Wiley & Sons, Inc. Copyright
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Figure 13.6.1 Preparation of bis[S-(4,4 -dimethoxytrityl)-2-mercaptoethyl]-N,N-diisopropylphosphoramidite. DMTr, 4,4 -dimethoxytrityl. Adapted from Aus´ın et al. (2005) with permission from the American Chemical Society. ◦
Silica gel (60 A, 230 to 400 mesh; EMD) Diisopropylethylamine (DIPEA; Aldrich) Anhydrous acetonitrile (MeCN; Glen Research) Diisopropylphosphoramidous dichloride (Aldrich) Aqueous saturated sodium hydrogen carbonate (NaHCO3 , Aldrich) Benzene (Aldrich) 25-, 50-, 100-, 250-, and 500-mL round-bottom flasks (Kontes) Rubber septa for 14/20- and 24/40-glass joints (Aldrich) 1-, 3-, and 10-mL plastic syringes (B-D) 100- and 250-mL separatory funnels (Kontes) 125-, 250-, and 500-mL Erlenmeyer flasks (Kimax) 100-mm funnels (Nalgene) Filter paper, no. 1 (Whatman) Rotary evaporator (B¨uchi) connected to a vacuum pump (KNF) 150-mL sintered-glass funnel (coarse porosity; Kontes) 500-mL vacuum Erlenmeyer flask Water aspirator 2.5 × 20–cm disposable Flex chromatography columns (Kontes) 2.5 × 7.5–cm TLC plates precoated with a 250-µm layer of silica gel 60 F254 (EMD) Vacuum desiccator High vacuum oil pump (Savant) Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) Prepare S-DMTr-2-mercaptoethanol S.1 1. Dissolve 20 mmol of 4,4 -dimethoxytrityl chloride in 40 mL anhydrous pyridine in a 250-mL round-bottom flask sealed with a rubber septum. 2. Cool the solution to ∼10◦ C in an ice bath.
Chemical Phosphorylation of Deoxyribonucleosides and Thermolytic DNA Oligonucleotides
3. While stirring the solution with a magnetic stirrer under a positive pressure of argon, add 18 mmol methyl thioglycolate using a 3-mL syringe. 4. Allow the reaction mixture to warm up to room temperature (∼25◦ C) and stir the solution for 14 hr.
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5. Dilute the reaction mixture by adding 150 mL diethyl ether. Add 20 mL water and transfer the two layers to a 250-mL separatory funnel. 6. Discard the aqueous layer and transfer the organic layer to a 250-mL Erlenmeyer flask. Add ∼10 g anhydrous sodium sulfate. Stir the suspension for 5 min and allow to stand for 10 min. 7. Remove the drying agent by gravity filtration through a 100-mm funnel fitted with a Whatman no. 1 filter paper. Wash the solid with 20 mL diethyl ether. Pool the filtrates in a 500-mL round-bottom flask and remove the solvent using a rotary evaporator connected to a vacuum pump. 8. Dissolve the material in 10 mL toluene and transfer the solution to a 100-mL roundbottom flask. Concentrate the solution under reduced pressure in a rotary evaporator (∼20 mmHg) and dry the crude material by coevaporation three times with 50 mL toluene. 9. Dissolve the solid in 40 mL anhydrous tetrahydrofuran. 10. Cool the solution to ∼5◦ C in an ice bath. 11. Add 21.6 mmol LiAlH4 in eight ∼100-mg portions. CAUTION: LiAlH4 reacts violently with water and liberates hydrogen, a very flammable gas. Keep the container tightly closed in a dry location.
12. Allow the reaction mixture to warm to room temperature and stir for 2 hr using a magnetic stirrer. 13. Quench the reaction by adding 10 mL ethyl acetate. Stir for 15 min and transfer the suspension to a 500-mL Erlenmeyer flask. 14. Add 100 mL diethyl ether followed by slow addition of 80 mL water over a period of 10 min. Stir for 1 hr. 15. Filter the suspension over a 2.5-cm-thick pad of Celite packed into a 150-mL sinteredglass funnel using a 500-mL vacuum Erlenmeyer flask connected to a water aspirator. Wash the Celite cake with 50 mL diethyl ether. 16. Transfer the filtrates to a 250-mL separatory funnel. 17. Decant the organic layer into a 500-mL Erlenmeyer flask and add ∼10 g anhydrous sodium sulfate. Stir the suspension for 5 min and allow to settle for 10 min. 18. Remove the drying agent by gravity filtration through a 100-mm funnel fitted with Whatman no. 1 filter paper. Wash the solid with 50 mL diethyl ether, combine the filtrates in a 500-mL round-bottom flask, and remove solvent using a rotary evaporator connected to a vacuum pump. 19. Dissolve the crude product in ∼5 mL of 6:3:1 (v/v/v) hexane/CH2 Cl2 /Et3 N and add the solution to a 2.5 × 20–cm disposable Flex chromatography column (APPENDIX 3E) containing ∼40 g silica gel that has been equilibrated in 6:3:1 (v/v/v) hexane/CH2 Cl2 /Et3 N. 20. Elute the column with 6:3:1 (v/v/v) hexane/CH2 Cl2 /Et3 N and collect 8-mL fractions. 21. Analyze fractions by TLC (APPENDIX 3D) on a 2.5 × 7.5–cm silica gel 60 F254 TLC plate using 6:3:1 (v/v/v) hexane/CH2 Cl2 /Et3 N as the eluent. Pool appropriate fractions in a 250-mL round-bottom flask and concentrate under reduced pressure (∼20 mmHg) until a light yellow oil is obtained in 90% yield (6.15 g, 16.2 mmol).
Nucleoside Phosphorylation and Related Modifications
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Supplement 26
TLC analysis of S.1 reveals one UV-visible spot exhibiting an Rf of 0.4 when 6:3:1 (v/v/v) hexane/CH2 Cl2 /Et3 N is used as the eluent. The spot is orange-colored when the TLC plate is heated on a hot plate. The yield of S.1 is similar to that reported in the literature (Rademann and Schmidt, 1997) for a similar compound. S-(4,4 -Dimethoxytrityl)-2-mercaptoethanol (S.1): 1 H NMR (300 MHz, DMSO-d6 ): δ 7.36-7.15 (m, 5H), 7.24 (d, J = 9.0 Hz, 4H), 6.87 (d, J = 9.0 Hz, 4H), 4.75 (bs, 1H), 3.72 (s, 6H), 3.33 (t, J = 6.9 Hz, 2H), 2.23 (t, J = 6.9 Hz, 2H). 13 C NMR (75 MHz, DMSO-d6 ): δ 157.5, 145.3, 136.7, 130.2, 128.8, 127.7, 126.3, 113.0, 64.8, 59.5, 54.8, 34.6.
Synthesize phosphorylating reagent S.4 22. Dissolve 3.10 mmol S.1 and 30.9 mmol diisopropylethylamine in 10 mL anhydrous MeCN in a 50-mL round-bottom flask sealed with a rubber septum. Stir the solution using a magnetic stirrer. 23. Under a positive pressure of argon, add 1.40 mmol diisopropylphosphoramidous dichloride dropwise over a 2-min period using a 1-mL syringe. The progress of the reaction is determined by monitoring the disappearance of the 31 P NMR signal corresponding to diisopropylphosphoramidous dichloride (δ P = 170 ppm) and the appearance of a new signal at 148 ppm. The phosphinylation of S.1 is complete within 2 hr at ∼25◦ C.
24. Dilute the reaction mixture by adding 30 mL CH2 Cl2 and transfer it to a 100-mL separatory funnel. 25. Wash the organic phase once with 30 mL water and then twice with 15 mL of an aqueous saturated solution of NaHCO3 . 26. Decant the organic layer into a 125-mL Erlenmeyer flask and add ∼10 g anhydrous sodium sulfate. Stir the suspension for 5 min and allow to settle for 10 min. 27. Remove the drying agent by gravity filtration through a funnel fitted with Whatman no. 1 filter paper. Wash the solid with 20 mL CH2 Cl2 , pool the filtrates in a 250-mL round-bottom flask, and remove the solvent using a rotary evaporator connected to a vacuum pump. 28. Dissolve the crude product in ∼5 mL of 9:1 (v/v) benzene/triethylamine and add the solution to a 2.5 × 20–cm disposable Flex chromatography column containing ∼40 g silica gel that has been equilibrated in 9:1 (v/v) benzene/triethylamine. 29. Elute the column with 9:1 (v/v) benzene/triethylamine and collect 8-mL fractions. 30. Analyze fractions by TLC on a 2.5 × 7.5–cm silica gel 60 F254 TLC plate using 9:1 (v/v) benzene/triethylamine as the eluent. Pool appropriate fractions in a 250-mL round-bottom flask and concentrate under reduced pressure (∼20 mmHg) until a yellow oil is obtained. TLC analysis of S.4 reveals one UV-visible spot exhibiting an Rf of 0.2 when 9:1 (v/v) benzene/triethylamine is used as the eluent. The spot is orange-colored when the TLC plate is heated on a hot plate.
31. Dissolve the oily material in ∼3 mL benzene and add the solution to ∼100 mL of vigorously stirring cold (−20◦ C) hexane in a 250-mL round-bottom flask. 32. Allow the suspension to settle and carefully decant off most of the supernatant. Chemical Phosphorylation of Deoxyribonucleosides and Thermolytic DNA Oligonucleotides
33. Evaporate the wet material to dryness under reduced pressure using a rotary evaporator. 34. Expose the oily product to high vacuum (1 mmHg) for ∼15 min and then dissolve it in ∼5 mL benzene.
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35. Transfer the solution to a 25-mL round-bottom flask and freeze it in a dry-ice/acetone bath. 36. Lyophilize the frozen solution under high vacuum to produce 1.02 g (1.15 mmol) triethylamine-free S.4 as a light yellow powder in 82% yield relative to diisopropylphosphoramidous dichloride. Bis[S-(4,4 -dimethoxytrityl)-2-mercaptoethyl]-N,N-diisopropylphosphoramidite (S.4): 1 H NMR (300 MHz, DMSO-d6 ): δ 7.38-7.16 (m, 10H), 7.19 (d, J = 7.9 Hz, 8H), 6.85 (d, J = 7.9 Hz, 8H), 3.71 (s, 12H), 3.36 (m, 6H), 2.30 (t, J = 6.6 Hz, 4H), 1.02 (d, J = 6.7 Hz, 12H). 13 C NMR (75 MHz, DMSO-d6 ): δ 157.5, 145.0, 136.5, 130.1, 128.7, 127.7, 126.4, 113.0, 65.0, 61.3, (d, 2 JPC = 17.9 Hz), 54.9, 42.3, 42.1, 32.9 (d, JPC = 7.2 Hz), 24.2, 24.1. 31 P NMR (121 MHz, DMSO-d6 ): δ 147.7. APESI-HRMS: calcd. for C52 H60 NNaO6 PS2 (M+Na)+ 912.3497, found 912.3493.
SOLID-PHASE SYNTHESIS OF 2 -DEOXYRIBONUCLEOSIDE 5 -/3 -PHOSPHATE OR -THIOPHOSPHATE MONOESTERS
BASIC PROTOCOL 2
Manual solid-phase synthesis of 2 -deoxyribonucleoside 5 -/3 -phosphate or -thiophosphate monoesters is performed by acid-mediated cleavage of the DMTr group from succinyl long-chain alkylamine controlled-pore glass support functionalized with a leader 5 - or 3 -O-DMTr-2 -deoxyribonucleoside followed by treatment of the free 5 -/3 -hydroxyl with phosphoramidite S.4 in the presence of 1Htetrazole. Following oxidation/sulfurization of the phosphite triester, the corresponding phosphate/thiophophosphate triester is exposed to acidic conditions to remove the SDMTr groups, and then to DL-dithiothreitol to maintain the resulting free thiols in a reduced state during production of the 5 -/3 -phosphate or -thiophosphate monoester. 2 Deoxyribonucleoside 5 -/3 -phosphate or -thiophosphate monoesters are released from the support by treatment with pressurized ammonia gas or concentrated ammonium hydroxide at an elevated temperature as outlined in Figure 13.6.2.
Materials Trichloroacetic acid (TCA; Aldrich) Dichloromethane (CH2 Cl2 , EM Science) Synthesis columns filled with succinyl long-chain alkylamine controlled-pore glass ◦ (Succ-LCAA-CPG, 500 A) support functionalized with a leader 5 -O-DMTr- or 3 -O-DMTr-2 -deoxyribonucleoside (T, CBz , ABz and Gi-Bu , 0.2 µmol; Glen Research) Acetonitrile (MeCN; Acros) Dry argon gas cylinder (Matheson) 0.45 M 1H-tetrazole in MeCN (Glen Research) 0.1 M S.4 (see Basic Protocol 1) in dry MeCN Oxidant: ∼32% wt. ethyl(methyl)dioxirane (2-butanone peroxide) in dimethyl phthalate (Aldrich; for 5 -/3 -phosphate monoesters) 0.05 M 3H-1,2-benzodithiol-3-one-1,1-dioxide in MeCN (for 5 -/3 -thiophosphate monoesters) DL-Dithiothreitol (DTT; Aldrich) Triethylamine (Et3 N; Aldrich) Concentrated ammonium hydroxide (NH4 OH; Fisher Scientific) 2 M triethylammonium acetate (TEAA) buffer, pH 7.0 (Applied Biosystems) 1-, 3-, and 10-mL syringes with Luer tips 20-G hypodermic needles with Luer-tip adapters 250-mL vacuum Erlenmeyer flasks with rubber septa Water aspirator Tygon tubing (Fisher)
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Figure 13.6.2 Solid-phase synthesis of 2 -deoxyribonucleoside-5 -phosphate/thiophosphate (A) or -3 phosphate/thiophosphate (B) monoesters. B: thymin-1-yl (a), cytosin-1-yl (b), adenin-9-yl (c), guanin-9-yl (d); BP : thymin-1-yl (a), 4-N-benzoylcytosin-1-yl (b), 6-N-benzoyladenin-9-yl (c), 2-N-isobutyrylguanin-9-yl (d); CPG-LCAA-Succ, succinyl long-chain alkylamine controlled-pore glass; DMTr, 4,4 -dimethoxytrityl; TCA, trichloroacetic acid. Adapted from Aus´ın et al. (2005) with permission from the American Chemical Society.
4-mL screw-capped glass vials (Wheaton), one with rubber septum 1-mL gas-tight glass syringe (Hamilton) 55◦ C heating block 5-µm Supelcosil LC-18S reversed-phase HPLC column (25 cm × 4.6 mm; Supelco)
Chemical Phosphorylation of Deoxyribonucleosides and Thermolytic DNA Oligonucleotides
Prepare 5 -/3 -phosphate and thiophosphate monoesters 1. Connect a Luer-tipped 3-mL syringe filled with 1.5 mL of 3% TCA in CH2 Cl2 to a synthesis column containing Succ-LCAA-CPG functionalized with 0.2 µmol 5 -/3 -O-DMTrT (S.5a/S.12a) or one of the N-protected 5 - or 3 -O-DMTr-2 deoxyribonucleosides S.5b-d/S.12b-d (Fig. 13.6.2). Push the liquid through the column over a 3-min period. Expel excess liquid by allowing it to drip off the column. 2. Connect a 20-G hypodermic needle to one end of the synthesis column using a Luer-tipped adapter and puncture the rubber septum capping a 250-mL vacuum Erlenmeyer flask attached to a water aspirator. Apply vacuum to aspirate residual acid, and then wash two times with 10 mL MeCN dispensed from a 10-mL syringe.
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3. Air dry the support using the water aspirator. Remove the synthesis column from the vacuum Erlenmeyer flask and flush the support with argon for 1 min through a 1-mL plastic syringe connected to the argon gas cylinder via Tygon tubing. 4. Add, by syringe, 0.3 mL of 0.45 M 1H-tetrazole in MeCN to 30 µmol of S.4 in a 4-mL vial that has been flushed with argon and capped with a rubber septum. 5. Connect one end of the synthesis column to a 1-mL gas-tight glass syringe and the other end to a 20-G hypodermic needle using a Luer-tipped adapter. 6. Draw the solution of activated S.4 through the synthesis column using the syringe. Agitate the support suspended in the solution for 3 min. 7. Expel excess activated S.4 and wash the support two times with 10 mL of MeCN as described in step 2. 8. Prepare a solution of 0.1 M ethyl(methyl)dioxirane in CH2 Cl2 and push 1 mL of the solution back and forth through the synthesis column over a 1-min period using a 1-mL syringe. For 5 -/3 -thiophosphate monoesters, push 1 mL of 0.05 M 3H-1,2-benzodithiol-3-one1,1-dioxide in MeCN through the synthesis column over a 2-min period using a 1-mL syringe.
9. Expel excess oxidant and wash the support two times with 10 mL CH2 Cl2 as described in step 2. 10. Push 10 mL of 3% TCA in CH2 Cl2 through the synthesis column over a 15-min period using a 10-mL syringe. 11. Expel excess acid and wash the support two times with 10 mL MeCN as described in step 2. 12. Make a solution of 1.2% (w/v) DTT and 5% (v/v) Et3 N in 3 mL water. Push all the solution back and forth through the synthesis column over a 60-min period using a 3-mL syringe. 13. Expel excess reagents and wash the support first with 10 mL water and then two times with 10 mL MeCN as described in step 2.
Release from support and deprotect nucleobases 14. Transfer the solid support of each synthesis column to individual 4-mL glass vials. 15. Pipet 2 mL conc. NH4 OH into each vial and leave the suspension standing 30 min at 25◦ C. 16. Pipet each ammoniacal solution into individual 4-mL glass vials and place the tightly capped containers in a heating block. Heat the solutions to 55◦ C for 10 hr. Allow the solutions to cool to room temperature. Solutions of the thymine derivatives (S.10a, S.11a, S.13a, and S.14a) do not require heating. Proceed to step 17.
17. Evaporate each ammoniacal solution using a stream of air. 18. Dissolve each crude 2 -deoxyribonucleoside 5 -/3 -phosphate monoester in 500 µL of 0.1 M TEAA buffer, pH 7.0, for analysis. Alternatively, release of the 2 -deoxyribonucleoside 5 -/3 -phosphate monoesters from the support and nucleobase deprotection can be achieved in one step by placing each synthesis column in a pressure vessel connected to an ammonia gas cylinder. The vessel is pressurized to ∼10 bar for 10 hr at 25◦ C. Upon release of the internal pressure, each monoester is eluted off the synthesis column with 500 µL of 0.1 M TEAA, pH 7.0, using a 1-mL syringe.
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Table 13.6.1 RP-HPLC Retention Times (tR ) of Crude 2 -Deoxyribonucleoside-5 -/3 Phosphate and 5 -/3 -Thiophosphate Monoestersa
Phosphates Compound
Thiophosphates tR (min)
Compound
tR (min)
S.10a
11.5
S.11a
12.3
S.10b
9.2
S.11b
10.2
S.10c
12.9
S.11c
13.4
S.10d
11.6
S.11d
12.1
S.13a
12.8
S.14a
13.7
S.13b
10.3
S.14b
11.3
S.13c
13.1
S.14c
13.9
S.13d
11.5
S.14d
12.4
a Chromatographic conditions are described in Basic Protocol 2. Adapted from Aus´ın et al. (2005) with
permission from the American Chemical Society.
Analyze crude monoesters by RP-HPLC 19. Inject 100 µL of the crude monoester solution through a 5-µm Supelcosil LC-18S HPLC column (4.6 mm × 25 cm). Pump a linear gradient of 1% MeCN/min starting from 0.1 M TEAA buffer, pH 7.0, at a flow rate of 1 mL/min for 40 min. The chromatographic profile of each 2 -deoxyribonucleoside 5 -/3 -phosphate monoester is compared with that of an authentic commercial sample. Table 13.6.1 lists the RP-HPLC retention times of S.10a-d and S.13a-d, which were identical to those of the commercial reference samples. RP-HPLC profiles of a representative 2 deoxyribonucleoside 5 -phosphate monoester and 2 -deoxyribonucleoside 3 -phosphate monoester are available in charts 2 and 6, respectively, in Aus´ın et al. (2005). Table 13.6.1 also provides the retention times of S.11a-d and S.14a-d. The identities of these compounds were directly derived from those of S.10a-d and S.13a-d. With the exception of a sulfurization step, which replaces step 8, the preparation of S.11a-d and S.14a-d is identical to that of S.10a-d and S.13a-d. RP-HPLC profiles of a representative 2 -deoxyribonucleoside 5 -thiophosphate monoester and 2 -deoxyribonucleoside 3 -thiophosphate monoester are available in charts 4 and 8, respectively, in Aus´ın et al. (2005). ALTERNATE PROTOCOL 1
SOLID-PHASE SYNTHESIS OF OLIGONUCLEOTIDE 5 -PHOSPHATE/THIOPHOSPHATE MONOESTERS The versatility of the phosphorylating reagent S.4 is demonstrated in the preparation of oligonucleotide 5 -phosphate/thiophosphate monoesters S.17/S.18 (see Fig. 13.6.3). Following synthesis of the oligonucleotide by standard solid-phase methods, phosphorylation is performed as described in Basic Protocol 2.
Additional Materials (also see Basic Protocol 2)
Chemical Phosphorylation of Deoxyribonucleosides and Thermolytic DNA Oligonucleotides
Reagents recommended for manual and automated solid-phase oligonucleotide synthesis including: Synthesis columns filled with succinyl long-chain alkylamine controlled-pore glass (Succ-LCAA-CPG) support functionalized with 0.2 µmol 5 -O-DMTrN4 -benzoyl-2 -deoxycytidine (Glen Research) Standard 5 -O-DMTr-2 -deoxyribonucleoside-3 -O-(2-cyanoethyl-N,Ndiisopropyl) phosphoramidites (T, CBz , ABz and Gi-Bu ; Glen Research) Activator solution: 1H-tetrazole in acetonitrile (Glen Research) Oxidation solution: 0.02 M iodine in THF/pyridine/water (Glen Research) Cap A solution: acetic anhydride in THF/pyridine (Glen Research)
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Figure 13.6.3 For solid-phase synthesis of oligonucleotide 5 -phosphate/thiophosphate monoesters, standard oligonucleotide synthesis is followed by phosphorylation as in Basic Protocol 2. B: thymin-1-yl (a), cytosin-1-yl (b), adenin-9-yl (c), guanin-9-yl (d); BP : thymin-1-yl (a), 4-N-benzoylcytosin-1-yl (b), 6-N-benzoyladenin-9-yl (c), 2-N-isobutyrylguanin-9-yl (d); CPG-LCAA-Succ, succinyl long-chain alkylamine controlled-pore glass; DMTr, 4,4 -dimethoxytrityl. Adapted from Aus´ın et al. (2005) with permission from the American Chemical Society.
Cap B solution: 1-methylimidazole in THF (Glen Research) Deblocking solution: trichloroacetic acid in dichloromethane (Glen Research) 1 M Tris·Cl buffer, pH 9.0 (APPENDIX 2A) 1.0 M magnesium chloride (MgCl2 ; Sigma) Bacterial alkaline phosphatase (E. coli; Sigma) DNA/RNA synthesizer (Applied Biosystems 392 DNA/RNA synthesizer) 1.5-mL microcentrifuge tubes 37◦ C water bath 90◦ C heating block Additional reagents and equipment for automated DNA synthesis (APPENDIX 3C), HPLC (UNIT 10.5), and MALDI-TOF-MS (UNIT 10.1) Synthesize oligonucleotide 5 -phosphate or 5 -thiophosphate monoester For oligonucleotide 5 -phosphate monoester S.17 1a. Perform two 0.2-µmol-scale solid-phase syntheses of d(AP TP CP CP GP TP AP GP CP TP AP AP GP GP TP CP AP TP GP C) using an Applied Biosystems 392 DNA/RNA synthesizer in trityl-off mode according to the manufacturer’s recommendations (also see APPENDIX 3C). Modify the synthetic cycle by extending each of the wait times for the coupling, oxidation, and capping steps to 1 min. All ancillary reagents required for the automated preparation of oligonucleotides were purchased from Glen Research and used as recommended by the manufacturers.
2a. Remove the synthesis columns from the synthesizer. With one synthesis column, repeat Basic Protocol 2, steps 3 through 7 and then steps 3 to 18 to give S.17. With the other column, repeat steps 14 to 18 to produce a crude oligonucleotide with a free 5 -hydroxyl group. 3a. To analyze the crude products, inject 25 µL of the oligonucleotide solution through a 5-µm Supelcosil LC-18S HPLC column (UNIT 10.5). Pump a linear gradient of 1% MeCN/min starting from 0.1 M TEAA buffer, pH 7.0, at a flow rate of 1 mL/min for 40 min. The chromatographic profile of the oligonucleotide 5 -phosphate monoester S.17 is compared with that of its 5 -unphosphorylated congener to assess the coupling efficiency of the phosphorylating reagent S.4 (chart 12, Aus´ın et al., 2005). In addition, both oligonucleotides were characterized by MALDI-TOF-MS (UNIT 10.1) in the negative ion mode.
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P d(AP TP CP CP GP TP AP GP CP TP AP AP GP GP TP CP AP TP GP C) calcd. for C195 H245 N75 O121 P20 (M−H)− 6195, found 6199.
(S.17): MALDI-TOF-MS:
d(AP TP CP CP GP TP AP GP CP TP AP AP GP GP TP CP AP TP GP C): MALDI-TOF-MS: calcd. for C195 H245 N75 O118 P19 (M−H)− 6116, found 6120.
For oligonucleotide 5 -thiophosphate monoester S.18 1b. Perform a 0.2-µmol-scale solid-phase synthesis of PS d(APS TPS CPS CPS GPS TPS APS GPS CPS TPS APS APS GPS GPS TPS CPS APS TPS GPS C) (S.18) using an Applied Biosystems 392 DNA/RNA synthesizer in trityl-on mode according to the manufacturer’s recommendations (also see APPENDIX 3C). Also perform a 0.2-µmol-scale solidphase synthesis of d(APS TPS CPS CPS GPS TPS APS GPS CPS TPS APS APS GPS GPS TPS CPS APS TPS GPS C) in the trityl-off mode. Modify the standard synthesis cycle as follows: a. Perform the capping step after the oxidative sulfurization reaction (Iyer et al., 1990). b. For the sulfurization steps, use 0.05 M 3H-1,2-benzodithiol-3-one-1,1-dioxide in MeCN. c. For the terminal 5 -phosphorylation reaction, use 0.1 M S.4 in dry MeCN. Connect the vial to the synthesizer through port no. 5 and identify the reagent as no. 5 in the synthetic sequence. d. Extend each wait time for the coupling, sulfurization, and capping steps to 1 min. Modification c does not apply to the synthesis of d(APS TPS CPS CPS GPS TPS APS GPS CPS TPS APS APS GPS GPS TPS CPS APS TPS GPS C). All ancillary reagents required for the automated preparation of oligonucleotides were purchased from Applied Biosystems and/or Glen Research, and used as recommended by the manufacturers.
2b. Remove the synthesis columns from the synthesizer. Repeat Basic Protocol 2, steps 10 to 18, to give S.18, or steps 14 to 18, to give the 5 -unphosphorylated oligonucleoside phosphorothioate. 3b. Analyze as in step 3a. The chromatographic profile of S.18 is compared with that of its 5 -unphosphorylated congener to further assess the coupling efficiency of the phosphorylating reagent S.4 (chart 14, Aus´ın et al., 2005). In addition, both oligonucleotides were characterized by MALDI-TOF-MS (UNIT 10.1) in the negative ion mode. PS d(APS TPS CPS CPS GPS TPS APS GPS CPS TPS APS APS GPS GPS TPS CPS APS TPS GPS C) (S.18): MALDI-TOF-MS: calcd. for C195 H245 N75 O101 P20 S20 (M−H)− 6516, found 6523.
d(APS TPS CPS CPS GPS TPS APS GPS CPS TPS APS APS GPS GPS TPS CPS APS TPS GPS C): MALDITOF-MS: calcd. for C195 H245 N75 O99 P19 S19 (M−H)− 6421, found 6422.
Convert S.17 to its 5 -unphosphorylated congener 4. To 1 OD260 of crude lyophilized S.17 in a 1.5-mL microcentrifuge tube, add: 6 µL of 1 M Tris·Cl buffer, pH 9.0 8 µL of 1.0 M MgCl2 80 µL ddH2 O 6 µL (0.7 U) bacterial alkaline phosphatase. Chemical Phosphorylation of Deoxyribonucleosides and Thermolytic DNA Oligonucleotides
Incubate 4 hr at 37◦ C. 5. Heat the digest 2 min at 90◦ C and microcentrifuge 5 min at 16,000 × g, 25◦ C.
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6. Analyze 50 µL of the centrifuged digest by HPLC as in step 3a. The chromatographic profile of the oligonucleotide 5 -phosphate monoester treated with bacterial alkaline phosphatase shows complete 5 -dephosphorylation (chart 13 in Aus´ın et al., 2005). These results, along with the MS data, further confirm the identity of oligonucleotide 5 -phosphate monoester S.17.
SYNTHESIS OF A DINUCLEOTIDE WITH A THERMOLYTIC 4-THIOPHOSPHATO-1-BUTYL THIOPHOSPHATE-PROTECTING GROUP
ALTERNATE PROTOCOL 2
The dinucleotide S.22 serves as a model to precisely assess the thermosensitivity properties of selected phosphate/thiophosphate protecting groups and their relevance toward the development of thermolytic oligonucleotide prodrugs. As shown in Figure 13.6.4, the preparation of dinucleotide S.22 is challenging in that the generation of a thiophosphate monoester will impart thermosentitive properties to the dinucleoside thiophosphateprotecting group. It is therefore critically important that formation of the thiophosphate monoester be achieved under conditions that will not induce premature thermolytic cleavage of the dinucleoside thiophosphate-protecting group. The phosphorylating reagent S.4 has been designed specifically for this purpose since commercially available phosphorylating reagents are not compatible with the conditions required for the preparation of such a specific class of thermolytic phosphate/thiophosphate-protecting groups.
Additional Materials (also see Basic Protocol 2) Deoxyribonucleoside phosphoramidite S.19 (see Support Protocol) Acetic acid (Aldrich) Anhydrous pyridine (Aldrich) Hydrazine monohydrate (NH2 NH2 ·H2 O; Aldrich) Ammonia gas cylinder (Aldrich) Pressure vessel 90◦ C heating block Additional reagents and equipment for HPLC (UNIT 10.5) and MALDI-TOF-MS (UNIT 10.1) Prepare thermolytic dinucleotide S.22 1. Connect a Luer-tipped 3-mL syringe filled with 1.5 mL of 3% TCA in CH2 Cl2 to a synthesis column containing Succ-LCAA-CPG functionalized with 0.2 µmol 5 -O-DMTrT (S.5a). Push the liquid through the column over a period of 3 min. 2. Perform Basic Protocol 2, steps 2 and 3. 3. Using a syringe, add 0.3 mL of 0.45 M 1-H-tetrazole in MeCN to 30 µmol of S.19 in a 4-mL vial that has been flushed with argon and capped with a rubber septum. 4. Connect one end of the synthesis column to a 1-mL gas-tight glass syringe and the other end to a 20-G hypodermic needle. 5. Draw the solution of activated S.19 through the synthesis column using the syringe attached to it. Agitate the support suspended in the solution for a period of 3 min. 6. Expel excess activated S.19 and wash the support two times with 10 mL MeCN as described in Basic Protocol 2, step 2. 7. Push 1 mL of 0.05 M 3H-1,2-benzodithiol-3-one-1,1-dioxide in MeCN through the synthesis column over a 2-min period using a 1-mL syringe. 8. Expel excess oxidant and wash the support two times with 10 mL MeCN as described in Basic Protocol 2, step 2.
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Figure 13.6.4 Solid-phase synthesis of dinucleotide S.22 as a thermolytic oligonucleotide prodrug model. DMTr, 4,4 dimethoxytrityl; DTT, DL-dithiothreitol; Lev, levulinyl; TCA, trichloroacetic acid; Thy, thymin-1-yl. Adapted from Aus´ın et al. (2005) with permission from the American Chemical Society.
9. Push 3 mL of 0.5 M NH2 NH2 ·H2 O in 3:2 (v/v) pyridine/acetic acid back and forth through the synthesis column over a 10-min period at 25◦ C. 10. Expel excess reagent and wash the support two times with 10 mL CH2 Cl2 and two times with 10 mL MeCN. 11. Perform Basic Protocol 2, steps 3 to 7. 12. Repeat treatment with oxidant as in steps 7 and 8 of this protocol. 13. Perform Basic Protocol 2, steps 10 to 13. 14. Place the synthesis column in a pressure vessel connected to an ammonia gas cylinder. Pressurize the vessel to ∼10 bar for a period of 30 min. 15. Upon release of the internal pressure of the vessel, elute the crude dinucleotide S.22 off the synthesis column with 500 µL of 0.1 M TEAA, pH 7.0, using a 1-mL syringe. 16. Characterize by 31 P NMR spectroscopy and by MALDI-TOF-MS (UNIT 10.1). P NMR (121 MHz, H2 O): δ 67.1, 66.9, 51.9. −MALDI-TOF-MS: calcd. for C24 H35 N4 O14 P2 S2 (M−H)− 729.6, found 729.7.
31
Chemical Phosphorylation of Deoxyribonucleosides and Thermolytic DNA Oligonucleotides
Convert to dinucleoside phosphorothioate TPS T 17. Heat the solution of crude S.22 in a tightly capped 4-mL glass vial for 1 hr at 90◦ C using a heating block. Allow the solution to cool to room temperature.
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Figure 13.6.5 RP-HPLC analysis of crude dinucleotide S.22 and its thermal conversion to the dinucleoside phosphorothioate diester TPS T. (A) Crude S.22 prepared and deprotected as in Alternate Protocol 2. (B) TPS T produced by heating S.22 in 0.1 M TEAA (pH 7.0) for 1 hr at 90◦ C. (C) Dinucleoside phosphotriester S.20 after removal of DMTr and levulinyl groups and release from the support. Chromatographic conditions described in Alternate Protocol 2, step 18. Adapted from Aus´ın et al. (2005) with permission from the American Chemical Society.
18. To analyze the conversion, inject 50 µL of the dinucleotide solution through a 5-µm Supelcosil LC-18S HPLC column. Pump a linear gradient of 1% MeCN/min starting from 0.1 M TEAA, pH 7.0, at a flow rate of 1 mL/min for 40 min. Chromatographic profiles of the crude dinucleotide S.22 and its thermolytic conversion to TPS T are shown in Figure 13.6.5.
PREPARATION OF DEOXYRIBONUCLEOSIDE PHOSPHORAMIDITE S.19 The synthesis of S.19 begins with the preparation of the phosphorodiamidite S.23 (Fig. 13.6.6), which was prepared from the reaction of commercial bis(N,Ndiisopropylamino)chlorophosphine and 4-hydroxy-1-butyl levulinate (Tsukamoto and Kondo, 2003). The reaction of S.23 with 5 -O-DMTr-2 -deoxythymidine in the presence of 1H-tetrazole gives S.19, which is purified by silica gel chromatography.
SUPPORT PROTOCOL
Additional Materials (also see Basic Protocol 1) Bis(N,N-diisopropylamino)chlorophosphine (Aldrich) 4-Hydroxy-1-butyl levulinate (Tsukamoto and Kondo, 2003) 5% (w/v) silver nitrate in 1:1 (v/v) EtOH/H2 O 5 -O-(4,4 -Dimethoxytrityl)-2 -deoxythymidine (Chem-Impex International) Sublimed 1H-tetrazole (Glen Research) Dry-ice/acetone bath 250-mL vacuum Erlenmeyer flask 7 × 15–cm glass chromatography column (Ace Glass) Cotton swab Hot plate 1- and 10-mL glass syringe 16-G needle Lyophilizer Prepare phosphorodiamidite S.23 1. Add 4.05 g (15.2 mmol) bis(N,N-diisopropylamino)chlorophosphine in ∼1-g portions over a 30-min period, to a magnetically stirred solution of 2.82 g (15.0 mmol) 4-hydroxy-1-butyl levulinate, 12.4 g (95.9 mmol) DIPEA, and 30 mL dry CH2 Cl2 in a 100-mL round-bottom flask capped with a rubber septum.
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Figure 13.6.6 Synthesis of the deoxyribonucleoside phosphoramidite S.19. DMTr, 4,4 dimethoxytrityl; Lev, levulinyl; Thy, thymin-1-yl.
2. Monitor the progress of the reaction by 31 P NMR spectroscopy. The 31 P NMR signal corresponding to bis(N,N-diisopropylamino)chlorophosphine (δ P 135.5) disappears completely and a new signal corresponding to S.23 appears (δ P 122).
3. Filter the suspension through a sintered-glass funnel of coarse porosity using a 250-mL vacuum Erlenmeyer flask and a water aspirator. Wash the solid with 30 mL benzene. 4. Pool the filtrates in a 250-mL round-bottom flask and remove the solvent using a rotary evaporator connected to a vacuum pump. 5. Suspend the crude phosphorodiamidite in ∼5 mL of 9:1 (v/v) benzene/triethylamine and apply the suspension to the top of a 7 × 15–cm glass column packed with ∼100 g of silica gel 60 that has been equilibrated in 9:1 (v/v) benzene/triethylamine. 6. Elute the column with 9:1 (v/v) benzene/triethylamine and collect 8-mL fractions. 7. Identify the fractions containing S.23 by silica gel TLC using 9:1 (v/v) benzene/ triethylamine as the eluent. Visualize the product by spreading a 5% (w/v) solution of silver nitrate in 1:1 (v/v) EtOH/H2 O over the surface of the TLC plate with a cotton swab, and heating the TLC plate on a hot plate until a black spot appears (Rf = 0.62). 8. Pool the appropriate fractions in a 250-mL round-bottom flask and evaporate the solvents under reduced pressure using a rotary evaporator to afford 4.50 g (10.8 mmol) S.23 as an oil in 72% yield. H NMR (300 MHz, C6 D6 ): δ 4.04 (t, J = 6.4 Hz, 2H), 3.54 (dt, J = 5.9 Hz, JHP = 7.4 Hz, 2H), 3.53 (sept, J = 6.8 Hz, 2H), 3.49 (sept, J = 6.8 Hz, 2H), 2.35 (t, J = 6.5 Hz, 2H), 2.15 (t, J = 6.5 Hz, 2H), 1.61 (s, 3H), 1.60 (m, 4H), 1.23 (d, J = 6.8 Hz, 12H), 1.19 (d, J = 6.8 Hz, 12H). 13 C NMR (75 MHz, C6 D6 ): 204.5, 172.3, 64.5, 64.2 (d, 2 JCP = 21.5 Hz), 44.7, 44.6, 37.6, 29.1, 28.4 (d, 3 JCP = 8.4 Hz), 28.1, 26.0, 24.8, 24.7, 24.1, 24.0. 31 P NMR (121 MHz, C6 D6 ): δ122.8. 1
Chemical Phosphorylation of Deoxyribonucleosides and Thermolytic DNA Oligonucleotides
Prepare 2 -deoxyribonucleoside phosphoramidite S.19 9. Dry 542 mg (1.00 mmol) commercial 5 -O-(4,4 -dimethoxytrityl)-2 -deoxythymidine by coevaporation, twice with 5 mL anhydrous pyridine and once with 5 mL dry toluene, in a 50-mL round-bottom flask using a rotary evaporator
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connected to a vacuum pump. Add a stir bar and seal the flask with a rubber septum. 10. Under an argon atmosphere, add 5 mL anhydrous MeCN using a 10-mL glass syringe, and then add 417 mg (1 mmol) phosphorodiamidite S.23 via a 1-mL glass syringe attached to a 16-G needle. Stir the solution with a magnetic stirrer. 11. While stirring under a positive pressure of argon, remove the rubber septum from the flask and quickly add 0.5 mmol of sublimed 1H-tetrazole using a spatula. Seal the flask with the rubber septum. Continue stirring 1 hr. 12. Add a second portion of 0.5 mmol sublimed 1H-tetrazole. Allow the reaction mixture to stir 3 hr at ∼25◦ C. 13. Add 1 mL triethylamine by syringe and immediately concentrate the reaction mixture to a syrup using a rotary evaporator connected to a vacuum pump. 14. Suspend the crude product in ∼3 mL of 9:1 (v/v) benzene/triethylamine and apply the suspension to a 2.5 × 20–cm disposable Flex chromatography column containing ∼30 g silica gel that has been equilibrated in 9:1 (v/v) benzene/triethylamine. 15. Elute the column with 9:1 (v/v) benzene/triethylamine and collect 10-mL fractions. 16. Analyze fractions by TLC on a 2.5 × 7.5–cm EMD silica gel 60 F254 TLC plate using 9:1 (v/v) benzene/triethylamine as the eluent. Visualize using a hand-held UV lamp. TLC analysis of diastereomeric S.19 shows two tight spots, the Rf of which is ∼0.45 when 9:1 (v/v) benzene/triethylamine is used as the eluent.
17. Pool appropriate fractions in a 250-mL round-bottom flask and rotary evaporate under reduced pressure (∼20 mmHg) until a white amorphous solid is obtained. 18. Dissolve the solid in ∼3 mL benzene and add the solution to ∼100 mL of vigorously stirred cold (−20◦ C) hexane in a 250-mL round-bottom flask. 19. Allow the suspension to settle and carefully decant off most of the supernatant. 20. Evaporate the wet material to dryness under reduced pressure using a rotary evaporator and then dissolve in ∼10 mL dry benzene. 21. Transfer the solution to a 100-mL round-bottom flask, freeze in a dry-ice/acetone bath, and lyophilize under high vacuum to afford (733 mg, 0.85 mmol) triethylaminefree S.19 as a white powder in 85% yield. P NMR (121 MHz, C6 H6 ): δ 148.5, 147.8. APESI-HRMS: calcd. for C46 H61 N3 O11 P (M+H+ ) 862.4044, found 862.4036.
31
COMMENTARY Background Information The development of oligonucleotide prodrugs has evolved considerably in recent years as a judicious approach to enhance cellular uptake of antisense oligonucleotides and improve the stability of these biopolymers to nucleases present in extracellular and intracellular environments. Oligonucleotide prodrugs are typically produced by temporarily neutralizing the negatively charged phosphates/thiophosphates of therapeutic oligonucleotides with bioreversible protecting groups. Given that cellular uptake of oligonucleotide prodrugs is
commensurate with their relative lipophilicity and postulated to proceed through a passive diffusion mechanism (Bologna et al., 2002), the neutralization of oligonucleotidic phosphodiesters with bioreversible acylthioethyl (Iyer et al., 1994; Barber et al., 1995; Tosquellas et al., 1998; Bologna et al., 2002), acyloxymethyl (Iyer et al., 1995), and 4acyloxybenzyl (Iyer et al., 1996, 1997) groups, or with derivatives of bis(hydroxymethyl)1,3-dicarbonyl compounds (Ora et al., 2001; Poij¨arvi et al., 2002, 2004, 2005), has been investigated as a process to improve cellular
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Chemical Phosphorylation of Deoxyribonucleosides and Thermolytic DNA Oligonucleotides
uptake of oligonucleotides. However, most of these groups are sensitive to the nucleophilic conditions routinely used for nucleobase deprotection. Thus, to prevent cleavage of bioreversible phosphate/thiophosphate protecting groups under such conditions, new strategies for nucleobase protection/deprotection have beee proposed (Alvarez et al., 1999; Spinelli et al., 2002; Guerlavais-Dagland et al., 2003; Ferreira et al., 2004). These strategies are arduous, however, and the requirement for intracellular carboxyesterase activity to convert neutral oligonucleotide prodrugs to their polyanionic bioactive state further underscores the limitations of these approaches to the preparation of oligonucleotide prodrugs. More recently, the preparation of oligonucleoside phosphorothioates functionalized with the thermolytic 2-(N-formyl-Nmethyl)aminoethyl group for thiophosphate protection has been reported (Grajkowski et al., 2005). These uncharged oligonucleotides exhibited the characteristics of oligonucleotide prodrugs and did not require intracellular esterases or other enzymes for prodrug-to-drug conversion; only a 37◦ C aqueous environment was necessary for their conversion to functional oligonucleoside phosphorothioate diesters. These thermolytic oligonucleotide prodrugs contained a CpG motif (Klinman et al., 2002; Krieg, 2002) and demonstrated immunotherapeutic properties in mice (Grajkowski et al., 2005). Moreover, the synthesis of these modified oligonucleotides did not require nucleobase-protectecting groups different than those commonly employed in solid-phase oligonucleotide synthesis. Depending on the nature of the phosphate/ thiophosphate protecting groups used to generate oligonucleotide prodrugs, the solubility of neutral oligonucleotides in biological media may be problematic. To avoid this problem, it was hypothesized that the functionalization of oligonucleotide prodrugs with one phosphomonoester group would impart these biomolecules with adequate solubility in biological media without significantly impairing cellular uptake attributes. To test this hypothesis, the thermolytic dinucleotide prodrug model S.22 was designed to undergo intramolecular cyclodeesterification and lead to the dinucleoside phosphorothioate diester TPS T under thermolytic conditions. The dinucleotide prodrug S.22 was prepared as shown in Figure 13.6.4 and described in Alternate Protocol 2, using deoxyribonucleoside phosphoramidite S.19
(see Support Protocol) via solid-phase techniques. Following hydrazinolysis of the levulinyl group from the solid-phase-linked dinucleotide S.20, thiophosphorylation of the 4-hydroxy-1-butyl thiophosphate protecting group was challenging in that S.22 must be produced without inducing premature thermolytic cleavage of the 4-thiophosphato-1butyl thiophosphate-protecting group. Several phosphorylating agents (S.24-S.29; see Fig. 13.6.7) have been reported in the literature for the preparation of 5 -/3 -phosphate or thiophosphate monoester derivatives of nucleosides, oligonucleotides, and polyols (Horn and Urdea, 1986; Uhlmann and Engels, 1986; Horn et al., 1987; Thuong and Chassignol, 1987; Guzaev et al., 1995; Lefebvre et al., 1995; Szczepanik et al., 1998; Tosquellas et al., 1998; Olesiak et al., 2002; Lartia and Asseline, 2004). Phosphorylating reagents S.24 (Uhlmann and Engels, 1986; Thuong et al., 1987; Szczepanik et al., 1998), S.26 (Horn et al., 1986, 1987), S.27 (Olesiak et al., 2002), and S.28 (Guzaev et al., 1995) require elevated temperature conditions for the generation of phosphate/thiophosphate monoesters within an acceptable period of time. Such conditions are unsuitable for the preparation of oligonucleotides functionalized with thermosensitive phosphotriester groups. Whereas the coupling efficiency of phosphorylating reagents S.24, S.25 (Lefebvre et al., 1995; Tosquellas et al., 1998), and S.27 cannot be easily monitored because they lack a reporter group, the H-phosphonate reagent S.29 (Lartia and Asseline, 2004) is incompatible with automated phosphoramidite chemistry. The limitations of these phosphorylation reagents toward the preparation of the heat-sensitive dinucleotide S.22 prompted the development of a phosphorylating agent compatible with automated phosphoramidite chemistry and capable of generating oligonucleotide phosphate/thiophosphate monoester derivatives under mild temperature conditions. To permit a relatively accurate assessment of its coupling efficiency, the phosphorylation reagent was functionalized with two reporter groups. Specifically, the bis[S(4,4 -dimethoxytrityl)-2-mercaptoethyl]-N,Ndiisopropylphosphoramidite S.4 was prepared and investigated for its potential application as an efficient reagent in the phosphorylation of DNA oligonucleotides and their thermolylic analogs (Aus´ın et al., 2005). The preparation of S.4 was achieved as shown in Figure 13.6.1 and as described in Basic Protocol 1. The parameters for optimal
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Figure 13.6.7 Phosphorylating reagents for nucleosides and oligonucleotides. DMTr, 4,4 -dimethoxytrityl. Adapted from Aus´ın et al. (2005) with permission from the American Chemical Society.
coupling efficiency of S.4 were determined by performing syntheses of 5 -/3 -phosphate or -thiophosphate monoester derivatives of commercial 2 -deoxyribonucleosides covalently linked through their 3 - or 5 -hydroxyl to long-chain alkylamine controlled-pore glass (LCAA-CPG) as shown in Figure 13.6.2 and as described in Basic Protocol 2. A 0.1 M solution of 1H-tetrazole-activated S.4 in dry acetonitrile was typically employed for the 5 -/3 -phosphitylation of detritylated S.5a-d or S.12a-d (see Fig. 13.6.2) and a solution of 0.1 M ethyl(methyl)dioxirane (Katakoa et al., 2001) in dichloromethane was selected to produce the solid-phase-linked 2 -deoxyribonucleoside 5 -/3 -phosphotriester derivatives. The selection of this oxidant, as opposed to the standard aqueous iodine solution, resulted in higher yields of the desired 2 -deoxyribonucleoside 5 -/3 phosphomonoester derivatives S.10a-d or S.13a-d. Alternatively, the sulfur transfer reagent 3H-1,2-benzodithiol-3-one-1,1-dioxide was used as an oxidant in the preparation of 2 -deoxyribonucleoside 5 -/3 -thiophosphate monoesters. Removal of the S-DMTr groups from solid-phase-bound 2 -deoxyribonucleoside 5 -/3 -phosphate or -thiophosphate triesters was accomplished under acidic conditions followed by treatment with aqueous dithiothreitol (DTT). The use of DTT is absolutely required for clean and optimal
production of 2 -deoxyribonucleoside 5 -/3 phosphate or -thiophosphate monoesters. The coupling efficiency of S.4 was compared with that of the commercial phosphorylating reagents S.26 and S.28 in the preparation of representative 2 -deoxyribonucleoside 5 -/3-phosphate or -thiophosphate monoesters. RP-HPLC analysis of the crude reaction products was performed (for chromatographic profiles of S.10d, S.11b, and S.14b, see charts 3, 5, and 9, respectively, in Aus´ın et al., 2005). The results indicate that the coupling efficiency of S.4 is comparable to that of S.26 and S.28 in the production of these compounds. When applied to the 5 -phosphorylation of a solid-phase-linked oligonucleotide (20-mer), the phosphorylating reagent S.4 was somewhat less efficient than S.26. Comparative polyacrylamide gel electrophoresis (PAGE) of the crude and deprotected oligonucleotide 5 -phosphate monoester produced using each phosphorylating reagent corroborates this assessment (data not shown) and is consistent with the RP-HPLC profile of the phosphorylation reaction (chart 12 in Aus´ın et al., 2005). The coupling yield of S.4 did not improve upon performing two consecutive 5 phosphorylation reactions or increasing the ◦ pore size of the CPG support to 1000 A. Thus, steric considerations related to the reagent and/or the size of the oligonucleotide chain are likely responsible for the relatively lower
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coupling efficiency (∼85%) of S.4 when applied to the 5 -phosphorylation of a 20-nt oligomer. Consistent with this assessment is the 5 -phosphorylation of a tetranucleotide via S.4, which proceeded in yields >95% (Aus´ın et al., 2005) and further underscored the significance of steric factors in the ability of S.4 to 5 -phosphorylate larger oligonucleotides. The phosphorylating reagent S.4 was particularly efficient in the preparation of the thermolytic dinucleotide prodrug model S.22 (Fig. 13.6.4). Specifically, a side-by-side phosphorylation of the solid-phase-linked dinucleoside 4-hydroxy-1-butyl thiophosphate triester was performed using either S.4 or S.28, after hydrazinolysis of S.20, to give S.22 at 25◦ C. RP-HPLC analysis of S.22 indicates that S.4 is by far superior to S.28 in the generation of S.22 under conditions that will not induce substantial thermolytic cleavage of the 4thiophosphato-1-butyl protecting group (chart 17 in Aus´ın et al., 2005). The phosphorylation of a 4-hydroxy-1butyl phosphotriester located in the middle of an oligonucleotide chain (20-mer) using S.4 was, however, less efficient than that achieved in the preparation of S.22, but comparable to the 5 -phosphorylation of an identical 20-mer under the same conditions. Purification of the 20-mer oligonucleotide functionalized with a 4-phosphato-1-butyl phosphate triester was accomplished by RP-HPLC; like its 4-hydroxy-1-butyl precursor, it was isolated as a mixture of two diastereomers (Fig. 13.6.8). In spite of its steric bulk, the phosphorylating agent S.4 is unique in its ability to produce phosphate/thiophosphate monoesters at 25◦ C. This attribute is critical to the phosphorylation
Chemical Phosphorylation of Deoxyribonucleosides and Thermolytic DNA Oligonucleotides
of thermosensitive oligonucleotide prodrugs and is recommended for such an application.
Critical Parameters and Troubleshooting Readers are referred to the Critical Parameters and Troubleshooting section of UNIT 2.7, which addresses issues pertinent to the preparation of phosphorodiamidite S.23. It is recommended that the phosphorylating reagent S.4 and 2 -deoxyribonucleoside phosphoramidite S.19 be precipitated from cold hexane following silica gel purification. This precipitation step removes contaminants, which often coelute with the desired phosphoramidites. A number of parameters including exclusion of residual triethylamine and moisture must be taken into consideration to ensure optimal coupling efficiency of phosphoramidites during automated and/or manual solid-phase oligonucleotide synthesis. These concerns are also addressed in UNIT 2.7. The use of oxidizers other than ethyl(methyl)dioxirane, when phosphorylation reagent S.4 is employed in the preparation of nucleoside 5 -/3 -phosphate monoesters or oligonucleotide 5 -phosphate monoesters, will result in lower yields and should therefore be avoided. Replacement of DTT with tris(2-carboxyethyl)phosphine hydrochloride in the preparation of nucleoside 5 -/3 -phosphates or -thiophosphates and oligonucleotide 5 phosphates or -thiophosphates, when using S.4 as the phosphorylation reagent, is not recommended as it will result in considerably lower yields of the desired phosphorylated/thiophosphorylated products. A freshly
Figure 13.6.8 Comparative RP-HPLC analysis of fully deprotected and purified d(AP TP CP CP GP TP AP GP CP TP(POB) AP AP GP GP TP CP AP TP GP C) (A) and its precursor d(AP TP CP CP GP TP AP GP CP TP(HBU) AP AP GP GP TP CP AP TP GP C) (B). P(POB), 4-phosphato-1-butyl phosphate triester; P(HBU), 4-hydroxy-1-butyl phosphate triester. Chromatographic conditions described in Basic Protocol 2, step 19. Peak heights normalized to highest peak, which is set to 1 arbitrary unit. (A) –MALDI-TOF-MS: calcd. for C199 H236 N75 O122 P20 (M–H)– 6249, found 6255. (B) –MALDI-TOF-MS: calcd. for C199 H235 N75 O119 P19 (M–H)– 6170, found 6178.
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prepared solution of DTT should always be used for optimal results. It is also recommended to prepare fresh TCA solution from solid TCA for the cleavage of both O- and S-DMTr protecting groups, as commercial TCA solutions contain variable amounts of phosgene (Cie´slak et al., 2005), potentially leading to the formation of unwanted side products. Removal of the levulinyl group from S.20 upon exposure to 0.5 M hydrazine hydrate in 3:2 (v/v) pyridine/AcOH under the conditions stated in Alternate Protocol 2 does not cleave the dinucleotide from the CPG support to a significant extent (Iwai and Ohtsuka, 1988a,b; Ueno et al., 2003). However, the duration of the hydrazinolysis step should be kept to a minimum to mitigate untimely loss of the N-benzoyl group from adenines and cytosines (Iwai and Ohtsuka, 1988a,b).
Anticipated Results The synthesis of the alcohol S.1 was performed as described in the literature (Rademann and Schmidt, 1997) for a similar compound. The yield of silica-gel-purified S.1 was 90%. The phosphorylating reagent S.4 was purified by silica gel chromatography and was isolated as a lyophilized, triethylaminefree powder in 82% yield. The stability of S.4 is noteworthy. 31 P NMR analysis of S.4 stored as a 0.1 M solution in dry MeCN over 25 days at 25◦ C revealed the formation of sideproducts accounting for only ∼15% of total peak area (Aus´ın et al., 2005). The purity of crude deoxyribonucleoside 5 -/3 -phosphate or -thiophosphate monoesters prepared using S.4 was ∼95% on the basis of RP-HPLC analyses of the phosphorylation reactions. of 5 -Phosphorylation/thiophosphorylation oligonucleotides (20-mers) using S.4 proceeded to the extent of ∼85% as judged by PAGE and RP-HPLC analyses of the 5 -phosphorylated and 5 -unphosphorylated oligonucleotides (chart 12 in Aus´ın et al., 2005). The phosphorodiamidite S.23 was isolated as an oil in 72% yield after silica gel chromatography. The deoxyribonucleoside phosphoramidite S.19 was prepared employing the diamidite S.23 and was isolated after silica gel chromatography in 85% yield as a lyophilized, triethylamine-free powder. Solid-phase synthesis of S.22 from phosphoramidite S.19 proceeded smoothly in yields exceeding 95% on the basis of RP-HPLC analysis of the crude dinucleotide
prior to and after thermolytic cleavage of the 4-thiophosphato-1-butyl thiophosphate protecting group (Fig. 13.6.5).
Time Considerations The preparation and purification of alcohol S.1 takes ∼2.5 days. The synthesis of phosphorylating reagent S.4 from S.1 and its subsequent purification require ∼1 day, including lyophilization. Solid-phase 5 -/3 phosphorylation or -thiophosphorylation of deoxyribonucleosides and 5 -phosphorylation/-thiophosphorylation of oligodeoxyribonucleotides in addition to complete nucleobase deprotection requires 2 to 12 hr to complete. Preparation, purification, and lyophilization of phosphorodiamidite S.23 are achieved within 1 day. Synthesis of deoxyribonucleoside phosphoramidite S.19 from S.23 along with its purification and lyophilization are also accomplished within 1 day. Solid-phase synthesis of the thermolytic dinucleotide prodrug model S.22, including release from the support, RP-HPLC analysis of the crude material, and that of its thermolytic conversion to the dinucleoside phosphorothioate diester TPS T, take 8 hr to complete.
Literature Cited Alvarez, K., Vasseur, J.-J., Beltran, T., and Imbach, J.-L. 1999. Photocleavable protecting groups as nucleobase protections allowed the solid-phase synthesis of base-sensitive SATEprooligonucleotides. J. Org. Chem. 64:63196328. Aus´ın, C., Grajkowski, A., Cie´sak, J., and Beaucage, S.L. 2005. An efficient reagent for the phosphorylation of deoxyribonucleosides, DNA oligonucleotides and their thermolytic analogues. Org. Lett. 7:4201-4204. Barber, I., Rayner, B., and Imbach, J.-L. 1995. The prooligonucleotide approach. I: Esterasemediated reversibility of dithymidine-S-alkylphosphorothioates. Bioorg. Med. Chem. Lett. 5:563-568. Bologna, J.-C., Viv`es, E., Imbach, J.-L., and Morvan, F. 2002. Uptake and quantification of intracellular concentration of lipophilic pro-oligonucleotides in HeLa cells. Antisense Nucleic Acid Drug Dev. 12:33-41. Cie´slak, J., Aus´ın, C., Chmielewski, M.K., Kauffman, J.S., Snyder, J., Del-Grosso, A., and Beaucage, S.L. 2005. 31 P NMR study of the desulfurization of oligonucleoside phosphorothioates effected by “aged” trichloroacetic acid solutions. J. Org. Chem. 70:3303-3306. Ferreira, F., Vasseur, J. J., and Morvan, F. 2004. Lewis acid deprotection of silyl-protected oligonucleotides and base-sensitive oligonucleotide analogues. Tetrahedron Lett. 45:62876290.
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Grajkowski, A., Pedras-Vasconcelos, J., Wang, V., Aus´ın, C., Hess, S., Verthelyi, D., and Beaucage, S.L. 2005. Thermolytic CpG-containing DNA oligonucleotides as potential immunotherapeutic prodrugs. Nucl. Acids Res. 33:3550-3560. Guerlavais-Dagland, T., Meyer, A., Imbach, J.-L., and Morvan, F. 2003. Fluoride-labile protecting groups for the synthesis of base-sensitive methyl-SATE oligonucleotide prodrugs. Eur. J. Org. Chem. 2327-2335. Guzaev, A., Salo, H., Azhayev, A., and L¨onnberg, H. 1995. A new approach for chemical phosphorylation of oligonucleotides at the 5 -terminus. Tetrahedron 51:9375-9384. Horn, T. and Urdea, M.S. 1986. A chemical 5 phosphorylation of oligodeoxyribonucleotides that can be monitored by trityl cation release. Tetrahedron Lett. 27:4705-4708. Horn, T., Allen, J.S., and Urdea, M.S. 1987. Solid supported chemical 5 -phosphorylation of oligodeoxyribonucleotides that can be monitored by trityl cation release—Application to gene synthesis. Nucleosides Nucleotides 6:335340. Iwai, S. and Ohtsuka, E. 1988a. Synthesis of oligoribonucleotides by the phosphoramidite approach using 5 -levulinyl and 2 -tetrahydrofuranyl protection. Tetrahedron Lett. 29:5383-5386. Iwai, S. and Ohtsuka, E. 1988b. 5 -Levulinyl and 2 -tetrahydrofuranyl protection for the synthesis of oligoribonucleotides by the phosphoramidite approach. Nucl. Acids Res. 16:9443-9456. Iyer, R.P., Phillips, L.R., Egan, W., Regan, J.B., and Beaucage, S.L. 1990. The automated synthesis of sulfur-containing oligodeoxyribonucleotides using 3H-1,2-benzodithiol-3-one 1,1-dioxide as a sulfur-transfer reagent. J. Org. Chem. 55:46934699. Iyer, R.P., Yu, D., and Agrawal, S. 1994. Stereospecific bio-reversibility of dinucleoside S-alkyl phosphorothiolates to dinucleoside phosphorothioates. Bioorg. Med. Chem. Lett. 4:2471-2476. Iyer, R.P., Yu, D., and Agrawal, S. 1995. Prodrugs of oligonucleotides—The acyloxyalkyl esters of oligodeoxyribonucleoside phosphorothioates. Bioorg. Chem. 23:1-21. Iyer, R.P., Yu, D., Devlin, T., Ho, N.-H., and Agrawal, S. 1996. Acyloxyaryl prodrugs of oligonucleoside phosphorothioates. Bioorg. Med. Chem. Lett. 6:1917-1922. Iyer, R.P., Ho, N.-H., Yu, D., and Agrawal, S. 1997. Bioreversible oligonucleotide conjugates by site-specific derivatization. Bioorg. Med. Chem. Lett. 7:871-876.
Chemical Phosphorylation of Deoxyribonucleosides and Thermolytic DNA Oligonucleotides
Katakoa, M., Hattori, A., Okino, S., Hyodo, M., Asano, M., Kawai, R., and Hayakawa, Y. 2001. Ethyl(methyl)dioxirane as an efficient reagent for the oxidation of nucleoside phosphites into phosphates under nonbasic anhydrous conditions. Org. Lett. 3:815-818. Klinman, D.M., Takeshita, F., Gursel, I., Leifer, C., Ishii, K.J., Verthelyi, D., and Gursel, M. 2002.
CpG DNA: Recognition by and activation of monocytes. Microbes Infect. 4:897-901. Krieg, A.M. 2002. CpG motifs in bacterial DNA and their immune effects. Annu. Rev. Immunol. 20:709-760. Lartia, R. and Asseline, U. 2004. New reagent for the preparation of oligonucleotides involving a 5 -thiophosphate or a 5 -phosphate group. Tetrahedron Lett. 45:5949-5952. Lefebvre, I., P´erigaud, C., Pompon, A., Aubertin, A.-M., Girardet, J.-L., Kirn, A., Gosselin, G., and Imbach, J.-L. 1995. Mononucleoside phosphotriester derivatives with S-acyl2-thioethyl bioreversible phosphate-protecting groups—Intracellular delivery of 3 -azido-2 ,3 dideoxythymidine 5 -monophosphate. J. Med. Chem. 38:3941-3950. Olesiak, M., Krajewska, D., Wasilewska, E., Korczy´nski, D., Baraniak, J., Okruszek, A., and Stec, W.J. 2002. Thiophosphorylation of biologically relevant alcohols by the oxathiaphospholane approach. Synlett 6:967-971. Ora, M., M¨aki, E., Poij¨arvi, P., Neuvonen, K., Oivanen, M., and L¨onnberg, H. 2001. Hydrolytic stability of nucleoside phosphotriesters derived from bis(hydroxymethyl)-1,3-dicarbonyl compounds and their congeners: Towards a novel pro-drug strategy for antisense oligonucleotides. J. Chem. Soc. Perkin Trans. 2881885. Poij¨arvi, P., M¨aki, E., Tomperi, J., Ora, M., Oivanen, M., and L¨onnberg, H. 2002. Towards nucleotide prodrugs derived from 2,2-bis(hydroxymethyl)malonate and its congeners: Hydrolytic cleavage of 2-cyano-2(hydroxymethyl)-3-methoxy-3-oxopropyl and 3-(alkylamino)-2-cyano-2-(hydroxymethyl)-3oxopropyl protections from the internucleosidic phosphodiester and phosphorothioate linkages. Helv. Chim. Acta 85:1869-1876. Poij¨arvi, P., Oivanen, M., and L¨onnberg, H. 2004. Towards oligonucleotide prodrugs: 2,2-bis(ethoxycarbonyl) and 2(alkylaminocarbonyl)-2-cyano substituted 3-(pivaloyloxy)propyl groups as biodegradable protecting groups for internucleosidic phosphoromonothioate. Lett. Org. Chem. 1:183-188. Poij¨arvi, P., Heinonen, P., Virta, P., and L¨onnberg, H. 2005. 2,2-Bis(ethoxycarbonyl)- and 2(alkylaminocarbonyl)-2-cyano-substituted 3(pivaloyloxy)propyl groups as biodegradable phosphate protections of oligonucleotides. Bioconjug. Chem. 16:1564-1571. Rademann, J. and Schmidt, R.R. 1997. Repetitive solid-phase glycosylation on an alkyl thiol polymer leading to sugar oligomers containing 1,2trans- and 1,2-cis-glycosidic linkages. J. Org. Chem. 62:3650-3653. Spinelli, N., Meyer, A., Hayakawa, Y., Imbach, J.-L., and Vasseur, J.-J. 2002. Use of allylic protecting groups for the synthesis of base-sensitive prooligonucleotides. Eur. J. Org. Chem. 1:4956.
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Szczepanik, M.B., D´esaubry, L., and Johnson, R.A. 1998. One-pot synthesis of deoxyadenosine 3 -thiophosphates. Tetrahedron Lett. 39:74557458. Thuong, N.T. and Chassignol, M. 1987. Synthesis and reactivity of oligothymidylates containing a substituted intercalating agent and thiophosphate group. Tetrahedron Lett. 28:4157-4160. Tosquellas, G., Alvarez, K., Dell’Aquila, C., Morvan, F., Vasseur, J.-J., Imbach, J.-L., and Rayner, B. 1998. The pro-oligonucleotide approach: Solid-phase synthesis and preliminary evaluation of model pro-dodecathymidylates. Nucl. Acids Res. 26:2069-2074. Tsukamoto, H. and Kondo, Y. 2003. Facile and selective cleavage of allyl ethers based on palladium(0)-catalyzed allylic alkylation of N,N -dimethylbarbituric acid. Synlett 7:10611063.
Ueno, Y., Shibata, A., Matsuda, A., and Kitade, Y. 2003. Synthesis of 3 -3 -linked oligonucleotides branched by a pentaerythritol linker and the thermal stabilities of the triplexes with singlestranded DNA or RNA. Bioconjug. Chem. 14:684-689. Uhlmann, E. and Engels, J. 1986. Chemical 5 phosphorylation of oligonucleotides valuable in automated DNA-synthesis. Tetrahedron Lett. 27:1023-1026.
Contributed by Cristina Aus´ın, Andrzej Grajkowski, Jacek Cie´slak, and Serge L. Beaucage Center for Drug Evaluation and Research Food and Drug Administration Bethesda, Maryland
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CHAPTER 14 Biologically Active Nucleosides INTRODUCTION ucleosides and their polymers are naturally occuring molecules with a wide area of functions in cell biology. Probably the best known functions are their roles in storage of information (DNA) and energy (ATP) in cells, in the use of this information via transcription and translation (RNA), as co-factors for enzymatic reactions (NAD) and in signal transduction systems (cAMP, ligands for purinergic receptors). It is not surprising, therefore, that chemists have considered nucleoside analogs as a potentially rich source of biologically active molecules. Three main fields of research that have been explored in the past involve the development of nucleoside analogs as antitumoral and antiviral agents and as ligands for membrane-bound receptors. The antitumoral and antiviral nucleosides are meant to kill cells or viruses, mainly via the interaction of their metabolites with DNA synthesis. Nucleosides developed as ligands for protein receptors are considered as agonists or antagonists for these receptors and may have broad applicability against human diseases. The first series of nucleoside analogs that will be described in this chapter are antiviral nucleosides, because this is the field that covers most of the nucleoside drugs that are on the market.
N
The first antiviral compounds that were discovered were methisazon and 5-iodo-2 deoxyuridine. In contrast to evolution in other therapeutic areas, antiviral research has developed rather slowly because the life cycle of viruses is closely associated with cellular metabolism, viruses are localized intracellularly and possess few virally encoded genes, and acute viral infections can be prevented by developing effective vaccines. This has changed since the discovery of a retrovirus as the causative agent of acquired immune deficiency syndrome (AIDS). Most antiviral agents have been discovered using screening programs. Only recently has “rational drug design” entered the field and this is, in most cases, based on a lead compound that was discovered by a trial-and-error approach. The classical antiviral compounds are nucleoside analogs with structures very similar to those of natural nucleosides: the structures of 5-iodo-2 -deoxyuridine (IDU) and 3 azido-3 -deoxythymidine (zidovudine) are very similar to that of thymidine (Fig. 14.0.1). IDU is an anti–herpes virus agent (HSV-1, HSV-2) and is used in clinics for the treatment of herpes keratitis. Zidovudine is an anti-HIV nucleoside.
Figure 14.0.1
Structural similarities between two nucleoside analogs and natural thymidine.
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Both nucleosides (IDU and zidovudine) have had an important impact on the development of antiviral drugs. IDU has long been the model compound for the development of new anti-herpes agents. The observation that a simple nucleoside (zidovudine) could slow down the evolution of AIDS has given an important stimulus to antiviral research. Since then, specific viral enzymes have been identified and used for the development of new selective antiviral agents. An important milestone in this field is the discovery of phosphonate nucleosides for the treatment of cytomegalovirus (CMV), human immunodeficiency virus (HIV), and hepatitis B virus (HBV) infections, to which Dr. Hol´y has made the most significant contribution. This class of compounds is described in UNIT 14.2. The discovery of phosphonate nucleosides as antiviral agents demonstrated that it is possible to circumvent the first step in the metabolic activation of nucleoside analogs (i.e., the conversion of the nucleoside to its monophosphate) and that such charged compounds can be taken up by cells (although not very efficiently). Antiviral nucleosides can be ranked in several categories. Base-modified pyrimidine nucleosides [IDU, 5-ethyl-2 -deoxyuridine (EDU), 5-(E)-bromovinyl-2 -deoxyuridine (BVDU), and 5-trifluoromethyl-2 -deoxyuridine (TFT); see Fig. 14.0.2] are all antiherpes agents. They are converted to their triphosphates by viral and cellular kinases. As several herpes viruses code for their own thymidine kinase, which has a broader substrate specificity than the human congener, their nucleosides are preferentially phosphorylated in virus-infected cells. This results in a selective antiviral activity. The triphosphates of the modified nucleosides function as substitutes for polymerases, and the modified nucleosides can be incorporated into DNA and interfere with normal transcription and replication processes. The monophosphate of TFT is also an inhibitor of thymidylate synthetase, which explains its strong antiviral activity. BVDU is active against HSV-1 and varicella-zoster virus (VZV). TFT is used topically for the treatment of herpes keratitis. 9-(β-D-Arabinofuranosyl)adenine (ara-A; Fig. 14.0.3) was the first example of a sugarmodified purine nucleoside that came on the market. It was used against HSV-1 encephalitis and herpes zoster in immunocompromised patients. However, ara-A was soon replaced by another series of sugar-modified purine nucleosides having an acyclic “sugar” chain. In UNIT 14.1, Dr. Hol´y describes the synthesis of representative acyclic nucleosides. Ara-A is rapidly deaminated in vivo by adenosine deaminase, and the search for deaminase inhibitors has led to the discovery of acyclovir or 9(2-hydroxyethoxymethyl)guanine. Acyclovir has remained the standard for treatment of HSV and varicella-zoster virus infection. Ganciclovir is being used for CMV infections. These acyclic nucleosides can be considered analogs of 2 -deoxyguanosine. Acyclovir is phosphorylated in virus-infected cells to its triphosphate. Although this compound
Introduction
Figure 14.0.2
Base-modified pyrimidine nucleosides (also see IDU in Fig. 14.0.1).
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Figure 14.0.3
Sugar- and base-modified purine nucleosides.
is a purine nucleoside, it is phosphorylated to its monophosphate by the viral-specific thymidine/deoxycytidine kinase. The triphosphate of acyclovir is a competitive inhibitor of dGTP for the viral DNA polymerase. The triphosphate may also function as substrate for the polymerase; upon incorporation of acyclovir into DNA, the triphosphate functions as chain terminator. Ganciclovir is phosphorylated in CMV-infected cells by the virally encoded (UL97) protein kinase. Although this agent has a broad activity spectrum (HSV-1, HSV-2, VZV, CMV, Epstein-Barr virus [EBV]), it is toxic for bone marrow (neutropenia) and its medical use is restricted to the treatment of CMV infections in immunocompromised patients. Penciclovir has the same antiviral spectrum as acyclovir. Its prolonged action is due to the higher intracellular stability of its triphosphate. Ribavirin (Fig. 14.0.3) is an example of a base-modified purine nucleoside. It has a broad antiviral spectrum against DNA as well as RNA viruses. It is mainly active against influenza virus and respiratory syncytial virus. Therapeutically, it is used for the treatment of respiratory syncytial virus and hepatitis C infection (in combination with interferons). HIV is a retrovirus, which means that the genomic RNA of the virus is first converted into DNA using reverse transcriptase. This enzyme has a broader substrate specificity than cellular DNA polymerases, and this property has been used as the basis for the development of nucleoside analogs as anti-HIV agents. The first nucleoside analog that came on the market for treatment of HIV was zidovudine (Fig. 14.0.1), and it was followed by didanosine, zalcitabine, stavudine, lamivudine, abacavir, and emtricitabine (Fig. 14.0.4). All these nucleoside analogs interfere with the function of reverse transcriptase once they are converted intracellularly to their triphosphates (in the case of didanosine and abacavir, the base moiety is also modified during intracellular metabolism into an adenine and guanine base, respectively). They all function as chain terminators after incorporation into DNA. Didanosine, zalcitabine, and stavudine are considered dideoxynucleoside analogs because they lack secondary hydroxyl groups on the sugar moiety. Abacavir is an example of a carbocyclic nucleoside; its synthesis is described in UNIT 14.4. Many carbocyclic nucleosides were synthesized in the 1980s and 1990s; however, only abacavir has reached
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Figure 14.0.4
Modified nucleosides as anti-HIV agents.
Figure 14.0.5
Prodrugs of purine nucleosides.
the market as an antiviral agent. Lamivudine and emtricitabine are L-nucleosides. These L-nucleosides are less toxic than their D-congeners because they are less effectively transported in mitochondria. Emtricitabine is four- to ten-fold more active than lamivudine in vitro against HIV, although the effect is cell-type dependent. In addition to lamuvidine (which is used against HIV and HBV), other L-nucleosides (L-thymidine and L-deoxycytidine) are under development for use against hepatitis B infections. The synthesis of L-thymidine in gram quantities is described in UNIT 14.3.
Introduction
The in vivo activity of a drug is dependent on the way the drug is administered. The bioavailability of nucleosides is often not very high, and therefore prodrugs are developed. The specific metabolism of nucleosides allows one to make use of enzymes involved in nucleoside metabolism to develop a prodrug. Xanthine oxidase is such a candidate. Guanine nucleosides are not very soluble in water. The 6-deoxy analog of the diacetyl derivative of penciclovir gives blood plasma concentrations that are ten times higher than with penciclovir when given orally. This prodrug (famciclovir; Fig. 14.0.5) is converted to penciclovir in vivo by esterase and xanthine oxidase. Valaciclovir (cfr. supra) is another example of a prodrug of a purine nucleoside. This L-valylester of acyclovir was developed as prodrug of acyclovir. It is absorbed readily in the intestines, making use
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Figure 14.0.6
Nucleoside phosphonates as antiviral agents.
of a stereospecific transport system, and is rapidly hydrolyzed after absorption. Both prodrugs (famciclovir and valaciclovir) are used for the treatment of systemic HSV and VZV infections. Nucleoside phosphonates (Fig. 14.0.6) are composed of a base, a phosphonate group, and a sugar mimic. The compounds that are currently on the market do not have a glycosidic bond and are enzymatically and chemically more stable than regular nucleosides. Nucleoside phosphonates need to be phosphorylated to their diphosphates by cellular enzymes before showing activity against a broad range of viruses. They are not well absorbed when given orally, and they are administered as prodrugs. Cidofovir is active against herpes, adeno, polyoma, papilloma, and pox viruses. It is used for treatment of CMV retinitis in AIDS. Adefovir is active against retroviruses, hepadnaviruses, and herpes viruses. It is used in its prodrug form (adefovir dipivoxil) for the treatment of HBV infections. Tenofovir is active against retroviruses and hepadnaviruses and is used in its prodrug form (tenofovir disoproxil) for treatment of HIV infection. These nucleoside phosphonates have the important characteristic that they have a long action time, which means that they require much less frequent dosing than other nucleosides. The field of antiviral nucleosides is still expanding and many new compounds are under development and/or in a research phase. Many infectious viruses are still waiting for a series of new nucleoside or nucleotide analogs for treatment. Perhaps the most striking examples are papilloma viruses and hepatitis C viruses. One of the few examples of nucleoside analogs that are active against HCV are 2 -C-methylated ribonucleosides. This finding has reactivated interests in synthetic protocols to obtain 2 - and 3 -branched-chain nucleosides. UNIT 14.5 describes a classical approach to obtain such nucleosides starting from the appropriate carbohydrate precursors. As several papilloma viruses can induce tumors, such nucleoside analogs may also be preventive for these cancers. Hepatitis C infection is a worldwide problem and also the most common cause of hepatocellular carcinoma. Another important direction for investigation is the treatment of pox virus infections (variola virus, the cause of smallpox, could be used as an efficient bioterrorist weapon). Hemorrhagic fever viruses may become a problem with climate changes. Ebola virus and rabies virus are among the most deadly viruses to infect humans. Influenza viruses cause epidemics or pandemics in humans. The SARS-associated coronavirus
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(severe acute respiratory syndrome) may lead to acute breathing problems and 10% mortality. Many viruses lead to gastrointestinal infections (e.g., reovirus) and cause diarrhea. Chemically, there are many opportunities for developing new antiviral agents in the nucleoside field. Most nucleoside analogs currently on the market have several drawbacks, and there is a lot of room for new and safer compounds. For example, one might ask how long acyclovir can remain the gold standard for treatment of herpes virus infection when its activity spectrum is not very broad and the compound is not very potent. The mode of action of most antiviral nucleosides is very similar, and new nucleoside analogs will emerge that target other intracellular enzymes to exert an antiviral effect. Many areas of nucleoside chemistry still need to be evaluated for their potential to generate new antiviral compounds. Although several compounds with a non-nucleosidic structure are being used as antiviral agents, nucleoside analogs will remain an important class of antivirals (as single therapy or in combination therapy) because of the simple reason that the most effective anti-infection agents are those that make and break covalent bonds. Piet Herdewijn
Introduction
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Synthesis of Acyclic Analogs of Adenosine
UNIT 14.1
This unit describes the synthesis of three acyclic analogs of adenosine. The first, 9-(S)(2,3-dihydroxypropyl)adenine (DHPA), is a compound useful for biological investigation of consequences induced by the reversible inhibition of cellular processes caused by a metabolically inert inhibitor of methylation reactions. The second, eritadenine, is a natural compound and is an extremely active irreversible inactivator of the same enzyme. The third is a somewhat less but still highly active inactivator, 3-(adenin-9yl)-2-hydroxypropanoic acid (AHPA). Contrary to the first two cases, the procedure for AHPA involves racemic compounds. Straightforward reactions leading to the target products are described in the basic protocols, while support protocols provide verified procedures for the preparation of key starting materials that have limited availability or are not available at all. The scales of these procedures are verified, and the syntheses can be performed in a standard organic chemistry laboratory with common equipment. A few special procedures and simple pieces of equipment are described in detail. If not stated otherwise, solutions are evaporated on a rotary evaporator at 1.5 to 2 kPa with a maximum bath temperature of 40◦ C. The compounds are dried at 1.5 Pa over phosphorus pentoxide. Analytical TLC is performed on Merck silica gel plates (UV254; 10-cm runs) in systems based on chloroform/methanol or toluene/ethyl acetate mixtures, with detection under UV light or (for non-UV-absorbing compounds) by iodine vapors or p-anisaldehyde reagent (see Reagents and Solutions). Silica gel supplied by Sigma is used for preparative column chromatography. Filtration of solutions is performed by suction through a Celite pad in a glass filter funnel. Crystallization of compounds is generally performed from saturated solutions made by refluxing in the appropriate solvent, which are then filtered while hot through glass or a tight cotton wool filter. Unless stated otherwise, crystallization is performed at 4◦ C. All flasks are equipped with NS 29/32 joints.
SYNTHESIS OF 9-(S)-(2,3-DIHYDROXYPROPYL)ADENINE (DHPA) The reaction scheme depicted in the Figure 14.1.1 illustrates the sequence of three reactions which compose the method of preparation: (a) formation of sodium salt of adenine by the reaction of adenine with sodium hydride, (b) predominantly regiospecific alkylation of adenine to the 1,3-dioxolane derivative (S.3) by the reaction of the above sodium salt with (S)-2,2-dimethyl-1,3-dioxolan-4-ylmethyl p-tolylsulfonate (S.2), and (c) deprotection of the reaction intermediate by acid hydrolysis. The scheme also shows synthesis of S.2, which is presented in Support Protocol 1.
BASIC PROTOCOL 1
The removal of the protecting isopropylidene group from S.3 can be achieved by boiling for 45 min in 80% acetic acid. However, due to the danger of racemization, the hydrolysis is performed instead with diluted aqueous sulfuric acid. After neutralization with barium hydroxide and filtration of barium sulfate, the product (S.4) is obtained by crystallization. It has a negligible content of ashes.
Materials Sodium hydride, 60% dispersion in paraffin oil Dimethylformamide (DMF), anhydrous Adenine (R)-2,2-Dimethyl-4-hydroxymethyl-1,3-dioxolane p-tolylsulfonate (S.2; see Support Protocol 1) Toluene, reaction grade Chloroform, reaction grade Contributed by Antonin Hol´y Current Protocols in Nucleic Acid Chemistry (2005) 14.1.1-14.1.21 C 2005 by John Wiley & Sons, Inc. Copyright
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Figure 14.1.1 tolylsulfonyl.
Synthesis of 9-(S)-(2,3-dihydroxypropyl)adenine (S-DHPA). Py, pyridine; Ts, p-
Celite Methanol, reaction grade Diethyl ether, reaction grade 0.25 M aqueous sulfuric acid Barium hydroxide octahydrate (or monohydrate), crystalline Saturated aqueous barium hydroxide Ethanol, reaction grade 1-L round-bottom flask Magnetic stirrer with heating plate and silicon oil bath Calcium chloride protecting tube Rotary evaporator with diaphragm vacuum pump Reflux condenser 8- to 10-cm-diameter glass filtration funnel with filter flask 500- and 1000-mL Erlenmeyer flasks 2-L glass or plastic beaker (flat bottomed) Plastic foil Water aspirator pH detection papers Additional reagents and equipment for TLC (APPENDIX 3D) Synthesize (S)-4-(adenin-9-yl)methyl-2,2-dimethyl-1,3-dioxolane 1. While stirring on a magnetic stirrer, add 12 g of 60% sodium hydride in paraffin oil (7.2 g, 0.3 mol NaH) portionwise to 400 mL DMF in a 1-L round-bottom flask placed in a silicon oil bath. Prior to the reaction, check the quality of the DMF by adding several milligrams of sodium hydride suspension to 2 mL DMF in a reagent tube. Do not use DMF if sodium hydride reacts with foaming, which indicates the decomposition of the solvent and the presence of formic acid or water. No special drying of the flask is necessary. The flask and other glassware used in the procedure can be rinsed with ethanol or acetone and air dried. Synthesis of Acyclic Analogs of Adenosine
CAUTION: Perform the reaction in a well-ventilated chemical fume hood. Wear protective gloves while working with sodium hydride. It is water sensitive. Wash everything that comes into contact with sodium hydride first with ethanol and then with water.
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2. Add 40.5 g (0.3 mol) adenine to the suspension in small portions (∼2 g each), always waiting until after the violent hydrogen evolution ceases before adding the next portion. 3. Close the flask with a calcium chloride protecting tube and heat the silicon oil bath slowly to 80◦ C. Continue stirring the reaction mixture at this temperature for 2 hr. At the end of this period the suspension turns very thick and obstructs the magnetic stirring. It will quickly dissolve in the next step.
4. Add a solution of 89 g (0.31 mol) (R)-2,2-dimethyl-4-hydroxymethyl-1,3-dioxolane p-tolylsulfonate in 100 mL DMF. Continue heating the mixture under magnetic stirring at 100◦ C for 20 hr. Heating and stirring can be discontinued overnight.
5. Evaporate the solvent in vacuo on a rotary evaporator at max. 45◦ C, and coevaporate the residue twice with 150 mL toluene. 6. Add 600 mL chloroform and bring the mixture to boil under a reflux condenser for 2 to 3 min. Decant the solution and filter it while hot through a glass filter with a 1-cm-thick Celite pad. Repeat the extraction and filtration twice more using 500 mL chloroform. Prepare the Celite pad by suspending the required quantity of Celite in methanol, filtering it through the glass filter, and washing with methanol. Do not let air pass through the layer of Celite during vacuum filtration!
7. Evaporate the combined chloroform filtrates in vacuo. The residue should be solid or semi-solid. Co-distill the residue once more with 100 mL toluene to remove residual dimethylformamide.
8. Crystallize the solid residue from boiling methanol. After cooling to room temperature, set the solution aside at 0◦ C overnight. 9. Filter the crystalline material, wash it with 50 to 100 mL methanol and then 100 mL diethyl ether, and dry in vacuo. 10. Evaporate the filtrate in vacuo and crystallize the residue from methanol (see steps 8 and 9). 11. Check the purity of the two crops by TLC (APPENDIX 3D) using 9:1 (v/v) chloroform/methanol, with detection under UV light (Rf = 0.15 for adenine, 0.40 for S.3). The yield of S.3 should be between 35 and 45 g (47% to 60%). There is a certain quantity of the material left in the mother liquors. It is contaminated mainly with other regioisomers (a long, more slowly moving spot on TLC) and unreacted adenine. As the mother liquor also contains paraffin oil (4.8 g), the product would have to be purified by silica gel column chromatography, and the quantity of product obtained is usually not worth the cost and effort. Compound characterization: white leaflets, m.p. 210◦ -211◦ C. C11 H15 N5 O2 (249.3): 52.99% C, 6.06% H, 28.09% N. 1 H NMR (DMSO-d6 ): δ 1.27, 1.31 (2 × s, 2 × 3H, 2 × CH3 , isopropylidene), 3.89 (m, 2H, J1 ,2 = 6.0 Hz, J1 ,2 = 5.5 Hz, Jgem = 9.0 Hz, H1 ), 4.29 (m, 2H, H3 ), 4.49 (m, 1H, H2 ), 7.10 (br s, NH2 ), 8.05, 8.15 (s, 2H, H2 + H8).
Synthesize 9-(S)-(2,3-dihydroxypropyl)adenine 12. In a glass or plastic 2-L beaker, dissolve 40 g of S.3 under stirring in 800 mL diluted (0.25 M) aqueous sulfuric acid. Cover the beaker with a plastic foil and leave the solution standing overnight at room temperature.
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13. Place the beaker in an ice bath and cool the solution down to 0◦ C. 14. Add ∼60 g crystalline barium hydroxide (octahydrate) in small portions under stirring, at such a rate as to keep the solution at or beneath 20◦ C. Monitor the pH of the suspension occasionally using pH paper. When the pH reaches 6, cease adding solid hydroxide and add a saturated solution of barium hydroxide in water to bring the pH to 7.0 (using a pH meter with glass electrode). If the pH of the solution exceeds the neutral point and goes into the alkaline range, it can be brought back to neutral by adding very dilute sulfuric acid or a few pieces of dry ice.
15. Warm the suspension to 80◦ C. After 30 min, filter through a thick pad of Celite (≥2 cm thick) and wash the precipitate with 1 L boiling water. It is convenient to use a filter funnel of larger diameter and to renew the surface of the Celite pad by gently scraping it with spatula.
16. Cool the filtrate down and evaporate it to dryness under vacuum. 17. Crystallize the residue from a minimum volume of boiling 80% aqueous ethanol (∼400 to 500 mL). Let the solution stand overnight in a refrigerator (4◦ C). 18. Collect the material by filtration, wash it on the filter with ethanol and diethyl ether, and dry the product under vacuum. The yield of S.4 should be between 30 and 32 g (90% to 95%). Compound characterization (S.4): m.p. 202◦ -203◦ C. [α]D 20 = –460 (c = 1, water). 1 H NMR (400 MHz, DMSO-d6 ): δ 3.31 (dd, 1H, Jgem = 11.1 Hz, J3 b,2 = 6.3 Hz, H3 b), 3.40 (dd, 1H, Jgem = 11.1 Hz, J3 a,2 = 5.2 Hz, H3 a), 3.83 (dddd, 1H, J2 ,1 = 8.2, 3.6 Hz, J2 ,3 = 6.3, 5.2 Hz, H2 ), 4.03 (dd, 1H, Jgem = 13.9 Hz, J1 b,2 = 8.2 Hz, H1 b), 4.32 (dd, 1H, Jgem = 13.9 Hz, J1 a,2 = 3.6 Hz, H1 a), 5.10 (vb, 2H, 2 × OH), 7.64 (bs, 2H, NH2 ), 8.12, 8.21 (2 × s, 2 × 1H, H2 + H8). SUPPORT PROTOCOL 1
PREPARATION OF (R)-[2,2-DIMETHYL-1,3-DIOXOLAN-4-YL]METHYL p-TOLYLSULFONATE Though this compound is now commercially available (Daiso, TCI America), it is sometimes practical (and certainly less expensive) to prepare it fresh from (S)-4hydroxymethyl-2,2-dimethyl-1,3-dioxolane (S.1), which is available from D-mannitol.
Materials p-Tolylsulfonyl chloride Pyridine, anhydrous (S)-4-Hydroxymethyl-2,2-dimethyl-1,3-dioxolane (S.1; Pfanstiehl Laboratories, Senn Chemicals) Ethanol Ethyl acetate Magnesium sulfate, anhydrous 2-L round-bottom flask Rotary evaporator 1. In a 2-L round-bottom flask, dissolve 250 g (1.31 mol) p-tolylsulfonyl chloride in 1 L pyridine and cool with ice. 2. Add dropwise, under cooling with ice, a solution of 141 g (1.065 mol) (S)-4hydroxymethyl-2,2-dimethyl-1,3-dioxolane in 250 mL pyridine. Synthesis of Acyclic Analogs of Adenosine
3. Stir the mixture 2 hr at 0◦ C and set it aside for 2 days at ambient temperature. 4. Add 50 mL ethanol and let stand 1 hr.
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5. Concentrate the solution in vacuo in a rotary evaporator at 30◦ C to give a thick syrup. 6. Dilute with 1 L ethyl acetate and wash three times with 500 mL water. 7. Dry with magnesium sulfate, filter, and evaporate. The yield of S.2 should be between 285 and 290 g (93% to 97%). It can be stored in a tightly closed bottle in a refrigerator (4◦ C).
SYNTHESIS OF 9-(RS)-(2,3-DIHYDROXYPROPYL)ADENINE In some cases, it is possible to use the racemic product instead of the more expensive pure enantiomer, even in those cases when only one of the enantiomers is responsible for the desired effect and the other is quite neutral. Since the thalidomide case, such use is prohibited for human medicine, but can be useful in synthetic procedures where, for instance, a racemate might be an intermediate that loses the asymmetric carbon atom in the subsequent reaction sequence. The reaction scheme depicted in Figure 14.1.2 illustrates the alkylation of adenine with 4-chloro-2,2-dimethyl-1,3-dioxolane (S.5) in the presence of potassium carbonate. The racemic reaction intermediate (S.6) can be hydrolyzed by diluted aqueous inorganic acid to afford the title compound (S.7).
ALTERNATE PROTOCOL
Materials Adenine Potassium carbonate 4-Chloromethyl-2,2-dimethyl-1,3-dioxolane (S.5) Dimethylformamide (DMF), anhydrous Ethanol, reaction grade Diethyl ether, reaction grade Methanol, reaction grade Activated charcoal Celite Chloroform Conc. sulfuric acid, reagent grade Aqueous ammonia Saturated barium hydroxide octahydrate (or monohydrate) 3-L three-neck round-bottom flask Mechanical KPG stirrer Reflux condenser Calcium chloride protecting tube Silicon oil bath Rotary evaporator with diaphragm vacuum pump 2-L round-bottom flask 8- to 10-cm-diameter glass filtration funnel with filter flask Additional reagents and equipment for TLC (APPENDIX 3D) Synthesize 4-(adenin-9-yl)methyl-2,2-dimethyl-1,3-dioxolane 1. Set up a 3-L three-neck round-bottom flask with a mechanical KPG stirrer, equipped with a reflux condenser with calcium chloride protecting tube, and placed in a silicon oil bath. 2. Add 175 g (1.3 mol) adenine, 375 g anhydrous potassium carbonate, 220 g 4-chloro2,2-dimethyl-1,3-dioxolane, and 1400 mL DMF. Stir the mixture under reflux (bath temperature, 160◦ C) for 40 hr (total heating time, with interruptions). Heating and stirring can be discontinued overnight.
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Figure 14.1.2
Synthesis of 9-(RS)-(2,3-dihydroxypropyl)adenine (RS-DHPA).
3. Filter while hot through a Celite pad and wash the material on the filter twice with 150 mL DMF. 4. Concentrate the filtrate to half its volume under vacuum in a rotary evaporator at 50◦ C, and leave it to stand at room temperature overnight. 5. Filter the first crop of product, and wash it with 300 mL ethanol in portions and then with 200 mL diethyl ether. Dry under vacuum. 6. Evaporate the filtrate to dryness (maximum bath temperature, 60◦ C) and dissolve the residue in 1 L methanol. 7. Add 5 g activated charcoal and filter the mixture while hot over a Celite pad. Leave it to stand overnight at 0◦ C. Collect the second crop of the product by filtration and wash it with 200 mL ethanol. 8. Check the identity and purity of these two crops (steps 5 and 7) by TLC (APPENDIX 3D) in 4:1 (v/v) chloroform/methanol, with detection under UV light (Rf = 0.15 for adenine, 0.40 for S.6).When acceptable, combine the two crops for the next step. 9. To recrystallize the product, first add the combined (or individual) portions of product to boiling methanol in a 2-L round-bottom flask in a preheated silicon oil bath, and dissolve completely with stirring under reflux. Then leave the solution to stand overnight in a refrigerator at 4◦ C. The volume of methanol required for crystallization of the crude material depends on the speed of dissolution. If the compound spontaneously crystallizes during the dissolution, the solvent volume ultimately needed might be up to twice as large as needed otherwise.
10. Recover the product as in step 5. The yield of S.6 should be between 170 and 180 g (53% to 57%). It can be stored for an indefinite period of time at room temperature. Compound characterization: white leaflets, m.p. 208◦ -209◦ C. C11 H15 N5 O2 (249.3): 52.99% C, 6.06% H, 28.09% N. 1 H NMR (DMSO-d6 ): δ 1.27, 1.31 (2 × s, 2 × 3H, 2 × CH3 , isopropylidene), 3.89 (m, 2H, J1 ,2 = 6.0 Hz, J1 ,2 = 5.5 Hz, Jgem = 9.0 Hz, H1 ), 4.29 (m, 2H, H3 ), 4.49 (m, 1H, H2 ), 7.10 (br s, NH2 ), 8.05, 8.15 (s, 2H, H2 + H8).
Synthesize 9-(RS)-(2,3-dihydroxypropyl)adenine 11. Add conc. sulfuric acid slowly (dropwise) to a suspension of 174.5 g (0.7 mol) S.6 in 1300 mL water to dissolve the product, and continue the addition until the pH reaches 1.7 to 1.8. Leave the solution to stand overnight at ambient temperature. Synthesis of Acyclic Analogs of Adenosine
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12. Check the composition of the mixture by TLC in 4:1 (v/v) chloroform/methanol. When the spots of faster-moving starting compound have disappeared, continue to the next step. After spotting the sample on the TLC plate, keep it over an open bottle of conc. aqueous ammonia to neutralize the acid before developing the plate. The Rf value of the product is much slower compared to the starting material.
13. Neutralize the solution by adding hot saturated barium hydroxide solution until the pH reaches 7 to 7.1. Leave the solution standing at ambient temperature for 4 hr or overnight. 14. Filter through a Celite pad and wash the precipitate on the filter with a total of 1.5 L boiling water. 15. Cool the filtrate and evaporate it to half its volume under vacuum. Cool the solution on ice and filter the product by suction. 16. Wash the product on the filter with ethanol and diethyl ether (200 mL each) and then dry under vacuum. 17. Evaporate the filtrate to dryness and crystallize the residue from 80% aqueous ethanol. Collect the crystalline product S.7 by filtration, wash successively with ethanol and diethyl ether (200 mL each), and dry under vacuum. The yield of S.7 should be between 127 and 140 g (53% to 57%). It can be stored in dry form in closed bottles for an indefinite period of time at 4◦ C. Compound characterization: white crystals, m.p. 207◦ -208◦ C. C8 H11 N5 O2 (209.2): 45.93% C, 5.30% H, 33.48% N. UV spectrum (pH 2): λmax 259 (εmax 13,000); (pH 7 and 12): λmax 261 (εmax 11800). 1 H NMR (DMSO-d6 ): δ 3.45 (m, 2H, H1 ), 3.82 (m, 1H, H2 ), 4.14 (m, 2H, J2 ,3 = 3.5 Hz, Jgem = 12.5 Hz, H3 ), 7.06 (br s, NH2 ), 8.01, 8.20 (s, 2H, H2 + H8).
SYNTHESIS OF 4-(ADENIN-9-YL)-(2R,3R)-DIHYDROXYBUTANOIC ACID (ERITADENINE)
BASIC PROTOCOL 2
Eritadenine is a relatively simple natural adenine derivative isolated from the edible mushroom Lentinus edodes shitake. It seems to be the only natural compound that can be classified as an acyclic nucleoside analog (except for willardiine, which belongs among the amino acid analogs). It was believed to cause hypocholesterolemic activity by increasing the penetration of cholesterol through the intestinal wall. Later it was shown that this effect is not general and does not occur in human volunteers. It was then discovered that eritadenine can completely inactivate S-adenosylhomocysteine (SAH) hydrolase, which is involved in the regulation mechanism of biological methylation via Sadenosylmethionine (SAM). The inactivation is complete and irreversible within seconds, and the enzyme does not recover. Thus, it is a useful tool for biological investigation. There are several ways to prepare eritadenine. The simplest method would appear to be alkylation of adenine by 2,3-O-protected (isopropylidene or cyclohexylidene) Derythronolactone. However, this lactone is not easily available and must be synthesized by a multistep procedure from D-ribonolactone. In contrast, the procedure presented here (Fig. 14.1.3) can easily be performed on a large scale. It consists of oxidative degradation of free aldoses in alkaline solution, the so-called Neff reaction. The degradation occurs at carbon atoms 1 and 2, of which C1 is degraded to a carbonate and C2 is oxidized to a carboxylate. Methyl 2,3-O-isopropylidene-D-ribofuranoside (S.8), which is easily available from D-ribose, is transformed to the protected 5-(adenin-9-yl)-5-deoxy-D-ribofuranoside (S.10) via the corresponding 5-O-(p-tolylsulfonyl) derivative (S.9). After deprotection,
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Figure 14.1.3
Synthesis of D-eritadenine.
the resulting 5-O-(adenin-9-yl)-5-deoxy-D-ribofuranose (S.11) is exposed to oxygen (or air oxygen) in a strongly alkaline solution. After deionization on Dowex 50 and Dowex 1 ion-exchange resins, the product (S.12) is obtained by elution of the latter resin with aqueous formic acid and recrystallized from aqueous ethanol. The reaction is not quite straightforward, and is regularly accompanied by deeper degradation involving carbon atom C3 and even C4. Thus, eritadenine is accompanied in the reaction mixture with the (R)-enantiomer of 3-(adenin-9-yl)-2-hydroxypropanoic acid (AHPA; S.13) and even with a small amount (∼1%) of (adenin-9-yl)acetic acid. After transformation to alkyl esters, the mixture can be separated by silica gel column chromatography.
Materials
Synthesis of Acyclic Analogs of Adenosine
Sodium hydride, 60% dispersion in paraffin oil Dimethylformamide (DMF), anhydrous Adenine Methyl 2,3-O-isopropylidene-5-O-(p-tolylsulfonyl)-D-ribofuranoside (S.9; see Support Protocol 2) Toluene, reaction grade Chloroform, reaction grade Celite Methanol, reaction grade Ethanol, reaction grade Diethyl ether, reaction grade Conc. and 0.25 M aqueous sulfuric acid Sodium hydroxide (NaOH), reagent grade, pellets and 0.1 mM solution Oxygen 2-Propanol Conc. aqueous ammonia Dowex 50 × 8, acid form (see recipe)
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Dowex 1 × 2, acetate form (see recipe) 0.02 M acetic acid Conc. formic acid, reagent grade Amberlite IR 45 resin, OH form (see recipe) Silica gel Conc. hydrochloric acid (HCl), reagent grade Calcium chloride protecting tube 500- and 1000-mL round-bottom flasks Magnetic stirrer with heating plate and silicon oil bath Calcium chloride protecting tube Rotary evaporator with diaphragm vacuum pump 2-L Erlenmeyer flask 8- to 10-cm-diameter glass filtration funnel with filter flask 2-L glass or plastic beakers (flat bottomed) Glass column: 4-cm diameter, 40-cm length Fraction collector (optional) UV detector for continuous-flow detection (optional) 200-mL dropping funnel Peristaltic pump (optional) Reflux condenser 250-, 500-, and 1000-mL Erlenmeyer flasks Water aspirator Additional reagents and equipment for TLC (APPENDIX 3D) Synthesize methyl 5-(adenin-9-yl)-5-deoxy-2,3-O-isopropylidene-D-ribofuranoside 1. While stirring with a magnetic stirrer, add 4.8 g of 60% sodium hydride in paraffin oil (2.88 g, 0.12 mol NaH) portionwise to 200 mL DMF in a 500-mL round-bottom flask placed in a silicon oil bath. Prior to the reaction, check the quality of the DMF by adding several milligrams of sodium hydride suspension to 2 mL DMF in a reagent tube. Do not use DMF if sodium hydride reacts with foaming, which indicates the decomposition of DMF and the presence of formic acid or water. No special drying of the flask is necessary. The flask and other glassware used in the procedure can be rinsed with ethanol or acetone and air dried. CAUTION: Perform the reaction in a well-ventilated chemical fume hood. Wear protective gloves while working with sodium hydride. It is water sensitive. Wash everything that comes into contact with sodium hydride first with ethanol and then with water.
2. Add 16.2 g (0.12 mol) adenine to the suspension in small portions (∼2 g each), always waiting until after the violent hydrogen evolution ceases before adding the next portion. 3. Close the flask with a calcium chloride protecting tube and heat the silicon oil bath to 70◦ C. Stir the reaction mixture at this temperature for 1 hr. At the end of this period the suspension turns very thick and obstructs the magnetic stirring. It will quickly dissolve in the next step.
4. Add 35.8 g (100 mmol) crystalline methyl 2,3-O-isopropylidene-5-O-(ptolylsulfonyl)-D-ribofuranoside (S.9) to the thick suspension and continue stirring for 16 hr at 100◦ C. Heating and stirring can be discontinued overnight.
5. Evaporate the volatiles under vacuum on a rotary evaporator at 60◦ C, and coevaporate the residue twice with 100 mL toluene.
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6. Extract the residue three times with 150 mL boiling chloroform, filtering the solution through a Celite pad. 7. Check the precipitate on the filter for the presence of product by TLC (APPENDIX 3D) using 4:1(v/v) chloroform/methanol as the eluent (the product is faster than adenine). If needed, repeat the extraction with boiling chloroform. 8. Combine the extracts and evaporate under vacuum. 9. Dissolve in a minimum volume of boiling methanol, transfer the solution to an Erlenmeyer flask, and let it stand overnight in a refrigerator at 4◦ C. 10. Filter the crystalline product by suction, wash it twice with 50 mL precooled (0◦ C) ethanol and then twice with 50 mL diethyl ether, and dry it under vacuum. The yield of S.10 should be between 22 and 24 g (67% to 72%). It can be stored in dry form in closed bottles at 0◦ C. H NMR (400 MHz, DMSO-d6 ): δ 1.21, 1.34 (2 × s, 2 × 3H, CH3 -isopropylidene), 3.26 (s, 3H, OCH3 ), 4.20, 4.30 (2 × dd, 2H, Jgem = 14.0 Hz, J5 ,4 = 7.5 Hz, H5 ), 4.61 (td, 1H, J4 ,5 = 7.5 Hz, J4 ,3 = 1.0 Hz, H4 ), 4.67 (d, 1H, J2 ,3 = 6.0 Hz, H2 ), 4.81 (dd, 1H, J3 ,2 = 6.0 Hz, J3 ,4 = 1.0 Hz, H3 ), 4.97 (s, 1H, H1 ), 7.29 (bs, 2H, NH2 ), 8.17 (s, 1H, H2), 8.19 (s, 1H, H8). 13 C NMR (100.6 MHz, DMSO-d6 ): 24.82, 26.38 (CH3 -isopropylidene), 46.32 (CH2 -5 ), 54.95 (OCH3 ), 81.47 (CH-3 ), 84.15 (CH-4 ), 84.75 (CH-2 ), 109.21 (CH-1 ), 111.92 (C-isopropylidene), 118.86 (C5), 141.21 (CH-8), 149.74 (C4), 152.77 (CH-2), 156.23 (C6).
1
Synthesize eritadenine (Neff’s degradation) 11. Add 5 mL conc. sulfuric acid dropwise to a suspension of 20 g S.10 in 300 mL water and keep the solution 8 hr at 60◦ C. Confirm that the starting material has disappeared by TLC in 3:1 (v/v) chloroform/methanol. 12. Neutralize the solution with 10% sodium hydroxide by adding NaOH pellets and filling with water to a final volume of 1 L. 13. Bubble oxygen gently through the vigorously stirred solution for 10 to 12 hr, until the starting material disappears. Check by TLC in 7:1:2 (v/v/v) 2-propanol/conc. aqueous ammonia/water (the product has a slower mobility than the starting material). Air can be used in place of oxygen, but should be passed through a 10% NaOH solution to remove carbon dioxide before being introduced to the solution.
14. Transfer the reaction mixture to a plastic 2-L beaker. Rinse the flask twice with 100 mL water and add to the beaker. 15. Add Dowex 50 × 8 (H+ form) under stirring to neutralize the mixture. Add conc. aqueous ammonia to bring the pH of the solution to 10, and stir for 1 hr. 16. Filter the resin and wash it with 500 mL water. Concentrate the filtrate under vacuum to ∼200 mL volume. 17. Adjust a 4 × 40–cm glass column over a fraction collector and connect it with a UV detector (optional). 18. Fill the column with 300 mL Dowex 1 × 2 resin (acetate form) and wash it with 300 mL of 0.02 M acetic acid.
Synthesis of Acyclic Analogs of Adenosine
19. Take the concentrate from step 16 and add a few drops of ammonia to alkalify the solution to pH 9 to 10. Place the solution in a 200-mL dropping funnel and let it pass through the Dowex 1 × 2 column. Wash the column thoroughly with at least 1 L water (optionally, using a peristaltic pump) until the UV absorption of the eluate drops.
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20. Transfer the resin to a 2-L beaker with the use of pressurized air. Rinse the column with 200 mL water and add to the beaker. 21. Add 1 L of 2 M formic acid and stir the suspension for 30 min. Collect the resin by filtration. 22. Set aside the eluate and repeat step 21 with the collected resin. 23. Combine the eluates and evaporate at a maximum of 40◦ C (bath temperature) on a rotary evaporator. Co-distill the residue three times with 50 mL water. 24. Dissolve the raw material in a minimum volume of boiling water, transfer the solution to a 500-mL Erlenmeyer flask, and carefully add an equal volume of ethanol. Scratch the wall of the flask with a spatula to initiate crystallization. Set the flask in a refrigerator for 1 to 2 days to complete crystallization. 25. Filter the crystalline material, wash with ethanol and diethyl ether (100 mL each), and dry under vacuum. 26. Evaporate the filtrate under vacuum. Repeat steps 24 to 26.
Purify crude eritadenine 27. Add 3 mL conc. sulfuric acid to 300 mL methanol in a 1-L round-bottom flask equipped with reflux condenser and magnetic stir bar. 28. Place the apparatus in a silicon oil bath on a magnetic stirrer with heater plate and bring the solution to boil. 29. Add the crude eritadenine (13 to 14 g) to the solution and boil for 4 hr under reflux. Let stand overnight at ambient temperature. 30. Neutralize the stirred mixture by adding the required amount of Amberlite IR 45 resin in OH form, filter, and wash with 200 mL methanol. Amberlite is used here instead of Dowex 1, as it is a much weaker base and does not bind carboxylic acids as strongly.
31. Evaporate the solution under vacuum. Add 50 mL toluene and evaporate the solution to dryness. 32. Prepare a column of silica gel (400 mL) in chloroform. 33. Dissolve the crude methyl ester in 150 mL methanol, add 25 to 30 g dry silica gel, and evaporate in vacuo to dryness. 34. Transfer the silica gel with adsorbed material onto the top of the column and elute the column by a stepwise gradient from 0% to 20% (v/v) methanol in chloroform, increasing by 2.5% increments of methanol. Collect 20-mL fractions (optionally in a fraction collector). Check the eluate by TLC in 85:15 (v/v) chloroform/methanol. 35. Collect fractions containing methyl (adenin-9-yl)acetate (Rf = 0. 40), methyl (R)3-(adenin-9-yl)-2-hydroxypropanoate (Rf = 0.30), and eritadenine methyl ester (the main fraction; Rf = 0.20). 36. Dissolve eritadenine methyl ester in a 250-mL flask by adding 100 mL of 0.1 mM NaOH. Heat it to a boil and add diluted (1:10) HCl to a pH of 3 to 4. Scratch the flask walls to induce crystallization, and leave the flask standing in the refrigerator overnight. 37. Filter eritadenine by suction. Wash with ice water, ethanol, and diethyl ether. Dry under vacuum.
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The yield of S.12 should be between 22 and 24 g (67% to 72%). Eritadenine is quite stable and can be kept at room temperature in white glass bottles without signs of change. Compound characterization: m.p. > 260◦ C, [α]D 20 = + 15.6◦ (c = 1, 1 M HCl). C9 H11 N5 O4 (253.2): 42.68% C, 4.38% H, 27.66% N. UV spectrum (pH 2, pH 12): λmax 261 (εmax 14500). 1 H NMR (400 MHz, DMSO-d6 ): δ 3.22 (d, 1H, J3 ,2 = 8.6 Hz, H3 ), 3.60 (td, 1H, J2 ,3 = 8.6 Hz, J2 ,1 = 8.4, 2.7 Hz, H2 ), 4.03 (dd, 1H, Jgem = 13.8 Hz, J1 b,2 = 8.4 Hz, H1 b), 4.40 (dd, 1H, Jgem = 13.8 Hz, J1 a,2 = 2.7 Hz, H1 a), 7.13 (bs, 2H, NH2 ), 8.04, 8.11 (2 × s, 2 × 1H, H2 + H8). SUPPORT PROTOCOL 2
PREPARATION OF METHYL 2,3-O-ISOPROPYLIDENE-5-O-(p-TOLYLSULFONYL)-D-RIBOFURANOSIDE This material is used in one of the synthetic approaches to eritadenine and structurally related compounds. Starting material is easily available from D-ribose by glycosidation followed by protection of the cis-diol of the methyl D-ribofuranoside. The free primary hydroxyl group is reactive and can be easily converted to the p-tolylsulfonyl ester. This in turn enables broad exploitation of the intermediate as a substrate of nucleophilic substitution.
Materials D-Ribose
Methanol Conc. sulfuric acid Fehling solutions (see recipe) Sodium methoxide Triethylamine Toluene, reaction grade Acetone 2,2-Dimethoxypropene 2-Methoxypropane Diethyl ether, reaction grade Celite Pyridine anhydrous, reagent grade p-Tolylsulfonyl chloride Ethyl acetate Diluted (1:20) aqueous sulfuric acid Saturated potassium hydrogen carbonate Chloroform, reaction grade Hexane 2-L three-neck round-bottom flask Calcium chloride protecting tubes Dropping funnels Magnetic stirrer with heating plate and silicon oil bath pH detection papers 8- to 10-cm-diameter glass filtration funnel with filter flask 1- and 2-L round-bottom flasks Rotary evaporator with diaphragm vacuum pump Reflux condenser Y connector Distillation apparatus: optionally according to Figure 14.1.4, consisting of a 500-mL distillation flask on a short condenser, attached to a vacuum oil pump Synthesis of Acyclic Analogs of Adenosine
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Figure 14.1.4 Sketch of a short water-cooled distillation apparatus for higher-boiling or viscous liquids. Applicable to round-bottom flasks of up to 1 L volume, with magnetic stirring. An internal thermometer can be mounted in a correct position via the central tubing, e.g., through a bored plastic stopper.
1-L separatory funnel 500 and 1000 mL Erlenmeyer flasks Water aspirator Glass column: 4 cm diameter, 40 cm length (optional) Fraction collector (optional) Additional reagents and equipment for TLC (APPENDIX 3D)
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Synthesize methyl 2,3-O-isopropylidene-D-ribofuranoside 1. Set up a 2-L three-neck round-bottom flask equipped with a stir bar, calcium chloride protecting tube, and dropping funnel, and place on a magnetic stirrer. 2. Add 150 g (1 mol) D-ribose and 1 L methanol, and then add 10 mL conc. sulfuric acid dropwise under stirring over 1 hr. Continue stirring until all solid material dissolves, and then leave the solution standing overnight at ambient temperature. 3. Check whether glycosidation is quantitative by using the Fehling assay for reducing sugars and hydroxyaldehydes. Mix 2 mL Fehling solution A with 2 mL Fehling solution B in a test tube, and add several mg (0.2 to 0.5 mL) of the reaction mixture. Boil briefly (10 sec) and observe the color of the mixture. In the presence of reducing sugars, the deep blue color of the solution changes to brown, and eventually yellow cuprous oxide precipitates out. If the glycosidation is complete, the Fehling reaction should be negative (no color change).
4. Neutralize the mixture to a pH near 6.5 to 7.0 by portionwise addition of solid sodium methoxide (∼21 to 22 g). Spot the solution on wet pH paper to determine the neutralization point. Then make the mixture slightly alkaline (pH 9 to 10) by addition of triethylamine. 5. Filter the mixture by suction through a glass filter with a Celite pad into a 2-L round-bottom flask and evaporate the filtrate under vacuum on a rotary evaporator. Co-distill the residue twice with 100 mL toluene. 6. Add 200 mL acetone and 300 mL of 2,2-dimethoxypropane to the residue and place the flask in an ice bath on a magnetic stirrer. While stirring, slowly add a few drops of sulfuric acid (or preferably HCl in ethanol) to make the mixture acidic. Check pH on a wet pH paper. 7. Attach a reflux condenser protected by calcium chloride tube and dropping funnel via a Y connector. While stirring and cooling with ice, carefully add 200 mL 2methoxypropene at a rate that keeps the exothermic reaction under control. After addition is complete, stir the solution overnight at ambient temperature. 8. Add 5 mL triethylamine to make the mixture alkaline, and evaporate the volatiles in vacuo. 9. Triturate the residue with 500 mL acetone and then three times with 200 mL diethyl ether. 10. Filter the extract by suction through a Celite pad and evaporate off the volatiles in vacuo. 11. Transfer the residue by diethyl ether to a 500-mL distillation flask, set on a short condenser (e.g., Fig. 14.1.4), and distill in an oil-pump vacuum. Collect the main fraction, which boils at 100◦ to 110◦ C/10 Pa. The yield of S.8 should be between 130 and 140 g (64% to 69%). The product is sufficiently stable to be kept for several weeks at room temperature in a closed white glass flask without signs of change. Compound characterization: 1 H NMR (400 MHz, CDCl3 ): δ 1.39, 1.49 (2 × s, 2 × 3H, CH3 -isopropylidene), 3.44 (s, 3H, OCH3 ), 3.62 (dd, 1H, Jgem = 12.6 Hz, J5b,4 = 3.4 Hz, H5b), 3.70 (dd, 1H, Jgem = 12.6 Hz, J5a,4 = 2.3 Hz, H5a), 4.44 (bt, 1H, J4,5 = 3.4, 2.3 Hz, H4), 4.59 (d, 1H, J2,3 = 5.9 Hz, H2), 4.84 (bd, 1H, J3,2 = 5.9 Hz, H3), 4.98 (s, 1H, H1). Synthesis of Acyclic Analogs of Adenosine
Synthesize methyl 2,3-O-isopropylidene-5-O-(p-tolylsulfonyl)-D-ribofuranoside 12. Set up a 1-L round-bottom flask equipped with stir bar, calcium chloride protecting tube, and dropping funnel, and place in an ice bath on a magnetic stirrer.
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13. Dissolve 40.8 g (0.2 mol) freshly distilled S.8 in 140 mL pyridine. 14. Under cooling with ice water and stirring, add dropwise over 1 hr a solution of 45.8 g (0.24 mol) p-tolylsulfonyl chloride in 100 mL pyridine. Stir for 2 hr at 0◦ C, then leave the mixture standing at ambient temperature overnight. 15. Add 10 mL water and let stand 30 min. 16. Concentrate under vacuum to half the volume (bath temperature not exceeding 30◦ C). 17. Add 500 mL ethyl acetate and 200 mL water, transfer the solution to 1-L separatory funnel, and discard the aqueous layer. 18. Wash the organic layer with diluted (1:20) sulfuric acid in 100-mL portions until the aqueous phase remains acid (∼pH 3 to 6). 19. Wash with 100 mL water and then with saturated potassium hydrogencarbonate solution in 100-mL portions until the aqueous phase remains alkaline (∼pH 9 to 10). 20. Wash twice with 100 mL water. 21. Evaporate the organic phase to dryness under vacuum and co-distill the residue with 100 mL toluene. Check the progress of the reaction by TLC (APPENDIX 3D) in chloroform (product: Rf = 0.58 with UV detection; starting sugar: Rf = 0.32 with iodine vapor detection). 22. Dissolve the sharply evaporated residue from the product-containing fractions in the same volume of warm (30◦ to 35◦ C) diethyl ether, add hexane, and stir occasionally until the mixture remains permanently turbid. Scratch the flask walls to induce spontaneous crystallization, and leave in the refrigerator overnight. 23. Filter the chromatographically pure crystalline product, wash it with 150 mL hexane, and dry under vacuum. Use directly for synthesis of eritadenine. The yield of S.9 should be between 50 and 52 g (70% to 73%). Additional compound can be obtained from the mother liquor by silica gel column chromatography, although the quantity obtained is usually not worth the effort. The product can be stored in dry form in a closed bottle at 4◦ C for several months without signs of decomposition. Compound characterization: white crystals, m.p. 79◦ C. C16 H22 O7 S (358.4). 1 H NMR (400 MHz, CDCl3 ): δ 1.29, 1.45 (2 × s, 2 × 3H, CH3 -isopropylidene), 2.46 (s, 3H, CH3 -Tos), 3.24 (s, 3H, OCH3 ), 4.00, 4.03 (2 × dd, 2H, Jgem = 12.0 Hz, J5,4 = 6.9 Hz, H5), 4.31 (td, 1H, J4,5 = 6.9 Hz, J4,3 = 1.2 Hz, H4), 4.54 (d, 1H, J2,3 = 5.9 Hz, H2), 4.60 (dd, 1H, J3,2 = 5.9 Hz, J3,4 = 1.2 Hz, H3), 4.93 (s, 1H, H1), 7.36 (m, 2H, H-m-Tos), 7.81 (m, 2H, H-o-Tos).
SYNTHESIS OF 3-(ADENIN-9-YL)-2-HYDROXYPROPANOIC ACID (AHPA) The title compound (Fig. 14.1.5) is a very powerful inhibitor of SAH hydrolase, and one should therefore expect its biological activity in systems requiring methylations. This was proven by its antiviral activity, which is most probably connected with capping of viral mRNA. Most notable is its activity (or still better, the activity of its esters; Votruba et al., 1990) against poxviruses, not excluding variola (smallpox) virus. This protocol exemplifies the applicability of "classical" organic reactions to the synthesis of acyclic nucleosides by modification of the acyclic side chain. The limiting factor is the character of the heterocyclic base, which in many cases is inert to the reaction conditions. In other cases, the functional groups of the base (e.g., NH2 , OH, NH) may participate in or disturb the reaction, which thus cannot be performed at all or requires proper protection prior to the reaction.
BASIC PROTOCOL 3
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Figure 14.1.5
Synthesis of 3-(adenin-9-yl)-2-hydroxypropanoic acid (AHPA).
The synthesis of racemic 3-(adenin-9-yl)-2-hydroxypropanoic acid described here involves four steps with only one intermediate (Fig. 14.1.5). It is applicable to large-scale synthesis, since it makes use of inexpensive materials and isolation procedures. Alkylation of adenine with bromoacetaldehyde diethylacetal results in 9-(2,2-diethoxyethyl)adenine (S.14). This is the key intermediate (applicable also to other syntheses), which acts as a precursor of substituted acetaldehyde. This compound can be liberated by acid hydrolysis and undergoes cyanohydrin synthesis with potassium cyanide to form a cyanohydrin intermediate, which is directly acid-hydrolyzed to give the hydroxy acid (S.15).
Materials
Synthesis of Acyclic Analogs of Adenosine
Adenine Potassium carbonate Dimethylformamide Bromoacetaldehyde diethylacetal Celite Ethanol Conc. hydrochloric acid (HCl), reagent grade Chloroform Methanol Sodium cyanide Acetic acid Diethyl ether 2.5-L three-neck round-bottomed flask Mechanical KPG stirrer Reflux condenser Calcium chloride protecting tube Dropping funnel Magnetic stirrer with heating element and silicon oil bath
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8- to 10-cm-diameter glass filtration funnel with filter flask Rotary evaporator Additional reagents and equipment for TLC (APPENDIX 3D) NOTE: All solvents and chemicals are reagent grade.
Synthesize 9-(2,2-diethoxyethyl)adenine 1. Add 100 g (0.74 mol) adenine and 120 g potassium carbonate to a 2.5-L three-neck round-bottom flask equiped with a mechanical KPG stirrer, reflux condenser with calcium chloride protecting tube, and dropping funnel, and set in a silicon oil bath. Pour in 1500 mL DMF. 2. Warm the stirred suspension to 140◦ C and add dropwise over 1 hr 190 g bromoacetaldehyde diethyl acetal. Stir at 140◦ C 16 hr. 3. Filter the hot suspension through a glass filter with a Celite pad, wash the precipitate with 150 mL DMF, and leave the filtrate to crystallize at room temperature overnight. 4. Filter the crystalline product, wash with 100 mL ethanol, and recrystallize from 1.5 L ethanol. 5. Combine all filtrates (ethanol and DMF) and evaporate under vacuum. Recrystallize the residue twice from ethanol. The yield of S.14 should be between 118 and 120 g (47% to 48%). It can be stored at room temperature without decomposition. Compound characterization: white leaflets; m.p. 218◦ -219◦ C. C11 H17 N5 O2 (251.3): 52.57% C, 6.82% H, 27.87% N. 1 H NMR (400 MHz, DMSO-d6 ): δ 1.02 (t, 6H, Jvic = 7.0 Hz, CH3 ), 3.41, 3.64 (2 × dq, 4H, Jgem = 9.6 Hz, Jvic = 7.0 Hz, CH2 -O), 4.22 (d, 2H, Jvic = 5.4 Hz, CH2 -N), 4.84 (t, 1H, Jvic = 5.4 Hz, CH), 7.23 (bs, 2H, NH2 ), 8.05, 8.15 (2 × s, 2 × 1H, H2 + H8).
Synthesize 3-(adenin-9-yl)-2-hydroxypropanoic acid (AHPA) 6. Add 16 mL conc. HCl to a suspension of 30 g (0.12 mol) S.14 in 600 mL water. CAUTION: These steps make use of and evolve strong poison. It is essential to work in a well-ventilated chemical fume hood and to wear gloves.
7. Heat the solution under reflux until the reaction is complete, as determined by TLC using 6:3 (v/v) chloroform/methanol (the product is substantially slower than the starting material). It is necessary to neutralize the sample on the TLC plate. To achieve this, hold the start line for several seconds over an open bottle of conc. aqueous ammonia solution.
8. Cool the mixture to –5◦ C, add 30 g (0.612 mol) sodium cyanide, and bring the pH of the mixture as quickly as possible to pH 6 to 7 by adding acetic acid. 9. Stir 5 hr at 0◦ C and then overnight at room temperature. Add 400 mL conc. HCl and reflux for 5 hr. 10. Evaporate under vacuum and coevaporate five to six times with 100 mL water to remove most of the HCl. 11. Add 100 mL ice-cold water and filter. Wash with 50 mL water, 50 mL ethanol, and finally 200 mL diethyl ether. Dry under vacuum. The yield of S.15 should be between 22 and 25 g (61% to 70%). The product can be stored at room temperature without decomposition.
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Compound characterization: white crystals, m.p. > 260◦ C. C8 H9 N5 O3 (223.23): 43.05% C, 4.06% H, 31.3% N. UV spectrum (pH 2): λmax 261 (εmax 13500). 1 H NMR (400 MHz, D2 O, ref(dioxane) = 3.75 ppm): δ 4.32 (dd, 1H, Jgem = 13.8 Hz, J1 b,2 = 7.2 Hz, H1 b), 4.38 (dd, 1H, J2 ,1 = 7.2, 3.1 Hz, H2 ), 4.44 (dd, 1H, Jgem = 13.8 Hz, J1 a,2 = 3.1 Hz, H1 a), 8.01, 8.06 (2 × s, 2 × 1H, H2 + H8).
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Amberlite IR 45 resin, OH form To convert Amberlite IR 45 to the hydroxyl form, stir it for 3 hr with 0.5 M NaOH, filter through a glass filter, and wash with water (carbon dioxide free) immediately before use. Convert the residual unused material also to the acetate form as for Dowex 1 (see recipe). p-Anisaldehyde reagent To prepare p-anisaldehyde reagent, dissolve 9.2 mL p-methoxybenzaldehyde, 37.5 mL acetic acid, and 12.5 mL sulfuric acid in 338 mL of 96% (v/v) ethanol. The reagent is stable for several months at room temperature. Spray uniformly on the developed and air-dried TLC plate. Dry the plate and heat briefly with a heat gun to visualize the brown-violet spots on a crimson background. CAUTION: Work in a well-ventilated chemical fume hood!
Dowex 1 × 2 resin, acetate form Suspend 500 g Dowex 1 × 2 resin (100-200 mesh) in 0.5 M NAOH and let it stand overnight. Wash with 2 L water, resuspend in 2 L water, and add acetic acid to ∼1 M. Let stand overnight. Wash briefly with water. Store at room temperature. To regenerate resin batchwise, wash with 2 L water and let stand overnight in 0.5 to 1 M NaOH. Wash briefly with water and let stand overnight in 2 M acetic acid. Wash briefly with water. Keep wet resin at room temperature (stable for years). The recycling can be repeated for years. It is essential not to wash the resin too extensively with water, as it will become sensitive to microbial infection, which can destroy it completely, particularly at higher room temperatures. It is recommended to wash the recycled resin occasionally with ethanol.
Dowex 50 × 8, acid form Wash 500 g Dowex 50 × 8 resin (100-200 mesh) briefly with 2 L water, suspend in 2 L of 2 M HCl, and let stand overnight in a chemical fume hood. Filter through a glass filter and then wash with water until the pH of the filtrate is neutral. Store at room temperature. The resin capacity is ∼1 mequiv/mL.
To regenerate resin batchwise, wash with 2 L water and let stand overnight in 0.5 to 1 M NaOH. Wash briefly with water and let stand overnight in 2 M HCl. Filter through a glass filter and then wash with water until the pH of the filtrate is neutral. Keep at room temperature (stable for years). The recycling can be repeated for years.
Synthesis of Acyclic Analogs of Adenosine
Fehling solutions Fehling solution A: 69.3 g/L copper sulfate pentahydrate in water. Fehling solution B: 364 g/L potassium sodium tartrate tetrahydrate and 100 g/L NaOH in water. Store both solutions several years at room temperature in tightly closed bottles.
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COMMENTARY Background Information Acyclic nucleosides are by definition nucleoside analogs in which the aldopentofuranose ring is replaced with an alkyl, alkenyl, or alkynyl chain bearing at least one hydroxyl group. This chain is attached to the heterocyclic base at nitrogen atoms and can be linear or branched. This broad definition can have additional limitations, including compounds wherein the heterocyclic base is linked directly to a cycloalkyl residue (carbocyclic nucleosides). The least precise but more natural limitation is the definition of compounds that behave like nucleosides, i.e., that interact with the enzymes of nucleic acid metabolism, undergo phosphorylation by nucleoside kinases, or replace nucleosides at an enzyme binding site to prevent the enzyme-catalyzed reaction or to undergo the appropriate transformation themselves. Thus, N-hydroxymethyl or N-(2-hydroxyethyl) derivatives can hardly be considered analogs of nucleosides in general. However, the classification becomes difficult with hydroxyl-bearing N-propyl derivatives, as some 9-(2-hydroxypropyl)purines clearly interact with herpesvirus thymidine kinase (Pospisil et al., 2002), and 9-(S)-(2,3dihydroxypropyl)adenine behaves unequivocally as an adenosine analog that interacts with S-adenosylhomocysteine (SAH) hydrolase by replacing adenosine at the enzyme binding site (Votruba and Hol´y, 1980). On the other hand, both the 7-substituted (S)and (R)-(2,3-dihydroxypropyl)theophyllines (diprophylline) with bronchodilatory activity behave simply as water-soluble coffeine derivatives in the interaction with cyclic AMP phosphodiesterases (Hol´y and Vanecek, 1979). Similarly, 9-erythro-(2hydroxynonyl)adenine (EHNA), a strong inhibitor of adenosine aminohydrolase, probably should not be considered an acyclic analog of adenosine, but rather an N-alkyladenine with a hydrophobic substituent. The attention paid to the study of acyclic nucleoside analogs is substantiated by their chemical and metabolic stability (Hol´y and Cih´ak, 1981; Votruba et al., 1983). The nucleoside linkage in classical base-modified nucleosides is labile, in particular in purine 2deoxyribonucleosides, which easily decompose in acidic media. Enzymatic degradation of nucleosides at the nucleoside linkage is catalyzed by hydrolases or phosphorylases, depending on the reaction mechanism. This process is massive but very undesirable, particu-
larly in the context of nucleoside-based drugs in cancer chemotherapy or in the treatment of viral diseases. Liberation of the base may create undesirable toxicity problems; lowering the level of active drug requires a more frequent application regimen and/or enhanced drug doses, and thus increases the patient’s body strain. Replacement of the aldopentose residue by an acyclic chain dramatically alters the character of the nucleoside linkage, which turns the labile hemiacetal bond to an enzymatically (and chemically) stable N-alkyl linkage (De Clercq and Hol´y, 1979). Contrary to the comparatively rigid conformation of the classical nucleoside, which must fit to the active site of the enzyme, the high flexibility of the C–C linkages (especially those of sp3 character) allows the acyclic nucleoside molecule to adopt the optimum conformation of the molecule to minimize the energy of its complex with the enzyme. There are two main groups of acyclic nucleosides: one where the side chain bears no asymmetry center and one where it contains one or more asymmetry centers. The wellknown group of acyclic guanine nucleoside antivirals (e.g., acycloguanosine, ganciclovir, penciclovir) belongs to the former category. The far-fetched similarity between guanosine and 9-[2-(hydroxyethoxy)methyl]guanine (acycloguanosine) is not sufficient for cellular nucleoside kinases to recognize this compound as a substrate for phosphorylation. The viral enzyme HSV-1 thymidine kinase, however, can recognize acycloguanosine and thereby initiates the sequence of events (intracellular processes) that leads to inhibition of DNA polymerases. Consequently, this activation takes place solely in the infected cells, so that drug toxicity is negligible. The latter (chiral) group comprises, among others, reversible inhibitors of SAH hydrolase such as 9-(S)-(2,3-dihydroxypropyl)adenine (DHPA; Hol´y 1975, 1978a,b), with broadspectrum biological (including antiviral) activity (De Clercq et al., 1978), and the irreversible SAH inactivators eritadenine (Hol´y et al., 1982, 1985a,b; Votruba and Hol´y, 1982) and 3-(adenin-9-yl)-2-hydroxypropanoic acid (AHPA; Hol´y, 1984; De Clercq and Hol´y, 1985). Contrary to the majority of modified nucleosides, which act by inhibiting nucleic acid synthesis, the mechanism of SAH hydrolase inhibitors is based on the inhibition of SAMmediated methylation in proliferating cells (De Clercq et al., 1981; Jelinek et al., 1981;
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Sl´ama et al., 1983). These three SAH inhibitors are the focus of this unit. To date, there are numerous SAH-hydrolase inhibitors, in particular compounds with carbocyclic nucleoside structure, fluorinated sugar and other sugar modifications, 3deazaadenine nucleosides, and so on. Some exhibit very powerful antiviral activity aimed at RNA viruses. There are also different modes of SAHase inhibition, including reversible inhibition, covalent binding, and suicide inhibition. The adenine acyclic nucleosides selected here also differ in mechanism. While the reversible inhibition and metabolic inertia of (S)-DHPA makes it a useful tool for investigation of processes in proliferating systems, the two adenine-derived carboxylic acids, Deritadenine and AHPA, are fast-acting inhibitors capable of depleting SAHase activity in the cell to the core level. Their capacity has not yet been fully exploited, though their preparation is much simpler compared to the above nucleoside analogs.
groscopic nature of the salt and by the large volume of highly inflammable solvents used for its preparation.
Anticipated Results The yields for the individual preparations are given at the appropriate sites in the text. It is essential to use dry DMF, and it is recommended to distill it under vacuum from phosphorus pentoxide leaving some distillation residue, and to store it over molecular sieves. With this treatment, the solvent is stable for a minimum of 5 months at room temperature. This procedure also removes dimethylamine. The alkylations (steps 1 in Basic Protocols 1, 2 and 3) proceed quickly at elevated temperatures, and the optimum temperature indicated should be reached as soon as possible. However, it is not recommended to exceed 120◦ C bath temperature. All the alkylation products are sufficiently stable and their isolation should not cause any problems.
Time Considerations Critical Parameters and Troubleshooting
Synthesis of Acyclic Analogs of Adenosine
All the methods described here were verified on a nominal or higher scale. The potential hazards are identified and discussed within the text. Except for the notorious sodium cyanide, there is no highly poisonous chemical used in the synthetic steps. The sodium salt of adenine, which is made by the action of sodium hydride on adenine, makes use of a comparatively safe reagent protected by mineral oil. This oil is not removed and thus remains in the reaction mixtures until removed by crystallization or chromatography. It has been mentioned that there may be difficulties caused by the physical behavior of the sodium salt of adenine in DMF. The slurry requires approximately twice the volume of the solvent to be stirrable. When the procedure should be scaled up, it is recommended to use the cesium salt of adenine instead. This is easily obtained by replacing sodium hydride with cesium carbonate (0.5 equiv with respect to adenine). This applies specifically to step 1 in Basic Protocols 1 and 2. The third alternative is to prepare the crystalline sodium salt of adenine separately by diethyl ether precipitation (3 vol) of a 1 M adenine solution in 1 M sodium methoxide, and then add the dried material directly to the solution of synthon in the corresponding volume of DMF. This access, though apparently very easy, is complicated by the highly hy-
Because some of these procedures are lengthy, time requirements are given both in calendar days and in terms of active work performed by laboratory personnel. DHPA. The approximate average time required to prepare the S isomer of DHPA (Basic Protocol 1) is 3-4 days for the dioxolane intermediate (depending on the heating regimen) and 2 days for the final product. The period requiring immediate attention is not higher than 5 hr and 3 hr, respectively. Preparation of the starting tosylated dioxolane reagent (Support Protocol 1) requires an additional 2 days (depending on the rate of hydrolysis), with not more than 5 hr of hands-on time. For the RS racemic mixture of DHPA (Alternate Protocol), preparation of the dioxolane intermediate requires a comparable amount of time as for Basic Protocol 1. Preparation of the final product requires 2 days (depending on the rate of hydrolysis), with up to 5 hr hands-on time. Eritadenine. Preparation of the 2 ,3 O-isopropylidene-5 -O-tosyl-protected ribose starting reagent from D-ribose (Support Protocol 2) requires an initial 3 days, with 9 to 10 hr of manual work. Once this reagent is in hand, preparation of the 2,3-O-protected nucleoside intermediate requires 2 days, with 5 hr hands-on time. Final preparation of eritadenine by Neff’s degradation and subsequent purification of the product require approximately 1 week, depending on the efficacy of evaporation of large water volumes and on the
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automatization of silica gel chromatography. As an alternative, it is recommended to use flash chromatography with original silica gel columns (e.g., ISCO). AHPA. Preparation of the intermediate 9-(2,2-diethoxyethyl)adenine and subsequent conversion to the final product require 2 days each, with hands-on time of no more than 4 and 5 hr, respectively.
Literature Cited De Clercq, E. and Hol´y, A. 1979. Antiviral activity of aliphatic nucleoside analogs: Structurefunction relationship. J. Med. Chem. 22:510513. De Clercq, E. and Hol´y, A. 1985. Alkyl esters of 3-adenin-9-yl-2-hydroxypropanoic acid: A new class of broad-spectrum antiviral agents. J. Med. Chem. 28:282-287. De Clercq, E., Descamps, J., De Somer, P., and Hol´y, A. 1978. (S)-9-(2,3-Dihydroxypropyl)adenine: An aliphatic nucleoside analog with broad-spectrum antiviral activity. Science 200:563-564. De Clercq, E., Leyten, R., Sobis, H., Matousek, J., Hol´y, A., and De Somer, P. 1981. Inhibitory effect of a broad-spectrum antiviral agent, (S)-9(2, 3-dihydroxypropyl)adenine, on spermatogenesis in mice. Toxicol. Appl. Pharmacol. 59:441451. Hol´y, A. 1975. Aliphatic analogs of nucleosides, nucleotides and oligonucleotides. Collect. Czech. Chem. Commun. 40:187-214. Hol´y, A. 1978a. Synthesis of N-(2,3dihydroxypropyl) derivatives of heterocyclic bases. Collect. Czech. Chem. Commun. 43:2054-2061. Hol´y, A. 1978b. Synthesis of some 2,3dihydroxypropyl derivatives of purine bases. Collect. Czech. Chem. Commun. 43:3103-3117. Hol´y, A. 1984. Preparation and synthetic utilization of 3-(adenin-9-yl)-2-hydroxyalkanoic acids and their derivatives. Collect. Czechoslov. Chem. Commun. 49:2148-2166. Hol´y, A. and Cih´ak, A. 1981. Metabolism of 9(S)-(2,3-dihydroxypropyl)adenine, an antiviral agent, in mice. Biochem. Pharmacol. 30:23592361. Hol´y, A. and Vanecek, M. 1979. Synthesis and pharmacological properties of enantiomeric derivatives of 7-(2,3-dihydroxypropyl)theophylline. Collect. Czech. Chem. Commun. 44:2550-2555.
Hol´y, A., Votruba, I., and De Clercq, E. 1982. Synthesis and antiviral properties of stereoisomeric eritadenines. Collect. Czech. Chem. Commun. 47:1392-1407. Hol´y, A., Votruba, I., and De Clercq, E. 1985a. Structure-activity studies on open-chain analogs of nucleosides: Inhibition of S-adenosyl-Lhomocysteine hydrolase and antiviral activity. 1. Collect. Czech. Chem. Commun. 50:245-261. Hol´y, A., Votruba, I., and De Clercq, E. 1985b. Structure-activity studies on open-chain analogs of nucleosides: Inhibition of S-adenosyl-Lhomocysteine hydrolase and antiviral activity. 2. Collect. Czech. Chem. Commun. 50:262-279. Jel´ınek, R., Hol´y, A., and Votruba, V. 1981. Embryotoxicity of 9-(S)-(2,3-dihydroxypropyl)adenine. Teratology 24:267-275. Pospisil, P., Pilger, B.D., Marveggio, S., Schelling, P., Wurth, C., Scapozza, L., and Folkers, G. 2002. Synthesis, kinetics, and molecular docking of novel 9-(2-hydroxypropyl)purine nucleoside analogs as ligands of herpesviral thymidine kinases. Helv. Chim. Acta 85:3237-3250. Sl´ama, K., Hol´y, A., and Votruba, I. 1983. Insect sterility induced by a broad-spectrum antiviral agent, 9-(2,3-dihydroxypropyl)adenine. Entomol. Exp. Appl. 33:9-14. Votruba, I. and Hol´y, A. 1980. Inhibition of S-adenosyl-L-homocysteine hydrolase by the aliphatic nucleoside analog 9-(S)(2,3-dihydroxypropyl)adenine. Collect. Czech. Chem. Commun. 45:3039-3044. Votruba, I. and Hol´y, A. 1982. Eritadenines— novel type of potent inhibitors of S-adenosyl-Lhomocysteine hydrolase. Collect. Czech. Chem. Commun. 47:167-174. Votruba, I., Hol´y, A., and De Clercq, E. 1983. Metabolism of the broad-spectrum antiviral agent, 9-(S)-(2,3-dihydroxypropyl)adenine, in different cell lines. Acta Virol. 27:273-276. Votruba, I., Hasobe, M., Hol´y, A., and Borchardt, R.T. 1990. 2-Methylpropyl ester of 3-(adenin9-yl)-2-hydroxypropanoic acid. Mechanism of antiviral action in vaccinia virus-infected L929 cells. Biochem. Pharmacol. 39:1573-1580.
Contributed by Antonin Hol´y Institute of Organic Chemistry and Biochemistry Academy of Sciences of the Czech Republic Prague, Czech Republic
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Synthesis of Acyclic Nucleoside Phosphonates
UNIT 14.2
In this unit, the group of N-(2-phosphonomethoxy)ethyl (PME) compounds is first represented by the procedure for preparation of a very efficient antiviral agent, PMEA (adefovir). The PME group is further represented by the 2,6-diaminopurine analog, which has strong antiretroviral and antineoplastic activity, and by the 2-amino-6-chloropurine derivative, which is a common intermediate for further compounds, represented here by the guanine derivative PMEG and by PME-N6 -cyclopropyl-2,6-diaminopurine. Following these related compounds, the so-called N-(R)-2-(phosphonomethoxy)propyl (PMP) derivatives are represented by the adenine derivative (tenofovir), which is the active component (Palmer et al., 2000) formed from the prodrug disoproxil tenofovir fumarate (the anti-AIDS drug Viread). Finally, a procedure is presented for preparation of N-(S)-[3hydroxy-2-(phosphonomethoxy)propyl]adenine (HPMPA); both HPMPA and HPMPC (cidofovir) exhibit strong antiviral activity directed against DNA viruses (De Clercq et al., 1986). Acyclic nucleoside phosphonates (ANPs) can be prepared by two distinct synthetic approaches: a two-step procedure in which alkylation of the nucleobase is followed by attachment of a phosphonomethyl ether group, and a synthon procedure in which attachment of the entire side chain is achieved in a single condensation step between the nucleobase and an appropriate synthon. The methods presented here for synthesis of PME derivatives all utilize a common PME synthon. PMPA synthesis is presented using both a synthon approach and a two-step approach. Support protocols are provided for preparation of the PME and PMP synthons. Synthesis of the HPMP derivatives uses a two-step approach. If not stated otherwise, the solutions are evaporated on rotary evaporator at 1.5 to 2 kPa with a maximum bath temperature of 40◦ C. The compounds are dried at 1.5 Pa over phosphorus pentoxide. Analytical TLC uses Merck silica gel plates (UV254; 10-cm runs) in systems based on chloroform/methanol or toluene/ethyl acetate mixtures, with detection under UV light or (for nonabsorbing compounds) by iodine vapors, p-anisaldehyde, or p-nitrobenzylpyridine (see Reagents and Solutions). Silica gel supplied by Sigma is used for preparative column chromatography. Filtration of solutions is performed by suction through a Celite pad in a glass funnel. Crystallization of the compounds is generally performed from saturated solutions made by refluxing in the appropriate solvent, which are then filtered while hot through glass or tight cottonwool filter. Unless stated otherwise, crystallization is performed at 4◦ C.
SYNTHESIS OF 9-[2-(PHOSPHONOMETHOXY)ETHYL]ADENINE (ADEFOVIR, PMEA) This compound belongs to the most important ANPs, at least from the viewpoint of drug development (De Clercq et al., 1986, 1987; Naesens and De Clercq, 1997). It had reached the point of advanced clinical studies before it was found that at the dose efficient for treatment of AIDS (Balzarini et al., 1989, 1991) certain values of characteristic clinical parameters were out of the tolerated errors. However, adefovir also very actively suppresses reverse transcription in the life cycle of hepatitis B virus (Naesens et al., 1994). The dosage for this application is so much lower that the negative symptoms do not appear at all. In the oral prodrug form, adefovir dipivoxil is approved for hepatitis B therapy, specifically for lamivudine-resistant patients (Hadziyannis et al., 2003).
Contributed by Antonin Hol´y Current Protocols in Nucleic Acid Chemistry (2005) 14.2.1-14.2.38 C 2005 by John Wiley & Sons, Inc. Copyright
BASIC PROTOCOL 1
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14.2.1 Supplement 22
Figure 14.2.1 Synthesis of (A) 9-[2-(phosphonomethoxy)ethyl]adenine (PMEA) and 9-[2-(phosphonomethoxy) ethyl]-2,6-diaminopurine (PMEDAP) (B). S.16 and S.19 are the PME and C1 synthons, respectively. X = OTs.
The synthesis described below (Fig. 14.2.1A; Hol´y et al., 1989a) is simple and uses the synthon approach, in which alkylation of adenine is performed in a single step using a PME synthon, diisopropyl 2-chloroethoxymethylphosphonate (S.16; prepared in Support Protocol 1). The sodium salt of adenine is heated with the synthon in DMF, and the diisopropyl ester intermediate (S.2) is purified and cleaved with bromotrimethylsilane. After hydrolysis and deionization, the product (S.3) is purified by anion-exchange chromatography or crystallized in the zwitter-ionic form (Hol´y et al., 1989a).
Materials
Synthesis of Acyclic Nucleoside Phosphonates
Sodium hydride, 60% dispersion in paraffin oil Dimethylformamide (DMF) Adenine (S.1) Diisopropyl 2-chloroethoxymethylphosphonate (S.16; see Support Protocol 1) Toluene Chloroform Celite Silica gel Methanol Ethyl acetate Hexane Bromotrimethylsilane
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Acetonitrile Conc. aqueous ammonia Dowex 50 × 8, acid form (see recipe) 1:1 (v/v) diluted HCl (from conc. hydrochloric acid and water) Ethanol Diethyl ether 500-mL and 1-L round-bottom flasks Magnetic stirrer with heating plate and silicon oil bath 250-mL dropping funnel Rotary evaporator with diaphragm pump Glass filter funnels with 1-L filter flasks Reflux condenser 6 × 40–cm and 5 × 40–cm chromatography columns Fraction collector (optional) UV detector (optional) 2-L thick-wall beaker Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Synthesize 9-[2-(diisopropylphosphonylmethoxy)ethyl]adenine 1. Place 4.0 g of 60% sodium hydride in paraffin oil (2.4 g, 0.1 mol NaH) in 350 mL DMF in a 1-L round-bottom flask, and add 13.5 g (0.1 mol) adenine while stirring. Continue to stir 1 hr at 80◦ C. Prior to the reaction, check the quality of the DMF by adding several milligrams of sodium hydride suspension to 2 mL DMF in a reagent tube. Do not use DMF if sodium hydride reacts with foaming, which indicates the decomposition of DMF and the presence of formic acid. No special drying of the flask is necessary. The flask and other glassware used in the procedure can be rinsed with ethanol or acetone and air dried. CAUTION: Perform the reaction in a well-ventilated chemical fume hood. Wear protective gloves while working with sodium hydride. It is water sensitive. Wash everything that comes into contact with sodium hydride first with ethanol and then with water.
2. While stirring, use a 250-mL dropping funnel to add dropwise (over 15 min) 28 g (0.11 mol) diisopropyl 2-chloroethoxymethylphosphonate (S.16) in 100 mL DMF. Stir 16 hr at 110◦ C. 3. Evaporate under vacuum on a rotary evaporator, and co-distill the residue twice with 100 mL toluene. 4. Add 300 mL chloroform and boil under reflux for 10 min. Decant and vacuum filter the extract through Celite. Repeat this step twice more. 5. Evaporate the combined extracts under vacuum. 6. Apply the concentrated chloroform solution onto a 6 × 40–cm column containing 300 mL silica gel in chloroform. Elute the column with 1 L chloroform and then with 95:5 (v/v) chloroform/methanol. Check the fractions by TLC (APPENDIX 3D) in the same solvent system (Rf = 0.47). 7. Combine the product-containing fractions and evaporate under vacuum. 8. Dissolve the residue in a minimum volume of boiling ethyl acetate, add hexane slowly until the solution remains turbid (i.e, does not clear with stirring). Scratch the wall of the flask with a spatula to initiate crystallization, and leave to stand overnight at 0◦ C.
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9. Filter, wash with 100 mL hexane, and dry under vacuum. The yield of S.2 should be between 22 and 25 g (61% to 70%). It can be stored at room temperature indefinitely without decomposition. Compound characterization: m.p. 138◦ C. C14 H24 N5 O4 P (357.3). 1 H NMR (DMSO-d6 ): δ 1.13, 1.18 (2 × d, 2 × 6H, 4CH3 ), 3.79 (d, 2H, J(P-CH) = 8.40 Hz, O-CH2 -P), 3.91 (t, 2H, H2 ), 4.34 (t, 2H, J1 ,2 = 5.0 Hz, H1 ), 4.46 (dsept, 2H, JCH,CH2 = 6.1 Hz, J(P-OCH) = 7.7 Hz, P-O-CH), 7.19 (br s, 2H, NH4.34 ), 8.09 (s, 1H, H8), 8.15 (s, 1H, H2).
Synthesize 9-[2-(phosphonomethoxy)ethyl]adenine 10. Add 20 mL bromotrimethylsilane to 25 g (70 mmol) S.2 in 150 mL acetonitrile in a 500-mL round-bottom flask. Quickly close with a stopper and dissolve by shaking the flask by hand. 11. Let the solution stand overnight at ambient temperature, and then evaporate under vacuum. 12. Add 100 mL water followed by conc. ammonia solution until a strongly alkaline pH is achieved (∼pH 10). Evaporate under vacuum. 13. Dissolve the residue in 300 mL water and add as much Dowex 50 × 8 (acid form) as needed to redissolve the originally precipitated material. 14. Pour the suspension onto a 5 × 40–cm column containing 300 mL of the same resin and wash with 2 L water. 15. Transfer the resin from the column to a 2-L thick-wall beaker and add 500 mL water followed by enough conc. ammonia solution to make the solution strongly alkaline. 16. Filter, wash the resin with twice with 200 mL water, and evaporate under vacuum. 17. Dissolve the residue in 200 mL boiling water and adjust to pH 3 with 1:1 (v/v) diluted HCl. Add 200 mL ethanol, scratch the wall of the flask with a spatula to initiate crystallization, and set in the refrigerator overnight. 18. Collect the product by filtration. Wash with ∼50 mL of 50% (v/v) aqueous ethanol followed by 100 mL ethanol and then 100 mL diethyl ether. Dry under vacuum. The yield of S.3 should be ∼17 g (90%). It can be stored at room temperature indefinitely without decomposition. Compound characterization: m.p. 301◦ C. C8 H12 N5 O4 P (273.2). 1 H NMR (500 MHz, D2 O+NaOD, ref(dioxane) = 3.75 ppm): δ 3.51 (d, 2H, JH,P = 8.5 Hz, H4 ), 3.94 (t, 2H, J2 ,1 = 4.9 Hz, H2 ), 4.40 (t, 2H, J1 ,2 = 4.9 Hz, H1 ), 8.17 (s, 1H, H2), 8.23 (s, 1H, H8). 13 C NMR (125.8 MHz, D2 O+NaOD, ref(dioxane) = 69.3 ppm): 46.23 (CH2 -1 ), 71.07 (d, JC,P = 152 Hz, CH2 -4 ), 72.85 (d, JC,P = 10 Hz, CH2 -2 ), 120.94 (C5), 145.80 (CH-8), 151.53 (C4), 154.91 (CH-2), 158.08 (C6). BASIC PROTOCOL 2
Synthesis of Acyclic Nucleoside Phosphonates
SYNTHESIS OF 2,6-DIAMINO9-[2-(PHOSPHONOMETHOXY)ETHYL]PURINE (PMEDAP) PMEDAP is renowned for its extraordinary efficacy against retroviruses and DNA viruses (Naesens et al., 1989, 1993), but its main target is antineoplastic activity. This activity has been proven both for the compound alone (Otov´a et al., 1997, 1999; Bobkov´a et al., 2000) and in combination with paclitaxel derivatives (Bobkov´a et al, 2001). The preparation (Fig. 14.2.1B) follows that of Basic Protocol 1 for PMEA, except for the use of cesium carbonate for the primary condensation. From the alkali metal carbonate series (sodium, potassium, rubidium, and cesium), cesium carbonate gives the best yields (Dvor´akov´a et al., 1993). However, it is also possible to use the sodium salt of 2,6-diaminopurine,
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prepared in situ in analogy with the sodium salt of adenine (as described in Basic Protocol 1 or in UNIT 14.1; Hol´y et al., 1999).
Materials 2,6-Diaminopurine (S.5; TCI America) Cesium carbonate Dimethylformamide (DMF) Diisopropyl 2-chloroethoxymethylphosphonate (S.16; see Support Protocol 1) Celite Toluene Chloroform Silica gel Methanol Ethanol Hexane Acetonitrile Bromotrimethylsilane Conc. aqueous ammonia 1:1 (v/v) diluted HCl (from conc. hydrochloric acid and water) Diethyl ether 1-L round-bottom flasks Magnetic stirrer with heating plate and silicon oil bath Glass filter funnels with 1-L filter flasks Rotary evaporator with diaphragm pump Reflux condenser 6 × 40–cm chromatography column Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Synthesize 2,6-diamino-9-[2-(diisopropylphosphonylmethoxy)ethyl]purine 1. Stir a mixture of 15 g (0.1 mol) 2,6-diaminopurine, 16.5 g (0.05 mol) cesium carbonate, and 250 mL DMF in a 1-L round-bottom flask for 30 min at 100◦ C. 2. Add 32.5 g (0.125 mol) diisopropyl 2-chloroethoxymethylphosphonate (S.16) and stir 10 hr at 120◦ C. 3. Filter while hot through a Celite pad and wash the insoluble material with 50 mL DMF. 4. Evaporate under vacuum on a rotary evaporator, and co-distill the residue twice with 100 mL toluene. 5. Add 300 mL chloroform and boil under reflux for 20 min. Decant and vacuum filter the extract through Celite. Repeat this step twice more. 6. Evaporate the combined extracts under vacuum. 7. Apply the concentrated chloroform solution onto a 6 × 40–cm column containing 300 mL silica gel in chloroform. Elute the column with 1 L chloroform and then continue with 95:5 (v/v) chloroform/methanol to elute the product. Check the fractions by TLC (APPENDIX 3D) in the same solvent system (Rf = ∼0.40). 8. Combine the product-containing fractions and evaporate under vacuum. Biologically Active Nucleosides
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9. Dissolve the purified product in ∼200 mL boiling ethanol, add hexane until the solution remains turbid (i.e, does not clear with stirring). Scratch the wall of the flask to initiate crystallization, and allow to stand overnight at 0◦ C. 10. Collect the product by filtration, wash with 100 mL hexane, and dry under vacuum. The yield of S.6 should be ∼20 to 25 g (60% to 75%). It can be stored at room temperature indefinitely without decomposition. Compound characterization: m.p. 193◦ -194◦ C. C14 H25 N6 O4 P (372.4). 1 H NMR (DMSOd6 ): δ 1.16, 1.18 (2 × d, 2 × 6H, 4CH3 ), 3.77 (d, 2H, J(P-CH) = 8.3 Hz, P-CH2 ), 3.82 (t, 2H, H2 ), 4.34 (t, 2H, J1 ,2 = 5.0 Hz, H1 ), 4.52 (dsept, 2H, JCH,CH2 = 6.2 Hz, J(P-OCH) = 7 Hz, P-O-CH), 5.78 (br s, 2H, NH2 ), 6.67 (br s, 2H, NH2 ), 7.65 (s, 1H, H8).
Synthesize 2,6-diamino-9-[2-(phosphonomethoxy)ethyl]purine 11. Place 18.6 g (0.05 mol) S.6 in a 1-L round-bottom flask and add 400 mL acetonitrile and 40 mL bromotrimethylsilane. Quickly close with a stopper and dissolve by manual shaking. 12. Let the solution stand overnight at room temperature and evaporate under vacuum. 13. Add 50 mL water and then add conc. aqueous ammonia to give a strongly alkaline reaction (∼pH 10 to 10.5). Evaporate under vacuum. 14. Co-distill with 50 mL water, and then dissolve in 600 mL boiling water. 15. Under magnetic stirring, bring pH to ∼3.5 with 1:1 diluted HCl, and let stand overnight at room temperature. 16. Collect the crystalline material by filtration. Wash with 50 mL water, then 50 mL ethanol, and then 100 mL diethyl ether. Dry under vacuum. The yield of S.7 should be ∼14 g (97%). It can be stored at room temperature indefinitely without decomposition. Compound characterization. m.p. >300◦ C. C8 H13 N6 O4 P (288.2). 1 H NMR (500 MHz, D2 O+NaOD, ref(dioxane) = 3.75 ppm): δ 3.55 (d, 2H, JH,P = 8.5 Hz, H4 ), 3.90 (t, 2H, J2 ,1 = 5.4 Hz, H2 ), 4.20 (t, 2H, J1 ,2 = 5.4 Hz, H1 ), 7.87 (s, 1H, H8). 13 C NMR (125.8 MHz, D2 O+NaOD, ref(dioxane) = 69.3 ppm): 45.75 (CH2 -1 ), 70.91 (d, JC,P = 153 Hz, CH2 -4 ), 72.88 (d, JC,P = 11, CH2 -2 ), 115.14 (C5), 143.23 (CH-8), 153.37 (C4), 158.02 (C6), 161.89 (C2). BASIC PROTOCOL 3
Synthesis of Acyclic Nucleoside Phosphonates
SYNTHESIS OF 9-[2-(PHOSPHONOMETHOXY)ETHYL]GUANINE (PMEG) Although all guanine-derived acyclic nucleotide analogs are too toxic to be of any specific therapeutic use, they continue to have scientific importance, particularly in cases where the selectivity ratio is less unfavorable (Andrei et al., 1998; De Clercq et al., 1999). PMEG has an important therapeutic potential in treatment of warts caused by papilloma viruses (HPV), which often become malignant (Kreider et al., 1990). To circumvent the problem of a narrow margin between toxic and therapeutic dosages, the structurally related derivative 2-amino-6-cyclopropylaminopurine (see Basic Protocol 4) can be used instead of PMEG (Compton et al., 1999). This compound is converted in the cell to PMEG and thus acts as a prodrug of the latter. Unlike adenine, guanine cannot be directly alkylated to afford the required N9 isomer; the main regioisomer obtained is the N7-alkylguanine. To circumvent this problem, it is possible to use 2-amino-6-chloropurine (S.8; Fig. 14.2.2) as an alkylation substrate. Alkylation of this base gives the N9 isomer (S.9) as the major product, though the amount of other regioisomers is considerable. The 2-amino-6-chloropurine intermediate can be then hydrolyzed in alkali or, preferably, by a mineral acid to give the guanine derivative (S.10). Deprotection gives the final product (S.11). The same intermediate can
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Figure 14.2.2 analog.
Synthesis of 9-[2-(phosphonomethoxy)ethyl]guanine (PMEG) and its 2-amino-6-cyclopropylamino
be used for preparation of related analogs, including N6 -substituted 2,6-diaminopurine derivatives of the PMEDAP series by the reaction with primary or secondary amines, or 2-amino-6-sulfanyl derivatives by the reaction with thiourea (Hol´y et al., 2001).
Materials 2-Amino-6-chloropurine (S.8) Cesium carbonate Diisopropyl 2-chloroethoxymethylphosphonate (S.16; see Support Protocol 1) Dimethylformamide (DMF) Toluene Chloroform Celite Silica gel Ethyl acetate Hexane Diethyl ether 1 M HCl Conc. aqueous ammonia Acetone Bromotrimethylsilane
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Acetonitrile 0.25 M sulfuric acid 500-mL and 1-L round-bottom flasks Magnetic stirrer with heating plate and silicon oil bath Rotary evaporator with diaphragm pump Glass filter funnels with 1-L filter flasks 6 × 40–cm chromatography column Reflux condenser Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Synthesize 2-amino-6-chloro-9-[2-(diisopropylphosphonylmethoxy)ethyl]purine 1. Prepare a mixture of 17.0 g (0.1 mol) 2-amino-6-chloropurine, 16.5 g (0.05 mol) cesium carbonate, and 28 g (0.11 mol) diisopropyl 2-chloroethoxymethylphosphonate in 350 mL DMF in a 1-L round-bottom flask. Stir for 3 hr at 120◦ C with exclusion of moisture. 2. Evaporate the volatiles under vacuum using a rotary evaporator, and co-distill the residue three times with 50 mL toluene. 3. Extract with 200 mL boiling chloroform. Decant and filter the extract through Celite. Repeat this step two more times. 4. Evaporate under vacuum and apply the residue onto a dry 6 × 40–cm column of 300 mL silica gel. Elute with chloroform and monitor fractions by TLC using 9:1 (v/v) chloroform/methanol as eluent. To prepare a dry silica gel column, set up the empty glass column in a perpendicular position and join the outflow tube to a running water aspirator. Begin shedding the silica gel powder through the funnel with 4- to 5-mm outflow diameter located at the open top of the column, occasionally tapping lightly on the glass tube wall. The direction of powder stream should be oriented alongside the axis of the column. A properly made dry column not only spares solvent (separation takes place during soaking), but often gives better separation than columns poured from a silica gel suspension.
5. Collect fractions containing the main (faster-moving) product and evaporate under vacuum. 6. Dissolve in 150 mL ethyl acetate and then add hexane until the solution remains turbid (i.e, does not clear with stirring). Scratch the wall of the flask with a spatula and leave overnight at 0◦ C to crystallize. 7. Collect crystals by filtration, wash with 50 mL of 1:2 (v/v) diethyl ether/hexane, and dry under vacuum. The yield of S.9 should be ∼23 g (59%). It can be stored in a brown glass bottle at room temperature indefinitely without decomposition. Compound characterization: m.p. 93◦ C. C8 H13 N6 O4 P (288.2). 1 H NMR (DMSO-d6 ): δ 1.11, 1.16 (2 × d, 2 × 6H, 4CH3 ), 3.77 (d, 2H, J(P-CH) = 8.3 Hz, P-CH2 ), 3.87 (t, 2H, H2 ), 4.24 (t, H, J1 ,2 = 5.0 Hz, H1 ), 4.52 (dsept, 2H, JCH,CH2 = 6.2 Hz, J(P-OCH) = 7 Hz, P-O-CH), 6.89 (br s, 2H, NH2 ), 8.06 (s, 1H, H8).
Synthesize 9-[2-(diisopropylphosphonylmethoxy)ethyl]guanine 8. Boil a solution of 9.8 g (0.025 mol) S.9 in 240 mL of 1 M HCl under reflux for 1 hr. Synthesis of Acyclic Nucleoside Phosphonates
9. Add conc. aqueous ammonia to the the warm mixture to make it alkaline. 10. Cool it down to room temperature and concentrate under vacuum on a rotary evaporator to a thick paste.
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11. Redissolve by heating to a boil, adding water as required. Allow to crystallize overnight at room temperature. 12. Collect by filtration. Wash with 50 mL cold water, then 100 mL acetone, and then 100 mL diethyl ether. Dry under vacuum. The yield of S.10 should be ∼7 g (78%). It can be stored at room temperature indefinitely without decomposition. Compound characterization: m.p. 228◦ C. C13 H22 N5 O5 P (359.3). 1 H NMR (DMSO-d6 ): δ 1.16, 1.19 (2 × d, 2 × 6H, 4CH3 ), 3.77 (d, 2H, J(P-CH) =8.3 Hz, P-CH2 ), 3.81 (t, 2H, H2 ), 4.11 (t, 2H, H1 ), 4.51 (m, 2H, P-O-CH), 6.49 (br s, 2H, NH2 ), 7.64 (s, 1H, H8), 10.70 (br s, 1H, NH).
Synthesize 9-[2-(phosphonomethoxy)ethyl]guanine (PMEG) 13. Add 15 mL bromotrimethylsilane to a suspension of 5.4 g (15 mmol) S.10 in 120 mL acetonitrile in a 500-mL flask. Quickly close the flask with a stopper and dissolve the compound by shaking the flask by hand. 14. Allow to stand overnight at room temperature. Evaporate the volatiles under vacuum on a rotary evaporator. 15. Add 50 mL water. After 20 min, make the solution alkaline (∼pH 10 to 11) with conc. aqueous ammonia. 16. Evaporate under vacuum on a rotary evaporator. 17. Redissolve the residue in 200 mL boiling water and acidify the hot solution with 0.25 M sulfuric acid to pH 3.5. 18. Leave overnight at room temperature to crystallize. 19. Collect the product by filtration. Wash it with 100 mL acetone and then 100 mL diethyl ether. Dry under vacuum. The yield of S.11 should be ∼3 to 4 g (74% to 97%). It can be stored at room temperature indefinitely without decomposition. Compound characterization: m.p. > 280◦ C. C7 H10 N5 O5 P (275.2). 1 H NMR (DMSO-d6 ): δ 3.62 (d, 2H, J(P-CH) = 8.5 Hz, P-CH2 ), 3.93 (t, 2H, H2 ), 4.21 (t, 2H, J1 ,2 = 5.1 Hz, H1 ), 7.87(s, 1H, H8).
SYNTHESIS OF 2-AMINO-N6 -(CYCLOPROPYLAMINO)9-[2-(PHOSPHONOMETHOXY)ETHYL]PURINE
BASIC PROTOCOL 4
The title compound (S.13; Fig. 14.2.2) behaves in vivo as a prodrug of the guanine derivative PMEG. It is most likely converted by an adenylate deaminase–like enzyme. Due to the high cytotoxicity of PMEG, the cyclopropylamino analog exhibits anticancer activity (Hatse et al., 1999; Naesens et al., 1999; Valeri´anov´a et al., 2001, 2003). The synthetic procedure makes use of the reactive 6-chloro intermediate described in Basic Protocol 3 (Hol´y et al., 1999).
Materials Cyclopropylamine 2-Amino-6-chloro-9-[2-(diisopropylphosphonylmethoxy)ethyl]purine (S.9; see Basic Protocol 3) Dioxane Chloroform Methanol Ethanol Dowex 50 × 8, acid form (see recipe)
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1:10 (v/v) aqueous ammonia in 20% aqueous methanol Acetonitrile Bromotrimethylsilane Dowex 1 × 2, acetate form (see recipe) 0.2 and 1 M acetic acid Acetone Diethyl ether 250-mL and 1-L round-bottom flasks Reflux condenser Soda lime protecting tube Magnetic stirrer with heating plate and silicon oil bath 4 × 40–cm glass chromatography column Peristaltic pump with column adapter UV detector Fraction collector (optional) Rotary evaporator with diaphragm pump Vacuum desiccator with P2 O5 1-L cylinder-shaped bottles Plastic tubing Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Synthesize 2-amino-6-cyclopropylamino-9-[2-(diisopropylphosphonylmethoxy)ethyl]purine 1. Set up a 250-mL round-bottom flask with a reflux condenser, protected by a soda lime protecting tube. Add 10 mL cyclopropylamine to a suspension of 7.8 g (20 mmol) S.9 in 120 mL dioxane and stir under reflux (120◦ C bath temperature). 2. Follow the course of reaction by TLC in 4:1 (v/v) chloroform/methanol. The reaction time should be 3 to 5 hr. The mobility of the product is slower than the starting material.
3. After the reaction is finished, evaporate the solution under vacuum and co-distill four times with 50 mL ethanol. 4. Dissolve the residue in 20% (v/v) aqueous methanol and then add Dowex 50 × 8 (acid form) to acidify the solution to pH 2 to 3. 5. Pour the suspension into a 4 × 40–cm glass chromatography column with 150 mL of the same resin, equilibrated with 20% aqueous methanol. 6. Wash the column with 20% aqueous methanol at 2 to 3 mL/min, following the fractions by UV absorption. 7. When the eluate is no longer acidic and the UV absorption has dropped, elute the column with a 1:10 (v/v) ammonia in 20% aqueous methanol. 8. Collect the main UV-absorbing fractions of the ammonia eluate and evaporate under vacuum on a rotary evaporator. 9. Dry the residue first by co-distilling it three times with 50 mL ethanol and then by drying overnight over P2 O5 at 10 to 14 Pa. Synthesis of Acyclic Nucleoside Phosphonates
Synthesize 2-amino-6-cyclopropylamino-9-[2-(phosphonomethoxy)ethyl]purine 10. Add 100 mL acetonitrile and 20 mL bromotrimethylsilane, close the flask tightly, and dissolve by shaking. Set the mixture aside overnight at room temperature. 11. Evaporate the volatiles on rotary evaporator under vacuum.
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12. Add 50 mL water and make the solution alkaline (∼pH 9 to 10) with conc. aqueous ammonia. Evaporate to dryness under vacuum. 13. Repeat steps 4 to 7, replacing 20% methanol with water. 14. Evaporate the ammonia eluate to a volume of ∼20 mL and make it slightly alkaline (pH 9 to 10) with conc. aqueous ammonia. For successful purification on strong anion exchangers, it is crucial to apply a weakly alkaline solution to the column to ensure complete dissociation of all acids.
15. Apply the solution onto a glass column with 200 mL Dowex 1 × 2 (acetate form) equilibrated with 0.02 M acetic acid. Wash the column with 0.02 M acetic acid until the UV absorption of the eluate drops. 16. Prepare 1 L of 1 M acetic acid and degas it under vacuum for 10 to 20 min. 17. Set up a linear gradient system of 0.02 to 1.0 M acetic acid, using two 1-L cylindershaped bottles connected by plastic tubing. Set the bottle with the lower concentration on a magnetic stirrer, and ensure that the bottoms of the two bottles are level. Use a peristaltic pump to pump solution from the bottle with the lower concentration (where the two solutions are being mixed). 18. Begin pumping the solution through the column at 2 to 3 mL/min, following the eluate by UV absorption. If desired, collect fractions automatically. 19. Collect the main UV-absorbing peak, evaporate under vacuum, and co-distill three times with 150 mL water. Crystallize the residue from a minimum amount of boiling water overnight at 0◦ C. 20. Collect the crystalline product by filtration. Wash with 100 mL acetone and then 100 mL diethyl ether, and dry under vacuum. Compound characterization (S.13): m.p. 263◦ C. C11 H17 N6 O4 P (318.4). 1 H NMR (D2 O + NaOD): δ 0.67 (m, 2H, H2" + H3"), 0.89 (m, 2H), 2.82 (m, 1H, H1”), 3.54 (d, 2H, J(P,CH) =8.6, P-CH), 3.91 (t, 2H, J1 ,2 = 4.6, H2 ), 4.24 (t, 2H, H1 ), 7.86 (s, 1H, H8).
PREPARATION OF THE PME SYNTHON: DIISOPROPYL 2-CHLOROETHOXYMETHYLPHOSPHONATE
SUPPORT PROTOCOL 1
The title compound is an example of a synthon for the second synthetic approach to ANPs, a reagent which introduces the whole side chain at the heterocyclic base at once in a single reaction. The PME synthon is used to synthesize PMEA, PMEDAP, and PMEG and its derivatives. To prepare the PME synthon (S.16; Fig. 14.2.3A), 2-chloroethyl chloromethyl ether (S.15) is first obtained by the reaction of 2-chloroethanol with HCl and 1,3,5-trioxane or paraformaldehyde. The synthon is then formed by Arbuzoff reaction of triisopropyl phosphite with S.15 (Hol´y et al., 1999). The Arbuzoff reaction is very easy to perform. It depends very much on the temperature, and the activation scope can be sharp and comparatively high. The reactivity of the two C–Cl linkages of S.15 with triisopropyl phosphite is entirely different, with a strong preference for the chloromethyl residue.
Materials 1,3,5-Trioxane 2-Chloroethanol (S.14) Dry ice/ethanol bath Hydrogen chloride (gas) Dry nitrogen (gas) Calcium chloride Celite
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Figure 14.2.3
Scheme for synthesis of the PME and C1 synthons. Ts, p-tolylsulfonyl.
Dichloromethane Conc. sulfuric acid Triisopropyl phosphite 3-L reaction flask with inlet/outlet system Thermometer Plastic tubing Calcium chloride protecting tube Magnetic stirrer with heating plate Hydrogen chloride drying system (composed of washing bottle with conc. sulfuric acid, protective flasks before and after, inlet tube with sinter, with all parts interconnected in-line by polypropylene tubing) 2-L separatory funnel Rotary evaporator with diaphragm pump 2-L distillation flask Sintered glass filter funnels with 2-L filter flasks 40-cm distillation columns with head and thermometer 1-L two-neck round-bottom flask 250-mL dropping funnel Diaphragm vacuum pump Oil vacuum pump 250-mL and 1-L round-bottom receiving flasks CAUTION: This procedure makes use of gaseous hydrogen chloride and must be performed in a well-ventilated fume hood. The product itself or the side products of the reaction may be carcinogens. All precautions must be taken to avoid inhalation of the vapors or contact with skin or eyes.
Synthesis of Acyclic Nucleoside Phosphonates
Synthesize 2-chloroethyl chloromethyl ether 1. Build a reactor apparatus consisting of a 3-L reaction flask with inlet system, thermometer for measuring inside temperature, and outlet tube with plastic tubing closed with calcium chloride protecting tube and reaching up to the vent. 2. Load the reactor with 210 g of 1,3,5-trioxane and 564 g of 2-chloroethanol, and stir until the reagents are completely dissolved.
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3. Cool the reactor in a dry ice/ethanol bath until the temperature inside the reaction mixture drops to –11◦ C. 4. Introduce hydrogen chloride through the inlet of the HCl drying system at −11◦ to −9◦ C for 10 to 12 hr. 5. Transfer the reaction mixture to a 2-L separatory funnel and separate the lower product-containing fraction from the upper layer (water). 6. Pour the liquid mixture to a 2-L distillation flask on a magnetic stirrer and bubble dry nitrogen through the liquid for 1 hr at room temperature with efficient stirring. 7. Dry overnight with ∼100 g calcium chloride, filter, and wash twice with 100 mL dichloromethane. 8. Evaporate the solvent and other volatiles at a maximum of 40◦ C/5 bar. 9. Mount a 40-cm distillation column (with head and thermometer) onto the flask and redistill the crude product. Take the fraction that distills at 50◦ to 55◦ C at 21 mbar (or 21 kPa). The yield of S.15 should be ∼600 g. This product can be used directly for the reaction with triisopropyl phosphite without delay. It is possible to keep it in tightly closed bottle, preferably in a refrigerator, for up to 3 to 4 months, but there is a risk of partial decomposition. Compound characterization: liquid of sharp odor, b.p. 52◦ C/21 kPa. 1 H NMR (400 MHz, CDCl3 ): δ 3.69 (t, 2H, Jvic = 5.4 Hz, CH2 Cl), 3.95 (t, 2H, Jvic = 5.4 Hz, CH2 O), 5.53 (s, 2H, OCH2 Cl). 13 C NMR (100.6 MHz, CDCl3 ): 41.89 (CH2 Cl), 70.16 (CH2 O), 82.39 (OCH2 Cl).
Synthesize diisopropyl 2-chloroethoxymethylphosphonate 10. Build an apparatus on a magnetic stir plate consisting of a 1-L two-neck roundbottom flask with a 250-mL dropping funnel and a 40-cm glass distillation column with a distillation head and thermometer. 11. Place 136 g (1.056 mol) triisopropyl phosphite in the flask and heat it to 130◦ C. 12. Add dropwise, under stirring, 246 mL (1 mol) S.15 at a rate where the column does not become overloaded with the isopropyl chloride (b.p. 36◦ C) that forms during the reaction. The reaction of chloromethyl ether with triisopropyl phosphite needs a certain critical mass to start. Heat the phosphite to the above temperature, add ∼1 to 2 mL of the chloro derivative, and wait ∼30 sec to see whether the reaction (accompanied by formation of isopropyl chloride) has started. Repeat this procedure until the reaction starts (usually at 140◦ to 145◦ C), then add the remaining chloromethyl ether at a rate where isopropyl chloride condenses easily in the receiver. Should the reaction not begin after addition of 5 mL chloromethyl ether, raise the bath temperature by 10◦ C.
13. After the reaction has subsided, heat an additional 2 to 3 hr at 140◦ C and distill in a diaphragm pump vacuum, then in an oil pump vacuum. 14. Take the fraction with a b.p. at 120◦ to 122◦ C /13 Pa. The yield of PME synthon (S.16) should be between 210 and 240 g (85% to 93%). It is stable indefinitely at room temperature. Compound characterization: oil, b.p. 120◦ to 122◦ C / 13 Pa. 1 H NMR (400 MHz, CDCl3 ): δ 1.347, 1.351 (2 × d, 12H, Jvic = 6.2 Hz, CH3 -iPr), 3.64 (t, 2H, Jvic = 5.7 Hz, CH2 Cl), 3.81 (d, 2H, JH,P = 8.3 Hz, CH2 P), 3.86 (t, 2H, Jvic = 5.7 Hz, CH2 O), 4.77 (dh, 2H, JH,P = 7.7 Hz, Jvic = 6.2 Hz, CH-iPr). 13 C NMR (100.6 MHz, CDCl3 ): 23.95, 24.06 (d, JC,P = 4 Hz, CH3 -iPr), 42.35 (CH2 Cl), 66.03 (d, JC,P = 168 Hz, CH2 P), 71.21 (d, JC,P = 7 Hz, CH-iPr), 72.99 (d, JC,P = 11 Hz, CH2 O).
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BASIC PROTOCOL 5
SYNTHON METHOD FOR SYNTHESIS OF 9-(R)-[2-(PHOSPHONOMETHOXY)PROPYL]ADENINE [TENOFOVIR, (R)-PMPA] The title compound (Fig. 14.2.4, S.33) is the active component of the succesful antiAIDS drug Viread. It can be prepared by three synthetic methods: (1) by condensation of the (R)-PMP synthon (S.32) with adenine (the synthon approach; Hol´y et al., 1995), (2) by reaction of 9-(R)-(2-hydroxypropyl)adenine (S.25) with diisopropyl ptolylsulfonyloxymethylphosphonate (the two-step approach; Hol´y and Masoj´ıdkov´a, 1995), or (3) by reduction of the hydroxymethyl group in HPMP derivatives (see S.38 in Fig. 14.2.6). While the third alternative is not practical, the first two approaches are comparable in labor and time consumption, with a slight preference for the stepwise approach in large-scale syntheses. The synthon method has an optimum application in the preparation of PMP derivatives derived from various heterocyclic bases on not too large a scale, or in cases where the alkylation of the base is difficult, or where the base does not withstand the conditions of the stepwise procedure. The synthon and two-step methods proceed through a common precursor, 9-(R)-[2diisopropylphosphonyl)methoxypropyl]adenine (S.27). This protocol describes the onestep condensation of the PMP synthon (S.32) with the heterocyclic base in DMF in the presence of equivalent sodium hydride or cesium carbonate to give the protected diester of the PMP compound (S.27), which is then deprotected by treatment with bromotrimethylsilane to give PMPA (S.33). Preparation of the (R)-PMP synthon is achieved by various routes described in Support Protocols 2 to 6. The two-step method for achieving S.27 is presented in the Alternate Protocol.
Materials
Synthesis of Acyclic Nucleoside Phosphonates
Sodium hydride, 60% dispersion in paraffin oil Dimethylformamide (DMF) Adenine (R)-PMP synthon (S.32; see Support Protocol 3 or 6) Toluene Chloroform Celite Methanol Silica gel Acetonitrile Bromotrimethylsilane Conc. aqueous ammonia Dowex 50 × 8, acid form (see recipe) Dowex 1 × 2, acetate form (see recipe) 1 M acetic acid Ethanol Diethyl ether 250-mL round-bottom flasks Magnetic stirrer with heating plate and silicon oil bath Rotary evaporator with diaphragm pump Glass filter funnel with filter flask 4 × 40–cm and 3 × 40–cm chromatography columns Fraction collector (optional) Gradient maker (see Basic Protocol 4) Calcium chloride protecting tube Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E)
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Figure 14.2.4 Scheme of (R)-PMPA preparation by the (R)-PMP synthon and stepwise procedures. Bn, benzyl; Pd/C, palladium-on-charcoal catalyst; Py, pyridine; Ts, p-tolylsulfonyl; X = Br, OTs.
Perform condensation 1. While stirring, add 0.8 g of 60% sodium hydride in paraffin oil (0.48 g, 20 mmol NaH) to 80 mL DMF in a 250-mL flask, followed by 2.7 g (20 mmol) adenine. Stir for 30 min at 80◦ C under a calcium chloride protecting tube. For precautions, see Basic Protocol 1, step 1.
2. Add in one portion 8.8 g (21.6 mmol) (R)-PMP synthon (S.32) in 40 mL DMF. Stir 16 hr at 100◦ C under exclusion of moisture. 3. Evaporate under vacuum on a rotary evaporator and then co-distill twice with 50 mL toluene. 4. Extract with 200 mL boiling chloroform. Decant and filter the extract through Celite. Repeat extraction and filtration until there is no more product (S.27) left in the material, as determined by TLC in 9:1 (v/v) chloroform/methanol (Rf S.27 = 0.60). 5. Combine the extracts, evaporate under vacuum, and purify on a 4 × 40–cm chromatography column containing 250 mL silica gel in chloroform. Elute with chloroform and monitor fractions by TLC. 6. Dry the amorphous product (5.0 g S.27) in a 250-mL round-bottom flask. The work-up can be interrupted after step 6 or 9.
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Compound characterization (S.27): C16 H26 N5 O4 P (371.4). 1 H NMR (DMSO-d6 ): δ 7.79 and 7.49 (d, 2H, arom. H), 4.56 (m, 2H, P-OCH), 4.05 (dd, 1H, J = 3.2, 10.6 Hz, O-CH2 ), 3.91 (dd, 1H, J = 5.7, 10.6 Hz, O-CH2 ), 3.75 (m, 1H, O-CH), 3.72 (dd, 1H, JP,CH = 9.2 Hz, Jgem = 13.8 Hz, P-CH2 ), 3.64 (dd, 1H, JP,CH = 8.9 Hz, Jgem = 13.8 Hz, P-CH2 ), 2.42 (s, 3H, CH3 arom.), 1.22 (d, 6H) and 1.205 (d, 3H) and 1.20 (d, 3H, JCH3,CH = 6.1 Hz, CH3 ), 1.04 (d, 3H, JCH3,CH = 6.4 Hz, CH3 ). 13 C NMR (DMSO-d6 ): 145.12, 132.42, 130.35 (2C) and 127.82 (2C, C arom.), 74.52 (d, JP,C = 12.2, O-CH), 72.28 (O-CH2 ), 70.37 (d), 70.34 (d, JP,C = 6.4, P-OCH), 62.89 (d, JP,C = 165.5, P-CH2 ), 23.99 (d, 2C, JP,C = 3.9) and 23.84 (d, 2C, JP,C = 4.4, CH3 ), 21.27 and 15.71 (CH3 ).
Perform deprotection 7. Add to the foamy residue 80 mL acetonitrile and 15 mL bromotrimethylsilane. Quickly close the flask and dissolve the compound by shaking. Allow to stand 15 to 18 hr in a closed flask at room temperature. 8. Evaporate under vacuum and then dissolve the residue in 100 mL water. Wait 20 min. Add conc. aqueous ammonia to make the reaction alkaline (∼pH 10 to 11) and evaporate again. 9. Deionize the residue on a 3 × 40–cm chromatography column containing 200 mL Dowex 50 × 8 (acid form). Wash with water until UV absorption of the eluate drops, then elute with 1 L of dilute (1:10) ammonia, and evaporate. 10. Dissolve the residue in 30 mL water. Make the solution alkaline (∼pH 10 to 10.5) by adding conc. aqueous ammonia and apply to a 3 × 40–cm chromatography column containing 150 mL Dowex 1 × 2 (acetate form). Elute with water until the UV absorption drops. 11. Continue eluting with a linear gradient of acetic acid as described (see Basic Protocol 4, steps 16 to 18), but using 1 L water and 1 L of 1 M acetic acid. 12. Collect the main UV-absorbing band and evaporate under vacuum. Co-distill the residue twice with 20 mL water, and dissolve in 20 mL boiling water. 13. Add an equal volume of ethanol and leave the solution standing overnight in the refrigerator (4◦ C) to crystallize. 14. Collect the product (S.33) by filtration, wash it with 50 mL ethanol and then 100 mL diethyl ether, and dry under vacuum. The yield of S.33 should be ∼3.5 g (60% relative to S.27). It can be stored indefinitely at room temperature. Additional pure product is contained in the filtrate and can be obtained by evaporation in vacuo and recrystallization under similar conditions. Compound characterization: m.p. > 260◦ C. C9 H14 N5 O4 P (287.2). 1 H NMR (500 MHz, D2 O+NaOD, ref(dioxane) = 3.75 ppm): δ 1.08 (d, 3H, JCH3 ,2 = 6.4 Hz, CH3 ), 3.42, 3.52 (2 × d, 2H, Jgem = 12.4 Hz, JH,P = 9.2 Hz, H4 ), 3.97 (m, 1H, H2 ), 4.25 (dd, 1H, Jgem = 14.7 Hz, J1 b,2 = 5.5 Hz, H1 b), 4.36 (dd, 1H, Jgem = 14.7 Hz, J1 a,2 = 4.0 Hz, H1 a), 8.17 (s, 1H, H2), 8.28 (s, 1H, H8). 13 C NMR (125.8 MHz, D2 O+NaOD, ref(dioxane) = 69.3 ppm): 18.94 (CH3 ), 50.12 (CH2 -1 ), 69.53 (d, JC,P = 151 Hz, CH2 -4 ), 78.04 (d, JC,P = 11 Hz, CH-2 ), 120.70 (C5), 146.35 (CH-8), 151.91 (C4), 154.98 (CH-2), 158.16 (C6). SUPPORT PROTOCOL 2
Synthesis of Acyclic Nucleoside Phosphonates
PREPARATION OF (R)-2-O-TETRAHYDROPYRANYLPROPANE-1,2-DIOL This compound (S.22; Fig. 14.2.4) can serve as a common intermediate for both the synthon and two-step approaches leading to PMP compounds. In the synthon approach, benzylation and acid hydrolysis are followed by linking of the phosphonomethylether function. The resulting intermediate (S.29) gives by hydrogenolysis the phosphonatesubstituted propanol derivative (S.31), which can be either applied directly for Mitsunobu alkylation or (even better) for tosylation, which affords the common synthon
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(S.32) applicable to one-step condensation leading to PMP derivatives. In the two-step alternative, the hydroxyl function of S.22 is transformed into a p-tolylsulfonyl derivative (S.23), which is then used to alkylate the base and gives, after acid deprotection, an N-(2-hydroxypropyl) derivative of the base (S.25) as the key intermediate for attachment of the phosphonomethyl ether group. This route is not presented in this unit, although another route to S.25 is given in the Alternate Protocol. The best available starting materials for synthesis of S.22 leading to optically pure enantiomers are undoubtedly lactic acid esters. The (R)- and (S)-enantiomers are commercially available as isobutyl and ethyl ester, respectively. Reaction with dihydropyran catalyzed with anhydrous HCl proceeds easily. It can be performed without solvent; though very exothermic, the reaction is quite safe even on a scale of several hundred grams. This compound could be easily transformed to the final product by reduction with LiAlH4 ; however, for safety reasons, the present method uses commercial Red-Al as a reducing agent.
Materials Isobutyl (R)-lactate (S.20) Dihydropyran 6 M HCl in DMF (see recipe) Silver oxide Celite Triethylamine Red-Al (Aldrich, Fluka) Diethyl ether Ethyl acetate Dioxane Magnesium sulfate, anhydrous 500-mL round-bottom flask Calcium chloride protecting tube Glass filter funnels (≥12 cm diameter) with 2-L filter flasks Distillation apparatus with 250-mL flask 500-mL three-neck flask Mechanical KPG stirrer Reflux condenser with calcium chloride protecting tube 200-mL dropping funnel Thermometer 600-mL glass beaker Broad glass funnel 1. In a 500-mL round-bottom flask, mix 100 mL (0.7 mol) isobutyl (R)-lactate with 70 mL dihydropyran. Add 1 mL of 6 M HCl in DMF and leave the mixture standing overnight at ambient temperature under a calcium chloride protecting tube. The strongly exothermic reaction that occurs is safe and does not require any external cooling.
2. Add 5 g silver oxide and stir magnetically for 7 hr. 3. Filter over Celite, add 2 mL triethylamine, and distill under vacuum. Take the fraction with a b.p. of 140◦ C/20 mbar. This protocol can be interrupted at steps 3 and 4. Biologically Active Nucleosides
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4. Set up a reaction apparatus consisting of a 500-mL three-neck flask with a mechanical KPG stirrer, a reflux condenser closed with calcium chloride protecting tube, a 200-mL dropping funnel, and a thermometer that reaches into the reaction mixture. 5. Quickly weigh ∼300 g (±10 g) of Red-Al into a 600-mL glass beaker and transfer it to the reactor using a broad glass funnel, followed (with caution) by 700 mL dry diethyl ether. CAUTION: Red-Al is a very reactive compound. Beware of contamination. Wash the beaker quickly with ethanol and then with water. Wear protective gloves.
6. Dilute the ester (step 3, 154 g) with 200 mL diethyl ether, and drop the solution under stirring and without cooling into the Red-Al solution at such a rate that the mixture boils moderately. Stir for an additional 2 hr at ambient temperature. 7. Cool the mixture in an ice-water bath and slowly drop in 120 mL ethyl acetate at a rate where reflux stays under control. 8. Slowly drop in 150 mL water and stir an additional 30 min. 9. Filter through a Celite pad, wash with 200 mL diethyl ether and then 200 mL dioxane, and evaporate the filtrate under vacuum. 10. Dissolve the residue in 1 L diethyl ether and dry overnight over magnesium sulfate. 11. Filter, evaporate the filtrate under vacuum, and transfer the residue to a 250-mL distillation flask. 12. Distill under vacuum, taking the fraction that boils at 87◦ to 94◦ C/2 kPa. The yield of S.22 should be ∼98 to 100 g (87% to 89% relative to the starting (R)-lactate). The product can be kept in a tightly closed bottle in the refrigerator for up to several months, although there is the risk of partial decomposition. Compound characterization: oil, b.p. 90◦ -94◦ C. 1 H NMR (500 MHz, DMSO-d6 ): δ 1.03, 1.07 (d, 3H, Jvic = 6.3 Hz, CH3 ), 1.37-1.50, 1.56-1.88 (2 × m, 6H, CH2 -THP), 3.24, 3.26 (dt, 1H, Jgem = 11.0 Hz, Jvic = 5.5 Hz, bCH2 OH), 3.31-3.50 (m, 2H, aCH2 OH + bCH2 OTHP), 3.63-3.71 (m, 1H, CH), 3.75-3.86 (m,1H, aCH2 O-THP), 4.52, 4.56 (t, 1H, Jvic = 5.5 Hz, OH), 4.66, 4.71 (dd, 1H, Jvic = 4.7, 2.8 Hz, CHO-THP). 13 C NMR (125.8 MHz, DMSO-d6 ): 16.61, 18.58 (CH3 ), 19.46, 19.57, 25.28, 25.32, 30.77, 30.93 (CH2 -THP), 61.64, 61.75 (CH2 O-THP), 65.04, 65.51 (CH2 -OH), 73.56, 74.08 (CH), 95.89, 98.04 (CHO-THP). Essentially identical conditions are used for synthesis of (S)-2-O-tetrahydropyranylpropane-1,2-diol from ethyl (S)-lactate. In that case, 120 g (1.05 mol) ethyl (S)-lactate and 100 mL dihydropyran give the protected ester (b.p. 90◦ –94◦ C/13 Pa) at a yield of 130 g (87%). Further steps were performed in an essentially identical manner with proportionally increased volumes, except for step 6, wherein the ester was added to the Red-Al solution undiluted. (S)-2-O-Tetrahydropyranylpropan-1,2-diol was obtained as a fraction with b.p. 86◦ –90◦ C/13 Pa. SUPPORT PROTOCOL 3
Synthesis of Acyclic Nucleoside Phosphonates
PREPARATION OF THE (R)-PMP SYNTHON: (R)-2-O(DIISOPROPYLPHOSPHONYLMETHOXY)PROPYL p-TOLYLSULFONATE In order to introduce the protected phosphonomethyl group at the secondary group in propane-1,2-diol, it is necessary to use suitable protection. 2-OTetrahydropyranylpropane-1,2-diol (S.22) can be benzylated and subsequently treated with acid (ion-exchange resin in acid form) to afford 1-O-benzylpropane-1,2diol (S.28). From this point there are two alternatives for phosphonomethylation: (1) chloromethylation followed by Arbuzoff reaction with triisopropyl phosphite (S.28-S.30-S.29), or (2) direct condensation (S.28-S.29) using either diisopropyl p-tolylsulfonyloxymethylphosphonate or diisopropyl bromomethylphosphonate. This
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protocol presents the chloromethylation/Arbuzoff route (via S.30), and direct etherification is presented in Support Protocol 4. The resulting O-benzyl-protected compound is then hydrogenated over palladium catalyst and subsequently tosylated to afford the title compound (S.32).
Materials Sodium hydride, 60% dispersion in paraffin oil Hexane Dimethylformamide (DMF), reagent grade (R)-2-O-Tetrahydropyranyl-1,2-propanediol (S.22; see Support Protocol 2) Benzyl bromide Saturated ammonia in methanol Ethyl acetate 80% (v/v) aqueous methanol Dowex 50 × 8, acid form (see recipe) Methanol Diethyl ether Magnesium sulfate, anhydrous 1,2-Dichloroethane 1,3,5-Trioxane Calcium chloride, anhydrous Toluene Triisopropyl phosphite Aluminum oxide, neutral Conc. aqueous HCl Argon Palladium-on-charcoal catalyst Celite Hydrogen Triethylamine Pyridine 4-Dimethylaminopyridine p-Tolylsulfonyl chloride Chloroform Glass receiver with glass filter (see Fig. 14.2.5) Diaphragm vacuum pump 2-L round-bottom reaction flasks Mechanical KPG stirrer 200-mL dropping funnel Magnetic stirrer Rotary evaporator with diaphragm pump Reflux condensers Glass filter funnels with 2-L filter flasks 250-mL distillation flask in distillation apparatus 500-mL three-neck round-bottom flask with inlet and outlet tubes HCl drying system with dry HCl from a cylinder and an inlet tube 8 × 40–cm chromatography columns Vacuum desiccator with P2 O5 Extraction funnel 2-L Erlenmeyer flask 1-L two-neck round-bottom flask Calcium chloride protecting tube at the end of polypropylene tubing reaching to the vent
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Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Synthesize (R)-1-O-benzylpropane-1,2-diol 1. Weigh 33 g of 60% sodium hydride in paraffin oil (19.8 g, 0.825 mol NaH) in a glass receiver with a glass filter (Fig. 14.2.5). Wash out the oil with 300 mL hexane, close with a glass stopper, and dry under vacuum for 10 min with exclusion of moisture using a diaphragm vacuum pump. For precautions, see Basic Protocol 1, step 1.
2. Transfer the dried material to a 2-L round-bottom reaction flask and add 800 mL DMF.
Synthesis of Acyclic Nucleoside Phosphonates
Figure 14.2.5 pounds.
Filtering device for filtration and drying of air- and/or moisture-sensitive com-
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3. Place the flask in an ice bath and begin stirring with a mechanical KPG stirrer to cool the solution. Using a 200-mL dropping funnel, add dropwise over 1 hr 130 g (0.812 mol) of (R)-2-O-tetrahydropyranylpropane-1,2-diol (S.22) and then stir for 30 min. 4. Slowly add 103 mL (0.866 mol) benzyl bromide. Stir the mixture for 2 hr with the KPG stirrer, and then remove the KPG stirrer and stir overnight at ambient temperature on a magnetic stirrer. 5. Add 20 mL methanolic ammonia and leave 1 hr at room temperature to decompose any residual benzyl bromide. Evaporate the volatiles under vacuum. The protocol can be interrupted at steps 5, 6, and 7.
6. Add 1 L ethyl acetate, transfer the solution to an extraction funnel, wash three times with 200 mL water, and evaporate the organic layer under vacuum. 7. Transfer the residue in 1 L of 80% aqueous methanol to a 2-L round-bottom flask and add ∼50 mL Dowex 50 × 8 (acid form). 8. Stir under reflux for 2 hr. Filter and then wash the resin with 200 mL methanol. Evaporate the organic solvents under vacuum. 9. Extract the remaining emulsion three times with 150 mL diethyl ether. Discard the aqueous phase, dry the extract with magnesium sulfate overnight, and evaporate under vacuum. 10. Transfer the oil to a 250-mL distilling flask and distill under vacuum. Take the fraction that boils at 92◦ to 96◦ C/100 Pa. The yield of S.28 should be ∼100 to 110 g (70% to 77% relative to S.22). The product can be kept in a tightly closed bottle in the refrigerator for several months without decomposition. Essentially identical conditions are used for the synthesis of (S)-1-O-benzylpropane-1,2diol (yield 81%, b.p. 86◦ -94◦ C/13 Pa).
Synthesize (R)-PMP synthon via chloromethylation 11. Place 200 mL 1,2-dichloroethane in a 500-mL three-neck round-bottom flask with inlet and outlet tubes. Add 80 g S.28, 25 g 1,3,5-trioxane, and 25 g calcium chloride. 12. While stirring and cooling with ice, saturate the solution with dry HCl gas through an inlet tube that reaches deep into the reaction flask. Leave the mixture to stand overnight at room temperature under a calcium chloride protecting tube. 13. Evaporate the solvent under vacuum and co-distill the residue three times with 50 mL toluene. 14. Add 104 g (0.5 mmol) triisopropyl phosphite and gradually heat the mixture with stirring under a reflux condenser. A vigorous reaction begins at 110◦ C. After it subsides, heat the mixture at 120◦ -140◦ C for an additional 2 hr and then allow it to cool. The protocol can be interrupted at steps 14, 15, and 16.
15. Apply the concentrated toluene solution of the crude reaction product (S.29) to an 8 × 40–cm chromatography column with 800 mL aluminum oxide, and elute the column with toluene, taking 250-mL fractions. 16. Analyze the fractions by TLC in 9:1 (v/v) toluene/ethyl acetate, with detection under UV light (Rf = 0.70 S.29; 0.60 S.28).
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17. Combine the product-containing fractions and evaporate them under vacuum. Dry the product overnight over phosphorus pentoxide at 13 Pa at room temperature. The yield of S.29 is ∼70% to 80% from S.28 when prepared via chloromethylation. The product should be used to prepare the PMP synthon as soon as possible. In the meantime, keep it in a deep freezer (–10◦ C).
18. Dissolve the residue (52.5 g, 0.15 mol) in 500 mL methanol in a 2-L round bottom flask, and add 1 mL conc. aqueous HCl. 19. Wash the flask with argon, add 2 g palladium-on-charcoal catalyst, and stir overnight under a hydrogen atmosphere at slight overpressure (100 kPa). 20. Filter through a Celite pad, wash with 200 mL methanol, and add triethylamine to make the mixture alkaline (∼pH 9 to 10). 21. Evaporate to dryness and co-distill three times with 50 mL pyridine. 22. Dissolve the residue in 300 mL pyridine, add 1.5 g 4-dimethylaminopyridine, and cool with ice. 23. While stirring the solution and cooling with ice, add 38 g (0.2 mol) p-tolylsulfonyl chloride in one portion. Stir 1 hr on ice and then continue stirring overnight at ambient temperature. 24. Cool the mixture on ice, add 20 mL methanol, and wait 30 min. 25. Add 20 ml water and wait another 1 hr. 26. Add 1 L ethyl acetate and extract the solution three times with 300 mL water. 27. Evaporate the organic phase under vacuum and co-distill five times with 50 mL toluene at a maximum 30◦ C bath temperature. 28. Apply the residue in a small volume of chloroform to an 8 × 40–cm chromatography column of 800 mL dry silica gel (see Basic Protocol 3, step 4) and elute it with chloroform. Monitor fractions by TLC in chloroform, with detection under UV light. 29. Combine the product-containing fractions and evaporate the solvent under vacuum. Dry the residue overnight over phosphorus pentoxide at 13 Pa at room temperature. The yield of S.32 should be ∼60 g (0.11 mol, 73% relative to S.29). The product should be used to prepare PMPA as soon as possible. In the meantime, keep it in a deep freezer (–10◦ C). Compound characterization (S.32): C17 H20 O4 PS (320.4). 1 H NMR (DMSO-d6 ): δ 8.15 and 8.07 (s, 2H, H2 + H8), 7.29 (brs, 2H, NH2 ), 4.49 (m, 2H, P-OCH), 4.26 (dd, 1H, J1 a,2 = 3.7 Hz, Jgem = 14.4 Hz, H1 a), 4.17 (dd, 1H, J1 b,2 = 6.6 Hz, Jgem = 14.4 Hz, H1 b), 3.96 (m, 1H, H2 ), 3.79 (dd, 1H, JP,CH = 9.2 Hz, Jgem = 13.7 Hz, P-CH2 ), 3.72 (dd, 1H, JP,CH = 9.5 Hz, Jgem = 13.7 Hz, P-CH2 ), 1.19 (d, 3H), 1.16 (d, 3H), 1.15 (d, 3H), 1.12 (d, 3H), 1.08 (d, 3H, JCH3,CH = 6.1 Hz, CH3 ). Essentially identical conditions are used for the synthesis of (S)-2-O-(diisopropylphosphonylmethoxy)propyl p-tolylsulfonate from (S)-1-O-benzylpropane-1,2-diol. SUPPORT PROTOCOL 4 Synthesis of Acyclic Nucleoside Phosphonates
PREPARATION OF 1-O-BENZYL(R)-2-O-(DIISOPROPYLPHOSPHONYLMETHYL)PROPANE-1,2-DIOL VIA DIRECT ETHERIFICATION This method makes use of diisopropyl p-tolylsulfonyloxymethylphosphonate (the C1 synthon; Fig. 14.2.3), which reacts with the sodium alkoxide formed from (R)-1-Obenzylpropane-1,2-diol in situ by the action of sodium hydride.
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Materials Sodium hydride, 60% dispersion in paraffin oil Hexane Tetrahydrofuran, freshly distilled from NaH (R)-1-O-Benzylpropane-1,2-diol (S.28; see Support Protocol 3) Diisopropyl p-tolylsulfonyloxymethylphosphonate (S.19; see Support Protocol 5) Celite Ethyl acetate Toluene Silica gel Glass receiver with glass filter (Fig. 14.2.5) Rotary evaporator with diaphragm pump 4-L three-neck round-bottom flask Magnetic stirrer 250-mL dropping funnel Calcium chloride protecting tube Thermometer Glass filter funnel with filter flask 8 × 40–cm glass chromatography column Vacuum desiccator with P2 O5 Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) 1. Weigh 20 g of 60% sodium hydride in paraffin oil (12 g, 0.5 mol NaH) in a glass receiver with glass filter (Fig. 14.2.5), wash it with 300 mL hexane, and close quickly and tightly. Dry under vacuum for 10 min using a rotary evaporator with a diaphragm pump. For precautions, see Basic Protocol 1, step 1.
2. Set up a 4-L three-neck round-bottom reaction flask with a magnetic stirrer, 250-mL dropping funnel, calcium chloride protecting tube, and thermometer. Cautiously transfer the washed sodium hydride (0.5 mol) to 500 mL tetrahydrofuran freshly distilled from NaH. 3. While stirring, use a 250-mL dropping funnel to add dropwise a solution of 75 g (0.45 mol) S.28 in 200 mL tetrahydrofuran. Continue stirring 1 hr. 4. Cool with ice and add dropwise, under stirring, a solution of 175 g (0.5 mol) diisopropyl p-tolylsulfonyloxymethylphosphonate (S.19) in 200 mL tetrahydrofuran. Stir for 3 to 4 hr, and then set the reaction aside for 2 days. CAUTION: Approximately 30 min after warming to room temperature, the foaming of the mixture becomes hardly controllable. This is the reason a large flask is used for the reaction. Be prepared to reduce the foam with ether vapors from a bottle of ether, or with a few drops of 5% dodecanol solution in ether.
5. Filter through a Celite pad, wash with 100 mL tetrahydrofuran, and evaporate under vacuum. CAUTION: The material on the filter may contain residual sodium hydride. Therefore, wash the Celite pad with ethanol before using water to wash it. The reaction can be interrupted at this point.
6. Dissolve the residue in 100 to 150 mL toluene and apply the solution onto an 8 × 40–cm column of dry silica gel (see Basic Protocol 3, step 4). Elute with toluene and monitor fractions by TLC in 95:5 (v/v) toluene/ethyl acetate. The first
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fraction contains 1-O-benzylpropane-1,2-diol 2-p-tolylsulfonate (total, 0.12 mol, 25%) followed by the product (Rf = 0.60 to 0.65). 7. Collect the fractions containing product, evaporate under vacuum, and dry under vacuum over P2 O5 . The yield of S.29 should be ∼97 to 100 g (0.29 mol, 58% relative to S.28). The product should be used as soon as possible to prepare the (R)-PMP synthon (see Support Protocol 6). In the meantime, keep it in a deep freezer (–10◦ C) to avoid decomposition. SUPPORT PROTOCOL 5
PREPARATION OF THE C1 SYNTHON: DIISOPROPYL p-TOLYLSULFONYLOXYMETHYLPHOSPHONATE This diester is one of the key compounds of ANP chemistry, as it is used to introduce the phosphonomethyl ether group onto an isolated hydroxyl function. An isopropyl ester group protects the phosphonate group without danger of alkylation of (mainly) acid NH functions, which otherwise occurs with ethyl and in particular methyl esters at elevated temperatures. The p-tolylsulfonyloxy group is an excellent leaving anion. In addition to suitable reactivity, the main advantages of this reagent are its low cost, simple preparation, and crystalline form and stability, which make it easy to prepare and handle. The preparation (Fig. 14.2.3B) consists of the reaction of diisopropyl phosphite (S.17) with paraformaldehyde in the presence of triethylamine (Abramoff-Pudovik reaction). The hydroxymethylphosphonate intermediate (S.18) is then transformed to the target compound (S.19) by reaction with p-tolylsulfonyl chloride and a tertiary base.
Materials
Synthesis of Acyclic Nucleoside Phosphonates
Triethylamine Diisopropyl phosphite Paraformadehyde, dried at atmospheric pressure over sulfuric acid in vacuum desiccator for ≥48 hr Ethyl acetate Ethanol p-Nitrobenzylpyridine reagent (see recipe) Acetonitrile p-Tolylsulfonyl chloride 4-Dimethylaminopyridine (optional) Diethyl ether Magnesium sulfate, anhydrous Toluene Silica gel (40-60 µm) Chloroform Methanol Cyclohexane Hexane 500 ml round-bottomed flask Reflux condenser with calcium chloride protecting tube Magnetic stirrer with heating plate and silicon oil bath 1-L Erlenmeyer flask Glass filter funnel with filter flask Rotary evaporator with diaphragm pump 8 × 20– to 8 × 22–cm glass chromatography column Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) NOTE: All solvents and chemicals should be reagent grade.
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Synthesize diisopropyl hydroxymethylphosphonate 1. Add 8.5 mL triethylamine to a mixture of 100 g (0.6 mol) diisopropyl phosphite and 23 g dried paraformaldehyde in a 500-mL round-bottom flask with a stir bar. 2. Connect a reflux condenser with a calcium chloride protecting tube and stir at 100◦ C (bath temperature) until the paraformaldehyde dissolves, and then continue stirring for 1 hr. 3. Follow the course of the reaction by TLC in 9:1 (v/v) ethyl acetate/ethanol with detection by p-nitrobenzylpyridine reagent. This crude material is used directly for tosylation.
Synthesize diisopropyl p-tolylsulfonyloxymethylphosphonate 4. Dissolve the reaction mixture in 500 mL acetonitrile in a 1-L flask. Add 126 g (0.66 mol) p-tolylsulfonyl chloride, 0.8 g 4-dimethylaminopyridine, and 94 mL (0.67 mol) triethylamine. Control the mildly exothermic reaction by cooling under cold tap water. Set the reaction aside overnight without cooling. 5. Cool the mixture with ice and add 50 mL water dropwise over 1 hr, with stirring, to decompose excess sulfonyl chloride. Let sit 6 hr at ambient temperature. 6. Add 250 mL water and evaporate acetonitrile under vacuum. 7. Extract the aqueous solution three times with 150 mL diethyl ether, dry the extract in a 1-L Erlenmeyer flask with anhydrous magnesium sulfate overnight. The workup can be interrupted (over the weekend) at this point or after step 9.
8. Filter the dried solution and evaporate it on a rotary evaporator. When the pump reaches its maximum, heat the residue for 30 min at 50◦ C. 9. Dissolve the oil in 100 mL toluene and filter it through an 8 × 20– to 8 × 22–cm glass chromatography column containing 500 mL dry silica gel (see Basic Protocol 3, step 4). Wash out impurities with chloroform and elute the product with ethyl acetate. Monitor fractions by TLC using 9:1 (v/v) chloroform/methanol. 10. Evaporate the eluate under vacuum and remove the last traces of solvent as described in step 8. 11. Add 25 to 50 mL cyclohexane to the residual oil and stir it on a magnetic stirrer (the oil crystallizes). 12. Filter the product. If necessary, dilute the thick suspension with hexane before filtration. 13. Wash with hexane and dry in vacuo. The yield of S.19 should be between 140 and 160 g (72% to 73%). It can be stored in dry form in closed bottles at 0◦ C.
PREPARATION OF THE (R)-PMP SYNTHON To reach the target following direct etherification (see Support Protocol 4), the 1-Obenzyl group in compound S.29 must be removed and the functional hydroxyl group of the resulting intermediate S.31 converted to p-tolylsulfonate. The last step must be performed under mild conditions, due to the presence of the adjacent phosphonateprotecting isopropyl ester group, which could be trans-esterified by the primary hydroxyl group.
SUPPORT PROTOCOL 6
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Materials (R)-1-O-Benzyl-2-O-(diisopropylphosphonylmethyl)propane-1,2-diol (S.29; see Support Protocol 4) Methanol, spectroscopically pure or HPLC pure Argon 10% palladium-on-charcoal catalyst Celite Toluene Pyridine 4-Dimethylaminopyridine p-Tolylsulfonyl chloride Ethyl acetate Diluted HCl: 1:20 (v/v) in water Saturated potassium hydrogencarbonate Magnesium sulfate, anhydrous Silica gel 2-L round-bottom flask Rotary evaporator with diaphragm pump Bladder as hydrogen reservoir Glass filter funnel with filter flask 6 × 40–cm glass chromatography column Remove benzyl group 1. Dissolve 69 g (0.2 mol) S.29 in 1 L methanol (spectroscopically pure or HPLC pure) in a 2-L round-bottom flask, and wash it repeatedly with argon. 2. Add 2 g palladium catalyst. 3. Evacuate the flask and fill it with hydrogen from a bladder. Stir overnight at room temperature, connected to the bladder at slight overpressure (100 kPa). 4. Filter through a Celite pad, wash it with 100 mL methanol, and evaporate under vacuum. CAUTION: For safety, always avoid the presence of air in the system when manipulating solvents and a hydrogenation catalyst. The reaction mixture could burst and/or go into flames. Always remove air from the system by washing it optimally with argon, eventually with nitrogen. When neither is available, at least evacuate the system long enough to fill it with the solvent vapors! CAUTION: Catalysts filtered from hydrogenation reactions are activated and dangerous. As soon as the catalysts are dry, they can self-ignite and burn in air. This applies in particular to catalysts on charcoal, which can reach high temperatures and break glass filtering funnels. Never let air pass through the pad when filtering by suction. When the filtration is complete, always pour water on the catalyst layer!
5. Evaporate the residue twice with 50 mL toluene and twice with 50 mL pyridine, and dissolve the residue in 300 mL pyridine. The protocol can be interrupted here or after step 9.
Synthesis of Acyclic Nucleoside Phosphonates
Characterization data (S.31): 1 H NMR (500 MHz, DMSO-d6 ): δ 1.05 (d, 3H, JCH3,2 = 6.3 Hz, 2CH3 ), 1.24, 1.25 (2 × d, 2 × 6H, Jvic = 6.2 Hz, CH3 -iPr), 3.30, 3.40 (2 × dt, Jgem = 11.0 Hz, J1,OH = 5.5 Hz, J1,2 = 5.5 Hz, H1), 3.50 (qt, 1H, J2,CH3 = 6.3 Hz, J2,1 = 5.5 Hz, H2), 3.76, 3.78 (2 × dd, Jgem = 13.8 Hz, JH,P = 8.7 Hz, H4), 4.59 (dh, 2H, JH,P = 7.8 Hz, Jvic = 6.2 Hz, CH-iPr), 4.61 (t, 1H, JOH,1 = 5.5 Hz, OH). 13 C NMR (125.8 MHz, DMSO-d6 ): 16.73 (CH3 -2), 23.89, 24.03 (d, JC,P = 4 Hz, CH3 -iPr), 63.18 (d, JC,P = 166 Hz, CH2 -4), 64.59 (CH2 -1), 70.21 (d, JC,P = 6 Hz, CH-iPr), 78.34 (d, JC,P = 12 Hz, CH-2).
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Perform tosylation 6. Cool the solution with ice and, while cooling and stirring, add 1 g of 4dimethylaminopyridine followed by 45 g (0.24 mol) p-tolylsulfonyl chloride. 7. Stir 2 hr at 0◦ C and let the reaction mixture warm up overnight to ambient temperature. 8. Add 20 mL methanol and stir for 2 hr. 9. Concentrate under vacuum to half of the original volume. 10. Add 600 mL ethyl acetate. Wash three times with 150 mL water, then wash repeatedly with 150-mL portions of diluted (1:20) HCl to acidify the reaction. Wash with 100 mL saturated potassium hydrogencarbonate, and finally with 100 mL water. 11. Dry with magnesium sulfate, filter, and evaporate under vacuum. 12. Purify by on a 6 × 40–cm glass chromatography column containing 500 mL dry silica gel (see Basic Protocol 3, step 4), eluting with toluene. Monitor fractions by TLC using 9:1 (v/v) toluene/ethyl acetate. 13. Dry the resulting colorless thick oil under vacuum at 50◦ C. The yield of S.32 should be ∼40 g (0.25 mol, 50% relative to S.29). The product should be used for the preparation of PMPA as soon as possible. In the meantime, keep it in a deep freezer (–10◦ C). Compound characterization (S.32): C17 H20 O4 PS (320.4). 1 H NMR (DMSO-d6 ): δ 8.15 and 8.07 (s, 2H, H2 + H8), 7.29 (brs, 2H, NH2 ), 4.49 (m, 2H, P-OCH), 4.26 (dd, 1H, J1 a,2 = 3.7 Hz, Jgem = 14.4 Hz, H1 a), 4.17 (dd, 1H, J1 b,2 = 6.6 Hz, Jgem = 14.4 Hz, H1 b), 3.96 (m, 1H, H2 ), 3.79 (dd, 1H, JP,CH = 9.2 Hz, Jgem = 13.7 Hz, P-CH2 ), 3.72 (dd, 1H, JP,CH = 9.5 Hz, Jgem = 13.7 Hz, P-CH2 ), 1.19 (d, 3H), 1.16 (d, 3H), 1.15 (d, 3H), 1.12 (d, 3H), 1.08 (d, 3H, JCH3,CH = 6.1 Hz, CH3 ). Essentially identical conditions are used for the synthesis of (S)-2-O-(diisopropylphosphonylmethoxy)propyl p-tolylsulfonate from (S)-1-O-benzylpropane-1,2-diol.
STEPWISE METHOD FOR TENOFOVIR SYNTHESIS This approach consists of three steps: (1) preparation of the acyclic nucleoside, (2) formation of the phosphonomethylether linkage, and (3) cleavage of the protecting ester groups. For the first reaction, it is convenient to use optically active propane-1,2-diol cyclic carbonate, which gives in quantitative conversion the required product, which needs only to be recrystallized from ethanol. For the second reaction, the linkage is made using the C1 synthon used in the direct etherification approach.
ALTERNATE PROTOCOL
Materials Adenine Cesium carbonate Dimethylformamide (DMF) (R)-Propanediol-1,2-carbonate (S.26; Fig. 14.2.4) Toluene Ethanol Diethyl ether Sodium hydride, 60% dispersion in paraffin oil Diisopropyl p-tolylsulfonyloxymethylphosphonate (S.19; see Support Protocol 5) Acetic acid Chloroform Dowex 50 × 8, acid form (see recipe)
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Conc. aqueous ammonia Methanol Silica gel 250-mL round-bottom flask Magnetic stirrer with heating plate and silicon oil bath Rotary evaporator with diaphragm pump Glass filter funnel with filter flask 3 × 40–cm and 4 × 40–cm glass chromatography columns UV detector Fraction collector (optional) Calcium chloride protecting tube Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Synthesize (R)-9-(2-hydroxypropyl)adenine 1. Prepare a solution of 6.75 g (50 mmol) adenine and 1 g cesium carbonate in 70 mL DMF in a 250-mL round-bottom flask, and stir at 120◦ C for 20 min under a calcium chloride protecting tube. 2. Add 5 mL (6.1 g, 60 mmol) (R)-propanediol-1,2-carbonate in one portion and stir another 5 hr at 120◦ C under exclusion of moisture. According to TLC in 4:1 (v/v) chloroform/methanol, the conversion of adenine is quantitative.
3. Evaporate the volatiles under vacuum on a rotary evaporator and co-distill the residue with 50 mL toluene. This work-up should not be interrupted.
4. Recrystallize the crude product from boiling ethanol, adding ether until the solution remains turbid (i.e, does not clear with stirring). Leave at 4◦ C for crystals to form. 5. Collect the crystals by filtration, wash them with 50 mL cold ethanol and then with 100 mL diethyl ether, and dry under vacuum. The yield of S.25 should be ∼5 to 6 g (51% to 62%). An additional crop of product can be obtained from the mother liquor by deionization on a Dowex 50 × 8 column and recrystallization of the deionized material. The product can be stored indefinitely at room temperature.
Synthesize 9-(R)-[2-(diisopropylphosphonylmethoxy)propyl]adenine 6. Add 3.0 g of 60% sodium hydride in paraffin oil (1.8 g, 75 mmol NaH) to a solution of 4.83 g (25 mmol) S.25 and 10.6 g S.19 precooled to −30◦ C. Stir for 2 days with exclusion of moisture. For precautions, see Basic Protocol 1, step 1.
7. Slowly add 3 mL acetic acid and evaporate to dryness. 8. Extract with 300 mL boiling chloroform and evaporate. 9. Deionize the residue on a 3 × 40–cm column containing 200 mL Dowex 50 × 8 (acid form), and elute with diluted ammonia (1:10) in 20% aqueous methanol. This work-up should not be interrupted. Synthesis of Acyclic Nucleoside Phosphonates
10. Purify the residue on a 4 × 40–cm chromatography column containing 250 mL silica gel, eluting with chloroform. Monitor fractions by TLC using 4:1 (v/v) chloroform/ methanol. Dry to give the pure product as a foam.
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The yield of S.27 should be about 5.3 g (57%). It can be stored in closed bottles indefinitely at room temperature. Compound characterization (S.27): C16 H26 N5 O4 P (371.4). 1 H NMR (DMSO-d6 ): δ 7.79 and 7.49 (d, 2H, arom. H), 4.56 (m, 2H, P-OCH), 4.05 (dd, 1H, J = 3.2, 10.6 Hz, O-CH2 ), 3.91 (dd, 1H, J = 5.7, 10.6 Hz, O-CH2 ), 3.75 (m, 1H, O-CH), 3.72 (dd, 1H, JP,CH = 9.2 Hz, Jgem = 13.8 Hz, P-CH2 ), 3.64 (dd, 1H, JP,CH = 8.9 Hz, Jgem = 13.8 Hz, P-CH2 ), 2.42 (s, 3H, CH3 arom.), 1.22 (d, 6H) and 1.205 (d, 3H) and 1.20 (d, 3H, JCH3,CH = 6.1 Hz, CH3 ), 1.04 (d, 3H, JCH3,CH = 6.4 Hz, CH3 ). 13 C NMR (DMSO-d6 ): 145.12, 132.42, 130.35 (2C) and 127.82 (2C, C arom.), 74.52 (d, JP,C = 12.2, O-CH), 72.28 (O-CH2 ), 70.37 (d), 70.34 (d, JP,C = 6.4, P-OCH), 62.89 (d, JP,C = 165.5, P-CH2 ), 23.99 (d, 2C, JP,C = 3.9) and 23.84 (d, 2C, JP,C = 4.4, CH3 ), 21.27 and 15.71 (CH3 ).
Synthesize 9-(R)-[2-(phosphonomethoxy)propyl]adenine 11. Perform as described (see Basic Protocol 5, steps 7 to 14). SYNTHESIS OF 9-(S)-[3-HYDROXY2-(PHOSPHONOMETHOXY)PROPYL]ADENINE (HPMPA)
BASIC PROTOCOL 6
The series of so-called HPMP derivatives belongs to the oldest representatives of the ANP group (De Clercq et al., 1986). The parent molecule HPMPA (Fig. 14.2.6A) is active against DNA viruses and higher cellular parasites (protozoa; Kaminsky et al., 1996). For the majority of heterocyclic bases, it is only the (S) form that is active. The selection depends on the enantiospecificity of the nucleotide kinases that anabolize these nucleotide analogs. The most attractive compounds are derivatives of adenine, 2,6-diaminopurine, guanine, and cytosine (HPMPC or cidofovir; Fig. 14.2.6B). Cidofovir is approved for use and is widely applied to treatment of DNA viral diseases (De Clercq, 1993, 1998, 2002; Bray et al., 2002). Though a common synthon for HPMP derivatives has been developed (Hol´y, 1987), it is not very easy to make and has little value for preparation of more then a small quantity of the desired material. Practical syntheses follow the stepwise approach, which makes use of dialkyl p-tolylsulfonyloxymethylphosphonates. On an industrial scale the diethyl ester is used, but this protocol uses the diisopropyl ester for the same purpose. The synthesis is based of alkylation of the heterocyclic base with chiral trityloxymethyloxiranes (tritylglycidols), which proceeds easily and affords protected N-(3-trityloxy2-hydroxypropyl) derivatives, compounds which can also be obtained by tritylation of the 2,3-dihydroxypropyl derivatives (Hol´y, 1975). Reaction of adenine with trityl-(S)oxirane affords the crucial intermediate S.35, which has one hydroxyl group available for substitution by the phosphonate group to give S.37. Deprotection and purification by ion-exchange chromatography give the target product, S.38 (Hol´y et al., 1989b; Hol´y, 1993). Protection of the exocyclic amino group in basic nucleosides (S.36) is not absolutely required for the etherification reaction leading to phosphonates; however, it diminishes the extent of side reactions. A general method for specific N-benzoylation in the presence of hydroxyl functions makes use of silylation in pyridine, which takes place at the hydroxyl groups only. The most sensitive step in this protocol is the workup of the benzoylation mixture, which must be completed quickly to remove the large excess of ammonia as fast as possible (by evaporation in a flask of large volume-to-surface ratio, or by continuous dropwise addition of the cooled reaction mixture to the evacuated flask on a rotary evaporator). There are substantial losses of N-benzoyl derivative when this approach is applied to a larger scale than that described here. This problem can easily be circumvented by using N6 -benzoyladenine instead of adenine in the first reaction with (S)-tritylglycidol. Although not described in step format, this is illustrated for HPMPC in Fig. 14.2.6B.
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Figure 14.2.6
Schemes of the synthesis of (S)-HPMPA and (S)-HPMPC.
Materials
Synthesis of Acyclic Nucleoside Phosphonates
Trityl-(S)-glycidol Adenine Dimethylformamide (DMF) Cesium carbonate Chloroform Silica gel Pyridine Chlorotrimethylsilane Benzoyl chloride Conc. aqueous ammonia Magnesium sulfate, anhydrous Tetrahydrofuran Sodium hydride, 60% dispersion in paraffin oil Diisopropyl p-tolylsulfonyloxymethylphosphonate (S.19; see Support Protocol 5) Acetic acid Ethyl acetate Methanol Sodium methoxide Dowex 50 × 8, acid form (see recipe) Triethylamine Diethyl ether Ethanol Toluene Bromotrimethylsilane Acetonitrile Dowex 1 × 2, acetate form (see recipe) 500-mL round-bottom flask Magnetic stirrer with heating plate and silicon oil bath Rotary evaporator with a diaphragm pump 5 × 50–cm glass chromatography column 2-L Erlenmeyer flask
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Glass filter funnel with filter flask Reflux condenser Calcium chloride protecting tube Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E) Synthesize 9-(S)-(3-trityloxy-2-hydroxypropyl)adenine 1. Prepare a solution of 21 g (66 mmol) trityl-(S)-glycidol and 8.1 g (60 mmol) adenine in 250 mL DMF in a 500-mL round-bottom flask. Heat to 110◦ C under a calcium chloride protecting tube. 2. Add, while stirring, 1.8 g cesium carbonate. Continue heating and stirring under exclusion of moisture for 12 hr. 3. Evaporate under vacuum on a rotary evaporator. Extract with 500 mL chloroform and apply the extract onto a 5 × 50–cm glass chromatography column containing 300 mL silica gel. 4. Elute the product with chloroform, monitoring fractions by TLC in 95:5 (v/v) chloroform/methanol (Rf = 0.50). Combine product-containing fractions and dry to a foam. Yield of S.35 is 20 g (73%). The workup can be interrupted in any of the intermediate isolation steps.
Synthesize 9-(S)-(3-trityloxy-2-hydroxypropyl)-N6 -benzoyladenine 5. Combine 21.6 g (48 mmol) S.35 with 240 mL pyridine in a 1-L flask. Add 40 mL chlorotrimethylsilane and stir 1 hr. 6. Add 32 mL benzoyl chloride and stir for 2 hr. 7. Cool with ice and add 50 mL water followed by 120 mL conc. ammonia. Stir 30 min at 0◦ C. 8. Evaporate the volatiles quickly under vacuum. Use a 2-L flask and do not increase bath temperature over 30◦ C. 9. Extract the residue three times with 200 mL chloroform, wash the extract three times with 100 mL water, and dry with magnesium sulfate. 10. Filter and evaporate. Dry to a foam under vacuum.
Synthesize 9-(S)-[3-hydroxy-2-(diisopropylphosphonylmethoxy)propyl]adenine 11. Combine 18.0 g (32.5 mmol) S.36 with 100 mL tetrahydrofuran. Add 4.0 g of 60% sodium hydride in paraffin oil (2.40 g, 0.1 mol NaH) and stir at 0◦ C for 1 hr. 12. Add 12.6 g (36 mmol) of the tolylsulfonyl derivative S.19 and stir two days in a tightly closed flask. 13. Add 3.5 mL acetic acid and 350 mL ethyl acetate, extract the mixture three times with 100 mL water, and dry the organic phase.
Synthesize 9-(S)-[3-hydroxy-2-(phosphonomethoxy)propyl]adenine (HPMPA) 14. Evaporate the solvents under vacuum, redissolve the residue of crude S.37 in 270 mL boiling methanol, and add 30 mL of 1 M sodium methoxide in methanol. Let stand overnight at room temperature. 15. Neutralize the solution by adding Dowex 50 × 8 (acid form) and then add triethylamine to make the reaction alkaline.
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16. Filter and wash with 200 mL methanol. Evaporate the filtrate to dryness under vacuum. 17. Add to the residue 200 mL of 80% acetic acid in a preheated bath (110◦ C) and boil under reflux 40 min. 18. Evaporate under vacuum, add 250 mL water, and extract three times with 100 mL diethyl ether. Discard the extract, evaporate the aqueous phase, and dry it by codistillation with ethanol and toluene (100 mL each). 19. Add 10 mL bromotrimethylsilane to a suspension of 6.7 g (13.8 mmol) S.37 diester in 100 mL acetonitrile. Quickly close with a stopper, dissolve by shaking manually, and let stand 20 hr at room temperature. 20. Evaporate the volatiles under vacuum, and decompose by adding 50 mL water and allowing to sit for 15 min. 21. Add conc. aqueous ammonia to make the reaction alkaline (∼pH 10 to 11). Evaporate under vacuum. 22. Deionize the crude product on a 3 × 40–cm column containing 200 mL Dowex 50 × 8 (acid form), and elute with dilute ammonia (1:10) in 20% aqueous methanol. 23. Evaporate the dilute ammonia eluate and apply it on a glass column containing 150 mL Dowex 1 × 2 (acetate form). Wash with water until the UV absorption drops, and then elute the product (HPMPA) with 1 M acetic acid until the UV absorption drops. 24. Evaporate the product-containing eluate, co-distill the residue with 100 mL water, and dissolve in 20 mL boiling water. Add 80 mL ethanol and set it in the refrigerator to crystallize overnight. 25. Collect the crystals by filtration, wash with 100 mL ethanol and then 100 mL diethyl ether, and dry under vacuum. The yield of this deprotection step should be ∼7.3 g (74%). The product (S.38) can be stored in closed bottles for several years at room temperature. An additional amount of the pure product is contained in the filtrate and can be obtained by evaporation in vacuo. Compound characterization. C9 H14 N5 O5 P (303.2). 1 H NMR (500 MHz, D2 O+NaOD, ref(dioxane) = 3.75 ppm): δ 3.45 (dd, 1H, Jgem = 12.5 Hz, JbCH2 OH,2 = 5.4 Hz, bCH2 OH), 3.46 (d, 1H, Jgem = 12.3 Hz, JH,P = 9.6 Hz, H4 b), 3.51 (d, 1H, Jgem = 12.3 Hz, JH,P = 8.8 Hz, H4 a), 3.47 (dd, 1H, Jgem = 12.5 Hz, JaCH2 OH,2 = 3.5 Hz, aCH2 OH), 3.83 (m, 1H, H2 ), 4.35 (dd, 1H, Jgem = 14.8 Hz, J1 b,2 = 6.4 Hz, H1 b), 4.42 (dd, 1H, Jgem = 14.8 Hz, J1 a,2 = 4.6 Hz, H1 a), 8.20 (s, 1H, H2), 8.27 (s, 1H, H8). 13 C NMR (125.8 MHz, D2 O+NaOD, ref(dioxane) = 69.3 ppm): 46.55 (CH2 -1 ), 63.24 (CH2 -OH), 70.84 (d, JC,P = 151 Hz, CH2 -4 ), 82.63 (d, JC,P = 11 Hz, CH-2 ), 120.86 (C5), 146.18 (CH-8), 151.81 (C4), 155.09 (CH-2), 158.20 (C6).
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Synthesis of Acyclic Nucleoside Phosphonates
p-Anisaldehyde reagent To prepare p-anisaldehyde reagent, dissolve 9.2 mL p-methoxybenzaldehyde, 37.5 mL acetic acid, and 12.5 mL sulfuric acid in 338 mL of 96% (v/v) ethanol. The reagent is stable for several months at room temperature. Spray uniformly on the developed and air-dried TLC plate. Dry the plate and heat briefly with a heat gun to visualize the brown-violet spots on a crimson background. CAUTION: Work in a well-ventilated chemical fume hood!
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Dowex 1 × 2, acetate form Suspend 500 g Dowex 1 × 2 resin (100-200 mesh) in 0.5 M NAOH and let it stand overnight. Wash with 2 L water, resuspend in 2 L water, and add acetic acid to ∼1 M. Let stand overnight. Wash briefly with water. Store at room temperature. To regenerate resin batchwise, wash with 2 L water and let stand overnight in 0.5 to 1 M NaOH. Wash briefly with water and let stand overnight in 2 M acetic acid. Wash briefly with water. Keep wet resin at room temperature (stable for years). The recycling can be repeated for years. It is essential not to wash the resin too extensively with water, as it will become sensitive to microbial infection, which can destroy it completely, particularly at higher room temperatures. It is recommended to wash the recycled resin occasionally with ethanol.
Dowex 50 × 8, acid form Wash 500 g Dowex 50 × 8 resin (100-200 mesh) briefly with 2 L water, suspend in 2 L of 2 M HCl, and let stand overnight in a chemical fume hood. Filter through a glass filter and then wash with water until the pH of the filtrate is neutral. Store at room temperature. The resin capacity is ∼1 mequiv/mL.
To regenerate resin batchwise, wash with 2 L water and let stand overnight in 3 vol of 0.5 to 1 M NaOH. Wash briefly with water and let stand overnight in 3 vol of 2 M HCl. Filter through a glass filter and then wash with water until the pH of the filtrate is neutral. Keep at room temperature (stable for years). The recycling can be repeated for years.
HCl in DMF, 6 M Introduce dry hydrogen chloride into 50 mL dimethylformamide (DMF) in a 250-mL flask in an ice bath until the mixture begins to crystallize. Remove the cooling bath, add DMF until the crystals dissolve, and determine the content of HCl in a 1 mL aliquot by titration. Add DMF to give a 6 M final concentration. Keep in a tightly closed bottle at 4◦ C (stable > 1 year). p-Nitrobenzylpyridine reagent Dissolve 2.0 g p-nitrobenzylpyridine in 96% ethanol (100 mL). Spray uniformly over the TLC plate, dry it with a heat gun air stream, and keep it briefly over an open flame to ignite the fumes. After 2 to 4 sec, quench the flame by blowing the stream of air, and place the hot TLC plate quickly in an ammonia atmosphere in a closed vessel. The spots of alkylating compounds (e.g., the alkyl ester of the ANP) capable of quarternizing the pyridine system give a dark blue color reaction. COMMENTARY Background Information In order to display antimetabolic activity, nucleoside analogs must in nearly all cases undergo intracellular phosphorylation. This is catalyzed by nucleoside kinases that ought to be present in all somatic cells. The product is a 5 -nucleotide (nucleoside-5 -phosphate). This molecule is transformed to a 5 -diphosphate or 5 -triphosphate in the two subsequent steps catalyzed by nucleotide kinases and nucle-
oside diphosphate kinases, respectively. The latter molecules are the so-called active antimetabolites, which can then inhibit DNA or RNA synthesis, etc. Thus, nucleoside kinases play a key role in the activity of numerous antiviral and/or anticancer drugs opening the sequence mentioned. When these enzymes are present at low levels or are completely absent, the respective therapeutic efficacy is low or the drug does not act at all.
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Synthesis of Acyclic Nucleoside Phosphonates
It is not possible to transport highly polar monoesters of phosphoric acid through the lipophilic cellular membrane. In any case, there would be a massive dephosphorylation, so that following passage through the membrane into the cytoplasm only the parent nucleoside would remain. [Recently, many examples have arisen for circumventing the problem by lipophilic nucleotide prodrugs which pass through the membrane and liberate the free biologically active nucleotide in the cytoplasm (Beadle et al., 2002; Bradbury, 2002; Keith et al., 2003).] Such a cleavage is made impossible in the phosphonomethyl ether analogs of nucleotides, which preserve the isopolarity principle and adopt conformation that is very similar to that of the corresponding nucleotides. The phosphonomethyl ethers of certain acyclic nucleosides are thus capable of getting inside the cell and simulating a nucleotide (Hol´y, 2003). After conversion by nucleotide kinases and nucleoside diphosphate kinases, they are transformed to 5 -diphosphate or 5 -triphosphate analogs, respectively (Krejcov´a et al., 2000). These anabolites interfere with de novo DNA synthesis by their action on DNA polymerase or, in the case of retroviruses, on reverse transcriptase (Hol´y et al., 1990; Kramata et al., 1996, 1998; Pisarev et al., 1997; Birkus et al., 1999, 2001). There are various classes of these so-called acyclic nucleoside phosphonates (ANP), which differ by the character of the parent acyclic nucleoside and by the heterocyclic base. The latter is predominantly but not exclusively adenine, guanine, 2,6-diaminopurine (Balzarini et al., 1993), or some aza (Hol´y et al., 1996) or deaza (Dvor´akov´a and Hol´y, 1993) analog of these bases. Synthesis of acyclic nucleoside phosphonates makes use of two general approaches: (1) stepwise synthesis, which consists in the preparation of a protected acyclic nucleoside with one free hydroxyl group followed by introduction of the phosphonomethyl ether residue to this hydroxyl, and (2) the synthon approach, wherein the heterocyclic base is alkylated by a reagent that contains all features of the future side-chain. While the latter approach is applicable mainly when the reaction must be performed on a small scale (e.g., rare base, labeled compounds, larger series of related compounds or small conversion, low regiospecificity of alkylation), the former approach is used mainly as a largerscale preparation procedure. All these reactions afford as a product the diisopropyl ester
of phosphonomethyl ether and, in general, the ester protecting groups can be removed simply by trans-silylation followed by hydrolysis. This procedure consists of treatment of the diester intermediate with bromotrimethylsilane in acetonitrile, dimethylformamide, or a mixture of these solvents, followed by dissolution in water. There are different ways to achieve the final isolation, but the highly preferred method is ion-exchange chromatography. For compounds containing a protonatable amino function, it is best to bind them on a cation exchanger, elute them with ammonia, and purify the desalted product by anion-exchange chromatography using a volatile acid (acetic acid, formic acid) as an eluent. In the absence of a protonatable group, it is recommended to repeatedly evaporate the residue of the reaction mixture with an inert solvent before hydrolysis, pass the aqueous solution through a cationexchange resin, and repeatedly co-evaporate with water to remove the volatile hydrobromic acid. The dried product can usually be isolated as a pure crystalline solid.
Critical Parameters and Troubleshooting All the methods described here were verified on a nominal or higher scale. Potential hazards were identified and have been commented on within the protocols. The sodium salt of adenine is made by the action of sodium hydride on adenine, and makes use of a comparatively safe reagent protected by mineral oil. In most cases, the oil is not removed prior to the reaction, and remains in the reaction mixtures until removed by crystallization or chromatography. In large preparations where the oil can cause difficulties or increase the total mass of material to be purified, it can be easily washed out by hexane prior to the reaction. This will uncover the dangerous material, which is extremely sensitive to humidity. In this case, it is advantageous to use a specific device as shown in Figure 14.2.5. The NaH suspension can be weighed directly into it. A device of dimensions given in the figure easily takes 40 g of suspension, which corresponds to 1 mol of sodium hydride. The material is washed well with hexane and, while it is still wet, the receiver is closed by a glass stopper and connected with a diaphragm pump. The powder of dry NaH is then easily trasferred to the reaction flask. This device is used in the author’s laboratory for filtration of products and their immediate drying as well.
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Anticipated Results The yields in the individual preparations are given at the appropriate sites in the text. It is essential to use dry DMF. It is recommended to distill the DMF under vacuum from phosphorus pentoxide, leaving some distillation residue, and to keep the distilled DMF over molecular sieves. When treated in this fashion, the solvent is stable for a minimum of 5 months at room temperature. This procedure also removes dimethylamine present in the solvent. The alkylations (step 1 in Basic Protocols 1, 2 and 3) proceed quickly at elevated temperatures, and the optimum temperature indicated should be reached as soon as possible. However, it is not recommended to exceed a bath temperature of 120◦ C. All the alkylation products are sufficiently stable, and their isolation should not cause any problems, except for 6chloropurine (and 2-amino-6-chloro) derivatives. While in most cases it is quite practical to apply the alkylation in the presence of cesium carbonate, with sensitive bases it is more advantageous to use sodium salts generated in situ at (or below) room temperature and to heat up the reaction mixture carefully. These sodium salts of the bases mentioned are usually soluble in DMF, so it is possible to compensate for the reduced reactivity due to temperature loss by increasing the concentration of reactants. In the preparation of N6 -substituted purine ANPs, the strategic question is whether to perform the synthesis of the 6-chloropurine phosphonate diester intermediately followed by its reaction with the amine and cleavage of the ester functions (Fig. 14.2.2), or to prepare the amino-substituted base from the 6-chloro derivative first and treat it by either of the two methods of ANP synthesis (stepwise or synthon procedure). The transformation of the base usually does not cause problems, and alkylation of such adenine derivatives is usually regiospecific and easy to perform. Therefore, for the synthesis of a large quantity of a single compound, the second approach is preferred. On the other hand, the first approach is ideal for making guanine derivatives, and has also been used successfully to prepare a large number of N6 -substituted derivatives of adenine and 2,6-diaminopurine. The 2-hydroxyethyl or (R)-2-hydroxypropyl residues are introduced by treatment of the base with 1,2-ethylene or (R)-1,2propylene carbonate; for introduction of the 3-O-protected 2,3-dihydroxypropyl residue, it is most practical to use enantiomers of trityloxymethyloxirane for N-alkylation
of the purine and pyrimidine bases. This gives the appropriate intermediates bearing an isolated secondary hydroxyl group. For introduction of the phosphonomethyl ether function, condensation typically uses dialkyl p-tolylsulfonyloxymethylphosphonate in the presence of NaH. The dialkyl bromomethylphosphonate can also be used, and gives better results in certain cases. However, this reagent is not easily accessible and it is too expensive for routine use. The character of the alkyl ester group is determined by its stability and its ability to N-alkylate the base in the side reaction. This makes the methyl and ethyl esters unsuitable, and the group used in most cases is isopropyl.
Time Considerations Because some of these procedures are lengthy, time requirements are given both in calendar days and in terms of real active work performed by laboratory personnel. PMEA and PMEDAP. On average, preparation of S.2, S.3, S.6, and S.7 requires 2 calendar days each, with hands-on work not exceeding 5 hr each. PMEG. On average, preparation of S.9 requires 2 calendar days; however, the total time consumption depends on the automation of column chromatography or flash chromatography. Most of this period requires only 4 hr of hands-on work. Preparation of S.10 and S.11 requires 2 calendar days each, with a maximum 3 hr hands-on time each. PME synthon. An approximate average time required for preparation of S.15 is 3 calendar days, with hands-on work not exceeding 5 hr. Most of this time will be spent watching the proper functioning of the cylinder-joined gas-inlet system or distillation. The work-up can be interrupted (over the weekend) in step 7. The average time required for preparation of S.16 should be 1 day. This procedure should not be interrupted. PMPA by synthon method. An approximate average time required for preparation of PMPA (S.33) from the (R)-PMP synthon (S.32) is 2 days, with actual work time that does not exceed 8 hr. PMP synthon. The time required for preparation of the synthon depends on the route used. The average time required for preparation of S.22 (Support Protocol 2) and S.28 (Support Protocol 3) is 2 to 3 calendar days and 2 calendar days, respectively. To proceed via chloromethylation (Support Protocol 3), preparation of the synthon S.32 requires 3 calendar days. To proceed via direct
Biologically Active Nucleosides
14.2.35 Current Protocols in Nucleic Acid Chemistry
Supplement 22
etherification, preparation of the C1 synthon (Support Protocol 5) requires 2 days (with not more than 5 hr hands-on time), etherification to give S.29 (Support Protocol 4) requires 3 calendar days, and final preparation of S.32 (Support Protocol 6) requires another 3 calendar days. PMPA by two-step method. The condensation reaction (preparation of S.25) takes 2 days, with handwork time that does not exceed 3 hr, and addition of the phosphonomethylether linkage (S.27) requires another 4 days, with handwork time that does not exceed 6 hr. HPMPA. An approximate average time required for the whole sequence of reactions leading to HPMPA is 8 to 9 days, with handson work not exceeding 6 hr.
Literature Cited Andrei, G., Snoeck, R., Piette, J., Delvenne, V., and De Clercq, E. 1998. Antiproliferative effects of acyclic nucleoside phosphonates on human papillomavirus (HPV)-harboring cell lines compared with HPV-negative cell lines. Oncol. Res. 10:523-531. Balzarini, J., Naesens, L., Herdewijn, P., Rosenberg, I., Hol´y, A., Pauwels, R., Baba, M., Johns, D.G., and De Clercq, E. 1989. Marked in vivo antiretrovirus activity of 9-(2-phosphonylmethoxyethyl)adenine, a selective anti-human immunodeficiency virus agent. Proc. Natl. Acad. Sci. U.S.A. 86:332336.
Bobkova, K., Otov´a, B., Marinov, I., Mandys, V., Panczak, A., Votruba, I., and Hol´y, A. 2000. Anticancer effect of PMEDAP—monitoring of apoptosis. Anticancer Res. 20:1041-1047. Bobkova, K., Gut, I., Mandys, V., Hol´y, A., Votruba, I., and Otov´a, B. 2001. Antitumour activity of a combined treatment with PMEDAP and docetaxel in the Prague inbred Sprague-Dawley/cub rat strain bearing T-cell lymphoma. Anticancer Res. 21:2725-2731. Bradbury, J. 2002. Orally available cidofovir derivative active against smallpox. LANCET 359:1041. Bray, M., Martinez, M., Kefauver, D., West, M., and Roy, C. 2002. Treatment of aerosolized cowpox virus infection in mice with aerosolized cidofovir. Antiviral Res. 54:129-142. Compton, M.L., Toole, J.J., and Paborsky, L.R. 1999. 9-(2-Phosphonylmethoxyethyl)-N6 – cyclopropyl-2,6-diaminopurine (cpr-PMEDAP) as a prodrug of 9-(2-phosphonylmethoxy ethyl)guanine (PMEG). Biochem. Pharmacol. 58:709-714. De Clercq, E. 1993. Therapeutic potential of HPMPC as an antiviral drug. Rev. Med. Virol. 3:85-96. De Clercq, E. 1998. Towards an effective chemotherapy of virus infections: Therapeutic potential of cidofovir [(S)-1-[3-hydroxy-2(phosphonomethoxy)propyl]cytosine, HPMPC] for the treatment of DNA virus infections. Collect. Czech. Chem. Commun. 63:480-506.
Balzarini, J., Naesens, L., Slachmuylders, J., Niphuis, H., Rosenberg, I., Hol´y, A., Schellekens, H., and De Clercq, E. 1991. 9-(2Phosphonylmethoxyethyl)adenine (PMEA) effectively inhibits retrovirus replication in vitro and simian immunodeficiency virus infection in rhesus monkeys. AIDS 5:21-28.
De Clercq, E. 2002. Cidofovir in the treatment of poxvirus infections. Antiviral Res. 55:1-13.
Balzarini, J., Hol´y, A., Jindrich, J., Naesens, L., Snoeck, R., Schols, D., and De Clercq, E. 1993. Differential antiherpesvirus and antiretrovirus effects of the (S) and (R) enantiomers of acyclic nucleoside phosphonates: Potent and selective in vitro and in vivo antiretrovirus activities of (R)-9-(2-phosphonomethoxypropyl)-2,6diaminopurine. Antimicrob. Agents Chemother. 37:332-338.
De Clercq, E., Sakuma, T., Baba, M., Pauwels, R., Balzarini, J., Rosenberg, I., and Hol´y, A. 1987. Antiviral activity of phosphonylmethoxyalkyl derivatives of purine and pyrimidines. Antiviral Res. 8:261-272.
Beadle, J.R., Hartline, C., Aldern, K.A., Rodriguez, N., Harden, E., Kern, E.R., and Hostetler, K.Y. 2002. Alkoxyalkyl esters of cidofovir and cyclic cidofovir exhibit multiple-log enhancement of antiviral activity against cytomegalovirus and herpesvirus replication in vitro. Antimicrob. Agents Chemother. 46:2381-2386.
Synthesis of Acyclic Nucleoside Phosphonates
Birkus, G., Votruba, I., Otmar, M., and Hol´y, A. 2001. Interactions of 1-[(S)-3-hydroxy2-(phosphonomethoxy)propyl]cytosine (cidofovir) diphosphate with DNA polymerases a, d and e∗ . Collect. Czech. Chem. Commun. 66:1698-1706.
Birkus, G., Votruba, I., Hol´y, A., and Otov´a, B. 1999. 9-[2-(Phosphonomethoxy)ethyl]adenine diphosphate (PMEApp) as a substrate toward replicative DNA polymerases alpha, delta, epsilon, and epsilon∗ . Biochem. Pharmacol. 58:487-492.
De Clercq, E., Hol´y, A., Rosenberg, I., Sakuma, T., Balzarini, J., and Maudgal, P.C. 1986. A novel selective broad-spectrum anti-DNA virus agent. Nature 323:464-467.
De Clercq, E., Andrei, G., Balzarini, J., Hatse, S., Liekens, S., Naesens, L., Neyts, J., and Snoeck, R. 1999. Antitumor potential of acyclic nucleoside phosphonates. Nucleosides Nucleotides 18:759-771. Dvor´akov´a, H. and Hol´y, A. 1993. Synthesis and biological effects of N-(2phosphonomethoxyethyl) derivatives of deazapurine bases. Collect. Czech. Chem. Commun. 58:1419-1429. Dvor´akov´a, H., Hol´y, A., Votruba, I., and Masoj´ıdkov´a, M. 1993. Synthesis and biological effects of acyclic analogs of deazapurine nucleosides. Collect. Czech. Chem. Commun. 58:629648.
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Hadziyannis, S.J., Tassopoulos, N.C., Heathcote, E.J., Chang, T.T., Kitis, G., Rizzetto, M., Marcellin, P., Lim, S.G., Goodman, Z., Wulfsohn, M.S., Xiong, S., Fry, J., and Brosgart, C.L. 2003. Adefovir dipivoxil for the treatment of hepatitis B e antigen-negative chronic hepatitis B. N. Engl. J. Med. 348:800-807. Hatse, S., Naesens, L., De Clercq, E., and Balzarini, J. 1999. N-6-Cyclopropyl-PMEDAP: A novel derivative of 9-(2-phosphonylmethoxyethyl)2,6-diaminopurine (PMEDAP) with distinct metabolic, antiproliferative, and differentiationinducing properties. Biochem. Pharmacol. 58:311-323. Hol´y, A. 1975. Aliphatic analogs of nucleosides, nucleotides and oligonucleotides. Collect. Czech. Chem. Commun. 40:187-214. Hol´y, A. 1987. Phosphonylmethyl analogs of nucleotides and their derivatives: Chemistry and biology. Nucleosides Nucleotides 6:147-155. Hol´y, A. 1993. Syntheses of enantiomeric N-(3hydroxy-2-phosphonomethoxypropyl) derivatives of purine and pyrimidine bases. Collect. Czech. Chem. Commun. 58:649-674. Hol´y, A. 2003. Phosphonomethoxyalkyl analogs of nucleotides. Curr. Pharm. Des. 9:2567-2592. Hol´y, A. and Masoj´ıdkov´a, M. 1995. Synthesis of enantiomeric N-(2-phosphonomethoxypropyl) derivatives of purine and pyrimidine bases. 1. The stepwise approach. Collect. Czech. Chem. Commun. 60:1196-1212.
the carbon atoms of the base. J. Med. Chem. 42:2064-2086. Hol´y, A., Votruba, I., Tloustov´a, E., and Masoj´ıdkov´a, M. 2001. Synthesis and cytostatic activity of N-[2-(phosphonomethoxy)alkyl] derivatives of N6 -substituted adenines, 2,6diaminopurines and related compounds. Collect. Czech. Chem. Commun. 66:1545-1592. Kaminsky, R., Schmid, C., Grether, Y., Hol´y, A., De Clercq, E., Naesens, L., and Brun, R. 1996. (S)-9-(3-Hydroxy-2-phosphonylmethoxypropyl)adenine [(S)-HPMPA]: A purine analog with trypanocidal activity in vitro and in vivo. Trop. Med. Int. Health 1:255-263. Keith, K.A., Hitchcock, M.J.M., Lee, W.A., Hol´y, A., and Kern, E.R. 2003. Evaluation of nucleoside phosphonates and their analogs and prodrugs for inhibition of orthopoxvirus replication. Antimicrob. Agents Chemother. 47:21932198. Kramata, P., Votruba, I., Otov´a, B., and Hol´y, A. 1996. Different inhibitory potencies of acyclic phosphonomethoxyalkyl nucleotide analogs toward DNA polymerases alpha, delta and epsilon. Mol. Pharmacol. 49:1005-1011. Kramata, P., Downey, K.M., and Paborsky, L.R. 1998. Incorporation and excision of 9-(2phosphonylmethoxyethyl)guanine (PMEG) by DNA polymerase delta and epsilon in vitro. J. Biol. Chem. 273:21966-21971.
Hol´y, A., Rosenberg, I., and Dvor´akov´a, H. 1989a. Synthesis of N-(2-phosphonylmethoxyethyl) derivatives of heterocyclic bases. Collect. Czech. Chem. Commun. 54:2190-2210.
Kreider, J.W., Balogh, K., Olson, R.O., and Martin, J.C. 1990. Treatment of latent rabbit and human papillomavirus infections with 9(2-phosphonylmethoxy)ethylguanine (PMEG). Antiviral Res. 14:51-58.
Hol´y, A., Rosenberg, I., and Dvor´akov´a, H. 1989b. Synthesis of (3-hydroxy-2-phosphonylmethoxypropyl) derivatives of heterocyclic bases. Collect. Czech. Chem. Commun. 54:2470-2501.
Krejcov´a, R., Horsk´a, K., Votruba, I., and Hol´y, A. 2000. Interaction of guanine phosphonomethoxyalkyl derivatives with GMP kinase isoenyzmes. Biochem. Pharmacol. 60:19071913.
Hol´y, A., Votruba, I., Merta, A., Cern´y, J., Vesel´y, J., Vlach, J., Sediv´a, K., Rosenberg, I., Otmar, M., and Hrebabeck´y, H. 1990. Acyclic nucleotide analogs: Synthesis, antiviral activity and inhibitory effects on some cellular and virus-encoded enzymes in vitro. Antiviral Res. 13:295-311.
Naesens, L. and De Clercq, E. 1997. Therapeutic potential of HPMPC (cidofovir), PMEA (adefovir) and related acyclic nucleoside phosphonate analogs as broad-spectrum antiviral agents. Nucleosides Nucleotides 16:983-992.
Hol´y, A., Dvor´akov´a, H., and Masoj´ıdkov´a, M. 1995. Synthesis of enantiomeric N-(2phosphonomethoxypropyl) derivatives of heterocyclic bases. 2. Synthon approach. Collect. Czech. Chem. Commun. 60:1390-1409. Hol´y, A., Dvor´akov´a, H., Jindrich, J., Masoj´ıdkov´a, M., Budesinsk´y, M., Balzarini, J., Andrei, G., and De Clercq, E. 1996. Acyclic nucleotide analogs derived from 8-azapurines: Synthesis and antiviral activity. J. Med. Chem. 39:40734088. Hol´y, A., Gunter, J., Dvor´akov´a, H., Masoj´ıdkov´a, M., Andrei, G., Snoeck, R., Balzarini, J., and De Clercq, E. 1999. Structure-antiviral activity relationship in the series of pyrimidine and purine N-[2-(2-phosphonomethoxy)ethyl] nucleotide analogs. 1. Derivatives substituted at
Naesens, L., Balzarini, J., Rosenberg, I., Hol´y, A., and De Clercq, E. 1989. 9-(2Phosphonylmethoxyethyl ) - 2,6 - diaminopurine (PMEDAP): A novel agent with anti-human immunodeficiency virus activity in vitro and potent anti-Moloney murine sarcoma virus activity in vivo. Eur. J. Clin. Microbiol. Infect. Dis. 8:1043-1047. Naesens, L., Neyts, J., Balzarini, J., Hol´y, A., Rosenberg, I., and De Clercq, E. 1993. Efficacy of oral 9-(2-phosphonylmethoxyethyl)2,6-diaminopurine (PMEDAP) in the treatment of retrovirus and cytomegalovirus infections in mice. J. Med. Virol. 39:167-172. Naesens, L., Balzarini, J., and De Clercq, E. 1994. Therapeutic potential of PMEA as an antiviral drug. Rev. Med. Virol. 4:147-159. Naesens, L., Hatse, S., Segers, C., Verbeken, E., Declercq, E., Waer, M., and Balzarini,
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J. 1999. 9-(2-Phosphonylmethoxyethyl)-N(6)cyclopropyl-2,6-diaminopurine: A novel prodrug of 9-(2-phosphonylmethoxyethyl)guanine with improved antitumor efficacy and selectivity in choriocarcinoma-bearing rats. Oncol. Res. 11:195-203. Otov´a, B., Z´ıdek, Z., Hol´y, A., Votruba, I., Sladk´a, M., Marinov, I., and Leskov´a, V. 1997. Antitumor activity of novel purine acyclic nucleotide analogs PMEA and PMEDAP. In Vivo 11:163167. Otov´a, B., Francov´a, K., Franek, F., Koutn´ık, P., Votruba, I., Hol´y, A., Sladk´a, M., and Schramlov´a, J. 1999. 9-[2(Phosphonomethoxy)ethyl]-2,6-diaminopurine (PMEDAP)—a potential drug against hematological malignancies—induces apoptosis. Anticancer Res. 19:3173-3182. Palmer, S., Buckheit, R.W., Gilbert, H., Shaw, N., and Miller, M.D. 2000. AntiHIV-1 activity of PMEA, PMPA (9-(2phosphonylmethoxypropyl)adenine) and AZT against HIV-1 Subtypes A, B, C, D, E, F, G and O. Antiviral Res. 46:21.
Pisarev, V.M., Lee, S.H., Connelly, M.C., and Fridland, A. 1997. Intracellular metabolism and action of acyclic nucleoside phosphonates on DNA replication. Mol. Pharmacol. 52:63-68. Valerianov´a, M., Votruba, I., Hol´y, A., Mandys, V., and Otov´a, B. 2001. Antitumour activity of N6-substituted PMEDAP derivatives against T-cell lymphoma. Anticancer Res. 21:20572064. Valerianov´a, M., Otov´a, B., Bil´a, V., Hanzalov´a, J., Votruba, I., Hol´y, A., Eckschlager, T., Krejci, O., and Trka, J. 2003. PMEDAP and its N6-substituted derivatives: Genotoxic effect and apoptosis in in vitro conditions. Anticancer Res. 23:4933-4939.
Contributed by Antonin Hol´y Institute of Organic Chemistry and Biochemistry Academy of Sciences of the Czech Republic Prague, Czech Republic
Synthesis of Acyclic Nucleoside Phosphonates
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Synthesis of β-L-2 -Deoxythymidine (L-dT) The “unnatural” L-nucleoside β-L-2 -deoxythymidine (L-dT) is a potent, specific, and selective inhibitor of the replication of hepatitis B virus (HBV) both in vitro and in vivo (Bryant et al., 2001; Han, 2005). This compound has been shown to exhibit an excellent safety profile in preclinical testing and is currently being evaluated in Phase III clinical trials. This unit describes in detail a novel strategy for the preparation of L-dT (Gosselin et al., 2004) involving a multi-step synthetic procedure which can be applied for the synthesis of β-L-2 -deoxythymidine on a 10-g scale (Fig. 14.3.1), quantities required to conduct preclinical in vitro and in vivo studies. This procedure (Gosselin et al., 2004) involves the direct condensation of thymine with properly acylated-β-L-ribofuranose (S.2; Recondo and Rinderknecht, 1959), followed by full deacylation, 5 ,3 -O-selective protection, radical 2 -deoxygenation, and finally removal of all protecting groups. 1-O-Acetyl-2,3,5-tri-O-benzoyl-β-L-ribofuranose (S.2) is not commercially available and requires synthesis from L-ribose using the three-step sequence developed for the corresponding D-enantiomer (Cimpoa et al., 1994). This procedure, which produces L-dT in six steps with an overall satisfactory yield, is general and may be applied to other 2 -deoxynucleosides incorporating different heterocyclic bases.
UNIT 14.3 BASIC PROTOCOL
CAUTION: All reactions must be run in a suitable fume hood with efficient ventilation. Safety glasses, a protective laboratory coat, and reagent-impermeable protective gloves should be worn.
Materials Argon gas source L-Ribose (S.1; Interchim), dried over phosphorous pentoxide (P2 O5 ) Methanol and anhydrous methanol 96% (v/v) sulfuric acid (H2 SO4 ) in water Chloroform and anhydrous chloroform 10% (v/v) sulfuric acid (H2 SO4 ) in 95% ethanol Pyridine and dry pyridine Benzoyl chloride, anhydrous (BzCl) Dichloromethane (CH2 Cl2 ) Saturated aqueous sodium bicarbonate (aq. NaHCO3 ) Sodium sulfate, anhydrous (Na2 SO4 ) Toluene Acetic acid (AcOH) Acetic anhydride (Ac2 O) Petroleum ether Diethyl ether Ethyl acetate (EtOAc) 95% ethanol (EtOH) Thymine Ammonium sulfate [(NH4 )2 SO4 ] Hexamethyldisilazane (HMDS) 1,2-Dichloroethane, dry Trimethylsilyl trifluoromethanesulfonate (TMSOTf) Silica gel (Merck 60H) Sand Biologically Active Nucleosides Contributed by Claire Pierra, David Dukhan, Gilles Gosselin, and Jean-Pierre Sommadossi Current Protocols in Nucleic Acid Chemistry (2006) 14.3.1-14.3.9 C 2006 by John Wiley & Sons, Inc. Copyright
14.3.1 Supplement 24
Figure 14.3.1 Synthesis scheme for the preparation of β-L-2 -deoxythymidine (L-dT) (S.7) from L-ribose (S.1). The expected yields are given in parentheses. Abbreviations: AIBN, α,α -azoisobutyronitrile; Bz, benzoyl; DMAP, 4dimethylaminopyridine; HMDS, hexamethyldisilazane; TIPSCl2 , 1,3-dichloro-1,1,3,3-tetramethyldisiloxane; TMSOTf, trimethylsilyl trifluoromethanesulfonate; TTMSS, tris(trimethylsilyl)silane.
Synthesis of β-L-2 Deoxythymidine
Methanol saturated with ammonia (MeOH/NH3 , see recipe) 1,3-Dichloro-1,1,3,3-tetraisopropyldisiloxane (TIPSCl2 ) Acetonitrile, anhydrous 4-Dimethylaminopyridine (DMAP) O-Phenyl chlorothionoformate [PhOC(S)Cl] 0.5 N hydrochloric acid (HCl) in water Dioxane, dry α,α -Azoisobutyronitrile (AIBN) Tris(trimethylsilyl)silane (TTMSS) Ammonium fluoride (NH4 F) Absolute ethanol 250-, 500-, 1000-, and 2000-mL round-bottom flasks, oven dried Magnetic stirrer and stir bars Silica gel thin-layer chromatography (TLC) plates (Merck, Kieselgel 60 F254 , 0.2-mm thickness) Heat gun Rotary evaporator equipped with a vacuum pump 200-mL dropping funnels 1-, 2-, and 3-L separatory funnels 500- and 1000-mL Erlenmeyer flasks Filter funnel Vacuum desiccator with P2 O5 80◦ and 100◦ C oil baths Reflux condensers Glass chromatography columns: 8 × 50–cm, 6 × 30–cm, 7 × 15–cm Steel cylinder Splash head Additional reagents and equipment for TLC (APPENDIX 3D) and column chromatography (APPENDIX 3E)
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Current Protocols in Nucleic Acid Chemistry
Prepare acylated β-L-ribose (S.2) 1. In an oven-dried 2-L round-bottom flask equipped with a stir bar and argon gas source, dissolve 50 g of dry L-ribose S.1 in 750 mL dry methanol. While stirring at 0◦ C (on an ice bath), add 4 mL of 96% H2 SO4 . 2. Stir overnight or until the reaction is complete. Monitor the reaction by TLC (APPENDIX 3D) on silica gel plates using 80:20 (v/v) CH2 Cl2 /MeOH. Visualize bands by dipping the plate in 10% (v/v) sulfuric acid in 95% EtOH and then heating with a heat gun (2000 W, 600◦ C). The starting material should be run alongside the reaction for comparison. Typically, this intermediate has an Rf value of 0.44.
3. Neutralize the reaction mixture with 200 mL pyridine and remove the solvents using a rotary evaporator equipped with a vacuum pump to give a yellow oil. 4. Co-evaporate the crude product three times with 50 mL dry pyridine. Dissolve the residue in a mixture of 350 mL anhydrous pyridine and 150 mL anhydrous chloroform, and immerse the 2-L reaction flask in an ice bath. Add 190 mL BzCl dropwise using a 200-mL dropping funnel, keeping the temperature below 4◦ C throughout the addition. The BzCl, which hydrolyzes when exposed to water, should be stored protected from moisture and flushed with argon after use. During the reaction, some precipitate (pyridinium salts) will appear and the reaction will become dark pink.
5. Warm the reaction mixture to room temperature and keep stirring for 4 hr. Monitor the reaction by TLC using 95:5 (v/v) CH2 Cl2 /MeOH, and visualize the bands by UV shadowing (APPENDIX 3D) and by dipping the plate in 10% (v/v) sulfuric acid in 95% EtOH and then heating with a heat gun (2000 W, 600◦ C). Apply both of these visualization methods to all subsequent TLC steps. Typically, this intermediate has an Rf value of 0.75 and the starting material (S.1) has an Rf of 0.06 under these conditions.
6. Dilute reaction with 200 mL chloroform and carefully add to 500 mL aq. NaHCO3 solution. Pour the mixture into a 2-L separatory funnel and allow the phases to separate. Wash the organic layer with 500 mL water and pour into a 1000-mL Erlenmeyer flask. Dry the organic layer over anhydrous Na2 SO4 . Filter under vacuum using a filter funnel, and evaporate the filtrate to dryness on a rotary evaporater equipped with a vacuum pump to give a brown syrup. Use ice-cold aq. NaHCO3 solution if the reaction is too exothermic.
7. Azeotrope (co-evaporate) the residue three times with 50 mL toluene. Dissolve the crude product in 80 mL acetic acid, place the flask in an ice bath under stirring, and then add 185 mL acetic anhydride. Finally, add 27 mL of 96% H2 SO4 slowly (over 30 min) using a 200-mL dropping funnel. 8. Warm the reaction to room temperature and keep stirring for 3 hr until the reaction is complete. Monitor by TLC using 1:1 (v/v) petroleum ether/diethyl ether, and visualize the bands by UV shadowing and with sulfuric acid/EtOH. Typically, the Rf value of the desired compound (S.2) is 0.21.
9. Quench by adding 50 mL water directly into the reaction flask, then dilute with 200 mL EtOAc. Pour the mixture into a 3-L separatory funnel, wash the organic
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layer with saturated aqueous NaHCO3 , dry the organic layer over anhydrous Na2 SO4 , filter, and evaporate to dryness in a rotary evaporator. CAUTION: Add the aq. NaHCO3 solution carefully, as the neutralization can be violent.
10. Crystallize the crude product from 500 mL of 95% ethanol for several days at 0◦ to 5◦ C, filter on a filter funnel under vacuum, and check purity of the product. 1-O-Acetyl-2,3,5-tri-O-benzoyl-β-L-ribofuranose (S.2): 44 g (44% yield from S.1). TLC (1:1 (v/v) petroleum ether/diethyl ether): Rf = 0.21. m.p. 122◦ -123◦ C (lit. for D-enantiomer: 121◦ to 123◦ C). 1 H NMR (400 MHz, DMSO-d6 ): δ 7.8-7.1 (15H, m, Bz), 6.1 (1H, s, H1), 5.6 (1H, m, H2), 5.5 (1H, d, J = 4.8 Hz, H3), 4.6 (1H, m, H4), 4.5-4.3 (2H, m, H5 and H5 ), 1.7 (3H, s, CH3 ). FAB-MS (GT-matrix): 597 [M+G+H]+ , 445 [M-CH3 COO]+ , 383 [M-C6 H5 COO]+ , 105 [C6 H5 CO]+ .
Silylate thymine 11. Dry 5.3 g thymine and 500 mg ammonium sulfate in an oven-dried 500-mL roundbottom flask in a vacuum desiccator over P2 O5 for 1 full day. 12. Dissolve dry mixture in 230 mL HMDS. 13. Put the flask in a 100◦ C oil bath and allow the reaction mixture to reflux overnight using a reflux condenser under an argon atmosphere. Once the reaction mixture becomes clear, the reaction is considered to be complete. The silylated base must be manipulated under an argon atmosphere to avoid removal of the silylated groups.
14. Let reaction cool to room temperature and evaporate excess solvents under reduced pressure using a rotary evaporator.
Condense S.2 with silylated thymine 15. Dissolve the silylated base residue in 80 mL anhydrous 1,2-dichloroethane. To this solution, add successively a solution of 17.6 g acylated L-ribose (S.2) in 190 mL anhydrous 1,2-dichloroethane, and 13.5 mL TMSOTf. Stir this solution for 3 hr at room temperature. 16. Monitor the reaction by TLC using 80:20 (v/v) CH2 Cl2 /EtOAc, and visualize the bands by UV shadowing and with sulfuric acid/ethanol. The starting sugar (S.2) should be run alongside the reaction to follow its disappearance. Typically, the desired product has an Rf value of 0.45 and the starting sugar has an Rf of 0.77 under these conditions.
17. Quench the reaction mixture with 10 mL water, dilute with 100 mL chloroform, and carefully wash with 300 mL aq. NaHCO3 in a 1-L separatory funnel. 18. Dry the organic layer over anhydrous Na2 SO4 , filter, and evaporate to dryness on a rotary evaporator. 19. Prepare a slurry of silica gel in chloroform. Pour slurry into an 8 × 50–cm chromatography column and carefully layer 2 cm sand on top of slurry (see APPENDIX 3E for column chromatography). 20. Dissolve the crude product in a minimal amount of CH2 Cl2 and carefully layer on top of the column.
Synthesis of β-L-2 Deoxythymidine
21. Start eluting with 2 L CH2 Cl2 , then with a linear gradient (0% to 10%) of EtOAc in CH2 Cl2 . Collect 500-mL fractions and combine those that contain pure product S.3 as determined by TLC.
14.3.4 Supplement 24
Current Protocols in Nucleic Acid Chemistry
22. Evaporate combined fractions to dryness in the rotary evaporator and dry overnight under vacuum. 23. Check the purity of the product. 1-[2,3,5-Tri-O-benzoyl-β-L-ribofuranosyl]thymine (S.3): 15 g (75% yield from S.2). TLC (80:20 (v/v) dichloromethane/ethyl acetate): Rf = 0.45. 1 H NMR (400 MHz, DMSO-d6 ): δ 11.48 (1H, s, NH, D2 O exchangeable), 8.0-7.4 (16H, m, Bz and H6), 6.19 (1H, d, J = 4.3 Hz, H1 ), 5.9 (2H, m, H2 and H3 ), 4.7-4.6 (3H, m, H4 , H5 and H5 ), 1.67 (3H, s, CH3 ). FAB-MS (GT-matrix): 571 [M+H]+ , 445 [M-B]+ , 105 [C6 H5 CO]+ ; 569 [M-H]– , 125 [B]– , 121 [C6 H5 COO]– .
Deprotect S.3 24. Treat 14.8 g of S.3 with 300 mL MeOH/NH3 for 3 days at room temperature in a steel cylinder. 25. Cool down the cylinder in an ice bath and carefully open it to check the reaction by TLC using 85:15 (v/v) CH2 Cl2 /MeOH. Visualize bands by UV shadowing and with sulfuric acid/EtOH. The protected nucleoside should be run alongside the reaction to follow its disappearance. Typically, the desired 1-[β-L-ribofuranosyl]thymine (S.4) has an Rf value of 0.35.
26. When deprotection is complete, bubble argon through the solution for 1 hr to remove most of the ammonia, then evaporate the resulting solution to dryness on a rotary evaporator. 27. Dissolve the crude product in a minimal amount of absolute ethanol and place the flask for several days at 0◦ to 5◦ C. Filter crystals (4.6 g; 69% yield from S.3) on a filter funnel under vacuum. 28. Analyze the product by TLC, melting-point determination, 1 H NMR, MS, UV spectroscopy, and optical rotation determination. 1-[β-L-Ribofuranosyl]thymine (S.4): TLC (85:15 (v/v) dichloromethane/methanol): Rf = 0.35. m.p. 122◦ -123◦ C. 1 H NMR (400 MHz, DMSO-d6 ): δ 11.29 (1H, s, NH, D2 O exchangeable), 7.72 (1H, d, J = 1.0 Hz, H6), 5.76 (1H, d, J = 5.6 Hz, H1 ), 5.32 (1H, d, J = 5.8 Hz, 2 -OH, D2 O exchangeable), 5.1 (1H, t, J = 5.2 Hz, 5 -OH, D2 O exchangeable), 5.1 (1H, d, J = 5.0 Hz, 3 -OH, D2 O exchangeable), 4.0 (1H, m, H2 ), 3.9 (1H, m, H3 ), 3.8 (m, H4 ), 3.6-3.3 (2H, m, H5 and H5 ), 1.76 (3H, s, CH3 ). FAB-MS (GT-matrix): 259 [M+H]+ , 127 [B+2H]+ ; 257 [M-H]– , 125[B]– . UV (H2 O): λmax = 267.0 nm (ε 9700), λmin = 235.0 nm (ε 2200). [α]D 20 +9.0 (c = 1.0, H2 O).
Perform 3 ,5 -O-silylation 29. Co-evaporate 4.5 g of S.4 two times with 30 mL anhydrous pyridine and then dissolve in 34 mL anhydrous pyridine. 30. Add 6.6 mL TIPSCl2 and stir the reaction overnight at room temperature. Monitor the reaction by TLC using 85:15 (v/v) CH2 Cl2 /MeOH, and visualize bands by UV shadowing and with sulfuric acid/EtOH. The TIPSCl2 , which hydrolyzes when exposed to water, should be stored protected from moisture and flushed with argon after use. These plates are developed using 85:15 (v/v) dichloromethane/methanol to see the disappearance of starting material as well as the formation of the desired product. Typically, the desired product S.5 has an Rf value of 0.77.
31. When the reaction is finished, quench with 100 mL water, dilute mixture with 200 mL EtOAc, and wash successively with 200 mL aq. NaHCO3 and 200 mL water. Dry the organic layer over Na2 SO4 , filter, and evaporate on a rotary evaporator to obtain the crude product.
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32. Prepare a 6 × 30–cm silica gel chromatography column as described in step 19. Apply the residue (dissolved in a minimum volume of toluene) to the column and elute with a mixture of 80:20 (v/v) toluene/EtOAc. Collect 250-mL fractions and combine those that contain pure product S.5 as determined by TLC using 70:30 toluene/EtOAc. 33. Check purity of the product. 1-[3,5-O-(1,1,3,3-Tetraisopropyl-1,3-disiloxanyl)-β-L-ribofuranosyl]thymine (S.5): 8.6 g (61% yield from S.4). TLC (70:30 (v/v) toluene/ethyl acetate): Rf = 0.14. 1 H NMR (400 MHz, DMSO-d6 ): δ 11.36 (1H, s, NH, D2 O exchangeable), 7.7(1H, s, H6), 5.43 (1H, d, J = 1.1 Hz, H1 ), 5.39 (1H, d, J = 4.9 Hz, 2 -OH, D2 O exchangeable), 4.1-3.8 (5H, 3m, H2 , H3 , H4 , H5 and H5 ), 1.64 (3H, s, CH3 ), 1.0-0.8 (24H, m, isopropyl). FAB-MS (GT-matrix): 501 [M+H]+ , 261 [TIPSO]+ , 127 [B+2H]+ ; 499 [M-H]– , 125 [B]– .
Perform 2 -deoxygenation 34. Dissolve 8.5 g of S.5 in 70 mL anhydrous acetonitrile. 35. Add 4.15 g of DMAP and 2.6 mL PhOC(S)Cl and stir 19 hr at room temperature. The PhOC(S)Cl, which hydrolyzes when exposed to water, should be stored protected from moisture and flushed with argon after use.
36. Analyze the reaction by TLC using 70:30 (v/v) toluene/EtOAc and visualize the bands by UV shadowing and with sulfuric acid/EtOH. The silylated nucleoside should be run alongside the reaction to follow its disappearance. Typically, the thiophenoxy intermediate has an Rf value of 0.37.
37. Remove the solvent under vacuum on a rotary evaporator, then dilute the residue with 200 mL EtOAc and add 200 mL of 0.5 N aq. HCl solution. 38. Pour the mixture into a clean 1-L separatory funnel and allow the phases to separate. Pour the organic phase into a 500-mL Erlenmeyer flask and dry it over anhydrous Na2 SO4 . Filter in a 250-mL round-bottom flask and evaporate the filtrate to dryness on the rotary evaporator. The residue is used for the next step without further purification.
39. Dissolve the residue in 50 mL of anhydrous dioxane. Add 830 mg AIBN and 13 mL TTMSS. 40. Equip the 250-mL round-bottom flask with a reflux condenser and stir bar, and reflux the reaction mixture 4 hr in a preheated 100◦ C oil bath. Monitor reaction by TLC using 60:40 (v/v) toluene/EtOAc. Cool mixture to room temperature and evaporate to dryness on a rotary evaporator. The Rf value for the desired 2 -deoxy derivative is 0.34.
41. Prepare a 6 × 30–cm silica gel chromatography column as described in step 19. Apply the residue (dissolved in a minimum amount of dichloromethane) to the column and elute with a linear gradient of 0% to 2% MeOH/CH2 Cl2 . Collect 250-mL fractions and combine those that contain pure product S.6 as determined by TLC (4% methanol in dichloromethane). 42. Check purity of the product.
Synthesis of β-L-2 Deoxythymidine
14.3.6 Supplement 24
1-[5,3-O-(1,1,3,3-tetraisopropyl-1,3-disiloxanyl)-2 -deoxy-β-L-ribofuranosyl]thymine (S.6): 7.1 g (86% yield from S.5). TLC (96:4 (v/v) dichloromethane/methanol): Rf = 0.46. 1 H NMR (400 MHz, DMSO-d6 ): δ 11.38 (1H, s, NH, D2 O exchangeable), 7.39 (1H, s, H6), 6.0 (1H, m, H1 ), 4.5 (1H, m, H3 ), 4.0-3.9 (2H, m, H5 and H5 ), 3.7 (1H, m, H4 ), 2.5-2.3 (2H, 2m, H2 and H2 ), 1.75 (1H, s, CH3 ), 1.0-0.8 (24H, m, isopropyl). FAB-MS (GT-matrix): 485 [M+H]+ , 261 [TIPSO]+ , 127 [B+2H]+ ; 967 [2M-H]– , 483 [M-H]– , 125 [B]– . Current Protocols in Nucleic Acid Chemistry
Remove 3 ,5 -O-silylation 43. In a 1-L round-bottom flask equipped with a magnetic stir bar, dissolve 6.85 g of S.6 in 300 mL methanol and add 7 g of ammonium fluoride. 44. Place flask in a 80◦ C oil bath and allow reaction mixture to reflux for 3 hr. 45. When the reaction is completed, allow the reaction mixture to cool to room temperature, then add 20 g of silica gel to the flask and evaporate the solvent to dryness in a rotary evaporator equipped with a splash head. 46. Apply the solid on top of a 7 × 15–cm silica gel chromatography column and elute the product with a linear gradient of 0% to 10% MeOH/CH2 Cl2 . Collect 100-mL fractions and combine those that contain pure product as determined by TLC (10% methanol in dichloromethane). 47. Evaporate combined fractions to dryness on the rotary evaporator to obtain 3.17 g of the desired product (90% yield from S.5). Crystallize from absolute ethanol for several days at 0◦ to 5◦ C to produce 2.5 g of white crystals. 48. Analyze the product by TLC, melting point determination, 1 H NMR, MS, UV spectroscopy, and optical rotation determination. 2 -Deoxy-β-L-thymidine (S.7): TLC (90:10 (v/v) dichloromethane/methanol): Rf = 0.37. m.p. = 186◦ -189◦ C. 1 H NMR (400 MHz, DMSO-d6 ): δ 11.26 (1H, s, NH, D2 O exchangeable), 7.83 (1H, s, H6), 6.08 (1H, m, H1 ), 5.14 (1H, d, J = 4.2 Hz, 2 -OH, D2 O exchangeable), 4.94 (1H, d, J = 5.1 Hz, 5 -OH, D2 O exchangeable), 4.1 (1H, m, H3 ), 3.7 (1H, m, H4 ), 3.5 (2H, m, H5 and H5 ), 2.0-1.9 (2H, m, H2 and H2 ), 1.68 (1H, d, J = 0.8 Hz, CH3 ). FAB-MS (GT-matrix): 243 [M+H]+ , 127 [B+2H]+ ; 241 [M-H]– , 125 [B]– . UV (EtOH): λmax = 267.0 nm (ε 10500), λmin = 234.0 nm (ε 1800). [a]D 20 –13.0 (c = 1.0, DMSO). Anal. calcd. for C10 H14 N2 O5 : C, 49.58; H, 5.83; N, 11.57; observed: C, 49.53; H, 5.94; N, 11.50.
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Methanol saturated with ammonia (MeOH/NH3 ) Bubble ammonia gas through a cold 1 L solution of anhydrous methanol (0◦ C, ice bath) until saturation (stabilization of the volume of the solution). The solution must be stored in a stoppered bottle no longer than 6 months at −20◦ C. COMMENTARY Background Information Morbidity and mortality of significant magnitude due to hepatitis B virus (HBV) infection have prompted intensive efforts in searching for potent and selective antiviral agents against this virus. Since the U.S. Food and Drug Administration approved lamivudine for the treatment of HIV infection in 1996 and for HBV in 1998, intensive studies on “unnatural” L-nucleosides as antiviral or anticancer agents have been conducted (Graciet and Schinazi, 1999; Gumina et al., 2002). Through an extensive structureactivity analysis, it appeared that the 3 -OH group of the β-L-2 -deoxyribose of the β-L-2 deoxynucleoside series confers unique speci-
ficity for anti-HBV activity (Bryant et al., 2001; Han, 2005). In this chemical series, L-dT was identified as the most potent, selective, and specific inhibitor of the replication of HBV in cell culture experiments (Bryant et al., 2001) without any cellular or mitochondrial toxicity. If the structure of L-dT is considered, this compound exhibits no chemical modifications and differs from its natural nucleoside only with respect to the stoichiometric relationship of its sugar and base moieties, the L-configuration versus the D-configuration. The selectivity of L-dT is a critical issue since long-term treatment is expected for chronic HBV infection. Thus, L-dT appeared
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14.3.7 Current Protocols in Nucleic Acid Chemistry
Supplement 24
as an investigational new drug for treatment of chronic HBV infection. To conduct preclinical in vivo antiviral studies in woodchucks and pharmacokinetic and toxicology studies in rodents and monkeys, semi-large-scale synthesis of L-dT was required. L-dT was prepared for the first time by Smejkal and Sorm (1964) by condensation of thyminyl mercury with the crystalline 3,5-diO-p-toluyl-2-deoxy-L-ribofuranosyl chloride (obtained in three steps from 2-deoxy-Lribose) in toluene, followed by deacylation of the resulting protected nucleoside and crystallization. A few years later, Hol´y developed another synthetic method for the preparation of L-dT. This alternative route consists of the preparation of 2 -deoxy-L-uridine starting from L-arabinose and its conversion into the thymidine counterpart by introduction of a methyl group into position 5 of the uracil ring (Hol´y, 1972). This key step was accomplished by hydroxyalkylation with formaldehyde and hydrogenolysis of the resulting 5-hydroxymethyl derivative. Both of the above procedures do not appear suitable for the preparation of L-dT on a larger scale as they are time consuming and involve tedious purification steps or difficult handling reactions. This led to the developement of the method presented here: a six-step sequence involving direct condensation of thymine with 1-O-acetyl2,3,5-tri-O-benzoyl-L-ribofuranose, followed by successive deacylation and selective 3 ,5 O-silylation, 2 -deoxygenation, and removal of the protecting groups. This approach can be efficiently used for the general synthesis of β-L-2 -deoxynucleosides incorporating different heterocyclic bases. Using this method, L-dT has been synthesized on a semi-large scale and its potency and safety have been evaluated in vivo using the woodchuck model of chronic HBV infection (Bryant et al., 2001; Han, 2005). The promising results of early in vitro and animal studies paved the way for Phase I/II human clinical trials. These studies demonstrated an excellent safety profile with favorable pharmacokinetics. On the basis of this promising data, a large, multi-center, international Phase III study of L-dT (telbivudine) was initiated and is ongoing (Han, 2005).
Critical Parameters Synthesis of β-L-2 Deoxythymidine
The most critical parameter in each reaction is purity of the reactants and reagents. Some reagents, like BzCl, PhOC(S)Cl, TIPSCl2 , and
TMSOTf, are sensitive to moisture and will deteriorate over time. As a consequence, the best results are obtained with freshly opened bottles of reagents. Alternatively, such reagents should be stored under a dry argon atmosphere.
Anticipated Results If the procedures are followed as described in this unit, the yields of isolated product should be comparable to those reported in the protocol steps. Silylation of the base can be carried out using N,Obis(trimethysilyl)acetamide (BSA) instead of hexamethyldisilazane (HMDS). To the same extent, the reagent tris(trimethylsilyl)silane (TTMSS) can be replaced by tri-n-butyltin hydride (tBu3 SnH) during the reductive deoxygenation step.
Time Considerations The time required for the synthesis of L-dT starting with L-ribose is 2 weeks. This estimated time does not include the time necessary to purify or dry solvents. All commercially available solvents were used without further purification and purchased as anhydrous solvent, when needed.
Literature Cited Bryant, M.L., Bridges, E.G., Placidi, L., Faraj, A., Loi, A.-G., Pierra, C., Dukhan, D., Gosselin, G., Imbach, J.-L., Hernandez, B., Juodawlkis, A., Tennant, B., Korba, B., Cote, P., Marion, P., Cretton-Scott, E., Schinazi, R., and Sommadossi, J.-P. 2001. Antiviral L-nucleosides specific for Hepatitis B Virus infection. Antimicrob. Agents Chemother. 45:229-235. Cimpoa, A.R., Hunter, P.J., and Evans, C.A. 1994. On the conversion of the arabino- and ribofuranosyl methyl glycosides to their 1-O-acetyl derivatives. J. Carbohydr. Chem. 13:1115-1119. Gosselin, G., Pierra, C., Benzaria, S., Dukhan, D., Imbach, J.L., Loi, A.-G., La Colla, P., Cretton-Scott, E., Bridges, E.G., Standring, D.N., and Sommadossi, J.P. 2004. β-L-2 -Deoxythymidine (L-dT) and β-L-2 deoxycytidine (L-dC): How simple structures can be potent, selective and specific anti-HBV drugs. In Frontiers in Nucleosides and Nucleic Acids (R.F. Schinazi and D.C. Liotta, eds.), pp. 309-317. IHL Press, Tucker, Georgia. Graciet, J.C.G. and Schinazi, R.F. 1999. From Dto L-nucleoside analogs as antiviral agents. Adv. Antivir. Drug Design 3:1-68. Gumina, G., Chong, Y., Choo, H., Song, G.-Y., and Chu, C.K. 2002. L-Nucleosides: Antiviral activity and molecular mechanism. Curr. Top. Med. Chem. 2:1065-1086. Han, S.-T.B. 2005. Telbivudine: A new nucleoside analogue for the treatment of chronic hepatitis B. Expert Opin. Investig. Drugs 14:511-519.
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Current Protocols in Nucleic Acid Chemistry
Hol´y, A. 1972. Nucleic acid components and their analogues. CLIII. Preparation of 2 -deoxy-Lribonucleosides of the pyrimidine series. Collect. Czech. Chem. Commun. 37:4072-4087. Recondo, E.F. and Rinderknecht, H. 1959. Eine neue, einfache synthese des 1-O-acetyl-2,3,5tri-O-β-D-ribofuranosides. Helv. Chim. Acta 42:1171-1173. Smejkal, J. and Sorm, F. 1964. Nucleic acids components and their analogues. LIII. Preparation of 1-2 -deoxy-β-L-ribofuranosylthymine, “L-thymine”. Collect. Czech. Chem. Commun. 29:2809-2813.
Contributed by Claire Pierra, David Dukhan, and Gilles Gosselin Universit´e Montpellier II Montpellier, France Jean-Pierre Sommadossi Idenix Pharmaceuticals, Inc. Cambridge, Massachusetts
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14.3.9 Current Protocols in Nucleic Acid Chemistry
Supplement 24
Synthesis of Carbovir and Abacavir from a Carbocyclic Precursor
UNIT 14.4
A facile method for the production of carbocyclic nucleosides from the versatile lactam 2-azabicyclo[2.2.1]hept-5-en-3-one was originated in the authors’ laboratory (Vince and Daluge, 1978). The lactam, subsequently coined the Vince lactam by Chemical and Engineering News (Stinson, 1994; Rouhi, 2003), is now commercially available either as a racemic mixture or as enantiomers. Following the identification of a retrovirus (human immunodeficiency virus or HIV) as the etiological agent of human acquired immunodeficiency syndrome (AIDS), an intense effort was made to identify drugs for the treatment of this disease. The lactam method was employed in the synthesis of a new class of anti-HIV nucleosides. These carbocyclic 2 ,3 -didehydro-2 ,3 -dideoxy-2-amino6-substituted purines were termed carbovirs (Vince et al., 1988; Vince and Hua, 1990). The parent compound, carbocyclic 2 ,3 -didehydro-2 ,3 -dideoxyguanosine (carbovir), is enzymatically converted to the active form, carbovir triphosphate, which inhibits HIV reverse transcriptase. All other active derivatives in this series are ultimately converted to the same active metabolite. The anti-HIV drug derived from this series, abacavir (Ziagen), obtains its activity via conversion to carbovir triphosphate. The preparation of the key intermediate, cis-[4-(2-amino-6-chloro-9H-purin-9-yl)-2-cyclopentenyl]carbinol, is described for the preparation of carbovir and abacavir.
PREPARATION OF CARBOVIR AND ITS 6-CHLOROPURINE PRECURSOR The synthetic procedure for the synthesis of carbovir and abacavir via the 6-chloropurine intermediate is outlined in Figure 14.4.1. The protocol described here includes the entire procedure from the starting Vince lactam to carbovir, and is based on the method using the racemic lactam. The individual enantiomers can be used in the same reactions to obtain the optical isomers. The starting lactam can be made by the procedure described by Vince and Daluge (1978) or can be obtained commercially from a supplier such as Sigma-Aldrich. The final step from the 6-chloropurine precursor (S.6) to carbovir (S.7) is presented as two different methods that depend on the desired synthesis scale. Synthesis of abacavir (S.8) from S.6 is described in Basic Protocol 2.
BASIC PROTOCOL 1
Materials 2-Azabicyclo[2.2.1]hept-5-en-3-one (Vince lactam; Sigma-Aldrich) 12 N HCl 1 N dry HCl/methanol Pyridine Acetic anhydride Ethyl acetate Anhydrous calcium chloride Sodium borohydride Tetrahydrofuran (THF) Barium hydroxide Dry ice Absolute ethanol 1-Butanol 2-Amino-4,6-dichloropyrimidine (Aldrich) Triethylamine Silica gel 60 (230 to 400 mesh; EM Science) Chloroform (CHCl3 ) Contributed by Robert Vince and Mei Hua Current Protocols in Nucleic Acid Chemistry (2006) 14.4.1-14.4.8 C 2006 by John Wiley & Sons, Inc. Copyright
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14.4.1 Supplement 25
Methanol (MeOH) p-Chloroaniline Ice-salt bath Sodium nitrite Acetic acid Sodium acetate trihydrate Acetone Zinc dust Nitrogen source Diethyl ether Triethyl orthoformate 3 N NaOH Reflux condenser Mechanical stirrer Rotary evaporator B¨uchner filter funnel Silicon oil bath 5.0 × 15–, 2.0 × 18–, 4.0 × 10–, 2.0 × 7.5–, and 5 × 5–cm columns Silica gel 60 F254 TLC plates (0.25-mm layer; EM Science) Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) Prepare cis-acetamidocyclopent-2-enemethyl acetate (S.1) 1. Dissolve 50 g Vince lactam in 750 mL of 1 N HCl and reflux for 1 hr (120◦ C). Concentrate solution to dryness on a rotary evaporator to obtain the carboxylic acid–amine salt (66 g; 88% yield). 2. Dissolve the carboxylic acid–amine salt in 750 mL of 1 N dry HCl/methanol and reflux for 1 hr (75◦ C). Remove the volatile material on a rotary evaporator to obtain the methyl ester–amine salt as a syrup. To prepare 1 N dry HCl/methanol, pass dry HCl gas into MeOH, under cooling, then adjust to a 1 N concentration.
3. Dissolve the syrup in 250 mL pyridine and 200 mL acetic anhydride, and stir overnight at room temperature. 4. Concentrate the solution on a rotary evaporator and crystallize from 200 mL ethyl acetate to yield the ester-acetamide compound (70 g; 83% yield; m.p. 64◦ -66◦ C). 5. Prepare the reducing agent by mixing 47.7 g ground calcium chloride and 32.55 g sodium borohydride in 900 mL THF, and stir 1 hr at room temperature. 6. Add 52.5 g of the ester-acetamide to 450 mL THF, and then add to the reducing agent. Stir 18 hr at ambient temperature. 7. Carefully and gradually add 550 mL ice-cold water to hydrolyze the complex. First add 3-mL portions, then gradually increase the portion size added. 8. Acidify the cold mixture to pH 1.5 by adding 6 N HCl. 9. Remove the solvent on a rotary evaporator, dissolve the residue in 370 mL pyridine and 370 mL acetic anhydride, and stir overnight at ambient temperature. Synthesis of Carbovir and Abacavir from a Carbocyclic Precursor
10. Concentrate the solution to dryness on a rotary evaporator and collect the pure product (S.1) by crystallization from 250 mL ethyl acetate overnight. On the next day, filter the product. cis-Acetamidocyclopent-2-enemethyl acetate (S.1): 41 g (73%). m.p. 65◦ C.
14.4.2 Supplement 25
Current Protocols in Nucleic Acid Chemistry
Figure 14.4.1 Preparation of abacavir and carbovir from a common 6-chloropurine precursor that is derived from Vince lactam.
Remove acetyl groups 11. Prepare a 0.5 N solution of barium hydroxide in 400 mL water. 12. Add 10.0 g (50 mmol ) S.1 to the barium hydroxide solution and reflux the mixture overnight (110◦ C). 13. Cool the mixture to room temperature. Neutralize the solution to pH 7 by adding dry ice with stirring. 14. Remove the precipitate through a B¨uchner filter funnel. 15. Concentrate the filtrate to dryness on a rotary evaporator. 16. Extract the residue three times with 150 mL absolute ethanol.
Biologically Active Nucleosides
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17. Concentrate the ethanol extract on a rotary evaporator to a colorless syrup of crude S.2. (±)-cis-(4-Aminocyclopent-2-enyl)methanol (S.2) is used directly for the next step without purification.
Perform pyrimidine heterocycle substitution 18. Dissolve S.2 in 200 mL of 1-butanol. 19. Add 12.3 g (75 mmol) 2-amino-4,6-dichloropyrimidine and 30 mL triethylamine to the 1-butanol solution and reflux the mixture on a silicon oil bath for 48 hr (130◦ C). 20. Remove the volatile materials on a rotary evaporator, <40◦ C, to yield a syrup. 21. Adsorb the syrup onto 30 g silica gel. Pack the silica gel sample onto a 5.0 × 15–cm column (APPENDIX 3E). 22. Elute the column three times with 300 mL each of 40:1, 30:1, and 20:1 (v/v) CHCl3 /MeOH, and check each fraction by TLC (APPENDIX 3D) using 10:1 (v/v) CHCl3 /MeOH (Rf = 0.31). The desired fractions are eluted with the 20:1 CHCl3 /MeOH fractions.
23. Collect product fractions and concentrate them on a rotary evaporator to obtain a thick syrup. 24. Crystallize the syrup from 150 mL ethyl acetate to yield white crystals. (±)-cis-[4-(2-Amino-6-chloropyrimidin-4-ylamino)cyclopent-2-enyl]methanol (S.3): 9.15 g (76%). m.p. 132◦ −134◦ C.
Derivatize 5 position of pyrimidine ring 25. Dissolve 1.47 g (11.5 mmol) p-chloroaniline in 25 mL of 3 N HCl and cool on an ice-salt bath. 26. Dissolve 870 mg (12.5 mmol) sodium nitrite in 10 mL water and cool on an ice-salt bath. 27. Add the cold sodium nitrite solution to the cold p-chloroaniline solution to form a cold diazonium salt solution. 28. Dissolve 2.40 g (10 mmol) S.3 into 50 mL water, 50 mL acetic acid, and 20 g sodium acetate trihydrate. 29. Add the cold diazonium salt solution to the S.3 solution and stir the mixture overnight at ambient temperature. 30. Remove the yellow precipitate by filtering with a B¨uchner funnel under vacuum, and wash the precipitate with 200 mL cold water to pH 7 (check with pH paper). 31. Air dry the solid in a well-ventilated chemical fume hood. 32. Collect the dried product (S.4). Crude (S.4): 3.60 g (94%). m.p. 229◦ C (dec).
33. Purify the crude product from 500 mL of 1:2 (v/v) acetone/methanol to yield a pure sample. Synthesis of Carbovir and Abacavir from a Carbocyclic Precursor
(±)-cis-{4-[2-Amino-6-chloro-5-(4-chloro-phenylazo)pyridin-4-ylamino]cyclopent-2enyl}methanol (S.4): m.p. 241◦ -243◦ C (dec). MS: m/e 378 and 380 (M+ and M+ +2). IR (KBr) (cm−1 ): 3600-3000, 1620, 1580. Anal. calcd. (C16 H16 Cl2 N8 O): C, 50.67; H, 4.25; N, 22.16; found: C, 50.59; H, 4.20; N, 21.99.
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Perform reductive cleavage of azo function 34. Mix 379 mg (1.0 mmol) S.4 with 0.65 g (10 mmol) zinc dust, 0.32 mL acetic acid, 15 mL water, and 15 mL ethanol. Reflux the mixture under nitrogen for 3 hr (130◦ C). This reaction can be scaled up five to ten times. For example, use 1.89 g (5.0 mmol) S.4 with 3.25 g (50 mmol) zinc dust, 1.6 mL acetic acid, 75 mL water, and 75 mL ethanol. Use 10 g silica gel with a 4 × 14–cm column (step 37), 200 mL of 1:1 methanol/diethyl ether to yield pink crystal product (step 40), and the resulting yield of S.5 is 0.85 g.
35. Remove the zinc dust on a B¨uchner funnel. 36. Evaporate filtrate to a thick syrup on a rotary evaporator with a water aspirator. 37. Adsorb the syrup onto 2 g silica gel. Pack the sample onto a 2.0 × 18–cm column. 38. Elute the column with 15:1 (v/v) CHCl3 /MeOH and check each fraction by TLC using 10:1 (v/v) CHCl3 /MeOH (Rf = 0.35). 39. Collect product fractions and concentrate to a pink syrup on a rotary evaporator with a water aspirator. 40. Purify the crude product from 40 mL of 1:1 (v/v) methanol/diethyl ether to yield a pink crystal product. (±)-cis-[4-(2,5-Diamino-6-chloro-pyrimidin-4-ylamino)cyclopent-2-enyl]methanol (S.5): 170 mg (66%). m.p. 168◦ -170◦ C. MS: m/e 255 and 257 (M+ and M+ +2). IR (KBr) (cm−1 ): 3600-3000, 1620, 1580. Anal. calcd. (C10 H14 ClN5 O): C, 46.97; H, 5.52; N, 27.39; found: C, 47.10; H, 5.56; N, 27.36.
Form purine heterocycle by ring closure 41. Mix 1.41 g (5.5 mmol) S.5, 30 mL triethyl orthoformate, and 1.40 mL of 12 N HCl. Stir the mixture overnight at ambient temperature. 42. Concentrate the mixture to dryness on a rotary evaporator. 43. Add 40 mL of 0.5 N HCl to the residue and stir for 1 hr at room temperature. 44. Neutralize the mixture to pH 8 by adding 1 N NaOH. 45. Concentrate the mixture on a rotary evaporator. 46. Adsorb the residue on 7.5 g silica gel. Pack the sample onto a 4.0 × 10–cm column. 47. Elute the column with 20:1 (v/v) CHCl3 /MeOH and check each fraction by TLC using 5:1 (v/v) CHCl3 /MeOH (Rf = 0.64). 48. Collect product fractions and concentrate to a white solid (S.6). Yield: 1.18 g (80%).
49. Recrystallize the crude product from 25 mL ethanol to yield the pure 6-chloropurine precursor. (±)-cis-[4-(2-Amino-6-chloropurin-9-yl)cyclopent-2-enyl]methanol (S.6): m.p. 145◦ 147◦ C. MS: m/e 265 and 267 (M+ and M+ +2). IR (KBr) (cm−1 ): 3600-2600, 1620, 1580. 1 H NMR (DMSO-d6 ): 8.01 (s, 1H, H8), 6.98-6.80 (s, 2H, NH2 ), 6.15-6.05 and 5.94-5.84 (dd, 2H, CH=CH, J = 5.0 Hz), 5.50-5.32 (m, 1H, H1), 4.79-4.60 (t, 1H, CH2 OH), 3.51-3.39 (d, 2H, CH2 OH), 2.95-2.71 (m, 1H, H4 ), 2.69-2.55 (m, 1H, CHH ), 1.72-1.50 (m, 1H, CHH ). Anal. calcd. (C11 H12 N5 OCl· 34 H2 O): C, 47.31; H, 4.87; N, 25.09; found: C, 47.40; H, 4.94; N, 25.21. Biologically Active Nucleosides
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Supplement 25
Prepare carbovir For a 100-mg scale 50a. Mix 266 mg (1.0 mmol) S.6 and 20 mL of 0.33 N NaOH, and reflux the mixture for 5 hr (120◦ C). 51a. Concentrate the solution on a rotary evaporator. 52a. Adsorb the residue onto 2 g silica gel. Pack the sample onto a 2.0 × 7.5–cm column. 53a. Elute the column with 5:1 (v/v) CHCl3 /MeOH and check each fraction by TLC using 5:1 (v/v) CHCl3 /MeOH (Rf = 0.27). 54a. Collect product fractions and concentrate to a white solid on a rotary evaporator with a water aspirator. 55a. Recrystallize the crude product from 20 mL of 1:4 (v/v) methanol/water to yield a white crystal product. (±)-cis-2-Amino-9-(4-hydroxymethylcyclopent-2-enyl)-9H-purin-6-ol (carbovir; S.7): 152 mg (61%). m.p. 254◦ -256◦ C (dec). MS: m/e 247(M+ ). IR (KBr) (cm−1 ): 3600-2600, 1600. 1 H NMR (DMSO-d6 ): 10.57-10.50 (s, 1H, 6-OH), 7.60-7.56 (s, 1H, H8), 6.50-6.35 (s, 2H, NH2 ), 6.14-6.06 and 5.89-5.81 (dd, 2H, CH=CH, J = 5.0 Hz), 5.38-5.26 (m, 1H, H1 ), 4.76-4.65 (t, 1H, CH2 OH), 3.47-3.39 (d, 2H, CH2 OH), 2.92-2.80 (m, 1H, H4 ), 2.65-2.55 (m, 1H, CHH ), 1.64-1.50 (m, 1H, CHH ). Anal. calcd. (C11 H13 N5 O2 · 43 H2 O): C, 50.66; H, 5.60; N, 26.86; found: C, 50.44; H, 5.59; N, 26.76.
For a 1- to 10-g scale 50b. Dissolve 4.3 g (16 mmol) S.6 into 220 mL of 1 N HCl and 60 mL methanol. Heat solution 9 hr on a 60◦ C silicon oil bath. When scaled up to 5 g, some polymer was obtained (sticky in appearance). Therefore, acidic conditions were used to avoid polymerization.
51b. Concentrate the solution on a rotary evaporator to 50 mL. 52b. Neutralize the solution to pH 7 by adding 3 N NaOH. Evaporate the solution. 53b. Adsorb the residue onto 10 g silica gel. Pack the sample onto a 5 × 5–cm column. 54b. Elute with 5:1 (v/v) CHCl3 /MeOH and check each fraction by TLC using 5:1 (v/v) CHCl3 /MeOH (Rf = 0.27). 55b. Collect product fractions to yield crude product (S.7). Yield: 3.1 g (77%).
56b. Recrystallize the crude product from 35 mL 1:4 (v/v) methanol/water to yield the pure compound. Carbovir (S.7): 2.4 g (60%). m.p. 254◦ -256◦ C (dec).
BASIC PROTOCOL 2
Synthesis of Carbovir and Abacavir from a Carbocyclic Precursor
PREPARATION OF ABACAVIR FROM THE 6-CHLOROPURINE PRECURSOR The synthesis of abacavir (S.8) from the 6-chloropurine intermediate (S.6) is illustrated in Figure 14.4.1. The scale presented is comparable to the smaller scale method in Basic Protocol 1. This reaction could be scaled up without any modifications except for the amounts of ethanol and cyclopropylamine. For example, a 10-fold scale up would require a 2.5-fold increase in the amounts of ethanol and cyclopropylene.
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Current Protocols in Nucleic Acid Chemistry
Materials 6-Chloropurine precursor (S.6; see Basic Protocol 1) Ethanol Cyclopropylamine 1 N NaOH Silica gel (230 to 400 mesh; EM Science) Chloroform (CHCl3 ) Methanol (MeOH) Acetonitrile Silicon oil bath Rotary evaporator 2 × 13–cm column Silica gel 60 F254 TLC plate (0.25-mm layer; EM Science) Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) 1. Mix 0.26 g (1.0 mmol) S.6, 5 mL ethanol, and 20 g cyclopropylamine. Reflux the mixture on a silicon oil bath for 25 hr (55◦ C). 2. Add 1 mL (1 mmol) of 1 N NaOH to the mixture. Concentrate the mixture on a rotary evaporator. 3. Adsorb onto 2 g silica gel. Pack the sample on a 2 × 13–cm column (APPENDIX 3E). 4. Elute the column with 50:1 (v/v) CHCl3 /MeOH and check each fraction by TLC (APPENDIX 3D) using 10:1 (v/v) CHCl3 /MeOH (Rf = 0.24). 5. Collect product fractions and concentrate to a yellow residue on a rotary evaporator. 6. Recrystallize the crude product from 12 mL acetonitrile to yield the white crystal product. (±)-cis-[4-(2-Amino-6-cyclopropylaminopurin-9-yl)cyclopent-2-enyl]methanol (abacavir; S.8): 0.19 g (68%). m.p. 74◦ C soften, 154◦ C melt. MS: m/e 287 (m+1)+ . IR (KBr) (cm−1 ): 3317, 3195, 1594, 1483, 1451. 1 H NMR (DMSO-d6 ): 7.60 (s, 1H, H8), 7.22 (s, 1H, NH), 6.10 and 5.92 (dd, 2H, CH=CH), 5.78 (s, 2H, NH2 ), 5.38 (s, 1H, H1 ), 4.00 (t, 1H, CH2 OH), 3.44 (d, 2H, CH2 OH), 3.00 (m, 1H, NHCH), 2.80 (m, 1H, H4 ), 2.60 (m, 1H, H5 ), 1.55 (m, 1H, H5 ), 1.55 (m, 4H, CH2 CH2 ). Anal. (C14 H18 N6 O) C,H,N.
COMMENTARY Background Information A major disadvantage of many clinically used nucleoside drugs is their ability to undergo metabolism or degradation to inactive forms. For example, most nucleosides are efficient substrates for nucleoside phosphorylases, which cleave the nucleosides at their N-glycosyl linkage and release the free base forms. The acid-labile glycosidic linkages are also easily cleaved by acid in the stomach. To circumvent these problems, carbocyclic nucleosides were designed in which the furanose ring oxygen of the sugar moiety was replaced by carbon. As predicted, these non-glycosidic nucleosides were resistant to both enzymatic and acid hydrolysis. As a result, many of the
carbocyclic nucleosides exhibit interesting biological properties and can be used in the areas of antitumor and antiviral chemotherapy. Most of the synthetic schemes to carbocyclic nucleosides utilize a cyclopentane or cyclopentene to provide the carbocyclic sugar. These starting materials are inexpensive and provide the advantage of having an intact five-membered ring. Several of these approaches have been described in recent reviews (Zhu, 2000; Jeong and Lee, 2004). Other approaches utilizing aristeromycin, ring closing metathesis, and carbohydrate precursors are also discussed with their advantages and disadvantages. For example, the natural products, aristeromycin and carbohydrates, yield
Biologically Active Nucleosides
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Supplement 25
chiral products, while the synthetic schemes are usually long. The use of cyclopentene precursors have the advantage of short synthetic schemes, but require a resolution step or the use of a chiral catalyst to obtain the pure enantiomers. The main advantage of using the Vince lactam is the relative positioning of the double bond and the two substituents on the cyclopentene ring. Opening of the lactam followed by reduction conveniently provides the amine group for purine or pyrimidine construction cis to the hydroxymethyl group. It also places the double bond at the 2,3 position, which is essential for anti-HIV activity. Other advantages include the commercial availability of both enantiomers from Sigma-Aldrich and the ease of conversion to the desired nucleosides.
Compound Characterization NMR spectra were recorded on a 300-MHz GE spectrometer. Chemical shifts (δ) are given in part per million (ppm) and referenced to tetramethylsilane as an internal standard. Coupling constants (J) are given in Hertz and refer to apparent multiplicities. Elemental analyses were performed by M-H-W Laboratories, Phoenix, AZ. Melting points were determined on a Mel-Temp 2 apparatus and are uncorrected. Electron impact mass spectra (EIMS) were obtained with a Kratos/AEI MS-30, and chemical impact (CI) MS were obtained with a Finnigan 4000. Thin-layer chromatography (TLC) was performed on EM Science silica gel 60 F254 (0.25-mm layer), and column chromatography was performed on EM Science silica gel 60 (230 to 400 mesh).
Critical Parameters and Troubleshooting Characterization of the products requires knowledge of 1 H NMR, UV, and MS. Prior experience with organic chemistry laboratory techniques such as recrystallization, solvent evaporation, extraction methods, TLC, HPLC, and vacuum filtration is required. All intermediates should be stored at 4◦ C and protected from moisture. Strict adherence to all out-
lined procedures is highly recommended for safety reasons and for optimum yields of pure products.
Anticipated Results These procedures are suitable for preparation of milligram to multi-gram amounts. The final products are very stable and can be stored for several years at room temperature under anhydrous conditions.
Time Considerations The synthesis of the final carbocyclic 6-substitued-2-aminopurine nucleosides from the Vince lactam starting material can be accomplished in 2 to 3 weeks.
Literature Cited Jeong, L.S. and Lee, J.A. 2004. Recent advances in the synthesis of the carbocyclic nucleosides as potential antiviral agents. Antiviral Chem. Chemother. 15:235-250. Rouhi, A.M. 2003. Cover Story. Chem. Eng. News 81:37-52. Stinson, S.C. 1994. Market, environmental pressures spur change in fine chemicals industry. Chem. Eng. News 72:12-26. Vince, R. and Daluge, S. 1978. Synthesis of carbocyclic aminonucleosides. J. Org. Chem. 43:2312-2320. Vince, R. and Hua, M. 1990. Synthesis and antiHIV activity of carbocyclic 2 ,3 -didehydro2 ,3 -dideoxy-2,6-disubstituted purine nucleosides. J. Med. Chem. 33:17-21. Vince, R., Hua, M., Brownell, J., Daluge, S., Lee, F.C., Shannon, W.M., Lavelle, G.C., Qualls, J., Weislow, O.S., Kiser, R., Canonico, C.G., Schultz, R.H., Narayanan, V.L., Mayo, L.G., Shoemaker, R.H., and Boyd, M.R. 1988. Potent and selective activity of a carbocyclic nucleoside analog (Carbovir: NSC 614846) against human immunodeficiency virus in vitro. Biochem. Biophys. Res. Commun. 156:1046-1053. Zhu, Z.-F. 2000. The latest progress in the synthesis of carbocyclic nucleosides. Nucleosides Nucleotides Nucleic Acids 19:651-690.
Contributed by Robert Vince and Mei Hua Center for Drug Design University of Minnesota Minneapolis, Minnesota
Synthesis of Carbovir and Abacavir from a Carbocyclic Precursor
14.4.8 Supplement 25
Current Protocols in Nucleic Acid Chemistry
Synthesis of 2 - and 3 -C-Methylribonucleosides
UNIT 14.5
Basic Protocols 1 and 2 describe the synthesis of the protected starting carbohydrates, 3-C-methyl-D-ribofuranose and 2-C-methyl-D-ribofuranose, respectively. Basic Protocols 3 and 4 describe full details of their use for the preparation of 3 - and 2 -Cmethylribonucleosides, respectively, using the uracil derivatives as examples. Modifications for preparation of the cytosine, adenine, and guanine derivatives are given in Alternate Protocols 1 and 2. CAUTION: Carry out all operations involving organic solvents and reagents in a wellventilated fume hood, and wear gloves and protective glasses.
PREPARATION OF THE PROTECTED 3-C-METHYL-D-RIBOFURANOSE The sequence of reactions outlined here (Fig. 14.5.1) illustrates a general approach for conversion of branched allofuranose S.1 into the fully protected ribofuranose S.6, which is suitable for subsequent incorporation of a nucleic acid base via a glycosylation reaction (Vorbr¨uggen and Ruh-Pohlenz, 2001). This five-step procedure can be applied for allofuranoses with different alkyl groups at C3 (Rosenthal and Mikhailov, 1980; Mikhailov et al., 1983). Compound S.1 may be easily prepared starting with commercially available 1,2:5,6-di-O-isopropylidene-α-D-glucofuranose by subsequent oxidation and Grignard reaction with methylmagnesium iodide. Several reliable procedures of this two-step conversion may be found in the literature (Bio et al., 2004).
BASIC PROTOCOL 1
Selective removal of the 5,6-O-isopropylidene group in S.1 is achieved by treatment with 75% AcOH, and the resulting triol S.2 is isolated cleanly by crystallization with a 90% yield. Periodate oxidation of the 5,6-diol group followed by sodium borohydride reduction
Figure 14.5.1 Synthesis of 1,2,3-tri-O-acetyl-5-O-benzoyl-3-C-methyl-α,β-D-ribofuranose (S.6) from S.1 in a 56% expected overall yield. AcOH, acetic acid; Ac2 O, acetic anhydride; BzCl, benzoyl chloride; DMAP, 4dimethylaminopyridine. Biologically Active Nucleosides Contributed by Leonid Beigelman and Sergey N. Mikhailov Current Protocols in Nucleic Acid Chemistry (2007) 14.5.1-14.5.26 C 2007 by John Wiley & Sons, Inc. Copyright
14.5.1 Supplement 28
provides crystalline ribofuranose S.3 in a 89% isolated yield. Selective benzoylation of the 5-OH in furanose S.3 is necessary to prevent undesirable formation of the pyranose form of 3-C-methyl-D-ribose, which could form during removal of the 1,2-isopropylidene group (from S.4 to S.5). Despite a significant difference in the reactivity of the primary and tertiary OH groups in furanose S.3, it is advisable to perform benzoylation at 0◦ C with a slow addition of BzCl to achieve high yields (85% to 90%) for crystalline monobenzoate S.4. The next two steps (removal of the 1,2-isopropylidene group from S.4 and subsequent full acylation of S.5) should be performed as one sequence, without isolating the intermediate triol S.5. It is important to remove all traces of trifluoroacetic acid via repeated coevaporation with toluene. Subsequent acylation is performed in the presence of the catalyst DMAP to achieve acetylation of the tertiary hydroxyl at C3. The product, l,2,3-tri-O-acetyl-5-O-benzoyl-3-C-methyl-α,β-D-ribofuranose (S.6), is usually obtained as a mixture of anomers (α,β) and requires column purification. The individual β-anomer can be isolated in crystalline form (∼25% yield), but since both α and β anomers can be utilized in the synthesis of 3 -C-methylribonucleosides, it is advisable to pull appropriate fractions to give a combined yield of 83% (Beigelman et al., 1988).
Materials 1,2:5,6-Di-O-isopropylidene-3-C-methyl-α-D-allofuranose (S.1; Bio et al., 2004) Acetic acid (AcOH) 1-Butanol, reagent grade Ethanol (EtOH), reagent grade Phosphorus pentoxide (P2 O5 ) Sodium periodate (NaIO4 ) Sodium borohydride (NaBH4 ) Chloroform (CHCl3 ), reagent grade Methanol (MeOH), analytical grade Hexane, reagent grade Anhydrous pyridine Benzoyl chloride Saturated sodium bicarbonate (NaHCO3 ) Anhydrous sodium sulfate (Na2 SO4 ) Toluene, reagent grade 90% trifluoroacetic acid (F3 CCOOH) Acetic anhydride, reagent grade 4-Dimethylaminopyridine (DMAP) Silica gel: Kieselgel 60 (0.06 to 0.20 mm; Merck) 250- and 500-mL round-bottom flasks Rotary evaporator equipped with a water aspirator and an oil pump Reflux condenser Vacuum filtration system with glass filters (porosity 3) Vacuum desiccator 250-mL and 1-L separatory funnels Dropping funnel connected to a CaCl2 protection tube Silica-coated aluminum TLC plates with fluorescent indicator (Merck silicagel 60 F254 ) 254-nm UV lamp 3 × 20–cm sintered glass chromatography column, porosity 3 Vacuum oil pump Synthesis of 2 - and 3 -C-Methylribonucleosides
Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E)
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Current Protocols in Nucleic Acid Chemistry
Prepare S.2 1. Weigh 12.6 g (46 mmol) S.1 into a 250-mL round-bottom flask. Add 100 mL of 75% acetic acid and keep the solution for 24 hr at room temperature. 2. Evaporate all volatile material to dryness in vacuo using a rotary evaporator. Add to the residue 50 mL of 1-butanol and again evaporate to dryness. Repeat the evaporation two times with 50 mL of 1-butanol. 3. Add 40 mL ethanol, attach a condenser, and reflux (78◦ C) until complete dissolution of the solid. Cool the flask to room temperature and keep the mixture 16 hr at 0◦ C. 4. Collect the precipitate (S.2) by vacuum filtration on a glass filter (porosity 3), wash the precipitate with 10 mL ethanol, and dry over P5 O5 in a vacuum desiccator overnight. 5. Characterize the compound by NMR. The compound is stable stored for at least 12 months at room temperature. 1,2-O-Isopropylidene-3-C-methyl-α-D-allofuranose (S.2). Yield of white crystals 9.6 g (89%). m.p. 133◦ -134◦ C (EtOH). [α]20 D +23.0◦ (c 1, chloroform), [α]20 D +40.1◦ (c 1, methanol). 1 H NMR (100 MHz, CDCl3 ): 5.71 d (1H, J1 ,2 = 4.0 Hz, H1), 4.16 d (1H, H2), 3.85-3.55 m (4H, H4, 5, 6a, 6b), 1.60 s (3H, Me), 1.37 s (3H, Me), 1.35 s (3H, C-Me-3). 13 C NMR (CDC13 ): 113.1 (Me2 C), 103.7 (C1), 85.3 (C2), 79.1 (C4), 78.1 (C3), 70.9 (C5), 64.9 (C6), 26.8, 26.6 (Me2 C), 19.4 (C-Me−3).
Prepare S.3 6. Weigh 8.3 g (38.8 mmol) S.2 into a 500-mL round-bottom flask containing a stir bar and dissolve in 70 mL water. 7. Add 8.3 g (38.8 mmol) NaIO4 in one portion and stir the reaction mixture 1 hr at room temperature. 8. Add 300 mL ethanol and filter NaIO3 by vacuum filtration on a glass filter (porosity 3). Wash the precipitate with 30 mL ethanol. 9. Add to the combined filtrates 3 g (80 mmol) NaBH4 in 0.5-g portions over 1 hr and keep the reaction mixture for 16 hr at room temperature. 10. Neutralize the reaction mixture to pH 7.0 with acetic acid. Add 100 mL water and 300 mL chloroform. Separate the organic layer using a 1-L separatory funnel and wash the aqueous layer with 300 mL chloroform. 11. Evaporate the combined organic layers in a 500-mL round-bottom flask to dryness in vacuo using a rotary evaporator connected to a vacuum system. 12. Add 20 mL methanol to the residue and evaporate all volatile material to dryness in vacuo using a rotary evaporator. Repeat coevaporation with 20 mL methanol. 13. Dissolve the residue in a minimal volume (∼20 mL) of chloroform at room temperature and add hexane (∼50 to 70 mL) until faintly turbid. Keep the mixture for 16 hr at 0◦ C. 14. Collect the hygroscopic precipitate (S.3) by vacuum filtration on a glass filter (porosity 3), wash the precipitate with 10 mL hexane, and dry over P2 O5 in a vacuum desiccator overnight. 15. Characterize the compound by NMR. The compound is stable stored for at least 12 months at room temperature.
Biologically Active Nucleosides
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1,2-O-Isopropylidene-3-C-methyl-α-D-ribofuranose (S.3). Yield of white hygroscopic crystals 7.0 g (89%). m.p. 90◦ -92◦ C (chloroform/hexane). [α]20 D +24.8◦ (c 1, methanol). 1 H NMR (100 MHz, CDCl3 ): 5.89 d (1H, J1 ,2 = 4.0 Hz, H1), 4.15 d (1H, H2), 4.04-3.68 m (3H, H4,5a,5b), 1.62 s (3H, Me), 1.38 s (3H, Me), 1.20 s (3H, C-Me-3). 13 C NMR (CDC13 ): 113.6 (Me2 C), 105.0 (C1), 86.1 (C2), 83.2 (C4), 77.9 (C3), 61.8 (C5), 27.0, 26.8 (Me2 C), 19.2 (C-Me−3).
Prepare S.4 16. Weigh 6.0 g (29.5 mmol) S.3 into a 250-mL round-bottom flask, add 50 mL anhydrous pyridine, and evaporate to dryness using a rotary evaporator equipped with an oil pump. 17. Add 50 mL anhydrous pyridine and a stir bar, close the flask with a dropping funnel connected to a CaCl2 tube. Cool the reaction mixture to 0◦ C using an ice-water bath and slowly add (over 20 min) 3.7 mL (32.0 mmol) benzoyl chloride with magnetic stirring. 18. Keep the reaction mixture for 16 hr at 0◦ C. 19. Add 30 mL sat. NaHCO3 and stir for 30 min. 20. Add 100 mL chloroform and separate the organic layer using a 250-mL separatory funnel. Wash the aqueous layer with 100 mL chloroform. 21. Wash the combined organic layers with 30 mL sat. NaHCO3 and then with 30 mL water. 22. Dry the organic layer over ∼20 g Na2 SO4 , filter off Na2 SO4 by gravity filtration, and wash the precipitate with 30 mL chloroform. Evaporate the combined filtrates using a rotary evaporator connected to a vacuum system. 23. Remove traces of pyridine by coevaporating three times with 30 mL toluene. 24. Dissolve the residue in a minimal volume (∼20 mL) of chloroform at room temperature and add hexane (∼100 to 120 mL) until faintly turbid. Keep the mixture for 16 hr at 0◦ C. 25. Collect the precipitate (S.4) by vacuum filtration on a glass filter (porosity 3), wash the precipitate with 10 mL hexane, and dry over P2 O5 in vacuum desiccator overnight. 26. Characterize the compound by NMR. The compound is stable stored for at least 12 months at room temperature. 5-O-Benzoyl-1,2-O-isopropylidene-3-C-methyl-α-D-ribofuranose (S.4). Yield of white crystals 7.7 g (85%). m.p. 109◦ -110◦ C (chloroform/hexane). [α]20 D +13.5◦ (c 1, chloroform). 1 H NMR (100 MHz, CDCl3 ): 8.10-7.30 m (5H, Bz), 5.77 d (1H, J1 ,2 = 4.0 Hz, H1), 4.60-4.00 m (3H, H4,5a,5b), 4.11 d (1H, H2), 2.70 s (1H, OH, exchanged with D2 O), 1.57 s (3H, Me), 1.34 s (3H, Me), 1.23 s (3H, C-Me-3). 13 C NMR (CDC13 ): 166.9, 133.4, 130.0,128.6 (Bz), 112.6 (Me2 C), 103.7 (C1), 84.2 (C2), 79.5 (C4), 77.0 (C3), 63.5 (C5), 26.6, 26.5 (Me2 C), 18.4 (C-Me-3).
Prepare S.6 27. Weigh 3.1 g (10 mmol) S.4 into a 250-mL round-bottom flask. Add 50 mL of 90% F3 CCOOH, stopper the flask, and keep the solution for 20 min at room temperature. 28. Monitor deprotection by TLC in 95:5 (v/v) CH3 Cl/EtOH. Synthesis of 2 - and 3 -C-Methylribonucleosides
The starting compound S.4 (Rf = 0.98) usually disappears after 20 min at 20◦ C. The product S.5 (Rf = 0.33) moves slower in the same solvent system.
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29. When the reaction is complete, add 30 mL toluene and evaporate all volatile material to dryness in vacuo using a rotary evaporator. Coevaporate two times with 30 mL toluene. 5-O-Benzoyl-3-C-methyl-α,β-D-ribofuranose (S.5) is obtained as a syrup in quantitative yield and may be used in the next steps without purification. H NMR (100 MHz, D2 O-Me2 SO-d6 ): 8.10-7.40 m (5H, Bz), 5.32 d (0.5H, J1,2 = 4.0 Hz, H1), 5.21 d (0.5H, J1,2 = 4.0 Hz, H1), 4.46-4.10 m (3H, H4,5a,5b), 3.82 d (0.5H, H2), 3.70 d (0.5H, H2), 1.30 s (1.5H, C-Me-3), 1.27 s (1.5H, C-Me-3). The ratio of α/β anomer (determined by p.m.r. spectroscopy) is 1:1.
1
30. Dissolve the obtained product S.5 in 20 mL anhydrous pyridine and evaporate to dryness using a rotary evaporator equipped with an oil pump. 31. Add 50 mL anhydrous pyridine and 10 mL acetic anhydride, stopper the flask, and keep the solution for 16 hr at room temperature. 32. Add 50 mg DMAP, stopper the flask, and keep the solution for 24 hr at room temperature. 33. Decompose excess acetic anhydride with 10 mL methanol and keep the solution for 30 min at room temperature. 34. Evaporate to dryness using a rotary evaporator connected to a vacuum system. 35. Add 100 mL chloroform and 100 mL water, separate the organic layer using a 250-mL separatory funnel, and wash the aqueous layer with 100 mL chloroform. 36. Wash the combined organic layers with 50 mL water, then two times with 50 mL sat. NaHCO3 , and then 50 mL water. 37. Dry the organic layer over ∼20 g Na2 SO4 , filter off Na2 SO4 by gravity filtration, and wash the precipitate with 30 mL chloroform. Evaporate the combined filtrates to dryness using a rotary evaporator connected to a vacuum system. 38. Remove traces of pyridine by coevaporating three times with 30 mL toluene using a rotary evaporator connected to a vacuum system. 39. Prepare a slurry of 50 g silica gel in chloroform and pour it into a 3 × 20–cm chromatography column. 40. Dissolve the obtained residue in a minimal amount of chloroform and layer it carefully on top of the silica gel. Wash and elute the column with 100 mL chloroform and collect 25-mL fractions. 41. Evaluate fractions by TLC in chloroform and combine the UV-absorbing fractions that contain S.6 only. 42. Evaporate the volatile materials using a rotary evaporator connected to a vacuum system, and dry the residual syrup for 2 to 3 hr under vacuum using an oil pump. First UV-absorbing fractions contain the pure 1,2,3-tri-O-acetyl-5-O-benzoyl-3-Cmethyl-β-D-ribofuranose. Yield of syrup, which slowly crystallizes while stored, 1.0 g (25%). Further elution with chloroform affords a mixture of 1,2,3-tri-O-acetyl-5-O-benzoyl-3C-methyl-α,β-D-ribofuranose. Yield of syrup 2.3 g (58%). Total yield 3.5 g (83%). Both fractions may be successfully used for the preparation of 3 -C-methylribonucleosides.
43. Characterize the product by TLC and NMR. The compound is stable stored for at least 12 months at ambient temperature.
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1,2,3-Tri-O-acetyl-5-O-benzoyl-3-C-methyl-β-D-ribofuranose (S.6). TLC (chloroform): Rf 0.49. m.p. 55◦ C. [α]20 D +16.3◦ (c 1.1, chloroform). 1 H NMR (200 MHz, CDCl3 ): 8.15-7.30 m (5H, Bz), 6.03 d (1H, J1,2 = 1.5 Hz, H1), 5.43 d (1H, H2), 4.62 dd (1H, J4,5a = 4.0, J4,5a = 7.5 Hz, H4), 4.56 dd (1H, J5a,5b = –12.0 Hz, H5a), 4.40 dd (1H, H5b), 2.10 s (3H, Ac), 2.04 s (3H, Ac), 2.03 s (3H, Ac), 1.72 s (3H, C-Me-3). From 1 H NMR spectra of the mixture of α,β-anomers, chemical shifts and coupling constants of 1,2,3-tri-O-acetyl-5-O-benzoyl-3-C-methyl-α-D-ribofuranose may be readily extracted. 1 H NMR (200 MHz, CDCl3 ): 8.15-7.30 m (5H, Bz), 6.36 d (1H, J1,2 = 4.5 Hz, H1), 5.37 d (1H, H2), 4.76-4.32 m (3H, H4, 5a, 5b), 2.12 s (3H, Ac), 2.08 s (3H, Ac), 2.05 s (3H, Ac), 1.64 s (3H, C-Me-3). BASIC PROTOCOL 2
PREPARATION OF THE PROTECTED 2-C-METHYL-D-RIBOFURANOSE This protocol, outlined in Figure 14.5.2, describes the preparation of the 2-C-methyl-Dribofuranose S.13 starting with the same allofuranose S.1 utilized in the synthesis of 3-C-methyl-D-ribofuranose S.6 (see Basic Protocol 1). The key element of this strategy (Beigelman et al., 1987) is the shortening of the carbon chain in S.1 between C1 and C2 rather than between C5 and C6 as in Basic Protocol 1.
Figure 14.5.2 Synthesis of 1,2,3-tri-O-acetyl-5-O-p-methylbenzoyl-2-C-methyl-β-D-ribofuranose (S.13) from S.1 in a 36% expected overall yield. AcOH, acetic acid; Ac2 O, acetic anhydride; BnCl, benzyl chloride; DMAP, 4-dimethylaminopyridine; DMSO, dimethyl sulfoxide; Pd(OH)2 /C, palladium hydroxide on carbon; TolCl, p-methylbenzoyl chloride (toluoyl chloride). Synthesis of 2 - and 3 -C-Methylribonucleosides
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To achieve this transformation and ensure formation of the ribofuranose ring with simultaneous retention of the methyl group and its configuration at C3 (which becomes C2 in S.11), it is necessary to protect the 6-OH and 3-OH in the starting compound S.1. The benzyl (Bn) group has been selected to protect the tertiary OH, taking into account possible migration of alternative acyl protecting groups during the acid-catalyzed removal of the 1,2-isopropylidene group at later stages (S.8 to S.9). The benzylation requires somewhat forced conditions (70◦ C), but proceeds smoothly, allowing for isolation of the protected allofuranose S.7 in a 90% yield in crystalline form. Selective removal of the 5,6-O-isopropylidene group with 75% AcOH leads to crystalline diol S.8 in a 90% yield. Among different options for selective introduction of the methylbenzoyl group at 6-OH, the best results were consistently obtained with stannyledene activation of the 5,6-diol according to the Moffat procedure (Wagner et al., 1974). The mono 6-O-methylbenzoyl derivative S.9 was isolated in an 80% yield after chromatography. It is recommended that the next four steps be performed without isolation to obtain the protected 2-C-methyl-D-ribofuranose S.11 in a 75% to 80% yield. Removal of the 1,2-O-isopropylidene group proceeds smoothly with 90% trifluoroacetic acid. Removal of residual acid by several coevaporations with toluene is recommended. The NaIO4 oxidation of the 1,2-diol group generates an intermediate furanose with CHO protection at C3. Subsequent removal of the formyl group with NaOMe/MeOH must be monitored carefully by TLC and quenched quickly with Dowex 50 (H+ ) resin when completed to prevent removal of the 5-O-methylbenzoyl group. This critical step directly influences the yield of the four-step sequence from S.9 to S.11. Standard acetylation of the deformylated furanose proceeds in high yield and provides the β-anomer S.11 in a 75% to 80% yield starting from S.9. Removal of the benzyl protecting group by homogeneous hydrogenation and subsequent acetylation in the presence of DMAP completes synthesis of the target precursor, l,2,3-tri-O-acetyl-5-O-p-methylbenzoyl-2-C-methyl-βD-ribofuranose (S.13), as a β-anomer in high yield.
Materials 1,2:5,6-Di-O-isopropylidene-3-C-methyl-α-D-allofuranose (S.1; Bio et al., 2004) Anhydrous dimethyl sulfoxide (DMSO) 80% (w/w) sodium hydride (NaH) in mineral oil Benzyl chloride, reagent grade Chloroform (CHCl3 ), reagent grade Anhydrous sodium sulfate (Na2 SO4 ) Silica gel: Kieselgel 60 (0.06 to 0.20 mm; Merck) 75% (v/v) acetic acid (AcOH) 1-Butanol, reagent grade Ethanol (EtOH), reagent grade Phosphorus pentoxide (P2 O5 ) Dibutylstannoxane Dry methanol (MeOH) Triethylamine, reagent grade p-Methylbenzoyl chloride (TolCl) Saturated sodium bicarbonate (NaHCO3 ) 90% (v/v) trifluoroacetic acid (F3 CCOOH) Toluene, reagent grade 1,4-Dioxane 1 M sodium periodate (NaIO4 ) Ethyl acetate (EtOAc), reagent grade 1 M sodium methoxide (NaOMe) in methanol Dowex 50 (H+ ) resin
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Anhydrous pyridine Acetic anhydride, reagent grade Cyclohexene 20% palladium hydroxide on carbon (Pd(OH)2 /C) Hyflo Super Cel 4-Dimethylaminopyridine (DMAP) 250-mL round-bottom flasks Reflux condensers equipped with a CaCl2 protection tube Oil bath with temperature control 250- and 500-mL separatory funnels Rotary evaporator equipped with a water aspirator and an oil pump 3 × 20– and 5 × 30–cm sintered glass chromatography columns, porosity 3 Silica-coated aluminum TLC plates with fluorescent indicator (Merck silicagel 60 F254 ) 254-nm UV lamp Vacuum oil pump Vacuum filtration system with glass filters (porosity 3) Vacuum desiccator 100-mL funnel with sintered disc (porosity 3) Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E) Prepare S.7 1. Weigh 11.2 g (40.8 mmol) S.1 into a 250-mL round-bottom flask with a stir bar. Add 120 mL anhydrous dimethyl sulfoxide and attach a condenser equipped with a CaCl2 protection tube. 2. Add 3.0 g (100 mmol) of 80% (w/w) sodium hydride in mineral oil in 1.0-g portions over a 20-min period with stirring at room temperature. Stir the reaction mixture until the complete evolution of hydrogen (20 to 30 min). 3. Add 12.0 mL (104 mmol) benzyl chloride in one portion and heat the mixture with stirring for 2.5 hr in a 70◦ C oil bath. 4. Cool the flask to room temperature, pour into a mixture of water and ice (400 mL), and stir for 20 min. 5. Add 200 mL chloroform and separate the organic layer using a 500-mL separatory funnel. Wash the aqueous layer with 200 mL chloroform. Wash the combined organic layers with 50 mL water. 6. Dry the organic layer over ∼40 g Na2 SO4 , filter off the Na2 SO4 by gravity filtration, and wash the precipitate with 30 mL chloroform. Evaporate the combined filtrates to dryness using a rotary evaporator connected to a vacuum system. 7. Prepare a slurry of 100 g silica gel in chloroform and pour it into a 5 × 30–cm chromatography column. 8. Dissolve the obtained residue in a minimal amount of chloroform and layer it carefully on top of the silica gel. Wash and elute the column with chloroform and collect 25-mL fractions.
Synthesis of 2 - and 3 -C-Methylribonucleosides
9. Evaluate fractions by TLC in chloroform and combine the fractions that contain S.7 only. The first fractions contain benzyl chloride.
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10. Evaporate the volatile materials from the combined fractions using a rotary evaporator connected to a vacuum system, and dry the residual syrup for 2 to 3 hr under a vacuum using an oil pump. S.7 is obtained as a slowly crystallized syrup (13.0 g) that may be used in the next step without further purification.
11. Characterize the compound by NMR. The compound is stable stored for at least 12 months at room temperature. 3-O-Benzyl-1,2:5,6-di-O-isopropylidene-3-C-methyl-α-D-allofuranose (S.7). Yield of slowly crystallized syrup 13.0 g (87%). 1 H NMR (100 MHz, CDCl3 ): 7.40-7.13 m (5H, PhCH2 ), 5.67 d (1H, J1 ,2 = 3.7 Hz, H1), 4.64 brs (2H, CH2 Ph), 4.29 d (1H, H2), 4.193.95 m (3H, H4, 5, 6a, 6b), 1.58 s (3H, Me), 1.39 s (3H, Me), 1.34 s (6H, 2Me), 1.27 s (3H, C-Me-3).
Prepare S.8 12. Weigh 10.9 g (30 mmol) S.7 into a 250-mL round-bottom flask. Add 100 mL of 75% acetic acid and keep the solution for 24 hr at room temperature. 13. Evaporate all volatile material to dryness in vacuo using a rotary evaporator. Add 50 mL of 1-butanol to the residue and evaporate again to dryness. Repeat evaporation two additional times with 50 mL of 1-butanol. 14. Add 40 mL ethanol, attach a condenser, and reflux (78◦ C) to complete dissolution of the solid. Cool the flask to room temperature and keep the mixture 16 hr at 0◦ C. 15. Collect the precipitate (S.8) by vacuum filtration on a glass filter (porosity 3), wash the precipitate with 10 mL ethanol, and dry over P2 O5 in a vacuum desiccator overnight. 16. Characterize the compound by NMR. The compound is stable stored for at least 12 months room temperature. 3-O-Benzyl-1,2-O-isopropylidene-3-C-methyl-α-D-allofuranose (S.8). Yield of white crystals 8.76 g (90%). m.p. 117◦ -118◦ C (ethanol). [α]20 D +44.8◦ (c 1, chloroform). 1 H NMR (100 MHz, DMSO-d6 ): 7.38-7.14 m (5H, PhCH2 ), 5.68 d (1H, J1 ,2 = 3.8 Hz, H1), 4.60 d (1H, JH,H = –11.5 Hz, CHHPh), 4.52 d (1H, CHHPh), 4.44 d (1H, H2), 4.28 t (1H, JOH,6a = JOH,6b = 5.0 Hz, 6-OH, exchanged with D2 O), 3.86 t (1H, JOH,5 = 7.0 Hz, 5-OH, exchanged with D2 O), 3.64-3.38 m (3H, H4, 5, 6a, 6b), 1.48 s (3H, Me), 1.30 s (6H, Me, C-Me-3).
Prepare S.9 17. Weigh 2.7 g (8.32 mmol) S.8 into a 250-mL round-bottom flask with a stir bar. Add 2.11 g (8.44 mmol) dibutylstannoxane and 100 mL dry MeOH, and attach a condenser equipped with a CaCl2 protection tube. 18. Reflux the mixture in a 70◦ C oil bath until complete dissolution (1 hr). 19. Cool the flask to room temperature. Add in one portion, with stirring, 3.55 mL (25.32 mmol) triethylamine and 3.91 g (25.32 mmol) p-methylbenzoyl chloride. Attach a condenser equipped with a CaCl2 protection tube and stir the reaction mixture for 2 hr at room temperature. 20. Collect the formed precipitate by vacuum filtration on a glass filter (porosity 3), wash the precipitate with 20 mL chloroform, and concentrate the combined filtrates to dryness using a rotary evaporator connected to a vacuum system. Biologically Active Nucleosides
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21. Add to the residue 70 mL chloroform and 20 mL water, separate the organic layer using a 250-mL separatory funnel, and wash the organic layer with 10 mL sat. NaHCO3 and 10 mL water. 22. Dry the organic layer over ∼10 g Na2 SO4 , filter off the Na2 SO4 by gravity filtration, and wash the precipitate with 20 mL chloroform. Evaporate the combined filtrates to dryness using a rotary evaporator connected to a vacuum system. 23. Prepare a slurry of 50 g silica gel in chloroform and pour it into a 3 × 20–cm chromatography column. 24. Dissolve the obtained residue in a minimal amount of chloroform and layer it carefully on top of the silica gel. Wash and elute the column with chloroform and collect 25-mL fractions. 25. Evaluate fractions by TLC in chloroform and combine the fractions that contain S.6 only. 26. Evaporate the volatile materials from the combined fractions using a rotary evaporator connected to a vacuum system, and dry the residual syrup for 2 to 3 hr under vacuum using an oil pump. 27. Characterize the compound by TLC and NMR. The compound is stable stored for at least 12 months at room temperature. 3-O-Benzyl-1,2-O-isopropylidene-3-C-methyl-6-O-methylbenzoyl-α-D-allofuranose (S.9). Yield of thick colorless syrup 3.0 g (81%). TLC (chloroform): Rf 0.50. [α]20 D +34.2◦ (c 1, chloroform). 1 H NMR (100 MHz, DMSO-d6 ): 7.86 d (2H, JH,H = 7.0 Hz, MeBz), 7.44-7.04 m (7H, MeBz, PhCH2 ), 5.71 d (1H, J1 ,2 = 3.7 Hz, H1), 4.98 d (1H, JOH,5 = 5.5 Hz, 5-OH, exchanged with D2 O), 4.64 d (1H, JH,H = –11.5 Hz, CHHPh), 4.54 d (1H, CHHPh), 4.48 d (1H, H2), 4.45-3.82 m (4H, H4,5, 6a, 6b), 2.38 s (3H, MeBz), 1.48 s (3H, Me), 1.32 s (6H, Me, C-Me-3). Anal. calcd. for C25 H30 O7 : C, 67.86; H, 6.83; found: C, 67.54; H, 6.79.
Prepare S.11 28. Weigh 4.4 g (10 mmol) S.9 into a 250-mL round-bottom flask. Add 40 mL of 90% F3 CCOOH, stopper the flask, and keep the solution for 20 min at room temperature. 29. Monitor deprotection by TLC in 95:5 (v/v) CH3 Cl/EtOH. The starting compound S.9 (Rf = 0.99) usually disappears after 20 min at 20◦ C. The product S.10 (Rf = 0.43) moves slower in the same solvent system.
30. When the reaction is complete, add 30 mL toluene and evaporate all volatile material to dryness in vacuo using a rotary evaporator. Coevaporate three times with 20 mL toluene. 31. Add to the residue 70 mL chloroform and 20 mL water, separate the organic layer using a 250-mL separatory funnel, and wash the organic layer with 10 mL sat. NaHCO3 and 10 mL water. 32. Dry the organic layer over ∼10 g Na2 SO4 , filter off the Na2 SO4 by gravity filtration, and wash the precipitate with 20 mL chloroform. Evaporate the combined filtrates to dryness using a rotary evaporator connected to a vacuum system. 3-O-Benzyl-3-C-methyl-6-O-methylbenzoyl-α,β-D-allofuranose (S.10) is obtained as a syrup in near quantitative yield and may be used in the next steps without purification. Synthesis of 2 - and 3 -C-Methylribonucleosides
33. Dissolve the obtained product S.10 in 60 mL of 1,4-dioxane and 20 mL water, and add in one portion 11 mL of 1 M NaIO4 . Stir for 16 hr at room temperature.
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34. Monitor periodate oxidation by TLC in 3:2 (v/v) toluene/ethyl acetate. The starting compound S.10 (Rf = 0.20) usually disappears after 8 to 10 hr at 20◦ C. The product (Rf = 0.69) moves faster in the same solvent system.
35. Add 80 mL ethanol and filter NaIO3 by vacuum filtration on a glass filter (porosity 3). Wash the precipitate with 30 mL ethanol, and evaporate the combined filtrates to dryness using a rotary evaporator connected to a vacuum system. 36. Add to the residue 70 mL chloroform and 20 mL water, separate the organic layer using a 250-mL separatory funnel, and wash the organic layer with 10 mL water. 37. Dry the organic layer over ∼10 g Na2 SO4 , filter off the Na2 SO4 by gravity filtration, and wash the precipitate with 20 mL chloroform. Evaporate the combined filtrates to dryness using a rotary evaporator connected to a vacuum system, and dry the residual syrup for 2 to 3 hr under vacuum using an oil pump. 38. Dissolve the residue in 50 mL dry methanol and add 0.3 mL freshly prepared 1 M NaOMe in methanol. Keep the solution for 20 to 30 min at room temperature. 39. Monitor deformylation by TLC in 3:2 (v/v) toluene/ethyl acetate. The starting compound (Rf = 0.69) usually disappears after 25 min at 20◦ C. The product (Rf = 0.58) moves slower in the same solvent system.
40. Neutralize the reaction mixture with ∼0.2 to 0.3 mL Dowex 50 (H+ ) resin. Check the pH of the solution by applying a small sample onto a piece of moistened pH paper.
41. Filter resin by vacuum filtration on a glass filter (porosity 3), wash the resin with 10 mL methanol, and evaporate the combined filtrates to dryness using a rotary evaporator connected to a vacuum system. 42. Dissolve the obtained syrup in 30 mL anhydrous pyridine and evaporate to dryness using a rotary evaporator equipped with an oil pump. 43. Add 50 mL anhydrous pyridine and 10 mL acetic anhydride, stopper the flask, and keep the solution for 16 hr at room temperature. 44. Decompose excess acetic anhydride with 10 mL methanol and keep the solution for 30 min at room temperature. Evaporate to dryness using a rotary evaporator connected to a vacuum system. 45. Add 100 mL chloroform and 100 mL water, separate the organic layer using a 250mL separatory funnel, and wash the aqueous layer with 100 mL chloroform. Wash the combined organic layers with 50 mL water, then two times with 50 mL sat. NaHCO3 , and finally with 50 mL water. 46. Dry the organic layer over ∼20 g Na2 SO4 , filter off the Na2 SO4 by gravity filtration, and wash the precipitate with 30 mL chloroform. Evaporate the combined filtrates to dryness using a rotary evaporator connected to a vacuum system. 47. Remove traces of pyridine by coevaporating three times with 30 mL toluene using a rotary evaporator connected to a vacuum system. 48. Prepare a slurry of 50 g of silica gel in chloroform and pour it into a 3 × 20–cm chromatography column. 49. Dissolve the obtained residue in a minimal amount of chloroform and layer it carefully on top of the silica gel. Wash and elute the column with chloroform and collect 25-mL fractions.
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50. Evaluate fractions by TLC in chloroform and combine the fractions that contain S.11 only. 51. Evaporate the volatile materials from the combined UV-absorbing fractions using a rotary evaporator connected to a vacuum system, and dry the residual syrup for 2 to 3 hr under vacuum using an oil pump. 52. Characterize the compound by TLC and NMR. The compound is stable stored for at least 12 months at room temperature. l,3-Di-O-acetyl-2-O-benzyl-2-C-methyl-5-O-p-methylbenzoyl-β-D-ribofuranose (S.11). Yield of thick colorless syrup 3.5 g (77%). TLC (chloroform): Rf 0.50. [α]20 D +11.8◦ (c 0.9, chloroform). 1 H NMR (100 MHz, CDCl3 ): 7.82 d (2H, JH,H = 7.0 Hz, MeBz), 7.22-7.02 m (7H, MeBz, PhCH2 ), 6.14 s (1H, H1), 5.29 d (1H, J3 ,4 = 7.8 Hz, H3), 4.54 s (2H, CH2 Ph), 4.52-4.20 m (3H, H4, 5a, 5b), 2.30 s (3H, MeBz), 2.02 s (3H, Ac), 1.90 s (3H, Ac), 1.34 s (3H, C-Me-2). Anal. calcd. for C25 H28 O8 : C, 65.78; H, 6.18; found: C, 65.42; H, 6.14.
Prepare S.12 53. Weigh 0.91 g (2 mmol) S.11 into a 250-mL round-bottom flask with a stir bar and dissolve in 70 mL dry EtOH. Add to the solution 50 mL cyclohexene and 400 mg of 20% Pd(OH)2 /C in one portion. 54. Attach a condenser equipped with a CaCl2 protection tube and reflux the mixture 2 to 3 hr in an 83◦ C oil bath. 55. Monitor debenzylation by TLC in chloroform. The starting compound S.11 (Rf = 0.50) usually disappears after 2.5 hr. The product (Rf = 0.21) moves slower in the same solvent system.
56. Place a 2- to 3-cm layer of Hyflo Super Cel onto a 100-mL funnel with sintered disc (porosity 3) and wash the layer with 10 mL ethanol. 57. Cool the flask to room temperature, filter the suspension using a vacuum pump, and wash the Hyflo Super Cel layer with 30 mL ethanol. 58. Evaporate the combined filtrates to dryness using a rotary evaporator connected to a vacuum system. 59. Prepare a slurry of 20 g silica gel in chloroform and pour it into a 3 × 20–cm chromatography column. 60. Dissolve the obtained residue in a minimal amount of chloroform and layer it carefully on top of the silica gel. Wash and elute the column with chloroform and collect 25-mL fractions. 61. Evaluate fractions by TLC in chloroform and combine the fractions that contain S.12 only. 62. Evaporate the volatile materials from the combined UV-absorbing fractions using a rotary evaporator connected to a vacuum system, and dry the residual syrup for 2 to 3 hr under vacuum using an oil pump. 63. Characterize the compound by TLC and NMR. The compound is stable stored for at least 12 months at room temperature. Synthesis of 2 - and 3 -C-Methylribonucleosides
1,3-Di-O-acetyl-2-C-methyl-5-O-p-methylbenzoyl-β-D-ribofuranose (S.12). Yield of thick colorless syrup 0.6 g (82%). TLC (chloroform): Rf 0.21. [α]20 D –25.6◦ (c 1, chloroform). 1 H NMR (100 MHz, CDCl3 ): 7.88 d (2H, JH,H = 7.0 Hz, MeBz), 7.15 d (2H, JH,H = 7.0 Hz, MeBz), 5.98 s (1H, H1), 5.28 d (1H, J3 ,4 = 7.2 Hz, H3), 4.64-4.24 m (3H, H4, 5a, 5b), 2.40 s (3H, MeBz), 2.14 s (3H, Ac), 2.00 s (3H, Ac), 1.32 s (3H, C-Me-2).
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Prepare S.13 64. Weigh 0.80 g (2.2 mmol) S.12 into a 250-mL round-bottom flask, dissolve it in 10 mL anhydrous pyridine, and evaporate to dryness using a rotary evaporator equipped with an oil pump. 65. Add 30 mL anhydrous pyridine, 5 mL acetic anhydride, and 50 mg DMAP. Stopper the flask and keep the solution for 24 hr at room temperature. 66. Decompose excess acetic anhydride with 5 mL methanol and keep the solution for 30 min at room temperature. Evaporate to dryness using a rotary evaporator connected to a vacuum system. 67. Add 50 mL chloroform and 50 mL water, separate the organic layer using a 250-mL separatory funnel, and wash the aqueous layer with 50 mL chloroform. Wash the combined organic layers with 20 mL water, then two times with 20 mL sat. NaHCO3 , and finally with 20 mL water. 68. Dry the organic layer over ∼ 20 g Na2 SO4 , filter off the Na2 SO4 by gravity filtration, and wash the precipitate with 20 mL chloroform. Evaporate the combined filtrates to dryness using a rotary evaporator connected to a vacuum system. 69. Remove traces of pyridine by coevaporating three times with 30 mL toluene using a rotary evaporator connected to a vacuum system. 70. Prepare a slurry of 30 g silica gel in chloroform and pour it into a 3 × 20–cm chromatography column. 71. Dissolve the obtained residue in a minimal amount of chloroform and layer it carefully on top of the silica gel. Wash and elute the column with chloroform and collect 25-mL fractions. 72. Evaluate fractions by TLC in chloroform and combine the UV-absorbing fractions that contain S.13 only. 73. Evaporate the volatile materials from the combined fractions using a rotary evaporator connected to a vacuum system, and dry the residual syrup for 2 to 3 hr under vacuum using an oil pump. 74. Characterize the compound by TLC and NMR. The compound is stable stored for at least 12 months at room temperature. 1,2,3-Tri-O-acetyl-2-C-methyl-5-O-p-methylbenzoyl-β-D-ribofuranose (S.13). Yield of thick colorless syrup 0.80 g (90%). TLC (chloroform): Rf 0.51. [α]20 D –1.3◦ (c 1.2, chloroform). 1 H NMR (100 MHz, CDCl3 ): 7.87 d (2H, JH,H = 7.0 Hz, MeBz), 7.17 d (2H, JH,H = 7.0 Hz, MeBz), 6.45 s (1H, H1), 5.40 d (1H, J3 ,4 = 7.3 Hz, H3), 4.60-4.20 m (3H, H4,5a,5b), 2.38 s (3H, MeBz), 2.10 s (3H, Ac), 2.08 s (3H, Ac), 1.98 s (3H, Ac), 1.60 s (3H, C-Me-2). Anal. calcd. for C20 H24 O9 : C, 58.82; H, 5.92; found: C, 58.70; H, 5.90.
PREPARATION OF 1-(3-C-METHYL-β-D-RIBOFURANOSYL)URACIL 3 -C-Methylribonucleosides
(S.15) are prepared in two steps starting from bistrimethylsilyl derivatives of nucleic acid bases and the protected 3-C-methyl-Dribofuranose (S.6) as outlined in Figure 14.5.3. The Vorbr¨uggen reaction (described in Vorbr¨uggen and Ruh-Pohlenz, 2001) has completely replaced the previously employed methods for synthesis of nucleosides and their analogs. The method involves glycosylation of trimethylsilyl derivatives of heterocyclic bases with peracylated sugars in aprotic solvents in the presence of Lewis acids such as SnCl4 or trimethylsilyl trifluoromethanesulfonate (TMSOTf). The use of this approach has significantly simplified the reaction procedure and increased the yields of the target products. The presence of a
BASIC PROTOCOL 3
Biologically Active Nucleosides
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Figure 14.5.3 Synthesis of 3 -C-methylribonucleosides (S.15). The expected overall yields are given in parentheses. DCE, 1,2-dichloroethane; TMS2 B, bis-trimethylsilyl derivatives of heterocyclic bases; TMSOTf, trimethylsilyl trifluoromethanesulfonate.
2-O-acyl group in a carbohydrate residue is believed to be crucial for the stereochemistry of this reaction, since it stabilizes the C1 carbonium ion generated via an intermediate 1,2-acyloxonim ion and yields 1,2-trans derivatives. In the first step of this procedure, fully protected 3 -C-methylribonucleosides (S.14) are prepared in good yields. In general, peracylated S.6 behaves more like an unsubstituted peracylated ribofuranose in this condensation reaction. The cleavage of the benzoyl and acetyl protecting groups is achieved using 5 M ammonia in methanol under mild conditions, and 3 -C-methylribonucleosides (S.15) are purified by crystallization (Mikhailov et al., 1983). This protocol describes the preparation of 3 -C-methyluridine (S.15a), which was obtained in a 60% overall yield.
Materials
Synthesis of 2 - and 3 -C-Methylribonucleosides
Uracil Anhydrous pyridine 1,1,1,3,3,3-Hexamethyldisilazane (HMDS), reagent grade Anhydrous toluene, reagent grade Anhydrous 1,2-dichloroethane 1,2,3-Tri-O-acetyl-5-O-benzoyl-3-C-methyl-α,β-D-ribofuranose (S.6; see Basic Protocol 1) 2 M trimethylsilyl trifluoromethanesulfonate (TMSOTf)
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Chloroform (CHCl3 ), reagent grade Ethanol (EtOH), reagent grade Saturated sodium bicarbonate (NaHCO3 ) Hyflo Super Cel Anhydrous sodium sulfate (Na2 SO4 ) Silica gel: Kieselgel 60 (0.06 to 0.20 mm; Merck) 5 M ammonia in methanol (half saturated at 0◦ C) Methylene chloride Acetone Diethyl ether, reagent grade Phosphorus pentoxide (P2 O5 ) 100- and 250-mL round-bottom flasks Reflux condenser equipped with a CaCl2 protection tube Oil bath with temperature control Rotary evaporator equipped with a water aspirator Silica-coated aluminum TLC plates with fluorescent indicator (Merck silica gel 60 F254 ) 254-nm UV lamp 100-mL funnel with a sintered disc (porosity 3) Vacuum pump for filtration 100- and 250-mL separatory funnels 3 × 20–cm sintered glass chromatography column, porosity 3 Vacuum oil pump Glass filters (porosity 3) Vacuum desiccator Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E) Perform glycosylation 1. Weigh 1.12 g (10 mmol) uracil into a 250-mL round-bottom flask, add 5 mL anhydrous pyridine and 10 mL HMDS (b.p. 125◦ C), and attach a condenser equipped with a CaCl2 protection tube. 2. Reflux the mixture in a 130◦ C oil bath until complete dissolution of uracil (3 to 4 hr). 3. Cool the flask, remove the condenser, and concentrate the solution to a viscous oil using a rotary evaporator connected to a vacuum system (bath temperature ∼30◦ to 35◦ C). The formed bis-O-trimethylsilyl uracil is very hygroscopic.
4. Dissolve the residue in 20 mL dry toluene and concentrate the solution to a viscous oil using a rotary evaporator (bath temperature ∼30◦ to 35◦ C). Remove traces of pyridine by coevaporating two times with 20 mL toluene. 5. Dissolve the residue in 20 mL anhydrous 1,2-dichloroethane, and add in one portion a solution of 2.6 g (6.6 mmol) S.6 in 30 mL anhydrous 1,2-dichloroethane, plus 6 mL of 2 M TMSOTf in anhydrous 1,2-dichloroethane. Stopper the flask and keep the solution for 16 hr at room temperature. 6. Monitor reaction by TLC in 98:2 (v/v) CH3 Cl/EtOH. The starting S.13 (Rf = 0.95) usually disappears after 10 to 16 hr at room temperature. The product S.14a (Rf = 0.49) moves slower in the same solvent.
Biologically Active Nucleosides
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7. When the reaction is complete, add 30 mL sat. NaHCO3 and stir the suspension for 20 min at 0◦ C. 8. Place a 2- to 3-cm layer of Hyflo Super Cel onto a 100-mL funnel with a sintered disc (porosity 3) and wash it with 20 mL chloroform. Filter the suspension through the layer using a vacuum pump and wash the layer with 50 mL chloroform. 9. Separate the organic layer using a 250-mL separatory funnel and wash the organic layer with 50 mL water. 10. Dry the organic layer over ∼10 g Na2 SO4 , filter off the Na2 SO4 by gravity filtration, and wash the precipitate with 20 mL chloroform. Concentrate the combined filtrates to a solid using a rotary evaporator connected to a vacuum system. 11. Prepare a slurry of 30 g silica gel in chloroform and pour it into a 3 × 20–cm chromatography column. 12. Dissolve the obtained residue in a minimal amount of chloroform and layer it carefully on top of the silica gel. Wash the column with 200 mL chloroform and 300 mL of 1% (v/v) ethanol in chloroform, and elute with 2% (v/v) ethanol in chloroform. Collect 25-mL fractions. 13. Evaluate fractions by TLC using 98:2 (v/v) CH3 Cl/EtOH and combine the fractions that contain S.14a only. 14. Evaporate the volatile materials from the combined fractions using a rotary evaporator connected to a vacuum system, and dry the residual foam for 2 to 3 hr under vacuum using an oil pump. 15. Grind the foam with a stainless steel spatula and dry the resulting powder 12 to 16 hr using a vacuum oil pump. 16. Characterize the product by TLC and NMR. The compound is stable stored for at least 12 months at ambient temperature. 1-(2,3-Di-O-acetyl-5-O-benzoyl-3-C-methyl-β-D-ribofuranosyl)uracil (S.14a). Yield of white amorphous solid 2.35 g (80%). TLC (98:2 v/v CH3 Cl/EtOH): Rf = 0.49.1 H NMR (200 MHz, CDCl3 ): 8.55 brs (1H, NH), 8.06-7.32 m (5H, Bz), 7.36 d (1H, J6,5 = 8.0 Hz, H6), 6.16 d (1H, J1 ,2 = 7.5 Hz, H1 ), 5.43 dd (1H, J5,NH = 2.0 Hz, H5; converted into doublet with J5,6 = 8.0 Hz, on addition of D2 O), 5.36 d (1H, H2 ), 4.88 dd (1H, J4 ,5 a = 3.3, J4 ,5 b = 3.6 Hz, H4 ), 4.70 dd (1H, J5 a,5 b = –12.8, H5 a), 4.50 dd (1H, H5 b), 2.16 s (6H, 2 Ac), 1.73 s (3H, C-Me-3 ).
Perform deprotection 17. Weigh 1.0 g (2.24 mmol) S.14a into a 100-mL round-bottom flask, add 15 ml of 5 M ammonia in methanol (half saturated at 0◦ C), stopper the flask, and keep the solution for 24 hr at room temperature. 18. Evaporate all volatile material under reduced pressure using a rotary evaporator. 19. Partition the residue between 20 mL chloroform and 30 mL water using a 100-mL separatory funnel. Separate the aqueous layer and wash it two times with 20 mL methylene chloride. Evaporate aqueous layer to dryness using a rotary evaporator connected to a vacuum system. 20. Dissolve the residue in a minimal amount of hot water (2 to 3 mL) and keep the mixture for 2 to 3 days at 0◦ C. Synthesis of 2 - and 3 -C-Methylribonucleosides
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21. Collect the precipitate by vacuum filtration on a glass filter (porosity 3), wash with 3 mL acetone and then 3 mL diethyl ether, and dry in a vacuum desiccator with P2 O5 for 24 hr at room temperature. 22. Characterize the compound by UV, TLC, and 1 H NMR. 1-(3-C-Methyl-β-D-ribofuranosyl)uracil (S.15a). Yield of white crystals 0.43 g (75%). TLC (8:2 v/v CH3 Cl/EtOH): Rf 0.30. m.p. 213◦ -214◦ C (water). UV λmax (ε): (pH 1-7) 262 nm (10500); (pH 13) 262 nm (7700). NMR (200 MHz, D2 O): 7.90 d (1H, J6 ,5 = 8.0 Hz, H6), 5.96 d (1H, J1 ,2 = 7.8 Hz, H1 ), 5.90 d (1H, H5), 4.18 d (1H, H2 ), 4.09 dd (1H, J4 ,5 a = 3.8, J4 ,5 b = 4.9 Hz, H4 ), 3.85 dd (1H, J5 a,5 b = –12.8 Hz, H5 a), 3.74 dd (1H, H5 b), 1.40 s (3H, C-Me-3 ). Anal. calcd. for C10 H14 N2 O6 : C, 46.51; H, 5.46; N, 10.85; found: C, 46.67; H, 5.62; N, 10.71.
PREPARATION OF 3-C-METHYL-β-D-RIBOFURANOSYL DERIVATIVES OF CYTOSINE, ADENINE, AND GUANINE
ALTERNATE PROTOCOL 1
1-(3-C-Methyl-β-D-ribofuranosyl)cytosine (S.15b), 9-(3-C-methyl-β-D-ribofuranosyl)adenine (S.15c), and 9-(3-C-methyl-β-D-ribofuranosyl)guanine (S.15d) are prepared in two steps starting with bis-trimethylsilyl nucleobases and S.6 using the steps outlined in Basic Protocol 3 (Fig. 14.5.3). The adenine derivative is prepared from bis-trimethylsilylN6 -benzoyladenine and is obtained in an overall yield of 56%. The guanine derivative is prepared from bis-trimethylsilyl-N2 -palmitoylguanine and is obtained in an overall yield of 32%. The cytosine derivative is prepared from bis-trimethylsilyl-cytosine and is obtained in an overall yield of 53%. N4 -Benzoylcytosine can also be used successfully in the condensation reaction with nearly the same overall yield. One procedural modification is used for the cytosine derivative S.15b. For crystallization after step 19 of Basic Protocol 3, the residue is dissolved in a minimal amount of 95% ethanol (4 to 5 mL), diethyl ether is added until the mixture is faintly turbid, and the mixture is kept for 2 to 3 days at 0◦ C. Crystalline compound S.15b is then collected by filtration and characterized as described in steps 21 and 22. 1-(2,3-Di-O-acetyl-5-O-benzoyl-3-C-methyl-β-D-ribofuranosyl)cytosine (S.14b). Yield of white amorphous solid 68%. TLC (9:1 v/v CH3 Cl/EtOH): Rf 0.42. 1 H NMR (100 MHz, CDCl3 ): 8.14-7.44 m (5H, Bz), 7.50 d (1H, J6,5 = 7.5 Hz, H6), 6.35 d (1H, J1 ,2 = 7.5 Hz, H1 ), 5.55 d (1H, H5), 5.38 d (1H, H2 ), 4.90-4.53 m (3H, H4 ,5 a,5 b), 2.15 s (6H, 2 Ac), 1.70 s (3H, C-Me-3 ). 1-(3-C-Methyl-β-D-ribofuranosyl)cytosine (S.15b). Yield of white crystals 68%. m.p. 222◦ 225◦ C (ethanol/ether). TLC (8:2 v/v CH3 Cl/EtOH): Rf 0.12. UV λmax (ε): (pH 7-13) 271 nm (8800); (pH 1) 280 nm (12700). 1 H NMR (200 MHz, D2 O): 7.82 d (1H, J6 ,5 = 7.5 Hz, H6), 6.04 d (1H, H5), 5.94 d (1H, J1 ,2 = 7.5 Hz, H1 ), 4.16 d (1H, H2 ), 4.10 dd (1H, J4 ,5 a = 3.6, J4 ,5 b = 4.9 Hz, H4 ), 3.82 dd (1H, J5 a,5 b = –12.6, H5 a), 3.71 dd (1H, H5 b), 1.38 s, (3H, C-Me-3 ). Anal. calcd. for C10 H15 N3 O5 : C, 46.69; H, 5.88; N, 16.33; found: C, 46.63; H, 5.83; N, 16.28. 9-(2,3-Di-O-acetyl-5-O-benzoyl-3-C-methyl-β-D-ribofuranosyl)-N6 -benzoyladenine (S.14c). Yield of white amorphous solid 78%. Rf = 0.45 (98:2 v/v CH3 Cl/EtOH).1 H NMR (200 MHz, CDCl3 ): 8.95 brs (1H, NH), 8.63 s (1H, H8), 8.07 s (1H, H2), 8.06-7.30 m (10H, Bz), 6.29 d (1H, J1 ,2 = 7.5 Hz, H1 ), 6.07 d (1H, H2 ), 4.97 dd (1H, J4 ,5 a = 3.5, J4 ,5 b = 4.5 Hz, H4 ), 4.80 dd (1H, J5 a,5 b = –12.5 Hz, H5 a), 4.56 dd (1H, H5 b), 2.17 s (3H, Ac), 2.09 s (3H, Ac), 1.83 s (3H, C-Me-3 ). 9-(3-C-Methyl-β-D-ribofuranosyl)adenine (S.15c). Yield of white crystals 72%. TLC (8:2 v/v CH3 Cl/EtOH): Rf 0.30. m.p. 207◦ -208◦ C (softening at 172◦ C) (water). [α]20 D –53.0◦ (c 1, water). UV λmax (ε): (pH 7-13) 260 nm (15000); (pH 1) 258 nm (14700). NMR (200 MHz, D2 O): 8.25 s (1H, H8), 8.12 s (1H, H2), 5.96 d (1H, J1 ,2 = 8.0 Hz, H1 ), 4.62 d (1H, H2 ), 4.20 dd (1H, J4 ,5 a = 2.8, J4 ,5 b = 3.2 Hz, H4 ), 3.89 dd (1H, J5 a,5 b = –12.5 Hz, H5 a), 3.80 dd (1H, H5 b), 1.49 s (3H, C-Me-3 ). Anal. calcd. for C11 H15 N5 O4 : C, 46.97; H, 5.38; N, 24.90; found: C, 46.83; H, 5.29; N, 24.78.
Biologically Active Nucleosides
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9-(2,3-Di-O-acetyl-5-O-benzoyl-3-C-methyl-β-D-ribofuranosyl)-N2 -palmitoylguanine (S.14d). Yield of white amorphous solid 46%. TLC (98:2 v/v CH3 Cl/EtOH): Rf 0.30.1 H NMR (100 MHz, CDCl3 ): 8.96 brs (1H, NH), 7.58 s (1H, H8), 8.06-7.33 m (5H, Bz), 6.03 d (1H, J1 ,2 = 7.5 Hz, H1 ), 5.88 d (1H, H2 ), 5.13 t (1H, J4 ,5 a = J4 ,5 b = 3.5 Hz, H4 ), 4.724.56 m (2H, H5 a,5 b), 2.18 s (3H, Ac), 2.08 s (3H, Ac), 1.27 s (3H, C-Me-3 ), 2.66-0.76 m (31H, palmitoyl). 9-(3-C-Methyl-β-D-ribofuranosyl)guanine (S.15d). Yield of white crystals 70%. TLC (8:2 v/v CH3 Cl/EtOH): Rf 0.17. m.p. 250◦ C (decomposition) (water). [α]20 D –16.0◦ (c 1, DMSO). UV λmax (ε): (pH 1) 255 nm (12000); (pH 13) 263 nm (11000). NMR (200 MHz, DMSO-d6 ): 10.65 brs (1H, NH), 7.87 s (1H, H8), 6.38 brs (1H, NH2 ), 5.66 d (1H, J1 ,2 = 8.0 Hz, H1 ), 5.32 d (1H, JOH,2 = 6.0 Hz, 2 -OH), 5.10 t (1H, JOH,5 = 5.0 Hz, 5 -OH), 4.66 s (1H, 3 -OH), 4.28 d (1H, H2 ), 3.90-3.50 m (3H, H4 ,5 a,5 b), 1.30 s (3H, C-Me-3 ). Anal. calcd. for C11 H15 N5 O5 : C, 44.44; H, 5.09; N, 23.56; found: C, 44.57; H, 5.21; N, 23.52. BASIC PROTOCOL 4
PREPARATION OF 1-(2-C-METHYL-β-D-RIBOFURANOSYL)URACIL 2 -C-Methylribonucleosides (S.17) are prepared in two steps starting from bistrimethylsilyl derivatives of nucleic acid bases and the protected 2-C-methyl-Dribofuranose S.13 as outlined in Figure 14.5.4. As in the synthesis of 3 -Cmethylribonucleosides, the Vorbr¨uggen reaction (described by Vorbr¨uggen and RuhPohlenz, 2001) was used, but with some modifications. Due to steric hindrance from the 2-C-methyl group, only limited conversion to the desired 2 -C-methylribonucleosides is observed under standard conditions (16 hr at room temperature). Raising the temperature and adding catalyst accelerates the reaction, but results in degradation of the starting 2-C-methyl-D-ribofuranose S.13 and lower overall yields (Beigelman et al., 1987). The best overall yields are obtained when condensation is performed with an excess of catalyst for 7 days at room temperature. Several later reports (Harry-O’kuru et al., 1997; Tang et al., 1999) confirmed that synthesis of 2 -C-methylribonucleosides by Vorbr¨uggen condensation from perbenzoylated 2-C-methyl sugar requires excess catalyst and higher temperatures. Cleavage of the benzoyl and acetyl protecting groups is achieved using 5 M ammonia in methanol under mild conditions, and 2 -C-methylribonucleosides (S.17) are purified by crystallization. This protocol describes preparation of 2 -C-methyluridine (S.17a), which is obtained in a 55% overall yield.
Materials
Synthesis of 2 - and 3 -C-Methylribonucleosides
Uracil Anhydrous pyridine 1,1,1,3,3,3-Hexamethyldisilazane (HMDS), reagent grade Anhydrous toluene, reagent grade Anhydrous 1,2-dichloroethane 1,2,3-Tri-O-acetyl-5-O-p-methylbenzoyl-2-C-methyl-β-D-ribofuranose (S.13; see Basic Protocol 2) 2 M Trimethylsilyl trifluoromethanesulfonate (TMSOTf) Chloroform (CHCl3 ), reagent grade Ethanol (EtOH), reagent grade Saturated sodium bicarbonate (NaHCO3 ) Hyflo Super Cel Anhydrous sodium sulfate (Na2 SO4 ) Silica gel: Kieselgel 60 (0.06 to 0.20 mm; Merck) 5 M ammonia in methanol (half saturated at 0◦ C) Acetone Diethyl ether, reagent grade Phosphorus pentoxide (P2 O5 )
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Figure 14.5.4 Synthesis of 2 -C-methylribonucleosides (S.17). The expected overall yields are given in parentheses. DCE, 1,2-dichloroethane; TMS2 B, bis-trimethylsilyl derivatives of heterocyclic bases; TMSOTf, trimethylsilyl trifluoromethanesulfonate.
100- and 250-mL round-bottom flasks Reflux condenser equipped with a CaCl2 protection tube Oil bath with temperature control Rotary evaporator equipped with a water aspirator Silica-coated aluminum TLC plates with fluorescent indicator (Merck silicagel 60 F254 ) 254-nm UV lamp 100-mL funnel with a sintered disc (porosity 3) Vacuum pump for filtration 100- and 250-mL separatory funnels 3 × 20–cm sintered glass chromatography column, porosity 3 Vacuum oil pump Glass filters (porosity 3) Vacuum desiccator Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) and column chromatography (APPENDIX 3E) Biologically Active Nucleosides
14.5.19 Current Protocols in Nucleic Acid Chemistry
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Prepare S.16a 1. Weigh 0.40 g (3.55 mmol) uracil into a 250-mL round-bottom flask, add 5 mL anhydrous pyridine and 10 mL HMDS (b.p. 125◦ C), and attach a condenser equipped with a CaCl2 protection tube. 2. Reflux the mixture in a 130◦ C oil bath until complete dissolution of uracil (3 to 4 hr). 3. Cool the flask to room temperature, remove the condenser, and concentrate the solution to a viscous oil using a rotary evaporator connected to a vacuum system (bath temperature ∼30◦ to 35◦ C). The formed bis-O-trimethylsilyluracil is very hygroscopic.
4. Dissolve the residue in 20 mL dry toluene and concentrate the solution to a viscous oil using a rotary evaporator (bath temperature ∼30◦ to 35◦ C). Remove traces of pyridine by coevaporating two times with 20 mL toluene. 5. Dissolve the residue in 20 mL anhydrous 1,2-dichloroethane, add in one portion a solution of 0.90 g (2.2 mmol) S.13 in 10 mL anhydrous 1,2-dichloroethane, plus 2 mL of 2 M TMSOTf in anhydrous 1,2-dichloroethane. Stopper the flask and keep the solution for 16 hr at room temperature. 6. Monitor reaction by TLC in 98:2 (v/v) CH3 Cl/EtOH. TLC reveals the presence of the starting S.13 (Rf = 0.96). The product S.16a (Rf = 0.48) moves slower in the same solvent.
7. Add an additional 0.75 mL of 2 M TMSOTf in anhydrous 1,2-dichloroethane, stopper the flask, and keep the solution for ∼6 days at room temperature until S.13 disappears completely. 8. When the reaction is complete, add 10 mL sat. NaHCO3 and stir the suspension for 20 min at 0◦ C. 9. Place a 2- to 3-cm layer of Hyflo Super Cel onto a 100-mL funnel with a sintered disc (porosity 3) and wash it with 20 mL chloroform. Filter the suspension through the 100-mL funnel using a vacuum pump and wash the layer with 50 mL chloroform. 10. Separate the organic layer using a 250-mL separatory funnel and wash the organic layer with 20 mL water. 11. Dry the organic layer over ∼10 g Na2 SO4 , filter off the Na2 SO4 by gravity filtration, and wash the precipitate with 20 mL chloroform. Concentrate the combined filtrates to a solid using a rotary evaporator connected to a vacuum system. 12. Prepare a slurry of 30 g silica gel in chloroform and pour it into a 3 × 20–cm chromatography column. 13. Dissolve the obtained residue in a minimal amount of chloroform and layer it carefully on top of the silica gel. Wash the column with 200 mL chloroform and 300 mL of 1% (v/v) ethanol in chloroform, and elute with 2% (v/v) ethanol in chloroform. Collect 25-mL fractions. 14. Evaluate fractions by TLC using 98:2 (v/v) CH3 Cl/EtOH and combine the fractions that contain S.16a only. Synthesis of 2 - and 3 -C-Methylribonucleosides
15. Evaporate the volatile materials from the combined fractions using a rotary evaporator connected to a vacuum system, and dry the residual foam 2 to 3 hr in a vacuum oil pump.
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16. Grind the foam with a stainless steel spatula and dry the resulting powder 12 to 16 hr using a vacuum oil pump. 17. Characterize the product by TLC and NMR. The compound is stable stored for at least 12 months at ambient temperature. 1-(2,3-Di-O-acetyl-5-O-p-methylbenzoyl-2-C-methyl-β-D-ribofuranosyl)uracil (S.16a). Yield of white amorphous solid 0.75 g (74%). TLC (98:2 v/v CH3 Cl/EtOH): Rf 0.49.1 H NMR (100 MHz, CDCl3 ): 8.14 brs (1H, NH), 7.84 d (2H, JH,H = 7.0 Hz, BzMe), 7.34 d (1H, J6,5 = 8.0 Hz, H6), 7.20 d (2H, JH,H = 7.0 Hz, BzMe), 6.21 s (1H, H1 ), 5.54 dd (1H, J5,NH = 2.0 Hz, H5; converted into doublet with J5,6 = 8.0 Hz, on addition of D2 O), 5.29 d (1H, J3 ,4 = 6.5 Hz, H3 ), 4.70-4.30 m (3H, H4 , 5 a, 5 b), 2.40 s (3H, BzMe), 2.09 s (6H, 2Ac), 1.53 s (3H, C-Me-2 ).
Prepare S.17a 18. Weigh 0.40 g (0.87 mmol) S.16a into a 100-mL round-bottom flask and add 12 ml of 5 M ammonia in methanol (half saturated at 0◦ C). Stopper the flask and keep the solution for 48 hr at room temperature. 19. Evaporate all volatile material under reduced pressure using a rotary evaporator. 20. Partition the residue between 10 mL chloroform and 20 mL water using a 100-mL separatory funnel. Separate the aqueous layer and wash it two times with 10 mL chloroform. Evaporate aqueous layer to dryness using a rotary evaporator connected to a vacuum system. 21. Dissolve the residue in a minimal amount of hot water (2 to 3 mL) and keep the mixture for 2 to 3 days at 0◦ C. 22. Collect the precipitate by vacuum filtration on a glass filter (porosity 3), wash with 3 mL acetone and then with 3 mL diethyl ether, and dry in a vacuum desiccator with P2 O5 for 24 hr at room temperature. 23. Characterize the compound by UV, TLC, and 1 H NMR. 1-(2-C-Methyl-β-D-ribofuranosyl)uracil (S.17a). Yield of white crystals 0.162 g (72%). TLC (8:2 v/v isopropanol/conc.NH4 OH/water): Rf 0.68. m.p. 118◦ -119◦ C (water, softening at 101◦ C). [α]20 D +82.0◦ (c 0.7, water). UV λmax (ε): (pH 1-7) 262 nm (10000); (pH 13) 262 nm (7660). 1 H NMR (400 MHz, D2 O): 7.87 d (1H, J6 ,5 = 8.0 Hz, H6), 5.98 s (1H, H1 ), 5.87 d (1H, H5), 4.01 m (2H, H4 , 5 a), 3.86 d (1H, J3 ,4 = 9.3 Hz, H3 a), 3.82 dd (1H, J5 b,4 = 4.2, J5 b,54 a = –13.0 Hz, H5 b), 1.18 s (3H, C-Me-2 ). Anal. calcd. for C10 H14 N2 O6 ×2 H2 O: C, 40.81; H, 6.17; N, 9.52; found: C, 40.63; H, 6.14; N, 9.23.
PREPARATION OF 2-C-METHYL-β-D-RIBOFURANOSYL DERIVATIVES OF CYTOSINE AND ADENINE
ALTERNATE PROTOCOL 2
1-(2-C-Methyl-β-D-ribofuranosyl)cytosine (S.17b) and 9-(2-C-methyl-β-D-ribofuranosyl)adenine (S.17c) are prepared in two steps starting with bis-trimethylsilyl-N-protected nucleobases and S.13 using the steps outlined in Basic Protocol 4 (Fig. 14.5.4). The cytosine derivative is prepared from bis-trimethylsilyl-N4 -benzoylcytosine and is obtained in an overall yield of 49%. The adenine derivative is prepared from bis-trimethylsilyl-N6 benzoyladenine and is obtained in an overall yield of 55%. As for preparation of S.17a, it is essential to use an excess of trimethylsilyl trifluoromethanesulfonate. After Basic Protocol 4, step 20, the residue is crystallized from a minimal amount of water. The reaction of bis-trimethylsilyl-N2 -palmitoylguanine with sugar S.13 under the conditions utilized for 2 -C-methyl-U, C, or A is rather slow at room temperature and results in a complicated mixture. It is possible to isolate the desired 2 -C-methyl-N2 palmitoylguanosine derivative from that mixture with low yield (∼10% to 15%) after
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tedious chromatography. Attempts to increase the reaction temperature result in decomposition of the starting sugar. A more reliable synthesis of the 2 -C-methyl-G derivative was described recently by Li and Piccirilli (2006). This method is based on condensation of a perbenzoylated derivative of S.13 that allows the Vorbr¨uggen condensation to be carried out at higher temperature with an excess of catalyst. 1-(2,3-Di-O-acetyl-5-O-p-methylbenzoyl-2-C-methyl-β-D-ribofuranosyl)cytosine (S.16b). Yield of white amorphous solid 65%. TLC (98:2 v/v CH3 Cl/EtOH): Rf 0.55.1 H NMR (100 MHz, CDCl3 ): 9.02 brs (1H, NH), 7.92-7.12 m (11H, BzMe, Bz, H5,6), 6.46 s (1H, H1 ), 5.32 d (1H, J3 ,4 = 6.2 Hz, H3 ), 4.72-4.40 m (3H, H4 , 5 a, 5 b), 2.42 s (3H, BzMe), 2.12 s (3H, Ac), 2.08 s (3H, Ac),1.50 s (3H, C-Me-2 ). 1-(2-C-Methyl-β-D-ribofuranosyl)cytosine (S.17b). Yield of white crystals 75%. m.p. 235◦ 238◦ C (water). TLC (8:2 v/v isopropanol/conc.NH4 OH/water): Rf 0.68. [α]20 D +132.0◦ (c 0.5, water). UV λmax (ε): (pH 7-13) 273 nm (8800); (pH 1) 281 nm (12500). 1 H NMR (200 MHz, D2 O): 7.77 d (1H, J6 ,5 = 7.6 Hz, H6), 6.00 s (1H, H1 ), 5.98 d (1H, H5), 3.96 m (2H, H3 ,4 ), 3.86 m (2H, H5 a,5 b), 1.14 s, (3H, C-Me-2 ). Anal. calcd. for C10 H15 N3 O5 : C, 46.69;H, 5.88; N, 16.33; found: C,46.64;H, 5.81; N, 16.29. 9-(2,3-Di-O-acetyl-5-O-p-methylbenzoyl-2-C-methyl-β-D-ribofuranosyl)-N6 -benzoyladenine (S.16c). Yield of white amorphous solid 72%. TLC (98:2 v/v CH3 Cl/EtOH): Rf 0.54.1 H NMR (100 MHz, CDCl3 ): 9.08 brs (1H, NH), 8.70 s (1H, H8), 8.08 s (1H, H2), 7.92-7.12 m (7H, BzMe, Bz), 7.18 d (2H, JH,H = 7.2 Hz, BzMe), 6.50 s (1H, H1 ), 5.84 d (1H, J3 ,4 = 6.5 Hz, H3 ), 4.72-4.40 m (3H, H4 ,5 a,5 b), 2.38 s (3H, BzMe), 2.15 s (6H, 2Ac), 1.42 s (3H, C-Me-2 ). 9-(2-C-Methyl-β-D-ribofuranosyl)adenine (S.17c). Yield of white crystals 76%. m.p. 248◦ 250◦ C (water). TLC (8:2 v/v isopropanol/conc.NH4 OH/water): Rf 0.71. [α]20 D –18.0◦ (c 0.8, water). UV λmax (ε): (pH 7-13) 260 nm (15000); (pH 1) 258 nm (14800). 1 H NMR (400 MHz, D2 O): 8.25 s (1H, H8), 8.08 s (1H, H2), 6.00 s (1H, H1 ), 4.08 d (1H, J3 ,4 = 9.1 Hz, H3 ), 4.01 ddd (lH, J4 ,5 a = 2.3, J4 ,5 b = 3.6 Hz, H4 ), 3.93 dd (1H, J5 a,5 b = –13.0 Hz, H5 a), 3.77 dd (1H, H5 b), 0.78 s (3H, C-Me-2 ). Anal. calcd. for C11 H15 N5 O4 : C, 46.97; H, 5.38; N, 24.90; found: C, 46.83; H, 5.27; N, 24.73.
COMMENTARY Background Information
Synthesis of 2 - and 3 -C-Methylribonucleosides
This unit describes details for preparation of 2 - and 3 -C-methylribonucleosides starting from common 2- and 3-C-methylribofuranose precursors. These conformationally restricted analogs of natural ribonucleosides proved to be valuable tools (Mikhailov et al., 1999) for the elucidation of the mechanism of action for several enzymes operating with nucleic acids and their components, including adenosine deaminase (Kalinitchenko et al., 1988), uridine and purine nucleoside phosphorylases (Zinchenko et al., 1987), various nucleases (Mikhailov et al., 1992; Moiseyev et al., 1997), and RNA polymerases (Aivazashvili et al., 1986; Savochkina et al., 1989; Pravdina et al., 1990; Mikhailov et al., 1991; Tunitskaya et al., 1997). Another application, developed by Piccirilli’s group, is an advanced strategy to probe the conformation of specific residues within functional RNA (ribozymes, aptamers) based on a strong preference for 3 -endo conformation in 2 -C-methylribonucleosides (Tang et al., 1999; Ye et al., 2005). Recently, several
groups identified 2 -C-methylribonucleosides as potent inhibitors of hepatitis C virus RNA replication (Eldrup et al., 2004a,b; Pierrra et al., 2005). It was also reported that 3 -Cmethylribonucleosides demonstrate promising anticancer activity (Franchetti et al., 2005). This confirmed the pioneering observation by Walton’s group in the 1960s on the high biological activity of 2 - and 3 -Cmethylribonucleosides (Jenkins et al., 1968; Nutt et al., 1968; Walton et al., 1969). In principle, synthesis of branched-chain nucleosides with 2 - and 3 -C-methyl substitution can be achieved by two approaches—one starting from ribonuclesides and another starting from appropriately protected branchedchain ribofuranoses that are subsequently coupled with a desired nucleic acid base. Both approaches have inherent advantages and disadvantages, and either can be used depending on the research objectives. If only one modified nucleoside is required, then synthesis starting from a ribonucleoside will be shorter with a possibly higher
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overall yield. This is particularly true for 2 -C-methylribonucleosides, where the short and relatively high-yielding synthesis of 2 C-Me-U and 2 -C-Me-C via Grignard addition to a 2 -keto intermediate was developed by Ueda’s group (Matsuda et al., 1990). Similar approaches for synthesis of 3 -Cmethylribonucleosides are less effective due to difficulties in achieving correct stereochemistry at C3 (Huss et al., 1991). The second approach, starting from branched-chain ribofuranoses, is very well developed (Harry-O’kuri et al., 1997; Franchetti et al., 2005; Ye et al., 2005), culminating recently in efficient multi-kilogram synthesis of 2 -C-methyl-7-deaza-adenosine (Bio et al., 2004). Synthesis from sugar precursors is usually longer, but allows preparation of different nucleosides from a common precursor for Vorbr¨uggen condensation. In addition, carbohydrate intermediates are quite often crystalline, thus shortening synthesis time by eliminating chromatographic isolations. Since 2 - and 3 -C-methylribonucleosides can be considered conformational antipodes from the standpoint of sugar pucker (Mikhailov, 1988), syntheses of both classes of target nucleosides from the same precursor have been developed. This approach provides a useful set of analogs for analysis of substrate specificity and conformational requirements for enzymes operating with nucleosides and nucleic acids (reviewed in Mikhailov et al., 1999). The readily available 3-C-methyl-Dallofuranose S.1 is converted to the key precursor for glycosylation, 1,2,3-tri-Oacetyl-5-O-benzoyl-3-C-methyl-α,β-D-ribofuranose (S.6), in five steps with an overall yield of 55% to 60%. All steps are straightforward and provide crystalline intermediates, so that only the last step requires column chromatography. The original publication (Mikhailov et al., 1983) describes two procedures for conversion of the 5-O-benzoyl ribofuranose intermediate S.4 to the triacetate S.6—one based on acetolysis with AcOH/Ac2 O/H2 SO4 and another based on removal of the 1,2-O-isopropylidene group with trifluoroacetic acid followed by acetylation using pyridine/Ac2 O/DMAP. Although the acetolysis reaction is faster and requires fewer manipulations, it was subsequently demonstrated that, under standard acetolysis conditions, even after initial acetylation of 3-OH in S.5, the resulting
3-C-methylribofuranose is a very good substrate for epimerization at C2, thus providing the undesired 3-C-methylarabinofuranose (Beigelman et al., 1988). It is possible to obtain only the desired fully acetylated D-ribofuranose derivative by varying the concentration of Ac2 O during acetolysis, but using the two-step procedure outlined in this unit is recommended to avoid any possibility for formation of undesired 3-C-methylarabinofuranose side products. Subsequent preparation of the 3 -Cmethylribonucleosides is quite reliable and is based on the widely used Vorbr¨uggen procedure (Vorbr¨uggen and Ruh-Pohlenz, 2001). Pyridine/HMDS provides a very reliable and efficient silylating reagent in preparation of bis-trimethylsilyl derivatives of nucleic acid bases required for the above condensation. To maintain a homogeneous reaction, an N6 -benzoyl derivative of adenine and N2 -palmitoyl derivative of guanine are used in the silylation step. These protecting groups are removed during the final ammonolysis step that leads to free nucleosides isolated by crystallization. In general, yields for 3 -C-methylribonucleosides obtained by the Vorbr¨uggen procedure are comparable to those for unmodified ribonucleosides. The lower yields for the guanosine derivative are reflective of N9/N7 regioselectivity under thermodynamic conditions (CF3 SO3 SiMe3 /DCE/reflux). It is possible that better yields can be obtained by using N2 -acetyl-6-O-diphenylcarbamoylguanine, which sterically hinders the N7 position (Robins et al., 1996), or by using higher boiling toluene as a solvent (Li and Piccirilli, 2006). In synthesis of 2 -C-methylribonucleosides, conversion of the same 3-C-methylD-allofuranose (S.1) to the protected 2-Cmethylribofuranose (S.13) follows a strategy introduced by J. Fox’s group for the synthesis of 2-deoxy-2-fluoro-D-arabino nucleosides from D-glucose (Reichman et al., 1975). The overall synthetic route for preparation of 2-Cmethylribofuranose is longer than the preparation of the related 3-C-methylribofuranose. This scheme was optimized to provide high yields in each step, so that the target compound (S.13) is obtained in a 36% overall yield from S.1. The most critical step in this synthesis is the “one-pot” conversion of allofuranose S.9 to ribofuranose S.11, and particularly the removal of the formyl group without effecting
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stability of the 5-O-methylbenzoyl protection. This procedure works very reproducibly but requires some training and attention. Interestingly, the intermediate 2-Cmethylribofuranose derivatives S.11-S.13 are obtained as pure β-anomers, which significantly simplifies the interpretation of corresponding NMR spectra. Removal of the benzyl group from furanose S.11 utilizes homogeneous hydrogenolysis over the Perlman catalyst in ethanol/cyclohexene (Hanessian et al., 1981), although other alternatives could be considered (Bio et al., 2004). When Vorbr¨uggen condensation is applied to 2-C-methylribofuranose, the best results are achieved with prolonged condensation times (7 days, room temperature) and excess triflate catalyst. With fully benzoylated 2-Cmethylribofuranose, more aggressive conditions (SnCl4 , 3 hr refluxing in acetonitrile) were reported with similar results (HarryO’kuru et al., 1997; Li and Piccirilli, 2003). In particular, the conditions described by Li and Piccirilli (2006) were recommended for synthesis of 2 -C-methylguanosine with high yield (78%), high stereoselectivity (β/α >99:1), and impressive regioselectivity (N9/N7 >99:1). However, triacetate S.13 is not very stable during Vorbr¨uggen condensation at such high temperatures. Standard ammonolysis and crystallization are utilized for isolation of the target 2 -C-methylribonucleosides.
Compound Characterization
Synthesis of 2 - and 3 -C-Methylribonucleosides
Physical and spectroscopic data for all products and intermediates have been reported (Mikhailov et al., 1983; Beigelman et al., 1987, 1988). The structure of compounds is supported by NMR spectroscopy, and elemental analysis and optical rotation were used for additional characterization. For 2-Cmethylribofuranoses (S.11-S.13), it was difficult to prove anomeric configuration because of the absence of J1,2 values. However, since J1,2 +J3,4 is constant for similar compounds (Davies, 1978), J3,4 may be used to determine the anomeric configuration. Based on literature reports, J3,4 values of >5 and <3 Hz are typical for β- and α-D-ribofuranoses, respectively (Kam et al., 1979). In the proton spectra of 2-C-furanoses S.11-S.13, the H3 proton was observed as a doublet (J3,4 = 7.2-7.8 Hz), whereas the corresponding signal in the α anomer appears as a broad singlet (Jenkins et al., 1968). Therefore, intermediates S.11S.13 are β anomers.
According to the 1 H NMR spectra, a large shift towards S-conformers occurs in the case of 3 -C-methylribonucleosides (S.15) in comparison with natural nucleosides. Coupling constants J1 ,2 in the range of 7.5 to 8.0 Hz are typical for 3 -Cmethylribonucleosides. In contrast, in the case of 2 -C-methylribonucleosides (S.17), S-N equiblibrium is shifted towards N-conformers with large J3 ,4 = 9.1 to 9.3 Hz coupling constants. In addition to NMR characterization, absolute configuration was confirmed by X-ray for 3 -C-methylcytidine (Mikhailov et al., 1983) and 2 -C-methyluridine (Beigelman et al., 1987), thus validating the original assignments by Walton’s group.
Critical Parameters and Troubleshooting The most critical parameter in each reaction is the purity of intermediates and reagents. It is highly advisable to dry all intermediates over P2 O5 in a desiccator and coevaporate with anhydrous pyridine or toluene before proceeding with the next reaction. Anhydrous solvents are usually critical for reaction success. Solvents should thus be freshly distilled and stored over molecular sieves, or should be taken from a freshly opened bottle of commercial anhydrous solvents. As mentioned above, some transformations require previous training, primarily in analyzing the extent of reactions by TLC. This skill can only be acquired by systematic practice and careful selection of TLC conditions (i.e., solvent development system), but it is essential to obtain optimal yields and avoid unnecessary chromatographic separations. The starting material and reaction mixture are typically compared during TLC evaluation. Approximately 1 optical unit of each starting compound and reaction mixture are placed on the marked baseline at equal distances. For closely moving compounds, co-spotting of both compounds (or reaction mixture) is highly recommended. After the chromatogram is developed in the appropriate solvent system, the end of the solvent run is marked and spots are identified by UV illuminaition (254 nm). In some cases, it may be necessary to develop the chromatogram two times to achieve better separation.
Anticipated Results If the procedures described in this unit are followed closely and without modification, the yields and purity of intermediates should be comparable to those reported here.
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Time Considerations The five-step preparation of the 3-Cmethylribofuranose S.6, including isolation by crystallization in the first three steps, usually takes 8 days. Subsequent preparation of each 3 -C-methylribonucleoside requires 3 days, including column chromatography (3 to 5 hr) and crystallization. The seven-step synthesis of the 2-C-methylribofuranose S.13 requires 7 to 8 days, including chromatography on all but one step (S.9). Synthesis of each 2 -Cmethylribonucleoside requires 10 days due to the long condensation step (1 week). Several operations can be staggered while waiting for compounds to be dried or fully crystallized.
Literature Cited Aivazashvili, V.A., Mikhailov, S.N., Padyukova, N.S., Karpeisky, M.Y., and Bibilashvili, R.S. 1986. 1-(3 -C-Methyl-β-D-ribofuranosyl)uracil 5 -triphosphate, a terminator of RNA synthesis catalyzed by E.coli RNA polymerase. Bioorg. Khim. 12:713-715. Beigelman, L.N., Ermolinsky, B.S., Gurskaya, G.V., Tsapkina, E.N., Karpeisky, M.Y., and Mikhailov, S.N. 1987. New syntheses of 2 -Cmethylnucleosides starting from D-glucose and D-ribose. Carbohydr. Res. 166:219-232. Beigelman, L.N., Gurskaya, G.V., Tsapkina, E.N., and Mikhailov, S.N. 1988. Epimerization during acetolysis of 3-O-acetyl-5-O-benzoyl-1,2isopropylidene-3-C-methyl-α-D-ribofuranose. Synthesis of 3 -C-methylnucleosides with the β-D-ribo and α-D-arabino configurations. Carbohydr. Res. 181:77-88. Bio, M.M., Xu, F., Waters, M., Williams, J.M., Savary, K.S., Cowden, C.J., Yang, C., Buck, E., Song, Z.J., Tschaen, D.M., Volante, R.P., Reamer, R.A., and Grabowski, E.J. 2004. Practical synthesis of a potent hepatatis C virus RNA replication inhibitor. J. Org .Chem. 69:62576266. Davies, D.B. 1978. Conformation of nucleosides and nucleotides. Progr. Nucl. Magn. Reson. Spectrosc. 12:135-225. Eldrup, A.B., Phravc, M., Brooks, J., Bhat, B., Prakash, T.P., Cook, P.D., Wolanski, B., and Olsen, D.B. 2004a. Structure-activity relationship of heterobase-modified 2 -C-methyl ribonucleosides as inhibitors of hepatatis C virus RNA replication. J. Med. Chem. 47:52845297. Eldrup, A.B., Phravc, M., Brooks, J., Bhat, B., Prakash, T.P., Cook, P.D., Wolanski, B., and Olsen, D.B. 2004b. Structure-activity relationship of purine ribonucleosides for inhibition of hepatatis C virus RNA-dependent RNA polymerase. J. Med. Chem. 47:2283-2295. Franchetti, P., Cappellacci, L., Pasqualini, M., Petrelli, R., Vita, P., Jayaram, H.N., Horvath, Z., Szekeres, T., and Grifantini, M. 2005. Antitumor activity of C-methyl-β-D-ribofuranosyladenine
nucleoside ribonucleotide reductase inhibitors. J. Med. Chem. 48:4983-4989. Hanessian, S., Liak, T.J., and Vanasse, B. 1981. Facile cleavage of benzyl ethers by catalytic transfer hydrogenation. Synthesis 5:396-397. Harry-O’kuru, R.E., Smith, J.M., and Wolfe, M.S. 1997. A short, flexible route toward 2 -C-branched ribonucleosides. J. Org. Chem. 62:1754-1759. Huss, S., De las Heras, F.G., and Camarasa, M.J. 1991. Synthesis of 3 -C-ethynylnucleosides of thymine. Tetrahedron 47:1727-1736. Jenkins, S.R., Arison, B., and Walton, E. 1968. Branched chain sugar nucleosides. II. 2 C-Methyladenosine. J. Org. Chem. 33:24902494. Kalinitchenko, E.N., Beigelman, L.N., Mikhailov, S.N., and Mikhailopulo, I.A. 1988. Substrate specificity of adenosine deaminase: Role of methyl groups at 2 -, 3 - and 5 -positions of adenosine. Bioorg. Khim. 14:1157-1161. Kam, B.L., Barascut, L.L, and Imbach, J.L. 1979. A general method of synthesis and isolation, and an NMR spectroscopic study, of tetra-O-acetylD-aldopentofuranose. Carbohydr. Res. 69:135142. Li, N.-S. and Piccirilli, J.A. 2003. Synthesis of the phosphoramidite derivative of 2 -deoxy 2 -C-βmethylcytidine. J. Org. Chem. 68:6799-6802. Li, N.-S. and Piccirilli, J.A. 2006. Efficient synthesis of 2 -C-β-methylguanosine. J. Org. Chem. 71:747-754. Matsuda, A., Itoh, H., Takenuki, K., Sasaki, T., and Ueda, T. 1990. Alkyladdition reaction of pyrimidine 2 -ketonucleosides: Synthesis of 2 branched-chain sugar pyrimidine nucleosides. Chem. Pharm. Bull. 38:2947-2953. Mikhailov, S.N. 1988. Synthesis and properties of C -methylnucleosides and their phosphoric esters. Nucleosides Nucleotides 7:679-682. Mikhailov, S.N., Beigelman, L.N., Gurskaya, G.V., Padyukova, N.S., Yakovlev, G.I., and Karpeisky, M.Y. 1983. Synthesis and properties of 3 -Cmethylnucleosides and their phosphoric esters. Carbohydr. Res. 124:75-96. Mikhailov, S.N., Padyukova, N.S., Lysov, Y.P., Savochkina, L.P., Chidgeavadze, Z.G., and Beabealashvilli, R.S. 1991. Substrate properties of C -methylnucleoside and C -methyl-2 deoxynucleoside 5 -triphosphates in RNA and DNA synthesis reactions catalysed by RNA and DNA polymerases. Nucleosides Nucleotides 10:339-343. Mikhailov, S.N., Oivanen, M., Oksman, P., and Lonnberg, H. 1992. Hydrolysis of 2 - and 3 -Cmethyluridine 2 ,3 -cyclic monophosphates and interconvesion and dephosphorylation of the resulting 2 - and 3 -monophosphates: Comparison with reactions of uridine monophosphates. J. Org. Chem. 57:4122-4126. Mikhailov, S.N., Lysov, Y.P., and Yakovlev, G.I. 1999. Use of functionally competent
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nucleosides and nucleotide analogs in studying enzyme-substrate interactions (review). Mol. Biol. (Russia) 33:393-407. Moiseyev, G.P., Yakovlev, G.I., Lysov, Y.P., Chernyi, A.A., Polyakov, K.M., Oivanen, M., Lonnberg, H., Beigelman, L.N., Efimtseva, E.V., and Mikhailov, S.N. 1997. Determination of the nucleotide conformation in the productive enzyme-substrate complexes of RNAdepolymerases. FEBS Lett. 404:169-172. Nutt, R.F., Dickinson, M.I., Holly, F.M., and Walton, E. 1968. Branched chain sugar nucleosides. III. 3 -C-Methyladenosine. J. Org. Chem. 33:1789-1795. Pierra, C., Benzaria, S., Amador, A., Moussa, A., Mathieu, S., Storer, R., and Gosselin, G. 2005. NM283, an efficient prodrug of potent antiHCV agent 2 -C-methylcytidine. Nucleosides Nucleotides Nucleic Acids 24:767-770. Pravdina, N.F., Mikhailov, S.N., Veselovskaya, T.V., Padyukova, N.S., and Galegov, G.A. 1990. Inhibition of RNA synthesis reactions catalysed by RNA polymerases from flue A virus by some nucleoside 5 -triphosphate analogs. Mol. Gen. Mikrobiol. Virusol. 22-25. Reichman, U., Watanabe, K.A., and Fox, J.J. 1975. A practical synthesis of 2-deoxy-2-fluoro-Darabinofuranose derivatives. Carbohydr. Res. 42:233-240. Robins, M.J., Zou, R., Guo, Z., and Wnuk, S.F. 1996. Nucleic acid related compounds. 93. A solution for the historic problem of regioselective sugar-base coupling to produce 9glycosylguanines or 7-glycosylguanines. J. Org. Chem. 61:9207-9212. Rosenthal, A. and Mikhailov, S.N. 1980. Branchedchain sugar nucleosides. Synthesis of 3 -C-ethyl (and 3 -butyl) uridine. Carbohydr. Res. 79:235242. Savochkina, L.P., Sviriyeva, T.V., Beigelman, L.N., Padyukova, N.S., Kuznetsov, D.A., Lysov, Y.P., Mikhailov, S.N., and Beabealashvilli, R.S. 1989. Substrate properties of C methylnucleoside triphosphates in RNA synthesis reactions catalysed by Escherichia coli RNA polymerase. Mol. Biol. 23:1700-1710.
Tang, X.Q., Liao, X., and Piccirilli, J.A. 1999. 2 -C-Branched ribonucleosides: Synthesis of the phosphoramidite derivatives of 2 -C-βmethylcytidine and their incorporation into oligonucleotides. J. Org. Chem. 64:747-754. Tunitskaya, V.L., Rusakova, E.E., Padyukova, N.S., Ermolinsky, B.S., Chernyi, A.A., Kochetkov, S.N., Lysov, Y.P., and Mikhailov, S.N. 1997. Substrate properties of C -methyl UTP derivatives in T7 RNA polymerase reactions. Evidence for N-type NTP conformation. FEBS Lett. 400:263-266. Vorbr¨uggen, H. and Ruh-Pohlenz, C. 2001. Handbook of Nucleoside Synthesis. John Wiley & Sons, Hoboken, N.J. Wagner, S., Verheyden, J.P.H., and Moffat, J.G. 1974. Preparation and synthetic utility of some organotin derivatives of nucleosides. J. Org. Chem. 39:24-30. Walton, E., Jenkins, S.R., Nutt, R.F., and Holly, F.M. 1969. Branched chain sugar nucleosides. V. Synthesis and antiviral properties of several branched-chain sugar nucleosides. J. Med. Chem. 12:306-309. Ye, J.-D., Liao, X., and Piccirilli, J.A. 2005. Synthesis of 2 -C-difluoromethylribonucleosides and their enzymatic incorporation into oligoribonucleotides. J. Org. Chem. 70:7902-7910. Zinchenko, A.I., Barai, V.N., Erosheskaya, L.A., Beigelman, L.N., Mikhailov, S.N., Karpeisky, M.Y., and Mikhailopulo, I.A. 1987. 2 -, 3 - and 5 -Methylderivatives of uridine in reactions of microbiological transglycosylation. Proc. USSR Acad. Sci. 297:731-734.
Contributed by Leonid Beigelman InterMune Brisbane, California Sergey N. Mikhailov Engelhardt Institute of Molecular Biology Russian Academy of Sciences Moscow, Russia
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CHAPTER 15 Nucleoside Prodrugs and Delivery Strategies INTRODUCTION prodrug is a derative of a biologically active compound that will be converted in vivo to the parent drug by chemical or enzymatic reactions. From this point of view, every modified nucleoside that needs to be converted into its monophosphate or triphosphate could be considered a prodrug. However, we use the name “prodrug” in a broader sense to describe derivatives of the modified nucleosides and nucleotides themselves. Development of a prodrug is essentially a rescue operation designed to circumvent some fundamental drawback of a selected drug. The activity of a compound is highly dependent on the form and the formulation in which the compound is presented. In the nucleoside and nucleotide field, prodrugs are developed, for example, to increase the solubility and bioavailability of the active compound.
A
In addition to the classical approach of making esters as prodrugs, the development of nucleoside and nucleotide prodrugs can benefit from our understanding of the different enzymes involved in nucleoside and nucleotide metabolism. One of the oldest examples in the nucleoside field is the use of the 5 -O-monophosphate of ara-A (Fig. 15.0.1). Ara-A has a very low solubility in water, and administration of ara-A by infusion requires large volumes. The greater solubility of the monophosphate of ara-A permits the use of a smaller infusion volume. Another enzyme that has proved valuable in prodrug design is xanthine oxidase. Guanine nucleosides are very insoluble in water because of the strong intermolecular associations through hydrogen bonding and π-stacking between the base moieties. As a consequence, drugs such as acyclovir are quite insoluble in body fluids. The 6-deoxy analog of acyclovir (desiclovir; Fig. 15.0.1) is more soluble and is metabolized in vivo by xanthine oxidase to acyclovir. When administered orally, 6-deoxy-acyclovir gives the same blood levels of acyclovir as those obtained with intravenous acyclovir. An analogous approach has also proved successful for penciclovir. At 1 hr after oral administration, the diacetyl derivative of 6-deoxy-penciclovir gives peak plasma concentrations that are ten-fold higher than those detected following an equivalent oral dose of penciclovir. This prodrug has been given the name of famciclovir. The potential therapeutic armamentarium of acyclic anti-HSV and anti-VZV nucleosides has been extended with the L-valyl ester of acyclovir (valacyclovir). This drug is a prodrug of acyclovir and has been developed because of the low oral bioavailability of the parent compound. The same approach has been used to improve the oral bioavailability of L-dC, an anti-HBV agent. Abacavir (Fig. 15.0.2) is an example of a prodrug from a carbocyclic purine nucleoside. Abacavir is intracellularly activated to carbovir triphosphate, which is active against HIV infections. This compound is described in UNIT 14.4. Classical examples of prodrugs in the anti-cancer field are to be found in the many fluorouracil prodrugs, of which tegafur (Fig. 15.0.2) is the best known. Most nucleosides need to be phosphorylated intracellularly to exert their biological effect. An advantage to using phosphorylated nucleosides as therapeutic agents is that they are Nucleoside Prodrugs and Delivery Strategies Contributed by Piet Herdewijn Current Protocols in Nucleic Acid Chemistry (2007) 15.0.1-15.0.4 C 2007 by John Wiley & Sons, Inc. Copyright
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Figure 15.0.1
Anti-HSV prodrugs based on enzymatic hydrolysis and oxidation reactions.
Figure 15.0.2 (tegafur).
An example of an anti-HIV prodrug (abacavir) and an antitumoral prodrug
active in cases where intracellular phosphorylation is difficult (i.e., when the modified nucleoside is a poor substrate for cellular and/or viral kinases). However, the use of phosphorylated nucleosides is hampered by the fact that nucleoside monophosphates are not taken up by cells and that they are easily dephosphorylated by esterases to give the parent nucleoside. Because the first phosphorylation by nucleoside kinase is the rate-limiting step in the metabolic activation of most modified nucleosides, several prodrug strategies have been designed to bypass the nucleoside kinase step. These prodrugs can penetrate cells and deliver the nucleoside 5 -O-monophosphate intracellularly. Striking examples of this group of pronucleotides are phosphoramidate pronucleotides, cycloSal Introduction
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Figure 15.0.3
Prodrug approaches to deliver monophosphates into cells.
pronucleotides, SATE pronucleotides, lipid ester prodrugs, and amino acid pronucleotides (Fig. 15.0.3). A logical pursuit of this approach is the development of nucleoside phosphonate derivatives. This approach has been very successful in the field of acyclic nucleosides. The poor bioavailability of these compounds has been circumvented by synthesizing phosphonate prodrugs, which can be given orally. Adefovir dipivoxil (for the treatment of HBV infections) and tenofovir disoxoproxil (for the treatment of HIV infections; Fig. 15.0.4) can be administered on a once-a-day dosing schedule. These compounds are described in UNIT 14.2. To open this chapter, UNIT 15.1 describes the synthesis of amino acid pronucleotides. UNIT 15.2 describes the synthesis of hexadecyloxypropyl esters of the phosphonate nucleosides HPMPC and HPMPA (for synthesis of the biologically active nucleosides, see UNIT 14.2). UNIT 15.3 describes the bis(SATE) prodrug approach, which is based on a combined enzymatic-chemical degradation pathway. One of the main issues with such
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Figure 15.0.4
Examples of phosphonate prodrugs to increase bioavailability.
prodrug approaches is the low stability of the prodrugs in vivo. This seems to be largely overcome in the phosphonate series, as the described prodrugs are readily absorbed and are orally active in mammals. The development of prodrugs from modified nucleosides and nucleotides is an expanding field of research that has recently been extended to nucleic acid prodrugs. This last concept is very complicated because of the multiple degradation reactions that are involved in liberating the original nucleic acid. Uptake enhancement for oligonucleotides can be achieved by conjugate chemistry, as demonstrated for peptide-oligonucleotide conjugates (e.g., UNIT 4.28) and cholesterol-oligonucleotide conjugates. The field of nucleic acid prodrugs is promising because cellular uptake is (and will remain) a bottleneck in the development of oligonucleotide therapeutics. Piet Herdewijn
Introduction
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Synthesis of Amino Acid Phosphoramidate Monoesters via H-Phosphonate Intermediates
UNIT 15.1
This unit describes a method for generating nucleoside phosphoramidate monoesters (S.3 and S.4; Fig. 15.1.1) from H-phosphonate intermediates (S.2). Two different methods are outlined for generating H-phosphonate monoesters. Both methods are simple and utilize readily available phosphitylating reagents. The nucleoside H-phosphonate monoesters are subsequently oxidized to the nucleoside phosphoramidates in the presence of iodine and the appropriate amino acid methyl ester. Phosphitylation. The Basic Protocol utilizes diphenyl phosphite as the phosphitylating agent. Diphenyl phosphite is inexpensive, is easy to handle, and produces high yields of the H-phosphonate monoester (S.2). This method involves reaction of diphenyl phosphite with a nucleoside followed by a basic work-up to yield the nucleoside H-phosphonate monoester. The Alternate Protocol utilizes bis(N,N-diisopropylamino)chlorophosphine as the phosphitylating agent. Bis(N,N-diisopropylamino)chlorophosphine is more reactive than diphenyl phosphite, is relatively inexpensive and easy to handle, and produces high yields of the H-phosphonate. Nucleosides are reacted with bis(N,N-diisopropylamino)chlorophosphine to generate a nucleoside bis(N,N-diisopropylamino)phosphine intermediate, followed by an acidic work-up to yield the nucleoside H-phosphonate monoesters.
Figure 15.1.1 Formation of AZT amino acid phosphoramidates. R = amino acid; (a) glycine; (b) L-alanine; (c) L-valine; (d) L-leucine; (e) L-tyrosine; (f) L-phenylalanine; (g) L-tryptophan; (h) D-phenylalanine; (i) D-tryptophan. Nucleoside Prodrugs and Delivery Strategies Contributed by Cindy J. Choy, Dan P. Drontle, and Carston R. Wagner Current Protocols in Nucleic Acid Chemistry (2006) 15.1.1-15.1.12 C 2006 by John Wiley & Sons, Inc. Copyright
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Oxidation. Nucleoside H-phosphonate monoesters are activated with trimethylsilyl chloride (TMSCl), to generate a bis-silylphosphite intermediate, which can be oxidized with iodine in pyridine. Addition of the amino acid methyl ester and triethylamine (TEA) yields the desired phosphoramidate. Amidation. Treatment of the phosphoramidate methyl ester with methylamine in methanol generates the corresponding phosphoramidate methyl amide. CAUTION: Carry out experimental procedures under an inert atmosphere, with dried glassware and anhydrous reagents. BASIC PROTOCOL
GENERATION OF AZT H-PHOSPHONATE WITH DIPHENYL PHOSPHITE AND ITS CONVERSION TO AZT AMINO ACID PHOSPHORAMIDATE METHYL AMIDES This methodology, originally developed by Stawinski and co-workers, utilizes diphenyl phosphite as an inexpensive phosphitylating agent (Jankowska et al., 1994) with minor modifications (Iyer et al., 2000). The resultant H-phosphonate monoester is oxidized using iodine and trimethylsilyl chloride (Kers and Kraszewski, 1998) and reacted with amino acid methyl esters. Finally, the phosphoramidate amino acid methyl esters are converted to the methyl amide using concentrated methylamine in methanol. Although this methodology yields phosphoramidate monoesters in high yield and is compatible with a variety of amine and nucleoside substrates, it does have limitations. The most significant limitation is the formation of significant amounts (5% to 15%) of phenyl H-phosphonate monoester during the phosphitylation step. One solution is to use the Alternate Protocol; for another solution, see Critical Parameters and Troubleshooting.
Materials
Synthesis of Amino Acid Phosphoramidate Monoesters
3 -Azido-3 -deoxythymidine (S.1; AZT; Toronto Research Chemicals) Argon (or nitrogen) gas Anhydrous pyridine (<50 ppm water; Acros) Diphenyl phosphite (85% pure; Sigma-Aldrich) Triethylamine (TEA, 99.5% pure; Sigma-Aldrich) Dichloromethane (CH2 Cl2 ), ACS reagent grade Silica gel 60 (230 to 400 mesh; EMD) Chloroform (CHCl3 ), ACS reagent grade Methanol (MeOH), ACS reagent grade Ammonium hydroxide (NH4 OH) Amino acid methyl ester hydrochloride for conversion (select one): Glycine methyl ester hydrochloride (a) L-Alanine methyl ester hydrochloride (b) L-Valine methyl ester hydrochloride (c) L-Leucine methyl ester hydrochloride (d) L-Tyrosine methyl ester hydrochloride (e) L-Phenylalanine methyl ester hydrochloride (f) L-Tryptophan methyl ester hydrochloride (g) D-Phenylalanine methyl ester hydrochloride (h) D-Tryptophan methyl ester hydrochloride (i) Trimethylsilyl chloride (TMSCl, 99% pure; Sigma-Aldrich) Iodine, sublimed Amberlite IRP 64 (100 to 400 wet mesh; Sigma-Aldrich) Methylamine solution (see recipe) Vacuum pump 10- and 50-mL pear-shaped flasks
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Cannula (Teflon tubing) Rotary evaporator equipped with a water aspirator 5 × 17–, 2 × 6–, 3 × 17–, and 1.8 × 8–cm chromatography columns TLC plates: silica-coated aluminum plates with fluorescent indicator (Merck silica gel 60 F254 ) 254-nm UV lamp 50-mL round-bottom flasks Capped vial Lyophilizer Additional reagents and equipment for column chromatography (APPENDIX 3E) and TLC (APPENDIX 3D) Prepare 5 -AZT-H-phosphonate (TEA salt) 1. Dry 755 mg (2.83 mmol) AZT (S.1) for at least 8 hr in a clean, dry 50-mL pear-shaped flask on a vacuum pump. Dry all other glassware in a 125◦ C oven. Items that cannot be placed in the oven should be desiccated.
2. Purge inert gas lines with argon for 5 min prior to starting reaction. 3. Attach a clean, dry 50-mL round-bottom flask with a magnetic stir bar to argon line and flush with argon for 5 min. Add 8 mL anhydrous pyridine to the flask, then add 541 µL (2.83 mmol) diphenyl phosphite. 4. Dissolve AZT in 7 mL anhydrous pyridine. Attach to argon line and flush with argon for several minutes. 5. Transfer AZT slowly into diphenyl phosphite via a cannula over a period of ∼40 min. Once this transfer is complete, let reaction stir 2 hr under argon at room temperature. The rate of transfer can be conveniently controlled if a syringe is used to create a vacuum in the flask containing diphenyl phosphite.
6. After 2 hr, treat reaction with 3 mL water and 3 mL TEA for 15 min. 7. Concentrate the reaction mixture on a rotary evaporator to a semi-viscous oil, then dissolve in 70 mL water and extract four times with 100 mL dichloromethane. Evaporate the aqueous layer on a rotary evaporator to form a viscous oil. 8. Pack a 5 × 17–cm silica gel column in 9:1 (v/v) CHCl3 /MeOH (APPENDIX 3E). Load the sample onto the column and elute the first 200 mL with the above solvent system, then switch to 5:2:0.25 (v/v/v) CHCl3 /MeOH/water with 0.5% NH4 OH to elute desired product. 9. Combine fractions containing the product, as determined by TLC (APPENDIX 3D) using the elution solvent and visualization with UV light (Rf = 0.2). Concentrate on a rotary evaporator. To minimize contamination with phenyl phosphite monoester, the first two to three UVactive fractions should be concentrated in a separate flask and checked by NMR.
10. Dry the resulting clear glass solid overnight in vacuo (in a vacuum pump at 0.05 to 0.5 Torr). The compound is very hygroscopic, necessitating excessive drying.
11. Characterize the compound by TLC, 1 H NMR, and 31 P NMR. ◦
This compound can be stored in a capped vial for several years at 4 C without significant decomposition.
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Triethylammonium 3 -azido-3 -deoxythymidine 5 -phosphite (S.2). Yield of colorless solid: 840 mg (69%). 1 H NMR (D2 O, 300 MHz): 7.522 (d, J = 1.1 Hz, 1H), 6.604 (d, J = 638 Hz, 1H), 6.090 (t, J = 6.7 Hz, 1H), 4.326 (m, 1H), 4.009 (m, 1H), 3.933 (m, 2H), 3.023 (q, J = 7.3 Hz, 6H), 2.328 (m, 2H), 1.736 (d, J = 1.1 Hz, 3H), 1.100 (t, J = 7.3 Hz, 9H). 31 P NMR (D2 O, 121 MHz): 6.856.
Oxidize AZT H-phosphonate and condense with amino acid 12. Dry 190 mg (0.439 mmol) S.2 at least 6 hr in vacuo (0.05 to 0.5 Torr) in a 50-mL round-bottom flask. 13. Dry 0.878 mmol (2 eq) amino acid methyl ester hydrochloride in a separate 50-mL pear-shaped flask at least 6 hr in vacuo (0.05 to 0.5 Torr). For S.3a: For S.3b: For S.3c: For S.3d: For S.3e: For S.3f: For S.3g: For S.3h: For S.3i:
110 mg glycine methyl ester hydrochloride 121 mg L-alanine methyl ester hydrochloride 148 mg L-valine methyl ester hydrochloride 160 mg L-leucine methyl ester hydrochloride 173 mg L-tyrosine methyl ester hydrochloride 189 mg L-phenylalanine methyl ester hydrochloride 223 mg L-tryptophan methyl ester hydrochloride 189 mg D-phenylalanine methyl ester hydrochloride 223 mg D-tryptophan methyl ester hydrochloride.
14. Purge inert gas lines with argon for 5 min prior to starting reaction. 15. Attach stoppered flask containing AZT H-phosphonate and a magnetic stir bar to argon line. Dissolve contents in 7 mL dry pyridine. 16. Add 167 µL (1.32 mmol) TMSCl to reaction mixture. 17. Attach a clean, dry 10-mL pear-shaped flask with 167 mg (0.659 mmol) iodine to argon line and dissolve in 3 mL dry pyridine. Since iodine should be carefully titrated into the reaction, using <3 mL pyridine is not recommended. However, using >5 mL causes unnecessary dilution.
18. Under vigorous stirring, use a cannula to transfer the iodine solution (∼1 equiv.) dropwise into the flask containing AZT H-phosphonate. Stop once the solution turns reddish-brown. Transfer of iodine should be carried out as slowly as possible to prevent overtitration with iodine. The rate of transfer can be conveniently controlled if a syringe is used to create a vacuum in the flask containing AZT H-phosphonate.
19. Allow reaction to stir under argon for 15 min. 20. Add the amino acid methyl ester hydrochloride and 0.43 mL (3.08 mmol) TEA to the reaction and let stir for 30 min under argon. 21. Evaporate the pyridine on a rotary evaporator to form a reddish-orange solid. 22. Dissolve solid in 50 mL of 1 N NH4 OH and extract four times with 10 mL CHCl3 . 23. Concentrate aqueous layer on a rotary evaporator to obtain an orange solid. 24. Prepare a 2 × 6 – cm amberlite IRP 64 column in water. Dissolve crude phosphoramidate monoester in ∼3 mL water and add to the top of the column. Elute with water. Synthesis of Amino Acid Phosphoramidate Monoesters
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Amberlite IRP 64 contains resin fines that can clog the column. To remove resin fines, suspend amberlite in water and allow it to settle for several minutes, then decant any amberlite suspended in the water. Repeat procedure until water is free of amberlite. After removing resin fines, amberlite can be stored for several years in water on the bench. Current Protocols in Nucleic Acid Chemistry
25. Combine fractions containing product, as determined by TLC and visualized with UV light. 26. Pack a 3 × 17 – cm silica gel column using 5:2:0.25 (v/v/v) CHCl3 /MeOH/water with 0.5% NH4 OH. Load crude phosphoramidate onto the column and elute with the same solvent. 27. Combine fractions containing the product, as determined by TLC using the elution solvent and visualization with UV light (Rf = 0.18). Evaporate on a rotary evaporator. 28. Characterize compound S.3a-i by 1 H and 31 P NMR. These derivatives can be stored in a capped vial for several years at 4◦ C without significant decomposition. 2-[3 -Deoxy-3 -azidothymidyl(hydroxy)phosphorylamino]acetic acid methyl ester (S.3a). Yield of slightly pink solid: 94 mg (49%). 1 H NMR (D2 O, 300 MHz): 7.550 (d, J = 1 Hz, 1H), 6.091 (t, J = 6.7 Hz, 1H), 4.311 (m, 1H), 3.978 (m, 1H), 3.935-3.787 (m, 2H), 3.505 (s, 3H), 3.465 (d, J = 11.5 Hz, 2H), 2.354-2.303 (m, 2H), 1.753 (d, J = 1 Hz, 3H). 31 P NMR (D2 O, 121 MHz): 7.855. 2-(S)-[3 -Deoxy-3 -azidothymidyl(hydroxy)phosphorylamino]proprionic acid methyl ester (S.3b). Yield of white solid: 79 mg (48%). 1 H NMR (D2 O, 300 MHz): 7.557 (d, J = 1.1 Hz, 1H), 6.085 (t, J = 6.7 Hz, 1H), 4.298 (m, 1H), 3.993 (m, 1H), 3.898-3.776 (m, 2H), 3.620 (m, 1H), 3.536 (s, 3H), 2.331 (m, 2H), 1.760 (d, J = 1.1 Hz, 3H), 1.141 (d, J = 6.9 Hz, 3H). 31 P NMR (D2 O, 121 MHz): 6.626. -(S)-[3 -Deoxy-3 -azidothymidyl(hydroxy)phosphorylamino]-3-methylbutyric acid methyl ester (S.3c). Yield of white solid: 154 mg (68%). 1 H NMR (D2 O, 300 MHz): 7.573 (d, J = 1.1 Hz, 1H), 6.077 (t, J = 6.7 Hz, 1H), 4.293 (m, 1H), 3.994 (m, 1H), 3.907-3.752 (m, 2H), 3.550 (s, 3H), 3.338 (m, 1H), 2.328 (m, 2H), 1.778 (d, J = 1.1 Hz, 3H), 1.772 (m, 1H), 0.725 (d, J = 6.8 Hz, 3H), 0.715 (d, J = 6.8 Hz, 3H). 31 P NMR (D2 O, 121 MHz): 7.166. 2-(S)-[3 -Deoxy-3 -azidothymidyl(hydroxy)phosphorylamino]-4-methylvaleric acid methyl ester (S.3d). Yield of white solid: 98 mg (31%). 1 H NMR (D2 O, 300 MHz): 7.600 (d, J = 1.1 Hz, 1H), 6.086 (t, J = 6.8 Hz, 1H), 4.303 (m, 1H), 3.987 (m, 1H), 3.895-3.747 (m, 2H), 3.538 (m, 1H), 3.538 (s, 3H), 2.324 (m, 2H), 1.771 (d, J = 1.1 Hz, 3H), 1.485 (m, 1H), 1.335-1.278 (m, 2H), 0.672 (d, J = 6.6 Hz, 6H). 31 P NMR (D2 O, 121 MHz): 6.701. 2-(S)-[3 -Deoxy-3 -azidothymidyl(hydroxy)phosphorylamino]-3-(4-hydroxyphenyl)proprionic acid methyl ester (S.3e). Yield of white solid: 149 mg (49%). 1 H NMR (D2 O, 300 MHz): 7.401 (s, 1H), 6.859 (d, J = 7.3 Hz, 2H), 6.547 (d, J = 8.2 Hz, 2H), 6.000 (t, J = 6.6 Hz, 1H), 4.119 (m, 1H), 3.845 (m, 1H), 3.647 (m, 1H), 3.620-3.540 (m, 2H), 3.486 (s, 3H), 2.718 (dd, J = 6.2, 13.5 Hz, 1H), 2.591 (dd, J = 7.7, 13.5 Hz, 1H), 2.230 (m, 1H), 2.105 (m, 1H), 1.662 (s, 3H). 31 P NMR (D2 O, 121 MHz): 6.487. 2-(S)-[3 -Deoxy-3 -azidothymidyl(hydroxy)phosphorylamino]-3-phenylproprionic acid methyl ester (S.3f). Yield of white solid: 216 mg (64%). 1 H NMR (D2 O, 300 MHz): 7.424 (s, 1H), 7.120-6.974 (m, 5H), 5.998 (t, J = 6.7 Hz, 1H), 4.153 (m, 1H), 3.843 (m, 1H), 3.728 (m, 1H), 3.585 (m, 2H), 3.491 (s, 3H), 2.794 (m, 1H), 2.699 (m, 1H), 2.219 (m, 1H), 2.116 (m, 1H), 1.675 (s, 3H), 1.141 (d, J = 6.9 Hz, 3H). 31 P NMR (D2 O, 121 MHz): 6.370. 2-(S)-[Hydroxy-(3 -deoxy-3 -azidothymidyl)phosphorylamino]-3-(3-indolyl)proprionic acid methyl ester (S.3g). Yield of white solid: 198 mg (70%). 1 H NMR (D2 O, 300 MHz): 7.222 (d, J = 7.9 Hz, 1H), 7.130 (d, J = 8.1 Hz, 1H), 7.072 (s, 1H), 6.880 (s, 1H), 6.857 (t, J = 7.7, 1 Hz, 1H), 6.729 (t, J = 7.5 Hz, 1H), 5.690 (t, J = 6.6 Hz, 1H), 3.883 (m, 1H), 3.763 (m, 1H), 3.635 (m, 1H), 3.527 (m, 2H), 3.448 (s, 3H), 2.899 (m, 1H), 2.765 (m, 1H), 1.857 (m, 1H), 1.682 (m, 1H), 1.493 (s, 3H). 31 P NMR (D2 O, 121 MHz): 6.412. 2-(R)-[3 -Deoxy-3 -azidothymidyl(hydroxy)phosphorylamino]-3-phenylproprionic acid methyl ester (S.3h). Yield of white solid: 282 mg (68%). 1 H NMR (D2 O, 300 MHz): 7.415 (s, 1H), 7.158-6.998 (m, 5H), 5.995 (t, J = 6.8 Hz, 1H), 4.118 (m, 1H), 3.808 (m, 1H), 3.738 (m, 1H), 3.661 (m, 1H), 3.467 (m, 1H), 3.403 (s, 3H), 2.754 (m, 2H), 2.212 (m, 2H), 1.679 (s, 3H), 1.141 (d, J = 6.9 Hz, 3H). 31 P NMR (D2 O, 121 MHz): 6.204.
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2-(R)-[Hydroxy-(3 -deoxy-3 -azidothymidyl)phosphorylamino]-3-(3-indolyl)proprionic acid methyl ester (S.3i). Yield of white solid: 256 mg (63%). 1 H NMR (D2 O, 300 MHz): 7.299 (d, J = 7.9 Hz, 1H), 7.182 (d, J = 8.1 Hz, 1H), 7.141 (s, 1H), 6.939 (s, 1H), 6.930 (dd, J = 7.0, 8.1, 1 Hz, 1H), 6.816 (t, J = 7.0, 7.9 Hz, 1H), 5.773 (t, J = 6.8 Hz, 1H), 3.851-3.741 (m, 2H), 3.691 (m, 1H), 3.605 (m, 1H), 3.357 (s, 3H), 3.332 (m, 1H), 2.882 (m, 2H), 2.002 (m, 1H), 1.815 (m, 1H), 1.504 (s, 3H). 31 P NMR (D2 O, 121 MHz): 6.396.
Convert to methyl amide 29. Dissolve 0.1 mmol S.3a-i in 2 mL of ∼10 M methylamine solution in a capped vial for 5 days at room temperature. Do not use a rubber septum because it will decompose under these conditions.
30. Concentrate reaction mixture on a rotary evaporator. 31. Pack a 1.8 × 8–cm silica gel column with 5:2:0.25 (v/v/v) CHCl3 /MeOH/water with 0.5% NH4 OH. Load crude phosphoramidate onto the column and elute with the same solvent. 32. Monitor fractions by TLC using the same solvent and UV visualization (Rf = 0.17). Combine fractions containing a single UV spot and evaporate on a rotary evaporator. 33. Lyophilize S.4a-i to yield a white, fluffy solid. 34. Characterize this compound by 1 H and 31 P NMR. The yields of this reaction vary from 65% to 100%. These derivatives can be stored in a capped vial for several years at 4◦ C without significant decomposition. (1-(S)-Methylcarbamoylmethyl)phosphoramidic acid mono-(3 -deoxy-3 -azidothymidyl)ester (S.4a). Yield of white solid: 32 mg (92%). 1 H NMR (D2 O, 300 MHz): 7.538 (d, J = 1.1 Hz, 1H), 6.089 (t, J = 6.7 Hz, 1H), 4.302 (m, 1H), 3.993 (m, 1H), 3.920-3.785 (m, 2H), 3.305 (d, J = 11.2 Hz, 2H), 2.571 (s, 3H), 2.323 (m, 2H), 1.735 (d, J = 1.1 Hz, 3H). 31 P NMR (D2 O, 121 MHz): 7.956. (1-(S)-Methylcarbamoylethyl)phosphoramidic acid mono-(3 -deoxy-3 -azidothymidyl)ester (S.4b). Yield of white solid: 41 mg (84%). 1 H NMR (D2 O, 300 MHz): 7.5487 (d, J = 1.1 Hz, 1H), 6.083 (t, J = 6.8 Hz, 1H), 4.283 (m, 1H), 3.988 (m, 1H), 3.889-3.757 (m, 2H), 3.440 (m, 1H), 2.566 (s, 3H), 2.313 (m, 2H), 1.733 (d, J = 1.1 Hz, 3H), 1.120 (d, J = 7.1 Hz, 3H). 31 P NMR (D2 O, 121 MHz): 6.685. (2-Methyl-1-(S)-methylcarbamoylpropyl)phosphoramidic acid mono-(3 -deoxy-3 azidothymidyl)ester (S.4c). Yield of white solid: 40 mg (65%). 1 H NMR (D2 O, 300 MHz): 7.567 (d, J = 1.1 Hz, 1H), 6.082 (t, J = 6.8 Hz, 1H), 4.257 (m, 1H), 3.999 (m, 1H), 3.895-3.738 (m, 2H), 3.220 (m, 1H), 2.583 (s, 3H), 2.318 (m, 2H), 1.828 (m, 1H), 1.762 (d, J = 1.1 Hz, 3H), 0.759 (d, J = 6.8 Hz, 3H), 0.692 (d, J = 6.8 Hz, 3H). 31 P NMR (D2 O, 121 MHz): 6.851. (3-Methyl-1-(S)-methylcarbamoylbutyl)phosphoramidic acid mono-(3 -deoxy-3 azidothymidyl)ester (S.4d). Yield of white solid: 25 mg (66%). 1 H NMR (D2 O, 300 MHz): 7.588 (d, J = 1.1 Hz, 1H), 6.081 (t, J = 6.8 Hz, 1H), 4.275 (m, 1H), 3.995 (m, 1H), 3.896-3.746 (m, 2H), 3.403 (m, 1H), 2.570 (s, 3H), 2.320 (m, 2H), 1.753 (d, J = 1.1 Hz, 3H), 1.504 (m, 1H), 1.352-1.225 (m, 2H), 0.693 (d, J = 6.6 Hz, 6H). 31 P NMR (D2 O, 121 MHz): 6.359.
Synthesis of Amino Acid Phosphoramidate Monoesters
[2-(4-Hydroxyphenyl)-1-(S)-methylcarbamoylethyl]phosphoramidic acid mono-(3 deoxy-3 -azidothymidyl)ester (S.4e). Yield of white solid: 36 mg (71%). 1 H NMR (D2 O, 300 MHz): 7.401 (s, 1H), 6.932 (d, J = 8.6 Hz, 2H), 6.581 (d, J = 8.6 Hz, 2H), 5.987 (t, J = 6.6 Hz, 1H), 4.040 (m, 1H), 3.807 (m, 1H), 3.550 (m, 1H), 3.483-3.400 (m, 2H), 2.799 (m, 1H), 2.542 (2, 3H), 2.534 (m, 1H), 2.240 (m, 1H), 2.093 (m, 1H), 1.696 (d, J = 1.0 Hz, 3H). 31 P NMR (D2 O, 121 MHz): 5.761.
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(2-Phenyl-1-(S)-methylcarbamoylethyl)phosphoramidic acid mono-(3 -deoxy-3 azidothymidyl)ester (S.4f). Yield of white solid: 67 mg (99%). 1 H NMR (D2 O, 300 MHz): 7.419 (d, J = 1.0 Hz, 1H), 7.161-7.047 (m, 5H), 5.978 (t, J = 6.7 Hz, 1H), 4.083 (m, 1H), 3.812 (m, 1H), 3.629 (m, 1H), 3.504-3.365 (m, 2H), 2.866 (ddd, J = 1.8, 5.3, 13.7 Hz, 1H), 2.669 (dd, J = 8.1, 13.7 Hz, 1H), 2.541 (s, 3H), 2.236 (ddd, J = 4.2, 6.4, 14.1 Hz, 1H), 2.112 (m, 1H), 1.707 (d, J = 1.0 Hz, 3H). 31 P NMR (D2 O, 121 MHz): 5.732. [2-(3-Indolyl)-1-(S)-methylcarbamoylethyl]phosphoramidic acid mono-(3 -deoxy-3 azidothymidyl)ester (S.4g). Yield of white solid: 62 mg (100%). 1 H NMR (D2 O, 300 MHz): 7.397 (d, J = 7.9 Hz, 1H), 7.187 (d, J = 8.2 Hz, 1H), 7.135 (s, 1H), 7.011 (s, 1H), 6.944 (dd, J = 7.0, 8.2 Hz, 1H), 6.836 (dd, J =7.0, 7.9 Hz, 1H), 5.736 (t, J = 6.7 Hz, 1H), 3.943 (m, 1H), 3.715-3.634 (m, 2H), 3.520-3.395 (m, 2H), 3.050 (ddd, J = 2.4, 4.4, 14.6 Hz, 1H), 2.780 (dd, J = 8.2, 14.6 Hz, 1H), 2.557 (s, 3H), 2.014 (m, 1H), 1.770 (m, 1H), 1.554 (s, 3H). 31 P NMR (D2 O, 121 MHz): 6.049. (2-Phenyl-1-(R)-methylcarbamoylethyl)phosphoramidic acid mono-(3 -deoxy-3 azidothymidyl)ester (S.4h). Yield of white solid: 56 mg (85%). 1 H NMR (D2 O, 300 MHz): 7.387 (d, J = 1.1 Hz, 1H), 7.193-7.032 (m, 5H), 6.022 (t, J = 6.7 Hz, 1H), 4.066 (m, 1H), 3.807 (m, 1H), 3.653-3.532 (m, 2H), 3.295 (m, 1H), 2.822-2.687 (m, 2H), 2.470 (s, 3H), 2.308-2.169 (m, 2H), 1.657 (d, 1.1 Hz, 3H). 31 P NMR (D2 O, 121 MHz): 5.713. [2-(3-Indolyl)-1-(R)-methylcarbamoylethyl]phosphoramidic acid mono-(3 -deoxy-3 azidothymidyl)ester (S.4i). Yield of white solid: 64 mg (99%). 1 H NMR (D2 O, 300 MHz): 7.425 (d, J = 7.9 Hz, 1H), 7.267 (d, J =7.3 Hz, 1H), 7.193 (d, J = 1.1 Hz, 1H), 7.020 (s, 1H), 7.007 (m, 1H), 6.885 (m, 1H), 5.860 (t, J = 6.8 Hz, 1H), 3.779 (m, 1H), 3.725 (m, 1H), 3.647-3.552 (m, 2H), 3.177 (m, 1H), 3.025 (m, 1H), 2.835 (dd, J = 7.5, 14.5 Hz, 1H), 2.472 (s, 3H), 2.121 (m, 1H), 1.862 (m, 1H), 1.525 (d, J = 1.1 Hz, 1H). 31 P NMR (D2 O, 121 MHz): 6.028.
FORMATION OF H-PHOSPHONATE MONOESTERS VIA BIS(N, N-DIISOPROPYLAMINO)CHLOROPHOSPHINE
ALTERNATE PROTOCOL
An alternate approach to the formation of nucleoside H-phosphonate monoesters utilizes bis(N, N-diisopropylamino)chlorophosphine as a phosphitylating agent (Fig. 15.1.2). The reaction is performed in two steps. The first step involves the formation of a nucleoside 5 phosphoramidite. In the second step, the phosphoramidite is reacted with acetic acid and water to form the H-phosphonate monoester. This method produces the H-phosphonate cleanly without the significant formation of side products.
Additional Materials (also see Basic Protocol) 2 -Acetyl-3 -deoxyadenosine (prepared as described by Zhang et al., 2003) Anhydrous N,N-diisopropylethylamine (DIPEA, 99% pure; Sigma-Aldrich) Bis(N,N-diisopropylamino)chlorophosphine (Acros) Anhydrous dioxane (99% pure; Acros) Glacial acetic acid Dowex 50WX4-50 (20 to 50 mesh; Sigma-Aldrich) Acetone/dry ice bath 2.5 × 15–cm chromatography columns 100-mL round-bottom flasks Prepare 2 -acetyl-3 -deoxyadenosine H-phosphonate 1. Dissolve 0.2285 g (0.779 mmol) 2 -acetyl-3 -deoxyadenosine in 10 mL dry CH2 Cl2 under argon gas. Add 0.177 mL (1.013 mmol) DIPEA and continue stirring for 30 min. 2. Dissolve bis(N,N-diisopropylamino)chlorophosphine in 3 mL dry CH2 Cl2 under argon gas and cool to 0◦ C. Add dropwise via a cannula to the solution of 2 -acetyl-3 deoxyadenosine. Allow reaction to warm to room temperature and continue stirring for 2 hr. Current Protocols in Nucleic Acid Chemistry
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15.1.7 Supplement 25
Figure 15.1.2
Alternate protocol for the formation of nucleoside H-phosphonates.
3. Evaporate reaction mixture on a rotary evaporator and then dry in vacuo overnight (in a vacuum pump at 0.05 to 0.5 Torr). 4. Dissolve crude reaction mixture in 5 mL dry dioxane under argon gas. 5. Add 0.27 mL (4.674 mmol) glacial acetic acid dropwise to crude reaction mixture. Continue stirring for 9 hr at room temperature. 6. Add 5 mL pyridine and 5 mL water. Stir for 30 min. 7. Concentrate reaction mixture on a rotary evaporator. 8. Dissolve crude mixture in 10 mL water. Extract aqueous layer three times with 10 mL CH2 Cl2 . 9. Collect aqueous layer and evaporate on a rotary evaporator. 10. Pack a 2.5 × 15–cm column with silica gel in 5:3:0.25 (v/v/v) CHCl3 /MeOH/water with 0.5% NH4 OH. Load the sample on the column and elute the desired compound with the same solvent. 11. Monitor fractions by TLC using the same solvent and UV visualization. Combine fractions containing a single UV spot and evaporate to dryness on a rotary evaporator. 12. Dry the resulting white solid 2 hr in vacuo. 13. To generate the TEA salt, pack a 2.5 × 15–cm column with Dowex 50WX4-50 in water. Wash column until it is pH neutral. Rinse resins with 1:3 (v/v) 10% aq. TEA/MeOH solution until basic (pH ∼10). Wash resins with water to a neutral pH. Dowex 50WX4-50 is acidic. Excess acid is washed off with water before adding TEA/MeOH solution. If the Dowex 50WX4-50 is not washed adequately, it would require more TEA/MeOH to form the TEA salt. Once the column is basic, excess TEA is removed by washing with water to pH ∼7.
14. Load the sample and elute with water. 15. Collect fractions containing UV activity in a 100-mL round-bottom flask. 16. Freeze in an acetone/dry ice bath and lyophilize. 17. Characterize the product by 1 H NMR.
Synthesis of Amino Acid Phosphoramidate Monoesters
2 -Acetyl-3 -deoxyadenosine H-phosphonate. Yield: 0.2174 g (55.3%). 1 H NMR (D2 O): 8.183 (s, 1H), 7.988 (s, 1H), 7.596-5.464 (d, JPH = 639.3, 1H), 6.050-6.044 (d, J = 1.8 Hz, 1H), 5.443-5.412 (m, 1H), 4.088-4.017 (m, 1H), 3.912-3.833 (m, 1H), 3.061-2.988 (q, J = 7.2 Hz, 6H), 2.445-2.343 (m, 1H), 2.237-2.162 (m, 1H), 2.008 (s, 3H), 1.129-1.080 (t, J = 7.5 Hz, 9H). 31 P NMR (D2 O): 7.807.
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To make the desired nucleoside H-phosphonate monoesters, use 200 mg (0.439 mmol) of this compound in step 12 of the Basic Protocol (i.e., 0.5 eq relative to the amino acid methyl ester hydrochloride).
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Methylamine solution Bubble methylamine gas (MeNH2 , 98% pure; Sigma-Aldrich) through anhydrous methanol until saturated. Dilute an aliquot with water and titrate with 1.0 N HCl using phenolphthalein as an indicator dye to determine the precise pH. The concentration of methylamine is then determined from the titration. The saturation point can be determined by weighing the methanol solution and stopping when the weight no longer changes.
COMMENTARY Background Information Nucleoside analogs are an important class of antiviral and anticancer agents. However, the utility of these compounds is often limited by poor pharmacokinetic properties. Nucleosides often possess low oral bioavailability, a relatively short plasma half-life, and low tissue distribution. Another problem that limits the efficacy of nucleosides is the dependence upon their phosphorylation by kinases to form the corresponding nucleoside phosphates, often to the triphosphate. In some cases, phosphorylation to the monophosphate is rate-limiting and may diminish the therapeutic efficacy of a nucleoside. Finally, drug resistance often develops through decreased phosphorylation of the nucleoside to the nucleoside monophosphate. To address these problems, nucleoside amino acid phosphoramidates have been synthesized as a prodrug approach to deliver phosphorylated nucleoside analogs as monoanions into the cell. These phosphoramidates are converted intracellularly by a phosphoramidase to the corresponding nucleoside monophosphate, thus overcoming the need for kinase activity to activate the nucleoside. Initially, AZT amino acid phosphoramidates were investigated as potential antiHIV and anti-breast cancer agents. The L-tryptophan methyl amide derivative was shown to possess anti-breast cancer activity in MCF-7 cells comparable to that of AZT (Iyer et al., 2000). Furthermore, this compound was found to be at least as effective as AZT in inhibiting the growth of NMU-induced rat mammary tumors (unpub. observ.). Development of this approach required a synthetic protocol that allows for variation of
both the amino acid and nucleoside positions. H-Phosphonate chemistry provides the most convenient method for synthesizing nucleoside phosphoramidates in moderate to high yield. Two methods are provided in this unit for the synthesis of the nucleoside H-phosphonate intermediate. Either method generates high yields of the intermediate, but the methodology utilized in the Basic Protocol also generates significant amounts (∼10% to 15%) of phenyl phosphite monoester as a contaminant. A solution to this problem, and considerations as to which protocol is more suitable, are addressed in greater depth in the Critical Parameters and Troubleshooting section. Although synthesis of phosphoramidates of ribonucleosides is beyond the scope of this unit, these protocols can be readily adapted to the formation of phosphoramidate monoesters of ribonucleosides. In the case of ribonucleosides, protection of the 2 - and 3 -hydroxyl groups prior to phosphitylation is necessary and easily accomplished through protection with an orthoformate ester. The protecting group can then be removed after formation of the H-phosphonate monoester but prior to the oxidative coupling of the amino acid (unpub. observ.). Synthesis of H-phosphonate intermediates H-Phosphonates are versatile intermediates and simple building blocks for synthesis of oligonucleotides and phosphoramidates. Various methods have been developed for generating nucleoside H-phosphonate monoesters by reactions of protected nucleosides with various phosphitylating reagents. Diphenyl phosphite. Diphenyl phosphite is an inexpensive and easy-to-handle alternative
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15.1.9 Current Protocols in Nucleic Acid Chemistry
Supplement 25
to other phosphitylating reagents. Nucleoside H-phosphonate monoesters are generated with this method in high yields (77% to 90%). The reaction involves transesterification with a protected nucleoside followed by basic hydrolysis to generate the nucleoside H-phosphonate monoester. Jankowska et al. (1994) have reported that symmetrical dinucleotides can be averted if excess phosphitylating reagent is used. Phenyl H-phosphonate monoester, which can have a similar Rf value to the desired nucleoside H-phosphonate, has been obtained when only one equivalent of nucleoside was used. The Alternate Protocol can be used to circumvent this problem. Bis(N,N - diisopropylamino)chlorophosphine. Bis(N,N-diisopropylamino)chlorophosphine is relatively inexpensive and has been used to generate nucleoside H-phosphonates in high yields (70% to 91%; Marugg et al., 1986a,b; Nahum et al., 2002). Reaction of bis(N,N-diisopropylamino)chlorophosphine with a nucleoside proceeds through a nucleoside phosphite intermediate, which can be converted to the corresponding nucleoside H-phosphonate monoester via acidolysis. This method is not compatible with nucleosides containing acid-labile protecting groups. PCl3 . A commonly used method is PCl3 /imidazole, which consistently generates good yields (75% to 90%) of the protected Hphosphonate monoesters (UNIT 2.6). Triazole can be substituted for imidazole with little effect on yields. However, PCl3 is highly reactive and difficult to handle. Salicylchlorophosphite. Salicylchlorophosphite is commercially available and stable. Reaction of a nucleoside with salicylchlorophosphite generates a nucleoside phosphite intermediate that can be converted to the corresponding H-phosphonate via hydrolysis. Marugg et al. (1986a) demonstrated that nucleoside H-phosphonates can be generated in very high yields (88% to 91%) using this method. Salicylchlorophosphite is preferred over bis(N,N-diisopropylamino)chlorophosphine due to the mild hydrolytic conditions. Though salicylchlorophosphite is very efficient, difficulties can occur in separating the H-phosphonate monoester from salicylic acid.
Synthesis of Amino Acid Phosphoramidate Monoesters
Synthesis of phosphoramidates via oxidation of H-phosphonates One commonly used oxidation method for H-phosphonate monoesters involves iodine, as described in this unit. This procedure is a convenient and mild oxidation that has been
employed in the synthesis of phosphoramidates from corresponding H-phosphonate monoesters. This method generates a reactive iodophosphate that is susceptible to nucleophilic addition by an amine to yield the desired phosphoramidate in good yields (50% to 70%; Garegg et al., 1987). Alternative synthetic approaches for phosphoramidates An alternative approach for the synthesis of phosphoramidates involves the generation of nucleoside monophosphates, followed by a coupling reaction to generate the desired compound. This approach has been used successfully by Abraham et al. (1996, 1997) to synthesize phosphoramidates of 1-β-arabinofuranosylcytosine (Ara-C), 9-[(2hydroxyethoxy)methyl]guanine (acyclovir, ACV, Zovirax), and 5-fluoro-2 -deoxyuridine (FUdR) via dicyclohexylcarbodiimide (DCC) coupling from their corresponding monophosphates. Synthesis of the nucleoside monophosphates from the protected nucleoside can be accomplished through reactions with 2-cyanoethyl phosphate or phosphorus oxychloride with modest yields (60% to 70%). The monophosphates are then coupled to the desired amino acid methyl ester via DCC coupling (50% to 60% yield). This approach is advantageous if the nucleoside monophosphate is commercially available. Another approach for synthesis of phosphoramidates utilizes phosphorochloridate chemistry. Abraham and Wagner (1994) has used this method to synthesize phosphoramidates of AZT, 3 -fluoro-3 -deoxythymidine (FLT), and 2 ,3 -dideoxy-didehydrothymidine (D4T). The nucleoside is treated with two equivalents of 2-cyanoethyl-N,N-diisopropylchlorophosphoramidite to generate the nucleoside phosphoramidite intermediate in good yields (79% to 96%). The phosphoramidite intermediate is treated with tetrazole and methanol to generate the methyl phosphite intermediate. Next, this intermediate is oxidized in the presence of iodine and an amino acid methyl ester in a onepot, two-step reaction to yield the desired phosphoramidate in good yields (79% to 100%). Although this approach gives good overall yields (30% to 77%) and does not require excess nucleoside, this method suffers from the need to use a tedious purification procedure. In addition, 2-cyanoethyl-N,Ndiisopropylchlorophosphoramidite is expensive and this method employs two equivalents.
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Current Protocols in Nucleic Acid Chemistry
Critical Parameters and Troubleshooting Nucleoside amino acid methyl amide phosphoramidate monoesters can be synthesized in a good to moderate yield in three steps using the protocols outlined in this unit. In addition to knowledge of basic organic chemistry skills such as TLC, solvent evaporation, extraction, and column chromatography, knowledge of 1 H and 31 P NMR is essential. Furthermore, techniques such as ESI-MS, mass analysis, and HPLC are useful for the characterization of the compounds and determination of purity. In addition to these common practices, the importance of maintaining anhydrous conditions during the reaction and scrupulous attention to anhydrous storage and usage of reagents cannot be overemphasized. Care should also be taken with respect to reaction times; reaction times should not be longer than stated in the protocols. The Basic Protocol for the formation of nucleoside H-phosphonate monoester involves a stoichiometric equivalence of diphenyl phosphite with the nucleoside. This protocol, however suffers from the formation of significant (5% to 15%) amounts of phenyl H-phosphonate monoester, which is difficult to remove from the desired product. This problem can be largely overcome by using 0.5 equiv of diphenyl phosphite rather than 1 equiv (unpub. observ.). Although this is not an optimal solution, the cost of using diphenyl phosphite as the limiting reagent can be partially mitigated by recovery of the unreacted nucleoside by column chromatography. The Alternate Protocol utilizes bis(N,Ndiisopropylamino)chlorophosphine. This protocol has the advantage that one can use a 1:1 equivalence of phosphitylating agent to nucleoside. Nevertheless, this reagent is more expensive than diphenyl phosphite and is less amenable to long-term storage. During the oxidation of the nucleoside H-phosphonate to the phosphoramidate with iodine, care should be taken to titrate one equivalent of iodine into the reaction. Excess iodine is difficult to remove and will cause the purified phosphoramidate to appear yellow in color. Unwanted recovery of the starting nucleoside H-phosphonate monoester after oxidizing with iodine can occur for one of two reasons: either insufficient iodine or excess water. The first problem can be avoided by titrating the reaction until it is reddish brown. Excess water can be removed through extensive drying or through the use of additional TMSCl.
Anticipated Results Yields of nucleoside phosphoramidate monoester vary from moderate to good when utilizing H-phosphonate chemistry as the key intermediate (48% for first two steps). These results are consistent with results from Kers and Kraszewski (1998), and are modest improvements over previous methods, such as DCC coupling (38% over first two steps) and phosphoramidite chemistry (30% for first three steps).
Time Considerations Synthesis of AZT amino acid phosphoramidate monoesters starting from AZT (S.1) can be accomplished in ∼10 days per amino acid methyl ester. To make the process more efficient, preparation of large-scale quantities of S.2 as a useful synthetic intermediate is recommended. These protocols do not differ substantially from those developed by Stawinski, and times required for isolation and purification were similar to those for standard organic compounds.
Literature Cited Abraham, T.W. and Wagner, C.R. 1994. A phosphoramidite-based synthesis of phosphoramidate amino acid diesters of antiviral nucleosides. Nucleosides Nucleotides 13:18911903. Abraham, T.W., Kalman, T.I., McIntee, E.J., and Wagner, C.R. 1996. Synthesis and biological activity of aromatic amino acid phosphoramidates of 5-fluoro-2 -deoxyuridine and 1β-arabinofuranosylcytosine: Evidence of phosphoramidase activity. J. Med. Chem. 39:45694575. Abraham, T.W., McIntee, E.J., Iyer, V.V., Schinazi, R.F., and Wagner, C.R. 1997. Synthesis, biological activity, and decomposition studies of amino acid phosphomonoester amidates of acyclovir. Nucleosides Nucleotides 16:2079-2092. Garegg, P.J., Regber, T., Stawinski, J., and Stromberg, R. 1987. Nucleoside phosphonates: Part 7. Studies on the oxidation of nucleoside phosphonate esters. J. Chem. Soc., Perkin Trans. 1 1269-1273. Iyer, V.V., Griesgraber, G.W., Radmer, M.R., McIntee, E.J., and Wagner, C.R. 2000. Synthesis, in vitro anti-breast cancer activity, and intracellular decomposition of amino acid methyl ester and alkyl amide phosphoramidate monoesters of 3 azido-3 -deoxythymidine (AZT). J. Med. Chem. 43:2266-2274. Jankowska, J., Sobkowski, M., Stawinski, J., and Kraszewski, A. 1994. Studies on aryl H-phosphonates. I. An efficient method for the preparation of deoxyribo- and ribonucleoside 3 -H-phosphonate monoesters by transesterification of diphenyl H-phosphonate. Tetrahedron Lett. 35:3355-3358.
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Kers, A. and Kraszewski, A. 1998. A new synthetic method for the preparation of nucleoside phosphoramidate analogues with the nitrogen atom in bridging positions of the phosphoramidate linkage. Tetrahedron Lett. 39:1219-1222. Marugg, J., Tromp, M., Kuyl-Yeheskiely, E., van der Marel, G.A., and van Boom, J.H. 1986a. A convenient and general approach to the synthesis of properly protected D-nucleoside-3 hydrohenphosphonates via phosphite intermediates. Tetrahedron Letters 27:2661-2664. Marugg, J., Tromp, M., van der Marel, G.A., and van Boom, J.H. 1986b. A new versatile approach to the preparation of valuable deoxynucleoside 3 -phosphite intermediates. Tetrahedron Lett. 27:2271-2274. Nahum, V., Zundorf, G., Levesque, S.A., Beaudoin, A.R., Reiser, G., and Fischer, B. 2002. Adenosine 5 -O-(1-boranotriphosphate) derivatives as novel P2Y(1) receptor agonists. J. Med. Chem. 45:5384-5396. Zhang, L., Cui, Z., and Zhang, B. 2003. An efficient synthesis of 3 -amino-3 -deoxyguanosine from guanosine. Helv. Chim. Acta 86:703-710.
Contributed by Cindy J. Choy, Dan P. Drontle, and Carston R. Wagner University of Minnesota Minneapolis, Minnesota
Synthesis of Amino Acid Phosphoramidate Monoesters
15.1.12 Supplement 25
Current Protocols in Nucleic Acid Chemistry
Synthesis of Cidofovir and (S)-HPMPA Ether Lipid Prodrugs
UNIT 15.2
This unit provides procedures for preparing alkoxyalkyl monoester prodrugs of two potent antiviral compounds: (S)-1-(3-hydroxy-2-phosphonomethoxypropyl)cytosine, i.e., (S)-HPMPC, also known as cidofovir or CDV, and (S)-9-(3-hydroxy-2phosphonomethoxypropyl)adenine, i.e., (S)-HPMPA. Without modification, negatively charged acyclic nucleoside phosphonates (ANPs) such as cidofovir and (S)-HPMPA are too polar to penetrate cell membranes effectively and, consequently, have low bioavailability upon oral administration. In the approach described here, the lipophilicity and intestinal absorption of nucleoside phosphonates is enhanced by masking one of the phosphonate charges and transforming the drugs to resemble lysophosphatidylcholine, a readily absorbed dietary phospholipid. Absorbed alkoxyalkyl prodrugs are efficiently taken up by cells and converted enzymatically to the parent phosphonate, a process that leads to high intracellular levels of ANP diphosphate, the active antiviral metabolite. Hexadecyloxypropyl-cidofovir (HDP-CDV) and hexadecyloxypropyl-(S)-HPMPA (HDP-(S)-HPMPA) are active when administered orally in several animal models of antiviral disease and are currently being evaluated for further development (Painter and Hostetler, 2004; Beadle et al., 2006). Two approaches for synthesizing the alkoxyalkyl ester prodrugs are described. Basic Protocol 1 utilizes the Mitsunobu reaction to achieve dehydrative coupling between cyclic cidofovir and 3-hexadecyloxy-1-propanol. Selective basic hydrolysis converts the cyclic phosphonodiester to HDP-CDV. Basic Protocol 2 illustrates an approach that incorporates the lipophilic ester early into a stepwise synthesis and yields HDP-(S)HPMPA directly. Two procedures for the preparation of 3-hexadecyloxy-1-propanol are also provided (see Support Protocols 1 and 2). Interested readers should also refer to UNIT 14.2 for methods related to the synthesis and characterization of acyclic nucleoside phosphonates. CAUTION: All reactions must be run in a suitable fume hood with efficient ventilation. Safety glasses and reagent-impermeable protective gloves should be worn. Products of Basic Protocols 1 and 2 exhibit potent biological activity and readily penetrate the skin and mucosal surfaces and should be handled with care. When handling powder, a particle mask should be worn. NOTE: All glassware should be oven dried, and all reactions should be performed under anhydrous conditions unless noted otherwise. All reactions are performed under a nitrogen atmosphere.
SYNTHESIS OF HEXADECYLOXYPROPYL-CIDOFOVIR Hexadecyloxypropyl-cidofovir (S.4; see Fig. 15.2.1) is a broad-spectrum antiviral agent for DNA viruses, active against herpesviruses (HSV-1, HSV-2, HCMV, VZV), adenoviruses, and orthopoxviruses. This protocol describes its preparation in two steps from cyclic cidofovir (S.1) and 3-hexadecyloxy-1-propanol (S.2). Cyclic cidofovir is obtained from cidofovir, which bears a primary hydroxyl group at the 3 position that readily forms the six-membered cyclic ester (Bischofberger et al., 1994). The Mitsunobu reaction (Mitsunobu, 1981) is used to couple S.1 and 3-hexadecyloxy-1-propanol. Finally, the cyclic diester is exposed to basic conditions that selectively hydrolyze the cyclic ester and yield hexadecyloxypropyl-cidofovir.
Contributed by James R. Beadle Current Protocols in Nucleic Acid Chemistry (2007) 15.2.1-15.2.16 C 2007 by John Wiley & Sons, Inc. Copyright
BASIC PROTOCOL 1
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Figure 15.2.1 Synthesis scheme for the preparation of hexadecyloxypropyl-cidofovir (HDP-CDV, S.4) from cyclic cidofovir (S.1) and 3-hexadecyloxy-1-propanol (S.2). DIAD, diisopropylazodicarboxylate.
Materials Cidofovir (CDV; WaterStone Technology; also see UNIT 14.2) N,N-Dimethylformamide, anhydrous (DMF; Aldrich) N,N -Dicyclohexyl-4-morpholinecarboxamidine (Aldrich) 1,3-Dicyclohexylcarbodiimide (DCC; Aldrich) Pyridine Diethyl ether Dichloromethane (CH2 Cl2 ) 1 N hydrochloric acid (HCl) in H2 O Acetone Toluene 3-Hexadecyloxy-1-propanol (S.2; see Support Protocol 1 or 2) Triphenylphosphine (Aldrich) Diisopropylazodicarboxylate (DIAD; Aldrich) Chloroform (CHCl3 ) Methanol (MeOH) Concentrated ammonium hydroxide (NH4 OH) Phospray TLC reagent (Supelco) Ethanol (EtOH) Silica gel 60 (230 to 400 mesh; EMD Chemicals) p-Dioxane 0.5 N sodium hydroxide (NaOH) in H2 O Nitrogen tank 50% (v/v) acetic acid in H2 O Isopropanol Synthesis of Cidofovir and (S)-HPMPA Ether Lipid Prodrugs
500-, 250-, and 100-mL round-bottom flasks, oven dried 60-mL addition funnel 60◦ and 40◦ C oil baths Rotary evaporator equipped with a vacuum pump
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Frit funnel (medium porosity, 10 to 16 µm) with vacuum Azeotropic distillation apparatus Dean-Stark trap Vacuum oven (70◦ C) Silica gel thin-layer chromatography (TLC) plates (e.g., 250-µm Silica Gel GF Uniplates; Analtech) Filter funnel with vacuum UV lamp Muffle furnace set to 400◦ C Glass flash chromatography columns: 50 × 457 mm Vacuum oil pump Synthesize cyclic cidofovir (S.1) 1. Place 2 g (7.2 mmol) cidofovir and 40 mL DMF in a 100-mL roundbottom flask containing a stir bar. Add 2.3 g (7.9 mmol) N,N -dicyclohexyl-4morpholinecarboxamidine and stir 12 hr at room temperature. The cidofovir should slowly dissolve to give a clear solution.
2. Add 3.7 g (1.8 mmol) DCC and 100 mL pyridine to a 500-mL round-bottom flask equipped with a 60-mL addition funnel and a stir bar. While stirring, heat the mixture to 60◦ C, and then slowly add the cidofovir/DMF solution over ∼15 min using the addition funnel. After addition is complete, continue stirring at 60◦ C for 16 hr. 3. Cool to room temperature and evaporate the solvents using a rotary evaporator connected to a vacuum pump. 4. Wash the residue with 60 mL diethyl ether and then dissolve in 40 mL water. 5. Wash the aqueous layer three times with 25 mL CH2 Cl2 . 6. Concentrate the aqueous layer to 20 mL using a rotary evaporator, and then acidify it to pH 3.5 using 1 N HCl. Cool the aqueous solution in a 4◦ C refrigerator for 12 hr. 7. Collect the precipitated S.1 dihydrate by vacuum filtration on a medium-porosity frit funnel and wash with 40 mL acetone. 8. Dry the resulting powder by azeotropic distillation with toluene. Transfer S.1 dihydrate to a 100-mL round-bottom flask and add a Dean-Stark trap. Heat the mixture to reflux and continue until no more water collects in the trap (∼15 min). 9. Cool to room temperature. Collect the solid by vacuum filtration and keep in a vacuum oven at 70◦ C and 10 mmHg for 12 hr to obtain the anhydrous compound. Cyclic cidofovir (cCDV; S.1): 1.55 g (83% yield). 1 H NMR: δ 7.83 (d, 1H), 6.15 (d, 1H), 3.70-4.27 (m, 7H). 31 P NMR: δ 9.35, singlet.
Synthesize HDP-cCDV (S.3) 10. In a 250-mL round-bottom flask equipped with a stir bar and nitrogen gas source, suspend the following in 50 mL dry DMF: 1 g (3.8 mmol) anhydrous cyclic cidofovir (S.1) 2.3 g (7.6 mmol) 3-hexadecyloxy-1-propanol (S.2) 2.0 g (7.6 mmol) triphenylphosphine. 11. While stirring at room temperature, add 1.5 g (7.6 mmol) DIAD in three portions over ∼15 min.
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12. Stir overnight or until the reaction is complete. Monitor the reaction by TLC on silica gel plates using 80:20:1:1 (v/v) CHCl3 /MeOH/conc. NH4 OH/H2 O. Visualize the spots under UV light, then by spraying with Phospray and placing in a muffle furnace at 400◦ C. The starting material should be run alongside the reaction for comparison. The product spot is easily identifiable by TLC: it is UV active, turns blue when treated with Phospray, and chars dark black at 400◦ C. The product typically has an Rf value of 0.7.
13. Quench the reaction with 1 mL water and remove the solvents using a rotary evaporator equipped with a vacuum pump to give a light brown oil. 14. Dissolve the residue in 75 mL CH2 Cl2 , add 10 g silica gel, and evaporate to dryness on the rotary evaporator. 15. Pour the adsorbed mixture onto the top of a 50 × 457–mm flash chromatography column packed with 60 g silica gel. Start eluting with CH2 Cl2 , then with a gradient from 0% to 15% EtOH in CH2 Cl2 . Collect 25-mL fractions. 16. Combine fractions that contain pure product S.3 as determined by TLC using 80:20:1:1 (v/v) CHCl3 /MeOH/conc. NH4 OH/H2 O. 17. Evaporate combined fractions to dryness on the rotary evaporator. 18. Crystallize the product from 10 mL p-dioxane, collect on a filter funnel under vacuum, and dry overnight under vacuum using a vacuum oil pump. CAUTION: p-Dioxane can form potentially explosive peroxides upon exposure to air. It should always be stored in a tightly closed container and used within 6 months after opening or tested for peroxides before use.
19. Check the purity of the product. Hexadecyloxypropyl-cyclic cidofovir (S.3): 1.2 g (58% yield from cCDV). TLC (80:20:1:1 (v/v) CHCl3 /MeOH/conc. NH4 OH/H2 O): Rf = 0.6 (UV and charring). 1 H and 31 P NMR spectroscopic analysis showed the presence of two diastereomeric (axial and equatorial) alkylation products. 1 H NMR (DMSO-d6 ): δ 0.85 (t, 3H), 1.23 (broad s, 28H), 1.47 (m, 2H), 1.84 (p, 2H), 3.55-4.20 (m, 11H), 5.65 (dd, 1H), 7.18 and 7.04 (broad s, 1H), 7.55 and 7.45 (d, 1H), 8.30 (broad s, 2H). 31 P NMR: +13.88 and +12.62. MS (ESI): m/z 544 (M+H)+ , 542 (M–H)– .
Synthesize HDP-CDV (S.4) 20. Place 2.62 g S.3 (4.8 mmol) and 25 mL of 0.5 M NaOH in a 100-mL round-bottom flask equipped with a stir bar. 21. Put the flask in a 40◦ C oil bath and stir the mixture under a nitrogen atmosphere for 4 hr or until hydrolysis is complete. Monitor the reaction by TLC using 80:20:1:1 (v/v) CHCl3 /MeOH/conc. NH4 OH/H2 O, and visualize the spots by UV shadowing, Phospray, and charring. The product appears at the origin. No charring or UV should remain from S.3 (Rf = 0.5-0.8). During the reaction the starting material will dissolve to give a clear solution.
22. When complete, cool the reaction mixture in an ice/water bath to 0◦ C. 23. While stirring, slowly neutralize the mixture with 50% acetic acid until a thick precipitate forms and the pH is less than 6, and then add 1 N HCl until the pH reaches ∼1. Collect the solid product on a filter funnel under vacuum and dry overnight under vacuum. Synthesis of Cidofovir and (S)-HPMPA Ether Lipid Prodrugs
24. Crystallize the crude product from 20 mL isopropanol, filter on a filter funnel under vacuum, and dry overnight under vacuum.
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25. Check the purity of the product. Hexadecyloxypropyl-cidofovir (S.4): 2.1 g (76% yield). 1 H NMR (DMSO-d6 ): δ 0.86 (t, 3H), 1.24 (broad s, 28H), 1.47 (m, 2H), 1.73 (p, 2H), 3.20-3.89 (m, 11H), 5.72 (m, 1H), 7.21 (d, 1H), 7.54 (d, 1H), 8.23 (broad s, 2H). 31 P NMR: +13.98. MS (ESI) m/z 584 (M+Na)+ , 560 (M–H)– . The product can be stored in an air-tight container for at least 12 months at –20◦ C.
SYNTHESIS OF HEXADECYLOXYPROPYL-(S)-HPMPA (S)-9-(3-Hydroxy-2-phosphonomethoxypropyl)adenine was the first acyclic nucleoside phosphonate to show potent, broad-spectrum antiviral activity (De Clercq et al., 1986). The alkoxyalkyl prodrug HDP-(S)-HPMPA inhibits the replication of DNA viruses, including herpes simplex virus (HSV) types 1 and 2, varicella zoster virus (VZV), thymidine kinase–deficient (TK– ) mutants of HSV and VZV, human cytomegalovirus (HCMV), Epstein-Barr virus, hepatitis B virus (HBV), adenoviruses, and various orthopoxviruses (Beadle et al., 2006). Unlike (S)-HPMPA, HDP-(S)-HPMPA shows significant activity against HIV-1 in vitro (Hostetler et al., 2006).
BASIC PROTOCOL 2
The method for preparing HDP-(S)-HPMPA differs from Basic Protocol 1 in that it is a stepwise approach in which the alkoxyalkyl ester is incorporated early into the synthesis. The key step is preparation of the phosphonate synthon, hexadecyloxypropyl p-toluenesulfonyloxymethylphosphonate (S.8; see Fig. 15.2.2) from commercially available diethyl toluenesulfonyloxymethylphosphonate and S.2. Alkylation of the secondary hydroxyl of (S)-9-(3-trityloxy-2-hydroxypropyl)-N6 -trityladenine (S.7) with S.8 affords the fully protected HPMPA analog (S.9). Detritylation under acidic conditions gives HDP-(S)-HPMPA.
Figure 15.2.2 Synthesis scheme for the preparation of hexadecyloxypropyl-(S)-HPMPA (HDP-(S)-HPMPA, S.10) from adenine. R, hexadecyloxypropyl; Tr, trityl. Nucleoside Prodrugs and Delivery Strategies
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Materials Adenine (S.5; Aldrich) (S)-Trityl glycidyl ether (Daiso) N,N-Dimethylformamide (DMF), anhydrous 1,8-Diazabicyclo[5,4,0]undec-7-ene (DBU; Aldrich) Chloroform (CHCl3 ) Methanol (MeOH) Toluene, dry Trityl chloride (Aldrich) N,N-Dimethylaminopyridine (DMAP; Aldrich) Pyridine (anhydrous, Aldrich) Ethyl acetate (EtOAc) Saturated sodium bicarbonate (NaHCO3 ), cold Anhydrous magnesium sulfate (MgSO4 ) Silica gel 60 (230 to 400 mesh; EMD Chemicals) Hexanes Diethyl toluenesulfonyloxymethylphosphonate (Biofine International) Dichloromethane (CH2 Cl2 ), anhydrous Bromotrimethylsilane (Aldrich) Oxalyl chloride (Acros Organics) Diethyl ether Triethylamine, dry Sodium t-butoxide (Aldrich) NaCl, cold saturated solution Ethanol (EtOH) 80% acetic acid in water Phospray TLC reagent (Supelco) Concentrated ammonium hydroxide (NH4 OH) 250-, 100-, and 50-mL round-bottom flasks, oven dried Reflux condensers 50◦ , 60◦ , and 100◦ C oil baths Silica gel thin-layer chromatography (TLC) plates (e.g., 250-µm Silica Gel GF Uniplates, Analtech) UV lamp Rotary evaporator and vacuum pump Filter funnels with vacuum Vacuum oil pump 250-mL and 1-L separatory funnels Glass flash chromatography columns: 50 × 457 mm and 64 × 457 mm 150-mL dropping funnel Muffle furnace set to 400◦ C Synthesize S.6 1. In a 250-mL round-bottom flask equipped with a magnetic stir bar and a reflux condenser place: 4.9 g (36 mmol) adenine 10.0 g (31.6 mmol) (S)-trityl glycidyl ether 100 mL anhydrous DMF. Synthesis of Cidofovir and (S)-HPMPA Ether Lipid Prodrugs
2. While stirring, add 1.1 g (7.2 mmol) DBU and then heat the suspension to 100◦ C in an oil bath. Continue stirring 2 hr or until the reaction is complete. Monitor the reaction by TLC using 9:1 (v/v) CHCl3 /MeOH, visualizing spots by UV light.
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The starting material should be run alongside the reaction for comparison. Adenine will be near the origin and the product will have Rf = 0.5.
3. When complete, allow the reaction to cool, then remove solvents using a rotary evaporator equipped with a vacuum pump until a thick, light-brown slurry remains. 4. Add 50 mL dry toluene to the flask and swirl or stir ∼30 min until crystallization is complete. Collect the solid on a filter funnel under vacuum, wash the solid with 50 mL toluene, and dry under vacuum overnight using a vacuum oil pump. 5. Crystallize the crude product from 50 mL toluene, collect on a filter funnel under vacuum, and dry overnight under vacuum. 6. Check the purity of the product. (S)-9-(3-Trityloxy-2-hydroxypropyl)adenine (S.6): 11.3 g (70% from adenine). 1 H NMR (CDCl3 ): δ 8.20 (s, 1H), 7.75 (s, 1H), 7.23-7.43 (m, 15H), 4.42 (dd, 1H), 4.27 (dd, 1H), 4.13-4.17 (m, 1H), 3.22 (dd, 1H), 3.10 (dd, 1H).
Synthesize S.7 7. In a 250-mL round-bottom flask equipped with a stir bar and a reflux condenser, dissolve the following in 120 mL dry pyridine: 10.8 g (24 mmol) S.6 6.6 g (24 mmol) trityl chloride 0.3 g (2.4 mmol) DMAP. 8. Put the flask in a 50◦ C oil bath and stir 16 hr. Monitor the reaction by TLC using EtOAc, visualizing the spots by UV light. S.6 will remain at the origin, trityl chloride has Rf = 0.9, and the product will appear at Rf = 0.4.
9. When complete, quench by adding 2 mL water, dilute with 200 mL EtOAc, and pour the mixture into a 1-L separatory funnel. Wash the organic layer with cold, saturated NaHCO3 . 10. Dry the organic layer over anhydrous MgSO4 , filter by gravity filtration, and evaporate to a brown oil using a rotary evaporator. 11. Adsorb the residue on 40 g silica gel and add to a 64 × 457–mm flash chromatography column packed with 400 g silica gel. Elute the column with hexanes, then with a linear gradient of 10% to 60% EtOAc/hexanes. Collect 50-mL fractions. 12. Combine fractions that contain pure product as determined by TLC using EtOAc. 13. Evaporate the combined fractions to dryness on a rotary evaporator and keep under vacuum overnight. 14. Check the purity of the product. (S)-9-(3-Trityloxy-2-hydroxypropyl)-N6 -trityladenine (S.7): ∼11.1 g of a glassy solid (66% from S.6).1 H NMR: δ 7.93 (s, 1H), 7.27 (s, 1H), 7.22-7.42 (m, 30H), 4.36 (dd, 1H), 4.24 (dd, 1H), 4.18 (m,1H), 3.21 (dd, 1H), 3.07 (dd, 1H).
Synthesize S.8 15. In a 250-mL round-bottom flask equipped with a magnetic stir bar, add 9.5 g (29.5 mmol) diethyl toluenesulfonyloxymethylphosphonate and 150 mL anhydrous CH2 Cl2 . Place the flask in an ice/water bath and cool to 0◦ C. 16. While stirring, add 27 g (175 mmol) bromotrimethylsilane. Allow the ice to melt slowly and let the mixture warm to room temp, and continue stirring for 18 hr.
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17. Evaporate the solvent using a rotary evaporator equipped with a vacuum pump to give a light brown oil. Co-evaporate the product three times with 25 mL dry toluene. 18. Dissolve the residue in 150 mL CH2 Cl2 , cool to 0◦ C with an ice bath, and add 0.5 mL DMF. 19. Equip the flask with a 150-mL dropping funnel and use it to slowly add a solution of 22 g (175 mmol) oxalyl chloride in 50 mL CH2 Cl2 over about 30 min. CAUTION: This results in evolution of CO2 .
20. Allow to warm to room temperature and stir 5 hr. 21. Evaporate the solvents using the rotary evaporator. Suspend the residue in 100 mL diethyl ether. 22. While stirring, use a dropping funnel to add a solution of
6.46 g (21.5 mmol) 3-hexadecyloxy-1-propanol (S.2) 10 mL dry pyridine 50 mL diethyl ether. 23. Stir for 3 hr or until TLC analysis using 1:1 (v/v) hexanes/EtOAc indicates complete phosphonylation of the alcohol. Visualize the spots by spraying with Phospray and charring in a muffle furnace at 400◦ C. The product should appear at the origin and the starting material S.2 typically has Rf = 0.5.
24. When complete, pour the reaction mixture into 250 mL cold saturated NaHCO3 and stir vigorously 1 hr. 25. Transfer the mixture to a separatory funnel and separate the organic layer. Extract the aqueous phase with 50 mL diethyl ether. 26. Combine the organic extracts, dry over MgSO4 , filter by gravity filtration, and evaporate to dryness using the rotary evaporator. 27. Adsorb the residue on 30 g silica gel and add to a 50 × 457–mm flash chromatography column packed with 200 g silica gel. Start eluting with 300 mL CH2 Cl2 , then with a linear gradient of 0% to 15% EtOH in CH2 Cl2 . Collect 50-mL fractions. 28. Combine fractions that contain pure product S.8 as determined by TLC using 9:1 (v/v) CHCl3 /MeOH. The product (Rf ≈ 0.4) should have slight UV shadowing, a blue color after spraying with Phospray, and dark black charring at 400◦ C.
29. Evaporate combined product fractions to dryness on the rotary evaporator and dry overnight under vacuum. 30. Crystallize the crude product from 50 mL EtOAc, filter on a filter funnel under vacuum, and dry overnight under vacuum. 31. Check the product for purity.
Synthesis of Cidofovir and (S)-HPMPA Ether Lipid Prodrugs
3-(Hexadecyloxy)propyl toluenesulfonyloxymethylphosphonate, sodium salt (S.8): Yield ∼11.1 g (67% from S.2). 1 H NMR (CDCl3 ): δ 0.88 (t, 3H, -CH3 ), 1.26 (br s, 26H, -OCH2 CH2 (CH2 )13 CH3 ), 1.54 (m, 2H, -OCH2 CH2 (CH2 )13 CH3 ), 1.83 (qt, 2H, OCH2 CH2 CH2 O-), 2.46 (s, 3H, toluyl-CH3 ), 3.38 (t, 2H, -CH2 -), 3.47 (t, 2H, -CH2 -), 3.91 (dt, 2H, -P(O)OCH2 -), 4.02 (d, 2H, -CH2 P(O)O-), 7.37 (d, 2H, aromatic), 7.80 (d, 2H, aromatic). MS (ES): m/z 571.32 [M+Na]+ , 593.27 [M–H+2Na]+ , 547.02 [M–H]– . Anal. calcd. for C27 H48 O7 NaPS·0.5H2 O: %C 55.94, %H 8.52; found %C 56.34, %H 8.56.
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Synthesize S.9 32. In a 100-mL round-bottom flask equipped with a stir bar, dissolve 2.1 g (3.0 mmol) S.7 in 50 mL dry triethylamine. While stirring, add 340 mg (3.5 mmol) sodium t-butoxide and stir 15 min. 33. Add 1.7 g (3.0 mmol) S.8 and then put the flask into a 50◦ C oil bath. Stir the reaction mixture 3 hr. Monitor the reaction by TLC using 9:1 (v/v) CHCl3 /MeOH. The starting material should be run alongside to aid in identifying the product. The product (Rf ≈ 0.7) is UV active, turns yellow after Phospray, and chars dark black at 400◦ C.
34. When complete, pour the mixture into cold saturated NaCl and transfer to a 250-mL separatory funnel. Extract the mixture three times with 30 mL EtOAc. 35. Dry the organic phase over anhydrous MgSO4 , filter by gravity filtration, and then evaporate the solvents using a rotary evaporator. 36. Adsorb the residue on 20 g silica gel and add to a 50 × 457–mm flash chromatography column packed with 100 g silica gel. Start eluting with 250 mL CH2 Cl2 , then with a linear gradient of 0% to 10% EtOH in CH2 Cl2 . Collect 25-mL fractions. 37. Combine fractions that contain pure product S.9 as determined by TLC using 9:1 (v/v) CHCl3 /MeOH. 38. Evaporate combined fractions to dryness using a rotary evaporator. 39. Check the product for purity. 3-(Hexadecyloxy)propyl-(S)-9-[3-trityloxy-2-(phosphonomethoxy)propyl]-N6 -trityladenine (S.9): Yield: ∼1.0 g (31% from S.7). 1 H NMR(CDCl3 ): δ 0.88 (t, 3H), 1.33 (br s, 26H), 1.46 (m, 2H), 1.77 (qt, 2H), 3.37-3.8 (m, 9H), 3.86 (m, 2H), 4.03 (m, 2H), 7.2-7.5 (br m, 30H), 7.82 (s, 1H), 8.20 (s, 1H). MS (ES): m/z 1071 [M+H]+ , 1069 [M–H]– .
Synthesize S.10 40. In a 50-mL round-bottom flask equipped with a stir bar, suspend 1.0 g S.9 in 20 mL 80% aqueous acetic acid. 41. Put the flask into a 60◦ C oil bath and stir for 1 hr or until detritylation is complete as determined by TLC using 9:1 (v/v) CHCl3 /MeOH. Visualize the spots by UV light, Phospray, and charring at 400◦ C. The product should appear close to the origin, be UV active, not yellow after Phospray, and char dark black at 400◦ C.
42. When complete, cool the reaction mixture to 4◦ C, filter by gravity filtration, then remove the solvents using a rotary evaporator and keep under vacuum overnight. 43. Adsorb the residue on 5 g of silica gel and add to a 50 × 457–mm flash chromatography column packed with 30 g silica gel. Elute the column with a linear gradient of 0% to 30% MeOH in CH2 Cl2 . Collect 20-mL fractions. 44. Combine fractions that contain pure S.10 as determined by TLC using 70:30:3:3 (v/v) CHCl3 /MeOH/conc. NH4 OH/H2 O, visualizing with UV light, Phospray, and charring at 400◦ C (product Rf = 0.5). 45. Evaporate combined fractions to dryness on the rotary evaporator and dry overnight under vacuum. Nucleoside Prodrugs and Delivery Strategies
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46. Check the product for purity. 3-(Hexadecyloxy)propyl-(S)-9-[3-hydroxy-2-(phosphonomethoxy)propyl]adenine, sodium salt (HDP-(S)-HPMPA, S.10): Yield ∼455 mg (82%). 1 H NMR (CD3 OD): δ 0.84 (t, 3H), 1.20 (br s, 18H), 1.23 (m, 2H), 1.63 (qt, 2H), 3.67 (m, 2H), 3.20-3.55 (m, 11H), 4.05-4.34 (pair dd, 2H), 8.13 (s, 1H), 8.17 (s, 1H). 31 P NMR: δ 15.18. MS (ES): m/z 586 [M+H]+ , 608 [M+Na]+ , 585 [M–H]– . Anal. calcd. for C28 H52 NaN5 O6 P·0.5H2 O: %C 54.53, %H 8.50, %N 11.36; found: %C 54.70, %H 8.50, %N 11.20. The product can be stored in an air-tight container for at least 12 months at –20◦ C. SUPPORT PROTOCOL 1
SYNTHESIS OF 3-HEXADECYLOXY-1-PROPANOL The in vitro antiviral activity of alkoxyalkyl ANPs strongly depends on the length of the aliphatic alkyl chain. Optimal antiviral activity and selectivity occurs when the alkoxyalkyl ester has a total of about 20 atoms in the lipid chain (Wan et al., 2005), as in the hexadecyloxypropyl group. This protocol describes the synthesis of 3-hexadecyloxy-1propanol (S.2, see Fig. 15.2.3) in two steps from commercial 1-hexadecanol via formation of the methanesulfonate (Crossland and Servis, 1970) and its reaction with the alkoxide anion generated from 1,3-propanediol. The procedure is a straightforward adaptation of the Williamson ether synthesis, one of the most widely used methods for the synthesis of ethers (Baumann, 1967). Formation of the dialkylated product is minimized by the use of excess of 1,3-propanediol.
Materials 1-Hexadecanol (Aldrich) Triethylamine (Aldrich), dry Methanesulfonyl chloride (Aldrich) Dichloromethane (CH2 Cl2 ), dry 0.1 N HCl, cold NaCl, saturated solution MgSO4 , anhydrous 95% ethanol (EtOH) Sodium hydride, 60% dispersion in paraffin oil (Aldrich) N,N-Dimethylformamide (DMF), dry 1,3-Propanediol (Shell Chemical or Aldrich) Hexanes 1-L round-bottom flasks, oven dried 250-mL dropping funnels 2-L beaker Glass filter funnels Separatory funnel Rotary evaporator with vacuum pump Vacuum oil pump 500-mL Erlenmeyer flask Synthesize hexadecyl methanesulfonate 1. In a 1-L round-bottom flask equipped with a stir bar, dissolve 30 g (124 mmol) 1-hexadecanol and 15.2 g (150 mmol) dry triethylamine in 250 mL dry CH2 Cl2 . Cool the mixture in an ice/water bath to 0◦ C.
Synthesis of Cidofovir and (S)-HPMPA Ether Lipid Prodrugs
2. Using a dropping funnel, slowly add a solution of 17.2 g methanesulfonyl chloride in 30 mL CH2 Cl2 and stir 3 hr.
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Figure 15.2.3
Synthesis schemes for the preparation of 3-hexadecyloxy-1-propanol (S.2).
3. Quench the reaction with 2 mL water, and then filter using a glass filter funnel. Rinse the solid with 50 mL CH2 Cl2 and transfer the filtrate into a separatory funnel. 4. Extract three times with 50 mL cold 0.1 N HCl, then one time with 50 mL saturated NaCl. 5. Dry the organic phase over MgSO4 , filter by gravity filtration, and remove the solvents using a rotary evaporator. 6. Crystallize the crude product from 150 mL of 95% EtOH. Collect the product on a filter funnel and keep under vacuum overnight using a vacuum oil pump. Hexadecyl methanesulfonate: Yield 32.4 g (82%). 1 H NMR (CDCl3 ): δ 4.22 (t, 2H), 3.00 (s, 1H), 1.75 (quintet, 2H), 1.26 (broad s, 26H), 0.86 (t, 3H).
Synthesize 3-hexadecyloxy-1-propanol 7. In a 1-L round-bottom flask equipped with a magnetic stir bar, suspend 6.0 g sodium hydride (10 g of a 60% dispersion in paraffin oil) in 500 mL dry DMF. 8. While stirring at room temperature, use a dropping funnel to add 20 mL (300 mmol) 1,3-propanediol over 15 min, then stir an additional 15 min. CAUTION: This results in evolution of H2 .
9. Add 20 g (57 mmol) hexadecyl methanesulfonate in one portion and stir the reaction mixture 18 hr. 10. Carefully add 1 mL water to destroy any excess sodium hydride. Pour the reaction mixture into a 2-L beaker containing 1 L ice water. 11. Collect the solid by filtration through a glass filter funnel and keep under vacuum overnight using a vacuum oil pump. 12. Transfer the crude product to a 500-mL Erlenmeyer flask equipped with a stir bar and add 200 mL hexanes. Stir and heat the mixture on a hot plate until the solid dissolves. Cool the final solution to 4◦ C and keep overnight. Collect the solid by vacuum filtration. 3-Hexadecyloxy-1-propanol (S.2): Yield ∼14.6 g (77%). 1 H NMR (CDCl3 ): δ 0.84 (t, 3H), 1.24 (bs, 26 H), 1.59 (m, 2H), 1.86 (m, 2H), 3.40 (t, 2H), 3.61 (t, 2H), 3.82 (t, 2H). The compound can be stored indefinitely in a tightly closed container at room temperature. Nucleoside Prodrugs and Delivery Strategies
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SUPPORT PROTOCOL 2
LARGE-SCALE SYNTHESIS OF 3-HEXADECYLOXY-1-PROPANOL Bromides can also be used as the alkylating agent in reactions with 1,3-propanediol to form lipid ethers. 1-Bromohexadecane is inexpensive and especially useful when performing larger-scale syntheses of S.2. This alternate procedure also avoids use of sodium hydride, which is difficult and hazardous to use in large-scale preparations. Instead, the alkoxide is generated by dissolving potassium hydroxide in neat 1,3-propanediol, and then distilling under vacuum to remove the water formed.
Materials 1,3-Propanediol (Shell Chemical or Aldrich) 85% potassium hydroxide (KOH; Fisher Scientific) 1-Bromohexadecane (Aldrich) Hexanes 1-L round-bottom flask, oven dried Heating mantle Vacuum distillation apparatus (distillation head) Reflux condenser Bent adapter Thermometer Vacuum pump Nitrogen tank 2-L three-neck flask Overhead mechanical stirrer 250-mL dropping funnel 3-L beaker Glass filter funnel with 2-L filter flask 1. In a 1-L round-bottom flask equipped with a magnetic stir bar, place 500 g 1,3propanediol. 2. While stirring, add 85% KOH and use a heating mantle to heat gently (50◦ C) until the KOH dissolves. Allow the flask to cool. 3. Add a distillation head, thermometer, reflux condenser, and bent adapter to the flask and attach to a vacuum pump. Apply vacuum (∼1 torr) and heat the flask until 1,3propanediol distills at ∼90◦ C. Collect ∼40 mL 1,3-propanediol, remove the heating mantle, and allow the 1,3-propanediol mixture to cool under a nitrogen atmosphere. 4. Transfer the solution to a 2-L three-neck flask equipped with an overhead mechanical stirrer, heating mantle, thermometer, and 250-mL dropping funnel. 5. Stir vigorously while heating the mixture to 125◦ C, and then use the dropping funnel to slowly add (over 30 min) 200 g (0.65 mol) 1-bromohexadecane. Continue stirring 4 hr at 125◦ C. 6. Allow to cool to room temperature. If the mixture solidifies, keep it slightly warm. 7. Without stirring, pour the reaction mixture into a 3-L beaker containing 1000 g crushed ice. Stir gently with a spatula to break up any clumps. 8. When the ice is melted, collect the solid by vacuum filtration on a glass filter funnel, then dry the product completely under vacuum using a vacuum oil pump. Synthesis of Cidofovir and (S)-HPMPA Ether Lipid Prodrugs
9. Transfer the crude product to a 1-L Erlenmeyer flask equipped with a stir bar and add 500 mL hexanes. Stir and heat the mixture on a hot plate until the solid dissolves.
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Cool the final solution to 4◦ C and keep overnight. Collect the solid by vacuum filtration. The anticipated yield of 3-hexadecyloxy-1-propanol (S.2) is ∼140 g (72%) as glistening, waxy plates. The compound can be stored indefinitely in a tightly closed container at room temperature.
COMMENTARY Background Information Significant progress continues to be made in the effort to discover small molecule nucleoside analogs useful for the treatment of viral diseases. One of the most exciting advancements of the past 20 years has been the development of the acyclic nucleoside phosphonates (ANPs), a family of broad-spectrum antiviral agents. ANPs are enzymatically stable nucleoside monophosphate analogs that need two phosphorylations to be converted to their diphosphate forms, which then inhibit viral replication via chain termination or competitive inhibition of viral polymerases. Extensive research efforts initiated by De Clerq and Hol´y’s discovery of (S)-HPMPA (De Clercq et al., 1986) so far have yielded three FDAapproved compounds: (S)-1-(3-hydroxy-2phosphonomethoxypropyl)cytosine (cidofovir, Vistide), 9-(2-phosphosphonomethoxyethyl)adenine (adefovir dipivoxil, Hepsera), and (R)-9-(2-phosphonomethoxypropyl)adenine (tenofovir disoproxil, Viread) for treatment of HCMV, HBV, and HIV, respectively. Excellent reviews covering the chemistry and biological properties of ANPs are available (Hol´y, 2003, 2006; De Clercq and Hol´y, 2005), and protocols for the preparation of several important examples are provided by Dr. Antonin Hol´y in UNIT 14.2. To be active when administered orally, ANPs are typically converted to prodrug forms that mask one or both of the phosphonate negative charges. Without masking, highly polar ANPs have very low oral bioavailability and concentrate in the kidney proximal tubule, leading to nephrotoxicity (Cundy et al., 1996). The marketed oral formulations adefovir dipivoxil (Hepsera) and tenofovir disoproxil fumarate (Viread) employ the bis-pivaloyloxymethyl and bis-(isopropoxycarbonyl)oxymethyl esters, respectively. Several other phosphonatemasking prodrug strategies, including aryl phosphonamidates (Ballatore et al., 2001) and cycloSal esters (Meier and Balzarini, 2006), have been applied to ANPs. HDP-CDV and HDP-(S)-HPMPA are examples of another useful prodrug strategy
where the phosphonate is masked by esterification with a long-chain alkoxyalkyl ether. This modification improves the oral bioavailability of poorly absorbed compounds, presumably because the prodrugs resemble lysophosphatidylcholine, a partially degraded dietary phospholipid (Scow et al., 1967), and are readily absorbed from the gastrointestinal tract into the plasma. The prodrugs are taken up in tissues and metabolized intracellularly, mainly by cleavage of the phosphonoester bond. The ether lipid prodrug strategy was previously applied to acyclovir monophosphate and phosphonoformate, and the results suggest that this approach increases the intestinal absorption of a variety of polar molecules and may lead to better antiviral therapies (Hostetler et al., 2000). Synthesis of HDP-CDV Application of the lipid ether approach to preparing a cidofovir prodrug was stimulated by recent interest in finding an oral therapy for smallpox (variola major), a potential agent of bioterrorism. Several ANPs, including cidofovir and (S)-HPMPA, were known to be active against the orthopoxviruses, but were not active orally (Bray et al., 2000). HDP-CDV was synthesized and its in vitro activity and selectivity against orthopoxviruses and cytomegalovirus was shown to be greatly enhanced relative to the unmodified phosphonate (Beadle et al., 2002; Kern et al., 2002). Further structure-activity studies indicated that optimal selectivity was found with 20-atom alkyl, alkenyl, alkylglyceryl, or alkoxyalkyl groups, such as hexadecyloxypropyl, esterified to CDV (Wan et al., 2005). Investigations into the mechanism of the antiviral enhancement showed that treatment of cells with HDP-CDV leads to much higher levels of intracellular CDV, and consequently to higher levels of cidofovir diphosphate than with unmodified CDV (Aldern et al., 2003). Oral dosing of HDP-CDV resulted in good bioavailability, greatly reduced accumulation of drug in the kidney (Ciesla et al., 2003), and anti-orthopoxvirus efficacy in three lethal animal models of orthopoxvirus
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disease, including ectromelia (Buller et al., 2004; Quenelle et al., 2004; Smee et al., 2004). On the basis of these results, HDP-CDV is currently under development for possible use as an oral treatment in case of a smallpox bioterror attack (Painter and Hostetler, 2004). Basic Protocol 1 involves a dehydrative coupling of 3-hexadecyloxy-1-propanol and cyclic cidofovir (cCDV) using the Mitsunobu reaction (Mitsunobu, 1981). In the second step, the cyclic ester is selectively opened by hydrolysis under mild basic conditions. cCDV must be prepared by using one of the published methods (Bischofberger et al., 1994; Louie and Chapman, 2001). An advantage to using cCDV as a starting material is that the coupled diester product is not charged, which allows simple purification by silica gel chromatography. A drawback is that both the axial and equatorial diastereoisomers are formed, which makes characterization somewhat more difficult. Initial attempts to alkylate cCDV with bromides gave only modest yields, and significant amounts of N4 -alkylated byproducts were also formed and were difficult to separate. An alternative approach was to use the Mitsunobu reaction (Fig. 15.2.1), which was expected to promote reaction of the alcohol with only the acidic pronucleophile (the phosphonate). Using standard Mitsunobu conditions (triphenylphosphine, DIAD), the yield of S.3 increased considerably, to almost 50%, and formation of the N4 -alkylated byproducts was prevented. This type of esterification procedure is most useful when a free phosphonate is available. For example, alkoxyalkyl prodrugs of PMEG, PMEA, PMEDAP, and others have been prepared using the protocols described in UNIT 14.2 followed by coupling with 3-hexadecyloxy-1propanol (Valiaeva et al., 2006).
Synthesis of Cidofovir and (S)-HPMPA Ether Lipid Prodrugs
Synthesis of HDP-(S)-HPMPA Basic Protocol 2, outlined in Figure 15.2.2, illustrates a second type of approach to the synthesis of alkoxyalkyl ANPs in which the lipophilic phosphonoester is incorporated early into a stepwise synthesis. The primary advantage is that any protecting groups used in assembling the ANP also assist in attaching the lipid. Other advantages are that this approach avoids problems of insolubility that are often encountered with highly polar ANPs and coupling methods, and that the presence of the lipophilic alkoxyalkyl moiety allows purification of intermediates and the final product to be accomplished using simple flash chro-
matography on silica gel. More complicated ion-exchange purifications are generally not required. A potential drawback to the stepwise approach is that the free phosphonate, (S)-HPMPA, is not an intermediate and must be synthesized separately if needed. The protocol for HDP-(S)-HPMPA is based on the previous (S)-HPMPA syntheses reported by Webb (1989) and Hol´y and Rosenberg (1987). Both groups prepared (S)-9-(2,3dihydroxypropyl)adenine derivatives with the 3-hydroxyl and 6-amino groups protected, then etherified the 2 -hydroxyl with dialkyl ptoluenesulfonyloxymethylphosphonate. (S)-9(3-Trityloxy-2-hydroxypropyl)-N6 -trityladenine (S.7), prepared by Webb’s procedure with slight modifications, proved to be a superior intermediate for alkylation with S.8, presumably because the trityl group is more stable than the N6 -benzoyl intermediate under the basic conditions of the subsequent alkylation step. Hexadecyloxypropyl toluenesulfonyloxymethylphosphonate is prepared in “one pot” from commercial diethyl toluenesulfonyloxymethylphosphonate by successively treating with TMSBr, oxalyl chloride, 3-hexadecyloxy-1-propanol/pyridine, and finally aqueous NaHCO3 . This stable phosphonate ester can be recrystallized to high purity, stored, and used to synthesize other ANP prodrugs, if desired. Interestingly, triethylamine, used as the solvent, was found to facilitate the alkylation reaction. Longer reaction times and lower yields resulted when the alkylation was attempted in either tetrahydrofuran or N,Ndimethylformamide. Reaction of S.7 and S.8 should provide satisfactory yields of the fully protected (S)HPMPA analog S.9. Occasionally the product may be contaminated with small amounts of unreacted S.8, since the Rf values on silica gel are similar. However, impure S.9 can be deprotected (80% aqueous acetic acid) and then purified easily during the final chromatography. Using Basic Protocol 2, HDP-(S)-HPMPA has been synthesized in 5-gram quantities and studies of its efficacy in antiviral models are underway. In addition to the examples described in this unit, the two synthetic approaches in Basic Protocols 1 and 2 have been adapted to prepare prodrugs of several other ANPs (Valiaeva et al., 2006). Esterification of ANPs with alkoxyalkyl moieties appears to represent a general approach that enhances both the in vitro activity and oral efficacy of this important class of antiviral.
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Critical Parameters and Troubleshooting The synthesis of 3-hexadecyloxy-1-propanol is straightforward and efficient. Careful attention is required during the final recrystallization step to ensure adequate yield. The crude product should be dissolved in a minimal amount of hot hexanes and then cooled and left at 4◦ C overnight for complete crystallization. The Mitsunobu reaction, in addition to requiring strictly anhydrous reagents, requires anhydrous cCDV. Note that both published methods for cCDV yield a crystalline product which is the dihydrate. It is essential to thoroughly remove water of hydration, preferably by azeotropic distillation with toluene followed by drying at 70◦ C in a vacuum oven. Careful TLC using the visualization techniques stated in the protocols is essential to monitoring both the progress of reactions as well as the chromatographic purifications. Knowledge of 31 P NMR is very helpful for final characterization of the compounds. Alkoxyalkyl esters of ANPs, including the ones described in this unit, are quite stable and can be stored for at least a year in a –20◦ C freezer if kept in an air-tight container.
Anticipated Results The yields that can be expected are stated in the protocols for each reaction step. Both Support Protocols provide good yields of the important lipid ether intermediate, 3hexadecyloxy-1-propanol (S.2, 72% to 77%). If large quantities of S.2 are needed, Support Protocol 2 can be used to quickly obtain 100 to 300 g using relatively small glassware. The synthesis of cCDV (S.1) can be accomplished in 82% yield starting from commercially available CDV using the DCC cyclization described in Basic Protocol 1. Alternatively, a procedure reported by Louie and Chapman (2001) claims up to 94% yield and uses inexpensive ethyl chloroformate to effect the cyclization. The coupling of cCDV to S.2 occurs in moderate yield (58%), and the ring-opening hydrolysis and final recrystallization can be accomplished in ∼72% yield. The synthesis of HDP-(S)-HPMPA is more laborious since it starts from adenine. Precursor S.7 can be synthesized from adenine in 46% overall yield in two steps, and the phosphonate synthon S.8 is made in a one-pot reaction from S.2 and diethyl p-toluenesulfonyloxymethylphosphonate in 67% yield. The alkylation of the 2 -hydroxyl of S.7 with S.8 is typically achieved in a Current Protocols in Nucleic Acid Chemistry
modest yield of ∼31% for the protected compound S.9, which yields HDP-(S)-HPMPA after a final deprotection step.
Time Considerations The synthesis of 3-hexadecyloxy-1propanol can be accomplished in 2 or 3 days. Coupling of the lipid to cCDV and hydrolysis to the corresponding alkoxyalkyl CDV usually requires an additional 4 days. Synthesis of the (S)-HPMPA analogs from adenine can be accomplished in about 10 days. If synthesis of more than one (S)-HPMPA analog is planned, it is helpful to prepare a large batch of S.7 and store it in a tightly closed container for subsequent use.
Literature Cited Aldern, K., Ciesla, S.L., Winegarden, K., and Hostetler, K.Y. 2003. Increased antiviral activity of 1-O-hexadecyloxypropyl-[2-14 C]cidofovir in MRC-5 human lung fibroblasts is explained by unique cellular uptake and metabolism. Mol. Pharmacol. 63:678-681. Ballatore, C., McGuigan, C., De Clercq, E., and Balzarini, J. 2001. Synthesis and evaluation of novel amidate prodrugs of PMEA and PMPA. Bioorg. Med. Chem. Lett. 11:1053-1056. Baumann, W.J., Schmid, H.H.O., Ulshofer, H.W., and Mangold, H.K. 1967. Alkoxy lipids. IV. Synthesis and characterization of naturally occurring ethers, esters and ether esters of diols. Biochim. Biophys. Acta 144:355-365. Beadle, J.R., Hartline, C., Aldern, K.A., Rodriguez, N., Harden, E., Kern, E.R., and Hostetler, K.Y. 2002. Alkoxyalkyl esters of cidofovir and cyclic cidofovir exhibit multiple-log enhancement of antiviral activity against cytomegalovirus and herpesvirus replication in vitro. Antimicrob. Agents Chemother. 46:2381-2386. Beadle, J.R., Wan, W.B., Ciesla, S.L., Keith, K.A., Hartline, C., Kern, E.R., and Hostetler, K.Y. 2006. Synthesis and antiviral evaluation of alkoxyalkyl derivatives of 9-(S)-(3-hydroxy-2phosphonomethoxypropyl)adenine against cytomegalovirus and orthopoxviruses. J. Med. Chem. 49:2010-2015. Bischofberger, N., Hitchcock, M.J., Chen, M.S., Barkhimer, D.B., Cundy, K.C., Kent, K.M., Lacy, S.A., Lee, W.A., Li, Z.H., and Mendel, D.B. 1994. 1-[((S)-2-hydroxy-2-oxo-1,4,2dioxaphosphorinan-5-yl)methyl]cytosine, an intracellular prodrug for (S)-1-(3-hydroxy2-phosphonylmethoxypropyl)cytosine with improved therapeutic index in vivo. Antimicrob. Agents Chemother. 38:2387-2391. Bray, M., Martinez, M., Smee, D.F., Kefauver, D., Thompson, E., and Huggins, J. 2000. Cidofovir protects mice against lethal aerosol or intranasal cowpox virus challenge. J. Infect. Dis. 181:1019. Buller, R.M., Owens, G., Schriewer, J., Melman, L., Beadle, J.R., and Hostetler, K.Y. 2004. Efficacy of oral active ether lipid analogs of cidofovir
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in a lethal mousepox model. Virology 318:474481. Ciesla, S.L., Trahan, J., Wan, W.B., Beadle, J.R., Aldern, K.A., Painter, G.R., and Hostetler, K.Y. 2003. Esterification of cidofovir with alkoxyalkanols increases oral bioavailability and diminishes drug accumulation in kidney. Antiviral Res. 59:163-171. Crossland, R. and Servis, K.L. 1970. Facile synthesis of methanesulfonate esters. J. Org. Chem. 35:3195-3196. Cundy, K.C., Bidgood, A.M., Lynch, G., Shaw, J.P., Griffin, L., and Lee, W.A. 1996. Pharmacokinetics, bioavailability, metabolism, and tissue distribution of cidofovir (HPMPC) and cyclic HPMPC in rats. Drug Metab. Dispos. 24:745-752. De Clercq, E. and Hol´y, A. 2005. Case history: Acyclic nucleoside phosphonates: A key class of antiviral drugs. Nat. Rev. Drug Discov. 4:928940. De Clercq, E., Hol´y, A., Rosenberg, I., Sakuma, T., Balzarini, J., and Maudgal, P.C. 1986. A novel selective broad-spectrum anti-DNA virus agent. Nature 323:464-467. Hol´y, A. 2003. Phosphonomethoxyalkyl analogs of nucleotides. Curr. Pharm. Des. 9:2567-2592. Hol´y, A. 2006. Antiviral acyclic nucleoside phosphonates structure activity studies. Antiviral Res. 71:248-253. Hol´y, A. and Rosenberg, I. 1987. Stereospecific syntheses of 9-(S)-(3-hydroxy-2phosphonylmethoxypropyl)adenine (HPMPA). Nucleic Acids Symp. Ser. 18:33-36. Hostetler, K.Y., Beadle, J.R., Hornbuckle, W.E., Bellezza, C.A., Tochkov, I.A., Cote, P.J., Gerin, J.L., Korba, B.E., and Tennant, B.C. 2000. Antiviral activities of oral 1-Ohexadecylpropanediol-3-phosphoacyclovir and acyclovir in woodchucks with chronic woodchuck hepatitis virus infection. Antimicrob. Agents Chemother. 44:1964-1969. Hostetler, K., Aldern, K., Wan, W.B., Ciesla, S.L., and Beadle, J.R. 2006. Alkoxyalkyl esters of (S)-9-[3-hydroxy-2(phosphonomethoxy)propyl]adenine are potent inhibitors of the replication of wild-type and drug-resistant human immunodeficiency virus type 1 in vitro. Antimicrob. Agents Chemother. 50:2857-2859. Kern, E.R., Hartline, C., Harden, E., Keith, K., Rodriguez, N., Beadle, J.R., and Hostetler, K.Y. 2002. Enhanced inhibition of orthopoxvirus replication in vitro by alkoxyalkyl esters of cidofovir and cyclic cidofovir. Antimicrob. Agents Chemother. 46:991-995.
Louie, M. and Chapman, H. 2001. An efficient process for the synthesis of cyclic HPMPC. Nucleosides Nucleotides Nucleic Acids 20:1099-1102. Meier, C. and Balzarini, J. 2006. Application of the cycloSal-prodrug approach for improving the biological potential of phosphorylated biomolecules. Antiviral Res. 71:282-292. Mitsunobu, O. 1981. The use of diethyl azodicarboxylate and triphenylphosphine in synthesis and transformation of natural products. Synthesis 1981:1-28. Painter, G. and Hostetler, K.Y. 2004. Design and development of oral drugs for the prophylaxis and treatment of smallpox infection. Trends Biotechnol. 22:423-427. Quenelle, D.C., Collins, D.J., Wan, W.B., Beadle, J.R., Hostetler, K.Y., and Kern, E.R. 2004. Oral treatment of cowpox virus and vaccinia virus infections in mice with ether lipid esters of cidofovir. Antimicrob. Agents Chemother. 48:404412. Scow, R.O., Stein, Y., and Stein, O. 1967. Incorporation of dietary lecithin and lysolecithin into lymph chylomicrons in the rat. J. Biol. Chem. 242:4919-4924. Smee, D.F., Wong, M., Bailey, K.W., Beadle, J.R., Hostetler, K.Y., and Sidwell, R.W. 2004. Effects of four antiviral substances on lethal vaccinia virus (IHD Strain) respiratory infections in mice. Int. J. Antimicrob. Agents 23:430-437. Valiaeva, N., Beadle, J.R., Aldern, K.A., Trahan, J., and Hostetler, K.Y. 2006. Synthesis and antiviral evaluation of alkoxyalkyl esters of acyclic purine and pyrimidine nucleoside phosphonates against HIV-1 in vitro. Antiviral Res. 72:1019. Wan, W.B., Beadle, J.R., Hartline, C., Kern, E.R., Ciesla, S.L., Valiaeva, N., and Hostetler, K.Y. 2005. Comparison of the antiviral activities of alkoxyalkyl and alkyl esters of cidofovir against human and murine cytomegalovirus replication in vitro. Antimicrob. Agents Chemother. 49:656662. Webb, R.R. II 1989. The bis-trityl route to (S)HPMPA. Nucleosides Nucleotides 8:619-624.
Contributed by James R. Beadle University of California, San Diego La Jolla, California
Synthesis of Cidofovir and (S)-HPMPA Ether Lipid Prodrugs
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Chemistry of bisSATE Mononucleotide Prodrugs
UNIT 15.3
The need to mask the two charged phosphate oxygens of a 5 -mononucleotide to obtain neutral and lipophilic prodrugs (pronucleotides) led to a study of mononucleoside phosphotriesters bearing phosphate-protecting groups. To provide the intracellular delivery of a 5 -mononucleotide from its prodrug, the design of such phosphate-protecting groups was based on general considerations related to enzymatic and chemical stabilities of phosphotriester derivatives. S-Acyl-2-thioethyl (SATE) groups have been studied extensively for efficient esterase-mediated transient phosphate protection of bioactive mononucleotide or phosphonate analogs (P´erigaud et al., 1996, 2000). This unit describes two strategies for preparation of bis(SATE) phosphotriester derivatives. Mononucleoside phosphotriester derivatives can be synthesized following two common strategies (Fig. 15.3.1) using either PIII (hydrogenphosphonate, phosphoramidite, etc.) or PV (phosphodiester, phosphomonoester, etc.) intermediates (Slotin, 1977; Reese, 1978). Initially developed using hydrogenphosphonate chemistry (P´erigaud et al., 1993), the synthesis of a wide range of mononucleoside bis(SATE) phosphotriesters has been carried out by coupling the nucleoside analog with an appropriate phosphoramidite reagent, followed by in situ oxidation (Lefebvre et al., 1995). This PIII strategy is described in Basic Protocol 2. As the phosphoramidite approach necessitates strictly anhydrous conditions, it may not be appropriate for synthesizing the large quantities of SATE pronucleotides required for in vivo studies. An alternative PV strategy (see Alternate Protocol 2) can also be used, where the corresponding 5 -mononucleotide is activated and coupled with the thioester precursor (Lannuzel et al., 1999). For either pathway, preliminary preparation of a thioester precursor is required. These precursors can be obtained from the corresponding
Figure 15.3.1 General scheme for the preparation of bis(SATE) phosphotriester derivatives of nucleoside analogs by the PIII (right) or PV (left) approach. THF, tetrahydrofuran.
Nucleoside Prodrugs and Delivery Strategies
Contributed by Suzanne Peyrottes and Christian P´erigaud
15.3.1
Current Protocols in Nucleic Acid Chemistry (2007) 15.3.1-15.3.13 C 2007 by John Wiley & Sons, Inc. Copyright
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Figure 15.3.2 Preparation of the SATE moiety (S.3a-c; see Basic Protocol 1 and Alternate Protocol 1). DBU, 1,8-diazabicyclo[5.4.0]undec-7-ene.
thioacids or acyl halides, depending on their commercial availability (Fig. 15.3.2; see Basic Protocol 1 and Alternate Protocol 1). CAUTION: Carry out all operations involving organic solvents and reagents in a wellventilated fume hood. Wear appropriate protective clothing and glasses. NOTE: All glassware should be oven dried, and all reactions should be performed under anhydrous conditions. BASIC PROTOCOL 1
TWO-STEP PREPARATION OF SATE MOIETY As very few thioacids are commercially available, a two-step procedure was initially developed for preparation of the SATE moiety using common acyl halides (S.1) as the starting material. These are converted to the corresponding thioacids (S.2) according to published methods (Loeliger and Fluckiger, 1976). The thioacids are then treated with 2-iodoethanol in the presence of 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU; Ono et al., 1978). Depending on the nature of the acyl moiety, the thioesters (S.3) are obtained in good yields after distillation or purification by silica gel chromatography.
Materials Pyridine distilled from KOH Acetone/dry ice bath Dihydrogen sulfide gas (H2 S) Pivaloyl chloride (S.1b) 5 N sulfuric acid Diethyl ether Anhydrous Na2 SO4 Thioacid: e.g., thioacetic acid (S.2a), thiopivaloic acid (S.2b), or thiobenzoic acid (S.2c) Toluene 1,8-Diazabicyclo[5.4.0]undec-7-ene (DBU) Iodoethanol Dichloromethane Ethyl acetate Chemistry of bis(SATE) Mononucleotide Prodrugs
3-L and 100-mL separatory funnels Glass funnel with a cotton bud Rotary evaporator
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High-vacuum distillation apparatus Additional reagents and equipment for column chromatography (APPENDIX 3E) and thin-layer chromatography (TLC; APPENDIX 3D) Prepare thioacids from acyl halides 1. Cool 480 mL pyridine to −30◦ C in an acetone/dry ice bath. While stirring at this temperature, flush with dihydrogen sulfide for 1.5 hr. 2. Add, dropwise, 120 mL of pivaloyl chloride (S.1b) at −30◦ C, then add ∼950 mL of 5 N sulfuric acid to reach a pH of ∼5. 3. Transfer to a 3-L separatory funnel, dilute the mixture with 200 mL diethyl ether, and separate the layers. 4. Dry the organic layer over anhydrous Na2 SO4 , filter using a glass funnel with a cotton bud, and evaporate to dryness on a rotary evaporator. 5. Distill under high vacuum. Thiopivaloic acid (S.2b): 99 g (86% from S.1b). b.p. 40◦ C (29 mmHg). 1 H NMR (300 MHz, CDCl3 ): δ 3.69 (1H, sl, SH), 1.27 (3H, s, CH3 ). FAB-MS (NBA-matrix): 117 [M–H]– .
Prepare thioesters from thioacids 6. Dissolve 1.15 eq of the appropriate thioacid (S.2a-c) in 0.5 mL toluene per mmol thioacid. 7. Cool to 0◦ C (ice bath) and, while stirring, add 1.15 eq DBU followed by 1 eq iodoethanol. 8. Stir the reaction mixture for 3 hr at room temperature. 9. Dilute with dichloromethane, transfer to a 100-mL separatory funnel, and separate the layers. Wash the organic layer twice with water. 10. Dry the organic layer over anhydrous Na2 SO4 , filter using a glass funnel with a cotton bud, and evaporate to dryness on a rotary evaporator. 11. Prepare slurry of silica gel in dichloromethane (for S.3a-b) or toluene (for S.3c). Pour the gel into a chromatography column and carefully layer 1 cm of sand on the top of the slurry. For a reaction performed on a 1-g scale of iodoethanol, use a 3 × 20–cm column.
12. Dissolve the crude product in a minimal amount of dichloromethane (or toluene) and carefully layer on top of the column. Elute with a stepwise gradient of:
0% to 5% ethyl acetate in dichloromethane (for S.3a) 0% to 10% ethyl acetate in dichloromethane (for S.3b) 0% to 10% ethyl acetate in toluene (for S.3c). 13. Monitor fractions by TLC, visualizing the bands using UV and/or iodine. Combine fractions containing pure product and evaporate to dryness on a rotary evaporator without heating the bath. Typically, the ester derivative (coming from the transposition of the corresponding thioester) has the higher Rf value. Thioesters are stable several weeks when stored at −20◦ C under argon atmosphere. S-Acetyl-2-thioethanol (S.3a): 65% from S.2a. TLC (11:7:2, v/v/v, isopropanol/water/ 30% ammonia): Rf = 0.57. 1 H NMR (300 MHz, DMSO-d6 ): δ 4.94 (1H, t, J = 5.5 Hz, OH), 3.45 (2H, m, CH2 O), 2.90 (2H, t, J = 6.6 Hz, SCH2 ), 2.31 (3H, s, CH3 ).
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S-Pivaloyl-2-thioethanol (S.3b): 89% from S.2b. TLC (5:95, v/v, MeOH/CH2 Cl2 ): Rf = 0.61. 1 H NMR (300 MHz, DMSO-d6 ): δ 4.94 (1H, t, J = 5.5 Hz, OH), 3.45 (2H, m, CH2 O), 2.88 (2H, t, J = 6.6 Hz, SCH2 ), 1.16 (9H, s, CH3 ). S-Phenyl-2-thioethanol (S.3c): 68% from S.2c. TLC (8:2, v/v, toluene/EtOAc): Rf = 0.3. H NMR (300 MHz, DMSO-d6 ): δ 7.91 (2H, m, HAr ), 7.68 (1H, m, HAr ), 7.54 (2H, m, HAr ), 5.05 (1H, t, J = 5.4 Hz, OH), 3.57 (2H, m, CH2 O), 3.14 (2H, t, J = 6.5 Hz, SCH2 ).
1
ALTERNATE PROTOCOL 1
ONE-POT PREPARATION OF SATE MOIETY Since the initial procedure (see Basic Protocol 1) presents several drawbacks, including the use of the gaseous and bad-smelling reagent dihydrogen sulfide, an alternative method was developed involving a one-pot reaction starting from commercially available acyl halides. The thiol derivative (i.e., 2-mercaptoethanol) is converted to its corresponding thiolate (Liu et al., 1979; Reibig and Scherer, 1980) in the presence of sodium hydride and then directly condensed with the acyl halide (S.1) to give rise to the thioester derivative (S.3).
Materials 2-Mercaptoethanol Anhydrous diethyl ether Sodium hydride (NaH), 80% suspension in oil (Riedel-de Ha¨en; http://www.riedeldehaen.de/gb/index.html) Pivaloyl chloride or benzoyl chloride Diethyl ether Anhydrous Na2 SO4 500-mL separatory funnel Glass funnel with a cotton bud Rotary evaporator High-vacuum distillation apparatus Additional reagents and equipment for column chromatography (APPENDIX 3E) Prepare thioesters from acyl halides 1. Mix 8.5 mL of 2-mercaptoethanol and 50 mL diethyl ether. Cool to 0◦ C (ice bath). 2. While stirring, add 3.04 g of NaH (80% suspension in oil) in portions of ∼0.5 g. CAUTION: Add the NaH carefully, as dihydrogen gas is produced. During the addition a white precipitate will appear.
3. Stir the reaction mixture for 3 hr, keeping the temperature <5◦ C. 4. Add, dropwise, 10 mL pivaloyl chloride (for S.1b) or 9.4 mL benzoyl chloride (for S.1c). 5. After 30 min stirring, dilute with 100 mL diethyl ether and pour into a 500-mL separatory funnel. 6. Wash the organic layer twice with 50 mL water. Dry the organic layer over anhydrous Na2 SO4 , filter using a glass funnel with a cotton bud, and evaporate to dryness on a rotary evaporator. 7. Purify by column chromatography (see Basic Protocol 1, steps 11 to 13) or distill under high vacuum. Chemistry of bis(SATE) Mononucleotide Prodrugs
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S-Pivaloyl-2-thioethanol (S.3b): 10.6 g (80% from S.1b). TLC (5:95, v/v, MeOH/CH2 Cl2 ): Rf = 0.61. b.p. 50◦ to 52◦ C (0.035 mbar). S-Phenyl-2-thioethanol (S.3c): 11.1 g (75% from S.1c). TLC (8:2, v/v, toluene/EtOAc): Rf = 0.3. b.p. 96◦ to 98◦ C (0.035 mbar). Current Protocols in Nucleic Acid Chemistry
PREPARATION OF BIS(SATE) PHOSPHOTRIESTER DERIVATIVES OF AZT USING PHOSPHORAMIDITE INTERMEDIATES
BASIC PROTOCOL 2
This protocol is based on the use of 3 -azido-2 ,3 -dideoxythymidine (AZT) as a nucleosidic model; however, similar pathways have been applied to various nucleoside analogs, and related data can be gleaned from the literature (Benzaria et al., 1996; Groschel et al., 1999; P´erigaud et al., 1999; Placidi et al., 2001; Egron et al., 2002; Bazzanini et al., 2005). Commonly used in oligonucleotide synthesis, PIII chemistry is the most efficient method to prepare mononucleoside phosphotriester derivatives, which are obtained in a single step under mild conditions and with good yields. The bis(SATE) phosphoramidite intermediates (Fig. 15.3.3, S.4a-c) are prepared from commercially available N,N-diisopropylphosphoramidous dichloride and the corresponding thioester precursors (S.3a-c). The nucleoside analog is introduced in the last step, precluding the loss of valuable material. Coupling of the nucleoside analog is performed in the presence of 1H-tetrazole and is followed by in situ oxidation to yield the product S.5a-c.
Materials N,N-Diisopropylphosphoramidous dichloride (Aldrich) Anhydrous tetrahydrofuran (THF, Fluka) Argon source Acetone/dry ice bath Thioester precursor (S.3a-c; see Basic Protocol 1 or Alternate Protocol 1) Triethylamine (TEA) distilled from CaH2 Cyclohexane Ethyl acetate (EtOAc) TLC spray reagents: 5% (v/v) sulfuric acid (H2 SO4 ) in ethanol (for nucleosidic derivatives) Molybden-blue solution (for phosphorus-containing derivatives; Sigma or see recipe) Silica gel (Macherey-Nagel Kieselgel 60): 0.063- to 0.20-mm (for phosphoramidite intermediates) and 0.040- to 0.063-mm (for phosphotriester products) 3 -Azido-2 ,3 -dideoxythymidine (AZT) Phosphorus pentoxide (P2 O5 ) Acetonitrile distilled from CaH2 0.45 M 1H-tetrazole in acetonitrile (Fluka) 3-Chloroperbenzoic acid Dichloromethane 10% (w/v) aq. sodium sulfite (NaHSO3 ) Saturated aq. sodium bicarbonate (NaHCO3 ) Anhydrous sodium sulfate (Na2 SO4 ) Toluene Methanol TLC plate: silica-coated aluminum plate with fluorescent indicator (Merck silica gel 60 F254 ) Cotton bud Glass funnel 3 × 10– and 3 × 20–cm chromatography columns Vacuum desiccator Rotary evaporator equipped with a vacuum pump 100-mL separatory funnel Additional reagents and equipment for column chromatography (APPENDIX 3E) and thin-layer chromatography (TLC; APPENDIX 3D)
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Figure 15.3.3 Preparation of bis(SATE) phosphotriester derivatives of AZT (see Basic Protocol 2). AZT, 3 -azido-2 ,3 -dideoxythymidine; mcPBA, 3-chloroperbenzoic acid; THF, tetrahydrofuran.
Prepare bis(SATE) phosphoramidite intermediates 1. Dissolve 1.01 g (5 mmol) of N,N-diisopropylphosphoramidous dichloride in 35 mL THF. Apply an argon atmosphere and cool to −78◦ C in an acetone/dry ice bath. 2. Prepare a solution of 10 mmol of the appropriate thioester precursor (S.3a-c) and 3.07 mL triethylamine in 25 mL THF and add it dropwise to the first reaction mixture. After addition is complete, stir 2 to 3 hr at room temperature. Monitor the reaction by TLC using 8:1:1 (v/v/v) cyclohexane/ethyl acetate/triethylamine. To avoid decomposition of the product on silica, elute TLC plates once before spotting. The desired compound has the upper Rf value.
3. Filter off the triethylammonium hydrochloride. Perform filtration through a cotton bud (using a glass funnel) under argon atmosphere to avoid decomposition of the product. 4. Evaporate the filtrate and coevaporate once with 50 mL cyclohexane to obtain an oily residue. 5. Prepare a slurry of silica gel for flash chromatography in 95:5 (v/v) cyclohexane/triethylamine. Pour the gel into a 3 × 10–cm chromatography column and carefully layer 1 cm of sand on the top of the slurry. 6. Dissolve the crude product in a minimal amount of cyclohexane and carefully layer on top of the column. Elute with a stepwise gradient of 0% to 5% ethyl acetate in cylohexane containing 1% triethylamine. 7. Monitor fractions by TLC, combine fractions containing pure product, and evaporate to dryness using a rotary evaporator to obtain the phosphoramidite as an oil. Chemistry of bis(SATE) Mononucleotide Prodrugs
Phosphoramidite intermediates are stable several months when stored at −20◦ C under argon atmosphere.
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Bis(S-acetyl-2-thioethyl) N,N-diisopropylphosphoramidite (S.4a): 1.33 g (72% from S.3a). TLC (9:1:1, v/v/v, cyclohexane/EtOAc/TEA): Rf = 0.48. 1 H NMR (300 MHz, DMSO-d6 ): δ 3.59 (6H, m, CH2 O, CHMe2 ), 3.04 (4H, t, J = 6.4 Hz, SCH2 ), 2.32 (6H, s, CH3 CO), 1.10 (12H, d, J = 6.8 Hz, CH3 ). 31 P NMR (DMSO-d6 ): δ 147.9. FAB-MS (thioglycerol-matrix): 370 [M+H]+ . Bis(S-pivaloyl-2-thioethyl) N,N-diisopropylphosphoramidite (S.4b): 1.68 g (74% from S.3b). TLC (8:1:1, v/v/v, cyclohexane/EtOAc/TEA): Rf = 0.53. 1 H NMR (300 MHz, DMSO-d6 ): δ 3.59 (6H, m, CH2 O, CHMe2 ), 3.01 (4H, t, J = 6.3 Hz, SCH2 ), 1.16 (18H, s, (CH3 )3 C), 1.10 (12H, d, J = 6.8 Hz, C(CH3 )2 ). 31 P NMR (DMSO-d6 ): δ 148. FAB-MS (thioglycerol-matrix): 454 [M+H]+ , 145 [tBuCOS(CH2 )2 ]+ . Bis(S-benzoyl-2-thioethyl) N,N-diisopropylphosphoramidite (S.4c): 0.99 g (40% from S.3c). TLC (8.5:0.5:1, v/v/v, cyclohexane/EtOAc/TEA): Rf = 0.45. 1 H NMR (300 MHz, DMSO-d6 ): δ 7.89 (4H, m, HAr ), 7.68 (2H, m, HAr ), 7.53 (4H, m, HAr ), 3.65 (6H, m, CH2 O, CHMe2 ), 3.27 (4H, t, J = 6.3 Hz, SCH2 ), 1.10 (12H, d, J = 6.8 Hz, C(CH3 )2 ). 31 P NMR (DMSO-d6 ): δ 148. FAB-MS (thioglycerol-matrix): 494 [M+H]+ .
Perform coupling reaction 8. Dissolve 0.27 g of AZT and 1.2 eq of the appropriate phosphoramidite (S.4a-c) in acetonitrile (2 mL/mmol of nucleoside analog). Apply argon atmosphere. Prior to use, the nucleoside derivative must be dried in a desiccator overnight under reduced pressure and in the presence of P2 O5 .
9. While stirring, add 3 eq of 1H-tetrazole (0.45 M solution in acetonitrile). Continue stirring 1 hr. 10. Cool the reaction mixture to −40◦ C in an acetone/dry ice bath and add 0.23 g of 3-chloroperbenzoic acid dissolved in 2.5 mL dichloromethane. 11. Allow to warm to room temperature and add 1.3 mL of 10% (w/v) aq. sodium sulfite. 12. Transfer the mixture to a 100-mL separatory funnel and separate the organic and aqueous layers. 13. Wash the aqueous layer twice with 10 mL dichloromethane. Combine the organic layers and wash once with 5 mL saturated aq. NaHCO3 , then twice with 5 mL water. 14. Dry the organic layer over anhydrous Na2 SO4 , filter using a glass funnel with a cotton bud, and evaporate to dryness on a rotary evaporator. 15. Prepare a slurry of silica gel in dichloromethane. Pour the gel into a 3 × 20–cm chromatography column and carefully layer 1 cm of sand on the top of the slurry. 16. Dissolve the crude product in a minimal amount of dichloromethane and carefully layer on top of the column. Elute with a stepwise gradient of:
0% to 3% methanol in dichloromethane (for S.5a,b) 0% to 2% methanol in dichloromethane (for S.5c). 17. Monitor fractions by TLC using 95:5 (v/v) dichloromethane/methanol and combine fractions containing pure product. 18. Evaporate combined fractions to dryness in a rotary evaporator and dry overnight under vacuum. Phosphotriester derivatives are stable several years when stored at −20◦ C under argon atmosphere. 3 -Azido-2 ,3 -dideoxythymidin-5 -yl bis(S-acetyl-2-thioethyl)phosphate (S.5a): 0.42 g (75% from AZT). TLC (95:5, v/v, CH2 Cl2 /MeOH): Rf = 0.43. 1 H NMR (300 MHz, DMSO-d6 ): δ 11.4 (1H, bs, NH), 7.46 (1H, s, H6), 6.12 (1H, t, J = 6.7 Hz, H1 ), 4.50-4.40
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(1H, m, H3 ), 4.23-4.17 (2H, m, H5 ), 4.10-4.00 ( 5H, m, CH2 O, H4 ), 3.12 (4H, t, J = 6.3 Hz, SCH2 ), 2.44-2.33 (8H, m, CH3 CO, H2 ), 1.78 (3H, s, CH3 ). 31 P NMR (DMSO-d6 ): δ0−0.72. FAB-MS (thioglycerol-matrix): 552 [M+H]+ , 1101 [2M–H]– , 550 [M–H]– . 3 -Azido-2 ,3 -dideoxythymidin-5 -yl bis(S-pivaloyl-2-thioethyl)phosphate (S.5b): 0.42 g (65% from AZT). TLC (95:5, v/v, CH2 Cl2 /MeOH): Rf = 0.42. 1 H NMR (300 MHz, DMSOd6 ): δ 11.4 (1H, bs, NH), 7.47 (1H, s, H6), 6.12 (1H, t, J = 6.6 Hz, H1 ), 4.50-4.43 (1H, m, H3 ), 4.20-4.17 (2H, m, H5 ), 4.08-3.98 ( 5H, m, CH2 O, H4 ), 3.10 (4H, t, J = 6.3 Hz, SCH2 ), 2.43-2.31 (2H, m, H2 ), 1.78 (3H, s, CH3 ), 1.16 (18H, s, C(CH3 )3 ). 31 P NMR (DMSO-d6 ): δ −0.70. FAB-MS (thioglycerol-matrix): 636 [M+H]+ , 634 [M–H]– . 3 -Azido-2 ,3 -dideoxythymidin-5 -yl bis(S-benzoyl-2-thioethyl)phosphate (S.5c): 0.40 g (58% from AZT). TLC (95:5, v/v, CH2 Cl2 /MeOH): Rf = 0.47. 1 H NMR (300 MHz, DMSOd6 ): δ 11.3 (1H, bs, NH), 7.90-7.50 (10H, m, HAr ), 7.45 (1H, s, H6), 6.09 (1H, t, J = 6.7 Hz, H1 ), 4.48-4.41 (1H, m, H3 ), 4.25-4.16 (6H, m, CH2 O, H5 ), 4.01-3.98 (1H, m, H4 ), 3.38-3.35 (4H, m, SCH2 ), 2.42-2.22 (2H, m, H2 ), 1.75 (3H, s, CH3 ). 31 P NMR (DMSO-d6 ): δ −0.63. FAB-MS (thioglycerol-matrix): 676 [M+H]+ , 1349 [2M–H]– , 674 [M–H]– . ALTERNATE PROTOCOL 2
PREPARATION OF BIS(tBuSATE) PHOSPHOTRIESTER DERIVATIVE OF AZT FROM THE 5 -MONOPHOSPHATE Preliminary results of the in vitro evaluation of bis(SATE) phosphotriesters prompted the development of a method for synthesizing larger quantities of a selected derivative for evaluation in animal models. Among the SATE transient phosphate-protecting groups, the S-pivaloyl-2-thioethyl (tBuSATE) group was selected for its resistance to enzymatic hydrolysis (Lefebvre et al., 1995). In this respect, the two different synthetic approaches using PIII and PV intermediates were compared (Lannuzel et al., 1999). For large-scale synthesis, the phosphoramidite pathway did not seem to be effective due to the strictly anhydrous conditions required as well as the formation of byproducts. On the other hand, the monophosphate approach appeared more appropriate. This method, illustrated in Figure 15.3.4, first requires the synthesis of the 5 -monophosphate of AZT (AZTMP, S.7), which is a stable intermediate. The mononucleotide is activated by TPS-Cl and then coupled with the thioester precursor S.3b, giving rise to a single product S.5b, which is easily purified by silica gel chromatography.
Materials
Chemistry of bis(SATE) Mononucleotide Prodrugs
3 -Azido-2 ,3 -dideoxythymidine (AZT; S.6) Pyridine Trimethyl phosphate distilled from BaO Phosphorus oxychloride, distilled 1 M triethylammonium bicarbonate (TEAB) buffer, pH 7 (see recipe) Toluene Diethyl ether DEAE-Sephadex A25 gel Dowex 50WX2 ion-exchange resin (H+ form) TLC spray reagents: 5% (v/v) sulfuric acid (H2 SO4 ) in ethanol (for nucleosidic derivatives) Molybden-blue solution (for phosphorus containing derivatives; Sigma or see recipe) S-Pivaloyl-2-thioethanol (S.3b; see Basic Protocol 1 or Alternate Protocol 1) Triisopropylbenzenesulfonyl chloride (TPS-Cl) Aq. sodium bicarbonate (NaHCO3 ) Dichloromethane Anhydrous Na2 SO4 Methanol
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Figure 15.3.4 Preparation of the S.5b AZT derivative using Alternate Protocol 2. TEAB, triethylammonium bicarbonate; TPS-Cl, triisopropylbenzenesulfonyl chloride.
100-mL and 500-mL separatory funnels Rotary evaporator 3 × 20–cm low-pressure chromatography column Lyophilizer 1 × 20– and 5 × 30–cm chromatography columns Vacuum oil pump Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) Prepare 5 -monophosphate of AZT 1. Coevaporate 0.2 g (0.75 mmol) AZT (S.6) twice with pyridine. 2. Dissolve in 7.5 mL trimethyl phosphate. Cool to 0◦ C (ice bath). 3. While stirring, add, dropwise, 0.15 g (0.98 mmol) phosphorus oxychloride. Stir for 3 hr at 0◦ C. 4. Quench the reaction by adding 20 mL of 1 M TEAB buffer, pH 7. 5. Pour the mixture into a 100-mL separatory funnel. 6. Extract the aqueous layer twice with 20 mL toluene and then twice with 20 mL diethyl ether. Evaporate the aqueous layer to dryness on a rotary evaporator. 7. Prepare a slurry of DEAE-Sephadex A25 gel in water. Pour the gel into a 3 × 20–cm low-pressure chromatography column. 8. Dissolve the crude product in a minimal amount of water and carefully layer on top of the column. Start eluting with 100 mL water, then with a linear gradient of 0% to 100% 0.2 M TEAB buffer. Collect 50-mL fractions and monitor by TLC.
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9. Combine fractions containing pure product, evaporate to remove most of the TEAB, and then lyophilize. 10. Prior to use in the coupling reaction, prepare a slurry of Dowex 50WX2 ion-exchange resin (H+ form) in water. Pour the gel into a 1 × 20–cm chromatography column. 11. Dissolve AZTMP (5 -monophosphate of AZT) in a minimal amount of water and carefully layer on top of the column. Elute with water and collect 10-mL fractions. Monitor by TLC, combine desired fractions, and lyophilize. 5 -O-Phosphoryl-3 -azido-2 ,3 -dideoxythymidine (S.7): TLC (11:7:2, v/v/v, isopropanol/water/30% ammonia): Rf = 0.57. 1 H NMR (300 MHz, D2 O): δ 7.82 (1H, s, H6), 6.30 (1H, t, J = 6.8 Hz, H1 ), 4.56-4.50 (1H, m, H3 ), 4.22-4.19 ( 1H, m, H4 ), 4.10-4.03 (2H, m, H5 ), 2.54-2.49 (2H, m, H2 ), 1.95 (3H, s, CH3 ). 31 P NMR (D2 O): δ 2.6. FAB-MS (thioglycerol): 346 [M–H]– .
Perform coupling reaction 12. Prepare a mixture of 0.67 g (1.93 mmol) S.7 and 1.57 g (9.65 mmol) S.3b and coevaporate twice with dry pyridine. Dissolve the residue in 15 mL dry pyridine. 13. While stirring, add 2.92 g (9.65 mmol) TPS-Cl. Continue stirring the reaction mixture overnight at room temperature. 14. Quench the reaction by adding 100 mL aq. NaHCO3 . Dilute with 200 mL dichloromethane and transfer to a 500-mL separatory funnel. 15. Wash the organic layer twice with 100 mL water. 16. Dry the organic layer over anhydrous Na2 SO4 , filter off the drying agent, and evaporate the filtrate to dryness on a rotary evaporator. Coevaporate with toluene. 17. Prepare a slurry of silica gel in dichloromethane. Pour the gel into a 5 × 30–cm chromatography column and carefully layer 1 cm of sand on the top of the slurry. 18. Dissolve the crude product in a minimal amount of dichloromethane and carefully layer on top of the column. Start eluting with 500 mL dichloromethane, then with 2% methanol in dichloromethane. Collect 100-mL fractions. 19. Combine fractions containing pure product S.5b as determined by TLC. 20. Evaporate the appropriate fractions to dryness in a rotary evaporator and dry overnight under high vacuum with a vacuum oil pump. Yield S.5b: 0.83 g (68% from S.7).
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Molybden-blue solution Solution A: Boil 40.11 g of molybdenum(VI) oxide (MoO3 ) in 1 L of 25 N sulfuric acid until the MoO3 is completely dissolved (3 to 4 hr). Let the light yellow solution slowly cool to ambient temperature overnight (the solution will turn light blue). Solution B: Boil 1.78 g of molybdenum powder and 500 mL of solution A for 15 min, cool, and decant from the remaining residue. Chemistry of bis(SATE) Mononucleotide Prodrugs
continued
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Spray reagent: Add equal volumes of solutions A and B to 4 volumes of water. Solutions A and B are stable for several months when stored in the dark; the spray reagent has to be prepared about weekly. This TLC spray reagent is prepared according to Dittmer and Lester (1964).
Triethylammonium bicarbonate (TEAB) buffer, pH 7 Prepare a 1 L aqueous solution of 1 M triethylamine, and chill by stirring on an ice bath. While stirring, bubble carbon dioxide gas through the solution until the pH reaches 7. Store in a stoppered bottle up to 6 months at 4◦ C. Monitor the pH after long periods of storage (>1 month).
COMMENTARY Background Information
Initially investigated with 3 -azido-2 ,3 dideoxythymidine (AZT) as a nucleoside model (Lefebvre et al., 1995), the SATE approach has been applied to anti–human immunodeficiency virus (anti-HIV) nucleoside analogs that are limited by the first phosphorylation step, either through a dependence on kinase-mediated phosphorylation or by a rate-limiting step in the anabolism pathway (P´erigaud et al., 1994; Girardet et al., 1995; Shirasaka et al., 1995; ThumannSchweitzer et al., 1996; Valette et al., 1996). In all cases, in vitro evaluation of the corresponding bis(SATE) phosphotriester derivatives shows increased antiviral activities compared to the parent nucleosides. For example, the SATE pronucleotides of 2 ,3 -dideoxyadenosine (ddA) exhibit a very potent antiretroviral effect in human lymphoblastoid cells (CEM-SS, MT-4) and in stimulated and unstimulated primary cultured human cell lines such as peripheral blood mononuclear (PBM) cells or monocytederived macrophages (P´erigaud et al., 1994; Shirasaka et al., 1995; Thumann-Schweitzer et al., 1996). Measured in terms of selectivity index (SI), these bis(SATE) phosphotriesters emerged as very potent and selective inhibitors of HIV and SIV replication in vitro. Thus, when applied to an antiviral nucleoside analog hampered at the first phosphorylation step, the bis(SATE) pronucleotide approach leads to enhanced in vitro antiviral efficiency. Investigation of SATE biolabile phosphate protection was also extended to the 5 -mononucleotide of 9-(2-hydroxyethoxymethyl)guanine (acyclovir, ACV). This nucleoside analog is currently used in the treatment of herpes simplex virus (HSV) and varicella zoster virus (VZV) infections. In order to exploit the potency of ACVTP as an anti–hepatitis B virus (anti-HBV) agent, two
bis(SATE) phosphotriester derivatives of ACV have been evaluated for their inhibitory effects on the replication of HBV in human hepatoblastoma-derived liver Hep-G2 cells (2.2.15 cells) transfected with human HBV DNA, as well as in primary duck hepatocyte cultures infected with the duck virus (DHBV; Hantz et al., 1999; P´erigaud et al., 1999). DHBV is a model used largely for the screening and evaluation of new anti-HBV agents both in vitro and in vivo in chronically infected animals. In contrast to ACV, the corresponding bis(SATE) phosphotriesters exhibited a potent and selective in vitro anti-HBV activity in these experiments, proving that the use of this pronucleotide approach can extend the antiviral spectrum of particular nucleoside analogs. Compared to the established 2.2.15 cell line, the greater activity observed in primary cells (in which a more dynamic replication occurs and high virus yields are achieved rapidly) may reflect an increased conversion of these derivatives to their phosphorylated forms. Differences in the in vitro antiviral activity of bis(SATE) pronucleotides have been associated with multiple factors including lipophilicity, the decomposition kinetics of the prodrug (in culture medium and inside cells), as well as the particular metabolism of the parent nucleoside (P´erigaud et al., 2000). In cell culture experiments, comparative evaluation of pronucleotides bearing different SATE groups have generally shown that phosphotriesters incorporating the lipophilic and enzymatically resistant S-pivaloyl-2-thioethyl (tBuSATE) group exhibit similar or superior antiviral effects compared to the more labile S-acetyl-2-thioethyl (MeSATE) phosphate group. Besides monomeric nucleotides, this prodrug approach could also be developed to modify numerous phosphorylated effectors such as oligomeric derivatives (i.e., oligonucleotides),
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as well as phosphopeptides. In this respect, the SATE approach has been applied to improve therapeutic applications of antisense oligonucleotides by temporary masking of the negative charges of the sugar-phosphate backbone with biolabile protecting groups. Such pro-oligonucleotides are hoped to enter cells, where esterases trigger the release of the parent oligonucleotide (Morvan et al., 2000). Phosphopeptide prodrugs, bearing O-phosphate protected by two SATE groups (tBuSATE and MeSATE; Math´e et al., 1998), were also designed and shown to circumvent the poor cellular permeability of the parent phosphopeptide in cellular assays (Liu et al., 2000).
Critical Parameters and Troubleshooting Phosphoramidite intermediates and the phosphorodichloridite precursor are sensitive to moisture; consequently, they should be stored and handled under a dry argon atmosphere. Experience with chemical laboratory techniques such as distillation, TLC, and column chromatography is needed. Due to the low stability of phosphoramidite intermediates as well as the thioester precursors on silica, their purification should be performed quickly and the products should not stand more than a day on the chromatography column itself. Concerning the thio-compounds, used glassware should be washed thoroughly with diluted bleach solution, and all aqueous residues should be quenched with bleach. Depending of the nucleoside analog, the corresponding prodrugs may be obtained as oils or solids. They should be stored at −20◦ C, as should their diluted solutions. BisSATE derivatives are not stable in basic or strongly acidic media.
Anticipated Results
Chemistry of bis(SATE) Mononucleotide Prodrugs
The protocols described here for AZT and phosphoramidite intermediates can be applied to a large variety of nucleoside analogs. Depending on the solubility of the nucleoside analogs, tetrahydrofuran, acetonitrile, or dimethylformamide could also be used during the coupling step. 3-Chloroperbenzoic acid can be replaced by tert-butyl hydroperoxide during the oxidation step. Using the PIII approach, the expected yield for the coupling step should be between 55% to 75%. Purity of the resulting prodrug should be checked using common spectroscopic methods (NMR, HPLC) as well as elemental analysis.
Time Considerations The synthesis of one mononucleotide prodrug can be accomplished in two weeks, including distillation of reagents and solvents. In order to save time, anhydrous solvents may be purchased.
Literature Cited Bazzanini, R., Gouy, M.H., Peyrottes, S., Gosselin, G., P´erigaud, C., and Manfredini, S. 2005. Synthetic approaches to a mononucleotide prodrug of cytarabine. Nucleosides Nucleotides Nucleic Acids 24:1635-1649. Benzaria, S., Pelicano, H., Johnson, R., Maury, G., Imbach, J.L., Aubertin, A.M., Obert, G., and Gosselin, G. 1996. Synthesis, in vitro antiviral evaluation, and stability studies of bis(S-acyl-2-thioethyl) ester derivatives of 9-[2(phosphonomethoxy)ethyl]adenine (PMEA) as potential PMEA prodrugs with improved oral bioavailability. J. Med. Chem. 39:4958-4965. Dittmer, J.C. and Lester, R.L. 1964. A simple, specific spray for the detection of phospholipids on thin-layer chromatograms. J. Lipid Res. 5:126127. Egron, D., P´erigaud, C., Gosselin, G., Aubertin, A.M., Gatanaga, H., Mitsuya, H., Zemlicka, J., and Imbach, J.L. 2002. Increase of the adenallene anti-HIV activity in cell culture using its bis(tBuSATE) phosphotriester derivative. Bioorg. Med. Chem. Lett. 12:265-266. Girardet, J.-L., P´erigaud, C., Aubertin, A.M., Gosselin, G., Kirn, A., and Imbach, J.-L. 1995. Increase of the anti-HIV activity of D4T in human T-cell culture by the use of the sate pronucleotide approach. Bioorg. Med. Chem. Lett. 5:2981-2984. Groschel, B., Himmel, N., Cinatl, J., Perigaud, C., Gosselin, G., Imbach, J.L., Doerr, H.W., and Cinatl, J. Jr. 1999. ddC- and 3TC-bis(SATE) monophosphate prodrugs overcome cellular resistance mechanisms to HIV-1 associated with cytidine kinase deficiency. Nucleosides Nucleotides 18:921-926. Hantz, O., P´erigaud, C., Borel, C., Jamard, C., Zoulim, F., Tr´epo, C., Imbach, J.-L., and Gosselin, G. 1999. The SATE pronucleotide approach applied to acyclovir: Part II. Effects of bis(SATE)phosphotriester derivatives of acyclovir on duck hepatitis B virus replication in vitro and in vivo. Antiviral Res. 40:179. Lannuzel, M., Egron, D., Imbach, J.L., Gosselin, G., and Perigaud, C. 1999. Synthesis of the tBuSATE pronucleotide of AZT by two different synthetic approaches. Nucleosides Nucleotides 18:1001-1002. Lefebvre, I., P´erigaud, C., Pompon, A., Aubertin, A.M., Girardet, J.L., Kirn, A., Gosselin, G., and Imbach, J.L. 1995. Mononucleoside phosphotriester derivatives with S-acyl-2-thioethyl bioreversible phosphate protecting groups. Intracellular delivery of 3 -azido-2 ,3 -dideoxythymidine 5 -monophosphate. J. Med. Chem. 38:39413950.
15.3.12 Supplement 29
Current Protocols in Nucleic Acid Chemistry
Liu, H.J., Lee, S.P., and Chan, W.H. 1979. A facile preparation of thiol esters from carboxylic acids. Synth. Comm. 9:91-96. Liu, W.Q., Vidal, M., Math´e, C., P´erigaud, C., and Garbay, C. 2000. Inhibition of the RasDependant mitogenic pathway by phosphopeptide prodrugs with antiproliferative properties. Bioorg. Med. Chem. Lett. 10:669-672. Loeliger, P. and Fluckiger, E. 1976. Sulfide contraction via alkylative coupling: 3-methyl-2,4heptanedione. Org. Synth. 55:127. Math´e, C., P´erigaud, C., Gosselin, G., and Imbach, J.L. 1998. Phosphopeptide prodrug bearing an Sacyl-2-thioethyl enzyme-labile phosphate protection. J. Org. Chem. 63:8547-8550. Morvan, F., Vasseur, J.J., Viv`es, E., Rayner, B., and Imbach, J.L. 2000. The oligonucleotide prodrug approach: The pro-oligonucleotides. In Pharmaceutical Aspects of Oligonucleotides (P. Couvrer and C. Malvy, eds.) pp. 79-97. Taylor & Francis, London. Ono, N., Yamada, T., Saito, T., Tanaka, K., and Kaji, A. 1978. A convenient procedure for esterification of carboxylic acids. Bull. Chem. Soc. Jpn. 51:2401-2404. P´erigaud, C., Gosselin, G., Lefebvre, I., Girardet, J.L., Benzaria, S., Barber, I., and Imbach, J.L. 1993. Rational design for cytosolic delivery of nucleoside monphosphates: “SATE” and “DTE” as enzyme-labile transient phosphate protecting groups. Bioorg. Med. Chem. Lett. 3:25212526. P´erigaud, C., Aubertin, A.-M., Benzaria, S., Pelicano, H., Girardet, J.-L., Maury, G., Gosselin, G., Kirn, A., and Imbach, J.-L. 1994. Equal inhibition of the replication of human immunodeficiency virus in human T-cell culture by ddA bis(SATE)phosphotriester and 3 -azido2 ,3 -dideoxythymidine. Biochem. Pharmacol. 48:11. P´erigaud, C., Girardet, J.-L., Gosselin, G., and Imbach, J.-L. 1996. Comments on nucleotide delivery forms. In Advances in Antiviral Drug Design (E. De Clercq, ed.) pp. 147-172. JAI Press, London. P´erigaud, C., Gosselin, G., Girardet, J.-L., Korba, B.E., and Imbach, J.-L. 1999. The S-acyl2-thioethyl pronucleotide approach applied to acyclovir: Part I. Synthesis and in vitro antihepatitis B virus activity of bis(S-acyl-2thioethyl)phosphotriester derivatives of acyclovir. Antiviral Res. 40:167. P´erigaud, C., Gosselin, G., and Imbach, J.-L. 2000. Anti-HIV phosphotriester pronucleotides:
Basis for the rational design of biolabile phosphate protecting groups. In Biomedical Chemistry. Applying Chemical Principles to the Understanding and Treatment of Disease (P.F. Torrence, ed.) pp. 115-141. John Wiley & Sons, New York. Placidi, L., Faraj, A., Loi, A.G., Pierra, C., Egron, D., Cretton-Scott, E., Gosselin, G., P´erigaud, C., Martin, L.T., Schinazi, R.F., Imbach, J.-L., El Kouni, M.H., Bryant, M.L., and Sommadossi, J.-P. 2001. Antiviral activity and intracellular metabolism of bis(tbutylSATE) phosphotriester of β-L-2 ,3 dideoxyadenosine, a potent inhibitor of HIV and HBV replication. Antivir. Chem. Chemother. 12:99-108. Reese, C.B. 1978. The chemical synthesis of oligoand poly-nucleotides by the phosphotriester approach. Tetrahedron 34:3143-3179. Reibig, H.V. and Scherer, B., 1980. A simple synthesis of thiol esters from copper-I-mercaptides and acyl chlorides. Tetrahedron Lett. 21:42594262. Shirasaka, T., Chokekijchai, S., Yamada, A., Gosselin, G., Imbach, J.-L., and Mitsuya, H. 1995. Comparative analysis of anti-human immunodeficiency virus type 1 activities of dideoxynucleoside analogs in resting and activated peripheral blood mononuclear cells. Antimicrob. Agents Chemother. 39:2555. Slotin, L.A. 1977. Current methods of phosphorylation of biological molecules. Synthesis 737752. Thumann-Schweitzer, C., Gosselin, G., P´erigaud, C., Benzaria, S., Girardet, J.-L., Lefebvre, I., Imbach, J.-L., Kirn, A., and Aubertin, A.-M., 1996. Anti-human immunodeficiency virus type 1 activities of dideoxynucleoside phosphotriester derivatives in primary monocytes/macrophages. Res. Virol. 147:155. Valette, G., Pompon, A., Girardet, J.-L., Cappellacci, L., Franchetti, P., Grifantini, M., La Colla, P., Loi, A.G., P´erigaud, C., Gosselin, G., and Imbach, J.-L. 1996. Decomposition pathways and in vitro HIV inhibitory effects of isoddA pronucleotides: Toward a rational approach for intracellular delivery of nucleoside 5 -monophosphates. J. Med. Chem. 39:1981-1990.
Contributed by Suzanne Peyrottes and Christian P´erigaud Universit´e Montpellier Montpellier, France
Nucleoside Prodrugs and Delivery Strategies
15.3.13 Current Protocols in Nucleic Acid Chemistry
Supplement 29
STANDARD NOMENCLATURE, DATA, AND ABBREVIATIONS Selected Abbreviations Used in This Manual A adenine; adenosine Ac acetyl ACE acetoxyethoxy; acetoxyethyl ADA adenosine deaminase ADP adenosine 5 -diphosphate ADTT 3-amino-1,2,4-dithiazoline5-thione AEEA 2-aminoethoxy-2-ethoxy acetic acid AFM atomic force microscopy AMP adenosine 5 -monophosphate AMV-RT avian myoblastosis virus reverse transcriptase ANA arabinonucleic acid AP alkaline phosphatase 3-APA 3-aminopicolinic acid APS ammonium persulfate Ar aryl ASY average stepwise yield ATP adenosine 5 -triphosphate ATPase adenosine triphosphatase ATT 6-aza-2-thiothymine AU absorbance units AUFS absorbance units full scale B base (nucleobase) BAP bacterial alkaline phosphatase BCIP 5-bromo-4-chloro-3-indolyl phosphate BDT 3H-1,2-benzodithiol3-one-1,1-dioxide BHOC benzhydryloxycarbonyl BIT benzimidazolium triflate BMPM 1,1-bis(4-methoxyphenyl)-1pyrenylmethyl Bn benzyl bNA branched nucleic acid BOC tert-butyloxycarbonyl BOMP 2-(benzotriazol-1-yloxy)1,1-dimethyl-2-(pyrrolidin-1-yl)-1,3,2diazaphospholidinium hexafluorophosphate BSA bovine serum albumin; bis(trimethylsilyl)acetamide Bu butyl Bz benzoyl C cytidine; cytosine
APPENDIX 1
APPENDIX 1A
cAMP adenosine 3 ,5 -cyclicmonophosphate CD circular dichroism CDI 1,1 -carbonyldiimidazole cDNA complementary deoxyribonucleic acid CDP cytidine 5 -diphosphate CE capillary electrophoresis; cyanoethyl Cex p excess heat capacity CHA α-cyano-4-hydroxycinnamic acid Ci curie CID collision-induced dissociation CMCT 1-cyclohexyl-3-(2morpholinoethyl)carbodiimide metho-p-toluene sulfonate CMP cytidine 5 -monophosphate COSY correlation spectroscopy CPG controlled-pore glass CPI cyclopropapyrroloindole CPK Corey-Pauling-Koltun (molecular models) cpm counts per minute CPMB Current Protocols in Molecular Biology CSA D-10-camphorsulfonic acid CSO (1S)-(+)(10-camphorsulfonyl)oxaziridine CSP calf spleen phosphodiesterase CTAB cetyltrimethylammonium Ctmp 1-(2-chloro-4-methylphenyl)-4methoxypiperidin-4-yl CTP cytidine 5 -triphosphate dA deoxyadenosine Dabcyl 4-[4-(dimethylamino)phenyl]azobenzoic acid dADP deoxyadenosine diphosphate dAMP deoxyadenosine monophosphate DAP diaminopurine DAST (diethylamino)sulfur trifluoride dATP deoxyadenosine triphosphate DBMB 2-(dibromomethyl)benzoyl DBPNC 2,6-dibromo-4-benzoquinoneN-chloroimine DBU 1,8-diazabicyclo[3.4.0]undecene7-ene dC deoxycytosine Abbreviations and Useful Data
Current Protocols in Nucleic Acid Chemistry (2004) A.1A.1-A.1A.5 C 2004 by John Wiley & Sons, Inc. Copyright
A.1A.1 Supplement 18
Selected Abbreviations Used in This Manual
DCA dichloroacetic acid DCC 1,3-dicyclohexylcarbodiimide dCDP deoxycytosine diphosphate DCHA dicyclohexylamine DCI 4,5-dicyanoimidazole DCM dichloromethane dCMP deoxycytosine monophosphate dCTP deoxycytosine triphosphate DDQ 2,3-dichloro-5,6dicyano-1,4-benzoquinone DEAD diethyl azodicarboxylate DEC 1-(3-dimethylaminopropyl)-3ethylcarbodiimide DEMA diethoxymethyl acetate DEPC diethylpyrocarbonate dG deoxyguanosine DG distance geometry dGDP deoxyguanosine diphosphate dGMP deoxyguanosine monophosphate dGTP deoxyguanosine triphosphate DHB dihydroxybenzoic acid DHPC dihexanoylphosphatidylcholine dI deoxyinosine DIAD diisopropyl azodicarboxylate DIP 4,7-diphenyl-1,10-phenanthroline DIPEA or DIEA diisopropylethylamine DMAEDE 2 -O-2-[2-(N,Ndimethylamino)ethyloxy]ethyl DMAoE 2 -O-[2-(N,Ndimethylaminooxy)ethyl DMAP 4-dimethylaminopyridine DMBOC 3 ,5 - dimethoxybenzoinoxycarbonyl DMEM Dulbecco’s minimal essential medium (or Dulbecco’s modified Eagle’s medium) Dmf dimethylaminomethylene DMF dimethylformamide DMS dimethyl suberimidate; dimethyl sulfate DMSO dimethylsulfoxide DMPC dimyristoylphosphatidylcholine DMTr 4,4 -dimethoxytrityl DNA deoxyribonucleic acid DNBSB 2-(2,4-dinitrophenylsulfenyloxymethyl)benzoyl dNDP deoxynucleoside diphosphate dNMP deoxynucleoside monophosphate DNP 2,4-dinitrophenyl DNPEOC 2-(2,4-dinitrophenyl)ethoxycarbonyl dNTP deoxynucleoside triphosphate DODC 3,3 -diethyloxadicarbocyanine DPC diphenylcarbamoyl DPI 1,2-dihydro-3H-pyrrolo[3,2-e]indolecarboxylate dpm disintegrations per minute
DSC differential scanning calorimetry; disuccinimidyl carbonate dsDNA double-stranded DNA dT deoxythymidine dTDP deoxythymidine diphosphate DTE dithioerythritol dTMP deoxythymidine monophosphate DTMT 2-(isopropylthiomethoxymethyl)benzoyl DTr 4-decyloxytrityl (C10 Tr) DTT dithiothreitol dTTP deoxythymidine triphosphate dUTP deoxyuridine triphosphate DVB divinylbenzene DX double cross-over EC Enzyme Commission EDC 1-[3-(dimethylamino)propyl]-3ethylcarbodiimide EDITH 3-ethoxy-1,2,4-dithiazoline-5-one EDTA ethylenediaminetetraacetic acid EI electron impact (mass spectrometry) ENU ethylnitrosourea ESI electrospray ionization (mass spectrometry) Et ethyl FAB fast atom bombardment (mass spectrometry) FAM 5-carboxyfluorescein FDA (United States) Food and Drug Administration FFT fast Fourier transform FLB formamide loading buffer Fm fluorenylmethyl FMN flavin mononucleotide FMOC 9-fluorenylmethoxycarbonyl For formyl FPLC fast protein liquid chromatography Fpmp 1-(2-fluorophenyl)4-methoxypiperidin-4-yl FRET fluorescence resonance energy transfer FTICR Fourier-transform ion-cyclotron resonance (mass spectrometry) G guanine; guanosine GDP guanosine 5 -diphosphate GMP guanosine 5 -monophosphate GTP guanosine 5 -triphosphate HATU O-(7-azabenzotriazol-1-yl)-1,1,3,3tetramethyluronium hexafluorophosphate HBTU 2-(1H-benzotriazol-1-yl)-1,1,3,3tetramethyluronium hexafluorophosphate HEC hydroxyethylcellulose HEG hexa(ethylene glycol) HEPES N-(2-hydroxyethyl)piperazine-N (2-ethanesulfonic acid) HEPPSO N-(2-hydroxyethyl)piperazineN -(2-hydroxypropanesulfonic acid)
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HMDS hexamethyldisilazane HMFS N-[9-(hydroxymethyl)-2fluorenyl]succinamic acid HMQC heteronuclear multiple quantum coherence HNA hexitol nucleic acid HOAt 7-aza-1-hydroxybenzotriazole HOBt or HOBT 1-hydroxybenzotriazole 3-HPA 3-hydroxypicolinic acid HPLC high-performance liquid chromatography HPMC hydroxypropylmethylcellulose HQDA hydroquinone-O,O -diacetic acid HRMS high-resolution mass spectrometry HSDIS hydroxystyryldiisopropylsilyl HSDMS hydroxystyryldimethylsilyl HSQC heteronuclear single quantum coherence HZ hydrazine I inosine i-Bu isobutyl; isobutyryl ICS isocarbostyril IEC ion-exchange chromatography IMT imidazolium triflate i-Pr isopropyl IR infrared IRAA internal reference amino acid ITC isothermal titration calorimetry IUB International Union of Biochemistry IUPAC International Union of Pure and Applied Chemistry JOE 6-carboxy-4 ,5 -dichloro-2 ,7 dimethoxyfluorescein LCAA long-chain alkylamine LNA locked nucleic acid LSIMS liquid secondary-ion mass spectrometry L-TOF linear time-of-flight (mass spectrometry) MALDI matrix-assisted laser desorption/ionization (mass spectrometry) MAST [bis(2-methoxyethyl)amino]sulfur trifluoride MB minor groove binder MB-ODN minor groove binder– oligodeoxyribonucleotide conjugate MBHA p-methylbenzhydrylamine MC Monte Carlo mCPBA meta-chloroperoxybenzoic acid (also 3-chloroperoxybenzoic acid) MD molecular dynamics MDMP 1,5-dimethoxylcarbonyl-3methoxypentan-3-yl Me methyl
MeNPOC methylnitropiperonyloxycarbonyl Me-P methylphosphonate MES 2-(N-morpholino)ethanesulfonic acid MICS methylisocarbostyril MM molecular mechanical MMI methylenemethylimino MMTr 4-monomethoxytrityl MOPS 3-(N-morpholino)propane sulfonic acid MOTr 4-methoxy-4 -octyloxytrityl MOX 9-(p-anisyl)xanthen-9-yl MPC N-methylpyrrolecarboxamide MPLC medium-pressure liquid chromatography mRNA messenger ribonucleic acid MS mass spectrometry MSNT 1-(mesitylene-2-sulfonyl)-3-nitro1,2,4-triazole MTHP 4-methoxytetrahydropyran-4-yl MTMB 2-(methylthiomethoxymethyl) butyryl MTMT 2-(methylthiomethoxymethyl) benzoyl MTPI methyltriphenoxyphosphonium iodide MWCO molecular weight cutoff NAB nucleic acid builder (computer program) NAIM nucleotide analog interference mapping NAIS nucleotide analog interference suppression NBOM 2-nitrobenzyloxymethyl NBT 5-nitrobenzimidazolium triflate; nitroblue tetrazolium n-Bu n-butyl NDB Nucleic Acid Database NDP nucleoside diphosphate NDPK nucleoside diphosphate kinase NHS N-hydroxysuccinimide NMI N-methylimidazole NMM N-methylmesoporphyrin NMP nucleoside monophosphate; N-methylpyrrolidinone NMR nuclear magnetic resonance NOE nuclear Overhauser effect NOESY nuclear Overhauser effect spectroscopy N-P phosphoramidate NP1 nuclease P1 NPE 2-(4-nitrophenyl)ethyl NPEOC 2-(4-nitrophenyl)ethoxycarbonyl NPOC nitrophenyloxycarbonyl Abbreviations and Useful Data
A.1A.3 Current Protocols in Nucleic Acid Chemistry
Supplement 18
Selected Abbreviations Used in This Manual
NPPOC 2-(2-nitrophenyl) propoxycarbonyl NRC Nuclear Regulatory Commission nt nucleotide NTP nucleoside triphosphate ODN oligodeoxyribonucleotide PA picolinic acid Pac or pac phenoxyacetyl PACE phosphonoacetate PAGE polyacrylamide gel electrophoresis PAL peptide amide linker (resin) PAM 4-hydroxymethylphenylamidomethyl (resin) PAPOC p-phenylazophenyloxycarbonyl PAPS 3 -phosphoadenosine5 -phosphosulfate PASS product-anchored sequential synthesis PBS phosphate-buffered saline PCR polymerase chain reaction PDITC phenylene diisothiocyanate PEG polyethylene glycol PEG-MME polyethylene glycol– monomethyl ether PEO polyethylene oxide PEP phosphoenolpyruvate PFP pentafluorophenyl PFPC bis(pentafluorophenyl) carbonate Ph phenyl PICS 7-propynyloxycarbostyril PIM 7-propynyl-3-methylisocarbostyril PIPES piperazine-N,N bis(2-ethanesulfonic acid) PMSF phenylmethylsulfonyl fluoride PMT photomultiplier tube pn or PN phosphoramidate (linkage) PNA peptide nucleic acid PNK polynucleotide kinase pnODN oligodeoxyribonucleotide N3 →P5 phosphoramidate PNP purine nucleoside phosphorylase po or PO phosphodiester (linkage) Pr propyl PPA or PPG pyrazolo[3,4-d]pyrimidine analog of dA or dG ps or PS phosphorothioate (linkage) PS polystyrene PTFE polytetrafluoroethylene PTMT 2-(isopropylthiomethoxymethyl) benzoyl Pu purine Pv pivaloyl Px 9-phenylxanthen-9-yl (pixyl) PX paranemic cross-over Py pyrimidine
PyAOP 7-aza-benzotriazol-1-yloxytripyrrolidinophosphonium hexafluorophosphate PyBOP 1H-benzotriazol-1-yloxytripyrrolidinophosphonium hexafluorophosphate QM quantum mechanics or quantum mechanical QNAIM quantitative nucleotide analog interference mapping R, R . . . alkyl groups Rf retention factor (TLC) RFP radiofrequency plasma RFLP restriction fragment length polymorphism rMD restrained molecular dynamics RMSD root mean squared deviation RNA ribonucleic acid RNAi RNA interference RNAP RNA polymerase RNase ribonuclease RNP ribonucleoprotein ROX 5-(and 6-)carboxy-X-rhodamine RP-HPLC reversed-phase high-performance liquid chromatography RT-PCR reverse transcription polymerase chain reaction SA sinapinic acid SDS sodium dodecyl sulfate sec-Bu sec-butyl SELEX Systematic Evolution of Ligands by Exponential Enrichment SET 5-ethylthio-1H-tetrazole siRNA small interfering RNA SNP single nucleotide polymorphism S-PACE thiophosphonoacetate 3SR self-sustained sequence replication SSC sodium chloride/sodium citrate (buffer) SSCP single-stranded conformational polymorphism ssDNA single-stranded DNA SSPE saline sodium phosphate/EDTA (buffer) STM scanning tunneling microscopy STR short tandem repeats SVPD or SVP snake venom phosphodiesterase T thymidine; thymine T-jump temperature-jump (relaxation) tac tert-butylphenoxyacetyl (also t-PAC) TAE Tris/acetate/EDTA (buffer) TAMRA 5-(and 6-)carboxy-N,N,N ,N tetramethylrhodamine TBAF tetrabutylammonium fluoride
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TBDMS tert-butyldimethylsilyl TBE Tris/borate/EDTA (buffer) TBF 4-[17-tetrabenzo(a,c,g,i) fluorenylmethyl] TBS Tris-buffered saline t-Bu tert-butyl TBAF tetrabutylammonium fluoride TBAP tetrabutylammonium phosphate (also pyrophosphate or dihydrogenphosphate) TBE Tris/borate/EDTA (buffer) TCA trichloroacetic acid TCEP tris-(2-carboxyethyl)phosphine TDP thymidine 5 -diphosphate TE Tris/EDTA (buffer) TEA triethylamine (also Et3 N) TEAA triethylammonium acetate TEAB triethylammonium bicarbonate TEMED N,N,N ,N -tetramethylethylenediamine TES N-tris(hydroxymethyl)methyl-2aminoethanesulfonic acid TFA trifluoroacetic acid TFC triplex-forming circle TFMSA trifluoromethanesulfonic acid TFO triplex-forming oligonucleotide TFP tetrafluorophenyl THAP 2 ,4 ,6 -trihydroxyacetophenone THF tetrahydrofuran THP tetrahydropyran-2-yl TIPS triisopropylsilyl; tetraisopropyldisiloxane TLC thin-layer chromatography TMP 3,4,7,8-tetramethyl-1,10phenanthroline; thymidine 5 -monophosphate; trisodium trimetaphosphate
TMS trimethylsilyl TMSE trimethylsilylethyl TMS-OTf trimethylsilyl trifluoromethanesulfonate TMTr trimethoxytrityl TNT terminal deoxyribonucleotide transferase TOCSY total correlation spectroscopy TOF time-of-flight (mass spectrometry) Tol toluoyl TOM [(triisopropylsilyl)oxy]methyl t-PAC tert-butylphenoxyacetyl (also tac) TPP triphenylphosphine Tr diphenylmethyl (trityl) tRNA transfer ribonucleic acid TREAT-HF triethylammonium trihydrofluoride Tris tris(hydroxymethyl)aminomethane TROSY transverse relaxation–optimized spectroscopy Ts p-toluenesulfonyl (tosyl) TsOH toluene-4-sulfonic acid TTP thymidine 5 -triphosphate TX triple cross-over U uracil; uridine UDP uridine 5 -diphosphate UMP uridine 5 -monophosphate UTP uridine 5 -triphosphate UV ultraviolet VNTR variable number tandem repeats XAL xanthen alkonic acid (linker/resin) yDBR yeast debranching enzyme
Abbreviations and Useful Data
A.1.5 Current Protocols in Nucleic Acid Chemistry
Supplement 18
Characteristics of Nucleic Acids
APPENDIX 1B
Knowledge of the structural features of nucleic acids is important to an understanding of their biological function. This section provides information about the major nucleoside/nucleotide constituents and structures for A-, B-, and Z-DNA Some experimentally useful properties of the nucleoside/nucleotide building blocks are listed in Table A.1B.1. The chemical structures of the nucleosides can be seen in Figure A.1B.1, and aspects of nucleotide stereochemistry that are important to an understanding of base pairing and secondary structure can be found in Figures A.1B.2 and A.1B.3 and in Tables A.1B.2 and A.1B.3. Although Watson-Crick pairings play a critical role in defining nucleic acid secondary structures, a wide variety of alternative base pairings can be important in higher-order conformations; some of these are shown in Figure A.1B.4. Figure A.1B.5 depicts the three best characterized helix structures for DNA: A-, B-, and Z-DNA. A comprehensive collection of reviews on all aspects of nucleic acid structure has recently been published (Neidle, 1999).
Table A.1B.1
Nucleotide ATP ADP AMP Adenosinec dATPd dAMPd dA CTP CDP CMP Cytidine dCTPd dCMP dC GTP GDP GMP Guanosinec dGTPd dGMPd dG UTP UDP UMP Uridine TTPd TMPd Thymidined
Physical Characteristics of the Nucleotidesa
Mol. wt. (g/mol)
λmax (nm)
λmin (nm)
εmax (mM−1 cm−1)
A280/A260
507.2 427.2 347.2 267.2 491.2 331.2 251.2 483.2 403.2 323.2 243.2 467.2 307.2 227.2 523.2 443.2 363.2 283.2 507.2 347.2 267.2 484.2 404.2 324.2 244.2
259 259 259 260 259 259 260 271 271 271 271 272 271 271 253 253 252 253 252 253 254 262 262 262 262
227 227 227 227 226 226 225 249 249 249 250 — 249 250 223 224 224 223 222 222 223 230 230 230 230
15.4 15.4 15.4 14.9 15.4 15.2 15.2 9.0 9.1 9.1 9.1 9.1 9.3 9.0 13.7 13.7 13.7 13.6 13.7 13.7 13.0 10.0 10.0 10.0 10.1
0.15 0.16 0.16 0.14 0.15 0.15 0.15 0.97 0.98 0.98 0.93 0.98 0.99 0.97 0.66 0.66 0.66 0.67 0.66 0.67 0.68 0.38 0.39 0.39 0.35
0 0 11 — 0 11 — 0 0 15 — 0 18 — 0 0 6 — 0 6 — 0 0 20 —
6 26 52 — — 52 — 11 33 64 — — 65 — 5 17 40 — — 41 — 14 41 75 —
34 54 65 — 35 — — 41 64 75 — 43 — — 25 45 51 — 26 — — 49 71 80 —
482.2 322.2 242.2
267 267 267
— 234 235
9.6 9.6 9.7
0.73 0.73 0.70
0 24 —
— 74 —
52 — —
TLC mobilityb A B C
a Spectral data are assembled from Fasman (1975) at pH 7.0 except where footnoted otherwise. b TLC mobility is expressed as the percent distance a given spot migrates relative to the solvent front (Rf) in three different
TLC systems using 0.5-mm polyethylenimine cellulose plates: “A” is 0.25 M LiCl, “B” is 1.0 M LiCl, and “C” is 1.6 M LiCl. c Spectral measurements taken at pH 6.0. d Spectral data assembled from Dawson et al. (1987).
Standard Nomenclature, Data, and Abbreviations
Current Protocols in Nucleic Acid Chemistry (2000) A.1B.1-A.1B.14 Copyright © 2000 by John Wiley & Sons, Inc.
A.1B.1
Table A.1B.2 Average Torsion Angles for A-, B-, and Z-DNA Backbone (Dickerson, 1992)a
Torsion Angle α (O3′-P-O5′-C5′) β (P-O5′-C5′-C4′) γ (O5′-C5′-C4′-C3′) δ (C5′-C4′-C3′-O3′) ε (C4′-C3′-O3′-P) ζ (C3′-O3′-P-O5′) χ (O4′-C1′-N-C2(4))
A-DNA
B-DNA
Z-DNA
287 173 64 78 209 283 195
295 167 51 129 203 240 257
48, 223 179, 221 190, 55 100, 138 256, 299 291, 80 67, 201
aRotations are given in degrees (°). The parameters are illustrated in Figure
A.1B.2.
Table A.1B.3
Mean Value of Helix Parameters for A-, B-, and Z-DNAa
Parameter X-displacement Y-displacement Inclination Tip Buckle Propeller Opening Shift Slide Rise Tilt Roll Twist
A-DNA B-DNA B-DNA mean (range) −4.1 — 12.0 11.0 −2.4 −8.3 — — −1.6 2.9 — 6.3 31.1
0.8 0.1 2.4 0 −0.2 −11.1 — — 0.4 3.4 0 0.6 36.1
0.5 (−0.6 to 1.5) — −1.2 (−10.9 to 11.0) 0.1 (−11.3 to 11.3) 0.9 (−10.6 to 16.0) −11.0 (−24 to 5.2) 1.5 (−8.4 to 8.1) 0 (−1.1 to 1.1) −0.1 (−1.1 to 1.1) 3.4 (3.2 to 4.0) −0.2 (−7.6 to 7.6) 1.8 (−12.4 to 16.0) 36.2 (21.8 to 50.6)
Z-DNA 3.0 (C), 3.0 (G) −2.3 (C), 2.3 (G) −6.2 (C), −6.2 (G) 2.9 (C), −2.9 (G) −6.2 (C), 6.2 (G) −1.3 (C), −1.3 (G) — — 5.4 (C-G), −1.1 (G-C) 3.9 (C-G), 3.5 (G-C) 0, 0 −5.8 (C-G), 5.8 (G-C) −9.4 (C-G), −50.6 (G-C)
aTranslations are given in angstroms (Å) and rotations in degrees (°). The parameters are illustrated in Figure A.1B.3. The values of the parameters for A-DNA, B-DNA, and Z-DNA are from Dickerson, 1992, Heinemann et al., 1994, and Hartman and Lavery, 1996.
Characteristics of Nucleic Acids
A.1B.2 Current Protocols in Nucleic Acid Chemistry
O
O
4
3
pKa = 9.4
NH
5
CH3
NH2 NH
pKa = 10.0
pKa = 4.4
N
2
6
N1 R
O
N R
uridine, R = ribosyl
pKa < 1
O
thymidine, R = deoxyribosyl
7
N
9
N
1
4
2
3N
O N
pKa = 3.8
N
8
cytidine, R = ribosyl deoxycytidine, R = deoxyribosyl
pKa = 2.4
NH2 5 6
O
N R
N
R
NH N
pKa = 9.4
NH2
R
adenosine, R = ribosyl deoxyadenosine, R = deoxyribosyl
guanosine, R = ribosyl deoxyguanosine, R = deoxyribosyl
5′
HO
O
4′
1′ 2′
3′
OH
ribosyl
OH
NH2
pKa = 12.3 -12.5 pKa = 6
O O O γ α β P P P O O O O O O O
N N O
N N
R = HO
pKa < 1
O OH
deoxyribosyl
OH
OH
adenosine triphosphate (ATP)
pKa = 14
Figure A.1B.1 Line drawings of the nucleosides. The chemical structure that predominates at neutral pH is shown. Drawings of the nucleotide bases and their associated sugars, either ribose or deoxyribose, are shown separately. In the representations of ribose (by itself and as a nucleoside triphosphate, ATP) and deoxyribose, the bold lines indicate that this portion of the sugar is coming out of the page toward the reader. In this view, the base is found above the plane of the sugar, while the 3′-hydroxyl group is found below the plane of the sugar. The pKa values for all groups are shown; pKa values above 7 imply proton association to the pictured structure, while pKa values below 7 imply proton dissociation from the pictured structure. The tautomeric form of a given base may
change at different pH values. The pKa values given are for nucleotide monophosphates and were taken from Dawson et al. (1987); a fuller discussion of the chemical basis for these values can be found in a review by T’so (1974). The small numbers adjacent to adenosine, uridine, and ribose indicate the nomenclature of the purines, pyrimidines, and sugars, respectively. Groups appended to a ring have the same numbering as the position to which they are linked; thus, the “O6” moiety of guanosine is the carbonyl oxygen bonded to C6 in the ring. Similarly, “O3′” on ribose or deoxyribose indicates the oxygen of the hydroxyl group bonded to C3′ in the ring. The α, β, and γ phosphates in a nucleoside triphosphate (adenosine triphosphate, ATP) are also indicated.
A.1B.3 Current Protocols in Nucleic Acid Chemistry
Supplement 4
Figure A.1B.2 Nucleotide stereochemistry. (A) Various torsion angles for DNA backbone. Average values for A-, B-, and Z-DNA are given in Table A.1B.2. (B) The torsion angle (θ) is defined as the angle between the two planes defined by the three bonds A-B-C and B-C-D. When bonds A-B and C-D are eclipsed, the torsion angle is 0. The direction of rotation shown in the figure corresponds to a positive value of θ. (C) Depending on the rotation about the bond between C1′ of the sugar and either N1 (for pyrimidines) or N9 (for purines), a nucleotide can be described as either “anti” or “syn.” Because of steric constraints, nucleotides are generally found in the “anti” configuration, with their Watson-Crick hydrogen bond donor-acceptors swung outward away from the plane of the sugar ring. However, guanosine is sometimes found in a “syn” configuration, both in polynucleotides
and in solution. In this form, the bulk of the purine ring is positioned directly over the plane of the sugar. (D) The sugar ring can also adopt different stereochemistries. These are labeled according to which group is bent out of the plane of the ring, and in which direction. If a portion of the ring is bent “upward” toward the base, this is known as “endo,” while if it is bent “downward” away from the base, this is known as “exo.” The majority of nucleotides cluster in two domains centered at C2′-endo and C3′-endo (Saenger, 1984). In the figure, plain lines represent bonds that are within the plane of the sugar, while bold lines indicate that the bond is bent out of the plane. Hence, “C3′ endo-C2′ exo” describes a furanose ring in which the 2′ and 3′ carbons have been twisted in opposite directions and the bond connecting them crosses the plane of the ring.
A.1B.4 Supplement 4
Current Protocols in Nucleic Acid Chemistry
Figure A.1B.3 Definitions of helix parameters for duplex DNA (Dickerson, 1989). The transformations shown are all in the positive direction.
Standard Nomenclature, Data, and Abbreviations
A.1B.5 Current Protocols in Nucleic Acid Chemistry
Figure A.1B.3
continued
A.1B.6 Current Protocols in Nucleic Acid Chemistry
H N H
N
N
O N
N
N
R
H N
R
N
R
A:T
N H
O
O H N
O
N
N N
H N
N
N R
G:C
N
N
R
N H H
R N
H
N
N
Watson-Crick pairings
O N
N H N
N O
H H N
O
R
H
O
N H
R
N N H
A:C
G:U
H
reverse wobble pairing
wobble pairing
H O
H
N
O
N
R
N N H R
N
N N
N
N
N H
N R
R
N
O
H
O T·A Hoogsteen (parallel chains)
Figure A.1B.4 Base pairing schemas. The chemical structures of the nucleotide bases determine the formation of secondary and tertiary structures in nucleic acids. A wide variety of hydrogen bonding schemas (indicated by dashed lines) are possible between different bases. Watson-Crick pairings are perhaps the most widely known and are the basis of the double-helical structure of complementary, anti-parallel DNA strands. Other base pairs can also be accommodated within the double helix, such as “wobble pairings,” in which the bases are slightly off-center with respect to each other. By using the N7 hydrogen bond acceptor of the
T·A
N
N
H
reverse Hoogsteen (antiparallel chains) purine bases adenosine and guanosine, an even wider variety of structures becomes possible. Bonds involving N7 of the purine bases allow tertiary structural interactions to occur in nucleic acids, including triple base pairs (Radhakrishnan and Patel, 1994) and “G quartet” (Sen and Gilbert, 1988). A compilation of possible base pairs has been published (Burkard et al., 1999). This information can also be viewed on the World Wide Web at http://www.imb-jena.de/IMAGE_BPDIR.html.
Standard Nomenclature, Data, and Abbreviations
A.1B.7 Current Protocols in Nucleic Acid Chemistry
R
R
N
N
N N
H N H
N
N
N
N
O
H N H
H
N
N
H H
N
R
N
N
N
A·A:T
O
H
H
N
N N
N
R
O H
G·G:C
H N H
R N
O N
O
N H
R
R
R N
H
N
N
N
N
O
O
N
N
H
H
O N
H
H H
N N
N
O
R N
O
N
H
H
O
N
H
N
N
R
R T·A:T
N N
N
H N H
N
R
O H
C+·G:C
R
H N
N
N N
N
H N
N
O
O
N N
H H N
H
H
N H H
N N
O
O H
R
N
R
H
N
H
N H
N
N N
N R
G-quartet Figure A.1B.4
continued
A.1B.8 Current Protocols in Nucleic Acid Chemistry
Figure A.1B.5 Nucleic acid secondary structures. The structural consequence of the ability of nucleotides to form Watson-Crick base pairs is nucleic acid double helices. In th is figure, the self-complementar y 12-mer CGCGAATTCGCG is shown as both A- and B-form helices. Two representations of the A helix have been shown in order to emphasize the depth of the major groove. The arrows and brackets in these figures are not drawn to scale.
While both of these helices are right-handed (in terms of anthropomorphic referents, if you were to point your thumb along a strand in a 5′ to 3′ manner, the twist of the helix would be the same as the curl of your right hand), their structural details are very different: B DNA has roughly 10 bases per full turn, while A DNA and A RNA have 11 to 12; the major groove of B-form helices is wide and the minor
A.1B.9 Current Protocols in Nucleic Acid Chemistry
groove is narrow, while for A-form helices this is reversed; in B-forms the base pairs are located close to the helix axis (as can be seen in end-on views), while in A-forms the base pairs are pushed out away from the long helical axis, leaving a “hole” in the middle of the polynucleotide coil (if one imagines DNA as a flat ribbon, then B DNA is twisted from its ends, while A DNA is coiled on itself).
in B DNA) and a 3′ endo deoxyribose (found in A DNA) are indicated. While there are a variety of other helical forms, the most striking is that found in Z DNA. The Z DNA coil is left- rather than right-handed and contains G:C base pairs where the G is in the “syn” conformation (shown in the inset).
Different helical forms are largely due to differences in sugar stereochemistry. Examples of a 2′ endo deoxyribose (found
A.1B.10 Current Protocols in Nucleic Acid Chemistry
The uneven progression, or zigzag, of Z DNA can be more easily seen when the polynucleotide backbone is shown in isolation; the inset shows the connectivity between phosphates by 5′ to 3′ vector arrows. Because of its odd shape,
base pairs actually protrude from what would be a cavity in A or B DNA; thus, Z DNA has a minor but no major groove. This diagram is based on the original structure of alternating C:G/G:C base pairs (Wang et al., 1979).
A.1B.11 Current Protocols in Nucleic Acid Chemistry
A.1B.12 Current Protocols in Nucleic Acid Chemistry
A.1B.13 Current Protocols in Nucleic Acid Chemistry
LITERATURE CITED Burkard, M.E., Turner, D.H., Tinoco, I. Jr. 1999. Structure of base pairs involving at least two hydrogen bonds. In RNA World, 2nd ed. (R.F. Gesteland, T.R. Cech, and J.F. Atkins, eds.) pp. 675-680, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Dawson, M.C., Elliott, D.C., Elliott, W.H., and Jones, K.M. (eds.). 1987. Data for Biochemical Research, 3rd ed. Clarendon Press, Oxford. Dickerson, R.E. 1989. Definitions and nomenclature of nucleic acid structure components. Nucl. Acids Res. 17:1797-1803. Dickerson, R.E. 1992. DNA structure from A to Z. Methods Enzymol. 211:67-111. Fasman, G. (ed.). 1975. Handbook of Biochemistry and Molecular Biology, Vol. 1: Nucleic Acids, 3rd ed. CRC Press, Boca Raton, Fla. Hartman, B. and Lavery, R., 1996. DNA structural forms. Q. Rev. Biophys. 29:309-368. Heinemann, U., Ailings, C., and Hahn, M., 1994. Crystallographic studies of DNA helix structure. Biophys. Chem. 50:157-176. Neidle, S. 1999. Oxford Handbook of Nucleic Acid Structure. Oxford University Press, Oxford. Radhakrishnan, I. and Patel, D.J. 1994. DNA triplexes: solution structures, hydration sites, energetics, interactions, and function. Biochemistry. 33:11405-11416. Saenger, W. 1984. Principles of Nucleic Acid Structure. Springer-Verlag, New York. Sen, D. and Gilbert, W. 1988. Formation of parallel four-stranded complexes by guanine-rich motifs in DNA and its implications for meiosis. Nature 334:364-366. T’so, P.O.P. 1974. Bases, nucleosides, and nucleotides. In Basic Principles in Nucleic Acid Chemistry, Vol. 1 (P.O.P. T’so, ed.) pp. 453-584. Academic Press, San Diego. Wang, A.H., Quigley, G.J., Kolpak, F.J., Crawford, J.L., van Boom, J.H., van der Marel, G., and Rich, A. 1979. Molecular structure of a left-handed double helical DNA fragment at atomic resolution. Nature 282:680-686.
Adapted by Donald E. Bergstrom, Purdue University, from Current Protocols in Molecular Biology
Characteristics of Nucleic Acids
A.1B.14 Current Protocols in Nucleic Acid Chemistry
IUPAC-IUB Joint Commission on Biochemical Nomenclature Abbreviations and Symbols for the Description of Conformations of Polynucleotide Chains The following rules are as close as possible to the originally published version [1] and are reproduced here with permission from the European Journal of Biochemistry. A World Wide Web version prepared by G.P. Moss can be found at http://www.chem.qmw.ac.uk/iupac/ misc/pnuc1.html. Several conventions and notations for polynucleotide conformation have been used by various authors [2-10]. To overcome this confusion, the Joint Commission on Biochemical Nomenclature (JCBN) appointed a panel of experts to review the problem and make recommendations. Their proposals, together with suggestions from the members of JCBN and other scientists, are presented here as recommendations that have been approved by the International Union of Pure and Applied Chemists (IUPAC) and the International Union of Biochemistry (IUB). The nomenclature proposed here is consistent with that recommended for polypeptide conformation [11] as well as with recommendations for polysaccharide conformation [13] and stereochemistry of synthetic polymers [14]. The recommendations on polypeptide conformation [11] also cover general problems of specifying the conformation of biopolymers. Nomenclature of nucleic acids and symbols for their constituents follow published recommendations [15].
1. GENERAL PRINCIPLES OF NOTATION 1.1. Chain Direction The atoms of the main chain are denoted in Figure A.1C.1. The direction of progress of a polynucleotide chain is from the 5′ end to the 3′ end of the sugar residue. Notes (a) The definition of chain direction is in accord with the definition of the nucleotide unit (see section 1.2). (b) The definition of chain direction with respect to the sugar carbon atoms of the nucleotide unit is consistent with the alternative description of polynucleotide sequences as pro-
APPENDIX 1C
Figure A.1C.1 Designation of chain direction and main chain atoms of i th unit in a polynucleotide chain.
gressing from the 3′ end of one unit to the 5′ end of the next through the phosphate group, i.e., in the chain sequence L-(3′→5′)-M(3′→5′)-N, etc. (written as LpMpN or L-M-N for a known sequence, or L, M, N for an unknown sequence).
1.2. Definition of a Nucleotide Unit A nucleotide unit is the repeating unit of a polynucleotide chain; it comprises three distinct parts: the D-ribose or 2-deoxy-D-ribose (2-deoxy-D-erythro-pentose) sugar ring, the phosphate group, and the purine or pyrimidine base. The sugar ring and the phosphate group form the backbone of the polynucleotide chain; the base ring linked to the sugar residue constitutes the side chain as shown in Figure A.1C.1. A nucleotide unit is defined by the sequence of atoms from the phosphorus atom at the 5′ end to the oxygen atom at the 3′ end of the pentose sugar; it includes all atoms of the sugar and base rings. Specific units (i, j,..., or 3, 4, 5, etc.) are designated by the letter or number in brackets. The units are numbered sequentially in the chain direction, starting at the first nucleotide residue, irrespective of the presence or absence of a phosphate group at the 5′-terminal unit. The same numbering, A(1), pU(2), pU(3), etc., would apply to the sequence ApUpUp- and pApUpUp-.
1.3. Atom Numbering The atom numbering of the constituents of the nucleotide unit is shown in Figure A.1C.1,
Current Protocols in Nucleic Acid Chemistry (2000) A.1C.1-A.1C.9 Copyright © 2000 by John Wiley & Sons, Inc.
Standard Nomenclature, Data, and Abbreviations
A.1C.1
Figure A.1C.2 The atom numbering for the bases of common nucleosides and nucleotides. Hydrogen atoms carry the same numbers as the heavy atoms to which they are attached. The name in parenthesis applies when the “d” in parenthesis in the formula is present.
A.1C.2, and A.1C.3. The numbering scheme for the bases shown in Figure A.1C.2 is the same as that recommended by IUPAC (Rule B-2.11 on page 58 in [16] and on page 5567 in [17]). The atoms belonging to the sugar moiety are distinguished from those of the base by the superscript prime mark on the atom number. Atoms are specified by the appropriate number after the symbol: e.g., C2, N3 (for base) and C1′, C5′, O5′ (for sugar). Atoms of a specific unit (i, j,..., or 3, 4, 5) may be designated by the letter or number of the unit in brackets: e.g., O3′(i), P(i + 1) and N1(3), C2′(4). Hydrogen atoms carry the same number as the heavy atoms to which they are attached: e.g., base ring H6 (pyrimidine) and H8 (purine) as shown in Figure A.1C.2; sugar ring H1′, H2′
The Description of Conformations of Polynucleotide Chains
Figure A.1C.3 Designation of sugar ring atoms and hydrogen atoms. (A) In β-D-nucleosides and nucleotides; (B) in their 2′-deoxy derivatives.
etc, as shown in Figure A.1C.3. Where there is more than one hydrogen atom (such as at C5′ of the sugar ring), the atoms are designated numerically (e.g., H5′1 and H5′2); H5′1 and H5′2 correspond to the pro-S and pro-R positions [18], respectively (Fig. A.1C.3). The atom-numbering scheme for the 2′-deoxyribonucleotide chain is the same as that for the ribonucleotide chain shown in Figure A.1C.1. The numbering for the sugar ring atoms of both D-ribose and 2-deoxy-D-ribose rings is shown in Figure A.1C.3. (Note the absence of a prime in “2-deoxy-D-ribose” in the previous sentence; C2′ of a nucleotide is C2 of its ribose residue.) The two hydrogen atoms attached to the C2′ atom of a nucleoside are denoted by H2′1 and H2′2, corresponding to the pro-S and pro-R positions respectively (Fig. A.1C.3). The hydrogen atoms of hydroxyl groups are specified as in O5′H, O3′H, and O2′H, where appropriate, whereas the hydroxyl groups are specified as OH5′, etc. Notes (a) Designation of the sugar-ring oxygen atom by O4′ conforms with chemical nomenclature; it has been widely but inaccurately denoted by O1′ in the past. (b) Detailed atom numbering for the modified nucleotides is not considered here. (c) The numerical designation of C5′ and C2′ methylene hydrogen atoms supersedes that introduced by Davies [10].
1.4. Bonds, Bond Lengths, and Interatomic Distances Covalent bonds are denoted by a hyphen between atoms: e.g., O5′-C5′, C5′-H5′1, and C2-N3. Atoms in specified nucleotide units are indicated by putting the number of the unit in parentheses: e.g., O5′(i) − C5′(i), O3′(i) − P(i + 1). Bond lengths are denoted by b(O5′, C5′) or b[O3′(i), P(i + 1). Use of the symbol l for bond length is avoided because it can be confused with the numeral 1 and because l is used for vibration amplitude in electron diffraction (section 1.4 of [11]). Hydrogen bonds are denoted by a dotted line, with the donor atom being written first, if it can be specified: e.g., intramolecular O5′...N3 hydrogen bonding in some purine derivatives or O5′(i)...N3(j) for intermolecular hydrogen bonding. The position of the hydrogen may also be indicated, as in O5′H...N3. Hydrogen-bonded base pairs are considered separately (see section 4.1). Distances
A.1C.2 Current Protocols in Nucleic Acid Chemistry
between nonbonded atoms are denoted by a dot: e.g., O5′(i).O3′(j ).
1.5. Bond Angles The bond angle included between three atoms A-B-C is written τ(A, B, C), which may be abbreviated to τ(B) if there is no ambiguity.
1.6. Torsion Angles [18] If a system of four atoms A-B-C-D is projected onto a plane normal to B-C, the angle between the projection of A-B and the projection of C-D is described as the torsion angle about bond B-C; this angle may also be described as the angle between the plane containing atoms A, B, and C, and the plane containing atoms B, C, and D. The torsion angle is written in full as θ(A, B, C, D), which may be abbreviated, if there is no ambiguity, to θ(B, C). In the statement of this rule, the angle θ is used as a general angle rather than as referring to any particular bond (see Rule 2 for designation of main-chain torsion angles). The zero-degree torsion angle (θ = 0°) is given by the conformation in which the projections of A-B and C-D coincide (this is also known as the eclipsed or cis conformation). When the sequence of atoms A-B-C-D is viewed along the central bond B-C, a torsion angle is considered positive when the bond to the front must be rotated clockwise in order that it may eclipse the bond to the rear as shown in Figure A.1C.4A. When the bond to the front must be rotated counterclockwise in order to eclipse the bond at the rear, the angle is considered negative, as shown in Figure A.1C.4B. Angles are usually measured from 0° to 360°, but they may be expressed as −180° to +180° when special relationships between conformers need to be emphasized. Illustrations of the definition of torsion angles are shown for positive and negative values of θ in Figure A.1C.4A and Figure A.1C.4B respectively. It should be noted that a clockwise turn of the bond containing the front atom about the central bond gives a positive value of θ from whichever end the system A-B-C-D is viewed in Figure A.1C.4A; similar considerations apply to the conformation with a negative torsion angle (Fig. A.1C.4B).
1.7. Conformational Regions If the precise torsion angle for a conformation is not known, it may be convenient to specify it roughly by naming a conformational region—i.e., a range in which the torsion angle lies. For this the Klyne-Prelog nomenclature
Figure A.1C.4 Newman projections illustrating positive and negative torsion angles. (A) A clockwise turn of the bond containing the front atom about the central bond is needed for it to eclipse the bond to the back regardless of the end from which the system is viewed; hence the value of θ is positive (+θ). (B) A counterclockwise turn of the bond containing the front atom is needed for it to eclipse the bond to the back atom regardless of the end from which the system is viewed; hence the value of θ is negative (−θ).
[18, 19], accepted in organic chemistry, is recommended. The relationship between the terms used, ±synperiplanar (±sp), ±synclinal (±sc), ± anticlinal (±ac), and ±antiperiplanar (±ap), and the magnitudes of the torsion angles are shown in Figure A.1C.5. The range 0° ± 90° is denoted as syn and the range 180° ± 90° is denoted as anti. Note In order that conformations described by the torsion angle defined in rule 1.6 be consistent in sign and magnitude with the conformational regions (±syn, ±anti) shown in Figure A.1C.5, it is necessary, when looking down the B→C (or C→B) bond, that the front bond A-B (or D-C) should define the zero (0°) position, and that the back bond should define the conformational region.
Examples (a) In Figure A.1C.3 the O5′-C5′ bond makes an angle of +60° to the C4′-C3′ bond and 300° (−60°) to C4′-O4′. With the O5′-C5′ bond defining the zero position, these conformations correspond to the +sc and -sc regions, respectively. (b) For the sequence of atoms A-B-C-D as shown in Figure A.1C.4A, the same conformation with the torsion angle θ being positive (+θ) is found when looking along either the B-C bond (A to the front) or the C-B bond (D to the front); this conformation is described as +sc.
Standard Nomenclature, Data, and Abbreviations
A.1C.3 Current Protocols in Nucleic Acid Chemistry
Figure A.1C.6 Section of a polynucleotide backbone showing the atom numbering and the notation for torsion angles. (A) Conventional representation; (B) absolute stereochemistry. Figure A.1C.5 Relationship between the synanti terminology for describing conformational regions [18,19], and the magnitude of the torsion angle 0° to 360° (or 0° ± 180°) with the front bond specifying the zero position. sp, synperiplanar; sc, synclinal, ap, antiperiplanar; ac, anticlinal. Other descriptions of particular torsion angles are also given for comparison: 0°, cis (c); 60° +gauche (g+); 180°, trans (t); 300°, −gauche (g−).
Similarly, the conformation designated by a negative value of θ (−θ), shown in Figure A.1C.4B, corresponds to the -sc region. (c) See section 2.3, notes (a) and (c), for the example in Figure A.1C.11.
2. THE NUCLEOTIDE UNIT The notations used to designate the various torsion angles in the nucleotide unit are indicated in three sections: sugar–phosphate backbone chain, sugar ring, and sugar–base side chain.
2.1. Sugar-Phosphate Backbone Chain (Main Chain)
The Description of Conformations of Polynucleotide Chains
Note The recommended α − ζ notation differs from the φ, ψ, ω notation [5, 8] adopted by many workers and from an earlier α − ζ notation [9]. A substantial majority of the subcommittee favored the α − ζ notation because it is convenient to remember for a backbone repeat of six bonds. The recommended α − ζ notation is the second of the systems proposed by Seeman et al. [9] and was chosen because it starts at the phosphorus atom which is the first atom of the nucleotide unit, has the highest atomic number, and is the only atom of its kind in the backbone.
2.2. Sugar Ring 2.2.1. Endocyclic torsion angles The sugar ring occupies a pivotal position in the nucleotide unit because it is part of both the backbone and the side chain. In order to provide a complete description of the ring conformation, it is necessary to specify the endocyclic torsion angles for the ring as well as the bond lengths and bond angles. The five endocyclic torsion angles for the bonds O4′-C1′,
The backbone of a polynucleotide chain consists of a repeating unit of six single bonds as shown in Figure A.1C.1: i.e., P-O5′, O5′-C5′, C5′-C4′, C4′-C3′, C3′-O3′, and O3′-P. The torsion angles about these bonds are denoted, respectively, by the symbols α, β, γ, δ, ε, and ζ. The symbols α(i) − ζ(i) are used to denote torsion angles of bonds within the ith nucleotide unit as shown in Figure A.1C.6 and Figure A.1C.7. The sequence of main-chain atoms used to define each backbone torsion angle is shown in Figure A.1C.7. Figure A.1C.7 Torsion angles for backbone conformations of the i th nucleotide in polynucleotide chains.
A.1C.4 Current Protocols in Nucleic Acid Chemistry
Figure A.1C.8 Torsion angles in sugar rings of β-D-nucleosides and nucleotides.
C1′-C2′, C2′-C3′, C3′-C4′, and C4′-O4′ are denoted by the symbols, ν0, νl, ν2, ν3, and ν4, respectively. The sequence of atoms used to define each backbone torsion angle is shown in Figure A.1C.8: e.g., ν0 refers to the torsion angle of the sequence of atoms C4′-O4′-C1′ -C2′, etc. Notes (a) The backbone torsion angle δ and the endocyclic torsion angle ν3 both refer to rotation about the same bond, C4′-C3′. Both angles are needed for complete description of the main-chain and sugar-ring conformations in some studies. (b) The notation τ0 − τ4 previously used [5, 20-22] to represent torsion angles about the bonds in the sugar ring is superseded by the present notation (ν), which is consistent with polysaccharide nomenclature [13]. The symbol τ is now used to denote a bond angle, which is consistent with polypeptide nomenclature [11]. 2.2.2. Puckered forms Since the sugar ring is generally nonplanar, its conformation may need designation. If four of its atoms lie in a plane, this is chosen as a reference plane, and the conformation is described as envelope (E); if they do not, the reference plane is that of the three atoms that are closest to the five-atom, least-squares plane, and the conformation is described as twist (T) [23, 24]. Atoms that lie on the side of the reference plane from which the numbering of the ring appears clockwise are written as superscripts and precede the letter (E or T); those on the other side are written as subscripts and follow the letter (Fig. A.1C.9). These definitions [24] mean that atoms on the same side of the plane as C5 in D-ribofuranose derivatives are written as preceding superscripts. Notes (a) The present E and T notations for puckered forms of the sugar ring conform to those recommended for the conformational nomenclature of five- and six-membered rings of monosaccharides and their derivatives [24].
Figure A.1C.9 Diagrammatic representation of sugar-ring conformations of β-D-nucleosides and their relation to the pseudorotational Ntype and S-type conformers (Section 2.2.3.). The purine or pyrimidine base is represented by B. (Figure adapted from that of Saenger [26].)
(b) The E/T notation has superseded the endo/exo description [25], in which atoms now designated by superscripts were called endo, and those now designated by subscripts were called exo. Figure A.1C.10 shows both systems of designation. Examples: C3′-endo/C2′-exo has become 3T2 C3′-endo has become 3E. (c) Symmetrical twist conformations, in which both atoms exhibit equal displacements with respect to the five-atom plane, are denoted by placing the superscript and subscript on the same side of the letter T: e.g., 23T, 43T, etc. 2.2.3. Pseudorotational analysis The sugar ring conformation has also been described by Altona and Sundaralingam [27] using the concept of pseudorotation, which has been found advantageous in describing the conformational dynamics of the sugar ring [28]. Each conformation of the furanose ring can be unequivocally described by two pseudorotational parameters: the phase angle of pseudorotation, P, and the degree of pucker, ψm. A standard conformation (P = 0°) is defined with a maximally positive C1′-C2′-C3′ -C4′ torsion angle [(.e., the symmetrical 23T form), and P has value 0° to 360°. Conformations in the upper, or northern, half of the circle (P = 0° ± 90°) are denoted N and those in the lower, or
Standard Nomenclature, Data, and Abbreviations
A.1C.5 Current Protocols in Nucleic Acid Chemistry
Figure A.1C.10 The pseudorotational pathway of the D-aldofuranose ring, showing the relation between phase angle of pseudorotation P (0° to −360°), the envelope (E) and twist (T) notations, and the endo and exo notations. N-type conformations correspond to the northern half (P = 0 ± 90°) and S-type correspond to the southern half of the pseudorotational cycle. The symbols “r” and “d” represent the usual range of P values for N and S conformations of ribo- (r) and 2′-deoxyribo- (d) furanose rings of β-D-nucleosides and nucleotides. (Diagram adapted from the work of Altona and Sundaralingam [27].)
southern, half of the circle (P = 180° ± 90°) are denoted S. The relationship between P and the endo/exo and T/E notations is illustrated in Figure A.1C.10. It may be seen that the symmetrical twist (T) conformations arise at even multiples of 18° of P and the symmetrical envelope (E) conformations arise at odd multiples of 18° of P. Note The present designation of the degree of pucker (ψm) differs from the original notation (τm) of Altona and Sundaralingam [27] in order to avoid confusion with the notation for bond angles.
2.3. N-Glycosidic Bond
The Description of Conformations of Polynucleotide Chains
The torsion angle about the N-glycosidic bond (N-C1′) that links the base to the sugar is denoted by the symbol χ; this is the same as the notation used to denote side-chain torsion angles in polypeptides [11]. χ(i) denotes the torsion angle in the ith nucleotide unit. The sequence of atoms chosen to define this angle is O4′-C1′-N9-C4 for purine and O4′C1′-N1-C2 for pyrimidine derivatives. Thus when χ = 0° the O4′-C1′ bond is eclipsed with the N9-C4 bond for purine and the N1-C2 bond
F igure A.1C.11 Diagrammatic representation of the N-glycosidic bond torsion angle χ and the syn and anti regions for purine and pyrimidine derivatives. The purine derivative (left) is viewed down the N9-C1′ bond and is shown in the +sc conformation. The pyrimidine derivative (right) is viewed down the N1-C1′ bond and is shown in the −ac conformation. The sugar ring is shown as a regular pentagon.
for pyrimidine derivatives. The definitions of torsion angles (section 1.6) of the N-glycosidic bond are illustrated, looking along the bond, in Figure A.1C.11. Notes (a) The choice of bond sequence to define χ is based on accepted chemical nomenclature [14] and, at the same time, the use of the terms syn and anti to describe different conformational regions of χ for purine and pyrimidine derivatives is now consistent with accepted chemical nomenclature (Rule 1.7): i.e., syn, χ = 0° ± 90° anti, χ = 180° ± 90°. Examples of syn and anti conformations are shown in Figure A.1C.11. In the new convention most conformations formerly described as syn and anti remain syn and anti respectively, except the high-anti region which may be described as -synclinal (-sc). Rule 1.7. (b) Many of the conventions for defining the torsion angle of a bond in nucleic acids [2-8, 29, 30] have been based on the sugar ring O4′ atom [5, 6, 26] or C2′ atom [4, 30] in conjunc-
A.1C.6 Current Protocols in Nucleic Acid Chemistry
tion with the base ring C8(Pur)/C6(Pyr) atoms [5, 6, 30] or base ring C4(Pur)/C2(Pyr) atoms [4, 29]. Approximate relationships between the definitions of torsion angles have been summarized by Sundaralingam [31], and these aid the comparison of conformations described in the older literature. A substantial portion of the literature has used the nomenclature based on sugar ring O4′-C1′ and base ring N9C8(purine) and N1-C6(pyrimidine). χold is related to the present definition χnew by the relation, χnew ∼ χold ± 180°. (c) Following the definitions of conformational regions in Rule 1.7 and Figure A.1C.5, the anti conformation of the pyrimidine example in Figure A.1C.11 corresponds to the -anticlinal (-ac) conformation and the syn conformation of the purine derivative corresponds to the +synclinal (+sc) conformation.
2.4. Orientation of Side Groups For the precise definition of the orientation of any pendant groups, specification of the torsion angle about the exocyclic bond is necessary. The exocyclic torsion angle may be denoted by the symbol η with a locant to indicate the atom to which it refers. Examples: Ribose rings. The symbol η2′ may be used to denote the torsion angle about the C2′-O2′ bond for the sequence of atoms Cl′-C2′-O2′ -X, where X = H, CH3, PO2− 3 , etc. If no confusion is possible, the symbol η (without any additional index) may be used for the C2′-O2′ bond. Base. Torsion angles for bonds in base rings such as C6-N6 in adenine, C2-N2 in guanine and C4-N4 in cytosine may be specified by η6, η2, and η4, respectively. When the groups are substituted by hydrogen atoms only (see Fig. A.1C.2), the relevant dihedral angles defined by the sequence rules are: η61 = N1-C6-N6-H61 η62 = N1-C6-N6-H62 η21 = N1-C2-N2-H21 η22 = N1-C2-N2-H22 η41 = N3-C4-N4-H41 η42 = N3-C4-N4-H42. The rules may be adapted for substituted base and sugar rings, such as those of minor components of tRNA. Examples: 1-Methyladenosine: use η11, η12, η13 for the C-CH3 group conformation. 2′-O-Methyladenosine: use η2′ for rotation about the C2′-O2′ bond and η2′1, η2′2, η2′3 for the OCH3 group.
Note Recommendations governing the description of conformations of side chains and derivatives follow the sequence rules (Rule 2) and the side-chain rules (Rule 4) of the recommendations for polypeptides [11].
3. HYDROGEN BONDS 3.1. Polarity of Hydrogen Bonds In specifying a hydrogen bond, the atom covalently linked to the hydrogen atom is mentioned first, as in X-H...Y. The polarity of a hydrogen bond is from the hydrogen-atom donor to the acceptor.
3.2. Geometry of Hydrogen Bonds The hydrogen bond may be described by extension of the nomenclature of sections 3.1, 1.4, 1.5, and 1.6, so that for the hydrogen bond in the system Ci) − X(i) − H(i)...Y(k) − C(k) the following symbols may be used: b[H(i)...Y(k)] or b[H(i),Y(k)] τ[X(i)-H(i)...Y(k)] or τ[X(i),H(i),Y(k)] τ[H(i)...Y(k)-C(k)] or τ[H(i),Y(k),C(k)]. Where the positions of hydrogen atoms are not available, the following may be used: b[X(i),Y(k)] and τ[C(i),X(i),Y(k)]
4. HELICAL SEGMENTS A regular helix is strictly of infinite length, with the torsion angles in a nucleotide unit the same for all units. Two or more polynucleotide chains may associate in a helical complex through hydrogen bonding between base pairs. Torsion angles for each residue may differ for different chains in the same double, triple, etc., complex. A helical segment of a polynucleotide chain may be described in terms of the torsion angles of the nucleotide units or in terms of the helix characteristics summarized in section 4.2.
4.1. Base Pairs Base pairs with different geometries have been observed. These geometries should be denoted by the appropriate hydrogen-bonding scheme specifying both the heterocyclic base (e.g., Ade, Ura, Gua, Cyt) and the heteroatoms involved in the hydrogen bonding. In some cases it is also desirable to specify the nucleotide unit (i, j, etc.). Typical examples are: For a Watson-Crick A:U base pair: AdeN6:O4Ura, UraN3:N1Ade (if necessary, Ade(i)N6:O4Ura(j), etc.). For a reversed Watson-Crick A:U base pair: AdeN6:O2Ura, UraN3:N1Ade.
Standard Nomenclature, Data, and Abbreviations
A.1C.7 Current Protocols in Nucleic Acid Chemistry
4.2. Helix Characteristics In the description of helices or helical segments the following symbols should be used: n = number of residues per turn h = unit height (translation per residue along the helix axis) t = 360°/n = unit twist (angle of rotation per residue about the helix axis) n = pitch height of helix = nh. A polynucleotide may be accurately described in terms of the polar atomic coordinates ri, φi, zi where for each atom i, ri is the radial distance from the helix axis and φi and zi are the angular and height differences, respectively, relative to a reference point. The reference point should be either a symmetry element, as in RNA and DNA, or the C1′ atom of a nucleotide if no symmetry element between polynucleotide chains is present.
LITERATURE CITED [1] IUPAC-IUB JCBN (International Union of Pure and Applied Chemists–International Union of Biochemistry Joint Commission on Biochemical Nomenclature. 1983. Eur. J. Biochem. 131:9-15. [Also in Proceedings of the 16th Jerusalem Symposium, Nucleic Acids, the Vectors of Life (1983; B. Pullman and J. Jortner, eds.) pp. 559565; Pure Appl. Chem. (1983) 55:1273-1280; Biochemical Nomenclature and Related Documents, 2nd ed. (1992) pp. 115-221, Portland Press.] [2] Donohue, J. and Trueblood, K.N. 1960. J. Mol. Biol. 2:363-371. [3] Sasisekharan, V., Lakshminarayanan, A.V., and Ramachandran, G.N. 1967. In Conformation of Biopolymers II (G.N. Ramachandran, ed.) pp. 641-654. Academic Press, New York. [4] Arnott, S. and Hukins, D.W.L. 1969. Nature 224:886-888. [5] Sundaralingam, M. 1969. Biopolymers 7:821860. [6] Lakshminarayanan, A.V. and Sasisekharan, V. 1970. Biochim. Biophys. Acta 204:49-59. [7] Olson, W.K. and Flory, P.J. 1972. Biopolymers 11:1-23. [8] Sundaralingam, M., Pullman, B., Saenger, W., Sasisekharan, V., and Wilson, H.R. 1973. In Conformations of Biological Molecules and Polymers (E.D. Bergman and B. Pullman, eds.) pp. 815-820. Academic Press, New York. [9] Seeman, N.C., Rosenburg, J.M., Suddath, F.L., Kim, J.J.P., and Rich, A. 1976. J. Mol. Biol. 104:142-143. [10] Davies, D.B. 1978. In NMR in Molecular Biology (B. Pullman, ed.) pp. 509-516. Reidel, Dordrecht. The Description of Conformations of Polynucleotide Chains
[11] IUPAC-IUB CBN (Commission on Biochemical Nomenclature). 1971. Abbreviations and symbols for the description of the conformation of polypeptide chains, Tentative rules 1969 (approved 1974). Arch. Biochem. Biophys. 145:405421. [Also in Biochem. J. 121:577-585 (1971); Biochemistry, 9:3471-3479 (1970); Biochim. Biophys. Acta 229:l-17 (1971); Eur. J. Biochem. 17:193-201 (1970); J. Biol. Chem. 245:64896497 (1970); Mol. Biol. (in Russian) 7:289-303 (1973); Pure Appl.Chem. 40:291-308 (1974); on pp. 94-102 in [11]; and in Biochemical Nomenclature and Related Documents, 2nd ed., 1992, pp. 73-81, Portland Press.] [12] International Union of Biochemistry. 1978/1992. Biochemical Nomenclature and Related Documents, 2nd ed. The Biochemical Society, London/Portland Press. [13] IUPAC-IUB JCBN (Joint Commission on Biochemical Nomenclature). 1983. Symbols for specifying the conformation of polysaccharide chains, Recommendations 1981. Eur. J. Biochem. 131:5-7. [Also in Pure Appl. Chem. (1983) 55:1269-1272 and in Biochemical Nomenclature and Related Documents, 2nd ed., 1992, pp. 177-179, Portland Press.] [14] IUPAC-CMN (Commission on Macromolecular Nomenclature). 1981. Stereochemical definitions and notations relating to polymers. Pure Appl. Chem. 53:733-752. [15] IUPAC-IUB CBN (Commission on Biochemical Nomenclature). 1971. Abbreviations and symbols for nucleic acids, polynucleotides and their constituents, Recommendations 1970. Arch. Biochem. Biophys. 145:425-436. [Also in Biochem. J. 120:449-454 (1970); Biochemistry 9:4022-4027 (1970); Biochim. Biophys. Acta 247:1-12 (1971); Eur. J. Biochem. 15:203-208 (1970), corrected 25:l (1972); Hoppe-Seyler’s Z. Physiol. Chem. (in German) 351:1055-1063 (1970); J. Biol. Chem. 245:5171-5176 (1970); Mol. Biol. (in Russian) 6:167-174 (1972); Pure Appl. Chem. 40:277-290 (1974); also on pp. 116-121 in [11].] [16] IUPAC (International Union of Pure and Applied Chemistry). 1979. Nomenclature of Organic Chemistry, Sections A, B, C, D, E, F and H (J. Rigandy and S.P. Klesney, eds). Pergamon Press, Oxford. [17] IUPAC-CNOC (Commission on the Nomenclature of Organic Chemistry). 1960. Definitive rules for nomenclature of organic chemistry. J. Am. Chem. Soc. 82:5545-5574. [18] IUPAC Commission on Nomenclature of Organic Chemistry (CNOC). 1976. Rules for the nomenclature of organic chemistry, Section E: Stereochemistry, Recommendations 1974. Pure Appl. Chem. 45:11-30. [Also on pp. 473-490 in [15], on pp. 1-18 in [11]; and in Biochemical Nomenclature and Related Documents, 2nd ed. (1992) pp. 1-18, Portland Press.] [19] Klyne, W. and Prelog, V. 1960. Experientia 16:521-523. [20] Arnott, S. and Hukins, D.W.L. 1972. Biochem. Biophys. Res. Commun. 47:1504-1509.
A.1C.8 Current Protocols in Nucleic Acid Chemistry
[21] Pullman, B. Perahia, D., and Saran, A. 1972. Biochim. Biophys. Acta. 269:1-14.
[26] Saenger, W. 1973. Angew. Chem. Intl. Ed. Engl. 12:591-601.
[22] Saran, A. and Govil, G. 1971. J. Theor. Biol. 33:407-418.
[27] Altona, C. and Sundaralingam, M. 1972. J. Am. Chem. Soc. 94:8205-8212.
[23] Hall, L.D. 1963. Chem. Ind. (Lond.) (1963):950-951.
[28] Altona, C. and Sundaralingam, M. 1973. J. Am. Chem. Soc. 95:2333-2334.
[24] IUPAC-IUB JCBN (Joint Commission on Biochemical Nomenclature). 1981. Conformational nomenclature for five- and six-membered ring forms of monosaccharides and their derivatives, Recommendations 1980. Arch. Biochem. Biophys. 207:469-472. [Also in Eur. J. Biochem. 111:;295-298 (1980); Pure Appl. Chem. 53:1901-1905 (1981).]
[29] Kang, S. 1971. J. Mol. Biol. 58:297-315. [30] Saenger, W. and Scheit, K.H. 1970. J. Mol. Biol. 50:153-169. [31] Sundaralingam, M. 1973. In Conformations of Biological Molecules and Polymers (E.D. Bergman and B. Pullman, eds.) pp. 417-455. Academic Press, New York.
[25] Jardetzky, C.D. 1960. J. Am. Chem. Soc. 82:229-233.
Standard Nomenclature, Data, and Abbreviations
A.1C.9 Current Protocols in Nucleic Acid Chemistry
Nucleoside and Nucleotide Nomenclature As with any area of chemistry, the utilization of proper nomenclature when referring to nucleosides, nucleotides, and nucleic acids provides a clarity that facilitates understanding by other scientists. This chemical direction began in the nineteenth century with the isolation and identification of various constituents of nucleic acids, and continues unabated today with, for example, the chemical synthesis of oligonucleotide analogs of various lengths and with all manner of modifications. The term “nucleotide” was first used by P.A. Levene (Levene and Mandel, 1908); “nucleoside” was introduced by the same author the following year (Levene and Jacobs, 1909). The specific components of nucleic acids, as they were isolated from natural sources, were given names related to those sources that have remained with them over the years. For example, guanine was given its name because it was first isolated from bird excrement, i.e., guano (Unger, 1846a,b). Adenine was named by Kossel (1885, 1886) after isolation from bovine pancreas, the root being derived from the Greek word for gland. Interestingly, the first nucleotide that was actually isolated was inosinic acid, by Liebig (1847), and the first nucleoside isolated was guanosine, which was initially given the name vernine (Schulze and Bosshard, 1885, 1886). The main purpose of this appendix is to provide pertinent references that will direct the reader to the relevant guidelines or evolving nomenclature as described in the literature. When additional suggestions or guidance are appropriate, those comments will be included as well. One of the beauties and challenges of science as it evolves is the process of keeping pace with accurate, concise, and flexible systems of nomenclature. Chemists have always been able to discover or synthesize compounds that render existing systems inadequate. Current nomenclature in the area of nucleosides, nucleotides, and nucleic acids comprises a mixture of (1) common names that have gained official recognition, (2) guidelines that have been derived and officially recommended by the International Union of Pure and Applied Chemistry (IUPAC)/International Union of Biochemistry and Molecular Biology (IUBMB), and (3) evolving usage that is derived by individual scientists and laboratories and subjected to peer review through publication. A working group has been commissioned
APPENDIX 1D
(1998) by IUBMB to review guidelines for nucleotide (including oligonucleotide) nomenclature. As those guidelines are developed and made available, they will be referenced in future updates of this appendix. As noted earlier, the first nomenclature arose out of natural product isolation and identification, and thus common names were utilized for adenosine, guanosine, cytidine, uridine, and thymidine. The purine numbering system arose from that era, and it has been kept intact (see below) with the usual site of carbohydrate attachment at N-9. The pyrimidine numbering system was changed, at least as applied to the standard nucleosides, and in the current system the site of attachment of the carbohydrate in normal nucleosides is N-1 (earlier literature has the numbering such that attachment is at N-3). The accepted system of numbering for both purines and pyrimidines is illustrated in Figure A.1D.1. Nomenclature systems for the carbohydrate and nitrogen base moieties in nucleosides and nucleotides remain unchanged, except that primed numbers are used to refer to the carbohydrate atoms when the entire molecule is being considered. The fact that DNA and RNA differ mainly in the presence or absence of a 2′-hydroxyl in the carbohydrate pentofuranosyl moiety has simplified nomenclature. The only complicating factor is that uridine, which occurs in RNA, is replaced by thymidine in DNA, and unfortunately, thymidine, isolated first from thymus DNA, was not given a 2′-deoxy designation when named by P.A. Levene (Levene and London, 1929; Levene and Tipson, 1935). Nomenclature guidelines have maintained the original usage, with thymidine referring to a 2′-deoxynucleoside, while in all other cases the 2′deoxy is included within the name for the DNA component. Abbreviations that are used, however, can take into account the 2′-deoxy nature of thymidine quite readily. For example, the abbreviations for 2′-deoxyadenosine, 2′-de-
A
B N1
6
5 2 3 4
N
N 7
9
8
N
N3
4
5 2 1 6
N
Figure A.1D.1 Numbering system for (A) purines and (B) pyrimidines.
Contributed by John A. Secrist III Current Protocols in Nucleic Acid Chemistry (2000) A.1D.1.1-A.1D.3 Copyright © 2000 by John Wiley & Sons, Inc.
Standard Nomenclature, Data, and Abbreviations
A.1D.1
Table A.1D.1
Available Nomenclature Recommendations on Nucleic Acids
Title Nomenclature of Carbohydrates Abbreviations and Symbols for Nucleic Acids, Polynucleotides, and their Constituents (published 1970) Abbreviations and Symbols for the Description of Conformations of Polynucleotide Chains (published 1982) Nomenclature for Incompletely Specified Bases in Nucleic Acid Sequences (published 1984) Nomenclature of Junctions and Branchpoints in Nucleic Acids (published 1994)
oxyguanosine, 2′-deoxycytidine, and thymidine are dA, dG, dC, and dT.
Nomenclature Documents
Nucleoside and Nucleotide Nomenclature
Two excellent Web sites are available for learning about nomenclature documents coming from IUPAC, IUBMB, and the Joint Commission on Biochemical Nomenclature. The documents mentioned below are also included on these websites: http://www.chem.qmw.ac.uk/ iubmb and http://www.chem.qmw.ac.uk/iupac/. Chemists wishing to learn about nucleoside nomenclature have several sources to consult. To obtain information on carbohydrate nomenclature, a recent revision/updating of the IUPAC recommendations “Nomenclature of Carbohydrates,” is available. For these recommendations as well as for others listed below, the literature citations will include all the journals that have reproduced them, so that readers can consult the one most readily available to them. For example, the carbohydrate nomenclature document is available in McNaught (1996, 1997a,b,c). The basic set of recommendations for nucleosides, nucleotides, and nucleic acids, entitled “Abbreviations and Symbols for Nucleic Acids, Polynucleotides, and Their Constituents,” was published in 1970 and is available in IUPAC-IUB (1970a,b, 1971a,b,c,d, 1972, 1974). This broadly-based document covers normal nucleosides and their abbreviations, as well as nucleotides, oligonucleotides, and polynucleotides. Information on names and abbreviations for modified nucleosides that occur in various nucleic acids, including tRNA, is also included. Available nomenclature recommendations on nucleic acids are listed in Table A.1D.1.
Available in McNaught (1996, 1997a,b,c) IUPAC-IUB (1970a,b, 1971a,b,c,d, 1972, 1974) IUPAC-IUB (1983a,b)
NC-IUB (1985a,b, 1986a,b,c)
NC-IUBMB (1995a,b, 1996)
A summary of the modified nucleosides in RNA is available, along with symbols and notes and references relevant to naming these nucleosides (Limbach et al., 1994). Updates of this reference are available through the following Web site: http://medlib.med.utah.edu/RNAmods/.
LITERATURE CITED IUPAC-IUB. 1970a. Abbreviations and symbols for nucleic acids, polynucleotides, and their constituents. Eur. J. Biochem. 15:203-208. IUPAC-IUB. 1970b. Abbreviations and symbols for nucleic acids, polynucleotides, and their constituents. J. Biol. Chem. 245:5171-5176. IUPAC-IUB. 1971a. Abbreviations and symbols for nucleic acids, polynucleotides, and their constituents. Arch. Biochem. Biophys. 145:425-436. IUPAC-IUB. 1971b. Abbreviations and symbols for nucleic acids, polynucleotides, and their constituents. Biochem. J. 120:449-454. IUPAC-IUB. 1971c. Abbreviations and symbols for nucleic acids, polynucleotides, and their constituents. Biochemistry 9:4022-4027. IUPAC-IUB. 1971d. Abbreviations and symbols for nucleic acids, polynucleotides, and their constituents. Biochim. Biophys. Acta 247:1-12. IUPAC-IUB. 1972. Abbreviations and symbols for nucleic acids, polynucleotides, and their constituents. Eur. J. Biochem. 25:1. IUPAC-IUB. 1974. Abbreviations and symbols for nucleic acids, polynucleotides, and their constituents. Pure Appl. Chem. 40:277-290. IUPAC-IUB. 1983a. Abbreviations and symbols for the description of conformations of polynucleotide chains. Eur. J. Biochem. 131:9-15. IUPAC-IUB. 1983b. Abbreviations and symbols for the description of conformations of polynucleotide chains. Pure Appl. Chem. 55:12731280. Kossel, A. 1885. Über eine neue Base aus dem Thierkörper. Ber. 18:79-81.
A.1D.2 Current Protocols in Nucleic Acid Chemistry
Kossel, A. 1886. Weitere Beiträge zur Chemie des Zellkerns. Z. Physiol. Chem. 10:248-264. Levene, P.A. and Mandel, J.A. 1908. Über die Konstitution der Thymo-nucleinsäure. Chem. Ber. 41:1905-1909. Levene, P.A. and Jacobs, W.A. 1909. Über die HefeNucleinsäure. Chem. Ber. 42:2474-2478. Levene, P.A. and London, E.S. 1929. The structure of thymonucleic acid. J. Biol. Chem. 83:793. Levene, P.A. and Tipson, R.S. 1935. The ring structure of thymidine. J.Biol. Chem. 109:623. Liebig, J. 1847. Über die Bestandtheile der Hüssigkeiten des Heisches. Ann. 62:257-369. Limbach, P.A., Crain, P.F., and McCloskey, J.A. 1994. Summary: The modified nucleosides of RNA. Nucl. Acids Res. 22:2183-2196. McNaught, A.D. 1996. Nomenclature of carbohydrates. Pure Appl. Chem. 68:1919-2008. McNaught, A.D. 1997a. Nomenclature of carbohydrates (recommendations 1996). Adv. Carbohydr. Chem. Biochem. 52:43-177. McNaught, A.D. 1997b. Nomenclature of carbohydrates. J. Carbohydr. Chem. 16:1191-1280. McNaught, A.D. 1997c. Nomenclature of carbohydrates. Carbohydr. Res. 297:1-92. NC-IUB. 1985a. Nomenclature for incompletely specified bases in nucleic acid sequences. Biochem. J. 229:281-286. NC-IUB. 1985b. Nomenclature for incompletely specified bases in nucleic acid sequences.Eur. J. Biochem. 150:1-5. NC-IUB. 1985c. Nomenclature for incompletely specified bases in nucleic acid sequences. Nucl. Acids Res. 13:3021-3030.
NC-IUB. 1986a. Nomenclature for incompletely specified bases in nucleic acid sequences. J. Biol. Chem. 261:13-17. NC-IUB. 1986b. Nomenclature for incompletely specified bases in nucleic acid sequences. Mol. Biol. Evol. 3:99-108. NC-IUB-1986c. Nomenclature for incompletely specified bases in nucleic acid sequences. Proc. Nat. Acad. Sci. U.S.A. 83:4-8. NC-IUBMB. 1995a. Nomenclature of junctions and branchpoints in nucleic acids. Eur. J. Biochem. 230:1-2. NC-IUBMB. 1995b. Nomenclature of junctions and branchpoints in nucleic acids. Nucleic Acid Res. 23:3363-3364. NC-IUBMB. 1996. Nomenclature of junctions and branchpoints in nucleic acids. J. Mol. Biol. 554555. Schulze, E. and Bosshard, E. 1885. Zur Kenntniss des Vorkommens von Allantoin, Asparagin, Hypoxanthin und Guanin in den Pflanzen. Z. Physiol. Chem. 9:420-444. Schulze, E. and Bosshard, E. 1886. Über einen neuen stickstoffhaltigen Pflanzenbestandtheil. Z. Physiol. Chem. 10:80-89. Unger, B. 1846a. Bemerkungen zu obiger Notiz. Ann. 58:18-20. Unger, B. 1846b. Das Guanin und seine Verbindungen. Ann. 59:58-68.
Contributed by John A. Secrist III Southern Research Institute Birmingham, Alabama
Standard Nomenclature, Data, and Abbreviations
A.1D.3 Current Protocols in Nucleic Acid Chemistry
A Convenient Stereochemical Notation for P-Chiral Nucleotide Analogs The configuration of P-chiral organic compounds is generally described by adaptation of the CIP (Cahn-Ingold-Prelog) convention for the phosphorus atom (Cahn et al., 1966; Prelog and Helmchen, 1982; Mata et al., 1993). Using this convention, it is possible to assign RP /SP configuration to tetra- and tri-coordinated phosphorus compounds (analogous to a chiral sp3 carbon center). The CIP system is reliable and universal. However, due to a formal character of its priority rules, stereochemically related compounds with the same spatial distribution of substituents often may have opposite configurations. This issue was recognized for amino acids and carbohydrates, for which the D/L convention remained in common use and is approved by IUPAC (IUPAC, 1971, 1984). By analogy, all P-chiral nucleotide analogs seem to fall into one category of molecules based on the same framework of dinucleoside monophosphates. Thus, a notation making use of these structural similarities, rather than formal priorities of ligands, should simplify assignment of stereochemical descriptors and readout of a configuration. While the convenience of this approach was noted by Lebedev and Wickstrom, their “pseudoaxial/pseudoequatorial” terms have never been developed into a comprehensive system (Lebedev and Wickstrom, 1996). Presented here is the DP /LP stereochemical notation for chiral dinucleotide monophosphate derivatives, which highlights a relationship between compounds possessing similar structural motifs (Sobkowski et al., 2005, 2006a,b). The notation allows direct comparison of physical and biochemical properties of compounds with the same spatial arrangement of ligands at the phosphorus center, irrespective of their RP /SP notation—properties such as susceptibility to enzymatic digestion, stability of formed aggregates, analysis of 31 P NMR chemical shifts, and tracing stereochemistry of reactions. Below, the rules for the assignment of the DP /LP configuration and several examples of their applications to P-chiral dinucleoside monophosphate analogs are presented. An extension of the DP /LP notation to analogs containing a single nucleoside unit is also presented.
APPENDIX 1E
The DP /LP nomenclature is not intended as a replacement for the absolute RP /SP notation in the field of nucleic acids, but rather as a complement. Although this notation works accurately for a wide range of P-chiral nucleotide analogs, its fundamentals are less rigid than those of the CIP convention. To avoid confusion or possible ambiguities, we suggest using the DP /LP descriptors together with the RP /SP descriptors.
GENERAL PRINCIPLES Definition of the DP /LP Notation For the assignment of the DP /LP configuration, a dinucleoside monophosphate fragment should be presented with a nucleosid3 -yl as an upper unit and a nucleosid-5 -yl as a lower one, while two remaining ligands should be placed horizontally according to the absolute configuration of a given compound. The spatial location of ligands at phosphorus should follow the Fischer projection, i.e., bonds drawn vertically should lie below the projection plane and bonds drawn horizontally should lie above the plane (Fig. A.1E.1). The left-hand side position of a single-bonded ligand defines the LP configuration of phosphorus, while its right-hand side position defines the DP configuration. In generic structures for dinucleoside monophosphates with DP and LP configurations depicted in Figure A.1E.1, the ligands at the phosphorus center are designated as G1 , G2 , X, and Z. Ligands G1 and G2 have fixed positions as the upper and lower units, respectively, while ligands X and Z can take either the left or right position, depending on absolute configuration at the phosphorus center. For DNA or RNA analogs, G1 stands for a nucleosid-3 -yl and G2 for a nucleosid-5 yl, and Y1 and Y2 usually are integral parts of these moieties. A ligand forming a double bond to the phosphorus (e.g., oxygen or sulfur in the P O and P S groups, respectively) is always designated as X.
Rules for Spatial Allocation of Ligands Rule 1 For dinucleoside phosphates presented in a form similar to the Fischer projection as shown
Contributed by Michal Sobkowski, Jacek Stawinski, and Adam Kraszewski Current Protocols in Nucleic Acid Chemistry (2007) A.1E.1-A.1E.16 C 2007 by John Wiley & Sons, Inc. Copyright
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Figure A.1E.1
Generic structures for definition of the DP /LP notation.
in Figure A.1E.1, i.e., with a nucleosid-3 -yl standing for G1 and a nucleosid-5 -yl standing for G2 moiety, DP configuration is defined as the one having a single-bonded ligand Z to the right and the P X group to left. For compounds with LP configuration, Z is to the left and P X is to the right. X is any atom doublebonded to the phosphorus or a free electron pair. Y1 and Y2 are atoms, or groups of atoms, that are integral parts of G1 and G2 .
Rule 2 For nucleotide analogs having carbohydrate residues other than ribo- or deoxyribofuranose, the assignment of ligands as G1 and G2 is done primarily with respect to their resemblance to the natural nucleosidic residues. If such an analogy is not obvious, the following rules should be used: A. A carbohydrate residue having a phosphorus center bound to a carbon atom of higher order is assigned as G1 . B. If carbon atoms to which the phosphorus center is attached are of the same order in both carbohydrate residues, the residue of higher CIP priority is assigned as G1 .
Rule 3
Stereochemical Notation for P-Chiral Nucleotide Analogs
If both X and Z can form a double bond with phosphorus, the following order of priority should be used for the assignment of ligand X (P X bond): A. P O > P S > P Se > P Te > P N. B. In other instances, the double bond should be set to an atom of lower CIP priority. Configurations with X and Z assigned opposite to that specified by Rule 3A or 3B should be referred to as pseudo-DP or pseudoLP .
Rule 4 If the G1 or G2 group is a non-nucleosidic residue (e.g., alkyl, aryl, acyl, sulfonyl, or phosphoryl), the nucleoside keeps its original position as G1 or G2 , while the assignment of non-nucleosidic ligands is done as follows: A. If only one ligand can form a double bond with phosphorus, that ligand should be designated as X (P X bond). Among the remaining ligands, the residue of lower CIP priority should be assigned as Z. B. If two ligands can form a double bond with phosphorus, they should be designated as X and Z according to Rule 3. C. If all three ligands can form a double bond with phosphorus, the assignment of X (P X bond) should be made according to Rule 3, and among the remaining ligands the residue of lower CIP priority should be designated as Z. In all three cases, the fourth ligand becomes the second G group.
Rule 5 If G1 and G2 groups are non-nucleosidic residues, ligands X and Z should be assigned according to the appropriate Rules 1 to 4, while the ligand allocation to G1 and G2 positions is governed by the CIP convention: the group with higher CIP priority should be designated as G1 , and the one with lower CIP priority as G2 .
APPLICATIONS OF DP /LP DESCRIPTORS Dinucleoside Monophosphate Analogs A comprehensive discussion on applications of the DP /LP notation to dinucleoside
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Figure A.1E.2 P O bond.
Examples of DP (A) and LP (B) notation for dinucleoside monophosphate analogs with a
monophosphate analogs can be found in Sobkowski et al. (2006a).
Dinucleoside monophosphate analogs containing the P O bond Diribo- (or deoxyribo-) nucleoside monophosphate derivatives contain two nucleosidic ligands that are defined as G1 and G2 , while X is an oxygen. In this case, DP /LP configuration can be read immediately from the position of the Z group: the compound has DP configuration when group Z is pointing to the right and LP configuration when Z is pointing to the left (Fig. A.1E.2). The following points should be noted. 1. A correlation between DP /LP and RP /SP configurations can be easily found using the following rule of thumb: DP = RP (and LP = SP ) if atomic mass of the Z atom (a single-bonded ligand) >16, DP = SP (and LP = RP ) if atomic mass of the Z atom (a single-bonded ligand) <16. This applies only to analogs containing the P=O bond. 2. The DP /LP notation can be applied to P-chiral internucleotide linkages of nucleic acids analogously as the RP /SP convention is used. However, in contrast to the absolute RP /SP configuration, the new notation unequivocally correlates configuration at the phosphorus center with a spatial position of the attached ligands in a nucleic acid framework.
For example, in the case of a double-stranded B-DNA form, a Z substituent of DP diastereomers is always located in the major groove, while a Z substituent for the LP counterparts always points away from the duplex. 3. Since phosphorus in a natural internucleotide bond is prochiral, for the purpose of stereochemical correlation analysis, the two nonbridging oxygens of the phosphate group can be designated as pro-DP and pro-LP . Contrary to pro-RP /pro-SP descriptors, these have absolute meaning: replacement of a pro-DP (or pro-LP ) oxygen by any other atom will result in an analog having DP or LP configuration, respectively (Fig. A.1E.3). In B-DNA, pro-DP oxygen thus points to the major groove, while pro-LP oxygen points to the bulk of solvent.
Dinucleoside monophosphate analogs without a P O bond There are many phosphate analogs in which the phosphoryl group P O has been replaced by a P X functionality, where X can be S, Se, N, C, and so on. A set of exemplary nonionic compounds is shown in Figure A.1E.4. Since bond orders for ligands attached to the phosphorus centers in these compounds are well defined, the assignment of DP /LP configuration is usually rather straightforward. For derivatives that are not fully esterified, a more careful analysis is required (see discussion of ambident anions).
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Figure A.1E.3
Figure A.1E.4
A structure for definition of pro-DP /pro-LP positions.
DP /LP notation applied to dinucleoside monophosphate analogs without a P O bond.
Dinucleoside phosphite analogs Although phosphite triesters and their analogs (e.g., phosphinites and phosphoramidites) contain only single-bonded ligands, they can be accommodated within the DP /LP system by treating a lone electron pair as ligand X and a non-nucleosidic moiety as ligand Z (Rule 1). Examples of the application of the DP /LP notation to tervalent phosphorus compounds are shown in Figure A.1E.5.
Dinucleoside monophosphate analogs containing modified ribose or other sugar moieties
Stereochemical Notation for P-Chiral Nucleotide Analogs
The DP /LP notation can be applied to systems containing various furanoses (e.g., modified ribose, arabinose, or xylose), hexoses, or acyclic nucleoside derivatives. Moreover, neither inversion of stereochemistry at the 1 position nor lack of an aglycone affects the DP /LP configuration, allowing application of this system to α-nucleotides, fragments contain-
ing abasic units, or carbohydrate phosphates (Fig. A.1E.6). For derivatives containing various sugar units, the assignment of ligands for the G1 and G2 positions is governed by Rule 2. The superior principle of this assignment is the structural similarity of a carbohydrate residue of a phosphodiester unit to the natural dinucleoside monophosphate skeleton. Only if such analogy is ambiguous should further steps of Rule 2 be invoked. Thus, in diester S.1 (Fig. A.1E.6), although the attachment point of 5,5 -dimethylribose is the 3◦ carbon, this unit should be designated as the G2 ligand due to its obvious resemblance to a ribose-5 -yl moiety. In compound S.2, the hexose-6 -yl residue was assigned as G2 due to its similarity to a ribose-5 -yl group. For structures containing a diester scaffold unsuitable for direct comparison with dinucleoside monophosphate, Rules 2A and 2B should be applied. First, if the attachment
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Figure A.1E.5
DP /LP notation for tervalent phosphorus dinucleoside analogs.
Figure A.1E.6
DP /LP notation for phosphoesters containing nucleoside analogs.
sites of a phosphate group to carbohydrate residues differ in carbon orders (1◦ , 2◦ , or 3◦ ), the G1 position (an upper unit) should be assigned to a group of higher order and the G2 position (a lower unit) to the ligand attached through a carbon of lower order (Rule 2A; e.g., S.3 in Fig. A.1E.6). If a phosphate moiety is bonded to two carbohydrate residues with attachment carbons of the same order, the unit of higher CIP priority should be assigned to position G1 according to Rule 2B. For
example, for the symmetric 3 -3 phosphorothioate S.4, adenine is superior to thymine according to the CIP priority rules, and thus the deoxyadenosine moiety is assigned to the G1 position. The DP /LP system can also be applied to nucleotide analogs containing any atom in place of oxygen in a bridging position of the phosphorus center (Y1 and/or Y2 , see Fig. A.1E.1). In contrast to the RP /SP system, such modifications do not affect DP /LP configuration.
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Stereochemical Notation for P-Chiral Nucleotide Analogs
Figure A.1E.7
Mesomeric forms of phosphorothioate diester analogs.
Figure A.1E.8
Tautomers of phosphorothioate diesters.
Figure A.1E.9
DP /LP notation for isotopomers.
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Dinucleoside monophosphate analogs containing ambident anions Partially esterified P-chiral derivatives are often drawn in their anionic forms. In such compounds, the negative charge is delocalized, resulting in fractional bond orders between phosphorus and both nonbridging atoms (e.g., phosphorothioate S.5, Fig. A.1E.7). In principle, the DP /LP notation is not applicable to compounds in which the charge is distributed over the phosphorus center, since none of the ligands can be assigned as
either X or Z. For such derivatives, one mesomeric structure with the negative charge localized on the Z atom should be chosen for the purpose of assigning a DP /LP configuration. Rule 3 controls which of the two heteroatoms should be treated as ligand X (double-bonded to phosphorus) and ligand Z (bearing a formal negative charge). The priority order for a P X residue was defined as follows: P O > P S > P Se > P Te > P N Such an arbitrarily chosen order of priority is congruent with the most common
Figure A.1E.10 An example of correlation between the DP /LP notation and the stereochemistry of reactions of dinucleoside monophosphate analogs (some reactions are hypothetical only).
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way of presenting ambident anions in the literature. Thus, phosphorothioates should be considered as phosphorothiolates (O P S− , e.g., as S.6-DP or S.6-LP , Fig. A.1E.7), phosphoroselenoates as phosphoroselenolates (O P Se− ), phosphoroselenothioates as phosphoroselenothionates (S P Se− ), and so on. If it is necessary to discuss separately two mesomeric forms of such ambident esters, the descriptors “pseudo-LP ” and “pseudo-DP ” may be used to indicate formulas that do not follow Rule 3A/3B (e.g., S.6-DP versus S.6pseudo-LP ). Tautomeric structures When ambident phosphodiester analogs are protonated and form neutral species, their single and double bonds are clearly distinguishable, allowing direct assignment of the DP /LP configuration. As charged diesters have two mesomeric forms, the neutral species may exist in two tautomeric (ono/olo) forms. Such tautomers should be treated as separate compounds (e.g., S.7 and S.8, Fig. A.1E.8) having opposite DP /LP configurations. Examples of these types of compounds include phosphorothioates, phosphoroselenoates, or other types of chiral ambident phosphodiesters. If the kind of a tautomeric form is unknown for a given compound, Rule 3 should be applied for the assignment of ligands Z and X. For example, for the tautomeric pair shown in Figure A.1E.8, structure S.7 should be used because of the priority of P O > P S.
Stereochemical Notation for P-Chiral Nucleotide Analogs
Figure A.1E.11
Dinucleoside monophosphate isotopomers Rule 3A for designation of ligands X and Z is sufficient for most cases encountered in nucleotide chemistry. When this is not decisive, Rule 3B should be used. It states that if the assignment of X and Z positions cannot be made on the basis of the previous rules, the double bond should be assigned to the atom of lower CIP priority. An example of the application of Rule 3B is the assignment of DP /LP descriptors to isotopomers (Fig. A.1E.9). For dinucleoside monophosphate S.9, which is labeled with 18 O at the nonbridging position of the phosphorus center, 16 O (lower CIP priority) should be chosen as the double-bonded ligand X and 18 O (higher CIP priority) as ligand Z.
Relationship between DP /LP notation and stereochemistry of reactions Apart from naming compounds, the DP /LP notation can be applied for conveniently tracing the stereochemistry at phosphorus during reactions. Contrary to the CIP notation, inversion of configuration at phosphorus in most cases causes a change in DP /LP stereochemical descriptors, while stereoretentive processes do not affect the DP /LP descriptors. More care is required during stereochemical analysis of reactions including ambident anions (see above), for which one should choose a canonical or “pseudo” form that fits the reaction pathway. An example of a reaction sequence involving a phosphorus center is shown in Figure A.1E.10. The H-phosphonothioate
DP /LP notation for compounds with one nucleoside residue.
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Figure A.1E.12 General structures of non-ionic phosphorus esters with one nucleoside residue (DP configurations shown; CIP notation may vary from case to case).
S.10-LP is oxidized with iodine to phosphoroiodidothioate S.11-LP with retention of configuration, which is reflected by retention of the LP descriptor. The CIP notation in this instance does not follow the stereochemistry, as the RP descriptor of S.10 changes to SP for S.11. In the next steps, diester S.11-LP is esterified with an alcohol to triester S.12DP , or is transformed into other halophosphate analogs, S.13-DP or S.14-DP . All these reactions are stereoinvertive and the DP /LP descriptors change accordingly. For the RP /SP notation, no such correlation can be found
as the SP descriptor changes into RP only for the reaction from S.11 to S.14. Finally, amination of phosphorochloridothioate S.14-DP yields amide S.15-LP , with proper correlation of DP /LP descriptors with the inversion.
Nucleotide Analogs Containing Only One Nucleoside Unit The DP /LP notation can also be used for compounds that have only one nucleosidic ligand G, for example, to investigate the stereochemistry of reactive intermediates (e.g., S.16, Fig. A.1E.11; Sobkowski et al., 2006b). The
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Figure A.1E.13 Examples of DP /LP notation for mononucleosidic derivatives with only one ligand that can form a P X bond.
assignment of DP /LP configuration of such mononucleotide derivatives is based on Rules 1 to 3, assisted by Rule 4, which refers to non-nucleosidic ligands. According to Rule 4, compounds having one nucleosidic residue are grouped into three classes depending on the number of ligands that can form a double bond with phosphorus. This rule can also be applied to P-chiral analogs of nucleoside polyphosphates and cyclic phosphates (see below).
Compounds with one ligand that can form a double bond to phosphorus Stereochemical Notation for P-Chiral Nucleotide Analogs
Nucleoside monophosphate analogs bearing one localized P=X bond comprise a broad range of fully substituted (i.e., non-ionic and without dissociable hydrogen) tri- or tetracoordinated phosphoesters, phosphoamides,
halophosphates, etc., along with their analogs bearing various heteroatoms in place of one or more oxygens at the phosphorus center. Figures A.1E.12 and A.1E.13 show several examples of such mononucleotide analogs. The procedure of assigning a DP /LP configuration starts with putting the nucleoside and the P=X moiety in proper positions of general structures (Fig. A.1E.1) according to Rules 1 to 3. Then, among the two remaining ligands, the one of lower CIP priority is designated as Z and the one of higher priority as the second G group (Rule 4A). For compounds S.19 to S.26 (Fig. A.1E.13), a nucleosidic residue always takes the G1 or G2 position (Rule 1), while X is designated to a double-bonded chalcogen or an electron pair. The other ligands attached to the phosphorus center are assigned as Z and
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the second G according to their CIP priority. For example, (1) since CH3 < Ph, Z = CH3 and G2 = Ph for phosphinite S.20; (2) since OEt < OtBu, Z = OEt and G2 = OtBu for triester S.23; and (3) since NH-Ar < O-Ar, Z = NH-Ar and G1 = O-Ar for phosphoramidate S.24.
Compounds with two ligands that can form a double bond to phosphorus For mononucleosidic derivatives bearing two ligands that can form a double bond to phosphorus, the assignment of DP /LP configuration can be made most conveniently by initial designation of the two single-bonded ligands
(one of them being a nucleosidic residue) as G1 and G2 according to Rules 1 and 2. Rule 3 then specifies the priority order for the remaining ambident ligands as X and Z. General structures of such ambident mono- and diesters are shown in Figure A.1E.14, and examples of the correct notation for this class of compounds are given in Figure A.1E.15. In each case, the designation of nucleosid-3 -yl as G1 is obvious, and there is no doubt about the assignment of the second ligand that cannot form a double bond as G2 . Both of the remaining moieties are able to form a double bond and they take horizontal positions in a graphical presentation. The position of a double bond (ligand X)
Figure A.1E.14 General structures of ambident phosphorus esters with one nucleoside residue (DP configuration shown).
Figure A.1E.15 Examples of DP /LP notation for mononucleosidic derivatives with two ligands able to form a P X bond.
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is designated according to priorities given by Rule 3: P=O > P=S (Rule 3A, S.27), P=O > P=Se (Rule 3A, S.28), P=S > P=N (Rule 3A, S.29), and P=32 S > P=35 S (Rule 3B, S.30).
Compounds with three ligands that can form a double bond to phosphorus Mononucleosidic derivatives bearing three ligands that can form a double bond to phosphorus have a general structure shown in Figure A.1E.16. Among these three ligands, the one of the highest priority for the formation of a double bond to phosphorus is found using Rule 3 and is assigned as X. Among the two remaining ligands, the one of lower CIP priority is assigned as Z, and the last one takes the remaining G2 position. For example, for phosphoroamidoselenoate S.31 (Fig. A.1E.17), the oxygen has a higher priority for the P=X bond
compared to selenium and nitrogen (Rule 3A). Among the last two, the NH2 moiety is designated as Z as its CIP priority is lower than that of Se. Similarly, for the doubly labeled phosphorodithioate S.32, the oxygen is assigned as X (Rule 3A), while 32 S is assigned as Z (CIP: 35 S > 32 S). In the case of phosphorothioate S.33, oxygen isotope 16 O of lower CIP priority is assigned as X (Rule 3B), while 17 O is assigned as Z. The sulfur is designated as the G2 ligand because its CIP priority is higher than oxygen.
P-Chiral nucleoside polyphosphate analogs To apply the DP /LP convention to P-chiral nucleoside di-, tri-, or polyphosphate analogs, the structures should be drawn with the polyphosphate chain placed vertically, and
Figure A.1E.16 General structures of ambident phosphorus esters with one nucleoside residue (DP configuration shown).
Stereochemical Notation for P-Chiral Nucleotide Analogs
Figure A.1E.17 Examples of DP /LP notation for mononucleosidic derivatives with three ligands able to form a P X bond.
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Figure A.1E.18
Examples of DP /LP notation for P-chiral nucleoside polyphosphate analogs.
each phosphate residue should be congruent to the Fischer projection. Such presentation forces designation of nucleosidic and phosphoric moieties as G1 and/or G2 groups. The remaining ligands should be assigned according to Rules 1 to 4. Three examples of P-chiral nucleoside diand triphosphate analogs are shown in Figure A.1E.18. For nucleoside diphosphate analog S.34, the chiral phosphoric residue is in the α position. The G1 and G2 groups should be assigned to the β-phosphate (Rule 4D) and nucleoside (Rule 1), respectively. The oxygen is obviously assigned as the double-bonded X ligand, and the methyl group as the Z ligand. For chiral β-phosphoramidate residue in triphosphate derivative S.35, the G1 group is assigned to the γ-phosphate and G2 to the nucleotide moiety (Rule 4D). The priority order given by Rule 3A then governs the designation of the oxygen as X and the NH2 moiety as Z. Finally, triphosphate derivative S.36 gives an example of a compound bearing three P-chiral residues. For the α and β moieties, the G1 group is assigned to a phosphate and the G2 group to the nucleoside or nucleotide residues, respectively. In the next step, the oxygen is assigned as X because it has priority for the P=X residue over the sulfur (Rule 3A), which becomes the Z ligand. The γ-phosphorothioamidate moiety has two
ligands (O, S) able to form a double bond to phosphorus, so assignment of its configuration is governed by Rule 4B. Thus, oxygen is assigned as X and sulfur as Z, while the imidazoyl and a nucleotide residues are G1 and G2 , respectively.
P-Chiral cyclic nucleotide analogs A prerequisite of using the DP /LP notation is a graphical presentation of a phosphate center of a given structure in the form of a Fisher projection with nucleosidic residues placed in vertical positions. Thus, in order to apply DP /LP convention to P-chiral derivatives of cyclic nucleotides, these have to be presented with the nucleoside attachment pointing in the upper and the lower positions of the phosphorus tetrahedron. The proper assignment of 2 , 3 , or 5 positions of the ribose moiety as G1 and G2 is governed by Rule 2. Examples of popular presentations of 3 ,5 (S.37 to S.39) and 2 ,3 - (S.41 to S.42) cyclic phosphate derivatives are shown in Figure A.1E.19. For the assignment of DP /LP configuration, RP cyclic phosphoramidate S.37 to S.39 should be presented in the form of S.40 (with nucleosidic residues placed in vertical positions). According to Rule 2A, the secondary attachment point (C3 ) is assigned as G1 and the primary attachment (C5 ) as G2 . Keeping the absolute configuration intact and applying
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Figure A.1E.19
Stereochemical Notation for P-Chiral Nucleotide Analogs
Various presentations of P-chiral cyclic nucleotide analogs.
Rule 3 to select X and Z ligands, this particular compound is found to be in the LP configuration. In the case of the 2 ,3 -cyclic phosphoroselenoate (S.41-S.42), the same procedure is applied except that Rule 2B is used instead of Rule 2A to determine the G1 and G2 positions. Since the 2 - and 3 -carbons are of the same order but 2 -C has a higher CIP priority, the former one is designated as G1 (S.43). Note that, for nucleoside cyclic phosphates, DP /LP configurations can also be determined directly from a “standard” graphical presentation. If the Z ligand points up (as for S.37, S.39, and S.42) or upward (as for S.38 and S.41), the configuration is LP . Otherwise, if Z points down or downward, the configuration is DP .
Relationship between DP /LP notation and stereochemistry of reactions The DP /LP notation can be used to trace the stereochemistry of reactions of mononucleosidic P-chiral derivatives, analogously as shown for dinucleosidic compounds (see above). For example, activation of nucleoside H-phosphonate S.44 with pivaloyl chloride yields a mixed anhydride. The fate of the LP -(SP ) diastereomer of this compound, S.45, is shown in Figure A.1E.20. The reaction of S.45 with pyridine yields DP -(RP ) adduct S.46 (an inversion of configuration). This can be esterified with phenol to afford LP -(SP ) diester S.47 (inversion), which in turn can be converted into DP -(RP ) H-phosphonoamidate S.48 (inversion) or transesterified into DP -(SP )
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Figure A.1E.20 An example of the correlation between the stereochemistry of reactions of mononucleoside monophosphate analogs and the DP /LP notation (some reactions are hypothetical only).
dinucleoside H-phosphonate S.49 (inversion). Stereoretentive sulfurization of S.49 yields DP -(RP ) phosphorothioate S.50. All transformations shown give perfect match of stereochemistry and its DP /LP notation. It should be noted, however, that this correlation does not hold to the same extent as for dinucleo-
sidic derivatives. Nevertheless, in most cases, the DP /LP notation properly reflects the stereochemistry of reactions, while such correlations using the CIP convention are incidental (e.g., stereoinvertive substitution of SP diester S.47 leads either to SP diester S.49 or to RP amidate S.48).
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P-Chiral Non-Nucleosidic Compounds The DP /LP convention can, in principle, be used for non-nucleosidic phosphorus compounds as prescribed by Rule 5. Such an extension of the DP /LP system could facilitate comparison of the stereochemistry of the phosphorus centers in nucleosidic and nonnucleosidic compounds, or could be used, for example, to trace stereochemistry during phosphorylation of nucleosides. This application is beyond the scope of this manual and will not be discussed here.
LITERATURE CITED Cahn, R.S., Ingold, C., and Prelog, V. 1966. Specification of molecular chirality. Angew. Chem., Int. Ed. 5:385-415. IUPAC Commission on the Nomenclature of Organic Chemistry (CNOC) and IUPAC-IUB Commission on Biochemical Nomenclature (CBN). 1971. Tentative rules for carbohydrate nomenclature. Eur. J. Biochem. 21:455477. IUPAC-IUB Joint Commission on Biochemical Nomenclature (JCBN). 1984. Nomenclature and symbolism for amino acids and peptides, recommendations 1983. Pure Appl. Chem. 56:595624. Lebedev, A.V. and Wickstrom, E. 1996. The chirality problem in P-substituted oligonucleotides. Perspect. Drug Discov. Design 4:17-40. Mata, P., Lobo, A.M., Marshall, C., and Johnson, A.P. 1993. The CIP sequence rules: Analysis and
proposal for a revision. Tetrahedron Asymmetry 4:657-668. Prelog, V. and Helmchen, G. 1982. Basic principles of the CIP-system and proposals for a revision. Angew. Chem., Int. Ed. 21:567-583. Sobkowski, M., Stawinski, J., and Kraszewski, A. 2005. A proposal for a new stereochemical notation for P-chiral nucleotide analogues and related compounds. Nucleosides Nucleotides Nucleic Acids 24:1301-1307. Sobkowski, M., Stawinski, J., and Kraszewski, A. 2006a. A proposal for a convenient notation for P-chiral nucleotide analogues. Part 2. Dinucleoside monophosphate analogues. Nucleosides Nucleotides Nucleic Acids 25:1363-1375. Sobkowski, M., Stawinski, J., and Kraszewski, A. 2006b. A proposal for a convenient notation for P-chiral nucleotide analogues. Part 3. Compounds with one nucleoside residue and non-nucleosidic derivatives. Nucleosides Nucleotides Nucleic Acids 25:1377-1389.
Contributed by Michal Sobkowski and Adam Kraszewski Institute of Bioorganic Chemistry Poznan, Poland Jacek Stawinski Stockholm University Stockholm, Sweden and Institute of Bioorganic Chemistry Poznan, Poland
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LABORATORY STOCK SOLUTIONS AND EQUIPMENT
APPENDIX 2
Common Buffers and Stock Solutions
APPENDIX 2A
This section describes the preparation of buffers and reagents used in the manipulation of nucleic acids. For preparation of acid and base stock solutions, see Tables A.2A.1 and A.2A.2 as well as individual recipes. GENERAL GUIDELINES When preparing solutions, use deionized, distilled water and (for most applications) reagents of the highest grade available. Sterilization is recommended for most applications and is generally accomplished by autoclaving. Materials with components that are volatile, altered or damaged by heat, or whose pH or concentration are critical should be sterilized by filtration through a 0.22-µm filter. In many cases such components are added from concentrated stocks after the solution has been autoclaved. Where specialized sterilization methods are required, this is indicated in the individual recipes. CAUTION: It is important to follow laboratory safety guidelines and heed manufacturers’ precautions when working with hazardous chemicals; consult institutional safety officers and appropriate references for further details. STORAGE Most simple stock solutions can be stored indefinitely at room temperature if reasonable care is exercised to keep them sterile; where more rigorous conditions are required, this is indicated in the individual recipes. Table A.2A.1
Molarities and Specific Gravities of Concentrated Acids and Bases a
Acid/base Acids Acetic acid (glacial) Formic acid Hydrochloric acid Nitric acid Perchloric acid Phosphoric acid Sulfuric acid Bases Ammonium hydroxide Potassium hydroxide Potassium hydroxide Sodium hydroxide
Molecular weight
% by weight
Molarity (approx.)
1 M solution (ml/liter)
Specific gravity
60.05
99.6
17.4
57.5
1.05
46.03
98.00 98.07
90 98 36 70 60 72 85 98
23.6 25.9 11.6 15.7 9.2 12.2 14.7 18.3
42.4 38.5 85.9 63.7 108.8 82.1 67.8 54.5
1.205 1.22 1.18 1.42 1.54 1.70 1.70 1.835
35.0
28
14.8
67.6
0.90
56.11 56.11 40.0
45 50 50
11.6 13.4 19.1
82.2 74.6 52.4
1.447 1.51 1.53
36.46 63.01 100.46
aCAUTION: Handle strong acids and bases carefully.
Current Protocols in Nucleic Acid Chemistry (2000) A.2A.1-A.2A.12 Copyright © 2000 by John Wiley & Sons, Inc.
Laboratory Stock Solutions and Equipment
A.2A.1
SPECIAL CONSIDERATIONS FOR WORKING WITH RNA RNA is susceptible to degradation by ribonucleases, which are ubiquitous, very stable, and generally require no cofactors to function. Therefore, it is very important when working with RNA to take precautions against RNase contamination. 1. Treat all water and salt solutions except those containing Tris with DEPC (diethylpyrocarbonate; see recipe below), which inactivates ribonucleases by covalent modification. 2. If possible, make separate stock solutions to use for working with RNA and keep separate to ensure that “dirty” pipets do not come in contact with them. 3. Bake glassware 4 hr at 150°C. Rinse plasticware in chloroform or use directly out of the package (when it is generally free from contamination). Autoclaving will not fully inactivate many RNases. 4. Wear clean disposable gloves that have not been worn in any potentially RNase-contaminated areas. SPECIAL CONSIDERATIONS FOR PCR EXPERIMENTS Because the polymerase chain reaction (PCR) is designed to detect very small amounts of DNA, only a few molecules of contaminating DNA will produce unwanted amplification products. Ideally, PCR should not be carried out in the same room where large quantities of DNA are handled. Even where such spatial separation is not practical, the following housekeeping procedures will help avoid contamination with extraneous DNA (H.D. Kay, pers. comm.). 1. Keep laboratory surfaces clean by swabbing with 5% to 10% chlorine bleach. Put fresh absorbent paper bench protectors on bench before beginning PCR. 2. Wear disposable gloves and change them frequently while setting up PCRs. 3. Use only sterile disposable plasticware. 4. Keep a separate set of pipetting devices for setting up PCRs. If possible, use these instruments only with cotton-plugged tips to minimize transfer of DNA by aerosol. A separate microcentrifuge for PCR work is also desirable. 5. Whenever possible, set up PCRs in a laminar-flow hood or Class II biological safety cabinet to help prevent contamination by airborne DNA particles. A UV light within the hood or cabinet will help inactivate contaminating DNA. 6. Handle microcentrifuge tubes aseptically. Do not touch the interior of the hinged cap; if this happens, discard the tube. Microcentrifuge tubes briefly before opening to pellet drops around the cap and help keep reagents and reaction mixtures away from potentially contaminating fingers. Have only one tube open at a time, and open each tube away from the remaining tubes. Hand-held microcentrifuge tube openers (e.g., USA/Scientific Plastics) are available to facilitate aseptic technique. 7. Include negative controls (i.e., no primer and no template) in all PCRs. SELECTION OF BUFFERS
Common Buffers and Stock Solutions
Table A.2A.2 reports pKa values for some common buffers. Note that polybasic buffers, such as phosphoric acid and citric acid, have more than one useful pKa value. When choosing a buffer, select a buffer material with a pKa close to the desired working pH (at the desired concentration and temperature for use). In general, effective buffers have a range of approximately 2 pH units centered about the pKa value. Ideally the dissociation
A.2A.2 Current Protocols in Nucleic Acid Chemistry
constant—and therefore the pH—should not shift with a change in concentration or temperature. If the shift is small, as for MES and HEPES, then a concentrated stock solution can be prepared and diluted without adjustment to the pH. Buffers containing phosphate or citrate, however, show a significant shift in pH with concentration change, and Tris buffers show a large change in pH with temperature. For convenience, concentrated stock solutions of these buffers can still be used, provided that a pH adjustment is made after any temperature and concentration adjustments. All adjustments to pH should be made using the appropriate base—usually NaOH or KOH, depending on the corresponding free counterion. Tetramethylammonium hydroxide can be used to prepare buffers without a mineral cation. Many common buffers are supplied both as a free acid or base and as the corresponding salt. By mixing precalculated amounts of each, a series of buffers with varying pH values can conveniently be prepared. Table A.2A.2
pKa Values and Molecular Weights for Some Common Biological Buffersa
Name
Chemical formula or IUPAC name
pKa
Useful pH MW range (g/mol)
Phosphoric acid Citric acidb Formic acid Succinic acid Citric acidb Acetic acid Citric acidb Succinic acid MES Bis-Tris
H3PO4 C6H8O7 (H3Cit) HCOOH C4H6O4 C6H7O7− (H2Cit−) CH3COOH C6H6O7− (HCit2−) C4H5O4− 2-(N-Morpholino]ethanesulfonic acid bis(2-Hydroxyethyl)iminotris (hydroxymethyl)methane N-(2-Acetamido)-2-iminodiacetic acid Piperazine-N,N′-bis(2-ethanesulfonic acid) N-(Carbamoylmethyl)-2-aminoethanesulfonic acid 1,3-Diaza-2,4-cyclopentadiene C7H12O4 3-(N-Morpholino)propanesulfonic acid NaH2PO4
2.12 (pKa1) 3.06 (pKa1) 3.75 4.19 (pKa1) 4.74 (pKa2) 4.75 5.40 (pKa3) 5.57 (pKa2) 6.15 6.50
— — — — — — — — 5.5-6.7 5.8-7.2
98.00 192.1 46.03 118.1
6.60
6.0-7.2
190.2
6.80
6.1-7.5
302.4
6.80
6.1-7.5
182.2
7.00 7.20 7.20
— — 6.5-7.9
68.08 160.2 209.3
7.21 (pKa2)
—
120.0
ADA PIPES ACES Imidazole Diethylmalonic acid MOPS Sodium phosphate, monobasic Potassium phosphate, monobasic TES HEPES HEPPSO Glycinamide HCl Tricine Glycylglycine Tris
60.05
195.2 209.2
KH2PO4
7.21 (pKa2)
N-tris(Hydroxymethyl)methyl-2aminoethanesulfonic acid N-(2-Hydroxyethyl)piperazine-N′(2-ethanesulfonic acid) N-(2-Hydroxyethyl)piperazine-N′(2-hydroxypropanesulfonic acid) C2H6N2O⋅HCl N-tris(Hydroxymethyl)methylglycine C4H8N2O3 Tris(hydroxymethyl)aminomethane
7.40
6.8-8.2
229.3
7.55
6.8-8.2
238.3
7.80
7.1-8.5
268.3
8.10 8.15 8.20 8.30
7.4-8.8 7.4-8.8 7.5-8.9 7.0-9.0
110.6 179.2 132.1 121.1
136.1
continued
A.2A.3 Current Protocols in Nucleic Acid Chemistry
Table A.2A.2
pKa Values and Molecular Weights for Some Common Biological Buffers a, continued
Name
Chemical formula or IUPAC name
pKa
Useful pH MW range (g/mol)
Bicine Boric acid CHES
N,N-bis(2-Hydroxyethyl)glycine H3BO3 2-(N-Cyclohexylamino)ethanesulfonic acid 3-(Cyclohexylamino)-1-propanesulfonic acid Na2HPO4
8.35 9.24 9.50
7.6-9.0 — 8.6-10.0
163.2 61.83 207.3
10.40
9.7-11.1
221.3
12.32 (pKa3)
—
142.0
K2HPO4
12.32 (pKa3)
—
174.2
CAPS Sodium phosphate, dibasic Potassium phosphate, dibasic
aSome data reproduced from Buffers: A Guide for the Preparation and Use of Buffers in Biological Systems (Mohan, 1997) with permission of Calbiochem. bAvailable as a variety of salts, e.g., ammonium, lithium, sodium.
RECIPES Ammonium acetate, 10 M Dissolve 385.4 g ammonium acetate in 150 mL H2O Add H2O to 500 mL Sterilize by filtration BCIP, 5% (w/v) Dissolve 0.5 g 5-bromo-4-chloro-3-indolyl phosphate disodium salt (stored at −20°C) in 10 mL of 100% dimethylformamide (DMF). Store wrapped in aluminum foil up to 6 months at 4°C. The BCIP may not dissolve completely. Vortex the solution immediately before use and pipet with a wide-mouth pipet tip. Discard solution if it turns pinkish.
DEPC (diethylpyrocarbonate)-treated solutions Add 0.2 mL DEPC to 100 mL of the solution to be treated. Shake vigorously to dissolve the DEPC. Autoclave the solution to inactivate the remaining DEPC. CAUTION: Wear gloves and use a fume hood when using DEPC, as it is a suspected carcinogen. Many investigators keep the solutions they use for RNA work separate to ensure that “dirty” pipets do not go into them. One may also try to have separate working area, pipettors, etc. to avoid contamination. Do not treat solutions containing Tris with DEPC, as Tris inactivates the DEPC.
Common Buffers and Stock Solutions
dNTPs: dATP, dTTP, dCTP, and dGTP Concentrated stocks: Purchase deoxyribonucleoside triphosphates (dNTPs) from a commercial supplier (Pharmacia Biotech is recommended) either as ready-made 100 mM solutions, the preferred form for shipping and storage, or in lyophilized form. If purchased lyophilized, dissolve dNTPs in deionized H2O to an expected concentration of 30 mM, then adjust to pH 7.0 with 1 M NaOH (to prevent acid-catalyzed hydrolysis). Determine the actual concentration of each dNTP by UV spectrophotometry at 260 nm, using the following extinction coefficients: adenine, εIM260nm = 15,200; cytosine, εIM260nm = 7,050; guanosine, εIM260nm = 12,010; thymine, εIM260nm = 8,400. continued
A.2A.4 Current Protocols in Nucleic Acid Chemistry
Working solutions: Prepare working solutions of desired concentration (commonly 2 mM) for each dNTP by diluting concentrated stocks appropriately. Remember that the molarity of the 3dNTP and 4dNTP mixes refers to the concentration of each precursor present in the solution. 4dNTP mixes: For use in various molecular biology applications, prepare mixed dNTP solutions containing equimolar amounts of all four DNA precursors; e.g.: 2 mM 4dNTP mix: 2 mM each dATP, dTTP, dCTP, dGTP 1.25 mM 4dNTP mix: 1.25 mM each of dATP, dTTP, dCTP, dGTP. 3dNTP mixes: For use in radioactive labeling procedures, prepare similar stocks lacking one particular dNTP but containing equimolar amounts of the remaining three precursors; e.g.: 2 mM 3dNTP mix (minus dATP): 2 mM each of dTTP, dCTP, dGTP. Store dNTPs and dNTP mixtures as aliquots at −20°C (stable for ≤1 year). DTT (dithiothreitol), 1 M Dissolve 1.55 g DTT in 10 mL water and filter sterilize. Store in aliquots at −20°C. Do not autoclave to sterilize.
EDTA (ethylenediaminetetraacetic acid), 0.5 M (pH 8.0) Dissolve 186.1 g disodium EDTA dihydrate in 700 mL water. Adjust pH to 8.0 with 10 M NaOH (∼50 mL; add slowly). Add water to 1 L and filter sterilize. Begin titrating before the sample is completely dissolved. EDTA, even in the disodium salt form, is difficult to dissolve at this concentration unless the pH is increased to between 7 and 8. Heating the solution may also help to dissolve EDTA.
Ethidium bromide solution Concentrated stock (10 mg/mL): Dissolve 0.2 g ethidium bromide in 20 mL H2O. Mix well and store at 4°C in dark or in a foil-wrapped bottle. Do not sterilize. Working solution: Dilute stock to 0.5 µg/mL or other desired concentration in electrophoresis buffer (e.g., 1× TBE or TAE) or water. To use: Ethidium bromide working solution is used to stain agarose gels to permit visualization of nucleic acids under UV light. Gels should be placed in a glass dish containing sufficient working solution to cover them and shaken gently or allowed to stand for 10 to 30 min. If necessary, gels can be destained by shaking in electrophoresis buffer or water for an equal length of time to reduce background fluorescence and facilitate visualization of small quantities of DNA. Alternatively, a gel can be run directly in ethidium bromide by using working solution (made with electrophoresis buffer) as the solvent and running buffer for the gel. CAUTION: Ethidium bromide is a toxic and powerful mutagen. Gloves should be worn when working with solution or gel and a mask should be worn when weighing out solid. Keep separate solid and liquid waste containers for disposal of ethidium bromide–contaminated material.
Formamide loading buffer, 2× Prepare in deionized formamide: 0.05% (w/v) bromphenol blue 0.05% (w/v) xylene cyanol FF 20 mM EDTA Do not sterilize Store at −20°C Laboratory Stock Solutions and Equipment
A.2A.5 Current Protocols in Nucleic Acid Chemistry
Gel loading buffer, 6× 0.25% (w/v) bromphenol blue 0.25% (w/v) xylene cyanol FF 40% (w/v) sucrose or 15% (w/v) Ficoll 400 or 30% (v/v) glycerol Store at 4°C (room temperature if Ficoll is used) This buffer does not need to be sterilized. Sucrose, Ficoll 400, and glycerol are essentially interchangeable in this recipe. Other concentrations (e.g., 10×) can be prepared if more convenient.
HCl, 1 M Mix in the following order: 913.8 mL H2O 86.2 mL concentrated HCl (Table A.2A.1) HEPES-buffered saline, 2× 90 mL H2O 1.6 g NaCl (0.27 M) 74.6 mg KCl (10 mM) 21.3 mg Na2HPO4 (1.5 mM) 0.18 g glucose (10 mM) 1.07 g HEPES (45 mM) Adjust pH to desired value (see Table A.2A.2) with 0.5 N NaOH Adjust volume to 100 mL with H2O Filter sterilize Store in aliquots indefinitely at −20°C KCl, 1 M 74.6 g KCl H2O to 1 L Kinase buffer, 10× 700 mM Tris⋅Cl, pH 7.4 (see recipe below) 100 mM MgCl2 (see recipe below) Store in aliquots indefinitely at room temperature Discard solution if a precipitate forms. A 10× kinase buffer is sold with commercially available kinase enzymes, but usually contains DTT. This buffer may be used as a substitute if the reaction must be done under nonreducing conditions.
MgCl2 , 1 M 20.3 g MgCl2⋅6H2O H2O to 100 mL MgCl2 is extremely hygroscopic. Do not store opened bottles for long periods of time.
MgSO4 , 1 M 24.6 g MgSO4⋅7H2O H2O to 100 mL NaCl, 5 M 292 g NaCl H2O to 1 L
Common Buffers and Stock Solutions
NaOH, 10 M Dissolve 400 g NaOH in 450 mL H2O Add H2O to 1 L
A.2A.6 Current Protocols in Nucleic Acid Chemistry
NBT (nitroblue tetrazolium chloride), 5% (w/v) Dissolve 0.5 g NBT in 10 mL of 70% dimethylformamide (DMF). Store wrapped in aluminum foil up to 1 year at 4°C. PBS (phosphate-buffered saline) 8.00 g NaCl (0.137 M) 0.20 g KCl (2.7 mM) 0.24 g KH2PO4 (1.4 mM) 1.44 g Na2HPO4 (0.01 M) H2O to 800 mL Adjust pH as desired (usually to pH 7.4 with 1 M HCl) Add H2O to 1 L Prepackaged PBS (pH 7.4), which is reconstituted by adding water, is commercially available from Sigma. This is very convenient if used in large quantities.
PCR amplification buffer, 10× 500 mM KCl 100 mM Tris⋅Cl, pH 8.3 (see recipe below) x mM MgCl2 0.1% (w/v) gelatin Store in aliquots at −20°C This solution can be sterilized by autoclaving. Alternatively, it can be made from sterile water and stock solutions, and the sterilization omitted. 15 mM MgCl2 is the concentration (x) used for most PCR reactions. However, the optimal concentration depends on the sequence and primer of interest and may have to be determined experimentally.
Phenol, buffered 8-Hydroxyquinoline Liquefied phenol, redistilled 50 mM Tris base (unadjusted pH ∼10.5) 50 mM Tris⋅Cl, pH 8.0 (see recipe below) TE buffer, pH 8.0 (see recipe below) Add 0.5 g of 8-hydroxyquinoline to a 2-L glass beaker containing a stir bar. Gently pour in 500 mL of liquefied phenol or melted crystals of redistilled phenol (melted in a water bath at 65°C). The phenol will turn yellow due to the 8-hydroxyquinoline, which is added as an antioxidant. Add 500 mL of 50 mM Tris base. Cover the beaker with aluminum foil and stir 10 min at low speed with magnetic stirrer at room temperature. Let phases separate at room temperature. Gently decant the top (aqueous) phase into a suitable waste receptacle. (Aspiration also works well to remove the majority of the top phase.) Remove what cannot be decanted with a 25-mL glass pipet and a suction bulb. Add 500 mL of 50 mM Tris⋅Cl, pH 8.0. Repeat equilibration with 500 mL of 50 mM Tris⋅Cl, pH 8.0, twice. The pH of the phenol phase can be checked with indicator paper and should be 8.0. If it is not, repeat equilibration until this pH is obtained. DNA will partition into organic phase at acidic pH; therefore pH must be >7.8. Add 250 mL of 50 mM Tris⋅Cl, pH 8.0, or TE buffer, pH 8.0, and store at 4°C in brown glass bottles or clear glass bottles wrapped in aluminum foil. Phenol prepared with 8-hydroxyquinoline as an antioxidant can be stored ≤2 months at 4°C. Phenol must be redistilled before use, because oxidation products of phenol can damage and introduce breaks into nucleic acid chains. Redistilled phenol is commercially available. Regardless of the source, the phenol must be buffered before use. If liquefied phenol is yellowish or pink when purchased, discard. continued
Laboratory Stock Solutions and Equipment
A.2A.7 Current Protocols in Nucleic Acid Chemistry
CAUTION: Phenol can cause severe burns to skin and damage clothing. If it contacts skin, wash with soap and water. Do NOT use ethanol. Gloves, safety glasses, and a lab coat should be worn whenever working with phenol, and all manipulations should be carried out in a fume hood. A glass receptacle should be available exclusively for disposing of used phenol and chloroform. Filter tips (available from Rainin Instruments) should be used when pipetting solutions to protect pipettors.
Phenol/chloroform/isoamyl alcohol, 25:24:1 (v/v/v) Mix 25 vol buffered phenol (bottom phase; see recipe above) with 24 vol chloroform and 1 vol isoamyl alcohol. Store in brown glass bottle or in clear glass bottle wrapped in aluminum foil ≤2 months at 4°C. To use: Mix 1:1 (v/v) with DNA sample solution to form an emulsion, then microcentrifuge 15 sec at maximum speed, room temperature. Assuming that the salt concentration in the DNA is <0.5 M, the DNA-containing aqueous phase will be on top; transfer this to a new tube, being careful not to disturb the interface. Reextract if necessary, and then extract a final time with 1 vol chloroform to remove residual phenol. Alternatively, isoamyl alcohol may be omitted and a 1:1 mix of phenol and chloroform used instead. The isoamyl alcohol enhances the separation of the phases and reduces foaming, however. Fuller details of phenol extraction are given in CPMB UNIT 2.1.
PMSF (phenylmethanesulfonyl fluoride), 10 mM Dissolve 1.74 mg/mL PMSF in isopropyl alcohol. Store aliquots indefinitely at −20°C. CAUTION: PMSF is extremely toxic to the mucous membranes of the respiratory tract, the eyes, and the skin. It may be fatal if inhaled, swallowed, or absorbed through the skin. Gloves, safety glasses, and a lab coat should be worn when working with PMSF. Wash any contacted areas immediately with large volumes of water and discard contaminated clothing. PMSF stock solutions can be made up to 17.4 mg/mL (100 mM) if necessary. Note that PMSF is inactivated in aqueous solutions; the rate of inactivation increases with both temperature and pH. Methods for disposal and detection of PMSF can be found in Lunn (2000).
Potassium acetate buffer, 0.1 M Solution A: 11.55 mL glacial acetic acid per L (0.2 M) in water. Solution B: 19.6 g potassium acetate (KC2H3O2) per L (0.2 M) in water.
continued
Table A.2A.3 Preparation of 0.1 M Sodium and Potassium Acetate Buffersa
Common Buffers and Stock Solutions
Desired pH
Solution A (ml)
Solution B (ml)
3.6 3.8 4.0 4.2 4.4 4.6 4.8 5.0 5.2 5.4 5.6
46.3 44.0 41.0 36.8 30.5 25.5 20.0 14.8 10.5 8.8 4.8
3.7 6.0 9.0 13.2 19.5 24.5 30.0 35.2 39.5 41.2 45.2
aAdapted by permission from CRC (1975).
A.2A.8 Current Protocols in Nucleic Acid Chemistry
Table A.2A.4 Preparation of 0.1 M Sodium and Potassium Phosphate Buffersa
Desired pH 5.7 5.8 5.9 6.0 6.1 6.2 6.3 6.4 6.5 6.6 6.7 6.8
Solution A (ml) 93.5 92.0 90.0 87.7 85.0 81.5 77.5 73.5 68.5 62.5 56.5 51.0
Solution B (ml) 6.5 8.0 10.0 12.3 15.0 18.5 22.5 26.5 31.5 37.5 43.5 49.0
Desired pH 6.9 7.0 7.1 7.2 7.3 7.4 7.5 7.6 7.7 7.8 7.9 8.0
Solution A (ml) 45.0 39.0 33.0 28.0 23.0 19.0 16.0 13.0 10.5 8.5 7.0 5.3
Solution B (ml) 55.0 61.0 67.0 72.0 77.0 81.0 84.0 87.0 90.5 91.5 93.0 94.7
aAdapted by permission from CRC (1975).
Potassium acetate buffer, 0.1 M Solution A: 11.55 mL glacial acetic acid per L (0.2 M) in water. Solution B: 19.6 g potassium acetate (KC2H3O2) per L (0.2 M) in water. Referring to Table A.2A.3 for desired pH, mix the indicated volumes of solutions A and B, then dilute with water to 100 mL. Filter sterilize if necessary. Store up to 3 months at room temperature. This may be made as a 5- or 10-fold concentrate by scaling up the amount of sodium acetate in the same volume. Acetate buffers show concentration-dependent pH changes, so check the pH by diluting an aliquot of concentrate to the final concentration. To prepare buffers with pH intermediate between the points listed in Table A.2A.3, prepare closest higher pH, then titrate with solution A.
Potassium phosphate buffer, 0.1 M Solution A: 27.2 g KH2PO4 per L (0.2 M final) in water. Solution B: 34.8 g K2HPO4 per L (0.2 M final) in water. Referring to Table A.2A.4 for desired pH, mix the indicated volumes of solutions A and B, then dilute with water to 200 mL. Filter sterilize if necessary. Store up to 3 months at room temperature. This buffer may be made as a 5- or 10-fold concentrate simply by scaling up the amount of potassium phosphate in the same final volume. Phosphate buffers show concentration-dependent changes in pH, so check the pH of the concentrate by diluting an aliquot to the final concentration. To prepare buffers with pH intermediate between the points listed in Table A.2A.4, prepare closest higher pH, then titrate with solution A.
SDS, 20% (w/v) Dissolve 20 g SDS (sodium dodecyl sulfate or sodium lauryl sulfate) in water to 100 mL total volume with stirring. Filter sterilize using a 0.45-µm filter. It may be necessary to heat the solution slightly to fully dissolve the powder.
SDS sample buffer See Table A.2A.5. Laboratory Stock Solutions and Equipment
A.2A.9 Current Protocols in Nucleic Acid Chemistry
Supplement 9
Table A.2A.5
Preparation of SDS Sample Buffera
Ingredient
2×
4×
Final conc. in 1× buffer
0.5 M Tris⋅Cl, pH 6.8b SDS Glycerol Bromphenol blue 2-Mercaptoethanolc,d H2O
2.5 mL 0.4 g 2.0 mL 20 mg 400 µL to 10 mL
5.0 mL 0.8 g 4.0 mL 40 mg 800 µL to 10 mL
62.5 mM 2% (w/v) 10% (v/v) 0.1% (w/v) ∼300 mM —
aCan be divided into aliquots and stored several months at room temperature (after addition of 2-ME); when solution
turns yellow/green, discard. bSee recipe below. cAlternatively, dithiothreitol (DTT), at a final concentration of 100 mM, can be substituted for 2-mercaptoethanol (2-ME). dAdd just before use.
Silanized glassware For smaller items: In a well-vented fume hood, place glassware or plasticware (e.g., tubes, tips) in a dedicated vacuum desiccator with an evaporating dish containing 1 mL dichlorodimethylsilane. Apply vacuum with an aspirator and allow ∼50% of the liquid to evaporate (several minutes). Turn off aspirator and allow items to remain under vacuum for 30 min. Remove the lid and allow fumes to vent into the hood for ∼30 min. If desired, autoclave silanized items. Do not leave the desiccator attached to the vacuum pump. This will suck away the silane, minimizing deposition and damaging the pump.
For larger items: Silanize items that do not fit in a desiccator by briefly rinsing with or soaking in a solution of ∼5% dichlorodimethylsilane in a volatile organic solvent (e.g., chloroform, heptane). Remove organic solvent by evaporation, allowing deposition of dichlorodimethylsilane. This approach is particularly useful for treating glass plates for denaturing polyacrylamide sequencing gels (APPENDIX 3B). Treatment of glassware, plasticware, or equipment with dichlorodimethylsilane introduces a short polymer of dimethylsiloxane onto its surface. Polydimethylsiloxane is silicone oil. Autoclaving or rinsing with water removes the reactive chlorosilane end of the dimethylsiloxane polymer generated by dichlorodimethylsilane. CAUTION:Dichlorodimethylsilane vapors are toxic and highly flammable. Always perform in a fume hood.
Sodium acetate, 3 M Dissolve 408 g sodium acetate trihydrate (NaC2H3O2⋅3H2O) in 800 mL H2O Adjust pH to 4.8, 5.0, or 5.2 (as desired) with 3 M acetic acid (see Table A.2A.1) Add H2O to 1 L Filter sterilize Sodium acetate buffer, 0.1 M Solution A: 11.55 mL glacial acetic acid per L (0.2 M) in water. Solution B: 27.2 g sodium acetate (NaC2H3O2⋅3H2O) per L (0.2 M) in water. Referring to Table A.2A.3 for desired pH, mix the indicated volumes of solutions A and B, then dilute with water to 100 mL. Filter sterilize if necessary. Store up to 3 months at room temperature. This may be made as a 5- or 10-fold concentrate by scaling up the amount of sodium acetate in the same volume. Acetate buffers show concentration-dependent pH changes, so check the pH by diluting an aliquot of concentrate to the final concentration. Common Buffers and Stock Solutions
To prepare buffers with pH intermediate between the points listed in Table A.2A.3, prepare closest higher pH, then titrate with solution A.
A.2A.10 Supplement 9
Current Protocols in Nucleic Acid Chemistry
Sodium phosphate buffer, 0.1 M Solution A: 27.6 g NaH2PO4⋅H2O per L (0.2 M final) in water. Solution B: 53.65 g Na2HPO4⋅7H2O per L (0.2 M) in water. Referring to Table A.2A.4 for desired pH, mix the indicated volumes of solutions A and B, then dilute with water to 200 mL. Filter sterilize if necessary. Store up to 3 months at room temperature. This buffer may be made as a 5- or 10-fold concentrate by scaling up the amount of sodium phosphate in the same final volume. Phosphate buffers show concentration-dependent changes in pH, so check the pH by diluting an aliquot of the concentrate to the final concentration. To prepare buffers with pH intermediate between the points listed in Table A.2A.4, prepare closest higher pH, then titrate with solution A.
SSC (sodium chloride/sodium citrate), 20× 3 M NaCl (175 g/L) 0.3 M Na3citrate⋅2H2O (88 g/L) Adjust pH to 7.0 with 1 M HCl SSPE (sodium chloride/sodium phosphate/EDTA), 20× 800 mL H2O 175 g NaCl (3 M) 7.4 g EDTA (20 mM) 24.0 g NaH2PO4 (0.20 mM) Adjust pH with 10 N NaOH (∼6.5 mL for pH 7.4) Adjust volume to 1 L with H2O TAE (Tris/acetate/EDTA) electrophoresis buffer, 10× 24.2 Tris base 5.71 mL glacial acetic acid 3.72 g Na2EDTA⋅2H2O H2O to 1 L TBE (Tris/borate/EDTA) electrophoresis buffer, 10× 108 g Tris base (890 mM) 55 g boric acid (890 mM) 960 mL H2O 40 mL 0.5 M EDTA, pH 8.0 (20 mM final; see recipe above) 10× and 5× TBE tend to precipitate over time. If convenient, dilute to 2× or 1× immediately, or stir continuously.
TBS (Tris-buffered saline) 800 mL H2O 8.00 g NaCl (0.137 M) 0.2 g KCl (2.7 mM) 3.0 g Tris base (24.8 mM) Adjust pH as desired (usually to pH 8) with 1 M HCl Adjust volume to 1 L with H2O Prepackaged TBS (pH 8.0), which is reconstituted by adding water, is commercially available from Sigma. This is very convenient if used in large quantities.
Laboratory Stock Solutions and Equipment
A.2A.11 Current Protocols in Nucleic Acid Chemistry
Supplement 9
TCA (trichloroacetic acid), 100% (w/v) 500 g TCA 227 mL H2O TE (Tris/EDTA) buffer 10 mM Tris⋅Cl, pH 7.4, 7.5, or 8.0 (or other pH; see recipe below) 1 mM EDTA, pH 8.0 (see recipe above) Tris⋅Cl, 1 M Dissolve 121 g Tris base in 800 mL H2O Adjust to desired pH with concentrated HCl Adjust volume to 1 L with H2O Filter sterilize if necessary Store up to 6 months at 4°C or room temperature Approximately 70 mL HCl is needed to achieve a pH 7.4 solution, and ∼42 mL for a solution that is pH 8.0. IMPORTANT NOTE: The pH of Tris buffers changes significantly with temperature, decreasing approximately 0.028 pH units per 1°C. Tris-buffered solutions should be adjusted to the desired pH at the temperature at which they will be used. Because the pKa of Tris is 8.08, Tris should not be used as a buffer below pH ∼7.2 or above pH ∼9.0. Always use high-quality Tris (lower-quality Tris can be recognized by its yellow appearance when dissolved).
Urea loading buffer, 2× 5 mg bromphenol blue (0.05% w/v) 5 mg (w/v) xylene cyanol FF (0.05% w/v) 4.8 g urea (8 M) 186 mg EDTA (50 mM) H2O to 10 mL Do not sterilize Store up to 6 months at room temperature ACKNOWLEDGEMENT The editorial board thanks Joanne Cleary (University of Michigan, Ann Arbor) for revising this appendix and adding recipes, including the general guidelines for buffer selection. LITERATURE CITED Chemical Rubber Company. 1975. CRC Handbook of Biochemistry and Molecular Biology, Physical and Chemical Data, 3d ed., Vol. 1. CRC Press, Boca Raton, Fla. Lunn, G. 2000. Laboratory Safety. In Current Protocols in Protein Science (Coligan, J.E., Dunn, B., Ploegh, H.L., Speicher, D.W., and Wingfield, P.T., eds.) pp. A.2A.1-A.2A.26. John Wiley & Sons, New York. Mohan, C. (ed.). 1997. Buffers: A Guide for the Preparation and Use of Buffers in Biological Systems. Calbiochem, San Diego, Calif.
Common Buffers and Stock Solutions
A.2A.12 Supplement 9
Current Protocols in Nucleic Acid Chemistry
COMMONLY USED TECHNIQUES
APPENDIX 3
References to Commonly Used Techniques
APPENDIX 3A
Many of the protocols in this manual assume a knowledge of basic (chemical and biochemical) techniques. Although these are outside the scope of Current Protocols in Nucleic Acid Chemistry (CPNC), a number of basic methods can be found in this volume — either as support protocols or as part of longer protocols. While tailored to the particular goals of the units in which they appear, such protocols can be adapted by the trained researcher to suit the needs of a particular laboratory. If additional explanation or details for molecular techniques are required, the reader is advised to consult Current Protocols in Molecular Biology (CPMB; Ausubel et al., 2000). To facilitate this cross-referencing, we have cited relevant CPMB units throughout the book. Alternatively, protocols from other published laboratory manuals can be used. For basic chemical methods, any number of college laboratory textbooks can be consulted. Table A.3A.1 lists some commonly used techniques described in the book; if a protocol is not listed here, check the index. Table A.3A.1
Locations of Techniques Used in CPNC
Technique
CPNC reference
Denaturing polyacrylamide gels DNA, genomic fragments, sonication DNA, genomic fragments, size fractionation DNA strand scission by piperidine DNA, ethanol precipitation DNA, phenol/chloroform extraction DNA, 5′ labeling DNA quantitation, spectrophotometrically Mutagenic PCR Nitrogen atmosphere, setup Oligonucleotide synthesis, general Oligonucleotides, deprotection Oligonucleotide extinction coefficient Oligomers, primer extension Oligonucleotide purification cation-exchange resin spin column C18 reversed-phase cartridge molecular-weight-cutoff filter electroelution from gel slice elution from gel slice Primer design RNA, 3′ labeling RNA, 5′ labeling RNA renaturation RNAse-free water, testing RNAse-free reagents RNA purification, NAP-25 Sephadex columns Transcription, in vitro Trityl assay Thin-layer chromatography
APPENDIX 3B UNIT 8.1 UNIT 8.1 UNIT 6.4 UNIT 6.1 APPENDIX 2A UNITS 6.1 & 9.2 UNIT 5.2 UNIT 9.4 UNITS 1.1 & 1.3 APPENDIX 3C APPENDIX 3C UNIT 7.3 UNIT 6.1
Current Protocols in Nucleic Acid Chemistry (2000) A.3A.1-A.3A.2 Copyright © 2000 by John Wiley & Sons, Inc.
UNIT 10.1 UNIT 10.1 UNIT 10.1 UNIT 10.1 UNIT 5.4 UNITS 5.2, 6.3, & 9.2 UNIT 9.2 UNITS 6.1 & 6.3 UNITS 6.1 & 6.3 UNIT 6.3 UNIT 6.1 APPENDIX 2A UNIT 5.2 UNIT 9.4 APPENDIX 3C UNIT 2.4
Commonly Used Techniques
A.3A.1
LITERATURE CITED Ausubel, F.A. Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (eds.) 2000. Current Protocols in Molecular Biology. John Wiley & Sons, Inc. New York.
References to Commonly Used Techniques
A.3A.2 Current Protocols in Nucleic Acid Chemistry
Denaturing Polyacrylamide Gel Electrophoresis
APPENDIX 3B
Thin polyacrylamide gels that contain a high concentration of urea as a denaturant are capable of resolving short (<500 nucleotides) single-stranded fragments of DNA or RNA that differ in length by as little as one nucleotide. Such gels are uniquely suited for nucleic acid sequence analysis, which is required for all of the footprinting protocols in Chapter 6. Thicker gels are often used to purify oligonucleotides. The following protocol describes the pouring, running, and processing of a typical “sequencing” gel which is 40-cm long with a uniform thickness of 0.4 mm, containing 7 M urea and 4% to 8% acrylamide. RNA samples are sometimes run on gels prepared with 8 M urea. A more detailed protocol with variations and extensive troubleshooting information is provided in CPMB UNIT 7.6. Some guidelines for pouring and running preparative gels are given in annotations to some steps. POURING, RUNNING, AND PROCESSING DENATURING POLYACRYLAMIDE GELS
BASIC PROTOCOL
Materials 70% ethanol or isopropanol in squirt bottle 5% (v/v) dimethyldichlorosilane (Sigma) in CHCl3 Denaturing acrylamide gel solution (see recipe) TEMED 10% (w/v) ammonium persulfate (make fresh weekly and store at 4°C) 1× TBE electrophoresis buffer, pH 8.3 to 8.9 (APPENDIX 2A) Samples for electrophoresis containing formamide and marker dyes 30 × 40–cm front and back gel plates 0.2- to 0.4-mm uniform-thickness spacers Large book-binder clamps 60-mL syringe 0.2- to 0.4-mm shark’s-tooth or preformed-well combs Sequencing gel electrophoresis apparatus Pasteur pipet or Beral thin stem (Beral Enterprises) Power supply with leads 95°C heating block or water bath 46 × 57–cm gel blotting paper (e.g., Whatman 3MM) Kodak XAR-5 X-ray film NOTE: Many companies provide equipment needed for sequencing experiments; a list of suppliers is provided in CPMB Table 7.6.1. Assemble the gel sandwiches 1. Meticulously wash front and back 30 × 40–cm gel plates with soap and water. Rinse well with deionized water and dry. Wet plates with 70% ethanol or isopropanol in a squirt bottle and wipe dry with Kimwipe or other lint-free paper towel. A typical preparative gel uses 20 × 16–cm plates.
2. Apply a film of 5% dimethyldichlorosilane in CHCl3 to one side of each plate by wetting a Kimwipe with the solution and wiping carefully. After the film dries, wipe plate with 70% ethanol or isopropanol and dry with a Kimwipe. Check plates for dust and other particulates. Commonly Used Techniques Contributed by Lisa M. Albright and Barton E. Slatko Current Protocols in Nucleic Acid Chemistry (2000) A.3B.1-A.3B.5 Copyright © 2000 by John Wiley & Sons, Inc.
A.3B.1
3. Assemble gel plates according to manufacturer’s instructions, with the silanized surfaces facing inward. Use 0.2- to 0.4-mm uniform-thickness spacers and large book-binder clamps, making certain side and bottom spacers fit tightly together. Use thicker spacers (e.g., 1.6-mm) for preparative gels.
Prepare and pour the gel 4. Prepare 60 mL of desired denaturing acrylamide gel solution in a 100-mL beaker. (See Table A.3B.1 for appropriate acrylamide concentrations for resolving singlestranded DNAs. Consult Table A.3B.2 for preparative gels.) Thoroughly mix 60 µL TEMED, then 0.6 mL of 10% ammonium persulfate, into acrylamide solution immediately before pouring gel. To speed dissolution of urea, the gel mix can be heated before adding TEMED and ammonium persulfate; however, to prevent degradation of acrylamide, do not heat over 55°C. Allow to cool to room temperature (≤25°C) before adding the TEMED and ammonium persulfate to prevent polymerization while pouring the gel. If particulate matter remains, filter through a Whatman no. 1 filter paper in a funnel. To achieve slower polymerization, reduce amounts of TEMED and ammonium persulfate to 40 ì L and 0.4 mL, respectively.
5. Pour gel immediately. Gently pull acrylamide solution into a 60-mL syringe, avoiding bubbles. With short plate on top, raise upper edge of gel sandwich to 45° angle from the benchtop and slowly expel acrylamide between plates along one side. Adjust angle of plates so gel solution flows slowly down one side. 6. When solution reaches top of short plate, lower gel sandwich so that the top edge is ∼5 cm above benchtop. Place an empty disposable pipet-tip rack or stopper underneath the sandwich to maintain the low angle. Insert flat side of a 0.2- to 0.4-mm shark’s-tooth comb into the solution 2 to 3 mm below top of short plate, being very careful to avoid bubbles. Use book-binder clamps to pinch combs between plates so that no solidified gel forms between combs and plates. Layer extra acrylamide gel solution onto comb to ensure full coverage. Alternatively, insert teeth of preformed-well comb into gel solution and clamp as above. The comb should be the same thickness as the spacers. Rinse syringe with water to remove acrylamide.
Set up the electrophoresis apparatus 7. When gel polymerizes, remove bottom spacer or tape at bottom of gel sandwich. Remove extraneous polyacrylamide from around combs with razor blade. Clean spilled urea and acrylamide solution from outer plate surfaces with water. Remove shark’s-tooth comb gently from gel sandwich without stretching or tearing top of gel. Clean comb with water so it will be ready to be reinserted in step 10.
Table A.3B.1 Migration of Oligodeoxynucleotides (Bases) in “Sequencing” Denaturing Polyacrylamide Gels Relative to Dye Markers
Denaturing Polyacrylamide Gel Electrophoresis
Polyacrylamide
Bromphenol blue
Xylene cyanol
5% 6% 8% 10%
35 b 26 b 19 b 12 b
130 b 106 b 75 b 55 b
A.3B.2 Current Protocols in Nucleic Acid Chemistry
If preformed-well comb was used, take care to prevent tearing of polyacrylamide wells. This comb will not be reinserted.
8. Fill bottom reservoir of gel apparatus with 1× TBE buffer so that gel plates will be submerged 2 to 3 cm in buffer. Place gel sandwich in electrophoresis apparatus and clamp plates to support. Sweep out any air bubbles at bottom of gel by squirting buffer between plates using syringe with a bent 20-G needle.
9. Pour 1× TBE buffer into top reservoir to ∼3 cm above top of gel. Rinse top of gel with 1× TBE buffer using a Pasteur pipet or Beral thin stem. 10. Reinsert teeth of cleaned shark’s-tooth comb into gel sandwich with points just barely sticking into gel. Using a Pasteur pipet or Beral thin stem, rinse wells thoroughly with 1× TBE buffer to remove stray fragments of polyacrylamide. If a preformed-well comb is used, this step is omitted.
11. Preheat gel ∼30 min by setting power supply to 45 V/cm, 1700 V, 70 W constant power. Preparative gels are usually preheated and run at 20 to 40 V/cm, constant voltage.
Load and run the gel 12. Rinse wells with 1× TBE buffer just prior to loading gels, to remove urea that has leached into them. 13. Heat samples 2 min at 95°C in covered microcentrifuge tubes, then place on ice. Load 2 to 3 µL sample per well. Rinse sequencing pipet tip twice in lower reservoir after dispensing from each reaction tube. 14. Run gels at 45 to 70 W constant power. Maintain a gel temperature of ∼65°C. Observe migration of marker dyes (Table A.3B.2) to determine length of electrophoresis. Temperatures >65°C can result in cracked plates or smeared bands; too low a temperature can lead to incomplete denaturation. To ensure even conduction of the heat generated during electrophoresis, an aluminum plate (0.4 cm thick, 34 × 22–cm) can be clamped onto the front glass plate with the same book-binder clamps used to hold the gel sandwich to the apparatus. The aluminum plate must be positioned so that it does not touch any buffer during electrophoresis.
Table A.3B.2 Concentrations of Acrylamide Giving Optimum Resolution for Purification of DNA Fragments Using Denaturing PAGEa
Acrylamide (%) 30 20 10 8 6 5 4
Fragment sizes separated (bases) 2-8 8-25 25-35 35-45 45-70 70-300 100-500
Migration of bromphenol blue (bases)
Migration of xylene cyanol (bases)
6 8 12 19 26 35 ∼50
20 28 55 75 105 130 ∼230
aData, from Maniatis et al. (1975), are for single-stranded DNA; RNA will migrate slightly more slowly than DNA of the same sequence and length. Taken from CPMB UNIT 2.12.
Commonly Used Techniques
A.3B.3 Current Protocols in Nucleic Acid Chemistry
Preparative gels are usually run at 20 to 40 V/cm, constant voltage. Turn off power when the position of the tracking dye indicates that the oligonucleotide has migrated sufficiently for isolation. Proceed with isolation method of choice.
Process and dry the gel 15. Fill dry-ice traps attached to gel dryer (if required) and preheat dryer to 80°C. 16. After electrophoresis is complete, drain buffer from upper and lower reservoirs of apparatus and discard liquid as radioactive waste. 17. Remove gel sandwich from apparatus and place under cold running tap water until surfaces of both glass plates are cool. Lay sandwich flat on paper towels with short plate up. Remove excess liquid and remaining clamps or tape. Remove one side spacer and insert long metal spatula between glass plates where spacer had been. Pry plates apart by gently rocking spatula. The gel should stick to the bottom plate. If it sticks to the top plate, flip sandwich over. Slowly lift top plate from the side with inserted spatula, gradually increasing the angle until the top plate is completely separated from gel.
18. Once plates are separated, remove second side spacer and any extraneous bits of polyacrylamide around gel. 19. Hold two pieces of dry 46 × 57–cm blotting paper together as one piece. Beginning at one end of gel and working slowly towards the other, lay paper on top of gel. Take care to prevent air bubbles from forming between paper and gel. 20. Peel blotting paper up; gel should come off plate with it. Gradually curl paper and gel away from plate as it is being pulled away. 21. Place paper and gel on preheated gel dryer. Cover with plastic wrap. Remove any bubbles between plastic wrap and gel by gently rubbing covered surface of gel from middle toward edges with a Kimwipe. Dry gel thoroughly 20 min to 1 hr at 80°C. When gel is completely dry, the plastic will easily peel off without sticking.
22. Remove plastic wrap and place dried gel in X-ray cassette with Kodak XAR-5 film in direct contact with gel. Autoradiograph at room temperature. After sufficient exposure time (usually overnight), remove X-ray film and process.
Denaturing Polyacrylamide Gel Electrophoresis
A.3B.4 Current Protocols in Nucleic Acid Chemistry
REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.
Denaturing acrylamide gel solution Reagent Urea (ultrapure; g)a 38% acrylamide/2% bisacrylamide (mL) 10× TBE buffer (APPENDIX 2A; mL) H2O (mL) Total volume (mL)
Acrylamide concentration 4%
6%
8%
25.2 6.0 6.0 27 60
25.2 9.0 6.0 24 60
25.2 12.0 6.0 21 60
a7 M final concentration.
Filter solution through Whatman no. 1 filter paper. Store 2 to 4 weeks at 4 °C. CAUTION: Acrylamide and bisacrylamide are hazardous. Solutions of acrylamide deteriorate quickly, especially when exposed to light or left at room temperature. RNA samples are often run on sequencing gels containing 8 M rather than 7 M urea. In this case, use 28.8 g urea for 60 mL of gel solution.
LITERATURE CITED Maniatis, T., Jeffrey, A., and deSande, H.U. 1985. Chain length determination of small double- and single-stranded DNA molecules by polyacrylamide gel electrophoresis. Biochemistry 14:3787-3794.
Contributed by Lisa M. Albright Allison Park, Pennsylvania Barton E. Slatko New England Biolabs Beverly, Massachusetts
Commonly Used Techniques
A.3B.5 Current Protocols in Nucleic Acid Chemistry
Introduction to the Synthesis and Purification of Oligonucleotides
APPENDIX 3C
Avery’s realization that DNA carries the genetic information lead chemists on a 40-year search that has culminated in efficient, automated oligonucleotide synthesis on solidphase supports. Modern nucleic acid synthesizers utilize phosphite triester chemistries that employ stable phosphoramidite monomers to build the growing polymer (see UNIT 3.3). These robust reactions allow both chemists and molecular biologists to easily generate specific ribo- and deoxyribo oligonucleotides with a variety of labels, modified linkages, and nonstandard bases attached throughout the chain. The synthesis of short (less than 40-nucleotide) probes and primers requires no more special expertise than the ability to read the synthesizer operator’s manual, and longer oligonucleotides (up to 150 nucleotides) can be synthesized with greater care. The introductory section of this appendix provides strategies for the maximization of synthetic yield, the generation of sequences containing site-specific modifications, and the isolation of synthetic oligonucleotides. Protocols describe monitoring the progress of synthesis via the trityl assay (see Basic Protocol 1 and Support Protocol) and methods for deprotection of DNA (see Basic Protocol 2) and RNA (see Basic Protocol 3) oligonucleotides. This appendix augments the detailed instructions provided by the manufacturers of oligonucleotide synthesizers. A functional understanding of the synthesis chemistries, coupled to insights on the mechanical operation of the synthesizer, will allow the user to minimize input time and maximize oligonucleotide output. INTRODUCTION TO CHEMICAL NUCLEIC ACID SYNTHESIS Many hydroxyl and amine moieties make nucleic acids very animated molecules with rich chemistries of their own that can interfere with the phosphite triester reactions used to couple the nucleotide monomers; therefore, protection strategies are necessary in chemical synthesis to mask the functional groups on the monomers so that the only significant reaction is the desired 3′ to 5′ sequential condensation of monomers to the growing oligonucleotide. These synthetic protecting groups must be chosen so that they can be removed easily to expose the natural nucleotides. The fully protected monomers for nucleic acid synthesis are generally called phosphoramidites (Fig. A.3C.1; also see recipe in Reagents and Solutions). In traditional protection schemes the nucleophilic amino moieties on the bases are protected (see UNIT 2.1) with either isobutyryl (N-2 of guanine) or benzoyl (N-6 of adenine and N-4 of cytidine) groups, both of which can be removed at the completion of synthesis by ammoniolysis. However, recent advances have lead to the widespread use of phenoxyacetyl (PAC) protection of adenosine, dimethylformadine (DmF) protection of guanosine, and acyl protection of cytosine to yield oligonucleotides that can be deprotected rapidly under very mild conditions (Reddy et al., 1994). The 5′ primary hydroxyl of the ribose sugar is protected (see UNIT 2.3) with a dimethoxytrityl (DMT) ether moiety which is removed by mild protic acids at the beginning of each coupling cycle. The efficiency of synthesis at each coupling cycle can be monitored by detecting the release of the chromophore trityl cation. To synthesize nucleic acids with the natural phosphodiester backbone, the 3′ secondary hydroxyl function of the ribose sugar is derivatized with a highly reactive phosphitylating agent. The phosphate oxygen on this moiety is usually masked by β-cyanoethoxy and diisopropylamine protecting groups. By insulating the phosphate oxygen with alternative groups, modified phosphate backbones may be accessed. Finally, Contributed by Andrew Ellington and Jack D. Pollard, Jr. Current Protocols in Nucleic Acid Chemistry (2000) A.3C.1-A.3C.22 Copyright © 2000 by John Wiley & Sons, Inc.
Commonly Used Techniques
A.3C.1
for ribonucleic acids the secondary 2′ hydroxyl of the ribose is shielded throughout the chemical synthesis by the tert-butyldimethylsilyl group (UNIT 2.2). The convenience of using these protecting groups for automated nucleic acid synthesis is that they yield nearly lesion-free natural nucleic acids with high efficiency through simple hydrolysis, nucleophilic displacement, and redox chemistries. In a standard synthesis cycle, the nucleotide chain grows from an initial protected nucleoside derivatized via its terminal 3′ hydroxyl to a solid support (see UNIT 3.1). Reagents and solvents are pumped through the support to induce the consecutive removal and addition of sugar protecting groups in order to isolate the reactivity of a specific chemical moiety on the monomer and effect its stepwise addition to the growing oligonucleotide chain. This design eliminates the need to purify synthetic intermediates or unreacted reagents, because they
OMe
O
MeO
base
O O
Z
5' dimethoxytrityl protection
O
O
O
O
Si
2' t-butyldimethylsilyl protection (Z)
O
base
O
base
O
HN O
Z
P N
N N
CN
O
β-cyanoethyl phosphorus protection
O
benzoylcytidine protection O
O N N
HN NH
O
N
isobutyrylguanine protection Introduction to the Synthesis and Purification of Oligonucleotides
N N
N N
benzoyladenine protection
Figure A.3C.1 Structure of nucleoside phosphoramidite units, showing traditional protection groups for 2′ hydroxyl, phosphate, and base moieties. For DNA, Z = H (no protection required); for RNA, Z = O-protection group.
A.3C.2 Current Protocols in Nucleic Acid Chemistry
are simply rinsed off the column at the end of each chemical step. Assembly of the protected oligonucleotide chain is carried out in four chemical steps: deblocking, activation/coupling, oxidation, and capping (Fig. A.3C.2). Cleavage and deprotection then reveal the single-stranded nucleic acid. Deblocking The synthesis cycle begins with the removal of the acid-labile DMT ether from the 5′ hydroxyl of the 3′ terminal nucleoside. This is usually accomplished by using dichloroacetic acid (DCA) in dichloromethane. The resulting trityl cation chromophore can be quantitated to determine coupling efficiency (see Basic Protocol 1). After deblocking, the 5′ hydroxyl is the only reactive nucleophile capable of participating in the subsequent coupling step. Since the nitrogenous bases of the growing DNA chain are susceptible to acid-catalyzed depurination, the deblocking step is short, and an acetonitrile rinse thoroughly removes the deblocking agent from the support. Also, coupling efficiency and accuracy are increased by this wash, since premature detritylation of the incoming phosphoramidite monomer is prevented. Activation/Coupling Following deblocking of the 5′ hydroxyl group, the next protected phosphoramidite is delivered to the reaction column along with the weakly acidic activator tetrazole (pKa = 4.8). Nucleophilic attack of the previously freed 5′ hydroxyl upon the incoming monomer elongates the nucleic acid chain. Because this protonated phosphoramidite is so reactive, the coupling reaction is usually complete within 30 sec. A molar excess of tetrazole over the phosphoramidite ensures complete activation, and a molar excess of phosphoramidite over free 5′ hydroxyls of the growing chain promotes efficient coupling. To optimize the coupling efficiency, the amounts of reagents injected and the coupling time can be varied (see Synthesizing Long Oligonucleotides). Capping In spite of these efficiency measures, a small percentage of the support-bound nucleoside’s 5′ hydroxyls do not couple to the incoming activated monomer. They must be rendered inactive to minimize deletion products and simplify the purification process. Usually, acetic anhydride and N-methylimidazole dissolved in pyridine and tetrahydrofuran (THF) act to create an acylating agent that “caps” the unextended 5′ hydroxyls. The 5′ acetyl ester cap is unreactive in all subsequent cycles and is removed during the final ammonia deprotection step. Additional acetonitrile washing subsequent to capping can increase synthetic yield. After coupling and capping, the internucleotide linkage is a trivalent phosphite triester that is extremely unstable and must be oxidized to a phosphotriester, which will ultimately yield natural DNA. Oxidation In the last step of the cycle, the unstable phosphite triester linkages are oxidized to a more stable phosphotriester by 0.02 M iodine dissolved in water/pyridine/THF (see recipe for oxidizer). An iodine-pyridine adduct forms to the phosphite triester and is subsequently displaced by water to yield phosphorus oxidized to the pentavalent state. Pyridine also neutralizes the hydrogen iodide byproduct. Because the oxidizer contains water, the support is rinsed several times with acetonitrile following this reaction. One cycle of monomer addition is then complete, and another cycle begins with the removal of the 5′ DMT from the previously added monomer. Cleavage/Deprotection At the end of the synthesis, the final trityl can either be removed with a final acid wash (“trityl-off”), or be left on for purification purposes (“trityl-on”). The oligonucleotide
Commonly Used Techniques
A.3C.3 Current Protocols in Nucleic Acid Chemistry
HO
Bn+1
O O O P
–
O
O
O
Bn
O
n
O –
O
P O
O
B final: cleavage/deprotection
OH DMT-O
O
B1′
1. deblocking O P
DMT-O
O
B2′
O O P O O
NC
HO-O
O
B1′
O O
B1′ P
O
2 activation/coupling
P
4. oxidation 3. capping
DMT-O
O
B2′
DMT-O
O
O O
O
B1′ NC
O
O P
O
O
B1′
NC
O
O P
B2′
N
O P
O P
P
= polymer support
Figure A.3C.2 Steps in the assembly of the protected oligonucleotide chain.
Introduction to the Synthesis and Purification of Oligonucleotides
A.3C.4 Current Protocols in Nucleic Acid Chemistry
itself is removed from the support with concentrated ammonium hydroxide. Additionally, this treatment deprotects the phosphorus by β-elimination of the cyanoethyl group and removes the protecting groups from the heterocyclic bases to yield a single-stranded nucleic acid. RNA Synthesis RNA chemical synthesis is identical to that used for DNA except for the need for an additional protecting group at the 2′ hydroxyl of ribose. This position is usually protected with tert-butyldimethyl silyl groups, which are stable throughout the synthesis (Fig. A.3C.1). They are removed at the final deprotection step by addition of a basic fluoride ion such as tetrabutylammonium fluoride (TBAF). The remaining positions on both the sugar and the bases are protected in the same fashion as for DNA. By adjusting several parameters in the DNA synthesis protocol—including the coupling times, monomer delivery rate, frequency of washing steps, and types of capping reagents—stepwise coupling efficiencies of up to 99% can be obtained (Wincott et al., 1995; G. Glick, pers. comm.). However, for the casual user this yield represents the exception rather than the norm, and only shorter oligoribonucleotides (<20 bases) should initially be attempted. STRATEGIES FOR NUCLEIC ACID SYNTHESIS A Checklist for Nucleic Acid Synthesis Consistency and planning are the keys to reliable nucleic acid synthesis. Organizing these repetitive tasks into a standard operating protocol will streamline efficiency and produce better-quality oligonucleotides. 1. Plan syntheses to optimize machine use. Oligonucleotides of similar size should be combined in parallel runs, since synthesizing many short oligonucleotides followed by a longer one is faster than mixing the sets on dual-column synthesizers. 2. Determine the total number of bases to be incorporated. Be sure there are enough of the required reagents and phosphoramidites available for the entire synthesis. Consult the synthesizer manual for the amount of reagent needed for each coupling (also see Troubleshooting). Remember that different synthetic scales (e.g., 0.25 µmol versus 1 µmol) require different amounts of reagents. Syntheses should be planned so that a phosphoramidite is almost completely exhausted. Also, phosphoramidites that have been dissolved for >2 weeks should be replaced. 3. Consider special programming requirements. Many synthesis options pertaining to the scale of the synthesis, backbone composition, and presence of protecting groups can be modified. Create a log for users to fill out detailing exact synthesis requirements, and check the log prior to synthesis. A computerized log book is especially useful and allows for an organized oligonucleotide nomenclature (e.g., R20.17 may refer to Rebecca’s 20-mer, the 17th 20-mer made on the system). 4. If the synthesizer lacks a trityl monitor, set up a fraction collector to monitor the chromophoric trityl cation release (see Basic Protocol 1). 5. Initialize the system. If previous oligonucleotides were cleaved from their supports automatically, rinse the columns for 30 sec with acetonitrile to remove any remaining traces of ammonium hydroxide. If the machine has been at rest for >6 hr, immediately before starting the synthesis (and following the addition of fresh reagents) remove any stale reagents or moisture from the lines by priming them. This will maximize Commonly Used Techniques
A.3C.5 Current Protocols in Nucleic Acid Chemistry
the first coupling step’s efficiency. Check the reagent and phosphoramidite flow rates to ensure that reagents are being properly delivered. 6. Start the synthesis. Confirm that the flow through the lines and columns is not obstructed and monitor the first few trityl releases. An abortive synthesis of a 60-mer sequence wastes much more material than a failed 3-mer run. Synthesizing Long Oligonucleotides (≥100 Bases) Modern synthesizers can routinely produce sequences of 150 or more nucleotides in usable amounts (≥10 µg). Several strategies can be employed to enhance the yield from syntheses longer than 100 bases (UNIT 3.3). 1. Exclude water from the system (the importance of this cannot be overstated; see Critical Parameters). Replace reagents on the machine with fresh ones before all long syntheses. This is particularly important for the phosphoramidites and especially for guanosine phosphoramidite, since it decomposes more quickly than the other two protected bases (Zon et al., 1985). 2. Monitor trityl releases for shorter runs prior to attempting the synthesis of long oligonucleotides to limit wastage of expensive reagents. In general, if the stepwise efficiency of synthesis is <99%, alter parameters to increase the efficiency on shorter sequences before attempting to synthesize a long oligonucleotide (see Basic Protocol 1). 3. Use dichloroacetic acid (DCA) rather than trichloroacetic acid (TCA) for deblocking if the synthesizer is compatible with this reagent. Depurination (cleavage of the glycosidic bond) under acidic conditions is a prominent side reaction that ultimately limits DNA synthesis. DCA tends to show much better synthetic yields than trichloroacetic acid, especially for longer oligonucleotides (R. Pon, pers. comm.). Use a 2% (v/v) DCA/1,2-dichloroethane mixture. 4. Modify the synthesis protocol to increase the coupling time of the phosphoramidite. Also, additional methylene chloride wash steps included prior and subsequent to deblocking, along with increased acetonitrile washing subsequent to capping, lead to increased yields (G. Glick, pers. comm.). 5. Increase the phosphoramidite concentration to enhance the coupling efficiency—e.g., use a concentration of 50 mg/ml (double the normal concentration; 20-fold molar excess over the synthetic polynucleotide chain) for longer sequences. 6. Using a support matrix such as control pore glass (CPG; UNIT 3.1) with a loading capacity of <40 µmol/g can greatly increase yields of long oligonucleotide. Furthermore, the pore size of the support should be 1000 Å for >100-mers and 2000 Å for 200-mers (Gait, 1984) to alleviate molecular crowding and steric effects. For a typical 1-µmol-scale synthesis of a 150-mer, 20 mg of a support with a loading capacity of 5 mmol/g is used.
Introduction to the Synthesis and Purification of Oligonucleotides
If it is too difficult to synthesize the desired sequence in a reasonable yield, or if oligomers >150 bases are desired, the nucleic acids may be made in segments and ligated (Bartel and Szostak, 1993) following PCR with a proofreading polymerase such as Pfu or Tth. Note, however, that since PCR is usually a mutagenic procedure, any products generated should be checked by sequencing. Also, very long synthetic oligonucleotides (300 to 600 bases) have also been synthesized directly, and in spite of incredibly low yields, rare full-length products have been successfully amplified by PCR (Ciccarelli et al., 1991).
A.3C.6 Current Protocols in Nucleic Acid Chemistry
Finally, mutually primed synthesis (e.g., CPMB UNIT 8.2) can also be a suitable option for oligomers >150 bases. Synthesizing RNA Since many structural and mechanistic studies are underway concerning catalytic RNAs, the catalog of commercially available modified RNA monomers has recently bloomed (Table A.3C.1). RNA chemical synthesis (see UNIT 3.5) has become as routine as that of DNA (see UNIT 3.3) and typically uses identical 5′-dimethoxytrityl-β-cyanoethyl-protected phosphoramidites except for an additional protecting group on the 2′ hydroxyl (see UNIT 2.2). tert-Butyldimethylsilyl protection of the 2′-hydroxyl group is the basis of most commercially available RNA phosphoramidites, since the silyl group is stable to both acid and base and can be removed with fluoride ion. Recently, however, RNA monomers with 2′-acetal groups (FPMP, CEE; also see UNIT 2.2) have appeared; these have the advantage of being conveniently removed at pH 2 just prior to use of the particular sample of RNA. Strategies for ribophosphoramidite protection are an active area of research, and recent work with 5′-silyl ethers in conjunction with 2′-orthoester protection has proven particularly interesting (S. Scaaringe, pers. comm.). For the casual user of RNA, it is often easier just to purchase small quantities of the required sequence from a ribo-oligonucleotide synthesis company such as Baron Consulting, Dharmacon Research, Genosys, or Peninsula Labs (see SUPPLIERS APPENDIX). Isomeric purity of the phosphoramidites is often variable because of the difficulties inherent in distinguishing between the vicinal hydroxyls of ribose. If a homogeneous population of 5′-to-3′-linked oligoribonucleotides is required (as in most cases), then a thin-layer chromatography (TLC) or 31P nuclear magnetic resonance (NMR) analysis of the starting phosphoramidites should be performed to establish their isomeric composition. While phosphorus NMR facilities are not generally available to molecular biologists, TLC is both inexpensive and straightforward. Recommended solvent systems to separate the 2′ and 3′ ribonucleotide phosphoramidites are 1:1 ether/chloroform, or 40:58:2 or 50:46:4 dichloromethane/hexane/triethylamine (Usman et al., 1987). With a few slight modifications, the procedures and precautions described for DNA synthesis chemistry apply to RNA as well (UNIT 3.5). Since stepwise coupling efficiency is lower than that of DNA, even greater care should be taken to exclude water completely from the closed system. Because the 2′ hydroxyl is often protected with sterically hindering protecting groups, reaction times for RNA reagents tend to be longer, and adjustments should be made to phosphoramidite concentrations and coupling times, as detailed below. As is true for all RNA work, equipment and reagents that will contact unprotected oligonucleotides should be RNase free to avoid degradation of the synthesized material. Depending on the synthesizer and coupling program used, RNA phosphoramidites are suspended in dry acetonitrile at a concentration of 0.1 to 1.0 M with a 6- to 10-fold excess of reagents delivered per 300 sec coupling. S-ethyltetrazole and DCI have also been found to be a more effective activators than the traditional tetrazole (Sproat et al., 1995; Vargeese et al, 1998). Also, additional methylene chloride washing steps included prior to and subsequent to deblocking, along with increased acetonitrile washing subsequent to capping, lead to increased yields (G. Glick, pers. comm.).
Commonly Used Techniques
A.3C.7 Current Protocols in Nucleic Acid Chemistry
Table A.3C.1
Suppliers of Unnatural and Modified Phosphoramidites
Phosphoramidite typea
Suppliersb
Fast/mild deprotecting DNA and RNA monomers
Cruachem Glen Research Perkin-Elmer BioGenex ChemGenes PerSeptive BioSystems Sigma
Modified DNA monomers
Appligene Interactiva BioTechnologie Glen ChemGenes Cruachem Sigma Perkin-Elmer Solid Phase Science
Convertible deoxynucleosides
Glen Research
Radiolabeled deoxyphosphoramidites
Cambridge Isotopes
Modified RNA monomers
ChemGenes BioGenex Boehringer Mannheim Glen Research Dalton Perkin-Elmer Sigma Cruachem
Labeling monomers
Clontech Cruachem Glen Research Solid Phase Science Perkin-Elmer BioGenex Sigma ChemGenes Boehringer Mannheim ABI Biotechnology PerSeptive BioSystems Genosys
Uniquely structured oligonucleotide branched or cyclic
Clontech Glen Research
Non-enzymatically extended 3′ ends
Glen Research
Alternative backbones
Glen Research ChemGenes
aAll phosphoramidites are 5′ DMT and 3′ cyanoethyl protected (CED) unless otherwise noted.
Introduction to the Synthesis and Purification of Oligonucleotides
bSee Suppliers Appendix for supplier contact information.
A.3C.8 Current Protocols in Nucleic Acid Chemistry
Incorporation of Modified Nucleosides Chemical nucleic acid synthesis allows for the incorporation of unnatural or modified bases, as well as a variety of labeling moieties, into an oligonucleotide (see Chapter 4). This can be extremely useful for testing models of structural interactions between enzymes and nucleic acids, selecting labeled molecules from a population of unlabeled ones, or gaining insights into the parameters that govern nucleic acid structure and chemistry. Modified backbone chemistries such as phosphorothioates, phosphoroamidates, and phosphotriesters are also readily available. In general, the bases themselves can be obtained commercially and are handled like any other phosphoramidite; however, consult the company that supplies the analog about necessary modifications to programs or reagents (see Table A.3C.1). Typically, the only adjustment needed is to dissolve the modified base at a somewhat higher concentration than normal to overcome problems associated with reactivity. Most of the methods used to increase the yield of long and ribo-oligonucleotides may be applied to the synthesis of modified nucleic acids. When synthesizing modified oligonucleotides, compatibilities of the chemistries, placement of modifications relative to other chemical groups, and 5′ to 3′ directionality are all factors to consider. Generally when an oligonucleotide is end-labeled/modified, a long flexible tether is added to allow greater accessibility. Stretches of four deoxythymines are often used for this purpose. Also, adding deoxythymines 5′ to the label (5′-TTTT-label-3′) can aid in separating labeled molecules from unlabeled ones by size. Note that some tagging phosphoramidites allow for the enzymatic extension or kinasing of the modified oligonucleotide, while others do not. Finally, oligonucleotides may also be synthesized directly on solid glass supports (Cohen et al., 1997). Terminal transferase can be used as an alternative means of incorporating modified bases at the 3′ end of an oligonucleotide (Ratliff, 1982; e.g., CPMB UNIT 3.6). This enzyme is tolerant of a variety of substrates, and has been used to add deoxynucleotide triphosphates derivatized at virtually every position (C-8 on adenine, any of the amino groups, C-5 on cytosine, O-6 on guanosine) to DNA. It also functions, though less well, with RNA bases. It can use any DNA oligonucleotide that is at least 2 bases long [d(pXpX)] and contains a free 3′ hydroxyl as a primer. A potential problem in preparing homogeneous polynucleotides using terminal transferase is that a statistically random number of bases is added to the 3′ end of the template (with the exception of molecules such as cordycepin, which act as chain terminators due to the absence of 3′ hydroxyl). If a single species is desired, it can be gel purified. Polynucleotide phosphorylase may also be used to incorporate modified bases at the 3′ end (Gillam and Smith, 1980). A more controlled means of introducing modified nucleotides relies on T4 RNA ligase and substrates of the form A(5′)ppX (where X can be virtually any molecule, including for example ribose or amino acids, in a pyrophosphate linkage with adenosine; Uhlenbeck and Gumport, 1982). The minimal template for reactions of this form is a trinucleoside containing a free 3′ hydroxyl. RNA reacts much better than DNA, and single-strand molecules act as better templates than double-stranded ones. Since 3′ hydroxyl groups are required, substrates of the form A(5′)ppXp will undergo only a single round of addition, unlike the similar reaction with terminal transferase. In some cases, the compound A(5′)ppXp can be generated directly by RNA ligase from pXp and ATP, although the substrate requirements for the X moiety are much more strict than in the ligation reaction. Thus, while virtually any dinucleotide of the form A(5′)ppX can be added to an oligonucleotide, only a few compounds (primarily sterically “small” derivatives of natural bases) can be used by the enzyme to form A(5′)ppXp from pXp. Commonly Used Techniques
A.3C.9 Current Protocols in Nucleic Acid Chemistry
T4 RNA ligase can catalyze the ligation of single-stranded oligonucleotides in the presence of ATP and various analogs (Kinoshita et al., 1997). Templates prepared by terminal transferase or by T4 RNA ligase that contain modified nucleotides (or other adducts) at their 3′ termini may be able to act as substrates in this reaction. This would allow modified nucleotides to be introduced into the middle of a longer chain. However, the substrate specificity of the enzyme for the 3′ hydroxyl donor is highly substrate dependent and will have to be determined empirically. Synthesizing Degenerate Oligonucleotides Current combinatorial and “irrational” nucleic acid design methodologies focus on the ability to create large pools of random sequences from which useful sequences may be culled (Szostak, 1992; also see Chapter 9). In addition, random mutagenesis using degenerate oligonucleotides allows for the exploration of “sequence space” surrounding a given protein or RNA structure. Sequences can be produced that give a completely random distribution of nucleotides at a given position or, alternatively, the sequence can be biased or “doped” toward a particular base with only a low level of randomization. Most synthesizers can be programmed for in-line degenerate mixing of bases, which is useful if only a few positions are to be randomized. A potential problem with this method is that, if mixing is incomplete, the sequence will be skewed toward whichever phosphoramidite enters the column first, since the reaction of the activated phosphoramidite with the free 5′ hydroxyl is extremely fast. Therefore, while in-line mixing will generate all base substitutions at a given position, the distribution of these substitutions may not be uniform. If a statistically random distribution of nucleotides is required or if long stretches of random sequence are to be made, it is better to manually mix the phosphoramidites together and use this mixture for the degenerate position. A true random distribution may be obtained by mixing A, C, G, and T phosphoramidites in a 3:3:2:2 molar ratio to compensate for the faster coupling times of G and T phosphoramidites (D. Bartel, pers. comm.). On synthesizers where phosphoramidites are loaded without detaching the bottle, the mixing generated by sequential loading of each phosphoramidite into the extra bottle is sufficient to generate randomized sequences. Oliphant and Struhl (Oliphant et al., 1986; Oliphant, 1989) have constructed degenerate oligonucleotides using mixed phosphoramidites, but have modified the synthesis protocol by deleting the capping step during the random additions. This increases the overall yield of long oligonucleotides, since sequences that fail to elongate are not terminated, and the size of the final product is more heterogeneous. This method is particularly useful if deletions, as well as randomized bases, in a given sequence are required. Hermes et al. (unpub. observ.) have developed a detailed protocol for producing statistically mutated oligonucleotides. This method employs in-line mixing between pure phosphoramidite contained in separate bottles and equimolar mixtures of the four bases contained in an additional bottle. The obvious advantage of such a method is that doped and clean sequences can be synthesized on the same oligonucleotide. Whether or not this method is employed, it is an example of how to dissect the chemistry of mixed-site oligonucleotide synthesis. Hermes et al. (1989) have shown that mutations introduced by this method are indeed statistically random.
Introduction to the Synthesis and Purification of Oligonucleotides
Regardless of the strategy employed, the level of misincorporation of an oligonucleotide should be decided in advance by the mutagenesis frequency desired. Quantitatively, this level is given by the probability distribution: P (n,m,x) = [m!/(m − n)!n!][x]n [1 − x]m×n
A.3C.10 Current Protocols in Nucleic Acid Chemistry
where P is the probability of finding n errors in an oligonucleotide m in length with x level of misincorporation (fraction “wrong” nucleotides delivered). This equation describes a Poisson distribution. If primarily single mutations are desired, then x should be maximized for n = 1; if multiple mutations (e.g., doubles, or triples in a single oligonucleotide) are necessary, x should be correspondingly higher. If the mix is optimized for n mutations, then n − 1 and n + 1 mutations will occur in roughly equal amounts and n mutations will be the most frequent. Cloning randomized oligonucleotides can be difficult, since a complementary wild-type sequence will generate mismatches that may result in biased correction in vivo. To avoid this problem, second-strand synthesis can be primed from a nonrandom portion of the sequence, or mutually primed synthesis (e.g., CPMB UNIT 8.2) can be utilized. Alternatively, Derbyshire et al. (1986) describe the direct cloning of doped single-strand material with “sticky ends” into a double-stranded cloning vector. Finally, Reidhaar-Olson and Sauer (1988) describe the synthesis of complementary oligonucleotides containing inosine (which can pair with any of the four natural bases) directly across from randomized codons. This method resulted in the successful introduction of a wide variety of mutations into the gene for lambda repressor, although there was a slight compositional bias in cloned sequences. STRATEGIES FOR OLIGONUCLEOTIDE PURIFICATION Deprotected nucleic acids may be purified and isolated by a variety of methods. The method of choice will depend on the availability of resources, the purity required—some methods cannot separate (n−1)-mers from n-mers—and time considerations. Any of the methods described below can be used to clean up crude material. Isolation Methods Precipitation Direct precipitation of the nucleic acids constitutes a fast and efficient purification from contaminating small molecules such as urea and phenol, but does not allow for purification of abortive synthesis products from the full-length one. If oligonucleotide size separation is required, this method should be used in conjunction with some of the other methods described. After deprotection, resuspend the whitish pellet obtained in water. Add MgCl2 to a final concentration of 10 mM and mix along with 5 vol ethanol. Precipitation should be immediate. Freeze the sample briefly at −20° or −70°C. Centrifuge the precipitated material, wash with 95% ethanol, dry, and resuspend in water. Precipitated deoxyoligonucleotides can be used for sequencing or cloning. They can also be used in PCR reactions, although the efficiency of amplification may be reduced as compared to gel-purified oligonucleotides. If the DNA is to be phosphorylated, a more thorough purification procedure is necessary, since T4 polynucleotide kinase is inhibited by lingering ammonium ions. Sizing columns Size-exclusion chromatography is extremely useful as a final purification step, especially when small-molecule contamination occurs with otherwise pure oligonucleotides, but (like precipitation) does not effect purification of abortive synthesis products from the full-length one. If oligonucleotide size separation is required, this method should be used in conjunction with some of the other methods described. Oligonucleotides purified by PAGE might contain small amounts of low-molecular-weight contaminants such as urea or phenol that might inhibit enzymatic reactions; sizing columns are a simple way to
Commonly Used Techniques
A.3C.11 Current Protocols in Nucleic Acid Chemistry
decontaminate these samples. Spin columns are simple to prepare and use, but gravityflow columns give better, more reproducible separation. The type of resin used should be adjusted based on the size of the oligonucleotide being purified (e.g., Sephadex G-25 for 25-mers or less, G-50 for longer sequences). Reversed-phase cartridges A hydrophobic matrix may be used to separate full-length from abortive synthesis products if the final trityl group is left on following the final monomer coupling reaction. The resulting hydrophobically tagged full-length “trityl-on” oligonucleotide can be separated easily from failure sequences, which lack trityl groups and do not efficiently bind the hydrophobic matrix. Several companies supply columns designed specifically for the purification of “trityl-on” oligonucleotides (e.g., Applied Biosystems oligonucleotide purification cartridges). The procedure is simple and can be performed on a number of samples in parallel within only a few minutes. The yield from such columns is excellent, often >80% of the applied sample. However, although a majority of failure sequences are removed using this method, many shorter sequences are co-purified with the desired full-length material. Some of these fragments are due to cleavage of depurinated DNA. These apurinic molecules can be eliminated prior to cleavage from the column by treatment with lysine (Horn and Urdea, 1988). In addition, if care is not taken to wash and elute samples from these columns slowly, some inhibitors (particularly of ligation reactions) may co-elute. Denaturing PAGE Denaturing polyacrylamide gel electrophoresis (APPENDIX 3B) separates oligonucleotides with single-residue resolution and is the method of choice for purifying full-length oligonucleotides. However, the compatibility of the chemistries of modified nucleotides incorporated into the nucleic acids and acrylamide matrix should be checked (thiolated oligonucleotides seem to undergo Michael addition to the acrylamide, which renders them irreversibly capped). HPLC Liquid chromatography can also separate oligonucleotides with single-residue resolution, but its chief advantage is speed. Total run time can be as short as 30 min. The use of alkali perchlorate salts has made ion exchange the HPLC method of choice given that long oligonucleotide (>40 residues) may be easily purified in large scales (≥25 µmol; Sproat et al., 1995; Warren and Vella, 1994). However, secondary structural migration anomalies are generally more severe than those found with PAGE. Depending on the system employed, the amount of oligonucleotide that can be purified in a single chromatographic run can be comparable to PAGE, and sample recovery is typically >70%. For laboratories with an HPLC system and a need to routinely purify short oligonucleotides with no secondary structure, this method is ideal. Oligonucleotides can be purified with HPLC by charge differences through ion exchange or hydrophobicity if the final trityl group is left on following the ultimate monomer coupling reaction. Only the “trityl-on” systems use buffers that can be lyophilized. A more complete treatment of the complexities of oligonucleotide purification by HPLC can be found in Applied Biosystems User Bulletin no. 13 on oligonucleotide purification (Applied Biosystems, 1988). Confirming the Oligonucleotide Sequence Introduction to the Synthesis and Purification of Oligonucleotides
Most oligonucleotides that are used for cloning need not be checked immediately after synthesis, since the clones themselves will be checked after biological or enzymatic amplification. However, in cases where a sequence will be used in a structural application
A.3C.12 Current Protocols in Nucleic Acid Chemistry
such as mobility-shift assays (e.g., CPMB UNIT 12.1), filter-binding assays (e.g., CPMB UNIT 12.8), or crystallography, it is desirable to confirm the sequence. For almost all oligonucleotides, this usually requires chemical sequencing (e.g., CPMB UNIT 7.5). Although sequencing will confirm that the correct product was made, it cannot determine the homogeneity. How unnatural or protected bases will react during chemical sequencing, or how they will affect mobility on a sequencing gel, is not predictable. In order to determine what fraction of molecules contain only the natural bases (A, T, C, and G), it is necessary to digest the DNA enzymatically to completion and to examine its composition by a comparison with standard bases separated by HPLC. At least one HPLC buffer system has been developed specifically for examining modified nucleosides in chemically synthesized oligonucleotides (Eadie et al., 1987). The insertion and deletion rates for chemical nucleic acid synthesis are non-negligible. The rate of insertions (presumably due to DMT cleavage via tetrazole) has been measured to be as high as 0.4% per position, and the rate of deletions (presumably due to incomplete capping) has been found to be as high as 0.5% per position (A. Keefe and D. Wilson, pers. comm.). Therefore, for an oligonucleotide 100 bases in length and assuming a random, noncorrelated mechanism of action, only about 40% of the sequences will be the intended one. Finally, matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS; UNIT 10.1), which has recently emerged as a new biotechnology tool, may be used to determine the sequence of deoxy and ribo-oligonucleotides of up to 60 bases (Zhu et al., 1997). MONITORING DNA SYNTHESIS USING THE TRITYL ASSAY A trityl cation is released from the 5′ end of the growing oligonucleotide during each synthesis cycle, and the yield of each step of the synthesis can be determined by spectrophotometrically measuring the amount of trityl cation liberated. This procedure is the simplest and most rapid means available for the identification of problems with synthesized oligonucleotides prior to deprotection and purification. Also, in-line quantitative trityl monitors can be interfaced to most synthesizers (Ana-Gen Technologies).
BASIC PROTOCOL 1
Materials 0.1 M para-toluene sulfonic acid (TSA; monohydrate) in acetonitrile (see recipe for dry acetonitrile) 15-ml glass tubes (graduated, if possible) 1. Collect the trityl cation solution in 15-ml glass tubes after each step. It is helpful to use graduated tubes so that a uniform final volume of acetonitrile can be attained before assay.
2. Dilute the first three and last three fractions to 10 ml with 0.1 M TSA in acetonitrile. Mix thoroughly. CAUTION: Handle the solution consisting of dichloroacetic acid (or trichloroacetic acid) in dichloromethane and acetonitrile with gloves, because it is corrosive as well as toxic. Avoid prolonged contact with toxic acetonitrile fumes. Do not pipet by mouth. Although 2.5 ml of deblocking solution is initially released, because acetonitrile is a volatile liquid the volume may change during the course of a synthesis. Fractions may sit for several days before being assayed without affecting the results. Samples that have evaporated to dryness must be thoroughly redissolved. The acid ensures protonation of the trityl groups, making them more strongly colored. It is misleading merely to visualize the yellow/orange color of the fractions, however, since variable volumes of differing acidity are often released.
3. Dilute each sample 20- to 50-fold in the same solution. Measure the absorbance at 498 nm versus acetonitrile/TSA.
Commonly Used Techniques
A.3C.13 Current Protocols in Nucleic Acid Chemistry
These dilutions are necessary because most spectrophotometers cannot accurately measure absorbances >2.
4. Calculate the stepwise coupling efficiency and absolute yield for the synthesis as a whole. The stepwise efficiency is given by: (stepwise efficiency) n =
average absorbance of last three fractions average absorbance of first three fractions
where n is the number of trityl nucleotides in the oligonucleotide (equal to the length of the oligonucleotide for trityl-off syntheses). The absolute yield of product is given by: µmol DMT =
(absorbance of last fraction) × (dilution factor) × (10 ml ) 70 ml / µmol
where 70 µmol is the extinction coefficient of DMT. The average stepwise efficiency is useful in determining the relative efficiency of each cycle of the synthesis. The absolute yield is useful for determining how many milligrams of product are present for subsequent purifications, although not all of the product will be of the desired length. The average absorbance of the first and last few steps is used to avoid discrepancies in individual trityl assays. Clearly, if the synthesis was performed on a 1-mmol column, the absolute yield in the first few fractions should be close to 1 mmol. A low absolute yield (absorbance) in the first trityl release, followed by higher absolute yields (absorbances) in the next few fractions, is sometimes observed. This is because some spontaneous detritylation occurs during storage of the columns and the trityls are subsequently rinsed off during the prime lines program. In this case, the average stepwise efficiency should be calculated with fractions 2 through 4. It should be noted that visual assessment of the trityl fractions cannot begin to detect subtle differences (<5%) which may be critical in terms of overall yield.
5. Perform troubleshooting assay (see Support Protocol) if average stepwise efficiency is low. SUPPORT PROTOCOL
USING THE TRITYL ASSAY FOR TROUBLESHOOTING If the average stepwise efficiency for the oligonucleotide synthesis is low, each fraction should be assayed in order to diagnose the problem (what counts as a “low” yield depends on the length and quantity of oligonucleotide desired). In general, synthetic efficiency should be >99% per step, although lower stepwise efficiencies can be tolerated for short oligonucleotides (<40 bases) or where yield is not critical. Three classes of failures can be detected by trityl assays. A low absolute yield at the first step followed by similarly low absorbances that results in a low overall yield is commonly due to inefficient purging of activator or phosphoramidite lines prior to the synthesis. Such a problem frequently occurs when a synthesizer has not been used for several days. Purge the lines with dry reagents prior to starting the run to avoid inefficient synthesis. Most machines have a priming program precisely for this purpose.
Introduction to the Synthesis and Purification of Oligonucleotides
If the stepwise efficiency of each step is low, there is a systematic problem with one of the common reagents, such as the acetonitrile. Often this is due to moisture in one or more of the reagents, and it is more likely if reagents have not been recently replaced. The phosphoramidites should be used until they are almost completely exhausted during a
A.3C.14 Current Protocols in Nucleic Acid Chemistry
series of syntheses, so that fresh chemicals will be diluted as little as possible by older, potentially wet material (see Critical Parameters). Finally, individual trityl assays are most useful in determining when phosphoramidites have become defective. In this case, drops in stepwise efficiency will only be seen at the coupling steps involving the phosphoramidites in question. Many problems, such as inefficient oxidation or product depurination, cannot be detected by the trityl cation assay. Therefore, the trityl assay procedure should be used in conjunction with HPLC or gel electrophoresis for product analysis, especially if a homogenous population of oligonucleotides is essential. DEPROTECTION OF DNA OLIGONUCLEOTIDES After synthesis is complete, the DNA may be cleaved from the column and protecting groups removed by treatment with ammonia. Although very extended treatment in base can harm DNA, hours of ammoniolysis are still preferred to ensure complete deprotection, since a homogeneous population of “natural” oligonucleotides at slightly lower yield is better than a mixed population of partially deprotected, “unnatural” molecules.
BASIC PROTOCOL 2
To cleave the DNA from the support matrix and remove protecting groups completely, the support-bound product must be treated with concentrated ammonia at 55° to 60°C overnight (≥12 hr). Even with such extended treatment, deoxyguanosine may not be completely deprotected (Schulhof et al., 1987). Raising the temperature at which oligonucleotides are deprotected has been reported to speed up the process (≥5 hr at 70°C). Phosphoramidites with more labile protecting groups such as phenoxyacetyl or dimethylformadine masking adenosine and guanosine have recently become commercially available (Table A.3C.1). These allow essentially complete deprotection within 30 to 60 min at 70°C (Reddy et al., 1994). Also, by replacing the traditional benzyl protection of cytosine with acetyl and using a 1:1 mixture of aqueous ammonium hydroxide and aqueous methylamine, oligonucleotides synthesized with traditional purine protections may be completely deprotected in 5 min at 65°C (Reddy et al., 1995). Finally, anhydrous ammonia gas-phase deprotection of oligonucleotides has recently been described; this provides a convenient method for parallel deprotection of as many columns will fit in a reactor. Since no water is present, the fully deprotected oligonucleotides remain adsorbed to the column matrix, thereby guaranteeing that no cross-contamination will occur. The oligonucleotides can then be eluted with water and desalted or further purified. Using PAC-protected monomers, the cleavage and deprotection processes can be completed in ∼30 min (Boal et al., 1996). Materials Concentrated (14.8 N) ammonium hydroxide (see recipe) Triethylamine 3:1 (v/v) concentrated ammonium hydroxide/ethanol n-Butanol Screw-cap plastic vial (preferably fitted with rubber O ring) Heat block or oven, 55° to 60°C 0.2-µm filter 1. In a screw-cap plastic vial, suspend the synthesis support matrix or the already support-cleaved oligonucleotide in ∼1.0 ml concentrated ammonium hydroxide for a 1-µmol synthesis.
Commonly Used Techniques
A.3C.15 Current Protocols in Molecular Biology
Supplement 47
Depending on the synthesizer or nucleic acid provider, the oligonucleotide may come attached to a support matrix or free in an ammonium hydroxide solution. The volume of ammonium hydroxide in which product is eluted from the synthesizer is variable. The ammonium hydroxide used should not have been diluted by excessive vapor loss.
2. Place the sample in heat block or oven for ≥12 hr at 55° to 60°C. Seal vial tightly with Parafilm (if not fitted with a rubber O ring).
3. After cleavage from the support and deprotection are complete, spin the sample briefly in a tabletop centrifuge to pool the ammonia and support that may have collected in the cap. Let the vial cool to room temperature before opening it to avoid sample boil-over. 4. Filter off the support by passing the liquid through an 0.2-µm filter and wash the filter with 0.3 ml of 3:1 (v/v) ammonium hydroxide/ethanol. 5. Precipitate the oligonucleotide from the resulting supernatant by adding 10 vol n-butanol and vortexing for 15 sec. Centrifuge 10 min at 16,000 × g, 4°C. 6. Remove and discard the single aqueous n-butanol phase to reveal the white oligonucleotide pellet. Oligonucleotides 20 residues or shorter with good trityl responses are typically suitable for use directly in DNA sequencing, PCR amplification, and gel-shift analysis. However, if more homogenous material is required, methods such as denaturing PAGE and HPLC may be employed to further purify the full length product.
7. If a yellowish liquid or crusty pellet remains, rather than a white powder, resuspend the pellet in 0.1 ml distilled water and precipitate again with n-butanol as described above. Generally, it is not necessary to add additional salt for precipitation. Further extraction will aid in removing any residual ammonia or volatile organics. If the yellow color does not disappear, it will ultimately be removed by almost any of the standard purification methods.
8. Lyophilize the sample to dryness in a Speedvac evaporator. This deprotection is primarily intended for oligonucleotides with the trityl group off. When the trityl group is left on, care must be taken that it is not prematurely hydrolyzed from the DNA by acid conditions. During lyophilization, a drop of triethylamine must be added regularly to the sample in order to maintain its basicity (Applied Biosystems, 1988). Heating of the samples should be avoided. BASIC PROTOCOL 3
Introduction to the Synthesis and Purification of Oligonucleotides
DEPROTECTION OF RNA OLIGONUCLEOTIDES Techniques and recommendations for deprotecting RNA are similar to those for DNA. Ammoniolysis cleaves the RNA from the support and frees the bases of their protecting groups. Additionally, the 2′ hydroxyl protecting group must be removed to reveal a functional RNA sequence. Reagents, water, and plasticware to which RNA is exposed must be sterile. The protocol detailed below is for use with nucleosides bearing the standard isobutyryl and benzyl protection on the bases and 2′ silyl protection. Protection options are becoming available that both increase yield and streamline deprotection time (Table A.3C.1). Materials 100% ethanol 3:1 (v/v) concentrated (14.8 N) ammonium hydroxide/ethanol 3 M sodium acetate, pH 5.2 ( APPENDIX 2A) Triethylamine trihydrofluoride Screw-cap plastic vial (preferably fitted with rubber O ring)
A.3C.16 Supplement 47
Current Protocols in Molecular Biology
Heat block or oven, 55° to 60°C 0.2-µm filter Sephadex G-25 column (Amersham Pharmacia Biotech) 1. In a screw-cap plastic vial, suspend the synthesis support matrix or the already support-cleaved oligonucleotide in ∼1.2 ml of 3:1 (v/v) ammonium hydroxide/ethanol for up to a 1-µmol scale. Depending on the synthesizer or nucleic acid provider, the oligo may come attached to a support matrix or free in an ammonium hydroxide solution. The volume of ammonium hydroxide in which product is eluted from the synthesizer is variable. The ammonium hydroxide used should not have been diluted by excessive vapor loss.
2. Place the sample in a heat block or oven for 12 to 16 hr at 55° to 60°C. Seal vial tightly with Parafilm (if not fitted with a rubber O ring). If fast-cleaving phosphoramidites such as PAC-protected purines and acyl-protected cytosine are used, the deprotection time of the bases can be a little as 10 min in methylamine at 65°C (Wincott et al., 1995).
3. After cleavage from the support and base deprotection are complete, spin the sample briefly in a tabletop centrifuge to pool the ammonia and support that may have collected in the cap. Let the vial cool to room temperature before opening it to avoid sample boil-over. 4. Filter off the support by passing the liquid through an 0.2-µm filter and wash the filter with 0.3 ml of 3:1 (v/v) ammonium hydroxide/ethanol. 5. Evaporate the combined solutions to dryness in the Speedvac evaporator without heating. Resuspend the pellet in 0.2 ml of 100% ethanol and evaporate to dryness without heating. 6. Treat the dried residue with triethylamine trihydrofluoride (0.3 ml for a 0.2-µmol or 0.5 ml for a 1-µmol synthesis). Allow to rotate in a foil-covered screw-cap vial in the dark for at least 24 but no more than 48 hr. Alternative methods exist to remove the 2′ silyl protecting groups either under dilute acidic conditions (Kawahara et al., 1996) or with anhydrous triethylamine/hydrogen flouride in N-methylpyrrolidinone (Wincott et al., 1995).
7. To desalt via ethanol precipitation, add an equal volume of water to the triethylamine trihydrofluoride solution and immediately dilute with 1⁄10 vol of 3.0 M sodium acetate, pH 5.2. Precipitate by adding 3 vol of 100% ethanol and chilling for ∼20 min at −80°C. Centrifuge 10 min at 16,000 × g, 4°C. An alternative method exists for isolation of the RNA by diethyl ether precipitation (Song and Jones, 1999).
8. Remove and discard the single ethanol layer to reveal the white oligonucleotide pellet. 9. If a yellowish liquid or crusty pellet remains, rather than a white powder, resuspend the pellet in 0.1 ml distilled water and repeat the sodium acetate/ethanol precipitation as described in step 7. If desired, desalt the RNA on a desalting matrix such as Sephadex G-25. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX. Commonly Used Techniques
A.3C.17 Current Protocols in Nucleic Acid Chemistry
Acetonitrile, dry Since large volumes are used for rinsing lines and dissolving other reagents, acetonitrile is one of the most costly reagents in nucleic acid synthesis. If the DNA synthesizer is used infrequently, it may be useful to produce small amounts of dry acetonitrile immediately prior to synthesis since the acetonitrile will accumulate water after the bottle has been opened. Moisture greatly diminishes synthetic yield. In this case acetonitrile can be prepared in any laboratory equipped for routine distillations. However, the time and effort involved in setting up and maintaining a still should be balanced against the cost of obtaining dry acetonitrile. The use of molecular sieving pouches that do not release metal ions is recommend; they are often available from the synthesizer manufacturer and should keep the acetonitrile to <20 ppm water. Some commercial suppliers now market bulk solvents specifically for nucleic acid synthesis (e.g., Baker “low-water” acetonitrile, 0.002% or 20 ppm water; Burdick and Jackson acetonitrile, 0.001% or 10 ppm water). The cost of these special dry reagents is about the same as that of HPLC-grade acetonitrile (which typically contains 0.01% or 100 ppm water), which is the starting material for a laboratory distillation. CAUTION: Acetonitrile vapor is poisonous.
Ammonium hydroxide Purchase concentrated (14.8 N) ammonium hydroxide in 0.5- to 1.0-liter bottles, and store it at 4°C to reduce the ammonia gas found in the vapor phase above the liquid. CAUTION: Concentrated ammonium hydroxide is extremely caustic. Breathing the vapors is harmful. Always use this compound in the fume hood, as it is possible to be quickly overcome by ammonia fumes and blinded. Ammonia is volatile and its concentration will decrease on repeated opening, until it is no longer completely functional as a deprotecting reagent. When this occurs, obtain a fresh supply.
Oxidizer Prepare 0.02 M iodine in 7:2:1 (v/v/v) THF/pyridine/water. Store ≤6 months at room temperature. CAUTION: Pyridine is toxic in both liquid and vapor forms. CAUTION: Iodine is harmful if inhaled. Beware of spills containing both iodine and ammonia since explosive compounds can be formed. Most commercial sources of THF contain BHT, a free-radical scavenger that prevents the buildup of explosive peroxides. This compound has no effect on synthesis chemistry. Any commercial grade of pyridine is acceptable; use resublimed iodine.
Phosphoramidites Prepare phosphoramidites in extremely dry, commercially sealed acetonitrile (see recipe) according to the procedure recommended by the synthesizer manufacturer. Store ≤6 months at 4°C and discard after 2 weeks. Some companies recommend dissolving phosphoramidites to give equal molarities of the four bases, while others recommend a standard weight/volume ratio that will give slightly different final molar concentrations. The actual phosphoramidite concentration may matter for some applications—e.g., when making mixed-site oligonucleotides.
COMMENTARY Introduction to the Synthesis and Purification of Oligonucleotides
Critical Parameters By optimizing the reagents and protocols used in oligonucleotide synthesis, it is reliably
possible to make products of greater length and yield, while minimizing the costs associated with unproductive runs.
A.3C.18 Current Protocols in Nucleic Acid Chemistry
Excluding water from solvents The most critical factor in any synthesis is how the reagents are handled to exclude water from the system. From the moment a bottle is opened, it is in contact with water in the air, and all the solvents used are hygroscopic and will absorb water vapor, which reduces yields. This problem is so severe that it is advisable to avoid large-scale, lengthy, or important runs on rainy or highly humid days. Special anhydrous reagents can be purchased from most manufacturers of nucleic acid synthesizers. Additionally, chemical suppliers are now beginning to market solvents specifically for DNA synthesis. Adding molecular sieving pouches to the activator and acetonitrile used before oxidation is extremely useful in preserving the anhydrous environment of the phosphoramidite coupling reaction. Phosphoramidites are the most sensitive to water contamination because they are easily hydrolyzed, which renders them unreactive. Precautions must be taken to avoid exposing them to even small amounts of water. Very dry acetonitrile (<0.003% water), kept as a separate stock and sealed under argon, should be used to dissolve the phosphoramidites. The acetonitrile should be introduced through the septum on the amidite bottles via a syringe. Glass syringes can be dried in a 300°C drying oven, then cooled in a desiccator prior to use. Plastic syringes can be dried in a 45°C vacuum oven. Disposable plastic syringes from air-tight sterile casings are dry enough to use with acetonitrile to be dispensed into phosphoramidites with no additional precautions. The syringe should be filled with argon or helium prior to drawing up the acetonitrile, so that wet air is not introduced into the system. On some synthesizers, this can be done directly via the phosphoramidite ports. Otherwise, an argon line should be used. An argon line is generally a helpful tool for DNA synthesis chemistry. It consists of an argon tank with a regulator connected to a piece of plastic (Tygon) tubing. The tubing is then fitted with either a Pasteur pipet or a syringe and needle. Gas flow can be roughly determined via an in-line “bubbler” (e.g., Aldrich). Argon from a tank is dry enough so that an in-line desiccator is unnecessary. It is necessary to flush the line for several seconds prior to use. Empty bottles can be dried under an argon stream, which will help to exclude condensation from the air. Partially used reagents should be sealed under an argon layer to prevent equilibration with water vapor in the air (the heavier argon will exclude air from the containers).
Choosing synthesis columns Automated DNA synthesis generally takes place on a solid support made of controlledpore glass (CPG; see UNITS 3.1 & 3.2). This is a porous, nonswelling particle, 125 to 177 µm in diameter, that is derivatized with a terminal nucleotide attached to a long spacer arm. The accessibility of the growing end of the oligonucleotide chain is influenced by the pore size of the particle. It has been recommended that oligonucleotides up to 50 bases in length should be synthesized on CPG with ∼500-Å pores, while longer oligonucleotides should be synthesized on CPG with ∼1000-Å or larger pores. Be sure that the column geometry is compatible with the trityl monitor. Also, when packing columns be sure to purge the column first with dry acetonitrile and then with dry argon to ensure that all CPG particles are caught between the filters, since loose particles can damage the synthesizer. CPG columns are available that contain ∼0.2 to 10.0 µmol of linked nucleotides. In general larger loadings are used for larger oligonucleotides since as the size of the oligonucleotide increases, the overall yield decreases. Also, subsequent purification steps invariably involve losses, and the amount of product of correct length must be kept high to ensure that there is enough material. Oligonucleotides >35 bases should be synthesized on 1.0-µmol CPG columns. See Synthesizing Long Oligonucleotides for discussion of choosing a column when synthesizing ≥100-base oligonucleotides. Alternatively, columns that are more heavily derivatized with the 3′ terminal nucleotide are available as solid supports (e.g., Fractosil, from Merck), and a higher yield of product per column (and per amount of reagent delivered) can be obtained using these columns. However, Fractosil is not recommended for synthesizing oligonucleotides >20 bases long. Synthesis chemistry Regardless of reagent and final product purity, there are inherent limitations in the chemistry used for the synthesis of oligonucleotides. Therefore, it is necessary to understand the differences between chemically synthesized and biologically derived DNA. Failure to produce “natural” DNA can be due to the synthesis chemistry used. Methylation of some bases can occur during deprotection with thiophenol in methyl phosphoramidite–based syntheses. Under standard synthesis conditions, these methylated bases can
Commonly Used Techniques
A.3C.19 Current Protocols in Nucleic Acid Chemistry
account for up to 7% of the nucleotides present, with N-3-methyl-dT being the primary modified base (Farrance et al., 1989). However, this problem does not always occur—some such syntheses have been reported to contain 99.9% dA, dG, dT, and dC. In general, this problem can be avoided by increasing the time of the thiophenol deprotection step to between 60 and 90 min. With both methyl and β-cyanoethyl chemistries, the glycosidic bond of N-protected purine phosphoramidites is subject to hydrolysis during DMT removal with acid. Such depurination eventually leads to strand cleavage during the ammonia deprotection step. N-protected adenosine is more sensitive to depurination than guanosine, and it is most sensitive when located at the 5′ end of an oligonucleotide chain (Tanaka and Letsinger, 1982). In order to maintain low levels of depurination, the deblocking step should not be longer than ∼1 min. Additionally, the mildest effective acid (DCA rather than TCA) should be used. Occasionally, however, protected nucleotides are supplied as monomethoxytrityl (MMT) esters, which are 10-fold more resistant to acidic detritylation. Oligonucleotides synthesized with MMT as a protecting group are more susceptible to depurination, since the deblocking step must be correspondingly lengthened to achieve complete detritylation and high yields. In cases where only one base is being added as the MMT compound, it is recommended that this step be performed manually, so that the nucleotides are subjected to only one lengthy acid treatment. Advances in oligonucleotide synthesis chemistry may mitigate the problems associated with depurination. Recently, nucleotides derivatized with protecting groups that render synthesized material less sensitive to depurination have become commercially available (Schulhof et al., 1987; Pharmacia, 1989).
Introduction to the Synthesis and Purification of Oligonucleotides
Reagents Most reagents may be purchased from the synthesizer manufacturer or from companies that specialize in reagents for nucleic acid synthesis; however, some labs choose to make their own reagents to reduce costs. Notes on the preparation and storage of certain reagents are provided above. Other reagents should probably be purchased from commercial sources, as the preparation of anhydrous materials is more difficult and expensive than most molecular biology labs can support. An appropriate text should be consulted on the medical dangers of all solutions and reagents used in DNA synthe-
sis (see Key References). Most suppliers of materials have a fairly extensive list of products available on-line.
Troubleshooting Rescue after reagent depletion Most synthesizer problems require a visit from the service engineer. However, syntheses during which reagent or solvent bottles run empty can sometimes be rescued. Certain instruments respond to depleted reagents during a synthesis by continuing the synthesis or stopping. For instruments that stop, it may be possible to recover oligonucleotides, depending on which reservoir was depleted. It is important that the lines be rinsed after an empty reservoir has been detected (some machines perform this step automatically). Otherwise, the oligonucleotide should be resynthesized. Many machines have the capacity to restart in the middle of a cycle, and if not, the cycle can be finished manually. The exact procedure will vary depending on which reagent has been exhausted. Acetonitrile. This is quite serious. The lines cannot be rinsed of whatever reagent they last contained; thus, determine whether to continue with the synthesis based on the reactivity of the last reagent in the lines. Deblocking reagent. Refill the reservoir. The machine can continue at the beginning of the cycle that was interrupted, but full-length yield will greatly suffer. Phosphoramidites. Refill the reservoir and perform manual coupling. In this way, the chains that did elongate will not form n + 1 products, and those that did not will have a chance to elongate. Programmed synthesis can be resumed with the next cycle. Capping reagent. Perform a manual capping and complete the cycle manually, at which point programmed synthesis can resume. This ensures that capping is efficient for this cycle and avoids accumulation of n − 1 sequences. Oxidizer. Running out of oxidizer is particularly dangerous, because the unstable phosphite linkages remain on the column for long periods. It is best to discard the material; otherwise, perform a manual oxidation and continue with the synthesis. Ammonium hydroxide. Some material may be left on the column if ammonium hydroxide runs out during the course of synthesis. Refill the bottle and pump more ammonium hydroxide through the column, or remove the column and treat with ammonium hydroxide.
A.3C.20 Current Protocols in Nucleic Acid Chemistry
Thiophenol. Refill the thiophenol reservoir. Restart from the deprotection step. At worst, a small amount of product may have cleaved because of inefficient methyl deprotection. Routine maintenance To avoid other purely mechanical problems with a DNA synthesizer, create a regular maintenance schedule to perform a few minor maintenance tasks. Change the various frits and filters that remove debris before they enter the system; furthermore, remove the deposits of salt and debris as these may affect flow rates and decrease volumes delivered. Change the filters on the acetonitrile bottle more often, because the flow of this reagent is much greater than any other. Also, change any rubber O rings every few months. Rinse all the instrument’s lines with base and organic solvents thoroughly every 500 hr of machine use or approximately every 3 months. Finally, rinse the lines extensively with dry acetonitrile. Finish the general maintenance by checking the flow rates of the instrument after cleaning and then synthesize a small oligonucleotide. Monitor the trityls carefully in order to confirm that the cleaning did not affect any aspect of the synthesis cycle. If this regimen is followed, many minor delays encountered in the synthesis schedule can be avoided.
Anticipated Results When the synthesizer is working properly (>98% coupling efficiency), typical yields of deoxyoligonucleotides 20 bases in length at the 1-µmol scale, as measured from the trityl response, are nearly 60%. Also, these shorter deoxyoligonucelotides are typically suitable for DNA sequencing, PCR amplification, and gel-shift analysis without extensive purification. However, if very homogenous material is required, further purification must be done, because many sequences will be truncated. Longer DNA sequences are generally obtained in lesser yield and quality and should be purified. Sequences of >120 bases are generally obtained in ≤10% yield as measured by the trityl response. Yields for RNA sequences are typically much lower than for their DNA counterparts of a given length, and the products must be extensively purified.
Time Considerations On a typical day when most of the bottles on the synthesizer need refilling, it may take 30 to 60 min to fill the bottles, empty waste, install columns, and rinse and prime the lines prior to starting a synthesis. The most time-consuming
syntheses are those involving doped oligonucleotides, which require a high level of precision in distributing the various phosphoramidites to the appropriate bottles. A computer interfaced with the instrument can increase synthesis accuracy and throughput. Instruments take varying amounts of time to synthesize oligonucleotides depending on the number of columns in use, the length of the oligonucleotide, and the synthesis program. Deprotection and isolation of a standard DNA oligonucleotide will probably take one working day, while RNA oligonucleotides take longer because of the extra deprotection and desalting steps.
Literature Cited Applied Biosystems. 1988. User Bulletin no. 13. Applied Biosystems, Foster City, Calif. Bartel, D. and Szostak, J.W. 1993. Isolation of new ribozymes from a pool of random sequences. Science 261:1411-1418. Boal, J.H., Wilk, A., Harindranath, N., Max, E.E., Kempe, T., and Beaucage, S.L. 1996. Cleavage of oligodeoxyribonucleotides from controlledpore glass supports and their rapid deprotection by gaseous amines. Nucl. Acids Res. 24:31153117. Ciccarelli, R.B., Gunyuzlu, P., Huang, J., Scott, C., and Oakes, F.T. 1991. Construction of synthetic genes using PCR after automated DNA synthesis of their entire top and bottom strands. Nucl. Acids Res. 19:6007-6013. Cohen, G., Deutsch, J., Fineberg, J., and Levine, A. 1997. Covalent attachment of DNA oligonucleotides to glass. Nucl. Acids Res. 25:911-912. Derbyshire, K.M., Salvo, J.J., and Grindley, N. 1986. A simple and efficient procedure for saturation mutagenesis using mixed oligodeoxynucleotides. Gene 46:145-152. Eadie, J.S., McBride, L.J., Efcavitch, J.W., Hoff, L.B., and Cathcart, R. 1987. High-performance liquid chromatographic analysis of oligodeoxyribonucleotide base composition. Anal. Biochem. 165:442-447. Farrance, I.K., Eadie, J.S., and Ivarie, R. 1989. Improved chemistry for oligodeoxyribonucleotide synthesis substantially improves restriction enzyme cleavage of a synthetic 35mer. Nucl. Acids Res. 17:1231-1245. Gait, M.J. (ed.). 1984. Oligonucleotide Synthesis: A Practical Approach. IRL Press, Washington, D.C. Gillam, S. and Smith, M. 1980. Use of E. coli polynucleotide phosphorylase for the synthesis of oligodeoxyribonucleotides of defined sequence. Methods Enzymol. 65:687-701. Hermes, J.D., Parekh, S.M., Blacklow, S.C., Kuster, H., and Knowles, J.R. 1989. A reliable method for random mutagenesis: The generation of mutant libraries using spiked deoxyribonucleotide primers. Gene 184:143-151.
Commonly Used Techniques
A.3C.21 Current Protocols in Nucleic Acid Chemistry
Horn, T. and Urdea, M.S. 1988. Solid supported hydrolysis of apurinic sites in synthetic oligonucleotides for rapid and efficient purification on reverse-phase cartridges. Nucl. Acids Res. 16:11559-11571.
lated ribonucleotide 3′-O-phosphoramidites on a controlled-pore glass support: Synthesis of a 43nucleotide sequence similar to the 3′ half molecule of an E. coli formylmethionine tRNA. J. Am. Chem. Soc. 109:7845-7854.
Kawahara, S., Wada. T., and Sekine, M. 1996. Unprecedented mild acid-catalyzed desilyation of the 2-O-tert-butyldimethylsilyl group from chemically synthesized oligoribonucleotides intermediates via neighboring group participation of the internucleotide phosphate residue. J. Am. Chem. Soc. 118:9461-9468.
Vargeese, C., Carter, J., Yegge, J., Karivjansky, S., Settle, A., Kropp, E., Peterson, K., and Peiken, W. 1998. Efficient activation of nucleoside phosphoramidites with 4,5-dicyanoimidazole during oligonucleotide synthesis. Nucl. Acids Res. 26:1046-1050.
Kinoshita, Y., Nishigaki K., and Husimi Y. 1997. Fluorescence-, isotope- or biotin-labeling of the 5′-end of single-stranded DNA/RNA using T4 RNA ligase. Nucl. Acids Res. 25:3747-3748. Oliphant, R. 1989. Functional Sequences from Random DNA. Harvard University Thesis, Boston, Mass. Oliphant, R., Nussbaum, A.L., and Struhl, K. 1986. Cloning of random-sequence oligodeoxynucleotides. Gene 44:177-183. Pharmacia. 1989. Analects Vol. 17, no. 2. Pharmacia Biotech, Piscataway, NJ. Ratliff, R.L. 1982. Terminal deoxynucleotidyltransferase. In The Enzymes, Vol. XV (P.D. Boyer, ed.) pp. 105-118. Academic Press, San Diego. Reddy, M.P., Hanna, N.B., and Farooqui, F. 1994. Fast cleavage and deprotection of oligonucleotides. Tetrahedron Lett. 35:4311-4314. Reddy, M.P., Farooqui, F., Hanna, N.B. 1995. Methylamine deprotection provides increased yield of oligoribonucleotides. Tetrahedron Lett. 36:8929-8932. Reidhaar-Olson, J.F. and Sauer, R.T. 1988. Combinatorial cassette mutagenesis as a probe of the informational content of protein sequences. Science 241:53-57. Schulhof, J.C., Molko, D., and Teoule, R. 1987. The final deprotection step in oligonucleotide synthesis is reduced to a mild and rapid ammonia treatment by using labile base-protecting groups. Nucl. Acids Res. 15:397. Song, Q. and Jones, R.A. 1999. Use of silyl-ethers as fluoride scavengers in RNA synthesis. Tetrahedron Lett. 40:4653-4654. Sproat, B., Colonna, F., Mullah, B., Tsou, D., Andrus, A., Hampel, A., and Vinayak, R. 1995. An efficient method for the isolation and purification of oligoribonucleotides. Nucleosides Nucleotides 14:255-273. Szostak, J. 1992. In vitro genetics. Trends Biochem. Sci. 17:89-93. Tanaka, T. and Letsinger, R.L. 1982. Syringe method for stepwise chemical synthesis of oligonucleotides. Nucl. Acids Res. 10:3249.
Introduction to the Synthesis and Purification of Oligonucleotides
Uhlenbeck, O.C. and Gumport, R.I. 1982. T4 RNA ligase. In The Enzymes, Vol. XV (P.D. Boyer, ed.) pp. 31-58. Academic Press, San Diego. Usman, N., Ogilvie, K.K., Jiang, M.Y., and Cederagren, R.J. 1987. Automated chemical synthesis of long oligoribonucleotides using 2′-O-sily-
Warren, W.J. and Vella, G. 1994. Analysis and purification of synthetic oligonucleotides by highperformance liquid chromatography. Methods Mol. Biol. 233-264. Wincott, F., DiRenzo, A., Shaffer, C., Grimm, S., Tracz, D., Workman, C., Sweedler, D., Gonzalez, C., Scaringe, S., and Usman, N. 1995. Synthesis, deprotection, analysis and purification of RNA and ribozymes. Nucl. Acids Res. 23:2677-2684. Zhu, Y., He, L., Srinivasan, R., and Lubman, D. 1997. Improved resolution in the detection of oligonucleotides up to 60-mers in matrix-assisted laser desorption/ionization time-of-flight mass spectrometry using pulsed-delayed extraction with a simple high voltage transistor switch. Rapid Commun. Mass Spectrom. 11:987-992. Zon, G., Gallo, K.A., Samson, C.J., Shao, K., Summers, M.F., and Byrd, R.A. 1985. Analytical studies of “mixed sequence” oligodeoxyribonucleotides synthesized by competitive coupling of either methyl or β-cyanoethyl-N,N-diisopropylamino phosphoramidite reagents, including 2′deoxyinosine. Nucl. Acids Res. 13:8181-8196.
Key References Applied Biosystems, 1988. See above. A well-organized overview of synthetic oligonucleotide synthesis, purification, and quantitation. Bretherick, L.,1986. Hazards in the Chemical Laboratory, 4th ed. Alden Press, Oxford. A guide to hazardous chemical handling. Gait, 1984. See above. The seminal text on synthetic oligonucleotide synthesis that provides critical insight.
Internet Resources http://www.interactiva.de/ Web site detailing synthesis chemistries, procedures, and reagents for solid-phase oligonucleotide chemistry.
Contributed by Andrew Ellington University of Texas Austin, Texas Jack D. Pollard, Jr. Harvard Medical School and Massachusetts General Hospital Boston, Massachusetts
A.3C.22 Current Protocols in Nucleic Acid Chemistry
Thin-Layer Chromatography
APPENDIX 3D
Thin-layer chromatography (TLC) is used as a tool for the determination of compound identity and purity. Most commonly, this technique is used to monitor the progress of a chemical reaction or to assay fractions collected from a larger chromatographic separation (e.g., column chromatography). TLC is typically performed using commercially available glass plates which are coated with a thin layer of adsorbent (TLC plates). Silica gel and alumina are polar adsorbents commonly used to separate compounds of low to medium polarity. Highly polar compounds will “stick” to polar adsorbents and may be purified alternatively using C18 reversed-phase TLC plates (glass plates coated with a highly lipophilic adsorbent). In reversed-phase separations, nonpolar compounds will “stick” to the adsorbent, making it possible to separate highly polar compounds. Regardless of the adsorbent used, TLC can be used to run test separations in preparation for column chromatography. This unit describes a method for spotting test compound onto a TLC plate and developing the plate in a suitable solvent system, as well as the various methods for visualizing the results, which are then used to calculate retention factor (Rf) values (see Basic Protocol). The Alternate Protocol describes co-spotting the plate with candidate compounds in order to identify the unknown sample without relying on Rf values. Instructions for cutting TLC plates are given in Support Protocol 1, and a procedure for preparing spotters is given in Support Protocol 2. THIN-LAYER CHROMATOGRAPHY The preparation of a TLC plate is accomplished by dissolving the mixture of compounds to be separated in a suitable solvent (e.g., acetone, chloroform, ethyl acetate), spotting the mixture onto a TLC plate, and allowing the solvent to evaporate. The TLC plate is then developed in a TLC chamber containing a suitable solvent system (Fig. A.3D.1). The solvent in the chamber is drawn upward onto the plate by capillary action and “carries” along the compounds present in the mixture. Separation of compounds occurs because each compound is retained differently on the adsorbent. When the solvent has eluted to the top of the plate, the TLC plate is removed from the chamber. Only colored compounds may be visualized by the naked eye following development of the TLC plate; as many compounds are not colored, an alternative method for visualizing the plate is necessary. There are numerous methods for visualizing TLC plates (see Zweig and Sherma, 1972, for more information). Three different methods: destructive, semidestructive, and nondestructive, are described here.
BASIC PROTOCOL
Following visualization, the retention factor of each spot is determined. The retention factor, or Rf value, measures how strongly each compound is retained on the adsorbent in a particular solvent system. Compounds with large Rf values migrate higher up the plate than compounds with small Rf values. Materials Test compound Appropriate test compound volatile solvent Appropriate solvent system (eluent) 50% to 98% sulfuric acid (destructive visualization) Solid iodine (semidestructive visualization) Thin-layer chromatography (TLC) plate, cut to size (see Support Protocol 1) Contributed by C.L.F. Meyers Current Protocols in Nucleic Acid Chemistry (2000) A.3D.1-A.3D.8 Copyright © 2000 by John Wiley & Sons, Inc.
Synthesis of Unmodified Oligonucleotides
A.3D.1 Supplement 3
Figure A.3D.1
Diagram of a TLC developing chamber.
TLC spotter (see Support Protocol 2) Wide-mouthed TLC chamber or beaker with lid (Fig. A.3D.1) Filter paper 110°C oven (destructive visualization) Hand-held UV light source (nondestructive visualization) Spot a TLC plate 1. Using a pencil, draw a faint line at the edge of the plate ∼1 cm from the bottom of a thin-layer chromatography (TLC) plate cut to an appropriate size. This is the baseline of the TLC plate. If spotting more than one compound along the bottom of the TLC plate, it may be helpful to make the appropriate number of notches (very faintly) on the baseline (~1 spot every 5 mm) to mark the position of each spot (Fig. A.3D.2). IMPORTANT NOTE: Do not draw heavily on the TLC plate or the adsorbent will be scratched away.
2. Dissolve the mixture of compounds to be analyzed in a volatile solvent (e.g., acetone, ethyl acetate, dichloromethane). If using a C18 reversed-phase TLC plate, the compound may dissolve only in water/organic solvent mixtures. Dissolve the compound in as little water as possible.
3. Draw a small amount of the dissolved mixture up into the tip of a TLC spotter and briefly touch the tip of the spotter onto the baseline of the TLC plate in the appropriate position to deposit a small amount of the sample in the form of a “spot.” Make the spot as small as possible (typically 2 to 3 mm). Thin-Layer Chromatography
IMPORTANT NOTE: The longer the tip of the spotter is held on the plate, the larger the spot will be.
A.3D.2 Supplement 3
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Figure A.3D.2 Spotting a TLC plate. The baseline is ~1 cm from the bottom and spots are ~5 mm apart.
4. Let the solvent in the spot evaporate and repeat the procedure (step 3) in the same place on the TLC plate in order to accumulate a sufficient amount of the compound at the baseline of the plate. IMPORTANT NOTE: Be careful not to make the spots too concentrated, as mixtures of compounds that have been overloaded onto the TLC plate will appear “streaky” and unresolved when developed. Compounds dissolved in water/organic solvent mixtures will take longer to dry. If the compound absorbs UV light, the plate can be visualized by UV absorption to determine whether enough compound has been loaded onto the plate (see step 9c).
5. Repeat step 4 along the baseline of the TLC plate until the desired number of mixtures to be analyzed have been spotted. IMPORTANT NOTE: Do not spot mixtures too closely together along the baseline. Overlapping spots will “bleed” into each other’s path of migration, making it difficult to identify the origin of each spot (see step 1).
Develop plate 6. Choose an appropriate solvent system (eluent) to develop the TLC plate. Pour the chosen eluent into a wide-mouthed TLC chamber or beaker to a depth of no more than 5 mm, and cover the chamber to prevent evaporation. If necessary, line the inside edges of the chamber with filter paper in order to saturate the chamber atmosphere with the eluent vapors and prevent solvent evaporation. Choosing an eluent is often more complicated than it sounds. Each separation carried out on a TLC plate will require a different solvent system and will depend upon the adsorbent chosen for the separation. In general, organic solvent systems are used for separations carried out on silica gel or alumina, while water/acetonitrile or water/alcohol solvent mixtures are used for separations carried out on C18 reversed-phase medium. Refer to the CRC Handbook of Chromatography (Zweig and Sherma, 1972) for guidance in choosing a solvent system for a given separation.
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7. Place the TLC plate into the chamber, making sure the baseline is above the level of the eluent in the chamber. Cover the chamber and allow the solvent to travel upward on the TLC plate (Fig. A.3D.1). Development of silica gel or alumina TLC plates may take only a few minutes, whereas the development of a C18 reversed-phase TLC plate may take considerably longer. Aqueous solvent systems elute up the TLC plate much more slowly than organic solvent systems.
8. When the eluent nears the top of the plate, remove the plate from the chamber, mark the position of the solvent front, and allow the eluent to evaporate in a fume hood. CAUTION: Do not breathe solvent vapors. C18 reversed-phase TLC plates developed with aqueous solvent systems will take several minutes to dry.
Visualize TLC plate 9a. Destructive visualization: Spray the TLC plate with 50% to 98% sulfuric acid, then heat the plate in a 110°C oven for several minutes. This method is termed destructive visualization because any compound present on the plate will decompose to give a dark spot. This is not the only destructive visualization method available. There are numerous “sprays” and “dips” that can be used to visualize compounds containing certain functional groups (Zweig and Sherma, 1972).
9b. Semidestructive visualization: Assemble a wide-mouthed, covered chamber containing a few crystals of iodine. Place the TLC plate inside the chamber and let it stand for several minutes. Remove the plate and circle the colored spots with a pencil. Most, but not all, compounds present on the plate will absorb iodine vapors and change color. Over time, the spots will disappear.
9c. Nondestructive visualization: Shine either long-wave UV light (dark spots against a green background) or short-wave UV light (glowing spots against a dark background) with a hand-held UV light source to visualize compounds. CAUTION: UV light is damaging to the eyes and exposed skin. Protective eyewear should be worn at all times while using a UV light source. UV light can be used to visualize certain compounds on TLC plates. This method is termed nondestructive visualization because non-photolabile compounds generally do not decompose under these conditions. Commercially available TLC plates contain a fluorescent additive that glows bright green when the plate is placed under long-wave UV light. Compounds containing a UV-absorbing chromophore are termed UV-active and can be visualized using this method. In general, these compounds will appear as dark spots on a fluorescent green background when viewed under long-wave UV light. When viewed under short-wave UV light, the plates remain dark and the compounds glow. In either case, visible spots should be circled with a pencil for future reference. Hand-held UV lamps with both long-wave and short-wave UV light are available commercially for this purpose.
Determine Rf value 10. Measure the distance from the baseline to the center of the spot, which is the distance the spot traveled on the TLC plate. 11. Measure the distance from the baseline to the solvent front, which is the distance the solvent traveled on the TLC plate. Thin-Layer Chromatography
12. Divide the distance the spot traveled (step 10) by the distance the solvent traveled (step 11). This ratio is the Rf value and should be in the range of 0.0 to 1.0.
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A
B
C
Figure A.3D.3 Results of TLC. (A) The spot has a Rf of 0.0. For silica gel adsorbent, this means that the eluent is not polar enough. For C18 reversed-phase adsorbent, this means that eluent is too polar (water content too high). (B) The spot has a Rf of 0.94. For silica gel adsorbent, this means that the eluent is too polar. For C18 reversed-phase adsorbent, this means that the eluent is not polar enough (organic content too high). (C) The mixture actually contains two compounds, as is made apparent when the TLC plate is developed using a suitable eluent. The spots have Rf values of 0.29 and 0.67, respectively.
Rf value = (distance the spot traveled)/(distance the solvent traveled) = 0.0 to 1.0
13. Evaluate the Rf value. Values less than 0.2 or greater than 0.8 are not particularly informative. These spots may contain mixtures of compounds that were not separated during the development of the plate (Fig. A.3D.3). Spots with Rf values of <0.2 or >0.8 should be reevaluated by adjusting the polarity of the eluent to obtain Rf values within the range of 0.2 to 0.8.
CO-SPOTTING A TLC PLATE Although the Rf value of a particular spot may give some information about the identity of a compound, it is not always exactly reproducible. For example, an Rf value may vary depending upon the amount of sample spotted on the plate. For this reason, an Rf value alone cannot be used as a certain indication of compound identity. However, the identity of a spot can be determined by co-spotting the unknown compound being analyzed with standards of what the compound may be, or “authentic” compounds. In this protocol, a method for determining the identity of an unknown compound as either authentic compound A or B is described (Fig. A.3D.4).
ALTERNATE PROTOCOL
Additional Materials (also see Basic Protocol) Unknown compound Authentic compounds A and B 2 × 5–cm TLC plate (see Support Protocol 1) 1. Cut a TLC plate that is wide enough to hold 4 spots at the baseline (~1 spot every 5 mm; see Support Protocol 1). The first three positions will be used to spot each compound by itself. The fourth position will be used to spot all three compounds together.
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Figure A.3D.4 Co-spotting a TLC plate (see Alternate Protocol). Lane (1) unknown compound. Lane (2) authentic compound A. Lane (3) authentic compound B. Lane (4) mixture of authentic compounds A and B and unknown compound. Notice that the unknown (lane 1) and authentic compound B (lane 3) have similar Rf values (0.44 and 0.40, respectively) when run side-by-side on the TLC plate. Upon closer examination of lane 4, it is shown that, in fact, the unknown and authentic compound B are identical.
2. Spot the unknown compound at the first and fourth positions on the TLC plate. Allow the solvent to evaporate completely. 3. Spot authentic compound A in the second and fourth positions. Again, allow the solvent to evaporate completely. 4. Spot authentic compound B at the third and fourth positions. The TLC plate should be spotted as in Figure A.3D.4.
5. Develop the TLC plate using a suitable eluent chosen previously for the unknown (see Basic Protocol, steps 6 to 8). 6. Visualize the developed TLC plate (see Basic Protocol, step 9 to 13). The TLC plate should demonstrate the results shown in Figure A.3D.4. Notice that the unknown and authentic compound B have similar Rf values when run side-by-side on the TLC plate. Upon closer examination of the fourth position, it is shown that, in fact, the unknown and authentic compound B are identical. SUPPORT PROTOCOL 1
Thin-Layer Chromatography
CUTTING TLC PLATES Precoated TLC plates (10 × 20 cm) are commercially available with a variety of adsorbents. Before performing TLC, the larger glass plates need to be cut down to a workable size. Prescored TLC plates are commercially available and can simply be broken at the right positions. Alternatively, cutting can be done manually using a cutting tool, as described here. This protocol produces approximately twenty 2 × 5–cm plates. Longer plates may be needed for applications with longer elution times, which are generally required for more difficult separations. The width of the plate can be adjusted to accommodate the number of spots that are to be loaded and the size of the chamber used to develop the plate. Materials 10 × 20–cm TLC plates Glass cutter: diamond cutters preferred
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1. Place a 10 × 20–cm TLC plate adsorbent-side-down on a large piece of filter paper or paper towel. IMPORTANT NOTE: Do not use printed paper as it contains dyes and chemicals that may be transferred to the adsorbent on the plate. IMPORTANT NOTE: When handling precoated glass plates, always wear gloves or be sure to handle with clean hands. Dirt or oil from the hands can be transferred to the adsorbent.
2. To guide the cut, hold a ruler down the center of the plate lengthwise and score the plate using a glass cutter (diamond cutters work the best). Break the plate along the score, producing two ~5 × 20 cm plates. CAUTION: Cut TLC plates are very sharp. IMPORTANT NOTE: C18 reversed-phase TLC plates may “flake” when broken along the scored line.
3. Place each plate adsorbent-side-down on the filter paper. Holding the ruler as a guide, make a series of scores across each plate, ∼2 cm apart. Break the plates along the scores to make approximately twenty 2 × 5–cm TLC plates. TLC plates of this size should be appropriate for 4 to 5 spots.
PREPARATION OF TLC SPOTTERS The spotter is the apparatus used to load the mixture of compounds to be separated onto the TLC plate. It is appropriately named because it makes a “spot” on the plate. They are easily prepared from capillary tubes, as described here. Alternatively, commercially available, disposable micropipets can be used as spotters. These are sold in packages of 100 and are available from most scientific supply companies (e.g., 1 µL; Drummond).
SUPPORT PROTOCOL 2
Materials ~1 × 100–mm capillary tubes, open-ended 1. Put the center of the open-ended capillary tube into a small, blue Bunsen burner flame. Hold it in the flame until the center of the capillary tube begins to liquefy. IMPORTANT NOTE: Do not leave the capillary tube in the flame too long or it will melt the edges of the tube together in the center.
2. Immediately remove the capillary tube from the flame and pull the ends in opposite directions (Fig. A.3D.5). 3. Break the capillary tube in the center to make two spotters. Several spotters may be needed for loading different compounds onto the TLC plate. It is good practice to use a different spotter for each sample; however, in some cases it may be necessary only to “wash” the spotter before using it again (e.g., for analyzing fractions collected during column chromatography). To wash a spotter, simply draw a suitable solvent (e.g., acetone) up into the thin end (tip) of the spotter. Release the acetone onto a paper towel by touching the tip of the spotter to the towel. Repeat several times to ensure that all of the previous sample has been removed from the spotter. Synthesis of Unmodified Oligonucleotides
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Figure A.3D.5 Preparation of TLC spotters.
LITERATURE CITED Zweig, G. and Sherma, J. 1972. CRC Handbook of Chromatography, Volumes I and II. CRC Press, Cleveland, Ohio.
Contributed by C.L.F. Meyers Purdue University West Lafayette, Indiana
Thin-Layer Chromatography
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Column Chromatography
APPENDIX 3E
Column chromatography is a technique in which a glass column filled with adsorbent (i.e., silica gel, alumina, or C18 reversed-phase packing) is used for the purification of large quantities (milligrams to grams) of reaction product mixtures. As with thin-layer chromatography (TLC; APPENDIX 3D), the adsorbent used varies depending upon the type of separation to be carried out. In any case, it is often useful to analyze the mixture by thin-layer chromatography prior to column chromatography. For example, if suitable conditions using silica gel TLC plates are found, then the silica gel should be used as the adsorbent for the column chromatography. Separations carried out using C18 reversedphase adsorbent are time-consuming because elution of water-containing solvent systems is generally slow. For these types of separations, it is best to use a medium- or high-pressure chromatography system (MPLC or HPLC). Consequently, this discussion will focus only on those separations carried out using silica gel or alumina. After the column has been packed with the appropriate adsorbent and eluent (see Support Protocol), the crude mixture of compounds to be separated is loaded onto the top of the adsorbent layer, and then flushed downward through the column (see Basic Protocol). The procedure can be carried out using gravity to pull the solvent down the column, or by forcing the solvent through the column using a technique known as flash chromatography. Microscale chromatography is also possible (see Alternate Protocol). As with TLC, each compound “sticks” to the adsorbent differently. Thus, the mixture is separated as the solvent elutes through the column. The individual compounds exit the end of the column and are collected in separate tubes or flasks, analyzed by TLC, and isolated. Generally, purifications of >95% can be achieved. COLUMN CHROMATOGRAPHY After the column is packed (see Support Protocol), it is time to load the crude mixture of compounds to be separated onto the top layer of the column. If a gradient eluent is being used to separate the mixture, dissolve the crude mixture in the least polar solvent used in the gradient, using as little solvent as possible. If the crude mixture will not dissolve completely in the chosen solvent system, it may be possible to dissolve the mixture in a different, more suitable solvent (consult someone skilled in this technique). Generally, loading the crude mixture onto the column using solvents other than the chosen eluent should be avoided, as it may affect the separation.
BASIC PROTOCOL
Materials Crude mixture Eluent Chromatography column, packed (see Support Protocol) Flash chromatography apparatus (optional): 250 mL solvent reservoir, 35/20 spherical joint with appropriate flow controller (Chemglass), and air source Additional reagents and equipment for thin-layer chromatography (TLC; APPENDIX 3D) Load column 1. Dissolve the crude mixture of compounds in the chosen eluent. Use a Pasteur pipet to load the mixture onto a packed chromatography column. With the stopcock closed, let the mixture run slowly down the inner side of the column, continually moving the Synthesis of Unmodified Oligonucleotides Contributed by C.L.F. Meyers Current Protocols in Nucleic Acid Chemistry (2000) A.3E.1-A.3E.7
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pipet tip around the inner circumference of the column as the mixture is released from the pipet. IMPORTANT NOTE: Be careful not to disturb the top layer of sand. Moving the pipet tip will ensure that the mixture is evenly deposited on the top of the column.
2. Open the stopcock and allow the solvent to drain into a flask until the upper level of solvent reaches the top layer of sand. IMPORTANT NOTE: Do not let the solvent level run below the top layer of sand.
3. Using a Pasteur pipet, rinse the sides of the column with a small amount of eluent (use as little solvent as necessary). Again, drain the solvent until the upper level of solvent reaches the top layer of sand. Assemble apparatus 4a. Conventional chromatography: Fill the column with the chosen eluent. IMPORTANT NOTE: Be careful not to disturb the top layer of sand.
4b. Flash chromatography: Attach the solvent reservoir to the top of the column using the appropriate clamp. Attach a compatible flow controller to the top of the solvent reservoir (Fig. A.3E.1). The use of a solvent reservoir is optional for this procedure. The flow controller is equipped with a hose that can be hooked up to a source of air or nitrogen. For most separations, the laboratory air hose is sufficient for supplying the appropriate amount of pressure. Air pumps can also be used to supply gas pressure.
Column Chromatography
Figure A.3E.1 Professional flash chromatography columns. (A) Conventional. (B) Screw-thread. Reprinted from Mayo et al. (1994) with permission from John Wiley & Sons.
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5. Assemble test tubes and rack for fraction collection from the column. Separate compounds and collect fractions 6a. Conventional chromatography: Open the stopcock and allow the solvent to elute by gravity from the bottom of the column. 6b. Flash chromatography: Open the stopcock and flow controller completely. Turn on the gas source and adjust the flow controller until the solvent elutes from the bottom of the column at the desired rate. Slower elution (~1 drop per sec) may be necessary for the separation of compounds with similar Rf values (∆Rf < 0.1). Compounds whose Rf values are very different (∆Rf > 0.1 to 0.15) can be separated easily with faster elution (5 to 10 drops per sec).
7. Let the eluent run through the column and collect the different fractions in test tubes. Do not let the solvent run below the top layer of sand. Add more solvent to the column by stopping the air pressure (flash chromatography), closing the stopcock, removing the flow controller (flash chromatography), and adding more solvent to the solvent reservoir. If using a particularly volatile solvent (e.g., diethyl ether or methylene chloride), the solvent may evaporate as it elutes from the column, leaving crystals of compound on the tip of the stopcock. If this occurs, simply dissolve/rinse the crystals into the test tube, and analyze the fraction by TLC.
8. As the solvent elutes down the column, collect fractions in test tubes or small flasks. Analyze each fraction by TLC (APPENDIX 3D) to determine the point at which the desired compound(s) elute from the column, using one TLC plate for 4 to 6 fractions, depending upon the width of the TLC plate and the size of the developing chamber. In most cases, the same spotter can be used during the entire chromatography, as long as it is cleaned after loading each fraction onto the TLC plate. The TLC for each fraction will also give an idea of the purity of the compound. If running a silica gel column, compounds with higher Rf values will elute from the column first. If the compounds are colored, the separation can be observed as it takes place down the column; however, many organic compounds are colorless, and the progress of the chromatography must be followed using TLC.
9. Use the Rf value to obtain a rough estimation of the amount of solvent it will take to elute a particular compound. For example, if a compound has an Rf of 0.5, it will migrate halfway down the column during the elution of one column volume of solvent. A column volume is the volume of solvent equal to the volume of adsorbent. Using the different Rf values of the various compounds that are being separated, the number of column volumes it will take to elute every compound using a particular solvent system can be estimated.
Optimize chromatography 10. Evaluate the need to change the polarity of the eluent. After the first compound has eluted from the silica gel or alumina column, the polarity of the solvent may need to be increased to avoid unnecessary use of large solvent volumes and long separation times. The composition and polarity of the new solvent system will depend upon the type of separation being carried out. TLC can be used as a way to predict which will be the best solvent to use. If it is decided that a gradient solvent system is to be used during the chromatography, the conversion from the initial solvent system to a more polar solvent system should be made gradually. For example, if adjusting the solvent system from 5% methanol in chloroform to 15% methanol in chloroform, the adjustment should be made incrementally. To do this, change the solvent system to 8% methanol in chloroform
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and elute one column volume of the new solvent system. Change the solvent system to 12% methanol in chloroform and, again, elute one column volume of solvent. Finally, change the solvent system to the desired 15% methanol in chloroform and pass enough solvent through the column to elute the desired compound from the column. The number of column volumes eluted during the conversion from one solvent system to another is dependent upon the nature of the separation.
11. To change the eluent, let the old solvent run down to the top sand layer and close the stopcock. Slowly add the new solvent to the top of the column. IMPORTANT NOTE: Be careful not to disturb the sand layer.
12. After analyzing each fraction by TLC, combine fractions containing the same pure compound and strip away the solvent using a rotary evaporator. CAUTION: Do not use a flame to evaporate organic solvents. ALTERNATE PROTOCOL
SUPPORT PROTOCOL
MICROSCALE CHROMATOGRAPHY Flash column chromatography can be carried out on small quantities (milligram amounts) of crude product mixtures using a Pasteur pipet (short tip) as a column. The column is packed (see Support Protocol) and loaded (see Basic Protocol) in the same manner as the large-scale column. Compound elution can be accomplished by pushing the eluent through the column using a pipet bulb or an air hose. PACKING A CHROMATOGRAPHY COLUMN Before carrying out column chromatography, the glass column must be packed with the appropriate adsorbent and a suitable solvent (Fig. A.3E.2). There are two methods for packing a column: wet packing and dry packing. Wet packing can be messy, but this technique works every time and is preferred for packing silica gel columns. Dry packing often results in the formation of air bubbles that can be difficult to remove. When this happens, the column must be repacked; however, when alumina is used as an adsorbent, the preferred method is dry packing, as wet packing with alumina requires making a slurry that is difficult to pour. Often, it is necessary to carry out column chromatography using a gradient eluent. This means that the polarity of the solvent system may need to be changed during the chromatography so that both nonpolar and polar compounds can be eluted from the column. To determine if this is necessary, analyze the mixture by TLC (APPENDIX 3D). If the compounds to be separated have widely different Rf values, more than one solvent system may be necessary in order to elute both compounds adequately. If chromatography using a gradient eluent is to be used, it will be necessary to pack a silica gel or alumina column using the least polar solvent in the gradient. Materials Eluent Adsorbent (e.g., mesh silica gel; Fisher) Glass wool Glass column with ground glass or threaded joint at the top and stopcock at the bottom Fine sand Stiff rubber tube or rubber stopper attached to a handle (e.g., wooden or glass rod)
Column Chromatography
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Figure A.3E.2 Packing a chromatography column.
Prepare column and adsorbent 1. Place a small wad of glass wool just above the closed stopcock in the glass column. Push the plug of glass wool down onto the top of the stopcock using a long rod. A glass column made specifically for column chromatography, or an ordinary buret, can be used as the column. If the plug is too loose, adsorbent will leak through. If the plug is too dense, the eluent will flow very slowly through the column, increasing the time it takes to carry out the column chromatography.
2. Clamp the column vertically to a ring stand inside a hood, or to the grid work inside the hood if possible. Leave enough room at the bottom of the column for a test tube rack or flask. 3. To insure that no adsorbent leaks from the column, add enough fine sand to cover the plug of glass wool. 4. Add the chosen eluent to a height of ∼1 in. (2.56 cm) above the layer of sand. Make sure that the stopcock is closed before adding the eluent. This will prevent a disturbance of the sand layer when the adsorbent is finally added.
5. Weigh out the appropriate amount of adsorbent into a beaker.
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For routine chromatographic separations, ∼40 g of adsorbent per 1 g of crude mixture works well. The exact amount of adsorbent to use will depend upon the degree of separation observed by TLC (Still et al., 1978). Commonly used adsorbents are commercially available from most scientific supply companies (e.g., Fisher). Many adsorbents are available in a variety of grades. The optimum adsorbent grade will depend upon the type of separation and adsorbent being used. Silica gel with a pore size of 230 to 400 mesh will be sufficient for routine chromatographic separations. If necessary, the adsorbent may be activated by heating (i.e., removing water) prior to use. Furthermore, silica gel is weakly acidic, so it may be necessary to deactivate it by treatment with dilute amine prior to and during the separation of weakly basic compounds such as aliphatic amines. For a complete guide to adsorbents and their particular applications, refer to the CRC Handbook of Chromatography (Zweig and Sherma, 1972).
Pour column Wet packing columns: 6a. Add enough of the chosen eluent to a preweighed amount of adsorbent (step 5) to make a slurry that is easy to pour (~5 mL eluent/g adsorbent). Use a spatula to mix the slurry. Be sure that all of the adsorbent is wet and that there are no air bubbles in the slurry. 7a. Slowly pour the slurry down the inner edge of the column. IMPORTANT NOTE: If the pouring is too quick, the layer of sand at the bottom of the column will be disturbed and the sand will mix with the adsorbent.
8a. Rinse the sides of the column with the eluent and allow the adsorbent to settle. Proceed to step 9. Dry packing alumina columns: 6b. Fill the column halfway with the chosen eluent. 7b. Slowly add the dry adsorbent to the column. As the adsorbent is added, gently tap the sides of the column with a stiff rubber tube or a rubber stopper attached to the end of a handle (i.e., glass or wooden rod). Tapping the sides of the column will ensure that the adsorbent settles to the bottom of the column evenly. If the adsorbent is added too quickly, heat may be liberated as the adsorbent absorbs the solvent. If this happens, the solvent may boil and result in a change in the solvent composition of the eluent (the eluent may be a mixture of two or more solvents). If this happens, the column must be repacked.
8b. Make sure there are no air bubbles or cracks in the column after dry packing. Remove air bubbles by tapping the sides of the column with a stiff rubber tube or a rubber stopper attached to the end of a handle until they rise to the top of the adsorbent layer. Some air bubbles are difficult to remove. Columns containing air bubbles or cracks will not be effective for separating mixtures of compounds. If air bubbles cannot be removed, the column should be repacked.
9. After the column has been packed, open the stopcock and allow the eluent to drain through the column into a clean flask. IMPORTANT NOTE: Do not let the solvent drop below the upper level of adsorbent. The eluent that is collected can be reused during the chromatography.
Column Chromatography
10. Tap the sides of the column with a stiff rubber tube or a rubber stopper attached to the end of a handle. This will result in additional settling of the adsorbent, leaving a small amount of solvent on the upper level of adsorbent. Drain this small amount of
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solvent into the flask until it reaches the upper level of adsorbent. Tap the column again and drain the residual solvent until the adsorbent is tightly packed. Tapping the column aids in tightly packing the adsorbant. This technique also ensures that the upper level of adsorbent (the “baseline” of the column) is even.
11. Add a small amount of fine sand to the upper level of adsorbent (∼0.5 cm). Using eluent, rinse down any dry sand sticking to the edges of the column. Tap the sides of the column with a stiff rubber tube or a rubber stopper attached to the end of a handle to ensure that the upper layer of sand is even. If the adsorbent is not tightly packed, the upper level of adsorbent and the top layer of sand may mix. This will produce an uneven “baseline” for the chromatography. If this occurs, the column should be repacked.
LITERATURE CITED Mayo, D.W., Pike, R.M., and Trumper, P.K. 1994. Microscale Organic Laboratory: With Multistep and Multiscale Syntheses, 3rd ed. John Wiley & Sons, New York. Still, C.W., Kahn, M., and Mitra, A. 1978. Rapid Chromatographic technique for preparative separations with moderate resolution. J. Org. Chem. 43:2923-2925. Zweig, G. and Sherma, J. 1972. CRC Handbook of Chromatography, Volumes I and II. CRC Press, Cleveland, Ohio.
Contributed by C.L.F. Meyers Purdue University West Lafayette, Indiana
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RESOURCES
APPENDIX 4
Useful Nucleic Acid Chemistry Web Sites
APPENDIX 4A
Below is a listing of selected internet resources found to be of particular value by the CPNC editorial board; it is not intended to be a complete listing of all web resources cited in this manual. In addition to the sites listed here, be sure to see the Internet Resources sections at the end of selected units for specialized sites pertaining to the contents of those units. Nucleic acid structure Nucleic Acid Database—structural information about nucleic acids http://ndbserver.rutgers.edu/NDB/ndb.html
National Center for Biotechnology (NCBI) database of 3-dimensional macromolecular structures http://www.ncbi.nlm.nih.gov/Structure
Nucleic Acid Nomenclature and Structure—good overview of important structural parameters including the Dickerson movements of bases in sequence-dependent structures (tip, inclination, opening, propeller, buckle, twist, roll, slide, rise, shift, and tilt) http://www.imb-jena.de/ImgLibDoc/nana/IMAGE_NANA.html#watson
IMB Jena Image Library of Biological Macromolecules—additional data on nucleic acid structure http://www.imb-jena.de/IMAGE.html
Overview of DNA and RNA structure http://info.bio.cmu.edu/Courses/BiochemMols/DNA/DNA.html
Protein Data Bank—includes RNA structures http://www.rcsb.org/pdb
Nomenclature IUPAC-IUB Commission on Biochemical Nomenclature (CBN) Abbreviations and Symbols for Nucleic Acids, Polynucleotides, and their Constituents—Recommendations 1970 http://www.chem.qmw.ac.uk/iupac/misc/naabb.html
IUPAC-IUB Joint Commission on Biochemical Nomenclature (JCBN) Abbreviations and Symbols for the Description of Conformations of Polynucleotide Chains—Recommendations 1982 (Also reproduced in this volume; see APPENDIX 1C) http://www.chem.qmw.ac.uk/iupac/misc/pnuc1.html
RNA resources RNA modification database http://medstat.med.utah.edu/RNAmods
Compilation of tRNA sequences and tRNA genes http://www.uni-bayreuth.de/departments/biochemie/trna Resources Current Protocols in Nucleic Acid Chemistry (2000) A.4A.1-A.4A.3 Copyright © 2000 by John Wiley & Sons, Inc.
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The RNA World—large compilation of RNA resources from the Institute of Molecular Biology at Jena. Includes links to databases and web tools for 3-D structure and sequences, as well as links for RNA viruses and a variety of RNA-related software, books, tutorials, and meetings. http://www.imb-jena.de/RNA.html
The RiboWeb Project—three-dimensional models of the E. coli 30S ribosomal subunit and 16S rRNA http://www-smi.stanford.edu/projects/helix/ribo3dmodels/index.html
Ribosomal Database Project http://www.cme.msu.edu/RDP/html/index.html
RNA Secondary Structures—group I introns and 16S and 23S rRNA http://pundit.colorado.edu:8080
RNase P Database http://jwbrown.mbio.ncsu.edu/RNaseP/home.html
Small RNA Database http://mbcr.bcm.tmc.edu/smallRNA/smallrna.html
The RNA Society Homepage http://www.pitt.edu/∼rna1/
Nucleic acid informatics RNA Modeling http://uracil.cmc.uab.edu/RNA-Modeling/
MFOLD WWW server—fold your own sequences on M. Zuker’s server, includes resources such as free energy and enthalpy tables for RNA folding http://www.ibc.wustl.edu/∼zuker/rna/form1.cgi
RNA thermodynamics software and links http://www.ibc.wustl.edu/∼zuker/rna/node3.html
RNADRAW—freeware program for RNA secondary structure calculation and analysis for Microsoft Windows http://mango.mef.ki.se/∼ole/rnadraw/rnadraw.html
MacMolecule (MacOS) and PCMolecule (Windows) for viewing pdb files http://www.molvent.com
Miscellaneous Nucleic Acids Research http://www3.oup.co.uk/nar/
Translational Active Regions Database (TARD) http://benpc.bionet.nsc.ru/SRCG/Translation/index.html Useful Nucleic Acid Chemistry Web Sites
Extinction coefficient calculation for DNA and RNA oligomers http://paris.chem.yale.edu/extinct.html
A.4A.2 Current Protocols in Nucleic Acid Chemistry
Glen Research—extensive collection of material for the synthesis of modified oligonucleotides http://www.glenres.com/
Oligonucleotide Calculator—Tm, MW, and OD http://www.embl-heidelberg.de/∼toldo/JaMBW/3/1/9/
Tm Determination—allows input of salt and oligonucleotide concentration http://alces.med.umn.edu/rawtm.html
Resources
A.4A.3 Current Protocols in Nucleic Acid Chemistry
SELECTED SUPPLIERS OF REAGENTS AND EQUIPMENT Listed below are addresses and phone numbers of commercial suppliers who have been recommended for particular items used in our manuals because: (1) the particular brand has actually been found to be of superior quality, or (2) the item is difficult to find in the marketplace. Consequently, this compilation may not include some important vendors of biological supplies. For comprehensive listings, see Linscott’s Directory of Immunological and Biological Reagents (Santa Rosa, CA), The Biotechnology Directory (Stockton Press, New York), the annual Buyers’ Guide supplement to the journal Bio/Technology, as well as various sites on the Internet.
A.C. Daniels 72-80 Akeman Street Tring, Hertfordshire, HP23 6AJ, UK (44) 1442 826881 FAX: (44) 1442 826880 A.D. Instruments 5111 Nations Crossing Road #8 Suite 2 Charlotte, NC 28217 (704) 522-8415 FAX: (704) 527-5005 http://www.us.endress.com A.J. Buck 11407 Cronhill Drive Owings Mill, MD 21117 (800) 638-8673 FAX: (410) 581-1809 (410) 581-1800 http://www.ajbuck.com A.M. Systems 131 Business Park Loop P.O. Box 850 Carlsborg, WA 98324 (800) 426-1306 FAX: (360) 683-3525 (360) 683-8300 http://www.a-msystems.com Aaron Medical Industries 7100 30th Avenue North St. Petersburg, FL 33710 (727) 384-2323 FAX: (727) 347-9144 http://www.aaronmed.com Abbott Laboratories 100 Abbott Park Road Abbott Park, IL 60064 (800) 323-9100 FAX: (847) 938-7424 http://www.abbott.com ABCO Dealers 55 Church Street Central Plaza Lowell, MA 01852 (800) 462-3326 (978) 459-6101 http://www.lomedco.com/abco.htm Aber Instruments 5 Science Park Aberystwyth, Wales SY23 3AH, UK (44) 1970 636300 FAX: (44) 1970 615455 http://www.aber-instruments.co.uk ABI Biotechnologies See Perkin-Elmer ABI Biotechnology See Apotex
Access Technologies Subsidiary of Norfolk Medical 7350 N. Ridgeway Skokie, IL 60076 (877) 674-7131 FAX: (847) 674-7066 (847) 674-7131 http://www.norfolkaccess.com
Adaptive Biosystems 15 Ribocon Way Progress Park Luton, Bedsfordshire LU4 9UR, UK (44)1 582-597676 FAX: (44)1 582-581495 http://www.adaptive.co.uk
Accurate Chemical and Scientific 300 Shames Drive Westbury, NY 11590 (800) 645-6264 FAX: (516) 997-4948 (516) 333-2221 http://www.accuratechemical.com
Adobe Systems 1585 Charleston Road P.O. Box 7900 Mountain View, CA 94039 (800) 833-6687 FAX: (415) 961-3769 (415) 961-4400 http://www.adobe.com
AccuScan Instruments 5090 Trabue Road Columbus, OH 43228 (800) 822-1344 FAX: (614) 878-3560 (614) 878-6644 http://www.accuscan-usa.com AccuStandard 125 Market Street New Haven, CT 06513 (800) 442-5290 FAX: (877) 786-5287 http://www.accustandard.com Ace Glass 1430 NW Boulevard Vineland, NJ 08360 (800) 223-4524 FAX: (800) 543-6752 (609) 692-3333 ACO Pacific 2604 Read Avenue Belmont, CA 94002 (650) 595-8588 FAX: (650) 591-2891 http://www.acopacific.com Acros Organic See Fisher Scientific Action Scientific P.O. Box 1369 Carolina Beach, NC 28428 (910) 458-0401 FAX: (910) 458-0407 AD Instruments 1949 Landings Drive Mountain View, CA 94043 (888) 965-6040 FAX: (650) 965-9293 (650) 965-9292 http://www.adinstruments.com
Advanced Bioscience Resources 1516 Oak Street, Suite 303 Alameda, CA 94501 (510) 865-5872 FAX: (510) 865-4090 Advanced Biotechnologies 9108 Guilford Road Columbia, MD 21046 (800) 426-0764 FAX: (301) 497-9773 (301) 470-3220 http://www.abionline.com Advanced ChemTech 5609 Fern Valley Road Louisville, KY 40228 (502) 969-0000 http://www.peptide.com Advanced Magnetics See PerSeptive Biosystems Advanced Process Supply See Naz-Dar-KC Chicago Advanced Separation Technologies 37 Leslie Court P.O. Box 297 Whippany, NJ 07981 (973) 428-9080 FAX: (973) 428-0152 http://www.astecusa.com Advanced Targeting Systems 11175-A Flintkote Avenue San Diego, CA 92121 (877) 889-2288 FAX: (858) 642-1989 (858) 642-1988 http://www.ATSbio.com Advent Research Materials Eynsham, Oxford OX29 4JA, UK (44) 1865-884440 FAX: (44) 1865-84460 http://www.advent-rm.com
Advet Industrivagen 24 S-972 54 Lulea, Sweden (46) 0920-211887 FAX: (46) 0920-13773 Aesculap 1000 Gateway Boulevard South San Francisco, CA 94080 (800) 282-9000 http://www.aesculap.com Affinity Chromatography 307 Huntingdon Road Girton, Cambridge CB3 OJX, UK (44) 1223 277192 FAX: (44) 1223 277502 http://www.affinity-chrom.com Affinity Sensors See Labsystems Affinity Sensors Affymetrix 3380 Central Expressway Santa Clara, CA 95051 (408) 731-5000 FAX: (408) 481-0422 (800) 362-2447 http://www.affymetrix.com Agar Scientific 66a Cambridge Road Stansted CM24 8DA, UK (44) 1279-813-519 FAX: (44) 1279-815-106 http://www.agarscientific.com A/G Technology 101 Hampton Avenue Needham, MA 02494 (800) AGT-2535 FAX: (781) 449-5786 (781) 449-5774 http://www.agtech.com Agen Biomedical Limited 11 Durbell Street P.O. Box 391 Acacia Ridge 4110 Brisbane, Australia 61-7-3370-6300 FAX: 61-7-3370-6370 http://www.agen.com Agilent Technologies 395 Page Mill Road P.O. Box 10395 Palo Alto, CA 94306 (650) 752-5000 http://www.agilent.com/chem
Suppliers
1 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Agouron Pharmaceuticals 10350 N. Torrey Pines Road La Jolla, CA 92037 (858) 622-3000 FAX: (858) 622-3298 http://www.agouron.com Agracetus 8520 University Green Middleton, WI 53562 (608) 836-7300 FAX: (608) 836-9710 http://www.monsanto.com AIDS Research and Reference Reagent Program U.S. Department of Health and Human Services 625 Lofstrand Lane Rockville, MD 20850 (301) 340-0245 FAX: (301) 340-9245 http://www.aidsreagent.org AIN Plastics 249 East Sanford Boulevard P.O. Box 151 Mt. Vernon, NY 10550 (914) 668-6800 FAX: (914) 668-8820 http://www.tincna.com Air Products and Chemicals 7201 Hamilton Boulevard Allentown, PA 18195 (800) 345-3148 FAX: (610) 481-4381 (610) 481-6799 http://www.airproducts.com ALA Scientific Instruments 1100 Shames Drive Westbury, NY 11590 (516) 997-5780 FAX: (516) 997-0528 http://www.alascience.com Aladin Enterprises 1255 23rd Avenue San Francisco, CA 94122 (415) 468-0433 FAX: (415) 468-5607 Aladdin Systems 165 Westridge Drive Watsonville, CA 95076 (831) 761-6200 FAX: (831) 761-6206 http://www.aladdinsys.com Alcide 8561 154th Avenue NE Redmond, WA 98052 (800) 543-2133 FAX: (425) 861-0173 (425) 882-2555 http://www.alcide.com Aldrich Chemical P.O. Box 2060 Milwaukee, WI 53201 (800) 558-9160 FAX: (800) 962-9591 (414) 273-3850 FAX: (414) 273-4979 http://www.aldrich.sial.com
Alexis Biochemicals 6181 Cornerstone Court East Suite 103 San Diego, CA 92121 (800) 900-0065 FAX: (858) 658-9224 (858) 658-0065 http://www.alexis-corp.com Alfa Laval Avenue de Ble 5 - Bazellaan 5 BE-1140 Brussels, Belgium 32(2) 728 3811 FAX: 32(2) 728 3917 or 32(2) 728 3985 http://www.alfalaval.com Alice King Chatham Medical Arts 11915-17 Inglewood Avenue Hawthorne, CA 90250 (310) 970-1834 FAX: (310) 970-0121 (310) 970-1063 Allegiance Healthcare 800-964-5227 http://www.allegiance.net Allelix Biopharmaceuticals 6850 Gorway Drive Mississauga, Ontario L4V 1V7 Canada (905) 677-0831 FAX: (905) 677-9595 http://www.allelix.com Allentown Caging Equipment Route 526, P.O. Box 698 Allentown, NJ 08501 (800) 762-CAGE FAX: (609) 259-0449 (609) 259-7951 http://www.acecaging.com Alltech Associates Applied Science Labs 2051 Waukegan Road P.O. Box 23 Deerfield, IL 60015 (800) 255-8324 FAX: (847) 948-1078 (847) 948-8600 http://www.alltechweb.com Alomone Labs HaMarpeh 5 P.O. Box 4287 Jerusalem 91042, Israel 972-2-587-2202 FAX: 972-2-587-1101 US: (800) 791-3904 FAX: (800) 791-3912 http://www.alomone.com Alpha Innotech 14743 Catalina Street San Leandro, CA 94577 (800) 795-5556 FAX: (510) 483-3227 (510) 483-9620 http://www.alphainnotech.com Altec Plastics 116 B Street Boston, MA 02127 (800) 477-8196 FAX: (617) 269-8484 (617) 269-1400
Alza 1900 Charleston Road P.O. Box 7210 Mountain View, CA 94043 (800) 692-2990 FAX: (650) 564-7070 (650) 564-5000 http://www.alza.com Amac 160B Larrabee Road Westbrook, ME 04092 (800) 458-5060 FAX: (207) 854-0116 (207) 854-0426 Amano Enzyme Company 2-7, 1-chome, Nishiki, Naka-ku, Nagoya, 460-8630 Japan (81) (0)52-211-3032 FAX: (81) (0)52-211-3054 http://www.amanoenzyme.co.jp/english/ Amaresco 30175 Solon Industrial Parkway Solon, Ohio 44139 (800) 366-1313 FAX: (440) 349-1182 (440) 349-1313 Ambion 2130 Woodward Street, Suite 200 Austin, TX 78744 (800) 888-8804 FAX: (512) 651-0190 (512) 651-0200 http://www.ambion.com
American Laboratory Supply See American Bioanalytical American Medical Systems 10700 Bren Road West Minnetonka, MN 55343 (800) 328-3881 FAX: (612) 930-6654 (612) 933-4666 http://www.visitams.com American Qualex 920-A Calle Negocio San Clemente, CA 92673 (949) 492-8298 FAX: (949) 492-6790 http://www.americanqualex.com American Radiolabeled Chemicals 11624 Bowling Green St. Louis, MO 63146 (800) 331-6661 FAX: (800) 999-9925 (314) 991-4545 FAX: (314) 991-4692 http://www.arc-inc.com American Scientific Products See VWR Scientific Products American Society for Histocompatibility and Immunogenetics P.O. Box 15804 Lenexa, KS 66285 (913) 541-0009 FAX: (913) 541-0156 http://www.swmed.edu/home pages/ ASHI/ashi.htm
American Association of Blood Banks College of American Pathologists 325 Waukegan Road Northfield, IL 60093 (800) 323-4040 FAX: (847) 8166 (847) 832-7000 http://www.cap.org
American Type Culture Collection (ATCC) 10801 University Boulevard Manassas, VA 20110 (800) 638-6597 FAX: (703) 365-2750 (703) 365-2700 http://www.atcc.org
American Bio-Technologies See Intracel Corporation
Amersham See Amersham Pharmacia Biotech
American Bioanalytical 15 Erie Drive Natick, MA 01760 (800) 443-0600 FAX: (508) 655-2754 (508) 655-4336 http://www.americanbio.com
Amersham International Amersham Place Little Chalfont, Buckinghamshire HP7 9NA, UK (44) 1494-544100 FAX: (44) 1494-544350 http://www.apbiotech.com
American Cyanamid P.O. Box 400 Princeton, NJ 08543 (609) 799-0400 FAX: (609) 275-3502 http://www.cyanamid.com American HistoLabs 7605-F Airpark Road Gaithersburg, MD 20879 (301) 330-1200 FAX: (301) 330-6059 American International Chemical 17 Strathmore Road Natick, MA 01760 (800) 238-0001 (508) 655-5805 http://www.aicma.com
Amersham Medi-Physics Also see Nycomed Amersham 3350 North Ridge Avenue Arlington Heights, IL 60004 (800) 292-8514 FAX: (800) 807-2382 http://www.nycomed-amersham.com Amersham Pharmacia Biotech 800 Centennial Avenue P.O. Box 1327 Piscataway, NJ 08855 (800) 526-3593 FAX: (877) 295-8102 (732) 457-8000 http://www.apbiotech.com
Suppliers
2 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
Amgen 1 Amgen Center Drive Thousand Oaks, CA 91320 (800) 926-4369 FAX: (805) 498-9377 (805) 447-5725 http://www.amgen.com
Analytical Genetics Testing Center 7808 Cherry Creek S. Drive, Suite 201 Denver, CO 80231 (800) 204-4721 FAX: (303) 750-2171 (303) 750-2023 http://www.geneticid.com
Apple Scientific 11711 Chillicothe Road, Unit 2 P.O. Box 778 Chesterland, OH 44026 (440) 729-3056 FAX: (440) 729-0928 http://www.applesci.com
Archimica Florida P.O. Box 1466 Gainesville, FL 32602 (800) 331-6313 FAX: (352) 371-6246 (352) 376-8246 http://www.archimica.com
Amicon Scientific Systems Division 72 Cherry Hill Drive Beverly, MA 01915 (800) 426-4266 FAX: (978) 777-6204 (978) 777-3622 http://www.amicon.com
AnaSpec 2149 O’Toole Avenue, Suite F San Jose, CA 95131 (800) 452-5530 FAX: (408) 452-5059 (408) 452-5055 http://www.anaspec.com
Applied Biosystems See PE Biosystems
Arcor Electronics 1845 Oak Street #15 Northfield, IL 60093 (847) 501-4848
Amika 8980F Route 108 Oakland Center Columbia, MD 21045 (800) 547-6766 FAX: (410) 997-7104 (410) 997-0100 http://www.amika.com Amoco Performance Products See BPAmoco AMPI See Pacer Scientific
Ancare 2647 Grand Avenue P.O. Box 814 Bellmore, NY 11710 (800) 645-6379 FAX: (516) 781-4937 (516) 781-0755 http://www.ancare.com Ancell 243 Third Street North P.O. Box 87 Bayport, MN 55033 (800) 374-9523 FAX: (651) 439-1940 (651) 439-0835 http://www.ancell.com
Amrad 576 Swan Street Richmond, Victoria 3121, Australia 613-9208-4000 FAX: 613-9208-4350 http://www.amrad.com.au
Anderson Instruments 500 Technology Court Smyrna, GA 30082 (800) 241-6898 FAX: (770) 319-5306 (770) 319-9999 http://www.graseby.com
Amresco 30175 Solon Industrial Parkway Solon, OH 44139 (800) 829-2805 FAX: (440) 349-1182 (440) 349-1199
Andreas Hettich Gartenstrasse 100 Postfach 260 D-78732 Tuttlingen, Germany (49) 7461 705 0 FAX: (49) 7461 705-122 http://www.hettich-centrifugen.de
Anachemia Chemicals 3 Lincoln Boulevard Rouses Point, NY 12979 (800) 323-1414 FAX: (518) 462-1952 (518) 462-1066 http://www.anachemia.com Ana-Gen Technologies 4015 Fabian Way Palo Alto, CA 94303 (800) 654-4671 FAX: (650) 494-3893 (650) 494-3894 http://www.ana-gen.com Analtech Inc. P.O. Box 7558 Newark, DE 19714 (800) 441-7540 FAX: (302) 737-7115 (302) 737-6960 http://www.analtech.com Analytical Biological Services Cornell Business Park 701-4 Wilmington, DE 19801 (800) 391-2391 FAX: (302) 654-8046 (302) 654-4492 http://www.ABSbioreagents.com
Anesthetic Vaporizer Services 10185 Main Street Clarence, NY 14031 (719) 759-8490 www.avapor.com Animal Identification and Marking Systems (AIMS) 13 Winchester Avenue Budd Lake, NJ 07828 (908) 684-9105 FAX: (908) 684-9106 http://www.animalid.com Annovis 34 Mount Pleasant Drive Aston, PA 19014 (800) EASY-DNA FAX: (610) 361-8255 (610) 361-9224 http://www.annovis.com Apotex 150 Signet Drive Weston, Ontario M9L 1T9, Canada (416) 749-9300 FAX: (416) 749-2646 http://www.apotex.com
Applied Imaging 2380 Walsh Avenue, Bldg. B Santa Clara, CA 95051 (800) 634-3622 FAX: (408) 562-0264 (408) 562-0250 http://www.aicorp.com Applied Photophysics 203-205 Kingston Road Leatherhead, Surrey, KT22 7PB UK (44) 1372-386537 Applied Precision 1040 12th Avenue Northwest Issaquah, Washington 98027 (425) 557-1000 FAX: (425) 557-1055 http://www.api.com/index.html Appligene Oncor Parc d’Innovation Rue Geiler de Kaysersberg, BP 72 67402 Illkirch Cedex, France (33) 88 67 22 67 FAX: (33) 88 67 19 45 http://www.oncor.com/prod-app.htm Applikon 1165 Chess Drive, Suite G Foster City, CA 94404 (650) 578-1396 FAX: (650) 578-8836 http://www.applikon.com Appropriate Technical Resources 9157 Whiskey Bottom Road Laurel, MD 20723 (800) 827-5931 FAX: (410) 792-2837 http://www.atrbiotech.com APV Gaulin 100 S. CP Avenue Lake Mills, WI 53551 (888) 278-4321 FAX: (888) 278-5329 http://www.apv.com Aqualon See Hercules Aqualon Aquebogue Machine and Repair Shop Box 2055 Main Road Aquebogue, NY 11931 (631) 722-3635 FAX: (631) 722-3106 Archer Daniels Midland 4666 Faries Parkway Decatur, IL 62525 (217) 424-5200 http://www.admworld.com
Arcturus Engineering 400 Logue Avenue Mountain View, CA 94043 (888) 446 7911 FAX: (650) 962 3039 (650) 962 3020 http://www.arctur.com Argonaut Technologies 887 Industrial Road, Suite G San Carlos, CA 94070 (650) 998-1350 FAX: (650) 598-1359 http://www.argotech.com Ariad Pharmaceuticals 26 Landsdowne Street Cambridge, MA 02139 (617) 494-0400 FAX: (617) 494-8144 http://www.ariad.com Armour Pharmaceuticals See Rhone-Poulenc Rorer Aronex Pharmaceuticals 8707 Technology Forest Place The Woodlands, TX 77381 (281) 367-1666 FAX: (281) 367-1676 http://www.aronex.com Artisan Industries 73 Pond Street Waltham, MA 02254 (617) 893-6800 http://www.artisanind.com ASI Instruments 12900 Ten Mile Road Warren, MI 48089 (800) 531-1105 FAX: (810) 756-9737 (810) 756-1222 http://www.asi-instruments.com Aspen Research Laboratories 1700 Buerkle Road White Bear Lake, MN 55140 (651) 264-6000 FAX: (651) 264-6270 http://www.aspenresearch.com Associates of Cape Cod 704 Main Street Falmouth, MA 02540 (800) LAL-TEST FAX: (508) 540-8680 (508) 540-3444 http://www.acciusa.com Astra Pharmaceuticals See AstraZeneca
Suppliers
3 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
AstraZeneca 1800 Concord Pike Wilmington, DE 19850 (302) 886-3000 FAX: (302) 886-2972 http://www.astrazeneca.com AT Biochem 30 Spring Mill Drive Malvern, PA 19355 (610) 889-9300 FAX: (610) 889-9304 ATC Diagnostics See Vysis ATCC See American Type Culture Collection Athens Research and Technology P.O. Box 5494 Athens, GA 30604 (706) 546-0207 FAX: (706) 546-7395 Atlanta Biologicals 1425-400 Oakbrook Drive Norcross, GA 30093 (800) 780-7788 or (770) 446-1404 FAX: (800) 780-7374 or (770) 446-1404 http://www.atlantabio.com Atomergic Chemical 71 Carolyn Boulevard Farmingdale, NY 11735 (631) 694-9000 FAX: (631) 694-9177 http://www.atomergic.com Atomic Energy of Canada 2251 Speakman Drive Mississauga, Ontario L5K 1B2 Canada (905) 823-9040 FAX: (905) 823-1290 http://www.aecl.ca ATR P.O. Box 460 Laurel, MD 20725 (800) 827-5931 FAX: (410) 792-2837 (301) 470-2799 http://www.atrbiotech.com
Aventis BP 67917 67917 Strasbourg Cedex 9, France 33 (0) 388 99 11 00 FAX: 33 (0) 388 99 11 01 http://www.aventis.com Aventis Pasteur 1 Discovery Drive Swiftwater, PA 18370 (800) 822-2463 FAX: (570) 839-0955 (570) 839-7187 http://www.aventispasteur.com/usa Avery Dennison 150 North Orange Grove Boulevard Pasadena, CA 91103 (800) 462-8379 FAX: (626) 792-7312 (626) 304-2000 http://www.averydennison.com Avestin 2450 Don Reid Drive Ottawa, Ontario K1H 1E1, Canada (888) AVESTIN FAX: (613) 736-8086 (613) 736-0019 http://www.avestin.com AVIV Instruments 750 Vassar Avenue Lakewood, NJ 08701 (732) 367-1663 FAX: (732) 370-0032 http://www.avivinst.com Axon Instruments 1101 Chess Drive Foster City, CA 94404 (650) 571-9400 FAX: (650) 571-9500 http://www.axon.com Azon 720 Azon Road Johnson City, NY 13790 (800) 847-9374 FAX: (800) 635-6042 (607) 797-2368 http://www.azon.com
Aurora Biosciences 11010 Torreyana Road San Diego, CA 92121 (858) 404-6600 FAX: (858) 404-6714 http://www.aurorabio.com
BAbCO 1223 South 47th Street Richmond, CA 94804 (800) 92-BABCO FAX: (510) 412-8940 (510) 412-8930 http://www.babco.com
Automatic Switch Company A Division of Emerson Electric 50 Hanover Road Florham Park, NJ 07932 (800) 937-2726 FAX: (973) 966-2628 (973) 966-2000 http://www.asco.com
Bacharach 625 Alpha Drive Pittsburgh, PA 15238 (800) 736-4666 FAX: (412) 963-2091 (412) 963-2000 http://www.bacharach-inc.com
Avanti Polar Lipids 700 Industrial Park Drive Alabaster, AL 35007 (800) 227-0651 FAX: (800) 229-1004 (205) 663-2494 FAX: (205) 663-0756 http://www.avantilipids.com
Bachem Bioscience 3700 Horizon Drive King of Prussia, PA 19406 (800) 634-3183 FAX: (610) 239-0800 (610) 239-0300 http://www.bachem.com
Bachem California 3132 Kashiwa Street P.O. Box 3426 Torrance, CA 90510 (800) 422-2436 FAX: (310) 530-1571 (310) 539-4171 http://www.bachem.com Baekon 18866 Allendale Avenue Saratoga, CA 95070 (408) 972-8779 FAX: (408) 741-0944 Baker Chemical See J.T. Baker Bangs Laboratories 9025 Technology Drive Fishers, IN 46038 (317) 570-7020 FAX: (317) 570-7034 http://www.bangslabs.com Bard Parker See Becton Dickinson Barnstead/Thermolyne P.O. Box 797 2555 Kerper Boulevard Dubuque, IA 52004 (800) 446-6060 FAX: (319) 589-0516 http://www.barnstead.com Barrskogen 4612 Laverock Place N Washington, DC 20007 (800) 237-9192 FAX: (301) 464-7347 BAS See Bioanalytical Systems BASF Specialty Products 3000 Continental Drive North Mt. Olive, NJ 07828 (800) 669-2273 FAX: (973) 426-2610 http://www.basf.com Baum, W.A. 620 Oak Street Copiague, NY 11726 (631) 226-3940 FAX: (631) 226-3969 http://www.wabaum.com Bausch & Lomb One Bausch & Lomb Place Rochester, NY 14604 (800) 344-8815 FAX: (716) 338-6007 (716) 338-6000 http://www.bausch.com Baxter Fenwal Division 1627 Lake Cook Road Deerfield, IL 60015 (800) 766-1077 FAX: (800) 395-3291 (847) 940-6599 FAX: (847) 940-5766 http://www.powerfulmedicine.com
Baxter Healthcare One Baxter Parkway Deerfield, IL 60015 (800) 777-2298 FAX: (847) 948-3948 (847) 948-2000 http://www.baxter.com Baxter Scientific Products See VWR Scientific Bayer Agricultural Division Animal Health Products 12707 Shawnee Mission Pkwy. Shawnee Mission, KS 66201 (800) 255-6517 FAX: (913) 268-2803 (913) 268-2000 http://www.bayerus.com Bayer Diagnostics Division (Order Services) P.O. Box 2009 Mishiwaka, IN 46546 (800) 248-2637 FAX: (800) 863-6882 (219) 256-3390 http://www.bayer.com Bayer Diagnostics 511 Benedict Avenue Tarrytown, NY 10591 (800) 255-3232 FAX: (914) 524-2132 (914) 631-8000 http://www.bayerdiag.com Bayer Plc Diagnostics Division Bayer House, Strawberry Hill Newbury, Berkshire RG14 1JA, UK (44) 1635-563000 FAX: (44) 1635-563393 http://www.bayer.co.uk BD Immunocytometry Systems 2350 Qume Drive San Jose, CA 95131 (800) 223-8226 FAX: (408) 954-BDIS http://www.bdfacs.com BD Labware Two Oak Park Bedford, MA 01730 (800) 343-2035 FAX: (800) 743-6200 http://www.bd.com/labware BD PharMingen 10975 Torreyana Road San Diego, CA 92121 (800) 848-6227 FAX: (858) 812-8888 (858) 812-8800 http://www.pharmingen.com BD Transduction Laboratories 133 Venture Court Lexington, KY 40511 (800) 227-4063 FAX: (606) 259-1413 (606) 259-1550 http://www.translab.com
Suppliers
4 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
BDH Chemicals Broom Road Poole, Dorset BH12 4NN, UK (44) 1202-745520 FAX: (44) 1202- 2413720 BDH Chemicals See Hoefer Scientific Instruments BDIS See BD Immunocytometry Systems Beckman Coulter 4300 North Harbor Boulevard Fullerton, CA 92834 (800) 233-4685 FAX: (800) 643-4366 (714) 871-4848 http://www.beckman-coulter.com Beckman Instruments Spinco Division/Bioproducts Operation 1050 Page Mill Road Palo Alto, CA 94304 (800) 742-2345 FAX: (415) 859-1550 (415) 857-1150 http://www.beckman-coulter.com Becton Dickinson Immunocytometry & Cellular Imaging 2350 Qume Drive San Jose, CA 95131 (800) 223-8226 FAX: (408) 954-2007 (408) 432-9475 http://www.bdfacs.com Becton Dickinson Labware 1 Becton Drive Franklin Lakes, NJ 07417 (888) 237-2762 FAX: (800) 847-2220 (201) 847-4222 http://www.bdfacs.com Becton Dickinson Labware 2 Bridgewater Lane Lincoln Park, NJ 07035 (800) 235-5953 FAX: (800) 847-2220 (201) 847-4222 http://www.bdfacs.com Becton Dickinson Primary Care Diagnostics 7 Loveton Circle Sparks, MD 21152 (800) 675-0908 FAX: (410) 316-4723 (410) 316-4000 http://www.bdfacs.com Behringwerke Diagnostika Hoechster Strasse 70 P-65835 Liederback, Germany (49) 69-30511 FAX: (49) 69-303-834 Bellco Glass 340 Edrudo Road Vineland, NJ 08360 (800) 257-7043 FAX: (856) 691-3247 (856) 691-1075 http://www.bellcoglass.com Bender Biosystems See Serva
Beral Enterprises See Garren Scientific Berkeley Antibody See BAbCO Bernsco Surgical Supply 25 Plant Avenue Hauppague, NY 11788 (800) TIEMANN FAX: (516) 273-6199 (516) 273-0005 http://www.bernsco.com Berry and Associates 2434 Bishop Circle East Dexter, MI 48130 (800) 357-1145 FAX: (734) 426-9077 (734) 426-3787 http://www.berryassoc.com Beta Medical and Scientific (Datesand Ltd.) 2 Ferndale Road Sale, Manchester M33 3GP, UK (44) 1612 317676 FAX: (44) 1612 313656 Bethesda Research Laboratories (BRL) See Life Technologies Biacore 200 Centennial Avenue, Suite 100 Piscataway, NJ 08854 (800) 242-2599 FAX: (732) 885-5669 (732) 885-5618 http://www.biacore.com Bilaney Consultants St. Julian’s Sevenoaks, Kent TN15 0RX, UK (44) 1732 450002 FAX: (44) 1732 450003 http://www.bilaney.com Binding Site 5889 Oberlin Drive, Suite 101 San Diego, CA 92121 (800) 633-4484 FAX: (619) 453-9189 (619) 453-9177 http://www.bindingsite.co.uk BIO 101 See Qbiogene Bio Image See Genomic Solutions Bioanalytical Systems 2701 Kent Avenue West Lafayette, IN 47906 (800) 845-4246 FAX: (765) 497-1102 (765) 463-4527 http://www.bioanalytical.com BioAutomation 8408 Kenning Court Plano, TX 75024 972-335-2525 FAX: 972-335-3731 http://www.bioautomation.com
Biocell 2001 University Drive Rancho Dominguez, CA 90220 (800) 222-8382 FAX: (310) 637-3927 (310) 537-3300 http://www.biocell.com Biocoat See BD Labware BioComp Instruments 650 Churchill Road Fredericton, New Brunswick E3B 1P6 Canada (800) 561-4221 FAX: (506) 453-3583 (506) 453-4812 http://131.202.97.21
Biological Detection Systems See Cellomics or Amersham Biomeda 1166 Triton Drive, Suite E P.O. Box 8045 Foster City, CA 94404 (800) 341-8787 FAX: (650) 341-2299 (650) 341-8787 http://www.biomeda.com BioMedic Data Systems 1 Silas Road Seaford, DE 19973 (800) 526-2637 FAX: (302) 628-4110 (302) 628-4100 http://www.bmds.com
BioDesign P.O. Box 1050 Carmel, NY 10512 (914) 454-6610 FAX: (914) 454-6077 http://www.biodesignofny.com
Biomedical Engineering P.O. Box 980694 Virginia Commonwealth University Richmond, VA 23298 (804) 828-9829 FAX: (804) 828-1008
Bioengineering AG Sagenrainstrasse 7 CH8636 Wald, Switzerland (41) 55-256-8-111 FAX: (41) 55-256-8-256
Biomedical Research Instruments 12264 Wilkins Avenue Rockville, MD 20852 (800) 327-9498 (301) 881-7911 http://www.biomedinstr.com
Biofine International Inc. P.O. Box 712 Blaine, WA 98231-0712 (866) 472-9441 FAX: (866) 623-7408 (604) 438-8181 FAX: (604) 438-8378 http://biofineintl.com Biofluids Division of Biosource International 1114 Taft Street Rockville, MD 20850 (800) 972-5200 FAX: (301) 424-3619 (301) 424-4140 http://www.biosource.com BioFX Laboratories 9633 Liberty Road, Suite S Randallstown, MD 21133 (800) 445-6447 FAX: (410) 498-6008 (410) 496-6006 http://www.biofx.com BioGenex Laboratories 4600 Norris Canyon Road San Ramon, CA 94583 (800) 421-4149 FAX: (925) 275-0580 (925) 275-0550 http://www.biogenex.com Bioline 2470 Wrondel Way Reno, NV 89502 (888) 257-5155 FAX: (775) 828-7676 (775) 828-0202 http://www.bioline.com Bio-Logic Research & Development 1, rue de l-Europe A.Z. de Font-Ratel 38640 CLAIX, France (33) 76-98-68-31 FAX: (33) 76-98-69-09
Bio/medical Specialties P.O. Box 1687 Santa Monica, CA 90406 (800) 269-1158 FAX: (800) 269-1158 (323) 938-7515 BioMerieux 100 Rodolphe Street Durham, North Carolina 27712 (919) 620-2000 http://www.biomerieux.com BioMetallics P.O. Box 2251 Princeton, NJ 08543 (800) 999-1961 FAX: (609) 275-9485 (609) 275-0133 http://www.microplate.com Biomol Research Laboratories 5100 Campus Drive Plymouth Meeting, PA 19462 (800) 942-0430 FAX: (610) 941-9252 (610) 941-0430 http://www.biomol.com Bionique Testing Labs Fay Brook Drive RR 1, Box 196 Saranac Lake, NY 12983 (518) 891-2356 FAX: (518) 891-5753 http://www.bionique.com Biopac Systems 42 Aero Camino Santa Barbara, CA 93117 (805) 685-0066 FAX: (805) 685-0067 http://www.biopac.com Bioproducts for Science See Harlan Bioproducts for Science
Suppliers
5 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Bioptechs 3560 Beck Road Butler, PA 16002 (877) 548-3235 FAX: (724) 282-0745 (724) 282-7145 http://www.bioptechs.com BIOQUANT-R&M Biometrics 5611 Ohio Ave Nashville, TN 37209 (800) 221-0549 FAX: (615) 350-7282 (615) 350-7866 http://www.bioquant.com Bio-Rad Laboratories 2000 Alfred Nobel Drive Hercules, CA 94547 (800) 424-6723 FAX: (800) 879-2289 (510) 741-1000 FAX: (510) 741-5800 http://www.bio-rad.com Bio-Rad Laboratories Maylands Avenue Hemel Hempstead, Herts HP2 7TD, UK http://www.bio-rad.com BioRobotics 3-4 Bennell Court Comberton, Cambridge CB3 7DS, UK (44) 1223-264345 FAX: (44) 1223-263933 http://www.biorobotics.co.uk BIOS Laboratories See Genaissance Pharmaceuticals Biosearch Technologies 81 Digital Drive Novato, CA 94949 (800) GENOME1 FAX: (415) 883-8488 (415) 883-8400 http://www.biosearchtech.com BioSepra 111 Locke Drive Marlborough, MA 01752 (800) 752-5277 FAX: (508) 357-7595 (508) 357-7500 http://www.biosepra.com Bio-Serv 1 8th Street, Suite 1 Frenchtown, NJ 08825 (908) 996-2155 FAX: (908) 996-4123 http://www.bio-serv.com BioSignal 1744 William Street, Suite 600 Montreal, Quebec H3J 1R4, Canada (800) 293-4501 FAX: (514) 937-0777 (514) 937-1010 http://www.biosignal.com Biosoft P.O. Box 10938 Ferguson, MO 63135 (314) 524-8029 FAX: (314) 524-8129 http://www.biosoft.com
Biosource International 820 Flynn Road Camarillo, CA 93012 (800) 242-0607 FAX: (805) 987-3385 (805) 987-0086 http://www.biosource.com
Bioventures P.O. Box 2561 848 Scott Street Murfreesboro, TN 37133 (800) 235-8938 FAX: (615) 896-4837 http://www.bioventures.com
Boekel Scientific 855 Pennsylvania Boulevard Feasterville, PA 19053 (800) 336-6929 FAX: (215) 396-8264 (215) 396-8200 http://www.boekelsci.com
BioSpec Products P.O. Box 788 Bartlesville, OK 74005 (800) 617-3363 FAX: (918) 336-3363 (918) 336-3363 http://www.biospec.com
BioWhittaker 8830 Biggs Ford Road P.O. Box 127 Walkersville, MD 21793 (800) 638-8174 FAX: (301) 845-8338 (301) 898-7025 http://www.biowhittaker.com
Bohdan Automation 1500 McCormack Boulevard Mundelein, IL 60060 (708) 680-3939 FAX: (708) 680-1199
Biosure See Riese Enterprises Biosym Technologies See Molecular Simulations Biosys 21 quai du Clos des Roses 602000 Compiegne, France (33) 03 4486 2275 FAX: (33) 03 4484 2297 Bio-Tech Research Laboratories NIAID Repository Rockville, MD 20850 http://www.niaid.nih.gov/ncn/repos.htm Biotech Instruments Biotech House 75A High Street Kimpton, Hertfordshire SG4 8PU, UK (44) 1438 832555 FAX: (44) 1438 833040 http://www.biotinst.demon.co.uk Biotech International 11 Durbell Street Acacia Ridge, Queensland 4110 Australia 61-7-3370-6396 FAX: 61-7-3370-6370 http://www.avianbiotech.com Biotech Source Inland Farm Drive South Windham, ME 04062 (207) 892-3266 FAX: (207) 892-6774
Biozyme Laboratories 9939 Hibert Street, Suite 101 San Diego, CA 92131 (800) 423-8199 FAX: (858) 549-0138 (858) 549-4484 http://www.biozyme.com Bird Products 1100 Bird Center Drive Palm Springs, CA 92262 (800) 328-4139 FAX: (760) 778-7274 (760) 778-7200 http://www.birdprod.com/bird B & K Universal 2403 Yale Way Fremont, CA 94538 (800) USA-MICE FAX: (510) 490-3036 BLS Ltd. Zselyi Aladar u. 31 1165 Budapest, Hungary (36) 1-407-2602 FAX: (36) 1-407-2896 http://www.bls-ltd.com Blue Sky Research 3047 Orchard Parkway San Jose, CA 95134 (408) 474-0988 FAX: (408) 474-0989 http://www.blueskyresearch.com Blumenthal Industries 7 West 36th Street, 13th floor New York, NY 10018 (212) 719-1251 FAX: (212) 594-8828 BOC Edwards One Edwards Park 301 Ballardvale Street Wilmington, MA 01887 (800) 848-9800 FAX: (978) 658-7969 (978) 658-5410 http://www.bocedwards.com
Biotecx Laboratories 6023 South Loop East Houston, TX 77033 (800) 535-6286 FAX: (713) 643-3143 (713) 643-0606 http://www.biotecx.com
Boehringer Ingelheim 900 Ridgebury Road P.O. Box 368 Ridgefield, CT 06877 (800) 243-0127 FAX: (203) 798-6234 (203) 798-9988 http://www.boehringer-ingelheim.com Boehringer Mannheim Biochemicals Division See Roche Diagnostics
Brain Research Laboratories Waban P.O. Box 88 Newton, MA 02468 (888) BRL-5544 FAX: (617) 965-6220 (617) 965-5544 http://www.brainresearchlab.com Braintree Scientific P.O. Box 850929 Braintree, MA 02185 (781) 843-1644 FAX: (781) 982-3160 http://www.braintreesci.com Brandel 8561 Atlas Drive Gaithersburg, MD 20877 (800) 948-6506 FAX: (301) 869-5570 (301) 948-6506 http://www.brandel.com
Bio-Tek Instruments Highland Industrial Park P.O. Box 998 Winooski, VT 05404 (800) 451-5172 FAX: (802) 655-7941 (802) 655-4040 http://www.biotek.com
BioTherm 3260 Wilson Boulevard Arlington, VA 22201 (703) 522-1705 FAX: (703) 522-2606
BPAmoco 4500 McGinnis Ferry Road Alpharetta, GA 30005 (800) 328-4537 FAX: (770) 772-8213 (770) 772-8200 http://www.bpamoco.com
Branson Ultrasonics 41 Eagle Road Danbury, CT 06813 (203) 796-0400 FAX: (203) 796-9838 http://www.plasticsnet.com/branson B. Braun Biotech 999 Postal Road Allentown, PA 18103 (800) 258-9000 FAX: (610) 266-9319 (610) 266-6262 http://www.bbraunbiotech.com B. Braun Biotech International Schwarzenberg Weg 73-79 P.O. Box 1120 D-34209 Melsungen, Germany (49) 5661-71-3400 FAX: (49) 5661-71-3702 http://www.bbraunbiotech.com B. Braun-McGaw 2525 McGaw Avenue Irvine, CA 92614 (800) BBRAUN-2 (800) 624-2963 http://www.bbraunusa.com B. Braun Medical Thorncliffe Park Sheffield S35 2PW, UK (44) 114-225-9000 FAX: (44) 114-225-9111 http://www.bbmuk.demon.co.uk
Suppliers
6 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
Brenntag P.O. Box 13788 Reading, PA 19612-3788 (610) 926-4151 FAX: (610) 926-4160 http://www.brenntagnortheast.com Bresatec See GeneWorks Bright/Hacker Instruments 17 Sherwood Lane Fairfield, NJ 07004 (973) 226-8450 FAX: (973) 808-8281 http://www.hackerinstruments.com Brinkmann Instruments Subsidiary of Sybron 1 Cantiague Road P.O. Box 1019 Westbury, NY 11590 (800) 645-3050 FAX: (516) 334-7521 (516) 334-7500 http://www.brinkmann.com Bristol-Meyers Squibb P.O. Box 4500 Princeton, NJ 08543 (800) 631-5244 FAX: (800) 523-2965 http://www.bms.com Broadley James 19 Thomas Irvine, CA 92618 (800) 288-2833 FAX: (949) 829-5560 (949) 829-5555 http://www.broadleyjames.com Brookhaven Instruments 750 Blue Point Road Holtsville, NY 11742 (631) 758-3200 FAX: (631) 758-3255 http://www.bic.com Brownlee Labs See Applied Biosystems Distributed by Pacer Scientific
BTX Division of Genetronics 11199 Sorrento Valley Road San Diego, CA 92121 (800) 289-2465 FAX: (858) 597-9594 (858) 597-6006 http://www.genetronics.com/btx Buchler Instruments See Baxter Scientific Products Buckshire 2025 Ridge Road Perkasie, PA 18944 (215) 257-0116 Chimera Biotec GmbH Dortmund BioMedicine Center Emil-Figge-Str. 76 A D-44227 Dortmund Germany 49 (0)231 9742-840 FAX: 49 (0)231 9742-844 http://www.chimerabiotec.de/index.htm Burdick and Jackson Division of Baxter Scientific Products 1953 S. Harvey Street Muskegon, MI 49442 (800) 368-0050 FAX: (231) 728-8226 (231) 726-3171 http://www.bandj.com/mainframe.htm Burleigh Instruments P.O. Box E Fishers, NY 14453 (716) 924-9355 FAX: (716) 924-9072 http://www.burleigh.com Burns Veterinary Supply 1900 Diplomat Drive Farmer’s Branch, TX 75234 (800) 92-BURNS FAX: (972) 243-6841 http://www.burnsvet.com Burroughs Wellcome See Glaxo Wellcome
Bruel & Kjaer Division of Spectris Technologies 2815 Colonnades Court Norcross, GA 30071 (800) 332-2040 FAX: (770) 847-8440 (770) 209-6907 http://www.bkhome.com
The Butler Company 5600 Blazer Parkway Dublin, OH 43017 (800) 551-3861 FAX: (614) 761-9096 (614) 761-9095 http://www.wabutler.com
Bruker Analytical X-Ray Systems 5465 East Cheryl Parkway Madison, WI 53711 (800) 234-XRAY FAX: (608) 276-3006 (608) 276-3000 http://www.bruker-axs.com
Butterworth Laboratories 54-56 Waldegrave Road Teddington, Middlesex TW11 8LG, UK (44)(0)20-8977-0750 FAX: (44)(0)28-8943-2624 http://www.butterworth-labs.co.uk
Bruker Instruments 19 Fortune Drive Billerica, MA 01821 (978) 667-9580 FAX: (978) 667-0985 http://www.bruker.com
Buxco Electronics 95 West Wood Road #2 Sharon, CT 06069 (860) 364-5558 FAX: (860) 364-5116 http://www.buxco.com
C/D/N Isotopes 88 Leacock Street Pointe-Claire, Quebec Canada H9R 1H1 (800) 697-6254 FAX: (514) 697-6148 C.M.A./Microdialysis AB 73 Princeton Street North Chelmsford, MA 01863 (800) 440-4980 FAX: (978) 251-1950 (978) 251-1940 http://www.microdialysis.com Calbiochem-Novabiochem P.O. Box 12087-2087 La Jolla, CA 92039 (800) 854-3417 FAX: (800) 776-0999 (858) 450-9600 http://www.calbiochem.com Calorimetry Sciences 155 West 2050 North Spanish Fork, UT 84660 (801) 794-2600 FAX: (801) 794-2700 http://www.calscorp.com Caltag Laboratories 1849 Bayshore Highway, Suite 200 Burlingame, CA 94010 (800) 874-4007 FAX: (650) 652-9030 (650) 652-0468 http://www.caltag.com Cambridge Electronic Design Science Park, Milton Road Cambridge CB4 0FE, UK 44 (0) 1223-420-186 FAX: 44 (0) 1223-420-488 http://www.ced.co.uk Cambridge Isotope Laboratories 50 Frontage Road Andover, MA 01810 (800) 322-1174 FAX: (978) 749-2768 (978) 749-8000 http://www.isotope.com Cambridge Research Biochemicals See Zeneca/CRB Cambridge Technology 109 Smith Place Cambridge, MA 02138 (6l7) 441-0600 FAX: (617) 497-8800 http://www.camtech.com Camlab Nuffield Road Cambridge CB4 1TH, UK (44) 122-3424222 FAX: (44) 122-3420856 http://www.camlab.co.uk/home.htm Campden Instruments Park Road Sileby Loughborough Leicestershire LE12 7TU, UK (44) 1509-814790 FAX: (44) 1509-816097 http://www.campdeninst.com/home.htm
Cappel Laboratories See Organon Teknika Cappel Carl Roth GmgH & Company Schoemperlenstrasse 1-5 76185 Karlsrube Germany (49) 72-156-06164 FAX: (49) 72-156-06264 http://www.carl-roth.de Carl Zeiss One Zeiss Drive Thornwood, NY 10594 (800) 233-2343 FAX: (914) 681-7446 (914) 747-1800 http://www.zeiss.com Carlo Erba Reagenti Via Winckelmann 1 20148 Milano Lombardia, Italy (39) 0-29-5231 FAX: (39) 0-29-5235-904 http://www.carloerbareagenti.com Carolina Biological Supply 2700 York Road Burlington, NC 27215 (800) 334-5551 FAX: (336) 584-76869 (336) 584-0381 http://www.carolina.com Carolina Fluid Components 9309 Stockport Place Charlotte, NC 28273 (704) 588-6101 FAX: (704) 588-6115 http://www.cfcsite.com Cartesian Technologies 17851 Skypark Circle, Suite C Irvine, CA 92614 (800) 935-8007 http://cartesiantech.com Cayman Chemical 1180 East Ellsworth Road Ann Arbor, MI 48108 (800) 364-9897 FAX: (734) 971-3640 (734) 971-3335 http://www.caymanchem.com CB Sciences One Washington Street, Suite 404 Dover, NH 03820 (800) 234-1757 FAX: (603) 742-2455 http://www.cbsci.com CBS Scientific P.O. Box 856 Del Mar, CA 92014 (800) 243-4959 FAX: (858) 755-0733 (858) 755-4959 http://www.cbssci.com CCR (Coriell Cell Repository) See Coriell Institute for Medical Research
Suppliers
7 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Cedarlane Laboratories 5516 8th Line, R.R. #2 Hornby, Ontario L0P 1E0, Canada (905) 878-8891 FAX: (905) 878-7800 http://www.cedarlanelabs.com CE Instruments Grand Avenue Parkway Austin, TX 78728 (800) 876-6711 FAX: (512) 251-1597 http://www.ceinstruments.com CEL Associates P.O. Box 721854 Houston, TX 77272 (800) 537-9339 FAX: (281) 933-0922 (281) 933-9339 http://www.cel-1.com Cel-Line Associates See Erie Scientific Celite World Minerals 130 Castilian Drive Santa Barbara, CA 93117 (805) 562-0200 FAX: (805) 562-0299 http://www.worldminerals.com/celite Cell Genesys 342 Lakeside Drive Foster City, CA 94404 (650) 425-4400 FAX: (650) 425-4457 http://www.cellgenesys.com Cell Systems 12815 NE 124th Street, Suite A Kirkland, WA 98034 (800) 697-1211 FAX: (425) 820-6762 (425) 823-1010 Cellmark Diagnostics 20271 Goldenrod Lane Germantown, MD 20876 (800) 872-5227 FAX: (301) 428-4877 (301) 428-4980 http://www.cellmark-labs.com Cellomics 635 William Pitt Way Pittsburgh, PA 15238 (888) 826-3857 FAX: (412) 826-3850 (412) 826-3600 http://www.cellomics.com Celltech 216 Bath Road Slough, Berkshire SL1 4EN, UK (44) 1753 534655 FAX: (44) 1753 536632 http://www.celltech.co.uk Cellular Products 872 Main Street Buffalo, NY 14202 (800) CPI-KITS FAX: (716) 882-0959 (716) 882-0920 http://www.zeptometrix.com CEM P.O. Box 200 Matthews, NC 28106 (800) 726-3331
Centers for Disease Control 1600 Clifton Road NE Atlanta, GA 30333 (800) 311-3435 FAX: (888) 232-3228 (404) 639-3311 http://www.cdc.gov CERJ Centre d’Elevage Roger Janvier 53940 Le Genest Saint Isle France Cetus See Chiron Chance Propper Warly, West Midlands B66 1NZ, UK (44)(0)121-553-5551 FAX: (44)(0)121-525-0139 Charles River Laboratories 251 Ballardvale Street Wilmington, MA 01887 (800) 522-7287 FAX: (978) 658-7132 (978) 658-6000 http://www.criver.com Charm Sciences 36 Franklin Street Malden, MA 02148 (800) 343-2170 FAX: (781) 322-3141 (781) 322-1523 http://www.charm.com Chase-Walton Elastomers 29 Apsley Street Hudson, MA 01749 (800) 448-6289 FAX: (978) 562-5178 (978) 568-0202 http://www.chase-walton.com ChemGenes Ashland Technology Center 200 Homer Avenue Ashland, MA 01721 (800) 762-9323 FAX: (508) 881-3443 (508) 881-5200 http://www.chemgenes.com Chemglass 3861 North Mill Road Vineland, NJ 08360 (800) 843-1794 FAX: (856) 696-9102 (800) 696-0014 http://www.chemglass.com
Chem Service P.O. Box 599 West Chester, PA 19381-0599 (610) 692-3026 FAX: (610) 692-8729 http://www.chemservice.com Chemsyn Laboratories 13605 West 96th Terrace Lenexa, Kansas 66215 (913) 541-0525 FAX: (913) 888-3582 http://www.tech.epcorp.com/ChemSyn/ chemsyn.htm Chemunex USA 1 Deer Park Drive, Suite H-2 Monmouth Junction, NJ 08852 (800) 411-6734 http://www.chemunex.com Cherwell Scientific Publishing The Magdalen Centre Oxford Science Park Oxford OX44GA, UK (44)(1) 865-784-800 FAX: (44)(1) 865-784-801 http://www.cherwell.com Chimera Biotec GmbH Dortmund BioMedicine Center Emil-Figge-Str. 76 A D-44227 Dortmund Germany 49 (0)231 9742-840 FAX: 49 (0)231 9742-844 http://www.chimerabiotec.de/index.htm
Chromatographie ZAC de Moulin No. 2 91160 Saulx les Chartreux France (33) 01-64-54-8969 FAX: (33) 01-69-0988091 http://www.chromatographie.com Chromogenix Taljegardsgatan 3 431-53 Mlndal, Sweden (46) 31-706-20-70 FAX: (46) 31-706-20-80 http://www.chromogenix.com Chrompack USA c/o Varian USA 2700 Mitchell Drive Walnut Creek, CA 94598 (800) 526-3687 FAX: (925) 945-2102 (925) 939-2400 http://www.chrompack.com Chugai Biopharmaceuticals 6275 Nancy Ridge Drive San Diego, CA 92121 (858) 535-5900 FAX: (858) 546-5973 http://www.chugaibio.com Ciba-Corning Diagnostics See Bayer Diagnostics Ciba-Geigy See Ciba Specialty Chemicals or Novartis Biotechnology
ChiRex Cauldron 383 Phoenixville Pike Malvern, PA 19355 (610) 727-2215 FAX: (610) 727-5762 http://www.chirex.com Chiron Diagnostics See Bayer Diagnostics Chiron Mimotopes Peptide Systems See Multiple Peptide Systems Chiron 4560 Horton Street Emeryville, CA 94608 (800) 244-7668 FAX: (510) 655-9910 (510) 655-8730 http://www.chiron.com
Chemicon International 28835 Single Oak Drive Temecula, CA 92590 (800) 437-7500 FAX: (909) 676-9209 (909) 676-8080 http://www.chemicon.com
Chrom Tech P.O. Box 24248 Apple Valley, MN 55124 (800) 822-5242 FAX: (952) 431-6345 http://www.chromtech.com
Chem-Impex International 935 Dillon Drive Wood Dale, IL 60191 (800) 869-9290 FAX: (630) 766-2218 (630) 766-2112 http://www.chemimpex.com
Chroma Technology 72 Cotton Mill Hill, Unit A-9 Brattleboro, VT 05301 (800) 824-7662 FAX: (802) 257-9400 (802) 257-1800 http://www.chroma.com
Ciba Specialty Chemicals 540 White Plains Road Tarrytown, NY 10591 (800) 431-1900 FAX: (914) 785-2183 (914) 785-2000 http://www.cibasc.com CIBA Vision Division of Novartis AG 11460 Johns Creek Parkway Duluth, GA 30097 (770) 476-3937 http://www.cvworld.com Cinna Scientific Subsidiary of Molecular Research Center 5645 Montgomery Road Cincinnati, OH 45212 (800) 462-9868 FAX: (513) 841-0080 (513) 841-0900 http://www.mrcgene.com Cistron Biotechnology 10 Bloomfield Avenue Pine Brook, NJ 07058 (800) 642-0167 FAX: (973) 575-4854 (973) 575-1700 http://www.cistronbio.com Clark Electromedical Instruments See Harvard Apparatus Clay Adam See Becton Dickinson Primary Care Diagnostics
Suppliers
8 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
CLB (Central Laboratory of the Netherlands) Blood Transfusion Service P.O. Box 9190 1006 AD Amsterdam, The Netherlands (31) 20-512-9222 FAX: (31) 20-512-3332 Cleveland Scientific P.O. Box 300 Bath, OH 44210 (800) 952-7315 FAX: (330) 666-2240 http:://www.clevelandscientific.com Clonetics Division of BioWhittaker http://www.clonetics.com Also see BioWhittaker Clontech Laboratories 1020 East Meadow Circle Palo Alto, CA 94303 (800) 662-2566 FAX: (800) 424-1350 (650) 424-8222 FAX: (650) 424-1088 http://www.clontech.com CMA Microdialysis AB 73 Princeton Street North Chelmsford, MA 01863 (800) 440-4980 FAX: (978) 251-1950 (978) 251-1940 http://www.microdialysis.com Cocalico Biologicals 449 Stevens Road P.O. Box 265 Reamstown, PA 17567 (717) 336-1990 FAX: (717) 336-1993 Coherent Laser 5100 Patrick Henry Drive Santa Clara, CA 95056 (800) 227-1955 FAX: (408) 764-4800 (408) 764-4000 http://www.cohr.com Cohu P.O. Box 85623 San Diego, CA 92186 (858) 277-6700 FAX: (858) 277-0221 http://www.COHU.com/cctv Cole-Parmer Instrument 625 East Bunker Court Vernon Hills, IL 60061 (800) 323-4340 FAX: (847) 247-2929 (847) 549-7600 http://www.coleparmer.com Collaborative Biomedical Products and Collaborative Research See Becton Dickinson Labware Collagen Aesthetics 1850 Embarcadero Road Palo Alto, CA 94303 (650) 856-0200 FAX: (650) 856-0533 http://www.collagen.com Collagen Corporation See Collagen Aesthetics
College of American Pathologists 325 Waukegan Road Northfield, IL 60093 (800) 323-4040 FAX: (847) 832-8000 (847) 446-8800 http://www.cap.org/index.cfm
Cooper Instruments & Systems P.O. Box 3048 Warrenton, VA 20188 (800) 344-3921 FAX: (540) 347-4755 (540) 349-4746 http://www.cooperinstruments.com
Colonial Medical Supply 504 Wells Road Franconia, NH 03580 (603) 823-9911 FAX: (603) 823-8799 http://www.colmedsupply.com
Cora Styles Needles ’N Blocks 56 Milton Street Arlington, MA 02474 (781) 648-6289 FAX: (781) 641-7917
Colorado Serum 4950 York Street Denver, CO 80216 (800) 525-2065 FAX: (303) 295-1923 http://www.colorado-serum.com Columbia Diagnostics 8001 Research Way Springfield, VA 22153 (800) 336-3081 FAX: (703) 569-2353 (703) 569-7511 http://www.columbiadiagnostics.com Columbus Instruments 950 North Hague Avenue Columbus, OH 43204 (800) 669-5011 FAX: (614) 276-0529 (614) 276-0861 http://www.columbusinstruments.com Computer Associates International One Computer Associates Plaza Islandia, NY 11749 (631) 342-6000 FAX: (631) 342-6800 http://www.cai.com Connaught Laboratories See Aventis Pasteur Connectix 2955 Campus Drive, Suite 100 San Mateo, CA 94403 (800) 950-5880 FAX: (650) 571-0850 (650) 571-5100 http://www.connectix.com Contech 99 Hartford Avenue Providence, RI 02909 (401) 351-4890 FAX: (401) 421-5072 http://www.iol.ie/∼burke/contech.html Continental Laboratory Products 5648 Copley Drive San Diego, CA 92111 (800) 456-7741 FAX: (858) 279-5465 (858) 279-5000 http://www.conlab.com ConvaTec Professional Services P.O. Box 5254 Princeton, NJ 08543 (800) 422-8811 http://www.convatec.com
Coriell Cell Repository (CCR) See Coriell Institute for Medical Research Coriell Institute for Medical Research Human Genetic Mutant Repository 401 Haddon Avenue Camden, NJ 08103 (856) 966-7377 FAX: (856) 964-0254 http://arginine.umdnj.edu Corion 8 East Forge Parkway Franklin, MA 02038 (508) 528-4411 FAX: (508) 520-7583 (800) 598-6783 http://www.corion.com Corning and Corning Science Products P.O. Box 5000 Corning, NY 14831 (800) 222-7740 FAX: (607) 974-0345 (607) 974-9000 http://www.corning.com Costar See Corning Coulbourn Instruments 7462 Penn Drive Allentown, PA 18106 (800) 424-3771 FAX: (610) 391-1333 (610) 395-3771 http://www.coulbourninst.com Coulter Cytometry See Beckman Coulter Covance Research Products 465 Swampbridge Road Denver, PA 17517 (800) 345-4114 FAX: (717) 336-5344 (717) 336-4921 http://www.covance.com
CPL Scientific 43 Kingfisher Court Hambridge Road Newbury RG14 5SJ, UK (44) 1635-574902 FAX: (44) 1635-529322 http://www.cplscientific.co.uk CraMar Technologies 8670 Wolff Court, #160 Westminster, CO 80030 (800) 4-TOMTEC http://www.cramar.com Crescent Chemical 1324 Motor Parkway Hauppauge, NY 11788 (800) 877-3225 FAX: (631) 348-0913 (631) 348-0333 http://www.creschem.com Crist Instrument P.O. Box 128 10200 Moxley Road Damascus, MD 20872 (301) 253-2184 FAX: (301) 253-0069 http://www.cristinstrument.com Cruachem See Annovis http://www.cruachem.com CS Bio 1300 Industrial Road San Carlos, CA 94070 (800) 627-2461 FAX: (415) 802-0944 (415) 802-0880 http://www.csbio.com CS-Chromatographie Service Am Parir 27 D-52379 Langerwehe, Germany (49) 2423-40493-0 FAX: (49) 2423-40493-49 http://www.cs-chromatographie.de Cuno 400 Research Parkway Meriden, CT 06450 (800) 231-2259 FAX: (203) 238-8716 (203) 237-5541 http://www.cuno.com Curtin Matheson Scientific 9999 Veterans Memorial Drive Houston, TX 77038 (800) 392-3353 FAX: (713) 878-3598 (713) 878-3500
Coy Laboratory Products 14500 Coy Drive Grass Lake, MI 49240 (734) 475-2200 FAX: (734) 475-1846 http://www.coylab.com
CWE 124 Sibley Ave Ardmore, PA 19003 (610) 642-7719 FAX: (610) 642-1532 http://www.cwe-inc.com
CPG 3 Borinski Road Lincoln Park, NJ 07035 (800) 362-2740 FAX: (973) 305-0884 (973) 305-8181
Cybex Computer Products 4991 Corporate Drive Huntsville, AL 35805 (800) 932-9239 FAX: (800) 462-9239 http://www.cybex.com
Suppliers
9 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Cygnus Technology P.O. Box 219 Delaware Water Gap, PA 18327 (570) 424-5701 FAX: (570) 424-5630 http://www.cygnustech.com Cymbus Biotechnology Eagle Class, Chandler’s Ford Hampshire SO53 4NF, UK (44) 1-703-267-676 FAX: (44) 1-703-267-677 http://www.biotech.cymbus.com Cytogen 600 College Road East Princeton, NJ 08540 (609) 987-8200 FAX: (609) 987-6450 http://www.cytogen.com Cytogen Research and Development 89 Bellevue Hill Road Boston, MA 02132 (617) 325-7774 FAX: (617) 327-2405 CytRx 154 Technology Parkway Norcross, GA 30092 (800) 345-2987 FAX: (770) 368-0622 (770) 368-9500 http://www.cytrx.com Dade Behring 1717 Deerfield Road Deerfield, IL 60015 (847) 267-5300 FAX: (847) 267-1066 http://www.dadebehring.com Dagan 2855 Park Avenue Minneapolis, MN 55407 (612) 827-5959 FAX: (612) 827-6535 http://www.dagan.com Daiso 807 Aldo Avenue Suite 104 Santa Clara, CA 95054 (408) 855-8789 FAX: (408) 855-8784 http://www.daiso.co.jp Dako 6392 Via Real Carpinteria, CA 93013 (800) 235-5763 FAX: (805) 566-6688 (805) 566-6655 http://www.dakousa.com Dako A/S 42 Produktionsvej P.O. Box 1359 DK-2600 Glostrup, Denmark (45) 4492-0044 FAX: (45) 4284-1822
Dan Kar Scientific 150 West Street Wilmington, MA 01887 (800) 942-5542 FAX: (978) 658-0380 (978) 988-9696 http://www.dan-kar.com DataCell Falcon Business Park 40 Ivanhoe Road Finchampstead, Berkshire RG40 4QQ, UK (44) 1189 324324 FAX: (44) 1189 324325 http://www.datacell.co.uk In the US: (408) 446-3575 FAX: (408) 446-3589 http://www.datacell.com
Damon, IEC See Thermoquest
Diagnostic Instruments 6540 Burroughs Sterling Heights, MI 48314 (810) 731-6000 FAX: (810) 731-6469 http://www.diaginc.com
Deneba Software 1150 NW 72nd Avenue Miami, FL 33126 (305) 596-5644 FAX: (305) 273-9069 http://www.deneba.com Deseret Medical 524 West 3615 South Salt Lake City, UT 84115 (801) 270-8440 FAX: (801) 293-9000
DataWave Technologies 380 Main Street, Suite 209 Longmont, CO 80501 (800) 736-9283 FAX: (303) 776-8531 (303) 776-8214
Devcon Plexus 30 Endicott Street Danvers, MA 01923 (800) 626-7226 FAX: (978) 774-0516 (978) 777-1100 http://www.devcon.com
Datex-Ohmeda 3030 Ohmeda Drive Madison, WI 53718 (800) 345-2700 FAX: (608) 222-9147 (608) 221-1551 http://www.us.datex-ohmeda.com
Developmental Studies Hybridoma Bank University of Iowa 436 Biology Building Iowa City, IA 52242 (319) 335-3826 FAX: (319) 335-2077 http://www.uiowa.edu/∼dshbwww
DATU 82 State Street Geneva, NY 14456 (315) 787-2240 FAX: (315) 787-2397 http://www.nysaes.cornell.edu/datu
DeVilbiss Division of Sunrise Medical Respiratory 100 DeVilbiss Drive P.O. Box 635 Somerset, PA 15501 (800) 338-1988 FAX: (814) 443-7572 (814) 443-4881 http://www.sunrisemedical.com
David Kopf Instruments 7324 Elmo Street P.O. Box 636 Tujunga, CA 91043 (818) 352-3274 FAX: (818) 352-3139
Dharmacon Research 3200 Valmont Road, #5 Boulder, CO 80301 (800) 235-9880 FAX: (303) 415-9879 (303) 415-9880 http://www.dharmacon.com
Decagon Devices P.O. Box 835 950 NE Nelson Court Pullman, WA 99163 (800) 755-2751 FAX: (509) 332-5158 (509) 332-2756 http://www.decagon.com
DiaCheM Triangle Biomedical Gardiners Place West Gillibrands, Lancashire WN8 9SP, UK (44) 1695-555581 FAX: (44) 1695-555518 http://www.diachem.co.uk
Decon Labs 890 Country Line Road Bryn Mawr, PA 19010 (800) 332-6647 FAX: (610) 964-0650 (610) 520-0610 http://www.deconlabs.com
Diagen Max-Volmer Strasse 4 D-40724 Hilden, Germany (49) 2103-892-230 FAX: (49) 2103-892-222
Decon Laboratories Conway Street Hove, Sussex BN3 3LY, UK (44) 1273 739241 FAX: (44) 1273 722088
Dakopatts See Dako A/S
Degussa Precious Metals Division 3900 South Clinton Avenue South Plainfield, NJ 07080 (800) DEGUSSA FAX: (908) 756-7176 (908) 561-1100 http://www.degussa-huls.com
Diagnostic Concepts 6104 Madison Court Morton Grove, IL 60053 (847) 604-0957 Diagnostic Developments See DiaCheM
Diamedix 2140 North Miami Avenue Miami, FL 33127 (800) 327-4565 FAX: (305) 324-2395 (305) 324-2300 DiaSorin 1990 Industrial Boulevard Stillwater, MN 55082 (800) 328-1482 FAX: (651) 779-7847 (651) 439-9719 http://www.diasorin.com Diatome US 321 Morris Road Fort Washington, PA 19034 (800) 523-5874 FAX: (215) 646-8931 (215) 646-1478 http://www.emsdiasum.com Difco Laboratories See Becton Dickinson Digene 1201 Clopper Road Gaithersburg, MD 20878 (301) 944-7000 (800) 344-3631 FAX: (301) 944-7121 http://www.digene.com Digi-Key 701 Brooks Avenue South Thief River Falls, MN 56701 (800) 344-4539 FAX: (218) 681-3380 (218) 681-6674 http://www.digi-key.com Digitimer 37 Hydeway Welwyn Garden City, Hertfordshire AL7 3BE, UK (44) 1707-328347 FAX: (44) 1707-373153 http://www.digitimer.com Dimco-Gray 8200 South Suburban Road Dayton, OH 45458 (800) 876-8353 FAX: (937) 433-0520 (937) 433-7600 http://www.dimco-gray.com Dionex 1228 Titan Way P.O. Box 3603 Sunnyvale, CA 94088 (408) 737-0700 FAX: (408) 730-9403 http://dionex2.promptu.com Display Systems Biotech 1260 Liberty Way, Suite B Vista, CA 92083 (800) 697-1111 FAX: (760) 599-9930 (760) 599-0598 http://www.displaysystems.com
Suppliers
10 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
Diversified Biotech 1208 VFW Parkway Boston, MA 02132 (617) 965-8557 FAX: (617) 323-5641 (800) 796-9199 http://www.divbio.com DNA ProScan P.O. Box 121585 Nashville, TN 37212 (800) 841-4362 FAX: (615) 292-1436 (615) 298-3524 http://www.dnapro.com DNAStar 1228 South Park Street Madison, WI 53715 (608) 258-7420 FAX: (608) 258-7439 http://www.dnastar.com DNAVIEW Attn: Charles Brenner http://www.wco.com ∼cbrenner/dnaview.htm Doall NYC 36-06 48th Avenue Long Island City, NY 11101 (718) 392-4595 FAX: (718) 392-6115 http://www.doall.com Dojindo Molecular Technologies 211 Perry Street Parkway, Suite 5 Gaitherbusburg, MD 20877 (877) 987-2667 http://www.dojindo.com Dolla Eastern See Doall NYC Dolan Jenner Industries 678 Andover Street Lawrence, MA 08143 (978) 681-8000 (978) 682-2500 http://www.dolan-jenner.com Dow Chemical Customer Service Center 2040 Willard H. Dow Center Midland, MI 48674 (800) 232-2436 FAX: (517) 832-1190 (409) 238-9321 http://www.dow.com Dow Corning Northern Europe Meriden Business Park Copse Drive Allesley, Coventry CV5 9RG, UK (44) 1676 528 000 FAX: (44) 1676 528 001 Dow Corning P.O. Box 994 Midland, MI 48686 (517) 496-4000 http://www.dowcorning.com Dow Corning (Lubricants) 2200 West Salzburg Road Auburn, MI 48611 (800) 248-2481 FAX: (517) 496-6974 (517) 496-6000
Dremel 4915 21st Street Racine, WI 53406 (414) 554-1390 http://www.dremel.com Drummond Scientific 500 Parkway P.O. Box 700 Broomall, PA 19008 (800) 523-7480 FAX: (610) 353-6204 (610) 353-0200 http://www.drummondsci.com Duchefa Biochemie BV P.O. Box 2281 2002 CG Haarlem, The Netherlands 31-0-23-5319093 FAX: 31-0-23-5318027 http://www.duchefa.com Duke Scientific 2463 Faber Place Palo Alto, CA 94303 (800) 334-3883 FAX: (650) 424-1158 (650) 424-1177 http://www.dukescientific.com DuPont Biotechnology Systems See NEN Life Science Products DuPont Medical Products See NEN Life Science Products DuPont Merck Pharmaceuticals 331 Treble Cove Road Billerica, MA 01862 (800) 225-1572 FAX: (508) 436-7501 http://www.dupontmerck.com DuPont NEN Products See NEN Life Science Products Dynal 5 Delaware Drive Lake Success, NY 11042 (800) 638-9416 FAX: (516) 326-3298 (516) 326-3270 http://www.dynal.net Dynal AS Ullernchausen 52, 0379 Oslo, Norway 47-22-06-10-00 FAX: 47-22-50-70-15 http://www.dynal.no Dynalab P.O. Box 112 Rochester, NY 14692 (800) 828-6595 FAX: (716) 334-9496 (716) 334-2060 http://www.dynalab.com Dynarex 1 International Boulevard Brewster, NY 10509 (888) DYNAREX FAX: (914) 279-9601 (914) 279-9600 http://www.dynarex.com Dynatech See Dynex Technologies
Dynex Technologies 14340 Sullyfield Circle Chantilly, VA 22021 (800) 336-4543 FAX: (703) 631-7816 (703) 631-7800 http://www.dynextechnologies.com Dyno Mill See Willy A. Bachofen E.S.A. 22 Alpha Road Chelmsford, MA 01824 (508) 250-7000 FAX: (508) 250-7090 E.W. Wright 760 Durham Road Guilford, CT 06437 (203) 453-6410 FAX: (203) 458-6901 http://www.ewwright.com E-Y Laboratories 107 N. Amphlett Boulevard San Mateo, CA 94401 (800) 821-0044 FAX: (650) 342-2648 (650) 342-3296 http://www.eylabs.com Eastman Kodak 1001 Lee Road Rochester, NY 14650 (800) 225-5352 FAX: (800) 879-4979 (716) 722-5780 FAX: (716) 477-8040 http://www.kodak.com ECACC See European Collection of Animal Cell Cultures EC Apparatus See Savant/EC Apparatus Ecogen, SRL Gensura Laboratories Ptge. Dos de Maig 9(08041) Barcelona, Spain (34) 3-450-2601 FAX: (34) 3-456-0607 http://www.ecogen.com Ecolab 370 North Wabasha Street St. Paul, MN 55102 (800) 35-CLEAN FAX: (651) 225-3098 (651) 352-5326 http://www.ecolab.com ECO PHYSICS 3915 Research Park Drive, Suite A-3 Ann Arbor, MI 48108 (734) 998-1600 FAX: (734) 998-1180 http://www.ecophysics.com Edge Biosystems 19208 Orbit Drive Gaithersburg, MD 20879-4149 (800) 326-2685 FAX: (301) 990-0881 (301) 990-2685 http://www.edgebio.com
Edmund Buhler GmbH & Co. Rottenburger Str. 3 P.O. Box 12 24 D-7454 Bodelshausen Germany (49) 74717070 FAX: (49) 747170788 http://www.edmund-buehler.de/ Edmund Scientific 101 E. Gloucester Pike Barrington, NJ 08007 (800) 728-6999 FAX: (856) 573-6263 (856) 573-6250 http://www.edsci.com EG&G See Perkin-Elmer Ekagen 969 C Industry Road San Carlos, CA 94070 (650) 592-4500 FAX: (650) 592-4500 Elcatech P.O. Box 10935 Winston-Salem, NC 27108 (336) 544-8613 FAX: (336) 777-3623 (910) 777-3624 http://www.elcatech.com Electron Microscopy Sciences 321 Morris Road Fort Washington, PA 19034 (800) 523-5874 FAX: (215) 646-8931 (215) 646-1566 http://www.emsdiasum.com Electron Tubes 100 Forge Way, Unit F Rockaway, NJ 07866 (800) 521-8382 FAX: (973) 586-9771 (973) 586-9594 http://www.electrontubes.com Elicay Laboratory Products, (UK) Ltd. 4 Manborough Mews Crockford Lane Basingstoke, Hampshire RG 248NA, England (256) 811-118 FAX: (256) 811-116 http://www.elkay-uk.co.uk Eli Lilly Lilly Corporate Center Indianapolis, IN 46285 (800) 545-5979 FAX: (317) 276-2095 (317) 276-2000 http://www.lilly.com ELISA Technologies See Neogen Elkins-Sinn See Wyeth-Ayerst EMBI See European Bioinformatics Institute
Suppliers
11 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
EM Science 480 Democrat Road Gibbstown, NJ 08027 (800) 222-0342 FAX: (856) 423-4389 (856) 423-6300 http://www.emscience.com EM Separations Technology See R & S Technology Endogen 30 Commerce Way Woburn, MA 01801 (800) 487-4885 FAX: (617) 439-0355 (781) 937-0890 http://www.endogen.com ENGEL-Loter HSGM Heatcutting Equipment & Machines 1865 E. Main Street, No. 5 Duncan, SC 29334 (888) 854-HSGM FAX: (864) 486-8383 (864) 486-8300 http://www.engelgmbh.com Enzo Diagnostics 60 Executive Boulevard Farmingdale, NY 11735 (800) 221-7705 FAX: (516) 694-7501 (516) 694-7070 http://www.enzo.com Enzogenetics 4197 NW Douglas Avenue Corvallis, OR 97330 (541) 757-0288 The Enzyme Center See Charm Sciences Enzyme Systems Products 486 Lindbergh Avenue Livermore, CA 94550 (888) 449-2664 FAX: (925) 449-1866 (925) 449-2664 http://www.enzymesys.com Epicentre Technologies 1402 Emil Street Madison, WI 53713 (800) 284-8474 FAX: (608) 258-3088 (608) 258-3080 http://www.epicentre.com Carlo Erba Reagents P.O. Box 63 I-20090 Limito Milan, Italy (39)0295325357 FAX: (39)0295325330 http://carloerbareagenti.com Erie Scientific 20 Post Road Portsmouth, NH 03801 (888) ERIE-SCI FAX: (603) 431-8996 (603) 431-8410 http://www.eriesci.com
ES Industries 701 South Route 73 West Berlin, NJ 08091 (800) 356-6140 FAX: (856) 753-8484 (856) 753-8400 http://www.esind.com ESA 22 Alpha Road Chelmsford, MA 01824 (800) 959-5095 FAX: (978) 250-7090 (978) 250-7000 http://www.esainc.com Ethicon Route 22, P.O. Box 151 Somerville, NJ 08876 (908) 218-0707 http://www.ethiconinc.com Ethicon Endo-Surgery 4545 Creek Road Cincinnati, OH 45242 (800) 766-9534 FAX: (513) 786-7080 Eurogentec 3347 Industrial Ct, Suite A San Diego, CA 92121 (877) 387-6436 FAX: (858) 793-2666 (858) 793-2661 http://www.eurogentec.com European Bioinformatics Institute Wellcome Trust Genomes Campus Hinxton, Cambridge CB10 1SD, UK (44) 1223-49444 FAX: (44) 1223-494468 European Collection of Animal Cell Cultures (ECACC) Centre for Applied Microbiology & Research Salisbury, Wiltshire SP4 0JG, UK (44) 1980-612 512 FAX: (44) 1980-611 315 http://www.camr.org.uk Evergreen Scientific 2254 E. 49th Street P.O. Box 58248 Los Angeles, CA 90058 (800) 421-6261 FAX: (323) 581-2503 (323) 583-1331 http://www.evergreensci.com Exalpha Biologicals 20 Hampden Street Boston, MA 02205 (800) 395-1137 FAX: (617) 969-3872 (617) 558-3625 http://www.exalpha.com Exciton P.O. Box 31126 Dayton, OH 45437 (937) 252-2989 FAX: (937) 258-3937 http://www.exciton.com
Extrasynthese ZI Lyon Nord SA-BP62 69730 Genay, France (33) 78-98-20-34 FAX: (33) 78-98-19-45
Fisher Scientific 2000 Park Lane Pittsburgh, PA 15275 (800) 766-7000 FAX: (800) 926-1166 (412) 562-8300 http://www3.fishersci.com
Factor II 1972 Forest Avenue P.O. Box 1339 Lakeside, AZ 85929 (800) 332-8688 FAX: (520) 537-8066 (520) 537-8387 http://www.factor2.com
Fitzco 5600 Pioneer Creek Drive Maple Plain, MN 55359 (800) 367-8760 FAX: (612) 479-2880 (612) 479-3489 http://www.fitzco.com 5 Prime → 3 Prime See 2000 Eppendorf-5 Prime http://www.5prime.com
Falcon See Becton Dickinson Labware Fenwal See Baxter Healthcare Fidelity Systems 7961 Cessna Avenue Gaithersburg, MD 20879-4117 (301) 527-0804 FAX: (301) 527-8250 http://www.fidelitysystems.com Filemaker 5201 Patrick Henry Drive Santa Clara, CA 95054 (408) 987-7000 (800) 325-2747 Fine Science Tools 202-277 Mountain Highway North Vancouver, British Columbia V7J 3P2 Canada (800) 665-5355 FAX: (800) 665 4544 (604) 980-2481 FAX: (604) 987-3299 Fine Science Tools 373-G Vintage Park Drive Foster City, CA 94404 (800) 521-2109 FAX: (800) 523-2109 (650) 349-1636 FAX: (630) 349-3729 Fine Science Tools Fahrtgasse 7-13 D-69117 Heidelberg, Germany (49) 6221 905050 FAX: (49) 6221 600001 http://www.finescience.com Finn Aqua AMSCO Finn Aqua Oy Teollisuustiez, FIN-04300 Tuusula, Finland 358 025851 FAX: 358 0276019
Fleisch (Rusch) 2450 Meadowbrook Parkway Duluth, GA 30096 (770) 623-0816 FAX: (770) 623-1829 http://ruschinc.com Flow Cytometry Standards P.O. Box 194344 San Juan, PR 00919 (800) 227-8143 FAX: (787) 758-3267 (787) 753-9341 http://www.fcstd.com Flow Labs See ICN Biomedicals Flow-Tech Supply P.O. Box 1388 Orange, TX 77631 (409) 882-0306 FAX: (409) 882-0254 http://www.flow-tech.com Fluid Marketing See Fluid Metering Fluid Metering 5 Aerial Way, Suite 500 Sayosett, NY 11791 (516) 922-6050 FAX: (516) 624-8261 http://www.fmipump.com Fluorochrome 1801 Williams, Suite 300 Denver, CO 80264 (303) 394-1000 FAX: (303) 321-1119 Fluka Chemical See Sigma-Aldrich
Finnigan 355 River Oaks Parkway San Jose, CA 95134 (408) 433-4800 FAX: (408) 433-4821 http://www.finnigan.com
FMC BioPolymer 1735 Market Street Philadelphia, PA 19103 (215) 299-6000 FAX: (215) 299-5809 http://www.fmc.com
Fisher Chemical Company Fisher Scientific Limited 112 Colonnade Road Nepean Ontario K2E 7L6, Canada (800) 234-7437 FAX: (800) 463-2996 http://www.fisherscientific.com
FMC BioProducts 191 Thomaston Street Rockland, ME 04841 (800) 521-0390 FAX: (800) 362-1133 (207) 594-3400 FAX: (207) 594-3426 http://www.bioproducts.com
Suppliers
12 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
Forma Scientific Milcreek Road P.O. Box 649 Marietta, OH 45750 (800) 848-3080 FAX: (740) 372-6770 (740) 373-4765 http://www.forma.com Fort Dodge Animal Health 800 5th Street NW Fort Dodge, IA 50501 (800) 685-5656 FAX: (515) 955-9193 (515) 955-4600 http://www.ahp.com Fotodyne 950 Walnut Ridge Drive Hartland, WI 53029 (800) 362-3686 FAX: (800) 362-3642 (262) 369-7000 FAX: (262) 369-7013 http://www.fotodyne.com Fresenius HemoCare 6675 185th Avenue NE, Suite 100 Redwood, WA 98052 (800) 909-3872 (425) 497-1197 http://www.freseniusht.com Fresenius Hemotechnology See Fresenius HemoCare Fuji Medical Systems 419 West Avenue P.O. Box 120035 Stamford, CT 06902 (800) 431-1850 FAX: (203) 353-0926 (203) 324-2000 http://www.fujimed.com Fuji Silysia Chemical Ltd. 2-1846 Kozoji-cho Kasugai-shi Aichi-ken, Japan 487-0013 (81) 568-51-2516 FAX: (81) 568-51-8557 http://www.fujisilysia.co.jp/english/index.html Fujisawa USA Parkway Center North Deerfield, IL 60015-2548 (847) 317-1088 FAX: (847) 317-7298 Ernest F. Fullam 900 Albany Shaker Road Latham, NY 12110 (800) 833-4024 FAX: (518) 785-8647 (518) 785-5533 http://www.fullam.com Gallard-Schlesinger Industries 777 Zechendorf Boulevard Garden City, NY 11530 (516) 229-4000 FAX: (516) 229-4015 http://www.gallard-schlessinger.com
Gambro Box 7373 SE 103 91 Stockholm, Sweden (46) 8 613 65 00 FAX: (46) 8 611 37 31 In the US: COBE Laboratories 225 Union Blvd. Lakewood, CO 80215 (303) 232-6800 FAX: (303) 231-4915 http://www.gambro.com Garner Glass 177 Indian Hill Boulevard Claremont, CA 91711 (909) 624-5071 FAX: (909) 625-0173 http://www.garnerglass.com Garon Plastics 16 Byre Avenue Somerton Park, South Australia 5044 (08) 8294-5126 FAX: (08) 8376-1487 http://www.apache.airnet.com.au/ ∼garon Garren Scientific 9400 Lurline Avenue, Unit E Chatsworth, CA 91311 (800) 342-3725 FAX: (818) 882-3229 (818) 882-6544 http://www.garren-scientific.com GATC Biotech AG Jakob-Stadler-Platz 7 D-78467 Constance, Germany (49) 07531-8160-0 FAX: (49) 07531-8160-81 http://www.gatc-biotech.com Gaussian Carnegie Office Park Building 6, Suite 230 Carnegie, PA 15106 (412) 279-6700 FAX: (412) 279-2118 http://www.gaussian.com G.C. Electronics/A.R.C. Electronics 431 Second Street Henderson, KY 42420 (270) 827-8981 FAX: (270) 827-8256 http://www.arcelectronics.com GDB (Genome Data Base Curation) 2024 East Monument Street Suite 1200 Baltimore, MD 21205 (410) 955-9705 FAX: (410) 614-0434 http://www.gdb.org GDB (Genome Data Base, Home) Hospital for Sick Children 555 University Avenue Toronto, Ontario M5G 1X8 Canada (416) 813-8744 FAX: (416) 813-8755 http://www.gdb.org Gelman Sciences See Pall-Gelman
Gemini BioProducts 5115-M Douglas Fir Road Calabasas, CA 90403 (818) 591-3530 FAX: (818) 591-7084 Gen Trak 5100 Campus Drive Plymouth Meeting, PA 19462 (800) 221-7407 FAX: (215) 941-9498 (215) 825-5115 http://www.informagen.com Genaissance Pharmaceuticals 5 Science Park New Haven, CT 06511 (800) 678-9487 FAX: (203) 562-9377 (203) 773-1450 http://www.genaissance.com GENAXIS Biotechnology Parc Technologique 10 Avenue Amp`ere Montigny le Bretoneux 78180 France (33) 01-30-14-00-20 FAX: (33) 01-30-14-00-15 http://www.genaxis.com GenBank National Center for Biotechnology Information National Library of Medicine/NIH Building 38A, Room 8N805 8600 Rockville Pike Bethesda, MD 20894 (301) 496-2475 FAX: (301) 480-9241 http://www.ncbi.nlm.nih.gov Gene Codes 640 Avis Drive Ann Arbor, MI 48108 (800) 497-4939 FAX: (734) 930-0145 (734) 769-7249 http://www.genecodes.com Genemachines 935 Washington Street San Carlos, CA 94070 (650) 508-1634 FAX: (650) 508-1644 (877) 855-4363 http://www.genemachines.com Genentech 1 DNA Way South San Francisco, CA 94080 (800) 551-2231 FAX: (650) 225-1600 (650) 225-1000 http://www.gene.com General Scanning/GSI Luminomics 500 Arsenal Street Watertown, MA 02172 (617) 924-1010 FAX: (617) 924-7327 http://www.genescan.com
General Valve Division of Parker Hannifin Pneutronics 19 Gloria Lane Fairfield, NJ 07004 (800) GVC-VALV FAX: (800) GVC-1-FAX http://www.pneutronics.com Genespan 19310 North Creek Parkway, Suite 100 Bothell, WA 98011 (800) 231-2215 FAX: (425) 482-3005 (425) 482-3003 http://www.genespan.com G´en´ethon Human Genome Research Center 1 bis rue de l’Internationale 91000 Evry, France (33) 169-472828 FAX: (33) 607-78698 http://www.genethon.fr Genetic Microsystems 34 Commerce Way Wobum, MA 01801 (781) 932-9333 FAX: (781) 932-9433 http://www.genticmicro.com Genetic Mutant Repository See Coriell Institute for Medical Research Genetic Research Instrumentation Gene House Queenborough Lane Rayne, Braintree, Essex CM7 8TF, UK (44) 1376 332900 FAX: (44) 1376 344724 http://www.gri.co.uk Genetics Computer Group 575 Science Drive Madison, WI 53711 (608) 231-5200 FAX: (608) 231-5202 http://www.gcg.com Genetics Institute/American Home Products 87 Cambridge Park Drive Cambridge, MA 02140 (617) 876-1170 FAX: (617) 876-0388 http://www.genetics.com Genetix 63-69 Somerford Road Christchurch, Dorset BH23 3QA, UK (44) (0) 1202 483900 FAX: (44)(0) 1202 480289 In the US: (877) 436 3849 US FAX: (888) 522 7499 http://www.genetix.co.uk Gene Tools One Summerton Way Philomath, OR 97370 (541) 929-7840 FAX: (541) 929-7841 http://www.gene-tools.com
Suppliers
13 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
GeneWorks P.O. Box 11, Rundle Mall Adelaide, South Australia 5000, Australia 1800 882 555 FAX: (08) 8234 2699 (08) 8234 2644 http://www.geneworks.com Genome Systems (INCYTE) 4633 World Parkway Circle St. Louis, MO 63134 (800) 430-0030 FAX: (314) 427-3324 (314) 427-3222 http://www.genomesystems.com Genomic Solutions 4355 Varsity Drive, Suite E Ann Arbor, MI 48108 (877) GENOMIC FAX: (734) 975-4808 (734) 975-4800 http://www.genomicsolutions.com Genomyx See Beckman Coulter Genosys Biotechnologies 1442 Lake Front Circle, Suite 185 The Woodlands, TX 77380 (281) 363-3693 FAX: (281) 363-2212 http://www.genosys.com Genotech 92 Weldon Parkway St. Louis, MO 63043 (800) 628-7730 FAX: (314) 991-1504 (314) 991-6034 GENSET 876 Prospect Street, Suite 206 La Jolla, CA 92037 (800) 551-5291 FAX: (619) 551-2041 (619) 515-3061 http://www.genset.fr Gensia Laboratories Ltd. 19 Hughes Irvine, CA 92718 (714) 455-4700 FAX: (714) 855-8210 Genta 99 Hayden Avenue, Suite 200 Lexington, MA 02421 (781) 860-5150 FAX: (781) 860-5137 http://www.genta.com GENTEST 6 Henshaw Street Woburn, MA 01801 (800) 334-5229 FAX: (888) 242-2226 (781) 935-5115 FAX: (781) 932-6855 http://www.gentest.com Gentra Systems 15200 25th Avenue N., Suite 104 Minneapolis, MN 55447 (800) 866-3039 FAX: (612) 476-5850 (612) 476-5858 http://www.gentra.com
Genzyme 1 Kendall Square Cambridge, MA 02139 (617) 252-7500 FAX: (617) 252-7600 http://www.genzyme.com See also R&D Systems Genzyme Genetics One Mountain Road Framingham, MA 01701 (800) 255-7357 FAX: (508) 872-9080 (508) 872-8400 http://www.genzyme.com George Tiemann & Co. 25 Plant Avenue Hauppauge, NY 11788 (516) 273-0005 FAX: (516) 273-6199 GIBCO/BRL A Division of Life Technologies 1 Kendall Square Grand Island, NY 14072 (800) 874-4226 FAX: (800) 352-1968 (716) 774-6700 http://www.lifetech.com Gilmont Instruments A Division of Barnant Company 28N092 Commercial Avenue Barrington, IL 60010 (800) 637-3739 FAX: (708) 381-7053 http://barnant.com Gilson 3000 West Beltline Highway P.O. Box 620027 Middletown, WI 53562 (800) 445-7661 (608) 836-1551 http://www.gilson.com Glas-Col Apparatus P.O. Box 2128 Terre Haute, IN 47802 (800) Glas-Col FAX: (812) 234-6975 (812) 235-6167 http://www.glascol.com Glaxo Wellcome Five Moore Drive Research Triangle Park, NC 27709 (800) SGL-AXO5 FAX: (919) 248-2386 (919) 248-2100 http://www.glaxowellcome.com Glen Mills 395 Allwood Road Clifton, NJ 07012 (973) 777-0777 FAX: (973) 777-0070 http://www.glenmills.com Glen Research 22825 Davis Drive Sterling, VA 20166 (800) 327-4536 FAX: (800) 934-2490 (703) 437-6191 FAX: (703) 435-9774 http://www.glenresearch.com
Glo Germ P.O. Box 189 Moab, UT 84532 (800) 842-6622 FAX: (435) 259-5930 http://www.glogerm.com Glyco 11 Pimentel Court Novato, CA 94949 (800) 722-2597 FAX: (415) 382-3511 (415) 884-6799 http://www.glyco.com Gould Instrument Systems 8333 Rockside Road Valley View, OH 44125 (216) 328-7000 FAX: (216) 328-7400 http://www.gould13.com Gralab Instruments See Dimco-Gray GraphPad Software 5755 Oberlin Drive #110 San Diego, CA 92121 (800) 388-4723 FAX: (558) 457-8141 (558) 457-3909 http://www.graphpad.com Graseby Anderson See Andersen Instruments http://www.graseby.com Grass Instrument A Division of Astro-Med 600 East Greenwich Avenue W. Warwick, RI 02893 (800) 225-5167 FAX: (877) 472-7749 http://www.grassinstruments.com Greenacre and Misac Instruments Misac Systems 27 Port Wood Road Ware, Hertfordshire SF12 9NJ, UK (44) 1920 463017 FAX: (44) 1920 465136 Greer Labs 639 Nuway Circle Lenois, NC 28645 (704) 754-5237 http://greerlabs.com Greiner Maybachestrasse 2 Postfach 1162 D-7443 Frickenhausen, Germany (49) 0 91 31/80 79 0 FAX: (49) 0 91 31/80 79 30 http://www.erlangen.com/greiner GSI Lumonics 130 Lombard Street Oxnard, CA 93030 (805) 485-5559 FAX: (805) 485-3310 http://www.gsilumonics.com GTE Internetworking 150 Cambridge Park Drive Cambridge, MA 02140 (800) 472-4565 FAX: (508) 694-4861 http://www.bbn.com
GW Instruments 35 Medford Street Somerville, MA 02143 (617) 625-4096 FAX: (617) 625-1322 http://www.gwinst.com H & H Woodworking 1002 Garfield Street Denver, CO 80206 (303) 394-3764 Hacker Instruments 17 Sherwood Lane P.O. Box 10033 Fairfield, NJ 07004 (800) 442-2537 FAX: (973) 808-8281 (973) 226-8450 http://www.hackerinstruments.com Haemenetics 400 Wood Road Braintree, MA 02184 (800) 225-5297 FAX: (781) 848-7921 (781) 848-7100 http://www.haemenetics.com Halocarbon Products P.O. Box 661 River Edge, NJ 07661 (201) 242-8899 FAX: (201) 262-0019 http://halocarbon.com Hamamatsu Photonic Systems A Division of Hamamatsu 360 Foothill Road P.O. Box 6910 Bridgewater, NJ 08807 (908) 231-1116 FAX: (908) 231-0852 http://www.photonicsonline.com Hamilton Company 4970 Energy Way P.O. Box 10030 Reno, NV 89520 (800) 648-5950 FAX: (775) 856-7259 (775) 858-3000 http://www.hamiltoncompany.com Hamilton Thorne Biosciences 100 Cummings Center, Suite 102C Beverly, MA 01915 http://www.hamiltonthorne.com Hampton Research 27631 El Lazo Road Laguna Niguel, CA 92677 (800) 452-3899 FAX: (949) 425-1611 (949) 425-6321 http://www.hamptonresearch.com Harlan Bioproducts for Science P.O. Box 29176 Indianapolis, IN 46229 (317) 894-7521 FAX: (317) 894-1840 http://www.hbps.com
Suppliers
14 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
Harlan Sera-Lab Hillcrest, Dodgeford Lane Belton, Loughborough Leicester LE12 9TE, UK (44) 1530 222123 FAX: (44) 1530 224970 http://www.harlan.com Harlan Teklad P.O. Box 44220 Madison, WI 53744 (608) 277-2070 FAX: (608) 277-2066 http://www.harlan.com Harrison Research 840 Moana Court Palo Alto, CA 94306 (650) 949-1565 FAX: (650) 948-0493 Harvard Apparatus 84 October Hill Road Holliston, MA 01746 (800) 272-2775 FAX: (508) 429-5732 (508) 893-8999 http://harvardapparatus.com Harvard Bioscience See Harvard Apparatus Haselton Biologics See JRH Biosciences Hazelton Research Products See Covance Research Products Health Products See Pierce Chemical Heat Systems-Ultrasonics 1938 New Highway Farmingdale, NY 11735 (800) 645-9846 FAX: (516) 694-9412 (516) 694-9555 Heidenhain Corp 333 East State Parkway Schaumberg, IL 60173 (847) 490-1191 FAX: (847) 490-3931 http://www.heidenhain.com Hellma Cells 11831 Queens Boulevard Forest Hills, NY 11375 (718) 544-9166 FAX: (718) 263-6910 http://www.helmaUSA.com Hellma Postfach 1163 D-79371 M¨ullheim/Baden, Germany (49) 7631-1820 FAX: (49) 7631-13546 http://www.hellma-worldwide.de Henry Schein 135 Duryea Road, Mail Room 150 Melville, NY 11747 (800) 472-4346 FAX: (516) 843-5652 http://www.henryschein.com
Heraeus Kulzer 4315 South Lafayette Boulevard South Bend, IN 46614 (800) 343-5336 (219) 291-0661 http://www.kulzer.com Heraeus Sepatech See Kendro Laboratory Products Hercules Aqualon Aqualon Division Hercules Research Center, Bldg. 8145 500 Hercules Road Wilmington, DE 19899 (800) 345-0447 FAX: (302) 995-4787 http://www.herc.com/aqualon/pharma Heto-Holten A/S Distributed by ATR Gydevang 17-19 DK-3450 Allerod, Denmark (45) 48-16-62-00 FAX: (45) 48-16-62-97 Hettich-Zentrifugen See Andreas Hettich Hewlett-Packard 3000 Hanover Street Mailstop 20B3 Palo Alto, CA 94304 (650) 857-1501 FAX: (650) 857-5518 http://www.hp.com HGS Hinimoto Plastics 1-10-24 Meguro-Honcho Megurouko Tokyo 152, Japan 3-3714-7226 FAX: 3-3714-4657 Hitachi Scientific Instruments Nissei Sangyo America 8100 N. First Street San Elsa, CA 95314 (800) 548-9001 FAX: (408) 432-0704 (408) 432-0520 http://www.hii.hitachi.com Hi-Tech Scientific Brunel Road Salisbury, Wiltshire, SP2 7PU UK (44) 1722-432320 (800) 344-0724 (US only) http://www.hi-techsci.co.uk Hoechst AG See Aventis Pharmaceutical Hoefer Scientific Instruments Division of Amersham-Pharmacia Biotech 800 Centennial Avenue Piscataway, NJ 08855 (800) 227-4750 FAX: (877) 295-8102 http://www.apbiotech.com
Hoffman-LaRoche 340 Kingsland Street Nutley, NJ 07110 (800) 526-0189 FAX: (973) 235-9605 (973) 235-5000 http://www.rocheUSA.com Holborn Surgical and Medical Instruments Westwood Industrial Estate Ramsgate Road Margate, Kent CT9 4JZ UK (44) 1843 296666 FAX: (44) 1843 295446 Honeywell 101 Columbia Road Morristown, NJ 07962 (973) 455-2000 FAX: (973) 455-4807 http://www.honeywell.com Hood Thermo-Pad Canada Comp. 20, Site 61A, RR2 Summerland, British Columbia V0H 1Z0 Canada (800) 665-9555 FAX: (250) 494-5003 (250) 494-5002 http://www.thermopad.com Horiba Instruments 17671 Armstrong Avenue Irvine, CA 92714 (949) 250-4811 FAX: (949) 250-0924 http://www.horiba.com Hoskins Manufacturing 10776 Hall Road P.O. Box 218 Hamburg, MI 48139 (810) 231-1900 FAX: (810) 231-4311 http://www.hoskinsmfgco.com Hosokawa Micron Powder Systems 10 Chatham Road Summit, NJ 07901 (800) 526-4491 FAX: (908) 273-7432 (908) 273-6360 http://www.hosokawamicron.com HT Biotechnology Unit 4 61 Ditton Walk Cambridge CB5 8QD, UK (44) 1223-412583 Hugo Sachs Electronik Postfach 138 7806 March-Hugstetten, Germany D-79229(49) 7665-92000 FAX: (49) 7665-920090 Human Biologics International 7150 East Camelback Road, Suite 245 Scottsdale, AZ 85251 (480) 990-2005 FAX: (480)-990-2155 http://www.humanbiological.com Human Genetic Mutant Cell Repository See Coriell Institute for Medical Research
HVS Image P.O. Box 100 Hampton, Middlesex TW12 2YD, UK FAX: (44) 208 783 1223 In the US: (800) 225-9261 FAX: (888) 483-8033 http://www.hvsimage.com Hybaid 111-113 Waldegrave Road Teddington, Middlesex TW11 8LL, UK (44) 0 1784 42500 FAX: (44) 0 1784 248085 http://www.hybaid.co.uk Hybaid Instruments 8 East Forge Parkway Franklin, MA 02028 (888)4-HYBAID FAX: (508) 541-3041 (508) 541-6918 http://www.hybaid.com Hybridon 155 Fortune Boulevard Milford, MA 01757 (508) 482-7500 FAX: (508) 482-7510 http://www.hybridon.com HyClone Laboratories 1725 South HyClone Road Logan, UT 84321 (800) HYCLONE FAX: (800) 533-9450 (801) 753-4584 FAX: (801) 750-0809 http://www.hyclone.com Hyseq 670 Almanor Avenue Sunnyvale, CA 94086 (408) 524-8100 FAX: (408) 524-8141 http://www.hyseq.com IBF Biotechnics See Sepracor IBI (International Biotechnologies) See Eastman Kodak For technical service (800) 243-2555 (203) 786-5600 ICN Biochemicals See ICN Biomedicals ICN Biomedicals 3300 Hyland Avenue Costa Mesa, CA 92626 (800) 854-0530 FAX: (800) 334-6999 (714) 545-0100 FAX: (714) 641-7275 http://www.icnbiomed.com ICN Flow and Pharmaceuticals See ICN Biomedicals ICN Immunobiochemicals See ICN Biomedicals ICN Radiochemicals See ICN Biomedicals ICONIX 100 King Street West, Suite 3825 Toronto, Ontario M5X 1E3 Canada (416) 410-2411 FAX: (416) 368-3089 http://www.iconix.com
Suppliers
15 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
ICRT (Imperial Cancer Research Technology) Sardinia House Sardinia Street London WC2A 3NL, UK (44) 1712-421136 FAX: (44) 1718-314991 Idea Scientific Company P.O. Box 13210 Minneapolis, MN 55414 (800) 433-2535 FAX: (612) 331-4217 http://www.ideascientific.com IEC See International Equipment Co.
Immunotech 130, av. Delattre de Tassigny B.P. 177 13276 Marseilles Cedex 9 France (33) 491-17-27-00 FAX: (33) 491-41-43-58 http://www.immunotech.fr Imperial Chemical Industries Imperial Chemical House Millbank, London SW1P 3JF, UK (44) 171-834-4444 FAX: (44)171-834-2042 http://www.ici.com
IKA Works 2635 N. Chase Parkway, SE Wilmington, NC 28405 (910) 452-7059 FAX: (910) 452-7693 http://www.ika.net
Inceltech See New Brunswick Scientific
Ikegami Electronics 37 Brook Avenue Maywood, NJ 07607 (201) 368-9171 FAX: (201) 569-1626
Incyte 6519 Dumbarton Circle Fremont, CA 94555 (510) 739-2100 FAX: (510) 739-2200 http://www.incyte.com
Ikemoto Scientific Technology 25-11 Hongo 3-chome, Bunkyo-ku Tokyo 101-0025, Japan (81) 3-3811-4181 FAX: (81) 3-3811-1960 Imagenetics See ATC Diagnostics Imaging Research c/o Brock University 500 Glenridge Avenue St. Catharines, Ontario L2S 3A1 Canada (905) 688-2040 FAX: (905) 685-5861 http://www.imaging.brocku.ca Imclone Systems 180 Varick Street New York, NY 10014 (212) 645-1405 FAX: (212) 645-2054 http://www.imclone.com IMCO Corporation LTD., AB P.O. Box 21195 SE-100 31 Stockholm, Sweden 46-8-33-53-09 FAX: 46-8-728-47-76 http://www.imcocorp.se IMICO Calle Vivero, No. 5-4a Planta E-28040, Madrid, Spain (34) 1-535-3960 FAX: (34) 1-535-2780 Immunex 51 University Street Seattle, WA 98101 (206) 587-0430 FAX: (206) 587-0606 http://www.immunex.com
Incstar See DiaSorin
Instrutech 20 Vanderventer Avenue, Suite 101E Port Washington, NY 11050 (516) 883-1300 FAX: (516) 883-1558 http://www.instrutech.com
Innogenetics N.V. Technologie Park 6 B-9052 Zwijnaarde Belgium (32) 9-329-1329 FAX: (32) 9-245-7623 http://www.innogenetics.com Innovative Medical Services 1725 Gillespie Way El Cajon, CA 92020 (619) 596-8600 FAX: (619) 596-8700 http://www.imspure.com Innovative Research 3025 Harbor Lane N, Suite 300 Plymouth, MN 55447 (612) 519-0105 FAX: (612) 519-0239 http://www.inres.com Innovative Research of America 2 N. Tamiami Trail, Suite 404 Sarasota, FL 34236 (800) 421-8171 FAX: (800) 643-4345 (941) 365-1406 FAX: (941) 365-1703 http://www.innovrsrch.com
Integrated DNA Technologies 1710 Commercial Park Coralville, Iowa 52241 (800) 328-2661 FAX: (319) 626-8444 http://www.idtdna.com Integrated Genetics See Genzyme Genetics Integrated Scientific Imaging Systems 3463 State Street, Suite 431 Santa Barbara, CA 93105 (805) 692-2390 FAX: (805) 692-2391 http://www.imagingsystems.com Integrated Separation Systems (ISS) See OWL Separation Systems IntelliGenetics See Oxford Molecular Group
Inotech Biosystems 15713 Crabbs Branch Way, #110 Rockville, MD 20855 (800) 635-4070 FAX: (301) 670-2859 (301) 670-2850 http://www.inotechintl.com
Interactiva BioTechnologie Sedanstrasse 10 D-89077 Ulm, Germany (49) 731-93579-290 FAX: (49) 731-93579-291 http://www.interactiva.de
Individual Monitoring Systems 6310 Harford Road Baltimore, MD 21214
INOVISION 22699 Old Canal Road Yorba Linda, CA 92887 (714) 998-9600 FAX: (714) 998-9666 http://www.inovision.com
Indo Fine Chemical P.O. Box 473 Somerville, NJ 08876 (888) 463-6346 FAX: (908) 359-1179 (908) 359-6778 http://www.indofinechemical.com
Instech Laboratories 5209 Militia Hill Road Plymouth Meeting, PA 19462 (800) 443-4227 FAX: (610) 941-0134 (610) 941-0132 http://www.instechlabs.com
Interchim 213 J.F. Kennedy Avenue B.P. 1140 Montlucon 03103 France (33) 04-70-03-83-55 FAX: (33) 04-70-03-93-60
Industrial Acoustics 1160 Commerce Avenue Bronx, NY 10462 (718) 931-8000 FAX: (718) 863-1138 http://www.industrialacoustics.com
Instron 100 Royall Street Canton, MA 02021 (800) 564-8378 FAX: (781) 575-5725 (781) 575-5000 http://www.instron.com
Inex Pharmaceuticals 100-8900 Glenlyon Parkway Glenlyon Business Park Burnaby, British Columbia V5J 5J8 Canada (604) 419-3200 FAX: (604) 419-3201 http://www.inexpharm.com
Instrumentarium P.O. Box 300 00031 Instrumentarium Helsinki, Finland (10) 394-5566 http://www.instrumentarium.fi
Incyte Pharmaceuticals 3160 Porter Drive Palo Alto, CA 94304 (877) 746-2983 FAX: (650) 855-0572 (650) 855-0555 http://www.incyte.com
Ingold, Mettler, Toledo 261 Ballardvale Street Wilmington, MA 01887 (800) 352-8763 FAX: (978) 658-0020 (978) 658-7615 http://www.mt.com
Instruments SA Division Jobin Yvon 16-18 Rue du Canal 91165 Longjumeau, Cedex, France (33)1 6454-1300 FAX: (33)1 6909-9319 http://www.isainc.com
Interfocus 14/15 Spring Rise Falcover Road Haverhill, Suffolk CB9 7XU, UK (44) 1440 703460 FAX: (44) 1440 704397 http://www.interfocus.ltd.uk Intergen 2 Manhattanville Road Purchase, NY 10577 (800) 431-4505 FAX: (800) 468-7436 (914) 694-1700 FAX: (914) 694-1429 http://www.intergenco.com Intermountain Scientific 420 N. Keys Drive Kaysville, UT 84037 (800) 999-2901 FAX: (800) 574-7892 (801) 547-5047 FAX: (801) 547-5051 http://www.bioexpress.com International Biotechnologies (IBI) See Eastman Kodak International Equipment Co. (IEC) See Thermoquest
Suppliers
16 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
International Institute for the Advancement of Medicine 1232 Mid-Valley Drive Jessup, PA 18434 (800) 486-IIAM FAX: (570) 343-6993 (570) 496-3400 http://www.iiam.org International Light 17 Graf Road Newburyport, MA 01950 (978) 465-5923 FAX: (978) 462-0759 International Market Supply (I.M.S.) Dane Mill Broadhurst Lane Congleton, Cheshire CW12 1LA, UK (44) 1260 275469 FAX: (44) 1260 276007 International Marketing Services See International Marketing Ventures International Marketing Ventures 6301 Ivy Lane, Suite 408 Greenbelt, MD 20770 (800) 373-0096 FAX: (301) 345-0631 (301) 345-2866 http://www.imvlimited.com International Products 201 Connecticut Drive Burlington, NJ 08016 (609) 386-8770 FAX: (609) 386-8438 http://www.mkt.ipcol.com Intracel Corporation Bartels Division 2005 Sammamish Road, Suite 107 Issaquah, WA 98027 (800) 542-2281 FAX: (425) 557-1894 (425) 392-2992 http://www.intracel.com Invitrogen 1600 Faraday Avenue Carlsbad, CA 92008 (800) 955-6288 FAX: (760) 603-7201 (760) 603-7200 http://www.invitrogen.com
ISC BioExpress 420 North Kays Drive Kaysville, UT 84037 (800) 999-2901 FAX: (800) 574-7892 (801) 547-5047 http://www.bioexpress.com ISCO P.O. Box 5347 4700 Superior Lincoln, NE 68505 (800) 228-4373 FAX: (402) 464-0318 (402) 464-0231 http://www.isco.com Isis Pharmaceuticals Carlsbad Research Center 2292 Faraday Avenue Carlsbad, CA 92008 (760) 931-9200 http://www.isip.com Isolabs See Wallac ISS See Integrated Separation Systems J & W Scientific See Agilent Technologies J.A. Webster 86 Leominster Road Sterling, MA 01564 (800) 225-7911 FAX: (978) 422-8959 http://www.jawebster.com J.T. Baker See Mallinckrodt Baker 222 Red School Lane Phillipsburg, NJ 08865 (800) JTBAKER FAX: (908) 859-6974 http://www.jtbaker.com Jackson ImmunoResearch Laboratories P.O. Box 9 872 W. Baltimore Pike West Grove, PA 19390 (800) 367-5296 FAX: (610) 869-0171 (610) 869-4024 http://www.jacksonimmuno.com
In Vivo Metric P.O. Box 249 Healdsburg, CA 95448 (707) 433-4819 FAX: (707) 433-2407
The Jackson Laboratory 600 Maine Street Bar Harbor, ME 04059 (800) 422-6423 FAX: (207) 288-5079 (207) 288-6000 http://www.jax.org
IRORI 9640 Towne Center Drive San Diego, CA 92121 (858) 546-1300 FAX: (858) 546-3083 http://www.irori.com
Jaece Industries 908 Niagara Falls Boulevard North Tonawanda, NY 14120 (716) 694-2811 FAX: (716) 694-2811 http://www.jaece.com
Irvine Scientific 2511 Daimler Street Santa Ana, CA 92705 (800) 577-6097 FAX: (949) 261-6522 (949) 261-7800 http://www.irvinesci.com
Janssen Pharmaceutica 1125 Trenton-Harbourton Road Titusville, NJ 09560 (609) 730-2577 FAX: (609) 730-2116 http://us.janssen.com Jasco 8649 Commerce Drive Easton, MD 21601 (800) 333-5272 FAX: (410) 822-7526 (410) 822-1220 http://www.jascoinc.com Jena Bioscience Loebstedter Str. 78 07749 Jena, Germany (49) 3641-464920 FAX: (49) 3641-464991 http://www.jenabioscience.com Jencons Scientific 800 Bursca Drive, Suite 801 Bridgeville, PA 15017 (800) 846-9959 FAX: (412) 257-8809 (412) 257-8861 http://www.jencons.co.uk JEOL Instruments 11 Dearborn Road Peabody, MA 01960 (978) 535-5900 FAX: (978) 536-2205 http://www.jeol.com/index.html Jewett 750 Grant Street Buffalo, NY 14213 (800) 879-7767 FAX: (716) 881-6092 (716) 881-0030 http://www.JewettInc.com John’s Scientific See VWR Scientific John Weiss and Sons 95 Alston Drive Bradwell Abbey Milton Keynes, Buckinghamshire MK1 4HF UK (44) 1908-318017 FAX: (44) 1908-318708 Johnson & Johnson Medical 2500 Arbrook Boulevard East Arlington, TX 76004 (800) 423-4018 http://www.jnjmedical.com Johnston Matthey Chemicals Orchard Road Royston, Hertfordshire SG8 5HE, UK (44) 1763-253000 FAX: (44) 1763-253466 http://www.chemicals.matthey.com
Janke & Kunkel See Ika Works
Jolley Consulting and Research 683 E. Center Street, Unit H Grayslake, IL 60030 (847) 548-2330 FAX: (847) 548-2984 http://www.jolley.com
Janssen Life Sciences Products See Amersham
Jordan Scientific See Shelton Scientific
Jandel Scientific See SPSS
Jorgensen Laboratories 1450 N. Van Buren Avenue Loveland, CO 80538 (800) 525-5614 FAX: (970) 663-5042 (970) 669-2500 http://www.jorvet.com JRH Biosciences and JR Scientific 13804 W. 107th Street Lenexa, KS 66215 (800) 231-3735 FAX: (913) 469-5584 (913) 469-5580 Jule Bio Technologies 25 Science Park, #14, Suite 695 New Haven, CT 06511 (800) 648-1772 FAX: (203) 786-5489 (203) 786-5490 http://hometown.aol.com/precastgel/ index.htm K.R. Anderson 2800 Bowers Avenue Santa Clara, CA 95051 (800) 538-8712 FAX: (408) 727-2959 (408) 727-2800 http://www.kranderson.com Kabi Pharmacia Diagnostics See Pharmacia Diagnostics Kapak 5305 Parkdale Drive St. Louis Park, MN 55416 (800) KAPAK-57 FAX: (612) 541-0735 (612) 541-0730 http://www.kapak.com Karl Hecht Stettener Str. 22-24 D-97647 Sondheim Rh¨on, Germany (49) 9779-8080 FAX: (49) 9779-80888 Karl Storz K¨oningin-Elisabeth Str. 60 D-14059 Berlin, Germany (49) 30-30 69 09-0 FAX: (49) 30-30 19 452 http://www.karlstorz.de KaVo EWL P.O. Box 1320 D-88293 Leutkirch im Allg¨au, Germany (49) 7561-86-0 FAX: (49) 7561-86-371 http://www.kavo.com/english/ startseite.htm Keithley Instruments 28775 Aurora Road Cleveland, OH 44139 (800) 552-1115 FAX: (440) 248-6168 (440) 248-0400 http://www.keithley.com Kemin 2100 Maury Street, Box 70 Des Moines, IA 50301 (515) 266-2111 FAX: (515) 266-8354 http://www.kemin.com
Suppliers
17 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Kemo 3 Brook Court, Blakeney Road Beckenham, Kent BR3 1HG, UK (44) 0181 658 3838 FAX: (44) 0181 658 4084 http://www.kemo.com Kendall 15 Hampshire Street Mansfield, MA 02048 (800) 962-9888 FAX: (800) 724-1324 http://www.kendallhq.com Kendro Laboratory Products 31 Pecks Lane Newtown, CT 06470 (800) 522-SPIN FAX: (203) 270-2166 (203) 270-2080 http://www.kendro.com Kendro Laboratory Products P.O. Box 1220 Am Kalkberg D-3360 Osterod, Germany (55) 22-316-213 FAX: (55) 22-316-202 http://www.heraeus-instruments.de Kent Laboratories 23404 NE 8th Street Redmond, WA 98053 (425) 868-6200 FAX: (425) 868-6335 http://www.kentlabs.com Kent Scientific 457 Bantam Road, #16 Litchfield, CT 06759 (888) 572-8887 FAX: (860) 567-4201 (860) 567-5496 http://www.kentscientific.com Keuffel & Esser See Azon Keystone Scientific Penn Eagle Industrial Park 320 Rolling Ridge Drive Bellefonte, PA 16823 (800) 437-2999 FAX: (814) 353-2305 (814) 353-2300 Ext 1 http://www.keystonescientific.com Kimble/Kontes Biotechnology 1022 Spruce Street P.O. Box 729 Vineland, NJ 08360 (888) 546-2531 FAX: (856) 794-9762 (856) 692-3600 http://www.kimble-kontes.com Kinematica AG Luzernerstrasse 147a CH-6014 Littau-Luzern, Switzerland (41) 41 2501257 FAX: (41) 41 2501460 http://www.kinematica.ch
Kin-Tek 504 Laurel Street LaMarque, TX 77568 (800) 326-3627 FAX: (409) 938-3710 http://www.kin-tek.com
Lab Safety Supply P.O. Box 1368 Janesville, WI 53547 (800) 356-0783 FAX: (800) 543-9910 (608) 754-7160 FAX: (608) 754-1806 http://www.labsafety.com
Kipp & Zonen 125 Wilbur Place Bohemia, NY 11716 (800) 645-2065 FAX: (516) 589-2068 (516) 589-2885 http://www.kippzonen.thomasregister. com/olc/kippzonen
Lab-Tek Products See Nalge Nunc International
Kirkegaard & Perry Laboratories 2 Cessna Court Gaithersburg, MD 20879 (800) 638-3167 FAX: (301) 948-0169 (301) 948-7755 http://www.kpl.com Kodak See Eastman Kodak Kontes Glass See Kimble/Kontes Biotechnology Kontron Instruments AG Postfach CH-8010 Zurich, Switzerland 41-1-733-5733 FAX: 41-1-733-5734 David Kopf Instruments P.O. Box 636 Tujunga, CA 91043 (818) 352-3274 FAX: (818) 352-3139 Kraft Apparatus See Glas-Col Apparatus Kramer Scientific Corporation 711 Executive Boulevard Valley Cottage, NY 10989 (845) 267-5050 FAX: (845) 267-5550 Kulite Semiconductor Products 1 Willow Tree Road Leonia, NJ 07605 (201) 461-0900 FAX: (201) 461-0990 http://www.kulite.com Lab-Line Instruments 15th & Bloomingdale Avenues Melrose Park, IL 60160 (800) LAB-LINE FAX: (800) 450-4LAB FAX: (708) 450-5830 http://www.labline.com Lab Products 742 Sussex Avenue P.O. Box 639 Seaford, DE 19973 (800) 526-0469 FAX: (302) 628-4309 (302) 628-4300 http://www.labproductsinc.com LabRepco 101 Witmer Road, Suite 700 Horsham, PA 19044 (800) 521-0754 FAX: (215) 442-9202 http://www.labrepco.com
Labconco 8811 Prospect Avenue Kansas City, MO 64132 (800) 821-5525 FAX: (816) 363-0130 (816) 333-8811 http://www.labconco.com Labindustries See Barnstead/Thermolyne Labnet International P.O. Box 841 Woodbridge, NJ 07095 (888) LAB-NET1 FAX: (732) 417-1750 (732) 417-0700 http://www.nationallabnet.com LABO-MODERNE 37 rue Dombasle Paris 75015 France (33) 01-45-32-62-54 FAX: (33) 01-45-32-01-09 http://www.labomoderne.com/fr Laboratory of Immunoregulation National Institute of Allergy and Infectious Diseases/NIH 9000 Rockville Pike Building 10, Room 11B13 Bethesda, MD 20892 (301) 496-1124 Laboratory Supplies 29 Jefry Lane Hicksville, NY 11801 (516) 681-7711
Labtronix Manufacturing 3200 Investment Boulevard Hayward, CA 94545 (510) 786-3200 FAX: (510) 786-3268 http://www.labtronix.com Lafayette Instrument 3700 Sagamore Parkway North P.O. Box 5729 Lafayette, IN 47903 (800) 428-7545 FAX: (765) 423-4111 (765) 423-1505 http://www.lafayetteinstrument.com Lambert Instruments Turfweg 4 9313 TH Leutingewolde The Netherlands (31) 50-5018461 FAX: (31) 50-5010034 http://www.lambert-instruments.com Lancaster Synthesis P.O. Box 1000 Windham, NH 03087 (800) 238-2324 FAX: (603) 889-3326 (603) 889-3306 http://www.lancastersynthesis-us.com Lancer 140 State Road 419 Winter Springs, FL 32708 (800) 332-1855 FAX: (407) 327-1229 (407) 327-8488 http://www.lancer.com LaVision GmbH Gerhard-Gerdes-Str. 3 D-37079 Goettingen, Germany (49) 551-50549-0 FAX: (49) 551-50549-11 http://www.lavision.de Lawshe See Advanced Process Supply LC Laboratories 165 New Boston Street Woburn, MA 01801 (781) 937-0777 FAX: (781) 938-5420 http://www.lclaboratories.com
Labscan Limited Stillorgan Industrial Park Stillorgan Dublin, Ireland (353) 1-295-2684 FAX: (353) 1-295-2685 http://www.labscan.ie
LC Packings 80 Carolina Street San Francisco, CA 94103 (415) 552-1855 FAX: (415) 552-1859 http://www.lcpackings.com
Labsystems See Thermo Labsystems Labsystems Affinity Sensors Saxon Way, Bar Hill Cambridge CB3 8SL, UK 44 (0) 1954 789976 FAX: 44 (0) 1954 789417 http://www.affinity-sensors.com
LC Services See LC Laboratories
Labtronics 546 Governors Road Guelph, Ontario N1K 1E3, Canada (519) 763-4930 FAX: (519) 836-4431 http://www.labtronics.com
LECO 3000 Lakeview Avenue St. Joseph, MI 49085 (800) 292-6141 FAX: (616) 982-8977 (616) 985-5496 http://www.leco.com Lederle Laboratories See Wyeth-Ayerst
Suppliers
18 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
Lee Biomolecular Research Laboratories 11211 Sorrento Valley Road, Suite M San Diego, CA 92121 (858) 452-7700 The Lee Company 2 Pettipaug Road P.O. Box 424 Westbrook, CT 06498 (800) LEE-PLUG FAX: (860) 399-7058 (860) 399-6281 http://www.theleeco.com Lee Laboratories 1475 Athens Highway Grayson, GA 30017 (800) 732-9150 FAX: (770) 979-9570 (770) 972-4450 http://www.leelabs.com Leica 111 Deer Lake Road Deerfield, IL 60015 (800) 248-0123 FAX: (847) 405-0147 (847) 405-0123 http://www.leica.com Leica Microsystems Imneuenheimer Feld 518 D-69120 Heidelberg, Germany (49) 6221-41480 FAX: (49) 6221-414833 http://www.leica-microsystems.com Leinco Technologies 359 Consort Drive St. Louis, MO 63011 (314) 230-9477 FAX: (314) 527-5545 http://www.leinco.com
Life Science Resources Two Corporate Center Drive Melville, NY 11747 (800) 747-9530 FAX: (516) 844-5114 (516) 844-5085 http://www.astrocam.com Life Sciences 2900 72nd Street North St. Petersburg, FL 33710 (800) 237-4323 FAX: (727) 347-2957 (727) 345-9371 http://www.lifesci.com Life Technologies 9800 Medical Center Drive P.O. Box 6482 Rockville, MD 20849 (800) 828-6686 FAX: (800) 331-2286 http://www.lifetech.com Lifecodes 550 West Avenue Stamford, CT 06902 (800) 543-3263 FAX: (203) 328-9599 (203) 328-9500 http://www.lifecodes.com Lightnin 135 Mt. Read Boulevard Rochester, NY 14611 (888) MIX-BEST FAX: (716) 527-1742 (716) 436-5550 http://www.lightnin-mixers.com Linear Drives Luckyn Lane, Pipps Hill Basildon, Essex SS14 3BW, UK (44) 1268-287070 FAX: (44) 1268-293344 http://www.lineardrives.com
Leitz U.S.A. See Leica
Linscott’s Directory 4877 Grange Road Santa Rosa, CA 95404 (707) 544-9555 FAX: (415) 389-6025 http://www.linscottsdirectory.co.uk
Letica Scientific Instruments Panlab s.i., c/Loreto 50 08029 Barcelona, Spain (34) 93-419-0709 FAX: (34) 93-419-7145 http://www.panlab-sl.com
Linton Instrumentation Unit 11, Forge Business Center Upper Rose Lane Palgrave, Diss, Norfolk IP22 1AP, UK (44) 1-379-651-344 FAX: (44) 1-379-650-970 http://www.lintoninst.co.uk
Leybold-Heraeus Trivac DZA 5700 Mellon Road Export, PA 15632 (412) 327-5700 LI-COR Biotechnology Division 4308 Progressive Avenue Lincoln, NE 68504 (800) 645-4267 FAX: (402) 467-0819 (402) 467-0700 http://www.licor.com Life Science Laboratories See Adaptive Biosystems
List Biological Laboratories 501-B Vandell Way Campbell, CA 95008 (800) 726-3213 FAX: (408) 866-6364 (408) 866-6363 http://www.listlabs.com LKB Instruments See Amersham Pharmacia Biotech Lloyd Laboratories 604 West Thomas Avenue Shenandoah, IA 51601 (800) 831-0004 FAX: (712) 246-5245 (712) 246-4000 http://www.lloydinc.com
Loctite 1001 Trout Brook Crossing Rocky Hill, CT 06067 (860) 571-5100 FAX: (860) 571-5465 http://www.loctite.com Lofstrand Labs 7961 Cessna Avenue Gaithersburg, MD 20879 (800) 541-0362 FAX: (301) 948-9214 (301) 330-0111 http://www.lofstrand.com Lomir Biochemical 99 East Main Street Malone, NY 12953 (877) 425-3604 FAX: (518) 483-8195 (518) 483-7697 http://www.lomir.com LSL Biolafitte 10 rue de Temara 7810C St.-Germain-en-Laye, France (33) 1-3061-5260 FAX: (33) 1-3061-5234 Ludl Electronic Products 171 Brady Avenue Hawthorne, NY 10532 (888) 769-6111 FAX: (914) 769-4759 (914) 769-6111 http://www.ludl.com Lumigen 24485 W. Ten Mile Road Southfield, MI 48034 (248) 351-5600 FAX: (248) 351-0518 http://www.lumigen.com Luminex 12212 Technology Boulevard Austin, TX 78727 (888) 219-8020 FAX: (512) 258-4173 (512) 219-8020 http://www.luminexcorp.com LYNX Therapeutics 25861 Industrial Boulevard Hayward, CA 94545 (510) 670-9300 FAX: (510) 670-9302 http://www.lynxgen.com Lyphomed 3 Parkway North Deerfield, IL 60015 (847) 317-8100 FAX: (847) 317-8600 M.E.D. Associates See Med Associates Macherey-Nagel 6 South Third Street, #402 Easton, PA 18042 (610) 559-9848 FAX: (610) 559-9878 http://www.macherey-nagel.com Macherey-Nagel Valencienner Strasse 11 P.O. Box 101352 D-52313 Dueren, Germany (49) 2421-969141 FAX: (49) 2421-969199 http://www.macherey-nagel.ch
Mac-Mod Analytical 127 Commons Court Chadds Ford, PA 19317 800-441-7508 FAX: (610) 358-5993 (610) 358-9696 http://www.mac-mod.com Mallinckrodt Baker 222 Red School Lane Phillipsburg, NJ 08865 (800) 582-2537 FAX: (908) 859-6974 (908) 859-2151 http://www.mallbaker.com Mallinckrodt Chemicals 16305 Swingley Ridge Drive Chesterfield, MD 63017 (314) 530-2172 FAX: (314) 530-2563 http://www.mallchem.com Malven Instruments Enigma Business Park Grovewood Road Malven, Worchestershire WR 141 XZ, United Kingdom Marinus 1500 Pier C Street Long Beach, CA 90813 (562) 435-6522 FAX: (562) 495-3120 Markson Science c/o Whatman Labs Sales P.O. Box 1359 Hillsboro, OR 97123 (800) 942-8626 FAX: (503) 640-9716 (503) 648-0762 Marsh Biomedical Products 565 Blossom Road Rochester, NY 14610 (800) 445-2812 FAX: (716) 654-4810 (716) 654-4800 http://www.biomar.com Marshall Farms USA 5800 Lake Bluff Road North Rose, NY 14516 (315) 587-2295 Martek 6480 Dobbin Road Columbia, MD 21045 (410) 740-0081 FAX: (410) 740-2985 http://www.martekbio.com Martin Supply Distributor of Gerber Scientific 2740 Loch Raven Road Baltimore, MD 21218 (800) 282-5440 FAX: (410) 366-0134 (410) 366-1696 Mast Immunosystems 630 Clyde Court Mountain View, CA 94043 (800) 233-MAST FAX: (650) 969-2745 (650) 961-5501 http://www.mastallergy.com
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19 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Matheson Gas Products P.O. Box 624 959 Route 46 East Parsippany, NJ 07054 (800) 416-2505 FAX: (973) 257-9393 (973) 257-1100 http://www.mathesongas.com Mathsoft 1700 Westlake Avenue N., Suite 500 Seattle, WA 98109 (800) 569-0123 FAX: (206) 283-8691 (206) 283-8802 http://www.mathsoft.com Matreya 500 Tressler Street Pleasant Gap, PA 16823 (814) 359-5060 FAX: (814) 359-5062 http://www.matreya.com Matrigel See Becton Dickinson Labware Matrix Technologies 22 Friars Drive Hudson, NH 03051 (800) 345-0206 FAX: (603) 595-0106 (603) 595-0505 http://www.matrixtechcorp.com MatTek Corp. 200 Homer Ave. Ashland, Massachusetts 01721 (508) 881-6771 FAX: (508) 879-1532 http://www.mattek.com Maxim Medical 89 Oxford Road Oxford OX2 9PD, UK 44 (0)1865-865943 FAX: 44 (0)1865-865291 http://www.maximmed.com Mayo Clinic Section on Engineering Project #ALA-1, 1982 200 1st Street SW Rochester, MN 55905 (507) 284-2511 FAX: (507) 284-5988 McGaw See B. Braun-McGaw McMaster-Carr 600 County Line Road Elmhurst, IL 60126 (630) 833-0300 FAX: (630) 834-9427 http://www.mcmaster.com McNeil Pharmaceutical See Ortho McNeil Pharmaceutical MCNC 3021 Cornwallis Road P.O. Box 12889 Research Triangle Park, NC 27709 (919) 248-1800 FAX: (919) 248-1455 http://www.mcnc.org
MD Industries 5 Revere Drive, Suite 415 Northbrook, IL 60062 (800) 421-8370 FAX: (847) 498-2627 (708) 339-6000 http://www.mdindustries.com MDS Nordion 447 March Road P.O. Box 13500 Kanata, Ontario K2K 1X8, Canada (800) 465-3666 FAX: (613) 592-6937 (613) 592-2790 http://www.mds.nordion.com MDS Sciex 71 Four Valley Drive Concord, Ontario Canada L4K 4V8 (905) 660-9005 FAX: (905) 660-2600 http://www.sciex.com Mead Johnson See Bristol-Meyers Squibb Med Associates P.O. Box 319 St. Albans, VT 05478 (802) 527-2343 FAX: (802) 527-5095 http://www.med-associates.com Medecell 239 Liverpool Road London N1 1LX, UK (44) 20-7607-2295 FAX: (44) 20-7700-4156 http://www.medicell.co.uk Media Cybernetics 8484 Georgia Avenue, Suite 200 Silver Spring, MD 20910 (301) 495-3305 FAX: (301) 495-5964 http://www.mediacy.com Mediatech 13884 Park Center Road Herndon, VA 20171 (800) cellgro (703) 471-5955 http://www.cellgro.com Medical Systems See Harvard Apparatus Medifor 647 Washington Street Port Townsend, WA 98368 (800) 366-3710 FAX: (360) 385-4402 (360) 385-0722 http://www.medifor.com MedImmune 35 W. Watkins Mill Road Gaithersburg, MD 20878 (301) 417-0770 FAX: (301) 527-4207 http://www.medimmune.com
MedProbe AS P.O. Box 2640 St. Hanshaugen N-0131 Oslo, Norway (47) 222 00137 FAX: (47) 222 00189 http://www.medprobe.com Megazyme Bray Business Park Bray, County Wicklow Ireland (353) 1-286-1220 FAX: (353) 1-286-1264 http://www.megazyme.com
Metachem Technologies 3547 Voyager Street, Bldg. 102 Torrance, CA 90503 (310) 793-2300 FAX: (310) 793-2304 http://www.metachem.com Metallhantering Box 47172 100-74 Stockholm, Sweden (46) 8-726-9696 MethylGene 7220 Frederick-Banting, Suite 200 Montreal, Quebec H4S 2A1, Canada http://www.methylgene.com
Melles Griot 4601 Nautilus Court South Boulder, CO 80301 (800) 326-4363 FAX: (303) 581-0960 (303) 581-0337 http://www.mellesgriot.com
Metro Scientific 475 Main Street, Suite 2A Farmingdale, NY 11735 (800) 788-6247 FAX: (516) 293-8549 (516) 293-9656
Menzel-Glaser Postfach 3157 D-38021 Braunschweig, Germany (49) 531 590080 FAX: (49) 531 509799
Metrowerks 980 Metric Boulevard Austin, TX 78758 (800) 377-5416 (512) 997-4700 http://www.metrowerks.com
E. Merck Frankfurterstrasse 250 D-64293 Darmstadt 1, Germany (49) 6151-720 Merck See EM Science Merck & Company Merck National Service Center P.O. Box 4 West Point, PA 19486 (800) NSC-MERCK (215) 652-5000 http://www.merck.com
Mettler Instruments Mettler-Toledo 1900 Polaris Parkway Columbus, OH 43240 (800) METTLER FAX: (614) 438-4900 http://www.mt.com Miami Serpentarium Labs 34879 Washington Loop Road Punta Gorda, FL 33982 (800) 248-5050 FAX: (813) 639-1811 (813) 639-8888 http://www.miamiserpentarium.com
Merck Research Laboratories See Merck & Company Merck Sharpe Human Health Division 300 Franklin Square Drive Somerset, NJ 08873 (800) 637-2579 FAX: (732) 805-3960 (732) 805-0300
Michrom BioResources 1945 Industrial Drive Auburn, CA 95603 (530) 888-6498 FAX: (530) 888-8295 http://www.michrom.com Mickle Laboratory Engineering Gomshall, Surrey, UK (44) 1483-202178
Merial Limited 115 Transtech Drive Athens, GA 30601 (800) MERIAL-1 FAX: (706) 548-0608 (706) 548-9292 http://www.merial.com
Micra Scientific A division of Eichrom Industries 8205 S. Cass Ave, Suite 111 Darien, IL 60561 (800) 283-4752 FAX: (630) 963-1928 (630) 963-0320 http://www.micrasci.com
Meridian Instruments P.O. Box 1204 Kent, WA 98035 (253) 854-9914 FAX: (253) 854-9902 http://www.minstrument.com
MicroBrightField 74 Hegman Avenue Colchester, VT 05446 (802) 655-9360 FAX: (802) 655-5245 http://www.microbrightfield.com
Meta Systems Group 32 Hammond Road Belmont, MA 02178 (617) 489-9950 FAX: (617) 489-9952
Micro Essential Laboratory 4224 Avenue H Brooklyn, NY 11210 (718) 338-3618 FAX: (718) 692-4491
Suppliers
20 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
Micro Filtration Systems 7-3-Chome, Honcho Nihonbashi, Tokyo, Japan (81) 3-270-3141 Micro-Metrics P.O. Box 13804 Atlanta, GA 30324 (770) 986-6015 FAX: (770) 986-9510 http://www.micro-metrics.com Micro-Tech Scientific 140 South Wolfe Road Sunnyvale, CA 94086 (408) 730-8324 FAX: (408) 730-3566 http://www.microlc.com MicroCal 22 Industrial Drive East Northampton, MA 01060 (800) 633-3115 FAX: (413) 586-0149 (413) 586-7720 www.microcalorimetry.com
Midwest Scientific 280 Vance Road Valley Park, MO 63088 (800) 227-9997 FAX: (636) 225-9998 (636) 225-9997 http://www.midsci.com Miles See Bayer Miles Laboratories See Serological Miles Scientific See Nunc Millar Instruments P.O. Box 230227 6001-A Gulf Freeway Houston, TX 77023 (713) 923-9171 FAX: (713) 923-7757 http://www.millarinstruments.com MilliGen/Biosearch See Millipore
Microfluidics 30 Ossipee Road P.O. Box 9101 Newton, MA 02164 (800) 370-5452 FAX: (617) 965-1213 (617) 969-5452 http://www.microfluidicscorp.com
Millipore 80 Ashbury Road P.O. Box 9125 Bedford, MA 01730 (800) 645-5476 FAX: (781) 533-3110 (781) 533-6000 http://www.millipore.com
Microgon See Spectrum Laboratories
Miltenyi Biotec 251 Auburn Ravine Road, Suite 208 Auburn, CA 95603 (800) 367-6227 FAX: (530) 888-8925 (530) 888-8871 http://www.miltenyibiotec.com
Microlase Optical Systems West of Scotland Science Park Kelvin Campus, Maryhill Road Glasgow G20 0SP, UK (44) 141-948-1000 FAX: (44) 141-946-6311 http://www.microlase.co.uk Micron Instruments 4509 Runway Street Simi Valley, CA 93063 (800) 638-3770 FAX: (805) 522-4982 (805) 552-4676 http://www.microninstruments.com Micron Separations See MSI Micro Photonics 4949 Liberty Lane, Suite 170 P.O. Box 3129 Allentown, PA 18106 (610) 366-7103 FAX: (610) 366-7105 http://www.microphotonics.com MicroTech 1420 Conchester Highway Boothwyn, PA 19061 (610) 459-3514 Midland Certified Reagent Company 3112-A West Cuthbert Avenue Midland, TX 79701 (800) 247-8766 FAX: (800) 359-5789 (915) 694-7950 FAX: (915) 694-2387 http://www.mcrc.com
Miltex 6 Ohio Drive Lake Success, NY 11042 (800) 645-8000 FAX: (516) 775-7185 (516) 349-0001 Milton Roy See Spectronic Instruments Mini-Instruments 15 Burnham Business Park Springfield Road Burnham-on-Crouch, Essex CM0 8TE, UK (44) 1621-783282 FAX: (44) 1621-783132 http://www.mini-instruments.co.uk Mini Mitter P.O. Box 3386 Sunriver, OR 97707 (800) 685-2999 FAX: (541) 593-5604 (541) 593-8639 http://www.minimitter.com Misonix 1938 New Highway Farmingdale, NY 11735 (800) 645-9846 FAX: (516) 694-9412 http://www.misonix.com Mitutoyo (MTI) See Dolla Eastern
MJ Research Waltham, MA 02451 (800) PELTIER FAX: (617) 923-8080 (617) 923-8000 http://www.mjr.com Modular Instruments 228 West Gay Street Westchester, PA 19380 (610) 738-1420 FAX: (610) 738-1421 http://www.mi2.com Molecular Biology Insights 8685 US Highway 24 Cascade, CO 80809-1333 (800) 747-4362 FAX: (719) 684-7989 (719) 684-7988 http://www.oligo.net Molecular Biosystems 10030 Barnes Canyon Road San Diego, CA 92121 (858) 452-0681 FAX: (858) 452-6187 http://www.mobi.com Molecular Devices 1312 Crossman Avenue Sunnyvale, CA 94089 (800) 635-5577 FAX: (408) 747-3602 (408) 747-1700 http://www.moldev.com Molecular Designs 1400 Catalina Street San Leandro, CA 94577 (510) 895-1313 FAX: (510) 614-3608 Molecular Dynamics 928 East Arques Avenue Sunnyvale, CA 94086 (800) 333-5703 FAX: (408) 773-1493 (408) 773-1222 http://www.apbiotech.com Molecular Probes 4849 Pitchford Avenue Eugene, OR 97402 (800) 438-2209 FAX: (800) 438-0228 (541) 465-8300 FAX: (541) 344-6504 http://www.probes.com Molecular Research Center 5645 Montgomery Road Cincinnati, OH 45212 (800) 462-9868 FAX: (513) 841-0080 (513) 841-0900 http://www.mrcgene.com Molecular Simulations 9685 Scranton Road San Diego, CA 92121 (800) 756-4674 FAX: (858) 458-0136 (858) 458-9990 http://www.msi.com Monoject Disposable Syringes & Needles/Syrvet 16200 Walnut Street Waukee, IA 50263 (800) 727-5203 FAX: (515) 987-5553 (515) 987-5554 http://www.syrvet.com
Monsanto Chemical 800 North Lindbergh Boulevard St. Louis, MO 63167 (314) 694-1000 FAX: (314) 694-7625 http://www.monsanto.com Moravek Biochemicals 577 Mercury Lane Brea, CA 92821 (800) 447-0100 FAX: (714) 990-1824 (714) 990-2018 http://www.moravek.com Moss P.O. Box 189 Pasadena, MD 21122 (800) 932-6677 FAX: (410) 768-3971 (410) 768-3442 http://www.mosssubstrates.com Motion Analysis 3617 Westwind Boulevard Santa Rosa, CA 95403 (707) 579-6500 FAX: (707) 526-0629 http://www.motionanalysis.com Mott Farmington Industrial Park 84 Spring Lane Farmington, CT 06032 (860) 747-6333 FAX: (860) 747-6739 http://www.mottcorp.com MSI (Micron Separations) See Osmonics Multiple Peptide Systems 3550 General Atomics Court San Diego, CA 92121 (800) 338-4965 FAX: (800) 654-5592 (858) 455-3710 FAX: (858) 455-3713 http://www.mps-sd.com Murex Diagnostics 3075 Northwoods Circle Norcross, GA 30071 (707) 662-0660 FAX: (770) 447-4989 MWG-Biotech Anzinger Str. 7 D-85560 Ebersberg, Germany (49) 8092-82890 FAX: (49) 8092-21084 http://www.mwg biotech.com Myriad Industries 3454 E Street San Diego, CA 92102 (800) 999-6777 FAX: (619) 232-4819 (619) 232-6700 http://www.myriadindustries.com Nacalai Tesque Nijo Karasuma, Nakagyo-ku Kyoto 604, Japan 81-75-251-1723 FAX: 81-75-251-1762 http://www.nacalai.co.jp
Suppliers
21 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Nalge Nunc International Subsidiary of Sybron International 75 Panorama Creek Drive P.O. Box 20365 Rochester, NY 14602 (800) 625-4327 FAX: (716) 586-8987 (716) 264-9346 http://www.nalgenunc.com Nanogen 10398 Pacific Center Court San Diego, CA 92121 (858) 410-4600 FAX: (858) 410-4848 http://www.nanogen.com Nanoprobes 95 Horse Block Road Yaphank, NY 11980 (877) 447-6266 FAX: (631) 205-9493 (631) 205-9490 http://www.nanoprobes.com Narishige USA 1710 Hempstead Turnpike East Meadow, NY 11554 (800) 445-7914 FAX: (516) 794-0066 (516) 794-8000 http://www.narishige.co.jp National Bag Company 2233 Old Mill Road Hudson, OH 44236 (800) 247-6000 FAX: (330) 425-9800 (330) 425-2600 http://www.nationalbag.com National Band and Tag Department X 35, Box 72430 Newport, KY 41032 (606) 261-2035 FAX: (800) 261-8247 https://www.nationalband.com National Biosciences See Molecular Biology Insights National Diagnostics 305 Patton Drive Atlanta, GA 30336 (800) 526-3867 FAX: (404) 699-2077 (404) 699-2121 http://www.nationaldiagnostics.com National Institute of Standards and Technology 100 Bureau Drive Gaithersburg, MD 20899 (301) 975-NIST FAX: (301) 926-1630 http://www.nist.gov National Instruments 11500 North Mopac Expressway Austin, TX 78759 (512) 794-0100 FAX: (512) 683-8411 http://www.ni.com National Labnet See Labnet International
National Scientific Instruments 975 Progress Circle Lawrenceville, GA 300243 (800) 332-3331 FAX: (404) 339-7173 http://www.nationalscientific.com National Scientific Supply 1111 Francisco Bouldvard East San Rafael, CA 94901 (800) 525-1779 FAX: (415) 459-2954 (415) 459-6070 http://www.nat-sci.com Naz-Dar-KC Chicago Nazdar 1087 N. North Branch Street Chicago, IL 60622 (800) 736-7636 FAX: (312) 943-8215 (312) 943-8338 http://www.nazdar.com NEB See New England Biolabs Nektar Therapeutics 150 Industrial Road San Carlos, CA 94070 (650) 631-3100 FAX: 650.631.3150 http://www.nektar.com NEN Life Science Products 549 Albany Street Boston, MA 02118 (800) 551-2121 FAX: (617) 451-8185 (617) 350-9075 http://www.nen.com NEN Research Products, Dupont (UK) Diagnostics and Biotechnology Systems Wedgewood Way Stevenage, Hertfordshire SG1 4QN, UK 44-1438-734831 44-1438-734000 FAX: 44-1438-734836 http://www.dupont.com Neogen 628 Winchester Road Lexington, KY 40505 (800) 477-8201 FAX: (606) 255-5532 (606) 254-1221 http://www.neogen.com Neosystems 380, 11012 Macleod Trail South Calgary, Alberta T2J 6A5 Canada (403) 225-9022 FAX: (403) 225-9025 http://www.neosystems.com The Nest Group 45 Valley Road Southborough, MA 01772 (800) 347-6378 FAX: (508) 485-5736 (508) 481-6223 http://world.std.com/∼nestgrp
Neuro Probe 16008 Industrial Drive Gaithersburg, MD 20877 (301) 417-0014 FAX: (301) 977-5711 http://www.neuroprobe.com Neurocrine Biosciences 10555 Science Center Drive San Diego, CA 92121 (619) 658-7600 FAX: (619) 658-7602 http://www.neurocrine.com Nevtek HCR03, Box 99 Burnsville, VA 24487 (540) 925-2322 FAX: (540) 925-2323 http://www.nevtek.com New Brunswick Scientific 44 Talmadge Road Edison, NJ 08818 (800) 631-5417 FAX: (732) 287-4222 (732) 287-1200 http://www.nbsc.com New England Biolabs (NEB) 32 Tozer Road Beverly, MA 01915 (800) 632-5227 FAX: (800) 632-7440 http://www.neb.com New England Nuclear (NEN) See NEN Life Science Products New MBR Gubelstrasse 48 CH8050 Zurich, Switzerland (41) 1-313-0703
Nichols Institute Diagnostics 33051 Calle Aviador San Juan Capistrano, CA 92675 (800) 286-4NID FAX: (949) 240-5273 (949) 728-4610 http://www.nicholsdiag.com Nichols Scientific Instruments 3334 Brown Station Road Columbia, MO 65202 (573) 474-5522 FAX: (603) 215-7274 http://home.beseen.com technology/nsi technology Nicolet Biomedical Instruments 5225 Verona Road, Building 2 Madison, WI 53711 (800) 356-0007 FAX: (608) 441-2002 (608) 273-5000 http://nicoletbiomedical.com N.I.G.M.S. (National Institute of General Medical Sciences) See Coriell Institute for Medical Research Nikon Science and Technologies Group 1300 Walt Whitman Road Melville, NY 11747 (516) 547-8500 FAX: (516) 547-4045 http://www.nikonusa.com Nippon Gene 1-29, Ton-ya-machi Toyama 930, Japan (81) 764-51-6548 FAX: (81) 764-51-6547
Newark Electronics 4801 N. Ravenswood Avenue Chicago, IL 60640 (800) 4-NEWARK FAX: (773) 907-5339 (773) 784-5100 http://www.newark.com Newell Rubbermaid 29 E. Stephenson Street Freeport, IL 61032 (815) 235-4171 FAX: (815) 233-8060 http://www.newellco.com Newport Biosystems 1860 Trainor Street Red Bluff, CA 96080 (530) 529-2448 FAX: (530) 529-2648 Newport 1791 Deere Avenue Irvine, CA 92606 (800) 222-6440 FAX: (949) 253-1800 (949) 253-1462 http://www.newport.com Nexin Research B.V. P.O. Box 16 4740 AA Hoeven, The Netherlands (31) 165-503172 FAX: (31) 165-502291 NIAID See Bio-Tech Research Laboratories
Noldus Information Technology 751 Miller Drive Suite E-5 Leesburg, VA 20175 (800) 355-9541 FAX: (703) 771-0441 (703) 771-0440 http://www.noldus.com Nordion International See MDS Nordion North American Biologicals (NABI) 16500 NW 15th Avenue Miami, FL 33169 (800) 327-7106 (305) 625-5305 http://www.nabi.com North American Reiss See Reiss Northwestern Bottle 24 Walpole Park South Walpole, MA 02081 (508) 668-8600 FAX: (508) 668-7790 NOVA Biomedical Nova Biomedical 200 Prospect Street Waltham, MA 02454 (800) 822-0911 FAX: (781) 894-5915 http://www.novabiomedical.com
Suppliers
22 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
Novagen 601 Science Drive Madison, WI 53711 (800) 526-7319 FAX: (608) 238-1388 (608) 238-6110 http://www.novagen.com Novartis 59 Route 10 East Hanover, NJ 07936 (800) 526-0175 FAX: (973) 781-6356 http://www.novartis.com Novartis Biotechnology 3054 Cornwallis Road Research Triangle Park, NC 27709 (888) 462-7288 FAX: (919) 541-8585 http://www.novartis.com Nova Sina AG Subsidiary of Airflow Lufttechnik GmbH Kleine Heeg 21 52259 Rheinbach, Germany (49) 02226 920-0 FAX: (49) 02226 9205-11 Novex/Invitrogen 1600 Faraday Carlsbad, CA 92008 (800) 955-6288 FAX: (760) 603-7201 http://www.novex.com Novo Nordisk Biochem 77 Perry Chapel Church Road Franklington, NC 27525 (800) 879-6686 FAX: (919) 494-3450 (919) 494-3000 http://www.novo.dk Novo Nordisk BioLabs See Novo Nordisk Biochem Novocastra Labs Balliol Business Park West Benton Lane Newcastle-upon-Tyne Tyne and Wear NE12 8EW, UK (44) 191-215-0567 FAX: (44) 191-215-1152 http://www.novocastra.co.uk Novus Biologicals P.O. Box 802 Littleton, CO 80160 (888) 506-6887 FAX: (303) 730-1966 http://www.novus-biologicals.com/ main.html NPI Electronic Hauptstrasse 96 D-71732 Tamm, Germany (49) 7141-601534 FAX: (49) 7141-601266 http://www.npielectronic.com NSG Precision Cells 195G Central Avenue Farmingdale, NY 11735 (516) 249-7474 FAX: (516) 249-8575 http://www.nsgpci.com
Nu Chek Prep 109 West Main P.O. Box 295 Elysian, MN 56028 (800) 521-7728 FAX: (507) 267-4790 (507) 267-4689 Nuclepore See Costar Numonics 101 Commerce Drive Montgomeryville, PA 18936 (800) 523-6716 FAX: (215) 361-0167 (215) 362-2766 http://www.interactivewhiteboards.com NYCOMED AS Pharma c/o Accurate Chemical & Scientific 300 Shames Drive Westbury, NY 11590 (800) 645-6524 FAX: (516) 997-4948 (516) 333-2221 http://www.accuratechemical.com Nycomed Amersham Health Care Division 101 Carnegie Center Princeton, NJ 08540 (800) 832-4633 FAX: (800) 807-2382 (609) 514-6000 http://www.nycomed-amersham.com Nyegaard Herserudsvagen 5254 S-122 06 Lidingo, Sweden (46) 8-765-2930 Ohmeda Catheter Products See Datex-Ohmeda Ohwa Tsusbo Hiby Dai Building 1-2-2 Uchi Saiwai-cho Chiyoda-ku Tokyo 100, Japan 03-3591-7348 FAX: 03-3501-9001 Oligos Etc. 29970 SW Town Centre Loop West, Suite B419 Wilsonville, OR 97070 (800) 888-2358 FAX: (800) 869-0813 Olis Instruments 130 Conway Drive Bogart, GA 30622 (706) 353-6547 (800) 852-3504 http://www.olisweb.com Olympus America 2 Corporate Center Drive Melville, NY 11747 (800) 645-8160 FAX: (516) 844-5959 (516) 844-5000 http://www.olympusamerica.com
Omega Engineering One Omega Drive P.O. Box 4047 Stamford, CT 06907 (800) 848-4286 FAX: (203) 359-7700 (203) 359-1660 http://www.omega.com Omega Optical 3 Grove Street P.O. Box 573 Brattleboro, VT 05302 (802) 254-2690 FAX: (802) 254-3937 http://www.omegafilters.com Omni International 6530 Commerce Court Warrenton, VA 20187 (800) 776-4431 FAX: (540) 347-5352 (540) 347-5331 http://www.omni-inc.com Omnion 2010 Energy Drive P.O. Box 879 East Troy, WI 53120 (262) 642-7200 FAX: (262) 642-7760 http://www.omnion.com Omnitech Electronics See AccuScan Instruments Oncogene Research Products P.O. Box Box 12087 La Jolla, CA 92039-2087 (800) 662-2616 FAX: (800) 766-0999 http://www.apoptosis.com Oncogene Science See OSI Pharmaceuticals Oncor See Intergen Operon Technologies 1000 Atlantic Avenue Alameda, CA 94501 (800) 688-2248 FAX: (510) 865-5225 (510) 865-8644 http://www.operon.com Optiscan P.O. Box 1066 Mount Waverly MDC, Victoria Australia 3149 61-3-9538 3333 FAX: 61-3-9562 7742 http://www.optiscan.com.au Optomax 9 Ash Street P.O. Box 840 Hollis, NH 03049 (603) 465-3385 FAX: (603) 465-2291 Opto-Line Associates 265 Ballardvale Street Wilmington, MA 01887 (978) 658-7255 FAX: (978) 658-7299 http://www.optoline.com
Orbigen 6827 Nancy Ridge Drive San Diego, CA 92121 (866) 672-4436 (858) 362-2030 (858) 362-2026 http://www.orbigen.com Oread BioSaftey 1501 Wakarusa Drive Lawrence, KS 66047 (800) 447-6501 FAX: (785) 749-1882 (785) 749-0034 http://www.oread.com Organomation Associates 266 River Road West Berlin, MA 01503 (888) 978-7300 FAX: (978)838-2786 (978) 838-7300 http://www.organomation.com Organon 375 Mount Pleasant Avenue West Orange, NJ 07052 (800) 241-8812 FAX: (973) 325-4589 (973) 325-4500 http://www.organon.com Organon Teknika (Canada) 30 North Wind Place Scarborough, Ontario M1S 3R5 Canada (416) 754-4344 FAX: (416) 754-4488 http://www.organonteknika.com Organon Teknika Cappel 100 Akzo Avenue Durham, NC 27712 (800) 682-2666 FAX: (800) 432-9682 (919) 620-2000 FAX: (919) 620-2107 http://www.organonteknika.com Oriel Corporation of America 150 Long Beach Blvd. Stratford, CT 06615 (203) 377-8282 FAX: (203) 378-2457 http://www.oriel.com OriGene Technologies 6 Taft Court, Suite 300 Rockville, MD 20850 (888) 267-4436 FAX: (301) 340-9254 (301) 340-3188 http://www.origene.com OriginLab One Roundhouse Plaza Northhampton, MA 01060 (800) 969-7720 FAX: (413) 585-0126 http://www.originlab.com Orion Research 500 Cummings Center Beverly, MA 01915 (800) 225-1480 FAX: (978) 232-6015 (978) 232-6000 http://www.orionres.com
Suppliers
23 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Orochem Technologies 331 Eisenhower Lane South Lombard, IL 60148 (630) 916-0225 FAX: 630-916-0250 http://www.orochem.com Ortho Diagnostic Systems Subsidiary of Johnson & Johnson 1001 U.S. Highway 202 P.O. Box 350 Raritan, NJ 08869 (800) 322-6374 FAX: (908) 218-8582 (908) 218-1300 Ortho McNeil Pharmaceutical Welsh & McKean Road Spring House, PA 19477 (800) 682-6532 (215) 628-5000 http://www.orthomcneil.com Oryza 200 Turnpike Road, Unit 5 Chelmsford, MA 01824 (978) 256-8183 FAX: (978) 256-7434 http://www.oryzalabs.com OSI Pharmaceuticals 106 Charles Lindbergh Blvd. Uniondale, NY 11553 (800) 662-2616 FAX: (516) 222-0114 (516) 222-0023 http://www.osip.com Osmonics 135 Flanders Road P.O. Box 1046 Westborough, MA 01581 (800) 444-8212 FAX: (508) 366-5840 (508) 366-8212 http://www.osmolabstore.com Oster Professional Products 150 Cadillac Lane McMinnville, TN 37110 (931) 668-4121 FAX: (931) 668-4125 http://www.sunbeam.com Out Patient Services 1260 Holm Road Petaluma, CA 94954 (800) 648-1666 FAX: (707) 762-7198 (707) 763-1581 OWL Scientific Plastics See OWL Separation Systems OWL Separation Systems 55 Heritage Avenue Portsmouth, NH 03801 (800) 242-5560 FAX: (603) 559-9258 (603) 559-9297 http://www.owlsci.com Oxford Biochemical Research P.O. Box 522 Oxford, MI 48371 (800) 692-4633 FAX: (248) 852-4466 http://www.oxfordbiomed.com Oxford GlycoSystems See Glyco
Oxford Instruments Old Station Way Eynsham Witney, Oxfordshire OX8 1TL, UK (44) 1865-881437 FAX: (44) 1865-881944 http://www.oxinst.com Oxford Labware See Kendall Oxford Molecular Group Oxford Science Park The Medawar Centre Oxford OX4 4GA, UK (44) 1865-784600 FAX: (44) 1865-784601 http://www.oxmol.co.uk Oxford Molecular Group 2105 South Bascom Avenue, Suite 200 Campbell, CA 95008 (800) 876-9994 FAX: (408) 879-6302 (408) 879-6300 http://www.oxmol.com OXIS International 6040 North Cutter Circle Suite 317 Portland, OR 97217 (800) 547-3686 FAX: (503) 283-4058 (503) 283-3911 http://www.oxis.com Oxoid 800 Proctor Avenue Ogdensburg, NY 13669 (800) 567-8378 FAX: (613) 226-3728 http://www.oxoid.ca Oxoid Wade Road Basingstoke, Hampshire RG24 8PW, UK (44) 1256-841144 FAX: (4) 1256-814626 http://www.oxoid.ca Oxyrase P.O. Box 1345 Mansfield, OH 44901 (419) 589-8800 FAX: (419) 589-9919 http://www.oxyrase.com Ozyme 10 Avenue Amp`ere Montigny de Bretoneux 78180 France (33) 13-46-02-424 FAX: (33) 13-46-09-212 http://www.ozyme.fr PAA Laboratories 2570 Route 724 P.O. Box 435 Parker Ford, PA 19457 (610) 495-9400 FAX: (610) 495-9410 http://www.paa-labs.com
Pacer Scientific 5649 Valley Oak Drive Los Angeles, CA 90068 (323) 462-0636 FAX: (323) 462-1430 http://www.pacersci.com Pacific Bio-Marine Labs P.O. Box 1348 Venice, CA 90294 (310) 677-1056 FAX: (310) 677-1207 Packard Instrument 800 Research Parkway Meriden, CT 06450 (800) 323-1891 FAX: (203) 639-2172 (203) 238-2351 http://www.packardinst.com
Pel-Freez Biologicals 219 N. Arkansas P.O. Box 68 Rogers, AR 72757 (800) 643-3426 FAX: (501) 636-3562 (501) 636-4361 http://www.pelfreez-bio.com Pel-Freez Clinical Systems Subsidiary of Pel-Freez Biologicals 9099 N. Deerbrook Trail Brown Deer, WI 53223 (800) 558-4511 FAX: (414) 357-4518 (414) 357-4500 http://www.pelfreez-bio.com
Padgett Instrument 1730 Walnut Street Kansas City, MO 64108 (816) 842-1029
Peninsula Laboratories 601 Taylor Way San Carlos, CA 94070 (800) 650-4442 FAX: (650) 595-4071 (650) 592-5392 http://www.penlabs.com
Pall Filtron 50 Bearfoot Road Northborough, MA 01532 (800) FILTRON FAX: (508) 393-1874 (508) 393-1800
Pentex 24562 Mando Drive Laguna Niguel, CA 92677 (800) 382-4667 FAX: (714) 643-2363 http://www.pentex.com
Pall-Gelman 25 Harbor Park Drive Port Washington, NY 11050 (800) 289-6255 FAX: (516) 484-2651 (516) 484-3600 http://www.pall.com
PeproTech 5 Crescent Avenue P.O. Box 275 Rocky Hill, NJ 08553 (800) 436-9910 FAX: (609) 497-0321 (609) 497-0253 http://www.peprotech.com
PanVera 545 Science Drive Madison, WI 53711 (800) 791-1400 FAX: (608) 233-3007 (608) 233-9450 http://www.panvera.com
Peptide Institute 4-1-2 Ina, Minoh-shi Osaka 562-8686, Japan 81-727-29-4121 FAX: 81-727-29-4124 http://www.peptide.co.jp
Parke-Davis See Warner-Lambert Parr Instrument 211 53rd Street Moline, IL 61265 (800) 872-7720 FAX: (309) 762-9453 (309) 762-7716 http://www.parrinst.com Partec Otto Hahn Strasse 32 D-48161 Munster, Germany (49) 2534-8008-0 FAX: (49) 2535-8008-90
Peptide Laboratory 4175 Lakeside Drive Richmond, CA 94806 (800) 858-7322 FAX: (510) 262-9127 (510) 262-0800 http://www.peptidelab.com Peptides International 11621 Electron Drive Louisville, KY 40299 (800) 777-4779 FAX: (502) 267-1329 (502) 266-8787 http://www.pepnet.com Perceptive Science Instruments 2525 South Shore Blvd., Suite 100 League City, TX 77573 (281) 334-3027 FAX: (281) 538-2222 http://www.persci.com
PCR See Archimica Florida PE Biosystems 850 Lincoln Centre Drive Foster City, CA 94404 (800) 345-5224 FAX: (650) 638-5884 (650) 638-5800 http://www.pebio.com
Perimed 4873 Princeton Drive North Royalton, OH 44133 (440) 877-0537 FAX: (440) 877-0534 http://www.perimed.se
Suppliers
24 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
Perkin-Elmer 761 Main Avenue Norwalk, CT 06859 (800) 762-4002 FAX: (203) 762-6000 (203) 762-1000 http://www.perkin-elmer.com See also PE Biosystems PerSeptive Bioresearch Products See PerSeptive BioSystems PerSeptive BioSystems 500 Old Connecticut Path Framingham, MA 01701 (800) 899-5858 FAX: (508) 383-7885 (508) 383-7700 http://www.pbio.com PerSeptive Diagnostic See PE Biosystems (800) 343-1346 Pettersson Elektronik AB Tallbacksvagen 51 S-756 45 Uppsala, Sweden (46) 1830-3880 FAX: (46) 1830-3840 http://www.bahnhof.se/∼pettersson Pfanstiehl Laboratories, Inc. 1219 Glen Rock Avenue Waukegan, IL 60085 (800) 383-0126 FAX: (847) 623-9173 http://www.pfanstiehl.com PGC Scientifics 7311 Governors Way Frederick, MD 21704 (800) 424-3300 FAX: (800) 662-1112 (301) 620-7777 FAX: (301) 620-7497 http://www.pgcscientifics.com Pharmacia Biotech See Amersham Pharmacia Biotech Pharmacia Diagnostics See Wallac Pharmacia LKB Biotech See Amersham Pharmacia Biotech Pharmacia LKB Biotechnology See Amersham Pharmacia Biotech Pharmacia LKB Nuclear See Wallac Pharmaderm Veterinary Products 60 Baylis Road Melville, NY 11747 (800) 432-6673 http://www.pharmaderm.com Pharmed (Norton) Norton Performance Plastics See Saint-Gobain Performance Plastics
PHLS Centre for Applied Microbiology and Research See European Collection of Animal Cell Cultures (ECACC) Phoenix Flow Systems 11575 Sorrento Valley Road, Suite 208 San Diego, CA 92121 (800) 886-3569 FAX: (619) 259-5268 (619) 453-5095 http://www.phnxflow.com Phoenix Pharmaceutical 4261 Easton Road, P.O. Box 6457 St. Joseph, MO 64506 (800) 759-3644 FAX: (816) 364-4969 (816) 364-5777 http://www.phoenixpharmaceutical.com Photometrics See Roper Scientific Photon Technology International 1 Deerpark Drive, Suite F Monmouth Junction, NJ 08852 (732) 329-0910 FAX: (732) 329-9069 http://www.pti-nj.com Physik Instrumente Polytec PI 23 Midstate Drive, Suite 212 Auburn, MA 01501 (508) 832-3456 FAX: (508) 832-0506 http://www.polytecpi.com Physitemp Instruments 154 Huron Avenue Clifton, NJ 07013 (800) 452-8510 FAX: (973) 779-5954 (973) 779-5577 http://www.physitemp.com Pico Technology The Mill House, Cambridge Street St. Neots, Cambridgeshire PE19 1QB, UK (44) 1480-396-395 FAX: (44) 1480-396-296 http://www.picotech.com Pierce Chemical P.O. Box 117 3747 Meridian Road Rockford, IL 61105 (800) 874-3723 FAX: (800) 842-5007 FAX: (815) 968-7316 http://www.piercenet.com
PharMingen See BD PharMingen
Pierce & Warriner 44, Upper Northgate Street Chester, Cheshire CH1 4EF, UK (44) 1244 382 525 FAX: (44) 1244 373 212 http://www.piercenet.com
Phenomex 2320 W. 205th Street Torrance, CA 90501 (310) 212-0555 FAX: (310) 328-7768 http://www.phenomex.com
Pilling Weck Surgical 420 Delaware Drive Fort Washington, PA 19034 (800) 523-2579 FAX: (800) 332-2308 http://www.pilling-weck.com
PixelVision A division of Cybex Computer Products 14964 NW Greenbrier Parkway Beaverton, OR 97006 (503) 629-3210 FAX: (503) 629-3211 http://www.pixelvision.com P.J. Noyes P.O. Box 381 89 Bridge Street Lancaster, NH 03584 (800) 522-2469 FAX: (603) 788-3873 (603) 788-4952 http://www.pjnoyes.com Plas-Labs 917 E. Chilson Street Lansing, MI 48906 (800) 866-7527 FAX: (517) 372-2857 (517) 372-7177 http://www.plas-labs.com Plastics One 6591 Merriman Road, Southwest P.O. Box 12004 Roanoke, VA 24018 (540) 772-7950 FAX: (540) 989-7519 http://www.plastics1.com Platt Electric Supply 2757 6th Avenue South Seattle, WA 98134 (206) 624-4083 FAX: (206) 343-6342 http://www.platt.com Polaroid 784 Memorial Drive Cambridge, MA 01239 (800) 225-1618 FAX: (800) 832-9003 (781) 386-2000 http://www.polaroid.com Polyfiltronics 136 Weymouth St. Rockland, MA 02370 (800) 434-7659 FAX: (781) 878-0822 (781) 878-1133 http://www.polyfiltronics.com
Polymicro Technologies 18019 North 25th Avenue Phoenix, AZ 85023 (602) 375-4100 FAX: (602) 375-4110 http://www.polymicro.com Polyphenols AS Hanabryggene Technology Centre Hanaveien 4-6 4327 Sandnes, Norway (47) 51-62-0990 FAX: (47) 51-62-51-82 http://www.polyphenols.com Polysciences 400 Valley Road Warrington, PA 18976 (800) 523-2575 FAX: (800) 343-3291 http://www.polysciences.com Polyscientific 70 Cleveland Avenue Bayshore, NY 11706 (516) 586-0400 FAX: (516) 254-0618 Polytech Products 285 Washington Street Somerville, MA 02143 (617) 666-5064 FAX: (617) 625-0975 Polytron 8585 Grovemont Circle Gaithersburg, MD 20877 (301) 208-6597 FAX: (301) 208-8691 http://www.polytron.com Popper and Sons 300 Denton Avenue P.O. Box 128 New Hyde Park, NY 11040 (888) 717-7677 FAX: (800) 557-6773 (516) 248-0300 FAX: (516) 747-1188 http://www.popperandsons.com Porphyrin Products P.O. Box 31 Logan, UT 84323 (435) 753-1901 FAX: (435) 753-6731 http://www.porphyrin.com
Polylabo Paul Block Parc Tertiare de la Meinau 10, rue de la Durance B.P. 36 67023 Strasbourg Cedex 1 Strasbourg, France 33-3-8865-8020 FAX: 33-3-8865-8039
Portex See SIMS Portex Limited
PolyLC 9151 Rumsey Road, Suite 180 Columbia, MD 21045 (410) 992-5400 FAX: (410) 730-8340
Praxair 810 Jorie Boulevard Oak Brook, IL 60521 (800) 621-7100 http://www.praxair.com
Polymer Laboratories Amherst Research Park 160 Old Farm Road Amherst, MA 01002 (800) 767-3963 FAX: (413) 253-2476 http://www.polymerlabs.com
Powderject Vaccines 585 Science Drive Madison, WI 53711 (608) 231-3150 FAX: (608) 231-6990 http://www.powderject.com
Precision Dynamics 13880 Del Sur Street San Fernando, CA 91340 (818) 897-1111 FAX: (818) 899-4045 http://www.pdcorp.com
Suppliers
25 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Precision Scientific Laboratory Equipment Division of Jouan 170 Marcel Drive Winchester, VA 22602 (800) 621-8820 FAX: (540) 869-0130 (540) 869-9892 http://www.precisionsci.com Primary Care Diagnostics See Becton Dickinson Primary Care Diagnostics Primate Products 1755 East Bayshore Road, Suite 28A Redwood City, CA 94063 (650) 368-0663 FAX: (650) 368-0665 http://www.primateproducts.com
Progen Biotechnik Maass-Str. 30 69123 Heidelberg, Germany (49) 6221-8278-0 FAX: (49) 6221-8278-23 http://www.progen.de
Purina Mills LabDiet P. O. Box 66812 St. Louis, MO 63166 (800) 227-8941 FAX: (314) 768-4894 http://www.purina-mills.com
R & D Systems 614 McKinley Place NE Minneapolis, MN 55413 (800) 343-7475 FAX: (612) 379-6580 (612) 379-2956 http://www.rndsystems.com
Prolabo A division of Merck Eurolab 54 rue Roger Salengro 94126 Fontenay Sous Bois Cedex France 33-1-4514-8500 FAX: 33-1-4514-8616 http://www.prolabo.fr
Qbiogene 2251 Rutherford Road Carlsbad, CA 92008 (800) 424-6101 FAX: (760) 918-9313 http://www.qbiogene.com
R & S Technology 350 Columbia Street Peacedale, RI 02880 (401) 789-5660 FAX: (401) 792-3890 http://www.septech.com
Qiagen 28159 Avenue Stanford Valencia, CA 91355 (800) 426-8157 FAX: (800) 718-2056 http://www.qiagen.com
RACAL Health and Safety See 3M 7305 Executive Way Frederick, MD 21704 (800) 692-9500 FAX: (301) 695-8200
Quality Biological 7581 Lindbergh Drive Gaithersburg, MD 20879 (800) 443-9331 FAX: (301) 840-5450 (301) 840-9331 http://www.qualitybiological.com
Radiometer America 811 Sharon Drive Westlake, OH 44145 (800) 736-0600 FAX: (440) 871-2633 (440) 871-8900 http://www.rameusa.com
5 Prime → 3 Prime See 2000 Eppendorf-5 Prime http://www.5prime.com
Proligo 2995 Wilderness Place Boulder, CO 80301 (888) 80-OLIGO FAX: (303) 801-1134 http://www.proligo.com
Prime Synthesis, Inc. 2 New Road, Ste. 126 Aston, PA 19014 (800) 743-PRIME FAX: (610) 558-5923 (610) 558-5920 http://www.primesynthesis.com
Promega 2800 Woods Hollow Road Madison, WI 53711 (800) 356-9526 FAX: (800) 356-1970 (608) 274-4330 FAX: (608) 277-2516 http://www.promega.com
Princeton Applied Research PerkinElmer Instr.: Electrochemistry 801 S. Illinois Oak Ridge, TN 37830 (800) 366-2741 FAX: (423) 425-1334 (423) 481-2442 http://www.eggpar.com
Protein Databases (PDI) 405 Oakwood Road Huntington Station, NY 11746 (800) 777-6834 FAX: (516) 673-4502 (516) 673-3939
Princeton Instruments A division of Roper Scientific 3660 Quakerbridge Road Trenton, NJ 08619 (609) 587-9797 FAX: (609) 587-1970 http://www.prinst.com Princeton Separations P.O. Box 300 Aldephia, NJ 07710 (800) 223-0902 FAX: (732) 431-3768 (732) 431-3338 Prior Scientific 80 Reservoir Park Drive Rockland, MA 02370 (781) 878-8442 FAX: (781) 878-8736 http://www.prior.com PRO Scientific P.O. Box 448 Monroe, CT 06468 (203) 452-9431 FAX: (203) 452-9753 http://www.proscientific.com Professional Compounding Centers of America 9901 South Wilcrest Drive Houston, TX 77099 (800) 331-2498 FAX: (281) 933-6227 (281) 933-6948 http://www.pccarx.com
Protein Polymer Technologies 10655 Sorrento Valley Road San Diego, CA 92121 (619) 558-6064 FAX: (619) 558-6477 http://www.ppti.com Protein Solutions 391 G Chipeta Way Salt Lake City, UT 84108 (801) 583-9301 FAX: (801) 583-4463 http://www.proteinsolutions.com Prozyme 1933 Davis Street, Suite 207 San Leandro, CA 94577 (800) 457-9444 FAX: (510) 638-6919 (510) 638-6900 http://www.prozyme.com PSI See Perceptive Science Instruments Pulmetrics Group 82 Beacon Street Chestnut Hill, MA 02167 (617) 353-3833 FAX: (617) 353-6766 Purdue Frederick 100 Connecticut Avenue Norwalk, CT 06850 (800) 633-4741 FAX: (203) 838-1576 (203) 853-0123 http://www.pharma.com
Quantum Appligene Parc d’Innovation Rue Geller de Kayserberg 67402 Illkirch, Cedex, France (33) 3-8867-5425 FAX: (33) 3-8867-1945 http://www.quantum-appligene.com Quantum Biotechnologies See Qbiogene Quantum Soft Postfach 6613 CH-8023 Z¨urich, Switzerland FAX: 41-1-481-69-51 profi[email protected] Questcor Pharmaceuticals 26118 Research Road Hayward, CA 94545 (510) 732-5551 FAX: (510) 732-7741 http://www.questcor.com Quidel 10165 McKellar Court San Diego, CA 92121 (800) 874-1517 FAX: (858) 546-8955 (858) 552-1100 http://www.quidel.com R-Biopharm 7950 Old US 27 South Marshall, MI 49068 (616) 789-3033 FAX: (616) 789-3070 http://www.r-biopharm.com R. C. Electronics 6464 Hollister Avenue Santa Barbara, CA 93117 (805) 685-7770 FAX: (805) 685-5853 http://www.rcelectronics.com
Radiometer A/S The Chemical Reference Laboratory kandevej 21 DK-2700 Brnshj, Denmark 45-3827-3827 FAX: 45-3827-2727 Radionics 22 Terry Avenue Burlington, MA 01803 (781) 272-1233 FAX: (781) 272-2428 http://www.radionics.com Radnoti Glass Technology 227 W. Maple Avenue Monrovia, CA 91016 (800) 428-1416 FAX: (626) 303-2998 (626) 357-8827 http://www.radnoti.com Rainin Instrument Rainin Road P.O. Box 4026 Woburn, MA 01888 (800)-4-RAININ FAX: (781) 938-1152 (781) 935-3050 http://www.rainin.com Rank Brothers 56 High Street Bottisham, Cambridge CB5 9DA UK (44) 1223 811369 FAX: (44) 1223 811441 http://www.rankbrothers.com Rapp Polymere Ernst-Simon Strasse 9 D 72072 T¨ubingen, Germany (49) 7071-763157 FAX: (49) 7071-763158 http://www.rapp-polymere.com
Suppliers
26 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
Rasayan, Inc. 2802 Crystal Ridge Road Encinitas, CA 92024 (760) 944-1541 FAX: (760) 944-1543 http://www.rasayan.us Raven Biological Laboratories 8607 Park Drive P.O. Box 27261 Omaha, NE 68127 (800) 728-5702 FAX: (402) 593-0995 (402) 593-0781 http://www.ravenlabs.com Raytek Corporation 1201 Shaffer Road P.O. Box 1820 Santa Cruz, CA 95061-1820 (800) 227-8074 FAX: (831) 425-4561 (831) 458-1110 www.raytek.com Razel Scientific Instruments 100 Research Drive Stamford, CT 06906 (203) 324-9914 FAX: (203) 324-5568 Reagents International See Biotech Source Receptor Biology 10000 Virginia Manor Road, Suite 360 Beltsville, MD 20705 (888) 707-4200 FAX: (301) 210-6266 (301) 210-4700 http://www.receptorbiology.com Regis Technologies 8210 N. Austin Avenue Morton Grove, IL 60053 (800) 323-8144 FAX: (847) 967-1214 (847) 967-6000 http://www.registech.com
Reming Bioinstruments 6680 County Route 17 Redfield, NY 13437 (315) 387-3414 FAX: (315) 387-3415 RepliGen 117 Fourth Avenue Needham, MA 02494 (800) 622-2259 FAX: (781) 453-0048 (781) 449-9560 http://www.repligen.com Research Biochemicals 1 Strathmore Road Natick, MA 01760 (800) 736-3690 FAX: (800) 736-2480 (508) 651-8151 FAX: (508) 655-1359 http://www.resbio.com Research Corporation Technologies 101 N. Wilmot Road, Suite 600 Tucson, AZ 85711 (520) 748-4400 FAX: (520) 748-0025 http://www.rctech.com Research Diagnostics Pleasant Hill Road Flanders, NJ 07836 (800) 631-9384 FAX: (973) 584-0210 (973) 584-7093 http://www.researchd.com Research Diets 121 Jersey Avenue New Brunswick, NJ 08901 (877) 486-2486 FAX: (732) 247-2340 (732) 247-2390 http://www.researchdiets.com
Reichert Ophthalmic Instruments P.O. Box 123 Buffalo, NY 14240 (716) 686-4500 FAX: (716) 686-4545 http://www.reichert.com
Research Genetics 2130 South Memorial Parkway Huntsville, AL 35801 (800) 533-4363 FAX: (256) 536-9016 (256) 533-4363 http://www.resgen.com
Reiss 1 Polymer Place P.O. Box 60 Blackstone, VA 23824 (800) 356-2829 FAX: (804) 292-1757 (804) 292-1600 http://www.reissmfg.com
Research Instruments Kernick Road Pernryn Cornwall TR10 9DQ, UK (44) 1326-372-753 FAX: (44) 1326-378-783 http://www.research-instruments.com
Reliable Biopharmaceutical Corporation 1945 Walton Road St. Louis, MO 63114 (314) 429-7700 FAX: (314) 429-0937 http://www.reliablebiopharm.com Remel 12076 Santa Fe Trail Drive P.O. Box 14428 Shawnee Mission, KS 66215 (800) 255-6730 FAX: (800) 621-8251 (913) 888-0939 FAX: (913) 888-5884 http://www.remelinc.com
Research Organics 4353 E. 49th Street Cleveland, OH 44125 (800) 321-0570 FAX: (216) 883-1576 (216) 883-8025 http://www.resorg.com Research Plus P.O. Box 324 Bayonne, NJ 07002 (800) 341-2296 FAX: (201) 823-9590 (201) 823-3592 http://www.researchplus.com
Research Products International 410 N. Business Center Drive Mount Prospect, IL 60056 (800) 323-9814 FAX: (847) 635-1177 (847) 635-7330 http://www.rpicorp.com Research Triangle Institute P.O. Box 12194 Research Triangle Park, NC 27709 (919) 541-6000 FAX: (919) 541-6515 http://www.rti.org Restek 110 Benner Circle Bellefonte, PA 16823 (800) 356-1688 FAX: (814) 353-1309 (814) 353-1300 http://www.restekcorp.com Rheodyne P.O. Box 1909 Rohnert Park, CA 94927 (707) 588-2000 FAX: (707) 588-2020 http://www.rheodyne.com Rhodia Organics 259 Prospect Plains Road-CN 7500 Cranbury, NJ 08512-7500 (609) 860-3891 FAX: (609) 860-1841 http://www.rhodia-ppa.com Rhone Merieux See Merial Limited Rhone-Poulenc 2 T W Alexander Drive P.O. Box 12014 Research Triangle Park, NC 08512 (919) 549-2000 FAX: (919) 549-2839 http://www.Rhone-Poulenc.com Also see Aventis Rhone-Poulenc Rorer 500 Arcola Road Collegeville, PA 19426 (800) 727-6737 FAX: (610) 454-8940 (610) 454-8975 http://www.rp-rorer.com Rhone-Poulenc Rorer Centre de Recherche de Vitry-Alfortville 13 Quai Jules Guesde, BP14 94403 Vitry Sur Seine, Cedex, France (33) 145-73-85-11 FAX: (33) 145-73-81-29 http://www.rp-rorer.com Ribi ImmunoChem Research 563 Old Corvallis Road Hamilton, MT 59840 (800) 548-7424 FAX: (406) 363-6129 (406) 363-3131 http://www.ribi.com RiboGene See Questcor Pharmaceuticals
Ricca Chemical 448 West Fork Drive Arlington, TX 76012 (888) GO-RICCA FAX: (800) RICCA-93 (817) 461-5601 http://www.riccachemical.com Richard-Allan Scientific 225 Parsons Street Kalamazoo, MI 49007 (800) 522-7270 FAX: (616) 345-3577 (616) 344-2400 http://www.rallansci.com Richelieu Biotechnologies 11 177 Hamon Montral, Quebec H3M 3E4 Canada (802) 863-2567 FAX: (802) 862-2909 http://www.richelieubio.com Richter Enterprises 20 Lake Shore Drive Wayland, MA 01778 (508) 655-7632 FAX: (508) 652-7264 http://www.richter-enterprises.com Riese Enterprises BioSure Division 12301 G Loma Rica Drive Grass Valley, CA 95945 (800) 345-2267 FAX: (916) 273-5097 (916) 273-5095 http://www.biosure.com Robbins Scientific 1250 Elko Drive Sunnyvale, CA 94086 (800) 752-8585 FAX: (408) 734-0300 (408) 734-8500 http://www.robsci.com Roboz Surgical Instruments 9210 Corporate Boulevard, Suite 220 Rockville, MD 20850 (800) 424-2984 FAX: (301) 590-1290 (301) 590-0055 Roche Diagnostics 9115 Hague Road P.O. Box 50457 Indianapolis, IN 46256 (800) 262-1640 FAX: (317) 845-7120 (317) 845-2000 http://www.roche.com Roche Molecular Systems See Roche Diagnostics Rocklabs P.O. Box 18-142 Auckland 6, New Zealand (64) 9-634-7696 FAX: (64) 9-634-7696 http://www.rocklabs.com
Suppliers
27 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Rockland P.O. Box 316 Gilbertsville, PA 19525 (800) 656-ROCK FAX: (610) 367-7825 (610) 369-1008 http://www.rockland-inc.com Rohm Chemische Fabrik Kirschenallee D-64293 Darmstadt, Germany (49) 6151-1801 FAX: (49) 6151-1802 http://www.roehm.com Roper Scientific 3440 East Brittania Drive, Suite 100 Tucson, AZ 85706 (520) 889-9933 FAX: (520) 573-1944 http://www.roperscientific.com Rosetta Inpharmatics 12040 115th Avenue NE Kirkland, WA 98034 (425) 820-8900 FAX: (425) 820-5757 http://www.rii.com ROTH-SOCHIEL 3 rue de la Chapelle Lauterbourg 67630 France (33) 03-88-94-82-42 FAX: (33) 03-88-54-63-93 Rotronic Instrument 160 E. Main Street Huntington, NY 11743 (631) 427-3898 FAX: (631) 427-3902 http://www.rotronic-usa.com Roundy’s 23000 Roundy Drive Pewaukee, WI 53072 (262) 953-7999 FAX: (262) 953-7989 http://www.roundys.com RS Components Birchington Road Weldon Industrial Estate Corby, Northants NN17 9RS, UK (44) 1536 201234 FAX: (44) 1536 405678 http://www.rs-components.com Rubbermaid See Newell Rubbermaid Safe Cells See Bionique Testing Labs Sage Instruments 240 Airport Boulevard Freedom, CA 95076 (831) 761-1000 FAX: (831) 761-1008 http://www.sageinst.com Sage Laboratories 11 Huron Drive Natick, MA 01760 (508) 653-0844 FAX: 508-653-5671 http://www.sagelabs.com
Sai Life Sciences 11344 E. San Raphael Driveway San Diego, CA 92130 (858) 793-0400 FAX: (858) 225-0421 http://www.sailifesciences.com
Sartorius 131 Heartsland Boulevard Edgewood, NY 11717 (800) 368-7178 FAX: (516) 254-4253 http://www.sartorius.com
Saint-Gobain Performance Plastics P.O. Box 3660 Akron, OH 44309 (330) 798-9240 FAX: (330) 798-6968 http://www.nortonplastics.com
SAS Institute Pacific Telesis Center One Montgomery Street San Francisco, CA 94104 (415) 421-2227 FAX: (415) 421-1213 http://www.sas.com
Samchully Pharmaceuticals 8F Samtan Building 947-7 Dechi-dong Gangnam-gu Seoul, Korea 135-735 82-(0)2-527-6300 FAX: 82-(0)2-561-6006 http://www.samchullypharm.com San Diego Instruments 7758 Arjons Drive San Diego, CA 92126 (858) 530-2600 FAX: (858) 530-2646 http://www.sd-inst.com Sandown Scientific Beards Lodge 25 Oldfield Road Hampden, Middlesex TW12 2AJ, UK (44) 2089 793300 FAX: (44) 2089 793311 http://www.sandownsci.com Sandoz Pharmaceuticals See Novartis Sanofi Recherche Centre de Montpellier 371 Rue du Professor Blayac 34184 Montpellier, Cedex 04 France (33) 67-10-67-10 FAX: (33) 67-10-67-67 Sanofi Winthrop Pharmaceuticals 90 Park Avenue New York, NY 10016 (800) 223-5511 FAX: (800) 933-3243 (212) 551-4000 http://www.sanofi-synthelabo.com/us Santa Cruz Biotechnology 2161 Delaware Avenue Santa Cruz, CA 95060 (800) 457-3801 FAX: (831) 457-3801 (831) 457-3800 http://www.scbt.com Sarasep (800) 605-0267 FAX: (408) 432-3231 (408) 432-3230 http://www.transgenomic.com Sarstedt P.O. Box 468 Newton, NC 28658 (800) 257-5101 FAX: (828) 465-4003 (828) 465-4000 http://www.sarstedt.com
Savant/EC Apparatus A ThermoQuest company 100 Colin Drive Holbrook, NY 11741 (800) 634-8886 FAX: (516) 244-0606 (516) 244-2929 http://www.savec.com Savillex 6133 Baker Road Minnetonka, MN 55345 (612) 935-5427 Scanalytics Division of CSP 8550 Lee Highway, Suite 400 Fairfax, VA 22031 (800) 325-3110 FAX: (703) 208-1960 (703) 208-2230 http://www.scanalytics.com Schering Laboratories See Schering-Plough Schering-Plough 1 Giralda Farms Madison, NJ 07940 (800) 222-7579 FAX: (973) 822-7048 (973) 822-7000 http://www.schering-plough.com Schleicher & Schuell 10 Optical Avenue Keene, NH 03431 (800) 245-4024 FAX: (603) 357-3627 (603) 352-3810 http://www.s-und-s.de/englishindex.html Science Technology Centre 1250 Herzberg Laboratories Carleton University 1125 Colonel Bay Drive Ottawa, Ontario, Canada K1S 5B6 (613) 520-4442 FAX: (613) 520-4445 http://www.carleton.ca/universities/stc Scientific Instruments 200 Saw Mill River Road Hawthorne, NY 10532 (800) 431-1956 FAX: (914) 769-5473 (914) 769-5700 http://www.scientificinstruments.com Scientific Solutions 9323 Hamilton Mentor, OH 44060 (440) 357-1400 FAX: (440) 357-1416 http://www.labmaster.com
Scion 82 Worman’s Mill Court, Suite H Frederick, MD 21701 (301) 695-7870 FAX: (301) 695-0035 http://www.scioncorp.com Scott Specialty Gases 6141 Easton Road P.O. Box 310 Plumsteadville, PA 18949 (800) 21-SCOTT FAX: (215) 766-2476 (215) 766-8861 http://www.scottgas.com Scripps Clinic and Research Foundation Instrumentation and Design Lab 10666 N. Torrey Pines Road La Jolla, CA 92037 (800) 992-9962 FAX: (858) 554-8986 (858) 455-9100 http://www.scrippsclinic.com SDI Sensor Devices 407 Pilot Court, 400A Waukesha, WI 53188 (414) 524-1000 FAX: (414) 524-1009 Sefar America 111 Calumet Street Depew, NY 14043 (716) 683-4050 FAX: (716) 683-4053 http://www.sefaramerica.com Seikagaku America Division of Associates of Cape Cod 704 Main Street Falmouth, MA 02540 (800) 237-4512 FAX: (508) 540-8680 (508) 540-3444 http://www.seikagaku.com Sellas Medizinische Gerate Hagener Str. 393 Gevelsberg-Vogelsang, 58285 Germany (49) 23-326-1225 Senn Chemicals AG 11189 Sorrento Valley Road #4 San Diego, CA 92121 (858) 450 6091 FAX: (858) 450 6096 http://www.sennchem.com Sensor Medics 22705 Savi Ranch Parkway Yorba Linda, CA 92887 (800) 231-2466 FAX: (714) 283-8439 (714) 283-2228 http://www.sensormedics.com Sensor Systems LLC 2800 Anvil Street, North Saint Petersburg, FL 33710 (800) 688-2181 FAX: (727) 347-3881 (727) 347-2181 http://www.vsensors.com
Suppliers
28 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
SenSym/Foxboro ICT 1804 McCarthy Boulevard Milpitas, CA 95035 (800) 392-9934 FAX: (408) 954-9458 (408) 954-6700 http://www.sensym.com
Shelton Scientific 230 Longhill Crossroads Shelton, CT 06484 (800) 222-2092 FAX: (203) 929-2175 (203) 929-8999 http://www.sheltonscientific.com
Separations Group See Vydac
Sherwood-Davis & Geck See Kendall
Sepracor 111 Locke Drive Marlboro, MA 01752 (877) SEPRACOR (508) 357-7300 http://www.sepracor.com
Sherwood Medical See Kendall
Sera-Lab See Harlan Sera-Lab Sermeter 925 Seton Court, #7 Wheeling, IL 60090 (847) 537-4747 Serological 195 W. Birch Street Kankakee, IL 60901 (800) 227-9412 FAX: (815) 937-8285 (815) 937-8270 Seromed Biochrom Leonorenstrasse 2-6 D-12247 Berlin, Germany (49) 030-779-9060 Serotec 22 Bankside Station Approach Kidlington, Oxford OX5 1JE, UK (44) 1865-852722 FAX: (44) 1865-373899 In the US: (800) 265-7376 http://www.serotec.co.uk Serva Biochemicals Distributed by Crescent Chemical S.F. Medical Pharmlast See Chase-Walton Elastomers SGE 2007 Kramer Lane Austin, TX 78758 (800) 945-6154 FAX: (512) 836-9159 (512) 837-7190 http://www.sge.com Shandon/Lipshaw 171 Industry Drive Pittsburgh, PA 15275 (800) 245-6212 FAX: (412) 788-1138 (412) 788-1133 http://www.shandon.com Sharpoint P.O. Box 2212 Taichung, Taiwan Republic of China (886) 4-3206320 FAX: (886) 4-3289879 http://www.sharpoint.com.tw
Shimadzu Scientific Instruments 7102 Riverwood Drive Columbia, MD 21046 (800) 477-1227 FAX: (410) 381-1222 (410) 381-1227 http://www.ssi.shimadzu.com Sialomed See Amika Siemens Analytical X-Ray Systems See Bruker Analytical X-Ray Systems Sievers Instruments Subsidiary of Ionics 6060 Spine Road Boulder, CO 80301 (800) 255-6964 FAX: (303) 444-6272 (303) 444-2009 http://www.sieversinst.com Sigma-Aldrich 3050 Spruce Street St. Louis, MO 63103 (800) 358-5287 FAX: (800) 962-9591 (800) 325-3101 FAX: (800) 325-5052 http://www.sigma-aldrich.com Silenus/Amrad 34 Wadhurst Drive Boronia, Victoria 3155 Australia (613)9887-3909 FAX: (613)9887-3912 http://www.amrad.com.au Silicon Genetics 2601 Spring Street Redwood City, CA 94063 (866) SIG-SOFT FAX: (650) 365-1735 (650) 367-9600 http://www.sigenetics.com SIMS Deltec 1265 Grey Fox Road St. Paul, Minnesota 55112 (800) 426-2448 FAX: (615) 628-7459 http://www.deltec.com SIMS Portex 10 Bowman Drive Keene, NH 03431 (800) 258-5361 FAX: (603) 352-3703 (603) 352-3812 http://www.simsmed.com SIMS Portex Limited Hythe, Kent CT21 6JL, UK (44)1303-260551 FAX: (44)1303-266761 http://www.portex.com
Siris Laboratories See Biosearch Technologies Skatron Instruments See Molecular Devices SLM Instruments See Spectronic Instruments SLM-AMINCO Instruments See Spectronic Instruments Small Parts 13980 NW 58th Court P.O. Box 4650 Miami Lakes, FL 33014 (800) 220-4242 FAX: (800) 423-9009 (305) 558-1038 FAX: (305) 558-0509 http://www.smallparts.com Smith & Nephew 11775 Starkey Road P.O. Box 1970 Largo, FL 33779 (800) 876-1261 http://www.smith-nephew.com SmithKline Beecham 1 Franklin Plaza, #1800 Philadelphia, PA 19102 (215) 751-4000 FAX: (215) 751-4992 http://www.sb.com Solid Phase Sciences See Biosearch Technologies SOMA Scientific Instruments 5319 University Drive, PMB #366 Irvine, CA 92612 (949) 854-0220 FAX: (949) 854-0223 http://somascientific.com Somatix Therapy See Cell Genesys Sonics & Materials 53 Church Hill Road Newtown, CT 06470 (800) 745-1105 FAX: (203) 270-4610 (203) 270-4600 http://www.sonicsandmaterials.com Sonosep Biotech See Triton Environmental Consultants Sorvall See Kendro Laboratory Products Southern Biotechnology Associates P.O. Box 26221 Birmingham, AL 35260 (800) 722-2255 FAX: (205) 945-8768 (205) 945-1774 http://SouthernBiotech.com SPAFAS 190 Route 165 Preston, CT 06365 (800) SPAFAS-1 FAX: (860) 889-1991 (860) 889-1389 http://www.spafas.com
Specialty Media Division of Cell & Molecular Technologies 580 Marshall Street Phillipsburg, NJ 08865 (800) 543-6029 FAX: (908) 387-1670 (908) 454-7774 http://www.specialtymedia.com Spectra Physics See Thermo Separation Products Spectramed See BOC Edwards SpectraSource Instruments 31324 Via Colinas, Suite 114 Westlake Village, CA 91362 (818) 707-2655 FAX: (818) 707-9035 http://www.spectrasource.com Spectronic Instruments 820 Linden Avenue Rochester, NY 14625 (800) 654-9955 FAX: (716) 248-4014 (716) 248-4000 http://www.spectronic.com Spectrum Medical Industries See Spectrum Laboratories Spectrum Laboratories 18617 Broadwick Street Rancho Dominguez, CA 90220 (800) 634-3300 FAX: (800) 445-7330 (310) 885-4601 FAX: (310) 885-4666 http://www.spectrumlabs.com Spherotech 1840 Industrial Drive, Suite 270 Libertyville, IL 60048 (800) 368-0822 FAX: (847) 680-8927 (847) 680-8922 http://www.spherotech.com SPSS 233 S. Wacker Drive, 11th floor Chicago, IL 60606 (800) 521-1337 FAX: (800) 841-0064 http://www.spss.com SS White Burs 1145 Towbin Avenue Lakewood, NJ 08701 (732) 905-1100 FAX: (732) 905-0987 http://www.sswhiteburs.com Stag Instruments 16 Monument Industrial Park Chalgrove, Oxon OX44 7RW, UK (44) 1865-891116 FAX: (44) 1865-890562 Standard Reference Materials Program National Institute of Standards and Technology Building 202, Room 204 Gaithersburg, MD 20899 (301) 975-6776 FAX: (301) 948-3730
Suppliers
29 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Starplex Scientific 50 Steinway Etobieoke, Ontario M9W 6Y3 Canada (800) 665-0954 FAX: (416) 674-6067 (416) 674-7474 http://www.starplexscientific.com State Laboratory Institute of Massachusetts 305 South Street Jamaica Plain, MA 02130 (617) 522-3700 FAX: (617) 522-8735 http://www.state.ma.us/dph Stedim Labs 1910 Mark Court, Suite 110 Concord, CA 94520 (800) 914-6644 FAX: (925) 689-6988 (925) 689-6650 http://www.stedim.com Stem Cell Technologies 777 West Broadway, Suite 808 Vancouver, British Columbia V5Z 4J7 Canada (800) 667-0322 FAX: (800) 567-2899 (604) 877-0713 FAX: (604) 877-0704 http://www.stemcell.com Stephens Scientific 107 Riverdale Road Riverdale, NJ 07457 (800) 831-8099 FAX: (201) 831-8009 (201) 831-9800 Steraloids P.O. Box 689 Newport, RI 02840 (401) 848-5422 FAX: (401) 848-5638 http://www.steraloids.com Sterling Medical 2091 Springdale Road, Ste. 2 Cherry Hill, NJ 08003 (800) 229-0900 FAX: (800) 229-7854 http://www.sterlingmedical.com Sterling Winthrop 90 Park Avenue New York, NY 10016 (212) 907-2000 FAX: (212) 907-3626 Sternberger Monoclonals 10 Burwood Court Lutherville, MD 21093 (410) 821-8505 FAX: (410) 821-8506 http://www.sternbergermonoclonals. com
Stratagene 11011 N. Torrey Pines Road La Jolla, CA 92037 (800) 424-5444 FAX: (888) 267-4010 (858) 535-5400 http://www.stratagene.com Strategic Applications 530A N. Milwaukee Avenue Libertyville, IL 60048 (847) 680-9385 FAX: (847) 680-9837 Strem Chemicals 7 Mulliken Way Newburyport, MA 01950 (800) 647-8736 FAX: (800) 517-8736 (978) 462-3191 FAX: (978) 465-3104 http://www.strem.com StressGen Biotechnologies Biochemicals Division 120-4243 Glanford Avenue Victoria, British Columbia V8Z 4B9 Canada (800) 661-4978 FAX: (250) 744-2877 (250) 744-2811 http://www.stressgen.com Structure Probe/SPI Supplies (Epon-Araldite) P.O. Box 656 West Chester, PA 19381 (800) 242-4774 FAX: (610) 436-5755 http://www.2spi.com Sud-Chemie ¨ Performance Packaging 101 Christine Drive Belen, NM 87002 (800) 989-3374 FAX: (505) 864-9296 http://www.uniteddesiccants.com Sumitomo Chemical Sumitomo Building 5-33, Kitahama 4-chome Chuo-ku, Osaka 541-8550, Japan (81) 6-6220-3891 FAX: (81)-6-6220-3345 http://www.sumitomo-chem.co.jp Sun Box 19217 Orbit Drive Gaithersburg, MD 20879 (800) 548-3968 FAX: (301) 977-2281 (301) 869-5980 http://www.sunboxco.com Sunbrokers See Sun International
Stoelting 502 Highway 67 Kiel, WI 53042 (920) 894-2293 FAX: (920) 894-7029 http://www.stoelting.com
Sun International 3700 Highway 421 North Wilmington, NC 28401 (800) LAB-VIAL FAX: (800) 231-7861 http://www.autosamplervial.com
Stovall Lifescience 206-G South Westgate Drive Greensboro, NC 27407 (800) 852-0102 FAX: (336) 852-3507 http://www.slscience.com
Sunox 1111 Franklin Boulevard, Unit 6 Cambridge, ON N1R 8B5, Canada (519) 624-4413 FAX: (519) 624-8378 http://www.sunox.ca
Supelco See Sigma-Aldrich SuperArray P.O. Box 34494 Bethesda, MD 20827 (888) 503-3187 FAX: (301) 765-9859 (301) 765-9888 http://www.superarray.com Surface Measurement Systems 3 Warple Mews, Warple Way London W3 ORF, UK (44) 20-8749-4900 FAX: (44) 20-8749-6749 http://www.smsuk.co.uk/index.htm SurgiVet N7 W22025 Johnson Road, Suite A Waukesha, WI 53186 (262) 513-8500 (888) 745-6562 FAX: (262) 513-9069 http://www.surgivet.com Sutter Instruments 51 Digital Drive Novato, CA 94949 (415) 883-0128 FAX: (415) 883-0572 http://www.sutter.com Swiss Precision Instruments 1555 Mittel Boulevard, Suite F Wooddale, IL 60191 (800) 221-0198 FAX: (800) 842-5164 Synaptosoft 3098 Anderson Place Decatur, GA 30033 (770) 939-4366 FAX: 770-939-9478 http://www.synaptosoft.com SynChrom See Micra Scientific Synergy Software 2457 Perkiomen Avenue Reading, PA 19606 (800) 876-8376 FAX: (610) 370-0548 (610) 779-0522 http://www.synergy.com Synteni See Incyte Synthetics Industry Lumite Division 2100A Atlantic Highway Gainesville, GA 30501 (404) 532-9756 FAX: (404) 531-1347 Systat See SPSS Systems Planning and Analysis (SPA) 2000 N. Beauregard Street Suite 400 Alexandria, VA 22311 (703) 931-3500 http://www.spa-inc.net
3M Bioapplications 3M Center Building 270-15-01 St. Paul, MN 55144 (800) 257-7459 FAX: (651) 737-5645 (651) 736-4946 T Cell Diagnostics and T Cell Sciences 38 Sidney Street Cambridge, MA 02139 (617) 621-1400 TAAB Laboratory Equipment 3 Minerva House Calleva Park Aldermaston, Berkshire RG7 8NA, UK (44) 118 9817775 FAX: (44) 118 9817881 Taconic 273 Hover Avenue Germantown, NY 12526 (800) TAC-ONIC FAX: (518) 537-7287 (518) 537-6208 http://www.taconic.com Tago See Biosource International TaKaRa Biochemical 719 Alliston Way Berkeley, CA 94710 (800) 544-9899 FAX: (510) 649-8933 (510) 649-9895 http://www.takara.co.jp/english Takara Shuzo Biomedical Group Division Seta 3-4-1 Otsu Shiga 520-21, Japan (81) 75-241-5100 FAX: (81) 77-543-9254 http://www.Takara.co.jp/english Takeda Chemical Products 101 Takeda Drive Wilmington, NC 28401 (800) 825-3328 FAX: (800) 825-0333 (910) 762-8666 FAX: (910) 762-6846 http://takeda-usa.com TAO Biomedical 73 Manassas Court Laurel Springs, NJ 08021 (609) 782-8622 FAX: (609) 782-8622 TCI America 9211 N. Harborgate Street Portland, OR 97203 (800) 423-8616 FAX: (888) 520-1075 (503) 283-1681 FAX: (503) 283-1987 http://www.tciamerica.com Tecan US P.O. Box 13953 Research Triangle Park, NC 27709 (800) 33-TECAN FAX: (919) 361-5201 (919) 361-5208 http://www.tecan-us.com
Suppliers
30 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
Techne University Park Plaza 743 Alexander Road Princeton, NJ 08540 (800) 225-9243 FAX: (609) 987-8177 (609) 452-9275 http://www.techneusa.com Technical Manufacturing 15 Centennial Drive Peabody, MA 01960 (978) 532-6330 FAX: (978) 531-8682 http://www.techmfg.com Technical Products International 5918 Evergreen St. Louis, MO 63134 (800) 729-4451 FAX: (314) 522-6360 (314) 522-8671 http://www.vibratome.com Technicon See Organon Teknika Cappel Techno-Aide P.O. Box 90763 Nashville, TN 37209 (800) 251-2629 FAX: (800) 554-6275 (615) 350-7030 http://www.techno-aid.com Ted Pella 4595 Mountain Lakes Boulevard P.O. Box 492477 Redding, CA 96049 (800) 237-3526 FAX: (530) 243-3761 (530) 243-2200 http://www.tedpella.com Tekmar-Dohrmann P.O. Box 429576 Cincinnati, OH 45242 (800) 543-4461 FAX: (800) 841-5262 (513) 247-7000 FAX: (513) 247-7050 Tektronix 142000 S.W. Karl Braun Drive Beaverton, OR 97077 (800) 621-1966 FAX: (503) 627-7995 (503) 627-7999 http://www.tek.com Tel-Test P.O. Box 1421 Friendswood, TX 77546 (800) 631-0600 FAX: (281) 482-1070 (281) 482-2672 http://www.isotex-diag.com TeleChem International 524 East Weddell Drive, Suite 3 Sunnyvale, CA 94089 (408) 744-1331 FAX: (408) 744-1711 http://www.gst.net/∼telechem Terrachem Mallaustrasse 57 D-68219 Mannheim, Germany 0621-876797-0 FAX: 0621-876797-19 http://www.terrachem.de
Terumo Medical 2101 Cottontail Lane Somerset, NJ 08873 (800) 283-7866 FAX: (732) 302-3083 (732) 302-4900 http://www.terumomedical.com Tetko 333 South Highland Manor Briarcliff, NY 10510 (800) 289-8385 FAX: (914) 941-1017 (914) 941-7767 http://www.tetko.com TetraLink 4240 Ridge Lea Road Suite 29 Amherst, NY 14226 (800) 747-5170 FAX: (800) 747-5171 http://www.tetra-link.com TEVA Pharmaceuticals USA 1090 Horsham Road P.O. Box 1090 North Wales, PA 19454 (215) 591-3000 FAX: (215) 721-9669 http://www.tevapharmusa.com Texas Fluorescence Labs 9503 Capitol View Drive Austin, TX 78747 (512) 280-5223 FAX: (512) 280-4997 http://www.teflabs.com ThermoCare P.O. Box 6069 Incline Village, NV 89450 (800) 262-4020 (775) 831-1201 Thermo Labsystems 8 East Forge Parkway Franklin, MA 02038 (800) 522-7763 FAX: (508) 520-2229 (508) 520-0009 http://www.finnpipette.com Thermometric Spjutvagen 5A S-175 61 Jarfalla, Sweden (46) 8-564-72-200 Thermoquest IEC Division 300 Second Avenue Needham Heights, MA 02194 (800) 843-1113 FAX: (781) 444-6743 (781) 449-0800 http://www.thermoquest.com Thermo Separation Products Thermoquest 355 River Oaks Parkway San Jose, CA 95134 (800) 538-7067 FAX: (408) 526-9810 (408) 526-1100 http://www.thermoquest.com
Thermo Shandon 171 Industry Drive Pittsburgh, PA 15275 (800) 547-7429 FAX: (412) 899-4045 http://www.thermoshandon.com Thomas Scientific 99 High Hill Road at I-295 Swedesboro, NJ 08085 (800) 345-2100 FAX: (800) 345-5232 (856) 467-2000 FAX: (856) 467-3087 http://www.wheatonsci.com/html/nt/ Thomas.html Thomson Instrument 354 Tyler Road Clearbrook, VA 22624 (800) 842-4752 FAX: (540) 667-6878 (800) 541-4792 FAX: (760) 757-9367 http://www.hplc.com Thorn EMI See Electron Tubes Thorlabs 435 Route 206 Newton, NJ 07860 (973) 579-7227 FAX: (973) 383-8406 http://www.thorlabs.com Tiemann See Bernsco Surgical Supply Timberline Instruments 1880 South Flatiron Court, H-2 P.O. Box 20356 Boulder, CO 80308 (800) 777-5996 FAX: (303) 440-8786 (303) 440-8779 http://www.timberlineinstruments.com Tissue-Tek A Division of Sakura Finetek USA 1750 West 214th Street Torrance, CA 90501 (800) 725-8723 FAX: (310) 972-7888 (310) 972-7800 http://www.sakuraus.com Tocris Cookson 114 Holloway Road, Suite 200 Ballwin, MO 63011 (800) 421-3701 FAX: (800) 483-1993 (636) 207-7651 FAX: (636) 207-7683 http://www.tocris.com Tocris Cookson Northpoint, Fourth Way Avonmouth, Bristol BS11 8TA, UK (44) 117-982-6551 FAX: (44) 117-982-6552 http://www.tocris.com Tokyo Kasei Kogyo Co., Ltd. (Tokyo Chemical Industry Co., Ltd.) 4-10-2 Nihonbashi-honcho Chuo-ku Tokyo 103-0023, Japan (81) 3-5640-8851 FAX: (81) 3-5640-8865 http://www.tokyokasei.co.jp/index e. html
Tomtec See CraMar Technologies TopoGen P.O. Box 20607 Columbus, OH 43220 (800) TOPOGEN FAX: (800) ADD-TOPO (614) 451-5810 FAX: (614) 451-5811 http://www.topogen.com Toray Industries, Japan Toray Building 2-1 Nihonbash-Muromach 2-Chome, Chuo-Ku Tokyo, Japan 103-8666 (03) 3245-5115 FAX: (03) 3245-5555 http://www.toray.co.jp Toray Industries, U.S.A. 600 Third Avenue New York, NY 10016 (212) 697-8150 FAX: (212) 972-4279 http://www.toray.com Toronto Research Chemicals 2 Brisbane Road North York, Ontario M3J 2J8, Canada (416) 665-9696 FAX: (416) 665-4439 http://www.trc-canada.com TosoHaas 156 Keystone Drive Montgomeryville, PA 18036 (800) 366-4875 FAX: (215) 283-5035 (215) 283-5000 http://www.tosohaas.com Towhill 647 Summer Street Boston, MA 02210 (617) 542-6636 FAX: (617) 464-0804 Toxin Technology 7165 Curtiss Avenue Sarasota, FL 34231 (941) 925-2032 FAX: (9413) 925-2130 http://www.toxintechnology.com Toyo Soda See TosoHaas Trace Analytical 3517-A Edison Way Menlo Park, CA 94025 (650) 364-6895 FAX: (650) 364-6897 http://www.traceanalytical.com Transduction Laboratories See BD Transduction Laboratories Transgenomic 2032 Concourse Drive San Jose, CA 95131 (408) 432-3230 FAX: (408) 432-3231 http://www.transgenomic.com Transonic Systems 34 Dutch Mill Road Ithaca, NY 14850 (800) 353-3569 FAX: (607) 257-7256 http://www.transonic.com
Suppliers
31 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Travenol Lab See Baxter Healthcare Tree Star Software 20 Winding Way San Carlos, CA 94070 800-366-6045 http://www.treestar.com Trevigen 8405 Helgerman Court Gaithersburg, MD 20877 (800) TREVIGEN FAX: (301) 216-2801 (301) 216-2800 http://www.trevigen.com Trilink Biotechnologies 6310 Nancy Ridge Drive San Diego, CA 92121 (800) 863-6801 FAX: (858) 546-0020 http://www.trilink.biotech.com Tripos Associates 1699 South Hanley Road, Suite 303 St. Louis, MO 63144 (800) 323-2960 FAX: (314) 647-9241 (314) 647-1099 http://www.tripos.com Triton Environmental Consultants 120-13511 Commerce Parkway Richmond, British Columbia V6V 2L1 Canada (604) 279-2093 FAX: (604) 279-2047 http://www.triton-env.com Tropix 47 Wiggins Avenue Bedford, MA 01730 (800) 542-2369 FAX: (617) 275-8581 (617) 271-0045 http://www.tropix.com
Ultrasonic Power 239 East Stephenson Street Freeport, IL 61032 (815) 235-6020 FAX: (815) 232-2150 http://www.upcorp.com Ultrasound Advice 23 Aberdeen Road London N52UG, UK (44) 020-7359-1718 FAX: (44) 020-7359-3650 http://www.ultrasoundadvice.co.uk UNELKO 14641 N. 74th Street Scottsdale, AZ 85260 (480) 991-7272 FAX: (480) 483-7674 http://www.unelko.com Unifab Corp. 5260 Lovers Lane Kalamazoo, MI 49002 (800) 648-9569 FAX: (616) 382-2825 (616) 382-2803
United Desiccants See S¨ud-Chemie Performance Packaging
V & P Scientific 9823 Pacific Heights Boulevard, Suite T San Diego, CA 92121 (800) 455-0644 FAX: (858) 455-0703 (858) 455-0643 http://www.vp-scientific.com
United States Biochemical See USB
2000 Eppendorf-5 Prime 5603 Arapahoe Avenue Boulder, CO 80303 (800) 533-5703 FAX: (303) 440-0835 (303) 440-3705
Universal Imaging 502 Brandywine Parkway West Chester, PA 19380 (610) 344-9410 FAX: (610) 344-6515 http://www.image1.com
Tyler Research 10328 73rd Avenue Edmonton, Alberta T6E 6N5 Canada (403) 448-1249 FAX: (403) 433-0479
Upchurch Scientific 619 West Oak Street P.O. Box 1529 Oak Harbor, WA 98277 (800) 426-0191 FAX: (800) 359-3460 (360) 679-2528 FAX: (360) 679-3830 http://www.upchurch.com
UltraPIX See Life Science Resources
USCI Bard Bard Interventional Products 129 Concord Road Billerica, MA 01821 (800) 225-1332 FAX: (978) 262-4805 http://www.bardinterventional.com UVP (Ultraviolet Products) 2066 W. 11th Street Upland, CA 91786 (800) 452-6788 FAX: (909) 946-3597 (909) 946-3197 http://www.uvp.com
TSI Center for Diagnostic Products See Intergen
Ugo Basile Biological Research Apparatus Via G. Borghi 43 21025 Comerio, Varese, Italy (39) 332 744 574 FAX: (39) 332 745 488 http://www.ugobasile.com
USB 26111 Miles Road P.O. Box 22400 Cleveland, OH 44122 (800) 321-9322 FAX: (800) 535-0898 FAX: (216) 464-5075 http://www.usbweb.com
Union Carbide 10235 West Little York Road, Suite 300 Houston, TX 77040 (800) 568-4000 FAX: (713) 849-7021 (713) 849-7000 http://www.unioncarbide.com
United States Biological (US Biological) P.O. Box 261 Swampscott, MA 01907 (800) 520-3011 FAX: (781) 639-1768 http://www.usbio.net
UBI See Upstate Biotechnology
USA/Scientific 346 SW 57th Avenue P.O. Box 3565 Ocala, FL 34478 (800) LAB-TIPS FAX: (352) 351-2057 (3524) 237-6288 http://www.usascientific.com
Upjohn Pharmacia & Upjohn http://www.pnu.com Upstate Biotechnology (UBI) 1100 Winter Street, Suite 2300 Waltham, MA 02451 (800) 233-3991 FAX: (781) 890-7738 (781) 890-8845 http://www.upstatebiotech.com
Valco Instruments P.O. Box 55603 Houston, TX 77255 (800) FOR-VICI FAX: (713) 688-8106 (713) 688-9345 http://www.vici.com Valpey Fisher 75 South Street Hopkin, MA 01748 (508) 435-6831 FAX: (508) 435-5289 http://www.valpeyfisher.com Value Plastics 3325 Timberline Road Fort Collins, CO 80525 (800) 404-LUER FAX: (970) 223-0953 (970) 223-8306 http://www.valueplastics.com Vangard International P.O. Box 308 3535 Rt. 66, Bldg. #4 Neptune, NJ 07754 (800) 922-0784 FAX: (732) 922-0557 (732) 922-4900 http://www.vangard1.com
Varian Analytical Instruments 2700 Mitchell Drive Walnut Creek, CA 94598 (800) 926-3000 FAX: (925) 945-2102 (925) 939-2400 http://www.varianinc.com Varian Associates 3050 Hansen Way Palo Alto, CA 94304 (800) 544-4636 FAX: (650) 424-5358 (650) 493-4000 http://www.varian.com Vector Core Laboratory/ National Gene Vector Labs University of Michigan 3560 E MSRB II 1150 West Medical Center Drive Ann Arbor, MI 48109 (734) 936-5843 FAX: (734) 764-3596 Vector Laboratories 30 Ingold Road Burlingame, CA 94010 (800) 227-6666 FAX: (650) 697-0339 (650) 697-3600 http://www.vectorlabs.com Vedco 2121 S.E. Bush Road St. Joseph, MO 64504 (888) 708-3326 FAX: (816) 238-1837 (816) 238-8840 http://database.vedco.com Ventana Medical Systems 3865 North Business Center Drive Tucson, AZ 85705 (800) 227-2155 FAX: (520) 887-2558 (520) 887-2155 http://www.ventanamed.com Verity Software House P.O. Box 247 45A Augusta Road Topsham, ME 04086 (207) 729-6767 FAX: (207) 729-5443 http://www.vsh.com Vernitron See Sensor Systems LLC Vertex Pharmaceuticals 130 Waverly Street Cambridge, MA 02139 (617) 577-6000 FAX: (617) 577-6680 http://www.vpharm.com Vetamac Route 7, Box 208 Frankfort, IN 46041 (317) 379-3621 Vet Drug Unit 8 Lakeside Industrial Estate Colnbrook, Slough SL3 0ED, UK Vetus Animal Health See Burns Veterinary Supply
Suppliers
32 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
Viamed 15 Station Road Cross Hills, Keighley W. Yorkshire BD20 7DT, UK (44) 1-535-634-542 FAX: (44) 1-535-635-582 http://www.viamed.co.uk Vical 9373 Town Center Drive, Suite 100 San Diego, CA 92121 (858) 646-1100 FAX: (858) 646-1150 http://www.vical.com Victor Medical 2349 North Watney Way, Suite D Fairfield, CA 94533 (800) 888-8908 FAX: (707) 425-6459 (707) 425-0294 Virion Systems 9610 Medical Center Drive, Suite 100 Rockville, MD 20850 (301) 309-1844 FAX: (301) 309-0471 http://www.radix.net/∼virion VirTis Company 815 Route 208 Gardiner, NY 12525 (800) 765-6198 FAX: (914) 255-5338 (914) 255-5000 http://www.virtis.com Visible Genetics 700 Bay Street, Suite 1000 Toronto, Ontario M5G 1Z6, Canada (888) 463-6844 (416) 813-3272 http://www.visgen.com Vitrocom 8 Morris Avenue Mountain Lakes, NJ 07046 (973) 402-1443 FAX: (973) 402-1445 VTI 7650 W. 26th Avenue Hialeah, FL 33106 (305) 828-4700 FAX: (305) 828-0299 http://www.vticorp.com VWR Scientific Products 200 Center Square Road Bridgeport, NJ 08014 (800) 932-5000 FAX: (609) 467-5499 (609) 467-2600 http://www.vwrsp.com Vydac 17434 Mojave Street P.O. Box 867 Hesperia, CA 92345 (800) 247-0924 FAX: (760) 244-1984 (760) 244-6107 http://www.vydac.com Vysis 3100 Woodcreek Drive Downers Grove, IL 60515 (800) 553-7042 FAX: (630) 271-7138 (630) 271-7000 http://www.vysis.com
W&H Dentalwerk Burmoos ¨ P.O. Box 1 A-5111 B¨urmoos, Austria (43) 6274-6236-0 FAX: (43) 6274-6236-55 http://www.wnhdent.com Wako BioProducts See Wako Chemicals USA Wako Chemicals USA 1600 Bellwood Road Richmond, VA 23237 (800) 992-9256 FAX: (804) 271-7791 (804) 271-7677 http://www.wakousa.com
WaterStone Technology 12202 Hancock Street Carmel, IN 46032 (317) 644-0862 FAX: (317) 569-8068 www.waterstonetech.com Watlow 12001 Lackland Road St. Louis, MO 63146 (314) 426-7431 FAX: (314) 447-8770 http://www.watlow.com Watson-Marlow 220 Ballardvale Street Wilmington, MA 01887 (978) 658-6168 FAX: (978) 988 0828 http://www.watson-marlow.co.uk
Wako Pure Chemicals 1-2, Doshomachi 3-chome Chuo-ku, Osaka 540-8605, Japan 81-6-6203-3741 FAX: 81-6-6222-1203 http://www.wako-chem.co.jp/egaiyo/ index.htm
Waukesha Fluid Handling 611 Sugar Creek Road Delavan, WI 53115 (800) 252-5200 FAX: (800) 252-5012 (414) 728-1900 FAX: (414) 728-4608 http://www.waukesha-cb.com
Wallac See Perkin-Elmer
WaveMetrics P.O. Box 2088 Lake Oswego, OR 97035 (503) 620-3001 FAX: (503) 620-6754 http://www.wavemetrics.com
Wallac A Division of Perkin-Elmer 3985 Eastern Road Norton, OH 44203 (800) 321-9632 FAX: (330) 825-8520 (330) 825-4525 http://www.wallac.com Waring Products 283 Main Street New Hartford, CT 06057 (800) 348-7195 FAX: (860) 738-9203 (860) 379-0731 http://www.waringproducts.com Warner Instrument 1141 Dixwell Avenue Hamden, CT 06514 (800) 599-4203 FAX: (203) 776-1278 (203) 776-0664 http://www.warnerinstrument.com Warner-Lambert Parke-Davis 201 Tabor Road Morris Plains, NJ 07950 (973) 540-2000 FAX: (973) 540-3761 http://www.warner-lambert.com
Weather Measure P.O. Box 41257 Sacramento, CA 95641 (916) 481-7565 Weber Scientific 2732 Kuser Road Hamilton, NJ 08691 (800) FAT-TEST FAX: (609) 584-8388 (609) 584-7677 http://www.weberscientific.com Weck, Edward & Company 1 Weck Drive Research Triangle Park, NC 27709 (919) 544-8000 Wellcome Diagnostics See Burroughs Wellcome Wellington Laboratories 398 Laird Road, Guelph Ontario, Canada, N1G 3X7 (800) 578-6985 FAX: (519) 822-2849 http://www.well-labs.com
Wheaton Science Products 1501 North 10th Street Millville, NJ 08332 (800) 225-1437 FAX: (800) 368-3108 (856) 825-1100 FAX: (856) 825-1368 http://www.algroupwheaton.com Whittaker Bioproducts See BioWhittaker Wild Heerbrugg Juerg Dedual Gaebrisstrasse 8 CH 9056 Gais, Switzerland (41) 71-793-2723 FAX: (41) 71-726-5957 http://www.homepage.swissonline.net/ dedual/wild heerbrugg Willy A. Bachofen AG Maschinenfabrik Utengasse 15/17 CH4005 Basel, Switzerland (41) 61-681-5151 FAX: (41) 61-681-5058 http://www.wab.ch Winthrop See Sterling Winthrop Wolfram Research 100 Trade Center Drive Champaign, IL 61820 (800) 965-3726 FAX: (217) 398-0747 (217) 398-0700 http://www.wolfram.com World Health Organization Microbiology and Immunology Support 20 Avenue Appia 1211 Geneva 27, Switzerland (41-22) 791-2602 FAX: (41-22) 791-0746 http://www.who.org World Precision Instruments 175 Sarasota Center Boulevard International Trade Center Sarasota, FL 34240 (941) 371-1003 FAX: (941) 377-5428 http://www.wpiinc.com Worthington Biochemical Halls Mill Road Freehold, NJ 07728 (800) 445-9603 FAX: (800) 368-3108 (732) 462-3838 FAX: (732) 308-4453 http://www.worthington-biochem.com WPI See World Precision Instruments
Washington University Machine Shop 615 South Taylor St. Louis, MO 63310 (314) 362-6186 FAX: (314) 362-6184
Wesbart Engineering Daux Road Billingshurst, West Sussex RH14 9EZ, UK (44) 1-403-782738 FAX: (44) 1-403-784180 http://www.wesbart.co.uk
Waters Chromatography 34 Maple Street Milford, MA 01757 (800) 252-HPLC FAX: (508) 478-1990 (508) 478-2000 http://www.waters.com
Whatman 9 Bridewell Place Clifton, NJ 07014 (800) 631-7290 FAX: (973) 773-3991 (973) 773-5800 http://www.whatman.com
Wyeth-Ayerst Laboratories P.O. Box 1773 Paoli, PA 19301 (800) 666-7248 FAX: (610) 889-9669 (610) 644-8000 http://www.ahp.com
Wyeth-Ayerst 2 Esterbrook Lane Cherry Hill, NJ 08003 (800) 568-9938 FAX: (858) 424-8747 (858) 424-3700
Suppliers
33 Current Protocols in Nucleic Acid Chemistry
CPNC Supplement 30
Xenotech 3800 Cambridge Street Kansas City, KS 66103 (913) 588-7930 FAX: (913) 588-7572 http://www.xenotechllc.com Xillix Technologies 300-13775 Commerce Parkway Richmond, British Columbia V6V 2V4 Canada (800) 665-2236 FAX: (604) 278-3356 (604) 278-5000 http://www.xillix.com Xomed Surgical Products 6743 Southpoint Drive N Jacksonville, FL 32216 (800) 874-5797 FAX: (800) 678-3995 (904) 296-9600 FAX: (904) 296-9666 http://www.xomed.com Yakult Honsha 1-19, Higashi-Shinbashi 1-chome Minato-ku Tokyo 105-8660, Japan 81-3-3574-8960
Yamasa Shoyu 23-8 Nihonbashi Kakigaracho 1-chome, Chuoku Tokyo, 103 Japan (81) 3-479 22 0095 FAX: (81) 3-479 22 3435
YSI 1725-1700 Brannum Lane Yellow Springs, OH 45387 (800) 765-9744 FAX: (937) 767-9353 (937) 767-7241 http://www.ysi.com
Zymed Laboratories 458 Carlton Court South San Francisco, CA 94080 (800) 874-4494 FAX: (650) 871-4499 (650) 871-4494 http://www.zymed.com
Yeast Genetic Stock Center See ATCC
Zeneca/CRB See AstraZeneca (800) 327-0125 FAX: (800) 321-4745
Zymo Research 625 W. Katella Avenue, Suite 30 Orange, CA 92867 (888) 882-9682 FAX: (714) 288-9643 (714) 288-9682 http://www.zymor.com
Yellow Spring Instruments See YSI
YMC YMC Karasuma-Gojo Building 284 Daigo-Cho, Karasuma Nisihiirr Gojo-dori Shimogyo-ku Kyoto, 600-8106, Japan (81) 75-342-4567 FAX: (81) 75-342-4568 http://www.ymc.co.jp
Zivic-Miller Laboratories 178 Toll Gate Road Zelienople, PA 16063 (800) 422-LABS FAX: (724) 452-4506 (800) MBM-RATS FAX: (724) 452-5200 http://zivicmiller.com
Zynaxis Cell Science See ChiRex Cauldron
Zymark Zymark Center Hopkinton, MA 01748 (508) 435-9500 FAX: (508) 435-3439 http://www.zymark.com
Suppliers
34 CPNC Supplement 30
Current Protocols in Nucleic Acid Chemistry
